Advances in MARINE BIOLOGY VOLUME 37
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Advances in MARINE BIOLOGY Edited by
A. J. SOUTHWARD Marine Biological Association, The Laboratoty, Citadel Hill, Plymouth, England, UK
I? A. TYLER School of Ocean and Earth Science, University of Southampton, England, UK
and
C . M. YOUNG Harbor Branch Oceanographic Institution, Fort Pierce, Florida, USA
ACADEMIC PRESS A Horcourt Science and Technology Company
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This book is printed on acid-free paper Copyright 0 2000 by ACADEMIC PRESS Chapter entitled 'Population Structure and Dynamics of Walleye Pollock, Theragru chalcogrumma' is a US Government work in the public domain and not subject to copyright. All Rights Reserved No part of this publication may be reproduced or transmitted in any form or by any means, electronic or mechanical, including photocopy, recording, or any information storage and retrieval system, without permission in writing from the publisher. Academic Press 24-28 Oval Road, London NW1 7DX, UK http:llwww.hbuk.co.uWap/ Academic Press A Harcourt Science and Technology Company 525 B Street, Suite 1900, San Diego, California 92101-4495, USA http:llwww.apnet.com ISBN 0-12-026137-5 A catalogue record for this book is available from the British Library
'Ehpeset by Keyset Composition, Colchester, Essex Printed in Great Britain by MPG Books Ltd., Bodmin, Cornwall
99 0001 02 03 04 MP 9 8 7 6 5 4 3 2 1
K . M . BAILEY,Resource Assessment and Conservation Engineering Division, Alaska Fisheries Science Center, 7600 Sand Point Way NE, Seatde W A 98115, USA R. BEIRAS, Area de Ecoloxia, Universidade de Vigo, 36200 Galicia, Spain P. BEmzm, Marine Molecular Biology Laboratory, School of Fisheries, University of Washington, Seattle W A 98195, USA W. S. GRANT,Conservation Biology Division, Northwest Fisheries Science Center, 2725 Montlake Blvd., Seattle W A 98112, USA E. H I S ,IFREMER, Quai Silhouette, 33120 Arcachon, France T. J. QUINN 11, Juneau Center, School of Fisheries and Ocean Sciences, University of Alaska Fairbanks, 1120 Glacier Highway, Juneau A K !?98018677, U S A M . N . L. SEAMAN, Institute of Marine Research, 24105 Kiel, Germally
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CONTENTS
CONTRIBUTORS TO VOLUME 37...........................................
v
The Assessment of Marine Pollution .Bioassays with Bivalve Embryos and Larvae E. His. R. Beiras and M. N. L. Seaman 1. Introduction .................................................................. 3 2. Relevant Aspects of Bivalve Biology....................................... 10 3. Bioassay Methodology ....................................................... 40 4. Testing the Toxicity of Marine Pollutants to Bivalve Embryos and Larvae 87 5 . Assessing Marine Environmental Quality with Bivalve Embryo and Larval Bioassays..................................................................... 125 130 6. Summary and Discussion.................................................... 138 Acknowledgements ........................................................... References .................................................................... 139
Population Structure and Dynamics of Walleye Polllock. Theragra chalcogramma K. M. Bailey. T. J. Quinn II. I? Bentzen and W. S. Grant 1. 2. 3. 4. 5.
Introduction .................................................................. Background The Fishery. Life History and Ecosystem Interactions ....... Population Ecology.......................................................... Population Structure ......................................................... Management Implications ................................................... Acknowledgements ........................................................... References ....................................................................
Taxonomic Index .............................................................. Subject Index .................................................................. Cumulative Index of Titles .................................................... Cumulative Index of Authors .................................................
180 184 189 206 238 242 242 257 261 269 278
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The Assessment of Marine Pollution . Bioassays with Bivalve Embryos and Larvae E. His'. R . Beiras' and M . N. L . Seaman3
'IFREMER. Quai Silhouette. 33120 Arcachon. France 2 h e a de Ecoloxia. Universidade de Ego. 36200 Galicia. Spain 31nstitute of Marine Research. 24105 Kiel. Germany Correspondence to: Edouard His. IFREMER. Quai Silhouette. 33120 Arcachon. France. Fax: 05 56 83 89 80. Tel. 05 56 83 78 17. e-mail:
[email protected]
1. Introduction ......................................................................... 3 1.1. Generalities on pollution ...................................................... 3 1.2. Bioassays: advantages and limitations ...................................... 4 6 1.3. Bivalve larvae and pollution .................................................. 2. Relevant Aspects of Bivalve Biology ............................................. 10 2.1. Species used in bioassays ................................................... 10 2.2. Reproduction .................................................................. 16 2.3. Larval rearing in the laboratory ............................................. 25 3. Bioassay Methodology ............................................................ 40 3.1. General methods .............................................................. 40 50 3.2. Bioassay procedures ......................................................... 3.3. Bioassay applications: toxicity tests and environmental bioassays ....... 74 82 3.4. Statistical methods ........................................................... 4. Testing the Toxicity of Marine Pollutants to Bivalve Embryos and Larvae .... 87 87 4.1. Pollutants ...................................................................... 117 4.2. Intrinsic (biological) factors affecting toxicity ............................... 4.3. Extrinsic (environmental) factors affecting toxicity ......................... 121 122 4.4. Interactions between different toxicants .................................... 5. Assessing Marine Environmental Quality with Bivalve Embryo and Larval 125 Bioassays ........................................................................... 5.1. Algal and bacterial toxins .................................................... 125 127 5.2. Urban and industrial effluents ............................................... 5.3. Receiving waters .............................................................. 128 129 5.4. Sediments ..................................................................... 6. Summary and Discussion ......................................................... 130
ADVANCES IN MARINE BIOLOGY VOL. 37 ISBN 0-12-026137-5
Copyright 0 1999 Academic Press All rights of reproduction in any form reserved
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E. HIS, R. BEIRAS AND M. N. L. SEAMAN
6.1. Sensitivity of bioassay organisms ........................................... 131 6.2. Assessment of the toxicity of various contaminants ....................... 132 6.3. Bivalve embryo and larval bioassay methodology ........................ 134 6.4. Perspectives in future research on bivalve larval bioassays .............. 137 6.5. Concluding remarks .......................................................... 138 Acknowledgements ................................................................... 138 References ............................................................................. 139
Tens of thousands of synthetic substances are in existence today and hundreds of new compounds are being introduced every year. Because of the complexity of the physico-chemical interactions between pollutants and the marine environment, the potential toxicity of contaminants can be assessed adequately only by means of bioassays with living organisms. From a practical point of view, a bioassay needs to be sensitive and scientiJically valid, yield rapid results at moderate cost, and the organism in question must be readily available. Ecotoxicological bioassays with bivalve embryos and larvae fulfil these criteria better than most other tests. They have increasingly come into use during the past three decades and are now commonly employed to ascertain the biological effects of pure chemicals, as well as to determine the quality of efluents, coastal waters and sediments sampled in the field. There do not appear to be very great differences between bivalve species with regard to larval sensitivity to toxicants. The principal species f o r bioassays are oysters (Crassostrea gigas and C. virginica), and mussels (Mytilus edulis and M . galloprovincialis). Bioassays are conducted with gametes and larvae of all ages: sperm and unfertilized eggs, embryos, young D-larvae, intermediate umboned larvae, and pediveligers towards the end of the pelagic period. Embryos are usually the most sensitive stage. Recent advances now also permit bioassays on metamorphosing pediveligers, a method particularly suited to investigate the effects of adsorbate-contaminated surfaces. There are various criteria for the assessment of toxic effects, including embryogenesis success (abnormalities), larval growth, mortality, physiology (e.g. feeding or swimming activity), and metamorphosis success. Chronic toxicity studies may be carried out over periods of several weeks, but larval rearing in the laboratory requires considerable effort (e.g. cultivation of algal food). The method of choice f o r investigations of acute toxicity and for routine monitoring studies is the embryo bioassay because it is very sensitive, relatively simple, and produces results within 24 or 48 hours. The data obtained by different investigators are often dificult to compare, however, because of differences in methodology. There is no firmly established procedure, and further simplification and standardization of techniques is required. In bioassays with a single pollutant, the effective toxic concentration may
THE ASSESSMENT OF MARINE POLLUTION
3
span several orders of magnitude, depending on bioassay procedures, larval stage and choice of response. Tributyl-tin (TBT) is the most toxic compound ever bioassayed with bivalve larvae, with effective concentrations (EC,,) as low as a few nanograms per litre (i.e. lo--’ ppb). Heavy metals (particularly mercury, silver and copper) are next in order of toxicity, with ECSovalues between a few micrograms per litre (ppb) and several hundred ppb. Chlorine and some organochlorine pesticides may also have ECso values of less than 100 ppb, while detergents and petroleum products are generally less toxic.
1. INTRODUCTION
1.1. Generalities on pollution
Although human activities have always impacted on coastal areas, it is only within the last two centuries that the effects of industrialization, intensive agriculture and coastal engineering (including tourism) have seriously begun to threaten marine life. Some of these impacts lead to environmental pollution, i.e. the introduction by man into the environment of substances or energy which may put human health or living resources at risk (Holdgate, 1979). Pollutants are defined as substances present in the natural environment which are (at least in part) of anthropogenic origin, and which may have deleterious effects on living organisms (Moriarty, 1990). According to this definition it is necessary to distinguish between contaminants, which encompass all substances of anthropogenic origin introduced into the environment, and pollutants, which consist of those contaminants with presumably negative biological effects. Moriarty recognizes, however, that the term “pollutant” is generally applied in the sense of “contaminant”. The assessment of marine pollution is not restricted to the study of the water. Most of the pollutants - heavy metals and hydrophobic organic compounds in particular - have a tendency to adsorb to sediment particles, and their concentrations in marine sediments may be several orders of magnitude higher than in the water column (Livett, 1988). The association between pollutants and sediments can be of very long duration and it may have deleterious effects on organisms living on, and within, the bottom. The assessment of the toxicity of marine bottom sediments has, therefore, also come into focus since the late 1970s. According to Moriarty (1990) some 63000 chemical products are presently in use, and 3000 of these constitute 90% of the total mass of
4
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
compounds being produced industrially. In addition, between 200 and 1000 new synthetic compounds are being introduced every year. It would obviously be desirable to screen their possible ecological effects. As already pointed out by Cunningham (1979), ecotoxicological studies require several levels of investigation. Depending on the mode of action of the pollutants in question, these may be at the subcellular, cellular, organismic, population or ecosystem level (see also Bayne, 1985; Haux and Forlin, 1988). 1.2. Bioassays: advantages and limitations
Anthropogenic impacts on the aquatic environment may be viewed from a physical, chemical or biological perspective. The biological effects of pollutants in the environment are more important than the mere presence of pollutants; with regard to environmental quality criteria, the data from chemical analyses of pollutants can only be interpreted within a biological context. It is therefore logical to use biological systems for the assessment of environmental quality (Anonymous, 1989). The following points apply to ecotoxicological bioassays in general:
1. Detection of new pollutants: reliance on chemical analyses alone would presuppose that the potentially important pollutants are known and already being monitored. Examples such as tributyl-tin (TBT) demonstrate that this is not always the case and that biological systems provide the means to detect and identify the presence of new or unexpected pollutants. 2. Bioavailability: chemical data often do not reflect the bioavailability of pollutants, e.g. owing to the speciation of organic compounds or the bonding state of metals. By definition, biological systems can only respond to what is effectively available to them, and test organisms therefore provide the best indication of bioavailable pollutants exceeding toxicity thresholds. 3. Integration of toxic effects: toxic contaminants typically do not occur singly and environmental quality is generally determined by their combined effects. Biological systems respond to the totality of environmental pollution, thus providing an integrated response to the totality of pollutants present, as well as to their interactions. 4. Cost: the continuous increase in the number of contaminants being introduced entails a constant rise in the cost of chemical monitoring programmes. This underscores the importance of using biological techniques as reconnaissance systems, because they help to focus the effort of chemical analysis on situations of demonstrated biological
THE ASSESSMENT OF MARINE POLLUTION
5
relevance. Therefore, “bioassays ... may be used before any other testing commences as a cost-effective screening tool” (Chapman and Long, 1983, p. 83). Any bioassay used for routine monitoring purposes should meet a variety of criteria (Moore, 1966; Butler et al., 1971; Bryan et al., 1980, 1985; Stebbing et al., 1980): it should be easy to learn, affordable, and not require very sophisticated equipment it should be of short duration (a few hours or days, rather than weeks) the organisms employed should, if possible, be available for bioassay purposes year-round the organisms should be of ecological or economic importance the organisms, whether they originate from laboratory cultures or from field sampling, should be identical as far as possible, in order to reduce variability resulting from age, size etc. genetically homogeneous, or cloned, organisms would be preferable, to reduce the effects of genetic variability the data should be of a type that can be analysed by standard statistical methods. Most bivalves of commercial interest generally fulfil these criteria, particularly those living in the marine and brackish water environment, such as oysters of the genus Crassostrea and mussels of the genus Mytilus. They are eurytypic organisms with higher tolerance to environmental fluctuations and therefore better suited for studies on the evaluation of environmental quality (Bayne, 1985), than stenotypic organisms with narrow tolerance. The different ontogenetic stages in marine species may differ in their sensitivity to pollutants (Coglianese and Martin, 1981). Among the many methods employed in bioassays, those using meroplanktonic stages (such as sea urchin plutei or bivalve veligers) appear to be the most promising to obtain reliable biological responses with regard to coastal water quality (Klockner et al., 1985). Many authors have proposed the use of early life stages of bivalves for marine toxicological studies, because it is clearly established that they are more sensitive to toxic substances than are the adults (Wisely and Blick, 1967; Granmo, 1972; Brereton et al., 1973; Hrs-Brenko et al., 1977; Coglianese and Martin, 1981; Bourne et al., 1981; His and Robert, 1981, 1982; Ringwood, 1991). The success of a species depends on its performance during passage through successive life stages, and it is therefore realistic to use the most sensitive stage for the purpose of testing environmental quality (Stebbing et al., 1980; Calabrese, 1984).
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
1.3. Bivalve larvae and pollution
Bivalve molluscs are an exclusively aquatic taxon of extremely wide distribution in fresh, brackish and marine waters around the globe. They are particularly abundant in the highly productive marine coastal areas. They have been exploited for food since prehistoric times and they have been cultivated systematically in various parts of the world for many centuries. The first problems in bivalve culture with regard to pollution were encountered at the beginning of the twentieth century. In studies on the American oyster, Crassostrea virginica,Prytherch (1924) asserted that “the rapid decline of this valuable industry has been brought about by a constant depletion of the oyster beds from various factors, such as pollution . . .” (p. 1). Prytherch was also the first to attempt, and to succeed in, the “artificial reproduction” of oysters. Besides intending to obtain larvae for culture purposes and thus offset the irregularities of natural recruitment, he also intended to conduct basic studies on larval biology and “to produce larvae for tests in regard to the effect upon them of various chemicals in solution” (op. cit., p. 2). It was already evident from Prytherch’s studies that bivalve larvae (veligers) are particularly suited for investigating environmental impacts, because in oyster cultivation areas the first environmental effects to be observed concerned the species’ natural reproduction. The same observation was made again 60 years later by Stanley and DeWitt (1983) who declared that the absence of a population of Mercenaria mevcenaria in the species’ normal area of distribution is an ecological indicator of environmental disturbance. The efforts of Loosanoff and his associates at the Milford Laboratory in Connecticut led to the development of practical and reliable methods for the culture of bivalve larvae (reviewed by Loosanoff and Davis, 1963) and the first toxicological bioassays with such larvae. A serious constraint in working with oyster larvae, however, was the limitation to the species’ reproductive period, from June to September. The invention of “conditioning”, i.e. techniques to obtain ripe adults at all seasons (Loosanoff, 1945) made it possible to study larval nutrition throughout the year. The work conducted by Cole (1937,1939) in Great Britain, made it possible to study the nutritional value of marine unicellular algae, first isolated by Parke at Plymouth; certain species, especially Isochrysis galbana, permitted the rearing of larvae to metamorphosis (Davis, 1953; Loosanoff, 1955; Walne, 1956; Davis and Guillard, 1958). It thus became possible to grow veligers in the laboratory as a matter of routine, and to study their principal environmental requirements. This was followed by studies on the effects of micropollutants on growth and survival in Crassostrea virginica larvae (Davis, 1961; Calabrese and Davis, 1967).
THE ASSESSMENT OF MARINE POLLUTION
7
At the same time, the concept of the “biological quality” of seawater was introduced by Wilson (1951) and Wilson and Armstrong (1961), who found that pluteus larvae of the sea urchin Echinus esculentus were capable of developing in seawater obtained from the Celtic Sea, but not in water from the vicinity of Plymouth. Subsequently Woelke (1961, 1966, 1967, 1972), Okubo and Okubo (1962) and Dimick and Breese (1965) suggested the use of oyster embryos and larvae to study the general effects of pollutants in water samples from the natural environment, and not just for toxicological laboratory investigations of specific pollutants. This ecotoxicological approach, i.e. the study of samples obtained from the natural environment, has since been expanded to include assays with marine sediments (Chapman and Morgan, 1983; Swartz, 1989; Phelps and Warner, 1990). Most bivalve embryo-larval bioassays can be conducted within a relatively short time period (24 to 48 hours after fertilization). Assessments of sublethal toxic effects of pollutants are based on the percentage of normal D-shaped larvae obtained at the end of embryogenesis (determination of the concentration that inhibits larval development in 50% of the fertilized eggs). These types of studies have been conducted by Woelke and coworkers (Woelke, 1960a,b, 1961, 1966, 1967, 1968, 1972; Cardwell et al., 1977a,b), Davis and coworkers (Davis, 1961; Calabrese and Davis, 1967), and many others. Experiments of longer duration, based on the assessment of larval growth, have also been conducted in order to study the effects of sublethal micropollutant concentrations (e.g. Davis and Hidu, 1969a; His and Robert, 1981 and 1982; Watling, 1982). These types of studies represent a better simulation of conditions prevailing in the natural environment, but they also necessitate the cultivation of algae to feed the larvae during the experiment, which may last several days, or several weeks if it is continued until the end of the veliger’s pelagic life stage. Finally, recent work has focused on the use of eyed larvae several days old, in order to investigate the effects of pollutants on the pediveliger stage and on metamorphosis, because these stages are considered to be especially sensitive (Stafford, 1913; also Watling, 1978, 1983; Sheffrin, 1982; Sheffrin et al., 1984; Beiras and His, 1994, 1995a; His et al., 1997b). Most bioassays with bivalve larvae and embryos have been conducted with various species of oysters (57% of the publications listed in Table l), followed by mussels (22%). Only two other species have been used regularly in larval bioassays - the quahog Mercenaria (Venus) rnercenaria and the coot clam Mulinia lateralis. The first step in bioassays with bivalve embryos and larvae is to obtain spawning adults and rear the fertilized gametes. We will therefore review the present state of knowledge on bivalve reproduction, methods to obtain
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
8
Table 1 Bivalve species used in embryo and larval bioassays. Mytilus
Crassosfrea
c gigas Beiras and His 1994, 1995b Beiras ef al., 1998 Bourne ef al., 1981 Boyden el al., 1975 Brereton el ol., 1973 Butler el ni., 1992 Cardwell, 1976,1978 Cardwell ef al., 1976, 1977a.b. 1979a.b Chang ef al., 1996 Chapman and Morgan, 1983 Chapman ef al., 1991, 1992 Chien and Chou, 1989 Cleary ef al., 1993 Coglianese, 1982 Coglianese and Martin, 1981 Connell ef al.. 1997 Coon era[., 1990 Crecelius, 1979 Eertman ef 01.. 1W3 Rtt el al., 1990 Garland el al., 1986 Geffard, 1997 Glickstein. 1978 Helm ef al., 1974 Hi%1996 His and Robert, 1980, 1981,1982,1987b His and Seaman, 1993 His ef al., 1983, 1996 Klockner ef al., 1985 Konar and Stephanson, 1995 Le Gore, 1974 Lourens el a/., 1995 McFadzen, 1992 McFadzen and Cleary. 1994 Maimstone el al., 1989 Martin ef al., 1981 Nelson ef al., 1983 Okubo and Okubo, 1962 Pbelps and Warner, 1990 Renard, 1991 Renzoni, 1973a,b Robert and His, 1981, 1985 Robert ef al., 1986 Smith, 1968 Stewart and Blogoslawski, 1985
C virginica Baker and Mann, 1992. 1994a.b Brown. 1981 Brown and Roland, 1984 Butler and Lowe, 1978 Calabrese, 1972 Calabrese and Davis, 1964,1%7,1970 Calabrese ef a[.. 1973, 1977a.b Capuzzo, 1979 Chapman et nl., 1987 Davis 1958,1960,1961 Davis and Calabrese,
1964 Davis and Hidu, 1%9a,b Dim, 1973 Hidu, 1965 Hidu el ni., 1974 Ho and Zubkoff, 1979, 1980 Mann and Rainer, 1990 Mchnes, 1981 McInnes and Calabrese, 1978,1979 Nelson ef al., 1983 Noyes ef al., 1978 Phelps and Mihursky, 1986 Phelps and Warner, 1990 Prytherch, 1931, 1934 Reuzoai, 1915 Richardson ef ol., 1982 Ringwood and Brouwer, 1995 Roberts 1980,1987 Roberts and Casey, 1985 Roberts and Gleeson, 1978 Roberts ei al., 1975,1977 Roesijadi ef al., 19% Roosenburg ef ~ l .1980b , Sigler and Leibovitch, 1982 Stewart er al., 1979 Stiles-Jewel],1994 Stiles and Blogoslawski, 1993 Tagatz and hey, 1981 Ukeles and Sweeney, 1969 Widdows ef al., 1989 Wikfors and Ukeles 1982 Wirtb eta/., 1996 Wolfe ef al., 1993
M. edulis Akberali ef a/., 1985 Armstrong and Millemann, 1974 Beaumont and Budd, 1982,1984 Beaumont and Tserpes, 1984 Beaumont el al., 1981 Breese eral., 1963 Brunetti ef a/., 1989 Butler ef al., 1990 Chapman and Long, 1983 Chapman ef al., 1993, 1996 Courtright el a/., 1971 Dimick and Breese, 1965 Dixon and Prosser, 1986 Granmd, 1972 Granmo and Jorgensen, 1975 Granmo ef 01.. 1988, 1989 Hansen ef 111.. 1997 Hoare er al., 1995a,b Johnson, 1988 Knezovicb ei al., 1996 Lapota er al., 1993 Le Pennec and Prieur, 1972 Le Pennec ef al., 1973 Long ef al., 1990 Magnusson el al., 1996 Martin ef d., 1981 Mitchell ef al., 1985 Morgan ef al., 1986 Okubo and Okubo, 1962 Pavicic, 1980 Sheffrin and Williams, 1984 Sheffrin ef al., 1984 Stewart ef al., 1%7 Strdmgren and Nielsen, 1991 main, 1983 Wisely and Blick, 1967 Wolfe er al.. 1995 Zhadan er al., 1992
M.galloprovincialis Beiras and Hq 1995a Brunetti ef ai., 1989 Bucaille and Kim, 1979 His and Beiras, 1995 €in-Brenko er a[., 1977 Ix Pennec and Le Roux, 1979 Le ROUK,1977 Lucu ef a/., 1980 Pagano ef ol., 1996 Pavicic, 1976 Pavicic and Pihlar,1982 Pavidc ef al., 1984a, 1994a,b Renzoni, 1973a Robert and His, 1981 Seaman el al., 1991
9
THE ASSESSMENT OF MARINE POLLUTION
Table 1 -(Continued). Crnsroslrea
C. gigas Slewart el al., 1967,1983, 1991 Thain and Watts, 1987 Thain eral., 1990 Utting and Helm, 1985 Van den Hurk, 1994 Van den Hurk el al., 1997 Wang ef 01.. 1985 Warder al., 1992a,b Watling, 1978,1981,1982, 1983 Wikfors el al., 1993 Williams el 01.. 1986 Woelke, 1960,1961,1967, 1968, 1972 Wolfe er el., 19% Zhadan ei a/., 1992
Mytilus
C. virginica
M. edulis
M. galloprovincialis
Wright el al., 1983 Zaroogian and Morrison, 1981
Other species Argopeclen irradians: Nelson and Siddall, 1988; Wright el al., 1983 Callisfa brevisiphonata: Zhadan el a/., 1992 Cerastodema edule: Tunmemans el al., 19% Chlamys asperrima: Krassoi, 1995; Krassoi el 01.. 1996, 1997: Pablo el al., 1997; Stauber er a/., 1996 Clinocardium nultalli: Stewart el al., 1967 Crassoszreu ungulata: Renzoni, 1973a Crassosrrea nrccullaru: Watling, 1981,1982 Crassosrrea iredalei: Ramachandran ef a/., 1997 Crassostrea margaritacea: Watling, 1981, 1982 Crassosrrea rhizophorae: Chung, 1980; Nascimento, 1989; Pereira er al., 1998 Isognomon californicurn: Ringwood, 1990,1991,1992a.h, 1993 Macoma balthica: Tunmemans el al., 1996 Macrru chinensix Zhadan el a/., 1992 Mercenaria mercenaria: Brown, 1974;Byme and Calder, 197R Calabrese, 1972; Calabrese and Davis 1966,1970; Calabrese and Nelson, 1974; Calabrese el al., 1977a.b Davis, 1958, 1%0; Davis and Calabrese, 1964; Davis and Hidu, 1969a.b:
Hidu, 1965;Huntington and Miller, 1989; Laughlin el a/., 1988.1989; Miller, 1989; Pavicic. 1980, Roberts, 1987; Roberts 1983; Wright el a/.. 1983 Meretrix lusoria: Tzong-Shean and Chen, 1993 Mizuhopecfen yessoensix Karaseva and Medvedeva, 1993; Zhadan era/., 1992 Mulinia larerdix Burgess and Momson, 1994, Calabrese, 1970b; Calabrese and Rhodes 1974; Diaz 1975; Gormly e1 al., 19%; Hall el a/., 1995; Ho and Zubkoff, 1980, 1982, 1983; Mann el al., 1991; Morrison and Petrocelli, 1990 Pelletier el al., 1997; Renzoni, 1975; Roberts 1980:Wright eral., 1983 Mya arenaria: Roosenburg el al., 1980a Mytilus califomianus: Cherr er al., 1990; Spangenberg and Cherr. 1996 Mytilus lrossulus: Karaseva and Medvedeva, 1993 Osrreu edulix Connor, 1972 Davis 1961;Davis and Hidu. 1969b Helm. 1971; Millar and Scott, 1968, Nottage and Birkbeck, 1987a.b Renzoni. 1973b: Smith, 1968; Thain. 1983 Osfreaplicalula: Xu el al., 1994 Peaen marimus: Beaumont and Budd, 1982; Beaumont el al., 1987 Prolothaca staminea: Cardwell el a/., 1979b Rangiu cuneala: Mann el al., 1983 Saccosfrea commercialis (= Crassostrea commercialis): Krassoi. 1995; Nell and Holliday, 1986; Wilson and Hynes 1997; Wiseley and Blick, 1967 Scrobicularia plma: Ruiz er al., 1994,1995a.b.c Spisula sukhalinensis: Zhadan el a l , 1992 SpisuIu solidissima: Thurberg er al., 1975; Calabrese et a/., 1982: Eyster and Morse, 1984; Mann et al., 1983; Wright er a/., 1983 Tapesphilippinarum ( = bjaponica): Cardwell e1 al., 1979b: Cleary el al., 1993; McFadzen, 1992; McFadzen and Cleary, 1994 Tresus capar and 7: nuttalli: Cardwell. 1976; Cardwell el a/., 1978, 1979a el al., 1975; Robinson,
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E. HIS, R. BEIRAS AND M . N. L. SEAMAN
and fertilize the gametes, and cultivation of the larvae to metamorphosis, with emphasis on the principal species used to date in toxicological assays and environmental monitoring.
2. RELEVANT ASPECTS
OF BIVALVE BIOLOGY
2.1. Species used in bioassays
The species used most often in bioassays is the Pacific or Japanese oyster Crassostrea gigas (30%), followed by the American or Eastern oyster C. virginica (23%), the blue mussel Mytilus edulis (15%), the quahog or hard clam Mercenaria mercenaria (13%), the Mediterranean mussel Mytilus galloprovincialis (7%), the European oyster Ostrea edulis (3%), and the coot clam Mulinia lateralis (3%) (Table 1). The biology of these species is reviewed below. 2.1 .l. The Japanese oyster Crassostrea gigas The Pacific or Japanese oyster originates from Eastern Asia, where it is distributed from the Sea of Okhotsk southward along the coasts of Korea and Japan to China (Ahmed, 1975; Andrews, 1980; Ventilla, 1984). Following the studies by Imai and Sakai (1961) four varieties or races are generally recognized; these are, distributed from north to south, Hokkaido, Miyagi, Hiroshima and Kumamoto (see Quayle, 1969). They are characterized, among other points, by different spawning temperatures. The Miyagi variety has repeatedly been introduced to the Pacific coast of North America since the beginning of the twentieth century, and it has formed an important natural population which spawns irregularly, but intensively, in British Columbia. Kumamoto oysters have been introduced in smaller numbers to Washington State (Quayle, 1969; Andrews, 1980). Owing to the scarcity of naturally setting oysters, the spat for aquaculture in British Columbia and Washington State are mostly produced by hatcheries. Miyagi, but also Hiroshima and Kumamoto oysters, have been introduced to Australia, and now form important natural populations on the southwest coast and around Tasmania (Thompson, 1951; Summer, 1980a,b). They have spread accidentally to the coast of New South Wales, where oyster farmers consider them a nuisance (Medcof and Wolf, 1975). They were also introduced accidentally to New Zealand in the 1960s, where they quickly formed abundant natural populations (Dinamani, 1971, 1987).
