Reviews of Environmental Contamination and Toxicology Volume 204

Reviews of Environmental Contamination and Toxicology VOLUME 204

For further volumes: http://www.springer.com/series/398

Reviews of Environmental Contamination and Toxicology Editor

David M. Whitacre

Editorial Board Maria F. Cavieres, Valparaiso, Chile • Charles P. Gerba, Tucson, Arizona, USA John Giesy, Saskatoon, Saskatchewan, Canada • O. Hutzinger, Bayreuth, Germany James B. Knaak, Getzville, New York, USA James T. Stevens, Winston-Salem, North Carolina, USA Ronald S. Tjeerdema, Davis, California, USA • Pim de Voogt, Amsterdam, The Netherlands George W. Ware, Tucson, Arizona, USA

Founding Editor Francis A. Gunther

VOLUME 204

123

Coordinating Board of Editors D R . DAVID M. W HITACRE , Editor Reviews of Environmental Contamination and Toxicology 5115 Bunch Road Summerfield North, Carolina 27358, USA (336) 634-2131 (PHONE and FAX) E-mail: [email protected] D R . H ERBERT N. N IGG , Editor Bulletin of Environmental Contamination and Toxicology University of Florida 700 Experiment Station Road Lake Alfred, Florida 33850, USA (863) 956-1151; FAX (941) 956-4631 E-mail: [email protected] D R . DANIEL R. D OERGE , Editor Archives of Environmental Contamination and Toxicology 7719 12th Street Paron, Arkansas 72122, USA (501) 821-1147; FAX (501) 821-1146 E-mail: [email protected]

ISSN 0179-5953 ISBN 978-1-4419-1439-2 e-ISBN 978-1-4419-1440-8 DOI 10.1007/978-1-4419-1440-8 Springer New York Dordrecht Heidelberg London Library of Congress Control Number: 2009911931 © Springer Science+Business Media, LLC 2010 All rights reserved. This work may not be translated or copied in whole or in part without the written permission of the publisher (Springer Science+Business Media, LLC, 233 Spring Street, New York, NY 10013, USA), except for brief excerpts in connection with reviews or scholarly analysis. Use in connection with any form of information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed is forbidden. The use in this publication of trade names, trademarks, service marks, and similar terms, even if they are not identified as such, is not to be taken as an expression of opinion as to whether or not they are subject to proprietary rights. Printed on acid-free paper Springer is part of Springer Science+Business Media (www.springer.com)

Foreword

International concern in scientific, industrial, and governmental communities over traces of xenobiotics in foods and in both abiotic and biotic environments has justified the present triumvirate of specialized publications in this field: comprehensive reviews, rapidly published research papers and progress reports, and archival documentations. These three international publications are integrated and scheduled to provide the coherency essential for nonduplicative and current progress in a field as dynamic and complex as environmental contamination and toxicology. This series is reserved exclusively for the diversified literature on “toxic” chemicals in our food, our feeds, our homes, recreational and working surroundings, our domestic animals, our wildlife, and ourselves. Tremendous efforts worldwide have been mobilized to evaluate the nature, presence, magnitude, fate, and toxicology of the chemicals loosed upon the Earth. Among the sequelae of this broad new emphasis is an undeniable need for an articulated set of authoritative publications, where one can find the latest important world literature produced by these emerging areas of science together with documentation of pertinent ancillary legislation. Research directors and legislative or administrative advisers do not have the time to scan the escalating number of technical publications that may contain articles important to current responsibility. Rather, these individuals need the background provided by detailed reviews and the assurance that the latest information is made available to them, all with minimal literature searching. Similarly, the scientist assigned or attracted to a new problem is required to glean all literature pertinent to the task, to publish new developments or important new experimental details quickly, to inform others of findings that might alter their own efforts, and eventually to publish all his/her supporting data and conclusions for archival purposes. In the fields of environmental contamination and toxicology, the sum of these concerns and responsibilities is decisively addressed by the uniform, encompassing, and timely publication format of the Springer triumvirate:

Reviews of Environmental Contamination and Toxicology [Vol. 1 through 97 (1962–1986) as Residue Reviews] for detailed review articles concerned with any aspects of chemical contaminants, including pesticides, in the total environment with toxicological considerations and consequences. v

vi

Foreword

Bulletin of Environmental Contamination and Toxicology (Vol. 1 in 1966) for rapid publication of short reports of significant advances and discoveries in the fields of air, soil, water, and food contamination and pollution as well as methodology and other disciplines concerned with the introduction, presence, and effects of toxicants in the total environment. Archives of Environmental Contamination and Toxicology (Vol. 1 in 1973) for important complete articles emphasizing and describing original experimental or theoretical research work pertaining to the scientific aspects of chemical contaminants in the environment. Manuscripts for Reviews and the Archives are in identical formats and are peer reviewed by scientists in the field for adequacy and value; manuscripts for the Bulletin are also reviewed, but are published by photo-offset from camera-ready copy to provide the latest results with minimum delay. The individual editors of these three publications comprise the joint Coordinating Board of Editors with referral within the board of manuscripts submitted to one publication but deemed by major emphasis or length more suitable for one of the others. Coordinating Board of Editors

Preface

The role of Reviews is to publish detailed scientific review articles on all aspects of environmental contamination and associated toxicological consequences. Such articles facilitate the often complex task of accessing and interpreting cogent scientific data within the confines of one or more closely related research fields. In the nearly 50 years since Reviews of Environmental Contamination and Toxicology (formerly Residue Reviews) was first published, the number, scope, and complexity of environmental pollution incidents have grown unabated. During this entire period, the emphasis has been on publishing articles that address the presence and toxicity of environmental contaminants. New research is published each year on a myriad of environmental pollution issues facing people worldwide. This fact, and the routine discovery and reporting of new environmental contamination cases, creates an increasingly important function for Reviews. The staggering volume of scientific literature demands remedy by which data can be synthesized and made available to readers in an abridged form. Reviews addresses this need and provides detailed reviews worldwide to key scientists and science or policy administrators, whether employed by government, universities, or the private sector. There is a panoply of environmental issues and concerns on which many scientists have focused their research in past years. The scope of this list is quite broad, encompassing environmental events globally that affect marine and terrestrial ecosystems; biotic and abiotic environments; impacts on plants, humans, and wildlife; and pollutants, both chemical and radioactive; as well as the ravages of environmental disease in virtually all environmental media (soil, water, air). New or enhanced safety and environmental concerns have emerged in the last decade to be added to incidents covered by the media, studied by scientists, and addressed by governmental and private institutions. Among these are events so striking that they are creating a paradigm shift. Two in particular are at the center of everincreasing media as well as scientific attention: bioterrorism and global warming. Unfortunately, these very worrisome issues are now superimposed on the already extensive list of ongoing environmental challenges. The ultimate role of publishing scientific research is to enhance understanding of the environment in ways that allow the public to be better informed. The term “informed public” as used by Thomas Jefferson in the age of enlightenment vii

viii

Preface

conveyed the thought of soundness and good judgment. In the modern sense, being “well informed” has the narrower meaning of having access to sufficient information. Because the public still gets most of its information on science and technology from TV news and reports, the role for scientists as interpreters and brokers of scientific information to the public will grow rather than diminish. Environmentalism is the newest global political force, resulting in the emergence of multinational consortia to control pollution and the evolution of the environmental ethic. Will the new politics of the 21st century involve a consortium of technologists and environmentalists, or a progressive confrontation? These matters are of genuine concern to governmental agencies and legislative bodies around the world. For those who make the decisions about how our planet is managed, there is an ongoing need for continual surveillance and intelligent controls to avoid endangering the environment, public health, and wildlife. Ensuring safety-in-use of the many chemicals involved in our highly industrialized culture is a dynamic challenge, for the old, established materials are continually being displaced by newly developed molecules more acceptable to federal and state regulatory agencies, public health officials, and environmentalists. Reviews publishes synoptic articles designed to treat the presence, fate, and, if possible, the safety of xenobiotics in any segment of the environment. These reviews can be either general or specific, but properly lie in the domains of analytical chemistry and its methodology, biochemistry, human and animal medicine, legislation, pharmacology, physiology, toxicology, and regulation. Certain affairs in food technology concerned specifically with pesticide and other food-additive problems may also be appropriate. Because manuscripts are published in the order in which they are received in final form, it may seem that some important aspects have been neglected at times. However, these apparent omissions are recognized, and pertinent manuscripts are likely in preparation or planned. The field is so very large and the interests in it are so varied that the editor and the editorial board earnestly solicit authors and suggestions of underrepresented topics to make this international book series yet more useful and worthwhile. Justification for the preparation of any review for this book series is that it deals with some aspect of the many real problems arising from the presence of foreign chemicals in our surroundings. Thus, manuscripts may encompass case studies from any country. Food additives, including pesticides, or their metabolites that may persist into human food and animal feeds are within this scope. Additionally, chemical contamination in any manner of air, water, soil, or plant or animal life is within these objectives and their purview. Manuscripts are often contributed by invitation. However, nominations for new topics or topics in areas that are rapidly advancing are welcome. Preliminary communication with the editor is recommended before volunteered review manuscripts are submitted. Summerfield, NC, USA

David M. Whitacre

Contents

Bioconcentration, Bioaccumulation, and Metabolism of Pesticides in Aquatic Organisms . . . . . . . . . . . . . . . . . . . . . Toshiyuki Katagi

1

Electron Transfer as a Potential Cause of Diacetyl Toxicity in Popcorn Lung Disease . . . . . . . . . . . . . . . . . . . . . . . . . . Peter Kovacic and Andrew L. Cooksy

133

Index . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

149

ix

Contributors

Andrew L. Cooksy Department of Chemistry and Biochemistry, San Diego State University, CA 92182-1030, USA; Centro de Graduados e Investigacion del Instituto Tecnológico de Tijuana, Tijuana, B.C. México, [email protected] Toshiyuki Katagi Environmental Health Science Laboratory, Sumitomo Chemical Co., Ltd., Takarazuka, Hyogo 665-8555, Japan, [email protected] Peter Kovacic Department of Chemistry and Biochemistry, San Diego State University, San Diego, CA 92182-1030, USA, [email protected]

xi

Bioconcentration, Bioaccumulation, and Metabolism of Pesticides in Aquatic Organisms Toshiyuki Katagi

Contents Introduction . . . . . . . . . . . . . . . . . . . . . . . Bioconcentration . . . . . . . . . . . . . . . . . . . . . 2.1 Controlling Factors . . . . . . . . . . . . . . . . . 2.2 Theoretical Approach . . . . . . . . . . . . . . . . 2.3 Pesticides and Other Chemicals . . . . . . . . . . . . 3 Bioaccumulation . . . . . . . . . . . . . . . . . . . . . 3.1 Controlling Factors . . . . . . . . . . . . . . . . . 3.2 Bioaccumulation of Pesticides and Theoretical Approach 4 Metabolism . . . . . . . . . . . . . . . . . . . . . . . . 4.1 Metabolic Patterns . . . . . . . . . . . . . . . . . . 4.2 Enzymes . . . . . . . . . . . . . . . . . . . . . . 4.3 Metabolism of Pesticides and Other Chemicals . . . . . 5 Behavior of Pesticides in Larger-Scale Systems . . . . . . . . 5.1 Model Ecosystems . . . . . . . . . . . . . . . . . 5.2 Microcosms and Mesocosms . . . . . . . . . . . . . 6 Summary . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . .

1 2

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . . .

1 3 3 19 23 42 42 49 51 51 52 79 91 92 96 97 103

1 Introduction From the viewpoint of protecting the natural environment, aquatic ecotoxicological assessment of new pesticides and many existing ones has increasingly become more important. To assess the impact of pesticides on aquatic organisms, international T. Katagi (B) Environmental Health Science Laboratory, Sumitomo Chemical Co., Ltd., Takarazuka, Hyogo, 665-8555, Japan e-mail: [email protected] D.M. Whitacre (ed.), Reviews of Environmental Contamination and Toxicology Volume 204, Reviews of Environmental Contamination and Toxicology 204, C Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-4419-1440-8_1, 

1

2

T. Katagi

authorities (utilizing OECD and USEPA testing guidelines) require completion of many acute and chronic ecotoxicological studies. Among such studies is testing to measure the potential for bioconcentration. In addition, the authorities in these agencies insist that physico-chemical properties and environmental fate be determined for each registered pesticide. The rationale for such testing is based on the concept that, even if used in conformance with good agricultural practices, pesticides may enter surface waters by several routes such as spray drift, surface runoff, and field drainage, and they may be partitioned to bottom sediments (Katagi 2006). The endpoints of such ecotoxicological testing include mortality and effects on hatching, development, and reproduction. Such endpoints are usually expressed as median-lethal or median-effect concentrations (LC50 and EC50 ) and no-observed-effect-concentrations (NOEC); such values can be compared with predicted environmental concentrations in exposure media for purposes of risk assessment (Miyamoto et al. 2008). Because aquatic organisms interact with each other in the food web, knowledge of their tendency to bioconcentrate residues in water and from dietary exposure is important when evaluating real environmental pesticide effects. In general, bioconcentration is the most popular term for describing the process by which pesticides enter organisms directly from water through the gills or through epithelial tissues. In contrast, bioaccumulation includes the effect of dietary uptake through food consumption or intake of bottom sediments (Miyamoto et al. 1990). When the levels of a pesticide, accumulated by organisms, are concentrated through two or more trophic levels in a food web, the process is referred to as biomagnification (Connell 1988). Although information on ecotoxicity and bioconcentration is required and is useful from the regulatory standpoint, such information is also important for its use in evaluating complex processes and their outcomes. Such processes include interactions of pesticide molecules with dissolved and particulate components in water, uptake of pesticides from water into organisms, followed by elimination or metabolic transformation and storage as original or altered residues in tissues (Barron 1990; Connell 1988; Miyamoto et al. 1990). Although considerable amounts of relevant ecotoxicological data for pesticides are available from the ECOTOX database (USEPA 2007), information on pesticides in non-target aquatic organisms, other than fish, is rather limited. Furthermore, research on the metabolic profiles of pesticides in aquatic organisms, together with data on enzymes responsible for pesticide catabolism, has been mainly conducted on fish (James 1994; Schlenk 2005). It is my goal in this chapter to first address the bioconcentration and bioaccumulation profiles of pesticides and selected other chemicals known to exist as pollutants in aquatic organisms (other than in fish). I will do this by addressing the theoretical approach necessary to understand the mechanism by which uptake that results in bioconcentration or bioaccumulation occurs. The methods used to determine bioconcentration will also be discussed. Second, I will summarize the metabolic profiles for several pesticides and other chemicals in aquatic organisms and include a review of the enzymes involved in pesticide metabolism. In this chapter, I will only briefly address the process of biomagnification via food web uptake,

Bioconcentration, Bioaccumulation, and Metabolism

3

and, when I do, it will be in the context of either dietary exposure or higher-tier studies (e.g., model ecosystems). The structures of selected chemicals and pesticides, whose impact on aquatic organisms are discussed in this chapter, are presented in Appendices 1–6.

2 Bioconcentration 2.1 Controlling Factors The bioconcentration of pesticides and other chemicals into aquatic organisms mainly proceeds by passive diffusion through gills, epithelial tissues, or the gastrointestinal tract as shown in Fig. 1a. Bioconcentration is primarily

Fig. 1 Flow and transfer of chemicals in an aquatic system in relation to bioconcentration and bioaccumulation. (a) Conceptual flow diagram. DOM, dissolved organic matter; POM, particulate organic matter. (b) Kinetic model of bioconcentration: C, concentration; k, first-order rate constant; p, parent molecule; m, metabolite; w, water phase; b, biota phase; s, slowly exchangeable peripheral sites in biota; U, uptake; M, metabolism; E, elimination; G, growth

4

T. Katagi

controlled by the physico-chemical properties of the chemicals involved, the physiological disposition of each organism, and the surrounding environmental conditions (Barron 1990; Connell 1988; Landrum and Fisher 1998; Miyamoto et al. 1990). 2.1.1 Physico-chemical Properties Because chemicals first pass through a diffusion barrier, such as mucus and biological membranes, to reach circulating fluids, relative solubilities of such chemicals in water and n-octanol may act as a surrogate for lipids. Another factor, also important, is molecular size, which may simulate partitioning and diffusion processes in lipid-containing tissues. Lipid solubility and molecular size both affect the tendency to bioconcentrate. In general, the bioconcentration factor (BCF) is defined by Eq. (1). Cpb and Cpw constitute the concentrations of a chemical in biota and water, respectively. In the traditional hydrophobicity model, the good relationship that exists between BCF and the n-octanol/water partition coefficient (Kow ) or water solubility (WS) has been established for various combinations of chemical classes and fish. This relationship is described in Eq. (2), wherein a and b are constants (Ellgehausen et al. 1980; Mackay 1982; Neely et al. 1974; Veith et al. 1979): BCF = Cpb /Cpw .

(1)

log BCF = a log Kow (or WS) + b.

(2)

The correlation between BCF and physico-chemical properties has been reported for many aquatic organisms and is presented in Table 1. Rather than Kow and WS, Govers et al. (1984) successfully applied the molecular connectivity index, which is known to encode structural features that correlate with hydrophobicity to describe BCF value of polycyclic aromatic hydrocarbon (PAH) substances in Daphnia pulex. The coefficient of correlation (r2 ) between the two approaches is generally greater than 0.7–0.8, but lower values are observed when more than one chemical class or aquatic species are taken into account (Axelman et al. 1995; Hawker and Connell 1986; Mailhot 1987; Zaroogian et al. 1985). Manthey et al. (1993) reported a moderate correlation for a series of urea herbicides in Chlorella fusca; both hydrophobicity and other factors such as metabolism played a role in this study. The strong relationship between BCF and log Kow is shown clearly in Fig. 2. This plot shows the correlation between fish bioconcentration and log Kow for various pesticides registered in the past decade, and is based primarily on data available from the European Food Safety Authority (EFSA 2008) and the European Commission (EU 2008). In aquatic macrophytes, Turgut (2005) reported the good relationship of the concentration factor of a chemical in shoots of parrot feather with its Kow value; good relationships are also found for the concentration factors in transpiration stream and root in terrestrial plants.

Organochlorines and pesticides Herbicides Insecticides Pesticides Chlorinated aromatics

Organochlorines and pesticides Chlorobenzenes Organochlorines and PAH PAH PAH Pesticides, PAH, PCB PAH

Molluscs (four species)

Crassostrea virginica

10 4

PAH PAH

Pontoporeia hoyi

3.4–6.0 4.5–6.0

4.5–7.0

na 3.3–6.1 0.9–6.7 3.3–5.8

6 7 52 11 6

4.1–5.7 1.8–6.2

PAH

0.69

0.844 log Kow − 1.235, k

0.88

1.1 log Kow − 1.8, k

0.93 0.99

0.89 0.85 0.91 0.89

4.82 3 χc v + 1.276 0.752 log Kow − 0.4362 0.850 log Kow − 1.10 0.7207 log Kow − 0.334

0.38 log Kow + 3.78 0.65 log Kow + 1.8

0.97 0.93

1.76 log Kow − 6.33 0.898 log Kow − 1.315, k

0.29

0.92 0.96 0.91 0.87

r2d

Equation (log BCF =)c

2–6 −0.58 log WS (ppb) + 4.5 3.4–6.0 0.66 log Kow − 0.05 1.7–6.2 0.858 log Kow − 0.808 0.1–4 −0.843 log WS (ppb) + 5.15 Crassostrea virginica 3.9–6.5 0.72 log Kow + 0.41

4.0–7.8

Rangeb

4 22

17

4 6 16 17

34

na

Daphnia magna Daphnia pulex and magna Asellus aquaticus

Lymnaea stagnalis Daphnia pulex

Mytilus edulis

Mytilus edulis

Chemical class(s)

Species

van Hattum and Cid Montanes (1999) Curto et al. (1993) Landrum (1988)

Legierse et al. (1998) Hawker and Connell (1986) Govers et al. (1984) Southworth et al. (1978) Geyer et al. (1991) Axelman et al. (1995)

Zaroogian et al. (1985)

Hawker and Connell (1986) Watanabe et al. (1985) Zaroogian et al. (1985) Geyer et al. (1982) Ernst (1977)

References

Table 1 Correlation of bioconcentration factor (BCF) with physico-chemical properties for pesticides and other chemicals

Bioconcentration, Bioaccumulation, and Metabolism 5

Chlorobenzenes and PCB Pesticides, PCB Chlorobenzenes and PCB Pesticides and organics Urea herbicides Pesticides

Organochlorines and pesticides Aromatics Various chemicals

Oligochaetes (two species) Lemna minor Myriophyllum spicatum

Selenastrum capricornutum

4.1–7.1 2.1–5.2 1–7 >7

5 8 694

0.6–6.4 1.5–4.3 1.7–6.4

0.3–6.6 4.0–8.3

10 9 41 15 8

4.5–7.1

Rangeb

15

na

0.46 log Kow + 2.36 0.77 log Kow − 0.70 +  Fi − 1.37 log Kow + 14.4 +  Fi

0.83 0.73

0.64

0.81 0.56 0.93

0.681 log Kow + 0.164 0.53 log kw + 0.99 0.70 log Kow − 0.26 0.28 log Kow + 2.6

0.91 0.97

0.96

−0.75 (log Kow – 6.84)2 + 6.25 0.491 log Kow + 0.0562 0.98 log Kow – 2.24

r2d

Equation (log BCF =)c

Casserly et al. (1983) Meylan et al. (1999)

Geyer et al. (1984) Manthey et al. (1993) Ellgehausen et al. (1980) Mailhot (1987)

Lockhart et al. (1983) Gobas et al. (1991)

Connell et al. (1988)

References

b Range

of chemicals used to derive the equation. of physico-chemical properties (WS, Kow , kw ) in a logarithm unit. c BCF, bioconcentration factor on a wet weight basis, with k meaning the kinetic value; WS, water solubility with a unit in the parentheses; K , 1-octanol-water ow partition coefficient; kw , HPLC capacity factor; 3 χc v , third-order molecular connectivity index;  Fi, empirical correction factor. d Coefficient of correlation. na: not available. PAH = Polycyclic aromatic hydrocarbons; PCB = Polychlorinated benzenes

a Number

Fish

Scenedesmus acutus

Chlorella fusca

Chemical class(s)

Species

Table 1 (continued)

6 T. Katagi

Bioconcentration, Bioaccumulation, and Metabolism

7

Fig. 2 Correlation of log BCF (bioconcentration factor) and log Kow (octanol to water coefficient) in fish for pesticides developed in the most recent 10-year period

Mackay (1982) was the first to discuss the correlation of fish BCF with Kow . He used the fugacity model at thermodynamic equilibrium, as expressed in Eq. (3), wherein yL , γ , and v define the volume fraction of lipid in the organism, activity coefficient, and phase molar volume, with the subscripts of O (n-octanol phase) and L (lipid phase), respectively: log BCF = log Kow + log [yL (γO vO )/(γL vL )].

(3)

The ratio of γ O /γ L generally does not greatly change; vO is a constant and vL depends on the lipid composition of each organism (Connell 1988). A lower degree of correlation is observed when Eq. (3) is applied to all of the BCF data from many different species together, which may be partly accounted for by the span of the different vL values encountered. If n-octanol is a valid surrogate for lipid, when the (γ O vO )/(γ L vL ) term is at unity, “a” and “b” in Eq. (2) become 1 and log yL , respectively. However, the “a” value practically approaches unity (Table 1). The primary barrier for a chemical involved in passive uptake from water is the biological membrane, which consists basically of lipid bilayers, e.g., as in gill epithelium. Steric parameters such as molecular size and shape sometimes also become important factors in passive uptake (Barron 1990; Landrum and Fisher

8

T. Katagi

1998). Shaw and Connell (1984) did not demonstrate a correlation between BCFs of polychlorinated biphenyl (PCB) compounds normalized to that of 2 ,3,4,4 ,5 pentachlorobiphenyl, with their log Kow values, in mullet and the polychaete Capitella capitata. By introducing a steric effect coefficient (SEC), which effect varies with the substitution pattern of chlorine atoms, a better correlation between BCF and log (Kow × SEC) is produced. A clearer effect of molecular shape on BCF was demonstrated by Opperhulzen et al. (1985) for the bioconcentration of aromatic hydrocarbons in guppys. When the log Kow value exceeded 4, the uptake rate plateaued and the rate of elimination decreased. The BCF value was nearly constant at log Kow > 4–6, and the more hydrophobic compounds, such as hepta- and octa-chloronaphthalenes, did not bioconcentrate. By considering the average packing order in natural lipid membranes, these authors proposed a threshold cross section of 9.5 Å for molecules that can successfully pass through the membrane. A similar size effect was reported by Stange and Swackhamer (1994) for the bioconcentration of PCBs in three algae. The speciesspecific log BCF vs. log Kow plot showed a plateau near a log Kow value of 6, wherein the molecular cross section was estimated to be approximately 9 Å. The normalization of BCF with phospholipid content reduced this specificity; therefore, membrane permeability was considered to be important for uptake. Curto et al. (1993) reported a deviation from the log BCF vs. log Kow relationship for PAHs having five to six aromatic rings in the isopod Asellus aquaticus; these authors speculated that the reduced membrane permeability resulted from steric hindrance. The hydrophobicity of a chemical is well known to correlate with molecular surface area and volume (Katagi et al. 1995). Del Vento and Dachs (2002) reported the parabolic nature of the BCF values of PCB congeners in bacteria and phytoplankton, when data were plotted against the total surface area (TSA) of a molecule. The maximal BCF was observed at the TSA value of 250–270 Å2 , which corresponds to the threshold diameter of 8.9–9.3 Å, when a spherical shape of a molecule is assumed. This diameter is very close to the threshold cross section of 9.5 Å reported by Opperhulzen et al. (1985). By examining the parabolic nature of log BCF vs. log Kow plots for chemicals having various flexible structures, Dimitrov et al. (2002) proposed a maximal cross-sectional diameter (Dmax ) of a molecule that was based on molecular orbital calculations of energetically favorable conformers. Chemicals having a Dmax of less than 15 Å showed a high BCF value with an increase of log Kow , whereas those with Dmax of >15 Å were found to be accumulated up to a log BCF value of 3.3 at most. This threshold value of 15 Å was close to one half of the lipid-bilayer thickness in biological membranes. As with an increase of a molecular dimension, the solubility of a chemical in a lipid phase is considered to change the structural orientation of the molecule. The reason for this is because solvation of a chemical by lipid molecules requires more energy to structurally re-orient the molecule for fit than occurs with small molecules such as n-octanol. Florey-Higgins theory shows that the solubility of a chemical

Bioconcentration, Bioaccumulation, and Metabolism

9

is a function of molar volume and solvent, as well as solute–solvent interactions. By examining the solubilities (SL ) of 79 organic chemicals to triolein and dimyristoyl phosphatidylcholine in relation to log Kow , Chessells et al. (1992) obtained the parabolic relationship of log SL = − 0.08 + 0.52 log Kow − 0.09 (log Kow )2 (r = 0.78). When chemical concentration in fish was expressed by a polynomial function of log Kow (Connell and Hawker 1988), log BCF better correlated with log SL and n-octanol was found not to be a good surrogate for lipids over a wide range of log Kow . Isomerism of a chemical sometimes plays a role in bioconcentration. Hexachlorocyclohexane (HCH) has four isomers (α–δ), and its γ -isomer in known as lindane (30). BCFs in clam varied by isomer as follows: δ > α > β ≈ γ . However, the order for elimination was different: γ > α » δ ≈ β (Yamato et al. 1983). The corresponding values in the mussel Mytilus edulis (Ernst 1979) and bluegreen alga Anabaena sp. (Mathur and Saxena 1986) showed yet a different order; these differences indicate that species differences exist in the bioconcentration of these isomers. Species differences existed in the uptake of chlordane (24) isomers (Moore et al. 1977), and the enantio-selective metabolism of γ (or trans)-isomer accounted for the major difference in Mysis relicta between the isomers (Warner and Wong 2006). In oysters, the enantio-selective bioconcentration of flucythrinate (86) has also been addressed (Schimmel et al. 1983). The α (or I)- and β (or II)-isomers of endosulfan (27) differ in their conformation in the cycloheptyl moiety. The β-isomer has been reported to bioconcentrate more than the α-isomer in Daphnia magna (DeLorenzo et al. 2002) and in crayfish Procambarus clarkii (Naqvi and Newton 1990), probably because the β-isomer has lower metabolic transformation to the corresponding sulfate. However, the α-isomer was reported to be more highly bioconcentrated in algae (DeLorenzo et al. 2002; Rao and Lal 1987).

2.1.2 Aquatic Organisms Among the range of physiological conditions that exist in aquatic organisms, lipid content is considered to be the most important determinant for bioconcentration. The influence of lipid content on BCF is followed by extent of metabolism and excretion and developmental stage of the tested organism. Lipids are normally extracted from aquatic organisms using a chloroform–methanol mixture (Folch et al. 1957; Bligh and Dyer 1959). Chemical characterization and identification of each lipid component is usually performed using TLC (thin layer chromatography), HPLC (high-pressure liquid chromatography), GC (gas chromatography), and MS (mass spectrometry). Instead of chloroform, dichloromethane may be used to extract lipids (Booij and van den Berg 1994). Gardner et al. (1985) further developed microextraction techniques to deal with tissue samples weighing less than 10 mg. A summary of the total lipid content, on a dry or wet weight basis, of aquatic organisms is presented in Table 2.

10

T. Katagi Table 2 Lipid content of aquatic species

Species

Lipid content (%)a

References

Mussel

0.6–3.9 (w)

Oyster Clam Cockle Scallop Snail

1.5–2.8 (w) 0.8–2.3 (w) 0.6 (w) 0.5–1.1 (w) 0.4–4.4 (w)

Green alga

10–45 (d)

Blue-green alga

2.2–22 (d)

Golden alga Brown alga Diatom

7–41 (d) 1.2–5.3 (d) 7.6–24 (d)

Aquatic macrophyte Water flea

4.0–24 (d), 0.2 (w)

Bedford and Zabik (1973); Booij and van den Berg (1994) Copeman and Parrish (2004); Ernst (1979), King et al. (1990) Lin et al. (2003); Murphy et al. (2002); Pirini et al. (2000) Renberg et al. (1978); Serrano et al. (1997a) King et al. (1990) Copeman and Parrish (2004); King et al. (1990) Copeman and Parrish (2004) Copeman and Parrish (2004); King et al. (1990) Lalah et al. (2003); Legierse et al. (1998); Takimoto et al. (1987a) Thybaud and Caquet (1991) Borowitzka (1988); Canton et al. (1977); Choi et al. (1987) Halling-Sørensen et al. (2000); Kent and Currie (1995) Koelmans and Sánchez Jiménez (1994); Lu and Metcalf (1975) Rathore et al. (1993); Stange and Swackhamer (1994) Borowitzka (1988); Rathore et al. (1993) Stange and Swackhamer (1994); Sukhija et al. (1979) Borowitzka (1988) Dembitsky et al. (1990) Borowitzka (1988); Darley (1977) Stange and Swackhamer (1994) Dembitsky et al. (1992); Gobas et al. (1991)

Shrimp and crab Isopod Amphipod

Copepod Rotifer Midge Mayfly

5.5–23 (d), 0.3–2.0 (w)

Bychek and Gushchina (1999); Cowgill et al. (1984) Heisig-Gunkel and Gunkel (1982); Herbes and Allen (1983) Lu and Metcalf (1975) 10–40 (d), 1.0–2.0 (w) Cavaletto and Gardner (1988); King et al. (1990) Warner and Wong (2006) 0.3–0.7 (w) Gaskell et al. (2007); Van Hattum and Cid Montanes (1999) 2.4–43 (d), 2.3–5.3 (w) Arts et al. (1995); Cavaletto and Gardner (1988) Gardner et al. (1985); Herbes and Allen (1983) Meyer et al. (2000); Nebeker et al. (1989); Quigley et al. (1989) 8.3 (d) Herbes and Allen (1983) 1.1–1.5 (w) Guisande and Serrano (1989) 1.1 (w) Gaskell et al. (2007) 2–20 (d), 1.4–2.7 (w) Cavaletto and Gardner (1988); Drouillard et al. (1996) Meyer et al. (2000)

Bioconcentration, Bioaccumulation, and Metabolism

11

Table 2 (continued) Species

Lipid content

Caddisfly Polychaete

8.0–32.5 (d) 0.85–1.7 (w)

Oligochaete

aw

(%)a

References

Herbes and Allen (1983); Meyer et al. (2000) Booij and van den Berg (1994); Driscoll and McElroy (1996) 8–25.5 (d), 1.1–5.2 (w) Egeler et al. (1997); Cavaletto and Gardner (1988) Gardner et al. (1985); Gaskell et al. (2007) Nebeker et al. (1989); Whitten and Goodnight (1966) You et al. (2006)

and d mean the wet and dry weight basis in the whole body, respectively.

The lipid content of aquatic organisms greatly varies with species, but generally ranges from 1% to 5% on a wet weight basis. HPLC analysis of fresh muscle from eight fish species showed phospholipids (PL) and triacylglycerols (TAG) as main components, followed by sterols (ST) and free fatty acids (FA) (Ewald and Larsson 1994). Since the PL content explained only 31% of the variance in the tissue concentration of 2,2 ,4,4 -tetrachlorobiphenyl, not only the lipid content but also the chemical structure of the FA portion in lipids were considered to be important for bioconcentration. Lipids have been extensively characterized for shellfish in the mollusca phylum. Murphy et al. (2002) reported that PL (57–79%) and TAG (10–25%) comprise the main lipid components in mussels, followed by free FA and ST, with trace amounts existing of wax esters (<0.3%). Among the 50 FAs and fatty aldehydes released by transmethylation, GC-FID (flame ionization detection) and MS analyses have revealed that C16:0 , C20:5n−3 (eicosapentaenoic acid, EPA), and C22:6n−3 (docosahexenoic acid, DHA) are the main components of polyunsaturated FAs (PUFA). The saturated palmitic acid is considered to originate from stored glycogen, whereas the unsaturated FAs may derive from dietary algae. About 20 STs, of which cholesterol was dominant, were detected by GCFID. Similar profiles exist for FA and ST, and were reported for oyster, clam, cockle, and scallop (Copeman and Parrish 2004; King et al. 1990; Pirini et al. 2000), but with lower amounts of hydrocarbons, steryl esters, diacylglycerols, and methyl ketones. Recently, LC (liquid chromatography)-MS in electrospray ionization mode, coupled with solid-phase extraction, has been successfully applied to examine the FA, TAG, and PL composition of these species (Lacaze et al. 2007). The hydrophilic head group of PL was also examined in the mussel Mytilus edulis by HPLC and GC (Lin et al. 2003); the main components identified were phosphatidylcholine (PC; 47–72% of PL) and phosphatidylethanolamine (PE; 7–40%). Seasonal variance in the content of PL appears to be smaller than that of TAG, which may be accounted for by the fact that PL is a structural component of cell membrane, and, in contrast, TAG is used for energy storage. Furthermore, in cold weather, either FA unsaturation or content of phosphatidyl-ethanolamines (PE), -inositols (PI), and -serines (PS) increased in PL; these factors are lined with membrane fluidity (Lin et al. 2003; Pirini et al. 2000). Increased FA unsaturation

12

T. Katagi

was induced when a freshwater mussel was acclimatized at a lower temperature (Boryslawskyj et al. 1988). The lipid composition in algae has been extensively investigated. Chlorophyll a and β-carotene are widely distributed in various algae, among which major ones are cyanophyta (blue-green algae), chlorophyta (green algae), bacillariophyta (diatoms), and rhodophyta (red algae) (Chapman and Chapman 1973). The blue-green algae are prokaryota; the others are eukaryota. There are several excellent reviews on the biochemical composition of lipids in algae (Ahlgren et al. 1990; Benson and Shibuya 1962; Borowitzka 1988; Lewin 1974; Miller 1962; Wood 1974), especially for green algae (Thompson 1996), brown algae (Dembitsky et al. 1990), and diatoms (Darley 1977). The main lipid components in algae are monogalactosyl (MGDG) and digalactosyl (DGDG) diglycerides, which, along with TAG, PL, and ST, are characteristic of photosynthetic species. Sulfoquinovosyldiacylglycerol (SL) is also a glycolipid characteristic of photosynthetic species, and has been reported as the single glycolipid in the plasma membrane of green alga Dunaliella salina. Crude lipids that contain pigments can be extracted and fractionated using column chromatography or TLC, or better by using a silica Sep-pak to characterize each lipid fraction (Tatsuzawa et al. 1996). The algal FAs are the even-numbered C12 –C24 acids, mainly C16 and C18 , which are biosynthesized via the β-oxidative fatty acid cycle. These FAs have a considerable degree of unsaturation and their unique distribution in each phylum is a known characteristic. The degree of FA unsaturation varies by lipid fraction; more saturation exists in neutral lipids of the green alga (Scenedesmus obliquus) such as TAG, and in glycolipids than exists in PL (Choi et al. 1987). Flagellates have a higher content of α-linoleic acid, EPA, and DHA, the latter two of which are common to fish and shellfish (Ahlgren et al. 1990). The distribution of neutral-, glyco-, and phospho-lipids is known to greatly vary with algal growth (Halling-Sørensen et al. 2000; Stange and Swackhamer 1994). PI and PE are major components in PL, and are mainly located in the extra-chloroplast membranes. A great variety of sterols such as sitosterol, fucosterol, and cholesterol are observed in algae. Hydrocarbons are the minor components, but with predominance of n-heneicosahexaene (C21:6 ), in many algae. Diatoms are unique in having siliceous cell walls and a large vacuole, but the nature of major lipid components in diatoms are similar to those of green algae and higher plants (Darley 1977). The lipids that occur in blue-green algae, Anabaena doliolum and Anacystis nidulans, mainly consist of polar species such as glycolipids and phosphatidyl glycerol (PG); higher levels of SL and PG were detected in blue-green than in green alga Chlorella vulgaris (Rathore et al. 1993; Sukhija et al. 1979). Brown algae retain MGDG, DGDG, and SL as glycolipids, with the proportion of general lipids and PLs being higher than those in green and red algae (Dembitsky et al. 1990). Dembitsky et al. (1992) reported that several freshwater macrophytes contain neutral- (34–49%), glyco- (34–81%), and phospholipids (14–24%), and PC is the main component in PL, together with PG and PI, which is characteristic of green grasses; macrophytes, however, do not contain diacyltrimethylhomoserine widely distributed in marine Chlorophyta macrophytes.

Bioconcentration, Bioaccumulation, and Metabolism

13

It is also known that certain environmental factors may influence the lipid content and composition of algae (Borowitzka 1988). In mollusca, temperature is one of the most important factors that determine the FA content profile. Acclimatization at a lower temperature caused an increased content of PUFA, which resulted in an increase of membrane fluidity (Thompson 1996). A pH effect on PL composition in Chlamydomonas sp. has also been reported by Tatsuzawa et al. (1996). The algae were adapted to a lower pH, not only through higher saturation in the FA portion of the lipids to reduce membrane fluidity but also through more storage of TAG as an energy source to cope with an osmotic imbalance. Less is known about the biochemical composition of other aquatic organisms than for mollusca and algae. Heisig-Gunkel and Gunkel (1982) examined the contents of fat and protein in Daphnia pilicaria at three different temperatures. The fat and protein content varied according to the acclimatization period, and fat content slightly increased as temperature increased. Neutral lipids such as TAG and ST (52–59%) are the main components in D. magna, followed by more polar lipids and glycolipids (Cowgill et al. 1984). The content of TAG as an energy store was maximized in juveniles and decreased during somatic growth, but recovered in adults, wherein wax ester content increased. Free ST content gradually decreased as development proceeded, probably from synthesis of steroids and molting hormones (Bychek and Gushchina 1999). The even-carbon numbered C16 and C18 FAs, with or without unsaturation, predominated; the lower amounts of odd-carbon numbered FAs possibly originated from bacteria in the daphnia’s intestine. The degree of unsaturation in FA increased with developmental stage, and with a decrease of C18:1 consumed as an energy substrate. The lipid content of various freshwater aquatic organisms have been compiled through modeling PCB contamination in the Housatonic River; lipid content generally ranged from 1% to 4% on a wet weight basis (Donigian et al. 2004). Seasonal variation in lipid content was reported for freshwater aquatic organisms by some researchers. The seasonal and spatial changes in TAG content of the benthic amphipod Pontoporeia hoyi are caused by a spring bloom of large diatoms, and the much higher content in adult females is related to the reproductive process (Cavaletto and Gardner 1988; Quigley et al. 1989). A similar seasonal variation in lipid content was observed in two other amphipods, Hyalella azteca and Gammarus lucustris (Arts et al. 1995). Cavaletto and Gardner (1988) found that PL, ST, and TAG are the main lipid components in Tubificidae, Chironomidae, Mysis, and Dreissena. Similar to the amphipods, Gammarus pulex retained higher lipid content in the spring, but no statistically significant variation was observed for other species that had TAG as a main lipid component, e.g., the mayfly Ephemeroptera and caddisfly Trichoptera (Meyer et al. 2000). The FA composition of TAG and PL has been examined by GC-MS in the larvae of benthic invertebrates (Sushchick et al. 2003). Mono- and polysaturated C16 FAs comprise the main component of TAG, indicating that diatoms are in the diet of these invertebrates. More branched-chain odd-numbered FAs (C15 and C17 ) serve as bacterial markers and are detected in mayfly larvae; such FAs are considered to originate from intensive ingestion of detritus and sediment particles. The lipid content of rotifers is approximately 1% and is considered to have primarily a

14

T. Katagi

structural rather than energy storage function; in rotifers, carbohydrates constitute the main reservoir of energy storage (Guisande and Serrano 1989). Many biological parameters including size, biochemical constituents, and metabolic activity will change as organisms grow, and may, therefore, affect uptake, elimination, and bioconcentration profiles. Veith et al. (1979) examined the effect of developmental stage on bioconcentration of hexachlorobenzene (31) in minnows; in this study no significant effect was related to the age of fish. In contrast, algae grow very fast and the rate of chemical uptake is expected to depend on growth stage. Nakamura and Mochida (1988) examined the uptake kinetics of organophosphorus pesticides in the green alga Selenastrum capricornutum, and showed a 1.3- to 2.5-fold increase in uptake rate when cell number was at a steady state in contrast with an exponential growth stage. Algal growth rate significantly affected the bioconcentration of PCB congeners in Scenedesmus sp. (Swackhamer and Skoglund 1993). The log BCF vs. log Kow plot showed a downward curvature at log Kow values >6 for a short incubation period; this trend was more pronounced for algae with higher growth rates. Slower partition of more hydrophobic chemicals into algae, as compared with their growth, may result in dilution from the increased biomass. As previously mentioned, the difference in organismal lipid content sometimes accounts for age-dependent BCF. About fourfold more bioconcentration of pentachlorophenol (32) was reported for D. magna adults in a 24-hr static exposure than occurred for neonates (Kukkonen and Oikari 1988). Since the metabolic activity necessary to form the sulfate conjugate is very similar between adults and neonates, the difference in lipid content was regarded to account for the difference in BCF values. More trans-chlordane (24) was bioconcentrated in midge adults (Chironomus decorus) than in larvae; constant residues were reported in 2nd to 4th instars (Harkey and Klaine 1992). In contrast, the stepwise decrease in bioconcentration of chlorpyrifos (47) from the 2nd to 4th instars was observed in C. riparius (Buchwalter et al. 2004). The biotransformation rate was independent of instar stage; the reduced uptake through the body surface per body weight, in later-stage organisms with smaller surface area to volume ratios, may account for the difference. A similar size effect on uptake of PAHs was briefly reported for mayfly nymphs (Hexagenia limbata) by Stehly et al. (1990). Size effect also exists for bioconcentration in algae. As the size of green algae and diatoms decrease, their surface to volume ratio increases, which resulted in reports of higher uptake of atrazine (127) and fenitrothion (41) (Kent and Currie 1995; Tang et al. 1998a; Weiner et al. 2004). Growth is also an important factor when discussing metabolism in the context of bioconcentration. By comparing experimental BCF values of chlorinated anilines in the guppy with the predicted ones derived from BCF data on non-biotransformed chlorobenzenes, De Wolf et al. (1992) found that lower observed values are caused by biotransformation of anilines in the fish via N-acetylation. Serrano et al. (1997b) examined the bioconcentration of organophosphorus pesticides in Mytilus galloprovincialis for 35 days and found much lower BCF values than expected from the predictive equation (Connell 1988). This may have resulted from metabolic biotransformation of the pesticides during the experiment. More concrete evidence for an effect on bioconcentration from metabolism was demonstrated in Chironomus

Bioconcentration, Bioaccumulation, and Metabolism

15

R riparius and Lumbriculus variegatus for two polycyclic musk fragrances, Tonalid R  and Galaxolide (Artola-Garicano et al. 2003). The observed BCF values in the midge were much lower than ones calculated for non-metabolized chlorobenzenes (Roghair et al. 1992); the BCF values approached the predicted ones when the midges were exposed in the presence of piperonyl butoxide (PBO). Since PBO is known to be an inhibitor of cytochrome P450 enzymes, its effect on metabolic transformation is a key factor that influences bioconcentration rate. In contrast, observed and predicted values are almost the same in the oligochaete, and significant metabolism is unlikely to be involved. Because most chemicals that are bioconcentrated are deposited in the lipid phase of aquatic organisms, and tissues and organelles have different levels and types of lipids, one normally assumes that the absorbed chemical is heterogeneously distributed in organisms. However, detailed analysis of chemical residues in mollusca tissues, such as clam, mussel, and oysters, indicates that pesticides are generally distributed in visceral mass, including digestive glands and gonads, rather than in mantle and gill (Bedford and Zabik 1973; Brodtmann 1970; Rajendran and Venugopalan 1991; Sathe et al. 2005; Uno et al. 2001). Takimoto et al. (1987a) examined the distribution of radioactivity from uptake of 14 C-fenitrothion (41) in the snail Physa acuta by autoradiographic techniques and found that most 14 C resided in liver. Few investigations exist on the distribution of pesticides in algae. However, the algal surface structure seems to be an important factor that governs distribution. Vance and Maki (1976) have separately analyzed residues of dibrom (63) in the green alga Stigeoclonium pachydermum; these authors found that most of (63) was distributed in the cell wall fraction as a result of a thick mucilage coat that covers this organism’s filaments. In contrast, Chlamydomonas and Dunaliella sp. were found to take up the α-isomer of HCH into the cell interior, which is rich in lipids (Canton et al. 1977). Through the investigation of benzo[a]pyrene (7) uptake by periphyton communities, including desmids and diatoms collected from streams, Bruno et al. (1981) concluded, by using autoradiography, that most of benzo[a]pyrene (7) was distributed in the extensive mucilaginous sheaths that surround the desmid cells. The absorption and translocation of pesticides have been studied in aquatic macrophytes and terrestrial plants by using radiolabeled pesticides. The extent of acropetal and basipetal movement in plants seems to depend not only on pesticide type but also on plant species involved (Anderson et al. 1981; Frank and Hodgson 1964; Funderburk and Lawrence 1963; Hinman and Klaine 1992; Thomas and Seaman 1968). Only limited work has been reported on chemical distribution in other aquatic organisms. Crosby and Tucker (1971) reported some distribution of DDT (33) in the carapaces of D. magna. Derr and Zabik (1972) found that up to 30% of DDE (34) bioconcentrated by Chironomus tentans was distributed in egg mass, with much less distributed to the exuviae. Autoradiography has clearly shown that absorbed pesticides in amphipods and ostracods are concentrated in lipid material (Arts et al. 1995; Kawatski and Schmulbach 1970). Existing information strongly suggests that, in general, absorbed xenobiotic chemicals are usually distributed in lipid-rich tissues and organs, unless the organism has an impervious surface (skin, integument, etc.) that acts as a barrier to entry.

