Shellfish Aquaculture and the Environment

Shellfish Aquaculture and the Environment Shellfish Aquaculture and the Environment Edited by Sandra E. Shumway A Joh...

Author: Sandra E. Shumway

279 downloads 1185 Views 218MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

Shellfish Aquaculture and the Environment

Shellfish Aquaculture and the Environment Edited by Sandra E. Shumway

A John Wiley & Sons, Inc., Publication

This edition first published 2011 © 2011by John Wiley & Sons, Inc. Wiley-Blackwell is an imprint of John Wiley & Sons, formed by the merger of Wiley’s global Scientific, Technical and Medical business with Blackwell Publishing. Registered office: John Wiley & Sons Ltd, The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK Editorial offices:

2121 State Avenue, Ames, Iowa 50014-8300, USA The Atrium, Southern Gate, Chichester, West Sussex, PO19 8SQ, UK 9600 Garsington Road, Oxford, OX4 2DQ, UK

For details of our global editorial offices, for customer services and for information about how to apply for permission to reuse the copyright material in this book please see our website at www.wiley.com/ wiley-blackwell. Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Blackwell Publishing, provided that the base fee is paid directly to the Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license by CCC, a separate system of payments has been arranged. The fee codes for users of the Transactional Reporting Service are ISBN-13: 978-0-8138-1413-1/2011. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloging-in-Publication Data Shellfish aquaculture and the environment / edited by Sandra Shumway. p. cm. Includes bibliographical references and index. ISBN 978-0-8138-1413-1 (hardcover : alk. paper) 1. Shellfish culture—Environmental aspects. I. Shumway, Sandra. SH370.S54 2011 639'.4—dc22 2011011547 A catalogue record for this book is available from the British Library. This book is published in the following electronic formats: ePDF 9780470960936; Wiley Online Library 9780470960967; ePub 9780470960943; Mobi 9780470960950 Set in 10 on 12.5 pt Sabon by Toppan Best-set Premedia Limited

Disclaimer The publisher and the author make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation warranties of fitness for a particular purpose. No warranty may be created or extended by sales or promotional materials. The advice and strategies contained herein may not be suitable for every situation. This work is sold with the understanding that the publisher is not engaged in rendering legal, accounting, or other professional services. If professional assistance is required, the services of a competent professional person should be sought. Neither the publisher nor the author shall be liable for damages arising herefrom. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the author or the publisher endorses the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. 1

2011

For my parents, who launched my career at The Hummocks with a clam rake, a skiff, a 9.8 Johnson, and all the freedom, support, encouragement, and common sense a kid could ever need.

Contents List of Contributors Foreword Preface 1

2

Classification of impacts BMPs Assurance labeling Pressures to participate in certification programs Perspectives on ecolabeling Aquaculture certification programs Critique of bivalve shellfish ecolabeling efforts in the United States Criticisms of certification programs Towards more meaningful labeling Concluding remarks Literature cited

The role of shellfish farms in provision of ecosystem goods and services 3 João G. Ferreira, Anthony J.S. Hawkins, and Suzanne B. Bricker Introduction Methods of study Ecosystem goods: biomass production Ecosystem services: environmental quality Literature cited

13

Shellfish aquaculture and the environment: an industry perspective William Dewey, Jonathan P. Davis, and Daniel C. Cheney

33

Introduction Shellfish farmers and harvesters history of water quality protection and stewardship roles BMPs, the shellfish industry, and the role of available research Conclusion Literature cited 3

xi xiii xv

Molluscan shellfish aquaculture and best management practices John A. Hargreaves Introduction Ecosystem change and shellfish aquaculture

3 6

17 26

4

35

51 51 53

65 67 68

70 73 75 77 78

Bivalve filter feeding: variability and limits of the aquaculture biofilter 81 Peter J. Cranford, J. Evan Ward, and Sandra E. Shumway Introduction 81 Constraints on maximum feeding activity 82 Shellfish feeding in nature 85 Emerging knowledge on ecosystem interactions with the bivalve biofilter 109 Conclusions 111 Literature cited 113

33

42 48 48

53 54 64

5

Trophic interactions between phytoplankton and bivalve aquaculture Gary H. Wikfors

125

The interdependence of bivalves and phytoplankton 125 Bivalve population density: farmed bivalves are naturally gregarious 127 vii

viii

Contents

Bivalves as consumers and cultivators of phytoplankton Summary and prospects Acknowledgments Literature cited 6

The application of dynamic modeling to prediction of production carrying capacity in shellfish farming Jon Grant and Ramón Filgueira Physical oceanographic models Filtration and seston depletion Single-box models Higher-order models Fully spatial models Population-based models Local models Optimization Application to management Modeling environmental impact Sustainability and ecosystem-based management Literature cited

7

Bivalve shellfish aquaculture and eutrophication JoAnn M. Burkholder and Sandra E. Shumway Summary Introduction Most commonly reported: localized changes associated with shellfish aquaculture Interpretations from an ecosystem approach Modeling efforts to assess relationships between bivalve aquaculture and eutrophication Eutrophication of coastal waters from land-based nutrients Ecological and economic benefit of bivalve aquaculture in combating eutrophication

127 130 131 131

Conclusions Literature cited 8

Introduction Mussel farming: open landscape feeding in the sea Estimating the environmental value of mussel farming Trading nutrient discharges Agricultural environmental aid program and mussel farming Added ecosystem services through mussel farming The city of lysekil, the first buyer of a nutrient emission quota Swedish mussel farming and its markets Mussel meal instead of fish meal in organic feeds Mussel meal in feeds for organic poultry The use of the mussel remainder as fertilizer and biogas production Risk assessment of mussels for seafood, feed, and fertilizer Conclusions of the Swedish experience Literature cited

135 139 140 140 142 143 145 146 147 148 149 150 151

155

155 156

158 179

187 192

195

Mussel farming as a tool for re-eutrophication of coastal waters: experiences from Sweden Odd Lindahl

9

200 201

217 217 217 219 222 224 226 226 227 229 230

232 233 234 235

Expanding shellfish aquaculture: a review of the ecological services provided by and impacts of native and cultured bivalves in shellfish-dominated ecosystems 239 Loren D. Coen, Brett R. Dumbauld, and Michael L. Judge Introduction Aquaculture-based systems Remaining questions Literature cited

239 249 272 274

Contents ix

10

Bivalves as bioturbators and bioirrigators Joanna Norkko and Sandra E. Shumway Bivalves are key species in soft-sediment habitats What are bioturbation and bioirrigation? How do healthy soft-sediment bivalve populations affect their surroundings? Summary Literature cited

11

12

13

297 298

303 311 312

Environmental impacts related to mechanical harvest of cultured shellfish 319 Kevin D.E. Stokesbury, Edward P. Baker, Bradley P. Harris, and Robert B. Rheault Introduction Literature review Experimental design Conclusions Acknowledgments Literature cited

319 320 329 334 335 335

Genetics of shellfish on a human-dominated planet Dennis Hedgecock

339

Introduction Domestication of shellfish Conservation Conclusions Literature cited

339 341 347 352 352

Shellfish diseases and health management Ralph A. Elston and Susan E. Ford Shellfish health management and infectious disease prevention

Interactions of bivalve shellfish and parasites with the natural environment Interactions of hosts and disease agents within the aquaculture environment Solutions: 1. Shellfish aquaculture development and health management Solutions: 2. Implementing health management for shellfish aquaculture Summary Literature cited

297

14

Marine invaders and bivalve aquaculture: sources, impacts, and consequences Dianna K. Padilla, Michael J. McCann, and Sandra E. Shumway Introduction Introduced shellfish from aquaculture Species moved with aquaculture Introduced species that impact aquaculture Recommendations for minimizing spread and impacts of introductions Future needs Acknowledgments Literature cited

15

359

Balancing economic development and conservation of living marine resources and habitats: the role of resource managers Tessa L. Getchis and Cori M. Rose

359

Introduction Regulatory framework for shellfish aquaculture in the United States

360

367

370

377 385 386

395

395 397 406 407

412 415 415 416

425

425

429

x

Contents

Environmental best management practices (BMPs) Environmental marketing and other incentive programs Conclusions Literature cited 16

17 440 440 442 443

Education Donald Webster

447

Skills Aquaculture-related disciplines K-12 education Undergraduate degree programs Graduate degree programs 4-H and youth programs Extension programs Technology transfer Conclusion Literature cited

447 449 451 452 453 454 455 457 458 459

The implications of global climate change for molluscan aquaculture 461 Edward H. Allison, Marie-Caroline Badjeck, and Kathrin Meinhold Introduction Climate change in the oceans and coastal zones The effects of climate change on shellfish aquaculture systems Adapting shellfish farming to climate change impacts Shellfish aquaculture and climate change mitigation Conclusion Acknowledgments Literature cited

Index

461 462 467 478 482 484 485 485 491

List of Contributors Edward H. Allison The WorldFish Center Penang, Malaysia

Jonathan P. Davis Taylor Shellfish Farms Shelton, WA

Marie-Caroline Badjeck The WorldFish Center Penang, Malaysia

William Dewey Taylor Shellfish Farms Shelton, WA

Edward P. Baker Graduate School of Oceanography University of Rhode Island Narragansett, RI

Brett R. Dumbauld United States Department of Agriculture– Agricultural Research Service Hatfield Marine Science Center Newport, OR

Suzanne B. Bricker National Oceanic and Atmospheric Administration National Ocean Service Silver Springs, MD JoAnn M. Burkholder Center of Applied Aquatic Ecology North Carolina State University Raleigh, NC Daniel C. Cheney Pacific Shellfish Institute Olympia, WA Loren D. Coen 16007 Waterleaf Lane Ft. Myers, FL Peter J. Cranford Fisheries and Oceans Canada Bedford Institute of Oceanography Dartmouth, NS Canada

Ralph A. Elston AquaTechnics, Inc. Sequim, WA João G. Ferreira Institute of Marine Research New University of Lisbon Portugal Ramón Filgueira Department of Oceanography Dalhousie University Canada Susan E. Ford Haskin Shellfish Research Laboratory Rutgers University Port Norris, NJ Jon Grant Department of Oceanography Dalhousie University Canada xi

xii

List of Contributors

Tessa L. Getchis Connecticut Sea Grant University of Connecticut Groton, CT

Dianna K. Padilla Department of Ecology and Evolution Stony Brook University Stony Brook, NY

John Hargreaves Aquaculture Assessments LLC Baton Rouge, LA

Robert B. Rheault East Coast Shellfish Growers Association Wakefield, RI

Bradley P. Harris Department of Fisheries Oceanography School for Marine Science and Technology University of Massachusetts–Dartmouth Fairhaven, MA

Cori M. Rose New England Division U.S. Army Corps of Engineers Concord, MA

Anthony J.S. Hawkins Plymouth Marine Laboratory Plymouth, United Kingdom Dennis Hedgecock Department of Biology University of Southern California Los Angeles, CA Michael L. Judge Department of Biology Manhattan College Riverdale, NY Odd Lindahl The Royal Swedish Academy of Sciences Stockholm, Sweden Michael J. McCann Department of Ecology and Evolution Stony Brook University Stony Brook, NY Kathrin Meinhold The WorldFish Center Penang, Malaysia Joanna Norkko Tvärminne Zoological Station University of Helsinki Finland

Sandra E. Shumway Department of Marine Sciences University of Connecticut Groton, CT Kevin D.E. Stokesbury Department of Fisheries Oceanography School for Marine Science and Technology University of Massachusetts–Dartmouth Fairhaven, MA J. Evan Ward Department of Marine Sciences University of Connecticut Groton, CT Donald Webster Wye Research and Education Center University of Maryland Queenstown, MD Gary H. Wikfors National Oceanic and Atmospheric Administration National Marine Fisheries Service Northeast Fisheries Science Center Milford, CT

Foreword The publication of Shellfish Aquaculture and the Environment could not be more timely. At present, myriad local, state, federal, and private partners are working in the Gulf of Mexico to respond to the largest oil spill in our nation’s history. While the aftereffects of this disaster are not yet fully known, we do know that the environmental and economic ramifications will have significant long-term implications. This event, however, has made Americans nationwide deeply aware of the importance of healthy ecosystems, safe and sustainable fisheries, and the degree to which our economy—and in places like Louisiana and Mississippi, our national culture—depends on our relationship with and management of natural resources. No more so than now is our country aware of the importance of the complex fabric that interweaves our oceans and their ecosystems with the economies in coastal communities and beyond. Shellfish play a central role in our marine ecosystems and coastal communities. Shellfish generate ecosystem benefits including water quality improvements and habitat and species restoration, while shellfish restoration and commercial shellfish aquaculture provide a local food supply and jobs that help to maintain working waterfronts. Conversely, shellfish farming and restoration can have negative environmental impacts, especially if best management practices are not followed. Shellfish Aquaculture and the Environment addresses the environmental implications of shellfish aquaculture. This work began with the 2008 “Symposium on Shellfish and the

Environment” in Warwick, Rhode Island, which brought together some of the finest researchers and policymakers from around the country to address the environmental benefits and challenges associated with shellfish aquaculture. Dr. Sandy Shumway worked with the National Oceanic and Atmospheric Administration’s (NOAA) Aquaculture and Habitat Programs to convene the symposium and invite the speakers. Many of those presentations were precursors to the chapters presented here. As the Editor of the Journal of Shellfish Research for 25 years, research professor of marine sciences at the University of Connecticut, and a past president of the National Shellfisheries Association, Dr. Shumway is uniquely poised to communicate the profound role that shellfish aquaculture can play in supplying a source of safe, healthy, domestically sourced seafood as well as the critical ecological functions that shellfish serve. Dr. Shumway has focused her own research efforts on commercially important shellfish and has collaborated with scientists, communities, nongovernmental organizations, natural resource managers, government officials, and the shellfish aquaculture industry to facilitate public policy and resource management decisions for shellfish culture in the United States. Her commitment stems from her decades-long dedication to examining the interactions between shellfish and the environment. She recognizes that shellfish aquaculture is poised to make a significant contribution to the 37 million tons of seafood needed by 2030 to feed the world’s xiii

xiv

Foreword

population—and do so in an ecologically sustainable manner. Shellfish Aquaculture and the Environment is important on many practicable levels. It serves to inform resource managers and policymakers regarding the best available science on the environmental effects of shellfish aquaculture. It provides insights for managers and policymakers to communicate to scientists the information needed to foster informed decision making. It fosters information exchange that allows scientists to tailor research to answer specific questions to address potential limitations in shellfish aquaculture. The NOAA Aquaculture Program has sought to advance the science of shellfish restoration and commercially viable, environmentally sustainable aquaculture. A robust domestic aquaculture industry promises to make a significant contribution to a safe, local, and healthy seafood supply. In a world in which the United States imports approximately 84% of its seafood—half of which comes from aquaculture—it is incumbent on us as consumers and stewards of the environment to take responsibility for our consumption decisions. Critical at this time especially, shellfish aquaculture serves as an economic engine, securing

jobs in coastal communities, maintaining the spirit and energy of our working waterfronts, and supporting an array of secondary industries. I would like to thank Dr. Shumway for being a visionary and conceiving of this book as well as doing the hard work of organizing, editing, and cajoling to bring it all to fruition. Sandy’s dedication to expanding scientific knowledge has strengthened our basis for making resource management decisions based on sound science. With Sandy’s help and that of so many others with a passion for ocean stewardship and sustainable fisheries, we continue to shine the national spotlight on the role that shellfish aquaculture can and should play in our nation’s seafood supply. Now comes the hard part of translating attention into action through commercial production, habitat restoration, focused research, economic incentives, and planning at the local, state, and federal levels. Shellfish Aquaculture and the Environment will be another valuable tool as for advancing the state and science of shellfish aquaculture. Michael Rubino Manager NOAA Aquaculture Program

Preface Aquaculture is the fastest growing sector of food production globally and has grown almost 10% annually for the past 50+ years. Aquaculture now provides half of the fisheries products consumed globally; 80% of the shellfish are cultured. It has been estimated that by 2050 food production will have to increase by 70% and there is little question that aquaculture—fish and shellfish and algae—will play a major role in that expansion. Only the rate, geographic distribution, and quality remain to be determined. Shellfish aquaculture is poised to contribute substantially to this global need for food production; however, the political and scientific scrutiny is unprecedented. While the bulk of this scrutiny is focused on fish and shrimp culture, all aquaculture, including shellfish, is being watched very carefully. Just a decade ago, words such as “sustainable” and “ecosystem services” were foreign. Today, the seafood industry has “gone green” and having a “sustainably certified” label is an indispensable marketing tool. Shellfish culture has, for many years, been unjustifiably grouped by the popular media with fish and other forms of aquaculture. Not only do the techniques differ extensively, but shellfish are primary consumers and thus no feed is used in the process. Shellfish culture, touted as the “green” culture, may have limited negative impacts in isolated and localized situations (usually associated with overstocking in suspended culture), but the bulk of available data demonstrate that, overall, the environmental impacts of shellfish aquaculture are

minimal and most often beneficial. Shellfish aquaculture maintains working waterfronts, creates jobs, provides habitat for other organisms, removes excess nutrients from the water, and provides a multitude of other environmental services. The task remains to change the perception of aquaculture among all stakeholders: researchers, managers, consumers, environmentalists, and policymakers. Sociopolitical issues such as multiuser conflicts, aesthetics, and recreational uses still prevail in many areas. Clearly, introducing large densities of filterfeeding bivalve molluscs to a habitat, be it in suspended or bottom culture, may result in changes in the ecosystem. Changes can include depletion of phytoplankton, zooplankton, and seston, and localized increase in sedimentation rates via biodeposition, which in turn may induce organic enrichment and change sediment geochemistry and benthic community characteristics. The majority of impacts are site- and species-specific, and the detection and assessment of the influence of bivalve farming on the surrounding environs is a complex process. Sustainable aquaculture provides a healthy source of protein, and is good for the environment and the economy; however, sustainability means different things to different groups and individuals. All farming and culture activities have environmental and social impacts. It is a complex array of interactions, and while ecosystem management approaches are key to addressing and solving the environmental xv

xvi

Preface

issues, the socioeconomic issues associated with advancing sustainable shellfish aquaculture and embracing it as an environmentally and economically sound form of food production for future generations need and deserve a greater focus and presence. The book consists of 17 chapters covering all aspects of shellfish aquaculture, and there was a concerted effort to engage scientists from other venues as well as those with a background in shellfish biology. All of the authors are experts in their respective fields; many are new to the shellfish arena and their willingness to participate in this project is deeply appreciated. Their participation has added appreciably to the overall substance of the book. Topics covered include the role of shellfish farms in provision of ecosystem goods and services; best management practices; filter feeding; trophic interactions between phytoplankton and bivalve aquaculture; the application of dynamic modeling to prediction of production carrying capacity in shellfish farming; eutrophication; mussel farming as a tool for re-eutrophication of coastal waters; bivalves as bioturbators and bioirrigators; environmental impacts of mechanical harvest of cultured shellfish; genetics; shellfish diseases and health management; marine invaders; economic development and conservation of living marine resources and habitats and the role of resource managers; education; implications of global climate change for molluscan aquaculture; and an industry perspective of future development of shellfish aquaculture. The chapters in this book are not intended to be all-inclusive review papers. They are meant to provide readable and understandable background information on key issues associated with shellfish aquaculture to resource managers and policymakers, to help translate the results of scientific research into sound policy, and ensure the continued growth of sustainable molluscan aquaculture. It is hoped that this book will provide background information necessary on key param-

eters to assist in new sitings and expansion of existing aquaculture operations, habitat management, and potential restoration or enhancement efforts. It will also provide baseline information to aid in the development and evaluation of best management practices critical to responsible environmental stewardship. It will allow policymakers and managers to reach informed and reasonable decisions in a timely fashion as sustainable molluscan aquaculture continues to expand and take its place in the global arena as the need for increased seafood production continues to grow. Shellfish aquaculture provides ecosystem services and a healthy source of protein. There are challenges ahead that require all stakeholders—scientists, managers, policymakers, citizens, and aquaculturists alike—to adopt a holistic, realistic, and integrated view toward assessing and weighing the impacts and benefits associated with shellfish aquaculture, and to make informed decisions regarding acceptable impacts. It is time to take the “big picture” approach. Policymakers and managers need to be acutely aware of the big picture to make informed decisions. The future of aquaculture relies on a balance between research and common sense. Overregulation and unrealistic or unnecessary restraints in the name of caution—often a synonym for “afraid to take a stand”—will only slow a necessary and sustainable process. If shellfish aquaculture is to flourish, all constituents need to work together—the future of molluscan shellfish aquaculture and increased food production depends on it. This book would not have been possible without the financial and intellectual support of many people and agencies. First and foremost, thank you to the National Oceanic and Atmospheric Administration (NOAA) Aquaculture Program, especially Michael Rubino, Kate Naughten, and Brian Fredieu for their unflagging support and guidance in moving shellfish aquaculture to the forefront of recognition and acceptance. Justin Jeffryes

Preface

of Wiley-Blackwell has provided continued support and great patience. Kari Heinonen was an invaluable source of technical expertise and performed endless thankless tasks and the book would not have materialized without her. The book is representative of a group effort and I extend my heartfelt thanks to all the authors for sharing their time and expertise, to the reviewers for their timely input, and to the industry members who have regularly tried to keep me grounded, especially Leroy Creswell, Chris Davis, Joth Davis, Bill Dewey, Robin Downey, Rick Karney, Carter Newell, and Bob Rheault. I hope the final product has done justice to their collective efforts.

xvii

And finally, a special thanks to my furry associates, Gus and Zeus, who made all those long evenings of editing and proofreading a little easier to endure. Sandra E. Shumway Groton, CT This book was prepared by Sandra Shumway under award number NA08OAR4170834 from the National Oceanographic and Atmospheric Administration (NOAA) Marine Aquaculture Program, U.S. Department of Commerce.

Chapter 1

The role of shellfish farms in provision of ecosystem goods and services João G. Ferreira, Anthony J.S. Hawkins, and Suzanne B. Bricker

Introduction What is a farm? Shellfish farms vary widely in type, situation, and size. The type of culture can vary according to species, and even within the same species various approaches may be used, depending on factors such as tradition, environmental conditions, and social acceptance. For instance, mussels are cultivated on rafts in Galicia (Spain), and on longlines in the Adriatic Sea (Fabi et al. 2009). But they are also grown on poles in the intertidal area in both France (bouchot) and China (muli zhuang), or dredged from the bottom in Carlingford Lough (Ireland) and in the Eastern Scheldt (the Netherlands).

It is not unusual to use different culture techniques for the same species at different stages of the growth cycle, or to rear benthic organisms off-bottom, taking advantage of a greater exposure to pelagic primary production, better oxygenation, and predator exclusion. In a similar way, shellfish can be grown in intertidal areas, competing for space with other uses (e.g., geoduck grown in PVC tubes in Puget Sound, USA; oysters on trestles in Dungarvan Harbour, Ireland), or subtidally (e.g., scallop off Zhangzidao Island, northeast China). Cultivation takes place within estuaries, coastal lagoons, and bays (e.g., Figure 1.1), and increasingly in offshore locations, where there are less conflicts with other stakeholders in the coastal zone. In many parts of

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 3

4

Shellfish Aquaculture and the Environment

Figure 1.1 Aquaculture in Sanggou Bay, northeast China. Longlines used for shellfish culture are clearly visible in satellite images.

the world, onshore cultivation is also a reality, as occurs in Guangdong province (China) and elsewhere for razor clams and oysters, frequently in multispecies combinations (e.g., Ferreira et al. 2008a; Zhang et al. 2009; Nobre et al. 2010). The size of farms may vary widely, given various constraints imposed by physical space, environmental conditions (which directly influence production), ecological effects, and social acceptability. An obvious constraint on the viability of a shellfish farm is the natural food supply, which in some areas of the world has a direct relationship to the lease units. For instance, in China, the aquaculture cultivation unit is the Culture Mu (Nunes et al. 2003); in a similar way to the medieval bushel, the actual area of this unit varies among different bays, depending on the typical carrying capacity per unit area of each bay, as exemplified in Table 1.1 for Shandong province.

Table 1.1 Dependency of Chinese lease units on carrying capacity. Bay or system

Unit name

Area (m2)

All land-based agriculture

Mu

666.66 (1/15 ha)

Sanggou Bay

Culture Mu

1600–1800

Jiaozhou Bay

Culture Mu

3000–5000

Laizhou Bay

Culture Mu

5000–8000

For the purposes of this text, a farm is therefore defined as an integrated production unit, typically allocated as a lease, subject to specific pressures with associated impacts (Fig. 1.2). This can be an area of sea bottom where molluscs are grown (e.g., mussel/oyster culture, abalone in pens), off-bottom (but overlying bottom space) such as oyster trestles, or an area of water where rafts or lines are placed (droppers off longlines, Chinese lanterns), or

Role of shellfish farms in the ecosystem

Pre ssu res

SEA

Ponds Fish, shellfish Reduced eutrophication

Nitrogen Phosphorus

LAND

5

Shellfish structures, Fish cages Inshore Conflicts Land use, social issues

Conflicts Wild species, space, social

Imp

acts

Figure 1.2 Aquaculture farms: illustration of pressures, activities, and impacts on the coastal fringe.

ponds fringing coastal areas (razor clams). Whether farms are located on the bottom, offbottom, or as suspended structures, they generally preclude the use of the sea bottom for other human activities, such as fishing or recreation, and raise controversial issues related to multiuser interactions, as discussed in Chapter 9 (in this book). This chapter examines the role of the shellfish farm as a provider of ecosystem goods and services. The focus is on farms located in open estuarine and marine waters, from the intertidal zone to offshore locations. Although this book is aimed at shellfish (sensu bivalve mollusc) aquaculture, it is impossible to address the current state of the art of shellfish farming without the inclusion of integrated multitrophic aquaculture (IMTA), an approach that has been practiced in Southeast Asia for thousands of years, both in ponds and in open systems (Ferreira et al. 2008a), and is currently attracting considerable interest (Neori et al. 2004; Ridler et al. 2007; Paltzata et al. 2008). IMTA combines, in the right proportions, the cultivation of fed aquaculture species (e.g., finfish) with organic extractive aquaculture species (e.g., molluscan shellfish) and inorganic extractive aquaculture species (e.g., seaweeds) for a balanced ecosystem management approach that takes into consideration site specificity, operational limits, and food safety guidelines and regulations (e.g., Neori et al.

2004). The aim of IMTA is to increase longterm sustainability and profitability per cultivation unit (rather than per species in isolation, as is done in monoculture), in which all the cultivation components have economic value, and each has a key role in services and recycling processes of the system. Many of the analyses presented in this chapter are carried out by means of mathematical models, so we begin with a review of methodologies that provide the necessary grounding for the development and application of such models. We then examine some of the available options in the area of predictive modeling, and the remainder of the chapter reviews two main aspects: 1. Ecosystem goods supplied by shellfish farms. The emphasis is on production, and its optimization, including IMTA. 2. Ecosystem services supplied by shellfish farms. The environmental role of shellfish farms extends well beyond the harvest of shellfish per se, and includes interactions such as top-down control of eutrophication symptoms (see Chapters 7 and 8 in this book). Case studies are used throughout to illustrate the practical application of principles and techniques in real-world situations, drawing from examples worldwide, including the

6

Shellfish Aquaculture and the Environment

European Union (EU), North America, and Southeast Asia.

Methods of study Definition of culture practice An accurate description of culture practice is a key factor in the implementation of successful aquaculture models. The parameters of interest may be divided into three groups, which will be examined in turn: 1. Spatial parameters: These include the farm dimensions, positioning (e.g., height above tidal datum for intertidal culture such as oyster trestles), orientation, internal partitioning, and crop rotation. 2. Temporal parameters: Data such as the periods of seeding and harvest, together with the seeding and harvest effort, are critical for accurate simulations of production. 3. Morphological and physiological parameters: The range of weights of seed or spat, the cutoff weight for harvest, mortality rates, and any relevant influences on growth (e.g., fouling) are the final elements of culture practice description that must be considered. Although these data appear, in general terms, easy to acquire and less challenging to interpret and simulate than measures of individual growth or biodeposition, experience shows that accurate validation of culture practice poses a major challenge and can be a significant liability for ecological models of aquaculture (Ferreira et al. 2008b). The main difficulties in obtaining consistent data are due to the following factors: 1. Commercial interests introduce an element of confidentiality that is difficult to overcome.

2. Landings statistics are often inaccurate due to underreporting, and in some cases (Watson and Pauly 2001), overreporting. 3. Farming practice, in the sea as on land, is adapted according to variations in growth, environmental conditions, and market requirements. Corresponding data with respect, for example, to the dimensions of target species, or timings of seed and harvest, are by nature fuzzy. Models that use a deterministic approach do not accommodate this type of information well. In cases where natural spatfall is used (e.g., by means of oyster spat collectors) as opposed to hatchery-purchased seed, there is an additional stochastic component of interannual variability. Because all these data are model forcing functions, they severely constrain simulation outputs. For instance, in Belfast Lough, the lease areas for bottom mussel culture are rotated over a 3-year cycle to allow an annual harvest for animals which grow to maturity over a period of 30 months (Fig. 1.3). Clearly, a failure to account for this will overestimate production, irrespective of the accuracy of the underlying growth models. As a final caveat, the determination of natural mortality (m) poses a particular challenge. In general, this is applied as an average

Harvest Year 4

Harvest Year 3 Seeding Year 1

Seeding Year 2

Seeding Year 3

Harvest Year 5 Figure 1.3 Crop rotation in northern Irish mussel bottom culture (Ferreira et al. 2007b).

Role of shellfish farms in the ecosystem

for the culture period, neglecting the fact that the mortality rate will depend both on the life stage of the organism (e.g., will be higher for small animals and postspawning) and on environmental factors such as temperature, dissolved oxygen (DO), and predation. Furthermore, unless a time step of 1 year is used, an annual mortality of 100% will not reduce the stock to zero if m is simulated by means of a first order decay, since C = Co e − mt ,

(1.1)

where C = stock (number of animals); t = time (year); and m = mortality rate (year−1). A 100% mortality coefficient (m = 1) is equivalent to a mortality factor of e−m, reducing a population of 1000 animals to about 370 over 1 year. For an effective 90% mortality, this corresponds to a coefficient of m = 2.3.

Eutrophication assessment An important consideration for aquaculture operations, particularly when addressing conflicts of use, is the potential environmental impact of shellfish farms (see Fig. 1.2). Of particular interest is the role of shellfish in topdown control of eutrophication symptoms (see Chapter 7 in this book). Assessment of eutrophication is done using the well-tested Assessment of Estuarine Trophic Status (ASSETS) model. The model has three components: (1) Influencing Factors (Pressure)—an evaluation based on the natural processing of a system combined with the level of human related nutrient loads; (2) Eutrophic Condition (EC) (State)—an evaluation based on the combined condition of five symptoms of nutrientrelated water quality problems (chlorophyll a [Chl a] macroalgae, DO, nuisance/toxic

7

algal blooms, losses of submerged aquatic vegetation [SAV]); and (3) Future Outlook (Response)—an evaluation of likely future conditions resulting from changes in nutrient load that are based on projected population and land use change and the success of current or new management measures implemented within the watershed. The three components are assessed separately and then combined to provide a single ASSETS rating (Bricker et al. 1999, 2003, 2007, 2008; Scavia and Bricker 2006; Whitall et al. 2007; Ferreira et al. 2007c; Xiao et al. 2007; Borja et al. 2008). However, for the local/farm-scale assessment, only the EC component is relevant. The assessment of EC includes evaluation of five indicators that are divided into two groups. One group, termed primary symptoms, consists of indicators of the beginnings of eutrophication impacts (Chl a, macroalgae). The other group, secondary symptoms, consists of indicators of more significant nutrient-related degradation (DO, nuisance/ toxic bloom occurrences, seagrass losses). An assessment rating is developed for each indicator based on a combination of characteristics: problem concentrations or conditions (e.g., 90th percentile concentration for Chl a and 10th percentile concentration for DO of annual samples), spatial area, and frequency of occurrence of problem conditions. The method applies the assessments by salinity zone which are combined to give an areaweighted average for each indicator at the system level. The average rating for the primary symptoms and the highest (worst) rating for the secondary symptoms, using a precautionary approach, are combined in a matrix to determine the EC for the system. The Farm Aquaculture Resource Management (FARM) model (Ferreira et al. 2007a) simulates processes at the farm scale (about 100–1000 m in length), considering advective water flow and the corresponding transport of

8

Shellfish Aquaculture and the Environment

relevant water properties, shellfish production, and biodeposition. The water properties include suspended particulate matter (TPM), phytoplankton, organic detritus, and dissolved substances such as ammonia and DO. FARM uses a modified version of ASSETS (Fig. 1.4) to determine the impact of shellfish farms on eutrophication (Ferreira et al. 2007a, 2009a). This impact is estimated by examining the difference in Chl a and DO in waters upstream and downstream of the shellfish farm, assuming primarily a one directional flow, knowing that filter-feeding bivalves reduce water column phytoplankton, represented by Chl a as a proxy (e.g., Ryther et al. 1972, Cloern 1982, Cohen et al. 1984; Shumway et al. 1985). This in turn reduces the potential for development of hypoxia, despite the shellfish respiratory need for DO. The spatial area and frequency of occurrence components of the Chl a and DO indices and the other three ASSETS symptom indicators are not considered in order to keep the method simple and the results clear. Although macroalgae are often observed growing on aquaculture structures which may promote growth and thus be considered a contributor to eutrophication symptoms, macroalgal uptake of nutrients excreted by shellfish may actually limit the local impacts of eutrophication. In some places, aquaculture structures may preclude growth of seagrasses, but at the same time, the bivalve filtration results in clearer water which may promote seagrass growth in surrounding areas. Because of the equivocal nature of interactions between aquaculture operations and macroalgae and seagrasses, as well as the difficulty in measuring up- and downstream differences at the local scale, these indicators are not included. The nuisance/toxic bloom indicator is also not included in the FARM model assessment of eutrophication because, though bivalves may limit these populations as for Chl a, the trigger for bloom occurrences is not known

with certainty and thus, like macroalgae and seagrasses, the results would not necessarily indicate the impact of the aquaculture operation. Upstream concentrations of Chl a and DO are based on daily values linearly interpolated from monthly FARM model input values. Downstream concentrations are determined from daily FARM model outputs from the most downstream section of the farm. Since the analysis is a comparison of water quality conditions in waters flowing into and out of the farm, the standard ASSETS salinity zone spatial framework is not used. Water quality is estimated as the 90th percentile Chl a and 10th percentile DO of daily values for the duration of the model run (typically 2–3 years for harvestable size, dependent upon species). Those values are assigned a rating based on the standard ASSETS thresholds (Bricker et al. 2003). The final Chl a and DO ratings for inflowing and outflowing waters are compared to examine changes that have occurred as waters travel through the farm, and thus the impacts on these eutrophication indicators that can be attributed to the operation of the shellfish farm.

Modeling of individual growth Feeding and metabolism in bivalve shellfish are highly responsive to environmental variables that include temperature, aerial exposure, salinity, DO, current speed, food availability, and food composition, all of which differ both spatially and temporally within near-shore waters (Hawkins and Bayne 1992; Gosling 2003; see also Chapter 4). To achieve integrated simulation of these interrelations at ever-decreasing scales required to help optimize individual growth in space and time, differential equations are normally used to define and integrate physiological responses as component processes in the prediction of

Figure 1.4 Calculation of the eutrophic condition (state) component of ASSETS. The salinity zoning and weighting procedures are simplified in the FARM model (see text).

9

10

Shellfish Aquaculture and the Environment

individual growth, the individual being treated as an input-output system with energy and mass as state variables (e.g., Ross and Nisbet 1990; Brylinski and Sephton 1991; Powell et al. 1992; Raillard et al. 1993; van Haren and Kooijman 1993; Barillé et al. 1997; Campbell and Newell 1998; Grant and Bacher 1998; Scholten and Smaal 1999; Pouvreau et al. 2000, 2006; Solidoro et al. 2000; Ren and Ross 2001, 2005; Hoffmann et al. 2006; Ren and Schiel 2008; Spillman et al. 2008). Stochastic simulations of shellfish growth, for instance by modifying von Bertalanffy’s model to account for seasonal effects of water temperature alone on growth and development (e.g., Melià and Gatto 2005; Griebeler and Seitz 2007), do not afford such resolution. Rapid progress is being made in resolving functional relations between those component processes. For example, responsive adjustments in differential efficiencies and resulting rates of particle retention and ingestion are best related to metabolizable components of the available seston, rather than to the total suspended load; whether measured using chlorophyll as a marker for living organic matter, and/or the remaining detritus that is increasingly recognized as important in the nutrition of bivalve shellfish (Hawkins et al. 2002; Navarro et al. 2009). Perhaps counterintuitively, those adjustments are especially relevant over the lower end of observed natural ranges in food availability, when responses are higher per unit change in food abundance, and which have the highest proportional impacts on net energy balance (Hawkins et al. 1999; Pascoe et al. 2009). When integrated with empirical allometric relations, including interrelated effects of the other main environmental drivers such as temperature and salinity, defined responses enable simulation of how the organic composition and energy content of ingested matter change with tidal, seasonal, or spatial differences in food availability and composition. Dependent

relations predict rates of energy absorption, energy expenditure, and excretion. By these means, it is possible to successfully simulate short-term adjustments in feeding, fecal production, excretion, reproduction, net energy balance, and resulting growth across relevant ranges of natural variability (Hawkins et al. 1999, 2002, unpublished). An alternative approach, based on principles of dynamic energy budget, does not use allometric relations, instead assuming that feeding is proportional to surface area, whereas maintenance costs scale to body volume (Ren and Ross 2005; Pouvreau et al. 2006; Ren and Schiel 2008). An associated advance has been to recognize that common relations between those physiological and developmental processes enable generic model structures that may be calibrated to predict responses across the full environmental range experienced by any given species. Pouvreau et al. (2006) described how a single model successfully simulated growth in Crassostrea gigas reared in one natural and two experimental regimes. Other work has shown how a common set of functional differential equations within the ShellSIM model (www.shellsim.com), run with a minimal set of environmental drivers (temperature, salinity, total particulate matter, particulate organic matter [POM], and Chl a), using a single standard set of parameters for each different species, those parameters having been optimized per species on the basis of calibrations undertaken to date, has effectively (±20%) simulated dynamic responses in growth to natural environmental changes experienced by C. gigas and Mytilus edulis under various culture practices at eight contrasting sites throughout Europe and Asia (Ferreira et al. 2007b, 2008a; Hawkins et al., unpublished). Both ShellSIM and the model of Pouvreau et al. (ibid) have been calibrated and validated at single sites for other species, pending further validation elsewhere (Cardoso et al. 2006) (www.shellsim.com).

Role of shellfish farms in the ecosystem

11

Farm length Width

Current

Current Shellfish

Chl a

Depth 1

Chl a 2

n-1

n

Sections POM

POM Biodeposition

Figure 1.5 Schematic diagram of the FARM model.

Integration and analysis The system and the farm The assessment of the role of aquaculture in coastal ecosystems should be based on a holistic definition of sustainable carrying capacity, integrating physical, production, ecological and social elements as suggested by Inglis et al. (2000) and McKindsey et al. (2006). A corollary of this is that carrying capacity should first be determined at the system scale, prior to scaling down. This allows for an appropriate application of marine spatial planning (Buck et al. 2004), by establishing which zones are available for shellfish cultivation, within the context of multiple water uses of interest to stakeholders within the coastal region.

Farm-scale models To model growth and environmental interactions at the farm scale, a number of key interrelations need to be addressed (Dowd 1997; Prins et al. 1998). First, suspension-feeding shellfish may deplete the water of seston, with associated limitation of shellfish growth (Incze et al.

1981; Cloern 1982; Heasman et al. 1998), and in the process reduce primary and secondary symptoms of eutrophication (Ferreira et al. 2009a; see also Chapters 7 and 8). Second, sedimentation of shellfish feces to the bottom may lead to hypoxic and sulfidic conditions, with negative consequences for benthic communities and processes (Grant et al. 2005); pseudofeces, however, may break up too quickly due to water currents to allow deposition on the bottom (Giles and Pilditch 2004). Third, nitrogen excreted from shellfish or regenerated from fecal deposits may stimulate phytoplankton production, for potential recycling and benefit to shellfish (Dame et al. 1991; Newell 2004). To simulate these interrelations, farm-scale models (Fig. 1.5) typically address hydrodynamics, biogeochemical processes, shellfish growth, and population dynamics. Translation to impacts depends in part on advection, dispersion, and/or settlement. Most commonly, the approach taken at the farm scale is to model relevant water properties such as salinity, temperature, Chl a and detrital organic material to drive shellfish growth for assessment of production, and disposal of dissolved materials such as ammonia and DO, and

Shellfish Aquaculture and the Environment

Eutrophication assessment screening model

Chl a, dissolved oxygen

Mortality

e.g., Food depletion

e.g., ASSETS grade, nutrient trading

Physics and biogeochemistry models

e.g., Chl a, POM

Shellfish individual growth models

Shellfish population dynamics model

Optimization of farm activities

e.g., MPP, VMP

12

Shellfish growth

e.g., Harvestable biomass, APP

Shellfish production screening model

Figure 1.6 Conceptual scheme of the various components of the FARM model. The model core is within the dotted rectangle, the two screening models are external (Ferreira et al. 2007a).

biodeposition of fecal material for evaluation of environmental effects (Fig. 1.6). To improve the spatial resolution of different culture layouts, a farm may be defined as a series of contiguous sections, each seeded with a proportion of fresh cohorts as appropriate. Interactive effects between properties of the water body and cultured shellfish can then be simulated as water passes through the farm. To simulate the biomass production of market-size organisms, each model of shellfish growth is integrated in a population dynamics framework based on a standard conservation equation for the number of individuals within weight classes, using growth simulated by each shellfish model to calculate transitions of individuals between classes, taking into account

user-defined seeding and harvesting regimes, plus mortality (e.g., Bacher et al. 2003; Aure et al. 2007; Ferreira et al. 2007a; Duarte et al. 2008; Spillman et al. 2008). Individual-based configurations may also be used to resolve effects of genetic variation and/or different stressors on population dynamics (Hoffmann et al. 2006; Morales et al. 2006). Just as for models of individual shellfish growth, generic approaches to the modeling of culture practice at the farm scale are now established. For example, the FARM model (Ferreira et al. 2007a) has been validated for four shellfish species reared using bottom (blue mussel, Pacific oyster, Manila clam), longline (Mediterranean mussel), and pole culture (blue mussel) in five systems throughout the EU

Role of shellfish farms in the ecosystem

Ecosystem goods: biomass production Limits to production The marketable cohort From a management perspective, the simulation of production in shellfish aquaculture cannot be addressed solely on the basis of biomass yield, for instance by determining the total carbon produced within a farm. Bacher et al. (1998) defined carrying capacity for shellfish culture as the standing stock at which the annual production of the marketable cohort is maximized. This definition, termed the exploitation carrying capacity by Smaal et al. (1998), does not include environmental effects (ecological carrying capacity) and social concerns. Nevertheless, it emphasizes the production of market-sized animals within a specific time frame, and encapsulates concepts such as food depletion (which would lead to smaller animals) and negative local environmental effects (e.g., oxygen depletion), both of which limit production. Solely from the viewpoint of production carrying capacity, considering identical salinity and temperature patterns, the main constraints

to production of the marketable cohort (Cm), are food supply and stocking density (D). The first term is conditioned by current velocity (V) and food concentration (F) (Eq. 1.2). Cm = f (V , F, D)

(1.2)

An increase in seeding density results in a standard Malthusian curve of diminishing returns as seen in the total physical product (TPP), that is, the potential harvestable biomass.

Harmful Algal Blooms (HABs) Shellfish farms can be affected by externalities associated with climatic (see Chapter 17), oceanographic, and anthropogenic effects. Perhaps the most important influence, due to its regularity and negative consequences for farm profitability and human health, is the worldwide increase in HABs, (Shumway 1990; Hallegraeff 1993) as shown in Figure 1.7 for Chinese coastal waters. While this increase may not be as dramatic in the United States or Europe as in Southeast Asia, one of the major concerns is that some HAB events occur naturally, are not yet well understood, and cannot be predicted. This is the case in both the northeast and northwest coasts of the United States, the northeast Atlantic, and the Irish Sea, as well as the Benguela upwelling, and South and East China Seas. 250 Bohai Sea Yellow Sea

200

East China Sea

Number

(Ferreira et al. 2009a). A significant benefit of this modular approach is the capacity to integrate further with models that assess farmrelated impacts on water quality, nutrient cycling, and benthic processes, as well as marginal analyses of farm production potential and profit maximization, while assessing potential credits (see Chapter 8 in this book) for carbon and nitrogen trading (Dowd 2005; Grant et al. 2005; Ferreira et al. 2007a, 2009a). Many of the examples discussed in the following sections are based on applications of the FARM model, illustrating a means of quantitatively evaluating ecosystem goods and services which result from shellfish farming in coastal environments.

13

150

South China Sea Total

100 50 0 1950

1960

1970

1980

1990

Figure 1.7 Number of red tide occurrences along the Chinese coast (Xiao et al. 2007).

14

Shellfish Aquaculture and the Environment

HABs that co-occur with cyanobacteria in the Baltic have been associated with landoriginated nutrient loading, but elsewhere, offshore events (i.e., those associated with upwelling relaxation) are a major issue for farm production. Detection is based on regular phytoplankton monitoring programs and in some cases operational modeling, and provides shellfish farmers with early warning only on the scale of days, and in few areas. Due to the serious consequences for human health from ingestion of contaminated shellfish, harvest interdiction for prolonged periods (weeks to months) has serious consequences for bivalve production, including direct loss of sales, and lower revenue when harvesting bans are lifted, due to potential changes in the condition (tissue weight/total fresh weight) of the animals.

curves, by means of the application of marginal analysis (Jolly and Clonts 1993),

Profit optimization

The first-order derivative of the production function provides the marginal physical product (MPP). For constant input unit cost Px and output unit price Py, profit will be maximized when the value of the marginal product (VMP) equals Px, VMP may be defined as

The Cobb–Douglas production function (TPP curve) shown in Figure 1.8 (e.g., McCausland et al. 2006) may be represented by Equation 1.3, and used to calculate a further two derived

Y = f ( x1 ,| x2 , x3 ,! xn ) , where

Y = output of harvestable shellfish; x1 = initial stocking density of seed, considered the only variable input; and x2–xn = other inputs, considered to be held constant. Measured production data or simulation models may be used to calculate the average physical product (APP) (Eq. 1.4): APPx1 =

120

TPP

(1.4)

2.5

APP

80

1.5 1.0

60 0.5 Optimization VMP = MPP.Po 90 t VMP = Pi For Pi = 15% Po, MPP = 0.15

20 Stage I

0 0

Stage II 50

MPP

–0.5 Stage III

100

0.0

APP and MPP (no units)

2.0

100 TPP (t TFW)

TPP x1

3.0

140

40

(1.3)

–1.0

150 Seed (t TFW)

Figure 1.8 Example application of marginal analysis using constant forcing (see text for explanation).

Role of shellfish farms in the ecosystem

15

Table 1.2 Application of FARM to different species and culture types in Europe. Landings (t TFW)

Model results (t TFW)

System

Species

Loch Creran, Scotland

Crassostrea gigas (Pacific oyster)

Pertuis Breton, France

Mytilus edulis (blue mussel)

Bay of Piran, Slovenia

Mytilus galloprovincialis (Mediterranean mussel)

200

244.6

+22.3

Chioggia, Italy

Mytilus galloprovincialis (Mediterranean mussel)

660

557.1

−15.6

Ria Formosa, Portugal

Ruditapes philippinarum (Manila clam)

1041

119.3

+14.7

1551 2304

134.4 2322

Difference (%)

−13.3 +0.78

Source: Adapted from Ferreira et al. (2009a). 1 Production data for this system were obtained using the EcoWin2000 ecological model.

VMP = MPP ⋅ Py = Px.

(1.5)

A shellfish farmer should clearly aim to cultivate at a stocking density somewhere between stage 1 (Fig. 1.8), where the first derivative of the TPP curve ≥ 1, and stage 3, where further increases in seed density result in lower harvests, and therefore income reduction. The ideal point on the production function may be determined by means of the VMP, for which both the MPP and financial data with respect to seed and product must be known. Repeated runs of models such as FARM (e.g., Ferreira et al. 2007a, 2009a) may be used to generate the outputs required for marginal analysis. Maximum profit will only occur at the maximum income point if MPP = 0. Under the (reasonable) assumption that Py > 0, MPP will only be zero if Px is zero, that is, if the seed is obtained at no cost. This can occur, as presently observed in the distribution of mussel seed in Ireland, which has the twin effects of (1) reducing the incentive for farmers practicing bottom culture to distribute seed evenly and reduce mortality (Ferreira et al. 2007b) and (2) encouraging overexploitation, resulting in lower APP. Frequently, the APP value approaches unity, the profit resulting only from the differential between sale price and

seed cost since there is no biomass multiple (Ferreira et al. 2009a).

Real-world applications Table 1.2 presents reported harvest yields and simulation results for five different European shellfish farms (Ferreira et al. 2009a), which cultivate (in monoculture) the four major species commercially produced in Europe: blue and Mediterranean mussels, Pacific oyster, and Manila clam. These farms, which range from western Scotland (Loch Creran) to the southern coast of Portugal (Ria Formosa), were studied in the ECASA project (Borja et al. 2009; www.ecasa.org.uk) and represent a range of culture types and habitats including pelagic and benthic deployments in intertidal and subtidal locations within coastal bays and offshore sites. The drivers for the FARM model were obtained from measured data, outputs of system-scale models, or a combination of both. The results (given in total fresh weight) show good agreement with reported annual production, with deviations ranging from −16% to +22%. A profit maximization scenario was tested for each farm (Table 1.3), based on the

16

Shellfish Aquaculture and the Environment

Table 1.3 Comparison of standard model and profit maximization scenarios for the five study sites. Farm location

Species Culture type Farm area (ha) Cultivation period (days) Present setup Seed (t) TPP (t) TPP (t ha−1) APP Harvest profit (k1) Harvest income (k1 year−1) Profit/income (annualized) Profit maximization Seed (t) TPP (t) TPP (t ha−1) APP Harvest profit (k1) Harvest income (k1 year−1) Profit/income (annualized) Profit ratio (scenario/standard)

Loch Creran

Pertuis Breton

Bay of Piran

Chioggia

Ria Formosa

Pacific oyster Trestles 16.5 730

Blue mussel Longlines 200.0 415

Mediterranean Longlines 1.8 490

Mussel Longlines 200.0 308

Manila clam Bottom 11.4 180

41.2 134.4 8.1 3.3 630.7 335.9 0.94

664.0 2322.0 11.6 4.1 3445.0 3076.7 0.98

43.1 244.6 135.9 5.7 184.2 154.9 0.89

660.0 557.1 2.8 0.8 131.1 429.1 0.36

15.3 119.3 10.5 7.8 1,177.0 2,418.2 0.99

322.2 440.2 26.7 1.4 1879.0 1100.5 0.85 3.0

1000.0 3413.0 17.1 3.4 4356.0 3902.0 0.98 1.3

45.5 247.2 137.3 5.4 185.1 156.5 0.88 1.0

396.0 405.8 2.0 1.0 125.2 312.6 0.47 1.0

340.8 909.9 79.8 2.7 8,758.0 18,450.5 0.96 7.4

Source: Adapted from Ferreira et al. (2009a).

marginal analysis approach described above. Three of the farms can potentially increase production to improve their profits, the mussel farm in Slovenia (Piran) appears to be working at optimal capacity, and the Chioggia farm in the Venice area is making less than optimum profit since it incurs excessive production costs for the seed density applied, with respect to cost-benefit optimization. In all the farms except Chioggia, seed is purchased at a very low (in some cases insignificant) cost. The annualized profit : income ratio hardly changes, though the profit itself increases significantly, particularly for Loch Creran and Ria Formosa. In Chioggia, the profit is practically identical, although the seed tonnage is reduced by 40%. The only financial variables in this analysis are the cost of seed and price of product. Other marginal costs of shellfish farming can be

included in this approach by, for example, increasing the seed cost as a proxy for variables such as labor and fuel (Ferreira et al. 2007a). Changes to fixed costs such as lease charges do not influence the decision of a producer on optimal use of the variable input since this is based on the comparison of values of marginal product and marginal input. Multiple input and output variables may also be considered using marginal analysis, or alternative methods may be applied (e.g., Sharma et al. 1999). Production enhancement will be possible through a reduction in food depletion, an inevitable consequence of density increases. One of the ways to achieve this and simultaneously generate significant positive externalities from an environmental perspective is through the use of IMTA.

Role of shellfish farms in the ecosystem

17

Table 1.4 Oyster monoculture and IMTA scenarios in Sanggou Bay. Scenario A

Scenario B

Oysters in monoculture all sections

Oyster and fish IMTA all sections

People TPP (t TFW) APP

7.5 0.22

219.7 6.54

Planet Chlorophyll a (P90) N removal (kg year−1) Population equivalents (PEQ) Organic detritus removal (kg C year−1)

9.4→6.2 356 108 7816

9.4→5.9 2468 748 39,973

Description

ASSETS Profit Income (shellfish; k1 year−1) Gross profit (aquaculture; k1)

4 22.9 4.0

4

4

4

668.4 1065 + 151 = 1080

1

Income due to finfish culture.

Production enhancement using multiple species Zen and the art of polyculture A number of authors (Neori et al. 2004; Reid et al. 2007) have reviewed the benefits of IMTA. From the point of view of production alone, the use of particulate organic waste from finfish culture by filter-feeders, and of dissolved waste from both finfish and filterfeeders by macroalgae, may provide significant yield improvements. Table 1.4 shows FARM model results for a 3.2-ha farm in Sanggou Bay, northeast China (Fig. 1.1), for oysters (ShellSIM individual C. gigas growth model) in monoculture and in combined culture with finfish. Oyster density is 210 ind. m−2, a total of about 6.7 × 106 animals; for the IMTA scenario 15 fish cages are distributed equally throughout the farm, each with 1000 fish. The oysters are able to use both the organic waste from the fish faeces and surplus fish food, and the downstream sections of the farm, which would in monoculture be subject to food depletion, show significantly enhanced production.

Production of the marketable cohort, in this case individuals with a total fresh weight (TFW) > 50 g, increases by two orders of magnitude in IMTA, and the biomass multiple (APP) increases by one order of magnitude. There is an order of magnitude increase in annualized income from shellfish alone, to which revenue from the sale of finfish must be added. This analysis can be extended to include externalities, discussed in the section below on ecosystem services.

Ecosystem services: environmental quality Biodeposition, conservation, and biodiversity The negative externalities of shellfish aquaculture are usually reported as (1) biodeposition; (2) competition with native (wild) species. Although some of these aspects, particularly those related to conservation of wild species (see Chapters 12 and 14 in this book), are best dealt with at the system scale, some brief considerations may be made on farm-scale effects.

18

Shellfish Aquaculture and the Environment

Table 1.5 Sedimentation associated with oyster monoculture in Sanggou Bay (sedimentation through an empty farm, fully stocked farm, and shellfish biodeposition). Parameter

Empty farm

Stocked farm

Biodeposit production

Total (t POC) Annualized (t POC year−1) Annualized per area (kg POC m−2 year−1) Sediment organic enrichment (Δ% POC year−1) Total (t POM) Annualized (t POM year−1) Annualized per area (kg POM m−2 year−1) Sediment accretion (mm year−1)

138.01 83.96 2.62 2.02 363.19 220.94 6.9 2.66

136.62 83.11 2.6 2 359.52 218.71 6.83 2.63

11.88 7.23 0.23 0.17 31.26 19.02 0.59 0.23

POC, particulate organic carbon.

Biodeposition Biodeposition of faecal material from shellfish farms may lead to changes in bottom sediment composition (Chapter 10 in this book) through the increase of organic material (the equivalent of sediment eutrophication) with secondary symptoms of hypoxia or anoxia, resulting in changes to benthic communities. It is widely recognized that effects are much less extreme than for finfish aquaculture, due to the absence of artificial feed (e.g., Giles et al. 2009; Weise et al. 2009), and stem from poor regulation (e.g., inappropriate siting with respect to current speed) and/or poor culture practice (e.g., excessive stocking density). Few effects are reported for bottom culture, where excessive biodeposits would be expected to have a direct effect on the survival of the farm itself, and the ecosystem engineering capabilities of mussels and oysters may enhance epifaunal diversity (Commito et al. 2008; see Chapter 9 in this book). Under appropriate conditions for suspended culture, such as the increasing use of offshore sites, few effects on native macrobenthos can be observed (Dumbauld et al. 2009; Fabi et al. 2009). Bivalve filter feeding is a net removal of particulate organic material that naturally exists in the water column. Therefore, organic enrichment of the environment will at worst be localized since it is clear from a simple mass

balance that a shellfish farm by definition reduces POM, converting it into harvestable live biomass. Problems will only occur through the differential settling speed of biodeposits, which form larger aggregates than the source particles. Table 1.5 shows a modeling analysis of biodeposition for oyster longline monoculture carried out by means of the FARM model. No vertical turbulence factor is applied, which would act to reduce sedimentation, restricting it to periods of low current speed, and no sediment erosion or diagenesis is considered. All sedimented material is considered to remain under the farm area, the worst possible scenario both in terms of accretion and organic enrichment. The left column in Table 1.5 shows results for sedimentation over the culture period for all POM (algae and detritus) transported across an unstocked farm. The middle column adds the biodeposits of cultivated animals to the POM in the water column. The net effect of adding cultivated shellfish is to slightly reduce sediment organic enrichment and accretion rate, although the settling speed used for the biodeposits is at the low end of the range reported by Weise et al. (2009), which for the blue mussel M. edulis varies between 0.1 and 1.8 cm s−1. The right column shows biodeposit production by the cultivated oysters. Even assuming

Role of shellfish farms in the ecosystem

all such biodeposits actually fall to the bottom within the same farm, with no removal due to horizontal advection and dispersion, the resulting accretion rate of 0.23 mm year−1 is very low, corresponding to a POC enrichment factor of only 0.17% year−1. The deposition of shells below suspended culture structures such as mussel droppers as a consequence of natural mortality is often considered a negative environmental effect of shellfish aquaculture. Empty shells have a variety of uses, as illustrated in Figure 1.9 for a hatchery in northeast China. CaCO3 may additionally be used as a source of minerals in agroindustry (see Chapter 8 in this book). Voluntary improvements to culture practice techniques and better regulation are appropriate instruments for mitigation of the potential environmental impact of shell debris from farms.

19

Wild species Finally, a brief note should be made about interactions with native species. While resource partitioning effects are best examined at the ecosystem level (Sequeira et al. 2008), GISbased marine spatial planning is appropriate for analysis of habitat preservation. Models such as FARM incorporate algorithms for analysis of biodeposition effects, and for reducing the food supply to cultivated shellfish as a function both of the natural distribution of benthic wild species in the farm area and their characteristic filtration rates. Simulations for intertidal trestle culture of C. gigas in Dungarvan Harbour (Ferreira et al. 2009b) showed no significant effects of wild species filtration on oyster growth. Nevertheless, modeling of such effects is important for: 1. Improving the accuracy of farm-scale models by partitioning the available resource 2. Determining the baseline food requirements for natural benthic populations prior to licensing shellfish farms 3. Establishing an upper limit to stocking to help ensure adequate food supply and habitat requirements for wild species.

Integrated catchment management A shellfish farm, like any other assemblage of filter-feeders, removes phytoplankton and organic detritus from the water column (Chapter 5 in this book). In doing so, it provides a key ecosystem service by reducing primary symptoms of eutrophication (Bricker et al. 2003; Xiao et al. 2007; see also Chapter 7 in this book). This reduction has two major consequences:

Figure 1.9 Shells of the Chinese scallop Chlamys farreri used as spat collectors. (Photo courtesy of Dr. Q. F. Gao, Ocean University of China.)

1. It alters the underwater light climate, enabling autotrophic production to occur at greater depths, and potentially enabling the recovery of SAV (e.g., Zostera sp.,

20

Shellfish Aquaculture and the Environment

Posidonia sp.) and long-lived macroalgae (e.g., Laminaria sp.). SAV provides further ecosystem services as a refuge and nursery for juvenile fish, as well as increased sediment stability (Yamamuro et al. 2006). 2. It shortens the cycling of suspended organic matter by removing the opportunity for bacterial mineralization, and therefore the onset of secondary eutrophication symptoms such as hypoxia or anoxia. This top-down control can be an important complement to land-based nutrient removal. Phytoplankton, whether in fringing ponds or coastal and estuarine water, acts as a catchment loading filter by removing the causative factors of eutrophication, that is, nitrogen and phosphorus. In turn, shellfish farms remove the primary eutrophication symptom (Fig. 1.2).

The duality of food depletion As cultivation density is increased, the law of diminishing returns leads to lower growth of harvestable animals. In the example shown in Table 1.4, if all oysters above 5 g TFW were collected, the overall harvest would increase from 7.5 t TFW to 95 t TFW. Clearly, from a production perspective, in this example, oyster monoculture is inefficient due to food depletion. Oysters, however, perform an environmental role of bioremediation, as evidenced

from phytoplankton depletion, practically identical for monoculture and IMTA (Table 1.4).

Eutrophication assessment Although the role of shellfish farms in reducing eutrophication symptoms is clear, it is helpful to apply a well-tested methodology such as the ASSETS model to translate quantitative concentrations into qualitative indices. Because this simplified application focuses on the eutrophication status at the inflow and outflow points, and is therefore a differential or spatially comparative approach, the role of the farm with respect to the typical water quality at the site becomes clear. Additionally, the use of a percentile-based approach increases confidence in the comparison since the natural variability of many water quality parameters can make it difficult to distinguish a trend signal. A meaningful comparison among farms becomes possible not only at the production level but also with respect to environmental services. In oligotrophic systems, the ASSETS results may suggest that too much POM is being removed, with respect to the supply required to maintain the natural background of wild species. Table 1.6 (Ferreira et al. 2009a) represents the ASSETS color grades (corresponding to the EU Water Framework Directive scale:

Table 1.6 ASSETS results obtained for the five farms in Europe (Ferreira et al. 2009a). System

Percentile 90 Chl a (mg L−1)

Percentile 10 O2 (mg L−1)

ASSETS score

−0.1

0.0

High

0.5

0.0

Good

Bay of Piran

−4.3

−0.1

Good

Chioggia

−0.2

−0.1

High

Ria Formosa

−0.1

−0.1

High

Loch Creran Pertuis Breton

Role of shellfish farms in the ecosystem

21

Table 1.7 Mass balance for modeled individual growth of the Pacific oyster Crassostrea gigas. Variable

Value (units: see left column)

Nitrogen (gN)

Net biomass production (g TFW) Clearance (m3) Phytoplankton removal (mg Chl m−3) Detrital POM removal (g POM m−3) POM removal (g POM m−3) Spawning losses (g POM m−3) POM biodeposition (g POM m−3) Ammonia excretion (mM m−3) Total N removal (model) Percentage of net biomass production

101.44 23.14 83.61 71.98 75.28 0.17 43.35 3.82

1.011 — 0.2 4.25 4.45 0.01 2.56 0.05 1.82 1.8%

1

Calculated as 1% of biomass production, after Lindahl et al. (2005).

blue—high; green—good; yellow—moderate; orange—poor; red—bad). The score for the symptom in the inflowing water is shown on the left, outflow on the right. The concentration changes reflective of eutrophication symptoms are shown in blue (better or neutral) or red (worse). In these five real-world examples, the effect of shellfish farming on the ASSETS eutrophication score only results in a status change in Piran, Slovenia. In general terms, using the Chl a and DO categories reported in Bricker et al. (2003), and the synthesis score for EC derived from them (Ferreira et al. 2007a), a management proposal might be to site shellfish farms in areas where the ASSETS score would fall into the moderate or good category, and where the farm might change that score to good (or the low end of high). Licensing of farms in areas where the ASSETS score is already high must be carefully considered since an excessive cultivation density might potentially create undesirable food depletion effects. These would, in any case, reflect on the production success of the farm since the food scarcity would lead to low harvests. This can be seen for Piran and Chioggia, where the APP is 5.7 and 0.8, respectively (Table 1.3), which would be expected from the ASSETS scores shown for the two systems in Table 1.6.

A final note from these examples is that, even at sites where the cultivation density is high, the effects on DO concentration are negligible, reinforcing the positive value of environmental externalities of shellfish culture with respect to eutrophication.

Trading and valuation of nutrient credits Table 1.7 shows example mass balance outputs during growth of a single animal, using the ShellSIM model for Pacific oyster. Models (i.e., FARM) extrapolate such budgets to the farm scale, as illustrated in Figure 1.10 for Ruditapes sp. in southern Portugal (Ferreira et al. 2009a). In the Ria Formosa, clam growth is determined mainly by detrital POM, which is a reasonable expectation for a system with a short water residence time (Nobre et al. 2005) in which an autochtonous phytoplankton bloom is unable to develop (Ferreira et al. 2005). In this particular case, a rather high mortality is involved, given that the nutrient loading to the area results in eutrophication symptoms expressed as overgrowth of opportunistic seaweeds such as Enteromorpha (Fig. 1.11), which can smother benthic macrofauna. Nevertheless, about 60% of the nitrogen removed from the system by filtration is

Shellfish Aquaculture and the Environment

Shellfish Filtration

22

Phytoplankton removal 3457 kg C year–1

Detritus removal 321,271 kg C year–1

N removal (kg year–1) Population equivalents 8748 PEQ year–1

Assets Chl a O2 Score

Algae Detritus Excretion Feces Mortality

–538 –49975 142 21405 100

Mass balance

–28867

Annual income

Parameters

Shellfish farming: Nutrient treatment:

2418.2 k€ year–1 262.4 k€ year–1

Density of 90 clams m–2 180-day cultivation period 66% mortality

Total income:

2680.6 k€ year–1

3.3 kg N year–1 PEQ

Figure 1.10 Mass balance and nutrient emissions trading for clam aquaculture in Ria Formosa, southern Portugal (Ferreira et al. 2009a).

Figure 1.11 Clam culture area in the Ria Formosa. (Courtesy of J. Dilão.)

retained by the clams; a proportion of these animals is of harvestable size, and therefore is physically taken from the farm. It is possible to estimate the environmental value of this service by comparing it with the

comparable cost of land-based treatment; in this Ria Formosa farm, shellfish filtration provides a gross removal of about 325 t C year−1, of which about 1% is phytoplankton. This equates to the emissions of 8748 population

Role of shellfish farms in the ecosystem

equivalents (PEQ) and a net annual nitrogen removal of 29 t year−1. Based on substitution costs, this ecosystem service is valued at 0.26 M1 year−1, about 10% of the direct income from shellfish culture. Nitrogen credit trading at the watershed scale (e.g., USEPA 2004) is now a reality in parts of the United States. In Connecticut, the Nitrogen Credit Exchange (NCE) has been applied since 2002 for improved management of Long Island Sound, with over US$30 million in economic activity in the first 4 years of trading (Stacey, pers. comm.). The dollar value per credit has increased from $1.65 (2002) to $4.36 (2007), an annualized growth of 33%, substantially outperforming both the Dow Jones and the NASDAQ indices. Nitrogen credit trading has more local appeal than carbon emissions trading since the cause and effect may be observed at the watershed scale. Shellfish farmers have an opportunity to participate in the nitrogen trading market through the sale of credits to other stakeholders, such as agriculture. This may be of particular social relevance in remote areas of Europe, the United States, and Canada, where agriculture is required by law to reduce the application of fertilizer, but may become economically uncompetitive, leading to desertification of rural areas (Ferreira et al. 2007a–d).

Caveats The present analysis with respect to environmental quality, both in terms of shellfish products and the water body in which they are farmed, does not address issues such as disease, microbiological contamination, xenobiotics, or bioaccumulation. In general terms, for this type of remediation model to work together with viable market production, a careful control of other types of pollutants, frequently discharged to coastal waters concurrently with nitrogen and phosphorus, must be taken into account.

23

Benefits of multitrophic farming Apart from production enhancement, the additional environmental benefits of IMTA are illustrated in Table 1.4 for Sanggou Bay under the “Planet” section. For monoculture, the annualized net nitrogen removal is over 350 kg for one farm alone, corresponding to the emissions of 108 PEQ. Shellfish filtration in oyster monoculture for this farm in China provides a gross removal of about 11 t C year−1, of which about 30% is phytoplankton, corresponding to a net nitrogen removal valued at 3.2 k1 year−1, about 15% of the direct income from shellfish culture. For IMTA, the total nitrogen removal increases sevenfold, to about 2500 kg N year−1, that is, a positive externality valued at 22.4 k1 year−1. However, since the addition of fish cages provides a significant source of food to the shellfish, derived from uneaten fish ration and fecal matter, the direct value of goods (i.e., harvestable shellfish) produced is about 30 times greater than the nitrogen removal value. In this example, the shellfish are reducing the negative externalities of finfish aquaculture, which would otherwise represent an environmental cost manifested through anoxic sediment conditions and mortality of benthic organisms beneath the cage areas. A sensitivity analysis performed in FARM is given in Table 1.8, considering different particle diameters (and thus settling speeds) for biodeposits. The analysis was carried out only for IMTA since the harvest yield in monoculture is already very low and does not change much with increasing rates of biodeposition. As before, this analysis considers a worstcase scenario, with no vertical turbulence (which acts to reduce particle sinking) and no sediment erosion or mineralization. Although the calculation algorithm is precautionary, the trend, as expected, is for an exponential decrease in production as the biodeposit particle size increases. This reduction reflects an inability to use biodeposits before they sink

24

Shellfish Aquaculture and the Environment

Table 1.8 Sensitivity analysis: IMTA production, nitrogen removal, and biodeposition.

Biodeposit diameter (mm)

Harvestable biomass (t TFW)

Nitrogen removal (kg year−1)

Total income (shellfish sale + nutrient removal) (k1 year−1)

Sediment accretion (mm year−1)

Sediment organic enrichment (Δ% POC year−1)

0.0186 0.0221 0.0263 0.0312 0.0372 0.0442

187.8 144.7 93.0 45.6 19.4 10.6

2188 1851 1473 1091 772 556

590.8 456.9 296.2 148.6 65.9 37.4

5.64 6.60 7.70 8.89 10.10 11.25

4.29 5.02 5.85 6.75 7.67 8.55

below the farm area, and leads to an increase of sediment effects. The natural accretion rate for the same drivers without any aquaculture, considering a POM particle diameter of 15.6 μm, is 2.66 mm year−1, and the sediment organic enrichment is 1.66 Δ% POC year−1. The inclusion of the effects of biodeposit resuspension and diagenesis in the simulation may result in a reduction of over 60% in accretion rate (Giles et al. 2009), although the erosion component will result in a more widely dispersed farm biodeposit footprint. IMTA set out in vertical layers, as occurs in Sanggou Bay, can optimize particle use, taking advantage of oyster droppers which continue well below the finfish cages, thus profiting both from a vertical food supply from the fish waste and a horizontal one due to advective transport of algae and detrital matter. Maximization of environmental benefits of IMTA must therefore consider a combination of appropriate species (ideally including macroalgae for dealing with dissolved waste), densities and positioning, in order to progress toward integrated systems with very low nutrient emissions.

Scaling Tools applied to assess the role of shellfish farms in the provision of ecosystem goods and services can help to understand the global role

of shellfish farming in the marine environment at scales of (1) the system, (2) economic blocks, and (3) the world as a whole. Examples for each of these will be briefly discussed. At the system scale, the determination of overall production may be obtained through landings data, which in effect corresponds to integrating the harvest declared by each individual farm. This can also be carried out using system scale models (e.g., Ferreira et al. 2008b), or by the application of models such as FARM to a subset of typical farms. In situations where shellfish farming was once an important activity, it may be useful to repopulate those systems with virtual shellfish farms, back-calculating densities and areas from historical data. In Chesapeake Bay, this was done by means of an ecosystem-scale model (Ferreira et al. 2007d), where production in historical oyster bars (Fig. 1.12) was simulated, using C. gigas as a proxy for Crassostrea virginica. Over the simulation period, a harvestable biomass of 15.8 × 106 bushels was obtained, with a combined effect on environmental quality resulting in a reduction in Chl a 90th percentile of about 30%, from 16 to 11 μg L−1, and a net removal of 26,600 t N year−1, a population equivalent of 8 × 106. The five EU farms simulated in Ferreira et al. (2009a) collectively represent the main species and culture practices in Europe, which allowed an indicative budget calculation for

Role of shellfish farms in the ecosystem

25

Figure 1.12 Chesapeake Bay historical oyster bars (only Maryland areas shown).

European shellfish aquaculture. A total production of 1051 × 103 t year−1 (FAO 1999) was estimated for the major cultivated bivalves. Of these, 70% are mussels (54% blue mussel and 16% Mediterranean mussel), 23% are oysters, and 7% are clams. Production and nutrient removal data were used to calculate the role of EU shellfish farms in removing nutrients, which corresponds to a removal of over 55,000 tons of nitrogen per year, that is, a population equivalent of 17 million people, or the population of the Netherlands. The substitution value for landbased nutrient removal is estimated to be 0.4 billion 1 y−1.

Similar estimates may be carried out based on the worldwide reported shellfish aquaculture landings (FAO 2009), modeling results of nitrogen removal for a typical range of cultivated species. Figure 1.13 shows some results from this analysis: the present consumption of shellfish corresponds to a per capita equivalent of one mussel per day, and molluscan aquaculture removes the equivalent of 3% of the waste nitrogen produced by the population of the world, a net uptake of slightly over 660,000 t year−1. This uptake, which takes place in the most sensitive area of the world ocean, that is, the

26

Shellfish Aquaculture and the Environment

3.3 kg N year–1

1.8 kg N year–1 Figure 1.13 The role of cultivated shellfish in the world nitrogen budget.

coastal fringe, plays a significant role in reducing eutrophication and improving the quality of life for communities inhabiting the coastal zone. At present, the net removal of primary symptoms of eutrophication already corresponds to an ecosystem service valued at 7.5 billion 1, or 3% of the GDP of Portugal. Since the 1950s, world aquaculture (including that of molluscan shellfish) has been expanding almost seven times faster than the world population (FAO 2009). This suggests that, together with increasing the world food production and providing jobs in coastal communities, particularly as wild fisheries contract, the relevance of cultured shellfish in mitigating the potential consequences of nutrient loading to the coastal zone will increase. In summary, shellfish farms provide a set of valuable ecosystem goods and services, which may be quantified using tools such as the

models applied herein. As shellfish farming develops both in semi-enclosed systems and offshore, and as IMTA becomes a reality for many coastal farmers, the importance of such models to assess sustainability and trade-offs in the context of marine spatial planning will increase. The models themselves will become increasingly realistic as the research that underpins them sheds new light on the physiology of cultivated species, interactions within the “managed” trophic web, and relevant ecosystem processes.

Literature cited Aure, J., Strohmeier, T., and Strand, Ø. 2007. Modelling current speed and carrying capacity in longline blue mussel (Mytilus edulis) farms. Aquaculture Research 38:304–312.

Role of shellfish farms in the ecosystem

Bacher, C., Duarte, P., Ferreira, J.G., Héral, M., and Raillard, O. 1998. Assessment and comparison of the Marennes-Oléron Bay (France) and Carlingford Lough (Ireland) carrying capacity with ecosystem models. Aquatic Ecology 31(4):379–394. Bacher, C., Grant, J., Hawkins, A.J.S., Fang, J., Zhu, M., and Besnard, M. 2003. Modelling the effect of food depletion on scallop growth in Sungo Bay (China). Aquatic Living Resources 16:10–24. Barillé, L., Héral, M., and Barillé-Boyer, A.-L. 1997. Modélisation de l’écophysiologie de l’huître Crassostrea gigas dans un environnement estuarien. Aquatic Living Resources 10:31–48. Borja, A., Bricker, S.B., Dauer, D.M., Demetriades, N.T., Ferreira, J.G., Forbes, A.T., Hutchings, P., Jia, X., I, Kenchington, R., Marques, J.C., and Zhu, C. 2008. Overview of integrative tools and methods in assessing ecological integrity in estuarine and coastal systems worldwide. Marine Pollution Bulletin 56:1519– 1537. Borja, A., Germán Rodríguez, J., Black, K., Bodoy, A., Emblow, C., Fernandes, T.F., Forte, J., Karakassis, I., Muxika, I., Nickell, T.D., Papageorgiou, N., Pranovi, F., Sevastou, K., Tomassetti, P., and Angel, D. 2009. Assessing the suitability of a range of benthic indices in the evaluation of environmental impact of fin and shellfish aquaculture located in sites across Europe. Aquaculture 293(3–4):231–240. Bricker, S.B., Clement, C.G., Pirhalla, D.E., Orlando, S.P., and Farrow, D.R.G. 1999. National Estuarine Eutrophication Assessment. Effects of Nutrient Enrichment in the Nation’s Estuaries. NOAA, National Ocean Service, Special Projects Office and National Centers for Coastal Ocean Science, Silver Spring, MD. specialprojects.nos.noaa.gov/projects/cads/nees/ Eutro_Report.pdf Bricker, S.B., Ferreira, J.G., and Simas, T. 2003. An Integrated methodology for assessment of estuarine trophic status. Ecological Modelling 169:39–60. Bricker, S., Longstaff, B., Dennison, W., Jones, A., Boicourt, K., Wicks, C., and Woerner, J. 2007. Effects of Nutrient Enrichment in the Nation’s Estuaries: A Decade of Change, National Estuarine Eutrophication Assessment Update.

27

NOAA Coastal Ocean Program Decision Analysis Series No. 26. National Centers for Coastal Ocean Science, Silver Spring, MD. ccma.nos. noaa.gov/news/feature/Eutroupdate.html Bricker, S.B., Longstaff, B., Dennison, W., Jones, A., Boicourt, K., Wicks, C., and Woerner, J. 2008. Effects of nutrient enrichment in the nation’s estuaries: a decade of change. Special issue of Harmful Algae 8:21–32. Brylinski, M., and Sephton, T.W. 1991. Development of a computer simulation model of a cultured blue mussel (Mytilus edulis) population. Canadian Technical Report of Fisheries and Aquatic Sciences 1805:viii + 81. Buck, B.H., Krause, G., and Rosenthal, H. 2004. Extensive open ocean aquaculture development within wind farms in Germany: the prospect of offshore co-management and legal constraints. Ocean & Coastal Management 47(3–4):95– 122. Campbell, D.E., and Newell, C.R. 1998. MUSMOD, a production model for bottom culture of the blue mussel, Mytilus edulis L. Journal of Experimental Marine Biology and Ecology 219:171–203. Cardoso, J.F.M.F., Witte, J.I.J., and van der Veer, H.W. 2006. Intra- and interspecies comparison of energy flow in bivalve species in Dutch coastal waters by means of the Dynamic Energy Budget (DEB) theory. Journal of Sea Research 56:182–197. Cloern, J.E. 1982. Does benthos control phytoplankton biomass in south San Francisco Bay? Marine Ecology Progress Series 9:191–202. Cohen, R.R.H., Dresler, P.V., Philips, E.J.P., and Cory, R.L. 1984. The effect of the Asiatic clam Corbicula fluminea on phytoplankton of the Potomac River. Limnology and Oceanography 29:170–180. Commito, J.A., Como, S., Grupe, B.M., and Dowa, W.E. 2008. Species diversity in the soft-bottom intertidal zone: biogenic structure, sediment, and macrofauna across mussel bed spatial scales. Journal of Experimental Marine Biology and Ecology 366:70–81. Dame, R., Dankers, N., Prins, T., Jongsma, H., and Smaal, A. 1991. The influence of mussel beds on nutrient cycling in the Dutch Wadden Sea and Eastern Scheldt estuaries. Estuaries 14(2): 130–138.

28

Shellfish Aquaculture and the Environment

Dowd, M. 1997. On predicting the growth of cultured bivalves. Ecological Modelling 104:113– 131. Dowd, M. 2005. A bio-physical coastal ecosystem model for assessing environmental effects of marine bivalve aquaculture. Ecological Modelling 183:323–345. Duarte, P., Labarta, U., and Fernández-Reiriz, M. 2008. Modelling local food depletion effects in mussel rafts of Galician Rias. Aquaculture 274:300–312. Dumbauld, B.R., Ruesink, J.L., and Rumrill, S.S. 2009. The ecological role of bivalve shellfish aquaculture in the estuarine environment: a review with application to oyster and clam culture in West Coast (USA) estuaries. Aquaculture 290(3–4):196–223. Fabi, G., Manoukian, S., and Spagnolo, A. 2009. Impact of an open-sea suspended mussel culture on macrobenthic community (Western Adriatic Sea). Aquaculture 289:54–63. FAO. 1999. Regional Review on Trends in Aquaculture Development—Europe. Food and Agricultural Organization of the United Nations, Fish Culture Research Institute, Szarvas, Hungary. FAO. 2009. The State of World Fisheries and Aquaculture 2008 (SOFIA). Food and Agriculture Organization of the United Nations, Rome. Ferreira, J.G., Wolff, W.J., Simas, T.C., and Bricker, S.B. 2005. Does biodiversity of estuarine phytoplankton depend on hydrology? Ecological Modelling 187(4):513–523. Ferreira, J.G., Hawkins, A.J.S., and Bricker, S.B. 2007a. Management of productivity, environmental effects and profitability of shellfish aquaculture—the Farm Aquaculture Resource Management (FARM) model. Aquaculture 264:160–174. Ferreira, J.G., Hawkins, A.J.S., Monteiro, P., Service, M., Moore, H., Edwards, A., Gowen, R., Lourenco, P., Mellor, A., Nunes, J.P., Pascoe, P.L., Ramos, L., Sequeira, A., Simas, T., and Strong, J. 2007b. SMILE—Sustainable Mariculture in Northern Irish Lough Ecosystems: Assessment of Carrying Capacities for Environmentally Sustainable Shellfish Culture in Carlingford Lough, Belfast Lough, Larne Lough and Lough Foyle. Institute of Marine Research, Lisbon, Portugal.

Ferreira, J.G., Bricker, S.B., and Simas, T.C. 2007c. Application and sensitivity testing of an eutrophication assessment method on coastal systems in the United States and European Union. Journal of Environmental Management 82(4): 433–445. Ferreira, J.G., Hawkins, A.J.S., Bricker, S.B., and Xiao, Y. 2007d. Virtual oysters in the tagus estuary and in chesapeake bay—a model analysis of shellfish productivity and eutrophication control. Presented at the 6th European Conference on Ecological Modelling, ECEM’07, Trieste, Italy, 27–30 November 2007. Ferreira, J.G., Andersson, H.C., Corner, R.A., Desmit, X., Fang, Q., de Goede, E.D., Groom, S.B., Gu, H., Gustafsson, B.G., Hawkins, A.J.S., Hutson, R., Jiao, H., Lan, D., Lencart-Silva, J., Li, R., Liu, X., Luo, Q., Musango, J.K., Nobre, A.M., Nunes, J.P., Pascoe, P.L., Smits, J.G.C., Stigebrandt, A., Telfer, T.C., de Wit, M.P., Yan, X., Zhang, X.L., Zhang, Z., Zhu, M.Y., Zhu, C.B., Bricker, S.B., Xiao, Y., Xu, S., Nauen, C.E., and Scalet, M. 2008a. Sustainable Options for People, Catchment and Aquatic Resources: The SPEAR Project, An International Collaboration on Integrated Coastal Zone Management. Institute of Marine Research, Lisbon, Portugal. Ferreira, J.G., Hawkins, A.J.S., Monteiro, P., Moore, H., Service, M., Pascoe, P.L., Ramos, L., and Sequeira, A. 2008b. Integrated assessment of ecosystem-scale carrying capacity in shellfish growing areas. Aquaculture 275:138–151. Ferreira, J.G., Sequeira, A., Hawkins, A.J., Newton, A., Nickell, T., Pastres, R., Forte, J., Bodoy, A., and Bricker, S.B. 2009a. Analysis of coastal and offshore aquaculture: application of the FARM™ model to multiple systems and shellfish species. Aquaculture 289:32–41. Ferreira, J.G., Bricker, S.B., and Nunes, J.P. 2009b. Application of the EcoWin2000 and FARM models to shellfish culture in Killary Harbour and Dungarvan Harbour. UISCE Final Project Report. BIM Ireland (in press). Giles, H., Broekhuizen, N., Bryan, K.R., and Pilditch, C.A. 2009. Modelling the dispersal of biodeposits from mussel farms: the importance of simulating biodeposit erosion and decay. Aquaculture 291:168–178. Giles, H., and Pilditch, C.A. 2004. Effects of diet on sinking rates and erosion thresholds of mussel

Role of shellfish farms in the ecosystem

Perna canaliculus biodeposits. Marine Ecology Progress Series 282:205–219. Gosling, E. (ed.). 2003. Bivalve Molluscs: Biology, Ecology and Culture. Blackwell Science, Oxford. Grant, J., and Bacher, C. 1998. Comparative models of mussel bioenergetics and their validation at field culture sites. Journal of Experimental Marine Biology and Ecology 219:21–44. Grant, J., Cranford, P., Hargrave, B., Carreau, M., Schofield, B., Armsworthy, S., Burdett-Coutts, V., and Ibarra, D. 2005. A model of aquaculture biodeposition for multiple estuaries and field validation at blue mussel (Mytilus edulis) culture sites in eastern Canada. Canadian Journal of Fisheries and Aquatic Sciences 62:1271–1285. Griebeler, E.M., and Seitz, A. 2007. Effects of increasing temperatures on population dynamics of the zebra mussel Dreissena polymorpha: implications from an individual-based model. Oecologia 151:530–543. Hallegraeff, G.M. 1993. A review of harmful algal blooms and their apparent global increase. Phycologia 32(22):79–99. Hawkins, A.J.S., and Bayne, B.L. 1992. Physiological interrelations, and the regulation of production. In: Gosling, E. (ed.), The Mussel Mytilus: Ecology, Physiology, Genetics and Culture. Elsevier, Amsterdam, pp. 171–222. Hawkins, A.J.S., James, M.R., Hickman, R.W., Hatton, S., and Weatherhead, M. 1999. Modelling of suspension-feeding and growth in the green-lipped mussel Perna canaliculus exposed to natural and experimental variations of seston availability in the Marlborough Sounds, New Zealand. Marine Ecology Progress Series 191:217–232. Hawkins, A.J.S., Duarte, P., Fang, J.G., Pascoe, P.L., Zhang, J.H., Zhang, X.L., and Zhu, M. 2002. A functional simulation of responsive filter-feeding and growth in bivalve shellfish, configured and validated for the scallop Chlamys farreri during culture in China. Journal of Experimental Marine Biology and Ecology 281:13–40. Heasman, K.G., Pitcher, G.C., McQuaid, C.D., and Hecht, T. 1998. Shellfish mariculture in the Benguela system: raft culture of Mytilus galloprovincialis and the effect of rope spacing on food extraction, growth rate, production and condition of mussels. Journal of Shellfish Research 17:33–39.

29

Hoffmann, E.E., Klinck, J.M., Kraeuter, J.N., Powell, E.N., Grizzle, R.E., Buckner, S.C., and Bricelj, V.M. 2006. A population dynamics model of the hard clam, Mercenaria mercenaria: development of the age- and length-frequency structure of the population. Journal of Shellfish Research 25:417–444. Incze, L.S., Lutz, R.A., and True, E. 1981. Modelling carrying capacities for bivalve molluscs in open suspended-culture systems. Journal of World Maricult Society 12:143–155. Inglis, G.J., Hayden, B.J., and Ross, A.H. 2000. An overview of factors affecting the carrying capacity of coastal embayments for mussel culture. NIWA Client Report CHC00/69, Christchurch, New Zealand. Jolly, C.M., and Clonts, H.A. 1993. Economics of Aquaculture. Food Products Press, New York. Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L.-O., Olrog, L., Rehnstam-Holm, A.-S., Svensson, J., Svensson, S., and Syversen, U. 2005. Improving marine water quality by mussel farming: a profitable solution for Swedish society. Ambio 34(2):131–138. McCausland, W.D., Mente, E., Pierce, G.J., and Theodossiou, I. 2006. A simulation model of sustainability of coastal communities: aquaculture, fishing, environment and labour markets. Ecological Modelling 193(3–4):271–294. McKindsey, C.W., Thetmeyer, H., Landry, T., and Silvert, W. 2006. Review of recent carrying capacity models for bivalve culture and recommendations for research and management. Aquaculture 261(2):451–462. Melià, P., and Gatto, M. 2005. A stochastic bioeconomic model for the management of clam farming. Ecological Modelling 184:163–174. Morales, Y., Weber, L.J., Mynett, A.E., and Newton, T.J. 2006. Mussel dynamics model: a hydroinformatics tool for analyzing of different stressors on the dynamics of freshwater mussel communities. Ecological Modelling 19:448– 460. Navarro, E., S. Méndez, S., Ibarrola, I., and Urrutia, M.B. 2009. Comparative utilization of phytoplankton and vascular plant detritus by the cockle Cerastoderma edule: digestive responses during diet acclimation. Aquatic Biology 6:247–269.

30

Shellfish Aquaculture and the Environment

Neori, A., Chopin, T., Troell, M., Buschmann, A.H., Kraemer, G.P., Halling, C., Shpigel, M., and Yarish, C. 2004. Integrated aquaculture: rationale, evolution and state of the art emphasizing seaweed biofiltration in modern mariculture. Aquaculture 231:361–391. Newell, R.I.E. 2004. Ecosystem influences of natural and cultivated populations of suspension feeding bivalve molluscs: a review. Journal of Shellfish Research 23(1):51–61. Nobre, A.M., Ferreira, J.G., Newton, A., Simas, T., Icely, J.D., and Neves, R. 2005. Management of coastal eutrophication: integration of field data, ecosystem-scale simulations and screening models. Journal of Marine Systems 56(3/4): 375–390. Nobre, A.M., Ferreira, J.G., Nunes, J.P., Yan, X., Bricker, S., Corner, R., Groom, S., Gu, H., Hawkins, A., Hutson, R., Lan, D., Lencart e Silva, J.D., Pascoe, P., Telfer, T., Zhang, X., and Zhu, M. 2010. Assessment of coastal management options by means of multilayered ecosystem models. Estuarine, Coastal and Shelf Science 87:43–62. Nunes, J.P., Ferreira, J.G., Gazeau, F., Lencart-Silva, J., Zhang, X.L., Zhu, M.Y., and Fang, J.G. 2003. A model for sustainable management of shellfish polyculture in coastal bays. Aquaculture 219(1–4):257–277. Paltzata, D.L., Pearce, C.M., Barnes, P.A., and McKinley, R.S. 2008. Growth and production of California sea cucumbers (Parastichopus californicus Stimpson) co-cultured with suspended Pacific oysters (Crassostrea gigas Thunberg). Aquaculture 275(1–4):124–137. Pascoe, P.L., Parry, H.E., and Hawkins, A.J.S. 2009. Observations on the measurement and interpretation of clearance rate variations in suspensionfeeding bivalve shellfish. Aquatic Biology 6:181–190. Pouvreau, S., Bacher, C., and Héral, M. 2000. Ecophysiological model of growth and reproduction of the black pearl oyster, Pinctada margaritifera: potential applications for pearl farming in French Polynesia. Aquaculture 186:117–144. Pouvreau, S., Bourles, Y., Lefebvre, S., Gangnery, A., and Alunno-Bruscia, M. 2006. Application of a dynamic energy budget model to the Pacific oyster, Crassostrea gigas, reared under various environmental conditions. Journal of Sea Research 56:156–167.

Powell, E.N., Hofmann, E.E., Klinck, J.M., and Ray, S.M. 1992. Modelling oyster populations. I. A commentary on filtration rate. Is faster always better? Journal of Shellfish Research 11:387–398. Prins, T.C., Smaal, A.C., and Dame, R.F. 1998. A review of the feedbacks between bivalve grazing and ecosystem processes. Aquatic Ecology 31:349–359. Raillard, O., Deslous-Paoli, J.M., Héral, M., and Razet, D. 1993. Modélisation du comportement nutritionnel et de la croissance de l’huître japonaise Crassostrea gigas. Oceanology Acta 16:73–82. Reid, G.K., Robinson, S., Chopin, T., Lander, T., MacDonald, B.A., Haya, K., Burridge, F., Page, F., Ridler, N., Justason, A., Sewuster, J., Powell, F., and Marvin, R. 2007. An interdisciplinary approach to the development of integrated multi-trophic aquaculture (IMTA): bioenergetics as a means to quantify the effectiveness of IMTA systems and ecosystem response. World Aquaculture Society. Aquaculture 2007 conference proceedings. Ren, J.S., and Ross, A.H. 2001. A dynamic energy budget model of the Pacific oyster Crassostrea gigas. Ecological Modelling 142:105–120. Ren, J.S., and Ross, A.H. 2005. Environmental influence on mussel growth: a dynamic energy budget model and its application to the greenshell mussel Perna canaliculus. Ecological Modelling 189:347–362. Ren, J.S., and Schiel, D.R. 2008. A dynamic energy budget model; parameterisation and application to the Pacific oyster Crassostrea gigas in New Zealand waters. Journal of Experimental Marine Biology and Ecology 361:42–48. Ridler, N., Wowchuk, M., Robinson, B., Barrington, K., Chopin, T., Robinson, S., Page, F., Reid, G., Szemerda, M., Sewuster, J., and Boyne-Travis, S. 2007. Integrated Multi-Trophic Aquaculture (IMTA): a potential strategic choice for farmers. Aquaculture Economics & Management 11(1): 99–110. Ross, A.H., and Nisbet, R.M. 1990. Dynamic models of growth and reproduction of the mussel Mytilus edulis L. Functional Ecology 4:777–787. Ryther, J.H., Dunstan, W.M., Tenore, K.R., and Huguenin, J.E. 1972. Controlled eutrophication:

Role of shellfish farms in the ecosystem

increased food production from the sea by recycling human wastes. Biology Science 22: 144–152. Scavia, D., and Bricker, S.B. 2006. Coastal eutrophication assessment in the United States. Biogeochemistry 79:187–208. Scholten, H., and Smaal, A.C. 1999. The ecophysiological response of mussels (Mytilus edulis) in mesocosms to a range of inorganic loads: simulations with the model EMMY. Aquatic Ecology 33:83–100. Sequeira, A., Ferreira, J.G., Hawkins, A.J., Nobre, A., Lourenço, P., Zhang, X.L., Yan, X., and Nickell, T. 2008. Trade-offs between shellfish aquaculture and benthic biodiversity: a modelling approach for sustainable management. Aquaculture 274:313–328. Sharma, K.R., Leung, P., Chen, H., and Peterson, A. 1999. Economic efficiency and optimum stocking densities in fish polyculture: an application of data envelopment analysis (DEA) to Chinese fish farms. Aquaculture 180:207– 221. Shumway, S.E. 1990. A review of the effects of algal blooms on shellfish and aquaculture. Journal of World Aquatic Society 21(2):65–104. Shumway, S.E., Cucci, T.L., Newell, R.C., and Yentch, T.M. 1985. Particle selection, ingestion and absorption in filter-feeding bivalves. Journal of Experimental Marine Biology and Ecology 91:77–92. Smaal, A.C., Prins, T.C., Dankers, N., and Ball, B. 1998. Minimum requirements for modelling bivalve carrying capacity. Aquatic Ecology 31:423–428. Solidoro, C., Pastres, R., Melaku Canu, D., Pellizzato, M., and Rossi, R. 2000. Modelling the growth of Tapes philippinarum in North Adriatic lagoons. Marine Ecology Progress Series 199:137–148. Spillman, C.M., Hamilton, D.P., Hipsey, M.R., and Imberger, J. 2008. A spatially resolved model of seasonal variations in phytoplankton and clam

31

(Tapes philippinarum) biomass in Barbamarco Lagoon, Italy. Estuarine, Coastal and Shelf Science 79:187–203. USEPA. 2004. Water Quality Trading Assessment Handbook. USEPA Office of Water, Washington, DC. Van Haren, R.J.F., and Kooijman, S.A.L.M. 1993. Application of a dynamic energy budget model to Mytilus edulis (L.). The Netherlands Journal of Sea Research 31:119–133. Watson, R., and Pauly, D. 2001. Systematic distortions in world fisheries catch trends. Nature 414:534–536. Weise, A.M., Cromey, C.J., Callier, M.D., Archambault, P., Chamberlain, J., and McKindsey, C.W. 2009. Shellfish-DEPOMOD: modelling the biodeposition from suspended shellfish aquaculture and assessing benthic effects. Aquaculture 288:239–253. Whitall, D., Bricker, S., Ferreira, J.G., Nobre, A., Simas, T., and Silva, M.C. 2007. Assessment of eutrophication in estuaries: pressurestate-response and source apportionment. Environmental Management 40:678–690. Xiao, Y., Ferreira, J.G., Bricker, S.B., Nunes, J.P., Zhu, M., and Zhang, X. 2007. Trophic assessment in Chinese coastal systems—review of methodologies and application to the Changjiang (Yangtze) Estuary and Jiaozhou Bay. Estuaries and Coasts 30(6):1–18. Yamamuro, M., Hiratsuka, J.I., Ishitobi, Y., Hosokawa, S., and Nakamura, Y. 2006. Ecosystem shift resulting from loss of eelgrass and other submerged aquatic vegetation in two estuarine lagoons, Lake Nakaumi and Lake Shinji Japan. Journal of Oceanography 62: 551–558. Zhang, J., Hansen, P.K., Fang, J.G., Wang, W., and Jiang, Z. 2009. Assessment of the local environmental impact of intensive marine shellfish and seaweed farming—application of the MOM system in the Sungo Bay China. Aquaculture 287:304–310.

Chapter 2

Shellfish aquaculture and the environment: an industry perspective William Dewey, Jonathan P. Davis, and Daniel C. Cheney

Introduction Marine aquaculture has become an increasingly important contributor to global food production necessary to support an expanding world population estimated at 6 billion today and expected to grow to 8 billion by 2028 (U.S. Census Bureau 2009). Global demand for seafood products alone is projected to increase by 70% in the next 30 years as harvests from traditional capture fisheries either remain stable or continue to decline. Currently, production of seafood from fish and shellfish including aquaculture provide 15% of average annual animal protein consumption to 2.9 billion people (FAO 2008a, 2008b). At the same time, coastal marine ecosystems worldwide that support wild shellfisheries are

threatened by pollution, habitat degradation, overharvesting, and a growing dependence on common-pool resources, among other concerns (Jackson et al. 2001; Lotze et al. 2006; Halpern et al. 2008; Beck et al. 2009; Smith et al. 2010), lending an increasingly important role for sustainably produced and managed intensive marine aquaculture to fill the widening gap in the world’s capacity for food production. The shift to increasingly intensive aquaculture operations where suitable coastal sites exist, coupled with either peak extraction or serial depletion of many fisheries stocks, has stimulated discussion about how humans utilize and ultimately manage aquatic resources in the future (Pauley et al. 1998; Marra 2005). What remains clear is that marine aquaculture will likely continue to provide an increasingly

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 33

34

Shellfish Aquaculture and the Environment

World Aquaculture Production of Shellfish 5,000,000 4,500,000

Production (MT)

4,000,000 3,500,000 3,000,000 Abalones Clams Mussels Oysters Scallops

2,500,000 2,000,000 1,500,000 1,000,000 500,000

19 5 19 0 5 19 2 5 19 4 5 19 6 5 19 8 6 19 0 1962 6 19 4 6 19 6 6 19 8 7 19 0 7 19 2 19 74 7 19 6 7 19 8 8 19 0 8 19 2 1984 8 19 6 8 19 8 9 19 0 9 19 2 9 19 4 9 19 6 9 20 8 0 20 0 2002 2004 06

0

Year Figure 2.1 Shellfish aquaculture production for abalone, clams, mussels, oysters, and scallops for years 1950–2006. (Source: FAO Fisheries Department, Fishery Information, Data and Statistics Unit. FishStat Plus: universal software for fishery statistical time series, Version 2.3, 2000.) MT, metric tons.

significant share of fish- and shellfish-based resources (Costa-Pierce 2002), such that combined fish and shellfish aquaculture will in 2009, for the first time, supply half of the world’s seafood supply (FAO 2008a, 2008b). This is an important milestone for marine aquaculture in general as the world’s attention for millennia has been on increasingly efficient extraction of marine resources without significant attention paid to effectively manage or replenish stocks until relatively recently, and mainly only in developed portions of the world (Worm et al. 2009). Policy debate flourishes over the array of potential solutions necessary to maintain overall ocean health and the ability of nearshore marine ecosystems to remain resilient to climate change, pollution including excess nutrient runoff, and other threats associated with anthropogenic inputs to near-shore habitats critical to marine plants and animals. Yet, marine ecosystems are counted upon to provide the vast ecological and economic benefits to Earth and its inhabitants. To that end, marine aquaculture, including shellfish culture, has the

potential to supply an increasingly valuable contribution of high-quality protein-based foods for humans in cost-effective and sustainable sea-based production systems. Production from shellfish culture alone has greatly expanded. Shellfish production through aquaculture has greatly increased (in particular for oysters and clams), with the majority produced in Asia (FAO 2008a, 2008b) (Fig. 2.1). An ecologically and economically sustainable shellfish industry large enough to supply growing human populations with seafood depends on four critical components. First, a viable industry depends on the maintenance of certified growing waters located in productive, sheltered waters with access to marine shorelines. Shellfish are best grown in areas that are free from pollution, rich in productivity, and readily accessible by a trained workforce. Second, a stable and predictable regulatory framework that is responsive to changes in industry practices remains an integral requirement for successful aquaculture. This includes industry participation in developing, in concert with regulators, environmental codes of prac-

An industry perspective

tice (ECOPs; e.g., best management practices [BMPs] [see Chapter 3 in this book]) that remain flexible to changes in policy, scientific information, industry innovation, and markets. Third, a strong infrastructure for processing, transport, marketing, and sales of product, and for monitoring pollutants and other factors that can affect shellfish quality and safety. Fourth, an educated public that embraces the quality and variety of seafood products produced through marine farming is critically important (see Chapter 16 in this book). Public awareness of ecological and regulatory issues affecting the marine environment is high and growing, and it is largely incumbent on the marine farming sector to demonstrate its commitment to maintaining the biological integrity of the environments they utilize for farming shellfish. Shellfish aquaculture has not historically been subject to the same level of public and regulatory scrutiny to which intensive fish and shrimp culture operations have become accustomed. This view is changing. In the developed world, public interest is often less on enabling food to be grown from seafarm-based production, and more on ensuring the public’s multiple use of shorelines, the maintenance of the ecological integrity of coastal marine environments to the exclusion of other uses, and the preservation of viewscapes. Attempts to increase utilization of marine and shoreline environments for shellfish aquaculture has resulted in both existing and proposed operations receiving greatly increased public scrutiny over the last 30 years. Much of the public opposition to shellfish farming has been expressed as either real or perceived impacts on the environment due to specific shellfish aquaculture and fishery operations. Recent reviews have focused on the suite of ecological effects associated with shellfish aquaculture for a variety of habitats (see Kaiser et al. 1998; Nizzoli et al. 2006; Dumbauld et al. 2009), and a voluminous literature has developed describing species-specific environmental and

35

ecological interactions associated with shellfish aquaculture. Much of this work is summarized in this book. This chapter provides a perspective on the role the shellfish industry has in maintaining the environmental integrity of coastal environments suitable for shellfish aquaculture and associated commercial shellfisheries. First, the evolution of the shellfish industry in the U.S. Pacific Northwest is described with specific reference to the advocacy and stewardship role shellfish growers have long held relative to the development and maintenance of marine water quality and land use standards necessary to support a viable industry. The second section considers the variety of BMPs (Chapter 3 in this book) that have been developed to help integrate culture practices with the best available science and other public concerns relative to environmental effects associated with shellfish aquaculture. Together, with a short discussion of third-party sustainability certification efforts, a perspective is provided to suggest that the shellfish industry will continue to play an integral role into the future in helping shape public policy with regard to sustained multiple use of near-shore marine environments.

Shellfish farmers and harvesters history of water quality protection and stewardship roles Shellfish growers as water quality advocates The shellfish industry has long been an advocate for protecting and restoring water quality. Simply speaking, clean water is the lifeblood of the industry. This necessity is due to the historical practice in Europe, North America, and Australasia of consuming shellfish live— the case in particular for oysters and clams. This long tradition has the potential to increase the public health risk and subsequently requires that the waters from which they are harvested

36

Shellfish Aquaculture and the Environment

be exceptionally clean. In the United States, public health controls for shellfish were initiated in 1925 with the creation of the National Shellfish Sanitation Program (NSSP) (2007). The development of the program was triggered by a large number of illnesses attributed to the consumption of raw oysters, clams, and mussels in the late nineteenth century and early twentieth century. A large typhoid fever outbreak in 1924 with illnesses in New York, Chicago, and Washington, DC, and linked to the consumption of sewage-contaminated oysters was the final impetus for the creation of the NSSP. Unable to assure consumers that their products were safe to eat, the shellfish industry joined with state and local public health officials to request the Surgeon General of the U.S. Public Health Service to develop necessary control measures to ensure a safe shellfish supply. The NSSP ensures that shellfish are harvested from waters that meet stringent water quality standards and that they are transported, handled, and processed in a sanitary manner. Since the inception of the NSSP, both shellfish harvesters and growers have been strong advocates for water quality. Shellfish growers are arguably more ardent water quality advocates and resource stewards than wild harvesters, with the key distinction being that shellfish growers own the shellfish they grow and either own or lease the land on which they farm. De Alessi (1996) explored this phenomenon in Washington State where shellfish growers typically either own or lease the tidelands they farm. De Alessi notes that “[o]yster growers have had a profound effect on Willapa Bay and elsewhere in Washington . . . Ownership ties oyster growers to a particular spot and gives them a vested interest in protecting the local environment; their livelihood depends on it.” In an effort to expand awareness and demand for clean marine waters, Taylor Shellfish Farms (TSF), Washington State’s largest producer of farmed shellfish, has joined

efforts in other regions of the state and country to encourage shellfish gardening. Through sponsored seed and gear sales, TSF provides shellfish seed, culture equipment, information, and encouragement to shoreline residents to create shellfish gardens. In the process, shoreline owners learn about the importance of ensuring their septic systems are functioning properly, controlling pet and domestic animal wastes and the fate of herbicides and pesticides placed onto lawns and gardens. Like commercial shellfish growers, private shellfish gardeners have become strong advocates for water quality within their communities. Similarly, community shellfish gardens are springing up in the Pacific Northwest as they have elsewhere in the United States and, for similar reasons, commercial shellfish growers have encouraged and supported these publicly managed ventures. Community shellfish gardens are usually managed by environmental nongovernmental organizations and provide similar water quality education opportunities. In many cases, when the shellfish crops are sold, a portion of the revenue is used for water quality-related projects. In the Chesapeake Bay and elsewhere on the U.S. East Coast where nutrient pollution is a demonstrable problem, shellfish gardening is encouraged for the ecological services shellfish provide through filtration activities and removal of excess nutrients at harvest (Chapters 1, 4, 9, and 8 in this book). Besides toxins, heavy metals and organic pollutants that can impact the safety of shellfish cultivated for human consumption, shellfish growers are also impacted directly by pollution that also directly affects shellfish health. Molluscan shellfish, and their larvae in particular, have been long demonstrated to be highly vulnerable to degraded water quality. Impacts to the health of cultured shellfish crops also stimulate environmental advocacy by shellfish growers. A classic example in Washington State illustrates the relationship between a viable shell-

An industry perspective

fish industry and clean water. During the first half of the twentieth century, pulp and paper industries developed and thrived in the Pacific Northwest during a period when shellfish growers were focused on the culture of both native oysters (Ostrea lurida) and the introduced Pacific oyster (Crassostrea gigas). Untreated sulfite liquor effluent and other waste discharge from the pulp mills fouled bays and destroyed oyster beds in various parts of Puget Sound. Particularly hard hit were native oyster beds in southern Puget Sound and around Bellingham in northern Puget Sound. To protect their crops and tidelands, oyster farmers responded by suing the pulp mills and lobbying the state legislature and Congress for laws and regulations to address regulations for pulp mill effluent. It was an unpopular fight in local communities as it generated conflicts among different sectors (e.g., timber producers, pulp and paper mills, and shellfish producers) whose livelihoods collectively depended on locally generated resources (Steele 1957). Oyster growers played an integral role in this dispute and as a group responded with programs designed to generate public awareness of the relationship between pulp mill effluent and declining oyster populations (Fig. 2.2). Significantly, continued lobbying by the shellfish industry of the state legislature in 1945 incited the creation of the Washington Pollution Control Commission (now the Washington Department of Ecology). Water quality standards were established at that time along with a control board to enforce regulations. Unfortunately, the pulp mills continued to pollute. A 1957 letter by oysterman Ed Gruble to the Seattle Post Intelligencer newspaper claiming that “Puget Sound has almost become a ‘marine desert’ . . . 75 percent of the raw liquor still goes directly into Bellingham Bay, and the bay for a considerable distance from the pulp mill is black as ink” (cited from Gordon et al. 2001). Gruble and other oystermen testified repeatedly before Congress in

37

Washington, DC. These efforts by Washington’s oystermen contributed to the enactment of the Clean Water Act (CWA) in 1972. While the CWA was largely effective at stemming effects of point source pollution, shellfish growers continue to be plagued by nonpoint source pollution. Today, shellfish growers remain active both individually and through the Pacific Coast Shellfish Grower’s Association (PCSGA) lobbying for more stringent storm-water and onsite sewage laws and regulations.

Solutions to resolve use conflicts associated with shellfish culture development and maintenance with an emphasis on U.S. West Coast examples Shorelines where shellfish are grown in the United States and around the world are also popular places to reside. Population growth in coastal counties has been dramatic. In the contiguous United States, coastal counties are home to 53% of the nation’s population, or more than 148 million people. By 2015 the coastal population in the United States is expected to reach 165 million people, or an average density of 327 people per square mile (NOAA 1998; USDC 2001). Globally, approximately 37% of the world’s population lives within 100 km (62 m) of the coastline and 50% reside within 200 km (124 m) (Cohen et al. 1997; Hinrichson 2001). The increased popularity of coastal living not only contributes to water quality problems, but it also results in escalating conflicts over land use. The land use conflicts facing shellfish farmers are similar to those that terrestrial farmers and foresters experience as historic resource production areas are increasingly used for both housing and recreation. Conflicts arise when new residents unaccustomed to working waterfronts move to shorelines having historic shellfish operations or when changes occur in

38

Shellfish Aquaculture and the Environment

Figure 2.2 Washington State shellfish growers were actively engaged in water quality initiatives in the 1950s. Pulp mill effluent and its effect on oysters was a galvanizing issue and helped focus oyster growers on threats to the marine environment. (Source: David Steele, The Immigrant Oyster (Ostrea gigas) Now Known as the Pacific Oyster.)

the species cultivated that result in the use of more intensive culture methods. Conflicts also arise when shellfish culturists expand to locate new operations adjacent to previously developed shorelines with other established waterdependent uses. Some common user conflicts include the following: aesthetic impacts; noise; lighting and

hours of operations; physical interference with recreational or commercial fishing; and recreational use of the shoreline. Comprehensive land use planning coupled with zoning can be useful to reduce use conflicts, preserve existing shellfish culture operations, and provide opportunities for growth of the shellfish aquaculture industry. Together,

An industry perspective

land use planning and zoning are critical tools used for addressing terrestrial land use conflicts. Unfortunately, these approaches have been almost universally underutilized when considering use conflicts in marine and shoreline environments. One consequence of this lack of planning and zoning for industrial uses of the marine environment is that both longstanding and new shellfish culture operations are often challenged by shoreline opponents, most usually during the permitting stage for any expanded farm operations or the establishment of new farms. Challenges to most farm operations usually result in significant delays and increased costs that have significantly impeded the growth of the shellfish industry in much of the developed world over the last 30 years. As has been the case elsewhere, opposition to shellfish aquaculture development is often generated at the local or regional level and federal support for shellfish aquaculture has been positive, but generally limited by inadequate funding. In 1990 Congress created in Section 309 of theCoastal Zone Management Act (CZMA), a program of Coastal Zone Enhancement Grants to enhance state programs relative to eight national priorities. In 1996 Congress added aquaculture as a ninth priority. The act provides funding for the “[a] doption of procedures and policies to evaluate and facilitate the siting of public and private aquaculture facilities in the coastal zone, which will enable States to formulate, administer, and implement strategic plans for marine aquaculture.” Coastal zone management programs are encouraged by the National Oceanic and Atmospheric Administration (NOAA) to use this funding to develop and enhance regulatory planning and intragovernmental coordination mechanisms to provide meaningful state participation in the management of aquaculture, to balance the uses of coastal and ocean resources, to coordinate with existing authorities, and to minimize user conflicts. Unfortunately, funding for this program has been limited and most states have opted to use

39

what funding is available for priorities other than aquaculture. In the U.S. Pacific Northwest where commercial shellfish culture has had a long, historical presence, extensive new residential shoreline development has been intense. As a consequence, use conflicts between shoreline owners and shellfish farmers have increased, especially in recent years as more intensive forms of shellfish culture have been adopted by the industry (Fig. 2.3). Shellfish growers in Washington State have taken both proactive and reactive measures at the federal, state, and local levels to address the conflicts. Washington State’s primary law for land use planning is the Growth Management Act (GMA). One of the main goals of GMA is to identify and preserve commercial resources including timber, agricultural, and mineral resource lands. Shellfish growers have made the case recently that commercial shellfish tidelands are also resource lands that are deserving of protection under the act. This measure was omitted when the law was passed and the growers have proposed legislation to rectify this problem and hope to bring commercially valuable shellfish tidelands under the umbrella of the GMA. If successful, use conflicts should diminish over time as purchasers of shoreline property are notified that the tidelands adjacent are zoned for commercial shellfish culture and what types of activities might be expected on those lands. Water quality should improve, as well as counties adopt more stringent septic and storm water regulations to protect tidelands zoned for commercial shellfish culture. At the local level, Washington’s Shoreline Management Act (SMA) regulates shoreline development through the implementation of Shoreline Master Programs (SMPs). To ensure that policies and regulations are both supportive and reasonable and that irresponsible growth does not place their livelihoods in jeopardy, shellfish growers have again made a conscious effort to participate with local planning commissions, conservation districts and service on boards of nongovernmental

40

Shellfish Aquaculture and the Environment

Figure 2.3 Intensive geoduck farming operations on intertidal beaches in close proximity to upland property owners have generated resource use conflicts in Washington State. (Photo credit: Jon Rowley.)

organizations, and other entities who share a common agenda of protecting water quality and conserving resource lands. Outreach and education by shellfish growers is also effective at reducing user conflicts. A number of farms and grower organizations provide information through websites, videos and flyers, host or sponsor tours, and shellfish festivals and receptions that inform the public and policymakers about the industry. Growers regularly contribute time, funding, and product to assist fundraisers for local not-for-profit civic or environmental organizations. Several salmonid fishes in the Pacific Northwest listed in recent decades as threatened or endangered under the Endangered Species Act occupy habitat for a part of their life cycle that overlaps intertidal shellfish beds. The Magnuson Stevens Act subsequently mandated that essential fish habitat (EFH) for federally managed fish species be identified and protected. While shellfish beds can provide critical and essential fish habitat by themselves, the activities associated with shellfish farming can potentially have adverse impacts on fishes utilizing shellfish beds for habitat. For these specific reasons the PCSGA pursued the devel-

opment and implementation of BMPs in the mid 1990s. BMPs, or ECOPs (see Chapter 3 in this book) as they are sometimes called, have helped circumvent the need for formal regulations and provided growers with a “social license” to operate where they have been successfully developed and implemented. Formal regulations and use conflicts emerge where growers have not developed or have failed to adequately implement BMPs. The section following further describes how BMPs have specifically helped integrate regulatory reform with shellfish farming practices. In March 2007, in an effort to improve consistency of federal permitting of shellfish culture, the U.S. Army Corps of Engineers established a new programmatic permit. Nationwide Permit (NWP) 48 covers all existing shellfish farms prior to March 12, 2007. This permit requires consultation by the National Marine Fisheries Service and U.S. Fish and Wildlife Service (USFWS), as well as CWA certification and approval by states that the permit is consistent with the coastal zone management programs before farms are technically covered. BMPs under development and knowledge of the science about environ-

An industry perspective

41

Figure 2.4 Shellfish growers in Washington State lead volunteer efforts for annual beach cleanup activities to collect wayward or discarded aquaculture derived and other debris from beaches in Puget Sound. (Photo credit: William Dewey.)

mental effects have proven beneficial in the NWP 48 consultation process with NOAA, USFWS, and the Army Corps. Farm equipment and debris lost from shellfish culture operations either during storms or from simple carelessness is a growing problem for growers. Materials escaping the confines of marine farms and subsequently deposited on public and private beaches significantly erode public perception of shellfish aquaculture. Growers recognize this as an issue and regularly patrol shorelines in the vicinity of their farms to collect loose materials (Fig. 2.4). Growers are also seeking better means to secure culture equipment, reduce the amount of plastics used, and employ recyclable and biodegradable materials. In the Pacific Northwest, growers organize geographically broad and coordinated beach cleanups twice a year. In southern Puget Sound, a large group of companies and representatives from local tribes participate in collecting debris from over 100 mi of beach during these tightly coordinated cleanup events and all materials coming

from aquaculture sources are sorted and quantified by type and area of origin. This information is used to identify and work with specific growers to stop the proliferation of debris at the source. Self-policing of practices that do not serve the industry’s collective benefit has proved to be an important tool for reducing use conflicts. The CZMA is due for reauthorization. At the federal level, growers are engaged to ensure that when the CZMA is reauthorized it includes incentives and/or requirements for aquaculture planning for state waters and the preservation of working waterfronts. There is increased pressure as well for expanding domestic aquaculture production, developing ocean energy alternatives (wind, wave, current, thermal), and expanding offshore oil production in the U.S. exclusive economic zone (EEZ). The potential for increased user conflicts over completing uses has resulted in policymakers to utilize marine spatial planning increasingly as a tool to identify synergies among user groups and avoid conflicts between proposed

42

Shellfish Aquaculture and the Environment

uses. In 2009, the Obama administration formed an Interagency Ocean Policy Task Force to develop recommendations for effective coastal and marine spatial planning. In March 2010, Washington’s Governor Gregoire signed a bill into law establishing a process to conduct marine spatial planning for the state’s marine waters. Shellfish growers in the U.S. Pacific Northwest believe these efforts will help address the user conflicts that are negatively impacting their businesses. The history of the shellfish industry as advocates for clean water and land use policy as it relates to shellfish aquaculture in the U.S. Pacific Northwest serves as a lens through which to view user conflicts elsewhere. The lessons learned in Washington State are similar to those in other parts of the developed world where conflicts over resources are both common and increasing. As the public expands into rural areas for first and second home development and increased recreational opportunities, traditional rural economies based on resource extraction and farming are often targeted for reform and greater regulation. To combat these trends, the shellfish industry has adopted the tactics described above to better engage the public. These efforts have a record of mixed success and are largely viewed as stopgap measures to enable marine farming to continue to the extent possible. It is likely that increased BMP development and implementation coupled with a greater emphasis on marine zoning will be necessary in the future to accommodate growth in the shellfish aquaculture industry.

BMPs, the shellfish industry, and the role of available research Description, development, and implementation of BMPs for shellfish aquaculture As described under Chapter 3, a BMP is a tool defining and/or prescribing types of activities

or operations to meet some type of production, environmental or engineering goal. BMPs describe methods or techniques found to be the most effective and practical means to achieve an objective, while making the optimum use of natural and human resources. When referencing environmental goals, BMPs are often used synonymously with ECOPs. BMPs and ECOPs are typically created by industry members, often in concert with nongovernmental organizations and regulatory agencies. Coincident with the expanded application of marine environmental policies and regulations coupled with recognition of the role BMPs have in helping conserve and manage aquatic resources, the development and incorporation of BMPs by the shellfish aquaculture industry is increasing. Welldesigned BMPs offer guidance to both the industry and regulatory agencies for a broad range of practices including regulatory compliance, training, farm siting, planting and harvesting, pest, predator and disease control, waste management, vessel/vehicle operations, and shipping/packaging. BMPs are often tailored to apply to industry-specific harvest, processing, and production practices to ensure shellfish products meet public health and safety standards (see Chapter 3 in this book). These practices are typically covered under Hazard Analysis and Critical Control Point (HAACP) protocols, usually administered by state government public health agencies in concert with the U.S. Food and Drug Administration (USFDA) through the Interstate Shellfish Sanitation Conference (ISSC). The ISSC was formed in 1982 to help foster and promote shellfish sanitation through the cooperation of state and federal control agencies, the shellfish industry, and the academic community. The program has been generally successful in bringing agencies and the industry together to help resolve problems relating to shellfish health and sanitation. Table 2.1 offers a list of representative BMP programs in North America, Europe, and

Table 2.1 Examples of best management practice and environmental code of practice guidance for the shellfish industry. Sponsoring organization

Title

Date

Comment

Federation of European Aquaculture Producers

Code of Conduct for Aquaculture

2000

Developed with EU and UNFAO input; general guidelines for all aquaculture products, largely targeting finfish producers1

New Zealand Mussel Industry Council

New Zealand Mussel Industry Environmental Code of Practice

2004

Outlines desired environmental outcomes for mussel producers, with voluntary compliance and continued development2

Pacific Coast Shellfish Growers Association (PCSGA)

Environmental Codes of Practice for the Pacific Coast Shellfish Industry

2002

Comprehensive application of an environmental policy for all bivalve shellfish products; updated regularly3

British Columbia Shellfish Growers Association (BCSGA)

Environmental Management System Code of Practice

2001

Intended to foster and develop good neighbor practices and a public attitude of commitment to working with growers in protecting marine resources4

Aquaculture Council of Western Australia (ACWA)

Western Australian Mussel Aquaculture Industry Environmental Code of Practice

2003

Assesses farm practice risk, suggests measures to reduce that risk, and explains applicable regulation/policy5

Washington State Department of Natural Resources

Best Management Practices for Geoduck Aquaculture on State Owned Aquatic Lands

2007

Relates to farm and crop management, and harvest on public lands; based on available literature and grower input6

Florida Department of Agriculture and Consumer Services, Division of Aquaculture

Aquaculture Best Management Practices Rule (Manual)

2007

Comprehensive, for all aquaculture; required for shellfish hatcheries, producers leasing submerged lands from the State of Florida, and processors7

Virginia Institute of Marine Science, College of William and Mary

Best Management Practices for the Virginia Shellfish Culture Industry

2008

Voluntary guidelines, with four major management areas: social, operational, biological, and food safety and quality8

Southeastern Massachusetts Aquaculture Center (SEMAC)

Best Management Practices for the Shellfish Culture Industry in Southeastern Massachusetts

2009

Voluntary procedures developed in collaboration with the shellfish industry to focus on improving production while preserving the environment9

1

www.feap.info/feap/code/default_en.asp New Zealand Mussel Industry Council. 2004. New Zealand Mussel Industry Environmental Code of Practice. New Zealand Mussel Industry Council Ltd., Blenheim, New Zealand. 3 www.pcsga.org/ 4 www.bcsga.ca/industry-resources/additional-reports-documents/environmental-management-code-of-practice 5 www.aquaculturecouncilwa.com/assets/files/pdf/October%202002%20Mussel%20ECoP.pdf 6 www.dnr.wa.gov/Publications/aqr_aqua_2007bmp.pdf 7 www.floridaaquaculture.com/publications/P-01499-booklet-07_BMP_RULE.pdf 8 web.vims.edu/adv/aqua/MRR%202008_10.pdf?svr=www 9 www.mass.gov/agr/aquaculture/docs/Shellfish_BMPs_v09-04a.pdf 2

43

44

Shellfish Aquaculture and the Environment

Australia/New Zealand. These BMPs offer examples of both voluntary and compulsory standards that provide growers, processors, regulators, and consumers with a uniform set of guidelines to ensure sustainable and environmentally sensitive farm practices. In some cases, growers complying with all or most of the BMPs receive recognition through their governing organization or industry group. Additional information on environmental standards applied at the national level is available on fact sheets at the United Nations Food and Agriculture Organization (UNFAO) Fisheries and Aquaculture Department, National Aquaculture Legislation Overview (NALO) website: www.fao.org/fishery/nalo/ search/en. Environmental BMPs developed for the shellfish industry address measures to reduce or minimize, or mitigate the effects of culture practices on aquatic and terrestrial resources and interactions with other users of marine resources. A common thread in environmental BMPs is the application of methods to comply with existing environmental norms and standards typically applied to land-based farms. Because shellfish farms occupy aquatic habitats where the water is in common public ownership, the detail and complexity of BMPs developed for aquatic uses tend to markedly exceed those associated with land-based operations. Key elements in environmental BMPs guiding shellfish culture are mainly related to culture practices (species and type of culture method, and associated activities) and farm site and size of the operation. Examples of environmental BMPs are listed in Table 2.2. Shellfish aquaculture is predominantly a near-shore practice utilizing intertidal bedlands and shallow coastal waters. The presence of shellfish and other structure creates habitat that is functionally similar to macroalgae and seagrass (Dumbauld et al. 2009) and culture practices involving the placement and transfer of seeded shell, culture in bags on or off the bottom, mechanical or hand harvest,

and other activities may have specific environmental benefits relative to the enhancement and restoration potential of the farm or culture site. Different growout methods have unique environmental effects and varying resource requirements. For example, bottom-cultured oysters require no external inputs other than seeding and crop tending, whereas oysters grown in bag-on-bottom, longline, or suspended systems require the use of more material, intensive containment, or support systems; however, ground culture may be dependent on larger vessels for bed planting and harvest and can require more active predator and pest control. Culture practice BMPs and ECOPs are generally prescriptive and typically directed to specific culture methods and addressing measures to avoid adverse impacts. For example, the Environmental Management System Code of Practice for the British Columbia Shellfish Growers Association recommends “[a]ny modification of tenure substrate (e.g., removal of rocks, gravelling) should be conducted in compliance with an approved management plan and should be planned to minimize impacts on other naturally occurring wildlife and fish habitat.” The extent and intensity of farmed areas are controlled both by the availability of growing space and the carrying capacity of the water body to sustainability support both cultivated shellfish and the other aquatic organisms found in or moving through the culture area. Expansion of existing shellfish farms and creation of new farms is subject to varying levels of government and public oversight, ranging from minimal involvement and permitting requirements in areas with a history of shellfish culture to an extensive public process, the need for multiple permits, and highly prescriptive permitting conditions. Unfortunately, as was discussed earlier, this process can overshadow important siting criteria related to the environmental suitability of farming areas, the economic and cultural values of the farming practice, and the need to protect water bodies

Table 2.2 Examples of environmental BMPs for shellfish farms with focus on specific issues associated with industry practices. Issue

Objective

Example of BMP

Source

Public education

Promote public commitment to protect and enhance marine resources

Set a positive public example of environmental stewardship during all production activities

1, 2

Waste management

Minimize amount of waste produced and released by shellfish production

Practice the principles of reduction, reuse, recycling, and recovery for farms, processors, and suppliers

1, 2, 3

Access and property rights

Ensure the legal rights of public and private access are maintained

Recognize the needs of other marine resource users and promote methods to minimize user conflicts

1, 2

Noise

Minimize noise impacts

Make every reasonable effort to minimize noise during regular farming activities

1, 2

Light

Minimize the impact of artificial lighting

Make every effort to minimize and shield the use of lights on site

1, 2

Odor

Minimize farmgenerated odors

Keep all vessels, equipment, and vehicles clean and well maintained at all times

1, 2

Chemicals and fuels

Minimize the impact of chemicals, fuels, and lubricants

Minimize the use of chemicals that could enter the marine environment; use biodegradable products where appropriate; establish a spill response plan to handle emergencies

1, 2, 3, 4

Site density

Maintain and enhance the productive capacity of shellfish growing waters

Monitor the quality of shellfish on the farm and respond accordingly with appropriate husbandry practices; support research into the development of appropriate carrying capacity models

1, 2, 3

Interaction with fish and wildlife

Minimize and avoid impact on fish and wildlife

Predator control practices should be targeted at specific animals and have limited additional impacts on fish and wildlife habitats

1, 2, 3

Biofouling control

Minimize the amount and impact of biofouling discarded at the farm site

Adopt operating and maintenance practices that reduce the potential for nontarget species to become a significant factor

1, 2, 4

Vehicle operations

Minimize or avoid the use of vehicles in intertidal areas

Restrict route selection in intertidal areas to hard surfaces along the upper intertidal zone

1, 2

Vessels and marine equipment

Minimize any negative impacts of marine equipment operations

Receive adequate and appropriate training in the operation and maintenance of all their marine equipment; avoid damaging marine life and sensitive habitat when operating vessels and equipment

1, 2, 4

Aesthetics

Minimize public aesthetic concerns

Maintain farm sites and infrastructure in a clean and orderly manner

1, 2, 3

Navigation safety

Enhance marine safety in farm operations

Promote public awareness of the need for caution when operating vessels around shellfish operations

1, 2

45

46

Shellfish Aquaculture and the Environment

Table 2.2 (Continued) Issue

Objective

Example of BMP

Source

Harvesting

Reduce the environmental impacts of harvesting operations

Minimize harvest impact on the marine environment, other marine resource users, and upland owners

1, 2

Upland facility operations

Minimize environmental effects

Design and operate land-based facilities in a manner that minimizes adverse impacts to the receiving waters, adjacent wetlands, and uplands

1, 4

Farm maintenance

Minimize environmental, navigation, and aesthetic issues of culture systems

Collect and properly dispose or recycle materials when they are removed during harvesting; remove old or unnecessary gear in a timely manner

1, 3, 4

Genetic protection

Avoid impacts to wild shellfish

For seed secured out-of-state sources, hatcheries must utilize broodstock from waters within the growing region in their genetic selection program

4

1

www.bcsga.ca/industry-resources/additional-reports-documents/environmental-management-code-of-practice Pacific Coast Shellfish Growers Association. 2002. Environmental Codes of Practices. Unpublished. 3 web.vims.edu/adv/aqua/MRR%202008_10.pdf?svr=www 4 www.floridaaquaculture.com/publications/P-01499-booklet-07_BMP_RULE.pdf 2

certified or suited for shellfish farming from adverse land use practices. BMPs created by regulatory agencies are typically directed at these siting issues (Table 2.1). The question of system carrying capacity has been the subject of a large body of research focusing on the capability of the culture environment to support a given biomass and the effects of varying culture density or biomass on a fixed rate of aquatic production (Ferreira et al. 2007a, b, 2008; Sequeira et al. 2008; see also Chapters 1 and 6 in this book). BMPs to address carrying capacity are in place for several shellfish species as is the development of models to assess interactive carrying capacity (Ferreira et al. 2007b). In addition, the use of bivalve shellfish culture to reduce water column nutrients and help remediate land-based nutrient inputs is a potentially important application for BMPs that address specific culture practices. This may be especially relevant when considering, for example, the expansion of shellfish farming specifi-

cally for ecosystem benefits derived from nutrient reduction (Gren et al. 2009; see also Chapter 1).

Efforts to establish a program for sustainability certification and Third-party accountability The limitation of current shellfish BMPs is that they are either prepared by grower/processor groups in response to or at the request of regulatory agencies or other stakeholders, or are generated by government agencies as specific regulatory or management tools. Over the last 10 years, however, several third-party resource certification programs have been developed, largely focused on environmental sustainability in land-based and aquatic applications. Aquatic certifications to date have been mainly directed toward wild-harvest capture fisheries, an example being the Marine Stewardship Council (MSC) certification for wild-caught

An industry perspective

Alaska salmon. Several organizations are, however, currently engaged in the development of certification and inspection standards for bivalve aquaculture at both regional and international levels. The World Wildlife Fund (WWF) has been working since 2007 to create a suite of global environmental certification standards under the umbrella of the Bivalve Aquaculture Dialogue (originally called the Mollusc Dialogue with a goal “. . . to create performance-based standards that will minimize the key social and environmental issues associated with bivalve farming.” The Bivalve Aquaculture Dialogue identified key environmental and social issues related to bivalve production as follows: “1) Ecosystem integrity: Habitat interactions and ecological community structure modifications; 2) Genetics: Gene transfer to wild populations, inbreeding, and escapes; 3) Biosecurity: Deliberate or inadvertent introduction of new exotic species, pests, and pathogens; 4) Disease and pest management: Transfer of disease and pests to and from the wild, within the wild, and within aquaculture systems; loading of pathogens; and the use of chemicals for preventing and controlling diseases and pests; 5) Farm maintenance: Management and disposal of debris (e.g., nets and bags), chemicals, and organic waste; processing of wastes; treatment of effluent; and maintenance of equipment; 6) Multiuser cooperation: Location, development, and aesthetics of aquaculture sites; and public access to aquaculture sites” (from http:// www.worldwildlife.org/what/globalmarkets/ aquaculture/dialogues-molluscs.html). When the standards are finalized, it is proposed to give them to a new organization, the Aquaculture Stewardship Council, to be cofounded by the WWF. This organization will be responsible for working with independent, third-party entities to certify farms that are in compliance with the standards (WWF 2009a, 2009b). A new program on the U.S. West Coast is in the process of developing farm- and crop-

47

specific certifications based on terrestrial farming practices to certify both environmentally appropriate and employment-friendly shellfish production practices. This certification program, managed by the Portland, Oregonbased Food Alliance (www.foodalliance.org) develops inspection criteria and guidance for farms, food handlers, processors, and distributors based on a set of certification standards. Environmentally specific standards include the following: (1) ensure the health and humane treatment of animals; (2) no use of nontherapeutic antibiotics; (3) no genetically modified animals; 4) management procedures to reduce pesticide use and toxicity; (5) protect soil and water quality; (6) protect and enhance aquatic and wildlife habitat; and (7) continuously improve management practices. To assess whether an operation meets the Food Alliance standards, independent third-party inspectors use the evaluation criteria to assess whether and how desired management outcomes are being achieved. The benefit to the bivalve shellfish producer and provider is that environmental certification provides independent verification of marketing claims for social and environmental responsibility, can differentiate and add value to products, and can protect and enhance branded shellfish products. Efforts to develop certification standards should consider the scale and how widely adopted a program will become if implemented. Namely, should certification programs that address only local or regional considerations be developed as opposed to programs that are designed from the outset to consider a broader geographical range that includes the variability in domestic and international shellfish farm operations, food handlers, processors, distributors, and markets for shellfish? This is an important distinction. The former approach may have significant value at the local or regional level and may satisfy local producers, buyers, and consumers but can lead to a proliferation of standards and practices that are piecemeal in both development and

48

Shellfish Aquaculture and the Environment

implementation, vary in scale and application, and may not be readily duplicated on regional and greater scales. The latter approach, while difficult to implement, could potentially offer greater uniformity of standards, better industry buy-in, and may incorporate a far greater range of national and international participants. Both approaches are likely important to pursue as the shellfish industry in many parts of the world grapples with increased public scrutiny and intensified interest in better defining the environmental and social costs of food production.

guard water quality and other environmental standards relating to land use and nonpoint pollution in the Pacific Northwest over the last century offer important lessons for the public. As BMPs and certification standards are coupled with a better understanding of the environmental costs and benefits of shellfish aquaculture, it is likely that increasingly sustainable shellfish culture practices will provide food, increased ecosystem benefits for the public at large, while contributing to the safeguarding of a viable shellfish industry into the future.

Conclusion

Literature cited

Shellfish aquaculture worldwide is growing, especially in regions where shellfish resources form an integral component of the human diet and an important means for producing biologically efficient, sustainable sources of highquality food coupled with economic vitality for coastal areas that are increasingly impacted by human development. Overall, the contribution of global supplies of shellfish to fisheries products have grown from 3.9% of total production (by weight) in 1970 to about 36% in 2006; on a per capita basis, global production supplied by aquaculture increased from 0.7 kg per capita in 1970 to 7.8 kg per capita in 2006, an 11-fold increase over 36 years (FAO 2008a, 2008b). Shellfish growers in developed countries where use conflicts are well established are increasingly turning to BMPs and ECOPs to respond to public concerns and help ensure sustainable production into the future. Appropriately sited and managed, shellfish aquaculture will likely continue to supply increasing quantities of high-quality seafood utilizing increasingly domesticated stocks to enable sustainable coastal development economic opportunity and better food security (Marra 2005). The long-term stewardship role provided by the shellfish industry to help safe-

Beck, M.B., Brumbaugh, R.D., Airoldi, L., Carranza, A., Coen, L.D., Crawford, C., Defeo, O., Edgar, G.J., Hancock, B., Kay, M., Lenihan, H., Luckenbach, M.W., Toropova, C.L., and Zhang, G. 2009. Shellfish Reefs at Risk: A Global Analysis of Problems and Solutions. The Nature Conservancy, Arlington, VA, p. 52. Cohen, J.E., Small, C., Malinger, A., Gallup, J., and Sachs, J. 1997. Estimates of coastal populations. Science 278:1211–1212. Costa-Pierce, B.A. 2002. Ecological Aquaculture: the Evolution of the Blue Revolution. Blackwell Science, Oxford, UK. De Alessi, M. 1996. Oysters and Willapa Bay. Center for Conservation Case Study. Competitive Enterprise Institute, Washington DC. Dumbauld, B., Ruesink, J., and Rumrill, S. 2009. The ecological role of bivalve shellfish aquaculture in the estuarine environment: A review with application to oyster and clam culture in West Coast (USA) estuaries. Aquaculture 290:196– 223. Ferreira, J., Hawkins, A., and Bricker, S. 2007a. Management of productivity, environmental effects and profitability of shellfish aquaculture— the farm aquaculture resource management (FARM) model. Aquaculture 264:160–174. Ferreira, J., Hawkins, A., Monteiro, P., Service, M., Moore, H., Edwards, A., Gowen, R., Lourenco, P., Meller, A., Nunes, J., Pascoe, P., Sequeira, A., Simas, T., and Strong, J. 2007b. SMILE-Sustainable Mariculture in

An industry perspective

Northern Irish Lough Ecosystems—Assessment of Carrying Capacity for Environmentally Sustainable Shellfish Culture in Carlingford Lough, Strangford Lough, Belfast Lough, Larne Lough and Lough Foyle. Ed. Institute of Marine Research (IMAR). Ferreira, J., Hawkins, A., Monteiro, P., Moore, H., Service, M., Pascoe, P., Ramos, L., and Sequeira, A. 2008. Integrated assessment of ecosystemscale carrying capacity in shellfish growing areas. Aquaculture 275(1–4):138–151. Food and Agriculture Organization of the United Nations (FAO). 2008a. The State of World Fisheries and Aquaculture 2008. Technical Fisheries Bulletin. Fisheries and Aquaculture Department. FAO Rome (2009). www.fao.org/ docrep/011/i0250e/i0250e00.htmFAO Food and Agriculture Organization of the United Nations. 2008b. FISHSTAT Plus: Universal Software for Fishery Statistical Time Series (Food and Agriculture Organization, Rome) Version 2.32. Gordon, D.G., Blanton, N.E., and Nosho, T.Y. 2001. Heaven on the Half Shell: the Story of the Northwest’s Love Affair with the Oyster. Washington Sea Grant Program and West Winds Press, Seattle. Gren, I., Lindahl, O., and Lindqvist, M. 2009. Values of mussel farming for combating eutrophication: An application to the Baltic Sea. Ecological Engineering 35:935–945. Halpern, B.S., Walbridge, S., Selkoe, K.A., Kappel, C.V., Micheli, F., C.D. Agrosa, Bruno J.F., Casey, K.S., Ebert, C., Fox, H.E., Fujita, R., Heinemann, D., Lenihan, H.S., Madin, E.M.P., Perry, M.T., Selig, E.R., Spalding, M., Steneck, R., and Watson, R. 2008. A global map of human impact on marine ecosystems. Science 319:948–952. Henrickson, S.E., Wong, T., Allen, P., Ford, T., and Epstein, P.R. 2001. Marine swimming-related illness: implications for monitoring and environmental policy. Environmental Health Perspectives 109(7):645–650. Jackson, J.B.C., Kirby, M.X., Berger, W.H., Bjorndal, K.A., Botsford, L.W., Bourque, B.J., Bradbury, R.H., Cooke, R., Erlandson, J., Estes, J.A., Hughes, T.P., Kidwell, S., Lange, C.B., Lenihan, H.S., Pandolfi, J.M., Peterson, C.H., Steneck, R.S., Tegner, M.J., and Warner, R.R.

49

2001. Historical overfishing and the recent collapse of coastal ecosystems. Science 293: 629–638. Kaiser, M.J., Laing, I., Utting, S.D., and Burnell, G.M. 1998. Environmental impacts of bivalve mariculture. Journal of Shellfish Research 17:59–66. Lotze, H., Lenihan, H., Bourque, B., Bradbury, R., Cooke, R., Kay, M., Kidwell, S., Kirby, M., Peterson, C., and Jackson, J. 2006. Depletion, degradation, and recovery potential of estuaries and coastal seas. Science 312:1806–1809. Marra, J. 2005. When will we tame the oceans? Nature 436:175–176. National Oceanic and Atmospheric Administration. 1998. Pressures on coastal environments—population: distribution, density and growth. State of the Coast Report. National Oceanic and Atmospheric Administration. Silver Spring, Maryland. p. 32. Nizzoli, D., Welsh, T.D., Fano, E.A., and Viaroli, P. 2006. Impact of clam and mussel farming on benthic metabolism and nitrogen cycling, with emphasis on nitrate reduction pathways. Marine Ecology Progress Series 315:151– 165. Pauley, D., Christensen, V., Dalsgaard, J., Froese, R., and Torres, F. 1998. Fishing down marine food webs. Science. 279:860–863. Sequeira, A., Ferreira, J., Hawkins, A., Nobre, A., Lourenço, P., Zhang, X., Yan, X., and Nickell, T. 2008. Trade-offs between shellfish aquaculture and benthic biodiversity: A modeling approach for sustainable management. Aquaculture 274:313–328. Smith, M.D., Roheim, C.A., Crowder, L.B., Halpern, B.S., Turnipseed, M., Anderson, J.L., Asche, F., Bourillon, L., Guttormsen, A.G., Kahn, A., Liguori, L.A., McNevin, A., O’Connor, M.I., Squires, D., Tyedmers, P., Brownstein, C., Carden, K., Klinger, D.H., Sagarin, R., and Selkoe, K.A. 2010. Sustainability and global seafood. Science 327:784–786. Steele, E.N. 1957. The Rise and Decline of the Olympia Oyster. Fulco Publication, Elma, WA. U.S. Census Bureau. 2009. Population Division. www.census.gov/ipc/www/idb/index.php U.S. Department of Commerce. 2001. Population Change and Distribution: 1990 to 2000. Publication No. C2KBR/01-2. U.S. Census

50

Shellfish Aquaculture and the Environment

Bureau, U.S. Department of Commerce, Washington, D.C. U.S. Department of Health and Human Services, Food and Drug Administration. 2007. Interstate Shellfish Sanitation Conference, Guide for the Control of Molluscan Shellfish. World Wildlife Fund. 2009a. Aquaculture dialogues overview fact sheet. World Wildlife Fund, Washington, DC. www.worldwildlife.org/what/ globalmarkets/aquaculture/WWFBinaryitem 10107.pdf p. 2. World Wildlife Fund. 2009b. Molluscan aquaculture dialogues. World Wildlife Fund, Washington,

DC. www.worldwildlife.org/what/globalmarkets/aquaculture/dialogues-molluscs.html Worm, B., Hilborn, R., Baum, J.K., Branch, T.A., Collie, J.S., Costello, C., Fogarty, M.J., Fulton, E.A., Hutchings, J.A., Jennings, S., Jensen, O.P., Lotze, H.K., Mace, P.M., McClanahan, T.R., Minto, C., Palumbi, S.R., Parma, A.M., Ricard, D., Rosenberg, A.A., Watson, R., and Zeller, D. 2009. Rebuilding global fisheries. Science 325:578–585.

Chapter 3

Molluscan shellfish aquaculture and best management practices John A. Hargreaves

Introduction The concept of sustainability has slowly made inroads into public consciousness. Acting on their understanding of the concept, an increasing number of consumers are now making informed choices about seafood consumption. Political consumption of ecolabeled seafood provides a market-based incentive for producers to adopt responsible production methods. A price premium is the reward to the producer. In theory, the broader commons also benefits from the adoption of production techniques that are more environmentally benign. Underpinning a large part of certification and labeling programs are best management practices (BMPs), combined in creative and site-specific ways by producers to achieve

compliance with process standards or to improve environmental performance and sustainability. Sustainability is easy to define as a concept, but it is more difficult to identify specific examples of sustainable human endeavor. In large measure, the move toward a more sustainable development trajectory has been a reaction to evidence of what is understood to be unsustainable development. In many respects, it is easier to define sustainability in contrast to the effects of what is considered unsustainable. Some examples of the environmental consequences of unsustainable development include global climate change, water pollution (eutrophication and ocean acidification), high rates of nonrenewable resource use, land use changes, and the effect of invasive

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 51

52

Shellfish Aquaculture and the Environment

species on biodiversity. In developed countries, conventional farming is seen as unsustainable. Such farming is often characterized by largescale, large-capital investment; concentrated ownership; monocultures; high-yielding hybrid seed; large quantities of chemical fertilizer, pesticide, and energy inputs; and “factory” farms for livestock. Aquaculture has not been immune to criticism for what are described as the effects of unsustainable practices. Media reports about aquaculture highlight the accumulation of toxins and contaminants in cultured products, use of banned antibiotics, general or specific negative environmental impacts, use of genetically modified organisms (GMOs) (Chapter 12 in this book), animal welfare abuses, and human rights abuses. Fortunately, shellfish aquaculture has been largely spared the most severe criticism, although critics of aquaculture often paint with a broad brush, leading to generally negative attitudes or confusion in the minds of the public about any products from aquaculture. Perhaps the main reason that shellfish aquaculture is perceived favorably is the difference between two broad types of aquaculture. The primary source of nutrition in extractive aquaculture, such as shellfish farming, is naturally occurring foods that develop in response to nutrient loading into coastal ecosystems. In fed aquaculture, the primary source of nutrition is intentionally applied manufactured feed or fertilizer. The global-scale environmental effects of feed manufacture and use far exceed those of extractive aquaculture, such as shellfish farming. Shellfish aquaculture is often held up as an example of sustainable aquaculture (Shumway et al. 2003). First, sustainable aquaculture must be economically viable in the appropriate market context, whether local or global. Sustainability implies a long-term time frame. There are numerous examples of shellfish aquaculture growing areas, facilities, and markets that span many generations, including

those located in coastal areas of the South China Sea and North Atlantic Ocean. Sustainable aquaculture minimizes dependence on purchased inputs and makes efficient use of the resources that are applied. Sustainable aquaculture protects and maintains the supporting environment by operating within the ecological carrying capacity, which is defined as the yield that can be produced without resulting in an irreversible change to ecosystem structure and function. Shellfish aquaculture provides numerous ecological services, described elsewhere in this book (Chapters 1 and 9), including food provision, water quality improvement, and nutrient cycling. Sustainable aquaculture must maintain ecosystem resilience; it must operate to maintain the capacity of the supporting ecosystem to withstand stress and shocks. Finally, sustainable aquaculture, such as shellfish farming, provides meaningful livelihoods and promotes human well-being. This, in turn, enhances the vibrancy and stability of coastal communities as a key part of coastal economic development. This chapter will discuss BMPs in the context of direct implementation or active manifestation of the concept of sustainability. Sustainability is not a specific end point, but rather a trajectory of constant improvement, as measured by sustainability indicators. Collectively, BMPs are a locally adapted, farmscale approach to reduce environmental impacts and increase resource use efficiency. BMPs, when incorporated into codes of conduct or environmental management systems, form the basis for ecolabeling or other product certification programs. The price premium or market access obtained through product certification can serve as an incentive to change producer behavior. BMPs can be an effective means of achieving environmental protection goals, but there are limitations. For some environmental effects of shellfish aquaculture (e.g., invasive species, biodiversity concerns, diseases), BMPs can play a role but are only effective in the context

Molluscan shellfish aquaculture 53

of broader-scale regional, national, and international agreements. BMPs can also play a role in addressing local stakeholder (i.e., NIMBY) concerns, but are not the sole effective approach.

Ecosystem change and shellfish aquaculture We live on a planet with human-dominated ecosystems and where human well-being is critically dependent on the ecosystem services of the biosphere. The capacity of ecosystems to provide these services is related to the type and intensity of various drivers of ecosystem change. In the coastal ecosystems where shellfish aquaculture facilities are embedded, the major direct drivers of ecosystem change are (1) eutrophication and, more generally, pollution from adjacent ecosystems; (2) habitat loss; (3) invasive species; and 4) climate change, especially as manifest through ocean acidification and changes in sea level (MEA 2005). As the intensity of these drivers increases, the finite capacity of ecosystem services to provide food, regulate water quality, and provide basic ecosystem support functions of nutrient cycling and primary production becomes limiting. In general, in human-dominated ecosystems, shellfish aquaculture enhances the environmental capacity to provide these needed ecosystem services (Chapters 1 and 9), primarily by removing nutrients in the form of filtered seston. Shellfish aquaculture can be impacted by some of the direct drivers of coastal ecosystem change (Chapter 17) but, in the case of invasive species (Chapter 14), may itself be the main contributor to ecosystem change. To the extent that direct drivers of ecosystem change affect the capacity of shellfish aquaculture to provide food, BMPs can ameliorate that change and enhance the capacity of shellfish aquaculture to provide food for humans. To the extent that shellfish aquacul-

ture contributes to these direct drivers, BMPs can reduce the effect of shellfish aquaculture as a direct driver of ecosystem change. Effective efforts to address the major drivers of ecosystem change must be comprehensive and holistic. Piecemeal efforts are not costeffective, especially if major sources are not given priority. For example, an effective effort to reduce coastal eutrophication would address all major contributors of nutrients, chemicals, and sediment in the watershed and embayment. In this case, watershed protection BMPs that address runoff from farms and forests is the appropriate focus of efforts to improve environmental quality. The large potential capacity of shellfish aquaculture facilities to remove nutrients from human-dominated coastal ecosystems has a net positive environmental impact with respect to coastal eutrophication.

Classification of impacts The environmental impacts of aquaculture depend on the trophic level of cultured animals, the type of culture system, production intensity, and the extent and concentration of aquaculture development in an area. In the case of molluscan shellfish, in general, animals feed at a low trophic level on natural seston. Culture systems are open to the environment but production intensity is low, with deliberate inputs generally limited to cultivation support structures and hatchery-produced seed. In some locations, particularly in relatively closed embayments, the concentration of shellfish aquaculture facilities can lead to localized depletion of dissolved oxygen concentration and phytoplankton, and accumulation of biodeposits (see Chapter 9). Thus, of the general characteristics of aquaculture production systems that affect the environmental impact of shellfish aquaculture, the extent of aquaculture development relative to ecosystem carrying capacity is the most salient aspect.

54

Shellfish Aquaculture and the Environment

The environmental impacts of shellfish aquaculture occur at a range of spatial and temporal scales. The spatial extent of point source impacts can be divided into three zones (CSTT 1997). Zone A is near the farm or facility, defined by the volume within which tidal currents disperse soluble metabolites during one tidal cycle, with hydraulic residence times of a few hours. Zone B is the local water-body scale or near-field area where water residence times range from a few days to several weeks and where nutrient enrichment can lead to phytoplankton growth if other conditions are favorable. Zone C is the regional or far-field area where water residence times of weeks to months and where higher-order and emergent ecosystem processes occur. In general, nearfield effects are better studied and, as a result, better understood. Near-field effects, such as accumulation of biodeposits and dissolved oxygen depletion, are reversible and the duration is generally short term. The relative importance of near-field environmental impacts is often based on site-specific characteristics, especially the extent of water exchange or renewal. Far-field effects, such as introduction of pathogens or invasive species or effects of shellfish aquaculture on biodiversity, are less amenable to investigation, less well understood, and in some cases the effects are binary (presence, absence) and irreversible. Near-field effects have been the focus of certification efforts and most BMPs are directed toward mitigating the effect of near-field impacts and are applied at the scale of farm or facility. Farfield effects may best be addressed by institutional or policy frameworks, such as international treaties or conventions, or governmental regulation. It is difficult to assign an objective value to specific environmental impacts of aquaculture. There is seldom consensus of opinion on the most important environmental effects. The relative importance of each impact will differ among and even within stakeholder groups. Furthermore, there are often trade-offs

between different environmental impacts. The proposed introduction of the Suminoe oyster Crassostrea ariakensis to the Chesapeake Bay is an example. Introduction and establishment has the potential to improve the health of the bay by increasing filtration capacity. Is this potential improvement worth the risk associated with the introduction of a nonnative species? The Food and Agriculture Organization (FAO) Code of Conduct for Responsible Fisheries indicates that introductions of nonnative species should be done using a precautionary approach. Introducing nonnative species can have deleterious unintended ecological consequences, including displacement of native species, with cascading effects through food webs, and introduction of associated pests and pathogens (hitchhikers) (Chapter 14). The environmental risks associated with nonnative species introduction can be considered relative to the potential benefits associated with food production and other beneficial ecosystem services provided by the introduced species. Is the potential benefit worth the potential cost? Which value is more important?

BMPs In the context of the Clean Water Act, BMPs are defined as “a schedule of activities, prohibitions or practices, maintenance procedures, and other management practices to prevent or reduce the pollution of waters of the United States” (Code of Federal Regulations, 40 CFR 122.2). BMPs were established to address nonpoint source pollution, which refers to pollutant sources that are diffuse in space, such as those from agriculture and forestry. The original goal of BMPs was to protect the quality of flowing and standing freshwater bodies. By extension, and given that eutrophication is a major driver of ecosystem change in coastal marine environments, these BMPs can also protect the quality of coastal waters. Thus,

Molluscan shellfish aquaculture 55

watershed protection BMPs are potentially beneficial to coastal water quality and shellfish aquaculture, perhaps to a greater extent than BMPs for shellfish aquaculture itself. BMPs are specific protocols, practices, or procedures to manage specific operations in a socially and environmentally responsible manner, and are typically based on risk analysis and the best available scientific information (Nash et al. 2001, cited in Jensen and Zajicek 2008). BMPs have been defined simply as the means of mitigating unacceptable environmental interactions (NRC 2010). The current meaning and intent of BMPs now transcends the original focus on water quality protection, having been extended to address animal health and welfare (Chapter 13), food safety, environmental sustainability, and socioeconomic considerations (FAO 2010). BMPs provide specific guidance and instructions for a broad range of construction, operational, and management practices in aquaculture (Table 3.1). These BMPs are adapted by producers to the specific circumstances, requirements, and conditions of a particular site, production system, and species cultured (Chapter 15). Some BMP programs have developed best husbandry practices, some of which also improve resource use efficiency, profitability, and environmental performance. However, not all best husbandry practices improve environmental performance. Many environmental BMPs improve profitability through better environmental quality that improves growth and survival, better resource use efficiency, and the price premium derived from ecolabeling of the products of aquaculture. The acronym BMP refers to “best” management practices but it is perhaps more appropriate to use the word “better” in naming these tools and techniques (Clay 2008). Better practices are those that result in improved environmental performance relative to standard practices. Better implies that the best is yet to come. The word better also implies a notion of continuous improvement. As technology

changes and uncertainty is reduced, the performance target is shifted to accommodate the new conditions. The process of continuous improvement is properly viewed in the context of the best available technology or best professional judgment at a particular time. As new technologies and practices are developed, these will come to replace current “best” practices. The notions of continuous improvement and adaptive management are formally embedded in the ISO 140001 standard for Environmental Management Systems. The benefit of continuous improvement is likely constrained by diminishing returns. From an ecological economics perspective, each practice has a certain cost-benefit ratio. The most readily adopted practices will be those with a low cost-benefit ratio. Over time, the cost to achieve each additional unit of benefit will escalate. At this point, the trade-off between improved environmental performance and reduced economic sustainability must be evaluated. Obviously, the relative importance of each of these dimensions of sustainability is subjective and value-laden, and the process of resolving the trade-off is political.

Codes of conduct BMPs are sometimes incorporated into codes of conduct or environmental management systems, such as the ISO 14000 series of standards. The ISO/IEC (2004) defines a code of practice as a “document that recommends practices or procedures for the design, manufacture, installation, maintenance or utilization of equipment, structures, or products.” The code can be a standard, part of a standard, or independent of a standard. Codes of practice often include general statements of principle for sustainable production. A code of conduct provides a framework for using BMPs to meet environmental management objectives. Implementation of the principles articulated in codes is achieved through BMPs.

Table 3.1 Domain areas and better management practices for bivalve molluscan aquaculture. BMPs for shellfish aquaculture are similar among guidance documents. (Creswell and McNevin 2008).

Site selection, access, and maintenance Select sites of appropriate depth, salinity, substrate, oxygen, and water flow characteristics Maintain benthic biodiversity Avoid sites with submerged aquatic vegetation Minimize risks to navigation when selecting sites Locate floating upwelling systems (FLUPSYs) to comply with navigational requirements and to minimize impacts on the benthic environment Select sites that do not impede public access Select sites for hatcheries and land-based nurseries with a sufficient supply of high-quality water Site land-based facilities in locations that minimize environmental impacts and public health concerns and that are aesthetically consistent with adjacent properties Facility construction Minimize erosion during construction of land-based facilities Properly mark site boundaries, structures, and equipment Secure shellfish culture structures properly Place nets and equipment in suitable areas Use durable materials for facility construction General facility operation Minimize disturbance of substrate in shellfish growing areas during bed preparation and harvesting When collecting wild seed, use equipment and methods that will minimize potential adverse effects on surrounding ecosystems Remove and properly dispose derelict shellfish culture equipment and other solid wastes Properly dispose of wastewater generated by shellfish culture activities Remove growout equipment from water during mechanical cleaning Maintain vessels and equipment in good working order Use fuels, lubricants, and other petroleum-based products in a responsible manner Maintain proper containment of hazardous materials Provide adequate training in waste management Biofouling control Develop and implement an integrated biofouling management plan Use mechanical methods to control biofouling Use brine or freshwater dips to control biofouling, especially by ascidians Use biological methods to control biofouling Use appropriate chemical methods to control biofouling Use caustic chemicals in accordance with manufacturers’ guidelines and neutralize the chemicals prior to disposal When practical, remove biofouling organisms from the farm site

56

Table 3.1 (Continued)

Pest and predator control Develop and implement an integrated pest and predator management plan Comply with regulations that protect marine mammals and threatened and endangered species Use and maintain predator exclusion devices Use predator exclusion nets with hardened coatings Avoid predator exclusion devices that can entangle waterbirds and other predators Use off-bottom culture technologies Use biological control methods where appropriate Use chemical control methods only when other methods are ineffective Invasive species Be knowledgeable of and comply with all regulations governing the importation and transport of bivalve molluscs Carefully inspect all shellfish seedstock received from hatcheries and remove nontarget species Assure that all nonlocal organisms are nonviable when disposed Use broodstock and seed from verified sources Disease prevention, control, and eradication Adhere to state regulations regarding importation of mollusc broodstock or seed Isolate the culture facility from sources of infection Minimize stress through good husbandry practices Properly quarantine or dispose of infected stock and contaminated materials Limit wet-storage activities Maintain good records Drugs and therapeutic agents Minimize drug and therapeutic use in the shellfish hatchery Store drugs and chemicals in appropriate locations Use and dispose drugs and therapeutic agents as labeled Neutralize disinfecting agents and wash water containing disinfecting agents prior to disposal Genetic diversity Plan genetic programs to ensure adequate genetic diversity of the cultured organisms Use performance improvement programs that reduce risks of negative impacts Use appropriate chemicals for polyploid induction Aesthetic values Avoid excessive visual disruption to sites Operate facilities to minimize noise Reduce smells and odors emanating from shellfish facilities Minimize lighting disturbances Community relations Raise awareness and educate the public with respect to farming activities Maintain regular communication with surrounding activities

57

58

Shellfish Aquaculture and the Environment

Codes of conduct or practice and environmental management systems that include BMPs are similar approaches to minimize farm-scale environmental impacts. The most prominent example is the FAO Code of Conduct for Responsible Fisheries, which provides guidelines to United Nations member states. Some codes provide guidance on BMPs that are used in code implementation. Aquaculture codes of practice are documents that describe recommended practices (BMPs) for the production of aquatic animals. In the case of shellfish aquaculture, most BMP programs have been incorporated into codes of conduct prepared and adopted by local or national industry associations (Table 3.2). Such codes typically have general statements of principle and some broadly applicable BMPs. The code of conduct program established by the Pacific Coast Shellfish Growers Association (PCSGA) is being implemented by developing farm-level BMP plans with producers using a standard template. A similar program has recently (2010) been established by the East Coast Shellfish Growers Association (ECSGA).

BMPs and other environmental policy options BMPs can be compared with other policy approaches that are designed and implemented to achieve environmental management and protection goals. These approaches include performance standards, taxes, performance bonds, zoning, and voluntary agreements, among others (Engle and Wossink 2008). Policy options can be combined to achieve environmental management goals. For example, the Marine Stewardship Council specifies a combination of management practices, performance standards, and general design standards in its certification program for marine fisheries. Performance standards are often considered as an alternative to management standards, as

exemplified by BMPs. In contrast to management standards, performance standards specify the goal but do not specify the methods that producers must use to achieve the standard. Considerable flexibility and discretion are given to producers in meeting the standard; producer innovation is encouraged. Performance standards (“achieve a certain objective”) can be contrasted to management standards (“do things this way”), such as BMP-based programs (NRC 2010). BMP programs specify practices that are seen as indicative of a certain effect or environmental performance level. From a compliance perspective, it is easier to verify the presence of a practice than to demonstrate a specific effect and so BMP programs may be preferred by regulators and producers alike. Performance standards may be best applied by local or regional public resource management agencies and considered at the embayment or regional management level. The standards would be applied to manage ecological carrying capacity issues (NRC 2010). The regulatory goal is to maintain ecological effects within the established performance limits. Resource managers can then develop programs for shellfish growers (and other contributors to an impact) that apply specific design standards, such as BMPs, to individual farms or facilities. These programs are adjusted to consider variation in species, culture system, and local conditions. To summarize this approach, performance standards are applied at the basin level and specific BMP programs are applied at the producer level (NRC 2010). Another environmental management policy option that deserves mention is zoning. Zoning, as applied to coastal habitats, is the classification and delineation of a water body according to its suitability for particular uses. Zoning can serve a planning and regulatory function. In the case of shellfish aquaculture, zoning allows planners and regulators to specify the extent of development to maintain the ecosystem carrying capacity and preserve ecosystem

Molluscan shellfish aquaculture 59 Table 3.2 Examples of environmental management programs for shellfish aquaculture. Entity

Program type

Reference

Pacific Coast Shellfish Growers Association

Environmental management system

PCSGA 2001, 2002

Maine Aquaculture Association

Code of practice

Maine Aquaculture Association 2002

Maryland Aquaculture Coordinating Council

Best management practices

Maryland Aquaculture Coordinating Council 2007

Florida

Best management practices (mandatory)

FDACS 2007

Virginia

Code of practice

Oesterling and Luckenbach 2008

United States

Best management practices Massachusetts

Best management practices

Leavitt 2009

East Coast Shellfish Growers Association

Code of practice

Flimlin et al. 2010

Best management practices

Canada Department of Fisheries and Oceans

Operational statements

British Columbia Shellfish Growers Association

Code of practice

BCSGA 2001; BCMAFF 2002

Environmental management system

Europe Irish Sea Fisheries Board

Environmental code of practice

BIM 2003

Association of Scottish Shellfish Growers

Code of good practice

Association of Scottish Shellfish Growers 2005

Seafood Shetland

Code of practice

Seafood Shetland 2007

Bantry Bay Aquaculture

Environmental code of practice

Bantry Bay Aquaculture 2009

Invasive Species Ireland

Code of practice

Kelly and Maguire 2009

Environment Protection Authority (South Australia)

Code of practice

Environment Protection Authority 2005

Aquaculture New Zealand

Environmental management system

Aquaculture New Zealand 2007

Oceania

Code of practice

resilience. Zoning can be a component of comprehensive, integrated coastal zone management plans. Such plans can achieve multiple goals such as maximizing yields from aquaculture, maintaining biodiversity, protecting

threatened and endangered species, protecting watersheds, and preserving access for navigation, recreation, and tourism. The Maine Aquaculture Association has implemented the mechanism of cooperative

60

Shellfish Aquaculture and the Environment

Local Area or Bay Management Agreements to address ecosystem carrying capacity issues with respect to shellfish and salmon farming. These represent cooperative agreements among growers to achieve common objectives, particularly those pertaining to the use of hatcheryreared seed, biosecurity, and pest management issues.

Indicators and compliance monitoring The environmental effect of an activity can be assessed by measuring indicators. For monitoring and certification programs, it is important to develop appropriate indicators or metrics for the most significant environmental effects. Ecosystem performance indicators must be practical, responsive, and easily interpreted. To be most useful, indicators must be easy to measure and meaningfully indicate responses to changes in production practices. The number of indicators should be restricted to a small number of core indicators and a few supplemental indicators. Some indicators are continuous variables and others are binary, indicating presence or absence. Binary indicators are common in product certification standards. Classes of performance indicators include economic, resource use, environmental, animal health and welfare, human resource use, and public perception. Economic indicators measure the effect of farm management on economic performance, as indicated by net profit and internal rate of return. Biophysical resource use indicators measure the gross amount, productivity, and efficiency of resources used. Environmental indicators measure effects of an operation on the environment. These can include the appropriation of carrying capacity, water quality effects, benthic effects, changes to biodiversity, and habitat transformations. Environmental performance standards for shellfish aquaculture

might set limits for criteria such as turbidity, sediment sulfide concentration, changes in benthic biodiversity, and minimum dissolved oxygen concentration. The process of using indicators begins with establishing goals or objectives of the desirable state or condition. Then, indicators of particular impacts are selected and the targets set. The target is the desirable state of the environment and is established as a point-of-reference or performance standard. Once this is done, then the indicators should be measured to establish a baseline. The indicator should be measured before and after implementation of a BMP. Based on the results of indicator measurement, further changes to management may be warranted to meet the indicator targets. Once the targets are achieved, the goals or objectives should be reassessed and readjusted to stimulate further improvement. BMP verification requires a program of compliance monitoring, which can be achieved through self-evaluation, producer group evaluation, or true independent (third-party) evaluation. Table 3.3 provides an example of a sustainability self-evaluation reporting form for shellfish aquaculture. A procedure for review and correction of noncompliance with BMP programs must also be in place. Environmental monitoring and documentation of compliance with performance standards and BMP programs places a considerable record-keeping burden on producers, a feature that is often not fully appreciated before initiating a program.

Incentives for adoption of BMPs There are many potential benefits for producers to implement BMPs voluntarily, especially if BMPs are adopted comprehensively and holistically as part of a code of practice or environmental management system that serves as the basis for a certification program. In general, adoption of BMPs is facilitated when

Molluscan shellfish aquaculture 61

Table 3.3 Example of sustainability reporting for a shellfish farm.

Environmental profile Production volume (t year−1) Diesel fuel used (L year−1) Eco-efficiency (L kg−1 shellfish) Greenhouse gas emissions (kg CO2 equiv) Eco-efficiency (kg kg−1 shellfish) Other wastes (lost gear, other solids) Fuel spills Effect on marine environment Benthic effects Depredation control—seabirds, marine mammals Biofouling control Social sustainability Number of employees Length of service or employee turnover Employee health and safety (number of accidents) Employee training activities Social activities supported by farm Charitable donations Economic sustainability Sales revenue Profits Net surplus Investor returns (IRR)

producers perceive that a particular practice is economically beneficial or represents a low risk to economic sustainability. In this case, adoption of BMPs is selective and the commitment to improved environmental performance is superficial. BMPs can reduce costs and improve operational performance by increasing the efficiency of bioresource use. This is probably more important for fed aquaculture than extractive

aquaculture such as shellfish farming. However, the general approach of systematically reducing environmental effects and improving profitability by increasing input efficiency is nearly always fruitful. This provides one of the most powerful incentives for BMP adoption by producers. Some producers adopt BMPs as a condition of licensing or permitting, such as is the case with the mandatory BMP program in Florida (FDACS 2007). Shellfish producers must have a BMP-based plan approved by the state as a condition of operation. In other cases, producers may achieve regulatory compliance and avoid prosecution and fines by adopting BMPs. Producers may adopt BMPs to avoid more stringent regulations, as exemplified by specific performance standards. In such cases, producers may be selective in adopting only those BMPs that are necessary to meet regulatory requirements. BMPs can also play a role in product marketing, especially when they are included explicitly in product certification standards. It is a way to differentiate one product from another in the market on the basis of production methods that result in improved environmental performance. Product certification can provide access to product markets that are not available to conventional producers, provide a price premium incentive to producers for improved practices, and result in consumer preference of certified product over alternatives. In contrast to regulatory BMP programs that are mandatory, participation in product certification programs is voluntary. Voluntary adoption options for BMPs include environmental sustainability certification programs, investment or credit screens, seafood buyers’ purchasing policies, and incorporation into codes of conduct (Clay 2008). Increasingly, seafood buyers are establishing purchasing policies that specify certain production practices intended to address environmental concerns, real and perceived, many of which are addressed at the farm level with BMPs. In the

62

Shellfish Aquaculture and the Environment

United States, Wal-Mart has committed to sourcing sustainably produced seafood through Marine Stewardship Council certification for wild products and Global Aquaculture Alliance (GAA)/Aquaculture Certification Council certification for cultured products. Whole Foods has established buying standards for some cultured seafood products. In Europe, large transnational grocery chains, such as Carrefour (France) and Tesco (United Kingdom), have active programs of buying and selling seafood that is certified according to standards developed in-house. To date, BMPs have not been used as the basis for shellfish ecolabeling programs. The World Wildlife Fund (WWF) Bivalve Dialogue process and the U.S. Department of Agriculture (USDA) organic standards development are under way but no shellfish have been certified to proposed standards. These will be reviewed below. By far, most of the BMP programs in the United States are producer group initiatives. Examples of these were developed by the PCSGA, ECSGA, and Maine Aquaculture Association (Table 3.2). Perhaps not surprisingly, the content of the technical recommendations (BMPs) among programs is similar (Table 3.1). Incorporating BMPs can also be part of an effective response to concerns raised by some stakeholders, such as members of local communities, who may perceive that they are negatively affected by shellfish aquaculture operations, and representatives of environmental nongovernmental organizations (NGOs), who advocate for practices that improve environmental protection. Adoption of BMPs and public promotion of BMP-based environmental management programs can go a long way to placate these stakeholders.

Roles of BMPs for different groups Producers are the inventors and implementers of BMPs. Through producer group efforts,

BMPs are incorporated into codes of conduct. Producers seek marketing advantage through product certification programs that include specification or recommendation of BMPs. Research scientists and their outreach counterparts facilitate the transfer of BMPs and adoption by producers. Monitoring programs established as a partnership between producers and research scientists can document the performance and effectiveness of BMPs. Seafood buyers are broadly interested in seafood that is produced sustainably. To the extent that BMP programs increase the supply of certified seafood, buyers support these programs. Reflecting consumer concerns, nearly all seafood buyers rank food safety among their top concerns, with other quality attributes, including sustainability, of far lower importance. Regulators use BMP programs as a proxy to achieve compliance with environmental and other regulations (Chapter 15). As is the case with the Florida program, participation in the program is a condition of license or permit to operate. Some regulatory agencies recognize that broad application of BMP programs (including watershed protection through agriculture and forestry BMP programs) can reduce nonpoint source pollution and other environmental impacts, thereby meeting environmental protection goals. The relatively low cost of administering such a program may be desirable for regulatory agencies, especially during periods of budget shortfalls, rather than supporting more costly regulatory and monitoring programs.

Limitations of BMPs BMPs are best applied at the farm scale to address reversible, near-field environmental impacts. At the farm level, BMPs must be adapted to site-specific characteristics and conditions. For this reason, general BMPs, such as those found in many codes of conduct, have

Molluscan shellfish aquaculture 63

limited utility because they lack sufficient detail and specificity. To be most effective, producers must adapt BMPs to the specific characteristics of their operation. For a given impact, the selection and implementation of BMPs will depend on the species cultured, the method of culture, and the scale of the operation. BMPs are often used as a proxy for performance. It is assumed, often on the basis of common sense, that the environmental performance resulting from certain practices is better than that of others. The environmental performance resulting from implementation of a certain practice is not predictable or guaranteed. There are large uncertainties associated with the environmental effects of BMPs. The effectiveness of BMPs is often assumed, but monitoring is required to verify performance. Comparing the environmental performance of conventional with improved practices has not been documented with sufficient rigor. Few formal studies have been conducted to compare the environmental effects of standard with improved culture practices. Specific BMPs are recommended in response to specific impacts of operation. Thus, some BMP programs are prescriptive in their use of BMPs. This approach encourages compliance, not innovation. Alternatively, if performance standards are established, producers have considerable flexibility in achieving that performance standard. In the case of performance standards, the focus is on the end result of improved performance and reduced impacts. In the case of prescriptive BMP programs, the focus is on the practices themselves, not the outcome. Setting performance standards gives producers considerable flexibility in developing innovative methods to achieve the standard. Over time, the standard can be adjusted in consideration of changes in uncertainty and to stimulate further improvement. However, from the standpoint of government regulators and auditors of ecolabeling standards programs, verifying BMPs is much easier than

verifying performance standards, for which there is a substantial requirement and cost for environmental monitoring. One limitation of BMPs is that adoption is largely voluntary and therefore BMPs are not universally adopted among producers within an aquaculture sector. BMPs may be adopted selectively, especially those that are known or perceived to be low cost or that are merely sufficient to achieve regulatory compliance. Most aquaculture producers, especially owners of small-scale operations, are notoriously poor record-keepers. Therefore, verification of BMP performance through self-evaluation should be considered suspect. Independent verification of environmental performance, such as is required through many ecolabeling or product certification schemes, is costly. The BMP approach often does not work well for small-scale producers, who often lack the technical knowledge to implement BMPs. On the other hand, BMP programs managed by producer organizations, with the assistance of research scientists and their extension service counterparts, can be an effective means of disseminating improved technology and practices to small-scale producers. Nonetheless, larger-scale producers are often in a better position to invest in technologies and implement practices to improve environmental performance, and thus are the chief beneficiaries of certification programs based on BMP implementation. Many important environmental effects of shellfish aquaculture occur at embayment level or regional scales. In places with many shellfish aquaculture facilities, embayment or regional agreements and zoning are more effective policy tools to address ecological carrying capacity or other local concerns. Impacts related to introduction of potentially invasive shellfish species can be addressed by BMPs only in part. Regional, governmental, or intergovernmental agreements and regulations are a more effective approach to deal with largerscale and longer-term environmental impacts.

64

Shellfish Aquaculture and the Environment

It is important to emphasize that BMPs are not a panacea for all problems. Some problems or effects are intractable or irreversible. In such cases, BMPs may contribute to the solution, but the main source of the problem or impact is best addressed through other mechanisms.

Assurance labeling With respect to cultured seafood, the focus of government regulators is on food safety and environmental protection, especially water quality. In the United States, the Food and Drug Administration is responsible for the safety of the food supply and the Environmental Protection Agency regulates water quality through permitting and monitoring programs. Despite the activities of these programs, periodic failures of the food safety system have led to consumer fear and uncertainty about consuming certain food products, including shellfish, and undermined trust and credibility in government regulatory mechanisms (GAO 2001). Some citizens and NGOs are also concerned about the capacity and willingness of government regulators to meet environmental protection goals. This segment of the public is increasingly willing to accept so-called “nonstate, market-driven environmental governance” in the form of ecolabeling programs, where responsibility for environmental governance is shifted from “the State” to marketbased actors, especially large transnational grocery chains and environmental NGOs (Cashore 2002). It is, in essence, the privatization of global environmental governance (Clapp 1998). Businesses increasingly prefer to adopt voluntary environmental standards rather than submit to mandatory government regulations. Ecolabels are seals of approval given to products that are deemed to have fewer negative impacts on the environment than functionally or competitively similar products

(Deere 1999). They provide a market-based incentive for responsible production practices by providing a price premium. Participation in ecolabeling programs is voluntary. An ecolabel intends to communicate assurance to consumers that products conform to certain standards, specifically standards for environmentally responsible production. Siggs (2007) suggests that market-based quality schemes, certification, organic labels, ecolabels, and retailer specifications or procurement policies should be collectively described as assurance labels. There are three categories of ecolabeling programs based primarily on the method of conformity assessment (Deere 1999). Firstparty labeling schemes are established by companies that assess their own conformity with standards. Second-party labeling schemes are established by producer associations for their members. Standards are developed by the association, with some outside input from aquaculture experts or representatives of environmental NGOs. Conformity assessment is conducted by the producer group or by contracting external certifiers. Independent, thirdparty labeling schemes are established by an organization, such as an environmental NGO or producer association, that may function as the facilitator of the standards development process. The standards development organization is independent of producers and seafood buyers. Producers are certified by accredited auditors that are independent of the standards setting agency. For most consumers, an ecolabel based on third-party certification is the most credible and preferred. According to the ISO, the word “certification” should be reserved for type III labeling schemes. The ISO has a typology of voluntary environmental performance labeling in the ISO 14020 family of standards:

• Type I ecolabels are a voluntary, multiplecriteria-based, third-party program that awards a license that authorizes the use of environmental labels on products indicating

Molluscan shellfish aquaculture 65

overall environmental preferability of a product within a particular product category based on life cycle considerations. Type I environmental labeling is the outcome of an environmental certification program that results in a seal of approval. • Type II ecolabels are informative environmental self-declaration claims. These are equivalent to the first-party labeling schemes described above. • Type III ecolabels are voluntary programs that provide quantified environmental data of a product, under preset categories of parameters set by a qualified third party and based on life cycle assessment (LCA), and verified by that or another qualified third party. These are environmental declarations or report cards. Despite the formal typologies of labeling schemes, they are incomplete and unnecessarily rigid. The reality of existing labeling schemes is much more complex. A comprehensive typology would consider the originating group, the nature of the standards development process, the focus of the certification scheme, and the auditing process. An ecolabel is obtained by a producer through a certification process based on criteria set through standards. The standards development process has greatest credibility when the process is open to all stakeholders and the proceedings and decision making are transparent. The ecolabeling process begins with the development of standards. Depending on the type of labeling scheme, these standards can be developed by producers, producer associations, or a transparent, multistakeholder process where divergent views are resolved through discussion and negotiation. Once standards are developed, products are certified through an auditing process. Again, depending on the labeling scheme, products can be selfcertified by producers, certified by producer associations, or certified by auditors independent of the standards development organiza-

tion. After a product is certified, it can carry a label or seal of approval indicating that it was produced according to a set of process standards with specification that often include BMPs. Product traceability through the value chain from producer to consumer is the key to the success of any assurance scheme. The ISO defines traceability broadly as the “ability to trace the history, application, or location of that which is under consideration.” In aquaculture, traceability refers to the ability to follow the movement of product from the hatchery through production, processing, and distribution. Traceability provides assurance to consumers that products are safe and produced according to certain standards, as signaled by a label, for example. Traceability also provides producers with a mechanism to demonstrate that products have been produced according to a set of standards. Traceability through a continuous chain of custody allows isolation of the source of a food safety problem. Traceability allows differentiation of labeled products from conventionally produced products, especially in cases where processors or distributors handle multiple products. Recordkeeping is a critical part of traceability. Each unit of production (batch, bag, individual animal) requires an accompanying record to provide evidence of compliance with production standards. Essentially record-keeping is critical to demonstrate traceability and traceability is critical to demonstrate compliance with the standards of ecolabeling programs.

Pressures to participate in certification programs There are several institutional actors that influence the adoption of improved environmental practices by producers (Darnall et al. 2008). These actors exert qualitatively different pressures on producers to adopt an environmental management system or code of conduct,

66

Shellfish Aquaculture and the Environment

and ultimately, participate in ecolabeling programs.

Market pressures Consumer demand for ecolabeled seafood is growing. Some consumers choose to make a political statement about their support for certain environmental protection measures by making an informed decision to consume ecolabeled seafood. However, even among political consumers, food safety and humane treatment of cultured animals are more often indicated as important concerns than environmentally responsible production methods. Nonetheless, some consumers are willing to pay a price premium for ecolabeled seafood. Typically, this amounts to 5–10% more than the price for conventionally produced seafood (Wessels et al. 2001). Increasingly, seafood buyers are playing a critical role in the proliferation of ecolabeling schemes by purchasing certified seafood from producers. The most prominent actors are large transnational supermarket chains, as exemplified by companies such as Wal-Mart, Carrefour, and Tesco. Unilever has partnered with the WWF to create the Marine Stewardship Council. Seafood buyers provide market pressure in the form of an incentive or “pull” for the responsible production of seafood. By creating a market for ecolabeled seafood, these companies provide an incentive for producers to produce seafood in an environmentally and socially responsible way. Ecolabeling efforts for aquaculture products have focused on shrimp and salmon, two relatively high-value products. The shrimp and salmon aquaculture sectors have been the object of critical environmental and consumer advocacy campaigns based on a broad range of real and perceived negative environmental and socioeconomic effects. In part, ecolabeling programs have been adopted as a response by seafood buyers to source shrimp and salmon

from producers adhering to practices that improve the nature and extent of environmental impacts. In contrast and in relative terms, shellfish aquaculture is seen as environmentally benign, and thus the market pressures on seafood buyers to source certified shellfish are not as intense as on buyers of shrimp and salmon. There is considerable uncertainty about the market demand for ecolabeled shellfish. In general, the market demand for shellfish is far weaker than that for shrimp and salmon, especially in markets in developed countries. With molluscan shellfish, species, culture method (e.g., moules de bouchot), meat fullness, and local provenance are more important as the basis for labeling or branding than sustainability per se. More importantly, food safety is the primary concern of shellfish consumers, especially for those products consumed raw. Specific concerns about food safety in shellfish include the presence of pathogens leading to foodborne illness and persistent bioaccumulative toxins that are ubiquitous in the environment.

Social pressures Numerous social groups provide social pressures in the form of a “push” for the responsible production of seafood. These groups include environmental NGOs, producer associations, and community groups concerned with local development issues. Many of these groups claim to speak on behalf of the public trust and consumers. Environmental NGOs advocate for broad goals of resource conservation and environmental protection and view environmental management systems and ecolabeling programs as a mechanism to achieve those goals. Environmental NGOs have been very active in advocating BMPs and the incremental and continuous improvement in the environmental performance of aquaculture. The WWF is an example of an environmental

Molluscan shellfish aquaculture 67

NGO that is actively engaged with the producer community and other stakeholders.

Regulatory pressures Government regulators have a mandate to protect environmental quality. Environmental management systems and the ecolabeling programs based on them are a way to achieve environmental protection goals. In the United States, with the current economic downturn and limitations in funding for enforcement of environmental laws, particularly at the local level, environmental management systems are seen as a way for producers to achieve regulatory compliance without the financial burden of monitoring falling to state regulators.

Ownership pressures Some aquaculture producers, especially larger companies with multiple farm locations, have embraced the notion that production in an environmentally responsible way is an ethical issue. It is simply the right thing to do. Shellfish producers in particular are intimately embedded in supporting ecosystems and understand how external factors are forcing environmental changes that threaten their livelihood and business survival. These companies are part of a broader movement in the business community called corporate social responsibility (CSR). Although not available as a standard for third-party verification, the ISO 26000 CSR standard provides guidance to businesses interested in pursuing the “triple bottom line” of people, planet, and profit.

Perspectives on ecolabeling Producers Through ecolabeling programs, producers seek to obtain a marketing advantage by

selling shellfish at a premium price relative to shellfish produced conventionally. Alternatively, producers of ecolabeled shellfish may not obtain a price premium but may gain access to markets that are not available to conventional producers. In many ecolabeling programs, the process standard often includes guidance about sustainable production methods that include specific BMPs. Conventional producers are concerned about efforts to develop ecolabeling programs for shellfish because they consider their production practices to be sustainable as currently conducted and are concerned that ecolabeling will be a condition of market access. They are concerned that, in contrast to ecolabeled shellfish, consumers will perceive conventionally cultured shellfish as unsustainable. On a related point, small-scale producers are unable or unwilling to bear the costs associated with certification and ecolabeling programs. The costs of participation in these programs are well defined and fall exclusively to the producer but the value or benefits of price premiums are shared with seafood buyers. Producers also view ecolabeling and other product certification programs as a way to simultaneously gain market advantage and satisfy compliance with regulatory requirements related to environmental performance. In both cases, producers may adopt BMPs selectively, choosing those that satisfy the requirements of certification programs and government regulations.

Consumers Consumers do not have a clear understanding of the concept of sustainability. Furthermore, a majority of consumers believe that sustainable product claims are a marketing tactic. Consequently, identification of specific seafood ecolabels is weak, with only the MSC label having any recognition by consumers (BBMG

68

Shellfish Aquaculture and the Environment

2009). The USDA Organic label is also recognized by consumers, although there are no ecolabeled shellfish certified under USDA organic standards because they remain in development. There are currently no MSCcertified cultured shellfish (although several areas of culture-based fisheries are under consideration) on the market. There are numerous efforts under way to develop ecolabels for seafood produced in aquaculture. There is certainly a risk that consumers may become confused about the proliferation of ecolabels, which is especially troublesome given that many of the efforts have converged on essentially the same set of concerns and standards. There is now competition among certification schemes, and those that educate consumers about sustainability and effectively communicate the value of their label are most likely to prevail. Ecolabeled seafood is an example of a credence good. Credence goods are a product type where the quality is unknown after purchase and consumption. An ecolabel and the associated production method is a so-called “nonverifiable expert property,” which means that it is impossible for consumers to detect product attributes such as a sustainable production process (as well as food safety and nutritional value). The ecolabel signals to consumers that the product is safe and was produced sustainably. Consumers place trust in producers of seafood carrying an ecolabel and the seafood buyers that market ecolabeled products. The consumer places trust in the parts of the value chain that are “upstream” of consumers. This trust relationship is asymmetric because the producer, processor, and buyer hold more information about the product than the consumer. The label is intended to convey information to consumers about the way seafood is produced. Breeches in trust can be extremely damaging to the integrity of the ecolabel and the organizations that support it.

Aquaculture certification programs The FAO Guidelines for Aquaculture Certification establishes four areas of “minimum substantive criteria” for any aquaculture certification program (FAO 2010). These include animal health and welfare, food safety and quality, environmental integrity, and social responsibility. Although, strictly speaking, BMPs apply directly to the environmental quality area, BMP-like approaches can be applied to address concerns associated with the other areas. For example, standard operating procedures are used in the Hazard Analysis and Critical Control Point (HACCP) program to assure food safety and ISO CSR guidelines can be applied to inform socially responsible practices and activities.

Animal health and welfare The main source of guidance for animal health and welfare is the World Organisation for Animal Health (OIE) Aquatic Animal Health Code (www.oie.int), the FAO Code of Conduct for Responsible Fisheries (CCRF) Technical Guidelines on Health Management for Responsible Movement of Live Aquatic Animals (FAO 2007), and the International Council for the Exploration of the Sea (ICES) Code of Practice on the Introductions and Transfers of Marine Organisms (ICES 2005). Limiting spread of pathogens with shellfish seed requires ongoing vigilance. To the extent that implementation of environmental BMPs reduces stress on cultured animals, BMPs can contribute to the maintenance and improvement of animal health and welfare.