THE ASSESSMENT OF MARINE POLLUTION
11
In Europe (discounting the probable introduction centuries ago of Crassostrea angulata, often considered the same species as C. gigas, but never used in bioassays, except by Renzoni (1973a,b), Pacific oysters were first introduced to Great Britain in the form of hatchery-produced spat from British Columbia in 1972 and 1975 (Gosling, 1981). Natural spatfall has been observed in Britain after warm summers (Askew, 1972). After the decline of the stocks of C. angulata in France, massive imports of Miyagi spat from Japan, as well as spat and spawners from British Columbia, quickly led to the formation of important natural stocks in the early 1970s (His, 1976). C. gigas reproduces very successfully in most oyster culture areas of southwest France, and occasionally in Brittany. Natural spatfall has also led to the establishment of significant wild populations in the Netherlands (Andrews, 1980) and Germany. The latter probably originated from spawning by the Dutch stocks (Seaman and Ruth, 1997), because successful local reproduction of the German stocks appears improbable (Neudecker, 198l), even though very ripe adults are found in some years. Finally, Japanese oysters also reproduce in the Adriatic part of the Mediterranean from Triest (Valli et al., 1979) to the Canal of Lim, where spawning occurs from July to October (Filic and Krajnovic-Ozretic, 1978; Hrs-Brenko, 1982). Andrews (1980) considers C. gigas to be the only species of oyster to have been introduced as a successful member of coastal communities around the world; “this oyster has adapted well to a wide range of environmental conditions and is probably the most globally widespread and ubiquitous oyster species in the world” (Chew, 1990). This explains why it is one of the species used most frequently in marine ecotoxicology. Gametogenesis in C. gigas is activated when the water temperature attains 14°C and continues as temperatures rise; Mann (1979) showed that not only are temperatures of 15°C to 18°C required, but that gametogenesis also depends on the duration at different temperatures (day-degrees). In addition, a temperature of at least 18°C to 20°C is necessary for spawning to occur (Fujiya, 1970; Mann, 1979). Buroker (1985), on the other hand, mentions a spawning temperature of 19°C to 24”C, and a salinity of 11 to 32 psu. With respect to the reproductive cycle in the natural environment, Imai and Sakai (1961) showed the differences between the four varieties in transplantation experiments at Miyagi province. The Hokkaido and Miyagi oysters spawned in late August to early September, those from Hiroshima in early or mid-September; those from Kumamoto spawned in late July when water temperatures attained 22°C to 23°C and began to mature again, with mature gametes still present in November and December. More generally, Koganezawa (1978) reports that Pacific oysters in Japan spawn from June to August. On the Pacific coast of Siberia (Vostok Bay,
12
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
Sea of Japan), they generally spawn in July (Yakovlev, 1978). In the United States there are no natural populations except at Washington State, where they spawn in July and August (Katkansky and Sparks, 1966; Perdue and Erickson, 1984). In Australia the oysters spawn from midJanuary or mid-February on, after water temperatures exceed 22°C (Thompson, 1951). In New Zealand the reproductive season may be very extended, with light spawning in mid-spring and successive spawnings in summer and early fall (Dinamani, 1987). In Europe, spawning is occasional in British, Dutch and German waters, occurring only in exceptionally warm summers (Askew, 1972; Andrews, 1980; Neudecker, 1981). Pacific oysters spawn intensively on the Atlantic coast of France from the Bidassoa estuary on the French-Spanish border, to the Loire estuary, which is the region with the most important centres of oyster culture. Further north, in Brittany and Normandy, the oysters attain sexual maturity, but spawn only in unusually warm summers; spatfall is light, except in the nearly closed bay of the Rade de Brest. Gametogenesis begins at the end of winter and spawning takes place after the water temperature reaches 20°C or more. At Arcachon the oysters spawn every 2 or 3 weeks from mid-June to mid-August with intermittent periods of gonad restoration (His, 1976); from mid-August to the end of September, bioassays can be conducted here without laboratory conditioning by using unspawned ripe oysters from Brittany and Normandy. 2.1.2. The Eastern oyster Crassostrea virginica The Eastern oyster has its natural distribution on the Atlantic coast of North America from the Gulf of St Lawrence to the Gulf of Mexico and the West Indies (Ahmed, 1975; Hayes and Menzel, 1981; Buroker, 1983 and 1985; Kennedy, 1983). It has repeatedly been introduced for aquaculture purposes to the Pacific coast of North America (Andrews, 1980) and to the Baltic coast of Denmark and Germany since the late nineteenth century (Seaman and Ruth, 1997), but has never become established in Europe. It has, however, been introduced successfully to Hawaii. Loosanoff and Engle (1942) already suspected the existence of different physiological races, because of variations in spawning behaviour. Stauber (1950) and Loosanoff and Nomejko (1951) subsequently established the existence of three different races in the northern, central and southern areas of its distribution, with spawning thresholds at 17"C, 20°C and 25"C, respectively. Similarly, C. virginica from Long Island Sound may mature at 12"C, whereas those from Florida do not ripen at 18°C (Loosanoff, 1968). The spawning season is in late June and early July in the Gulf of St Lawrence, and from mid-June to early August at New Haven (Stiles and
THE ASSESSMENT OF MARINE POLLUTION
13
Longwell, 1973). In Chesapeake Bay it may last until the end of August (Kennedy and Krantz, 1982), and in Florida it lasts from March to October (Ingle, 1951). In Hawaii, C. virginica reproduces from February to November (Sakuda, 1966). 2.1.3. The European oyster Ostrea edulis The European oyster was originally distributed from Norway to Morocco and the Mediterranean (Walne, 1965; Ahmed, 1975; see also Korringa, 1941, 1958; Cole, 1941, 1942). It was introduced experimentally in 1949 to various estuaries in Maine (United States), where it reproduced, notably establishing itself at Boothbay Harbor (Loosanoff, 1955). It was also transplanted to the western coast of the United States with moderate success (Matthiessen, 1970). European oysters generally reproduce at temperatures of 15°C to 20°C (Buroker, 1985). According to the differences in spawning temperature, various physiological races exist (Korringa, 1958), but their distribution is not necessarily linked to geographic latitude. In Galicia (northwest Spain) some stocks spawn at temperatures as low as 12°C to 13”C, whereas in certain Norwegian fjords (where a superficial freshwater layer creates a greenhouse effect) the European oyster spawns at 25°C (Yonge, 1960; Wilson and Simons, 1985). Along the Atlantic coast the spawning season may last from early spring to November or December (Yonge, 1960). In the Adriatic, its larvae are encountered from April to October (HrsBrenko, 1982). 2.1.4. The mussels Mytilus edulis and M. galloprovincialis The genus Mytilus is widely distributed throughout the world (see Figure 1.6 in Gosling, 1992); its taxonomic status has often been investigated and discussed (Seed, 1971, 1976; Gosling, 1984, 1992; Lubet et al., 1984; Brock, 1985; MacDonald et al., 1991). Mytilus edulis and M . galloprovincialis are often sympatric, characterized by considerable morphological variability. They frequently interbreed, giving rise to intermediate phenotypes; where they coexist, the former is more frequently found in exposed, and the latter in more protected areas (Lubet, 1959). Their spawnings rarely coincide (Lubet, 1973; Seed, 1976). According to Gosling (1984) morphological, cytological, immunological, electrophoretic and hybridization studies show that M. galloprovincialis is “a form or ecotype” of M . edulis. There are no major differences between the chromosomes of the two forms (Dixon and Flavell, 1986). For reasons of convenience, however, they are still often referred to as separate species in the literature.
14
E. HIS,
R. BEIRAS AND M. N. L. SEAMAN
Both forms are very widely distributed. M. edufis is boreal, but eurythermic (Lubet, 1973), and is distributed from northern Norway (where summer temperatures hardly surpass 4°C to 5°C) to the Bay of Biscay (where summer temperatures may reach 23°C at Arcachon). It is found on both coasts of the North Atlantic, including the North and Baltic Seas (Gosling, 1992), but it is rare in Spain and Portugal, and very infrequent in the Mediterranean (Lubet et af.,1984; McDonald et af.,1991; Gosling, 1992). M. galloprovincialis is distributed from southern Britain to the northern coast of Africa, and the Mediterranean and Black Seas. According to Gosling (1992) M. trossufus,another ecotype of M. edufis, has a more northerly distribution than either M. edufis or M. gaffoprovincialis, and is found only in the northern hemisphere. The range of M. cafifornianus, considered to be a true species distinct from the M. edufis group, is restricted to the northeast Pacific, but is continuous over a latitudinal range of about 30 degrees. M. trossufus and M. cafifornianusare almost never used in bioassays. In many areas mussels may maintain sexual maturity and spawn repeatedly throughout the year. It is generally accepted that the reproductive period is shorter in cooler regions, and that it is more extended in M . galloprovincialis than in M . edulis (Seed, 1971, 1976). Spawning may generally be obtained in M. galloprovincialis throughout the year, except in unusually cold winters, when reproductive activity may cease altogether. The spawnings of greatest relevance for recruitment are in early spring and in fall, although the fall spawning may be late, or lacking altogether, in the northern range of its distribution (Lubet, 1973). In M. edufis on the coasts of Europe, the principal spawnings take place from March to early June. This is followed by a non-reproductive period during the summer, and gametogenesis resumes in October or November and continues through the winter (Bayne et af., 1975). Mussels from warmer waters generally spawn earlier, although this was not found to be the case in seven M. edulis populations on the east coast of the United States; Bayne’s rule on day-degrees did not apply either (Newel1 et af., 1982). 2.1.5. The hard clam Mercenaria mercenaria The northern quahog, or hard clam, is distributed on the Atlantic coast of North America from the Gulf of St Lawrence to the northern Gulf of Mexico, and it is particularly abundant from Maine to Virginia. In the southern part of its range, it is sympatric with M. campechiensis, with which it may interbreed. M. mercenaria has been introduced to the coast of California (Ansell, 1968; Stanley and DeWitt, 1983). It has also been introduced to Europe, forming wild stocks in the areas of Portsmouth and Southampton in Britain (Mitchell, 1974), but despite various introductions
THE ASSESSMENT OF MARINE POLLUTION
15
to France, significant populations have only become established in the Marennes-OlCron area and in Brittany. Small stocks also exist on the Dutch and Belgian coasts (Tebble, 1966). The reproductive cycle of this species has been reviewed by Stanley and DeWitt (1983), with additional descriptions by Keck et al. (1975), Dalton and Menzel (1983), Knaub and Eversole (1988), Manzi et al. (1985), and Walker and Heffernan (1995). The sexual cycle has been described for populations of Long Island Sound (Loosanoff, 1937a,b), Delaware Bay (Keck et al., 1975), North Carolina (Porter, 1964), South Carolina (Eversole et aE., 1980), and northwest Florida (Dalton and Menzel, 1983; Eversole, 1989). M . rnercenaria generally has two breeding peaks in North and South Carolina, where spawning occurs at 27°C to 30°C (Porter, 1964; Eversole et al., 1980). This bimodal pattern is not found at Delaware Bay, where spawning takes place at 25°C to 27°C (Keck et al., 1975), nor at Long Island Sound, where the temperature for spawning is 23°C to 25°C (Loosanoff, 1937a,b). In Britain the species has adapted to the colder climate and spawns at 18°C to 20°C (Mitchell, 1974). In contrast, Dalton and Menzel (1983) describe a trimodal spawning pattern in northern Florida, with peaks in autumn, winter and spring. Laughlin et al. (1988) assert that reproduction occurs year-round and that mature spawners are available without laboratory conditioning throughout the year. The existence of different spawning temperatures for different stocks has led Porter (1964) and Keck et al. (1975) to hypothesize the existence of different races. In summary, M. mercenaria does not seem to have a sexually inactive season, and individuals at various stages of gametogenesis can be found at all times, although the proportions vary greatly between locations. 2.1.6. The coot clam Mulinia lateralis The coot clam is also distributed on the eastern coast of North America from Canada to the eastern Gulf of Mexico (Kennedy and Mihursky, 1971; Calabrese and Rhodes, 1974; Morrison and Petrocelli, 1990; Burgess and Morrison, 1994). The species is not very abundant, except at certain favourable locations (Calabrese, 1969a,b, 1970a). According to the presence of the larvae in the plankton, the reproductive period is from mid-July to early December at Prince Edward Island, Canada (Sullivan, 1948); the larvae are found from May to November at Tred Avon River, Maryland (Shaw, 1965). At Long Island Sound, gametogenic activity continues throughout the year, with principal spawning peaks in late July and mid- to late August, the first peak being at temperatures near 20°C (Calabrese, 1970a; Calabrese and Rhodes, 1974).
16
E. HIS, R. BEIRAS AND M. N.
L. SEAMAN
This confirms the observations of Sullivan (1948) and Loosanoff et al. (1966).
2.1.7. Other species At least 25 other bivalve species have also been used in bioassays (see Table l), but most of them do not fully satisfy the criteria required (Stebbing et al., 1980). Although some of these species are of economic importance, their geographic distribution is generally limited. This applies in particular to the mussels Mytilus californianus and M . trossulus, the oysters Crassostrea angulata, C. cuccullata, C. margaritacea, C. rhizophorae, Saccostrea commercialis and Ostrea plicatula, as well as to the Hawalan species, Zsognomon californicum, which has been studied in detail by Ringwood (1990, 1991, 1992a,b, 1993) with regard to the micropollutant sensitivity of adults and larval stages.
2.2. Reproduction
Experiments with larvae depend first of all on the availability of spawning individuals of both sexes. The bivalve species commonly in use for bioassays have a very wide geographical distribution, but the availability of mature adults may nevertheless be highly variable. Following the studies by Orton (1927) on Ostrea edulis, Coe (1932a,b) on Crassostrea virginica and Loosanoff (1937a,b) on Mercenaria mercenaria, there have been numerous publications on bivalve reproduction (reviews by Galtsoff, 1964; Raven, 1964; Purchon, 1968; Sastry, 1975,1979; Seed, 1976; Andrews, 1979; Mackie, 1984; Gosling, 1992). 2.2.1. Gametogenesis The reproductive anatomy of bivalves is rather simple. The gonad consists of a mass of follicles which develop fully at the period of sexual maturity, at which time the sexual products make up a significant part of the body. The sexual products grow within genital ducts, the diameters of which increase progressively during gametogenesis; the various small ducts converge on larger gonoducts, through which the gametes are expelled into the exhalent part of the pallial cavity at the time of spawning. The simplicity of the reproductive organs, notably the absence of specialized structures (such as a penis in the male or accessory glands in the female) facilitates the change of sex observed in many bivalve species. In oysters, during the winter, when there is no reproductive activity, the
THE ASSESSMENT OF MARINE POLLUTION
17
gonad mass is replaced by a mass of connective tissue with vesicle cells containing lipids and glycogen. Embedded in this and close to the internal organs is a duplicate system of branching tubules, one on each side, beginning near the anterior end of the body, uniting into a single tube on either side and ending in the genital pore. These pores open into the suprabranchial chamber at the posterior base of the adductor muscle, in close proximity to the opening of the urinal ducts (Quayle, 1969). During the breeding season the reproductive organs form at least 50% of the body volume. In a fully ripe oyster, the gonadal tubules, small in diameter at the anterior end of the oyster and thickening as they approach the genital opening, may be clearly seen on the surface of the soft body of the oyster. At this time, the two gonadal systems are almost completely inseparable, except at the genital pores. In mussels the gonad extends in a diffuse manner throughout the mantle. It is made up of a multitude of acini which are grouped around tubules, which are in turn clustered around the gonoducts. These fuse to form the terminal gonoducts on either side of the body, which then unite and open into the genital papilla near the posterior adductor muscle. In venerid clams such as Mercenaria mercenuria, the gonad is very diffuse and located around the digestive gland, sometimes covering it entirely and extending into the foot by the time maturity is attained. The genital pores open into the exhalent part of the pallial cavity and the gametes are released by the exhalent siphon. In Muliniu lateralis, the gonad of ripe animals forms a uniform and continuous mass around the digestive tube and gland (Calabrese, 1970a, b). Most lamellibranchs (96%) are gonochoric, i.e. they have separate sexes (Coe, 1943), but there are various hermaphroditic species, including oysters. With regard to the hermaphrodites, the following classifications have been proposed (Table 2). Crussostrea gigus and C. virginica are both alternative hermaphrodites (Amemiya, 1929; Coe, 1932a,b), changes in sex occurring irregularly during the course of life. Most individuals only emit sexual products corresponding to one sex in the course of the reproductive season, but there is a small proportion of simultaneous hermaphrodites (Loosanoff, 1965a). Crassosfreu is usually protandric, about 70% of the individuals male in their first year, and about half in the second year, with females dominant in the older age groups. In Ostreu edulis, which is larviparous, protandric sex changes are the rule (Sparks, 1925; Orton, 1927). The time lag between the maturation of spermatocytes and oocytes is small, however, and European oysters may alternately function as males and females in the course of one reproductive season. The frequency of sex changes increases with temperature, so
18
E. HIS, R. BEIRAS AND M . N. L. SEAMAN
Table 2 nYo systems of classifying hermaphroditisms in Bivalvia (from Fretter and Graham, 1964).
Coe Functional or simultaneous Consecutive Rhythmical consecutive Alternative
Bacci
Simultaneous (with synchronous ripening) Successive (with asynchronous ripening) or consecutive Successive (with asynchronous ripening) or alternate Successive (with separate ripening) or alternate
it is related to latitude (Cole, 1942). The change from the female sex to the male is more rapid than vice versa. Mytilus edulis and M. galloprovincialis are gonochoric, even though rare cases of simultaneous hermaphroditism have been reported by Coe (1943) and Lubet (1959). According to the latter, hermaphroditism occurs in about 0.02 to 0.1% of the population. Changes of sex by the same individual have never been observed. Mercenaria rnercenaria is a protandric hermaphrodite (Eversole, 1989), but the simultaneous development of oocytes and spermatocytes is common during the first year of life (Loosanoff, 1937a,b), and a small percentage of simultaneous hermaphrodites is always present in the population. Nevertheless, 98% of the individuals are male during the first year, but the sex ratio becomes even with advancing age. Mulinia lateralis is gonochoric. The reproductive cycle is very brief, the egg-to-egg cycle being only 39 to 135 days with an average generation time of 60 days (Calabrese, 1970a). Individuals as small as 2.7 mm in length are already capable of spawning (Calabrese and Rhodes, 1974). Another practical aspect of this species is that the sexes are readily distinguishable, because the orange-red female gonad and the whitish male gonad are discernible through the shell near the umbo. Galtsoff (1964), Bayne (1976) and Mackie (1984) have described the structure of bivalve gametes in detail. In the males the chromosomal reduction (first order spermatocytes with 2n to second order spermatocytes with n chromosomes) is achieved by an equal division during spermatogenesis. In the female, however, the gametes are spawned at the germinal vesicle stage and only develop to the metaphase of the first meiotic division: meiosis is arrested until the egg is activated by sperm. Penetration of the ovum by the sperm is made possible by the acrosomal reaction (i.e. dissolution of the egg membrane by the sperm’s acrosome), after which the sperm cytoplasm fuses with the egg cytoplasm; this occurs within about 5 minutes in Mytilus edulis. Meiosis in the egg (Figure 1)then
THE ASSESSMENT OF MARINE POLLUTION
19
Figure 1 Normal development in the bivalve egg. For simplification only one pair of chromosomes is shown. (a) egg at release at metaphase of meiosis I, activation by sperm; (b) meiosis I is complete, first polar body extruded, sperm nucleus has entered egg; (c) meiosis I1 completed, second polar body extruded, male and female pronucleus unite; (d) first cleavage perpendicular to point of polar body extrusion. (From Beaumont and Fairbrother, 1991.)
continues with the formation of the first and second polar bodies, after which the egg’s chromatin forms the chromosome vesicle. This is followed by the fusion of the male and female chromosome vesicles and the formation of the 2n pronucleus (Longo and Anderson, 1969a,b; Bayne, 1976; Mackie, 1984; Cherr et al., 1990). 2.2.2. Sexual maturation in the field Spawning in marine invertebrates may occur year-round in regions with little seasonal variability (Sanders and Hessler, 1969, cited after Dalton and Menzel, 1983). According to Rand (1973), cool climates are characterized by species with a single annual spawning season, temperate zones by species with two separate spawning periods, and tropical areas by species with year-round spawning. The determination of sexual cycles has been based on direct observation of spawning activity in the natural environment, on determinations of the state of maturation of the gonad, on the appearance in the plankton of larvae of the species in question, and on the settlement of the juveniles (Seed, 1976). The reproductive cycle of bivalves can generally be divided into three phases: gametogenesis and vitellogenesis, spawning and fertilization, and larval growth and development (Newel1 et al., 1982). With regard to the maturation state of the gonad, a variety of scales have been proposed by Chipperfield (1953), Lubet (1959), Seed (1969) in mussels, Loosanoff (1942) and Kennedy and Battle (1964) in C. virginica, and Keck et al. (1975) and Eversole et al. (1980) in Mercenaria mercenaria. 2.2.2.1. Environmental factors influencing gametogenesis. Mackie (1984) has reviewed the principal factors governing sexual maturation in bivalves. They are either exogenous, such as temperature, lunar cycle, and,
20
E. HIS, R. BEIRAS AND
M. N.
L. SEAMAN
particularly, nutritional factors, or endogenous, such as genetic or hormonal factors. Among the external factors, Orton (1920) considered temperature the most important influence. Loosanoff and Nomejko (1951) were the first to advance a hypothesis on the existence of physiological races in Crassostrea virginica with different temperature requirements for spawning. The issue was taken up again by Ahmed (1975) with respect to oysters. As will become apparent in the next section, laboratory “conditioning” of tachydictic species, i.e. the possibility of inducing sexual maturation out of season, largely depends on an elevation of the temperature. Even though this applies to most bivalve species used in bioassays, it must be emphasized that mussels, to the contrary, reproduce at low temperatures. Bayne (1975) established a day-degree function for the relationship between temperature and duration of gametogenesis in Mytilus edulis. The same observation was made by Mann (1979) with regard to C. gigas, and by Knaub and Eversole (1988) in Mercenaria rnercenaria. In eurythermic species with high temperature tolerance, the duration of the reproductive cycle often varies with geographical latitude. Nutritional factors are also of paramount importance. Various authors have shown that bivalves accumulate nutritional reserves in their connective tissue at the time of sexual inactivity, in order to mobilize them towards the formation of sexual products during gametogenesis (see Mackie, 1984). The nutritional needs of mussels increase during gonad maturation (Bayne, 1975); stress and bad nutritional conditions, on the other hand, are accompanied by a reduction in fertility, with a decrease in the number of gametes produced and a modification of their biochemical composition (Bayne et al., 1975,1978; Bayne, 1985). A rise in temperature and stress from a simultaneous decrease in available food, and the influence of pollutants such as hydrocarbons also lead to lower gamete quality and larval viability. In C.virginica starvation leads to a decrease in the proportion of females (Bahr and Hillman, 1967). An increase in the proportion of males in a population of oysters may therefore be an indication of environmental disturbance (Kennedy, 1983). Among the endogenous factors, neurosecretion was first shown to be important in mussels by Lubet (1959), and in oysters by Nagabhushanam (1963). Genetic factors may also be important (Lannan et al., 1980). 2.2.2.2. Environmental factors influencing spawning behaviour. Male oysters are more sensitive to stimulation and usually spawn first. The first males induce spawning in their male neighbours, then the first females follow suit, and finally the entire population spawns simultaneously. This results in the formation of “spawn streaks” described by Quayle (1969), milky masses of water several hundred metres in length spreading in the channels of bays and estuaries. Among the environmental factors which
THE ASSESSMENT OF MARINE POLLUTION
21
induce spawning behaviour are the thermal stimulation resulting from abrupt differences in temperature between the water masses of the falling and the rising tide, changes in salinity during the tidal cycle, mechanical agitation by waves and currents, and differences in pressure between high and low water and during the falling tide. Spawning in the field is never provoked by one single stimulus; when an appropriate combination of stimuli occurs, spawning is most common during the 3 hours following high tide (58% of all spawnings), and it is more frequent (69%) and more prolonged (76% of the duration of all spawnings) at spring tides (His, 1976). In flat oysters, “swarming” (i.e. the release of larvae incubated within the mother’s pallial cavity) is also related to the lunar cycle: from July to September larvae are most abundant in the plankton following a full moon (Orton, 1926), or 10 days after a full or new moon (Korringa, 1941; Knight-Jones, 1952).
2.2.3. Cleavage, embryogenesis and larval development Bioassays may be conducted with gametes and larvae of all stages, from sperm and unfertilized eggs to metamorphosis of the pediveliger at the end of pelagic life. Many researchers indicate the time passed between fertilization and the fertilized gametes’ first exposure to the substance being investigated, as this may have a bearing on the interpretation of the results. The rates of development and cleavage, and the duration of the successive embryonic and larval stages differ widely between the various bivalve species, and they also depend largely on environmental factors, notably on the temperature. The development of bivalves consists of an embryonic phase followed by a larval phase, and has been described by various authors (Erdmann, 1935 in 0. edulis; Cahn, 1950 in C. gigas; Galtsoff, 1964 in C. virginica; Lubet, 1973 and Bayne, 1976 in M. edulis). More recent work by Elston (1980) on C. virginica and by Waller (1981) on 0. edulis and C. gigas have provided additional details of larval morphology, particularly demonstrating the complexity of various larval organs. The common terminology employed for the various development stages is shown in Figure 2. 2.2.3.1. Embryonal stages. The fertilized egg (see Figure 1) completes meiosis by expelling the first and second polar bodies, and then begins to divide. The first division is unequal, giving rise to the blastomeres AB and CD, as well as a polar lobe which fuses with CD. The second division (4-cell stage) leads to the formation of three blastomeres (A, B and C) at the animal pole and one very large blastomere (D) at the vegetal pole. During the third division, the cleavage becomes spiral and subsequent
22
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
TERM EGG BLASTULA GASTRULA TROCHOPHORE VELIGER 0-SHAPED STAGE UMBO STAGE PEDIVELIGER SPAT
PRESHELLED PRODISSOCONCH I STAGE (less (about 7 days) than 1 day)
DISSOCONCH (remainder of life)
PRODISSOCONCH II (10-15 days)
--
--...
...
--...
....
....-
-
Relationship of stages of development of the prodissoconch to other common terms used to describe the larval shell and body of Ostrea edulis (dashed lines indicate uncertainty or transition; duration of stages may be highly variable). (From Waller, 1981.)
Figure2
divisions lead to the formation of the morula. Cilia appear, and the embryo becomes motile, gyrating slowly. Gastrulation begins as the micromeres of the animal pole cover the blastula, followed by a slight invagination that gives rise to the archenteron and to a small blastopore. 2.2.3.2. Larval stages. The first larval stage is the trochophore, which is covered by short cilia; in mussels and some other species it also has a flagellum. A dorsal thickening of the ectodermis is secreted by the shell gland, forming the initial organic cuticle which eventually spreads to cover the entire body. A circular band of cilia, the prototroch, also forms, enveloping the apex. The second larval stage is the veliger, which is formed 24 hours after fertilization in Crussostrea, and after 48 hours in Mytilus. It has a straight dorsal hinge giving the larva the characteristic shape of a capital letter D (hence the synonymous terms “straight-hinge larva” and “D-larva”), and it is some 60 to 70 pm in size in most bioassay species (see below). At this stage the larva begins to feed. The shell, consisting of two valves, begins to calcify, forming the prodissoconch I. The prototroch has continued its development to form the velum. As the larva grows beyond 100 pm, secretion of the secondary shell (the prodissoconch 11) begins and the umbo begins to form, extending over the dorsal hinge (hence the terms “veliconcha” and ‘‘urnboned larva”). In oysters and mussels, as the larva grows to a size of 250 or 300 pm, an “eye spot” appears within the shell and the foot is formed (hence the terms “eyed larva” and “pediveliger” for this stage). As the ciliated foot grows and becomes functional, the larva becomes capable of both pelagic and benthic
THE ASSESSMENT OF MARINE POLLUTION
23
modes of life, either swimming by use of the velum (e.g. to feed), or creeping along a hard substrate with its foot (e.g. to explore for a suitable site for settlement). Once it has attained this stage it is ready to metamorphose. All of the various stages mentioned above are used in bioassays. 2.2.3.3. Settlement and metamorphosis. The first photographs of settlement and metamorphosis in Crassostrea virginica were made by Prytherch during the 1930s, and some of his images were reproduced by Medcof (1961). The fixation of pediveligers of Ostrea edulis has been described by Cranfield (1973). The first attachment stage after metamorphosis is the plantigrade or juvenile (Figure 3). Oysters settle by glueing their shell to the substratum with a proteinaceous cement produced by the pallial gland (Figure 4). Mussels and clams attach themselves to the substratum with the help of byssal threads produced by the byssal gland at the base of the foot. During metamorphosis the velum disappears, and in oysters the foot is resorbed as well; in mussels and clams, the foot continues to develop and specialize to the adult form. Labial palps and gills are developed to replace the larval feeding apparatus, the velum, and the final shell, the dissoconch, is secreted. The anatomical changes of metamorphosis may make mussel larvae unable to feed for 1 to 3 days (Bayne, 1976). In Crassostrea virginica, Baker and Mann (1994b, p. 239) reported that “velar feeding occurred during the searching and crawling stages, but not during cementation”, and subsequently, Baker and Mann (1994a, see Figure 1, p. 94) identified four separate phases of metamorphosis: settler, prodissoconch postlarva, dissoconch postlarva and juvenile. Various studies have been devoted to the problems of settlement and metamorphosis in bivalves (see Ritchie and Menzel, 1969;Lutz et al., 1970; Hidu and Haskin, 1971; Keck et al., 1971; Veitch and Hidu, 1971; Andrews, 1979; Mackie, 1984; Baker and Mann, 1994a,b). Prytherch (1924,1931 and 1934) found that copper induces metamorphosis in larvae of C. virginica. Since then, a great number of publications have documented natural and artificial chemical substances capable of inducing metamorphosis (see Hadfield, 1984; Pawlick, 1992). The special role of neuroactive compounds (L-DOPA, dopamine, serotonin, epinephrine, norepinephrine) has been shown by Coon and Bonar (19861, Coon et al. (1985, 1986, 1990), Shpigel et al. (1989) and Beiras and Widdows (1995), and the importance of biofilms has been demonstrated by Weiner et al. (1989). In oysters, neurotransmitters may induce metamorphosis without fixation, producing “cultchless spat” which are anatomically identical to naturally set spat (see photographs in Coon et al., 1985, p. 217; Coon and Bonar, 1986). These advances are of considerable practical significance in toxicological bioassays with pediveligers (see Section 3.2.4.5).
24
E. HIS, R. BEIRAS AND M .
N. L. SEAMAN
Figure 3 Crassostrea gigas pediveliger metamorphosing (a), and metamorphosed post-larva (b). Gr: gill rudiment; F foot; P d prodissoconch shell; D: dissoconch shell. (Experiments by His et al., 1997b.) Metamorphosis was induced by epinephrine.
25
THE ASSESSMENT OF MARINE POLLUTION
2 Larva begins swimlcrawl
Larva searches for surface cue
Larva detects
Cementation to substratum
Metamorphosis to juvenile oyster
Figure 4 Two-cue model of microbial induction of oyster settlement (1-3) and metamorphosis (45). A soluble cue induces searching behaviour and then a surface cue induces attachment. (From Weiner et al., 1989.)