16

T. Katagi

2.1.3 Environment Temperature and water chemistry, such as salinity, pH, and content of dissolved or adsorbed particulate organic matter, also may affect bioconcentration. When the temperature increased from 5◦ C to 20◦ C, BCF values of hexachlorobenzene (31) increased by a factor of 7–10 in three fish species (Veith et al. 1979). Through the thermodynamic analysis of the bioconcentration of several chlorobenzenes in guppies at different temperatures, Opperhulzen et al. (1988) demonstrated that n-octanol is a poor surrogate for fish lipid. The transfer process from water to fish was governed by a positive entropy change from loss of structuring of water molecules that surrounded chlorobenzene, whereas the transfer process for n-octanol was accompanied by exothermic enthalpy changes and small entropy changes. A similar temperature dependency for bioconcentration was observed in algae. Several green algae species bioconcentrated higher amounts of ioxynil (145) and 2,4-D (75), when temperatures were higher (Neumann et al. 1987; Valentine and Bingham 1974). Koelmans and Sánchez Jiménez (1994) reported on the bioconcentration of chlorobenzenes in Scenedesmus sp. and its relationship to an entropy gain that dominated the transfer process from water to algae, and found it to be similar to what was observed in fish. A slight increase in the BCF for pentachlorophenol (32) was reported in zebra mussel when temperatures were higher; however, BCF changes for dieldrin (22), lindane (30), and pentachlorophenol (32) were insignificant in other mussels (Boryslawskyj et al. 1988; Ernst 1979; Fisher et al. 1999). Therefore, the temperature effect on BCF appears to be less important for bivalva. Ambient temperature affected the lipid content and respiration rate in water fleas, which was considered to partly influence rates of bioconcentration. At a higher temperature, a higher correlation was observed between BCF and the lipid content in D. magna for atrazine (127); in addition, there was a temperature-dependent increase in the relative amount of lipid vs. protein at a higher temperature (HeisigGunkel and Gunkel 1982). More DDE (34) was bioconcentrated in D. pulex at higher temperatures (Nawaz and Kirk 1995). The authors believed that at higher temperatures respiration increased, and the enhanced gas exchange at the tissue surface of the branchial chamber reduced the thickness of a diffusive boundary layer, which resulted in the facilitated uptake. Abiotic and metabolic transformation sometimes encumbers determining how much BCF values are affected by temperature. Carbaryl (101) is more susceptible to hydrolysis and metabolism at higher pH and temperature. Therefore, carbaryl BCF values were not significantly changed in Chironomus riparius (Lohner and Fisher 1990). Parathion (39) was metabolically transformed to the corresponding oxon and phenol in the midge, which resulted in reduced bioconcentration of (39) (Lydy et al. 1990). The effect of illumination levels on bioconcentration was reported in algae and aquatic macrophytes. Weinberger and Greenhalgh (1985) observed higher uptake of aminocarb (98) by axenic aquatic hornwort (Ceratophyllum demersum), and much reduced uptake by dead organisms, which indicated the presence of active uptake processes in macrophytes. Similarly, more glufosinate (73) was taken up under similar circumstances by Lemna gibba (Ullrich et al. 1990). In contrast, ioxynil (145) was absorbed fourfold less by the green alga Ankistrodesmus braunii under bright

Bioconcentration, Bioaccumulation, and Metabolism

17

light conditions; this may be related to the change of membrane potential caused by more intense lighting (Neumann et al. 1987). For chemicals having a dissociable functional group, bioconcentration may be affected by the pH level in the medium and also by the internal or cytoplasmic pH of an aquatic organism. The pKa value of pentachlorophenol (32) is approximately 5, and the dissociated chemical species predominates under ambient environmental conditions. Therefore, bioconcentration is generally independent of the pH of the medium, as has been observed for bioconcentration in zebra mussels (Fisher et al. 1999). In contrast, the bioconcentration of ioxynil (145) in the green alga (Ankistrodesmus sp.), which has a pKa value of 3.96, showed a clear pH-dependency at pH 5.5–7.3 (Neumann et al. 1987). There was insignificant uptake at pH 7.3; hence, pH-dependency under acidic conditions portends more favorable partitioning of the undissociated chemical species into this alga. Similar pH profiles were reported for the bioconcentration of 2,4,5-trichlorophenol in Lemna gibba (Tront and Saunders 2006). However, the degree of dissociation is not always the dominant factor that controls uptake of a chemical. Kenney-Wallace and Blackman (1972) examined uptake by Lemna minor of several benzoic acids, each with a different chlorine substitution; these authors reported a 90-fold difference in relative uptake, which could not be explained on the basis of proportion of undissociated species at the tested pH (1000-fold). It was concluded that hydrophobicity most likely governed the partition. Glufosinate (73) was taken up not only by passive diffusion but also by an active process that involved a proton co-transport mechanism in L. gibba (Ullrich et al. 1990). Furthermore, bioconcentration pH-dependency seems to differ among algae species. More uptake in Scenedesmus sp. was observed under acid conditions for 2,4-D (75), which has a pKa value of 3; however, no such dependency was noted in Chlorella and Chlamydomonas sp. (Valentine and Bingham 1974). Küsel et al. (1990) monitored the in vivo cytoplasmic pH value in Chlorella sp. by NMR; by using the 31 P signal of an inorganic phosphate, the pH was almost constant at 7.2–7.8 in air, even at an external pH of 3–10. The dissociation constant (pKa ) of sulfonylurea herbicides is approximately 3–5. Therefore, a higher uptake of undissociated chemical species through biological membranes is expected at a lower pH. Such a pH-dependent BCF profile was observed in Chlorella fusca with chlorsulfuron (122) and metsulfuron-methyl (123), and was determined by trapping the ionized sulfonylurea molecule at the cytoplasmic pH (Fahl et al. 1995). Similarly, less non-dissociative lindane (30) was taken up by midge larvae (Chironomus riparius) at a lower pH (Fisher 1985). Wildi et al. (1994) reported a similar profile in bioconcentration of pyrene (5) by midge larvae, and speculated that the mucus that covers the body surface at lower pH levels may reduce diffusion of a chemical into the larvae. Micro-sensors have been applied to study the gut environment of larvae of the midge, Chironomus plumosus (Stief and Eller 2006). In vivo measurements, using such sensors, showed lower oxygen concentration and redox potential and slightly higher pH in the food bolus of the larvae. Neither diffusion of O2 from hemolymph nor O2 uptake during ingestion of food was sufficient to oxygenate the food bolus; hence, aerobic metabolism in the gut was considered to be unlikely. Hardness of water as a medium was found to affect both uptake and elimination processes of

18

T. Katagi

3-trifluoromethyl-4-nitrophenol (148) in C. tentans larvae, which was explained by the difference in dissociation rendered by the pH change linked to water hardness (Kawatski and Bittner 1975). In natural waters, there are usually agents, e.g., organic carbonaceous matter or dissolved, colloidal, or suspended particulates, that alter the bioavailability of chemicals to aquatic organisms (Fig 1a; Farrington 1991; Haitzer et al. 1998; Katagi 2006). Humic substances including fulvic, humic, and hydrophilic acids are usually the predominant constituents of dissolved organic matter (DOM). When measuring or estimating the bioconcentration or bioaccumulation potential of a chemical in sediment-dwelling organisms, the type and content of colloidal organic matter that exists in interstitial water should be considered. If an adsorption/desorption equilibrium has been reached between a chemical and DOM, the apparent BCF in the presence of DOM (BCFd ) can be expressed as BCFd = BCF0 /(1 + m × foc × α × Kdoc ),

(4)

where BCF0 = the BCF value in the absence of DOM; M = concentration of DOM (kg L−1 ); foc = organic carbon fraction in DOM (kg DOM kg−1 organic C); α = a factor measuring the degree of bioavailability of a DOM-associated chemical to organism; α = 1 means no bioavailability; Kdoc = partition coefficient (L kg−1 organic C) between dissolved organic carbon and water. The presence of dissolved humic acids reduced BCF of benzo[a]pyrene (7) from water for D. magna (McCarthy 1983). Kukkonen and Oikari (1991) have examined the 24-hr bioconcentration of organic pollutants including (7) in D. magna, in 20 natural Finnish freshwater bodies, having different humic composition. BCF values gradually decreased with increasing concentration of dissolved organic carbon (DOC); Eq. (4), where α = 1 illustrates such a profile. Kukkonen and Oikari (1991) also reported good correlation between BCF and DOM content that had been enriched with aromatic constituents. Since the Kdoc value is larger for most hydrophobic chemicals, it is anticipated that DOM will retard uptake of pyrethroids. Day (1991) reported that the presence of DOC at 10–15 ppm caused about a fourfold reduction of BCF in D. magna for deltamethrin (83), lambda-cyhalothrin (84), and fenvalerate (85); approximately 20–40% of these pesticides were present as associated forms, based on Kdoc values of approximately 105 L kg−1 . In bioconcentration and 96-hr acute toxicity studies with D. magna and Ceriodaphnia dubia, a lack of bioavailability of associated forms was recently demonstrated by Yang et al. (2006). By analyzing the BCF and LC50 values with Eq. (4), the α values were statistically estimated to be equal to unity. Haitzer et al. (1998) reviewed the effects of DOM on bioconcentration rates in aquatic organisms and reported two possible mechanisms. Reduced

Bioconcentration, Bioaccumulation, and Metabolism

19

bioconcentration was considered to originate from less chemical dissolved in water, because the chemical was associated with DOM, whereas at lower DOM levels (<10 ppm) bioconcentration was enhanced, possibly because of increased transfer of chemical to organisms. Haitzer et al. (1998) emphasized the importance of the chemical composition of humic substances, as well as contact period with a chemical, to explain degree of association. In fact, Schramm et al. (1998) demonstrated less retardation of BCF than expected from Eq. (4), for various chemicals, in the presence of aquatic humic acids that are more hydrophilic than terrestrial ones. In one case, Wang and Lay (1989) demonstrated lower BCF values for the hydrophilic salicylic acid in Lemna minor and two green algae in the presence of river fulvic acid; presence of fulvic acid also resulted in reduced acute and chronic toxicity of salicylic acid to D. magna. Moreover, much lower BCF values of triorganotin compounds in Chironomus riparius were reported in the presence of Aldrich humic acid (Looser et al. 2000). In contrast, a slightly enhanced bioconcentration of atrazine (127) was observed for D. magna in natural freshwaters having a DOC content of less than 10 ppm (Nikkilä et al. 2001). Since fulvic acid is an aromatic polymer that has carboxyl and hydroxyl groups, either the π–π interaction between aromatic rings or hydrogen bonding might operate to enhance bioconcentration, whereas electrostatic repulsion resulted in reduced bioconcentration. In another study, DOC content in an algal medium remained higher than that in natural water during incubation, and it was for this reason that lower BCF values than expected (from the log BCF vs. log Kow regression) were observed for the more hydrophobic PCB congeners (Stange and Swackhamer 1994). In shallow water bodies, such as freshwater streams, bottom sediments play a role as xenobiotic chemical adsorbents (Katagi 2006). If adsorption obeys a linear isotherm, with an organic carbon-normalized coefficient of Koc , the denominator of Eq. (4) becomes “1 + foc × Koc ”, and as a result less bioconcentration would occur for aquatic organisms exposed to overlying water. Kusk (1996) has reported a reduction in BCF for D. magna for pirimicarb (108) from 50 to 31–37, in the presence of sediment. In contrast, the bioconcentration of terbutryn (130) and fluridone (131) in Chironomus tentans was not affected by sediment (Muir et al. 1982).

2.2 Theoretical Approach The bioconcentration rate of a chemical into an organism from water is mainly governed by uptake and elimination processes. However, additional processes such as metabolic transformation (Cowan-Ellsberry et al. 2008; Lydy et al. 2000; Nuutinen et al. 2003; Schuler et al. 2003; van der Linde et al. 2001), transfer to slowly exchangeable peripheral phases in organisms (Lydy et al. 1994; Spacie and Hamelink 1982; Thybaud and Caquet 1991; Van Hattum and Cid Montanes 1999), and organism growth (Halling-Sørensen et al. 2000; Skoglund et al. 1996) may be important in some cases, as represented by the compartment model shown in Fig. 1b (Barron 1990; Del Vento and Dachs 2002; Looser et al. 2000; Wang et al. 1996). The concentration of a chemical in an organism (Cpb ) can be expressed as

20

T. Katagi

a general differential form (Eq. (5)), assuming first-order kinetics for each process (terms defined in the legend of Fig. 1 b): dCpd /dt = kU × Cpw − (kPE + kPS + kM + kG ) × Cpb + kP/S × Cpbs,

(5)

dCPb /dt = kU × Cpw − kPE × Cpb.

(6)

For not easily metabolized chemicals, this equation can be simplified as expressed in Eq. (6). This comprises a first-order one-compartment model if either the growth or the presence of a peripheral phase in organism is negligible during exposure. When a chemical in water is kept at a constant concentration in a flowthrough system (Cpw = constant), Cpb can be expressed as a function of time (t) as follows (Connell 1988; Ernst 1977; Farrington 1991):   Cpb = (kU /kPE ) × Cpw × 1 − exp (−kPE × t) .

(7)

Furthermore, by transferring the exposed organisms to a non-contaminated system, the elimination profiles can be directly observed. When the dissipation of a chemical in an organism obeys first-order kinetics, the following equation is derived: (8) dCpb /dt = −kPE × Cpb. By solving Eq. (8), the elimination half-life (CL50 ) and the time for an effective equilibrium (teq ), with 99% elimination, can be obtained and presented as 0.693/kPE and 4.605/kPE , respectively. The corresponding equations can also be derived for metabolites. When a biphasic process is observed, the double-exponential equation, which assumes a two-compartment model, is usually used. Examples of such a process were reported for the bioconcentration of atrazine (127) (Nikkilä et al. 2001) and benzo[a]pyrene (7) (McCarthy 1983) in D. magna, mirex (29) in Hyalella azteca (Jessiman and Qadri 1983), and PAHs in Asellus aquaticus (van Hattum and Cid Montanes 1999). Based on the estimated first-order rate constants, the kinetic BCF can be calculated as BCF =Cpb /Cpw = kU /kPE.

(9)

When equilibrium is rapidly reached for bioconcentration in algae, the Cpw and Cpb values are known to obey the Freundlich adsorption isotherm expressed in Eq. (10), where KF and n are the adsorption coefficient and a constant: 1/n

Cpb = KF × Cpw .

(10)

This relationship was utilized to analyze the bioconcentration of various pesticides in algae and daphnids (Canton et al. 1977; Ellgehausen et al. 1980; Hansen 1979; King et al. 1969; Paris and Lewis 1976). The kinetic model used assumes

Bioconcentration, Bioaccumulation, and Metabolism

21

the adsorption to the algal surface and uptake into the matrix of green algae, and was applied to examine the log Kow dependency in the bioconcentration of the PCB congeners (Skoglund et al. 1996). The poor fit of the regression curve by this kinetic model for the lipid-normalized BCF may show more affinity of PCB for lipids than n-octanol; or, there may be possible interactions with algal components other than lipids. Similarly to BCF, the log kU and log (1/kPE ) values are known to correlate with log Kow , as demonstrated in fish, algae, daphnids, and molluscs (Connell 1988; Ellgehausen et al. 1980; Hawker and Connell 1986). When hydrophobic chemicals with log Kow >6 are analyzed, the log kU vs. log Kow line showed a downward curvature as shown for the uptake of chlorinated hydrocarbons in fish (Connell and Hawker 1988), oligochaetes (Connell et al. 1988), and aquatic macrophytes (Gobas et al. 1991). Because the process of bioconcentration proceeds via adsorption, diffusion, and metabolism in organs with different biochemical composition, the two-compartment model is sometimes more suitable to describe uptake and elimination processes. The bioconcentration of lindane (30) in the snail Lymnaea palustris was analyzed by using a first-order two-compartment model, and assumed foot and visceral mass as being the central and peripheral compartments (Thybaud and Caquet 1991). BCF values, similar to the ones obtained with the one-compartment model, were obtained. The rapid elimination from foot, followed by the slower one from the visceral mass, could account for the biphasic profiles. Lydy et al. (1994) analyzed the uptake and elimination processes of pentachlorophenol (32) and 5,5 ,6trichlorobiphenyl (TCB) in Chironomus riparius using a two-compartment model. The authors speculated that the exoskeleton is a central compartment, judging from its apparent volume, which was almost the same for both compounds. In addition, internal tissues that have much more lipid were considered to be peripheral ones, because their apparent volumes retain more of the hydrophobic TCB. A twocompartment model can successfully describe the bioconcentration profiles of the parent and metabolite phases for 2-chlorobiphenyl (CB) in Chironumus tentans (Lydy et al. 2000). Rapid 14 C dissipation in the midge resulted from elimination of both CB and its metabolites. A similar approach was undertaken to analyze the bioconcentration profiles for tributyltin chloride (16), which was debutylated stepwise in the body of larval midges. Different BCF values between (16) and its triphenyl analog originated from differing degrees of metabolism (Looser et al. 2000). Much less metabolic activity existed in the oligochaete Lumbriculus variegatus than in larval midges for benzo[a]pyrene (7) in a study by Schuler et al. (2003). Nuutinen et al. (2003) scrutinized each estimated rate constant in Eq. (5) for effects of metabolism on bioconcentration values for fluoranthene (6), pentachlorophenol (32), and methyl parathion (39) in the amphipod Hyalella azteca. In this study, the larger value for kM and smaller kME showed the rapid metabolism of (39) through involvement of various enzymes and association of the oxon metabolite with acetylcholinesterase. The moderate but larger kM of (32) than for (6) implied that the former substance went through a phase-II conjugation reaction; the small kME of (6) portended less permeability of its oxidized metabolite through membranes.

22

T. Katagi

The elimination from aquatic organisms was kinetically examined for chlorinated hydrocarbons, PAHs, and PCBs that had a wide range of log Kows (van der Linde et al. 2001). The extent of metabolism was evaluated and results showed that the PAHs were metabolized at a moderate rate. Recently, Cowan-Ellsberry et al. (2008) successfully applied in vitro metabolism data, using fish liver hepatocytes or S9 fractions, to refine the BCF values of pesticides and surfactants in fish. When the growth rate of an organism becomes comparable to, or higher than, uptake and elimination rates, the kG value should be taken into account in any analysis of bioconcentration. Since the growth of algae is very rapid during exposure to a chemical, the exponential model for time-dependent increase of algal mass was introduced to examine the bioconcentration of organophosphorus pesticides in green algae (Nakamura and Mochida 1988; Jonsson et al. 2001). Instead of the exponential model, the Boltzman equation (Wang et al. 1996) and the allometric model (van der Linde et al. 2001) were successfully applied to bioconcentration in various aquatic organisms. Halling-Sørensen et al. (2000), by analyzing the bioconcentration of chlorinated hydrocarbons in Selenastrum sp., confirmed that algal growth was significantly retarded by nitrogen depletion in the growth medium. Approaches have been developed, other than using a simple kinetic model, for examining bioconcentration. For example, the fugacity model has been extensively applied to the theoretical investigation of bioconcentration (Mackay and Hughes 1984; Gobas and Mackay 1987). Gobas et al. (1986) utilized the diffusion rates through membrane-diffusion layer barriers to analyze uptake and elimination processes in fish. The uptake and elimination can be expressed by the following equations:   kU = (A/F) × Km × Dm × Dd / (δm × Dd + Km × δd × Dm ) ,   kPE = (A/F) × {(1 − a) + a × Km }   × Km × Dm × Dd / (δm × Dd + Km × δd × Dm ) ,

(11) (12)

where A = diffusion area; δ = diffusion layer thickness; D = diffusion coefficient; α = lipid fraction; F = fish weight; Km = a partition coefficient of a chemical between water and membrane; Subscripts m and d = membrane and diffusion layer, respectively. Membrane-controlled diffusion is considered to be dominant for chemicals of low hydrophobicity (log Kow < 3–4), and then the first term (δ m × Dd ) in the denominator of Eqs. (11) and (12) becomes much larger than the second one (Km × δ d × Dm ). In contrast, the diffusion-layer-controlled process becomes dominant for hydrophobic chemicals (log Kow > 3–4), meaning the opposite condition prevails

Bioconcentration, Bioaccumulation, and Metabolism

23

for δ m × Dd « Km × δ d × Dm . When log [(Dm × A)/(δ m × F)] and log [(Dd × A)/(δ d × F)] are defined as β m and β d , Eqs. (11) and (12) can be simplified as follows:   LogKow < 3 − 4; logkU = βm + logKm ; logkPE = βm − log (1 − α) /Km + α ; LogKow > 3 − 4; logkU = βd ; logkPE = βd − log[ (1 − α) + α × Km ]. Since Km can be approximated as Kow and β d is considered to increase with molecular size, the log kU and log kPE vs. log Kow plots would show parabolic curves. The log Kow dependency in the bioconcentration of non-metabolized chemicals with log Kow >3 could be well described by the above model. Legierse et al. (1998) applied this model to the bioconcentration of chlorthion (43) and several chlorobenzenes in snail Lymnaea stagnalis, by using the parameters from guppies; however, this approach failed to predict kU , KPE , and BCF values well, perhaps because of differences in the specific physiology of the snail. By using the diffusion theory, Wolf et al. (1991) successfully examined the partition of chlorobenzenes to aquatic macrophytes.

2.3 Pesticides and Other Chemicals The BCF and elimination clearance times (CL50 ) for pesticides and simple organic chemicals in aquatic organisms such as molluscs, algae, crustacean, and insecta (other than fish) are listed in Tables 3, 4, 5, and 6, together with brief descriptions of the study designs which yielded these values. For members of Insecta, data are included on the larvae and nymphs and on habitants in the aquatic environment. In these studies, algae were kept in a sterile medium under illumination and were mostly evaluated in the exponential growth phase. When pesticides and other chemicals are tested, exposures are conducted by using static or flow-through systems. Which system is used depends on the water solubility and hydrolytic stability of the chemical. The log BCF values determined for such xenobiotics mostly range between 0 and 6 for the 287 chemicals that have log Kow values between 3 and 7, as measured or estimated by the EPI-suite software and (USEPA 2008). Even for an individual chemical, BCF values significantly scatter among species tested. Values for the organochlorine pesticides, listed in Table 3, primarily show log BCF value of 3–5, with a variation within twofold, among species. A larger variance was observed for endosulfan (27), hexachlorobenzene (31), and DDT (33). The latter two insecticides have large log Kow values (5.73 and 6.91), and the difference in the lipid content of each organism is likely to result in these variations. By analogy to the metabolism of (27) in D. magna (DeLorenzo et al. 2002), the crayfish Procambarus clarkii may more significantly metabolize (27) xenobiotics to their sulfate forms, which results in much reduced BCF values. Aldrin (21) has a very high hydrophobicity (log Kow = 6.5), but its elimination is rapid in ostracods, which results from its oxidation to dieldrin (22) (Kawatski and Schmulbach 1972).

23

22

21

No.

Experimental conditionsc

Aldrin (6.5)

B: Anabaena cylindrica Aulosira fertilissima Anacystis nidulans Os: Chlamydotheca arcuata W: Daphnia magna I: Hexagenia bilineata Chironomus sp. C: Rangia cuneata Corbicula manilensis Mu: Lampsilis siliquoidea

n, st, 1 ppb, 23◦ C, 7 days/na n, st, 0.1–1 ppm, 29◦ C, 2 days/na n, st, 1 ppb, 23◦ C, 7 days/na n, st, 8.4 ppb, 22◦ C, 4 days/2 days n, fl, 0.02 ppb, 21◦ C, 3 days/na n, fl, 0.02 ppb, 21◦ C, 3 days/na n, fl, 0.02 ppb, 21◦ C, 3 days/na Dieldrin (5.2–5.4) n, fl, 0.55 ppb, 25◦ C, 72 hr/na y, fl, 0.89 ppb, na, 72 days/na n, fl, 0.57 ppb, 20◦ C, 3 weeks/3 weeks Sphaerium corneum n, st, 2.4–2.6 ppb, 10–19◦ C, 1 day/na Oy: Crassostrea virginica n, fl, 0.5 and 9 ppb, 23◦ C, 7 days/na G: Scenedesmus obliquus n, st, 1–20 ppb, 25◦ C, 36 hr/na B: Anabaena cylindrica n, st, 1 ppb, 23◦ C, 7 days/na Aulosira fertilissima n, st, 0.1–1 ppm, 27◦ C, 5 days/na Anacystis nidulans n, st, 1 ppb, 23◦ C, 7 days/na Nostoc muscorum n, st, 1 ppb, 23◦ C, 7 days/na W: Daphia magna n, st, 2–13 ppb, 21◦ C, 6 days/na Os: Chlamydotheca arcuata n, st, 6.6 ppb, 22◦ C, 4 days/2 days Photodieldrin (4.13) G: Ankistrodesmus amalloides n, st, 0.72 ppb, na, 2 days/na W: Daphnia pulex n, st, 3.3 ppb, na, 1 day/4 days

Pesticide (log Kow a ) Speciesb na na na 1 day na na na na na 4.7 days na na na na na na na na 1 day na 4 days

3.1∗ 2.3–2.6∗ 3.0∗ 3.9∗ 5.1 4.5 4.4 2.9–3.3 3.5 3.0–3.1 2.8 3.3–3.5 3.1∗ 2.3∗ 1.4–2.5∗ 2.7∗ 3.3∗ 4.1∗ 3.4∗ 1.6–1.7∗ 1.9∗

log BCFd CL50 e

Reinert (1972) Schauberger and Wildman (1977) Kumar and Lal (1988) Schauberger and Wildman (1977) Schauberger and Wildman (1977) Reinert (1972) Kawatski and Schmulbach (1972) Neudorf and Khan (1975) Khan et al. (1975)

Mason and Rowe (1976)

Boryslawskyj et al. (1988)

Schauberger and Wildman (1977) Dhanaraj et al. (1989) Schauberger and Wildman (1977) Kawatski and Schmulbach (1972) Johnson et al. (1971) Johnson et al. (1971) Johnson et al. (1971) Petrocelli et al. (1973) Hartley and Johnston (1983) Bedford and Zabik (1973)

References

Table 3 A summary of bioconcentration studies performed on organochlorine pesticides in aquatic organisms

24 T. Katagi

Endrin (5.2–5.4)

Endosulfan (3.83)

25

27

na na na na

n, fl, 1.5 ppb, 10◦ C, 20 days/na n, st, 0.1–1 ppm, 29◦ C, 2 days/na n, st, 0.1 ppm, 25◦ C, 16 hr/na n, st, 0.1 ppm, 8 weeks/8 weeks n, st, 0.1 ppm, 25◦ C, 24 hr/na

Mu: Anodonta piscinalis B: Anabaena sp. ARM310 G: Selenastrum capricornutum Cr: Procambarus clarkii

W: Daphnia magna

2.8 2.7–3.7∗ 3.4 – 0.7 (α), 0.3 (β) 3.5

na

n, fl, 0.1–1.4 ppb, 28◦ C, 10 days/na 1.0–1.8

C: Katelysia opima

na

na na na na na

Oy: Crassostrea virginica Mu: Mytilus edulis I: Pteronarcys dorsata

Ankistrodesmus amalloides Mc: Hydrilla verticillata W: Daphnia pulex I: Chironomus decorus 1 day na 1 day na

na na

na

G: Scenedesmus quadricauda Oy: Crassostrea madrasersis

n, fl, 1.5 ppb, 10◦ C, 20 days/na n, st, 0.1–100 ppb, na, 1 day/na

Mu: Anodonta piscinalis G: Scenedesmus quadricauda

3.7 (α), 3.6 (γ) 3.0 3.7–4.1 (α) 3.8–4.2 (γ) 3.7∗ 3.0 4.4∗ 2.3–2.5 (α) 3.2–3.4 3.4 2.9

log BCFd CL50 e

n, st, 0.72 ppb, 25◦ C, 1 day/na n, st, 5 ppb, 25◦ C, 6 days/na n, st, 0.5 ppb, 25◦ C, 1 day/3 days n, rn, 0.7–1.4 ppt, 20◦ C, 50 days/na n, fl, 0.1–50 ppb, 23◦ C, 7 days/na n, rn, na, 15◦ C, 7 days/na n, fl, 0.03–0.15 ppb, 15◦ C, 28 days/na n, st, 1 ppm, 25◦ C, 7 days/na 2.2∗ n, fl, 0.1–1.4 ppb, 28◦ C, 10 days/na 1.0–1.9

y, fl, 0.3–0.4 ppb, na, 72 days/na

Chlordane (6.1–6.22)

24

Experimental conditionsc

C: Corbicula manilensis

Pesticide (log Kow a ) Speciesb

No.

Table 3 (continued)

DeLorenzo et al. (2002)

Vance and Drummond (1969) Rajendran and Venugopalan (1991) Rajendran and Venugopalan (1991) Sabali¯unas et al. (1998) Narayana Rao and Lal (1987) DeLorenzo et al. (2002) Naqvi and Newton (1990)

Mason and Rowe (1976) Donkin et al. (1997) Anderson and DeFoe (1980)

Moore et al. (1977) Hinman and Klaine (1992) Moore et al. (1977) Harkey and Klaine (1992)

Sabali¯unas et al. (1998) Glooschenko et al. (1979)

Hartley and Johnston (1983)

References

Bioconcentration, Bioaccumulation, and Metabolism 25

Lindane (3.72–4.14) C: Corbicula manilensis Mu: Mytilus edulis

30

α-isomer

Hexachlorobenzene C: Corbicula manilensis (5.73) Mu: Mytilus edulis Sn: Lymnaea palustris O: Lumbriculus variegates

–

31

A: Hyalella azteca Gammarus lacustris

Mc: Hydrilla verticillata G: Chlorella pyrenoidosa Chlamydomonas sp. Dunaliella sp. W: Daphnia magna

Hexachlorohexane (4.52)

–

Is: Asellus aquaticus G: Chlorella pyrenoidosa C: Venerupis japonica

P: Lanice conchilega

Sn: Lymnaea palustris

Pesticide (log Kow

Speciesb

No.

a)

n, rn, 0.5 ppb, 10◦ C, 103 hr/na y, st, 0.5–5 ppb, na, 21 days/na y, fl, 0.7–4 ppb, 20◦ C, 28–44 days/na y, fl, 1.2–7.7 ppb, 20◦ C, 28 days/na n, fl, 0.3–5 ppb, 20◦ C, 28 days/na

4.1–4.4 4.3–4.5

>3.0 5.0–5.4 3.7–4.3

na na

na na na

na

na na na na na

1–2 days na na

4.7 days

na 0.7 hr

2.5 1.6–1.7 3.1

na na 22 hr

3.4 2.4–2.6 2.1

log BCFd CL50 e

1.7 2.6–3.1 2.2 (α), 2.1 (β) 2.1 (γ), 2.4 (δ) n, st, 0.12 ppm, 25◦ C, 6 days/na 1.6 n, st, 0.01–0.8 ppm, 28◦ C, 3 hr/na 2.2–2.4 n, st, 0.1–0.8 ppm, na, 2–3 hr/na 3.4∗ n, st, 0.1–0.8 ppm, na, 2–3 hr/na 3.2∗ n, st, 0.01–0.8 ppm, 20◦ C, 1.8–2.5 2 days/na y, fl, 0.59 ppb, na, 72 days/na 3.4

y, fl, 0.3 ppb, na, 72 days/na n, st, 0.9–1.8 ppb, 22◦ C, 8 days/na n, st, 2–5 ppb, 10◦ C, 8 days/18 days n, rn, na, 15◦ C, 7 days/na n, st, 6–600 ppb, 20◦ C, 10 days/7 days n, st, 2–5 ppb, 10◦ C, 8 days/18 days n, st, 2 ppb, 18◦ C, 5 days/3 days n,. st, 0.01–1 ppm, 20◦ C, 6 days/na n, fl, 1–2 ppb, na, 10 days/4 days

Experimental conditionsc

Table 3 (continued)

Schuytema et al. (1988) Nebeker et al. (1989)

Bauer et al. (1989) Baturo and Lagadic (1996) Schuytema et al. (1988)

Hartley and Johnston (1983)

Hinman and Klaine (1992) Canton et al. (1975) Canton et al. (1977) Canton et al. (1977) Canton et al. (1975)

Thybaud and Le Bras (1988) Hansen (1979) Yamato et al. (1983)

Ernst (1979)

Donkin et al. (1997) Thybaud and Caquet (1991)

Hartley and Johnston (1983) Renberg et al. (1978) Ernst (1979)

References

26 T. Katagi

32

A: Gammarus pulex Hyalella azteca Pontoporeia hoyi S: Mysis relicta I: Chironomus riparius W: Daphnia magna

Mc: Eichhornia crassipes

Anodonta anatina Oy: Crassostrea gigas Ab: Haliotis rufescens Haliotis fulgens P: Lanice conchilega

Dreissena polymorpha

Hexachlorobenzene G: Scenedesmus sp. (5.73) Mc: Myriophyllum spicatum I: Chironomus decoras Toxaphene (5.78) Oy: Crassostrea virginica S: Penaeus duorarum Palamonetes pugio Pentachlorophenol C: Corbicula fluminea (5.12) Mu: Mytilus edulis

31

28

Pesticide (log Kow a ) Speciesb

No.

2.2–2.5 1.4–1.7 1.2–1.3 1.6–1.8 3.6

0.4 day 3.6 hr 8.8 days 15 days 15 hr na

na

na <0.5 day <0.5 day <0.5 day 21 days

0.6 day

3.2 days

2.6 2.6–3.2

8.5 days na na na na 1 day

3.0 2.9 3.9–4.2 2.6–2.9 2.9–3.1 1.9–2.0

2.1 (leaf), 2.2 (root) n, st, 70 ppb, 16◦ C, 3 days/3 days 1.7 n, st, 132 ppb, 21◦ C, 1 day/3 days 2.1 n, fl, na, 4◦ C, 6 hr/7 days 3.0 n, st, na, 4◦ C, 6 hr/7 days 2.1 n, st, 0.9 ppb, 20◦ C, 16 hr/36 hr 2.7 n, st, 20 ppb, 20◦ C, 1 day/na 2.2–2.8

n, fl, na, 21◦ C, 21 days/133 days y, fl, 0.1 ppb, 20◦ C, 2 days/na n, fl, 5–46 ppb, 28◦ C, 4 days/na n, fl, 0.8–4 ppb, 26◦ C, 4 days/na n, fl, 3–10 ppb, 21◦ C, 4 days/na n, st, 30–100 ppb, 20◦ C, 72 hr/72 hr n, st, 2–5 ppb, 10◦ C, 8 days/18 days n, st, 3 ppb–11 ppm, 10–25◦ C, 6 hr/7 days n, st, 7–14 ppb, 13◦ C, 1 day/na n, fl, 0.8 ppm, 15◦ C, 5 hr/13 h n, fl, 1.2 ppm, 14◦ C, 5 hr/13 h n, fl, 0.5 ppm, 15◦ C, 5 hr/13 h n, st, 2–5 ppb, 10◦ C, 8 days/18 days n, st, 0.5 ppm, 26◦ C, 2 days/na

na

log BCFd CL50 e

n, st, 0.08 ppm, 5–39◦ C, 2 days/na 1.6–2.2∗

Experimental conditionsc

Table 3 (continued)

Ashauer et al. (2006) Nuutinen et al. (2003) Landrum and Dupuis (1990) Landrum and Dupuis (1990) Lydy et al. (1994) Kukkonen and Oikari (1988)

Roy and Hänninen (1994)

Mäkelä and Oikari (1990) Shofer and Tjeerdema (1993) Tjeerdema and Crosby (1992) Shofer and Tjeerdema (1993) Ernst (1979)

Fisher et al. (1999)

Ernst (1979)

Koelmans and Sánchez Jiménez (1994) Gobas et al. (1991) Knezovich and Harrison (1988) Schimmel et al. (1977) Schimmel et al. (1977) Schimmel et al. (1977) Basack et al. (1997)

References

Bioconcentration, Bioaccumulation, and Metabolism 27

DDT (6.91)

Dicofol (5.02)

36

Experimental conditionsc

Cr: Orconectes nais S: Palaemonetes kadiakensis Artemia nauplii I: Chironomus sp. Ephemera danica Ci: Blepharisma intermedium A: Hyalella azteca

Mu: Anodonta grandis

n, fl, 0.1 ppb, 21◦ C, 3 days/na n, fl, 0.08 ppb, 21◦ C, 3 days/na n, rn, 0.5–1 ppb, 20◦ C, 1 day/na n, fl, 0.05 ppb, 21◦ C, 3 days/na n, fl, 0.76 ppb, 14◦ C, 9 days/na n, st, 1 ppm, 24◦ C, 10 days/na n, fl, 1.6–1.8 ppm, 15◦ C, 28 days/na

n, fl, 0.62 ppb, 20◦ C, 3 weeks/4 weeks Sn: Vivipara heliciformis n, st, 5–50 ppb, na, 22 days/19 days C: Indonaia caerulea n, st, 5–50 ppb, na, 19 days/11 days G: Selenastrum capricornutum n, st, 16 ppb, 22◦ C, 2 hr/na Chlorella vularis n, st, 10 ppb, 23◦ C, 2 days/2 days Scenedesmus obliquus n, st, 1 ppm, 26◦ C, 7 days/na B: Anacystis nidulans n, st, 1 ppm, 26◦ C, 7 days/na D: Nitzschia closterium n, st, 10 ppb, 23◦ C, 2 days/2 days Cylindrotheca closterium n, st, 0.1 ppm, na, 21 days/na E: Euglena gracilis n, st, 1 ppm, 26◦ C, 7 days/na W: Daphnia magna n, st, 8–16 ppb, 21◦ C, 26 hr/na A: Gammarus fasciatus n, fl, 0.08 ppb, 21◦ C, 3 days/na Hyalella azteca n, rn, 0.02–0.12 ppb, 20◦ C, 2 days/8 days Diporeia sp. n, rn, 0.2–3.3 ppb, 4◦ C, 2 d/90 d

Pesticide (log Kow a ) Speciesb

33

No.

Table 3 (continued)

4–5 days na 17 hr na na 8.4 days na na na na 3–4 days 3–5 month na na na na na na na

2.5–2.8 4.6–5.5∗ 2.6∗ 2.8∗ 2.9∗ 4.9∗ 2.3∗ 2.0∗ 4.2–4.4 4.3 4.4–4.6 5.3–5.5 3.4 3.7 2.4 4.7 2.6–3.9 4.8 3.8–4.1

<1 day

1.9–2.5

Johnson et al. (1971) Johnson et al. (1971) Wang and Simpson (1996) Johnson et al. (1971) Södergren and Svensson (1973) Saxena et al. (1982) Spehar et al. (1982)

Lotufo et al. (2000)

Halling-Sørensen et al. (2000) Kikuchi et al. (1984) Gregory et al. (1969) Gregory et al. (1969) Kikuchi et al. (1984) Keil and Priester (1969) Gregory et al. (1969) Crosby and Tucker (1971) Johnson et al. (1971) Lotufo et al. (2000)

Pillai et al. (1980)

Yadav et al. (1978)

13.6 days Bedford and Zabik (1973)

References

3.4

log BCFd CL50 e

28 T. Katagi

Methoxychlor (5.08) Mu: Mytilus edulis Sn: Physa integra

37

G: Chlorella pyrenoidosa I: Pteronarcys dorsata

Diporeia sp.

n, st, 0.11 ppt, 5–25◦ C, 1 day/na n, rn, 0.5–1 ppb, 20◦ C, 1 day/na n, fl, 0.1–1 ppb, 21◦ C, 30 days/na n, rn, 1.12 ppb, 20◦ C, 2 days/8 days n, rn, 2.3–20 ppb, 4◦ C, 2 days/90 days n, fl, 9–12 ppb, 22◦ C, 21 days/na n, fl, 0.42–4.2 ppb, 15◦ C, 28 days/na n, st, 8–50 ppb, 15◦ C, 200 hr/na n, fl, 0.15–4.2 ppb, 15◦ C, 28 days/na

Experimental conditionsc

2.8

3.9∗

4.1 3.8

5.5–5.7

3.7–4.3 1.6–1.7 4.1–4.3 4.6

na na

na na

144 days

na na na 2.3 days

log BCFd CL50 e

Paris and Lewis (1976) Anderson and DeFoe (1980)

Renberg et al. (1978) Anderson and DeFoe (1980)

Lotufo et al. (2000)

Nawaz and Kirk (1995) Wang and Simpson (1996) Derr and Zabik (1972) Lotufo et al. (2000)

References

b Designation

by EPI-Suite (USEPA 2008) or experimental data therein. of species. A, amphipod; Ab, abalone; B, blue-green alga; C, clam; Ci, ciliate; Cr, crayfish; D, diatom; E, Euglenophyta; G, green alga; I, aquatic insect; Is, isopod; Mc, macrophyte; Mu, mussel; O, oligochaete; Os, ostracod; Oy, oyster; P, polychaete; S, shrimp; Sn, snail; W, water flea. c Application of pesticide to water; presence (y) or absence (n) of sediment in the system, exposure condition (static (st)/renewal (rn)/flow-through (fl)), concentration, temperature (◦ C), periods of exposure/elimination. d Experimentally obtained bioconcentration factor based on the overlying water concentration in the whole body (or each tissue). The asterisk means dryweight-basis. e 50% clearance time. If the biphasic elimination is observed, the CL value of the faster elimination is included. 50 na: Not available.

a Estimated

DDE (6.51)

34

W: Daphnia pulex S: Artemia nauplii I: Chironomus tentans A: Hyalella azteca

Pesticide (log Kow a ) Speciesb

No.

Table 3 (continued)

Bioconcentration, Bioaccumulation, and Metabolism 29

Pesticide (log Kow a )

Parathion (3.83)

Fenitrothion (3.30)

Fenitrothion (3.30)

No.

39

41

41

G: Chlamydomonas reinhardtii Chlorella vularis Scenedesmus quadricauda B: Anabaena flos-aquae Aulosira fertilissima Microcystis aeruginosa D: Aulacpseira granulata Nitzschia closterium Mc: Lemna minor W: Daphnia pulex S: Palaemon paucidens S: Penaeus japonicus

Sn: Cipangopaludina japonica Physa acuta

Anodonta cataractae

C: Mya arenaria

G: Scenedesmus obliquus B: Anacystis nidulans E: Euglena gracilis A: Hyalella azteca Mu: Mytilus edulis

Speciesb

1.6∗ 1.0–1.5∗ 1.7∗ 2.0–2.9∗ 1.5–2.4∗ 1.1–1.7∗ 2.0∗ 1.3 1.9 0.8 2.1

n, st, 10 ppb, 23◦ C, 2 days/2 days n, st, 1–50 ppb, 25◦ C, 1 day/na n, st, 10 ppb, 23◦ C, 2 days/2 days n, st, 1–10 ppm, 29◦ C, 5 days/na n, st, 1–50 ppb, 25◦ C, 1 day/na n, st, 1–50 ppb, 20◦ C, 1 day/na n, st, 10 ppb, 23◦ C, 2 days/2 days n, st, 3.9 ppb, 25◦ C, 5 days/25 days n, fl, 1 ppb, 18◦ C, 1 day/1 days n, fl, 1 ppb, 25◦ C, 3 days/1 days n, st, 0.5 ppb, 25◦ C, 1 day/na

0.4 days

1.7

0.9 hr na 2.6 hr na na na 0.9 hr 1.3 days 5 hr 1.5 hr na

na

0.4 days

1.3

2 days

1–2 days

1.3–2.5 1.0

na na na 0.7 hr 1–2 days

CL50 e

1.9∗ 1.7∗ 1.8∗ 1.1 1.9–2.1

2.5∗

log BCFd

n, st, 1 ppm, 26◦ C, 7 days/na n, st, 1 ppm, 26◦ C, 7 days/na n, st, 1 ppm, 26◦ C, 7 days/na n, st, 2.3 ppb, 22◦ C, 1 day/3 days n, fl, 0.2–13 ppb, 15◦ C, 14 days/ 28 days n, fl, 0.2–13 ppb, 15◦ C, 14 days/ 28 days n, fl, 0.8 ppb, 12◦ C, 14 days/ 28 days n, fl, 0.1 ppm, 25◦ C, 3 days/ 1–3 days n, fl, 0.1 ppm, 25◦ C, 3 days/ 1–3 days n, st, 1 ppm, 20◦ C, 1 day/na

Experimental conditionsc

Kikuchi et al. (1984) Guanzon et al. (1996) Kikuchi et al. (1984) Lal et al. (1987) Guanzon et al. (1996) Guanzon et al. (1996) Kikuchi et al. (1984) Lockhart et al. (1984) Takimoto et al. (1987b) Takimoto et al. (1987b) Kobayashi et al. (1985a)

Kent and Currie (1995)

Takimoto et al. (1987a)

Takimoto et al. (1987a)

McLeese et al. (1979)

McLeese et al. (1979)

Gregory et al. (1969) Gregory et al. (1969) Gregory et al. (1969) Nuutinen et al. (2003) McLeese et al. (1979)

References

Table 4 A summary of bioconcentration studies performed on organophosphorus pesticides in aquatic organisms

30 T. Katagi

Pesticide (log Kow a )

Fenitrooxon (1.69) Chlorothion (3.45) Fenthion (4.84)

Chlorpyrifos (4.7)

No.

42 43 44

47

I: Hydropsyche sp. Stenacrn sp.