Food safety and quality The FAO/World Health Organization (WHO) Codex Alimentarius is the main source of guidance for food safety and quality. The risk of contamination of a culture site is

Molluscan shellfish aquaculture 69

largely related to the type and extent of land-based sources of pollutants. Watershed protection BMPs for agriculture and forestry can reduce the risk of site contamination associated with runoff. Implementation of standard operating procedures for wastewater treatment plants and other industrial facilities can also reduce the risk of site contamination. The key to food safety is product traceability through the value chain. For shellfish (with the notable exception of New Zealand greenlipped mussels), value chains are often short, with local or regional markets predominating over global distribution networks. Traceability with shellfish is straightforward and well established because of long-standing public health concerns associated with consuming raw shellfish. In the case of molluscan shellfish, HACCP through the value chain and, in the United States, the National Shellfish Sanitation Program are responsible for assuring the public health of shellfish consumers. Product handling according to HACCP guidelines explicitly requires record keeping and product labeling to assure traceability in the event of the need for a product recall.

Environmental integrity The FAO uses the term environmental “integrity” but does not define it. Integrity can refer to the normative structure and function of ecological communities in particular habitats. Integrity implies intact structure and function and emphasizes the role of resilience, or the ability of ecosystems to withstand shocks and stress. Environmental standards emphasize the importance of site selection, including effects on sensitive habitats. Of importance to shellfish aquaculture, environmental standards should consider the cumulative effect of multiple farms. An aquaculture certification program should consider the key environmen-

tal impacts and develop and implement BMPs to address them. The main environmental impact types include biodiversity, genetic issues, invasive species, and water quality. BMPs to safeguard biodiversity in a humandominated coastal environment will be very different from those for an operation sited in a relatively pristine environment with a high abundance and diversity of native shellfish and other species. BMPs for concerns about genetic interactions with native shellfish will focus on the use of hatchery seed, using native species or low-risk nonnative species, and methods to minimize escape or loss. BMPs to address solid wastes (e.g., lost gear) and responsible use of energy and other resources are also appropriate for shellfish aquaculture.

Social responsibility The FAO suggests that social responsibility be included in any aquaculture certification program. Standards for social responsibility consider worker safety and welfare, compliance with child labor laws, and general effects and participation in local communities. Aquaculture businesses are embedded in communities that are affected by their presence. People in those communities have a reasonable expectation that aquaculture operations will not have undesirable effects on the local quality of life. Workers from those communities may be employed at the operation and expect a fair wage. The business might have investors that provide financial capital and technology and expect a reasonable return on investment. Consumers trust the producer to provide a safe and nutritious product. As a central guiding principle, all of the people directly connected to an aquaculture business should be treated fairly and responsibly. It is difficult to achieve consensus on measurable and meaningful indicators of socially responsible behavior. Developing BMPs for socially responsible behavior stretches the

70

Shellfish Aquaculture and the Environment

original meaning and intent of that policy tool. CSR guidelines are useful but, like similar codes of conduct, provide only general statements of ethical behavior.

Critique of bivalve shellfish ecolabeling efforts in the United States Compared with other products from capture fisheries or aquaculture, efforts to develop ecolabeling programs for bivalve shellfish lag behind. In part, this is related to a higher level of concern about the deleterious environmental effects of certain forms of aquaculture, especially shrimp farming in coastal ponds and salmon farming in net pens. In part, this is also related to a perception that the environmental effects of shellfish aquaculture are relatively harmless and thus do not require the same level of attention or reform of production practices as shrimp or salmon farming. Shellfish ecolabeling efforts have not progressed to nearly the same degree or extent as ecolabeling of other aquatic products. The GAA has established a Best Aquaculture Practices standard, with certification provided by the Aquaculture Certification Council. The program has resulted in the certification of shrimp, tilapia, and catfish (Ictalurus and Pangasius) farms. Of the assurance schemes that are currently in effect, more products have been certified by the GAA program than any other. To date, there are no bivalve shellfish farms certified according to the GAA Best Aquaculture Practices standards. Shellfish aquaculture, BMPs to minimize environmental impacts, and seafood ecolabeling efforts occur in the context of humandominated coastal ecosystems. Humans are part of ecosystems (for better or usually worse) and a variety of human activities are responsible for observed environmental changes. Certification efforts rarely, if at all, deal with environmental problems and specific effects holistically. Efforts at improving environmen-

tal quality are rarely proportional to the relative contribution of sources of specific impacts. Standards that are proposed for environmental certification of shellfish aquaculture seem likely to provide only marginal improvement in environmental performance relative to conventional production methods. Two shellfish ecolabeling efforts are reviewed here, revealing the unique challenges associated with ecolabeling bivalve shellfish.

The WWF bivalve aquaculture dialogue As part of its broader effort to develop standards for the main cultured species in global trade, the WWF began a Bivalve Aquaculture Dialogue in 2004. The purpose was to engage stakeholders in an open and transparent process “to create measurable, performancebased standards that will help minimize the key environmental and social impacts” of bivalve aquaculture. It was intended that these standards serve as the basis for a certification program similar to the effort for which WWF played a central role to develop the Marine Stewardship Council to certify seafood from capture fisheries. Unquestionably, this is a laudable goal and the WWF-supported aquaculture dialogues for other species, especially shrimp and salmon, are likely to lead to a marked reduction in environmental and socioeconomic impacts of the culture of those animals. Common to many standards development efforts, including the other WWF aquaculture dialogues, the focus is on minimizing near-field impacts. Arguably, the major near-field impacts of bivalve aquaculture—localized depletions of dissolved oxygen and phytoplankton and sediment organic matter accumulation downcurrent of bivalve culture facilities—are reversible, highly localized, and easily addressed by consideration of ecosystem carrying capacity using zoning or bay-level agreements. However, the unit of certification is an individual farm,

Molluscan shellfish aquaculture 71

and thus important carrying capacity issues are only indirectly addressed by the standards because nonparticipating farms are not considered. This issue could be addressed by collectively assessing and monitoring a cluster of farms as the unit of certification. For some of the larger-scale, far-field effects—such as effects on biodiversity, wild shellfish populations, or disease and pest management practices—the standards (WWF 2010) require documentation or evidence of meeting the standard or following BMPs. However, providing evidence and documentation of efforts to mitigate large-scale effects does not necessarily equate to effective reduction in the risk of adverse impacts. Developing indicators for effects and criterion that extend over a spatial scale that is larger than the immediate footprint of the farm is difficult, especially because other shellfish farms in an area and other activities may combine as drivers of ecosystem change. The WWF standards explicitly state that the focus is on environmental and social sustainability. There is no mention of food safety concerns. The FAO Technical Guidelines on Aquaculture Certification indicate that food safety and quality should be considered as one of the minimum substantive criteria of a certification scheme for aquaculture. Food safety is a critical concern of consumers, processers, and producers of a highly perishable product such as bivalve shellfish. Although food safety for shellfish is addressed by government agencies and programs in the developed world, in theory it will be possible to obtain WWF certification for bivalve shellfish cultured in waters with an unacceptable quality rating by government shellfish-safety regulators. The WWF standards include a requirement to adhere to existing laws, which presumably include laws governing shellfish sanitation and food safety, although no explicit mention is made of these. Finally, the Bivalve Dialogue took place over 6 years, yet there was very limited engagement with producers and other stakeholders in

developing countries. Six meetings were held in North America, one in Europe, and one in New Zealand. It was only after the Global Steering Committee of the Bivalve Dialogue had developed draft standards that “outreach” workshops were held in Vietnam, China, and Australia. For context, it is important to emphasize that, in 2008, 85% of the world’s oysters were produced in China, and the AsiaPacific region accounts for 95% of global oyster production (FAO 2010). Although the process has been open and transparent, participation in the WWF Bivalve Dialogue by stakeholders in developing countries has been weak. The course of the WWF Bivalve Dialogue process suggests that the primary beneficiaries of the dialogue process will be producers (and consumers) in developed countries. Regulators and government officials in the Asia-Pacific region are apparently hesitant to participate in the dialogue process on the basis of concern over “social issues,” specifically the potential loss of sovereignty associated with a shift in governance over these issues from the state to standards-setting and certifying bodies.

USDA organic standards Attempts to develop organic standards for aquaculture through the USDA’s National Organic Program have been under way for nearly a decade. Although draft standards for finfish aquaculture have been recommended for approval, the standards development process for organic bivalve aquaculture lags behind and remains in review. Organic standards for cultured shellfish, mostly mussels, have been developed and approved by organizations based in Europe and New Zealand (Table 3.4). However, there is considerable confusion about the terms used for organic labeling in Europe, where “bio,” “eco,” and “organic” are considered synonymous. The establishment of organic standards for aquaculture in the United States has been

72

Shellfish Aquaculture and the Environment

Table 3.4 Examples of assurance schemes that have developed process standards for shellfish aquaculture.

Entity

Country of program origin

BioGro New Zealand

Certification type

Shellfish type/location

New Zealand

Organic (non-IFOAM certified)

Shellfish

BioSuisse

Switzerland

Organic

Mussels; using Naturland standards

Conseil des Apellations Agroalimentaire du Quebec

Canada

Organic

Oysters, mussels

Filière Qualité Carrefour

France

Quality

Oysters

Irish Quality Mussel Program

Ireland

Quality

Mussels

Irish Quality Oyster Program

Oysters

Irish Quality Eco-Mussels

Environmental management

Mussels

KRAV

Sweden

Organic (non-IFOAM certified)

Blue mussels

Label Rouge

France

Comprehensive quality

Oysters and scallops

MSC

various

Sustainable

Suspended mussel culture (pending)

Naturland

Germany

Organic (IFOAM certified)

Rope culture of blue mussels

Safe Quality Food

Australia

Food safety

Oysters in Canada and Chile

Soil Association

United Kingdom

Organic

Bivalves

World Wildlife Fund

United States

Sustainable

Bivalves

Source: Corsin et al. 2007. IFOAM, International Federation of Organic Agriculture Movements.

complicated by the difficulty in accommodating the inherent characteristics of aquatic animal production within the philosophy and principles of organic agriculture. These principles are rooted in the idea that the key to producing organic food is improving soil tilth and fertility, accomplished by adding organic matter. There is no aquatic analogue to this terrestrial model and the addition of organic matter to water is seen as polluting. Soil is a fixed and solid matrix, whereas water is a dynamic fluid. Differences in the inherent properties of soil and water and how they should be managed to produce crops present philosophical difficulties to the organic com-

munity. Thus far, there continues to be stout resistance from many in the terrestrially oriented organic community to accommodate aquatic farming by extending the paradigm of organic agriculture to aquaculture. Resistance to organic certification is also related to the notion of control over production. Uncontrolled or wild nature is not considered organic, even though natural ecosystems are models for organic agroecosystems. Organic agroecosystems are actively managed, not passively natural or wild. The prevailing view in the organic agriculture community is that a facility cannot be “organic by neglect,” that there must be some kind of active control

Molluscan shellfish aquaculture 73

over the production environment, something that is especially true with respect to foods for cultured animals. The lack of control over the natural foods that passively flow around cultured bivalves is problematic. Although some wild crops, notably honey and seaweed, have been certified as organic, holding up these examples has not been sufficiently persuasive as an argument to allow certification of bivalve shellfish aquaculture as organic. The same arguments that have been made to preclude the use of fishmeal and fish oil derived from wild, pelagic forage fish in diets for cultured finfish that are eligible for organic certification have been made to rule out organic certification of bivalve shellfish.

Criticisms of certification programs The proliferation of certification programs has been portrayed as broadly beneficial: The public trust benefits by improved environmental performance of producers, seafood buyers benefit from healthy profit margins on marketing ecolabeled seafood, producers benefit from improved production efficiency and price premiums on certified product, and consumers benefit by access to and consumption of a health-supporting product that satisfies a psychological desire to “help the planet” by consuming seafood produced sustainably. Beyond these overly simplistic claims, the proliferation of certification schemes has unintended consequences and negative features.

Beneficiaries of ecolabeling The main beneficiaries of ecolabeling programs are the high-volume seafood buyers in the form of large transnational grocery chains. These chains are increasingly buying seafood from certified sources. Although producers bear the costs of making the transition to more sustainable practices and product certification,

the benefits are shared between seafood buyers and producers. This reinforces the existing power relationships in the seafood supply chain where seafood buyers dominate producers by setting price. Among producers, large-scale producers have more resources to commit to participation in certification programs than small-scale producers. Therefore, the benefits of such programs may accrue disproportionately to largescale producers. Finally, support for certification programs by relatively wealthy and educated consumers within developed countries is stronger than in developing countries. These wealthy consumers presumably obtain some nutritional benefit from consuming certified fish but poor consumers in developed countries obtain no such benefit. In general, the main beneficiaries of ecolabeling programs are transnational grocery chains and other seafood buyers, large-scale producers who can bear the cost of certification, and wealthy consumers in developed countries.

Small-scale producers and barriers to trade Most aquaculture producers in the world operate on a small scale in rural areas with limited resources. This is especially true in developing countries. It also applies to shellfish producers globally. The participation in markets for certified seafood by small-scale producers, particularly in developing countries, is extremely limited. These producers lack access to technical knowledge (e.g., BMPs) to make the transition to more sustainable practices. Small-scale producers also lack the financial resources to invest in more efficient technologies. The cost of certification is a significant barrier for participation in ecolabeling programs by small-scale producers in developing countries.

74

Shellfish Aquaculture and the Environment

Nearly all of the market for certified seafood is in developed countries. Ecolabels can restrict access to these markets by small-scale producers in developing countries. On a global macroeconomic scale, the certification system might have the unintended consequence of functioning as a nontarriff trade barrier. This is a concern but it is difficult to demonstrate because there is a lack of hard information about trade flows of ecolabeled seafood. Involving small-scale producers in certification programs will require special efforts. Organizations such as FAO and the Network of Aquaculture Centres in Asia-Pacific (NACA) recommend that small-scale producers be certified as a collective “cluster” where a group of farms sharing the same water resources would be the unit of certification. This approach is actually more broadly applicable because it takes account of ecosystem carrying capacity considerations, unlike programs where the unit of certification is a single farm. Another approach that may benefit small-scale producers is the development of CSR programs by seafood buyers that explicitly involve small-scale producers in meeting the supply requirements for certified seafood.

Ecolabels and information asymmetry As mentioned previously, consumer understanding of the meaning of sustainability is rather superficial and confused. Ecolabels do not adequately communicate information to consumers about the production process and product quality, a feature described as information asymmetry (van Amstel et al. 2008), which describes the gap between sellers and buyers regarding their understanding of the environmental attributes of a labeled product. Labeled products are examples of credence goods, where product attributes cannot be ascertained prior to purchase. Consumers do not observe the production process and therefore do not have a basis for

believing the process claims made by the label. The ecolabel embodies a trust relationship between producer and consumer. To be effective, consumers must understand the meaning of the label. Consumers must also find the label credible. Thus, an ecolabel that is a producer claim will have much different credibility among consumers from an ecolabel based on standards developed through a transparent and inclusive process and certified through a third party.

Proliferation of ecolabels and consumer confusion As of 2009, there were more than 400 labels and certification programs related to environmental attributes of products (BBMG 2009). In part, the proliferation of these programs is a response to the financial incentives associated with labeling. There are no checks and balances on the proliferation of ecolabels. With label proliferation, consumers have become overloaded with information and confused about the meaning, significance, and legitimacy of ecolabels and seals. When consumers were asked if they were familiar with particular seals or labels, 89% had never seen the Marine Stewardship Council label and only 10% made a decision to purchase seafood based on the presence of the label. The USDA organic seal was familiar to 62% of consumers. This suggests that consumer awareness of seafood ecolabels is rather weak and superficial. It also suggests that consumers are not sure what the label signifies. For seafood, especially in Europe, there are organic labels (e.g., Naturland, KRAV), quality labels (e.g., GlobalGAP, Label Rouge), and environmental sustainability labels (e.g., Marine Stewardship Council). Only savvy, well-informed consumers are able to differentiate among products certified according to the different standards. In aquaculture, at least two environmental certification schemes are in competition. One

Molluscan shellfish aquaculture 75

is the industry-supported program developed by the GAA and the other is an NGOsupported program by the WWF. Differences between standards developed by the GAA and the WWF Aquaculture Dialogue process are rather minor, despite the different pathways followed in the development of the standards. To date, there have been no aquaculture products certified according to the standards developed through the WWF dialogues. In contrast, significant quantities of seafood from aquaculture have been certified by the Aquaculture Certification Council according to the GAA Best Aquaculture Practice standards. Darden Foods, a major supporter of the GAA effort, only buys shrimp certified according to the GAA standards. Wal-Mart buys seafood that is certified by the Marine Stewardship Council (wild product) or the Aquaculture Certification Council (cultured product). There is a clear need to harmonize the existing standards.

Towards more meaningful labeling Comprehensive quality The WWF and USDA organic certification schemes described previously are examples of ecolabeling programs that focus rather narrowly on environmental considerations. These labels represent claims made about the production process, that the seafood was produced according to a set of environmentally favorable production standards. The label makes no claim about seafood safety, a major concern of consumers and seafood buyers. Seafood safety and production method are examples of intrinsic quality characteristics known as credence attributes. Credence attributes are not known even after purchase and consumption. Extrinsic product attributes such as labels or seals can communicate assurance to consumers that products are safe or produced in such a way to minimize environ-

mental impacts. Labeling can change credence attributes to search attributes, which are the intrinsic quality attributes that can be evaluated before purchase. Consumers are interested in a range of product quality attributes in making a purchasing decision (Fig. 3.1). Thus, for shellfish, consumers might be interested in intrinsic quality attributes such as safety, freshness, taste, and environmentally sustainable production methods, as well as extrinsic quality attributes such as price and the producer’s reputation or brand. To meet consumer demand for seafood with these product attributes requires certification and labeling programs that consider a more comprehensive definition of quality than certification schemes that focus narrowly on environmental and social considerations. The point is to consider the totality of the features and characteristics that comprise quality and to use consumer requirements as a starting point for developing certification programs. Under so-called total quality management (TQM) programs, specifications are made by seafood buyers and standards are specific to the market. Safety is an essential and critical part of a TQM system for seafood, especially shellfish. Although TQM programs focus on production, marketing and business policy are also included. TQM explicitly involves the workforce by making each worker responsible for all stages of the production cycle. Similar to ISO environmental management systems, TQM programs embrace continuous systematic improvement of processes and operations, including adapting and refining BMPs. One example of a comprehensive quality certification program in aquaculture is the Label Rouge program as applied to oysters cultured in the Marennes Oléron region of southwest France since 1989. Oysters certified by the program must meet very stringent organoleptic quality criteria, such as taste, odor, saltiness, texture, and absence of “milk”; high-quality appearance, including a regular,

76

Shellfish Aquaculture and the Environment

Quality Attributes (Intrinsic)

Food Safety Attributes Foodborne pathogens Bioaccumulative toxins Pesticide residues Other aquatic contaminants Food additives, preservatives Physical hazards (e.g., grit) Spoilage Irradiation Nutritional Attributes Calories Fat and cholesterol Sodium and minerals Carbohydrate and fiber Protein Vitamins Sensory/Organoleptic Attributes Taste and aftertaste Color Appearance Freshness Firmness Smell/aroma Value/Function Attributes Compositional integrity Size (meat-to-shell ratio) Style Preparation/convenience Packaging materials Shelf life

Quality Indicators and Cues (Extrinsic)

Test/Measurement Indicators Quality management systems Certification Records Labeling Minimum quality standards Occupational licensing Cues

Price Past purchase experience Processor name Reputation Brand Store name Packaging Advertising Country of origin Distribution outlet Warranty Other information provided

Process Attributes Traceability Place of origin (provenance) Biotechnology (GMOs) Organic Environmental impact Worker safety

Figure 3.1 Intrinsic quality attributes and extrinsic quality indicators and cues for molluscan shellfish. (Adapted from Caswell 2006.)

deeply cupped shell, green meat color, minimum size, and high shell fullness; specifications on duration and density in finishing ponds (claires); and other husbandry and environmental practices. There are requirements for transport, processing, and maximum shelf life. The label includes a rigorous traceability program to assure food safety. The label is

regulated by the French Ministry of Agriculture and Fisheries.

Including LCAs Two of the more pressing environmental problems facing humanity that have potential

Molluscan shellfish aquaculture 77

Table 3.5 Examples of life cycle assessment-derived impacts of examples of extractive (mussel) and fed (salmon net pen, salmon recirculating system) aquaculture systems. Units of impact categories are equivalents per 1000 kg live weight. System

Eutrophication potential

Global warming potential

Acidification potential

Mussel Salmon net pen Recirculating system

0.5 35 12

400 2,073 10,300

5 18 63

Sources: Ayer and Tyedmers 2009; Lozano et al. 2009; Iribarren et al. 2010.

negative implications for shellfish aquaculture are global climate change and ocean acidification. An LCA is a method to provide a holistic accounting of bioresource flows and globalscale environmental impacts associated with providing a product to society. The assessment is not restricted solely to the production process, but includes the full value chain and considers the impacts of bringing products from the farm to the consumer. An LCA is an analytical tool to evaluate large-scale global impacts, including global warming potential, acidification potential, eutrophication potential, marine ecotoxicity, energy use, and biotic resource use. Total energy use per unit output is a good surrogate for overall impact and can be used to compare methods of protein production. Given that shellfish aquaculture is a form of extractive aquaculture, an LCA of shellfish aquaculture is quite favorable in comparison with fed aquaculture in most impact categories (Table 3.5). In many cases, the production phase has a relatively minor effect relative to other parts of the value chain, such as seed production in hatcheries, processing, cold storage, transport, and distribution. For example, a study of the mussel aquaculture sector in Galicia, Spain, indicated that onshore processing (e.g., washing, grading, packaging) made the largest contribution to global environmental impacts (Iribarren et al. 2010). In such cases, efforts that focus on the production phase as a means of reducing larger-scale environmental impacts are misplaced and the focus should be shifted elsewhere in the value chain.

The LCA is a useful tool to measure “hot spots” of environmental impact or resource use inefficiencies in the value chain that can be addressed through BMPs or other management approaches. Nearly all certification schemes emphasize mitigation of farm-level effects and near-field impacts, with some consideration of regional impacts, such as effects on biodiversity and the effects of introduction of pathogens and invasive species. Larger-scale global impacts are rarely considered in certification programs. The LCA can be the basis for a more meaningful certification. One example is the Swedish organic certification program established by KRAV, which explicitly includes life cycle considerations. The ISO 14040 standard for LCA considers environmental impacts throughout a product’s value chain. For shellfish, this may include hatchery production of seed, growout, processing (including depuration), packaging, refrigerated storage, and transport. Significant energy use and other environmental impacts may occur after the production step. Furthermore, shellfish are highly perishable and significant losses may occur through the value chain from spoilage, further adding to environmental costs.

Concluding remarks BMP programs have excellent potential to improve the environmental performance of aquaculture and accelerate the adoption of

78

Shellfish Aquaculture and the Environment

better technology and practices by producers. The current trend suggests that BMP programs will become more widespread and extend to producers of cultured seafoods who have not yet had pressures or incentives to change. BMPs are not a panacea for reducing the negative environmental effects of aquaculture; a variety of policy options are needed. At the farm level, BMPs incorporated as part of an environmental management system or code of conduct can address the most important nearfield effects. At the embayment level, performance standards or zoning are likely to be the most effective policy approaches to resolve ecosystem carrying capacity issues. At the regional or international level, negotiated agreements may be the most effective mechanism to address nonnative species and disease transmission issues. The focus of BMP programs and sustainable product certification will continue to emphasize seafood items that are traded as global commodities. Products marketed locally are less likely to be certified but nonetheless may be produced following guidelines in voluntary BMP programs or codes of conduct established by governments or producer associations. With the possible exception of mussels, most cultured bivalves are marketed locally or regionally and there is thus less pressure to implement BMP programs that lead to product certification. For shellfish, the overriding consideration in product certification programs is rightly focused on food safety because this is the principal concern of seafood consumers. Sustainable production methods are important to consumers, but are much lower priority. Meaningful certification should take a broad view of the elements of product quality, with special emphasis on food safety. Product certification for sustainability in aquaculture is a work in progress. Although many such efforts are flawed, they represent good-faith efforts to improve the status quo and provide incentives to change producer

behavior. Product certification mostly benefits large-scale operations that have the resources to access the technology and knowledge required to improve performance and to pay for conformity assessment. However, most shellfish producers operate at a small scale and many uncertainties remain about their capacity to fully participate in certification programs. Considerable uncertainty also remains about the price premium, market access, or other value that can be obtained by producers of ecolabeled bivalve shellfish. Although consumers desire seafood that has been produced sustainably, they are wary of certification and are confused by the proliferation of ecolabels. Consumers think certification is mostly a marketing ploy or at worst a mechanism to “greenwash” products that are not produced sustainably. Consumer trust in labels and the reputation of seafood buyers is fragile. Given the current proliferation of ecolabels, it seems reasonable to assume that there will be a shakeout among competing labels, especially because many of these labels have converged on similar standards. The winning label might not be the best in terms of components of the standard or the degree of transparency in standards development, but is likely to be the best in terms of label marketing and acceptance by seafood buyers.

Literature cited Aquaculture New Zealand. 2007. New Zealand Greenshell Environmental Code of Practice. Aquaculture New Zealand, Nelson, New Zealand. Association of Scottish Shellfish Growers. 2005. Code of Good Practice. Association of Scottish Shellfish Growers, Connel, Scotland, UK. Ayer, N.W., and Tyedmers, P.H. 2009. Assessing alternative aquaculture technologies: life cycle assessment of salmonid culture systems in Canada. Journal of Cleaner Production 17: 362–373.

Molluscan shellfish aquaculture 79

Bantry Bay Aquaculture. 2009. Bantry Bay Aquaculture Code of Practice. Bantry Bay Aquaculture, Bantry Bay, Ireland. BBMG. 2009. Conscious Consumer Report: Redefining Value in A New Economy. BBMG, New York. BCMAFF (British Columbia Ministry of Agriculture, Food, and Fisheries). 2002. BC Shellfish Aquaculture Code of Practice. BCMAFF, Vancouver, BC. BCSGA (British Columbia Shellfish Growers Association). 2001. Environmental Management System Code of Practice. British Columbia Shellfish Growers Association, Duncan, BC. BIM (Bord Iascaigh Mhara—Irish Sea Fisheries Board). 2003. Ecopact—Environmental Code of Practice for Irish Aquaculture Companies and Traders. Irish Sea Fisheries Board, Dublin, Ireland. Cashore, B. 2002. Legitimization and the privitization of environment governance: how non-state market-driven (NSMD) governance systems gain rule-making authority. Governance 15:503– 529. Caswell, J.A. 2006. Quality assurance, information tracking, and consumer labeling. Marine Pollution Bulletin 53:650–656. Clapp, J. 1998. The privatization of global environmental governance: ISO 14000 and the developing world. Global Governance 4:295–316. Clay, J. 2008. The role of better management practices in environmental management. In: Tucker, C.S., and Hargreaves, J.A. (eds.), Environmental Best Management Practices for Aquaculture. Blackwell Publishing, Ames, IA, pp. 55–72. Corsin, F., Funge-Smith, S., and Clausen, J. 2007. A Qualitative Assessment of Standards and Certification Schemes Applicable to Aquaculture in the Asia-Pacific Region. Asia-Pacific Fishery Commission, FAO Regional Office for Asia and the Pacific, Bangkok, Thailand. Creswell, R.L., and McNevin, A.A. 2008. Better management practices for bivalve molluscan aquaculture. In: Tucker, C.S., and Hargreaves, J.A. (eds.), Environmental Best Management Practices for Aquaculture. Blackwell Publishing, Ames, IA, pp. 427–486. CSTT (Comprehensive Studies Task Team). 1997. Comprehensive studies for the purposes of

Article 6 & 8.5 of DIR 91/271 EEC, the Urban Waste Water Treatment Directive, 2nd edition. Marine Pollution Monitoring Management Group, Comprehensive Studies Task Team. Dept. of the Environment, Northern Ireland, Environment Agency, Scottish Environment Protection Agency and Water Services Association, 13 January 1997. Darnall, N., Henriques, I., and Sadorsky, P. 2008. Do environmental management systems improve business performance in an international setting? Journal of International Management 14:364– 376. Deere, C. 1999. Eco-Labelling and Sustainable Fisheries. IUCN—The World Conservation Union and the Food and Agriculture Organization of the United Nations (FAO), Washington and Rome. Engle, C.R., and Wossink, A. 2008. Economics of aquaculture better management practices. In: Tucker, C.S., and Hargreaves, J.A. (eds.), Environmental Best Management Practices for Aquaculture. Blackwell Publishing, Ames, IA, pp. 519–551. Environment Protection Authority. 2005. Code of Practice for the Environmental Management of the South Australian Oyster Farming Industry. Environment Protection Authority, Adelaide, Australia. FAO (Food and Agriculture Organization of the United Nations). 2007. Aquaculture Development. 2. Health Management for Responsible Movement of Live Aquatic Animals. FAO, Rome, Italy. FAO (Food and Agriculture Organization of the United Nations). 2010. 2008 Fishery and Aquaculture Statistics. FAO, Rome, Italy. FDACS (Florida Department of Agriculture and Consumer Services). 2007. Aquaculture Best Management Practices Rule, January 2007. DACS-P-01499. Florida Department of Agriculture and Consumer Services, Division of Aquaculture. Tallahassee, FL. Flimlin, G., Macfarlane, S., Rhodes, E., and Rhodes, K. 2010. Best Management Practices for the East Coast Shellfish Aquaculture Industry. USDANIFA, NRAC, NOAA. GAO (General Accounting Office). 2001. Report to the Committee on Agriculture, Nutrition, and Forestry, U.S. Senate. Food Safety–Federal

80

Shellfish Aquaculture and the Environment

Oversight of Shellfish Safety Needs Improvement. GAO-01-702. United States General Accounting Office, Washington, DC. ICES (International Council for Exploration of the Sea). 2005. ICES Code of Practice on the Introductions and Transfers of Marine Organisms 2005. ICES, Copenhagen. Iribarren, D., Moreira, M.T., and Feijoo, G. 2010. Revisiting the life cycle assessment of mussels from a sectorial perspective. Journal of Cleaner Production 18:101–111. ISO/IEC (International Organization for Standardization, International Electrochemical Commission). 2004. ISO/IEC Guide 2— Standardization and Related Activities—General Vocabulary. ISO/IEC, Geneva, Switzerland. Jensen, G.L., and Zajicek, P.W. 2008. Best management practice programs and initiatives in the United States. In: Tucker, C.S., and Hargreaves, J.A. (eds.), Environmental Best Management Practices for Aquaculture. Blackwell Publishing, Ames, IA, pp. 91–128. Kelly, J., and Maguire, C.M. 2009. Marine aquaculture code of practice: draft. Prepared for NIEA and NPWS as part of Invasive Species Ireland. Leavitt, D.F. (ed.). 2009. Best Management Practices for the Shellfish Culture Industry in Southeastern Massachusetts, Version 09-04a. SouthEastern Massachusetts Aquaculture Center, Bristol, RI. Lozano, S., Iribarren, D., Moreira, M.T., and Feijoo, G. 2009. The link between operational efficiency and environmental impacts. A joint application of life cycle assessment and data envelopment analysis. Science of the Total Environment 407:1744–1754. Maine Aquaculture Association. 2002. Recommended code of practice for aquaculture in Maine. Maryland Aquaculture Coordinating Council. 2007. Best management practices, a manual for Maryland aquaculture, July 2007. MEA (Millenium Ecosystem Assessment). 2005. Ecosystems and Human Well-Being: Synthesis. Island Press, Washington, DC. Nash, C.E., ed. 2001. The net pen salmon farming industry in the Pacific Northwest. NOAA Technical Memorandum NMFS-NWFSC-49.

Silver Springs, MD: National Oceanic and Atmospheric Administration. NRC (National Research Council). 2010. Ecosystem Concepts for Sustainable Bivalve Mariculture. Committee on Best Practices for Shellfish Mariculture and the Effects of Commercial Activities in Drakes Estero, Pt. Reyes National Seashore, California. National Academies Press, Washington, DC. Oesterling, M.J., and Luckenbach, M. 2008. Best Management Practices for the Virginia Shellfish Culture Industry. VIMS Marine Resource Report Number 2008-10. PCSGA (Pacific Coast Shellfish Growers Association). 2001. Environmental Policy. Pacific Coast Shellfish Growers Association, Olympia, WA. PCSGA (Pacific Coast Shellfish Growers Association). 2002. Environmental Codes of Practice for the Pacific Coast Shellfish Industry, June 2002. Pacific Coast Shellfish Growers Association, Olympia, WA. Seafood Shetland. 2007. Code of Practice for Shetland’s Shellfish Growers. Seafood Shetland, Lerwick, Shetland, UK. Shumway, S.E., Davis, C., Downey, R., Karney, R., Kraeuter, J., Parsons, J., Rheault, R., and Wikfors, G. 2003. Shellfish aquaculture—in praise of sustainable economies and environments. World Aquaculture 34(4):8–10. Siggs, M. 2007. Consumer assurance: market-based quality schemes, certification, organic labels, ecolabelling, retailer specifications. In: Arthur, R., and Nierentz, J. (eds.), Global Trade Conference on Aquaculture. FAO Fisheries Proceedings 9. FAO, Rome, Italy, pp. 89– 108. van Amstel, M., Driessen, P., and Glasbergen, P. 2008. Eco-labeling and information asymmetry: a comparison of five eco-labels in the Netherlands. Journal of Cleaner Production 16:263–276. Wessels, C.R., Cochrane, K., Deere, C., Wallis, P., and Willman, R. 2001. Product Certification and Ecolabelling for Fisheries Sustainability. FAO Fisheries Technical Paper 422. FAO, Rome. WWF (World Wildlife Fund). 2010. Bivalve Aquaculture Dialogue Standards. World Wildlife Fund, Inc., Washington, DC.

Chapter 4

Bivalve filter feeding: variability and limits of the aquaculture biofilter Peter J. Cranford, J. Evan Ward, and Sandra E. Shumway

Introduction A fundamental knowledge of bivalve feeding behavior is a minimum requirement for understanding how aquaculture interacts with the surrounding ecosystem. The potential environmental effects and ecological services of bivalve culture (Chapters 1 and 9 in this book) are related, in large part, to how the cultured population interacts with the ecosystem by means of suspension feeding. A close interplay between water filtration activity, primary production, seston availability, and hydrodynamics defines the magnitude of many of the ecological services provided by bivalves, as well as the sustainable level of aquaculture for a given area. Shellfish growth is limited primarily by a species’ capacity for nutrient

acquisition, which is regulated by feeding activity (Hawkins et al. 1999). Predictions of bivalve growth and the maximum aquaculture yield that can be produced within an area (production carrying capacity) often involve ecophysiological modeling, which includes equations describing how feeding processes (particle capture, selection, and ingestion) are related to population dynamics and environmental changes. Uncertainty or inaccuracy in feeding parameter estimates strongly influence model predictions of bivalve growth and carrying capacity (Dowd 1997). Suspension feeding always results in some local food depletion. The ecological costs of seston depletion by bivalve aquaculture are of concern only when the depletion zone is persistent and of an ecologically significant

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 81

82

Shellfish Aquaculture and the Environment

magnitude and spatial scale. Accurate information on how fast resident bivalve stocks can filter a body of water is required to assess the ecological carrying capacity of a region, which is the level of aquaculture that can be supported in the growing environment without leading to significant changes to ecological processes, species, populations, or communities (Gibbs 2007). Beyond the need to understand the potential direct effects of biofiltration, indirect effects on ecosystem processes and structure may result from the by-products of suspension feeding, including ammonia excretion and the egestion of particulate organic materials (feces and pseudofeces) on marine particle transport, energy flow, and nutrient cycling. Bivalve feeding activity has been studied across a wide range of laboratory and natural conditions and there is a vast literature, particularly on clearance rate, which is the volume of water cleared of particles of a certain size in a period of time. Despite this large research effort, there remains uncertainty in the measurement of clearance rate, which affects our confidence in predictions of individual to community-level feeding rates (Doering and Oviatt 1986; Cranford and Hill 1999; Riisgård 2001a, 2001b, 2004 Cranford 2001; Widdows 2001; Bayne 2004; Petersen et al. 2004). A fundamental understanding of the feeding behavior of individual bivalves provides the foundation for estimating population clearance rate, which is critical to understanding the ecological role of bivalves and environmental interactions with shellfish culture, including the capacity of dense bivalve assemblages to control the phytoplankton at the coastal ecosystem scale (Chapter 5 in this book). Feeding rate measurements must reflect the actual responses of specific species and cultured populations to the multiple physical, chemical, and biological factors that can influence feeding behavior in the natural environment. In this review, we summarize the available literature pertaining to feeding behavior with a focus on ascertaining the pres-

ence of predictive relationships for the clearance rate of key aquaculture species that can be used to help understand both positive and negative interactions between bivalve aquaculture activities and the environment. Our goal was to synthesize knowledge on specific aspects of bivalve feeding behavior relevant to the aquaculture issues outlined above. Several previously published reviews have addressed divergent hypotheses on physiological regulation of feeding activities and autonomous behavior (Jørgenson 1996; Bayne 1998; Riisgård 2001b). Such considerations are not the focus of this review, but it is not possible to address our task without contributing to this long-standing theoretical debate.

Constraints on maximum feeding activity Suspension-feeding lamellibranchiate bivalves rely on ciliated structures to capture and transport suspended particulate matter for selection and ingestion. The particle capture organ is the ctenidium (Ward et al. 1998), which also serves as the respiratory organ (gill). The asynchronous beating of lateral cilia on gill filaments serves as a pump that creates a water current which flows into the inhalant siphon or aperture, through the spaces between the gill filaments (or ostia) and then out of the exhalent siphon or aperture. Particles suspended in the feeding current are captured on the gill. An in-depth review of food capture, transport, and processing mechanisms in bivalve molluscs is provided by Ward and Shumway (2004). The ciliary activity of the gill delivers a flow that can be measured as a pumping (= ventilation) rate (P; L h−1) by various means such as the constant-level apparatus developed by Galtsoff (1926), the delivery of exhaled seawater into a constant flow of fresh water (Davenport and Woolmington 1982), the laser apparatus developed by Famme et al. (1986),

Bivalve filter feeding 83

the use of micro-flow meters (Meyhöfer 1985; Jones et al. 1992), and by the application of particle image velocimetry (Frank et al. 2008). Pumping rate scales with the size of the gill, which is somewhat proportional to the square of shell length (L2). Tissue dry weight (W) is proportional to L3 so pumping rate can, at least in theory, be expected to scale with W2/3 (=W0.67). This translates into a large increase in pumping rate for a small increase in body size (L or W). These relationships are described by the allometric equations P = aW b and P = aLb,

(4.1)

where a and b are fitted parameters. The allometric exponent (b) describes how fast the rate increases relative to body size. Jones et al. (1992) reported that maximum pumping rate for Mytilus edulis scaled with L2.1 (Fig. 4.1) and W0.70, and showed that that these allometric coefficients were not significantly different from the predicted values. Clearance rate (C; L h−1) is the more generally used measure of water processing than pumping rate, although the two measures are closely related. Some studies use the terms

Pumping Rate (L h–1)

3.5 3.0

2.09

Pmax = 0.0002 (L

)

2.19

Pmean = 0.0004 (L

)

2.5 2.0 1.5 1.0 0.5 0.0 10

20

30

40 50 60 Length (mm)

70

80

Figure 4.1 Allometric relationship between pumping rate (P) of the mussel Mytilus edulis and shell length (L). Multiple individual measurements for each mussel are shown, including the maximum (Pmax; 䉫). Regression lines and equations are shown for average P (Pmean; broken line) and Pmax (solid line) values. (Redrawn from Jones et al. 1992.)

clearance rate and filtration rate interchangeably, but the latter term is more often used to define the mass of particles cleared per unit time (e.g., mg h−1). Clearance and pumping rates are equal if all particles in the inhalant current are removed from suspension. Small particles (<2-μm diameter) in natural waters are not effectively retained by most species, but these particles can account for a high proportion of the total suspended particulate matter. Clearance rate, therefore, is measured for a particle size range that is retained by the species with 100% efficiency (>4- to 7-μm diameter, depending on the presence of eu- or pro-laterofrontal cilia, respectively; reviewed by Riisgård 2001a). While pumping rate can be measured directly by a number of methods, clearance rate is most often determined using indirect methods that involve measuring changes in particle abundance or concentration due to suspension-feeding activity of the bivalves. Under laboratory conditions, optimal food concentrations have been identified that stimulate the full exploitation of water pumping and particle clearance capacity in many species of bivalves (e.g., 1000–6000 Rhodomonas sp. cells mL−1 for Mytilus edulis; Kittner and Riisgård 2005). Riisgård et al. (2003) reported that under these conditions, the valves of Cardium edule, Mytilus edulis, and Mya arenaria are fully open within an hour after a starvation period of at least 24 h, and maximum clearance rate (Cmax) is thereafter maintained. The results of Cmax measurements on 13 bivalve species of different sizes from eight studies have been summarized by Riisgård (2001a, 2001b). Reanalysis of the parameters of the allometric relationships reported in these eight studies provide the following average (±standard deviation [SD]) relationships for dry tissue weight (W) and shell length (L): Cmax (L h−1 ) = 6.54 ± 2.41W 0.72 ± 0.09

(4.2)

Cmax (L h−1 ) = 0.0036 ± 0.10L1.60 ± 0.45

(4.3)

84

Shellfish Aquaculture and the Environment

As noted above, clearance rate is theoretically expected to scale with body geometry (L2 and W0.67), and the above equations generally confirm the assumed constraints of body size on maximum feeding activity. The exponent (b) in Equation 4.3 is lower than expected owing largely to the results of Meyhöfer (1985), which were conducted over a narrow size range. The median b-value of L1.81 may therefore be a more accurate estimate. Fluid dynamic forces can control the ability of shellfish to clear food particles. Uncertainty in the literature regarding the effect of current speed on bivalve feeding (positive vs. negative relationship) appears to stem from experimental conditions (Ackerman 1999). Turbulent conditions tend to result in a negative relationship between feeding activity and current speed, whereas the opposite is generally observed under laminar flow conditions. In general, there appears to be a unimodal functional response to flow: Moderate current speed and laminar flow conditions promote particle clearance rate, whereas high speeds and turbulent conditions inhibit clearance (reviewed by Ackerman 1999). The range of velocities permitting maximal feeding and shellfish growth depends on the species. For example, maximum clearance rate of Mytilus edulis was reported at ∼25 cm s−1 (Wildish and Miyares 1990) and 80 cm s−1 (Widdows et al. 2002), whereas Mytilus trossulus and Mytilus californianus exhibited peak rates at ∼18 and 12 cm s−1, respectively (Ackerman and Nishizaki 2004). The scallop Placopecten magellanicus exhibited flow inhibition at 20–25 cm s−1 (Wildish et al. 1987; Pilditch and Grant 1999), while the cockle Cerastoderma edule showed no inhibitory effects at 35 cm s−1 (Widdows and Navarro 2007). The reduction in feeding rate with increasing flow velocity appears to be a response of epifaunal bivalves to a flow-induced pressure differential between the pressure field in the mantle cavity, created by the ciliary pump, and

the external pressure field caused by flow velocity (Wildish and Kristmanson 1997). Back-pressure on the ciliary pump results in reduced pumping efficiency and valve closure (Jørgenson 1990; Wildish and Saulnier 1993). Similar effects of flow velocity have been observed for infaunal bivalves (Cole et al. 1992), and appear to result from the reduced ability of the bivalve to draw water into the mantle cavity when there is a high cross-flow. Temperature and salinity also serve as constraints on the maximum feeding rate of suspension-feeders. Temperature effects are related to a combination of mechanical (fluid dynamic) and physiological effects. At a scale of individual cilia, viscous forces dominate cilia and water motion. Water viscosity is inversely related to temperature, and the higher viscosity at lower temperatures has been shown to account for a large fraction of the effect of temperature on water pumping by ciliary filter-feeders, including bivalves (Jørgenson et al. 1990; Podolsky 1994). Viscosity affects the resistance of water flow within the shellfish pump, and the viscosity/ temperature relationship therefore limits their maximum clearance rate (Kittner and Riisgård 2005 and references cited therein). Salinity fluctuations also can severely disrupt normal feeding physiology (e.g., Navarro and González 1998; Gardner and Thompson 2001). Some species within the same taxon exhibit increased tolerance to low-salinity environments, resulting in differentiation of some species distributions (Seed 1992). The maximum capacity for food intake is ultimately limited by the morphological constraint of a limited gut volume and the time required to digest food. Both factors impose a major bottleneck on food uptake that morphological and physiological adaptations (e.g., active regulation of feeding rates) may help to mitigate (Hawkins and Bayne 1992; Bayne 1998).

Bivalve filter feeding 85

Shellfish feeding in nature Shellfish are opportunistic feeders that exploit the diverse nature of suspended particulate matter (the seston). The types of food utilized by shellfish include phytoplankton, ciliates, flagellates, zooplankton, and detritus, which occur within a spatially and temporally diverse mixture that includes inorganic materials (e.g., Trottet et al. 2008). Studies of the natural food available to suspension-feeding bivalves in coastal waters reveal marked short-term to seasonal variations in the concentration, composition, and nutritional value of the seston (e.g., Fegley et al. 1992; Cranford et al. 1998, 2005; Cranford and Hill 1999). Long-term changes in seston abundance and composition in temperate waters arise primarily from the seasonal cycle of primary production. Variability on a scale of days to weeks can result from algal blooms, horizontal phytoplankton patchiness, storm-induced resuspension of bottom deposits, or the spring/neap tidal cycle. In many coastal systems, fine-scale fluctuations in the seston are superimposed on these longer-term trends and result largely from tide-induced resuspension and deposition of bottom deposits and associated organic and inorganic constituents. The increase in the proportion of inorganic particles in the water column during resuspension events has a “dilution” effect on food quality, which is generally defined in terms of the organic and/or elemental content of seston. The challenge to shellfish ecophysiologists has been to characterize feeding responses over this wide range of feeding conditions, as well as to other potential mediating factors. The intent of such studies is to provide accurate predictive relationships between changes in food quantity and quality and feeding activity. In this section, we will characterize the range of temporal and spatial variability in clearance rate responses measured across a wide range of natural and experimental conditions prior to discussing

potential reasons for this variability in the following section.

Temporal variability in clearance rate Water pumping and clearance rate may be actively controlled by changing the activity of the lateral cilia of the gill that create the water flow, and by controlling various musculature that affects shell gape (valve opening), exhalent siphon area (Jørgenson et al. 1988; Newell et al. 2001), and interfilamentary distance of the gill (Jørgensen 1990; Medler and Silverman 2001; Gainey et al. 2003). These mechanisms appear to be uncoupled (Newell et al. 2001; Maire et al. 2007), possibly responding to different stimuli and/or providing different degrees of control over water flow and food acquisition. The capacity of shellfish to actively control clearance rate is a key factor in determining food acquisition, and individual shellfish appear to vary feeding activity continuously. Direct continuous measurements of water pumping rate in Mytilus edulis by Davenport and Woolmington (1982) showed that under optimal laboratory conditions (constant algal cell diet) mussels demonstrate relatively low temporal variability in pumping rate, but with irregular and sometimes large interruptions (Fig 4.2A). Variable diets (periodic starvation), however, induced large fluctuations in pumping rate. The large degree of variability in individual pumping rates is also shown in Figure 4.1B, which includes data on the full range of measurements from mussels of different size, including the maximum rate discussed above (Jones et al. 1992). Studies utilizing natural seston as the food source typically show large short-term variations in clearance rate over time periods ranging from minutes to hours, including periodic cessation of all feeding activity (Fig. 4.2B and Fig. 4.2C). Strohmeier et al. (2009) showed that large variations in

Shellfish Aquaculture and the Environment

(A)

60 50 40 30 20 10

0

0

2

4

6

8

Clearance Rate (L ind. h–1)

Pumping Rate (mL min–1)

86

(C)

60

40

20

0

0

6

12

18

24

0.4

0.2

0

0

20

40

60

80 100 120 140 160 180 200

Time (min)

Siphon Diameter (% maximum)

Clearance Rate (L min–1 mg–1)

(B) 0.6

30

36

42

48

52

Time (h)

Time (h) (D)

100 80

Feeding Chamber

60 40 20 In situ Benthic

0

0

6

12

18

24

30

Time (h)

Figure 4.2 Examples of short-term variations in shellfish feeding activity. (A) Continuous direct measurements of pumping rate of one Mytilus edulis during constant supply of an algal cell diet (5 × 103 cells mL−1) (Davenport and Woolmington 1982). (B) M. edulis clearance rate of natural seston (•, 5.5 mg L−1; 䊐, 2.5 mg L−1) in flow-through feeding chambers (redrawn from Fréchette and Bourget 1987). (C) Changes in clearance rate of 12 individual queen scallops (Pecten maximus; the symbols identify each scallop) feeding on natural seston in flow-through feeding chambers (redrawn from Strohmeier et al. 2009). (D) Percent maximum exhalent siphon area of M. edulis measured in situ (•) and in feeding chambers (䊏) (Newell et al. 2005).

short-term clearance rate responses of individual Mytilus edulis and Pecten maximus occur under relatively constant environmental conditions and that this variation was not synchronized with other individuals (Fig. 4.2C). Observations of shell gape and exhalent siphon area have been used to study feeding behavior under both controlled laboratory conditions (Newell et al. 1998; Riisgård et al. 2003; Frank et al. 2007; Maire et al. 2007) and naturally variable environmental conditions (Newell et al. 1998, 2005; Dolmer 2000; Saurel et al. 2007). Frank et al. (2007) showed that valve gape accounts for an average of only 25% (range of 2–82%) of the variation in clearance rate in oysters (Frank et al. 2007),

and this relationship can differ between individuals and within an individual over different periods of time. MacDonald et al. (2009) reported that exhalent siphon area explained a maximum of 53% of clearance rate variations in Mytilus edulis and that the correlation appeared to differ with mussel size. Valve gape and exhalent siphon area do influence water filtration to varying degrees, but these measurements should not be interpreted as being synonymous with clearance rate. The true value of these measurements is in their utility to provide semicontinuous measurements that can reveal general trends in feeding behavior (Frank et al. 2007; MacDonald et al. 2009).

Bivalve filter feeding 87

Newell et al. (2001) demonstrated that exhalent siphon area is more closely related to clearance rate than valve gape (see also Maire et al. 2007), and in situ video observations in a benthic mussel population in Maine, USA, showed large short-term variations in mussel feeding activity and feeding rates that were typically less than 50% of the maximum (Fig. 4.2D). Analysis of underwater photos of a mussel population in Limfjorden, Denmark, over a 4-day period showed that between 17% and 69% of the mussels were inactive (closed; Dolmer 2000). The highest percentage of closed mussels corresponded with a period of low food availability (<0.5 μg chlorophyll a [Chl] L−1), but a significant fraction of the population was inactive even when sufficient food was present. This study estimated that the gaping mussels were feeding at rates averaging 55% (range: 27–98%) of their maximum capacity. Saurel et al. (2007) used a similar approach to observe mussel feeding behavior in the Menai Strait, UK, and showed that individuals continuously varied feeding activity between zero and maximum over a 48-h period, with an average feeding rate of between 39% and 46% of maximum capacity. Field studies of temporal variations in clearance rate show a high degree of feeding variability over tidal to seasonal time scales (Fig. 4.3), with the transition between low, medium, and maximum clearance rates sometimes occurring over very short periods (e.g., Bayne and Widdows 1978; Thompson 1984; Deslous-Paoli et al. 1987; Fréchette and Bourget 1987; Cranford and Hargrave 1994; Smaal and Vonck 1997; Smaal et al. 1997; Newell et al. 1998, Cranford et al. 1998; Cranford and Hill 1999; James et al. 2001; Wong and Cheung 2001b; MacDonald and Ward 2009; Strohmeier et al. 2009). These studies, as well as the preponderance of research on bivalve feeding physiology, point to the high flexibility inherent within bivalve species to alter feeding behavior over different temporal scales.

Average standardized clearance rates Rate standardization and allometries For comparative purposes, water processing rates of bivalves have generally been standardized to a 1-g (dry tissue weight) individual using the following formula: Ys = Ye (1/We )b,

(4.4)

where Ys and Ye are the corrected and experimental physiological rates, respectively, We is the weight of the experimental animal, and b is the allometric exponent. Determining the appropriate exponent to use is a critical decision as the accuracy of the standardized value degrades exponentially as the b-value is progressively over- or underestimated. Selection of a b-value from the literature and comparisons between reported values requires some precautions (see below). Utilizing body mass to standardize clearance rate is merely a practical proxy for describing the effect of gill size (surface area). A complication is that seasonal changes in body mass are unrelated to gill size (fluctuations in internal nutrient stores and reproductive tissues fluctuates) and this introduces standardization errors that can complicate clearance rate comparisons both within and between studies. Shellfish condition index, which reflects seasonal cycles in nutritional and reproductive states, should be considered when comparing weight-standardized feeding rates. The true relationship between water processing rate and gill size may be more accurately related to shell length than to body weight (e.g., Filgueira et al. 2008). However, mean annual length/weight relationships can vary greatly between geographic locations and, under this scenario, gill size may be more accurately indicated, for comparative purposes, by animal weight than by shell length. The majority of results presented on the maximum clearance capacity of different shellfish species (above) come from populations in

88

Shellfish Aquaculture and the Environment

6

(A) Mytilus edulis

4

2

0 A

M

J

J

A

S

O

N

D

O

N

D

Clearance Rate (L g–1 h–1)

6 (B) Placopecten magellanicus 4

2

0

A

M

J

J

A

S

14 (C) Crassostrea gigas

12

Cerastoderma edule

10 8 6 4 2 0 M

A

M

J

J

A

S

O

N

D

J

F

Figure 4.3 Seasonal variation in standardized clearance rate of (A) the blue mussel Mytilus edulis, (B) the sea scallop (Placopecten magellanicus), and (C) the cockle Cerastoderma edule and oyster Crassostrea gigas. Solid lines in panels A and B are calculated from data reported in Cranford and Hill (1999). Discrete measurements are from Smaal et al. (1997; 䊏) and Smaal and Vonck (1997; •) for M. edulis, MacDonald and Thompson (1986; 䉬) for P. magellanicus, and Deslous-Paoli et al. (1987) for C. edule and C. gigas.

Denmark, which may help to explain the relatively low variability in allometric parameters given in Equations 4.2 and 4.3. We recommend that the selection of a standardization method be based on the specific goals of the study (e.g., understanding spatial vs. temporal variation), and that both the experimental animal weight and length be reported so that

other researchers can more effectively make comparisons between studies. The allometric relationship between clearance rate and body size of 21 species is summarized in Table 4.1. This summary only includes studies not previously reviewed by Winter (1978) and Riisgård (2001a, 2001b). The values of the allometric exponent for dry

Table 4.1 Clearance and pumping rate (C and P; L h−1) as a function of the size of different shellfish species according to the allometric equation Y = aXb, where Y is C or P, and X is dry tissue weight (W; g) or shell length (L; mm). Species and reference

Size

T (°C)

n

Weight relationship

0.1–5.7 g

12

15

C = 2.45W 0.80

Stuart (1982)

0.005–2.1 g

12.5

86

C = 1.48W 0.59

Van Erkon Schurink and Griffiths (1992)

10–60 mm

15

—

C = 0.89W 0.81

Smaal et al. (1986)*

—

9.5–15

53

C = 2.59 ± 0.12W 0.51 ± 0.02

Smaal et al. (1997)*

0.03–1.4 g

1.4–18.5

134

C = 1.44W 0.69

Urrutia et al. (1996)

—

—

—

C = aW 0.57

Newell and Bayne (1980)*

10–40 cm

—

68

C = 0.60W 0.51

0.01–7.0 g

3.4

21

C = 3.9W 0.60

Length relationship

Argopecten purpuratus Navarro and González (1998) Aulacomya ater C = 8.4 × 10−5L2.29

Cerastoderma edule

Chlamys islandica Vahl (1980)

Choromytilus meridionalis Griffiths (1980)

0.02–3.2 g

12, 18

64

C = 5.37W 0.60

C = 0.0064L1.58

Van Erkon Schurink and Griffiths (1992)

10–60 mm

15

—

C = 3.49W 1.00

C = 2.0 × 10−5L2.93

Bougrier et al. (1995)*

0.1–3.0 g

5–25

316

C = 3.92 ± 0.84W 0.50 ± 0.17

Ren et al. (2000)

0.02–0.4 g

10–13

38

C = aL0.57

Gerdes (1983)

0.05–0.81 g

20

10

C = 2.28L0.73

32–107 mm

13–21

41

Jones et al. (1992); maximum

0.01–2.1 g

10–12

24

P = 3.16W 0.72

P = 0.0037L2.09

Jones et al. (1992); mean

0.01–2.1 g

10–12

24

P = 1.78W 0.70

P = 0.0024L2.19

Smaal et al. (1997)*

0.02–3.90 g

0.4–19

139

C = 1.66 ± 0.55W 0.57 ± 0.17

Crassostrea gigas

C = 0.016L1.46

Mercenaria mercenaria Doering and Oviatt 1986)

C = 0.0307L0.97

Mytilus edulis

89

Table 4.1 (Continued) Species and reference

Size

T (°C)

n

Weight relationship

Hawkins et al. (1985)

45–57 mm

—

—

C = aW 0.45

Thompson (1984)*

0.07–4.20 g

0–15

128

C = 1.73 ± 0.19W 0.41 ± 0.08

Smaal et al. (1986)*

0.01–1.17g

9–15

20

C = 1.65 ± 0.79W 0.61 ± 0.20

Widdows (1978)

0.01–2.4 g

15

50

C = 2.65W 0.38

Lyner*

0.15–2.9 g

7–21

122

C = 1.98 ± 0.49W 0.45 ± 0.24

Cattewater*

0.13–2.8 g

10–17

105

C = 1.57 ± 0.36W 0.47 ± 0.11

Length relationship

Bayne and Widdows (1978)

Mytilus edulis platensis Dellatorre et al. (2007) Wild mussels

30–85 mm

20

49

C = 3.89W 0.56

C = 0.0040L1.72

Cultured mussels

16–73 mm

20

52

C = 5.74W 0.58

C = 0.0022L1.97

0.02–3 g

12

34

C = 1.13W 0.60

Mytilus chilensis Navarro and Winter (1982)

Mytilus galloprovincialis Filgueira et al. (2008); experiment 1

27–85 mm

—

59

C = 5.80W 0.60

C = 0.0035L1.72

Filgueira et al. (2008); experiment 2

24–86 mm

—

93

C = 5.02W 0.50

C = 0.0039L1.72

Van Erkon Schurink and Griffiths (1992)

10–60 mm

15

—

C = 4.08W 1.06

C = 1.7 × 10−5L3.02

Haure et al. (1998)*

0.1–2.7 g

10–30

149

C = 1.38 ± 1.15W 0.83 ± 0.07

Rodhouse (1978)

0.1–2 g

—

—

C = aW 0.48

Winter et al. (1984)

0.05–2.3 g

12

16

C = 1.32W 0.63

Winter et al. (1984)

0.05–2.3 g

12

12

C = 0.62W 0.60

20–100 mm

11–17

53

C = 2.29 ± 1.05W 0.75 ± 0.31

Ostrea edulis

Ostrea chilensis

Perna canaliculus James et al. (2001)*

90

C = 0.001 ± 0.016L1.99 ± 0.64

Bivalve filter feeding 91

Table 4.1 (Continued) Species and reference

Size

T (°C)

n

Weight relationship

Length relationship

25–85 mm

18–30

512

C = 0.81 ± 0.56W 0.61 ± 0.29

Berry and Schleyer (1983)

10–120 mm

20

97

C = 8.85W 0.66

C = 0.0027L1.86

Van Erkon Schurink and Griffiths (1992)

10–60 mm

15

—

C = 2.55W 0.88

C = 6.5 × 10−5L2.54

Pouvreau et al. (1999)

0.2–7.5 g

28

43

C = 25.88W 0.57

Yukihira et al. (1998)

0.05–20 g

25

52

C = 12.34W 0.60

0.05–20 g

25

54

C = 10.73W 0.62

Perna viridis Wong and Cheung (2001b)* Perna perna

Pinctada margaritifera

Pinctada maxima Yukihira et al. (1998)

Placopecten magellanicus MacDonald and Thompson (1986); 10 m

5–40 g

5.5–12

61

C = 0.942W 0.67

MacDonald and Thompson (1986); 31 m

5–20 g

1–11

56

C = 0.828W 0.69

0.005–3 g

22–25

120

C = 1.89W 0.32

0.03–0.87 g

13–18

54

C = 1.90 − 3.62W 0.61

Saccostrea commercialis KesarcodiWatson et al. (2001) Venerupis corrugatus Stenton-Dozey and Brown (1994)

Results are from studies not included in the previous reviews identified in the text. * = mean ± SD of significant regressions; a = intercept not reported.

body weight averaged 0.62 ± 0.05 (±2standard error [SE], n = 43). This b-value may be somewhat overestimated owing to the anomalously high values reported by Van Erkon Schurink and Griffiths (1992), which averaged

0.94. When these outlier data are excluded, the mean b-value falls to 0.58 ± 0.04 (n = 39), which is equal to the average reported by Winter (1978) for 12 species of bivalves (0.58 ± 0.07, n = 25). Combining these data

92

Shellfish Aquaculture and the Environment

sets gives an overall average b-value of 0.58 ± 0.04. This exponent is less than the theoretical value of 0.67 that assumes proportionality with gill surface area (see above) and is considerably less than the value of 0.72 reported for bivalves feeding at maximum capacity (Eq. 4.2). Although fewer studies have reported relationships between clearance rate and shell length, the allometries summarized in Table 4.1 indicate a mean b-value of 2.04 ± 0.54 (n = 14). This average is also weighted toward the high values reported by Van Erkon Schurink and Griffiths (1992), and a more generally applicable b-value is 1.78 ± 0.34 (n = 10). Although there is considerable variability between studies, there appears to be a tendency for b-values to be less than the theoretical proportionality between gill area and shell length (L2).

Meta-analysis of clearance rate responses for different species groups A meta-analysis was conducted on average clearance rate measurements extracted from 61, 30, 25, and 17 published studies on mytilid, pectinid, oyster, and cockle species, respectively. This was conducted to investigate central tendencies in values reported over a range of experimental conditions and to provide a foundation for discussing methodological viewpoints and the multiple factors potentially controlling the feeding responses of shellfish. Average rates reported using a wide range of direct and indirect clearance rate methodologies are incorporated in this analysis, including laboratory, field, and in situ studies with algal cell, seston, and/or mixed diets. The size of the literature dictated setting some limitations, and the studies identified in Table 4.2 (mussels) and Table 4.3 (scallops, oysters, and cockles) include only work with diet concentrations between 0 and 10 mg L−1, salinity greater than 20% ppt, and flow conditions less than 15 cm s−1. Potential food concentration, salinity, and water flow effects on

clearance rate are addressed separately. The food concentration limits were set to permit comparisons within a relatively narrow, but commonly studied range of food concentrations. Although shellfish feeding is responsive to low levels of many common anthropogenic contaminants and to toxic phytoplankton, these topics are outside the context of this review and such studies were not included in the meta-analysis, with the exception of utilizing data designated as representing reference (control) conditions. The studies reported represent an extensive, but not an exhaustive, search of the literature. Studies were randomly selected and the results are believed to be representative of the full range of published clearance rate values. Clearance rates reported in graphical form were digitized using Didger 4 software (Golden Software, Golden, CO). The literature on mussel feeding behavior is particularly extensive and frequency distributions of published average clearance rates for different mussel species of standard dry body weight (1 g) and shell length (60 mm) are presented in Figure 4.4 and summary statistics are given in Table 4.4. Results have generally been presented for mussels of standard weight (n = 401), but 72 average values were also reported for mussels of standard length (Fig. 4.4B). Reported clearance rates averaged 2.98 L g−1 h−1 and 3.23 L h−1 or a 60-mm mussel, but ranged widely with a maximum value of 17.5 L g−1 h−1 (not shown) reported for Mytilus galloprovincialis by Maire et al. (2007). The weight standardized distribution is skewed toward lower values, and the median of 2.32 L g−1 h−1 is substantially lower than the predicted mean of 6.54 L g−1 h−1 calculated using Equation 4.2 for animals feeding at maximum rates under optimal conditions. In fact, 95% of the reported mean values are below this predicted rate. Standardization by length gave a broad distribution of data (Fig. 4.4B) and the predicted maximum clearance rate of 5.95 L h−1 for a 60-mm mussel (Eq. 4.3 using b = 1.81) is still higher than the mean of

Table 4.2 Studies included in the meta-analysis of average clearance rate estimates for mytilid species subdivided by methodologies employed.

Flow-through chamber (aEq. 4.5 or bEq. 4.6 in text)

Closed system (aclearance, bchemostat, c static, or dflume)

Biodeposition (achamber or bin situ)

Consumption (atunnel, b chamber, or csuction)

Aulacomya ater Bayne et al. (1984)a, A

Stuart (1982)c, A

Choromytilus merdionalis Bayne et al. (1984)a, A

Griffiths (1980)c, A

Mytilus chilensis Navarro et al. (2003)a, A

Velasco and Navarro (2003)a, A, S

Mytilus edulis Okumus and Stirling (1994)a, S

Kittner and Riisgård (2005)a, A

Newell et al. (2005)a, S

Prins et al. 1994 a, S

Smaal and Vonck (1997)a, S

Newell et al. (1989)c, S

Petersen et al. (2004)a, S

Prins et al. (1996)a, S

Fréchette et al. (1991)a, S

Riisgård (1991)a, A

Hawkins et al. (1996)a, S

Bayne et al. (1989)b, A

Prins et al. (1996)a, S

Riisgård et al. (2003)a, A

Cranford and Hill (1999)b, S

Smaal et al. (1986)a, S

Newell et al. (2005)a, S

Riisgård and Møhlenberg (1979)b, A

Hawkins et al. (1997)a, A

Zurburg et al. (1994)a, S

Widdows et al. (1979)a, S

Widdows et al. (2002)d, A

Dellatorre et al. (2007)a,

Møhlenberg and Riisgård (1979)c, A

Petersen et al. (2004)a, b, S

S

Petersen et al. (2004)c,

Kiørboe et al. (1980)c, A

S

Smaal et al. (1997)b, S

Riisgård et al. 2003)a, A

Thompson (1984)a, S

Lucas et al. (1987)c, S

Bayne et al. (1987)a, S

Kiørboe et al. (1981)b, A

Bayne et al. (1985)a, b, A

Clausen and Riisgård (1996)a, A

Widdows (1978)a, A

Bayne et al. (1985)c,

A

Widdows and Johnson (1988)a, A Smaal and Twisk (1997)a, A Widdows et al. (1984)a, S Newell et al. (1998)b, S Vismann (1990)b, S MacDonald and Ward (2009)a, S Bayne and Widdows (1978)a, S

93

94

Shellfish Aquaculture and the Environment

Table 4.2 (Continued)

Flow-through chamber (aEq. 4.5 or bEq. 4.6 in text)

Closed system (aclearance, bchemostat, c static, or dflume)

Biodeposition (achamber or bin situ)

Maire et al. (2007)a, A

Iglesias et al. 1996)a, S

Consumption (atunnel, b chamber, or csuction)

Mytilus galloprovincialis Filgueira et al. (2006)a, A, S a, A

Filgueira et al. (2008)

a,

Navarro et al. (1996)

Navarro et al. (1991)a, S

A

Labarta et al. (1997)a, A

Pérez Camacho et al. (2000)a, S

Navarro et al. (1991)a, S Pérez Camacho et al. (2000)b, A Mytilus trossulus Ackerman and Nishizaki (2004)c, S Mytilus californianus Ackerman and Nishizaki (2004)c, S Perna canaliculus James et al. (2001)a, S

Zeldis et al. (2004)c, S

Hawkins et al. (1999)a, S

Perna perna Bayne et al. (1984)a, A

Resgalla et al. (2007)c, A

Berry and Schleyer (1983)c, A Perna viridis Wong and Cheung (1999)a, A Wong and Cheung (2001a)a, S Wong and Cheung (2001b)a, S Note that some studies used several methods and/or species. Superscripts indicate the method and use of algae cell (A) and/or seston diets (S).

3.23 L h−1 (median = 3.18 L h−1). The literature clearly shows that the maximum feeding response has only occasionally been observed in mussels and is an extreme condition within a wide and highly variable range of average rates.

One factor contributing to the shape of clearance rate frequency distributions is the variation in the allometric exponent employed in each study, which ranged from 0.2 to 1.1. Relatively few studies determined the b-value; instead using a value selected from the litera-

Table 4.3 Studies included in the meta-analysis of average clearance rate estimates for scallop, oyster and cockle species. Scallop

Oyster

Argopecten irradians

Cockle

Crassostrea gigas

Bauder et al. (2001)

1

Bayne (2004)

Cerastoderma edule

2, 4

Deslous-Paoli et al. (1987)4

Li et al. (2009); low density only)1

Bayne (1999)4

Foster-Smith (1975)4

Palmer (1980)4

Bougrier et al. (1995)4

Ibarrola et al. (2000)2

Deslous-Paoli et al. (1987)4

Ibarrola et al. (1998)4

Navarro et al. (2000)4

Zurburg et al. (1994)5

Iglesias et al. 1996)2

Navarro and González 1998)4

Dupuy et al. (2000)4

Iglesias et al. (1998)4

Lefebvre et al. (2000)2

Møhlenberg and Riisgård (1979)5

Argopecten purpuratus

Argopecten ventricosus-circularis 4

1

Sicard et al. (1999) Argopecten nucleus Velasco (2007)1 Chlamys nobilis Li et al. (2001)4 Chlamys opercularis

Ren et al. (2000)

Navarro et al. (1992)4

Haure et al. (2003)1

Navarro and Widdows (1997)1

Smaal and Zurburg (1997)5

Newell and Bayne (1980)1

Barillé et al. (2003)1

Prins and Smaal (1989)1

Gerdes (1983)4

Prins et al. (1991)2

1

McLusky (1973)

Hawkins et al. (1998)

Chlamys farreri

Crassostrea iradelei

Jihong et al. (2004)4

Hawkins et al. (1998)1

Zhou et al. (2006)2

Crassostrea virginica

Smaal et al. (1997)1 Urrutia et al. (2001)2 Widdows and Navarro (2007)4 Widdows and Shick (1985)1

Palmer (1980)4

Chlamys hastata Meyhöfer (1985)5

Pernet et al. (2008)4 Riisgård (1988)4

Chlamys islandica 1

Vahl (1980)

Ostrea edulis

Nodipecten nodosus

Haure et al. (1998)1

Velasco (2007)1

Newell et al. (1977)4 Rodhouse (1978)4

Placopecten magellanicus Bacon et al. (1998)1

Wilson (1983)5

Cranford and Grant (1990)4

Buxton et al. (1981)4 1

Cranford and Gordon (1992) Cranford et al. (1998)

Riisgård et al. (2003)3

Crassostrea belcheri

4

Ostrea chilensis

2

Dunphy et al. (2006)4

Cranford and Hill (1999)2

Chaparro and Thompson (1998)4

Cranford et al. (2005)2

Saccostrea commercialis 1

MacDonald and Thompson (1986)

Bayne et al. (1999)4

MacDonald and Ward (2009)1

Kesarcodi-Watson et al. (2001)1

Ward et al. (1992)4

95

96

Shellfish Aquaculture and the Environment

Table 4.3 (Continued) Scallop

Oyster

Cockle

MacDonald and Ward (1994)1 Pilditch and Grant (1999)4 Wildish et al. (1992)4 Pecten maximus Strohmeier et al. (2009)1 Pecten furtivus Møhlenberg and Riisgård (1979)5 Pecten opercularis Møhlenberg and Riisgård (1979)5 Pecten irradians Chipman and Hopkins (1954)5 Superscripts indicate the method employed (1 = flow-through; 2 = biodeposition; 3 = clearance; 4 = static; and 5 = other).

ture. Where sufficient data were reported on mussel weight and/or length, the standardized results were converted back to experimental rates using the reported b-value and then standardized again using the average weight and length allometric exponents of 0.58 and 1.8, respectively, determined above. Studies reporting nonstandardized rates were also standardized by this method, which increased the number of length-standardized values from 72 to 361. This procedure increased the median clearance rate to 2.46 L g−1 h−1 (Table 4.4; Fig. 4.4C) and 3.26 L h−1 for a 60-mm mussel (Table 4.4; Fig. 4.4D). The resulting distribution for weight-normalized rates is bimodal with a smaller peak at ∼5 L g−1 h−1 that is attributed to the results of studies designed to stimulate maximum clearance rate through the use of an optimal algal-cell diet. Comparison of Figure 4.4C and Figure 4.4D shows that the distribution of reported clearance rates of mussels of standard weight is more closely constrained than for animals of standard length. This suggests that some of the variability between published rates results from geographic differences in mussel length–weight

relationships and the fact that length is not a particularly good surrogate for weight (Hilbish 1986). Summaries of mean standardized clearance rates for scallops, oysters, and cockles are presented in Table 4.4 along with the mussel data that were shown in Figure 4.4. Given the bimodal distribution of these data, differences between species groups are examined using median values. Based on reported rates, mussels appear to feed at slightly lower rates relative to the other three groups. Standardization using a constant b-value of 0.58 indicated that oysters appear to have the lowest median clearance rate. In both cases, cockles exhibited the highest clearance rates. A factor having a large effect on the mean rate obtained for each species group is the type of experimental diet provided. Studies utilizing an algal cell-based diet resulted in clearance rates of mussels, scallops, and cockles that were on average 1.7 times higher than rates obtained utilizing a natural seston-based diet (Table 4.4). The only exception was for oysters, where data for seston diets was too sparse (n = 10) for an objective comparison. The

Bivalve filter feeding 97

12 60

(A) n = 401

(B) n = 72

10

50 8

Frequency

40

6

30 20

4

10

2

0 60

0 40

50

(C)

(D)

n = 357

n = 361 30

Frequency

40 20

30 20

10 10 0 0

2

4

6

8

Clearance rate (L g–1 h–1)

10

0

0

2

4

6

8

10

Clearance rate (L ind. h–1)

Figure 4.4 Frequency distributions of mussel clearance rate results from the 61 publications listed in Table 4.2. Average rates are standardized by dry weight (A and C; L g−1 h−1) and shell length (B and D; L h−1 for 60 mm individual). Plots A and B summarize results as published and plots C and D show the distributions after restandardization using b-values of 0.58 and 1.8 for weight and length, respectively. The arrows indicate median values, and the vertical dashed lines represent the maximum rates predicted by Equations 4.2 and 4.3 for weight- and length-standardized animals, respectively.

generally higher rates obtained for the highnutritional-quality algal-cell diets was the main reason for the bimodal frequency distribution observed for these species groups. For most aquaculture and wild population predictions requiring clearance rate data, the sestonbased results are a logical choice, with mussels, scallops, and cockles feeding at 2.43, 3.49, and 2.81 L g−1 h−1, respectively. The average

rate for oysters feeding on both types of diets (2.54 L g−1 h−1) is comparable with mussels, and cockles and scallops appear to have a relatively high rate of feeding. The above comparisons of clearance rates across bivalve species groups are not meant to be conclusive given possible differences in the range of experimental conditions employed in the study of each group. In fact, it may be

98

Shellfish Aquaculture and the Environment

Table 4.4 Summary statistics on average (±2SE) standardized (1 g dry tissue weight and 60 mm shell length or height) clearance rate values reported in papers listed in Tables 4.2 and 4.3 and after restandardization using average b-values (n = number of mean values applicable). Mussel

Scallop

Oyster

Cockle

Dry tissue weight as reported (L g−1 h−1) n

401

123

123

111

Median

2.32

2.63

3.00

3.37

Mean ± 2SE

2.98 ± 0.23

3.63 ± 0.59

3.47 ± 0.49

3.53 ± 0.44

−1

−1

Shell length (or height) as reported (L ind. h ) n

72

—

—

—

Median

3.18

—

—

—

Mean ± 2SE

3.23 ± 0.34

—

—

—

−1

−1

Dry tissue weight using b = 0.58 (L g h ) n

357

172

68

90

Median

2.46

3.21

2.05

3.57

Mean ± 2SE

3.24 ± 0.26

5.10 ± 0.76

2.54 ± 0.48

3.58 ± 0.38

Seston-based diet mean ± 2SE (n)

2.43 ± 0.28 (161)

3.49 ± 0.86 (79)

4.78 ± 0.56 (10)

2.81 ± 0.66 (50)

Algae-based diet mean ± 2SE (n)

4.18 ± 0.41 (178)

6.66 ± 1.27 (79)

2.15 ± 0.49 (58)

4.12 ± 0.54 (61)

60 mm shell length (height) using b = 1.8 (L ind.−1 h−1) n

361

130

49

67

Median

3.26

5.18

1.09

5.30

Mean ± 2SE

3.82 ± 0.28

7.89 ± 1.37

1.39 ± 0.28

6.03 ± 0.81

expected that mussels would have a higher feeding rate than cockles based on the observation of Hawkins et al. (1990) that the gill area of Mytilus edulis was more than four times greater per unit tissue mass than Cerastoderma edule. However, it is important to note that this morphological characteristic can be population specific, with the gill area-to-dry tissue mass ratio varying considerably within a species. For example, Mytilus edulis from Denmark and Spain exhibited gill area-to-dry weight ratios of 2458 mm2 g−1 (Møhlenberg and Riisgård 1979) and 4322 mm2 g−1 (Hawkins et al. 1990), respectively. Morphology by itself does not fully explain inter- and intraspecific

variations in clearance rate. The tropical oyster Pinctada margaritifera consistently exhibited the highest clearance rates of any bivalve studied (averaged 21.3 L g−1 h−1; Pouvreau et al. 1999, 2000; Yukihira et al. 1998) and had an intermediate gill area of 3502 mm2 g−1 (Pouvreau et al. 1999). This was the only species omitted from the meta-analysis owing to the large outlier effect on calculations of the mean clearance rate response. The average clearance rates reported in Table 4.4 for different bivalve species are useful for the first-order approximation of population and community clearance values. However, because of the very wide range of

Bivalve filter feeding 99

clearance rates and b-values reported for individual bivalve populations, we stress the importance of measuring actual clearance rates at the study site during the time(s) of year that is (are) pertinent to the specific application of the data. The effect of the allometric exponent employed can be seen in the difference between the reported mean and the restandardized mean clearance rate (Fig. 4.4 and Table 4.4). In addition, a sensitivity analysis of a scallop (Chlamys farreri) growth model showed that a 10% change in the weight exponent caused a 13% change in tissue growth predictions (Hawkins et al. 2002). This effect is relatively small owing to the small size of the scallops studied. The magnitude of the growth effect will increase with the size of the animal, such that the difference in b-values (e.g., 0.76 in Eq. 4.2 vs. 0.58 calculated from data in Table 4.1) would have a substantial impact on population food removal and aquaculture production predictions.

Precision and accuracy of clearance rate measurements The usual consequence of a high degree of intraindividual variability in feeding activity (see above) is that a large number of replicate measurements are required to obtain sufficient precision (i.e., the degree of agreement between replicated measurements) to protect the analyst from reaching erroneous conclusions when conducting statistical hypothesis tests. Although seldom reported, zero clearance rate values are often discarded prior to statistical analysis. This is a subjective practice that will increase the precision of the mean response, but which will cause overestimation when extrapolating mean individual rates to the population level. The periodic cessation of feeding activity is a normal occurrence (Fig. 4.2) that should not be ignored. Analysis of clearance rate data collected by Strohmeier et al. (2009), who collected 18 replicate measurements to ensure maximum precision, indi-

cate that the highest precision obtainable was ∼10% (SE/mean) and that achieving this level required a minimum of 12 replicate measurements. The studies listed in Table 4.2 (mytilid species) were examined and 30 studies provided sufficient data to calculate a precision of 15% for 254 average clearance rate estimates (average sample size was 11.6 replicates). The following illustrates the importance of this degree of intraindividual variability for studies attempting to detect significant differences in average clearance rate responses using inferential statistics. Power analysis was conducted for a paired t-test and one-way analysis of variance (ANOVA) (Power and Precision Ver. 2 software) based on the typical error variance observed across these clearance rate studies at n = 10. To achieve an acceptable statistical power of 80% at α = 0.05, the minimum “effect size” (the standardized mean difference in clearance rate between experimental treatments) that can be detected is 50%. This is a relatively large effect size for biological measurements, making it difficult to detect differences in average clearance rates between experimental treatments. When measuring clearance rate, the following quote by Nakagawa and Cuthill (2007) is particularly relevant: “. . . all biologists should be ultimately interested in biological importance, which may be assessed using the magnitude of an effect, but not its statistical significance.” Other relevant statistical considerations that require more attention include the use of regression analysis in cases where both the dependent and independent variables are based on the same data (e.g., clearance rate vs. seston organic content) and the need to use repeated-measures designs when clearance rate values are not independent (MacDonald and Ward 2009). Time series measures on the same individuals cannot be avoided in some cases (e.g., in situ observations of shell gape and biodeposition) so the independence assumption needs to be tested prior to the use inferential statistics. Strohmeier et al. (2009)

100

Shellfish Aquaculture and the Environment

employed autocorrelation analysis to show that mussel and scallop clearance rates were independent when repeated measurements were taken at time intervals greater than 1 day. Although clearance rate measurements of individual animals dominate the literature, an alternative approach is to measure population responses. Population responses tend to require fewer measurements to achieve a higher level of precision due to the “built-in” averaging of interindividual variations (Cranford et al. 1998; Iglesias et al. 1998). Individual and population measures do not necessarily provide comparable information, and it should not be assumed that average weight-standardized individual estimates are equal to similarly standardized population estimates. Interactions between individuals (e.g., disturbance, crowding, flow alteration, refiltration of water) may influence some population-based methodologies if they are not controlled. In an attempt to justify the general application of maximum clearance rate measurements in nature and related theories on shellfish feeding nonresponsiveness to environmental forcing, Riisgård (2001a; see also Jørgensen 1996) concluded that “. . . conflicting data on filtration [clearance] rates seem partially due to the incorrect use of methods, and partly to be caused by differences in experimental conditions.” Bayne (2001) has already provided sufficient rationale why this conclusion is unacceptable, but statements such as this continue to impede the objective discussion of the extrinsic factors that affect shellfish feeding behavior (e.g., Cranford 2001; Widdows 2001; Filgueira et al. 2006) and severely affect the peer review process. There is a need for accuracy in any biological measurement and this need is particularly critical when the intent is to extrapolate the responses of a few animals to predict the biofiltration capacity of extremely dense populations. Any significant errors in estimating the individual response will be magnified immensely at the population level. Consequently, an assessment of the accuracy

of clearance rate measurements continually requires attention, including discussion of the validity of assumptions inherent with available clearance rate methodologies, assumptions regarding the true feeding rate, and the intercalibration of clearance rate methodologies. A wide range of indirect clearance rate methodologies exist and can be classified based on the measurement of bivalve particle removal (depletion) or egestion (biodeposition). Particle removal methodologies include open (flowthrough chambers, in situ tunnels, and suction methods) and closed (chambers, aquaria, and mesocosms) systems. Examples of the closed method include the “static method” and the so-called “clearance method” described by Riisgård (2001a). Both determine clearance rate from the exponential decline in particle concentration as a function of time (Coughlan 1969), but the latter may be differentiated by the use of a bivalve population, an optimal algal cell diet, and an algal dosing pump to somewhat maintain the algal concentration. The flow-through chamber method has been the most commonly employed method and clearance rate is calculated using either of the following two equations: Flow-through equation: C = U ((Fin − Fout )/Fin ) (4.5) Steady-state equation: C = U ((Fin − Fout )/Fout ), (4.6) where U is the flow through the chamber (L h−1), and Fin and Fout are food particle concentrations in the inflow and outflow of the chamber, respectively. The “flow-through” equation applies to systems where all the particles in the inflow are available to the animals and recirculation of filtered water is absent. The “steady-state” equation represents an alternative for systems with low flow and water recirculation. The importance of ensuring that the flow rate is adequate to meet assumptions inherent with using the

Bivalve filter feeding 101

10 (B) 14

17

(A)

Other

Static

17

33

43 Flow-through (Eq. 4.5)

103 Biodeposition

Flow-through (Eq. 4.6)

Clearance

30 40 Static

Other

72

0

Clearance

2

Biodeposition

4

156 Flow-through (Eq. 4.5)

32

6

Flow-through (Eq. 4.6)

Clearance rate (L g–1 h–1)

9

8

Method Figure 4.5 Average (+standard error [SE], with sample size shown) weight-standardized clearance rates estimated for mytilid species by the displayed methods. Data are from studies listed in Table 4.2. Plot A summarizes reported standardized rates and plot B includes restandardized rates assuming b = 0.58. Arrows on the top of each plot connect methods that provided statistically equivalent results (ANOVA, P < 0.05).

flow-through equation has long been recognized (e.g., Riisgård 1977; Bayne et al. 1985) and can be demonstrated simply by measuring no change in clearance rate with increasing flow rate (U). Riisgård (2001a) stated, however, that “the prerequisites for using the method have often been disregarded” resulting in underestimated rates. Filgueira et al. (2006) and Pascoe et al. (2009) critically examined two different flow-through feeding chamber designs and the application of Equation 4.5 and concluded that the method complies with all theoretical requirements provided that the percentage of particles cleared from the flow is less than 25–30% (achieved at U > 150 mL min−1). This maximum percentage of particles cleared will vary somewhat with the geometry of different feeding chambers and is not a substitute for examining the relationship between clearance values and flow

rates; however, most published studies have reported less than 30% reduction of particles. Although several of the reviewed studies on mytilid species (Table 4.2) report levels of food depletion up to 40%, it is particularly important to note that even at 40% particle depletion, the potential underestimation of true clearance values resulting from insufficient chamber flow is relatively small compared with the large difference between the median and maximal clearance rates shown in Figure 4.4 (see figs. 4.4 and 4.5 in Filgueira et al. 2006). The steady-state equation (Eq. 4.6) was employed to estimate clearance rate in 10% of the studies listed in Table 4.2. However, Filgueira et al. (2006) and Pascoe et al. (2009) showed that this equation resulted in reduced precision and overestimation of clearance rate and concluded that Equation 4.5 provided a more accurate representation of true clearance

102

Shellfish Aquaculture and the Environment

rate. Petersen et al. (2004) reported similar results, but nonetheless promoted the general application of the steady-state equation. Feeding rate methodologies based on a combination of shellfish biodeposit production and suspended particle measurements have become increasingly prominent (Table 4.2) since the development of the pseudofeces/ feces biodeposition method (Navarro et al. 1991) and the related in situ biodeposition method (Cranford and Hargrave 1994). Both methods are similar except that the former approach permits measurements to be made at higher food concentrations where pseudofeces production becomes a significant fraction of filtration rate. As with all other methods, there is a need to ensure that certain assumptions are met for the proper application of these techniques. These assumptions include the need for accurate characterization of suspended particles retained by the gill (a common assumption for all indirect methods) and quantitative biodeposit collection. Assessment of these assumptions has become routine with the application of the in situ biodeposition method (Cranford and Hargrave 1994; Cranford et al. 1998), and a sensitivity analysis has shown that potential errors in seston diet characterization using standard water filtration methods could only result in underestimating feeding rates by less than 12% (Cranford and Hill 1999). A fundamental difference between biodeposition and particle depletion methodologies for measuring clearance rate is that they provide temporally integrated and near-instantaneous responses, respectively. In addition, the biodeposition method often provides an integrated response for several animals. Short-lived extreme clearance rate responses (maximum and zero) that may be of little ecological relevance can greatly influence the short-term mean response, but have little effect on integrated clearance rates obtained with the biodeposition method. As a result, the direct comparison of results obtained with these fun-

damentally different methodologies is not strictly justified, but such comparisons have been made and the results deserve some discussion. A number of studies have shown that clearance rate values obtained with the biodeposition and flow-through chamber methods provide statistically equivalent clearance rate estimates (Urrutia et al. 1996; Iglesias et al. 1998; Cranford and Hill 1999; Bayne 2004; Newell et al. 2005). Comparison of rates obtained with the biodeposition and indirect methods also gave similar results for both Mytilus edulis and Mulinia edulis (Navarro and Velasco 2003). Conversely, Petersen et al. (2004) concluded that the biodeposition method resulted in significantly lower clearance rate estimates relative to other methods tested. This intercalibration study and the results reported have been questioned based on (1) the failure to sample biodeposits quantitatively (Bayne 2004), (2) the lack of any validation that the experimental conditions actually conform with the basic requirements of the deposition method (Riisgård 2004), and (3) the erroneously high rates that can be obtained when using Equation 4.6 in conjunction with the flow-through method (Filgueira et al. 2006; Pascoe et al. 2009). Average weight-standardized clearance rate data summarized in Figure 4.4 (Table 4.2) were further analyzed to examine possible effects of different methodologies on mean clearance rates of mytilid species. Reported (Fig. 4.4A) and restandardized clearance rates (Fig. 4.4C) were analyzed separately by means of one-way ANOVAs, which indicated a significant difference in mean results obtained by the different methods (degree of freedom [d.f.] = 341 and 213, respectively; P < 0.001). Tukey post hoc comparisons of reported rates showed that most methods provided similar results. The exception was the clearance method, which provided an overall mean rate that was significantly higher by about a factor of two (Fig. 4.5A). When reported results were restandardized, the flow-through method

Bivalve filter feeding 103

(Eq. 4.5) gave similar results as the biodeposition and static chamber methods (Fig. 4.5B), but the clearance and steady-state flow-through method (Eq. 4.6) provided significantly (P < 0.05) higher clearance rates (Fig. 4.5). Data from these two methods are largely responsible for the second smaller peak in the frequency distribution (Fig. 4.4C). The majority of published comparisons of indirect clearance rate methodologies, as well as the results of our meta-analysis, shows that similar mean clearance rate results are obtained with most methods when they are employed properly under the same range of experimental conditions. An exception is that the application of Equation 4.6 with the flow-through method can cause overestimated values, particularly when chamber flow rates are relatively low (Pascoe et al. 2009). The clearance method has only been employed under a narrow range of conditions (optimal algal cell rations) and the anomalously high clearance rates that have been obtained reflect these conditions as opposed to any methodological error. This method is not compatible with the use of more complex diets (i.e., natural seston) and a direct comparison is not possible. Questions regarding the accuracy of clearance rate measurements have generated a rather fruitless debate in the literature because we cannot arbitrarily assume that any rate is more accurate than another. Accuracy may best be assessed by determining the ability of reported clearance values to predict measured tissue growth or the magnitude of phytoplankton depletion resulting from the feeding activity of a given bivalve population. Assessing feeding rates by examining tissue growth over time requires numerous assumptions regarding measurement accuracy for all the physiological components of growth as well as for any applicable energy conversion factors. Further complicating such an approach are the possible effects of resource limitation (selection of energy, carbon or nitrogen budget approaches) and problems comparing a time-integrated

measure (growth) with instantaneous physiological and dietary measures. This approach, however, has been used on several occasions. For example, Fréchette and Bacher (1998) used published data on seston clearance rates and tuned the parameters of the allometric relationship until a growth model provided estimates that matched the observed growth of mussels. The clearance relationship that yielded accurate growth predictions (2.12W0.408) was similar to the equation taken from the literature (Thompson 1984; 1.72W0.413). In this case, a relatively low clearance rate appeared to provide accurate predictions. Cranford and Hill (1999) showed that the growth of Mytilus edulis and Placopecten magellanicus was similar to predictions based on seasonal clearance rates measured using the in situ biodeposition method. Clausen and Riisgård (1996) attempted the only validation of the application of maximum clearance rates (Cmax) to a natural population of bivalves and concluded that Mytilus edulis must fully exploit their clearance capacity to explain the growth observed in nature. These authors, however, assumed that phytoplankton were the sole food resource. If the other available detrital and living food resources had been included in these growth calculations, a lower clearance rate would also calculate the observed growth. A somewhat more direct approach to assessing clearance measurement accuracy (i.e., fewer assumptions) is to back-calculate individual clearance rates required to explain the measured population effects on particle sedimentation or depletion in the surrounding water. Doering and Oviatt (1986) showed that clearance rates of Mercenaria mercinaria measured using a natural diet and the flow-through method gave estimates of gross sedimentation that agreed well with observations of gross sedimentation in a mesocosm. They noted that the use of maximum clearance rates (Eq. 4.2) would overestimate sedimentation by up to an order of magnitude. Results of a

Shellfish Aquaculture and the Environment

high-resolution spatial model of seston depletion in dense Mytilus edulis culture (Grant et al. 2008), which assumed an average clearance rate of 2.4 L g−1 h−1 (similar to median values reported herein), predicted bay-scale phytoplankton depletion levels that correspond well with measurements obtained using rapid, high-resolution phytoplankton mapping surveys with a towed sensor vehicle. Petersen et al. (2008) utilized an advection depletion model to determine particle depletion rates for a raft-culture unit containing 40-mm shell length mussels. They concluded that the measured depletion rates could be estimated using clearance rates between 0.6 and 0.9 L ind.−1 h−1. This is lower than would be predicted using clearance rates reported in Table 4.4, but this is expected given water refiltration under such dense culture conditions. Assumptions regarding the above back-calculation approach to assessing clearance rate accuracy include: No refiltration of water by bivalves located downcurrent, and no feedback effect on clearance rate from changing food concentration (depletion). Although both assumptions can compromise the validity of this approach, the general similarity of measurements and predictions across these studies tends to validate the high accuracy of most clearance rate methodologies when they are conducted under ecologically relevant conditions.

Controls on bivalve clearance rate Accurate descriptors of shellfish feeding behavior, which are needed to predict growth and environmental interactions, have to incorporate responses to the major endogenous and exogenous factors and must reflect the net response to multiple, simultaneous forcing parameters. Temperature is one factor that is known to limit the maximum feeding response in shellfish (e.g., Kittner and Riisgård 2005; see above). However, temperature has not been identified as an important control on

feeding behavior under more natural conditions for several mytilid and pectinid species (Widdows and Bayne 1971; Widdows 1973, 1976, 1978; Widdows et al. 1979; Thompson 1984; Thompson and Newell 1985; Prins et al. 1994; Smaal et al. 1997; Cranford and Hill 1999; Cranford et al. 2005; Strohmeier et al. 2009). For example, a time series of in situ sea scallop (Placopecten magellanicus) clearance rates measured at 3°C showed that feeding activity was variable and that the maximum clearance rate can be achieved even at this low temperature (Cranford et al. 2005). In situ measurements of mussel (Mytilus edulis) clearance rates at different times of the year also revealed little dependence of feeding on temperature (Fig. 4.6; Ward and MacDonald, unpublished data). An important consideration when exploring the effect of temperature on bivalve feeding is the need to distinguish between the welldocumented effects of temperature on the maximum clearance rates of fully open bivalves and the potential effects on animals that feed at submaximal rates. Animals stimulated to exhibit the maximum clearance response are essentially physiological slaves to the constraints that limit bivalve feeding capacity

6 Clearance rate (L g–1 h–1)

104

5 4 3 2 1 0

0

2

4

6 8 10 12 Temperature (°C)

14

16

Figure 4.6 Average (±SD) standardized clearance rates of Mytilus edulis measured in the field over a wide range of water temperatures using the flow-through chamber method (J.E. Ward and B.A. MacDonald, unpublished data obtained in Newfoundland, Canada).

7 6 5 4

(A) High seston Hawkins et al. (1999)

Perna canaliculus

3 2 1 0

0

5

50 Clearance rate (L h–1)

(body size, fluid dynamics, and related temperature effects). Although no physiological mechanisms can override these fundamental constraints on clearance rate, a number of studies have nonetheless employed optimal laboratory conditions as a basis for concluding that evidence for the regulation of feeding performance is lacking in shellfish (Jørgensen 1996; Riisgård 2001b; Kittner and Riisgård 2005). Geometric and physical factors only constrain the upper limits of filtration activity and do not rule out flexibility in feeding performance at reduced rates. Riisgård (2001b) concluded that it is important that future research on bivalve compensatory responses to variations in the environment be made under optimal conditions that result in maximum feeding activity. This would perpetuate an expected result that could only be extrapolated to cultured or wild shellfish populations if bivalves in nature always exhibited maximal feeding rates such as those stimulated by a controlled, artificial diet. The above discussion on feeding variability and range of mean responses clearly shows that it is unacceptable to limit observations to an artificial condition that does not exist in nature. Seston concentration has a strong influence on bivalve clearance rate and explains a large fraction of the variance in clearance rate measurements (Table 4.4). Although the shape of the relationship varies within and between species, clearance eventually declines as seston concentration increases over a narrow to broad range (e.g., Hawkins et al. 2001; James et al. 2001; Wong and Cheung 2001b; Hewitt and Pilditch 2004; Velasco and Navarro 2003). Numerous studies have shown that clearance rates exhibit an initial peak at relatively low concentrations, followed by a slow decline (Hawkins et al. 1999; James et al. 2001; Hewitt and Pilditch 2004). The example illustrated in Figure 4.7A from Hawkins et al. (1999) shows that variations in clearance rate of Perna canaliculus exposed to a wide turbidity range could largely be explained by a com-

Clearance rate (L g–1 h–1)

Bivalve filter feeding 105

10

15

20

(B) Low seston

Strohmeier et al. (1999) 6

40

Mytilus edulis

30

4

20 2 10 0

Pecten maximus

0 0.0

1.0

2.0

3.0

Chlorophyll a (µg L–1) Figure 4.7 Clearance rate in relation to food concentration (chlorophyll a; μg L−1) from studies on bivalve species from high (A) and low (B) seston environments. The curves are described by regression equations given in Table 4.4.

bination of Chl and total particulate matter (TPM) concentrations. Rates peaked around 2 μg Chl L−1 and declined at lower and higher concentrations (Fig. 4.7A). James et al. (2001) showed a similar result for this species, except that clearance rates of a population acclimated to low food availability peaked at a considerably lower concentration (∼0.4 μg Chl L−1). This unimodal response to food concentration maximizes food intake during periods of low food availability (Bayne et al. 1987) and can benefit energy intake at higher concentrations by preventing saturation of preingestive particle sorting mechanisms on the ctenidia and labial palps (Iglesias et al. 1992). The majority of bivalve feeding measurements have been conducted in regions where seston concentrations exceed 1 μg Chl L−1 for most of the year, and the general consensus from these studies is that feeding ceases at Chl concentrations between 0.5 and 1 μg L−1

106

Shellfish Aquaculture and the Environment

(Riisgård and Randløv 1981; Newell et al. 2001; Riisgård et al. 2003). The cessation of feeding at low Chl concentrations (<0.86 μg L−1) resulted in tissue loss (Hawkins et al. 1999). A study of Pecten maximus and Mytilus edulis from an area with naturally low seston levels (Norwegian coast), however, showed that both species did not cease feeding even when Chl was as low as 0.01 μg L−1 (Strohmeier et al. 2009; Fig. 4.7B). This indicates that some bivalve species can functionally adapt to prolonged exposure to low-seston conditions such that their biogeographical distribution is not constrained simply by their ability to clear dilute particles from suspension. Studies on bivalves residing in oligotrophic waters have reported some of the highest feeding rates for the given species measured under natural dietary conditions (Essink et al. 1989; Hawkins et al. 1998; Yukihira et al. 1998; Loret et al. 2000; Pouvreau et al. 1999, 2000; Maire et al. 2007; Strohmeier et al. 2009). This is apparently a necessary adaptation for growth and survival in low seston conditions (Pouvreau et al. 1999). Controlled laboratory experiments provide valuable insights into the responses of molluscan shellfish to specific conditions but have a limited capacity to duplicate the natural environment where food quantity and quality, and the ambient flow regime, can vary dramatically over different timescales. Although observations under laboratory conditions are needed to isolate specific responses, the knowledge required to predict temporal changes in food acquisition by bivalves must also come from field studies where feeding behavior may be affected by multiple variables. An example of the importance of obtaining clearance rate estimates under strictly in situ conditions is illustrated by the results shown in Figure 4.2D. Simultaneous time series of the feeding activity (siphon area) of mussels were markedly different for animals in feeding chambers compared with those in a natural benthic setting (Newell et al. 2005). Mussels in the chambers fed at a

consistent and relatively high rate, while the animals on the bottom exhibited large tidal variations in feeding activity. This may have resulted from location-specific differences in the food supply, exposure to flow conditions in the chambers that were unrepresentative of ambient conditions, and/or a response of the animals to experimental manipulation. In a review that highlighted the high plasticity of bivalve physiological processes, Bayne (1998) postulated “. . . the a priori expectation (and the hypotheses that flow from it) is that flexibility will lead to some measurable level of compensation in feeding behaviour, where compensation is defined as a change in a physiological process, in response to a change in feeding conditions, which results in an increase in nutritional uptake over the rate that would apply if no compensatory change occurred.” An endogenous factor potentially influencing clearance rate in nature, therefore, is the instinctive feeding strategy of a given bivalve species. There appears to exist a continuum of strategies ranging from species that primarily regulate clearance rate (e.g., Venerupis pullastra, Mercenaria mercenaria, Mya arenaria, and Pinctada margaritifera; Foster-Smith 1975; Bricelj and Malouf 1984; Bacon et al. 1998; Hawkins et al. 1998), to those that mainly regulate the production of pseudofeces (e.g., Mytilus edulis and Cardium edule; Foster-Smith 1975; Bayne 1993; Navarro et al. 1994; Hawkins et al. 1996), to others that employ both mechanisms to varying degrees (e.g., Atrina zelandica, Cerastoderma edule, Crassostrea gigas, Mulinia edulis, Mytilus chilensis, Placopecten magellanicus, and Perna viridis; Cranford and Gordon 1992; Navarro et al. 1992; Soletchnik et al. 1996; Urrutia et al. 1996; Barillé et al. 1997; Iglesias et al. 1996; Bacon et al. 1998; Hawkins et al. 1998; Velasco and Navarro 2002; Hewitt and Pilditch 2004). The strategy employed also appears to vary between geographic locations for the same species (e.g., Crassostrea gigas; Ren et al. 2000). In areas where the nutritional

Bivalve filter feeding 107

quality of the seston is generally high, there is little bioenergetic benefit to regulating particle selection (pseudofeces production), so the regulation of clearance rate with changing food abundance is the more effective strategy. Bivalves residing in more turbid regions with variable seston conditions benefit from regulating pseudofeces production, but those species that are best suited to high turbidity also regulate clearance rate to maximize energy intake (see Velasco and Navarro 2002). The so-called “bivalve functional response” to diet variability also depends on qualitative aspects of the available seston, which make it difficult to categorize the feeding strategy of many species from the available literature when only a limited range of seston qualities and quantities have been employed. A good indication of the important effect of diet quality on clearance rate can be seen in the difference between average rates measured using algal cell- and seston-based diets. For mussels, scallops, and cockles, laboratory measurements using algal cell cultures resulted in average clearance rates that are 60% higher than rates obtained using seston-based diets (Table 4.4). Feeding behavior is highly stimulated by specific nutritive compounds present in the diet (Navarro et al. 2000) owing, at least in part, to the capacity for chemoreception (Ward et al. 1992). Statistical descriptions of clearance rate responses to natural dietary conditions have been developed for several bivalve species and are summarized in Table 4.5. These models indicate that diet quality, which is generally expressed as seston organic content, is an important factor. However, clearance rate tends to be better explained by changes in quantitative rather than qualitative aspects of the diet (see also Hewitt and Pilditch 2004 and cited papers). Despite an abundance of knowledge on the responses of bivalves to a range of environmental forcing functions, no clear relationships are apparent between seasonal variations in feeding activity and these variables (e.g.,

Deslous-Paoli et al. 1987; Smaal et al. 1997; Cranford and Hill 1999; Hewitt and Pilditch 2004; Strohmeier et al. 2009). Additionally, bivalves residing in adjacent areas with apparently similar conditions can show marked differences in their feeding behavior (Hewitt and Pilditch 2004). The general lack of correlation between clearance rate and ambient environmental conditions may stem from a number of factors including 1. the limited ability of the statistical methods employed to quantify complex interactions between multivariate forcing functions, many of which may cause a nonlinear response; 2. the measurement of bulk seston variables that inadequately characterize food quantity and quality and the stimulatory/ inhibitory properties of seston components on feeding behavior; 3. the inability to continuously measure many potentially important environmental parameters, particularly diet quality; 4. the presence of time-averaged behavior in which the bivalves do not compensate for relatively short-term environmental variations but remain adapted to longer-term conditions; 5. feedback from the regulation of digestive processes that effect feeding behavior; and 6. the additional influence of seasonally variable endogenous nutrient demands on the animal’s energy balance and compensatory feeding responses. The high energy costs of gametogenesis are an obvious, albeit poorly understood, seasonally variable factor potentially forcing the need for compensatory feeding adjustments (Deslous-Paoli et al. 1987; Cranford and Hill 1999). Whereas the majority of bivalve feeding studies have been concerned primarily with determining instantaneous responses to environmental change, the role of past feeding history (e.g., the persistence of feeding behavior in the face

Table 4.5 Significant (P < 0.05) regressions between bivalve clearance rate (C) and the total particulate matter concentration (TPM) and organic content (OC; proportion) of TPM. Species and source

r2

Equation

Applicable range

C (mL g−1 h−1) = 337 × e−0.5(log(TPM-100)/1.1) × 2

50–580 mg TPM L−1

0.81

−1

0.88

−1

0.85

−1

50–580 mg TPM L

0.65

C (L g−1 h−1) = −0.649(TPM) + 6.364

0–10 mg algae L−1

0.80

C (L g−1 h−1) = e(0.3968 − 0.2118(TPM) × OC)

5–22 mg TPM L−1

0.50

−1

Atrina zelandica Hewitt and Pilditch (2004) (Four determinations)

−1

−1

−0.5(log(TPM-86)/0.81) × 2

−1

−1

−0.5(log(TPM-1.06)/160) × 2

−1

−1

−0.5(log(TPM-1.3)/93) × 2

C (mL g h ) = 193 × e C (mL g h ) = 428 × e C (mL g h ) = 360 × e

18–550 mg TPM L 70–600 mg TPM L

Argopecten irradians Palmer (1980) Cerastoderma edule Iglesias et al. (1996) Navarro et al. (1992)

−1

−1

(−0.0409 × Vol + 0.560)

C (L g h ) = e

3–30 mg TPM L

0.66

C (L g−1 h−1) = [91 × (Vol + 1)1.2 × (Chl + 1)2.1 × e(−1.7 × (Vol + 1)) + (−1.4 × (Chl + 1)) + (0.21 × (Vol + 1) × (Chl + 1))] − 1

3.2–105 mg TPM L−1

0.58

C (L h−1 cm−1) = 0.17 × (TPM/(TPM + 0.39)) × e(−0.00378 × TPM)

∼1–650 mg TPM L−1

0.79

C (L g−1 h−1) = 10(2.95 − 0.59 × logTPM − 1.53 × logOC)

2–203 mg TPM L−1

0.82

−1

0.85

Chlamys farreri Hawkins et al. (2001) Crassostrea gigas Ren et al. (2000) Mulinia edulis Velasco and Navarro (2002)

−1

−1

(−0.0054 × TPM)

C (L g h ) = 2.3841 × e

3–200 mg TPM L

Hawkins et al. (1996)

C (L g−1 h−1) = 0.60 + 0.039(TPM)

5–120 mg TPM L−1

0.54

Strohmeier et al. (2009)

C (L h−1) = 5.35 − 0.67(Chl) + 0.56(lnChl) + 0.001/(Chl)

0–3 μg Chl L−1

0.34

C (l g−1 h−1) = 10(2.94 − 0.61 × logTPM − 1.33 × logOC)

2–203 mg TPM L−1

0.83

−1

Velasco and Navarro (2003) Mytilus edulis

Mytilus chilensis Velasco and Navarro (2002)

−1

−1

(−0.0064 × TPM)

C (L g h ) = 1.2423 × e

3–200 mg TPM L

0.92

Hawkins et al. (1999)

C (L g−1 h−1) = 6.81 × (lnChl/1.1)2.71 × e(−2.71 × ((lnChl/1.11) − 1)) × e(−0.19 × lnTPM)

2–3790 mg TPM L−1

0.53

James et al. (2001)*

C (L h−1) = (a × (e(b × Chl)))/(1 + (Chl/c)d)

0.3–0.51 μg Chl L−1

0.83

C (L g−1 h−1) = 12.2 × POM−0.90

2–17 mg POM L−1

0.33

Velasco and Navarro (2003) Perna canaliculus

Perna viridis Hawkins et al. (1998) Wong and Cheung (2001b) Wong and Cheung (2001b)

−1

−1

−1.140

−1

−1

−1.462

C (L g h ) = 4.351 × TPM C (L g h ) = 3.114 × TPM

1–18 mg TPM L

−1

0.68

1–18 mg TPM L

−1

0.64

Pecten maximus Strohmeier et al. (2009)

C (L h−1) = 44.12 − 13.96(Chl) + 5.02(lnChl) + 0.018/(Chl)

0–3 μg Chl L−1

0.44

Seston abundance was measured as chlorophyll a (Chl; μg L−1), total particulate matter (TPM; mg L−1), particle volume (Vol; mm3 L−1), and/or particulate organic matter (POM; mg L−1). *Equation constants not reported.

108

Bivalve filter feeding 109

of further change) and time-dependent physiological acclimation processes in defining food acquisition processes has seldom been addressed (Bayne 1993). It has been suggested that at least some bivalves do not regulate food utilization over the short term, but utilize timeaveraged optimization behavior (Hawkins et al. 1985). For example, if food availability fluctuates rapidly over short-time scales, the animal may waste valuable internal resources responding to every change. Although clearing particles requires minimal energetic cost, the higher costs of digestion and the need to make continuous digestive enzyme adjustments may make continous adaptation an overly wasteful strategy. The time-averaged optimization strategy was supported by time series data on sea scallop feeding responses to storm-induced dietary changes (Cranford et al. 1998).

Emerging knowledge on ecosystem interactions with the bivalve biofilter Research on ecosystem-level interactions with the feeding activity of bivalve populations has focused on the potential control of phytoplankton assemblages and seston via particle depletion, and the consequences of excreted and egested waste products to energy flow and nutrient cycling. Two emerging research topics are briefly summarized here that emphasize the need for consideration of additional effects of biofiltration on coastal ecosystems. These effects further contribute to the characterization of some bivalve molluscs as “ecosystem engineers.”

Transparent exopolymer particle (TEP) production The feeding processes of bivalves can directly affect the water column through removal of suspended particulate matter, that is, biofiltra-

tion. Suspension feeding, however, might have indirect effects on suspended particles as well, further contributing to benthic-pelagic coupling. Studies by Ward and coworkers demonstrate that bivalves and other suspension-feeders can produce significant quantities of TEPs; Mckee et al. 2005; Heinonen et al. 2007; Li et al. 2007). TEPs are discrete, gel-like particles that can be found in both marine and freshwater systems (Alldredge et al. 1998; Passow et al. 2001; Passow 2002). They are composed of high-molecular-weight mucopolysaccharides that are released into the water column by a variety of microorganisms including phytoplankton and bacteria (Decho 1990; Kiørboe and Hansen 1993; Passow and Alldredge 1994; Alldredge et al. 1998). The presence of TEPs can have a substantial impact on coagulation efficiency and flocculation of suspended matter into marine snow (Alldredge et al. 1993; Passow 2002). TEPs can act as a glue that binds together small organic and inorganic material, resulting in the formation of larger aggregates that sink rapidly to the benthos (Alldredge and Silver 1988; Alldredge et al. 1993; Kiørboe et al. 1994; Passow and Alldredge 1995). The abundance and distribution of TEPs in the marine environment has been measured in numerous studies (measured as gum xanthan [GX] equivalents). In general, mean TEP concentrations are lower in open-ocean waters (e.g., 29–512 μg GX L−1, Engel 2004; Wurl and Holmes 2008) and higher in coastal waters (e.g., 100–3500 μg GX L−1; Passow 2002; Wetz et al. 2009). Peak concentrations are associated with phytoplankton blooms, and high concentrations have been measured in near-shore waters inhabited by dense assemblages of corals, macroalgae, mangroves, seagrasses, and suspension-feeders (e.g., 290–>8000 μg GX L−1, Ramaiah et al. 2001, Fabriciusa et al. 2003; Mckee et al. 2005; Wurl and Holmes 2008; Wetz et al. 2009). The concentration of TEPs at any specific location, however, can vary widely depending on

110

Shellfish Aquaculture and the Environment

local physical and biological conditions. One potentially significant source of TEPs in nearshore waters is assemblages of suspensionfeeding bivalves. In laboratory studies, biomass-specific production rates of TEPs from suspension-feeders fall within the range of 0.8–6.7 μg GX g−1 h−1 (Mckee et al. 2005; Heinonen et al. 2007; Li et al. 2007), but can vary due to differences in water pumping activity of the animals. Species-specific TEP production rates under natural conditions, however, may be higher. For example, in a field study using benthic chambers, Mckee et al. (2005) measured a production rate for eastern oysters of 34 μg GX g−1 h−1. Using this field production rate, an oyster with a dry tissue mass of 1 g could produce about 816 μg of TEPs in 24 h. Although the fate and turnover time of TEPs is not entirely known (hours to months; Passow 2002), some of this material is likely transported both vertically and horizontally away from shellfish beds. Bivalve-derived TEPs are known to enhance the aggregation of suspended particulate matter (Li et al. 2007). Therefore, such processes could increase deposition of particulate matter, and enhance benthic-pelagic coupling, in areas devoid of suspension-feeders.

Size-dependent particle retention Grazing of phytoplankton by dense assemblages of bivalves not only has the potential to change phytoplankton biomass but may also affect community composition (Prins et al. 1998; Norén et al. 1999). Bivalves effectively retain particles larger than ∼2–8 μm, depending on the species, and therefore some nanoplankton and all picoplankton (both photoautotrophic and heterotrophic) are not effectively captured and consumed. Picoplankton cells essentially exist within a size range that represents refugia from capture by bivalves and where they may benefit from bivalves depleting

their major competitors and predators (ciliates and flagellates), the increased light availability that accompanies bivalve-mediated seston depletion, and the excretion of ammonia directly into nutrient-depleted summer surface waters by the suspended culture. Picoplankton may become available to the bivalves, however, through particle aggregation processes (Cranford et al. 2005; Kach and Ward 2008), or through linkage to higher tropic levels via the microzooplankton (Loret et al. 2000). Mesocosm studies show that mussel grazing can change phytoplankton species composition, shifting the community to one dominated by picoplankton (Olsson et al. 1992; Prins et al. 1997). Size-selective feeding by extensive bivalve culture is thought to be an important reason why picoplankton cells constitute the most abundant component of the phytoplankton community in the Thau Lagoon in France (Courties et al. 1994; Vaquer et al. 1996; Souchu et al. 2001), and in Tracadie Bay, Canada (Cranford et al. 2008). Picoplankton species have also been shown to dominate the phytoplankton biomass in several embayments supporting extensive suspended mussel culture in Prince Edward Island (50–80% of total Chl), but not in an adjacent unfarmed bay where the microphytoplankton dominate (Cranford et al. 2008). That study showed a close relationship between the average baywide picophytoplankton contribution and an index of bay-scale seston depletion by mussel culture. Future research is needed to assess the ecosystem consequences of this top-down destabilization of the food chain, including possible changes in predator–prey relationships and competitive interactions that could result in trophic regime shifts. A shift from microphytoplankton to picophytoplankton biomass could affect particle transport dynamics via reduced settling velocity and altered flocculation processes. The latter is dependent on the production of sticky exopolymers that can be produced in large quantities by diatoms

Bivalve filter feeding 111

(see also previous section). These potential ecological effects of shellfish culture need to be better understood and considered in the determination of carrying capacity predictions and when considering implementing schemes that use bivalve farms for combating eutrophication (Lindahl et al. 2005).

Conclusions Forty years ago, Brian Morton wrote: “The view that bivalve molluscs are ideally adapted to fulfilling a mode of life in which the processes of feeding and digestion are continuous has long been accepted by most zoologists. An increasing amount of evidence, however, is being put forward to suggest that the opposite is true, that is, that bivalves are discontinuous in their feeding and digestive habits” (Morton 1970). This evidence has since increased dramatically and, with few exceptions, reveals a remarkable capacity of bivalve suspensionfeeders to finely adjust clearance rate as opposed to simply switching between feeding and nonfeeding states (see Fig. 4.8). The above meta-analysis of the contemporary literature demonstrates that accurate and comparable estimates of clearance rate can easily be obtained using many direct and indirect methodologies using familiar precautions. A major methodological pitfall stems from the application of artificial dietary conditions that stimulate a predetermined (e.g., maximal) feeding response as a basis for developing theories on bivalve feeding behavior in nature. This approach represents an experimental bias that defies basic scientific principles, but has been a prerequisite for all studies that continue to support the autonomous view of bivalve feeding. Whether or not the exhibited high flexibility in clearance rates constitutes a homeostatic strategy to maximize the individuals net energy balance is outside the scope of this review. However, the preponderance of

the literature demonstrates that the opposing theory of autonomous feeding is obsolete. Bivalve feeding behavior has important implications for aquaculture, including the optimization of farm location and layout, the forecasting of bivalve growth and carrying capacity, and the determination of potential ecological services and impacts. An abundance of information is available on the clearance rate responses of bivalves in nature that can support these applications. Clearance rates are highly variable across short to long temporal scales and large near- and far-field differences in feeding behavior occur within the same species. Predicting wild and farmed bivalve population responses to environmental variability over different scales is a major challenge. For clearance rate measurements to be relevant to addressing aquaculture issues, they need to address scales of variation that are relevant to the specific question being addressed. Incorporating additional elements of environmentally induced physiological regulation into bivalve growth models is challenging as it is difficut for the models to deal with fine-scale temporal variations in the seston (Grant 1996). Despite this shortcoming, the current modeling capacity appears sufficient for accurately replicating observed seasonal growth responses of bivalves (Chapter 6 in this book), and also appears acceptable for accurately predicting effects of feeding on ambient food supplies (see above). Nonetheless, siteand time-specific measurements of clearance rate are encouraged whenever possible to help improve or to test model applications. These measurements will increase confidence among aquaculture stakeholders on the practical and regulatory applications of population-level clearance calculations. They will also improve ecophysiological and ecosystem model predictions and will increase capacity to address more specific questions related to fine-scale changes in feeding behavior. For example, greater spatial resolution within models would

112 Figure 4.8 Blue mussels, Mytilus edulis, actively filtering seawater. (Courtesy of Tore Strohmeier and Øivind Strand.)

Bivalve filter feeding 113

permit more quantitative assessments of optimal farm site location/layout and multifarm interactions, whereas increased temporal resolution will aid in predicting seasonally variable bivalve controls on the phytoplankton. Additional work is needed across a wide range of biogeographical settings, timescales, and species to further resolve details on the interactions between bivalves and their environment, and the way in which feeding responds to quantitative and qualitative dietary stimuli.

Literature cited Ackerman, J.D. 1999. Effect of velocity on the filter feeding of dreissenid mussels (Dreissena polymorpha and Dreissena bugensis): implications for trophic dynamics. Canadian Journal of Fisheries and Aquatic Sciences 56:1551–1561. Ackerman, J.D., and Nishizaki, M.T. 2004. The effect of velocity on the suspension feeding and growth of the marine mussels Mytilus trossulus and M. californianus: implications for niche separation. Journal of Marine Systems 49:195– 207. Alldredge, A.L., and Silver, M.W. 1988. Characteristics, dynamics and significance of marine snow. Progress in Oceanography 20: 41–82. Alldredge, A.L., Passow, U., and Logan, B.E. 1993. The abundance and significance of a class of large, transparent organic particles in the ocean. Deep-Sea Res 40:1131–1140. Alldredge, A.L., Passow, U., and Haddock, S.H.D. 1998. The characteristics and transparent exopolymer particle (TEP) content of marine snow formed from thecate dinoflagellates. Journal of Plankton Research 20:393–406. Bacon, G.S., MacDonald, B.A., and Ward, J.E. 1998. Physiological responses of infaunal (Mya arenaria) and epifaunal (Placopecten magellanicus) bivalves to variations in the concentration and quality of suspended particles I. Feeding activity and selection. Journal of Experimental Marine Biology and Ecology 219:105–125. Barillé, L., Prou, J., Heral, M., and Razet, D. 1997. Effects of high natural seston concentrations on

the feeding, selection, and absorption of the oyster Crassostrea gigas. Journal of Experimental Marine Biology and Ecology 212:149–172. Barillé, L., Haure, J., Pales-Espinosa, E., and Morançais, M. 2003. Finding new diatoms for intensive rearing of the pacific oyster (Crassostrea gigas): energy budget as a selective tool. Aquaculture 217:501–514. Bauder, A.G., Cembella, A.D., Bricelj, V.M., and Quilliam, M.A. 2001. Uptake and fate of diarrhetic shellfish poisoning toxins from the dinoflagellate Procentrum lima in the bay scallop Argopecten irradians. Marine Ecology Progress Series 213:39–52. Bayne, B.L. 1993. Feeding physiology of bivalves: time-dependence and compensation for changes in food availability. In: Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Marine Ecosystem Processes, NATO ASI Series, Vol. G 33. SpringerVerlag, Berlin, pp. 1–24. Bayne, B.L. 1998. The physiology of suspension feeding bivalve molluscs: an introduction to the Plymouth “TROPHEE” workshop. Journal of Experimental Marine Biology and Ecology 219:1–19. Bayne, B.L. 1999. Physiological components of growth differences between individual oysters (Crassostrea gigas) and a comparison with Saccostrea commercialis. Physiological and Biochemical Zoology 72:705–713. Bayne, B.L. 2001. Reply to comment by H.U. Riisgård. Ophelia 54:211. Bayne, B.L. 2004. Comparisons of measurements of clearance rates in bivalve molluscs. Marine Ecology Progress Series 276:305–306. Bayne, B.L., and Widdows, J. 1978. The physiological ecology of two populations of Mytilus edulis L. Oecologia 37:137–162. Berry, P.F., and Schleyer, M.H. 1983. The brown mussel Perna perna on the Natal coast, South Africa: utilization of available food and energy budget. Marine Ecology Progress Series 13:201–210. Bayne, B.L., Klumpp, D.W., and Clarke, K.R. 1984. Aspects of feeding, including estimates of gut residence time, in three mytilid species (Bivalvia, Mollusca) at two contrasting sites in the Cape Peninsula, South Africa. Oecologia 64:26–33. Bayne, B.L., Brown, D.A., Burns, K., Dixon, D.R., Ivanovici, A., Livingstone, D.R., Lowe, D.M.,

114

Shellfish Aquaculture and the Environment

Moore, M.N., Stebbing, A.R.D., and Widdows, J. 1985. The Effects of Stress and Pollution on Marine Animals. Chapter 7: Physiological Proceedures. Praeger, New York, pp. 161–178. Bayne, B.L., Hawkins, A.J.S., and Navarro, E. 1987. Feeding and digestion by the mussel Mytilus edulis L. (Bivalvia: Mollusca) in mixtures of silt and algal cells at low concentrations. Journal of Experimental Marine Biology and Ecology 111:1–22. Bayne, B.L., Hawkins, A.J.S., Navarro, E., and Iglesias, I.P. 1989. Effects of seston concentration on feeding, digestion and growth in the mussel Mytilus edulis. Marine Ecology Progress Series 55:47–54. Bayne, B.L., Svensson, S., and Nell, J.A. 1999. The physiological basis for faster growth in the Sydney rock oyster, Saccostrea commercialis. The Biological Bulletin 197:377–387. Bougrier, S., Geairon, P., Deslous-Paoli, J.M., Bacher, C., and Jonquières, G. 1995. Allometric relationships and effects of temperature on clearance and oxygen consumption rates of Crassostrea gigas (Thunberg). Aquaculture 134:143–154. Bricelj, V.M., and Malouf, R.E. 1984. Influence of algal and suspended sediment concentrations on the feeding physiology of the hard clam Mercenaria mercenaria. Marine Biology 84:155–165. Buxton, C.D., Newell, R.C., and Field, J.G. 1981. Response-surface analysis of the combined effects of exposure and acclimation temperatures on filtration, oxygen consumption and scope for growth in the oyster Ostrea edulis. Marine Ecology Progress Series 6:73–82. Chaparro, O.R., and Thompson, R.J. 1998. Physiological energetics of brooding in Chilean oyster Ostrea chilensis. Marine Ecology Progress Series 171:151–163. Chipman, W.A., and Hopkins, J.G. 1954. Water filtration by the bay scallop, Pecten irradians, as observed with the use of radioactive plankton. Biological Bulletin, Marine Biological Laboratory, Woods Hole 107:80–91. Clausen, I., and Riisgård, H.U. 1996. Growth, filtration and respiration in the blue mussel, Mytilus edulis: no evidence for physiological regulation of the filter-pump. Marine Ecology Progress Series 141:34–45.

Cole, B.E., Thompson, J.K., and Cloern, J.E. 1992. Measurement of filtration rates by infaunal bivalves in a recirculating flume. Marine Biology 113:219–225. Coughlan, J. 1969. The estimation of filtration rate from the clearance of suspensions. Marine Biology 2:256–358. Courties, C., Vaquer, A., Troussellier, M., Lautier, J., Chrétiennot-Dinet, M.J., Neveux, J., Machado, C., and Claustre, H. 1994. Smallest eukaryotic organism. Nature 370:255. Cranford, P.J. 2001. Evaluating the “reliability” of filtration rate measurements in bivalves. Marine Ecology Progress Series 215:303–305. Cranford, P.J., and Grant, J. 1990. Particle clearance and absorption of phytoplankton and detritus by the sea scallop Placopecten magellanicus (Gmelin). Journal of Experimental Marine Biology and Ecology 137:105–121. Cranford, P.J., and Gordon, D.C. 1992. The influence of dilute clay suspensions on sea scallop (Placopecten magellanicus) feeding activity and tissue growth. Netherlands Journal of Sea Research 30:107–120. Cranford, P.J., and Hargrave, B.T. 1994. In situ time-series measurement of ingestion and absorption rates of suspension-feeding bivalves: Placopecten magellanicus (Gmelin). Limnology and Oceanography 39:730–738. Cranford, P.J., and Hill, P.S. 1999. Seasonal variation in food utilization by the suspension-feeding bivlave molluscs Mytilus edulis and Placopecten magellanicus. Marine Ecology Progress Series 190:223–239. Cranford, P.J., Emerson, C.W., Hargrave, B.T., and Milligan, T.G. 1998. In situ feeding and absorption responses of sea scallops Placopecten magellanicus (Gmelin) to storm-induced changes in the quantity and composition of the seston. Journal of Experimental Marine Biology and Ecology 219:5–70. Cranford, P.J., Armsworthy, S.L., Mikkelsen, O., and Milligan, T.G. 2005. Food acquisition responses of the suspension-feeding bivalve Placopecten magellanicus to the flocculation and settlement of a phytoplankton bloom. Journal of Experimental Marine Biology and Ecology 326:128–143. Cranford, P.J., Li, W., Strand, Ø., and Strohmeier, T. 2008. Phytoplankton depletion by mussel

Bivalve filter feeding 115

aquaculture: high resolution mapping, ecosystem modeling and potential indicators of ecological carrying capacity. ICES CM Document 2008/H:12. 5 p. www.ices.dk/products/CMdocs/ CM-2008/H/H1208.pdf Davenport, J., and Woolmington, A.D. 1982. A new method of monitoring ventilatory activity in mussels and its use in a study of the ventilatory patterns of Mytilus edulis L. Journal of Experimental Marine Biology and Ecology 62:55–67. Decho, A.W. 1990. Microbial exopolymer secretion in ocean environments: their role in food webs and marine processes. Oceanography Marine Biology Annual Annual Review 28:73–153. Dellatorre, F.G., Pascual, M.S., and Barón, P.J. 2007. Feeding physiology of the Argentine mussel Mytilus edulis platensis (d’Orbigny, 1846): does it feed faster in suspended culture systems? Aquaculture International 15:415– 424. Deslous-Paoli, J.-M., Héral, M., Goulletquer, P., Boromthanarat, W., Razet, D., Garnier, J., Prou, J., and Barillet, L. 1987. Evolution saisonniere de la filtration de bivalves intertidaux dans des conditions naturelles. Océanis 13(4–5):575– 579. Doering, P.H., and Oviatt, C.A. 1986. Application of filtration rate models to field populations of bivalves: an assessment using experimental mesocosms. Marine Ecology Progress Series 31:265–275. Dolmer, P. 2000. Feeding activity of mussels Mytilus edulis related to near-bed currents and phytoplankton biomass. Journal of Sea Research 44:221–231. Dowd, M. 1997. On predicting the growth of cultured bivalves. Ecological Modelling 104:113– 131. Dunphy, B.J., Hall, J.A., Jeffs, A.G., and Wells, R.M.G. 2006. Selective particle feeding by the Chilean oyster, Ostrea chilensis; implications for nursery culture and broodstock conditioning. Aquaculture 261:594–602. Dupuy, C., Vaquer, A., Lam-Höai, T., Rougier, C., Mazouni, N., Lautier, J., Collos, Y., and Le Gall, S. 2000. Feeding rate of the oyster Crassostrea gigas in a natural phytoplankton community of the Mediterranean Thau Lagoon. Marine Ecology Progress Series 205:171–184.

Engel, A. 2004. Distribution of transparent exopolymer particles (TEP) in the Northeast Atlantic Ocean and their potential significance for aggregation processes. Deep-Sea Research 51:83– 92. Essink, K., Tydeman, P., de Koning, F., and Kleef, H.L. 1989. On the adaptation of the mussel Mytilus edulis L. to different environmental suspended matter concentrations. In: Klekowski, R.Z., Styczynska-Jurewicz, E., and Falkowski, L. (eds.), Proc. 21st Eur. Mar. Biol. Symp. Gdansk, Poland, 1988. Ossolineum, Gdansk, 41–51. Fabriciusa, K.E., Wildb, C., Wolanskia, E., and Abelec, D. 2003. Effects of transparent exopolymer particles and muddy terrigenous sediments on the survival of hard coral recruits. Estuarine, Coastal and Shelf Science 57:613–621. Famme, P., Riisgård, H.U., and Jorgensen, C.B. 1986. On direct measurements of pumping rates in the mussel Mytilus edulis. Marine Biology 92:323–327. Fegley, S.R., MacDonald, B.A., and Jacobsen, T.R. 1992. Short-term variation in the quantity and quality of seston available to benthic suspension feeders. Estuarine, Coastal and Shelf Science 34:393–412. Filgueira, R., Labarta, U., and Fernandez-Reiriz, M.J. 2006. Flow-through chamber method for clearance rate measurements in bivalves: design and validation of individual chambers and mesocosm. Limnology and Oceanography Methods 4:284–292. Filgueira, R., Labarta, U., and Fernández-Reiríz, M.J. 2008. Effect of condition index on allometric relationships of clearance rate in Mytilus galloprovincialis Lamarck, 1819. Revista de Biología Marina y Oceanografia 43(2):391– 398. Foster-Smith, R.L. 1975. The effect of concentration of suspension on the filtration rates and pseudofaecal production for Mytilus edulis (L.), Cerastoderma edule (L.), and Venerupis pullastra (Montaga). Journal of Experimental Marine Biology and Ecology 17:1–22. Frank, D.M., Hamilton, J.F., Ward, J.E., and Shumway, S. 2007. A fiber optic sensor for high resolution measurement and continuous monitoring of valve gape in bivalve molluscs. Journal of Shellfish Research 26:575–580.

116

Shellfish Aquaculture and the Environment

Frank, D.M., Ward, J.E., Shumway, S.E., Gray, C., and Holohan, B.A. 2008. Application of particle image velocimetry to the study of suspension feeding in marine invertebrates. Marine and Freshwater Behaviour and Physiology 41:1–18. Fréchette, M., and Bacher, C. 1998. A modelling study of optimal stocking density of mussel populations kept in experimental tanks. Journal of Experimental Marine Biology and Ecology 219:241–255. Fréchette, M., and Bourget, E. 1987. Significance of small-scale spatio-temporal heterogeneity in phytoplankton abundance for energy flow in Mytilus edulis. Marine Biology 94:231–240. Fréchette, M., Booth, D.A., Myrand, B., and Bérard, H. 1991. Variability and transport of organic seston near a mussel aquaculture site. ICES Marine Science Symposium 192:24–32. Gainey, L.F., Walton, J.C., and Greenberg, M.J. 2003. Branchial musculature of a venerid clam: pharmacology, distribution, and innervation. The Biological Bulletin 204:81–95. Galtsoff, P.S. 1926. New methods to measure the rate of flow produced by the gills of oysters and other Mollusca. Science 63:233–234. Gardner, J.P.A., and Thompson, R.J. 2001. The effects of coastal and estuarine conditions on the physiology and survivorship of the mussel Mytilus edulis and M. trossulus and their hybrids. Journal of Experimental Marine Biology and Ecology 265:119–140. Gerdes, D. 1983. The Pacific oyster Crassostrea gigas Part I. Feeding behaviour of larvae and adults. Aquaculture 31:195–219. Gibbs, M.T. 2007. Sustainability performance indicators for suspended bivalve aquaculture activities. Ecological Indicators 7:94–107. Grant, J. 1996. The relationship of bioenergetics and the environment to the field growth of cultured bivalves. Journal of Experimental Marine Biology and Ecology 200:239–256. Grant, J., Bacher, C., Cranford, P.J., Guyondet, T., and Carreau, M. 2008. A spatially explicit ecosystem model of seston depletion in dense mussel culture. Journal of Marine Systems 73:155– 168. Griffiths, R.J. 1980. Filtration, respiration and assimilation in the black mussel Choromytilus meridionalis. Marine Ecology Progress Series 3:63–70.

Haure, J., Penisson, C., Bougrier, S., and Baud, J.P. 1998. Influence of temperature on clearance and oxygen consumption rates of the flat oyster Ostrea edulis: determination of allometric coefficients. Aquaculture 169:211–224. Haure, J., Huvet, A., Palvadeau, H., Nourry, M., Penisson, C., Martin, J.L.Y., and Boudry, P. 2003. Feeding and respiratory time activities in the cupped oysters Crassostrea gigas, Crassostrea angulata and their hybrids. Aquaculture 218:539–551. Hawkins, A.J.S., and Bayne, B.L. 1992. Physiological interrelations, and the regulation of production. In: Gosling, E. (ed.), The Mussel Mytilus: Ecology, Physiology, Genetics and Culture. Elsevier, Amsterdam, pp. 171–222. Hawkins, A.J.S., Salkeld, P.N., Bayne, B.L., Gnaiger, E., and Lowe, D.M. 1985. Feeding and resource allocation in the mussel Mytilus edulis: evidence for time-averaged optimization. Marine Ecology Progress Series 20:273–287. Hawkins, A.J.S., Navarro, E., and Iglesias, J.I.P. 1990. Comparative allometries of gut-passage time, gut content and metabolic faecal loss in Mytilus edulis and Cerastoderms edule. Marine Biology 105:197–204. Hawkins, A.J.S., Smith, R.F.M., Bayne, B.L., and Héral, M. 1996. Novel observations underlying the fast growth of suspension-feeding shellfish in turbid environments: Mytilus edulis. Marine Ecology Progress Series 131:179–190. Hawkins, A.J.S., Smith, R.F.M., Bougrier, S., Bayne, B.L., and Héral, M. 1997. Manipulation of dietary conditions for maximal growth in mussels, Mytilus edulis, from the MarennesOléron Bay, France. Aquatic Living Resources 10:13–22. Hawkins, A.J.S., Smith, R.F.M., Tan, S.H., and Yasin, Z.B. 1998. Suspension-feeding behaviour in tropical bivalve molluscs: Perna viridis, Crassostrea belcheri, Crassostrea iradelei, Saccostrea cucculata and Pinctada margarifera. Marine Ecology Progress Series 166:173– 185. Hawkins, A.J.S., James, M.R., Hickman, R.W., Hatton, S., and Weatherhead, M. 1999. Modelling of suspension-feeding and growth in the green-lipped mussel Perna canaliculus exposed to natural and experimental variations of seston availability in the Marlborough Sounds,

Bivalve filter feeding 117

New Zealand. Marine Ecology Progress Series 191:217–232. Hawkins, A.J.S., Fang, J.G., Pascoe, P.L., Zhang, J.H., Zhang, X.L., and Zhu, M.Y. 2001. Modelling short-term responsive adjustments in particle clearance rate among bivalve sispensionfeeders: separate unimodal effects of seston volume and composition in the scallop Chlamys farreri. Journal of Experimental Marine Biology and Ecology 262:61–73. Hawkins, A.J.S., Duarte, P., Fang, J.G., Pascoe, P.L., Zhang, J.H., Zhang, X.L., and Zhu, M.Y. 2002. A functional model of responsive suspensionfeeding and growth in bivalve shellfish, configured and validated for the scallop Chlamys farreri during culture in China. Journal of Experimental Marine Biology and Ecology 281:13–40. Heinonen, K.B., Ward, J.E., and Holohan, B.A. 2007. Production of transparent exopolymer particles (TEP) by benthic suspension feeders in coastal systems. Journal of Experimental Marine Biology and Ecology 341:184–195. Hewitt, J.E., and Pilditch, C.A. 2004. Environmental history and physiological state influence feeding responses of Atrina zelandica to suspended sediment concentrations. Journal of Experimental Marine Biology and Ecology 306:95–112. Hilbish, T.J. 1986. Growth trajectories of shell and soft tissue in bivalves: seasonal variation in Mytilus edulis L. Journal of Experimental Marine Biology and Ecology 96:103–113. Ibarrola, I., Navarro, E., and Iglesias, J.I.P. 1998. Short-term adaptation of digestive processes in the cockle Cerastoderma edule exposed to different food quantity and quality. Journal of Comparative Physiology B 168:32–40. Ibarrola, I., Navarro, E., and Urrutia, M.B. 2000. Acute and acclimated digestive responses of the cockle Cerastoderma edule (L.) to changes in food quality and quantity. Journal of Experimental Marine Biology and Ecology 252:181–198. Iglesias, J.I.P., Navarro, E., Alvarez, P.J., and Armentia, Y. 1992. Feeding, particle selection and absorption in cockles Cerastoderma edule (L) exposed to variable conditions of food concentration and quality. Journal of Experimental Marine Biology and Ecology 162:177–198.

Iglesias, J.I.P., Urrutia, M.B., Navarro, E., AlvarezJorna, P., Larretxea, X., Bougrier, S., and Héral, M. 1996. Variability of feeding processes in the cockle Cerastoderma edule (L.) in response to changes in seston concentration and composition. Journal of Experimental Marine Biology and Ecology 197:121–143. Iglesias, J., Urrutia, M., Navarro, E., and Ibarrola, I. 1998. Measuring feeding and absorption in suspension-feeding bivalves: an appraisal of the biodeposition method. Journal of Experimental Marine Biology and Ecology 219:71–86. James, M.R., Weatherhead, M.A., and Ross, A.H. 2001. Size-specific clearance, excretion, and respiration rates, and phytoplankton selectivity for the mussel Perna canaliculus at low levels of natural food. New Zealand Journal of Marine and Freshwater Research 35:73–86. Jihong, Z., Fang, J.G., Hawkins, A.J.S., and Pascoe, P.L. 2004. The effect of temperature on clearance rate and oxygen consumption of scallops, Chlamys ferreri. Journal of Shellfish Research 23:715–721. Jones, H.D., Richards, O.G., and Southern, T.A. 1992. Gill dimensions, water pumping rate and body size in the mussel Mytilus edulis. Journal of Experimental Marine Biology and Ecology 155:213–237. Jørgenson, C.B. 1990. Bivalve Filter Feeding: Hydrodynamics, Bioenergetics, Physiology and Ecology. Olsen and Olsen, Fredensborg, Denmark. Jørgenson, C.B. 1996. Bivalve filter feeding revisited. Marine Ecology Progress Series 142:287– 302. Jørgenson, C.B., Larsen, P.S., Møhlenberg, M., and Riisgård, H.U. 1988. The bivalve pump: properties and modelling. Marine Ecology Progress Series 45:205–216. Jørgenson, C.B., Larsen, P.S., and Riisgård, H.U. 1990. Effects of temperature on the mussel pump. Marine Ecology Progress Series 64:89– 97. Kach, D.J., and Ward, J.E. 2008. The role of marine aggragates in the ingestion of picoplankton-size particles by suspension-feeding molluscs. Marine Biology 153:797–805. Kesarcodi-Watson, A., Lucas, J.S., and Klumpp, D.W. 2001. Comparative feeding and physiolog-

118

Shellfish Aquaculture and the Environment

ical energetics of diploid and triploid Sydney rock oysters, Saccostrea commercialis I. Effects of oyster size. Aquaculture 203:177–193. Kiørboe, T., and Hansen, J.L.S. 1993. Phytoplankton aggregate formation: observations of patterns and mechanisms of cell sticking and the significance of exopolymer material. Journal of Plankton Research 15:993–1018. Kiϕrboe, T., Mϕhlenberg, F., and Nϕhr, O. 1980. Feeding, particle selection and carbon absorption in Mytilus edulis in different mixtures of algae and resuspended bottom material. Ophelia 19:193–205. Kiϕrboe, T., Mϕhlenberg, F., and Nϕhr, O. 1981. Effect of suspended bottom material on growth and energetics in Mytilus edulis. Marine Biology 61:283–288. Kiørboe, T., Lundsgaard, C., Olesen, M., and Hansen, J.L.S. 1994. Aggregation and sedimentation processes during a spring phytoplankton bloom: a field experiment to test coagulation theory. Journal of Marine Research 52:297– 323. Kittner, C., and Riisgård, H.U. 2005. Effect of temperature on filtration rate in the mussel Mytilus edulis: no evidence for temperature compensation. Marine Ecology Progress Series 305:147– 152. Labarta, U., Fernández-Reiríz, M.J., and Babarro, J.M.F. 1997. Differences in physiological energetics between intertidal and raft cultivated mussels Mytilus galloprovincialis. Marine Ecology Progress Series 152:167–173. Lefebvre, S., Barillé, L., and Clerc, M. 2000. Pacific oyster (Crassostrea gigas) feeding responses to a fish-farm effluent. Aquaculture 187:185– 198. Li, S.-C., Wang, W.-X., and Hsieh, D.P.H. 2001. Feeding and absorption of the toxic dinoflagellate Alexandrium tamarense by two marine bivalves from the south China Sea. Marine Biology 139:617–624. Li, B.L., Ward, J.E., and Holohan, B.A. 2007. Effect of transparent exopolymer particles (TEP) from suspension feeders on particle aggregation. Marine Ecology Progress Series 357:67–77. Li, Y., Veilleux, D.J., and Wikfors, G.H. 2009. Particle removal by Northern bay scallops Argopecten irradians irradians in a semi-natural

setting: application of a flow-cytometric technique. Aquaculture 296:237–245. Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L.-O., Olrog, L., Rehnstam-Holm, A.-S., Svensson, J., Svensson, S., and Syversen, U. 2005. Improving marine water quality by mussel farming—a profitable solution for Swedish society. Ambio 34:131–138. Loret, P., Gall, S.L., Dupuy, C., Blanchot, J., Pastoureaud, A., Delesalle, B., Caisey, X., and Jonquières, G. 2000. Heterotrophic protists as a trophic link between picocyanobacteria and the pearl oyster Pincada margaritifera in the Takapoto lagoon (Tuamotu Archipelago, French Polynesia). Aquatic Microbial Ecology 22:215–226. Lucas, M.I., Newell, R.C., Shumway, S.E., Seiderer, L.J., and Bally, R. 1987. Particle clearance and yield in relation to bacterioplankton and suspended particulate availability in estuarine and open coast populations of the mussel Mytilus edulis. Marine Ecology Progress Series 36: 215–224. MacDonald, B.A., and Thompson, R.J. 1986. Influence of temperature and food availability on the ecological energetics of the giant scallop Placopecten magellanicus. III. Physiological ecology, the gametogenic cycle and scope for growth. Marine Biology 93:37–48. MacDonald, B.A., and Ward, J.E. 1994. Variation in food quality and particle selectivity in the sea scallop Placopecten magellanicus (Mollusca: Bivalvia). Marine Ecology Progress Series 108:251–264. MacDonald, B.A., and Ward, J.E. 2009. Feeding activity of scallops and mussels measured simultaneously in the field: repeated measures sampling and implications for modelling. Journal of Experimental Marine Biology and Ecology 371:42–50. MacDonald, B.A., Robinson, S.M.C., and Barrington, K.A. 2009. Evaluating the use of exhalent siphon area in estimationg feeding activity of blue mussels, Mytilus edulis. Journal of Shellfish Research 28:289–297. Maire, O., Amouroux, J.-M., Duchêne, J.-C., and Grémare, A. 2007. Relationship between filtration activity and food availability in the Mediterranean mussel Mytilus galloprovincialis. Marine Biology 152:1293–1307.

Bivalve filter feeding 119

Mckee, M.P., Ward, J.E., MacDonald, B.A., and Holohan, B.A. 2005. Production of transparent exopolymer particles (TEP) by the eastern oyster, Crassostrea virginica. Marine Ecology Progress Series 288:141–149. McLusky, D.S. 1973. The effect of temperature on the oxygen consumption and filtration rate of Chlamys (Aequipecten) opercularis (L.) (Bivalvia). Ophelia 10:141–154. Medler, S., and Silverman, H. 2001. Muscular alteration of gill geometry in vitro: implications for bivalve pumping processes. The Biological Bulletin 200:77–86. Meyhöfer, E. 1985. Comparative pumping rates in suspension-feeding bivalves. Marine Biology 85:137–142. Møhlenberg, F., and Riisgård, H.U. 1979. Filtration rate, using a new indirect technique, in thirteen species of suspension-feeding bivalves. Marine Biology 54:143–147. Morton, B. 1970. The tidal rhythm and rhythm of feeding and digestion in Cardium edule. Journal of the Marine Biological Association of the United Kingdom 50:499–512. Nakagawa, S., and Cuthill, I.C. 2007. Effect size, confidence interval and statistical significance: a practical guide for biologists. Biology Reviews of the Cambridge Philosophical Society 82: 591–605. Navarro, J.M., and Widdows, J. 1997. Feeding physiology of Cerastoderma edule in response to a wide range of seston concentrations. Marine Ecology Progress Series 152:175–186. Navarro, J.M., and Winter, J.E. 1982. Ingestion rate, assimilation efficiency and energy balance in Mytilus chilensis in relation to body size and different algal concentrations. Marine Biology 67:255–266. Navarro, J.M., and Velasco, L.A. 2003. Comparison of two methods for measuring filtration rate in filter feeding bivalves. Journal of the Marine Biological Association of the United Kingdom 83:553–558. Navarro, E., Iglesias, J.I.P., Perez Camacho, A., Labarta, U., and Beiras, R. 1991. The physiological energetics of mussels (Mytilus galloprovincialis Lmk) from different cultivation rafts in the Ria de Arosa (Galicia, N.W. Spain). Aquaculture 94:197–212.

Navarro, E., Iglesias, J.I.P., and Ortega, M.M. 1992. Natural sediment as a food source for the cockle Cerastoderma edule (L.): effect of variable particle concentration on feeding, digestion and the scope for growth. Journal of Experimental Marine Biology and Ecology 156:69–87. Navarro, E., Iglesias, J.I.P., Ortega, M.M., and Larretxea, X. 1994. The basis for a functional response to variable food quantity and quality in cockles Cerastoderma edule (Bivalvia Cardiidae). Physiological Zoology 67:468– 496. Navarro, E., Iglesias, J.I.P., Perez Camacho, A., and Labarta, U. 1996. The effect of diets of phytoplankton and suspended bottom material on feeding and absorption of raft mussels (Mytilus galloprovincialis Lmk). Journal of Experimental Marine Biology and Ecology 198:175–189. Navarro, J.M., and González, C.M. 1998. Physiological responses of the Chilean scallop Argopecten purpuratus to decreasing salinities. Aquaculture 167:315–327. Navarro, J.M., Leiva, G.E., Martinez, G., and Aguilera, C. 2000. Interactive effects of diet and temperature on the scope for growth of the scallop Argopecten purpuratus during reproductive conditioning. Journal of Experimental Marine Biology and Ecology 247:67–83. Navarro, E., Labarta, U., Fernández-Reiríz, M.J., and Velasco, A. 2003. Feeding behaviour and differential absorption of biochemical components by the infaunal bivalve Mulinia edulis and the epibenthic Mytilus chilensis in response to changes in food regimes. Journal of Experimental Marine Biology and Ecology 287:13–35. Newell, R.I.E., and Bayne, B.L. 1980. Seasonal changes in the physiology, reproductive condition and carbohydrate content of the cockle Cardium (=Cerastoderma) edule (Bivalvia: Cardiidae). Marine Biology 56:11–19. Newell, R.C., Johnson, L.G., and Kofoed, L.H. 1977. Adjustment of the components of energy balance in response to temperature change in Ostrea edulis. Oecologia (Berl.) 30:97–110. Newell, C.R., Shumway, S.E., Cucci, T.L., and Selvin, R. 1989. The effect of natural seston particle size and type on feeding rates, feeding selectivity and food resource availability for the

120

Shellfish Aquaculture and the Environment

mussel Mytilus edulis Linnaeus, 1758 at bottom cuture sites in Maine. Journal of Shellfish Research 8(1):187–196. Newell, C.R., Campbell, D.E., and Gallagher, S.M. 1998. Development of the mussel aquaculture lease site model MUSMOD©: a field program to calibrate model formulations. Journal of Experimental Marine Biology and Ecology 219:143–169. Newell, C.R., Pilskaln, C.H., Robinson, S.M., and MacDonald, B.A. 2005. The contribution of marine snow to the particle food supply of the benthic suspension feeder, Mytilus edulis. Journal of Experimental Marine Biology and Ecology 321:109–124. Newell, C.R., Wildish, D.J., and MacDonald, B.A. 2001. The effects of velocity and seston concentration on the exhalent siphon area, valve gape and filtration rate of the mussel Mytilus edulis. Journal of Experimental Marine Biology and Ecology 262:91–111. Norén, F., Haamer, J., and Lindahl, O. 1999. Changes in the plankton community passing a Mytilus edulis bed. Mar. Ecol. Prog. Ser. 191:187–194. Okumus, I., and Stirling, H.P. 1994. Physiological energetics of cultivated mussel (Mytilus edulis) populations in two Scottish west coast sea lochs. Marine Biology 119:125–131. Olsson, P., Graneli, E., Carlsson, P., and Abreu, P. 1992. Structuring of a postspring phytoplankton community by manipulation of trophic interactions. Journal of Experimental Marine Biology and Ecology 158:249–266. Palmer, R.E. 1980. Behaviour and rhythmic aspects of filtration in the bay scallop, Argopecten irradians concentricus (Say) and the oyster, Crassostrea virginica (Gmelin). Journal of Experimental Marine Biology and Ecology 45:273–295. Pascoe, P.L., Parry, H.E., and Hawkins, A.J.S. 2009. Observations on the measurement and interpretation of clearance rate variations in suspensionfeeding bivalve shellfish. Aquatic Biology 6:181–190. Passow, U. 2002. Transparent Exopolymer Particles, TEP, in aquatic environments. Progress in Oceanography 55:287–333. Passow, U., and Alldredge, A.L. 1994. Distribution, size and bacterial colonization of transparent

exopolymer particles (TEP) in the ocean. Marine Ecology Progress Series 113:185–198. Passow, U., and Alldredge, A.L. 1995. Aggregation of a diatom bloom in a mesocosm: the role of transparent exopolymer particles (TEP). DeepSea Research. Part II, Topical Studies in Oceanography 42:99–109. Passow, U., Shipe, R.F., Murray, A., Pak, D.K., and Brzezinski, M.A. 2001. Origin of transparent exopolymer particles (TEP) and their role in the sedimentation of particulate matter. Continental Shelf Research 21:327–346. Pérez Camacho, A., Labarta, U., and Navarro, E. 2000. Energy balance of mussels Mytilus galloprovincialis: the effect of length and age. Marine Ecology Progress Series 199:149–158. Pernet, F., Tremblay, R., Redjah, I., Sévigny, J.-M., and Gionet, C. 2008. Physiological and biochemical traits correlate with differences in growth rate and temperature adaptation among groups of the eastern oyster Crassostrea virginica. Journal of Experimental Biology 211: 969–977. Petersen, J.K., Bougrier, S., Small, A.C., Garen, P., Robert, S., Larsen, J.E.N., and Brummelhuis, E. 2004. Intercalibration of mussel Mytilus edulis clearance rate measurements. Marine Ecology Progress Series 267:197–194. Petersen, J.K., Nielsen, T.G., van Duren, L., and Maar, M. 2008. Depletion of plankton in a raft culture of Mytilus galloprovincialis in Ría de Vigo, NW Spain. I. Phytoplankton. Aquatic Biology 4:113–125. Pilditch, C.A., and Grant, J. 1999. Effect of variations in flow velocity and phytoplankton concentration on sea scallop (Placopecten magellanicus) grazing rates. Journal of Experimental Marine Biology and Ecology 240:111–136. Podolsky, R.D. 1994. Temperature and water viscosity: physiological versus mechanical effects on suspension feeding. Science 265:100– 103. Pouvreau, S., Jonquières, G., and Buestel, D. 1999. Filtration by the pearl oyster, Pinctada margaritifera, under conditions of low seston load and small particle size in a tropical lagoon habitat. Aquaculture 176:295–314. Pouvreau, S., Bodoy, A., and Buestel, D. 2000. In situ suspension feeding behaviour of the pearl oyster, Pinctada margaritifera: combined effects

Bivalve filter feeding 121

of body size and weather-related seston composition. Aquaculture 181:91–113. Prins, T.C., and Smaal, A.C. 1989. Carbon and nitrogen budgets of the mussel Mytilus edulis L. and the cockle Cerastoderma edule (L.) in relation to food quality. Scientia Marina 53:477– 482. Prins, T.C., Smaal, A.C., and Power, A.J. 1991. Selective ingestion of phytoplankton by the bivalves Mytilus edulis L. and Cerastoderma edule (L.). Hydrobiological Bulletin 25:93– 100. Prins, T.C., Dankers, N., and Smaal, A.C. 1994. Seasonal variation in the filtration rates of a semi-natural mussel bed in relation to seston composition. Journal of Experimental Marine Biology and Ecology 176:69–86. Prins, T.C., Escaravage, V., Smaal, A.C., and Peeters, J.C.H. 1995. Nutrient cycling and phytoplankton dynamics in relation to mussel grazing in a mesocosm experiment. Ophelia 41:289–315. Prins, T.C., Smaal, A., and Dame, R. 1997. A review of the feedbacks between grazing and ecosystem processes. Aquatic Ecology 31: 349–359. Prins, T.C., Smaal, A.C., Pouwer, A.J., and Dankers, N. 1996. Filtration and resuspension of particulate matter and phytoplankton on an intertidal mussel bed in the Oosterschelde estuary (SW Netherlands). Marine Ecology Progress Series 142:121–134. Ramaiah, N., Yoshikawa, T., and Furuya, K. 2001. Temporal variations in transparent exopolymer particles (TEP) associated with a diatom spring bloom in a subarctic ria in Japan. Marine Ecology Progress Series 212:79–88. Ren, J.S., Ross, A.H., and Schiel, D.R. 2000. Functional descriptions of feeding and energetics of the Pacific oyster Crassostrea gigas in New Zealand. Marine Ecology Progress Series 208: 119–130. Resgalla, C., Brasil, E.S., Laitano, K.S., and Reis Filho, R.W. 2007. Physioecology of the mussel Perna perna (Mytilidae) in Southern Brasil. Aquaculture 207:464–474. Riisgård, H.U. 1977. On measurements of the filtration rates of suspension feeding bivalves in a flow system. Ophelia 16:167–173. Riisgård, H.U. 1988. Efficiency of particle retention and filtration rate in 6 species of Northeast

American bivalves. Marine Ecology Progress Series 45:217–223. Riisgård, H.U. 1991. Filtration rate and growth in the blue mussel, Mytilus edulis Linnaeus, 1758: dependence on algal concentration. Journal of Shellfish Research 10:29–35. Riisgård, H.U. 2001a. On measurement of filtration rates in bivalves—the stony road to reliable data: review and interpretation. Marine Ecology Progress Series 211:275–291. Riisgård, H.U. 2001b. Physiological regulation versus autonomous filtration in filter-feeding bivalves: starting points for progress. Ophelia 54:193–209. Riisgård, H.U. 2004. Intercalibration of methods for measurement of bivalve filtration rates—a turning point. Marine Ecology Progress Series 276:307–308. Riisgård, H.U., and Møhlenberg, F. 1979. An improved automatic recording apparatus for determining the filtration rate of Mytilus edulis as a function of size and algal concentration. Marine Biology 52:61–67. Riisgård, H.U., and Randløv, A. 1981. Energy budgets, growth and filtration rates in Mytilus edulis at different algal concentration. Marine Biology 61:227–234. Riisgård, H.U., Kittner, C., and Seerup, D.F. 2003. Regulation of the opening state and filtration rate in filter-feeding bivlaves (Cardium edule, Mytilus edulis, Mya arenaria) in response to low algal concentration. Journal of Experimental Marine Biology and Ecology 284:105–127. Rodhouse, P.G. 1978. Energy transformations by the oyster Ostrea edulis L. in a temperate estuary. Journal of Experimental Marine Biology and Ecology 34:1–22. Saurel, C., Gascoigne, J.C., Palmer, M.R., and Kaiser, M.J. 2007. In situ mussel feeding behaviour in relation to multiple environmental factors: regulation through food concentration and tidal conditions. Limnology and Oceanography 52(5):1919–1929. Seed, R. 1992. Systematics, evolution and distribution of mussels belonging to the genus Mytilus: an overview. American Malacological Bulletin 9:123–137. Sicard, M.T., Maeda-Martinez, A.N., Ormart, P., Reynoso-Granados, T., and Carvalho, L. 1999.

122

Shellfish Aquaculture and the Environment

Optimum temperature for growth in the Catarina scallop (Argopecten ventricosus-circularis, Sowerby II, 1842). Journal of Shellfish Research 18:385–199. Smaal, A.C., and Twisk, F. 1997. Filtration and absorption of Phaeocystis cf. globosa by the mussel Mytilus edulis L. Journal of Experimental Marine Biology and Ecology 209:33–46. Smaal, A.C., and Vonck, A.P.M.A. 1997. Seasonal variation in C, N and P budgets and tissue composition of the mussel Mytilus edulis. Marine Ecology Progress Series 153:167–179. Smaal, A.C., and Zurburg, W. 1997. The uptake and release of suspended and dissolved material by oysters and mussels in Marennes-Oléron Bay. Aquatic Living Resources 10:23–30. Smaal, A.C., Vonck, A.P.M.A., and Bakker, M. 1997. Seasonal variation in physiological energetics of Mytilus edulis and Cerastoderma edule of different size classes. Journal of the Marine Biological Association of the United Kingdom 77:817–838. Smaal, A.C., Verhagen, J.H.G., Coosen, J., and Haas, H.A. 1986. Interaction between seston quantity and quality and benthic suspension feeders in the Oosterschelde, The Netherlands. Ophelia 26:385–399. Soletchnik, P., Goulletquer, P., Héral, M., Razet, D., and Geairon, P. 1996. Evaluation di bilan énergétique de l’huître creuse, Crassostrea gigas, en baie de Marennes-Oléron (France). Aquatic Living Resources 9:65–73. Souchu, P., Vaquer, A., Collos, Y., Landrein, S., Deslous-Paoli, J.-M., and Bibent, B. 2001. Influence of shellfish farming activities on the biogeochemical composition of the water column in Thau Lagoon. Marine Ecology Progress Series 218:141–152. Stenton-Dozey, J.M.E., and Brown, A.C. 1994. Short-term changes in the energy balance of Venerupis corrugatus (Bivalvia) in relation to tidal availability of natural suspended particles. Marine Ecology Progress Series 103:57–64. Strohmeier, T., Strand, Ø., and Cranford, P. 2009. Clearance rates of the great scallop (Pecten maximus) and blue mussel (Mytilus edulis) at low seston concentrations. Marine Biology 156:1781–1795. Stuart, V. 1982. Absorbed ration, respiratory cost and resultant scope for growth in the mussel

Aulacomya ater (Molina) fed on a diet of kelp detritus of different ages. Marine Biology Letters 3:289–306. Thompson, R.J. 1984. The reproductive cycle and physiological ecology of the mussel Mytilus edulis in a subarctic, non-estuarine environment. Marine Biology 79:277–288. Thompson, R.J., and Newell, R.I.E. 1985. Physiological responses to temperature in two latitudinally separated populations of the mussel, Mytilus edulis. In: Gibbs, P.E. (ed.), Proceedings of the 19th European Marine Biology Symposium. Cambridge University Press, Cambridge, pp. 481–495. Trottet, A., Roy, S., Tamigneaux, E., Lovejoy, C., and Tremblay, R. 2008. Impact of suspended mussels (Mytilus edulis L.) on plankton communities in a Magdalen Islands lagoon (Québec, Canada): a mesocosm approach. Journal of Experimental Marine Biology and Ecology 365:103–115. Urrutia, M.B., Iglesias, J.I.P., Navarro, E., and Prou, J. 1996. Feeding and absorption in Cerastoderma edule under environmental conditions in the Bay of Marennes-Oleron (Western France). Journal of the Marine Biological Association of the United Kingdom 76:431–450. Urrutia, M.B., Navarro, E., Iglesias, J.I.P., and Iglesias, J.I.P. 2001. Preingestive selection processes in the cockle Cerastoderma edule: mucas production related to rejection of pseudofaeces. Marine Ecology Progress Series 209:177– 187. Vahl, O. 1980. Seasonal variations in seston and the growth rate of the Iceland Scallop, Chlamys islandica (O.F. Müller) from Balsfjord, 70°N. Journal of Experimental Marine Biology and Ecology 48:195–204. Van Erkon Schurink, C., and Griffiths, C.L. 1992. Physiological energetics of four South African mussel species in relation to body size, ration and temperature. Comparative Biochemistry and Physiology 101A:779–789. Vaquer, A., Troussellier, M., Courtues, C., and Bibent, B. 1996. tandind stock and dynamics of picophytoplankton in the Thau Lagoon (northwest Mediterranean coast). Limnology and Oceanography 41:1821–1828. Velasco, L.A. 2007. Energetic physiology of the Caribbean scallops Argopecten nucleus and

Bivalve filter feeding 123

Nodipecten nodosus fed with different microalgal diets. Aquaculture 270:299–311. Velasco, L.A., and Navarro, J.M. 2002. Feeding physiology of infaunal (Mulinia edulis) and epifaunal (Mytilus chilensis) bivalves under a wide range of concentrations and qualities of seston. Marine Ecology Progress Series 240:143– 155. Velasco, L.A., and Navarro, J.M. 2003. Feeding physiology of two bivalves under laboratory and field conditions in response to variable food concentrations. Marine Ecology Progress Series 291:115–124. Vismann, B. 1990. Field measurements of filtration and respiration rates in Mytilus edulis L. An assessment of methods. Sarsia 75:213–216. Ward, J.E., and Shumway, S.E. 2004. Separating the grain from the chaff: particle selection in suspension- and deposit-feeding bivalves. Journal of Experimental Marine Biology and Ecology 300:83–130. Ward, J.E., Cassell, H.K., and MacDonald, B.A. 1992. Chemoreception in the sea scallop Placopecten magellanicus (Gmelin). I. Stimulatory effects of phytoplankton metabolites on clearance and ingestion rates. Journal of Experimental Marine Biology and Ecology 163:235–250. Ward, J.E., Sanford, L.P., Newell, R.I.E., and MacDonald, B.A. 1998. A new explanation of particle capture in suspension-feeding bivalve molluscs. Limnology and Oceanography 43:741–752. Wetz, M.S., Robbins, M.C., and Paerl, H.W. 2009. Transparent exopolymer particles (TEP) in a river-dominated estuary: spatial-temporal distributions and an assessment of controls upon TEP formation. Estuaries and Coasts 32:447– 455. Widdows, J. 1973. The effects of temperature on the metabolism and activity of Mytilus edulis L. Netherlands Journal of Sea Research 7:387–398. Widdows, J. 1976. Physiological adaptation of Mytilus edulis to cyclic temperatures. Journal of Comparative Physiology 105(2):115–128. Widdows, J. 1978. Combined effects of body size, food concentration and season on the physiology of Mytilus edulis. Journal of the Marine Biological Association of the United Kingdom 58:109–124.

Widdows, J. 2001. Bivalve clearance rates: inaccurate measurements or inaccurate reviews and misrepresentation? Marine Ecology Progress Series 221:303–305. Widdows, J., and Bayne, B.L. 1971. Temperature acclimation of Mytilus edulis with reference to its energy budget. Journal of the Marine Biological Association of the United Kingdom 51:827–843. Widdows, J., and Johnson, D. 1988. Physiological energetics of Mytilus edulis: scope for growth. Marine Ecology Progress Series 46:113–121. Widdows, J., and Navarro, J.M. 2007. Influence of current speed on clearance rate, algal cell depletion in the water column and resuspension of biodeposits of cockles (Cerastoderma edule). Journal of Experimental Marine Biology and Ecology 343:44–51. Widdows, J., and Shick, J.M. 1985. Physiological responses of Mytilus edulis and Cardium edule to aerial exposure. Marine Biology 85:217–232. Widdows, J., Fieth, P., and Worrall, C.M. 1979. Relationships between seston, available food and feeding activity in the common mussel Mytilus edulis. Marine Biology 50:195–207. Widdows, J., Donkin, P., Salkeld, P.N., Cleary, J.J., Lowe, D.M., Evans, S.V., and Thomson, P.E. 1984. Relative importance of environmental factors in determining physiological differences between two populations of mussels (Mytilus edulis). Marine Ecology Progress Series 17:33–47. Widdows, J., Lucas, J.S., Brinsley, M.D., Salkeld, P.N., and Staff, F.J. 2002. Investigation of the effects of current velocity on mussel feeding and mussel bed stability using an annular flume. Helgoland Marine Research 56:3–12. Wildish, D.J., and Kristmanson, D.D. 1997. Benthic Suspension Feeders and Flow. Cambridge University Press, Cambridge. Wildish, D.J., and Miyares, M.P. 1990. Filtration rate of blue mussels as a function of flow velocity: preliminary experiments. Journal of Experimental Marine Biology and Ecology 142:213–219. Wildish, D.J., and Saulnier, A.M. 1993. Hydrodynamic control of filtration in Placopecten magellanicus. Journal of Experimental Marine Biology and Ecology 174:65–82. Wildish, D.J., Kristmanson, D.D., Hoar, R.L., DeCoste, A.M., McCormick, S.D., and White,

124

Shellfish Aquaculture and the Environment

A.W. 1987. Giant scallop feeding and growth responses to flow. Journal of Experimental Marine Biology and Ecology 113:207–220. Wildish, D.J., Kristmanson, D.D., and Saulnier, A.M. 1992. Interactive effect of velocity and seston concentration on giant scallop feeding inhibition. J. Exp. Mar. Biol. Ecol. 155: 161–168. Wilson, J.H. 1983. Retention efficiency and pumping rate of Ostrea edulis in suspensions of Isochrysis galbana. Marine Ecology Progress Series 12:51–58. Winter, J.E. 1978. A review of knowledge of suspension-feeding in lamellibranchiate bivalves, with special reference to artificial aquaculture systems. Aquaculture 13:1–33. Winter, J.E., Acevedo, M.A., and Navarro, J.M. 1984. Quempillén estuary, an experimental oyster cultivation station in southern Chile. Energy balance in Ostrea chilensis. Marine Ecology Progress Series 20:151–164. Wong, W.H., and Cheung, S.G. 1999. Feeding behaviour of the green mussel, Perna viridis (L.): responses to variation in seston quantity and quality. Journal of Experimental Marine Biology and Ecology 236:191–207. Wong, W.H., and Cheung, S.G. 2001a. Feeding rhythms of the green-lipped mussel, Perna viridis (Linnaeus, 1758) (Bivalvia: Mytilidae) during spring and neap tidal cycles. Journal of Experimental Marine Biology and Ecology 257:13–36.

Wong, W.H., and Cheung, S.G. 2001b. Feeding rates and scope for growth of green mussel, Perna viridis (L.) and their relationship with food availability in Kat O, Hong Kong. Aquaculture 193:123–137. Wurl, O., and Holmes, M. 2008. The gelatinous nature of the sea-surface microlayer. Marine Chemistry 110:89–97. Yukihira, H., Klumpp, D.W., and Lucas, J.S. 1998. Effects of body size on suspension feeding and energy budgets of the pearl oysters Pinctada margaritifera and P. maxima. Marine Ecology Progress Series 170:119–130. Zeldis, J., Robinson, K., Ross, A., and Hayden, B. 2004. First observations of predation by New Zealand Greenshell mussels (Perna canaliculus) on zooplankton. Journal of Experimental Marine Biology and Ecology 311:287–299. Zhou, Y., Yang, H., Zhang, T., Liu, S., Zhang, S., Liu, Q., Xiang, J., and Zhang, F. 2006. Influence of filtering and biodeposition by the cultured scallop Chlamys farreri on benthic-pelagic coupling in a eutrophic bay in China. Marine Ecology Progress Series 317:127–141. Zurburg, W., Smaal, A., Héral, M., and Dankers, N. 1994. Seston dynamics and bivalve feeding in the Bay of Marennes-Oléron (France). Netherlands Journal of Aquatic Ecology 28(3–4):459–466.

Chapter 5

Trophic interactions between phytoplankton and bivalve aquaculture Gary H. Wikfors

The interdependence of bivalves and phytoplankton The term “trophic interactions” used here is purposely broader than the concept of one thing feeding on another. In the case of the bivalve molluscs that are suspension-feeders and the microalgae—chiefly phytoplankton— that constitute a large fraction of the living component of the suspended seston upon which molluscs feed, the most obvious interaction is bivalves eating algae. Increasingly, however, the reverse trophic interaction is being recognized; dissolved inorganic and organic waste compounds released by metabolically active bivalves can supply microalgae with nutrient and energy requirements for their growth (Officer et al. 1982; Boucher and

Boucher-Rodoni 1988; Smaal and Prins 1993; Prins et al. 1998; Newell 2004). This two-way interaction can be viewed as a type of community symbiosis developed over long evolutionary timescales (Fig. 5.1). How does aquaculture of molluscan shellfish fit into this long-established symbiosis between molluscs and phytoplankton? One could assume that, as in most monoculture cultivation scenarios, farming of suspensionfeeding molluscs should intensify the interactions in time and space, possibly upsetting critical environmental equilibria. I argue, to the contrary, that natural populations of most molluscan species that are farmed intensively tend to be highly gregarious, naturally forming dense assemblages akin to those created through aquaculture practices. Further,

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 125

126

Shellfish Aquaculture and the Environment

Harvested (pOrg. N, pOrg. C)

pOrg. N

Water column

CO2

dOrg. N

Oyster nursery

Phytoplankton (pOrg. N,pOrg. C)

NH4+

(pOrg. N, pOrg. C)

Tidal flow

NO2– –

NO3

Fecal matter deposition

Biotic uptake

O2 Advective/

diffusive exchange pOrg. N, pOrg. C

N2

NO3

–

NH4+

dOrg. N

pOrg. N

Toxic sediments

Burial Figure 5.1 “Box model” of a suspension-culture, oyster nursery, with arrows depicting exchanges of carbon, nitrogen, and phosphorus between the oyster nursery and environmental compartments. Of particular note are the arrows indicating return of respiratory carbon (CO2) and excreted nitrogen (NH4+) to the phytoplankton community, thereby recycling resources not assimilated immediately by the oysters. (Figure original, S.L. Meseck and G.H. Wikfors.)

reef-building characteristics of some species, such as oysters, have served under natural conditions to transfer benthic organisms into the pelagic realm where they are within the primary productivity maximum near the water surface and less vulnerable to stress from siltation and hypoxia (Lenihan 1999). I argue that bivalve aquaculture can restore trophic balance between the bivalves and phytoplankton communities that may have existed before habitat modifications caused by other human activities, such as harvesting (dragging former reefs flat), channel dredging for boat transportation, and bulkheading (Rothschild et al. 1994; Hargis and Haven 1999). Restoring bivalve–phytoplankton trophic interactions through shellfish aquaculture has the potential to mitigate ecosystem imbalances attributed to nutrient overenrichment and to help reverse the cycle of ecosystem degradation in coastal waters resulting from

both eutrophication (see Chapter 7 in this book) and food web disruption (Ulanowicz and Tuttle 1992; Cerco and Noel 2007). The main challenge in the harmonization of bivalve aquaculture and coastal ecosystembased management of water quality involves scaling. In development of a natural system, the highly adaptive reproductive strategy of most bivalves, that is, high fecundity but low investment in offspring, allows bivalve populations to respond on decadal timescales to changes in the quantity and quality of trophic resources available from phytoplankton primary productivity, as well as other ecological changes (Dekshenieks et al. 2000). In siting and scaling of shellfish aquaculture, one does not often have the benefit of knowing past carrying capacity. Even if such knowledge was available, it is likely that bottom-up forcing functions of nutrient inputs, as well as competition for primary production from

Phytoplankton and bivalve aquaculture

zooplankton, have changed as a result of other environmental changes (Peterson and Lipcius 2003). Thus, quantitative knowledge of bivalve–phytoplankton trophic interactions in coastal waters will inform bivalve aquaculture development to effectively serve the needs of both seafood production and ecosystem restoration.

Bivalve population density: farmed bivalves are naturally gregarious The characteristics of bivalves that make physical proximity of individuals necessary are external fertilization of gametes coupled with effective nonmotility. If spawning is to lead to offspring, individuals contributing gametes must be close enough to each other for fertilization to occur before gametes are lost to physical dilution or consumption by grazers (Andre and Lindegarth 1995). Populations of bivalves often, therefore, are referred to as “reefs” or “beds,” depending on vertical structure of the aggregation (Korringa 1946; Wells 1957; Hargis and Haven 1999; Lawrie and McQuaid 2001). Impaired spawning success has been attributed to shellfish overharvest or depletion of natural populations below “critical densities” (Kraeuter et al. 2005). Particle clearance, excretion, and biodeposition intensities of dense, farmed populations of bivalves, therefore, cannot be considered as “unnatural,” as bivalve species farmed are not typically distributed widely at low density; they occur by biological necessity as concentrated aggregations. Oysters have long been considered to be “ecosystem engineers,” more for their reefbuilding activities modifying benthic habitat than for their trophic interactions (Lenihan and Peterson 1998). Only recently are the particle clearance and nutrient recycling activities of oysters being considered in oyster restoration efforts (Coen et al. 2007). Similarly, mussels attach to any hard substrate in the

127

environment and then to each other, forming three-dimensional aggregations that change shape as byssal threads are formed and broken by waves and tidal currents (Dolmer 2000; Lawrie and McQuaid 2001). Clams are the main infaunal bivalve group farmed. Subsurface aggregations of clams have been hypothesized to result from water current influences upon larval settlement and provision of food to the benthic boundary layer (Wells 1957), as well as from substrate refugia from predation, for example, under shell hash or between rocks. With scallops, it is more difficult to conform to the “shellfish are gregarious” generalization. Most scallop species farmed are at least somewhat motile, able to adjust their location vertically and spatially according to changes in conditions. In intensive culture, some scallop species appear to attempt to adjust their spatial density to minimize intraspecific competition for food (Rhodes and Widman 1984). Accordingly, scallop aquaculture tends to occur at somewhat lower intensity, employing ear-hanging or lantern-net methods to achieve spatial distribution in three dimensions. In general, though, the argument that bivalve aquaculture is analogous to “monoculture agriculture” ignores the natural distribution patterns of bivalve species farmed.

Bivalves as consumers and cultivators of phytoplankton When bivalves feed, they remove suspended particulate matter from the water as the first step in the process by which they acquire the energy and materials they need to live and to grow. The elemental composition of suspended organic material in the sea often is estimated by the “Redfield ratio” or the “extended Redfield ratio” (Twining et al. 2004). Two important points that limit the accuracy of these ratios to represent seston or phytoplankton composition are as follows: (1) these ratios are based on, and averaged for, samples of

128

Shellfish Aquaculture and the Environment

suspended solids collected from the sea, not based on an internal, chemical stoichiometry imposed by the living organisms in the samples (Falkowski 2000); and (2) average values reported mainly are from open-ocean samples, not from coastal, estuarine, or brackish waters where most molluscan aquaculture is practiced (Arrigo 2005). A large amount of variance, often on diurnal timescales, in the relative carbon content of seston results from assimilation of this element into microalgal sugars by photosynthesis during daylight and subsequent catabolism of these sugars and release of carbon dioxide in darkness (Paerl and Mackenzie 1977). As the dietary energy available to a feeding bivalve is modified by the carbon status of the phytoplankton, feeding over the course of the day will present bivalves with a range of energy contents within ingested food. Similarly, the protein, hence nitrogen, content of phytoplankton is dependent on the availability of this nutrient and sufficient energy for anabolic protein synthesis. Under conditions of energy limitation, phytoplankton will assimilate more nitrogen than needed for cell division (Geider and La Roche 2002)— a process referred to as “luxury consumption.” Thus, the two most important nutritional inputs for bivalves, energy and protein, can be expected to vary considerably over diurnal and seasonal cycles. Whether or not this variation averages over annual timescales to the Redfield ratio is not certain and probably very different from place to place. Less attention is paid to the phosphorus content in seston consumed by bivalves, as this element generally is thought to be in excess supply in coastal waters—an assumption that may not be valid in coastal waters used for shellfish cultivation (Howarth and Marino 2006). Many coastal waters are considered to be negatively impacted by eutrophication (see Chapter 7 in this book)—an overabundance of nutrients (chiefly biologically available nitrogen) leading to “excess” phytoplankton

primary productivity (Cloern 2001). Negative ecosystem consequences of high planktonic primary productivity are attributed to shading of submerged aquatic vegetation and especially to benthic hypoxia and anoxia resulting from bacterial respiration of unassimilated phytoplankton biomass. Although water quality criteria may value “clear” water devoid of phytoplankton, filter-feeding bivalves will starve in the absence of suspended food particles. Thus, bivalves exploit as a resource the phytoplankton production viewed by water quality managers as a nuisance (Lindahl et al. 2005). Indeed, phytoplankton standing stock is defined as the difference between primary production and consumption; therefore, coastal waters discolored by phytoplankton can be considered a consequence of both increased fertilization with nutrients (bottomup) and reduced grazing pressure (top-down) if historic bivalve populations have been depleted. In this respect, implementing bivalve aquaculture in eutrophic coastal waters can be considered as “restoration” of the particleclearing function to the impacted ecosystem (Nelson et al. 2004). As with any ecological interaction, too much of a good thing is still too much. It is generally agreed that the so-called “pseudofeces threshold” for bivalves is in the range of 2–10 mg L−1 suspended particles (Bayne and Newell 1983). Above this threshold, a portion of the particles captured is rejected before ingestion because the respiratory activities of the gills take precedence over food capture. In the opposite direction of this trophic interaction, “too much” bivalve filtration has been considered as a possible risk to pelagic food webs that support fisheries. Thus, the concept of “carrying capacity” for bivalves must be considered at several levels (see Chapter 6 in this book). First, the physical carrying capacity within the spatial domain must be considered—how many shellfish will fit in a defined area (Smaal et al. 2001)? Using

Phytoplankton and bivalve aquaculture

suspension culture, physical carrying capacity often can be increased to a point where other considerations become limiting (Fréchette and Bacher 1998). Next, production carrying capacity is defined as the density above which production is limited by lack of resources such as food or oxygen. Production carrying capacity has been exceeded in some shellfish cultivation situations, such as raft culture of mussels in Spain (Blanton et al. 1987). It can be argued, though, that natural recruitment sometimes exceeds production carrying capacity, leading to mortality of individuals in populations (Peterson and Black 1988). Ecological carrying capacity next considers the overall impact of the shellfish cultivation activity upon the other living components of the ecosystem (Dame and Prins 1998; Grant et al. 2007). It seems unrealistic to define this limit as the density at which any ecosystem effects are apparent, although it often is. Instead, I would argue that ecosystem carrying capacity should be defined as the point at which the positive consequences are balanced by the negatives. Agreeing on a definition of ecosystem carrying capacity overlaps, thus, with the highest-order determination—social carrying capacity. This can be defined as the level of activity that will be tolerated by human societies interested in the ecosystems. One cannot posit a scientific definition for this beyond what has been stated. Moving from gross measures of food quantity to food quality, there is a general consensus that high protein contents in phytoplankton cells, and consequently in seston of coastal waters, generally are able to provide nutritional needs of bivalves for dietary protein (Brown et al. 1997). In contrast, specific lipids, especially long-chain, polyunsaturated fatty acids (PUFAs) and certain sterols, may be limiting in phytoplankton and seston food sources of bivalves (Trider and Castell 1980). These lipids are required as structural membrane components in bivalve cells, rather than for

129

their energy content (Delaporte et al. 2005). Dietary PUFAs and sterols are dependent on both the energy status and the taxonomic composition of the phytoplankton community (Sargent et al. 1985), with some microalgal classes being devoid of these compounds (e.g., chlorophytes have no PUFAs longer than 18 carbons, but 20- and 22-carbon PUFAs are considered to be essential). Despite this knowledge concerning qualitative nutritional needs of bivalves derived from laboratory feeding studies (Knauer and Southgate 1999), there is no evidence yet that biodeposits—the rejected portion of seston filtered but not assimilated— is selectively stripped of PUFAs or biologically useful sterols. Indeed, evidence from analysis of shellfish tissues indicates that some lipids not used in construction of new cell membranes are accumulated in the tissues, in a way like parts that do not fit in the construction (Goad 1981). Thus, although much evidence exists for selective retention and ingestion of particles based on both physical and chemical properties (Ward and Shumway 2004), it is not clear that selection is based on nutritional criteria (see Chapter 4 in this book). This area of research needs further work to clarify selectivity in bivalve feeding and how this may impact planktonic communities and nutrient cycling (Ward et al. 1994). One clear change in nutrient chemistry that can occur as a consequence of bivalve feeding and elimination processes involves the nutrient silica (Si). Silica is a macronutrient for one microalgal class, the Bacillariophyceae, or the diatoms. When a bivalve consumes diatom biomass, portions of the nitrogen and phosphorus components are assimilated into bivalve tissues, and remaining portions are returned to the environment in relatively labile forms. Complex, organic molecules in biodeposits can be recycled rapidly by bacterial decomposition, and nitrogenous wastes in the form of ammonia and urea are available immediately for phytoplankton reuse. Silica in

130

Shellfish Aquaculture and the Environment

diatom frustules, however, can be returned to the environment in a form, the mineral opal, that is only slowly remineralized under conditions found within biodeposits (Nelson et al. 1995). Thus, molluscan shellfish can be considered efficient recyclers of nitrogen and phosphorus in the environment (Smaal and Zurburg 1997; Souchu et al. 2001; Newell et al. 2005), but they may represent a sink for silica. Thus, intense feeding by bivalves can be considered an activity that encourages the growth of nondiatom microalgae on recycled nitrogen and phosphorus. This is the process characterized as the “cultivation” of a flagellate food source by bivalve populations (Prins et al. 1998). Nondiatom taxa do include flagellates in several taxonomic groups that are useful nutritionally, for example, prasinophytes, prymnesiophytes, and cryptophytes. Harmful dinoflagellate and cyanobacterial taxa, however, may also benefit from bivalve selective recycling of nitrogen and phosphorus. The possible contribution of intensive bivalve aquaculture to increases in harmful algal blooms can be theorized, but there is no evidence, even circumstantial, that this has occurred. Indeed, bivalves tend to filter and partially degrade many harmful algal species (Cerrato et al. 2004; Hégaret et al. 2007, 2008) and seem to serve mainly as a vector transporting possibly viable cells from pelagic to benthic compartments in the environment.

Summary and prospects Trophic interactions between shellfish aquaculture and phytoplankton fundamentally involve feeding and nutrient recycling activities of bivalve molluscs, which tend to sustain primary production locally, but favor nondiatom taxa. This sustained primary productivity during summer may benefit other planktivorous animals, for example, micro- and meiozooplankton that can contribute to the finfish food chain. In shallow, coastal ecosystems

where historical bivalve populations have been reduced by fishing and other human activities, restoration of bivalve feeding and nutrient recycling activities could serve to restore or “rebalance” trophic structure. Fundamentally, though, assimilation of nitrogen and phosphorus into shellfish tissues provides an opportunity to remove these nutrients from the environment during harvest. Only recently has this environmental benefit been recognized (Officer et al. 1982; Rice 1999; Lindahl et al. 2005). The implementation of nitrogen trading mechanisms to manage coastal eutrophication is providing an opportunity for shellfish farmers to realize a portion of the ecosystem service value of their harvests (Ferreira et al. 2007) (see Chapters 1 and 8 of in this book). Finally, to project into the future, the filterfeeding activities of cultivated bivalve molluscs can be viewed as a natural solution to a current problem in microalgal technology. Large investments are being made into development of technologies to mass-culture microalgae for the purpose of producing biofuels (Chisti 2007). One of the main technical and economic challenges in this technology is removing the microalgal biomass from the water in which it is suspended. Bivalve molluscs do this very efficiently, and eutrophic estuaries are already growing more phytoplankton than the largest planned bioreactor of pond-based microalgal farms. Even if converting bivalve biomass into biodiesel is not practical, harvest of cultivated bivalves from eutrophic estuaries, planted for the purpose of eutrophication mitigation, is also a way to extract marine protein and lipids from microalgae in a form with other possible uses. In one ongoing project in Sweden, mussel meal and oils are being used in poultry feeds as replacements for fish meal (Lindahl et al. 2005). A major controversy stalling development of carnivorous fish aquaculture expansion is the concern that “feeding fish to fish” will further deplete the forage base for natural fish stocks and cause a global imbalance in fisher-

Phytoplankton and bivalve aquaculture

ies trophic structure (Naylor et al. 2000). Rather than “microalgal biomass-to-biofuel” technologies, one can envision development of “coastal eutrophication-to-fish feed” technologies that solve two problems simultaneously using trophic interactions between phytoplankton and aquacultured bivalve molluscs as the functional “clutch.”

Acknowledgments Many thanks to the following colleagues for discussions stimulating the thoughts expressed in this paper: Sandra Shumway, Chris Brown, Mark Tedesco, Robert Rheault, Odd Lindahl, Michael Rubino, Arthur Glowka, Loy Wilkinson, and Frank Trainor.