2.3. Larval rearing in the laboratory
2.3.1. Laboratory conditioning of spawners To conduct routine bioassays with bivalve embryos and larvae it is desirable to be able to obtain ripe adults year-round. In some species, mature adults can be found in the field during many months, and in others it is possible to condition them out of season. This was first discovered by Loosanoff (1945), when he attempted to clear Eastern oysters from a growth of sponges by keeping them in running seawater at elevated temperatures and found that by doing so he had induced gametogenesis in winter. He subsequently demonstrated that the duration of conditioning can be prolonged or shortened by manipulating the temperature, and that the technique can be applied to Mercenaria mercenaria as well (Loosanoff and Davis, 1950, 1951, 1952, reviewed 1963). In the case of Crassostrea virginica,gonad maturation can be obtained in the winter by gradually raising the water temperature to 20°C or 25°C; it is even possible to transfer oysters directly from a temperature of 5°C in the field to 20°C in the laboratory. The duration of conditioning depends on the temperature, as long as it is maintained within the range of 15°C to 30°C. Prolonged incubation at 27°C to 28°C can, however, inhibit or delay spawning (Ruddy et al., 1975). At the end of the reproductive season, after the adult oysters have used up their reserves, it is difficult or impossible to induce them to mature
26
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
again immediately. To avoid this inconvenience, Loosanoff and Davis (1963) took maturing oysters from Long Island Sound and transferred them to the cooler waters of Maine, where their maturation was arrested. They were subsequently able to induce final maturation and spawning by manipulating the temperature in the laboratory. Spawning can be delayed this way for 6 to 8 weeks; beyond this time limit, progressive resorption of the gonad makes spawning impossible. To circumvent the problem, Loosanoff and Davis induced oysters to spawn early and then transferred them also to Maine, where they resorbed the residual sexual products and reconstituted their glycogen reserves, thus beginning a new reproductive cycle and becoming susceptible to conditioning again. This two-sided approach to conditioning (acceleration and inhibition of gametogenesis) can be applied to C. virginica and M. mercenaria (Davis and Chanley, 1955a; Loosanoff and Davis, 1963), and also to C. gigus, and it is of great practical relevance to toxicological bioassays. In Mufinia faterafis it is equally possible to obtain mature adults throughout the year by delaying or accelerating sexual maturation with the same manipulations of temperature (Calabrese and Rhodes, 1974); apparently it works even better with the coot clam than with the other species (Burgess and Morrison, 1994). Conditioning has also been achieved with Ostrea edufis (Dannevig, 1951; Abou-Ela, 1960), where adults can be transferred from environmental temperatures of 1°C to 5°C to conditioning tanks at 21°C (Walne, 1966). Supplementary feeding with cultured algae (Tetrasefmis suecica) during conditioning improves the subsequent survival of the larvae (Helm et af., 1973). It is equally possible to delay spawning in mussels after initiation of gametogenesis by transferring them to 4°C or 5°C one month before the time of spawning and by supplementing their nutrition with cultured phytoplankton (Bayne, 1965, 1975; Riisgard et af., 1980; Dixon and Prosser, 1986). Mussels have been thermally conditioned by Mitchell et al. (1985) by maintaining them for 3 weeks in unfiltered seawater at 14°C. Spawning was obtained by Bayne (1975) one month early in adults which had initiated gametogenesis by keeping them at 15°C (5°C higher than the ambient temperature) and by feeding them with Tetrasefmis suecica (at least 2.2% of the meat weight per day). The time required for gonad maturation depends on the stage of maturation at the onset of conditioning as well as on the conditioning temperature. On the other hand, stress and sub-optimum rearing conditions (particularly with respect to nutrition) reduce the fertility of mussels, as well as the quality of the gametes and larvae (Bayne, 1975, 1985). The methods for conditioning scallops, oysters and clams in hatcheries have been reviewed by Utting and Millican (1997): “of particular
27
THE ASSESSMENT OF MARINE POLLUTION
Table 3 The fecundity of bivalves used in bioassays. Species
Crassostrea gigas Crassostrea virginica Mytilus edulis
Mytilus galloprovincialis Mercenaria mercenaria
Mulinia lateralis Ostrea edulis
Fecundity (eggs per female; x106) 15 to 114 15 to 114.8 10 to 66.4 (M = 28.8) 10 >0.5 1.2 to 7.6 2 no data 0.38 to 18.83 0.6 to 13.2 (A4 = 6.3) 8 to 39.5 ( M = 24.6) 1.4 >7 (M = 3 to 4) 0.09 to >1 0.616 to 1.155 0.1 (1year old) to 1.5 (7 years old)
Reference Galtsoff, 1964 Galtsoff, 1964 Davis and Chanley, 1955a Lubet, 1959 Bayne, 1975 Bayne et al., 1978 Thompson, 1979 Lubet, 1973 Ansell, 1967 Bricelj and Malouf, 1980 Davis and Chanley, 1955a Knaub et al., 1987 Calabrese, 1969a,b Cole, 1941 Millar, 1961 Walne. 1974b*
M: mean value; *estimated from the number of incubated larvae.
importance is the optimization of the quantity and the quality of microalgae diets that are provided during the broodstock conditioning”. Flow-through systems are generally better than recirculation systems, because “natural phytoplankton is a valuable component of the diet during broodstock conditioning” (p. 46). Gametogenesis is improved by preconditioning in high food regimes before the actual conditioning at elevated temperature. Egg quality and larval viability depend largely on an adequate accumulation of lipid reserves in the ova. 2.3.2. Spawning
To have a suitable number of replicates, or to be able to study a range of concentrations in toxicity tests, a great number of larvae are often required in bioassays. Most of the bivalves used in bioassays are characterized by high fertility. Flat oysters are an exception; being larviparous, not only are they less fecund, but they also have much larger oocytes (100 to 150pm in diameter in Ostrea edulis) than the other species. The data on fecundity (Table 3) are incomplete, especially with regard to Mytilus galloprovincialis. The data on oysters have been reviewed by Galtsoff (1964), those on Mercenaria mercenaria by Davis and Chanley
28
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
(1955a) and by Eversole (1989), Mytilus edulis by Thompson (1979), and Mulinia lateralis by Calabrese (1969a,b). The number of eggs released by any individual always depends on its size and physiological condition, as evidenced by significant correlation between size and number of eggs spawned in Crassostrea virginica and M. mercenaria. Stress and pollution, however, are known to reduce the fertility in mussels (Bayne, 1972; Zaroogian et al., 1979; Zaroogian and Morrison, 1981). The reproductive physiology of Crassostrea virginica has been studied in detail by Galtsoff (1938a,b, 1940, 1964). He determined that spawning in female oysters involves complex mechanisms with the participation of the nervous system, the gills, the mantle and the adductor muscle. The genital pore is situated near the anus. Oocytes are expelled into the exhalent part of the pallial cavity, then pass through the gills and accumulate between the gills and the mantle before being expelled by muscular action through the inhalant part of the pallial cavity into the surrounding water. The shell movements associated with spawning are “so characteristic that they cannot be mistaken for any other type of muscular activity” (Galtsoff, 1964), and they can serve to monitor the frequency and duration of reproduction in the field (His, 1975, 1976). Ova are violently expelled at regular intervals, forming whitish clouds near the spawning female. In the male, however, the sperm leave as a steady stream with the exhalent current, almost without contraction of the adductor muscle (Galtsoff, 1938a,b). In the water column, the gametes emit pheromones (termed “fertilizins” by Galtsoff, 1938a,b, 1940) which induce spawning in members of the opposite sex. Sperm also release a nucleoprotein termed diantline which facilitates spawning in both sexes by relaxing the adductor muscle, enlarging the gill ostiae, augmenting ciliary activity, and thus increasing the pumping rate (Nelson and Allison, 1940). These observations probably apply to all oysters of the genus Crassostrea (Galtsoff, 1964). The presence of similar pheromones has been demonstrated in mussels by Lubet (1959). In Ostrea edulis the ova also pass through the gills into the inhalant part of the pallial cavity, but this does not give rise to any visible change in behaviour (Yonge, 1960). The ova are fertilized by sperm inhaled by the female and are subsequently retained within the pallial cavity for about one week, after which they are released (a phenomenon termed “swarming” by Dutch and German authors). In Mercenaria mercenaria and in Mytilus, sperm are liberated with the exhalent current in a whitish thread-like stream which rapidly dissipates and gives the water a milky appearance. The ova are similarly liberated by the females without particular valve movements. In the laboratory, spawning can be induced in mature bivalves by a
THE ASSESSMENT OF MARINE POLLUTION
29
variety of physical and chemical stimuli (Table 4, after Le Pennec, 1981; see also Mackie, 1984). In the case of Eastern oysters from South Carolina and Florida it is not possible to induce spawning in the laboratory by the usual methods (thermal stimulation and addition of sperm suspension), because they are often subjected to these stimuli for extended periods in their habitat (Maurer and Price, 1968). Gametes may be obtained from individuals that refuse to spawn naturally by “stripping” (i.e. teasing the gonad with a forceps), however this is not generally recommended (Woelke, 1961, 1966; ASTM, 1989; Widdows, 1993). These methods are discussed in Sections 3.1.3.2 and 6.3.2. 2.3.3. Fertilization In the oviparous species, with the notable exception of Mercenaria mercenaria, the eggs are somewhat irregular in shape at the time of release, becoming spherical after a few minutes. Egg sizes (Table 5) do not vary much among species, with diameters of 50 to 55 pm in Crassostrea, 60 to 70 pm in Mytilus, 70 to 73 pm in M. mercenaria, and about 50 pm in Mulinia lateralis (Lubet, 1959; Yonge, 1960; Loosanoff and Davis, 1963; Galtsoff, 1964; Fretter and Graham, 1964; Calabrese and Rhodes, 1974; Purchon, 1968; Mackie, 1984; Eversole, 1989). In M. mercenaria the egg has a gelatinous envelope which swells to a diameter of 163 to 170 pm,and which may persist to the blastula stage (Loosanoff and Davis, 1950). Fertilization is practically instantaneous when the egg meets the spermatozoa (Allen et al., 1988). According to Lu (1986, in Allen et al., 1988) in Crassostrea gigas the second polar body is extruded 50 minutes after fertilization at a temperature of 18”C, after 43 minutes at 2VC, and after 32 minutes at 25°C. 2.3.3.1. Polyspermy. Galtsoff (1964, p. 343) states, “A few seconds after the sperm head touches the egg surface a thin transparent fertilization membrane is elevated”. Alliegro and Wright (1983) have, however, questioned the existence of this phenomenon: “It is possible that the fertilization envelope reported by Galtsoff can be attributed to the refractile nature of the fertilized egg surface seen with light microscopy.” Galtsoff also notes that two or more sperm (“supernumeraries”) may enter the egg before the barrier is developed when sperm suspensions are thick; this leads to polyspermy and results in an abnormal embryo (Turner, 1958; Loosanoff and Davis, 1963; Galtsoff, 1964). Determination and use of appropriate sperm to egg ratios significantly enhance the yields of viable larvae (Stephano and Gould, 1988). According to Stiles and Longwell (1973), in C. virginica the number of sperm has to approach 20 per egg before the incidence of chromosome and division abnormalities increases significantly, but this has been questioned by
30
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
Table 4 Experimental methods used to induce spawning in marine bivalves (completed after LE Pennec, 1981). Method
Species
Reference
Stimulation due to transport
Crassostrea gigas
Imai, 1967
Unspecified temperature fluctuations
Crassostrea virginica C. virginica Mercenaria mercenarin Mulinia lateralis
Prytherch, 1924 Wells, 1927 Loosanoff, 1937b Calabrese, 1969b
Specified temperature fluctuations
Mytilus edulis C. gigas Mytilus galloprovincialis M. edulis
Bayne, 1965 Imai, 1967 Masson, 1975 Lutz and Hidu, 1979
Temperature fluctuations and addition of gametes
C. virginica C. virginica, M. mercenaria M. mercenaria several bivalves M. lateralis M. lateralis
Davis, 1953
Constant temperature and addition of gametes
C. virginica, C. gigas
Galtsoff. 1930
Salinity fluctuations
tsognomon califomicum
Ringwood, 1990,1991,1992
Electrical stimulation
M. edulis M. edulis M. edulis
Aboul-Ela, 1960 Iwata, 1950 Sugiura, 1962
M. edulis M. edulis
Iwata, 1951a,b Morse et al., 1977; Garland et al., 1986 Loosanoff and Davis, 1963 Loosanoff and Davis, 1963 Castagna et al., 1985
Chemical stimulation KCI 0.5 M
HzOz NbOH Injection of serotonin Pricking the adductor muscle
M. mercenaria C. virginica M. mercenaria, M. edulis M. edulis
Loosanoff, 1954 Chesnut et al., 1957 Loosanoff and Davis, 1963 Kennedy et al., 1974 Calabrese and Rhodes, 1974
M. galloprovincialis
Hrs-Brenko and Calabrese, 1969 Masson, 1975
Use of "Kraft mill effluent"
M. edulis
Breese et al., 1963
Addition of algae
M. edulis, Mytilus californianus, C. gigas, M. rnercenaria M. californianus
Breese and Robinson, 1981 Smith and Strehlow, 1983
Table 5 Sue of egg, size of early veliger, sue at time of metamorphosis and duration of larval development under laboratory conditions.
Species Crassostrea gigas Crassostrea virginica Mytilus edulis Mytilus galloprovincialis Mercenaria mercenaria Mulinia lateralis Ostrea edulis
Temperature ("C) 25 24 22 20 30-32.5 16 (11)
Egg diameter
Size of early veliger
Size at time of metamorphosis
(Pm)
(Pm)
(CLm)
-
76.8 68 68-75
303-320 310 275-315
96 100-120 76
260 215-300 298 ? 23 258
50-55
?
?
18 24*
77,8 ? 4
18 ?
20-25 20-22 18-20
-
73-75 70-73 ? 50 114-126
86 60 ?
170-190*** 208
175-240 21Cb230 20fA220 240-350 280-300
Duration of larval development (4 1619 17-21 12** 36-40 10-12 1 6 2 0 (34-38) ? ?
17-19 16 (6-8**) 6 6-8 10-16 15-16
Reference Gerdes, 1983b Beiras and His, 1994 Loosanoff and Davis, 1963 Davis and Calabrese, 1964 Davis and Calabrese, 1964 Bayne, 1965 Loosanoff and Davis, 1963 Sprung, 1984 Beiras and His, 1995a Aguirre, 1979 Loosanoff and Davis, 1963 Loosanoff and Davis, 1963 Calabrese and Rhodes, 1974 Walne, 1974b Loosanoff and Davis, 1963
*egg incubation at 20°C; **33"C. Data from references in italics. ***larval sue at liberation. Mytilus edulis (Bayne, 1965): temperature and corresponding duration of pelagic life in parenthesis.
32
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
Staeger and Horton (1976) in C. gigas and Alliegro and Wright (1983) in C. virginica. In the Japanese oyster, the mean percentages of larvae developing to the D-shape stage increased until 7.3 X 10’ s p e d 1 0 0 eggs were used (Staeger and Horton, 1976), and in C. virginica the number of sperm entering eggs is restricted to one per fertilized egg at a sperm:egg ratio as high as 1OOO:l (Alliegro and Wright, 1983). In both cases gametes were obtained by stripping the gonads. In mussels, viable larvae can be obtained with variable sperm to egg ratios. The recommended sperm to egg ratio is in the order of 103:1in M. edulis (Sprung and Bayne, 1984) and M . galloprovincialis (Sedan0 et al., 1995). In M . californianus it varies between 25 and 200 sperm per egg, depending on the physiological state of the adults of both sexes (Cherr et al., 1990). In Mercenaria mercenaria the optimum ratio is between 1.3 and 2.5 X 103:1 (Bricelj and Malouf, 1980). Finally, Calabrese (1984) states that polyspermy is to be avoided in Mulinia lateralis, but without speclfying sperm densities. In their study on polyspermy in C. gigas Stephano and Gould (1988) found that, compared to eggs spawned naturally, those obtained by stripping are very susceptible to polyspermy upon immediate fertilization. This susceptibility decreases greatly when the eggs are held in seawater for 90 minutes before fertilization. The authors conclude that stripped eggs lack a “sperm block” present in naturally spawned eggs. Togo et al. (1995) obtained normal (monospermic) fertilization in Mytilus edulis when the eggs were fertilized at a sperm ratio of 5 X 103:1 within 30minutes after spawning. They recognized three mechanisms for blocking polyspermy in mussels. The first depends on a rapid depolarization of the egg’s plasma membrane, the second consists of the suppression of the acrosomal reaction, and the third is the blocking of contact or fusion between the plasma membranes of sperm and egg. 2.3.3.2. Ageing of gametes affer spawning. One of the prerequisites for bivalve embryo and larva bioassays is to begin with the best possible biological material, i.e. with a proportion of normal larvae in the controls as close to 100% as possible. “Oyster eggs undergo aging and lose their ability to be fertilized” (Galtsoff, 1964). Except in Mulinia lateralis (see Section 2.1.6), it is impossible to know in advance to which sex the individual adult belongs. Successful fertilization requires simultaneous spawning of at least one member of each sex; in general, however, one or more males spawn first, and only very rarely does a female spawn before any of the males has done so. According to Galtsoff (1964), concentrated sperm (presumably obtained by stripping) of C. virginica conserves its fertilizing capacity for at least 24 hours at lO”C,whereas diluted sperm at room temperature [probably 18°C to 20°C] loses its capacity within 4 or 5 hours. The number of dividing eggs drops to 60% after 5 hours, and to 20% after 10 hours. In C. gigas the sperm
33
THE ASSESSMENT OF MARINE POLLUTION 6-
I
0
I
20
I
I
40
L
I
I
M)
I
80
I
I
100
Hours from fertilization
Figure 5 The rates of cleavage and early development of embryos of Mytilus edulis at different temperatures. A,8°C (Bayne, 1965); 0, 18°C (Bayne, 1965); X , 20°C (Field, 1922, cited by Bayne, 1976); U, 19-22°C (Rattenbury and Berg, 1954, cited by Bayne, 1976). Stages of development as follows: 1, first polar lobe; 2, first cilia; 3, trochophore; 4,appearance of velar cilia; 5, appearance of shell gland; 6, prodissoconch I. (From Bayne, 1976.)
remain very motile for 1.5 hours after release at temperatures of 21°C to 25°C (Stephano and Gould, 1988);however, "if fertilization was delayed for more than 60 to 90 minutes after gamete liberation, the proportion of larvae which developed was greatly reduced" (Walne and Helm, 1974, p. 1; Helm and Millican, 1977, p. 2). In Mytilus edulis, diluted sperm lose their motility within 1 or 2 hours at room temperature, but maintain their fertilizing capacity for several hours at 15°C;fertilization of eggs is possible after 4 to 6 hours at 18"C, but at 15°C the fertilization rate drops to 40% within 6 to 11 hours after spawning (Sprung and Bayne, 1984). In M. galloprovincialis, fertilization was still possible within 8,7 and 4 hours after spawning at 10°C, 14°C and 18"C, respectively (Sedan0 et al., 1995).
2.3.4. Physical requirements of bivalve larvae The range of environmental conditions over which embryogenesis is possible will depend on the location of the adult population (Loosanoff, 1954; Bayne, 1976). The duration of the embryonal and larval phases depends mainly on temperature (Figure 5, Tables 5 and 6), and also on salinity and other environmental factors, including food availability
Table 6 Embryonic and larval development event times of oviparous marine bivalves. Crassostrea gigas, 25°C (Cahnn, 1950); in parenthesis, 20°C (Tazawa et al., 1985). Crassostrea virginica, 23-25°C (Galtsoff, 1964). Mytilus edulis, 17°C and 2 5 0 ~(Armstrong and Millemann, 1974). Mytilus galloprovincialis, 10-17°C (Aguirre, 1979); in parenthesis, 20°C (Masson, 1975). Mercenaria mercenaria, 22°C (Loosanoff and Davis, 1950). Mulinia lateralis, 20-25°C (Calabrese and modes, 1974).
Developmental stage
Crassostrea gigas
Crassostrea virginica
Mytilus edulis
Fertilized egg 1st polar body 2nd polar body + polar lobe 2-cell 4-cell 8-cell 16-cell 32-cell 64-cell ciliated blastula trochophore veliger
0 min 50-60 min 70 rnin
Omin 25-52 rnin 40-65 min
20 min
100 min
45-72 min 52-120 rnin 55-195 min
(420) 480 min 24h (15) 48 h
390 min 8-9 h 24 h
Omin
Mytilus galloprovincialis
Mercenaria mercenaria
Mulinia lateralis
0 min (80 min)
0 min
Omin
90 min 180-210 min
45 min 90 min
(24 h) 48 h
360 rnin 12h 24-36 h
40 min
65 min 90 min 120 min 150 rnin 180 min 220 min 450 min 16-19 h 40 h
9h 15h
Table 7 Effects of temperature, salinity and combined effects of temperature and salinity on survival and growth of bivalve 1arvae. Salinity (psu)
Temperature ("C) Species
Min.
Max.
Crassostrea gigas
15"
34"
Crassostrea virginica
17.7"
Mytilus edulis
10" 14" 5"
Opt.
Min.
Max.
15
39
30" 30-32.5"
15
15-20'
15 24 15
Opt. 25 19-27 30 2&27
T/S opt.
30"/30 30"/18-35
Mytilus galloprovincialis
22" 20" 20"
Mercenaria mercenaria
20" 12.5" 25-27.5" 2CL25" 10.5-15 25-30" 12.5"
Mulinia lateralis
20"
30"
27.5"
30"
30" 25-27"
20"
Ostrea edulis 17.5"
40 33 40
30-32 25-30 25-30 35 30-35 27
20
30
7
33
32.5 25-30
20
22.5-27
18"/27 20"/25-30 20'135 20°/30 21.5-33'122-31 22.5-27.5"/20-35 20-26'123-32 25"130
References Helm and Millican, 1977 Nell and Holliday, 1988 His et al., 1989 Davis and Calabrese, 1964 Lough, 1975 Bayne, 1965 Lough, 1974 Hrs-Brenko and Calabrese, 1969 His et al., 1989 Hrs-Brenko, 1977 Davis and Calabrese, 1964 Lough, 1975 Calabrese, 1969c Lough, 1975 Morrison and Petrocelli, 1990 Robert et al., 1989 Davis and Calabrese, 1969 Davis and Ansell, 1962
36
E. HIS, R . BEIRAS AND
M. N. L. SEAMAN
(Calabrese, 1969c; Mackie, 1984; Widdows, 1991). The time required to attain the D stage in different species is shown in Table 6. It must be pointed out that the data of Cahnn (1950) are not very accurate, because C. gigas attains the veliger stage within 24 hours, the same as C. virginica, at least at 25°C (see also Loosanoff et al., 1966; Helm and Millican, 1977; Tazawa et al., 1985). Moreover, it should be kept in mind that in bivalves, particularly in oysters, development is not highly synchronous (Stephano and Gould, 1988). The influence of various environmental effects on Crussostrea virginica larvae has been the subject of a modelling study by Dekshenieks et al. (1993). 2.3.4.1. Temperature. Embryos are generally more sensitive to environmental factors than larvae (see Section 4.2.2). Thus with regard to temperature the cleavage stages in M . edulis require a slightly narrower temperature range than the shelled larvae (Bayne, 1976; see Sastry, 1979); similar observations have been made concerning Mercenaria mercenaria (Loosanoff, 1954; Kennedy et al., 1974). In Mytilus galloprovincialis the embryos do not develop normally above 20"C, although this temperature represents the optimum for larval growth (Hrs-Brenko, 1977; His et al., 1989). In general, the rate of development increases with temperature as long as it remains within the range of tolerance (Sastry, 1979). 2.3.4.2. Salinity. As in the case of temperature, development of the eggs requires a narrower salinity range than survival and growth of the larvae (Bayne, 1976,1983). In the case of C. virginica the minimum salinity at which the eggs will develop is determined by the salinity at which the broodstock was kept prior to spawning (Davis, 1958; Davis and Calabrese, 1964). Salinity tolerance also depends on genetic factors (Newkirk et al., 1977; Newkirk, 1978; Widdows, 1991) and on interactions with temperature (Table 7). Studies on the combined effects of temperature and salinity show that, compared to nutrition and temperature, salinity has relatively little effect on larval growth and development. Significant interactions occur only at the extreme limits of the tolerance ranges of temperature and salinity. Within these limits, growth depends on temperature and food (Widdows, 1991). 2.3.4.3. Oxygen. The respiration of bivalve larvae has been studied by Gerdes (1983b), Tazawa et al. (1985) and Hoegh-Guldberg and Manahan (1995) in Crassostrea gigas, by MacInnes and Thurberg (1973), Widdows et al. (1989), and Baker and Mann (l992,1994a,b) in Crassostrea virginica, by Riisgard et al. (1980, 1981), and Wang and Widdows (1991) in Mytilus edulis, and by Morrison (1971) in Mercenaria mercenaria. According to Morrison (1971), hypoxia does not affect larval growth except at oxygen concentrations below 4 mg 1-I. Hyperoxia, on the other hand, negatively affects growth at 13.7 mg 1-I (180% saturation) in larvae
THE ASSESSMENT OF MARINE POLLUTION
37
of Mercenaria mercenaria (Huntington and Miller, 1989). Settlement was reduced significantly in hypoxic treatments, as compared to normoxic treatments, and no settlement took place in anoxic conditions in Crassostrea virginica larvae (Baker and Mann, 1992, 1994a,b). In mussels, both embryos and early prodissoconch larvae developed and grew normally at Po, values greater than or equal to 3.16 kPa (Wang and Widdows, 1991; see also Widdows, 1991, p. 158). Aeration in the presence of an antibiotic (1 mg 1-' erythromycin) increased larval survival in Crassostrea rhizophorae larvae at high temperature (30"C), but reduced individual growth and total biomass (Lemos et al., 1994). Hoegh-Guldberg and Manahan (1995) found that the conditions to which bivalve larvae are subjected in the small respirometers used in respiration studies negatively affect larval metabolism. From a practical point of view, artificial oxygenation of the small chambers used in these tests is not recommended, because resulting turbulence inhibits larval growth and survival (Helm and Spencer, 1972; see also Loosanoff and Davis, 1963, p. 38). 2.3.4.4. Turbidity. The effects of turbidity on bivalve larvae have been studied by Davis (1960), Robinson (1983), and Huntington and Miller (1989) in Mercenaria mercenaria; by Davis and Hidu (1969b) and Dekshenieks el al. (1993) in C. virginica; by Davis and Hidu (1969b) in Ostrea edulis; and by Seaman et al. (1991, and unpublished data) in Mytilus edulis, M. galloprovincialis, Crassostrea gigas and 0. edulis. Natural turbidity values, up to a few hundred mg 1-' tend to be beneficial to larval growth (excepting polluted sediments), because adsorption and desorption processes on the surface of suspended particles may buffer and stabilize the water (Koke, 1993). In bioassays, however, the water used is usually filtered (mesh of 1pm, or less), and turbidity is not a factor of influence in most experiments (excepting sediment bioassays). 2.3.4.5. p H . The data on pH requirements of larvae are scarce. The pH should not be less than 6.75 for C. virginica, and 7 for Mercenaria mercenaria, and both species are unable to reproduce when the pH remains above 9. M. lateralis requires a pH between 7.25 and 8.25 for reproduction (Calabrese and Davis, 1966,1970; Calabrese, 1970b). Krassoi et al. (1996) maintained the pH between 7.8 and 8.4 in tests on Chlamys asperrima.
2.3.5. Nutritional requirements of bivalve larvae Nutrition plays a decisive role in the development of bivalve larvae, even though they are capable of surviving a few days without food (Loosanoff, 1954; Bayne, 1965; Millar and Scott, 1967; Ukeles, 1975; His and Seaman, 1992). With regard to the combined effects of salinity, temperature and
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N. L.
SEAMAN
nutrition, the last has the greatest effect, explaining 85 to 88% of the variance in growth in Ostrea edulis larvae (Robert et al., 1989), 64 to 75% in Mytilus galloprovincialis, and 54 to 70% in Crassostrea gigas (His et al., 1989). A great number of studies have been devoted to the nutrition of bivalve larvae (reviews by Ukeles, 1975; Bayne 1976, 1983; Sastry, 1979; Widdows, 1991; Boidron-Metarion, 1995). As far as bivalve larval bioassays are concerned, nutrition is a topic of paramount importance, because although it is not difficult to maintain adequate temperature and salinity ranges for an experiment, providing adequate food may be more difficult to put into practice. Imai et al. (1950) and Imai and Sakai (1961), for instance, succeeded in rearing larvae of C. gigas to metamorphosis in tanks by feeding them Monas sp., a colourless flagellate, at Milford. However, Loosanoff (1969) was unable to obtain any growth in larvae fed with the strain he received from Imai, and this species is no longer used in larval culture. 2.3.5.1. Phytoplankton. Marine bivalve larvae feed mostly on nanoplankton and are thus termed planktotrophic (Thorson, 1950). Veligers pass through three different trophic stages: the endotrophic period, during which nutrition depends exclusively on vitelline reserves; the mixotrophic period, in which both vitelline reserves and exogenous resources are used; and the exotrophic period with exclusive use of external food sources (Lucas et al., 1986; Boidron-Metairon, 1995). Despite studies with other feeds, it has become clear since the pioneering work by Loosanoff and Davis (1963) and Walne (1963) that phytoplankton is the principal food source for bivalve larvae; no food other than unicellular algae has been found to be entirely satisfactory for bivalve cultures (Webb and Chu, 1981). Food algae for culture of veligers must meet three criteria: adequate size (limited by the diameter of the mouth and oesophagus of the larvae), good nutritional quality and ease of cultivation. Some 50 species of algae have been tested for the purpose but only a dozen are generally used in bivalve larval culture (ChrCtiennot-Dinet et al., 1986). Not only does the nutritional quality vary among different algal species, but food quality can also vary during cultivation, and some may produce substances which are toxic to the larvae (Davis and Chanley, 1955b). For instance, Nannochloris cells may excrete a growth-inhibiting substance in great concentrations during the stationary phase of algal growth (Bayne, 1965). Methods to feed bivalve larvae were developed at Milford by Davis (1953) and Davis and Guillard (1958), and reviewed by Loosanoff and Davis (1963). The food consisted of a mixture of live algal flagellates (Calabrese and Davis, 1966) given at a rate of 0.01 ml packed cell volume per litre culture per day (Hidu, 1965). According to Calabrese
THE ASSESSMENT OF MARINE POLLUTION
39
(1970a) the algae used were Zsochrysis galbana, Monochrysis Zutheri and ChZoreZZa sp. Veligers are capable of feeding selectively (Davis, 1953). The work of Davis and Guillard (1958) and Walne (1963) has shown that bivalve larvae grow best when they are fed a mixed diet of two or more algal species (see also Bayne, 1983). Their quantitative and qualitative dietary requirements may change in the course of development (Davis and Guillard, 1958 Loosanoff and Davis, 1963). Veligers of Crassostrea virginica, for instance, are incapable of using Chlorella sp. during the first days of life (Babinchak and Ukeles, 1979), although they do utilize them at the age of about 1 week, at a size of 110pm (Loosanoff and Davis, 1963). Oyster larvae of the genus Crassostrea have particularly narrow requirements, and the number of algal species suitable as food for them is limited, Mytilus and Mercenaria are rather tolerant and Ostrea is intermediate. The methods of rearing larvae of Crassostrea gigas most frequently used nowadays follow Walne and Helm (1974), Helm and Millican (1977), and Utting and Spencer (1991). These methods may be used for rearing most species of bivalve larvae used in ecotoxicological bioassays. The best results were obtained by rearing D-larvae without aeration at densities of about 10 per ml in 1 litre of filtered and ultraviolet-sterilized seawater, and feeding them with an algal mixture of Zsochrysis galbana Parke and Chaetoceros calcitrans (Paulsen) Takano (50 cells of each per pl of larval incubation volume). The water is changed every 2 days and larvae up to a size of 120 pm are retained on a 45 pm nylon mesh-based PVC sieve, rinsed with freshly filtered seawater and put back into incubation. After the umbo has developed (usually at the age of 6 to 8 days) the cultivation densities may be reduced to 5000 or 6000 per litre and the feeding regime is changed to 3.3 cells per pl of culture of Tetraselmis suecica (Kylin) Butch plus 33 cells per p1 each of 1. galbana and C. calcitrans (representing approximately equal volumes of all three species of algae). At Arcachon we follow the same methods, but we have found that ultraviolet sterilization of the water is not indispensable, and we use stainless steel sieves which are sterilized at 180°C. 2.3.5.2. Bacteria and microencapsulated diets. Davis (1953), Hidu and Tubiash (1963), Millar and Scott (1967), Mengus (1978) and Prieur (1981), among others, have studied the importance of bacteria for larval nutrition. Widdows (1991, p. 151) concluded “there is little evidence that they play a significant role in meeting the nutritional requirements during larval growth”. Douillet (1993) found that oyster larvae fed on a bacterial strain received only 41% of their carbon requirements. There have also been experiments with inert foods (see Robert and Trintignac, 1997). The first investigations by Ukeles (1975) were not very encouraging. Chu er al. (1982, 1987) have been able to rear larvae of Crassostrea
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E. HIS, R. BEIRAS AND M. N. L. SEAMAN
virginica to metamorphosis, but in their experiments only 2 to 25% of the larvae attained the eyed stage, and 2 to 20% achieved metamorphosis. The development of a reliable artificial diet would represent an enormous progress in bivalve larval culture, because it would vastly reduce the time and effort required (Ukeles, 1975), while standardizing feeding of the larvae between laboratories. 2.3.5.3. Dissolved organic matter. Finally, bivalve embryos and larvae are also able to take up dissolved organic substances (review by Widdows, 1991). This may well be beneficial, but Widdows (1991) concludes that there is no evidence that larvae are able to grow and develop solely on dissolved organics. The issue of the uptake of dissolved organic matter is important with regard to bioassays because it may also be a factor in the action of various pollutants.