C: Venus gallina B: Anabaena sp. ARM310 Aulosira fertilissima S: Artemia parthenogenetica A: Gammarus pulex Is: Asellus aquaticus

Mytilus galloprovincialis Oy: Crassostrea virginica

Mu: Mytilus edulis

Artemia salina Cr: Procambarus clarkii Co: Sinocalanus tenellus R: Brachionus plicatilis S: Penaeus japonicus Sn: Lymnaea stagnalis C: Marcia hiantina

Speciesb

n, st, 0.1 ppb, 12◦ C, 3 days/3 days y, fl, 0.7–5 ppb, 12–25◦ C, 23 days/na n, st, 3 ppb, 20◦ C, 6 hr/na n, st, 3 ppb, 20◦ C, 6 hr/na

n, st, 0.3 ppb, 25◦ C, 12 hr/72 hr n, st, 0.1 ppm, 25◦ C, 3 days/na n, st, 20 ppb, 22◦ C, 2 days/21 days n, st, 0.1 ppm, 25◦ C, 3 days/na n, st, 0.1 ppm, 25◦ C, 3 days/na n, st, 3 ppb, 25◦ C, 12 hr/72 hr n, fl, 27 ppb, 20◦ C, 10 days/na n, rn, 0.02–0.2 ppm, 28◦ C, 15 days/15 days n, rn, 1–3.2 ppm, 18◦ C, 24–38 days/38 days n, rn, 1 ppm, 18◦ C, 35 days/na n, fl, 0.7 ppb, 22◦ C, 28 days/ 14 days n, st, 1–56 ppm, 18◦ C, 4 days/na n, st, 1–10 ppm, 29◦ C, 5 days/na n, st, 1–10 ppm, 29◦ C, 5 days/na n, st, 0.5–100 ppb, 20◦ C, 2 days/na

Experimental conditionsc

Table 4 (continued)

1.6 1.1

na na

1.5 days 1–4 days

na na na na

0.5 0.8–2.8∗ 1.7–2.6∗ 3.0–3.8 3.2 5.4

na 2.5 days

5 days

6 hr na na na na 12 hr 2h na

CL50 e

2.6 2.8

2.4–2.7

2.1 3.7 2.1 3.6 3.3 0.5 1.5 1.0–2.3

log BCFd

Ashauer et al. (2006) Cid Montañés and van Hattum (1995) Tang and Siegfried (1996) Tang and Siegfried (1996)

Serrano et al. (1995) Lal et al. (1987) Lal et al. (1987) Varó et al. (2000)

Serrano et al. (1997b) Woodburn et al. (2003)

Serrano et al. (1997a)

Kobayashi et al. (1990) Kashiwada et al. (1985b) Escartin and Porte (1996) Kashiwada et al. (1995b) Kashiwada et al. (1995b) Kobayashi et al. (1990) Legierse et al. (1998) Sathe et al. (2005)

References

Bioconcentration, Bioaccumulation, and Metabolism 31

Diazinon (3.81)

Pyridaphenthion (3.2) Demeton-S-methyl (1.02)

48

50

Phorate (3.56)

Dimethoate (0.78)

Malathion (2.36)

Malathion (2.36) Methidathion (2.20)

52

55

57

57 58

51

Pesticide (log Kow

No.

a)

Myriophyllum aquaticum Elodea Canadensis B: Anabaena sp. ARM310 Aulosira fertilissima Mu: Mytilus galloprovincialis B: Anabaena sp. ARM310 Aulosira fertilissima B: Anabaena sp. ARM310 Aulosira fertilissima Mc: Spirodela oligorrhizu Myriophyllum aquaticum Elodea Canadensis Co: Sinocalanus tenellus S: Artemia salina R: Brachionus plicatilis Mu: Mytilus galloprovincialis C: Venus gallina

Mc: Spirodela oligorrhizu

Sn: Cipangopoludina malleata Cr: Procambarus clarkii S: Artemia salina Co: Sinocalanus tenellus R: Brachionus plicatilis G: Chlorella saccharophila

Speciesb

log BCFd 0.8 0.7 2.6 3.5 2.7 1.4 1.1 –1.6 –0.4 0.5–1.1∗ 0.9–1.1∗ –0.5 0.3–1.9∗ 0.0–2.1∗ 1.9–2.5∗ 2.0–2.3∗ 1.4 0.5 0.08 3.7 2.3 4.6 2.3 –0.03

Experimental conditionsc n, fl, 10 ppb, 20◦ C, 7 days/na n, fl, 10 ppb, 20◦ C, 7 days/na n, st, 1 ppm, 25◦ C, 3 days/na n, st, 1 ppm, 25◦ C, 3 days/na n, st, 1 ppm, 25◦ C, 3 days/na n, st, 10 ppm, 22◦ C, 7 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 0.1–1 ppm, 29◦ C, 2 days/na n, st, 0.1–1 ppm, 29◦ C, 2 days/na n, st, 3–56 ppm, 18◦ C, 4 days/na n, st, 0.1–1 ppm, 27◦ C, 5 days/na n, st, 0.1–1 ppm, 27◦ C, 5 days/na n, st, 0.1–1 ppm, 29◦ C, 2 days/na n, st, 0.1–1 ppm, 29◦ C, 2 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 15 ppb, 20◦ C, 3 days/na n, st, 0.3 ppm, 25◦ C, 3 days/na n, st, 0.3 ppm, 25◦ C, 3 days/na n, rn, 1 ppm, 18◦ C, 35 days/na n, st, 3–56 ppm, 18◦ C, 4 days/na

Table 4 (continued)

na

na na na na na na na na na na na

na na na na na

na

na na na na na

na

CL50 e

Serrano et al. (1995)

Kumar and Lal (1988) Kumar and Lal (1988) Narayana Rao and Lal (1987) Narayana Rao and Lal (1987) Gao et al. (2000a) Gao et al. (2000a) Gao et al. (2000a) Kashiwada et al. (1995a) Kashiwada et al. (1995b) Kashiwada et al. (1995b) Serrano et al. (1997b)

Gao et al. (2000a) Gao et al. (2000a) Dhanaraj et al. (1989) Dhanaraj et al. (1989) Serrano et al. (1995)

Gao et al. (2000a)

Kanazawa (1978) Kashiwada et al. (1995b) Kashiwada et al. (1995b) Kashiwada et al. (1995b) Jonsson et al. (2001)

Kanazawa (1978)

References

32 T. Katagi

Phosmet (2.78)

Dichlorvos (1.47) Chlorfenvinphos (3.8) Crufomate (3.42)

59

62 65

W: Daphnia magna I: Chironomus plumosus Cr: Orconectes nais Mu: Mytilus edulis Mu: Mytilus galloprovincialis Mc: Spirodela oligorrhizu Myriophyllum aquaticum Elodea Canadensis

Speciesb

log BCFd 0.3 0.8 0.8 0.04 2.4 0.4 0.4 –1.3

Experimental conditionsc n, fl, 1.2 ppb, 20◦ C, 2 days/na n, fl, 1.2 ppb, 20◦ C, 2 days/na n, fl, 1.2 ppb, 20◦ C, 2 days/na n, st, na, 15◦ C, 3–7 days/na n, rn, 1 ppm, 18◦ C, 35 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 1 ppm, 22◦ C, 8 days/na n, st, 1 ppm, 22◦ C, 8 days/na

na na na

na na na na na

CL50 e

Gao et al. (2000a) Gao et al. (2000a) Gao et al. (2000a)

Julin and Sanders (1977) Julin and Sanders (1977) Julin and Sanders (1977) Donkin et al. (1997) Serrano et al. (1997b)

References

b Designation

by EPI-Suite (USEPA 2008) or experimental data therein. of species. A, amphipod; Ab, abalone; B, blue-green alga; C, clam; Ci, ciliate; Cr, crayfish; D, diatom; E, Euglenophyta; G, green alga; I, aquatic insect; Is, isopod; Mc, macrophyte; Mu, mussel; O, oligochaete; Os, ostracod; Oy, oyster; P, polychaete; S, shrimp; Sn, snail; W, water flea. c Application of pesticide to water; presence (y) or absence (n) of sediment in the system, exposure condition (static (st)/renewal (rn)/flow-through (fl)), concentration, temperature (◦ C), periods of exposure/elimination. d Experimentally obtained bioconcentration factor based on the overlying water concentration in the whole body (or each tissue). The asterisk means dryweight-basis. e 50% clearance time. If the biphasic elimination is observed, the CL value of the faster elimination is included. 50 na, not available; Co, copepod; R, rotifer.

a Estimated

66

Pesticide (log Kow a )

No.

Table 4 (continued)

Bioconcentration, Bioaccumulation, and Metabolism 33

Lambda-cyhalothrin (6.8–7.0)

Fenvalerate (6.2)

Flucythrinate (6.2)

84

85

86

Oy: Crassostrea virginica

Mu: Anodonta piscinalis Oy: Crassostrea virginica Sn: Helisoma trivolvis G: Chlamydomonas reinhardtii W: Daphnia galeata Mu: Mytilus edulis

Mu: Anodonta piscinalis Oy: Crassostrea virginica Sn: Helisoma trivolvis B: Anabaena sp. ARM310 Aulosira fertilissima Mc: Lemna minor I: Hydropsyche sp. Stenacrn sp. Pteronarcys dorsata I: Chironomus riparius y, st, 0.2 ppb, 23◦ C, 4 days/na n, fl, 1.5 ppb, 10◦ C, 20 days/na n, fl, 1 ppb, 29◦ C, 28 days/37 days n, fl, 0.02–0.8 ppb, 15◦ C, 28 days/na n, st, 16 ppb, 20◦ C, 1 hr/na n, 0.1–0.5 ppb, 20◦ C, 2 days/na n, rn, 0.04–0.4 ppm, 15◦ C, 3–7 days/na n, fl, 1 ppb, 29◦ C, 28 days/52 days

n, n, n, n, n, n, n, n, n, n,

fl, 1.5 ppb, 10◦ C, 20 days/na fl, 1 ppb, 28◦ C, 31 days/40 days fl, 16 ppb, 15◦ C, 28 days/na st, 0.1 ppm, 27◦ C, 5 days/na st, 0.1 ppm, 27◦ C, 5 days/na st, 2.3–3 ppb, 25◦ C, 5 days/15 days st, 3 ppb, 20◦ C, 6 hr/na st, 3 ppb, 20◦ C, 6 hr/na fl, 0.03–0.4 ppb, 15◦ C, 4 days/na st, 0.2 ppb, 23◦ C, 4 days/na

Experimental conditionsc

na na <1 week na na na na 10 days

3.4

na <1 week na na na 18 days na na na na

CL50 e

3.1–3.5 2.5 3.7 2.6–3.1 2.3 4.0–4.6 2.9–3.6

2.3 3.3 2.9 1.8–2.9 1.6–3.4 2.3 (trans), 2.2 (cis) 1.5 1.4 2.3 3.2–3.3

log BCFd

Schimmel et al. (1983)

Hamer et al. (1999) Sabali¯unas et al. (1988) Schimmel et al. (1983) Anderson (1982) Day and Kaushik (1987) Day and Kaushik (1987) Donkin et al. (1997)

Sabali¯unas et al. (1988) Schimmel et al. (1983) Spehar et al. (1983) Kumar and Lal (1988) Kumar and Lal (1988) Lockhart et al. (1984) Tang and Siegfried (1996) Tang and Siegfried (1996) Anderson (1982) Hamer et al. (1999)

References

b Designation

by EPI-Suite (USEPA 2008) or experimental data therein. of species. A, amphipod; Ab, abalone; B, blue-green alga; C, clam; Ci, ciliate; Cr, crayfish; D, diatom; E, Euglenophyta; G, green alga; I, aquatic insect; Is, isopod; Mc, macrophyte; Mu, mussel; O, oligochaete; Os, ostracod; Oy, oyster; P, polychaete; S, shrimp; Sn, snail; W, water flea. c Application of pesticide to water; presence (y) or absence (n) of sediment in the system, exposure condition (static (st)/renewal (rn)/flow-through (fl)), concentration, temperature (◦ C), periods of exposure/elimination. d Experimentally obtained bioconcentration factor based on the overlying water concentration in the whole body (or each tissue). The asterisk means dry-weight-basis. e 50% clearance time. If the biphasic elimination is observed, the CL value of the faster elimination is included. 50

a Estimated

Allethrin (4.78) Permethrin (6.5)

80 81

Speciesb

Table 5 A summary of bioconcentration studies performed on pyrethroid insecticides in aquatic organisms

Pesticide (log Kow a )

No.

34 T. Katagi

G: Scenedesmus acutus G: Scenedesmus acutus Cr: Procambarus clarkii

Acids 75 2,4-D (2.81) 76 2,4,5-T (3.31) 77 Triclopyr (2.53)

95 CNA (2.52) Carbamates 98 Aminocarb (1.9)

n, st, 0.4 ppm, 15◦ C, 4 days/8 days n, st, 0.04–1 ppm, 12–24◦ C, 1–3 days/ 6 days n, st, 6.3 ppb, 25◦ C, 5 days/15 days

S: Macrobrachium rosenbergii Sn: Biomphalaria glabrata

Mu: Mytilus edulis Is: Caecidolea racovitzai racovitzai Mc: Lemna minor

1.9–2.6

n, fl, 0.1–0.2 ppm, 24◦ C, 22 day/na

References

0.5 days <1 days 24 days

1.7

na

1.9 days

1.5 days

na

na na

na na na na na

na

Lockhart et al. (1984)

McLeese et al. (1981) Richardson et al. (1983)

Duncan et al. (1977)

Wang et al. (1992b)

Wang et al. (1992b)

Aly et al. (1984)

Guanzon et al. (1996) Guanzon et al. (1996)

Wofford et al. (1981) Mayer and Sanders (1973) Mayer and Sanders (1973) Mayer and Sanders (1973) Guanzon et al. (1996)

Wofford et al. (1981)

na Böhm and Müller (1976) na Böhm and Müller (1976) 7–15 days Barron et al. (1991)

CL50 e

0–0.3 1.2–1.4

2.6–2.7

–1.5

0.04–0.8

2.9–3.3∗ 1.8–2.3∗

n, st, 1–50 ppb, 25◦ C, 1 day/na n, st, 1–50 ppb, 20◦ C, 1 day/na

n, rn, 0.01–0.1 ppm, 22◦ C, 30 days/ 30 days n, rn, 0.01–0.1 ppm, 22◦ C, 30 days/ 30 days n, fl, 1%, 24◦ C, 18–32 hr/na

0.5–1.5 2.6 2.9 2.6 1.7–2.2∗

n, st, 0.1–0.5 ppm, na, 1 day/na n, st, 80 ppb, na, 14 days/na n, st, 0.18 ppm, na, 7 days/na n, st, 80 ppb, na, 7 days/na n, st, 1–50 ppb, 25◦ C, 1 day/na

S: Penaeus aztecus W: Daphnia magna I: Chironomus plumosus Hexagenia bilineata 88 Isoprothiolane (2.88) G: Scenedesmus quadricauda B: Microcystis aeruginosa D: Aulacpseira granulata Amides 90 Dicryl (3.29) G: Scenedesmus quadricauda 93 Butachlor (4.5) C: Corbicula fluminea

1.3–1.6

0.3 0.5 –0.3 to 0.04

log BCFd

n, st, 0.1–0.5 ppm, na, 1 day/na

n, st, 11 ppb, 30◦ C, 8 hr/na n, st, 13 ppb, 30◦ C, 8 hr/na n, st, 1–2.5 ppm, 22◦ C, 11 days/36 days

Experimental conditionsc

Oy: Crassostrea virginica

Esters 13 Di-n-butyl phthalate (4.5)

Speciesb

No. Pesticide (log Kow a )

Table 6 A summary of bioconcentration studies performed on other pesticides and other chemicals in aquatic organisms

Bioconcentration, Bioaccumulation, and Metabolism 35

G: Chlorella fusca G: Chlorella fusca Mc: Lemna minor Elodea densa 120 Diflubenzuron (3.88) G: Scenedesmus subspicatus

Ureas 116 Diuron (2.68) 118 Isoproturon (2.87)

Mu: Mytilus edulis C: Corbicula japonica Corbicula fluminea

109 Thiobencarb (3.4)

Sn: Cipangopludina chinensis W: Daphnia magna A: Gammarus pseudolimnaeus S: Macrobrachium rosenbergii I: Chironomus plumosus Hexagenia bilineata

Mu: Mytilus edulis W: Daphnia magna

n, st, 0.16 ppm, 22◦ C, 0.5 hr/na n, st, 0.14 ppm, 22◦ C, 0.5 hr/na n, st, 53 ppb, 23◦ C, 21 days/na y, st, 2 ppb, 21◦ C, 22 days/na n, st, 0.2 ppm, na, 7 days/4 days

na na na na 3 days

na na

1.4–1.9 1.5–1.8 3.4 2.6 1.2 1.8 2.9–3.6

1.8 days

na na

3 days

<1 day na na 0.7 day 14 days 2.3 days

32 days

na 144 days

CL50 e

–2.0

1.2–1.4 1.9–2.2

n, fl, 28–75 ppb, 20–22◦ C, 5 days/na n, fl, 28–75 ppb, 20–22◦ C, 5 days/na n, rn, 0.02–0.2 ppm, 22◦ C, 30 days/ 30 days n, fl, 28–75 ppb, 20–22◦ C, 5 days/na n, fl, 28–75 ppb, 20–22◦ C, 5 days/na

2.1

1.0 1.6–1.7 1.5–1.6 2.0 3.5 0.5–0.9

n, fl, 3–35 ppb, 10–15◦ C, 9 days/20 days n, st, 2 ppb, na, 2 days/na y, st, 10–40 ppb, na, 3 days/na n, fl, 3–30 ppb, 10–15◦ C, 9 days/20 days n, fl, 4.2 ppb, 22◦ C, 14 days/15 days n, rn, 0.02–0.2 ppm, 22◦ C, 30 days/ 30 days n, fl, 4.2 ppb, 22◦ C, 14 days/15 days

2.2

n, st, na, 4◦ C, 6 hr/7 days

S: Mysis relicta

107 Molinate (3.21) 108 Pirimicarb (1.7)

0.4 4.3

n, rn, 1–9 ppm, 15◦ C, 3–7 days/na n, fl, na, 4◦ C, 6 hr/7 days

Mu: Mytilus edulis A: Pontoporeia hoyi

101 Carbaryl (2.36)

log BCFd

Experimental conditionsc

Speciesb

No. Pesticide (log Kow a )

Table 6 (continued)

Manthey et al. (1993) Manthey et al. (1993) Böttcher and Schroll (2007) Feurtet-Mazel et al. (1996) Yu-yun et al. (1993)

Sanders and Hunn (1982) Sanders and Hunn (1982)

Wang et al. (1992b)

Sanders and Hunn (1982) Sanders and Hunn (1982)

Uno et al. (1997)

Donkin et al. (1997) Landrum and Dupuis (1990) Landrum and Dupuis (1990) Watanabe et al. (1985) Kusk (1996) Kusk (1996) Watanabe et al. (1985) Uno et al. (1997) Wang et al. (1992b)

References

36 T. Katagi

Triazines 127 Atrazine (2.61)

125 Chlomethoxyfen (4.4)

Diphenyl ethers 124 Chlornitrofen (CNP) (4.97)

123 Metsulfuron-methyl (2.2)

122 Chlorsulfuron (2.0)

121 CCU (3.84)

No. Pesticide (log Kow

a)

Sn: Lymnaes palustris G: Scenedesmus acutus Chlorella sp. Pediastrum sp. D: Cyclotella gamma

C: Corbicula fluminea S: Macrobrachium rosenbergii

C: Corbicula japonica Sn: Cipangopludina chinensis G: Scenedesmus quadricauda B: Microcystis aeruginosa D: Aulacpseira granulata Mu: Mytilus edulis

Mu: Mytilus edulis

G: Chlorella fusca

G: Scenedesmus subspicatus G: Chlorella fusca

Speciesb

1.9–2.1∗ 1.8–2.2∗ 2.7 1.8 1.1

n, st, 1–50 ppb, 25◦ C, 1 day/na n, st, 1–50 ppb, 20◦ C, 1 day/na n, fl, 2–17 ppb, 10–15◦ C, 9 days/20 days n, rn, 0.3 ppm, 22◦ C, 30 days/30 days n, rn, 2–20 ppb, 22◦ C, 30 days/30 days

0.6–0.9 1.5 2.2∗ 2.5∗ 1.6∗

1.6–2.5∗

n, st, 1–50 ppb, 25◦ C, 1 day/na

y, fl, 5–125 ppb, na, 21 days/na n, st, 11 ppb, 30◦ C, 8 hr/na n, st, 40 ppb, 20◦ C, 1 day/na n, st, 40 ppb, 20◦ C, 1 day/na n, st, 40 ppb, 20◦ C, 1 day/na

3.8 2.6

3.4

0.9 (pH 6), 1.7 (pH 5) 0.0 (pH 6), 1.2 (pH 5)

2.5–3.8

log BCFd

n, fl, 0.8–12 ppb, 10–15◦ C, 9 days/ 20 days n, fl, 1.7 ppb, 22◦ C, 14 days/15 days n, fl, 1.7 ppb, 22◦ C, 14 days/15 days

n, st, 7.6 ppm, 22◦ C, 2 hr/na

n, st, 7.2 ppm, 22◦ C, 2 hr/na

n, st, 0.2 ppm, na, 7 days/4 days

Experimental conditionsc

Table 6 (continued)

na na na na na

4 days <5 days

na na 1.6 days

na

14 days 3 days

4–8 days

na

na

1 days

CL50 e

Baturo and Lagadic (1996) Böhm and Müller (1976) Tang et al. (1998a) Tang et al. (1998a) Tang et al. (1998a)

Wang et al. (1992b) Wang et al. (1992b)

Guanzon et al. (1996) Guanzon et al. (1996) Watanabe et al. (1985)

Guanzon et al. (1996)

Uno et al. (1997) Uno et al. (1997)

Watanabe et al. (1985)

Fahl et al. (1995)

Fahl et al. (1995)

Yu-yun et al. (1993)

References

Bioconcentration, Bioaccumulation, and Metabolism 37

4-Chloroaniline (1.83) Phenol (1.46)

Salicylic acid (2.26)

o-Toluidine (1.32)

Trinitrotoluene (1.6)

Tri-n-butyltin (7.35)

–

–

–

15

16

–

Miscellaneous – Aniline (0.9)

130 Terbutryn (3.74)

129 Cyanazine (2.22)

No. Pesticide (log Kow

a)

1.6∗ 0.98 2.3–2.5 0.6–0.7 0.3–1.0 2.3–4.1 1.7 1.1 1.5

n, st, 40 ppb, 20◦ C, 1 day/na n, st, 57 ppb, 25◦ C, 6 days/na n, st, 0.27 ppm, 10◦ C, 1 day/na n, st, 23 ppb, 20◦ C, 1 day/na n, fl, 0.17–0.3 ppm, 8–20◦ C, 7–8 days/na n, fl, 10–40 ppb, 24◦ C, 22 days/na n, st, 14 ppb, 25◦ C, 4 hr/3 days n, st, 0.67 ppm, 15◦ C, 2 days/2 days n, st, 0.62 ppm, 15◦ C, 2 days/2 days

Synedra acus Mc: Hydrilla verticillata Periphyton community W: Daphnia magna Daphnia pulicaria

ca 3 ca 3 0.6 0.7

n, st, 50 ppb, 20◦ C, 4 days/na n, st, 50 ppb, 20◦ C, 4 days/na n, st, 5 ppm, 15◦ C, 1 day/1 days n, st, 5 ppm, 15◦ C, 1 day/1 days

Oy: Crassostrea gigas

0.4 1.2 0.6 4.5∗

0.5

n, st, 0.4–2 ppm, 20–22◦ C, 1 day/1 days

n, st, 50 ppb, na, 1 day/na n, rn, 0.28 ppm, 23◦ C, 6 days/4 days n, st, 50 ppb, na, 1 day/na n, st, 20 ppb, 20◦ C, 4 weeks/na

2.3–2.9∗

n, st, 5.7 ppb, 22◦ C, 2 hr/na

I: Chironomus tentans A: Hyalella azteca O: Lumbriculus variegatus G: Ankistrodesmus falcatus

1.9

n, st, 0.5–1.3 ppm, 20–22◦ C, 1 day/1 day

G: Scenedesmus quadricauda G: Selenastrum capricornutum G: Scenedesmus quadricauda G: Scenedesmus subspicatus Mc: Lemna minor Mu: Mytilus edulis

G: Scenedesmus quadricauda I: Chironomus tentans A: Gammarus fossarum Is: Asellus aquaticus

log BCFd

Experimental conditionsc

Table 6 (continued)

Speciesb

na >4 days na na

0.4 hr

na 0.4 hr

na

<1 hr

na

<1 hr

<8 hr 10 h 7 hr

na

na na na na na

CL50 e

Wang and Lay (1989) Knezovich and Crosby (1985) Knezovich and Crosby (1985) Belden et al. (2005) Sims and Steevens (2008) Belden et al. (2005) Maguire et al. (1984)

Wang and Lay (1989)

Halling-Sørensen et al. (2000) Hardy et al. (1985)

Hardy et al. (1985)

Muir et al. (1982) Richter and Nagel (2007) Richter and Nagel (2007)

Tang et al. (1998a) Hinman and Klaine (1992) Nikkilä et al. (2001) Nikkilä et al. (2001) Heisig-Gunkel and Gunkel (1982) Aly et al. (1984)

References

38 T. Katagi

MSMA (–3.1)

I: Chironomus tentans C: Tapes philippinarum W: Daphnia pulex Co: Acanthocyclops robustus Os: Eucypris virens I: Chironomus annularius W: Daphnia magna G: Nannochloris oculata W: Daphnia magna

Mc: Eichhornia crassipes Ceratophyllum demersum Cr: Procambarus clarkii Mc: Potamogeton pectinatus

Speciesb

n, st, 0.4 ppb–1 ppm, na, 2 days/na n, st, 0.4 ppb–1 ppm, na, 2 days/na n, st, 0.1 ppb, 22◦ C, 150 hr/na n, st, 1.5–4.4 ppm, 22◦ C, 4 days/na n, st, 1.5–4.4 ppm, 22◦ C, 4 days/na

n, st, 7.8 ppb, 25◦ C, 4 hr/3 days n, st, 2 ppb, 22–23◦ C, 12 days/12 days n, st, 0.4 ppb–1 ppm, na, 2 days/na n, st, 0.4 ppb–1 ppm, na, 2 days/na

n, st, 5–10 ppm, 25◦ C, 6 weeks/na n, st, 5–10 ppm, 25◦ C, 3 weeks/na n, rn, 0.5–50 ppm, 22◦ C, 8 weeks/8 weeks n, st, 1 ppm, 20◦ C, 2 weeks/na

Experimental conditionsc

Table 6 (continued)

1.0 1.8 2.3 2.0 1.7

1.4 1.6 –0.2 to 0.9 2.0 (shoot), 1.6 (root) 2.1 1.7 1.4 1.7

log BCFd

na na na na na

<8 hr 2 days na na

na na >8 weeks na

CL50 e

Chaton et al. (2002) Chaton et al. (2002) Liu et al. (1996) Ferrando et al. (1996) Ferrando et al. (1996)

Muir et al. (1982) Ueda et al. (1988) Chaton et al. (2002) Chaton et al. (2002)

Anderson et al. (1981) Anderson et al. (1981) Naqvi et al. (1990) Marguis et al. (1981)

References

b Designation

by EPI-Suite (USEPA 2008) or experimental data therein. of species. A, amphipod; Ab, abalone; B, blue-green alga; C, clam; Ci, ciliate; Cr, crayfish; D, diatom; E, Euglenophyta; G, green alga; I, aquatic insect; Is, isopod; Mc, macrophyte; Mu, mussel; O, oligochaete; Os, ostracod; Oy, oyster; P, polychaete; S, shrimp; Sn, snail; W, water flea. c Application of pesticide to water; presence (y) or absence (n) of sediment in the system, exposure condition (static (st)/renewal (rn)/flow-through (fl)), concentration, temperature (◦ C), periods of exposure/elimination. d Experimentally obtained bioconcentration factor based on the overlying water concentration in the whole body (or each tissue). The asterisk means dryweight-basis. e 50% clearance time. If the biphasic elimination is observed, the CL value of the faster elimination is included. 50 na, Not available; Co, copepod; R, rotifer; CNA, 4-Chloronicotinanilide; CCU, 1-(4-chlorophenyl)-3-(2-chlorobenzoyl)urea; MSMA, monosodium methanearsonate; MET, monochloro derivative of diclobutrazole.

a Estimated

134 MET (3.2) 136 Tetradifon (4.61)

132 Pyrazolate (3.9) 133 Fipronil (4.0)

131 Fluridone (3.16)

17

No. Pesticide (log Kow

a)

Bioconcentration, Bioaccumulation, and Metabolism 39

40

T. Katagi

The elimination of some pesticides appears to be highly dependent on species. Lindane (30) showed a higher bioconcentration and longer CL50 values in polychaetes than in bivalves and isopods (Ernst 1979; Thybaud and Caquet 1991; Thybaud and Le Bras 1988). Significant differences in elimination were observed for the more hydrophobic pesticides, pentachlorophenol (32), DDT (33), and its metabolite DDE (34). Molluscs rapidly eliminated (32), which has a CL50 of less than 1 day (Shofer and Tjeerdema 1993; Tjeerdema and Crosby 1992), whereas 2–3 weeks is necessary for elimination from polychaetes and shrimp (Ernst 1979; Landrum and Dupuis 1990). Significant differences in CL50 (by 100 days) were reported for (33) and (34), among amphipods (Lotufo et al. 2000). Lower BCF values primarily ranging from 1 to 2 were reported for organophosphorus pesticides having moderate hydrophobicity (Table 4). Chlorpyrifos (47) has larger BCF values, by an order of 1–2, in the isopods Asellus aquaticus and in Artemia sp., compared with other aquatic organisms (Cid Montañés and van Hattum 1995; Varó et al. 2000). A similar observation was reported for fenitrothion (41), diazinon (48), and malathion (57) in shrimp and copepods (Kashiwada et al. 1995b). Higher bioconcentration rates in these species are likely to originate from lower metabolic activities. The clearance of organophosphorus pesticides is rather rapid, with CL50 values being less than 4 days. The pyrethroid insecticides, listed in Table 5, show log BCF values of 2–4, which is smaller by about one order of magnitude than would be expected from the regression equations in Table 1. Organochlorine insecticides such as toxaphene (28) and hexachlorobenzene (31) have log Kow values of approximately 6. Generally, organochlorine pesticides are considered to be resistant to metabolism although they show log BCF values of 4–6 in molluscs, which are near predicted levels, but these values are higher than those seen for pyrethroids (2–4). Therefore, the pyrethroids are probably metabolized through ester cleavage and hydroxylation, which renders their end residues more hydrophilic than their parent compounds. Bioconcentration values for pesticides other than those mentioned above are presented in Table 6. BCF values of less than 1000 have been reported for ionizable acids, esters, amides, carbamates, ureas, and triazines. Many pesticides, with such functionalities, have chemical groups that assist in hydrogen bonding and are more easily metabolized. Such groups produce BCF values that are generally lower than would be predicted from the regression equations in Table 1. In contrast, the log BCF value of carbaryl (101) (log Kow = 1.85) in the amphipod Pontoporeia hoyi is 4.3, much higher than the corresponding value (2.2) in Mysis relicta. The difference is explained by the very slow elimination and from the lower metabolic activity (Landrum and Dupuis 1990). The ionizable salicylic acid showed unexpectedly high bioconcentration (log BCF = 3) in green algae, which may be related to its effect on algal growth (Wang and Lay 1989). Furthermore, the very hydrophilic MSMA (17) was reported to be moderately bioconcentrated in aquatic plants and retained more residues in roots as compared with stem and leaves; it is thought that some mechanism, other than partition to a lipid fraction, produced this behavior (Anderson et al. 1981). From the foregoing, one can conclude that two factors govern what actual BCF values are achieved: (a) one is the inherent log Kow value of a tested substance

Bioconcentration, Bioaccumulation, and Metabolism

41

Mollusks

6

Crustacea 6 4

log BCF

log BCF

4

2

2 0

–4

0 0

2

4

6

–2

0

–4

–2 log Kow

Algae

4

2

4

6

8

0

log Kow

Insecta

6

log BCF

log BCF

6

2

–2

8

4

2

0

0

2

4 log Kow

6

8

0

2

4 log Kow

6

8

Fig. 3 Relationship between BCF and log Kow in aquatic organisms for pesticides

and (b) the other is the balance struck between a chemical’s hydrophobicity and other factors, predominant of which is ease of metabolism under test conditions. Irrespective of test chemical class and species of aquatic organism, all BCF values correlate with log Kow values, which is reflected in the equation of log BCF = 0.427 log Kow + 0.462 (r2 = 0.337). This means that 34% of the variance in BCF can be accounted for by hydrophobicity of a chemical. Since each organism has a specific physiology that significantly affects uptake and elimination processes, each organism’s profile portends what probable bioconcentration levels may be reached. This can be demonstrated through the following analysis of BCF values for several phyla and classes of organisms (Fig. 3). The result of each linear regression analysis is described below:

Molluscs Algae Crustacea Insecta

log BCF = 0.579 log Kow log BCF = 0.305 log Kow log BCF = 0.537 log Kow log BCF = 0.493 log Kow

− 0.391 + 1.05 + 0.150 − 0.0407

(r2 = 0.442, n = 79) (r2 = 0.279, n = 77) (r2 = 0.381, n = 74) (r2 = 0.411, n = 26)

The higher correlations in molluscs and crustacea agree with less variation in the regression lines reported for mussels, bivalves, daphnids, and amphipods (Table 1). In contrast, the variance in regression seems larger in algae that exhibit a significant variety of shape, size, and surface structure; these physical features may explain the lower correlation for algae.

42

T. Katagi

To examine the effect of a chemical class on BCF, the correlation of BCF with log Kow was analyzed for chlorinated hydrocarbons (Table 3) and organophosphorus pesticides (Table 4). Correlations were moderately good only for crustaceans in the former class and molluscs in the latter, as described below. The other organisms only exhibited a very weak correlation (r2 < 0.1). Chlorinated hydrocarbons (log Kow = 3.83−6.91), crustacean: log BCF = 0.835 log Kow − 1.47 (r2 = 0.39, n = 31) log BCF = −0.173 (log Kow − 7.98)2 + 4.36 (r2 = 0.405, n = 31) Organophosphorus pesticides (log Kow = 0.78−4.96), molluscs: log BCF = 0.543 log Kow − 0.436 (r2 = 0.438, n = 17) log BCF = −0.106 (log Kow − 5.66)2 + 2.11 (r2 = 0.471, n = 17) The linear regressions are, respectively, close to those for organochlorines, PAHs in Daphnia pulex (Hawker and Connel 1986), and insecticides in Mytilus edulis (Zaroogian et al. 1985). The quadratic equation provided slightly better correlation in both cases, which is in accord with the theoretical considerations discussed in the previous section. These analyses show that the BCF values for pesticides and other simple organic chemicals are highly dependent not only on the species of aquatic organism but also on the chemical class under test.

3 Bioaccumulation 3.1 Controlling Factors The presence of bottom sediments and food webs in the real aquatic environment complicates the bioconcentration process. Bottom sediments serve as important habitats for many aquatic organisms such as molluscs, larvae of chironomids, and oligochaetes. It is important to consider a host of factors when generating bioaccumulation data, including exposure levels of a chemical in interstitial water, presence of sediment particles or detritus, dietary habits of each tested organism, etc. (Connell 1988; Landrum and Fisher 1998; Miyamoto et al. 1990). BAF is similar to BCF, but the denominator Cpm (see Eq. (13)) includes the accumulating effect of chemical exposure not only in media such as water and sediment but from the diet as well: BAF = Cpb /Cpm .

(13)

Fewer BAF than BCF values are available in the literature for pesticides. Where BAF values are reported, most have been performed on sediment dwellers (Table 7). BAF values, when available, are generally less than 10, but tend to be higher for persistent organic substances, e.g., benzo[a]pyrene (7), in oligochaetes. Such persistent organics are generally not easily metabolized and have longer CL50 values as well. Similarl to the relationship that exists between BCF values and log Kow values for any substance, the log Kow value of a chemical also influences what its BAF, calculated as Cpb /Cps using its concentration in sediment (Cps ), is likely to be. Oliver

Photo-dieldrin Lindane α-isomer of (31) Hexachlorobenzene Pentachlorophenol DDT DDE Terbutryn

Bentazone Pendimethalin Ioxynil

23 30

31 32 33 34 130

137 143 145

Hyalella azteca I: Chironomus tentans O: Lumbriculus variegates W: Daphnia pulex O: Tubifex tubifex W: Daphnia magna O: Tubifex tubifex O: Lumbriculus variegatus O: Tubifex tubifex O: Tubifex tubifex Is: Asellus aquaticus A: Gammarus fossarum O: Lumbriculus variegatus O: Lumbriculus variegatus O: Lumbriculus variegatus

A: Gammarus fossarum

s, st, 0.4 ppm, 11 days/15 days s, st, 0.4 ppm, 11 days/15 days f (leaf), st/fl, 0.4–0.7 ppb, 15◦ C, 2 days/ 2 days f (leaf), st/fl, 0.4–0.7 ppb, 15◦ C, 2 days/ 2 days s, st, 13.5 ppt, 20◦ C, 3 days/3 days s, st, 13.6 ppt, 20◦ C, 3 days/3 days s, st, 14 ppt, 20◦ C, 3 days/3 days f (algae), st, 1 ppb, na, 36 hr/7 days s, st, 0.74 ppm, 8◦ C, 79 days/84 days f (algae), st, 10 ppb, 20◦ C, 2 days/na s, st, 0.9 ppm, 8◦ C, 79 days/84 days s, st, 0.4 ppm, 20◦ C, 14 days/na s, st, 0.1 ppm, 8◦ C, 79 days/84 days s, st, 0.29 ppm, 8◦ C, 79 days/84 days f (leaf), st/fl, na, 15◦ C, 2 days/1 days f (leaf), st/fl, na, 15◦ C, 2 days/3 days s, st, 0.4–1.2 ppb, 20◦ C, 10 days/na s, st, 0.1–0.3 ppb, 20◦ C, 10 days/na s, st, 0.1–0.4 ppb, 20◦ C, 10 days/na

15◦ C, 15◦ C,

Experimental conditionsb 0.8 days 1.4 days 2.5 h 8.7 h 1.2 days 4.8 hr 2.9 days 4 days < 5 days na 24 days na 53 days 80 days 4.3 hr 2.9 hr na na na

–1.52 –0.68 –0.8 –0.02 1.2 0.88 –0.5 0.49 1.46 –0.3 0.63 –0.37 –1.4 –0.3 to 0.6 –1.0–0.9 0.4–1.7

CL50 d

1.32 0.69 –1.4

log BAFc

Schuler et al. (2003) Schuler et al. (2003) Schuler et al. (2003) Khan et al. (1975) Oliver (1987) Canton et al. (1975) Oliver (1987) Nikkilä et al. (2003) Oliver (1987) Oliver (1987) Richter and Nagel (2007) Richter and Nagel (2007) Mäenpää et al. (2003) Mäenpää et al. (2003) Mäenpää et al. (2003)

Richter and Nagel (2007)

Christensen et al. (2002) Christensen et al. (2002) Richter and Nagel (2007)

References

b Application

by EPI-Suite (USEPA 2008) or experimental data therein. of pesticide to sediment (s) or food (f), exposure condition (static (st)/flow-through (fl)), concentration, temperature (◦ C), periods of exposure/elimination. c Bioaccumulation factor based on the sediment or food concentration in the whole body. d 50% clearance time. na, Not available.

a Estimated

Benzo[a]pyrene

7

P: Arenicola marina Nereis diversicolor Is: Asellus aquaticus

Speciesa

Table 7 A summary of bioaccumulation studies of selected pesticides and other chemicals in aquatic organisms

Pyrene

Pesticide

5

No.

Bioconcentration, Bioaccumulation, and Metabolism 43

44

T. Katagi

(1984, 1987), using spiked sediments, investigated the bioaccumulation of various chlorinated aromatic compounds and PCBs in oligochaetes. BAF vs. log Kow plots of tested chemicals showed a parabolic profile, with maximum values approaching 6–10 for substances having log Kow values of 6. Generally, as worms eliminated less chemical, and the log Kow of a chemical increased, higher accumulation occurred. However, further increases of log Kow above 6 significantly reduced the uptake of a bioavailable chemical dissolved in water, because it became more tightly bound. The chemical adsorbed to sediment and the BAF value decreased; therefore, a dynamic balance exists between rates of adsorption/desorption of a chemical in sediment and how rapidly the worms can eliminate the chemical. 3.1.1 Bottom Sediment Because many invertebrates and their larvae live in and on bottom sediments, the bioaccumulation of chemicals sourced either from the organic carbon or from interstitial water phase should be considered. Bartlett et al. (2004) demonstrated that the primary route for bioaccumulation for tributyltin (16) into the amphipod Hyalella azteca was exposure to the chemical dissolved in water. The authors observed similar BAF values, whether the amphipods were kept on sediment or were isolated in overlying water by a cage. Burgess and McKinney (1999) separately examined the bioaccumulation of PCB congeners in two bivalves from whole sediment or in overlying or interstitial waters; there was better correlation of BAF with the PCB concentration in the overlying water, which was believed to have resulted from bivalve-specific behavior. A similar trend was reported for the bioaccumulation of chlorobenzenes in Chironomus decorus, and was explained by reduced bioavailability caused by DOC content of interstitial water, or adsorption onto sediment particles (Knezovich and Harrison 1988). Tenfold higher concentrations of chlorpyrifos (47) were observed in interstitial water than those predicted by partition onto sediments; such adsorption to sediments reduced the toxicity to C. tentans, but the correction for association with DOC gave similar LC50 values as those observed in the wateronly system (Ankley et al. 1994). In a very hydrophobic pyrethroid, the immediate adsorption to river and pond sediments caused reduction of its concentration in the overlying water, which resulted in an increase of apparent BAF values in C. tentans larvae. However, a similar degree of bioaccumulation to that in water-only exposure was obtained if residues of the pyrethroids in interstitial water were instead used to estimate BAF (Muir et al. 1985b). Paraquat (149) is a cationic herbicide. Because humic substances are usually negatively charged, electrostatic attraction would facilitate the formation of (149)humic substance complexes. Wiegand et al. (2007) demonstrated that reduced uptake of (149) into oligochaetes, by addition of lake dissolved organic matter, decreased the bioavailability of (149). When the effect of sediment adsorption is mitigated by maintaining the water concentration of a chemical constant in water, BAF values for hexachlorobenzene (31) were not significantly decreased in amphipods (Schuytema et al. 1990). Both organic carbon content and particle size distribution in sediments affected levels of bioaccumulation. Fine particles are known to adsorb

Bioconcentration, Bioaccumulation, and Metabolism

45

more chemical than do coarse fractions (Katagi 2006). This fact explained the different BAF values of four herbicides seen for Lumbriculus variegatus in four sediments (Mäenpää et al. 2003). These observations strongly suggest that only the chemical freely dissolved in an overlying or interstitial water phase is bioavailable to aquatic organisms. Ecological factors associated with aquatic organisms have been reported to sometimes affect bioaccumulation. Fluoranthene (6) was more highly bioaccumulated in the marine polychaete Nereis virens, when this organism co-existed with higher numbers of the amphipod Corophium volutator; the increase in bioaccumulation of (6) was explained to result from bioturbation of bottom sediment by the amphipods (Ciarelli et al. 2000). Among copepods, the deeper burrowing Amphiascus tenuiremis showed higher bioaccumulation of azinphos-methyl (61) than the epibenthic Microarthridion littorale, due to a different frequency of exposure (Klosterhaus et al. 2003). DiToro et al. (1991) developed a methodology to assess bioaccumulation based on the equilibrium partitioning of a chemical between sediment carbon and interstitial water phases. The fraction of a chemical dissolved in interstitial water at concentration Cd is considered to be the only bioavailable amount; this amount can be estimated from information on sediment and DOC content (from foc , Koc , m, and Kdoc terms in Eq. (4)). The concentrations of a chemical in an organism (Cpb ) and in sediment (Cps ) are expressed below, and allow the biota-sediment accumulation factor (BSAF) that is based on the normalization to fb and foc to be defined as presented in Eq. (14):     BSAF = Cpb /fb / Cps /foc ,

(14)

where Cpb = KL (lipid-water partition coefficient); fb = weight fraction of lipid in an organism × Cd ; Cps = foc × Koc × Cd (linear isotherm is assumed). When dietary uptake by ingestion is neglected, BASF is equal to KL /Koc . Since the KL and Koc values can be expressed by a function of Kow , the BSAF value would theoretically be a constant that is independent of organism and a chemical. The following relationship of Kow is known with Koc and KL for a chemical with a log Kow of less than 6: Koc = 0.41 × Kow (Karickhoff 1981) and KL = 0.048 × Kow (Mackay 1982). When the specific density of sediment and the lipid fraction of an organism are respectively assumed to be 1.5 kg L–1 and 1–5%, KL is equal to BCF/(0.01−0.05) and then the BSAF value is calculated to be 1.6–7.8. The BASF values determined for pesticides in aquatic organisms are quite variable. Such variability exists not only within each study but also among species and chemical classes (Table 8). Ingersoll et al. (2003) examined the bioaccumulation of DDT (33), its metabolites, and PAHs, with log Kow values ranging from 3 to 8 in oligochaetes. These authors reported that the plateau BSAF values were 1–8 for (33)

Pyrene Benzo[a]pyrene

Lindane Hexachlorobenzene DDT

DDE TCBP

Chlorpyrifos Azinphos-methyl

Permethrin Cypermethrin

Bentazone Pendimethalin

5 7

30 31 33

34 –

47 61

81 82

137 143

O: Lumbriculus variegates P: Nereis diversicolor Leitoscolopos fragilis O: Tubifex tubifex O: Tubifex tubifex A: Hyalella azteca Leptocherirus plumulosus O: Lumbriculus variegates O: Lumbriculus variegates A: Hyallela azteca I: Chironomus tentans O: Lumbriculus variegates Co: Amphiascus tenuiremis Microarthridion littorale O: Lumbriculus variegates W: Daphnia magna I: Chironomus tentans O: Lumbriculus variegates O: Lumbriculus variegates

Speciesa NA 1.2–1.3 1.4–1.7 2.9–5.2 2.9–5.2 0.7–1.8 1.9 ± 0.2 1.1–2.3 1.0 ± 0.3 NA NA 1.5–2.3 2.2 5.6 1.5–2.6 NA NA 1.23 1.23

fb b

b Wet

by EPI-Suite (USEPA 2008) or experimental data therein. weight fraction of lipid in an organism. C Source of sediment or soil. d Organic carbon %. NA, not available. TCBP, 3,4,3 ,4 -tetrachlorobiphenyl

a Estimated

Pesticide

No.

foc d 1.4–3.0 0.6 0.6 2.0 2.0 0.4–0.6 1.78 1.3–7.9 0.29 0.29 0.29 1.3–7.9 3.85 3.85 1.3–7.9 1–13 1–13 0.54–24 0.54–24

Sourcec Lake × 2 Marine Marine Artificial soil Artificial soil Lake Marine Lake × 2 Creek Creek Creek Lake × 2 Estuarine Estuarine Lake × 2 River × 3 River × 3 Lake × 4 Lake × 4 0.42–0.59 0.028 1.40 2.81–2.93 3.13–4.96 0.44–2.08 2.51 ± 0.54 2.00–5.31 0.05–0.33 1.73–3.85 0.47–1.33 1.49–6.54 26.8 2.2 2.17–3.79 0.08–0.31 0.08–0.63 2.9–14.9 2.2–5.5

BSAF

Leppänen and Kukkonen (2006) Driscoll and McElroy (1996) Driscoll and McElroy (1996) Egeler et al. (1997) Egeler et al. (1997) Lotufo et al. (2001a) Lotufo et al. (2001b) You et al. (2006) Leppänen et al. (2003) Leppänen et al. (2003) Leppänen et al. (2003) You et al. (2006) Klosterhaus et al. (2003) Klosterhaus et al. (2003) You et al. (2006) Maund et al. (2002) Maund et al. (2002) Mäenpää et al. (2003) Mäenpää et al. (2003)

References

Table 8 Biota-sediment accumulation factors (BSAF) of pesticides and chemicals in aquatic organisms

46 T. Katagi

Bioconcentration, Bioaccumulation, and Metabolism

47

and its metabolites and 0.4–3 for PAHs. These values were somewhat proportional to the hydrophobicity of the studied chemicals and were almost constant at log Kow of 3–8. Values that were lower than expected probably originated from metabolism effects or reduced bioavailability from association with DOC. Low metabolic activity in the isopod Asellus aquaticus was used by Van Hattum et al. (1998) to explain lower than expected BASF values (0.1–1) in PAH-contaminated littoral sediments. Maund et al. (2002) estimated the BASF values of cypermethrin (82) in D. magna and Chironomus tentans to, respectively, be 0.08–0.31 and 0.08–0.63, in the three river sediments with different foc values. Similar BSAF values were obtained for the two species, but these values decreased as bioavailability was reduced in sediment from association with organic carbon. A higher correlation of BSAF with Kdoc than Koc for dieldrin (22) in oligochaetes indicated the importance of DOC in determining the bioavailable fraction (Standley 1997). Nikkilä et al. (2003) reported reduced bioavailability of phenol derivatives in Lumbricilus variegatus from sediment with higher organic carbon. Because the soluble fraction of a chemical in interstitial water is considered to be in equilibrium with DOC, and sediment particle phases (DiToro et al. 1991; Kraaij et al. 2003), one can correct for the BSAF in the bioavailable fraction by using Tenax resin or solid-phase micro-extraction (SPME) glass fiber. Leppänen et al. (2003) have examined the bioaccumulation of 3,4,3 ,4 -tetrachlorobiphenyl and desorption profiles in worms, chironomids, and amphipods from sediment by using Tenax resin. The correction of the BSAF values in Lumbriculus variegatus failed when the rapid and slower desorption fractions were used; this may be caused by the non-linearity of desorption and limits in using the steady-state lipid concentration. The good correlation between the concentration of DDE (33), chlorpyrifos (47), and permethrin (81) in the worm and the fraction of fast desorption (for 6 hr) from sediment, by using Tenax, was reported by You et al. (2006). They also found a linear correlation between the concentrations in the worm and SPME fibers. However, the slower equilibration in fibers than in biota resulted in a slope in the regression line of less than one. Similar profiles for (81) in daphnids have been reported by Yang et al. (2006). 3.1.2 Dietary Route Aquatic organisms usually ingest prey, sediment particles, and detritus contaminated by chemicals, and this may affect bioaccumulation rates. In benthic organisms, the water ventilation rate is usually several orders of magnitude higher than the ingestion rate. Although uptake via water is important for most chemicals, the dietary route is also important for many hydrophobic compounds. Furthermore, the gut clearance rate is one of the important factors that control bioaccumulation in benthic invertebrates. Brook et al. (1996) gravimetrically examined the gut clearance rate in mayfly nymph, larval midge, and a freshwater worm, and reported 75%, 90%, and 100% depletion, respectively, during a 12-hr period. To evaluate the major uptake route and the contribution of metabolism, Richter and Nagel (2007) examined bioaccumulation via water uptake and dietary routes of terbutryn (130) and benzo[a]pyrene (7) in the amphipod Gammarus fossarum, and in the isopod Asellus

48

T. Katagi

aquaticus. Much higher BCF than BAF values were observed in all cases, which shows that water exposure is the main route of uptake. The more hydrophobic (7) was more highly metabolized when taken in via dietary exposure. The importance of the dietary route was demonstrated in four benthic invertebrates by Gaskell et al. (2007), who studied the bioaccumulation of the very hydrophobic dimethyldidodecylammonium chloride in treated clay. The feeding rate could not explain the variation of BAF among species, but both gut passage time and gut surfactancy appeared to contribute to the differences. By separately exposing chlorpyrifos (47) and aqueous solutions of it to several kinds of particulates, Bejarano et al. (2003) estimated the tissue absorption efficiency in clams. Results expressed as absorption efficiency in percent (AE%) were 23–31% and 7–17% for particulates and dissolved fractions, respectively. Algal cells constituted the best food for clams, indicating that bioaccumulation proceeds via ingestion of the contaminated particles rather than via filtration of dissolved chemical. The contribution of ingestion to bioaccumulation has been elaborately examined in the oligochaete Lumbriculus variegatus by utilizing their reproduction mode, wherein the posterior end of the organism does not ingest sediment for about 1 week after segmentation (Leppänen and Kukkonen 1998). By comparing the accumulated residues of pyrene (5) in oligochaetes that either normally ingest or do not ingest food, the authors estimated the contribution of ingestion to bioaccumulation to be approximately 61% on a total radioactivity basis (Conrad et al. 2002). When metabolic transformation is taken into account, the corresponding contribution for (5) and benzo[a]pyrene (7), respectively, becomes 34–38% and 73–75%, indicating less transformation of (7) (Leppänen and Kukkonen 2000). By using a similar method, Conrad et al. (2000) observed the reduction in bioaccumulation of dichlorophenol (DCP) by 20% in non-ingesting oligochaetes; the lower contribution of ingestion is considered to stem from less adsorption of DCP to sediment. These authors also investigated the effect of sediment aging on a dietary uptake route of (5). The result was that an increased fraction of sequestered (5) had a reduced contribution to bioaccumulation after aging. Higher metabolism in an organism generally reduces bioaccumulation. The substance (7), when spiked with sediment, was taken up by several marine species including polychaetes, amphipods, and bivalves, and the uptake showed an inverse correlation between BAF and extent of metabolism (Driscoll and McElroy 1996; Rust et al. 2004). In another example, (5) was metabolized by polychaetes during rapid bioaccumulation from spiked sediment (Christensen et al. 2002). The longer incubation resulted in higher BAF values, probably because of body size; there was no increase in feeding rate. By using a brook sediment contaminated with creosote, Lyytikäinen et al. (2007) investigated the time course of BAF in Lumbriculus variegatus for PAHs. The highest BAF values were observed just after exposure, followed by a PAH-specific decrease, which was explained by the extent of elimination and biotransformation. In a field monitoring experiment, higher residues of PCB than PAH were detected in fish, crustaceans, polychaetes, and molluscs, and residue levels increased with increasing activity of mixed-function oxidases; this

Bioconcentration, Bioaccumulation, and Metabolism

49

was understood to indicate some contribution from metabolism on bioaccumulation rates (Connor 1984). The effect on bioaccumulation of consuming contaminated food has also been investigated. Daphnids were reported to have increased BAF values (by a factor of 1–2) after exposure to tetradifon (136) (Ferrando et al. 1996), photo-dieldrin (23) (Khan et al. 1975), and α-hexachlorocyclohexane, when the water fleas were fed freshwater algae contaminated with these compounds. The bioaccumulation of atrazine (127) in Daphnia pulicaria was increased when they were fed contaminated food at 8◦ C; no clear dependency on food amount was observed, and no effect occurred at higher temperatures (Heisig-Gunkel and Gunkel 1982). In contrast, the bioaccumulation of dieldrin (22) (Reinert 1972), endosulfan (27) (DeLorenzo et al. 2002), and fenvalerate (85) (Day and Kaushik 1987) in water fleas was not significantly affected by the co-existence of contaminated algae; the explanation for fenvalerate might be accounted for by the reduced intake of algae, which resulted from toxicity. These results show the low probability of biomagnification for most pesticides, possibly as a result of their metabolic transformation (Connell 1988; Ellgehausen et al. 1980).