Literature cited Andre, C., and Lindegarth, M. 1995. Fertilization efficiency and gamete viability of a sessile, freespawning bivalve, Cerastoderma edule. Ophelia 43(3):215–227. Arrigo, K.R. 2005. Marine microorganisms and global nutrient cycles. Nature 437:349–355. Bayne, B.L., and Newell, R.C. 1983. Physiological energetics of marine molluscs. In: Saleuddin, A.S.M., and Wilbur, K.M. (eds.), The Mollusca, Vol. 4. Academic Press, New York, pp. 407–515. Blanton, J.O., Tenore, K.R., Castillejo, F., Atkinson, L.P., Schwing, F.B., and Lavin, A. 1987. The relationship of upwelling to mussel production in the rias on the western coast of Spain. Journal of Marine Research 45(2):497–511. Boucher, G., and Boucher-Rodoni, R. 1988. In situ measurement of respiratory metabolism and nitrogen fluxes at the interface of oyster beds. Marine Ecology Progress Series 44:229–238. Brown, M.R., Jeffrey, S.W., Volkman, J.K., and Dunstan, G.A. 1997. Nutritional properties of microalgae for mariculture. Aquaculture 151:315–331. Cerco, C.F., and Noel, M.R. 2007. Can oyster restoration reverse cultural eutrophication in

131

Chesapeake Bay? Estuaries and Coasts 30(2):331–343. Cerrato, R.M., Caron, D.A., Lonsdale, D.J., Rose, J.M., and Schaffner, R.A. 2004. Effect of the northern quahog Mercenaria mercenaria on the development of blooms of the brown tide alga Aureococcus anophagefferens. Marine Ecology Progress Series 281:93–108. Chisti, Y. 2007. Biodiesel from microalgae. Biotechnology Advances 25:294–306. Cloern, J.E. 2001. Our evolving conceptual model of the coastal eutrophication problem. Marine Ecology Progress Series 210:223–253. Coen, L.D., Brumbaugh, R.D., Bushek, D., Grizzle, R., Luckenbach, M.W., Posey, M.H., Powers, S.P., and Tolley, S.G. 2007. Ecosystem services related to oyster restoration. Marine Ecology Progress Series 341:303–307. Dame, R.F., and Prins, T.C. 1998. Bivalve carrying capacity in coastal ecosystems. Aquatic Ecology 31:409–421. Dekshenieks, M.M., Hofmann, E.E., Klinck, J.M., and Powell, E.N. 2000. Quantifying the effects of environmental change on an oyster population: a modeling study. Estuaries 23(5): 593–610. Delaporte, M., Soudant, P., Moal, J., Kraffe, E., Marty, Y., and Samain, J.-F. 2005. Incorporation and modification of dietary fatty acids in gill polar lipids by two bivalve species Crassostrea gigas and Ruditapes philippinarum. Comparative Biochemistry and Physiology. Part A, Molecular & Integrative Physiology 140(4):460–470. Dolmer, P. 2000. Feeding activity of mussels Mytilus edulis related to near-bed currents and phytoplankton biomass. Journal of Sea Research 44:221–231. Falkowski, P. 2000. Rationalizing nutrient ratios in unicellular algae. Journal of Phycology 36:3–6. Ferreira, J.G., Hawkins, A.J.S., and Bricker, S.B. 2007. Management of productivity, environmental effects and profitability of shellfish aquaculture—the Farm Aquaculture Resource Management (FARM) model. Aquaculture 264(1–4):160–174. Fréchette, M., and Bacher, C. 1998. A modeling study of optimal stocking density of mussel populations kept in experimental tanks. Journal of Experimental Marine Biology and Ecology 219:241–255.

132

Shellfish Aquaculture and the Environment

Geider, R.J., and La Roche, J. 2002. Redfield revisited: variability of C : N : P in marine microalgae and its biochemical basis. European Journal of Phycology 37(1):1–17. Goad, L.J. 1981. Sterol biosynthesis and metabolism in marine invertebrates. Pure and Applied Chemistry 51:837–852. Grant, J., Curran, K.J., Guyondet, T.L., Tita, G., Bacher, C., Koutitonsky, V., and Dowd, M. 2007. A box model of carrying capacity for suspended mussel aquaculture in Lagune de la Grande-Entrée, Iles-de-la-Madeleine, Québec. Ecological Modelling 200(1–2):193–206. Hargis, W.J., Jr., and Haven, D.S. 1999. Oyster reef habitat restoration: a synopsis and synthesis of approaches. In: Luckenbach, M.W., Mann, R., Wesson, J.A., and Gloucester Point, V.A. (eds.), Chesapeake Oyster Reefs, Their Importance, Destruction and Guidelines for Restoring Them. Virginia Institute of Marine Science Press, Gloucester Point, VA, Chapter 23. Hégaret, H., Wikfors, G.H., and Shumway, S.E. 2007. Diverse feeding responses of five species of bivalve mollusc when exposed to three species of harmful algae. Journal of Shellfish Research 26(2):549–559. Hégaret, H., Shumway, S.E., Wikfors, G.H., Pate, S., and Burkholder, J.A.M. 2008. Potential transport of harmful algae via relocation of bivalve molluscs. Marine Ecology Progress Series 361:169–179. Howarth, R.W., and Marino, R. 2006. Nitrogen as the limiting nutrient for eutrophication in coastal marine ecosystems: evolving views over three decades. Limnology and Oceanography 51(1 Pt 2):364–376. Knauer, J., and Southgate, P.C. 1999. A review of the nutritional requirements of bivalves and the development of alternative and artificial diets for bivalve aquaculture. Reviews in Fisheries Science 7(3–4):241–280. Korringa, P. 1946. A revival of natural oyster beds? Nature 158:586–587. Kraeuter, J.N., Buckner, S., and Powell, E.N. 2005. A note on a spawner-recruit relationship for a heavily exploited bivalve: the case of northern quahogs (hard clams), Mercenaria mercenaria in Great South Bay New York. Journal of Shellfish Research 24(4):1043–1052.

Lawrie, S.M., and McQuaid, C.D. 2001. Scales of mussel bed complexity: structure, associated biota and recruitment. Journal of Experimental Marine Biology and Ecology 257:135–161. Lenihan, H.S. 1999. Physical–biological coupling on oyster reefs: how habitat structure influences individual performance. Ecological Monographs 69(3):251–275. Lenihan, H.S., and Peterson, C.H. 1998. How habitat degradation through fishery disturbance enhances impacts of hypoxia on oyster reefs. Ecological Applications 8:128–140. Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L.-O., Olrog, L., Rehnstam-Holm, A.-S., Svensson, J., Svensson, S., and Syversen, U. 2005. Improving marine water quality by mussel farming: a profitable solution for swedish society. Ambio 34(2):131–138. Naylor, R.L., Goldburg, R.J., Primavera, J.H., Kautsky, N., Beveridge, M.C.M., Clay, J., Folke, C., Lubchenco, J., Mooney, H., and Troell, M. 2000. Effect of aquaculture on world fish supplies. Nature 405:1017–1024. Nelson, D.M., Tréguer, P., Brzezinski, M.A., Leynaert, A., and Quéguiner, B. 1995. Production and dissolution of biogenic silica in the ocean: revised global estimates, comparison with regional data and relationship to biogenic sedimentation. Global Biogeochemical Cycles 9(3):359–372. Nelson, K.A., Leonard, L.A., Posey, M.H., Alphin, T.D., and Mallin, M.A. 2004. Using transplanted oyster (Crassostrea virginica) beds to improve water quality in small tidal creeks: a pilot study. Journal of Experimental Marine Biology and Ecology 298:347–368. Newell, R.I.E. 2004. Ecosystem influences of natural and cultivated populations of suspension-feeding bivalve molluscs: a review. Journal of Shellfish Research 23(1):51–61. Newell, R.I.E., Fisher, T.R., Holyoke, R.R., and Cornwell, J.C. 2005. Influence of eastern oysters on nitrogen and phosphorus regeneration in Chesapeake Bay, USA. In: Dame, R.F., and Olenin, S. (eds.), The Comparative Roles of Suspension-Feeders in Ecosystems. Springer, Dordrecht, The Netherlands, pp. 93–120. Officer, C.B., Smayda, T.J., and Mann, R. 1982. Benthic filter feeding: a natural eutrophication

Phytoplankton and bivalve aquaculture

control. Marine Ecology Progress Series 9:203–210. Paerl, H.W., and Mackenzie, L.A. 1977. A comparative study of the diurnal carbon fixation patterns of nanoplankton and net plankton. Limnology and Oceanography 22(4):732–738. Peterson, C.H., and Black, R. 1988. Densitydependent mortality caused by physical stress interacting with biotic history. The American Naturalist 132:257–270. Peterson, C.H., and Lipcius, R.N. 2003. Conceptual progress towards predicting quantitative ecosystem benefits of ecological restorations. Marine Ecology Progress Series 264:297–307. Prins, T.C., Smaal, A.C., and Dame, R.F. 1998. A review of the feedbacks between bivalve grazing and ecosystem processes. Aquatic Ecology 31:349–359. Rhodes, E.W., and Widman, J.C. 1984. Densitydependent growth of the bay scallop Argopecten irradians irradians, in suspension culture. International Council for the Exploration of the Sea. 18:1–8. Rice, M.A. 1999. Control of eutrophication by bivalves: filtration of particulates and removal of nitrogen through harvest of rapidly growing stocks. Journal of Shellfish Research 18(1):275. Rothschild, B., Ault, J., Goulletquer, P., and Heral, M. 1994. Decline of the Chesapeake Bay oyster population: a century of habitat destruction and overfishing. Marine Ecology Progress Series 111:29–39. Sargent, J.R., Eilertsen, H.C., Falk-Petersen, S., and Taasen, J.P. 1985. Carbon assimilation and lipid production in phytoplankton in northern Norwegian fjords. Marine Biology 85(2): 109–116. Smaal, A.C., and Prins, T.C. 1993. The uptake of organic matter and the release of inorganic nutrients by bivalve suspension feeder beds. In: Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Coastal Ecosystem Processes. Springer-Verlag, Berlin, pp. 271–298.

133

Smaal, A.C., and Zurburg, W. 1997. The uptake and release of suspended and dissolved material by oysters and mussels in Marennes-Oléron Bay. Aquatic Living Resources 10:23–30. Smaal, A., van Stralen, M., and Schuiling, E. 2001. The interaction between shellfish culture and ecosystem processes. Canadian Journal of Fisheries and Aquatic Sciences 58(5): 991–1002. Souchu, P., Vaquer, A., Collos, Y., Landrein, S., Deslous-Paoli, J.-M., and Bibent, B. 2001. Influence of shellfish farming activities on the biogeochemical composition of the water column in Thau lagoon. Marine Ecology Progress Series 218:141–152. Trider, D.J., and Castell, J.D. 1980. Effect of dietary lipids on growth, tissue composition and metabolism of the oyster (Crassostrea virginica). Journal of Nutrition 110(7):1303–1309. Twining, B.S., Baines, S.B., and Fisher, N.S. 2004. Element stoichiometries of individual plankton cells collected during the southern ocean iron experiment (SOFeX). Limnology and Oceanography 49(6):2115–2128. Ulanowicz, R., and Tuttle, J. 1992. The trophic consequences of oyster stock rehabilitation in Chesapeake Bay. Estuaries and Coasts 15:298–306. Ward, J.E., and Shumway, S.E. 2004. Separating the grain from the chaff: particle selection in suspension- and deposit-feeding bivalves. Journal of Experimental Marine Biology and Ecology 300(1–2):83–130. Ward, J.E., Newell, R.I.E., Thompson, R.J., and Macdonald, B.A. 1994. In Vivo studies of suspension-feeding processes in the Eastern Oyster, Crassostrea virginica (Gmelin). The Biological Bulletin 186(2):221–240. Wells, H.W. 1957. Abundance of the Hard Clam Mercenaria mercenaria in relation to environmental factors. Ecology 38:123–128.

Chapter 6

The application of dynamic modeling to prediction of production carrying capacity in shellfish farming Jon Grant and Ramón Filgueira

Bivalve aquaculture is well known to be increasing worldwide. Among the “greenest” of seafarming activities, bivalves utilize natural seston as food, requiring no organic subsidy as in fish culture. Natural food particles can, however, be limiting to bivalve culture yield, and in order to maintain a sustainable resource, it is necessary to balance the biomass of farmed animals with the amount of food available and production of waste. Marine systems are complex, with suspended food being a dynamic quantity subject to various diffusion–advection processes due to wind, tides, and hydrography. Field measurements alone make it to difficult to capture these dynamics, particularly with respect to forecasting. Simulation models are therefore an important approach to predicting shellfish growth as a result of environmental

conditions. Simulation models have long been operational in agricultural ecosystems (Cabrera et al. 2006), where this approach is more feasible since inputs, outputs, and transports are all managed, and system boundaries are rigidly defined. As suggested above, the most important concept in production carrying capacity (CC) is seston depletion, in which bivalves limit their own food supply. High biomass of bivalves, either naturally occurring or in culture, can reduce stocks of phytoplankton and thus exert top-down control of primary production. This is a two-way street; in addition to controlling seston, bivalve growth is dependent on seston quality and quantity. Other studies have determined that it is the flux of seston rather than the concentration

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 135

136

Shellfish Aquaculture and the Environment

that is important in this limitation (Grizzle and Lutz 1989). In marine aquaculture, bivalve biomass and grazing pressure are inserted into the natural trophic structure. This is in contrast to agriculture, which is thoroughly managed on private property, and the agroecosystem entirely constructed. Models of shellfish ecosystems are most often trophically based, with a minimum of phytoplankton, nutrients, and seston compartment. Although one might categorize the structure of CC models based on trophic level alone, there is a more fundamental division that defines the components of these models as follows:

• Biogeochemical model: A catch-all name for phytoplankton, nutrients, other grazers (zooplankton), and benthos that interact as the receiving ecosystem where bivalve culture is inserted. Fluxes within this component are to a large extent dependent on the physical submodel since the planktonic components are transported in suspension in the model domain. • Bivalve ecophysiology model: The details of the bivalve energy budget determine ingestion, absorption, and egestion, and thus the growth rate of the animal, its impact on seston, and biodeposition to sediments. There have been several approaches to this component including traditional scope for growth (Grant and Bacher 1998) and dynamic energy budgets (DEBs; Rosland et al. 2009). • Physical oceanographic model: Although this may be incorporated into the larger CC model, it often stands alone, in some cases yielding averaged conditions to the biogeochemistry. The level of complexity of this model is dependent on a variety of factors as discussed below. One advantage of using bivalve growth as a predictive variable is that it is readily ground truthed. When field studies are conducted as

part of a CC research program, it is possible to establish growth trials from which animals can be sampled in time series. Although this is not uncommon, it is rarely carried out in different spatial locations, as we have done in previous work (Waite et al. 2005). Although we partition model efforts above, there is the added dimension of spatial scale at which the model is targeted. Feeding and removal of seston occur at the level of an individual bivalve, perhaps on a centimeter scale. This is conceptualized in self-thinning models (Frechette and Bacher 1998). Bivalves such as mussels in suspended culture, however, are also aggregated in socks, and socks into longlines or rafts (see below). The spatial scale at which seston depletion is defined is a major distinction between model examples, ranging from single boxes representing a whole bay to hundreds of boxes representing multiple locations on a geolocated map. Increased spatial resolution requires greater model complexity and computational resources. Moreover, physical circulation models must also be able to specify exchange coefficients on a spatial scale similar to the ecosystem model. An explicit definition of CC is essential in seeking to focus the goals of directed models. This has been explored as a contrast between economic, production, ecological, and social CC (Gibbs 2009). The production definition relates to cultured biomass and/or growth rate that can be sustained by available food. It has been particularly difficult to define food limitation, as we discuss below. In this case, models are oriented toward the industry as well as economic CC (i.e., considering both yield and price) since bivalve growth rate is involved. Ecological CC seeks to minimize impacts, most often to benthic habitats via organic loading (see Chapter 9 in this book). This type of model is used by regulators to manage these aspects of aquaculture. In both cases, the criterion or threshold for whether CC has been reached is subjective. For example, increasing stocked biomass may result in

Dynamic modeling and production carrying capacity

137

1. New shellfish farming Release assessment 2. Filtration within the farm

3. Food reduction within the farm

4. Food limitation within the farm

Exposure assessment

6. Food limitation in distant farms Consequence assessment

5. Effect on growth within the farm

7. Effect on growth in distant farm

Carrying capacity

End point of concern

Figure 6.1 Stages of assessment in a risk analysis approach. (Redrawn from Bacher and Black 2008.)

reduced individual weight, but the acceptability of this growth detriment (economic CC) is a judgment call for the farmer. Similarly, if biodeposition pushes the benthos toward hypoxia, how much is too much? Various classification schemes have been used to define these steps in a risk analysis approach (Bacher

and Black 2008) (Fig. 6.1), but more objective criteria for these thresholds are desirable. There are two further fundamental approaches to CC models: (1) prediction of individual growth and, by extension, biomass that can be supported. In this approach, the assumption is that if the growth trajectory of

138

Shellfish Aquaculture and the Environment

a single animal can be predicted, then population biomass can be increased in the model until growth rate declines to a certain level. In this case, biomass in culture increases as the animals grow. We note that the ground thruthing is this case is very robust since mussel growth can be quantified with field growth trials. (2) Biomass is prescribed as a forcing on seston and nutrients. Seston may be depleted in this setup, but declining seston does not alter shellfish biomass. Changing food conditions caused by grazing can, however, be used to examine individual growth rate in a bioenergetic submodel. Models of CC may be of several trophic types, but in our synopsis they share in common a reliance on food as a limiting factor. Most of the models considered in this chapter are of the type involving phytoplankton– nutrients–zooplankton (PNZ; Fig. 6.2), that is, trophic webs, but population models are also examined. One of the main distinctions between models is spatial resolution, ranging from box to map-compliant models. In addi-

tion, the major emphasis is on farm production rather than environmental impact. In all cases, we do not attempt to review numerous examples, but rather consider the generalities of each model type, followed by highlighting selected case studies. We pose the following questions with a focus on mussel aquaculture while recognizing the applicability to other bivalve species:

• What types of models exist and how do they vary in spatial resolution and ecosystem structure? • What kind of outputs do these models provide relative to farm management? • How can model results be used as advice? • What is the relationship of CC models to ecosystem-based management? In this chapter, we emphasize the utility and application of models in farm management, as a subset of the application of ecosystembased management to coastal environments. Although the models discussed below are spe-

Coastal waters

Seston

River

Microalgae

Mussels

Nutrients

Sediment O2, H2S, fauna

Figure 6.2 PNZ model with the inclusion of the mussel submodel as well as connections with coastal waters, river, and sediment compartments. Dashed arrows represent advective or diffusive exchange, while solid arrows represent biogeochemical processes.

Dynamic modeling and production carrying capacity

cific to shellfish culture, they are in some cases generic enough to include effects in any coastal ecosystem. Their continued development and application represents real progress in coastal zone management.

Physical oceanographic models Seston depletion through suspension feeding constitutes a demand on phytoplankton and other particulates, whereas advection-diffusion renews this supply. These opposing fluxes underline the importance of physical models in the overall determination of CC for coastal aquaculture. A full discussion of this topic is beyond the scope of the present chapter, but some generalizations are provided. In the case of a zero-dimensional (0D) model (volumetric exchange as a single bulk parameter), simple physical models based on tidal prism may be employed. These models may be configured to include the role of river input as an additional flushing term via salinity gradients (Luketina 1998). Increased requirements for circulation in CC models require corresponding spatial resolution and hydrodynamic linkage between regions of a given estuary or bay. A finite element or finite difference grid of boundaries and bathymetry is often employed for this purpose. Tidal harmonics and wind forcing are then applied according to the Navier– Stokes equations to produce a numerical model of circulation. In cases of temperature or salinity stratification, it is necessary to set up a three-dimensional (3D) model, but in shallow and/or well-mixed waters, a twodimensional (2D) model is sufficient. The latter is simpler since vertical diffusivity is not explicit. There are many expressions that have been applied to the renewal of water in coastal ecosystems, including turnover time, exchange, flushing, e-folding time, and so on (Monsen et al. 2002). The latter authors calculate flushing time Tf as the “volume of water in a defined

139

(bounded) system (V) divided by the volumetric flow rate (Q) through the system.” Tf =

V Q

(6.1)

In the case of triangular or other grids, the volume exchange in a grid element may be calculated from water velocity and crosssectional area (Filgueira et al. submitted). In this example, we use the inverse of flushing time, expressed as exchange rate (t−1), which may then be multiplied by any scalar to determine mass flux. As Monsen et al. (2002) point out, there may be assumptions of complete renewal, whereas quantities such as e-folding assume less complete (e.g., 1/e) exchange. We note that the there is a difference between localized exchange (between adjacent elements) and renewal time of water with the coastal ocean (Koutitonsky et al. 2004), the latter concept embodied in residence time (RT) (Monsen et al. 2002). As models progress from a one-box tidal prism that includes a spatially averaged RT to a finite element grid, the concept of localized exchange becomes formalized. Temporal averaging is also important in the application of physical forcing to CC models. Many biogeochemical models operate on daily time steps, whereas hydrodynamics change on an hourly or shorter timescale. In this case, it may be necessary to average over tides, so that the higher frequency temporal variation is smoothed. Coupling of physical models with biogeochemical components may occur in real time, that is, the hydrodynamic model is unified with the ecosystem portions, driving the latter on the same timescale. This arrangement is common in PNZ models, and applied in cases of specialized software (see below). In contrast, the output of hydrodynamic models may be saved as a matrix of flows or exchange coefficients to be used within the ecosystem model. In either case, the topology of spatial elements must be maintained, that is, location and

140

Shellfish Aquaculture and the Environment

arrangement of cells, so that fluxes of materials and solutes are correctly prescribed.

Filtration and seston depletion The basis for quantification of seston depletion and its use in CC models is an understanding of the mechanisms involved in suspension feeding (see Chapter 4 in this book). Bivalve feeding activity has been studied extensively and shown to depend on endogenous factors such as body size or reproductive stage as well as exogenous forces such as temperature and seston quality (e.g., Bayne 1976; Shumway and Parsons 2006). The effect of seston characteristics on bivalve physiological behavior has stimulated a remarkable degree of controversy. Filtration has been considered to be subject to physiological regulation (Bayne 1998) with the purpose of maximizing energy uptake. Alternatively, filtration may be understood as an “automatized” process (Jørgensen 1996), whereby under optimal conditions (pressure, temperature) the filtration pump system is designed to function at maximum capacity (see discussion in Ward and Shumway 2004). These arguments notwithstanding, from an ecological point of view it is hard to deny that particulate food has a significant influence on bivalve feeding physiology (Grant 1996) and consequently on CC estimations. The supply of food in marine environments is directly related to hydrodynamics insofar as primary production depends heavily on physical processes that transport cells and nutrients (Cloern 1996; Lucas et al. 1999). At small scales, the effect of flow on bivalve feeding activity and growth has been examined on mussel beds. For example, Frechette et al. (1989) observed food depletion in dense mussel beds when turbulent renewal was low, a result confirmed in later studies (Muschenheim and Newell 1992; Nielsen and Maar 2007). At larger scales, water quality may be controlled by mussel populations (Møhlenberg

et al. 2007), making advection a limiting factor in mussel bed production in shallow sites such as the Oosterschelde (van Stralen and Dijkema 1994). Dolmer (2000) observed benthic mussels to reduce algal concentration below a threshold where valves close and filtration ceases, causing a vertical gradient in chlorophyll concentration. A subsequent increase in near-bed current velocities homogenized the water column in terms of chlorophyll and increased valve opening. At still larger scales, Blanton et al. (1987) described for the Ria de Arosa (northwest Spain) a significant positive relationship between mussel condition and upwelling as a result of nutrient addition and consequent increase in primary production. In the same ecosystem, Figueiras et al. (2002) demonstrated that mussel growth occurs mainly during the upwelling season. Smaal et al. (2001) showed how the construction of a barrier in the Oosterschelde, and the subsequent alteration of hydrodynamics, caused a change in the relationship between condition index and standing stock, suggesting overstocking. We conclude from natural systems that food is limiting in bivalve production, and that water renewal is critical on a variety of spatial scales in providing food. As promulgated by previous authors, these are the foundations of CC estimations (Dame 1996; Wildish and Kristmanson 1997).

Single-box models As a consequence of the above concepts, the importance of water exchange to bivalve food supplies was the cornerstone of the first CC models. These were constructed as “one-box” models, that is, no spatial domain, and were characterized by a set of scaling arguments in a spreadsheet-type motif (e.g., Carver and Mallet 1990). Incze et al. (1981) modeled CC for mussel aquaculture at lease scale in open systems by comparing gross energy needed by

Dynamic modeling and production carrying capacity

a cultivated population with available energy flow through the culture area. Officer et al. (1982) studied the control on phytoplankton populations comparing the time that benthic filter feeders need to clear the water with phytoplankton growth rate. Newell and Shumway (1993) combined oceanographic and ecophysiological data to study the sensitivity of mussel growth and harvest yield in mussel beds depending on factors controlled by the farmers, seeding density and site location. In addition, the later study demonstrated the effect of cultured biomass on phytoplankton depletion, which was suggested as a tool to manage culture. Herman (in Heip et al. 1995) integrated water RT, primary production time (PPT), and bivalve filter-feeder clearance rates to suggest a model to explore CC. These types of rates were used by Dame and Prins (1998) to examine the CC of suspension feeding bivalves in 11 coastal and estuarine ecosystems. They defined the following turnover times: (1) water RT is the time it takes for the volume or mass of water within a basin to be replaced with water from outside the system; (2) PPT is the ratio of yearly averages of phytoplankton biomass to phytoplankton primary production within the particular ecosystem; and (3) bivalve clearance time (CT) is the time needed for total bivalve filter-feeder biomass within an ecosystem to filter particles from total system volume. The comparison of these indices provides a simple way to explore ecosystem processes related to the role that bivalve populations exert on phytoplankton communities and therefore CC. Gibbs (2007) reviewed these indices, with special emphasis on CC analysis as well as functional changes in the system and their ecological meaning. This analysis of CC corresponds to 0D models, in which the ecosystem is contained within a single box but with no spatial resolution. We note that these models are static in time and have no predictive power. Guyondet et al. (2005) produced a sophisticated version of this type of model in which

141

they used a 3D circulation model to characterize subregions of an estuary. They then applied a clearance index as described above for Dame and Prins (1998) to demonstrate the importance of physical exchange in dominating food supplied to cultured oysters. This paper also showed the inadequacy of tidal prism calculations for interior subregions. There is temptation to use 0D models and phytoplankton depletion as a form of impact assessment, relative to ecological CC. However, the lack of physical rigor associated with tidal prism calculations in complex estuaries, and absence of water quality standards based on chlorophyll, decry this application. Higher-order spatial models are more appropriate to this end as detailed below. In order to describe ecosystem processes more fully, it is necessary to move to dynamic models using ordinary differential equations to describe the trophic components. This approach is classically applied within PNZ models (Kremer and Nixon 1978). The PNZ model also incorporates mussels (M) and detritus (D) submodels as follows: dP = + Pgrowth − Pmortality − Mgrazing − Zgrazing dt ± Pmixing (6.2) dN = + N river + Mexcretion + Zexcretion − Puptake dt (6.3) ± N mixing dZ = + Znet growth − Zmortality − Mgrazing ± Zmixing dt (6.4) dD = + Dresuspension + Mfeces + Pmortality + Zmortality dt − Dsinking − Mgrazing ± Dmixing (6.5) dM = + Mnet growth + Mseeding − Mmortality dt − Mharvesting

(6.6)

142

Shellfish Aquaculture and the Environment

The mixing term includes water exchange in the study area. Although the single-box model traditionally uses simple physics, for example, tidal prism, examples such as Guyondet et al. 2005 (described above) expand the spatial realism of the simple spatial domain. Similarly, Dowd (2003) represented exchange within a bay using a one-dimensional (1D) tracer equation, allowing spatially dependent estimations of CC for mussel culture. A special case of the one-box model is that of Ecopath, an ecological model based on trophic mass balance (www.ecopath.org). In this approach, the net biomass production of a population is equal to the sum of consumption by predators, growth, reproduction, and other components, including harvest in the case of a commercial species. Ecopath is graphical user interface (GUI)-based and has a long history in marine applications. It was used to predict the production and ecological CC of bivalve shellfish in two bays in New Zealand (Jiang and Gibbs 2005) based on the analysis of the whole food web, integrating 23 components. A comparison between Ecopath and dynamic models is beyond the context of this chapter. Although PNZ models are criticized by these authors, we maintain that assessment of CC mandates explicit parameterization of physical particle renewal and bivalve ecophysiology as detailed in the PNZ approach.

Higher-order models The PNZ approach provides the basis for models of increasing spatial detail. Greater spatial resolution conveys the ability to define fluxes in at least two dimensions, usually XY (2D); 3D models are distinguished by the inclusion of depth, necessary where there is density or temperature stratification and/or where vertical layers of suspended food must be addressed. In both cases, their spatial definition may be classified into two groups, box

models and fully spatial (map-compliant) models. The difference between them is related to spatial resolution and how physics is included in the model. The spatial resolution of box models is lower than the fully coupled case, and the results can be only crudely mapped. Physical forcing from a numerical model must be condensed to correspond to the boxes. Fully spatial models are often built on a grid, in some case identical to the hydrodynamic grid, and the physics may have the same resolution as the ecosystem model. The physical submodel is then either coupled directly to the biogeochemical submodel, running circulation in real time, or delivers a matrix of velocity or exchange coefficients as a forcing function. Grant et al. (2007) developed a box model of CC for Lagune de la Grande-Entrée (Québec, Canada), in which the domain was divided into four discrete connected boxes (Fig. 6.3). The structure of a typical box model includes (1) relationships between different compartments as well as (2) boundary conditions, that is, the time series from outside of the model domain that force the results, and (3) spatial connections, in the form of hydrodynamics related to the described spatial arrangement. This study places special emphasis on three important aspects related to modeling: ground truthing, parameter sensitivity, and predictive management capabilities. The ground truthing process was carried out by comparison of observed data versus model predictions for two variables: (1) mussel growth, and (2) nitrate and phytoplankton concentrations in the water column. This paper also includes a sensitivity analysis with subsets of parameter combinations (Plackett and Burman 1946) providing essential information on the relative importance of each parameter in model outputs (Hamby 1994). In addition, this study shows the potential utility of ecological models in aquaculture management. The authors ran different model scenarios based on comparing mussel size with

Dynamic modeling and production carrying capacity

(A)

Depth (m)

(B) –67.583˚ –65.147˚ –62.750˚ –60.333˚ –57.917˚

143

8 7 6 5

Newfoundland 48.333˚

48.333˚

4

New Brunswick

3

45.917˚

2

45.917˚ Nova Scotia

–62.750˚

–61.750˚

–61.500˚

House Harbour 1 Lagoon

–61.333˚

3 47.583˚

2

Gulf of St. Lawrence 47.500˚ –61.750˚

47.500˚ –61.500˚

(C)

Model Structure Zooplankton

Mussels

Phytoplankton

Detrital Matter Nutrients

Spatial Connections

4 47.583˚

0

–57.917˚

Boundary Conditions

–67.583˚

1

Figure 6.3 Lagune de la Grande-Entrée, Iles-de-la-Madeleine, location and site map with modeled boxes (A), bathymetry of the model domain (B), and simplified model structure (C). (Adapted from Grant et al. 2007.)

stocking biomass in the various boxes in order to optimize the production. In this case, the CC criterion was not explicit, but biomass– size curves allow selection of yield based on stocking density. Box models entail coarse spatial resolution, but their computational simplicity and condensed hydrodynamics constitute a strong point, resulting in a valuable initial approach to culture management. Optimization routines (see below) applied to parameterization also run more easily since boxes are few in number.

Fully spatial models Despite the utility of box models, their low spatial resolution limits the potential for managing culture placement. More fully spatial models are consistent with ecosystem-based

management goals that involve marine spatial planning (Douvere 2008). Spatial resolution also affects the results of the model especially when processes are concentration dependent (Fennel and Neumann 2004). A fully coupled model requires the linking of a physical submodel of water exchange within the study area to an ecophysiological submodel describing organisms and their interactions. Organization and tracking of the model topology requires a computation scheme for handling the resultant matrices. One of the most common approaches is via high-level programming languages such as MATLAB (www.mathworks.com), for example, in Fennel and Neumann (2004). In some cases, biological models are linked directly to physical submodels within the same software environment, such as DHI’s MIKE physical models joined with their ECO Lab solver

144

Shellfish Aquaculture and the Environment

63˚02’

63˚00’

63˚02’

63˚00’ mg C m–3

46˚25’

50

1200

100 46˚24’

1000 800

200

200

100

500

300

600

900

1000

46˚23’

600

600 400 300 200 (A) 46˚22’

(B)

63˚02’

100

Spring –3 10 mussels m

Spring –3 0 mussels m 63˚00’

63˚02’

0 63˚00’

Figure 6.4 Modeled maps of chlorophyll in Tracadie Bay for spring conditions of (A) zero mussels and (B) 10 mussels m−3 in the farm areas. (Source: Grant et al. 2008.)

(www.dhigroup.com). This is also the case for the freeware GEMSS (Generalized Environmental Modeling System for Surfacewaters; www.erm-smg.com) and ROMS (Regional Ocean Modeling System; www.myroms.org) (Fennel et al. 2006). In the context of bivalve aquaculture, there are several examples of fully coupled models applied to CC of different sites, for example, Pastres et al. (2001), Duarte et al. (2003), and Grant et al. (2008). In the latter paper, maps of seston depletion at a mussel aquaculture site in eastern Canada indicate the effects of dense mussel farming, including mapped output in these types of models (Fig. 6.4). In other cases, separate physical and biological models have been joined to create fully spatial models for bivalve aquaculture. Among these tools is EcoWin2000 (www. ecowin2000.com), which provides a platform for integration of various other models, and adds functionality of its own (Ferreira et al. 2008) (Fig. 6.5). In this example, a 3D physical model (Delft3D; www.sobek.nl) pro-

vides the spatial arrangement of the study area and the simulation of hydrodynamic transport. The shellfish physiology is simulated by means of an individual bivalve growth model called ShellSIM (www.shellsim.com) (Hawkins et al. 2002). Finally, EcoWin2000 acts as a core, coupling both models and providing other necessary submodels to run different management scenarios. Filgueira et al. (submitted) developed a modeling approach to create fully spatial physical-biological models based on the coupling of output from Aquadyn (www.synexusglobal.com), a 2D hydrodynamic model, and Simile (www.simulistics. com), an object-oriented modeling package. These types of models have a long tradition in ecosystem simulation, made famous by Stella (www.iseesystems.com). However, Simile has special capabilities for creating and maintaining spatial topology. Filgueira et al. (submitted) cite advantages of this approach as follows: (1) the coupling procedure is open source and only minor changes are necessary to adapt the model to different locations;

Dynamic modeling and production carrying capacity

Delft3D

Run Delft 3D for large domain (Western Irish Sea and four loughs) using a fine grid (each lough has hundreds of cells)

Define larger boxes (<50) within the loughs with GIS for E2K using (І) current and bathymetry data (іі) WFD (ііі) aquaculture distribution

Use D3D to calculate water fluxes across these larger boxes at 30m intervals, and at the seaward boundary of the lough domain – supply these offline as spreadsheets

Hydrodynamic transport simulated in E2K by reading these spreadsheet files during an E2K model run

145

ShellSIM EcoWin2000

Enter larger box areas, volumes etc from GIS into E2K

Implement individual growth model in E2K, test, and then add population dynamics

Experimentally simulate individual shellfish physiology using raceways

Use results to conceptualise an energy budget model for individual growth

Run full E2K model, calibrate and validate

Management scenarios

Extract E2K initial conditions for state variables in each box, boundary conditions and calibration data

Implement and validate individual growth model in Stella or other simple modelling package

Prepare a water quality database including both the individual lough and inputs from freshwater and ocean boundaries (measurements)

Measure individual shellfish growth rates in the field

Figure 6.5 Scheme of physical–ecological coupling carried out by EcoWin2000. (Source: Ferreira et al. 2008.)

(2) the biogeochemical model can be modified using a GUI interface; (3) the results are easily plotted, mapped, and exported to a geographic information system (GIS); and (4) optimization tools are included in the package.

Population-based models In addition to ecosystem process models, there are approaches that explore CC in terms of population dynamics models. These take into account the age/size structure of a cultured population to simulate temporal variation of abundance in each weight class, total standing stock, and system production. (Gangnery et al. 2001). This model design allows a detailed description of size- and time-dependent processes such as mortality, seeding, and harvest-

ing. Different seeding events provide new individuals that will grow according to an empirical growth function based on available food supplies (Fig. 6.6). A detailed description related to seeding and harvesting processes provides scenarios to explore the effect of different farming procedures, culture methods, and seeding time as applied to Thau Lagoon (France). Another utility related to farm management involves the effect of different ecological events on bivalve populations, for example, temporary closure of oyster harvesting due to a toxic algal bloom in Thau Lagoon (Gangnery et al. 2004). Although the model is based on population structure, it can be used to estimate the biological component of Dame and Prins’s (1998) ratios. For example, Gangnery et al. (2001) estimated population clearance rate in

146

Shellfish Aquaculture and the Environment

r nn

r Minimum harvest weight

n3 m

G

n2 m m e

n4

n4

m

m

n3 m

n3 m

n2

n2

t+1

Weight

n1 m

e

r

m

n1 m

e t

nk m

m

n1

n1

m

nk m

m e

t+2

t+n

Time Figure 6.6 Conceptual scheme of the population dynamic model, where n is the number of individuals whose weight is in the range w to w + dw at time t, G is the growth rate (g day−1), related to the individual growth variance, m is the mortality rate (day−1), r is the harvest rate (g day−1), and e is the seeding. (Source: Gangnery et al. 2001.)

comparison to lagoon volume, providing a measurement of bivalve CT (see above). Assigning the above precautions to system-wide estimates of clearance and flushing, this approach can be used to manage cultured populations via CC criteria as in 0D models described previously. In addition, this kind of model can be coupled to an ecosystem model allowing a detailed description of demography within 1D, 2D, or 3D models (Ferreira et al. 1998). Although not involving aquaculture, there is a prolific series of simulation models for American oyster populations dealing with disease susceptibility, harmful algal blooms, and climate change scenarios (freshwater flow, temperature). Many of these studies involve 3D hydrodynamic models coupled to ecophysiology and population models of oysters and clams, representing a significant contribution to shellfish modeling over the last 20 years (e.g., Klinck et al. 2002; Hofmann et al. 2006).

Local models Although the term “ecosystem” implies bay or estuary length scales, the entire trophic system is realized at farm scales, especially since this is the level at which seston depletion may be most obvious (Grant et al. 2007). In this case, seston depletion is viewed as a succession of decreasing particle concentrations, offset by a physical renewal term. One of the first quantitative models of shellfish CC is based on this scale (Incze et al. 1981). The concept implied herein is readily depicted (Fig. 6.7). Pilditch et al. (2001) established one of the more quantitative models of this type since it included diffusionbased renewal of seston. They showed how the increase in lease area at a site in Nova Scotia could cause a seston depletion of 20–50%, inducing a decrease in scallop growth. One of the most important advantages of farm-scale models is related to farm

Dynamic modeling and production carrying capacity

Farm length

Width

Current Chl a POM

Current

Shellfish Depth

147

1

2

3

n–1

n

Sections

Chl a POM

Figure 6.7 Farm-scale model layout. (Source: Ferreira et al. 2008.)

management. For example, in the same study, Pilditch et al. (2001) suggested that food to scallops could be enhanced by altering the lease geometry, potentially increasing economic profit without further cost. Similarly, Aure et al. (2007) employed a minimum chlorophyll concentration (filtration threshold) to examine the effect of farm aspect ratio on mussel seston depletion. This approach has been made more widely available via the FARM model (Farm Aquaculture Resource Management; www. farmscale.org; Ferreira et al. 2007), a local scale seston depletion model in which a Cobb–Douglas function was added to determine the optimal stocking density in terms of maximum economic profit. In order to carry out environmental assessment, FARM applies a subset of ASSETS methodology (Assessment of Estuarine Trophic Status; Bricker et al. 2003). Given that FARM includes both production tools and environmental assessment related to eutrophication, the model could be used by farmers and regulatory agencies. Duarte et al. (2008) developed an analytical model at raft scale (Spanish rias) to study CC (maximize growth rate) dependent on mussel density and current velocity. The novelty of this work is related to inclusion of sizedependent bivalve distribution within the farm (seed, juveniles, adults). Similar to the above papers, their results suggest that a change in culture practices (raft area, rope density) could optimize production, and consequently profit of the farm, without additional investment.

Duarte et al. (2008) also emphasized that farm size and spatial arrangement could be important variables for increasing production at bay scale. This approach combines both local and bay scales, an approach with obvious advantages, but rarely applied.

Optimization The information used to set up the ecosystem models described above is sometimes difficult to obtain. These models require multiple parameters that in some cases are uncertain or not available (Smaal et al. 1997). This introduces unknown errors in model predictions and thus less confidence in the results of simulations. Optimization tools such as nonlinear parameter estimation take control of the model, running it multiple times in order to determine the optimal set of parameters that minimize the discrepancies between model output and corresponding measurements. Rosland et al. (2009) used a nonlinear optimization process (Nelder–Mead) to estimate parameters of a DEB model applied to blue musels under low-seston conditions. In the same way, Filgueira et al. (2010) used PEST (Parameter ESTimation; www.pesthomepage. org), an optimization tool that uses the Gauss–Marquardt–Levenberg algorithm to estimate water exchange in Lysefjord (Norway) using chlorophyll as a conservative tracer.

148

Shellfish Aquaculture and the Environment

Optimization tools can also be used in ecosystem management to determine the most suitable scenario according to a given CC criterion. For example, Filgueira et al. (2010) used this approach to estimate mussel biomass that could be grown in a Norwegian fjord based on maintaining seston depletion within limits set by natural variation in chlorophyll at the ocean boundary. The use of optimization tools is growing in applied ecological modeling (Reis and Nitter 2008). Its application to aquaculture models has been limited, but it is clearly a powerful method in the estimation of poorly known parameters and the mathematical expression of CC.

Application to management Ecological system models are robust decisionmaking tools because they simulate system organization, function, and change (Odum

(A)

and Odum 2000) within complex manipulated ecosystems (Dowd 2005). Specifically, the potential to run alternative scenarios allows exploration of the consequences of changing the standing stock of bivalves or their distribution within a bay. The management potential of ecosystem models is explicit in some of the examples already provided, but we delve into further specifics here. Pastres et al. (2001) developed a fully coupled spatial model of the Venice Lagoon based on a 3D physical model and a PNZ trophic model with the addition of Tapes philippinarum and seston submodels. They calculated a Potential Growth Index (PGI), which relates phytoplankton availability to bivalve energetic requirements. Using this index as well as indices of sediment health and bathymetry, the authors created a map of potential rearing areas using GIS techniques (Fig. 6.8A). Given that PGI is a static index based on the energetic requirements of an individual, it

(B) Density (# m–3) 100 20

20

20 10

70 80 60

60

70

50 40 30

30

0.75 < E < 1 <0.75

70

70

50

40

>1

80

60

30

40

50

10

60

30

30

90

20 10

Figure 6.8 (A) Classification of the potential rearing areas in Venice Lagoon. (Source: Pastres et al. 2001.) (B) Chlamys farreri density map maximized by accepting a 10% penalty in final total weight. (Source: Bacher et al. 2003.)

Dynamic modeling and production carrying capacity

does not take into account population density, and thus top-down regulation of phytoplankton. The authors carried out three simulations using different seeding densities observing that transport processes were more important for bivalve growth than indicated by the PGI value. In addition, scenario analysis showed differential growth depending on the spatial location, which could be used to adjust seeding density and optimize shellfish production. Bacher et al. (2003) used a 2D spatial model to examine the effect of food depletion on Chlamys farreri growth in Sungo Bay (China). A detailed spatial mesh was necessary to track food concentration and determine in each grid cell the standing stock biomass that could be supported according to an established criterion. The chosen metric was a growth decrease of 10% compared with growth determined without density effects. The results appear as a map of projected culture density where the 10% growth criterion is conserved, providing a spatial decision support tool for scallop management (Fig. 6.8B). Both of the above examples are related to production CC; however, other criteria can be applied in the same way to estimate ecological CC.

Modeling environmental impact As stated previously, one aspect of CC models involves environmental impact, but this area of model development lags behind that of production CC based on food limitation. Although it could be argued that seston depletion is a form of environmental impact, there has been greater emphasis on benthic effects, namely organic loading due to biodeposition. These processes involve production of feces and pseudofeces, sedimentation and resuspension rates (see Chapter 10 in this book), particle aggregation, and a whole suite of benthic metabolic processes including aerobic and anaerobic respiration.

149

In this regard, modeling of culture impact involves particle dynamics. The properties of complex organic-mineral aggregates are poorly known, and must be understood in the context of a shallow water benthic boundary layer typical of aquaculture sites. In light of these parameterization difficulties, it is not surprising that suspended sediment transport models of aquaculture effects are uncommon. One of the best examples of this genre comes from a study of salmon cage impacts in which waste feces and food were dispersed within a 3D hydrodynamic model (Panchang et al. 1997). More commonly, aquaculture waste has been modeled locally using tools such as DEPOMOD specifically applied to shellfish (Weise et al. 2009) or customized models (Giles et al. 2009). Improvements in empirical understanding of the behavior of shellfish biodeposits (Giles and Pilditch 2004) have led to greater potential to develop these models at the ecosystem scale. In general, carbon loading has been related empirically to benthic biodiversity as a means of establishing acceptable biodeposition thresholds (Weise et al. 2009) Taking a somewhat different approach, Grant et al. (2005) calculated a ratio between the mass of mussel biodeposits produced and the ability of estuaries to flush them. This model obviated some of the particle behavior since deposition was not simulated. Instead, particle loading was compared with the background of suspended solids to determine if mussel input could be “noticed” as a source. Diagenetic models are well developed in general for marine sediments, and this progress has translated into aquaculture-specific models of organic loading. For example, Thau Lagoon is subject to urban eutrophication as well as extensive oyster and mussel culture (Bacher et al. 1995). These types of shallow environments may be somewhat complex to model since they contain both macroalgae and benthic microalgae that are influenced by nutrient loading. A wide range of models of nitrogen regeneration, oxygen dynamics,

150

Shellfish Aquaculture and the Environment

primary production, shellfish growth, and populations have been completed for this area. Chapelle et al. (2001) used a coupled physicalbiogeochemical model to examine anoxia in Thau Lagoon and suggest ways that change in oyster culture location and density could be used to improve oxygen conditions.

Sustainability and ecosystem-based management As models emerge from research to operational tools, they incorporate GIS, decision support, habitat suitability, and a variety of approaches associated with increased spatial resolution (Goulletquer and Le Moine 2002; Vincenzi et al. 2006; Nobre and Ferreira 2009). In addition, a number of research projects have addressed modeling of shellfish aquaculture, for example, ECASA (Ecosystem Approach for Sustainable Aquaculture; www.ecasa.org.uk/), which is a comprehensive examination of tools, indicators, and models for assessing various types of aquaculture from an ecosystem perspective. It is obvious that an approach based on entire ecosystems has the capacity to provide broadly based information on aquaculture management. Because trophic structure and other fundamentals of ecosystem structure are included in these models, they could easily be used to manage other aspects of coastal systems such as eutrophication (Nobre and Ferreira 2009). The term ecosystem-based management has many dimensions including capture fisheries, coastal community involvement, and so on. In our context, this term may be defined as decision making based on maintaining ecosystem functioning (stocks and fluxes within ecosystem models) within the bounds of natural variation.

This standard could be applied to the effects of seston depletion as well as to benthic impacts. In field studies, both seston depletion

and benthic impacts tend to have a focus at the local scale because measurements are feasible. One reason that ecosystem models are so useful in studies of CC is that they can make predictions about the far field (Grant et al. 2008); we suggest that the ecosystem scale is more relevant to sustainability than the local scale. Ferreira et al. (2008) examined culture location, effects of climate change (temperature increase), and interaction with wild bivalves at the ecosystem scale using modeling as described above (Fig. 6.5). The approaches to sustainability associated with marine farming include integrated multitrophic aquaculture (IMTA) or polyculture. This practice has been modeled for shellfish culture in which macroalgae and bivalves are grown together (Duarte et al. 2003); however, the benefits of managing culture wastes are probably less than for fed aquaculture where exogenous nitrogen is introduced. It is clear that sustainable aquaculture will not be managed by dynamic modeling alone, but by a combination of GIS, modeling, and coastal zone management that includes input from coastal communities. Decision support is key to bringing science into policy including risk analysis (Bacher and Black 2008) and the DPSIR (Driving Forces-Pressures-StateImpacts-Responses) approach that links economic and ecological factors at the ecosystem scale (Nobre 2009). Gibbs (2009) is skeptical that dynamic PNZ models can reliably be used for CC, a view that is obviated by the successful what-if management scenarios provided by application of validated models in multiple locations (Marinov et al. 2007; Bacher and Black 2008; Ferreira et al. 2008; Filgueira and Grant 2009). Therefore, dynamic simulation models constitute a powerful technique in exploring complex ecosystem processes and consequently CC of shellfish aquaculture. Additionally, they are an important approach in examining aquaculture in the context of ecosystem-based management.

Dynamic modeling and production carrying capacity

Literature cited Aure, J., Strohmeier, T., and Strand, Ø. 2007. Modelling current speed and carrying capacity in long-line blue mussel (Mytilus edulis) farms. Aquaculture Research 38:304–312. Bacher, C., and Black, E. 2008. Risk assessment of the potential decrease of carrying capacity by shellfish farming. In: Assessment and communication of environmental risks in coastal aquaculture. GESAMP Reports and Studies No 76. FAO, Rome. pp. 90–111. Bacher, C., Bioteau, H., and Chapelle, A. 1995. The impact of a cultivated oyster population on the nitrogen dynamics: the Thau Lagoon case (France). Ophelia 42:29–54. Bacher, C., Grant, J., Hawkins, A., Fang, J., Zhu, P., and Duarte, P. 2003. Modeling the effect of food depletion on scallop growth in Sungo Bay (China). Aquatic Living Resources 16:10–24. Bayne, B.L. 1976. Marine Mussels: Their Ecology and Physiology. Cambridge University Press, London. Bayne, B.L. 1998. The physiology of suspension feeding by bivalve molluscs: an introduction to the Plymouth “TROPHEE” workshop. Journal of Experimental Marine Biology and Ecology 219:1–19. Blanton, J.O., Tenore, K.R., Castillejo, F., Atkonson, L.P., Schwing, F.B., and Lavin, A. 1987. The relationship of upwelling to mussel production rias on the western coast of Spain. Journal of Marine Research 45:497–511. Bricker, S.B., Ferreira, J.G., and Simas, T. 2003. An integrated methodology for assessment of estuarine trophic status. Ecological Modelling 169(1):39–60. Cabrera, V., Hildebrand, P., Jones, J.W., Letson, D., and de Vries, A. 2006. An integrated North Florida dairy farm model to reduce environmental impacts under seasonal climate variability. Agriculture, Ecosystems, and Environment 113:82–97. Carver, C.E.A., and Mallet, A.L. 1990. Estimating the carrying capacity of a coastal inlet for mussel culture. Aquaculture 88(1):39–53. Chapelle, A., Lazure, P., and Souchu, P. 2001. Modelling anoxia in the Thau Lagoon (France). Oceanologica Acta 24:87–97.

151

Cloern, J.E. 1996. Phytoplankton bloom dynamics in coastal ecosystems: a review with some general lessons from sustained investigation of San Francisco Bay. Reviews of Geophysics 34(2):127–168. Dame, R.F. 1996. Ecology of Marine Bivalves: An Ecosystem Approach. CRC Press, Boca Raton, FL. Dame, R.F., and Prins, T.C. 1998. Bivalve carrying capacity in coastal ecosystems. Aquatic Ecology 31:409–421. Dolmer, P. 2000. Feeding activity of mussels Mytilus edulis related to near-bed currents and phytoplankton biomass. Journal of Sea Research 22:221–231. Douvere, F. 2008. The importance of marine spatial planning in advancing ecosystem-based sea use management. Marine Policy 32:762– 771. Dowd, M. 2003. Seston dynamics in a tidal inlet with shellfish aquaculture: a model study using tracer equations. Estuarine, Coastal and Shelf Science 57:523–537. Dowd, M. 2005. A bio-physical coastal ecosystem model for assessing environmental effects of marine bivalve aquaculture. Ecological Modelling 183:323–346. Duarte, P., Meneses, R., Hawkins, A.J.S., Zhu, M., Fang, J., and Grant, J. 2003. Mathematical modelling to assess the carrying capacity for multispecies culture within coastal waters. Ecological Modelling 168:109–143. Duarte, P., Labarta, U., and Fernández-Reiriz, M.J. 2008. Modelling local food depletion effects in mussel rafts of Galician Rias. Aquaculture 274:300–312. Fennel, W., and Neumann, T. 2004. Introduction to the Modelling of Marine Ecosystems. Elsevier Oceanography Series, 72. Elsevier, Amsterdam. Fennel, K., Wilkin, J., Levin, J., Moisan, J., O’Reilly, J., and Haidvogel, D. 2006. Nitrogen cycling in the Middle Atlantic Bight: results from a threedimensional model and implications for the North Atlantic nitrogen budget. Global Biogeochemical Cycles 20:GB3007. doi: 10.1029/2005GB002456 Ferreira, J.G., Duarte, P., and Ball, B. 1998. Trophic capacity of Carlingford Lough for oyster culture—analysis by ecological modelling. Aquatic Ecology 31:361–378.

152

Shellfish Aquaculture and the Environment

Ferreira, J.G., Hawkins, A.J.S., and Bricker, S.B. 2007. Management of productivity, environmental effects and profitability of shellfish aquaculture—the Farm Aquaculture Resource Management (FARM) model. Aquaculture 264:160–174. Ferreira, J.G., Hawkins, A.J.S., Monteiro, P., Moore, H., Service, M., Pascoe, P.L., Ramos, L., and Sequeira, A. 2008. Integrated assessment of ecosystem-scale carrying capacity in shelfish growing areas. Aquaculture 275:138–151. Filgueira, R., and Grant, J. 2009. A box model for ecosystem-level management of mussel culture carrying capacity in a coastal bay. Ecosystems 12:1222–1233. Figueiras, F.G., Labarta, U., and Fernández-Reiriz, M.J. 2002. Coastal upwelling, primary production and mussel growth in the Rias Baixas of Galicia. Hydrobiologia 484:121–131. Filgueira, R., Grant, J., Strand, Ø., Asplin, L., and Aure, L. 2010. A simulation model of carrying capacity for mussel aquaculture in a Norwegian fjord: role of artificial-induced upwelling. Aquaculture 308:20–27. Filgueira, R., Grant, J., Bacher, C., and Carreau, C. submitted. A physical-biogeochemical coupling scheme for studying long-term processes in shallow coastal ecosystems. Environmental Modelling and Software. Frechette, M., and Bacher, C. 1998. A modelling study of optimal stocking density of mussel populations kept in experimental tanks. Journal of Experimental Marine Biology and Ecology 219:241–255. Frechette, M., Butman, C.A., and Geyer, W.R. 1989. The importance of boundary-layer flow in supplying phytoplankton to the benthic suspension feeder, Mytilus edulis L. Limnology and Oceanography 34(1):19–36. Gangnery, A., Bacher, C., and Buestel, D. 2001. Assessing the production and the impact of cultivated oysters in the Thau lagoon (Mediterranee, France) with a population dynamics model. Canadian Journal of Fisheries and Aquatic Sciences 58:1012–1020. Gangnery, A., Bacher, C., and Buestel, D. 2004. Modelling oyster population dynamics in a Mediterranean coastal lagoon Thau, France): sensitivity of marketable production to environmental conditions. Aquaculture 230:323–347.

Gibbs, M.T. 2007. Sustainability performance indicators for suspended bivalve aquaculture activities. Ecological Indicators 7:94–107. Gibbs, M.T. 2009. Implementation barriers to establishing a sustainable coastal aquaculture sector. Marine Policy 33:83–89. Giles, H., and Pilditch, C.A. 2004. Effects of diet on sinking rates and erosion thresholds of mussel Perna canaliculus biodeposits. Marine Ecology Progress Series 282:205–219. Giles, H., Broekhuizen, N., Bryan, K.R., and Pilditch, C.A. 2009. Modelling the dispersal of biodeposits from mussel farms: the importance of simulating biodeposits erosion and decay. Aquaculture 291:168–178. Goulletquer, P., and Le Moine, O. 2002. Shellfish farming and coastal zone management (CZM) development in the Marennes-Oleron bay and Charentais Sounds (Charente Maritime, France): a review of recent developments. Aquaculture International 10(6):507–525. Grant, J. 1996. The relationship of bioenergetics and the environment to the field growth of cultured bivalves. Journal of Experimental Marine Biology and Ecology 200:239–256. Grant, J., and Bacher, C. 1998. Comparative models of mussel bioenergetics and their validation at field culture sites. Journal of Experimental Marine Biology and Ecology 219:21–44. Grant, J., Cranford, P.J., Hargrave, B., Carreau, M., Schofield, B., Armsworthy, S., Burdett-Coutts, V., and Ibarra, D. 2005. A model of aquaculture biodeposition for multiple estuaries and field validation at blue mussel (Mytilus edulis) culture sites in eastern Canada. Canadian Journal of Fisheries and Aquatic Sciences 62(6):1271– 1285. Grant, J., Curran, K.J., Guyondet, T.L., Tita, G., Bacher, C., Koutitonsky, V., and Dowd, M. 2007. A box model of carrying capacity for suspended mussel aquaculture in Lagune de la Grande-Entrée, Iles-de-la-Madeleine, Québec. Ecological Modelling 200:193–206. Grant, J., Bacher, C., Cranford, P.J., Guyondet, T., and Carreau, M. 2008. A spatially explicit ecosystem model of seston depletion in dense mussel culture. Journal of Marine Systems 73:155– 168. Grizzle, R.E., and Lutz, R.A. 1989. A statistical model relating horizontal seston fluxes and

Dynamic modeling and production carrying capacity

bottom sediment characteristics to growth of Mercenaria mercenaria. Marine Biology 102:95–105. Guyondet, T., Koutitonsky, V.G., and Roy, S. 2005. Effects of water renewal estimates on the oyster aquaculture potential of an inshore area. Journal of Marine Systems 58:35–51. Hamby, D.M. 1994. A review of techniques for parameter sensitivity analysis of environmental models. Environmental Monitoring and Assessment 32(2):135–154. Hawkins, A.J.S., Duarte, P., Fang, J.G., Pascoe, P.L., Zhang, J.H., Zhang, X.L., and Zhu, M.Y. 2002. A functional model of responsive suspensionfeeding and growth in bivalve shellfish, configured and validated for the scallop Chlamys farreri during culture in China. Journal of Experimental Marine Biology and Ecology 281:13–40. Heip, C.H.R., Goosen, N.K., Herman, P.M.J., Kromkamp, J., Middelburg, L., and Soetaert, K. 1995. Production and consumption of biological particles in temperate tidal estuaries. Oceanography and Marine Biology: An Annual Review 33:1–149. Hofmann, E.E., Klinck, J.M., Kraeuter, J.N., Powell, E.N., Grizzle, R.E., Buckner, S.C., and Bricelj, V.M. 2006. A population dynamics model of the hard clam, Mercenaria mercenaria: development of the age-and length-frequency structure of the population. Journal of Shellfish Research 25(2):417–444. Incze, L.S., Lutz, R.A., and True, E. 1981. Modeling carrying capacity for bivalve molluscs in open, suspended-culture systems. Journal of the World Mariculture Society 12(1):143–155. Jiang, W., and Gibbs, M.T. 2005. Predicting the carrying capacity of bivalve shellfish culture using a steady, linear food web model. Aquaculture 244:171–185. Jørgensen, C.B. 1996. Bivalve filter feeding revisited. Marine Ecology Progress Series 142:287–302. Klinck, J.M., Hofmann, E.E., Powell, E.N., and Dekshenieks, M.M. 2002. Impact of channelization on oyster production: a hydrodynamic-oyster population model for Galveston Bay, Texas. Environmental Monitoring and Assessment 7(4):273–289. Koutitonsky, V.G., Guyondet, T.S., Hilaire, A., Courtenay, S.C., and Bohgen, A. 2004. Water

153

renewal estimates for aquaculture developments in the Richibucto estuary, Canada. Estuaries 27(5):839–850. Kremer, J., and Nixon, S.W. 1978. A Coastal Marine Ecosystem: Simulation and Analysis. SpringerVerlag, New York. Lucas, L.V., Koseff, J.R., Cloern, J.E., Monismith, S.G., and Thompson, J.K. 1999. Processes governing phytoplankton blooms in estuaries. I: the local production-loss balance. Marine Ecology Progress Series 187:1–15. Luketina, D. 1998. Simple tidal prism models revisited. Estuarine, Coastal and Shelf Science 46:77–84. Marinov, D., Galbiati, L., Giordani, G., Viaroji, P., Norro, A., Bencivelli, S., and Zaldivar, J.M. 2007. An integrated modelling approach for the management of clam farming in coastal lagoons. Aquaculture 269:306–320. Møhlenberg, F., Petersen, S., Petersen, A.H., and Gameiro, C. 2007. Long-term trends and shortterm variability of water quality in Skive Fjord, Denmark—nutrient load and mussels are the primary pressures and drivers that influence water quality. Environmental Monitoring and Assessment 127:503–521. Monsen, N.E., Cloern, J.E., Lucas, L.V., and Monismith, S.G. 2002. A comment on the use of flushing time, residence time, and age as transport time scales. Limnology and Oceanography 47(5):1545–1553. Muschenheim, D.K., and Newell, C.R. 1992. Utilization of seston flux over a mussel bed. Marine Ecology Progress Series 85:131– 136. Newell, C.R., and Shumway, S.E. 1993. Grazing of natural particles by bivalve molluscs: a spatial and temporal perspective. In: Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Coastal Ecosystems. Springer-Verlag, New York, pp. 85–148 Nielsen, T.G., and Maar, M. 2007. Effects of a blue mussel Mytilus edulis bed on vertical distribution and composition of the pelagic food web. Marine Ecology Progress Series 339: 185–198. Nobre, A.M. 2009. An ecological and economic assessment methodology for coastal ecosystem management. Environmental Management 44: 185–204.

154

Shellfish Aquaculture and the Environment

Nobre, A.M., and Ferreira, J.G. 2009. Integration of ecosystem-based tools to support coastal zone management. Journal of Coastal Research SI 56:1676–1680. Odum, H.T., and Odum, E.C. 2000. Modeling for All Scales. An Introduction to System Simulation. Academic Press, San Diego. Officer, C.B., Smayda, T.J., and Mann, R. 1982. Benthic filter feeding: a natural eutrophication control. Marine Ecology Progress Series 9:203–210. Panchang, V., Cheng, G., and Newell, C. 1997. Modeling hydrodynamics and aquaculture waste transport in coastal. Maine Estuaries 20: 14–41. Pastres, R., Solidoro, C., Cossarini, G., Melaku Canu, D., and Dejak, C. 2001. Managing the rearing of Tapes philippinarum in the lagoon of Venice: a decision support system. Ecological Modelling 138:231–245. Pilditch, C.A., Grant, J., and Bryan, K.R. 2001. Seston supply to sea scallops (Placopecten magellanicus) in suspended culture. Canadian Journal of Fisheries and Aquatic Sciences 58:214–253. Plackett, R.L., and Burman, J.P. 1946. The design of optimum multifactorial experiments. Biometrika 33:305–325. Reis, S., and Nitter, A. 2008. Ecological models, optimization. In: Jorgensen, S.E., and Fath, B.D. (eds.), Encyclopedia of Ecology, Vol. 2. Elsevier, Oxford, pp. 1058–1064. Rosland, R., Strand, Ø., Alunno-Bruscia, M., Bacher, C., and Strohmeier, T. 2009. Appliyng Dynamic Energy Budget (DEB) theory to simulate growth and bio-energetics of blue mussels under low seston conditions. Journal of Sea Research 62:49–61. Shumway, S.E., and Parsons, G.J. 2006. Scallops: Biology, Ecology and Aquaculture. Elsevier, Amsterdam.

Smaal, A.C., Prins, T.C., Dankers, N., and Ball, B. 1997. Minimum requirements for modelling bivalve carrying capacity. Aquatic Ecology 31(4):423–428. Smaal, A., van Stralen, M., and Schuiling, E. 2001. The interaction between shellfish culture and ecosystem processes. Canadian Journal of Fisheries and Aquatic Sciences 58:991–1002. van Stralen, M.R., and Dijkema, R.D. 1994. Mussel culture in a changing environment—the effects of a coastal engineering project on mussel culture (Mytilus edulis L.) in the Oosterschelde Estuary (SW Netherlands). Hydrobiologia 283:359– 379. Vincenzi, S., Caramori, G., Rossi, R., and De Leo, G.A. 2006. A GIS-based habitat suitability model for commercial yield estimation of Tapes philippinarum in a Mediterranean coastal lagoon (Sacca di Goro, Italy). Ecological Modelling 193:90–104. Waite, L., Grant, J., and Davidson, J. 2005. Bayscale spatial growth variation of mussels Mytilus edulis in suspended culture, Prince Edward Island, Canada. Marine Ecology Progress Series 297:157–167. Ward, J.E., and Shumway, S.E. 2004. Separating the grain from the chaff: particle selection in suspension- and deposit-feeding bivalves. Journal of Experimental Marine Biology and Ecology 300:83–130. Weise, A.M., Cromey, C.J., Callier, M.D., Archambault, P., Chamberlain, J., and McKindsey, C.W. 2009. Shellsh-DEPOMOD: modelling the biodeposition from suspended shellsh aquaculture and assessing benthic effects. Aquaculture 288:239–253. Wildish, D., and Kristmanson, D.D. 1997. Benthic Suspension Feeders and Flow. Cambridge University Press, New York.

Chapter 7

Bivalve shellfish aquaculture and eutrophication JoAnn M. Burkholder and Sandra E. Shumway

Summary Increased nutrient supplies and associated pollutants from land-based human activities have pervasively degraded coastal ecosystems worldwide, destroying habitats, causing finfish and shellfish disease and death, and promoting harmful algal blooms. While these effects of cultural eutrophication (anthropogenic nutrient overenrichment) from land-based human activities are well known, recent controversy has focused on another source of nutrient enrichment, aquaculture, as potentially a major contributor to eutrophication. Here we evaluate the significance of bivalve shellfish aquaculture in the eutrophication of coastal waters based on the available evidence and,

conversely, the impacts of land-based nutrient pollution and associated pollutants on bivalve aquaculture. Of the 62 ecosystems reviewed here, ∼7% or four ecosystems have sustained system-level adverse impacts from large, intensive bivalve culture operations. The other 93% have sustained either negligible or only localized significant adverse effects contributing to eutrophication from bivalve shellfish aquaculture. Thus, the great majority of ecosystems with bivalve aquaculture studied to date have been described as sustaining minimal or only localized significant eutrophication effects from shellfish farming. Instead, the utility of bivalve aquaculture in effectively reducing phytoplankton and the nutrients available for

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 155

156

Shellfish Aquaculture and the Environment

blooms is being harnessed by some nations for economic benefit to offset nutrient overenrichment from land-based sources. The effects of bivalve culture on the surrounding environment are site-specific and especially depend on the hydrography (flushing and water exchange) and shellfish density. Exceptions to minimal or localized effects have been documented at the ecosystem scale, mostly in poorly flushed lagoons with highdensity shellfish culture. These exceptions underscore the need to consider the ecosystem’s carrying capacity, rather than only the carrying capacity for maximal shellfish production, in bivalve aquaculture over large areas within a given system. Such considerations increasingly are assisted by models such as the Farm Aquaculture Resource Management (FARM) model and the shellfish aquaculture waste model DEPOMOD. In contrast to the localized effects generally reported for bivalve aquaculture, land-based sources of eutrophication have overwhelmed many coastal ecosystems worldwide, and coastal population growth and associated nutrient pollution are continuing to increase rapidly. The acute, obvious effects of urban and agricultural nutrient pollution, often accompanied by loadings of suspended sediments, microbial pathogens, and toxic substances, are fish kills and high-biomass algal blooms. Much more serious chronic impacts, however, include long-term shifts in nutrient supplies, increased “dead zones” of lowoxygen bottom water, loss of critical habitat such as seagrass meadows, stimulation of harmful algal species that are low in food quality, reduction of shellfish recruitment and grazing, and increased shellfish physiological stress, disease, and death. Increasing temperatures from present warming trends in climate change can stress shellfish, and would be expected to interact with pollution to weaken shellfish hosts further and facilitate pathogen attack. Overall, relative to land-based pollution sources, bivalve aquaculture has been found to

contribute little to eutrophication except in some poorly flushed areas with high shellfish density, and aquaculturists should strive to maintain cultures below ecological carrying capacity to prevent such ecosystem-level adverse effects. In contrast to the generally minimal effects of bivalve aquaculture on eutrophication, major, pervasive nutrient pollution from many urban and agricultural sources is seriously affecting shellfish populations and shellfish aquaculture in many coastal waters of the world, and these impacts are expected to increase with rapidly expanding coastal development. Considering that shellfish aquaculture is vital to meet the seafood demands of the rapidly increasing global human population, there is a pressing need for resource managers and policymakers to increase protection of shellfish aquaculture operations from land-based nutrient pollution.

Introduction Cultural eutrophication, the process of overenriching of surface waters with excessive nutrients from human activities, is among the most serious recognized threats to present-day coastal ecosystems (National Research Council [NRC] 2000; GEOHAB, Global Ecology and Oceanography of Harmful Algal Blooms Programme 2006). Increased nutrient supplies from land-based human activities have pervasively degraded coastal ecosystems to the extent that more than two-thirds of coastal rivers and bays in the United States are now moderately to severely degraded from cultural eutrophication, exacerbated by poor flushing and significant human population growth in coastal areas (Bricker et al. 1999, NRC 2000). The excessive nutrients can stimulate blooms of noxious and toxic algae, and increase water column turbidity from the algal overgrowth together with pollutants such as suspended sediments that accompany nutrient loading. The reduced light causes beneficial seagrass

Bivalve shellfish aquaculture and eutrophication

157

Land-based atmospheric P

Sunlight Harvest

5 N, P,

C

Tidal exchange

Phytoplankton

Grazers, bacteria, fungi

N, P, O2 Exchange

Seston

Anaerobic sediment

Bivalves

Nutrients

Sediment

Aerobic sediment

4

1

Detritus

3

2 Biodeposits NH4+

Benthic 1∞ Producers NO3–

NO3–

Nitrification

NO2–

N2

NH4+

Mixing

Land-based N, runoff

PO4–3 DOC

Organic N, P,C Burial

NO2– Denitrification

N2

Figure 7.1 Conceptual diagram of land-based nutrient sources from watershed runoff and atmospheric pollution (nitrogen [N] as inorganic forms [Ni ≡ nitrate, ammonium], particulate N, dissolved organic N [DON]; phosphorus [P]; and carbon, especially considering dissolved organic carbon [DOC]), and bivalve shellfish aquaculture (bivalves) interactions with nutrient supplies in coastal ecosystems as related to (1) removal of seston (suspended particulate material) during filter feeding, (2) biodeposition of feces and pseudofeces, (3) excretion of nutrients (especially ammonia, which is ionized to ammonium; also phosphate and various organic nutrient forms), (4) removal of N, P, and organic carbon in bivalve harvest, and (5) resuspension of nutrients, detritus, and sediments into the water column during some harvesting activities. (Modified from Cranford et al. 2006.) Note that 1° = primary; and that this diagram depicts oxygenated (aerobic) sediments beneath the shellfish cultures, although localized impacts often include anoxic sediments beneath farms in poorly flushed areas.

meadows to die because of depressed photosynthesis (Burkholder et al. 2007 and references therein). In addition, while the excess algae photosynthesize and increase the dissolved oxygen (DO) during the day, at night their respiration can deplete most or all of the available oxygen so that fish suffocate to death (Breitburg 2002). As the algal blooms senesce and die, their decomposition contributes to this oxygen “sag.” Harmful microbial pathogens—viruses, bacteria, fungi, and protozoans—also frequently are added along with nutrient overenrichment (Burkholder et al. 1997, and references therein; Paul and Meyer 2001). As the ecosystem is driven out

of balance, these multiple stressors interact to cause the disease and death of higher trophic level organisms such as wild finfish and shellfish (Wiegner et al. 2003). While the effects of eutrophication from land-based human activities are well known, recent controversy has focused on another source of nutrient overenrichment, aquaculture, as potentially a major contributor to cultural eutrophication (Goldberg and Triplett 1997; Kaiser et al. 1998; Newell 2004; Richard 2004) (Fig. 7.1). Finfish culture requires direct inputs of nutrient-rich feed, whereas shellfish culture relies mostly on naturally occurring phytoplankton and suspended matter, with no

158

Shellfish Aquaculture and the Environment

supplementary food added. Adverse effects of shellfish farming on the water column and benthic environments under and near subtidal mussel farms have been described as comparatively much lower than those around salmon farms (Mirto et al. 2000; La Rosa et al. 2002; Yokoyama 2002; Crawford et al. 2003). How does shellfish aquaculture affect nutrient enrichment of coastal areas? The general perception is that shellfish aquaculture is benign or beneficial because it relies on ambient primary production, can improve water clarity and reduce nutrients and phytoplankton concentrations through shellfish filter feeding, and does not require addition of fish or other food (Folke and Kautsky 1989; Crawford et al. 2003; Shumway et al. 2003). Shellfish filter feeding can depress phytoplankton biomass and alter phytoplankton assemblage structure, and the shellfish also can access carbon from the microbial loop through consumption of heterotrophic and mixotrophic bacteriovores (Lucas et al. 1987; Dupuy et al. 2000). In addition, large macroinvertebrates and benthic fishes sometimes respond positively to shellfish cultures (D’Amours et al. 2008a; see also Chapter 9 in this book). Nevertheless, the effects of shellfish culture on nutrient cycling and food web dynamics have received mixed reports. Increased nutrient supplies from shellfish biodeposits can promote phytoplankton and benthic algal growth, and the increased food supply can enhance shellfish growth (Weiss et al. 2002). In turn, the removal of phytoplankton by filter-feeding shellfish can effect a strong “topdown” control of eutrophication symptoms (sensu Bricker et al. 2007), and the shellfish can also influence water column biogeochemistry (Souchu et al. 2001). Some authors have reported that high densities of molluscs beneficially control eutrophication despite their addition of organic-rich biodeposits as feces and pseudofeces to the bottom sediments, and that mussel culture represents basically a selfregulated aquaculture system (Folke and

Kautsky 1989). Others describe extreme adverse impacts on beneficial naturally occurring macrofauna and plankton (Mattsson and Lindén 1983, da Costa and Nalesso 2006). It has been suggested by some authors that, because of high regeneration of nutrients by shellfish, shellfish cultures may increase eutrophication (Baudinet et al. 1990) by reducing nutrient limitation and stimulating algal growth rates (Prins et al. 1998). Decreased water movement and current velocity caused by the structural features of shellfish cultures (e.g., Nugues et al. 1996) would be expected to exacerbate these effects in the localized area. On the other hand, high densities of filter-feeding shellfish can depress phytoplankton biomass while promoting higher turnover rates (Doering et al. 1986; Sterner 1986; Doering 1989, Asmus and Asmus 1991), and it has been argued that this control on phytoplankton biomass can stabilize the ecosystem (Herman and Scholten 1990) as long as the algal assemblage does not escape control by shifts to species that are inefficiently filtered (Prins and Smaal 1994). This chapter addresses two questions: How significant is bivalve shellfish aquaculture in the eutrophication (nutrient pollution, oxygen deficits) of coastal waters, based on present evidence? Conversely, what are the impacts of land-based nutrient pollution and association pollutants on bivalve aquaculture?

Most commonly reported: localized changes associated with shellfish aquaculture General effects In moderation, nutrient enrichment—regardless of the source—promotes beneficial increases in phytoplankton and benthic algal production and, in turn, higher production of zooplankton, macroinvertebrates, finfish, and shellfish that use the primary (photosynthetic)

Bivalve shellfish aquaculture and eutrophication

production directly or indirectly for food (Reitan et al. 1999; Paterson et al. 2003; Zeldis et al. 2008; Burkholder and Glibert 2011). But when added in excess, nutrient pollution can cause algal overgrowth. Nighttime respiration of the excess algal growth can cause oxygen depletion in bottom waters and sometimes throughout the water column. Decomposition of this excess production by aerobic bacteria and fungi can also lead to oxygen depletion. As more chronic, long-term effects, nutrient overenrichment promotes major shifts in the structure of plant and animal communities, often resulting in high biomass of a few tolerant species and loss of overall biodiversity. Where aerobic surface sediments overlay deeper anaerobic sediments, microbially mediated, coupled nitrification-denitrification can convert organic and inorganic N from animal wastes, detritus, and other sources to nitrogen gas (N2), which can effectively reduce the N available for most primary producers—until the microbial consortium depletes the sediment oxygen content. At that point, the coupled nitrification-denitrification is inhibited, more phosphorus can be released to the water column, and toxic hydrogen sulfide can begin to accumulate (see Newell 2004, and references therein). Shellfish beds take up chlorophyll a, seston, and particulate matter (particulate organic carbon, particulate organic nitrogen, and particulate organic phosphorus—POC, PON, and POP, respectively), and tend to release ammonium, orthophosphate, and silicate (Dame et al. 1991; Prins and Smaal 1994). Filtration and biodeposition of shellfish is considered beneficial to water quality by controlling phytoplankton densities and sequestering nutrients that are removed from the system when shellfish are harvested, buried in the sediments, or lost through denitrification (Kaspar et al. 1985; Newell et al. 2002). Bivalve shellfish enhance benthic/pelagic coupling through filter-feeding of phytoplankton, deposition of feces and pseudofeces to the sediments, and

159

increase of nutrient remineralization rates (Hatcher et al. 1994; Prins and Smaal 1994; Dame 1996). Thus, large densities of bivalves cultured in poorly flushed coastal waters can alter the pelagic-benthic energy fluxes by depleting phytoplankton, zooplankton, and seston in the water column through filter feeding; by increasing sedimentation rates from biodeposition of feces and pseudofeces; and by decreasing oxygen, thereby changing sediment characteristics and benthic community composition (Callier et al. 2008) (Table 7.1). Nutrients such as nitrogen and phosphorus are excreted by shellfish and buried in the sediments; a portion of this nutrient supply is also regenerated from the biodeposits and recycled back to the water column where it can support phytoplankton production (see Newell 2004). For example, bivalve molluscs have been estimated to digest and absorb about 50% of the particulate N that they filter from the water column (Newell and Jordan 1983); much of the absorbed fraction is used for tissue growth, but some is excreted, mostly as ammonium (Bayne and Hawkins 1992). Excretion rates of nutrients to the water column by shellfish can be substantial (e.g., Table 7.2), and shellfish have been reported to play a major role in benthic nutrient regeneration in coastal ecosystems through rapid and efficient recycling of inorganic N and P to primary producers (Magni et al. 2000). Algal production, including growth of certain harmful species, can be stimulated by excreta from some bivalve species. Intensive bivalve cultivation can alter the N:P nutrient stoichiometry and change the major N species to reduced forms, especially ammonia as well as certain organic forms, and these N forms are preferred by various harmful algae (e.g. Berg et al. 1997; Arzul et al. 2001; Glibert et al. 2005). If the biodeposits from the cultured bivalves settle into aerobic sediments overlying anaerobic sediments, coupled nitrification-

Table 7.1 Localized effects related to eutrophication that have been reported from bivalve shellfish culture, with examples of references.

Reduction or depletion of nanophytoplankton, zooplankton, and/or seston

Escaravage et al. (1989); Navarro et al. (1991); Perez Camacho et al. (1991); Newell and Shumway (1993); Dankers and Zuidema (1995); Boyd and Heasman (1998), Heasman et al. (1998); Meeuwig et al. (1998); Pitcher and Calder (1998); Ogilvie et al. (2000); Pilditch et al. (2001); Souchu et al. (2001); Cranford et al. (2003); Dowd (2003); Condon (2005); Strohmeier et al. (2005); Banas et al. (2007) Low risk of reduced food resources for filter feeders (Crawford et al. 2003) Effects not found (Fréchette et al. 1991; Mojica and Nelson 1993; Murdoch and Oliver 1995; Danovaro et al. 2004)

Increased water clarity that has promoted growth of beneficial seagrasses

Deslous-Paoli et al. (1998)

Nutrient replenishment and/or enhanced phytoplankton productivity

Kaspar et al. (1985); Doering et al. (1986); Barranguet et al. (1994); Ball et al. (1997) Songsangjinda et al. (2000)

Increased abundance of cyanobacteria under shellfish cultures

Mirto et al. (2000)

Higher POM under shellfish cultures than in a control site; and/or higher C : N ratios under the rafts indicated accumulation of refractory POM mostly from feces and decomposing mussels

Mojica and Nelson (1993), Nugues et al. (1996), Chivilev and Ivanov (1997), Mirto et al. (2000), Stenton-Dozey et al. (2001), Bendell-Young (2006), Metzger et al. (2007), Lu and Grant (2008)

Higher total N, organic N, dissolved organic C, chlorophyll a, and/or phaeopigment concentrations in surficial sediments from bivalve biodeposition

Ito and Imai (1955); Grenz et al. (1991); Nugues et al. (1996); Mirto et al. (2000); Condon (2005); Giles et al. (2006); Munroe and McKinley (2007) Not found (Mojica and Nelson 1993)

Increased sedimentation from biodeposition of shellfish feces and pseudofeces, increased organic carbon content of sediments, and/or altered sediment geochemistry

Ito and Imai (1955); Mattsson and Lindén (1983); Rosenberg and Loo (1983); Escaravage et al. (1989); Baudinet et al. (1990); Grenz et al. (1991); Perez-Camacho et al. (1991); Hatcher et al. (1994); Grant et al. (1995); Nugues et al. (1996); Spencer et al. (1996, 1998); Songsangjinda et al. (2000); Chamberlain et al. (2001); Jie et al. (2001); Christensen et al. (2003); Crawford et al. (2003); Danovaro et al. (2004); Hartstein and Rowden (2004); Callier et al. (2006); Giles et al. (2006, 2009); Mallet et al. (2006); Callier et al. (2007, 2008); Weise et al. (2009)

Increased deposition of fecal matter and increased oxygen depletion from substantial biomass of fouling organisms such as ascidians on culture gear and shellfish stock

Stenton-Dozey et al. (1999)

Altered redox values and/or higher benthic respiration

Baudinet et al. (1990—respiration; but exceeded oxygen production only in 1 month over the annual study); Stenton-Dozey et al. (2001); Crawford et al. (2003)

Not found (no effect on DO) (Mojica and Nelson 1993)

Mallet et al. (2006)—no significant differences between low-density culture and reference sites

160

Table 7.1 (Continued)

Increased bottom-water turbidity

Crawford et al. (2003)

Increased abundance of resident benthic infauna

Kaiser et al. (1996); Spencer et al. 1996)

Depressed abundance or biomass, lower species richness, and/or mortality of benthic macrofauna

Tenore et al. (1982); Mattsson and Lindén (1983); Jaramillo et al. (1992); Barranguet et al. (1994); Grant et al. (1995—although macrofaunal biomass sometimes was higher under the mussel cultures); Beadman et al. (2004) Not found (Mojica and Nelson 1993; Yokoyama 2002; Danovaro et al. 2004; Whiteley and Bendell-Young 2007)

Reduced macrofaunal biomass, alteration of trophic structure, and/or competition with indigenous species

Sauriau et al. (1989); Stenton-Dozey et al. (2001); Bendell-Young (2006)

Modification of current patterns and circulation

Ottmann and Sornin (1985); Nugues et al. (1996); Boyd and Heasman (1998); Crawford et al. (2003)

Changes in benthic macrofaunal community composition and/or diversity; often, a decrease in dominance of native bivalve molluscs and sea urchins offset by an increase in opportunistic polychaetes

Mattsson and Lindén (1983); Kaspar et al. (1985); Castel et al. (1989); Baudinet et al. (1990); Grant et al. (1995); Spencer et al. (1996, 1998); Mirto et al. (2000); Chamberlain et al. (2001); Beadman et al. (2004); Miron et al. (2005); Bendell-Young (2006); da Costa and Nalesso (2006) Not found (Kaiser et al. 1996; Danovaro et al. 2004)

Higher abundance of benthic predatory shellfish such as crabs, demersal fishes, or starfish which were attracted to cultured shellfish that fell to the bottom sediments beneath cultures, and/or to increased abundance of deposit-feeding prey organisms

Iglesias (1981); Romero et al. (1982); Rosenberg and Loo (1983); López-Jamar et al. (1984); González-Gurriarán (1986); Freire et al. (1990); Grant et al. (1995); Inglis and Gust (2003); Smith and Shackley (2004); D’Amours et al. (2008b)

Higher average diversity in benthic macrofauna under a mussel site than a reference site; higher abundance of individuals at the reference site; overall, no negative effect

da Costa and Nalesso (2006)

Higher diversity of macrofauna per unit area of oyster cage culture areas than in reference areas with or without aquatic vegetation

Dealteris et al. (2004)

Higher bacterial abundance under cultures, and/or higher activities of bacterial exoenzymes under cultures; higher density and biomass of microbial assemblages under cultures

Grenz et al. (1990); Danovaro et al. (2004); La Rosa et al. (2002)

Extensive bacterial mats under cultures

Dahlbäck and Gunnarsson (1981)

Elevated anoxia, anaerobic metabolism (seasonal), and/or higher oxygen consumption

Barranguet et al. (1994); Grant et al. (1995); Chivilev and Ivanov (1997); Chamberlain et al. (2001); Stenton-Dozey et al. (2001); Giles et al. (2006—higher oxygen consumption in summer)

Elevated sediment sulfide concentrations

Ito and Imai (1955); Mattsson and Lindén (1983); Mariojouls and Sornin (1987); Barranguet et al. (1994); Stenton-Dozey et al. (2001); Crawford et al. (2003)

161

162

Shellfish Aquaculture and the Environment

Table 7.1 (Continued)

Accelerated sulfur cycle (higher sulfate reduction and sulfur oxidation rates) under intensive shellfish cultures

Asami et al. (2005)

Localized elevated rates of ammonium release (seasonal)

Kaspar et al. (1985); Baudinet et al. (1990); Barranguet et al. (1994); Hatcher et al. (1994); Grant et al. (1995); Stenton-Dozey et al. (2001); Giles et al. (2006); Nizzoli et al. (2007)

In well-flushed areas, significantly higher nitrate fluxes under cultures

Giles et al. (2006)

Higher rates of N remineralization in surficial sediments under cultures

Grenz et al. (1990); Grenz et al. (1991); Hatcher et al. (1994); Gilbert et al. (1997)—dissimilatory ammonium production (98% of the nitrate in the farming area was reduced to NH4+ and 2% to N2O; thus, most of the Ni remained available in the ecosystem)

Higher denitrification capacity in mussel farm sediments; lower nitrification (oxidative process) in mussel farm sediments

Kaspar et al. (1985); Hatcher et al. (1994); Gilbert et al. (1997)

Higher phosphate and/or silicate fluxes under cultures

Baudinet et al. (1990—P, Si); Magni et al. (2000— P); Souchu et al. (2001—P) Note: not found by Kaspar et al. (1985—P) or by Hatcher et al. (1994—P)

Mixed effects, apparently depending on local current patterns: at one large-scale mussel farm, but not at a second farm, high sedimentation and organic enrichment of the benthos, and reduced diversity of benthic infauna

denitrification also removes nitrogen from the sediments as N2 gas. Thus, if the surface sediments contain some oxygen, biodeposits from shellfish aquaculture promote net ecosystem losses of nitrogen and phosphorus not only by sediment burial but also by microbial nitrification-denitrification. This important process is inhibited, however, if copious biodeposits from high bivalve densities cause anoxia of the surface sediments (Newell 2004). Filtration of particulate matter by bivalves also reduces turbidity so that more light is available for benthic microalgae which take up ammonium, nitrate, and phosphorus, effectively reducing the substrates needed for nitrificationdenitrification but also helping to control regeneration of sediment nutrients to the water column (Newell 2004).

Chamberlain et al. (2001)

These effects are minimized in sites with more rapid flushing and water exchange, so that hydrography is a key factor influencing the environmental effects of shellfish aquaculture (Boyd and Heasman 1998; Crawford et al. 2003; Hartstein and Rowden 2004). For example, comparison of two mussel (Mytilus edulis) farms in sites with high versus low current velocity along the coast of southwestern Ireland revealed significant alterations in benthic community structure at the site with low current velocity, but not at the highervelocity site (Chamberlain et al. 2001). Similarly, open-sea mussel cultures had minimal detrimental effect on benthic fauna along the western Adriatic coast (Fabi et al. 2009). Smaal and Zurburg (1997) found no significant release of nutrients from oysters

Table 7.2 Comparison of nutrient excretion rates by different species of clams, mussels, oysters, and scallops based on some examples from the published literature.

Approach

NH4+

NO3− + NO2−

PO4−3

Temperature (°C) or period

Algoa Bay, South Africa (μg NH4+ ind.−1 h−1; means)

Lab./field

0.31–5.20

nd

nd

15–19

Cockcroft (1990)

Maitland River, South Africa (μmol NH4N g DW−1 h−1)

Field

0.35–8.10

nd

nd

na

Prosch and McLachlan (1984)

Lab./field

2.9

nd

nd

15–19

Cockcroft (1990)

Lab.

0.1

neg

nd

13–15

Henriksen et al. (1983)

Delaware Bay, USA (μmol g DW−1 h−1)

Lab.

0.9–1.5

neg

nd

20

Srna and Baggaley (1976)

Cultured seed clams (from Mook Sea Farms, Inc., Damariscotta, ME) (μg NH4N g clam−1 day−1)

Lab.

20.0–89.4

nd

nd

20

Pfeiffer et al. (1999)

Hatchery, Ireland (μmol g DW−1 h−1)

Lab.

0.16–1

nd

nd

18.8

Xie and Burnell (1995)

Marennes-Oléron, France (μmol g DW−1 h−1)

Lab.

0.5–13

nd

nd

5–25

Goulletquer et al. (1989)

International Shellfish Enterprises, Moss Landing, USA (as Tapes japonica; μg NH4+N g live wt.−1 day−1)

Lab.

27.84–72.24

nd

nd

12, 14, 16, 18

Mann and Glomb (1978)

Seto Inland Sea, Japan (μmol g DW−1 h−1)

Lab.

3.8–10.6

0–12.3

0.7–3.9

19.6–21.6

Magni et al. (2000)

Seto Inland Sea, Japan (mmol m−2 day−1)

Field ext.

1.2–14

0.6–6.8

0.03– 3.6

Annual

Magni et al. (2000)

Virgin Islands, USA (μmol g DW−1 h−1)

Lab.

1.9–4.9

nd

nd

20.1

Langton et al. (1977)

Species, study area, units of nutrient excretion

Source

Clams Donax serra

Donax sordidus Sundays River, South Africa (μmol g DW−1 h−1) Macoma balthica Wadden Sea, Denmark (μmol g WW−1 h−1) Mercenaria mercenaria

Tapes [Ruditapes] philippinarum

163

Table 7.2 (Continued) Species, study area, units of nutrient excretion

Approach

NH4+

NO3− + NO2−

PO4−3

Temperature (°C) or period

Lab.

3.49–16.22

nd

nd

12 (winter)

Navarro (1988)

Great Sippewissett, MA, USA (μmol g−1 DW h−1)

Lab.

2.5

nd

nd

Annual

Jordan and Valiela (1982)

Near Duke University Marine Laboratory, Beaufort, NC, USA (μmol g−1 day−1; as Modiolus demissus)

Lab.

1.66–4.46

nd

nd

∼23

Lum and Hammen (1964)

Narragansett Bay, RI, USA (μmol g DW−1 h−1)

Lab.

0.26

nd

nd

21

Nixon et al. (1976)

Eastern Scheldt, the Netherlands (AFDW)

Field

0–13.9

neg

0.35– 1.7

Jun–Oct

Dame et al. (1991)

Eastern Wadden Sea, Germany (mmol m−2 h−1)

Field

0.32–5.5

neg

0.85

Apr–Jun

Asmus et al. (1990)

Linher River, UK (μmol g DW−1 h−1)

Lab.

4.9–34.6

nd

nd

11–21

Bayne and Scullard (1977)

Lynher River near Plymouth, UK (μg N g DW−1 h−1)

Lab.

∼6–18

nd

nd

8, 12, 15

Livingstone et al. (1979)

Lynher Estuary, southwestern UK (means; μg N g DW−1 h−1)

Lab.

∼8–39

nd

nd

5, 10, 15, 20 (up to 25)

Widdows (1978)

Narragansett Bay, RI, USA (μmol g DW−1 h−1)

Lab.

3.1

nd

nd

15

Nixon et al. (1976)

Sound, Denmark (μmol g DW−1 h−1)

Field

0.14–3.1

nd

0.10– 0.53

0.7–18

Schlüter and Josefsen (1994)

Swansea Bay, UK (μg N g DW−1 h−1)

Field

22.7–29.3

nd

nd

∼Annual

Bayne et al. (1979)

Western Scheldt, the Netherlands (μmol g−1 DW h−1)

Field

1.1

nd

nd

12

Smaal et al. (1997)

Western Wadden Sea, the Netherlands (AFDW)

Field

0–13.9

neg

0.35– 1.7

Jun-Oct

Dame et al. (1991)

Source

Mussels Choromytilus chorus Queule Estuary, southern Chile (μg g DW−1 h−1) Guekensia demissus

Mytilus edulis

164

Table 7.2 (Continued) Species, study area, units of nutrient excretion

Temperature (°C) or period

Approach

NH4+

NO3− + NO2−

PO4−3

Field

1–3.7

neg

0.05– 0.43

Apr–Sep

Prins and Smaal (1994)

Ría de Arousa, Galicia, northwest Spain (μmol g−1 day−1; intertidal and raft culture habitats)

Lab.

1.66–4.46

nd

nd

July

Lum and Hammen (1964)

Ría de Arousa, Galicia, northwest Spain (μg NH4+N g DW−1 h−1; means, intertidal, and raft culture habitats, days 1 and 15)

Lab.

4.92–8.17

nd

nd

14–15 (collected in July)

Labarta et al. (1997)

Seto Inland Sea, Japan (μmol g DW−1 h−1)

Lab.

9.3–16.9

0–1.9

1.3–5.5

19.6–21.6

Magni et al. (2000)

Seto Inland Sea, Japan (mmol m−2 day−1)

Field ext.

0.23–24

0.03–2.7

0.06– 6.5

Annual

Magni et al. (2000)

Tidal creeks, North Inlet Estuary, SC, USA (mg N g oyster−1 h−1)

Field

0.39

nd

nd

July–Aug

Dame and Libes (1993)

Charlestown Pond, RI, USA (μmol g tissue−1 day−1)

Lab.

1.56–2.19

nd

nd

na

Hammen et al. (1966)

Chesapeake Bay, USA (mg N g oyster C−1 day−1)

Modeled

1.43

nd

nd

Summer average

Cerco and Noel (2007)

Delaware Bay, USA (μmol g DW−1 day−1)

Lab.

0.28

nd

nd

20

Snra and Baggaley (1976)

Near Duke University Marine Laboratory, NC (μmol g tissue −1 day−1)

Lab.

0.298–0.978

nd

nd

22.0–25.5

Hammen (1968)

North Inlet Estuary, SC, USA (μmol m−2 h−1)

Field

2825– 15,304

0–0.2

nd

28–30

Dame et al. (1985)

Western Wadden Sea, the Netherlands (g N or P m−2 h−1)

Field

0.04–0.11

nd

0.05– 0.08

June

Dame and Dankers (1988)

Field

0.28–6.7

nd

nd

∼7.5–17

Boucher and BoucherRodoni (1988)

Western Wadden Sea, the Netherlands (mmol m−2 h−1)

Source

Mytilus galloprovincialis

Musculista senhousia

Oysters Crassostrea virginica

Crassostrea gigas Bay of Pempoul (Bay of Morlaix), North Brittany, France (μmol g total WW−1 h−1)

165

166

Shellfish Aquaculture and the Environment

Table 7.2 (Continued) Species, study area, units of nutrient excretion

Temperature (°C) or period

Approach

NH4+

NO3− + NO2−

PO4−3

Sanggou Bay, north China (μmol h−1 g DW−1; NH4+N, PO4−3P)

Field ext.

0.51–5.40

nd

0.11– 0.64

Jan., Jul.

Mao et al. (2006)

Sea Salter Shellfish, Ltd., Whitstable, UK (juveniles; μg NH3N g live wt.−1 day)

Lab.

9–38.8

nd

nd

12, 15, 18, 21

Mann (1979)

Lab.

11.6–21.2

nd

nd

12, 15, 18, 21

Mann (1979)

Anclote Estuary, Tarpon Springs, FL, USA (μg NH3N g DW−1 h−1; means)

Field

72–140

nd

nd

21.5–31.7 (May–Nov)

Barber and Blake (1985)

Homosassa, FL, USA (μg N mg AFDW−1 h−1, as means; larvae and juveniles)

Lab.

0.125–0.384

nd

nd

25

Lu et al. (1999)

Lab.

19.7–41.9

nd

nd

12 (annual mean)

Navarro and Gonzalez (1998)

Lab.

178–147

nd

nd

17, 23

Yang et al. (1999)

Source

Ostrea edulis International Shellfish Enterprises, Moss Landing, CA, USA (juveniles; μg NH3N g live wt.−1 day) Scallops Argopecten irradians concentricus

Argopecten purpuratus Bay of Hueihue, Chloé, Chile (μg NH4+N g DW−1 h−1; means) Chlamys farreri Xujia Maidao, Qingda, China (μg g DW−1 h−1)

Modified from Magni et al. (2000). AFDW, ash-free dry weight; DW, dry weight; field ext., extrapolation of laboratory experiments to a field community situation; lab., laboratory experiments; live wt., live weight; na, not available; neg, negligible excretion found; nd, not determined; WW, wet weight.

(Crassostrea gigas) and mussels (Mytilus edulis) cultured on intertidal tables in Marennes-Oléron Bay along the coast of southwestern France, and reasoned that most of the biodeposits had been flushed away so that mineralization occurred elsewhere.

Localized effects of biodeposits were reported in sheltered sites of Marlborough Sounds, New Zealand, within 30–50 m from aquaculture operations, but at times wave action resuspended and dispersed the biodeposits over wide areas so that there was little overall

Bivalve shellfish aquaculture and eutrophication

effect on the sediments beneath the farms (Hartstein and Stevens 2005). Thus, the effects of shellfish aquaculture on the surrounding environment are site-specific (Chamberlain et al. 2001). Accordingly, the detectable farm footprint (term from Giles et al. 2009) is mostly localized and can be minimal, slight, or severe, depending on an array of factors such as background enrichment, sediment composition and porosity, hydrographic features, depth, bottom topography, water exchange, abundance of bioturbator fauna, and culture density (Pietros and Rice 2003; Newell 2004; Gren et al. 2009). The degree and spatial extent of environmental impacts is related to dispersal of the biodeposits (Chamberlain et al. 2001; Newell 2004), so it has been suggested that benthic effects of shellfish farms in high-energy environments should extend over larger areas (Giles et al. 2009). The substantial dilution in highenergy environments would “lose the signal,” however, so that detectable effects would likely remain localized. The choice of sampling stations and parameters of focus strongly influences interpretations about how shellfish aquaculture affects coastal environments, as shown in an evaluation of two mussel farms versus control sites in Great-Entry (GE; area 2.5 km2, annual harvest 180 tonnes) and Havre-aux-Maisons (HAM; area 1.25 km2, annual harvest 160 tonnes) lagoons in the Magdalen Islands, Quebec, Canada (Callier et al. 2008). These two farms were the same age, and the lagoons where each was sited had similar average depth, current velocity, tidal range, temperature, and salinity. In both farms, mussels were cultivated on suspended longlines over a 2-year growout cycle. Yet contrasting patterns were found: GE, which had previously been described as a more naturally enriched environment (Bourget and Messier 1982), had low benthic species diversity, abundance and biomass, and sediment characteristics. The mussel operation there was assessed as having little apparent localized effect, possibly because

167

it was a more naturally enriched environment (Bourget and Messier 1982). In contrast, at HAM, clear localized effects included an increase in the percent organic matter of the sediment, and a decrease in benthic macrofaunal diversity and abundance under the mussel lines. Other features of the study design, such as the position sampled within the shellfish farm, could also have a major influence on data interpretations; for example, sites within areas of older mussels can differ substantially from sites with cultures of younger animals (Callier et al. 2007). Considering sediment characteristics, Sundbäck et al. (2000) reported that rates of nitrification-denitrification were about 10-fold higher on an annual basis in fine-grained sediments with abundant bioturbator fauna than in sediments with higher porosity and lower bioturbator biomass (see Chapter 10 in this book). Anoxia in even the surface sediments from overenrichment by bivalve deposits in shellfish culture areas can be a cumulative, chronic effect—that is, the longer the shellfish are cultivated in a given location, the more frequent and sustained the anoxic sediment conditions (e.g., Ito and Imai 1955). As a significant consequence, the high hydrogen sulfide concentrations in anoxic sediments can kill nitrifying bacteria, and even if the surface sediments can be reoxygenated, this microbial consortium must be restored before nitrification can resume (Sloth et al. 1995). Some researchers have reported no significant localized impacts from suspended shellfish cultures on the underlying benthic environment. For example, Shaw (1998) found negligible effects of suspended mussel cultures in Prince Edward Island, Canada, based on a benthic sediment survey of organic matter and oxygen levels in 20 estuaries with culture and reference sites. Most published studies, in contrast, have described significant but localized effects from suspended bivalve aquaculture on the underlying area and a limited area surrounding the farms (Tables 7.1 and 7.3).

Table 7.3 Examples of locations and characteristics of bivalve culture systems for which effects have been reported. Location

Culture, wild stocks (information given)

Reference(s)

Mussels (Mytilus galloprovincialis) in intensive suspended culture ongoing for 10 years; culture area 80 ha; annual marketable yield 1880–2720 tonnes (2000–3000 tons) wet weight with shell; 320 ropes per raft

Stenton-Dozey et al. (1999, 2001); also Boyd and Heasman (1998); Heasman et al. (1998); Pitcher and Calder (1998)

Lagoon of Takapoto Atoll, French Polynesia—within a nearly enclosed reef rim; mean depth 25 m; water residence time ∼4 years

“Extensive” pearl oyster (Pinctada margaritifera) culture for 20 years; ∼2 million cultured animals on down lines suspended on subsurface longlines

Niquil et al. (2001)

Mangoku-ura Inlet, Miyagi Prefecture, northern Japan— experimental suspended oyster farms—strong winds often suspended the bottom sediments

Harinohama farm (depth 4 m) and Sawada farm (depth 2.6 m)

Ito and Imai (1955)

Matsushima Bay, northern coast of Katsurashima Island, Japan—mean depth 4 m (at mean tide)

Suspended oyster cultures—40 rafts maintained for 6 years, 150 rafts maintained for 3 years, and 20 racks maintained for 2 years

Ito and Imai (1955)

Nikolskaya Inlet, Kandalaksha Bay, White Sea—semi-enclosed and relatively shallow (mean depth 60 m); water residence time 5–6 years (Cobelo-García et al. 2006)

Two suspended mussel farms (Mytilus edulis), each 15,000 m2, maintained for ∼10 years; depth 26–28 m or 13–16 m

Chivilev and Ivanov (1997)

Seto Inland Sea, southwestern Japan—sheltered waters that had sustained substantial land-based pollution (nutrients, other)

Clam (Tapes philippinarum) and mussel (Musculista senhousia) cultures

Magni et al. (2000)

Xuejiadao intertidal area in Jiaozhou Bay, eastern China—tidal range ∼1.4 m

Manila clam (Tapes philippinarum) farming transects (472 animals m−2; ∼197 g wet weight m−2)

Jie et al. (2001)

Yamada Bay, Sanriku coast, northeastern Japan—semienclosed; area 30 km2

Oysters (∼4,535 tonnes year−1) and scallops—intensive aquaculture

Asami et al. (2005)

Beatrix Bay (sheltered area; area 24 km2; water current 6–12 cm s−1, depth ∼19 m at mid-tide, tidal range up to 4 m)

Longline cultures of greenshell mussels (Perna canaliculus); 45 farms cover ∼8.4% of the total bay area, 50–200 m from shore; total annual harvest ∼3630 tonnes (4000 tons)

Christensen et al. (2003)

Catherine Cove (CC; sheltered area; depth 25–42 m, area 1.8 ha)

Longline mussel cultures (Perna canaliculus) CC, EB—11–12 longlines (length 3.5–5 km); cultures maintained for 15 years; BP—seven longlines; area 1.95 ha; cultures maintained for ∼3 years

Murdoch and Oliver (1995), Ogilvie et al. (2000)

Localized effects Africa (1) Small Bay within Saldanha Bay, South Africa (Benguela system)— current velocity averaged 7.5 cm s−1 between rafts and 1.25 cm s−1 within rafts; depth 12–15 m; water column strongly stratified in summer Asia–Orient (7)

Australia–New Zealand (10)

Elaine Bay (EB; moderately exposed; depth ∼30 m) Blowhole Point (BP; exposed; depth 8–14 m, area 1.15 ha)

168

Table 7.3 (Continued) Location

Culture, wild stocks (information given)

Reference(s)

Elie Bay, Laverique Bay (sheltered; depth 15–25 m)

Longline mussel cultures (Perna canaliculus); two farms in Elie Bay had been in production 1–10 years; two farms in Laverique Bay had been in production 10 years

Inglis and Gust (2003)

Kenepuru Sound, a side-arm of Pelorus Sound—mean depth ∼11 m, 3–4 m tidal fluctuation, maximum current speed ∼4 km h−1

Longline mussel cultures (Perna canaliculus) in the center of a row of 5 similar farms that had been maintained during the past five years

Kaspar et al. (1985)

Port Esperance (PE), St. Helens (SH), and Eaglehawk Bay (EB), Tasmania, Australia

Crawford et al. (2003)

PE—depth 8–12 m

PE—mostly oyster suspended cultures (Crassostrea gigas), also mussels (Mytilus planulatis), maintained since 1984; total culture area 5.6 ha; average annual harvest ∼108 tonnes

SH—depth 7.5–9 m, average flow 3.8 cm s−1 at 2–6 m above the seabed, and ∼18 cm s−1 in the upper water column

SH—oyster and mussel cultures maintained since the mid-1990s, expanded from 3 to 6 ha in 1999; average annual harvest ∼73 tonnes

EB—depth mostly 8–10 m (in some areas, 4 m), average flow 3.4 cm s−1 at 4–8 m above the seabed

EB—mostly oysters with some mussels, maintained since the late 1970s; total culture area 13.8 ha; average annual harvest ∼220 tonnes

Western Firth of Thames, New Zealand—high-energy environment; current speeds from 3.1 to 32.8 cm s−1 during flood tide and from 3.7 to 31.7 cm s−1 during ebb tide

Mussel culture (Perna canaliculus), area 45 ha; maintained since 1994; 145 longlines in the upper 6 m of the water column; each of three blocks of longlines ∼1000 m long × 108–186 m wide; production 1520 tonnes (1680 tons) per harvest at 15-month intervals

Giles et al. (2006)

Arcachon Bay, southwestern France—total bay area 155 km2, mostly intertidal and channels; tidal range from 2.0 to 3.9 m

Oyster parks (Crassostrea gigas) covered 10 km2 of the 40-km2 subtidal area

Castel et al. (1989); Escaravage et al. (1989)

Bangor Pier in the Menai Strait, north Wales

Mussels (Mytilus edulis) cultivated in the specific study area since 2000, by laying directly onto the substratum (“seabed”); large-scale experiment at four densities (2, 3, 5, 7.5 kg m−2) on 400-m2 plots

Beadman et al. (2004)

Bay of Aiguillon and the Marennes-Oleron basin, western France

Oyster tables with bouchots (poles 4–6 m in height, sunk halfway into the sediments) in a tidal zone

Ottmann and Sornin (1985)

Europe (17)

169

Table 7.3 (Continued) Location

Culture, wild stocks (information given)

Reference(s)

Canale, Bay of Mont-Saint-Michel, English Channel, northern France

Oyster tables with bouchots in a tidal zone

Ottmann and Sornin (1985)

Carlingford Lough—small embayment along the east coast of Ireland—shallow (mostly 2–10 m), with extensive areas exposed at low tide; maximum depth 35 m, maximum current speed 0.35– 0.87 m s−1, tidal range 3.7–4.0 m

Intertidal oyster and clam cultures (Crassostrea gigas—400 tonnes; Tapes semidecussata—had not yet been harvested); and wild mussels (Mytilus edulis—1000 tonnes dredged in 1992)

Ball et al. (1997)

Carteau Bay in the Gulf of Fos, Mediterranean Sea—sheltered; mean depth 4.5 m, surface area ∼150 km2

Mussels (Mytilus galloprovincialis)— intensive culture for 5 years (70 tables, each 15 m × 50 m, and each supporting 1000–1500 mussel ropes 3–4 m in length); cultivation area covered 0.4% of total cove area

Grenz (1989); Baudinet et al. (1990); Grenz et al. (1990, 1991); Barranguet et al. (1994); Barranguet (1997)

Cattolica-Rimini, Middle Adriatic Sea—∼2.41–3.22 km from the coast (average depth 11 m)

Large longline mussel culture (species not given); farm area 2 km2

Danovaro et al. (2004)

Coast of southwestern Ireland— low-energy environments (depth 12–15 m, mean current velocity 0.4–1.25 cm s−1)

Two longline mussel farms (Mytilus edulis) that had been in production for 8–14 years; each produced ∼90–135 tonnes (100–150 tons) of mussels year−1

Chamberlain et al. (2001)

Gaeta Gulf, Tyrrhenian Sea, western Mediterranean Sea— microtidal (∼30 cm), sheltered site

Mussel cultures (annual production ∼360 tonnes [400 tons])

La Rosa et al. (2002)

Longline mussel cultures; annual biomass production ∼360 tonnes (400 tons)

Mirto et al. (2000)

Northwestern coast of Sweden— Island of Tjärnö, northwestern coast of Sweden (depth 8–13 m, tide ∼0.3 m, generally weak currents averaging ∼3 cm s−1)

Longline mussel cultures (Mytilus edulis); total area 2800 m2; yielded 100 tonnes in 18 months

Dahlbäck and Gunnarsson (1981)

Skagerrak archipelago northwestern coast of Sweden

Longline mussel culture (Mytilus edulis), 4500 m2 (lines 180 m in length)

Rosenberg and Loo (1983)

Northern archipelago of the Swedish west coast—currents weak (∼3 cm s−1)

Longline mussel cultures

Mattsson and Lindén (1983)

Lysefjorden, southern Norway

Mussel farm (Mytilus edulis), 200 m × 15 m

Strohmeier et al. (2005)

Ría de Arosa (Arousa) within Rías Bajas, Galicia, northwestern Spain—the cultivation area was sheltered by a natural sand bank

Mussels (Mytilus edulis)—more than 90,700 tonnes (100,000 tons) total wet weight year−1 over a 230-km2 area (20 m × 20 m rafts; more than 2000 rafts in more than 30 areas, each with an average of 600 hanging ropes)

Iglesias (1981); Marino et al. (1982); Romero et al. (1982); Tenore et al. (1982); López-Jamar et al. (1984); González-Gurriarán (1986); Pérez Camacho et al. (1991)

Gaeta Gulf, Tyrrhenian Sea— sheltered area, microtidal (∼30 cm), turbid

170

Table 7.3 (Continued) Location

Culture, wild stocks (information given)

Reference(s)

Mussel rafts (Mytilus galloprovincialis)—modeling study (2000 rafts)

Navarro et al. (1991)

Ría de Muros e Noia, Galicia, northwestern Spain—mean depth 25 m, area 90 km2

Longline mussel cultures (∼70 rafts in two areas)

González-Gurriarán (1986)

River Exe Estuary, Devon, UK—sheltered intertidal area; current velocity ∼30 cm s−1, current velocity outside the culture area ∼39 and ∼54 cm s−1 in ebb tide and flood tide, respectively

Oysters (Crassostrea gigas) in intensive but relatively small-scale trestle cultivation; ∼110–120 oysters per bag (individual animal weight ∼25–40 g); depth less than 1 m

Nugues et al. (1996)

Southend-on-Sea, Whitstable, Kent, southeast England—shallow shelving mudflat

Benthic cultures of Manila clams (Tapes philippinarum), harvested by suction dredging; ∼3500 clams m−2

Kaiser et al. (1996)

Western inner Swansea Bay, Wales, UK—shallow, high-tidalenergy environment (maximum depth 20 m)

Mussel cultures (Mytilus edulis) since 1998; maximum abundance ∼600 m−2

Smith and Shackley (2004)

Western Wadden Sea, the Netherlands—tidal range 0.9– 1.9 m; 50% of the total area was exposed at low tide; water mass turned over every 5–15 days

Mussels (Mytilus edulis) cultures on subtidal flats since 1949; total culture area 70 km2 (=5% of the total area); annual production ∼33–136 × 106 kg fresh weight

Dame et al. (1991); Dankers and Zuidema (1995, and references therein)

Baynes Sound, British Columbia, Canada—on the eastern side of Vancouver Island, separated from the Strait of Georgia by Denman Island

Intensive culture of manila clams (Tapes [Venerupis] philippinarum) and oysters (Crassostrea gigas) along 90% of the intertidal beaches; clams were seeded on the beaches at a density of ∼400 animals m−2

Jamieson et al. (2001); Bendell-Young (2006); Munroe and McKinley (2007)

Great-Entry Lagoon (GE) (Havreaux-Maisons Lagoon [HAM]), Magdalen Islands, southern Quebec, Canada—average tidal range ∼0.6 m; typically weak currents (5–18 cm s−1); strong winds effected a well-mixed water column; depth in culture areas ∼6 m

Suspended mussel farms (Mytilus edulis) in operation since the 1980s; at GE, cultures cover 2.5 km2 (400 91-m longlines) and yields ∼180 tonnes of mussels year−1; at HAM, cultures cover 1.25 km2 (200 76-m longlines)

Callier et al. (2006, 2007, 2008)

Great-Entry Lagoon (GE) (HouseHarbour Lagoon [HH]) in the Magdalen Islands; and Cascapedia Bay (CAS), southern Quebec, Canada (generally more exposed and energetic); mean tidal range 1.9 m; typical current velocity ∼10 cm s−1 (note: HAM, above, is synonymous with HH)

Suspended mussel farms, as above for GE and HH; at the CAS site, the 1.4-km2 mussel culture area was ∼3.5 km offshore (depth 20 m) and consisted of 150 backlines (each 142 m long and supporting 1100 m of mussel line seeded at a density of 820 mussels m−2)

Weise et al. (2009)

Sheltered sites in the inner estuary of Ría de Arosa

North America (21)

171

Table 7.3 (Continued) Location

Culture, wild stocks (information given)

Reference(s)

Indian River Lagoon, FL—shallow, nontidal, wind-mixed lagoon; mean depth 1.5 m

Hard clam (Mercenaria mercenaria) aquaculture (benthic)

Mojica and Nelson (1993)

Malpeque Bay, Cascumpec Bay, and New London Bay, northern Prince Edward Island, Canada— sheltered areas, depth 6–15 m; mean height of high tide 0.9 m, mean height of low tide 0.2 m

Longline mussel cultures (Mytilus edulis); four farms, each consisting of a 100-m horizontal surface rope from which mussel socks were suspended at depths from 3–10 m

D’Amours et al. (2008a)

Point Judith Pond, RI, USA—depth 2.4–3.0 m

Oyster cultures (Crassostrea virginica)—culture area 1 ha; more than 600 cages, each 0.6 m × 0.6 m and each holding 12 mesh bags of shellfish

Dealteris et al. (2004)

Sites along the coasts of Prince Edward Island, Canada

Longline mussel cultures (Mytilus edulis)

Meeuwig et al. (1998)

Brudenell River: mean depth 14.5 m, water residence time 44 days

2.03 × 107 g dry weight standing stock

Broughton’s Creek: mean depth 18.0 m, water residence time 332 days

7.29 × 107 g dry weight standing stock

Cardigan Bay: mean depth 17.6 m, water residence time 255 days

6.75 × 107 g dry weight standing stock

Murray River: mean depth 10.0 m, water residence time 356 days

9.63 × 107 g dry weight standing stock

Rustico Bay: mean depth 7.1 m, water residence time 83 days

2.7 × 107 g dry weight standing stock

St. Peters Bay: mean depth 10.3 m, water residence time 61 days

6.39 × 107 g dry weight standing stock

St.-Simon Bay, New Brunswick, Canada—a shallow open bay with excellent water exchange; mean depth 0.6 m at low tide

Eastern oyster cultures (Crassostrea virginica; maximum density equivalent to 4000 oyster bags ha−1; final oyster biomass 8 kg m−2); sampling area was a 35-ha oyster lease used for bottom culture (1982–1997), then for floating bag and oyster table culture

Mallet et al. (2006); Lu and Grant (2008)

Tracadie Bay, northern Prince Edward Island, Canada—sheltered areas; mean tidal range ∼1 m

Modeling effort in an area of intensive longline mussel culture (Mytilus edulis)

Dowd (2003)

172

Table 7.3 (Continued) Location

Culture, wild stocks (information given)

Reference(s)

Tracadie Bay (TB) and Winter Bay (WB), northern shore of Prince Edward Island, Canada, facing the Gulf of St. Lawrence—shallow, enclosed tidal lagoon; mean depth 3 m, maximum depth 6 m

Longline mussel cultures (Mytilus edulis)—∼10 leases in WB; more than 40 plots in TB; overall, spat and market-sized mussel mussel production (year, 2000) was more than 210 tonnes (WB) and 4600 tonnes (TB)

Miron et al. (2005)

Upper South Cove, Lunenburg, Nova Scotia, Canada (Corkum’s Island Mussel Farm)—mean depth 1–2 m, maximum depth 10 m; protected area, with maximum current speeds 15 cm s−1 at 1 m above the seabed

Longline mussel culture (Mytilus edulis, also Mytilus trossulus) in the upper 3 m of water column; culture area ∼4000 m2; ∼400 mussels m−2

Hatcher et al. (1994); Grant et al. (1995)

Whitehaven Harbour, northeastern Nova Scotia, Canada—mean tidal range 1.4 m; experimental site 0.5 km southeast of Munroe Point (mean depth 19 m)

Suspended cultures of sea scallops (Placopecten magellanicus)— cultivation area 80 m × 50 m; 10 longlines rigged with vertically hanging ropes spaced 1 m apart; average scallop density 12.2 ind. m−3

Pilditch et al. (2001)

Willapa Bay, WA—shallow, coastal-plain, upwellinginfluenced estuary; more than 50% of the surface area and volume were in the intertidal zone; tidal range 3.5 m; half of the bay volume entered and left with every tide, and 30% of the intertidal volume was replaced with new water on every tide; yet in the landward reach of the estuary, the average water age was 3–5 weeks

Intensive culture of Pacific oysters (Crassostrea gigas)

Banas et al. (2007)

Coqueiro’s Beach, Anchieta, southeastern Brazil—average depth 3 m; tidal currents up to 25 cm s−1 in winter and 35 cm s−1 in summer

Longline mussel cultures (Perna perna), maintained since 1998 (100 lines; estimated annual production 24 tonnes)

da Costa and Nalesso (2006)

Queule River Estuary, southern Chile—central portion, depth ∼4 m

Mussel cultures (Choromytilus chorus, Mytilus chilensis, up to 250–300 adults m−2)

Jaramillo et al. (1992)

Hard clam (Mercenaria mercenaria) benthic cultures for ∼25 years (4.5 × 107 adults, at or near carrying capacity); annual harvest 2 × 107 clams (average shell length 60 mm)

Kuo et al. (1998); Luckenbach and Wang (2004a,b); Condon (2005)

South America (2)

Ecosystem-scale effects (4) Cherrystone Inlet, eastern Chesapeake Bay, USA—small, shallow embayment (6 km2, mean depth 1 m; volume 1.54 × 107 m3 at high tide; average volume [time-averaged tidal prism] 5.8 × 106 m3)

173

174

Shellfish Aquaculture and the Environment

Table 7.3 (Continued) Location

Culture, wild stocks (information given)

Reference(s)

Marennes-Oléron Bay, western Atlantic coast of France—shallow macrotidal bay with high turbidity from suspended sediments; area ∼136 km2 (Dame (1996); mean depth ∼5 m; water residence time 5–10 days; bivalve populations in the bay could filter the water volume in ∼2.7 days (Smaal and Prins 1993)

Mudflats (700–800 m in width) with mussel culture structures (bouchots) and oyster racks (Mytilus edulis, Crassostrea gigas), operated since 1850; shellfish cultures covered ∼60% of the bay area (∼60 km2 mussel cultures, 32 km2 oyster cultures) and had added high levels of biodeposits; on at least two occasions, this system had been overstocked with shellfish and overexploited; standing stock of Crassostrea gigas was ∼100,000 tonnes fresh weight; annual production of Crassostrea gigas was ∼30,000 tonnes fresh weight

Sornin et al. (1983); Sornin et al. (1986); Sauriau et al. (1989); Héral (1993); Raillard and Ménesguen (1994); Smaal and Zurburg (1997); Bacher et al. (1998); Dame and Prins (1998); Gouleau et al. (2000)

Sacca di Goro Lagoon, Northern Adriatic Sea, Italy—had two 900-m openings; mean depth 1.5 m area 26 km2; water residence time up to 25 days; freshwater flows were highly managed with dredging

Intensive suspended clam (Tapes philippinarum) and mussel (Mytilus galloprovincialis) culture; as of 2004, approximately one-third (∼8 km2) of the total lagoon area was licensed for clam culture and 0.4 km2 was licensed for mussel culture; annual harvest ∼13,600 tonnes (15,000 tons) of clams and 900 tonnes (1000 tons) of mussels

Mazouni et al. (1996); Viaroli et al. (1996); Mazouni et al. (1998); Bartoli et al. (2001); Mazouni et al. (2001); Viaroli et al. (2003); Mazouni (2004); Nizzoli et al. (2005, 2006a, 2006b, 2007)

Thau Lagoon, France—had two small openings; mean depth 4.5 m; had excessive nutrient inputs from the watershed, very low water renewal (water residence time ∼3–5 months; limited rainfall, warm temperatures; wind-mixed and microtidal with bottom-water anoxia in summer)

Intensive suspended oyster (Crassostrea gigas—80%) and mussel (Mytilus galloprovincialis—20%) culture covered ∼20% (∼15 km2) of the total lagoon area (40 oysters m−2; total standing stock 22,670 tonnes [25,000 tons]; annual harvest ∼13,600 tonnes [15,000 tons])

Deslous-Paoli et al. (1993, 1998); Mazouni et al. (1996, 1998, 2001); De Casabianca et al. (1997); Gilbert et al. (1997); Souchu et al. (2001); Mazouni (2004); Mesnage et al. (2007); Metzger et al. (2007)

Note that the number in parentheses after each region indicates different ecosystems studied. Note that some studies that are described in the text are not included here (e.g., Prins and Smaal 1994) because specifics about the aquaculture were not provided.

Typical findings are illustrated by Baudinet et al.’s (1990) or Grant et al.’s (1995) comparisons of an area beneath a suspended mussel culture (Mytilus galloprovincialis) versus a reference site. Working in a well-mixed area of the Gulf of Fos, France, Baudinet et al. (1990) examined upward fluxes of nitrate, nitrite, ammonia, silicate, and phosphate as well as oxygen at the sediment–water interface. Both

the culture area and the reference site had organic-rich sediments, especially the culture area because of high deposition of fecal matter. Transformation of biodeposited organic matter seasonally increased phosphate, silicate, and ammonia fluxes, whereas there were negligible or minor differences in nitrate/nitrite fluxes and the oxygen production/consumption equilibrium. The ratio of Si : P fluxes (as

Bivalve shellfish aquaculture and eutrophication

Si(OH)4 : PO4−3) ranged from 6 to 15, much lower than the ratio of the concentrations of these nutrients in the water column (18–70), suggesting that the flux values mainly reflected decomposition of recently sedimented phytoplankton. Benthic respiration and consumption of oxygen exceeded oxygen production under the mussel cultures only in 1 of 5 months when surveys were conducted. Reference site macrofauna (adult invertebrates larger than 0.5 mm) were poorly diversified (Shannon Index between 2 and 3) and consisted mostly of polychaete detritivores (40% Cirratulidae, 30% Capitellidae—mainly Mediomastus sp.), with maximal densities at 50,000 m−2. The redox break line (aerobic/anaerobic boundary) was fairly deep and occurred at ca. 10 mm. Under the mussel cultures, there was yet lower diversity of benthic macrofauna (Shannon Index between 1.5 and 2). Nearly all of the fauna were polychaetes that are considered to be indicative of high organic pollution— for example, 10,000 Capitella capitata m−2, and 60,000 Ophryotrocha sp. m−2, small species which exist in the surficial sediments and consume biodeposits. The redox break line was much shallower, only ca. 2 mm from the sediment surface. The authors’ overall interpretation from their data was that biodeposits from high-density mussel cultures cause localized rather than ecosystem-level effects. Grant et al. (1995) compared the area under a suspended mussel culture (Mytilus edulis, Mytilus trossulus) versus a reference site of similar sediment texture in a small Nova Scotia cove (7-m depth): Sedimentation rates of mussel feces and pseudofeces were higher under the mussel culture lines; there was a shift toward more anaerobic metabolism at the mussel site; and maximum rates of ammonium release at the mussel site were twofold higher than maximal rates at the reference site. Benthic macrofauna also differed in abundance and community structure at the two sites; the mussel site was dominated by mol-

175

luscs Ilyanassa spp. and Nuca tenusucata, which apparently were attracted to mussels that were lost from the culture, and to enriched organic matter in the biodeposits. The authors’ overall assessment was that there were minimal adverse impacts on the surrounding area from bivalve aquaculture. Severe localized negative effects of bivalve shellfish farming are infrequently reported. For example, Dahlbäck and Gunnarsson (1981) reported severe localized effects from intensive blue mussel farming in a poorly flushed area along the northern Swedish coast (water depth 8–13 m, tide ca. 0.3 m, weak currents at ∼3 cm s−1). The sedimentation rate (3 g C m−2 day−1) under the culture area was about threefold higher than at a nearby reference site. There was accumulation of sediment rich in organic matter and sulfide: at 15°C, 30.5 mmol sulfate m−2 day−1 in the upper 10 cm of sediment under mussel cultures, and much higher sulfide seasonally. As a second example, Stenton-Dozey et al. (2001) measured significantly higher particulate organic matter (POM) and sediment anoxia under mussel (Mytilus galloprovincialis) rafts than in a control reference site in a South African bay (fine to medium sand sediments, average current velocity 1.25 cm s−1 within rafts and 7.5 cm s−1 between rafts). The refractory POM from mussel feces, decomposing dead mussels, and biofouling organisms caused elevated C : N ratios. Total reducible sulfides also increased threefold downcore in sediments under mussel rafts. Highest and most variable rates of ammonium efflux occurred under the mussel cultures (825 ± 500 mmol NH4+ m−2 h−1) as well. Other localized impacts can include depressed biomass and altered community structure of benthic macrofauna (Table 7.1). Localized effects can persist for some time after farming is discontinued, especially in poorly flushed areas. Martin et al. (1991) reported that biodeposits accumulated in sandy sediments beneath oyster tables, but were no longer detected 2 months after the

176

Shellfish Aquaculture and the Environment

(A)

(B)

(C)

Figure 7.2 (A, B) Geoduck farming operations (nogeoduckfarm.com/_wsn/page2.html and www.protectourshoreline.org); (C) geoduck (www.pac.dfo-mpo.gc.ca/…/geopath/intro-eng.htm).

cultures were removed. On the other hand, Mattsson and Lindén (1983) found that the benthic fauna at a site along the Swedish west coast was dominated in number by the clam Nucula nitidosa, and in biomass by the heart urchin Echinocardium cordatum and brittle stars Ophiura spp. prior to initiation of mussel farming. After 6–15 months of mussel aquaculture, these species were replaced by opportunistic polychaetes (Capitella capitata, Scolelepis fuliginosa, Microphthalmus sczelkowii), localized within a zone under and 5–20 m around the cultures. After the mussels were harvested, only limited recovery was observed in the localized affected area after 1.5 years. In Saldanha Bay, South Africa, StentonDozey et al. (1999, 2001) documented accumulation of refractory organic matter under mussel (Mytilus galloprovincialis) rafts and adverse effects on benthic macrofauna, with only marginal recovery 4 years after the cultures were removed.

Special mention is merited of a major controversy in Puget Sound of the northwestern United States concerning the environmental impacts of the Pacific geoduck (Panopea abrupta; Fig. 7.2). Geoducks are the largest known burrowing clams; they range from 0.5 to 1.5 kg at maturity, but can weigh up to 7.5 kg with a siphon length of more than 1 m (Hilborn et al. 2004). Market size in Washington State is just under 1 kg (Hilborn et al. 2004). Geoduck aquaculturists use polyvinyl chloride (PVC) pipes (length ca. 360 mm, diameter ca. 152 mm) pushed into intertidal sediments as predator exclusion devices or “nursery tubes” to protect the seed stock, which are set in high densities (ca. 20,000 to 43,500 PVC pipes per acre, or ∼8100 to ∼17,600 pipes per hectare; W. Dewey, Taylor Shellfish Farms, Inc., Shelton, WA, pers. comm.). Considering their size and the typical density of culture, geoducks could be significant contributors to nutrient supplies in

Bivalve shellfish aquaculture and eutrophication

farm areas, as evidenced by the abundant macroalgae that commonly thrive in the localized areas (Fig. 7.2). However, the environmental effects of geoduck aquaculture remain to be determined—a white paper commissioned by the state of Washington to determine environmental impacts of geoduck aquaculture reported that there was no peer-reviewed research on eutrophication effects as of late 2007 (Straus et al. 2008), and information is still not available. The available research about the effects of shellfish aquaculture on eutrophication mostly has focused on oysters or mussels that are suspended in the water column or attached to supports. Influences of cultured shellfish such as clams, which are grown directly within the bottom sediments, are logistically more difficult to assess, but similarly have been reported to include displacement of the natural benthic macrofauna and loss or replacement of their associated functions (e.g., sediment mixing or oxygen transport into anoxic sediments), increased oxygen consumption, excretion of nitrogenous organic wastes, enhanced sedimentation of suspended particulate matter, and concentration of biodeposits in the farm areas that promotes sulfide accumulation in the surficial sediments (Sorokin et al. 1999; Bartoli et al. 2001). Clam harvesting techniques such as dredging alter sediment stratification and can increase nutrient fluxes to the overlying water (Kaiser et al. 1998; Bartoli et al. 2001; see also Chapter 11 of this book). Despite major emphasis on assessing the role of suspended mussel farming in eutrophication, the approach of most studies has been described as consisting mostly of “intuitive considerations about the relationships between mollusc culture and eutrophication processes: the collection and removal from the system of organisms that feed on microalgae and particulate matter should result in a net reduction of nutrient loads.… This consideration is based on the assumption that, contrary to fish farming systems in which food (and so, N and

177

P) [is] continuously supplied from outside, molluscs meet their food requirements in situ” (Bartoli et al. 2001). Although there are localized increases in nutrient supplies in shellfish aquaculture sites, the total amount of nutrients regenerated from bivalve biodeposits is comparable with the nutrients that would be released from other means of phytoplankton decomposition. Thus, whereas finfish aquaculture is driven by the addition of large quantities of nutrients within fish food, in shellfish aquaculture sites, the maximum phytoplankton biomass supported by nutrients that are regenerated from shellfish biodeposits cannot exceed the level that would be sustained by ambient nutrients (see Newell et al. 2005).

Localized effects on nutrient dynamics The most commonly reported influence of bivalve shellfish aquaculture on nutrient supplies is high rates of ammonium flux (direct excretion plus regeneration from biodeposits in the sediments). Maximal rates of 5 mmol N m−2 h−1 have been reported, especially during warmer seasons (see review in Newell 2004). Sources of the nitrogen released include not only N-rich phytoplankton but also bacteria and heterotrophic flagellates (Asmus and Asmus 1991; Kreeger and Newell 2001). Considering these high levels of ammonium regeneration, a common interpretation has been that bivalve populations rapidly recycle nutrients and thus enhance phytoplankton production. For example, Magni et al. (2000) assessed the effects of organic loads from biodeposits of a mussel farm (Tapes philippinarum) in the Tyrrhenian Sea, a poorly flushed area along the western Mediterranean. Their analysis indicated that direct excretion of N and P by the mussels accounted for up to 90% of benthic nutrient fluxes in this system over an

178

Shellfish Aquaculture and the Environment

annual cycle. In other work, a small suspended mussel culture operation (Perna canaliculus, 45 ha) in a well-flushed area of the Firth of Thames, New Zealand, was estimated to contribute substantially to the inorganic N supplies needed by primary producers in the localized area (Giles et al. 2006) (Table 7.1). At a reference site, sediment dissolved inorganic N (DIN) release was calculated to supply ∼74% of the N requirements of the primary producers, indicating that benthic nutrient regeneration was important in supporting primary production for this system (Zeldis 2005; Giles et al. 2006). Under the mussel farm, in contrast, DIN release would have met an estimated 94% of the N requirements for primary producers. In addition to nutrient recycling, burial of N and P with accumulating sediments (Kaspar et al. 1985; Hatcher et al.

1994), and removal of N from the ecosystem by denitrification under suitable conditions, can be enhanced by bivalve biodeposition (Newell et al. 2002; Newell et al. 2005). Beyond a commonly reported increase in ammonia flux and organic carbon concentrations beneath suspended bivalve shellfish aquaculture sites, and estimates of effects on primary producers in the culture area, few attempts have been made to evaluate localized effects of bivalve aquaculture on nutrient pools and transformations. Among the most detailed studies is that of Kaspar et al. (1985), who attempted to assess the nitrogen cycle seasonally over an annual cycle at a mussel (Perna canaliculus) farm versus a nearby reference site in Kenepuru Sound, New Zealand (Table 7.4). Nitrate + nitrite pools were similar at the two sites, but sediment ammonium was

Table 7.4 Comparison of nitrogen dynamics at a mussel culture site versus a nearby reference site in Kenepuru Sound, New Zealand, based on seasonal measurements over an annual cycle (from Kaspar et al. 1985). Parameter

Reference site

Mussel culture site

Organic N (top 12 cm of sediment)

7.4–10.8 mol m−2

6.1–8.9 mol m−2

Nitrate + nitrite pools

- - - - - - Similar at both sites - - - - - -

Sediment ammonium (surface; 12 cm)

86 nm cm−3; 112 nm cm−3

418 nm cm−3; 149 nm cm−3

Molar C : N ratio of sediment organic matter

7.9–10.0

6.2–7.2

Molar N : P ratio of sediment organic matter

3.3–6.1

4.3–7.2 −2

−1

21.7–37.1 mmol m−2 day−1

Total N mineralization rate (top 12 cm of sediment)

8.5–25.0 mmol m day

Sediment denitrification

0.1–0.9 mmol m−2 day−1

0.7–6.1 mmol m−2 day−1

Percentage of reduced nitrate that was denitrified (sediment)

93%

76%

Ammonium excretion by mussels

—

4.7% (summer) and 7.4% (winter) of the combined N mineralization by mussels + sediment

Chlorophyll a (surface sediments)

∼75 mg m−2

∼75 mg m−2

Phaeophytin levels (surface sediments)

52 mg m−2

137 mg m−2

Phytoplankton productivity (comparable except in winter [shown]—14C-CO2 fixation)

232.5 mmol m−2 day−1

1183 mmol m−2 day−1

Bivalve shellfish aquaculture and eutrophication

up to fivefold higher in the mussel farm site; in addition, total denitrification was 21% higher, and total loss of N through mussel harvest and denitrification was 68% higher. In Upper South Cove, Nova Scotia, Hatcher et al. (1994) examined the effects of enhanced sedimentation under suspended mussel cultures (Mytilus edulis, Mytilus trossulus) on the respiration and nutrient fluxes of the benthic communities and benthic/pelagic coupling over an annual cycle. The reference site sediments were a net sink for total dissolved N, whereas the sediments under the mussel lines were a source. A significant relationship between water-column chlorophyll and sedimentation rates was enhanced under the mussel lines, and ammonium release was significantly higher under the mussel cultures throughout the year. However, most of the POM trapped in sediment traps at both the mussel farm site and the reference site did not contribute significantly to C or N flux or longterm burial in the underlying sediments. The authors suggested that most of the POM likely had been transported from both sites as tidally driven horizontal flux.

Interpretations from an ecosystem approach Logically, the potential for ecosystem-scale effects from bivalve aquaculture on nutrient supplies, phytoplankton blooms, oxygen deficits, and other factors associated with eutrophication would be expected to increase in quiet, poorly flushed lagoons and embayments as the relative proportion of total area covered by the cultures increased. Although an ecosystem approach is needed to evaluate the effects of shellfish aquaculture on the surrounding ecosystem, few such studies exist in the published literature, likely because of the massive effort and cost involved in adequate assessment. Complex nutrient loading patterns and

179

fates often characterize lagoons and embayments where most shellfish aquaculture occurs. Concentrations are relatively easy to measure, but mass water transport, needed to estimate nutrient loadings (Burkholder et al. 2004, 2006), is more difficult to assess accurately. In addition, an array of biogeochemical processes (assimilation, sedimentation, adsorption, denitrification, chemical reduction, bioturbation)—each a challenge to assess at the ecosystem level—influence nutrient transport and fate. While the net removal of nutrients from the system with shellfish harvest is relatively easy to quantify, the biogeochemical processes involved in nutrient assimilation, regeneration, and mobilization rates associated with the metabolic activities of the shellfish are more difficult to assess (Bartoli et al. 2001). Several attempts to evaluate the role of shellfish in eutrophication at the ecosystem scale are reviewed here. Thus far, as would be expected, the data indicate that the effects of bivalve shellfish aquaculture on coastal ecosystems depend on biogeochemical processes as well as the characteristics of the shellfish species being cultured (e.g., the species, size, age, health, density of animals per unit area, nutrient assimilation efficiency—Bayne et al. 1976), the areal extent of the cultures in comparison with the total system area, the water residence time and mixing characteristics, the balance between effective nutrient removal (via particle filtering) from the water column versus nutrient additions from sediment biodeposits, and the amount and types of other potential nutrient sources. Rather than attempting to rigorously assess each of the many variables—which would be costprohibitive for most scientists—the relatively few ecosystem-scale assessments have used a more simplistic or “first cut” approach. It is generally accepted that natural populations of bivalve filter-feeders can be important in nitrogen cycling in coastal ecosystems, but

180

Shellfish Aquaculture and the Environment

Table 7.5 Impact of mussel farming area (0.0525 km2) fluxes on Carteau Cove (total area, 13 km2).

Nutrient input by sediments in the site outside the mussel farm (mol h−1) Nutrient input by sediments under the mussel farm (mol h−1) Influence of mussel cultivation zone on Carteau Cove

NH4+

PO4−3

Si(OH)4

612.3 14.0 2.3%

169.0 3.0 1.8%

2012 28.7 1.4%

From Baudinet et al. (1990). Average input values to the water column were estimated from fluxes of three surveys.

their effects on P cycling and on N and P stoichiometry appear to be more variable (Nizzoli et al. 2006a, 2006b). Some authors have not found a significant effect of bivalve shellfish on P recycling (Kaspar et al. 1985; Dame et al. 1991; Hatcher et al. 1994; Condon 2005); others have reported a preferential recycling of N over P in high-density oyster reefs (Sornin et al. 1986) and mussel beds (Prins and Smaal 1994), or a balanced release of N and P (based on measurements of single Tapes animals; Magni et al. 2000). A decrease in the N : P ratio of seston with increasing mussel biomasss was documented by Prins et al. (1995) but, in general, sediment oxygenation and iron, sulfur, and calcium cycling are considered to control P net efflux from sediments more strongly than bivalve shellfish (Krom and Berner 1981; Hatcher et al. 1994; Nizzoli et al. 2006a). Effects of natural and cultured populations on nitrogen cycling are much clearer, especially regarding ammonium regeneration.

Negligible effects of many bivalve farms Although regeneration of nutrients from the sediments to the water column is commonly reported to be stimulated by shellfish aquaculture (Doering et al. 1987; Hatcher et al. 1994; Chapelle et al. 2000), ecosystem-scale effects have been evaluated as negligible for many bivalve culture operations because the culture area is small in comparison with the total eco-

system area, and/or the culture is sited in moderately flushed to well-flushed conditions. For example, Baudinet et al. (1990) used localized flux data collected at a reference site versus beneath suspended mussel cultures (Mytilus galloprovincialis; n = 3 hemispheric bell jars, 17 L in volume, per site) to estimate the effect of mussel farming on Carteau Cove, a small embayment in southern France along the Mediterranean Sea. The bivalve farm covered only about 0.4% of the total 13 km2 area of the cove, and consisted of 70 rope hanging structures that had been in place for 3 years; each 15 × 50 m table supported 1000–1500 mussel ropes that were 3–4 m in length. The area was somewhat sheltered by a natural sandbank. The flux data were collected on five dates during spring, summer, and/or fall of 2 years. There were significant localized effects of the mussel farms on nutrient fluxes and benthic infauna, but on an areal basis, the influence of the mussel cultivation area on all of the cove ranged from 1.4% increase (in Si(OH)4) to 2.3% increase (in NH4+) (Table 7.5). As another example, Niquil et al. (2001) estimated the effects of farmed pearl oysters (Pinctada margaritifera) and associated bivalves (pen shells Pinctada maculata) on the lagoon of Takapoto Atoll (Takapoto, French Polynesia), which has a relatively long water residence time of 4 years. Pearl oyster farming was described as extensively developed in the lagoon for 20 years, including stock of ∼2 million P. margaritifera and ∼10 million P. maculata on the culture systems, but the

Bivalve shellfish aquaculture and eutrophication

Significant effects of intensive bivalve culture on nutrient inputs, especially in poorly flushed areas In contrast to the above findings are data for systems with bivalve cultures that are at or near exploitation carrying capacity, especially in poorly flushed systems. Many studies from such areas have focused on naturally occurring shellfish beds or individual cultured animals and used the data to make inferences about the importance of bivalve shellfish beds to the total system. For example, in the Seto Inland Sea, Japan, Magni et al. (2000) extrapolated data from individual incubations of Manila (short-necked) clams (Tapes philippinarum) and the Japan (Asian) mussel (Musculista senhousia) to estimate that benthic nutrient fluxes were 10-fold higher from shellfish areas than diffusive fluxes modeled from sediment profiles. In this poorly flushed system, the authors also suggested that cultured shellfish caused elevated rates of ammonium production and oxygen consumption at the ecosystem scale. As another example, over a 2-year period, Prins and Smaal (1994) used a benthic ecosystem tunnel device to examine fluxes of particulate materials and dissolved substances between bivalve beds and the water column in the Oosterschelde Estuary along the coast of the Netherlands. The authors compared rates of N and P regeneration from mussels, predominantly consisting of blue mussel cultures (5.3 g C m−2), with estimates of the total nitrogen (TN) mineralization (pelagic + benthic) as calculated from a model (Simulation Model

5

Mussels Total

4 N mineralization in ton day−1

animals on the culture systems were only a small percentage of the total bivalve shellfish populations present. While Niquil et al. (2001) did not estimate the effects of the shellfish on the culture systems on nutrient supplies, their model indicated that those shellfish consumed only 0.24% of the planktonic gross primary production in the Takapoto Atoll lagoon.

181

3

2

1

0 Apr June Sept 1988

Apr June Sept 1989

Figure 7.3 The amount of N mineralized by mussel beds in the central part of the Oosterschelde Estuary, compared with estimates of TN mineralization (benthic + pelagic) from the Oosterschelde ecosystem model SMOES. The bar represents the 10–90% range of model estimates, and the line shows the median (Prins and Smaal 1994).

Oosterschelde EcoSystem; SMOES). Excretion by the mussels was estimated to contribute 31–85% of the total phosphate flux from the mussel bed and, on average, only about twothirds of the particulate organic P taken in by the mussels was recycled as phosphate. Estimated ammonium regeneration by the mussels was about 50% of the median value of the model result for total N mineralization. Ammonium excretion by the mussels accounted for 17–94% of the total ammonium flux from the shellfish bed (Fig. 7.3), supporting the authors’ hypothesis that the mussel population played a major role in N recycling in the central part of the Oosterschelde ecosystem. The case of shallow but fairly well-flushed systems with intensive bivalve cultures at

182

Shellfish Aquaculture and the Environment

Figure 7.4 Hard clam (Mercenaria mercenaria) cultures in Cherrystone Inlet, tributary to Chesapeake Bay along the eastern shore of Virginia, USA. Each rectangular net (∼4 m × 18 m) covers ∼50,000 clams (Luckenbach and Wang 2004b).

or near exploitation carrying capacity is exemplified by the northern quahog (Mercenaria mercenaria) benthic cultures in Cherrystone Inlet, on the Delmarva Peninsula near the mouth of Chesapeake Bay (6 km2, mean depth 1 m, salinity 14–23) (Condon 2005). For the past ∼25 years, juvenile clams (shell height 10–15 mm) have been planted in rows (∼4 m × 18 m) at densities of 550– 1650 clams m−2, covered by polyethylene netting to reduce predation over a ∼18–30month growout period (Luckenbach and Wang 2004b) (Fig. 7.4). Estimates indicated that the bivalve cultures largely control phytoplankton dynamics in Cherrystone Inlet; the clams filtered ∼10.1–81.9% of the tidal creek volume per day, depending on the time of year (Condon 2005, and references therein). Work by Condon (2005) also supported the premise that the cultured clams also significantly influence nitrogen cycling in this ecosystem. Available literature values for clam metabolism were used to develop a model (described below) to assess the role of clams as sources

versus sinks for the N pool in Cherrystone Inlet during 2003. Clam harvest that year (∼20 × 106 clams; shell height ∼60 mm) was estimated to remove 18,000 kg N. Twice that amount (∼36,000 kg N) was removed to the atmosphere by denitrification of biodeposits, whereas 900 kg N day−1 were released to the water column by clam excretion (Luckenbach and Wang 2004). In contrast, during the higher-harvest year 2004 (57.3 × 106 clams, shell height 43 mm), Condon (2005) estimated that ∼11,600–13,000 kg N were removed through denitrification of biodeposits. A possible reason given for this discrepancy was that different literature values were used to calculate clam biodeposition rates. The total estimated annual removal of N by harvesting was 2360 kg N year−1 in 2003, versus 5450 kg N year−1 in 2004. Estimates of N recycling by clam excretion differed between the two studies (12–30 kg N day−1 estimated by Condon 2005, versus 900 kg N day−1 given above), in part because the 2005 study, but not the earlier work, included ammonification of clam biode-

Bivalve shellfish aquaculture and eutrophication

posits in estimating N release by clams. Nitrification-denitrification and ammonification of clam biodeposits accounted for more of the total clam-induced N flux than clam excretion and harvest, especially during summer, and total N recycling by clams exceeded N removal by clams in both 2003 and 2004 (Condon 2005). The modeling effort also indicated that the cultured clam population was large enough to dominate carbon and nitrogen processes in Cherrystone Inlet, and large enough to have significant effects on the phytoplankton assemblage. Clams were estimated to consume more than 50% of the gross primary production in spring and fall months when temperatures were optimal for clam growth. The large DIN fluxes from the clam cultures were suspected to support not only phytoplankton and benthic microalgal growth but also the development of thick macroalgal mats that covered the clam nets throughout much of the growing season. In poorly flushed conditions, perhaps the best studied ecosystems with intensive bivalve culture operations are the Thau Lagoon along the Mediterranean coast of southern France, and the Sacca di Goro Lagoon along the northern coast of Italy. The findings from these systems are described in detail here to provide examples of “worst-case” effects of highdensity bivalve cultures in poorly flushed conditions: In the Thau Lagoon, areas of intensive Japanese oyster culture (Crassostrea gigas) were compared with reference sites without shellfish culture. The Thau Lagoon is a shallow microtidal, wind-mixed system that sustains bottom-water anoxia during calm summer conditions (Deslous-Paoli et al. 1993). It is eutrophic from excessive land-based nutrient inputs as well as shellfish culture (Mesnage et al. 2007). Culture operations were described to cover ∼20% of the lagoon with 40 oysters m−2 area (total standing stock 22,675 tonnes, annual harvest ∼13,600 tonnes) (Deslous-Paoli et al. 1993, 1998; Souchu et al. 2001; Mesnage

183

et al. 2007). De Casabianca et al. (1997) described the lagoon as having sustained “shellfish farming-dominant eutrophication,” with abundant macroalgae Ulva rigida and Gracilaria bursa-pastoris in or near the bivalve culture areas. Mazouni et al. (1996, 1998) reported a significant influence of oyster culture on water-column DO and dissolved N concentrations, less so on water-column phosphate. Oxygen uptake in the culture area ranged from 0 μmol m−2 h−1 (January) to 11,823 ± 377 μmol m−2 h−1 (July). Ammonium and nitrate+nitrite were released from the culture areas during the summer season at 2905 ± 327 μmol m−2 h−1 and 891 ± 88 μmol m−2 h−1, respectively. In that season, the nitrate+nitrite flux represented about 20% of the total DIN production. During winter, ammonium flux was negligible, whereas nitrate+nitrite was released at 177 ± 97 μmol m−2 h−1. Phosphate release was low except in 2 months, May and November (∼1700 to ∼2700 μmol m−2 h−1). Mazouni et al. (1998) suggested that in this lagoon, oyster cultures (i.e., oysters and their epibiota) produced 2 × 107 mol N year−1 and “play a central role in N renewal in the water column.” Souchu et al. (2001) reported that grazing by the cultured oysters controlled phytoplankton biomass during all seasons except summer. Thus, for most of the year, the ammonium regenerated from the shellfish cultures was not used by phytoplankton for new production but, rather, was oxidized to nitrate by nitrifying pelagic bacteria. The authors suggested that at least a portion of this nitrate likely diffused into the sediments where it was denitrified to N2 gas and effectively removed from the ecosystem. Culture of the mussel Mytilus galloprovincialis and the Manila clam Tapes philippinarum dominates the Sacca di Goro. The lagoon is 26 km2 in area with mean depth of 1.5 m, and it receives freshwater inflows from the Po di Volano and the Po di Goro deltaic branches (Nizzoli et al. 2005). As of 2004, about one-third of the lagoon area (∼8 km2)

184

Shellfish Aquaculture and the Environment

was licensed for clam culture and 0.4 km2 was licensed for mussel culture. The mussel farms consisted of five parallel 1-km lines of trellis separated by ∼50 m of water. Mussels were cultivated on ropes (1–1.5 m in length) suspended from the trellis (mean density, 5 ropes m−2), and at maturity the biomass on each rope ranged from 9 to 12 kg fresh weight. Maximum annual production was ∼1360– 1815 tonnes (1500–2000 tons), but part of the culturing was moved to areas outside of the lagoon because of mussel death from high summer temperatures coupled with episodic anoxia. Thus, mussel production within the lagoon had declined to less than 900 tonnes year−1 (Nizzoli et al. 2005). Study of oxygen, nitrogen, and phosphorus fluxes from the mussel farm area versus a reference site revealed that the mussel farm contributed “intense biodeposition” of organic matter that stimulated sediment oxygen demand and inorganic N and P regeneration rates (Nizzoli et al. 2005). The overall benthic fluxes measured (−11.4 ± 6.5 mmol O2 m−2 h−1; 1.59 ± 0.47 mmol NH4+ m−2 h−1 and 94 ± 42 μmol PO4−3 m−2 h−1) were among the highest ever recorded from a bivalve shellfish aquaculture site (Nizzoli et al. 2005). The mussel rope community was described as “an enormous sink” for oxygen and POM, and a large source of dissolved inorganic N and P to the water column: The authors estimated that 1 m2 of mussel farm had an oxygen demand of 46.8 mmol m−2 h−1, and released inorganic N and P at 8.5 mmol m−2 h−1 and 0.3 mmol m−2 h−1, respectively. Thus, the mussel ropes accounted for 70% to more than 90% of the overall oxygen and nutrient fluxes. Nizzoli et al. (2005) suggested that the net effect of the mussel farm on phytoplankton might be to increase phytoplankton turnover and overall production, rather than to limit phytoplankton biomass through filter feeding. Manila clam farms were reported to cover about one-third of the area of the Sacca di Goro Lagoon, at densities up to 2500 clams m−2

(Bartoli et al. 2001; Nizzoli et al. 2007). Bartoli et al. (2001) described an example of more adverse ecosystem-scale effects from shellfish aquaculture: The clam farms were estimated to have stimulated whole-lagoon dark oxygen consumption and ammonium recycling by a factor of 1.8 and 6.5, respectively. The authors assessed the effects of dense clam cultures (up to ∼2300 adults m−2) that covered about onethird of the area of the lagoon. Shortly following initiation of these cultures and their harvest by sediment dredging, extensive macroalgal overgrowth (the chlorophyte Ulva rigida, which is common in nutrient overenriched areas) developed along with anoxic events and mass-death of the cultured shellfish. The anoxic events generally occurred during summer when the massive Ulva growth (up to 800 g dry weight m−2) suddenly died and decomposed, promoting high fluxes of sulfide from the sediments (Viaroli et al. 1996). Bartoli et al. (2001) compared a reference site with few clams versus a culture area for benthic nutrient fluxes, oxygen and carbon dioxide concentrations, and extrapolated the data to estimate ecosystem-scale effects. The data were based on five sediment cores (height 40 cm, inner diameter 20 cm) collected from both the control site and the culture site during one summer season. Densities of other macrofauna were low in the cores from both sites. Oxygen, CO2, NH4+, reactive silica, and PO4−3 fluxes were significantly higher in the culture areas, attributed to clam metabolism and to the reducing conditions in the surficial sediments (Table 7.6). Phytoplankton removal or chlorophyll a flux to the sediments was highly variable, with the highest net flux (630 μg chlorophyll a m−2 h−1) in the core with the highest clam density. These data were used to make interpretations about the influence of clam aquaculture in the Sacca di Goro at the ecosystem scale. When the data were extrapolated to the lagoonal system, average whole-lagoon dark sediment oxygen demand and CO2 production

Bivalve shellfish aquaculture and eutrophication

185

Table 7.6 Average environmental conditions in a reference site with few short-necked clams (Tapes [Ruditapes] philippinarum) versus a dense culture site. Parameter

Reference site

Clam culture area

Dark oxygen consumption*

−2.67 ± 0.58 mmol O2 m−2 h−1

−12.49 ± 4.54 mmol O2 m−2 h−1 (4.7× higher)

CO2 production rates*

1.22 ± 0.81 mmol CO2 m−2 h−1

10.42 ± 5.61 mmol CO2 m−2 h−1 (8.5× higher)

O2 flux*

Estimated oxygen demand 2.78 mmol O2 m−2 h−1

Estimated oxygen demand 4.72 mmol O2 m−2 h−1 (1.7× higher)

NH4+ flux from sediments*

0.15 ± 0.22 mmol NH4+ m−2 h−1

2.65 ± 0.22 mmol NH4+ m−2 h−1 (17.7× higher; 4.13 mmol NH4+ m−2 h−1 in the core with highest clam density)

Oxidized N (NO3−, NO2−), urea fluxes

Highly variable; not statistically different between sites

PO4−3 flux from sediments*

Negligible

0.15 ± 0.02 mmol PO4−3 m−2 h−1

Silicate flux from sediments*

0.04 ± 0.08 mmol SiO2 m−2 h−1

0.37 ± 0.14 mmol SiO2 m−2 h−1 (9× higher)

Potential O2 consumption* from resuspension of sediments (0–5 cm in depth), simulating harvest

27 ± 9 mmol O2 m−2 h−1

56 ± 12 mmol O2 m−2 h−1 (2× higher)

Compiled from Bartoli et al. (2001). An asterisk (*) indicates that the difference between the two sites was statistically significant (P < 0.001); all of these parameters except PO4−3 were significantly correlated with clam biomass.

were stimulated by a factor of 1.8 and 3.3, respectively, and nutrient release was 6.5-fold higher for NH4+N and 4.6-fold higher for PO4−3 (Bartoli et al. 2001). The highly biodegradable clam feces and pseudofeces could potentially promote rapid nutrient recycling— up to 4000 μmol NH4+ m−2 h−1 and 150 μmol PO4−3 m−2 h−1—that would stimulate macroalgal growth. It was estimated that at maximal production (13,600 tonnes [15,000 tons], although 18,000–22,700 tonnes was suggested to be more realistic) 41.7 tonnes of N and 9 tonnes of P were removed with clam harvest. Annual nutrient loads from land-based (freshwater) inputs were estimated at 1179 tonnes of N and 36 tonnes of P. Thus, clam harvest was estimated to have removed about 4% of the N and 25% of the P in external, land-based inputs. On the other hand, the clams stimulated inorganic N and P regeneration from the

sediments to the overlying water: During midMay to mid-September (water temperature >20°C, dark period 10 h day−1), an estimated 457 tonnes of NH4+ and 50.8 tonnes of PO4−3 were regenerated to the water column in the clam culture areas. By comparison, the amount of N and P removed by clam harvest was small. The authors’ overall interpretation was that the cultured clams had significantly affected this lagoonal ecosystem through increased sediment and water column anoxia and high nutrient fluxes to the water column, and that the premise of clam aquaculture functioning as a control for eutrophication is unrealistic in the Sacca di Goro because of the high densities of clams and extensive culture area. In later work in the same lagoon (Nizzoli et al. 2005, 2006a, 2006b, 2007), Nizzoli and coworkers developed N and P budgets for a control area (1600 m−2) with low density of

186

Shellfish Aquaculture and the Environment

Manila clams (30 ind. m−2), a low-density culture area (∼400 m2; 300 young individuals seeded m−2), and a high-density culture area (∼110 m2; ∼800 young individuals seeded m−2). External freshwater nutrient loads were estimated for comparison with excretion and filtration activity of the clams, deposition of particulate matter, and nutrient recycling including light and dark fluxes at 1, 3, 5, and 7 months after April seeding. Relative to the control area, in the culture areas there was a significant increase in the downward fluxes of particulate nutrients coincident with an enhancement of dissolved nutrient forms (ammonium, soluble reactive phosphate) that were released (effluxed) to the water column (Table 7.7). Estimated particulate N and par-

ticulate P uptake was five- (low-density culture) to ninefold (high-density culture) higher than at the control site, indicating that a major fraction of the suspended particulate matter was retained by the bivalves. Over the entire farming cycle, the total dissolved phosphorus (TDP) internal loading at the low- and high-density culture sites was two- to fourfold higher than at the control site. Whereas the sediments in the control site were a net sink for dissolved N, substantial N was released, mostly as ammonium, at the culture sites. The contribution of the bivalves to N and P loads at the lagoon level through filtration, assimilation, regeneration, and burial pathways was estimated assuming a clam market

Table 7.7 Estimated influence of short-necked clam cultures (Tapes philippinarum) on nutrient cycling and removal in the Sacca di Goro Lagoon (compiled from Nizzoli et al. 2006a). Parameter (entire farming cycle)

Control

Low density

High density

Entire farming cycle (mol m−2) Particulate N uptake by clams from the water

1.7

9.1

16.3

−0.3

1.6

6.9

Particulate P uptake

0.1

0.6

1.0

TDP efflux

0.2

0.5

0.8

Total dissolved N (TDN) flux (mostly as ammonium)

End of farming cycle Harvested N as mollusc flesh

∼0

0.4

1.8

Harvested P as mollusc flesh

∼0

0.02

0.04

Fraction of biodeposited N exported as commercial product

—

1.2%

6%

Fraction of biodeposited P exported as commercial product

—

0.75%

3%

Fraction of biodeposited N recycled as dissolved inorganic or organic N

—

∼7.5%

Fraction of biodeposited P recycled as dissolved inorganic or organic P

—

∼2%

30% 3%

Overall (annual cycle—5440 tonnes of clams produced in the lagoon over the study duration) Removal of nutrients by Tapes philippinarum: 0.23 g N, 0.03 g P per individual → 124 tonnes PN, 17 tonnes PP Recycling of nutrients by Tapes philippinarum: 0.15 g N, 0.02 g P per individual → 83 tonnes TDN, 11 tonnes TDP Removal of nutrients by harvesting Tapes philippinarum: 15 tonnes N, 0.8 tonnes P

Bivalve shellfish aquaculture and eutrophication

size of 10 g wet weight and a crop of 5440 tonnes (6000 tons), roughly half of the total biomass produced over an annual cycle in the Sacca di Goro Lagoon from aquaculture of Tapes philippinarum (Table 7.7). Comparison of the bivalve contribution to loadings delivered to the lagoon by freshwater sources indicated that the amount of particulate matter processed by the clams was within the same order of magnitude as the land-based loadings. The regenerated fraction was about 30% of the external TDN load, and about 90% of the external TDP load. Overall, Nizzoli et al. (2006a) estimated that in this poorly flushed lagoon, there were elevated rates of ammonium production, phosphorus enrichment, and oxygen consumption at the ecosystem scale. This study indicated rapid coupling between sedimented bivalve biodeposits and benthic recycling. Nizzoli et al. (2006a) suggested that the mollusc cultures likely reduced the export of particulate matter from the lagoon to the open sea. The authors’ overall interpretation was that in this lagoon, a significant fraction of particulate N and P external loads is retained and recycled as dissolved nutrients by clam aquaculture. They suggested that the retention of particulate matter in culture areas, and the alteration of the particulate-to-dissolved nutrient ratio, could negatively affect water and sediment quality and stimulate the growth of nuisance macroalgae in the lagoon ecosystem. The authors acknowledged that their comparison did not consider nutrients supplied by phytoplankton growth, or nutrients contributed to the lagoon by tidal currents from the sea. They also recognized that their extrapolation to the ecosystem level from the small experimental control and aquaculture plots should be considered with caution. Of the 72 ecosystems reviewed here (Table 7.3), only ∼6% or four ecosystems have sustained system-level adverse impacts from large, intensive bivalve culture operations (Table 7.3). The other 94% have sustained negligible

187

or only localized significant adverse effects contributing to eutrophication from bivalve shellfish aquaculture. This analysis is based upon peer-reviewed, published data. Nevertheless, merits mention that Pawlowski et al. (accepted; see below) described some coastal waters along the Orient such as China as having sustained system-level adversed impacts from bivalve aquaculture, based on review of unpublished data from the Food and Agriculture Organization of the United Nations.

Modeling efforts to assess relationships between bivalve aquaculture and eutrophication Nutrient enrichment interactions with bivalve aquaculture only recently began to be assessed through modeling efforts, mostly within the past decade. Earlier production models focused on growth and development of bivalves under different environmental conditions (Bayne and Warwick 1998; Henderson et al. 2001). Models of the population dynamics of a given cultured bivalve species were developed that considered the number of individuals, growth, food availability, population renewal through seeding, marketable size, water residence time, certain ecophysiological traits, and other variables (e.g., Bacher et al. 1998) to estimate the carrying capacity of an ecosystem for bivalve production, that is, the maximum biomass of cultured shellfish that a farm or waterway could sustain without a decrease in production (Dame and Prins 1998; Smaal et al. 2001; Duarte et al. 2003). Some of these models included spatial features of the embayment based on a hydrodynamic model, and also depicted the nitrogen or carbon cycling among phytoplankton, cultured oysters, and detritus. Some models were constructed with a twodimensional, coupled physical-biogeochemical framework that considered more than one shellfish species as polycultures (Duarte et al.

188

Shellfish Aquaculture and the Environment

2003). Other efforts assessed ecosystem effects of shellfish culture using a carbon-based food web model that examined benthic/pelagic coupling by forcing a shift from pelagic filterfeeders to benthic consumers (Leguerrier et al. 2004). Eutrophication began to be more directly considered in carrying capacity models that were developed for shellfish culture at local scales or sites. For example, a mussel production model (MUSMOD©—Campbell and Newell 1998) was created to guide seeding of bottom culture lease sites in Maine, USA, to optimal carrying capacity, and it predicted mussel production based on physical and biological variables. Some carrying capacity models have considered shellfish growth as a function of ecosystem characteristics related to nutrient enrichment, such as DO deficits or primary production. EMMY (Ecophysiological Model of Mytilus edulis), an early model by Scholten and Smaal (1999), simulated growth and reproduction of individual mussels and examined the effects of eutrophication reduction scenarios on mussel growth under controlled experiments over a range of nutrient loads to mesocosms. The model was designed for application as a management tool to estimate carrying capacity. As other examples, Inglis et al. (2000) described a carrying capacity model for mussel growth and condition that consists of three integrated submodels including (1) a hydrodynamics model that simulates effects of tides, freshwater inputs, and weather on current flows, flushing rates, and water column structure; (2) an “ecosystem model” to simulate phytoplankton abundance, which includes water stratification, light penetration/intensity, nutrient supplies and recycling within both the water column and sediments, as well as mortality, sedimentation, and predation of phytoplankton; and (3) a mussel energetics mode that considers filtration rates, the amount of food ingested and assimilated, and the proportions allocated to growth and reproduction. Condon (2005) developed a model to assess

the effects of hard clam cultures on carbon and nitrogen cycling in Cherrystone Inlet based on data for the clam population and water quality, using published values for clam feeding and respiration rates. The model was used to evaluate the potential influence of clam cultures on the particulate carbon pool through feeding and respiratory demands, and on carbon and nitrogen cycling via feeding and biodeposition, excretion, microbial processing of wastes, and clam harvest. This model indicated that Cherrystone Inlet was at or near exploitation carrying capacity for clam aquaculture. Numeric models are becoming increasingly popular as management tools to assist in the rapid expansion of shellfish aquaculture worldwide by refining site selection, defining site limitations, optimizing production, and designing and implementing monitoring programs (Chamberlain et al. 2006; Giles et al. 2009). Some of these models, mostly adapted from finfish operations, are being used to assess the magnitude and spatial extent of environmental effects from shellfish aquaculture. It is important to note, however, that shellfish aquaculture operations generally are larger and more diffuse than finfish farms, characteristics that would result in different nutrient dispersal patterns (Hartstein and Stevens 2005). These operations can also attenuate flow over substantial areas (Plew et al. 2005). In addition, sinking speeds of fecal and pseudofecal material from shellfish culture would be expected to be slower than that of fecal material from finfish operations because the shellfish feces are derived from phytoplankton, whereas the finfish feces consist of the remains of relatively more dense fish food (Cromey et al. 2002). More recently, various modeling efforts have aimed to link watershed nutrient loading and ecosystem carrying capacity for shellfish aquaculture (e.g., Fig. 7.5). Luckenbach and Wang (2004a,b) described work to link a watershed-based loading model with a physical transport-based water quality model to simulate primary production and predict car-

Bivalve shellfish aquaculture and eutrophication

189

Meteorological data Meteorological data Ulva (on/off) Po River nutrient data scenarios

Lagoon Model Watershed Model

Flows nutrients

Open sea: flows, nutrients

Aquaculture: area, initial conditions, and seeding densities

Figure 7.5 Integrated model for the Sacca di Gorro Lagoon and its watershed. (Redrawn from Marinov et al. 2007.)

rying capacity for intensive hard clam aquaculture in the Chesapeake Bay area. Their water quality model realistically simulated primary production and various water quality parameters. They also developed and tested watershed loading models that predict surface and groundwater inputs to coastal waters. Planned efforts included coupling the water quality and watershed loading models, developing clam physiology and populationlevel submodels and a sediment deposition/ resuspension submodel, and then linking all of these components to estimate exploitation carrying capacity for clam production in selected areas such as Cherrystone Inlet. The ultimate goal is to use the coupled models to predict how land use changes will affect water quality, primary production, and shellfish carrying capacity in this system and, with parameter modifications, in other coastal waters. Marinov et al. (2007) developed a coupled watershed and three-dimensional biogeochemical model for the Sacca di Goro Lagoon. It considered clam productivity with versus without macroalgal blooms, tied to nutrient enrichment and influences of climatic variability as dry, average, and wet years. Chapelle et al. (2000) developed an ecosystem model for the Thau Lagoon, based on nitrogen cycling and DO concentrations, toward evaluating the effects of intensive oyster aquaculture versus watershed inputs on the lagoonal ecosystem. The watershed includes substantial agriculture, industries, and urbanized areas. The model used data from the OxyThau program

(Deslous-Paoli et al. 1993), which collected 5 years of data to assess interactions among oyster culture, water column/sediment nitrogen cycling, and land-based (watershed) versus climatic influences on the lagoonal ecosystem. It coupled hydrodynamics from a twodimensional model with nutrient cycling integrated into a box model. Simulations indicated that nitrogen cycling and oxygen deficits were driven by meteorological forcing during wet seasons, especially precipitation events which caused land-based nutrient inputs that stimulated new primary production. During the dry summer season, oyster excretion/sediment release and microzooplankton excretion/ mineralization produced substantial ammonium that stimulated “regenerated” primary production, so that the ecosystem remained highly productive without land-based inputs. Thus, depending on the season, both landbased inputs and shellfish cultures were important in the nitrogen dynamics of this poorly flushed lagoon. The model suggested that biodeposition from the oyster cultures and subsequent sediment release was a major source of N for the lagoonal ecosystem, and was linked to oxygen reduction and localized hypoxia. Giles et al. (2009) noted that previous numerical models of biodeposition from shellfish farms have overlooked biodeposit decay and most erosion features, which could substantially affect estimates. They used two particle tracking models, one to simulate initial dispersal of fecal pellets and the other for initial dispersal and erosion, to estimate

190

Shellfish Aquaculture and the Environment

biodeposition from a suspended mussel farm of the greenshell mussel Perna canaliculus. In sheltered areas, represented by the initial dispersal model, biodeposit decay mostly affected fecal pellet density on the seafloor. In highenergy areas, represented by the erosion model, decay more strongly affected the spatial extent of the detectable farm footprint. The model’s predicted fluxes underestimated measured rates by about 50%. Two other models, DEPOMOD and FARM, are briefly described here to illustrate the utility of modeling approaches in assessing interactions between shellfish aquaculture and eutrophication, and also the economic benefit of bivalve farms in mitigating land-based eutrophication (below). First, the finfish aquaculture waste model DEPOMOD (models the deposition and biological effects of waste solids from marine salmon cage farms; Cromey et al. 2002) was adapted for suspended mussel aquaculture by Weise et al. (2009), considering field data for species-specific biodeposition rates and particle settling velocities, along with several finfish model parameters. ShellfishDEPOMOD was tested at three Mytilus edulis farms in Quebec, Canada, that differed in hydrologic regime. Model predictions for sedimentation rates were compared with data for deposition rates from sediment traps. Localized effects of sedimentation rates were reported at ∼two to five fold higher than rates in corresponding control sites without mussel aquaculture. The model accurately predicted accumulation of sediments within 30 m to more than 90 m for farms in shallow versus deeper sites, respectively, except for a site in House-Harbor Lagoon. The authors attributed the disparity between model predictions and observed sedimentation rates to resuspension and advection of nonfarm-derived materials and complex hydrodynamics. The model also correctly predicted patterns of waste disposal at one site while underestimating biodeposition, attributed to the fact that biodeposits from biofouling communities were

not considered. Highest biodeposition rates (>15 g m−2 day−1) coincided with localized changes in benthic community structure, as indicated by the Infaunal Trophic Index (Maurer et al. 1999) and the Marine Biotic Index (Borja et al. 2000). The Farm Aquaculture Resource Management (FARM) model (Fig. 7.6) was designed for prospective analyses of culture location and species selection; ecological and economic optimization of culture practices; and environmental assessment of farm-related eutrophication effects, including mitigation (Ferreira et al. 2007; see also Chapter 1 in this book). FARM can be used to screen various water quality effects and to examine nutrient mass balance, so it provides a valuation methodology for integrated nutrient management. The modeling framework applies a combination of physical and biogeochemical models, bivalve growth models, and screening models to assess shellfish production and eutrophication. It originally was parameterized for five species, alone or mixed, including the Pacific oyster (Crassostrea gigas), the blue mussel (Mytilus edulis), the Manila clam (Tapes philippinarum), the cockle (Cerastoderma edule), and the Chinese scallop (Chlamys farreri). FARM adapts the Assessment of Estuarine Trophic Status (ASSETS) model (Bricker et al. 2008) for use at local scales, considering chlorophyll a and DO. Its eutrophication simulation includes a sustainability metric for carrying capacity, allowing users to test for thresholds of low DO and to assess potential consequences for water quality and stock mortality. Ferreira et al. (2007) used the FARM model to assess quantitatively the role of shellfish culture in controlling nutrient emissions to coastal waters. They developed a mass balance for nutrients in a 6000-m2 bottom-culture oyster (Crassostrea gigas) operation, using data from various cultivated coastal ecosystems and realistic oyster densities. The simulation was for a 45-day period, using seed

Bivalve shellfish aquaculture and eutrophication

191

Farm length Width

Chl a

Current

Shellfish Depth

POM

1

2

3

Chl a

n

POM

Sections

Physics and biogeochemistry models

Mortality

e.g., food depletion

e.g., Chl a, POM

Shellfish individual growth models

Shellfish population dynamics model

Optimization of farm activities

Shellfish growth

e.g., harvestable biomass, APP

e.g., ASSETS grade Nutrient trading

Eutrophication assessment screening model

Chl a, dissolved oxygen

Shellfish production screening model

e.g., MPP, VMP

Current

Figure 7.6 Conceptual scheme of the FARM model. POM, particulate organic matter; MPP, marginal physical product; VMP, value of the MPP; APP, average physical production. (Modified from Ferreira et al. 2007).

densities of 25, 100, and 500 individuals m−3. The model predicted that the lowest shellfish density would reduce chlorophyll a by 15%, while DO remained at or above ∼6 mg/L. The moderate shellfish density was predicted to reduce chlorophyll a by 45%, but DO decreased to ∼4 mg/L or more. While the high shellfish density would have effected a 92% reduction in chlorophyll a, it also was predicted to cause hypoxic conditions from a DO sag to ∼1.8 mg/L (Ferreira et al. 2007). Among the most ambitious efforts to date to assess contributions of N and P from

shellfish aquaculture was contributed by Pawlowski et al. (accepted), who developed a model that estimates shellfish culture inputs worldwide. Most of the data used for the model simulations were obtained from the Food and Agricultural Organization of the United Nations (2008). Pawlowski et al. acknowledged major uncertainties in their approach because of its global scale and lack of sufficient information on some parameters. Noting the major increase in shellfish aquaculture that is both underway and anticipated, Pawlowski et al. assessed nutrient release from

192

Shellfish Aquaculture and the Environment

marine shellfish culture (bivalves, crustaceans, gastropods, collectively) as an important contributor to dissolved and particulate nutrients in coastal marine ecosystems, especially in eastern Asia. The authors projected dramatic increases in nutrient contributions from shellfish cultures by 2050. In some areas, such as coastal waters of China, marine shellfish aquaculture was evaluated as already contributing significantly to total nutrient inputs: As of 2006, shellfish aquaculture was estimated to have contributed 18% and 30% of all marine aquaculture + river exports of N and P, respectively. Nevertheless, the overall contribution of shellfish aquaculture to nutrient inputs was projected to increase from ∼1% of total river exports in 2006 to, at a maximum, ∼6% by 2050. Thus, Pawlowski et al.’s model indicated that projected as well as recent contributions of shellfish aquaculture to global N and P loading of coastal marine ecosystems are small in comparison with global river N and P exports.

Eutrophication of coastal waters from land-based nutrients In comparison with the generally localized effects of bivalve aquaculture on nutrient supplies, the following information depicts largescale, extensive, ubiquitous impacts of land-based nutrient pollution on coastal ecosystems: By the turn of the twenty-first century, about 60% of U.S. coastal rivers and bays already were moderately to severely degraded from land-based nutrient pollution (National Research Council 2000). Nitrogen is the primary nutrient that limits phytoplankton growth in many estuarine and coastal ecosystems and, thus, is a key nutrient in eutrophication (Burkholder and Glibert 2011). Global consumption of nitrogen fertilizer has dramatically increased over the past 70 years, and much of this increase is in the form of urea fertilizer which is an organic N form that has been linked to increased growth of various harmful algal species (Glibert et al. 2006) (Fig.

7.7). Phosphorus, another major nutrient that can stimulate phytoplankton overgrowth, has shown a more modest increase as well (Glibert and Burkholder 2006).

Nutrient or ecological stoichiometry The ratio of N to P, or the nutrient stoichiometry, has also been greatly altered by landbased, anthropogenic nutrient additions. Nutrient stoichiometry relates changes in the relative composition of N and P in cells and tissues of aquatic organisms versus the water column. Changes in the relative proportion of N and P have promoted major alterations in metabolism, species composition, and food web structure (Sterner et al. 2002; Elser et al. 2007). Overall, as Howarth (2008) wrote, The past few decades have seen a massive increase in coastal eutrophication globally, leading to widespread hypoxia and anoxia, habitat degradation, alteration of food web structure, loss of biodiversity, and increased frequency, spatial extent, and duration of harmful algal blooms.… Agricultural sources are the largest source of nitrogen pollution to many of the planet’s coastal marine ecosystems. The rate of change in nitrogen use in agriculture is incredible, and over half of the synthetic nitrogen fertilizer that has ever been produced has been used in the past 15 years. Atmospheric deposition of nitrogen from fossil fuel combustion [urban source] also contributes . . . and is the largest single source of nitrogen pollution in some regions.

Estuaries and coastal waters are now the most nutrient overenriched ecosystems in the world (Wassmann 2005), attributed primarily to land-based nutrient sources, and coastal human population growth and nutrient loading from land-based pollution sources are projected to increase exponentially over the next two decades (Howarth et al. 2002) (Fig. 7.8). Regions of large-scale nutrient overenrichment from land-based sources include (among many examples) the Kattagat/Baltic

Bivalve shellfish aquaculture and eutrophication

Million tonnes N

250

193

(A)

200 150 100 50

20 20

20 10

00 20

19 90

0 19 8

19 6

19 7

0

0

0

7.0 (B) Riverine export (Tg N year−1)

6.5 6.0 5.5 5.0 4.5 4.0 3.5 3.0 2.5 1960

1970

1980

1990

2000

2010

2020

2030

Figure 7.7 (A) The change in world consumption (million metric tons of N) of total synthetic nitrogen fertilizers (solid line) and urea consumption (solid bars) since 1960. (From Glibert et al. 2006 in Biogeochemistry, with permission.) (B) Global riverine (land-based) export of nitrogen through the 1990s, and estimated through 2030. (Redrawn from Howarth et al. 2002.)

Sea and the eastern North Sea in northern Europe; the northern Adriatic Sea and northwestern Black Sea in southern Europe; the Seto Inland Sea, Yellow Sea, and East China Sea in the Orient; and Long Island Sound, Chesapeake Bay, the Albemarle-Pamlico Estuarine System, and the northern Gulf of Mexico in the United States (Boesch 2002). Many estuaries, coastal embayments, and coastal lagoons in Europe (Crouzet et al. 1999; Conley et al. 2000), Japan (Suzuki 2001), Australia (McComb 1995), and the United States (Bricker et al. 2008) have been adversely affected by landbased nutrient pollution. A recent assessment of nutrient-related impacts in U.S. estuaries indicated that nearly two-thirds of the assessed systems are moderately to seriously degraded by land-based eutrophication, considering

noxious/toxic blooms of phytoplankton and macroalgae, oxygen deficits, loss of beneficial submersed aquatic vegetation, and other factors (Bricker et al. 2007, 2008). Conditions are predicted to worsen in nearly two-thirds of these estuaries within the next decade (Bricker et al. 2008). Increased phytoplankton biomass from nutrient overenrichment may be beneficial to bivalve shellfish aquaculture until noxious or toxic algal species begin to be directly or indirectly stimulated by excessive nutrient inputs (Burkholder 2001; Burkholder et al. 2008). Altered nutrient supplies and supply ratios from land-based sources have been directly related to the loss of beneficial algal food species, and their replacement by undesirable algae that are toxic, not readily filtered, and/

194

Shellfish Aquaculture and the Environment

P inputs (mmol m–2 year–1)

10,000

1000

100

10 Freshwater wetlands and lakes Forests

1

Agroecosystems Estuaries and coastal waters

0.1 10

100

1000

N inputs (mmol m

10,000 –2

100,000

–1

year )

Figure 7.8 Phosphorus and nitrogen inputs to various types of ecosystems, showing the highest nutrient enrichment for estuaries and coastal waters (Wassmann 2005).

or not as nutritious (Ryther 1954; Smayda 1989; Cloern 2001). Scientists have now reached consensus that land-based anthropogenic nutrient enrichment is an important cause of many harmful algal blooms worldwide (Heisler et al. 2008). Beyond depressed food quality, the toxins from some algal species can bioaccumulate in shellfish and decrease fecundity, promote disease and death, and render shellfish unsafe for human consumption (see reviews in Shumway 1990; Burkholder 1998). Shellfish cultures in many regions of the world must be carefully monitored for algal toxins, and this problem is apparently increasing in some areas (Shumway 1990; Curtis et al. 2000). In some areas such as along the coast of Sweden, biotoxin accumulation is now considered the largest impediment to further expansion of commercial shellfish operations (Lindahl et al. 2005). A positive relationship between nitrogen loading and harmful algal blooms is evident from comparison of the global distribution of land-based N export and the documented occurrences of several major harmful algal species (Glibert and Burkholder 2006). Back to consideration of geoduck culture in

the northwestern United States, a strong correlation has been established between harmful algal blooms—including species that thrive in nutrient overenriched waters (Burkholder et al. 2008; Heisler et al. 2008)—and human population density, rather than geoducks, in the Northwest (GEOHAB 2006). While the comparative role of geoduck culture in contributing to eutrophication is not yet known, the role of other anthropogenic inputs, collectively considered, is clear. The nutrient overenrichment and associated pollutants from land-based eutrophication, such as suspended sediments, microbial pathogens, and pesticides and other toxic substances, directly or indirectly have increased oxygen deficits, reduced or eliminated habitat for wild and cultured shellfish, contaminated coastal waters with fecal bacteria and microbial pathogens, depressed recruitment and survival of shellfish larvae and juveniles, and increased physiological stress and disease (Bricker et al. 1999; Mallin et al. 2000; Breitburg 2002; Wiegner et al. 2003; Glasoe and Christy 2004; Bricker et al. 2008, and references therein). Accordingly, as an example among many such studies, Scott et al. (1996) compared 60 sta-

Bivalve shellfish aquaculture and eutrophication

tions in two tidal creeks, one of which drained a highly urbanized watershed and the other, a relatively undisturbed watershed in the southeastern United States. Nearly 70% of the sampling sites from the tidal creek in the urbanized watershed were closed to shellfish harvesting because of excess fecal coliform densities, and mortality rates of juvenile and adult eastern oysters (Crassostrea virginica) were much higher than in a tidal creek that drained the undisturbed watershed. Monitoring of shellfish meats indicated that more than 50% of stations in both tidal creeks exceeded the Interstate Shellfish Sanitation Conference Depuration Meat Standard. The pervasive, major impacts of land-based eutrophication overwhelm the mostly localized effects of bivalve aquaculture. During the past several decades, for example, catastrophic losses of seagrass meadows have been documented worldwide, especially in quiet, poorly flushed estuaries and coastal embayments and lagoons with reduced tidal flushing where land-based nutrient loads are both concentrated and frequent (see review in Burkholder et al. 2007). Along with inland watershed inputs of nutrients transported to marine coasts by rivers and estuaries (Caraco 1995; Vitousek et al. 1997), rapidly increasing human population density on coastlands is more than double that in inland areas (Nicholls and Small 2002, McGranahan et al. 2007). Seagrass decline in favor of macroalgae or phytoplankton is a typical response, and cultural eutrophication from land-based sources has been invoked as a major cause of seagrass disappearance worldwide (Burkholder et al. 2007). The loss of seagrass meadows has destroyed habitat for wild shellfish and many other beneficial fauna. By contrast, minimal seagrass loss from bivalve shellfish culture generally has been reported (e.g., Crawford et al. 2003), mostly in localized areas from physical disturbance during placement and harvest (e.g., Everett et al. 1995), and rarely from nutrient inputs (e.g., De Casabianca et al.

195

1997). Instead, bivalve shellfish culture generally reduces phytoplankton and other turbidity, thus affording more light for seagrass growth (Newell 2004). Another compelling example of the fact that land-based nutrients, in most coastal waters, represent the overwhelming cause of eutrophication was given by Páez-Osuna et al. (1998) for shrimp aquaculture (white and blue shrimp—Penaeus vannarnei and Penaeus stylirostris, respectively) in coastal waters of Mexico. Shrimp culture is generally considered to cause substantially more environmental degradation than bivalve culture (Naylor et al. 1998). About 2 kg of feed are needed to produce 1 kg of shrimp in semi-intensive and intensive production systems, as much as onethird of the feed is not consumed, and pond draining during shrimp harvest releases about 90% of all of the nutrients that are produced (see references in Páez-Osuna et al. 1998). The authors linked the ∼250 shrimp farms along the northwestern coast of Mexico to significant localized impacts. Nevertheless, the intensive shrimp cultures were estimated to contribute only about 1.5% of the land-based anthropogenic N and about 0.9% of the landbased anthropogenic P to the coastal waters of Mexico.