3. BIOASSAY METHODOLOGY As Calabrese (1984) has pointed out, the various techniques and methods of ecotoxicological investigation with embryos, larvae and adult marine bivalves have resulted from advances in aquaculture. There are various manuals and reviews describing in detail the technical precautions necessary for conducting ecotoxicological tests with seawater (types of material, cleaning and sterilization, preparatory steps, etc.). Recommended literature on bioassays with bivalve larvae includes the work of Woelke (1972), the standards of ASTM (1980, 1989), Calabrese (1984), and more recently Widdows (1993) and Krassoi et al. (1996). The general principle is to eliminate the influence of toxic materials and any type of hazardous conditions, which could invalidate the bioassay, other than the condition under study. A broad theoretical and practical knowledge of larval rearing and larval biology is an important prerequisite for performing embryo and larval bioassays. Valid ecotoxicological studies and proper interpretation of their results presuppose an understanding of the normal development of the test species and a mastery of the technical procedures for optimal laboratory rearing. 3.1. General methods
3.1.1. Seawater quality 3.1.1.1. Natural seawater. One common element for all types of toxicity tests is the need for seawater of excellent quality. The first studies with marine bivalve embryos and larvae were carried out at biological
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41
laboratories specialized in shellfish aquaculture and located on bays and estuaries where running seawater was obtained from an area in which oyster populations reproduced naturally. This was notably the case at Milford (Connecticut), where bioassays were pioneered by Davis (1961), Loosanoff and Davis (1963), and Hidu (1965). As pointed out by Loosanoff and Davis (1963) and Woelke (1972), the source of seawater must be located at a site which is not contaminated in any way, and where the salinity is stable year-round (or can be kept stable by pumping water at appropriate periods of the tidal cycle). In addition, materials that might release toxic substances must be scrupulously avoided. Most coastal areas are nowadays increasingly subject to anthropogenic impacts resulting in a deterioration of the “biological quality” of the seawater. One of the first examples was noted by Millar and Scott (1968) at the laboratory of Millport (U.K.), where an unknown toxic substance slowed the growth of Ostrea edulis larvae. Shortly thereafter, Helm (1971, p. 8) remarked that “at Conway, sea water varies considerably in quality both in the short term and seasonally”, and the biological quality of the seawater was improved by adding 1mg 1-l of EDTA (Utting and Helm, 1985). Brereton et al. (1973) and Boyden et al. (1975) showed that the poor growth of Crassostrea gigas larvae and the irregularities in oyster recruitment resulted from the presence of zinc in the water, the source of which was mines in the catchment area that drain into the bay at which the biological station was located. This corroborated the observations of Wilson (1951) and Wilson and Armstong (1961) concerning the variability of the “biological quality” of coastal waters (cf. Section 1.3). The volume of the vessels employed in bioassays has been greatly reduced and can now be as small as 3ml (see 3.2.3), so that large quantities of seawater are no longer required and bioassays are not restricted to laboratories located on the seashore. In addition, seawater may be obtained offshore beyond the region of anthropogenic impact in order to obtain it “from an area known to support a healthy naturally reproducing population of bivalves” (ASTM, 1989, p. 339). Seawater filtration. Following the work of Loosanoff and Davis (1963) and Walne (1966, 1974b), most authors agree that bioassays should be conducted only with filtered seawater sterilized with ultraviolet light (Beaumont and Budd, 1982; Beaumont and Terpes, 1984; ASTM, 1989; Hoare et al., 1995a,b; Krassoi et al., 1996). The porosity of the filter is usually equal to, or less than, 1 pm. The ASTM proposes 0.45 pm, but many authors prefer 0.2 pm (Utting and Spencer, 1991; Ringwood, 1991; Hoare et al., 1995a,b; studies by His and coworkers at Arcachon). Filters of 0.1 pm have been used as well (Lemos et al., 1994). Simple filtration systems may prove equally useful. For example, Armstrong and
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M. N. L. SEAMAN
Millemann (1974) were able to cultivate various larval stages of Mytilus edulis in seawater which had merely been passed through a sand filter. In contrast to almost all other workers, Garland et al. (1986) found that 0.2 pm filtered seawater led to poor growth in bivalve larvae. lbenty years of experience with bivalve embryos and larvae of Crassostrea gigas, Ostrea edulis and Mytilus galloprovincialis at Arcachon have shown clearly, however, that 0.2 pm Htration of seawater is perfectly adequate for toxicity bioassays of 24 or 48 hours duration, as well as for tests of larval growth over periods of several weeks, and for studies of metamorphosis, as long as the seawater is obtained from adequate sites. Filtration of the water should be carried out immediately prior to the incubation, no more than 1 or 2 hours before the test is initiated. In the case of monitoring of in situ water quality, on the other hand, “ideally, filtration is not recommended, because it may remove some of the dissolved contaminants” (Thain, 1991, p. 4), and because it will definitely remove fine suspended particles to which pollutants are likely to adsorb. In such cases good results can be obtained by simply sieving the water through a 32 pm stainless steel sieve in order to eliminate the largest suspended particles and possible predators of the larvae (His and Beiras, 1995). Bacteria and antibiotics. Because the comparatively small volume of the incubation vessels (usually 1 litre at most) favours bacterial growth, and the bacterial populations eliminated by filtration are rapidly reconstituted (Walne, 1958; Lemos et al., 1994), many authors have added antibiotics to the seawater even after filtration and ultraviolet sterilization (cf. Davis and Chanley, 1955b; Walne, 1958). Hidu and Tubiash (1963) recommended the systematic addition of antibiotics (dihydrostreptomycin-streptomycin sulphate = Combistrep at 2 ppm) in larval cultures of Crassostrea virginica and Merceaaria mercenaria. They thought (p. 25, op. cit.), “an antibioticinduced bacterial flora. . . may be utilized by larvae as a food source” (see also Section 2.3.5.2). Similarly, Calabrese and Davis (1967, p. 12) used sulfamethazine soluble powder at 0.33 ppm, Brereton et al. (1973) added 0.3g1-1 of penicillin to the seawater, and Lemos et al. (1994) added 1mg 1-’ of erythromycin. The latter, however, found that although growth is thus improved by antibiotics in Crassostrea rhizophorae cultured at high temperatures and low salinity (30°C and 12 to 20 psu), this is later followed by increases in mortality. Chloramphenicol (5 mg 1-1 of 1pm filtered seawater) was used in larval cultures of Spisula solidissima by Thurberg et al. (1975). Le Pennec and coworkers (Le Pennec and Prieur, 1972; Le Pennec et al., 1973) have shown that antibiotics (aureomycin, erythromycin, chloramphenicol and sulfamethazine) may influence the metabolism and growth of bivalve larvae. This may affect the accuracy of
THE ASSESSMENT OF MARINE POLLUTION
43
bioassays designed to test a pollutant’s possible reduction of larval growth. Although Woelke (1966) stated that the seawater should be used within 3 hours of sampling, we have obtained excellent results at Arcachon with seawater sampled offshore and stocked for 24 hours at ambient temperature (18°C to 19OC), but filtered immediately before use (see above). On the other hand, Thain (1991) apparently did not obtain good results with seawater sampled offshore during winter and stocked at -18°C in acid-washed bottles, as he states that “even 60 percent abnormalities - in controls - is acceptable” (Thain, 1991, p. 7). We shall see below that most workers do not accept such high levels of abnormalities in the controls. Klockner et al. (1985) also encountered high levels of abnormality in the controls when they reared larvae in “aged (4years, filtered - Seitz K 15) and pasteurized (8OOC for 1 h) sea water” (p. 2). 3.1.1.2. Artificial seawater. Seawater quality problems can be overcome with artificial seawater made by dissolving the mineral constituents of oceanic water in distilled water. This also prevents the complex interactions between pollutants and organic matter that may affect toxicity. Contrary to the opinion of Thain (1991), artscial seawater is perfectly suited for bivalve embryo bioassays (e.g. Krassoi, 1995; His et al., 1997a). The seawater formula of Zaroogian et al. (1979), used successfully by Calabrese et al. (1973), Calabrese and Nelson (1974), Utting and Helm (1985) and recommended by ASTM (1989), gives excellent results in embryo-larva bioassays of 24 hours duration in Crassostrea gigas, or 48 hours in Mytilus galloprovincialis. It may be prepared using reagent-grade chemicals and deionized water aerated before conducting the test. EDTA, which chelates metals and some pollutants, may be omitted (His et al., 1997b, p. 352). Besides Zaroogian’s formulation, various other seawater media have been used to rear bivalve larvae (Table 8), and Krassoi (1995) has recently used several different formulations to grow larvae of Chlamys asperrima and Saccostrea commercialis. Kester et al. (1967), who proposed a modification of the formula by Lyman and Fleming, stressed that artificial seawater should be made up with reagents of known composition; this is not always the case, as the composition of the salt used by Courtright ef al. (1971) is unknown. The formula used by Chang et al. (1996) is a simplification of the medium of Zaroogian et al. (1979), containing only six mineral salts (NaC1, KC1, CaCl,, MgCl,, MgS04 and NaHCO,), instead of eleven. 3.1.2. Broodstock Spawners in good physiological condition are indispensable; they must be able to mature in an unpolluted environment and in excellent nutritional
Table 8 Artificial seawater (ASW) formulations used for the rearing of bivalve larvae.
Artificial seawater Lyman and Fleming, in Sverdrup et al. (1949) Wood (1961) Zaroogian et al. (1969)
Leslie coarse hide salt Instant Ocean sea salt mixture Instant Ocean Aquarium Systems, Mentor OH Different ASW formulations
Bivalve species Ostrea edulis 0. edulis Crassostrea virginica Mercenaria mercenaria Crassostrea gigas C. gigas Mytilus galloprovincialis C. virginica Mytilus edulis C. virginica C. gigas C. virginica Chlamys asperrima Saccostrea commercialis
Reference Helm (1971) Millar and Scott (1968) Calabrese et al. (1973) Calabrese and Nelson (1974) Utting and Helm (1985) His et al. (1997a) His et al. (1997a) La Roche et al. (1970) Courtright et al. (1971) Sigler and Leibovitz (1982) Chang et al. (1996) Chang et al. (1996) Krassoi (1995) Krassoi et al. (1996)
Remarks Rearing Rearing Bioassay Bioassay Rearing Bioassay Bioassay Bioassay Bioassay Bioassay Bioassay Bioassay Bioassay Bioassay
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conditions, whether in the field or in the laboratory (ASTM, 1980, 1989; Krassoi et al., 1996). The best results are usually obtained during the regular spawning season by using adults from areas where bivalves are cultivated (see Section 2.2). On the other hand, when bioassays need to be conducted out of season, ripe adults may be obtained by conditioning techniques (see Section 2.3.1). Most commercial hatcheries are capable of supplying mature oysters, Crassostrea spp., which are the ones used most extensively in bioassays (see Table l ) , to laboratories lacking the facilities for conditioning bivalves. A word of caution is in order, however, because the adults from commercial hatcheries are sometimes overripe, i.e. they have been conditioned for too long without spawning. “. . . Oysters resorbing their unspawned products . . . result in poor bioassay material” (Woelke, 1972, p. 31), with abnormality levels near, or above 20% in the controls, thus generating results of questionable validity (see Section 3.1.3.2). As noted by Eertman et al. (1993, p. 38) in the case of C. gigas embryos and larvae, “despite the oysters being conditioned for spawning, the autumn and winter periods seem less suitable for performing toxicity tests” as larval abnormalities reached 40% in October and March. Moreover, many hatcheries clean out their facilities in autumn and winter and are therefore unable to provide spawners during this time. If the spawners are subjected to unfavourable environmental conditions, this will affect the quality of the gametes and influence the results of the test (Widdows, 1993; see Section 2.2.2). In Mytilus rrossulus and Mizuhopecten yessoensis, exposure of the adults to sublethal doses of heavy metals (Cu and Zn) prevents the formation of gametes capable of normal embryonic development (Karaseva and Medvedeva, 1993). On the other hand, adaptive mechanisms such as the formation of metalloproteins have also been found, particularly in the case of cadmium (see Webb, 1979), and these metalloproteins are later found in the larvae upon their exposure to the contaminant (e.g. Pavicic et at., 1984, 1994a,b; Roesijadi et al., 1996, 1997; Ringwood and Brouwer, 1995). Similarly, Hoare et al. (1995a,b) found that Mytilus edulis embryos obtained from spawners originating from a site polluted by heavy metals had higher tolerance to copper than those spawned by adults from a non-polluted area; tolerance appeared to be “maternally determined”. There have been attempts to cryopreserve bivalve larvae in order to dispense with the need to obtain mature adults, but little progress has been made since the first experiments by Lannan (1971; see also Renard, 1991). It has usually not been possible to obtain adequate percentages of normal D-larvae (80% or higher) from cryopreserved gametes. Nevertheless, McFadzen (1992), Cleary et al. (1993) and McFadzen and Cleary (1994) have performed bioassays with veligers of C. gigas cryopreserved 24 hours
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after fertilization, and of Tapes philippinarum 48 hours old. Contrary to the authors’ contention, it is not at all certain that 48-hour cryopreserved larvae are more sensitive as test organisms than the embryonic stages favoured by most other workers. More recently, Chao et al. (1997) have proposed the use of C. gigas and Meretrix lusoria cryopreserved late embryos and early larvae in aquaculture management programmes on the basis of survival rates from 62 to 84%. Future progress may yet facilitate the development of routine bioassays with cryopreserved material, making it possible, for example, to exchange genetically homogeneous material between laboratories. At present, however, cryopreservation techniques are not sufficientlyreliable and widespread for routine employment. We agree with Widdows (1993, p. 153), that the use of cryopreserved D-larvae “still remains to be fully evaluated”. 3.1.3. Spawning and fertilization Spawners must be used within 24 hours after being obtained from the field or the hatchery; they need to be carefully cleaned of fouling organisms and scrubbed in seawater. Two-year-old animals are preferable in the case of oysters, in order to have a fair chance of obtaining both males and females (see Section 2.2.1). 3.1.3.1. Induction of spawning. Gametes may be obtained either by stimulating the animals to spawn, or by “stripping” the gonad. The former method, recommended by Woelke (1972), has been adopted by ASTM (1989) in Crassostrea gigas, C. virginica, Mytifus spp. and Mercenaria mercenaria. Mature adults are placed in 1 litre beakers with filtered seawater and stimulated to spawn by varying the temperature. According to ASTM (1989, Table 2, p. 341) the temperature may be raised by 5°C to 10°C above the temperature of conditioning, but it is not supposed to exceed 20°C for mussels. On the other hand, bivalves may be subjected in the field to temperatures which fluctuate from more than 30°C (at low tide and strong solar irradiation) to 20°C at high tide. His and Beiras (1995) and His et al. (1997a) have found that 30 minute periods at temperatures alternating between 18°C and 28°C serve very well to induce spawning in both C. gigas and Mytilus galloprovincialis. Dixon and Prosser (1987) induced M . edufis to spawn at 30”C, and C. Bittkau (personal communication) obtained healthy gametes of the same species from the Baltic in winter (ambient temperature 5°C) after exposing them to fluctuations between 18°C and 28°C immediately after field sampling. An additional spawning stimulus may be provided by adding a suspension of sperm in filtered seawater to the beakers containing the mature adults (obtained by stripping the gonads of a male). To avoid
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undesired fertilizations in such a case, it is also possible to use a suspension of previously frozen sperm which has been brought to ambient temperature. Ringwood (1992b) used heat-killed sperm to stimulate spawning in Zsognomon californicum. 3.1.3.2. Stripping the gonad. With regard to the production of gametes by stripping the gonad, Allen et al. (1988, p. 7) states, “during our research, we investigated the conventional wisdom that stripped eggs of Pacific oysters are inferior to naturally spawned ones. We found nothing to support that belief”. Alliegro and Wright (1983) stress that stripping avoids the often tedious induction of spawning by other methods, which may take hours. According to Thain (1991), there is no important difference between the use of larvae obtained by stripping or by natural spawning in the conduction of ecotoxicological bioassays. Most other authors, however, disagree (Woelke, 1961, 1966; Loosanoff and Davis, 1963; Loosanoff, 1965b, 1969; Wilson, 1981; ASTM, 1989; Widdows, 1993). Although stripping of mature females can provide some uninjured ripe eggs that can be fertilized and develop normally, among the eggs obtained by induced spawning there are always considerably fewer abnormal ones than among eggs obtained by stripping (Loosanoff and Davis, 1963). As early as 1961, Woelke stated that “this practice [stripping] was dropped” because of “the frequency with which unsatisfactory results were achieved, apparently due to immature eggs” (Woelke, 1961, p. 115). Moreover, in Mercenaria mercenaria it is not possible to fertilize stripped eggs until they have been treated by a weak ammonia solution “to cause the germinal vesicle to break and the eggs become physically prepared for fertilization” (Loosanoff, 1969, p. 21). In Mytilus edulis “attempts to obtain mature (fertilizable) eggs by removing the ovary of apparently ripe females have met with unsatisfactory results, because it is only during their stay in the ovary, and during and following the act of spawning, that the ova become physiologically mature” (Longo and Anderson, 1969a, p. 73). In C. gigas, spawned oocytes are considerably less susceptible to polyspermy (fertilization of one egg by more than one sperm) than oocytes mechanically removed from the ovary (Stephano and Gould, 1988; Konar and Stephenson, 1995). In short, we agree with ASTM, that “use of eggs stripped from female bivalves is not recommended because it often results in an excess of poorly developed and malformed embryos” (ASTM, 1989, p. 343). 3.1.3.3. Number of parents. Opinions differ with regard to the number of parents whose spawn should be used in bioassays. Thain (1991) states that, for Crassostrea gigas, after stripping the gonads, “three batches of eggs are pooled and two batches of sperm are also pooled”. Johnson (1988), in Mytilus edulis, and Krassoi et al. (1996) in Chlamys
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E. HIS, R. BEIRAS AND M. N. L. SEAMAN
asperrima mixed the gametes of three males and three females; Beaumont et al. (1987) also preferred to use gametes of several spawners, both in M. edulis and Pecten maximus, “in order to avoid working with the very restricted genetic variation inherent in full or half sibs” (p. 300), as did Spangenberg and Cherr (1996) in M. californianus and Mercenaria mercenaria, and Laughlin et al. (1988, 1989) in M. mercenaria. Cherr et al. (1990) conducted separate fertilizations of several females of Mytilus californianus with sperm of two or three males, and Roberts (1987), in Crassostrea virginica and Mercenaria mercenaria, fertilized the eggs of several females with the sperm of one or several males. Finally, Ringwood (1991, 1992a,b) mixed the gametes of several males and females in Zsognomon californicum. ASTM (1980) recommended using gametes from two individuals of each sex in order to obtain a genetic mix of embryos. In contrast, ASTM (1989) recommends in order to verify differences in sensitivity between parental couples “ideally, the test should be conducted by subjecting progeny from each of at least three individual male-female pairings to each of the one or more control treatments” (p. 242). Woelke (1972) and Chapman et al. (1992) in Crassostrea gigas, and Granmo et al. (1988, 1989) in Mytilus edulis, recommended the use of a single male and a single female in a given bioassay; the same approach was chosen by Butler et al. (1990) in M. edulis: “eggs and sperm from two individuals were selected to give a single pairing, to minimize genetic variability in the test embryos” (p. 214). 3.1.3.4. Elimination of impurities in the spawn. Loosanoff and Davis (1963) recommended sieving the gametes to eliminate tissue fragments emitted by the bivalves along with the gametes. Butler et al. (1990) passed the gametes through a 1OOpm mesh, and Thain (1991) through 90pm. ASTM (1989) recommends a mesh of 75pm for the eggs and 37 pm for the sperm, the fertilized eggs being retained on a sieve of 37 pm for rinsing and counting. Krassoi et al. (1996) retained the eggs on a sieve of 45 pm, after passing them through a mesh of 100 pm. Granmo (1972), on the other hand, rinsed the eggs three times, allowing them to decant between rinsings, prior to fertilization. Spangenberg and Cherr (1996) simply rinsed the eggs in 0.45 pm filtered seawater after fertilization, and Ringwood (1992a,b) eliminated tissue debris by rinsing and gentle centrifugation. His and Beiras (1995) and His et al. (1997a) obtained excellent results by passing the eggs and sperm through sieves of 100 p m and 32 pm respectively, before proceeding to fertilize, without further rinsing or sieving after fertilization. 3.1.3.5. Counting the eggs. After spawning, and sieving or rinsing, the first step in a bioassay is to count the eggs. The method proposed for
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this purpose by Loosanoff and Davis (1963) has been adopted by most of the workers in the field (e.g. Woelke, 1972). It consists in placing the fertilized eggs in a graduated cylinder of 1 litre volume filled with filtered seawater and gently mixing the water column with a “plunger constructed by drilling holes in a disc of acrylic plastic or fibreglass of suitable diameter and attaching it to a PVC or acrylic rod of suitable length” (ASTM, 1989, p. 343; see Figure 7 in Woelke, 1972). Several samples are taken by pipette and counted under the microscope in a haematocytometer or well slide. A sample (usually 100 p1, or less) should consist of about 50 eggs, and, as the number of eggs in the cylinder is initially unknown, it may be necessary to adjust the volume of the samples accordingly, or to take a large sample from the graduated cylinder and dilute it according to need. The total number of eggs is computed by extrapolating the average number of eggs in the samples to the volume of the cylinder. 3.1.3.6. Fertilization. The sperm suspension should be obtained at approximately the same time as the eggs are collected: “Fresh actively-moving sperm are used to assure a normal fertilization of eggs” (Loosanoff and Davis, 1963, p. 35). To obtain a concentrated suspension, males should be made to spawn in the smallest possible volume of filtered seawater; some authors propose to count the sperm with a haematocytometer (Cherr et al., 1990; Ringwood, 1992b; Konar and Stephenson, 1995; Krassoi et al., 1996), particle counter (Thain, 1991), or by “turbidimeter readings previously calibrated with a hematocytometer” (Eyster and Morse, 1984, p. 642). ASTM (1989) admits that “precise sperm counts are unnecessary after one gains a little experience” (p. 343), even though optimum fertilization depends on the ratio of eggs to sperm (Section 2.3.3.1). This differs between species (and between authors), and it is also strongly influenced by gamete quality (potency of the sperm and receptiveness of the eggs, which are difficult to appraise). Most authors add a few millilitres of dense, freshly spawned sperm suspension and verify fertilization success under the microscope; 5 to 10 sperm (24 at most) should be visibly attached to each egg membrane (Konar and Stephenson, 1995; His and Beiras, 1995; His et al., 1997a). Because the biological quality of the gametes deteriorates rapidly after spawning (Section 2.3.3.2), fertilizations should preferably be undertaken within 30 minutes. After eggs and sperm have been united in one container, the actual fertilization takes place within minutes and the polar bodies are expelled within half an hour (Allen et al., 1988; see also Section 2.2.3). As a rule, fertilization of the gametes for bioassays should be undertaken as soon as the first females in the batch have spawned.
E. HIS, R. BEIRAS AND M. N.
L. SEAMAN
3.2. Bioassay procedures In assays with bivalve embryos and larvae, we may distinguish between acute toxicity tests, when response is recorded after short-term exposure (a few days or hours), and chronic toxicity tests, when the exposure period covers the majority of the larval life span (several weeks). The former tests frequently record embryogenesis success or larval mortality as the response, whereas the latter record sub-lethal responses, usually larval growth. Both kinds of tests are complementary. Acute toxicity tests are easier to standardize and provide a rapid assessment, whereas chronic toxicity tests are more sensitive and better approximate the environmental conditions. The execution of longer-term sub-lethal tests requires perfect capabilities in larval rearing - particularly with regard to the culture of food algae - as well as the availability of relatively sophisticated equipment, including a unit for the cultivation of algae. Sub-lethal tests cannot, therefore, be performed as a routine method in most marine biological laboratories, yet they remain an integral part of more detailed studies on particular chemicals or environmental hot spots. The first step in all bioassays - whether they are acute tests with embryos, chronic exposure studies lasting several days with larvae, or investigations on pediveligers and toxic effects on metamorphosis - is the production of gametes, embryos and D-larvae of excellent quality and the elimination of all artefacts resulting from any type of deficiency in the biological material. We have already reviewed the methods to obtain mature adults, induce spawning, and perform in vitro fertilization in Section 2.3. The principal causes of failure in bioassays with Chlamys asperrima have been summarized by Krassoi et al. (1996, Table 7). These include loss of broodstock, spawning of broodstock in holding tanks, poor spawning, poor fertilization success, low percentage of normal larval development (<70%), termination of development at the trochophore stage, and finally poor preservation of larvae preventing proper assessment of the biological response. These problems are common to all bivalve species used in marine ecotoxicological bioassays. 3.2.1. Spermiotoxicity test The use of sperm, previously exposed to a contaminant, to fertilize eggs in the absence of that contaminant has mostly been tested using sea urchins (e.g. Dinnel et al., 1982, 1987; Dinnel, 1995). Spermiotoxicity tests have rarely been used with bivalves. Renzoni (1973a, p. 11, experiment 4) “inseminated Crassostrea gigas and C. angulata eggs in clean filtered sea water with sperm previously maintained (3 h) in sea water with various concentrations of oil/emulsilier mixtures”. This author also studied the
THE ASSESSMENT OF MARINE POLLUTION
51
effects of water soluble extracts of three crude oils on sperm of Mulinia lateralis by exposing them for 3 hours and then using them to fertilize untreated eggs (Renzoni, 1975). His and Robert (1980) subjected sperm of C. gigas for 30 minutes to various concentrations of tributyl-tin (TBT), and subsequently fertilized eggs with that sperm in a toxicant-free medium. Eyster and Morse (1984) separately exposed sperm and eggs of the surf clam, Spisula solidissima, to silver salts, and then conducted fertilization experiments with untreated eggs and treated sperm, and with untreated sperm and treated eggs. Finally, Ringwood (1992b) studied the effects of 1 hour exposure to TBT, and to cadmium and copper salts on sperm of Isognomon californianum, using the method of Dinnel et al. (1987). Unlike sea urchin tests, spermiotoxicity tests with bivalves are not more sensitive than embryo-larval tests (see also Section 4.2.2).
3.2.2. Embryo-larval bioassay The progress of embryogenesis from fertilized egg to straight-hinge D-larva is the biological response most frequently used in bivalve bioassays. Therefore, the methodological details of the embryo development bioassays will be discussed in depth. 3.2.2.1. Age of the fertilized eggs at the beginning of the test. There is some disagreement as to the time that should pass between fertilization and the beginning of the bioassay. The ASTM standard states, “embryos should be tested within an hour of spawning and kept suspended during the period by frequent agitation with a perforated plunger” (ASTM, 1989, p. 252). Woelke (1972), Cardwell et al. (1977b), Morgan et al. (1986), Johnson (1988) and Thain (1991) used an interval of between 1 and 2 hours, and Laughlin et al. (1988) used an interval of 4 hours between fertilization and exposure to the toxicant. Widdows (1993) also proposes waiting 4 hours. Some authors wait until the first polar body is expelled (Chien and Chou, 1989; Spangenberg and Cherr, 1996). Ringwood (1992a,b) initiates the test within 30 minutes after fertilization, as is the case at the Arcachon laboratory (e.g. His and Beiras, 1995; His et al., 1997a). In our view, two ideas should be highlighted. First, the rapidity of embryonic development to the D-larva stage allows restriction of the duration of the test to less than 24 hours in Crassostrea spp., and less than 48 hours in Mytilus spp. (see next Section). Second, the ideal test (that closest to natural environmental conditions) would perform fertilization already in the presence of the toxicant. As explained below (see Section 3.2.3), in practice it is difficult to perform simultaneous fertilizations in separate treatments, and this difficulty may be overcome by beginning incubation as soon as possible after the sperm suspension
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E. HIS, R. BEIRAS A N D M. N. L. SEAMAN
has been added to the eggs, i.e. after about 15 minutes, in order to maximize the exposure period. 3.2.2.2. Incubation conditions: duration, temperature and egg density. The duration of bioassays also varies between authors, and it cannot be discussed independently of the temperature, which determines the rapidity of the development. Duration depends on the species used as well. We shall also review here another basic incubation parameter: egg density. In the case of Crassosfrea gigas, ASTM (1989) recommends incubation at 20°C for 48 hours. The same values were suggested by Woelke (1967, 1972), Cardwell et al. (1977a, 1979a) and Konar and Stephenson (1995), whereas Thain (1991) and His et al. (1997a) propose incubating the eggs at 24°C for 24 hours. Van den Hurk (1994) followed the incubation temperature and period recommended by ASTM, but subjected the embryos to a light-dark cycle of 16 hours of light to 8 hours of dark. For Crassostrea virginica, ASTM (1989) also recommends that embryos be incubated at 20°C to 25°C for 48 hours. Calabrese and Davis (1967), Calabrese et al. (1973) and Calabrese and Nelson (1974) used temperatures of 20°C to 26"C, salinity of 25 psu and incubation periods of 48 hours in C. virginica and Mercenaria mercenaria. Roberts (1987) conducted the tests at 28"C, and obtained D-larvae in 48 hours. For Mytilus edulis, ASTM (1989) recommended 16"C, Granmo (1972) and Granmo et al. (1988) used 20"C, while Johnson (1988) incubated at 15°C for 72 hours. Butler et al. (1990) incubated for 48 hours in the dark at 20°C. For M. galloprovincialis, His and Beiras (1995) incubated embryos at 25 psu salinity and 18°C 2 1°C for 48 hours, while Brunetti et al. (1989), used 20°C for 40 hours. Cherr et al. (1990) incubated M. californianus in the dark for 96 hours at 12°C and Spangenberg and Cherr (1996) for 48 hours to 64 hours at 15°C. With Mulinia lateralis, Calabrese (1970a,b), Calabrese and Rhodes (1974) and Morrison and Petrocelli (1990) performed their experiments at 21°C for 48 hours, while Burgess and Morrison (1994) used 25°C to 26°C for 48 hours. Finally, in Zsognomon californicurn, Ringwood (1992b) conducted 48 hour incubations, and Krassoi et at. (1996) recommended incubation of Chlamys asperrima at 18°C 5 05°C for 48 hours. Regarding egg density, Calabrese and coworkers (Calabrese and Davis, 1967; Calabrese, 1970b; Calabrese et al., 1973; Calabrese and Nelson, 1974; Calabrese and Rhodes, 1974) used 10 to 20 eggs per ml, while Woelke (1967, 1972), Cardwell et al. (1977a, 1979a) and ASTM (1989) recommended a maximum density of 30 embryos per ml, even though normal development is possible at densities of 100 embryos per ml. Granmo (1972), Granmo et al. (1988), Johnson (1988), Burgess and Morrison (1994), Butler et al. (1990), Cherr et al. (1990), Ringwood (1992b), His and coworkers (His and Beiras, 1995; His et al., 1997a), Konar and Stephenson (1995) and Krassoi et al. (1996) followed the ASTM recommendation,
THE ASSESSMENT OF MARINE POLLUTION
53
whereas Roberts (1987) and Brunetti et al. (1989) conducted the tests with 40 embryos per ml. Thain (1991) used 50 embryos per ml, and Morrison and Petrocelli (1990) used up to 75 embryos per ml. As a rule, we think embryo density should not exceed 30 embryos per ml. The incubation period depends on the species; bioassays should not last longer than the shortest possible period required to complete embryogenesis. At optimal temperature, an exposure period of 24 hours is sufficient for most species, with some exceptions, such as Mytilus species, where 48 hours are usually necessary. 3.2.2.3. Volume of the test vessels. Acute toxicity tests are frequently used in biological monitoring of waters and sediments, often requiring numerous vials for numerous replicates to permit adequate statistical evaluation. There has been a continuing trend towards simplification of the bioassay procedures, with a concomitant reduction of incubation volumes. The first toxicity bioassays by Davis (1961), Hidu (1965), Calabrese and Davis (1966) and Woelke (1960 to 1972) were conducted in glass and polypropylene beakers of 1 to 2 litre volumes. Since 1980 ASTM has found that “very small vessels, containing as little as 10 to 30 ml may be used if data are provided demonstrating these systems have negligible effects on water quality” (ASTM, 1980, p. 249). Similarly, Byrne and Calder (1977) studied the toxicity of the water-soluble fraction of various petroleum products on larvae of Mercenaria rnercenaria in 10ml tubes, Morrison and Petrocelli (1990) used scintillation vials of 10 ml, and Ringwood (1992b) incubated lsognomon californicum embryos in tubes of 12 ml. Volumes of 100 ml have been used by Watling (1981) and Johnson (1988); Laughlin et al. (1988) used 50ml glass finger bowls; Butler et al. (1990) and Spangenberg and Cherr (1996) conducted their studies with glass jars of 30ml (cf. Widdows, 1993), whereas His and Beiras (1995), Beiras and His (1995a,b) and His et al. (1997a) carried out their incubations in transparent polypropylene Coulter Counter accuvettes of 25ml permitting the assessment of the larvae under an inverted microscope without further manipulation of the test organisms. Finally, Cherr et al. (1990), Chang et al. (1996) and Gormly et al. (1996) conducted tests in 3 ml wells commonly used for tissue cultures, which also permit analysis under an inverted microscope. Transparent flat-bottomed multi-well tissue culture plates of 3ml volume make it possible to study simultaneously four samples, with five to six replicates each, under an inverted microscope; the results are absolutely comparable to those obtained with 30 ml expendable vials or with beakers of 1 litre (His and Beiras, unpublished data). It is important to carefully define the criteria and end points upon which an assessment of toxicity is based, regardless of the physiological activity being studied (feeding activity, respiration, swimming behaviour,
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L. SEAMAN
(4 Figure 6 Effects of a contaminant (CuS04)on Crassostreagigas larval growth. (a) 1%-day-oldlarvae in controls. (b) 12-day-old larvae at 50 pg Cu 1-I; only a few larvae grew normally. Growth inhibition increased with increasing copper sulphate concentrations;some larvae which had not reached the umboned stage were still alive on day 12. (From His and Robert, 1982.)