3.2 Bioaccumulation of Pesticides and Theoretical Approach Aquatic organisms may absorb dissolved chemicals directly from water through respiratory organs (e.g., gills), through the body surface, or may ingest chemicals through intake of contaminated sediments or prey. Therefore, any kinetic analysis of bioaccumulation should consider each of these exposure processes. The simplest model for bioaccumulation is that of a non-metabolized chemical dissolved in interstitial water or sediment. Egeler et al. (1997) utilized such a one-compartment model (Eqs. (6)–(8)) by using the initial concentration in sediment (Cps 0 ), rather than the Cpw , to analyze the uptake and elimination processes in oligochaetes. Under conditions in which a chemical is gradually degraded in sediment, Eq. (15) can be derived from Eq. (7) by conveniently incorporating the degradation rate (λ) as follows: 0 × [exp( − λ × t) − exp( − kPE × t)]. Cpb = [kU /(kPE − λ)] × Cps

(15)

This approach has been successfully applied to correct for the effect of timedependent change of chemical concentration in sediment, when the bioaccumulation of pentachlorophenol (32) and three herbicides were analyzed in oligochaetes and larval chironomids (Mäenpää et al. 2003, 2008). By using this method, Landrum (1989) has shown a negative linear correlation with log Kow in elimination rates for PAHs. Furthermore, the parabolic nature of sediment-based BAF was observed vs. log Kow that had its maximum at log Kow of 5. Markwell et al. (1989) applied a three-compartment model that assumed equilibrium among 15 lipophilic chlorinated organics, with benthic fauna and interstitial water. In this study, concentrations of these xenobiotics in interstitial water were in equilibrium with sediment. These authors found that the sediment-based BAF was weakly dependent on the Kow

50

T. Katagi

values for each chemical. However, BAF values, estimated from interstitial water concentrations, showed a significant correlation with Kow , i.e., log BAF = 1.11 × log Kow − 1.0 (r = 0.98). The differential equations that describe this model have been mathematically solved, and the corresponding parameters, such as adsorption and desorption rates between sediment and water and kU in Fig. 1b, were estimated by fitting the equation to experimental data from the oligochaete study (Gabric et al. 1990). A similar relationship of log BAF = 1.4 × log Kow − 5.5 (n = 27, r = 0.95) was obtained on an interstitial water basis. Shaw and Connell (1987) have introduced the use of the Freundlich isotherm, Eq. (10), between sediment and interstitial water in a kinetic analysis of PCB’s accumulation in mullet and polychaetes. Resulting estimates portended higher uptake rates of PCBs for polychaetes than for mullet, but elimination rates for the two species were comparable. The relationships of kU and kPE to log Kow have been further investigated for oligochaetes in the context of uptake from interstitial water and elimination of 15 chlorinated aromatics (Connell et al. 1988). The kU values showed a parabolic profile with a maximum at log Kow of 7.4, whereas kPE changed linearly with only weak correlation to log Kow . As a result, log BAF also displayed a parabolic shape when plotted against log Kow , and had a good correlation coefficient (r = 0.98). The obtained profiles were very similar to those in bioconcentration of a chemical from water. These kinetic analyses show that uptake of dissolved chemical from interstitial water is of importance for sediment-dwelling organisms in watersediment systems. Predominance of the water exposure route was also demonstrated for chironomids and amphipods. Muir et al. (1983) considered the additional term describing the uptake of four pesticides and two chemicals from sediment (Eq. (6)) to estimate the rate of each process in bioaccumulation by larval Chironomus tentans. The larger apparent kU values, when quartz sand was used instead of natural sediments, demonstrated the importance of adsorption onto sediments. However, the uptake rates on a wet weight basis were found to be insignificantly affected by the presence of sediment in the accumulation of anthracene (3) in Hyalella azteca (Landrum and Scavia 1983). Much slower uptake rates from sediment (one- to threefold) were observed in both studies, but a high water-sediment ratio (100:1) in the natural environment may reduce the importance of the sediment uptake route even though the concentration of a lipophilic chemical in sediment is much higher than in water. When aquatic organisms ingest contaminated sediments, both metabolism and gut clearance of the contaminating chemicals should be considered when characterizing dissipation from the body. Schuler et al. (2003) used a three-compartment model to estimate the kU , kM , kPE , and kME values (Fig 1b) in studying the bioaccumulation of sediment-spiked benzo[a]pyrene (7) in amphipods, larval midges, and worms. Only the larval midge showed significant metabolism (75% after 3 days), with comparable changes to kPE and kME values. Elimination of some chemicals sometimes display biphasic profiles in aquatic organisms (Egeler et al. 1997; Landrum and Scavia 1983; Muir et al. 1982; Richter and Nagel 2007; Shaw and Connell 1987). Such profiles result from various mechanisms that include reduced elimination of metabolites or formation of bound

Bioconcentration, Bioaccumulation, and Metabolism

51

residues. Furthermore, gut contents may also affect the shape of chemical elimination curves, because such contents may be rapidly or slowly eliminated from the body. Bartlett et al. (2004) demonstrated higher body residues of tributyltin (16) at an early stage of exposure, when Hyalella azteca were kept in a sediment-spiked water-sediment system, as compared with those kept in a cage above the sediment. The kinetic analysis of biphasic elimination showed that gut contents contributed 20–30% of total body burden, and the time for 50% clearance via gut elimination was estimated to be 6 h. The elimination of a chemical from the body was much slower, with the half-life of 8 days. If all processes relating to uptake and dissipation are included (i.e., uptake via gills, ingestion of food or sediment, metabolism and elimination), the kinetic and fugacity models may function to adequately elucidate the bioaccumulation profiles of a chemical in aquatic organisms. Gobas et al. (1988) developed the fugacity model for fish and proposed feeding rates and transport parameters in water and in lipid phases that lead to gastrointestinal absorption. For the filter feeding and detritivorous benthic invertebrates, the fugacity of PCB congeners in biota (fB ) was theoretically derived, and its applicability was checked by using monitoring data on sediment, caddisfly larvae, gammarus, mussel, and crayfish (Morrison et al. 1996). The ratio of fB to the fugacity in sediment (fS ) ranged from 0.3 to 7.7, and the BASF values were calculated to be 0.5–12.4 by using the relationship of fB /fS = 0.62 × BASF. The fB /fS values showed a tendency to follow a parabolic relationship with log Kow , and served to statistically better predict bioaccumulation than did the equilibrium partition approach. The most sensitive parameter was the fugacity ratio of digestivity and absorption of food and fraction of organic carbon or lipid in the diet that was removed by digestion. Furthermore, the fugacity model was applied to estimate bioaccumulation at a different trophic level (Hendriks et al. 2001). This model only weakly predicted the absorption process (r2 = 0.39), but was better in predicting elimination (r2 = 0.70) in several aquatic organisms that interact via food web. The estimated bioaccumulation values were generally in accord with observed ones for the compounds retaining log Kow values of 2–7. To analyze low BSAF values (0.01–0.1) seen in crayfish that were exposed to various PAHs in a creek, Thomann and Komlos (1999) successfully introduced a kinetic model and found that the relative contribution of the dietary exposure route toward water gradually increased at the log Kow values of PAHs above approximately 5–6, with its extent emphasized by alkyl substitution in the ring structures.

4 Metabolism 4.1 Metabolic Patterns Metabolism is one of the most important factors that govern the bioconcentration, bioaccumulation, and detoxification of pesticides. The metabolism of pesticides in fish has been thoroughly investigated by many researchers, and excellent reviews

52

T. Katagi

are available thereon (Edwards and Millburn 1985; Huckle and Millburn 1990; Schlenk 2005). There are commonly defined stages of metabolism. These are generally classified as phase-I and -II reactions that proceed successively to facilitate elimination of pesticides that have been taken up by aquatic organisms. As phase-I metabolic reactions proceed, hydrophilicity of the xenobiotic metabolites generally increases. This results from enzymatic oxidation and hydrolysis that produce metabolites with OH, COOH and NH2 , SH functional groups. These functional groups are then subject to conjugation with carbohydrates, glutathione, sulfate, and amino acids. Such metabolic profiles are also common in aquatic organisms as well as fish, but the contribution of each reaction is dependent on species (James 1986, 1987, 1994; James and Pritchard 1990; Lech and Vodicnik 1985; Miyamoto et al. 1990). The typical metabolic reactions reported for aquatic organisms (except fish) are presented in Table 9. Oxidation is usually catalyzed by mixed function oxidases, including cytochrome P450 enzymes. Such enzymes produce stepwise alkyl transformation that ultimately results in carboxyl groups, aromatic ring hydroxylation, desulfuration, and/or S-oxidation. Reductive dehalogenation and dehydrohalogenation, typical reactions for e.g., DDT (33), as well as reduction of nitro and S-oxide groups are also known. Hydrolysis, catalyzed by various esterases, is common in the metabolism of organophosphorus and pyrethroid pesticides. Transferase-catalyzed conjugation with glucose and its derivatives predominates in algae and macrophytes, and is also reported for other organisms, as is conjugation with glucuronic acid and sulfate. The acetylation, formylation, and conjugation of amino acids mostly proceed for amino and carboxyl groups after reduction of a nitro group or ester cleavage. Glutathione conjugation of pesticides in aquatic organisms is not commonly reported. In addition to the common reactions that have been characterized above, there are unique metabolic routes that have been reported, mainly for algae. Megharaj et al. (1994) found that the nitrite ion was produced after incubation of parathion (39) with green and blue-green algae; this interaction indicates fission of theC–N bond. The C–P bond was probably cleaved by the blue-green alga Anabaena sp. under phosphorus starvation conditions, since the addition of glyphosate (72) to a medium significantly increased the activity of extracellular alkaline phosphatase (Ravi and Balakumar 1998). The green algae Ankistrodesmus sp. was found to metabolize about half of available tributyltin (16) to the corresponding dibutyl derivative, with trace amounts of monobutyl one and inorganic tin, during a 1-month incubation; such action indicates cleavage of the C–Sn bond (Maguire et al. 1984). Greca et al. (1996b, 2003) reported the unique ring rearrangement reactions for androsta-1,4-dien derivatives and sinapic acid in green algae.

4.2 Enzymes Many kinds of enzymes play key roles in detoxifying pesticides and chemicals that are taken up by aquatic organisms from water and bottom sediments. In this section, the profiles of the reactions catalyzed by each enzyme type are described,

Bioconcentration, Bioaccumulation, and Metabolism

53

Table 9 Typical metabolic pathways of pesticides in aquatic organisms Metabolism

Type Reaction scheme

Oxidation

S-oxidation Desulfuration Others

O1 O2 O3 O4 O5 O6 O7 O8

R–CH3 → R–CH2 OH → R–CHO → RCOOH –O{N}–CH3 → [–O{N}–CH2 OH] → –O{N}H Ar–H → Ar–OH Ar–H → Ar–(OH)2 Quinone –S– → –S(O) – → –S(O2 ) – P=S → P=O, C–SO3 H → C–OH C=C → epoxide, ketone, NH2 →NHOH→NO, R–X→R–OH [X=NO2 ]

Dehalogenation Dehydrohalogenation Nitro group Multiple bond

R1 R2 R3 R4

Sulfone, sulfoxide Carboxyl ester Phosphoryl (sulfonyl) ester

R5 H1 H2

Amide Carbamate Urea Others

H3 H4 H5 H6

RX → R–H + X– [X = halogen] CH–CX → C=C + HX [X = halogen] –NO2 → [–NO]→ –NHOH → –NH2 –N=N– → –HN–NH–, –C≡C– → –C=C– → –CH–CH– –S(O2 ) – → –S(O) – → –S–Hydrolysis –C(=O)OR → –COOH + R–OH –P(=X) –YR → –P(=X) –OH + RYH [X, Y = O, S] ROS(=O)OR → ROH + R SO3 H –C(=O)NR– → –COOH + –NHR –NC(=O)O(or S)R → –NH + RO(or S)H –NHC(=O)NR– → –NH2 + RNH– –CN → –CONH2 → –COOH, R–X → R–OH [X = halogen]

Glucuronidation Glucosidation

C1 C2

Alkyl oxidation O (N)-dealkylation Ring hydroxylation

Reduction

Conjugation

Sulfation C3 Glutathione conjugation C4 N-acylation C5 Amino acid conjugation C6 Methylation C7 Miscellaneous M

R–XH → R–X–Gla [X = O, COO, S, NH] R–XH → R–X–Glu → R–X–(6O–R )Glu [X = O, COO, S, NH; R = acetyl, malonyl, pentosyl] R–X–H → R–X–SO3 H [X = O, NH] R–X + GSH → R–SG → R–Cys–Gly → R–Cys → R– (N–acetyl–Cys) –NH2 → –NHCHO, –NHC(=O)CH3 R–COOH → RC(=O)NHC(R )COOH R–O(or NH)H → R–O(or NH)CH3 Isomerization, rearrangement, etc

R: alkyl or aryl. Ar: aryl. Cys: cysteine. Gly: glycine. Gla: glucuronic acid. Glu: glucose. GSH: glutathione. Gla: glucuronic acid.

HOOC HO HO

Glu: glucose.

O OH OH

HO HO

OH

GSH: glutathione. O

O OH OH

SH

O

HO NH2

N H

O H N

O

OH

54

T. Katagi

as are the endogeneous and exogeneous substrates. The methods for measuring enzyme activity are briefly given, as are the pH and temperature conditions that usually give optimal activity. In addition, taxonomic implications and tissue distribution of enzyme activity are described. The catalytic mechanism of each enzyme is also treated in relation to induction and inhibition of chemical or pesticide activity. Finally, certain basic enzyme profile information such as molecular weight, presence of subunits, and possible isoforms is also commented upon, some in the context of immunochemical assays or gene analyses. 4.2.1 Oxidases Mixed-function oxidases are among the most important and widely distributed phase-I enzymes in aquatic organisms; mixed-function oxidases include an array of cytochrome P450 enzymes (James 1994; Lech and Vodicnik 1985). Oxidases are mainly located in the microsomal fraction, and hepatic tissues are rich in mixedfunction oxidases. The range of reactions catalyzed by this class of enzymes include those presented in Table 9. The catalytic oxidation of substrates by P450 enzymes has been extensively investigated; this reaction proceeds via the reaction mechanism presented in Fig. 4 a(Watanabe 2000). P450-enzyme forms have been assembled into a superfamily of enzymes that have a common character, and the CYP nomenclature is used to identify one isoform from another. One molecular oxygen and two electrons are supplied in reactions that involve CYP enzymes. The electron supply

ROH

RH

P(Fe3+)

(RH)P(Fe3+)

(R·)P(Fe–OH)3+

e−

H2O2 (RH)P(Fe–O)3+

(RH)P(Fe2+)

H2 O 2H+

O2

H2O 2+

3+

(RH)P(Fe

e−

Fig. 4 Reaction schemes for cytochrome P450 and esterase enzymes. (a) P, porphyrin; RH, substrate. The dashed lines refer to shunt pathways. (b) R, R , R1–3 ; alkyl or aryl groups. X = S 0 or O

O R

O22−)

(RH)P(Fe O2)

(RH)P(Fe3+O2−·)

O

Carboxyesterases

O

+ R'OH

R' H2 O

R

OH

X P OR 3 R1O OR2

X

Phosphotriester hydrolases

H2O

R1O

P OH + R3OH OR2

Bioconcentration, Bioaccumulation, and Metabolism

55

comes from nicotinamide adenine dinucleotide phosphate (NADPH) or nicotinamide adenine dinucleotide (NADH), which are indispensable for inserting these oxygen atoms into many kinds of substrates; the reactive species involved in the insertion process is an oxo-ferryl porphyrin π-cation radical. This reactive species can be conveniently prepared by using peroxide via a so-called shunt pathway. P450 enzymes usually operate as terminal oxidases in multicomponent electron-transfer chains that contain one or more fundamental redox domains. Such domains include flavoprotein-containing flavin adenine dinucleotide (FAD), flavin mononucleotide (FMN), ferredoxin, and cytochrome b5 . Solé and Livingstone (2005) measured the levels of the components of P450-dependent monooxygenase systems, including NADPH (or NADH)-dependent cytochrome c reductase and NADH-dependent ferricyanide (b5 ) reductase, and found them in most marine invertebrates; total P450 levels in marine invertebrates are generally in the 50–60 pmol mg–1 protein. The activities of P450 enzymes were estimated to be approximately 30 pmol mg−1 for marine algae (Pflugmacher and Sandermann 1998b) and 8–600 pmol mg−1 protein for annelida (Lee 1998). The P450 activity recorded for aquatic species, other than fish, are lower than comparable activity in mammals (Snyder 2000). Because most benzo[a]pyrene (7) metabolites are quinones when NADPH is present, Livingstone et al. (1989) proposed a P450-mediated peroxidation mechanism via one-electron oxidation for molluscs. Zebra mussel metabolized thiometon, the P=S derivative of demeton-S-methyl (51), via stepwise sulfur oxidation in the absence of NADPH, indicating probable one-electron oxidation (Dauberschmidt et al. 1997). FAD monooxygenase (FMO) also plays a role in metabolizing xenobiotics in aquatic organisms (James 1994; Lech and Vodicnik 1985). Chemicals that contain nucleophilic sulfur, nitrogen, and phosphorus atoms are the typical substrates for FMO. Kurelec (1985) reported FMO activity in the microsomal fraction of the mussel Mytilus edulis toward its typical substrate N,N-dimethylaniline by monitoring the oxidation of NADPH. Optimal FMO activity occurred at a pH of 8.4; this pH is in accord with that of pig FMO. The presence of FMO in the visceral mass and gills of Pacific Oyster was supported by evidence that optimal activity toward 2-aminofluorene was observed at pH 8.4; moreover, significant amounts of molecular oxygen were taken up in the presence of NADPH via metabolism of N- and S-containing substrates (Schlenk and Buhler 1989b). The residual activity for hydroxylation of benzo[a]pyrene (7), in the absence of NADPH, was also detected, where 7-ethoxyresorufin-O-deethylase (EROD) activity was not detected. Activity was enhanced by adding tert-butyl hydroperoxide, although its reduction by reduced glutathione suggested a co-oxidation pathway possibly involving a oneelectron transfer of oxygen from a lipid hydroperoxide. Recently, Boutet et al. (2004) characterized the full cDNA encoding of two non-P450 enzymes in the digestive glands of Pacific oyster. One of these was a monoamine oxidase A consisting of 521 amino acids, which was induced by hydrocarbons and some pesticides. The other was FMO and consisted of 451 amino acids. FMO, which was membranebound and found in the smooth endoplasmic reticulum, catalyzed a four-electron reduction of molecular oxygen. These enzymes had the SGGCY and GAGPAG motif, characteristic of FAD.

56

T. Katagi

P450 activity can be estimated in the presence of NADPH and molecular oxygen by analyzing substrate dissipation or product formation using radiolabels or spectrophotometric methods. Various substrates have been utilized to estimate P450 activity, because each P450 family is known to be metabolically different. Examples of chemicals that have differential metabolic character include: benzo[a]pyrene (7) (CYP1), ethoxyresorfin (CYP1, Fig. 5a), ethoxycoumarin (CYP1 and 2, Fig. 5a), and aldrin (21) (CYP 6 and 12) (Rewitz et al. 2006). The chemical (7) is determined by radioassay of a radiolabeled substrate or fluorometry (Schlenk and Buhler 1988). EROD and ECOD (7-ethoxycoumarin-O-deethylase) activities are estimated by measuring the fluorescence intensity of hydroxy-resorufin and -coumarin, respectively. P450 content is usually estimated spectrophotometrically, using the difference in molar extinction coefficients at 450 and 490 nm (91 M−1 cm−1 ) for the reduced CO-complex (Omura and Sato 1964). However, instead of the typical peak at 450 nm, the time-dependent lower-wavelength peak corresponding to so-called P420 enzyme species was observed at 420 nm for microsomal fractions prepared from the digestive glands of molluscs (Livingstone et al. 1989). The addition of NADPH to the CO-treated microsomal fraction, prepared from hepatopancreas of spiny lobster, showed a λmax of 420 nm, which resulted from a low NADPH cytochrome c reductase : P450 ratio (James 1989). The lower content than expected of P450 was considered to originate from inhibition or denaturation by digestive enzymes or detergent-like molecules during preparation of the microsomal fraction. Lindström-Seppä et al. (1983) also reported the apparently low P450 activity in the hepatopancreas of crayfish, which was caused by the presence of heat-labile proteases. To prevent denaturation of the P450 system, an esterase inhibitor such as phenyl methyl sulfonyl fluoride (PMSF, Fig. 5b) is added to the enzyme assay medium. For example, the intestinal microsomal fractions of polychaetes, prepared in the presence of PMSF, showed a typical difference spectrum of P450 without any peak originating from P420 (McElroy 1990). A typical difference spectrum, with a λmax of 450 nm, was obtained for CO-treated microsomes prepared from marine algae, but a P420 peak was also present. The CO-difference spectrum of P450 could not always be measured because of interference from certain pigments (Pflugmacher et al. 1999; Pflugmacher and Sandermann 1998b; Thies et al. 1996). The two types of difference spectra are well known to actually occur (Kominami 1993) as a result of the reaction of a substrate with P450 ((RH)P(Fe3+ ) in Fig. 4a). The type-I spectrum is characterized by a peak at 385–390 nm, and a trough at 425–435 nm is obtained when a substrate is loosely bound to its active site. In contrast, direct coordination of a substrate to iron as a sixth ligand results in the difference spectrum with a peak at 425–435 nm and a trough at 390–410 nm (type-II). In molluscan microsomal fractions, type-II compounds such as pyridine directly coordinating the central iron of P450 showed typical spectra with a λmax of 424–427 nm and a λmin of 384–389 nm (Livingstone et al. 1989). However, type-I compounds such as phenobarbital (PB, Fig. 5a) and benzo[a]pyrene (7) having no specific atom coordinating to the iron gave the reverse type-I- or type-II-like difference spectra with a λmax of 420–430 nm and a λmin of 380–410 nm. Similar profiles of the difference spectra were obtained for the pond snail (Wilbrink et al. 1991b). The microsomal P450

Bioconcentration, Bioaccumulation, and Metabolism

57

Fig. 5 Typical substrates, inducers, and inhibitors for different enzymes capable of catalyzing the metabolism of pesticides and other chemicals

58

T. Katagi

in freshwater crayfish showed no clear type-I difference spectrum toward ethymorphine but the typical type-II spectrum, having a λmax of 426–431 nm, was obtained for aniline (Lindström-Seppä et al. 1983). In the polychaete Nereis virens, P450 showed a clear type-I spectrum with a peak at 390 nm and a trough at 420 nm with a usual type-II spectrum (Lee 1998). Oxidases such as P450 are widely distributed in freshwater and marine invertebrates including Arthropoda, Echinodermata, Annelida, and Mollusca (ArtolaGaricano et al. 2003; James and Boyle 1998; Lee 1981, 1998; Snyder 2000; Solé and Livingstone 2005). Livingstone et al. (1989) reported the seasonal variation of oxidase activity in the digestive glands of the mussel Mytilus edulis. The existence of P450 was also reported for marine green, red, and brown algae (Pflugmacher et al. 1999; Pflugmacher and Sandermann 1998b) and Euglena gracilis (Briand et al. 1993). EROD and ECOD activities are known to exist in emergent and subemergent aquatic macrophytes (Pflugmacher and Steinberg 1997) and in green algae (Thies et al. 1996; Thies and Grimme 1995). Among these aquatic species, P450s are generally distributed in all tissues, but with high levels in hepatic-like organs such as hepatopancreas and in steroidogenic tissues such as the digestive glands of molluscs, gonads of echinoderms and hepatopancreas, or green glands of crustaceans (Arun et al. 2006; James 1989; Johnston and Corbett 1986a; Livingstone et al. 1989; Rewitz et al. 2006; Schlenk and Buhler 1988; Snyder 2000). Lee (1981) observed a high specific activity in the green glands of the blue crab and the activity was dependent on maturity and molting cycle. P450 activity in organs can be induced by many kinds of substrates, as shown in Fig. 5a (Snyder 2000). P450 activity was induced in molluscs by PAHs and 3-methylcholanthrene (3MC), but molluscs did not respond to phenobarbital (PB) or β-naphthoflavone (BNF) (Livingstone et al. 1989). Wilbrink et al. (1991b) did not observe induction of P450 activity by either PB or 3MC in the digestive glands of pond snails. Both P450 and NADPH ferricyanide reductase activity was induced in microsomal fractions from the digestive glands of freshwater mussels by exposure to 2,4-dichlorophenol (Petushok et al. 2002). Benzo[a]pyrene (7), Aroclor 1254 (PCB congeners), and crude oil, including PAHs, acted to significantly induce P450 activity in the polychaete Nereis virens (Lee 1981). N. virens was found to metabolize benz[a]anthracene by the aid of benzo[a]pyrene hydroxylase, the activity of which was only slightly induced by exposure to 3MC; this indicated metabolic involvement of constitutive P450s (McElroy 1990). P450 was not induced by BNF and 3MC in spiny lobster, crayfish, or crab (James 1989). Arun et al. (2006) measured the activity of each component in the P450-dependent monooxygenase system in freshwater prawns and found increased activity not only for P450 but also for NADPH (NADH), cytochrome c reductase, and cytochrome b5 , which activity increase resulted from exposure to oil effluent at a ppt level. Treatment of Hyalella azteca with BNF increased the metabolism of 2,4,6-trinitrotoluene (15) through aryl oxidation by P450 and also resulted in reduction of nitro groups by NADPH-P450-reductases (Sims and Steevens 2008). BNF significantly increased the activity of benzo[a]pyrene hydroxylase in gumboot chiton (Schlenk and Buhler 1988). Co-exposure of Chironomus tentans to atrazine (127) increased the acute toxicity of organophosphorus pesticides

Bioconcentration, Bioaccumulation, and Metabolism

59

having the P=S moiety, but such activity was reduced for omethoate (56) (Anderson and Zhu 2004). The significant induction of P450 by (127) was confirmed by the ECOD assay, which inducted resulted in more bioactivation of P=S to P=O as well as increased metabolism of (56). Belden and Lydy (2000) also proposed involvement of P450 from observed reductions in the oxon of chlorpyrifos (47), when NADPH was absent or piperonyl butoxide (PBO) was present. Both (47) and (127) were also confirmed to act as P450 inducers, although no induction was observed for DDT (33) in C. tentans (Rakontondravelo et al. 2006). Moita et al. (2000) further observed increased epoxidase activity toward aldrin (21) by exposure to (127), with a concomitant increased intensity of the 45 kDa band in the SDSPAGE (sodium dodecyl sulfate polyacrylamide gel electrophoresis) analysis of the microsomal fraction. In addition to the ECOD activity, C. riparius was found to have either the naphthalene-induced MROD (7-methoxyresorufin-O-demethylase) and EROD activities, in contrast to Daphnia magna (Sturm and Hansen 1999), or atrazine-induced MROD activity (Londoño et al. 2004). The EROD activity in C. riparius was enhanced by exposure to PB and permethrin (81) and increased when this species was fed double the normal ration, possibly from increased amounts of energy for detoxification becoming available (Fisher et al. 2003). Enzyme activity may also be inhibited by specific substrates (Fig. 5a). The PROD (7-pentoxyresorufin-O-depentylase) activity in the digestive glands of pond snails was inhibited by SKF-525A (Wilbrink et al. 1991b). The exposure of D. magna to either PBO, at concentrations well below its LC50 value, or SKF-525A caused a significant dose-dependent reduction in formation of metabolites from pyrene (5); this indicated involvement of P450 (Akkanen and Kukkonen 2003; Ikenaka et al. 2006). PBO completely inhibited aldrin epoxidase activity in the microsomal fraction of C. riparius (Estenik and Collins 1979). Epoxidation of aldrin (21) was inhibited by exposure of caddisfly larvae to carbon monoxide or SKF 525A, showing the involvement of P450-catalyzed oxidation (Krieger and Lee 1976). The mixed-function oxidase activity in microsomes of polychaetes was also inhibited by SKF-525A (Lee 1998). The toxicity of Hyalella azteca and C. tentans to organophosphorus pesticides was greatly reduced in the presence of PBO, because PBO inhibited P450-catalyzed bioactivation to form the corresponding oxon derivatives. In contrast, there was an insignificant effect in the worm Lumbriculus variegatus because of its lower sensitivity to these pesticides (Ankley and Collyard 1995). A similar mechanism was found to operate either in the reduced toxicity of methyl parathion (39) or in the increased activity of permethrin (81) in H. azteca by the P450 inhibitor cimetidine or PBO, respectively (Amweg et al. 2006; Ortíz et al. 2004). PBO was found to significantly inhibit the activities of P450-catalyzed O- and N-dealkylation in green algae (Thies and Grimme 1995; Thies et al. 1996). Less information is available in the literature on pH and temperature optimas for enzyme activity in aquatic species compared with similar information on metabolism. Through metabolism studies with aldrin (21) in bivalva, crustacea, and insecta, optimal pH (7–8) and temperature (–30◦ C) for epoxidase activity were estimated (Estenik and Collins 1979; Khan et al. 1972; Krieger and Lee 1976; Moita et al. 2000). The best studied member of the P450 superfamily in marine

60

T. Katagi

invertebrates is CYP1A1, which is induced by dioxins, PAHs, and PCBs, though many CYP isoforms have been reported (Snyder 2000). Among different phyla, many immunological studies have shown the presence of multiple etiotopes such as CYP1A, 2,3A and 4A, and the cross-reactivity against vertebrate xenobiotic metabolizing P450 s (CYP1A, 2B, 2E, 3A, and 4A) (Rewitz et al. 2006). Enzyme activity for many oxidases has been characterized for many substrates in molluscs. Microsomal fractions from the digestive glands of Mytilus edulis contained two isoforms of oxidases as evidenced by two peaks in DEAE-sepharose ion-exchange column chromatography (Livingstone et al. 1989). SDS-PAGE electrophoresis of these oxidases showed a molecular weight of 53 kDa and the low-spin state of iron was characterized by the absorption spectrum, when the enzymes were oxidized. Peters et al. (1998) have partially purified P450 from the digestive glands of M. edulis by using polyclonal antibodies toward several known P450 isoforms. Through Western blot analysis, the authors showed that the mussel P450 consisted of at least five different CYP forms having molecular weights of 42–53 kDa, each of which reacted with anti-CYP1A, 2B, 2E, 3A, and 4A antibodies. Chaty et al. (2004) examined the amplification of CYP1A and CYP4 sequences on cDNA from the digestive glands of the mussel Unio tumidus and found that exposure to phthalate (DEHP (14)) amplifies the CYP4 transcripts; however, no change was observed from exposure to the PCB congeners Aroclor 1254, which is a CYP1A inducer. In the pond snail Lymnaea stagnalis, either dealkylation of 7-pentoxyresorufin and aminopyrine or hydroxylation of biphenyl (1) proceeded, but not in the presence of EROD activity (Wilbrink et al. 1991b). The difference spectra and these enzyme activities showed the involvement of isozymes P450 b/e. CYP2 and CYP3 families are the most abundant forms in crustacean hepatopancreas microsomes, as demonstrated by antibody studies (James and Boyle 1998). Monooxygenation by CYP2 and CYP3 is faster than that of CYP 1. Moreover, the specific oxygenation sites on the benzo[a]pyrene (7) moiety is highly dependent on what CYP family mediates the process. Baldwin and Lebranc (1994b) have identified at least ten polar metabolites, including six hydroxylated ones in the metabolism of testosterone by Daphnia magna. The NADPH-dependent formation of metabolites and inhibition of in vitro metabolism in the microsomal fractions by carbon monoxide, or metabolite-specific inhibition by PBO, strongly suggested involvement of several isozymes. Diethylstilbestrol decreased the hydroxylase activity when the metabolism of testosterone in D. magna was studied; however, only two metabolites were formed, which indicated involvement of multiple enzymes (Baldwin et al. 1995). In vitro metabolism of fenitrothion (41) in a cell suspension of hepatopancreas from the crayfish Procambarus clarkii demonstrated involvement of several P450 s; this conclusion was based not only on reduced degradation of (41), which produced slightly more oxon in the presence of the CYP1A inhibitor, α-naphthoflavone, but also on increased EROD activity from exposure to (41) (Birmelin et al. 1998; Escartin and Porte 1996). The molecular weight of CYP 2 in spiny lobster was estimated to be 52.5 kDa by SDS-PAGE assay (James and Boyle 1998). Western Blot analysis of the microsomal fractions from the digestive glands of gumboot chiton toward antibodies of trout P450 LM2 and LM4b showed

Bioconcentration, Bioaccumulation, and Metabolism

61

two bands at 54 kDa and a single one at 60 kDa, respectively (Schlenk and Buhler 1989a). The intensity of the former band increased by exposure to BNF, but less intensity was observed for the latter, indicating the presence of at least two P450 isozymes. The in vitro metabolism of aldrin (21) was studied; SDS-PAGE analysis of microsomal proteins prepared from Chironomus tentans showed a single band at 45 kDa, which cross-reacted with a Drosophila anti-P450 polyclonal antibody in the immunoblot assay (Moita et al. 2000). The genomic DNA from the whole body of C. tentans was amplified by polymerase chain reaction, followed by purification, and was then subjected to sequence analysis (Londoño et al. 2004). The 444-bp product showed a nucleotide sequence similar to that of CYP4 genes. When algal N-dealkylation of metflurazon (138) was studied, the microsomal fractions of two Chlorella sp. were subjected to Western Blot analysis using polyclonal antisera (Thies et al. 1996). Differences in cross-reactions of the algal proteins (46–67 kDa) from the two species toward antisera indicated the presence of different isozymes, which may also explain why the Michaelis–Menten constants were different. Metabolism investigations with three long-alkyl chain fatty acids and 3-chlorobiphenyl in nine marine algae gave the corresponding hydroxylated metabolites; these results suggested involvement of several isozymes (Pflugmacher et al. 1999). Briand et al. (1993), employing immunochemical methods, identified three isozymes in the microsomes of Euglena gracilis, using the antibodies for rat liver CYP2C11, 2E, and 2B. The three isozymes had molecular weights of 50, 50.5, and 52 kDa and exhibited the different modulation by PB. Although researchers have extensively studied oxidase-catalyzed metabolism of various compounds and associated enzymology, investigations of reaction mechanisms in aquatic species have emphasized algal metabolism of PAHs. Given the nature of metabolites identified, both monooxygenases and dioxygenases are likely to play a role in algal metabolism of various aromatic compounds, e.g., PAHs and dioxins, (Chan et al. 2006; Schoeny et al. 1988; Todd et al. 2002). Cerniglia et al. (1979, 1980c) examined the metabolism of naphthalene (2) in three blue-green algae and found both 4-hydroxy-1-tetralone and cis-1,2-dihydroxy-1,2dihydronaphthalene, in addition to 1-naphthol, whose oxygen originated from O2 . A monooxygenase was considered to be involved in the stepwise oxygenation from 1-naphthol to 4-hydroxy-1-tetralone via 1,2-arene oxide, whereas the formation of the cis-diol was probably catalyzed by dioxygenases. Furthermore, Narro et al. (1992b) used MS to examine metabolites from (2) that had been deuterated at the 1- or 2-position. About 70% retention of deuterium, with insignificant primary isotope effect, suggested that the 1,2-arene oxide formation followed an NIH-shift. The same mechanism was observed when 1-naphthalenesulfonic acid was metabolized in Scenedesmus obliquus, primarily to 1-hydroxy-2-naphthalenesulfonic acid, with only small amounts of 1-naphthol being formed (Kneifel et al. 1997). The same metabolites were identified in 18 different algal cultures including green, red, and brown algae and diatoms, with no formation of trans-diol identified by using co-chromatography and authentic standards (Cerniglia et al. 1980a). The cis configuration (with oxygen incorporation from O2 ) was also confirmed from metabolism studies in green algae, by the identification of 4,5-, 7,8-, and 11,12-diol

62

T. Katagi

of benzo[a]pyrene (7) using HPLC, MS, and fluorescence spectroscopy (Lindquist and Warshawsky 1985a, b; Warshawsky et al. 1988). In contrast, the marine bluegreen alga Armenellum quadruplicatum PR-6 appeared to metabolize phenanthrene (4) via 1,2-arene oxide, by a monooxygenase and epoxide hydrolase, to form the trans-9,10-dihydroxy-9,10-dihydrophenanthrene (Narro et al. 1992a). Incorporation of an oxygen atom from O2 was demonstrated by MS, and its configuration was confirmed by H-NMR and separation of the diastereomers using HPLC. Semple and Cain (1996) reported the mineralization of phenol by the golden brown alga Ochromonas danica; these authors disclosed that the phenol oxidation to catechol by monooxygenase was followed by catechol 2,3-dioxygenase-catalyzed meta-cleavage of the phenyl ring. 4.2.2 Reductases Although reductive metabolism has been reported in many aquatic organisms, most relevant information on its mechanism and the enzymes involved is limited to algae and aquatic macrophytes. In some cases, an enzyme cofactor may participate in reductive transformations. In the degradation of mexacarbate (99) by blue-green algae, a flavoprotein fraction prepared by gel-filtration chromatography of an axenic cell-free suspension showed higher metabolic activity in the presence of FMN (Matsumura and Esaac 1979). Garrison et al. (2000) prepared a fraction of powdered Elodea canadensis, with a molecular weight cut-off at about 1200, by size-exclusion chromatography for degradation of DDT (33). The authors supposed some enzyme cofactor responsible for the formation of DDD (35) in the presence of ascorbic acid. NADH or NADPH was indispensable as co-factors for reductive degradation of azo dyes to the corresponding anilines by Chlorella and Oscillatoria sp. as evidenced by IR analysis (Jingi and Houtian 1992). The relevant enzymes were induced by acclimatizing algae to azo dyes, and their activities were significantly increased in the presence of these dinucleotides. The dechlorinating reductive activity in the marine diatom Thalassiosira sp. for various chlorophenols was found to also depend on NADH (Lovell et al. 2002). The regio- and stereo-selective reduction of keto groups in sterols has been reported for various algae including Chlorella sp., although its mechanism is unclear (Pollio et al. 1994; Greca et al. 1996a). Sims and Steevens (2008) reported the reduction of 2,4,6-trinitrotoluene (15) in the amphipod Hyalella azteca, and the reduction was considered to be catalyzed by NADPH-P450 reductase. The possible involvement of dehalogenases in the degradation of chlorinated hydrocarbons, such as carbon tetrachloride, hexachloroethane, and trichloroethene, was reported for algae and aquatic macrophytes (Nzengung et al. 2003; Wolfe and Hoehamer 2003). Recently, Kuritz et al. (1997) examined the metabolism of lindane (30) by using the wild and transpositional mutants of Anabaena sp. PCC7120. This alga possesses genes organized in an operon as 5-nir A-nrt ABCD-nar B-3 , and they encode for nitrite reductase, nitrate transport proteins, and nitrate reductase. There was no direct correlation of degradation to individual enzyme activity from the encoding nir operon, although some function of a membrane nitrate–nitrite reduction complex was considered to play a role in the degradation of (30). A similar

Bioconcentration, Bioaccumulation, and Metabolism

63

approach was undertaken by using Anabaena sp. to examine the reduction mechanism of methyl parathion (39) (Barton et al. 2004). A gene mutation relating to nitrate reduction did not affect the degradation of (39), and the transient formation of the nitroso derivative in the presence of light probability proceeded by the action of ferredoxin-NADP+ reductase. Kuritz and Wolk (1995) successfully increased the metabolic activity of Anabaena sp. for lindane (30) and 4-halogenated benzoic acids by using the genetically engineered algae that bore plasmids capable of enhanced degradation. 4.2.3 Esterases Esterases are classified into three categories based on their reaction profiles (Basack et al. 1998; Thompson 1999; Wheelock et al. 2005). A-esterases that include phosphotriester hydrolases can hydrolyze organophosphorus (OP) compounds as shown in Fig. 4b, and are not inhibited by OPs. B-esterases, including cholinesterases (ChE) and carboxyesterases (CaE), are typically inhibited by OPs as a result of the extremely slow dephosphorylation of tetrahedral intermediates formed between OPs and a serine residue at their active sites (Fukuto 1990). CaEs are well known to hydrolyze pyrethroids and carbamates, as shown in Tables 10 and 11. Among ChEs, acetylcholine (AchE) and butyrylcholine (BchE) esterases found in neuromuscular junctions are common in aquatic organisms that have genetic and molecular polymorphism (Galloway et al. 2002). In contrast, CaEs are usually distributed in all tissues and play a role in hydrolyzing a wide variety of endogeneous and exogeneous esters. ChE activity was distributed both in the supernatant and in pellet fractions of Corbicula fluminea but was affected differently by OPs and inhibitors such as PMSF and esserin (Fig. 5b) (Basack et al. 1998). Finally, C-esterases (acetylesterases) are not inhibited by OPs. In general, the enzyme activity of esterases is spectrophotometrically determined by reacting the product formed via enzymatic hydrolysis of substrates such as thiocholines or thioacetate (Fig. 5b) with the chromogenic reagent DTNB (5,5 -dithio-bis(2-nitrobenzoic acid)) (Basack et al. 1998). For CaE, a simple UV determination can be used instead of using 1-naphthyl acetate as a substrate (Belden and Lydy 2000). Among inhibitors other than pesticides (Fig. 5b), PMSF and eserine generally inhibit most esterases and ChEs, but BW, iso-OMPA, and CBDP specifically inhibit AchE, BchE, and CaE, respectively (Damásio et al. 2007; Escartin and Porte 1997; Vioque-Fernández et al. 2007). In algal metabolism of pesticides, the involvement of phosphatases has been previously reported. Subramanian et al. (1994) found that phosphorous starvation increased activity of extracellular alkaline phosphatases in blue-green algae, whereas exposure to organophosphorus pesticides induced acid phosphatase activity. In studying the metabolism of glyphosate (72) in blue-green algae, the extent of induction was dependent on species (Ravi and Balakumar 1998). The speciesdependent metabolism of soman (74) was observed for duckweed (Hoskin et al. 1999). Giant duckweed could hydrolyze (74) with release of a fluoride ion with the aid of acid anhydrolase, whereas no similar metabolism was observed by Lemna

Pesticide/structure

Parathion

Methyl parathion

Fenitrothion

No.