Ecological and economic benefit of bivalve aquaculture in combating eutrophication Native shellfish additions commonly have been considered as a means of helping to reverse cultural eutrophication effects in shallow waters. As a recent example, Cerco and Noel (2007) added an oyster module to a predictive eutrophication model of Chesapeake Bay to assess the potential utility of native oyster restoration (Crassostrea virginica) on DO, phytoplankton biomass as chlorophyll a, light attenuation, and submersed aquatic vegetation. The model predicted that a

196

Shellfish Aquaculture and the Environment

10-fold increase in the existing oyster biomass would reduce the summer surface chlorophyll a, system-wide, by about 1 mg m−3, increase the summer average deep-water DO by 0.25 g m−3, substantially increase beneficial submersed aquatic vegetation, and remove 30,000 kg N day−1 through enhanced denitrification. This latter amount of N removal was estimated to be more than the nitrogen added to the bay by direct atmospheric deposition, or about 10% of the total system loading. Similarly, most shellfish aquaculture is thought to have an overall positive effect on water quality, primary production, and biodiversity except, as mentioned, for intensive culture in localized, poorly flushed waters (Naylor et al. 2000; Gibbs 2004; McKindsey et al. 2006). Thus, shellfish aquaculture has been considered as bioremediation tool for polluted sites, not only for reducing nutrient loads and phytoplankton blooms but also for removing toxic contaminants and reducing concentrations of microbial pathogens (Rice 2001; Gifford et al. 2004). The grazing role of shellfish in removing phytoplankton from the water by filter feeding can offset phytoplankton stimulation by nutrient overenrichment from land-based sources. Shellfish culture has also been considered as a means to reduce the primary symptoms of eutrophication such as increased chlorophyll a, and associated oxygen deficits that are caused by high phytoplankton respiration at night and bloom senescence, death, and decomposition (Newell 2004). Shellfish removal of excess phytoplankton and other particulate matter can also increase the light available for growth of seagrasses that provide beneficial habitat (Burkholder et al. 2007). Harvest of the cultured bivalves additionally removes nutrients from the system (Songsangjinda et al. 2000), although this effect may be minor depending on the system (e.g., Bartoli et al. 2001). The harvested animals could be used for seafood, fodder, and agricultural fertilizers, “thus recycling nutrients from sea to land” as a cost-effective method to improve coastal water quality

(Lindahl et al. 2005). Integration of shellfish culture with some forms of agriculture for overall reduction of nutrient inputs appears to be increasingly justified considering nutrient discharge regulations and increasing effluent treatment costs (Andrew and Frank 2004). Beyond qualitative or localized studies, few analyses are available on the economic value of bivalve aquaculture in reducing nutrient supplies to coastal waters, but the concept is promising and the knowledge base is beginning to rapidly expand, as the following examples illustrate. Hart (2003) applied a dynamic linearquadratic model to test the effectiveness of two control measures on N pollution to coastal waters along western Sweden—one upstream as agricultural abatement, and the other downstream as mussel aquaculture. With respect to mussel aquaculture, the model considered harvest and removal of the mussels as the main N reduction measure. In the northwestern fjords, mussel aquaculture was about 2000 tonnes per year, but was estimated at potentially 15,000 tonnes (Haamer 1996, Kollberg 1999 in Hart 2003), which would correspond to removal of 150 tonnes of N per year or about 20% of the inflow to these waters from land-based sources (Haamer 1996). The overall interpretation was that mussel cultivation could be a cost-effective measure against N pollution along the west coast of Sweden, although Hart (2003) cautioned that various assumptions used to model the costs of mussel cultivation needed to be further tested. Also focusing on the Swedish west coast, Lindahl et al. (2005) modeled the potential effects of blue mussel aquaculture on N cycling within the Gullmar Fjord. It was assumed that the outflow water from the mussel culture area had unchanged concentrations of phosphate and nitrate, but an 18% increase in ammonium and a 17% increase in detrital particles (from mussel intake of N). It was also assumed that when the concentration of plankton in the water column was greater than 4 μg chloro-

Bivalve shellfish aquaculture and eutrophication

ORGANIC FERTILIZER

ORGANIC FODDER

197

HUMAN CONSUMPTION

MUSSEL FARM NUTRIENTS

PHYTOPLANKTON

BIODEPOSITS Figure 7.9 The Agro-Aqua recycling system of nutrients from sea to land. (Redrawn from Lindahl et al. 2005; see also Chapter 8 in this book.)

phyll a L−1, the mussels would not be able to digest all of the food and would reject some filtered plankton as pseudofeces that sank as detritus. The model output indicated that net transport of dissolved and particulate N at the mouth of the fjord was reduced by 20% through mussel farming. Lindahl et al. (2005) suggested that nutrient trading systems involving mussel aquaculture should be introduced to improve coastal water quality, augmenting N reduction by the sewage treatment plant in the Lysekil community. As of 2004 (most recent available data), the plant was releasing more than 36 tonnes (nearly 40 tons) of N per year to the Gullmar Fjord, with plans to reduce that number by 25 tonnes (28 tons) in accordance with European Union regulation 91/271/ EEG. Realistic expansion of mussel farming

was estimated potentially to accomplish removal of 25 tonnes of N through harvest of 2540 tonnes (2800 tons) of mussel biomass with N content of ∼1%. About two-thirds of the harvested mussels could be used for human consumption, and the remaining small or damaged mussels could be used for Agro-Aqua recycling of nutrients (Fig. 7.9). In a field study, 4.5–18.1 tonnes (5–20 tons) of mussel tissue per hectare were applied as organic fertilizer to grow barley. As a second example, mussel meat was fed to laying hens and resulted in higher egg yield and improved taste. Overall, this effort has led to a test of nutrient trading at the local scale wherein the sewage treatment plant is allowed to trade N cleaning with a mussel farm (Lindahl and Kollberg 2009). Depending on the outcome, shellfish farming

198

Shellfish Aquaculture and the Environment

in some areas, together with nutrient emission trading, may be applied to other areas in various countries. The potential value of mussel farming for alleviating the effects of land-based eutrophication is being explored in the Baltic Sea as well, wherein the “replacement value of nutrient cleaning” by mussel cultures has been estimated using a nonlinear programming model that compares costs and impacts of the mussel farms with other abatement measures such as sewage treatment plants, changes in land use and fertilizer practices, and increased cleaning by households and industries not connected to municipal sewage treatment (Gren et al. 2009). The recently developed cost minimization model considered 20 abatement measures that affect agriculture, industry, transport, and households in 24 basins of the Baltic Sea. Under multiple scenarios, mussel aquaculture was evaluated to be a cost-effective method to alleviate eutrophication, even when mussels from some basins of the Baltic Sea were too small for seafood harvest (Gren et al. 2009). The cost-effectiveness of mussel culture in nutrient removal from the water column depended on mussel growth, sales options, assumptions about mussel farming capacity, and the nutrient load targets (Gren et al. 2009). Estimated marginal cleaning costs of nutrients by mussel aquaculture, calculated as the difference in minimum costs for given nutrient reduction targets with versus without mussel farms as a cleaning option, ranged from 20–138 million euros per year. Mussel culture had a positive value for a large range of nutrient conditions but also varied greatly, from 0.1 to 1.1 billion euros per year. Evaluation of mussel culture as a cleaning device under the Helcom Baltic Sea Action Plan (Helcom 2007) indicated that inclusion of mussel aquaculture could decrease the total abatement cost by ca. 5%, corresponding to a value of 0.22 euro kg−1 live mussel under favorable cost and growth conditions. Moreover, the value from contributions of mussel cul-

tures to savings of control costs for achieving the Baltic Sea Action Plan by Helcom (2007) would range from 5–60% of the market price of live mussels as seafood. The large range for the estimated value of mussel cultures in combating eutrophication underscored the need for more empirical research on mussel growth parameters, nutrient concentrations under different salinity and current conditions, and locations of mussel operations. The options of selling mussels, which is influenced by toxin and pathogen content, would also be important in determining the marginal cleaning cost of nutrient by mussel culture. Possible increase in nutrient regeneration from mussel filtration was not considered in Gren et al.’s (2009) model. The rationale given was that although mussel farming affects biogeochemistry and the benthic ecosystem below the longlines through biodeposits, dropped mussels, and other detritus, the negative effects are known to be localized near the farms, “and have to be judged in relation to the overall positive effects of using mussels to improve coastal ecosystem quality” (Gren et al. 2009, p. 8). Gren et al. emphasized that the focus of their study was to assess the potential for mussel farming as a cost-effective environmental measure in the Baltic Sea, and recommended further work to examine whether the “eventual negative effects on the Baltic ecosystem (from mussel aquaculture) can be kept local and be acceptable.” The authors supported Lindahl and Kollbergs’s (2009) proposed use of mussel farming as a compensation measure for agricultural nutrient emissions in a trade bidding system, and suggested that the utility of mussel farming in combination with nutrient emission trading could also be extended to an international scale. For all such endeavors, it is important that bivalve stocking densities be sufficiently constrained to maintain aerobic conditions in the surficial sediments overlying anaerobic sediments, so that coupled nitrificationdenitrification can occur (Newell 2004). Areas

Bivalve shellfish aquaculture and eutrophication

Net N removal from a ~0.61 hectare (1.5 acre) oyster farm would correspond to the amount of N from untreated wastewater discharge from more than 3000 people, or treated sewage of about 18,000 people

199

Phytoplankton removal 31,000 kg C year–1 Net removal 9.7 tonnes year–1

Detritus removal 84,540 kg C year–1

Net N removal (kg year–1)

Population equivalents 3237 PEQ year–1

ASSETS Chl a O2 Score

Phytoplankton Detritus Excretion Feces Mass balance

INCOME Shellfish farming: Sewage treatment:

2300 k€ year–1 2000 k€ year–1

Total income:

4300 k€ year–1

−4822 –13,151 3745 ⎫ 3545 ⎬ ⎭ –10,683

40% of ingested N returned to ecosystem

PARAMETERS Density of 500 oysters m–3 180-day cultivation period 11 μg L−1 Chl a 3.3 kg N year–1 PEQ

Figure 7.10 Use of the FARM model to assess mass balance of N and the nutrient emissions trading potential of bivalve aquaculture, here, a 1.5-acre oyster farm versus sewage treatment (Modified from Ferreira et al. 2007).

with moderate current flow would continually add oxygenated water to culture areas and help to keep the surface sediments aerobic, while additionally dispersing the biodeposits across a larger bottom area and effectively diluting their oxygen demand in decomposition (Haven and Morales-Alamo 1966; Newell 1994). Nevertheless, bivalve aquaculture holds promise for effective application in mitigating the effects of land-based nutrient pollution with economic benefit. For example, Ferreira et al. (2007) used the FARM model to assess the role of bottom culture of oysters in nutrient removal over about half a year, including an integration analysis of revenue sources. The model indicated that a ∼0.61 hectare (1.5 acre) oyster farm would achieve a net removal of 9.7 tonnes of N per year, equivalent to the amount of N contributed from untreated wastewater discharge of more than 3000 people or the treated sewage of about 18,000 people (Fig. 7.10). As yet there has been no attempt to estimate the value of bivalve aquaculture (specifically, mussel farms) with a replacement cost method to combat eutrophication within a broader context that considers alternative abatement measures, spatial scales, and different nutrient load targets (Gren et al.

2009). Gren et al. (2009) suggested that bivalve aquaculture could be so harnessed similarly as wetlands have been valuated as nutrient sinks, or forests as carbon sinks. The potential utility of shellfish aquaculture combined with seaweed culture for N removal and improved DO were assessed by Miller and Wands (2009) in Long Island Sound. A mechanistic numerical model of eutrophication processes in the Sound, the System Wide Eutrophication Model (SWEM), was modified to include empirical data for filtration of particulate organic nutrients by bivalves and uptake of dissolved inorganic nutrients by cultured marine macroalgae. Model simulations indicated that bivalve mollusc cultures combined with seaweed cultures could increase minimum DO by as much as 2 mg/L, to at least 3.5 mg/L. The SWEM results additionally suggested that bivalve culture and macroalgal harvesting would be more effective than additional reductions in land-based N loadings, beyond the reductions already mandated by an existing total maximum daily load for N, to attain DO standards and provide improved habitat for beneficial aquatic life. There is also strong interest in the potential for cultured shellfish, in polyculture with

200

Shellfish Aquaculture and the Environment

Salmon cultures 300 tonnes in 12 closed production units, each 500 m3; total water flow, 60 m3 min–1 standard high-energy dry feed Outflow 15 tonnes of N (87% dissolved),

Water 2.4 tonnes of P (29% dissolved)

Blue mussel cultures 112.5 tonnes of mussels (wet weight) needed to filter 60 m3 min−1 If all particles were filtered, mussels would retain 25% of the N Outflow 25−30% N released as feces; (total 13.9 tonnes of dissolved N;

Water 40–50% released as dissolved N including 0.9 tonnes from mussels)

Seaweed cultures Closed units; 1000 m3; assume 4% N in dry weight; dry weight 20% of wet weight; Estimated growth rate of 10% day−1 45 tonnes of seaweed needed to take up all dissolved N from salmon and mussel production Figure 7.11 Theoretical model linking production of salmon, blue mussels, and macroalgae. (Based on Bodvin et al. 1996.)

finfish and with macroalgae, to alleviate the nutrient pollution from finfish aquaculture (Folke and Kautsky 1989; Shpigel et al. 1993; Buschmann et al. 1996; Parsons et al. 2002). For example, Bodvin et al. (1996) developed a theoretical model linking the production of salmon, blue mussels, and macroalgae (based on small-scale culture information for the kelp, Laminaria digitata, since data were not available for mass-culture of appropriate seaweeds in systems other than “two-dimensional” shallow basins) (Fig. 7.11). The authors acknowledged that such theoretical models are a modest first step, as some of the critical assumptions require further development. For example, the mussels were assumed to consume all incoming particles in the outflow from salmon aquaculture. Although mussels have been shown to use particulate wastes from fish or shrimp culture as a food resource (Hopkins et al. 1993; Kwei Lin et al. 1993; Shpigel et al. 1993), they cannot be expected to be 100% efficient in removing such wastes. It is doubtful that seaweeds (whatever the species) could be maintained with the assumed uniform growth

in a deep, three-dimensional production system for extended periods, since high light likely would be required, and since light, temperature, and other parameters that vary with depth would affect macroalgal growth. The authors suggested that if the concept can be developed to be commercially viable, an obvious application would be to assist the aquaculture industry along the southern coast of Norway, and that similar systems could be used in sheltered locations of other regions that have access to deeper waters.

Conclusions This chapter addressed two questions: How significant is bivalve shellfish aquaculture in the eutrophication (nutrient pollution, oxygen deficits) of coastal waters, based on present evidence? Conversely, what are the impacts of land-based nutrient pollution and association pollutants on bivalve aquaculture? In response to the first question, four, or ∼7%, of the 62 ecosystems described in the many publica-

Bivalve shellfish aquaculture and eutrophication

tions reviewed here have sustained ecosystemlevel eutrophication from bivalve shellfish aquaculture. These impacts mostly have occurred in poorly flushed systems with extremely high densities cultured shellfish that exceeded the ecological carrying capacity. The remaining 93%, or the great majority, of the ecosystems thus far have sustained negligible or only localized eutrophication effects from bivalve culture. The four exceptions underscore the need to consider ecosystem carrying capacity rather than the carrying capacity for maximal shellfish production to minimize adverse effects. The response to the second question is also clear: Land-based sources of eutrophication have seriously degraded most estuaries and coastal waters throughout the world. Their major, pervasive influence overwhelms the mostly localized impacts that have been documented from bivalve shellfish aquaculture. Bivalve aquaculture is projected to increase significantly during the coming decades (Shumway et al. 2003; Shumway and Kraeuter 2004, Food and Agricultural Organization of the United Nations 2006; Pawłowski et al. accepted). Coastal human population growth, already comprising more than half of the ∼six billion people on the Earth, is increasing in many regions and projected to continue to rapidly increase. Land-based sources of eutrophication are expected to continue to be the clear, dominant force driving eutrophication of most estuarine and coastal marine ecosystems worldwide. The acute, obvious effects of urban and land-based agricultural nutrient pollution and associated pollutants are fish kills and high-biomass algal blooms, but serious, more insidious chronic impacts include long-term shifts in nutrient supplies, large areas of hypoxic and anoxic bottom habitats, loss of beneficial submersed aquatic vegetation, reduction in shellfish recruitment and grazing, and increased shellfish physiological stress, disease, and death. Increasing temperatures from warming trends in climate change can

201

stress shellfish (Philippart et al. 2003), and are expected to interact with nutrient overenrichment and related pollution to weaken shellfish and make them more prone to disease. In summary, relative to land-based pollution sources, bivalve aquaculture has been found to contribute little to eutrophication except in some poorly flushed areas with high shellfish density. Aquaculturists should strive to maintain cultures below ecological carrying capacity to prevent such ecosystem-level adverse effects. Within the constraints of ecosystem carrying capacity, the beneficial effects of bivalve shellfish aquaculture in effectively reducing phytoplankton and the water-column nutrients available for blooms are beginning to be harnessed for economic benefit to offset nutrient overenrichment from land-based sources in coastal zones. In contrast to the generally minimal effects of bivalve aquaculture on eutrophication, major, pervasive nutrient pollution from many urban and agricultural sources is seriously affecting shellfish populations and shellfish aquaculture in many coastal waters of the world, and these impacts are expected to increase with rapidly expanding coastal development. Considering that shellfish aquaculture is vital to meet the seafood demands of the rapidly increasing global human population, there is a pressing need for resource managers and policymakers to increase protection of shellfish aquaculture operations from landbased nutrient pollution.

Literature cited Andrew, M.L., and Frank, L. 2004. Integrated aquaculture system for nutrient reduction in agricultural wastewater: potential and challenges. Bulletin of Fisheries Research Agency (Japan) Suppl. 1:143–152. Arzul, G., Seguel, M., and Clement, A. 2001. Effect of marine animal excretions on differential growth of phytoplankton species. International Council for the Exploration of the Sea (ICES). Journal of Marine Science 58:386–390.

202

Shellfish Aquaculture and the Environment

Asami, H., Aida, M., and Watanabe, K. 2005. Accelerated sulfur cycle in coastal marine sediment beneath areas of intensive shellfish aquaculture. Applied and Environmental Microbiology 71:2925–2933. Asmus, R.M., and Asmus, H. 1991. Mussel beds, limiting or promoting phytoplankton. Journal of Experimental Marine Biology and Ecology 148:215–232. Asmus, H., Asmus, R.M., and Reise, K. 1990. Exchange processes in an intertidal mussel bed: a sylt-flume study in the Wadden Sea. Berichte der Biologische Anstalt Helgoland 6:1–79. Bacher, C., Duarte, P., Ferreira, J.G., Héral, M., and Raillard, O. 1998. Assessment and comparison of the Marennes-Oléron Bay (France) and Carlingford Lough (Ireland) carrying capacity with ecosystem models. Aquatic Ecology 31:379–394. Ball, B., Raine, R., and Douglas, D. 1997. Phytoplankton and particulate matter in Carlingford Lough, Ireland. An assessment of food availability and the impact of bivalve culture. Estuaries 20:430–440. Banas, N.S., Hickey, B.M., Newton, J.A., and Ruesink, J.L. 2007. Tidal exchange, bivalve grazing, and patterns of primary production in Willapa Bay, Washington, USA. Marine Ecology Progress Series 341:123–139. Barber, B.J., and Blake, N.J. 1985. Substrate catabolism related to reproduction in the bay scallop Argopecten irradians concentricus, as determined by O/N and RQ physiological indices. Marine Biology 87:13–18. Barranguet, C. 1997. The role of microphytobenthic primary production in a Mediterranean mussel culture area. Estuarine, Coastal and Shelf Science 44:753–765. Barranguet, C., Alliot, E., and Plante-Cluny, M.-R. 1994. Benthic microphytic activity at two Mediterranean shellfish cultivation sites with reference to benthic fluxes. Oceanologica Acta 17:211–221. Bartoli, M., Nizzoli, D., Viaroli, P., Turolla, E., Castaldelli, G., Fano, E.A., and Rossi, R. 2001. Impact of Ruditapes philippinarum farming on nutrient dynamics and benthic respiration in the Sacca di Goro. Hydrobiologia 455:203–212. Baudinet, D., Alliot, E., Berland, B., Grenz, C., Plante-Cuny, M.R., Plante, R., and Salen-Picard,

C. 1990. Incidence of a mussel culture on biogeochemical fluxes at the sediment water interface. Hydrobiologia 207:187–196. Bayne, B.L., and Hawkins, A.J.S. 1992. Ecological and physiological aspects of herbivory in benthic suspension-feeding molluscs. In: John, D.M., Hawkins, S.J., and Price, J.H. (eds.), PlantAnimal Interactions in the Marine Benthos. Systematics Association, Special Volume No. 46. Clarendon Press, Oxford, UK, pp. 265–288. Bayne, B.L., and Scullard, C. 1977. Rates of nitrogen excretion by species of Mytilus (Bivalvia: Mollusca). Journal of the Marine Biological Association of the United Kingdom 57: 355–369. Bayne, B.L., and Warwick, R.M. (eds.). 1998. Feeding and growth of bivalves: observations and models. Journal of Experimental Marine Biology and Ecology 219(Special Issue):1– 262. Bayne, B.L., Thompson, R.J., and Widdows, J. 1976. Physiology: I. In: Bayne, B.L. (ed.), Marine Mussels: Their Ecology and Physiology. Cambridge University Press, Cambridge, UK, pp. 121–206. Bayne, B.L., Moore, M.N., Widdow, J., Livingstone, D.R., and Salkeld, P. 1979. Measurement of the responses of individuals to environmental stress and pollution: studies with bivalve molluscs. Philosophical Transactions of the Royal Society of London. Series B, Biological Sciences 286:563–581. Beadman, H.A., Kaiser, M.J., Galanidi, M., Shucksmith, R., and Willows, R.I. 2004. Changes in species richness with stocking density of marine bivalves. Journal of Applied Ecology 41:464–475. Bendell-Young, L.I. 2006. Contrasting the community structure and select geochemical characteristics of three intertidal regions in relation to shellfish farming. Environmental Conservation 33:21–27. Bodvin, T., Indergaard, M., Norgaard, E., Jensen, A., and Skaar, A. 1996. Clean technology in aquaculture—a production without waste products? Hydrobiologia 326/327:83–86. Boesch, D.F. 2002. Challenges and opportunities for science in reducing nutrient overenrichment of coastal ecosystems. Estuaries 25: 886–900.

Bivalve shellfish aquaculture and eutrophication

Borja, A., Franco, J., and Pérez, V. 2000. A marine biotic index to establish the ecological quality of soft-bottom benthos within European estuarine and coastal environments. Marine Pollution Bulletin 40:1100–1114. Boucher, G., and Boucher-Rodoni, R. 1988. In situ measurement of respiratory metabolism and nitrogen fluxes at the interface of oyster beds. Marine Ecology Progress Series 44:229–238. Bourget, E., and Messier, D. 1982. Macrobenthic density, biomass, and fauna of intertidal and subtidal sand in a Magdalen Islands lagoon, Gulf of St. Lawrence. Canadian Journal of Zoology 61:2509–2518. Boyd, A.J., and Heasman, K.G. 1998. Shellfish mariculture in the Benguela system: water flow patterns within a mussel farm in Saldanha Bay, South Africa. Journal of Shellfish Research 17:25–32. Breitburg, D.L. 2002. Effects of hypoxia, and the balance between hypoxia and enrichment, on coastal fishes and fisheries. Estuaries 25:767–781. Bricker, S.B., Clement, C.G., Pirhalla, D.E., Orland, S.P., and Farrow, D.G.G. 1999. National Estuarine Eutrophication Assessment: A Summary of Conditions. National Oceanic and Atmospheric Association, Silver Spring, MD. Bricker, S.B., Longstaff, B., Dennison, W., Jones, A., Boicourt, K., Wicks, C., and Woerner, J. 2007. Effects of Nutrient Enrichment in the Nation’s Estuaries: A Decade of Change. NOAA Coastal Ocean Program Decision Analysis Series No. 26. National Centers for Coastal Ocean Service, Silver Spring, MD. Bricker, S.B., Longstaff, B., Dennison, W., Jones, A., Boicourt, K., Wicks, C., and Woerner, J. 2008. Effects of nutrient enrichment in the nation’s estuaries: a decade of change. Harmful Algae 8:21–32. Burkholder, J.M. 1998. Implications of harmful marine microalgae and heterotrophic dinoflagellates in management of sustainable marine fisheries. Ecological Applications 8:S37–S62. Burkholder, J.M. 2001. Eutrophication and oligotrophication. In: Levin, S. (ed.), Encyclopedia of Biodiversity, Vol. 2. Academic Press, New York, pp. 649–670. Burkholder, J.M., Mallin, M.A., Glasgow, H.B., Larsen, L.M., McIver, M.R., Shank, G.C.,

203

Deamer-Melia, N., Briley, D.S., Springer, J., Touchette, B.W., and Hannon, E.K. 1997. Impacts to a coastal river and estuary from rupture of a large swine waste holding lagoon. Journal of Environmental Quality 26:1451–1466. Burkholder, J., Eggleston, D., Glasgow, H., Brownie, C., Reed, R., Melia, G., Kinder, C., Janowitz, G., Corbett, R., Posey, M., Alphin, T., Toms, D., Deamer, N., and Springer, J. 2004. Comparative impacts of two major hurricane seasons on the Neuse River and western Pamlico Sound. Proceedings of the National Academy of Sciences of the United States of America 101:9291– 9296. Burkholder, J.M., Dickey, D.A., Kinder, C., Reed, R.E., Mallin, M.A., Melia, G., McIver, M.R., Cahoon, L.B., Brownie, C., Deamer, N., Springer, J., Glasgow, H., Toms, D., and Smith, J. 2006. Comprehensive trend analysis of nutrients and related variables in a large eutrophic estuary: a decadal study of anthropogenic and climatic influences. Limnology and Oceanography 51:463–487. Burkholder, J.M., Tomasko, D., and Touchette, B.W. 2007. Seagrasses and eutrophication. Journal of Experimental Marine Biology and Ecology 350:46–72. Burkholder, J.M., Glibert, P.M., and Skelton, H.M. 2008. Mixotrophy, a major mode of nutrition for harmful algal species in eutrophic waters. Harmful Algae 8:77–93. Buschmann, A.H., López, D.A., and Medina, A. 1996. A review of environmental effects and alternative production strategies of marine aquaculture in Chile. Aquacultural Engineering 15:397–421. Callier, M.D., Weise, A.M., McKindsey, C.W., and Desrosiers, G. 2006. Sedimentation rates in a suspended mussel farm (Great-Entry Lagoon, Canada): biodeposit production and dispersion. Marine Ecology Progress Series 322:129– 141. Callier, M.D., McKindsey, C.W., and Desrosiers, G. 2007. Multi-scale spatial variations in benthic sediment geochemistry and macrofaunal communities under a suspended mussel culture. Marine Ecology Progress Series 348:103–115. Callier, M.D., McKindsey, C.W., and Desrosiers, G. 2008. Evaluation of indicators used to detect mussel farm influence on the benthos: two case

204

Shellfish Aquaculture and the Environment

studies in the Magdalen Islands, Eastern Canada. Aquaculture 278:77–88. Campbell, D.E., and Newell, C.R. 1998. MUSMOD©, a production model for bottom culture of the blue mussel, Mytilus edulis L. Journal of Experimental Marine Biology and Ecology 219:171–203. Caraco, N. 1995. Influence of human populations on P transfers to aquatic systems: a regional scale study using large rivers. In: Tiessen, H. (ed.), Phosphorus in the Global Environment. SCOPE 54. John Wiley & Sons Ltd., New York, pp. 235–247. Castel, J., Labourge, J.P., Escaravage, V., Auby, I., and Garcia, M.E. 1989. Influence of seagrass and oyster parks on the abundance and biomass patterns of meio- and macrobenthos in tidal flats. Estuarine, Coastal and Shelf Science 28:71–85. Cerco, C.F., and Noel, M.R. 2007. Can oyster restoration reverse cultural eutrophication in Chesapeake Bay? Estuaries and Coasts 30:331–343. Chamberlain, J., Fernandes, T.F., Read, P., Nickell, T.D., and Davies, I.M. 2001. Impacts of biodeposits from suspended mussel (Mytilus edulis L.) culture on the surrounding surficial sediments. International Council for the Exploration of the Sea 58:411–416. Chamberlain, J., Weise, A.M., Grant, J., and Dowd, M. 2006. Modeling the effects of biodeposition from shellfish farms on the near field benthic environment. Modeling Approaches to Assess the Potential Effects of Shellfish Aquaculture on the Marine Environment. Division of Fisheries and Oceans—Canadian Science Advisory Secretariat Research Document 2006/032. Chapelle, A., Menesguen, A., Paoli, J., Souchu, P., Mazouni, N., Vaquer, A., and Millet, B. 2000. Modelling nitrogen, primary production and oxygen in a Mediterranean lagoon. Impact of oysters farming and inputs from the watershed. Ecological Modeling 127:161–181. Chivilev, S., and Ivanov, M. 1997. Response of the Arctic benthic community to excessive amounts of nontoxic organic matter. Marine Pollution Bulletin 35:280–286. Christensen, P.B., Glud, R.N., Dalsgaard, T., and Gillespie, P. 2003. Impacts of longline mussel farming on oxygen and nitrogen dynamics and

biological communities of coastal sediments. Aquaculture 218:567–588. Cloern, J.E. 1982. Does the benthos control phytoplankton biomass in South San Francisco Bay? Marine Ecology Progress Series 9:191– 202. Cloern, J.E. 2001. Our evolving conceptual model of the coastal eutrophication problem. Marine Ecology Progress Series 210:223– 253. Cobelo-García, A., Millward, G.E., Prego, R., and Lukashin, V. 2006. Metal concentrations in Kandalaksha Bay, White Sea (Russia) following the spring snowmelt. Environmental Pollution 1433:89–99. Cockcroft, A.C. 1990. Nitrogen excretion by the surf zone bivalves Donax serra and D. sordidus. Marine Ecology Progress Series 60:57–65. Condon, E.D. 2005. Physiological ecology of the cultured hard clam. Mercenaria mercenaria. M.S. Thesis, College of William and Mary, Gloucester Point, VA, 209pp. Conley, D.J., Kaas, H., Møhlenberg, E., Rasmussen, B., and Wildolf, J. 2000. Characteristics of Danish estuaries. Estuaries 23:848–861. Cranford, P., Dowd, M., Grant, J., Hargrave, B., and McGladdery, S. 2003. Ecosystem level effects of marine bivalve aquaculture. In: Fisheries and Oceans Canada (ed.), In: A Scientific Review of the Potential Environmental Effects of Aquaculture in Aquatic Systems, Vol. 1. Canadian Technical Report of Fisheries and Aquatic Science, Fisheries and Oceans Canada, Burlington, ON, Canada, pp. 51–95. Cranford, P., Anderson, R., Archambault, P., Balch, T., Bates, S., Bugden, G., Callier, M.D., Carver, C., Comeau, L., Hargrave, B., Harrison, G., Horne, E., Kepay, P.E., Li, W.K.W., Mallet, A., Ouellette, M., and Strain, P. 2006. Indicators and Thresholds for Use in Assessing Shellfish Aquaculture Impacts on Fish Habitat. Canadian Science Advisory Secretariat Research Document 2006/034. Department of Fish and Oceans, Ottawa, ON, Canada. Crawford, C.M., Macleod, C.K.A., and Mitchell, I.M. 2003. Effects of shellfish farming on the benthic environment. Aquaculture 224:117– 140. Cromey, C.J., Nickell, T.D., and Black, K.D. 2002. DEPOMOD—modeling the deposition and bio-

Bivalve shellfish aquaculture and eutrophication

logical effects of waste solids from marine cage farms. Aquaculture 214:211–239. Crouzet, P., Leonard, J., Nixon, S., Rees, Y., Parr, W., Laffon, L., Bøgestrand, J., Kristensen, P., Lallana, C., Izzo, G., Bokn, T., and Bak, J. 1999. Nutrients in European ecosystems. Environmental Assessment Report 4. European Environmental Agency, Copenhagen, Denmar. Curtis, K.M., Trainer, V.L., and Shumway, S.E. 2000. Paralytic shellfish toxins in geoduck clams (Panope abrupta): variability, anatomical distribution, and comparison of two toxin detection methods. Journal of Shellfish Research 19: 313–319. D’Amours, O., Archambault, P., McKindsey, C.W., and Robichaud, L. 2008a. The influence of bivalve aquaculture on ecosystem productivity. World Aquaculture Magazine, September:26– 29. D’Amours, O., Archambault, P., McKindsey, C., and Johnson, L.E. 2008b. Local enhancement of epibenthic macrofauna by aquaculture activities. Marine Ecology Progress Series 371:73–84. da Costa, K.G., and Nalesso, R.C. 2006. Effects of mussel farming on macrobenthic community structure in Southeastern Brazil. Aquaculture 258:655–663. Dahlbäck, B., and Gunnarsson, L.A.H. 1981. Sedimentation and sulfate reduction under a mussel culture. Marine Biology 63:269–275. Dame, R.F. 1996. Ecology of Marine Bivalves: An Ecosystem Approach. CRC Press, Boca Raton, FL. Dame, R.F., and Dankers, N. 1988. Uptake and release of materials by a Wadden Sea mussel bed. Journal of Experimental Marine Biology and Ecology 118:207–216. Dame, R., and Libes, S. 1993. Oyster reefs and nutrient retention in tidal creeks. Journal of Experimental Marine Biology and Ecology 171:251–258. Dame, R.F., and Prins, T.C. 1998. Bivalve carrying capacity in coastal ecosystems. Aquatic Ecology 31:409–421. Dame, R.F., Wolaver, T.G., and Libes, S.M. 1985. The summer uptake and release of nitrogen by an intertidal oyster reef. Netherlands Journal of Sea Research 19:265–268. Dame, R., Dankers, N., Prins, T., Jongsma, H., and Smaal, A. 1991. The influence of mussel beds on

205

nutrients in the western Wadden Sea and eastern Scheldt. Estuaries 14:130–138. Dankers, N., and Zuidema, D.R. 1995. The role of the mussel (Mytilus edulis L.) and mussel culture in the Dutch Wadden Sea. Estuaries 18:71–80. Danovaro, R., Gambi, C., Luna, G.M., and Mirto, S. 2004. Sustainable impact of mussel farming in the Adriatic Sea (Mediterranean Sea): evidence from biochemical, microbial and meiofaunal indicators. Marine Pollution Bulletin 49:325–333. De Casabianca, M.-L., Laugier, T., and Collart, D. 1997. Impact of shellfish farming eutrophication on benthic macrophyte communities in the Thau lagoon, France. Aquaculture International 5:301–314. Dealteris, J.T., Kilpatrick, B.D., and Rheault, R.R. 2004. A comparative evaluation of the habitat value of shellfish aquaculture gear, submerged aquatic vegetation and a non-vegetated seabed. Journal of Shellfish Research 23:867–874. Deslous-Paoli, J.-M., Mazouni, N., Souchu, P., Landrein, S., Pichot, P., and Juge, C. 1993. Oyster farming impact on the environment of a Mediterranean lagoon (Thau). Preliminary results of the OXYTHAU program. NATO ASI Series 33:519–520. Deslous-Paoli, J.-M., Souchu, P., Mazouni, N., Juge, C., and Dagault, F. 1998. Relations milieuressources: impact de la conchyliculture sur un environnement lagunaire méditerranéen (Thau). Oceanologica Acta 21:831–843. Doering, P.H. 1989. On the contribution of the benthos to pelagic production. Journal of Marine Research 47:371–383. Doering, P.H., Oviatt, C.A., and Kelly, J.R. 1986. The effects of the filter-feeding clam Mercenaria mercenaria on carbon cycling in experimental marine mesocosms. Journal of Marine Research 44:839–861. Doering, P.H., Kelly, J.R., Oviatt, C.A., and Sowers, T. 1987. Effect of the hard clam Mercenaria mercenaria on benthic fluxes of inorganic nutrients and gases. Marine Biology 94:377–383. Dowd, M. 2003. Seston dynamics in a tidal inlet with shellfish aquaculture: a model study using tracer equations. Estuarine, Coastal Shelf Science 57:523–537. Duarte, P., Meneses, R., Hawkins, A.J.S., Zhu, M., Fang, J., and Grant, J. 2003. Mathematical mod-

206

Shellfish Aquaculture and the Environment

elling to assess the carrying capacity for multispecies culture within coastal waters. Ecological Modelling 168:109–143. Dupuy, C., Pastoureaud, A., Ryckaert, M., Sauriau, P.G., and Montanie, H. 2000. Impact of the oyster Crassostrea gigas on a microbial community in Atlantic coastal ponds near La Rochelle. Aquatic Microbial Ecology 22:227–242. Escaravage, V., Garcia, M.E., and Castel, J. 1989. The distribution of meiofauna and its contribution to detritic pathways in tidal flats (Arcachon Bay, France). Scientia Marina 53:551–559. Everett, R., Ruiz, G., and Carlton, J. 1995. Effect of oyster mariculture on submerged aquatic vegetation: an experimental test in a Pacific Northwest estuary. Marine Ecology Progress Series 125:205–217. Fabi, G., Manoukian, S., and Spagnolo, A. 2009. Impact of an open-sea suspended mussel culture on macrobenthic community (Western Adriatic Sea). Aquaculture 289:54–63. Ferreira, J.G., Hawkins, A.J.S., and Bricker, S.B. 2007. Management of productivity, environmental effects and profitability of shellfish aquaculture—the Farm Aquaculture Resource Management (FARM) model. Aquaculture 264:160–174. Folke, C., and Kautsky, N. 1989. The role of ecosystems for sustainable development of aquaculture. Ambio 18:234–243. Food and Agricultural Organization of the United Nations (FAO). 2006. The State of the World Fisheries and Aquaculture. FAO Fisheries Department, Rome, Italy. Food and Agricultural Organization of the United Nations (FAO). 2008. Fisheries and aquaculture information and statistics service: aquaculture production 1950–2006. FishStat Plus; Rome, Food and Agriculture Organization of the United Nations (www.fao.org/fishery/statistics/ software/fishstat/en). Fréchette, M., Booth, D.A., Myrand, B., and Bermard, H. 1991. Variability and transport of organic seston near a mussel aquaculture site. ICES Marine Science Symposium 192:24–32. Freire, J., Fernandez, L., and Gonzalez-Gurriara, E. 1990. Influence of mussel raft culture on the diet of Liocarcinus arcuatus (Leach) (Brachyura: Portunidae) in the Ria de Arosa (Galicia, NW Spain). Journal of Shellfish Research 9:45–57.

GEOHAB, Global Ecology and Oceanography of Harmful Algal Blooms Programme. 2006. In: Glibert, P. (ed.), Habs in Eutrophic Systems. IOC and SCOR, Paris and Baltimore, p. 74. Gibbs, M.T. 2004. Interactions between bivalve shellfish farms and fishery resources. Aquaculture 240:367–396. Gifford, S., Dunstan, R.H., O’Connor, W., Roberts T., and Toia, R. 2004. Pearl aquaculture—profitable environmental remediation? The Science of the Total Environment 319:27–37. Gilbert, F., Souchu, P., Bianchi, M., and Bonin, P. 1997. Influence of shellfish farming activities on nitrification, nitrate reduction to ammonium and denitrification at the water-sediment interface of the Thau lagoon, France. Marine Ecology Progress Series 151:143–153. Giles, H., Pilditch, C.A., and Bell, D.G. 2006. Sedimentation from mussel (Perna canaliculus) culture in the Firth of Thames, New Zealand: impacts on sediment oxygen and nutrient fluxes. Aquaculture 261:125–140. Giles, H., Broekhuizen, N., Bryan, K.R., and Pilditch, C.A. 2009. Modelling the dispersal of biodeposits from mussel farms: the importance of simulating biodeposit erosion and decay. Aquaculture 291:168–178. Glasoe, S., and Christy, A. 2004. Literature Review and Analysis: Coastal Urbanization and Microbial Contamination of Shellfish Growing Areas. Puget Sound Action Team, Olympia, WA. Glibert, P.M., and Burkholder, J.M. 2006. The complex relationships between increasing fertilization of the Earth, coastal eutrophication, and HAB proliferation. In: Granéli, E., and Turner, J. (eds.), The Ecology of Harmful Algae. Springer-Verlag, New York, pp. 341–354. Glibert, P.M., Harrison, J., Heil, C., and Seitzinger, S. 2006. Escalating worldwide use of urea—a global change contributing to coastal eutrophication. Biogeochemistry 77:441–463. Goldberg, R., and Triplett, T. 1997. Murky Waters: Environmental Effects of Aquaculture in the United States. Environmental Defense Fund, Washington, DC. González-Gurriarán, E. 1986. Seasonal changes of benthic megafauna in the Ria de Muros e Noia (Galicia, North-West Spain): II. Decapod crustaceans (Brachyura). Marine Biology 92:201– 210.

Bivalve shellfish aquaculture and eutrophication

Gouleau, D., Jouanneau, J.M., Weber, O., and Sauriau, P.G. 2000. Short- and long-term sedimentation on Montportail-Brouage intertidal mudflat, Marennes-Oleron Bay (France). Continental Shelf Research 20:1513–1530. Goulletquer, P., Heral, M., Deslous-Paoli, J.M., Prou, J., Garnier, J., Razet, D., and Boromthanarat, W. 1989. Ecophysiologie et bilan énergétique de la palourde japonaise d’élevage Ruditapes philippinarum. Journal of Experimental Marine Biology and Ecology 132:85–108. Grant, J., Hatcher, A., Scott, D.B., Pocklington, P., Schafer, C.T., and Winters, G.V. 1995. A multidisciplinary approach to evaluating impacts of shellfish aquaculture on benthic communities. Estuaries 18:124–144. Gren, I.-M., Lindahl, O., and Lindqvist, M. 2009. Values of mussel farming for combating eutrophication. Ecological Engineering 35:935–945. Grenz, C. 1989. Quantification et de la Biodeposition en Zones de Production Conchylicole Intensive en Mediterranee. Ph.D. thesis, Universite d’Aix- Marseille II, 144pp. Grenz, C., Hermin, M., Baudinet, D., and Daumas, R. 1990. In situ biochemical and bacterial variation of sediments enriched with mussel biodeposits. Hydrobiologia 207:153–160. Grenz, C., Plante-Cuny, M.R., Plante, R., Alliot, E., Baudinet, D., and Berland, B. 1991. Measurements of benthic nutrient fluxes in Mediterranean shellfish farms: a methodological approach. Oceanologica Acta 14:195–201. Haamer, J. 1996. Improving water quality in a eutrophied fjord system with mussel farming. Ambio 25:356–362. Hammen, C.S. 1968. Aminotransferase activities and amino acid excretion of bivalve molluscs and brachiopods. Comparative Biochemistry and Physiology 26:697–705. Hammen, C.S., Miller, H.F., Jr., and Geer, W.H. 1966. Nitrogen excretion of Crassostrea virginica. Comparative Biochemistry and Physiology 17:1199–1200. Hart, R. 2003. Dynamic pollution control. Ecological Economics 47:79–93. Hartstein, N.D., and Rowden, A.A. 2004. Effect of biodeposits from mussel culture on macroinvertebrate assemblages at sites of different hydrodynamic regime. Marine Environmental Research 57:339–357.

207

Hartstein, N.D., and Stevens, C.L. 2005. Deposition beneath long-line mussel farms. Aquacultural Engineering 33:192–213. Hatcher, A., Grant, J., and Schofield, B. 1994. Effects of suspended mussel culture (Mytilus spp.) on sedimentation, benthic respiration and sediment nutrient dynamics in a coastal bay. Marine Ecology Progress Series 115:219–235. Haven, D.S., and Morales-Alamo, R. 1966. Aspects of biodeposition by oysters and other invertebrate filter feeders. Limnology and Oceanography 11:487–498. Heasman, K.G.M., Pitcher, G.C., McQuaid, C.D., and Hecht, T. 1998. Shellfish mariculture in the Benguela system: raft culture of Mytilus galloprovincialis and the effect of rope spacing on food extraction, growth rate, production, and condition of mussels. Journal of Shellfish Research 17:33–39. Heisler, J., Glibert, P., Burkholder, J., Anderson, D., Cochlan, W., Dennison, W., Gobler, C., Dortch, Q., Heil, C., Humphries, E., Lewitus, A., Magnien, R., Marshall, H., Stockwell, D., and Suddleson, M. 2008. Eutrophication and harmful algal blooms: a scientific consensus. Harmful Algae 8:3–13. Helcom. 2007. An Approach to Set Country-Wise Nutrient Reduction Allocations to Reach Good Marine Environment of the Baltic Sea. Helcom BSAP Eutro Expo/2007. Helsinki Commission, Helsinki, Finland. Henderson, A., Gamito, S., Karakassis, I., Pederson, P., and Smaal, A. 2001. Use of hydrodynamic and benthic models for managing environmental impacts of marine aquaculture. Journal of Applied Ichthyology 17:163–172. Henriksen, K., Rasmussen, M.B., and Jensen, A. 1983. Effect of bioturbation on microbial nitrogen transformations in the sediment and fluxes of ammonium and nitrate to the overlaying water. Environmental Biogeochemistry 35:193– 205. Héral, M. 1993. Why carrying capacity models are useful tools for management of bivalve culture. In: Dam, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Coastal Ecosystem Processes. Springer Verlag, Heidelberg, Germany, pp. 455–477. Herman, P.M., and Scholten, H. 1990. Can suspension feeders stabilize estuarine ecosystems? In:

208

Shellfish Aquaculture and the Environment

Gibson, R.N. (ed.), Trophic Relationships in the Marine Environment. Aberdeen University Press, Aberdeen, Scotland, pp. 104–116. Hilborn, R., Armstrong, D., Friedman, C., Naish, K., Orensanz, J., Ruesink, J., Vadopalas, B., Feldman, K., Valero, J., Cheney, D., Suhrbier, A., Christy, A., and Davis, J. 2004. Comprehensive literature review and synopsis of issues relating to geoduck (Panopea abrupta) ecology and aquaculture production. Draft of Deliverable, January 12. Prepared for the Washington State Department of Natural Resources, Olympia, WA, 123pp. Hopkins, J.S., Hamilton, R.D., II, Sandifer, P.A., and Browdy, C.L. 1993. The production of bivalve molluscs in intensive shrimp ponds and their effect on shrimp production and water quality. World Aquaculture 24:74–77. Howarth, R.W. 2008. Coastal nitrogen pollution: a review of sources and trends globally and regionally. Harmful Algae 8:14–20. Howarth, R.W., Boyer, E.W., Pabich, W.J., and Galloway, J.N. 2002. Nitrogen use in the United States from 1961–2000 and potential future trends. Ambio 31:88–96. Iglesias, J. 1981. Spatial and temporal changes in the demersal fish community of the Ria de Arosa (NW Spain). Marine Biology 65:199–208. Inglis, G.J., and Gust, N. 2003. Potential indirect effects of shellfish culture on the reproductive success of benthic predators. Journal of Applied Ecology 40:1077–1089. Inglis, G.J., Hayden, B.J., and Ross, A.H. 2000. An overview of factors affecting the carrying capacity of coastal embayments for mussel culture. Report NIWA (National Institute of Water and Atmospheric Research, Ltd.), New Zealand. Ito, S., and Imai, T. 1955. Ecology of oyster bed I. On the decline of productivity due to repeated cultures. Tohoku Journal of Agricultural Research 4:9–26. Jamieson, G.S., Chew, L., Gillespie, G., Robinson, A., Bendell-Young, L., Heath, W., Bravender, B., Tompkins, A., Nishimura, D., and Doucette, P. 2001. Phase 0 review of the environmental impacts of intertidal shellfish aquaculture in Baynes Sound. Canadian Science Advisory Secretariat Research Document 2001/125, ISSN 1480-4883, Canada (www document). www. dfo-mpo.gc.ca/csas/

Jaramillo, E., Beltran, C., and Bravo, A. 1992. Mussel biodeposition in an estuary in southern Chile. Marine Ecology Progress Series 82:85– 94. Jie, H., Zhang, Z., Zishan, Y., and Widdows, J. 2001. Differences in the benthic-pelagic particle flux (biodeposition and sediment erosion) at intertidal sites with and without clam (Ruditapes philippinarum) cultivation in Eastern China. Journal of Experimental Marine Biology and Ecology 261:245–261. Jordan, T.E., and Valiela, I. 1982. A nitrogen budget of the ribbed mussel, Geukensia demissa, and its significance in nitrogen flow in a New England salt marsh. Limnology and Oceanography 27:75–90. Kaiser, M.J., Spencer, B.E., and Edwards, D.B. 1996. Infaunal community changes as a result of commercial clam cultivation and harvesting. Aquatic Living Resources 9:57–63. Kaiser, M.J., Laing, I., Utting, S.D., and Burnell, G.M. 1998. Environmental impacts of bivalve mariculture. Journal of Shellfish Research 17:59–66. Kaspar, H.F., Gillespie, P.A., Boyer, I.C., and MacKenzie, A.L. 1985. Effects of mussel aquaculture on the nitrogen cycle and benthic communities of Kenepuru Sound, Marlborough Sounds, New Zealand. Marine Biology 85: 127–136. Kreeger, D.A., and Newell, R.I.E. 2001. Seasonal utilization of different seston carbon sources by the ribbed mussel, Geukensia demissa (Dillwyn) in a mid-Atlantic salt marsh. Journal of Experimental Marine Biology and Ecology 260:71–91. Krom, M.D., and Berner, R.A. 1981. The diagenesis of phosphorus in a nearshore marine sediment. Geochimica et Cosmochimica Acta 45:207–216. Kwei Lin, C., Ruamthaveesub, P., and Wanuchsoontom, P. 1993. Integrated culture of the green mussel (Perna viridis) in waste water from an intensive shrimp pond: concept and practice. World Aquaculture 24:68–73. La Rosa, T., Mirto, S., Favaloro, E., Savona, B., Sara, G., Danovaro, R., and Mazzola, A. 2002. Impact on the water column biogeochemistry of a Mediterranean mussel and fish farm. Water Research 36:713–721. Labarta, U., Fernandez-Reiniz, M.J., and Babarro, J.M.F. 1997. Differences in physiological ener-

Bivalve shellfish aquaculture and eutrophication

getics between intertidal and raft cultivated mussels Mytilus galloprovincialis. Marine Ecology Progress Series 152:167–173. Langton, R.W., Haines, K.C., and Lyon, R.E. 1977. Ammonia-nitrogen production by the bivalve mollusc Tapes japonica and its recovery by the red seaweed Hypnea musciformis in a tropical mariculture system. Helgolander Wissenschaftliche Meeresuntersuchungen 30: 217–229. Leguerrier, D., Niquil, N., Petiau, A., and Bodoy, A. 2004. Modeling the impact of oyster culture on a mudflat food web in Marennes-Oleron Bay (France). Marine Ecology Progress Series 273:147–162. Lindahl, O., and Kollberg, S. 2009. Can the EU agri-environmental aid program be extended into the coastal zone to combat eutrophication? Hydrobiologia 629:59–64. Lindahl, O., Hart, R., and Hernoth, B. 2005. Improving marine water quality by mussel farming: a profitable solution for Swedish society. Ambio 34:131–138. Livingstone, D.R., Widdows, J., and Fieth, P. 1979. Aspects of nitrogen metabolism of the common mussel Mytilus edulis: adaptation to abrupt and fluctuating changes in salinity. Marine Biology 53:41–55. López-Jamar, E., Iglesias, J., and Otero, J.J. 1984. Contribution of infauna and mussel-raft epifauna to demersal fish diets. Marine Ecology Progress Series 15:13–18. Lu, L., and Grant, J. 2008. Recolonization of intertidal infauna in relation to organic deposition at an oyster farm in Atlantic Canada—a field experiment. Estuaries and Coasts 31:767–775. Lu, Y., Blake, N.J., and Torres, J.J. 1999. Oxygen consumption and ammonia excretion of larvae and juveniles of the bay scallop Argopecten irradians concentricus (Say). Journal of Shellfish Research 18:419–424. Lucas, M.I., Newell, R.C., Shumway, S.E., Seiderer, L.J., and Bally, R. 1987. Particle clearance and yield in relation to bacterioplankton and suspended particulate availability in estuarine and open coast populations of the mussel Mytilus edulis. Marine Ecology Progress Series 36: 215–224. Luckenbach, M.W., and Harry, V. 2004. Linking watershed loading and basin-level carrying capacity models to evaluate the effects of land

209

use on primary production and shellfish aquaculture. Bulletin of the Fisheries Research Agency (Japan) Suppl. 1:123–132. Luckenbach, M.W., and Wang, H.V. 2004. Linking watershed loading and basin-level carrying capacity models to evaluate the effects of land use on primary production and shellfish aquaculture. Bulletin of the Fisheries Research Agency Suppl. 1:123–132. Lum, S.C., and Hammen, C.S. 1964. Ammonia excretion of Lingula. Comparative Biochemistry and Physiology 12:185–190. Magni, P., Montani, S., Takada, C., and Tsutsumi, H. 2000. Temporal scaling and relevance of bivalve nutrient excretion on a tidal flat of the Seto Inland Sea, Japan. Marine Ecology Progress Series 198:139–155. Mallet, A.O., Carver, C.E., and Landry, T. 2006. Impact of suspended and off-bottom Eastern oyster culture on the benthic environment in eastern Canada. Aquaculture 255:362–373. Mallin, M.A., Williams, K.E., Esham, E.C., and Lowe, R.P. 2000. Effect of human development on bacteriological water quality in coastal watersheds. Ecological Applications 10:1047– 1056. Mann, R. 1979. Some biochemical and physiological aspects of growth and gametogenesis in Crassostrea gigas and Ostrea edulis grown at sustained elevated temperatures. Journal of the Marine Biological Association of the United Kingdom 59:95–110. Mann, R., and Glomb, S. 1978. The effect of temperature on growth and ammonia excretion of the Manila clam Tapes japonica. Estuarine and Coastal Marine Science 6:335–339. Mao, Y., Zhou, Y., Yang, H., and Wang, R. 2006. Seasonal variation in metabolism of cultured Pacific oyster, Crassostrea gigas, in Sanggou Bay, China. Aquaculture 253:322–333. Marino, J., Pérez, A., and Roma, G. 1982. The mussel culture Mytilus edulis in the Ría de Arosa Northwestern Spain. Boletin Instituto Español de Oceanografia 7:297–308. Marinov, D., Galbiati, L., Giordani, G., Viaroji, P., Norro, A., Bencivelli, S., and Zaldivar, J.M. 2007. An integrated modelling approach for the management of clam farming in coastal lagoons. Aquaculture 269:306–320. Mariojouls, C., and Sornin, J.-M. 1987. Sur exploitation et détérioration de la qualité des terrains

210

Shellfish Aquaculture and the Environment

conchylicoles: conséquences sur les systèmes d’exploitation—exemples en France et Japon. Norois 34:51–61. Martin, J.L.M., Sornin, J.-M., Marchand, M., Depauw, N., and Joyce, J. 1991. The significance of oyster biodeposition in concentrating organic matter and contaminants in the sediment. In: De Pauw, N., and Joyce, J. (eds.), Aquaculture and the Environment. Reviews of the International Conference Aquaculture Europe ’91, Dublin, Ireland, June 10–12, 1991. Special Publication of the European Aquaculture Society, Vol. 14. Ostend, Belgium, p. 207 (abstract). Mattsson, J., and Lindén, O. 1983. Benthic macrofauna succession under mussels, Mytilus edulis L. (Bivalvia), cultured on hanging long-lines. Sarsia 68:97–102. Maurer, D., Nguyen, H., Robertson, G., and Gerlinger, T. 1999. The Infaunal Trophic Index (ITI): Its suitability for marine environmental monitoring. Ecological Applications 9:699– 713. Mazouni, N. 2004. Influence of suspended oyster cultures on nitrogen regeneration in a coastal lagoon (Thau, France). Marine Ecology Progress Series 276:103–113. Mazouni, N., Gaertner, J.C., Deslous-Paoli, J.M., Landrein, S., and Geringer d’Oedenberg, M. 1996. Nutrient and oxygen exchanges at the water-sediment interface in a shellfish farming lagoon (Thau, France). Journal of Experimental Marine Biology and Ecology 203:92–113. Mazouni, N., Gaertner, J.C., and Deslous-Paoli, J.M. 1998. Influence of oyster culture on water column characteristics in a coastal lagoon (Thau, France). Hydrobiologia 373–374:149–156. Mazouni, N., Gaertner, J.-C., and Deslous-Paoli, J.-M. 2001. Composition of biofouling communities on suspended oyster cultures: an in situ study of their interactions with the water column. Marine Ecology Progress Series 214:93–102. McComb, A.J. (ed.). 1995. Eutrophic Shallow Estuaries Lagoons. CRC Press, Boca Raton, FL. McGranahan, G., Balk, D., and Anderson, B. 2007. The rising tide: assessing the risks of climate change and human settlements in low elevation coastal zones. Environment and Urbanization 19:17–37. McKindsey, C.W., Anderson, M.R., Barnes, P., Courtenay, S., Landry, T., and Skinner, M. 2006.

Effects of Shellfish Aquaculture on Fish Habitat. Department of Fisheries and Oceans (DFO)— Canadian Science Advisory Secretariat Research Document 2006/011, Fisheries and Oceans Canada, Ottawa, ON, Canada. Meeuwig, J.J., Rasmussen, J.B., and Peters, R.H. 1998. Turbid waters and clarifying mussels: their moderation of empirical chl : nutrient relations in estuaries in Prince Edward Island, Canada. Marine Ecology Progress Series 171:139–150. Mesnage, V., Ogier, S., Bally, G., Disnar, J.R., Lottier, N., Dedieu, K., Rabouille, C., and Copard, Y. 2007. Nutrient dynamics at the sediment-water interface in a Mediterranean lagoon (Thau, France): influence of biodeposition by shellfish farming activities. Marine Environmental Research 63:257–277. Metzger, E., Simonucci, C., Viollier, E., Sarazin, G., Prevot, F., and Jezequel, D. 2007. Benthic response to shellfish farming in Thau lagoon: pore water signature. Estuarine, Coastal and Shelf Science 72:406–419. Miller, R.E., and Wands, J.R. 2009. Applying the System Wide Eutrophication Model (SWEM) for A Preliminary Quantitative Evaluation of Biomass Harvesting As A Nutrient Control Strategy for Long Island Sound. Hydroqual, Inc., Mahwah, NJ. Miron, G., Landry, T., Archambault, P., and Frenette, B. 2005. Effects of mussel culture husbandry practices on various benthic characteristics. Aquaculture 250:138–154. Mirto, S., La Rosa, T., Danavaro, R., and Mazzola, A. 2000. Microbial and meiofaunal response to intensive mussel-farm biodeposition in coastal sediments of the western Mediterranean. Marine Pollution Bulletin 40:244–252. Mojica, R., Jr., and Nelson, W.G. 1993. Environmental effects of a hard clam (Mercenaria mercenaria) aquaculture site in the Indian River Lagoon, Florida. Aquaculture 113:313– 329. Munroe, D., and McKinley, R.S. 2007. Commercial Manila clam (Tapes philippinarum) culture in British Columbia, Canada: the effects of predator netting on intertidal sediment characteristics. Estuarine, Coastal and Shelf Science 72: 319–328. Murdoch, R., and Oliver, M. 1995. Study of Chlorophyll Concentrations Within and Around

Bivalve shellfish aquaculture and eutrophication

Mussel Farms: Beatrix Bay, Pelorus Sound. 1995/6-WN, National Institute of Water and Atmospheric Research, Wellington, New Zealand. National Research Council (NRC). 2000. Clean Coastal Waters—Understanding and Reducing the Effects of Nutrient Pollution. National Academy Press, Washington, DC. Navarro, J. 1988. The effects of salinity on the physiological ecology of Choromytilus chorus Molina, 1782. Bivalvia: Mytilidae. Journal of Experimental Marine Biology and Ecology 122:19–33. Navarro, J.M., and Gonzalez, C.M. 1998. Physiological responses of the Chilean scallop Argopecten purpuratus to decreasing salinities. Aquaculture 167:315–327. Navarro, E., Iglesias, J.I.P., Camacho, A.P., Labarta, U., and Beiras, R. 1991. The physiological energetics of mussels (Mytilus galloprovincialis Lmk) from different cultivation rafts in the Ría de Arosa (Galicia, N.W. Spain). Aquaculture 94: 197–212. Naylor, R.L., Goldberg, R.J., Mooney, H., Beveridge, M.C., Clay, J., Folk, C., Kautsky, N., Lubchenco, J., Primavera, J., and Williams, M. 1998. Nature’s subsidies to shrimp and salmon farming. Nature 282:883–884. Naylor, R.L., Goldberg, R.J., Primavera, J.H., Kautsky, N., Beveridge, M.C., Clay, J., Folk, C., Lubchenco, J., Mooney, H., and Troell, M. 2000. Effect of aquaculture on world fish supplies. Nature 405:1017–1024. Newell, R.I.E. 2004. Ecosystem influences of natural and cultivated populations of suspension-feeding bivalve molluscs: a review. Journal of Shellfish Research 23:51–61. Newell, R.I.E., and Jordan, S.J. 1983. Preferential ingestion of organic material by the American oyster, Crassostrea virginica. Marine Ecology Progress Series 13:47–53. Newell, C.R., and Shumway, S.E. 1993. Grazing of natural particulates by bivalve molluscs: a spatial and temporal perspective. In: Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Coastal Ecosystem Processes. Springer, Berlin/Heidelberg, Germany, pp. 85–148. Newell, R.I.E., Cornwell, J.C., and Owens, M.S. 2002. Influence of simulated bivalve biodeposition and microphytobenthos on sediment nitro-

211

gen dynamics: a laboratory study. Limnology and Oceanography 47:1367–1379. Newell, R.I.E., Fisher, T.R., Holyoke, R.R., and Cornwell, J.C. 2005. Influence of eastern oysters on nitrogen and phosphorus regeneration in Chesapeake Bay. In: Dame, R., and Olenin, S. (eds.), The Comparative Roles of Suspension Feeders in Ecosystems. 47, NATO Science Series: IV—Earth and Environmental Sciences. Springer, Dordrecht, The Netherlands, pp. 93–120. Nicholls, R.J., and Small, C. 2002. Improved estimates of coastal populations and exposure to hazards released. EOS 83:301–307. Niquil, N., Pouvreau, S., Sakka, A., Legendre, L., Addessi, L., Borgne, R.L., Charpy, L., and Delesalle, B. 2001. Trophic web and carrying capacity in a pearl oyster farming lagoon (Takapoto, French Polynesia). Aquatic Living Resources 14:165–174. Nixon, S.W., Oviatt, C.A., Garber, J., and Lee, V. 1976. Die1 metabolism and nutrient dynamics in a salt marsh embayment. Ecology 57: 740–750. Nizzoli, D., Welsh, D.T., Bartoli, M., and Viaroli, P. 2005. Impacts of mussel (Mytilus galloprovincialis) farming on oxygen consumption and nutrient recycling in a eutrophic coastal lagoon. Hydrobiologia 550:183–198. Nizzoli, D., Bartoli, M., and Viaroli, P. 2006a. Nitrogen and phosphorous budgets during a farming cycle of the Manila clam Ruditapes philippinarum: an in situ experiment. Aquaculture 261:98–108. Nizzoli, D., Welsh, D.T., Fano, E.A., and Viaroli, P. 2006b. Impact of clam and mussel farming on benthic metabolism and nitrogen cycling, with emphasis on nitrate reduction pathways. Marine Ecology Progress Series 315:151–165. Nizzoli, D., Bartoli, M., and Viaroli, P. 2007. Oxygen and ammonium dynamics during a farming cycle of the bivalve Tapes philippinarum. Hydrobiologia 587:25–36. Nugues, M.M., Kaiser, M.J., Spencer, B.E., and Edwards, D.B. 1996. Benthic community changes associated with intertidal oyster cultivation. Aquaculture Research 27:913–924. Ogilvie, S.C., Ross, A.H., and Schiel, D.R. 2000. Phytoplankton biomass associated with mussel farms in Beatrix Bay, New Zealand. Aquaculture 181:71–80.

212

Shellfish Aquaculture and the Environment

Ottmann, F., and Sornin, J.-M. 1985. Observations on sediment accumulation as a result of mollusc culture systems in France. In: Labish Chao, N., and Kirby-Smith, W. (eds.), Proceedings of the International Symposium on Utilization of Coastal Ecosystems: Planning, Pollution, and Productivity, Vol. 1. University of Rio Grande do Sul, Brazil, pp. 329–337. Páez-Osuna, F., Guerrero-Galván, S.R., and RuizFernández, A.C. 1998. The environmental impact of shrimp aquaculture and the coastal pollution in Mexico. Marine Pollution Bulletin 36:65–75. Parsons, G.J., Shumway, S.E., Kuenstner, S., and Gryska, A. 2002. Polyculture of sea scallops (Placopecten magellanicus) suspended from salmon cages. Aquaculture International 10: 65–77. Paterson, K.J., Schreider, M.J., and Zimmerman, K.D. 2003. Anthropogenic effects on seston quality and quantity and the growth and survival of Sydney rock oyster (Saccostrea glomerata) in two estuaries in NSW, Australia. Aquaculture 221:407–426. Paul, M.J., and Meyer, J.L. 2001. Streams in the urban landscape. Annual Review of Ecology and Systematics 32:333–365. Pawlowski, M., Bouwman, L., Beusen, A., and Overbeek, C. Past and future nitrogen and phosphorus balances and feed use in global aquaculture: I. Shellfish and aquatic plants. Reviews in Fisheries Science. Pérez-Camacho, A., Gonzalez, R., and Fuentes, J. 1991. Mussel culture in Galicia (N.W. Spain). Aquaculture 94:263–278. Pfeiffer, T.J., Lawson, T.B., and Rusch, K.A. 1999. Northern quahog, Mercenaria mercenaria, seed clam waste characterization study: precursor to a recirculating culture system design. Aquacultural Engineering 20:149–161. Philippart, C.J.M., van Aken, H.M., Beukema, J.J., Bos, O.G., Cadee, G.C., and Dekker, R. 2003. Climate-related changes in recruitment of the bivalve Macoma balthica. Limnology and Oceanography 48:2171–2185. Pietros, J.M., and Rice, M.A. 2003. The impacts of aquacultured oysters, Crassostrea virginica (Gmelin, 1791) on water column nitrogen and sedimentation: results of a mesocosm study. Aquaculture 220:407–422.

Pilditch, C.A., Grant, J., and Bryan, K.R. 2001. Seston supply to sea scallops (Placopecten magellanicus) in suspended culture. Canadian Journal of Fisheries and Aquatic Science 58:241–253. Pitcher, G.C., and Calder, D. 1998. Shellfish culture in the Benguela system: phytoplankton and the availability of food for commercial mussel farms in Saldhana Bay, South Africa. Journal of Shellfish Research 17:15–24. Plew, D.R., Stevens, C.L., Spigel, R.H., and Hartstein, N.D. 2005. Hydrodynamic implications of large offshore mussel farms. IEEE Journal of Oceanic Engineering 31:95– 108. Prins, T.C., and Smaal, C.S. 1994. The role of the blue mussel Mytilus edulis in the cycling of nutrients in the Oosterschelde estuary (The Netherlands). Hydrobiologia 282/283:413– 429. Prins, T.C., Escaravage, V., Smaal, A.C., and Peeters, J.C.H. 1995. Nutrient cycling and phytoplankton dynamics in relation to mussel grazing in a mesocosm experiment. Ophelia 41:289–315. Prins, T.C., Smaal, C.S., and Dame, R.F. 1998. A review of the feedbacks between bivalve grazing and ecosystem processes. Aquatic Ecology 31:349–359. Prosch, R.M., and McLachlan, A. 1984. The regeneration of surf-zone nutrients by the sand mussel, Donax serra Röding. Journal of Experimental Marine Biology and Ecology l80:221–233. Raillard, O., and Ménesguen, A. 1994. An ecosystem model for estimating the carrying capacity of a macrotidal shellfish system. Marine Ecology Progress Series 115:117–130. Reitan, K.I., Oeie, G., Olsen, Y., and Reinerstsen, H. 1999. Effect of increased primary production in a fjord on growth of blue mussels and scallops. Journal of Shellfish Research 18:726. (abstract). Rice, M.A. 2001. Environmental impacts of shellfish aquaculture: filter feeding to control eutrophication. In: Tlusty, M., Bengtson, D., Halvorson, H.O., Oktay, S., Pearce, J., and Rheault, R.B. Jr. Marine Aquaculture and the Environment: A Meeting for Stakeholders in the Northeast. Cape Cod Press, Falmouth, MA, pp. 76–86.

Bivalve shellfish aquaculture and eutrophication

Richard, L. 2004. Balancing marine aquaculture inputs and extraction: combined culture of finfish and bivalve molluscs in the open ocean. Bulletin of Fisheries Research Agency (Japan) Suppl. 1:51–58. Romero, P., González-Gurriarán, E., and Penas, E. 1982. Influence of mussel rafts on spatial and seasonal abundance of crabs in the Ría de Arosa, North-West Spain. Marine Biology 72:201– 210. Rosenberg, R., and Loo, L.-O. 1983. Energy flow in a Mytilus edulis culture in western Sweden. Aquaculture 35:151–161. Ryther, J.H. 1954. The ecology of phytoplankton blooms in Moriches Bay and Great South Bay, Long Island, New York. Biological Bulletin 106:198–209. Sauriau, P.G., Mouret, V., and Rincé, J.-P. 1989. Organisation trophique de la malacofaune benthique non cultivée du bassin ostréicole de Marennes-Oléron. Oceanologica Acta 12:193– 204. Schlüter, L., and Josefsen, S.B. 1994. Annual variation in condition, respiration and remineralisation of Mytilus edulis L. in the Sound, Denmark. Helgolander Meeresunters 48:419–430. Scholten, H., and Smaal, A.C. 1999. The ecophysiological response of mussels (Mytilus edulis) in mesocosms to a range of inorganic nutrient loads: simulations with the model EMMY. Aquatic Ecology 33:83–100. Scott, G.I., Fulton, M.H., Strozier, E.D., Key, P.B., Daugomah, J.W., Porter, D., and Strozier, S. 1996. The effects of urbanization on the American oyster, Crassostrea virginica (Gmelin). Journal of Shellfish Research 15:523– 524. Shaw, K.R. 1998. Prince Edward Island benthic survey. Technical Report of Environmental Science, Vol. 4. Department of Fisheries and Environment, Montague, Prince Edward Island, Canada, 94pp. . Shpigel, M., Neori, A., Popper, D.M., and Giordin, H. 1993. A proposed model for “environmentally clean” land-based culture of fish, bivalves and seaweeds. Aquaculture 117:115– 128. Shumway, S.E. 1990. A review of the effects of algal blooms on shellfish and aquaculture. Journal of the World Aquaculture Society 21:65–104.

213

Shumway, S.E., and Kraeuter, J.N. (eds.). 2004. Molluscan shellfish research and management: charting a course for the future. Final Proceedings from the Cooperative research and Information Institute (CRII) Workshop, Charleston, SC, January 2000, 156pp. Shumway, S.E., Davis, C., Downey, R., Karney, R., Kraeuter, J., Parsons, J., Rheault, R., and Wikfors, G. 2003. Shellfish aquaculture—in praise of sustainable economies and environments. World Aquaculture 34:15–17. Sloth, N.P., Blackburn, T.H., Hansen, L.S., Risgaard-Petersen, N., and Lomstein, B.A. 1995. Nitrogen cycling in sediments with different organic loadings. Marine Ecology Progress Series 116:163–170. Smaal, A., and Prins, T.C. 1993. The uptake of organic matter and the release of inorganic nutrients by bivalve suspension feeder beds. In: Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Coastal Ecosystem Processes. Springer-Verlag, Heidelberg, Germany, pp. 273–298. Smaal, A.C., and Zurburg, W. 1997. The uptake and release of suspended and dissolved material by oysters and mussels in Marennes-Oléron Bay. Aquatic Living Resources 10:23–30. Smaal, A.C., Vonck, A.P.M.A., and Bakker, M. 1997. Seasonal variation in physiological energetics of Mytilus edulis and Cerastoderma edule of different size classes. Journal of the Marine Biological Association of the United Kingdom 77:817–838. Smaal, A., van Stralen, M., and Schuiling, E. 2001. The interaction between shellfish culture and ecosystem processes. Canadian Journal of Fisheries and Aquatic Sciences 58:991–1002. Smayda, T.J. 1989. Primary production and the global epidemic of phytoplankton blooms in the sea: a linkage? In: Cosper, E.M., Bricelj, V.M., and Carpenter, E.J. (eds.), Coastal and Estuarine Studies No. 35. Novel Phytoplankton Blooms. Springer-Verlag, New York, pp. 449–484. Smith, J., and Shackley, S.E. 2004. Effects of a commercial mussel Mytilus edulis lay on a sublittoral, soft sediment benthic community. Marine Ecology Progress Series 282:185–191. Snra, R.F., and Baggaley, A. 1976. Rate of excretion of ammonia by the hard clam Mercenaria

214

Shellfish Aquaculture and the Environment

mercenaria and the American oyster Crassostrea virginica. Marine Biology 36:251–258. Songsangjinda, P., Matsuda, O., Yamamoto, T., Rajendran, N., and Maeda, H. 2000. The role of suspended oyster culture on nitrogen cycle in Hiroshima Bay. Journal of Oceanography 56:223–231. Sornin, J.-M., Feuillet, M., Héral, M., and DeslousPaoli, J.-M. 1983. Effets des biodépôts de l’huître Crassostrea gigas (Thunberg) sur l’accumulation de matières organiques dans les parcs du bassin de Marennes-Oléron. Journal of Molluscan Studies Supplement 12A:185–197. Sornin, J.-M., Feulillet, M., Héral, M., and Fardeu, J.C. 1986. Influence descultures D’huitres Crassostrea gigas sur le cycle du phosphore en zone intertidale: role de la biodeposition. Oceanologica Acta 9:313–322. Sorokin, I.I., Giovanardi, O., Pranovi, F., and Sorokin, P.I. 1999. Need for restricting bivalve culture in the southern basin of the Lagoon of Venice. Hydrobiologia 400:141–148. Souchu, P., Vaquer, A., Collos, Y., Landrein, S., Deslous-Paoli, J.M., and Bibent, B. 2001. Influence of shellfish farming activities on the biogeochemical composition of the water column in Thau lagoon. Marine Ecology Progress Series 218:141–152. Spencer, B.E., Kaiser, M.J., and Edwards, D.B. 1996. The effect of Manila clam cultivation on an intertidal benthic community: the early cultivation phase. Aquaculture Research 27:261– 276. Spencer, B.E., Kaiser, M.J., and Edwards, D.B. 1998. Ecological effects of intertidal Manila clam cultivation: observations at the end of the cultivation phase. Journal of Applied Ecology 34:444–452. Srna, R., and Baggaley, A. 1976. Rate of excretion of ammonia by the hard clam Mercenaria mercenaria and the American oyster Crassostrea virginica. Marine Biology 36:251–258. Stenton-Dozey, J.M.E., Jackson, L.F., and Busby, A.J. 1999. Impact of mussel culture on macrobenthic community structure in Saldahana Bay, South Africa. Marine Pollution Bulletin 39: 357–366. Stenton-Dozey, J., Probyn, T., and Busby, A. 2001. Impact of mussel (Mytilus galloprovincialis) raft culture on benthic macrofauna, in

situ oxygen uptake, and nutrient fluxes in Saldanha Bay, South Africa. Canadian Journal of Fisheries and Aquatic Sciences 58:1021– 1031. Sterner, R.W. 1986. Herbivores’ direct and indirect effects on algal populations. Science 231:605– 607. Straus, K.M., Crosson, L.M., and Vadopalas, B. 2008. Effects of Geoduck Aquaculture on the Environment: A Synthesis of Current Knowledge. Washington Sea Grant, University of Washington, Seattle, WA. Strohmeier, T., Aure, J., Duinker, A., Castberg, T., Svardal, A., and Strand, Ø. 2005. Flow reduction, seston depletion, meat content and distribution of diarrhetic shellfish toxins in a long-line blue mussel (Mytilus edulis) farm. Journal of Shellfish Research 24:15–23. Sundbäck, K., Miles, A., and Goransson, E. 2000. Nitrogen fluxes, denitrification and the role of microphytobenthos in microtidal shallow-water sediments, an annual study. Marine Ecology Progress Series 200:59–76. Suzuki, T. 2001. Oxygen-deficient waters along the Japanese coast and their effects upon the estuarine ecosystem. Journal of Environmental Quality 30:291–302. Tenore, K.R., Boyer, L.G., Call, R.M., Corral, J., Garcia-Fernandez, C., Gonzalez, N., GonzalezGurriaran, E., Hanson, R.B., Iglesias, J., Krom, M., López-Jamar, E., McClain, J., Pamatmat, M.M., Perez, A., Rhoads, D.C., de Santiago, G., Tietjen, J., Westrich, J., and Windom, H.L. 1982. Coastal upwelling in the Rias Bajas, NW Spain: contrasting the benthic regimes of the Rias de Arosa and de Muros. Journal of Marine Research 40:701–772. Viaroli, P., Bartoli, M., Bondavalli, C., Christian, R.R., Giordani, G., and Naldi, M. 1996. Macrophyte communities and their impact on benthic fluxes of oxygen, sulphides and nutrients in shallow eutrophic environments. In: Caumette, P.J., Castel, J., and Herbert, R. (eds.), Developments in Hydrobiology 117. Coastal Lagoon Eutrophication and Anaerobic Processes: C.L.E.A.N.. Kluwer Academic Publishers, Dordrecht, Germany, pp. 105–119. Viaroli, P., Bartoli, M., Giordani, G., Azzoni, R., and Nizzoli, D. 2003. Short term changes of benthic fluxes during clam harvesting in a coastal

Bivalve shellfish aquaculture and eutrophication

lagoon (Sacca Di Goro, Po River Delta). Chemistry and Ecology 19:189–206. Vitousek, P.M., Aber, J., Howarth, R.W., Likens, G.E., Matson, P.A., Schindler, D.W., Schlesinger, W.H., and Tilman, G.D. 1997. Human alteration of the global nitrogen cycle: causes and consequences. Ecological Applications 7:737– 750. Wassmann, P. 2005. Cultural eutrophication: perspectives and prospects. In: Wassmann, P., and Olli, K. (eds.), Drainage Basin Inputs and Eutrophication: An Integrated Approach. University of Tromsø, Tromsø, Norway, pp. 224–234. www.ut.ee/∼olli/eutr/ Weise, A.M., Cromey, C.J., Callier, M.D., Archambault, P., Chamberlain, J., and McKindsey, C.W. 2009. Shellfish-DEPOMOD: modelling the biodeposition from suspended shellfish. Aquaculture 288:239–253. Weiss, E.T., Carmichael, R.H., and Valiela, I. 2002. The effect of nitrogen loading on the growth rates of quahogs (Mercenaria mercenaria) and soft-shell clams (Mya arenaria) through changes in food supply. Aquaculture 211:275–289. Whiteley, J., and Bendell-Young, L. 2007. Ecological implications of intertidal mariculture: observed differences in bivalve community structure between farm and reference sites. Journal of Applied Ecology 44:495–505. Widdows, J. 1978. Combined effects of body size, food concentration and season on the physiology of Mytilus edulis. Journal of the Marine Biological Association of the United Kingdom 58:109–124.

215

Wiegner, T.N., Seitzinger, S.P., Breitburg, D.L., and Sanders, J.G. 2003. The effects of multiple stressors on the balance between autotrophic and heterotrophic processes in an estuarine system. Estuaries 26:352–364. Xie, Q., and Burnell, G.M. 1995. The effect of activity on the physiological rates of two clam species, Tapes philippinarum (Adams & Reeve) and Tapes decussatus (Linnaeus). Biology and Environment: Proceedings of the Royal Irish Academy 95B:217–223. Yang, H., Wang, P., Zhang, T., Wang, J., He, Y., and Zhang, F. 1999. Effects of reduced salinity on oxygen consumption and ammonia-N excretion of Chlamys farreri. Chinese Journal of Oceanology and Limnology 17:208–211. Yokoyama, H. 2002. Impact of fish and pearl farming on the benthic environments in Gokasho Bay: evaluation from seasonal fluctuations of the macrobenthos. Fisheries Science 68:258–268. Zeldis, J.R. 2005. Magnitudes of natural and mussel farm-derived fluxes of carbon and nitrogen in the firth of Thames. CHC2005-048. Environment Waikato Technical Report 2005/30, 42pp. Zeldis, J.R., Howard-Williams, C., Carter, C.M., and Schiel, D.R. 2008. ENSO and riverine control of nutrient loading, phytoplankton biomass and mussel aquaculture yield in Pelorus Sound, NZ. Marine Ecology Progress Series 371:131–142.

Chapter 8

Mussel farming as a tool for re-eutrophication of coastal waters: experiences from Sweden Odd Lindahl

Introduction Many coastal areas are affected by eutrophication due to nutrient leakage from agriculture operations, sewage discharges, and other human activities. Blue mussels are not only good at harvesting nutrients through their food intake of phytoplankton but they also provide valuable seafood and raw material for production of feedstuff or use as fertilizer. Studies in Sweden have demonstrated that mussel farming can be used as a management tool for society to compensate for nutrient discharges in nutrient trading schemes. This concept most likely has a great potential on a global scale in temperate waters to counteract the negative effects of coastal eutrophication.

Mussel farming: open landscape feeding in the sea Eutrophication of coastal waters is causing anoxic bottom conditions and the formation of algal mats in shallow bays (Diaz and Rosenberg 1995; Cloern 2001; Diaz and Rosenberg 2008). The idea of farming blue mussels in order to reduce the amount of phytoplankton, and thereby the negative effects of the eutrophication, was introduced in the 1990s (Haamer 1995; see also Chapter 7 in this book). This was a new concept that regarded the increasing amounts of nutrients and plankton in coastal waters as a resource which should be recycled to land and reused. The blue mussel (Mytilus edulis) is, like many

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 217

218

Shellfish Aquaculture and the Environment

Figure 8.1 The principle of recycling nutrients from sea to land by mussel farming where the farming and harvest can be regarded as a recycling engine.

other marine organisms, a filter-feeding animal. They live by pumping in the surrounding water and filtering out particles, mainly phytoplankton, for food (see Chapter 5 in this book). New food particles are constantly brought to the sedentary mussels because the seawater is in continuous motion. The potential for ecological and environmental benefits of mussel farming to improve coastal water quality are scientifically well known, as pointed out by, for example, Ryther et al. (1972); Haamer et al. (1999); Edebo et al. (2000); Newell (2004). The largest proportion, ca. 80%, of the nutrients which are discharged to Swedish coastal waters comes from diffuse emissions such as runoff from forests and farm land, atmospheric deposition, and rural living. The remaining 20% comes from point sources such as sewage treatment plants and industries (Anonymous 2003). Since the agriculture

operations are one of the main sources of the emissions, it seemed logical that the nutrients harvested in the sea should be reused in agricultural operations (Fig. 8.1). The expression “agro-aqua recycling” has been introduced, and refers to mussel farming that can be used as a recycling engine of the nutrients (Haamer et al. 1999). There is interest from both an environmental and a socioeconomic point of view to try to recapture and reuse the excess nutrients because production of nitrogen as a fertilizer is an energy-demanding and climatenegative process, and phosphate is a limited resource on a global scale. These nutrients are assimilated into mussel biomass, which in turn can be used as seafood, as feedstuff, or as fertilizer in agricultural operations. One of the environmental issues with the highest priority as expressed by the Swedish Parliament is decreasing the supply of nutrients to the coastal zone as well as the amounts

Mussel farming and re-eutrophication in Sweden

to the open sea (Anonymous 2003). Mussel farming has been recognized as a possible measure to improve coastal water quality since the 1980s. Research on how to use mussel farming as a strategic environmental management tool has been ongoing since the late 1990s. It has been concluded that trading nutrient discharges is a necessary strategic tool for society in order to use mussel farming for recycling nutrients from sea to land (Lindahl et al. 2005). One kilogram of fresh mussels contains 8.5–12 g of nitrogen, 0.6–0.8 g of phosphorous, and about 40–50 g of carbon (Lutz 1980; Petersen and Loo 2004). Mussels are also a valuable and healthy marine food product, high in protein and with a fat content of only about 2%, 40% of which are Ω3 longchain fatty acid molecules (Berge and Austreng 1989). Longline farming is the most common method for mussel production in Sweden (Fig. 8.2). The mussels are mostly grown on vertical suspenders attached to horizontal longlines (Fig. 8.3). Approximately 300 t of high seafood quality mussels may be produced per hectare of sea surface in 12–18 months on the Swedish west coast with its marine conditions. It has been calculated that on average, 1 ha of mussel farming after 1–1.5 years of growth, resources 20 ha of the annual phytoplankton production to provide for the mussels’ food intake (Lindahl and Kollberg 2009). This calculation was made by using a long-term mean from 1985 to 2006 of annual primary productivity in Swedish marine coastal waters of 240 g C m−2 year−1 (Lindahl 2007), a carbon content of 4% in the live mussel, and a gross growth efficiency of 0.2 (Riisgård and Randløv 1981). A similar calculation for the brackish Baltic Sea area emphasized that 7.5 ha of primary production is required for each hectare of mussel farming (Lindahl and Kollberg 2009). This estimate was based on production of 120–150 t of longline farmed mussels grown over 2–3 years per hectare, and a phytoplankton production of 160 g C m−2 year−1 (Elmgren

219

1984). It was assumed that the carbon content and gross growth efficiency was the same as above. Mussel farming, from an environmental point of view, can be regarded as an activity similar to open landscape feeding on land. In the case of mussels, the result is clearer water and the fact that the phytoplankton biomass has been harvested and utilized. Thus, the excess nutrient resource in the coastal water is turned into mussel biomass, which can be used as seafood, as feedstuff, or as fertilizer instead of causing negative environmental effects.

Estimating the environmental value of mussel farming It has been shown that there is a willingness among common people to spend a significant amount of money to reduce the negative effects of eutrophication of coastal waters (Huang et al. 1997; Söderqvist 1998; Markowska and Zylicz 1999). According to the scant literature on assessment of the value of removing nutrients by mussel farming, there are simple comparisons of unit abatement costs with other abatement measures (Lindahl et al. 2005). This is an appropriate approach for a point source such as a sewage treatment plant and where there are no alternative cleaning options at the local scale. However, at a larger scale and concerning diffuse emissions, there are a number of additional abatement measures available. These include improvements of agricultural operations in order to reduce nutrient emissions, increased and improved sewage cleaning by rural households and industries not connected to common sewage treatment, the use of wetlands, and others. The value of mussel farming as an abatement measure is then determined by the cost savings obtained by the replacement of other measures that have a higher cleaning cost (Gren et al. 2009). For Northern Europe, there is only one study estimating the value of mussel farms for

Lulea Hailuoto

Gu lf o fB oth nia

Oulu

Trondheim

Vaasa

Sundsvall

NORWAY

SWEDEN

FINLAND

Turku

Oslo

Stavanger Stockholm

f of Gul

nd

a Finl

rg sbu eter St. P

Tallinn

Hiiumaa

RUSSIA

ESTONIA

Lysekil

k erra

Copenhagen Malmo

cS

Oland

Gulf of Riga

LITHUANIA

Rugen Gdansk

GERMANY NORTH 0 50 100 200

LATVIA

Ba

Bornholm Kiel

Riga

lti

gat

e Katt

Gotland

DENMARK

Funen Irodotou

ea

Saaremaa

g Ska

Kaliningrad (RUSSIA)

Vilnius Minsk

BELARUS POLAND

Kilometres Figure 8.2 Map of northwest Europe.

Figure 8.3 The principle of farming mussels using longlines.

220

Helsinki

Aland Islands

Mussel farming and re-eutrophication in Sweden

combating eutrophication in a wider context with respect to alternative abatement measures, spatial scale, and different nutrient load targets. This study applied the replacement cost method to areas ranging from the Swedish west coast with its marine conditions (20– 30 PSU), to the Sound (15–20 PSU), along the southern coast (10–15 PSU), and to the southern part of the east coast (5–10 PSU) in the Baltic Sea with its brackish conditions (Gren et al. 2009). Mussels have a comparative advantage when considering multipollutant abatement capacity, but mussel farming is a relatively recent innovative technology for cleaning, and there is a lack of data with respect to production cost, mussel sales options for human or animal consumption, and growth under variable conditions. The study calculated the marginal cleaning cost of mussel farms with and without mussel sales options, high and low mussel growth rates, and contents of nutrient in mussels. The calculated constant marginal cost then varied between no cost per kilogram nutrient cleaning, USD 100 kg−1, and USD 1000 kg−1 for nitrogen and phosphorous cleaning respectively (SEK 1 = USD 0.13). The low marginal cleaning cost occurred for the Kattegat and the Sound marine basins, whereas the largest costs were found in the Northern Baltic Proper brackish basin. The estimated marginal costs were highly dependent on the growth rate of the mussels, which is strongly connected to salinity. Another important factor noted was whether the main bulk of the mussel production was going to be marketed as seafood or as less valuable products such as feedstuff or fertilizer (Table 8.1). For example, there was no marginal cost for nitrogen removal along the Swedish west coast when the mussels were sold as seafood. The estimated marginal cost was about USD 8 kg−1 nitrogen when the mussels were used for feedstuff and about USD 190 kg−1 nitrogen when only nutrient harvest was performed and the harvested biomass was given no commercial

221

Table 8.1 Estimated marginal costs using mussel farming for nitrogen and phosphorus harvest along the Swedish coasts (Gren et al. 2009).

Skagerak/Kattegat Öresund Strait Southern Baltic Northern Baltic

USD kg−1 nitrogen

USD kg−1 phosphorus

0–42 0–47 8–44 17–100

0–420 0–470 80–440 170–1000

The cost span depended on the value of the mussels as seafood, feedstuff, or fertilizer for harvesting nutrients. SEK 1 = USD 0.13.

value (Lindqvist 2008; Gren et al. 2009). For phosphorous, the costs was approximately 10 times higher due to the fact that the content of phosphorus in a mussel is about one-tenth that of nitrogen. A special focus of the Gren et al. (2009) study was using mussel farming for environmental purposes in the brackish and eutrophic Baltic Sea. When comparing the marginal cleaning costs of mussel farming with those of 20 alternative abatement measures in 24 different drainage basins of the sea, it was found that mussel farming has a positive value for a large range of nutrient reductions. The results indicated that there was a large span in the calculated marginal costs for recycling nutrients from sea to land. Mussel farming had lower marginal cost for nutrient harvest compared with many other abatement measures in the Baltic, but at the same time the cost was much higher compared with marine areas depending on slower growth, reduced sales options due to the small size of the Baltic mussels (<3 cm). The estimated values, calculated as the difference in minimum costs for given nutrient reduction targets with and without the inclusion of mussel farming as a cleaning option, ranged between approximately USD 0.14 and USD 1.54 billion per year, which corresponded to a cost savings between 2% and 11% (Gren et al. 2009). It should be pointed out, however, that mussel

222

Shellfish Aquaculture and the Environment

farming in the Baltic Sea is a rather special case and mussels from this area are not suitable for use as seafood due to the brackish conditions. Alternatives for the use of the small Baltic mussels are as organic feedstuff replacing fish meal in feed for, for example, laying hens and chicken poultry (see below). The large range of values obtained pointed to the need for more empirical research on growth of mussels, nutrient concentration under different physical environmental conditions, and also on possible locations of mussel farms from a large scale perspective. Furthermore, the results of Gren et al. (2009) indicated that the options of selling mussels are crucial for the marginal cleaning cost of nutrients by mussel farming. Selling mussels also requires further exploration regarding the eventual content of toxins and pathogenic microbes in mussel tissue. Marginal cleaning costs are also affected by the type of technology employed in mussel farming. In the present study, only longline technology was assumed. This is a suitable technology for mussel farms in relatively wind-protected coastal zones, but may not be appropriate for open coastal zones prevailing in parts of the Baltic Proper. Instead, polyethylene pipes for buoyancy and nets as mussel collectors or longlines lowered below surface might be more useful. Another phenomenon that was not included in Gren et al. (2009) was the possible increase in nutrient regeneration from mussels’ filtration. The value of mussel farms as an abatement option is then zero if the nutrient regeneration exceeds the decrease in nutrients from harvesting of mussels. Gren et al. (2009) concluded that the potential values and technological development of mussel farming as a cleaning device are dependent not only on research but also on policies for combating eutrophication in the Baltic Sea. In general, economic instruments create the incentives for making the best use of scarce cleaning resources and for stimulating techno-

logical development (see, e.g., Baumol and Oates 1988).

Trading nutrient discharges A collaborative project between Norway and Sweden in the beginning of the 2000s pioneered commercial mussel farming as a compensating measure for nutrient discharges causing eutrophication (Lindahl et al. 2005). If the right tools and support were given in order to stimulate, and thus increase, mussel farming, the negative effects of coastal marine eutrophication could be reduced. This idea was a parallel to the global trading that is currently being used for carbon dioxide effluents into the atmosphere, but used on a local or regional scale. Such emissions trading schemes, or tradable quota schemes, were first introduced in 1970 in amendments to the U.S. Clean Air Act (Tietenberg 2003). Any activity that releases nutrients to the environment should also be responsible for the emission or discharge being mitigated. Quotas can be traded if it is too complicated or too expensive for the nutrient emitter to purify the effluent at the source. A compensation measure, which removes the corresponding amount of nutrients, can be carried out through a variety of different enterprises. In this case, it is a mussel farmer who gets paid for farming and harvesting a certain amount of mussels and the ecosystem services the farming provides (Fig. 8.4). The released nutrients give rise to a natural production of phytoplankton which in turn, is the main feed for mussels. The nutrients are assimilated into mussel meat and when the mussels are harvested, the nutrients are recycled back to land in the form of mussel biomass. The nutrient emitter who buys an emission quota thus connects environmental economy, where funding is used to combat eutrophication, with market economy, in which the mussel farm enterprise is running its business. Nutrient

Mussel farming and re-eutrophication in Sweden

223

nutrient emission trading eutrophication combat

mussel farming enterprise

nutrient emitter

Figure 8.4 The trading of nutrient emissions involves that the emitter buys a compensation quota from, for example, a mussel farm. The trading connects market economy, under which the mussel farm enterprise is acting, with environmental economy improving coastal water quality.

trading systems are becoming an important means of improving coastal waters in the United States, where the Environmental Protection Agency is mandated by the Clean Water Act to improve water quality (e.g., Chesapeake Bay Program 2000; Newell 2004; Pennsylvania Nutrient Management program 2009). In Sweden the use of trading schemes for nutrient discharges has been evaluated (Anonymous 2006a) and was approved at the highest political level for first practical trials in 2009 (Anonymous 2009). A prerequisite for trading schemes to work smoothly is that a market be available where sellers and buyers of nutrient quotas can easily do business. There is still no organization for handling nutrient trading in Sweden, but local or regional administrations will likely play a key role in this new environment of business networking and trading. For example, the first attempt in Sweden (the Lysekil case, see below) to trade a nitrogen discharge using mussel farming was under the supervision of the environmental officers from the local County Board in order to ensure that the agreed amounts of nutrients were recycled from sea to land as contracted. At a larger scale, one option could be a regionally licensed nutrient

trading enterprise that organizes the buying and selling of nutrient quotas. The payment, which should be based on bidding in competition with other available nutrient abatement measures, should be made per unit of nitrogen and/or phosphorous that is removed out of the coastal water. This approach will improve the cost-efficiency of nutrient trading and be beneficial not only to the involved enterprises but also to society as a whole. It is generally easy to identify the “owner” and to measure and calculate the amount of released nutrients if the nutrients are discharged from a point source. On the Swedish west coast and in many other coastal areas, more than 80% of the nutrients are released from diffuse sources (Anonymous 2003). The situation becomes more complex and reduces the possibility of easily trading the nutrient quotas. Who is to be blamed and thus who will have to pay for the nutrient emission if the emission has no “owner”? For example, it is possible to estimate the total nutrient emission from agriculture operations of an area, but it is much more difficult to point out the emission coming from each of the farm enterprises within the area. At present there is no statutory demand on the agricultural sector as a whole to do

224

Shellfish Aquaculture and the Environment

common measures in order to counteract their emissions. It should also be possible to trade the diffuse emissions, however, by combining different available options. An example of where increased mussel farm operations could be used as compensation for nutrient discharges is from rural living in coastal areas. The cost for single households to connect to municipal sewage plants is, in many cases, too high especially along the rocky Swedish and Norwegian Skagerrak coasts with its many islands. More or less effective infiltration systems are generally used, but these are simply not very efficient for nitrogen. A compensation for the nitrogen discharge could be through mussel farming where the households pay a mussel farm enterprise for the harvest of an agreed amount of nitrogen and/or phosphorous. The nitrogen discharge from a “rural” person is about 3.5–4 kg year−1 (Anonymous 2003), which means that an annual harvest of 350–400 kg of mussels would compensate for 100% of the emission from one person for both the nitrogen and the phosphorus supplies, provided the household is using low- or nonphosphorus detergents. Trading of such local solutions would be a task for the coastal communities to organize, but supervision is probably best carried out at a regional level.

Agricultural environmental aid program and mussel farming A simple way to utilize the potential of mussel farming for nutrient harvest would be to extend the existing European agricultural environmental aid program (EEC 2078/92 and 1257/1999) to include the coastal zone and aquaculture operations as suggested by Lindahl and Kollberg (2009). The legal framework is quite complex but some of the measures could easily be compared with mussel farming from an environmental point of view. From the administrative and legal point of view, however,

this has not been possible since this program specifically is designated only for farmland and is not present beyond the shoreline. Support has been given for the establishment of wetlands, spring cultivation, and catch crops in the agricultural environmental aid program, in order to decrease the nutrient releases from farmland into the environment. If aquaculture operations are to be included, it should involve paid support to mussel farming enterprises through their harvest of mussels (and thus their harvest of nutrients) in the same way that support is being paid to agricultural farmers for operations that reduce nutrient leakage from their farmland. This would be a simple, cost-effective, and straightforward way of improving coastal water quality at many coastal sites, but will require that the legal framework defines “farm water” equally as farmland (Fig. 8.5). The mussels feed on the natural phytoplankton community, which means that the phytoplankton biomass can be considered as a type of catch crop. In the previous agricultural environmental aid program, which was running between 2000 and 2006, catch crops in Sweden were compensated at USD 130 per hectare (Anonymous 2006b). The annual compensation for spring cultivation of a catch crop was USD 50 per hectare. Catch crops were farmed on 180,000 ha of farmland, and spring cultivation was used on 90,000 ha during the 2000s. The total annual cost for these measures was USD 28 million. The Swedish Commission for the Environment of the Seas (Anonymous 2003) has calculated that catch crops and spring cultivation together decreased the nitrogen release by 2000 t annually, which means a cost of USD 14 per kilogram of retained nitrogen. If a mussel farm enterprise were to be compensated at the same price, it means a subsidy of about USD 0.14 per kilogram of live mussels. This is roughly 25% of what the enterprise needs as a gross income for the mussels as estimated by Lindahl and Kollberg (2009).

Mussel farming and re-eutrophication in Sweden

225

spring cultivation catch crops

mussel farming

harvest

wetland

Figure 8.5 Wetlands, spring cultivation, and catch crops are measures in agriculture environmental operations that can be compared with mussel farming and harvest in the sea. Both measures combat eutrophication and should be able to organize under a common environmental aid program improving coastal water quality.

Farmers in Sweden pay environmental tax when using fertilizer. Another option to fund the recycling of nutrient discharges coming from agriculture operations would be to use this environmental tax. The tax was introduced to decrease the elevated levels of fertilizer spread in the fields and could have a double effect if in coastal areas some of it was used to further decrease nutrient levels in the sea through subsidizing, for example, extended mussel culture operations. This could, in part, pay for the increased nutrient discharge caused by the fertilizer through subsidizing the nutrient removal service mussel farming provides for the environment. There has not been any political interest in Sweden so far to transfer this kind of subsidy to the shellfish industry. During the years 1995–2001, about 2500 ha of wetlands were established in southern Sweden (Anonymous 2003). The investment costs seen over a 20-year period were calculated to be USD 4350 per hectare per year (Anonymous 2006b). With a calculated reduction of 120 kg of nitrogen per hectare of wetland per year, the cost of reduced nitrogen would be USD 36 per kilogram. If this cost is transformed to nitrogen uptake by mussels it would correspond to about USD 0.30 per kilogram mussel. A full comparison must include

the low running costs of wetlands compared with the much higher costs for mussel farming. Harvesting costs alone amount to USD 0.13– 0.19 per kilogram mussel. On the other hand, a wetland does not produce a commodity of commercial value, while the first-hand value of seafood mussels is between USD 0.4–0.6 per kilogram. When comparing the uptake of nutrients by agricultural catch crops and wetlands with that of the mussels, the temporal aspect is important. The uptake is far higher when temperatures are high (during summer). The primary production in the sea and also the mussel filtration capacity is less temperature dependent (Loo and Rosenberg 1983), and the phytoplankton growth in the stratified Swedish coastal waters is regulated by light if nutrients are not the limiting factor. In Swedish waters, the phytoplankton growth season generally lasts from February/March to October (Lindahl 1995). This means that nutrient discharge to coastal recipients during 8 months of the year is assimilated by the phytoplankton and then grazed by the mussels or other filtrating organisms. Thus, nutrient harvest in the sea by shellfish has a considerably longer annual active period compared with the agricultural activities on land.

226

Shellfish Aquaculture and the Environment

Added ecosystem services through mussel farming There is much information in the literature on the effects of the rich biosedimentation from a floating mussel farm (e.g., Hatcher et al. 1994; Grant et al. 1995; Mirtho et al. 2000; Newell 2004). The degradation of the increased supply of organic material will increase the oxygen consumption at the seabed surface, and may cause negative effects on the benthic ecosystem. In the worst case, it may cause the development of hydrogen sulfide and a more or less “dead bottom” may occur. To avoid such negative effects, the sediment surface must stay oxygenated, allowing the natural denitrification processes to continue. Denitrification is important since through this process different nitrogen substances, for example, ammonium, are transformed into biologically inactive nitrogen gas (Newell 2004). At sites with favorable bottom and water circulation conditions, the natural denitrification process may increase and has been estimated to add up to 25% of the nitrogen harvested by the mussels. In principle, it would also be possible to include this nitrogen flux to the atmosphere into a nitrogen trading scheme. It is a comprehensive task, however, to make good estimates of the flux and consequently also to trade it. Therefore, this option has not yet been utilized in Sweden. When mussels are farmed with an environmental perspective added to the commercial value, all of the farmed mussels and other attached biota should be brought to land in order to optimize the positive effect on the environment. By this principle, the amount of harvested and recycled nutrients is maximized. The remainder resource, mainly made up of small and crushed mussels, cannot be used as seafood. The remainder resource also contains large amounts of byssus, which is high in raw protein (55%), and thus also a valuable resource (Lindahl, unpublished data). Biofouling sea squirts (Ciona intestinalis) may also make up a considerable part of the harvest,

but are less valuable since the raw protein content is only 23%. Increased mussel production carried out with an environmental perspective will require that the quantity of the remainder will also increase and may, in the future, constitute up to 10,000 t or more along the Swedish west coast. It is essential that this protein and nutrient rich resource is utilized in a sustainable way, and if possible, is also traded. Two different products utilizing the remainder resource are (1) a feed additive for poultry production, and (2) as fertilizer on farmland. In both cases, the focus has been on the organic agricultural industry.

The city of lysekil, the first buyer of a nutrient emission quota The first Swedish case of trading a nutrient discharge started in the small town of Lysekil, situated on the west coast. In 2004, this community discovered the possibility of compensating mussel farming for the annual discharge of 39 t of nitrogen from its main sewage treatment plant. The Community Board of Lysekil was permitted by the County Board, as a trial between 2005 and 2011, to continue to emit nitrogen from the sewage plant, presupposing the same amount of nitrogen was annually “harvested” and brought ashore by 3500 t of blue mussel (Fig. 8.6). It was possible in accordance with the European Community sewage directive to exchange nitrogen removal in the sewage treatment plant by mussel farming in accordance with an EC-legal assessment (Prof. S. Westerlund, pers. comn.). The community of Lysekil bought this ecosystem service after bidding from a mussel farming enterprise, which ensured that the nitrogen removal would take place. Since the mussels are produced mainly as seafood for human consumption, farms cannot be located too close to the sewage discharge point. A monitoring program assures that the quality standard of harvested mussels for human con-

Mussel farming and re-eutrophication in Sweden

227

§:70% N-removal

< ~ 100% N-removal

Figure 8.6 Top panel shows the principle of required (70%) removal of nitrogen according to the sewage treatment regulation. Bottom panel shows how up to 100% of the nitrogen emission can be recycled by using mussel cultivation and harvest.

sumption will not be affected by such things as pathogenic microbes. The cost of USD 200,000 for the Lysekil Community is far below the price for constructing and running a traditional nitrogen removal step within the sewage plant. Additionally, a sewage plant of the size in Lysekil (>10 000 population equivalents [p.e.]) has, according to the sewage directive, to remove only 70% of the nitrogen. In this case Lysekil is currently heading for 100% removal in order to show that the mussels can do a better job compared with nitrogen removal using traditional techniques. As an extra bonus, about 3 t of phosphorus is also recycled back to land. The winners of the nutrient trading in Lysekil are thus both the community and the environment! The pristine benthic ecosystems and bottom sediments were characterized and described prior to the establishment of about 50 longlines in the Lysekil area in 2005 and then

compared with the situation after 2 years of mussel farming. The results demonstrated that the type of sediment and the species composition of the benthic community played an important role (Svanberg 2009). It was also found that the most important factor was locating the longlines on sites with continuous exchange of bottom water. This counteracted the risk of development of anoxic sediments.

Swedish mussel farming and its markets The economic basis of mussel farming and harvesting is to produce food for human consumption. The world production of mussels today exceeds 1.5 million t, of which half is produced and consumed in Europe. The demand is steadily increasing, but the main production areas in Europe have reached a

228

Shellfish Aquaculture and the Environment

level where they can no longer expand due to the shortage of suitable farm areas, and there has occasionally been a shortage of mussels on the market (Smaal 2002). The domestic consumption of fresh mussels in Sweden is only about 1000 t. There are estimates that an annual production of 50,000 t of mussels on the Swedish west coast alone is a realistic figure (Lindahl et al. 2005). This activity also is desirable for improving coastal water quality (Anonymous 2003). Thus, an increase in production and export of Swedish mussels over the next 15–20 years is quite realistic. The volume of Swedish mussel production has remained low (between 1000 and 2000 tons of mussels per year) since the beginning of the 1980s. The reasons for this are rather complex, but can best be described as a combination of shortage of capital, toxin problems, poor cooperation between farmers, and too low marketing or export efforts. The diarrheic shellfish toxin (DST) toxic events were earlier regarded as one of the strongest factors hampering an expansion of the mussel farm industry in Sweden. Food safety of Swedish mussels

is regulated under the European Community Directive 91/492 EEG, and since the 1990s, both the knowledge and experience exist to overcome the periods of toxicity. A reliable method for detoxification would be useful for mussel farmers to guarantee a more continuous supply of seafood mussels. Improved knowledge regarding factors affecting the elimination rate of these toxins is required to develop practical and cost-effective detoxification methods (Svensson 2003a, 2003b). Mussel farming as an environmental measure requires a reliable market for the harvested mussels in order to able to carry through with the agreed nutrient harvest quotas. This is especially important when nutrient trading contracts are to be fulfilled. As in the case of Sweden, the domestic seafood market is too small for an increase of the mussel production. One option is increased export which, at least so far, has not turned out to be a reliable option. Consequently, alternative strategies must be explored in order to increase interest and support of mussel farming in Sweden as an environmental measure (Fig. 8.7).

feedstuff

compost

Figure 8.7 The three main areas using mussels as seafood, for feedstuff production, or as fertilizer, the latter preferably first after composting.

Mussel farming and re-eutrophication in Sweden

Mussel farming entirely for feedstuff production has not been undertaken in Sweden, nor anywhere else to our knowledge. The “feed mussels” will have a considerably lower commercial value but, on the other hand, probably can be produced at roughly half or less the cost compared with seafood mussels. Still, the price to be able to process the feed mussel biomass into mussel meal to compete with and replace fish meal in, for example, organic feed (see below) may be too high. This will certainly be the case in the Baltic Sea, with the slower growth and smaller biomass, but also in marine areas. To overcome this economic difficulty, the income for the nutrient trading should be included in the final accounting of the feed mussel production. A combination of seafood, feed mussel production, and the environmental subsidies suggested above is a likely solution for the development of future mussel farm enterprises in Sweden and elsewhere.

Mussel meal instead of fish meal in organic feeds Due to their plumage, hens and chicken have a greater need than any other food producing animals for their feed to contain sulfur-rich amino acids, especially methionine and cysteine (Jakobsen and Hermansen 2001). Most feedstuffs used in poultry feed have a content below the requirements of the birds for these essential amino acids. The feedstuff enterprises often use synthetically produced methionine in order to enrich the feed used for laying hens. It is also common to include fish meal since it contains these important amino acids. None of these protein sources is suitable for organic production. According to the Swedish KRAV and International Federation of Agricultural Movements (IFOAM) in 2003, no more than 15% nonorganic components are allowed in feed given that egg or poultry production is regarded as organic.

229

Methionine-, cysteine-, and lysine-rich ingredients in organic egg and broiler production will be a scarce resource with a future shortage of allowed feed components (Jakobsen and Hermansen 2001). This will result in increased competition for protein feed components due to the reduced amounts of fishmeal and an EU ban against adding synthetic amino acids. In 2012, any nonorganically produced feed ingredient (EEG 2092/91 and 1294/2005) will also be banned. The nonorganic share in which fish meal constitutes a considerable part should decrease to 10% in 2008, 5% in 2010, and 0% in 2012. Since synthetically produced methionine not will be allowed, and it has been difficult to find organically produced feedstuff containing enough of the essential amino acids, this suggested regulation was changed in spring 2008 allowing for the continued use of fish meal. There is a growing resistance to the use of fish meal in feed and the research to find useful replacement products is ongoing. The shortage of good quality protein sources also jeopardizes animal health. It is a presumption that access to good feedstuffs has to be guaranteed if organic poultry production shall survive in the future (Elwinger and Wahlström 2000; Elwinger et al. 2002). The expansion of organic production for other monogastric animals, for example, pigs and fish, is also a likely development, especially in the industrial world. Thus, another option could be the use of mussel meal as an ingredient in aquafeed where fish meal and fish oil have traditionally been the primary ingredients. Currently, global production of these products is insufficient to support further growth of aquaculture at high levels in feed formulations (Hardy 2009). Sustainable feeds that support rapid, economical growth of farmed fish are required to achieve sustainable aquaculture. The blue mussel has a high content of the essential sulfur-rich amino acids methionine, cysteine, and lysine, matching the content in fishmeal (Table 8.2) (Lutz 1980; Berge and

230

Shellfish Aquaculture and the Environment

Table 8.2 The content of protein and share of sulfur-rich amino acids and lysine in mussel and fish meal and some other commonly used feedstuff products (Berge and Austreng 1989; Johansen 2008)

Protein (g kg−1 DW) Methionine (% of protein) Methionine + cystine (% of protein) Lysine (% of protein)

Mussel meat

Mussel meal

Fish meal

Rape cake

Peas

Soy cake

Wheat

645 1.8 2.6 6.0

764 2.5 4.2 7.7

670 2.8 3.7 7.4

237 2.0 4.5 5.6

265 1.0 2.4 7.1

520 1.4 2.9 6.2

120 1.6 3.9 2.8

Austreng 1989) and can, to the extent shells are included in the feed, also provide calcium carbonate. Since mussels are at the second step of the marine food chain, the use of mussels instead of fish for feed production is of large ecological significance at a time when many fish stocks are overexploited on local, regional, and global scales.

Mussel meal in feeds for organic poultry Mussels as feed for laying hens was first tested at a small scale in Norway in the mid-1980s (Berge 1986). The first pilot experiment in Sweden was carried out in 2003 and showed that the hens preferred steamed mussel meat over the ordinary standard feed for laying hens (Kollberg 2008). The yolks became strongly colored and no bad taste was perceived in the eggs. These eggs were found useful for cooking, baking, making mayonnaise, or direct consumption (K. Johansen, pers. comm.). Also, the meat of the hens was tasty and untainted. Steamed mussel meat is not practical to handle as a feedstuff at a large scale, and the next step was to dry the meat and grind it into mussel meal and mix with other components into a complete feed. A fresh live blue mussel roughly consists of three equal parts: shell, meat, and water. The raw material used for the meal production must be fresh and the whole process must be carried out under the same hygienic conditions

as for seafood production. In order to separate the meat from the shell, the mussels are steamed quickly and are thereafter spread on a shaking grid where the coagulated meat comes loose from the shell. It is then simple to separate meat and shell by using a density bath where the meat floats and the shell sinks. This is a standard technique used by the seafood industry to separate shell and meat. The meat is dried at between 85 and 90°C to about 5% water content and then ground (Kollberg and Lindahl 2006). This high temperature is also required to eliminate pathogenic microbes according to the regulations for feed production. Regardless of the heating procedure, it is required that each batch of meal is tested for occurrence of Salmonella before it is allowed to be delivered to the feed factory to be included in the complete feed. The dried mussel meal had a fat content of ∼8% with a high share of long-chained polyunsaturated fatty acids (Duinker et al. 2005; Kollberg and Lindahl 2006). The protein content is about 65%. One thousand kilograms of raw mussels results in approximately 250 kg of meat, which in turn provides 50 kg of dried mussel meal. The most commonly used feed mix to this point is 5% mussel meal, which means that 50 kg of mussel meal is sufficient to produce 1000 kg of feed (Table 8.3). There will also be 350 kg of shell, of which about 20% can be used as a calcium source in feed for poultry. The rest of the shells have a value of about 3 US cents per kilogram for liming including its

Mussel farming and re-eutrophication in Sweden

Table 8.3 Transformation of live mussels into mussel meat, meal, and feed.

Live mussels

Content of mussel meat

Dried mussel meat = mussel meal

5% mix in final feed

1000

250

50

1000

All in kilograms.

content of microelements. Farms with organic production have already shown an interest in the product (Olrog, pers. comm.). The Department of Animal Nutrition and Behavior at the Swedish Agricultural University continued the studies using mussel meal at a much larger scale and over a full production cycle. One of the studies evaluated the effects of mussel meal on laying hen diets. The study included 96 hens (Lohmann Selected Leghorn). Four diets with 0%, 3%, 6%, or 9% inclusion of mussel meal, replacing the same quantities of fish meal, were served (Jönsson and Elwinger 2009). At 26 weeks of age, five eggs from each treatment were collected and analyzed for internal egg quality. The different contents of mussel meal had no significant effect on production performance or egg quality parameters except from laying percentage and egg yolk pigmentation. Mussel meal concentration up to 6% tended to improve laying percentage compared with the 0% group. Yolk pigmentation increased significantly with increasing levels of mussel meal. There were no differences between the diets in fatty acid pattern regarding omega3 fatty acids docosahexaenoic acid (DHA) and dipicolinic acid (DPA) and linolenic acid in the egg yolk. Eicosapentaenoic acid (EPA) decreased significantly when fishmeal was replaced by mussel meal. Mussel meal was also tested as a high-quality protein source in broiler chicken diets, and it was found to be a good substitute for equal levels of fish meal (Waldenstedt and Jönsson 2006). It was concluded that mussel meal was a good protein source for poultry production.

231

It is well known that shellfish temporarily may be contaminated with a whole range of different biotoxins through their food intake of phytoplankton. In Swedish coastal waters the toxins belonging to the DST group are the most frequently occurring (Karlson et al. 2007). When mussel farming is used as an environmental measure it is essential that harvest can be performed regardless of any toxicity in the mussels in order to fulfill nutrient trading schemes (Lindahl et al. 2005; Jönsson and Holm 2009). A small-scale study was therefore carried out to evaluate the effects on animal health and morphology of the digestive tract of laying hens fed with three different diets: (1) a commercial feed, and diets containing (2) 15% normal nontoxic mussel meal, and (3) 15% toxic mussel meal corresponding to a concentration of 198.6 μg okadaic acid (OA) kg−1 feed. Twelve laying hens were divided into six groups and fed the experimental diets for 8 weeks. Animal health, production performance and egg quality were recorded. At sacrifice, tissue samples were prepared for histological evaluation using light microscopy (Jönsson and Holm 2009). OA did not have any adverse effects on animal health or production parameters, and no histological changes indicating disturbances in the digestive tract were observed. Internal egg quality was improved and eggs from hens fed either of the two mussel meal diets had an increased yolk color. The results above showed that mussels containing OA at a level far above the allowed limit for human consumption (160 μg kg−1) may be included in the feed without negative effects on parameters evaluated in this study. Mussels containing more that 10 times the toxin concentration used in the experiment can be used as feedstuff due to the small portion of mussel meal used in the feed. Such high toxicity in mussels is very rare in Swedish waters (Karlson et al. 2007). Results showed that some of the toxin content was lost during the drying process, probably because some of

232

Shellfish Aquaculture and the Environment

the fat was lost and with it also some of the DST toxicity (Lindahl, unpublished data). The market for organic eggs has increased considerably during recent years (Odelros 2008). Assuming that mussel meal can be produced at a large scale and at a reasonable cost compared with fish meal, there will be a market for it. Only about 5% of the meal has to be used in the poultry feeds, which means that a higher price will not necessarily become a large obstacle. Swedish organic egg producers already ask for feed containing mussel meal. Based on this great interest from the organic feed market, it has been calculated that the remainder from seafood mussel production will very soon not be enough to fill the market requirements. This has in turn triggered some of the Swedish mussel farm enterprises to start to investigate the economic and technical prerequisites for large-scale production of feed mussels. Mussel meal has been demonstrated to be as good as or even better than fish meal when incorporated as a high protein in feed, depending on its high content of protein and the amino acids methionine, cysteine. It is technically a rather straightforward process to produce mussel meal. The remaining problem seems to be the price on the feedstuff market since mussel meal, at least within the foreseeable future, will be more expensive to produce compared with fish meal. This is due to the higher cost of the mussels as a raw material for the meal production compared with fish for the fish meal production.

The use of the mussel remainder as fertilizer and biogas production The nitrogen, phosphorus, and potassium levels in the mussel remainder make it suitable to use as a fertilizer for cultivating grain (Lindahl et al. 2005). The easily decomposed shells have a liming effect and a number of micronutrients, for example, selenium, copper,

and zinc, are added to the soil. The discarded mussels used as fertilizer on farmland have given good results and are of special interest for organic farmers who cannot use commercial fertilizers. Crops increased between 25 and 50% compared with land that was not fertilized. The mussel remainder had approximately the same effect as the same amount of manure fertilizer. Since blue mussels are marine organisms and live in saline water and ions of both sodium and chloride have a negative effect on growing crops, that is, potatoes, it is important that the water inside the mussels is well drained before the remainder is spread on the farmland. Two major obstacles in using the mussel remainder as fertilizer are that there is a bad odor generated during the deterioration, and the agricultural farmers only need the mussel fertilizer during certain times of the year, while the mussel industry produces the remainder more or less continuously. To overcome these problems, composting experiments have been carried out in order to produce a “fertilizer of the sea,” which can be stored and used as needed; one which lacks the bad smell of decomposing organic material (Olrog and Christensson 2008). The mussel remainder was composted with straw or bark and the results were positive, with a shortened period of bad odor. The composted product could be stored and it was possible to guarantee quality assurance. The bark compost was visibly appealing with its dark bark and shiny shell pieces, and gardens and greenhouses could also be a future market. It has often been suggested that mussels not used as seafood or feed could be used for the production of biogas. A study by Lim et al. (2008) showed that anaerobic biodegradation is a feasible technique for the solubilization and methanogenesis of blue mussels, and seeded batch reactors of low salinity (<10 g L−1) can be employed to solve the problem of treatment and disposal of mussel wastes when this is the case. Gröndahl et al. (2009) have

Mussel farming and re-eutrophication in Sweden

233

Table 8.4 The content of some heavy metals and harmful organic substances in blue mussels in relation to limits in food, feed, fertilizer, and the Norwegian classification for environmental state (Kollberg and Ljungqvist 2007).

Substance

Food (mg kg−1 DW)

Feed (mg kg−1 DW)

Fertilizer (mg kg−1 DW)

Norwegian classification (mg kg−1 DW)

Lead Cadmium Nickel PCB (7) Dioxin

10 6.7 — — 27 × 10−6

11.4 2.3 — 0.23 1.4 × 10−6

100 2 50 0.4 —

<3 <2 <5 <0.33 <1.33 × 10−6

Sum DDT

—

—

<0.013

0.06

Blue mussel (mg kg−1 DW)

n

1.9 1.25 1.4 0.016 9.3 × 10−6* 0.83 × 10−6 0.004

72 87 73 38 8 2 8

*Older data.

evaluated the sustainability of ecological engineering methods to recover biomass nutrient resources from the Baltic Sea. They assessed the nutrient removal efficiency of algae and reed harvesting along with mussel farming in the brackish Baltic Sea and found that the mussels not were suitable for biogas production due to the high energy demand in harvesting, transportation, and biogas production, which will result in a low net energy balance. Obviously, mussels can be used for biogas production, but not just for energy production.

Risk assessment of mussels for seafood, feed, and fertilizer A comprehensive literature search was carried out on the content of harmful substances in benthic dwelling mussels sampled by a number of different monitoring programs from along the Swedish west coast (Kollberg and Ljungqvist 2007). The study included a group of heavy metals, arsenic, and some different organic substances with and without halogens. The technique analyzing organic substances has improved considerably over time and therefore only data from recent years were included. The concentrations found were compared with the limits used by Swedish authorities to control and regulate the food and feed

industry and for the use of fertilizers on farmland. Moreover, a comparison was made with the Norwegian classification limits on the environmental state, which has been set up by the Norwegian Pollution Control Authority to assess the condition of marine areas. In short, it was found that the concentrations of heavy metals were well below recommended limits for food, feed, and fertilizer for all of the investigated elements (Kollberg and Ljungqvist 2007) (Table 8.4). A likely explanation is that mussels mainly feed on phytoplankton and are thus at the second step of the food chain where the effects of any eventual biomagnification are at a low level and/or that heavy metals are not present. Cadmium is the only metal that eventually could create a risk when the mussel remainder was used as fertilizer on farmland, when the mussel remainder is used as an organic fertilizer. With the existing limits about 10 tons of mussel fertilizer per hectare of farmland, mussels could be used without any risk of biomagnification. The grain farming experiments showed that the optimal use of mussel fertilizer was in the range 5–10 t ha−1. There are many substances that have not been assigned limits for use as food, feed, and fertilizer. In this case, the Norwegian environmental classification was used and one substance, without limits that was not approved,

234

Shellfish Aquaculture and the Environment

was tributyltin (originally used to prevent fouling in bottom paint of yachts and ships). Only commercial vessels in Sweden are currently allowed to use this highly toxic compound. Concentrations of PCBs (sum 7) and dioxin in mussels were far below the suggested limits when older and analytically unsafe values were discarded. The only remaining substance in the blue mussel that may exceed the environmental limits was toxaphen, and this calls for continued attention, but again there are no regulatory limits for toxaphen in food or feed. Remains from the large variety of pharmaceuticals that enter the coastal waters through sewage have lately been given attention as a potential problem. There are, however, neither data on occurrence or concentrations of harmful pharmaceuticals in mussels or other marine organisms available, nor any limits for seafood or feedstuff. One important outcome of the risk assessment study was the recognition that a uniform system for the concentrations of the different substances found in the marine organisms and their corresponding limits for different uses (i.e., food, feed, and fertilizer) should be developed (Kollberg and Ljungqvist 2007). Condition and meat content of the blue mussel may vary considerably during the year and this must be taken into account with regard to the concentrations of harmful substances in a risk assessment. Thus, it has been recommended that a condition factor of the sampled mussels always should be reported together with the data on harmful substances (Lutz 1980). Health risks associated with consumption of virally contaminated shellfish are well documented (e.g., Lees 2000; Rippey, 1994). The symptoms of viral food poisoning are similar to those of diarrheic shellfish poisoning, but no attempts have been made to distinguish between the sources of the illnesses. The gramnegative bacteria Escherichia coli has been used as an indicator organism for fecal contamination in bivalves; however, the shellfish

industry still needs a more reliable viral indicator system (Hernroth et al. 2002). Another study was carried out to investigate human enteric virus contaminants in mussels from three sites along the Swedish west coast, forming a gradient of anthropogenic influence. Escherichia coli counts exceeded the limit for category A for shellfish sanitary safety in 40% of the samples from the in-fjord sites and it was concluded that the Escherichia coli standard for human fecal contamination seems to be an acceptable indicator of only local sanitary contamination and was not a reliable indicator of viral contaminants in the mussels. Enteric viruses were found in 50–60% of the mussel samples, and there were no pronounced differences between the samples from the three sites. To protect consumers and verify “clean” mussels, it was necessary to analyze for viruses as well. The study showed that, for risk assessment, separate modeling should be done for every specific area, with special emphasis on environmental factors such as land runoff (Hernroth et al. 2002). One way for the consumer to reduce the risk of viral food poisoning through eating mussels is of course to prepare the mussel meat well. With regard to mussels used for feed production, viral contamination will be eliminated when the mussel meat is heated, as is required for all feedstuff ingredients, during the drying procedure.

Conclusions of the Swedish experience Mussel farming is a simple, flexible, costeffective, and straightforward concept for improving coastal water quality at many coastal sites. Mussel farming can be compared with, and regarded as, open landscape feeding—only in the sea. At the same time, it is a sustainable production of valuable seafood and feedstuff products and provides coastal jobs. Since mussels are primary consumers,

Mussel farming and re-eutrophication in Sweden

and longline mussel farms are akin to floating reefs, the ecological threats are small and can be handled by careful localization of farming sites and implementation of best management practices (see Chapter 3 in this book). The use of mussel farming to combat the negative effects of eutrophication has a large potential in many coastal areas and may become a new commodity on a global scale. Trading nutrient emissions is an effective and useful tool for society to connect market economy with environmental economy and thus combat eutrophication. The complete concept of re-eutrophication, nutrient trading and mussel farming is a win-win solution for society and the environment as well as for all the involved enterprises, and consideration in other geographical regions is highly encouraged.

Literature cited Anonymous. 2003. Havet—tid för en ny strategi. Betänkande av Havsmiljökommissionen. Statens offentliga utredningar. SOU 2003:72. (In Swedish with English summary.) Anonymous. 2006a. Evaluation of Economic Instruments in the Environmental Field. The Swedish Environmental Protection Agency, ISBN 91-620-5616-6. (In Swedish with English summary.) Anonymous. 2006b. Yearbook of Agricultural Statistics. www.sjv.se (In Swedish with English summary). Anonymous. 2009. Regeringen inför ett särskilt statligt bidrag till lokala vattenvårdssatsningar som förbättrar havsmiljön (LOVA). Government Offices of Sweden, Ministry of the Environment, Memorandum 2009-03-05. (In Swedish.) Baumol, W., and Oates, W. 1988. The Theory of Environmental Policy: Externalities, Public Outlay and the Quality of Life, 2nd ed. PrenticeHall, Englewood Cliffs, NJ. Berge, G.M. 1986. Blåskjel som formiddel til regnbogeaure og verpehøner. Master thesis at Norwegian University of Life Sciences. (In Norwegian.)

235

Berge, G.M., and Austreng, E. 1989. Blue mussel in feed for rainbow trout. Aquaculture 81:79–90. Chesapeake Bay Program. 2000. www.chesapeakebay.net/content/publications/. Cloern, J.E. 2001. Our evolving conceptual model of the coastal eutrophication problem. Marine Ecology Progress Series 210:223–253. Diaz, R.J., and Rosenberg, R. 1995. Marine benthic hypoxia. A review of its ecological effects and the behavioral responses of benthic macrofauna. Oceanography and Marine Biology: An Annual Review 33:245–303. Diaz, R.J., and Rosenberg, R. 2008. Spreading dead zones and consequences for marine ecosystems. Science 321:926–929. Duinker, A., Moen, A.-G., Nortvedt, R., and Sveier, H. 2005. Utvidet kunnskap om blåskjell som fiskefôrressurs. Final report to the Norwegian Research Council, project nr.: 150109. (In Norwegian.) Edebo, L., Haamer, J., Lindahl, O., Loo, L.-O., and Piriz, L. 2000. Recycling of macronutrients from sea to land using mussel cultivation. Environment and Pollution 13:190–207. Elmgren, R. 1984. Trophic dynamics in the enclosed, brackish Baltic Sea. Rapports et Procésverbaux des Réunions. Conseil International por l’Éxploration de la Mer 183:152–169. Elwinger, K., Tauson, R., Tufvesson, M., and Hartmann, C. 2002. Feeding of layers kept in an organic feed environment. In: Wpsa, G. (ed.), Proceedings 11th European Poultry Conference, Bremen, Germany, CD-Rom. Pharma Service, Hannover, pp. 1–12. Elwinger, K., and Wahlström, A. 2000. Effects of methionine deficiency on performing of laying hens in an aviary system. Proc. XXI World’s Poultry Congress. CD rom, wpsa.com. SLU, Montreal, Canada. Grant, J., Hatcher, A., Scott, D.B., Pocklington, P., Schafer, C.T., and Winters, G.V. 1995. A multidisciplinary approach to evaluating impacts of shellfish aquaculture on benthic communities. Estuaries 18:124–144. Gren, I.-M., Lindahl, O., and Lindqvist, M. 2009. Values of mussel farming for combating eutrophication: an application to the Baltic Sea. Ecological Engineering 35(5):935–945. Gröndahl, F., Brandt, N., Karlsson, S., and Malmström, M.E. 2009. Sustainable use of

236

Shellfish Aquaculture and the Environment

Baltic Sea natural resources based on ecological engineering and biogas production. Proceeding Wessex Institute of Technology, ECOSUD 2009 8-10 July 2009 Chianciano Terme, Italy (in press). Haamer, J. 1995. Presence of the phycotoxin okadaic acid in mussels (Mytilus edulis) in relation to nutrient composition in Swedish coastal water. Journal of Shellfish Research 14:209–216. Haamer, J., Holm, A.S., Edebo, L., Lindahl, O., Norén, F., and Hernroth, B. 1999. Strategisk musselodling för att skapa kretslopp och balans i ekosystemet—kunskapsöversikt och förslag till åtgärder. Fishery Board of Sweden 6:5–29. (English summary). Hardy, R. 2009. Aquaculture feeds and ingredients: an overview. In: Burnell, G., and Allan, G. (eds.), New Technologies in Aquaculture. CRC Press, Woodhead Publishing Limited, New York, pp. 370–386. Hatcher, A., Grant, J., and Schofield, B. 1994. Effects of suspended mussel culture (Mytilus spp.) on sedimentation, benthic respiration and sediment nutrient dynamics in a coastal bay. Marine Ecology Progress Series 115:219–235. Hernroth, B.E., Conden-Hansson, A.-C., RehnstamHolm, A.-S., Girones, R., and Allard, A.K. 2002. Environmental Factors Influencing Human Viral Pathogens and Their Potential Indicator Organisms in the Blue Mussel, Mytilus edulis: the First Scandinavian Report. Applied and Environmental Microbiology 68(9):4523– 4533. Huang, J.H., Haab, T.C., and Whitehead, J.C. 1997. Willingness to pay for quality improvements: should revealed and stated preferences data be combined? Journal of Environmental Economics and Management 34(3):240–255. Jakobsen, K., and Hermansen, J.E. 2001. Organic farming—a challenge to nutritionists. Journal of Animal and Feed Sciences 10:29–42. Johansen, N.F. 2008. Ekologiskt foder—Hur kan man klara det? In Lindahl, O. (ed.), Muslingemel i stedet for fiskemel i økologiske foder till æglæggende høns, kylling og andre husdyr. Nordic Council, Copenhagen, p. 57. Jönsson, L., and Elwinger, K. 2009. Mussel meal as a replacement for fish meal in feeds for organic poultry—a pilot short term study. Acta

Agriculturae Scandinavica, Section A, Animal Sciences 59(1):22–27. Jönsson, L., and Holm, L. 2009. Effects of toxic and non-toxic blue mussel meal on health and product quality of laying hens. Journal of Animal Physiology and Animal Nutrition 94:405–412. Karlson, B., Rehnstam-Holm, A.S., and Loo, L.O. 2007. Temporal and spatial distribution of diarrhetic shellfish toxins in blue mussels, Mytilus edulis (L.), on the Swedish west coast, NE Atlantic, 1988-2005. Swedish Meteorological and Hydrological Institute, Reports Oceanography, no 35, 40. Kollberg, S. 2008. Produktion av musselmjöl för foderförsök. In: Lindahl, O. (ed.), Muslingemel i stedet for fiskemel i økologiske foder till æglæggende høns, kylling og andre husdyr. Nordic Council, Copenhagen, p. 57. Kollberg, S., and Lindahl, O. 2006. Musselmjöl istället för fiskmjöl i ekologiskt foder. The Ekhaga Foundation, www.ekhagastiftelsen.se/ eng/, project no. 2004-55. (In Swedish). Kollberg, S., and Ljungqvist, L. 2007. Musslor som livsmedel och råvara inom lantbruket. The Ekhaga Foundation, www.ekhagastiftelsen.se/ eng/, project no. 2005-48. (In Swedish.) Lees, D. 2000. Viruses in bivalve shellfish. International Journal of Food Microbiology 59:81–116. Lim, Y.-G., Niwa, C., Nagao, N., and Toda, T. 2008. Solubilization and methanogenesis of blue mussels in saline mesophilic anaerobic biodegradation. International Biodeterioation & Biogradation 61:251–260. Lindahl, O. 1995. Long-term studies of primary phytoplankton production in the Gullmar fjord, Sweden. In: Skjoldal, H.R., Hopkins, C., Erikstad, K.E., and Leinaas, H.P. (eds.), Ecology of Fjords and Coastal Waters. Elsevier Science Publishers B.V., Amsterdam, pp. 105–112. Lindahl, O. 2007. Primary production in the Gullmar Fjord. Swedish National Report on Eutrophication Status in the Kattegat and the Skagerrak. OSPAR Assessment 2007:41– 43. Lindahl, O., and Kollberg, S. 2009. Can the EU Agri-Environmental Aid Program be Extended into the Coastal Zone to Combat Eutrophication? Hydrobiologia 629(1):59–64.

Mussel farming and re-eutrophication in Sweden

Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L.-O., Olrog, L., Rehnstam-Holm, A.-S., Svensson, J., Svensson, S., and Syversen, U. 2005. Improving marine water quality by mussel farming—a profitable measure for Swedish society. Ambio 34(2):131–138. Lindqvist, M. 2008. The value of using blue mussel sea farming as a measure to remove nitrogen and phosphorous from the Baltic Sea, under the condition that the reductions are conducted in a cost effective way. Exam work no 517, The Swedish University for Agriculture, Institution of Economy. (In Swedish with English abstract.) Loo, L.-O., and Rosenberg, R. 1983. Mytilus edulis culture: growth and production in western Sweden. Aquaculture 35:137–150. Lutz, R.A. (ed.). 1980. Mussel Culture and Harvest: A North American Perspective. Elsevier Scientific Publishing Company, Amsterdam. Markowska, A., and Zylicz, T. 1999. Costing an international public good. Ecological Economics 30(2):301–316. Mirtho, S., La Rosa, T., Danavaro, R., and Mazzola, A. 2000. Microbial and meiofaunal response to intensive mussel-farm biodeposition in coastal sediments of the western Mediterranean. Marine Pollution Bulletin 40:244–252. Newell, R.I.E. 2004. Ecosystem influences of natural and cultivated populations of suspension feeding bivalve molluscs: a review. Journal of Shellfish Research 23:51–61. Odelros, Å. 2008. Ekologisk Fjäderfäproduktion. In: Lindahl, O. (ed.), Muslingemel i stedet for fiskemel i økologiske foder till æglæggende høns, kylling og andre husdyr. Nordic Council, Copenhagen, p. 57. Olrog, L., and Christensson, E. 2008. Användning av musslor och musselrester som gödselmedel i jordbruket. The Rural Economy and Agricultural Societies Report 2008:1. (In Swedish with English summary.) Pennsylvania Nutrient Management program. 2009. panutrientmgmt.cas.psu.edu/. Petersen, J.K., and Loo, L.-O. 2004. Miljøkonsekvenser af dyrking av blåmuslinger.

237

Interreg project III-A nr. GS 3041-45-02 “Blåskjellanlegg og nitrogenkvoter.” Final report 31.8.2004. (In Danish.) Riisgård, H.U., and Randløv, A. 1981. Energy budgets, growth and filtration rates in Mytilus edulis at different algal concentrations. Marine Biology 61:227–234. Rippey, S.R. 1994. Infectious diseases associated with molluscan shellfish consumption. Clinical Microbiology Review 7:419–425. Ryther, J.H., Dunstan, W.M., Tenore, K.R., and Huguenin, J.E. 1972. Controlled eutrophication: increased food production from the sea by recycling human wastes. BioScience 22:144– 152. Smaal, A.C. 2002. European mussel cultivation along the Atlantic coast: production status, problems and perspectives. Hydrobiologia 484:89–98. Söderqvist, T. 1998. Why give up money for the Baltic Sea? Motives for peoples willingness (or reluctance) to pay. Environmental and Resource Economics 12(2):141–153. Svanberg, L. 2009. Effects of mussel farming on sedimentation rates, oxygen consumption and carbon content in the underlying sediment— a case study. Master’s thesis, Department of Marine Ecology. Gothenburg University, Sweden. Contribution 354. Svensson, S. 2003a. Effects, dynamics and management of okadaic acid in blue mussels, Mytilus edulis. PhD thesis, Göteborg University, Depart. of Zoology/Zoophysiology, Sweden. Svensson, S. 2003b. Depuration of Okadaic acid (Diarrhetic Shellfish Toxin) in mussels, Mytilus edulis, feeding on different quantities of nontoxic algae. Aquaculture 218:277–291. Tietenberg, T. 2003. Environmental and Natural Resource Economics 6/E. Addison-Wesley, Reading, MA. Waldenstedt, L., and Jönsson, L. 2006. Mussel meal as a high quality protein source for broiler chickens. Proc. 12th European Poultry Conference, Verona 10–14 Sept., p. 349.

Chapter 9

Expanding shellfish aquaculture: a review of the ecological services provided by and impacts of native and cultured bivalves in shellfishdominated ecosystems Loren D. Coen, Brett R. Dumbauld, and Michael L. Judge

Introduction Aquaculture is making an ever-increasing contribution to the worldwide demand for shellfish at the same time that native “wild stock” populations have been or are significantly declining (Naylor et al. 2000; Beck et al. 2009, in press). At the same time, there has been increasing awareness of the ecosystem services that native bivalves provide as habitat, biofilters, and shoreline stabilizers (reviewed in Newell 2004; Coen et al. 2007; Grabowski

and Peterson 2007; see also Chapter 1 in this book). Habitat-forming bivalve species as those that are (1) reef-forming; (2) aggregationforming; or (3) shell-accumulating species (ASMFC 2007). Given (1) the worldwide decline of native species’ populations (Kirby 2004; Airoldi and Beck 2007; Beck et al. 2009; Gillespie 2009); (2) the associated loss of critical (“nursery”) habitats; and (3) the everincreasing worldwide contribution of cultured native and nonnative species, the beneficial services and negative impacts associated with

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 239

240

Shellfish Aquaculture and the Environment

native shellfish should be examined. This evaluation is critical since expanding aquaculture may provide many of the same services as native shellfish populations, while reducing fishing pressure on native “wild stocks” (e.g., Shumway and Kraeuter 2004; NRC 2010). Most or all native oyster species on the U.S. West Coast are significantly depressed or extirpated (e.g., U.S. West Coast, Canada, and Mexico oysters, Ostrea lurida and Ostrea conchaphila, reviewed in McGraw 2009), whereas Crassostrea virginica is either still harvested directly on the Gulf of Mexico and eastern U.S. coasts because of sufficient larval supply and settlement substrates (e.g., Coen et al. 2007; Brumbaugh and Coen 2009). In contrast to finfish aquaculture (numerous reviews), the use of molluscs as “farmed” species is generally perceived as benign, except perhaps with regard to the production of mussels on an industrial scale where significant impacts have been demonstrated quite often (e.g., Grant et al. 1995; Black 2001; Alongi et al. 2003; Asmus and Asmus 2005; Cranford et al. 2007; McKindsey et al. 2009; Paraskevi et al. 2009). In addition to the above-mentioned services, there are numerous examples where the shellfish aquaculture industry has fought for improved water quality standards, and helped to regulate and enhance waste water treatment, septic system upgrades (see also Lindahl et al. 2005; Gren et al. 2009) and the impacts of upland runoff (Chapters 8 and 9 in this book). Although numerous studies have focused on mussels grown in rope culture, associated carrying capacities, and related community and ecosystem effects, few if any directed studies have assessed molluscan aquaculture’s impacts or its parallel services to native-dominated systems (cf. NRC 2009, 2010). In this chapter, we summarize what is known about ecosystem services provided by bivalves, particularly in the United States, as well as any potential ecosystem impacts (both negative and positive), resulting from their

commercial culture. We use indirect evidence where available to gauge these effects for other cultured species, especially oysters (see also Heffernan 1999; Deal 2005; Dumbauld et al. 2009; Forrest et al. 2009; NRC 2009, 2010) and provide a brief overview of the state of our knowledge regarding natural bivalve communities, reefs and aggregations, “ecosystem services,” justification for shellfish protection, enhancement, and restoration efforts. We then include several case studies of shellfish aquaculture and discuss the direct and indirect, biotic and abiotic, and both positive and negative effects in an estuarine or marine context. For shellfish these impacts vary by (1) location of habitat (intertidal vs. subtidal); (2) extent of cultured versus natural acreage (scale); (3) other elements associated with the proposed area to be farmed (e.g., existing seagrasses, competitors, benthos, available hard substrate, exotics); and (4) latitudinal/geographical differences (cf. native or introduced species, regulations, western Atlantic, Gulf of Mexico, or Eastern Pacific). For some systems there is a wealth of general information that is transferable in part to specific sites (e.g., carrying capacity, lost habitat structure); for others sitespecific sampling and related research is required (e.g., marine mammals, flow, competitors, submerged aquatic vegetation [SAV]). All of these factors should be addressed before deciding whether bivalve aquaculture can partly or fully replace lost natural systems and the resultant effects (NRC 2009, 2010), especially given our limited understanding of these natural, and in many cases, “relic” native shellfish populations (Beck et al. 2009, in press).

Shellfish habitats, ecosystem services, and engineers Shellfish habitats—whether a living assemblage or an accumulation of dead shells— provide the necessary hard substrate for the

Impacts of native and cultured bivalves

attachment of many epifaunal species that would not be present otherwise in areas consisting of soft sediments . Along most of the world’s temperate to tropical shores, shellfish populations currently occur or once occurred in estuaries, as well as near-shore and offshore seafloor of the continental shelf (e.g., Kirby 2004; Shumway and Kraeuter 2004; Airoldi and Beck 2007; Beck et al. 2009, in press). One of the most well studied and widely occurring, habitat-forming species is the eastern oyster, Crassostrea virginica. Crassostrea virginica occurs from the St. Lawrence River, Canada, to the Atlantic coast of Argentina (Kennedy et al. 1996). Human activities, in concert with natural phenomena, have significantly affected the distribution and abundance of this oyster throughout its range as with many native habitat-forming bivalve species that are also widely harvested as a resource (see reviews in Kirby 2004; Kirby and Miller 2005; NRC 2007). Thus, they are unique in that they form “the habitat,” but are a “resource” and are impacted significantly by harvesting (e.g., Kaufman and Dayton 1997; Breitburg et al. 2000; Coen and Luckenbach 2000; Lenihan and Micheli 2000; Smyth et al. 2009). Such bivalve-generated habitats have declined precipitously due to many causes including (1) overharvesting; (2) physical disturbance by natural and anthropogenic impacts including waves and boat wakes; (3) diseases; (4) nutrient enrichment through runoff; (5) predation; (6) alteration of natural flow regimes and salinity patterns; and (7) loss of appropriate substrate for new recruits (e.g., Kennedy et al. 1996; Luckenbach et al. 1999; Coen and Luckenbach 2000; Beck et al. 2009; Volety et al. 2009). In contrast to other estuarine and marine habitats such as seagrasses, marshes, and mangroves, whose importance, functioning, and protection has been long studied (e.g., Nixon 1980; Boesch and Turner 1984; Costanza et al. 1997; Beck et al. 2001, 2003; Williams and Heck 2001; Faunce and Serafy 2006), the

241

biology and ecology of most shellfish species has primarily focused, until recently, on resource-related questions (e.g., Coen et al. 1999a, 1999b; Lenihan 1999; Luckenbach et al. 1999; Coen and Luckenbach 2000; Beck et al. 2003). Shellfish, traditionally, have only been considered a resource to be extracted (e.g., Lenihan and Peterson 1998; Coen et al. 1999b, 2007; Beck et al. 2009), and most state and federal fisheries management agencies did not view shell bottom habitats as essential fish habitat (EFH) until the late 1990s (e.g., Thayer 1992; Luckenbach et al. 1999; Coen et al. 1999b). The broader ecological value of shellfish recently has begun to gain more universal recognition (Luckenbach et al. 1999; Coen et al. 2007; Grabowski and Peterson 2007; Beck et al. 2009). The result has been new and broadened shellfish research and restoration perspectives (Luckenbach et al. 1999; Coen and Luckenbach 2000; Thayer et al. 2003, 2005; Grabowski and Peterson 2007; Schulte et al. 2009).

Complex, habitat-forming shellfish species For bivalves and gastropod molluscs, the calcareous shell functions as the protective exoskeleton forming the foundation for the above-outlined ecosystem services. Some shellfish (e.g., oysters and mussels) support significant commercial and recreational fisheries, with a subset creating unique and important three-dimensional habitats, particularly when in high densities. Using a previously developed classification (ASMFC 2007), we segregated these shellfish-dominated habitats into one of three major habitat-forming “morphotypes”: (1) reefs (a three-dimensional structure with a veneer of living and dead animals only paralleling coral reefs); (2) shell aggregations (living and dead shell); and (3) shell accumulations (dead; often referred to as shell hash) (reviewed in ASMFC 2007; Beck et al. 2009). Both

242

Shellfish Aquaculture and the Environment

bivalve and gastropod molluscs form these types of shellfish habitats, with some species falling into one or more of the above (classes 2 or 3) depending on the relative abundance of “dead” shell versus live individuals. A fourth “assemblage” type that potentially parallels many of the above-mentioned attributes are the high-density shellfish farms and associated structural hardware utilized by the aquaculture industry, along with its extensive netting and anchoring structures and devices. Members of two widespread genera, Crassostrea and Mytilus, are examples of reefforming shellfish that occur globally throughout many coastal areas (Galtsoff 1964; Sellers and Stanley 1984; Seed 1992; Seed and Suchanek 1992). Before hand and mechanized harvesting, subtidal oyster reefs in many estuaries extended meters to tens of meters above the bottom, forming extensive and complex three-dimensional reef structures that provided thousands to tens of thousands of hectares of habitat for various finfish and invertebrates (reviewed in Hargis and Haven 1999; Coen et al. 1999b; Peterson et al. 2003; ASMFC 2007; Grabowski and Peterson 2007; Dumbauld et al. 2009). Many bivalves (especially oysters such as Crassostrea virginica) have even more direct impacts, in addition to their significant filtering at higher turbidity levels (discussed in Coen 1995; Harsh and Luckenbach 1999; Hewitt and Norkko 2007; NRC 2010). These often very localized impacts influence one or all of the following: (1) recruitment; (2) growth; and (3) other organisms that live on or near the reef that seek shelter or food (Suchanek 1985; Zimmerman et al. 1989; Breitburg et al. 1995; Cummings et al. 1998; Kuhlmann 1998; Breitburg 1999; Kennedy and Sanford 1999; Lenihan 1999; Coen et al. 1999b; Beadman et al. 2004; Luckenbach et al. 2005; Tolley and Volety 2005; Rodney and Paynter 2006; ASMFC 2007; Munguia 2007; Stunz et al. 2010). Other habitat-forming species are those in the “Shell Aggregation” group. These may be

represented by scallops such as Placopecten magellanicus, a species that although not a true reef-former in the strictest sense, often occurs in sufficient densities to generate scallop shell-dominated habitats utilized by many other species (Langton and Robinson 1990; Stokesbury 2002; Talman et al. 2004). In many areas, pen shells (e.g., genera Atrina, Pinna), and other densely aggregating infaunal bivalves whose shells often protrude above the substrate, generate complex habitats with numerous species using both living animals or dead articulated shells as habitat (e.g., Perry 1936; Keough 1984; Cummings et al. 1998; Kuhlmann 1998; Norkko et al. 2006; Munguia 2007). Many estuarine and brackish water bivalve clams also occur in dense accumulations to form unique habitat assemblages (e.g., Rangia, Polymesoda, and other members of the corbiculid family). Mussels and pen shells also affect the abiotic realm. By protruding a few centimeters above the bottom, these shells create significant topographic roughness elements that substantially affect flow, creating larger areas of turbulence, thereby changing local hydrodynamics and particle transport (e.g., Officer et al. 1982; Keough 1984; Dame 1996; Cummings et al. 1998; Coen et al. 1999b; Norkko et al. 2006; ASMFC 2007; see also Chapter 10 in this book). A third shellfish habitat type, the “Shell Accumulation” group includes ocean quahogs (Arctica islandica), surf clams (Spisula solidissima), and other abundant large bivalves (e.g., Mercenaria spp., Tresus spp., and Mya arenaria) whose shells often persist long after the living organism is gone (NRC 2010). Shells of these species often accumulate on the seabed of the continental shelf and in estuaries in quantities sufficient to provide significant structure and habitat for a variety of organisms (e.g., Dumbauld et al. 1993, 2000; Auster et al. 1995; Palacios et al. 2000; Steimle and Zetlin 2000; Stoner and Titgen 2003). Recent studies and modeling efforts have focused on the decrease of available shell stocks in areas

Impacts of native and cultured bivalves

where shell is “mined” for restoration efforts. These studies have shown that extant shellfish populations are no longer generating enough new shell, and that ocean acidification may be impacting the survival of live animals and the longevity of nonliving shell (discussed in Powell et al. 2006; Mann and Powell 2007; Powell and Klinck 2007; NRC 2010). Aquaculture may be one solution for supplying the increasing amounts of shell required for restoration (NRC 2010), addressing buffering by shell dissolution from increasing global CO2 emissions, as well as declining estuarine pH (NRC 2010; Waldbusser et al. 2011). Accumulated shell also has important consequences for critical habitat of many offshore and inshore species. One hypothesis for the crash of Florida’s calico scallop (Argopecten gibbus) fishery in the 1980s is the loss of formerly significant shell accumulations at offshore nursery sites (ASMFC 2007; W. Arnold and W. Lyons, pers. comm.). A fourth habitat category (ASMFC 2007), “Shellfish Aquaculture,” has received little or no previous attention, but potential services and associated ecosystem values include most of those generated by wild stock bivalve populations (summarized in Coen et al. 2007; Grabowski and Peterson 2007; and discussed further below), and may include other “potential” and “realized” services or negative impacts that natural shellfish populations do not provide or cause. The diversity of bivalve shellfish species cultured worldwide is relatively high, with Crassostrea gigas (often referred to as the Pacific or Japanese oyster) having the widest global distribution through both deliberate and accidental introductions (reviewed in Ruesink et al. 2005; Molnar et al. 2008; Dumbauld et al. 2009; Smaal et al. 2009; Wrange et al. 2010; Chapter 14). Given the widespread occurrence and ever-increasing scale of molluscan aquaculture in intertidal and subtidal waters (see Naylor et al. 2000; NRC 2010; FAO 2009a), and the significant structural elements they provide in coastal and

243

estuarine waters, this fourth habitat type needs to be assessed in much greater detail as pointed out in several recent reviews (Dumbauld et al. 2009; NRC 2009, 2010, and references therein). Together, these four groups of shellfish habitat provide significant amounts of “abovebottom” structure not only for many mobile fish and invertebrate species but also for sessile species (Forrest et al. 2007; McKindsey et al. 2007; D’Amours et al. 2008; Mallet et al. 2009; Watson et al. 2009; NRC 2010). All four shellfish habitat reefs (aggregations, shell accumulations, and cultured species with associated “gear”) have been poorly studied (e.g., Safriel 1975; Suchanek 1985; Sebens 1991; Holt et al. 1998; ASMFC 2007), especially since many of these habitats are today only found as relics of past and widely occurring distributions (e.g., Kirby 2004; Lotze et al. 2006; Beck et al. 2009, in press). In addition to introducing new individuals to the system, bivalve mariculture operations utilize a great diversity of habitat-forming gear (i.e., ropes, rebar, netting; Figs. 9.1 and 9.2; see also Dealteris et al. 2004; Powers et al. 2007; Tallman and Forrester 2007; Munroe and McKinley 2007a; Coen et al. 2010; NRC 2010). Bivalve mariculture has other implications, such as changes in sedimentation and flow rates that are less understood (reviewed in Dumbauld et al. 2009; NRC 2009, 2010). For the remainder of this review, we focus on systems dominated by either native or cultured bivalves. We highlight parallels between native and cultured ecosystem services, while stressing direct and indirect impacts, as well as potential or demonstrated positive and negative effects, on the surrounding systems.

Shellfish ecosystem services Recently, scientists have begun to attribute numerous important ecological functions to molluscan shellfish-dominated systems

244

Shellfish Aquaculture and the Environment

Figure 9.1 Fouled subtidal clam growout cages in North Carolina. (Adapted from M. Power, Dauphin Island Sea Lab, and Powers et al. 2007.)

(A)

(B)

Figure 9.2 (A) A planted oyster restoration site in Charleston, SC. Oyster shell (Crassostrea virginica) stabilized with polypropylene mesh and rebar enforcing rods. (B) Note grunt (Orthopristis chrysopterus) caught in the deployed stabilizing mesh in Beaufort County, SC; a negative side effect of the use of larger meshes.

(reviewed in Newell 1988, 2004; Coen et al. 1999b, 2007; Grabowski and Peterson 2007; Beck et al. 2009, in press). The ecological processes that depend on the shellfish habitat described above can be thought of as “ecosystem services” (sensu Jones et al. 1994, 1997). In addition to their direct habitat-related value, shellfish provide important services for the ecosystem as a whole. These “ecosystem ser-

vices” include (1) significant filtering capacity; (2) enhanced benthic-pelagic coupling (Porter et al. 2004); (3) sediment/shoreline stabilization; (4) indirect habitat provisioning; and (5) juvenile and adult feeding areas (e.g., nursery habitats, areas of intense predation), among others. Perhaps three of the most important of these services include refuges for juvenile species (nursery areas), benthic-pelagic

Impacts of native and cultured bivalves

coupling, and shoreline protection through the reduction of erosion where operations/habitats are intertidal. Often only a single or a limited number of bivalve species form a dominant feature of inshore (estuarine) or near-shore (coastal) marine ecosystems. For example, the eastern oyster Crassostrea virginica forms living subtidal and intertidal reefs in many Atlantic and Gulf coast estuaries. Although Crassostrea virginica can occur to a depth of 30 m, they are found primarily in shallow waters less than 6 m deep to intertidal (Galtsoff 1964; Bahr and Lanier 1981; Burrell 1986; Kennedy et al. 1996; Coen et al. 1999b; ASMFC 2007). Oyster reefs, like many other reef-forming invertebrate species, are unique in that they form the actual living reef structure (e.g., Lenihan et al. 2001; J.B.C. Jackson et al. 2001; Newell 2004; Coen et al. 1999b, 2007) in estuaries, supporting a host of other organisms generally not found in surrounding sand or mud habitats (e.g., Wells 1961; Stanley and Sellers 1986; Coen et al. 1999b; ASMFC 2007; Stunz et al. 2010). We know very little about the unique ecosystem that native oysters on the Pacific coast of the United States (Olympia oyster, Ostrea equestris or Ostrea lurida) supported when they were sufficiently abundant, that is, prior to the late 1800s and early 1900s (McGraw 2009, and related papers in this Special Issue of the Journal of Shellfish Research vol. [28]). Recent work has attempted to quantify the contribution of oyster habitat to ecosystem functioning (e.g., Peterson et al. 2003; Peterson and Lipcius 2003; Grabowski and Peterson 2007). Oysters create complex habitats utilized by fish, crustaceans, bivalves, numerous other invertebrates, birds, and mammals (reviewed in Coen et al. 1999b; ASMFC 2007), potentially rivaling the role of salt marshes, seagrasses, and mangroves in nursery habitats (e.g., Glancy et al. 2003; Coen et al. 2006, unpublished data; Stunz et al. 2010). Wells (1961) surveyed oyster reefs in North Carolina and observed the presence

245

of over 300 species of plants and animals. Coen et al. (2006, unpublished data) observed over 140 finfish and macroinvertebrates species on South Carolina intertidal reefs. In the southeastern United States (portions of South Carolina, North Carolina, Georgia, and Florida), oyster reefs are a conspicuous intertidal rather than subtidal feature in most estuaries (e.g., Dame 1979; Bahr and Lanier 1981; Dame et al. 1984a, 1984b; Stanley and Sellers 1986; Coen et al. 1999a, 1999b; Van Dolah et al. 1999; Coen et al. 2004; Stunz et al. 2010). Shellfish also provide significant habitat structure for often larger, mobile (“transient”) species (e.g., Coen et al. 1999a, 1999b; Nestlerode 2004; ASMFC 2007). Given the total absence of any seagrasses in many of these southeastern U.S. estuaries, the presence of oyster reefs and related functioning is even more critical in these productive systems (Beck et al. 2001, 2003). In most estuaries, shellfish habitats exist in a landscape of more than one habitat (e.g., oysters, seagrasses, and salt marsh), together potentially providing complex utilization patterns and interactions among the numerous species (e.g., Bell et al. 1991; Eggleston et al. 1999; Micheli and Peterson 1999). We are just beginning to understand the broader ecological importance of this association (Valentine and Heck 1993; Peterson and Heck 1999, 2001a, 2001b; Wall et al. 2008; Booth and Heck 2009; NRC 2009). For example, oysters can filter up to 200 L of water per individual per day through direct ingestion, feces ejection, and particle rejection (pseudofeces). Oysters remove both algae and suspended particles while processing water, thereby improving water clarity and quality (Dame et al. 1984b; Cressman et al. 2003; Nelson et al. 2004; Newell 2004; Grizzle et al. 2006, 2008; Newell et al. 2007; NRC 2010). As a result, bivalves can positively impact light penetration and seagrass photosynthetic rates, which can enhance nutrients and potentially benefit seagrasses in shallow waters (Peterson and

246

Shellfish Aquaculture and the Environment

Heck 1999, 2001a, 2001b; Newell and Koch 2004; Valentine and Heck 1993; Cerrato et al. 2004; Wall et al. 2008; Booth and Heck 2009).

2004; Bain et al. 2007; Brumbaugh and Coen 2009).

Importance of oyster reef habitats Habitat considerations In contrast to the extensive subtidal oyster beds in estuaries such as in the Chesapeake Bay, Delaware Bay, off the Mississippi River (LA), or Apalachicola Bay (FL) in the southeastern United States (e.g., southern North Carolina, South Carolina, Georgia), oysters are found primarily in the intertidal zone to depths of 1–3 m (3–9 ft), which is in part dictated by local tidal ranges, among other factors (Bahr 1976; Bahr and Lanier 1981; Burrell 1986). Intertidal oyster beds typically become established where salinity is moderately high, food supply is sufficient, and siltation is not excessive. Intertidal oyster reefs consist of vertical clusters built upon a fragile matrix of live and dead shell surrounded by fine sediments (Anderson et al. 1979; Bahr and Lanier 1981; Dame et al. 1984a, 1984b; Burrell 1986; Coen et al. 1999a; Lenihan and Micheli 2000; Lenihan and Peterson 2004). Since many oyster larvae remain in the water column for up to 3 weeks after spawning occurs, larvae are subject to distribution throughout an estuary by tidal currents. Since the final larval stage of reef-forming species crawl on the bottom to search for suitable hard substrate upon which to permanently attach, its location is critical to the continued survival and regeneration of these complex reef habitats. Attachment sites include almost any hard surface, for example, other living oysters, oyster shell, rocks, docks, tree limbs, and pilings. It is during this final process that aquaculture can flourish in areas where sufficient natural larval supplies are available. However, in many areas, settlement is insufficient and requires hatcheries to supplement or totally supply larval recruits (Kennedy 1996; Luckenbach et al. 1999; Mann and Evans

The eastern oyster Crassostrea virginica, once a dominant feature of most Atlantic and Gulf coast estuaries, has drastically declined in many areas across the United States (Lenihan 1999; J.B.C. Jackson et al. 2001; Kirby 2004). Once valued primarily as a resource, oysters are now also recognized as key elements of many estuarine ecosystems (Coen et al. 1999a, 1999b; Luckenbach et al. 1999; Grabowski and Peterson 2007). Oysters create complex habitats utilized by fish, crustaceans, bivalves, numerous other invertebrates, birds, and mammals (Coen et al. 1999b; Lehnert and Allen 2002; Tolley and Volety 2005; ASMFC 2007; Stunz et al. 2010). During feeding, oysters and other bivalve molluscs can filter large quantities of water (Lindahl et al. 2005; Borthagaray and Carranza 2007; Buschbaum et al. 2009; Gren et al. 2009), improving water clarity and quality while transferring nutrients from the water column to the benthos (Newell 1988, 2004; Dame 1996; Dame et al. 2001). Declines in oyster populations are associated with decline in critical habitat, shifts from benthic to pelagically dominated communities, adverse effects on other species, reduced water quality, and changes in ecosystem dynamics (Rothschild et al. 1994; Newell et al. 2007). Complex reef-forming invertebrate species have been centers of biodiversity throughout geologic time (Kiessling et al. 2010). It is now widely recognized that oyster reefs are valuable habitat for a wide variety of organisms (e.g., Luckenbach et al. 1999; Coen et al. 1999b, 2007; Rodney and Paynter 2006; ASMFC 2007; Grabowski and Peterson 2007; Stunz et al. 2010) and that oyster resources/ habitats can be enhanced through directed oyster reef restoration programs (e.g., Coen and Luckenbach 2000; Luckenbach et al.

Impacts of native and cultured bivalves

2005). As a result, oyster reef restoration programs have expanded rapidly throughout the United States in recent years. Unfortunately, the shell hash needed to rebuild reefs is generally scarce in those areas where restoration is needed most (Bushek et al. 2004; Coen et al. 2007). Intertidal oysters bordering salt marshes can prevent erosion of tidal edge habitats (discussed in Meyer et al. 1997; Piazza et al. 2005; ASMFC 2007; Grabowski and Peterson 2007; Beck et al. 2009, in press; Coen et al., unpublished data) such as mangroves or marshes reducing wave impacts in tropical to subtropical zones (e.g., Danielsen et al. 2005; Kathiresan and Rajendran 2005; NRC 2007; Feigin et al. 2009). For example, in the southeastern United States, reefs often fringe Spartina and Juncus marsh edges in creeks and rivers, forming a unique association that protects these critical marsh habitats from natural (e.g., tidal or wind-driven waves) and anthropogenically derived (e.g., boat wakes) erosion (e.g., Meyer et al. 1997; Meyer and Townsend 2000; Piazza et al. 2005; ASMFC 2007; Goodwin 2007; Beck et al. 2009, in press; Coen et al., unpublished data).

Use of shellfish habitats by invertebrates and finfish We now know that shellfish-generated reefs have significantly greater vertical, threedimensional relief when compared with the surrounding two-dimensional bottom, thereby greatly enhancing biodiversity (reviewed in ASMFC 2007; NRC 2010). As discussed above, eastern oyster Crassostrea virginica reefs harbor diverse communities largely restricted to the reef structure or other hardbottom habitats, versus adjacent nonreef habitat. This enhanced vertical relief is of major importance, with implications for assessing habitat value for managed species, and is responsible for a revised management policy regarding these important biogenic reefs

247

(see ASMFC 2007; Grabowski and Peterson 2007; Beck et al. 2009, in press). While there is a dearth of information on vertebrate or invertebrate use of shellfishdominated reefs (reviewed in Coen et al. 1999b; ASMFC 2007; Grabowski and Peterson 2007; NRC 2009, 2010), new quantitative information on species associated with natural reefs or other complex, three-dimensional structure is growing (Luckenbach et al. 2005; NRC 2010; Coen et al., unpublished data). Observational data from intertidal and subtidal shellfish habitats in North America and Europe has also been summarized into the worldwide status of oysters (Airoldi and Beck 2007; Beck et al. 2009, 2011; Buschbaum et al. 2009; Reise et al. 2009; Smaal et al. 2009). More information is currently available regarding the impacts of avian predators and associated negative impacts on finfish aquaculture than for intertidal or subtidal shellfish operations (see Comeau et al. 2009).

Impacts to shorebirds and mammals Several studies have been conducted to examine the effects of shorebird foraging or human disturbance in intertidal habitats on mariculture. In Canada, Yasue (2005) found that human disturbance of migratory shorebirds causes subtle, but important, changes in foraging by semipalmated plovers (Charadrius semipalmatus) and least sandpipers (Calidris minutilla). In NewJersey, Burger (1994) has found that human disturbance can have negative effects on piping plover (Charadrius melodus) foraging and suggested that it is important to maintain the historical diversity of coastal habitats in order to alleviate the negative effects of humans on piping plover foraging. In northern California, Oregon, and Washington, seasonal monitoring of shorebird and waterfowl densities has provided invaluable baseline information, which allows evaluation of the effects of humans on these species (e.g., Colwell 1993;

248

Shellfish Aquaculture and the Environment

Colwell and Landrum 1993; Dodd and Colwell 1996; Warnock et al. 1998; Stempien 2007). Other studies have attempted to associate prey availability with shorebird abundances on intertidal mudflats in South America, and throughout Europe (e.g., Pienkowski 1983; Backwell et al. 1998; Triplet et al. 1999; Sanchez et al. 2006). Numerous birds use intertidal sand- or mudflats as feeding and resting sites during their normal activities or migrations (e.g., Quammen 1981, 1982, 1984; Baird et al. 1985; Norris et al. 1998; Caldow et al. 2004; Goss-Custard et al. 2004; Connolly and Colwell 2005; Stempien 2007; Stillman et al. 2007; Kraan et al. 2009). For shorebirds associated with aquaculture, concerns have included one or more of the following: (1) limited access to feeding due to cages and other gear; (2) changes in benthic sediments and associated communities; (3) potential entanglements (see Fig. 9.2B above) or ingestion of mesh or other materials; (4) noise related to activities during growout (i.e., planting, harvesting, and related maintenance); and (5) potential for migratory disruptions (e.g., Kelly et al. 1996; Kaiser et al. 1998; Hilgerloh et al. 2001; Connolly and Colwell 2005; Forrest et al. 2009; Godet et al. 2009; Atkinson et al. 2010). Unfortunately, too little is known at this point to draw any general conclusions. For marine and terrestrial mammals, we know even less about their positive or negative impacts related to aquaculture than for shorebirds as discussed above (e.g., Nash et al. 2000; Kemper et al. 2003; Lloyd 2003; Read et al. 2006; Roycroft et al. 2006; Forrest et al. 2009; NRC 2010). Recently, this has become an important consideration related to permitting, as National Park Service and National Research Council studies were undertaken to assess the impacts of intertidal oyster mariculture in California (NRC 2009, 2010), where marine mammals were an important factor in the deliberations. This rarely has been a con-

sideration for the eastern oyster on the U.S. East and Gulf coasts, perhaps because oyster aquaculture is only now being taken up in these areas. These potential issues need to be addressed as part of the Environmental Impact Statement (EIS) assessment and permitting process in areas where a potential “conflict” may arise between naturally occurring species and bivalve aquaculture (FAO 2009b). The ecology of these systems is too variable and the diversity of impacts too diverse and complex to transpose the limited results from other areas to new sites without additional assessments, especially in areas where “user” conflicts may occur (e.g., Forrest et al. 2009; NRC 2010).

Restoration of natural habitats and aquaculture’s potential role Most subtidal oyster reefs currently found in “Approved” shellfish harvesting areas (i.e., open to direct harvesting based on human health standards) rarely extend even a meter off the bottom while still providing one or more services for economically and ecologically important species (Bahr and Lanier 1981; Sellers and Stanley 1984; Dame 1996; Breitburg 1999; Coen et al. 1999a, 1999b; Lenihan 1999; Posey et al. 1999; Harding and Mann 2001; Peterson et al. 2003; Nestlerode 2004; Powell et al. 2008; Schulte et al. 2009). Intertidal oyster reefs are still common in many areas, in contrast to subtidal populations of conspecifics within the same ecoregion (sensu Beck et al. 2009, in press). This is mostly due to the fact that individuals from these intertidal habitats are often not sought after for the raw bar trade. Large- and small-scale restoration of both subtidal and intertidal oyster habitats is ongoing along most Atlantic and Gulf coast states (discussed in Coen and Luckenbach 2000; Burrows et al. 2005; Beck et al. 2009, in press; Brumbaugh and Coen 2009; ORET

Impacts of native and cultured bivalves

2009; Powers et al. 2009; Schulte et al. 2009). Determining the success of these restoration efforts is critical to optimizing our use of the requisite limited shell resource, and to be costeffective for restoring reef habitats. Consensus, however, on what constitutes a successful reef restoration project currently does not exist (e.g., Coen and Luckenbach 2000; Thayer et al. 2003, 2005; Coen et al. 2004; Brumbaugh et al. 2006; Powers et al. 2009; Hadley et al. 2010). The most commonly used metric of success has been the presence of market-sized (e.g., 3-in.-long by shell height) oysters (Luckenbach et al. 2005; Schulte et al. 2009). Use of a size metric derived from a commercial harvesting focus (and associated fishery regulation) is inappropriate if the goal of restoration is to restore ecological function and not a marketable resource (see Breitburg et al. 2000; Coen et al. 2004; Luckenbach et al. 2005; Thayer et al. 2005). State agencies have planted shell or relayed oysters to enhance fisheries for many years (Coen and Luckenbach 2000; Coen et al. 2004, 2007, 2010), but current large-scale (e.g., Army Corps of Engineers in the Chesapeake Bay) to smallscale community restoration efforts are nearly all targeted at reestablishing the associated ecological value of oyster reefs. Recent reef restoration efforts have also focused on the educational benefits derived from the process at the community-based scale and not the numbers of bushels that may be harvested from the reefs at some later time. In the future, restoration efforts need to focus on success criteria that are connected explicitly to the implicit goals of the restoration effort (Coen et al., unpublished data). Small- or large-scale oyster enhancement/ restoration approaches can create tens to hundreds of hectares of shellfish beds by adding material above an otherwise shell-less, softbottom habitat. This may increase settlement of larvae when recruits are not limiting, and can be enhanced by relaying “cultch” with juvenile oyster spat that is already attached, or

249

by adding adult broodstock to reefs. Additionally, oysters are moved or “relayed” into growout areas with lower disease or higher growth and survival rates as in the Delaware Bay (Kennedy 1996; Southworth and Mann 1998; Luckenbach et al. 1999; Coen and Luckenbach 2000; Smith et al. 2005; Brumbaugh et al. 2006; Powell et al. 2008; Brumbaugh and Coen 2009).

Aquaculture-based systems Bivalve species commonly cultured in the United States Current west coast intertidal oyster and clam mariculture Bivalve shellfish have been harvested for subsistence from estuaries along the U.S. West Coast for thousands of years (Naylor et al. 2000; Beck et al. 2009, in press), but extensive harvest of native oysters (Ostrea lurida) began with European colonialism in the mid-1800s (Baker 1995). Similar to the fate of the East Coast’s Crassostrea virginica, these oysters were gradually depleted due to a combination of overharvesting, extreme natural events, and the lack of shell replacement to maintain a substrate for natural recruitment (e.g., Kirby 2004; Ruesink et al. 2005). Eastern oysters (Crassostrea virginica) and later Pacific oysters (Crassostrea gigas) were introduced from the U.S. East Coast and Japan, respectively, in order to overcome this decline (Townsend 1896; Steele 1964; Lindsay and Simons 1997; Robinson 1997; Shaw 1997). Of particular interest for this chapter and related ones (see Chapter 14 in this book) is the potential impact (both obvious and more subtle) of a novel oyster’s (e.g., Pacific oyster) ecosystem services and resulting positive and negative effects after its introduction and expansion locally, as well as worldwide (Reise 1998; Drinkwaard 1999; Black 2001; Naylor et al. 2001; Ruesink et al.

250

Shellfish Aquaculture and the Environment

2005; Thieltges et al. 2006; McKindsey et al. 2007; Kochmann et al. 2008; Molnar et al. 2008; Sousa et al. 2009; Markert et al. 2010). On the U.S. West Coast, the introduction of Pacific oysters initiated widespread aquaculture and farming operations for this species and other bivalves. Recognizing the importance of the shellfish industry in the state of Washington, tidelands were deeded directly to growers for use. This encouraged potentially “sustainable” farming instead of a traditional fishery and Washington became the leading shellfish producer on the West Coast. Eastern oysters never reproduced successfully and were essentially replaced with the Pacific oyster (Crassostrea gigas). Juvenile Pacific oysters or seed were imported annually from Japan (NRC 2004), but this oyster also became naturalized in some locations where water temperatures were sufficient for spawning (e.g., Willapa and Dabob Bays in Washington, USA; Pendrell Sound and Ladysmith Harbor in British Columbia, Canada). Oyster farming operations became integrated with the advent of hatchery technology in the early 1980s (Nosho and Chew 1991; Conte et al. 1994). The Manila clam, Tapes philippinarum, which was inadvertently introduced with the Pacific oyster, and the much larger native geoduck, Panope generosa, are currently two of the most actively farmed clams on the West Coast. Both species are raised in hatcheries and seeded into estuarine intertidal culture grounds. It should be noted that mussels are also farmed on the West Coast, but here we restrict our discussion to oysters and clams because they co-occur in intertidal estuarine areas and mussels are mostly grown in suspended culture. The functional role of these cultured intertidal bivalves is similar to that of native bivalves in West Coast estuaries, and those described for Eastern oysters and others elsewhere, that is, they influence material processes and physical structure. Culture practices shape both the spatial and temporal scale of

this role as well as their interactions with and impacts upon other estuarine resources (reviewed in Dumbauld et al. 2009). The native oyster Ostrea lurida prefers to live in low intertidal and subtidal areas, where it is less exposed to extreme temperatures and probably formed extensive low relief aggregations before it succumbed to overexploitation and other anthropogenic influences (see Fig. 9.3; Baker 1995; Gillespie 2009; White et al. 2009). In contrast, Crassostrea gigas tolerates a wider range of temperatures and can withstand greater turbidity resulting in survival at much higher tidal elevations and over a broader spatial range, from more riverine areas to the estuary mouth (Fig. 9.3). Cultured Crassostrea gigas successfully grow where they have been planted. In locations such as Willapa Bay, they cover expansive areas of the upper intertidal zone (over 4625 ha or 20.4% of the bay’s intertidal area). Growers typically plant seed at relatively low densities and do not allow individuals to aggregate through thinning clusters in order to achieve optimal growth for later harvesting (Fig. 9.4C). Nevertheless, this oyster still forms extensive reeflike habitat in areas where it is left to “naturalize.” The resulting reefs are three-dimensionally complex with some vertical relief, much like the reefs formed by Crassostrea virginica on the East and Gulf coast estuaries of the United States (Fig. 9.4G). Oysters are harvested in a 2- to 4-year cycle, with harvesting practices themselves also directly influencing the environment (Tallis et al. 2009). Culture practices for this oyster also involve deploying artificial structures within which the deployed oysters are placed in bags on racks, clustered on stakes, or strung along polypropylene ropes supported by PVC stakes in the expansive tidal flats (Fig. 9.4D). These structures can enhance erosion and/or sediment deposition depending on local water flow conditions. They also provide numerous attachment sites for other sessile fouling organisms (see NRC 2009, 2010).

Impacts of native and cultured bivalves

1800

2010

251

Ostrea lurida

Crassostrea gigas Zostera japonica Zostera marina

1800 Eelgrass Oyster reef Oyster culture

2010

Figure 9.3 Intertidal distribution of eelgrass and oysters in a hypothetical U.S. West Coast estuary in 1800 versus present. Native oysters Ostrea lurida formed low-profile reefs in low intertidal and subtidal areas in 1800 where native eelgrass Zostera marina was also distributed (top). Pacific oysters Crassostrea gigas are now typically spread out in beds slightly higher in the intertidal where they intermingle with native eelgrass and in some cases the introduced eelgrass Zostera japonica at a higher elevation. This is depicted at the estuarine landscape scale (bottom) in an estuary where Pacific oysters have become naturalized and also form reefs closer to the head of the estuary where their larvae are retained.

Native littleneck (Leucoma staminea) and butter clams (Saxidomus gigantea) are also generally found at lower intertidal elevation than the cultured, nonnative Manila clams (Tapes philippinarum). All infaunal clams influence material processes in a similar way, but in contrast to oysters, they provide significantly less, physical structure above the sediment. A significant difference in cultured clam operations is the added physical feature provided by predator-exclusion devices

(e.g., netting and tubes; Fig. 9.4E and 9.4F; Powers et al. 2007; Tallman and Forrester 2007), substrate modification (beach gravelling), and the recurring disturbance related to planting, maintenance, and harvesting operations (Simenstad and Fresh 1995; NRC 2010). Numerous models have been built to assess the carrying capacity of localized systems for bivalve aquaculture (see Chapters 1 and 6 in this book), and where they have been tested

(A)

(B)

(C)

(D)

(E)

(F)

(G)

Figures 9.4 Intertidal habitats and shellfish culture in U.S. West Coast estuaries. (A) Open unstructured mudflat often dominated by burrowing shrimp; (B) structured eelgrass, Zostera marina, habitat; (C) bottom culture of Pacific oysters Crassostrea gigas forming structured habitat intermingled with eelgrass; (D) longline culture of Pacific oysters; (E) Manila clam Tapes philippinarum culture with predator netting; (F) geoduck Panope generosa culture with tubes for predator protection; and (G) Pacific oyster reef where they have become naturalized in Willapa Bay, WA.

252

Impacts of native and cultured bivalves

and verified, results generally suggest that cultured bivalves have measurable effects on water properties (e.g., Heral 1993; Grant et al. 1995, 2005; Leguerrier et al. 2004; Dowd 2005; Duarte et al. 2005, 2008; Drapeau et al. 2006; McKindsey et al. 2006; Ferreira et al. 2007; Fulford et al. 2007, 2010; Newell 2007; Ferreira et al. 2009; North et al. 2010; NRC 2010). The presence and magnitude of this effect depends on filtration ability of the bivalves (i.e., clearance rates of individuals and population size), location, water properties, and residence time of the water mass being filtered. These models have rarely been applied to intertidal shellfish aquaculture in West Coast estuaries (primarily Oregon and Washington, USA, and Canada) where water properties are largely driven by the coastal ocean and tidal forcing. These estuaries generally have large tidal prisms and relatively short resident times (i.e., are rapidly flushed) relative to the filtration capacity of the native bivalves present or for nonnative, cultured bivalves, and thus only very localized effects of filter feeding have been demonstrated (reviewed in NRC 2010). An exception is Willapa Bay, where the extensive culture of oysters and clams has the potential to regulate phytoplankton production in the system (Banas et al. 2007; Wheat et al. in review). Hydrography and corresponding phytoplankton concentrations in Willapa Bay and in many other West Coast estuaries are different from those of larger and perhaps more typical river-dominated systems on the U.S. East Coast. This results in a gradient of high food concentrations near the mouth of the estuary and lower values in the upper reaches of that estuary. Three competing hypotheses for these West Coast systems could explain this pattern: (1) physical mixing of nutrient-rich oceanic and poorer, river waters; (2) a longer residence time in the upper estuary relative to estuaries on the Gulf or East coasts; or (3) significant grazing by farmed oysters localized near the mouth of these estuaries

253

(Dumbauld et al. 2009). Banas et al. (2007) found that phytoplankton concentrations declined more rapidly with distance from the mouth of Willapa Bay than would be expected due to simple mixing, and that the difference was consistent with the capacity of oysters to filter out significant portions of the phytoplankton community. This is possible because a large portion (>80%) of the water never fully circulates over the intertidal flat, but is exported and reimported into the estuary due to the local complexities of tidal circulation, and the extent of the tidal flats in this estuary (Banas et al. 2007). More recent field measurements confirm that oysters are in fact able to remove >10% of the phytoplankton that circulated over a 100-m distance across a tidal flat versus a control, where no oysters were present (W. Wheat and J.L. Ruesink, unpublished data). Thus, hydrography and plankton dynamics of the estuarine system must be accounted for when defining carrying capacity and quantitatively characterizing the impact of adding or maintaining bivalve aquaculture. Comparing the historical situation in Willapa Bay with current conditions is also interesting as native Olympia oysters were less abundant or at least represented a lower biomass (i.e., 2.5× lower than current Crassostrea gigas populations; Ruesink et al. 2005) and occupied a different location within the estuary (Fig. 9.3, lower intertidal and generally further within the estuary). Another unknown component is the abundance of native clams and other filterfeeders that were historically present on the tidal flats where Crassostrea gigas is currently cultivated. Anecdotal information suggests past densities of native clams were also much higher than current population densities (L. Bennett and R. Sheldon, pers. comm.). Recent anthropogenic nutrient inputs have caused significant eutrophication within many estuaries such as the Chesapeake Bay, making the filtering capability of bivalves a potentially attractive “ecosystem service” (Chapters 8 and 9 in

254

Shellfish Aquaculture and the Environment

this book; Coen et al. 2007; Newell et al. 2007; Huang et al. 2008). To date, bivalve filtration has not been demonstrated as a direct benefit of aquaculture on the West Coast (Harbin-Ireland 2004; NRC 2010), in part due to high flushing rates, and also the lack of overlap between aquaculture and areas where eutrophication is now becoming an issue. This could change as the services provided by filterfeeders become more valuable, allowing aquaculture to be extended into areas whose waters are currently classified as nonshellfish-growing areas (see Brumbaugh and Toropova 2008; Beck et al. 2009, and discussed elsewhere here). Measurable effects of cultured bivalves on sediment properties depend on the density of the organisms and local system characteristics. Effects are expected to be local, depending on water flow, which disperses biodeposits (e.g., Callier et al. 2006; Callier et al. 2008; Weise et al. 2009), and on existing sediment characteristics at the aquaculture site (e.g., grain size; see Nizzoli et al. 2006; Mesnage et al. 2007; see also Chapter 10). Intertidal culture of clams and oysters typically occurs in areas with sandier substrates and relatively high flow, but it is more difficult to quantify the effects of biodeposition and to distinguish it from other sources of sediment deposition and nutrient addition than in suspended culture. This is in part due to the fact that most existing studies are based on suspended culture. Minimal evidence collected to date suggests that the mechanisms are similar to those documented for suspended culture and that high densities of oysters can cause reduced grain size and increased organic content (Rumrill and Poulton 2004; E.L. Wagner, unpublished data). Geoduck clams have been shown to raise pore water ammonia concentrations (J.L. Ruesink and K. Rowell, unpublished data). When evaluating the effects of localized biodeposition on the surrounding benthic invertebrate community, it is difficult to separate these effects from those of simply adding the

shellfish and associated structures, as well as the effects of “pulsed” disturbances such as harvesting- or maintenance-related activities. A significant amount of research has been conducted on the effects of adding shellfish, particularly oysters, to open, unstructured tidal flats in U.S. West Coast estuaries or on SAV such as eelgrass (Zostera marina; Dumbauld et al. 2009). The direct impact of such activities on eelgrass is a primary concern to management and regulatory agencies because this habitat has been shown to be so important for numerous estuarine finfish and invertebrates, and because seagrass densities are declining on a global basis (E.L. Jackson et al. 2001; Heck et al. 2003; Orth et al. 2006; Heck et al. 2008; Hughes et al. 2009; Waycott et al. 2009). That said, the ecosystem services of shellfish aquaculture, both as a structured habitat and other attendant services, have yet to be examined sufficiently (but see HarbinIreland 2004; Brumbaugh and Toropova 2008; Beck et al. 2009; Dumbauld et al. 2009; Forrest et al. 2009; NRC 2010). However, given time and more rigorous experimental studies, we may see more examples that elucidate these positive ecosystem services for the shellfish aquaculture industry, especially given shellfish growout sites may also provide optimal growth conditions for eelgrasses by adding sufficient nutrients in otherwise oligotrophic estuaries or by removing sufficient seston to improve light conditions in areas too turbid to allow seagrasses to take hold and thrive (reviewed in NRC 2010, and references therein). Although enhanced eelgrass (Zostera marina) growth due to enhanced bivalve biodeposition has been documented along the U.S. East Coast (cf. Peterson and Heck 2001a, 2001b; Wall et al. 2008), this same positive result does not seem to be the case for what has been observed in aquaculture systems on the West Coast, at least as has been examined to date. In Willapa Bay, the presence of oysters appears to have no effect on eelgrass growth (Ruesink et al. 2009; Tallis et al. 2009; Wagner

Impacts of native and cultured bivalves

et al., unpublished data), in part because nutrients are already close to optimum levels for seagrasses, but also because oysters do not substantially influence light levels, which have been shown to be the primary limiting factor for eelgrass in most West Coast estuaries studied (Thom et al. 2008). In another study, Japanese littleneck clam (Tapes philippinarum) production areas are generally found higher in the intertidal zone than native eelgrass. However, the introduced eelgrass Zostera japonica does interact with the cultured clams in Willapa Bay, WA (Tsai et al. 2010). There the presence of Zostera japonica reduces clam “condition” (or the ratio of meat weight to shell size) by altering water flow and thus food supply, but clams have no demonstrable reciprocal effect on eelgrass growth. Another cultured bivalve, the geoduck (Panope generosa), does have seasonal effects on eelgrass growth in south Puget Sound, WA. This observed impact may be due to the reduced seasonal density of eelgrass in the presence of clams during the summer and not biodeposition from clams per se (Ruesink and Rowell 2010). Some harvest techniques associated with oyster aquaculture such as harvest-related dredging reduce eelgrass densities directly by removing plant biomass. Additionally, observed seagrass recovery rates vary spatially within these estuaries, with slower rates in areas with softer, muddy sediments (up to 4 years) than for those in areas that have naturally sandier (coarser) sediments or for that matter those that are left undisturbed by dredging. Surveys and manipulative dredging studies suggest a gradient of impact from harvesting (recovery) technique on eelgrass. Ranked from worst to least impact: harvest dredging > narrow-spaced longlines > widely spaced longlines > handpicked ground culture (Tallis et al. 2009). Additionally, simply planting oysters at dense concentrations (>20% ground cover) will displace some eelgrass and change both cover and density, but results appear to be site specific (e.g., Tallis et al.

255

2009; E.L. Wagner, J.L. Ruesink, and B.R. Dumbauld, unpublished data). Studies have shown that recruitment of eelgrass seedlings varies by location within Willapa Bay and with different aquaculture methods (Wisehart et al. 2007). Seedling recruitment was lowest in areas that had just been dredged, low in longline culture areas, but surprisingly high in dredged areas 1 year later, suggesting fairly robust recovery rates and that competition with adult plants may likely play a role in seedling growth and survival. Evidence suggests that asexual reproduction (rhizome branching), which is important for local-scale recovery from disturbance, was relatively high at most sites, whereas sexual reproduction, which could be important for recovery of larger areas, varied substantially in locations across Washington State (see Ruesink et al. 2009). Eelgrass is less abundant overall in aquaculture beds (from 30% to 35% cover; Tallis et al. 2009); however, based on either individual shoot or ramet metrics and at the coarser population level, it appears to be able to coexist and survive with oyster aquaculture at multiple sites and over multiple years in Willapa Bay. Use of U.S. West Coast estuarine habitats by fish and invertebrates is much more limited than that published for both the East and Gulf coasts (reviewed in Coen et al. 1999b; ASMFC 2007, and papers therein) and appears to depend on spatial scale and organismal mobility. Researchers have generally found abundant and diverse communities of both benthic and epibenthic invertebrates associated with on-bottom oyster aquaculture (Fig. 9.4C), similar to those documented in adjacent natural eelgrass habitats (Fig. 9.4B). In Pacific Northwest estuaries, aquaculture-associated assemblages are also more diverse than those found in open, unstructured mud- or sand-flat areas, which are often dominated by burrowing shrimp (Fig. 9.4A; Simenstad and Fresh 1995; Trianni 1995; Hosack et al. 2006; Ferraro and Cole 2007).