calcium uptake, metamorphosis). The criteria used most frequently in embryo bioassays are mortality and larval abnormalities. 3.2.2.4. Larval mortality as the end point. It is not always easy for an inexperienced biologist to recognize moribund and recently dead larvae, particularly when the samples have been treated with formalin for later evaluation. On the other hand, it is very easy to obtain mortality in the larval and algal cultures unless various precautions are taken. Mass mortality is usually caused by bacteria, which can affect several incubations of an experimental series without relation to pollutant concentration. The addition of antibiotics is ineffective once mass mortality has begun. In contrast, mortality as a result of a contaminant appears gradually over several days and increases progressively with pollutant concentration; there are always some larvae which remain apparently unaffected. At certain concentrations growth may be inhibited and D-larvae, for instance, will not attain the umboned stage except for some individuals whose number decreases with increasing concentration. The small larvae are continuously decimated in the course of the experiment (e.g. His and Robert, 1982; see Figure 6). Loosanoff and Davis (1963, pp. 39-40, Figure 18), noted that in C. virginica and M. mercenaria, “it has
THE ASSESSMENT
OF MARINE POLLUTION
55
been clearly demonstrated that even though larvae originate from the same spawning and sometimes from the same parents and are kept in the same vessel under identical conditions, individuals grow at widely different rates . . .”. This is particularly true when larvae are exposed to contaminants, and in experiments with algae which do not satisfy the nutritional needs of the larvae very well (e.g. His et al., 1985). In both cases the size distributions of the larvae are heterogeneous, even though the mortality after 10 or 12 days is rarely more than 10% higher than in the controls. Various authors have described dead larvae. In fresh material, “because decomposition of larval tissue is slow in clean culture conditions, it is not easy to distinguish between moribund and recently dead bivalve larvae. In general, in larvae approaching death, velum cilia cease beating first and oesophagae cilia last” (Beaumont and Budd, 1984, p. 403). Similarly, “criteria for death in setting M y a arenaria larvae (and in straight-hinge larvae) included lack of internal movement and disrupted internal organization” (Roosenburg et al., 1980a, p. 107, and 1980b, p. 94). Larvae which have been dead for more than 24 hours, however, are easily distinguishable, because bacterial degradation and feeding of protozoa on the dead tissue leave nothing but a transparent empty shell. Various authors have devised methods to identify dead individuals in larval cultures. Byrne and Calder (1977) stained larvae of Mercenaria rnercenaria with neutral red, following the work of Loosanoff and Davis (1947; see also ASTM, 1980, p. 254). Test tubes of 10ml containing 300 larvae were then refrigerated at 4°C (or even frozen) and conserved for several days prior to evaluation. Richardson et al. (1982) noted in C. virginica that “the neutral red dye . . - was picked up by the tissue of living larvae, causing them to appear pink. This aided determination of live and dead larvae” (p. 102). Phelps and Warner (1990) distinguished between live and dead pediveligers “by cilia movement, clarity of internal organs and transparency. Dead larvae are often gray and opaque with no evidence of internal organization or movement.” They also facilitated identification by staining with neutral red at a final concentration of 1/10000 for 3 hours. In “lethal” embryotoxicity bioassays lasting 24 to 48 hours it is unnecessary to take mortality into account, because the contaminants affect embryogenesis and either prevent its completion or induce abnormalities. The appropriate criterion is thus the percentage of normal straight-hinge D-larvae obtained. If mortality occurs in the D-larvae, it may well be owing to causes other than the contaminant being investigated. Mortality, growth and larval behaviour are, in our opinion, valid criteria to be used in sub-lethal bioassays lasting several days. 3.2.2.5. Abnormal development as the end point. In embryo-larva
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E. HIS, R. BEIRAS AND M. N. L. SEAMAN
bioassays, the end point of the experiment is attainment of the straighthinge D-larva stage. A precise definition of what constitutes a normal D-larva, i.e. the correct identification of abnormalities, is therefore of paramount importance. Loosanoff and Davis (1963) have pointed out that it is difficult to distinguish among D-larvae of different species simply by observing their external morphology. The photographs of about 20 species reared by Loosanoff et al. (1966), including the species commonly used in bioassays (C. virginica, C. gigas, Mytilus edulis, Mercenaria mercenaria and Mulinia lateralis), show that straight-hinge veligers of all species have the same general shape of a capital D. Identification of the species at this stage is generally possible only by observation of the hinge (position of the ligament, number and form of the teeth; cf. Le Pennec, 1978) under an electron microscope. The larval abnormalities in oyster and mussel larvae described below generally apply to all bivalve species employed in bioassays, Several authors have described abnormalities, but the criteria they employed differ. Loosanoff and Davis (1963), in descriptions applying mostly to C. virginica and M. mercenaria, attributed the presence of “winged” larvae (velum protruding from the shell; p. 31), and of “saddleback” and “humpback” larvae (concave and convex hinges; p. 33) in their cultures to a variety of factors, including excessive densities, poor gamete quality and presence of contaminants. Courtright et al. (1971), while attempting to devise an artificial seawater adequate for larval rearing, also defined various abnormalities in D-larvae of Mytilus edulis (hollow type, ear muffed, black spotted, exploded but swimming embryos; see their Figure 3, p. 88, op. cit.). In the same species, Le Pennec and Le Roux (1979) described six types of abnormalities of the prodissoconch identified by scanning electron microscopy (Photograph 3, p. 5). Renzoni (1975, Figure 1, p. 127) described the abnormalities induced by petroleum products in embryos and D-larvae of C. virginica and M. lateralis as “protruding velum and small larvae without a shell”. Larval abnormalities caused by lead salts have been described by Hrs-Brenko et al. (1977, see Figure 2, p. 110) in M. galloprovincialis and by Karaseva and Medvedeva (1993) in M. trossulus. Finally, Eyster and Morse (1984) photographed various abnormalities resulting from the exposure of gametes and embryos of Spisula solidissima to silver, under an optical microscope (their Figures 1 , 2 and 3) and scanning electron microscope (Figures 4 and 5, p. 643). The criteria employed by Eyster and Morse (1984) to distinguish between normal and abnormal larvae are nearly identical to those used at Arcachon, illustrated in Figures 7 and 8, for Crassostrea gigas (His and Beiras, 1995; His et al., 1997a). In contrast, the D-larvae of Mulinia lateralis photographed by Gormly et al. (1996, see Figure l A , p. 231) are abnormal,
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Figure 7 The different abnormalities observed in D-larvae of oysters, Crassostrea gigus; (a) normal D-larva; (b) convex hinge; (c) indented shell margin; (d) incomplete shell; (e) protruding mantle. (From His et al., 1997a.)
because the visceral mass is not completely retracted into the shell and the hinge is not completely straight. In order to achieve scientifically accurate assessments of embryo-larval bioassays, certain prerequisites must be met:
1. Larval abnormalities must be clearly defined and the definition must not be based upon observations of in vitro larvae, but rather on studies of larvae isolated from the natural environment (compare Figures 7 and 8). 2. The experimental protocol (duration of the bioassay and evaluation methods) must not introduce any biases with regard to the results. 3. The maximum percentage of abnormalities permissible in the control incubations (i.e. resulting from experimental artefacts) must be specified. (a) Definition of abnormalities. Woelke (1972) states, “normal larvae as referred to here, are those which are fully shelled, even though many may be misshapen or undersized”. D-larvae defined in this manner are a far cry from those sampled in situ, e.g. in Figure 8. Woelke himself admitted that he adopted these criteria in order to simplify the evaluation, but his definition greatly diminishes the quality of the bioassay. Similarly, Cardwell et al. (1977a,b, 1979b)
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E. HIS,
R. BEIRAS AND M . N.
L. SEAMAN
Figure 8 One-day-old D-larvae isolated from the plankton of Arcachon Bay in July 1985, after an intense spawning of Crassostrea gigas oysters. The growth and chemical composition of the larvae were followed through to metamorphosis. (His and Maurer, 1988.)
determined larvae developing abnormally, as those failing to complete shell development typical of the prodissoconch I stage (p. 5). Abnormality, as defined by ASTM (1980, p. 254), is: “incomplete or abnormal development is based upon the proportion of larvae failing to develop a complete shell after 48 hours (abnormal), relative to the total number of normal plus abnormal larvae. Larvae possessing misshapen or otherwise malformed shells are considered as normal, provided shell development has been completed” (emphasis added), and the same is the case in ASTM (1989, p. 346): “all larvae with complete developed shells containing meat must be considered as normal”. This obviously classifies as normal the “winged”, “saddleback” and “humpback” larvae considered abnormal by Loosanoff and Davis (1963, p. 33), and the various embryonal stages (normal or not) failing to complete development to the D-larva stage are excluded from the analysis. In contrast, Thain (1991, p. 51) defined as abnormal “those eggs which were not
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59
fertilized and those which died at an early stage of development or became malformed”, without, however, defining precisely what constitutes an abnormality. We concur with Klockner et al. (1985, p. 7), who defined normal veligers of C. gigas as follows: “24-hour-old normal D-shaped larvae exhibited (1) a fully developed symmetrical shell and (2) complete soft structures including the velum. Any retardation resulting in incomplete soft parts or deformed shells (asymmetrical shape, concave D-line, carved shells) were considered abnormal.” Similarly, Krassoi et al. (1996) in Chlamys asperrima, considered that 48 hours after fertilization “normal larvae are defined as those possessing a perfect D-veliger shell. Those larvae without the D-veliger shell are defined as abnormal (including trochophores, blastopores, larvae with misshapen shell plates)” (7.16, p. 29). At Arcachon (e.g. His and Beiras, 1995; His et al., 1997a), we have defined abnormality on the basis of a study of larvae of Crassostrea gigas isolated from the natural environment (His and Maurer, 1988). Figure 8 shows one-day-old D-larvae from the Bay of Arcachon after a mass spawning of the in situ oyster population, isolated from the plankton in the waters of the bay for laboratory investigations on growth and biochemical composition of wild larvae (His and Maurer, 1988). All of these larvae may be classified as perfectly normal straight-hinge D-larvae (compare with Figure 7b, c, d and e). Because of its narrow connection with the open sea the Bay of Arcachon represents a veritable natural hatchery (cf. Yonge, 1960, p. 165) with mass spawning of Pacific oysters occurring almost every year in mid-July. In more than 30 years of observations of larvae from the natural environment, we have never discovered any of the abnormalities shown in Figure 7. These observations form the basis for the definition of abnormality in Mytilus galloprovincialis (His and Beiras, 1995) and C. gigas (His et al., 1997a). (b) Recording the response. With regard to the mathematical assessment of abnormality levels, Davis (1960) and Davis and Hidu (1969a,b), referring to Loosanoff and Davis (1963), determined the response (R) simply as the percentage of eggs which develop into D-larvae:
R = Average number of larvae in experimental cultures x100 (1) Average number of larvae in control cultures
The larvae were counted after 48 hours by retaining them on a 44pm sieve, re-suspending them in a graduated cylinder with
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E. HIS, R. BEIRAS AND M. N. L. SEAMAN
filtered seawater, and taking sub-samples by pipette after mixing the water column in the cylinder. This procedure has been adopted by numerous authors, with or without use of equation 1,particularly by Calabrese in his studies on Mulinia lateralis. Woelke (1960a,b), working with C. gigas, also adopted the methods developed at Milford, retaining the larvae on a 37 pm sieve after 48 hours and counting sub-samples of about 150 to 250 larvae for the determination of the percentage of abnormalities (Woelke, 1967, p. 115). Cardwell et al. (1977a,b, 1979a,b) basically used the same methods, but omitted the sieving procedure after 1977, asserting that “concentration of veliger-stage bivalve larvae with nylon screening having a porosity of 37 pm or greater may bias toxicity test results significantly” (Cardwell ef al., 1978, p. 132; see also Bourne et al., 1981, and Chapman and Morgan, 1983). The recommendations of ASTM (1980, 1989) are based on the work of Cardwell et al. (1977a,b, 1979a,b). This requires counting the number of fertilized gametes placed into the containers, to compare with the number of “normal” D-larvae obtained after 48 hours. These counts are based on sub-samples of 10ml taken directly from the containers upon mixing, and sieving is thus avoided. ASTM (1989) proposes an alternative method with preparation of three beakers in addition to the experimental incubations for the sole purpose of making the initial counts. Klockner et al. (1985) sub-sampled 400 to 500 D-larvae of C. gigas and directly determined the percentage of abnormalities, whereas Thain (1991) sub-sampled aliquots of 2 ml from his 30 ml recipients, counting the larvae after addition of two drops of 8% formalin. Finally, Krassoi et al. (1996) added formalin to their 30 ml cultures 48 hours after fertilization; after decantation of the larvae they eliminated the top 28.5 ml by pipette and transferred the remaining 1.5 ml to a Sedgewick-Rafter cell for microscopic examination. None of these authors practised sieving, nor did they compare the number of D-larvae obtained to the original number of fertilized gametes introduced. In contrast His and Beiras (1995) and His et al. (1997a), have adopted a technique using transparent polypropylene vials and 3 ml tissue culture microwells (after Cherr et al., 1990), eliminating any need for sub-sampling or preliminary counts, because all of the biological material introduced in the test is assessed at the end of the experiment. Moreover, sieving and sub-sampling even with an automatic pipette may introduce errors, because some of the material originally introduced (e.g. decomposing segmented eggs and malformed embryos) may be destroyed by these procedures (cf. the observations by Cardwell et al., 1978).
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THE ASSESSMENT OF MARINE POLLUTION
(c) Permissible percentages of larval abnormalities in control cultures. Woelke (1966, 1972) considered that abnormalities in control incubations should be less than 2.5%. They state, “between 5 and 10% we are extremely cautious in making firm conclusions; we totally reject any bioassay where our controls have in excess of 10% abnormal” (Woelke, 1961, p. 119 and 1966, p. 34). It must be kept in mind, however, that he sieved his larvae after 48 hours and used a very restricted definition of abnormality. Woelke expressed his results as a percentage of abnormalities when his controls exhibited less than 2.5% abnormal larvae. When his controls had a higher percentage of abnormalities, he calculated “percent net risk” (sic) - originally percentage net response (PNR) - by subtracting the percentage of abnormal controls from the percentage of abnormalities in the test incubations (originally Abbot’s equation in Emmens, 1948; Finney, 1971): PNR =
(% treatment abnormal - % control abnormal) X 100
100 - YOcontrol abnormal
(2)
Cardwell et a/. (1977a,b, 1979a’b) and ASTM (1980,1989) adopted Woelke’s definition of abnormality and Abbot’s equation (equation 2), but using an acceptable level of 10% abnormal larvae in the controls. Finally, Klockner et al. (1985) were much more restrictive in their definition of normality and based their evaluation on the Net Risk (NR): NR (%)
= (NT-NR) X
100NR
(3)
where “NR [is the] net risk - value of the test water sample (YO), NT [is the] mean relative number of normally developed larvae in replicates of test water samples, [and] NR [is the] mean relative number of normally developed larvae in replicates of reference water samples. NT and NR refer to the rates of normal larvae (raw data) on the basis of the total number o f . . . normal and abnormal larvae hatched together” (p. 9). The validity of their assessments is limited, however, by the fact that the percentage of normal larvae in their controls was merely 65.3% 5 3.3%. In our direct counts in culture vials (His and Beiras, 1995; His et al., 1997a), conducted 24 hours after fertilization in oysters and after 48 hours in mussels, we considered a test invalid when larval abnormalities attained 20% in the controls, whereas Thain (1991, p. 7) judged that “control abnormalities up to 50 or even 60 per cent are acceptable”.
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M. N. L. SEAMAN
In publications of bivalve embryo-larval bioassays, we think it is indispensable to mention the number of abnormally developed larvae in the controls (including the 95% confidence interval), to enable readers to appreciate the validity of the test. These data are usually absent in publications where the data are evaluated according to equation 2. 3.2.3. Tests of larval growth
These tests consist of determining sub-lethal responses by measuring the growth of the larval shell (cf. Widdows, 1993). It is interesting to note that the first anomalies detected in the field or in hatcheries did not concern embryogenesis, which proceeded normally, but larval growth. As mentioned above, toxicants usually induce a highly heterogeneous size distribution compared to healthy growing larvae (see Figure 6). The tests of larval growth are necessarily of longer duration (a few days, to several weeks if the pediveliger stage is to be studied) and require feeding the larvae. They are usually performed in beakers of 1litre (static tests), even though flow-through tests in running seawater are preferable (Calabrese, 1984, p. 460; cf Roberts and Gleeson, 1978; Roberts, 1980). These tests are even less standardized than embryo-larval bioassays. The size or growth of larvae during toxicological experiments is assessed by measuring the prodissoconch I and 11. The anatomically correct definition of these measurements (Figure 9) has been described by Galtsoff (1964) for adult oysters, and it applies equally to bivalve larvae (see also Bayne, 1976, p. 83). The following definitions are given by Galtsoff (1964, p. 18): “The height is the distance between the umbo and the ventral valve margin. The length is the maximum distance between the anterior and posterior margin measured parallel with the hinge axis. The width is the greatest distance between the outsides of the closed valves measured at right angles to the place of the shell commissure.” These measurements are nevertheless often specified incorrectly in the literature, even to this day (e.g. shell height is termed “width” by Loosanoff et al. (1966), and it is commonly termed “length” by many authors). It is therefore very important that authors, when reporting results using shell dimensions, do not limit themselves to using these terms (whether correctly or incorrectly); they must unequivocally specify how the determination was made in order to avoid confusion. In tests of larval survival and larval growth, it is necessary to distinguish between studies in which fertilization and embryogenesis have taken place in unpolluted conditions (the investigation is initiated exclusively with healthy D-larvae), and studies in which fertilization has taken place in the
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height height Prodissoconch I Urnboned (Prodissoconch 11)
height Dissoconch
Figure 9 The dimensions of the bivalve larval shell (prodissoconch I - D-larva and prodissoconch I1 - umboned larva), and of the adult shell (dissoconch). C-C’ = hinge axis. (From Le Pennec, 1978.)
presence of the contaminant. The importance of such differences to sensitivity has been recognized by all authors who have conducted ecotoxicological studies of growth in bivalve larvae. We have found toxic effects on larval growth and survival at copper concentrations five times lower when larvae were reared in the presence of metal from fertilization on, compared to larvae developed from unexposed gametes (His and Robert, 1981, 1982). An intermediate possibility is to begin exposure to the toxicant minutes after the start of fertilization. 3.2.3.1. Studies on growth inhibition with previously unexposed larvae. The first studies of bivalve larval growth in the presence of pollutants were conducted by Davis and Chanley (1955b), who investigated the effects of various antibiotics, and by Davis (1961), who demonstrated the toxicity of 32 pesticides on larvae of Crassostrea virginica and Mercenaria mercenaria. These experiments were concerned first with embryogenesis and the formation of D-larvae, and then with the subsequent development of D-larvae obtained in the absence of contaminants during a further 12 days of contaminant exposure. Growth and mortality were determined only on the twelfth day. Further laboratory studies were conducted at Milford by Hidu (1965) on the effects of synthetic surfactants on M. mercenaria; Calabrese and Davis (1966,1967) studied the effects of pH on C. virginica and M. mercenaria and of linear alkylate sulphonate (LAS), a surfactant, on C..virginica; Calabrese (1972), Calabrese and Davis (1967), Calabrese and Nelson (1974) and Calabrese et al. (1977a) then investigated the effects of heavy metals and various other pollutants such as detergents and pesticides. Finally, Calabrese and Rhodes (1974) applied the same techniques in studies with embryos and larvae of the coot clam, Mulinia lateralis, demonstrating the advantages of using this species in bioassays. The results obtained at Milford laboratory were summarized by Calabrese et al. (1977b, 1982). In studies of the effects of contaminants on larval growth using previously unexposed (healthy) larvae, the embryos were raised in an
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uncontaminated mass culture, collected on a sieve 48 hours after fertilization, counted and redistributed into the experimental beakers at concentrations of 5000 to 15 000 per litre. Incubation generally lasted about 10 days, except in the case of M. lateralis, which attains settlement competence after 6 to 8 days (cf. Calabrese, 1970b). The incubation water was changed every 2 days, always maintaining the appropriate contaminant concentration. To this end, the larvae were first collected on a sieve and sampled to determine mortality, possible abnormalities and shell size. Larval size was measured by ocular micrometer to a precision of 2 5 pm (cf. Calabrese, 1970b). Measurements can also be made by photographing samples of larvae and later measuring the diameter of the shell under a binocular microscope, either on the photographs or on the negatives (e.g. His and Robert, 1981, 1982). This is usually done with a video imaging system (e.g. Widdows et al., 1989; McFadzen, 1992). Larval weight is rarely measured (Widdows et al., 1989; Beiras and His, 1994, 1995a) despite the fact that it is required to express lethal concentrations on a mass-specific basis, enabling comparison of data among larvae of different sizes (see Section 4.2.3). In the ecotoxicological studies with Mytilus edulis and Pecten maximus by Beaumont et al. (1987), and those with Mercenaria mercenaria by Byrne and Calder (1977), larval feeding was based on the methods reviewed by Loosanoff and Davis (1963) (see Section 2.3.5). Renzoni (1975) studied the effects of the water soluble fraction of three petroleum products on growth in Mulinia lateralis during a period of 2 weeks, changing the water (and renewing the pollutant concentration) every 2 days and feeding the same algae as used by Calabrese (1970b). Laughlin et al. (1989) in studies of the effects of TBT on Mercenaria mercenaria fed the larvae exclusively with Isochrysis galbana (Tahiti strain). Finally, Gormly et al. (1996) studied the resistance of Mulinia lateralis to microbial pest control agents during 48 hours in microwells of 3 ml (15 to 20 larvae per well); the larvae were fed during these brief experiments on mortality and growth of the larvae. Some authors obtain their larvae from commercial hatcheries and this may pose various problems. Such larvae are generally the progeny of a haphazard mix of parents, and they may also represent a mix of incubations of different ages which have all been passed through the same sieves to obtain a homogeneous size; i.e., they may be the result of the culling procedure employed by commercial hatcheries to rid themselves of slow growers. Watling (1978, p. 126, and 1982) mentions that in the hatchery-reared larvae used “the range of individual sizes was restricted at the beginning of the experiments”, which may pose a problem in studies of larval growth.
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3.2.3.2. Studies on growth inhibition with D-larvae already exposed to contaminants. Studies in which fertilization and embryogenesis were conducted in the presence of the contaminant being bioassayed are rare, even though they represent the best approximation of in situ conditions. Granmo (1972) first distributed eggs of Mytilus edulis at a density of 10 per ml into vials with various concentrations of LAS (a surfactant) before proceeding to fertilize them. The water was subsequently changed daily during the first 8 days, the larvae being retained on a mesh of 60 pm at each change of water. Fertilization success was assessed after 20 to 24 hours, and larval mortality and size (“measured microscopically, the greatest length attained being recorded”, p. 356) were subsequently determined after 72 hours, 96 hours and 240 hours. Brereton et al. (1973) studied the effects of several concentrations of zinc sulphate on freshly fertilized eggs of Crassostrea gigus (15 000 per beaker of 1 litre) at a temperature of 20°C to 22°C and a salinity of 29 psu. Larval abnormalities were counted after 48 hours, and the D-larvae were subsequently passed through a 100 pm sieve to remove impurities, retained on a 45 pm sieve and incubated at a density of 5000 per litre at 26°C. They were grown for 5 days in the presence of the contaminant and another 5 days in unpolluted seawater, and fed daily with Zsochrysis galbana, Chaetoceros calcitrans, Monochrysis lutheri and Cyclotella nana at 100 cells per pl of larval culture volume. The water was changed every 48 hours, and at this time swimming behaviour was recorded, and mortality and growth were assessed. Shell size (“width across the valve”) was determined in 25 individuals of each of four replicates of each concentration. Laughlin er al. (1988) studied the toxicity of TBT to Mercenaria mercenaria; the eggs were exposed to the contaminant 4 hours after fertilization at densities of 150 per bowl of 50 ml, and the incubation lasted a total of 14 days. The solutions were renewed daily except during the first 48 hours, and the larvae were fed with Zsochrysis galbana (Tahiti strain) at 40 000 cells per ml. Size was determined daily in sub-samples of 25 larvae per incubation. The first studies of larval growth undertaken at Arcachon followed the methodology of Brereton et al. (1973) and Helm and Millican (1977). They were initially applied to investigate the effects of TBT on Crassostrea gigas and Mytilus galloprovincialis, showing the threat posed by this compound to coastal shellfish culture areas (His and Robert, 1980; Robert and His, 1981). These experiments were carried out with larvae that had been reared through embryogenesis in the presence of the toxicant, as well as with larvae reared to the D-larva stage in unpolluted water. The eggs were distributed into 2 litre beakers at a density of 60 per ml; oyster embryos were incubated for 24 hours at 20°C and mussel embryos for 48 hours at 19°C. After this period the contents of each beaker were passed through a 100 pm sieve to remove impurities, and the larvae were collected on a
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sieve with 40 pm mesh. After assessment of larval abnormalities the larvae were re-suspended in 250 ml graduated cylinders for recounting and incubation then continued at densities of eight individuals per ml for 7 days (only 5 days in the case of TBT, owing to the high toxicity of the concentrations tested). The water (0.2 pm filtered seawater) was changed daily and the larvae were fed daily as well. Mortality was assessed in each incubation, and larval size was determined with an ocular micrometer in sub-samples of 50 individuals. Studies of growth were subsequently extended to 10 or 12 days, and the investigations with larvae formed in unpolluted water were abandoned because of their lesser sensitivity. The feeding regimes of Helm and Millican (1977) were used at all times. The water was changed only every second day, and mortality and growth were therefore equally assessed in intervals of 2 days; shell size was determined by photographing sub-samples of larvae, rather than directly under the microscope. After studying the effects of TBT on larvae of Crassostrea gigas at concentrations around 1pg per litre, further studies were conducted on the effects of copper sulphate, cadmium chloride and salinity, and atrazine-simazine, a herbicide (His and Robert, 1982; His et al., 1983; Robert and His, 1985; Robert et al., 1986). The method has lately been modified further because of the impossibility of conducting numerous fertilizations in many different contaminant concentrations almost simultaneously. We therefore now prefer to add the sperm to the graduated cylinder immediately after determining the density of the egg suspension, distributing the gametes into the experimental vials 15 to 30 minutes after fertilization (Section 3.1.3.6). This method has been used to study the effects on C. gigas of 12 pesticides (His and Seaman, 1993), of mercury (Beiras and His, 1994) and of aluminium-plant effluents (His et al., 1996), as well as the effects of mercury on Mytilus galloprovincialis (Beiras and His, 1995a). 3.2.4. Tests on larval physiology These bioassays concern the effects of pollutants and physico-chemical environmental impacts on larval behaviour or activity, such as feeding activity, respiration, resistance to hypoxia, swimming behaviour and metamorphosis. Physiological analyses of bivalve larvae are not easy to conduct and may produce doubtful results, particularly in toxicity assays accompanied by larval mortalities and ensuing bacterial proliferation; for instance, in a study on the feeding behaviour of oyster larvae (Seaman and Ueberschar, unpublished), the samples in which all the larvae were dead had by far the highest activities of trypsin, a digestive enzyme produced by animals and bacteria alike.
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3.2.4.1. Feeding activity. After studying the effects of copper chloride, cadmium chloride and zinc chloride on the growth of algae (Monochrysis lutheri, Isochrysis galbana, Dunaliella euchlora and Phaeodactylum tricornutum), Wikfors and Ukeles (1982) proceeded to study the transfer of these contaminants to the next trophic level by feeding larvae of Crassostrea virginica with axenic cultures of 1. galbana grown in sub-lethal concentrations of copper chloride (10 mg% = 47.3 ppm Cu) and zinc chloride (2.5 mg% = 15.3ppm). The digestive tract of the larvae was observed by epifluorescence microscopy (study of the fluorescence of the algae ingested, by the method of Babinchak and Ukeles, 1979), showing that the larvae fed normally and digested the algal cells. Larval growth, however, was strongly affected, showing the potential for bioaccumulation of pollutants beginning with the level of primary production: “The phytoplankton can exhibit a limited increase in tolerance and adaptation to metals. These populations are then potentially toxic for grazing species at higher trophic levels” (Babinchak and Ukeles, 1979, p. 191). Wikfors et al. (1993) and Wikfors and Smolowitz (1995) also studied the effects of the toxic dinoflagellate Prorocentrum minimum on larval nutrition in C. virginica, again using epifluorescence microscopy. They were unable to conclude from their results that the reduction of larval growth and survival were a direct result of the toxic effect of the dinoflagellate. Widdows et al. (1989) used fluorescent microspheres (3.4 pm diameter) derived from autoclaved algal extracts to study the effects of hypoxia and anoxia on the ingestion rate of prodissoconch larvae (73 to 140 pm) and pediveligers (300 to 376 pm); the microspheres ingested were counted under an inverted epifluorescence microscope. In contrast to the pediveligers, the prodissoconch larvae continued to feed under anoxic conditions. 3.2.4.2. Respiration. There are few studies on respiration in bivalve larvae beyond those of Morrison (1971) and the investigations of hypoxia resistance by Widdows et al. (1989) and Baker and Mann (1992, 1994a), particularly with regard to the effects of pollutants. Thurberg et al. (1975), using a microrespirometer developed by MacInnes and Thurberg (1973), studied respiration rates in larvae of Spisula solidissima exposed to 50 ppb of silver (added as AgN03) at 2-day intervals during a 15-day period (initial and final size of the controls 73 and 243 pm, respectively). Larvae exposed to the pollutant had respiration rates one-third higher than the controls. Pavicic (1980) studied the combined effects of cadmium and zinc on respiration in embryos and larvae of Mytilus galloprovincialis. Oxygen consumption was measured during embryogenesis and into the D-larva stage at temperatures ranging from 10°C to 20°C. Fertilized eggs were placed in hermetically sealed incubation vessels of 500 ml volume at
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densities of 30 to 100 eggs per ml, and dissolved oxygen levels were determined by the Winkler method after 48 hours to 67 hours. The results were inconclusive “due to the difficulties of measuring low respiratory rates of bivalve larvae with sufficient accuracy to detect physiological changes resulting from the different treatments” (p. 63). 3.2.4.3. Swimming activity. Chang et al. (1996) studied the effects of copper and zinc fixed on biofilms in cell culture dishes (1.5 ml, each with 20 to 30 larvae) by observing under the microscope “speed of swimming, ciliary activity and the number of quiescent or inactive larvae” (p. 590) at different metal concentrations. The most sophisticated work on swimming behaviour of bivalve larvae has been undertaken by Mann and Rainer (1990) in larvae of C. virginica subjected to low oxygen concentrations, and by Mann et al. (1991) who studied the response of larvae of Spisula solidissima, Mulinia lateralis and Rangia cuneata to salinity discontinuities. These authors observed the behaviour of veligers in glass cylinders 30 cm high and 0.6 cm in diameter (with an approximate volume of 10.8ml), either by recording the motion of the larvae with a high-resolution video camera (Mann and Rainer, 1990), or by simple observation under a binocular microscope and recording the movements on graph paper (Mann et at., 1991). Van den Hurk (1994) studied the distribution of larvae of Crassostrea gigas in the water column of 1litre jars, in order to compare the behaviour of larvae exposed to a toxic sediment with that of larvae in unpolluted filtered seawater. In the absence of sediment the larvae remained at the bottom of the beakers, but moved towards the surface in the jars with sediment. The author concluded that this represented a strategy for avoidance of the sediment. We conducted preliminary trials to study the response of C. gigas oyster larvae to mercury. Setting up eight 25 ml vessels with a sub-lethal mercury concentration of 16 pg 1-1 and eight controls with seawater, we added 5000 larvae to each vessel and frequently sampled the upper water layer with a Id pipette. Figure 10 shows the mean numbers of larvae in these samples. During the first few hours there was a marked difference in the vertical distribution of mercury-exposed and unexposed veligers. The former tended to concentrate in the upper part of the experimental vessels, perhaps as a result of increased swimming activity. However, this kind of result must be interpreted with caution. Healthy oyster larvae (C. gigas as well as Ostrea edulis) often have a tendency to remain in the upper part of the water column when kept in beakers of 1 or 2 litres, as shown by our own observations in 20 years of larval culture and those of Wilson (1981). The umboned stages tend to float at the water-air interface until the appearance of the eye spot, forming characteristic dense mats. This phenomenon, termed “rafting” by Wilson, is “an
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140
120
F
100
Q)
2
Y-
80
O
b
Q
E 3
2
60 40
20
0 I
I
I
I
I
0
10
20
30
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I
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Time (h) Figure 10 Density (I&’) of Crassostrea gigus larvae actively swimming near the surface of the 25 ml vessels when exposed to 16 pg Hg 1-’ (0) and in seawater controls (0).Values are means ( 2 SD) of eight samples from each different vessel. Note the sigmficantly increased upward swimming activity in the mercury-exposed larvae during the first hours of the experiment.
indication of optimum conditions, and in our experiments such cultures grow and settle well” (Wilson, 1981, p. 21). On the other hand, moribund D-larvae and umboned larvae of oysters tend to accumulate at the bottom of the incubation vessels (cf. Galtsoff, 1964, p. 376). These observations do not apply to mussel larvae, which tend to remain at the bottom of the beakers when they are doing well. Therefore, little is known on the sub-lethal effects of toxicants on the swimming activity of veliger larvae, but this is a promising field for future studies. 3.2.4.4. Calcium uptake. Ho and Zubkoff (1979, 1980, 1982) have proposed “a sensitive and rapid bioassay”, using 45Caisotope uptake by
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larvae (Ho and Zubkoff, 1982, p. 409). Their first study (1979) concerned calcium uptake in larvae of C. virginica over a period of 68 hours in the presence of the toxic dinoflagellate Cochlodinium heterolobatum, following a phytoplankton bloom. The algae caused mortality in the larvae, as well as a reduction in calcium uptake. The method, described in detail in Ho and Zubkoff (1980, p. 144) was also used to study the effects of mercury, copper and zinc on Mercenaria mercenaria, showing that copper has the highest, and zinc the lowest toxicity of the three metals, both with respect to mortality and to calcium uptake; they suggested that the method be used for toxicity monitoring. In another study (Ho and Zubkoff, 1982), they found that calcium uptake by larvae was depressed by nickel, but stimulated by cadmium. 3.2.4.5. Metamorphosis. The last larval stage of bivalves, the pediveliger, considered particularly sensitive to pollutants (Watling, 1983), has been used in some 20 ecotoxicological studies. In the 1930s Prytherch was the first to record metamorphosis in Crassostrea virginicu on film; some of his films were recovered by Medcof (1961). Prytherch (1931,1934) showed that concentrations of 0.15 to 0.50 mg Cu 1-1 induce settlement in competent larvae of C. virginica. He interpreted this as an avoidance reaction to the contaminant, thus making the first ecotoxicological observation using pediveligers. Most studies with pediveligers have investigated the effects of heavy metals (Boyden et al., 1975; Watling, 1978, 1983; Nell and Holliday, 1986; Beiras and His, 1994). Sheffrin et al. (1984) worked with plantigrades, the first post-metamorphic stage. Chang et al. (1996) and Phelps and Mihursky (1986) investigated the effects of metals in the microbial layer of settlement surfaces. Ruiz et al. (1995b) studied the effects of TBT on metamorphosis in Scrobicularia plana. Smith and Hackney (1989) conducted a field study on the effects of hydrocarbons on in situ settlement in C. virginica. Various bioassays have been conducted on the environmental impacts of nuclear-energy plants, particularly with regard to the various treatments employed for the prevention of biofouling in tubes and canals. Studies focused mainly on the toxic effects of chlorination and ozonation on oyster and mussel pediveligers (Bucaille and Kim, 1979; Roosenburg et al., 1980a,b; Richardson et al., 1982; Roberts and Casey, 1985). The ability of the pediveligers to undergo metamorphosis has also been employed to assess the quality of marine sediments (e.g. Phelps and Warner, 1990), since the pediveliger is the earliest and most sensitive phase of the oyster that comes in contact with the substratum. The same idea motivated His et al. (1997b) to investigate the negative effects of a sediment polluted by polycyclic aromatic hydrocarbons (PAH) on recruitment in C. gigus.