39

40

41

O2 (OCH3 ), O7 (oxon), H2, R3 (NH2 ), C2 (O-β-Glu), C3 O2 (OCH3 ), H2, O7 (oxon) O7, H2, M (thiono-thiolo rearrangement) O2 (OCH3 ), H2, O7 (oxon) O2 (OCH3 ), H2, O7 (oxon) O7, H2, M (thiono-thiolo rearrangement) O2 (OCH3 ), O7 (oxon), H2, C3 O7, H2, M (thiono-thiolo rearrangement) O2 (OCH3 ), O7 (oxon), H2, C2 (O-β-Glu), C3

Physa acuta

G: Chlorella vulgaris Oedogonium cardiacum B: Anabaena flos-aquae D: Nitzschia closterium Mc: Myriophyllum sp. W: Daphnia pulex

I: Chironomus riparius S: Penaeus japonicus

Sn: Cipangopaludina japonica

O7, R3 (NH2 ), H2 R3 (NH2 ), H2, 1 unknown O7, R3 (NH2 ), H2 R3 (NH2 ), H2 R3 (NO, NH2 ) R3 (NH2 ), H2 O7 (oxon) R3 (NH2 ), H2 O2 (OCH3 ), H2, O7 (oxon), C1 (O-β-Gla) C2 (O-β-Glu), C3 O2 (OCH3 ), O7 (oxon), H2, R3 (NH2 ), C3

Typeb

Sn: Physa sp. G: Chlorella pyrenoidosa Oedogonium cardiacum B,G: Nostoc and Tolypothrix sp. B: Anabaena sp. strain PCC7120 W: Daphnia magna I: Chironomus riparius B,G: Six species Cr: Procambarus clarkii

Speciesa

Table 10 Metabolism of organophosphate insecticides in aquatic organisms

Kikuchi et al. (1984) Fisher (1985)E Kikuchi et al. (1984) Kikuchi et al. (1984) Weinberger et al. (1982)E Miyamoto et al. (1979)E , Takimoto et al. (1987b) Fisher (1985)E Kobayashi et al. (1985a, b, 1990)

Miyamoto et al. (1979)E , Takimoto et al. (1987a) Takimoto et al. (1987a)

Francis et al. (1980)E Mackiewicz et al. (1969) Francis et al. (1980)E Mostafa et al. (1991) Barton et al. (2004) Francis et al. (1980)E Lydy et al. (1990) Megharaj et al. (1994) Foster and Crosby (1986)

Referencesc

64 T. Katagi

Pesticide/structure

Fenitrothion

DCDEP Temephos

Chlorpyrifos

Quinalphos Demeton-S-methyl Phorate Disulfoton

No.

41

45 46

47

49 51 52 53

H2 H2 H2 O7 (oxon), H2 C4 (SCH3 substitution at C6) 2 unknowns O6 (sulfoxide) NM O6 (sulfoxide) O6 (sulfone)

O6 (sulfoxide, sulfone), O7, H2

Sn: Physa sp.

G: Oedogonium cardiacum Sn: Physa sp. C: Katalysia opima Mu: Mytilus edulis Oy: Crassostrea virginica B,G: Five species I: Chironomus tentans B: Anabaena and Aulosira sp. I: Chironomus tentans Mu: Dreissena polymorpha

C2 (O-β-Glu), C3 H2, O7 (oxon) O6 (sulfoxide, sulfone), O7, H2

O2 (OCH3 ), O7 (oxon), H2, C2 (O-β-Glu) O2 (OCH3 ), H2, O7 (oxon) H2, O7 (oxon) O2 (OCH3 ), H2, O7 (oxon) O2 (OCH3 ), H2, O7 (oxon) O2 (OCH3 ), H2, O7 (oxon), R3 (NH2 ),

Typeb

Oy: Crassostrea gigas G: Oedogonium cardiacum

Palaemon paucidens Artemia salina Cr: Procambarus clarkii Co: Sinocalenus and Oithona sp. R: Brachionus plicatilis Cb: Callinectes sapidus

Speciesa

Table 10 (continued)

Hale (1989) Metcalf and Sanborn (1975)E Metcalf and Sanborn (1975)E Metcalf (1976)E Metcalf (1976)E Kale et al. (1999bE , 2002) Serrano et al. (1997a, c) Woodburn et al. (2003) Megharaj et al. (1987) Anderson and Zhu (2004) Dhanaraj et al. (1989) Anderson and Zhu (2004) Dauberschmidt et al. (1997)

Takimoto et al. (1987b) Kashiwada et al. (1998) Birmelin et al. (1998) Kashiwada et al. (1998) Kashiwada et al. (1998) Johnston and Corbett (1986a, b)

Referencesc

Bioconcentration, Bioaccumulation, and Metabolism 65

Terbufos

Dimethoate Malathion

Monocrotophos Fenamiphos EPN

Leptophos

Glyphosate

54

55 57

64 67 70

71

72

O7 (oxon) H1 (β) H1 (α, β), H2 (P-S, C-S) H1 (α, β), H2 (P-S, C-S), O7 (oxon) 4 unknowns O6 (sulfoxide), H2 O7, R3 (NH2 ), H2 O7, R3 (NH2 ), H2 O7, R3 (NH2 ), H2 R1 (Br), O7 R1 (Br), O7, M (thiono-thiolo rearrangement) H2 (C-P)

B: Anabaena oryzae Phormidium fragile B,G: Five species G: Pseudokirchneriella subcapitata Sn: Physa sp. G: Oedogonium cardiacum W: Daphnia magna Sn: Physa sp. G: Oedogonium cardiacum B: Anabaena variabilis

O7

Sn: Physa sp.

I: Chironomus tentans G: Chlorella sp.

O6 (sulfoxide, sulfone), O7

Typeb

G: Oedogonium cardiacum

Speciesa

Metcalf and Sanborn (1975)E Metcalf and Sanborn (1975)E Anderson and Zhu (2004) O’Kelley and Deason (1976) Khalil and Mostafa (1987) Khalil and Mostafa (1987) Megharaj et al. (1987) Cáceres et al. (2008) Francis et al. (1980)E Francis et al. (1980)E Francis et al. (1980)E Francis et al. (1980)E Francis et al. (1980)E Ravi and Balakumar (1998)

Referencesc

b Type

by EPI-Suite (USEPA 2008) or experimental data therein. of metabolism is defined and listed in Table 9. The term in parentheses refers to the metabolic position and associated details. c Superscript “E” means a model ecosystem. Cb, crab ns: metabolic position not specified. NM, not metabolized. DCDEP, O,O-diethyl O-(2,4-dichlorophenyl) phosphorothioate.

a Estimated

Pesticide/structure

No.

Table 10 (continued)

66 T. Katagi

Pesticide/structure

2,4,5-T Triclopyr

Diclofop-methyl Cypermethrin Deltamethrin

Lambda-cyhalothrin Fenvalerate (2S-isomer) Methoprene

Propanil

Carboxin

76 77

Esters 78 82 83

84 85 87

Amides 89

94

Phenoxyacetic acids 75 2,4-D

No.

H3 H3, C5 (N-acetyl) O6 (sulfoxide) O3 (ns)

Mc: Lemna minor G: Chlorella vulgaris

B: Anabaena sp.

H1 H1 H1 H6 (CONH2 , COOH) O2 O2, H1

Mc: Lemna sp. Potamogeton berchtoldi Mc: Ceratophyllum demersum Sn: Cipangopaludina japonica G: Oedogonium cardiacum Sn: Physa sp.

Three G and seven B species

H1, O1, M (ether cleavage) ns, trans>cis H1, M (isomerization)

O3 (3, 5) C6 (taurine) C6 (taurine) O1, C7 (O) O1, C7 (O) C6 (taurine) O1, C7 (O)

Typeb

G: Chlorella and Scenedesmus sp. Mu: Mytilus edulis G: Dunaliella bioculata

G: Scenedesmus quadricauda L: Panulirus argus L: Panulirus argus Mc: Myriophyllum spicatum C: Lampsilis siliquoidea Cr: Procambarus clarkii Orocnectes sp.

Speciesa

Table 11 Metabolism of miscellaneous pesticides in aquatic organisms

Wright and Manule (1982) Wright et al. (1977) Mitsou et al. (2006) Balasubramanya and Patil (1980) Balasubramanya and Patil (1980)

Cai et al. (2007, 2008) Gowland et al. (2002) Baeza-Squibqn et al. (1988, 1990) Muir et al. (1985a)M Muir et al. (1985a)M Hand et al. (2001)E Ohkawa et al. (1980)E Metcalf and Sanborn (1975)E Metcalf and Sanborn (1975)E

Valentine and Bingham (1974) James (1982) James (1982) Getsinger et al. (2000)M Getsinger et al. (2000)M Barron et al. (1991) Getsinger et al. (2000)M

Referencesc

Bioconcentration, Bioaccumulation, and Metabolism 67

G: Chlorella pyrenoidosa G: Chlorella fusca

Monolinuron Buturon

Isoproturon

115 117

118

G: Chlorella and G: Anabaena sp. Mc: Lemna minor Mc: Ceratophyllum demersum

O2 O2 (CH3 & CH3 O), H5, C5 (N-acetyl) O2 (CH3 ), C2 (O-β-Glu) O2, O8 (C=C, ns), R4 (C≡C), H5, C5 (N-acetyl) O1 (i-Pr), O2, H5 ns O1 (i-Pr), O2, H5, C4 (N & i-Pr-O)

Carbofuran

102

15 G and two B species G: Chlorella vulgaris

Carbaryl

101

H3, C7 (O), conjugation H3, C7 (O), O3 (6), O5 (1,4), M (C–O cleavage)

105 Methomyl 109 Thiobencarb 110 Chlorpropham Ureas and benzoylureas 112 Flumeturon 113 Metobromuron

Aminocarb Mexacarbate Propoxur

Carbamates 98 99 100

Sn: Cipangopaludina chinesis Mc: Spirodela polyrhiza

Typeb

O2 (4-N, phenyl ring) O2 (4-N, phenyl ring), H4 O2 (NHCH2 OH) O2, H4 H4 H4 H4 H4, O1 (3, furan ring) O1 (NCH3 ; 3, furan ring) H4 H4 O3 (2), H4, O6 (sulfoxide, sulfone) H4

Naproanilide

96

Speciesa

Cr: Orconetes limosus Mc: Typha latifolia G: Oedogonium cardiacum Sn: Physa sp. Mc: Ceratophyllum demersum Sn: Indoplanorbis exustus I: Chironomus riparius B: Anabaena and Phormidium sp. Mc: Elodea Canadensis C: Corbicula sp. B: Phormidium fragile G: Oedogonium cardiacum Three G and five B species

Pesticide/structure

No.

Table 11 (continued)

Mostafa and Helling (2001) Böttcher and Schroll (2007) Pietsch et al. (2006)

Schuphan (1974) Tsorbatzoudi et al. (1976)

Zablotowicz et al. (1988) Tweedy et al. (1970)

Sundaram and Szeto (1979) Sundaram (1995)M Metcalf (1976)E Metcalf (1976)E Tsuge et al. (1976)E Tsuge et al. (1976)E Lohner and Fisher (1990) Khalil and Mostafa (1987) Yu et al. (1974b)E Metcalf and Sanborn (1975)E Khalil and Mostafa (1986) Chen et al. (1982)E Wright and Manule (1982)

Wang et al. (1992a)E Wang et al. (1992a)E

Referencesc

68 T. Katagi

Thidiazuron

Photo-thidiazuon

Diflubenzuron

119

–

120

Chlornitrofen Chlomethoxyfen Bifenox Atrazine

Cyanazine Pyrazolate Chlordimeform

Bentazone

Metflurazon

124 125 126 127

129 132 135

137

138

121 CCU Miscellaneous 16 Tri-n-butyltin

Pesticide/structure

No.

G: Chlorella fusca and sorokiniana

Mu: Mytilus edulis Sn: Physa sp. Sn: Physa sp. G: Four species Oedogonium cardiacum Sn: Physa sp. Cb: Uca minax C: Tapes philippinarum G: Chlorella and B: Oscillatoria sp. Cb: Uca minax

G: Ankistrodesmus falcatus D: Skeletonema costatum

G: Chlorella and B: Oscillatoria sp. G: Chlorella and B: Oscillatoria sp. G: Scenedesmus subspicatus Oedogonium cardiacum B: Plectonema sp. G: Scenedesmus subspicatus

Speciesa

M (cleavage of S–N or amide linkage) O2

M (cleavage of Sn–C bonds) O1 (β, γ , δ), M (cleavage of Sn–C bonds) R3 (NH2 ), C5 (formyl, acetyl) M (C–O cleavage) R3 (NH2 ), H1 O2 (ethyl) O2 (ethyl, i-Pr) O2 (ethyl, i-Pr) O2 (ethyl) O1, O2 (N–CH3 ), H2 (sulfonyl) O1, O2, H6 (formamidine)

ns (2 metabolites) H5, C7 (N) H5 ns (6 metabolites)

O3 (4), H5

H5

Typeb

Table 11 (continued)

Tantawy et al. (1984); Thies et al. (1996)

Booth et al. (1973)E

Watanabe et al. (1987) Lee et al. (1976)E Lee et al. (1976)E Zablotowicz et al. (1988) Metcalf and Sanborn (1975)E Metcalf and Sanborn (1975)E Yu (1975d)E Ueda et al. (1988) Benezet and Knowles (1981)

Maguire et al. (1984) Lee et al. (1989)

Yu-yun et al. (1993) Metcalf et al. (1975)E Schooby and Quinstad (1979) Yu-yun et al. (1993)

Benezet and Knowles (1981)

Benezet and Knowles (1981)

Referencesc

Bioconcentration, Bioaccumulation, and Metabolism 69

Oxadiazon

R-20458

Trifluralin

Acrolein

139

140

142

144

O: Lumbriculus variegates I: Chironomus riparius C: Elliptio complanata Cr: Orconectes virilis

W: Daphnia magna

C: Sphaerium corneum Sn: Helisoma sp.

G: Oedogonium cardiacum

Pond snail and blue mussel G: Oedogonium cardiacum Sn: Helisoma sp. G: Chlorella sp. Clamydomonas sp.

Speciesa O1 (t-Bu) O2 (i-Pr) O2 (i-Pr) O1, H6 (epoxide) O1, O8 & H6 (epoxide), M (C-O cleavage) O2, R3 (NH2 ; 2,4), M (benzimidazole) NM O2, R3 (NH2 ; 2,4), M (benzimidazole) O2, R3 (NH2 ; 2,4), M (benzimidazole) NM ns (2 metabolites) O1, O8 (epoxide), R4 O1, O8 (epoxide), R4

Typeb

Guerrero et al. (2002) Guerrero et al. (2002) Nordone et al. (1998) Nordone et al. (1998)

Isensee et al. (1979b)E

Guerrero et al. (2002) Isensee et al. (1979b)E

Isensee et al. (1979b)E

Murakami et al. (1993) Ambrosi et al. (1978)E Ambrosi et al. (1978)E Schooley and Quinstad (1979) Schooley and Quinstad (1979)

Referencesc

b Type

by EPI-Suite (USEPA 2008) or experimental data therein. of metabolism is defined and listed in Table 9. The term in parentheses refers to the metabolic position and associated details. c Superscript “E” means a model ecosystem. Ch, gumboot chiton; L, lobster; CCU, 1-(4-chlorophenyl)-3-(2-chlorobenzoyl)urea

a Estimated

Pesticide/structure

No.

Table 11 (continued)

70 T. Katagi

Bioconcentration, Bioaccumulation, and Metabolism

71

minor; this was taken as evidence for the absence of this enzyme. In metabolism studies with organophosphorus pesticides, various organophosphorus hydrolases were detected in several aquatic macrophytes by Wolfe and Hoehamer (2003). The presence of these enzymes indicated the potential to cleave not only susceptible P–O and P–S bonds but also the stronger P–C, P–N, and P–F ones. The location of alkaline phosphatases in tissues was studied by using a histochemical technique; alkaline phosphatases were demonstrated to reside in the cell walls of Chlorella vulgaris and various marine macrophytes (Brandes and Elston 1956; Hernández et al. 1994). In the mussel Mytilus edulis, CaE, in contrast to AChE, has been shown to be mainly distributed in tissue homogenates rather than in hemolymph. CaE was significantly inhibited by paraoxon (oxon derivative of (39)), but only slightly by eserine (Galloway et al. 2002). A similar inhibition of CaE by fenitrooxon (42) was reported in M. galloprovincialis, whose AchE is mainly distributed in gills; BchE and CaE are distributed in digestive glands (Escartin and Porte 1997). A seasonal variation in activities of these enzymes was observed. The lowest activity was observed in the spring, and was thought to result from water contamination from OPs, together with elevated metabolic activation caused by a higher temperature. In crayfish (Procambarus clarkii), CaE was mainly distributed in digestive glands and showed different sensitivity to different pesticides (Vioque-Fernández et al. 2007). Barata et al. (2004) found that CaE had a significant involvement in detoxifying chlorpyriphos (47), malathion (57), and carbofuran (102) in D. magna; this effect was recognized because of increased toxicity in the presence of CaE inhibitors. CaE activity, in D. magna resistant to fenitrothion (41), was slightly higher than activity in sensitive D. magna. However, these differences were not enough to explain the sensitivity (Damásio et al. 2007). Decapoda (e.g., lobster, shrimp, and crab) hydrolyzed methyl farnesoate, with esterase activity mainly located in their hepatopancreas (Homola and Chang 1997). Wheelock et al. (2005) reported that mammals and insects preferentially metabolized the trans-isomer of cypermethrin (82), and this moiety was possibly metabolized by CaE in mussel Mytilus edulis (Gowland et al. 2002). Baeza-Squiban et al. (1988, 1990) have shown that the green alga Dunaliella sp. can hydrolyze deltamethrin (83) to the corresponding acid and alcohol by CaE released into medium. Propanil (89) was hydrolyzed by various green and blue-green algae to 3,4-dichloroaniline, but only Anacystis sp. could metabolize chlorpropham (110), indicating some substrate specificity of CaEs (Wright and Maule 1982). Murthy and Sarena (1980) examined the hepatopancreatic esterases of freshwater prawn by using 2-naphthyl acetate as a substrate. The pH and temperature for optimal activity were found to be 7.4 and 40◦ C, respectively, comparable to values reported for other crustaceans. Esterase heterogeneity in the mussel Mytilus sp. has been confirmed by Ozreti´c and Krajnovi´c-Ozreti´c (1992). The enzymatic hydrolysis of 1-naphthyl acetate reached a maximum at pH 7.0–8.5, with the highest activity located in hepatopancreas. Electrophoresis of tissue homogenates showed five anodic zones and one cathodic zone, with a slightly tissue-dependent distribution; three CaEs were identified from their substrate specificity and inhibitory profiles. Multiple general esterases in Chironomus tentans were also confirmed by electrophoresis (Rakotondravelo et al. 2006). Using either inhibition studies or electrophoresis analysis, the soluble fraction of marine dinoflagellates was shown to

72

T. Katagi

contain many esterases, with a predominance of A-esterases (Barbier et al. 2000). Immunoblotting techniques helped characterize entities with a molecular weight of 80 kDa, and cytochemical analysis showed no specific localization of esterases in the cells. 4.2.4 Glucose and Glucuronic Acid Transferases These enzymes catalyze the transfer of glucose or glucuronic acid from the corresponding uridine diphosphate derivatives (UDPG and UDPGA) to oxygen, nitrogen, or sulfur atoms of a substrate as shown in Fig. 6a. Among aquatic organisms, glucuronidation is the main conjugation pathway in fish, whereas many invertebrates predominantly rely on glucosidation for conjugate production (James 1987, 1994). The glucose conjugate can be further metabolized by acetylation or conjugation with malonic acid or other saccharides. Enzyme activity can be estimated using spectrophotometric or radiochemical quantitation techniques. Glucuronidation has been reported to occur during the metabolism of PAHs, phenols, and pesticides by crustacea (Sanborn and Malins 1980; Reichert et al. 1985), bivalva (Borchert et al. 1997; Lalah et al. 2003; Michel et al. 1995), larval chironomids (Borchert et al. 1997; Kawatski and Bittner 1975; Leversee et al. 1982), and gumboot chiton (De Busk et al. 2000; Schlenk and Buhler 1988). The enzyme activity seems to be located in the microsomal fraction, as evidenced by results of an enzyme assay on the digestive glands of gumboot chiton. In the presence of UDPGA, phenol derivatives are the most popular substrates, and lansol (3,4-dibromo-5hydroxymethylpyrocatechol) has been reported to slightly enhance the enzyme’s activity (De Busk et al. 2000). The microsomal enzyme activity in gumboot chiton was determined to be 28–38 nmol mg–1 protein min−1 comparable to that in rats. More information on glucosidation has been accumulated through studies on the metabolism of phenols, anilines, PAHs, and pesticides in algae (Kneifel et al. 1997; Petroutsos et al. 2007; Schoeny et al. 1988; Schuphan 1974; Soeder et al. 1987; Warshawsky et al. 1988, 1990), aquatic macrophytes (Barber et al. 1995; Ben-Tal and Cleland 1982; Day and Saunders 2004; Ensley et al. 1994; Fujisawa et al. 2006; Nakajima et al. 2004; Pascal-Lorber et al. 2004; Roy and Hänninen 1994; Sharma et al. 1997; Tront and Saunders 2007), crustacea (Foster and Crosby 1986; Johnston and Corbett 1986a, b; Kashiwada et al. 1998; Kobayashi et al. 1985a, b, 1990; Kukkonen and Oikari 1988; Sanborn and Malins 1980; Schell and James 1989; Takimoto et al. 1987b), bivalva (Shofer and Tjeerdema 1993; Takimoto et al. 1987a; Tjeerdema and Crosby 1992), and gumboot chiton (Landrum and Crosby 1981). Pridham (1964) first reported the insignificant activity of UDPG transferases in aquatic macrophytes and algae using quinol and resorcinol, while the usage of various substrates has later revealed the wide distribution of O-glucosyltransferases in the soluble enzyme fraction of these species, but with the limited distribution of N- and S-glucosyltransferases (Pflugmacher and Sandermann 1998a; Pflugmacher et al. 1999). However, the marine diatom Skeletonema sp. has insignificant UDPG transferase activity (Yang et al. 2002). The enzyme activity of some crustaceans is localized in the microsomal fraction of hepatopancreas or green glands (Elmamlouk

Bioconcentration, Bioaccumulation, and Metabolism

73

and Gessner 1978; Hänninen et al. 1984; Li and James 1993; Schell and James 1989; Väisänen et al. 1983). Exposure of D. magna to diethylstilbestrol enhanced the formation of glucose conjugates of testosterone, but its presence was found to also slightly inhibit enzyme activity (Baldwin et al. 1995). UDPG was demonstrated to be an essential substrate for transfer of glucose in terrestrial molluscs, and the enzyme’s activity was optimal at pH 9.3 (Dutton 1965, 1966). Väisänen et al. (1983) reported significant activation of the enzyme’s activity by Mg2+ . Later, Elmamlouk and Gessner (1978) examined the UDPG transferases in the hepatopancreas of the lobster Homarus americanus and found that among several carbohydrates, only glucose is transferred from the corresponding UDP to p-nitrophenol (NP) at an optimal pH of about 8. A linear Lineweaver-Burk plot was obtained for NP while non-linearity was observed for UDPG with two estimated Km (Michaelis-Menten constant) values. Li and James (1993) have also found different pH values for optimal enzyme activity between antennal glands (pH 7.6) and hepatopancreas (pH 10.5) of the lobster when 2-naphthol was used as a substrate. Similar enzyme profiles were reported for the spiny lobster Pannulirus argus, wherein the optimal pH and temperature were 8.0–8.5 and 30–35◦ C, respectively; the reaction did not obey simple Michaelis–Menten kinetics, and Km and Vmax (maximum reaction velocity) values were highly dependent on substrates (Schell and James 1989). Based on these results, the presence of more than one isozyme is probable for UDPG transferases; this conclusion was supported by the different distribution in the glucosyl conjugates of testosterone metabolites in D. magna when treated with 20-hydroxyecdysone (Baldwin and LeBlanc 1994a). Other enzymes capable of transferring a carbohydrate to a xenobiotic appear to exist in Lemna minor, because phenol derivatives were successively conjugated by glucose and apiose in this species (Day and Saunders 2004).

4.2.5 Sulfotransferases The enzymatic reaction scheme for this enzyme type is shown in Fig. 6b. The SO3 moiety is transferred from phosphoadenosyl phosphosulfate (PAPS) to oxygen or nitrogen atoms of a substrate. The enzyme activity is usually estimated through the quantification of a sulfate metabolite by using [35 S]-PAPS or radiolabeled substrate. Sulfate conjugation is known to have high affinity but a low capacity, whereas carbohydrate conjugation is characterized as low affinity and high capacity (James 1994). Because the amount of endogeneous PAPS is less than that of UDPG(A), a larger amount of xenobiotic may limit the extent of sulfate conjugation. Sulfotransferase enzyme activity has been demonstrated to be mainly located in cytosolic fractions; tissues rich in this enzyme include mid-gut gland of clam (Kobayashi et al. 1987), digestive gland and gonad of mussel (Lavado et al. 2006), digestive gland of gumboot chiton (De Busk et al. 2000), antennal glands and hepatopancreas of lobster (Elmamlouk and Gessner 1978; Li and James 1993; Schell and James 1989), digestive tube of sea urchin (Janer et al. 2005a), and in dinoflagellates (Yoshida et al. 2002).

74

T. Katagi (a) Glucosylation & glucuronidation OH HOOC HO HO

O

HO HO

HO

Dehydrogenase

O HO

O UDP

Uridine diphosphate glucose (UDPG) Glucosyl transferase

O UDP

Uridine diphosphate glucuronic acid (UDPGA) Glucuronyl transferase

R' Y H (Y=O, NH,S) OH

UDPG +

O

HO HO

HOOC HO HO

Y R'

O

+ UDPGA

Y R'

O

Malonyl-CoA

NH

UDP =

O

OO

HO

Acyltransferase

O

OH

OH

P O

HO HO

O O

P

O

Y R'

N

-

O

O

O

O

OH

OH

OH

(b) Sulfation NH2

+ PAPS

R' Y H (Y = O, NH)

R’OSO3H

Sulfotransferase

N

PAPS (Phosphoadenosyl phosphosulfate) =

-

O

S O

O

P

O

N

O

O OH

O

(c) Glutathione conjugation

N

N

O-

O

O

-

X

P

OH

O-

Glutathione R-X + GSH

S-transferase

R-SG GSH (glutathione) =

γ-Glutamyl transpeptidase Aminopeptidase

R-Cys-Gly

O HO

R-Cys

SH

O

NH2

O

H N

N H

OH

O

COOH

N-acetyl transferase

R

S

NHCOCH3

(d) Acylation RCOOH

CoA-SH / ATP Ligase

RC(=O)S-CoA

(R=CH3, Acetyl-CoA) CoA-SH =

HS

R’NH2 Acyltransferase OH

H N

H N O

O

RC(=O)NHR’

NH2 N

OOO P O P O O O

N

N N

O

O OH O P OH O-

Fig. 6 A survey of conjugation reactions known to occur in aquatic organisms

Phenols, either intact or produced via phase-I metabolism such as hydroxylation, demethylation, or ester cleavage, are the main substrates. Enzyme activity has been reported in various aquatic organisms such as bivalva, crustacea, green algae, larval chironomids, and gumboot chiton. The Km and Vmax values in each species,

Bioconcentration, Bioaccumulation, and Metabolism

75

estimated from the Lineweaver-Burk plots, showed not only substrate specificity but also a substrate-dependent affinity of PAPS to sulfotransferase, together with sex differences (Li and James 1993; Schell and James 1989). Enzyme activity was induced by exposing short-necked clams to phenol derivatives (Kobayashi et al. 1987) and the mussel Mytilus edulis to dispersed crude oils and PAHs (Lavado et al. 2006), whereas its reduction was reported for gumboot chiton exposed to lanosol (De Busk et al. 2000). Bivalves living in phenol-contaminated seawaters near industrial sites have a significantly higher sulfotransferase activity than those that reside in non-polluted areas, providing evidence that the enzyme is induced by contaminants (Oshima et al. 1994). Baldwin and LeBlanc (1994a) reported enhanced enzyme activity in D. magna in the metabolism of testosterone by 20-hydroxyecdysone; the enhancement effect is limited to a few phase-I metabolites. In contrast, sulfotransferase was partly involved in the elimination of testosterone from D. magna and insignificantly affected by the presence of diethylstilbestrol (Baldwin et al. 1995). Sulfotransferase activity in the cytosol fraction of the spiny lobster hepatopancreas was found to be insignificant, and this fraction had decreased activity in the antennal glands, indicating the presence of some inhibitor (Schell and James 1989). Janer et al. (2005b) reported high activity of cytosolic sulfatase in the digestive glands of eastern oysters, where the Km value of p-nitrophenol to that in spiny lobster was comparable, but had a much lower estimated Vmax value for sulfotransferase. Therefore, the presence of sulfatase may account for the low or reduced sulfotransferase activity. The optima for pH and temperature of this enzyme are approximately 6–9 and ≥35◦ C, respectively (Elmamlouk and Gessner 1978; Lavado et al. 2006; Li and James 1993; Schell and James 1989; Yoshida et al. 2002), and are highly speciesdependent. The range of these pH and temperature optima profiles indicate the presence of isozymes, which has been demonstrated by Yoshida et al. (2002). Using successive column chromatography, these authors purified two isozymes that catalyze the sulfation of 11-hydroxy saxitoxin from cytosolic fractions of dinoflagellates. From SDS-PAGE and gel filtration analyses, Yoshida et al. (2002) demonstrated that one of the isozymes is monomeric and had a molecular weight of 65 kDa.

4.2.6 Glutathione-S-Transferases (GST) These enzymes are widely distributed in terrestrial and aquatic organisms and are known to catalyze the transfer of tripeptide glutathione to electrophilic chemicals such as epoxides, halides, and arene oxides that are formed via phase-I oxidation (James 1986, 1987, 1994; Lech and Vodicnik 1985). The formed conjugate undergoes further metabolism through catalysis by peptidases and N-acetyltransferase via two intermediates and finally to a mercapturic acid conjugate (Fig. 6c). Wilbrink et al. (1991a) identified the mercapturic acid conjugate of 1-chloro-2,4dinitrobenzene (CDNB) in water depurated from CDNB-exposed pond snails. The activities of these enzymes are spectrophotometrically determined by using reduced

76

T. Katagi

glutathione and electrophilic substrates (Fig. 5c) and are known to vary according to whether the substrates originate in crustaceans, bivalves, larval chironomids, oligochaetes, or aquatic macrophytes (Clark 1989; LeBlanc and Cochrane 1985; Pflugmacher et al. 2000; Rakotondravelo et al. 2006; Schrenk et al. 1998; Stenersen et al. 1987; Teisseire and Vernet 2001; Vidal and Narbonne 2000). CDNB is the most popular substrate for GST enzymes, although aquatic organisms appear to have relatively low GST activity. GST activity in aquatic species has mainly been found in the cytosol fraction of hepatopancreas, gills, green glands, gonads and digestive glands, and levels found are variable (James 1986). To clarify the relative distribution of GST between microsomal and cytosolic fractions, thorough washing of the microsomal fraction is necessary; such washing excludes contamination resulting from the residual cytosolic GST (Wilbrink et al. 1991a). In bivalve, GST activity is predominantly located in digestive glands and visceral mass (Fitzpatrick and Sheehan 1993; Petushok et al. 2002; Vidal and Narbonne 2000; Wilbrink et al. 1991a). In gumboot chiton, more activity was observed in intestine than in digestive glands (Schlenk and Buhler 1988). The cytosolic fraction (from hepatopancreas of crayfish, prawn, and crab) of crustaceans has a much higher content of GST. However, the microsomal fraction from freshwater crayfish has been reported to contain about one fourth as much GST activity as does the cytosolic one (Arun et al. 2006; Johnson and Corbett 1986a; LeBlanc and Cochrane 1985; Lindström-Seppä et al. 1983). A lower, but moderate GST activity has been reported in the microsomal fraction from the oligochaete Lumbriculus variegatus (ContardoJara and Wiegand 2008). GST activity was also observed in the cytosol fraction of larval chironomids (Yuen and Ho 2001). In emergent and submergent aquatic macrophytes and various algae, GST activity toward typical substrates, such as CDNB and 1,2-dichloro-4-nitrobenzene (DCNB), were detected in both microsomal and cytosolic factions (Pflugmacher and Steinberg 1997; Pflugmacher et al. 1999, 2000). The GST activity in freshwater green algae and diatoms was significantly lower for atrazine (127) than for CDNB (Tang et al. 1998b). As for mammals, which have at least seven multi-gene enzyme families ( α-, μ-, and π-classes), there are many GST isozymes in aquatic organisms that can conjugate CDNB. These generally consist of two subunits that have a molecular weight of 23–28 kDa (Clark 1989; James 1994; Stenersen et al. 1987). GSTs that foment the conjugation of ethacrynic acid (ETHA), DCNB, and 1,2-epoxy3-(4-nitrophenoxy)propane (ENPP) are categorized into π-, μ-, and θ-classes, respectively (Vidal et al. 2002). The range of pH values that are optimal for GST activity in aquatic invertebrates has been reported to be 7.5–8.0; however, the optimal temperature varies with species. The optimal pH and temperature in bivalves were 8.2 and 30◦ C, respectively, in the pond snail (Wilbrink et al. 1991a), 6.5 and 45–50◦ C, respectively, in a tropical freshwater snail (Abdalla et al. 2006), and approximately 7, with a linear increase of activity up to 40◦ C, respectively, in clam (Vidal and Narbonne 2000). Freshwater green algae and diatoms showed an optimal pH of 7–9, with a linear increase of activity up to 45◦ C (Tang et al. 1998b). In larval chironomids, the optima values for pH and temperature were 8 and 30–40◦ C, respectively (Yuen and Ho 2001).

Bioconcentration, Bioaccumulation, and Metabolism

77

The enzymology of GSTs and their isozymes has been examined in various aquatic species by purification using chromatography, followed by electrophoretic and immunochemical analyses. GSTs of the mussels Mytilus edulis and M. galloprovincialis were partially purified by affinity and anion-exchange chromatography, and showed the presence of four or five isozymes consisting of homodimers whose subunits had molecular weights of 24.5, 26.5, and 27.3 kDa from SDS-PAGE analyses (Fitzpatrick and Sheehan 1993). In the cytosolic fraction of clam, 24 and 26 kDa bands were recognized by the antibody (Hoarau et al. 2001). Vidal et al. (2002) analyzed the purified isozymes from the cytosolic fraction of visceral mass in Corbicula fluminea by SDS- and non-denaturing PAGE, and identified three dimeric proteins with molecular weights of 45, 55, and 64 kDa, each consisting of four different subunits (27–30 kDa). The immunoblot analysis, using the antiserum for the rat GST subunit, allowed successful categorization of the major subunits to π-class. The freshwater tropical snail Bulinus truncatus has four subunits, two of which are major ones, each having a molecular weight of 23.6 kDa (Abdalla et al. 2006). For crustaceans, each of four fractions, prepared from cytosolic GSTs of daphnids by saturation with ammonium sulfate, demonstrated different activities toward CDNB and ETHA. Such behavior indicates the presence of two isozymes (LeBlanc and Cochrane 1985). Dierickx (1987), using column chromatography, isolated four cationic isozymes and one neutral isozyme, with different activities toward CDNB, from Daphnia magna. Furthermore, by using affinity chromatography, LeBlanc and Cochrane (1987) purified six anionic GSTs from D. magna that had molecular weights of 55–61.7 kDa. These GSTs were responsible for about 40% of cytosolic GST activity, and it was shown by SDS-PAGE analysis that at least three monomers (27.5, 28, and 30.2 kDa) exist. In addition, these GSTs were characterized by several methods that included enzyme multiplicity, molecular weight, isoelectric points, and immunochemical distinction. At least seven glutathione-binding proteins were identified from application of these methods, four of which could conjugate CDNB (Baldwin and LeBlanc 1996). In the amphipod Gammarus italicus, a heterodimer consisting of subunits having molecular weights of 27 and 28 kDa was confirmed, in addition to a homodimer of 28 kDa subunits (Aceto et al. 1991). The heavier heterodimer was identified as a π-class subunit by Western Blot analysis. The other heterodimer did not interact with antisera from α-, μ-, and π-GST. Through the successive column chromatographic purification of the cytosolic fraction, five proteins with GST activities toward CDNB, comparable to those in mammals, were identified in larval chironomids (Yuen and Ho 2001). SDS-PAGE analysis showed the homogeneity of these proteins; the subunits each had molecular weights of 23 kDa and an isoelectric point of 5.5. Amino acid sequences of these isozymes were highly conserved at the N-terminus. Cytosolic GSTs, isolated from marine rotifers, have been identified by electrophoresis to be monomers with three isozymes having a molecular weight of 30, 31.4, and 33.7 kDa, in Brachionus plicatilis, and two isozymes of 26.3 and 28.5 kDa in B. calyciflorus; these isozymes may be primitive types of GST (Bowman et al. 1990). Exposure of the Mediterranean clam to DDE (34) induced isozymes that could catalyze conjugation of ETHA and CDNB (Hoarau et al. 2001). The

78

T. Katagi

CDNB-conjugating GST was also induced by benzo[a]pyrene (7) and was categorized as π-class by Western Blot analysis. The substance 1-iodo-2,4-dinitrobenzene was a better substrate for GST than CDNB and DCNB in aquatic macrophytes, and enzyme activity increased in aquatic plants at sites contaminated with PAH (Schrenk et al. 1998). CDNB and pentachlorophenol (32) enhanced GST activity toward CDNB in daphnids, but CDNB did not modulate activity toward EHTA (LeBlanc and Cochrane 1985). 2,4-Dichlorophenol also increased its activity toward CDNB in the digestive glands of freshwater mussels (Petushok et al. 2002) and in marine diatoms (Yang et al. 2002). PB enhanced the GST activity in D. magna by a factor of two, while the conjugation rate with glutathione for CDNB increased by only 30% (Baldwin and LeBlanc 1996). In contrast, exposure to neither tributyltin (16) nor PCB altered the activity of cytosolic GSTs in D. magna (Schmidt et al. 2006). The exposure of D. magna and crayfish to fenitrothion (41) at the ppb level did not significantly affect GST activity (Damásio et al. 2007; Escartin and Porte 1996), but increased activity was observed at ppm levels in crayfish, possibly because of GST-catalyzed O-demethylation (Birmelin et al. 1998). This same metabolic pathway was confirmed for (41) in crabs by Johnston and Corbett (1986a). These authors reported either no inhibition by SKF-525A or enhanced formation of a key metabolite of (41) in the cytosolic fraction, in proportion to the concentration of glutathione. Meems et al. (2004) reported increased cytosolic GST activity toward CDNB when D. magna was exposed to cypermethrin (82) or natural organic matter isolated from rivers. The cytosolic GST activity of Chironomus tentans was increased by 34% in the presence of DDT (33), when CDNB was used as a substrate (Rakotondravelo et al. 2006). Exposure to contaminated sediments has also increased GST activity in the oligochaete Lumbriculus variegatus. Atrazine (127), in water and sediments, increased GST activity in the worm toward CDNB; the increase was more extensive in microsomal GSTs, which were minor components, than in cytosolic ones (Contardo-Jara and Wiegand 2008). Paraquat (149) slightly increased the cytosolic GST activity in this same oligochaete at 5 ppb but significantly inhibited activity at 0.5 ppm (Wiegand et al. 2007). Many chemicals are known to inhibit a GST activity. Chlorophenol derivatives were found to inhibit the GST activity of D. magna toward CDNB, with extent of inhibition being dependent on the lipophilicity of the inhibitors (LeBlanc and Cochrane 1987). The mechanism of GST inhibition for phenols was investigated in gumboot chiton by De Busk et al. (2000), using the brominated phenol, lanosol. Lineweaver-Burk plots were used to determine how CDNB reacted to reduced glutathione at varied concentration of lanosol. The tested phenol was considered not to be a competitive inhibitor and was regarded, from its electrophilic nature, to be likely to bind near the active site of GST. The activities of the GST isozymes isolated from D. magna were also inhibited by benzoquinone and 2,4-D (75), neither of which competed with glutathione (Dierickx 1987). The changes in Km and Vmax values, estimated from Lineweaver-Burk plots, indicated that benzoquinone is a competitive inhibitor of CDNB, but 2,4-D was bound to the site in a manner different than standard substrate-binding.

Bioconcentration, Bioaccumulation, and Metabolism

79

4.2.7 Acyltransferases Carboxylic acid, intact or as a phase-I metabolite, is considered to be formed from ligase-catalyzation of the corresponding coenzyme A (CoA) complex. This complex reacts with amino derivatives to form amides with the aid of acyltransferase (Fig. 6d); however, this enzymatic profile has not been extensively investigated in aquatic organisms. Acyltransferase activity is usually estimated by monitoring the formation of the metabolites it foments. Janer et al. (2005a) reported acyltransferase activity in the microsomal fraction from gonads and digestive tubes of the sea urchin and amphipod Hyalella azteca. In so doing, these authors used palmitoyl-CoA as a co-substrate in the metabolism of testosterone; tributyltin (16) was found to be an inhibitor. In studying the metabolism of testosterone and estradiol, higher levels of enzyme activity were found in gonads of Mytilus edulis; enzyme induction also occurred and resulted from exposure to dispersed crude oils and PAHs (Lavado et al. 2006). Conjugation with taurine has been observed in the metabolism of aryloxyacetic acids by spiny lobster (James 1982) and crayfish (Barron et al. 1991). The conjugate was located in the hepatopancreas of these organisms and was eliminated via urine. Acetylation and formylation reactions are also known to occur mostly in algae, macrophytes, and bivalves. Such reactions have been reported for many aniline derivatives, either intact ones or ones that result from phase-I products via reduction of a nitro group or cleavage of an N(H)–C(=O) bond (Tables 11 and 12). O- and Nmethylation reactions have been commonly observed for phase-I and -II metabolism of nitro, amino, phenol, and carboxylic acid derivatives, but the relevant enzymology is obscure. The successive acylation of glucose conjugation, mainly at 6-O position, is also known to occur in algae and macrophytes (Day and Saunders 2004; Fujisawa et al. 2006; Pascal-Lorber et al. 2004; Petroutsos et al. 2007; Tront and Saunders 2007); this reaction also is catalyzed by acyltransferases.

4.3 Metabolism of Pesticides and Other Chemicals 4.3.1 Polycyclic Aromatic Hydrocarbons and Other Simple Chemicals Metabolic pathways known for PAHs and similar chemicals in aquatic organisms, other than fish, are listed in Table 12. One of the most common pathways, which affects PAHs and similar compounds, is ring hydroxylation by oxidases, followed by conjugation with sulfate, glucose, and/or glucuronic acid. The dihydroxydihydro derivatives generally have a cis configuration from catalytic conversion by dioxygenases, but the trans derivatives of phenanthrene and pyrene have also been reported (Narro et al. 1992a; Wildi et al. 1994). Carbohydrate conjugates usually show a β-linkage at the ether moiety, but Warshawsky et al. (1990) found that conjugates were hydrolyzed by α-glucosidase in the metabolism of benzo[a]pyrene (7) by the green alga Selenastrum capricornutum. Reductive transformation generally proceeds for simple aromatics that have nitro groups, and such transformation is followed by N-acetylation and formylation. Among aquatic macrophytes and some

Biphenyl

Naphthalene

PAH 1

2

S: Pandalus platyceros

B: Oscillatoria sp. Sn: Lymnaea stagnalis B, D, G + brown and red algae

Speciesa

Anthracene

Phenanthrene

Pyrene

Fluoranthene

3

4

5

6

O3 (1-methoxy), O4 (trans-9,10) O3 (ns), O4 (ns) O3 (ns) O3 (1), C3 (1) + unknown conjugates O4 (ns, trans) O3 (1) O3 (1) O3 (ns), O4 (ns) O3 (3, 8)

B: Armenellum quadruplicatum G: Selenastrum capricornutum W: Daphnia magna

I: Chironomus riparius P: Nereis diversicolor O: Lumbriculus variegatus G: Selenastrum capricornutum P: Capitella capitata

O3 (ns), O4 (ns)

O3 (ns)

O3 (1), O4 (1,2), O5 (1,4), C1 (O-β-Gla), C2 (O-β-Glu), C3 O3 (1, NIH shift), O7, C2 (O-β-Glu)

O3 (4; 2 and 3, traces), O4 (4,4 ) O3 (4, 2) O3 (1), O4 (cis-1,2), O5 (4-OH-1-tetralone)

Typeb

G: Selenastrum and Scenedesmus sp.

W: Daphnia magna and pulex

1-Naphthalenesulfonic acid G (six species)

Pesticide/structure

No.

Kneifel et al. (1997); Soeder et al. (11987) Herbes and Risi (1978); McCarthy (1983) Chan et al. (2006); Safonova et al. (2005) Narro et al. (1992a) Chan et al. (2006) Akkanen and Kukkonen (2003) Ikenaka et al. (2006) Wildi et al. (1994) Christensen et al. (2002) Lyytikäinen et al. (2007) Chan et al. (2006) Forbes et al. (2001)

Narro et al. (1992b); Todd et al. (2002) Sanborn and Malins (1980)

Cerniglia et al. (1980b) Wilbrink et al. (1991b) Cerniglia et al. (1979, 1980a,c),

Referencesc

Table 12 Metabolism of polycyclic aromatic hydrocarbons and selected other chemicals in aquatic organisms

80 T. Katagi

Pesticide/structure

Benzo[a]pyrene

Dibenzofuran Dibenzo-p-dioxin

Chlorobenzene

Nitrobenzene

No.

7

8 9

–

–

G: Oedogonium cardiacum Sn: Physa sp. W: Daphnia magna G: Oedogonium cardiacum Sn: Physa sp. W: Daphnia magna

Selected other chemicals

L: Panulirus argus G: Ankistrodesmus sp. G: Scenedesmus sp.

A: Rhepoxynius abronius

O3 (2) O3 (2,4), O4 (3,4) O3 (2,4) O3, R3 (NH2 ), C5 (acetyl) O3, R3 (NH2 ), C5 (acetyl) O3, R3 (NH2 )

O3 (9, 1, 3), O4 (9,10; 4,5; 7,8), O5 (1,6; 3,6; 6,12), C1 (O-β-Gla), C3 O3 (3), C1 (O-β-Gla) or C3 O4 (7,8; 9,10) O3 (12,6, 9, 3), O4 (7,8; 9,10) O3 (3), O4 (9,10; 7,8), O5 (ns) O3 (ns) O3 (3, 7), O4 (9,10; 7,8), C1 (O-β-Gla) or C3 O3 (3, 9), O4 (9,10; 7,8), O5 (ns), C1 or C3 O3 (3), C2 (O-β-Glu, 3), C3 (3) O3 (2, 3, 4), O4 (ns) O3 (2), O4 (ns)

Mu: Mytilus galloprovinciallis

C: Spaherium corneum Mercenaria mercenaria Oy: Crassostrea virginica Ch: Cryptochiton stelleri W: Daphnia magna I: Chironomus riparius

O4 (cis-11,12; cis -9,10; cis -7,8; cis -4,5) O5 (3,6), C2 (O-α, β-Glu; 4,5), C3 (4,5)

Typeb

Table 12 (continued)

G (five species)

Speciesa

Lu and Metcalf (1975)E Lu and Metcalf (1975)E Lu and Metcalf (1975)E Lu and Metcalf (1975)E Lu and Metcalf (1975)E Lu and Metcalf (1975)E

Schell and James (1989) Todd et al. (2002) Todd et al. (2002)

Borchert et al. (1997) Anderson (1985) Anderson (1985) Schlenk and Buhler (1988) McCarthy (1983) Borchert et al. (1997); Leversee et al. (1982) Reichert et al. (1985)

Warshawsky et al. (1988, 1990) Michel et al. (1995)

Schoeny et al. (1988)

Lindquist and Warshawsky (1985a, b)

Referencesc

Bioconcentration, Bioaccumulation, and Metabolism 81

Pesticide/structure

Phenol

Catechol

4-Cl-phenol 2,4-Cl2 -phenol

2,4-(CH3 )2 -phenol 4-NO2 -phenol 3-CH3 -4-NO2 -phenol

3-CF3 -4-NO2 -phenol Bisphenol A

No.