256

Shellfish Aquaculture and the Environment

A special case exists in Willapa Bay and Grays Harbor, WA, where the pesticide carbaryl (better known as “Sevin,” 1-naphthyl methylcarbamate) is currently used to control two species of burrowing shrimp that are viewed as significant “pests” by the shellfish aquaculture industry. This practice has been shown to act as a significant, short-term disturbance by causing direct mortality to many of the benthic invertebrates present, but having few long-term (>60 days) effects, other than removing the shrimp. Burrowing shrimp themselves may be viewed as “ecosystem engineers” because they significantly rework the structure of the associated benthic community via their extensive “bioturbation” (e.g., Posey 1986; Dumbauld et al. 2001). When treated with pesticide, the shrimp-dominated community is then replaced, at least for a culture cycle or two, with above-sediment structure in the form of oysters and even eelgrass (e.g., Dumbauld and Wyllie-Echeverria 2003). Among large, more mobile nekton, some species show loose habitat associations due to the presence of aquaculture operations, while others appear to be uneffected. In Willapa Bay, juvenile Chinook salmon (Oncorhynchus tshawytscha) and English sole (Parophrys vetulis) were found across habitats (eelgrass, oyster aquaculture, and mudflats), while other finfish such as tubesnouts (Aulorhynchus flavidus) were clearly associated with eelgrass (e.g., Hosack et al. 2006; Dumbauld et al. 2009). Separate studies conducted in Willapa Bay and Grays Harbor estuaries have shown that mature Dungeness crab (Metacarcinus magister) utilize unstructured muddy areas to feed and rock crab (Cancer productus) utilized cultured oysters, while young crab of both species prefer shell deposits and oyster aquaculture areas over eelgrass and especially unstructured, shrimp-dominated habitat for protection (e.g., Dumbauld et al. 2000; Holsman et al. 2006). Data from field enclosures and laboratory mesocosms suggests that juvenile salmon seek refuge in eelgrass habitats

(e.g., Semmens 2008; Dumbauld et al. 2009), but these functional associations with intertidal benthic habitat, especially shellfish aquaculture habitat, are less studied for other species (reviewed in NRC 2010). Structured habitat, such as aquaculture sites, can be used for feeding sites or as protection from larger predators (e.g., Bell et al. 1991; Heck and Crowder 1991; Coen et al. 1999b). As introduced above, both oyster and clam aquaculture involve the addition of significant structures such as longlines, poles, and bags for raising the shellfish off-bottom and tubes and netting for predator protection (e.g., Hecht and Britz 1992; Everett et al. 1995; Kaiser et al. 1998; Heise and Bortone 1999; Dealteris et al. 2004; O’Beirn et al. 2004; Munroe and McKinley 2007a; Powers et al. 2007; Tallman and Forrester 2007). Studies suggest that these affect water flow and biodeposition, while providing novel attachment sites for organisms, increasing the shading of seagrasses, and influencing the behavior of nekton, particularly those that are structure oriented and/or feed on fouling organisms (e.g., pipefish, pile perch, kelp surfperch; Everett et al. 1995; Simenstad and Fresh 1995; Thompson 1995; Rumrill and Poulton 2004; Weschler 2004; Munroe and McKinley 2007a, 2007b; Whiteley and Bendell-Young 2007; D’Amours et al. 2008; Dumbauld et al. 2009). Bivalves have been actively cultured in many U.S. West Coast estuaries such as Willapa Bay for nearly 100 years or more. At the present scale, shellfish aquaculture seems more sustainable than other human activities such as coastal development and pollution, which degrade and can even eliminate estuarine function and potentially undermine resilience (reviewed in Dumbauld et al. 2009). Management decisions about how to classify estuaries and maintain ecosystem services as provided by both native and cultured bivalves in West Coast estuaries and other areas should therefore consider both temporal and spatial scales. Cultured Crassostrea gigas have

Impacts of native and cultured bivalves

assumed a similar role to that played by the native Ostrea lurida prior to human intervention, but key differences exist, including planting and harvesting cycles, associated management practices and structures, as well as scale and location of these culture operations in the estuary. While there have been few landscape-level approaches to studies of bivalve shellfish aquaculture in West Coast estuaries (but see Carswell et al. 2006), we hypothesize about the case for Willapa Bay, WA, where shellfish culture operations are relatively extensive at nearly 13% of the total estuarine area (Dumbauld et al. 2009). In comparison, native (Olympia) oysters covered 7.5% of the total estuarine area (based on historical maps; Collins 1892), but their extrapolated occurrence was at a lower tidal elevation and distance further from the mouth of the estuary than the majority of the current shellfish aquaculture areas (Fig. 9.5A). Though they likely formed vertical aggregations of “clustered” oysters that might be called “reefs,” native oysters may have had a similar profile to current aquaculture beds because individuals and even clusters were much smaller. Thus, the role of cultured bivalves as material processors is potentially similar to that of native oysters (NRC 2009, 2010). However, their location closer to the estuary’s mouth and higher within the intertidal zone suggests that they may be processing phytoplankton before it reaches locations further up the estuary where native oysters were once abundant (Fig. 9.5B). The role of other filterfeeders such as native clams and burrowing shrimp, which exclude bivalves at some locations due to their “bioturbation,” is unknown but likely fluctuated over time as well. Cultured oyster habitat overlaps with approximately 43% of eelgrass habitat (mostly native but also nonnative Zostera japonica). This potentially provides more habitat for recruitment of species, such as juvenile Dungeness crab and English sole (Parophrys vetulus), than did native oysters that also overlapped with eel-

257

grass. Forty-five percent of the area currently covered by eelgrass was once native oyster habitat (Fig. 9.5A). These hypotheses assume that estuarine bathymetry has not changed. Borde et al. (2003) suggest that the area where eelgrass can now grow has increased by 22% through time. The functional value of large, undisturbed eelgrass meadows and unvegetated tidal flats versus “mixed” landscapes of patchy habitats, including shellfish beds with edges and corridors, needs to be examined at the appropriate landscape scale. This may be an area where innovative practices and best management practices (BMPs) developed by growers in association with scientists can be applied to conserve and even enhance the functional value of these shrinking estuarine habitats.

Associated impacts (positive and negative) Most mariculture operations remove all or most of the shell, along with the meats derived from the waters in which the bivalves were farmed. On the U.S. West Coast, the aquaculture industry operates nearly all bivalve harvesting operations, with little or no wild stock-based operations, so that shell is most often recycled efficiently for reuse. In contrast, during most of the nineteenth through twentieth centuries a great deal of the Crassostrea virginica shellstock was removed from estuaries in the Gulf of Mexico and Atlantic coast of the United States and not necessarily replanted. These operations removed the critical carbonate-based shell habitat (oysters, scallops, clams, etc.) that, under pre-European settlement accumulated, dissolved or was reduced through normal taphonomic processes (discussed in Tevesz and McCall 1983; Donovan 1991; Andersson et al. 2003; Gutiérrez et al. 2003; Powell et al. 2006; NRC 2010). In many ways, harvesting of the entire

258

Shellfish Aquaculture and the Environment

(A)

(B)

Native oysters 1892

(C)

Oyster and clam culture 2005

Eelgrass 2005

Figures 9.5 Estuarine habitat maps for Willapa Bay, WA, showing (A) the distribution of native oysters from a map created in 1892; (B) the distribution of cultivated Pacific oyster beds in 2006; and (C) the distribution of eelgrass (mostly Zostera marina, but some Zostera japonica) in 2005.

organism makes this a “put and take fishery” without returning a significant portion of the shell back into the system. Since bivalve molluscs are harvested as both the habitat and the resource, its removal significantly impacts future generations that require clean new substrate for recruiting larvae to settle upon. One of the most critical findings from the many restoration efforts conducted with Crassostrea virginica in the last decade is that the most successfully restored subtidal reefs are those that have sufficient vertical relief off the bottom for both food, flow, and fewer extended hypoxic events (discussed in Lenihan and Peterson 1998; Lenihan 1999; Coen and Luckenbach 2000; Coen et al. 2007; Schulte et al. 2009).

One critical discussion regarding both natural and cultivated bivalves that needs to be further addressed is whether the carbonaterich deposits consisting of both live and dead animals are carbon sinks or carbon sources. This issue needs to be addressed in greater detail before it can be resolved to everyone’s satisfaction as we discuss carbon sequestration and reduction of ocean acidification as a potentially important ecosystem service of shellfish habitats (e.g., Brewer 1997; Andersson et al. 2003; Caldeira and Wickett 2003; Feely et al. 2004; Gazeau et al. 2007; Salisbury et al. 2008; Doney et al. 2009; Borges and Gypens 2010; Hopkins et al. 2010; NRC 2010; R. Newell, pers. comm.). One major ecosystem service that aquaculture may fulfill

Impacts of native and cultured bivalves

is required shell for wild stock and related restoration efforts (NRC 2010). Ocean acidification may become a serious impediment to potential shell “stores” in estuarine waters (Waldbusser et al. 2011). Bivalve aquaculture often has positive effects for the surrounding seagrass communities, enhancing light through reduction in turbidity, and adding nutrient-rich biodeposits (reviewed in Haven and Morales-Alamo 1966; Luckenbach and Orth 1999; Williams and Heck 2001; Newell et al. 2002, 2005; Mallet et al. 2006, 2009; ASMFC 2007; NRC 2009, 2010, although some disagree with this conclusion; Smith et al. 2009). Seagrasses alone can enhance water clarity through removal of suspended sediments as water is baffled through blades (e.g., Hemminga and Duarte 2000; Beck et al. 2001, 2003; Williams and Heck 2001; Agawin and Duarte 2002; Larkum et al. 2006; McGlathery et al. 2007). Through their efficient filtering process, bivalve molluscs can remove significant portions of all of the total suspended sediments carried in the overlying water column. However, in some estuaries where suspended sediments are low relative to colored dissolved organic matter (CDOM), the impact of filter-feeding bivalves may have little or no impact on light levels (Corbett 2007; L. Coen and E. Milbrandt, pers. obs.). Additionally, seagrass seeds or vegetative shoots may be entrained directly or indirectly captured in both natural bivalve systems or in and around aquaculture operations where currents are often reduced. The presence of native bivalve populations also has been shown to have positive effects on submerged vegetation in temperate and subtropical-tropical systems (discussed in Castel et al. 1989; ASMFC 2007; Dumbauld et al. 2009; NRC 2009, 2010) where many seagrass-epiphyte communities are phosphorus and nitrogen limited (e.g., Johnson et al. 2006). Shellfish may enhance the supply of these nutrients through processes such as biodeposition. It has been suggested that the

259

positive effects of bivalve biodeposits on seagrass “fertilization” are more likely to play a role in oligotrophic waters than on those with relatively high available nutrients (Castel et al. 1989; Reusch et al. 1994; ASMFC 2007; Carroll et al. 2008; Dumbauld et al. 2009; Tallis et al. 2009; NRC 2010). Mariculture operations generally take place in “Approved” waters, away from coastal development and are associated high bacterial and nutrient levels. These waters are approved for direct shellfish harvest, that is, where shellfish are primarily grown for sale, while restoration for other “ecosystem services” such as filtration and habitat and broodstock enhancement can take place often in more eutrophic locations (cf. the Nature Conservancy’s Long Island Sound and Great South Bay goals; C. LoBue, pers. comm.). Though there are exceptions, on the U.S. East Coast where rivers supply most of the nutrients, mariculture locations occur in more oligotrophic waters where the fertilization effect could take place, while on the West Coast background nutrients are already high and supplied by the near-shore coastal ocean so the fertilization effect is less likely. Although many positive interactions have already been discussed for shellfish mariculture and associated seagrasses, and to a lesser extent macroalgae, many negative impacts or concerns have been expressed by seagrass researchers and permitting agencies charged with the protection of “potential” or “realized” seagrass populations. These have been discussed previously for shellfish aquaculture in general (Pillay 1992; Kaiser et al. 1998; Heffernan 1999; Kaiser 2000, 2001; Black 2001; E.L. Jackson et al. 2001; Ruesink et al. 2005, 2006; Hosack et al. 2006; ASMFC 2007; Tallman and Forrester 2007; Wisehart et al. 2007; Richardson et al. 2008; Dumbauld et al. 2009; Forrest et al. 2009; NRC 2009, 2010; Tallis et al. 2009) and include (1) co-opting space where seagrasses might otherwise expand their coverage through time (Everett et al. 1995); (2) extreme reduction of

260

Shellfish Aquaculture and the Environment

fertilization rates caused by cultured bivalve biodeposition (Huang et al. 2008) and epiphytic fouling or excessive macroalgal growth associated with enhanced nutrient availability (e.g., DeCasabianca et al. 1997; Hauxwell et al. 2001; Munroe and McKinley 2007a, 2007b; Powers et al. 2007); (3) physical disturbance from planting and harvesting associated with normal fishing or mariculture operations (Tallis et al. 2009); (4) enhanced competition from biofouling and other species as a result of the introduction of novel hard substrates related to aquaculture operations and deployment or accumulation of dead shell from these extended operations (Beal and Kraus 2002; Costa-Pierce and Bridger 2002; Orth et al. 2002; Erbland and Ozbay 2008; Lu and Grant 2008; Kimbro et al. 2009); (5) changes in both physical and chemical sediment characteristics resulting from changes in flow due to the extensive outplanting of aquaculture-related gear (Soniat et al. 2004; Kelly et al. 2008; see also Fig. 9.4 and the case study in this chapter); (6) past and present spraying of insecticides related to the control of burrowing decapod shrimp populations (e.g., reviewed in Feldman et al. 2000; Dumbauld et al. 2001, 2006, 2009); (7) introduction of direct seagrass competitors (Zostera japonica, probably the only documented invasive seagrass; Harrison and Bigley 1982; Posey 1988; Baldwin and Lovvorn 1994); (8) enhanced structure may cause trophic cascades and negatively affect seagrass populations (e.g., Heck et al. 2000a, 2000b, 2006; Duffy et al. 2003; Inglis and Gust 2003); (9) introduction of novel species through the use of oyster shell (discussed in Bushek et al. 2004; Cohen and Zabin 2009); (10) relocation of shellfish stocks and transport of harmful algae (e.g., Carriker 1992; Hégaret et al. 2008; Heil 2009; Lewitus and Coen, pers. obs.); (11) impacting access of birds, marine mammals, and other species to areas that would be simple, two-dimensional mudflats otherwise ; and (12) potential shifts in dominant ecosys-

tem structure (e.g., Crassostrea Gigas; see “Uniqueness of West Coast Aquaculture” section above; see also Michael and Chew 1976; Chew 1990; Carlton and Mann 1996; Reise 1998; Ruiz et al. 2000; Black 2001; Ruesink et al. 2005; Thieltges et al. 2006; McKindsey et al. 2007; Kochmann et al. 2008; Molnar et al. 2008; Cohen and Zabin 2009; Dumbauld et al. 2009; Sousa et al. 2009; Markert et al. 2010). Problems related to biodeposits (as first noted and reviewed by Dame 1996) have generally been associated with subtidal mussel operations in areas with poor flushing rates (e.g., Grant et al. 1995; Miller et al. 2002; Crawford et al. 2003; Cranford et al. 2007; Mallet et al. 2009; McKindsey et al. 2009; Chapter 10 in this book). Rarely have biodeposit accumulations been a problem in intertidal growout activities in well-flushed areas (e.g., discussed in the “Uniqueness of West Coast Aquaculture” section above; Dumbauld et al. 2009; NRC 2010). Given the extensive umbrella of knowledge under which largescale aquaculture now operates, site selection criteria and permitting most likely reduce the likelihood of such negative effects (e.g., Tenore and Gonzalez 1975; Tenore et al. 1982; Dame 1996; Wildish and Kristmanson 1997; Callier et al. 2008, but see Deal 2005), with most mariculturists striving to operate under more sustainable practices (see Chapter 3 in this book).

Case study: potential short- and long-term impacts of high-density intertidal hard clam aquaculture in southeastern U.S. tidal creeks Background At present, culture of the northern quahog (Mercenaria mercenaria) occurs in more U.S. states than any other native bivalve species under culture. Although market prices fluctuate greatly, overall U.S. market value for these clams is currently among the highest. Direct

Impacts of native and cultured bivalves

and indirect ecosystem effects of raising these clams at the elevated densities necessary for commercial success have not been well studied (e.g., Doering et al. 1986; Nugues et al. 1996; Cerrato et al. 2004; Tyler 2007). Some studies suggest that elevated bivalve densities can have important impacts on many of the abovementioned system attributes (e.g., Cloern 1982; Officer et al. 1982; Cohen et al. 1984; Asmus and Asmus 1991, 2005; Dame, 1993, 1996; Dame and Libes 1993; Newell 2004; ASMFC 2007; Cerco and Noel 2007). This species is typically grown in shallow subtidal or intertidal seafloor areas that are highly visible, and often adjacent to seagrass habitats, which impacts site selection, local permitting, and conflicts over seagrass issues (NRC 2009, 2010). Intertidal aquaculture areas with tidal ranges ≥1–2 m are complex benthic systems and difficult to study since sediments are subject to daily aerial exposure where microphytobenthos respond to light and other cues. For example, intertidal marine sediments are “biostabilized” by diatom films within the surface (Stal 2010). Although not the primary goal of a past research effort in South Carolina, one objective was to examine the direct and indirect effects of high-density hard clam mariculture on the adjacent inshore benthic ecosystem (Coen et al. 2000). Results showed that the deployment of hundreds to thousands of clam culture pens, each with tens of thousands of clams, had the potential to affect (1) local hydrodynamics, (2) sediment characteristics, (3) associated benthos, (4) food quality and quantity, and (5) the carrying capacity of the surrounding ecosystem. At that time, hard clam culture occurred on low intertidal mudflats in small tidal creeks where dense oyster reefs (Crassostrea virginica) co-occurred (Coen et al. 2000; Judge et al. 2000). In addition to clam growth experiments, several other experimental field studies were conducted to (1) evaluate food and flow regimes and the effects of localized food deple-

261

tion on observed clam growth; (2) examine how clam culture pens and related activities directly and indirectly affect ecosystem attributes of tidal creeks (e.g., C/N ratios, total organic carbon [TOC], sediment % organics, sediment grain size); and (3) examine how clam culture might affect creek communities (e.g., water column and benthic chlorophyll a, infauna, and stable isotope ratios of deployed seed clams over time). The study site was located just off the Kiawah River, Kiawah, SC (32°37.41′N, 80°06.60′W; Fig. 9.6) in an intertidal creek with shallow mudflats (<0.5 m in depth during low tide), interspersed with intertidal oysters on either side of the creek bank. Experimental plots were adjacent to and within a large commercial operation with approximately 600,000 clams in 400 circular predatorexclusion pens (0.30-m high, diameter = 2.44 m, area = 4.68 m2). They were typically exposed during mean low tide. At the site overall water depths at high tide often exceeded 1.8 m or more. Free stream tidal (flow) rates were typically high, driven by a 100% change every 12 h. Pens were approximately 0.5–2.5 m from one another within a given area. Densities of newly outplanted seed clams (5–10 mm in shell length) were approximately 4274 clams m−2 (approximately 20,000 individuals per pen). At a size of around 20 mm, clams were typically thinned out to 855 clams m−2 (4000 individuals per pen). During the study, the larger overall area often had over 41 million clams in 2300 pens, with other areas having up to 17 million additional clams in over 800 pens. Overall, Atlantic LittleNeck ClamFarms, Inc. (referred to subsequently as “ALC”) at one time had more than 7000 pens in South Carolina waters, with over 70 million clams planted within them (Judge et al. 2000; J. Manzi, pers. comm., 1997). This study focused on four locations with increasing distance from the above ALC site: (1) between and within pens; (2) 1–50 m away (three sites seaward and three sites

262

Shellfish Aquaculture and the Environment

Figure 9.6 Large-scale hard clam growout in South Carolina. Note hundreds of pens at one site near Kiawah, SC (each 4.68 m2 in area).

Figure 9.7 Small experimental cages (0.25 m2, with 6.4-mm mesh) adjacent to clam pens (see also Fig. 9.6).

downstream); (3) a control site at the creek entrance that was 340 m upstream of the above ALC site; and (4) a reference site without any clam aquaculture. At locations 1 and 2 (listed above), small experimental cages (0.25 m2, with 6.4-mm mesh) were also deployed to examine clam growth at three different densities.

Effects on sediment and infauna Sediment attributes were impacted by the presence of the pens, but the effects tended to be evident only within pens (Figs. 9.7 and 9.8). Even at short distances from the pens, sediment changes were not evident or predictable. Sediment organics (% at ignition), % silt, and

Impacts of native and cultured bivalves

263

Sediment Grain Size Analysis % Organics ± 1 SE

16

(A)

A

12 8

B

4 0

% Sand ± 1 SE

100 90 80 70 60 50 40 30 20 10 0

% Silt ± 1 SE

Inside pens

50 45 40 35 30 25 20 15 10 5 0

Outside pens

(B)

B A

Inside pens

(C)

Outside pens

A

B

Inside pens

% Clay ± 1SE

16

(D)

Outside pens

A

12

B

8 4 0 Inside pens

Outside pens

Location Figure 9.8 Results of a July 7, 1998, sediment grain size analysis across study sites; data combined across sites. Sediment cores were 3.5 cm in diameter and 5 cm deep. Inside pens (n = 16) were significantly different from all combined nonpen locations (n = 29) for each attribute (P < 0.001). SE, standard error.

sometimes % clay were much higher inside pens, while % sand tended to be higher outside pens. Absolute values of sediment parameters varied slightly over time, but the rank order of the results over time was quite similar. For example, organic content inside the pens aver-

aged 8.8% in year 1 collections, and 13.2% in the year 2 collections, a difference of 30% from those samples collected outside pens. In comparison, Mojica and Nelson (1993) found an increase in silt- and clay-sized particles, as well as an increase in organic content in close proximity to commercial hard clam growout

264

Shellfish Aquaculture and the Environment

bags. The shift in particle size may affect clam growth indirectly, as Grizzle (1988) noted that growth was slowest in finer sediments and fastest in coarser, sand-dominated sediments. Although nutrient characteristics of the sediments were not examined, investigators have found both positive and negative impacts from various shellfish farming practices (e.g., Holmer 1991; Gilbert et al. 1997). Within a given location, no small-scale (<50 cm) differences in total benthic biomass were detected between insides and outsides of pens. To examine larger-scale differences in total benthic abundance, data were pooled to compare among locations. One-way analysis of variance (ANOVA) tests revealed no significant differences among locations (P = 0.58, degrees of freedom [d.f.] = 38). Linear regression analysis, examining the relationship between total biomass and distance from pens, also revealed no significant relationship (r2 = 0.02, P = 0.43). Although total biomass of benthos does not appear to be related to proximity to pens, difference in the abundance of faunal taxa may be influenced by the presence of pens or associated changes in local microhabitat (e.g., sediment type or organic content, flow), but they were beyond the scope of this project (see Weston 1990; Hammel and Webb 1999). Chlorophyll a levels of intertidal sediment were several orders of magnitude higher than water column values. Results from both sediment chlorophyll samplings revealed significantly higher levels of chlorophyll a inside versus outside the pen area. Levels were approximately 20% higher inside pens than outside pens in May (6000 μg mL−1 sediment) and 50% higher in July (11,000 μg mL−1 samples). This difference corresponds to the time period during which site water column temperature and salinity were highest and dissolved oxygen levels were lowest. Seasonal variability in water column chlorophyll levels was observed with chlorophyll peaking in May, probably a result of initial spring blooms. Overall, levels of chlorophyll were highest in sediments with a high

proportion of silts or clays and organic matter as reported by Light and Beardall (1998, see also Cahoon et al. 1999).

Effects on phytoplankton abundance Concentrations of chlorophyll a in the water column were used as a surrogate for food quantity. Water samples were collected at slack high tide at all of the study sites at approximately monthly intervals using a horizontal Wildco water sampler, then filtered following standard fluorometric methods for quantifying chlorophyll a (μg L−1; see Coen et al. 2000; Grizzle et al. 2006, 2008). Water column chlorophyll a levels among sites increased and decreased with seasonal spring and summer phytoplankton blooms. Highest chlorophyll a levels were observed during the summer at all sites. Despite small-scale spatial variability, no consistent differences among adjacent locations were found. In order to determine within-site effects on variability of suspended food, the potential for depletion of water column phytoplankton at four different spatial scales was assessed by collecting water from different locations simultaneously within our field site using a vertical sampling apparatus or “sipper” (Judge et al. 1993). Experiments included (1) food depletion (or enhancement) as a parcel of water traveled the entire creek length from the control (creek entrance) site to within the center of the pens (340 m downstream) and as sampled at fixed heights above the substrate (1, 5, 15, and 45 cm); (2) food depletion as a parcel of water traveled 150 m across the entire ALC clam pen site (over approximately 400,000 clams); (3) food depletion as a parcel of water traveled through an individual clam pen housing approximately 3000 clams (sampled 1 m upstream from a pen and within that same pen); and finally, (4) small-scale (ca. 50 cm) food depletion, sampling over our smaller growth cages at densities of 5, 50, and 200 clams per 0.25-m2 cage (Coen et al. 2000; Judge et al. 2000).

Impacts of native and cultured bivalves

Across the entire creek, only a transitory chlorophyll a spike was observed during a flooding tide in June. Chlorophyll a concentrations were temporally elevated at very shallow water depths, independent of location. As the water depth increased, chlorophyll a levels declined and were not different among sites or sampling heights.

265

Simultaneous water samples were collected upstream to examine changes in chlorophyll a across the ALC pen site, within and downstream of pens at the four sampling heights mentioned above. Overall, there were few differences between upstream and downstream locations, or vertically among the four heights above the bottom (Fig. 9.9).

Food Depletion Experiments: 8/14/97 Sipper Results 10

Chl a (μg L–1) ± 1 SD

9

10 1st collection, 2 h after low tide

8

8

7

7

6

6

5

5

4

4

3

3

2

2

1

1

0 10 9

Chl a (μg L–1) ± 1 SD

9

2nd collection, 3 h after low tide

0 1.0

5.0 15.0

3rd collection, 4 h after low tide

10 9

8

8

7

7

6

6

5

5

4

4

3

3

2

2

1

1

0

1

5

4th collection, 5 h after low tide

15

45 West 50 m Between pens East 23 m

0 1 5 15 45 Height above bottom (cm)

1

5 15 45 Height above bottom (cm)

Figure 9.9 Summary of results of “sipper” sampling, Kiawah study sites, August 14, 1997. Four sampling heights were used for the four collections on a flooding tide. (Nota bene: some sampling heights absent due to the shallow water of the early flood.) Chlorophyll a (Chl a) concentrations were quantified fluorometrically. SD, standard deviation.

266

Shellfish Aquaculture and the Environment

A second experiment, conducted in May, quantified chlorophyll a concentrations (1) within a single pen; (2) in the pen area; and (3) upstream of pens. As before, no consistent depletion of chlorophyll a concentrations was observed. A third comparison of food levels within a pen and an area 1 m upstream of that pen was run in June and only a transitory elevation of chlorophyll a was documented during the initial flood. In general, no sustained reduction within a pen or among multiple sampling heights was observed. A fourth comparison, based on samples collected in June, measured potential food reduction across the smaller experimental cages (5, 50, or 200 clams per 0.25 m2), and similar to the experiments described above results exhibited no upstream versus downstream differences. In all cases, no consistent depletion of food levels arose at any of these scales, with the exception of the transitory elevation during the initial flood, most likely due to a resuspension of surficial sediments and benthic microalgae (Coen et al. 2000; Judge et al. 2000).

Effects on water flow Flow regimes in aquatic systems can be complex, especially when associated with threedimensional structures (reviewed in Nowell and Jumars 1984; Denny 1988; Denny and Wethey 2001). Water flow in the field was measured at two temporal scales. First, a Sontek (YSI, Inc., San Diego, CA) acoustic Doppler velocimeter (ADV; field model) was used to characterize current speed and direction at our field site via high frequency (≤25 MHz) readings of water flow and vertical mixing (x-, y-, z-axes). Current velocity was measured inside individual clam pens, aisles between pens, and at various distances upstream of pens. The ADV sampled variations in water flow speed along each dimensional axis up to 25 per second, within a small sampling cylinder (9-mm length × 6-mm diameter) whose height above the substratum could be positioned by the experimenter.

Water flow speed measurements were generally conducted on flooding tides, occasionally extending into the early ebb tide. On each sampling date, up to 13 heights above the bottom were sampled to characterize the benthic boundary layer conditions. These vertical velocity profiles allowed for the generation of shear velocity (u*) calculations via logarithmic profiling (Eq. 9.1) and Reynolds shear stress (Eq. 9.2) methods. When the flow direction varied within a benthic boundary layer profile (e.g., adjacent to obstruction created by the pens), only the Reynolds shear stress method could be calculated. u ( s ) = u∗ / k ln [( s − do ) / so ]

(9.1)

u∗ = [ τ R / ρ] , where τ R = ρu ′ w ′

(9.2)

0 .5

Benthic boundary layer sampling was conducted at two locations (340 m and 50 m) upstream of the ALC study site for 3 h during a flooding tide. Maximum near-surface water flow speeds approached or exceeded 30 cm s−1 at both locations. Vertical velocity profiles were constructed at the control creek entrance site and at a location 50 m away. At both locations, flow direction did not vary with sampling height above the bottom. Shear velocities (u*) for the control creek location at 5 cm above the bottom (Reynolds shear stress method) ranged from 1.205 cm s−1 (initial flood) to 1.715 cm s−1 (peak flood). Shear velocities (u*) at the site 50 m upstream (log profile method) ranged from 0.306 cm s−1 (late flood) to 1.557 cm s−1 (peak flood). Benthic boundary layer sampling between the pens (i.e., within the ALC study site) was conducted on three occasions for a total of 5.5 h during two flooding tides and an additional 6.5 h from the initial flood through early ebb tide. Maximum near-surface water approached 20 cm s−1 at this site. Vertical velocity profiles were constructed for three components of the tidal cycle, including peak flood. At this location, flow direction varied dramatically with

Impacts of native and cultured bivalves

267

Characterization of Turbulent Flow via Reynolds Shear Stress

Sampling Height (cm)

100

Control Site Pen Aisles Inside Pens

80 60 40

Top of Pens 20 0 0.0

0.5

1.0

2.0

1.5

2.5

3.0

–1

Shear Velocity (u ; cm s ) *

Figure 9.10 Comparison of turbulent flows at three sites (control, pen aisle, and within pens). Shear velocities calculated at several heights above the substrate via Reynolds shear stress method. ALC pens were 30 cm tall and indicated by dashed lines (---).

sampling height above the bottom, especially at the height of the clam pens. Shear velocities (u*) for this location at 5 cm above the bottom (Reynolds shear stress method) ranged from 0.388 cm s−1 (initial flood) to 1.123 cm s−1 (peak flood) (Coen et al. 2000; Judge et al. 2000). Benthic boundary layer sampling within a caged pen (within the ALC study site) was conducted for 3 h during a flooding tide. Maximum near-surface water flow velocities surpassed 20 cm s−1 above the top of the pens. Vertical velocity profiles extending into the ALC pens were constructed for peak flood components of the tidal cycle. At this location, flow direction again varied with sampling height above the bottom with noted shifts at the approximate ALC pen height (Fig. 9.10). Shear velocities (u*) for this location at 5 cm above the bottom varied from 0.225 cm s−1 (initial flood) to 0.197 cm s−1 (peak flood). Overall, the high-frequency, short-term flow characterization suggested that the presence of the pen structure alters both the mixing intensity and the horizontal direction of the flow patterns. Relative to the control location away from any pens (which forces the water

to move around or over the pens), the shear velocity (u*) near the bottom is reduced by 50% in the aisle between pens or reduced by 90% within a pen (Fig. 9.10). Thus, at the height of a feeding clam, pens may drastically reduce the food particle flux. Second, dental plaster (molded into 1.5-cm diameter × 10-cm length) cylinders or “clods” were deployed at different locations upstream, downstream, and within our field site. These plaster elements dissolve slowly, so they can be deployed for 7–10-day periods, integrating differences in flow rates over a longer timescale than measured with the current meter (weeks vs. minutes) could be made. This method allowed for the simultaneous monitoring of flow patterns among several growth locations not possible with a single ADV (Yund et al. 1991; Judge et al. 1992, 1993; Judge and Craig 1997; Giotta 1999, but see Porter et al. 2000). Sides of cylinders were coated with a polyurethane varnish to inhibit erosion, thereby ensuring that all dissolution would occur solely on the surface area of the cylinder top. Thus, mass loss was approximated by a linear function.

268

Shellfish Aquaculture and the Environment

Long-term patterns differences in water flow among locations where clam growth was also measured were estimated by dissolution of dental plaster cylinders. A total of 270 clods were deployed at multiple locations upstream, downstream, and within our field site, including within clam pens. Replicate clod cards were always deployed in pairs, glued with silicone cement atop a rectangular PVC plate (10 cm × 5 cm) and positioned 5 cm above the substrata. After retrieval, cylinders were removed from their holders, rinsed of sediment, dried at 60°C for 48 h, allowed to acclimate to ambient lab temperature and humidity for 48 h, and reweighed for mass lost (Coen et al. 2000; Judge et al. 2000). Variability in flow among locations (Fig. 9.11) were analyzed as an incomplete block ANOVA with deployment date as the blocking variable and mass lost as the dependent variable. Overall, there was a significant difference in mass loss among locations (ANOVA, P < 0.001), although several intermediate locations overlapped in plaster loss. The three locations common to all (control, 50 m, and between pens), food supply, sediment analysis, and ADV measurements proved to be significantly different (Coen et al. 2000; Judge et al. 2000). Moreover, the rank orders of these sites both qualitatively and quantitatively matched the shear velocity differences illustrated by the ADV. In addition, the plaster dissolution within pens exhibited a 25–33% reduction relative to outside (ANOVA, P < 0.001), demonstrating the extent of flow speed loss by the cage mesh itself. The use of the ADV and the plaster cylinders allowed water flow patterns to be quantified at two different temporal scales (minutes vs. weeks) and both measures exhibited strong concordance. Both current meter and clod card data revealed that clam farm pens significantly altered flow (and other site characteristics) within the tidal creek. Overall, the structural presence of cages imparted profound changes in the hydrody-

namic regimes within and around clam pens, thereby altering numerous sediment attributes (i.e., grain size and chlorophyll a concentrations). Cage-induced mixing caused a localized decoupling of the benthic boundary layer, which can dramatically affect the temporal variation of resuspended algal food supplies. Sediments generally were enriched in the organic fraction, but lower in sand, silt, and clay adjacent to the aquaculture pens. Sediment chlorophyll a (an indirect metric for food quantity) was significantly enhanced within the pens. Despite differences in mixing and sediment attributes induced by the clam pens, however, total sediment infaunal biomass (exclusive of Mercenaria) was not adversely impacted by high-density clam pens. Availability of suspended food varied among sampling dates, but there was little evidence for food depletion within the water column or across the growth cages. The only regular pattern was a transitory chlorophyll a spike via resuspension during the initial stages of a flooding tide (i.e., water depths less than 20 cm). Hydrodynamic patterns (measured at timescales from seconds to weeks) affecting delivery of suspended food showed that clam farm pens reduced mean water speed and mixing by an order of magnitude within a cage, while the caging material itself created localized regions of intense shear stress (Coen et al. 2000; Judge et al. 2000).

Longer-term and larger-scale concerns The case study was completed in 2000 at which time ALC had deployed more than 7000 pens on local South Carolina mudflats, with over 7 × 107 clams planted among them (Coen et al. 2000; ALC, pers. comm., 1997). The relevant findings included differences in sediment inside versus outside of growout pens, with interiors generally enriched in the organic fraction, but with lower coarse sands

Overall Cold Card Results 0.6

Deployed 8/12/97–8/19/97 (#1) n = 4 Aisle and Within Pens Combined n = 2 All Other Locations

0.5 0.4 0.3 0.2 0.1 0.0

Mean Dissolution Rate (g day–1) ± 1 SD

s s k 1 5 ol 57 en en st st ee ntr st Cr fP We ide P We o Co We nt Ins Fro

0.4

le

Ais

st

Ea

1

st

Ea

5

st

Ea

23

Deployed 10/31/97–11/13/97 (Control 10/31/97–12/8/97) #2 n = 4 Control and Inside Pens n = 2 All Other Locations Combines

0.3 0.2 0.1

*

0.0 s s k 1 5 ol 57 en en st st ee ntr st Cr fP We ide P We Co We to s n In Fro

0.4

le

Ais

st

Ea

1

st

Ea

5

st

Ea

23

Deployed 4/8/98–4/22/98 #3 n = 8 for West 57, West 5, and Inside Pens n = 7 for Control n = 4 for Creek

0.3 0.2 0.1 0.0

s k 1 5 ol ns 57 en st st ee ntr st Pe Cr We ide P We of Co We t n Ins Fro

le

Ais

0.4

st

Ea

1

st

Ea

5

st

Ea

23

Deployed 6/24/98–7/7/98 #4 (Control 6/24/98–7/8/98) n = 26 for Pens n = 7 for Control n = 6 for Aisle and West 57 n = 5 for Front n = 4 for West 5

0.3 0.2 0.1 0.0 s s k 1 5 ol 57 en en st st ee ntr st Cr fP We ide P We o Co We nt Ins Fro

le

Ais

st

Ea

1

st

Ea

5

st

Ea

23

Location Figure 9.11 Overview of results of dental plaster dissolution from four deployments. A new batch of “clods cards” was used for each deployment. *Control clod cards for deployment #2 were not discovered until ca. 20 days after the retrieval of the other clods.

269

270

Shellfish Aquaculture and the Environment

Figure 9.12 Abandoned cage pens left out for over six years near Kaiwah, SC. These intertidal substrates were heavily fouled by Crassostrea virginica and other bivalve species.

and more silt-clay fractions versus areas adjacent to the aquaculture pens. Disturbance of mudflats during sampling, removal of pens, human disturbance (e.g., noise, viewscape impacts), or other resident organisms had been expressed as a potential concern (see discussions and reviews by Brenchley 1981; Mojica and Nelson 1993; Addessi 1994; Brosnan and Crumrine 1994; Burger 1994; Hall 1994; Rodgers and Smith 1997; DeGrave et al. 1998; Dernie et al. 2003; Yasue 2005; Kimbro and Grosholz 2006; Becker et al. 2009; NRC 2009, 2010). Although a lot of work has been expended on the impacts of predators (e.g., rays, Orth 1975), other disturbances or bioturbation (e.g., Brenchley 1981; Hall 1994; Dahlgren et al. 1999), and human events (i.e., fishing; Glude and Landers 1953; Godcharles 1971; Mojica and Nelson 1993; Addessi 1994; Brosnan and Crumrine 1994; Coen 1995; Brown and Wilson 1997; Eckrich and Holmquist 2000; Kaiser 2000, 2001; Neckles et al. 2005; Forrest et al. 2009; Macleod et al. 2009), little direct research has simulated and assessed the impacts related to intertidal or subtidal shellfish culture (NRC 2009, 2010). As discussed above and elsewhere, growout structures and gear are generally cleaned or removed on an annual or multiyear basis. If left exposed over longer timescales (years) without attention, the cage pens themselves might provide additional hard substrate

habitat for nontarget species (e.g., oysters, mussels; see Figs. 9.12 and 9.13; see also Costa-Pierce and Bridger 2002; Dealteris et al. 2004; Munroe and McKinley 2007a; Powers et al. 2007; D’Amours et al. 2008; Erbland and Ozbay 2008; Dumbauld et al. 2009). The maintenance and ultimate removal of such materials presents an additional concern as does the possible impacts of excess nutrients related to death and decay of the associated fouling organisms removed in situ and left to decompose. In 2004, the South Carolina Department of Natural Resources (SCDNR) (through L. Coen) provided a research plan to address questions related to the remaining 4000, 8-ftdiameter pens abandoned in South Carolina’s waters. Given that insufficient escrow funds were set aside with the two South Carolina state agencies (SCDNR and the South Carolina Department of Health and Environmental Control [SCDHEC]) tasked with pen removal and related materials (Figs. 9.12 and 9.13) in the event of a financial failure, these two agencies were required to determine how to remove all of the remaining abandoned pens and associated materials. The plaintiff suggested that they be left as functioning critical habitat for numerous fish and invertebrate species. The abandonment of aquaculture-related gear is a potentially major problem and one that needs to be addressed more explicitly (see Fig. 9.14).

Figure 9.13 Abandoned cages and other pen parts (mesh, PVC, etc.) left on the leased mudflats in South Carolina, USA.

Figure 9.14 High-resolution aerial imagery of an area near Kiawah, SC (lease M194). Clam pens (see lower-altitude and ground-level images, Figs. 9.6 and 9.12) were removed in 2004. Note the cage impacts still recognizable (delineated by red lines) years after their removal. (Courtesy of K. Schulte, SCDNR and GeoVantage and PhotoScience, Inc.)

271

272

Shellfish Aquaculture and the Environment

Remaining questions Aquaculture will continue to increase worldwide while native “wild stock” populations decline to near unsustainable levels (Beck et al. 2009, in press). Living shoreline projects and native shellfish restoration projects conducted to provide one or more of the ecosystem services discussed above will also continue to expand, especially given sea level rise and the search for more ecofriendly approaches. One question that needs to be further addressed is whether structure alone is sufficient to enhance biodiversity around either natural biogenic shellfish reefs with multiple age classes of molluscs (e.g., Luckenbach et al. 2005), or whether the act of substrate placement alone is sufficient? Recent work with intertidal reefs suggests that significant resident and transient associates (e.g., Harding and Mann 1999, 2001; Coen et al. 2000; Coen et al. 2006) are attracted to shell even without live recruited oysters (Coen et al. 2006, unpublished data). More is known about the impacts of water column-based aquaculture (e.g., mussel culture) than we do about other bivalve species (e.g., oysters). While the number of large-scale shellfish aquaculture operations throughout the world, especially in Europe, the northwestern United States, and the Indo-West Pacific, continues to increase, few studies in those areas have examined the impacts on ecosystem properties at the landscape level. One exception is the spread and impact of Crassostrea gigas replacing soft-sediment dominated communities with intertidal oyster reefs (e.g., see Chapter 14 in this book and discussion above), which will continue to be critically debated and evaluated for the beneficial services and associated negative impacts of aquaculture practices. The underpinnings of an economic basis for restoration need to be resolved: (1) identification and quantification of ecosystem services (direct, indirect, and existence use); (2) determination of a clear definition of associated valuation and supporting services;

(3) development of necessary markets for attendant shellfish services; and (4) related nonmarket challenges (e.g., Freeman 1995; Costanza et al. 1997; Daily 1997; Meyer et al. 1997; Heal 2000; Meyer and Townsend 2000; Farber et al. 2002; Millennium Ecosystem Assessment 2003; Peterson et al. 2003; Cerrato et al. 2004; Lipton 2004; Newell 2004; NRC 2004; Newell et al. 2005; Cerco and Noel 2007; Grabowski and Peterson 2007; Brumbaugh and Toropova 2008; Wei et al. 2009). The continued use and expansion of BMPs (see Chapter 3) in the first, second, and third world, coupled with the application of research findings, will better focus shellfish mariculture on (1) where it can be commercially and ecologically sustainable; (2) site selection; (3) appropriate densities; (4) multiuse operations; and (5) where restoration using scaled-up aquaculture may be applied to areas where harvesting cannot occur as a result of administrative closures, (e.g., initiatives in places such as New York City) but where “farmed” species on an industrial scale may be used to treat eutrophication (e.g., Lindahl et al. 2005; Gren et al. 2009; Chapter 8 in this book). When one compares finfish aquaculture, which requires large inputs of feed often with associated negative impacts (e.g., Grant et al. 1995; Hastings and Heinle 1995; Kaiser et al. 1998; Heffernan 1999; Kaiser 2000, 2001; Crawford et al. 2003; Forrest et al. 2009; NRC 2010), use of bivalve (especially oyster) aquaculture is generally considered to be more sustainable, itself requiring excellent water quality, often with few demonstrable negative impacts at this time (e.g., Pillay 1992; Rice 2000; Read and Fernandes 2003; Newell 2004; Whiteley and Bendell-Young 2007; Dumbauld et al. 2009; NRC 2009, 2010). There is less consensus, however, among researchers on the use of nonnative species regardless of one’s following of the International Council for the Exploration of the Sea (ICES) protocols (e.g., Simenstad and Fresh 1995; Trianni 1995; Rumrill and

Impacts of native and cultured bivalves

Poulton 2004; Ruesink et al. 2005; Kochmann et al. 2008; Molnar et al. 2008; NRC 2009, 2010). Although there is insufficient space to review the issues related to the recent fishery options (including the potential introduction of Crassostrea ariakensis, a nonnative oyster from China and Japan) for the nearly lost native oyster population in the Chesapeake Bay (U.S. states of Maryland and Virginia) and North Carolina, there is much to be learned from the associated 5-year draft EIS process (discussed in depth in NRC 2004; a recent dedicated Journal of Shellfish Research vol. 27[3] 2008; Blankenship 2009). The NRC (2004) report was over 300 pages long, and the final EIS document exceeded 1500 pages, within which numerous potential oyster “management” options were examined, such as 2n or 3n Crassostrea ariakensis, and numerous outcomes and fruitful discussions resulted. Comments from individual state and local governments, federal agencies, the scientific community, industry groups, lawmakers, and countless others drew diverse opinions. In contrast to the numerous and earlier worldwide decisions to introduce (directed) nonnative bivalves into local waters (reviewed in Carlton and Mann 1996; NRC 2004; Ruesink et al. 2005; McKindsey et al. 2007; Molnar et al. 2008), the decision for the Chesapeake Bay was quite different. There was nearly a consensus conclusion to not introduce a nonnative oyster into the bay given the perceived or demonstrated risks versus benefits; whether this will follow elsewhere, only time will tell. Since all shellfish habitats consist of living and dead individuals, and given the lack of sufficient information currently available for any one site or multiple sites, it is necessary to consider the impacts of harvesting, aquaculture, and activities that ultimately impact estuarine and marine habitats. This chapter provides an overview of the state of our knowledge regarding natural bivalve communities, reefs and aggregations, ecosystem services, jus-

273

tification for shellfish protection, enhancement, and restoration efforts using several specific case studies that discuss the direct and indirect, biotic and abiotic, and positive and negative effects of shellfish aquaculture in estuarine and marine contexts. For the various shellfish species, these impacts vary by location of habitat (intertidal vs. subtidal), extent of cultured versus natural acreage (scale), other elements (e.g., existing seagrasses, competitors, benthos, available hard substrate, exotics) associated with the proposed area to be farmed, and by latitudinal/geographical differences (cf. native species, regulations, western Atlantic, Gulf of Mexico, or Eastern Pacific). For some systems there is a wealth of general information that is particularly transferable in part to specific sites (e.g., carrying capacity, lost habitat structure); for others site-specific sampling and related research is required (e.g., interactions with marine mammals, flow, competitors, SAV). All of these factors need to be addressed to assess whether bivalve aquaculture can even partly replace lost natural systems. Finally, general statements as to whether aquaculture has benign, relatively neutral, or even negative effects (NRC 2009, 2010) is still unclear, especially given our limited understanding of the natural, often “relic,” or “functionally extinct” populations historically dominating estuarine and marine ecosystems (Beck et al. 2009, in press). Although polyculture has been practiced for thousands of years in Asia, and the concept of producing seaweed and bivalves as dual products in estuaries is also well entrenched, recent interest in integrated multitrophic aquaculture has revolved around specifically using these organisms to process effluents produced by shrimp and fish culture in both onshore and offshore systems where feed is added (discussed in Folke and Kautsky 1989, 1992; McVey et al. 2002; Tomasso 2002; Neori et al. 2004; Shumway and Kraeuter 2004; Barrington et al. 2009; Pitta et al. 2009). Shellfish have been cocultured with other

274

Shellfish Aquaculture and the Environment

invertebrates or algae in numerous marine systems, though this is usually more the case in suspended culture operations (e.g., Tenore et al. 1974; Paltzat et al. 2008). In other examples, it is simply important to recognize and quantify the role that bivalves play in providing these services for existing near-shore ecosystems (see McVey et al. 2002; Nelson et al. 2004; Lindahl et al. 2005; Gren et al. 2009) and yet recognize variability in system characteristics, bivalve species being cultured, and the possibility of overloading systems with bivalves that can then become serious sources of self-pollution (e.g., Cranford et al. 2007; Yuan et al. 2010). New technology such as remote sensing and geographic information system (GIS) will continue to be used to examining aquaculture effects at the ecosystem level (e.g., Ward et al. 2003; Carswell et al. 2006; Grant et al. 2007). Despite the fact that numerous studies have focused on mussels grown in rope culture, the associated carrying capacities, and related community and ecosystem effects (above cited references), more directed studies need to be conducted that assess the impact of molluscan aquaculture, along with its potential parallel “services” with native (“wild stock”) reef systems (cf. NRC 2009, 2010). Currently, most information is derived from the results of studies in other areas and only indirect evidence is generally available, often for other species, thereby making it difficult to evaluate positive and negative potential (positive or negative) or actual impacts (see also Heffernan 1999; Deal 2005; Dumbauld et al. 2009; Forrest et al. 2009; NRC 2009, 2010). In conclusion, we suggest that aquaculture can provide enhanced structure paralleling that observed for natural communities. The use of native species is desirable because it reduces the chance of other unwanted effects while potentially restoring native bivalve ecosystem functioning, reduces the fishing pressure on native “wild stocks,” and at least partially contributes to native species restora-

tion (e.g., Olympia oyster, Polson et al. 2009; use of Crassostrea virginica triploids in Chesapeake Bay and elsewhere, Beck et al. 2009). A great deal more site-specific research (e.g., NRC 2009, 2010) needs to be directed at examining potential sustainable practices (FAO 2009b) and a greater emphasis needs to be placed on the impacts, both positive and negative, of large-scale bivalve aquaculture at the landscape or ecosystem scales.

Literature cited Addessi, L. 1994. Human disturbance and longterm changes on a rocky intertidal community. Ecological Applications 4:786–797. Agawin, N.S.R., and Duarte, C.M. 2002. Evidence of direct particle trapping by a tropical seagrass meadow. Estuaries 25:1205–1209. Airoldi, L., and Beck, M.W. 2007. Loss, status and trends for coastal marine habitats of Europe. Oceanography Marine Biology: An Annual Review 45:345–405. Alongi, D.M., Chong, V.C., Dixon, P., Sasekumar, A., and Tirendi, F. 2003. The influence of cage aquaculture on pelagic carbon flow and water chemistry in tidally dominated mangrove estuaries of peninsular Malaysia. Marine Environmental Research 55:313–333. Anderson, W.D., Keith, W.J., Tuten, W.R., and Mills, F.H. 1979. A survey of South Carolina’s washed shell resource. South Carolina Wildlife and Marine Resources Department Technical Rpt. No. 36. Andersson, A.J., Mackenzie, F.T., and Ver, L.M. 2003. Solution of shallow-water carbonates. Geology 31:513–516. ASMFC. 2007. The importance of habitat created by shellfish and shell beds along the Atlantic coast of the U.S.. Prepared by L.D. Coen, and R. Grizzle, with contributions by J. Lowery and K.T. Paynter, Jr. Habitat Management Series #8. Asmus, R.M., and Asmus, H. 1991. Mussel beds: limiting or promoting phytoplankton? Journal of Experimental Marine Biology and Ecology 148:215–232. Asmus, H., and Asmus, R.M. 2005. Significance of suspension-feeders systems on different spatial

Impacts of native and cultured bivalves

scales. In: Dame, R.F., and Olenin, S. (eds.), The Comparative Roles of Suspension-Feeders in Ecosystems. Springer-Verlag, Dordrecht, Netherlands, pp. 199–219. Atkinson, P.W., Maclean, I.M.D., and Clark, N.A. 2010. Impacts of shellfisheries and nutrient inputs on waterbird communities in the Wash, England. The Journal of Applied Ecology 47:191–199. Auster, P.J., Malatesta, R.J., and LaRosa, S.C. 1995. Patterns of microhabitat utilization by mobile megafauna on the southern New England (USA) continental shelf and slope. Marine Ecology Progress Series 127:77–85. Backwell, P.R.Y., P.D.O. Hara, and Christy J.H. 1998. Prey availability and selective foraging in shorebirds. Animal Behaviour 55:1659– 1667. Bahr, L.M., Jr. 1976. Energetic aspects of the intertidal oyster reef community at Sapelo Island, Georgia, USA. Ecology 57:121–131. Bahr, L.M., and Lanier, W.P. 1981. The ecology of intertidal oyster reefs of the South Atlantic coast: a community profile. U.S. Fish and Widlife Service, FWS/OBS-81/15, Washington, D.C. Bain, M., Suszkowski, D., Lodge, J., and Xu, L. 2007. Setting Targets for Restoration of the Hudson-Raritan Estuary. Hudson River Foundation, New York. Baird, D., Evans, P.R., Milne, H., and Pienkowski, M.W. 1985. Utilization by shorebirds of benthic invertebrate production in intertidal areas. Oceanography and Marine Biology: An Annual Review 23:573–597. Baker, P. 1995. Review of ecology and fishery of the Olympia oyster, Ostrea lurida with annotated bibliography. Journal of Shellfish Research 14:501–518. Baldwin, J.R., and Lovvorn, J.R. 1994. Expansion of seagrass habitat by the exotic Zostera japonica, and its use by dabbling ducks and brant in Boundary Bay, British Columbia. Marine Ecology Progress Series 103:119–127. Banas, N.S., Hickey, B.M., Newton, J.A., and Ruesink, J.L. 2007. Tidal exchange, bivalve grazing, and patterns of primary production in Willapa Bay, Washington, USA. Marine Ecology Progress Series 341:123–139. Barrington, K., Chopin, T., and Robinson, S. 2009. Integrated multitrophic aquaculture (IMTA) in

275

marine temperate waters. In: Soto, D. (ed.), Integrated Mariculture: A Global Review. FAO Fisheries and Aquaculture Technical Paper No. 529. FAO, Rome, pp. 7–46. Beadman, H.A., Kaiser, M.J., Galanidi, M., Shucksmith, R., and Willows, R.I. 2004. Changes in species richness with stocking density of marine bivalves. The Journal of Applied Ecology 41:464–475. Beal, B.F., and Kraus, M.G. 2002. Interactive effects of initial size, stocking density, and type of predator deterrent netting on survival and growth of cultured juveniles of the soft-shell clam, Mya arenaria L., in eastern Maine. Aquaculture 208:81–111. Beck, M.W., Heck, K.L., Able, K.W., Childers, D.L., Eggleston, D.B., Gillanders, B.M., Halpern, B., Hays, C.G., Hoshino, K., Minello, T.J., Orth, R.J., Sheridan, P.F., and Weinstein, M.R. 2001. The identification, conservation, and management of estuarine and marine nurseries for fish and invertebrates. Bioscience 51:633–641. Beck, M.W., Heck, K.L., Jr., Able, K.W., Childers, D.L., Eggleston, D.B., Gillanders, B.M., Halpern, B.S., Hays, C.G., Hoshino, K., Minello, T.J., Orth, R.J., Sheridan, P.F., and Weinstein, M.P. 2003. The Role of Nearshore Ecosystems as Fish and Shellfish Nurseries. Issues in Ecology, Number 11. Ecological Society of America, Washington, DC. Beck, M.W., Brumbaugh, R.D., Airoldi, L., Carranza, A., Coen, L.D., Crawford, C., Defeo, O., Edgar, G.J., Hancock, B., Kay, M., Lenihan, H., Luckenbach, M.W., Toropova, C.L., and Zhang, G. 2009. Shellfish reefs at risk: a global analysis of problems and solutions. The Nature Conservancy, Arlington, VA, p. 52. Beck, M., Brumbaugh, R., Airoldi, L., Carranza, A., Coen, L., Crawford, C., Defeo, O., Edgar, G., Hancock, B., Kay, M., Lenihan, H., Luckenbach, M., Toropova, C., Shepard, C., and Zhang, G. 2011. Oyster reefs at risk globally and recommendations for ecosystem revitalization. Bioscience 61:107–116. Becker, B.H., Press, D.T., and Allen, S.G. 2009. Modeling the effects of El Niño, density dependence, and disturbance on harbor seal (Phoca vitulina) counts in Drakes Estero, California: 1997–2007. Marine Mammal Science 25:1– 18.

276

Shellfish Aquaculture and the Environment

Bell, S.S., McCoy, E.D., and Mushinsky, H.R. (eds.). 1991. Habitat Structure: The Physical Arrangement of Objects in Space. Chapman and Hall, London. Black, K.D. 2001. Environmental Impacts of Aquaculture. Sheffield Academic Press, Sheffield. Blankenship, K. 2009. All sides weigh in on C. ariakensis introduction to Bay. February 2009 issue. Boesch, D.F., and Turner, R.E. 1984. Dependence of fishery species on salt marshes: the role of food and refuge. Estuaries 7:460–468. Booth, D.M., and Heck, K.L., Jr. 2009. Impacts of the American oyster (Crassostrea virginica) on growth rates of the seagrass Halodule wrightii. Marine Ecology Progress Series 389:117–126. Borde, A.B., Thom, R.M., Rumrill, S., and Miller, L.M. 2003. Geospatial habitat change analysis in Pacific Northwest coastal estuaries. Estuaries 26:1104–1116. Borges, A.V., and Gypens, N. 2010. Carbonate chemistry in the coastal zone responds more strongly to eutrophication than to ocean acidification. Limnology and Oceanography 55:346–353. Borthagaray, A., and Carranza, A. 2007. Mussels as ecosystem engineers: their contribution to species richness in a rocky littoral community. Acta Oecologica 31:243–250. Breitburg, D.L. 1999. Are 3-dimensional structure and healthy oyster populations the keys to an ecologically interesting and important fish community? Luckenbach, M.W., Mann, R., and Wesson, J.E. (eds.), Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. VIMS Press, Gloucester Point, VA, pp. 239–249. Breitburg, D.L., Palmer, M.A., and Loher, T. 1995. Larval distributions and the spatial patterns of settlement of an oyster reef fish: responses to flow and structure. Marine Ecology Progress Series 125:45–60. Breitburg, D., Coen, L.D., Luckenbach, M.W., Mann, R., Posey, M., and Wesson, J.A. 2000. Oyster reef restoration: convergence of harvest and conservation strategies. Journal of Shellfish Research 19:371–377. Brenchley, G.A. 1981. Disturbance and community structure: an experimental study of bioturbation in marine soft-bottom environments. Journal of Marine Research 39:767–790.

Brewer, P.G. 1997. Ocean chemistry of the fossil fuel CO2 signal: the haline signature of “business as usual.” Geophysical Research Letters 24:1367–1369. Brosnan, D.M., and Crumrine, L.L. 1994. Effects of human trampling on marine rocky shore communities. Journal Experimental Marine Biology and Ecology 177:79–97. Brown, B., and Wilson, W.H., Jr. 1997. The role of commercial digging of mudflats as an agent for change of infaunal intertidal populations. Journal Experimental Marine Biology and Ecology 218:49–61. Brumbaugh, R.D., and Coen, L.D. 2009. Contemporary approaches for small-scale oyster reef restoration to address substrate versus recruitment limitation: a review and comments relevant for the Olympia oyster, Ostrea lurida (Carpenter, 1864). Journal of Shellfish Research 28:147–161. Brumbaugh, R.D., and Toropova, C. 2008. Economic valuation of ecosystem services: a new impetus for shellfish restoration. Basins and Coasts 2:8–15. Brumbaugh, R.D., Beck, M.W., Coen, L.D., Craig, L., and Hicks, P. 2006. A Practitioners’ Guide to the Design and Monitoring of Shellfish Restoration Projects: An Ecosystem Services Approach. The Nature Conservancy, Arlington, VA. Burger, J. 1994. The effect of human disturbance on foraging behavior and habitat use in Piping Plover (Charadrius melodus). Estuaries 17:695–701. Burrell, V.G., Jr. 1986. Species profiles: life histories and environmental requirements of coastal fishes and invertebrates (South Atlantic)—American oyster. U.S. Fish Wildl. Serv. Biological Report 82 (11.57), U.S. Army Corps of Engineers TR EL-82-4, 17pp. Burrows, F., Harding, J.M., Mann, R., Dame, R., and Coen, L. 2005. Chapter 4, Restoration monitoring of oyster reefs. In: Thayer, G.W., McTigue, T.A., Salz, R.J., Merkey, D.H., Burrows, F.M., and Gayaldo, P.F. (eds.), ScienceBased Restoration Monitoring of Coastal Habitats, Volume Two: Tools for Monitoring Coastal Habitats. NOAA Coastal Ocean Program Decision Analysis Series No. 23. NOAA National Centers for Coastal Ocean Science, Silver Spring, MD, pp. 4.2–4.73.

Impacts of native and cultured bivalves

Buschbaum, C., Dittmann, S., Hong, J.S., Hwang, I.S., Strasser, M., Thiel, M., Valdivia, N., Yoon, S.P., and Reise, K. 2009. Mytilid mussels: global habitat engineers in coastal sediments. Helgoland Marine Research 63:47–58. Bushek, D., Richardson, D., Bobo, M.Y., and Coen, L.D. 2004. Short-term shell pile quarantine reduces the abundance of Perkinsus marinus remaining in tissues attached to oyster shell. Journal of Shellfish Research 23:369–373. Cahoon, L.B., Nearhoof, J.E., and Tilton, C.L. 1999. Sediment grain size effect on benthic microalgal biomass in shallow aquatic ecosystems. Estuaries 22:735–741. Caldeira, K., and Wickett, M.E. 2003. Oceanography: anthropogenic carbon and ocean pH. Nature 425:365–365. Caldow, R.W.G., Beadman, H.A., McGrorty, S., Stillman, R.A., Goss-Custard, J.D., le V. dit Durell, S.E.A., West, A.D., Kaiser, M.J., Mould, K., and Wilson, A. 2004. A behavior-based modeling approach to reducing shorebird-shellfish conflicts. Ecological Applications 14:1411– 1427. Callier, M.D., Weise, A.M., McKindsey, C.W., and Desrosiers, G. 2006. Sedimentation rates in a suspended mussel farm (Great-Entry Lagoon, Canada): biodeposit production and dispersion. Marine Ecology Progress Series 322:129–141. Callier, M.D., McKindsey, C.W., and Desrosiers, G. 2008. Evaluation of indicators used to detect mussel farm influence on the benthos: two case studies in the Magdalen Islands, Eastern Canada. Aquaculture 278:77–88. Carlton, J.T., and Mann, R.H. 1996. Transfers and world wide introductions. In: Kennedy, V.S., Newell, R.I.E., and Eble, A.F. (eds.), The Eastern Oyster Crassostrea Virginica. Maryland Sea Grant, College Park, MD, pp. 691–705. Carriker, M.R. 1992. Introductions and transfers of molluscs: risk consideration and implications. Journal of Shellfish Research 11:507–510. Carroll, J., Gobler, C.J., and Peterson, B.J. 2008. Resource limitation of eelgrass in New York estuaries: light limitation and nutrient stress alleviation by hard clams. Marine Ecology Progress Series 369:39–50. Carswell, B., Cheesman, S., and Anderson, J. 2006. The use of spatial analysis for environmental assessment of shellfish aquaculture in Baynes

277

Sound, Vancouver Island, British Columbia, Canada. Aquaculture 253:408–414. Castel, J., Labourg, P.J., Escaravage, V., Auby, I., and Garcia, M.E. 1989. Influence of seagrass beds and oyster parks on the abundance and biomass patterns of meio- and macrobenthos intertidal flats. Estuarine, Coastal and Shelf Science 28:71–85. Cerco, C.F., and Noel, M.R. 2007. Can oyster restoration reverse cultural eutrophication in Chesapeake Bay? Estuaries and Coasts 30: 331–343. Cerrato, R.M., Caron, D.A., Lonsdale, D.J., Rose, J.M., and Schaffner, R.A. 2004. Effect of the northern quahog Mercenaria mercenaria on the development of blooms of the brown tide alga Aureococcus anophagefferens. Marine Ecology Progress Series 281:93–108. Chew, K.K. 1990. Global bivalve shellfish introductions. World Aquaculture 21:9–22. Cloern, J.E. 1982. Does the benthos control phytoplankton biomass in south San Francisco Bay? Marine Ecology Progress Series 9:191–202. Coen, L.D. 1995. A review of the potential impacts of mechanical harvesting on subtidal and intertidal shellfish resources. SCDNR-MRRI, 46 pp. + three Appendices. Coen, L.D., and Luckenbach, M.W. 2000. Developing success criteria and goals for evaluating oyster reef restoration: ecological function or resource exploitation? Ecological Engineering 15:323–343. Coen, L.D., Knott, D.M., Wenner, E.L., Hadley, N.H., and Ringwood, A.H. 1999a. Intertidal oyster reef studies in South Carolina: design, sampling and experimental focus for evaluating habitat value and function. In: Luckenbach, M.W., Mann, R., and Wesson, J.A. (eds.), Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. Virginia Institute of Marine Science Press, Gloucester Point, VA, pp. 131–156. Coen, L.D., Luckenbach, M.W., and Breitburg, D.L. 1999b. The role of oyster reefs as essential fish habitat: a review of current knowledge and some new perspectives. In: Benaka, L.R. (ed.), Fish Habitat: Essential Fish Habitat and Rehabilitation. American Fisheries Society, Symposium 22, Bethesda, MD, pp. 438– 454.

278

Shellfish Aquaculture and the Environment

Coen, L.D., Judge, M., Moncreiff, C., and Hammerstrom, K. 2000. Final S-K Project Report, Hard clam (Mercenaria mercenaria) mariculture in U.S. waters: evaluating the effects of large-scale field grow-out practices on clam growth, nutrition and inshore estuarine creek communities. 119. Coen, L.D., Walters, K., Wilber, D., and Hadley, N. 2004. A South Carolina Sea Grant Report of A 2004 workshop to examine and evaluate oyster restoration metrics to assess ecological function, sustainability and success: results and related information. Report to the South Carolina Sea Grant Consortium, 27pp. www.oyster-restoration.org/scsg04/SCSG04.pdf Coen, L.D., Bolton-Warberg, M., and Stephen, J.A. 2006. An examination of oyster reefs as “biologically-critical” estuarine ecosystems. Final Report, Grant R/ER-10, Submitted to the South Carolina Sea Grant Consortium, 214 pages, plus appendices. Coen, L.D., Brumbaugh, R.D., Bushek, D., Grizzle, R., Luckenbach, M.W., Posey, M.H., Powers, S.P., and Tolley, G. 2007. As we see it. A broader view of ecosystem services related to oyster restoration. Marine Ecology Progress Series 341:303–307. Coen, L.D., Hadley, N., Shervette, V., and Anderson, W. 2010. Managing oysters in South Carolina: a five year program to enhance/restore shellfish stocks and reef habitats on through shell planting and technology improvements. SC Saltwater Recreational Fisheries License Program Final Report, 124pp. Cohen, A.N., and Zabin, C.J. 2009. Oyster shells as vectors for exotic organisms. Journal of Shellfish Research 28:163–167. Cohen, R.R.H., Dresler, P.V., Philips, E.J.P., and Cory, R.L. 1984. The effect of the Asiatic clam, Corbicula fluminea on phytoplankton of the Potomac River, Maryland. Limnology and Oceanography 29:170–180. Collins, J.W. 1892. Report on the fisheries of the Pacific coast of the United States. Report of the Commisioner for 1888, pp. 3–209. Washington, DC: United States Commission of Fish and Fisheries. Colwell, M.A. 1993. Shorebird community patterns in a seasonally dynamic estuary. The Condor 95:104–114.

Colwell, M.A., and Landrum, S.L. 1993. Nonrandom shorebird distribution and finescale variation in prey abundance. The Condor 95:94–103. Comeau, L.A., St.-Onge, P., Pernet, F., and Lanteigne, L. 2009. Deterring coastal birds from roosting on oyster culture gear in eastern New Brunswick, Canada. Aquacultural Engineering 40:87–94. Connolly, L.M., and Colwell, M.A. 2005. Comparative use of longline oysterbeds and adjacent tidal flats by waterbirds. Bird Conservation International 15:237–255. Conte, F.S., Harbell, S.C., and RaLonde, R.L. 1994. Oyster Culture: Fundamentals and Technology of the West Coast Industry. Western Regional Aquaculture Center, Seattle, WA. Corbett, C.A. 2007. Colored dissolved organic matter (CDOM), workshop summary. Technical Document 07-3, CHNEP, FL, 82pp. Costanza, R., d’Arge, R., de Groot, R., Farber, S., Grasso, M., Hannon, B., Naeem, S., Limburg, K., Paruelo, J., O’Neill, R.V., Raskin, R., Sutton, P., and van den Belt, M. 1997. The value of the world’s ecosystem services and natural capital. Nature 387:253–260. Costa-Pierce, B.A., and Bridger, C.J. 2002. The role of marine aquaculture facilities as habitats and ecosystems. In: Stickney, R.R., and McVey, J.P. (eds.), Ch. 8, Responsible Marine Aquaculture. CABI Publishing, Wallingford, UK, pp. 105–144. Cranford, P.J., Strain, P.M., Dowd, M., Hargrave, B.T., Grant, J., and Archambault, M.C. 2007. Influence of mussel aquaculture on nitrogen dynamics in a nutrient enriched coastal embayment. Marine Ecology Progress Series 347: 61–78. Crawford, C.M., Macleod, C.K.A., and Mitchell, I.M. 2003. Effects of shellfish farming on the benthic environment. Aquaculture 224:117– 140. Cressman, K.A., Posey, M.H., Mallin, M.A., Leonard, L.A., and Alphin, T.D. 2003. Effects of oyster reefs on water quality in a tidal creek estuary. Journal of Shellfish Research 22: 753–762. Cummings, V.J., Thrush, S.F., Hewitt, J.E., and Turner, S.J. 1998. The influence of the pinnid

Impacts of native and cultured bivalves

bivalve Atrina zelandica (Gray) on benthic macroinvertebrate communities on soft-sediment habitats. Journal of Experimental Marine Biology and Ecology 228:227–240. D’Amours, O., Archambault, P., Christopher, W., McKindsey, C.W., and Johnson, L.E. 2008. Local enhancement of epibenthic macrofauna by aquaculture activities. Marine Ecology Progress Series 371:73–84. Dahlgren, C.P., Posey, M.H., and Hulbert, A.W. 1999. The effects of bioturbation on the infaunal community adjacent to an offhore hardbottom reef. Bulletin of Marine Science 64:21–34. Daily, G.E. 1997. Nature’s Services—Societal Dependence on Natural Ecosystems. Island Press, Washington, DC. Dame, R.F. 1979. The abundance, diversity, and biomass of macrobenthos on North Inlet, South Carolina, intertidal oyster reefs. Proceedings of the National Shellfish Association 69:6– 10. Dame, R.F. 1993. Bivalve Filter Feeders and Coastal and Estuarine Ecosystem Processes. SpringerVerlag, Heidelberg, Germany. Dame, R. 1996. Ecology of Marine Bivalves: An Ecosystem Approach. CRC Marine Science Series, Boca Raton, FL. Dame, R.F., and Libes, S. 1993. Oyster reefs and nutrient retention in tidal creeks. Journal of Experimental Marine Biology and Ecology 171:251–258. Dame, R.F., Haskin, E., and Kjerfve, B. 1984a. Water flow over an intertidal oyster reef and its relationship to nutrient dynamics. Journal of Shellfish Research 4:86–87. Dame, R.F., Zingmark, R.G., and Haskin, E. 1984b. Oyster reefs as processors of estuarine materials. Journal of Experimental Marine Biology and Ecology 83:239–247. Dame, R., Bushek, D., and Prins, T. 2001. The role of suspension feeders as ecosystem transformers in shallow coastal environments. In: Reise, K. (ed.), The Ecology of Sedimentary Coasts. Springer-Verlag, Berlin, pp. 11–37. Danielsen, F., Sørensen, M.K., Olwig, M.F., Vaithilingam, S., Parish, F., Burgess, N.D., Hiraishi, T., Karunagaran, V.M., Rasmussen, M.S., Hansen, L.B., Quarto, A., and Suryadiputra, N. 2005. The Asian tsunami: a protective role for coastal vegetation. Science 310:643.

279

Deal, H. 2005. Sustainable shellfish: recommendations for responsible aquaculture. Report for the David Suzuki Foundation, BC, Canada, 41pp. Dealteris, J.T., Kilpatrick, B.D., and Rheault, R.B. 2004. A comparative evaluation of the habitat value of shellfish aquaculture gear, submerged aquatic vegetation and a non-vegetated seabed. Journal of Shellfish Research 23:867–874. DeCasabianca, M.-L., Laugier, T., and Collart, D. 1997. Impact of shellfish farming eutrophication on benthic macrophyte communities in the Thau lagoon, France. Aquaculture International 5:301–314. DeGrave, S., Moore, S.J., and Burnell, G. 1998. Changes in benthic macrofauna associated with intertidal oyster, Crassostrea gigas (Thunberg) culture. Journal of Shellfish Research 17:1137– 1142. Denny, M.W. 1988. Biology and the Mechanics of the Wave-Swept Environment. Princeton University Press, Princeton, NJ. Denny, M., and Wethey, D. 2001. Physical processes that generate patterns in marine communities. In: Bertness, M.D., Gaines, S.M., and Hixon, M.E. (eds.), Marine Community Ecology. Sinauer Associates, Sunderland, MA, USA, pp. 3–37. Dernie, K.M., Kaiser, M.J., Richardson, E.A., and Warwick, R.M. 2003. Recovery of soft sediment communities and habitats following physical disturbance. Journal of Experimental Marine Biology and Ecology 285:415–434. Check citation. Dodd, S.L., and Colwell, M.A. 1996. Seasonal variation in diurnal and nocturnal distributions of nonbreeding shorebirds at North Humboldt Bay, California. The Condor 98:196–207. Doering, P.H., Oviatt, C.A., and Kelly, J.R. 1986. The effects of the filter-feeding clam Mercenaria mercenaria on carbon cycling in experimental marine mesocosms. Journal of Marine Research 44:839–861. Doney, S.C., Fabry, V.J., Feely, R.A., and Kleypas, J.A. 2009. Ocean acidification: the other CO2 problem. Annual Review of Marine Science 1:169–192. Donovan, S.K. 1991. The Processes of Fossilization. Columbia University Press, New York. Dowd, M. 2005. A bio-physical coastal ecosystem model for assessing environment effects of

280

Shellfish Aquaculture and the Environment

marine bivalve aquaculture. Ecological Modelling 183:323–346. Drapeau, A., Comeau, L.A., Landry, T., Stryhn, H., and Davidson, J. 2006. Association between longline design and mussel productivity in Prince Edward Island, Canada. Aquaculture 261:879– 889. Drinkwaard, A.C. 1999. Introductions and developments of oysters in the North Sea area: a review. Helgoländer Meeresunters 52:301–308. Duarte, P., Hawkins, A.J.S., and Pereira, A. 2005. How does estimation of environmental carrying capacity for bivalve culture depend upon spatial and temporal scales? In Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Marine Ecosystem Processes, NATO ASI Series, Vol. 47, Springer, Dordrecht, The Netherlands. Duarte, P., Labarta, U., and Fernández-Reiriz, M.J. 2008. Modelling local food depletion effects in mussel rafts of Galician Rias. Aquaculture 274:300–312. Duffy, J.E., Richardson, J.P., and Canuel, E.A. 2003. Grazer diversity effects on ecosystem functioning in seagrass beds. Ecology Letters 6:637–645. Dumbauld, B.R., and Wyllie-Echeverria, S. 2003. The influence of burrowing thalassinid shrimps on the distribution of intertidal seagrasses in Willapa Bay, Washington, USA. Aquatic Botany 77:27–42. Dumbauld, B.R., Armstrong, D.A., and McDonald, T.L. 1993. Use of oyster shell to enhance intertidal habitat and mitigate loss of dungenous crab (Cancer magister) caused by dredging. Canadian Journal of Fisheries and Aquatic Sciences 50:381–390. Dumbauld, B., Visser, E., Armstrong, D.A., ColeWarner, L., Feldman, K., and Kauffman, B. 2000. Use of oyster shell to create habitat for juvenile Dungeness crab in Washington coastal estuaries: status and prospects. Journal of Shellfish Research 19:379–386. Dumbauld, B.R., Brooks, K.M., and Posey, M.H. 2001. Response of an estuarine benthic community to application of the pesticide carbaryl and cultivation of Pacific oysters (Crassostrea gigas) in Willapa Bay, Washington. Marine Pollution Bulletin 42:826–844. Dumbauld, B.R., Booth, S., Cheney, D., Suhrbier, A., and Beltran, H. 2006. An integrated pest

management program for burrowing shrimp control in oyster aquaculture. Aquaculture 261:976–992. Dumbauld, B.R., Ruesink, J.L., and Rumrill, S.S. 2009. The ecological role of bivalve shellfish aquaculture in the estuarine environment: a review with application to oyster and clam culture in West Coast (USA) estuaries. Aquaculture 290:196–223. Eckrich, C.E., and Holmquist, J.G. 2000. Trampling in a seagrass assemblage: direct effects, response of associated fauna, and the role of substrate characteristics. Marine Ecology Progress Series 201:199–209. Tomasso, J.R. (ed.) 2002. Aquaculture and the environment in the United States. U.S. Aquaculture Society, A chapter of the World Aquaculture Society, Baton Rouge, LA. U.S.A., 277pp. Eggleston, D.B., Elis, W.E., Etherington, L.L., Dahlgren, C.P., and Posey, M.H. 1999. Organism responses to habitat fragmentation and diversity: habitat colonization by estuarine macrofauna. Journal of Experimental Marine Biology and Ecology 236:107–132. Erbland, P.J., and Ozbay, G. 2008. Comparison of the macrofaunal communities inhabiting a Crassostrea virginica oyster reef and oyster aquaculture gear in Indian River Bay, Delaware. Journal of Shellfish Research 27:757–768. Everett, R.A., Ruiz, G.M., and Carlton, J.T. 1995. Effect of oyster mariculture on submerged aquatic vegetation: an experimental test in a Pacific Northwest estuary. Marine Ecology Progress Series 125:205–217. FAO. 2009a. The State of World Fisheries and Aquaculture 2008. Food and Agriculture Organization of the United Nations, Rome, Italy. FAO. 2009b. Environmental impact assessment and monitoring in aquaculture. FAO Fisheries and Aquaculture Technical Paper No. 527. Rome, FAO. 2009. 57pp (CDROM full document is 648 pp). Farber, S.C., Costanza, R., and Wilson, M.A. 2002. Economic and ecological concepts for valuing ecosystem services. Ecological Economics 41:375–392. Faunce, C.H., and Serafy, J.E. 2006. Mangroves as fish habitat: fifty years of field studies. Marine Ecology Progress Series 318:1–18.

Impacts of native and cultured bivalves

Feely, R.A., Sabine, C.L., Lee, K., Berelson, W., Kleypas, J., Fabry, V.J., and Millero, F.J. 2004. Impact of anthropogenic CO2 on the CaCO3 system in the oceans. Science 305: 362–366. Feigin, R.A., Lozada-Bernhard, S.M., Ravens, T.M., Möller, I., Yeager, K.M., and Baird, A.H. 2009. Does vegetation prevent wave erosion of salt marsh edges? PNAS 106:10109–10113. Feldman, K.L., Armstrong, D.A., Dumbauld, B.R., DeWitt, T.H., and Doty, D.C. 2000. Oysters, crabs, and burrowing shrimp: review of an environmental conflict over aquatic resources and pesticide use in Washington State’s (USA) coastal estuaries. Estuaries 23:141–176. Ferraro, S.P., and Cole, F.A. 2007. Benthic macrofauna-habitat associations in Willapa Bay, Washington, USA. Estuarine, Coastal and Shelf Science 71:491–507. Ferreira, J.G., Hawkins, A.J.S., and Bricker, S.B. 2007. Management of productivity, environmental effects and profitability of shellfish aquaculture—the Farm Aquaculture Resource Management (FARM) model. Aquaculture 264:160–174. Ferreira, J.G., Sequeira, A., Hawkins, A.J.S., Newton, A., Nickell, T.D., Pastres, R., Forte, J., Bodoy, A., and Bricker, S.B. 2009. Analysis of coastal and offshore aquaculture: application of the FARM model to multiple systems and shellfish species. Aquaculture 289:32–41. Folke, C., and Kautsky, N. 1989. The role of ecosystem for a sustainable development of aquaculture. Ambio 18:234–243. Folke, C., and Kautsky, N. 1992. Aquaculture with environment: prospects for sustainability. Ocean Coastal Management 17:5–24. Forrest, B., Keeley, N., Gillespie, P., Hopkins, G., Knight, B., and Govier, D. 2007. Review of the ecological effects of marine finfish aquaculture. Cawthron Report 1285:1–73. Forrest, B.M., Keeley, N.B., Hopkins, G.A., Webb, S.C., and Clement, D.M. 2009. Bivalve aquaculture in estuaries: review and synthesis of oyster cultivation effects. Aquaculture 298:1– 15. Freeman, M. 1995. The benefits of water quality improvements for the marine recreation: a review of the empirical evidence. Marine Resource Economics 10:385–406.

281

Fulford, R.S., Breitburg, D.L., Newell, R.I.E., Kemp, W.M., and Luckenbach, M.W. 2007. Effects of oyster population restoration strategies on phytoplankton biomass in Chesapeake Bay: a flexible modeling approach. Marine Ecology Progress Series 336:43–61. Fulford, R.S., Breitburg, D.L., Luckenbach, M.W., and Newell, R.I.E. 2010. Evaluating responses of estuarine food webs to oyster restoration and nutrient load reduction with a multi-species bioenergetics model. Ecological Applications 20:915–934. Galtsoff, P.S. 1964. The American oyster Crassostrea virginica Gmelin. U.S Fish Widl Serv Fishery Bulletin 64:1–480. Gazeau, F., Quiblier, C., Jansen, J.M., Gattuso, J.-P., Middelburg, J.J., and Heip, C.H.R. 2007. Impact of elevated CO2 on shellfish calcification. Geophysical Research Letters 34:L07603. doi:10.1029/2006GL028554 Gilbert, F., Souchu, P., Bianchi, M., and Bonin, P. 1997. Influence of shellfish farming activities on nitrification, nitrate reduction to ammonium and denitrification at the water sediment interface of the Thau lagoon, France. Marine Ecology Progress Series 151:143–153. Gillespie, G. 2009. Status of the Olympia oyster, Ostrea lurida, (Carpenter 1864) in British Columbia, Canada. Journal of Shellfish Research 28:59–68. Giotta, R.E. 1999. Distribution of the American oyster (Crassostrea virginica) in South Carolina: interacting effects of predation, sedimentation and water flow at varying tidal elevations. M.S. Thesis, College of Charleston, South Carolina, 98pp. Glancy, T.P., Frazer, K.T.K., Cichra, C.E., and Lindberg, W.J. 2003. Comparative patterns of occupancy by decapod crustaceans in seagrass, oyster, and marsh-edge habitats in a northeast Gulf of Mexico estuary. Estuaries 26:1291– 1301. Glude, J.B., and Landers, W.S. 1953. Biological effects on hard clams of hand raking and power dredging. U.S. Fish Wildl. Serv. Spec. Rep. Fish. 110, 43pp. Godcharles, M.F. 1971. A study of the effects of a commercial hydraulic clam dredge on benthic communities in estuarine areas. State of Fla. Dept. of Nat. Res. Tech. Ser. No. 64:1–51.

282

Shellfish Aquaculture and the Environment

Godet, L., Toupoint, N., Fournier, J., Le Mao, P., Retiere, C., and Olivier, F. 2009. Clam farmers and oystercatchers: effects of the degradation of Lanice conchilega beds by shellfish farming on the spatial distribution of shorebirds. Marine Pollution Bulletin 58:589–595. Goodwin, L. 2007. Evaluating the impacts of environmental parameters on shoreline erosion and related aspects: assessing the current status of vegetation, sediments, and biota. M.S. thesis, College of Charleston, SC 117pp. Goss-Custard, J.D., Stillman, R.A., West, A.D., Caldow, R.W.G., Triplet, P., le V. dit Durell, S.E.A., and McGrorty, S. 2004. When enough is not enough: shorebirds and shellfishing. Proceedings of the Royal Society of London. Series B. Biological Sciences 271:233–237. Grabowski, J.H., and Peterson, C.H. 2007. Restoring oyster reefs to recover ecosystem services. In: Ch. 15, Cuddington, K., Byers, J.E., Wilson, W.G., and Hastings, A. (eds.), Ecosystem Engineers: Concepts, Theory and Applications. Elsevier/Academic Press, Burlington, MA, pp. 281–298. Grant, J., Hatcher, A., Scott, D.B., Pocklington, P., Schafer, C.T., and Winters, G.V. 1995. A multidisciplinary approach to evaluating impacts of shellfish aquaculture on benthic communities. Estuaries 18:124–144. Grant, J., Cranford, P., Hargrave, B., Carreau, M., Schofield, B., Armsworthy, S., Burdett-Coutts, V., and Ibarra, D. 2005. A model of aquaculture biodeposition for multiple estuaries and field validation at blue mussel (Mytilus edulis) culture sites in eastern Canada. Canadian Journal of Fisheries and Aquatic Sciences 62:1271– 1285. Grant, J., Bugden, G., Horne, E., Archambault, M.C., and Carreau, M. 2007. Remote sensing of particle depletion by coastal suspension-feeders. Canadian Journal of Fisheries and Aquatic Sciences 64:387–390. Gren, I.-M., Lindahl, O., and Lindqvist, M. 2009. Values of mussel farming for combating eutrophication: an application to the Baltic Sea. Ecological Engineering 35:935–945. Grizzle, R.E. 1988. The relative effects of seston flux and sediments on individual growth rates of Mercenaria mercenaria: results of a factorial field experiment. Journal of Shellfish Research 7:160–161.

Grizzle, R.E., Greene, J.K., Luckenbach, M.W., and Coen, L.D. 2006. Measuring and modeling seston uptake by suspension feeding bivalve molluscs. Journal of Shellfish Research 25:643–649. Grizzle, R.E., Greene, J.K., and Coen, L.D. 2008. Seston removal by natural and constructed intertidal eastern oyster (Crassostrea virginica) reefs: a comparison with previous laboratory studies, and the value of in situ methods. Estuaries and Coasts 31:1208–1220. Gutiérrez, J.L., Jones, C.G., Strayer, D.L., and Iribarne, O.O. 2003. Molluscs as ecosystem engineers: the role of shell production in aquatic habitats. Oikos 101:79–90. Hadley, N.H., Hodges, M., Wilber, D.H., and Coen, L.D. 2010. Evaluating intertidal oyster reef development in South Carolina using associated faunal indicators. Restoration Ecology 18:691–701. Hall, S.J. 1994. Physical disturbance and marine benthic communities: life in unconsolidated sediments. Oceanography and Marine Biology Annual Review 32:179–239. Hammel, A., and Webb, D. 1999. Ecological implications of Mercenaria aquaculture in the Cape Cod National Seashore. Abstract and poster presented at the Annual Benthic Ecology Meeting, Baton Rouge, LA. Harbin-Ireland, A.C. 2004. Effects of oyster mariculture on the benthic invertebrate community in Drakes Estero, Pt. Reyes Peninsula, California. M.S. thesis, University of California, Davis, California. Harding, J.M., and Mann, R. 1999. Fish species richness in relation to restored oyster reefs, Piankatank River, Virginia. Bulletin of Marine Science 65:289–300. Harding, J.M., and Mann, R. 2001. Oyster reefs as fish habitat: opportunistic use of restored reefs by transient fishes. Journal of of Shellfish Research 20:951–959. Hargis, W.J., Jr., and Haven, D.S. 1999. Chesapeake oyster reefs, their importance, destruction and guidelines for restoring them. In: Luckenbach, M.W., Mann, R., and Wesson, J.A. (eds.), Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. Virginia Institute of Marine Science Press, Gloucester Point, VA, pp. 329–358. Harrison, P.G., and Bigley, R.E. 1982. The recent introduction of the seagrass Zostera japonica to

Impacts of native and cultured bivalves

the Pacific coast of North America. Canadian Journal of Fisheries and Aquatic Sciences 39:1642–1648. Harsh, D.A., and Luckenbach, M.W. 1999. Materials processing by oysters in patches: interactive roles of current speed and seston composition. In: Luckenbach, M.W., Mann, R., and Wesson, J.A. (eds.), Oyster Reef Habitat Restoration. A Synopsis and Synthesis of Approaches. Virginia Institute of Marine Science Press, Gloucester Point, VA, pp. 251– 265. Hastings, R.W., and Heinle, D.R. 1995. The effects of aquaculture in estuarine environments: introduction to the dedicated issue. Estuaries 18:1–284. Hauxwell, J., Cebrian, J., Furlong, C., and Valiela, I. 2001. Macroalgal canopies contribute to eelgrass (Zostera marina) decline in temperate estuarine ecosystems. Ecology 82:1007–1022. Haven, D.S., and Morales-Alamo, R. 1966. Aspects of biodeposition by oysters and other invertebrate filter feeders. Limnology and Oceanography 11:487–498. Heal, G. 2000. Nature and the Marketplace: Capturing the Value of Ecosystem Services. Island Press, Washington, DC. Hecht, T., and Britz, P. 1992. The current status, future prospects and environmental implications of mariculture in South Africa. South African Journal of Science 88:335–342. Heck, K.L., Jr., and Crowder, L.B. 1991. Habitat structure and predator-prey interactions in vegetated aquatic systems. In: Bell, S.S., McCoy, E.D., and Mushinsky, H.R. (eds.), Habitat Structure: the Physical Arrangement of Objects in Space. Chapman and Hall, London, pp. 281–295. Heck, K.L., Jr., Pennock, J.R., Valentine, J.F., Coen, L.D., and Sklenar, S.A. 2000a. Effects of nutrient enrichment and large predator removal on seagrass nursery habitats: an experimental assessment. Limnology and Oceanography 45: 1041–1057. Heck, K.L., Jr., Pennock, J.R., Valentine, J.F., Coen, L.D., and Sklenar, S.A. 2000b. Effects of nutrient enrichment and overfishing on seagrass ecosystems: an experimental assessment. Biologia Marina Mediterranea 7:220–222. Heck, K.L., Jr., Hays, C., and Orth, R.J. 2003. A critical evaluation of the nursery role hypothesis

283

for seagrass meadows. Marine Ecology Progress Series 253:123–136. Heck, K.L., Jr., Valentine, J.F., Pennock, J.R., Chaplin, G., and Spitzer, P.M. 2006. Effects of nutrient enrichment and grazing on shoal grass (Halodule wrightii) and its epiphytes: results of a field experiment. Marine Ecology Progress Series 326:145–156. Heck, K.L., Carruthers, T.J.B., Duarte, C.M., Hughes, A.R., Kendrick, G., Orth, R.J., and Williams, S.W. 2008. Trophic transfers from seagrass meadows subsidize diverse marine and terrestrial consumers. Ecosystems 11:1198– 1210. Heffernan, M.L. 1999. A review of the ecological implications of mariculture and intertidal harvesting in Ireland. Irish Wildlife Manuals, No. 7. Dúchas, The Heritage Service, Department of Arts, Heritage, Gaeltacht and the Islands, Dublin, Ireland, 156pp. www.npws.ie/en/media/ Media,3762,en.pdf Hégaret, H., Shumway, S.E., Wikfors, G.H., Pate, S., and Burkholder, J.M. 2008. Potential transport of harmful algae through relocation of bivalve molluscs. Marine Ecology Progress Series 361:169–179. Heil, D.C. 2009. Karenia brevis monitoring, management, and mitigation for Florida molluscan shellfish harvesting areas. Harmful Algae 8:608–610. Heise, R.J., and Bortone, S.A. 1999. Estuarine artificial reefs to enhance seagrass planting and provide fish habitat. Gulf of Mexico Science 17:59–74. Hemminga, M.A., and Duarte, C.M. 2000. Seagrass Ecology. Cambridge University Press, Cambridge. Heral, M. 1993. Why carrying capacity models are useful tools for management of bivalve molluscs culture. In: Dame, R.F. (ed.), Bivalve Filter Feeders in Estuarine and Coastal Ecosystem Processes. Springer-Verlag, Berlin, Heidelberg, pp. 455–477. Hewitt, J.E., and Norkko, J. 2007. Incorporating temporal variability of stressors into studies: an example using suspension-feeding bivalves and elevated suspended sediment concentrations. Journal of Experimental Marine Biology and Ecology 341:131–141. Hilgerloh, G., O’Halloran, J., Kelly, T.C., and Burnell, G.M. 2001. A preliminary study on the

284

Shellfish Aquaculture and the Environment

effects of oyster culturing structures on birds in a sheltered Irish estuary. Hydrobiologia 465:175–180. Holmer, M. 1991. Impacts of aquaculture on surrounding sediments: generation of organic-rich sediments. In: De Pauw, N., and Joyce, J. (eds.), Aquaculture and the Environment, Vol. 16. European Aquaculture Soc. Spec. Publ., pp. 155–175. Holsman, K.K., McDonald, P.S., and Armstrong, D.A. 2006. Intertidal migration and habitat use by subadult Dungeness crab Cancer magister in a NE Pacific estuary. Marine Ecology Progress Series 308:183–195. Holt, T.J., Rees, E.I., Hawkins, S.J., and Seed, R. 1998. Biogenic reefs, Volume 9: an overview of dynamic and sensitivity characteristics for conservation management of marine SACs. Scottish Association for Marine Science, Port Erin Marine Laboratory, University of Liverpool, Scotland. Hopkins, F.E., Turnera, S.M., Nightingale, P.D., Steinke, M., Bakkera, D., and Lissa, P.S. 2010. Ocean acidification and marine trace gas emissions. PNAS 107:760–765. Hosack, G.R., Dumbauld, B.R., Ruesink, J.L., and Armstrong, D.A. 2006. Habitat associations of estuarine species: comparisons of intertidal mudflat, seagrass (Zostera marina), and oyster (Crassostrea gigas) habitats. Estuaries Coasts 29:1150–1160. Huang, C.H., Lin, H.J., Huang, T.C., Su, H.M., and Hung, J.J. 2008. Responses of phytoplankton and periphyton to system-scale removal of oyster-culture racks from a eutrophic tropical lagoon. Marine Ecology Progress Series 358:1–12. Hughes, A.R., Williams, S.L., Duarte, C.M., Heck, K.L., and Waycott, M. 2009. Associations of concern: declining seagrasses and threatened dependent species. Frontiers in Ecology and the Environment 7:242–246. Inglis, G.J., and Gust, N. 2003. Potential indirect effects of shellfish culture on the reproductive success of benthic predators. The Journal of Applied Ecology 40:1077–1089. Jackson, J.B.C., Kirby, M.X., Berger, W.H., Bjorndal, K.A., Botsford, L.W., Bourque, B.J., Bradbury, R.H., Cooke, R., Erlandson, J., Estes, J.A., Hughes, T.P., Kidwell, S., Lange, C.B., Lenihan, H.S., Pandolfi, J.M., Peterson, C.H.,

Steneck, R.S., Tegner, M.J., and Warner, R.R. 2001. Historical overfishing and the recent collapse of coastal ecosystems. Science 293:629– 638. Jackson, E.L., Rowden, A.A., Attrill, M.J., Bossey, S.J., and Jones, M.B. 2001. The importance of seagrass beds as a habitat for fishery species. Oceanography Marine Biology: An Annual Review 39:269–303. Johnson, M.W., Heck, K.L., Jr., and Fourqurean, J.W. 2006. Nutrient content of seagrasses and epiphytes in the northern Gulf of Mexico: evidence of phosphorus and nitrogen limitation. Aquatic Botany 85:103–111. Jones, C.G., Lawton, J.H., and Shachak, M. 1994. Organisms as ecosystem engineers. Oikos 69:373–386. Jones, C.G., Lawton, J.H., and Shackak, M. 1997. Positive and negative effects of organisms as physical ecosystem engineers. Ecology 78: 1946–1957. Judge, M.L., and Craig, S.F. 1997. Positive flow dependence in the initial colonization of a fouling community: results from in situ water current manipulations. Journal of Experimental Marine Biology and Ecology 210:209–222. Judge, M.L., Coen, L.D., and Heck, K.L., Jr. 1992. The effect of long-term alteration of in situ currents on the growth of Mercenaria mercenaria in the northern Gulf of Mexico. Limnology and Oceanography 37:1550–1559. Judge, M.L., Coen, L.D., and Heck, K.L., Jr. 1993. Does Mercenaria mercenaria encounter elevated food levels in seagrass beds? Results from a novel technique to collect suspended food resources. Marine Ecology Progress Series 92:141–150. Judge, M.L., Coen, L.D., and Hammerstrom, K. 2000. The ecological implications of high density hard clam (Mercenaria mercenaria) mariculture on tidal creek environments. Journal of Shellfish Research 19:609–610. Kaiser, M.J. 2000. Ecological effects of shellfish cultivation. In: Black, K.D. (ed.), Environmental Impacts of Aquaculture. Sheffield Academic Press, Sheffield, UK, pp. 51–75. Kaiser, M.J. 2001. Ecological effects of shellfish cultivation. In: Black, K.D. (ed.), Environmental Impacts of Aquaculture. CRC Press, Boca Raton, FL.

Impacts of native and cultured bivalves

Kaiser, M.J., Laing, I., Utting, S.D., and Burnell, G.M. 1998. Environmental impacts of bivalve mariculture. Journal of Shellfish Research 17: 59–66. Kathiresan, K., and Rajendran, N. 2005. Coastal mangrove forests mitigated tsunami. Estuarine, Coastal and Shelf Science 65:601–606. Kaufman, L., and Dayton, P.K. 1997. Impacts of marine resource extraction on ecosystem services and sustainability. In: Daily, G.C. (ed.), Nature’s Services: Societal Dependence on Natural Ecosystems. Island Press, Washington, DC, pp. 275–293. Kelly, J.P., Evens, J.G., Stallcup, R.W., and Wimpfheimer, D. 1996. Effects of oyster culture on habitat use by wintering shorebirds in Tomales Bay, California. California Fish Game 82:160–174. Kelly, J.R., Proctor, H., and Volpe, J.P. 2008. Intertidal community structure differs significantly between substrates dominated by native eelgrass (Zostera marina L.) and adjacent to the introduced oyster Crassostrea gigas (Thunberg) in British Columbia, Canada. Hydrobiologia 596:57–66. Kemper, C., Pemberton, D., Cawthorn, M., Heinrich, S., Mann, J., Würsig, B., Shaughnessy, P.D., and Gales, R. 2003. Aquaculture and marine mammals: coexistence or conflict? Gales, N., Hindell, M., and Kirkwood, R. (eds.), Marine Mammals: Fisheries, Tourism and Management Issues. CSIRO Publishing, Collingwood, NSW, Australia. Kennedy, V.S. 1996. Biology of larvae and spat. In: Kennedy, V.S., Newell, R.I.E., and Eble, A.F. (eds.), The Eastern Oyster: Crassostrea Virginica. Maryland Sea Grant, College Park, MD, pp. 371–421. Kennedy, V.S., and Sanford, L.P. 1999. Characteristics of relatively unexploited beds of the eastern oyster, Crassostrea virginica, and early restoration programs. In: Luckenbach, M.W., Mann, R., and Wesson, J.A. (eds.), Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. Virginia Institute of Marine Science Press, Gloucester Point, VA, pp. 25–46. Kennedy, V.S., Newell, R.I.E., and Eble, A.F. (eds.). 1996. The Eastern Oyster: Crassostrea Virginica. Maryland Sea Grant, College Park, MD.

285

Keough, M.J. 1984. Dynamics of the epifauna of the bivalve Pinna bicolor: interactions among recruitment, predation and competition. Ecology 65:677–688. Kiessling, W., Simpson, C., and Foote, M. 2010. Reefs as cradles of evolution and sources of biodiversity in the Phanerozoic. Science 327:196– 198. Kimbro, D.L., and Grosholz, E.D. 2006. Disturbance influences oyster community richness and evenness, but not diversity. Ecology 87:2378– 2388. Kimbro, D.L., Largier, J., and Grosholz, E.D. 2009. Coastal oceanographic processes influence the growth and size of a key estuarine species, the Olympia oyster. Limnology and Oceanography 54:1425–1437. Kirby, M.X. 2004. Fishing down the coast: historical expansion and collapse of oyster fisheries along coastal margins. Proceedings of the National Shellfish Association 101:13096– 13099. Kirby, M.X., and Miller, H.M. 2005. Response of a benthic suspension feeder (Crassostrea virginica Gmelin) to three centuries of anthropogenic eutrophication in Chesapeake Bay. Estuarine, Coastal and Shelf Science 62:679–689. Kochmann, J., Buschbaum, C., Volkenborn, N., and Reise, K. 2008. Shift from native mussels to alien oysters: differential effects of ecosystem engineers. Journal of Experimental Marine Biology and Ecology 364:1–10. Kraan, C., van Gils, J.A., Spaans, B., Dekinga, A., Bijleveld, A.I., van Roomen, M., Kleefstra, R., and Piersma, T. 2009. Landscape-scale experiment demonstrates that Wadden Sea intertidal flats are used to capacity by molluscivore migrant shorebirds. The Journal of Animal Ecology 78:1259–1268. Kuhlmann, M.L. 1998. Spatial and temporal patterns in the dynamics and use of pen shells (Atrina rigida) as shelters in St. Joseph Bay, Florida. Bulletin of Marine Science 62:157– 179. Langton, R.W., and Robinson, W.E. 1990. Faunal associations on scallop grounds in the western Gulf of Maine. Journal of Experimental Marine Biology and Ecology 144:157–171. Larkum, A.W.D., Orth, R.J., and Duarte, C. (eds.). 2006. Seagrasses: Biology, Ecology and

286

Shellfish Aquaculture and the Environment

Conservation. Springer, Dordrecht, The Netherlands. Leguerrier, D., Niquil, N., Petiau, A., and Bodoy, A. 2004. Modeling the impact of oyster culture on a mudflat food web in Marennes-Oleron Bay (France). Marine Ecology Progress Series 273: 147–161. Lehnert, R.L., and Allen, D.M. 2002. Nekton use of subtidal oyster shell habitats in a southeastern U.S. estuary. Estuaries 25:1015–1024. Lenihan, H.S. 1999. Physical-biological coupling on oyster reefs: how habitat structure influences individual performance. Ecological Monographs 69:251–275. Lenihan, H.S., and Micheli, F. 2000. Biological effects of shellfish harvesting on oyster reefs: resolving a fishery conflict by ecological experimentation. Fishery Bulletin 98:86–95. Lenihan, H.S., and Peterson, C.H. 1998. How habitat degradation through fishery disturbance enhances impacts of hypoxia on oyster reefs. Ecological Applications 8:128–140. Lenihan, H.S., and Peterson, C.H. 2004. Conserving oyster reef habitat by switching from dredging and tonging to diver hand-harvesting. Fishery Bulletin 102:298–305. Lenihan, H.S., Peterson, C.H., Byers, J.E., Grabowski, J.H., Thayer, G.W., and Colby, D.R. 2001. Cascading of habitat degradation: oyster reefs invaded by refugee fishes escaping stress. Ecological Applications 11:748–764. Light, B.R., and Beardall, J. 1998. Distribution and spatial variation of benthic microalgal biomass in a temperate, shallow-water marine system. Aquatic Botany 61:39–54. Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L., Olrog, L., and Rehnstam-Holm, A. 2005. Improving marine water quality by mussel farming: a profitable solution for Swedish Society. Ambio 34:131–138. Lindsay, C.E., and Simons, D. 1997. The fisheries for Olympia oysters, Ostreola conchaphila; Pacific oysters, Crassostrea gigas; and Pacific razor clams, Siliqua patula, in the State of Washington. In: Mackenzie, C.L.J., Burrell, V.G.J., Rosenfield, A., and Hobart, W.L. (eds.), The History, Present Condition, and Future of the Molluscan Fisheries of North and Central America and Europe, Vol. 2, Pacific Coast and Supplemental Topics. Seattle, WA: NOAA, U.S. Department of Commerce, pp. 89–113.

Lipton, D. 2004. The value of improved water quality to Chesapeake Bay boaters. Marine Resource Economics 19:265–270. Lloyd, B.D. 2003. Potential Effects of Mussel Farming on New Zealand’s Marine Mammals and Seabirds: A Discussion Paper. Department of Conservation, Wellington, New Zealand. Lotze, H.K., Leniham, H.S., Bourque, B.J., Bradbury, R.H., Cooke, R.G., Kay, M.C., Kidwell, S.M., Kirby, M.X., Peterson, C.H., and Jackson, J.B.C. 2006. Depletion, degredation, and recovery potential of estuaries and coastal seas. Science 312:1806–1809. Lu, L., and Grant, J. 2008. Recolonization of intertidal infauna in relation to organic deposition at an oyster farm in Atlantic Canada—a field experiment. Estuaries and Coasts 31:767– 775. Luckenbach, M.W., and Orth, R.J. 1999. Interactions between benthic infauna and the transport and burial of Zostera marina seeds. Aquatic Botany 62:235–247. Luckenbach, M.W., Mann, R., and Wesson, J.A. (eds.). 1999. Oyster Reef Habitat Restoration. A Synopsis and Synthesis of Approaches. Virginia Institute of Marine Science Press, Gloucester Point, VA. Luckenbach, M.W., Coen, L.D., Ross, P.G., Jr., and Stephen, J.A. 2005. Oyster reef habitat restoration: relationships between oyster abundance and community development based on two studies in Virginia and South Carolina. Journal of Coastal Research Special Issue (40):64–78. Macleod, C.K., Forbes, S.E., Shepherd, C.J., and Crawford, C. 2009. Effects of oyster farming service vehicles on an intertidal sand flat. Aquaculture Research 40:772–780. Mallet, A.L., Carver, C.E., and Landry, T. 2006. Impact of suspended and off-bottom eastern oyster culture on the benthic environment in eastern Canada. Aquaculture 255:362–373. Mallet, A.L., Carver, C.E., and Hardy, M. 2009. The effect of floating bag management strategies on biofouling, oyster growth and biodeposition levels. Aquaculture 287:315–323. Mann, R., and Evans, E.A. 2004. Site selection for oyster habitat rehabilitation in the Virginia portion of the Chesapeake Bay: a commentary. Journal of Shellfish Research 23:41–49. Mann, R., and Powell, E.N. 2007. Why oyster restoration goals in the Chesapeake Bay are not and

Impacts of native and cultured bivalves

probably cannot be achieved. Journal of Shellfish Research 26:905–917. Markert, A., Wehrmann, A., and Kröncke1, I. 2010. Recently established Crassostrea-reefs versus native Mytilus-beds: differences in ecosystem engineering affects the macrofaunal communities (Wadden Sea of Lower Saxony, southern German Bight). Biological Invasions 12:15–32. McGlathery, K.J., Sundbäck, K., and Anderson, I.C. 2007. Eutrophication in shallow coastal bays and lagoons: the role of plants in the coastal filter. Marine Ecology Progress Series 348:1–18. McGraw, K.A. 2009. The Olympia oyster, Ostrea lurida Carpenter, 1864 along the west coast of North America. Journal of Shellfish Research 28:5–10. McKindsey, C.W., Thetmeyer, H., Landry, T., and Silvert, W. 2006. Review of recent carrying capacity models for bivalve culture and recommendations for research and management. Aquaculture 261:451–462. McKindsey, C.W., Landry, T., O’Beirn, F.X., and Davies, I.M. 2007. Bivalve aquaculture and exotic species: a review of ecological considerations and management issues. Journal of Shellfish Research 26:281–294. McKindsey, C.W., Lecuona, M., Huot, M., and Weise, A.M. 2009. Biodeposit production and benthic loading by farmed mussels and associated tunicate epifauna in Prince Edward Island. Aquaculture 295:44–51. McVey, J.P., Stickney, R., Yarish, C., and Chopin, T. 2002. Aquatic polyculture and balanced ecosystem management: new paradigms for seafood production. In: Stickney, R.R., and McVey, J.P. (eds.), Responsible Marine Aquaculture. CABI Publishing, Oxon, UK, pp. 91–104. Mesnage, V., Ogier, S., Bally, G., Disnar, J.R., Lottier, N., Dedieu, K., Rabouille, C., and Copard, Y. 2007. Nutrient dynamics at the sediment-water interface in a Mediterranean lagoon (Thau, France): influence of biodeposition by shellfish farming activities. Marine Environmental Research 63:257–277. Meyer, D.L., and Townsend, E.C. 2000. Faunal utilization of created intertidal eastern oyster (Crassostrea virginica) reefs in the southeastern United States. Estuaries 23:34–45. Meyer, D.L., Townsend, E.C., and Thayer, G.W. 1997. Stabilization and erosion control value of

287

oyster cultch for intertidal marsh. Restoration Ecology 5:93–99. Michael, P., and Chew, K.K. 1976. Growth of Pacific oyster Crassostrea gigas and related fouling problems under tray culture at Seabeck Bay, Washington. Proceedings of the National Shellfish Association 66:36–41. Micheli, F., and Peterson, C.H. 1999. Estuarine vegetated habitats as corridors for predator movements. Conservation Biology 13:869– 881. Millennium Ecosystem Assessment (MA) (ed.) 2003. Ecosystems and Human Well-Being: A Framework for Assessment. Island Press, Washington, DC. Miller, D.C., Norkko, A., and Pilditch, C.A. 2002. Influence of diet on dispersal of horse mussel Atrina zelandica biodeposits. Marine Ecology Progress Series 242:153–167. Mojica, R., Jr., and Nelson, W.G. 1993. Environmental effects of a hard clam (Mercenaria mercenaria) aquaculture site in the Indian River Lagoon, Florida. Aquaculture 113:313– 329. Molnar, J.L., Gamboa, R.L., Revenga, C., and Spalding, M.D. 2008. Assessing the global threat of invasive species to marine biodiversity. Frontiers in Ecology and the Environment 6:485–492. Munguia, P. 2007. Spatial structure of pen shell (Atrina rigida) communities. Marine Biology 152:149–156. Munroe, D., and McKinley, R.S. 2007a. Effects of predator netting on recruitment and growth of Manila clams (Venerupis philippinarum) on soft substrate intertidal plots in British Columbia, Canada. Journal of Shellfish Research 26:1035–1044. Munroe, D., and McKinley, R.S. 2007b. Commercial Manila clam (Tapes philippinarum) culture in British Columbia, Canada: the effects of predator netting on intertidal sediment characteristics. Estuarine, Coastal and Shelf Science 72: 319–328. Nash, C.E., Iwamoto, R.N., and Mahnken, C.V.W. 2000. Aquaculture risk management and marine mammal interactions in the Pacific Northwest. Aquaculture 183:307–323. National Research Council (NRC). 2004. Nonnative Oysters in the Chesapeake Bay. National Academies Press, Washington, DC.

288

Shellfish Aquaculture and the Environment

National Research Council (NRC). 2007. Mitigating Shore Erosion along Sheltered Coasts. National Academies Press, Washington, DC. National Research Council (NRC). 2009. Shellfish mariculture in Drakes Estero, Point Reyes National Seashore, California. Committee on Best Practices for Shellfish Mariculture and the Effects of Commercial Activities in Drakes Estero, Pt. Reyes National Seashore, California, 139pp. National Research Council (NRC). 2010. Ecosystem concepts for sustainable bivalve mariculture. Committee on Best Practices for Shellfish Mariculture and the Effects of Commercial Activities in Drakes Estero, Pt. Reyes National Seashore, California, 179pp. Naylor, R.L., Goldburg, R.J., Primavera, J.H., Kautsky, N., Beveridge, M.C.M., Clay, J., Folke, C., Lubchenco, J., Mooney, H., and Troell, M. 2000. Effect of aquaculture on world fish supplies. Nature 405:1017–1024. Naylor, R.L., Williams, S.L., and Strong, D.R. 2001. Aquaculture—a gateway for exotic species. Science 294:1655–1666. Neckles, H.A., Short, F.T., Barker, S., and Kopp, B.S. 2005. Disturbance of eelgrass Zostera marina by commercial mussel Mytilus edulis harvesting in Maine: dragging impacts and habitat recovery. Marine Ecology Progress Series 285:57–73. Nelson, K.A., Leonard, L.A., Posey, M.H., Alphin, T.D., and Mallin, M.A. 2004. Using transplanted oyster beds to improve water quality in small tidal creeks: a pilot study. Journal of Experimental Marine Biology and Ecology 298:347–368. Neori, A., Chopin, T., Troell, M., Buschmann, A.H., Kraemer, G.P., Halling, C., Shpigel, M., and Yarish, C. 2004. Integrated aquaculture: rationale, evolution and state of the art emphasizing seaweed biofiltration in modern mariculture. Aquaculture 231:361–391. Nestlerode, J.A. 2004. Evaluating restored oyster reefs in Chesapeake Bay: how habitat structure influences ecological function. Ph.D. Dissertation, VIMS, 262pp. Newell, R.I.E. 1988. Ecological changes in Chesapeake Bay: are they the result of overharvesting the eastern oyster (Crassostrea virginica)? Lynch, M.P., and Krome, E.C. (eds.), Understanding the Estuary: Advances in Chesapeake Bay Research. Chesapeake Research

Consortium Publication 129, Gloucester Point, VA, pp. 536–546. Newell, R.I.E. 2004. Ecosystem influences of natural and cultivated populations of suspension-feeding bivalve molluscs: a review. Journal of Shellfish Research 23:51–61. Newell, R.I.E. 2007. A framework for developing “ecological carrying capacity” mathematical models for bivalve mollusc aquaculture. Bulletin of Fisheries Research Agency 19:41–52. Newell, R.I.E., and Koch, E.W. 2004. Modeling seagrass density and distribution in response to changes in turbidity stemming from bivalve filtration and seagrass sediment stabilization. Estuaries 27:793–806. Newell, R.I.E., Cornwell, J.C., and Owens, M.S. 2002. Influence of simulated bivalve biodeposition and microphytobenthos on sediment nitrogen dynamics: a laboratory study. Limnology and Oceanography 47:1367–1379. Newell, R., Fisher, T., Holyoke, R., and Cornwell, J. 2005. Influence of Eastern oysters on nitrogen and phosphorus regeneration in Chesapeake Bay, USA. In: Dame, R., and Olenin, S. (eds.), The Comparative Roles of Suspension Feeders in Ecosystems, Vol. 47, NATO Science Series: IV— Earth and Environmental Sciences. Springer, Dordrecht, the Netherlands, pp. 93–120. Newell, R.I.E., Kemp, W.M., Hagy, J.D., III, Cerco, C.F., Testa, J.M., and Boynton, W.R. 2007. Topdown control of phytoplankton by oysters in Chesapeake Bay, USA: comment on Pomeroy et al. (2006). Marine Ecology Progress Series 341:293–298. Nixon, S.W. 1980. Between coastal marshes and coastal waters: a review of twenty years of speculation and research on the role of salt marshes in estuarine productivity and water chemistry. In: Hamilton, P., MacDonald, P., and MacDonald, K.B. (eds.), Estuarine and Wetland Processes. Plenum Press, New York, pp. 437–525. Nizzoli, D., Welsh, D.T., Fano, E.A., and Viaroli, P. 2006. Impact of clam and mussel farming on benthic metabolism and nitrogen cycling, with emphasis on nitrate reduction pathways. Marine Ecology Progress Series 315:151–165. Norkko, A., Hewitt, J.E., Thrush, S.F., and Funnell, G.A. 2006. Conditional outcomes of facilitation by a habitat-modifying subtidal bivalve. Ecology 87:226–234.

Impacts of native and cultured bivalves

Norris, K., Bannister, R.C.A., and Walker, P.W. 1998. Changes in the number of oystercatchers Haematopus ostralegus wintering in the Burry Inlet in relation to the biomass of cockles Cerastoderma edule and its commercial exploitation. The Journal of Applied Ecology 35:75–85. North, E.W., King, D.M., Xu, J., Hood, R.R., Newell, R.I.E., Paynter, K.T., Kellogg, M.L., Liddel, M.K., and Boesch, D.F. 2010. Linking optimization and ecological models in a decision support tool for oyster restoration and management. Ecological Applications 20:851–866. Nosho, T.Y., and Chew, K.K. 1991. Remote Setting and Nursery Culture for Shellfish Growers, Workshop Record, February 19, 1991. Washington Sea Grant, Seattle, WA. Nowell, A.R.M., and Jumars, P.A. 1984. Flow environments of aquatic benthos. Annual Review of Ecology and Systematics 15:303–328. Nugues, M.M., Kaiser, M.J., Spencer, B.E., and Edwards, D.B. 1996. Benthic community changes associated with intertidal oyster cultivation. Aquaculture Research 27:913–924. O’Beirn, F.X., Ross, P.G., and Luckenbach, M.W. 2004. Organisms associated with oysters cultured in floating systems in Virginia, USA. Journal of Shellfish Research 23:825–829. Officer, C.B., Smayda, T.J., and Mann, R. 1982. Benthic filter feeding: a natural eutrophication control. Marine Ecology Progress Series 9:203–210. ORET (Oyster Restoration Evaluation Team). 2009. Metadata analysis of restoration and monitoring activity database. Kramer, J.G., and Sellner, K.G. (eds.), Native Oyster (Crassostrea virginica) Restoration in Maryland and Virginia. An evaluation of lessons learned 1990–2007. Maryland Sea Grant Publication #UM-SG-TS-2009-02; CRC Publ. No. 09-168. College Park, MD. Available at http://www. oyster-restoration.org/reports/OysterResReport_ web.pdf. Orth, R.J. 1975. Destruction of eelgrass, Zostera marina, by the cownose ray, Rhinoptera bonasus, in the Chesapeake Bay. Chesapeake Science 16:205–208. Orth, R.J., Fishman, J.R., Wilcox, D.J., and Moore, K.A. 2002. Identification and management of fishing gear impacts in a recovering seagrass

289

system in the coastal bays of the Delmarva Peninsula, USA. Journal of Coastal Research, Special Issue (37):111–129. Orth, R.J., Carruthers, T.J.B., Dennison, W.C., Duarte, C.M., Fourqurean, J.W., Heck, K.L., Jr., Hughes, R., Kendrick, G., Kenworthy, W.J., Olyarnik, S., Short, F.T., Waycott, M., and Williams, S.L. 2006. A global crisis for seagrass ecosystems. Bioscience 56:987–996. Palacios, R., Armstrong, D.A., and Orensanz, J. 2000. Fate and legacy of an invasion: extinct and extant populations of the soft-shell clam (Mya arenaria) in Grays Harbor (Washington). Aquatic Conservation: Marine and Freshwater Ecosystems 10:279–303. Paltzat, D.L., Pearce, C.M., Barnes, P.A., and McKinley, R.S. 2008. Growth and production of California sea cucumbers (Parastichopus californicus Stimpson) co-cultured with suspended Pacific oysters (Crassostrea gigas Thunberg). Aquaculture 275:124–137. Paraskevi, P., Tsapkias, M., Apostolaki, E.T., Tsagaraki, T., Holmer, M., and Karakassis, I. 2009. Ghost nutrients from fish farms are transferred up the food web by phytoplankton grazers. Marine Ecology Progress Series 374:1–6. Perry, L.M. 1936. A marine tenement. Science 84:156–157. Peterson, B.J., and Heck, K.L. 1999. The potential for suspension feeding bivalves to increase seagrass productivity. Journal of Experimental Marine Biology and Ecology 240:37–52. Peterson, B.J., and Heck, K.L., Jr. 2001a. Positive interactions between suspension-feeding bivalves and seagrass-a facultative mutualism. Marine Ecology Progress Series 213:143–155. Peterson, B.J., and Heck, K.L. 2001b. An experimental test of the mechanism by which suspension feeding bivalves elevate seagrass productivity. Marine Ecology Progress Series 218:115–125. Peterson, C.H., and Lipcius, R.N. 2003. Conceptual progress towards predicting quantitative ecosystem benefits of ecological restorations. Marine Ecology Progress Series 264:297–307. Peterson, C.H., Grabowski, J.H., and Powers, S.P. 2003. Estimated enhancement of fish production resulting from restoring oyster reef habitat: quantitative valuation. Marine Ecology Progress Series 264:251–256.

290

Shellfish Aquaculture and the Environment

Piazza, B.P., Banks, P.D., and La Peyre, M.K. 2005. The potential for created oyster shell reefs as a sustainable shoreline protection strategy in Louisiana. Restoration Ecology 13:499–506. Pienkowski, M.W. 1983. Surface activity of some intertidal invertebrates in relation to temperature and the foraging behaviour of their shorebird predators. Marine Ecology Progress Series 11:141–150. Pillay, T.V.R. 1992. Aquaculture and the Environment. Halsted Press, New York. Pitta, P., Manolis, T., Apostolaki, E.T., Tsagaraki, T., Holmer, M., and Karakassis, I. 2009. “Ghost nutrients” from fish farms are transferred up the food web by phytoplankton grazers. Marine Ecology Progress Series 374:1–6. Polson, M., Hewson, W.E., Eemisse, D.J., Baker, P.K., and Zacherl, D.C. 2009. You say conchaphila, I say lurida: molecular evidence for restricting the Olympia oyster (Ostrea lurida Carpenter 1864) to temperate Western North America. Journal of Shellfish Research 28:11–21. Porter, E.T., Sanford, L.P., and Suttles, S.E. 2000. Gypsum dissolution is not a universal integrator of “water motion”. Limnology and Oceanography 45:145–158. Porter, E.T., Cornwell, J.C., Sanford, L.P., and Newell, R.I.E. 2004. Effect of oysters Crassostrea virginica and bottom shear velocity on benthicpelagic coupling and estuarine water quality. Marine Ecology Progress Series 271:61– 75. Posey, M.H. 1986. Predation on a burrowing shrimp: distribution and community consequences. Journal of Experimental Marine Biology and Ecology 103:143–161. Posey, M.H. 1988. Community changes associated with the spread of an introduced seagrass, Zostera japonica. Ecology 69:974–983. Posey, M.H., Alphin, T.D., Powell, C.M., and Townsend, E. 1999. Use of oyster reefs as habitat for epibenthic fish and decapods. In: Luckenbach, M.W., Mann, R., and Wesson, J.E. (eds.), Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. VIMS Press, Gloucester Point, VA, pp. 229–237. Powell, E.N., and Klinck, J.M. 2007. Is oyster shell a sustainable estuarine resource? Journal of Shellfish Research 26:181–194.

Powell, E.N., Kraeuter, J.N., and Ashton-Alcox, K.A. 2006. How long does oyster shell last on an oyster reef? Estuarine, Coastal and Shelf Science 69:531–542. Powell, E.N., Ashton-Alcox, K.A., Kraeuter, J.N., Ford, S.E., and Bushek, D. 2008. Long-term trends in oyster population dynamics in Delaware Bay: regime shifts and response to disease. Journal of Shellfish Research 27:729–755. Powers, M.J., Peterson, C.H., Summerson, H.C., and Powers, S.P. 2007. Macroalgal growth on bivalve aquaculture netting enhances nursery habitat for mobile invertebrates and juvenile fishes. Marine Ecology Progress Series 339: 109–122. Powers, S.P., Peterson, C.H., Grabowski, J.H., and Lenihan, H.S. 2009. Success of constructed oyster reefs in no-harvest sanctuaries: implications for restoration. Marine Ecology Progress Series 389:159–170. Quammen, M.L. 1981. Use of exclosures in studies of predation by shorebirds on intertidal mudflats. The Auk 98:812–817. Quammen, M.L. 1982. Influence of subtle substrate differences on feeding by shorebirds on intertidal mudflats. Marine Biology 71:339–343. Quammen, M.L. 1984. Predation by shorebirds, fish, and crabs on invertebrates in intertidal mudflats: an experimental test. Ecology 65:529–537. Read, P., and Fernandes, T.F. 2003. Management of environmental impacts of marine aquaculture in Europe. Aquaculture 226:139–163. Read, A.J., Drinker, P., and Northridge, S. 2006. Bycatch of marine mammals in U.S. and global fisheries. Conservation Biology 20:163–169. Reise, K. 1998. Pacific oysters invade mussel beds in the European Wadden Sea. Senckenbergiana Maritima 28:167–175. Reise, K., Bouma, T.J., Olenin, S., and Ysebaert, T. 2009. Coastal habitat engineers and the biodiversity in marine sediments. Helgoland Marine Research 63:1–2. Reusch, T.B.H., Chapman, A.R.O., and Groger, J.P. 1994. Blue mussels Mytilus edulis do not interfere with eelgrass Zostera marina but fertilize shoot growth through biodeposition. Marine Ecology Progress Series 108:265–282. Rice, M.A. 2000. A review of shellfish restoration as a tool for coastal water quality management. Environment Cape Cod 3:1–8.

Impacts of native and cultured bivalves

Richardson, N.F., Ruesink, J.L., Naeem, S., Hacker, S.D., Tallis, H.M., Dumbauld, B.R., and Wisehart, L.M. 2008. Bacterial abundance and aerobic microbial activity across natural and oyster aquaculture habitats during summer conditions in a northeastern Pacific estuary. Hydrobiologia 596:269–278. Robinson, A.M. 1997. Molluscan fisheries in Oregon: past, present and future. In: Mackenzie, C.L.J., Burrell, V.G.J., Rosenfield, A., and Hobart, W.L. (eds.), The History, Present Condition, and Future of the Molluscan Fisheries of North and Central America and Europe, Vol. 2, Pacific Coast and Supplemental Topics. U.S. Dept. Commerce, NOAA Tech. Rep. NMFS 128. Seattle, WA: NOAA, U.S. Department of Commerce, pp. 75–87. Rodgers, J.A., and Smith, H.T. 1997. Buffer zone distances to protect foraging and loafing waterbirds from human disturbance in Florida. Wildlife Society Bulletin 25:139–145. Rodney, W.S., and Paynter, K.T. 2006. Comparisons of macrofaunal assemblages on restored and non-restored oyster reefs in mesohaline regions of Chesapeake Bay in Maryland. Journal of Experimental Marine Biology and Ecology 335:39–51. Rothschild, B.J., Ault, J.S., Goulletquer, P., and Héral, M. 1994. Decline of the Chesapeake Bay oyster population: a century of habitat destruction and overfishing. Marine Ecology Progress Series 111:29–39. Roycroft, D., Kelly, T.C., and Lewis, L.J. 2006. Behavioural interactions of seabirds with suspended mussel longlines. Aquaculture International 15:25–36. Ruesink, J.L., Lenihan, H.S., Trimble, A.C., Heiman, K.W., Micheli, F., Byers, J.E., and Kay, M.C. 2005. Introduction of non-native oysters: ecosystem effects and restoration implications. Annual Review of Ecology and Systematics 36:643–689. Ruesink, J.L., Feist, B.E., Harvey, C.J., Hong, J.S., Trimble, A.C., and Wisehart, L.M. 2006. Changes in productivity associated with four introduced species: ecosystem transformation of a “pristine” estuary. Marine Ecology Progress Series 311:203–215. Ruesink, J.L., Hacker, S.D., Dumbauld, B.R., and Harbell, S. 2009. Scale-dependent and indirect

291

effects of filter feeders on eelgrass: understanding complex ecological interactions to improve environmental impacts of aquaculture. Dept. of Biology, University of Washington. Final Report to the Western Regional Aquaculture Center Seattle, Washington, 25pp. Ruiz, G.M., Fofonoff, P.W., Carlton, J.T., Wonham, W.J., and Hines, A.H. 2000. Invasion of coastal marine communities in North America: apparent patterns, processes, and biases. Annual Review of Ecology and Systematics 31:481–531. Rumrill, S.S., and Poulton, V.K. 2004. Ecological role and potential impacts of molluscan shellfish culture in the estuarine environment of Humboldt Bay, Ca. Oregon Department of State Lands, Final Annual Report to the Western Regional Aquaculture Center, 22pp. Safriel, U.N. 1975. The role of vermetid gastropods in the formation of Mediterranean and Atlantic reefs. Oecologia 20:85–101. Salisbury, J., Green, M., Hunt, C., and Campbell, J. 2008. Coastal acidification by rivers: a new threat to shellfish? Eos Transactions AGU 89:513–514. Sanchez, M.I., Green, A.J., and Castellanos, E.M. 2006. Spatial and temporal fluctuations in presence and use of chironomid prey by shorebirds in the Odiel saltpans, south-west Spain. Hydrobiologia 567:329–340. Schulte, D.M., Burke, R.P., and Lipcius, R.N. 2009. Unprecedented restoration of a native oyster metapopulation. Science 325:1124–1128. Sebens, K.P. 1991. Habitat structure and community dynamics in marine benthic systems. In: Bell, S.S., McCoy, E.D., and Mushinsky, H.R. (eds.), Habitat Structure: the Physical Arrangement of Objects in Space. Chapman and Hall, London, pp. 211–234. Seed, R. 1992. Systematics evolution and distribution of mussels belonging to the genus Mytilus: an overview. American Malacological Bulletin 9:123–137. Seed, R., and Suchanek, T.H. 1992. Population and community ecology of Mytilus. In: Gosling, E.M. (ed.), The Mussel Mytilus: Ecology, Physiology, Genetics, and Culture. Elsevier Science, Amsterdam, pp. 87–169. Sellers, M.A., and Stanley, J.G. 1984. Species profiles: life histories and environmental requirements of coastal fishes and invertebrates (north

292

Shellfish Aquaculture and the Environment

Atlantic)-American Oyster. U.S. F.W.S. Office of Biological Services Report No. FWS/OBS82/11.23, and USACE Report No. TR EL-82-4, Washington, DC. Semmens, B.X. 2008. Acoustically derived finescale behaviors of juvenile Chinook salmon associated with intertidal benthic habitats in an estuary. Canadian Journal of Fisheries and Aquatic Sciences 65:2053–2062. Shaw, W.N. 1997. The shellfish industry of California- Past, present and future. In: Mackenzie, C.L.J., Burrell, V.G.J., Rosenfield, A., and Hobart, W.L. (eds.), The History, Present Condition, and Future of the Molluscan Fisheries of North and Central America and Europe, Vol. 2, Pacific Coast and Supplemental Topics. U.S. Dept. Commerce, NOAA Tech. Rep. NMFS 128. Seattle, WA: NOAA, U.S. Department of Commerce, pp. 57–74. Shumway, S.E., and Kraeuter, J.N. (eds.) 2004. Molluscan shellfish research and management: charting a course for the future. Final proceedings from the Cooperative Research and Information Institute (CRII) Workshop, Charleston, SC, 156pp. Simenstad, C.A., and Fresh, K.L. 1995. Influence of intertidal aquaculture on benthic communities in Pacific Northwest estuaries: scales of disturbance. Estuaries 18:43–70. Smaal, A.C., Kater, B.J., and Wijsman, J. 2009. Introduction, establishment and expansion of the Pacific oyster Crassostrea gigas in the Oosterschelde (SW Netherlands). Helgoland Marine Research 63:75–83. Smith, G.F., Bruce, D.G., Roach, E.B., Hansen, A., Newell, R.I.E., and McManus, A.M. 2005. Habitat conditions of mesohaline oyster bars in the Maryland Chesapeake Bay: an assessment of 40 years of oyster management. North American Journal of Fisheries Management 25:1569– 1590. Smith, K.A., North, E.W., Shi, F., Chen, S., Hood, R.R., Koch, E.W., and Newell, R.I.E. 2009. Modeling the effects of oyster reefs and breakwaters on seagrass growth. Estuaries and Coasts 32:748–757. Smyth, D., Roberts, D., and Browne, L. 2009. Impacts of unregulated harvesting on a recovering stock of native oysters (Ostrea edulis). Marine Pollution Bulletin 58:916–922.

Soniat, T.M., Finelli, C.M., and Ruiz, J.T. 2004. Vertical structure and predator refuge mediate oyster reef development and community dynamics. Journal of Experimental Marine Biology and Ecology 310:163–182. Sousa, R., Gutiérrez, J.L., and Aldridge, D.C. 2009. Non-indigenous invasive bivalves as ecosystem engineers. Biology Invasions 11:2367–2385. Southworth, M., and Mann, R. 1998. Oyster reef broodstock enhancement in the Great Wicomico River, Virginia, Journal of Shellfish Research 17:1101–1114. Stal, L.J. 2010. Microphytobenthis as a biogeomorphological force in intertidal sediment stabilization. Ecological Engineering 36:236–245. Stanley, D.W., and Sellers, M.A. 1986. Species profile: life histories and environmental requirements of coastal fishes and invertebrates (Gulf of Mexico)-American Oyster. U.S. Fish Wildl. Serv. Biol Rep. 82(11.64) U.S. Army Corps of Engineers, TR EL-82-4, 25pp. Steele, E.N. 1964. The Immigrant Oyster (Ostrea Gigas) Now Known as the Pacific Oyster. Warren’s Quick Print, Olympia, WA. Steimle, F.W., and Zetlin, C. 2000. Reef habitats in the Middle Atlantic Bight: abundance, distribution, associated biological communities, and fishery resource use. Marine Fisheries Review 62:24–42. Stempien, J.A. 2007. Detecting avian predation on bivalve assemblages using indirect methods. Journal of Shellfish Research 26:271–280. Stillman, R.A., West, A.D., Caldow, R.W.G., and le V. dit Durell, S.E.A. 2007. Predicting the, effect of disturbance on coastal birds. Ibis 149:73–81. Stokesbury, K.D.E. 2002. Estimation of sea scallop abundance in closed areas of Georges Bank, USA. Transactions of the American Fisheries Society 131:1081–1092. Stoner, A.W., and Titgen, R.H. 2003. Biological structures and bottom type influence habitat choices made by Alaska flatfishes. Journal of Experimental Marine Biology and Ecology 292:43–59. Stunz, G.W., Minello, T.J., and Rozas, L.R. 2010. Relative value of oyster reef as habitat for estuarine nekton in Galveston Bay, Texas. Marine Ecology Progress Series 406:147– 159.

Impacts of native and cultured bivalves

Suchanek, T.H. 1985. Mussels and their role in structuring rocky shore communities. In: Moore, P.G., and Seed, R. (eds.), The Ecology of Rocky Coasts: Essays Presented to J.R. Lewis. Columbia Press, New York, pp. 70–96. Tallis, H.M., Ruesink, J.L., Dumbauld, B., Hacker, S., and Wisehart, L.M. 2009. Oysters and aquaculture practices affect eelgrass density and productivity in a Pacific Northwest estuary. Journal of Shellfish Research 28:251–261. Tallman, J.C., and Forrester, G.E. 2007. Oyster grow-out cages function as artificial reefs for temperate fishes. Transactions of the American Fisheries Society 136:790–799. Talman, S.G., Norkko, A., Thrush, S.F., and Hewitt, J.E. 2004. Habitat structure and the survival of juvenile scallops Pecten novaezelandiae: comparing predation in habitats with varying complexity. Marine Ecology Progress Series 269:197–207. Tenore, K.R., and Gonzalez, N. 1975. Food chain patterns in the Ría de Arosa, Spain: an area of intense mussel aquaculture. In: Persoone, G., and Jaspers, E. (eds.), 10th European Symposium on Marine Biology, Vol. 2. Universal Press, Wetteren, Belgium, pp. 601–619. Tenore, K.R., Browne, M.G., and Chesney, E.J. 1974. Polyspecies aquaculture systems: the detrital trophic level. Journal of Marine Research 32:425–432. Tenore, K.R., Boyer, L.F., Cal, R.M., Corral, J., Garcia-Fernandez, C., Gonzalez, N., GonzalezGurriaran, E., Hanson, R.B., Iglesias, J., Krom, M., Lopez-Jamar, E., McClain, J., Pamatmat, M.M., Perez, A., Rhoads, D.C., de Santiago, G., Tietjen, J., Westrich, J., and Windom, H.L. 1982. Coastal upwelling in the Rias Bajas, NW Spain: contrasting the benthic regimes of the Rias de Arosa and de Muros. Journal of Marine Research 40:701–772. Tevesz, M.J.S., and McCall, P.L. (eds.). 1983. Biotic Interactions in Recent and Fossil Benthic Communities. Plenum Press, New York. Thayer, G.W. (ed.) 1992. Restoring the Nation’s Marine Environment. Maryland Sea Grant College Program, College Park, MD. Thayer, G.W., McTigue, T.A., Bellmer, R.J., Burrows, F.M., Merkey, D.H., Nickens, A.D., Lozano, S.J., Gayaldo, P.F., Polmateer, P.J., and Pinit, P.T. 2003. Science-Based Restoration

293

Monitoring of Coastal Habitats, Volume 1: A Framework for Monitoring Plans under the Estuaries and Clean Waters Act of 2000 (Public Law 160-457). NOAA Coastal Ocean Program Decision Analysis Series No. 23. NOAA National Centers for Coastal Ocean Science, Silver Spring, MD. Thayer, G.W., McTigue, T.A., Salz, R.J., Merkey, D.H., Burrows, F.M., and Gayaldo, P.F. (eds.) 2005. Science-Based Restoration Monitoring of Coastal Habitats, Volume 2: Tools for Monitoring Coastal Habitats. NOAA Coastal Ocean Program Decision Analysis Series No. 23. NOAA National Centers for Coastal Ocean Science, Silver Spring, MD. Thieltges, D.W., Strasser, M., and Reise, K. 2006. How bad are invaders in coastal waters? The case of the American slipper limpet Crepidula fornicata in western Europe. Biological Invasions 8:1673–1680. Thom, R.M., Southard, S.L., Borde, A.B., and Stoltz, P. 2008. Light requirements for growth and survival of eelgrass (Zostera marina L.) in Pacific Northwest (USA) estuaries. Estuaries and Coasts 31:969–980. Thompson, D.S. 1995. Substrate additive studies for the development of hardshell clam habitat in Washington State: an analysis of effects on recruitment, growth and survival of the Manila clam, Tapes philippinarum, and on the species diversity and abundance of existing benthic organisms. Estuaries 18:91–107. Tolley, S.G., and Volety, A.K. 2005. The role of oysters in habitat use of oyster reefs by resident fishes and decapod crustaceans. Journal of Shellfish Research 24:1007–1012. Townsend, C.H. 1896. The transplanting of eastern oysters to Willapa Bay, Washington with notes on the native oyster industry. U.S. Commisioner of Fisheries, 193–202pp. Trianni, M.S. 1995. The influence of commercial oyster culture activities on the benthic infauna of Arcata Bay. M.S. thesis, Humboldt State University, Arcata, CA. Triplet, P., Stillman, R.A., and Goss-Custard, J.D. 1999. Prey abundance and the strength of interference in a foraging shorebird. The Journal of Animal Ecology 68:254–265. Tsai, C., Trimble, A.C., and Ruesink, J.L. 2010. Interactions between two introduced species:

294

Shellfish Aquaculture and the Environment

Zostera japonica (dwarf eelgrass) facilitates itself and reduces condition of Ruditapes phillipinarum (Manila clam) on intertidal flats. Marine Biology 157:1929–1936. Tyler, R.M. 2007. Effects of coverage by benthic seaweed mats on (northern quahog = hard clam) Mercenaria mercenaria in a eutrophic estuary. Journal of Shellfish Research 26:1021–1028. Valentine, J.F., and Heck, K.L., Jr. 1993. Mussels in seagrass meadows: their influence on macroinvertebrate abundance and secondary production in the Northern Gulf of Mexico. Marine Ecology Progress Series 96:63–74. Van Dolah, R.F., Holland, A.F., Coen, L.D., Ringwood, A.H., Levisen, M.V., Maier, P.P., Scott, G.I., Leight, A.K., Bobo, Y., and Richardson, D. 1999. Biological resources, report on the status of Broad Creek/Okatee River Systems. DHEC-MRRI-NOAA-Charleston. Volety, A.K., Savarese, M., Tolley, S.G., Arnold, W.S., Sime, P., Goodman, P., Chamberlain, R.H., and Doering, P.H. 2009. Eastern oysters (Crassostrea virginica) as an indicator for restoration of Everglades ecosystems. Ecological Indicators 9(Suppl. 1):S120–S136. Waldbusser, G.G., Voigt, E.P., Bergschneider, H., Green, M.A., and Newell, R.I.E. 2011. Biocalcification in the Eastern oyster (Crassostrea virginica) in relation to long-term trends in Chesapeake Bay pH. Estuaries and Coasts 34:221–231. Wall, C.C., Peterson, B.J., and Gobler, C.J. 2008. Facilitation of seagrass Zostera marina productivity by suspension-feeding bivalves. Marine Ecology Progress Series 357:165–174. Ward, D.H., Morton, A., Tibbitts, T.L., Douglas, D.C., and Carrera-Gonzalez, E. 2003. Longterm change in eelgrass distribution at Bahia San Quintin, Baja California, Mexico, using satellite imagery. Estuaries 26:1529–1539. Warnock, N., Haig, S.M., and Oring, L.W. 1998. Monitoring species richness and abundance of shorebirds in the Western Great Basin. The Condor 100:589–600. Watson, D., Shumway, S.E., and Whitlatch, R.B. 2009. Biofouling and the shellfish industry. In: Shumway, S.E., and Rodrick, G.E. (eds.), Shellfish Quality and Safety. Woodhead Publishing, Cambridge, England, pp. 317–336.

Waycott, M., Duarte, C.M., Carruthers, T.J.B., Orth, R.J., Dennison, W.C., Olyarnik, S., Calladine, A., Fourqurean, J.W., Heck, K.L., Jr., Hughes, A.R., Kendrick, G.A., Kenworthy, J.W., Short, F.T., and Williams, S.L. 2009. Accelerating loss of seagrasses across the globe threatens coastal ecosystems. PNAS 106:12377–12381. Wei, Z., Shi, H., Chen, S., and Zhu, M. 2009. Benefit and cost analysis of mariculture based on ecosystem services. Ecological Economics 68:1626–1632. Weise, A.M., Cromey, C.J., Callier, M.D., Archambault, P., Chamberlain, J., and McKindsey, C.W. 2009. Shell fish-DEPOMOD: modelling the biodeposition from suspended shellfish aquaculture and assessing benthic effects. Aquaculture 288:239–253. Wells, H.W. 1961. The fauna of oyster beds, with special reference to the salinity factor. Ecological Monographs 31:266–329. Weschler, J.F. 2004. Assessing the relationship between the ichthyofauna and oyster mariculture in a shallow embayment, Drakes Estero, Point Reyes National Seashore. M.S. thesis, University of California, Davis. Weston, D.P. 1990. Quantitative examination of macrobenthic community changes along an organic enrichment gradient. Marine Ecology Progress Series 61:233–244. White, J., Ruesink, J.L., and Trimble, A.C. 2009. The nearly forgotten oyster: Ostrea lurida Carpenter 1864 (Olympia oyster) history and management in Washington state. Journal of Shellfish Research 28:43–49. Whiteley, J., and Bendell-Young, L. 2007. Ecological implications of intertidal mariculture: observed differences in bivalve community structure between farm and reference sites. The Journal of Applied Ecology 44:495–505. Wildish, D.J., and Kristmanson, D.D. 1997. Benthic Suspension Feeders and Flow. Cambridge University Press, New York. Williams, S.W., and Heck, K.L., Jr. 2001. Seagrass communities. In: Bertness, M., Gaines, S., and Hay, M. (eds.), Marine Community Ecology. Sinauer Press, Sunderland, MA, pp. 317–337. Wisehart, L.M., Dumbauld, B.R., Ruesink, J.L., and Hacker, S.D. 2007. Importance of eelgrass early life history stages in response to oyster

Impacts of native and cultured bivalves

aquaculture disturbance. Marine Ecology Progress Series 344:71–80. Wrange, A.-L.J.V., Harkestad, L.S., Strand, Ø., Lindegarth, S., Christensen, H.T., Dolmer, P., Kristensen, P.S., and Mortensen, S. 2010. Massive settlements of the Pacific oyster, Crassostrea gigas, in Scandinavia. Biology Invasions 12:1145–1152. Yasue, M. 2005. The effects of human presence, flock size and prey density on shorebird foraging rates. Journal of Ethology 23:199–204. Yuan, X., Zhang, M., Liang, Y., Liu, D., and Guan, D. 2010. Self-pollutant loading from a suspen-

295

sion aquaculture system of Japanese scallop (Patinopecten yessoensis) in the Changhai sea area, Northern Yellow Sea of China. Aquaculture 304:79–87. Yund, P.O., Gaines, S.D., and Bertness, M.D. 1991. Cylindrical tube traps for larval sampling. Limnology and Oceanography 36:1167– 1177. Zimmerman, R., Minello, T., Baumer, T., and Castiglione, M. 1989. Oyster reef as habitat for estuarine macrofauna. NOAA Tech. Mem. NMFS-SEFC-249, 16pp.

Chapter 10

Bivalves as bioturbators and bioirrigators Joanna Norkko and Sandra E. Shumway

Bivalves are key species in soft-sediment habitats Both natural and cultured populations of bivalve molluscs provide a range of important ecosystem functions and services (see Chapters 1, 4, 8, and 9 in this book), some of which stem from the activities of different bivalve species living buried in the sediments. Bioturbation and bioirrigation are the mixing and flushing of sediments that are the result of the burrowing, feeding, and other activities of the animals in the sediment. Thus, bioturbation and bioirrigation are integral to a healthy soft-sediment ecosystem and, in general, infaunal bivalves such as clams have a positive, desirable influence on the sand or mud in

which they live, just as the earthworms in a vegetable patch or garden compost. Soft sediments are the most widespread habitats in the sea, extending from coastal estuaries and sandy shores, to continental shelves and the open ocean abyssal plains. Bivalve molluscs are often dominant in terms of biomass and/or abundance in estuarine and coastal soft-sediment habitats. Due to their habitat engineering and their influence on benthic-pelagic coupling and nutrient cycling, bivalve molluscs are key species in many habitats (Chapters 4 and 9 in this book; Jones et al. 1994; Dame et al. 2002). Consequently, changes in the condition, abundance, or distribution of different bivalve species may have cascading effects on both benthic and pelagic

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 297

298

Shellfish Aquaculture and the Environment

ecosystems (Vaughn and Hakenkamp 2001; Dahlhoff et al. 2002; Newell 2004; Norkko et al. 2006). In addition, bivalves are relatively long-lived and are therefore often used as indicator species when monitoring environmental change. Bivalves are linked to their environment through a multitude of processes that operate over different spatial and temporal scales. Large-scale processes, such as oceanographic conditions, and hydrodynamic and nutrient regimes, define the more general settings and the environmental framework that defines, for example, species distribution ranges and the level of primary production. Small-scale processes, such as individual physiology or behavior, are easier to study and amenable to experimental manipulation, but the extrapolation and scaling up of the results to the scale of an estuary or ecosystem are much more difficult. Therefore, generalizations based on small-scale studies are often not accurate at the larger scales of relevance to environmental problems and management practices. The way a pattern or a process is perceived, and therefore its accuracy, depends to a large extent on the scale at which the phenomenon is studied (Hewitt et al. 2007). This highlights the importance of explicitly incorporating scale in all environmental issues, for example, in the way the role of bivalves is assessed. This includes considering which processes operate at which scales, how they are linked, and which ones are most relevant for a particular issue, such as the most efficient and sustainable siting of aquaculture installations in different types of estuaries (Dumbauld et al. 2009). This chapter focuses on the mechanisms, dynamics, and effects of the bivalve bioturbation and bioirrigation processes and how these influence the health and stability of marine ecosystems. Particular attention is given to aspects of bioturbation/irrigation and other related sediment processes that are relevant for sustainable management of bivalve aquaculture operations.

What are bioturbation and bioirrigation? Soft-sediment habitats are greatly influenced by the activities of different taxa living within the sediment, including bivalves, crustaceans, and polychaete worms. The combination of their movements, feeding, ventilation, burrowing, tube construction, and biodeposit production results in structuring, mixing, and flushing of the sediment (Fig. 10.1, Table 10.1). The term “bioturbation” refers to the reworking of aquatic sediments by the organisms in the sediments and, in its broadest sense, includes the structuring activities of both burrowing animals and rooting plants, as well as microbes. In addition, animals living on the sediment surface (epifauna) and animals just visiting the sediment in search of food, such as rays and dugongs, may have bioturbating effects. The corresponding processes also occur in terrestrial soils, where earthworms are the most commonly used example. “Bioirrigation,” on the other hand, refers to the enhanced transport of solutes between the sediment and the overlaying water. This is the flushing of burrows that stems from the suspension feeding of the animals and their ventilation activities to facilitate transport of oxygen and excretory products. Bivalves are major players in the modification of sediments, and the effects we observe are a combination of both bioturbation and bioirrigation. In the following sections, “bioturbation” will be used to describe both phenomena, that is, to include both particle and solute transport. Bioturbation fundamentally affects the physical and chemical characteristics of softsediment habitats (Rhoads 1974; Aller 1988; Kristensen 1988; Levinton 1995; Aller 2001; Reise 2002; Meysman et al. 2006b) and can transform a homogenous sandy or muddy bottom into a heterogeneous landscape with physical structures such as burrows, pits, and mounds. Also, the mere physical presence of, for example, shells of both live and dead

Suspension and/or deposit feeding consumption of O2

Excretion of nutrients and CO2 biodeposition of feces and pseudofeces

Sediment mixing, flushing, and oxygenation

Nutrients

Water column

Added structure

Oxidized sediment RPD layer

Reduced sediment

Figure 10.1 Schematic diagram of the processes around a bivalve buried in the sediment.

Table 10.1 Combined effects of bivalve physical structure, bioturbation, bioirrigation, and feeding activities in natural bivalve populations in soft-sediment habitats. In addition, the relevance of these processes to aquaculture is summarized. Process

Effects on local environment

Relevance to aquaculture

Physical structure

Added heterogeneity; burrows, pits, mounds, shell material

Promotes overall higher diversity

Racks, bags, and longlines also provide structure; at high stocking densities, the habitat is homogenized and the diversity reduced

Oxygenation of the sediment; oxygen penetrates deeper into the sediment

Only applies if bivalves are able to bury into the sediment

Sediment mixing and sorting affects grain size

Racks, bags, and longlines may increase siltation

Individual bivalves provide substrate and habitat

Bioturbation; sediment reworking

Sediment destabilization, resuspension, and erosion Prevents siltation Bioirrigation; ventilation/feeding

Flushing of sediments with overlaying water

Biodeposition of feces and pseudofeces on the sediment surface

Benthic-pelagic coupling

Transport of oxygen into the sediment and excretory products out of the sediment

Increased organic content of the sediment; may provide food for other benthic species

Only applies if bivalves are able to bury into the sediment

Potential negative effects are highly dependent on flow, depth, and stocking density

-Higher microbial activity and oxygen demand in the sediment; potential for hypoxia Altered nutrient fluxes at the sediment-water interface

299

300

Shellfish Aquaculture and the Environment

bivalves in the sediment creates added structure and heterogeneity. Reworking of the sediment affects the sorting of the sediment, its grain size distribution, porosity, and the vertical profiles of both solids and solutes. It changes the rates and pathways of reaction processes within the sediment, and between the sediment and the overlaying water (Table 10.1). This subsequently affects biodiversity and ecosystem function in soft-sediment habitats. In general, bioturbation increases all fluxes at the sediment–water interface, including both particulates and solutes. Efficient fluxes are a prerequisite for any wellfunctioning ecosystem and are at the base of all ecosystem services society needs from the environment. Understanding these fluxes will help manage our coastal ecosystems efficiently and sustainably. Bioturbation increases the depth of oxygen penetration into the sediment from the overlaying water. The presence or absence of oxygen governs a multitude of both chemical and biological processes in the sediment, and therefore bioturbation affects the composition of both the invertebrate and microbial communities in the sediment (Glud 2008). The burrowing, mixing, and tube construction of different invertebrate species substantially increases the area of the sediment–water interface, by extending this layer deeper into the sediment as an irregular surface. In combination with active suspension feeding and burrow ventilation, this allows oxygen-rich water to penetrate deeper into the sediment, thereby oxygenating otherwise reduced and anoxic sediments. Bioturbation thus affects the area available for aerobic microbial activity, which affects the functioning of the whole system, since the sediment microbial community performs essential functions such as decomposition of deposited organic material and mineralization of nutrients. The depth of oxygen penetration may be seen as an upper, lighter oxidized sediment layer, with a sometimes relatively sharp transi-

tion to the underlying darker reduced sediment layer (Fig. 10.1). Depending on sediment type (oxygen penetrates deeper into porous sandy sediments compared with cohesive mud; Volkenborn et al. 2007) and the species composition and abundance of the benthic community, this lighter layer may range from just a millimeter to several centimeters thick. The border between oxidative and reducing reactions is called the redox potential discontinuity layer (RPD). Bioturbators may substantially extend the depth of this layer, and the depth of the RPD layer is often used as an indicator of environmental status, with deeper oxidized sediment layers indicating the presence of more animals and thus better conditions. This can roughly be measured in sediment cores taken with a see-through corer or quantified, for example, using sediment profile imagery (SPI), where vertical images are taken into the sediment with a prism mounted on a digital camera and subsequently analyzed using image analysis software (Grizzle and Penniman 1991). On a finer scale, the depth profiles of pH, oxygen, sulfides, and other compounds can be measured with millimeter accuracy using microsensors penetrating the sediment (Glud 2008). Present-day bioturbation generally does not affect more than about the top 30 cm of the sediment, that is, the sediment layers where most of the infaunal species live (although there are exceptions, such as deep-burrowing geoducks). Where sediment accumulates over time, however, some traces of bioturbation activities are preserved and can be seen in thick sediment layers that are thousands of years old (Erwin 2008). Similarly, absence of fauna is also recorded in the sedimentary record in the form of layers upon layers of undisturbed, laminated sediments, which are formed during periods of either natural or eutrophicationinduced anoxic conditions (Schaffner et al. 1992; Karlson et al. 2002; Fig. 10.2). The mechanisms of bioturbation and bioirrigation have been studied through a number

Bivalves as bioturbators and bioirrigators

(A)

301

(B)

Figure 10.2 Photos of laminated sediments, indicating absence of bioturbation (A), and bioturbated sediments, with an oxidized lighter layer several centimeters thick (B). (Courtesy of H.C. Nillson.)

of different methods since a combination of methods is required to clarify the small-scale mechanisms in the bioturbation processes (Quintana et al. 2007). Each method has its limitations and the choice of method or combination of methods depends on the question being posed and the timescale of interest; however, the necessity of always also conducting field experiments and field measurements needs to be emphasized, as they often yield contradicting results compared with laboratory and modeling studies (Table 10.2). All benthic species are different in terms of their bioturbation behavior and thus the net effects of bioturbation depend on the suite of species present and their abundances. This necessitates consideration of species-specific

patterns, processes, and types and magnitudes of effects. Bioturbating species have been divided into a range of different functional groups, but five broadly defined groups commonly used are biodiffusers, upward conveyors, downward conveyors, gallery diffusers, and regenerators (François et al. 2002; Gerino et al. 2003). Biodiffusers randomly move particles over short distances. This frequent, small-scale particle movement results in diffusive mixing and transport of sediment, and most bivalve species belong to this group. On the other hand, upward and downward conveyors, for example, many bioturbating polychaetes, transport particles through their guts, resulting in a nonlocal transport from deeper sediment layers up to the sediment

302

Shellfish Aquaculture and the Environment

Table 10.2 Examples of methodological approaches used for studying bioturbation and bioirrigation (not restricted to only bivalves), the spatial scale these methods apply to (individual organism, laboratory studies, field experiments, field surveys and monitoring, ecosystem level effects), and selected references to studies utilizing these methods.

Method

Bioturbation-related effect/ process studied

Spatial scale

Example reference

Rhoads and Young (1971)

Visual assessment, photographs, video

Tracks, pits, burrows, mounds

Laboratory, field experiments, surveys

Visual assessment of RPD layer depth

Net effects of bioturbation; oxygen penetration in the sediment

Laboratory, field experiments, surveys

Sediment analysis (including slicing)

Depth distribution of sediment grain size, water content, pelletization, organic content

Laboratory, field experiments, surveys

Maire et al. (2008)

Inert particle addition (e.g., luminophores, glass beads) or natural tracers (radionuclides)

Particle mixing, sediment reworking

Laboratory, field experiments

Gerino et al. (1998); Montserrat et al. (2009)

Flume studies

Sediment resuspension and erosion

Laboratory

Willows et al. (1998)

Inert solute addition (e.g., bromide)

Pore water irrigation

Laboratory, field experiments

Martin and Banta (1992)

Microelectrodes, planar optodes

Depth profiles of nutrients, O2, S, H2S

Very fine-scaled; laboratory, field experiments

Glud (2008)

Sediment incubations

Fluxes of nutrients, solutes and gasses across the water–sediment interface

Laboratory, field experiments

Asmus et al. (1998); Michaud et al. (2006)

Sediment profile imaging

Two-dimensional structures and oxygen penetration in the sediment

Surveys

Grizzle and Penniman (1991); Solan et al. (2004)

Computer-aided tomography

Three-dimensional structures in the sediment

Individual cores (slow/expensive)

Perez et al. (1999); Bouchet et al. (2009)

Models

Nutrient profiles and fluxes, sediment reworking and transport

Individual bivalves, cores, ecosystems

Willows et al. (1998); Meysman et al. (2006a)

See Maire et al. (2008) for a review of methods for quantifying sediment reworking rates.

surface or from the sediment surface down, respectively. Gallery diffusers create tube systems and mix sediment in the process. Regenerators also dig tube systems, with additional release of large amounts of sediment into the overlaying water during digging and transport of sediment from the surface to the

bottom of the burrow after it has been deserted. Common to all of these groups is that they facilitate fluxes of particulate matter and solutes between the sediment and the overlaying water, but models of bioturbation need to consider the different types of bioturbation (Michaud et al. 2006).

Bivalves as bioturbators and bioirrigators

How do healthy soft-sediment bivalve populations affect their surroundings? Bivalves have profound influences on surrounding macrobenthic communities. In comparison with many other bioturbating species such as highly active crabs, shrimp, and polychaete worms, bivalves may seem rather stationary and, at first glance, not seem to be doing much, at least not in terms of bioturbation. Due to their frequently dominating abundances and biomasses, however, and active filtration of large volumes of water, the net impact of their bioturbation and bioirrigation activities on the system are considerable (Jackson et al. 2001; Vaughn and Hakenkamp 2001; Jaramillo et al. 2007; Petersen et al. 2008). Although most soft-sediment bivalves can be classified as biodiffusers in terms of their bioturbation mode, different species have different burrowing depths (Table 10.3), which in turn influences their impact on sediment mixing, irrigation, and nutrient fluxes, and their effects on the surrounding communities. Moreover, the juveniles of most soft-sediment bivalves live at or close to the sediment surface, and move successively deeper into the sediment as they grow. Burrowing depth depends on siphon length (Zwarts and Wanink 1989); bivalves with short siphons live just underneath the sediment surfaces (e.g., hard clams or northern quahog, Mercenaria mercenaria), while species with longer siphons may be buried up to 40 cm down in the sediment (e.g., soft-shelled clams, Mya arenaria) or even 80 cm (geoducks, Panopea abrupta); however, shallow-burying bivalves (such as clams) tend to move more, thereby bioturbating the uppermost sediment layers more than the deep burrowers. Sediment reworking rates may further differ with season and food availability, even within a bivalve species (Stead and Thompson 2006; Maire et al. 2007). Thus, the patterns of bivalve bioturbation and facilitation effects on surrounding communities are temporally

303

variable and dependent on species, and the density and size structure of the population. The basic increased fluxes of particulates and solutes that bioturbation induces are rather straightforward, but they have a range of complex and context-dependent effects. In the following sections these different mechanisms by which soft-sediment bivalves influence their surroundings are described. In particular the aim is to illustrate the numerous multiway interactions between bioturbation, benthic biodiversity, and ecosystem function.

Provide structure, heterogeneity in sediments, stabilize/destabilize the sediment Bivalve beds physically influence the diversity and functioning of surrounding communities in a number of ways (see Chapter 9 in this book for an in-depth discussion on reefforming bivalves as ecosystem engineers). Although infaunal species such as clams do not provide the same type of three-dimensional habitat as do mussels and oysters on hard substrates, the presence of bivalves nevertheless provides increased structural heterogeneity within the sediments, with potential for a larger number of different favorable microhabitats in the interstitial spaces and thus increased diversity (Gutiérrez et al. 2003). Even the very small-scale movements around individual bioturbating bivalves are important and add heterogeneity to the soft-sediment system. Further, infaunal species have either siphons, feeding shafts, or parts of their shells reaching the sediment surface, which create additional bed roughness and heterogeneity. Bivalves protruding above the sediment surface will also affect flow in the benthic boundary layer, both due to their physical structure and to their feeding currents (Green et al. 1998). Shells provide hard substrate for recruitment in otherwise soft, unconsolidated sediments, which increases the diversity of epifaunal

304

Shellfish Aquaculture and the Environment

Table 10.3 Adult burrowing depth in natural populations of selected bivalve species and the type of shellfish aquaculture they are used in. In addition, their potential to actively bioturbate sediments in different types of culture is indicated with (+) or (−).

Species

Burrowing depth

Type of culture

Potential for bioturbation in culture

Mercenaria mercenaria (northern quahog, hard clam)

1–3 cm (up to 20 cm)

Bottom (“free”)

+

On-bottom bags, cover nets

+

Mya arenaria (soft-shelled clam)

10–20 cm (up to 40 cm)

Bottom

+

Panopea abrupta (geoduck clam)

60–80 cm

Bottom

+

Crassostrea gigas (Pacific oyster)

0

Bottom

−

Longlines, off-bottom bags and racks

−

Crassostrea virginica (eastern oyster)

0

Bottom

−

Suspended

−

Mytilus edulis (blue mussel)

0

Bottom

−

Suspended

−

Bottom

−

Suspended

−

Bottom

+

Mytilus galloprovincialis (Mediterranean mussel)

0

Ruditapes philippinarum (Manila clam)

0–5 cm

For many species, the burrowing depth differs between summer and winter (Zwarts and Wanink 1989). In cold climates, bivalves move deeper into the sediment in winter to avoid freezing, whereas in hot climates, bivalves may move deeper in summer to avoid overheating.

species in the system (sponges, bryozoans, tunicates, algae, etc.) (Gutiérrez et al. 2003). The bivalves affect the sedimentary habitat even after they die, as the shell debris continues to provide structure in the habitat (Hewitt et al. 2005; Erwin 2008) with positive influences on biodiversity. The effect of bivalves on the physical heterogeneity of the habitat is naturally dependent on bivalve density and aggregation. For example, at very high bivalve densities, competition for space becomes

important and diversity no longer increases (Whitlatch et al. 1997). Bivalve bioturbation has the potential to modify significantly the coastal sea and landscape. The prevalence of different types of soft-sediment habitats in coastal areas depends on the stability of the sediments, that is, the balance between rates of sediment deposition and sediment erosion. These processes are ultimately governed by hydrodynamic forcing (currents, waves) transporting the sediment

Bivalves as bioturbators and bioirrigators

and affecting its sorting and grain size, but biota significantly influences the formation and persistence of different habitat types by affecting the stability of the sediment (Widdows et al. 2004; Le Hir et al. 2007). In aquatic ecosystems some species act as sediment stabilizers (enhance sediment deposition, inhibit erosion), while other species act as sediment destabilizers (enhance erosion and sediment transport) (Widdows et al. 2000; Bouma et al. 2009). By providing physical protective structures, for example, mussel beds, oyster reefs, seagrass meadows, salt marsh plants, and tube-building polychaetes stabilize the sediment, modify the local hydrodynamics, and enhance sediment deposition. Also, microphytobenthos (i.e., microalgae/microorganisms living on the sediment surface) enhance sediment cohesiveness through the extracellular polymeric substances they secrete, which form a protective biofilm on the sediment surface. Other species act as sediment destabilizers by disrupting the cohesiveness of the sediment, which decreases the sediment mud content and increases erosion rates. The sediment is destabilized, for example, by clams, crustaceans, and lugworms that bioturbate or mix the sediment and facilitate resuspension (Widdows and Brinsley 2002). The sediment is also destabilized by grazers removing the protective microphytobenthic biofilm. Further, increased bed roughness and scouring around patches of animals or plants can destabilize sediments and enhance erosion locally. Bioturbating bivalves affect resuspension rates, both directly and indirectly, actively and passively (Graf and Rosenberg 1997). Experiments in nonaquaculture settings have demonstrated a positive relationship between bivalve density and sediment erosion, which is likely a combination of the bivalve-induced bioturbation and increased bed roughness, both of which promote sediment erosion. These patterns have been found for a number of species, including cockles (Cerastoderma edule), Manila clams (Ruditapes

305

philippinarum), and Baltic clams (Macoma balthica) (Widdows and Brinsley 2002; Sgro et al. 2005; Ciutat et al. 2007). Although natural clam populations may act to destabilize sediments, enhance erosion rates, and prevent siltation, the nets used to keep clams in place in aquaculture installations increase the rates of sedimentation and increase the sediment silt content (Kaiser et al. 1996; Chapter 9 in this book). Problems with sedimentation can, however, be avoided with development of better culture methods, such as the bags used for clam culture in Cedar Key, Florida (http:// shellfish.ifas.ufl.edu). These bags also provide habitat for other species and enhance diversity, provided the siting and layout of the bags is appropriate and coordinated. To predict sediment transport patterns, the critical thresholds for sediment erosion need to be identified (Le Hir et al. 2007). Faunal effects on sediment erosion, transport, and deposition have mainly been studied in intertidal areas, and our knowledge about these effects in deeper areas is much more limited. The interactions between sediment stabilizing and destabilizing processes are complex, dynamic, and continually at work. While bioturbating bivalves may destabilize the sediment, their bioturbation may simultaneously enhance nutrient fluxes from the sediment, which promotes the growth of sediment stabilizing microphytobenthos (Sandwell et al. 2009). Within a bivalve bed the continuous reworking of the sediment, which is characteristic for biodiffusers such as most infaunal bivalves, homogenizes the sediment and prevents formation of permanent structures and may prevent establishment of, for example, invasive salt marsh species. Conversely, spreading salt marshes reduce the habitat available for shellfish (Neira et al. 2006). Sediment stabilizers and destabilizers have spatially and temporally variable effects on the community, usually favoring their own type, which facilitates the persistence of patches (Volkenborn

306

Shellfish Aquaculture and the Environment

et al. 2009). These processes thus act both to modify and to maintain specific habitats. The antagonistic interactions between bioturbators and habitat-forming ecosystem engineers such as seagrasses, salt marshes, or oyster reefs create patchiness and habitat diversity, thereby favoring an overall higher biodiversity at the estuary scale (Jones et al. 1994; Bouma et al. 2009). To minimize disruption of sediment transport patterns in an estuary and thus maintain the current extent of the existing habitat types in the estuary, the complex interactions between sediment stabilizers and destabilizers need to be considered. This has implications for where bivalve aquaculture plots should be located in an estuary and what the spatial scale, type of culture, and layout of the installations should be, for example, continuous cover or several separate sections (Dumbauld et al. 2009).

Biodeposit production and degradation: effects on sediment biogeochemistry Bivalves affect fluxes of particles to and from the seafloor, both by directly intercepting and ejecting particles as part of their feeding, and by physically altering the hydrodynamic conditions that govern sedimentation and erosion rates (Graf and Rosenberg 1997; Widdows et al. 1998) (Chapter 9 in this book). Through their feeding, bivalves produce feces and pseudofeces, collectively termed biodeposits when they reach the seafloor. Chapter 4 (in this book) on “Bivalves as biofilters” provides details on regulation of feeding and production of feces and pseudofeces. In this section the focus is on biodeposits after they have been deposited on the seafloor and their context-dependent effects on benthic nutrient fluxes and surrounding benthic habitats. In addition to the ongrowing phase considered here, bivalve aquaculture may

impact benthic ecosystems also during seed collection and harvesting. These other aspects, however, concern management practices and are dealt with in Chapters 3 and 11 in this book. Bivalves have a dominating organizational role in ecosystems as they shunt a lot of the production from the pelagic to the benthic system in the form of biodeposits (Dame 1993; Newell 2004). Both feces and pseudofeces are ejected as mucus-bound aggregates, which sink faster than nonaggregated particles, thus aiding the vertical downward flux and increasing the deposition locally above background levels of sedimentation (Chapter 4 in this book; Ward and Shumway 2004). Rates of biodeposit production, and the ratio of feces to pseudofeces depends on the concentrations of food particles (phytoplankton, some detritus) and suspended sediment (inorganic matter), all of which vary seasonally, and also on the bivalve species, as different species have different clearance rates and abilities to select particles preferentially (Kautsky and Evans 1987; Jaramillo et al. 1992; Newell 2004; Ward and Shumway 2004; Giles et al. 2006). High loads of suspended sediments in the water (e.g., in sheltered, turbid estuaries) lead to larger amounts of pseudofeces being produced, which also increases the downward flux of particulate matter to the sediment (Iglesias et al. 1996; Norkko et al. 2001).

Effects of water flow Patterns of biodeposit accumulation or erosion in bivalve beds or underneath suspended bivalve cultures are mainly driven by the water flow at the site, with higher current velocities promoting erosion and efficient dispersal of the biodeposits (Chamberlain et al. 2001; Hartstein and Rowden 2004; Weise et al. 2009; Chapters 4, 6, and 9 in this book). With greater tidal currents and wave action, resuspension of biodeposits is more likely in intertidal or shallow bivalve beds, compared

Bivalves as bioturbators and bioirrigators

with bivalve beds or suspended aquaculture systems in deeper waters. In enclosed embayments, however, the tidal flushing may not be sufficient and biodeposits accumulate underneath shellfish farms (Cranford et al. 2009; Chapters 4 and 6 in this book) (Fig. 10.3). In deeper waters the biodeposits from suspended culture may be dispersed over a larger area before reaching the seafloor and thus have no detectable impact on the benthic habitat. The rates of biodeposit accumulation or erosion are further affected by the density of bioturbating infaunal bivalves and other fauna as they enhance resuspension and dispersal of biodeposits by acting as sediment destabilizers (see previous section).

Organic enrichment Bivalve biodeposition represents an organic enrichment of the sediment and this additional organic matter provides an important food

307

subsidy to benthic systems (Wotton and Malmqvist 2001). As the biodeposits may be rich in carbon and nitrogen, surrounding deposit-feeding macro-, meio-, and microfauna can benefit from increased food quality (Kautsky and Evans 1987; Norkko et al. 2001; Giles et al. 2006). The effects of increasing biodeposition follow the general hump-shaped benthic responses to organic enrichment, with increasing benthic diversity, abundance, and biomass at early stages of enrichment, and increasing sediment oxygen demand and anoxic conditions with benthic community disruption after severe enrichment (Pearson and Rosenberg 1978) (Fig. 10.3). The effects are directly related to the amounts of organic matter deposited and remaining on the seafloor, and as noted earlier, these amounts are dependent on bivalve species and density, the quality and quantity of their food supply, and the hydrodynamic setting, water flow, and depth at the site (Fig. 10.3).

Figure 10.3 Schematic diagram of the context and density dependence of the effects of bivalve populations and their biodeposits on surrounding benthic communities. Panels A–C represent increasing densities of bivalves in either natural or bottom-cultured populations, while panel D represents a high-density suspended culture, where the bivalves are not able to bioturbate the sediments. Computational model by Alf Norkko.

308

Shellfish Aquaculture and the Environment

Effects of biodeposition on sediment biogeochemistry Biodeposits affect the biogeochemistry in the sediment and at the sediment–water interface. Although bivalves also directly excrete inorganic nutrients, the remineralization processes of the biodeposits in the sediment are usually more important for nutrient regeneration and fluxes (Prins et al. 1998; Newell 2004). Accumulation of biodeposits increases sediment organic content, promotes high microbial activity and increases the sediment oxygen demand (Rodhouse and Roden 1987; Wotton and Malmqvist 2001; Nizzoli et al. 2006; Hargrave et al. 2008; Cranford et al. 2009), but the effects of biodeposition are site specific (Chamberlain et al. 2001; Hartstein and Rowden 2004). The decomposition of organic material consumes oxygen and increased sediment oxygen demand is commonly recorded, for example, under mussel culture longlines. High rates of biodeposition may adversely affect the sediment microbial community by shifting it from an aerobic to an anaerobic metabolism. The changing oxygen conditions subsequently affect nutrient fluxes in and out of the sediment. Under anoxic conditions phosphorus is released from iron compounds in the sediment. Biodeposition also affects the cycling, retention, and loss of nitrogen from the sediment (Cranford et al. 2007). Coupled nitrificationdenitrification removes nitrogen from the system in the form of N2 gas, but requires adjacent oxygenated sediments with nitrifying microbes and anoxic sediments with denitrifying microbes (Newell et al. 2002). Thus, the process is inhibited if the overlying water turns anoxic. Sediments underlying intensive shellfish aquaculture release ammonium since the coupled nitrification-denitrification has been disrupted (Hatcher et al. 1994; Newell et al. 2002). If the sediments turn anoxic, concentrations of H2S may build up to toxic levels. White sulfur bacterial mats (Beggiatoa spp.)

may also develop in areas of high biodeposition and high sediment oxygen demand. Since Beggiatoa requires both oxygen and H2S for metabolism, its presence indicates that the reducing conditions in the sediment have reached the sediment surface.

Effects of bioturbation on benthic nutrient fluxes and the processing of biodeposits The biogeochemical effects of biodeposition from shellfish aquaculture are strongly dependent on the quantity and quality of the material deposited. Efficient bivalve bioturbation does, however, ameliorate the potentially negative effects of excessive biodeposition since particle and solute transport by bioturbating fauna significantly influences rates and pathways of organic matter mineralization, particularly by increasing the oxygen penetration depth in the sediments. For example, Mya arenaria burrows are sites of enhanced microbial activity and high rates of sulphate reduction (Hansen et al. 1996) and burrowing Mercenaria mercenaria increase benthic fluxes of oxygen and nutrients (Doering et al. 1987). Bioturbation enhances coupled nitrificationdenitrification, and under fully oxygenated conditions any remaining phosphorus in the biodeposits is also permanently buried with the accumulating sediments. Infaunal bivalves both produce biodeposits and facilitate the remineralization of them in the sediment (Figs. 10.1 and 10.3). Thus, the combination of biodeposition and bioturbation by clams maintains a balanced benthic metabolism (Nizzoli et al. 2006), with the benthic nutrient fluxes being dependent on bivalve density (Sandwell et al. 2009). In natural infaunal bivalve beds and bottom cultures of clams and mussels, rates of biodeposition are not likely to reach high-enough levels to induce anoxic conditions and high levels of sulfide in sediments due to the negative feedback excessive biodeposition would have on

Bivalves as bioturbators and bioirrigators

bivalve densities. Nizzoli et al. (2006) compared mussel and clam farming and found that overall impacts of suspended mussel farming on oxygen and nutrient dynamics were greater than those of infaunal clam farming, and attributed this to the fact that infaunal, bioturbating bivalves stimulate transfer of both organic matter and oxygen in to the sediment, whereas suspended bivalves only enhance transfer of organic matter to the sediment via biodeposition. This also affected the pathways of nitrogen cycling. The feedback mechanisms between sediment organic content, oxygen conditions, and bioturbation are complex. For example, hard clams do not burrow as deep under hypoxic conditions, with a potential for cascading negative effects on sediment oxygenation (Weissberger et al. 2009). It is therefore important to limit rates of biodeposit accumulation to a level at which the bioturbators can still process effectively. Model calculations indicate that even though the harvest represents a net removal of phosphorus and nitrogen from the ecosystem, the deposition of feces and pseudofeces from a mussel farm increases the retention time of nutrients in some coastal areas (Cranford et al. 2007; Brigolin et al. 2009). With careful scaling and placement of farms in areas with sufficient flow, and by maintaining sufficient levels of bioturbation under the farms, mussel aquaculture can be used to remove nutrients from the system and thus combat eutrophication (Newell 2004; Lindahl et al. 2005; Gren et al. 2009; Chapter 8 in this book).

Effects of biodeposits on benthic community composition Shellfish aquaculture influences the surrounding benthic habitat through three main mechanisms: increased input of organic matter through biodeposition; increasing hypoxic and sulfidic sediments due to increasing oxygen demand from decomposition of biodeposits;

309

and added physical structure from either bivalve shells or farming installations. The effects of organic enrichment from biodeposition, however, have proven to be highly site specific and context dependent, and range from significant positive effects (Inglis and Gust 2003; D’Amours et al. 2008), to no or minimal effects (Hatcher et al. 1994; Grant et al. 1995; Kaiser et al. 1996; Chamberlain et al. 2001; Crawford et al. 2003), to significant negative effects with impoverished benthic communities under/near the farms (Smith and Shackley 2004; Hargrave et al. 2008; Cranford et al. 2009). Inside bivalve beds and under lines of suspended bivalves, the benthic infaunal species composition generally shifts toward species smaller, opportunistic detritivores, more tolerant to higher organic content, such as Oligochaeta and capitellid polychaetes (Beadman et al. 2004; Commito et al. 2005; Norling and Kautsky 2007; Ysebaert et al. 2009). Shells falling from mussel lines may also provide additional habitat and heterogeneity, with positive effects on the surrounding fauna (Grant et al. 1995). At very high rates of biodeposit accumulation, however, the sediment becomes anoxic and the macrofauna disappears. The net effects will depend both on the scale, stocking densities, and intensities of farming, as well as the hydrodynamic regime at the site (water depth, currents, sedimentation rates), which will interact to determine the amount of waste products accumulating on the seabed (Fig. 10.3). It has, however, repeatedly been reported that effects of, for example, mussel beds are relatively local and depend on bivalve density (Beadman et al. 2004; Hartstein and Rowden 2004; Ysebaert et al. 2009). Therefore, using a somewhat lower stocking density in combination with sufficient flow can reduce the potential negative effects on the benthic environment (Beadman et al. 2004). Since shellfish aquaculture is based on natural food sources, no additional nutrients or organic matter are added to the system. Therefore, the amount of biodeposition, and

310

Shellfish Aquaculture and the Environment

thus organic enrichment and potential subsequent negative effects, is significantly less under shellfish aquaculture installations compared with finfish aquaculture (Folke and Kautsky 1989; Naylor et al. 2000; Newell 2004). In finfish farming stocking densities are higher and the fish are fed artificial diets, resulting in a net addition of nutrients and organic matter to the site, which may result in waste accumulation and anoxic bottoms under the fish cages, disturbed benthic communities, and eutrophication of the surrounding ecosystem (Gowen and Bradbury 1987; Wu 1995; Kalantzi and Karakassis 2006). Due to this fundamental difference, it is important and essential to consider finfish and shellfish aquaculture as two separate industries.

Interactions between bivalve bioturbation and seagrasses In coastal areas, seagrass meadows are important structured habitats, providing nursery areas for fish and shellfish (Heck et al. 2003). Suspension-feeding bivalves reduce water turbidity, which enhances light penetration and thereby potentially promotes benthic primary producers, such as seagrasses and microphytobenthos. For example, successful recruitment of Mya arenaria was instrumental in turning a Danish lagoon from a turbid state to a clearwater state (Petersen et al. 2008). In some areas, improved water clarity resulting from large beds of cultured bivalves has even led to the reestablishment of seagrasses (Shumway and Kraeuter 2004). The effects are, however, species dependent and, for example, eastern oysters (Crassostrea virginica) have higher clearance rates than hard clams and therefore promote seagrass production more (Newell and Koch 2004). Sediment nutrient enrichment by biodeposits from natural bivalve populations can also elevate seagrass productivity (Peterson and Heck 2001a; Peterson and

Heck, 2001b). Bioturbating bivalves inside seagrass meadows also renew the water at the sediment surface and thereby reduce the risk of oxygen depletion. Nevertheless, particularly in eutrophic areas, high levels of biodeposition may lead to toxic sulfide concentrations in the sediment and increase the ammonium efflux from the sediment, which facilitates epiphyte growth, resulting in impaired growth of seagrass (Vinther and Holmer 2008; Vinther et al. 2008). Also, rack and stake oyster culture negatively influence seagrasses through physical disturbance and shading (Everett et al. 1995). If densities of aquaculture gear are sufficiently low, however, there are only minimal effects on seagrasses of, for example, oyster depuration gear (Vaudrey et al. 2009). Therefore, the multiple interactions between seagrass habitats and shellfish aquaculture need to be considered and managed at appropriate scales (Dumbauld et al. 2009).

Effects of bioturbation on contaminants and resting stages in the sediments Bivalve bioturbation also affects the mixing and depth distribution of contaminants and has the potential to either bury these elements deeper or to bring them back into circulation. For example, Ciutat et al. (2006) found a positive correlation between the density of cockles (Cerastoderma edule) and the resuspension of both sediment and polycyclic aromatic hydrocarbons (PAHs). Similarly, bioturbation by the bivalve Tellina deltoidalis in metal contaminated sediments caused metal release from the pore waters and higher concentrations of iron and manganese in overlying waters (Atkinson et al. 2007). On the other hand, bioturbation by Baltic clams increased the retention of cadmium and a hydrophobic organic pollutant in the sediment (Hedman et al. 2008). Thus, the fate of contaminant is dependent both on

Bivalves as bioturbators and bioirrigators

the type of compound and the rates of bioturbation in the habitat. Many planktonic species form dormant cysts or resting stages under adverse environmental conditions. The bioturbation-induced mixing processes apply to these cysts as well and the burial and reemergence of these is part of the natural cycle for many species. For example, Baltic clams extend the distribution of cladoceran resting eggs deeper into the sediment (Viitasalo 2007) and also differentially affect the emergence of several zooplankton species from the sediment (Viitasalo et al. 2007).

Summary Bioturbation and bioirrigation are very smallscale activities by individual bivalves, but the net effects of a whole bivalve population may have consequences for the long-term development of an entire coastal system, both in terms of nutrient dynamics and sediment stability. This highlights the importance of considering the appropriate spatial and temporal scales for both studying and managing any particular process, installation, or environmental effect, including shellfish aquaculture operations. Bivalve bioturbation facilitates transport of particulate and dissolved material between the sediment and the overlying water, oxygenates the sediment, and produces complex and favorable microhabitats, which facilitate benthic communities with higher overall diversity. It is commonly assumed that there is a general positive relationship between high diversity and well-functioning, healthy ecosystems. One way to assess ecosystem function is to estimate rates of nutrient fluxes in and out of the sediments (Lohrer et al. 2004). There are numerous studies on nutrient fluxes and bivalves, and we know that they are influenced, for example, by the interactions between bivalve density and species, sediment type,

311

organic content, hydrodynamic conditions, and environmental stress. Nevertheless, we are lacking in our understanding of the generality and ecosystem-scale consequences of these fluxes. Since the positive or negative effects of bivalve biodeposits are quality, quantity, and context dependent, and modified by bivalve bioturbation, studies of the effects of organic enrichment from shellfish aquaculture on sediment biogeochemistry, benthic communities, and seagrass beds have yielded site-specific and sometimes nonconclusive results. The associated complex biogeochemical processes are difficult to measure over larger scales in the field, but at the same time results gained from laboratory experiments are difficult to generalize to, for example, sites with different hydrodynamic regimes (Porter et al. 2004). Therefore, an integrated approach consisting of welldesigned laboratory experiments, field studies across different sites and along gradients within sites, and modeling is required to understand these processes and to apply them at the appropriate scale to sustainable management of shellfish aquaculture (Dumbauld et al. 2009). The benthic effects of bivalve aquaculture differ significantly between cultured species and also between different types of culture of the same species (Nizzoli et al. 2006; Ysebaert et al. 2009). Suspended culture generally only affects benthic environments through their biodeposition, while on-bottom culture affects benthic habitats through both biodeposition, altered physical structure at the sediment surface, and bioturbation of the sediments. Therefore, for example, clam farming results in a more balanced benthic metabolism, although it may be limited by available space. As the effects of biodeposition on the benthic environment are relatively local and dependent on the hydrodynamic regime, in order to minimize negative impacts on the benthic communities, farms should be sited in areas with relatively high current velocities and sufficient

312

Shellfish Aquaculture and the Environment

water exchange (Hartstein and Rowden 2004; Dumbauld et al. 2009) since in enclosed embayments the dispersive capacity may be surpassed (Cranford et al. 2009). Finally, homogenization of habitats removes the small-scale heterogeneity that sustains diversity, and loss of biodiversity will ultimately lower the ocean’s capacity to provide food for human consumption (Worm et al. 2006). Thus, shellfish aquaculture should be managed in the light of minimizing negative impacts on diversity.

Literature cited Aller, R.C. 1988. Benthic fauna and biogeochemical processes in marine sediments: the role of burrow structures. In: Blackburn, T.H., and Sørensen, J. (eds.), Nitrogen Cycling in Coastal Marine Environments. Wiley, New York, pp. 301–338. Aller, R.C. 2001. Transport and reactions in the bioirrigated zone. In: Boudreau, B., and Jørgensen, B.B. (eds.), The Benthic Boundary Layer: Transport Processes and Biogeochemistry. Oxford University Press, Oxford, pp. 269–301. Asmus, R.M., Jensen, M.H., Jensen, K.M., Kristensen, E., Asmus, H., and Wille, A. 1998. The role of water movement and spatial scaling for measurement of dissolved inorganic nitrogen fluxes in intertidal sediments. Estuarine, Coastal and Shelf Science 46:221–232. Atkinson, C.A., Jolley, D.F., and Simpson, S.L. 2007. Effect of overlying water pH, dissolved oxygen, salinity and sediment disturbances on metal release and sequestration from metal contaminated marine sediments. Chemosphere 69:1428–1437. Beadman, H.A., Kaiser, M.J., Galanidi, M., Shucksmith, R., and Willows, R.I. 2004. Changes in species richness with stocking density of marine bivalves. Journal of Applied Ecology 41:464–475. Bouchet, V.M.P., Sauriau, P.-G., Debenay, J.-P., Mermillod-Blondin, F., Schmidt, S., Amiard, J.C., and Dupas, B. 2009. Influence of the mode of macrofauna-mediated bioturbation on the vertical distribution of living benthic foraminifera: first insight from axial tomodensitometry.

Journal of Experimental Marine Biology and Ecology 371:20–33. Bouma, T.J., Olenin, S., Reise, K., and Ysebaert, T. 2009. Ecosystem engineering and biodiversity in coastal sediments: posing hypotheses. Helgoland Marine Research 63:95–106. Brigolin, D., Dal Maschio, G., Rampazzo, F., Giani, M., and Pastres, R. 2009. An individual-based population dynamic model for estimating biomass yield and nutrient fluxes through an off-shore mussel (Mytilus galloprovincialis) farm. Estuarine, Coastal and Shelf Science 82:365–376. Chamberlain, J., Fernandes, T.F., Read, P., Nickell, T.D., and Davies, I.M. 2001. Impacts of biodeposits from suspended mussel (Mytilus edulis L.) culture on the surrounding surficial sediments. ICES Journal of Marine Science 58:411–416. Ciutat, A., Widdows, J., and Readman, J.W. 2006. Influence of cockle Cerastoderma edule bioturbation and tidal-current cycles on resuspension of sediment and polycyclic aromatic hydrocarbons. Marine Ecology Progress Series 328: 51–64. Ciutat, A., Widdows, J., and Pope, N.D. 2007. Effect of Cerastoderma edule density on nearbed hydrodynamics and stability of cohesive muddy sediments. Journal of Experimental Marine Biology and Ecology 346:114–126. Commito, J.A., Celano, E.A., Celico, H.J., Como, S., and Johnson, C.P. 2005. Mussels matter: postlarval dispersal dynamics altered by a spatially complex ecosystem engineer. Journal of Experimental Marine Biology and Ecology 316:133–147. Crawford, C.M., Macleod, C.K.A., and Mitchell, I.M. 2003. Effects of shellfish farming on the benthic environment. Aquaculture 224:117– 140. Cranford, P.J., Strain, P.M., Dowd, M., Hargrave, B.T., Grant, J., and Archambault, M.-C. 2007. Influence of mussel aquaculture on nitrogen dynamics in a nutrient enriched coastal embayment. Marine Ecology Progress Series 347:61–78. Cranford, P.J., Hargrave, B.T., and Doucette, L.I. 2009. Benthic organic enrichment from suspended mussel (Mytilus edulis) culture in Prince Edward Island, Canada. Aquaculture 292:189–196.