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Two types of studies may be distinguished with regard to their methodology: those conducted before the importance of chemical inducers of metamorphosis had been unravelled, and those which were able to profit from the work of Coon et al. (1985,1986,1990), Coon and Bonar (1986) on metamorphosis in C. gigas and C. virginica, and of Shpigel et al. (1989) on Ostrea edulis. Metamorphosis is induced by L-Dopa (dihydroxyphenylalanine), and epinephrine, making it possible to obtain cultchless spat (metamorphosis without attachment to a settlement surface) in oysters. Cultchless spat are important in commercial and laboratory culture of oysters, because it is easier to collect post-larvae from the water column than from the walls of the incubation vessel. It should be pointed out that various authors (Phelps and Mihursky, 1986; His et al., 1997b) have conducted their studies with late veligers obtained from a commercial hatchery and raised within days to the pediveliger stage in the laboratory. Some authors (e.g. Watling, 1983) have worked with hatchery larvae having settlement competence, as these (eyed) larvae can easily be “shipped by overnight express under moist and cold conditions” (Chang et al., 1996, p. 590). Tests on pediveligers without chemical induction of metamorphosis. Boyden et al. (1975) and Watling (1978,1983) exposed hatchery larvae of Crassostrea gigas (16 to 23 days old, 240 pm to 300 pm in size) to heavy metals for 5 days at densities of 1000 to 15 000 larvae per 12 litres. The bottom of each beaker was covered by a ground glass plate and “a black PVC disc slightly arched to make available both upper and lower surfaces for settlement” (Boyden et al., 1975, p. 228; Watling, 1983, p. 345). The collectors were placed in unpolluted flowing seawater after 5 days, and the number of metamorphosed larvae and the mortality 30 days after contaminant exposure were assessed. Phelps and Mihursky (1986) placed 30000 eyed larvae of C. virginica in 1.5 litre beakers with 0.2pm filtered seawater. The bottom was covered with shell debris and coppercontaminated microbial films. After 4 hours the larvae were transferred to flowing estuarine water, filtered at 0.5 pm to allow the passage of food algae. The number of spat was assessed after 4 to 7 days. Richardson et al. (1982) reared their own larvae to the pediveliger stage in 9 to 21 days (sic). As soon as most of them had attained the eyed stage and the first settlements were observed, the larvae were placed in “shallow setting trays lined with black polyethylene sheeting and the entire bottom covered with numbered 2.5 by 7.6 cm glass microscope slides which are used as experimental substrate” (p. 101). They were exposed to ozone produced oxidants (OPO) and chlorine produced oxidants (CPO) as soon as 100 to 200 larvae “in the late crawling stage - not definitively fixed” were observed on the slides. Mortality was determined after 12, 24, 48 and 96 hours by staining the
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larvae with neutral red (only live larvae acquire the stain). They do not mention data on metamorphosis success. The effects of CPO on Mya arenariu and C. virginicu larvae were also studied by Roosenburg et al. (1980a,b) using a similar experimental protocol. D-larvae and pediveligers were exposed to CPO in a flowthrough setup. The latter were exposed to the contaminant after having settled; as in the case of Richardson et al. (1982), the larvae were collected when they attained the “swim-crawl phase prior to settlement” and placed at densities of 500 per ml in 8 litre fibreglass tanks, the bottom of which was covered with microscope slides. The tanks were filled with filtered seawater, and the larvae were allowed to settle on the slides. The number of fixed spat was then counted and the slides were cleaned except for sections of 4 cm*containing an average of 100 to 162 spat. The slides were then exposed to CPO in a flow-through system. Dependent variables were larval resistance to the pollutant and detachment of the spat from the settlement surfaces, as a function of concentration and duration of the exposure (8,24,48,72 and 96 hours in oysters; 4 hours, 6 hours, and the same intervals to 72 hours in clams). The authors mention percentage mortality and detachment, but they do not give clear data on the percentage of metamorphosed larvae. Studies on the effects of chlorinated seawater on fixation and detachment of mussels (Mytilus sp.) have been conducted by Bucaille and Kim (1979). Their experimental protocol is not described in detail, but they too worked with a flow-through system with continuous and discontinuous chlorination, first observing settlement in comparison to the controls (which had 90% settlement), and then the detachment of larvae which had initially settled in the absence of chlorination. Nell and Holliday (1986), working with Saccostrea commercialis in an experimental hatchery, investigated settlement behaviour in the presence of K+ and Cu2+in larvae retained on a 300 pm mesh (diagonal measurement) and placed at a density of 2000 larvae per litre in 8 litre aquaria painted black (four tanks per treatment). Grey polyvinyl sheets (300 X 150 X 0.8 mm) held 5 mm from the bottom of the tanks were used as a settlement surface. The aquaria were dimly lit at 100 lux, and weakly aerated. The water was changed daily, and the larvae were fed daily with 75 000 cells per ml of Zsochrysis gatbana and Monochrysis Eutheri. “The number of live settled spat that could not be washed off the sheets by gentle rinsing with sea water was counted and the viability of the larvae checked daily” (p. 264). Finally, Ruiz et ul. (1995a), attempting to demonstrate the effects of TBT on pediveligers of Scrobicularia plana, confronted technical difficulties which make their method hardly applicable in routine bioassays. The rates of successful embryogenesis were only about 10%. One
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advantage of S. plana, on the other hand, is that pediveligers of the “swimming and crawling” stage can be obtained in the laboratory in little less than a week, making it possible to conduct metamorphosis assays more rapidly than with other species. Competent larvae require sediment in order to metamorphose, and the authors used “acid washed sand ground with a mortar and pestle until it passed through a 1OOpm pore mesh, and autoclaved” (Ruig et al., 1995a, p. 120), deposited as a layer 3mm in thickness at the bottom of the experimental vessels of 90ml volume. They placed 70 larvae (10-day pediveligers about 240pm in length) in each vessel. The spat were exposed to four concentrations of TBT (ranging from 10 to 100 ng Sn 1-‘) for 30 days, during which the water was changed every 2 days. Some of the difficulties encountered by the authors concerned the use of sediment: “the definitive acceptance of sand as settling substrate seemed to be delayed in controls” (p. 121), “and the sand may not have been optimum substratum” (p. 123). It was also difficult to separate the larvae and the sand, and the alcohol used as a solvent for TBT provoked an excessive secretion of mucus leading to aggregation of the larvae and ensuing mortality. The use of solvents is sometimes unavoidable, but at least in the case of oyster larvae, alcohol is much more toxic than dimethylsulphide (DMSO) and acetone (His, unpublished data; cf. His and Seaman, 1993). The authors based their assessment on the growth and survival of the larvae and spat of S. plana. Tests on pediveligers with chemical induction of metamorphosis. The chemically mediated induction of settlement in oyster larvae pioneered by Coon and coworkers (op. cit.) has opened new possibilities for bioassays involving metamorphosis of pediveligers. Thus, Phelps and Warner (1990) studied the potential toxicity of contaminated sediments on metamorphosis of hatchery-reared C. gigus and C. Virginia. The stock of veligers was kept in aerated 2.7 litre beakers with artificial seawater (ASW) treated with antibiotics; the water was changed every 2 days (larvae retained on 149 pm sieves) and the larvae fed daily with Monochrysis and Zsochrysis. For the bioassays the larvae were placed in ASW with 0.001mol epinephrine for 20 hours, and the sediments were tested at 21°C in 2.75 ml wells with 1ml of sieved sediment and 1ml of water in tissue culture plates (24 wells per plate). After 96 hours the contents of each well were removed by automatic pipette and sieved through a 149 pm mesh. Larvae were classified as live, metamorphosed or dead after staining with neutral red: “Dead larvae were often grey and opaque with no evidence of internal organization or movement. Some dead larvae showed retraction or partial decomposition of tissues and some had invasion by protozoa. Metamorphosis was determined by the transition from rounded veliger to flat shape with new shell growth” (p. 199).
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The same method was used by Beiras and His (1994) to determine the effects of mercury on metamorphosis in C. gigas, with epinephrine being used at a final concentration of lop4M. Following the methods of Beiras and Widdows (1995), competent eyed larvae (size greater than 290 pm) were placed in 3 ml wells of tissue culture plates at a density of 11 per ml. The number of post-larvae (see Figure 3) was assessed under the microscope after 2 days of exposure to epinephrine and toxicant. In the same way, the toxicity of sediment contaminated with PAH was evaluated (His et al., 1997b). These two tests are prime examples of sub-lethal bioassays, as the larval mortality was extremely low (less than 3% in all cases). The criteria used to determine metamorphosis success were: loss of the velum and foot; plainly visible outline of the gills; secretion of the dissoconch, which was often slight (presumably owing to pollutant effects) but nevertheless visible as a translucent veil, as opposed to the strongly coloured prodissoconch I1 (see Figure 3).
3.3. Bioassay applications: Toxicity tests and environmental bioassays
3.3.1. Toxicity tests with pure chemicals Toxicity is primarily a function of the amount of toxicant available to the organism, i.e. the dose, and the duration of exposure. In aquatic organisms the toxicant is often present in solution and its concentration is taken as the dose. Toxicity can then be evaluated either by fixing an exposure time and varying the concentration or by fixing a concentration and varying exposure time. In the first case the percentage of test individuals exhibiting a biological response at each concentration is determined after the exposure period, and the theoretical concentration that would produce a response in 50% of the individuals is calculated (see below). This is the median effective concentration or EC50.In the second case the biological response after different periods of exposure is measured and the theoretical time at which a response occurs in 50% of the individuals at a particular concentration is calculated, this is the median effective time or ETSO.When lethal toxicity is studied the equivalent parameters obtained are the LC50 and LT,,, respectively. The difficulties associated with holding marine animals in the laboratory for long periods and maintaining a constant concentration of toxicant dissolved in the water have made the and LCS0)more popular. first approach (calculation of The toxicity of pure chemicals to marine organisms has been investigated in the laboratory by measuring a biological response, e.g. successful embryogenesis, at different concentrations of a toxic compound dissolved
THE ASSESSMENT OF MARINE POLLUTION
75
in seawater (see Section 3.4.1). The results can be represented by a concentration-response curve. For non-polar chemicals a non-toxic carrier (e.g. acetone or DMSO) is recommended and carrier controls must be included in the experimental design. Acute toxicity studies are those studies which investigate the effects of toxicants after short exposure periods (from a few hours to a few days), and which usually assess fertilization, embryo development or mortality; whereas chronic toxicity studies are those which use exposure periods lasting for most of the larval life span (at least one week), and which assess larval growth as the measure of toxicity. When data on the toxicity of the chemical to be studied are not available, it is advisable to conduct preliminary trials, testing concentrations in log" steps to find the range of no effect to 100% mortality. Once this range is known Concentrations should be tested in smaller steps, depending on the reliability of the nominal concentrations (i.e. measured actual concentrations must be significantly different among treatments). Since extrapolation in regression models is strongly ill-advised, a good test should always include concentrations causing both more than 50%, and less than 50%, response, even though concentrations giving 0% and 100% response are not to be included in the calculations of the LC50and related parameters. On the other hand, partial responses are necessary for most methods of calculation of the LCSo(see Section 3.4.1). Many toxicants tend to disappear from the water column during the bioassay because they are volatile, adsorb to the walls of the test vessels, or react with other compounds in the water. Therefore, the actual concentrations of the pollutants must always be measured at the beginning and at the end of the test, to make sure they do not deviate from the nominal concentrations. Dilution water should be 35psu artificial seawater made up under axenic conditions, or filtered (0.45 pm or lower) seawater of oceanic characteristics (see Section 3.1.1). When natural oceanic seawater is used, the concentrations of the main chemical toxicants should be analysed first. The containers used to make up the toxicant solution and to conduct the bioassays depend on the chemical properties of the toxicant and should be selected to minimize possible losses. Plastic containers (polypropylene or PTFE) should be used when assaying metals, while glass is preferred for organic compounds. Containers should be capped with an airtight seal. If the density of the test organisms is too high in static bioassays, spurious toxicity may result from low oxygen concentrations or ammonia excretion by the larvae (see Section 3.2.2.2). Experimental concentrations should be made up from a stock solution of the toxicant dissolved in distilled water at a concentration at least ten times higher than the maximum concentration to be tested. This limits the
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reduction in salinity to 10% or less when the required amount of stock solution is added to the incubation seawater. A 10% reduction from oceanic salinity (i.e. to 31.5 psu) does not affect the gametes, embryos or larvae of the marine bivalves reviewed here. The rest of the experimental concentrations can be prepared by diluting in seawater the higher concentration, using volumetric flasks. Replicates without animals should be run to permit the removal of samples for chemical analyses. The toxicity tests described here aim to ascertain the EC5* of the chemical being investigated. The EC5o has several applications:
1. It directly describes the degree of toxicity of a single chemical; the lower the EGO,the more toxic the chemical. 2. It can be used to study interactions between two chemicals, providing a common unit to compare the effects of toxicants acting singly or in combination. One Toxicity Unit (TU) is the concentration of the chemical divided by its EC5o. If a mixture of a certain number of TUs of two chemicals causes the same toxicity as the total number of TUs of each single chemical alone, then their interaction is additive. If the toxicity of the mixture is greater, the interaction is synergistic, and if the toxicity of the mixture is significantly lower, the interaction is termed antagonistic (see also Section 4.4). 3. Toxicity tests are an integral part of environmental risk assessment of chemicals likely to be released into the environment. Safe concentrations can be determined by multiplying the EC50 by a safety factor (generally 0.01). The safety factor is designed to account for the disparity between the results of an acute toxicity test in the laboratory and the situation in the natural environment, including possible effects of long-term (chronic) toxicity, and possible synergistic interactions with other pollutants in the natural environment.
3.3.2. Bioassays with samples from the natural environment. Rationale for the use of bioassuys in monitoring marine pollution Most contaminants originating from human activities enter the marine environment via atmospheric deposition, fluvial inputs, or direct spillage into the sea. Field samples (e.g. of water or sediments) generally contain a wide range of unidentified potential pollutants and their complete individual quantification is unrealistic. Even when chemical identification of the main toxicants included in the ever-growing “black lists” is achieved, this does not allow an inference on actual toxicity, because chemical speciation, complexation, interaction and other physico-chemical processes affect bioavailability. Therefore, biological assays are strongly
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recommended as tools to provide biological criteria for the assessment of pollution effects. The toxicity of bioassayed field samples may be classified according to the arbitrary scale recommended by Woelke (1966) who considers a sample which gives less than 5% abnormal larvae in excess of the controls as non-toxic, 5 to 15% abnormalities as slightly toxic, and over 15% as toxic. Irrespective of the source, contaminants in the oceans may accumulate in four compartments: water surface microlayer, water column, sediments and biota. Accumulation of pollutants within organisms is beyond the scope of this review. 3.3.2.1. Water quality bioassays. For assessing water quality, bulk seawater may be used immediately after collection for incubation of the fertilized eggs, as described by Woelke (1967). He recommended this biological procedure for the monitoring of estuaries, and pointed out that a chemical criterion (the Pearl-Benson index for pulp-mill wastes) was not correlated with toxicity to the test organisms. In order to allow comparisons between experiments performed with different biological material, Woelke (1972) calibrated the experimental data with a control incubated in clean seawater, thereby calculating the net response (see Section 3.2.2.5). This general approach has been followed by Lloyd and Thain (1981), Nelson et al. (1983), Klockner et al. (1985), Nascimento (1989), Thain (1992), Thain and Kirby (1994), and Konar and Stephenson (1995). Konar and Stephenson (1995) did not find significant differences in toxicity between sub-surface and bottom waters. The sea surface is an important concentration area of pollutants from the atmosphere (such as lead), as well as for hydrophobic substances, including crude oil and its derivatives, PAH, and chlorinated hydrocarbons. The surface microlayer therefore contains higher concentrations of metal and organometal contaminants than the sub-surface bulkwater (Hardy and Cleary, 1992), and is usually more toxic to bivalve larvae (McFadzen, 1992). This has motivated the development of methods for sampling the top few micrometres of the sea surface. While these methods are extremely useful for the detection of pollutants which can be below detection limits, the ecological relevance of the derived toxicity data is questionable (Thain, 1992). The use of euryhaline species allows the extension of the bivalve embryogenesis bioassay to fresh and brackish water. Morrison and Petrocelli (1990) conducted a bioassay using Mulinia lateralis at salinities as low as 1Opsu. The species commonly used in Europe, Mytilus galloprovincialis and C. gigas are also suitable for assaying coastal waters of different salinities after salinity is adjusted to 25 psu by adding either 0.2 pm filtered clean oceanic seawater or deionized water to the samples (His and Beiras, 1995; His et al., 1997a).
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3.3.2.2. Sediment quality bioassays. Pollutants are deposited and concentrated in sea-floor sediments by precipitation of dissolved chemicals, and by adsorption to the surface of particles which then sink to the sea floor (either directly, or after passing through the digestive system of an animal). Organic pollutants may persist in sea-floor sediments for long time periods owing to anoxic conditions which reduce the rate of microbial degradation. While bulk sediment analytical chemistry may provide an estimate of the degree of pollution, the fraction of the total that is bioavailable and affects the organisms can only be determined by using biological assays. Sediment characteristics such as particle size distribution, organic content, redox potential and acid volatile sulphides (AVS), affect the quantity of bioavailable pollutants (Anderson et al., 1987; Lamberson et al., 1992; Ankley et al., 1996b). The concentration of metals in the interstitial water, rather than total dry weight metal content of the sediment, is often correlated with toxicity (see Ankley et al., 1996a, and citations therein). Interstitial water can be obtained from the bulk sediment, thereby providing the opportunity to evaluate sediment quality by means of water-column organisms such as bivalve embryos and larvae. Sediment toxicity bioassays are widespread tools in environmental quality assessment (reviewed by Lamberson et al., 1992; Luoma and Ho, 1993), and most international agencies concerned with environmental management are developing marine bioassay methods and will promote their use to a greater extent in the future. These tests, according to Long and Chapman (1985), are one of three steps in a complete evaluation of the aquatic environment: (1) analytical chemistry, (2) toxicity bioassays, and (3) community structure (e.g. diversity indices) and other field studies (see Malueg et al., 1986; Chapman, 1990). The bivalve embryo bioassay has been adapted to deal with sediments by Chapman and Morgan (1983) and it has subsequently been validated for oceanic, coastal and estuarine environmental quality assessment (Williams et al., 1986; Long et al., 1990; US-EPA, 1991; Butler et al., 1992; Chapman et al., 1992; His et al., 1996; Van den Hurk et al., 1997). Carr (1998) found that an embryo-larval liquid phase bioassay is, on average, one order of magnitude more sensitive than the widely used whole-sediment adult amphipod bioassay. However, sediment bioassays have important limitations because of problems associated with the collection, storage and manipulation of the samples, which may affect the final toxicity (Anderson et al., 1987; Lee and Jones, 1987; Samoiloff, 1989; Swartz, 1989; Burton, 1992; Luoma and Ho, 1993; Beiras et al., 1998; Carr, 1998). Standardization of collection and storage procedures is overdue (Butler et al., 1992; Chapman et al., 1992; Lamberson et al., 1992; Beiras and His, 1995b). Different methods (including different test species) may lead to large differences between the
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estimates of toxicity for the same samples (e.g. Long et al., 1996; but see also Van den Hurk et al., 1997), and even contradicting results have been found (Phelps and Warner, 1990; Butler et al., 1992). Variability of the results is markedly reduced when the techniques to obtain the experimental matrix are standardized. Carr et al. (1996) found excellent agreement (10 of 11 cases) among the results for the same sediment pore waters assayed with three different tests (sea urchin embryogenesis, polychaete reproduction and copepod nauplii survival). The optimum method for the collection of samples depends on the purpose of the study. For biological assessment of the present state of pollution, only the surface layer (e.g. the top 2cm) should be collected. For heavy metal and hydrocarbon analyses, wooden sampling equipment is preferred. For more detailed studies on the speciation, sorption or mobilization of a toxicant, the integrity of the sediment's vertical profile should be preserved and Teflon-lined box-cores may be necessary. Further research is needed to determine the optimum preservation methods and the maximum period during which a sediment may be stored without affecting its toxicity. Fresh storage (4°C) should be limited to few days (2 to 7 days according to Anderson et al., 1987; Swartz, 1989; Phelps and Warner, 1990; Burton, 1992; Luoma and Ho, 1993). A significant change, often a decrease in sediment toxicity, may occur during prolonged storage (Phelps and Warner, 1990; Ho and Quinn, 1993; Beiras and His, 1995b; Beiras et al., 1998), presumably resulting from one or more of the following processes: (1) biodegradation of organics (e.g. Ho, cited by Luoma and Ho, 1993), (2) complexation with organic matter of inorganics and coagulation of the complexes (Landrum et al., 1987), (3) oxidation of reduced irodmanganese species to insoluble oxides and co-precipitation (scavenging) of trace metals (Lee and Jones, 1987; Luoma and Ho, 1993), and (4) reaction of metal ions with acid-volatile sulphides and precipitation of the resulting insoluble sulphides (e.g. CdS) (Di Tor0 et al., 1990). An optimum sample preservation method should prevent these processes from taking place. Drying or freezing the sediments is generally not advised because the structure of the sediment matrix is presumed to affect the toxicity. Unpredictable changes in sediment toxicity resulting from freezing have been reported by Schuytema et al. (1989). We have also found spurious toxicity (false positives) in control sediment samples deep-frozen at -200°C and frozen at -2O"C, but in the long term, freezing was found to be preferable to fresh storage (Beiras et al., 1998). More attention has been focused on the manipulation of the sediment prior to exposure to the biological material. Two basic techniques have been developed: exposure to bulk sediment, and incubation in a liquid phase. The liquid phase may consist of the interstitial (pore) water or it can be obtained by mixing the sediment with control water, i.e. elutriation,
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with subsequent decantation, filtration or centrifugation. Extraction of the sediment with an organic solvent has also been performed (Wolfe et al., 1995). Bulk sediments are generally more toxic than their elutriates (Van den Hurk, 1994; Geffard, 1997). Even in bulk sediment bioassays, water must be added and some sort of agitation provided because bivalve embryos and larvae are free-swimming organisms. Therefore these methods will be discussed along with elutriate bioassays. The difference is the presence or absence of the sediment during the incubations. Pore water bioassays. In these tests the liquid matrix to be assayed is obtained directly from the sediment. Interstitial water is separated by filtration under pressure (Carr et al., 1989; Long et al., 1996), or by centrifugation (Ankley and Schubauer-Berigan, 1994; Magnusson et al., 1996). The toxicity may depend on the method chosen (Carr, 1998). These are the “softest” methods to obtain a liquid phase, and therefore those which are supposed to represent the actual toxicity of the sediment in the natural environment with the highest accuracy. Nevertheless, elutriation has been used more frequently. Elutriate and sediment bioassays. In these tests, the assayed liquid matrix consists of unpolluted water which is mixed with the sediment. Several different mixing techniques have been used; rotary shaking (Cardwell et al., 1976; Chapman and Morgan, 1983; Williams et al., 1986; Becker et al., 1990), orbital shaking (Thain, 1992), magnetic stirring (Beiras and His, 1995b; Fichet et al., 1998), mechanical plunging (His et al., 1996), mechanical rolling (Butler et al., 1992; Van den Hurk, 1994; Van den Hurk et al., 1997), even hand shaking (Long et al., 1990). Daniels et al. (1989) claim rotary mixing produces the most consistent results. The proportion of sediment to water may vary by more than two orders of magnitude, between 2 and about 500 g 1-’ (sediment weight to total volume). The duration of mixing should be sufficient to saturate the liquid phase with the mobilizable fraction of the toxicants, and this is apparently achieved in as little as 3 minutes of rotation in stoppered bottles at 4rpm (Cardwell et al., 1976), or, according to Van den Hurk (1994), even just hand shaking for a few seconds. Van den Hurk (1994) did not find a significant effect of duration of mixing on the toxicity of the elutriate. Fertilized eggs can be incubated with the bulk sediment providing continuous agitation (e.g. Cardwell et al., 1976), but in this case deleterious effects resulting merely from physical abrasion cannot be ignored. Turbulence and turbidity generally reduce survival of bivalve larvae by 1 to 5 % , depending on the method of agitation (Davis, 1960; Davis and Hidu, 1969b; Seaman et al., 1991). The effects of turbulence and turbidity on larval growth rates are more complex, and depend on biological species, sediment type, and incubation method. At concentrations up to 1g 1-’, the growth rates may be increased or reduced by as
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much as 20% on average, but the variability is very high and may attain between 5% and 260% of the growth rate in controls (Davis, 1960, Davis and Hidu, 1969b; Seaman et al., 1991). At sediment concentrations of 2 to 5 g 1-' average growth rates may be comparable to those of the controls (Seaman et al., 1991), or they may be 40 to 60% lower (Davis, 1960; Davis and Hidu, 1969b). To assess and compare the degree of pollution in different sediments directly from toxicity data, the general geo-physico-chemical characteristics (such as particle size and organic matter content) in the sediments to be compared should be similar. These general characteristics affect the quantity and bioavailability of pollutants bound in the sediment matrix (Anderson et al., 1987; Lamberson et al., 1992). Without such similarity, inter-site toxicity comparisons have very limited value. Fine-silt sediments have a high surface-to-volume ratio, thus facilitating the sorption of a wide range of micropollutants, and their usually high organic content further facilitates the sorption of hydrophobic pollutants (DeGroot, 1973; Adams, 1987; Lee and Jones, 1987; Lamberson et ai., 1992). Long et al. (1990) found that clay and total organic carbon content of the sediments are more strongly correlated with sediment toxicity than heavy metal or hydrocarbon concentrations. Furthermore, it is desirable to record general water quality parameters, such as temperature, salinity, pH and dissolved oxygen in the liquid phase assayed, to guarantee suitable values for larval development. The optimum conditions of temperature, salinity, oxygen and pH for larval rearing in the laboratory have been discussed above (Section 2.3.4). Bivalve larvae are quite resistant to hypoxic conditions. There is no increase in mortality with decreases in oxygen concentration between 3 and 16% saturation (Huntington and Miller, 1989; Wang and Widdows, 1991; Bittkau, 1993). However, embryos are more sensitive to hypoxia than larvae (Bittkau, 1993). Highly hypoxic or nearly anoxic conditions do cause larval mortality, particularly in younger larvae. At a dissolved oxygen concentration of 0.2% saturation, Wang and Widdows (1991) found that survival decreases to 16%. We have found that such extremely low levels of dissolved oxygen may occur in elutriates from reduced sediments with a high organic content, despite thorough stirring during elutriation (R. Beiras, unpublished data). Other potential sources of false-negatives are ammonia and, in particular, sulphide, especially when highly reduced sediments and/or anoxic elutriation conditions are used. Reported no-observed-effect concentrations for ammonia and hydrogen sulphide are 4.7 and 0.1mg1-' respectively (SAIC, cited by Van den Hurk, 1994), while Cardwell and coworkers (1979b) reported ECS0of 15 and <0.01 mg 1-l respectively, and Bittkau (1993) working with 15- and 30-hour-old mussel
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embryos found 50% abnormal development at c. 0.3 mg H2S1-l. Knezovich et al. (1996) found the 48-hour LCso for mussel embryos to be between 29 and 205pg total sulphide per litre, and they expected hydrogen sulphide to be approximately 10% of total sulphide at the pH of the test solutions. Sulphides showed the strongest correlation with toxicity among a number of physico-chemical variables (including heavy metals and oils) in the experimental samples, with sulphide concentration accounting for 81% of the variation in larval mortality (Cardwell et al., 1976). Matthiessen et al. (1998) found toxic effects on C. gigas embryos in elutriates from sediment collected upstream of industrial effluent inputs, and they estimated H2S or NH3 toxicity as a source of false positives in evaluation programmes with elutriate bioassays (see also Sections 4.2 and 4.3). 3.4. Statistical methods
3.4.1. Dose-response curves The relationship between the dose of a potential toxicant and the biological response of the test organism is generally described by doseresponse curves. Preferred mathematical methods to describe this relationship are regression models, in which dose is the independent variable and the biological response is the dependent variable. When we are concerned with lethal toxicity, the response is the proportion of dead individuals (p), a binary discontinuous variable with a binomial distribution. In the case of sub-lethal toxicity, the response is often a normally distributed continuous variable such as growth, although in the context of the present review discontinuous binomially distributed responses are common. Examples include proportion of embryos developing into normal larvae or proportion of metamorphosed larvae. In both cases, a linear model is generally desirable to simplify curve fitting, model fit checking and prediction of the concentration causing a given biological response (Forbes, 1993). For proportion data, linearization of the dose-response curve is achieved by plotting concentrations on a log scale and transforming response values (p) into arcsine pin, normal equivalent deviates ( Z ) , probits (2 5 ) or logits (ln[pl(l - p ) ] ) . Transformed data can be fitted to a straight line, for example,
+
arcsine pl"
=a
+ b log C
(4)
where C is the concentration of toxicant. As mentioned above, all 0% and 100% responses are generally discarded. It may, however, prove impossible to find a mathematical formula that will yield a linear relationship.
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Once the data have been fitted to a linear regression model, effective concentrations (EC, - values corresponding to an X% reduction in the biological response), or lethal concentrations (LC,), can be calculated by interpolation. LCsois the most commonly used summary statistic, because according to the laws of regression its confidence interval (CI) is narrower than that for any other value of X.The LCso95% confidence intervals can be calculated according to the Litchfield-Wilcoxon method (Newman, 1995) as: Upper 95% CI limit = LCsof (5) Lower 95% CI limit = LCS0.f-' where
f=
LC84 I LC5012,77'" LC50 LC16
and N' is the total number of results between the 16% and the 84% predicted responses. The Litchfield-Wilcoxon method requires partial responses (between 16% and 84%). When such responses are not available the SpearmanKarber method can be applied (Finney, 1964). This procedure, whose advantages have been pointed out by Hamilton et al. (1977), is nonparametric (model-free) and only requires symmetry of the theoretical dose-response curve. This last method, though, tends to underestimate the LCso when all experimental concentrations cause mortalities higher than the control. Guidance for choice between these and other statistical methods is provided by Gelber et al. (1985). Despite efforts to standardize methods, biological material still carries a considerable amount of variability. Differences among stocks of adults and differences in gamete quality are well-known sources of undesired variability in bivalve larval bioassays, and unexposed control replicates are indispensable in the experimental design. In order to overcome this difficulty, the use of a positive control exposed to a reference toxicant such as sodium chloride, sodium dodecyl sulphate (SDS) or dissolved heavy metals, has also been suggested (e.g. Krassoi et al., 1996, 1997). Cardwell and co-workers discussed this subject in the context of bivalve larvae (Cardwell et al., 1977b) and advocate the use of a natural substance such as sodium chloride (Cardwell et al., 1979a). A reference toxicant should be easily available and measurable, highly soluble in water, chemically stable, and should elicit a universal response. Mercury chloride has all of the former properties except for ease of analysis, and, in contrast with sodium chloride, it can be used with euryhaline estuarine animals (see also Beiras and His, 1994, 1995a).