–

–

– –

– – –

– 11

C2 (O-β-Glu, 6O-malonyl- or pentosyl-β-Glu) et al. (2004); Tront and Saunders (2007) C2 (O-β-Glu, 6O-acetyl-β-Glu) C2 (O-β-Glu, 6O-malonyl-β-Glu) C2 (O-β-Glu, 6O-malonyl-β-Glu) C2 (6O-pentosyl-β-Glu, 6O-malonyl-β-Glu) C2 (O-β-Glu, 6O-malonyl-β-Glu) NM NM NM O1 (4), O3 (6), R4 R3 (NH2 ), C3, C5 (acetyl), C7 (N) C2 (O-β-Glu) O1, C2 (O-β-Glu) R3 (NH2 ), C1 (O-β-Gla), C3 C2 (O-β-Glu)

Lemna minor

Mentha aquatica C: Sphaerium corneum O: Lumbriculus variegates I: Chironomus riparius G: Chlorella and Scenedesmus sp. Sn: Tegula funebralis Mc: Lemna gibba Cr: Procambarus clarkii I: Chironomus tentans Mc: Lemna minor

Salvinia natans Myriophyllum spicatum Hippuris vulgaris Glyceria maxima

O3 (2), O8, M (ring cleavage) C2 (O-β-Glu) ns O8, M (ring cleavage) C2 (O-β-Glu, 6O-acetyl-β-Glu) C2 (O-β-Glu)

Typeb

Table 12 (continued)

Go: Ochromonas danica Mc: Lemna gibba Six freshwater microalgae Go: Ochromonas danica G: Tetraselmis marina Mc: Lemna gibba

Speciesa

Pascal-Lorber et al. (2004) Guerrero et al. (2002) Guerrero et al. (2002) Guerrero et al. (2002) Klekner and Kosaric (1992) Williamson et al. (1995) Fujisawa et al. (2006) Foster and Crosby (1986) Kawatski and Bittner (1975) Nakajima et al. (2004)

Pascal-Lorber et al. (2004) Pascal-Lorber et al. (2004) Pascal-Lorber et al. (2004) Pascal-Lorber et al. (2004)

Semple and Cain (1996) Barber et al. (1995) Ellis (1977) Semple and Cain (1996) Petroutsos et al. (2007) Ensley et al. (1994); Sharma et al. (1997) Day and Saunders (2004); PascalLorber

Referencesc

82 T. Katagi

Pesticide/structure

Aniline

4-NH2 -toluene 2-NH2 -toluene

3,4-Cl2 -aniline Benzidine 2-(CH3 CONH)-toluene Anisole

4-NO2 -anisole

Trinitrotoluene

Phenoxyacetic acid

No.

–

– –

– 12 – –

–

15

–

I: Chironomus tentans A: Hyallela azteca O: Tubifex tubifex Lumbriculus variegates L: Panulirus argus

Myriophyllum aquaticum

Cr: Procambarus clarkii B: Anabaena sp. Mc: Myriophyllum spicatum

B: Oscillatoria and Agmenellum spp. G: Oedogonium cardiacum Su: Strongylocentrotus sp. Mu: Mytilus edulis Oy: Crassostrea gigas Mc: Lemna gibba Sn: Physa sp. Mu: Mytilus edulis G: Oedogonium cardiacum Sn: Physa sp. Sn: Physa and Lymnaea sp. Su: Strongylocentrotus sp. Ch: Cryptochiton stelleri

Speciesa Cerniglia et al. (1981) Lu and Metcalf (1975)E Landrum and Crosby (1981) Knezovich and Crosby (1985) Knezovich and Crosby (1985) Fujisawa et al. (2006) Lu et al. (1977)E Knezovich and Crosby (1985) Lu and Metcalf (1975)E Lu and Metcalf (1975)E Hansen et al. (1972) Landrum and Crosby (1981) Landrum and Crosby (1981)

Referencesc

Foster and Crosby (1986) Pavlostathis and Jackson (1999) Hughes et al. (1997), Pavlostathis et al. (1998) O1, R3 (NH2 , 4<2; azoxy), C5 (2), O8 Bhadra et al. (1999); Hughes et al. (2, OH) (1997) Vanderford et al. (1997) R3 (NH2 ; 2, 4) Belden et al. (2005) R3 (NH2 ; 4>2), O1 Sims and Steevens (2008) R3 (NH2 ; 4>2) Conder et al. (2004) R3 (NH2 ; 2, 4) Belden et al. (2005) C6 (taurine) James (1982)

O3 (4), C5 (formyl, acetyl) C7 (N) O1, C5 (acetyl) O8 (NO, NHOH), C5 (formyl), C7 (N) O8 (NO), C5 (formyl), C7 (N) C2 (N-β-Glu), C6 (glutamic acid) C5 (acetyl), C7 (N) H3 O3 (2), conjugation O2, conjugation O2 O2, R3 (NH2 ), C5 (acetyl) O2, R3 (NH2 ), C2 (O-β-Glu), C3, C5 (acetyl) O2, C1 (O-β-Gla), C2 (O-β-Glu), C3 R3 (NHOH; azoxy) R3 (NH2 , 4>2; 4-NHOH; azo)

Typeb

Table 12 (continued)

Bioconcentration, Bioaccumulation, and Metabolism 83

Benzoic acid

Salicylic acid 4-Cl(I)-benzoic acid 4-Cl-3,5-(NO2 )2 -benzoic acid 3-Phenoxybenzoic acid

Phenylalanine Di-n-butyl phthalate

Di-2-ethylhexyl phthalate

Antipyrine Nonylphenol isomer Azo dyes

–

– – –

– 13

14

19 20 –

13 marine algae Oy: Crassotrea virginica S: Penaeus azecus G: Oedogonium cardiacum Sn: Physa sp. Mu: Mytilus califorianus Sn: Lymnaea stagnalis G: Chlorella and B: Oscillatoria sp.

Mc: Lemna gibba

Sn: Physa sp. W: Daphnia magna Mc: Lemna gibba B: Anabaena and Nostoc sp. G: Chlamydomonas reinhardtii

Speciesa

C2 (6O-malonyl-β-Glu), C6 (maleic acid) Mineralization of the phenyl ring H1 H1, conjugates H1, M (anhydride) H1, M (anhydride) O3 (4) O3 (3), C1 (O-β-Gla) R4, M (Cleavage of N-N bond)

O3 (2,3,4), O4 (1,2), C6 (glycine) O3 (2,3,4), O4 (1,2), C6 (glycine) C2 (O-β-Glu) H6 (4, halogen substitution) H6 (4, halogen substitution)

Typeb

Vose et al. (1971) Wofford et al. (1981) Wofford et al. (1981) Metcalf (1976)E Metcalf (1976)E Krieger et al. (1979) Lalah et al. (2003) Jingi and Houtian (1992)

Fujisawa et al. (2006)

Lu and Metcalf (1975)E Lu and Metcalf (1975)E Ben-Tal and Cleland (1982) Kuritz and Wolk (1995) Gutenkauf et al. (1998)

Referencesc

b Type

by EPI-Suite (USEPA 2008) or experimental data therein. of metabolism is defined and listed in Table 9. The term in parentheses refers to the metabolic position and associated details. c Superscript “E” means a model ecosystem. Go = golden algae; Su = sea urchin; Nonylphenol isomer: 4-(3 ,6 -dimethyl-3 -heptyl)phenol

a Estimated

–

Pesticide/structure

No.

Table 12 (continued)

84 T. Katagi

Bioconcentration, Bioaccumulation, and Metabolism

85

algae, N-glucosidation has also been reported for aniline derivatives (Fujisawa et al. 2006; Pflugmacher and Sandermann 1998a, b). Phenols are known to inhibit algal growth, but their extensive metabolism in some green and blue-green algae, where intermediate catechols were not bioaccumulated, resulted in no inhibitory effect (Pinto et al. 2002). Phenol is thought to be primarily oxidized by monooxygenases to the corresponding catechol derivative, whose ring is then cleaved by 2,3-dioxygenases (Semple et al. 1999). Another typical transformation for phenols is extensive conjugation with glucose and organic acids, especially in aquatic macrophytes (Pascal-Lorber et al. 2004). N- and O-methylation sometimes occurs as well, especially in bivalva (Knezovich and Crosby 1985; Lu et al. 1977; Williamson et al. 1995). Conjugation with amino and simple organic acids has been reported for benzoic acid derivatives (Fujisawa et al. 2006; Lu and Metcalf 1975). 4.3.2 Organochlorine Pesticides Metabolic reactions common to many organisms for organochlorine pesticides are summarized in Table 13. Molluscs and algae commonly metabolize cyclodiene insecticides by oxidation at the bridgehead carbon, and may form epoxides at the C=C bond as well. Epoxidation reactions also occur in ciliates and flagellates (Lal and Saxena 1982). Ring oxidation to form the corresponding ether, lactone, and/or aldehyde was observed for endosulfan (27). Reductive dechlorination and dehydrochlorination are among the most typical reactions for DDT (33) in various aquatic organisms, and the latter transformation was reported for lindane (30) (Goulding and Adams 1985). Mytilus edulis exhibited a unique metabolism of hexachlorobenzene (31) (Bauer et al. 1989). The main metabolite in this mussel was the thiomethyl derivative, which may be catalyzed by GST. In bivalva, pentachlorophenol (32) underwent conjugation at the phenol oxygen, with either sulfate or β-glucose. Oxidative dechlorination to form 1,4-dihydroquinone was observed for (32) in abalone (Shofer and Tjeerdema 1993; Tjeerdema and Crosby 1992). Elodea canadensis has been found to dechlorinate trichloroethene with the aid of a dehalogenase (Wolfe and Hoehamer 2003). 4.3.3 Organophosphorus Pesticides The metabolic profiles common for OPs in aquatic organisms are presented in Table 10. A common metabolic pathway is hydrolysis of the P–O bond, catalyzed by phosphotriester hydrolases, to form the corresponding phenols and alcohols (Fukuto 1990; Wheelock et al. 2005). In some cases, this reaction may proceed concomitantly via oxidative desulfuration by mixed-function oxidases (James and Boyle 1998). The carboxyl esters in malathion (57) and in other organophosphates (Khalil and Mostafa 1987) are hydrolyzed by CaE. Wolfe and Hoehamer (2003) revealed the involvement of acid phosphatases in hydrolysis of organophosphorus pesticides such as (57) in aquatic plants. GST catalyzes O-dealkylation at the P–O– alkyl moiety (Birmelin et al. 1998). Sulfur oxidation to the corresponding sulfoxide and sulfone is likely to be catalyzed by FAD-monooxygenases (James 1994; Lech

Pesticide/structure

Aldrin

Dieldrin

Chlordane

Heptachlor

Endosulfan

Lindane

No.

21

22

24

26

27

30

W: Daphnia magna Sn: Physa sp. G: Two Chlorella sp. B: Anabaena and Nostoc sp.

G: Scenedesmus sp.

O1 (9–OH, C=O) O1 (9–OH, C=O) O1 (9–OH, C=O) O1 (1), O8 (epoxide) O1 (1), O8 (epoxide) O1 (1), O8 (epoxide) H6 (1), O8 (epoxide) O7 (ether), O1 (lactone) O7 (diol),O1(hydroxyether, lactone), O6, MI(β) O7 (diol), O1 (ether, aldehyde), O6, MI (β) O6 R2 R2 R2

O8 (epoxide and diol) O1 (9–OH, C=O), O8 (epoxide) O8 (epoxide) O8 (epoxide) O8 (epoxide)

G: Dunaliella sp. Oedogonium cardiacum G, D: marine species I: Chironomus riparius Os: Chlamydotheca arcuata

G: Oedogonium cardiacum Sn: Physa sp. C: Corbicula manilensis G: Oedogonium cardiacum Sn: Physa sp. G: Oedogonium cardiacum Sn: Physa sp. B: Anabaena and Aulosira spp. B: Two Anabaena sp.

O8 (epoxide) O1 (9–OH, C=O), O8 (epoxide) O8 (epoxide)

Typeb

Mu: Mytilus califorianus Sn: Physa sp. B: Five species

Speciesa

Table 13 Metabolism of organochlorine pesticides in aquatic organisms

DeLorenzo et al. (2002) Metcalf et al. (1973b)E Sweeney (1969) Kuritz and Wolk (1995); Kuritz et al. (1997)

Sethunathan et al. (2004)

Krieger et al. (1979) Metcalf et al. (1973b)E Dhanaraj et al. (1989); Schauberger and Wildman (1977) Patil et al. (1972) Metcalf et al. (1973b)E Bowes (1972); Rice and Sikka (1973) Estenik and Collins (1979) Kawatski and Schmulbach (1971, 1972) Metcalf et al. (1973b)E Metcalf et al. (1973b)E Sanborn and Yu (1973)E Lu et al. (1975)E Lu et al. (1975)E Lu et al. (1975)E Lu et al. (1975)E Rao and Lal (1987) Lee et al. (2003)

Referencesc

86 T. Katagi

Pesticide/structure

Hexachlorobenzene

Pentachlorophenol

DDT

No.

31

32

33

O8 (OH) Substitution with SCH3 (1,4), SCH2 COOH (1) C3 C3 C3 C2 (O-β-Glu), C3, O5 (1,4-dihydro) C2 (O-β-Glu), C3, C7 (O), O5 (1,4-dihydro) C2 (O-β-Glu) R1, O3, C2 (O-β-Glu), C4, C7 (O) C3, C2 (O-β-Glu) NM ns (2 metabolites) R1, R2 R1, R2 R1, R2 R1, R2 R1, R2 R1, R2 R1, R2 R1, R2, O1 R1, R2 R2

Typeb

Table 13 (continued)

Mc: Lemna gibba Eichhornia crassipes W: Daphnia magna O: Lumbriculus variegates I: Chironomus riparius C: Indonaia caerula Katelysia opima Sn: Vivipara heliciformis Physa sp. Cipangopaludina japonica Oy: Crassostrea virginica G: Ankistrodesmus amalloides G: Dunaliella sp. B: Anabaena and Aulosira sp. D: Eleven species

C: Tapes philippinarum Mu: Mytilus edulis Oy: Crassostrea gigas Ab: Haliotis fulgens Haliotis rufescens

G: Oedogonium cardiacum Mu: Mytilus edulis

Speciesa

Sharma et al. (1997) Roy and Hänninen (1994) Kukkonen and Oikari (1988) Guerrero et al. (2002) Guerrero et al. (2002) Pillai et al. (1980) Kale et al. (1999a)E Yadav et al. (1978) Metcalf et al. (1971)E Ohkawa et al. (1980)E Brodtmann (1970) Neudorf and Khan (1975) Lal et al. (1987); Patil et al. (1972) Goulding and Ellis (1981) Keil and Priester (1969), Miyazaki and Thorsteinson (1972)

Kobayashi et al. (1970a, b) Ernst (1979) Shofer and Tjeerdema (1993) Shofer and Tjeerdema (1993) Tjeerdema and Crosby (1992)

Lu and Metcalf (1975)E Bauer et al. (1989)

Referencesc

Bioconcentration, Bioaccumulation, and Metabolism 87

DDT

DDE Methoxychlor

Dichlobenil

33

34 37

38

R1, R2, O1 R2 R2 R1 R1, R2 R2 R1, R4 (C=C), O8 (ketone) O2 O2, R2

S: Palaemonetes kadiakensis A: Gammarus fasciatus Hyalella azteca Diporeia sp. I: Ephemera danica Chironomus riparius C: Arcid blood clam Sn: Physa and Lymnaea sp. Physa sp. O3 (3), H6 (cyano), O-conjugation

R2

W: Duphnia pulex and magna

Mc: Myriophyllum brasiliense

R2 R1, R2 R1, R2 R1, R2

Typeb

Ci: Blepharisma intermedium Mc: Myriophyllum aquaticum Spirodela oligorrhiza Elodea canadensis

Speciesa

Saxena et al. (1982) Gao et al. (2000b) Gao et al. (2000b) Garrison et al. (2000); Gao et al. (2000b) Johnson et al. (1971), Neudorf and Khan (1975) Johnson et al. (1971) Johnson et al. (1971) Lotufo et al. (2000) Lotufo et al. (2000) Södergren and Svensson (1973) Johnson et al. (1971) Mehetre et al. (2002) Hansen et al. (1972) Kapoor et al. (1970)E ; Metcalf et al. (1971)E Sikka et al. (1974)

Referencesc

b Type

by EPI-Suite (USEPA 2008) or experimental data therein. of metabolism is defined and listed in Table 9. The term in parentheses refers to the metabolic position and associated details. c Superscript “E” means a model ecosystem.

a Estimated

Pesticide/structure

No.

Table 13 (continued)

88 T. Katagi

Bioconcentration, Bioaccumulation, and Metabolism

89

and Vodicnik 1985), and has been reported for demeton-S-methyl (51), disulfoton (53), and fenamiphos (67) (Anderson and Zhu 2004; Cáceres et al. 2008; Dauberschmidt et al. 1997). OP metabolites that result from phase-I metabolic reactions are further conjugated with carbohydrates and sulfate. Glucoside conjugation is the predominant pathway for OP metabolism in snails, shrimp, and crabs (Johnston and Corbett 1986a, b; Takimoto et al. 1987a, b; Kobayashi et al. 1990), but glucuronidation was additionally observed in crayfish (Foster and Crosby 1986). Such metabolic profiles are common to most aquatic organisms as demonstrated by the extensive studies performed on fenitrothion (41) in many species. The reductive transformation of organophosphates that have nitro group, such as parathion (39), (41), and EPN (70), takes place in snails, crabs, Daphnia, and algae (Francis et al. 1980; Megharaj et al. 1994; Mostafa et al. 1991; Takimoto et al. 1987a). Barton et al. (2004) confirmed that the nitroso derivative of (39) served as a reductive intermediate when metabolized by the blue-green alga Anabaena sp. Reductive debromination of leptophos (71) was reported by Francis et al. (1980) in a model ecosystem study. In the metabolism of chlorpyrifos (47) by oysters, the 6-thiomethyl derivative was identified by GC-MS (Woodburn et al. 2003). The mercapturic acid or cysteine derivative of (47), produced by GST catalysis and successive enzyme action, is considered to be further transformed successively by C–S β-lyase and thiol methyltransferase. 4.3.4 Other Pesticides Members of other pesticide classes are diverse and include esters, amides, carbamates, and ureas, together with others not classified as to a common chemical structure. The metabolic profiles of many of these “other” pesticides are often similar to those of PAHs, organochlorines, and organophosphates (Table 11). Organic acids are formed from some pesticides by enzymatic hydrolysis of esters and may be subsequently methylated (Getsinger et al. 2000; Wang et al. 1992a) or conjugated with amino acids such as taurine (Barron et al. 1991; James 1982). Amides, carbamates, and ureas are enzymatically hydrolyzed, and the resulting anilines are subjected to N-acetylation (Mitsou et al. 2006; Tsorbatzoudi et al. 1976; Tweedy et al. 1970). Acetylation, formylation, and methylation at either an oxygen or nitrogen, together with O-glucosidation, are sometimes reported in green algae (Metcalf et al. 1975; Schuphan 1974; Watanabe et al. 1987). However, oxidative transformation at the N-alkyl group and phenyl ring appears to occur more often in aquatic species. The successive oxidation of the N-alkyl group ultimately produces the NH2 derivative; the intermediate N-hydroxymethyl form has been identified for propoxur (100) in green alga (Metcalf 1976). Similar to what occurs with OP compounds, carboxin (94) and thiobencarb (109) underwent S-oxidation (Balasubramanya and Patil 1980; Chen et al. 1982). Glutathione conjugation was observed in the metabolism of isoproturon (118), an aquatic macrophyte, whose metabolites were identified by LC-MS (Pietsch et al. 2006). In addition to the usual cleavage of C–O and C–N bonds in esters and amides, a unique cleavage is known for the ether linkage in diphenyl ethers (Cai et al. 2007, 2008; Lee et al. 1976), the S–N bond of bentazone

90

T. Katagi

(137) (Booth et al. 1973), and the Sn–C bond of tributyltin (16) (Lee et al. 1989; Maguire et al. 1984). 4.3.5 Species Differences There are many examples that disclose metabolic differences among species. Such differences may result from differential uptake mechanisms or from different titers of relevant enzyme systems in any particular organism. Even among the same phylum and class, different metabolic profiles are sometimes observed. A similar oxidative metabolism of benzo[a]pyrene (7) that had been added to sediment was observed for the marine amphipods, Rhepoxynium abronius and Eohaustorius washingtonianus. However, higher metabolic activity was observed in the former species, which formed the metabolite 4,5-dihydrodiol (Reichert et al. 1985). Lotufo et al. (2000) identified significant differences in metabolic activity between the two freshwater amphipods Hyalella azteca and Diporeia sp., when these species were exposed to DDT (33) in water. The former amphipod metabolized 64% of (33) to DDE (34) over a period of 10 days, but did not form DDD (35); the latter amphipod slightly transformed (33), but only to (35) over a 28-day period. In another case, metabolic oxidation was involved; more O-demethylation of methoxychlor (37) and p-nitroanisole proceeded in Physa elliptica as compared with Lymnaea pallustris (Hansen et al. 1972). The blue-green alga Anabaena sp. predominantly metabolized endosulfan (27) via ring opening to form the corresponding diol; however, A. flos-aquae produced a significant amount of an unknown metabolite other than the hydroxyether, lactone, and sulfate derivatives (Lee et al. 2003). Even among green algae, a slightly different metabolism profile was reported for (27). Both Scenedesmus and Chlorococcum sp. mainly produced the sulfate metabolite from the α-isomer of (27), but much lower amounts of the β-isomer was converted to the aldehyde metabolite by the former alga (Sethunathan et al. 2004). Greca et al. (2008) utilized eleven green algae to examine the metabolic profiles of ethinylestradiol. Insignificant metabolism was observed in eight algae, whereas the remaining ones showed enantio- and region-selective hydroxylation. Ring hydroxylation at the 10βand 6α-positions was confirmed in Scenedesmus quadricauda and Ankistrodesmus braunii, respectively, and Selenastrum capricornutum mainly produced a glucose conjugate of the parent molecule, together with minor hydroxylated metabolites at 2- and 6β-position. Among different species, it is common to observe significantly different metabolic profiles of the same substance. Takimoto et al. (1987b) compared the metabolism of fenitrothion (41) among fish, molluscs, crustaceans, and algae. Reduction of the nitro group predominated in molluscs, while oxidative desulfuration was mainly observed in crustaceans and algae. Both ester cleavage and O-demethylation were commonly detected, but the rates and degree of following conjugation reactions were variable. p-Nitroanisole (NA) has been extensively metabolized by several aquatic organisms, but with variable profiles (Landrum and Crosby 1981). Mussels could not metabolize this chemical, whereas oxidative O-demethylation proceeded in starfish, sea cucumber, and gumboot chiton,

Bioconcentration, Bioaccumulation, and Metabolism

91

followed by conjugation with sulfate and glucose. In sea urchin, NA was reduced to the corresponding aniline, followed by acetylation. A major secondary metabolic pathway for acrolein (144) differed between clam and crayfish (Nordone et al. 1998). In clams, acrylic acid was formed via oxidation and then was predominantly reduced to propiolic acid. In crayfish, the main pathway was reduction of (144) to allyl alcohol followed by epoxidation. More metabolic dissipation of DDT (33) was observed in Daphnia pulex than in the alga Ankistrodesmus amalloides, although reductive dehydrochlorination to DDE (34) and DDD (35) via dechlorination was also detected in this freshwater alga (Neudorf and Khan 1975). Among benthic organisms, the shrimp Pandalus platyceros almost completely metabolized benzo[a]pyrene (7), but lower to insignificant metabolic activity was observed in two amphipods and a clam, respectively (Varanasi et al. 1985). The substance (7) is hydroxylated at the 3-position in larval Chironomus riparius, followed by conjugation with glucuronic acid and sulfate in an 83% conversion over a 20-hr period; the transformed percentage was less than 25% in snails and no conjugation occurred (Borchert et al. 1997). Oxidative metabolism of isoproturon (118) was observed to be a main pathway in algae. The extent of such oxidative metabolism was larger in Anabaena sp. than in Chlorella sp. and proceeded via stepwise N-demethylation, followed by the C–N bond cleavage in the urea moiety (Mostafa and Helling 2001). Lee et al. (1989) examined the metabolic activity of several estuarine algae toward tributyltin (16). Golden and green algae had no, or insignificant, activity in producing the corresponding dibutyl metabolite. However, significant metabolism of (16) was observed for two diatoms and a dinoflagellate, wherein there was light-induced formation of hydroxylated metabolites. Under lighting conditions that caused no growth inhibition, the metabolic activity of various algae was examined for benzo[a]pyrene (7). Activity was observed to decrease in the following order: green > blue-green > golden algae; the major resulting metabolites formed were the 3,6-dione and 9,10-dihydrodiol (Schoeny et al. 1988). The alligator weed, an aquatic plant, could not metabolize dichlobenil (38), whereas ring hydroxylation at the 3-position predominantly proceeded in parrot feather. Thereafter, conjugation and stepwise hydrolysis of the cyano group proceeded for parrot feather (Sikka et al. 1974). A different conjugation pathway for 2,4-dichlorophenol has been reported in aquatic plants (Pascal-Lorber et al. 2004). Although 2,4-D was primarily conjugated with glucose in all tested macrophytes, successive reactions produced O-acetylation of the hydroxymethyl group at the 6-position in Salvinia natans and O-malonylation in four other macrophytes. In Lemna minor and Glyceria maxima, a unique conjugation with pentose was found.

5 Behavior of Pesticides in Larger-Scale Systems Aquatic organisms are exposed by many different means to pesticides and industrial chemicals that reach aquatic bodies (Katagi 2006). The residue levels of such substances that enter organisms are controlled by rates of uptake, metabolism,

92

T. Katagi

and excretion (Fig. 1). Simply designed laboratory studies can clarify the relative importance of each of these processes, regarding the fate of a chemical, but such studies are incapable of fully emulating processes that occur in the natural environment. The reason for this is that there are various abiotic and biotic processes that take place as biomagnification through the food web proceeds. Moreover, field studies that incorporate all processes are generally difficult to control, and generate enormous amounts of complex data, which is costly and difficult to analyze. Therefore, many researchers first rely on laboratory-scale studies or more complex model ecosystems; such studies may utilize multiple aquatic species that are normally part of the food web (Metcalf and Sanborn 1975; Miyamoto et al. 1985). In addition, many kinds of field microcosm and mesocosm studies have been conducted. In many cases, the purposes of such studies are focused on addressing specific issues such as ecotoxicological effects and how to mitigate them (Miyamoto et al. 2008).

5.1 Model Ecosystems Depending on study design, model ecosystems can address quite a range of environmental circumstances. Soil or bottom sediments may be present or absent. A radiolabeled pesticide may be applied to any compartment of an ecosystem (such as water or bottom sediment) to simulate spray drift or a runoff event, etc. Aquatic model ecosystems can be classified as being terrestrial-aquatic (Metcalf et al. 1971; Metcalf and Sanborn 1975), aquatic (Isensee 1976; Schuth et al. 1974), and/or recirculating static (Ambrosi et al. 1978; Isensee et al. 1976, 1979a,b), and sitespecific models have been developed that emulate river (Lynch et al. 1982; Maki and Johnson 1977; Rose and McIntire 1970) or paddy field (Chen et al. 1982; Lee et al. 1976; Wang et al. 1992a) conditions. In Table 14 a long list of substances is presented that have been studied in model systems of one design or another. Lu and Metcalf (1975) developed a flask-scale system of the simplest design. This system included only a few aquatic species and was intended to examine bioaccumulation and metabolism of simple aromatic compounds. A terrestrial-aquatic model system simulates real agronomical practices of pesticide application to fields that are followed by runoff events. This system consists of a sloping shelf of sand in water with several aquatic organisms. The terrestrial end of the sloping shelf that is above water is flattened, and a radio-labeled pesticide is applied to seedlings of the sort planted therein. A pesticide and its metabolites enter the water phase from feces of insect larvae on plants or leaf frass. In contrast, the aquatic model directly simulates runoff events, because the pesticide is applied directly to the bottom sediment. The recirculation system adds the effect of water motion to the aquatic model ecosystem. Some researchers have applied pesticides in a continuous-flow system to examine the effect of pesticide adsorption to bottom sediment and the long-term effects on organisms to simulate worst conditions (Virtanen and Hattula 1982; Yockim et al. 1980).

Bioconcentration, Bioaccumulation, and Metabolism

93

Table 14 A listing of pesticides and some related chemicals that have been tested in model ecosystem studies No. Pesticide/structure Organochlorines 21 Aldrin 22 Dieldrin

24 25 26 28 29 30 31 33

Chlordane Endrin Heptachlor Toxaphene Mirex Lindane Hexachlorobenzene DDT

34/35 DDE and DDD 37 Methoxychlor – Methoxychlor analogs – Ethoxyanilineb – 2,5,2 ,4 ,5 -Cl5 BPb – TCDDb Organophosphates 39 Parathion 41 Fenitrothion 46 Temephos 47 Chlorpyrifos 54 Terbufos 57 Malathion 60 Phosalone 68 Acephate 69 Fonofos 70 EPN 71 Leptophos Acids and esters 75 2,4-D 76 2,4,5-T 84 Lambda-cyhalothrin 85 Fenvalerate (2S-isomer) Carbamates 99 Mexacarbate 100 Propoxur 101 Carbaryl

102 Carbofuran

Typea /references T: Metcalf (1976); Metcalf et al. (1973b) T: Metcalf et al. (1973b); Sanborn and Yu (1973); F: Rose and McIntire (1970) T: Lu et al. (1975); Sanborn et al. (1976) T: Metcalf (1976); Metcalf et al. (1973b) T: Lu et al. (1975) T: Sanborn et al. (1976); R: Isensee et al. (1979a) T: Metcalf et al. (1973b); A: Isensee (1976) T: Metcalf et al. (1973b); A: Matsumura (1977) T: Metcalf et al. (1973b); A: Isensee et al. (1976) T: Metcalf et al. (1971); Tsuge et al. (1976) A: Bajet and Navarro (2002); Kale et al. (1999a); Matsumura (1977), Wandiga et al. (2002); R: Miyamoto et al. (1979); Ohkawa et al. (1980) T: Metcalf et al. (1971); A: Bajet and Navarro (2002) T: Kapoor et al. (1970); Metcalf et al. (1971) T: Kapoor et al. (1970, 1972, 1973); Coats et al. (1974) T: Hirwe et al. (1972) T: Metcalf (1976) R: Yockim et al. (1978) T: Francis et al. (1980); Yu and Sanborn (1975) T: Metcalf and Sanborn (1975); A: Weinberger et al. (1982), A : Fisher (1985); R: Miyamoto et al. (1979) T: Metcalf and Sanborn (1975) T: Metcalf (1976); A: Kale et al. (1999b) T: Metcalf and Sanborn (1975) T: Metcalf and Sanborn (1975) R: Ambrosi et al. (1978) T: Metcalf and Sanborn (1975) T: Metcalf and Sanborn (1975); F/A: Huckins et al. (1986) T: Francis et al. (1980) T: Metcalf and Sanborn (1975); Francis et al. (1980) T: Metcalf and Sanborn (1975) R: Yockim et al. (1978) A : Hand et al. (2001) R: Ohkawa et al. (1980) A: Matsumura (1977) T: Metcalf (1976); Metcalf and Sanborn (1975) A: Kanazawa et al. (1975); Tsuge et al. (1976); A : Fisher and Lohner (1986) T: Metcalf and Sanborn (1975) T: Yu et al. (1974b); Metcalf and Sanborn (1975)

94

T. Katagi Table 14 (continued)

No. Pesticide/structure

Typea /references

103 XMCb 104 Bufencarb 106 Aldicarb 109 Thiobencarb 111 Phenmedipham Amides and ureas 91 Propachlor 92 Alachlor 96 Naproanilide 120 Diflubenzuron Miscellaneous – Benzene derivatives – 2,4,6-Cl3 -phenol 7 Benzo[a]pyrene 12 Benzidine 14 Di-2-ethylhexyl phthalate 18 Cacodylic acid 79 Dicamba 87 Methoprene 97 Captan 127 Atrazine

A: Kanazawa et al. (1975); Kazano et al. (1976) T: Yu et al. (1974a) T: Metcalf and Sanborn (1975); A : Suora and Fisher (1986) P: Chen et al. (1982) T: Metcalf and Sanborn (1975)

– N-Nitroso-atrazine 129 Cyanazine – Diphenyl ether herbicides 137 Bentazone 139 Oxadiazon – Dinitroaniline herbicides 142 Trifluralin 143 146 147 148 150 a T,

Pendimethalin Metribuzin Pyrazon TFM Diquat

T: Yu et al. (1975c) T: Yu et al. (1975c) P: Wang et al. (1992a) T: Metcalf et al. (1975) A : Lu and Metcalf (1975) F: Virtanen and Hattula (1982) T,A: Lu et al. (1977) T,A: Lu et al. (1977) T: Metcalf (1976); Metcalf et al. (1973a) A: Schuth et al. (1974) T: Yu et al. (1975a) T: Metcalf and Sanborn (1975) T: Metcalf and Sanborn (1975) T: Metcalf and Sanborn (1975); F/A: Huckins et al. (1986); Lynch et al. (1982) A: Kearney et al. (1977b) T: Yu et al. (1975d) P: Lee et al. (1976) T: Booth et al. (1973) R: Ambrosi et al. (1978) A: Kearney et al. (1977a) R: Isensee et al. (1979b); F/A: Huckins et al. (1986); Yockim et al. (1980) A: Isensee and Durbey (1983) T: Metcalf and Sanborn (1975) T: Yu et al. (1975b) F: Maki and Johnson (1977) A : Shaw and Hopke (1975)

terrestrial-aquatic model ecosystem developed by Metcalf et al., in which the chemical was applied to a planted seedling; A, aquatic model ecosystem developed by Kearney et al., in which the chemical was applied to the bottom sediment (or soil) and; A , A wherein application was made to water in the presence or absence of sediment; R, recirculating static model ecosystem developed by Isensee et al.; F, flow-through system; P, paddy model. b Abbreviations: Ethoxyaniline, N-(α-trichloromethyl-4-ethoxybenzyl)-4-ethoxyaniline; TCDD, 2,3,7,8-Tetrachlorodibenzo-p-dioxins; TFM, 3-trifluoromethyl-4-nitrophenol. XMC, 3,5-dimethylphenyl N-methylcarbamate.

Bioconcentration, Bioaccumulation, and Metabolism

95

Using an aquatic model ecosystem in the presence of sand, Matusmura (1977) scrutinized critical experimental factors that affected the accumulation of pesticides in organisms. Based on the distribution of pesticides in the system, both a larger relative ratio of biomass to medium and a very shallow medium were found to cause significant effects on bioconcentration. The effect of medium pH was examined on carbaryl (101) and aldicarb (106) in model ecosystems (Fisher and Lohner 1986; Suora and Fisher 1986). The abiotic degradation profiles were affected by pH, but the distribution of the pesticides and their metabolites was not significantly changed for these carbamates. Because such models are usually open systems, poor material balance is a common problem; such losses result from volatilization or azeotropic distillation (Kanazawa et al. 1975; Yu et al. 1974b). Although closed systems equipped with volatile traps have been used to improve the recovery of 14 C, there was a failure to demonstrate the advantage of the modified system either because of insufficient volatile trapping or because of inadequate bottom sediment analyses (Kazano et al. 1976; Tsuge et al. 1976). Most model ecosystem studies have utilized single applications, although Miyamoto et al. (1979) examined the effect of multiple applications of fenitrothion (41) in a recirculating system; these authors reported no increase in bioaccumulation as compared with a single application. Although model systems may allow collection of valuable information on the interactions among several species, linked by a food web, obtaining time-dependent distribution data for tested pesticides and their metabolites is difficult. A main reason for this difficulty stems from sampling during the study, which deteriorates the balance of the system. Model ecosystems are more complex than single species studies and show a considerably larger variance of data. Using statistical analysis, Isensee (1976) found that response data from several aquatic species in the aquatic model ecosystem studies were one half to one fourth as variable as data coming from microcosm population responses. A coefficient of variance for BCF values was 30–40%; replication was therefore highly recommended. In model ecosystems, two other indices were introduced to assess ecological magnification (EM) and biodegradability (BI) of a chemical in aquatic organisms. The EM index is defined as the concentration ratio of the chemical between organism and water. The EM index is considered to be a bioconcentration factor in non-equilibrium systems and has been demonstrated in fish to inversely correlate with either water solubility of a pesticide or percentage of unextractable fractions in the organism (Lu and Metcalf 1975; Metcalf et al. 1973b; Metcalf and Sanborn 1975). The correlation of log EM with log Kow in fish (Lu and Metcalf 1975), based on the former observations, is in accordance with bioconcentration profiles of the tested compounds under equilibrium conditions. However, either the negative correlation between lipid contents of several species and EM or the positive correlation between the percent of the remaining compound and Hammett σ indicates that metabolism is more important than passive partitioning. The other index, BI, is defined by the concentration ratio of polar products toward non-polar ones in organisms. The index was found to correlate with water solubility for DDT-like pesticides (Kooper et al. 1973). The metabolic profiles in each species introduced into model ecosystems are presented in Tables 10–13. These profiles appear to agree with those

96

T. Katagi

observed in single species studies, but may be less definitive due not only to possible reabsorption of metabolites produced by other species but also to the contribution of microbial and abiotic degradation, especially in a water-sediment system.

5.2 Microcosms and Mesocosms There are many limitations in assessing the distribution in and biological effects of pesticides in natural water bodies. Such limitations have resulted in the introduction of larger-scale systems such as microcosms and mesocosms that are better designed to cope with experimental limitations (Caquet et al. 2000; Møhlenberg et al. 2001). Field studies, conducted in a limited area, are known to provide a better understanding of a xenobiotic’s abiotic dissipation (i.e., photolysis and volatilization) than more simple studies do. However, such field studies, with pesticides and their metabolites, result in complex data from interactions among many aquatic species, and therefore require evaluation with sophisticated statistical methods. Because concentrations of pesticides and their metabolites change with time and dissipation profiles are highly site- and season-specific (Arts et al. 1995; Getsinger et al. 2000; Merlin et al. 2002; Muir et al. 1991, 1992), normal conceptions of bioconcentration at equilibrium cannot be used. Field-derived BCF values vary with time and are often different from those obtained under laboratory conditions (Day 1990; Murakami et al. 1990; Ohyama et al. 1987; Uno et al. 1997). Rather than using the usual BCF approach, investigation of the biological effects of pesticide residues in the body is considered to be more important (DiToro et al. 1991; Lotufo et al. 2001a). Therefore, many field microcosm and mesocosm studies are focused on assessing the biological effects of studied xenobiotics, with less emphasis on evaluating the distribution and metabolism of tested substances (Katagi 2006). Furthermore, obtaining sufficient numbers of biological samples for analysis of metabolites is often difficult because of the smaller body mass and rather low residue levels that exist in many aquatic species. An outdoor pond microcosm study with isoproturon (118) showed variable half-lives in the water column; differences depended on the coverage of the water surface by aquatic macrophytes, wherein much higher residues were detected (Merlin et al. 2002). In this experiment, residues of (118) in sediment and molluscs also showed large variability. A similar effect on distribution was observed for 14 C-diflubenzuron (120) that had been applied to littoral enclosures. Results showed a lower total 14 C mass balance, (53–82%) even after 3 hr (Knuth and Heinis 1995). Rapid dissipation in water, followed by a gradual decrease with a half-life of 2–3 weeks, concomitant with delayed accumulation in sediment, was reported for lindane (30) after it was sprayed on the surface of pond mesocosms (Caquet et al. 1992). Lindane residues in snails and aquatic macrophytes finally declined to levels below the detection limit after 10 weeks. The maximum apparent BCF values for (30) were calculated to be 230 and 56, respectively, for these two species. In the same study, the maximum apparent BCF values for deltamethrin (83) were estimated to be 400 and 1460 in snails and macrophytes, respectively. These values appear to be in the range of laboratoryderived ones, but are inadequate for use in assessing the possible bioconcentration because of concentration changes of the pesticide that occurred in the medium.

Bioconcentration, Bioaccumulation, and Metabolism

97

The duration of sunlight exposure is an indirect factor that controls the dissipation rate of pesticides. Alachlor (92), applied to pond microcosms, was found to more slowly dissipate under reduced sunlight exposure. The lower dissipation was explained by reduced growth of bacterial communities that were capable of co-metabolizing (92) via GST conjugation (Ensz et al. 2003). Both temperature and pH may affect the dissipation of a pesticide in the water column. The effect of temperature and pH has been demonstrated in a ditch mesocosm study conducted on 14 C-linuorn (114) (Crum et al. 1998). Higher temperature and pH caused more rapid dissipation of (114) but did not affect the distribution of 14 C, where only a very minor fraction was accumulated in macrophytes. Only limited information on metabolic degradation of pesticides in mesocosm studies is available (Table 11). A large-scale indoor microcosm study performed on lambda-cyhalothrin (84) has shown formation of the corresponding chrysanthemic acid derivative, which was mainly distributed in the water column (Hand et al. 2001). Abiotic hydrolysis and metabolic transformation in macrophytes, or aufwuchs in the system were proposed as possible mechanisms; the low 14 C recovery was thought to be possibly derived from mineralization processes. By monitoring stream water after an aerial application of fenitrothion (41), Moody et al. (1978) detected an amino derivative of the parent compound, which may have been produced by bacterial reduction. Also found were trace amounts of an S-isomer produced via a sunlight-induced thiono-thiolo rearrangement. Residues in some macrophytes of (41) quickly dissipated within 3 days. Reabsorption of a triclopyr (77) metabolite from the water column was reported by Getsinger et al. (2000) in a field monitoring test, which (77) was applied to lake water. The highest residues of (77) were detected in macrophytes; also detected was the corresponding phenol. However, the 3,6-dihydroxy derivative, the typical photoproduct of (77), was not detected in any biological samples. The methoxy derivative of the phenol, unlikely to be found in aquatic organisms, was significantly accumulated in clams and crayfish. The reason given for this was that the metabolite was microbially produced and reabsorbed from water. Sundaram (1995) identified N-demethylated metabolites and phenol, in aquatic macrophytes, through field monitoring after aerial application of mexacarbate (99) to forests.

6 Summary The ecotoxicological assessment of pesticide effects in the aquatic environment should normally be based on a deep knowledge of not only the concentration of pesticides and metabolites found but also on the influence of key abiotic and biotic processes that effect rates of dissipation. Although the bioconcentration and bioaccumulation potentials of pesticides in aquatic organisms are conveniently estimated from their hydrophobicity (represented by log Kow ), it is still indispensable to factor in the effects of key abiotic and biotic processes on such pesticides to gain a more precise understanding of how they may behave in the natural environment. Relying only on pesticide hydrophobicity may produce an erroneous environmental impact assessment. Several factors affect rates of pesticide dissipation and accumulation in the aquatic environment. Such factors include the amount and type of sediment

98

T. Katagi

present in the water and type of diet available to water-dwelling organisms. The particular physiological behavior profiles of aquatic organisms in water, such as capacity for uptake, metabolism, and elimination, are also compelling factors, as is the chemistry of the water. When evaluating pesticide uptake and bioconcentration processes, it is important to know the amount and nature of bottom sediments present and the propensity that the studied aquatic organisms have to absorb and process xenobiotics. Extremely hydrophobic pesticides such as the organochlorines and pyrethroids are susceptible to adsorb strongly to dissolved organic matter associated with bottom sediment. Such absorption reduces the bioavailable fraction of pesticide dissolved in the water column and reduces the probable ecotoxicological impact on aquatic organisms living in water. In contrast, sediment dwellers may suffer from higher levels of direct exposure to a pesticide, unless it is rapidly degraded in sediment. Metabolism is important to bioconcentration and bioaccumulation processes, as is detoxification and bioactivation. Hydrophobic pesticides that are expected to be highly stored in tissues would not be bioconcentrated if susceptible to biotic transformation by aquatic organisms to more hydrophilic metabolites. In mammalian toxicology, those xenobiotics that are more rapidly metabolized to hydrophilic entities are generally less toxic. By analogy, pesticides that are metabolized to similar entities by aquatic species surely are less ecotoxicologically significant. One feature of fish and other aquatic species that makes them more relevant as targets of environmental studies and of regulation is that they may not only become contaminated by pesticides or other chemicals, but that they constitute an important part of the human diet In this chapter, we provide an overview of the enzymes that are capable of metabolizing or otherwise assisting in the removal of xenobiotics from aquatic species. Many studies have been performed on the enzymes that are responsible for metabolizing xenobiotics. In addition to the use of conventional biochemical methods, such studies on enzymes are increasingly being conducted using immunochemical methods and amino acid or gene sequences analysis. Such studies have been performed in algae, in some aquatic macrophytes, and in bivalva, but less information is available for other aquatic species such as crustacea, annelids, aquatic insecta, and other species. Although their catabolizing activity is often lower than in mammals, oxidases, especially cytochrome P450 enzymes, play a central role in transforming pesticides in aquatic organisms. Primary metabolites, formed from such initial enzymatic action, are further conjugated with natural components such as carbohydrates, and this aids removal from the organisms. The pesticides that are susceptible to abiotic hydrolysis are generally also biotically degraded by various esterases to form hydrophilic conjugates. Reductive transformation is the main metabolic pathway for organochlorine pesticides, but less information on reductive enzymology processes is available. The information on aquatic species, other than fish, that pertains to bioconcentration factors, metabolism, and elimination is rather limited in the literature. The kinds of basic information that is unavailable but is needed on important aquatic species includes biochemistry, physiology, position in food web, habitat, life cycle,

Bioconcentration, Bioaccumulation, and Metabolism

99

etc. Such information is very important to obtaining improved ecotoxicology risk assessments for many pesticides and other chemicals. More research attention on the behavior of pesticides in, and affect on many standard aquatic test species (e.g., daphnids, chironomids, oligochaetes and some bivalves) would particularly be welcome. In addition to improving ecotoxicology risk assessments on target species, such information would also assist in better delineating affects on species at higher trophic levels that are predaceous on the target species. There is also need for designing and employing more realistic approaches to measure bioconcentration and bioaccumulation, and ecotoxicology effects of pesticides in natural environment. The currently employed steady-state laboratory exposure studies are insufficient to deal with the complexity of parameters that control the behavior or concentration of pesticide components extant in the environment. In contrast to the abiotic processes of pesticide investigated under the strictly controlled conditions, each process is significantly affected in the natural environment not only by the site-specific chemistry of water and sediment but also by climate. From this viewpoint, ecotoxicological assessment should be conducted, together with the detailed analyses of abiotic processes, when higher-tier mesocosm studies are performed. Moreover, in-depth investigation is needed to better understand the relationship between pesticide residues in organisms and associated ecotoxicological endpoints. The usual exposure assessment is based on apparent (nominal) concentrations of pesticides, and the residues of pesticides or their metabolites in the organisms are not considered in to the context of ecotoxicological endpoints. Therefore, more metabolic and tissue distribution information for terminal pesticide residues is needed for aquatic species both in laboratory settings and in higher-tier (microcosm, mesocosm) studies.