Bivalves as bioturbators and bioirrigators

Dahlhoff, E.P., Stillman, J.H., and Menge, B.A. 2002. Physiological community ecology: variation in metabolic activity of ecologically important rocky intertidal invertebrates along environmental gradients. Integrative and Comparative Biology 42:862–871. Dame, R.F. 1993. Bivalve Filter Feeders and Coastal and Estuarine Ecosystem Processes. Springer Verlag, Heidelberg. Dame, R., Bushek, D., Allen, D., Lewitus, A., Edwards, D., Koepfler, E., and Gregory, L. 2002. Ecosystem response to bivalve density reduction: management implications. Aquatic Ecology 36:51–65. D’Amours, O., Archambault, P., McKindsey, C.W., and Johnson, L.E. 2008. Local enhancement of epibenthic macrofauna by aquaculture activities. Marine Ecology Progress Series 371:73–84. Doering, P.H., Kelly, J.R., Oviatt, C.A., and Sowers, T. 1987. Effect of the hard clam Mercenaria mercenaria on benthic fluxes of inorganic nutrients and gases. Marine Biology 94:377–383. Dumbauld, B.R., Ruesink, J.L., and Rumrill, S.S. 2009. The ecological role of bivalve shellfish aquaculture in the estuarine environment: a review with application to oyster and clam culture in West Coast (USA) estuaries. Aquaculture 290:196–223. Erwin, D.H. 2008. Macroevolution of ecosystem engineering, niche construction and diversity. Trends in Ecology and Evolution 23:304– 310. Everett, R.A., Ruiz, G.M., and Carlton, J.T. 1995. Effect of oyster mariculture on submerged aquatic vegetation: an experimental test in a Pacific Northwest estuary. Marine Ecology Progress Series 125:205–217. Folke, C., and Kautsky, N. 1989. The role of ecosystems for a sustainable development of aquaculture. Ambio 18:234–243. François, F., Gerino, M., Stora, G., Durbec, J.-P., and Poggiale, J.-C. 2002. Functional approach to sediment reworking by gallery-forming macrobenthic organisms: modeling and application with the polychaete Nereis diversicolor. Marine Ecology Progress Series 229:127– 136. Gerino, M., Aller, R.C., Lee, C., Cochran, J.K., Aller, J.Y., Green, M.A., and Hirschberg, D. 1998. Comparison of different tracers and

313

methods used to quantify bioturbation during a spring bloom: 234-thorium, luminophores and chlorophyll a. Estuarine, Coastal and Shelf Science 46:531–547. Gerino, M., Stora, G., François, F., Gilbert, J.C., Poggiale, J.-C., Mermillod-Blondin, F., Desrosiers, G., and Vervier, P. 2003. Macroinvertebrate functional groups in freshwater and marine sediments: a common mechanistic classification. Vie Milieu 53:222–231. Giles, H., Pilditch, C.A., and Bell, D.G. 2006. Sedimentation from mussel (Perna canaliculus) culture in the Firth of Thames, New Zealand: impacts on sediment oxygen and nutrient fluxes. Aquaculture 261:125–140. Glud, R.N. 2008. Oxygen dynamics of marine sediments. Marine Biology Research 4:243–289. Gowen, R.J., and Bradbury, N.B. 1987. The ecological impact of salmonid farming in coastal waters: a review. Oceanography and Marine Biology: An Annual Review 25:563–575. Graf, G., and Rosenberg, R. 1997. Bioresuspension and biodeposition: a review. Journal of Marine Systems 11:269–278. Grant, J., Hatcher, A., Scott, D.B., Pocklington, P., Schafer, C.T., and Winters, G.V. 1995. A multidisciplinary approach to evaluating impacts of shellfish aquaculture on benthic communities. Estuaries 18:124–144. Green, M.O., Hewitt, J.E., and Thrush, S.F. 1998. Seabed drag coefficient over natural beds of horse mussels (Atrina zelandica). Journal of Marine Research 56:613–637. Gren, I.-M., Lindahl, O., and Lindqvist, M. 2009. Values of mussel farming for combating eutrophication: an application to the Baltic Sea. Ecological Engineering 35:935–945. Grizzle, R.E., and Penniman, C.A. 1991. Effects of organic enrichment on estuarine macrofaunal benthos: a comparison of sediment profile imaging and traditional methods. Marine Ecology Progress Series 74:249–262. Gutiérrez, J.L., Jones, C.G., Strayer, D.L., and Iribarne, O.O. 2003. Molluscs as ecosystem engineers: the role of shell production in aquatic habitats. Oikos 101:79–90. Hansen, K., King, G.M., and Kristensen, E. 1996. Impact of the soft-shell clam Mya arenaria on sulfate reduction in an intertidal sediment. Aquatic Microbial Ecology 10:181–194.

314

Shellfish Aquaculture and the Environment

Hargrave, B.T., Doucette, L.I., Cranford, P.J., Law, B.A., and Milligan, T.G. 2008. Influence of mussel aquaculture on sediment organic enrichment in a nutrient-rich coastal embayment. Marine Ecology Progress Series 365:137–149. Hartstein, N.D., and Rowden, A.A. 2004. Effect of biodeposits from mussel culture on macroinvertebrate assemblages at sites of different hydrodynamic regime. Marine Environmental Research 57:339–357. Hatcher, A., Grant, J., and Schofield, B. 1994. Effects of suspended mussel culture (Mytilus spp.) on sedimentation, benthic respiration and sediment nutrient dynamics in a coastal bay. Marine Ecology Progress Series 115:219– 235. Heck, K.L., Jr., Hays, G., and Orth, R.J. 2003. Critical evaluation of the nursery role hypothesis for seagrass meadows. Marine Ecology Progress Series 253:123–136. Hedman, J.E., Bradshaw, C., Thorsson, M.H., Gilek, M., and Gunnarsson, J.S. 2008. Fate of contaminants in Baltic Sea sediments: role of bioturbation and settling organic matter. Marine Ecology Progress Series 356:25–38. Hewitt, J.E., Thrush, S.F., Halliday, J., and Duffy, C. 2005. The importance of small-scale habitat structure for maintaining beta diversity. Ecology 86:1619–1626. Hewitt, J.E., Thrush, S.F., Dayton, P.K., and Bonsdorff, E. 2007. The effect of spatial and temporal heterogeneity on the design and analysis of empirical studies of scale-dependent systems. The American Naturalist 169:398– 408. Iglesias, J.I.P., Urrutia, M.B., Navarro, E., AlvarezJorna, P., Larretxea, X., Bougrier, S., and Heral, M. 1996. Variability of feeding processes in the cockle Cerastoderma edule (L.) in response to changes in seston concentration and composition. Journal of Experimental Marine Biology and Ecology 197:121–143. Inglis, G.J., and Gust, N. 2003. Potential indirect effects of shellfish culture on the reproductive success of benthic predators. Journal of Applied Ecology 40:1077–1089. Jackson, J.B.C., Kirby, M.X., Berger, W.H., Bjorndal, K.A., Botsford, L.W., Bourque, B.J., Bradbury, R.H., Cooke, R., Erlandson, J., Estes, J.A., Hughes, T.P., Kidwell, S., Lange, C.B.,

Lenihan, H.S., Pandolfi, J.M., Peterson, C.H., Steneck, R.S., Tegner, M.J., and Warner, R.R. 2001. Historical overfishing and the recent collapse of coastal ecosystems. Science 293:629– 638. Jaramillo, E., Bertrán, C., and Bravo, A. 1992. Mussel biodeposition in an estuary in southern Chile. Marine Ecology Progress Series 82:85–94. Jaramillo, E., Contreras, H., and Duarte, C. 2007. Community structure of the macroinfauna inhabiting tidal flats characterized by the presence of different species of burrowing bivalves in Southern Chile. Hydrobiologia 580:85– 96. Jones, C.G., Lawton, J.H., and Shachak, M. 1994. Organisms as ecosystem engineers. Oikos 69: 373–386. Kaiser, M.J., Edwards, D.B., and Spencer, B.E. 1996. Infaunal community changes as a result of commercial clam cultivation and harvesting. Aquatic Living Resources 9:57–63. Kalantzi, I., and Karakassis, I. 2006. Benthic impacts of fish farming: meta-analysis of community and geochemical data. Marine Pollution Bulletin 52:484–493. Karlson, K., Rosenberg, R., and Bonsdorff, E. 2002. Temporal and spatial large-scale effects of eutrophication and oxygen deficiency on benthic fauna in Scandinavian and Baltic waters—a review. Oceanography and Marine Biology: An Annual Review 40:427–489. Kautsky, N., and Evans, S. 1987. Role of biodeposition by Mytilus edulis in the circulation of matter and nutrients in a Baltic coastal ecosystem. Marine Ecology Progress Series 38:201–212. Kristensen, E. 1988. Benthic fauna and biogeochemical processes in marine sediments: microbial activities and fluxes. In: Blackburn, T.H., and Sørensen, J. (eds.), Nitrogen Cycling in Coastal Marine Environments. Wiley, New York, pp. 275–299. Le Hir, P., Monbet, Y., and Orvain, F. 2007. Sediment erodability in sediment transport modelling: can we account for biota effects? Continental Shelf Research 27:1116–1142. Levinton, J. 1995. Bioturbators as ecosystem engineers: control of the sediment fabric, inter-individual interactions, and material fluxes. In: Jones, C.G., and Lawton, J.H. (eds.), Linking

Bivalves as bioturbators and bioirrigators

Species and Ecosystems. Chapman & Hall, New York, pp. 29–38. Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L.-O., Olrog, L., Rehnstam-Holm, A.-S., Svensson, J., Svensson, S., and Syversen, U. 2005. Improving marine water quality by mussel farming: a profitable solution for Swedish society. Ambio 34:131–138. Lohrer, A.M., Thrush, S.F., and Gibbs, M.M. 2004. Bioturbators enhance ecosystem function through complex biogeochemical interactions. Nature 431:1092–1095. Maire, O., Duchêne, J.C., Grémare, A., Malyuga, V.S., and Meysman, F.J.R. 2007. A comparison of sediment reworking rates by the surface deposit-feeding bivalve Abra ovata during summertime and wintertime, with a comparison between two models of sediment reworking. Journal of Experimental Marine Biology and Ecology 343:21–36. Maire, O., Lecroart, P., Meysman, F., Rosenberg, R., Duchêne, J.-C., and Grémare, A. 2008. Quantification of sediment reworking rates in bioturbation research: a review. Aquatic Biology 2:219–238. Martin, W.R., and Banta, G.T. 1992. The measurement of sediment irrigation rates: a comparison of the BR− tracer and 222RN/226RA disequilibrium techniques. Journal of Marine Research 50: 125–154. Meysman, F.J.R., Galaktionov, O.S., Gribsholt, B., and Middelburg, J.J. 2006a. Bio-irrigation in permeable sediments: an assessment of model complexity. Journal of Marine Research 64:589–627. Meysman, F.J.R., Middelburg, J.J., and Heip, C.H.R. 2006b. Bioturbation: a fresh look at Darwin’s last idea. Trends in Ecology and Evolution 21:688–695. Michaud, E., Desrosiers, G., Mermillod-Blondin, F., Sundby, B., and Stora, G. 2006. The functional group approach to bioturbation: II. The effects of the Macoma balthica community on fluxes of nutrients and dissolved organic carbon across the sediment–water interface. Journal of Experimental Marine Biology and Ecology 337:178–189. Montserrat, F., Van Colen, C., Provoost, P., Milla, M., Ponti, M., Van den Meersche, K., Ysebaert, T., and Herman, P.M.J. 2009. Sediment segrega-

315

tion by biodiffusing bivalves. Estuarine, Coastal and Shelf Science 83:379–391. Naylor, R.L., Goldburg, R.J., Primavera, J.H., Kautsky, N., Beveridge, M.C.M., Clay, J., Folke, C., Lubchenco, J., Mooney, H., and Troell, M. 2000. Effect of aquaculture on world fish supplies. Nature 405:1017–1024. Neira, C., Grosholz, E.D., Levin, L.A., and Blake, R. 2006. Mechanisms generating modification of benthos following tidal flat invasion by a Spartina hybrid. Ecological Applications 16:1391–1404. Newell, R.I.E. 2004. Ecosystem influences of natural and cultivated populations of suspension-feeding bivalve molluscs: a review. Journal of Shellfish Research 23:51–61. Newell, R.I.E., Cornwell, J.C., and Owens, M.S. 2002. Influence of simulated bivalve biodeposition and microphytobenthos on sediment nitrogen dynamics: a laboratory study. Limnology and Oceanography 47:1367–1379. Newell, R.I.E., and Koch, E.W. 2004. Modeling seagrass density and distribution in response to changes in turbidity stemming from bivalve filtration and seagrass sediment stabilization. Estuaries 27:793–806. Nizzoli, D., Welsh, D.T., Fano, E.A., and Viaroli, P. 2006. Impact of clam and mussel farming on benthic metabolism and nitrogen cycling, with emphasis on nitrate reduction pathways. Marine Ecology Progress Series 315:151–165. Norkko, A., Hewitt, J.E., Thrush, S.F., and Funnell, G.A. 2001. Benthic-pelagic coupling and suspension-feeding bivalves: linking site-specific sediment flux and biodeposition to benthic community structure. Limnology and Oceanography 46:2067–2072. Norkko, A., Hewitt, J.E., Thrush, S.F., and Funnell, G.A. 2006. Conditional outcomes of facilitation by a habitat-modifying subtidal bivalve. Ecology 87:226–234. Norling, P., and Kautsky, N. 2007. Structural and functional effects of Mytilus edulis on diversity of associated species and ecosystem functioning. Marine Ecology Progress Series 351:163–175. Pearson, T.H., and Rosenberg, R. 1978. Macrobenthic succession in relation to organic enrichment and pollution of the marine environment. Oceanography and Marine Biology: An Annual Review 16:229–311.

316

Shellfish Aquaculture and the Environment

Perez, K.T., Davey, E.W., Moore, R.H., Burn, P.R., Rosol, M.S., Cardin, J.A., Johnson, R.L., and Kopans, D.N. 1999. Application of computeraided tomography (CT) to the study of estuarine benthic communities. Ecological Applications 9:1050–1058. Petersen, J.K., Hansen, J.W., Laursen, M.B., Clausen, P., Carstensen, J., and Conley, D.J. 2008. Regime shift in a coastal marine ecosystem. Ecological Applications 18:497–510. Peterson, B.J., and Heck, K.L., Jr. 2001a. An experimental test of the mechanism by which suspension feeding bivalves elevate seagrass productivity. Marine Ecology Progress Series 218:115– 125. Peterson, B.J., and Heck, K.L., Jr. 2001b. Positive interactions between suspension-feeding bivalves and seagrass—a facultative mutualism. Marine Ecology Progress Series 213:143–155. Porter, E.T., Cornwell, J.C., Sanford, L.P., and Newell, R.I.E. 2004. Effect of oysters Crassostrea virginica and bottom shear velocity on benthic– pelagic coupling and estuarine water quality. Marine Ecology Progress Series 271:61–75. Prins, T.C., Smaal, A.C., and Dame, R.F. 1998. A review of the feedbacks between bivalve grazing and ecosystem processes. Aquatic Ecology 31:349–359. Quintana, C.O., Tang, M., and Kristensen, E. 2007. Simultaneous study of particle reworking, irrigation transport and reaction rates in sediment bioturbated by the polychaetes Heteromastus and Marenzelleria. Journal of Experimental Marine Biology and Ecology 352:392–406. Reise, K. 2002. Sediment mediated species interactions in coastal waters. Journal of Sea Research 48:127–141. Rhoads, D.C. 1974. Organism-sediment relations on the muddy sea floor. Oceanography and Marine Biology: An Annual Review 12:223– 300. Rhoads, D.C., and Young, D.K. 1971. Animalsediment relations in Cape Cod Bay, Massachusetts II. Reworking by Molpadia oolitica (Holothuroidea). Marine Biology 11: 255–261. Rodhouse, P.G., and Roden, C.M. 1987. Carbon budget for a coastal inlet in relation to intensive cultivation of suspension-feeding bivalve molluscs. Marine Ecology Progress Series 36:225–236.

Sandwell, D.R., Pilditch, C.A., and Lohrer, A.M. 2009. Density dependent effects of an infaunal suspension-feeding bivalve (Austrovenus stutchburyi) on sandflat nutrient fluxes and microphytobenthic productivity. Journal of Experimental Marine Biology and Ecology 373:16–25. Schaffner, L.C., Jonsson, P., Diaz, R.J., Rosenberg, R., and Gapcynski, P. 1992. Benthic communities and bioturbation history of estuarine and coastal systems: effects of hypoxia and anoxia. In: Vollenweider, R.A., Marchetti, R., and Viviani, R. (eds.), Marine Coastal Eutrophication. Elsevier, Amsterdam, pp. 1001–1016. Sgro, L., Mistri, M., and Widdows, J. 2005. Impact of infaunal Manila clam, Ruditapes philippinarum, on sediment stability. Hydrobiologia 550:175–182. Shumway, S.E., and Kraeuter, J.N. (eds.). 2004. Molluscan shellfish research and management: charting a course for the future. Final Proceedings from the Cooperative Research and Information Institute (CRII) Workshop, Charleston, SC, January 2000. Smith, J., and Shackley, S.E. 2004. Effects of a commercial mussel Mytilus edulis lay on a sublittoral, soft sediment benthic community. Marine Ecology Progress Series 282:185–191. Solan, M., Wigham, B.D., Hudson, I.R., Kennedy, R., Coulon, C.H., Norling, K., Nilsson, H.C., and Rosenberg, R. 2004. In situ quantification of bioturbation using time-lapse fluorescent sediment profile imaging (f-SPI), luminophore tracers and model simulation. Marine Ecology Progress Series 271:1–12. Stead, R.A., and Thompson, R.J. 2006. The influence of an intermittent food supply on the feeding behaviour of Yoldia hyperborea (Bivalvia: Nuculanidae). Journal of Experimental Marine Biology and Ecology 332:37– 48. Vaudrey, J.M.P., Getchis, T., Shaw, K., Markow, J., Britton, R., and Kremer, J.N. 2009. Effects of oyster depuration gear on eelgrass (Zostera marina L.) in a low density aquaculture site in Long Island Sound. Journal of Shellfish Research 28:243–250. Vaughn, C.C., and Hakenkamp, C.C. 2001. The functional role of burrowing bivalves in freshwater ecosystems. Freshwater Biology 46: 1431–1446.

Bivalves as bioturbators and bioirrigators

Viitasalo, S. 2007. Effects of bioturbation by three macrozoobenthic species and predation by necto-benthic mysids on cladoceran benthic eggs. Marine Ecology Progress Series 336: 131–140. Viitasalo, S., Katajisto, T., and Viitasalo, M. 2007. Bioturbation changes the patterns of benthic emergence in zooplankton. Limnology and Oceanography 52:2325–2339. Vinther, H.F., and Holmer, M. 2008. Experimental test of biodeposition and ammonium excretion from blue mussels (Mytilus edulis) on eelgrass (Zostera marina) performance. Journal of Experimental Marine Biology and Ecology 364:72–79. Vinther, H.F., Laursen, J.S., and Holmer, M. 2008. Negative effects of blue mussel (Mytilus edulis) presence in eelgrass (Zostera marina) beds in Flensborg fjord, Denmark. Estuarine, Coastal and Shelf Science 77:91–103. Volkenborn, N., Polerecky, L., Hedtkamp, S.I.C., van Beusekom, J.E.E., and de Beer, D. 2007. Bioturbation and bioirrigation extend the open exchange regions in permeable sediments. Limnology and Oceanography 52:1898– 1909. Volkenborn, N., Robertson, D.M., and Reise, K. 2009. Sediment destabilizing and stabilizing bioengineers on tidal flats: cascading effects of experimental exclusion. Helgoland Marine Research 63:27–35. Ward, J.E., and Shumway, S.E. 2004. Separating the grain from the chaff: particle selection in suspension- and deposit-feeding bivalves. Journal of Experimental Marine Biology and Ecology 300:83–130. Weise, A.M., Cromey, C.J., Callier, M.D., Archambault, P., Chamberlain, J., and McKindsey, C.W. 2009. Shellfish-DEPOMOD: modelling the biodeposition from suspended shellfish aquaculture and assessing benthic effects. Aquaculture 288:239–253. Weissberger, E.J., Coiro, L.L., and Davey, E.W. 2009. Effects of hypoxia on animal burrow construction and consequent effects on sediment redox profiles. Journal of Experimental Marine Biology and Ecology 371:60–67. Whitlatch, R.B., Hines, A.H., Thrush, S.F., Hewitt, J.E., and Cummings, V. 1997. Benthic faunal responses to variations in patch density and patch size of a suspension-feeding bivalve.

317

Journal of Experimental Marine Biology and Ecology 216:171–189. Widdows, J., and Brinsley, M. 2002. Impact of biotic and abiotic processes on sediment dynamics and the consequences to the structure and functioning of the intertidal zone. Journal of Sea Research 48:143–156. Widdows, J., Brinsley, M.D., Salkeld, P.N., and Elliott, M. 1998. Use of annular flumes to determine the influence of current velocity and bivalves on material flux at the sediment-water interface. Estuaries 21:552–559. Willows, R.I., Widdows, J., and Wood, R.G. 1998. Influence of an infaunal bivalve on the erosion of an intertidal cohesive sediment: a flume and modeling study. Limnology and Oceanography 43:1332–1343. Widdows, J., Brown, S., Brinsley, M.D., Salkeld, P.N., and Elliott, M. 2000. Temporal changes in intertidal sediment erodability: influence of biological and climatic factors. Continental Shelf Research 20:1275–1289. Widdows, J., Blauw, A., Heip, C.H.R., Herman, P.M.J., Lucas, C.H., Middelburg, J.J., Schmidt, S., Brinsley, M.D., Twisk, F., and Verbeek, H. 2004. Role of physical and biological processes in sediment dynamics of a tidal flat in Westerschelde Estuary, SW Netherlands. Marine Ecology Progress Series 274:41–56. Worm, B., Barbier, E.B., Beaumont, N., Duffy, J.E., Folke, C., Halpern, B.S., Jackson, J.B.C., Lotze, H.K., Micheli, F., Palumbi, S.R., Sala, E., Selkoe, K.A., Stachowicz, J.J., and Watson, R. 2006. Impacts of biodiversity loss on ocean ecosystem services. Science 314:787–790. Wotton, R.S., and Malmqvist, B. 2001. Feces in aquatic ecosystems. BioScience 51:537– 544. Wu, R.S.S. 1995. The environmental impact of marine fish culture: towards a sustainable future. Marine Pollution Bulletin 31:159–166. Ysebaert, T., Hart, M., and Herman, P.M.J. 2009. Impacts of bottom and suspended cultures of mussels Mytilus spp. on the surrounding sedimentary environment and macrobenthic biodiversity. Helgoland Marine Research 63: 59–74. Zwarts, L., and Wanink, J. 1989. Siphon size and burying depth in deposit- and suspension-feeding benthic bivalves. Marine Biology 100: 227–240.

Chapter 11

Environmental impacts related to mechanical harvest of cultured shellfish Kevin D.E. Stokesbury, Edward P. Baker, Bradley P. Harris, and Robert B. Rheault

Introduction Shellfish are important to the health of marine estuaries. High shellfish biomass is associated with a robust healthy estuarine ecosystem where the entire water volume of an estuary is filtered every few days. Low shellfish biomass is associated with a compromised ecosystem. Healthy estuaries support economically healthy tourism, recreation, and fishing industries. Few would debate these assertions (generalizations); however, the topic of shellfish aquaculture often stirs an adverse response that may or may not be based in fact. Some believe shellfish aquaculture offers tremendous social benefit arguing it is quintessentially sustainable, helps keep estuaries healthy, provides

jobs, increases the supply of highly nutritious food, and increases the diversity of the nation’s food supply. If this line of thinking is true and shellfish aquaculture is valuable, it is important to address objections scientifically. We present here an overview of the current scientific thought on the environmental impacts of shellfish harvesting and a method for scientifically assessing impact of shellfish harvesting. The number of shellfish farms located in public trust waters of the United States has grown steadily over the past two decades. The 2005 Census of Aquaculture reports 1222 shellfish farms nationwide and total product sales exceeding $185 million. Along with an increase in the number of farms is an increase

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 319

320

Shellfish Aquaculture and the Environment

in the number of objections to shellfish farming. Mechanical harvesting (e.g., dredging or raking), which disturbs the sediment, is a leading complaint. Shellfish farms typically operate in shallow coastal or estuarine environments and are exposed to natural disturbance from both terrestrial and marine sources. Heavy rainfall hundreds of miles inland can result in a pronounced drop in salinity of estuary headwaters; rainy years and dry years cause unusually long periods of low or high salinities; low or high seasonal temperatures can effect species survival; wave action from storms resuspends sediment turning entire embayments brown with turbidity and settling everywhere. Exceptionally high wave action can physically alter the benthos and the benthic community. Anoxic and hypoxic events may cause wholesale destruction of animal communities. Coastal waters are dynamic and make detection of significant impacts from mechanical harvesting of shellfish difficult. Accurately assessing impact requires a formal experimental design with a clear hypothesis and subsequent gathering of appropriate data and statistical analysis to determine if harvest impact on the environment is significant or not. The before-after control-impact (BACI) experimental design provides the framework for testing harvest impact hypotheses. Over the past decade, we have conducted cooperative research of the biology and ecology of sea scallops (Placopecten magellanicus) on Georges Bank. Some of this research examined the impact of scallop dredges on the benthos and benthic organisms and for this research the BACI experimental design (Stokesbury and Harris 2006) was employed. In fact, the method of this research serves as a precedent to resolve disputes about harvest impact of shellfish. When it was determined that an unprecedented number of scallops worth more than $70 million in the Nantucket Lightship Closed Area were going to be lost due to oldage mortality, an effort to open the closed area

was put forth. The initiative was opposed and a lawsuit was filed in U.S. Federal Court. The case, Conservation Law Foundation et al. v. Donald Evans, Secretary of Commerce et al., was heard in the nation’s capital and after reviewing the scientific findings of the assessment of dredge impact on the benthic community, using the BACI method, the court supported the opening of the closed area to a scallop harvest and commended the industry for its support and participation in the research.

Literature review Overview The topic of environmental impacts of fishing gear has generated a great deal of debate and numerous published studies. The impacts have been compared with clear-cutting: “trawling gear devastates the world’s continental shelves” (Baulch 1999) and generating “watery wastelands” (Levy 1998). The reality of the situation is more nuanced and requires an understanding of the physical and biological environments, the type of gear being used, and how these variables interact. The following is an excerpt from a review “Effects of Trawling and Dredging on Seafloor Habitat” by the Ocean Studies Board (OSB) (Dorsey and Pederson 1998). Many experimental studies have documented the acute, gear-specific effects of trawling and dredging on various types of habitat. The results confirm predictions based on the ecological principle that stable communities of low mobility, long-lived species will be more vulnerable to acute and chronic physical disturbance than will short-lived species in changeable environments. Trawling and dredging can reduce habitat complexity by removing or damaging the biological and physical structures of the seafloor. The extent of the initial effects and the rate of recovery

Environmental impacts of mechanical harvest

depend on the habitat stability. The more stable biogenic (i.e., of biological origin), gravel, and mud habitats experience the greatest changes and have the slowest recovery rates. In contrast, less consolidated coarse sediments in areas of high natural disturbance show fewer initial effects. Because those habitats tend to be populated by opportunistic species that re-colonize more rapidly, recovery is faster as well.

The direct effects of fishing gear are the most obvious and the vast majority of the literature addresses these effects. They include

• Reduction of complexity and diversity or shifts in community structure following large-scale removal of target species as well as bycatch or capture of nontarget species • Loss of vertical structure which provides critical habitat and refuge for juvenile fish and crustaceans • Reduction in productivity or biomass following harvest Potential indirect effects (from Dorsey and Pederson 1998) include the following:

• Nutrient cycling. Seafloor trawling and dredging could increase or decrease the exchange rate of nutrients between the sediment and water column and introduce pulses of productivity in addition to pulses from the natural seasonal cycle. • Hypoxia. Suspension and oxidation of sulfides can scavenge oxygen and cause local areas of low oxygen or hypoxia. • Increased susceptibility to other stressors. Loss of physical structure in a habitat can expose organisms to other stressors, such as predation. • Turbidity. Suspension of sediments can limit visibility and decrease light penetration. This might impact feeding behavior, impair respiration, or slow the growth of algae or submerged aquatic vegetation (SAV).

321

Numerous authors have compiled litanies of potential impacts of dredge harvesting (for these, see Bradstock and Gordon 1983; Thrush et al. 1995; Collie et al. 1997; Auster and Langton 1999). Others have suggested that dredging impacts are transient and mimic natural disturbances such as storms, and some even suggest reworking the sediment stimulates benthic productivity (MacKenzie 1982; Van Dolah et al. 1991; Currie and Parry 1996; De Alteris et al. 1999). This chapter will examine the literature as it pertains to the impacts of mechanical harvest methods for cultured shellfish on leased beds, pointing out the differences between these methods and the use of offshore trawls and dredges used to capture wild fish and shellfish.

Significant impacts What is an “impact” and what is “significant”? Since no activity can be done without an impact (Newton’s third law), the important questions are as follows: What is a significant impact? What can be considered a positive or beneficial impact (Langan 1998)? What temporal scale is being considered? Are the impacts significant if the effects are undetectable after a few days, weeks, or months or if the impacts pale in comparison with frequent natural disturbances such as storms (De Alteris et al. 1999)? Are the impacts significant if they are restricted to a small area or if they result in ecosystem benefits? The nature of these questions demands a subjective response that must balance a host of issues. Clearly, there are impacts to be avoided. These include wholesale changes in ecosystem structure or trophic energy flow; long-term changes that are not recoverable within a reasonable time frame after the practice ceases (such as introductions of diseases or nonnative species); and negative impacts on threatened or endangered species. These are all clear

322

Shellfish Aquaculture and the Environment

examples of “significant” impacts. Beyond these examples are vast areas of subjective interpretation where the answer one gets often depends on the spatial or temporal scale of the question.

Descriptions of dredges Dredges, typically consisting of a net on a frame towed behind a boat, have been designed to harvest epifauna or infauna, and the configuration of the gear varies greatly depending on the target species and the substrate. Some dredges skim the surface of the seabed while fishing as exemplified by offshore sea scallop (Placopecten magellanicus) fishery. Other dredges utilize hydraulic jets, toothed rakes, or suction apparatus to harvest shellfish located within the sediment. Mechanical harvest of cultured shellfish is typically done with a rakelike device with a trailing bag to collect the catch. They remove molluscan shellfish from the seabed, and they are also used to harvest crustacea, finfish, and echinoderms. None of this should be confused with channel dredging, which is used to deepen or widen waterways by removing sediment. Dredges used to harvest shellfish are designed to capture shellfish, leaving the sediment behind. In estuarine waters, dredges are used to collect clams, mussels, oysters, conch, and crabs (Conner and Simon 1979; Ruffin 1995). Beds of mussels are incredibly dense and can range from the intertidal to subtidal. Most mussel beds are surveyed at low tide and marked with buoys where they are to be dredged. Mussel dredges are considerably lighter than most other dredges, and although the tows are very short in duration (2–5 min), they are very efficient, yielding 10,000 lb in 2–3 h with little to no bycatch. A typical oyster dredge consists of a steel frame, 0.3–2.0 m wide, with a toothed blade (Fig. 11.1). The tow chain or wire and a bag for the catch are attached to the frame. The dredge is towed

Figure 11.1 Oyster dredge consists of a steel frame, 0.3– 2.0 m wide, with a toothed blade. (Photo by Steve Allen.)

slowly (<1 m s−1), from vessels that are 7–15 m long. Similar dredges are used to catch blue crabs in the mid-Atlantic region during the winter. These dredges can have teeth as long as 8 cm that penetrate soft bottoms to capture partially buried shellfish. To capture shellfish that live deeper than a few centimeters in the sediment, fishermen fit dredges or rakes with longer teeth or occasionally use water jets to loosen the sediment and bring the shellfish to the surface. This is commonly called a “hydraulic dredge” (Fig. 11.2). Large hydraulic clam dredges are used offshore to collect surfclams (Spisula solida) and ocean quahogs (Arctica islandica), while smaller (1–2 m) dredges are used inshore to collect soft- and hard-shell clams.

Environmental impacts of mechanical harvest

323

Figure 11.2 Hydro clam dredge using water jets to loosen the sediment and bring the shellfish to the surface. (Photo by Tessa Getchis.)

In soft clam fishery, which occurs in shallow estuarine waters, the dredge head (manifold and blade) is sometimes attached to a conveyor or “escalator” that carries the shellfish to the working deck of the vessel. Offshore, larger heavier dredges are used to catch sea scallops, which live on the surface in sand–gravel seabed. Sea scallop dredges have a steel frame, sweep chains, and a bag constructed of steel rings (Fig. 11.3). The width or mouth opening of the dredge ranges from 3.0 to 4.5 m, and dredge weight varies from 500 kg to 1000 kg. The dredge is designed to ride on shoes placed on the outer edges of the frame so that scallops are swept up into the bag by the sweep chains.

Differences between wild harvest fishing and the harvest of cultured shellfish To better understand the range of impacts associated with dredge harvesting, it is helpful

to differentiate dredge harvesting of cultured crops on leased bottom from dredge harvesting of wild shellfish. The two practices differ greatly in the frequency and scale of dredging activity, with important ramifications on the degree of impact (Kaiser et al. 1996; Collie et al. 2000). Wild harvest fishermen do not know the spatial distribution of shellfish density or where other fishermen have been fishing. Consequently, they typically tow in areas based on their experience. They will sample grounds that were historically productive and occasionally try new areas to see if populations have shifted, concentrating their effort on areas with high yields until the densities have been reduced. Since fishermen rarely coordinate their efforts collectively, they are not systematic or efficient and may dredge the same areas repeatedly while searching for a good tow. In contrast, when a grower uses a dredge on his lease, he knows exactly where to go to maximize the catch of market size shellfish (he seeded the area), and when an area is

324

Shellfish Aquaculture and the Environment

Figure 11.3 The New Bedford offshore sea scallop dredge used in this fishery weighs about 1870 kg, was 4.5 m wide, has a series of vertical and horizontal sweep chains, which prevent large rocks from entering the bag, a 20.3-mm diamond mesh twine top, a 4.5 × 0.8 m bag knit of 89-mm steel rings, and rubber chaffing gear. (Photo by B. Harris.)

harvested the farmer knows not to waste further effort there. These tows are usually much shorter because the density of the shellfish is much higher on leased grounds than in wild populations. Because the tows are shorter, the mortality of nontarget organisms that might be caught in the net is greatly reduced. Most shellfish farming takes place in shallow coastal waters with sandy or silty bottom. The species that live in these waters are well adapted to periodic disturbances from storm events and wave action (De Alteris et al. 1999). Species in these environments tend to be

opportunists that rapidly recolonize disturbed bottom and are tolerant of high loads of suspended sediment. These environments recover from major disturbances within a few weeks or months (Coen 1995). One of the main impacts attributed to dredge harvesting is the flattening of vertical structure and so reducing habitat complexity (Bradstock and Gordon 1983). Dredges can damage “emergent epifauna” such as sponges or corals that provide important habitat for juvenile fish and other species. Most inshore shellfish lease sites are devoid of such vertical structure. Further, shellfish farmers reseed their crops annually. In the case of oyster farms using extensive culture practice on hard bottom, seed may be broadcast directly on the bottom or as spat-on-shell. Either technique greatly enhances vertical structure. By planting shell and shellfish seed, growers replace the vertical structure that serves as vital habitat for many species. A final difference between wild harvest dredging and dredging on leased bottom is that shellfish farmers need to allow their crops to grow undisturbed for many months, in some cases up to 3 years, before harvesting. At any given time, a farmer will harvest from only a small portion of his grounds while the rest is left to grow. As a result, cultured bottom may be more diverse and productive because of the influence of high shellfish biomass and the culture activities.

Impacts on species diversity and productivity In the marine environment, stable substrates are often in limited supply and the addition of any firm substrate or vertical relief attracts organisms increasing local diversity. The removal of shells, bivalves, and other structures by harvest gear of any type will have the opposite effect. In oyster aquaculture, unlike the wild fishery, the shell and juvenile shellfish

Environmental impacts of mechanical harvest

are replanted after harvest so the vertical structure is replaced. In clam aquaculture grounds, there is typically little structure to begin with, the disturbance of harvest activity is short term, and recovery is rapid. Seed clams are replanted following harvest so productivity and biomass recover quickly. The following is excerpted from a review by Coen (1995): Most studies agree that dredging causes some mortality to small and large infaunal and epifaunal organisms in the direct path of the device (Godcharles 1971, Kyte et al. 1975, Kyte and Chew 1975, Vining 1978, Meyer et al. 1981, Mackenzie 1982, Peterson et al. 1987, Barnes et al. 1991). However, since many of these small benthic organisms (crustaceans, polychaetes, molluscs) have rapid generation times, high fecundities and excellent recolonization capacities, it is generally accepted that this community effect is only short-term (e.g., Godcharles 1971, Peterson et al. 1987, Bennett et al. 1990, Hall et al. 1990). Hall et al. (1990) suggest that the effects will be apparent and protracted only if the fauna are primarily immobile or if the affected area is large relative to remainder of the habitat.

In addition to the removal of the target species, and some nontarget species, diversity is temporarily impacted by the attraction of predatory species to the dredge tracks to eat uncovered or damaged prey. Within 1 h of scallop dredging, Caddy (1973) observed predators at densities up to 30 times those outside sea scallop dredge tracks, especially winter flounder, and also sculpin and rock crabs. Similarly, Eleftheriou and Robertson (1992) noted congregations of fish (primarily pleuronectids, gadoids, and gobies) feeding in scallop dredge tracks, as well as seastars and a large variety of crustaceans. Where hoes were used for commercial digging for soft-shell clams in Maine, the species richness and density of three poly-

325

chaete species were significantly reduced (Brown and Wilson 1997). Dolmer et al. (1999) noted reduced densities of small polychaetes found after dredging for bottom cultures mussels, while infaunal abundance and diversity decreased immediately following suction dredge harvesting of Manila clams from an area of muddy sand bottom in Northern Europe (Spencer et al. 1998). Bycatch mortality is an issue with offshore dredge fisheries. Nontarget species are often killed because they were crushed in the bottom of a full net bag, or in the case of fish because their swim bladder expands when exposed to low pressures at the surface. Shellfish farmers rarely see these problems because (1) they tow slowly, allowing finfish to avoid the dredge; (2) they focus fishing in areas with very high densities of the target species; (3) tows are very short; and (4) they are working shallow waters.

Timescales of recovery Most studies indicate that, especially in shallow high-energy environments that are adapted to frequent disturbances, the communities tend to recover quickly from dredging impacts. While significant impacts are often observed immediately following bottom culture harvesting in unvegetated, soft-sediment habitat, quick recovery of invertebrate communities appears quite common (Kaiser et al. 1998a, 1998b). This resiliency of the benthos is characteristic of shallow-water coastal and estuarine systems, which are subjected to continual disturbances (Turner et al. 1995). In addition to controlled experiments, there are dozens of anecdotal reports of massive sets of clams (Mya, Mercenaria, or Spisula) following dredging, significant storms, or oil spills (Visel, pers. comm.). These species have adapted to rapidly colonize empty niches. Perhaps one of the most complete studies on faunal impact was done in Florida by Godcharles (1971), who discovered no lasting

326

Shellfish Aquaculture and the Environment

impacts on the benthic populations. Using three gear types (a benthic corer, a trynet trawl, and a hydraulic escalator dredge) to sample both infauna and epifauna, they reported little difference between control and experimental dredging sites. Recovery was slowest in vegetated areas, which were completely stripped of plants by the dredge and had a maximum recovery time of 13 months. Godcharles reported no significant faunal differences between control and experimental plots (including the vegetated stations) in the trynet samples. Tarnowski (2001) reviewed recovery times from various sources of man-made impacts and found that with few exceptions recovery was rapid, in most cases on the order of months. Goodwin and Shaul (1978) evaluated the impact of a mechanical escalator harvester on a subtidal clam bed in Puget Sound, WA. The abundance of attached kelp was reduced and dredging left large amounts of old clamshell and sand at the substrate surface. The harvest had little effect on the number of benthic animal species, but did reduce the number of individuals and the weight per unit area of some organisms. Most species had recovered to the control plot levels in 1 year. Hall et al. (1990) evaluated the impact of hydraulic dredging for razor clams (Ensis sp.) on an infaunal community in a Scottish Sea Loch. Infaunal samples from replicate fished and unfished plots were examined at 1 and 40 days postharvest. After 40 days, no effects of fishing could be detected and no visible signs of fishing remained on the seabed. They concluded that “…hydraulic dredging is unlikely to have persistent effects on most of the infaunal community in most habitats.” Hall and Harding (1997) examined the effects of hydraulic suction dredging and tractor dredging of cockles (Cerastostema edule) on nontarget benthic infauna. Although the suction dredge experiment revealed some

statistically significant effects, the faunal structure in disturbed plots recovered (i.e., approached that of the undisturbed controls) after only 56 days. They conclude “that mechanical harvesting methods impose high levels of mortality on nontarget benthic fauna, but that recovery of disturbed sites is rapid and the overall effects on populations is probably low.” Kaiser et al. (1996) examined infaunal community changes as a result of commercial Manila clam (Tapes philippinarum) cultivation and harvesting in England. Harvesting by suction dredge altered sediment composition and reduced the density and the total number of species. Seven months later, however, no significant differences were found between the infaunal community in the harvested clam bed and either of the control areas, indicating that the practice of clam cultivation has no longterm effects on the environment or benthic community at this site. Several researchers have pointed to the importance of evaluating fishing impacts in light of the impacts of storm disturbances (De Alteris et al. 1999; Stokesbury and Harris 2006). Unfortunately, relatively little is known about the impacts of severe storm events, and it is difficult to conduct controlled experiments to measure their effects. While most experimental studies are able to detect short-term impacts, long-term changes are probably restricted to long-lived fragile species or communities found in environments that are infrequently disturbed by natural phenomena (Kaiser 1998).

Impacts on vertical structure Benthic organisms (plants, corals, and sponges) and sediment forms (mud burrows and gravel) add structure to the seafloor and increase habitat complexity (Freese et al. 1999; Kreiger 2001). Seafloor structures serve as nurseries

Environmental impacts of mechanical harvest

for juvenile fish and provide refuge and food for adults. Even small structures, such as cobbles and clamshells, can form important habitat. Areas of the seafloor that lack these structures do not support the variety of fish populations observed in more complex regions (Collie et al. 1997; Kaiser et al. 1999). With repeated trawling, the physical relief of the seafloor is reduced and juveniles of many fish species which aggregate near seabed structure are depleted and there is an overall reduction in benthic production (Jennings et al. 2001). Also, removal of physical structure in a habitat can force some species into less optimal environments. For instance, the dredging of natural oyster reefs in North Carolina has lowered the reefs’ vertical relief, forcing oysters to settle in deeper areas that are more prone to hypoxia (Lenihan and Peterson 1998).

Impacts on SAV Submerged aquatic vegetation (SAV) is important to numerous species and is often protected under federal law as essential fish habitat (EFH). There is no question that dredges designed to penetrate the substrate will damage SAV. Eelgrass (Zostera marina) is an important component of estuarine areas from Nova Scotia to North Carolina, and is the primary habitat for the economically important bay scallop (Argopecten irradians). Tarnowski (2001) characterized the impact of hydraulic dredges on eelgrass as “catastrophic.” In many U.S. jurisdictions, shellfish leases are rarely granted in areas with significant eelgrass; however, there are some. Peterson et al. (1987) compared various methods of clam harvesting (Mercenaria mercenaria). Harvesting with clam kicking boats (a practice that uses propellers aimed downward to blow sediment off wide areas to expose buried clams) caused a 65% reduction in seagrass biomass relative to

327

controls, and recovery was only partial after 4 years. Further, in areas harvested by hand raking, which is perceived to have a relatively lower disturbance level, seagrass biomass decreased 25%. Oyster culture leases harvested by dredging displayed decreased eelgrass shoot density, shoot length, and biomass compared with reference plots. Biomass was reduced 30% after 1 year and 96% after 4 years, with effects persisting up to 2 years posttreatment. Hilary et al. (2005) found that commercial dragging for blue mussels posed a severe threat to eelgrass, with a recovery rate of 6 years in favorable environments and more than 20 years in less conducive conditions.

Sediment resuspension: turbidity impacts Sediment resuspension is an impact of many activities including channel dredging, shellfish harvesting, boat traffic, and land runoff. Storms also stir up sediments in shallow waters and can turn embayments brown with suspended sediment; for this reason most estuarine species are predictably tolerant of high suspended sediment loads (O’Connor et al. 1976, 1977; Thistle 1981). The observed effects are site specific as a consequence of sediment grain size and type and hydrological conditions. (Barnes et al. 1991). Harvest dredges (especially those that use water jets) will resuspend sediment in the water column. Most of the larger particles settle almost immediately, but fine silt can remain suspended for days. Most studies show that over 95% of the sediment sinks to the bottom within a few tens of meters of the source (reviewed in Coen 1995; Black and Parry 1999). The persistence of fine particles in the water column increases turbidity and reduces light levels. This can temporarily decrease predator feeding success and enhance prey survival in

328

Shellfish Aquaculture and the Environment

some cases (Vinyard and O’Brien 1976; Gregory 1990); however, results are system or species dependent, variable, and often conflicting (Johnston and Wildish 1982; Boehlert and Morgan 1985; Hanson and Walton 1990). Turbidity can affect eggs, larval, juvenile and adult fishes, and shellfish in estuarine and marine ecosystems (reviewed in Peddicord et al. 1975 for invertebrates, Simenstad 1990 for fish). Hoffman and Dolmer (2000) studied the effects of a large mussel dredging fishery in Denmark on the distribution of fish and epibenthic invertebrates and found no longterm effects. Tarnowski (2001) reported that hydraulic dredge impacts to turbidity are worst in fine silty clay sediments because the particles remain suspended longest. The maximum distance of detectable deposits resulting from hydraulic dredging was 22.9 m (75 ft), while another study found negligible sedimentation at 4.6 m (15 ft) from a dredging site. Values as high as 584 mg L−1 of suspended solids were recorded at the conveyor belt of an escalator dredge working in a silt/clay mud flat (Kyte and Chew 1975). This value rapidly dropped to 89 mg L−1 at a distance of 61 m (200 ft) from the dredge, although a plume was still visible. Background silt loadings at the site varied from 4 to 441 mg L−1. The following is excerpted from Coen (1995): Although the effects of shellfish dredging on turbidity levels have not been studied, the organisms that live in these highly variable, estuarine ecosystems typically encounter elevated and highly variable suspended sediment loads, with ambient seston levels often varying by several orders of magnitude over short durations (e.g., daily, Kyte et al. 1975, Settlemyre and Gardiner 1977, Auld and Schubel 1978, Barnes et al. 1991). Hence, they are generally considered tolerant of short-term perturbations (Kyte et al. 1975). Also, most of the fishes and crustaceans (with the exception of barnacles) are highly mobile.

Simenstad (1990) concluded that most estuarine fishes move out or are adapted to elevated suspended sediments and that most behavioral or sublethal effects seen in the lab are even more ambiguous when extrapolated to the field. Auld and Schubel (1978) concluded the same for eggs and larvae of six Chesapeake Bay species. Thus, while the effects remain unknown, it is unlikely that the limited turbidity plumes created by subtidal or intertidal shellfish dredging operations have a major impact on the biological resources in those habitats.

Turbidity can reduce light levels and the resultant shading could have an impact on plant growth. If dredging activities occur to such an extent that light levels are reduced below ambient levels for extended periods or over wide areas then eelgrass, benthic macroalgae and phytoplankton might be affected; however, this impact has not been documented.

Other impacts of resuspended sediments Marine sediments typically become anoxic below the surface due to the consumption of oxygen by bacteria that are decomposing the organic matter that collects on the bottom. The greater the flux of organic matter to the bottom, the greater the rate of oxygen consumption. In deeper sediment pore waters, oxygen becomes depleted and sulfate is reduced to hydrogen sulfide (Nixon 1981). This gives the sediment its characteristic black color and “rotten egg” odor. The depth at which the sediment turns black is called the redox layer (short for reduction-oxidation). In very productive environments, the anoxic black layer of sediment is very close to the surface because organic matter is being deposited faster than it can be consumed by the bacteria. If you stir these sediments into oxy-

Environmental impacts of mechanical harvest

genated water, the hydrogen sulfide (H2S) will be oxidized (sediment oxygen demand). Organic matter also contains nutrients such as nitrogen and phosphate. As organic matter decomposes, these nutrients which are bound in tissues are remineralized into soluble forms and released into the sediment pore water (Nixon 1981). Stirring the sediments can temporarily cause an acceleration of the release of nutrients, but the total amount released over time is not affected by disrupting the sediments unless the dredge digs deep enough to stir up nutrients that have been “buried” or sediments where pore waters no longer travel (Barnes et al. 1991). Kyte et al. (1975) found that harvesting had little long-term effect on the local water chemistry. Ambient seston levels (6.9–441 mg L−1) often exceeded those associated with harvesting, thus obscuring any potential shortterm effects. Few persistent effects on water column chemistry (e.g., nutrients, dissolved oxygen [DO], H2S) were observed. Most of the literature that describes the potential impact of sediment resuspension on water chemistry is related to dredging for channel maintenance. The magnitude of sediment resuspension when one is excavating and then dumping tons of sediments is clearly far greater than the disturbances caused by a hydraulic dredge. Few studies document this impact from fishing gear. Krost (1990, 1993) estimated an annual oxygen demand of 491–2656 t O2 due to the release of H2S by sediment resuspension caused by global trawling. Dispersed over the estimated area being fished annually, this is minimal; however, locally, the impacts might be measurable.

Conclusions Both studies designed to assess the impacts of hydraulic harvesters in shallow estuaries (Coen 1995; Tarnowski 2001) concluded that the

329

impacts were reversible, short term, and, due to the limited scale of the activity, unlikely to have significant adverse impacts except in the case of dredging in eelgrass beds, which is prohibited in most states. The following is excerpted from Coen (1995): Overall, findings consistently support the same conclusion: the short-term effects of subtidal escalator harvesters are minimal, with no long-term chronic effects, even under worst case scenarios. Observed effects are often indistinguishable from ambient levels or natural variability. These conclusions are based on field experimentation and knowledge of natural estuarine variation (physical, chemical and biological). The most obvious effects (e.g., sediment plume) cease when operations are halted, but natural events are continuous. Naturally high turbidities and variable river discharges are common to South Carolina, hence it is predictable that direct effects are probably within previously observed norms. Estuarine communities appear, in general, to be tolerant of the short-term harvester effects including resuspension/turbidity, direct burial/smothering, nutrient release and decreased water quality due to elevated biochemical oxyen demand (BOD), and direct disturbance or removal of infauna.

Experimental design Overview Creating a valid experimental design in the marine environment can be difficult. It can be difficult to collect data and the high degree of spatial and temporal variation (patchiness) can necessitate large collections. It is important to keep the design basic and simple in order to produce high-quality results. Proper use of statistics requires several assumptions about the data being collected. We review these concepts

330

Shellfish Aquaculture and the Environment

in a cursory fashion here, but proper use of powerful statistical tools presumes that certain assumptions about the data have been met. For instance, data must be randomly sampled, normally distributed, and not skewed. Adequate numbers of samples must be used to account for natural variability. If these concepts are foreign, then it is advisable to recruit the assistance of an extension agent or university personnel to help. Statisticians can do remarkable things with data. As Disraeli (Green 1979) said, “there are lies, damn lies and statistics.” It is best to limit harvest impact questions to a small number of variables (species of interest) and determine the presence or absence of an impact based on simple observations and understandable statistics. The most important step is to establish a simple testable hypothesis. Whether a shellfish farming site has been established for years or is just starting, it is best to decide what will be monitored and begin gathering data right away. Here we can make an important distinction. Examining the conditions of the marine environment before a harvest event serves as a critical control even if the site has been established for several years. The argument that “you did not sample before your aquaculture site was established” is not valid as only the impact of harvesting is being tested. The impact of the aquaculture site as a whole on the environment is a much bigger question, which this experi-

ment might help with, but will not answer completely.

BACI experimental design The BACI design assumes that the control and impact areas have similar environments and communities, and that these communities will change over time in the same fashion, except for any harvest-caused disturbances in the impact areas (Green 1979). The optimal BACI design uses a two-way analysis of variance (ANOVA) where the interaction between the control site and time is compared with the interaction between the impact site and time to statistically detect an impact (Green 1979). The two-way ANOVA is only reliable if populations (of organisms or water quality conditions) in the control and impact areas are equal. This is rarely the case in marine field studies (especially when farm sites have enhanced populations of shellfish), and the statistics required to deal with this inequality are complex and controversial (for an example, see Black and Miller 1991; Rangeley 1994; Black and Miller 1994). Several researchers have suggested using only graphs and tables to indicate environmental impacts, while others recommend statistical tests which are usually limited to t-tests and one-way ANOVAs (Green 1979; Stewart-Oaten et al. 1986; Underwood 1994). The sampling protocol is straightforward (Table 11.1).

Table 11.1 Sampling design for a before-after control-impact experiment where the impact is mechanical harvesting of shellfish. Preimpact

1 2

Control

Impact

x x

x x

Postimpact

Impact Impact

Recovery 1

Control

Impact

Control

Impact

x x

x x

x x

x x

1 and 2 are experimental plots. X = at least 30 random samples measuring number of species and number of individuals per species.

Environmental impacts of mechanical harvest

Control site The control is important for two reasons: (1) the data gathered at the control site are what is compared with the data gathered at the impact (harvest) site in order to demonstrate the presence or absence of an impact; and (2) the control site accounts for natural variation. Ideally, the control site should not be influenced by the harvest site. The control and impact sites should have similar physical, chemical, and biological characteristics. The assumption is that naturally occurring changes in these characteristics will affect both sites in the same manner; for example, in the event of a storm, the control and impact sites would both experience the same level of wave action, turbidity, and the same drop in salinity from rain water.

What to measure? Measured variables can be physical, chemical, or biological; and they can be in the water column, on the seabed surface, or in the sediment. Typically, biological surveys of macrofauna (organisms visible to the naked eye) are conducted with quadrat samples in which the number of each species is recorded per a standard unit area (e.g., 1 m2 quadrat). There are also methods for sampling the benthic infauna by taking core samples and sifting them or using a suction pump to empty a box core and collect the infauna in a mesh bag (usually 1–5mm mesh). The bags are usually labeled, placed in a cooler, and analyzed later in the laboratory with a dissecting microscope; identification of benthic infauna is labor intensive and often requires specific training (if this type of analysis is required, seek professional assistance). An off-the-shelf guide book such as the Peterson Field Guides Atlantic Seashore will serve as a preliminary identification text for macrofauna. Also, a great deal of identification information is available at websites such

331

as Wikipedia, but information on the Web should always be validated with published literature.

When to measure? Determining when to sample after a harvest event is one of the most important design features and speaks to the subjective nature of determining impacts. Taking samples a day after the harvest will almost certainly show a difference when compared with the control site. Sampling 2 months later may reveal no differences between the impact and control sites. Determining this time interval should be done on a case-by-case basis and centers on what variable is being used to assess impacts (macroinvertebrates, aquatic vegetation, polychaetes), considering factors such as harvesting frequency, species life cycles, and the residence time of water in the estuary.

Random sampling It is difficult to implement truly random sampling. Consider dividing the control and impact areas into equal numbered units and using a random number generator to determine where to sample. Try and sample at least 30 locations. Random number generators can be found in standard spreadsheet software or free online at www.random.org.

Determining sample size It is better to take many small quadrats than a few large quadrats (Krebs 1999). The golden rule of sampling is that if the optimal number of samples is unknown, then take at least 30. According to the central limit theorem, as long as sampling is random and there are suitable numbers of the species of interest, 30 samples will produce an accurate representation of the

332

Shellfish Aquaculture and the Environment

U

H

Unharvested

Harvested

Figure 11.4 In a hypothetical experimental design for an aquaculture impact, there are two sites: one is harvested (H), which is the impact site, and one is unharvested (U), which is the control site. In each area, 30 random quadrat samples are collected (black dots).

population. If you want to learn more, we recommend Kreb’s Ecological Methodology (2nd ed., 1998), which is an excellent guide to sampling. A hypothetical BACI study design with 30 quadrats randomly placed in the control and impact study areas is presented in Figure 11.4 and produces the summary data presented in Table 11.2. In this example, the quadrat samples are taken with a digital still camera on a frame providing a 1.3-m2 sample area (Fig. 11.5). Four macrofaunal species were counted in each quadrat: mussels, sea stars, sea scallop, and rock crabs. The summary data includes the total counts observed for the four species, the average, and the standard deviation (Table 11.2). The standard deviation indicates how much each quadrat count varies from the average count. These four species had similar densities in the control and impact areas before the harvest. After the harvest, the densities in the control area remained the same. However, in the

Table 11.2 Summary data from 30 randomly located quadrats in the impact (harvested) and control (unharvested) areas; the sum, mean average, standard deviation of the mean (SD), coefficient of variance (CV = Mean/SD), the number of samples required to obtain a 25% precision (n), and the 95% confidence limits for the means (CL) are shown. Control Quadrats

Before Sum Average SD CV n 95% CL After Sum Average SD CV n 95% CL

Harvested

Mussels

Scallops

Sea stars

Rock crabs

Mussels

Scallops

Sea stars

Rock crabs

368 12.3 7.90 0.64 27 2.83

99 3.3 2.39 0.73 34 0.86

149 5.0 3.03 0.61 24 1.09

41 1.4 1.25 0.91 53 0.45

383 12.8 7.99 0.63 25 2.86

116 3.9 1.81 0.47 14 0.65

156 5.2 3.18 0.61 24 1.14

46 1.5 1.14 0.74 35 0.41

362 12.1 7.82 0.65 27 2.80

98 3.3 2.20 0.67 29 0.79

129 4.3 3.37 0.78 39 1.21

46 1.5 1.11 0.72 33 0.40

72 2.4 1.73 0.72 33 0.62

96 3.2 2.30 0.72 33 0.82

893 29.8 12.26 0.41 11 4.39

163 5.4 3.33 0.61 24 1.19

Environmental impacts of mechanical harvest

333

Figure 11.5 Two 1.3-m2 quadrat samples from the Great South Channel off of Cape Cod, MA. The upper quadrat contains horse mussels, shell hash, and granular pebble substrate; the lower quadrat contains sea stars, a sea scallop, rock crab, shell hash, and a fine sand substrate.

harvest area, the number of mussels decreases significantly (after harvest average of 2.4 ± 0.62 compared with 12.8 ± 2.86 mussels per quadrat), sea stars and rock crabs increased significantly, while scallops have remained the same. To conduct a two-way ANOVA of the data in Table 11.2, we used Sigmastat® (Systat Software, Chicago, IL) version 3.5. It is user-

friendly and provides excellent summary tables. The two-way ANOVA indicated significantly less (P < 0.001) mussels at the harvest impact site when compared with the control site. This indicates there is less than a one in one thousand chance that this kind of difference would occur by chance. In this example, we sampled before and after a single impact event. Creating a time

334

Shellfish Aquaculture and the Environment

series of samples before and after several harvest events will provide more powerful harvest impact statistics. Often, in wild fishery BACI experiments, an impact is immediately apparent but undetectable after a few weeks or months depending on the natural dynamics of the environment, that is, an area of high tidal currents or frequent storms.

Shellfish farmer as marine scientist We recommend shellfish farmers develop a data gathering and analysis program to monitor abundance of carefully selected species and/or water quality parameters. There are many benefits to gathering data at appropriate times and places (temporal and spatial intervals) at a shellfish farm site and control site. Mainly, the analysis of the data will provide the grower with a better understanding of the ecology of their farm site. Second, with an eye toward good stewardship and environmental sustainability, science-based conclusions will reveal whether a shellfish farm operation or the shellfish harvest activities significantly impact the environment. If there is significant negative impact, the farm may have to change harvesting practices. If the data indicate there is no impact, the farm can defend itself from misguided or spurious objections. If the data show a positive impact, it may win over more shellfish farm supporters by showing that shellfish culture is good for the marine environment. A data gathering program will demonstrate a proactive and responsible approach to farming. It will also create a remarkable data set that may be used by scientists studying trends in the marine environment. Farm and control site water quality monitoring may include 1. Secchi disk measurements; once a week 2. Surface and bottom water temperature measurements; once a week

3. Surface and bottom water DO measurements; eight times a year Estimated time to acquire a set of measurements = 30 min Farm and control site measurements of species may include 1. Placing length of rope at various random locations and tallying fouling organisms on 2 in. of rope; four times a year Estimated time to gathering data per event = 2 h; 8 h per year 2. Survey of epifaunal and epifloral communities; four times a year Estimated time to gather data per event = 4 h; 16 h per year 3. Camera survey of macrovertebrates (fish, crustaceans, etc.) using cheap underwater camera with computer download capacity; four times a year. Estimated time to gathering data per event = 2 h Total water quality and species measurements = approximately 40 h per year

Conclusions Harvesting the produce of a shellfish aquaculture operation is, in many ways, the most rewarding part of the business. Growth in the industry has resulted in more frequent sightings of men and machines hauling product out of the water, and concerns have been raised about the impacts of harvesting on the environment. While these concerns parallel those associated with harvesting wild shellfish (e.g., scallop dredging), they are not comparable. There have been a number of bibliographies and reviews of the current literature on the impacts of harvesting shellfish (see Hopkins and McKinney 1976; Coen 1995; Dorsey and Pederson 1998; Jennings and Kaiser 1998;

Environmental impacts of mechanical harvest

Kaiser et al. 1998a, 1998b; Rester 2000; Barnette 2001; Dieter et al. 2003; McKindsey et al. 2006). The literature is diverse and often controversial, and it can be argued either way depending on the point of view being put forward. In our work with the scallop industry of New England, we conducted a large BACI experiment following the procedure outlined above. The experiment was presented in federal court and helped to support the NMFS decision to allow limited scallop harvest in closed areas of Georges Bank, which the judge ultimately upheld. Having data specific to the impacts of harvesting aquaculture sites collected under a simple experimental design will greatly reduce the arguments against aquaculture operations by reducing the amount of uncertainty surrounding the question of environmental impacts.

Acknowledgments We thank Sandy Shumway for asking us to contribute this chapter. Mike Dadswell provided a review of an early version and offered many helpful suggestions.

Literature cited Auld, A.H., and Schubel, J.R. 1978. Effects of suspended sediment on fish eggs and larvae: a laboratory assessment. Estuarine, Coastal and Marine Science 6:153–164. Auster, P.J., and Langton, R.W. 1999. The effects of fishing on fish habitat. In: Benaka, L.R. (ed.), Fish Habitats: Essential Fish Habitat and Rehabilitation. American Fisheries Society Symposium 22, Bethesda, MD. Barnes, D., Chytalo, K., and Hendrickson, S. 1991. Final policy and generic environmental impact statement on management of shellfish in uncertified areas program. New York Department of Environment and Conservation, 79 pp. Barnette, M.C. 2001. A review of the fishing gear utilized within the Southeast Region and their

335

potential impacts on essential fish habitat. NOAA Technical Memorandum NMFS-SEFSC449. Baulch, H. 1999. Clear-cutting the ocean floor: trawling gear devastates the world’s continental shelves. Alternatives Journal 25(3):7. Bennett, D.H., Chandler, J.A., Dunsmoor, L.K., and Barila, T. 1990. Use of dredged material to enhance fish habitat in Lower Granite reservoir, Idaho-Washington. In: Simenstad, C.A. (ed.), Effects of Dredging on Anadromous Pacific Coast Fishes. Seattle, WA, September 8–9, 1988, pp. 132–143. Workshop Proceedings, University of Washington and WA Sea Grant Program. Black, R., and Miller, R.J. 1991. Use of the intertidal zone by fish in Nova Scotia. Environmental Biology of Fishes 31:109–121. Black, R., and Miller, R.J. 1994. The effects of seaweed harvesting on fishes: a response. Environmental Biology of Fishes 39:325– 328. Black, K.P., and Parry, G.D. 1999. Entrainment, dispersal, and settlement of scallop dredge sediment plumes: field measurements and numerical modeling. Canadian Journal of Fisheries and Aquatic Sciences 56(12):2271–2281. Boehlert, G.W., and Morgan, J.B. 1985. Turbidity enhances feeding abilities of larval pacific herring, Clupea harengus pallasi. Hydrobiologia 123:161–170. Bradstock, M., and Gordon, D.P. 1983. Coral-like bryozoan growths in Tasman Bay, and their protection to conserve commercial fish stocks. New Zealand Journal of Marine and Freshwater Research 17(2):159–163. Brown, B., and Wilson, Jr., W.H. 1997. The role of commercial digging of mudflats as an agent for change of infaunal intertidal populations. Journal of Experimental Marine Biology and Ecology 218(1):49–61. Caddy, J.F. 1973. Underwater observations on tracks of dredges and trawls and some effects of dredging on a scallop ground. Journal of Fisheries Research Board of Cananda 30:173–180. Coen, L.D. 1995. A review of the potential impacts of Mechanical Harvesting on subtidal and intertidal shellfish resources. Prepared for the SCDNR 46pp. Collie, J.S., Escanero, G.A., and Valentine, P.C. 1997. Effects of bottom fishing on the benthic

336

Shellfish Aquaculture and the Environment

megafauna of Georges Bank. Marine Ecology Progress Series 155:159–172. Collie, J.S., Hall, S.J., Kaiser, M.J., and Poiner, I.R. 2000. A quantitative analysis of fishing impacts on shelf-sea benthos. The Journal of Animal Ecology 69(5):785–798. Conner, W.G., and Simon, J.L. 1979. The effects of oyster shell dredging on an estuarine benthic community. Estuarine, Coastal and Marine Science 9:749–758. Conservation Law Foundation et al. v. Donald Evans, Secretary of Commerce et al., 209 F.Supp.2d 1 (D.D.C. 2001) No. 00CV1134 (GK). December 28, 2001. Currie, D.R., and Parry, G.D. 1996. Effects of scallop dredging on a soft sediment community: a large-scale experimental study. Marine Ecology Progress Series 134(1–3):131–150. De Alteris, J., Skrobe, L., and Lipsky, C. 1999. The significance of seabed disturbance by mobile fishing gear relative to natural processes: a case study in Narragansett Bay, Rhode Island. In: Benaka, L.R. (ed.), Fish Habitats: Essential Fish Habitat and Rehabilitation. American Fisheries Society, Symposium 22, Bethesda, MD, pp. 224–237. Dieter, B.E., Wion, D.A., and McConnaughey, R.A. (eds.). 2003. Mobile fishing gear effects on benthic habitats: a bibliography. 2nd ed. NOAA Technical Memorandum NMFS-AFSC-135. Dolmer, P., Kristensen, P.S., and Hoffmann, E. 1999. Dredging of blue mussels (Mytilus edulis L.) in a Danish sound: stock sizes and fisheryeffects on mussel population dynamic. Fisheries Research 40:73–80. Dorsey, E.M., and Pederson, J. (eds.), 1998. Effects of fishing gear on the sea floor of New England. In: Effects of Trawling and Dredging on Seafloor Habitat (2002) Ocean Studies Board (OSB). MIT Sea Grant Publication 98-4, Boston, MA. Eleftheriou, A., and Robertson, M.R. 1992. The effects of experimental scallop dredging on the fauna and physical environment of a shallow sandy community. Netherlands Journal of Sea Research 30:289–299. Freese, L., Auster, P.J., Heifetz, J., and Wing, B.L. 1999. Effects of trawling on seafloor habitat and associated invertebrate taxa in the Gulf of Alaska. Marine Ecology Progress Series 182:119–126.

Godcharles, M.F. 1971. A study of the effects of a commercial hydraulic clam dredge on benthic communities in estuarine areas. State of Florida Department of Natural Resources, Marine Resources Laboratory. Technical Series No. 64. Goodwin, L., and Shaul, W. 1978. Studies of the mechanical escalator harvester on a subtidal clam bed in Puget Sound, Washington. Progress Report No. 53. State of Washington Department of Fisheries. 23 p. Green, R.H. 1979. Sampling Design and Statistical Methods for Environmental Biologists. John Wiley & Sons, New York. Gregory, R.S. 1990. Effects of turbidity on benthic foraging and predation risk in juvenile chinook salmon. In: Simenstad, C.A. (ed.), Effects of Dredging on Anadromous Pacific Coast Fishes. Seattle, WA, September 8–9, 1988, pp. 64–73. Workshop proceedings, Seattle, September 8–9, Washington Sea Grant Report WSG-WO 90-01. Hall, S.J., and Harding, M.J.C. 1997. Physical disturbance and marine benthic communities: the effects of mechanical harvesting of cockles on non-target benthic infauna. The Journal of Applied Ecology 34(2):497–517. Hall, S.J., Basford, D.J., and Robertson, M.R. 1990. The impact of hydraulic dredging for razor clams Ensis sp. on an infaunal community. Netherlands Journal of Sea Research. 27(1):119–125. Hanson, C.H., and Walton, C.P. 1990. Potential effects of dredging on early life stages of striped bass (Morone saxatilis) in the San Francisco Bay area: an overview. In: Simenstad, C.A. (ed.), Effects of Dredging on Anadromous Pacific Coast Fishes. Seattle, WA, September 8–9, 1988, pp. 38–56. Workshop proceedings, Seattle, September 8–9, Washington Sea Grant Report WSG-WO 90-01. Hilary, N.A., Short, F.T., Barker, S., and Kopp, B.S. 2005. Disturbance of eel grass Zostera marina by commercial mussel Mytilus edulis harvesting in Maine: dragging impacts and habitat recovery. Marine Ecology Progress Series 285: 57–73. Hoffman, E., and Dolmer, P. 2000. Effect of closed areas on distribution of fish and epibenthos. ICES Journal of Marine Science 57(5):1310– 1314.

Environmental impacts of mechanical harvest

Hopkins, S.H., and McKinney, L.D. 1976. A review of the literature pertaining to the effects of dredging on oyster reefs and their associated faunas. In: Bouma, A.H. (ed.), Shell Dredging and Its Influence on Gulf Coast Environments. Gulf Publishing Company, Houston, TX, pp. 3–12. Jennings, S., and Kaiser, M.J. 1998. The effects of fishing on marine ecosystems. Advances in Marine Biology 34:201–352. Jennings, S., Pinnegar, J.K., Polunin, N.V.C., and Warr, K.J. 2001. Impacts of trawling disturbance on the trophic structure of benthic invertebrate communities. Mar Ecol Prog Ser 213:127–142. Johnston, D.D., and Wildish, D.J. 1982. Effects of suspended sediment on feeding by larval herring (Clupea harengus harengus L.). Bulletin of Environmental Contamination and Toxicology 29:261–267. Kaiser, M.J. 1998. Significance of bottom-fishing disturbance. Conservation Biology 12(6):1230– 1235. Kaiser, M.J., Edwards, D.B., and Spencer, B.E. 1996. Infaunal community changes as a result of commercial clam cultivation and harvesting. Aquatic Living Resources 9:57–63. Kaiser, M.J., Armstrong, P.J., Dare, P.J., and Flatt, R.P. 1998a. Benthic communities associated with a heavily fished scallop ground in the English Channel. Journal of the Marine Biological Association of the United Kingdom 78(4):1045–1059. Kaiser, M.J., Laing, I., Utting, S.D., and Burnell, G.M. 1998b. Environmental impacts of bivalve mariculture. Journal of Shellfish Research 17(1):59–66. Kaiser, M.J., Rogers, S.I., and Ellis, J.R. 1999. Importance of benthic habitat complexity for demersal fish assemblages. In: Benaka, L.R. (ed.), Fish Habitat: Essential Fish Habitat and Rehabilitation. American Fisheries Society, Symposium 22, Bethesda, MD, pp. 212–223. Krebs, C.J. 1999. Ecological Methodology, 2nd ed.. Benjamin/Cummings, Addison-Welsey Educational publishers Inc., Menlo Park, CA. Kreiger, K.J. 2001. Coral impacted by fishing gear in the Gulf of Alaska. In: Willison, J., Hall, J., Gass, S., Kenchington, E., Butler, M., and Doherty, P. (eds.), Proceedings of the First International Symposium on Deep-Sea Corals.

337

Proceedings of a Symposium held at Dalhousie University, Halifax, Nova Scotia, Canada, July 30–August 2, 2000. Ecology Action Centre and Nova Scotia Museum, Halifax, NS, pp. 106–117. Krost, P. 1990. The impact of otter-trawl fishery on nutrient release from the sediment and macrofauna of Kieler Bucht (western Baltic). Ph.D. dissertation. Berichte aus dem Institut fur Meereskunde an der Christian-AlbrechtsUniversitat Kiel. Kiel. 200 160 pp. Krost, P. 1993. The significance of the bottom trawl fishery for the sediment, its exchange processes, and the benthic communities in the Bay of Kiel. Arbeiten des Deutschen Fischerei-Verbandes. NA(57):43–60. Kyte, M.A., and Chew, K.K. 1975. A review of the hydraulic escalator shellfish harvester and its known effects in relation to the soft-shell clam, Mya arenaria. Washington Sea Grant Publication WSG 75-2, 1–32 pp. Kyte, M., Averill, P., and Hendershott, T. 1975. The impact of the hydraulic escalator shellfish harvester on an intertidal soft-shell clam flat in the Harraseeket River, Maine. Dep. Mar. Res., Augusta, Maine, Project Completion Report, 54 pp. Langan, R. 1998. The effect of dredge harvesting on eastern oysters and the associated benthic community. In: Dorsey, E.M., and Pederson, J. (eds.), Effects of Fishing Gear on the Sea Floor of New England. MIT Sea Grant Pub. 98-4, Boston, MA. Lenihan, H.S., and Peterson, C.H. 1998. How habitat degradation through fishery disturbance enhances impacts of hypoxia on oyster reefs. Ecological Applications 8(1):128– 140. Levy, S. 1998. Watery wastelands. New Scientist 158(2134):40–44. MacKenzie, C.L. 1982. Compatibility of invertebrate populations and commercial fishing for ocean quahogs. North American Journal of Fisheries Management 2:270–275. McKindsey, C.W., Anderson, M.R., Barnes, P., Courtenay, S., Landry, T., and Skinner, M. 2006. Effects of shellfish aquaculture on fish habitat (2006) DFO Canadian science advisory secretariat. www.dfo-mpo.gc.ca/csas/ Research Document 2006/011.

338

Shellfish Aquaculture and the Environment

Meyer, T.L., Cooper, R.A., and Pecci, K.J. 1981. The performance and environmental effects of a hydraulic clam dredge. Marine Fisheries Review 43:14–22. Nixon, S.W. 1981. Remineralization and nutrient cycling in estuarine ecosystems. In: Neilson, B.J., and Cronin, L.E. (eds.), Estuaries and Nutrients. Humana Press, Clifton, NJ, pp. 111–138. O’Connor, J.M., Neumann, D.A., and Sherk, J.A. 1976. Lethal effects of suspended sediments on estuarine fish. U.S. Coastal Engineering Research Technical Paper 76(20):1–38. O’Connor, J.M., Neumann, D.A., and Sherk, J.A. 1977. Sublethal effects of suspended sediments on estuarine fish. U.S. Coastal Engineering Research Technical Paper 77(3):1–90. Peddicord, R.K., McFarland, V.A., Belfiori, D.P., and Byrd, T.E. 1975. Effects of suspended solids on San Francisco Bay organisms. USACOE Dredge Disposal Study, San Francisco Bay and estuary, 1-158. Peterson, C.H., Summerson, H.C., and Fegley, S.R. 1987. Ecological consequences of mechanical harvesting of clams. Fisheries Bulletin 85:281–298. Rangeley, R.W. 1994. The effects of seaweed harvesting on fishes: a critique. Environmental Biology of Fishes 39:319–323. Rester, J.K. 2000. Annotated bibliography of fishing impacts on habitat. Ocean Springs, Mississippi, Gulf States Marine Fisheries Commission 73: 178 pp. Ruffin, K.K. 1995. The effects of hydraulic clam dredging on nearshore turbidity and light attenuation in Chesapeake Bay, Maryland. MS Thesis, University of Maryland. 97 p. Settlemyre, J.L., and Gardiner, L.R. 1977. Suspended sediment flux through a salt marsh drainage basin. Estuarine Coastal Marine Science 5:653–663. Simenstad, C.A. (ed.). 1990. Effects of dredging on anadromous Pacific coast fishes. Workshop Proceedings, University of Washington and WA Sea Grant Program, 160 pp. Spencer, B.E., Kaiser, M.J., and Edwards, D.B. 1998. Intertidal clam harvesting: benthic com-

munity change and recovery. Aquaculture Research 29(6):429–437. Stewart-Oaten, A., Murdoch, W.W., and Parker, K.R. 1986. Environmental impact assessment: “Pseudoreplication” in time? Ecology 67: 929–940. Stokesbury, K.D.E., and Harris, B.P. 2006. Impact of limited short-term sea scallop fishery on epibenthic community of Georges Bank closed areas. Marine Ecology Progress Series 307:85–100. Tarnowski, M. 2001. revised 2006) A literature review of the ecological effects of hydraulic escalator dredging. MDDNR Fisheries Technical Report Series No. 48. Thistle, D. 1981. Natural physical disturbances and communities of marine soft bottoms. Marine Ecology Progress Series. 6:223–228. Thrush, S.F., Hewitt, J.E., Cummings, V.J., and Dayton, P.K. 1995. The impact of habitat disturbance by scallop dredging on marine benthic communities: what can be predicted from the results of experiments? Marine Ecology Progress Series 129:141–150. Turner, S.J., Thrush, S.F., Pridmore, R.D., Hewitt, J.E., Cummings, V.J., and Maskery, M. 1995. Are soft-sediment communities stable? An example from a windy harbor. Marine Ecology Progress Series 120:219–230. Underwood, A.J. 1994. On beyond BACI: sampling designs that might reliably detect environmental disturbances. Ecological Applications 4:3– 15. Van Dolah, R.F., Wendt, P.H., and Vonlevisen, M. 1991. A study of the effects of shrimp trawling on benthic communities in 2 South Carolina sounds. Fisheries Research 12(2):139–156. Vining, R. 1978. Final environmental impact statement for the commercial harvesting of subtidal hardshellclams with a hydraulic escalator shellfish harvester. State of Washington, Department of Fisheries, DNR, 57 pp. Vinyard, G.L., and O’Brien, W.J. 1976. Effects of light and turbidity on reactive distance of bluegill (Lepomis macrochirus). Journal of Fisheries Research Board of Canada 33:2845– 2849.

Chapter 12

Genetics of shellfish on a human-dominated planet Dennis Hedgecock

Introduction Consideration of the genetics of shellfish in the context of aquaculture and the environment requires one to pull focus back from the current scene on the ecological stage, in which humans have realized global ecological dominance, to the broader coevolutionary play, in which shellfish and humans find themselves. This chapter reviews and synthesizes the science behind three issues pertinent to the genetics of shellfish in aquaculture and the environment: (1) genetic impacts of translocations or introductions of shellfish; (2) the genetic impacts of interbreeding between hatchery stocks and wild populations, such as might arise in either shellfish restoration

programs or commercial aquaculture; and (3) domestication and genetic improvement of shellfish for aquaculture.

Shellfish aquaculture and conservation A long view of global mollusc aquaculture suggests that we tackle these issues in reverse order, starting with domestication. Over the past three decades, world supply of living marine resources has kept pace with human consumption only because of an exponential growth in aquaculture production (FAO 2009a, 2009b). Shellfish aquaculture is a substantial part of this global growth, though it

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 339

340

Shellfish Aquaculture and the Environment

Million metric tons

50 40

30 Molluscs 20 10

0 1975

1980

1985

1990 Year

1995

2000

2005

Figure 12.1 World aquaculture production by FAO categories of aquatic animals (FAO 2009b). At top and labeled are molluscs exclusive of cephalopods, followed in order beneath by freshwater and diadromous fish, demersal marine fish, and crustaceans. All other categories (miscellaneous aquatic animals, pelagic marine fish, marine fish, and cephalopods) have contributions too small to be seen. Molluscs, primarily bivalves, comprise a substantial though decreasing share of global production.

has been slowly losing ground to finfish production (Fig. 12.1). In 1975, mollusc production was 1.5 million metric tons (Mt), 42% of total aquatic animal production of 3.6 Mt. Mollusc production had double-digit annual growth during the early 1990s, peaking at 25% in 1993, but has average a more modest annual growth rate of 4.9% over the past decade. Finfish production over the past decade, on the other hand, has expanded at an average annual rate of 6.0%, with the result that mollusc production comprised only 26% of aquatic animal production in 2007, the latest year for which statistics are available (FAO 2009b). Unless mollusc aquaculture moves offshore into the open ocean (Buck 2007; Stevens et al. 2008), future expansion will be constrained by competition for alternative uses of coastal regions, where most humans live, and by the degradation of coastal and estuarine habitats. Thus, shellfish aquaculture will likely need to become more efficient, producing more from less area. Genetic improvement and domestication are proven routes to increasing the efficiency of agricultural production.

At the same time and for many of the same reasons, there is increasing concern about conservation of natural shellfish diversity and the preservation of the ecological services that shellfish provide. The issues of humanmediated translocations and introductions and of interbreeding of hatchery and wild populations will be treated together, as impacts on shellfish conservation.

Genetics of wild and cultivated shellfish The genetics of bivalve mollusc populations has a long history, pertinent pieces of which have been reviewed (Gaffney 1996; Gosling 2003). Early work on molluscan genetics focused on the geographic structure of natural populations (e.g., Koehn et al. 1976; Buroker 1983; Karl and Avise 1992; Cunningham and Collins 1994; McDonald et al. 1996), on positive correlations of fitness-related traits, such as growth and survival, with allozyme heterozygosity (“allozyme-associated heterosis,” e.g., Zouros et al. 1980; Fujio 1982; Gaffney 1994; Zouros

Genetics of shellfish on a human-dominated planet

and Pogson 1994; David 1998; Launey and Hedgecock 2001), on quantitative genetics of complex traits, such as growth, survival, and disease resistance (Lannan 1980; Newkirk 1980; Sheridan 1997; Langdon et al. 2003; Dégremont et al. 2007; Hedgecock and Davis 2007), and most recently on the development and application of genomic tools and resources (Hedgecock et al. 2005, 2007a; Cunningham et al. 2006; Saavedra and Bachere 2006; Gaffney 2008; Tanguy et al. 2008). Genetic evidence for population subdivision has a bearing on the issues of translocations, introductions, and interactions of hatchery and wild populations to be discussed. The causes of allozymeassociated heterosis, the heritability of production characteristics, and the development of genomic approaches to understanding complex traits and physiological ecology are relevant to discussion of domestication and genetic improvement. More recent genetics research has focused on temporal genetic variation in natural shellfish populations (e.g., Hedgecock 1994; Li and Hedgecock 1998; Hedgecock et al. 2007b), as a consequence, potentially, of sweepstakes reproductive success (SRS) (Hedgecock 1994; Hedrick 2005; Sargsyan and Wakeley 2008). Large variance in reproductive success both in natural and hatchery-propagated populations must be considered in evaluating the potential genetic interaction between the two (Gaffney et al. 1993; Boudry et al. 2002).

Domestication of shellfish Domestication of terrestrial plants and animals was based on a remarkably small number of species (Diamond 1997). Wheat, corn, and rice, for example, provide more than 60% of edible dry matter and 50% of protein consumed by humans (Harlan 1995). Compared with the rather restricted diversity of terrestrial animal domesticates (i.e., birds and mammals), the phylogenetic diversity of cultivated aquatic animals encompasses a full range of bilaterally

341

symmetric animal life, from the Lophotrochozoa (gastropods and molluscs) and the Ecdysozoa (crustaceans) to the Deuterostomia (sea urchins, sea cucumbers, and teleost fish). In part, this diversity simply reflects the greater phyletic diversity of aquatic life, but it also arises because domestication of shellfish, despite cultivation dating back to Roman times (Günther 1897), is in its infancy.

Shellfish are protodomesticates Only a handful of aquatic species—all finfish (goldfish, common carp, and perhaps the aquarium fishes, guppies, and neon tetras; Balon 1995, 2004)—have been domesticated, in the sense that they are profoundly changed from wild progenitors. Under this evolutionary definition of domestication, no shellfish species can be considered domesticated. Recently, Duarte et al. (2007) claimed that aquatic species, including over 60 species of bivalve molluscs, were being rapidly domesticated, but their definition of domestication— “breeding, care, and feeding of organisms are controlled by humans”—falls far short of the profound-change criterion that most students of domestication set. Shellfish aquaculture is clearly in the protodomestication phase (Harris and Hilman 1989), in which diverse species of shellfish are no more than exploited captives (Clutton-Brock 1981). The trend documented by Duarte et al. (2007) merely reflects the growth of aquaculture globally and of interest in culturing diverse bivalves. Although the history of shellfish domestication will likely take another century to write, the history of terrestrial domestication suggests that the number of shellfish domesticates will be much smaller than the 60 species of bivalves identified by Duarte et al. (2007). Obvious species on which domestication efforts might be focused are the seven bivalve molluscs that are among the top 40 aquaculture species in the world (Fig. 12.2).

342

Shellfish Aquaculture and the Environment

Million metric tons 0

5

10

15

20

25

30

35

40

45

50

1998 1999 2000 2001 2002 2003 2004 2005 2006 2007 a

b

c

*

d e

f

g

Figure 12.2 Aquaculture production by species (data from FAO 2009b). Segments in each annual bar are coded as follows: cross-hatched segments, filter-feeding bivalve molluscs (a = Crassostrea gigas; b = Ruditapes philippinarum; c = Patinopecten yessoensis; d = Sinonovacula constricta; e = Mytilidae; f = Anadara granosa; g = Perna viridis; * = miscellaneous marine molluscs); unfilled segments, herbivorous or omnivorous freshwater fishes; filled segments, carnivorous fish or crustaceans; dark gray, miscellaneous freshwater fishes; light gray, other species. Species, by rank: (1) Pacific cupped oyster; (2) silver carp; (3) grass carp (white amur); (4) Japanese carpet shell (Manila clam); (5) common carp; (6) whiteleg shrimp; (7) catla; (8) bighead carp; (9) Nile tilapia; (10) freshwater fishes; (11) crucian carp; (12) Atlantic salmon; (13) yesso scallop; (14) Pangasius catfishes; (15) marine molluscs; (16) roho labeo; (17) milkfish; (18) constricted tagelus (razor clam); (19) sea mussels; (20) rainbow trout; (21) giant tiger prawn; (22) white amur bream; (23) Chinese mitten crab; (24) channel catfish; (25) blood cockle; (26) black carp; (27) tilapias; (28) marine fishes; (29) amur catfish; (30) red swamp crawfish; (31) snakehead; (32) green mussel; (33) mrigal carp; (34) cyprinids; (35) Japanese eel; (36) flathead gray mullet; (37) Japanese sea bass; (38) sea snails; (39) giant river prawn; (40) Mandarin fish; (41) other species.

At present, the literature on bivalve genetics, breeding, and genomics for the top seven species (Crassostrea gigas, Ruditapes philippinarum, Patinopecten yessoensis, Sinonovacula constricta, Mytilidae, Anadara granosa, and Perna viridis) is extremely thin compared with knowledge of terrestrial domesticates (Table 12.1). With the exception of the Pacific oyster, for which 72 papers were found and for which genomic resources are accumulating at a rapid pace (Hedgecock et al. 2005; Gaffney 2008), little attention is being paid to the other top bivalve species (Saavedra and Bachere 2006; Tanguy et al. 2008). Mussels are a distant second, with 29 papers, but the greater proportion of papers in the genomic resources category than in the quantitative genetics and marker development cat-

egories reflects, in part, greater interest in the ecology rather than culture of these bivalves. Thus, the knowledge base for domesticating top-producing shellfish species is quite narrow.

Advantages of domestication Why domesticate shellfish? Improvements in the characteristics and yields of domesticated species are well known and widely appreciated. Physiological changes in shellfish species will likely be just as profound as those in terrestrial domesticates and perhaps more dramatic for their rapidity since the principles of breeding are now well developed, widespread, and aided by modern molecular and genomic approaches. More importantly, substantial

Genetics of shellfish on a human-dominated planet

343

Table 12.1 Number of papers retrieved from the Institute for Scientific Information (ISI) Web of Science pertaining to breeding, genetics, and genomics for the top seven globally cultured bivalve species (FAO 2009b).

Species1

Quantitative genetics and breeding2

Molecular markers3

Genomic resources4

Union

Crassostrea gigas Ruditapes philippinarum Patinopecten yessoensis Sinonovacula constricta Mytilidae Anadara granosa Perna viridis

32 0 2 0 7 0 0

43 1 7 1 8 0 2

26 1 1 0 17 0 0

72 2 7 1 29 0 2

1

Searches done with an OR statement of common and scientific names (e.g., “Ruditapes philippinarum” OR “Manila clam” OR “Japanese carpet shell”). 2 Separate searches with topics “heritability,” “selective breeding,” and “genetic improvement” combined. 3 Separate searches with topics “microsatellite*” and “SNPs” combined. 4 Separate searches with topics “linkage map*,” “QTL,” “Marker assisted selection,” “ESTs.” “Microarray*,” “BAC,” “Transcriptomic*,” “Genomic Resources” combined.

levels of genetic diversity still exist in extant natural populations, from which most aquaculture species continue to be derived. Improvements in yield of ∼10–15% per generation have been obtained for fish (Bentsen et al. 1998; Gjedrem 2000) and for the Pacific oyster (Langdon et al. 2003). Insofar as domestication and genetic improvement of aquatic species will increase the efficiency and sustainability of aquaculture, long-term research toward these ends must be encouraged. Much of the initial improvement in production characteristics of aquatic species, such as the Atlantic salmon and the Pacific oyster, has been based on additive genetic variance and realized by standard methods of individual or family selection. In this form of breeding, animals with desired characters are bred, and they pass on to their progeny alleles that contribute positively to the desired traits; such selection steadily enriches the population with beneficial alleles, increasing the proportion of animals with desired traits and the population mean. Family selection has nearly doubled the yield of Pacific oysters over that of wild stocks in large-scale field trials (C. Langdon, pers. comm.; Fig. 12.3). There is considerable evidence that selection is effective in improving

shellfish, but there is also evidence that inbreeding and inbreeding depression can rather easily wipe out gains from selection. Nonadditive variance is also likely to be important in shellfish species. Nonadditive variance causes offspring to deviate from the average of the parental trait values. It is relatively more important in the breeding of common carp, perhaps because additive genetic variance has already been fixed during the longer history of carp domestication (Wohlfarth 1993). Nonadditive variance appears to be relatively more important in highly fecund marine shellfish. The Pacific oyster, for example, shows hybrid vigor (heterosis) for yield (Fig. 12.4) that is as dramatic as that in maize (Shull 1908; Crow 1998), particularly since it emerges, not from crosses among major land races, but from crosses among partially inbred lines derived from a single wild population (Hedgecock and Davis 2007). Dramatic heterosis for yield in oysters is associated with equally dramatic levels of inbreeding depression (Evans et al. 2004), which results from a large load of deleterious recessive mutations (Bierne et al. 1998; Launey and Hedgecock 2001). A large mutational load in shellfish is consistent with the

344

Shellfish Aquaculture and the Environment

% Difference in Yield from Industry Controls + 1 SD

275% 250% 225% 200%

177%

175% 134%

150% 125%

98%

105%

100%

100% 75% Inbred

Wild

MBP Average All Families

MBP Top Five Families

Industry Controls (100%)

Figure 12.3 Yield of families after two generations of selection by the Molluscan Broodstock Program (MBP) (Langdon et al. 2003; C. Langdon, pers. comm.), compared with yields from inbred families, wild families, and industry controls, to which all values are scaled. The average of all MBP selected families and of the top five families suggest that substantial improvements in yield can be achieved with family selection.

hypothesis that heterosis results from dominance of wild type over deleterious recessive alleles in hybrids (Crow 1998). A large mutational load was also predicted by G. C. Williams’ (1975) elm-oyster model for the advantages of sexual reproduction in species with high fecundity and high early mortality. Since high fecundity and high early mortality are the dominant life history features among marine fish (Winemiller and Rose 1992) and invertebrates (Thorson 1950), we might expect considerable scope for genetic improvement of shellfish to come from crossbreeding of inbred lines derived from natural populations of such species.

Commercial crossbreeding for higher shellfish yield A detailed analysis of four factorial crosses of inbred lines of the Pacific oyster showed that both additive and nonadditive components of variance are important contributors to oyster yield (Hedgecock and Davis 2007). Yield generally increased with the general combining ability (GCA) of inbred lines, a measure of

additive genetic variance, as expected from the response to family selection reported by Langdon et al. (2003). However, high-yielding hybrids with high, positive special combining ability (SCA), a measure of nonadditive genetic variance, and little GCA were also observed. The nonadditive genetic component was often the largest component of variance in yield, accounting for a remarkable 88–96% of yield variance at the seed stage in one cross. Even more remarkable in this study was the finding of large differences in yield between reciprocal hybrids (AB ≠ BA), which accounted for 21– 51% of yield variance and comprised maternal as well as nonmaternal components of variance. The bases of these significant reciprocal effects merit further study, but their existence clearly suggests that the direction of crossbreeding makes a substantial difference in yield. Shellfish breeding should clearly take advantage of both additive genetic variance, through selection among inbred lines, and nonadditive genetic variance, by identification of elite inbred lines for crossbreeding (Hedgecock and Davis 2007). A breeding scheme to accomplish these dual goals can be conceptualized in the

Genetics of shellfish on a human-dominated planet

345

(A)

(B) Adjusted mean seed weight (g)

700 600 500 400 300 200 100 0 7

×4

47

5

×3

35

ild

W

ild

W

5

×3

47

2

×9

92

7 2 5 2 7 ×4 ×9 ×3 ×4 ×9 92 35 92 35 47

Figure 12.4 Pacific oysters show dramatic growth heterosis (hybrid vigor): (A) Inbred and hybrid Pacific oysters produced by a factorial cross of two partially inbred lines, 6 and 7; (B) comparison of bulk seed weight (adjusted for numbers, ∼100, 90-day-old seed, and for initial weight) for top hybrids (cross-hatched bars), their elite inbred parents (vertically striped bars), and the wild stock typically cultured on the U.S. West Coast (solid bars). Five of six hybrids yield significantly more than the current stock cultured, with the best hybrid (47 × 92) yielding nearly twice what the current stock yields.

form of a biomass pyramid, at the top of which is an intense breeding program to develop and select among new inbred lines, to measure the crossbreeding potential of these lines, in order to identify elite lines for further commercial testing (Fig. 12.5). All these activi-

ties require minimal biomass but maximal value per individual. A broodstock oyster, which has resulted from at least two generations of breeding, with a pedigree verified by typing of molecular markers (see below), and which has been conditioned for spawning in a

346

Shellfish Aquaculture and the Environment

at In

fo

s

rm

as om

Propagation of elite inbred lines and validation of crosses at commercial scale

Bi

ion

Breeding Program

Commercial production

Figure 12.5 A conceptual breeding program for shellfish that captures both additive and nonadditive genetic variance for yield. At the top of the biomass pyramid, breeders develop inbred lines, select among them, cross them and test their hybrids, and make chemical triploids of elite hybrids. The middle tier of the program tests elite crosses on farms, collects yield data, propagates elite inbred lines, and makes and distributes high-yielding seed to production farms. Information flows up the pyramid; biomass increases going down the pyramid.

hatchery, may be worth hundreds to thousands of dollars. At the farm level, on the other hand, each oyster may be worth $0.25. To get from the research level to the production level requires an intermediate step for amplifying the numbers of inbred broodstock to that required for commercial spawns. Also, at this level, hybrid seed would be produced for largescale field trials and tracked and evaluated through the production system. A uniform nomenclature for lines and crosses is required to track hybrid stocks through the production system. Information but not broodstock would flow from the middle tier to the top tier of the biomass pyramid, affecting selection among inbred lines. A bioeconomic framework is essential but does not yet exist for determining the value of a commercial shellfish breeding program. Also required for any shellfish breeding program are genotyping to confirm pedigrees. Hedgecock and Davis (2007) reported that 10.5% of prospective parents for their crosses had multilocus, microsatellite marker genotypes that were incompatible with their parents; another 13.8% were excluded because

of missing information or uncertainty in the genotyping. Curole and Hedgecock (2007) report slightly lower rates of contamination (8.1%) and of uncertain or incomplete genotyping (6.6%) over a larger sampling period. These figures show the necessity of validating parent pedigrees for controlled crosses. Contamination of experimental bivalve populations has been reported previously (e.g., Mallet et al. 1985; Foltz 1986; Zouros et al. 1992; Li and Guo 2004) and must be confronted in all shellfish breeding and experimental research. As the oyster industry has shifted heavily in the past decade toward production of triploid oysters (oysters carrying three rather than the normal two sets of chromosomes; Nell 2002), a commercial crossbreeding program must also focus on production of triploid as well as diploid hybrid seed (Hedgecock and Davis 2007). Increasing production of triploids, which are effectively sterile (Allen and Downing 1986), is a welcome trend toward sustainability, as discussed in the next section, since it isolates genetically improved farmed stock from natural shellfish populations.

Genetics of shellfish on a human-dominated planet

Triploid seed is currently produced by fertilizing diploid eggs with sperm from tetraploid males (Guo et al. 1996). Existing tetraploid stocks of the Pacific oyster were derived haphazardly from a rather narrow genetic base of wild diploid oysters. To take full advantage of nonadditive genetic variance for yield, commercial shellfish breeding programs will need to build new tetraploid lines that incorporate general and specific combining abilities. This can be done by chemically inducing triploidy in eggs from diploid hybrid females fertilized with sperm from an unrelated inbred line and using the resulting three-line triploids and an additional unrelated inbred line to found fourline tetraploid stocks. The suggestion from the maize literature is that heterosis compounds with higher ploidy levels: just as the hybrid AB is better than inbred AA at the diploid level, ABC > AAB at the triploid level, and ABCD > AABC at the tetraploid level (Birchler et al. 2003). While methods for husbandry and breeding of bivalve molluscs have been steadily improving, genomic resources have also developed at a rapid rate. Development of genomic resources promises to accelerate discovery of phenotypicgenotypic associations, the genes underlying economically important traits, and methods for determining the breeding or crossbreeding values of broodstock at early larval stages (Pace et al. 2006; Hedgecock et al. 2007a).

Conservation The challenges in conserving while utilizing sustainably the planet’s imperiled aquatic biodiversity are enormous and global in nature (Jackson et al. 2001; Dulvy et al. 2003). Conservation of shellfish biodiversity is particularly challenging since diversity is highest in coastal and estuarine habitats, which are heavily impacted by human populations. To counteract or reverse the impacts of overfishing and habitat degradation, resource manag-

347

ers in the past have used introductions (see Chapter 14, in this book) or translocations as a tool for shellfish remediation. The ecological and genetic risks of these management options are well described but poorly quantified (NRC 2004). Today, those concerned with decimated shellfish populations are turning increasingly to restoration programs for native species, which often involve supplementing natural stocks of shellfish with hatchery-propagated seed. The genetic implications of allowing interaction of wild and hatchery-propagated shellfish populations have been considered (Allen and Hilbish 2000; Gaffney 2006) and are reviewed below.

Introductions of shellfish Coastal ecosystems are characterized as heavily invaded by nonindigenous species (Grosholz 2002). Most biological invasions are attributable to shipping, via the transport and exchange of ballast water, though purposeful, accidental, or inadvertent introductions of nonnative species, owing to aquaculture (Chapter 14) or the aquarium trade, have also caused profound changes in coastal ecosystems (Ruiz et al. 2000). Although introductions of nonnative shellfish into North America for aquaculture have been portrayed as a current threat (e.g., Naylor et al. 2001), such introductions are largely, if not entirely, historical events. For example, the eastern oyster was introduced into the San Francisco Bay from 1869 until 1940, when that particular “gateway for exotic species” closed forever (Miller et al. 2007). The difficulty of predicting the ecological consequences of shellfish introductions is well illustrated by the diverse effects that introductions of the Pacific oyster Crassostrea gigas have had on different continents and regions (Mann 1979; NRC 2004). MSX (multinucleated sphere unknown), a disease that caused epizootic mortalities of eastern oysters in the Chesapeake

348

Shellfish Aquaculture and the Environment

and Delaware Bays in the late 1950s and early 1960s and has now spread along the entire U.S. East Coast (NRC 2004), was introduced from Asia, where it infects Pacific oysters (Burreson et al. 2000); however, the means of MSX introduction—whether from illegal introduction of Pacific oysters for aquaculture, fouling of oysters on ship bottoms, or from ballast water—is unknown (NRC 2004). Purposeful introductions can be curtailed, in principle, with regulations and enforcement, but often these are lacking or insufficient to prevent some intentional or accidental introductions, as amply illustrated by the controversy over the proposed introduction of the Asian oyster Crassostrea ariakensis into the Chesapeake Bay (NRC 2004). The International Council for the Exploration of the Sea (ICES 2005) has developed a code of practice governing introductions and transfers of marine organisms, which is supposed to be enforced by its 20 member countries and to guide policy in nonmember countries. Currently, introductions of exotic shellfish and of interstate transfers of native and naturalized shellfish species are strictly regulated by state agencies along the U.S. West Coast. For example, a recent reintroduction of the Kumamoto oyster Crassostrea sikamea, a species that has been cultivated on the U.S. West Coast for six decades, required rearing in closed quarantine, with extensive disease testing, of both the adult oysters imported from Japan and their first-generation offspring (Camara et al. 2008), one generation beyond ICES codes of practice. While introduction of nonnative species raise large ecological concerns, their impact on genetic diversity of local species or of locally established conspecific populations is less well understood but is likely to be less important than their direct impact on ecosystems.

Translocations of shellfish Translocations of conspecific stocks from one area to another are problematic for disease

transmission and other ecological reasons (see Chapter 13 in this book; Hegaret et al. 2008; NRC 2004). Direct genetic implications of translocations depend on the amount of genetic divergence between local and translocated populations. Like other marine animals with planktonically dispersing larvae, marine bivalves tend to show minimal genetic divergence or high connectivity over oceanic basin scales (Hedgecock et al. 2007c), which makes it difficult to detect any translocation events. The eastern oyster is a notable exception, having a major genetic divergence between Gulf of Mexico and Atlantic populations (Buroker 1983; Reeb and Avise 1990; Karl and Avise 1992; Cunningham and Collins 1994; McDonald et al. 1996). Regional subpopulations in the northern and southern Atlantic areas, which have also been identified with molecular markers (Hoover and Gaffney 2005; Gaffney 2006), may correspond with physiological races identified earlier on the basis of a latitudinal gradient in spawning season (Loosanoff and Nomejko 1951; Barber et al. 1991). Still, genetic impacts from historical translocations of eastern oysters have yet to be documented. Milbury et al.’s (2004) study of recruitment from 4 million Gulf of Mexico eastern oysters intentionally planted in the Choptank River for restoration purposes detected only three of 3545 spat of Gulf origin. This number was smaller than, but not inconsistent with, the potential reproductive contributions of the planted oysters based on their survival and sex ratio and the size of the natural population. Alternatively, these three spat with a Gulf of Mexico genetic marker could have resulted from previous translocations, a hypothesis supported by detection of a South Atlantic genetic marker in 5% of the spat screened. Despite the difficulty of predicting impacts on local adaptation, any proposed shellfish translocation ought to be preceded, at least, by a determination of the population genetic structure of the target species (Bell et al. 2005; Ward 2006).

Genetics of shellfish on a human-dominated planet

Interest in shellfish restoration has grown substantially over the last decade or more, engendering, for example, the establishment by the Nature Conservancy of the Shellfish Restoration Network and the International Conference on Shellfish Restoration (ICSR) that has convened 11 times. Insofar as restoration projects commonly utilize translocation or hatchery enhancement as tools, they have genetic implications for the natural populations being restored. The genetic impacts of translocations have already been covered, so we turn attention, next, to the potential impacts of hatchery propagation on shellfish genetic diversity and fitness. We need, first, to review the reproductive biology of natural bivalve populations, in particular variance in reproductive success and its genetic consequences.

SRS of shellfish The majority of marine shellfish (and marine fish) share a suite of life history traits— relatively late maturation, high fecundity, small eggs, long-lasting and widely dispersing plankton-feeding larvae, and broad geographic ranges (Thorson 1950; Winemiller and Rose 1992; Palumbi and Hedgecock 2005)—that renders them more vulnerable to loss of variation and extinction than might be expected from their great abundance. These life history traits appear to be adaptations to a biphasic life cycle, in which larvae disperse to planktonic habitats, far from potentially cannibalistic adults, but face tremendously high, though variable, early mortality. Conservation of such species depends, therefore, not only on protection of adult forms but also on the preservation of a vast, poorly delimited and understood planktonic environment. High fecundity, on the order of a million or more eggs per female per spawning event, and early mortality, typically in the range of 10– 20% per day, make possible a high variance among individuals in reproductive success, the number of offspring contributed to the next

349

generation. Successful reproduction for most marine shellfish requires success at each step in a complex chain of events, from reproductive maturation and spawning of adults to external fertilization of gametes, development, growth and survival of larval forms, metamorphosis and recruitment into the adult habitat, and survival and growth of juveniles to maturity. Reproductive activity, though tuned to the annual seasonal cycle, must still match highly variable local conditions for this chain of events to be complete. The chances of successful reproduction may thus vary dramatically for individuals, perhaps even among individuals adjacent to one another in space but spawning at slightly different times. Consequently, reproductive success in marine organisms might, at times, resemble a sweepstakes lottery, in which there are a few big winners and many losers (Hedgecock 1994). The hypothesis of SRS, which makes testable predictions about temporal genetic change in populations and variance in the genetic composition of new recruits, has received support from both empirical (e.g., Li and Hedgecock 1998; Hauser et al. 2002; Turner et al. 2002; Lee and Boulding 2007, 2009; Hedgecock et al. 2007b) and theoretical studies (Waples 2002; Hedrick 2005; Eldon and Wakeley 2006; Sargsyan and Wakeley 2008). The implication of the SRS hypothesis for conservation is that these seemingly inexhaustible living marine resources may have effective population sizes that are orders of magnitude smaller than census sizes, and thus rates of genetic drift and inbreeding that can erode biodiversity on ecological timescales.

Interaction of hatchery-propagated and natural shellfish populations Adverse interactions of wild and hatcherypropagated stocks are likely growing with the global expansion of aquaculture (McGinnity et al. 2003; Hindar et al. 2006) and stock enhancement programs, including shellfish

350

Shellfish Aquaculture and the Environment

restoration efforts (Born et al. 2004; Gaffney 2006). High fecundity creates the risk that reproduction and spread of hatcherypropagated shellfish stocks will dilute the genetic diversity of wild populations. Ryman and Laikre (1991) provided a model for this problem, in which the effective size of a population comprising both naturally (wild) and artificially (hatchery or captive) propagated components is given by 1 x 2 (1 − x ) = + , N e Nc Nw 2

(12.1)

where Ne is the effective size of the mixed population, Nc is the effective size of the captive or hatchery population, Nw is the effective size of the wild component, and x is the proportion of the spawning population of hatchery origin.

Hatchery effects on genetic diversity and adaptedness For highly fecund shellfish, the risk that an enhancement program might dilute genetic diversity and reduce the effective size of a wild population is potentially great, but there are few cases in which a genetic impact has been rigorously quantified (Gaffney 2006). Initial enthusiasm for the intuitive simplicity of the Ryman–Laikre model can give way to skepticism about its application, when difficulties or uncertainties in measuring its parameters are fully appreciated. Nc is knowable, certainly, given current capabilities for high-throughput analysis of genetic markers and statistical tools for estimating the effective number of hatchery parents; this parameter can and should be determined. The other two parameters, x and Nw, are much more difficult to quantify, however, since they depend on information about the natural population and on proper spatial scales of measurement. One temptation is to estimate x, for example, by estimating the proportion of seed that are of hatchery origin;

such an estimate is limited by uncertainty in estimating wild seed production at an appropriate spatial scale. Moreover, since x should be determined at spawning, the relative rates of wild and hatchery seed survival to reproduction need to be known. Another genetic risk of hatchery-based shellfish restoration programs is the effect on fitness or adaptedness of natural populations. A hatchery stock is almost inevitably subject to intentional or unintentional artificial selection (“domestication” selection) in the hatchery environment. For example, fine-mesh screens are used universally in shellfish hatcheries to cull small individuals from larval cultures. This practice could select for size-at-age and therefore rapid larval development. If this trait were maladaptive in natural environments and if, through a restoration program, a hatchery stock were to swamp a local population, then the reproductive success of the “enhanced” population could, in principle, be reduced. Unfortunately, there are almost no data on the genetic impacts of such hatchery practices; indeed, it would be challenging to measure genotype-by-environment interaction for larval traits across both hatchery and natural habitats. Nevertheless, the risk of domestication selection can be mitigated to a large extent by continual replacement of hatchery broodstock with wild adults and exclusion of hatchery-bred adults from the hatchery broodstock, which would prevent cumulative effects of domestication selection across generations. Risks from hatchery enhancements on genetic diversity or adaptation are manageable with appropriate designs and monitoring (Hedgecock and Coykendall 2007).

Restoration of disease-ravaged shellfish populations Heavy disease pressure, such as experienced in the Chesapeake and Delaware Bays, poses further challenges for a traditional

Genetics of shellfish on a human-dominated planet

hatchery-based approach to shellfish restoration (Mann and Powell 2007). Planting disease-susceptible hatchery seed is unlikely to increase the adult population of oysters, except in low-salinity refuges where disease pressure is weak or absent. Still, hatcheries could conceivably play a role in restoration of natural populations by seeding stock selected for disease resistance or disease tolerance, a concept termed genetic rehabilitation (Allen and Hilbish 2000). The goal of genetic rehabilitation is not to enhance populations directly but to increase the frequencies of diseaseresistant or disease-tolerant genotypes to the point at which natural selection can amplify and spread them further. At the same time, however, the effective size of the selected stock needs to be large enough to minimize inbreeding depression, a decline in fitness that might offset any advantage of disease tolerance. Unfortunately, closed shellfish populations, including some selected for disease resistance, tend to have small effective sizes and inbreeding depression (Vrijenhoek et al. 1990; Hedgecock et al. 1992; Carlsson et al. 2006). Genetic analyses of juvenile oysters in the Great Wicomico River, Virginia, showed that less than 10% came from a diseaseresistant line that had been planted there (Hare et al. 2006); in a second genetic study of hatchery-supplemented Virginia populations, no significant contribution of selectively bred oysters was observed over a 4-year period (Carlsson et al. 2008). Thus, the fitness of selectively bred oysters in the natural environment must be taken into account in assessing the viability of a genetic rehabilitation strategy.

Geoduck aquaculture: a case history The risk of interaction between wild and hatchery-propagated farmed stocks has been raised as an issue in the culture of geoduck clams (Panopea generosa, formerly Panopea abrupta; Vadopalas et al. 2010) in the U.S.

351

Pacific Northwest. Washington State House Bill (HB) 2220, passed in April 2007, directed the Washington Sea Grant College Program to measure and assess, among other things, “[g] enetic interactions between cultured and wild geoduck, including measurements of differences between cultured geoducks and wild geoducks in terms of genetics and reproductive status.” Lack of demonstrable population genetic structure for geoduck (Vadopalas et al. 2004; Miller et al. 2006), as in most marine bivalves, suggests that mixing of divergent gene pools through translocation of farmed stock is unlikely to be a great risk. Dilution of genetic diversity or affects of intentional or inadvertent selection on cultured clams are potential issues, but several factors appear to diminish these risks. Current hatchery practices—to collect local broodstock, to rotate broodstock within spawning seasons and between years, and to avoid using cultured geoduck as broodstock—eliminate the possibility of cumulative genetic change from domestication selection and reduce the risk of random genetic changes from use or reuse of too few broodstock. What part of geoduck reproduction in the Puget Sound might be attributable to spawning of hatchery-propagated stocks (x in the Ryman–Laikre model) is unknown but likely small for the foreseeable future. Presently, geoduck aquaculture occupies a little more than 140 ha of intertidal area in Washington State and could grow to as much as 245 ha, which would be 1.4% of the nearly 18,000 ha of subtidal geoduck beds available for commercial harvest in Washington State. Relative to the reproductive output of an exploitable geoduck biomass of 73,848 t in 1999 (Sizemore and Ulrich 1999), spawning by a few hundred metric tons of farmed geoduck (annual farm harvest is ∼400 t in Washington State) is unlikely to be a significant contribution. Moreover, farmed geoduck clams are harvested at 5–7 years of age, when they are

352

Shellfish Aquaculture and the Environment

just reaching reproductive maturity and are mostly male (Straus et al. 2008). The age and size of farmed geoducks at harvest, about equal to the minimum size of clams in the exploitable biomass, is well below the size or age at which the clams reach maximum fecundity. While it seems improbable that geoduck aquaculture could have a measurable impact on natural diversity, precautionary research on the interaction of wild and farmed geoduck clams is underway with funding from HB 2220.

Conclusions

Sterilizing farmed shellfish stocks through triploidy

Literature cited

One way to eliminate much of the risk of interaction between wild and hatchery stocks is to render farmed stocks sterile. Triploidy is commonly induced in shellfish to reduce reproductive effort, divert energy to growth, and improve meat quality during the normal spawning season (Allen and Downing 1986; Nell 2002; Vadopalas and Davis 2004). Because triploids are effectively sterile, their use in shellfish aquaculture dramatically reduces the risk of spawning and mixing with local native or naturalized stocks. At the same time, triploidy offers only a temporary reduction in the risk of an introduction, if the farmed species is a nonnative (NRC 2004). Gene knockout technology offers another route to sterilization (Grewe et al. 2007; Wong and Van Eenennaam 2008), although widespread public resistance to genetically modified organisms is likely to make this a nonviable strategy. The ability to culture sterile stocks frees aquaculture to pursue the benefits of domestication while minimizing or preventing the interactions of farmed and wild stocks. Biosecurity of reproductively competent tetraploid stocks in the environment is just beginning to be addressed (Piferrer et al. 2009); early experience with tetraploid Pacific oysters suggests that they are not robust enough, at present, to outcompete diploid stocks.

Allen, S.K., and Downing, S.L. 1986. Performance of triploid Pacific oysters, Crassostrea gigas (Thunberg). 1. Survival, growth, glycogen-content, and sexual-maturation in yearlings. Journal of Experimental Marine Biology and Ecology 102:197–208. Allen, S.K., and Hilbish, T.J. 2000. Genetic Considerations for Hatchery-Based Restoration of Oyster Reefs. Workshop summary, September 21–22, 2000. Virginia Institute of Marine Science, Gloucester Point, VA. Balon, E.K. 1995. Origin and domestication of the wild carp, Cyprinus carpio—from Roman gourmets to the swimming flowers. Aquaculture 129:3–48. Balon, E.K. 2004. About the oldest domesticates among fishes. Journal of Fish Biology 65(Suppl. A):1–27. Barber, B.J., Ford, S.E., and Wargo, R.N. 1991. Genetic variation in the timing of gonadal maturation and spawning of the eastern oyster, Crassostrea virginica (Gmelin). The Biological Bulletin 181:216–221. Bell, J.D., Rothlisberg, P.C., Munro, J.L., Loneragan, N.R., Nash, W.J., Ward, R.D., and Andrew, N.L. 2005. Advances in Marine Biology 49, Restocking and Stock Enhancement of Marine Invertebrate Fisheries. Academic Press, San Diego, CA. Bentsen, H.B., Eknath, A.E., Palada-de Vera, M.S., Danting, J.C., Bolivar, H.L., Reyes, R.A., Dionisio, E.E., Longalong, F.M., Circa, A.V., Tayamen, M.M., and Gjerde, B. 1998. Genetic improvement of farmed tilapias: growth perfor-

Long-term research on developing and improving domesticated shellfish stocks is needed to make shellfish farming more efficient and to relieve fishing pressure on natural populations. Such research should be coupled with research on reducing or eliminating interactions between wild and farmed populations (e.g., by inducing triploidy in hatchery-propagated stocks).

Genetics of shellfish on a human-dominated planet

mance in a complete diallel cross experiment with eight strains of Oreochromis niloticus. Aquaculture 160:145–173. Bierne, N., Launey, S., Naciri-Graven, Y., and Bonhomme, F. 1998. Early effect of inbreeding as revealed by microsatellite analyses on Ostrea edulis larvae. Genetics 148:1893–1906. Birchler, J.A., Auger, D.L., and Riddle, N.C. 2003. In search of the molecular basis of heterosis. The Plant Cell 15:2236–2239. Born, A.F., Immink, A.J., and Bartley, D.M. 2004. Marine and coastal stocking: global status and information needs. In: Bartley, D.M., and Leber, K.M. (eds.), Marine Ranching. FAO Fisheries Technical Paper. No. 429. FAO, Rome, pp. 1– 12. www.fao.org/docrep/008/y4783e/y4783e00. htm Boudry, P., Collet, B., Cornette, F., Hervouet, V., and Bonhomme, F. 2002. High variance in reproductive success of the Pacific oyster (Crassostrea gigas, Thunberg) revealed by microsatellite-based parentage analysis of multifactorial crosses. Aquaculture 204:283–296. Buck, B.H. 2007. Experimental trials on the feasibility of offshore seed production of the mussel Mytilus edulis in the German Bight: installation, technical requirements and environmental conditions. Helgoland Marine Research 61:87– 101. Buroker, N.E. 1983. Population genetics of the American oyster Crassostrea virginica along the Atlantic coast and the Gulf of Mexico. Marine Biology 75:99–112. Burreson, E.M., Stokes, N.A., and Friedman, C.S. 2000. Increased virulence in an introduced pathogen: Haplosporidium nelsoni (MSX) in the eastern oyster Crassostrea virginica. Journal of Aquatic Animal Health 12:1–8. Camara, M.D., Davis, J.P., Sekino, M., Hedgecock, D., Li, G., Langdon, C.J., and Evans, S. 2008. The Kumamoto oyster Crassostrea sikamea is neither rare nor threatened by hybridization in the northern Ariake Sea, Japan. Journal of Shellfish Research 27:313–322. Carlsson, J., Carnegie, R.B., Cordes, J.F., Hare, M.P., Leggett, A.T., and Reece, K.S. 2008. Evaluating recruitment contribution of a selectively bred aquaculture line of the oyster, Crassostrea virginica used in restoration efforts. Journal of Shellfish Research 27:1117–1124.

353

Carlsson, J., Morrison, C.L., and Reece, K.S. 2006. Wild and aquaculture populations of the eastern oyster compared using microsatellites. Journal of Heredity 97:595–598. Clutton-Brock, J. 1981. Domesticated Animals from Early Times. University of Texas Press, Austin, TX. Crow, J.F. 1998. 90 Years ago: the beginning of hybrid maize. Genetics 148:923–928. Cunningham, C.W., and Collins, T.M. 1994. Developing model systems for molecular biogeography: vicariance and interchange in marine invertebrates. In: Schierwater, B., Streit, B., Wagner, G.P., and DeSalle, R. (eds.), Molecular Ecology and Evolution: Approaches and Applications. Birkhauser Verlag, Basel, pp. 405–433. Cunningham, C., Hikima, J.I., Jenny, M.J., Chapman, R.W., Fang, G.C., Saski, C., Lundqvist, M.L., Wing, R.A., Cupit, P.M., Gross, P.S., Warr, G.W., and Tomkins, J.P. 2006. New resources for marine genomics: bacterial artificial chromosome libraries for the eastern and Pacific oysters (Crassostrea virginica and C. gigas). Marine Biotechnology 8:521–533. Curole, J.P., and Hedgecock, D. 2007. Chapter 29. Bivalve genomics: complications, challenges, and future perspectives. In: Liu, Z. (ed.), Aquaculture Genome Technologies. Blackwell Publishing, Ames, IA, pp. 525–543. David, P. 1998. Heterozygosity-fitness correlations: new perspectives on old problems. Heredity 80:531–537. Dégremont, L., Ernande, B., Bedier, E., and Boudry, P. 2007. Summer mortality of hatchery-produced Pacific oyster spat (Crassostrea gigas). I. Estimation of genetic parameters for survival and growth. Aquaculture 262:41–53. Diamond, J. 1997. Guns, Germs, and Steel. WW Norton & Company, New York. Duarte, C.M., Marba, N., and Holmer, M. 2007. Rapid domestication of marine species. Science 316:382–383. Dulvy, N.K., Sadovy, Y., and Reynolds, J.D. 2003. Extinction vulnerability in marine populations. Fish and Fisheries 4:25–64. Eldon, B., and Wakeley, J. 2006. Coalescent processes when the distribution of offspring number among individuals is highly skewed. Genetics 172:2621–2633.

354

Shellfish Aquaculture and the Environment

Evans, F., Matson, S., Brake, J., and Langdon, C. 2004. The effects of inbreeding on performance traits of adult Pacific oysters (Crassostrea gigas). Aquaculture 230:89–98. FAO (Food and Agriculture Organization of the United Nations). 2009a. The State of World Fisheries and Aquaculture (SOFIA) 2008. FAO, Rome.www.fao.org/docrep/009/A0699e/A0699 e00.htm FAO (Food and Agriculture Organization of the United Nations) Fisheries Department, Fishery Information, Data, and Statistics Unit. 2009b. Aquaculture database release February 23, 2009, analyzed with FISHSTAT+ Version 2.32, 2000. Foltz, D.W. 1986. Null alleles as a possible cause of heterozygote deficiencies in the oyster Crassostrea virginica and other bivalves. Evolution 40:869–870. Fujio, Y. 1982. A correlation of heterozygosity with growth rate in the Pacific oyster, Crassostrea gigas. Tohuku Journal of Agricultural Research 33:66–75. Gaffney, P.M. 1994. Heteosis and heterozygote deficiencies in marine bivalves: more light? In: Beaumont, A.R. (ed.), Genetics and Evolution of Aquatic Organisms. Chapman & Hall, London, pp. 146–153. Gaffney, P.M. 1996. Biochemical and population genetics. In: Kennedy, V.S., Newell, R.I.E., and Eble, A.F. (eds.), The Eastern Oyster. Maryland Sea Grant College, University of Maryland, College Park, MD, pp. 423–441. Gaffney, P.M. 2006. The role of genetics in shellfish restoration. Aquatic Living Resources 19:277– 282. Gaffney, P.M. 2008. A BAC-based physical map for the Pacific oyster genome. Journal of Shellfish Research 27:1009. Gaffney, P.M., Bernat, C.M., and Allen, S.K. 1993. Gametic incompatibility in wild and cultured populations of the eastern oyster, Crassostrea virginica (Gmelin). Aquaculture 115:273–284. Gjedrem, T. 2000. Genetic improvement of coldwater fish species. Aquaculture Research 31: 25–33. Gosling, E. 2003. Bivalve Molluscs: Biology, Ecology and Culture. Fishing News Books, Blackwell Publishing, Oxford. Grewe, P.M., Patil, J.G., McGoldrick, D.J., Rothlisberg, P.C., Whyard, S., Hinds, L.A.,

Hardy, C.M., Vignarajan, S., and Thresher, R.E. 2007. Preventing genetic pollution and the establishment of feral populations: a molecular solution. In: Bert, T.M. (ed.), Ecological and Genetic Implications of Aquaculture Activities. Springer, Dordrecht, pp. 103–114. Grosholz, E. 2002. Ecological and evolutionary consequences of coastal invasions. Trends in Ecology & Evolution 17:22–27. Günther, R.T. 1897. The oyster culture of the ancient Romans. Journal of the Marine Biological Association of the United Kingdom 4:360–365. Guo, X.M., DeBrosse, G.A., and Allen, S.K. 1996. All-triploid Pacific oysters (Crassostrea gigas Thunberg) produced by mating tetraploids and diploids. Aquaculture 142:149–161. Hare, M.P., Allen, S.K., Bloomer, P., Camara, M.D., Carnegie, R.B., Murfree, J., Luckenbach, M., Meritt, D., Morrison, C., Paynter, K., Reece, K.S., and Rose, C.G. 2006. A genetic test for recruitment enhancement in Chesapeake Bay oysters, Crassostrea virginica, after population supplementation with a disease tolerant strain. Conservation Genetics 7:717–734. Harlan, J.R. 1995. The Living Fields. Cambridge University Press, Cambridge. Harris, D.R., and Hilman, G.C. 1989. Foraging and Farming: The Evolution of Plant Exploitation. Unwin Hyman, London. Hauser, L., Adcock, G.J., Smith, P.J., Bernal, J.H., and Carvalho, G.R. 2002. Loss of microsatellite diversity and low effective population size in an overexploited population of New Zealand snapper (Pagrus auratus). Proceedings of the National Academy of Science USA 99:11724– 11747. Hedgecock, D. 1994. Does variance in reproductive success limit effective population sizes of marine organisms? Beaumont, A.R. (ed.), Genetics and Evolution of Aquatic Organisms. Chapman & Hall, London, pp. 122–134. Hedgecock, D., and Davis, J.P. 2007. Heterosis for yield and crossbreeding of the Pacific oyster Crassostrea gigas. Aquaculture 272S1:S17–S29. Hedgecock, D., Chow, V., and Waples, R. 1992. Effective population numbers of shellfish broodstocks estimated from temporal variance in allelic frequencies. Aquaculture 108:215–232. Hedgecock, D., and Coykendall, K. 2007. Genetic risks of hatchery enhancement: the good, the

Genetics of shellfish on a human-dominated planet

bad, and the unknown. In: Bert, T.M. (ed.), Ecological and Genetic Implications of Aquaculture Activities. Springer, Dordrecht, pp. 85–101. Hedgecock, D., Gaffney, P.M., Goulletquer, P., Guo, X., Reece, K., and Warr, G.W. 2005. The case for sequencing the Pacific oyster genome. Journal of Shellfish Research 24:429–441. Hedgecock, D., Lin, J.Z., DeCola, S., Haudenschild, C.D., Meyer, E., Manahan, D.T., and Bowen, B. 2007a. Transcriptomic analysis of growth heterosis in larval Pacific oysters (Crassostrea gigas). Proceedings of the National Academy of Science, USA 104:2313–2318. Hedgecock, D., Launey, S., Pudovkin, A.I., Naciri, Y., S. Lapègue, and Bonhomme F. 2007b. Small effective number of parents (Nb) inferred for a naturally spawned cohort of juvenile European flat oysters Ostrea edulis. Marine Biology 150:1173–1182. Hedgecock, D., Edmands, S., and Barber, P. 2007c. Genetic approaches to measuring connectivity. Oceanography 20:70–79. Hedrick, P. 2005. Large variance in reproductive success and the Ne/N ratio. Evolution 59: 1596–1599. Hegaret, H., Shumway, S.E., Wikfors, G.H., Pate, S., and Burkholder, J.M. 2008. Potential transport of harmful algae via relocation of bivalve molluscs. Marine Ecology Progress Series 361:169–179. Hindar, K., Fleming, I.A., McGinnity, P., and Diserud, A. 2006. Genetic and ecological effects of salmon farming on wild salmon: modelling from experimental results. International Council for the Exploration of the Sea Journal of Marine Science 63:1234–1247. Hoover, C.A., and Gaffney, P.M. 2005. Geographic variation in nuclear genes of the eastern oyster, Crassostrea virginica Gmelin. Journal of Shellfish Research 24:103–112. ICES (International Council for Exploration of the Sea). 2005. ICES Code of Practice on the Introductions and Transfers of Marine Organisms 2005. 30 pp. Copenhagen: ICES. Jackson, J.B.C., Kirby, M.X., Berger, W.H., Bjorndal, K.A., Botsford, L.W., Bourque, B.J., Bradbury, R.H., Cooke, R., Erlandson, J., Estes, J.A., Hughes, T.P., Kidwell, S., Lange, C.B., Lenihan, H.S., Pandolfi, J.M., Peterson, C.H.,

355

Steneck, R.S., Tegner, M.J., and Warner, R.R. 2001. Historical overfishing and the recent collapse of coastal ecosystems. Science 293: 629–638. Karl, S.A., and Avise, J.C. 1992. Balancing selection at allozyme loci in oysters—implications from nuclear RFLPs. Science 256:100–102. Koehn, R.K., Milkman, R., and Mitton, J.B. 1976. Population genetics of marine pelecypods. 4. Selection, migration and genetic differentiation in blue mussel Mytilus edulis. Evolution 30:2–32. Langdon, C., Evans, F., Jacobson, D., and Blouin, M. 2003. Improved family yields of Pacific oysters Crassostrea gigas Thunberg derived from selected parents. Aquaculture 220:227–244. Lannan, J.E. 1980. Broodstock management of Crassostrea gigas. 3. Selective breeding for improved larval survival. Aquaculture 21: 347–351. Launey, S., and Hedgecock, D. 2001. High genetic load in the Pacific oyster. Genetics 159: 255–265. Lee, H.J., and Boulding, E.G. 2007. Mitochondrial DNA variation in space and time in the northeastern Pacific gastropod, Littorina keenae. Molecular Ecology 16:3084–3103. Lee, H.J., and Boulding, E.G. 2009. Spatial and temporal population genetic structure of four northeastern Pacific littorinid gastropods: the effect of mode of larval development on variation at one mitochondrial and two nuclear DNA markers. Molecular Ecology 18:2165–2184. Li, L., and Guo, X.M. 2004. AFLP-based genetic linkage maps of the Pacific oyster Crassostrea gigas Thunberg. Marine Biotechnology 6:26– 36. Li, G., and Hedgecock, D. 1998. Genetic heterogeneity detected by PCR-SSCP, among samples of larval Pacific oysters (Crassostrea gigas Thunberg), supports the hypothesis of large variance in reproductive success. Canadian Journal of Fisheries and Aquatic Sciences 55: 1025–1033. Loosanoff, V.L., and Nomejko, C.A. 1951. Existence of physiologically different races of oysters, Crassostrea virginica. Biological Bulletin 101:151–156. Mallet, A.L., Zouros, E., Gartner-Kepkay, K.E., Freeman, K.R., and Dickie, L.M. 1985. Larval

356

Shellfish Aquaculture and the Environment

viability and heterozygote deficiency in populations of marine bivalves—evidence from pair matings of mussels. Marine Biology 87: 165–172. Mann, R. (ed.). 1979. Exotic Species in Mariculture. The MIT Press, Cambridge. Mann, R., and Powell, E.N. 2007. Why oyster restoration goals in the Chesapeake Bay are not and probably cannot be achieved. Journal of Shellfish Research 26:905–917. McDonald, J.H., Verrelli, B.C., and Geyer, L.B. 1996. Lack of geographic variation in anonymous nuclear polymorphisms in the American oyster, Crassostrea virginica. Molecular Biology and Evolution 13:1114–1118. McGinnity, P., Prodohl, P., Ferguson, K., Hynes, R., Maoileidigh, N.O., Baker N., Cotter, D., O’Hea, B., Cooke, D., Rogan, G., Taggart, J., and Cross, T. 2003. Fitness reduction and potential extinction of wild populations of Atlantic salmon, Salmo salar, as a result of interactions with escaped farm salmon. Proceedings of the Royal Society of London Series B, Biological Sciences 270:2443–2450. Milbury, C.A., Meritt, D.W., Newell, R.I.E., and Gaffney, P.M. 2004. Mitochondrial DNA markers allow monitoring of oyster stock enhancement in the Chesapeake Bay. Marine Biology 145:351–359. Miller, K.M., Supernault, K.J., Li, S., and Withler, R. 2006. Population structure in two marine invertebrate species (Panopea abrupta and Strongylocentrotus franciscanus) targeted for aquaculture and enhancement in British Columbia. Journal of Shellfish Research 25:33–42. Miller, A.M., Ruiz, G.M., Minton, M.S., and Ambrose, R.F. 2007. Differentiating successful and failed molluscan invaders in estuarine ecosystems. Marine Ecology Progress Series 332:41–51. National Research Council (NRC). 2004. NonNative Oysters in the Chesapeake Bay. National Academies Press, Washington, D.C. Naylor, R.L., Williams, S.R., and Strong, D.R. 2001. Aquaculture—a gateway for exotic species. Science 294:1655–1656. Nell, J.A. 2002. Farming triploid oysters. Aquaculture 210:69–88. Newkirk, G.F. 1980. Review of the genetics and the potential for selective breeding of commer-

cially important bivalves. Aquaculture 19:209– 228. Pace, D.A., Marsh, A.G., Leong, P.K., Green, A.J., Hedgecock, D., and Manahan, D.T. 2006. Physiological bases of genetically determined variation in growth of marine invertebrate larvae: a study of growth heterosis in the bivalve Crassostrea gigas. Journal of Experimental Marine Biology and Ecology 335:188– 209. Palumbi, S.R., and Hedgecock, D. 2005. The life of the sea: implications of marine population biology to conservation policy. In: Norris, E.A., and Crowder, L.B. (eds.), Marine Conservation Biology. Island Press, Washington, D.C., pp. 33–46. Piferrer, F., Beaumont, A., Falguière, J.-C., Flajšhans, M., Haffray, P., and Colombo, L. 2009. Polyploid fish and shellfish: production, biology and applications to aquaculture for performance improvement and genetic containment. Aquaculture 293:125–156. Reeb, C.A., and Avise, J.C. 1990. A genetic discontinuity in a continuously distributed species— mitochondrial-DNA in the American oyster, Crassostrea virginica. Genetics 124:397– 406. Ruiz, G.M., Fofonoff, P.W., Carlton, J.T., Wonham, M.J., and Hines, A.H. 2000. Invasion of coastal marine communities in North America: apparent patterns, processes, and biases. Annual Review of Ecology and Systematics 31:481–531. Ryman, N.R., and Laikre, L. 1991. Effects of supportive breeding on the genetically effective population size. Conservation Biology 5:325– 329. Saavedra, C., and Bachere, E. 2006. Bivalve genomics. Aquaculture 256:1–14. Sargsyan, O., and Wakeley, J. 2008. A coalescent process with simultaneous multiple mergers for approximating the gene genealogies of many marine organisms. Theoretical Population Biology 74:104–114. Sheridan, A.K. 1997. Genetic improvement of oyster production—a critique. Aquaculture 153:165–179. Shull, G.H. 1908. The composition of a field of maize. American Breeders Association Reports 4:296–301. Sizemore, B., and Ulrich, M. 1999. 1999 Geoduck Atlas: Atlas of Major Geoduck Tracts of Puget

Genetics of shellfish on a human-dominated planet

Sound. Washington Department of Fish and Wildlife, Olympia. Stevens, C., Plew, D., Hartstein, N., and Fredriksson, D. 2008. The physics of open-water shellfish aquaculture. Aquacultural Engineering 38:145– 160. Straus, K.M., Crosson, L.M., and Vadopalas, B. 2008. Effects of geoduck aquaculture on the environment: a synthesis of current knowledge. Washington Sea Grant Program, Seattle, 64 p. www.wsg.washington.edu/research/geoduck/literature_review.html Tanguy, A., Bierne, N., Saavedra, C., Pina, B., Bachere, E., Kube, M., Bazin, E., Bonhomme, F., Boudry, P., Boulo, V., Boutet, I., Cancela, L., Dossat, C., Favrel, P., Huvet, A., Jarque, S., Jollivet, D., Klages, S., Lapegue, S., Leite, R., Moal, J., Moraga, D., Reinhardt, R., Samain, J.F., Zouros, E., and Canario, A. 2008. Increasing genomic information in bivalves through new EST collections in four species: development of new genetic markers for environmental studies and genome evolution. Gene 408:27–36. Thorson, G. 1950. Reproductive and larval ecology of marine bottom invertebrates. Biological Reviews 25:1–45. Turner, T.F., Wares, J.P., and Gold, J.R. 2002. Genetic effective size is three orders of magnitude smaller than adult census size in an abundant, estuarine-dependent marine fish (Sciaenops ocellatus). Genetics 162:1329– 1339. Vadopalas, B., and Davis, J. 2004. Optimal chemical triploid induction in geoduck clams, Panopea abrupta (Conrad, 1849), by 6-dimethylaminopurine. Aquaculture 230:29–40. Vadopalas, B., Leclair, L.L., and Bentzen, P. 2004. Microsatellite and allozyme analyses reveal few genetic differences among spatially distinct aggregations of geoduck clams (Panopea abrupta, Conrad 1849). Journal of Shellfish Research 23:693–706.

357

Vadopalas, B., Pietsch, T.W., and Friedman, C.S. 2010. The proper name for the geoduck: resurrection of Panopea generosa Gould, 1850, from the synonymy of Panopea abrupta (Conrad, 1849) (Bivalvia: Myoida: Hiatellidae). Malacalogia 52(1):169–173. Vrijenhoek, R.C., Ford, S.E., and Haskin, H.H. 1990. Maintenance of heterozygosity during selective breeding of oysters for resistance to MSX disease. Journal of Heredity 81:418–423. Waples, R.S. 2002. Evaluating the effect of stagespecific survivorship on the Ne/N ratio. Molecular Ecology 11:1029–1037. Ward, R.D. 2006. The importance of identifying spatial population structure in restocking and stock enhancement programmes. Fisheries Research 80(1):9–18. Williams, G.C. 1975. Sex and Evolution. Princeton University Press, Princeton. Winemiller, K.O., and Rose, K.A. 1992. Patterns of life-history diversification in North American fishes: implications for population regulation. Canadian Journal of Fisheries and Aquatic Sciences 49:2196–2218. Wohlfarth, G.W. 1993. Heterosis for growth-rate in common carp. Aquaculture 113:31–46. Wong, A.C., and Van Eenennaam, A.L. 2008. Transgenic approaches for the reproductive containment of genetically engineered fish. Aquaculture 275:1–12. Zouros, E., and Pogson, G.H. 1994. The present status of the relationship between heterozygosity and heterosis. In: Beaumont, A.R. (ed.), Genetics and Evolution of Aquatic Organisms. Chapman & Hall, London, pp. 146–153. Zouros, E., Singh, S.M., and Miles, H.E. 1980. Growth-rate in oysters—an overdominant phenotype and its possible explanations. Evolution 34:856–867. Zouros, E., Freeman, K.R., Ball, A.O., and Pogson, G.H. 1992. Direct evidence for extensive paternal mitochondrial-DNA inheritance in the marine mussel Mytilus. Nature 359:412–414.

Chapter 13

Shellfish diseases and health management Ralph A. Elston and Susan E. Ford

Shellfish health management and infectious disease prevention The interactions between the environment and infectious diseases of cultured shellfish are many and varied. The natural environment is the source of infectious agents and provides external forces such as temperature and salinity that modify the host–parasite relationship. The aquaculture system is a special environment itself, and one for which management of shellfish health must be learned and practiced. Shellfish infectious disease agents can be transferred geographically by human activities and also through natural pathways. In addition, shellfish diseases can emerge from endemic or resident pathogens after a

change in environmental conditions or host susceptibility. The following sections illustrate the interactions of the natural and shellfish aquaculture environments with infectious disease agents using selected case studies and then discuss solutions for maintaining and enhancing shellfish disease prevention and health management. We provide a detailed discussion of health management in regard to shellfish transfers that are used to facilitate stock seeding or augmentation for harvest of edible shellfish because such transfers can be a mechanism for the movement of shellfish infectious disease agents. Intercontinental or interocean transfers of shellfish have demonstrated that severe negative effects can result from introducing

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 359

Shellfish Aquaculture and the Environment

l ca gi

Drivers and eff ec nd interactions a to s in rs a ch

ate lim :c

Physiological homeostasis Microorganism

tivity n ac a m hu

Organism

: rs

Immunity and stress response

al physical forc es natur and

Effec tor s: bio lo

360

Dr ive Figure 13.1 Natural and human forces directly affect shellfish health and also drive a complex web of physical and biological effectors that impinge on the shellfish, its physiological equilibrium, its immunity and stress response, and on its associated microorganisms.

infectious agents along with shellfish stocks (Chapter 14 in this book), and that these can affect wild as well as cultured shellfish. On the other hand, production of seed from shellfish hatcheries and nurseries has become a reliable, efficient, and necessary means of providing sustainable edible shellfish harvests on a worldwide basis. While inherent risk of transfer of infectious shellfish diseases cannot be absolutely avoided in the practice of shellfish aquaculture, risk from such transfers can and should be greatly reduced by application of health management principles. Additionally, culture of shellfish can become even more efficient than it has been over the last half century with the application of health management principles that have been initiated for shellfish aquaculture. Such measures will minimize the potential dangers not only to aquacultured

species but also to wild stocks in the vicinity of culture operations.

Interactions of bivalve shellfish and parasites with the natural environment Figure 13.1 shows a diagrammatic representation of natural and human forces acting as root causes or “drivers” of changes in shellfish health and disease status. These drivers can change the characteristics of the physical environment and thus its ability to maintain a healthy condition. They can also cause direct introduction of infectious shellfish disease agents such as bacteria, viruses, or parasites from one geographic area to another, directly affecting the shellfish, the microorganism, or

Shellfish diseases and health management 361

both. Alternatively and more commonly, physical and climatic drivers act through a complex chain or web of “effectors,” which means that the natural and human-caused forces act indirectly to start a cascade of physical or biological events that affect the health of the shellfish. For example, unusually warm ocean water, as a result of cyclic or other climatic alterations, may change ocean chemistry and may also indirectly affect the shellfish by altering conditions that favor the proliferation of opportunistically pathogenic bacteria, such as is believed to have occurred on the Pacific coast of the United States in 2007 (Elston et al. 2008). Temperature and salinity (salt content) of ocean and estuarine water are two of the better understood influences on shellfish disease as will be discussed in case examples, but there are many other less well-understood physical and water chemistry factors that affect the health and infectious disease status of shellfish.

Seasonal cycles influenced by temperature In general, parasite infection cycles in bivalve molluscs follow seasonal temperature cycles, the distribution of parasites within estuaries is influenced by salinity, and the geographic range of parasites is largely constrained by local temperature regimes. Organisms in the genera Haplosporidium, Perkinsus, Marteilia, and Bonamia, which parasitize oysters, clams, and mussels, typically infect their hosts, and proliferate in them most rapidly, during the warm months (Ford and Haskin 1982; Montes et al. 1991; Burreson and Ragone Calvo 1996; Villalba et al. 2005). Regression during the colder part of the year, caused by the death of parasites or of heavily infected hosts, or both, followed by renewed proliferation and reinfection as the temperature warms again, produces distinct seasonal cycles (Fig. 13.2) (Culloty and Mulcahy 1996; Oliver et al. 1998;

Carnegie et al. 2008; Tun et al. 2008). Mortality, with a certain lag time, typically follows the infection pattern (Fig. 13.3). The temperature response of a diseasecausing organism may be inversely related (e.g., higher disease rates at lower temperatures), or complex with specific peaks of growth responding to specific temperatures (Elston et al. 1982). Infection of the Pacific oyster with the protozoan parasite Mikrocytos mackini is an example of a disease that is exacerbated at colder temperatures. It was reported that the disease requires prolonged exposure of oysters to a low temperature (10°C) for development of a disease condition (Hervio et al. 1996). In lower latitudes near the edge of the range of the parasite, disease prevalence and severity are extremely low and the disease is of no biological consequence to aquaculture because temperatures warmer than 15°C prevent the development of Mikrocytos mackini disease(Bower et al. 1997). Thus, this disease is also subject to shifts in prevalence and severity, but the magnitude of these parameters will decrease with increasing temperature.

Local distribution of parasites influenced by salinity Marine and estuarine parasites, like their hosts, are sensitive to salinity. Interactions between salinity and two important parasites of the eastern oyster, Crassostrea virginica, have been well documented both in vitro and in vivo. The eastern oyster, an estuarine organism, lives over a salinity range of about 5–30 PSU. Haplosporidium nelsoni, which causes MSX disease in this species, is strongly inhibited by low salinity and rarely survives below 10 PSU (Ford 1985). Perkinsus marinus, which is responsible for dermo disease in the same host, is also inhibited by low salinity, although to a much lesser extent (Fig. 13.4), and can survive exposure in vivo at 3 PSU

Shellfish Aquaculture and the Environment

70

B

60

30

B

(A)

B J

J

B

J

50

B J B

B B

J

40

J B

J

20 15

B J

30

J

20

B

J

B

J

10

5

B

J

J

0

100

B

Percent Prevalence

(B)

80

J J

J

B

B

B

B

B

J 60

J B

B

15

B J J

B

10

B 5

J J

B

B

J

0

60

30 (C)

50

J

J

J

J

J

B

B

Prevalence

J

Temperature

15

J B

B

B

B

10

10

J J

J 5 B

Apr

May

0 Jul

B

Jun

B Mar

Jan

Dec

Oct

Nov

Sep

Jul

Aug

B

Jun

B B

25 20

B

20

0

B J

J

J

30

B J

J

Feb

40

25 20

J

0

J

30

J

B

J

40 20

10

J

B 0

25

Temperature (°C)

362

Figure 13.2 Examples of water temperature-associated seasonal prevalence cycles for protozoan parasites of oysters. (A) Haplosporidium nelsoni (MSX) in eastern oysters in Delaware Bay, NJ; (B) Perkinsus marinus (dermo) in eastern oysters in Delaware Bay, NJ; (C) Marteiliodes chungnuenis infecting Pacific oysters in Okayama Prefecture, Japan. Marteiliodes chungnuenis data from Tun et al. (2008), with permission.

(Chu et al. 1993) Beginning in the nineteenth century, a practice of extensive aquaculture developed in Delaware and lower Chesapeake Bays, USA, which involved transplanting small seed oysters from natural beds in the upper,

low-salinity portions of the estuary to private growing and fattening grounds in the highersalinity regions (Ford 1997; MacKenzie 1997). After the initial outbreak of MSX disease in lower Delaware Bay in the late 1950s,

Shellfish diseases and health management 363

40

3 J B

Sample Intensity

J

Mortality

B

B J

Temperature

B

35 30

Sample Intensity

B 2

B

25

1.5

20 15

J

1

B

B

10

B J

0.5

5

B 0 Mar

J

Apr

May

J

J

J

Jun

Percent Mortality and Temperature (°C)

2.5

Jul

Aug

Sep

Oct

0 Nov

Figure 13.3 Example of the time delay between temperature, Perkinsus marinus (dermo) infection development, and mortality for a group of eastern oysters in Delaware Bay, NJ.

100 90

Percent Prevalence

80 70 60 50 40 30 20

MSX

10

Dermo

0 8

10

12

14

16

18

20

22

24

Mean Midtide Salinity Figure 13.4 Long-term mean prevalence of Haplosporidium nelsoni (MSX) (1958–1980) and Perkinsus marinus (dermo) (1990–2007) in eastern oysters in Delaware Bay, NJ.

the industry was able to persist because the low-salinity environment of the upper bay provided a refuge from disease that did not significantly diminish the supply of seed oysters. After Perkinsus marinus began causing oyster

deaths in the Delaware Bay in 1990, however, seed oysters were no longer protected from disease, and the practice of “planting” oysters gave way to direct marketing from the upper bay natural beds. Thus, the differing responses of these two pathogens to the same environmental gradient had vastly different consequences for the practice of extensive oyster culture. Many clams and mussels, as well as other oyster species are cultured in higher-salinity waters: ∼25 PSU to full-strength seawater (32– 35 PSU) and a similarly strong influence of salinity on the spatial distribution of infections by their parasites is less well documented, although Perkinsus olseni and Perkinsus chesapeaki, which infect a variety of molluscs, are all sensitive to low salinity when tested in vitro (La Peyre et al. 2006). A newly discovered species of Bonamia which infects the noncommercial cupped oyster Ostrea equestris and the experimentally introduced oyster Crassostrea ariakensis (see below) is clearly limited in its distribution by low salinity. Experimental field deployments indicated that Crassostrea

364

Shellfish Aquaculture and the Environment

ariakensis developed infections at sites with mean salinities of 28–33 PSU, but not at sites with lower salinities (Bishop et al. 2006). Previously infected Crassostrea ariakensis placed experimentally at 30, 20, and 10 PSU all lost infections, but parasites disappeared fastest at 10 and 20 PSU and histological examination of these two groups showed evidence of parasite degradation, suggesting “that low to moderate salinities detrimentally affected the parasite” (Audemard et al. 2008).

Geographic distribution of parasites influenced by temperature The geographic range of parasites is undoubtedly influenced by their tolerance for high or low temperatures. A good example is Quahog Parasite Unknown (QPX), a parasite of the hard clam, Mercenaria mercenaria, in northeastern North America. It has been found in clams from the Canadian Maritime provinces south to the mouth of Chesapeake Bay (Lyons et al. 2007) and is considered to be an opportunistic pathogen. It has not been reported from locations farther south even though hard clam culture is widespread along the southeastern coast of the United States. The hypothesis that high temperature inhibits the parasite in the southern United States is supported by evidence from laboratory experiments in which QPX was cultured in vitro at different temperatures. Both Brothers et al. (Brothers et al. 2000) and Perrigault et al. (2010) found that growth in culture was maximum in the 20–25°C range and decreased rapidly above 30°C. The high-temperature sensitivity may explain why hard clams from southern stocks become heavily infected when they are exposed in enzootic waters, whereas adjacent clams from northern stocks acquire few detectable infections (Ford et al. 2002; Ragone Calvo et al. 2007). Two hypotheses have been offered for this disparity, and in both, temperature is the major driver (Ragone Calvo

et al. 2007). These hypotheses are that (1) high temperatures restrict or prohibit the presence of QPX in the south and therefore southern clams have never experienced selective mortality and thus never developed genetically based resistance to QPX; and (2) southern clam stocks are not adapted to the colder temperatures of the northeast and mid-Atlantic and thus are physiologically disadvantaged in that environment, making them more susceptible to opportunistic disease agents of any kind. These hypotheses are not mutually exclusive, and will require focused testing to define their possible influence on QPX disease development in different stocks.

Temperature influences on the expression of disease Many pathogens remain at very low abundance—perhaps even undetectable—in bivalves until they are stimulated to proliferate by an environmental trigger. An example is the causative agent of withering syndrome in both wild and cultured abalone of several species on the U.S. West Coast. The organism, a Rickettsiales-like prokaryote (Candidatus Xenohaliotis californiensis) (Friedman et al. 2000) is strongly influenced by temperature and in the field, outbreaks of withering syndrome have been linked to ocean warming during El Niño cycles (Moore et al. 2000; Vilchis et al. 2005). Not only is transmission enhanced at elevated temperature (Braid et al. 2005) but cryptic or very low-intensity infections that cause no clinical signs can develop into lethal infections if the abalone is placed at high temperature (≥18°C) (Moore et al. 2001) (Fig. 13.5). Vibriosis is an opportunistic, but occasionally highly virulent, disease of shellfish larvae and juveniles. Outbreaks in shellfish hatcheries are often linked to conditions within the hatchery but increased ocean temperature and inshore nutrient enrichment by upwelling have

Shellfish diseases and health management 365

35 BB B

14.7 °C B

20

B B

15 10

60 Body Burden

40

Standard Hemolymph

20 BB

Aug

Apr May Jun Jul

Jan

Feb Mar

Weeks of Exposure

0 Nov Dec

J JJJJ JJJJ JJJJ JJJ JJJJ JJJJJJJJ JJJJ J 0 BBBBBBBBBBBBBBBBBBB 0 2 4 6 8 10 12 14 16 18 20 22 24 26 28 30 32

Sep Oct

5

BBB

80

Aug

J

18.5 °C

Jun Jul

25

B

100

Percent Prevalence

Percent Mortality

30

BBB

Figure 13.5 Induction of withering syndrome by elevated temperature. Cumulative mortality of red abalone Haliotis rufescens exposed to high (18.5°C) and low (14.7°C) temperatures for a period of 32 weeks. Both groups of abalone originated from a farm with no history of withering syndrome; only the elevated temperature group developed the withering syndrome signs and experienced associated mortality. (Redrawn from Moore et al. 2000.)

Figure 13.6 Comparison of methods for Perkinsus marinus (dermo) in eastern oysters. All methods employed incubation of oyster tissue in Ray’s fluid thioglycollate medium. Both the standard detection assay (pieces of mantle and rectum) and the hemolymph assay showed a pronounced dip in prevalence in late winter; however, the total body burden assay showed that most oysters remained infected during this period, although the abundance of parasites decreased. (Redrawn from Bushek et al. 1994.)

also been linked to a severe outbreak of the disease (Elston et al. 2008). Similarly, infection by oyster herpes virus (OsHV) disease requires a permissive environmental temperature greater than about 25– 26°C to cause productive infections and morbidity in hatchery reared oyster larvae (LeDeuff et al. 1996). Perkinsus marinus, the agent of dermo disease, is another good example of how environment can influence the expression of disease by inhibiting or by favoring proliferation of the parasite. In vitro experiments clearly demonstrate that the parasite replicates fastest at high temperature and salinity, and that its growth is inhibited at low temperatures and salinities (Dungan and Hamilton 1995). The prevalence of Perkinsus marinus has long been reported to decline in vivo over the winter and spring in the mid-Atlantic and northeastern United States (Andrews and Hewatt 1957; Ford and Smolowitz 2007); however, this observation is simply a manifestation of lessened infection intensity as parasites die, are

eliminated from the oyster, and can no longer reliably be detected by the traditional culture method (Bushek et al. 1994) (Fig. 13.6). Small numbers of parasites persist in most oysters and serve as foci for renewed proliferation when the temperature increases again in spring (Ragone Calvo and Burreson 1994). Similarly, transplanting oysters with light or undetectable Perkinsus marinus infections from low to high salinity can stimulate rapid infection development (Andrews and Ray 1988). The history of Perkinsus marinus further illustrates, on a much larger scale, how temperature, combined with a probable introduction, can trigger a disease outbreak. Although Perkinsus marinus was first described and associated with oyster mortalities in the Gulf of Mexico in the late 1940s, it had probably infected oysters in the southern United States for decades, at the very least (Mackin and Hopkins 1962). In 1949, the parasite was detected in oysters from the lower portion of Chesapeake Bay (Andrews and Hewatt

366

Shellfish Aquaculture and the Environment

1957) and was found in oysters from other sites along the southeastern United States around the same time (Ray 1954). Early studies documented that the parasite multiplied and killed oysters most readily at temperatures above 20°C and was especially virulent when the temperature exceeded 25°C. This temperature dependency was thought to explain why the parasite did not become established further north (Mackin 1962). Even when large numbers of infected oysters were brought into Delaware Bay from the lower Chesapeake in the mid-1950s, Perkinsus marinus, which spread to nearby native oysters, caused no noticeable mortality (Ford 1996). The parasite effectively disappeared in the bay after an embargo was placed on all oyster imports at the start of a period of below-average winter temperatures in the 1960s. Numerous surveys of oysters in Delaware Bay and sites farther north failed to detect the parasite in the succeeding years (Newman 1971; Meyers 1981; Ford 1996). In 1990, however, a severe epizootic began in Delaware Bay. Over the next 2 years, during a period of significant warming, especially in the winter, dermo disease outbreaks occurred in both wild and cultured eastern oysters along the coast from Delaware Bay to Cape Cod, MA, a 500-km distance (Cook et al. 1998). Although the range extension was not associated with known contemporary transplantation of oysters, historical transfers of oysters from the south to overfished regions of the north, dating back more than a century, would have undoubtedly introduced Perkinsus marinus many times (Ford 1992). In this colder region, the parasite may have persisted at low and undetectable levels, as apparently occurred in Delaware Bay, until the temperature became warm enough to stimulate an epizootic. Thus, a combination of introductions by oyster growers and a changing environment that became more favorable for the parasite is a likely explanation for the rapid range extension of dermo disease

outbreaks in the northeastern United States (Ford 1996).

Other forces in the natural environment Although temperature and salinity are the two best documented environmental influences on shellfish disease dynamics, there are clearly many chemical constituents and physical parameters of seawater that likely affect shellfish infectious disease agents or host defense and physiological mechanisms. A detailed review of all such mechanisms is beyond the scope of this chapter, notwithstanding the limited knowledge base in this area, but we provide some examples of other physical and chemical environmental factors that directly affect shellfish health and may interact with shellfish infectious agents and shellfish to promote disease conditions. Clearly, in estuarine environments, high rates of freshwater input can enrich the estuary with nutrients and such conditions are closely linked to phytoplankton blooms (Margalef 1978; Smayda 1997; Anderson et al. 2008) that in turn are linked to bacterial blooms. And as a more specific example, certain pathogenic bacteria require iron and may proliferate in estuaries during periods of iron enrichment or by using a competitive advantage for the sequestration of iron (Wang and Newton 1969). Acid sulfate soils are another example of environmental factors that may affect shellfish both directly and indirectly. Disturbance of acid sulfate soils can cause estuarine acidification, as documented in many areas of eastern Australia. It was reported in that region that acid sulfate soil outflows have low pH and elevated metal concentrations, primarily of iron and aluminum (Dove and Jesmond 2007). Declines in the production of Sydney rock oysters Saccostrea glomerata have been linked to estuarine acidification. It was reported that reduced oyster valve activity and

Shellfish diseases and health management 367

filtration rates occurred at pH 5.5 and that experimental acidic treatments (pH 5.1) containing 7.64 mg L−1 of aluminum or acid sulfate soil affected water caused changes in the mantle and gill soft tissues after short-term (6 h) exposure (Dove and Jesmond 2007). While these pH affects may directly effect larval shellfish survival, they may also promote the proliferation of opportunistic pathogens and render the developing larvae more susceptible to infectious agents. Similarly, with regard to low pH, ocean acidification, resulting from increased carbon dioxide concentrations in ocean water, may directly affect the ability of larval shellfish to deposit aragonite (Chapter 17 in this book), the first calcareous material of shell building in larvae (Anthony et al. 2009; Doney et al. 2009; Watson et al. 2009), and may also interact with infectious agents to reduce survival of larvae. While such complex interactions may be common phenomena, knowledge of the actual processes is poorly developed. Water circulation patterns and estuarine flushing rates that would dilute or concentrate infective elements clearly can affect infectious dose, exposure duration, and a variety of factors that affect shellfish health. It was proposed that dilution of Perkinsus marinus by fresh water inflow, as well as salinity per se, could act to diminish infection pressure in lowsalinity regions (Mackin 1956). Ford and Haskin (1982) described a time gradient in the onset of Haplosporidium nelsoni infections over a section of lower Delaware Bay where the salinity was entirely favorable for the parasite. They hypothesized that the center of this region was the major source of infective stages, which were diluted as they spread outward by both fresh- and saltwater. Fuentes et al. (1995) found a clear gradient of infection by Marteilia refringens in blue mussels, Mytilus galloprovincialis, in the Ria de Arousa in northwestern Spain. They could not clearly link it to salinity and speculated that poorer flushing might contribute to the higher infection levels they

recorded at the head of the estuary compared with its mouth. In a study that measured the abundance of Perkinsus marinus in the water column as a function of tidal flow, Ellin (Ellin 2000) was able to show that regular tidal flushing and episodic events such as hurricanes influenced the movements and concentrations of planktonic Perkinsus marinus.

Interactions of hosts and disease agents within the aquaculture environment Shellfish culture activities can potentially affect disease expression by altering the condition of the shellfish, the disease agent, or the production environment. Movement of broodstock or juveniles can introduce a new disease agent to a naive native population or introduce a naive host to an enzootic disease agent. Production systems change the environment by growing animals under conditions that are substantively different from those that they encounter in their natural environment. Health management is required to minimize the chances for the culture environment to facilitate proliferation and transmission of pathogens or stress the shellfish so that its natural defenses are compromised.

Introduction of a disease agent Although transport of bivalves, primarily as juveniles for growout, is often stated to be responsible for the spread of shellfish infectious disease agents (Rosenfield and Kern 1979), good documentation of this mechanism is rare. The introduction of Haplosporidium nelsoni to the U.S. East Coast is often blamed on the unauthorized importation of infected Pacific oysters, Crassostrea gigas, the presumed natural host, sometime before epizootic mortalities occurred in the mid-Atlantic estuaries in the late 1950s (Burreson et al. 2000).

368

Shellfish Aquaculture and the Environment

Such an introduction of infected oysters could have occurred as there are anecdotal reports of importation of Pacific oysters for shellfish culture around this time (Andrews 1980). However, the only documented imports occurred in other regions and/or well before or after the outbreaks (Burreson et al. 2000; National Research Council 2004). Other means of introduction should not be discounted. For instance, immediately before and after World War II, the number of ships transiting the Panama Canal from the Pacific to east coast ports increased dramatically (Ruiz et al. 2006). The parasite might have arrived in Pacific oysters attached to the hulls of ships, as spores in ballast water, or even in a hypothesized alternate or intermediate host (Burreson and Ford 2004). In contrast, the movement of Bonamia ostreae, a highly lethal parasite of the European flat oyster, Ostrea edulis, from the U.S. West Coast to Europe through oyster shipments is relatively well documented. Microcell parasites were first noted during trials of Ostrea edulis in California in the 1960s (Katkansky et al. 1969). These microcells and their effect on the host appeared to be identical to the parasite later described as Bonamia ostreae. Subsequent to the California report, microcells and disease manifestations that also appeared identical to Bonamia ostreae were found in Ostrea edulis in Washington State and California (Elston et al. 1986; Friedman et al. 1989). The source of the infected Washington oysters was traced to a hatchery in Elkhorn Slough, CA, which was also the source of oysters shipped to France in the 1970s (Elston et al. 1986). High mortalities caused by Bonamia ostreae began in Brittany, France, in 1979 (Balouet et al. 1983) and around the same time in northern Spain, where oysters had also been shipped from the California facility (Elston et al. 1986; Figueras 1991; Cigarría and Elston 1997). After the initial introductions, the parasite spread to many

other European sites, with a devastating impact on Ostrea edulis culture (Carnegie and Cochennec-Laureau 2004). During the 1970s, numerous lots of Ostrea edulis were sent to Maine from California, and Bonamia ostreae was subsequently discovered in naturalized flat oysters there (Friedman and Perkins 1994), although prevalence and intensity were low (Zabaleta and Barber 1996). The argument that Bonamia ostreae was introduced to new areas in shipments of infected oysters is strengthened by the fact that the parasite is directly transmissible between oysters (Poder et al. 1982). Shipments of infected oysters from Elkhorn Slough, CA, is the most plausible explanation for the spread of Bonamia ostreae to Washington State and Europe, and probably also to some locations in Maine (Friedman and Perkins 1994), although some embayments in Maine remained Bonamia-free in the late 1980s (Elston et al. 1987). The possibility of an introduction to California from the mid-Atlantic (Farley et al. 1988) is less likely. Ostrea edulis was originally introduced to the U.S. East Coast in a shipment from the Netherlands to the Bureau of Commercial Fisheries laboratory in Milford, CT, in 1949 (Loosanoff 1955). In the early 1960s, offspring were transferred to Chincoteague Bay, VA, and to several sites in California. Farley et al. (Farley et al. 1988) described a microcell parasite in the Chincoteague Bay group and later in oysters transferred to California sites, including a hatchery in Elkhorn Slough. They concluded that the microcell was Bonamia ostreae, based on morphological and pathological similarities to the French parasite. More recent molecular evidence, however, suggests that the Chincoteague Bay microcell is probably not Bonamia ostreae, but is similar to the Bonamia exitiosa-like parasite that may be a natural parasite of the crested oyster, Ostreola equestris, south of Cape Hatteras (see the following section) (R. Carnegie, Virginia Institute of Marine Science, pers. comm., September 2009).

Shellfish diseases and health management 369

Bonamia ostreae is thus an example of a disease agent, known to be transmitted directly between oysters, with a well-documented linkage to the movement of the host oyster. It is also an example of an avoidable introduction into Europe, as the presence of the parasite in the California flat oysters and the high mortalities there were known. This information was available prior to the shipment of the infected oysters to Europe (Katkansky and Warner 1974), although the oyster seed was reported to have been examined by histology and declared disease-free in regard to at least some of the shipments to Spain (Cigarría and Elston 1997). More structured health management and more sensitive detection methods are in place today and, if used properly, would prevent such an introduction.

Introduction of naive host Bonamiosis in Crassostrea ariakensis Heavy losses of the native eastern oyster, Crassostrea virginica, due to MSX and dermo diseases during the 1980s and 1990s in the Chesapeake Bay led to interest in the possibility of introducing another species of oyster that would not be susceptible to either disease. The Asian oyster, Crassostrea ariakensis, appeared to be a good candidate. Following the International Council on the Exploration of the Sea (ICES) protocol (ICES 1994), with the additional precaution of using triploid oysters to diminish the potential for reproduction and possible naturalization, the oysters were tested in waters enzootic for both parasites in Virginia (Calvo et al. 2001). These oysters did not develop detectable MSX infections. Perkinsus marinus infections did occur, although they were not sufficiently severe to kill the infected oysters. However, a later challenge test found that the oysters developed severe lethal infections in a laboratory hatchery (Moss et al. 2006). Subsequently, during a

test of growth and survival of Crassostrea ariakensis in waters just south of Chesapeake Bay in North Carolina, sudden high mortality occurred that was associated with infection by a previously undescribed species of Bonamia (Burreson et al. 2004). A survey of local fauna immediately detected the parasite, along with another Bonamia sp., in a small, noncommercial species, the crested oyster, Ostrea equestris. Although season peak prevalence of infection by this newly discovered Bonamia sp. in the nonnative Asian oyster exceeded 90% (Burreson et al. 2004; Carnegie et al. 2008), that in the native oyster rarely exceeded 2% (R. Carnegie, Virginia Institute of Marine Science, pers. comm., 2009). Although Bonamia sp. is restricted to highsalinity water, there is concern about the possible spread of the parasite into the estuarine waters of Chesapeake Bay (where it has not been observed in histological assays) if culture of the nonnative were to be initiated on a large scale (Carnegie et al. 2008).

Vibrio tapetis (causative agent of brown ring disease) Brown ring disease primarily affects the Japanese littleneck clam (also known as the Manila clam), Venerupis (=Tapes, =Ruditapes) philippinarum in Western Europe (Paillard et al. 1994). The disease name derives from the characteristic ring of organic material (periostracum) deposited by affected clams on the inner edge of each valve. The host clams were accidentally introduced to the northwestern United States in shipments of Pacific oysters in the 1930s, and then deliberately from there to France in the late 1970s for aquaculture, due to their more favorable growth rate compared with their native cousin, Venerupis decussatus (Flassch and Leborgne 1992). For a nearly a decade, the species appeared to thrive in its new environment and became established in the wild, but in 1987, heavy mortalities associated with the brown ring condition were

370

Shellfish Aquaculture and the Environment

reported in culture parks of northwestern Brittany. As culture of the clam spread to other European countries, brown ring disease outbreaks followed and the disease is now found in England, Spain, Portugal, and Italy, as well as France (Paillard 2004). The disease affects naturalized populations of Venerupis philippinarum in Europe, as well as those under culture. The native European littleneck clam, Venerupis decussatus, is highly resistant to brown ring disease, even though it inhabits the same waters (Maes and Paillard 1992; Allam et al. 2001). Interestingly, the condition has not been reported in northwestern North America, the source of clams brought to Europe. Very light conchiolin deposits were recently reported in Venerupis philippinarum in Korea, near the northern portion of the clam’s native range. A bacterium genetically similar to Vibrio tapetis was isolated from affected clams (Park et al. 2006). The seriousness of the disease outbreak in northern France may illustrate another case in which a naive, introduced host was much more susceptible than a closely related native host to an enzootic pathogen.

Solutions: 1. Shellfish aquaculture development and health management Annual world production of molluscan shellfish is estimated to exceed 12 million metric tons (FAO 2007). While much of this critical portion of the world food supply is still derived from wild-catch fisheries, increasingly intensive production technology provides a more reliable and higher-quality source of shellfish seed that may be genetically adapted or selected for specific growout locations. Starting with research and development in the 1950s and commercialization in the 1970s, the molluscan (oysters, clams, scallops, abalone, and related species) aquaculture industry has become increasing intensified on a worldwide basis. Wild reproduction and recruitment cannot

sustain the demands of quality, product specialization, and quantity that the growing world population and market requires. In addition, natural reproduction of edible molluscs has declined in many developed shoreline areas due to shoreline urbanization, industrial and residential development, and consequent degradation of habitat and water quality. The bivalve shellfish industry is increasingly based on a sustainable intensive agricultural production model, with multiple modes and methods for growing marketable product (Fig. 13.7). Intensification requires the selection and management of broodstocks and the controlled production of larval and seedstocks. This intensification provides an opportunity for selecting seedstocks with desirable traits and adaptability for optimal growth and quality in specific environments. The availability of high-quality seedstocks provides opportunities for producers to purchase seed and grow market-size product without the necessity of building and operating a complex and capital-intensive seed production facility. As a result, sales of shellfish seed are routinely made on a regional, and occasionally an international, basis and support growout industries in many regions of the world. Good animal husbandry and health management principles are thus essential to establish the high health status of such shellfish seed and to minimize the risks of disease outbreaks and disease agent transfers as described in previous sections of this chapter.

Achieving healthy farmed shellfish stocks Prevention of infectious animal disease and thus ensuring the health of food animals must be managed by a system of farming protocols and management procedures that maintain a healthy rearing environment, and testing for disease-causing agents. Where geographically

Shellfish diseases and health management 371

(1) Brood stock spawning

(2) Larval culture (3a) Seed cultivation in upweller

(3c) Seed set on cultch (4) Growout to market-sized oyster

(3b) Seed cultivation in floating upweller (“flupsie”)

Figure 13.7 Diagram showing vertical integration of shellfish farming including broodstock management, hatchery production of larvae, and growout of the final product. There are many varieties of culture and growout methods that are particular to species and locations. (From Elston 1999.)

restricted diseases (often referred to as reportable diseases) are concerned, government oversight and regulation is universally recognized as necessary. Systematic and effective management of food animal health results in reduction of infectious disease outbreaks, ability to minimize effects of disease outbreaks that do occur, reduction of animal losses and suffering, reduction of wasted resources, and more efficient farming production systems. These goals of animal health management are widely agreed upon. The details of how such goals are achieved may be debated vigorously, but with regard to the containment of reportable disease agents, some combination of producer expertise and responsibility, and independent government oversight is clearly needed.

Biosecurity Biosecurity is the word applied to the overall objective of maintaining animal health by preventing the spread of infectious diseases. While the word biosecurity has taken on a variety of additional definitions beyond the scope of animal health, it is defined in animal health management as precautions taken to minimize the risk of introducing an infectious disease agent into an animal population. Such precautions, to be effective, must be organized, comprehensive, and systematically applied. A biosecurity plan or “high health” plan must be developed for the specific animal husbandry system and effectively integrated into the regional governmental system of animal disease notification. A biosecurity plan must

372

Shellfish Aquaculture and the Environment

identify pathways for the introduction and spread of infectious disease and define steps that are applied to mitigate risks of introducing and spreading infectious diseases through such pathways. The biosecurity plan should be accountable to independent oversight and ensure that infectious disease risks are regularly reassessed and biosecurity measures are adjusted accordingly.

Health management and biosecurity: Shellfish High Health Program On the U.S. West Coast, the Pacific Coast Shellfish Growers Association developed and adopted a voluntary Shellfish High Health Plan (SHHP) (Elston 2004), initially designed for seed producers but applicable to all shellfish producers. This plan was subsequently adopted as one requirement for shellfish farms applying for U.S. Department of Agriculture, Animal and Plant Health Inspection Service health certificates for export of live shellfish product. The SHHP requires participating producers to establish the following basic elements for animal health management: 1. A system of shellfish health surveillance, certifications, records, and documentation 2. A schedule of examination for certifiable or reportable shellfish diseases 3. A plan for broodstock management 4. Hatchery and nursery operations protocols 5. Characterization of reportable disease status of water sources and shellfish culture areas 6. Maintenance of broodstock integrity for production facilities and hatcheries 7. Operations protocols that pertain to health management 8. A response plan for infectious disease outbreaks Once shellfish growers establish and practice a customized animal health management plan

for their farms that incorporates these elements, they will have markedly reduced the risks associated with shellfish infectious disease outbreaks.

Awareness, education, and effectiveness of biosecurity measures It is clear that the quality of biosecurity and maintenance of animal health is directly related to the awareness and practice of animal health management by farm operators. Education and awareness (Chapter 16 in this book) of infectious animal health risks by aquatic farmers is the most effective and important component of animal health management, and particularly so within countries where animal transfers are easily implemented in that animals do not have to cross international borders. Self-governance in matters of animal disease control does not constitute a complete and sufficient system of animal health management but without committed action by an educated animal farming industry, infectious disease control efforts are extremely difficult, generally ineffective, and highly cost-inefficient.

Governmental and intergovernmental systems for shellfish disease control The World Organization for Animal Health (still referred to as OIE, in reference to its former name, Office Internationale Epizooties) was formed in 1924 as a response to the spread of rinderpest, an infectious viral disease of cattle and other species, occurring in Belgium. The organization became the leading world animal health advisory organization (composed of 174 member countries in 2009) and became even more important with the official agreement between the OIE and the World Trade Organization in 1998 under which OIE became the official arbiter and rulemaker in regard to international live aquatic animal transports. In 1995, OIE adopted its first

Shellfish diseases and health management 373

version of the Aquatic Animal Health Code and the Manual of Diagnostic Tests for Aquatic Animals, which included molluscs, among other aquatic species groups. Both the Code and the Manual of Diagnostic Tests (OIE 2009b, 2009c) have been refined and improved through a process of expert working groups, committees, and adoption by the General Session. These documents are in a state of constant evolution based on disease surveillance, emerging diseases, new technology, and to some extent, political will of member countries. These documents are used by member country competent authorities to develop individual country standards for recognition of reportable animal diseases, methods for risk analysis, import/export procedures, contingency plans, sanitation, surveillance and detection, and all matters related to import and export of aquatic products that carry risk of aquatic infectious disease agents. The term competent authority refers to the designated governmental department or entity that has authority and responsibility for animal disease control in the particular country. The OIE, based on advice from its expert committees, specifies diseases that are reportable or notifiable to the OIE. Member countries are obligated to report the occurrence of these notifiable disease agents to the OIE, through their competent authority. The notifiable diseases for molluscan shellfish as of 2009 are the following: Infection with Bonamia ostreae Infection with Bonamia exitiosa Infection with Marteilia refringens Infection with Perkinsus marinus Infection with Perkinsus olseni Infection with Xenohaliotis californiensis Abalone viral mortality Each member country of the OIE has an official delegate who represents the competent authority for that country. While not all countries belong to the OIE and reporting systems

may not be consistent with OIE standards in all member countries, the OIE is generally considered the reference authority for animal diseases. However, under OIE guidelines, member countries may establish higher standards (i.e., additional controlled diseases for their country) if they demonstrate freedom from such additional disease agents. A detailed discussion of individual country disease control systems is beyond the scope of this chapter. National regulatory authorities and regional, provincial, or state jurisdictions, to the extent they are present in a given country, are responsible for implementing a notifiable disease agent surveillance and reporting system.

Surveillance to support animal health management Shellfish health surveillance, which provides the knowledge of where, in what species, and under what environmental conditions specific infectious animal diseases occur is a fundamental requirement for effective biosecurity and animal health management. Surveillance for infectious diseases requires a commitment of resources and must be done on a regular and ongoing basis to provide meaningful health management information. Existing shellfish health surveillance information may be intensive for particular time periods in that effort is often related to outbreaks of disease, or to finite-length surveillance projects. Surveillance efforts tend to be focal in geographic area and intensive for cultured species but more limited or nonexistent for noncultured or nonharvest species that may be disease vectors or carriers. Such spotty information is the nature of the geographic focus of shellfish culture and the seemingly always limited resources available for surveillance. In the western United States, shellfish farms have undertaken the expense of shellfish health surveillance for over two decades in order to facilitate interstate movements of shellfish and

374

Shellfish Aquaculture and the Environment

export of shellfish seed. Even such well-planned and well-implemented surveillance programs may not be comprehensive in regard to potential, and unknown, vector species. Thus, shellfish health management is always a work in progress. Initiation of shellfish disease surveillance is likely to reveal infectious conditions and diseases present in shellfish populations. Most often, such observations are of existing conditions that may fluctuate in prevalence and intensity. In some cases, these observations represent newly introduced diseases. However, it is a common mistake to regard a newly discovered shellfish disease as a disease new to the area of discovery. Only long-term surveillance can reveal changes in geographic distribution of diseases and their prevalence and intensity.

if any shellfish or live animal transfers are to be made, there is never a zero risk. Each transfer must be evaluated on the basis of best available knowledge. In some cases, further information may be needed to make an informed decision. This issue also is often framed in the question of whether the presence of a microorganism constitutes the presence of a disease. The answer is clearly no, as a disease state must be initiated in a susceptible host and, in many cases, shellfish and other animal species live in some balance with infectious agents, such that disease expression is suppressed. In some cases, answers are not clearly intuitive and some level of risk assessment is used to better frame the level of risk, the key unknown information and a methodology to manage risk for particular cases.

Assessing impacts of disease

Establishing geographic biosecurity for shellfish culture

A critically important area of shellfish infectious disease management is the assessment of the seriousness and consequences of particular infectious disease agents. Several issues repeatedly arise in regard to such assessments. While there are clearly infectious disease agents that cause serious losses of shellfish, as discussed in this chapter, there are also examples of bacteria, viruses, and parasites that occur at apparently low prevalence, low severity, and overall low health impact. Whether or not such microorganisms constitute a geographic transfer hazard may be debated, and clear resolution of such debate is difficult. While it is possible that such low-impact and low-prevalence microorganisms might have limited impact in a known shellfish host species, they could have a high and significant impact in other host species or in a different environment. However, all shellfish, like all animal species, have some associated microorganisms, so that all transfers of live animal stocks necessarily involve some transfer of microorganisms. As a result,

Biosecurity must be effectively implemented in order to allow the operation of animal husbandry, including the movement of broodstocks and seedstocks of animals between facilities and between coastal growout areas that are part of the larger marine environment. In contrast to the requirements for marine aquaculture biosecurity, a freshwater fish culture facility, operating at an inland site using groundwater, is more easily managed as a controlled facility for which the potential pathways for infectious diseases are viewed as more clearly identified and managed. The primary difference in relative ease of management for such a facility is a known and often pathogen-free water supply, accomplished through the use of groundwater or water sterilization methods, which are better known for freshwater than for saltwater. Other vector pathways, such as through air and by insects and rodents may be more challenging.

Shellfish diseases and health management 375

As the market for marine shellfish increases and the technology of culture advances, the industry is becoming increasingly dependent on hatchery production of shellfish larvae. Biosecurity for marine shellfish hatcheries is not so easily managed as biosecurity for freshwater fish hatcheries because most shellfish hatcheries rely on seawater drawn from estuarine environments. Hatcheries usually process such seawater in various ways to remove marine organisms, debris, and a portion of the microbiological component of the incoming seawater. However, no systems are available that are definitively demonstrated to effectively remove shellfish infectious disease agents without degrading water quality by removing essential components or introducing toxic byproducts of water treatment. Other pathways to minimize infectious disease entry (such as controlled introduction of broodstocks and seedstocks, and good sanitation) can be effectively implemented but with current technology, a given facility is usually not isolated from the infectious disease agents in the marine water it uses. While effective water treatment for the removal of infectious agents that does not degrade water quality needs to be developed, the present approach is to survey the surrounding waters for the presence of known infectious disease agents of concern. This surveillance must be done on a regular basis and scheduled to effectively determine the infectious disease status of water and animals that are used in the hatchery. Similarly, open nurseries for shellfish seed and broodstock holding and conditioning areas are managed by surveillance and knowledge of the infectious disease status of the waters in which they operate and by the other biosecurity measures mentioned, including control of intentional transfers of potential disease-carrying stocks into the water bodies where seedstock or broodstock are held. In order to develop a biosecurity plan and provide surveillance of marine environments where shellfish hatcheries, nurseries, and

broodstock maintenance areas are operated, a system of geographic “compartmentalization” or a larger-scale system of geographic “zoning” are suggested by the World Organization for Animal Health. These approaches are the basis for establishing disease-specific, disease-free management areas, compartments, and zones. The specific disease-free nature of such geographic compartments or zones is maintained by ongoing surveillance and ensured, to the extent possible, by implementation of the biosecurity plan.

Biosecurity established by compartmentalization and zoning The following brief description of zoning and compartmentalization is based on World Organization for Animal Health Aquatic Animal Health Code, 2009, Chapter 4.1 (OIE 2009a). Providing for compartmentalization or zoning recognizes the potential difficulty of establishing and maintaining freedom from a particular disease for an entire country, especially for diseases whose entry is difficult to control. Animal subpopulations may be separated by natural or artificial geographical barriers or, in certain situations, by the application of appropriate management practices. Zoning and compartmentalization are thus procedures implemented by a country to define subpopulations with distinct aquatic animal health status for the purpose of disease control or international trade. Compartmentalization applies to a subpopulation when management practices related to biosecurity are the defining factors, while zoning applies when a subpopulation is defined on a geographical basis. In practice, geographic considerations and good management are important in the application of both compartmentalization and zoning. Guidelines for applying these concepts are recommended in the relevant disease chapters of the OIE (2009a, 2009c) and are summarized here, based on those documents. According to these guidelines, before trade

376

Shellfish Aquaculture and the Environment

in live aquatic animals or aquatic animal products may occur, an importing country needs to be satisfied that its aquatic animal health status will be protected. In addition to contributing to the safety of international trade, zoning and compartmentalization may assist disease control or eradication within countries that produce aquaculture products. Implementation of compartmentalization or zoning will also assist in disease control following any disease outbreaks that may occur and in establishing the resumption of trade after such an outbreak. Separate requirements need to be developed for each disease for which the application of zoning or compartmentalization is used as a disease prevention tool. Thus, zones and compartments are disease specific, although some disease control and biosecurity measures can overlap if more than one disease agent is subject to the measures. The OIE Aquatic Code provides guidelines and specific steps for establishing compartments or zones. The competent authority (as defined previously) with jurisdiction in the producing country defines the animal subpopulation, the requirements for surveillance, and details of animal identification and traceability of aquatic animals. Such jurisdictions may require registration of all the aquaculture establishments and the establishment and maintenance of aquatic animal health status through a common biosecurity plan. The biosecurity measures used may include movement controls, use of natural and artificial boundaries, spatial separation of aquatic animals, and commercial management and husbandry practices related to animal health and surveillance for significant disease-causing agents. In addition to information on aquatic animal movements, the biosecurity plan should include production and stock records, feed sources, traceability of components of production related to animal health, surveillance results, a visitor logbook, morbidity and mortality history, medications, documentation of training, and any other criteria

necessary for evaluation of risk mitigation. The biosecurity plan should also describe how the measures will be audited to ensure that the risks are regularly reassessed and the measures adjusted accordingly. The exporting country must have the resources needed and available to establish and maintain a zone or compartment for international trade purposes including human and financial resources and the technical capability of the competent authority and the industry. The procedures used and details of implementation must be transparent and publicly available.

Shellfish health management: a veterinary approach Interest in shellfish infectious diseases increased after the outbreak of MSX disease in eastern oysters in the Delaware and Chesapeake Bays in the United States in the late 1950s. This interest was heightened by the outbreaks of marteiliasis and bonamiasis in Europe in the 1970s. A further marked interest in infectious shellfish diseases has occurred since about the mid-1980s, sparked by the growth of the shellfish industry and development of hatchery technology, continuing effects of previously as well as more recently discovered infectious diseases of shellfish, and the development of molecular biology. Much of this research work has been basic in nature and aimed at elucidating biology of infectious organisms, their phylogenetic relationships, their natural cycles, mechanisms of infectivity, and host response. With the growth of the farmed shellfish production industry, including the export of live shellfish products, a need for the application of infectious disease management has emerged. This is a need for application of health management, in addition to the ongoing need for basic and applied research. The application of animal health principles in terrestrial animal husbandry is implemented by veterinary medi-

Shellfish diseases and health management 377

cine. The veterinary medical profession has shown an interest in aquatic animal medicine over the last three decades, but training and attracting veterinarians to marine invertebrate animal health management is a slow process because relatively few well-defined jobs exist in the field. Nonetheless, growth of vertically integrated shellfish farming industries, development of intensive animal husbandry systems, development of governmental systems for aquatic disease control, and the legal requirements for both chemotherapeutant usage and veterinary attestation of export health certificates provides the basis of a need for a contingent of trained, qualified, and dedicated veterinarians to assist in shellfish health management. There is no doubt that a veterinary background in animal health management is extremely useful and applicable to shellfish husbandry. However, a veterinarian wishing to be useful to the shellfish industry requires substantial knowledge and experience outside of that offered in the routine curriculum of most colleges of veterinary medicine. Recognition of this need and its fulfillment by qualified veterinarians will advance the shellfish production industry. It is helpful to visualize the three components of shellfish health management in terms of regulation, animal health management, and research and recognize that these are three distinct components of animal health but that the individuals and institutions involved in these three components must interact effectively to yield the benefits of animal health management (Fig. 13.8).

Solutions: 2. Implementing health management for shellfish aquaculture With the advent of shellfish hatcheries in the last half of the twentieth century, shellfish culture began to emerge as a vertically integrated food production enterprise, similar in structure to traditional forms of agriculture.

Regulatory Structure

Practice of Medicine

Research and Development Figure13.8 The “animal health management house” depicts the relationship of health management components. Basic and applied research (the foundation) provide discovery of technical components of health management. Such discoveries are converted into the practice and management of animal health (veterinary medicine), and a regulatory or governmental structure (the protective roof) is in place to provide a system for protection from reportable diseases.

As a result, opportunities emerged to better control shellfish disease dissemination, and coincidentally, new opportunities for the accidental dissemination of infectious shellfish diseases emerged. The development of shellfish hatchery and nursery technology has occurred gradually and is still evolving. As this evolution occurs, production becomes more efficient and resources are more efficiently utilized. Shellfish hatcheries and nurseries, to be cost-effective, usually must produce shellfish larvae and juveniles in high concentrations. Such high concentrations may subject the early life stage shellfish to higher concentrations of opportunistic but virulent infectious agents than would occur in nature. Examples of diseases caused by such agents include vibriosis caused by Vibrio tubiashii and other bacterial species, and viral diseases such as oyster velar virus disease (OVVD) and OsHV disease, as well as noninfectious disease conditions (e.g., toxins from natural algal blooms). While much has been learned about conditions, such as opportunistic vibrioses and their management, there is still a great need and opportunity to improve the rearing environment in shellfish

378

Shellfish Aquaculture and the Environment

hatcheries, for example, by understanding and implementing appropriate water treatment systems and by understanding and managing the bacterial ecology of the hatchery. There is also a need and opportunity to improve conditions in shellfish nurseries, which usually are in a more open environment than hatcheries, given that as shellfish seed increase in size, their food requirement increases exponentially and for large-scale production they must be transferred to open environments with natural food production. The consequences of failure to manage animal health and failure to have an animal health management program in place were detailed in earlier sections of this discussion regarding the consequences of introduction of disease agents and introduction of naive hosts. We now provide some detailed examples regarding the mechanics of establishment of an effective animal health management system. This discussion is intended to provide examples of some of the specific challenges that have been documented in shellfish hatchery and nursery health management in the context of components of a vertically integrated shellfish production system. Many of the infectious agents that affect shellfish in hatcheries, nurseries, and intensive growout systems are opportunistic organisms that become pathogenic to shellfish because of the culture system in which they are held. Improving the environment of that system is a critical component of animal health management.