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Comparisons between experiments are only permissible after correcting the percentages of biological responses with the blanks (without toxicant) using the following equation (Emmens, 1948; Newman, 1995, p. 136, Abbot’s formula): P = (Po - P,)*100/(100 - Pc)
where P is the corrected response (%), Po is the non-corrected original response (%) and P, is the control response (%). This expression has been used to compare responses from experimental and control samples in seawater quality assessment bioassays (see also equation 2, Section 3.2.2.5). 3.4.2. Qualitative assessment of environmental samples
When environmental samples of unknown composition are tested (either water or sediment liquid phases) we can either attempt a quantitative assessment as explained above or we can obtain a qualitative “yes or no” criterion of toxicity, this criterion being a significant reduction in the biological response (e.g. 24-hour embryogenesis success) compared to a non-toxic control. First approaches were based on arbitrary scales such as that by Woelke (1966) (described in Section 3.3.2) who considered as toxic all samples which increased control mortality by more than 15%. We can proceed with a more consistent approach by means of a statistical comparison between a control of water (or sediment liquid phase) from well-known unpolluted sources and the samples whose biological quality we wish to assess. Statistical analysis should then include the following steps: 1. Angular transformation when the biological response is measured in percentages (this is usually the case). 2. Checking the distribution for normality (e.g. chi-square test) and homogeneity of variance (e.g. Bartlett’s test). 3. If the distribution is normal and the variances are homogeneous, ANOVA test for significance of the differences in biological response and a posteriori multiple range tests for identification of problem samples significantly different to the controls (often Dunnett’s test). 4. In case assumptions for ANOVA (2) are clearly unwarranted, nonparametric rank tests (e.g. Wilcoxon) should be applied. When this procedure is used in toxicity tests with known concentrations of single toxicants, it is usually referred to as the NOEC (no observed effect concentration)/ANOVA approach. Strong criticism of the
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NOEC/ANOVA approach arises from the fact that NOEC must be one of the experimental treatments, thus depending to a large extent on the experimenter’s choice of concentrations. Therefore, when toxicity of isolated identified compounds is studied, the dose-response approach and ECx calculation described above are preferred (OECD, 1998). When comprehensive analytical chemistry data for the main pollutants of concern at the different sampling sites are available, identification of the chemicals causing the observed toxicity may be attempted. One possible procedure is to identify toxicants responsible for the observed toxicity by non-parametric Spearman correlation between chemical (concentration) and biological (toxicity) data. Non-parametric rank correlation is preferred because the amount of pollutant is not expected to be normally distributed along the sampling sites. Rather a skewed distribution with a few extremely polluted sites is often the case. Another more ambitious approach would use step-wise multiple linear regression aiming to describe toxicity as a function of the relevant toxicant concentrations weighed by the corresponding coefficients. However, nonlinear interactions between different toxicants (among other causes) prevent these deterministic models from having any predictive value. 3.4.3. Minimum sample size When designing a biological assay with bivalve larvae, two questions arise regarding sample size: how many larvae should be observed in each experimental vessel, and how many replicates (vessels) should be set up for each experimental treatment. To answer these questions adequately we need a preliminary estimate of the variability of the biological response we are to record, both within replicates and among replicates. Let us take first the question of number of observations per vessel. In the case of a variable with a theoretical normal distribution, such as larval size, we can use the following iterative procedure (Zar, 1984). Given an estimation S2 of the population variance obtained from n data, the number of observations, m,needed to estimate the parametric mean with an error s d is:
where 1- a is the confidence level and 1- p is the power of the test. When consulting the statistical tables the t value should be obtained from the two-tailed column while the F corresponds to single-tailed values. In the case of discontinuous variables with only two possible values (normal D-larvae versus other forms, or alive versus dead individuals), we take from a population of N individuals a sample of size n, and record the
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number of individuals, c, with a certain characteristic (e.g. normal D-larvae). The proportion p ’ = c/n is an estimate of the parametric proportion, p , of individuals in the whole population with that characteristic. Burstein (1971) provides us with tables and explanations appropriate for knowing the sample size (n) necessary to estimate p with a certain error margin at a given confidence level. The tables are based on a Poisson approximation when p ‘ 0.25 and a normal approximation when 0.25 < p ‘ s 0.5. (When p ‘ > 0.5, simply find sample size for q‘ = 1- p ‘ . ) This procedure assumes sampling without replacement from an infinite population. In the particular case when sample size is a non-negligible fraction of the total population (>5% N), then n is overestimated, and a correction for finite populations can be applied. The corrected sample size (nF)is in that case (Burstein, 1971):
==
nF= nN/(n+ N - 1)
(10)
For example, given a 25 ml vessel with 20 embryos per ml ( N = 500), the minimum sample size ( n ) necessary to estimate the proportion of normal D-larvae (p) with an error of 0.05 at 95% confidence level, expecting p values of 1, 0.90, 0.75, 0.5, 0.25, 0.10 and 0, would be respectively n = 58, n = 200, n = 334, n = 400, n = 334, n = 200, and n = 58. After correcting for finite populations (since n > 5%N in all cases), then nF = 52, nF = 143, nF = 200, nF= 222, nF= 200, nF= 143, nF = 52. Therefore a sample size of 200 usually allows 95% confidence in our estimate with a low (0.05) error. Moreover, in embryo-larval bioassays, responses often follow a “yes or no” pattern; i.e., extreme values demanding lower sample sizes are more common than proportions close to 0.5. Concerning the question of number of replicates per treatment, we are interested now in the minimum sample size required to detect differences between two population means. Each population consists of the mean for each vessel; therefore they can be assumed to be normally distributed, irrespective of the recorded response (either percentage of mortality or larval size). Again, given an estimation, S2, of the population variance needed obtained from n data, the number of replicates per treatment (m), to estimate the parametric mean with an error s d is (Sokal and Rohlf, 1969):
where 1- a is the confidence level and P is the power of the test. When consulting the statistical tables, the t values should be obtained from the two-tailed column. Applying the methods just described, and aiming for P = 90% and
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a = 0.05, we recommend an experimental design for the bivalve embryolarval bioassays with five replicates per treatment. When replicates are 25 ml accuvettes, 200 observations per vessel to record percentages, and 25 observations per vessel to record normally distributed variables such as size are recommended. This enables estimation of the responses with errors as low as 5% in percentages and 7.5 pm in sizes.
4. TESTING THE TOXICITY OF MARINE POLLUTANTS TO BIVALVE EMBRYOS AND LARVAE 4.1. Pollutants 4.1.lMetals .
Tables 9 and 10 show that the most sensitive biological response to the most toxic heavy metals, such as mercury, silver and copper, is the inhibition of normal embryogenesis, followed by growth reduction. Sub-lethal responses are generally more sensitive than larval mortality. For instance, in the case of copper, the metal most frequently studied, the average values for EC5* are: 39.8 pg 1-' (sd 51.0, n = 26) for embryogenesis inhibition, 63.6 pg 1-l (sd 60.0, n = 17) for growth reduction, and 86.5 pg 1-l (sd 76.1, n = 16) for larval mortality. The heterogeneity of the data, however, prevents these differences from reaching stathical significance. Pediveliger settlement as a criterion of toxicity appears similar in sensitivity to embryogenesis, but data are scarce. According to Watling (1983) about 40 pg Cu 1-l reduces settlement of Crassostrea by half, and Chang et al. (1996) estimate that the value is 5 to 15 pg 1-1 (but see also Phelps and Mihursky, 1986). Other responses, such as respiration and genotoxicity, require a considerable experimental effort and are not suited for routine studies. 4.1.1.1. Embryogenesis. Table 9 summarizes the data on metal concentrations which cause 50% inhibition of normal embryogenesis (ECS0). Despite important variation in test procedures among the different studies, two conclusions can be drawn from a detailed inspection of the table: (1) there is a remarkable consistency in the ranking of the toxicity of the different metals, and (2) the species most frequently used in the bioassays display a similar sensitivity to the various pollutants. For instance, the EC50of mercury to C. gigas is 10.4 pg 1F' (26.7, n = 5) and to Mytilus spp. it is 10.1 (k7.8, n = 4). The average ECS0in pg of metal per litre are (in order from most toxic to least toxic) the following. Mercury, 10 (55.7 sd, n = 14); silver, 19 (29.3, n = 11); copper, 39 (k50, n = 27); barium, 189 (n = 1); zinc, 320 (5497,
Table 9 Toxicity of heavy metals to bivalve embryos. ECS0:toxicant concentration causing 50% abnormal embryogenesis. Chlorides were used for Hg, Cu, Zn, Ni, Cd,nitrates for Ag, Pb, oxide for As and chromate for Cr, except when otherwise stated. Test species
Initial stage (time after fertilization)
Exposure conditions (time, temperature, salinity, density, food, seawater)
EC.
(pg metal ion I-')
HE
Ae
Cu
Zn
Ni
Pb
Cd
Reference and notes As
Cr
Mn ~~
C. gigar
fertilized egg
27°C
C. gigas
fertilized egg
C. gigas
fertilized egg
C. gigas (and 3 other SPP.)
fertilized egg
48 h, 2&2TC, 5 pm FSW uv sterilized 48 h, 2O"C, Mpsu. 24-28ml-l, 5.7 (5.5*) 1 pm FSW uv sterilized 48 h, 20% %25psu, 35 d-', FSW uv sterilized
C. gigas
ferrilized egg
48 b, 20°C 33 psu, 2 M 6 I&', 1 pm FSW 48 h, 20°C. 3Opsu. H)ml-*, low FSW uv sterilized 48 b. 20°C. 24-28 I&'. 1um FSW uv sterilized ' 48h, 25'C, 3635psu. 5 p m FSW
c.g1glLF
fertilized egg
C. gigas
fertilized egg
C. gigas
fertilized egg (15 min)
C margariracea C. cucullafa C. gigas
egg
C gigas C. gigas
egg
C. gigas
fertilized egg
C. gigas (and 5 other
fertilized egg
32-100 1 O C G 3200 -250
1&32
>2500,920* 13900. 1290' 370,60* 1700.590* 1618
C. virginica
fertilized egg (15 mi.) fertilized egg
C. virginica
fertilized egg
(1h) C virginica
fertilized egg (2 h)
Coglianese and Martin, 1981; Cu nitrate; abnormal larvae excluded Knezovich ef a/., 1981: *humic matter 2mgI-', **EDTA 1 0 - 6 ~ .
10 12, 40,* 100"
6.7
48 h. 26°C 28 psu, 30 d-', 0.8 pm FSW 24, 48 h, 24% 32psu, 20 n P , 0.2 um FSW 48 h, b C %36 d-', 33psu 1 pm FSW uv sterilized 23psu 24 h, 24T, 20, 25,3Opsu, 0.2 pm FSW 24 to 72 h, methods not given
22
180*
>loo >loo >loo >loo
>loo
Martin ef al.. 1981; Cu,Zn and Ni sulphates; abnormal larvae excluded Wathng, 1981; *calculated by
160* 130* 13
1100 >loo >1w >loo >loo >loo >loo >loo
>1W >100
His and Robert, 1981; abnormal larvae
5.3
119
349
758
611
326
4538
excluded
larvae excluded Coghanese, 1982; Ag nitrate; abnormal larvae excluded
218 5-6.5 W13.5 5 4 . 5
>so 44
1&100
24 b, 24°C 24 d-', 0.2 pm FSW
13
42-48 h, 26°C 15-17 ml-', ASW 44.48 h, 26psu. 20-22d-', 1 pm FSW uv sterilized 48h, 20. 25, 30°C 22.5, 27.5psu, 1 pm FSW
5.6
5.8
103
310
20°C 11.4
24.2 35.3 32.2
15.1 18.7 18.3
206 325 230
25'C 12.6 30°C 10.2
1&20
extrapolation.
His and Robert, 1982; sulphate; abnormal
10
SPP.)
C. gigas
Okubo and Okubo. 1%2; Cu and Zn sulphates Brereton d al., 1973: most larvae abnormal from Zn 150. Glickstem, 1978; abnormal larvae excluded Se decreases Hg toxicity; *nitrate. Cardwell ef ul., 1979a; all sulphate; 'fully shelled regardless of shape.
5w1000
1180 2450 3800 7500* 10300
Robert and His, 1985; abnormal larvae excluded Zhadan ef al., 1992; approximate ranges; sulphates; abnormal larvae excluded Beuas and His,1994; abnormal larvae excluded 16000 Calabrese ef al., 1973; *arsenite MacInnes and Calabrese. 1978; abnormal larvae excluded MacInnes and Catabrese. 1979; 220 at 2% 27.5 psu; abnormal larvae excluded
48h, 25psu. 60ml-, FSW sterilized
C. iredalei
fertilized egg
M. edulis
fertilized egg
1?-17"C
M. edulis
fertilized egg
M. edulrr
M. galloprovincialis
fertilized egg (2 h) fertilized egg (0.5-2 h) fertilized egg
48 b, 17°C. 2 4 2 8 ml-l,1 pm FSW uv sterilized 48 h, 19°C. 26psu, 151&', 5 pm FSW uv sterilized 2 d. W C , 35 psu, 30 I&'
M. galloprovincialis
fertilized egg
M.galloprovincialk
fertilized egg
M. mercenaria
fertilized egg
M. galloprovmcialis
S.soltdissimo
I. califomicum
fertllued egg (1h) fertilized egg (2 h) fertilized egg (4 h) fertilized egg (15min) sperm + egg (1 h later)
I. califomicum
fertilized egg
M. lateralis
S. plano I. californicum
(M Chlamys asperrima Chlamys aspemma
fertilized egg (>I h) fertilized egg (30 min)
10-32 5.8
14
32-100 3.2-10* 5.8
24
3.5
891
476
13'7
us*
4426 4743**
145
4055
Hrs-Brenko el al., 1977; abnormal larvae excluded Pavicic. 1980: Z n sulohate. *60h at 16°C; "60h at 1 6 k abnormal larvae excluded Pavicic er al... 1994a:. abnormal larvae excluded Beiras and Hi$ 1995a; abnormal larvae excluded Calabrese and Nelson, 1974
10 4.8
21
166
310
780
Eyster and Morse, 19W, abnormal larvae excluded
6.4-9.5 c. 16
48 h, 21°C. 10, 3Opsu, 75 I&'
18,17
48 h, ZOS'C, 24psu. 25 I&', 0.45 pm FSW 48 h, 2 4 T, 34psu (24psu), 2 ml-', 0.22 wm FSW 2 h. temnerature not eiven. 34psd 200 I&', 0.8 FSW 48 h, temperature not given, 0.45 pm 34psu, 17 d-', FSW 48 h, 18°C 33psu, 30ml-', 1pm FSW uv sterilized 48h, 18'C, 33psu, U)ml-', 5 sm FSW uv sterilized
10-20
&
320loo0 175
250-500
48 h, 20"C, 37-38psu 48 h, 20°C 37-38 psu, 3 W m1-l. marselv FSW 48 h, 20°C, ml-', 0:2 pm FSW 42-48 h, 26"C, 15-17 m1-I. ASW 48 h, 20"C, 3Opsu, 30 ml-l, 0.22 pm FSW uv sterilized
Ramachandran er al., 1997; Cu sulphate; abnormal larvae excluded 3 2 W >loo00 Okubo and Okuho. 1962; Cu,Zn and too00 Cr sulphate; *acetate. 1200 >3000 4469 Martin el al., 1981; Cu,Zn and Ni sulphate; abnormal larvae excluded 30000 Morgan er al., 1986. 459
81
Momson and Petrocelli, 1990. sulphate. Ruiz el 111.. 1995~:nitrate; abnormal larvae excluded, control embryogenesis success <4% Ringwood, 1990; results for 24 psu salinity in brackets
55
Ringwood, 1992b; endpoint: cleavage. Ringwood, 1992b; abnormal larvae excluded
7
48*
39'
266
I26
4.5.
45
295
I350
Krassoi et al., 1996; Cu and Zn sulphate; abnormal larvae excluded Krassoi er al., 1997, Cu sulphate: abnormal larvae excluded
Bold 95% CI provided. ASW artificial seawater; FSW filtered seawater. Other metals: Al C.virginica 42-48 h EC50>7500 (Calabrese er 01.. 1973: chloride); M.galloprovincialis 48 h E C , 54-162 (Pagano ef al.. 1996, sulphate): Saccosrrea commercialis 48 h E C , 200 (Wilson and Hyne, 1997; sulphate). E a Mytilus colifominnus 48 h E G O= 189 (abnormal larvae as judged by polarization o p t i q Spangenberg and Cherr, 1996). Sr Mytilus californianus 48 h ECso >50o00 (abnormal larvae as judged by polarization o p t i q Spangenberg and Cherr, 1996). Se C. gigas and M. edulis 48 h ECS, >loo00 (Martin er al., 1981); C. gigar 48 h EC, >loo00 for SeOl and Na2Se03(Glickstein. 1978). Fe M. edulis E C , 1OMxI-32000 (Okubo and Okubo, 1962). M o M. edulis 48 h E C , = 147000 (Morgan er al., 1986).
Table 10 Effects of heavy metals on mortality, growth and metamorphosis of bivalve larvae. LC50:toxicant concentration causing 50% mortality; otherwise EC50: toxicant concentration causing 50% reduction in the end-point. Nitrate for Ag and chlorides for the other metals were used except when otherwise stated. No attempt has been made to calculate accurate LCsOvalues from original data, because of variability between trials within an experiment and, particularly, differences between nominal and actual concentrations. Rather, concentrations above andlor below 50% effect are provided. LC, or EC, fw metal ion 1.')
Exposure conditions hitial agekize
Test species
C. gigas C gigas C gigas
21 d, 250-320 pm 5d,88pm 16 d, 260 pm 6d,135~ 16 d, 309 pm
(time, temperature, salinity,
density, food, seawater) 5 d, 21°C 1ID-', mix algal food. 2-5 pm FSW uv
sterilized 4 d. U25"C 2.5 ml-l (young) 1.5 ID-' (old), mix algal food, 5 pm FSW 4 d, 22-23'C 2.5 ml" (young) 1.5 ID-' (old), mix algal food, 5 pm FSW
End-point
Hg
Ag
Cu
Zn
50
80
>loo
>loo
>lW
85 1100 75 >lo0
mortality
>lW >loo
C cucullata
3d,75pm 13d, 254 pm 24 h
60 >1M) 2447
>loo
c.gigas
1d,Mpm 211 pm 3 1 0 ~
C. virginica
2d
C. virginica
2d
C virginicn
48 h, 70 mn
0. edulis
1-3 d
12 d. 25°C. 10-12 I&', mix algal food, 1pm FSW 48h, 17.5.22.5,27.Jpsu, 20 d-'mix , algal food, 1pmFSW 8 d. 26°C. 27-28 psu, 20 I&', monoalgal food, 1 pm carbon FSW uv sterilized 4.2 h, 15°C0.8 ID-',no food
Cr
Referenceandnotes
Watling, 1978
200
60 >loo
monoalgal food. 0.8pm FSW 48 h, 24°C 24 I&', no food, 0.2 pm FSW
Cd
Boyden er d., 1975; sulpbate
mortality
3d,80pm 13d, 239 pm
5 d, 26°C. 28psu. 8 DIP,
Pb
>500
mortality
C margaritam
C gigas
Ni
mortality mortality (tissue degeneration)
m
mortality
12.0
Watling, 1982
80
>loo
>lo0
His and Robert, 1981 1994 & Beiras and €I
33
115 25.0
32.8
1200
Calabrese el a/., 1977a
>90 60-90
mortality
Macnes and Calabrese, 1979: *3060 at 1 7 . 5 salinity ~ ~ ~
&!XI*
mortality
mortality (no response touching with needle)
>86
3.3
227
Wikfors and Ukeles 1982
Connor, 1972
M. edulis
3d
15 d, 15% 32psu. 5 ID-'.mix algal food, 0.2 pm FSW
M. edulis
150 pm
10d, 1SoCc, 32psu, l o r n - ' , monoalgal food
M.galloprovincialis 2 d. 76 um 141 pm 225 pm 258 p m
48 h, 20"C, 24 m1-l. no food, 0.2 rn FSW
mortality (tissue degeneration) mortality
M. rnercenaria
2d
b 1 0 d. 25°C. 10-12 ID-',mix algal food, 1 pm FSW
I. californicurn
3d 10 d 24 d 36 d
48 h. 24'C. 34psu ( 2 4 ~ s ~ ) . mortality 1ID-],no fbod,'O.ZZ pm FSW
P mnxirnus
3d
C gigas
2 d, 50-70 pm
C. gigas
5d,88pm 16 d, 260 pm
15 d, 15°C. 32 psu. 5 ml-'. mix mortality (inactive algal food, 0.2 pm FSW oesophageal cilia) growth (length) 5 d + 5 d recovery, 26"C, 5 ID-',mix algal food, 5 pro FSW uv sterilized 7 d , 2 5 T , 2.5 d-' growth (length) (young), 5 d, 23°C 1.5 ID-' (old), mix algal food, 5 pm FSW growth (length) 4 d, 22-23", 2.5 ID-' (young) 1.5ml-l (old), mix algal food, 5 pm FSW
C. gigus
6 d, 135 pm
C cucullnra
51 164 322 383 14.7 32.4
Stromgren and Nielsen. 1991; sulphate; silicon-coated culture jars Beiras and His,1995a
16.4
195
Calahrese e t a / . , 1977a
5700
Ringwood, 1990.results at 24psu salinity in brackets
100
Beaumont er 01.. 1987 Brereton er a/., 1973; sulphate
-150
>1M)
75 120
3d,@Jw 13d, 239 pm
35 85
45
40
85
1W
3 d, 75 pm 13 d, 254 pm
40 85
50
45 120
egg 24 h
C. gigas
egg 24 h fertilized egg
7 d, 26°C 28psu. 6 ml-', growth (height) monoalgal food, 0.8 pm FSW mix growth (height) 12 d, 24% 32psu. 8 d-', algal food, 0.2 pm FSW 10d, 24°C 20,25,30psu, growth (height) 8 mix algal food, 0.2 pm FSW
I&',
Watling, 1978
1W7.00 80
C. gigas
C gigas
10
50 75
16d,309~ C margariracee
Beaumont er al., 1981
2W500
mortality (inactive oesophageal cilia) mortality
95
85
Watling, 1982
-12 12-24
His and Robert, 1981
613 -64
His and Robert, 1982: sulphate Robert and His, 1985
250
~~
~
Exposure wnditions (time, temperature, salinity, density, food, seawater)
Initial age/size
Test species
LC, or ECx (pg metal ion I-') End-point
Hg
c. gigar
ld,60&m
10 d, 24°C 8 I&', mix algal food 0.2 pm FSW
growth (height)
7.1
C virginica
2d
growth (length)
11.8
C. virginica
48 h, 70 ~ L I J
, 12 d, 25°C 10-12 d-'mix algal food, 1f l FSW 8 d, 26'C. 27-28psu. 20 ml-', monoalgal food, 1 pm carbon FSW uv sterilized
growth (length)
0.edulis
179,181pm
%h
3d
M. edulis
150 m
15 d, 15% 32psu. 5 d-l,mix growth (height) algal food 0.2 ~ L IFSW J 10d, 15°C32psu. lOml-', growth (length) monoalgal food
M.mercenaria
2d
b 1 0 d, 25°C. 10-12 d-'mix , algal food, 1pm FSW
growth (length)
21.4
fl maximur
3d
15 d. 15°C 3 2 ~ s5~d.-'. mix growth (height) algal food, 0.2 pm FSW
1. californicum
3d.85~1~
28 d, 24°C 34psu. l O d - ' , mix algal food, 0.45 pm FSW 4 d. temperature not given, 34 psu. 10 d-', mix algal food, 0.45 pm F'SW
growth (dry weight) (tissue dry weight)
72 h. 2C-22pC. 34psu. Mml-', 1~ L I FSW J
Ca uptake
5 d, 21°C. 1ml-', mix algal
settlement
21 d, 250-320 eyed
~LIJ,
food. 2-5 sterilized
~ L I FSW J uv
Cd
Cr
Referenceandnotes
30.2
47.1
Calabrese ei al., 1977a
1078 >27
Wikfors and Ukeles 1982
Stromgreu and Nielsen, 1991: sulphate; siliwnsoated culture jan 7.4
C. gigas
Pb
Walne. 1970
growth (height)
9d, 118pm
Ni
Beaumont et al., 1981
10d, 20°C. 8 d-'mix . algal food, 0.2 pm FSW
M. laiernlir
Zn
>86
M. galloprovincialis 2d,75fim
3d
Cu
Beiras and His 1994
growth (length)
M. edulis
I. californicum
Ag
Beiras and His 1995a 42.2
16.9
232
<110 0
Calabrese et a[.. 1977a
20-100
Beaumont el al., 1987 >20
Ringwood, 1992a
-20 -100
>20
growth
Ringwood, 1992b
(dry weight) 26.5
18.5
176 <125* >500'*
Ho and Zuhkoff, 1982 Boyden et al., 1975; sulphate; ' 5 d continuous exposure; **5 d exposure C5 d rewvery in clean FSW; at Z n 125 and 250 no marked reduction in settler number but 2 d delayed settlement
C. gigas
16 d
C. gigas
19 d
c. gigas
2290 pm, eyed
C. gigas
2280. eyed
C. virginica
2230, eyed
C virginica
eyed
6 d exposure f 3 d recovery, 22-u"C 1.5 ml-'. mix algal food, 5 pm FSW 20 d
3445
settlement*
1&20*
idem
4 d, 24T, 11 n f ' , no food, 0.2 pm FSW, eplnephrine lO-'mol. added d 2 4 d, 37 psu, ASW, epinephrine mol. added at different tunes
metamorphosis* metamorphosis
c. 1wo
c. loo0
4 d, 25 psu, ASW, mol. epinephrine added at different times 20 I&', flowing 0.5 p m FSW
metamorphosis
c. 500
c. 5M)
settlement (4 h) +spat mortality (4-7d)*
534 ppm**
ASW, artificial seawater; FSW, filtered seawater.
2&25
3c-35
3244
1&20 1&20*
1c-20
Watling, 1983; *fixation to bard substrate 1&20
Watling, 1983 (only approximate figures given); '1 d delayed settlement Beiras and Hi$1994 *adulr shell formation G a n g ef ul., 1996: nominal concentrations; estimated actual concentrations 2 orders of magnitude lower for cu
Phelps and Mihursky, 1986: *fan-shaped non-empty shells; **Cuconcentration in the setting surface micro-organism laver; nitrate
94
E. HIS, R. BEIRAS AND M. N. L. SEAMAN
n = 16); nickel, 683 (2425, n = 4); lead, 968 (2847, n = 5); cadmium, 2219 (23312, n = 18); arsenic 3913 (25073, n = 2); chromium, 4564 (23663, n = 6); and manganese, 23 000 (*9899, n = 2). In the case of copper toxicity the estimation of the EC5,, is much lower and more precise if the EC5o values extrapolated from Watling's (1981) data are considered as outliers and disregarded: 24 2 28 (n = 24). According to their embryotoxicity, metals can be classified into three groups. Dissolved mercury, silver and copper are toxic at the 10ppb (i.e. lo-*) level, zinc, nickel and lead are usually toxic at levels from 100 to 1000ppb to and cadmium, arsenic and chromium often show embryotoxicity to level only. Other bivalves at concentrations above the 1 ppm elements occasionally tested and found virtually non-toxic are selenium (EC50 > 10ppm), iron (ECSO> 10 ppm), strontium (ECS0> 50 ppm), and molybdenum (EC50 > 50 ppm) (Okubo and Okubo, 1962; Glickstein, 1978; Martin et al., 1981; Morgan et al., 1986; Spangenberg and Cherr, 1996). Data on aluminium toxicity (not included in Table 9) to bivalve embryos are scarce and contradictory. Calabrese et al. (1973) found no effects on embryogenesis at concentrations up to 7500 pg A1 1-', while recently Pagano et al. (1996) found toxic effects between 54 and 162 pg 1-', and Wilson and Hyne (1997) report ECSovalues of about 200 pg 1-l. In all three cases pH was within the limits of tolerance for bivalve embryos and larvae (27, Calabrese and Davis, 1966,Wilson and Hyne, 1997). However, Calabrese et al. (1973) used aluminium chloride, whereas Pagano et al. (1996) and Wilson and Hyne (1997) used aluminium sulphate; the differences in toxicity may therefore be a result of differences in speciation (see Section 4.3) of the salts in seawater. 4.1.1.2. Larval mortality and growth. The lethal and effective concentrations shown in Table 10 are even more difficult to compare than those for embryos (see Table 9) because of two additional variables, age of the larvae and duration of the experiment. As discussed below, larval tolerance increases with size in terms of survival (Watling, 1978;Beiras and His, 1994, 1995a) and growth (Watling, 1982). Also, detectable effective concentrations impairing larval growth depend on the duration of the experiment, and in the studies reviewed here this ranged from 4 to 28 days. In a number of cases, larval growth is a more sensitive response to toxicants than inhibition of embryogenesis (e.g. Beiras and His, 1994, 1995a). Cadmium, for instance, has been shown to inhibit larval growth at significantly lower concentrations than those which inhibit embryogenesis (compare Tables 9 and 10). Irrespective of its sensitivity, however, a bioassay using growth as the biological endpoint is neither simple nor quick thus, it is not suited for routine screening tests. Nevertheless, effective pollutant concentrations at which larval growth is significantly reduced are still worth studying because of their ecological relevance.
THE ASSESSMENT OF MARINE POLLUTION
95
Ho and Zubkoff (1982) studied the effective concentrations of some heavy metals which reduce calcium uptake in bivalve larvae, and found these concentrations to be similar to those affecting growth (see Table 10). Therefore, calcification of the organic matrix of the shell may be one of the processes impaired by heavy metals and can therefore be used as a measure of sub-lethal toxicity to bivalve larvae. 4.1.1.3. Settlement. Data on the effects of heavy metals on settlement and metamorphosis in competent pediveligers are scarce (Table lo), but they do indicate that the sensitivity of this ecologically relevant stage is similar to that of embryos and larvae. Bioassays with settlement as the endpoint might be particularly relevant for testing the toxicity of surfaceassociated toxicants, such as those that adsorb readily to sediment particles. A shortcoming for the development of settlement bioassays is the limited knowledge of the physico-chemical factors that naturally promote attachment to the substratum and morphogenic change in bivalves. Some advances have been made with the use of epinephrine, a potent inductor of metamorphosis in bivalve larvae, allowing massive and synchronous settlement in the controls (see Section 3.2.4.5). On the other hand, heavy metals do not necessarily inhibit larval settlement. Sub-lethal concentrations of copper are known to stimulate settlement in oyster larvae (Prytherch, 1934, in C. virginica; Nell and Holliday, 1986, in Saccostrea commercialis; P. Salkeld, Plymouth Marine Laboratory, personal communication, in C. gigas). 4.1.1.4. Respiration. Heavy metals are known to increase respiration in all developmental stages of bivalves, including eggs and larvae. Oxygen consumption increased by 1.2 times to 2.7 in Spisula solidissima larvae reared at 50 pg Ag 1-' (Thurberg et al., 1975) and by about 2 times in unfertilized eggs of Mytilus edulis exposed to 30 000 pg Cu 1-' (Akberali et al., 1985). Mercury also increased oxygen consumption of 5-day-old C. virginica larvae after 24-hour exposure to 1-100 pg 1-' (Cunningham, 1972, cited by Cunningham, 1979). However, zinc (100 pg 1-') and cadmium (2000 pg 1-l) have been reported to cause slight (1620%) decreases in respiration in mussel embryos (Pavicic, 1980). The sensitivity of this response is not substantially different from the test of embryogenesis inhibition, whereas the latter is much more simple, rapid and accurate to measure. Moreover, these tests have yielded contradictory results (generally an increase, but also some cases of decrease in respiration) which are difficult to interpret. Therefore respiration does not seem to be a suitable response for embryo and larval bioassays. 4.1.1.5. Genotoxicity. Mercury has been shown to be genotoxic, as exposure to 30 pg HgC121-1 doubles the mean number of sister-cromatid exchanges (SCE, a sensitive indicator of chromosomal damage), in fertilized mussel eggs (Brunetti et al., 1986). Nitrilotriacetate (a com-
96
E. HIS, R.
BEIRAS AND M. N. L. SEAMAN
ponent of some phosphate-free detergents), which is suspected of having mutagenic effects, did not increase SCE at the concentration tested (5mg1-'), nor did it interact with the genotoxicity of mercury chloride (Brunetti et al., 1986). TBT-oxide (up to 1pgl-') was not found to be genotoxic to 12-hour-old mussel larvae either, based on the results of SCE and chromosomal aberrations (Dixon and Prosser, 1986). Fractions extracted with an organic solvent from oil-polluted sediments have also been assayed for genotoxicity in bivalve larvae (Wolfe et at., 1995), but the results are difficult to interpret, because there was significant SCE induction by the reference sediment as well. 4.1.2. Biocides
The available data on biocide concentrations causing 50% reduction in normal embryogenesis, larval survival, growth and settlement are summarized in Table 11. Chemicals are listed in tentative order of decreasing toxicity, based on the highest toxicity values reported. In order to focus on the most relevant environmental pollutants, we are limiting this review to the most toxic biocides for bivalve larvae, i.e. those with ECS0values of less than 10 ppm (10 000 pg 1-') in at least one experiment. Substances of lesser toxicity, for which all published studies give an ECS0higher than lOppm, are not considered here regardless of the fact that they may be highly toxic to organisms other than bivalve larvae. Tributyl-tin (TBT) is the most toxic chemical assayed to date with bivalve embryos and larvae, with an EC50 below the unit ppb level in several instances. This is not surprising, considering that oysters and other molluscs are target species for this active component of antifouling paints. Halogens (chlorine, bromine) and halogen-produced oxidants (such as monochloramine, chloroform, bromamines, bromide, bromate, halomethanes, etc.) are also extremely toxic. Accurate measurement of EC50 values is difficult owing to the high volatility of these substances, and experimental designs should employ flow-through systems. Identification of individual halogen compounds is usually not attempted, but Stewart et at. (1979) found high toxicity of bromate and comparatively low toxicity of chloroform and bromoform. Data on ozone-produced oxidants are shown in Table 11 for comparison. Again, bivalve larvae are target species of these compounds, used in energy plants to prevent their cooling-water pipes clogging. Cyanide (CN-) is highly toxic to all aerobic organisms, and early developmental stages of bivalves are no exception, with ECsovalues in the 10-100 pg 1-' range (Okubo and Okubo, 1962; Pavicic and Pihlar, 1982; Pablo et al., 1997).