Appendix 1: Chemical Structures of Organic Chemicals and PAHs (Polyaromatic Hydrocarbons)

100

T. Katagi

Appendix 2: Chemical Structures of Organochlorine Pesticides and Related Compounds

Bioconcentration, Bioaccumulation, and Metabolism

Appendix 3: Chemical Structures of Organophosphorus Pesticides

101

102

T. Katagi

Appendix 4: Chemical Structures of Acid, Ester, and Amide Pesticides and Related Compounds

Appendix 5: Chemical Structures of Carbamate and Urea Pesticides and Related Compounds

Bioconcentration, Bioaccumulation, and Metabolism

103

Appendix 6: Chemical Structures of Miscellaneous Pesticides

References Abdalla AM, El-Mogy M, Farid NM, El-Sharabasy M (2006) Two glutathione S-transferase isoenzymes purified from Bulinus truncates (Gastropoda: Planorbidae). Comp Biochem Physiol 143B: 76–84. Aceto A, Di Ilio C, Bucciarelli T, Pantani C, Dell’Agata M, Pannunzio G, Federiu G (1991) Characterization of glutathione transferase from Gammarus Italicus. Comp Biochem Physiol 99B: 523–527. Ahlgren G, Lundstedt L, Brett M, Forsberg C (1990) Lipid composition and food quality of some freshwater phytoplankton for cladoceran zooplankters. J Plankton Res 12: 809–818. Akkanen J, Kukkonen JVK (2003) Biotransformation and bioconcentration of pyrene in Daphnia magna. Aquat Toxicol 64: 53–61. Aly OA, Shehata SA, Farag H (1984) Uptake and accumulation of selected herbicides by the fresh water alga Scenedesmus. Arch Environ Contam Toxicol 13: 701–705. Ambrosi D, Isensee AR, Macchia JA (1978) Distribution of oxadiazon and phosalone in an aquatic model ecosystem. J Agric Food Chem 26: 50–53. Amweg EL, Weston DP, Johnason CS, You J, Lydy MJ (2006) Effect of piperonyl butoxide on permethrin toxicity in the amphipod Hyalella azteca. Environ Toxicol Chem 25: 1817–1825. Anderson AC, Abdelghani AA, McDonnell D, Craig L (1981) Uptake of monosodium methanearsonate (MSMA) by vascular aquatic plants. J Plant Nutr 3: 193–201. Anderson RL (1982) Toxicity of fenvalerate and permethrin to several nontarget aquatic invertebrates. Environ Entmol 11: 1251–1257. Anderson RL, DeFoe DL (1980) Toxicity and bioaccumulation of endrin and methoxychlor in aquatic invertebrates and fish. Environ Pollut 22A: 111–121. Anderson RS (1985) Metabolism of a model environmental carcinogen by bivalve molluscs. Mar Environ Res 17: 137–140.

104

T. Katagi

Anderson TD, Zhu KY (2004) Synergistic and antagonistic effects of atrazine on the toxicity of organophosphorodithioate and organophosphorothioate insecticides to Chironomus tentans (Diptera: Chironomidae). Pestic Biochem Physiol 80: 54–64. Ankley GT, Call DJ, Cox JS, Kahl MD, Hoke RA, Kosian PA (1994) Organic carbon partitioning as a basis for predicting the toxicity of chlorpyrifos in sediments. Environ Toxicol Chem 13: 621–626. Ankley GT, Collyard SA (1995) Influence of piperonyl butoxide on the toxicity of organophosphate insecticides to three species of freshwater benthic invertebrates. Comp Biochem Physiol 110C: 149–155. Artola-Garicano E, Sinnige TL, van Holstein I, Vaes WHJ, Hermens JLM (2003) Bioconcentration and acute toxicity of polycyclic musks in two benthic organisms (Chironomus riparius and Lumbriculus variegatus). Environ Toxicol Chem 22: 1086–1092. Arts M, Ferguson ME, Glozier NE, Robarts RD, Donald DB (1995) Spatial and temporal variability in lipid dynamics of common amphipods: Assessing the potential for uptake of lipophilic contaminants. Ecotoxicol 4: 91–113. Arun S, Rajendran A, Subramanian P (2006) Subcellular/tissue distribution and responses to oil exposure of the cytochrome P450-dependent monooxygenase system and glutathione S-transferase in freshwater prawns (Macrobrachium malcolmsonii, M. lamarrei lamarrei). Ecotoxicology 15: 341–346. Ashauer R, Boxall A, Brown C (2006) Uptake and elimination of chlorpyrifos and pentachlorophenol into the freshwater amphipod Gammarus pulex. Arch Environ Contam Toxicol 51: 542–548. Axelman J, Broman D, Näf C, Pettersen H (1995) Compound dependence of the relationship log Kow and log BCFL . Environ Sci Pollut Res 2: 33–36. Baeza-Squiban A, Bouaicha N, Santa-Maria A, Marano F (1990) Demonstration of the excretion by Dunaliella bioculata of esterases implicated in the metabolism of deltamethrin, a pyrethroid insecticide. Bull Environ Contam Toxicol 45: 39–45. Baeza-Squiban A, Meinard C, Marano F (1988) Metabolism of deltamethrin in two cell types in vitro. Pestic Biochem Physiol 32: 253–261. Bajet CM, Navarro MP (2002) Degradation, release and bioavailability of 14 C DDT and 14 C DDE sediment residues to oysters and mussels. Environ Technol 23: 1293–1302. Balasubramanya RH, Patil RB (1980) Degradation of carboxin and oxycarboxin by microorganisms. Plant Soil 57: 457–462. Baldwin WS, LeBlanc GA (1994a) In vivo biotransformation of testosterone by phase I and II detoxication enzymes and their modulation by 20-hydroxy-ecdysone in Daphnia magna. Aquat Toxicol 29: 103–117. Baldwin WS, LeBlanc GA (1994b) Identification of multiple steroid hydroxylases in Daphnia magna and their modulation by xenobiotics. Environ Toxicol Chem 13: 1013–1021. Baldwin WS, LeBlanc GA (1996) Expression and induction of an immunochemically related class of glutathione S-transferases in Daphnia magna. Comp Biochem Physiol 113B: 261–267. Baldwin WS, Milam DL, LeBlanc GA (1995) Physiological and biochemical perturbations in Daphnia magna following exposure to the model environmental estrogen diethylstilbestrol. Environ Toxicol Chem 14: 945–952. Barata C, Solayan A, Porte C (2004) Role of B-esterases in assessing toxicity of organophosphorus (chlorpyrifos, malathion) and carbamate (carbofuran) pesticides to Daphnia magna. Aquat Toxicol 66: 125–139. Barber JT, Sharma HA, Ensley HE, Polito MA, Thomas DA (1995) Detoxification of phenol by the aquatic angiosperm, Lemna gibba. Chemosphere 31: 3567–3574. Barbier M, Prevot P, Soyer-Gobillard MO (2000) Esterases in marine dinoflagellates and resistance to the organophosphate insecticide parathion. Int Microbiol 3: 117–123. Barron MG (1990) Bioconcentration (will water-borne organic chemicals accumulate in aquatic animals?). Environ Sci Technol 24: 1612–1618.

Bioconcentration, Bioaccumulation, and Metabolism

105

Barron MG, Hansen SC, Ball T (1991) Pharmacokinetics and metabolism of triclopyr in the crayfish (Procambarus clarkii). Drug Metab Disp 19: 163–167. Bartlett AJ, Borgmann U, Dixon DG, Batcher SP, Maguire RJ (2004) Tributyltin uptake and depuration in Hyalella azteca: Implications for experimental design. Environ Toxicol Chem 23: 426–434. Barton JW, Kuritz T, O’Connor LE, Ma CY, Maskarinec MP, Davison BH (2004) Reductive transformation of methyl parathion by the cyanobacterium Anabaena sp. strain PCC7120. Appl Microbiol Biotechnol 65: 330–335. Basack SB, Oneto ML, Fucks JS, Wood EJ, Kesten EM (1998) Esterases of Corbicula fluminea as biomarkers of exposure to organophosphorus pesticides. Bull Environ Contam Toxicol 61: 569–576. Basack SB, Oneto ML, Verrengia Guerrero NR, Kesten EM (1997) Accumulation and elimination of pentachlorophenol in the freshwater bivalve Corbicula fluminea. Bull Environ Contam Toxicol 58: 498–503. Baturo W, Lagadic L (1996) Benzo[a]pyrene hydroxylase and glutathione s-transferase activities as biomarkers in Lymnaea palustris (mollusca, gastropoda) exposed to atrazine and hexachlorobenzene in freshwater mesocosms. Environ Toxicol Chem 15: 771–781. Bauer I, Weigelt S, Ernst W (1989) Biotransformation of hexachlorobenzene in the blue mussel (Mytilus edulis). Chemosphere 19: 1701–1707. Bedford JW, Zabik MJ (1973) Bioactive compounds in the aquatic environment: Uptake and loss of DDT and dieldrin by freshwater mussels. Arch Environ Contam Toxicol 1: 97–111. Bejarano AC, Widenfalk A, Decho AW, Chandler GT (2003) Bioavailability of the organophosphorus insecticide chlorpyrifos to the suspension-feeding bivalve, Mercenaria mercenaria, following exposure to dissolved and particulate matter. Environ Toxicol Chem 22: 2100–2105. Belden JB, Lydy MJ (2000) Impact of atrazine on organophosphate insecticide toxicity. Environ Toxicol Chem 19: 2266–2274. Belden JB, Ownby DR, Lotufo GR, Lydy MJ (2005) Accumulation of trinitrotoluene (TNT) in aquatic organisms: Part 2 – Bioconcentration in aquatic invertebrates and potential for trophic transfer to channel catfish (Ictalurus punctatus). Chemosphere 58: 1161–1168. Benezet HJ, Knowles CO (1981) Degradation of chlordimeform by algae. Chemosphere 10: 909–917. Benson AA, Shibuya I (1962) Surfactant lipids. In: Physiology and Biochemistry of Algae. Lewin RA (ed) Academic Press, New York, Chapter 22, pp 371–383. Ben-Tal Y, Cleland CF (1982) Uptake and metabolism of [14 C]salicylic acid in Lemna gibba G3. Plant Physiol 70: 291–296. Bhadra R, Spanggord RJ, Wayment DG, Hughes JB, Shanks JV (1999) Characterization of oxidation products of TNT metabolism in aquatic phytoremediation systems of Myriophyllum aquaticum. Environ Sci Technol 33: 3354–3361. Birmelin C, Escartin E, Goldfarb PS, Livingstone DR, Porte C (1998) Enzyme effects and metabolism of fenitrothion in primary cell culture of the red swamp crayfish Procambarus clarkii. Mar Environ Res 46: 375–378. Bligh EG, Dyer WJ (1959) A rapid method of total lipid extraction and purification. Can J Biochem Physiol 37: 911–917. Böhm HH, Müller H (1976) Model studies on the accumulation of herbicides by microalgae. Naturwissenschaften 63: 296. Booij K, van den Berg C (1994) Comparison of techniques for the extraction of lipids and PCBs from benthic invertebrates. Bull Environ Contam Toxicol 53: 71–76. Booth GM, Yu CC, Hansen DJ (1973) Fate, metabolism, and toxicity of 3-isopropyl-1H-2,1,3benzothiadiazin-4(3H)-1,2,2-dioxide in a model ecosystem. J Environ Qual 2: 408–411. Borchert J, Karbe L, Westendorf J (1997) Uptake and metabolism of benzo[a]pyrene absorbed to sediment by the freshwater invertebrate species Chironomus riparius and Sphaerium corneum. Bull Environ Contam Toxicol 58: 158–165.

106

T. Katagi

Borowitzka MA (1988) Fats, oils and hydrocarbons. In: Microalgal Biotechnology. Borowitzka MA (ed) Cambridge University Press, Cambridge, Chapter 10, pp 257–287. Boryslawskyj M, Garrood T, Stanger M, Pearson T, Woodhead DW (1988) Role of lipid/water partitioning and membrane composition in the uptake of organochlorine pesticides into a freshwater mussel. Mar Environ Res 24: 57–61. Böttcher T, Schroll R (2007) The fate of isoproturon in a freshwater microcosm with Lemna minor as a model organism. Chemosphere 66: 684–689. Boutet I, Tanguy A, Moraga D (2004) Molecular identification and expression of two non-P450 enzymes, monoamine oxidase A and flavin-containing monooxygenase 2 involved in phase I of xenobiotic biotransformation in the Pacific oyster, Crasspstrea gigas. Biochim Biophys Acta 1679: 29–36. Bowes GW (1972) Uptake and metabolism of 2,2 -bis-(p-chlorophenyl)-1,1,1-trichloroethane (DDT) by marine phytoplankton and its effect on growth and chloroplast electron transport. Plant Physiol 49: 172–176. Bowman BP, Snell TW, Cochrane BJ (1990) Isolation and purification of glutathione-S-transferases from Brachionus plicatilis and B. calyciflorus (Rotifera). Comp Biochem Physiol 95B: 619–624. Brandes D, Elston RN (1956) An electron microscopical study of the histochemical localization of alkaline phosphatase in the cell wall of Chlorella vulgaris. Nature 177: 274–275. Briand J, Julistiono H, Beaune P, Flinois JP, deWaziers I, Leroux JP (1993) Presence of proteins recognized by mammalian cytochrome P450 antibodies in Euglena gracilis. Biochim Biophys Acta 1203: 199–204. Brodtmann Jr NV (1970) Studies on the assimilation of 1,1,1-trichloro-2,2-bis(pchlorophenyl)ethane (DDT) by Crassostrea virginica Gmelin. Bull Environ Contam Toxicol 5: 455–462. Brook LT, Ankley GT, Call DJ, Cook PM (1996) Gut content weight and clearance rate for three species of freshwater invertebrates. Environ Toxicol Chem 15: 223–228. Bruno MG, Fannin TE, Leversee GJ (1981) The disposition of benzo[a]pyrene in the periphyton communities of two South Carolina streams: Uptake and biotransformation. Can J Bot 60: 2084–2091. Buchwalter DB, Sandahl JF, Jenkins JJ, Curtis LR (2004) Role of uptake, biotransformation, and target site sensitivity in determining the differential toxicity of chlorpyrifos to second to fourth instar Chironomus riparius (Meigen). Aquat Toxicol 66: 149–157. Burgess RM, McKinney RA (1999) Importance of interstitial water, overlying water and whole sediment exposures to bioaccumulation by marine bivalves. Environ Pollut 104: 373–382. Bychek EA, Gushchina IA (1999) Age-dependent changes of lipid composition in Daphnia magna. Biochemistry (Moscow) 64: 543–545. Cáceres T, Megharaj M, Naidu R (2008) Toxicity and transformation of fenamiphos and its metabolites by two microalgae Pseudikirchneriella subcapitata and Chlorococcum sp. Sci Total Environ 398: 53–59. Cai X, Liu W, Jin M, Lin K (2007) Relation of doclofop-methyl toxicity and degradation in algae cultures. Environ Toxicol Chem 26: 970–975. Cai X, Liu W, Sheng G (2008) Enantioselective degradation and ecotoxicity of the chiral herbicide diclofop in three freshwater algae cultures. J Agric Food Chem 56: 2139–2146. Canton JH, Greve PA, Slooff W, van Esch GJ (1975) Toxicity, accumulation and elimination studies of α-hexachlorocyclohexane (α-HCH) with freshwater organisms of different trophic levels. Water Res 9: 1163–1169. Canton JH, van Esch GJ, Greve PA, van Hellemond ABAM (1977) Accumulation and elimination of α-hexachlorocyclohexane (α-HCH) by the marine algae Chlamydomonas and Dunaliella. Water Res 11: 111–115. Caquet T, Lagadic L, Sheffield SR (2000) Mesocosms in ecotoxicology (1): Outdoor aquatic systems. Rev Environ Contam Toxicol 165:1–38.

Bioconcentration, Bioaccumulation, and Metabolism

107

Caquet T, Thybaud E, LeBras S, Jonot O, Ramada F (1992) Fate and biological effects of lindane and deltamethrin in freshwater mesocosms. Aquat Toxicol 23: 261–278. Casserly DM, Davis EM, Downs TD, Guthrie RK (1983) Sorption of organics by Selenastrum capricornutum. Water Res 17: 1591–1594. Cavaletto JF, Gardner WS (1988) Seasonal dynamics of lipids in freshwater benthic invertebrates. In: Lipids in Freshwater Ecosystem. Arts MT, Wainman BC (eds) Springer, New York, Chapter 6, pp 109–131. Cerniglia CE, Freeman JP, van Baalen C (1981) Biotransformation and toxicity of aniline and aniline derivatives in cyanobacteria. Arch Microbiol 130: 272–275. Cerniglia CE, Gibson DT, van Baalen C (1979) Algal oxidation of aromatic hydrocarbon: Formation of 1-naphthol from naphthalene by Agmenellum quadruplicatum, strain PR-6. Biochem Biophys Res Commun 88: 50–58. Cerniglia CE, Gibson DT, van Baalen C (1980a) Oxidation of naphthalene by cyanobacteria and microalgae. J Gen Microbiol 116: 495–500. Cerniglia CE, van Baalen C, Gibson DT (1980b) Oxidation of biphenyl by the cyanobacterium. Oscillatoria sp., straing JCM. Arch Microbiol 125: 203–207. Cerniglia CE, van Baalen C, Gibson DT (1980c) Metabolism of naphthalene by the Oscillatoria sp., strain JCM. J Gen Microbiol 116: 485–494. Chan SMN, Luan T, Wong MH, Tam NFY (2006) Removal and biodegradation of polycyclic aromatic hydrocarbons by Selenastrum capricornutum. Environ Toxicol Chem 25: 1772–1779. Chapman VJ, Chapman DJ (1973) In: The Algae. Macmillan Co., Ltd., London, 2nd Ed. Chaton PF, Ravanel P, Tissut M, Meyran JC (2002) Toxicity and bioaccumulation of fipronil in the nontarget arthropodan fauna associated with Subalpine mosquito breeding sites. Ecotoxicol Environ Safety 52: 8–12. Chaty S, Rodius F, Vasseur P (2004) A comparative study of the expression of CYP1A and CYP4 genes in aquatic invertebrate (freshwater mussel, Unio tumidus) and vertebrate (rainbow trout, Oncorhynchus mykiss). Aquat Toxicol 69: 81–93. Chen SJ, Hsu EL, Chen YL (1982) Fate of herbicide benthiocarb (thiobencarb) in a rice paddy model ecosystem. J Pestic Sci 7: 335–340. Chessells M, Hawker DW, Connell DW (1992) Influence of solubility in lipid on bioconcentration of hydrophobic compounds. Ecotoxicol Environ Safety 23: 260–273. Choi KJ, Nakhost Z, Barzana E, Karel M (1987) Lipid content and fatty acid composition of green algae Scenedesmus obliquus grown in a constant cell density apparatus. Food Biotechnol 1: 117–128. Christensen M, Andersen O, Banta GT (2002) Metabolism of pyrene by the polychaetes Nereis diversicolor and Arenicola marina. Aquat Toxicol 58: 15–25. Ciarelli S, Kater BJ, van Straalen NM (2000) Influence of bioturbation by the amphipod Corophium volutator on fluoranthene uptake in the marine polychaete Nereis Virens. Environ Toxicol Chem 19: 1575–1581. Cid Montañés JF, van Hattum B (1995) Bioconcentration of chlorpyrifos by the freshwater isopod Asellus aquaticus (L.) in outdoor experimental ditches. Environ Pollut 88: 137–146. Clark AG (1989) The comparative enzymology of the glutathione S-transferases from nonvertebrate organisms. Comp Biochem Physiol 92B: 419–446. Coats JR, Metcalf RL, Kapoor IP (1974) Metabolism of the methoxychlor isostere, dianisylneopentene, in mouse, insects, and a model ecosystem. Pestic Biochem Physiol 4: 201–211. Conder JM, LaPoint TW, Bowen AT (2004) Preliminary kinetics and metabolism of 2,4,6trinitrotoluene and its reduced metabolites in an aquatic oligochaete. Aquat Toxicol 69: 199–213. Connell DW (1988) Bioaccumulation behavior of persistent organic chemicals with aquatic organisms. Rev Environ Contam Toxicol 102: 117–154. Connell DW, Hawker DW (1988) Use of polynomial expressions to describe the bioconcentartion of hydrophobic chemicals in fish. Ecotoxicol Environ Safety 16: 242–257.

108

T. Katagi

Connell DW, Bowman M, Hawker DW (1988) Bioconcentration of chlorinated hydrocarbons from sediment by oligochaetes. Ecotoxicol Environ Safety 16: 293–302. Connor (1984) Fish/sediment concentration ratios for organic compounds. Environ Sci Technol 18: 31–35. Conrad AU, Comber SD, Simkiss K (2000) New method for the assessment of contaminant uptake routes in the oligochaete Lumbriculus variegatus. Bull Environ Contam Toxicol 65: 16–21. Conrad AU, Comber SD, Simkiss K (2002) Pyrene bioavailability; effect of sediment-chemical contact time on routes of uptake in an oligochaete worm. Chemosphere 49: 447–454. Contardo-Jara V, Wiegand C (2008) Biotransformation and antioxidized enzymes of Lumbriculus variegatus as biomarkers of contaminated sediment exposure. Chemosphere 70: 1879–1888. Copeman LA, Parrish CC (2004) Lipids classes, fatty acids, and sterols in seafood from Gilbert Bay, Southern Labrador. J Agric Food Chem 52: 4872–4881. Cowan-Ellsberry CE, Dyer SD, Erhardt S, Bernhard MJ, Roe AL, Dowty ME, Weisbrod AV (2008) Approach for extrapolating in vitro metabolism data to refine bioconcentration factor estimates. Chemosphere 70: 1804–1817. Cowgill UM, Williams DM, Esquivel JB (1984) Effect of maternal nutrition on fat and longevity of neonates of Daphnia magna. J Crustacean Biol 4: 173–190. Crosby DG, Tucker RK (1971) Accumulation of DDT by Daphnia magna. Environ Sci Technol 5: 714–716. Crum SJH, Aalderink GH, Brock TCM (1998) Fate of the herbicide linuron in outdoor experimental ditches. Chemosphere 36:2175–2190. Curto MJ, van Hattum B, Cid JF (1993) In situ bioaccumulation of PAH in freshwater isopods in relation to partitioning between sediments and water. Polycycl Aromat Compd 3: 1031–1038. Damásio J, Guilhermino L, Soares AMVM, Riva MC, Barata C (2007) Biochemical mechanisms of resistance in Daphnia magna exposed to the insecticide fenitrothion. Chemosphere 70: 74–82. Darley WM (1977) Biochemical composition. In: The Biology of Diatoms. Werner D (ed) Botanical Monographs vol 13, Blackwell Scientific Publications, Oxford, Chapter 7, pp 198–223. Dauberschmidt C, Dietrich DR, Schlatter C (1997) Investigation on the biotransformation capacity of organophosphates in the mollusc Dreissena polymorpha P. Aquat Toxicol 37: 283–294. Day JA, Saunders FM (2004) Glycosidation of chlorophenols by Lemna minor. Environ Toxicol Chem 23: 613–620. Day KE (1990) Pesticides residues in freshwater and marine zooplankton: A review. Environ Pollut 67: 205–222. Day KE (1991) Effects of dissolved organic carbon on accumulation and acute toxicity of fenvalerate, deltamethrin and cyhalothrin to Daphnia magna (Straus). Environ Toxicol Chem 10: 91–101. Day KE, Kaushik NK (1987) The adsorption of fenvalerate to laboratory glassware and the alfa Chlamydomonas reinhardtii, and its effect on uptake of the pesticide by Daphnia galeata mendotae. Aquat Toxicol 10: 131–142. De Busk BC, Chimote SS, Rimoldi JM, Schenk D (2000) Effect of the dietary brominated phenol, lanosol, on chemical biotransformation enzymes in the gumboot chiton Cryptochiton stelleri (Middendorf, 1846). Comp Biochem Physiol 127C: 133–142. Del Vento S, Dachs J (2002) Predcition of uptake dynamics of persistent organic pollutants by bacteria and phytoplankton. Environ Toxicol Chem 21: 2099–2107. Della Greca M, Pinto G, Pollio A, Previtera L, Temussi F (2003) Biotransformation of sinapic acid by the green algae Stichococcus bacillaris 155LTAP and Ankistrodesmus braunii C202.7a. Tetrahedron Lett 44: 2779–2780. Della Greca M, Previtera L, Fiorentino A, Pinto G, Pollio A (1996) Bioconversion of 17β-hydroxy17α- methylandrosta-1,4-dien-3-one and androsta-1,4-dien-3,17-dione in cultures of the green alga T76 Scenedesmus quadricauda. Tetrahedron 52: 13981–13990.

Bioconcentration, Bioaccumulation, and Metabolism

109

DeLorenzo ME, Taylor LA, Lund SA, Pennington PL, Strozier ED, Fulton MH (2002) Toxicity and bioconcentration potential of the agricultural pesticide endosulfan in phytoplankton and zooplankton. Arch Environ Contam Toxicol 42: 173–181. Dembitsky VM, Rozentsvet OA, Pechenkina EE (1990) Glycolipids, phospholipids and fatty acids of brown algae species. Phytochem 29: 3417–3421. Dembitsky VM, Rozentsvet OA, Zhuikova VS, Vasilenko RF, Kashin AG (1992) Lipid composition of freshwater macrophytes from the Volga river estuary. Phytochem 31: 3259–3261. Derr SK, Zabik MJ (1972) Biologically active compounds in the aquatic environment: The uptake and distribution of [1,1-dichloro-2,2-bis(p-chlorophenyl)ethylene], DDE by Chironomus tentans Fabricius (Diptera: Chironomidae). Trans Am Fish Soc 323–329. De Wolf W, de Bruijn JHM, Seinen W, Hermens JLM (1992) Influence of biotransformation on the relationship between bioconcentration factors and octanol-water partition coefficients. Environ Sci Technol 26: 1197–1201. Dhanaraj PS, Kumar S, Lal R (1989) Bioconcentration and metabolism of aldrin and phorate by the blue-green algae Anabaena (ARM310) and Aulosira fertilissima (ARM68). Agric Ecosys Environ 25: 187–193. Dierickx PJ (1987) Soluble glutathione transferase isoenzymes in Daphnia magna strauss and their interactions with 2,4-dichlorophenoxyacetic acid and 1,4-benzoquinone. Insect Biochem 17: 1–6. Dimitrov SD, Dimitorova NC, Walker JD, Veith GD, Mekenyan OG (2002) Predicting bioconcentration factors of highly hydrophobic chemicals. Effects of molecular size. Pure Appl Chem 74: 1823–1830. DiToro DM, Zarba CS, Hansen DJ, Berry WJ, Swartz RC, Cowan CE, Pavlou SP, Allen HE, Thomas NA, Paquin PR (1991) Technical basis for establishing sediment quality criteria for nonionic organic chemicals using equilibrium partitioning. Environ Toxicol Chem 10: 1541–1583. Donigian T, Lawrence G, Garland E, Paquin P, DiNitto R, McGrath R, Svirsky S, Campbell S, Clough J (2004) Model calibration: Modeling study of PCB contamination in the Housatonic river. Environmental remediation contract, General Electric (GE)/Housatonic river project, Pittsfield, Massachusetts, Contract No. DACW33-00-D-0006, Task Order No. 0003, DCN: GE-122304-ACMG. Volume 4, Appendix C, Attachment C-4, pp 1–546. Donkin P, Widdows J, Evans SV, Staff FJ, Yan T (1997) Effect of neurotoxic pesticides on the feeding rate of marine mussels (Mytilus edulis). Pestic Sci 49: 196–209. Driscoll SK, McElroy AE (1996) Bioaccumulation and metabolism of benzo[a]pyrene in the three species of polychaete worms. Environ Toxicol Chem 15: 1401–1410. Drouillard KD, Ciborowski JJH, Lazar R, Haffner GD (1996) Estimation of the uptake of organochlorines by the mayfly Hexagenia limbata (Ephemeroptera: Ephemeridae). J Great Lakes Res 22: 26–35. Duncan J, Brown N, Dunlop RW (1977) The uptake of the molluscicide, 4 -chloronicotinanilide into Biomphalaria glabrata (Say) in a glowing water system. Pestic Sci 8: 345–353. Dutton GJ (1965) Conjugation of phenols in molluscs: Determination of glucosyl, not glucuronyl, transfer from uridine diphosphate nucleotides. Biochem J 96: 36–37. Dutton GJ (1966) Uridine diphosphate glucose and the synthesis of phenolic glucosides by molluscs. Arch Biochem Biophys 116: 399–405. Edwards R, Millburn P (1985) The metabolism and toxicity of insecticides in fish. In: Progress in Pesticide Biochemistry and Toxicology, Insecticides. Hutson DH, Roberts TR (eds) John Wiley & Sons, Ltd. New York, vol 5, chap 6, pp 249–274. EFSA (2008) Conclusions of the risk assessment on active substances. Pesticide Risk Assessment Peer Review Unit (PRAPeR), Conclusions, European Food Safety Authority, http://www.efsa.europa.eu /EFSA/ScientificPanels/PRAPER/efsa_locale-1178620753812_ Conclusions494.htm.

110

T. Katagi

Egeler P, Römbke J, Meller M, Knacker T, Franke C, Studinger G, Nagel R (1997) Bioaccumulation of lindane and hexachlorobenzene by turbificid sludgeworms (oligochaeta) under standardized laboratory conditions. Chemosphere 35: 835–852. Ellgehausen H, Guth JA, Esser HO (1980) Factors determining the bioaccumulation potential of pesticides in the individual compartments of aquatic food chains. Ecotox Environ Safety 4: 134–157. Ellis BE (1977) Degradation of phenolic compounds by fresh-water algae. Plant Sci Lett 8: 213–216. Elmamlouk TH, Gessner T (1978) Carbohydrate and sulfate conjugations of p-nitrophenol by hepatopancreas of Homarus Americanus. Comp Biochem Physiol 61C: 363–367. Ensley HE, Barber JT, Polito MA, Oliver AI (1994) Toxicity and metabolism of 2,4-dichlorophenol by the aquatic angiosperm Lemna gibba. Environ Toxicol Chem 13: 325–331. Ensz AP, Knapp CW, Graham DW (2003) Influence of autochthonous dissolved organic carbon and nutrient limitation on alachlor biotransformation in aerobic aquatic systems. Environ Sci Technol 37:4157–4162. Ernst W (1977) Determination of the bioconcentration potential of marine organisms – A steady state approach. I. Bioconcentration data for seven chlorinated pesticides in mussels (Mytilus edulis) and their relation to solubility data. Chemosphere 6: 731–740. Ernst W (1979) Factors affecting the evaluation of chemicals in laboratory experiments using marine organisms. Ecotox Environ Safety 3: 90–98. Escartin E, Porte C (1996) Bioaccumulation, metabolism, and biochemical effects of the organophosphorus pesticide fenitrothion in Procambarus clarkii. Environ Toxicol Chem 15: 915–920. Escartin E, Porte C (1997) The use of cholinesterases and carboxylesterase activities from Mytilus Galloprovincialis in pollution monitoring. Environ Toxicol Chem 16: 2090–2095. Estenik JF, Collins WJ (1979) In vitro and in vivo studies of mixed-function oxidase in an aquatic insect, Chironomus riparius. In: Pesticide and Xenobiotic Metabolism in Aquatic Organisms. Khan MAQ, Lech JJ, Menn JJ (eds) ACS Symp Ser 99, American Chemical Society, Washington, DC, Chapter 21, pp 349–370. EU (2008) Plant Protection Products – New active substances – Decisions and review reports. Health & Consumer Protection Directorate-General, European Commission. http://ec.europa.eu/food/plant/protection/evaluation/new_subs_rep_en.htm . Ewald G, Larsson P (1994) Partitioning of 14 C-labeled 2,2 ,4,4 -tetrachlorobiphenyl between water and fish lipids. Environ Toxicol Chem 13: 1577–1580. Fahl GM, Kreft L, Altenburger R, Faust M, Boedeker W, Grimme LH (1995) pH-Dependent sorption, bioconcentration and algal toxicity of sulfonylurea herbicides. Aquat Toxicol 31: 175–187. Farrington JW (1991) Biogeochemical processes governing exposure and uptake of organic pollutant compounds in aquatic organisms. Environ Health Perspect 90: 75–84. Ferrando MD, Sancho E, Andreu-Moliner E (1996) Accumulation of tetradifon in an algae (Nannochloris oculata) and the cladoceran, Daphnia magna. Bull Environ Contam Toxicol 57: 139–145. Feurtet-Mazel A, Grollier T, Grouselle M, Ribeyre F, Boudou A (1996) Experimental study of bioaccumulation and effects of the herbicide isoproturon on freshwater rooted macrophytes. Chemosphere 32: 1499–1512. Fisher SW (1985) Effects of pH upon the environmental fate of [14 C]fenitrothion in an aquatic microcosm. Ecotoxicol Environ Saf 10: 53–62. Fisher SW, Hwang H, Atanasoff M, Landrum PF (1999) Lethal body residues for pentachlorophenol in Zebra mussels (Dreissena polymorpha) under varying condition of temperature and pH. Ecotoxicol Environ Saf 43: 274–283. Fisher SW, Lohner TW (1986) Studies on the environmental fate of carbaryl as a function of pH. Arch Environ Contam Toxicol 15: 661–667. Fisher T, Craane M, Callaghan A (2003) Induction of cytochrome P-450 activity in individual Chironomus riparius Meigen larvae exposed to xenobiotics. Ecotoxicol Environ Saf 54: 1–6.

Bioconcentration, Bioaccumulation, and Metabolism

111

Fitzpatrick PJ, Sheehan D (1993) Separation of multiple forms of glutathione-S-transferase from the blue mussel, Mytilus edulis. Xenobiotica 23: 851–861. Folch J, Lees M, Stanley HS (1957) A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem 226: 497–509. Forbes VE, Andreassen MSH, Christensen L (2001) Metabolism of the polycyclic aromatic hydrocarbon fluoranthene by the polychaete Capitata species I. Environ Toxicol Chem 20: 1012–1021. Foster GD, Crosby DG (1986) Xenobiotic metabolism of p-nitrophenol derivatives by the rice field crayfish (Procambarus clarkii). Environ Toxicol Chem 5: 1059–1070. Francis BM, Hansen LG, Fukuto TR, Lu PY, Metcalf RL (1980) Ecotoxicology of phenylphosphono- thioates. Environ Health Persp 36: 187–195. Frank PA, Hodgson RH (1964) A technique for studying absorption and translocation in submerged plants. Weeds 12: 80–82. Fujisawa T, Kurosawa M, Katagi T (2006) Uptake and transformation of pesticide metabolites by duckweed (Lemna gibba). J Agric Food Chem 54: 6286–6293. Fukuto TR (1990) Mechanism of action of organophosphorus and carbamate insecticides. Environ Health Persp 87: 245–254. Funderburk Jr HH, Lawrence JM (1963) Absorption and translocation of radioactive herbicides in submersed and emersed aquatic weeds. Weed Res 3: 304–311. Gabric AJ, Connell DW, Bell PF (1990) A kinetic model for bioconcentration of lipophilic compounds by oligochaetes. Water Res 24: 1225–1231. Galloway TS, Millward N, Browne MA, Depledge MH (2002) Rapid assessment of organophosphorus/carbamate exposure in the bivalve, mollusc Mytilus edulis using combined esterase activities as biomarkers. Aquat Toxicol 61: 169–180. Gao J, Garrison AW, Hoehamer C, Mazur CS, Wolfe NL (2000a) Uptake and phytotransformation of organophosphorus pesticides by axenically cultivated aquatic plants. J Agric Food Chem 48: 6114–6120. Gao J, Garrison AW, Hoehamer C, Mazur CS, Wolfe NL (2000b) Uptake and phytotransformation of o,p -DDT and p,p -DDT by axenically cultivated aquatic plants. J Agric Food Chem 48: 6121–6127. Gardner WS, Frez WA, Cichocki EA, Panish CC (1985) Micromethod for lipids in aquatic invertebrates. Limnol Oceanogr 30: 1099–1105. Garrison AW, Nzengung VA, Avants JK, Ellington JJ, Jones WJ, Rennels D, Wolfe NL (2000) Phytodegradation of p,p -DDT and the enantiomers of o,p -DDT. Environ Sci Technol 34: 1663–1670. Gaskell PN, Brooks AC, Maltby L (2007) Variations in the bioaccumulation of a sediment-sorbed hydrophobic compound by benthic macroinvertebrates: Patterns and mechanisms. Environ Sci Technol 41: 1783–1789. Getsinger KD, Petty DG, Madsen JD, Skogerboe JG, Houtman BA, Haller WT, Fox AM (2000) Aquatic dissipation of the herbicide triclopyr in Lake Minnetonka, Minnesota. Pest Manag Sci 56: 388–400. Geyer H, Politzki G, Freitag D (1984) Prediction of ecotoxicological behavior of chemicals: Relationship between n-octanol/water partition coefficient and bioaccumulation of organic chemicals by alga Chlorella. Chemosphere 13: 169–284. Geyer H, Sheeham P, Kotzios D, Freitag D, Korte F (1982) Prediction of ecotoxicological behaviour of chemicals: Relationship between physico-chemical properties and bioaccumulation of organic chemicals in the mussel Mytilus edulis. Chemosphere 11: 1121–1134. Geyer HJ, Scheunert I, Brilggemann R, Steinberg C, Korte F, Kettrup A (1991) QSAR for organic chemical bioconcentration in daphnia, algae and mussels. Sci Total Environ 109/110: 387–394. Glooschenko V, Holdrinet M, Lett JN, Frank R (1979) Bioconcentration of chlordane by the green alga Scenedesmus quadricauda. Bull Environ Contam Toxicol 21: 515–520. Gobas FAPC, Mackay D (1987) Dynamics of hydrophobic organic chemical bioconcentration in fish. Environ Toxicol Chem 6: 495–504.

112

T. Katagi

Gobas FAPC, McNeil EJ, Lovett-Doust L, Heffner GD (1991) Bioconcentration of chlorinated aromatic hydrocarbons in aquatic macrophytes. Environ Sci Technol 25: 924–929. Gobas FAPC, Muir DCG, Mackay D (1988) Dynamics of dietary bioaccumulation and faecal elimination of hydrophobic organic chemicals in fish. Chemosphere 17: 943–962. Gobas FAPC, Opperhuizen A, Hutzinger O (1986) Bioconcentration of hydrophobic chemicals in fish: Relationship with membrane permeation. Environ Toxicol Chem 5: 637–646. Goulding KH, Adams N (1985) The effects of pesticides on algae. Rev Environ Contam Toxicol 5: 199–253. Goulding KH, Ellis SW (1981) The interaction of DDT with two species of freshwater algae. Environ Pollut (Ser A) 25: 271–290. Govers H, Ruepert C, Aiking H (1984) Quantitative structure-activity relationships for polycyclic aromatic hydrocarbons: Correlation between molecular connectivity, physicochemical properties, bioconcentration and toxicity in Daphnia pulex. Chemosphere 13: 227–236. Gowland B, Webster L, Fryer R, Davies I, Moffat C, Stagg R (2002) Uptake and effects of R in the marine mussel, Mytilus edulis. the cypermethrin-containing sea lice treatment Excis Environ Pollut 120: 805–811. Greca MD, Fiorentino A, Pinto G, Pollio A, Previtera L (1996a) Biotransformation of progesterone by the green alga Chlorella emersoni C211-8H. Phytochem 41: 1527–1529. Greca MD, Pinto G, Pistillo P, Pollio A, Previtera L, Temussi F (2008) Biotransformation of ethinylestradiol by microalgae. Chemosphere 70: 2047–2053. Greca MD, Pinto G, Pollio A, Previtera L, Temussi F (2003) Biotransformation of sinapic acid by the green algae Stichococcus bacillaris 155LTAP and Ankistrodesmus braunii C202.7a. Tetrahedron Lett 44: 2779–2780. Greca MD, Previtera L, Fiorentino A, Pinto G, Pollio A (1996b) Bioconversion of 17β-hydroxy17α- methylandrosta-1,4-dien-3-one and androsta-1,4-dien-3,17-dione in cultures of the green alga T76 Scenedesmus quadricauda. Tetrahedron 52: 13981–13990. Gregory Jr WW, Reed JK, Priester Jr LE (1969) Accumulation of parathion and DDT by some algae and protozoa. J Protozool 16: 69–71. Guanzon Jr NG, Fukuda M, Nakahara H (1996) Accumulation of agricultural pesticides by three freshwater microalgae. Fisheries Sci 62: 690–697. Guerrero NR, Taylor MG, Davies NA, Lawrence MAM, Edwards PA, Simkiss K, Wider EA (2002) Evidence of differences in the biotransformation of organic contaminants in three species of freshwater invertebrates. Environ Pollut 117: 523–530. Guisande C, Serrano L (1989) Analysis of protein, carbohydrate and lipid in rotifers. Hydrobiologia 186/187: 330–346. Gutenkauf A, Düker A, Fock HP (1998) Fatae of substituted benzoate in the freshwater green algae, Chlamydomonas reinhardtii 11-32b. Biodegrad 9: 359–368. Haitzer M, Höss S, Traunspurger W, Steinberg C (1998) Effects of dissolved organic matter (DOM) on the bioconcentration of organic chemicals in aquatic organisms – A review. Chemosphere 37: 1335–1362. Hale RC (1989) Accumulation and biotransformation of an organophosphorus pesticide in fish and bivalves. Mar Environ Res 28: 67–71. Halling-Sørensen B, Nyholm N, Kusk KO, Jacobson E (2000) Influence of nitrogen status on the bioconcentration of hydrophobic organic compounds to Selenastrum capricornutum. Ecotoxicol Environ Saf 45: 33–42. Hamer MJ, Goggin UM, Muller K, Maund SJ (1999) Bioavailability of lambda-cthalothrin to Chironomus riparius in sediment-water and water-only systems. Aquat Ecosys Health Manag 2: 403–412. Hand LH, Kuet SF, Lane MCG, Maund SJ, Warinton JS, Hill IR (2001) Influences of aquatic plants on the fate of the pyrethroid insecticide lambda-cyhalothrin in aquatic environments. Environ Toxicol Chem 20: 1740–1745.

Bioconcentration, Bioaccumulation, and Metabolism

113

Hänninen O, Lindstöm-Seppä P, Koivusaari U, Väisänen M, Julkunen A, Juvonen R (1984) Glucuronidation and glucosidation reactions in aquatic species in boreal regions. Biochem Soc Trans 12: 13–17. Hansen LG, Kapoor IP, Metcalf RL (1972) Biochemistry of selective toxicity and biodegradability: Comparative O-dealkylation by aquatic organisms. Comp Gen Pharmac 3: 339–344. Hansen PD (1979) Experiments on the accumulation of lindane by the primary procedures Chlorella spec and Chlorella pyrenoidosa. Arch Environ Contam Toxicol 8: 721–731. Hardy JT, Dauble DD, Felice LJ (1985) Aquatic fate of synfuel residuals: Bioaccumulation of aniline and phenol by the freshwater phytoplankter Scenedesmus quadricauda. Environ Toxicol Chem 4: 29–35. Harkey GA, Klaine SJ (1992) Bioconcentration of trans-chlordane by the midge, Chironomus decorus. Chemosphere 24: 1911–1919. Hartley DM, Johnston JB (1983) Use of the freshwater clam Corbicula manilensis as a monitor for organochlorine pesticides. Bull Environ Contam Toxicol 31: 33–40. Hawker DW, Connell DW (1986) Bioconcentration of lipophilic compounds by some aquatic organisms. Ecotoxicol Environ Saf 11: 184–197. Heisig-Gunkel G, Gunkel G (1982) Distribution of a herbicide (atrazine, s-triazine) in Daphnia pulicaria: A new approach to determination. Arch Hydrobiol Suppl 59: 359–376. Hendriks AJ, van der Linde A, Comelissen G, Sijm DTHM (2001) The power of size. 1. Rate constants and equilibrium ratios for accumulation of organic substrates related to octanol-water partition ratio and species weight. Environ Toxicol Chem 20: 1399–1420. Herbes SE, Allen CP (1983) Lipid quantitation of freshwater invertebrates: Method modification for microquantitation. Can J Fish Aquat Sci 40: 1315–1317. Herbes SE, Risi GF (1978) Metabolic alteration and excretion of anthracene by Daphnia pulex. Bull Environ Contam Toxicol 19: 147–155. Hernández I, Niell FX, Fernández JA (1994) Alkaline phosphatase activity in marine macrophytes: Histochemical localization in some widespread species in southern Spain. Mar Biol 120: 501–509. Hinman ML, Klaine SJ (1992) Uptake and translocation of selected organic pesticides by the rooted aquatic plant Hydrilla verticillata Royle. Environ Sci Technol 26: 609–613. Hirwe AS, Metcalf RL, Kapoor IP (1972) α-Trichloromethylbenzylanilines and αtrichloromethylbenzyl phenyl ethers with DDT-like insecticidal actions. J Agric Food Chem 20: 818–824. Hoarau P, Gnassia-Barelli M, Romeo M, Girard JP (2001) Differential induction of glutathione Stransferases in the clam Ruditapes Decussatus exposed to organic compounds. Environ Toxicol Chem 20: 523–529. Homola E, Chang ES (1997) Distribution and regulation of esterases that hydrolyze methyl farnesoate in Homarus americanus and other crustaceans. Gen Comp Endocrinol 106: 62–72. Hoskin FCG, Walker JE, Mello CM (1999) Organophosphorus acid anhydrolase in slime mold, duckweed and mung bean: A continuing search for a physiological role and a natural substrate. Chemico-Biol Interact 119/120: 399–404. Huckins JN, Petty JD, England DC (1986) Distribution and impact of trifluralin, atrazine and fonofos residues in microcosms simulating a northern prairie wetland. Chemosphere 15: 563–588. Huckle KR, Millburn P (1990) Metabolism, bioconcentration and toxicity of pesticides in fish. In: Progress in Pesticide Biochemistry and Toxicology. Environmental Fate of Pesticides. Hutson DH, Roberts TR (eds). John Wiley & Sons, Ltd. New York, vol 7, Chapter 8, pp 176–243. Hughes JB, Shanks J, Vanderford M, Lauvitzen J, Bhadra R (1997) Transformation of TNT by aquatic plants and tissue cultures. Environ Sci Technol 31: 266–271. Ikenaka Y, Eun H, Ishizaka M, Miyabara Y (2006) Metabolism of pyrene by aquatic crustacean, Daphnia magna. Aquat Toxicol 80: 158–165.