Broodstock management Broodstocks that are used in a particular hatchery must be certified to be free of OIEreportable and other infectious diseases of regional concern, unless the hatchery operates within an established specific disease-endemic area and provides seedstocks only to other such areas. Broodstock health surveillance must be implemented on a regular and ongoing

basis with annual or more frequent examinations. Broodstock that does not have such an established health history cannot be brought into the hatchery. The broodstock holding area, where the broodstock are maintained while not being used in the hatchery, must be similarly protected from the introduction of any other shellfish stocks unless they are similarly certified free of infectious diseases of concern.

Dissemination of larval and seedstocks of shellfish Larval and seedstocks that are produced in a hatchery or nursery system may be moved to distant growout locations. To minimize the possibility that there are infectious disease agents of concern in seedstocks, they must be examined for the presence of such infectious diseases. These examinations provide reassurance of the absence of infectious diseases because the broodstock should already have been examined. However, there are some known infectious diseases that are believed to occur only in early life stages, and examination of seedstocks is a check to ensure that no disease-causing microorganism of concern was introduced during the production cycle. In addition to preventing the dissemination of such infectious diseases, health and disease examinations of hatchery and nursery products also help insure the successful survival and growth of the larvae and seed at the outplant location and provide an opportunity to evaluate the condition of such seed prior to its use in field sites.

Dissemination of microalgae used as shellfish feed Microalgal cultures are widely disseminated for research and shellfish production purposes. There are few known examples of the

Shellfish diseases and health management 379

dissemination of significant infectious diseases with microalgae, but opportunistic bacterial infections, such as that caused by widely distributed bacterial shellfish pathogen Vibrio tubiashii, are able to coexist cryptically in some microalgal cultures and the transfer of this shellfish pathogen has been documented in algal transfers (Elston et al. 2008). An examination of such cultures for these pathogenic bacteria and other known detectable pathogens should be conducted prior to the use of new strains of such algae.

Management of infectious diseases in shellfish hatcheries Shellfish hatcheries provide a highly concentrated environment in which control of critical factors for the health of shellfish larvae is only a partially developed technology, and opportunistic disease agents can and do often become established in such systems and markedly reduce production. Although the protozoan disease agents that are dangerous to adult bivalves are not known to infect larvae, the latter are highly susceptible to bacterial and viral infections (Elston 1984, 1993, 1997; Elston et al. 1999, 2008; Paillard et al. 2004). Members of the genus Vibrio are the principal known bacterial pathogens of larvae in the hatchery, where they can cause extensive losses when uncontrolled blooms and infections occur (Tubiash 1970; Brown and Losee 1978; Brown 1981; Elston et al. 1981, 2008). Species in the genera Pseudomonas, Alteromonas, and Aeromonas have also been implicated in hatchery mortalities of essentially all species of bivalves under culture (Brown 1974; Elston et al. 1999; Paillard et al. 2004). Such opportunistically pathogenic bacteria may be introduced from the ambient water, by broodstock transfer, or through algal food sources into the hatchery where the environment, if unmanaged, can facilitate their proliferation. Bacterial concentrations can be magnified by the tem-

peratures at which larvae are typically grown in high-density cultures. Species of shellfish pathogenic vibrios generally proliferate rapidly at optimal shellfish larval-rearing temperatures. Bacterial management consists of regular routine monitoring for known shellfish pathogenic vibrios (e.g., Vibrio tubiashii) and isolation and disinfection of contaminated algal foodstocks and larval cultures. Using routine monitoring, the source of pathogenic bacteria and their reservoirs in the culture environment must be indentified and eliminated. Known methods for management of such infections have been discussed in detail (Elston et al. 2008) but further development of water treatment systems, understanding and management of bacterial ecology in hatcheries, and the judicious use of antimicrobial compounds could greatly increase the efficiency of hatcheries and reduce resources wasted by the effects of such opportunistic infections. OVVD was the first viral infection found in larval bivalves (Elston 1979). The infection, caused by an iridovirus, has been found sporadically in Pacific oyster larvae from hatcheries. The infection seems to be effectively managed by destruction of infected stocks and thorough sanitation of affected parts of the hatchery system, but the source or reservoir of the virus, presumably adult oysters, has not been confirmed and there are no reports of the infection in juvenile oysters (Elston and Wilkinson 1985). However, other iridoviruses have been reported in several species of adult bivalves in Europe (Comps 1970, 1988; Comps and Duthoit 1976; Comps and Bonami 1977). OsHV has been found in Europe, New Zealand, and Australia in conjunction with mortalities of hatchery-produced larvae and juvenile bivalves of numerous species (Hine et al. 1992; Nicolas et al. 1992; Renault et al. 1994; Arzul et al. 2001a; Renault et al. 2001a, 2001b; Friedman et al. 2005; Burge et al. 2006, 2007). This infection has been found in geographically limited locations in shellfish juveniles in the United States but not in

380

Shellfish Aquaculture and the Environment

shellfish hatcheries, despite intensive investigation. OsHV is associated with juvenile oyster mortality but defining the exact role of the virus in such mortalities has been an elusive goal to date (Friedman et al. 2005; Burge et al. 2006, 2007). Both intra- and interspecies transmission has been demonstrated and the finding of OsHV in adults suggests the possibility of vertical transmission (Arzul et al. 2001b). Infections are temperature dependent, and productive infections for Pacific oyster OsHV occur above 25–26°C, although incomplete replication of viral particles may occur at about 22–23°C (LeDeuff et al. 1996). Other shellfish herpes viruses may replicate at lower temperatures (Meyers et al. 2009). Invasive protozoa have been found in hatchery-reared larval bivalves including a flagellated amoeba (Kent et al. 1987) in geoduck clams (Panope abrupta) and hard clams (Mercenaria mercenaria) (Ragone Calvo et al. 2002), and an invasive ciliate and a gregarine-like infection in Pacific oysters (Elston 1999; Elston et al. 1999). Little is known about the biology and management of these conditions.

crassostreae, the disease condition has been renamed “Roseovarius oyster disease (ROD)” (Boettcher et al. 2005; Maloy et al. 2007). In 1990, a severe outbreak occurred in cultured oysters on the north shore of Long Island, NY, and the disease has periodically affected oyster culture operations in several locations from New York to Maine. The disease has not been reported in oysters growing in the wild. ROD affects primarily young, rapidly growing oysters in a variety of nursery containers from floating trays to upwellers (Ford and Borrero 2001). Mortalities, which can exceed 90% within a few weeks, are largely restricted to juveniles with a shell height less than 25 mm, although the organic deposits are found in larger juveniles and are occasionally reported in adult oysters. Experimental studies demonstrate that high animal density, poor flushing, and the likely buildup of bacteria at culture sites (Bricelj et al. 1992; Rivara and Czyzyk 1995; Boettcher et al. 2006) are risk factors for this disease in shellfish nurseries (Fig. 13.9). Temperatures reaching and exceeding 21–22°C are permissive for the disease. Phytoplankton and vibrio blooms were associated with the onset of ROD at one site (Lee et al. 1996). Outbreaks of this disease are

Management of infectious diseases in shellfish nurseries

90 80 70

Percent Mortality

Roseovarius oyster disease (ROD) was first reported in the mid and late 1980s by oyster growers with nursery sites in New England states of the United States. The disease affected juvenile eastern oysters, Crassostrea virginica. It was characterized by a severe cupping of the lower valve and a ring of organic material deposited on the inner side of both valves, much the same as that found in brown ring disease. Originally, the syndrome was given the generic name “juvenile oyster disease (JOD).” To differentiate this condition from other mortalities of juvenile oysters, and because it is now established to be associated with a specific bacterial infection, Roseovarius

60 50 40 30 20 10 0 LD MD HD High Flow

LD MD HD Medium Flow

LD MD HD Low Flow

Figure 13.9 Effect of water flow through upwellers on mortality associated with Roseovarius oyster disease of juvenile eastern oysters. (Redrawn from unpublished data of G. Rivara, New York Sea Grant.) LD, MD, HD: low, medium, and high density.

Shellfish diseases and health management 381

managed and prevented by early-season production of larvae in hatcheries. Such production during the winter or early spring permits juveniles to grow beyond the 25-mm window of susceptibility before water temperatures reaches the 21–22°C threshold (Davis and Barber 1994; Ford and Borrero 2001). In addition, use of genetically selected stocks can significantly reduce losses associated with the disease (Barber et al. 1998; Lewis 2001). Vibriosis has also been documented as a significant source of losses of juvenile bivalves including nursery reared shellfish seed (Tubiash et al. 1965; Elston et al. 1982, 1999), and in the case of juvenile Pacific oysters, Vibrio tubiashii has been documented as the diseasecausing bacterium (Elston et al. 2008). The pathological manifestations of vibriosis are similar but distinctive from those of larval oysters. Focal infections tend to localize in the mantle or space between the mantle and the shell (Fig. 13.10). While the disease can be fatal in juvenile shellfish, they display some degree of host defense and presumably the infection can be resolved under favorable conditions. The disease can be transferred from

infected groups of larvae to the developing juveniles but may also infect juveniles held in high density with insufficient water exchange (Elston et al. 2008). While a high environmental concentration of the pathogen can be a management challenge, real-time detection, reduction of animal density, and enhanced water flow and sanitation can be used to manage the disease during such high-risk periods. Hinge ligament disease is an important opportunistic disease of many species of cultured bivalves (Elston 1984; Dungan and Elston 1988). The causative bacterium has been isolated and, although not fully characterized, appears to be associated with the group of gliding bacteria formerly known as Flexibacter spp. and now known as Tenacibaculum spp. (Dungan et al. 1989). The infection erodes the hinge ligament that attaches the two valves of bivalves. Light infections appear to have limited impact but severe infections can markedly decrease the normal function of the ligament. The disease can be managed to some extent with freshwater rinses and sodium hypochlorite treatments but

Figure 13.10 Histology section showing an aggressive opportunistic infection by Vibrio tubiashii bacteria, causing an abscess and localized between the internal shell surface and mantle of a juvenile Pacific oyster (abscess area indicated by oval). Mn, mantle. (From Elston et al. 2008.)

382

Shellfish Aquaculture and the Environment

neither treatment has been rigorously evaluated for effectiveness and the use of chemicals to treat the disease is subject to regulatory approval in some locations (Elston 1999). Reduction of culture temperature retards the enzymatic degradation of the ligament (Dungan et al. 1989). Several invasive protozoa have been found in nursery-reared juvenile bivalves including invasive ciliates in Pacific oyster juveniles and a gregarine-like infection in the same species (Elston et al. 1998; Elston 1999). These appear to be highly opportunistic infections that infect juvenile bivalves that are not in an optimal condition of health. These infections should be distinguished from end-stage colonization of dead tissue by ciliated protozoa such as Uronema marinum (Plunkett and Hidu 1978). Little is known about the prevention and management of these sporadically occurring infections.

Examples of infectious diseases and their management in shellfish growout systems: QPX disease and “summer mortality” QPX (infecting Mercenaria mercenaria) was discussed earlier as an example of a parasite with a geographic range that appears to be limited by high temperature, but it is an excellent example of how two very different aquaculture practices have an impact on disease outbreaks. QPX is a member of the phylum Labyrinthulomycota, a group of microorganisms that lives in marine and estuarine environments on micro- and macroalgae, and detritus (Porter 1990). It can become pathogenic to molluscs held in captivity or culture (Polglase 1980; Jones and O’Dor 1983; Bower 1987). The disease-causing agent is considered to be facultative and opportunistic, and can live outside, as well as inside, the clam and can take advantage of the clam host that is compromised in some way. Most QPX outbreaks

have been in culture situations, but the parasite has been detected, generally at low prevalence (rarely greater than 10%), in wild juvenile clams over most of its known range, from the Canadian Maritime provinces to the lower Chesapeake Bay, in the United States. Serious mortalities caused by QPX were documented in adult and juvenile hard clams in a Canadian hatchery in 1989 (Whyte et al. 1994). Subsequently, the organism was found in cultured clams in Massachusetts, New Jersey, and Virginia (Ragone Calvo et al. 1998; Smolowitz et al. 1998; Ford et al. 2002). Initial surveys (Ragone Calvo et al. 1998; Smolowitz et al. 1998; MacCallum and McGladdery 2000) led to the suspicion that QPX outbreaks might be associated with aquaculture practices. Evidence from a study in New Jersey supported this idea (Ford et al. 2002). Growers from New Jersey had purchased and planted seed from southern U.S. states because they could obtain it earlier in the season and they found that it grew faster than local seed. There is no evidence to suggest that the southern seed clams were infected when they arrived— indeed, QPX has not been observed in clams growing south of Virginia—or in seed coming from hatcheries (Ford et al. 1997). In subsequent experimental trials in Virginia and New Jersey, clams from Florida and South Carolina acquired numerous and heavy QPX infections and suffered high mortalities (Ragone Calvo et al. 2007). Those from Massachusetts and New Jersey had few infections and low mortality. Clams from Virginia exhibited intermediate values. Another risk factor may be the high density at which clams are planted within plots as well as the number of plots in growout sites. Experimental manipulation of density suggested this as a possibility (Ford et al. 2002). Clams (of South Carolina origin) were planted at three densities: (215, 430, and 860 clams m−2) and sampled for QPX infections and for clam mortality over the following 10 months. A weak positive correlation was found between clam density and QPX levels (Fig. 13.11), but

Shellfish diseases and health management 383

Percent Prevalence

70 60

10/3/1997

50

11/29/1997

40 30 20 10 0 6

Sample Intensity

5 4 3 2 1 0

215

430 Clams (m–2)

860

Figure 13.11 Effect of host density on a hard clam disease. (A) Prevalence and (B) sample intensity of Quahog Parasite Unknown (QPX) infections in hard clams field planted at three densities. Typical commercial planting density approximates the 215 m−2 figure. The clams had been deployed for 27 and 29 months when sampled. Error bars represent the standard error of the mean. (Redrawn from Ford et al. 2002.)

the 1-m2 plots may have been too small to significantly disadvantage clams or to have much of an effect on transmission. Another probable association with density was recorded when an outbreak of QPX disease occurred in an unusually dense bed of wild-set hard clams in Raritan Bay, on the border between New York and New Jersey. Subsequent histological surveys of other clam beds in the bay found few detectable infections. Prevalences were rarely above 10%, although occasional values as high as 30% were found and these were

generally in the most densely set beds (Allam et al. 2005). Thus, density, whether in culture or in the wild, appears to favor QPX outbreaks, but the most serious losses have been associated with the importation of nonnative seed clams from southern regions, which may be disadvantaged by low temperatures in northern growout locations or may have developed no inherent resistance to the disease through natural selection, or both. Clam growers now avoid using stocks from outside their region, and severe outbreaks of QPX disease have dropped significantly. A syndrome known as “summer mortality” of cultured bivalves has been reported in the literature for many years (Imai et al. 1965; Mori et al. 1965; Lipovsky and Chew 1972; Glude 1975; Koganezawa 1975; Mori 1979). The first reported occurrence of summer mortality was in the late 1940s and affected Pacific oysters (Crassostrea gigas) cultured in Japan (Koganezawa 1975). A similar syndrome in this oyster was subsequently described on the U.S. West Coast (Glude 1975; Cheney et al. 2000) and in France (Soletchnik et al. 2005). Warm-season losses have also been reported in blue mussels, Mytilus edulis, in Canada and the United States (Mallet et al. 1990; Newell and Lutz 1991; Myrand and Gaudreault 1995) and in zhikong scallops (Chlamys farreri) in China (Xiao et al. 2005). In the case of oysters and scallops, mortalities can reach a high level but the severity, and even the occurrence, of summer mortalities is highly variable from year to year. By definition, mortalities are initiated as water temperatures pass a temperature threshold (∼19–20°C for Pacific oysters). Summer mortalities sometimes occur during periods of low dissolved oxygen or during phytoplankton blooms (Cheney et al. 2000) and are greater in oysters reared near the sediment than in those placed off bottom on racks (Soletchnik et al. 2005). These losses are usually associated with reproductive development, which occurs during the warmer months for the affected species (Perdue et al. 1981;

384

Shellfish Aquaculture and the Environment

60 50 Percent Mortality

B B

40

B

B

B

B

30 B

20 10 0 10

B

100

1000

10,000

Number of Oysters per Bag Figure 13.12 Effect of stocking density of Pacific oysters on mortality associated with the summer mortality syndrome in France. The oysters were contained in mesh bags and held on racks. Error bars represent 95% confidence interval (CI). (Redrawn from Samain et al. 2008.)

Lambert et al. 2008). No single infectious agent has been implicated in the mortalities but Nocardia crassostreae (Friedman et al. 1991, 1998) is occasionally associated with later summer losses of Pacific oysters (Elston et al. 1987; Friedman and Hedrick 1991). Additionally, OsHV is found in some locations in seed oysters undergoing warm-season mortality losses (Renault et al. 2000; Arzul et al. 2001b; Friedman et al. 2005; Burge et al. 2006, 2007; Sauvage et al. 2009). However, it would be inaccurate and simplistic to attribute the cause of all or even most of summer mortality losses to either of these oyster pathogens. Scallops experiencing warm-temperature mortality in China were infested with many symbionts in their mantle cavities, a finding that was interpreted as indicating poor water circulation through the lantern nets in which they were being grown (Xiao et al. 2005). Algae and other encrusting epibionts readily foul the nets, bags, and cages in which cultured bivalves are grown. These effects reduce water flow, allowing the buildup of sediment and microorganisms within the containers.

Crowding and restricted water flow may then impair the ability of the bivalves within the containers to maintain adequate water flow in their gill cavity. Consistent with this hypothesis, decreasing the density of oysters in bags (Figs. 13.12 and 13.13) and scallops in lantern nets (Xiao et al. 2005) decreased the rate and extent of losses. Management practices (Chapter 3 in this book) can thus be implemented to reduce losses and the culturist’s challenge is to find cost-effective solutions that maximize sustainable production in a given unit area while minimizing costs and producing a high-quality product. A variety of solutions including site management and development of better technology will advance the health of cultured shellfish. Resistance and susceptibility to summer mortality losses in Pacific oysters, for example, are heritable traits (Boudry et al. 2008) and thus selective breeding (Chapter 12 in this book) should minimize mortality. Triploid oysters, which do not produce the quantity of gametes found in diploids, are used to produce high-quality product during summer periods in some locations but also

Shellfish diseases and health management 385

28 24 20 16 12 8 4 28 24 20 16 12 8 4 01

BVD

Mean Temperature (°C)

28 24 20 16 12 8 4

AURAY

BMO

02

03

04

05

06

07

08

09

10

11

12

Figure 13.13 Effect of temperature on the outbreak of summer mortality in a single cohort of Pacific oysters deployed as juveniles at three locations on the French coast in 2003. Shaded areas denote the mortality period. Note that the start of mortality at all three sites coincided with a temperature threshold of 19°C, despite the fact that they are geographically widely separated. BDV, Baie des Veys on the English Channel; AURAY, Riviere d’Auray on the northern Atlantic coast; BMO, Marennes Oléron Bassin on the mid-Atlantic coast. (Redrawn from Samain and McCombie 2008.)

may not survive as well as diploid oysters in many environments (Cheney et al. 2000; Gagnaire et al. 2006).

Summary In this chapter, we have illustrated the many types of interactions between shellfish aquaculture, disease, and the environment. We have presented case studies of environmental factors, both physical and biological, that have important influences on the health of cultured shellfish and the productivity of shellfish farms. On the other hand, concern about the impact of aquaculture on the environment (Chapter 9 in this book) has increased in recent years. There is little evidence that shellfish culture has had a negative impact on the environment; indeed, because adult molluscan shellfish are

largely grown in natural waters, those waters must be of the very highest quality. One area in which shellfish aquaculture has been known to have had a detrimental effect on the environment, however, is through the introduction of disease agents into new areas (i.e., the likely movement of Bonamia ostreae from the United States to Europe in oyster shipments). Thus, we have described the biosecurity structures put in place to minimize such unintentional introductions (Chapter 14 in this book), which are being implemented from the individual farm to the international level. The maintenance of a healthy culture environment, knowledge of how environment interacts with disease agents, and efforts to prevent the spread of infectious disease into new environments are all critical aspects of managing the interaction of shellfish aquaculture and the environment.

386

Shellfish Aquaculture and the Environment

Literature cited Allam, B., Ashton-Alcox, K., and Ford, S.E. 2001. Hemocyte activities associated with resistance to brown ring disease in Ruditapes spp. clams. Developmental and Comparative Immunology 25:365–375. Allam, B., Bushek, D., Pawagi, S., Ragone-Calvo, L., Dove, A., Normant, J., Thiel, J., Joseph, J., Barnes, D., and Ford, S. 2005. Monitoring QPX disease in Raritan—Sandy Hook Bays: results of a 3-year survey program. Journal of Shellfish Research 24(2):637. Anderson, D.M., Burkholder, J.M., Cochlan, W.P., Glibert, P.M., Gobler, C.J., Heil, C.A., Kudela, R.M., Parsons, M.L., Rensel, J.E.J., Townsend, D.W., Trainer, V.L., and Vargo, G.A. 2008. Harmful algal blooms and eutrophication: examining linkages from selected coastal regions of the United States. Harmful Algae 8(1): 39–53. Andrews, J.D. 1980. A review of introductions of exotic oysters and biological planning for new importations. U.S. National Marine Fisheries Service Marine Fisheries Review 42(12):1–11. Andrews, J.D., and Hewatt, W.G. 1957. Oyster mortality studies in Virginia II. The fungus disease caused by Dermocystidium marinum in oysters of Chesapeake Bay. Ecological Monographs 27:1–26. Andrews, J.D., and Ray, S.M. 1988. Management strategies to control the disease caused by Perkinsus marinus. Disease Processes in Marine Bivalve Molluscs. W. S. Fisher. Bethesda, MD, American Fisheries Society 18:257–264. Anthony, K., Causey, B., Conklin, E., Cros, A., Feely, R.A., Guinotte, J., Hofmann, G., Hoffman, J., Jokiel, P., Kleypas, J., Marshall, P., McLeod, E., Salm, R., and Veron, J. 2009. The Honolulu Declaration on ocean acidification and reef management. Prepared and adopted by participants of the Ocean Acidification Workshop, convened by The Nature Conservancy, Hawaii. Arzul, I., Nicolas, J.L., Davison, A.J., and Renault, T. 2001a. French scallops: a new host for ostreid herpesvirus-1. Virology 290(2):342–349. Arzul, I., Renault, T., and Lipart, C. 2001b. Experimental herpes-like viral infections in marine bivalves: demonstration of interspecies

transmission. Diseases of Aquatic Organisms 46(1):1–6. Audemard, C., Carnegie, R.B., Stokes, N.A., Bishop, M.J., Peterson, C.H., and Burreson, E.M. 2008. Effects of salinity on Bonamia sp survival in the Asian oyster Crassostrea ariakensis. Journal of Shellfish Research 27(3): 535–540. Balouet, G., Poder, M., and Cahour, A. 1983. Haemocytic parasitosis: morphology and pathology of lesions in the French flat oyster, Ostrea edulis L. Aquaculture 34:1–14. Barber, B.J., Davis, C.V., and Crosby, M.A. 1998. Cultured oysters, Crassostrea virginica, genetically selected for fast growth in the Damariscotta River, Maine, are resistant to mortality caused by Juvenile Oyster Disease (JOD). Journal of Shellfish Research 17(4):1171–1175. Bishop, M.J., Carnegie, R.B., Stokes, N.A., Peterson, C.H., and Burreson, E.M. 2006. Complications of a non-native oyster introduction: facilitation of a local parasite. Marine Ecology Progress Series 325:145–152. Boettcher, K.J., Geaghan, K.K., Maloy, A.P., and Barber, B.J. 2005. Roseovarius crassostreae sp. nov., a member of the Roseobacter clade and the apparent cause of juvenile oyster disease (JOD) in cultured Eastern oysters. International Journal of Systematic and Evolutionary Microbiology 55:1531–1537. Boettcher, K., Smolowitz, R., Lewis, E.J., Allam, B., Dickerson, H., Ford, S., Hug, A., Reece, K., Rivara, G., and Woodley, C.M. 2006. Juvenile Oyster Disease (JOD) in Crassostrea virginica: synthesis of knowledge and recommendations. Journal of Shellfish Research 25(2):683– 686. Boudry, P., Dégremont, L., and Haffray, P. 2008. Chapter 4.The genetic basis of summer mortality in Pacific oyster spat and potential for improving survival by selective breeding in France. In: Samain, J.F., and McCombie, H. (eds.), Summer Mortality of Pacific Oyster Crassostrea Gigas: The Morest Project. Editions Quae, Versailles, pp. 153–196. Bower, S.M. 1987. Labyrinthuloides haliotidis n. sp. (Protozoa: Labyrinthomorpha), a pathogenic parasite of small juvenile abalone in a British Columbia mariculture facility. Canadian Journal of Zoology 65:1996–2007.

Shellfish diseases and health management 387

Bower, S., Hervio, D., and Meyer, G. 1997. Infectivity of Mikrocytos mackini, the causative agent of Denman Island disease in Pacific oysters Crassostrea gigas, to various species of oysters. Diseases of Aquatic Organisms 29:11–116. Braid, B.A., Moore, J.D., Robbins, T.T., Hedrick, R.P., Tjeerdema, R.S., and Friedman, C.S. 2005. Health and survival of red abalone, Haliotis rufescens, under varying temperature, food supply, and exposure to the agent of withering syndrome. Journal of Invertebrate Pathology 89(3):219–231. Bricelj, V.M., Ford, S.E., Borrero, F.J., Perkins, F.O., Rivara, G., Hillman, R.E., Elston, R.A., and Chang, J. 1992. Unexplained mortalities of hatchery-reared, juvenile oysters, Crassostrea virginica (Gmelin). Journal of Shellfish Research 11(2):331–347. Brothers, C., Marks, E., and Smolowitz, R. 2000. Conditions affecting the growth and zoosporulation of the protistan parasite QPX in culture. Biological Bulletin 199(2):200–201. Brown, C. 1974. A pigment producing pseudomonad which discolors culture containers of embryos of a bivalve molluscs. Chesapeake Science 15(1):17–21. Brown, C. 1981. A study of two shellfish-pathogenic strains isolated from a Long Island hatchery during a recent outbreak of disease. Journal of Shellfish Research 1:83–87. Brown, C., and Losee, E. 1978. Observations on natural and induced epizootics of vibriosis in Crassostrea virginica larvae. Journal of Invertebrate Pathology 31:41–47. Burge, C.A., Griffin, F.J., and Friedman, C.S. 2006. Mortality and herpesvirus infections of the Pacific oyster Crassostrea gigas in Tomales Bay, California, USA. Diseases of Aquatic Organisms 72(1):31–43. Burge, C.A., Judah, L.R., Conquest, L.L., Griffin, F.J., Cheney, D.P., Suhrbier, A., Vadopalas, B., Olin, P.G., Renault, T., and Friedman, C.S. 2007. Summer seed mortality of the Pacific oyster, Crassostrea gigas Thunberg grown in Tomales Bay, California, USA: the influence of oyster stock, planting time, pathogens, and environmental stressors. Journal of Shellfish Research 26(1):163(10). Burreson, E.M., and Ford, S.E. 2004. A review of recent information on the Haplosporidia, with

special reference to Haplosporidium nelsoni (MSX disease). Aquatic Living Resources 17(4):499–517. Burreson, E.M., and Ragone Calvo, L.M. 1996. Epizootiology of Perkinsus marinus disease of oysters in Chesapeake Bay, with emphasis on data since 1985. Journal of Shellfish Research 15(1):17–34. Burreson, E.M., Stokes, N.A., and Friedman, C.S. 2000. Increased virulence in an introduced pathogen: Haplosporidium nelsoni (MSX) in the eastern oyster Crassostrea virginica. Journal of Aquatic Animal Health 12:1–8. Burreson, E.M., Stokes, N.A., Carnegie, R.B., and Bishop, M.J. 2004. Bonamia sp. (Haplosporidia) found in nonnative oysters Crassostrea ariakensis in Bogue Sound, North Carolina. Journal of Aquatic Animal Health 16(1):1–9. Bushek, D., Ford, S.E., and Allen, S.K., Jr. 1994. Evaluation of methods using Ray’s fluid thioglycollate medium for diagnosis of Perkinsus marinus infection in the eastern oyster, Crassostrea virginica. Annual Review of Fish Diseases 4:201–217. Calvo, G.W., Luckenbach, M.W., Allen, S.K., and Burreson, E.M. 2001. A comparative field study of Crassostrea ariakensis (Fujita 1913) and Crassostrea virginica (Gmelin 1791) in relation to salinity in Virginia. Journal of Shellfish Research 20(1):221–229. Carnegie, R.B., and Cochennec-Laureau, N. 2004. Microcell parasites of oysters: recent insights and future trends. Aquatic Living Resources 17(4):519–528. Carnegie, R.B., Stokes, N.A., Audemard, C., Bishop, M.J., Wilbur, A.E., Alphin, T.D., Posey, M.H., Peterson, C.H., and Burreson, E.M. 2008. Strong seasonality of Bonamia sp infection and induced Crassostrea ariakensis mortality in Bogue and Masonboro Sounds, North Carolina, USA. Journal of Invertebrate Pathology 98(3):335–343. Cheney, D.P., MacDonald, B.F., and Elston, R.A. 2000. Summer mortality of Pacific oysters, Crassostrea gigas (Thunberg): initial findings on multiple environmental stressors in Puget Sound, Washington, 1998. Journal of Shellfish Research 19(1):353–359. Chu, F.-L.E., La, J.F., and Burreson, C.S. 1993. Perkinsus marinus susceptibility and

388

Shellfish Aquaculture and the Environment

defense-related activities in eastern oysters, Crassostrea virginica: salinity effects. Journal of Invertebrate Pathology 62:226–232. Cigarría, J., and Elston, R. 1997. Independent introduction of Bonamia ostreae, a parasite of Ostrea edulis, to Spain. Diseases of Aquatic Organisms 29(2):157–158. Comps, M. 1970. La maladie des branchies chez les huitres du genre Crassostrea, cracteristiques et evolutions des alterations, precessus de cictrisation. Revue des Travaux de l’Institut des Peches Maritimes 34(1):23–44. Comps, M. 1988. Epizootic Disease of Oysters Associated with Viral Infections. Disease Processes in Marine Bivalve Molluscs. W. S. Fisher.American Fisheries Society, Special Publication 18. Bethesda, MA. Comps, M., and Bonami, J.R. 1977. Viral infection associated with mortality in the oyster Crassostrea gigas Thunberg. Comptes Rendus Hebdomadaires des Seances de l’Academie des Sciences. Serie D: Sciences Naturelles 285(11): 1139–1140. Comps, M., and Duthoit, J.L. 1976. Infection virale associee a la “maladie de branchies” de l’Huitre portugaise Crassostrea angulata Lmk. Comptes Rendus Hebdomadaire des Seances de l’Academie des Sciences, Serie D, Sciences Naturelles 283:1595–1597. Cook, T., Folli, M., Klinck, J., Ford, S., and Miller, J. 1998. The relationship between increasing sea surface temperature and the northward spread of Perkinsus marinus (Dermo) disease epizootics in oysters. Estuarine, Coastal and Shelf Science 46(4):587–597. Culloty, S.C., and Mulcahy, M.F. 1996. Season-, age-, and sex-related variation in the prevalence of bonamiasis in flat oysters (Ostrea edulis L.) on the south coast of Ireland. Aquaculture 144:53–63. Davis, C.V., and Barber, B.J. 1994. Size-dependent mortality in hatchery-reared populations of oysters, Crassostrea virginica (Gmelin 1791), affected by juvenile oyster disease. Journal of Shellfish Research 13(1):137–142. Doney, S.C., Fabry, V.J., Feely, R.A., and Kleypas, J.A. 2009. Ocean acidification: the other CO2 problem. Annual Review of Marine Science 1(1):169–192.

Dove, M.C., and Jesmond, S. 2007. Histological and feeding response of Sydney Rock oysters, Saccostrea glomerata, to acid sulfate soil outflows. Journal of Shellfish Research 26(2):509–518. Dungan, C.F., and Elston, R.A. 1988. Histopathological and ultrastructural characteristics of bacterial destruction of hinge ligaments in cultrued juvenile Pacific oysters, Crassostrea gigas. Aquaculture 72:1–14. Dungan, C.F., and Hamilton, R.M. 1995. Use of a tetrazolium-based cell proliferation assay to measure effects of in vitro conditions on Perkinsus marinus (Apicomplexa) proliferation. Journal of Eukaryotic Microbiology 42(4): 379–388. Dungan, C.F., Elston, R., and Schiewe, M. 1989. Evidence for colonization and destruction of hinge ligaments of cultured juvenile Pacific oysters (Crassostrea gigas) by cytophaga like bacteria. Applied Environmental Microbiology 55:1128–1135. Ellin, R.C. 2000. Planktonic Concentration and Dispersal of the Oyster Prasite Perkinsus Marinus. Marine Science. University of South Carolina, Columbia, Master of Science, p. 95. Elston, R. 1979. Virus-like particles associated with lesions in larval Pacific oysters, Crassostrea gigas. Journal of Invertebrate Pathology 33:71–74. Elston, R.A. 1984. Prevention and management of infectious diseases in intensive mollusc husbandry. Journal of World Mariculture Society 15:284–300. Elston, R.A. 1993. Infectious diseases of the Pacific Oyster, Crassostrea gigas. Annual Review of Fish Diseases 3:259–276. Elston, R.A. 1997. Bivalve mollusc viruses. World Journal of Microbiology & Biotechnology 13(4):393–403. Elston, R.A. 1999. Health Management, Development and Histology of Seed Oysters. World Aquaculture Society, Baton Rouge, LA, USA. Elston, R.A. 2004. Shellfish High Health Program. Shellfish Mariculture in Drakes Estero, Point Reyes National Seashore, California, The National Academies Press, Washington, DC, pp. 119–123.

Shellfish diseases and health management 389

Elston, R.A., and Wilkinson, M.T. 1985. Pathology, management and diagnosis of oyster velar virus diseass (OVVD). Aquaculture 48:189– 210. Elston, R.A., Leibovitz, L., Relyea, D., and Zatila, J. 1981. Diagnosis of Vibriosis in a commercial oyster hatchery epizootic, a Case History. Aquaculture 24:53–62. Elston, R.A., Elliott, E., and Colwell, R.R. 1982. Conchiolin infection and surface coating Vibrio: shell fragility, growth, depression and mortalities in cultured oysters and clams (Crassostrea gigas). Journal of Fish Diseases 5:265–284. Elston, R.A., Farley, C.A., and Kent, M.L. 1986. Occurrence and significance of bonamiasis in European flat oysters Ostrea edulis in North America. Diseases of Aquatic Organisms 2:49–54. Elston, R.A., Kent, M.L., and Wilkinson, M.T. 1987. Resistance of Ostrea edulis to Bonamia ostreae infection. Aquaculture 64:237–242. Elston, R.A., Frelier, P.F., and Cheney, D.P. 1998. Systemic gregarien-like protozoa in juvenile Pacific oysters, Crassostrea gigas (Thunberg 1973). Journal of Shellfish Research 17(4): 1177–1181. Elston, R.A., Frelier, P.F., Cheney, D.P., and Lynn, D. 1999. Invasive orchitophryid ciliate infections in juvenile Pacific and Kumomoto oysters, Crassostrea gigas and C. sikamea. Aquaculture 174:1–14. Elston, R.A., Hasegawa, H., Humphrey, K.L., Polyak, I.K., and Hase, C.C. 2008. Re-emergence of Vibrio tubiashii in bivalve shellfish aquaculture: severity, environmental drivers, geographic extent and management. Diseases of Aquatic Organisms 82(2):119–134. FAO. (2007). World aquaculture production by species groups. ftp://ftp.fao.org/fi/stat/summary/ b-1.pdf Farley, C.A., Wolf, P.H., and Elston, R.A. 1988. A long term study of “Microcell” Disease in oysters with a description of a new genus—Mikrocytos (g.n.) and two new species—Mikrocytos mackini (sp. n.) and Mikrocytos roughleyi (sp. n.). Fishery Bulletin 86(3):581–593. Figueras, A.J. 1991. Bonamia status and its effects in cultured flat oysters in the Ria de Vigo, Galicia (N.W. Spain). Aquaculture 93:225–233.

Flassch, J.P., and Leborgne, Y. 1992. Introduction in Europe, from 1972 to 1980 of the Japanese Manila clam, Tapes philippinarum and the effects on the aquaculture production and natural settlements. ICES Marine Science Symposia, Copenhagen 194:92–96. Ford, S.E. 1985. Effects of salinity on survival of the MSX parasite Haplosporidium nelsoni (Haskin, Stauber, and Mackin) in oysters. Journal of Shellfish Research 2:85–90. Ford, S.E. 1992. Avoiding the spread of disease in commercial culture of molluscs, with special reference to Perkinsus marinus (Dermo) and Haplosporidium nelsoni (MSX). Journal of Shellfish Research 11(2):539–546. Ford, S.E. 1996. Range extension by the oyster parasite Perkinsus marinus into the northeastern US: response to climate change? Journal of Shellfish Research 15(1):45–56. Ford, S.E. 1997. History and present status of molluscan shellfisheries from Barnegat Bay to Delaware Bay. In: MacKenzie, C.L., Jr., Burrell, V.G., Jr., Rosenfield, A., and Hobart, W.L. (eds.), The History, Present Condition, and Future of the Molluscan Fisheries of North and Central America and Europe, Vol. 1, North America, Vol. 127. Department of Commerce, Washington, U.S., pp. 119–140. Ford, S.E., and Borrero, F.J. 2001. Epizootiology and pathology of juvenile oyster disease in the eastern oyster, Crassostrea virginica. Journal of Invertebrate Pathology 78(3):141–154. Ford, S.E., and Haskin, H.H. 1982. History and epizootiology of Haplosporidium nelsoni (MSX), an oyster pathogen, in Delaware Bay, 1957– 1980. Journal of Invertebrate Pathology 40:118–141. Ford, S.E., and Smolowitz, R. 2007. Infection dynamics of an oyster parasite in its newly expanded range. Marine Biology 151:119–133. Ford, S.E., Smolowitz, R., Ragone-Calvo, L., Barber, R.D., and Kraeuter, J.N. 1997. Evidence that QPX (Quahog Parasite Unknown) is not present in hatchery-produced hard clam seed. Journal of Shellfish Research 16(2):519–521. Ford, S.E., Kraeuter, J.N., Barber, R.D., and Mathis, G. 2002. Aquaculture associated factors in QPX disease of hard clams: density and seed source. Aquaculture 208:23–38.

390

Shellfish Aquaculture and the Environment

Friedman, C.S., and Hedrick, R.P. 1991. Pacific oyster nocardiosis: isolation of the bacterium and induction of laboratory infections. Journal of Invertebrate Pathology 57:109–120. Friedman, C.S., and Perkins, F.O. 1994. Range extension of Bonamia ostreae to Maine, U.S.A. Journal of Invertebrate Pathology 64(3):179– 181. Friedman, C.S., McDowell, T., Groff, J.M., Hollibaugh, J.T., Manzer, D., and Hedrick, R.P. 1989. Presence of Bonamia ostrea among populations of the European flat oyster, Ostrea edulis Linne, in California, USA. Journal of Shellfish Research 8:133–137. Friedman, C.S., Beattie, J.H., Elston, R.A., and Hedrick, R.P. 1991. Investigation of the relationship between the presence of a Gram-positive bacterial infection and summer mortality of the Pacific oyster, Crassostrea gigas Thunberg. Aquaculture 94:1–15. Friedman, C.S., Beaman, B.L., Chun, J., Goodfellow, M., Gee, A., and Hedrick, R.P. 1998. Nocardia crassostreae sp. nov., the causal agent of nocardiosis in Pacific oysters. International Journal of Systematic and Evolutionary Microbiology 48:237–246. Friedman, C.S., Andree, K.B., Beauchamp, K.A., Moore, J.D., Robbins, T.T., Shields, J.D., and Hedrick, R.P. 2000. Candidatus Xenohaliotis californiensis, a newly described pathogen of abalone, Haliotis spp., along the west coast of North America. International Journal of Systematic and Evolutionary Microbiology 50:847–855. Friedman, C.S., Estes, R.M., Stokes, N.A., Burge, C.A., Hargove, J.S., Barber, B.J., Elston, R.A., Burreson, E.M., and Reece, K.S. 2005. Herpes virus in juvenile Pacific oysters Crassostrea gigas from Tomales Bay, California, coincides with summer mortality episodes. Diseases of Aquatic Organisms 63(1):33–41. Fuentes, J., Villalba, A., Zapata, C., and Alvarez, G. 1995. Effects of Stock and Culture Environment on Infections by Marteilia refringens and Mytilicola intestinalis in the Mussel Mytilus-galloprovincialis Cultured in Galicia (Nw Spain). Diseases of Aquatic Organisms 21(3):221–226. Gagnaire, B., Soletchnik, P., Madec, P., Gealron, P., Moine, O.L., and Renault, T. 2006. Diploid and

triploid Pacific oysters, Crassostrea gigas (Thunberg), reared at two heights above sediment in Marennes-Oleron Basin, France: difference in mortality, sexual maturation and hemocyte parameters. Aquaculture 254(1–4): 606–616. Glude, J.B. 1975. A summary report of Pacific coast oyster mortality investigations (1965–1972). Proceedings of the 3rd U.S. Japan meeting on aquaculture, Tokyo, Japan. Hervio, D., Bower, S.M., and Meyer, G.R. 1996. Detection, isolation and experimental transmission of Mikrocytos mackini, a microcell parasite of Pacific oysters Crassostrea gigas (Thunberg). Journal of Invertebrate Pathology 67:72– 79. Hine, P.M., Wesney, B., and Hay, B.E. 1992. Herpesvirus associated with mortalities among hatchery-reared larval Pacific oysters, Crassostrea gigas. Diseases of Aquatic Organisms 12(2):135–142. ICES. 1994. ICES Code of Practice on the Introductions and Transfers of Marine Organisms 1994. I. C. f. t. E. o. t. Seas. Copenhagen, Denmark. Imai, T., Numachi, K., Oizumi, J., and Sato, S. 1965. Studies on the mass mortality of the oyster in Matsushima Bay. II Search for the cause of mass mortality and the possibility to prevent it by transplantation experiment. Bull. Tohoku Teg. Fish Res. Lab 25:27–38. Jones, G., and O’Dor, R.K. 1983. Ultrastructural observations on a Thraustochytrid fungus parasitic in the gills of squid (Illex illecebrosus Lesueur). Journal of Parasitology 69(5):903– 911. Katkansky, S.C., and Warner, R.W. 1974. Pacific oyster disease and mortality studies in California. Marine Resources Technical Report. Long Beach, California Department of Fish and Game. Katkansky, S.C., Dahlstrom, W.A., and Warner, R.W. 1969. Observations on survival and growth of European flat oyster, Ostrea edulis, in California. California Department of Fish Game 55(1):69–74. Kent, M.L., Elston, R.A., Nerad, T.A., and Sawyer, T.K. 1987. An Isonema-like flagellate (protozoa: mastigophora) infection in larval geoduck clams, Panope abrupta. Journal of Invertebrate Pathology 50:221–229.

Shellfish diseases and health management 391

Koganezawa, A. 1975. Present status of studies on the mass mortality of cultured oysters in Japan and its prevention. Proceedings of the 3rd U.S. Japan Meeting on Aquaculture, Tokyo, Japan, Fishery Agency, Japanese Government, and Japan Sea Regional Fisheries Research Laboratory. La Peyre, M., Casas, S., and Peyre, J.L. 2006. Salinity effects on viability, metabolic activity and proliferation of three Perkinsus species. Diseases of Aquatic Organisms 71(1):59–74. Lambert, C., Moal, J., Moullac, G.L., and Pouvreau, S. 2008. Chapter 2. Mortality risks associated with physiological traits of oysters during reproduction. In: Samain, J.F., and McCombie, H. (eds.), Summer Mortality of Pacific Oyster Crassostrea Gigas: the Morest Project. Editions Quae, Versailles, pp. 63–106. LeDeuff, R.M., Renault, T., and Gerard, A. 1996. Effects of temperature on herpes-like virus detection among hatchery-reared larval Pacific oyster Crassostrea gigas. Diseases of Aquatic Organisms 24(2):149–157. Lee, M., Taylor, G.T., Bricelj, V.M., Ford, S.E., and Zahn, S. 1996. Evaluation of Vibrio spp. and microplankton blooms as causative agents of Juvenile Oyster Disease in Crassostrea virginica (Gmelin). Journal of Shellfish Research 15(2):319–329. Lewis, E.J. 2001. Juvenile oyster disease (JOD) and management strategies: a review. Bulletin of the National Research Institute of Aquaculture Supplement 5:101–109. Lipovsky, V.P., and Chew, K.K. 1972. Mortality of Pacific oysters, C. gigas: the influence of temperature and enriched seawater on survival. Proceedings of the National Shellfisheries Association 62: 128–140. Loosanoff, V.L. 1955. The European oyster in American waters. Science 121:119–121. Lyons, M.M., Smolowitz, R., Gomez-Chiarri, M., and Ward, J.E. 2007. Epizootiology of Quahog Parasite unknown (QPX) disease in northern quahogs (= hard clams) Mercenaria mercenaria. Journal of Shellfish Research 26(2):371–381. MacCallum, G.S., and McGladdery, S.E. 2000. Quahog Parasite Unknown (QPX) in the northern quahog Mercenaria mercenaria (Linnaeus, 1758) and M. mercenaria var. notata from Atlantic Canada, survey results from three mari-

time provinces. Journal of Shellfish Research 19(1):43–50. MacKenzie, C.L.J. 1997. The molluscan fisheries of Chesapeake Bay. In: MacKenzie, C.L., Jr., Burrell, V.G., Jr., Rosenfield, A., and Hobart, W.L. (eds.), The History, Present Condition, and Future of the Molluscan Fisheries of North and Central America and Europe, Vol. 1, North America, Vol. 127. Department of Commerce, Washington, U.S.. Mackin, J.G. 1956. Dermocystidium marinum and salinity. Proceedings of the National Shellfisheries Association 46: 116–133. Mackin, J.G. 1962. Oyster diseases caused by Dermocystidium marinum and other microorganisms in Louisiana. In: Mackin, J.G., and Hopkins, S.H. (eds.), Studies on Oysters in Relation to the Oil Industry, Vol. 7. Publication of the Institute of Marine Science, Texas A & M University, pp. 132–229. Mackin, J.G., and Hopkins, S.H. 1962. Studies on oyster mortality in relation to natural environments and to oil fields in Louisiana. In: Mackin, J.G., and Hopkins, S.H. (eds.), Studies on Oysters in Relation to the Oil Industry, Vol. 7. Publication of the Institute of Marine Science, Texas A & M University, pp. 1–131. Maes, P., and Paillard, C. 1992. Effect du Vibrio P1, pathogene de Ruditapes philippinarum, sur d’autres espèces de bivalves. Les Molluscques Marins, Biologies et Aquaculture. IFREMER, Actes de Colloques 14:141–148. Mallet, A.L., Carver, C.E.A., and Freeman, K.R. 1990. Summer mortality of the blue mussel in eastern Canada: spacial, temporal, stock and age variation. Marine Ecology Progress Series 67:35–41. Maloy, A.P., Ford, S.E., Karney, R., and Boettcher, K.J. 2007. Roseovarius crassostreae, the etiological agent of Juvenile Oyster Disease in Crassostrea virginica. Aquaculture 269:71– 83. Margalef, R. 1978. Life-forms of phytoplankton as survival alternatives in an unstable environment. Oceanologica Acta 1:493–590. Meyers, T.R. 1981. Endemic diseases of cultured shellfish of Long Island, New York: adult and juvenile American oysters (Crassostrea virginica) and hard clams (Mercenaria mercenaria). Aquaculture 22:305–330.

392

Shellfish Aquaculture and the Environment

Meyers, T.R., Burton, R., Evans, W., and Starkey, N. 2009. Detection of viruses and virus-like particles in four species of wild and farmed bivalve molluscs in Alaska, USA from 1987 to 2009. Diseases of Aquatic Organisms 88: 1–12. Montes, J., Villalba, A., Lopez, M.C., Carballal, M.J., and Mourelle, S.G. 1991. Bonamiasis in native flat oysters (Ostrea edulis L) from 2 intertidal beds of the Ortigueira Estuary (Galicia, NW Spain) with different histories of oyster culture. Aquaculture 93(3):213–224. Moore, J.D., Robbins, T.T., and Friedman, C.S. 2000. Withering syndrome in farmed red abalone Haliotis rufescens: thermal induction and association with a gastrointestinal Rickettsiales-like prokaryote. Journal of Aquatic Animal Health 12(1):26–34. Moore, J.D., Robbins, T.T., Hedrick, R.P., and Friedman, C.S. 2001. Transmission of the Rickettsiales-like prokaryote “Candidatus xenohaliotis californiensis” and its role in Withering syndrome of California abalone, Haliotis spp. Journal of Shellfish Research 20(2):867– 874. Mori, K. 1979. Effects of artificial eutrophication on the metabolism of the Japanese oyster C. gigas. Marine Biology 53:361–369. Mori, K., Tamate, H., Imai, T., and Itikawa, O. 1965. Changes in the metabolism of lipids and glycogen of the oysters during the stages of sexual maturation and spawning. Bull. Tohoku Teg. Fish Res. Lab 25:65–88. Moss, J.A., Burreson, E.M., and Reece, K.S. 2006. Advanced Perkinsus marinus infections in Crassostrea ariakensis maintained under laboratory conditions. Journal of Shellfish Research 25(1):65–72. Myrand, B., and Gaudreault, J. 1995. Summer mortality of blue mussels (Mytilus edulis Linneaus, 1758) in the Magdalen Islands (Southern Gulf of St Lawrence, Canada). Journal of Shellfish Research 14(2):395–404. National Research Council. 2004. Nonnative Oysters in the Chesapeake Bay. National Academy Press, Washington, DC. Newell, C., and Lutz, R. 1991. Growth and survival of cultured mussels in Maine. Bulletin of the Aquaculture Association of Canada 91(2): 47–53.

Newman, M.W. 1971. A parasite and disease survey of Connecticut oysters. Proceedings of the National Shellfisheries Association 61:59–63. Nicolas, J.L., Comps, M., and Cochennec, N. 1992. Herpes-like virus infecting Pacific oyster larvae, Crassostrea gigas. Bulletin of the European Association of Fish Pathologists 12(1):11. OIE. (2009a). Aquatic animal health code. Zoning and Compartmentalisation. www.oie.int/eng/ normes/fcode/en_chapitre_1.4.1.htm OIE. (2009b). Aquatic animal health code. www. oie.int/eng/normes/fmanual/A_summry.htm OIE. (2009c). Manual of diagnostic tests for aquatic animals. www.oie.int/eng/normes/fcode/en_sommaire.htm Oliver, L.M., Fisher, W.S., Ford, S.E., Burreson, E.M., Calvo, L.M., Sutton, E.B., and Gandy, J.A. 1998. Perkinsus marinus tissue distribution and seasonal variation in oysters (Crassostrea virginica) from Florida, Virginia, and New York. Diseases of Aquatic Organisms 34:51–61. Paillard, C. 2004. A short review of brown ring disease, a vibriosis affecting clams, Ruditapes philippinarum and Ruditapes decussatus. Aquatic Living Resources 17:467–475. Paillard, C., Maes, P., and Oubella, R. 1994. Brown Ring Disease in Clams. Annual Review of Fish Diseases 4:219–240. Paillard, C., Roux, F.L., and Borreg, J.J. 2004. Bacterial disease in marine bivalves, a review of recent studies: trends and evolution. Aquatic Living Resources 17(4):477–498. Park, K.I., Paillard, C., Chevalier, P.L., and Choi, K.S. 2006. Report on the occurrence of brown ring disease (BRD) in Manila clam, Ruditapes philippinarum, on the west coast of Korea. Aquaculture 255(1–4):610–613. Perdue, J.A., Beattie, J.H., and Chew, K.K. 1981. Some relationships between gametogenic cycle and summer mortality phenomenon in the Pacific oyster (Crassostrea gigas) in Washington state. Journal of Shellfish Research 1:9–16. Perrigault, M., Bugge, D.M., and Allam, B. 2010. Effect of environmental factors on survival and growth of quahog parasite unknown (QPX) in vitro. Journal of Invertebrate Pathology 104:83–89. Plunkett, L., and Hidu, H. 1978. The role of Uronema marinum (Protozoa) in oyster hatchery production. Aquaculture 15:219–235.

Shellfish diseases and health management 393

Poder, M., Cahour, A., and Balouet, G. 1982. Hemocytic parasitosis in European oyster Ostrea edulis L.: pathology and contamination. Proceedings, IIIrd International Colloquium on Invertebrate Pathology, Proc. 15th Annual Meeting of the Society for Invertebrate Pathology, University of Sussex, Brighton, U.K. Polglase, J.L. 1980. A preliminary report on the Thraustochytrid(s) and Labyrinthulid(s) associated with a pathological condition in the lesser octopus Eledone cirrhosa. Botanica Marina 23:699–706. Porter, D. 1990. Phylum Labyrinthomycota. In: Margulis, L., Corliss, J.O., Melkonian, M., and Chapman, D.J. (eds.), Handbook of Protoctista. Jones and Bartlett, Boston, pp. 388–398. Ragone Calvo, L., and Burreson, E.M. 1994. Characterization of overwintering infections of Perkinsus marinus (Apicomplexa) in Chesapeake Bay oysters. Journal of Shellfish Research 13(1):123–130. Ragone Calvo, L.M., Walker, J.G., and Burreson, E.M. 1998. Prevalence and distribution of QPX, Quahog Parasite Unknown, in hard clams, Mercenaria mercenaria, in Virginia, USA. Diseases of Aquatic Organisms 33:209– 219. Ragone Calvo, L.M., Elston, R.A., Reece, K.S., and Burreson, E.M. 2002. Investigations of An Isonema-like Flageggate Causing Mortality in Larval Hard Clams, Mercenaria Mercenaria. Fourth International Symposium on Aquatic Animal Health, New Orleans, LA, USA. Ragone Calvo, L.M., Ford, S.E., Kraeuter, J.N., Leavitt, D.F., Smolowitz, R., and Burreson, E.M. 2007. Influence of host genetic origin and geographic location on QPX disease in hard clams, Mercenaria mercenaria. Journal of Shellfish Research 26(1):109–119. Ray, S.M. 1954. Biological Studies of Dermocystidium Marinum, A Fungus Parasite of Oysters. Rice Institute, Houston, TX. Renault, T., Cochennec, N., Le, R.M., and Chollet, B. 1994. Herpes-like virus infecting Japanese oyster (Crassostrea gigas) spat. Bulletin of the European Association of Fish Pathologists 14:64–66. Renault, T., Deuff, R.M.L., Chollet, B., Cochennec, N., and Gerard, A. 2000. Concomitant herpeslike infections in hatchery-reared larvae and

nursery-cultured spat Crassostrea gigas and Ostrea edulis. Diseases of Aquatic Organisms 42:173–183. Renault, T., Lipart, C., and Arzul, I. 2001a. A herpes-like virus infecting Crassostrea gigas and Ruditapes philippinarum larvae in France. Journal of Fish Diseases 24(6):369–376. Renault, T., Lipart, C., and Arzul, I. 2001b. A herpes-like virus infects a non-ostreid bivalve species: virus replication in Ruditapes philippinarum larvae. Diseases of Aquatic Organisms 45(1):1–7. Rivara, G., and Czyzyk, S. 1995. Effects of flow and stocking density on growth rates and survival of eastern oysters with JOD. Journal of Shellfish Research 14(1):247. Rosenfield, A., and Kern, F.G. 1979. Molluscan imports and the potential for introduction of disease organisms. In: Mann, R. (ed.), Exotic Species in Mariculture. MIT Press, Cambridge, MA and London, England, pp. 165–191. Ruiz, G.M., Lorda, J., Arnwine, A., and Lion, K. 2006. Shipping patterns associated with the Panama Canal: effects on biotic exchange? Gollasch, S., Galil, B.S., and Cohen, A.N. (eds.), Bridging Divides: Maritime Canals As Invasion Corridors. Kluwer Press, Dordrecht, pp. 113–126. Samain, J.F., Ropert, M., Bedier, E., Soletchnick, P., Mazurié, J., Le Coz, F., Blin, J.L., Costil, K., Mille, D., Trintignac, P., Boudry, P., Haffray, P., Bacher, C., Grangeré, K., Pouvreau, S., Bourles, Y., Sylvand, B., Misko, P., Gohin, F., and Woerther, P. 2008. A synthesis of the Mortest project and recommendations for forecasting and managing oyster summer mortalities. In: Samain, J.F., and McCombie, H. (eds.), Summer Mortality of Pacific Oyster Crassostrea Gigas: The Morest Project. Editions Quae, Versailles, pp. 307–348. Sauvage, C., Pepin, J.F., Lapegue, S., Boudry, P., and Renault, T. 2009. Ostreid herpes virus-1 infection in families of the Pacific oyster, Crassostrea gigas, during a summer mortality outbreak: differences in viral DNA detection and quantification using real-time PCR. Virus Research 142:181–187. Smayda, T.J. 1997. Harmful algal blooms: their ecophysiology and general relevance to phyto-

394

Shellfish Aquaculture and the Environment

plankton blooms in the sea. Limnology and Oceanography 42(5):1137–1153. Smolowitz, R., Leavitt, D., and Perkins, F. 1998. Observations of a protistan disease similar to QPX in Mercenaria mercenaria (hard clams) from the coast of Massachusetts. Journal of Invertebrate Pathology 71:9–25. Soletchnik, P., Lambert, C., and Costil, K. 2005. Summer mortality of Crassostrea gigas (Thunberg) in relation to environmental rearing conditions. Journal of Shellfish Research 24(1):197–207. Tubiash, H.S., Chanley, P.E., and Leifson, E. 1965. Bacillary necrosis, a disease of larval and juvenile bivalve molluscs. Journal of Bacteriology 103(1):272–273. Tubiash, H.S., Colwell, R.R., and Sakazaki, R. 1970. Marine vibrios associated with bacillary necrosis, a disease of larval and juvenile bivalve molluscs. Journal of Bacteriology 103:272– 273. Tun, K.L., Shimizu, Y., Yamanoi, H., Yoshinaga, T., and Ogawa, K. 2008. Seasonality in the infection and invasion of Marteilioides chungnuensis in the Pacific oyster Crassostrea gigas. Diseases of Aquatic Organisms 80(2):157–165. Vilchis, L.I., Tegner, M.J., Moore, J.D., Friedman, C.S., Riser, K.L., Robbins, T.T., and Dayton, P.K. 2005. Ocean warming effects on growth, reproduction, and survivorship of Southern California abalone. Ecological Applications 15(2):469– 480.

Villalba, A., Casas, S.M., Lopez, C., and Carballal, M.J. 2005. Study of perkinsosis in the carpet shell clam Tapes decussatus in Galicia (NW Spain). II. Temporal pattern of disease dynamics and association with clam mortality. Diseases of Aquatic Organisms 65(3):257–267. Wang, C.C., and Newton, A. 1969. Iron transport in Escherichia coli: roles of energy-dependent uptake and 2,3-dihydroxybenzoylserine. Journal of Bacteriology 98(3):1142–1150. Watson, S.-A., Southgate, P.C., Tyler, P.A., and Peck, L.S. 2009. Early Larval Development of the Sydney Rock Oyster Saccostrea glomerata Under Near-Future Predictions of CO2-Driven Ocean Acidification. Journal of Shellfish Research 28(3):431–437. Whyte, S.K., Cawthorn, R.J., and McGladdery, S.E. 1994. QPX (Quahaug Parasite X) a pathogen of northern quahaug Mercenaria mercenaria from the Gulf of St. Lawrence, Canada. Diseases of Aquatic Organisms 19(2):129–136. Xiao, J., Ford, S.E., Yang, H., Zhang, G., Zhang, F., and Guo, X. 2005. Studies on mass summer mortality of cultured Zhikong scallops (Chlamys farreri Jones et Preston) in China. Aquaculture 250:602–615. Zabaleta, A.I., and Barber, B.J. 1996. Prevalence, intensity, and detection of Bonamia ostrea in Ostrea edulis L. in the Damariscotta River area, Mine. Journal of Shellfish Research 15:395– 400.

Chapter 14

Marine invaders and bivalve aquaculture: sources, impacts, and consequences Dianna K. Padilla, Michael J. McCann, and Sandra E. Shumway

Introduction Since the earliest times of human travel, people have transported species beyond their native ranges. Some introductions have been deliberate for food and trade, while others have been an unintentional consequence of species being transported with the movement of humans and goods. Historically, this was in large part due to a lack of knowledge or understanding of the potential dangers of such introductions, especially the ecological and economic damage that can occur when species are transported to areas outside of communities and systems with which they share a long-term evolutionary history. For example, starlings were deliberately introduced to Central Park in 1890 as

homage to Shakespeare, but are now a major pest, causing $800 million in agricultural damages annually (Pimentel et al. 2005). The eastern gray squirrel (Sciurus carolinensis) was first introduced to Britain in 1876 as a living garden ornament, and became major pests both in Britain and Europe. This introduction is now responsible for economic losses and extensive ecological damage across Britain and Europe (Gurnell et al. 2004). With globalization and increased trade, there has been an increase in the spread and impacts of species that have been introduced outside of their native range. As these introductions begin to have major impacts on ecosystem services on which we depend, such as species cultivated in aquaculture, greater

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 395

396

Shellfish Aquaculture and the Environment

interest has been generated on controlling the spread and impacts of nonnative species. The heightened awareness of the problems associated with nonnative introductions has resulted in more care and often regulations to prevent the unintentional spread of macrospecies beyond their native ranges. The introduction and spread of nonnative species has garnered much recent attention from scientists, managers, and industry on virtually every continent, as it is one of the most pressing environmental issues around the globe. The environmental and economic costs of introductions of nonnative species introductions are enormous and increasing daily. A recent estimate of the total annual cost of nonnative species introductions in the United States, in terms of losses and damages as well as control costs, exceeds $120 billion (Pimentel et al. 2005). In spite of awareness and regulations, new species introductions and the spread of existing pest species continue. Several terms have been used to describe such species of concern. They have been variously described as exotic species, alien species, introduced species, and invasive species (Colautti and Richardson 2009). Some authors have tried to make distinctions between species that are known to have economic impacts or those that behave differently in new areas as compared with their native habitats (Valéry et al. 2008). The economic and environmental impacts of most species have not, however, been assessed, and there are often long lag times between initial introductions and spread and the measured impacts for many species (e.g., Klinger et al. 2006; Blanchard 2009; Karatayev et al. 2009). Here were are considering all species that are out of place in time and space relative to their natural distribution, and will refer to them as introduced species or invaders. These species have been transported beyond their native range by human activities, rather than through natural dispersal. In marine waters, aquaculture has a long history, dating back to oyster cultivation by

the Romans (Gunther 1897; Chew 1990). Oysters and other marine species have been transported for aquaculture since at least the 1700s (reviewed in Food and Agriculture Organization (FAO) 1997; Chew 1990; Ruesink et al. 2005; McKindsey et al. 2007). Along with the transport of species for aquaculture there has been the unintentional transport of associated species (e.g., commensals, attached organisms, parasites, and diseases) to new waters. Charles Elton (1958) noted that aquaculture, especially oyster aquaculture, was particularly important for the spread of nonnative species. He states that, “But the greatest agency of all that spreads marine animals to new quarters of the world must be the business of oyster culture.” He goes on to say that, “The moving about, without particularly stringent precautions, of masses of oysters was bound to spread other species as well.” Why should we care about introduced species? As mentioned above, these species can cause considerable direct economic and environmental damage. They can impact human activities or structures (e.g., fouling humanmade and human-used structures), impact species of concern to humans (e.g., commercial species), or disrupt natural communities and the ecosystem services they provide. The spread of nonnative species can also have indirect effects. They can facilitate the spread of harmful algae or disease agents that are toxic or harmful to humans or other important species (e.g., Lilly et al. 2002; Cohen and Zabin 2009; Hégaret et al. 2009; Shumway, unpublished data; see also Chapter 13 in this book). For example, the shellfish disease caused by the protozoan Bonamia has been introduced to different shores with the transport of shellfish (Friedman and Perkins 1994; Cigarría and Elston 1997; Bishop et al. 2006), and this disease can have devastating impacts on local native fisheries where it is introduced (McArdle et al. 1991). Presently, shipping, ballast water, and hull fouling are the primary sources of new species

Marine invaders and bivalve aquaculture

introductions and the spread of introduced species in aquatic habitats (Cohen and Carlton 1998). Many species introduced through shipping activities have large impacts on aquaculture species and activities associated with aquaculture. Shipping is not, however, the only source of introduced pests. Aquaculture is the second leading source of introduced species (Ruiz et al. 1997). In many cases, other species are transported outside of their native range with aquaculture species (Mann 1979; Critchley and Dijkema 1984; Blanchard 1997; Cohen and Zabin 2009). In other cases, the aquaculture species themselves can escape culture and spread from where they are initially introduced (reviewed in McKindsey et al. 2007). Rather than providing a comprehensive history of species introductions, this chapter is intended to provide an overview of the practices surrounding introductions and the impacts of introductions on shellfish aquaculture. This chapter will also keep an eye toward improved management decisions and aquaculture practices that will reduce the likelihood of unwanted species introductions and their cascading negative impacts. Here we present information on shellfish aquaculture species and species transported in association with aquaculture activities that have become introduced outside of their native range, as well as introduced species that impact shellfish aquaculture. We also address management and policy strategies and needs to minimize the role of aquaculture as a source of nonnative introductions, as well as strategies that will help the aquaculture industry by reducing the introduction and spread of species that impact shellfish aquaculture.

Introduced shellfish from aquaculture The “Blue Revolution” of shellfish aquaculture is widely recognized as an important source of food and an important tool for pro-

397

tecting wild populations of commercial species, which often fall victim to overharvest (Muir 2005; Costa-Pierce 2010). Many authors have reviewed the history of the development of aquaculture (e.g., Shatkin et al. 1997, and references therein; Kurlansky 2006). Initially, the majority of shellfish introductions were deliberate, for replacement fisheries for a collapsed native species fishery or to develop a new industry, such as the introduction of Argopecten irradians to China (see Chew 1990). Shellfish represent one-quarter of all aquaculture production worldwide (FAO 1997; USDA 2009). In the U.S. Department of Agriculture (USDA) census of aquaculture, $203 million of the $1.1 billion aquaculture industry was associated with molluscan shellfish aquaculture (USDA 2009). By far, the fastest growing sector of the aquaculture industry is the culture of molluscs, including primarily oysters, clams, and scallops, which increased ∼130% from 1998 to 2005 (USDA 2009). As the shellfish industry has grown and hatchery and husbandry techniques have been developed, aquaculture has spread, and the number of species cultured and places where aquaculture has developed has grown accordingly. The species most widely introduced around the world for shellfish aquaculture is the Pacific oyster, Crassostrea gigas (Kurlansky 2006; McKindsey et al. 2007). This species was initially introduced to the Pacific coast of North America from Japan in 1903 to replace the fishery for the native Olympic oyster, Ostrea lurida, which was depleted by the late 1800s (Chew 1990; see also the Journal of Shellfish Research 28[1] and reviews therein). Olympic oysters form reefs, but are small bodied. They are slow growing, and have relatively low fecundity and limited dispersal potential (Baker 1995). Commercial harvesting rapidly decimated local populations and a replacement oyster was sought to maintain the growing oyster industry, and to supply oysters to the U.S. East Coast where populations of the native Crassostrea virginica were suffering

398

Shellfish Aquaculture and the Environment

from overharvest (Mann 1979; Quayle 1988). Similar stories have played out around the world as populations of native shellfish are overharvested and the demand for shellfish has increased. With growing demand, there has been a steady increase in efforts to indentify new shellfish species for commercial cultures. At present, there are at least 63 species of bivalves in aquaculture somewhere in the world. Table 14.1 provides a list of bivalve species grown in aquaculture, their native range, and the countries or continents in which they are presently grown in culture. Of the 63 bivalve species cultured, 15 (24%) are grown on continents outside of their native range (Table 14.1). Many of the species that are grown only on their native continent are grown in areas outside of their native range, where natural dispersal would never carry them. Of the 15 species grown on continents outside of their native range, 33% (5 of 15) have been documented to have established feral populations and are having negative impacts on the systems where they have invaded (Table 14.2). Activities associated with shellfish aquaculture have been responsible for the introduction of 48 additional noncultured species to new regions of the world (Table 14.3). Once species are introduced outside of their native range, they may continue to spread via the dispersal of adults or larvae. The new feral populations that are created may then serve as source populations for introductions through other vectors. For example, although Crassostrea gigas was deliberately introduced for culture to many areas in Europe, it appears also to have spread to some areas in Europe via fouled ship hulls (Fletcher and Manfredi 1995; Eno et al. 1997; Eno 1998). Many of the bivalve species grown in aquaculture are important ecosystem engineers, capable of having large impacts on communities and ecosystems where they are found by modifying the physical habitat or ecosystem processes in a way that changes the habitat for

other species (Jones et al. 1994; Cuddington et al. 2007). Oysters are especially important ecosystem engineers and, when in high density, have the ability to modify ecosystems dramatically and alter habitat suitability for other species (see Chapter 9 in this book) Among the aquaculture species that have become introduced outside of their native range, bivalves are by far the most studied. In some cases, these species have been deliberately introduced to establish feral populations for seeding aquaculture, such as Crassostrea gigas in British Columbia, Canada (Quayle 1988), which is frequently considered among the 100 worst invaders worldwide (DAISIE 2008). In other cases, species have escaped aquaculture operations and are now spreading through larval transport, such as Perna viridis (Rajagopal et al. 2006). In the Wadden Sea and in France, recent warming of local waters is believed to be facilitating the spread of Crassostrea gigas (Diederich et al. 2005; Schmidt et al. 2008; Thieltges et al. 2009) and increased invasion, especially through larval transport, is expected with continued climatic change. These feral populations can have large impacts on the systems they invade and can impact other ecologically and commercially important species (e.g., Dubois et al. 2006; Nehls et al. 2006; Rajagopal et al. 2006; Kochmann et al. 2008; Markert et al. 2010). These invasions can also impact marine reserves, a major tool for the protection of marine biodiversity and wild-caught fisheries. Klinger et al. (2006) found that invading Crassostrea gigas in the San Juan Archipelago in Washington State was more abundant in marine reserves than paired control areas outside of reserves. This invasion appears to be impacting biodiversity, especially when oysters are dense (D.K. Padilla, pers. obs.). Surprisingly, few quantitative or experimental studies have examined the impacts of aquaculture escapees on the systems they invade (Table 14.2). This is clearly an area where more research is needed before we can draw

Table 14.1 Bivalve species in aquaculture. Family

Species

Common name

Native range

Nonnative

Escaped

Arcidae

Anadara granosa

Blood cockle

Asia

Scapharca broughtonii

Inflated ark

Asia

Scapharca subcrenata

Half-crenate ark

Asia

Cardiidae

Cerastoderma edule

Common edible cockle

Africa, Europe, Former USSR

Hiatellidae

Panopea abrupta

Pacific geoduck

North America

Mactridae

Mactra glabrata

Smooth mactra

Africa

Mactra veneriformis

Globose clam

Asia

Spisula solidissima

Atlantic surf clam

North America

Myidae

Mya arenaria

Sand gaper

North America, Asia, Europe, Former USSR

North America

Yes

Mytilidae

Aulacomya ater

Cholga mussel

Africa, South America, Oceania

Choromytilus chorus

Choro mussel

South America

Mytilus californianus

Californian mussel

North America

Mytilus chilensis

Chilean mussel

South America

Mytilus coruscus

Korean mussel

Asia, Former USSR

Mytilus edulis

Blue mussel

Africa, North America, South America, Asia, Europe

Mytilus galloprovincialis

Mediterranean mussel

Africa, Asia, Europe, Former USSR

Africa, North America, Asia

Yes

Mytilus planulatus

Australian mussel

Oceania

Perna canaliculus

New Zealand mussel

Oceania

Perna indica

Indian brown mussel

Asia

Perna perna

South American rock mussel

Africa, South America

North America

Perna viridis

Green mussel

Asia, Oceania

North America, Oceania

Yes

399

Table 14.1 (Continued) Family

Species

Common name

Native range

Pectinidae

Aequipecten opercularis

Queen scallop

Africa, Europe

Argopecten irradians

Atlantic bay scallop

North America, South America

Argopecten purpuratus

Peruvian calico scallop

North America, South America

Argopecten ventricosus

Pacific calico scallop

North America, South America

Chlamys farreri

Farrer’s scallop

Asia, Former USSR

Chlamys islandica

Iceland scallop

North America, Europe, Former USSR

Chlamys nobilis

Noble scallop

Asia

Patinopecten yessoensis

Yesso scallop

Asia, Former USSR

Pecten fumatus

Australian southern scallop

Oceania

Pecten maximus

Great Atlantic scallop

Africa, Europe

Pecten novaezelandiae

New Zealand scallop

Oceania

Placopecten magellanicus

Sea scallop

North America

Pinctada fucata

Japanese pearl oyster

Africa, Asia, Oceania

Pinctada margaritifera

Black-lip pearl oyster

Africa, North America, Asia, Oceania

Pinctada maxima

Silver-lip pearl oyster

Asia, Oceania

Pteria penguin

Penguin wing oyster

Asia

Crassostrea belcheri

Lugubrious cupped oyster

Asia

Crassostrea corteziensis

Cortez oyster

North America, South America

Crassostrea gigas

Pacific oyster

Asia

Crassostrea iredalei

Slipper cupped oyster

Asia

Crassostrea madrasensis

Indian backwater oyster

Asia

Pteriidae

Ostreidae

400

Nonnative

Escaped

Asia

North America

Oceania

Africa, North America, South America, Europe, Oceania

Yes

Table 14.1 (Continued) Family

Species

Common name

Native range

Nonnative

Crassostrea rhizophorae

Mangrove cupped oyster

North America, South America

Oceania

Crassostrea rivularis

Suminoe oyster

Asia

North America

Crassostrea virginica

Eastern oyster

North America, South America

North America

Ostrea chilensis

Chilean flat oyster

South America

Ostrea edulis

European flat oyster

Europe

Ostrea lurida

Edible (flat) oyster

North America

Saccostrea commercialis

Sidney rock oyster

Oceania

Saccostrea cuccullata

Hooded oyster

Africa, Asia

Saccostrea echinata

Spiny oyster

Asia, Oceania

Solecurtidae

Sinonovacula constricta

Constricted tagelus

Asia

Tridacnidae

Tridacna derasa

Smooth giant clam

Asia, Oceania

Tridacna gigas

Giant clam

Asia, Oceania

Mercenaria mercenaria

Northern quahog

North America

Meretrix lusoria

Japanese hard clam

Asia

Meretrix meretrix

Asiatic hard clam

Asia

Paphia undulata

Undulate venus

Asia

Protothaca staminea

Pacific littleneck clam

North America

Venerupis decussatus

Grooved carpet shell

Africa, Asia, Europe

Venerupis philippinarum (Ruditapes philippinarum)

Japanese littleneck

North America, Asia

Saxidomus giganteus

Butter clam

North America

Venerupis pullastra

Pullet carpet shell

Africa, Asia, Europe

Veneridae

Escaped

Yes

North America

Europe

Europe

Europe

Species are grouped by family. Columns include the continents where each species is naturally found and grown in aquaculture (“Native range”) and where it has been introduced for aquaculture outside of that native range (“Nonnative”). For both the native range and where they are grown, species are not always found or grown in all areas within each geographic area. “Escaped” designates those species that have been documented to spread beyond where they are grown for aquaculture or have established feral populations. Information on all species from FAO (1996). Information on Mya arenaria, Mytilus galloprovincialis, Perna viridis and Crassostrea gigas also from the U.S. Geological Survey (USGS 2009) and data for Crassostrea virginica also from Coles et al. (1999).

401

Table 14.2 Bivalve aquaculture species that have escaped in regions outside of their native range, and documented impacts of feral populations. Scientific name

Common name

Impact

Species/systems impacted

Reference

Crassostrea gigas

Pacific oyster

Ecosystem engineer, change substrate available for other species, competition for space with benthic species, competition for food with suspension-feeders, overgrowth of benthic species, affect suspended particle concentrations and quality, decrease turbidity and increase light penetration, provide refuge for invertebrates from predators, affect flow and sedimentation, foul water systems

Soft sediment communities, fouling community (ascidians, bryozoans, sponges, hydrozoans, algae), native intertidal mussels, cultivated Crassostrea gigas, other engineering and reef building species including honeycomb worm (Sabellaria alveolata)

Diederich et al. (2005); Cognie et al. (2006); Decottignies et al. (2007a, 2007b); Rodriguez and Ibarra-Obando (2008); Sousa et al. (2009; reviewed in Ruesink et al. 2005)

Crassostrea virginica

Eastern oyster

Ecosystem engineer, affect phytoplankton species composition, biodeposition of waste materials, affect flow and sedimentation, decrease turbidity, increase light penetration, provide refuge for invertebrates from predators, foul water systems

Phytoplankton, benthic invertebrate communities

Mugg Pietros and Rice (2003); Sousa et al. (2009; reviewed in Ruesink et al. 2005)

Mytilus galloprovincialis

Mediterranean mussel

Ecosystem engineer, increase habitat for infaunal species, provide hard substrate, provide refuge for invertebrates, increase recruitment and species richness in some habitats, affect flow and sedimentation, empty shells block softsediment burrowing organisms, increase food supply for intertidal predators, competitive displacement of native species, hybridize with native mussels

Mytilus trossulus, soft-sediment burrowers, tube-building polychaete Gunnarea capensis, limpet Soutellastra granularis, limpet Soutellastra argenwillei, other benthic invertebrates, African black oystercatchers Haematopus moquini

Branch and Steffani (2004); Wonham (2004); Steffani and Branch (2005); Robinson et al. (2005, 2007); Coleman and Hockey (2008); Sousa et al. (2009); Branch et al. (2010)

402

Marine invaders and bivalve aquaculture

403

Table 14.2 (Continued) Scientific name

Common name

Impact

Species/systems impacted

Reference

Ostrea edulis

Edible (flat) oyster

Ecosystem engineer, provide refuge for invertebrates from predators, spread disease

Other populations of Ostrea edulis

da Silva et al. (2005); Sousa et al. (2009); reviewed in Ruesink et al. 2005)

Perna perna

Brown mussel

Ecosystem engineer, provide hard substrate in soft-sediment habitats, provide refuge for invertebrates from predators, affect flow and sedimentation, foul navigation buoys, foul water systems

Benthic invertebrate communities

Rajagopal et al. (2003); Sousa et al. (2009)

Perna viridis

Green mussel

Ecosystem engineer, provide hard substrate, affect water flow and sedimentation, foul power plant heat interchangers, foul water systems, clog crab traps and clam culture bags, foul vessels

Algae, hydroids, tubiculous polychaetes, barnacles, and ascidians, free-living polychaetes, and amphipods

Masilamoni et al. (2003, 2002); Rajagopal et al. (2003); Sousa et al. (2009; reviewed in Rajagopal et al. 2006)

Venerupis philippinarum

Japanese littleneck

Ecosystem engineer, provide refuge from predators, provide hard substrate, affect flow and sedimentation, enhancement of oxygen and solute penetration because of burrowing, increase filtration capacity of bivalves in ecosystem, increase food supply for intertidal predators

Eurasian oystercatcher, Haematopus ostralegus ostralegus, Polydora spp., other benthic invertebrates

Gosling (2003); Pranovi et al. (2006); Caldow et al. (2007); Sousa et al. (2009)

generalizations. Thus far, the majority of impacts appear to be due to the ecosystem engineering effects of these invaders (Padilla 2010). The types of impacts of invasion by feral aquaculture species that have been documented include fouling, overgrowth, or dis-

placement of native benthic species (Reise et al. 2005; Diederich 2006; Nehls et al. 2006; Buttger et al. 2008; Kochmann et al. 2008; Krassoi et al. 2008; Markert et al. 2010) and reduced recruitment of native species and negative impacts on populations of other

Table 14.3 Taxa that have been introduced through bivalve aquaculture, either intentionally or accidentally. Group

Scientific name

Alga (brown)

Introduction

Reference

Rugulopteryx okamurae

Accidental

Verlaque et al. (2009)

Sargassum muticum

Accidental

Critchley and Dijkema (1984); FAO (2005)

Intentional and accidental

Fletcher and Manfredi (1995); Curiel et al. (2001); Global Invasive Species Database (GISD) (2005)

Undaria pinnatifida

Common name

Asian kelp

Alga (diatom)

Coscinodiscus wailesii

Accidental

Laing and Gollasch (2002)

Alga (dinoflagellate)

Alexandrium catenella

Accidental

Lilly et al. (2002)

Accidental

GISD (2005)

Accidental

Langeland et al. (1984)

Accidental

GISD (2005)

Accidental

Langeland et al. (1984)

Accidental

GISD (2005)

Alexandrium minutum

Red tide dinoflagellate

Gonyaulax excavata Gymnodinium catenatum

Chain-forming dinoflagellate

Procentrum minimum Alga (green)

Codium fragile tomentosoides

Dead man’s fingers

Alga (red)

Heterosiphonia japonica

Accidental

Sjøtun et al. (2008); Moore and Harries (2009)

Annelid

Boccardia proboscidea

Accidental

Bailey-Brock (2000)

Polydora sp.

Accidental

FAO (2007)

Polydora nuchalis

Accidental

Bailey-Brock (1990)

Bacteria

Vibrio cholerae

Asiatic cholera

Accidental

GISD (2005)

Bryozoan

Bugula neritina

Brown bryozoan

Accidental

GISD (2005)

Schizoporella errata

Branching bryozoan

Accidental

GISD (2005)

Schizoporella unicornis

Single-horn bryozoan

Accidental

GISD (2005)

Mytilicola orientalis

Parasitic copepod

Accidental

McKindsey et al. (2007)

Rhithropanopeus harrisii

Estuarine mud crab

Accidental

GISD (2005)

Asterias amurensis

Flatbottom sea star

Accidental

GISD (2005)

Crustacean

Echinoderm

404

Table 14.3 (Continued) Group

Scientific name

Common name

Introduction

Mollusc

Batillaria attramentaria

Asian hornsnail

Accidental

Boonea bisuturalis

Two-groove odostome

Accidental

GISD (2005)

Crassostrea gigas

Pacific oyster

Intentional

GISD (2005)

Crassostrea virginica

Eastern oyster

Intentional

McKindsey et al. (2007)

Crepidula fornicata

Common Atlantic slippersnail

Accidental

Minchin et al. (1995); Blanchard (1997); GISD (2005)

Cyclope neritea

Accidental

Le Duff et al. (2009)

Fusinus rostratus

Accidental

Le Duff et al. (2009)

Accidental

GISD (2005)

Gemma gemma

Amethyst gem clam

Reference

Accidental

Le Duff et al. 2009

Mercenaria mercenaria

Northern quahog

Intentional

FAO (2005); Le Duff et al. 2009

Musculista senhousia

Senhouse mussel

Accidental

Bachelet et al. (2009); GISD (2005)

Mya arenaria

Eastern soft-shell clam

Accidental

GISD (2005); Le Duff et al. 2009

Mytilopsis sallei

Black-striped mussel

Accidental

GISD (2005)

Mytilus galloprovincialis

Mediterranean mussel

Accidental

GISD (2005)

Ocinebrellus inornatus

Japanese oyster drill

Accidental

Faasse and Ligthart (2009); Le Duff et al. (2009)

Ostrea edulis

Edible (flat) oyster

Intentional

GISD (2005)

Perna perna

Brown mussel

Intentional and accidental

FAO (2005)

Perna viridis

Green mussel

Intentional and accidental

GISD (2005)

Rangia cuneata

Atlantic rangia

Accidental

GISD (2005)

Rapana venosa

Asian rapa whelk

Accidental

GISD (2005); Le Duff et al. (2009)

Urosalpinx cinerea

Eastern oyster drill

Accidental

Faasse and Ligthart (2009); GISD (2005)

Venerupis philippinarum

Manila clam

Accidental

Cloern (1982); FAO (2005); Le Duff et al. (2009)

Plant

Spartina alterniflora

Smooth cordgrass

Accidental

Chew (1998); Civille et al. (2005)

Protozoan

Perkinsus marinus

Dermo

Accidental

Cohen and Zabin (2009)

Tunicate

Ascidiella aspersa

European sea squirt

Accidental

GISD (2005)

Styela clava

Asian tunicate

Accidental

GISD (2005)

Styela plicata

Leathery tunicate

Accidental

GISD (2005)

Gibbula albida

405

406

Shellfish Aquaculture and the Environment

important native ecosystem engineers (Dubois et al. 2006; Kelly and Volpe 2007; Kelly et al. 2008). Feral populations of aquaculture species have also been found to function as an ecological trap for important native species (Trimble et al. 2009) or to hybridize with native species, disrupting local populations and aquaculture (e.g., Rawson et al. 1999; Wonham 2004; Dias et al. 2009; Shields et al. 2010).

Species moved with aquaculture The movement of species for aquaculture has also resulted in the introduction of a wide range of additional species associated with the target species (Mann 1979; Carlton 1989, 1996, 1999). This has included all types of organisms, from macro animal and algal species to microparasites, including disease agents (Chapter 13), and harmful microalgae (Table 14.3). Over 40 species of bivalves, gastropods, arthropods, echinoderms, crustaceans, and algae have been introduced through activities associated with bivalve shellfish aquaculture. In some cases, these organisms were introduced historically, when the aquaculture industry was in its infancy, and when the ecological and economic dangers of transporting nonnative species to new environments were not known or considered unimportant (Elton 1958). For example, in the early days of oyster aquaculture, large quantities of oysters were collected from natural environments and sent by train or ship to new parts of the world (Mann 1979). As ecosystem engineers, oysters provide habitat for a wide range of both sessile and mobile species. Thus, when oysters were placed in new habitats with the hopes that they would grow and thrive, all of their associates were also moved and introduced. In some cases, the species that were transferred became significant pests of the aquaculture industry that was established. Smooth cordgrass, Spartina alterniflora, is the

primary salt marsh grass on the Atlantic coast of North America. However, this species was accidentally introduced on the West Coast, most likely with oysters transferred from the Atlantic seaboard (Civille et al. 2005). The spread of this invader greatly reduces essential habitat for shellfish and aquaculture, and thus, although it is a protected species in much of its native habitat, it has become a major pest where introduced, especially when it hybridizes with native congeners (Brusati and Grosholz 2008). The Atlantic slipper snail, Crepidula fornicata, is another such species. This snail is a suspension feeder that occurs in very high population densities. It was carried to Europe with the transfer of the eastern oyster, Crassostrea virginica, which was imported as a replacement for the severely overharvested European flat oyster, Ostrea edulis. Although the eastern oyster did not prove to be a good species for aquaculture in Europe, and was replaced by importation of the Pacific oyster, Crassostrea gigas, introduced populations of Crepidula fornicata have flourished (Minchin et al. 1995; Blanchard 1997). This snail is currently a major problem for shellfish aquaculture in many areas of Europe, where it competes with aquaculture species for suspended food. The Japanese littleneck (=Manila clam), Venerupis (Ruditapes) philippinarum, is another species accidentally introduced with the Pacific oyster. Rather than becoming a pest like Crepidula fornicata, the Japanese littleneck has become an important commercial species (Chew 1990). Japanese littlenecks have primarily supported a wild-caught fishery on the U.S. West Coast and Canada, but there has been a recent expansion of aquaculture for this species in North America and Europe (Flassch and Leborgne 1990; Zhu et al. 1999; Ferreira et al. 2009). With the development of hatchery techniques, the movement of wild-caught animals for aquaculture has been greatly reduced as growers are able to reliably produce seed for

Marine invaders and bivalve aquaculture

transplant in the lab, and no longer need to rely on reproduction in the wild or constant restocking from native regions (Quayle 1988). In some cases, live animals or their shells are still moved for aquaculture and restoration purposes (Luckenbach et al. 1999; Hégaret et al. 2008; Cohen and Zabin 2009). Awareness of the problems associated with nonnative introductions has resulted in more care, and in some cases, regulations to prevent the unintentional spread of macrospecies. Microspecies, including harmful algae (see Hégaret et al. 2008), protozoans, and viruses, including some that cause disease, remain a challenge, especially when species are moved within a country with the transfer of aquaculture species. This was the case with the spread of the oyster parasite Perkinsus marinus (dermo), which was spread to Delaware Bay with oysters from the Chesapeake (Ford 1996). Advances are being made to determine best practices to minimize the likelihood of unintentional transfers of species (e.g., Mineur et al. 2007; Hégaret et al. 2008; see section below). Illicit trade in some species continues, and it continues to introduce species to new areas (Verlaque and Latala 1996).

Introduced species that impact aquaculture Many nonnative species that have been introduced in marine and estuarine waters around the world have deleterious impacts on shellfish aquaculture (e.g., Anon 2005; Castilla et al. 2005; Bullard et al. 2007; Lambert 2007) (Table 14.4). Ironically, as mentioned above, some of these species are those that were introduced via aquaculture (Table 14.3). As mentioned above, Crepidula fornicata is very abundant now in its introduced range and competes with native blue mussels, as well as Crassostrea gigas, which is now the major shellfish grown in aquaculture in Europe (Beninger et al. 2007; Blanchard et al. 2008;

407

Blanchard 2009). The oyster drill, Urosalpinx cinerea, was introduced with shellfish aquaculture from the Atlantic coast of North America to the Pacific coast and now is an important predator for Crassostrea gigas aquaculture and is impacting restoration efforts for the native Olympia oyster, Ostrea lurida, in Washington State (Buhle and Ruesink 2009). The green crab, Carcinus maenas, a European native that was introduced to the North American Atlantic coast over 100 years ago, was recently introduced to the Pacific coast of North America in 1995. This invader spread north with warm waters during an El Niño Southern Oscillation (ENSO) event, and populations have persisted as far north as British Columbia, Canada (Behrens Yamada et al. 2005, 2008a). Research is under way to determine the impacts of this invader on shellfish and aquaculture (Behrens Yamada et al. 2008a, 2008b; Behrens Yamada and Kosro 2010). A large number of marine invaders that impact shellfish aquaculture are fouling species (Table 14.4). They include a variety of macroalgae, ascidians (both colonial and solitary), and bryozoans. Ascidians are of special concern as they are increasing in abundance globally and can spread very quickly once introduced (Anon 2005; Bullard et al. 2007; Lambert 2007). They are large bodied and foul cages and other gear, reducing water flow necessary for healthy shellfish (Bullard et al. 2005; Ramsay et al. 2008). They also are very effective suspension-feeders, can directly compete with bivalves for suspended food (Currie et al. 1998), and may serve as a vector for transfer of harmful algal species (Shumway, unpublished). In a recent survey assessing the impacts of fouling species on the economics of aqauculture, Adams et al. (2011) found that efforts to control biofouling cost ∼15% of the total operating costs for individual aquaculture businesses and over 40% of businesses feel that fouling decreases the marketability of their product. With the increased number of

Table 14.4 Introduced species that have been documented to impact bivalve aquaculture, and which aquaculture species or industry they are known to impact. Group

Scientific name

Alga (brown)

Alga (dinoflagellate)

Common name

Species impacted

Impact

Reference

Sargassum muticum

Ostrea edulis; mussel long line

Foul organisms, may foul gear

Critchley and Dijkema (1984); Harries et al. (2007)

Alexandrium minutum

Mussels

Toxic, PSP

Hallegraeff et al. (1988); Delgado et al. (1990)

Gonyaulax excayata

Mussels

Toxic, PSP

Langeland et al. (1984)

Gymnodinium catenatum

Mytilus galloprovincialis

Toxic, PSP

Hallegraeff et al. (1988); Laiño (1991)

Procentrum lima

Mussels

Toxic, PSP

Levasseur et al. (2003)

Procentrum mexicanum

Mussels

Toxic, PSP

Levasseur et al. (2003)

Procentrum minimum

Mussels

Toxic, PSP

Langeland et al. (1984)

Mytilus galloprovincialis; oysters

Increase recruitment and survival, overgrowth, decrease abundance

Trowbridge (1999); Bulleri et al. (2006)

Alga (green)

Codium fragile

Annelid

Imogine mcgrathi

Pincta imbricata

Predation

O’Connor and Newman (2001)

Boccardia proboscidea

Oysters

Bore into shell, cause blisters, increase parasitism risk

Bailey-Brock (2000); National Introduced Marine Pest Information System (NIMPIS) (2002)

Ostrea angasi, Patinopecten yessoensis, Saccostrea commercialis, Saccostrea glomerata

Bore into shell

FAO (2007); Ogburn et al. (2007)

Mytilus edulis

Bore into shell

FAO (2007)

Accumulate in culture ponds

Bailey-Brock (1990)

Bore into shell

Bailey-Brock (1990); Nell (2007)

Polydora spp.

Polydora ciliata

Dead man’s fingers

Mudworm

Polydora nuchalis Polydora websteri

408

Crassostrea gigas, Mytilus galloprovincialis, Ostrea angasi, Pecten fumatus

Table 14.4 (Continued) Group

Scientific name

Bacteria

Nocardia crassostreae

Bryozoan

Common name

Species impacted

Impact

Reference

Crassostrea gigas

Disease

Friedman et al. (1998)

Proteobacteria

Juvenile oyster disease

Crassostrea angulata, Crassostrea virginica

Disease

Renault et al. (2002); Renault and Novoa (2004)

Vibrio cholerae

Asiatic cholera

Shellfish

Disease

Dalsgaard (1998)

Scallop pearl nets

Foul gear

Dumont et al. (2009)

Bugula neritina Schizoporella errata

Branching bryozoan

Foul gear

GISD (2005)

Schizoporella unicornis

Single-horn bryozoan

Foul gear

GISD (2005)

Pecten maximus

Foul gear

Ross et al. (2004)

Cnidarian

Tubularia crocea

Crustacean

Carcinus maenus

European green crab

Crassostrea gigas, Mya arenaria, Mytilus edulis

Predation

Behrens Yamada et al. (2008a,b); Glude (1955); Dare et al. (1983); Floyd and Williams (2004); Murray et al. (2007); Behrens Yamada and Kosro (2010)

Mytilicola orientalis

Parasitic copepod

Mytilus edulis, other mussels, Ostrea gigas, Ostrea lurida, Paphia staminea

Parasitism

Odlaug (1946); Cole and Savage (1951); Gee et al. (1977)

Crassostrea gigas

Foul gear

Sala and Lucchetti (2008)

Pilumnus spinifer Echinoderm

Asterias amurensis

Flatbottom sea star

Fulvia tenuicostata, oysters

Predation

NIMPIS (2002); Ross et al. (2002)

Mollusc

Anadara demirii

Arcid clam

Crassostrea gigas

Foul gear, foul organisms

Morello et al. (2004); Sala and Lucchetti (2008)

Crepidula fornicata

Common Atlantic slippersnail

Crassostrea gigas, Mytilus edulis, and other oysters and mussels

Competition for food, interference competition, exclude other species, foul organisms

Blanchard (1997); Barton and Heard (2005); Decottignies et al. (2007a, 2007b); Thieltges (2005)

409

Table 14.4 (Continued) Group

Scientific name

Common name

Mytilopsis adamsi

Impact

Reference

False mussel

Exclude other species, foul organisms, foul gear

Wangkulangkul and Lheknim (2008)

Mytilopsis sallei

Black-stripped mussel

Decrease species richness

NIMPIS (2002)

Mytilus galloprovincialis

Mediterranean mussel

Crassostrea gigas, Mytilus californianus, Mytilus edulis

Decrease species richness, foul gear, overgrowth

Harger (1968); Geller (1999); Sala and Lucchetti (2008)

Ocinebrellus inornatus

Japanese oyster drill

Mussels, Ostrea lurida

Predation

Buhle and Ruesink (2009); Faasse and Ligthart (2009)

Rapana venosa

Asian rapa whelk

Anadara inaequivalvis, Crassostrea gigas, Mytilus edulis, Mytilus galloprovincialis, Tapes philippinarum

Predation

Kerckhof et al. (2006); Savini and OcchipintiAmbrogi (2006)

Urosalpinx cinerea

Eastern oyster drill

Mussels, Ostrea lurida, other oysters

Predation

Barton and Heard (2005); Buhle and Ruesink (2009); Faasse and Ligthart (2009)

Plant

Spartina alterniflora

Smooth cordgrass

Oysters

Habitat alteration, hybridization with native species

Chew (1998); Brusati and Grosholz (2008)

Protozoan

Bonamia exitiosa

Tiostrea chilensis

Disease

Ruesink et al. (2005)

Bonamia osteae

Crassostrea sikamea, Ostrea angasi, Ostrea edulis, Tiostrea chilensis

Disease

Ruesink et al. (2005)

Crassostrea virginica

Disease

Ruesink et al. (2005)

Ostrea angasi, Ostrea edulis, Tiostrea chilensis

Disease

Ruesink et al. (2005)

Saccostrea commercialis

Disease

Ruesink et al. (2005)

Haplosporidium nelsoni

MSX

Marteilia refringens

Marteilia sydneyi

410

QX

Species impacted

Table 14.4 (Continued) Group

Tunicate

Scientific name

Common name

Species impacted

Impact

Reference

Perkinsus marinus

Dermo

Crassostrea ariakensis, Crassostrea gigas, Crassostrea virginica

Disease

Ruesink et al. (2005)

Ascidiella aspersa

European sea squirt

Mussels, oysters, scallops

Competition for food, foul gear

Braithwaite et al. (2006); Currie et al. (1998)

Scallop ropes

Foul gear

Castilla et al. (2005)

Asterocarpa humilis Botrylloides violaceus

Orange sheath tunicate

Mytilus edulis

Foul organisms

Ramsay et al. (2008)

Botryllus schlosseri

Golden star tunicate

Crassostrea gigas, Mytilus edulis

Foul gear, foul organisms

Ramsay et al. (2008); Sala and Lucchetti (2008)

Ciona intestinalis

Vase tunicate

Mytilus edulis, other mussels, scallop pearl nets

Foul gear, foul organisms

Bullard et al. (2005); Castilla et al. (2005); Braithwaite et al. (2006); Blum et al. (2007); Ramsay et al. (2008); Dumont et al. (2009)

Didemnum sp.

Carpet tunicate

Mussels, mussel cages, oyster farms, scallops

Foul gear, overgrowth

Bullard et al. (2007)

Foul gear

Bullard et al. (2005)

Foul gear

Castilla et al. (2005)

Diplosoma listerianum Scallop ropes

Molgula ficus Molgula manhattensis

Sea grape

Styela clava

Asian tunicate

Styela plicata

Leathery tunicate

Foul gear Mytilus edulis

Foul organisms, overgrowth

Bullard et al. (2005); LeBlanc et al. (2007); Ramsay et al. (2008)

Decrease species richness, competition for space, foul organisms, slough off and remove other species

Sutherland (1978)

411

412

Shellfish Aquaculture and the Environment

Table 14.4 (Continued) Group

Scientific name

Virus

Common name

Species impacted

Impact

Reference

Iridolike viruses

Crassostrea angulata, Crassostrea gigas

Disease

Ruesink et al. (2005)

Oyster herpesvirus

Crassostrea gigas, Crassostrea sikamea, Ostrea edulis,

Disease

Ruesink et al. (2005)

Oyster virus velar disease

Crassostrea gigas

Disease

Ruesink et al. (2005)

Many of these species are likely to have additional ecological or economic impacts. PSP, paralytic shellfish poisoning.

introduced fouling species, especially ascidians, these costs are expected to escalate. The risk of human-mediated spread of shellfish diseases is also of concern. Increasingly, diseases and parasites are spread to new areas that can have large impacts on native shellfisheries as well as aquaculture (Renault et al. 2002; Renault and Novoa 2004; reviewed in Ruesink et al. 2005). We are also seeing increased spread of bloom-forming harmful algae (HABs). These algae can be spread with transported shellfish. Many HABs can have devastating impacts on shellfish populations (Matsuyama and Shumway 2009). In some cases, HABs cause shellfish mortality, while in others, they result in shellfish closures. Hégaret et al. (2008) examined the potential for bivalve aquaculture to spread harmful algal species and developed methods to minimize the likely transport of HAB species by holding the shellfish either in filtered seawater, free of algae, for 48 h or out of the water for the same time period.

Recommendations for minimizing spread and impacts of introductions It is clear that reducing the introduction and spread of nonnative species will be in the best interest of aquaculturists as well as natural

resource managers, and will simultaneously protect biodiversity and help conservation efforts. The large economic costs of unwanted species introductions are being faced by aquaculture facilities and shellfish farmers on a daily basis. Many nonnative species including those introduced by aquaculture itself (Table 14.3) have well-documented impacts on aquaculture (Table 14.4). The documented impacts of unwanted nonnative species are often on aquaculture species and include predation and competition, as well as disease and toxic algae. For others, the costs are seen through fouling of gear, all of which increases the costs to aquaculture, and reduces production (Watson et al. 2009; Adams et al. 2011). The large economic costs and environmental impacts associated with introductions of species outside of their native range has also put international pressure on scientists and managers to develop methodologies and policies that will minimize the spread of these species (Firestone and Corbett 2005), including reducing the likelihood of aquaculture species themselves becoming pests (Read and Fernandes 2003; ICES 2005) (Table 14.2). New laws are being implemented to reduce the transfer of species via shipping ballast (Firestone and Corbett 2005; Gregg et al. 2009; Vander Zanden et al. 2010) as well as

Marine invaders and bivalve aquaculture

deliberate introductions of species known to be harmful elsewhere or potentially harmful in the proposed site (Lacey Act 1900). Because it is difficult, if not impossible, to predict which species will become future problem invaders and because the potential costs of invasion are high, in most cases, risk-averse strategies are used that ban or minimize the transfer of any nonnative species. Natural resource managers should be concerned and careful about allowing introductions of nonnative species to new areas, even if those species are grown in other regions of the same country or state. Small differences in local environmental conditions can be the difference between a species that is locally contained and one that is spreading and impacting other species or systems of economic or environmental interest, as has been seen with the spread of Crassostrea gigas in France and the Netherlands (Diederich et al. 2005; Nehls et al. 2006; Kochmann et al. 2008; Schmidt et al. 2008; Thieltges et al. 2009; Markert et al. 2010). This is an important challenge for the introduction of all species with larvae that disperse or those that can readily be moved with gear or boats. In marine systems, although humans recognize political and geographic boundaries, other species do not. Species with long-distance dispersal larvae can travel hundreds of kilometers and readily cross political boundaries. This issue came into sharp focus for states surrounding the Chesapeake Bay with regard to the proposed wide-scale introduction of the nonnative oyster, Crassostrea ariakensis (Committee on Nonnative Oysters in the Chesapeake Bay, National Research Council 2004; Kingsley-Smith et al. 2009). Ultimately, the decision was made to not allow this largescale introduction. The potential gains were seen as too few and the potential costs due to the introduction of a nonnative too high. In addition, it was determined that the proposed economic gains of this introduction would not be realized.

413

Any policy (or lack of policy) in one state or country will affect its neighbors. Such awareness has led to new European Union standards of conformity among member states, recognizing the need to protect aquaculture from unwanted introductions, as well as public resources and private industry from the spread of species introduced through aquaculture activities. Council Regulation (EC) No. 708/2007 of 11 June 2007 created a framework governing aquacultural practices to protect aquatic environments from risks associated with the use of nonnative species of animals and plants (including microscopic species) in aquaculture. It controls the movement of any species that is locally absent for use in all types of aquaculture. It also requires all member states to take all appropriate measures to avoid risk to native species and communities resulting from the movement of nonnative species for aquaculture. The regulation requires that neighboring states are informed when any permits for nonnative aquaculture species are granted. Within the United States, similar types of issues are of concern. The regulation of aquaculture permitting is determined within states, and there is no uniform means by which states allow, or do not allow, establishment of new species for aquaculture. In addition, there is no requirement for cooperation between adjacent states that share waterways on decisions regarding allowing, or disallowing, the introduction of new species for aquaculture. To address these issues, large-scale national and international efforts have begun to develop best management practices (Chapter 3 in this book), standards, and certification for the development of sustainable aquaculture and to minimize the social and environmental issues associated with bivalve farming. One clear example is the leadership taken by the World Wildlife Fund (WWF) in developing the Bivalve Dialogue (WWF 2011), initiated in 2004, which focuses on oysters, mussels, clams, and scallops. Since 2004, there have been

414

Shellfish Aquaculture and the Environment

meetings in North America, Europe, New Zealand, and China, with more than 300 participants, with the goal of providing a framework for the development of criteria, indicators, and standards for responsible molluscan shellfish farming. The WWF dialogue has identified several environmental and social issues related to molluscan production, which include several related to the introduction of nonnative species. These include concerns regarding gene transfer to wild populations and the consequences of escapes and deliberate or inadvertent introduction of new nonnative species, including pests and pathogens. This dialogue has produced a list of principles, which includes those that will reduce the transfer and risk of nonnative species introductions. These include as follows: (1) obey the law and comply with all national and local regulations; (2) conserve natural habitat and local biodiversity; (3) protect the health and genetic integrity of wild populations; (4) manage disease and pests in an environmentally responsible manner; (5) use resources efficiently; (6) be a good neighbor and conscientious coastal citizen; (7) continually improve practices over time; and (8) develop and operate farms in a socially responsible manner. The WWF dialogues have led to the development of an international Aquaculture Stewardship Council (ASC) with the aim of developing independent third-party accreditation and certification of aquaculture operations of species for which standards have been developed. The ASC will provide a consumer label that can be used by processors and distributors that certify that shellfish products were grown in ways that meet a set of standards. Individual countries are now starting to develop aquaculture codes of practice that include minimizing the risks and impacts of the spread of introduced species and reducing the likelihood of aquaculture escapees. For example, Ireland is developing such a code that

includes examples of actions individual farmers can take, especially for dealing with fouling organisms (www.invasivespeciesireland.com/ cops/aquaculture). The Irish codes are based on those developed by the International Council for the Exploration of the Sea (ICES) Code of Practice on the Introductions and Transfers of Marine Organisms developed in 2005 (ICES 2005) and other international working groups developed to stop the spread of unwanted invaders. The goal of the ICES code is to reduce the ecological, environmental, economic, and genetic impacts associated with the transfer of species associated with aquaculture activities. It includes things such as cleaning all boats and equipment that comes in contact with the water, removing all living matter, detritus, and sediment from equipment, and the removal of all water trapped in equipment, including rinse water used to clean boats and gear, in order to prevent accidental transfers of unwanted species. They also focus on preventing the movement of fouled vessels or equipment from one area to another and the use of anitfouling technologies. Another important part of reducing the spread of unwanted invaders is reporting all suspicious organisms found, which can greatly help identify new invaders and allow rapid response efforts to eliminate new invasions. There are clear economic incentives for the aquaculture sector to continue to develop management and industry practices that reduce the impacts of nonnative species on products and equipment. These practices will also insure long-term sustainability of the industry. Such actions include selecting sites for aquaculture and methods that will minimize potential fouling, regular boiling (i.e., cleaning) of gear or cycling gear to reduce the abundance of nonnative species in an area, and disposing of unwanted animals and cleaning water in a way that prevents further spread of these nuisance species. Another effort to help reduce the spread and impacts of unwanted nonnatives

Marine invaders and bivalve aquaculture

is a project by the Collective Research on Aquaculture Biofouling (CRAB; www. crabproject.com). Because of the great cost of gear and animal fouling, especially the growing impact of nonnatives, this is a case where it is clearly in the best interest of farmers to take measures to stop the spread invaders. This project is a particularly good example of how to get education and outreach to farmers, and provide them with recommendations that will reduce their costs and reduce the spread of invaders. In general, gear cycling, as recommended by the USDA (www.mrc.state.va.us/ CRD/VA_706ajs.pdf) is one mechanism for reducing the impacts and spread biofouling while keeping costs minimal for farmers.

Future needs There are many parallels between aquaculture and agriculture. Both are important industries that provide essential food resources; however, agriculture relies on domesticated species. These species have been bred for characteristics that enhance production and have been transformed through breeding, often for thousands of years, making them very different than their wild ancestors. Their success is generally dependent on the humans that grow them. Aquaculture, on the other hand, relies on the captive culture of essentially wild species, and like many natural species, when transplanted to new environments these animals can reproduce and spread, often with unintended and harmful results. Continued efforts to develop regulations and oversight will simultaneously allow the benefits of aquaculture to be realized and protect both natural marine systems and shellfish aquaculture from the unwanted impacts of nonnative species introductions. In addition, we need to focus attention to practices that can result in shellfish aquaculture species themselves from becoming

415

unwanted invaders. The development of best management practices that prevent the release of unwanted or unneeded individuals to the environment are necessary. The release of larvae or gametes and leaving leftover individuals in the environment after desirable individuals are harvested can lead to the establishment of feral populations that can spread and cause unwanted environmental damage. Similarly, regulations that prevent deliberate attempts to establish populations of nonnative species for harvest or allow nonnative species in aquaculture to reproduce where they can spread are greatly needed. We also need mechanisms that will minimize the risks of the spread of nonnative species and the escape of aquaculture species. This is especially important when there is consideration of introducing new species for aquaculture outside of their native range. One factor that will influence the spread and impacts of species introductions is climatic change. Species that were once contained in a region, with temperature-driven physiological limits or limits on reproduction, are able to reproduce now that waters are warmer. As a consequence of climatic change, more species may spread to areas thought to be safe from these species (Diederich et al. 2005; Klinger et al. 2006; Kochmann et al. 2008; Thieltges et al. 2009). Planning and consideration of new aquaculture species or the transport of species also needs to include a consideration of projected changes in climate (Chapter 17 in this book) and the potential for the spread of nonnatives when considering the risks of species introductions.

Acknowledgments We would like to thank Robinson Herrera and Geoff Bolen for help with assembling the data for the tables in this chapter.

416

Shellfish Aquaculture and the Environment

Literature cited Adams, C.M., Shumway, S.E., Whitlach, R.B., and Getchis, T. 2011. Biofouling in marine molluscan shellfish aquaculture: a survey assessing the business and economic implications of mitigation. Journal of the World Aquaculture Society 42:242–252. Anon. 2005. Invasive ascidian biofouling in aquaculture: an increasing problem and can it be controlled? Journal of Shellfish Research 24:682–683. Bachelet, G., Blanchet, H., Cottet, M., Dang, C., de Montaudouin, X., de Moura Queirós, A., Gouilieux, B., and Lavesque, N. 2009. A roundthe-world tour almost completed: first records of the invasive mussel Musculista senhousia in the north-east Atlantic (southern Bay of Biscay). Marine Biodiversity Records 2:e119. Bailey-Brock, J.H. 1990. Polydora nuchalis (Polychaeta: Spionidae), a new Hawaiian record from aquaculture ponds. Pacific Science 44: 81–87. Bailey-Brock, J.H. 2000. A new record of the polychaete Boccardia proboscidea (Family Spionidae), imported to Hawai’i with oysters. Pacific Science 54:27–30. Baker, P. 1995. Review of ecology and fishery of the Olympia oyster, Ostrea lurida with annotated bibliography. Journal of Shellfish Research 14:501–518. Barton, E., and Heard, J. 2005. The Marine Biological Association of the United Kingdom and The Marine Life Information Network for Britain and Ireland. www.marlin.ac.uk/PDF/ MLTN_alien_non_natives.pdf Behrens Yamada, S., Dumbauld, B.R., Kalin, A., Hunt, C.E., Figlar-Barnes, R., and Randall, A. 2005. Growth and persistence of a recent invader Carcinus maenas in estuaries of the northeastern Pacific. Biological Invasions 7:309–321. Behrens Yamada, S., Hardege, J.D., and Bublitz, R. 2008a. Sex pheromones: new tools for controlling European green crabs? Journal of Shellfish Research 27:463. Behrens Yamada, S., Randall, A., and Gillespie, G.E. 2008b. European green crab status in 2006. Journal of Shellfish Research 27:464.

Behrens Yamada, S., and Kosro, P.M. 2010. Linking ocean conditions to year class strength of the invasive European green crab, Carcinus maenas. Biological Invasions 12:1791–1804. Beninger, P.G., Decottignies, P., Guiheneuf, F., Barille, L., and Rince, Y. 2007. Comparison of particle processing by two introduced suspension feeders: selection in Crepidula fornicata and Crassostrea gigas. Marine Ecology Progress Series 334:165–177. Bishop, M.J., Carnegie, R.B., Stokes, N.A., Peterson, C.H., and Burreson, E.M. 2006. Complications of a non-native oyster introduction: Facilitation of a local parasite. Marine Ecology Progress Series 325:145–152. Blanchard, M. 1997. Spread of the slipper limpet Crepidula fornicata (L. 1758) in Europe. Current state and consequences. Scientia Marina 61(Suppl. 2):109–118. Blanchard, M. 2009. Recent expansion of the slipper limpet population (Crepidula fornicata) in the Bay of Mont-Saint-Michel (Western Channel, France). Aquatic Living Resources 22:11–19. Blanchard, M., Pechenik, J.A., Giudicelli, E., Connan, J.P., and Robert, R. 2008. Competition for food in the larvae of two marine molluscs, Crepidula fornicata and Crassostrea gigas. Aquatic Living Resources 21:197–205. Blum, J.C., Change, A.L., Liljesthröm, M., Schenk, M.E., Steinberg, M.K., and Ruiz, G.M. 2007. The non-native solitary ascidian Ciona intestinalis (L.) depresses species richness. Journal of Experimental Marine Biology and Ecology 342:5–14. Braithwaite, R.A., Cadavid Carrascosa, M.C., and McEvoy, L.A. 2006. Biofouling of salmon cage netting and the efficacy of a typical copper-based antifoulant. Aquaculture 262:219–226. Branch, G.M., and Steffani, C.N. 2004. Can we predict the effects of alien species? A case-history of the invasion of South Africa by Mytilus galloprovincialis (Lamarck). Journal of Experimental Marine Biology and Ecology 300:189–215. Branch, G.M., Odendaal, F., and Robinson, T.B. 2010. Competition and facilitation between the alien mussel Mytilus galloprovincialis and indigenous species: moderation by wave action. Journal of Experimental Marine Biology and Ecology 383:65–78.

Marine invaders and bivalve aquaculture

Brusati, E.D., and Grosholz, E.D. 2008. Does invasion of hybrid cordgrass change estuarine food webs? Biological Invasions 11:917–926. Buhle, E.R., and Ruesink, J.L. 2009. Impacts of invasive oyster drils on Olympia oyster (Ostrea lurida Carpenter 1864) recovery in Willapa Bay, Washington, United States. Journal of Shellfish Research 28:87–96. Bullard, S., Whitlach, R.B., Shumway, S., and Osman, R. 2005. Scientists crying “foul”: sea squirts invade Long Island Sound. Wrack Lines, Fall/Winter. Connecticut Sea Grant. Bullard, S.G., Lambert, G., Carman, M.R., Byrnes, J., Whitlatch, R.B., Ruiz, G., Miller, R.J., Harris, L., Valentine, P.C., Collie, J.S., Pederson, J., McNaught, D.C., Cohen, A.N., Asch, R.G., Dijkstra, J., and Heinonen, K. 2007. The colonial ascidian Didemnum sp. A: current distribution, basic biology and potential threat to marine communities of the northeast and west coasts of North America. Journal of Experimental Marine Biology and Ecology 342:Special Issue:99– 108. Bulleri, F., Airoldi, L., Branca, G.M., and Abbiati, M. 2006. Positive effects of the introduced green alga, Codium fragile ssp. tomentosoides, on recruitment and survival of mussels. Marine Biology 148:1213–1220. Buttger, H., Asmus, H., Asmus, R., Buschbaum, C., Dittmann, S., and Nehls, G. 2008. Community dynamics of intertidal soft-bottom mussel beds over two decades. Helgoland Marine Research 62:23–36. Caldow, R.W.G., Stillman, R.A., le Vdit Durell, S.E.A., West, A.D., McGrorty, S., Goss-Custard, J.D., Wood, P.J., and Humphreys, J. 2007. Benefits to shorebirds from invasion of a nonnative shellfish. Proceedings of the Royal Society B, Biological Sciences 274:1449–1455. Carlton, J.T. 1989. Man’s role in changing the face of the ocean: biological invasions and implications for conservation of near- shore environments. Conservation Biology 3:265– 273. Carlton, J.T. 1996. Marine bioinvasions: the alteration of marine ecosystems by nonindigenous species. Oceanography 9:36–43. Carlton, J.T. 1999. Molluscan invasions in marine and estuarine communities. Malacologia 41:439–454.

417

Castilla, J.C., Uribe, M., Bahamonde, N., Clarke, M., Desqueyroux-Faundez, R., Kong, I., Moyano, H., Rozbaczylo, N., Santelices, B., Valdovinos, C., and Zavala, P. 2005. Down under the southeastern Pacific: marine nonindigenous species in Chile. Biological Invasions 7:213–232. Chew, K.K. 1990. Global bivalve shellfish introductions. World Aquaculture 21:9–22. Chew, K.K. 1998. Growing impact of smooth cordgrass, Spartina alterniflora, in Washington State. Aquaculture Magazine 24:93–97. Cigarría, J., and Elston, R. 1997. Independent introduction of Bonamia ostreae, a parasite of Ostrea edulis, to Spain. Diseases of Aquatic Organisms 29:157–158. Civille, J.C., Sayce, K., Smith, S.D., and Strong, D.R. 2005. Reconstructing a century of Spartina alterniflora invasion with historical records and contemporary remote sensing. Ecoscience 12:330–338. Cloern, J.E. 1982. Does the benthos control phytoplankton biomass in South San Francisco bay? Marine Ecology Progress Series 9:191–202. Cognie, B., Haure, J., and Barillé, L. 2006. Spatial distrubtion in a temperate coastal ecosystem of the wild stock of the farmed oyster Crassostrea gigas (Thunberg). Aquaculture 259:249–259. Cohen, A.N., and Carlton, J.T. 1998. Accelerating invasion rate in a highly invaded estuary. Science 279:555–558. Cohen, A.N., and Zabin, C.J. 2009. Oyster shells as vectors for exotic organisms. Journal of Shellfish Research 28:163–167. Colautti, R.I., and Richardson, D.M. 2009. Subjectivity and flexibility in invasion terminology: too much of a good thing? Biological Invasions 11:1225–1229. Cole, H.A., and Savage, R.E. 1951. The effect of the parasitic copepod, Mytilicola intestinalis (Steuer) upon the condition of mussels. Parasitology 41:151–161. Coleman, R.A., and Hockey, P.A.R. 2008. Effects of an alien invertebrate species and wave action on prey selection by African black oystercatchers (Haematopus moquini). Austral Ecology 33:232–240. Coles, S.L., DeFelice, R.C., Eldredge, L.G., and Carlton, J.T. 1999. Historical and recent intro-

418

Shellfish Aquaculture and the Environment

ductions of nonindigenous marine species into Pearl Harbor, Oahu, Hawaiian Islands. Marine Biology. 135:147–158. Committee on Nonnative Oysters in the Chesapeake Bay, National Research Council. 2004. Nonnative Oysters in the Chesapeake Bay. The National Academies Press, Washington, D.C. Costa-Pierce, B.A. 2010. Sustainable ecological aquaculture systems: the need for a new social contract for aquaculture development. Marine Technology Society Journal 44:88–112. Critchley, A.T., and Dijkema, R. 1984. On the presence of the introduced brown alga Sargassum muticum, attached to commercially imported Ostrea edulis in the S. W. Netherlands. Botanica Marina 27:211–216. Cuddington, K., Byers, J.E., Wilson, W.G., and Hastings, A. 2007. Ecosystem Engineers. Plants to Protists. Elsevier Press, Amsterdam. Curiel, D.P., Guidetti, B., Scattolin, M., and Marzocchi, M. 2001. The introduced algae Undaria pinnatifida (Laminariales, Alariaceae) in the lagoon of Venice. Hydrobiologia 477:209–219. Currie, D.R., McArthur, M.A., and Cohen, B.F. 1998. Exotic Marine Pests in the Port of Geelong, Victoria, Marine and Freshwater Resources Institute. Report No. 8:1–57. da Silva, P.M., Fuentes, J., and Villalba, A. 2005. Growth, mortality and disease suceptibility of oyster Ostrea edulis families obtained from brood stocks of different geographical origins, through on-growing in the Ría de Arousa (Galicia, NW Spain). Marine Biology 147: 965–977. DAISIE European Invasive Alien Species Gateway. 2008. Crassostrea gigas. www.europe-aliens. org/ (accessed on August 1, 2010). Dalsgaard, A. 1998. The occurrence of human pathogenic Vibri spp. and Salmonella in aquaculture. International Journal of Food Science and Technology 33:127–138. Dare, P.J., Davies, G., and Edwards, D.B. 1983. Predation on juvenile Pacific oysters (Crassostrea gigas Thunberg) and mussels (Mytilus edulis L.) by shore crabs (Carcinus maenas L.). Fisheries Ressearch Technical Report No. 73. MAFF Direct. Fish. Res., Lowestoft. Decottignies, P., Beninger, P.G., Rincé, Y., and Riera, P. 2007a. Trophic interactions between

two introduced suspension-feeders, Crepidula fornicata and Crassostrea gigas are influenced by seasonal effects and qualitative selection capacity. Journal of Experimental Marine Biology and Ecology 342:231–241. Decottignies, P., Beninger, P.G., Rincé, Y., Robbins, R.J., and Riera, P. 2007b. Exploitation of natural food sources by two sympatric, invasive suspension-feeders: Crassostrea gigas and Crepidula fornicata. Marine Ecology Progress Series 334:179–192. Delgado, M., Estrada, M., Camp, J., Fernández, J.V., Santmart, M., and Lletí, C. 1990. Development of a toxic Alexandrium minutum Halim (Dinophyceae) bloom in the harbour of Sant Carles de la Ràpita (Ebro Delta, northwestern Mediterranean). Scientia Marina 54:1–7. Dias, P.J., Batista, F.M., Shanks, A.M., Beaumont, A.R., Davies, I.M., and Snow, M. 2009. Gametogenic asynchrony of mussels Mytilus in a mixed-species area: implications for management. Aquaculture 295:175–182. Diederich, S. 2006. High survival and growth rates of introduced Pacific oysters may cause restrictions on habitat use by native mussels in the Wadden Sea. Journal of Experimental Marine Biology and Ecology 328:211–227. Diederich, S., Nehls, G., van Beusekom, J.E.E., and Reise, K. 2005. Introduced Pacific oysters (Crassostrea gigas) in the northern Wadden Sea: invasion accelerated by warm summers? Helgoland Marine Research 59:97–106. Dubois, S., Commito, J.A., Olivier, F., and Retière, C. 2006. Effects of epibionts on Sabellaria alveolata (L.) biogenic reefs and their associated fauna in the Bay of Mont SaintMichel. Estuarine, Coastal and Shelf Science 68:635–646. Dumont, C.P., Urriago, J.D., Abarca, A., Gaymer, C.F., and Thiel, M. 2009. The native rock shrimp Rhynchocinetes typus as a biological control of fouling in suspended scallop cultures. Aquaculture 292:74–79. Elton, C.S. 1958. The Ecology of Invasions by Animals and Plants. Methuen and Co. Ltd, London. Eno, N.C. 1998. The introduction to British waters of nonnative marine molluscs and the implications to nature conservation interests. Journal

Marine invaders and bivalve aquaculture

of Conchology Special Publication (No. 2): 287–294. Eno, N.C., Clark, R.A., and Sanderson, W.G. 1997. Non-Native Marine Species in British Waters: A Review and Directory. Joint Nature Conservation Committee, Peterborough. Faasse, M., and Ligthart, M. 2009. American (Urosalpinx cinerea) and Japanese oyster drill (Ocinebrellus inornatus) (Gastropoda: Muricidae) flourish near shellfish culture plots in the Netherlands. Aquatic Invasions 4:321–326. Ferreira, J.G., Sequeira, A., Hawkins, A.J.S., Newton, A., Nickell, T.D., Pastres, R., Forte, J., Bodoy, A., and Bricker, S.B. 2009. Analysis of coastal and offshore aquaculture: application of the FARM model to multiple systems and shellfish species. Aquaculture 289:32–41. Firestone, J., and Corbett, J.J. 2005. Coastal and port environments: international legal and policy responses to reduce ballast water introductions of potentially invasive species. Ocean Development and International Law 36:291– 316. Flassch, J.P., and Leborgne, Y. 1990. Introduction in Europe, from 1972 to 1980, of the Japanese Manila clam (Tapes philippinarum) and the effects on aquaculture production and natural settlement. ICES Marine Science Symposia 194:92–96. Fletcher, R.L., and Manfredi, C. 1995. The occurrence of Undaria pinnatifida (Phaeophyceae, Laminarales) on the south coast of England. Botanica Marina 38:355–358. Floyd, T., and Williams, J. 2004. Impact of green crab (Carcinus maenas L.) predation on a population of softshell clams (Mya arenaria L.) in the Southern Gulf of St. Lawrence. Journal of Shellfish Research 23:457–462. Food and Agriculture Organization (FAO). 1996. List of Animal Species Used in Aquaculture. FAO Fisheries Circular No. 914, Rome, Italy. Food and Agriculture Organization (FAO). 1997. Aquaculture development. FAO Technical Guidelines for Responsible Fisheries No. 5. FAO, Rome, Italy. 40 pp. Food and Agriculture Organization (FAO). 2005. Database on Introductions of Aquatic Species (DIAS). In: FAO Fisheries and Aquaculture Department [online]. www.fao.org/fishery/ (accessed on March 30, 2010).

419

Food and Agriculture Organization (FAO). 2007. Cultured Aquatic Species Information Programme. In: FAO Fisheries and Aquaculture Department [online]. www.fao.org/fishery/ (accessed on March 30, 2010). Ford, S.E. 1996. Range extension by the oyster parasite Perkinsus marinus into the northeastern United States: response to climate change? Journal of Shellfish Research 15:45–56. Friedman, C.S., and Perkins, F.O. 1994. Range extension of Bonamia ostreae to Maine, U.S.A. Journal of Invertebrate Pathology 64:179– 181. Friedman, C.S., Beaman, B.L., Chun, J., Goodfellow, M., Gee, A., and Hedrick, R.P. 1998. Nocardia crassostrea sp. nov., the causal agent of nocardiosis in Pacific oysters. International Journal of Systematic Bacteriology 48:237–246. Gee, J.M., Maddock, L., and Davey, J.T. 1977. The relationship between infestation by Mytilicola intestinalis, Steuer (Copepoda, Cyclopoidea) and the condition index of Mytilus edulis in southwest England. ICES Journal of Marine Sciences 37:300–308. Geller, J. 1999. Decline of a native mussel masked by sibling species invasion. Conservation Biology 13:661–664. Global Invasive Species Database (GISD). 2005. www.issg.org/ (accessed on January 28, 2010). Glude, J.B. 1955. The effects of temperature and predators on the abundance of the soft-shell clam, Mya arenaria, in New England. Transactions of the American Fisheries Society 84:13–26. Gosling, E. 2003. Bivalve Molluscs: Biology, Ecology and Culture. Fishing New Books, Oxford. Gregg, M., Rigby, G., and Hallegraeff, G.M. 2009. Review of two decades of progress in the development of management options for reducing or eradicating phytoplankton, zooplankton and bacteria in ship’s ballast water. Aquatic Invasions 4:521–565. Gunther, R.T. 1897. The oyster culture of the ancient Romans. Journal of the Marine Biological Association of the United Kingdom 4:360– 365. Gurnell, J., Wauters, L.A., Lurz, P.W.W., and Tosi, G. 2004. Alien species and interspecific competition: effects of introduced eastern grey squirrels

420

Shellfish Aquaculture and the Environment

on red squirrel population dynamics. Journal of Animal Ecology 73:26–35. Hallegraeff, G.M., Steffensen, D.A., and Wetherbee, R. 1988. Three estuarine Australian dinoflagellates that can produce paralytic shellfish toxins. Journal of Plankton Research 10:553–541. Harger, J.R.E. 1968. The role of behavioral traits in influencing the distribution of two species of sea mussel, Mytilus edulis and Mytilus californianus. The Veliger 11:45–49. Harries, D.B., Cook, E., Donnan, D.W., Mar, J.M., Harrow, S., and Wilson, J.R. 2007. The establishment of the invasive alga Sargassum muticum on the east coast of Scotland: rapid northwards spread and identification of potential new areas for colonisation. Aquatic Invasions 2:367– 377. Hégaret, H., Shumway, S.E., Wikfors, G.H., Pate, S., and Burkholder, J.M. 2008. Potential transport of harmful algae via relocation of bivalve molluscs. Marine Ecology Progress Series 361:169–179. Hégaret, H., Wikfors, G.H., and Shumway, S.E. 2009. Biotoxin contamination and shellfish safety. In: Shumway, S.E., and Rodrick, G.E. (eds.), Shellfish Quality and Safety. Woodhead Publishing, Cambridge, pp. 43–80. International Council for the Exploration of the Sea (ICES). 2005. ICES Code of Practice on the Introductions and Transfers of Marine Organisms 2005. 30 pp. Jones, C.G., Lawton, J.H., and Shachak, M. 1994. Organisms as ecosystem engineers. Oikos 69:373–386. Karatayev, A.Y., Burlakova, L.E., Padilla, D.K., Mastitsky, S.E., and Olenin, S. 2009. Invaders are not a random selection of species. Biological Invasions 11:2009–2019. Kelly, J.R., and Volpe, J.P. 2007. Native eelgrass (Zostera marina L.) survival and growth adjacent to non-native oysters (Crassostrea gigas Thunberg) in the Strait of Georgia, British Columbia. Botanica Marina 50:143–150. Kelly, J.R., Proctor, H., and Volpe, J.P. 2008. Intertidal community structure differs significantly between substrates dominated by native eelgrass (Zostera marina L.) and adjacent to the introduced oyster Crassostrea gigas (Thunberg) in British Columbia, Canada. Hydrobiologia 596:57–66.

Kerckhof, F., Vink, R.J., Nieweg, D.C., and Post, N.J. 2006. The veined whelk Rapana venosa has reached the North Sea. Aquatic Invasions 1:35–37. Kingsley-Smith, P.R., Harwell, H.D., Kellogg, M.L., Allen, S.M., Allen, S.K., Meritt, D.W., Paynter, K.T., and Luckenbach, M.W. 2009. Survival and growth of triploid Crassostrea virginica (Gmelin, 1791) and C. ariakensis (Fujita, 1913) in bottom environments of Chesapeake Bay: implications for an introduction. Journal of Shellfish Research 28:169–184. Klinger, T., Padilla, D.K., and Britton-Simmons, K. 2006. Two invaders achieve higher densities in marine reserves. Aquatic Conservation—Marine and Freshwater Ecosystems 16:301–311. Kochmann, J., Buschbaum, C., Volkenborn, N., and Reise, K. 2008. Shift from native mussels to alien oysters: differential effects of ecosystem engineers. Journal of Experimental Marine Biology and Ecology 364:1–10. Krassoi, F.R., Brown, K.R., Bishop, M.J., Kelaher, B.P., and Summerhayes, S. 2008. Conditionspecific competition allows coexistence of competitively superior exotic oysters with native oysters. Journal of Animal Ecology 77:5–15. Kurlansky, M. 2006. The Big Oyster: History on the Half Shell. Ballantine Books, New York. Lacey Act. 1900. An Act to Enlarge the Powers of the Department of Agriculture, Prohibit the Transportation by Interstate Commerce of Game Killed in Violation of Local Laws, and for Other Purposes. United States Statutes at Large 31: chapter 553, 187–89. US Government Printing Office, Washington, DC. Laing, I., and Gollasch, S. 2002. Coscinodiscus wailesii: a nuisance diatom in European waters. In: Leppäkoski, E., Gollasch, S., Olenin, S. (eds.), Invasive Aquatic Species of Europe: Distribution, Impacts, and Management. Kluwer Academic Publishers, Boston, MA, pp. 53–55. Laiño, C.C. 1991. Gymnodinium catenatum toxins from mussels (Mytilus galloprovincialis). Environmental Technology 12:33–40. Lambert, G. 2007. Invasive sea squirts: a growing global problem. Journal of Experimental Marine Biology and Ecology 342 Special Issue:3–4. Langeland, G., Hasselgård, T., Tangen, K., Skulberg, O.M., and Hjelle, A. 1984. An outbreak of para-

Marine invaders and bivalve aquaculture

lytic shellfish poisoning in western Norway. Sarsia 69:185–193. LeBlanc, N., Davidson, J., Tremblay, R., McNiven, M., and Landry, T. 2007. The effect of antifouling treatments for the clubbed tunicate on the blue mussel, Mytilus edulis. Aquaculture 264:205–213. Le Duff, L., Grall, M.J., and Quiniou, L. 2009. First record of the gastropod Fusinus rostratus (Mollusca: Fasciolaridae) on the northern coast of Brittany (Western Channel, France). Marine Biodiversity Records 2:e63, 1–3. Levasseur, M., Couture, J., Weise, A.M., Michaud, S., Elbrächter, M., Sauvé, G., and Bonneau, E. 2003. Pelagic and epiphytic summer distributions of Prorocentrum lima and P. mexicanum at two mussel farms in the Gulf of St. Lawrence, Canada. Aquatic Microbial Ecology 30:283– 293. Lilly, E.L., Kulis, D.M., Gentien, P., and Anderson, D.M. 2002. Paralytic shellfish poisoning toxins in France linked to a human-introduced strain of Alexandrium catenella from the western Pacific: evidence from DNA and toxin analysis. Journal of Plankton Research 24:443–452. Luckenbach, M.W., Mann, R., and Wesson, J. 1999. Oyster Reef Habitat Restoration: A Synopsis and Synthesis of Approaches. Virginia Instituted of Marine Science Press, Gloucester Point, VA. Mann, R. 1979. Exotic Species in Mariculture. Proceedings of Symposium on Exotic Species in Mariculture: Case Histories of the Japanese Oyster, Crassostrea Gigas (Thunberg), with Implications for Other Fisheries. Woods Hole Oceanographic Institution, Woods Hole, September 18–20, 1978. MIT Press, Cambridge, MA. Markert, A., Wehrmann, A., and Kroncke, I. 2010. Recently established Crassostrea-reefs versus native Mytilus-beds: differences in ecosystem engineering affects the macrofaunal communities (Wadden Sea of Lower Saxony, southern German Bight). Biological Invasions 12:15–32. Masilamoni, J.G., Azariah, J., Nandakumar, K., Jesudoss, K.S., Satpathy, K.K., and Nair, K.V.K. 2001. Excretory products of green mussel Perna viridis L. and their implications on power plant operation. Turkish Journal of Zoology 25: 117–125.

421

Masilamoni, G., Jesudoss, K.S., Nandakumar, K., Satapathy, K.K., Azariah, J., and Nair, K.V.K. 2002. Lethal and sub-lethal effects of chlorination on green mussel Perna viridis in the context of biofouling control in a power plant cooling water system. Marine Environment Research 53:65–76. Matsuyama, Y., and Shumway, S. 2009. Impacts of harmful algal blooms on shellfisheries aquaculture. In: Burnell, G., and Allan, G. (eds.), New Technologies in Aquaculture: Improving Production Efficiency, Quality and Environmental Management. Woodhead Publishing Limited, Oxford, pp. 580–609. McArdle, J.F., McKiernan, F., Foley, H., and Jones, D.H. 1991. The current status of Bonamia disease in Ireland. Aquaculture 93:273–278. McKindsey, D.W., Landry, T., O’Beirn, F.X., and Davies, I.M. 2007. Bivalve aquaculture and exotic species: a review of ecological considerations and management issues. Journal of Shellfish Research 26:281–294. Minchin, D., McGrath, D., and Duggan, C.B. 1995. The slipper limpet, Crepidula fornicata (L.) in Irish Waters, with a review of its occurrence in the north-eastern Atlantic. Journal of Conchology 35:247–254. Mineur, F., Belsher, T., Johnson, M.P., Maggs, C.A., and Verlaque, M. 2007. Experimental assessment of oyster transfers as a vector for macroalgal introductions. Biological Conservation 137: 237–247. Moore, C.G., and Harries, D.B. 2009. Appearance of Heterosiphonia japonica (Ceramiales: Rhodophyceae) on the west coast of Scoatland, with notes on Sargassum muticum (Fucales: Heterokontophyta). Marine Biodiversity Records 2(e131):1–5. Morello, E.B., Solustri, C., and Froglia, C. 2004. The alien bivalve Anadara demiri (Arcidae): a new invader of the Adriatic Sea. Italy. Journal of the Marin Biological Association of the United Kingdom 84:1057–1064. Mugg Pietros, J., and Rice, M.A. 2003. The impacts of aquacultured oysters, Crassostrea virginica (Gmelin, 1791) on water column nitrogen and sedimentation: results of a mesocosm study. Aquaculture 220:407–422. Muir, J. 2005. Managing to harvest? Perspectives on the potential of aquaculture. Philosophical

422

Shellfish Aquaculture and the Environment

Transactions of the Royal Society B 360: 191–218. Murray, L.G., Seed, R., and Jones, T. 2007. Predicting the impacts of Carcinus maenas predation on cultivated Mytilus edulis beds. Journal of Shellfish Research 26:1089– 1098. National Introduced Marine Pest Information System (NIMPIS). 2002. Eds: Hewitt C.L., Martin R.B., Sliwa C.,McEnnulty F.R., Murphy N.E., Jones T. & Cooper S. crimp.marine.csiro. au/nimpis (accessed on January 25, 2010). Nehls, G., Diederich, S., Thieltges, D.W., and Strasser, M. 2006. Wadden Sea mussel beds invaded by oysters and slipper limpets— competition or climate control? Helgoland Marine. Research 60:135–143. Nell, J.C. 2007. Dieseases of Sydney rock oysters. New South Wales (NSW) Department of Primary Industries Primefact 589:1–4. O’Connor, W.A., and Newman, L.J. 2001. Halotolerance of the oyster predator, Imogine mcgrathi, a stylochid flatworm from Port Stephens, New South Wales, Australia. Hydrobiologia 459:157–163. Odlaug, T.O. 1946. The effect of the copepod, Mytilicola orientalis, upon the Olympia oyster, Ostrea lurida. Transactions of the American Microscopical Society 65:311. Ogburn, D.M., White, I., and Mcphee, D.P. 2007. The disappearance of oyster reefs from Eastern Australian estuaries—impact of colonial settlement or mudworm invasion? Coastal Management 35:271–287. Padilla, D.K. 2010. Impacts and spread of a nonnative ecosystem engineer, the Pacific oyster Crassostrea gigas. Integrative and Comparative Biology 50:213–225. Pimentel, D., Zuniga, R., and Morrison, D. 2005. Update on the environmental and economic costs associated with alien-invasive species in the United States. Ecological Economics 52:273– 288. Pranovi, F., Franceschini, G., Casale, M., Zucchetta, M., Torricelli, P., and Giovanardi, O. 2006. An ecological imbalance induced by a non-native species: the Manila clam in the Venice Lagoon. Biological Invasions 8:595–609. Quayle, D.B. 1988. Pacific oyster culture in British Columbia. Canadian Bulletin of Fisheries and

Aquattic Science No. 218. Canada Fisheries Research Board of Canada, Ottawa. Rajagopal, S., Venugopalan, V.P., Van der Velde, G., and Jenner, H.A. 2003. Tolerance of five species of tropical marine mussels to continuous chlorination. Marine Environmental Research 55: 277–291. Rajagopal, S., Venugopalan, V.P., Van der Velde, G., and Jenner, H.A. 2006. Greening of the coasts: a review of the Perna viridis success story. Aquatic Ecology 40:273–297. Ramsay, A., Davidson, J., Landry, T., and Arsenault, G. 2008. Process of invasiveness among exotic tunicates in Prince Edward Island, Canada. Biological Invasions 10:1311–1316. Rawson, P.D., Agrawal, V., and Hilbish, T.J. 1999. Evidence of a steep genetic break among blue mussel (Mytilus spp.) populations associated with Cape Mendocino, CA, USA. Marine Biology 134:201–211. Read, P., and Fernandes, T. 2003. Management of environmental impacts of marine aquaculture in Europe. Aquaculture 226:139–163. Reise, K., Olenin, S., and Thieltges, D.W. 2005. Are aliens threatening European aquatic coastal ecosystems? Helgoland Marine Research 60:77–83. Renault, T., and Novoa, B. 2004. Viruses infecting bivalve molluscs. Aquatic Living Resources 17:397–409. Renault, T., Chollet, B., Cochennec, N., and Gerard, A. 2002. Shell disease in eastern oysters, Crassostrea virginica, reared in France. Journal of Invertebrate Pathology 79:1–6. Robinson, T.B., Griffiths, C.L., McQuaid, C.D., and Ruis, M. 2005. Marine alien species of South Africa—status and impacts. African Journal of Marine Science 27:297–306. Robinson, T.B., Griffiths, C.L., Branch, G.M., and Govender, A. 2007. The invasion and subsequent die-off of Mytilus galloprovincialis in Langebaan Lagoon, South Africa: effects on natural communities. Marine Biolgoy 152:225– 232. Rodriguez, L.F., and Ibarra-Obando, S.E. 2008. Cover and colonization of commercial oyster (Crassostrea gigas) shells by fouling organisms in San Quintin Bay, Mexico. Journal of Shellfish Research 27:337–343. Ross, D.J., Johnson, C.R., and Hewitt, C.L. 2002. Impact of introduced seastars Asterias amurensis

Marine invaders and bivalve aquaculture

on survivorship of juvenile commercial bivavles Fulvia tenuicostata. Marine Ecology Progress Series 241:99–112. Ross, K.A., Thorpe, J.P., and Brand, A.R. 2004. Biological control of fouling in suspended scallop cultivation. Aquaculture 229:99–116. Ruesink, J.L., Lenihan, H.S., Trimble, A.C., Heiman, K.W., Micheli, F., Byers, J.E., and Kay, M.D. 2005. Introduction of non-native oysters: ecosystem effects and restoration implications. Annual Review of Ecology Evolution and Systematics 36:643–689. Ruiz, G., Carlton, J., Grosholz, E., and Hines, A.H. 1997. Global invasions of marine and estuarine habitats by non-indigenous species: mechanisms, extent and consequences. American Zoologist 37:621–632. Sala, A., and Lucchetti, A. 2008. Low-cost tool to reduce biofouling in oyster longline culture. Aquacultural Engineering 39:53–58. Savini, D., and Occhipinti-Ambrogi, A. 2006. Consumption rates and prey prefences of the invasive gastropod Rapana venosa in the Northern Adriatic Sea. Helgoland Marine Research 60:153–159. Schmidt, A., Wehrmann, A., and Dittmann, S. 2008. Population dynamics of the invasive Pacific oyster Crassostrea gigas during the early stages of an outbreak in the Wadden Sea (Germany). Helgoland Marine. Research 62:367–376. Shatkin, G., Shumway, S.E., and Hawes, R. 1997. Considerations regarding the possible introduction of the Pacific oyster (Crassostrea gigas) to the Gulf of Maine: a review of global experience. Journal of Shellfish Research 16:463–477. Shields, J.L., Heath, J.W., and Heath, D.D. 2010. Marine landscape shapes hybrid zone in a broadcast spawning bivalve: introgression and genetic structure in Canadian west coast Mytilus. Marine Ecology Progress Series 399:211–223. Sjøtun, K., Husa, V., and Peña, V. 2008. Present distribution and possible vectors of introductions of the alga Heterosiphonia japonica (Ceramiales, Rhodophyta) in Europe. Aquatic Invasions 3:377–394. Sousa, R., Gutierrez, J.L., and Aldridge, D.C. 2009. Non-indigenous invasive bivalves as ecosystem engineers. Biological Invasions 11:2367– 2385.

423

Steffani, C.N., and Branch, G.M. 2005. Mechanisms and consequences of competition between an alien mussel, Mytilus galloprovincialis, and an indigenous limpet, Scutellastra argenvillei. Journal of Experimental Marine Biology and Ecology 317:127–142. Sutherland, J.P. 1978. Functional roles of Schizoporella and Styela in the fouling community at Beaufort, North Carolina. Ecology 59:257–264. Thieltges, D.W. 2005. Impact of an invader: epizootic American slipper limpet Crepidula fornicata reduces survival and growth in European mussels. Marine Ecology Progress Series 286:13–19. Thieltges, D.W., Reise, K., Prinz, K., and Jensen, K.T. 2009. Invaders interfere with native parasite–host interactions. Biological Invasions 11:1421–1429. Trimble, A.C., Ruesink, J.L., and Dumbauld, B.R. 2009. Factors preventing the recovery of a historically overexploited shellfish species, Ostrea lurida Carpenter 1864. Journal of Shellfish Research 28:97–106. Trowbridge, C.D. 1999. An assessment of the potential spread and options for control of the introduced green macroalga Codium fragile spp. tomentosoides on Australian shores. Center for Research on Introduced Marine Pests and CSIRO Marine Research. United States Department of Agriculture (USDA). 2009. The Census of Agriculture. www.agcensus.usda.gov (accessed on January 2010). United States Geological Survey (USGS). 2009. Nonindigenous Aquatic Species. nas.er.usgs.gov/ (accessed on January 2010). Valéry, L., Fritz, H., Lefeuvre, J., and Simberloff, D. 2008. In search of a real definition of the biological invasion phenomenon itself. Biological Invasions 10:1345–1351. Vander Zanden, M.J., Hansen, G.J.A., Higgins, S.N., and Kornis, M.S. 2010. A pound of prevention, plus a pound of cure: early detection and eradication of invasive species in the Laurentian Great Lakes. Journal of Great Lakes Research 36:199–205. Verlaque, M., and Latala, A. 1996. Sur une espe`ce japonaise de Chondrus (Gigartinaceae, Rhodophyta) accidentellement introduite dans l’e′tang de Thau (France, Me′diterrane′e). Cryptogamie Algologie 17:153–164.

424

Shellfish Aquaculture and the Environment

Verlaque, M., Steen, F., and De Clerck, O. 2009. Regulopteryx (Dictyotales, Phaeophyceae), a genus recently introduced to the Mediterranean. Phycologia 48:536–542. Wangkulangkul, K., and Lheknim, V. 2008. The occurrence of an invasive alien mussel Mytilopsis adamsi Morrison, 1946 (Bivalvia: Dreissenidae) in estuaries and lagoons of the lower south of the Gulf of Thailand with comments on their establishments. Aquatic Invasions 3:325–330. Watson, D.I., Shumway, S.E., and Whitlach, R.B. 2009. Biofouling and the shellfish industry. In: Burnell, G., and Allan, G. (eds.), New Technologies in Aquaculture: Improving Production Efficiency, Quality and Environmental

Management. Woodhead Publishing Limited, Oxford, pp. 317–336. Wonham, M.J. 2004. Mini-review: distribution of the Mediterranean mussel Mytilus galloprovincialis (Bivalvia : Mytilidae) and hybrids in the Northeast Pacific. Journal of Shellfish Research 23:535–543. World Wildlife Federation (WWF). 2011. www. worldwildlife.org/what/globalmarkets/aquaculture/dialogues-molluscs.html Zhu, S.M., Saucier, B., Durfey, J., Chen, S.L., and Dewey, B. 1999. Waste excretion characteristics of Manila clams (Tapes philippinarum) under different temperature conditions. Aquacultural Engineering 20:231–244.

Chapter 15

Balancing economic development and conservation of living marine resources and habitats: the role of resource managers Tessa L. Getchis and Cori M. Rose

Introduction Bivalve shellfish production represents a large and growing segment of the U.S. and global seafood industry, with nearly 20% of domestic and 27% of worldwide aquaculture production being attributed to shellfish aquaculture (U.S. Department of Agriculture National Agricultural Statistics Service 2006; Food and Agricultural Organization of the United Nations 2008). Although production is increasing, there is uncertainty and public concern with respect to the ecological effects of aquaculture practices, which threatens to constrain further development of the industry (Food and Agricultural Organization of the United Nations 2008; National Research Council

2010). This ambiguity has led to both local and sweeping national changes to the manner in which shellfish aquaculture is regulated and the husbandry standards by which shellfish producers must adhere to, in the United States and elsewhere. Evaluation of the effect of bivalve shellfish aquaculture on the marine and estuarine nearshore environment has, until recently, predominantly focused on near-field assessment of the effects of highly intensive operations outside of the United States (Cranford et al. 2003). It has been assumed that because, unlike finfish aquaculture, shellfish cultivation occurs in open water without the addition of feed, the likelihood for adverse effects was low. This incomplete rendering led to a general mindset

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 425

426

Shellfish Aquaculture and the Environment

Figure 15.1 Longline mussel culture.

that shellfish cultivation is a “benign” use of the marine environment. However, the continuing controversy over the effects of marine fish farming and the common misconception that shellfish aquaculture incorporates similar husbandry techniques has driven environmentalists, scientists, regulators, and industry in the United States and elsewhere to reconsider this view. Consequently, there has been more of an impetus to fully investigate and properly document the potential for immediate and local effects resulting from shellfish farming, as well as the probability for longer-term and estuary-wide effects to occur. The purpose of this chapter is to highlight the current framework for and emphasize the national trends in the regulation of bivalve shellfish aquaculture within the United States.

Environmental effects The principal concerns with respect to the interaction of shellfish aquaculture and the environment are generally considered to be

water quality degradation (Chapter 7); changes in sediment chemistry and composition (Chapter 10); habitat degradation, altered biodiversity and community structure (Chapters 5 and 9); the introduction of nonnative species including predators, pests, and disease; the spread of harmful algal blooms (Chapters 13 and 14); and the loss of genetic diversity in wild shellfish populations (Chapter 12). Some studies have indicated that intensive shellfish production can result in ecosystemlevel changes in water quality (Souchu et al. 2001), phytoplankton biomass (Nichols et al. 1990; Banas et al. 2007), and benthic sediment chemistry and composition (Dahlback and Gunnarsson 1981; Kaspar et al. 1985; Figueras 1989; Grant et al. 1995; Cranford et al. 2003) (see Fig. 15.1). Direct and indirect impacts to critical habitats and changes in biodiversity have been linked to operations that utilize submerged or floating cultivation structures, intertidal rearing methodologies such as predator netting, or harvesting gear such as dredges. Physical disturbance such as scouring and

Development and conservation: Role of resource managers

427

Figure 15.2 Aquaculture operations in eelgrass beds.

sedimentation can indirectly impact biodiversity by altering or removing essential habitat. Of particular interest to resource managers is the potential for adverse effect to threatened or endangered species (e.g., mammals, birds, turtles, and migratory fish) and protected habitats such as submerged aquatic vegetation (SAV) (see Fig. 15.2). Such disturbances to federally protected species such as Pacific salmon (Simenstad and Fresh 1995; Thom 2009) as well as negative impacts to special habitat such as seagrasses have been documented (Everett et al. 1995; Tallis et al. 2009). However, these impacts are very site and gear specific. In addition, intentional and accidental introductions of aquatic organisms have resulted in significant and often undesirable consequences to aquatic environments (Nichols et al. 1990). Aquaculture, in general, is now the leading cause of nonnative introductions (Chapter 14 in this book) to inland aquatic ecosystems (Welcome 1988) and introductions to marine waters have also been documented. In addition to the unintended spread of the target culture species, the proliferation of predators, pests,

and disease-causing organisms is a significant ecological and economic problem (McKindsey et al. 2007). As such, most states now require a shellfish transport permit that certifies that the product (larvae, seed, or adult shellfish) is a native species, disease/toxin-free, and will not pose harm to existing shellfish populations. Recent studies have demonstrated the potential for a loss of genetic diversity when hatchery stocks interbreed with wild populations of shellfish (Arnold et al. 2004, 2009). Interbreeding is especially difficult to prevent or control as many molluscan shellfish are broadcast spawners and larvae can be carried long distances away from their reproductive populations (Chapter 12 in this book). This is not to suggest that the beneficial aspects of shellfish culture should be overlooked or underestimated. The current body of knowledge and scientific documentation is replete with the value of shellfish and shellfish aquaculture for their ability to improve water quality and clarity making the environment more suitable for aquatic organisms.

428

Shellfish Aquaculture and the Environment

Filter-feeding molluscs achieve this by reducing excess phytoplankton associated with eutrophication and other fine particulate matter, and transforming inorganic nutrients into bioavailable forms (Reusch et al. 1994; Peterson and Heck 2001a, 2001b; Newell 2004; Newell and Koch 2004). In addition, the shellfish, the physical aquaculture structures themselves, and in some cases the associated shell deposits can all serve to provide a

level of three-dimensional complexity and hard substrate that results in enhancement of the diversity and abundance of biota that is similar or greater than natural marine habitats (Brehmer et al. 2003; Crawford et al. 2003a; DeAlteris et al. 2004; O’Beirn et al. 2004; Roycroft et al. 2004; Lindahl et al. 2005; Pinnix et al. 2005; Wechsler 2006; Powers et al. 2007; Clynick et al. 2008; Erbland and Ozbay 2008) (see Fig. 15.3).

(A)

(B)

Figure 15.3 Physical aquaculture structures as enhanced three-dimensional structure and hard substrate for other organisms. (A) A lobster (inset) in a cage full of oysters; (B) a godwit foraging among cages.

Development and conservation: Role of resource managers

The effects of aquaculture on the estuarine environment, whether beneficial or detrimental, are highly dependent on the species cultured and the type, scale, intensity, and frequency of the activity (Kaiser et al. 1998; Folke et al. 2004; Vaudrey et al. 2009; see also Chapter 9 in this book). In addition, the intensity and temporal and spatial extent of the effects may vary in response to the local environment’s resistance to change (Bradbury et al. 1983; Simenstad and Fresh 1995). Most experts agree that scale and intensity are two of the more important management considerations when permitting aquaculture and that understanding the effect of scale is critical to planning for sustainable aquaculture production (Dumbauld et al. 2009). Unfortunately, information on these effects is lacking as few studies in the United States until recently have assessed the cumulative and additive effects of multiple farms and estuary-wide effects of shellfish aquaculture.

Social effects In addition to environmental effects, other major considerations in the siting of aquaculture operations include the impact to navigation and other competing uses of the coastal zone, and concern that the activity will result in undesirable aesthetic impacts such as visual and noise pollution. Societal issues such as these also have the potential to impede the growth of the bivalve shellfish aquaculture industry along populated coastlines (see Fig. 15.4). Although these impacts are difficult to assess and mostly due to subjective interpretation, their consideration is nonetheless necessary as coastal waters are generally held in “public trust” for the benefit of all (Fernandez 1996). Although it is important to mention that societal concerns can be critical to the successful siting of shellfish aquaculture projects, we note that their discussion is beyond the scope of this chapter.

429

The uncertainty with respect to environmental interactions and social conflicts associated with shellfish aquaculture has led to an increasingly complex review process for shellfish aquaculture projects. The permitting process is not only daunting for the industry, but extremely challenging for the regulatory community that is responsible for balancing economic development, conservation of living marine resource and habitats, and other uses of the coastal zone. Hence, regulators often take a conservative approach to siting aquaculture projects rather than accepting the responsibility for unknown or unforeseen risks (Dumbauld et al. 2009). There are many competing issues that resource agencies must often consider with respect to shellfish aquaculture and the environment. The main focus of this manuscript is to elaborate on the potential effects (beneficial or adverse) that bivalve culture may present to marine species and habitats. However, resource managers not only regulate the shellfishing industry but they also on the other hand promote its existence. As such, resource managers must also monitor water quality and enforce no-discharge zones that can impact shellfish production and harvesting. There are also regulations such as health and sanitation standards and requirements that growers recognize as critical to production (Aspen Systems Corporation 1981). In addition, both resource managers and producer associations are in the process of or have developed management practices that address public concern and will foster sustainable growth of the industry.

Regulatory framework for shellfish aquaculture in the United States Regulation of the shellfish aquaculture industry in the United States often includes review and agency oversight at one or more local, county, state, and federal levels. Consequently, a permit from several of these government

430

Shellfish Aquaculture and the Environment

(A)

(B)

Figure 15.4 Geoduck aquaculture. (A) Predator exclusion devices; (B) geoduck farm in a tideland.

bodies may be required for establishment of an aquaculture operation. It may even be necessary to obtain multiple permits from a single agency, particularly at the state level. In addition to securing leases or deeds for shellfish

cultivation grounds, permits or licenses may be required for the operation of aquaculture vessels or land-based facilities, for placing and marking aquaculture gear, and for transplanting or harvesting product, among others.

Development and conservation: Role of resource managers

Programmatic agreements, joint procedures, or other administrative processes are sometimes used at the state level to reduce the burden of redundant agency review of aquaculture activities considered to have negligible or minor impact on natural resources and/or navigation. Still, the complexity of the regulatory structure is considered the primary constraint to the development of the aquaculture industry. The uncertainty related to the interaction of shellfish aquaculture and the environment further complicates this already confusing regulatory process.

Near-shore aquaculture Currently, the majority of shellfish culture in the United States occurs within the boundaries of state waters, less than 3 NM from shore (Dewey et al. 2007). Oversight for this activity is administered through an amalgam of federal and state coastal management law and regulation not always directly applicable, or easily adaptable, to the needs of the shellfish industry. A continuing industry-wide hurdle is the fact that commercial aquaculture is largely a private sector initiative that seeks to benefit from resources commonly held by states and regulated by government entities for the public trust. Because some forms of shellfish culture systems, such as surface or submerged gear, can be exclusionary in terms of their ability to displace or otherwise impede a variety of other recreational and commercial uses of the coastal zone, the siting of near-shore aquaculture activity can be subject to a substantial amount of controversy (National Research Council 1992; Cicin-Sain et al. 2004). As such, applications for this type of aquaculture draw in public governmental bodies that play a role in protecting the environment and managing the public trust through policies such as coastal zone management and the navigable servitude, which again may not be designed to foster development of the aquaculture industry.

431

Historically, aquaculture was considered and regulated in many states as an activity related to fisheries (Rubino and Wilson 1993), or recognized by states as an agricultural activity with oversight under their respective agricultural agencies. In many instances, this approach resulted in a streamlined state review process through which all permits and licenses could be acquired from a single agency. However, the simplicity of this strategy was thwarted with the passage of federal legislation such as the Coastal Zone Management Act (CZMA) of 1972. The CZMA encouraged states to exercise their responsibilities with respect to the use of land and water resources of the coastal zone and required the development and implementation of management programs to balance competing uses of the U.S. coast and near-shore waters. As such, any activity in coastal waters with the potential to affect the area’s use, resources, or its character is required to be evaluated for consistency with the policies of a particular state’s coastal zone management program. The paradox being that coastal zone management usually does not fall within the auspices of a state’s department of agriculture, therefore requiring the engagement of more than one state agency in the review of applications for aquaculture activity. To address this disjointed process, the CZMA was amended in 1990 and again in 1996 to encourage planning and provide funding (1) for the adoption of procedures and policies to evaluate and facilitate the siting of public and private aquaculture in the coastal zone; (2) to enable states to formulate, administer, and implement strategic plans for marine aquaculture; and (3) to develop a coordinated process among multiple state agencies to regulate and ultimately permit aquaculture facilities in the coastal zone. Unfortunately, the impact of these amendments on aquaculture development varied and, in many states, these amendments did little to unravel the complexity of the permitting process.

432

Shellfish Aquaculture and the Environment

For a traditional shellfish aquaculture project (not requiring water diversion, withdrawal, or discharges), there are two main components to authorization. They are the leasing (or purchasing) of cultivation grounds and the permitting of aquaculture structures and/or resultant benthic modifications, in tidal or navigable waters. Leasing of public lands in the near-shore environment is almost always handled at some combination of the municipal, county, or state level, in accordance with statute or regulation. However, the federal government is charged by Congress with regulating the installation of all structures, including aquaculture gear, in coastal and navigable waterways to ensure protection of their course, condition, location, or capacity for commercerelated purposes. The federal government is also required to issue permits for any activity that involves a discharge of dredged or fill material, which includes the deposition of shellfish and or cultch, in waters of the United States (defined by regulation to include resources such navigable waters, inland rivers, lakes, streams, and wetlands at 33 CFR 328).

Permitting agencies and authorization The U.S. Army Corps of Engineers (USACE), through its Regulatory Program (33 CFR 320– 332), administers the laws passed by Congress to regulate work in waters and wetlands, and in most cases manages the coordination process for application review with federal resource agencies and state regulatory agencies. The USACE’ pertinent authorities are Section 10 of the Rivers and Harbors Act of 1899 (RHA) (33 USC 403) for structures and preservation of navigation, and Section 404 of the Clean Water Act (CWA) (33 USC 1344) for the discharge of dredged or fill material in waters or wetlands. The agency is also required by the Council of Environmental Quality

(CEQ), which is the White House coordinator of federal environmental review, to implement the National Environmental Policy Act (NEPA) (42 USC 4321) for its permit actions as well as to comply with other environmental directives such as Executive Orders and Code of Federal Regulation. NEPA requires federal agencies such as the USACE to consider environmental values into their decision-making processes and to take into account the environmental impacts of proposed activities and reasonable alternatives to those activities. To meet NEPA requirements, the USACE must prepare an Environmental Impact Statement, which is reviewed and filed by the U.S. Environmental Protection Agency (USEPA). In addition to environmental impacts, the USACE must consider the effects of proposed projects on other factors such as water quality, navigation, public use, and enjoyment of navigable waters, and aquatic health management. The USACE is required to comply with a myriad of other federal agency regulations. These include Section 401 of the CWA that addresses discharges or effluents with the potential to impact water quality, Section 307 (c) (3) (A) of the aforementioned CZMA (16 USC 1456) that mandates that the USACE coordinate with the state coastal zone management agency to ensure that the proposed activity is consistent with the state’s coastal zone management plan, the Endangered Species Act (ESA), the Marine Mammal Protection Act (16 USC 1361), the National Historic Preservation Act (16 USC 470), Section 302 of the Marine Protection, Research, and Sanctuaries Act, and the Magnuson–Stevens Fishery Conservation and Management Act (MSFCMA) (50 CFR 229.2). The USACE administers the regulatory review process and generally coordinates with three key federal agencies during the routine review of applications for shellfish aquaculture in near-shore waters. They are the National Oceanic and Atmospheric Administration’s National Marine Fisheries Service (NOAA

Development and conservation: Role of resource managers

NMFS) within the U.S. Department of Commerce, the USEPA, and the U.S. Fish and Wildlife Service (USFWS) within the U.S. Department of Interior. Depending on the potential for environmental and social effects, the agency may coordinate with any number of other federal agencies including, but not limited to, the U.S. Coast Guard, the Advisory Council on Historic Preservation, and various agencies with the Departments of Commerce and Interior to address impacts relevant to the expertise of that agency during a review. An example of a “roadmap” for the permitting process for near-shore shellfish aquaculture is provided in Figure 15.5. The USEPA’s involvement in the review of shellfish aquaculture activity stems from its shared oversight and development of policy under the CWA. The CWA is the legislation that authorizes the Secretary of the Army; acting through the Chief of Engineers, to issue permits, after notice and opportunity for public hearing, for the discharge of dredged or fill material into the waters of the United States (33 CFR 323). The selection and use of “disposal” sites are identified in accordance with guidelines developed by the Administrator of USEPA in conjunction with the Secretary of the Army and published in 40 CFR 230. While the USACE administers many of the day-today elements of the CWA 404 program, the responsibilities of USEPA under this section include developing and interpreting environmental criteria used in evaluating permit applications; determining scope of geographic jurisdiction, approving, and overseeing state assumption of the 404 Program identifying, activities that are exempt; reviewing/ commenting on individual permit applications; and enforcing Section 404 provisions. The Fish and Wildlife Coordination Act of 1934 (FWCA) (16 USC 661) provides the basic authority for the involvement of the USFWS and NOAA NMFS in evaluating impacts to fish and wildlife from proposed projects and provides the avenue to fulfill the

433

will of Congress to protect the quality of the aquatic environment as it affects the conservation, improvement and enjoyment of fish and wildlife resources. Reorganization Plan No. 4 of 1970 transferred certain functions, including certain fish and wildlife-water resources coordination responsibilities, from the Secretary of the Interior to the Secretary of Commerce. Under the FWCA, federal agency activities that propose to modify any body of water/wetlands must first consult with the USFWS or NOAA NMFS, as appropriate. Both of these federal agencies also have consultation responsibilities as it pertains to the issuance of a permit where the action has the potential to result in adverse effect to listed species or their critical habitat under the ESA. The MSFCMA requires that the USACE consult with the Department of Commerce (delegated to NOAA NMFS) whenever the agency proposes to authorize, fund, or undertake an action that may adversely affect any essential fish habitat (EFH), such as seagrasses. Further, the act requires that NOAA NMFS recommend conservation recommendations to avoid, mitigate, or offset the impact of any activity that may adversely affect EFH. In the event that the USACE receives conservation recommendations from NOAA NMFS, it is not required to follow the recommendations, but it must provide a detailed response to the agency explaining the reasons for not following the recommendations and describing the measures it will take to avoid, mitigate, or offset the impact of the activity on EFH. Needless to say, both the USFWS and NOAA NMFS play an important advisory role in the USACE’s permit review process for shellfish aquaculture activity. It is this complex, sometimes inconsistent, and often unpredictable and overlapping collection of state and federal policy and regulatory review highlighted above, which is often cited as the main reason why the goal for sustainable and profitable shellfish aquaculture

APPLICANT Contacts State Department of Agriculture, receives and submits application package

MUNICIPAL SHELLFISH COMMISSION Provides lease documentation and comments if project is to be located in town waters USACE Seeks input from various agencies; conducts a technical review, reports concerns at Joint Permit Processing Screening Meeting

DEPARTMENT OF AGRICULTURE Reviews application, confirms receipt with applicant, forwards application to various agencies

DEPARTMENT OF ENVIRONMENTAL PROTECTION Office of Long Island Sound Programs (DEP OLISP) Determines if project is exempt or not exempt from DEP/OUSP permits; sends letter notify applicant

DEP (various divisions) Issue DEP permits (if applicable)

NOAA NMFS USFWS

DEP/Fisheries, DEP/Boating Determine if the project results in substantial, minimal or no concerns, reports back to DEP/OLISP

Joint Permit Processing Screening Meeting With substantial concerns:

DEP (various divisions) Send letter to applicant stating if DEP permits are required

USEPA

USACE Begins Individual Permit* process

DEP/OLISP Conducts Federal Coastal Zone Management Plan Consistency Review

With minimal concerns:

With no concerns:

DEP/OLISP Provides sign-off for Programmatic General Permit (PGP)**

DEP/OLISP Provides sign-off for Programmatic General Permit (PGP)**

USACE Issues PGP with conditions to applicant

USACE Issues PGP to applicant

DEP/OLISP Identifies if activity concurs with Coastal Zone Management Plan Consistency, applicant notified that activity is approved

USACE Issues Individual Permit to applicant

DEPARTMENT OF AGRICULTURE Issues Certificate for Aquaculture Operations Figure 15.5 An example roadmap of the permitting process for near-shore shellfish aquaculture. Source: Getchis et al. 2008.

434

Development and conservation: Role of resource managers

production is unlikely to be reached in the United States (Aspen Systems Corporation 1981; Duff et al. 2003).

Permit types Permits for shellfish aquaculture from the USACE can take many forms depending on which region, division, or district that the proposed project is to be located. The agency has 8 regions, 11 divisions, and 36 district offices and is decentralized in its operation to allow the various state and federal regulatory programs to complement one another and reduce duplication of effort (U.S. Army Corps of Engineers 1999). In many cases, it is the scope of the project and nature of the potential impacts that will determine which review procedure is most applicable. The majority of new shellfish aquaculture activities in the nation have been historically, and continue to be, reviewed under an Individual or “Standard” Section 10 permit process that requires submission of an application directly to the USACE. This procedure is a common form of review for activities that may have more than minimal adverse impact on the nation’s waters. An application of this nature generally requires a full public interest review involving coordination with other agencies, interested parties, and the general public through issuance of a public notice and an opportunity for public hearing, as well as consideration of a broad range of potential impacts (individual and cumulative) such as the effect of the facility/structure on recreation, fish, and other wildlife, pollution, economic factors, safety, aesthetics, protection of navigational integrity, cultural values, and water quality. This review is usually commensurate with the potential level of impact to the aquatic resource. Consequently, it may also involve consideration of less environmentally damaging configuration alternatives or other locations and it will often result in a more

435

thorough review of the potential environmental and socioeconomic effects of the proposed activity than other types of permits. A final determination to issue a permit for an activity will ultimately result in an Environmental Assessment and Statement of Findings, and the USACE cannot issue a permit for a project if the proposed work is not in compliance with other laws such as Section 401 of the CWA and the CZMA. Depending on where in the country the proposed project is to be located, abbreviated processes termed Letters of Permission (LOPs) or General Permits (including Nationwide Permits [NWPs], Regional Permits, or State Programmatic Permits) may also be available. An LOP is a type of permit usually for work under Section 10 of the RHA issued at a regional level, that includes coordination with federal and state fish and wildlife agencies and a public interest evaluation, but without the publishing of an individual public notice (33 CFR 325.2). An LOP procedure is an alternative process for evaluating standard permit applications. It requires that the LOP procedure itself is advertised through public notice that allows agencies, interested parties, and the general public to provide comment and have an opportunity for public hearing on establishment of the process. It is designed to reduce administrative procedure and to expedite permit decisions for cases that include only minor work in waters of the United States; do not have significant individual or cumulative environmental impacts; and should encounter no appreciable opposition. However, similar to a standard permit, a CZMA consistency concurrence must be obtained or presumed. LOPs may also include general conditions and appropriate case-specific provisions necessary to protect the environment, including natural and cultural resources or other aspects of the public interest. NWPs are those permits issued at a national level to authorize specific types of minor

436

Shellfish Aquaculture and the Environment

activities and minimize evaluation time. NWPs are authorizations that can be used throughout the United States, with some exceptions. Their purpose is to reduce the regulatory reporting burden for work with no more than minimal impact to the aquatic environment while maintaining adequate protection of aquatic resources. The thresholds for the impacts and the types of activities allowed under the NWPs are established as national policy, and therefore are not regionally flexible. Consequently, some USACE districts and divisions have suspended or revoked these permits and replaced them with alternate forms of abbreviated processes. Where NWPs are used, regional conditions are often established to ensure that their implementation is consistent with meeting the program’s goals of streamlining federal and state review for those activities that are demonstrated to have no more than minimal impact to the aquatic environment. There is only one NWP specifically designed to address shellfish aquaculture activities. It is NWP Number 48 for authorization of existing structures for rearing of shellfish within a permitted project boundary and the discharge of fill material, which may be necessary for shellfish seeding, rearing, cultivating, or transplanting, and the related discharges associated with harvesting activities. The permit is applicable only to existing commercial shellfish aquaculture projects and does not apply to new operations or expansion of an existing facilities, to the cultivation of additional species (not previously cultivated by the enterprise), to the construction of structures such as docks and piers, or to the deposition of shell material into the water as waste. New shellfish aquaculture operations and expansion of existing facilities continue to require review through one of the other federal review processes. This activityspecific permit for shellfish culture reflects a 2002 modification to the USACE regulatory definition of “fill material,” whereby the

measure for regulation of a discharge under Section 404 of the CWA (33 USC 404) was changed from one of a “primary purpose” test to that of an “effects-based” test. Consequently, existing shellfish aquaculture projects resulting in a discharge of fill material, including the placement of shellfish or cultch material that would change the elevation of the bottom below the high tide line, could be regulated as a discharge where previously regulatory review was not required. The 2002 USACE change in definition combined with the perception that this case-by-case evaluation results in inconsistent application of CWA 404 jurisdiction for shellfish aquaculture has created significant confusion within the industry (Dewey 2005). The last category of authorization is Regional or State Programmatic General Permits. These abbreviated processes are sometimes developed for general categories of activities or for a state program when the activities being evaluated are similar in nature and cause minimal environmental impact (individually and cumulatively). The intent of this process is to streamline review and reduce the duplication of regulatory control by state and federal agencies. Several states have established the beginning of, or continue to improve upon, a regulatory framework for the leasing and permitting of aquaculture in near-shore waters. These include, but are not limited to, California under the California Sustainable Oceans Act, Oregon under its Territorial Sea Plan Part II, Hawaii under Ocean and Submerged Lands Leasing (Chapter 190D), Washington under Revised Code of Washington Aquaculture Leasing Statutory and Regulatory Framework, Maine under Maine Conservation Marine Resources state statute, and New Jersey under the New Jersey Aquaculture Development Act. These efforts to streamline the permitting process should ultimately reduce the effort and time required to complete the

Development and conservation: Role of resource managers

437

BASELINE STATE WATERS (0–3 NM) TERRITORIAL SEA (0–12 NM) CONTIGUOUS ZONE (12–24 NM) EXCLUSIVE ECONOMIC ZONE (12–200 NM) EDGE OF CONTINENTAL MARGIN CONTINENTAL SHELF CONTINENTAL SLOPE RISE

Figure 15.6 Schematic of state and federal maritime zones in the United States.

application and review process for shellfish aquaculture.

Offshore aquaculture Currently, only a small percentage of production actually results from shellfish cultivation between 3 and 200 mi offshore in the U.S. Exclusive Economic Zone (EEZ) (see Fig. 15.6), and the activities that contribute to this production are predominantly experimental in nature. The majority of open-water offshore bivalve culture occurring in recent years has been in the form of pilot projects, predominantly for mussels, seeking to lay the groundwork for competitive technically and economically feasible commercial enterprises. Much of this activity has taken place offshore in, or adjacent to, waters of the states of California, Massachusetts, Rhode Island, and New Hampshire. At least one of these demonstration projects has led to a subsequent

commercial-scale operation in the offshore oceanic environment (R. Barnaby, pers. comm.). The University of New Hampshire’s Open Ocean Aquaculture Demonstration Project in the Gulf of Maine with the Portsmouth, New Hampshire, Commercial Fishermen’s Cooperative is one of the longest-running offshore examples, developed in 1997, with the intent to demonstrate the biological, engineering, operational, and economic feasibility of culturing both finfish and shellfish in unprotected, oceanic environments (Ward et al. 2007). Today the project consists of an ocean spar submerged sea grid cage system manufactured by Ocean Spar Technologies, LLC, for native finfish such as halibut, and haddock, an associated remote controlled feeding buoy, and submerged longlines for suspended ropes or socks for the rearing of mussels. The project has over 10 years of quantitative and qualitative assessment including physical substrate characteristics, water quality monitoring, benthic community infauna and epifauna

438

Shellfish Aquaculture and the Environment

analysis, and practical engineering associated with a very high-energy offshore environment that exhibits rapid flushing and a high carrying capacity (Langdon 2002). Reportedly, bivalve mollusc efforts have been thwarted by the status of marine-based aquaculture, in general, in that the activity is not directly addressed in federal statute in a holistic manner that gives a single agency authority for the siting of facilities and the ability to manage for a sustainable industry with intent to maximize the production of seafood (Fletcher and Neyrey 2003; Mittal 2008). The legal and regulatory environment surrounding the offshore aquaculture industry is cited consistently as one of the major hurdles to its development in the United States (Fletcher and Neyrey 2002; Langdon 2008). In 1978, the National Research Council Committee on Aquaculture found that the procedures required to obtain authorizations for offshore aquaculture were a “severe deterrent” to the development of the industry (National Research Council 1978). Subsequently, U.S. Congress cited the “diffused legal jurisdiction” and “lack of supportive Government policies” when it encouraged the development of a U.S. national aquaculture policy in 1980. Not surprisingly, an inadequate regulatory regime, that is, one comprising laws adopted to address problems or industries other than aquaculture, continues to hinder aquaculture across the globe and in the United States (Browdy and Hargreaves 2009). Under the National Aquaculture Act (16 USC 2801), the Department of Agriculture is the lead federal agency to provide for the development of aquaculture in the United States. However, by virtue of its regulatory authority for structures and other activities in near-shore waters, the USACE is the lead federal agency with primary jurisdiction for the evaluation of impacts and the issuance of permits in offshore waters under the Outer Continental Shelf Lands Act (OCSLA) (43

USC 1333). The act provides for the regulation of structures and devices temporarily or permanently attached to the seabed in the EEZ for the purpose of exploring for, developing, or producing resources from the outer continental shelf. A permit from the U.S. Coast Guard under Aids to Navigation (33 CFR 62) may also be required. Other authorizations that may be necessary for marine-based offshore facilities, but not normally applicable to molluscan shellfish systems, include approval from the USEPA activities that would result in a discharge from a point source into ocean waters under the National Pollutant Discharge Elimination System (NPDES) pursuant to Section 318 of the CWA (40 CFR 122.24), or in the case of a discharge that is categorized as a waste, the Ocean Dumping Act (33 USC 1412). Similar to the near-shore regulatory review process, federal agencies with some level of oversight, but no actual permitting authority, include NOAA NMFS and the USFWS, the Regional Fishery Management Councils under the MSFCMA, the Minerals Management Service (MMS), and the U.S. Department of Interior. NOAA NMFS is responsible for the evaluation of proposals for new facilities in the marine environment that are regulated by other agencies, such as those for aquaculture or oil exploration. Evaluation of these activities is undertaken to ensure that marine mammals (Marine Mammal Act), endangered species (ESA), and national marine sanctuary resources (National Marine Sanctuaries Act) are protected. The agency also coordinates with eight regional fishery management councils to manage fishing activity and protect EFH in federal waters. Projects are reviewed to assess the potential for impact to EFH under the MSFCMA as a result of aquaculture activity. The MMS is given authority under the OCSLA for management of lease sites on submerged lands of the outer continental shelf. However, their participation within

Development and conservation: Role of resource managers

marine-based aquaculture and shellfish aquaculture in particular is likely to be minimal unless the facility is located proximal to an existing lease or attached to an oil or gas platform. Depending on where a proposed project is to be located within the country, concurrence from the U.S. Department of Interior, through the Secretary of Interior, may be required if the offshore aquaculture facilities may be situated on leases or easements, approved under the OCSLA or within 1 mi of any other permitted facility or for which a plan has been approved under the OCSLA. Although the number of federal agencies, their narrow regulatory focus, and what appears to be a lack of continuity appears to be disjointed for open ocean aquaculture, the detailed interaction and detailed coordination between agencies within the existing procedures still has the potential to develop into a comprehensive, streamlined permitting and monitoring program for the future, if the various issues of underwater land management can be resolved. NOAA is the lead federal agency for marine aquaculture policy and is pursuing the development of a comprehensive national policy and regulatory framework for sustainable marine aquaculture in federal waters, a task initiated in 2005 as heretofore unenacted legislation entitled The National Offshore Aquaculture Act (U.S. Congress, Senate 2005, S6238; U.S. Congress, Senate 2007, S7665). The agency’s priorities for program development can be found within its 2007 10-year plan for aquaculture development (NOAA 2007). If this or similar subsequent legislation is enacted, NOAA would become the lead agency responsible for coordination and evaluation of offshore aquaculture activities in close coordination with other federal resource and regulatory agencies (including USACE and USEPA), state agencies, tribes, and stakeholders. The National Sustainable Offshore Aquaculture Act was introduced into the U.S.

439

Congress House of Representatives in 2009 (U.S. Congress, House 2009, H4363). Its purpose is to establish an Office of Sustainable Offshore Aquaculture within NOAA NMFS that would assume responsibility for implementation of regulation for offshore aquaculture in the EEZ. The legislation proposes a comprehensive approach to the development and execution of a regulatory process and promotes the identification of an integrated framework to set research priorities for the promotion of sustainable offshore aquaculture in the EEZ. In response to a Government Accounting Office review of this proposed legislation and with consideration of the Obama administration’s establishment of a Council on Environmental Quality Interagency Ocean Policy Task Force (IAOP Task Force), NOAA is reassessing existing Department of Commerce and NOAA Aquaculture policies to address the current administration’s goals and to enhance opportunities for the establishment of economically and environmentally sustainable U.S. aquaculture. The essence of this directive prevails today as the basis for future regulation of offshore aquaculture. The IAOP Task Force released an interim report in 2009 and posted its Interim Framework for Effective Coastal and Marine Spatial Planning (Interim Framework) for public review and comment. Under the proposed process, coastal and marine spatial planning would be regional in scope, and developed cooperatively among federal, state, tribal, local authorities, and regional governance structures, with substantial stakeholder and public input. Along these lines, NOAA has scheduled a series of “listening sessions” around the country in 2010, in which the agency will obtain comments and hear recommendations from the public pertaining to all forms of marine aquaculture (all species, nearshore, and offshore). These sessions will provide a basis for identifying the scope and objectives of a draft national policy for

440

Shellfish Aquaculture and the Environment

sustainable marine aquaculture in the United States.

Environmental best management practices (BMPs) BMPs have been developed and implemented in an effort to reduce or minimize adverse environmental and social effects, food safety issues, and other public concerns resulting from proposed shellfish aquaculture activity in near-shore and offshore waters. BMPs are general overarching principles and specific procedures or methodologies used to guide the day-to-day operation of aquaculture businesses. Compliance with BMPs can be voluntary or mandated and can by driven by industry, regulatory agencies, environmental groups, or other nongovernmental organizations. Industry-driven and governmentmandated BMPs have been reviewed by Creswell and McNevin (2008) and Jensen and Zajicek (2008), respectively. An overview on BMPs directly related to shellfish aquaculture is provided in Chapter 3, and as such, we provide only a brief summary of relevant programs in the following sections.

Industry-driven BMPs Industry-driven BMPs have been developed by state industry groups and regional associations as a proactive measure to solicit public support for sustainable industry development. The Pacific Coast Shellfish Growers Association (PCSGA) and East Coast Shellfish Growers Association (ECSGA) have developed environmental codes of practice and BMPs, respectively. Both organizations are currently conducting outreach projects to instruct industry members on how to adopt and implement such practices. Participation is voluntary, but peer pressure may influence producers to embrace these measures in order to secure additional market share.

Government-facilitated BMPs Driven by concerns that aquaculture may pose adverse effects to the environment, restrict traditional uses of the coastal zone, result in food safety issues, or other public concerns, several states have established either voluntary or mandatory best management or codes of practice for shellfish aquaculture. Voluntary BMP programs now exist in Maine,1 Massachusetts,2 Maryland,3 New Jersey,4 Virginia,5 and Washington,6 while the state of Florida7 has implemented the first government-mandated shellfish BMP program (Jensen and Zajicek 2008).

Environmental marketing and other incentive programs Both within and outside of the United States, government agencies, environmental groups, and other nongovernmental organizations have developed formal policies encouraging or

1

Maine Aquaculture Association, Maine Aquaculture Code of Practice: www.maineaquaculture.com/Code_of_Practice_ v1.pdf. 2 D.F. Leavitt, Best Management Practices for the Shellfish Industry in Southeastern Massachusetts: www.mass.gov/agr/ aquaculture/docs/Shellfish_BMPs_v09-04a.pdf. 3 Maryland Aquaculture Coordinating Council, Best Management Practices: A Manual for Maryland Aquaculture: www.marylandseafood.org/pdf/best_management_practices_ manual.pdf. 4 Rutgers Cooperative Extension, Recommended Management Practices for Aquatic Farms: Agricultural Management Practices (AMPS) Aquatic Organism Health Management Plan: www.jerseyseafood.nj.gov/aquacultureamp.pdf. 5 M. Oesterling and M. Luckenbach, Environmental Code of Practices for the Virginia Shellfish Culture Industry: www. vims.edu/adv/aqua/MRR 2008_9.pdf. 6 Washington State Department of Natural Resources, 2007 Best Management Practices (BMP’s) for Geoduck Aquaculture on State Owned Aquatic Lands in Washington State: www. dnr.wa.gov/Publications/aqr_aqua_2007bmp.pdf. 7 Florida Department of Agriculture and Consumer Services, Aquaculture Best Management Practices Rule: www. floridaaquaculture.com/publications/P-01499-booklet-07_ BMP_RULE.pdf.

Development and conservation: Role of resource managers

mandating environmentally sustainable aquaculture development through the implementation of BMPs, and certification guidelines to ensure compliance with such policies and practices. Many programs initially focused on the development of guidelines for finfish and crustacean culture but have recently redirected their efforts to address shellfish production. The Food and Agriculture Organization of the United Nations (FAO) has developed both a Code of Conduct for Responsible Fisheries8 and Technical Guidelines on Aquaculture Certification.9 Other programs include the Global Good Agricultural Practice Standards,10 the Codex Principles for Food Import and Export Inspection and Certification,11 and the World Organisation for Animal Health’s Aquatic Animal Health Code.12 The World Wildlife Fund (WWF), through its molluscan shellfish dialogues, is leading a global effort to develop performance-based standards for farmed clams, mussels, scallops, and oysters that will minimize social and environmental issues resulting from aquaculture activity. When complete, the standards will be turned over to the Aquaculture Stewardship Council, which will be responsible for working with independent, third-party entities to certify producers that are in compliance with the standards (World Wildlife Fund 2010). The standards are undergoing review and expected to be finalized in 2010. The certification process will allow for producers to use an 8

Food and Agriculture Organization of the United Nations, Code of Conduct for Responsible Fisheries: ftp://ftp.fao.org/ docrep/fao/005/v9878e/V9878E00.PDF. 9 Food and Agriculture Organization of the United Nations, Technical Guidelines on Aquaculture Certification: library. enaca.org/certification/publications/aquaculture-certificationguidelines-final.pdf. 10 Global Good Agricultural Practice Standards: www.globalgap.org/cms/front_content.php?idart=34. 11 Codex Alimentarius Commission, The Principles for Food Import and Export Inspection and Certification: www.fao.org/ docrep/009/y6396e/Y6396E01.htm. 12 World Organisation for Animal Health, Aquatic Animal Health Code: www.oie.int/international-standard-setting/ aquatic-code/access-online.