THE ASSESSMENT OF MARINE POLLUTION
97
Intensive agriculture increasingly relies on the use of phyto-sanitary chemical products for pest control. In 1992 the world consumption of these products was approximately 1.4 million tons (Andral, 1996). In coastal areas these products are leached from the soil by precipitation and drained into the sea. The toxicity of a great number of insecticides, herbicides and other biocides has been assayed with early developmental stages of bivalves, but the most important contribution still remains the pioneering study by Davis and Hidu (1969a). More recently, His and Seaman (1993) extended these studies of lethal and sub-lethal toxicity in embryos and larvae to 12 commercially available pesticides. Some organochlorinated pesticides such as trichlorocarbanilide (TCC), 2,3dichloro-l,4-naphthoquinone(Phygon), dichlorodiphenyltrichloroethane (DDT), pentachlorophenol (PCP), dehydroabyetilamine compounds (Delrad, Rosin Amine D), and cyanide can be toxic at concentrations below 100 pg 1-l. Toxicity of chlorophenols increases with the number of chlorine substituents, from an ECsO in the order of 20000pg1-' for 4-chlorophenol (4-CP, one chlorine atom) (Krassoi et al., 1997), to values around 1000 for tetrachlorophenol (TCP, four chlorine atoms) (Davis and Hidu, 1969a), and 27-55 pg 1-' for pentachlorophenol (PCP) (Woelke, 1972). Another group of persistent chlorinated insecticides (dieldrin, endrin, lindane, methoxychlor, toxaphene, aldrin) and copepodicides (dichlorvos) can be toxic at 100-500ppb. Dichlorvos, a highly toxic pesticide used in salmon farming for the control of ectoparasitic copepods, is particularly relevant in marine toxicology, because it is deliberately introduced into the marine environment in appreciable quantities. Crustacean larvae are ten times more sensitive to Dichlorvos than molluscs (LCs0: 5-50 pg 1-' for lobster larvae, according to McHenery et al., 1991). Organo-phosphorated pesticides are generally less toxic, with EC50values in the range of 500-1000 pg 1-' (Parathion, Fenitrothion, Guthion, etc.), or higher (Malathion). Herbicides (e.g. 2,4-D, atrazine-simazine, Neburon, Diuron, etc.) are usually toxic to bivalve embryos and larvae only at concentrations above the ppm level (e.g. Robert ef al., 1986), with the exception of Dinoterbe, a highly toxic phenolic herbicide used in maize culture, which has an ECsOof 50-100 pg 1-'. As in the case of metals, pesticides affect embryonal development more than larval survival. Growth reduction may take place at concentrations below those inhibiting embryogenesis, although this is not always the case (see Table 11). The sensitivity of larval growth is particularly dramatic in the case of TBT, which reduces the rate of growth at concentrations in the order of 1ng 1-l (His et al., 1983; Lapota et al., 1993). In those cases in which there are sufficient data for comparison, the various bivalve species (oysters, mussels and clams)
Table 11 Toxicity of pesticides, biocides and miscellaneous chemicals to early developmental stages of bivalves. LCso: toxicant concentration causing 50% mortality; otherwise ECSo:toxicant concentration causing 50% reduction in the end-point. Chemicals are tentatively ranked in decreasing toxicity order. Except for TBT,data on other toxicants are mostly from the classic work by Davis and Hidu (1969a); these authors refer to Loosanoff and Davis (1963) for details on experimental procedures. Davis and Hidu (1969a) and His and Seaman (1993) used acetone as a carrier when appropriate. Pesticide TBT
Test sp.
Initial age/&
Exposure conditions
End-voint
Notes
C. gigas
30 min fertilized egg
24h, F W 24% ZEpsu, 0.8 pm
embryogenesis
<5
His and Robert, 1980
C gigas
30 min fertilized egg
24 h, 24°C 28psu. 0.8 pm
embryogenesis
(1
Robert and fig1981
c gigas
30 min fertilized egg
FSW 48 h, 24'C 28psu, 0.2 pm FSW
embryogenesis
0.5-1.
C. virginica
4-16 cell embryos
M.galloprovincialis 3 0 m h fertilized egg M. mercenaria
4-16 cell embryos
I. cnlifomicum
sperm + egg (1 b later) fertilized egg
1. difomicum
- - ..
48 h, 20-24'C, 1&22psu, 40,'&I 1pm carbon FSW 24 b, 1YC 28psu, 0.8 pm FSW 48 b, 20-24T, 1&22psu, 40d-', 1pm carbon FSW 2 b, 34psu, ux)ml-', 0.45 pm FSW 48 h, 34psu, 17d-', 0.45 @n FSW 48 h, 24'C 28psu, 0.8 pm FSW 24% 28psu. mix algal food, 0.8 @n FSW
His ef al., 1983
embryogenesis
1.30.
embryogenesis
<1
Robert and Hig1981
embryogenesis
1.13
Roberts. 1987
cleavage
4.0
Ringwwd, 1992b
Roberts. 1987
(acetate) (abnormal larvae excluded) (acetate) (abnormal larvae excluded) (acetate) (abnormal larvae excluded except visceral mass abnonnalities) '(0.71 acetone as carrier) (acetate) (abnormal larvae excluded)
embryogenesis
0.5
Ringwood,1992b
(abnormal larvae excluded)
mortality
<5
His and Robert, 1980
mortality
M (4 d)
(acetate, glacial acetic acid as carrier) (acetate, glacial acetic acid as carrier)
C gigas
egg
24°C 28psu, 8 d-l,mix algal food, 0.2 w FSW
mortality
c gigas
not given
48 h, up to 1I&'
mortality
C virginicn
24 h
mortality
M. edulis
not given
48 h. 2&24T, lS-22psu. 10ml-', mix algal food, 1 pm carbon FSW 48 h. up to 1
M. edulis
7 d, 133J.UU
mortality
15 d, WC. 33psu. 4 d-', mortality (oesophageal mix algal food, 0.2 pm FSW uv sterilized cilia ceased heating)
Robert and His 1981 1-3 (5 d) <1 (9 d) 0.5-1 (6 d) His ef al., 1983 0.24.5 (8 d) 0.0210.05 (12 d) Thain, 1983 1.6 3.%
Roberts 1987
2.3
Tbain, 1983
0.100
Beaumont and Budd. 1984
(acetate, glacial acetic acid as carrier) (bisTBT oxide, acetic acid 01 ethanol as carrier) (cbloride, glacial acetic acid as camer) (bisTBT oxide, acetic acid or ethanol as carrier) (bisTBT oxide, carrier glacial acetic acid)
M. edulis
8d, 140pm
M. galloprovincialis 1d(?), 73 pm M. mercenaria
24 h
M. mercenaria
24-48 h
S. plana
10 d, 240 pm
C. gigas
24 h, 61pn
c. gigm M. edulis
7 d, 133pm
M.edulis
8 d , 140pm
M.galloprovincialis 24 h
CI
(?), 73 pm
M. mercenaria
4 h fertilized egg
S. plana
2d, loOpm
s. p l M 4
10d, 239pn
I. californicum
3d
C. virginica
gastrulaltrochophore
M.mercennria
25 d, 19T, 2 d-', mix algal food, 50 pm FSW
mortality (empty valves)
4 d, lPC, 28psu, mix algal mortality food, 0.8 pm FSW mortality 48 h, 2&24°C, 18-22psu, 10 ml-', mix algal food, 1 pm carbon FSW mortality 8 d, temperature not given. 3Oosu. 0.1 dP.monodeal food 30 d, 20.5"C, 24psu. 0.8 d-',mortality' monoalgal food, 0.45 wm FSW 48 h, W C , Bpsu, mix algal growth (height) food, 0.8 pm FSW 8 d, 24°C. 28psu. 8 d-'. mix growth (height) algal food, 0.2 pm FSW 15d, WC, 33psu, 4 d - ' , growth (length) mix algal food, 0.2 pm FSW uv sterilized 25 d, 19T. 2 mi-', mix algal growth (length) food, 50 pm FSW growth (height) 24 h, 19'C, 28psu, mix algal food, 0.8 prn FSW growth (length) 14 d, 25"C, 32psu, 3 ml-', monoalgal food 6 d, 22.5"C, 24psu, growth (length) monoalgal 0.4-3 d-', food, 0.45 pn FSW 30 d, 20.5"C24psu. 0.8 d-',growth (length) monoalgal food, 0.45 pm FSW 4 d, temperature not given, growth 34psu, lOd-', mix algal (dry weight) food, 0.45 pn FSW 17-28'C*, lMOpsu
mortality (48 h)
>0.13* 5
1.65 -0.6. 0.13-0.31
0.1-1
Lapota er 01.. 1993
Robert and Hi$1981 Roberts 1987 Laughlin ef al., 1989
*1-2.5 at 48 h @isTBT oxide in acetone)
Ruiz ef al.. 199%
(bisTBT oxide, ethanol as camer) 'nowexistent or inactive soft puts inside gaping valves (acetate, glacial acetic acid as carrier) [acetate, glacial acetic acid as carrier) (hisTBT oxide. glacial acetic acid as camer)
Robert and Hi% 1981
His et al., 1983 Beaumont and Budd. 1984
0.W64.050 Lapota ef al., 1993 <1 0.0~.10
Robert and
His 1981
Laughlin el al., 1988
(TBT salt and camer not given) [acetate, glacial acetic acid as carrier) (hisTBT odde, acetone as carrier) (bisTBT oxide, ethanol as carrier)
<0.13
Ruiz ef al., 1995a
<0.13
Ruiz er al., 199%
(bisTBT oxide, ethanol as carrier)
>0.5
Ringwood, 1 W b
(TBT chloride)
Roberts er a/., 1975
'those ambient depending on season, no more than e2'C within a single experiment **depending on methods
<s-110** <5
(TBT salt and camer not given) 'making allowance of control mortality (acetate, glacial acetic acid as carrier) (chloride, glacial acetic acid as carrier)
Test sp.
Pesticide
Phygon
Delrad
End-point
ECsa or L% I-')
Reference
7 d larva
open-flow system
C gig-
0.5-2 h 1st polar 2onl-'
19, 22, 25,ZB"C 18,26,34psu
3-7 h blastula 20 tn-' 6 1 6 b trocophore 20 tn-' 1 6 3 0 b larva 5 d-', monoalgal diet 24-50 h lama 5 mlP, monoalrral diet pediveliger
19, 22, 25, 28'C 18,26,34psu 19.22, 25. 28°C 18,26, 34psu 19.22, 25, 28'C 18,26, 34psu 19, 22, 25, 28°C 18, 26, 34mu
1B"c
120 (20°C) Capulzo, 1979 80 (U'C') 10 (ZOT)** <10(25T*)** no cleavage (1 h) 1031, 1303, Chien and Chou, 1989 1322.1070 852, 1273, 1425 456, 524,606, 654 mortality 458, 571,651 (1 hY 384,362,502,926 mortality 430, 524, 677 (1 h)' 556,729. 867, 950 mortality 688,766,872 (1 b)' 607,589,569,550 mortality 525, 573,638 (1 h)* 1500' Bucaille and Kim, 1979 settlement
fertilized egg (usually 2-cellstage) 2d
48h
embryogenesis
10 d, 24T, mix algal food
mortality growth
C. virginica
fertilized egg (usually 2ceU stage) 2d
48h
embryogenesis
14
Davis and Hidu, 1%9a
12 d, 24°C mix algal food
M. mercenaria
fertilized egg 2d
48 h 10 d. ZaT, mix algal food
mortality growth embryogenesis mortality growth
41 5-10 14 1750 25-50
Davis and Hidu, 1%9a
C.virginica
fertilized egg 2d fertilized egg 2d
12 d, 24°C mix algal food
M. mercenaria
M. mercenaria
CN-(NaCN)
Exposure conditions
C. virginica
Mytilus sp.
TCC
Initial agdsize
10 d, 24T, mix algal food
mortality (48 h, 30 min exposure)
mortality
grad,
mortality growth embryogenesis
Cgigas M. edulis M. galloprovincialis fertilized egg D-larva D-larva
48 h, 37-38psu
Chlamys arperrimn 30-min-old embryo
lVC, 32psu, 30d-', 5 pm FSW uv sterilized
embryogenesis* mortality (48 h) growth (48 h)
embryogenesis (48 h)
32
Notes 'back to U1"C after exposure *+a added as chloramine
*no movement, immobile cilia *no movement, immobile d i a
'no movement, immobile cilia *no movement, immobile cilia 'continuous chlorination
Davis and Hidu, 1%9a
37 2.5-5
31 50 72
Davis and Hidu, 1%9a Davis and Hidu, 1%9a
10-50
32-100 32-100
lo.@* w
Okuho and Okubo, 1962 Pavicic and Pihlar, 1982
'abnormal larvae excluded **l5,11.7 with Cd and zinc cyanide ***c.16,10 with Cd and zinc cyanide
Pablo er al., 1997
*U8,686with iron-cyanide complexes
c. ll***
286*
Rosin A m i e D C. virginica
fertilized egg 2d
48 h 12 d, ?AT, mix algal food
embryogenesis mortality
<250 27-55
Woelke, 1972
1250 71
Davis and Hidu, 1%9a
PCP'
C. gigus
f e r t i i i d egg
20T, P25psu, 2&30ml-', FSW uv sterilized
embryogenesis (48 h) (fully shelled*)
Pcp
C. virginica
fertilized egg 2d
48h 12 d, 24°C. mix algal food
embryogenesis mortality growth
C virginica
fertilized egg 2d
48h 12 d, 24°C. mix algal food
embryogenesis mortality growth
4250 <25
PCP'
M. edulis
fertilized egg
48 h, 24 psu
embryogenesis
300-400
CI-produced oxidants
C. Virginia
W
C Virginia
eyed, >14 d fertilized egg
24°C. 12-14psu, no food, 1pn mortality' FSW, flow-through system (48, .% h) Bow-through system mortahty (48 h)
M. lateralis C. virginica
M. arenaria
97-117 pm 308 pn 100% eyed
27-28°C. 15-16psu. diluted sewage, no food*, flow-through system
3648 h
17-20-C 13psu, 1wm FSW, flow-through system
18 d
Davis and Hidu, 1969a
ZpentachJornphenyl acetate
R. Brown, cited by Dimick and Breese, 1965
'Na pentachloropheuate
Roosenhurg et a!., 1980b
*no movement and disturbed
>25
300, 64 -300, -300 27,46** two separate experiments
internal organization *author mentions monochloramine, bromamine. bromide, halomethanes and bromate
Roberts 1980
38 mortality (96 h) <20 Roberts and Casey, 1985 metamorphosis 30-60** (every 24 h until end of spatfall) mortality (no c. 500 (8 h) Roosenburg et al., 1980a movement and c. 300 (16 h) >500 (16 h) disturbed internal W 5 0 0 (24 h) <200 (48 h) organization)
BrCl-produced oxidants
C. virginica
4 h fertilized egg
ZOpsu, flow-through system
mortality (48 h)
100, 210** two separate experiments
CIBr-produced oxidants*
C virginica
fertilized egg
flow-through system
mortality (48 h)
220
03-produced oxidants
C virginica
48 h D-larva 9-21d pediveliger
25°C. 12psu. 1pm FSW. flow-through system
mortality* (48, 96 h)
DDT
C. gigas
fertilized egg
2O"C, XSpsu, 2&30 F'SW uv sterilized
I&',
*even misshapen or undersized 'Na pentachlorophenate
>so
Pcp2
h
Davis and Hidu, 1%9a
<25
embryogenesis (48 h) (h11~ shelled*)
-300, 125
Roberts and Gleeson, 1978
Roberts 1980
Richardson
ef
>300, >300 100-1WO
'mixed diet for mortality exp. **1&30 in other experiments with younger pediveligers
'author mentions bromamine, bromide, halomethanes and bromate a[., 1982
*no vital stain, empty shells lack of movement in D-larvae
Wnelke, 1972
'even misshapen or undersized
EC, or LG, Pesticide
Test sp.
Initial agelsize
Exposure conditions
End-point
(fig 1-7
Reference
C virginica
2d
12 d. 24°C mix algal food
mortality growth
34 >25
Bromate
C. Virginia
fertilized egg
48 b, ZMPC ZSpsu, 15 ml-',
mortality
50-100
Stewart el d..1979
Dinoterbe'
C gigm
fertilized egg
24!%?!,!!8~l-1, FSW
embryogenesis (24 h)
5C-100
His and Beirns
70
Lucu cf d.,1980
78 340
Davis and Hidu. 1%9a
0.2 pm
Slimicide'
M.g d o p r o v i n c i d u fertilized egg
96 h, 13°C37psu
embryogenesis
Omazene
C Virginia
48 h 12 d, 24°C mix algal food
embryogenesis mortality growth embryogenesis mortality
M.merenaria
Co-Ral
Dichlorvos
fertilid egg 2d fertilized egg
Zd
48h 10 d, W C , mix algal food
C virginiua
fertilized egg 2d
48h 12 d, 24°C mix algal food
M. mercenaria
fertilized egg 2d
48h 10 d. 24T, mxi algal food
C gigar
fertilized egg (1-2 h)
48 h, U'C 50 n P , 0.45 pm
Notes
Davis and Hidu, 1%9a
*phenolic herbicide
(unpublished data)
growth
1lx-250 81 378 250-500
embryogenesis mortality growth embryogenesis mortality growth
110 >loo0 1lx-250 9120 5210 500-5000
embryogenesis
165
*bis-trichloromethyl-sulphone (20%), methylene bis-thiocyanate(5%)
Davis and Hidu, 1969a
Davis and Hidu, 1%9a Davis and Hidu, 1%9a
ll~ainel a/., 1990
FSW Dieldrin
C virginiua
fertilized egg 2d
48h 12 d, 24°C mix algal food
embryogenesis mortality growth
640 >10 000 1lx-250
Davis and Hidu. 1%9a
Endrin
C. Virginia
fertilized egg 2d
48h 12 d, 24'C mix algal food
embryogenesis mortality growth
>lo 000
790
Davis and Hidu, 1%9a
Lindane
C. gigm
fertilized egg
20% 325psu, 20-30 ml-', FSW uv sterilized
embryogenesis
5000
C gigm
fertilized egg (30 min)
9d, 24°C Bpsu, 3 O d - l (egg), 8 d-' (larva), mix algal food, 0.2 pm FSW
(48 h)
1Cn-250 Woelke, 1972
*even misshapen or undersized
His and Seaman, 1993
'6 d
(fUW
shelled') mortahty growth
170 100-250*
M. mercstaria
fertilized egg fertilized egg 2d
c5 P
fertilized egg
C virginica
Methoxychlor
48h 48h 10 d. 24°C mix algal food
embryogenesis embryogenesis mortality growth
9100 >loo00 >loo00 >loo00
Davis and Hidu. 1%9a
embryogenesis
Mo (430) 170 (260) 250 (440) 210 (160)
Cardwell er al., 1979a
190
Davis and Hidu, 1969a
(and 3 other spp.)
Roccal
M.mercenaria
fertilized egg 2d
48h 10 d, 24"C, mix algal food
embryogenesis mortality growth
140 >lo0
Dowcide G
M. rnercenarin
fertilized egg 2d
48h 10 d. 24°C. mix algal food
embryogenesis mortality
<250 <250
Davis and Hidu, 1%9a
Toxaphene
M.memenaria
fertilized egg 2d
48h 10 d, 24°C mix algal food
embryogenesis mortality growth
1120
Davis and Hidu. 1%9a
<250
fertilized egg 2d
48h 10 d, 2 4 T mix algal food
embryogenesis mortality
>loo00 410
Aldrin
M. mercenada
<250
growth
<250
Davis and Hidu, 1%9a
Griseofulvin
M.mercenaria
fertilized egg 2d
48h 10 d. 24T, mix algal food
embryogenesis mortality growth
<250 <1wO >lwO
Davis and Hidu, 1%9a
Propen-1-01-3
M.mercenarin
fertilized egg 2d
48h 10 d. 24'C, mix algal food
embryogenesis mortality
t250
1030
Davis and Hidu, 1%9a
C. virginica
fertilized egg 2d
48h 10 d 24°C mix algal food
embryogenesis mortality growth
5900
Davis and Hidu. 1%9a
25C-5500
fertilized egg 2d
48h 12 d, 24°C mix algal food
embryogenesis mortality growth
25C-5500
Silvex
Nemagon
Isoproturon
N-3452
M. mercenada
C gigas
C virginica
fertilized egg (30 min)
9 d, 24°C 28psu. 30 ml-l (egg). 8 tn-' (larva). mix algal food, 0.2 c ~ FSW m
mortality growth
fertilized egg 2d
48h 12 d. 24°C mix algal food
embryogenesis mortality
710 10 Ooo 780 370
Davis and Hidu, 1969a
His and Seaman. 1993
2500
Davis and Hidu. 1969a
'fully shelled regardless of shape (total)
Pesticide
Test sp.
Initial agehize
Exposure conditions
End-point
EC, or LC& ( M1.')
Reference
Notes
Nabam
C virginica M . rnercenana
fertilized egg fertilized egg 2d
48 h 48h 10 d, 24°C mix algal food
embryogenesis embryogenesis mortality growth
Davis and Hidu, 1969a Davis and Hidu. 1969a
TCP
C. virginica
fertilized egg 2d
48h 12 d. 24% mix algal food
embryogenesis mortality growth
Davis and Hidu, 1969a
Parathion'
C gigas
fertilized egg (30 d)
9d, 24% 28psu, 3Oml-' (egg), 8 d - ' (larva), mix algal food, 0.2 I . L ~FSW
mortality
His and Seaman, 1993
'methyl-parathion
C gigas
fertilized egg
20°C. *25psu, 20,30ml-', FSW uv sterilized
Woelke, 1972
*even misshapen or undersized
C virginica M. edulis
2d fertilized egg
12 d, 24% mix algal food 1>17"C
embryogenesis (48 b) (fully shelled) growth embryogenesis
Davis and Hidu, 1%9a Okubo and Okubo, 1962
Fenitrothion
c gigar
fertilized egg (30 min)
9d, 24°C 28psu, 30~1.' (egg). 8ml" (larva). mix algal food, 0.2 pm FSW
growth
His and Seaman. 1993
Guthion
C. virginica M.mercenaria
fertilized egg fertilized egg 2d
48h 48h 10 d, 24T, mix algal food
embryogenesis embryogenesis mortality
Davis and Hidu, 1%9a Davis and Hidu. 1%9a
2;J-D ester
C. virginica
fertilized egg 2d
48 h 12 d, 24°C mix algal food
embryogenesis mortality growth
Davis and Hidu. 1%9a
Dowcide A
M. rnercenana
fertilized egg 2d
48 h 10 d, 24°C mix algal food
embryogenesis mortality growth
Davis and Hidu, 1969a
C. virginica
fertilized egg 2d fertilized egg 2d
48h 12 d, 24°C mix algal food 48h 10 d, 2 f C . mix algal food
embryogenesis mortality embryogenesis mortality
Davis and Hidu, 1%9a
fertilized egg 2d
48 h 12 d, 24% mix algal food
mortality
Davis and Hidu, 1969a
fertilized egg
48 h, 2&29"C, ZSpsu, 15In-', 0.45 m FSW
Parathion
N-3514
M. mercenaria Dipterex
C. virginica
Uranin (stain)
M.edulis
Bromoform
C virginica
Chloroform
growth
Davis and Hidu, 1%9a
growth embryogenesis
Okubo and Okubo, 1962
mortality
Stewart et af., 1979
Di-syston
C. virginica
fertilized egg 2d
48 h 12 d, 24°C. mix algal food
M. mercenaria
fertilized egg 2d
48 h 10 d, 24°C mix algal food
AtrazineSimazine'
c. grga
Mecoprop
C gigas
Sevin
24 h, 2 4 T 28psu. 81n-' (larva), mix algal food (larva), 0.2 pm FSW
embryogenesis* mortality (10 d) growth** (5 d)
fertilized egg (30 min)
9 d, 24°C 28psu, 30 d-l (egg), 8ml-' (larva), mix algal food, 0.2 pm FSW
mortality growth
48 h, 20°C. 25psu. 5 0 n t ' emhryogenesis (Dimick and Breese, 1965) 2 0 PZPSU. ~ 2 6 3 0 ml-1, embryogenesis FSW uv sterilized (48 h) (fully shelled') 48 h embryogenesis 12 d, 24% mix algal food mortality prowth 48 h, 20°C 25psu. 5 0 d P embryogenesis 1 h exposure + 48 h recovery, embryogenesis 17°C. 25psu, 1 0 d - ' , (abnormal sand-FSW, no food larvae excluded)
c gigm
egg
C. gigas
fertilized egg
C. virginica
fertilized egg 2d
M. edulu
M. edulis
M. mercenario
embryogenesis mortality growth embryogenesis mortality growth
5860 3670
Davis and Hidu, 1969a
1W2500
5280 1390 1W2500 5W10000
Davis and Hidu, 1969a
Robert ef al., 1986
<2500 1CQO-2500 4200 1W2500 2200 (803:) lEO&3700 wx)
'25% Atrazine-25% Simazine *abnormal larvae excluded **height
His and Seaman,1993
Stewart ef 01.. 1967 Woelke, 1972
*values for I-naphthol, Sevin hydrolytic product *even misshapen or undersized
Davis and Hidu, 1%9a
3000
>m
egg egg 1st polar body (25 min) 2ceU (1h) 64cell or more (4 h) blastula (8 h) trocophore (20 h) veliger (32 h) fertilized egg 2d
48h 10 d, 24°C. mix algal food
embryogenesis mortality growth
3820 2500 1W2500
2300 (1300:) Stewart ef al., 1967 20 700 (24 5 0 0 3 Armstrong and 5300 (5200') Millemann, 1974
7000 8300 16 000 19 000
2Aooo Davis and Hidu, 1969a
Fenvalerate
C. virginica
not given
48 h, not given
emhryogenesis
>loo0
P. W. Borthwick, cited by
Neburon
M. mercenan'a
fertilized egg 2d
48 h 10 d, 24°C. mix algal food
embryogenesis mortality
<2400 <2m
Davis and Hidu, 1%9a
Diuron
M. mercenaria
fertilized egg 2d
48h 10 d, 24°C. mix algal food
emhryogenesis mortality growth
2530
Davis and Hidu, 1969a
>5000 >So00
Tagatz and hey. 1981
*values for I-naphthol, Sevin hydrolytic product
Exposure conditions
End-point
C. virginica
fertilized egg 2d
48h 12 d. 24"C, mix algal food
embryogenesis mortality growth
Davis and Hidu. 1%9a
C. virginica M. mercenaria
fertilized egg fertilized egg 2d
48 h 48h 10 d, 24"C, mix algal food
embryogenesis embryogenesis mortality growth
Davis and Hidu, 1%9a Davis and Hidu, 1%9a
embryogenesis
Okubo and Okubo, 1%2
Test sp.
Pesticide Malathion
Rhodamine B (stain)
M. edulis
T m i c acid
M. edulis
Initial age/size
20°C ==25psu,20-30 ml-', FSW uv sterilized
C gigas KKIO,
c
embryogenesis embryogenesis (48 h*)
3200-10000 210 000
Okuho and Okubo, 1962 Cardwell er 01.. 1979a
embryogenesis
3200-10 000 3200-10 000
Okubo and Okubo, 1962
M. edulis
Dicapthon
Phosphamidon Monuron
M. mercenarin
C. gigas M. mercenaria
Reference
fertilized egg 2d
48 h
10 d, 24°C mix algal food
embryogenesis mortality growth
3340 5740 >2000
fertilized egg
20% ~ 2 5 p s u 20-30 . ml-', FSW uv sterilized
embryogenesis
4lmo-mm
48h 10 d, 24'C. mix algal food
embryogenesis mortality
fertilized egg 2d
Notes
*fully shelled, even misshapen or undersized
Davis and Hidu, 1%
Woelke, 1972
(48h*)
*fully shelled, even misshapen or undersized
Davis and Hidu. 1%9a
growth
Fenuron
M. mercenaria
fertilized egg 2d
48h 10 d, 24°C mix algal food
embryogenesis mortality
Davis and Hidu, 1%9a
Carbofuran
C. gigas
fertilized egg (30 min)
9d. 24% Bpsu, 3 O d P (egg), 6 d - l (larva), mix algal food, 0.2 p m FSW
mortality growth
His and Seaman, 1993
Bromoxynil
c gigas
fertilized egg (30 min)
9 d, 24°C 28psu, 30 I&' (egg), E d - ' (larva), m h algal food, 0.2 pm FSW
mortality growth
7000 5000-10000
His and Seaman, 1993
Metaldehide
C. gigas
fertilized egg (30 min)
9 d, 24% 28psu, 30 m1-l (egg), 8 d P (larva), mix algal food, 0.2 pm FSW
mortality growth
7400 5000-10 000*
His and Seaman. 1993
Carbetamide
c. gigas
fertilized egg (30 min)
9 d, 24°C 28psu. 30 ml" (larva), mix (egg), 6 d-' algal food, 0.2 FSW
mortality growth
93w
His and Seaman, 1993
210000
Bold: 95% CI provided. FSW, filtered seawater.
*6 d
*6 d
107
THE ASSESSMENT OF MARINE POLLUTION
Table 12 Interspecific comparison of sensitivity of bivalve embryos to Sevin insecticide. Median effective concentrations (EC,,) inhibiting normal embryogenesisare given.
Species
EGO(mg 1-7 ~~
Crassostrea gigas C. virginica Mytilus edulis
Mercenaria mercenaria
2.2 1.8-3.7 2.04.0
1.6-3.2 2.3 2.5-4.0
Reference ~
~~
Stewart et al., 1967 Woelke, 1972 Davis and Hidu, 1969b Dimick and Breese, 1965 Stewart et al., 1967 Davis and Hidu, 1969b
show similar sensitivity to biocides (e.g. Sevin, Table 12); this corresponds to the similarity in their sensitivity to metals. The effects of biocides on larval settlement are poorly documented. DDT did not greatly affect settlement in C. gigas (Loosanoff, 1954), but DDT and other insecticides did affect settlement in Ostrea edulis (Waugh et al., 1952). Cardwell et al. (1979b), working with C. gigas embryos at 20°C, reported a 48-hour EC50ammonia of 15 mg NH3-N1-’. Gormly et al. (1996) assayed the toxicity of microbial pest-control agents (Bacillus thuringiensis, B. alvei, Metharyzium anisopliae) and of a virus of the gipsy moth to D-larvae of Mulinia lateralis, finding strong effects of B. thuringiensis upon this non-target species. 4.1.3. Detergents and oil Data on the toxicity of detergents and oil to bivalve embryos and larvae are given in Table 13. Detergents are usually a mixture containing at least one surfactant and one builder. The surfactant is the active component that reduces the surface tension of the water and dissolves the organic molecules of a stain. The first synthetic detergents included non-linear surfactants, which were poorly degradable. Owing to environmental concerns, these so-called “hard” detergents have largely been replaced by “soft” detergents, which use readily degradable compounds with a linear alkyl group, such as the linear alkylate sulphonates (LAS) as surfactants. The builder of a detergent contributes to the cleaning action by sequestering cations that would otherwise interfere in the action of the surfactant. Polyphosphates used to be the typical cation-sequestering agents in detergents, but concerns about their role in water eutrophication have prompted their replacement. Nitrilotriacetate (NTA) was one of the alternative builders, but its innocuity to mammals has been under discussion (Stoker and Seager, 1976), and zeolites (sodium aluminium
Table 13 Toxicity of detergents, oil and detergent-oil mixtures to early developmental stages of bivalves. LC5,,: toxicant concentration causing 50% mortality; otherwise ECS0:toxicant concentration causing 50% reduction in the end-point. Several detergents were assayed as mixtures: in these cases, only the active components are mentioned and their percentages in the assayed mixture provided. Concentrations are expressed as p,g I-’ of active cornponenth except when otherwise stated. Detergent!Oil
Test species
Cationic surfactants para diisobutyl C.virginicn phenoxyethoxy ethyl dimethyl benzyl ammonium chloride