441

“ecolabel” identifying the product as environmentally sustainable. Through the efforts of the WWF, the certifying parties it collaborates with, and the growers that comply with the standards, the environmental impacts of certain sectors of the aquaculture industry (e.g., shrimp production) have been drastically reduced and the resulting environmental marketing programs have helped to improve public perception of those sectors. In addition to promoting industry adherence to BMPs, government agencies have offered incentive programs to producers that modify their cultivation practices in a way that reduces adverse and/or promotes beneficial environmental effects of their activity. For example, the U.S. Department of Agriculture’s Natural Resource Conservation Service (USDA NRCS) administers the Environmental Quality Incentives Program (EQIP), which offers technical expertise for planning and designing conservation practices that protect natural resources while maintaining or enhancing productivity. The EQIP program offers cost-share funds to shellfish producers to make such practices affordable. In an effort to improve shellfish health and prevent disease, Massachusetts producers utilized EQIP funds to create buffers or “no-harvest zones” between cultivation and wild harvest areas. In Rhode Island, producers established oyster reefs in an effort to provide new and/or expanded habitat for aquatic organisms and improve water quality. Additionally, several government agencies and nongovernmental organizations within the United States and elsewhere have proposed the use of shellfish to manage eutrophication of coastal waters and incentive programs such as nitrogen credit exchange programs for producers (Chapters 7 and 8 in this book). At least one practical application has been explored by Lindahl et al. (2005), who examined the potential for using intensive mussel culture as an alternative to traditional wastewater treatment plants in an effort to reduce excess nitrogen entering a Swedish estuary. The

442

Shellfish Aquaculture and the Environment

project was successful in terms of removing the targeted level of nitrogen from the system; however, due to unforeseen shellfish market barriers, the operation failed and the wastewater treatment plants were ultimately erected. While shellfish convert, and when harvested, may contribute to the removal of large quantities of nitrogenous wastes, the exact amount of nitrogen removed is highly variable and dependent on a number of factors. Many coastal systems are so heavily eutrophic that it is improbable that shellfish alone can remediate the problem. Even so, a number of U.S states and countries outside of the United States are considering the use of shellfish in addition to other nitrogen reduction measures. As such, researchers are applying theoretical models to assess the nitrogen reduction potential of shellfish farms and for the valuation of nitrogen credits, which may be traded as part of a comprehensive nutrient abatement plan (Ferreira et al. 2009). The adoption of BMPs through environmental marketing and incentive programs may prove valuable in terms of their benefits to both the environment and the shellfish industry. However, some producers still question whether program standards are attainable and affordable for the average grower (R. Rheault, pers. comm.). The costs associated with participating in and adhering to such programs should be incurred by the industry; however, to ensure producer participation such programs should not be cost prohibitive.

Conclusions Sustainable development of the U.S. shellfish aquaculture industry is unlikely unless an industry-specific process of regulatory control that maintains environmental integrity and upholds the important social aspects of the public trust is maintained. While it is essential

to understand how cultivation practices and gear interact with the marine environment, the preservation of cultural and historical or other traditional uses of the coastal zone must be considered in the review of aquaculture applications. Measures intended to manage and regulate activities conducted within the public resource base will continue to have a significant influence on the pace of growth and competitiveness within the commercial aquaculture sector. It is ultimately up to the public to decide what trade-offs they are willing to accept in order to have a sustainable aquaculture industry and seafood supply. The identification and implementation of BMPs as well as the use of marketing and incentive programs to inform the consumer of industry adherence to such practices may allow for the creation of new or expansion of existing markets and lead to greater acceptance of any trade-offs associated with shellfish aquaculture. Additionally, to foster expansion in this sector, the regulatory review process must be transparent and easily navigable for both industry and regulators. This may be achieved, in part, through education and outreach on the permitting processes and government agency responsibilities for regulation of nearshore aquaculture in coastal states and offshore aquaculture within the U.S. EEZ. Ultimately, it may benefit a state’s regulatory bodies, as well as the industry members within that state, to conduct a formal stakeholder review to learn how to best streamline their permitting process. Shellfish cultivation plays an important role in our economy, society, and environment. It fulfills the demand for both staple and highend seafood products, as well as provides a source of jobs and revenue, and is an important part of the cultural heritage of many coastal states in the United States. Finally, shellfish and shellfish aquaculture play a critical and beneficial role in the marine environment that should not be overlooked.

Development and conservation: Role of resource managers

Literature cited Arnold, W., Walters, S., Fajans, J., Peters, S., and Bert, T. 2004. Influence of congeneric aquaculture on hard clam (Mercenaria spp.) population genetic structure. Aquaculture International 12:139–160. Arnold, W.S., Geiger, S.P., and Stephenson, S.P. 2009. Mercenaria mercenaria introductions into Florida, USA, waters: duration, not size of introduction, influences genetic outcomes. Aquatic Biology 5:49–62. Aspen Systems Corporation. 1981. Aquaculture in the United States: Regulatory Constraints. Final report. Rockville, MD: Fish & Wildlife Service, U.S. Department of the Interior. Banas, N.S., Hickey, B.M., Newton, J.A., and Ruesink, J.L. 2007. Tidal exchange, bivalve grazing, and patterns of primary production in Willapa Bay, Washington. Marine Ecology Progress Series 341:123–139. Bradbury, R.H., Hammond, L.S., Reichelt, R.E., and Young, P.C. 1983. Prediction versus explanation in environmental impact assessment. Search 14(1):323–325. Brehmer, P., Gerlotto, F., Guillard, J., Sanguinède, F., Guénnegan, Y., and Buestel, D. 2003. New applications of hydroacoustic methods for monitoring shallow water aquatic ecosystems: the case of mussel culture grounds. Aquatic Living Resources 16:333–338. Browdy, C.L., and Hargreaves, J.A. (eds.). 2009. Overcoming technical barriers to the sustainable development of competitive marine aquaculture in the United States. U.S. Department of Commerce, Silver Spring, MD USA. NOAA Technical Memo NMFS F/SPO-100. 114pp. Cicin-Sain, B., Bunsick, S.M., DeVoe, R., Eichenberg, T., Ewart, J., Halvorson, H., Knecht, R.W., and Rheault, R. 2004. Development of A Policy Framework for Offshore Marine Aquaculture in the 3-200 Mile U.S. Ocean Zone. Delaware Sea Grant, Newark, DE. Clynick, B.G., McKindsey, C.W., and Archambault, P. 2008. Distribution and productivity of fish and macroinvertebrates in mussel aquaculture sites in the Magdalen Islands (Quebec, Canada). Aquaculture 283:203–210.

443

Cranford, P., Dowd, M., Grant, J., Hargrave, B., and McGladdery, S. 2003. Ecosystem level effects of marine bivalve aquaculture. In Fisheries and Oceans Canada, A Scientific Review of the Potential Environmental Effects of Aquaculture in Aquatic Ecosystems—Vol. 1, Canadian Technical Report of Fisheries and Aquatic Sciences. 2450:iX + 131 p. Crawford, C.M., Macleod, C.K.A., and Mitchell, I.M. 2003a. Effects of shellfish farming on the benthic environment. Aquaculture 224:117– 140. Creswell, R.L., and McNevin, A.A. 2008. Better management practices for bivalve molluscan aquaculture. In: Tucker, C.S., and Hargreaves, J.A. (eds.), Environmental Best Management Practices for Aquaculture. Blackwell Publishing, Ames, IA, pp. 427–486. Dahlback, B., and Gunnarsson, L.A.H. 1981. Sedimentation and sulfate reduction under a mussel culture. Marine Biology 63:269–275. DeAlteris, J.T., Kilpatrick, B.D., and Rheault, R.B. 2004. A comparative evaluation of the habitat value of shellfish aquaculture gear, submerged aquatic vegetation and a non-vegetated seabed. Journal of Shellfish Research 23(3):867–874. Dewey, W. 2005. Army corps of engineering shellfish culture permitting issue. East Coast Shellfish Growers Association. East Coast Shellfish Association, 12 Dec 2005. Retrieved 3 May 2010 from: www.ecsga.org/Pages/Issues/Army_ Corps/Permitting_Update Dewey, W., Bunsick, S., Moyer, J., and Plauché, S. 2007. National trends in shellfish aquaculture production, policy and regulation—planning and regulatory tools for coastal planners. In Proceedings of Coastal Zone 07, Portland Oregon, July 22 to 26, 2007. Duff, J.A., Getchis, T.S., and Hoagland, P. 2003. A review of legal and policy constraints to aquaculture in the US Northeast. Northeast Regional Aquaculture Center, Aquaculture White Paper No. 5, NRAC Publication No. 03-005. Dumbauld, B.R., Ruesink, J.L., and Rumrill, S.S. 2009. The ecological role of bivalve shellfish aquaculture in the estuarine environment: a review with application to oyster and clam culture in West Coast (USA) estuaries. Aquaculture 290:196–223.

444

Shellfish Aquaculture and the Environment

Erbland, P.J., and Ozbay, G. 2008. Comparison of the macrofaunal communities inhabiting a Crassostrea virginica oyster reef and oyster aquaculture gear in Indian River Bay, Delaware. Journal of Shellfish Research 27:757–768. Everett, R., Ruiz, G., and Carlton, J.T. 1995. Effect of oyster mariculture on submerged aquatic vegetation: an experimental test in a Pacific Northwest estuary. Marine Ecology Progress Series 125:205–217. Fernandez, J.L. 1996. Public trust, riparian rights, and aquaculture: a storm brewing in the Ocean State, William & Mary. Environmental Law and Policy Review 20:293–331. Ferreira, J.G., Sequeira, A., Hawkins, A.J.S., Newton, A., Nickell, T.D., Pastres, R., Forte, J., Bodoy, A., and Bricker, S.B. 2009. Analysis of coastal and offshore aquaculture: application of the FARM model to multiple systems and shellfish species. Aquaculture 292:129–138. Figueras, A.J. 1989. Mussel culture in Spain and France. World Aquaculture 20(4):8–17. Fletcher, K.M., and Neyrey, E. 2002. Legal Hurdles to Offshore Aquaculture Leasing. In: Bridger, C.J., and Reid, T.H. (eds.), June 17–20, 2001. St. Andrews, NB, Canada. Open Ocean Aquaculture IV, Symposium Program and Book of Abstracts. Mississippi-Alabama Sea Grant Consortium, Ocean Springs, MS, pp. 45– 46. Fletcher, K.M., and Neyrey, E. 2003. Marine aquaculture zoning: a sustainable approach in the growth of offshore aquaculture. In: Bridger, C.J., and Cost-Pierce, B.A. (eds.), Open Ocean Aquaculture: From Research to Commercial Reality. The World Aquaculture Society, Baton Rouge, LA. Folke, C., Carpenter, S., Walker, B., Scheffer, M., Elmqvist, T., Gunderson, L., and Holling, C.S. 2004. Regime shifts, resilience, and biodiversity in ecosystem management. Annual Review of Ecology, Evolution, and Systematics 35:557– 581. Food and Agricultural Organization of the United Nations (FAO). 2008. The State of the World Fisheries and Aquaculture. FAO Fisheries Department, Rome, Italy. Getchis, T.S., Rose, C.M., Carey, D., Kelly, S., Bellantuono, K., and Francis, P. 2008. A guide to marine aquaculture permitting in Connecticut.

Connecticut Sea Grant College Program. CTSG08-02. 140pp. Grant, J., Hatcher, A., Scott, D.B., Pocklington, P., Schafer, T., and Winters, G.V. 1995. A multidisciplinary approach to evaluating impacts of shellfish aquaculture on benthic communities. Estuaries 18(1A):124–144. Jensen, G., and Zajicek, P.W. 2008. Best management practice programs and initiatives in the United States. In: Tucker, C.S., and Hargreaves, J.A. (eds.), Environmental Best Management Practices for Aquaculture. Blackwell Publishing, Ames, IA, pp. 91–128. Kaiser, M.J., Laing, I., Utting, S.D., and Burnell, G.M. 1998. Environmental impacts of bivalve mariculture. Journal of Shellfish Research 17(1):59–66. Kaspar, H.F., Gillespie, P.A., Boyer, I.C., and MacKenzie, A.L. 1985. Effects of mussel aquaculture on the nitrogen cycle and benthic communities in Kenepuru Sound, Marlborough Sounds, New Zealand. Marine Biology 85:127–136. Langdon, C. 2002. Advances in submerged longline culture of blue mussels Mytilis edulis in the Open Ocean. In: Bridger, C.J., and Reid, T.H. (eds.), Open Ocean Aquaculture IV, Symposium Program and Book of Abstracts. MississippiAlabama Sea Grant Consortium, Ocean Springs, MS, pp. 73–74. June 17–20, 2001. St. Andrews, NB, Canada. Langdon, C. (ed.). 2008. Offshore Aquaculture in the Pacific Northwest ORESU-W-08-001. Oregon Sea Grant, Corvallis, OR. Lindahl, O., Hart, R., Hernroth, B., Kollberg, S., Loo, L.-O., Olrog, L., Rehnstam-Holm, A.S., Svensson, J., Svensson, S., and Syversen, U. 2005. Improving marine water quality by mussel farming: a profitable solution for Swedish society. Ambio 34:131–138. McKindsey, C.W., Landry, T., O’Beirn, F.X., and Davies, I.M. 2007. Bivalve aquaculture and exotic species: a review of ecological considerations and management issues. Journal of Shellfish Research 26(2):281–294. Mittal, A. 2008. Offshore marine aquaculture: multiple administrative and environmental issues need to be addressed in establishing a U.S. regulatory framework (GAO-08-594). U.S. Government Accountability Office, Report to

Development and conservation: Role of resource managers

the Chairman, Committee on Natural Resources, House of Representatives. Washington, DC: Government Accountability Office. National Oceanic and Atmospheric Administration. 2007. NOAA 10-year plan for marine aquaculture. NOAA Aquaculture Program. National Research Council. 1978. Committee on Aquaculture, National. Aquaculture in the United States. National Academy of Science, Washington, DC. National Research Council. 1992. Marine aquaculture: opportunities for growth. Report of the Committee on Assessment of Technology and Opportunities for Marine Aquaculture in the United States, Marine Board, Commission on Engineering and Technical Systems. National Research Council. 2010. Ecosystem concepts for sustainable bivalve mariculture, ocean studies board, committee on best practices for shellfish mariculture and the effects of commercial activities in Drakes Estero, Pt. Reyes National Seashore, California. Newell, R.I.E. 2004. Ecosystem influences of natural and cultured populations of suspension feeding bivalves: a review. Journal of Shellfish Research 23(1):51–61. Newell, R.I.E., and Koch, E.W. 2004. Modeling seagrass density and distribution in response to changes in turbidity stemming from bivalve filtration. Estuaries 27:793–806. Nichols, F.H., Thompson, J.K., and Schemel, L.E. 1990. Remarkable invasion of San Francisco Bay (California, USA) by the asian clam Potamocorbula amurensis, II, Displacement of a former community. Marine Ecology Progress Series 66:95–101. O’Beirn, F.X., Ross, P.G., and Luckenbach, M.W. 2004. Organisms associated with oysters cultured in floating systems in Virginia, USA. Journal of Shellfish Research 23(3):825–829. Peterson, B.J., and Heck, K.L. 2001a. Positive interactions between suspension-feeding bivalves and seagrass—a facultative mutualism. Marine Ecology Progress Series 213:143–155. Peterson, B.J., and Heck, K.L. 2001b. An experimental test of the mechanism by which suspension feeding bivalves elevate seagrass productivity. Marine Ecology Progress Series 218:115–125. Pinnix, W.D., Shaw, T.A., Acker, K.C., and Hetrick, N.J. 2005. Fish communities in eelgrass, oyster

445

culture, and mudflat habitats of North Humboldt Bay, California. Arcata Fish. Technical Report #TR2005-02. 55pp. Powers, M.J., Peterson, C.H., Summerson, H.C., and Powers, S.P. 2007. Macroalgal growth on bivalve aquaculture netting enhances nursery habitat for mobile invertebrates and juvenile fishes. Marine Ecology Progress Series 339:109–122. Reusch, T.B.H., Chapman, A.R.O., and Groger, J.P. 1994. Blue mussels Mytilus edulis do not interfere with eelgrass Zostera marina but fertilize shoot growth through biodeposition. Marine Ecology Progress Series 108:265– 282. Roycroft, D., Kelly, T.C., and Lewis, L.J. 2004. Birds, seals and the suspension culture of mussels in Bantry Bay, a non-seaduck area in Southwest Ireland. Estuarine, Coastal and Shelf Science 61(4):703–712. Rubino, M.C., and Wilson, C.A. 1993. Issues in Aquaculture Regulation. Bluewaters, Inc., Bethesda, MD. Simenstad, C.A., and Fresh, K.L. 1995. Influence of intertidal aquaculture on benthic communities in Pacific Northwest estuaries: scales of disturbance. Estuaries 18(1A):43–70. Souchu, P., Vaquer, A., Collos, Y., Landrein, S., Deslous-Paoli, J.-M., and Bibent, B. 2001. Influence of shellfish farming activities on the biogeochemical composition of the water column in Thau lagoon, France. Marine Ecology Progress Series 218:141–152. Tallis, H.M., Ruesink, J.L., Dumbauld, B., Hacker, S., and Wisehart, L.M. 2009. Oysters and aquaculture practices affect eelgrass density and productivity in a Pacific Northwest Estuary. Journal of Shellfish Research 28(2):251–261. Thom, B. 2009. National marine fisheries service. Endangered Species Act—Section 7 Programmatic Consultation Biological and Conference Opinion Nationwide Permit 48, 28 Apr 2009. Seattle, Washington. U.S. Army Corps of Engineers. 1999. Water resources policies and authorities—digest of water resources policies and authorities. Engineer Pamphlet 1165-2-1, Chapter 21. U.S. Department of Agriculture National Agricultural Statistics Service (USDA NASS). 2006. Census of aquaculture 2005, Volume 3,

446

Shellfish Aquaculture and the Environment

Special Studies Part 2 (AC-02-SP-2). National Agricultural Statistics Service. Washington, DC. Vaudrey, J.M.P., Getchis, T.S., Shaw, K., Markow, J., Britton, R., and Kremer, J.M. 2009. Effects of oyster depuration gear on eelgrass (Zostera marina L.) in a low density aquaculture site in Long Island Sound. Journal of Shellfish Research 28(2):243–250. Ward, L., Grizzle, R., Irish, J., and Langan, R. 2007. Environmental monitoring of an experimental open ocean aquaculture facility in the Western Gulf of Maine, pursuing and advancing technologies for offshore aquaculture in the Western Gulf of Maine in an environmentally responsible manner. University of New Hampshire presented at AFS Symposium,

Environmental Impacts of Coastal Ocean Aquaculture, San Francisco, CA, September 4, 2007. Wechsler, J.F. 2006. Assessing the relationship between the ichthyofauna and oyster mariculture in a shallow coastal embayment, drakes estero, point reyes national seashore. Masters Thesis. University of California, Davis. Welcome, R.L. 1988. International introductions of inland aquatic species. FAO Fisheries Technical Paper 294. FAO Fisheries Department, Rome. World Wildlife Fund. 2010. Bivalve shellfish dialogue. Retrieved 20 May 2009 from: www. worldwildlife.org/what/globalmarkets/aquaculture/dialogues-molluscs.html

Chapter 16

Education Donald Webster

Skills Many aspects of education affect success in shellfish aquaculture. First, there are formal academic disciplines that can be used by practitioners to gain the knowledge required for building and managing shellfish aquaculture operations. There are educational programs that use shellfish aquaculture as a tool to encourage scientific learning and the development of skills. Then there are educational programs that provide outreach and technology transfer to shellfish aquaculturists in offcampus, noncredit programs designed to solve identified needs of producers. Groups involved in industry development, including the United Nations Food and Agriculture Organization (UNFAO), have long recognized educational programs as important in establishing successful and sustainable resource-based industries. Shellfish aquaculture can be divided into several components, each of which incorpo-

rates varying levels of training and technical competence that require different levels of training and education (Pillay 1974):

• • • •

Hatchery Setting Nursery Growout

Hatcheries (see Fig. 16.1) may acquire broodstock from natural populations or develop lines through organized breeding programs. The skills required in operating hatcheries include conditioning of the animals, spawning and larval care, as well as the production of phytoplankton associated with nutrition. In some instances, as with oysters, the hatchery may sell the larvae, while in others the product will be shellfish seed after metamorphosis. In the latter, it would combine both hatchery and setting procedures. Although there are maintenance jobs that use more limited skill sets, the

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 447

448

Shellfish Aquaculture and the Environment

Figure 16.1 Shellfish hatchery worker cares for geoduck clam seed on the Pacific coast of the United States. (Photo credit: D. Webster, University of Maryland.)

hatchery phase tends to require more technically trained workers due to the critical care requirements during the early life stages of the animals. Algology is also an important component of many hatcheries. Skilled algologists must be well trained in establishing and maintaining viable cultures and adept at producing sufficient quantities of suitable phytoplankton for the production volume and timing needed by the hatchery. Often this requires the culture of several species due to the dietary needs of the culture species. Hatcheries that delve into the development of shellfish lines should have expert assistance from geneticists to avoid problems that can lead to “bottlenecking” of genetic material and the resulting loss of beneficial attributes in the lines.

The setting phase includes care of animals through metamorphosis. Oysters may be exposed to cultch during this process for them to attach to. Clams are provided protection until they settle and begin to develop shells. Mussels may be gathered from the introduction of suspended cultch and the resultant capture of wild seed from natural spawning areas (see Fig. 16.2). In oysters, the setting process is normally accomplished by introducing larvae to shell or shell fragments, although chemical methods for inducing metamorphosis have been used to produce spat that have no cultch attachment. The animals proceed through metamorphosis and begin growing. The setting phase usually involves material handling and makes use of less highly skilled labor than that required in hatcheries. In commercial production, worker skills include training to properly prepare cultch by ensuring that it is clean and washed. Workers must also know how to install and maintain the setting units and understand physical factors such as temperature and salinity that need to be maintained for the juvenile animals to survive. A nursery phase may or may not be included, depending on the species and its intended use. If included, the small, newly metamorphosed animals will be cared for until they reach a size deemed necessary for planting, while in others they may be deployed immediately, bypassing the nursery phase. There is usually a trade-off between cost and ultimate survival of the animals in this decision since it is widely regarded that the deployment of larger animals generally results in higher final survival. Nursery operations may be carried out in open water or in containment devices, such as upwellers or downwellers. These devices are designed to protect small animals while providing them with ample food supplies for enhanced growth. Nursery operators require skills that include the handling of small shellfish and an understanding of the periodic

Education 449

Figure 16.2 Field crew placing mussels in socking material on rafts in Maine where they will be grown. (Photo credit: D. Webster, University of Maryland.)

cleaning necessary to maximize their growth and survival. Growout is the ultimate goal of commercial operations. Animals are introduced into the growing areas using the method chosen by the producer. This may entail deployment on bottom, which is a traditional technique for species such as oysters or clams, or the animals may be protected in a variety of trays, cages, or other enclosures that are designed to protect them from predators and aggregate them for ultimate retrieval (see Fig. 16.3). Shellfish aquaculture can also be subdivided by production motivation. Commercial aquaculture has the goal of producing animals for sale, generally for consumption, although some sales exist for aquaria, test animals, or other purposes. Restoration aquaculture is used for introducing animals into depleted populations for the purpose of environmental enhancement or harvest. These harvests may be for either recreational or commercial purposes.

While many commercial shellfish producers have become successful without formal academic training in shellfish-related disciplines, a key factor that helps ensure success is an appreciation for science and the ability to acquire skills that can aid in identifying problems and developing solutions. Knowledge of the scientific method is therefore very useful.

Aquaculture-related disciplines Many disciplines are used in the overall conduct of shellfish aquaculture. From designing hatcheries and production equipment to solving growout problems, conducting successful businesses, and carrying out scientific investigations, there are useful areas of study that can aid the ultimate success of prospective culturists (Cole and Hall 1973). Among these are

450

Shellfish Aquaculture and the Environment

Figure 16.3 Cages are used to help protect oysters from predators such as crabs and rays while they are growing. (Photo credit: D. Webster, University of Maryland.)

• Biological sciences (including microbiology, ecology, and related subjects) • Chemistry (including organic chemistry) • Environmental science • Engineering (including mechanical, civil, agricultural, ocean, and others) • Business administration (management, finance, marketing, and related subjects) While it is apparent that biological sciences would be favored due to the need to understand the life cycle of the animals being cultured, other disciplines can be highly useful. A well-grounded understanding of the culture requirements of the animals being raised is critical for success. Chemistry and organic chemistry are physical sciences that are beneficial for understanding the nature of reactions that occur within the environment. An understanding of the nature of salinity, acid and base relationships, and ionization as they relate to water quality and the impact that these have on the cultured organisms is important in many aspects of aquaculture.

Engineering subdivisions include civil, mechanical, and electrical, each of which have applications in aquaculture. The specialized field of agricultural engineering includes training in many of the other areas. Agricultural engineers are largely concerned with applying their skills in solving problems dealing with the production of food and fiber. Therefore, it makes them well suited for shellfish aquaculture. Likewise, ocean engineers are trained to design systems that can withstand the stresses that occur in the dynamic marine environment, which makes them particularly useful when developing systems in open waters. Engineering contributes greatly in areas such as hatchery design and in developing production equipment with the ability to withstand stresses in severe environments or when problems of material handling arise in large-scale operations (see Fig. 16.4). Since aquaculture involves so many important areas of knowledge, it is also useful for teaching a broad range of audiences. Involvement at an early age can often stimulate students to follow it as a vocation.

Education 451

Figure 16.4 Floating upwellers are engineered with a rotating paddle wheel to move large quantities of water for seed oysters. (Photo credit: D. Webster, University of Maryland.)

K-12 education Shellfish aquaculture has the ability to attract young people, who often develop an interest in the animals as they learn about them. Encouraging young people to become interested in shellfish aquaculture at an early age can lead to starting them on career paths while providing them with useful knowledge about the role of shellfish in the environment. That makes shellfish and the culture of them particularly attractive for use in educational programs. Aquaculture education programs have been developed in many areas and are used to teach a range of subjects such as biology, chemistry, mathematics, and marketing. In the early years during primary grades, students may learn about the various ways in which shellfish are raised. If they are fortunate to live near the coast, they may be able to participate in a field trip to a hatchery or production site. Many producers understand the benefits of having young people learn about their businesses and are quick to welcome

them to their operations. For those not near the coast, shellfish have been used in aquaria to demonstrate their ability to clear water by biofiltration. In these early years, they are normally used in conjunction with basic science curricula. Middle grades often find students learning about the life cycle of the animals and being introduced to science labs. Here, they may participate in dissecting shellfish to learn about their digestive, respiratory, circulatory, and reproductive systems. They may also be introduced to environmental science that teaches them about the relationship between species, as well as their places and functions within ecosystems. Educational resources are available to teachers interested in using aquaculture in the classroom, with many of these now being accessible through the Internet. High school programs (see Fig. 16.5) include more advanced studies where students learn scientific method by investigating existing literature, developing hypotheses, designing experiments to test them, analyzing results,

452

Shellfish Aquaculture and the Environment

Figure 16.5 Dr. Joseph Buttner works with high school youths to use shellfish aquaculture as a teaching tool. (Photo credit: J. Buttner, Salem State University.)

and reaching conclusions. In this manner, shellfish aquaculture can be used to teach many skills. Mathematics can aid students in calculating production and determining efficiency of shellfish setting, as well as determining flow rates and comparing diets. The study of commensal organisms can lead to a better understanding of the relationships that shellfish have with other creatures in the environment. Engineering studies aid students in designing culture systems and determining how to solve production problems. While many school programs integrate aquaculture into traditional courses, there are formal programs that use aquaculture as a focus for teaching (http://bigquil.blogspot. com). One example is the Sound School in Connecticut (USA) (www.soundschool.com), where students design, build, and maintain culture systems and develop research projects. Some examples have led students to monitor oysters in nearby areas to determine parameters affecting growth and survival. Building

and maintaining small-scale hatcheries provides an excellent way to combine engineering with biology while learning production techniques. The products from these experiments are sometimes used for stocking natural areas, which leads to an appreciation of the use of aquaculture for restoration.

Undergraduate degree programs At the university level, degree programs in aquaculture, often including shellfish, have become established in many countries (University of Hawaii, Manoa n.d.). Institutions in Canada, Australia, New Zealand, Scotland, and the United States offer programs of this type. Some institutions have developed 2-year programs that are typically more vocational in nature and provide significant hands-on training to students. These may grant Associate of Science (AS) degrees or offer certificates to graduates.

Education 453

Two-year programs are often created in areas where aquaculture industries exist and are frequently meant to produce graduates with the skills needed by local businesses. This relationship provides a ready source of trained technicians for companies while offering attractive employment potential to program graduates. However, to be genuinely beneficial for lifelong applicability, these programs should offer training that provides students with the skills needed to earn a living across a spectrum of aquaculture industries rather than just provide training in local production techniques or systems. Studies of vocational programs have found that students gain up to a 5-year knowledge advantage by attending these programs compared with those who gain their training by directly entering employment and learning while on the job. Institutions with two-year programs may also offer four year courses of study leading to the Bachelor of Science (BS) degree. Those who complete the 2-year program may be allowed to transfer most, if not all, of the credits earned into the 4-year programs. This is beneficial for students who are interested in studying aquaculture but (1) may not be able to commit the time needed to a 4-year program, (2) may initially be undecided about how far they wish to pursue a formal degree program, or (3) are financially unable to afford more than 2 years of initial training. In this case, it may be possible for the student to apply for the 4-year program at a later date after gaining employment and working in the industry, where they would find their employment experiences valuable for background. Courses included in BS degree program will likely include basic sciences, communications, and economics at the core, while adding a range of electives tailored to the course of study and interests of the student. Labs connected with many of the courses are designed to teach skills that will be required by students for gaining employment in industry.

Graduate degree programs Many students who wish to become proficient in shellfish aquaculture pursue training through the graduate school level. Graduate programs offer students the opportunity for advanced study with professors who have experience and reputation in the field. A key factor in graduate programs is that the student will be taught to think critically and further develop skills in the scientific method. In applying for a program leading to a graduate degree, the student should consider the reputation of the institution and the major professor with whom the student seeks to become associated. Choosing an educational institution and a major professor cannot be taken lightly or decided upon without significant investigation. Once accepted, the professor will assist the student in assembling a graduate committee, choosing a course of study, and developing a research topic while mentoring the student during the course of his or her graduate study. There are institutions offering graduate degrees in aquaculture or aquaculture-related studies throughout the world. These include Master of Science (MS) or Doctor of Philosophy (PhD) programs, depending on the institution. Many of these are connected with colleges of biological or life sciences. A few engineering programs also have aquaculture options. Master of Business Administration (MBA) programs can be useful to those interested in the business aspect of aquaculture. It has been noted many times that more aquaculture businesses fail because of poor business management decisions than because of technical problems. Advanced or graduate degrees may open up more potential pathways for employment since they are required for many jobs in contemporary society, especially in developed nations. Positions in academic institutions for research faculty now generally require the PhD

454

Shellfish Aquaculture and the Environment

degree, while the MS is considered as minimum for associate staff and extension faculty. This is also true for many government positions, although an undergraduate degree may be accepted by many more state and local agencies. In many instances, an undergraduate degree may be the minimum necessary to begin employment with agencies; successful candidates, however, often see the benefit of pursuing advanced degrees throughout their careers. This allows them to open additional pathways for advancement or pursue opportunities for other positions. In addition to formal classroom training and degree-granting programs, shellfish aquaculture has other options that can provide learning experiences in off-campus and noncredit environments. These programs can be extremely important to the long-term development of the industry and have the ability to provide continuing support to producers.

4-H and youth programs In order to change the future, it has been recognized as advantageous to train future generations. Youth programs provide a way to change attitudes and instill values. Youth vocational programs developed for agriculture over the past century have been particularly successful in developing that industry. These programs provide a useful guide for teaching shellfish aquaculture skills and techniques to new generations of young people. One of the most successful is the 4-H program that has been operated by the U. S. Department of Agriculture. It became a principal cause for the application of modern agricultural techniques and is a major component of extension programs in the United States, where 4-H exists with the other major program areas of Agricultural Science and Family and Consumer Science. Today, 4-H includes over 6 million young people who are organized to

learn leadership, citizenship, and life skills through local clubs and activities (National 4-H Council 2010). 4-H principally relies on clubs led by local leaders, but with a strong national support network. The 4-H community has over half a million volunteers and has been a key part of agricultural leadership and skill development for decades. Learning activities of clubs are supported by the latest research from Land Grant universities. These are focused in (1) science and technology, (2) healthy living, and (3) citizenship. This tie with Extension provides programs in more than 3000 counties within the United States through local offices. 4-H programs have traditionally involved farm or rural youth but have been expanded to new audiences. One of these is shellfish aquaculture. Some local projects across the nation have used shellfish topics with the most focused program currently based in Washington State. The Jefferson County 4-H club known as “Big Quil Enterprises” provides students with training to manage a youth-operated oyster business on the Hood Canal, a wellknown shellfish production area (http:// bigquil.blogspot.com). The club has been supported by a local oyster company, which buys the product from the club for processing and sale (Big Quil Enterprises 2006). Typical of 4-H programs, the club includes about 50 students who learn the skills needed to produce oysters while acquiring an ability to manage the operation as a business. As in any business, the goal of the club is to make a profit and become self-sustaining. While most of their production is sold to a local shellfish company, club members participate in festivals throughout the state where they distribute information about their project while selling shellfish. This provides them with the opportunity to educate others about the benefits of shellfish aquaculture. 4-H can provide a suitable model for other regions seeking to introduce both the skills and attitude necessary to make a living in

Education 455

Figure 16.6 Florida fishermen became successful clam farmers through the application of research and extension programs. (Photo credit: D. Webster, University of Maryland.)

shellfish aquaculture. While other programs exist that provide youth with skills, the ties to university research gives 4-H a unique ability to provide training that can translate into future industry expansion using state-of-theart technology. Related 4-H projects in areas such as leadership development and public speaking aids in developing articulate industry spokespersons who can become politically involved to help bring about positive change (http://4-H.org).

Extension programs Extension education has many important uses in establishing, supporting, and developing shellfish aquaculture (see Fig. 16.6). Since many who decide to go into business may not have had formal training, there is a need to provide educational services to teach skills needed for success. Extension programs are often used for this purpose. These efforts extend or apply research done by universities at campuses and experiment or field stations

to production sites. Extension specialists are usually university-trained graduates in a biological science (Kensler 1989). Extension education consists of off-campus, noncredit programs that provide researchbased solutions to industry problems (Severs et al. 1997). This has been recognized as an essential element in developing agriculture, fisheries, and aquaculture by agencies involved in both national and international development. While many terms are used for extension work around the world, one of the most descriptive is the Dutch “Voorlichting,” which means “lighting the path.” Essentially, extension workers provide guidance that helps people identify and solve problems. In the United States, this mission has been part of the Land Grant and Sea Grant college programs. The Land Grant system has roots that extend back to 1862, while the Sea Grant system that was envisioned to provide similar services in the marine environment was established in 1966. Land Grant includes academic institutions in all U.S. states, while Sea Grant colleges are found in coastal and Great Lakes

456

Shellfish Aquaculture and the Environment

states. Both include ties to island commonwealths, republics, and territories. The system provides linkage between research, education, and extension. In the operational model, research conducted to enhance production and profitability is disseminated to producers through extension programs, while problems identified at the field level are transmitted to researchers for future investigation. The body of knowledge that results is included in courses for the education programs that train the next generation of producers and scientists in campus educational programs. Land Grant colleges were established to research problems in agriculture production while providing campus-based education for future farmers and scientists. It was recognized that a great deal of agricultural research needed to be transferred from academia to farm fields for successful implementation. The system that was established provided funds for offices at the county level staffed with extension agents who worked directly with farmers to apply research for solving defined problems. These offices were jointly funded by federal, state, and local governments (van den Ban and Hawkins 1985). Internationally, the UNFAO has long recognized the importance of extension programs to the success of development projects. While interest in extension programs waned during the 1990s, they have recently increased and are again recognized as important in international development projects (Marine Technical Assistance Group 1982). UNFAO models have varied over the years. The Training and Visitation (T&V) system was used for many years. It was based on extension agents making specific recommendations to farmers about practices that were determined should be adopted by them. As such, it was top-down and paternalistic in nature, and did not always result in long-term success. It has been replaced by systems that create empowerment and experiential learning

where farmers are urged to make decisions on their own. Farmer-to-farmer exchanges are also favored as a means of spreading knowledge. This new model is known as Participatory Technology Development (PTD). Extension is often misunderstood by those who think of it only as providing fact sheets or single programs to potential growers in response to specific questions. In fact, a successful extension program includes a series of educational events targeting specific problems and must involve the audience in the process of development and implementation. A well-designed program will work with aquatic farmers to help them identify their problem. It will then help with providing alternatives for solution while aiding them in choosing the most practical path for implementation. The final phase is to assist in evaluating results and deciding whether to remain with that solution or choose another. This evaluation and assessment process is critical for success. The participatory system has been recognized as more effective for continued success than top-down models that provide farmers with only a single course of action. In essence, extension educators act as mentors, similar to professors who teach students to think and act as scientists. Extension programs are designed in several phases. The planning phase identifies goals, often by conducting a formal “needs assessment.” Here also is where project priorities and objectives are determined and the target audience identified. The design and implementation phase includes selecting and developing program content, delivery methods, and resource materials and determining the project timeline. At the conclusion, the evaluation phase measures the success of the program and judges its impacts upon the participants as well as the other factors it was intended to influence. In creating programs, it is vitally important that the needs of the community and the society being targeted are taken into account.

Education 457

Factors that are considered important include the social, historical, economic, educational, and political. In some instances, especially in rural areas or developing nations, there may be audience members who are illiterate. A determination will need to be made when creating educational materials and methods that will allow them to learn important concepts. In these instances, pictorial designs for printed material could be used to illustrate concepts. Modern technology such as the use of portable computers has allowed the use of audio and visual training aids to be used as well. Political factors are frequently important since they may include laws and regulations that would prevent the use or implementation of new or creative production techniques that might otherwise prove beneficial. Extension programs are planned, implemented, and evaluated in a seven-step process known as “Bennett’s hierarchy.” This involves delineating 1. Inputs (time, resources) 2. Activities (content and methods) 3. People (numbers, characteristics, contact frequency) 4. Reactions (interest, satisfaction levels) 5. KASA change(knowledge, attitudes, skills, aspirations) 6. Practice (adoption and application of knowledge) 7. Results (economic, environmental, and social actions) In order to properly judge success in reaching goals, the evaluation of results is critical and should be determined in the three areas of social, economic, and environmental change (Sea Grant Editorial Board 2000). Properly evaluating results is not an easy task. Many people make the mistake of stopping after only a few steps and reporting results based on number of attendees at a program or on the dissemination of written materials. In creating new extension organizations, especially in developing nations, it is impor-

tant that extension educators not be given tasks such as enforcing regulations or collecting fees or taxes. They must have the trust of the farmers they are working with in order to build relationships that will bring support to the extension programs. Governments frequently invest in extension since they see it as a way to aid national goals of increasing food production and creating economic growth, especially among poorer areas. They often recognize the ability of these programs to increase the welfare of rural people and to promote sustainable production practices. Shellfish are well suited to sustainability and are regarded as “green” businesses in many nations. In some cases aquaculture extension activities may be combined within larger agricultural units because of their better equipment and larger numbers (FAO Technical Guidelines 1997). Extension education is an accepted and well-documented way of aiding shellfish farmers in their businesses. In many cases, educators also have appointments as research faculty. These joint appointments can be beneficial in establishing important links between those who identify problems and the research that helps find solutions. However, it must also be stated that joint appointments create multiple responsibilities for those who have them, which may affect the overall efficiency or performance of the program. Joint appointments often mean that the person will be evaluated by two different groups or with different criteria for the research and extension portions of their job. This may lead to difficulty if those involved in the evaluation process lack understanding about the differences.

Technology transfer Another route for disseminating information is through technology transfer. This system seeks to share knowledge, skills, methods, equipment, and processes that have already

458

Shellfish Aquaculture and the Environment

Figure 16.7 Computerized information system guiding towed dredge samples to assess bottom for shellfish culture suitability. (Photo credit: D. Webster, University of Maryland.)

been developed, often for other industries. The application of these is made accessible to those who it is believed can best integrate them into their production processes. Government agencies in some countries use technology transfer to provide new ideas for production methods and equipment to industry. This is an accepted practice in many nations where a designated agency will keep abreast of new technology, evaluating it for application to industry needs, and providing the resulting information to appropriate businesses for that application. Agencies may also invest in projects that are designed to help purchase, install, and evaluate technology under field conditions. If the application is successful, they will publicize it to others in the industry, trying to gain as much acceptance as possible. It is not unusual to find an Office of Technology Transfer in agencies, companies, or institutions that are tasked with finding applications for the new methods and equipment that these bodies have developed. Since many academic institutions rely on patents or the licensing of technology developed by their

faculty for income, they may be constantly on the watch for areas in which their technology could be applied. Perhaps the most widespread use of technology transfer has been in the commercial application of devices developed for the military. The use of radar, sonar, and position determining equipment such as the global positioning system (GPS) is now widespread in aquaculture as well as many other industries (see Fig. 16.7). The use of side-scan sonar and subbottom profiling has been applied in shellfish aquaculture to map beds that were previously carried out by laborious physical methods and with time-consuming manual chart plotting. The development of computers and creation of the World Wide Web have brought information management within the reach of all but the most isolated areas.

Conclusion Educational needs within the shellfish aquaculture industry are many and varied. However,

Education 459

they are important for the growth of it and certainly cannot succeed without it. The growth of computer and communication technology has made it possible for information to be obtained throughout the world in a short time. It can link learners with educators across continents and provide televised demonstrations of equipment and methods that would otherwise have take days to see. Perhaps one of the most important aspects has been the ability to magnify the effort of teachers and provide information to broader audiences than were previously able to be reached. This has also led to dissemination of results in a timely manner to those who have the ability to use them. It is now possible to link educators in several nations around the world with students, essentially bringing the world into the classroom. Computer-based information sources such as listservs and social networks have made it possible to share ideas with people of similar interest in far-flung areas. Websites serve as information hubs and can include text, video, voice-over presentations, and interactive technology. The information that used to take days or weeks to get across a town now can travel around the world in a matter of seconds. The structure that combines research and education with extension services has been effective in getting many shellfish aquaculture businesses established. Continuing to provide this support to those in production is critical for the overall success of shellfish aquaculture.

Literature cited Big Quil Enterprises. 2006. http://bigquil.blogspot. com/search/label/Big%20Quil%20Enterprises. Accessed June 23, 2011.

Cole, R.C., and Hall, D.N.F. 1973. Guide to fishery education and training. FAO Fisheries Technical Paper, Rome. FAO Technical Guidelines for Responsible Fisheries Aquaculture Development-5. 1997. Food and Agriculture Organization of the United Nations, Rome, Italy. Kensler, C. 1989. A Regional Survey of the Aquaculture Sector in North America (Including Canada, Greenland and the United States of America). United Nations Development Programme, FAO, Rome, Italy. Marine Technical Assistance Group. 1982. An Evaluation of Fishery and Aquaculture Programs of the Agency for International Development. National Academy Press, Washington, D.C. National 4-H Council. 2010. About 4-H: Who We Are. http://www.4-h.org/about/youthdevelopment-organization/. Accessed June 23, 2011. Pillay, T.V.R. 1974. Planning of Aquaculture Development—an Introductory Guide. Fishing News Books, Surrey, England. Sea Grant Editorial Board B. Wilkins, Leader Emeritus. 2000. Fundamentals of a Sea Grant Extension Program. Cornell University, Ithaca, NY. Severs, B., Graham, D., Gamon, J., and Conklin, N. 1997. Education through Cooperative Extension. Delmar Publishers, Albany NY. University of Hawaii, Manoa. Education Opportunities—University Degree Programs. (http://praise.manoa.hawaii.edu/ed.php?univ). van den Ban, A.W., and Hawkins, H.S. 1985. Agricultural Extension, 2nd ed. Blackwell Science, Cambridge, MA.

Chapter 17

The implications of global climate change for molluscan aquaculture Edward H. Allison, Marie-Caroline Badjeck, and Kathrin Meinhold

Introduction The overwhelming majority of the world’s climate scientists agree that human activity is changing the global climate (Anderegg et al. 2010). Rising CO2 and other greenhouse gas emissions from industry, domestic use and agriculture, and the degradation of carbon sinks through deforestation and other land use and coastal ecosystem conversions are together leading to rapid warming of the earth’s atmosphere (IPCC 2007a). Public awareness of the changing climate is increasing, although people in industrialized countries remain divided on its cause and distracted by more immediate concerns (Lorenzoni and Pidgeon 2006; Dunlap and McCright 2008). Commitment by

individuals, civil society, businesses, and states to reduce emissions and invest in adapting to change is diffuse or limited, hence the continuing difficulties in reaching a global accord on emissions targets (Lazarus 2009). While humanity debates the issues and how to deal with them, global warming is leading to changes in the basic biophysical processes that determine the structure and function of the oceans (Harley et al. 2006). This, in turn, has impacts on the coastal and marine ecosystems within which molluscan shellfish are farmed. Climate change should therefore be an issue of concern to shellfish farmers, processors, and traders, as well as to researchers and regulators and to all of us who enjoy eating shellfish. All these activities will be affected by

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 461

462

Shellfish Aquaculture and the Environment

the changing climate, and some are already being affected. In this chapter, we review the relevant elements of climate change and their likely impact on shellfish farming operations and value chains, and we propose measures to respond to them, so that we may retain productive industries and access to a highly valued source of food. Molluscs are cultured in both temperate and tropical environments, in marine and brackish water, in a variety of farming systems, ranging from extensive seabed cultivation of oysters and clams, through semi-intensive line and raft cultivation of mussels and scallops, through to intensive recirculation systems for abalone (Chapter 1 in this book). They also contribute to economy and society in various ways—from providing low-input food production systems for small-scale farmers and fisherfolk, by being part of a diversified coastal economy sustaining small communities, to being major commercial concerns (Chapter 2 in this book). As we will show, even the effects of a single climate change-related variable— ocean acidification due to increasing concentration of CO2 in the atmosphere—cannot be accurately predicted. When we consider the multiple pathways through which climate change (and societal responses to it) will take place, prediction becomes difficult on the basis of current knowledge. In this chapter, rather than attempting to make ill-founded model predictions of future harvests, and their distribution and value, we limit ourselves to identifying the pathways of potential impact of climate change on molluscan shellfish aquaculture, reviewing the evidence for impact so far, and giving some examples of the nature and scale of predicted impacts from recent case studies. We also review possible adaptive responses, mindful that investment in such adaptation will have to contend with continuing uncertainty about the nature and extent of impacts. Finally, we consider how shellfish farms can become part of the solution to a

lower carbon future for our societies, by exploring their role in producing food that has a small “carbon footprint” (Bunting and Pretty 2007; Hickey 2008), and by evaluating the proposition that shellfish aquaculture, through the production of calcium carbonate shells, can act as a “carbon sink.”

Climate change in the oceans and coastal zones As recently reviewed by the Intergovernmental Panel on Climate Change (IPCC) (IPCC 2007a), global warming leads to changes in physical processes in aquatic systems, such as alternations in the major ocean currents and in the local circulation patterns in coastal systems; changes in the frequency and severity of the El Niño Southern Oscillation (ENSO; driven by the Humboldt Current in the Pacific); and thermal expansion and addition of ice melt to the oceans, leading to sea level rise (Fig. 17.1). In freshwater, rainfall and riverflow patterns and rates of evapotranspiration are changing. Storms, floods, and droughts are changing in frequency and intensity. Chemical changes brought about by rising levels of atmospheric CO2 are also leading to an acidification of the oceans, with consequences for ocean food chains and for coral reefs. Drawing on recent reviews by the IPCC (2007a) and by Barange and Perry (2009), supplemented by reference to other specific studies, the main chemical and physical elements of global change affecting the oceans can be summarized as follows.

Heat content and temperature The oceans are warming, but there are geographical differences in the extent of that warming, and there continue to be decadal

Implications of global climate change

463

Increased greenhouse gas concentrations

Human activities

Increased UV Increased air temperature

Intensified atmospheric pressure gradients

Increased storm frequency

Sea level rise

Intensified upwelling (?)

Increased CO2

Increased water temperature

Decreased pH

Figure 17.1 The main physical changes in the oceans attributable to anthropogenic global warming. (Reproduced from Harley et al. 2006.)

cycles which can mean short-term cooling in some areas, despite the overall warming trend. Warming is not exclusive to surface waters, with deep warming being seen particularly clearly in the Atlantic Ocean. While coastal large marine ecosystems (LMEs) on the West Coast of the Americas have shown slight cooling, most other areas have warmed, with enclosed shelf seas (e.g., Mediterranean, Black Sea) showing particularly large increases. Increases in temperature (and salinity; see below) from changes in precipitation, evaporation, river runoff, and ice melt are likely to lead to increased vertical stratification and water column stability in oceans and lakes, reducing nutrient availability to the euphotic zone and thus primary and secondary production. Ecosystem productivity is likely to be reduced in most tropical and subtropical oceans, seas, and lakes, and increased in high latitudes.

Ocean salinity, density, and stratification Overall indications are that, globally, oceans are becoming less saline due to greater precipitation, higher runoff, ice melting, and advection, but that there are large regional differences. Salinity is increasing in the surface of the subtropical North Atlantic Ocean (15– 42°N), while further north and in the Southern Ocean waters are becoming less saline. Lower salinity is also reported in the Pacific, except in the upper 300 m and in the subtropical gyre, where salinity is increasing, and the Indian Ocean is generally showing increases in salinity in the upper layers. Salinity affects species distribution, and the growth and reproductive success of individual shellfish species, and will therefore affect the composition and productivity of both natural and farmed shellfish populations.

464

Shellfish Aquaculture and the Environment

Ocean circulation and coastal upwelling Observed and predicted changes in the ocean’s heat content and salinity are and will continue to affect circulation patterns. These are expected to decline, but at present there is conflicting and unclear evidence as to whether climate change will increase or decrease coastal upwelling. This is important in terms of its implications for biological production in the major fisheries and shellfish farms of upwelling areas.

Sea level rise Global average sea level has been rising at an average rate of 1.8 mm per year since 1961. The rate has accelerated since 1993 to about 3.1 mm per year. Higher rates in coming decades are likely, and recent research (e.g., Rahmstorf et al. 2007) indicates that they may be higher than even the most pessimistic (IPCC 2007a) scenario. Sea level change is not geographically uniform, however, because it is controlled by regional ocean circulation processes. The largest losses of land and impact on populations and economies are likely to be in East and Southeast Asia, although proportional impacts may be even higher on low-lying small-island developing states in the Pacific and Indian Oceans in particular. Rises in sea levels may affect coastal bays, lagoons, and wetlands, which provide sites for shellfish cultivation, and coastal onshore infrastructure used by fisherfolk (e.g., housing, harbors).

Land–ocean exchanges Land use change contributing to and resulting from climate change, particularly deforestation and hydrological modifications, has downstream impacts, especially in terms of erosion in catchment areas and increased sus-

pended sediment loads. In contrast, damming and channelization have greatly reduced the supply of sediments to the coast from other rivers through retention of sediment by dams (many of which are themselves now being built as adaptations to climate change—e.g., to increase water storage, regulate floods, or reduce emissions from fossil fuel-based power generation). Levels of sedimentation have the potential to have significant impacts on shellfish farming through changes in turbidity, salinity, stratification, and nutrient availability, all of which affect estuarine and coastal ecosystems and the productivity of the phytoplankton community.

Changes in low-frequency climate variability patterns Natural climate variability takes place through intermittent 1–2-year duration events (e.g., ENSO) and intrinsic variability operating at decadal and longer timescales. ENSO events are associated with many atmospheric and oceanic patterns, including abnormal patterns of rainfall over the tropics, Australia, southern Africa and India, and parts of the Americas, easterly winds across the entire tropical Pacific, air pressure patterns throughout the tropics, and sea surface temperatures. Coincident ecological changes are both vast and global through their impacts in coastal upwelling systems: They increase coastal temperatures, reduce plankton production by lowering the thermocline (which inhibits upwelling of nutrients), and change trophodynamic relationships; they also have knock-on effects on the hydrological cycle and therefore on agriculture and even on forest fire severity, as in Indonesia following the 1997 ENSO event. Examples of aquaculture impacts of El Niño are given in the next section. Some studies expect stronger and more frequent ENSO events as a result of global warming, while others suggest that the evidence is still lacking because ENSO is not well-

Implications of global climate change

enough simulated in climate models to have full confidence in these projected changes (Cane 2005). Other naturally occurring climatic variability being impacted by climate change relates to atmospheric associations or teleconnections; changes in the position and intensity of atmospheric convection in one area result in adjustments in pressure cells in adjacent areas and can lead to altered wind and ocean current patterns on a global scale. Understanding these connections and how climate change will affect them is a research frontier for climate scientists.

Increased frequency and severity of extreme climate events Most major tropical storms are generated around latitudes 8° and 35° north and south of the equator. There are ongoing scientific controversies around the oft-stated perception that storm frequency and severity are increasing due to climate change, and that there may be changes in seasonality and typical track of storm events. An emerging consensus around recent trends and predictions for hurricane energy (a measure of storm severity) indicate no evidence for increased frequency of storms, but strong evidence for a trend for increasing severity and therefore destructiveness of storms over the last 20–40 years (i.e., exceeding the periodicity of decadal cycles in the ocean climate system and attributable to anthropogenic warming); this trend is particularly evident in the western Atlantic (Emanuel et al. 2008).

Ocean acidification and changes in chemical properties Continued uptake of atmospheric CO2 by the oceans has decreased the pH of surface seawater by 0.1 units in the last 200 years. Model estimates predict further reduction of 0.3 to 0.5 pH units over the next 100 years. It is

465

expected that pH reduction will change the depth below which calcium carbonate dissolves, increasing the volume of ocean that is undersaturated with respect to aragonite and calcite, which are used by marine organisms to build their shells and by corals to build their skeletons. The impacts of these changes will be greater for some regions and ecosystems and may be most severe for shell-bearing organisms, tropical coral reefs, and cold-water corals in the Southern Ocean. Ocean acidification has been portrayed in apocalyptic terms. The conclusions of two recent reviews (Doney et al. 2009; Hendriks et al. 2010) are more measured but still point to the need for urgent action on curbing emissions to prevent rapid change. The direct implications for shellfish have galvanized much recent research, which is reviewed in the next section. Impacts of the above physical and chemical changes in coastal and oceanic waters have potentially profound ecological impacts. Drawing on Barange and Perry (2009) and Brierley and Kingsford (2009), these impacts include as follows.

Timing and success of physiological, spawning, and recruitment processes All living organisms have specific ranges of environmental conditions to which they are adapted and within which they perform optimally. Physiological performance may degrade and cause stress at temperatures outside the normal limits. Likely impacts of climate change include changes in timing and success of fertilization, survival and growth of shellfish, timing and extent of migrations, and structure of food webs.

Primary production Satellite observations suggest a 6% reduction in global oceanic primary production between

466

Shellfish Aquaculture and the Environment

the early 1980s and late 1990s, but with substantial regional differences. The climate– plankton link in the ocean is found most strongly in the tropics and mid latitudes, where the typically low levels of surface nutrients limit phytoplankton growth. Climate warming further inhibits mixing, reducing the upward nutrient supply and lowering productivity (Doney 2006). Reduced primary production is likely to mean reduced fish and shellfish production in many aquatic ecosystems, although the relationship may not be straightforward as much productivity is derived from the “microbial loop” and dependent on influx and cycling of organic material from terrestrial areas for continental shelf seas.

larval survival rates, disease immunity, etc.) are not negatively affected by climate change. Thus, it cannot be expected that warming will bring more and faster-growing shellfish.

Phenomenological changes More than half of all terrestrial, freshwater, or marine species studies have exhibited measurable changes in their phonologies (timing of life cycle events) over the past 20–140 years, in line with climate changes. In the oceans, this includes changes in the timing and extent of seasonal phytoplankton blooms which have not been tracked by zooplankton, suggesting an emerging mismatch between food supply and predator life cycles.

Changes in distributions Climate change is expected to drive most terrestrial and marine species ranges toward the poles, expanding the range of warmer-water species and contracting that of colder-water species. Such changes have already been documented in the North Sea and North Atlantic for finfish. A recent study modeling “climate envelopes” based on existing species distributions of both fish and exploited invertebrates suggests that there will be substantive changes in distribution of fish and invertebrate communities, and large numbers of extinctions of species with restricted habitat ranges (Cheung et al. 2009).

Species invasions and diseases Pathogens are spreading to higher latitudes and harmful algal blooms (HABs) are becoming more common (although these may also be due to localized nutrient enrichment). Ecosystems may become more liable to invasive species as they change conditions from those that may have prevented warmer water species from breeding, for example. Thermally stressed shellfish in aquaculture systems may become more liable to viral, fungal, and bacterial infections.

Regime shifts and extreme events Abundance changes At the simplest level, biological production processes occur faster at higher temperatures. However, increased growth rates of shellfish (and hence increased biomass and production) will only occur in response to higher temperatures when food supply is adequate to these increased demands, and when other life cycle processes (spawning migrations, fertilization,

Gradual and variable climate changes can provoke sudden and perhaps irreversible biological responses as ecosystems shift from one state to another (de Young et al. 2008). The altered state may provide an alternative fishery, or it may simply be less productive. Examples of regime shifts are the alternations between sardine and anchovy-dominated pelagic fisheries in some upwelling areas, and arguably, the

Implications of global climate change

collapse of the Newfoundland cod fishery signaled a regime shift driven by the combined stressors of overfishing and climate variability. The links between global warming, these physical and chemical changes in hydrology and oceanography, and the ecological changes in aquatic food webs are thus complex, and when we then add the operations and economics of the shellfish farming sector, the complexity increases further. In order to identify issues that can be evaluated and acted upon in the context of mollusc culture, the next section is structured around a simplified schema that draws on the mass of observation and prediction that underlies the above brief review, and illustrates some the main potential pathways of impact from climate change to shellfish specifically.

The effects of climate change on shellfish aquaculture systems Drawing on a review of the extensive literature on existing and potential climate change impacts on the oceans, fisheries, and on aquaculture systems (e.g., Handisyde et al. 2006; Brander 2007; Brierley and Kingsford 2009; De Silva and Soto 2009; Badjeck et al. 2010), the main pathways through which climate variability and change are hypothesized to impact shellfish are illustrated in Figure 17.2. After a brief explanation of the diagram, we review the available evidence for each of the illustrated pathways. First, it is important to remind ourselves that climate change is not the only “driver” affecting the operation of shellfish farms and may not be the most influential one. Nonclimate drivers of change could include pollution that affects the location and operating costs of shellfish farms; competing uses for the foreshore and coastal waters that constrain the expansion of the sector—for example, industry and urbanization, tourism and residential

467

development, and protected areas; and changes in coastal land use and associated marine habitats which affect key life cycle stages of shellfish populations or water quality. Markets change, too: People no longer eat the quantity of oysters that New Yorkers did in the eighteenth and nineteenth centuries (Kurlansky 2006) but expectations on quality and variety are likely to increase. Although farming techniques for many shellfish remain simple, technology also evolves, notably in the development of hatcheries, in the selective breeding that this makes possible, and in the development of closed or recirculation systems. The legislative and policy context for shellfish farming, processing, and trade also adapts and responds to new threats and opportunities. Climate change is thus an additional driver of change in a sector in which environmental management and food safety already have a strong influence. Both of these drivers will themselves be affected by climate change, as we will discuss in the next section. Human activities are generating increased emissions of greenhouse gasses—principally methane and carbon dioxide. The changing chemical composition of the earth’s atmosphere is leading to a global change in the climate. There is an overall warming trend, accompanied by changing weather patterns (which may include localized or seasonal cooling). Increased CO2 emissions are also responsible for the “other CO2 problem”— ocean acidification, which has far-reaching implications for all calcifying marine organisms, of obvious concern to shellfish farming (hence the large number of experimental studies, reviewed below) and with particularly alarming prognoses for the future of coral reefs (Doney et al. 2009). Warming and changing weather are leading to rising sea surface temperatures and changes in the flow and variability of ocean currents (the two are linked). These changes lead to linked sets of physical, chemical, and biological effects, which include reduced oxygenation in waters

468

Shellfish Aquaculture and the Environment

Human activity

Increased emissions: other GHGS

Increased CO2 emissions

Atmospheric warming and changing weather patterns

Rising sea surface temperature

Nonclimate drivers of change: e.g., pollution, competing uses for coastal waters, fishing pressure, land use and habitat change, changing patterns of demand, technology development, legislation and policy

Increased CO2 concentration in oceans

Ocean acidification (reduced pH)

Reduced oxygenation of heated, enclosed waters Changes in primary production and food web structure and function Changes in natural spatfall Increased frequency of pathogenic infections

Changing ocean currents

Changes in distribution of pests and alien species

Sea level rise

Increased frequency and severity of extreme weather events

Coastal flooding Extreme rainfall events High winds and wave energy

Reduced rates of calcification in shellfish Decreased growth and production Increased mortality, altered recruitment patterns Shifting sites of production Changes in species cultured

Increased losses and direct damage to aquaculture installations and coastal infrastructure

Increased costs of production Changes in production volume and value Needs and opportunities for adaptation in culture locations, species grown, technology, system management, transportation and marketing

Figure 17.2 Multiple pathways by which climate change potentially impacts molluscan shellfish aquaculture. GHGs, greenhouse gasses.

where circulation is limited, changes in primary production and food web structure and function, and in life history processes such as spatfall of shellfish. Warming and changes in currents are also associated with increased frequency and poleward range expansion of HABs, as well as facilitating the spread of pests and alien species. These are issues that already plague the shellfish farming sector. In combination, the changes outlined above are likely to lead to l physiological stresses that decrease growth and production, although warming could improve growth rates by speeding up metabolism, if primary production is not limiting. There will certainly be changes in where particular species can be successfully farmed, or in which species are farmed in areas traditionally used for shellfish cultivation.

The increasing heat energy in the oceans and atmosphere and change in currents also affect the formation of storm events. The warming sea and melting polar ice caps are leading to a rise in sea level, which, when taken together with potentially increased storm intensity, are likely to increase the severity of coastal flooding, extreme rainfall events, and stormy weather. This combination of changes threatens to bring increased losses of stock and direct damage to aquaculture installations and service facilities, to coastal infrastructure, and indeed to people’s homes. In countries without effective storm early warning and evacuation systems, and with large, vulnerable coastal populations, hundreds or even thousands of human lives are lost with every major storm, as was the case with the major

Implications of global climate change

cyclones in Bangladesh and Myanmar in the last 3 years (ESCAP and UNISDR 2010). Through these pathways of impact, shellfish farming systems and the value chains associated with them are likely to face increased costs of production and changes in the volume, species composition and value of their production. These costs will be passed on to consumers, affecting demand. There will be a need—or, more optimistically, an opportunity—to adapt. Adaptation options are discussed in Section 4. What, then, do we know of the likelihood and potential magnitude of the impacts described above? In Tables 17.1–17.3, we summarize research on the relationship between climate variables and impacts on farmed shellfish. First, a caveat: Our review is not comprehensive as it is confined to studies in which the authors explicitly make reference to climate change. For this reason, it is largely recent. There is of course an extensive literature, dating back more than a century, linking shellfish population dynamics to climate variability but as it predates concern for anthropogenic warming it makes no reference to climate change and no prediction of likely impacts of such change. A more comprehensive review— beyond the scope of this one—is required to make use of this data set to search for relationships that can help predict future change. The use of “climate analogues” is standard in climate change research applied to other agricultural systems. Using temporal analogues involves investigating the effects of past climatic events, including climate variability, to predict future changes and inform adaptation (Glantz 1988; Easterling et al. 1992). Spatial analogues refer to conducting research in one region and identifying parallels to how another might be affected (Ford et al. 2010). In agricultural research, this involves looking at what species and farming systems will be suitable for the anticipated climate, based on studying areas of the world where such conditions are the current norm. To take an example, condi-

469

tions in Chesapeake Bay in a warm future (Paolisso et al. 2010) might look like presentday conditions in the Gulf of Mexico, so the shellfish culture conditions on the Gulf coast may be a spatial analogue for those that can be expected in Chesapeake Bay at the end of the twenty-first century. Literature that makes reference to the potential impacts of climate change on shellfish aquaculture is concentrated on looking at two of the pathways illustrated in Figure 17.2: ocean acidification and its potential impacts on shell formation and growth rate; and the spread and impact of HABs and its effects (Tables 17.1 and 17.2). Studies of the direct and indirect effects of warming, and of multiple, interacting pathways are rarer (Table 17.3). While there are many reports of storm and flood damage to shellfish farming operations, there are none in the peer-reviewed literature which demonstrate or claim increased frequency or severity of such events, linked with global warming. Studies of the impacts of ocean acidification of shellfish have burgeoned in the last 5 years, since ocean acidification became a global concern. While some early studies used a range of pH outside values even the gloomiest prognosis for the future, the studies reviewed in Table 17.1 have confined experimental manipulation of pH to levels corresponding to values predicted by 2050 or 2100 under IPCC emissions scenarios. In general, the 0.5 pH unit decrease in seawater forecast by 2100 is associated with decreased growth and calcification rates in juvenile and adult oysters, mussels, and scallops, although the results for mussels are ambiguous. Waldbusser et al. (2011) point to the complexity of the relationship, which is mediated by salinity and temperature changes that will occur concurrently with acidification. Talmage and Gobler (2010) include preindustrial CO2 levels and associated seawater pH in their experiments, which demonstrate that bivalve development may already have been impacted by the changes that have taken place

Table 17.1 Evidence for impact of increased CO2 concentration and ocean acidification on shellfish species and culture operations. Species and location

Impact

Nature of evidence

Reference

Oyster (Crassostrea virginica); Chesapeake Bay, USA, laboratory studies

Estuarine waters are more susceptible to acidification because they are subject to multiple acid sources and are less buffered than marine waters. Biocalcification declined significantly with a reduction of 0.5 pH units and higher temperature, but salinity mitigated the decrease in biocalcification. Complex relationships between water chemistry, eutrophication, pH, and biocalcifiation do not allow simple prediction of reduced biocalcification with increasing CO2 concentration in the atmosphere.

Field water quality monitoring in Chesapeake Bay, and experimental studies on oyster larvae

Waldbusser et al. (2011)

Hard-shell clams and scallops: Mercenaria mercenaria and Argopecten irradians; laboratory studies, USA

Compared with present CO2 conditions (390 ppm), larvae grown under near preindustrial CO2 concentrations (250 ppm) had significantly faster growth and metamorphosis, higher survival and lipid accumulation rates, as well as having thicker, more robust shells. Bivalves exposed to CO2 levels expected later this century (750 ppm) had shells that were malformed and eroded.

Experimental studies on larvae, with link to past, current, and projected oceanic CO2 concentrations

Talmage and Gobler (2010)

Eastern oyster (Crassostrea virginica) and the Suminoe oyster (Crassostrea ariakensis); laboratory studies, USA

Oyster larvae grown in estuarine water under four pCO2 regimes, representing preindustrial, present, and projected concentrations. Crassostrea virginica experienced a 16% decrease in shell area and a 42% reduction in calcium content when preindustrial and end of twenty-first-century pCO2 treatments were compared. Crassostrea ariakensis showed no change to either growth or calcification. Both species demonstrated net calcification and growth, even when aragonite was undersaturated.

Experimental studies on larvae, with link to past, current, and projected oceanic CO2 concentrations

Miller et al. (2009)

Mussels (Mytilus edulis); laboratory studies, Norway

Virtually no growth at pH 6.7 and reduced growth at pH 7.1, but no significant difference between growth at pH 8.1 (current) and pH 7.4 and 7.6

Short-term experimental studies on juveniles and adults

Berge et al. (2006)

Mussels (Mytilus edulis) and oysters (Crassostrea gigas); laboratory studies, the Netherlands

The calcification rates of the edible mussel and Pacific oyster decline linearly with increasing pCO2. Mussel and oyster calcification may decrease by 25% and 10%, respectively, by the end of the century, following one of the IPCC emissions scenarios (740 ppm in 2100).

Experimental studies with juvenile and adult organisms

Gazeau et al. (2007)

470

Table 17.2 Changed distribution, increased frequency, and/or virulence of pathogenic infections and alien species affecting shellfish. Species and location

Impact

Nature of evidence

Reference

Oysters (Crassostrea virginica); northeast USA

Statistically significant links between increased winter water temperature and the northward movement of outbreaks of protozoan Perkinsus marinus, the causative agent of dermo disease in oysters.

Observation of recent sea surface temperature trend and northward range expansion of pathogen

Cook et al. (1998)

Shellfish; USA and globally.

Increased economic impact of warming and increased climate variability on foodborne and waterborne pathogens, with shellfish singled out as a key source of concern. For example, seasonal warming of sea surface temperatures enhances plankton blooms of copepods that serve as reservoirs for Vibrio cholerae. These blooms have been followed by a lagged increase in cholera cases that generally occur in the wake of El Niño events.

Observational; literature review using correlations with past climate variability to infer impacts of future climate change

Rose et al. (2001)

Harmful microalgae species; the North Sea

The risk of harmful dinoflagellate and raphidophyte blooms will increase rather than decrease due to climate change, as growth of these species increases under predicted 2100 ocean conditions, as simulated experimentally.

Laboratory studies on the culture of various harmful and nonharmful microalgae at elevated temperatures

Peperzak (2003)

Unspecified shellfish species; North America

During the 1987 El Niño, a bloom of the dinoflagellate Karenia brevis, previously confined to the Gulf of Mexico, extended northward after warm Gulf Stream water reached far up the east coast, resulting in human neurological shellfish poisoning (NSP) in North Carolina. An outbreak of amnesic shellfish poisoning (ASP) also occurred on Prince Edward Island when warm eddies of the Gulf Stream neared the shore, and heavy rains increased nutrient-rich runoff, resulting in a bloom of the diatom, Pseudonitzchia

Observational; case study using correlations with past climate variability (e.g., ENSO events) to infer impact of future climate change

Patz et al. (2006)

Oysters, (Crassostrea virginica); Louisiana, USA

Analyses of a 10-year time series of disease prevalence and environmental parameters showed a teleconnection between the ENSO and oyster disease—Perkinsus (=Dermocystidium) marinus—in the northern Gulf of Mexico. Salinity increases precede increased disease prevalence by several months; these increases are strongly driven by ENSO events. This relationship could be used to inform the management of oyster populations under predicted climate change.

Observational; case study using correlations with past climate variability (ENSO events) to infer impact of future climate change

Soniat et al. (2006)

Pacific oyster (Crassostrea gigas) introduced to the Netherlands

Spatfall of introduced Crassostrea gigas on natural mussel beds in the northern Wadden Sea is linked to high late-summer water temperatures, forecast to be more frequent under climate change, with potential impact on future mussel harvests.

Observational; links an 18-year recruitment data set with temperature records

Diederich et al. (2010)

471

Table 17.3 Evidence for other direct and indirect impacts of climate change on coastal waters that affect shellfish species and culture operations. Climate variable

Species and location

Impact

Nature of evidence

Reference

ENSO cycles

Oysters (Crassostrea virginica); Gulf of Mexico, USA

In a National Oceanic and Atmospheric Administration study of chemical contaminant burdens in oysters (trace metal, polynuclear aromatic hydrocarbon, and pesticides), ENSO cycles were hypothesized to be important in establishing the interannual variability in contaminant body burden. The implications are that climate variability and change may affect levels of contaminants in shellfish.

Observational; correlations between interannual patterns of contaminant loads and large-scale climate cycles, 1986–1992

Kim et al. (1999)

Spatfall and/or recruitment to natural populations

Macoma baltica; the Wadden Sea, the Netherlands

Rising seawater temperatures affect recruitment by a decrease in reproductive output and by spring advancement of bivalve spawning. Apparently, global warming upsets the evolved reproductive strategy of this marine bivalve to tune its reproduction to the most optimal environmental conditions for the first vulnerable life stages, most importantly the match/ mismatch of time of spawning with that of the phytoplankton bloom and the settlement of juvenile shrimps on the tidal flats

Field observation, 1973–2001

Philippart et al. (2003)

Ocean warming

Red abalone, Haliotis rufescens, and green abalone, Haliotis fulgens; California

For red abalone, warm temperatures increased the onset of withering syndrome, a fatal abalone disease, and halted growth and reproduction. In contrast, green abalone survivorship, growth, and reproduction were relatively robust irrespective of temperature, while their growth and reproduction were most strongly influenced by food quantity. Cool-water red abalone suffer stronger consequences in warm water than do green abalone.

Experimental study of synergistic effects of temperature and food quantity and quality on survivorship, growth, and reproduction

Vilchis et al. (2005)

472

Table 17.3 (Continued) Climate variable

Species and location

Impact

Nature of evidence

Reference

Changes in primary production

Phytoplankton; global

Reduced primary production in the post-1999 warming period (likely to have impact on shellfish production).

Emergent relationships between observed warming and productivity, and future predicted warming

Behrenfeld et al. (2006)

Ocean warming

Bivalve species; global

Greater tolerance for temperature variability in temperate-zone bivalves means that warming will have proportionately greater impact on survival, range, and productivity of tropical species than temperate ones.

Literature review of upper thermal tolerance limits

Compton et al. (2007)

Increased frequency of extreme rainfall events and river flows

Mussels (Mytilus edulis); Conwy Estuary, Wales, UK

Climate change will bring more frequent extreme rainfall and high flow events in rivers, which can impact shellfish populations and farms in estuaries through decreased salinity and/or increased transport of sediments, organic matter, and nutrients. One extreme event impacted condition index, while the other reduced total hemocyte count, suggesting impact on health and resilience of estuarine mussel populations.

Evidence from sampling over 18 months at control and impacted sites, covering two extreme flow events, with a hypothesized link to climate change

Oliver et al. (2008)

Coastal hypoxia, driven by eutrophication and sea temperature rise

Review of 206 species of benthic metazoans; globally

Molluscs are the benthic organisms most tolerant to hypoxia, crustaceans the least. This is therefore not likely to be an important way in which global warming affects molluscan aquaculture.

Review of 872 experimental studies of hypoxia tolerance

VaquerSunyer and Duarte (2008)

473

474

Shellfish Aquaculture and the Environment

Table 17.3 (Continued) Climate variable

Species and location

Impact

Nature of evidence

Reference

Reduced upwelling in coastal waters and increased water retention times in bays

Mussels (Mytilus edulis); Galicia, northwest Spain

The extent and intensity of the upwelling season has decreased by 30% and 45% over the last 40 years, respectively. This has led to a 240% increase in water retention time in the four large coastal inlets where 15% of the world’s harvest of blue mussels come from, which is causing increasing occurrence of harmful microalgae, with dramatic negative effects on raft cultivation of mussels, confining harvests to progressively fewer number of days per year.

Hydrographic, biological oceanography and aquaculture production studies over a 40-year period

ÁlvarezSalgado et al. (2008)

Ocean warming and acidification (synergistic effects)

Sydney rock oyster, Saccostrea glomerata; southeast Australia

Fertilization rate, embryonic development rate, and frequency of abnormal veligers increased with temperature and CO2 increase. Predicted changes in ocean acidification and temperature over the next century may have severe implications for the distribution and abundance of this commerciallyimportant species.

Laboratory studies, both short term and prolonged

Parker et al. (2009)

to date. The species and life stage specificity of acidification impacts makes easy generalizations impossible. Recent meta-analysis of the experimental studies of potential impacts of acidification on marine organisms remains disputed on whether the oceans face a crisis or not (Dupont et al. 2010; Hendriks and Duarte 2010; Hendriks et al. 2010). The predicted impacts of warming on pathogens and parasites of shellfish (Chapter 13 in this book) and of invasive species (Chapter 14 in this book) are apparently more clear-cut than those of acidification (Table 17.2). Correlating past variability and warming

with the spread of diseases and alien species leads to a prediction that such problems will be more frequent in the future. This claim is strengthened by being rooted in observation on change that has already happened, particularly concerning HABs. The most important harmful algae and their poisoning syndromes (in brackets) relevant to shellfish are diatoms from the genus Pseudonitzschia (amnesic shellfish poisoning), and species of dinoflagellates from the genera Alexandrium, Pyrodinium, and Gymnodinium (paralytic shellfish poisoning), Karenia (neurotoxic shellfish poisoning and aerosolized

Implications of global climate change

475

Sea surface temperature

191 more d °C ay +6 s 7 more 2 1 °C da +4 ys more d 9 6 ay °C s +2

Temperature (°C)

20

15 13°C threshold

10

Historical window of increased opportunity 68 days

J

F

M

A

M

J

J

A

S

O

N

D

Month Figure 17.3 Potential climate change impacts on Puget Sound shellfish toxicity. Climatological monthly means of reconstructed sea surface temperature (SST) in Sequim Bay, Puget Sound, from 1921 to 2007 indicate an average 68-day duration of the 13°C threshold for accelerated growth of the paralytic shellfish poisoning agent Alexandrium catenella (shaded period). Scenarios for warmer SST conditions by 2, 4, and 6°C are shown in gray with the associated widening of the window of increased opportunity for Alexandrium catenella growth. (Reproduced from Moore et al. 2008.)

Florida red tide respiratory syndrome), Dinophysis and Prorocentrum (diarrhetic shellfish poisoning). In a global review of health issues relating to climate change and oceans, Moore et al. (2008) state that, “. . . evidence that climate change has influenced the frequency, duration, and geographical range of HABs is emerging as monitoring data . . . accumulate.” Similarly, Marcogliese (2008) links past climate variability with the dynamics of pathogens and parasites to hypothesize that “transmission rates of parasites and pathogens are expected to increase with increasing temperature. Evidence suggests that the virulence of some pathogens and parasites may also increase with global warming.” Despite this apparent weight of evidence, we have limited understanding of marine ecosystem responses to the multiple, interacting pathways of potential impact of climate change, and lack of any knowledge of whether the marine biota can

adapt genetically and phenotypically to the pace of current climate change. This makes prediction of the future distribution “fraught with difficulties” (Hallegraeff 2010). Of course, this should not stop anyone from trying ,and the presentation of future scenarios, even if of unknown probability, can be useful to galvanize responsive action. For example, Moore et al. (2008) predict that future warming will lengthen the season in which oysters from Puget Sound will be dangerous to consume due to possible phytotoxin contamination (Fig. 17.3). This communicates clearly to growers, traders, and consumers the likely practical implications of climate change. So far, the news is all bad. However, climate change can bring positive benefits in certain places and to some shellfish fisheries and farming systems. When we consider the range of potential pathways of impact of variables

476

Shellfish Aquaculture and the Environment

associated with ocean warming and changes in currents (Table 17.3), the outlook becomes more mixed. While some field observations and experimental studies show negative population-level impacts of warming on clam and abalone populations (Philippart et al. 2003; Vilchis et al. 2005), studies of wild,

exploited scallop populations in the middle of their current species range show an increase in recruitment—associated with warm years in a variable climate in the case of Peruvian scallops and with both warm years and an overall warming trend in the case of European scallops (Fig. 17.4).

(A) 2.5

140 120

2

Temp PCA

1.5

100

1

80

0.5

60

0

Temp PCA

R (# age–2 scallops tow–1)

R

–0.5

40

–1 20 0 1990

–1.5 1995

2000

2005

–2 2010

Year

(B) Landings

Sea Surface Temperatures

20

4000

19

3500

18 17

3000

16 2500 15 2000

14

1500

13

1000

12

500

11

Sea Surface Temperatures (°C)

Landings (10 of tons)

4500

10

0

82 83 84 85 86 87 88 89 90 91 92 93 94 95 96 97 98 99 00 01 02 03 Figure 17.4 Warmer waters may favor shellfish reproduction: evidence from recruitment to scallop fisheries. (A) Time series of mean spring sea temperature (a principal component derived from four time series of water temperature) in the North Irish Sea, and scallop (Pecten maximus) recruitment (R) 2 years later (reproduced from Shephard et al. 2010). (B) Annual catch of the Peruvian scallop (Argopecten purpuratus) and sea surface temperature changes in Pisco, Peru, 1982–2003 (Badjeck et al. 2009). During the 1982–1983 El Niño, a 2-year time lag is observed between temperature peaks and increases in landings as fishing effort peaked in 1984–1985 when the 1982–1983 cohort reached its marketable size. In the 1997–1998 El Niño, the scallop stock again increased but instead of waiting for the cohort biomass to build up, small juvenile scallops of low market value were extracted immediately, leading to both a shorter lag (1 year) and to lower landings. Temp PCA, temperature principle component analysis.

Implications of global climate change

Elevated temperatures of coastal waters could also lead to increased production of aquaculture species by expanding their range. These species could be cultivated in higher latitudes as well as in existing farms as a result of a longer warm season during which water temperature will be near optimal. A decrease in sea ice cover could widen the geographical boundaries, allowing cultivation of commercially valuable species in areas hitherto unsuitable (McCarthy et al. 2001). However, tropical species are often already near their thermal limits and a slight temperature change might have significant negative effects on their physiology. In the case of bivalves, tropical species live closer to their upper thermal lethal limits than temperate species, which are better adapted to a wider temperature range fluctuation (Compton et al. 2007). This has potentially serious consequences for tropical shellfish farmers, especially as overall vulnerability of aquaculture systems to climate change is higher in those developing countries in the tropics that have a high risk of exposure to multiple climate stressors; a relatively greater economic and dietary dependence on fish and shellfish resources than wealthier countries with larger, more diverse economies; and limited government and personal resources (such as savings and insurance) with which to adapt to change (Handisyde et al. 2006; Allison et al. 2009). Complex, multifactorial impacts have also been demonstrated from review of recent change, monitoring of the impacts of extreme events, and from laboratory studies that manipulate more than one variable (Table 17.3). Again, the prognosis is mixed. Reduced oxygen concentration in warmer, more stratified waters will impact molluscs less than other coastal shellfish, such as crabs (Vaquer-Sunyer and Duarte 2008). Resilience of mussel populations in estuaries may decrease if extreme flow events become more frequent or more intense (Oliver et al. 2008). For particular species and culture systems, predictions that integrate a range of possible

477

stressors have been made, although usually these are based on inductive reasoning rather than empirical evidence or formal prediction. This is not a criticism: Plausible scenarios of climate change’s potential impacts are required to inform policy decisions, and expert judgement based on a broad knowledge of pathways and relationships can be useful. For example, according to Bell et al. (2010), pearl farming in the tropical Pacific faces risks from increased acidification of the ocean as the shells of black-lipped pearl oysters will be weaker. This is likely to lead to higher rates of predation of juveniles and lower rates of collection of wild spat. Large-scale farms may be forced to rely more heavily on hatcheries to produce spat, increasing production costs. Acidification may also impair the ability of pearl oysters to form nacre. If so, pearl quality may decline progressively, reducing the value of pearls produced in the future. More severe cyclones can also be expected to increase the risk of damage to the infrastructure of pearl farms in subtropical Pacific Island countries. Major shellfish culture sites also merit a holistic view of the potential impacts of climate change. Paolisso et al. (2010) have conducted an overall diagnosis of the potential impact of climate change on Chesapeake Bay that serves as a good example. Scenarios for CO2 emissions indicate that by the end of the twentyfirst century the bay region will experience increases in CO2 concentrations, sea level, and water temperature of the order 50–160%, 0.7–1.6 m, and 2–6°C, respectively. Also likely are increases in precipitation amount (very likely in the winter and spring) and intensity, and the intensity of cyclones (though their frequency may decrease), and in sea level variability. The greatest uncertainty is associated with changes in annual streamflow, though it is likely that winter and spring flows will increase. Climate change alone will cause the bay to function very differently in the future. The authors conclude that likely changes include increases in coastal flooding and submergence of estuarine wetlands, increases in

478

Shellfish Aquaculture and the Environment

salinity variability, in HABs, and the extent and duration of hypoxia. Eelgrass, the dominant submerged aquatic vegetation in the bay, will decrease in abundance and extent. There are also likely to be alterations in interactions among trophic levels, with subtropical fish and shellfish species ultimately being favored in the bay. The magnitude of these changes is sensitive to the CO2 emission trajectory, so that actions taken now to reduce CO2 emissions will reduce longer-term climate impacts on the bay. More studies of this type will be useful to the shellfish farming sector in other major growing areas. Almost all the studies in the literature focus on the biological impacts of climate change on production systems, but climate change will impact all links in the value chain linking producers and consumers. For example, any increases in the intensity and frequency of extreme climatic events such as storms, floods, and droughts will negatively impact aquaculture production and may also result in significant damage mainly related to decreased farming capacity (loss of infrastructure) or decreased access to markets (damaged roads). This may translate into economic losses that small-scale farmers are unable to cope with. In Indian River Lagoon, FL, the Florida Department of Agriculture estimated Hurricanes Frances and Charley (2004) caused $8.7 million in losses for clam and oyster farmers (Bierschenk 2004). This estimate pertained only to the value of the stock and to lost incomes and to traded value; it did not include infrastructure losses such as to buildings, docks, vessels, and nursery and hatchery facilities. Climate change may also act through indirect pathways which may be difficult to foresee. For example, Yeoman and McMahonBeattie (2006), in a study of the potential impacts of climate change on tourism to Scotland, included a concern that removing local shellfish from menus, as a result of increased risk of disease associated with warming, could detract from visitor experi-

ence and act as one among a range of disincentives for them to return. In aquaculture, problems with disease management, limited availability of suitable sites, and a lack of stable and equitable access to input and output markets mean that it can be a high-risk venture in many parts of the world. The addition of climate-associated risks could slow the growth of aquaculture in developing countries and reduce recent gains in closing the “supply gap” through provision of lower-cost food to lower-income consumers, and providing millions of small-holder farmers in developing countries with a new cash crop for export and a nutritious supplement to staple crops (De Silva and Soto 2009).

Adapting shellfish farming to climate change impacts Projected impacts of climate change tend to sound apocalyptic when they do not address the possibility of adaptation. In this case, we are considering adaptation as the actions of people and their institutions, rather than in the Darwinian sense of genetic change by organisms in response to changing environments, although that, too, may occur under the selective pressure of rapid climate change. Humans—even conservative ones—are highly adaptable, and adaptation is possible at multiple scales and in multiple domains. Figure 17.5 provides a framework through which to summarize the possibilities for adaptation in the shellfish farming sector. Modifying slightly the ideas of Preston and Stafford-Smith (2009) and Grafton (2010), we consider: Who can adapt? What elements of the system can be adapted? How can resources and capacity required to adapt be mobilized, and what processes support these adaptation options? Adaptive actions can be taken by individual shellfish farmers, processors, and traders, or even individual consumers (“Who”?). An

Implications of global climate change

479

What to Adapt? Biophysical System

Habitat

Water quality

Species

Social and Economic system

Employment

Market

Governance

Scale

Local Regional National

Global

Who Adapts? Individuals

Groups

How to Adapt?

Government

Farmers Firms Regulatory bodies Traders Communities Advisory services Processors Producer associations Retailers Consumers

Drawing on Capital

Human Social Natural Physical Financial Approaches

Autonomous Palnned Flexible Mandated Responses

Social

Economic

Technical

Figure 17.5 The adaptation process. (Adapted from Preston and Stafford-Smith 2009 and Grafton 2010.)

example of autonomous adaptation is described by Nell (2001) in New South Wales, Australia, where summer temperatures on exposed leases can rise to 30–40°C and kill both Sydney rock and Pacific oysters. The problem is most acute when low tides coincide with the warmest period on a hot summer’s day. Some farmers have installed irrigation sprinkler systems on their leases and spray saltwater over the oysters to keep them cool. Other farmers use shade cloth over trays to keep oysters cool and predators out, but this practice restricts water flow over the trays and increases the risk of mudworm infestations. Spraying oysters with seawater also keeps the oysters cooler than the shade cloth does. Individuals can be adaptive in adjusting to short-run changes of the type described above and to short-lived “shocks,” while governments and communities have a comparative advantage in large-scale adaptation that requires substantial investment over longer

time periods, such as rezoning a coastal area to reflect habitat changes. In conjunction with the private sector, governments can help shellfish farmers cope with climate change impacts through risk sharing and transfer mechanisms such as insurance schemes and emergency funds. For instance in the United States, clam growers can be insured for losses that occur because of decrease in salinity, disease, freeze, hurricane, ice floe, oxygen depletion, storm surge, and tidal wave under the Cultivated Clam Pilot Crop Insurance Program, part of the Federal Crop Insurance Program (USDA 2010). However, state-sponsored crop and income insurance is only a partial solution; it does not prevent the loss of lives or assets, and has been criticized as being a new mechanism for transferring subsidies (van Anrooy et al. 2006). Governments have also an important role in creating an enabling environment for adaptation to take place, and a regulatory

480

Shellfish Aquaculture and the Environment

environment that prevents reactive adaptation that has negative environmental, social, and economic consequences. At national level, climate change adaptation should be mainstreamed in aquaculture development plans. In Peru, where scallops represented 43% of the total value aquaculture products exported in 2009(Produce 2010a), the new aquaculture development plan 2010– 2021 (Produce 2010b) considers climate changes and natural disasters but no explicit risk management and adaptation plans are put forward for scallops, a resource highly sensitive to climate fluctuations (Wolff and Mendo 2000). Similarly, national adaptation programs of action (NAPAs) prepared by least developing countries should include the aquaculture sector since they are a basis for channeling the adaptation funds promised in recent global climate change agreements (UNFCCC 2009). Some NAPAs contain provision for adaptation of fishing and fish farmingdependent communities, but in light of their high vulnerability, more needs to be done to support adaptation through capacity building, finance, technology, and innovation. In terms of what actions can be taken (“What to adapt”?), adaptations to ensure that farming systems are suited to the changing biophysical conditions may include choice of species cultivated or location chosen, or, at a larger scale, water quality or habitat management to maintain good growing conditions in the face of change. As an example of adaptation through species change, due to increased sea surface temperatures attributed to global warming, abalone (Haliotis), are now present in England’s West Country and the local government has been promoting this new industry to rejuvenate the ailing fishing sector (Brown and Sutton 2002). Maintaining a livelihood or a community may entail adjustments in livelihood strategy and employment, including diversification out of shellfish farming if conditions become unsuitable. Changes in species grown may

require new markets to be developed, or existing consumer preferences to be adjusted. In today’s globalized seafood markets, such adjustments are relatively minor and take place quickly. Governance arrangements may also have to adapt to new conditions—this can mean anything from extending the coverage of food safety inspections to changing regulations to allow species translocations, changing legal designation of shellfish growing areas, or renegotiating leases to the seabed to allow for changing distribution of cultivated shellfish. Adaptive measures can be implemented (“How to adapt”?) by deploying available capital, whether that comprises knowledge and skills (human capital), or involves mobilizing farmers’ groups to lobby for changing policy or legislation, or simply getting support from friends and neighbors to deal with loss of assets during a storm or flood (social capital). Land or water rights (natural capital) can be sold, leased, or deployed for different uses (e.g., coastal leisure tourism). Processing plants can be adjusted to new product lines and small boats switched from tending fish farms to inshore fishing (physical capital). Loans, savings, and insurance (financial capital) can all be mobilized to invest in new ventures, ease temporary production turndowns, or recover from shocks such as flood damage of closure due to shellfish poisoning outbreaks. Improved depuration technologies can be brought into use to deal with increased microbial or HAB concerns. Adaptation can be autonomous (bottomup), planned (top-down), negotiated between stakeholders (flexible), or mandated in law, depending on the nature of required response. Autonomous decisions usually take place at farm or household level, while planned responses, such as investment in disaster preparedness and response strategies, require multilevel planning and coordination. An example of autonomous adaptation to frequent storms is found among the oyster farmers of the western coast of Taiwan (Chen

Implications of global climate change

2008): Information available on the size and timing of typhoons (the provision of which is a planned adaptive response) is used by farmers to make decisions on dates of setting and harvesting oysters. Farmers also adapt to the increased frequency and severity of storms by selling small-size or medium-size oysters that can be harvested between typhoon seasons, rather than growing to a larger size and risking loss in the typhoon season. Responses to climate change can be technical (e.g., early warning systems, “climateproofing” of infrastructure, improved means of testing for shellfish toxicity and depuration), economic (e.g., weather-linked insurance, planned economic restructuring to reduce dependence on climate-sensitive industries), or social/institutional (e.g., community self-help programs, participation in grassroots adaptation knowledge networks). Adaptation is also a political process. Climate change justice is a term usually heard with reference to the moral obligation of highemissions wealthy nations to assist adaptation by low-emission poor nations that are vulnerable to climate change, but it may apply on a microscale as well. Shellfish farmers are likely to be in the vanguard of climate change as they often experience the impacts of changes on both land (and freshwater) and sea. Issues such as investment in sea defenses versus managed retreat in the face of sea level rise are not purely economic ones, as they involve preservation or loss of culturally valued landscapes and of people’s homes and livelihoods. For example, in Norfolk, England, the principle of social justice has been invoked by local residents and politicians to challenge a policy decision to discontinue maintenance of coastal defenses (McKenna et al. 2009).

Costs of adaptation To maintain the flow of benefits to society and the economy from fisheries, governments are

481

likely to have to increase their investments in developing coherent “climate-proof” sectoral policy and legislation, management, and development, including some of the elements described above. The following costs are likely to arise to address adaptation, or in response to fisheries and aquaculture decline due to climate change:

• Investments in “climate-proofing” infrastructure (e.g., coastal defenses, design of harbors); • Social and economic costs of redundancies in fishing and aquaculture sector; • Weather-linked unemployment insurance for fisherfolk (mostly used in developed countries; few developing countries have this kind of formal social security system); • Costs of ecosystem maintenance and repair—for example, marine protected areas, restoration of salt marshes and mangroves, artificial reef construction, beach replenishment, maintaining “environmental flows” in rivers; and • There will also be a need to consider the costs to fisheries and aquaculture of adaptation in other sectors; for instance, the use of “soft” coastal defenses with fish and shellfish habitat value (mangroves, artificial reefs, protecting existing reefs) where fisheries and aquaculture are important to coastal livelihoods, rather than “hard” defenses such as sea walls, which prioritize land values over coastal habitat values. Many of the adaptation options and processes described above are not specific to the shellfish sector, so, to conclude this section, it seems appropriate to consider the sorts of sectorspecific technical adaptations we foresee. First, we think there will be widespread changes in species cultured in existing shellfish-growing areas. Second, the increased frequency of HABs could close off fisheries and farming areas more frequently, and there will be a need for improved surveillance systems to prevent

482

Shellfish Aquaculture and the Environment

health incidents to consumers and economic losses to growers. Third, given the unpredictable nature of climate change impacts, we can foresee a shift toward adoption of more “climate-proof” closed or recirculation systems in aquaculture—akin to intensive poultry production—where the environment is under more direct human control. Aquaculture dependent on the collection from the wild of climate-sensitive natural larvae, spat, or “seed” could begin to disappear in their current locations. An obvious adaptive measure is to close the life cycle of these species by developing hatcheries, but this can be both technologically demanding and expensive. With strong markets providing an incentive for continued innovation, we are confident that the sector will adapt and persist.

Shellfish aquaculture and climate change mitigation Aquaculture is not a large contributor to agricultural greenhouse gasses, and neither, therefore, does it have enormous global potential for halting or reversing climate change, but, nevertheless, it is responsible for a small (and currently unquantified) proportion of the greenhouse gas emissions from agriculture, which formed around 13.5% of total anthropogenic emissions in 2004 (IPCC 2007b). There is also potential for making aquaculture a lower-carbon food production system. There have even been discussions on the role that aquaculture could play in carbon capture and storage. Molluscan shellfish aquaculture features in both the debate on low-carbon food futures and on aquaculture’s potential for mitigation. Aquaculture systems are responsible for greenhouse gas emissions, caused by direct energy consumption, indirect or embodied energy consumption, land conversion, and as a result of soil, water, and waste management (Bunting and Pretty 2007). Direct energy con-

sumption constitutes the most obvious and widely assessed source of carbon emissions from aquatic farming (Bunting and Pretty 2007). They are generated through a range of activities, for example, the collection/ production of juveniles, general system operations, harvesting, processing, and distribution of the product. The dependence of an aquaculture system on industrial energy varies with the means of production, the intensity of the operation, the degree of mechanization, and the quality and quantity of feed used (Troell et al. 2004). Indirect or embodied energy inputs necessary for aquaculture production include, for example, the energy required to sustain human labor and to build and maintain fixed capital assets such as farm infrastructure, processing facilities, harvesting machinery, and transportation equipment. Depending on the aquaculture system, the scale and form of these inputs will vary widely (Tyedmers and Pelletier 2007). In different studies, the differentiation between direct and indirect energy use may vary. Most importantly, the energetic costs of producing, harvesting, processing, and transporting feed components are sometimes counted as direct, and in other cases as indirect, energy consumption. This needs to be kept in mind when comparing the energy efficiencies and carbon footprints of different food production systems estimated by different studies. In Table 17.4, the global warming potential (GWP) and cumulative energy demand of different aquaculture systems is presented, highlighting the relatively low GWP and energy inputs for shellfish aquaculture. Various possibilities exist to reduce greenhouse gas emissions in aquaculture still further, at both farm level and higher levels. At the farm level, these include the reduction of energy and fuel use, the generation and application of renewable energies, as well as the adoption of more resource-efficient culture practices. Direct and indirect fossil fuel use on farms can be reduced by use of energy efficient machinery, minimizing waste, using energy-

Table 17.4 Global warming potential (GWP) and cumulated energy demand (CED) for aquatic products derived from aquaculture, shellfish in boldface. Species

System

GWP (kg CO2e t−1)

CED (MJ t−1)

Source

Turbot

Inland recirculating system; France

6017

290,986

Aubin et al. (2009)

Arctic char

Land-based freshwater recirculating system; Canada

10,300

233,000

Ayer and Tyedmers (2009)

Atlantic salmon

Land-based saltwater flowthrough system; Canada

5410

132,000

Rainbow trout

Freshwater raceways; France

2753

78,229

Aubin et al. (2009)

Prawns

Extensive polyculture system (prawn, tilapia, milkfish and crab) Philippines

5108

67,000

Baruthio et al. (2009)

Trout

Experimental recirculating system; France

1602

57,659

d’Orbcastel et al. (2009)

Sea bass

Sea cages; Greece

3601

54,656

Aubin et al. (2009)

Pangasius

Pump feed; Vietnam

ca. 1320

54,411

Henriksson (2009)

Salmon

United Kingdom

3270

47,900

Pelletier et al. (2009)

Polyculture

Extensive polyculture system (prawn, tilapia, milkfish, and crab); Philippines

3553

46,000

Baruthio et al. (2009)

Atlantic salmon

Marine floating bag system; Canada

2250

37,300

Ayer and Tyedmers (2009)

Trout

Flow-through system; France

2015

34,869

d’Orbcastel et al. (2009)

Salmon

Chile

2300

33,200

Pelletier et al. (2009)

Salmon

Canada

2370

31,200

Atlantic salmon

Conventional marine net pen system; Canada

2073

26,900

Ayer and Tyedmers (2009)

Tilapia

Pond based; Indonesia

2100

26,500

Pelletier and Tyedmers (2010)

Salmon

Norway

1790

26,200

Pelletier et al. (2009)

Tilapia

Lake based; Indonesia

1520

18,200

Pelletier and Tyedmers (2010)

Milkfish

Semi-intensive; Philippines

ca. 1050

14,879

Henriksson (2009)

Pangasius

Pond based, intensive; Vietnam

8,930

13,200

Bosma et al. (2009)

—

12,000

Troell et al. (2004) Henriksson (2009)

Mussel Milkfish

Intensive; Philippines

ca. 920

11,547

Milkfish

Extensive; Philippines

ca. 500

10,799

Pangasius

Tidal feed; Vietnam

ca. 1030

9861

Mussel

Longline system

—

4000

Tlusty and Lagueux (2009)

Oysters

Low-maintenance aquaculture system; Thailand

ca. 80

580

Henriksson (2009)

483

484

Shellfish Aquaculture and the Environment

efficient lighting, using low-carbon and/or recycled building materials, sourcing inputs locally (feed, seed, fertilizers, etc.) and selling to local markets (Bunting and Pretty 2007). As well as emissions reductions, the coastal environment is also receiving attention as a potential site of carbon sequestration, defined as “the process of increasing the carbon content of a reservoir other than the atmosphere” (UNEP 2006). Much of this interest relates to maintaining vegetation-dominated coastal ecosystems that are net carbon sinks, such as salt marshes, mangroves, and eelgrass beds (Laffoley and Grimsditch 2009). There is also growing interest in using microalgae and seaweed (macroalgae) cultivation as a means to sequester carbon, as they can do so more efficiently then terrestrial plants (Kaladharan et al. 2009; Kumar et al. 2010). They could potentially be grown with shellfish in integrated farming systems, to make such systems “carbon neutral.” The big unresolved question is whether shellfish farms can sequester carbon—a proposal made due to the fact that the shell consists to a large extent of calcium carbonate— CaCO3 (Bunting and Pretty 2007; Hickey 2008). Hickey (2008) demonstrated that the potential of oysters to sequester carbon could even be competitive with some plant species. Based on production data relating to the culture of mussels (Mytilus edulis) on rafts in Killary Harbour, Ireland, Rodhouse and Roden (1987) estimated that 10.8 t C year−1 would be assimilated in mussel production and that the removal rate of carbon during harvest was 0.008 t C m−2 year−1, equating to 80 t C ha−1 year−1. However, the results of this study are based, most importantly, on the shell carbon content without considering factors such as respiration or the calcification process. Calcification induces shifts in the seawater carbonate equilibrium to generate dissolved CO2 (Chauvaud et al. 2003):

Ca2 + + 2HCO3− → CaCO3 + CO2 + H 2O (17.1) The calcification process thus actually releases CO2 into the water, rather than contributing to its net removal from the atmosphere (Chauvaud et al. 2003; Laffoley and Grimsditch 2009). Therefore, shellfish farming does not seem an appropriate means for effective carbon sequestration. However, the carbon cycles in marine systems are very complex involving tightly coupled processes such as photosynthesis, respiration, calcification, and calcite dissolution (Chauvaud et al. 2003), and further research may lead to different understanding of carbon cycling in coastal waters and the potential role of shellfish. We do not, however, expect piles of oyster shells to be eligible for carbon payments anytime soon. The most useful contributions of shellfish farming to climate change mitigation is the compatibility of such farming with the maintenance of coastal environments that do act as carbon sinks, and the energy efficiency with which they can produce high-quality food for people.

Conclusion Molluscan shellfish fishing and farming is a small and climate-sensitive sector of the world economy, but one that is both locally important to sustain livelihoods and coastal cultures and environments, and globally valued for its contribution to many distinctive food cultures—from New Orleans’ oysters Rockefeller, to the cockles that flavor Penang’s renowned street food noodle dish char kuay teow, and from the quotidian moules-frites of Europe’s pays-bas to the upmarket, icesculpted displays of fruits de mer outside Brussels’ Rue des Bouchers restaurants. Shellfish aquaculture is also a dynamic and responsive sector which already adapts to rapidly changing environments and markets. The recent reorientation of environmental

Implications of global climate change

research on shellfish toward understanding the impacts of climate change is building the knowledge base required for planned adaptation but we still face a daunting complexity of multiple interacting pathways of impact linking climate with the operation and economics of shellfish culture and the marketing of shellfish. The three main areas of adaptive response—social, economic, and technical— will all have a part to play. Their relative importance and ultimate success overall in terms of resilient shellfish value chains will be very much dependent on context: location, species, technologies, vulnerabilities of producers, and flexibility of markets. Apart from some scattered anecdotes, at present there is little documented experience of the adaptive strategies pursued by shellfish farmers and other actors in the shellfish value chain, but this is unsurprising as climate change adaptation is a relatively recent field of applied research. What several authors have noted is that individual small-scale producers and traditional production environments are vulnerable, while major growers and retailers are resilient as they have the capital and national or global reach to invest in sourcing from new areas, developing new markets, and switching to new products. Understanding processes of adaptation at multiple scales should be prioritized as an area of research on which future adaptive responses can be developed. There are also surprising gaps in our knowledge of carbon cycling in coastal waters and oceans—an omission being addressed by the United Nations Environment Programme’s “Blue Carbon” initiative (Nellemann et al. 2009). Climate change will not destroy the shellfish farming sector, but adapting will impose further costs on it. In some places, however, there may be gains rather than losses as warming increases spatfall and growth rates and potentially extends the range of farming toward the poles. Close environmental health

485

monitoring is already vital to the sector so the potential increases in harmful algae populations, and on other pests and pathogens, are likely to be rapidly detected, which will help with timely response and adaptation. Shellfish farming associations also have an opportunity to market their products as “low-carbon” food sources. Finally, maintaining clean coastal environments for growing shellfish may also contribute to the maintenance of other coastal ecosystem services, including carbon sequestration. Adaptable growers, buoyant market demand, effective monitoring and regulatory systems, and synergies between shellfish farming, coastal environmental protection, and low-carbon food production systems will all help ensure a future for shellfish farming in a changing climate. It would all be so much simpler, though, if we could agree to cut emissions to slow and reverse the warming trend.

Acknowledgments Thanks to Malcolm Beveridge for early guidance on the structure and scope of this chapter, and for helpful comments and suggestions on a later draft. This review was made possible by funding from the U.K. Natural Environment Research Council QUEST thematic program (QUEST_Fish) and from the German Academic Exchange Services (DAAD)

Literature cited Allison, E.H., Perry, A.L., Badjeck, M-C., Adger, W.N., Andrew, N.A., Brown, K., Conway, D., Halls, A., Pilling, G.M., Reynolds, J.D., and Dulvy, N.K. 2009. Vulnerability of national economies to potential impacts of climate change on fisheries. Fish and Fisheries 10:173–196. Álvarez-Salgado, X.A., Labarta, U., FernándezReiriz, M.J., Figueiras, F.G., Rosón, G., Piedracoba, S., Filgueira, R., and Cabanas, J.M. 2008. Renewal time and the impact of harmful algal blooms on the extensive mussel raft culture

486

Shellfish Aquaculture and the Environment

of the Iberian coastal upwelling system (SW Europe). Harmful Algae 7(6):849–855. Anderegg, W.R.L., Prall, J.W., Harold, J., and Schneider, S.H. 2010. Expert credibility in climate change. Proceedings of the National Academy of Sciences of the United States of America 107(27):12107–12109. Aubin, J., Papatryphon, E., van der Werf, H.M.G., and Chatzifotis, S. 2009. Assessment of the environmental impact of carnivorous finfish production systems using life cycle assessment. The sustainability of seafood production and consumption. Journal of Cleaner Production 17:354–361. Ayer, N.W., and Tyedmers, P.H. 2009. Assessing alternative aquaculture technologies: life cycle assessment of salmonid culture systems in Canada. The sustainability of seafood production and consumption. Journal of Cleaner Production 17:362–373. Badjeck, M.-C., Mendo, J., Wolff, M., and Lange, H. 2009. Climate variability and the Peruvian scallop fishery: the role of formal institutions in resilience building. Climatic Change 94(1–2): 211–232. Badjeck, M.-C., Allison, E.H., Halls, A.S., and Dulvy, N.K. 2010. Impacts of climate variability and change on fishing-based livelihoods. Marine Policy 34(3):375–383. Barange, M., and Perry, I. 2009. Physical and ecological impacts of climate change relevant to marine and inland capture fisheries and aquaculture. In: Cochrane, K., De Young, C., Soto, D., and Bahri, T. (eds.), Climate Change Implications for Fisheries and Aquaculture. Overview of Current Scientific Knowledge. Food and Agriculture Organization of the United Nations, Rome, pp. 7–106. Baruthio, A., Aubin, J., Mungkung, R.L.J., and van der Werf, H.M. 2009. Environmental assessment of Filipino fish/prawn polyculture using Life Cycle Assessment. In: Nemecek, T., and Gaillard, G. (eds.), Proceedings of the 6th International Conference on LCA in the Agri-Food Sector—Towards A Sustainable Management of the Food Chain. November 12–14, 2008, Zürich, Switzerland, pp. 242–247. Behrenfeld, M.J., O’Malley, R.T., Siegel, D.A., McClain, C.R., Sarmiento, J.L., Feldman, G.C.,

Milligan, A.J., Falkowski, P.G., Letelier, R.M., and Boss, E.S. 2006. Climate-driven trends in contemporary ocean productivity. Nature 444:752–755. PG(2046; Bell, Batty, Ganachaud, Gehrke, Hobday, Hoegh-Guldberg, Johnson, Le Borgne, Lehodey, Lough, Pickering, Pratchett, Sheaves, Waycott; 2010)Bell, J., Batty, M., Ganachaud, A., Gehrke, P., Hobday, A., HoeghGuldberg, O., Johnson, J., Le Borgne, R., Lehodey, P., Lough, J., Pickering, T., Pratchett, M., Sheaves, M., and Waycott, M. 2010. Preliminary assessment of the effects of climate change on fisheries and aquaculture in the Pacific. In: The Contribution of Fisheries to the Economies of Pacific Island Countries and Territories. Pacific Studies Series, Asian Development Bank, Manila, Philippines. Berge, J.A., Bjerkeng, B., Pettersen, O., Schaanning, M.T., and Oxnevad, S. 2006. Effects of increased seawater concentrations of CO2 on growth of the bivalve Mytilus edulis L. Chemosphere 62(4):681–687. Biershenck, E. 2004. Storm damages shock local shellfish industry. Victoria, Australia, GROWfish, Gippsland Aquaculture Industry NetworkGAIN (17/09/2004). Bosma, R.H., Hanh, C.T.T., and Potting, J. 2009. Environmental impact assessment of the pangasius sector in the Mekong Delta. Wageningen University. Brander, K.M. 2007. Global fish production and climate change. Proceedings of the National Academy of Sciences of the United States of America 104(50):19709–19714. Brierley, A.S., and Kingsford, M.J. 2009. Impacts of climate change on marine organisms and ecosystems. Current Biology 19:R602–R614. Brown, P., and Sutton, T. 2002. Global warming brings new cash crop for fishermen. The Guardian (10 December 2002), London. Bunting, S.W., and Pretty, J. 2007. Global carbon budgets and aquaculture—emissions, sequestration and management options. Centre for Environment and Society Occasional Paper 2007-1. University of Essex, UK. Cane, M.A. 2005. The evolution of El Nino, past and future. Earth and Planetary Science Letters 230:227–240. Chauvaud, L., Thompson, J.K., Cloern, J.E., and Thouzeau, G. 2003. Clams as CO2 generators:

Implications of global climate change

the Potamocorbula amurensis example in San Francisco Bay. Limnology and Oceanography 48:2086–2092. Chen, Y.L. 2008. The adaptive strategies of oyster farmers to climate variation: typhoon in Tainan, Taiwan. Masters thesis, Institute of Marine Aquaculture, University of Taiwan, 121 pp (in Chinese—English abstract accessed via etd.lib. nsysu.edu.tw/ETD-db/ETD-search/view_etd?UR N=etd-0907109-173345 Cheung, W.W.L., Lam, V.W.Y., Sarmiento, J.L., Kearney, K., Watson, R., and Pauly, D. 2009. Projecting global marine biodiversity impacts under climate change scenarios. Fish and Fisheries 10(3):235–251. Compton, T.J., Rijkenberg, M.J.A., Drent, J., and Piersma, T. 2007. Thermal tolerance ranges and climate variability: a comparison between bivalves from differing climates. Journal of Experimental Marine Biology and Ecology. 352:200–211. Cook, T., Folli, M., Klinck, J., Ford, S., and Miller, J. 1998. The relationship between increasing sea-surface temperature and the northward spread of perkinsus marinus (dermo) disease epizootics in oysters. Estuarine, Coastal and Shelf Science 46:587–597. De Silva, S.S., and Soto, D. 2009. Climate change and aquaculture: potential impacts, adaptation and mitigation. In: Cochrane, K., De Young, C., Soto, D., and Bahri, T. (eds.), Climate Change Implications for Fisheries and Aquaculture. Overview of Current Scientific Knowledge. Food and Agriculture Organization of the United Nations, Rome, pp. 151–212. de Young, B., Barange, M., Beaugrand, G., Harris, R., Perry, R.I., Scheffer, M., and Werner, F. 2008. Regime shifts in marine ecosystems: detection, prediction and management. Trends in Ecology and Evolution 23(7):402–409. Diederich, S., Nehls, G., van Beusekom, J.E.E., and Reise, K. 2010. Introduced Pacific oysters (Crassostrea gigas) in the northern Wadden Sea: invasion accelerated by warm summers? Helgoland Marine Research 59(2): 97–106. Doney, S.C. 2006. Plankton in a warmer world. Nature 444:695–696. Doney, S.C., Fabry, V.J., Feely, R.A., and Kleypas, J.A. 2009. Ocean acidification: the other CO2

487

problem. Annual Review of Marine Science 1:169–192. d’Orbcastel, E.R., Blancheton, J.-P., and Aubin, J. 2009. Towards environmentally sustainable aquaculture: comparison between two trout farming systems using life cycle assessment. Aquacultural Engineering 40:113–119. Dunlap, R.E., and McCright, A.M. 2008. A widening gap: republican and democratic views on climate change. Environment 50: 26–35. Dupont, S., Dorey, N., and Thorndyke, M. 2010. What can meta-analysis tell us about vulnerability of marine biodiversity to ocean acidification? Estuarine, Coastal and Shelf Science 89(2): 182–185. Easterling, W.E., Rosenberg, N.J., McKenney, M.S., and Allan Jones, C. 1992. An introduction to the methodology, the region of study, and a historical analog of climate change. Agricultural and Forest Meteorology 59(1–2): 3–15. Emanuel, K., Sundararajan, R., and Williams, J. 2008. Hurricanes and global warming; results from downscaling IPCC AR4 simulations. Bulletin of the American Metrological Society 89(3):347–367. ESCAP and UNISDR. 2010. Protecting development gains, reducing disaster vulnerability and building resilience in Asia and the Pacific. The Asia-Pacific Disaster Report, 2010. Economic and Social Commission for Asia and the Pacific, United Nations International Strategy for Disaster Reduction, Bangkok, Thailand. 129 pp. Ford, J.D., Keskitalo, E.C.H., Smith, T., Pearce, T., Berrang-Ford, L., Duerden, F., and Smit, B. 2010. Case study and analogue methodologies in climate change vulnerability research. Wiley Interdisciplinary Reviews: Climate Change 1(3):374–392. Gazeau, F., Quiblier, C., Jansen, J.M., Gattuso, J.-P., Middelburg, J.J., and Heip, C.H.R. 2007. Impact of elevated CO2 on shellfish calcification. Geophysical Research Letters 34:L07603. Glantz, M. 1988. Societal Responses to Regional Climatic Change: Forecasting by Analogy. Westview Press, Boulder, CO. Grafton, R.Q. 2010. Adaptation to climate change in marine capture fisheries. Marine Policy 34:606–615.

488

Shellfish Aquaculture and the Environment

Hallegraeff, G.M. 2010. Ocean climate change, phytoplankton community responses and harmful algal blooms; a formidable predictive challenge. Journal of Phycology 46(2):220– 235. Handisyde, N.T., Ross, L.G., Badjeck, M.-C., and Allison, E.H. 2006. The effects of climate change on world aquaculture: a global perspective. Final Technical Report. DFID Aquaculture and Fish Genetics Research Programme, Stirling Institute of Aquaculture, Stirling, U.K., 151 pp. www. aqua.stir.ac.uk/GISAP/climate/index.htm Harley, C.D.G., Hughes, A.R., Hultgren, K.M., Miner, B.G., Sorte, C.J.B., Thornber, C.S., Rodriguez, L.F., Tomanek, L., and Williams, S.L. 2006. The impacts of climate change in coastal marine systems. Ecology Letters 9:228–241. Hendriks, I.E., and Duarte, C.M. 2010. Ocean acidification: seperating evidence from judgment— —A reply to Dupont et al. Estuarine, Coastal and Shelf Science 89(2):186–190. Hendriks, I.E., Duarte, C.M., and Álvarez, M. 2010. Vulnerability of marine biodiversity to ocean acidification: a meta-analysis. Estuarine, Coastal and Shelf Science 86(2): 157–164. Henriksson, P. 2009. Energy intensity in tropical aquaculture. Cumulative energy demand of milkfish, pangasius and oyster production in SE Asia. Master’s Thesis, Department of Systems Ecology, Stockholm University,Sweden. Hickey, J.P. 2008. Carbon sequestration potential of shellfish. Seminars on Sustainability. University of South Australia. Accessed via: www.oysterssa. com.au/media/files/755.pdf IPCC. 2007a. Climate Change 2007: The Physical Science Basis. Summary for Policymakers. Intergovernmental Panel on Climate Change, UNEP, Geneva. IPCC. 2007b. Climate Change 2007: Mitigation of Climate Change. Working Group III Contribution to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change. UNEP, Geneva. Kaladharan, P., Veena, S., and Vivekanandan, E. 2009. Carbon sequestration by a few marine algae: observation and projection. Journal of the Marine Biological Association of India 51:107–110.

Kim, Y., Powell, E.N., Wade, T.L., Presley, B.J., and Brooks, J.M. 1999. Influence of climate change on interannual variation in contaminant body burden in Gulf of Mexico oysters. Marine Environmental Research 48(4–5):459–488. Kumar, A., Ergas, S., Yuan, X., Sahu, A., Zhang, Q., Dewulf, J., Malcata, F.X., and van Langenhove, H. 2010. Enhanced CO2 fixation and biofuel production via microalgae: recent developments and future directions. Trends in Biotechnology 28(7):371–380. Kurlansky, M. 2006. The Big Oyster: History on the Half Shell. Random House, New York. Laffoley, D.d’A., and Grimsditch, G. 2009. The Management of Natural Coastal Carbon Sinks. IUCN, Gland, Switzerland. Lazarus, R.J. 2009. Super wicked problems and climate change: restraining the present to liberate the future. Cornell Law Review 94:1153– 1234. Lorenzoni, I., and Pidgeon, N. 2006. Public views on climate change: european and USA perspectives. Climatic Change 77:73–95. Marcogliese, D.J. 2008. The impact of climate change on the parasites and infectious diseases of aquatic animals. Revue scientifique et technique 27(2):467–484. McCarthy, J., Canziani, O.S., Learly, N., Dokken, D., and White, K. (eds.). 2001. Climate Change 2001: Impacts, Adaptation, and Vulnerability. Cambridge University Press, Cambridge. McKenna, J., Cooper, J.A.G., and O’Hagan, A.M. 2009. Coastal erosion management and the European principles of ICZM: local versus strategic perspectives. Journal of Coastal Conservation 13(2–3):165–173. Miller, A.W., Reynolds, A.C., Sobrino, C., and Riedel, G.F. 2009. Shellfish face uncertain future in high CO2 world: influence of acidification on oyster larvae calcification and growth in estuaries. Public Library of Science One 4(5):e5661. doi:10.1371/journal.pone.0005661 Moore, S.K., Trainer, V.L., Mantua, N.J., Parker, M.S., Laws, E.A., Backer, L.C., and Fleming, L.E. 2008. Impacts of climate variability and future climate change on harmful algal blooms and human health. Environmental Health 7(Suppl. 2):S4. Nell, J.A. 2001. The history of oyster farming in Australia. Marine Fisheries Review 3:14–25.

Implications of global climate change

Nellemann, C., Corcoran, E., Duarte, C.M., Valdes, L., DeYoung, C., Fonseca, L., and Grimsditch, G. (eds.). 2009. Blue Carbon—The Role of Healthy Oceans in Binding Carbon—A Rapid Response Assessment. UNEP, Nairobi, Kenya. Oliver, L.R., Seed, R., and Reynolds, B. 2008. The effect of high flow events on mussels (Mytilus edulis) in the Conway estuary, North Wales, UK. Hydrobiologia 597(1):117–127. Paolisso, M., Secor, D., Sellner, K., Wardrop, D., and Wood, R. 2010. Potential climate-change impacts on the Chesapeake Bay. Estuarine, Coastal and Shelf Science 86(1):1–20. Parker, L.M., Ross, P.M., and O’Connor, W.A. 2009. The effect of ocean acidification and temperature on the fertilization and embryonic development of the Sydney rock oyster Saccostrea glomerata (Gould 1850). Global Change Biology 15:2123–2136. Patz, J.A., Olson, S.H., and Gray, A.L. 2006. Climate change, oceans, and human health. Oceanography 19(2):52–59. Pelletier, N., and Tyedmers, P. 2010. Life cycle assessment of frozen tilapia fillets from Indonesian lake-based and pond-based intensive aquaculture systems. Journal of Industrial Ecology. 14(3):467–481. Pelletier, N., Tyedmers, P., Sonesson, U., Scholz, A., Ziegler, F., Flysjo, A., Kruse, S., Cancino, B., and Silverman, H. 2009. Not all salmon are created equal: life cycle assessment (LCA) of global salmon farming systems. Environmental Science & Technology 43:8730–8736. Peperzak, L. 2003. Climate change and harmful algal blooms in the North Sea. Acta Oecologica 24(S1):S139–S144. Philippart, C.J.M., van Aken, H.M., Beukema, J.J., Bos, O.G., Cadée, G.C., and Dekker, R. 2003. Climate-related changes in recruitment of the bivalve Macoma balthica. Limnology & Oceanography 48(6):2171–2185. Preston, B., and Stafford-Smith, M. 2009. Framing vulnerability and adaptive capacity assessment: discussion paper. CSIRO Climate Adaptation Flagship Working Group Paper No 2, CSIRO, Canberra. www.csiro.au/org/climateadaptationflagship.html Produce. 2010a. Peru: Valor de la exportacion de los productos hidrobiologicos procedentes de la

489

actividad de acuicultura por especie segun pais de destino 2000–2009. www.produce.gob.pe/ portal/portal/apsportalproduce/internapesqueria ?ARE=3&JER=460 (accessed on September 12, 2010). PRODUCE, Lima, Peru. Produce. 2010b. National plan for aquaculture development—PNDA. General Aquaculture Directorate Deputy Minister’s Office for Fisheries Ministry of Production, Lima, Peru. Rahmstorf, S., Cazenave, A., Church, J.A., Hansen, J.E., Keeling, R.F., Parker, D.E., and Sommervile, R.C.J. 2007. Recent climate observations compared to projections. Science 316(5825):709. Rodhouse, P.G., and Roden, C.M. 1987. Carbon budget for a coastal inlet in relation to intensive cultivation of suspension-feeding bivalve molluscs. Marine Ecology Progress Series 36: 225–236. Rose, J.B., Epstein, P.R., Lipp, E.K., Sherman, B.H., Bernard, S.M., and Patz, J.M. 2001. Climate variability and change in the United States: potential impacts on water- and foodborne diseases caused by microbiologic agents. Environmental Health Perspectives 109(2):211–220. Shephard, S., Beukers-Stewart, B., Hiddink, J., Brand, A., and Kaiser, M. 2010. Strengthening recruitment of exploited scallops Pecten maximus with ocean warming. Marine Biology 157(1): 91–97. Soniat, T.M., Klinck, J.M., Powell, E.N., and Hofmann, E.E. 2006. Understanding the success and failure of oyster populations: climatic cycles and Perkinsus marinus. Journal of Shellfish Research 25(1):83–93. Talmage, S.C., and Gobler, C.J. 2010. Effects of past, present, and future ocean carbon dioxide concentrations on the growth and survival of larval shellfish. Proceedings of the National Academy of Sciences of the United States of America 107(40):17246–17251. Tlusty, M.F., and Lagueux, K. 2009. Isolines as a new tool to assess the energy costs of the production and distribution of multiple sources of seafood. The sustainability of seafood production and consumption. Journal of Cleaner Production 17:408–415. Troell, M., Tyedmers, P., Kautsky, N., and Rönnbäck, P. 2004. Aquaculture and energy use. In: Cleveland, C.J. (ed.), Encyclopedia of Energy. Elsevier, New York, pp. 97–108.

490

Shellfish Aquaculture and the Environment

Tyedmers, P., and Pelletier, N. 2007. Biophysical accounting in aquaculture: insights from current practice and the need for methodological development. In: Bartley, D.M., Brugère, C., Soto, D., Gerber, P., and Harvey, B. (eds.), Comparative Assessment of the Environmental Costs of Aquaculture and Other Food Production Sectors. Methods for Meaningful Comparisons. FAO/ WFT Expert Workshop, 24–28 April 2006, Vancouver, Canada. Food and Agriculture Organization of the United Nations, Rome. UNEP. 2006. Marine and coastal ecosystems and human well-being. A synthesis report based on the findings of the Millennium Ecosystem Assessment. United National Environment Programme, Nairobi, Kenya. UNFCCC. 2009. Copenhagen accord. Advance unedited version. Decision -/CP.15 18 December 2009. unfccc.int/files/meetings/cop_15/application/pdf/cop15_cph_auv.pdf. United Nations Framework Convention on Climate Change. (accessed on Octorber 31, 2010). USDA. 2010. Cultivated clam pilot crop insurance underwriting guide. 2010 and succeeding crop years FCIC-24100 (08-2010). United States Department of Agriculture, Federal Crop Insurance Corporation, Product Development Division, Kansas City. 36 pp. van Anrooy, R., Secretan, P.A.D., Lou, Y., Roberts, R., and Upare, M. 2006. Review of the current state of world aquaculture insurance. FAO

Fisheries Technical Paper. No. 493. FAO, Rome. 92 pp. Vaquer-Sunyer, R., and Duarte, C.M. 2008. Thresholds of hypoxia for marine biodiversity. Proceedings of the National Academy of Sciences of the United States of America 105(40): 15452–15457. Vilchis, L.I., Tegner, M.J., Moore, J.D., Friedman, C.S., Riser, K.L., Robbins, T.T., and Dayton, P.K. 2005. Ocean warming effects on growth, reproduction, and survivorship of southern California abalone. Ecological Applications 15(2):469– 480. Waldbusser, G.G., Voigt, E.P., Bergschneider, H., Green, M.A., and Newell, R.I.E. 2011. Biocalcification in the Eastern Oyster (Crassostrea virginica) in relation to long-term trends in Chesapeake Bay pH. Estuaries and Coasts 34(2):221–231. Wolff, M., and Mendo, J. 2000. Management of the Peruvian bay scallop metapopulation with regard to environmental change. Aquatic Conservation Marine Freshwater Ecosystems 10:117–126. Yeoman, I., and McMahon-Beattie, U. 2006. Understanding the impact of climate change on Scottish tourism. Journal of Vacation Marketing 12(4):371–379.

Index Note: Page numbers in italics refer to figures, those in bold to tables. abalone, 365, 472 abalone viral mortality, 373 abundance of shellfish, changes in as result of climate change, 466 acid sulfate soils, and shellfish diseases, 366–367 acidification of ocean as result of climate change, 465, 467, 469, 470, 474, 474, 477 and shellfish diseases, 367 adaptedness, effect of hatcheries on, 350 additive variance, 344, 346 Aequipecten opercularis (queen scallop), 400 Aeromonas sp., 379 agricultural environmental aid programs, 224–225, 225 agro-aqua recycling, 197, 197, 218, 218 Alexandrium catenella, 404 Alexandrium minutum (red tide dinoflagellate), 404, 408 algae impact on bivalve aquaculture, 408 introduced from bivalve aquaculture, 404 algal growth. See harmful algal blooms algology, 448 Alteromonas sp., 379 amethyst gem clam (Gemma gemma), 405 ammonia flux, and eutrophication, 177–178 Anadara demirii (arcid clam), 409 Anadara granosa (blood cockle), 342, 342, 343, 399 Anadara inaequivalvis, 410 animal health/welfare, in aquaculture certification programs, 68 annelids impact on bivalve aquaculture, 408 introduced from bivalve aquaculture, 404 anoxia. See oxygen conditions APP (average physical product), 14, 15

Aquaculture Stewardship Council, 414, 441 aquaculture-related disciplines, 449–450, 451 Aquatic Animal Health Code, 373, 376 Arcidae, 399 arcid clam (Anadara demirii), 409 Arctic char, 483 Arctica islandica (ocean quahog), 242, 322 Argentine mussel (Mytilus edulis platensis), 90 Argopecten gibbus (calico scallop), 243 Argopecten irradians (Atlantic bay scallop), 95, 108, 327, 397, 400, 470 Argopecten irradians concentricus, 166 Argopecten nucleus, 95 Argopecten purpuratus (Peruvian calico scallop), 89, 95, 166, 400, 476 Argopecten ventricosus (Pacific calico scallop), 400 Argopecten ventricosus-circularis, 95 ASC (Aquaculture Stewardship Council), 414, 441 ascidians, 407, 412 Ascidiella aspersa (European sea squirt), 405, 411 Asian hornsnail (Batillaria attramentaria), 405 Asian kelp (Undaria pinnatifida), 404 Asian mussel (Musculista senhousia), 165, 168, 181, 405 Asian oyster. See Crassostrea ariakensis Asian rapa whelk (Rapana venosa), 405, 410 Asian tunicate (Styela clava), 405, 411 Asiatic cholera (Vibrio cholerae), 404, 409 Asiatic hard clam (Meretrix meretrix), 401 Assessment of Estuarine Trophic Status model, 7, 8, 9, 20–21, 20, 147 ASSETS model. See Assessment of Estuarine Trophic Status model

Shellfish Aquaculture and the Environment, First Edition. Edited by Sandra E. Shumway. © 2011 John Wiley & Sons, Inc. Published 2011 by John Wiley & Sons, Inc. 491

492

Index

assurance labeling, 61–62, 64–65, 70, 72, 73, 78 Aquaculture Stewardship Council, 414, 441 as barrier to trade, 73–74 beneficiaries of, 73 comprehensive quality approach, 75–76, 76 and consumers, 67–68 FAO Guidelines for Aquaculture Certification, 68–70, 71 GAA/Aquaculture Certification Council Best Aquaculture Practices standard, 70, 75 and information asymmetry, 74 and life cycle assessments, 76–77, 77 organic standards, 71–73, 72 pressures to participate, 65–67 and producers, 67, 73–74 proliferation of ecolabels, 68, 74–75, 78 and standards, 65 third-party certification programs, 46–48, 64–65 and traceability, 65, 69 Type I/II/III ecolabels, 64–65 World Wildlife Fund Bivalve Aquaculture Dialogue, 47, 70–71, 75, 413–414, 441 Asterias amurensis (flatbottom sea star), 404, 409 Asterocarpa humilis, 411 Atlantic bay scallop. See Argopecten irradians Atlantic rangia (Rangia cuneata), 405 Atlantic salmon, 342, 343, 483 Atlantic slipper snail (Crepidula fornicata), 405, 406, 407, 409 Atlantic surf clam (Spisula solidissima), 242, 322, 325, 399 Atrina sp., 242 Atrina zelandica, 106, 108 Aulacomya ater (cholga mussel), 89, 93, 399 Aulorhynchus flavidus (tubesnout), 256 Australian mussel (Mytilus planulatis), 169, 399 Australian southern scallop (Pecten fumatus), 400, 408 average physical product, 14, 15 BACI (before-after control-impact) experimental design, 320, 330, 330 Bacillariophyceae, 129 bacteria impact on bivalve aquaculture, 409 introduced from bivalve aquaculture, 404 Baltic clam. See Macoma balthica Baltic Sea, mussel farming in, 198, 221–222 Batillaria attramentaria (Asian hornsnail), 405 Belfast Lough, crop rotation in, 6, 6 benthic community composition, and biodeposition, 309–310

benthic nutrient fluxes, and bioturbation, 308–309 best husbandry practices, 55 best management practices, 40, 42, 43, 44, 45–46, 46, 48, 51, 52–53, 54–55, 56–57, 77–78, 272, 440, 442 and assurance labeling programs, 62 for carrying capacity, 46 and codes of conduct, 55, 58, 59, 441 and compliance monitoring, 60, 61 and continuous improvement, 55 for culture practice, 44 and diminishing returns, 55 environmental, 44, 45–46, 55, 440–442 and environmental management systems, 55, 58, 59 government-facilitated, 440 incentive programs, 52–54, 441–442 industry-driven, 440 limitations of, 62–64 and nutrient reduction, 46 performance indicators, 60 performance standards, 58, 63 and producers, 62 and product marketing, 61–62 and regulators, 62 and regulatory requirements, 61 and research scientists, 62 and seafood buyers, 62 state programs, 440 watershed protection, 53, 54–55 and zoning, 58–60 biodeposition, 18–19, 18, 19, 189, 254, 259, 260, 311–312 and benthic community composition, 309–310 and integrated multitrophic aquaculture, 23–24, 24 methodology for clearance rate, 101, 102 and organic enrichment, 307, 307 and oxygen conditions, 308–309 and sediment biogeochemistry, 308 and soft sediment habitats, 306–310 and water flow, 306–307, 307 biodiversity, 347–352, 426–427 biofuel, microalgae as, 130 biogas, mussel remainder as, 232–233 biogeochemical models of carrying capacity, 136 bioirrigation, 297, 298, 299, 299, 302, 311. See also bioturbation biological sciences, as aquaculture-related discipline, 450

Index

biomass production, 13–17, 13, 15, 16 production enhancement using multiple species, 17, 17 profit optimization, 14–16, 14, 15 biosecurity, 371–372 and compartmentalization, 375–376 geographic, 374–376 and zoning, 375–376 bioturbation, 297, 298, 299, 299, 300–303, 301, 302, 304, 311–312 and benthic nutrient fluxes, 308–309 and contaminants, 310–311 and oxygen conditions, 299, 300, 308–309 and resting stages in sediments, 311 and seagrasses, 310 and structural heterogeneity of soft sediments, 303–306 Bivalve Aquaculture Dialogue (World Wildlife Fund), 47, 70–71, 75, 413–414, 441 bivalve ecophysiology models of carrying capacity, 136 bivalve species grown in aquaculture, 399–403 black mussel (Choromytilus meridionalis), 89, 93 black-lip pearl oyster. See Pinctada margaritifera black-striped mussel (Mytilopsis sallei), 405, 410 blood cockle (Anadara granosa), 342, 342, 343, 399 blue mussel. See Mytilus edulis BMPs. See best management practices Boccardia proboscidea, 404, 408 body size and clearance rate, 87–88, 89–91, 91–92, 94, 96, 97–99 and feeding activity, 83, 84 Bonamia exitiosa, 373, 410 Bonamia ostreae, 368–369, 373, 385, 410 Bonamia sp., 361, 363–364, 369, 396 bonamiosis, 369 Boonea bisuturalis (two-groove odostome), 405 Botrylloides violaceus (orange sheath tunicate), 411 Botryllus schlosseri (golden star tunicate), 411 box models of carrying capacity, 142–143, 143 branching bryozoan (Schizoporella errata), 404, 409 broodstock management, 378 brown bryozoan (Bugula neritina), 404, 409 brown mussel. See Perna perna brown ring disease, 369–370 bryozoans impact on bivalve aquaculture, 409 introduced from bivalve aquaculture, 404 Bugula neritina (brown bryozoan), 404, 409 burrowing shrimp, 256, 257 butter clam (Saxidomus gigantean), 251, 401

493

calico scallop (Argopecten gibbus), 243 Calidris minutilla (least sandpiper), 247 California mussel (Mytilus californianus), 84, 94, 399, 410 Cancer productus (rock crab), 256 Candidatus Xenohaliotis californiensi, 364, 373 Capitella capitata, 175, 176 Capitellidae, 175 carbon sequestration, and aquaculture, 484 Carcinus maenas (green crab), 407, 409 Cardiidae, 399 Cardium edule, 83, 106 carp, 342 carpet tunicate (Didemnum sp.), 411 carrying capacity, 11, 13, 70, 128–129, 201 best management practices for, 46 ecological, 52, 82, 129, 136–137, 137 economic, 136, 137 exploitation, 13 modeling of, 135–150, 137, 138, 143, 144, 145, 146, 147, 148, 188, 189 physical, 128–129 production, 13, 81, 129, 135–149, 137, 138, 143, 144, 145, 146, 147, 148, 150 and seston depletion, 135–136, 138, 140 social, 129 catch crops, and reduction in eutrophication, 224, 225, 225 catfish, 342 catla, 342 Cerastoderma edule (common edible cockle), 84, 88, 89, 95, 98, 106, 108, 190, 305, 310, 326, 399 certification programs. See assurance labeling chain-forming dinoflagellate (Gymnodinium catenatum), 404, 408 Charadrius melodus (piping plover), 247 Charadrius semipalmatus (semipalmated plover), 247 chemistry, as aquaculture-related discipline, 450 Cherrystone Inlet (Chesapeake Bay, U.S.), 181–183, 182, 188, 189 Chesapeake Bay (U.S.), 24, 25, 181–183, 182, 188, 189, 273, 369, 413, 469, 477 Chilean flat oyster (Ostrea chilensis), 90, 95, 401 Chilean mussel. See Mytilus chilensis Chinese mitten crab, 342 Chinese scallop. See Chlamys farreri Chinook salmon (Oncorhynchus tshawytscha), 256 Chl a. See chlorophyll a Chlamys farreri (Chinese scallop; Farrer’s scallop), 19, 95, 99, 108, 148, 149, 166, 190, 383, 400 Chlamys hastata, 95 Chlamys islandica (Iceland scallop), 89, 95, 400

494

Index

Chlamys nobilis (noble scallop), 95, 400 Chlamys opercularis, 95 chlorophyll a, 7, 10, 17, 87, 105, 159, 160, 178, 184, 190, 191, 195, 196, 261, 264, 265, 265, 266, 268 cholga mussel (Aulacomya ater), 89, 93, 399 choro mussel (Choromytilus chorus), 164, 173, 399 Choromytilus chorus (choro mussel), 164, 173, 399 Choromytilus meridionalis (black mussel), 89, 93 Ciona intestinalis, 226, 411 circulation/coastal upwelling changes, as result of climate change, 464, 474 Cirratulidae, 175 clam aquaculture, 127 and eutrophication, 168, 170–174, 177 and increased CO2 concentration/ocean acidification, 470 in tidal creeks (southeastern U.S.), 260–268, 262, 263, 265, 267, 269, 270, 270, 271 on West Coast (U.S.), 250, 251 clam dredges, 323 clams, nutrient excretion rates, 163 Clean Water Act, 37, 433 clearance rate, 82, 83–84, 111 biodeposition methodology, 101, 102 and body size, 87–88, 89–91, 91–92, 94, 96, 97–99 controls on, 104–107, 104, 105, 108, 109 and current speed, 84 and exhalent siphon area, 86–87 and flow conditions, 84 methodologies for, 100–104, 101 populations, measurement of, 100 precision/accuracy of measurements, 99–104, 101 rate standardization/allometries, 87–88, 89–91, 91–92 and salinity, 84 and seston, 105–107, 105, 108 species groups, responses for, 92, 93–94, 94, 95–96, 96–99, 97, 98 and temperature, 84, 104–105, 104 temporal variability in, 85–87, 86, 88 and valve gape, 86–87 climate analogues, 469 climate change, 461–462, 484–485 and abundance of shellfish, 466 and acidification of ocean, 465, 467, 469, 470, 474, 474, 477 adapting shellfish farming to impacts of, 478–482, 479 and circulation changes, 464, 474 and climate variability patterns, 464–465

and coastal upwelling changes, 464, 474 and diseases, 466, 471, 474–475, 475 and distributions of species, 466, 471, 477 and extreme climate events, 465, 473, 478 and heat content/temperature changes in oceans/ coastal zones, 462–463, 472, 473, 474, 475– 477, 476 and hypoxia, 473 and land-ocean exchanges, 464 mitigation of, 482, 483, 484 and oceans, 462–467, 463 and phenomenological changes, 466 and physiological processes, 465 and primary production, 465–466, 473 and regime shifts, 466–467 and salinity of oceans, 463 and sea level rise, 464 and shellfish aquaculture systems, 467–469, 468, 470, 471, 472–474, 474–478, 475, 476 and spatfal, 472 and species invasions, 415, 466, 471 climate variability pattern changes, as result of climate change, 464–465 cnidarians, impact on bivalve aquaculture, 409 coastal upwelling changes, as result of climate change, 464, 474 Coastal Zone Management Act, 39, 41, 431 coastal zones, and land use conflicts, 37–42, 40, 41 Cobb-Douglas production function, 14, 15 cockle. See Cerastoderma edule codes of conduct, 55, 58, 59, 441 Codium fragile sp. (dead man’s fingers), 404, 408 Collective Research on Aquaculture Biofouling (CRAB), 414–415 commercial aquaculture, 449 commercial crossbreeding, 344–347, 346 common edible cockle. See Cerastoderma edule community shellfish gardens, 36 compartmentalization, for establishing biosecurity, 375–376 compliance monitoring, and best management practices, 60, 61 comprehensive quality approach to assurance labeling, 75–76, 76 constricted tagelus (Sinovacula constricta), 342, 342, 343, 401 contaminants, and bioturbation, 310–311 continuous improvement, and best management practices, 55 Corbiculidae, 242 corporate social responsibility, 67, 69–70 Cortez oyster (Crassostrea corteziensis), 400

Index

Coscinodiscus wailesii, 404 Council on Environmental Quality Interagency Ocean Policy Task Force, 42, 439 CRAB (Collective Research on Aquaculture Biofouling), 414–415 crab dredges, 322 Crassostrea sp., 242 Crassostrea angulata, 409, 412 Crassostrea ariakensis (Asian oyster), 54, 273, 348, 363–364, 369, 411, 413, 470 Crassostrea belcheri (lugubrious cupped oyster), 95, 400 Crassostrea corteziensis (Cortez oyster), 400 Crassostrea gigas (Pacific oyster), 10, 12, 15, 15, 16, 19, 21, 21, 24, 25, 37, 88, 89, 95, 106, 108, 162–163, 165–166, 169, 170, 171, 173, 174, 183, 190, 191, 243, 249, 250, 251, 252, 253, 256–257, 258, 260, 272, 304, 342, 342, 343, 343, 344, 345, 347–348, 361, 367–368, 380, 381, 383, 384, 384, 385, 397, 398, 400, 402, 405, 406, 407, 408–412, 413, 470, 471, 479 Crassostrea iradelei (slipper-cupped oyster), 95, 400 Crassostrea madrasensis (Indian backwater oyster), 400 Crassostrea rhizophorae (mangrove cupped oyster), 401 Crassostrea rivularis (Suminoe oyster), 401 Crassostrea sikamea (Kumamoto oyster), 348, 410, 412 Crassostrea virginica (eastern oyster), 24, 25, 95, 110, 165, 172, 195, 240, 241, 244, 245, 246, 247, 248, 249, 250, 257, 258, 261, 270, 274, 304, 310, 347, 348, 361–363, 362, 363, 365, 366, 369, 376, 380, 380, 397–398, 401, 402, 405, 406, 409–411, 470, 471, 472 credence attributes/goods, 68, 74, 75 Crepidula fornicata (Atlantic slipper snail), 405, 406, 407, 409 crop insurance, 479 crossbreeding, commercial, 344–347, 346 crustaceans impact on bivalve aquaculture, 409 introduced from bivalve aquaculture, 404 CSR (corporate social responsibility), 67, 69–70 culture practice, 6–7, 6, 44 current speed and clearance rate, 84 in tidal creek clam aquaculture (southeastern U.S.), 266–268, 267, 269 CWA (Clean Water Act), 37, 433 Cyclope neritea, 405 cyprinids, 342

495

CZMA (Coastal Zone Management Act), 39, 41, 431 dead man’s fingers (Codium fragile sp.), 404, 408 debris from shellfish operations, cleanup of, 41, 41 denitrification. See nitrification-denitrification DEPOMOD model, 190 dermo. See Perkinsus marinus dermo disease, 365–366, 365 diarrheic shellfish toxin (DST), 228, 231–232 Didemnum sp. (carpet tunicate), 411 diminishing returns, and best management practices, 55 Diplosoma listerianum, 411 diseases of shellfish, 359–361, 360, 367–370, 382– 385, 383, 412 abalone viral mortality, 373 and acid sulfate soils, 366–367 and acidification of ocean, 367 bonamiosis, 369 brown ring disease, 369–370 and climate change, 466, 471, 474–475, 475 dermo, 365–366, 365 diarrheic shellfish toxin (DST), 228, 231–232 disease agents, introduction of, 367–369 and estuarine flushing rates, 367 and freshwater input, 366 geographic distribution of parasites, 364 governmental/intergovernmental systems for control of, 372–373 in hatcheries, management of, 379–380 hinge ligament disease, 381–382 impacts of, assessing, 374 iridolike viruses, 412 iridoviruses, 379 juvenile oyster disease (proteobacteria), 409 MSX (Haplosporidium nelsoni), 347–348, 361–363, 362, 363, 367–368, 410 native host, introduction of, 369–370 in natural environment, 360–367, 360, 362, 363, 365 in nurseries, management of, 380–382, 380, 381 oyster herpes virus (OsHV) disease, 365, 379–380, 412 oyster velar virus disease (OVVD), 379, 412 protozoan infections, 380, 382 QPX, 364, 382–383, 383 reportable diseases, 370–371, 373 restoration of populations after disease, 350–351 Roseovarius oyster disease, 380–381, 380 and salinity, 361–364, 363 summer mortality syndrome, 383–385, 384, 385

496

Index

and temperature, 361, 362, 363, 364–366 vibriosis, 364–365, 381, 381 and water circulation patterns, 367 withering syndrome, 364, 365 See also health management of shellfish distributions of species, changes in as result of climate change, 466, 471, 477 diversity genetic, effect of hatcheries on, 350, 427 and mechanical harvesting, 321, 324–325 domestication, and shellfish genetics, 341–344, 342, 343, 344, 345 Donax serra, 163 Donax sordidus, 163 dredges, 320, 321, 322–323, 322, 323, 324 DST (diarrheic shellfish toxin), 228, 231–232 Dungeness crab (Metacarcinus magister), 256, 257 East Coast Shellfish Growers Association, 440 eastern oyster. See Crassostrea virginica eastern soft-shell clam. See Mya arenaria Echinocardium cordatum, 176 echinoderms impact on bivalve aquaculture, 409 introduced from bivalve aquaculture, 404 ecolabeling programs. See assurance labeling ecological carrying capacity, 52, 82, 129, 136–137, 137 economic carrying capacity, 136, 137 Ecopath, 142 ECOPs (environmental codes of practice). See best management practices ecosystem change, and shellfish aquaculture, 53 ecosystem-based management, 150 EcoWin2000, 144, 145 ECSGA (East Coast Shellfish Growers Association), 440 edible (flat) oyster. See Ostrea lurida education, 447, 458–459 algology, 448 aquaculture-related disciplines, 449–450, 451 extension programs, 455–457, 455 4-H programs, 454–455 graduate degree programs, 453–454 growout phase, skills required in, 449, 450 hatchery phase, skills required in, 447–448, 448 and health management, 372 K-12, 451–452, 452 and land use conflicts, 40 nursery phase, skills required in, 448–449 setting phase, skills required in, 448, 449

technology transfer, 457–458, 458 undergraduate degree programs, 452–453 eelgrass (Zostera sp.), 19–20, 251, 252, 254–256, 257, 258, 327, 427 EFH (essential fish habitat), and shellfish beds, 40 El Niño Southern Oscillation, 462, 464–465, 472 engineering, as aquaculture-related discipline, 450, 451 English sole (Parophrys vetulis), 256, 257 Ensis sp. (razor clam), 326, 342 ENSO (El Niño Southern Oscillation), 462, 464–465, 472 Enteromorpha, 21 environmental codes of practice. See best management practices environmental impacts of shellfish aquaculture, 53–54 altered biodiversity, 426–427 and far-field operations, 54, 71 genetic diversity, loss of, 427 habitat degradation, 426–427 and mechanical shellfish harvesting, 319–335, 322, 323, 324, 330, 331, 331, 332 modeling of, 149–150 and near-field operations, 54, 70–71 nonnative species, introduction of, 54, 427 environmental integrity, in aquaculture certification programs, 69 environmental management systems, 55, 58, 59 Environmental Quality Incentives Program, 441 EPA (U.S. Environmental Protection Agency), 433 EQIP (Environmental Quality Incentives Program), 441 Escherichia coli, 234 essential fish habitat, and shellfish beds, 40 estuarine flushing rates, and shellfish disease, 367 estuarine mud crab (Rhithropanopeus harrisii), 404 European flat oyster. See Ostrea edulis European sea squirt (Ascidiella aspersa), 405, 411 European Union rules for introduction of nonnative species, 413 eutrophication, 53, 128 and algal growth, 159 and ammonia flux, 177–178 from aquaculture, 155–156, 157–159, 157, 160–162, 162, 167, 168–174, 174–177, 179–187, 180, 181, 182, 185, 200–201 assessment/modeling of, 7–8, 9, 20–21, 20, 187–192, 189, 191, 196–200, 197, 199, 200 and hydrography, 156, 162, 166 from land-based activities, 155, 156–157, 157, 192–195, 193, 194, 201, 217

Index

nitrification-denitrification, 159–160, 162, 167 nutrient dynamics, 177–179, 178 nutrient excretion rates, 159, 163–166 oxygen depletion, 157, 159, 167 and phytoplankton, 128, 158, 177, 196 reduction of, 19–20, 195–200, 197, 199, 200, 217–219, 218, 221–235, 221, 223, 225, 227, 228, 230, 231, 233 in Sacca de Goro Lagoon (Italy), 183–187, 185, 186, 189, 189 and shrimp culture, 195 in Thau Lagoon (France), 183, 189 exhalent siphon area, and clearance rate, 86–87 exploitation carrying capacity, 13 extended Redfield ratio, 127–128 extension programs, 455–457, 455 extreme climate events, as result of climate change, 465, 473, 478 false mussel (Mytilopsis adamsi), 410 FAO Guidelines for Aquaculture Certification, 68–70, 71 far-field/offshore operations, 54, 71, 437–440, 437 Farm Aquaculture Resource Management model, 7–8, 11, 12, 12–13, 15, 15, 147, 190–191, 191 FARM model. See Farm Aquaculture Resource Management model farms, shellfish, 3–5, 4, 4, 5, 12, 242, 243 farm-scale models, 11–13, 11, 12 Farrer’s scallop. See Chlamys farreri feeding activity, 18, 81–82, 85, 109, 111–112 and body size, 83, 84 pumping rate, 82–83, 83, 85, 86 and seston, 81–82, 85 and size-dependent particle retention, 110–111 and transparent exopolymer particle (TEP) production, 109–110 and viscosity of water, 84 See also clearance rate; trophic interactions of phytoplankton and bivalves fertilizer, mussel remainder as, 232–234, 233 filtration, and seston depletion, 140 Fish and Wildlife Coordination Act of 1934, 433 fishing gear, environmental impact of, 320–321 flatbottom sea star (Asterias amurensis), 404, 409 flathead gray mullet, 342 flow conditions, and clearance rate, 84 Food Alliance, 47 food depletion, 20, 21 food safety/quality, 36, 66, 68–69, 71, 228, 234 food supply, and shellfish aquaculture, xiii–xiv, xv, 33–34, 34, 48

497

fouling, 407, 412, 414–415 4-H programs, 454–455 freshwater input, and shellfish diseases, 366 fully spatial models of carrying capacity, 143–145, 144, 145 Fulvia tenuicostata, 409 Fusinus rostratus, 405 FWCA (Fish and Wildlife Coordination Act) of 1934, 433 GAA/Aquaculture Certification Council Best Aquaculture Practices standard, 70, 75 gardens, shellfish, 36 Gemma gemma (amethyst gem clam), 405 genetics of shellfish, 339, 340–341 and additive variance, 344, 346 and biodiversity conservation, 347–352 commercial crossbreeding, 344–347, 346 and domestication, 341–344, 342, 343, 344, 345 and genotyping, 346 interaction of hatchery-propagated and native populations, 349–350, 351–352 and introductions of nonnative shellfish, 347–348 and nonadditive variance, 343–344, 346 rehabilitation, 350–351 and sterilization, 352 sweepstakes reproductive success, 349 translocation, impact of, 348–349 and triploidy, 346–347, 352 genotyping, 346 geoduck. See Panopea abrupta; Panopea generosa geographic biosecurity, 374–376 giant clam (Tridacna gigas), 401 Gibbula albida, 405 Global Aquaculture Alliance/Aquaculture Certification Council Best Aquaculture Practices standard, 70, 75 globose clam (Mactra veneriformis), 399 GMA (Growth Management Act; Washington State), 39 golden star tunicate (Botryllus schlosseri), 411 Gonyaulax excavata, 404, 408 goods/services from aquaculture, xiii, 5, 24, 25, 26, 427–428, 428 biomass production, 13–17, 13, 14, 15, 16, 17 economic, xiii, xiv, xv ecosystem, xiii, xv seagrasses, growth of, 254 See also eutrophication: reduction of; nitrogen: removal goods/services from native shellfish, 243–246 Gracilaria bursa-pastoris, 183

498

Index

graduate degree programs, 453–454 Grays Harbor (Washington State, U.S.), 256 great Atlantic scallop. See Pecten maximus green crab (Carcinus maenas), 407, 409 green mussel. See Perna viridis greenhouse gas emissions from aquaculture, 482, 483, 484 grooved carpet shell (Venerupis decussatus), 369, 370, 401 growout phase, skills required in, 449, 450 growout structures/gear, abandoned, 270, 270, 271 Growth Management Act (Washington State), 39 growth of aquaculture, 339–340, 340, 397, 425 grunt (Orthopristis chrysopterus), 244 Guekensia demissus, 164 Gunnarea capensis, 402 Gymnodinium catenatum (chain-forming dinoflagellate), 404, 408 habitats of shellfish, 240–243, 244, 245–246, 257–259, 270, 270, 271, 273, 326–327 degradation of, 426–427 farms, 242, 243 and mammals, 248 oyster reefs, 245–247 reef-forming shellfish, 241, 242 restoration of, 243, 244, 246–247, 248–249, 272 shell-accumulation, 241, 242–243 shell-aggregation, 241, 242 and shorebirds, 247–248 soft-sediment, 297, 298, 299, 299, 300, 301, 302, 303–310, 307 use by invertebrates/finfish, 247 HABs. See harmful algal blooms HACCP (Hazard Analysis and Critical Control Point) protocols, 42, 69 Haematopus moquini, 402 Haematopus ostralegus ostralegus, 403 half-crenate ark (Scapharca subcrenata), 399 Haliotis fulgens (green abalone), 472 Haliotis rufescens (red abalone), 365, 472 Haliotis sp., 480 Haplosporidium nelsoni (MSX), 347–348, 361–363, 362, 363, 367–368, 410 Haplosporidium sp., 361 harmful algal blooms, 13–14, 13, 130, 159, 193–194, 412, 471, 474–475 harvesting, mechanical, 319–320, 329, 334–335 differences in harvesting native and cultured shellfish, 323–324 and diversity, 321, 324–325 dredges, 320, 321, 322–323, 322, 323, 324

fishing gear, 320–321 and nutrient cycling, 321, 329 and oxygen conditions, 321, 328–329 and productivity, 321, 324–325 recovery timescales, 325–326 research on harvesting impact, 329–334, 330, 332, 332, 333 and sediment resuspension, 327–329 significance of impacts, 321–322 and submerged aquatic vegetation, 327 trawling, 320 and turbidity, 321, 327–328 and vertical structure, 321, 324, 326–327 and water chemistry, 328–329 hatcheries and adaptedness, 350 and genetic diversity, 350 hatchery-propagated and native populations, interaction of, 349–350, 351–352 management of infectious disease in, 379–380 hatchery phase, skills required in, 447–448, 448 Hazard Analysis and Critical Control Point protocols, 42, 69 health management of shellfish, 359–361, 360, 367, 377, 377–378, 384–385 biosecurity, 371–372, 374–376 broodstock management, 378 and education, 372 governmental/intergovernmental systems for disease control, 372–373 hatcheries, management of infectious disease in, 379–380 healthy farmed shellfish stocks, achieving, 370–371 and intensive aquaculture production model, 370, 371 larval and seedstocks, dissemination of, 378 and low-impact/low-prevalence microorganisms, 374 microalgae used as feed, dissemination of, 378–379 nurseries, management of infectious disease in, 380–382, 380, 381 surveillance in support of, 373–374 veterinary approach, 376–377, 377 See also diseases of shellfish heat content/temperature changes in oceans, as result of climate change, 462–463, 472, 473, 474, 475–477, 476 Heterosigma akashiwo, 159 Heterosiphonia japonica, 404 Hiatellidae, 399 higher-order models of carrying capacity, 142–143

Index

hinge ligament disease, 381–382 honeycomb worm (Sabellaria alveolata), 402 hooded oyster (Saccostrea cuccullata), 401 H2S (hydrogen sulfide), 329 hydraulic dredges, 322, 323 hydrogen sulfide (H2S), 329 hydrography, and eutrophication, 156, 162, 166 hypoxia, as result of climate change, 473 IAOP (Interagency Ocean Policy) Task Force, 42, 439 Iceland scallop (Chlamys islandica), 89, 95, 400 ICES 2005 (International Council for the Exploration of the Sea Code of Practice on the Introductions and Transfers of Marine Organisms), 414 Imogine mcgrathi, 408 IMTA. See integrated multitrophic aquaculture incentive programs, and best management practices, 52–54, 441–442 Indian backwater oyster (Crassostrea madrasensis), 400 Indian brown mussel (Perna indica), 399 individual growth, modeling, 8, 10 infauna, effects on, from clam aquaculture in tidal creeks, 264, 268 inflated ark (Scapharca broughtonii), 399 information asymmetry, and assurance labeling, 74 integrated multitrophic aquaculture, 5, 17, 17, 23–24, 24, 150, 273–274, 483 intensive aquaculture production model, 370, 371 Interagency Ocean Policy Task Force, 42, 439 International Council for the Exploration of the Sea Code of Practice on the Introductions and Transfers of Marine Organisms (ICES 2005), 414 Interstate Shellfish Sanitation Conference, 42 introductions of nonnative species. See nonnative species, introductions of iridolike viruses, 412 iridoviruses, 379 ISO 14000 standards, 55 14020, 64–65 14040, 77 ISSC (Interstate Shellfish Sanitation Conference), 42 Japanese Japanese Japanese Japanese Japanese 410 Japanese

carpet shell, 342 eel, 342 hard clam (Meretrix lusoria), 401 littleneck clam. See Tapes philippinarum oyster drill (Ocinebrellus inornatus), 405, pearl oyster (Pinctada fucata), 400

499

Japanese sea bass, 342 juvenile oyster disease (proteobacteria), 409 K-12 education, 451–452, 452 Kiawah River (South Carolina, U.S.), 261–268, 262, 263, 265, 267, 269, 270, 271 Korean mussel (Mytilus coruscus), 399 Kumamoto oyster (Crassostrea sikamea), 348, 410, 412 Label Rouge program (France), 75–76 Laminaria sp., 20 Land Grant college program, 455–456 land use conflicts, 37–42, 40, 41 land-based activities, and eutrophication, 155, 156– 157, 157, 192–195, 193, 194, 201, 217 land-ocean exchanges, changes in as result of climate change, 464 larvae, dissemination of, 378 LCAs (life cycle assessments), and assurance labeling, 76–77, 77 least sandpiper (Calidris minutilla), 247 leathery tunicate (Styela plicata), 405, 411 Letters of Permission, 435 Leucoma staminea (littleneck clam), 251 life cycle assessments, and assurance labeling, 76–77, 77 littleneck clam (Leucoma staminea), 251 local models of carrying capacity, 147–148, 148 longline method of mussel farming, 219, 220, 222, 426 LOPs (Letters of Permission), 435 lugubrious cupped oyster (Crassostrea belcheri), 95, 400 Lysekil, Sweden, and nutrient trading, 196–198, 197, 223, 226–227, 227 Macoma balthica (Baltic clam), 163, 305, 310, 311, 472 Mactra glabrata (smooth mactra), 399 Mactra veneriformis (globose clam), 399 Mactridae, 399 Magnuson-Stevens Fishery Conservation and Management Act, 432, 433 Maine Aquaculture Association Local Area/Bay Management Agreements, 59–60 Mandarin fish, 342 mangrove cupped oyster (Crassostrea rhizophorae), 401 Manila clam. See Tapes philippinarum Manual of Diagnostic Tests for Aquatic Animals, 373

500

Index

marginal physical product, 14, 15 market pressures to participate in assurance labeling programs, 66 Marteilia, 361 Marteilia refringens, 367, 373, 410 Marteilia sydneyi (QX), 410 marteiliasis, 376 Marteiliodes chungnuenis, 362 mechanical harvesting. See harvesting, mechanical Mediomastus sp., 175 Mediterranean mussel. See Mytilus galloprovincialis Mercenaria sp., 242 Mercenaria mercenaria (northern quahog), 89, 103, 106, 163, 172, 173, 181–183, 182, 260–261, 268, 303, 304, 308, 325, 327, 364, 380, 382, 401, 405, 470 Meretrix lusoria (Japanese hard clam), 401 Meretrix meretrix (Asiatic hard clam), 401 Metacarcinus magister (Dungeness crab), 256, 257 microalgae used as biofuel, 130 used as feed, 378–379 Microphthalmus sczelkowii, 176 Mikrocytos mackini, 361 milkfish, 342, 483 Minerals Management Service, 438–439 MMS (Minerals Management Service), 438–439 models for aquaculture, 26 Assessment of Estuarine Trophic Status (ASSETS), 7, 8, 9, 20–21, 20, 147 and culture practice, 6–7, 6 for eutrophication assessment, 7–8, 9, 187–192, 189, 191 Farm Aquaculture Resource Management (FARM), 7–8, 11, 12, 12–13, 15, 15, 147, 190–191, 191 farm-scale, 11–13, 11, 12 of individual growth, 8, 10 ShellSIM, 10 modeling of carrying capacity, 135–139, 137, 138, 150, 188, 189 application to management, 148–149, 148 biogeochemical models, 136 bivalve ecophysiology models, 136 box models, 142–143, 143 fully spatial models, 143–145, 144, 145 higher-order models, 142–143 local models, 147–148, 148 optimization of models, 147–148 physical oceanographic models, 136, 139–140 phytoplankton-nutrients-zooplankton (PNZ) models, 138, 138, 141–142 population-based models, 145–146, 146

single-box models, 140–142 and spatial scale, 136 modeling of environmental impact of shellfish aquaculture, 149–150 Modiolus demissus, 164 Molgula ficus, 411 Molgula manhattensis (sea grape), 411 Mollusc Dialogue. See Bivalve Aquaculture Dialogue (World Wildlife Fund) mollusks impact on bivalve aquaculture, 409–410 introduced from bivalve aquaculture, 405 morphological parameters of culture practice, 6 mortality, 6–7 MPP (marginal physical product), 14, 15 MSFCMA (Magnuson-Stevens Fishery Conservation and Management Act), 432, 433 MSX. See Haplosporidium nelsoni mudworm (Polydora sp.), 403, 404, 408 Mulinia edulis, 102, 106, 108 multitrophic farming, 5, 17, 17, 23–24, 24, 150, 273–274 Musculista senhousia (Asian mussel; Senhouse mussel), 165, 168, 181, 405 mussel dredges, 322 mussel farming, 3, 127, 449 and agricultural environmental aid programs, 224–225, 225 in Baltic Sea, 221–222 and climate change, 473, 474, 483 and denitrification, 226 and diarrheic shellfish toxin, 228, 231–232 environmental value of, 219, 221–222, 221, 226 and eutrophication, 168–174, 180, 217–219, 218, 221–235, 221, 223, 225, 227, 228, 230, 231, 233, 441–442 and food safety, 228, 234 and increased CO2 concentration/ocean acidification, 470 and introduced species, 408–411 longline method, 219, 220, 222, 426 markets, in Sweden, 227–229, 228 mussel meal, use in organic feed, 229–232, 230, 231, 233–234, 233 and nutrient discharges, trading, 222–224, 223, 226–227, 227 remainder resource, 226, 232–234, 233 mussels, nutrient excretion rates, 164–165 Mya arenaria (eastern soft-shell clam; sand gaper), 83, 106, 242, 303, 304, 308, 310, 325, 399, 405, 409 Myidae, 399

Index

Mytilicola orientalis (parasitic copepod), 404, 409 Mytilidae, 242, 342, 342, 343, 399 Mytilopsis adamsi (false mussel), 410 Mytilopsis sallei (black-striped mussel), 405, 410 Mytilus californianus (California mussel), 84, 94, 399, 410 Mytilus chilensis (Chilean mussel), 90, 93, 106, 108, 173, 399 Mytilus coruscus (Korean mussel), 399 Mytilus edulis (blue mussel), 10, 12, 15, 15, 16, 18, 25, 83–86, 83, 86, 88, 89–90, 93, 98, 102, 103, 104, 104, 105, 106, 108, 112, 147, 162–163, 164–165, 166, 168, 169, 170, 171, 172, 173, 174, 175, 179, 181, 181, 190, 196–197, 200, 200, 217–218, 229–230, 230, 232, 233, 234, 304, 327, 383, 399, 407, 408–411, 470, 473, 474, 484 Mytilus edulis platensis (Argentine mussel), 90 Mytilus galloprovincialis (Mediterranean mussel), 12, 15, 15, 16, 25, 90, 92, 94, 165, 168, 170, 171, 174, 175, 176, 180, 183–184, 304, 367, 399, 402, 405, 408, 410 Mytilus planulatis (Australian mussel), 169, 399 Mytilus trossulus, 84, 94, 173, 175, 179, 402 NAPAs (national adaptation programs of action), 480 national adaptation programs of action, 480 National Environmental Policy Act, 432 National Marine Fisheries Service, 432–433, 438 National Oceanic and Atmospheric Administration, 439–440 National Shellfish Sanitation Program, 36 National Sustainable Offshore Aquaculture Act, 439 Nationwide Permits, 40–41, 435–436 native shellfish species, 19, 239–240, 272, 273, 274 and cultured shellfish, 239–240 habitats, 241–243 harvesting, 323–324 and hatchery-propagated populations, interaction of, 349–350, 351–352 NCE (Nitrogen Credit Exchange), 23 near-field operations, 54, 70–71, 431–432 NEPA (National Environmental Policy Act), 432 New Zealand green-lipped mussel. See Perna canaliculus New Zealand scallop (Pecten novaezelandiae), 400 nitrification-denitrification, 159–160, 162, 167, 226, 308 nitrogen credits, trading of, 23, 130, 196–198, 197, 222–224, 223, 226–227, 227

501

pollution, 192, 193, 194 removal, 21–26, 21, 22, 24, 26, 130, 441–442 See also nutrients Nitrogen Credit Exchange, 23 NOAA (National Oceanic and Atmospheric Administration), 439–440 NOAA NMFS (National Marine Fisheries Service), 432–433, 438 noble scallop (Chlamys nobilis), 95, 400 Nocardia crassostreae, 384, 409 Nodipecten nodusus, 95 nonadditive variance, 343–344, 346 nonnative species, introductions of, 54, 347–348, 395–397, 415, 427 and climate change, 415 European Union rules, 413 minimizing spread/impact, 412–415 and shipping activities, 396–397 species introduced from aquaculture, 397–398, 399–401, 402–403, 403, 404–405, 406 species moved with aquaculture, 406–407 species that impact aquaculture, 407, 408–412, 412 nonpoint pollution sources, 54 northern quahog. See Mercenaria mercenaria NSSP (National Shellfish Sanitation Program), 36 Nucula nitidosa, 176 nurseries, management of infectious disease in, 380–382, 380, 381 nursery phase, skills required in, 448–449 nutrients cycling of, impact of mechanical harvesting on, 321, 329 dynamics of, 177–179, 178 enrichment, 159–160 excretion rates, 159, 163–166 overenrichment. See eutrophication reduction of, and best management practices, 46 trading/valuation of credits for, 21–23, 21, 22, 196–198, 197, 222–224, 223, 226–227, 227 See also nitrogen NWPs (Nationwide Permits), 40–41, 435–436 ocean quahog (Arctica islandica), 242, 322 Ocinebrellus inornatus (Japanese oyster drill), 405, 410 OCSLA (Outer Continental Shelf Lands Act), 438, 439 offshore/far-field operations, 54, 71, 437–440, 437 OIE (World Organization for Animal Health), 372–373 Olympic oyster. See Ostrea lurida

502

Index

Oncorhynchus tshawytscha (Chinook salmon), 256 Open Ocean Aquaculture Demonstration Project, 437–438 Ophiura sp., 176 Ophryotrocha sp., 175 orange sheath tunicate (Botrylloides violaceus), 411 organic agriculture, and aquaculture, 72–73 organic enrichment, 307, 307 organic standards, 71–73, 72 Orthopristis chrysopterus (grunt), 244 OsHV (oyster herpes virus) disease, 365, 379–380, 412 Ostrea angasi, 408, 410 Ostrea chilensis (Chilean flat oyster), 90, 95, 401 Ostrea conchapila, 240 Ostrea edulis (European flat oyster), 90, 95, 166, 368–369, 401, 403, 405, 406, 408, 410, 412 Ostrea equestris, 245, 363, 368 Ostrea gigas, 409 Ostrea lurida (edible [flat] oyster; Olympic oyster), 37, 240, 245, 249, 250, 251, 257, 258, 397, 401, 407, 409–410 Ostreidae, 400 Outer Continental Shelf Lands Act, 438, 439 OVVD (oyster velar virus disease), 379, 412 ownership pressures to participate in assurance labeling programs, 67 oxygen conditions and biodeposition, 308–309 and bioturbation, 299, 300, 308–309 and eutrophication, 157, 159, 167 and mechanical harvesting, 321, 328–329 oyster aquaculture, 127, 249–250, 252, 273, 396, 450, 451 and climate change, 483 and eutrophication, 168–174 and increased CO2 concentration/ocean acidification, 470 and introduced species, 408–412 oyster dredges, 322, 322 oyster drill (Urosalpinx cinerea), 405, 407, 410 oyster herpes virus (OsHV) disease, 365, 377, 379–380, 412 oyster reef habitats, 245–247 oyster velar virus disease (OVVD), 377, 379, 412 oysters, nutrient excretion rates, 165–166 Pacific calico scallop (Argopecten ventricosus), 400 Pacific Coast Shellfish Growers Association, 40, 372, 440 Pacific cupped oyster, 342

Pacific geoduck. See Panopea abrupta; Panopea generosa Pacific littleneck clam (Protothaca staminea), 401 Pacific Northwest (U.S.) certification programs, 47 and land use planning, 39–40, 40, 41, 41, 42 and water quality efforts, 36–37, 38 See also Puget Sound; Washington State Pacific oyster. See Crassostrea gigas Pacific salmon, 427 Pangasius, 70, 342, 483 Panopea abrupta (Pacific geoduck), 176–177, 176, 303, 304, 351, 380, 399 Panopea generosa (Pacific geoduck), 40, 250, 252, 254, 255, 300, 351–352, 430, 448 Paphia staminea, 409 Paphia undulata (undulate venus), 401 parasites Bonamia ostreae, 368–369, 373, 385, 410 geographic distribution of, and temperature, 364 Haplosporidium nelsoni (MSX), 347–348, 361– 363, 362, 363, 367–368, 410 Mikrocytos mackini, 361 Perkinsus marinus, 361–362, 362, 363, 365–366, 365, 367, 369, 373, 405, 407, 411, 471 Quahog Parasite Unknown (QPX), 364, 382–383, 383 and salinity, 361–364, 363 and seasonal temperature cycles, 361, 362, 363 parasitic copepod (Mytilicola orientalis), 404, 409 Parophrys vetulis (English sole), 256, 257 particle removal methodology for clearance rate, 100–102, 101 Patinopecten yessoensis (Yesso scallop), 342, 342, 343, 400, 408 PCSGA (Pacific Coast Shellfish Growers Association), 40, 372, 440 pearl farming, 477 pearl oyster. See Pinctada margaritifera Pecten fumatus (Australian southern scallop), 400, 408 Pecten furtivus, 96 Pecten irradians, 96 Pecten maximus (great Atlantic scallop), 86, 86, 96, 106, 108, 400, 409, 476 Pecten novaezelandiae (New Zealand scallop), 400 Pecten opercularis, 96 Pectinidae, 400 Penaeus stylirostris, 195 Penaeus vannarnei, 195 penguin wing oyster (Pteria penguin), 400

Index

performance indicators, and best management practices, 60 performance standards, and best management practices, 58, 63 Perkinsus chesapeaki, 363 Perkinsus marinus (dermo), 361–362, 362, 363, 365–366, 365, 367, 369, 373, 405, 407, 411, 471 Perkinsus olseni, 363, 373 Perkinsus sp., 361 permits. See regulatory framework Perna canaliculus (New Zealand green-lipped mussel), 69, 90, 94, 105, 105, 108, 168, 169, 178–179, 178, 190, 399 Perna indica (Indian brown mussel), 399 Perna perna (brown mussel; South American rock mussel), 91, 94, 173, 399, 403, 405 Perna viridis (green mussel), 91, 94, 106, 108, 342, 342, 343, 398, 399, 403, 405 Peruvian calico scallop. See Argopecten purpuratus phenomenological changes, as result of climate change, 466 physical carrying capacity, 128–129 physical oceanographic models of carrying capacity, 136, 139–140 physiological parameters of culture practice, 6 physiological processes, effects of climate change on, 465 phytoplankton and climate change, 473 effects on, from clam aquaculture in tidal creeks, 264–266, 265, 268 and eutrophication, 128, 158, 177, 196 grazing of, 110–111 trophic interactions with bivalves, 125–131, 126 phytoplankton-nutrients-zooplankton (PNZ) carrying capacity models, 138, 138, 141–142 Pilumnus spinifer, 409 Pincta imbricata, 408 Pinctada fucata (Japanese pearl oyster), 400 Pinctada maculata, 180–181 Pinctada margaritifera (black-lip pearl oyster), 91, 98, 106, 168, 180–181, 400 Pinctada maxima (silver-lip pearl oyster), 91, 400 Pinna sp., 242 piping plover (Charadrius melodus), 247 Placopecten magellanicus (sea scallop), 84, 88, 91, 95–96, 102, 104, 106, 173, 242, 320, 322, 323, 324, 325, 400 plants impact on bivalve aquaculture, 410 introduced from bivalve aquaculture, 405

503

polyculture. See integrated multitrophic aquaculture Polydora ciliata, 408 Polydora nuchalis, 404, 408 Polydora sp. (mudworm), 403, 404, 408 Polydora websteri, 408 Polymesoda sp., 242 population density of bivalves, 127 population-based models of carrying capacity, 145– 146, 146 Posidonia sp., 20 poultry, use of mussel meal as feed for, 229–232, 230, 231 prawns, 342, 483 primary production, effects of climate change on, 465–466, 473 Procentrum lima, 408 Procentrum mexicanum, 408 Procentrum minimum, 404, 408 producers and assurance labeling, 67, 73–74 and best management practices, 62 as water quality advocates, 35–37, 38 product marketing, and best management practices, 61–62 production carrying capacity, 13, 81, 129, 135–149, 137, 138, 143, 144, 145, 146, 147, 148, 150 production enhancement using multiple species, 17, 17 productivity, impact of mechanical harvesting on, 321, 324–325 profit optimization, 14–16, 14, 15 proteobacteria (juvenile oyster disease), 409 Protothaca staminea (Pacific littleneck clam), 401 protozoans impact on bivalve aquaculture, 410–411 infections, 380, 382 introduced from bivalve aquaculture, 405 Pseudomonas sp., 379 Pteria penguin (penguin wing oyster), 400 Pteriidae, 400 public concerns about shellfish aquaculture, 35, 52 Puget Sound (U.S.) climate change impacts, 475 and Panopea generosa (Pacific geoduck), 176–177, 176, 255, 351–352 and water quality efforts, 37, 38, 41, 41 pullet carpet shell (Venerupis pullastra), 106, 401 pulp/paper mills, effect on water quality, 37, 38 pumping rate, in feeding activity, 82–83, 83, 85, 86 QPX (Quahog Parasite Unknown), 364, 382–383, 383

504

Index

Quahog Parasite Unknown, 364, 382–383, 383 queen scallop (Aequipecten opercularis), 400 QX (Marteilia sydneyi), 410 rainbow trout, 342, 483 Rangia cuneata (Atlantic rangia), 405 Rangia sp., 242 Rapana venosa (Asian rapa whelk), 405, 410 razor clam (Ensis sp.), 326, 342 recovery timescales, and mechanical harvesting, 325–326 red swamp crawfish, 342 red tide, 13. See also harmful algal blooms red tide dinoflagellate (Alexandrium minutum), 404 Redfield ratio, 127–128 redox potential discontinuity layer, 300 reef-forming shellfish, 241, 242 re-eutrophication. See eutrophication: reduction of regime shifts, as result of climate change, 466–467 Regional Permits, 436 regulators, and best management practices, 62 regulatory framework, 429–431, 442 Letters of Permission, 435 Nationwide Permits, 40–41, 435–436 and near-shore aquaculture, 431–432 and offshore aquaculture, 437–440, 437 permitting agencies, 432–433, 434, 435 as requirement for successful aquaculture, 34–35 Section 10 permit process, 435 types of, 435–437 regulatory pressures to participate in assurance labeling programs, 67 regulatory requirements, and best management practices, 61 remainder resource, and mussel farming, 226, 232– 234, 233 reportable diseases, 370–371, 373 research scientists, and best management practices, 62 resource managers and introductions of nonnative species, 413 role in balancing economic development and conservation, 429 resting stages in sediments, and bioturbation, 311 restoration aquaculture, 243, 244, 246–247, 248– 249, 272, 349–351, 449 Rhithropanopeus harrisii (estuarine mud crab), 404 Rhodomonas sp., 83 risk analysis, 137, 137 rock crab (Cancer productus), 256 ROD (Roseovarius oyster disease), 380–381, 380 roho labeo, 342

Roseovarius crassostreae, 380 Roseovarius oyster disease, 380–381, 380 RPD (redox potential discontinuity) layer, 300 Ruditapes philippinarum. See Tapes philippinarum Rugulopteryx okamurae, 404 Sabellaria alveolata (honeycomb worm), 402 Sacca de Goro Lagoon (Italy), eutrophication in, 183–187, 185, 186, 189, 189 Saccostrea commercialis, 91, 95, 401, 408, 410 Saccostrea cuccullata (hooded oyster), 401 Saccostrea echinata (spiny oyster), 401 Saccostrea glomerata (Sydney rock oyster), 366–367, 408, 474, 479 salinity and clearance rate, 84 of oceans, changes in as result of climate change, 463 and shellfish diseases, 361–364, 363 salmon, 40, 47, 66, 77, 200, 200, 256, 342, 343, 427, 483 sand gaper. See Mya arenaria Sargassum muticum, 404, 408 SAV (submerged aquatic vegetation), 19–20, 327, 427, 427 Saxidomus gigantean (butter clam), 251, 401 scaling of research, 24–26, 25, 26, 298 of shellfish aquaculture, 126–127 scallop aquaculture and climate change, 476, 476 and eutrophication, 173 and increased CO2 concentration/ocean acidification, 470 and introduced species, 409, 411 scallop dredges, 323, 324 scallops nutrient excretion rates, 166 populations of, 127 Scapharca broughtonii (inflated ark), 399 Scapharca subcrenata (half-crenate ark), 399 Schizoporella errata (branching bryozoan), 404, 409 Schizoporella unicornis (single-horn bryozoan), 404, 409 Scolelepis fuliginosa, 176 sea bass, 342, 483 Sea Grant college program, 455–456 sea grape (Molgula manhattensis), 411 sea level rise, as result of climate change, 464 sea mussels, 342 sea scallop. See Placopecten magellanicus sea snails, 342

Index

seafood buyers, and best management practices, 62 seagrasses and bioturbation, 310 growth of, 8, 259–260 loss of, and eutrophication, 195 See also eelgrass Section 10 permit process, 435 sediment biogeochemistry, and biodeposition, 308 sediment resuspension, impact of mechanical harvesting, 327–329 sedimentation, 18, 18–19, 254, 262–264, 263, 268 seedstock, dissemination of, 378 semipalmated plover (Charadrius semipalmatus), 247 Senhouse mussel (Musculista senhousia), 165, 168, 181, 405 services/goods from aquaculture, xiii, 5, 24, 25, 26, 427–428, 428 biomass production, 13–17, 13, 14, 15, 16, 17 economic, xiii, xiv, xv ecosystem, xiii, xv seagrasses, growth of, 254 See also eutrophication: reduction of; nitrogen: removal services/goods from native shellfish, 243–246 seston and clearance rate, 105–107, 105, 108 depletion of, 81–82, 135–136, 138, 140, 146–147 and feeding activity, 81–82, 85, 140 nutritional content of, 128, 129 setting phase, skills required in, 448, 449 shell debris, 19, 19 shell-accumulation habitat, 241, 242–243 shell-aggregation habitat, 241, 242 shellfish gardens, 36 shellfish growers. See producers Shellfish High Health Plan, Pacific Coast Shellfish Growers Association, 372 ShellSIM model, 10 SHHP (Shellfish High Health Plan), Pacific Coast Shellfish Growers Association, 372 shipping activities, and introductions of nonnative species, 396–397 Shoreline Management Act (Washington State), 39 shrimp, 66, 70, 75, 195, 200, 342, 472 silica, in nutrient chemistry, 129–130 silver-lip pearl oyster (Pinctada maxima), 91, 400 single-box models of carrying capacity, 140–142 single-horn bryozoan (Schizoporella unicornis), 404, 409 Sinovacula constricta (constricted tagelus), 342, 342, 343, 401

505

siting of shellfish aquaculture, 34, 45–46, 126–127, 429, 431 size-dependent particle retention, 110–111 slipper-cupped oyster (Crassostrea iradelei), 95, 400 SMA (Shoreline Management Act; Washington State), 39 smooth cordgrass (Spartina alterniflora), 405, 406, 410 smooth giant clam (Tridacna derasa), 401 smooth mactra (Mactra glabrata), 399 snakehead, 342 social carrying capacity, 129 social effects of aquaculture, 429, 430 social pressures to participate in assurance labeling programs, 66–67 social responsibility, 67, 69–70 soft-sediment habitats, 297, 298, 299, 299, 300, 301, 302, 303 and biodeposition, 306–310 and organic enrichment, 307, 307 structural heterogeneity of, 303–306 and water flow, 306–307, 307 Solecurtidae, 401 Soutellastra argenwillei, 402 Soutellastra granularis, 402 South American rock mussel. See Perna perna Spartina alterniflora (smooth cordgrass), 405, 406, 410 spatfal, and climate change, 472 spatial parameters of culture practice, 6 species invasions, as result of climate change, 415, 466, 471 spiny oyster (Saccostrea echinata), 401 Spisula solidissima (surf clam), 242, 322, 325, 399 spring cultivation, and reduction in eutrophication, 224, 225 SRS (sweepstakes reproductive success), 349 standards in assurance labeling, 65 organic, 71–73, 72 performance, 58, 63 State Programmatic General Permits, 436 sterilization, 352 structural heterogeneity of soft sediments, and bioturbation, 303–306 Styela clava (Asian tunicate), 405, 411 Styela plicata (leathery tunicate), 405, 411 submerged aquatic vegetation, 19–20, 327, 427, 427 Suminoe oyster (Crassostrea rivularis), 401 summer mortality syndrome, 383–385, 384, 385 surf clam (Spisula solidissima), 242, 322, 325, 399 surveillance, in health management, 373–374

506

Index

sustainability, 43–44, 52, 150 sweepstakes reproductive success, 349 SWEM (System Wide Eutrophication Model), 199 Sydney rock oyster. See Saccostrea glomerata System Wide Eutrophication Model (SWEM), 199 Tapes japonica, 163 Tapes philippinarum (Ruditapes philippinarum, Venerupis philippinarum, Manila clam, Japanese littleneck clam), 12, 15, 15, 16, 21, 22, 22–23, 148–149, 148, 163, 168, 171, 174, 177–178, 181, 183–187, 185, 186, 190, 250, 251, 252, 255, 304, 305, 326, 342, 342, 343, 369–370, 401, 403, 405, 406, 410 Tapes semidecussata, 170 technology transfer, 457–458, 458 Tellina deltoidalis, 310 temperature and clearance rate, 84, 104–105, 104 and geographic distribution of parasites, 364 and shellfish diseases, 361, 362, 363, 364–366 temporal parameters of culture practice, 6 temporal variability in clearance rate, 85–87, 86, 88 Tenacibaculum sp., 381 TEP (transparent exopolymer particle) production, 109–110 Thau Lagoon (France), eutrophication in, 183, 189 third-party certification programs, 46–48, 64–65 tidal creeks (southeastern U.S.), clam aquaculture in, 260–268, 262, 263, 265, 267, 270, 270, 271 tilapia, 70, 342, 483 Tiostrea chilensis, 410 Tomentosoides, 404 total quality management programs, and assurance labeling, 75 TPP curve and profit optimization, 14, 15 TQM (total quality management) programs, and assurance labeling, 75 traceability, and assurance labeling, 65, 69 trade barrier, assurance labeling as, 73–74 translocation, impact of, 348–349, 367–369. See also nonnative species, introductions of transparent exopolymer particle production, 109–110 trawling, 320 Tresus sp., 242 Tridacna derasa (smooth giant clam), 401 Tridacna gigas (giant clam), 401 Tridacnidae, 401 triploidy, 346–347, 352 trophic interactions of phytoplankton and bivalves, 125–131, 126. See also feeding activity trout, 342, 483

tubesnout (Aulorhynchus flavidus), 256 Tubularia crocea, 409 tunicates impact on bivalve aquaculture, 411 introduced from bivalve aquaculture, 405 turbidity, impact of mechanical harvesting, 321, 327–328 turbot, 483 two-groove odostome (Boonea bisuturalis), 405 Type I/II/III ecolabels, 64–65 Ulva rigida, 183 Undaria pinnatifida (Asian kelp), 404 undergraduate degree programs, 452–453 undulate venus (Paphia undulata), 401 unsustainable development, 43–44 Uronema marinum, 382 Urosalpinx cinerea (oyster drill), 405, 407, 410 U.S. Army Corps of Engineers, 40, 432–433, 434, 435 U.S. Department of Interior, 439 U.S. Environmental Protection Agency, 433 U.S. Fish and Wildlife Service, 433 USACE. See U.S. Army Corps of Engineers USDA National Organic Program, 71 USFWS (U.S. Fish and Wildlife Service), 433 value of marginal product, 14–15 valve gape, and clearance rate, 86–87 Veneridae, 401 Venerupis corrugatus, 91 Venerupis decussatus (grooved carpet shell), 369, 370, 401 Venerupis philippinarum. See Tapes philippinarum Venerupis pullastra (pullet carpet shell), 106, 401 vertical structure, impact of mechanical harvesting, 321, 324, 326–327 veterinary approach to shellfish health management, 376–377, 377 Vibrio cholerae (Asiatic cholera), 404, 409 Vibro sp., 379 Vibrio tapetis, 369–370 Vibrio tubiashii, 377, 379, 381, 381 vibriosis, 364–365, 381, 381 viruses, impact on bivalve aquaculture, 379, 411, 412 viscosity of water, and feeding activity, 84 VMP (value of marginal product), 14–15 Washington State (U.S.), and water quality efforts, 36–37, 38, 39–40, 40, 41, 41, 42 Grays Harbor (Washington State, U.S.), 256 Growth Management Act, 39

Index

Shoreline Management Act, 39 Willapa Bay, 253, 254–255, 256, 257, 258 water chemistry, impact of mechanical harvesting, 328–329 water circulation patterns, and shellfish diseases, 367 water flow and biodeposition, 306–307, 307 and clam aquaculture in tidal creeks, 266–268, 267, 269 and eutrophication, 156, 162, 166 and soft-sediment habitats, 306–307, 307 water quality assessment of, 8 role of producers in protecting, 35–37, 38 watershed protection, best management practices, 53, 54–55 wetlands, and reduction in eutrophication, 225, 225 white amur bream, 342 wild species. See native shellfish species

507

Willapa Bay (Washington State, U.S.), 253, 254–255, 256, 257, 258 withering syndrome, 364, 365 World Organization for Animal Health (OIE), 372–373, 375 World Wildlife Fund Bivalve Aquaculture Dialogue, 47, 70–71, 75, 413–414, 441 WWF Bivalve Aquaculture Dialogue. See World Wildlife Fund Bivalve Aquaculture Dialogue Xenohaliotis californiensis, 364, 373 Yesso scallop. See Patinopecten yessoensis zoning and best management practices, 58–60 for establishing biosecurity, 375–376 Zostera sp. See eelgrass