114

T. Katagi

Ingersoll CG, Brunson EL, Wang N, Dwyer FJ, Ankley GT, Mount DR, Huckins J, Petty J, Landrum PF (2003) Uptake and depuration of nonionic organic contaminants from sediment by the oligochaete, Lumbriculus variegatus. Environ Toxicol Chem 23: 872–885. Isensee AR (1976) Variability of aquatic model ecosystem-derived data. Intern J Environ Stud 10: 35–41. Isensee AR, Durbey PS (1983) Distribution of pendimethalin in an aquatic microecosystem. Bull Environ Contam Toxicol 30: 239–244. Isensee AR, Holden ER, Woolson EA, Jones GE (1976) Soil persistence and aquatic bioaccumulation potential of hexachlorobenzene (HCB). J Agric Food Chem 24: 1210–1214. Isensee AR, Jones GE, McCann JA, Pitcher FG (1979a) Toxicity and fate of nine toxaphene fractions in an aquatic model ecosystem. J Agric Food Chem 27: 1041–1046. Isensee AR, Kearney PC, Jones GE (1979b) Modeling aquatic ecosystems for metabolic studies. In: Pesticide and Xenobiotic Metabolism in Aquatic Organisms. Khan MAQ, Lech JJ, Menn JJ (eds) ACS Symp Ser 99, American Chemical Society, Washington, DC, Chapter 13, pp 195–216. James MO (1982) Disposition and taurine conjugation of 2,4-dichlorophenoxyacetic acid, 2,4,5trichlorophenoxyacetic acid, bis(4-chlorophenyl)acetic acid and phenylacetic acid in the spiny lobster, Panulirus argus. Drug Metab Disp 10: 516–522. James MO (1986) Xenobiotic conjugation in fish and other aquatic species. In: Xenobiotic Conjugation Chemistry. Paulson GD, Caldwell J, Hutson DH, Menn JJ (eds), ACS Symp Ser 299, American Chemical Society, Washington, DC, Chapter 2, pp 29–47. James MO (1987) Conjugation of organic pollutants in aquatic species. Environ Health Perspect 71: 97–103. James MO (1989) Cytochrome P450 monooxygenases in crustaceans. Xenobiotica 19: 1063–1076. James MO (1994) Pesticide metabolism in aquatic organisms. Chem Plant Prot 9: 153–189. James MO, Boyle SM (1998) Cytochrome P450 in crustacean. Comp Biochem Physiol 121C: 157–172. James MO, Pritchard JB (1990) Pesticide metabolism in aquatic organisms. In: Pesticide Chemistry, Advances in International Research, Development, and Legislation. Proc 7th Intl Cong Pest Chem (IUPAC), Hamburg, Frechse VCH (ed), Weinheim, pp 277–286. Janer G, LeBlanc GA, Porte C (2005a) Androgen metabolism in invertebrates and its modulation by xenoandrogens. Ann NY Acad Sci 1040: 354–356. Janer G, Mesia-Vela S, Kauffman FC, Porte C (2005b) Sulfatase activity in the oyster Crassostrea virginica: Its potential interference with sulfptransferase determination. Aquat Toxicol 74: 92–95. Jessiman BJ, Qadri SU (1983) Bioaccumulation kinetics of the organochlorine pesticide mirex in amphipods. Ecotoxicol Environ Saf 7: 295–305. Jingi L, Houtian L (1992) Degradation of azo dyes by algae. Environ Pollut 75: 273–278. Johnson BT, Saunders CR, Sanders HO, Campbell RS (1971) Biological magnification and degradation of DDT and aldrin by freshwater invertebrates. J Fish Res Bd Canada 28: 705–709. Johnston JJ, Corbett MD (1986a) The effects of salinity and temperature on the in vitro metabolism of the organophosphorus insecticide fenitrothion by the blue crab, Callinectes sapidus. Pestic Biochem Physiol 26: 193–201. Johnston JJ, Corbett MD (1986b) The uptake and in vivo metabolism of the organophosphate insecticide fenitrothion by the blue crab, Callinectes sapidus. Toxicol Appl Pharmacol 85: 181–188. Jonsson CM, Paraiba LC, Mendoza MT, Sabater C, Carrasco JM (2001) Bioconcentration of the insecticide pyridaphenthion by the green algae Chlorella saccharophila. Chemosphere 43: 321–325. Julin AM, Sanders HO (1977) Toxicity and accumulation of the insecticide imidan in freshwater invertebrates and fishes. Trans Am Fish Soc 106: 386–392. Kale SP, Carvalho FP, Raghu K, Sherkhane PD, Pandit GG, Mohan Rao A, Mukherjee PK, Murthy NBK (1999b) Studies on degradation of 14 C-chlorpyrifos in the marine environment. Chemosphere 39: 969–976.

Bioconcentration, Bioaccumulation, and Metabolism

115

Kale SP, Murthy NBK, Raghu K, Sherkhane PD, Carvalho FP (1999a) Studies on degradation of 14 C-DDT in the marine environment. Chemosphere 39: 959–968. Kale SP, Sherkhane PD, Murthy NBK (2002) Uptake of 14 C-chlorpyrifos by clams. Environ Technol 23: 1309–1311. Kanazawa J (1978) Bioconcentration ratio of diazinon by freshwater fish and snail. Bull Environ Contam Toxicol 20: 613–617. Kanazawa J, Isensee AR, Kearney PC (1975) Distribution of carbaryl and 3,5-xylyl methylcarbamate in an aquatic model ecosystem. J Agric Food Chem 23: 760–763. Kapoor IP, Metcalf RL, Hirwe AS, Coats JR, Khalsa MS (1973) Structure activity correlations of biodegradability of DDT analogs. J Agric Food Chem 21: 310–315. Kapoor IP, Metcalf RL, Hirwe AS, Lu PY, Coats JR, Nystrom RE (1972) Comparative metabolism of DDT, methylchlor, and ethoxychlor in mouse, insects, and in a model ecosystem. J Agric Food Chem 20: 1–6. Kapoor IP, Metcalf RL, Nystrom RF, Sangha GK (1970) Comparative metabolism of methoxychlor, methiochlor and DDT in mouse, insects, and in a model ecosystem. J Agric Food Chem 18: 1145–1152. Karickhoff SW (1981) Semi-empirical estimation of sorption of hydrophobic pollutants on natural sediments and soils. Chemosphere 10: 833–846. Kashiwada S, Mochida K, Ozoe Y, Nakamura T (1995a) Malathion – tolerance and degrading abilities of brakish zooplankton, Sinocalanus tenellus and Oithona davisae. J Pestic Sci 20: 161–164. Kashiwada S, Mochida K, Ozoe Y, Nakamura T (1995b) Contribution of zooplankton to disappearance of organophosphorus insecticides in environmental water. J Pestic Sci 20: 503–512. Kashiwada S, Mochida K, Ozoe Y, Nakamura T (1998) Metabolism of fenitrothion in several brackish and marine zooplankton species. J Pestic Sci 23: 308–311. Katagi (2006) Behavior of pesticides in water-sediment systems. Rev Environ Contam Toxicol 187: 133–251. Katagi T, Miyakado M, Takayama C, Tanaka S (1995) Theoretical estimation of octanol-water partition coefficient for organophosphorus pesticides. In: Classical and Three-Dimensional QSAR in Agrochemistry. Hansch C, Fujita T (eds) ACS Symp Ser 606, American Chemical Society, Washington, DC, Chapter 4, pp 48–61. Kawatski JA, Bittner MA (1975) Uptake, elimination and biotransformation of the lampricide 3-trifluoromethyl-4-nitrophenol (TFM) by larvae of the aquatic midge Chironomus tentans. Toxicology 4: 183–194. Kawatski JA, Schmulbach JC (1970) Chlamydotheca arcuata: Autoradiographic localization of 14 C-aldrin and 14C-dieldrin. Trans Am Microsc Soc 89: 424–427. Kawatski JA, Schmulbach JC (1971) Epoxidation of aldrin by a freshwater ostracod. J Econ Entmol 64: 316–317. Kawatski JA, Schmulbach JC (1972) Uptake and elimination of 14 C-aldrin and 14 C-dieldrin by the ostracod Chlamydotheca arcuata (Sars). Int J Environ Anal Chem 1: 283–291. Kazano H, Asakawa M, Tomizawa C (1976) Balance study of 14 C derived from N-CH3 -(14 C)labeled 3,5-xylyl methylcarbamate insecticide (XMC) applied on a model ecosystem. Appl Ent Zool 11: 263–266. Kearney PC, Isensee AR, Kontson A (1977a) Distribution and degradation of dinitroaniline herbicides in an aquatic ecosystem. Pestic Biochem Physiol 7: 242–248. Kearney PC, Oliver JE, Helling CS, Isensee AR, Kontson A (1977b) Distribution, movement, persistence and metabolism of N-nitrosoatrazine in soils and a model aquatic ecosystem. J Agric Food Chem 25: 1177–1181. Keil JE, Priester LE (1969) DDT uptake and metabolism by a marine diatom. Bull Environ Contam Toxicol 4: 169–173. Kenney-Wallace G, Blackman GE (1972) The uptake of growth substances. XIV. Patterns of uptake by lemna minor of phenoxyacetic and benzoic acids following progressive chlorination. J Exp Botany 23: 114–127.

116

T. Katagi

Kent RA, Currie D (1995) Predicting algal sensitivity to a pesticide stress. Environ Toxicol Chem 14: 983–991. Khalil Z, Mostafa IY (1986) Interaction of pesticides with freshwater algae. I. Effect of methomyl and its possible degradation by Phormidium fragile. J Environ Sci Health B21: 289–301. Khalil Z, Mostafa IY (1987) Interaction of pesticides with freshwater algae. II. Degradation of 14 C-labelled carbofuran by Anabaena oryzae and Phormidium fragile. Isotope Rad Res 19: 35–41. Khan HM, Neudoff S, Khan MAQ (1975) Absorption and elimination of photodieldrin by daphnia and goldfish. Bull Environ Contam Toxicol 13: 582–587. Khan MAQ, Kamel A, Wolin RJ, Runnels J (1972) In vivo and in vitro epoxidation of aldrin by aquatic food chain organisms. Bull Environ Contam Toxicol 8: 219–228. Kikuchi R, Yasutaniya T, Takimoto Y, Yamada H, Miyamoto J (1984) Accumulation and metabolism of fenitrothion in three species of algae. J Pestic Sci 9: 331–337. King I, Childs MT, Dorsett C, Ostrander JG, Monsen ER (1990) Shellfish: Proximate composition, minerals, fatty acids and sterols. J Am Diet Assoc 90: 677–685. King PH, Yeh HH, Warren PS, Randall CW (1969) Distribution of pesticides in surface waters. J Am Water Works Assoc 61: 483–486. Klekner V, Kosaric N (1992) Degradation of phenols by algae. Environ Technol 13: 493–501. Klosterhaus SL, DiPinto LM, Chandler GT (2003) A comparative assessment of azinphos methyl bioaccumulation and toxicity in two estuarine meiobenthic harpacticoid copepods. Environ Toxicol Chem 22: 2960–2968. Kneifel H, Elmendorff K, Hegewald E, Soeder CJ (1997) Biotransformation of 1naphthalenesulfonic acid by the green alga Scenedesmus obliquus. Arch Microbiol 167: 32–37. Knezovich JP, Crosby DG (1985) Fate and metabolism of o-toluidine in the marine bivalve molluscs Mytilus edulis and Crassostrea gigas. Environ Toxicol Chem 4: 435–446. Knezovich JP, Harrison FL (1988) The bioavailability of sediment-sorbed chlorobenzenes to larvae of the midge, Chironomus decorus. Ecotoxicol Environ Saf 15: 226–241. Knuth ML, Heinis LJ (1995) Distribution and persistence of diflubenzuron within littoral enclosure mesocosms. J Agric Food Chem 43:1087–1097. Kobayashi K, Akitake H, Tomiyama T (1970a) Studies on the metabolism of pentacjlorophenate, a herbicide, in aquatic organisms – II. Biochemical change of PCP in sea water by detoxication mechanism of Tapes philippinarum. Bull Jpn Soc Sci Fish 36: 96–102. Kobayashi K, Akitake H, Tomiyama T (1970b) Studies on the metabolism of pentachlorophenate, a herbicide, in aquatic organisms – III. Isolation and identification of a conjugated PCP yielded by a shell-fish, Tapes philippinarum. Bull Jpn Soc Sci Fish 36: 103–108. Kobayashi K, Nakamura Y, Imada N (1985a) Metabolism of an organophosphorus insecticide, fenitrothion, in tiger shrimp Penaeus japonicus. Bull Jpn Soc Sci Fish 51: 599–603. Kobayashi K, Nakamura Y, Imada N (1985b) Formation of the sulfate and glucoside metabolites of fenitrothion in tiger shrimp Penaeus japonicus. Bull Jpn Soc Sci Fish 51: 2013–2017. Kobayashi K, Oshima Y, Hamada S, Taguchi C (1987) Induction of phenol-sulfate conjugating activity by exposure to phenols and duration of its induced activity in short-necked clam. Bull Jpn Soc Sci Fish 53: 2073–2076. Kobayashi K, Rompas RM, Oshima Y, Imada N (1990) A comparative study on the toxicity, absorption and depuration of fenitrothion and its oxon in Japanese tiger shrimp. Bull Jpn Soc Sci Fish 56: 923–928. Koelmans AA, Sánchez Jiménez C (1994) Temperature dependency of chlorobenzene bioaccumulation in phytoplankton. Chemosphere 28: 2041–2048. Kominami S (1993) Substrate binding and the reduction of cytochrome P-450. In: Cytochrome P-450. 2nd Ed. Omura T, Ishimura Y, Fujii-Kuriyama Y (eds) VCH Publishers Inc., New York, pp 64–80. Kooper IP, Metcalf RL, Hirwe AS, Coats JR, Khalsa MS (1973) Structure activity correlations of biodegradability of DDT analogs. J Agric Food Chem 21: 310–315.

Bioconcentration, Bioaccumulation, and Metabolism

117

Kraaij R, Maayer P, Busser FJM, Bolscher MVH, Seinen W, Tolls J (2003) Measured pore-water concentration make equilibrium partitioning work – A data analysis. Environ Sci Technol 37: 268–274. Krieger RI, Gee SJ, Lim LO, Ross JH, Wilson A (1979) Disposition of toxic substances in mussels (Mytilus califorianus): Preliminary metabolic and histologic studies. In: Pesticide and Xenobiotic Metabolism in Aquatic Organisms. Khan MAQ, Lech JJ, Menn JJ (eds) ACS Symp Ser 99, American Chemical Society, Washington, DC, Chapter 16, pp 259–277. Krieger RI, Lee PW (1976) Properties of the aldrin epoxidase system in gut and fat body of a caddisfly larva. J Eco Entmol 66: 1–6. Kukkonen J, Oikari A (1988) Sulphate conjugation is the main route of pentachlorophenol metabolism in Daphnia magna. Comp Biochem Physiol 91C: 465–468. Kukkonen J, Oikari A (1991) Bioavailability of organic pollutants in boreal waters with varying levels of dissolved organic material. Water Res 25: 455–463. Kumar S, Lal R (1988) Uptake of dieldrin, dimethoate and permethrin by cyanobacteria, Anabaena sp. and Aulosira fertilissima. Environ Pollut 54: 55–61. Kurelec (1985) Exclusive activation of aromatic amines in the marine mussel Mytilus edulis by FAD-containing monooxygenase. Biochem Biophys Res Commun 127: 773–778. Kuritz T, Bocanera LV, Rivera NS (1997) Dechlorination of lindane by the cyanobacterium Anabaena sp. strain PCC7120 depends on the function of the nir operon. J Bacteriol 179: 3368–3370. Kuritz T, Wolk CP (1995) Use of filamentous cyanobacteria for biodegradation of organic pollutants. Appl Environ Microbiol 61: 234–238. Küsel AC, Sianoudis J, Leibfritz D, Grimme LH, Mayer A (1990) The dependence of the cytoplasmic pH in aerobic and anaerobic cells of the green algae Chlorella fusca and Chlorella vulgaris on the pH of the medium as determined by 31 P in vivo NMR spectroscopy. Arch Microbiol 153: 254–258. Kusk KO (1996) Bioavailability and effect of pirimicarb on Daphnia magna in a laboratory freshwater/sediment system. Arch Environ Contam Toxicol 31: 252–255. Lacaze JPCL, Stobo LA, Turrell EA, Quilliam MA (2007) Solid-phase extraction and liquid chromatography – mass spectroscopy for the determination of free fatty acids in shellfish. J Chromatogr A 1145: 51–57. Lal R, Saxena DM (1982) Accumulation, metabolism, and effects of organochlorine insecticides on microorganisms. Microbial Rev 46: 95–127. Lal S, Lal R, Saxena DM (1987) Bioconcentration and metabolism of DDT, fenitrothion and chlorpyrifos by the blue-green algae Anabaena sp. and Aulosira fertilissima. Environ Pollut 46: 187–196. Lalah JO, Behechti A, Severin GF, Lenoir D, Günther K, Kettrup A, Schramm KW (2003) The bioaccumulation and fate of a branched 14 C-nonylphenol isomer in Lymnaea stagnalis L. Environ Toxicol Chem 22: 1428–1436. Landrum PF (1988) Toxicokinetics of organic xenobiotics in the amphipod, Pontoporeia hoyi: Role of physiological and environmental variables. Aquat Toxicol 12: 245–271. Landrum PF (1989) Bioavailability and toxicokinetics of polycyclic aromatic hydrocarbons sorbed to sediments for the amphipod Pontoporeia hoyi. Environ Sci Technol 23: 588–595. Landrum PF, Crosby DG (1981) Comparison of the disposition of several nitrogen-containing compounds in the sea urchin and other marine invertebrates. Xenobiotica 11: 351–361. Landrum PF, Dupuis WS (1990) Toxicity and toxicokinetics of pentachlorophenol and carbaryl to Pontporeia hoyi and Mysis relicta. In: Aquatic Toxicology and Risk Assessment: Thirteenth Volume. Landis WG, van der Schalie WH (eds) STP 1096, ASTM, Philadelphia, pp 278–289. Landrum PF, Fisher SW (1998) Influence of lipids on the bioaccumulation and trophic transfer of organic contaminants in aquatic organisms. In: Lipids in Freshwater Ecosystems. Arts MT, Wainman BC (eds) Springer, New York, Chapter 9, pp 203–234. Landrum PF, Scavia D (1983) Influence of sediment on anthracene uptake, depuration and biotransformation by the smphipod Hyalella azteca. Can J Fish Aquat Sci 40: 298–305.

118

T. Katagi

Lavado R, Janer G, Porte C (2006) Steroid levels and steroid metabolism in the mussel Mytilus edulis: The modulating effect of dispersed crude oil and alkylphenols. Aquat Toxicol 78S: S65–S72. LeBlanc GA, Cochrane BJ (1985) Modulation of substrate-specific glutathione S-transferase activity in Daphnia magna with concomitant effects on toxicity tolerance. Comp Biochem Physiol 82C: 37–42. LeBlanc GA, Cochrane BJ (1987) Identification of multiple glutathione-S-transferases from Daphnia magna. Comp Biochem Physiol 88B: 39–45. Lech JJ, Vodicnik MJ (1985) Biotransformation. In: Fundamentals of aquatic toxicology. Rand GM, Petrocelli SR (eds) Hemisphere Publishing Co., New York, Chapter 18, pp 520–557. Lee AH, Lu PY, Metcalf RL, Hsu EL (1976) The environmental fate of three dichlorophenyl nitrophenyl ether herbicides in a rice paddy model ecosystem. J Environ Qual 5: 482–486. Lee RF (1981) Mixed function oxygenases (MFO) in marine invertebrates. Biol Lett 2: 87–105. Lee RF (1998) Annelid cytochrome P450. Comp Biochem Physiol 121C: 173–179. Lee RF, Valkirs AO, Seligman PF (1989) Importance of microalgae in the biodegradation of tributyltin in estuarine waters. Environ Sci Technol 23: 1515–1518. Lee SE, Kim JS, Kennedy IR, Park JW, Kwon GS, Koh SC, Kim JE (2003) Biotransformation of an organochlorine insecticide, endosulfan, by Anabaena species. J Agric Food Chem 51: 1336–1340. Legierse KCHM, Sijm DTHM, van Leeuwen CJ, Seinen W, Hermens JLM (1998) Bioconcentration kinetics of chlorobenzenes and the organophosphorus pesticide chlorthion in the pond snail Lymnaea stagnalis – a comparison with the guppy Poecilia veticulata. Aquat Toxicol 41: 301–323. Leppänen MT, Kukkonen JVK (1998) Relative importance of ingested sediment and pore water as bioaccumulation routes for pyrene to oligochaete (Lumbriculus variegates, Müller). Environ Sci Technol 32: 1503–1508. Leppänen MT, Kukkonen JVK (2000) Fate of sediment-associated pyrene and benzo[a]pyrene in the freshwater oligochaete Lumbriculus variegatus (Müller). Aquat Toxicol 49: 199–212. Leppänen MT, Kukkonen JVK (2006) Evaluating the role of desorption in bioavailability of sediment-associated contaminants using oligochaetes, semipermeable membrane devices and Tenax extraction. Environ Pollut 140: 150–163. Leppänen MT, Landrum PF, Kukkonen JVK, Greenberg MS, Burton GA, Robinson SD, Gossiaux DC (2003) Investigating the role of desorption on the bioavailability of sedimentassociated 3,4,3 ,4 -tetrachlorobiphenyl in benthic invertebrates. Environ Toxicol Chem 22: 2861–2871. Leversee GJ, Giesy JP, Landrum PF, Gerould S, Bowling JW, Fannin TE, Haddock JD, Bartell SM (1982) Kinetics and biotransformation of benzo[a]pyrene in Chironomus riparius. Arch Environ Contam Toxicol 11: 25–31. Lewin RA (1974) Biochemical taxonomy. In: Algal Physiology and Biochemistry. Stewart WDP (ed) Botanical Monographs vol 10, Blackwell Scientific Publications, Oxford, Chapter 1, pp 1–39. Li CLJ, James MO (1993) Glucose and sulfate conjugations of phenol, β-naphthol and 3hydroxybenzo[a]pyrene by the American lobster (Homarus americanus). Aquat Toxicol 26: 57–72. Lin H, Jiang J, Xue CH, Zhang B, Xu JC (2003) Seasonal changes in phospholipids of mussels (Mytilus edulis Linne). J Sci Food Agric 83: 133–135. Lindquist B, Warshawsky D (1985a) Identification of the 11,12-dihydro-11,12dihydroxybenzo[a]pyrene as a major metabolite produced by the green algae, Selenastrum capricornutum. Biochem Biophys Res Commun 130: 71–75. Lindquist B, Warshawsky D (1985b) Stereospecificity in algal oxidation of the carcinogen benzo[a]pyrene. Experientia 41: 767–769. Lindström-Seppä P, Koivusaari U, Hänninen O (1983) Metabolism of foreign compounds in freshwater crayfish (Astacus astacus L) tissues. Aquat Toxicol 3: 35–46.

Bioconcentration, Bioaccumulation, and Metabolism

119

Liu ZT, Kong ZM, Zhou F, Wang LS (1996) Bioconcentration and toxicity effect on lipid content of aquatic organisms. Bull Environ Contam Toxicol 56: 135–142. Livingstone DR, Kirchin MA, Wiseman A (1989) Cytochrome P-450 and oxidative metabolism in molluscs. Xenobiotica 19: 1041–1062. Lockhart WL, Billeck BN, de March GE, Muir DCG (1983) Uptake and toxicity of organic compounds: Studies with an aquatic macrophyte (Lemna minor). ASTM Spec Tech Publ 802: 460–468. Lockhart WL, Metner DA, Billeck BN, Rawn GP, Muir DCG (1984) Bioaccumulation of some forestry pesticides in fish and aquatic plants. In: Chemical and Biological Controls in Forestry, Garner WY, Harvey Jr J (eds), ACS Symposium Ser 238, American Chemical Society, Washington, DC, pp 297–315. Lohner TW, Fisher SW (1990) Effects of pH and temperature on the acute toxicity and uptake of carbaryl in the midge, Chironomus riparius. Aquat Toxicol 16: 335–354. Londoño DK, Siegfried BD, Lydy MJ (2004) Atrazine induction of a family 4 cytochrome P450 gene in Chironomus tentans (Diptera: Chironomidae). Chemosphere 56: 701–706. Looser PW, Fent K, Berg M, Goudsmit GH, Schwarzenbach RP (2000) Uptake and elimination of triorganotin compounds by larval midge Chironomus riparius in the absence and presence of Aldrich humic acid. Environ Sci Technol 34: 5165–5171. Lotufo GR, Farrar JD, Duke BM, Bridges TS (2001b) DDT toxicity and critical body residue in the amphipod Leptocherirus plumulosus in exposures to spiked sediment. Arch Environ Contam Toxicol 41: 142–150. Lotufo GR, Landrum PF, Gedeon ML (2001a) Toxicity and bioaccumulation of DDT in freshwater amphipods in exposures to spiked sediments. Environ Toxicol Chem 20: 810–825. Lotufo GR, Landrum PF, Gedeon ML, Tique EA, Herche LR (2000) Comparative toxicity and toxicokinetics of DDT and its major metabolites in freshwater amphipods. Environ Toxicol Chem 19: 368–379. Lovell CR, Eriksen NT, Lewitus AJ (2002) Resistance of the marine diatom Thalassiosira sp. to toxicity of phenolic compounds. Mar Ecol Prog Ser 229: 11–18. Lu PY, Metcalf RL (1975) Environmental fate and biodegradability of benzene derivatives as studies in a model aquatic ecosystem. Environ Health Perspect 10: 269–284. Lu PY, Metcalf RL, Hirwe AS, Williams JW (1975) Evaluation of environmental distribution and fate of hexachlorocyclopentadiene, chlordane, heptachlor, and heptachlor epoxide in a laboratory model ecosystem. J Agric Food Chem 23: 967–973. Lu PY, Metcalf RL, Plummer N, Mandel D (1977) The environmental fate of three carcinogens: Benzo[a]pyrene, benzidine and vinyl chloride evaluated in laboratory model ecosystems. Arch Environ Contam Toxicol 6: 129–142. Lydy MJ, Hayton WL, Staubus AE, Fisher SW (1994) Bioconcentration of 5,5 ,6-trichlorobiphenyl and pentachlorophenol in the midge, Chironomus riparius, as measured by a pharmacokinetic model. Arch Environ Contam Toxicol 26: 251–256. Lydy MJ, Lasater JL, Landrum PF (2000) Toxicokinetics of DDE and 2-chlorobiphenyl in Chironomus tentans. Arch Environ Contam Toxicol 38: 163–168. Lydy MJ, Lohner TW, Fisher SW (1990) Influence of pH, temperature and sediment type on the toxicity, accumulation and degradation of parathion in aquatic systems. Aquat Toxicol 17: 27–44. Lynch TR, Johnson HE, Adams WJ (1982) The fate of atrazine and a hexachlorobiphenyl isomer in naturally-derived model stream ecosystems. Environ Toxicol Chem 1: 179–192. Lyytikäinen M, Pehkonen S, Akkanen J, Leppänen M, Kukkonen VK (2007) Bioaccumulation and biotransformation of polycyclic aromatic hydrocarbons during sediment tests with oligochaetes (Lumbriculus variegatus). Environ Toxicol Chem 26: 2660–2666. Mackay D (1982) Correlation of bioconcentration factors. Environ Sci Technol 16: 274–278. Mackay D, Hughes AI (1984) Three-parameter equation describing the uptake of organic compounds by fish. Environ Sci Technol 18: 439–444.

120

T. Katagi

Mackiewicz M, Deubert KH, Gunner HB, Zuckerman BM (1969) Study of parathion biodegradation using gnotobiotic techniques. J Agric Food Chem 17: 129–130. Mäenpää KA, Sormunen AJ, Kukkonen JVK (2003) Bioaccumulation and toxicity of sediment associated herbicides (ioxynil, pendimethalin and bentazone) in Lumbriculus variegatus (Oligochaete) and Chironomus riparius (Insecta). Ecotoxicol Environ Saf 56: 398–410. Mäenpää KA, Sorsa K, Lyytikäinen M, Leppänen MT, Kukkonen JVK (2008) Bioaccumulation, sublethal toxicity, and biotransformation of sediment-associated pentachlorophenol in Lumbriculus variegatus (oligochaeta). Ecotoxicol Environ Saf 69: 121–129. Maguire RJ, Wong PTS, Rhamey JS (1984) Accumulation and metabolism of tri-n-butyltin cation by a green algae, Ankistrodesmus falcatus. Can J Fish Aquat Sci 41: 537–540. Mailhot H (1987) Prediction of algal bioaccumulation and uptake rate of nine organic compounds by ten physicochemical properties. Environ Sci Technol 21: 1009–1013. Mäkelä P, Oikari OJ (1990) Uptake and body distribution of chlorinated phenolics in the freshwater mussel, Anodonta anatine L. Ecotoxicol Environ Saf 20: 354–362. Maki AW, Johnson HE (1977) Kinetics of lampricide (TFM, 3-trifluoromethyl-4-nitrophenol) residues in model stream communities. J Fish Res Board Can 34: 276–281. Manthey M, Faust M, Smolk S, Grimme LH (1993) Herbicide bioconcentration in algae: Studies on lipophilicity–activity relationships (LSAR) with Chlorella fusca. Sci Total Environ (Suppl.) 1: 453–459. Marguis LY, Comes RD, Yang CY (1981) Absorption and translocation of fluridone and glyphosate in submerged vascular plants. Weed Sci 29: 229–236. Markwell RD, Connell DW, Gabric AJ (1989) Bioaccumulation of lipophilic compounds from sediments by oligochaetes. Water Res 23: 1443–1450. Mason JW, Rowe DR (1976) The accumulation and loss of dieldrin and endrin in the eastern oyster. Arch Environ Contam Toxicol 4: 349–360. Mathur R, Saxena DM (1986) Effect of hexachlorohexane (HCH) isomers on growth of and their accumulation in the blue-green alga, Anabaena sp. (ARM 310). Environ Biol 7: 239–251. Matsumura F (1977) Absorption, accumulation, and elimination of pesticides by aquatic organisms. Environ Sci Res 10: 77–105. Matsumura F, Esaac EG (1979) Degradation of pesticides by algae and aquatic microorganisms. In: Pesticide and Xenobiotic Metabolism in Aquatic Organisms. Khan MAQ, Lech JJ, Menn JJ (eds) ACS Symp Ser 99, American Chemical Society, Washington, DC, Chapter 22, pp 371–387. Maund SJ, Hamer MJ, Lane MCG, Farrelly E, Rapley JH, Goggin UM, Gentle WE (2002) Partitioning, bioavailability, and toxicity of the pyrethroid insecticide cypermethrin in sediments. Environ Toxicol Chem 21: 9–15. Mayer Jr FL, Sanders HO (1973) Toxicology of phthalic acid esters in aquatic organisms. Environ Health Persp 3: 153–157. McCarthy JF (1983) Role of particulate organic matter in decreasing accumulation of polynuclear aromatic hydrocarbons by Daphnia magna. Arch Environ Contam Toxicol 12: 559–568. McElroy AE (1990) Polycyclic aromatic hydrocarbon metabolism in the polychaete Nereis virens. Aquat Toxicol 18: 35–50. McLeese DW, Sergeant DB, Metcalf CD, Zitko V, Burridge LE (1981) Uptake and excretion of aminocarb and pesticide diluent 585 by mussels (Mytilus edulis). Bull Environ Contam Toxicol 24: 575–581. McLeese DW, Zitko V, Sergeant DB (1979) Uptake and excretion of fenitrothion by clams and mussels. Bull Environ Contam Toxicol 22: 800–806. Meems N, Steinberg CEW, Wiegand C (2004) Direct and interacting toxicological effects on the waterflea (Daphnia magna) by natural organic matter, synthetic humic substances and cypermethrin. Sci Total Environ 319: 123–136. Megharaj M, Madhavi DR, Sreenivasulu C, Umamaheswari A, Venkateswarlu K (1994) Biodegradation of methyl parathion by soil isolate of microalgae and cyanobacteria. Bull Environ Contam Toxicol 53: 292–297.

Bioconcentration, Bioaccumulation, and Metabolism

121

Megharaj M, Venkateswarlu K, Rao AS (1987) Metabolism of monocrotophos and quinalophos by algae isolated from soil. Bull Environ Contam Toxicol 39: 251–256. Mehetre ST, Kale SP, Sherkhane PD, Murthy NBK (2002) Uptake of 14 C-DDE by marine clams. J Nuclear Agric Biol 31: 105–109. Merlin G, Vuillod M, Lissolo T, Clement B (2002) Fate and bioaccumulation of isoproturon in outdoor aquatic microcosms. Environ Toxicol Chem 21: 1236–1242. Metcalf RL (1976) Laboratory model ecosystem evaluation of the chemical and biological behavior of radiolabeled micropollutants. Environ Qual Saf 5: 141–151. Metcalf RL, Booth GM, Schuth CK, Hansen DJ, Lu PY (1973a) Uptake and fate of 2ethylhexylphthalate in aquatic organisms and in a model ecosystem. Environ Health Perspect 4: 27–34. Metcalf RL, Kapoor IP, Lu PY, Schuth CK, Sherman P (1973b) Model ecosystem studies of the environmental fate of six organochlorine pesticides. Environ Health Perspect 4: 35–44. Metcalf RL, Lu PY, Bowlus S (1975) Degradation and environmental fate of 1-(2,6difluorobenzoyl)-3- (4-chlorophenyl)urea. J Agric Food Chem 23: 359–364. Metcalf RL, Sanborn JR (1975) Pesticides and environmental quality in Illinois. Ill Nat Hist Surv Bull 31: 381–436. Metcalf RL, Sangha GK, Kapoor IP (1971) Model ecosystem for the evaluation of pesticide biodegradability and ecological magnification. Environ Sci Technol 5: 709–713. Meyer GM, Meyer EI, Meyns S (2000) Lipid content of stream macroinvertebrates. Arch Hydrobiol 147: 447–463. Meylan WM, Howard PH, Boethling RS, Aronson D, Printup H, Gonchie S (1999) Improved method for estimating bioconcentration/bioaccumulation factor from octanol/water partition coefficient. Environ Toxicol Chem 18: 664–672. Michel XR, Beasse C, Narbonne JF (1995) In vivo metabolism of benzo[a]pyrene in the mussel Mytilus galloprovinciallis. Arch Environ Contam Toxicol 28: 215–222. Miller JDA (1962) Fats and steroids. In: Physiology and Biochemistry of Algae. Lewin RA (ed) Academic Press, New York, Chapter 21, pp 357–370. Mitsou K, Koulianou A, Lambropoulou D, Pappas P, Albanis T, Lekka M (2006) Growth rate effects, responses of antioxidant enzymes and metabolic fate of the herbicide propanil in the aquatic plant Lemna minor. Chemosphere 62: 275–284. Miyamoto J, Klein W, Takimoto Y, Roberts TR (1985) Critical evaluation of model ecosystems. Pure Appl Chem 57: 1523–2536. Miyamoto J, Mikami N, Takimoto Y (1990) The fate of pesticides in aquatic ecosystems. In: Progress in Pesticide Biochemistry and Toxicology. Environmental Fate of Pesticides. Hutson DH, Roberts TR (eds) John Wiley & Sons, Ltd., New York, vol 7, Chapter 6, pp 123–147. Miyamoto J, Takimoto Y, Mihara K (1979) Metabolism of organophosphorus insecticides in aquatic organisms, with special emphasis on fenitrothion. In: Pesticide and Xenobiotic Metabolism in Aquatic Organisms. Khan MAQ, Lech JJ, Menn JJ (eds) ACS Symp Ser 99, American Chemical Society, Washington, DC, Chapter 1, pp 3–20. Miyamoto M, Tanaka H, Katagi T (2008) Ecotoxicological risk assessment of pesticides in aquatic ecosystems. Sumitomo Kagaku R & D Report, vol 1, pp 1–18. Miyazaki S, Thorsteinson AJ (1972) Metabolism of DDT by freshwater diatom. Bull Environ Contam Toxicol 8: 81–83. Møhlenberg F, Petersen S, Gustavsson K, Lauridsen T, Friberg N (2001) Mesocosm experiments in the approval procedure for pesticides – a literature study on effects of mesocosm characteristics and validity of extrapolation methods to protect sensitive species. Pesticide Research No. 56, Danish Environmental Protection Agency, Denmark, pp 107. Moita F, Siegfried BD, Scharf ME, Lydy MJ (2000) Atrazine induction of cytochrome P450 in Chironomus tentans larvae. Chemosphere 40: 285–291. Moody RP, Greenhalgh R, Lockhart L, Weinberger P (1978) The fate of fenitrothion in an aquatic ecosystem. Bull Environ Contam Toxicol 19: 8–14.

122

T. Katagi

Moore R, Toro E, Stanton M, Khan MAQ (1977) Absorption and elimination of 14 C-alpha and gamma chlordane by a freshwater alga, daphnid and goldfish. Arch Environ Contam Toxicol 6: 411–420. Morrison HA, Gobas FAPC, Lazar R, Haffner GD (1996) Development and verification of a bioaccumulation model for organic contaminants in benthic invertebrates. Environ Sci Technol 30: 3377–3384. Mostafa FIY, Helling CD (2001) Isoproturon degradation as affected by the growth of two algal species at different concentrations and pH values. J Environ Sci Health B36: 709–727. Mostafa IY, Shabana EF, Khalil Z, Mostafa FIY (1991) The metabolic fate of 14 C-parathion by some fresh water phytoplankton and its possible effects on the algal metabolism. J Environ Sci Health B26: 499–512. Muir DCG, Grift NP, Townsend BE, Metner DA, Lockhart WL (1982) Comparison of the uptake and bioconcentration of fluridone and terbutryn by rainbow trout and Chironomus tentans in sediment and water systems. Arch Environ Contam Toixicol 11: 595–602. Muir DCG, Kenny DF, Grift NP, Robinson RD, Titman RD, Murkin HR (1991) Fate and acute toxicity of bromoxynil esters in an experimental prairie wetland. Environ Toxicol Chem 10:395–406. Muir DCG, Rawn GP, Grift NP (1985a) Fate of the pyrethroid insecticide deltamethrin in small ponds: A mass balance study. J Agric Food Chem 33: 603–609. Muir DCG, Rawn GP, Townsend BE, Lockhart WL, Greenhalgh R (1985b) Bioconcentration of cypermethrin, deltamethrin, fenvalerate and permethrin by Chironomus tentans larvae in sediment and water. Environ Toxicol Chem 4: 51–61. Muir DCG, Townsend BE, Lockhart WL (1983) Bioavailability of six organic chemicals to Chironomus tentans larvae in sediment and water. Environ Toxicol Chem 2: 269–281. Muir DCG, Yarechewski AL, Neal BR (1992) Influence of surface films on the fate of deltamethrin following aerial application to prairie ponds. Environ Toxicol Chem 11:581–591. Murakami Y, Fukushima S, Nishimune T, Sueki K, Tanaka R (1990) Examination of pollution by organochlorine herbicides and fungicides in Yodo River and Osaka Bay (Studies on environmental contaminants in food. 3) J Food Hyg Soc Jpn 31: 36–43. Murakami Y, Nishimura T, Sueki K (1993) Studies on a rice plant pesticide – Oxadiazon and its metabolites in shellfishes. Toxicol Environ Chem 38: 119–129. Murphy KJ, Mooney BD, Mann NJ, Nichols PD, Sinclair AJ (2002) Lipid, FA, and sterol composition of New Zealand green lipped mussel (Perna canaliculus) and Tasmanian blue mussel (Mytilus edulis). Lipids 37: 587–595. Murthy RC, Sarena P (1980) Esterase activity in the hepatopancreas of Macrobrachium lamarrei (Crustacea: Decapoda). Experientia 36: 323–325. Nakajima N, Ohshima Y, Edmonds JS, Tamaoki M, Kubo A, Aono H, Saji H, Morita M (2004) Glucosylation of bisphenol a by various plant species. Phyton (Austria) 45: 471–476. Nakamura T, Mochida K (1988) Kinetic evaluation of the uptake and decomposition of pesticides by algae (1). J Pestic Sci 13: 93–98. Naqvi SM, Flagge CT, Hawkins RL (1990) Arsenic uptake and depuration by red crayfish, Procambarus clarkii, exposed to various concentration of monosodium methanearsonate (MSMA) herbicide. Bull Environ Contam Toxicol 45: 94–100. Naqvi SM, Newton DJ (1990) Bioaccumulation of endosulfan (Thiodan insecticide) in the tissues of Luisiana crayfish, Procambarus clarkii. J Environ Sci Health B25: 511–526. Narayana Rao VVS, Lal R (1987) Uptake and metabolism of insecticides by blue-green algae Anabaena and Aulosira fertilissima. Microbios Lett 36: 143–147. Narro ML, Cerniglia CE, van Baalen C, Gibson DT (1992a) Metabolism of phenanthrene by the marine cyanobacterium Armenellum quadruplicatum PR-6. Appl Environ Microbiol 58: 1351–1359. Narro ML, Cerniglia CE, van Baalen C, Gibson DT (1992b) Evidence for an NIH shift in oxidation of naphthalene by the marine cyanobacterium Oscillatoria sp. strain JCM. Appl Environ Microbiol 58: 1360–1363.

Bioconcentration, Bioaccumulation, and Metabolism

123

Nawaz S, Kirk KL (1995) Temperature effects on bioconcentration of DDE by Daphnia. Freshwater Biol 34: 173–178. Nebeker AV, Griffis WL, Wise CM, Hopkins E, Barbitta JA (1989) Survival, reproduction and bioconcentration in invertebrates and fish exposed to hexachlorobenzene. Environ Toxicol Chem 8: 601–611. Neely WB, Branson DR, Blau GE (1974) Partition coefficient to measure bioconcentration potential of organic chemicals in fish . Environ Sci Technol 8: 1113–1115. Neudorf S, Khan MAQ (1975) Pick-up and metabolism of DDT, dieldrin and photodieldrin by a freshwater akga (Ankistrodesmus amalloides) and a microcrustacean (Daphnia pulex). Bull Environ Contam Toxicol 13: 443–450. Neumann W, Leasch H, Urbach W (1987) Mechanisms of herbicide sorption in microalgae and the influence of environmental factors. Pestic Biochem Physiol 27: 189–200. Nikkilä A, Halme A, Kukkonen JVK (2003) Toxicokinetics, toxicity and lethal body residues of two chlorophenols in the oligochaete worm, Lumbriculus variegatus, in different sediments. Chemosphere 51: 35–46. Nikkilä A, Paulsson M, Almgren, Blanck H, Kukkonen JVK (2001) Atrazine uptake, elimination, and bioconcentration by periphyton communities and Daphnia magna: Effects of dissolved organic carbon. Environ Toxicol Chem 20: 1003–1011. Nordone AJ, Dotson TA, Kovacs MF, Doane R, Biever RC (1998) Metabolism of [14 C]acrolein R Herbicide): Nature and magnitude of residues in freshwater fish and shellfish. (Magnacide H Environ Toxicol Chem 17: 276–281. Nuutinen S, Landrum PF, Schuler LJ, Kukkonen JVK, Lydy MJ (2003) Toxicokinetics of organic contaminants in Hyalella azteca. Arch Environ Contam Toxicol 44: 467–475. Nzengung VA, O’Neill WL, McCutcheon SC, Wolfe NL (2003) Sequestration and transformation of water soluble halogenated organic compounds using aquatic plants, algae, and microbial mats. In: Phytoremediation: Transformation and Control of Contaminants. John Wiley & Sons, Inc., Chichester, Chapter 16, pp 499–528. Ohkawa H, Kikuchi R, Miyamoto J (1980) Bioaccumulation and biodegradation of the (S)-acid R isomer of fenvalerate (Sumicidin ) in an aquatic model ecosystem. J Pestic Sci 5: 11–22. Ohyama T, Jin K, Katoh Y, Chiba Y, Inoue K (1987) Fate and behavior of herbicides, butachlor, CNP, chlomethoxynil and simetryn in river water, shellfish and sediments of the Ishikari River. Bull Environ Contam Toxicol 39: 555–562. O’Kelley JC, Deason TR (1976) Degradation of pesticides by algae. EPA-600/3-76-022, PB 251933, Office of Research and Development, US Environmental Protection Agency, Washington, DC, pp 41. Oliver BG (1984) Uptake of chlorinated organics from anthropogenically contaminated sediments by oligochaete worms. Can J Fish Aquat Sci 41: 878–883. Oliver BG (1987) Biouptake of chlorinated hydrocarbons from laboratory-spiked and field sediments by oligochaete worms. Environ Sci Technol 21: 785–790. Omura T, Sato R (1964) The carbon-monoide-binding pigment of liver microsomes. I. Exidence for its hemoprotein nature. J Biol Chem 239: 2370–2378. Opperhulzen A, Serné P, Van der Steen JMD (1988) Thermodynamics of fish/water and octa-1-ol/water partitioning of some chlorinated benzenes. Environ Sci Technol 22: 286–292. Opperhulzen A, Velde Ewvd, Gobas FAPC, Leim DAK, Steen JMDvd, Hutzinger O (1985) Relationship between bioconcentration in fish and steric factors of hydrophobic chemicals. Chemosphere 14: 1871–1896. Ortíz RG, Martínez-Tabche L, Terrón SO (2004) Effects of cimetidine and phenobarbital on methyl parathion metabolism in Hyalella azteca. Bull Environ Contam Toxicol 72: 1247–1252. Oshima Y, Kobayashi K, Hidaka C, Izu S, Imada N (1994) Differences in the drug-metabolizing enzyme activities among fish and bivalves living in waters near industrial and non-industrial areas. Bull Environ Contam Toxicol 53: 106–112.

124

T. Katagi

Ozreti´c B, Krajnovi´c-Ozreti´c M (1992) Esterase heterogeneity in mussel Mytilus galloprovincialis: Effects of organophosphate and carbamate insecticides in vitro. Comp Biochem Physiol 103C: 221–225. Paris DF, Lewis DL (1976) Accumulation of methoxychlor by microorganisms isolated from aqueous systems. Bull Environ Contam Toxicol 15: 24–32. Pascal-Lorber S, Rathahao E, Cravedi JP, Laurent F (2004) Metabolic fate of [14 C]-2,4dichlorophenol in macrophytes. Chemosphere 56: 275–284. Patil KC, Matsumura F, Boush GM (1972) Metabolic transformation of DDT, dieldrin, aldrin and endrin by marine microorganisms. Environ Sci Technol 6: 629–632. Pavlostathis SG, Comstock KK, Jacobson ME, Saunders FM (1998) Transformation of 2,4,6tronitrotoluene by the aquatic plant Myriophyllum spicatum. Environ Toxicol Chem 17: 2266–2273. Pavlostathis SG, Jackson GH (1999) Biotransformation of 2,4,6-trinitrotoluene in Anabaena sp. cultures. Environ Toxicol Chem 18: 412–419. Peters LD, Nasci C, Livingstone DR (1998) Immunochemical investigations of cytochrome P450 forms/epitopes (CYP1A, 2B, 2E, 3A and 4A) in digestive gland of Mytilus sp. Comp Biochem Physiol 121C: 361–369. Petrocelli SR, Hanks AR, Andersen J (1973) Uptake and accumulation of an organochlorine insecticide (dieldrin) by an estuarine mollusc, Rangia cuneata. Bull Environ Contam Toxicol 10: 315–320. Petroutsos D, Wang J, Katapodis P, Kekos D, Sommerfeld M, Hu Q (2007) Toxicity and metabolism of p-chlorophenol in the marine microalga Tetraselmis marina. Aquat Toxicol 85: 192–201. Petushok N, Gabryelak T, Patecz D, Zavodnik L, Szollosi Varga I, Deér KA (2002) Comparative study of the xenobiotic metabolizing system in the digestive gland of the bivalve molluscs in different aquatic ecosyetms and in aquaria experiments. Aquat Toxicol 61: 65–72. Pflugmacher S, Sandermann Jr H (1998a) Taxonomic distribution of plant glucosyltransferses activity on xenobiotics. Phytochemistry 49: 507–511. Pflugmacher S, Sandermann Jr H (1998b) Cytochrome P450 monooxygenases for fatty acid and xenobiotics in marine macroalgae. Plant Physiol 117: 123–128. Pflugmacher S, Schröder P, Sandermann Jr H (2000) Taxonomic distribution of plant glutathione S-transferases acting on xenobiotics. Phytochem 54: 267–273. Pflugmacher S, Steinberg C (1997) Activity of phase I and phase II detoxication enzymes in aquatic macrophytes. Angew Bot 71: 144–146. Pflugmacher S, Wiencke C, Sandermann Jr H (1999) Activity of phase I and phase II detoxication enzymes in Antarctic and Arctic macroalgae. Mar Environ Res 48: 23–36. Pietsch C, Krause E, Burnison BK, Steinberg CEW, Pflugmacher S (2006) Effects and metabolism of the phenylurea herbicide isoproturon in the submerged macrophyte Ceratophyllum demersum L. J Appl Bot Food Qual 80: 25–30. Pillai MKK, Mittal PK, Agarwal HC (1980) Bioaccumulation, metabolism & elimination of DDT by the freshwater clam Indonaia caerulea (Lea). Indian J Exp Biol 18: 1439–1442. Pinto G, Pollio A, Previtera L, Temussi F (2002) Biodegradation of phenols by microalgae. Biotechnol Lett 24: 2047–2051. Pirini M, Trigari G, Manuzzi MP, Ventrella V, Pagliarani A, Trombetti F, Borgatti AR (2000) Seasonal changes in the lipid pattern of Mytilus galloprovincialis and evaluation of nutritional quality. Italian J Biochem 49: 99–100. Pollio A, Pinto G, Greca MD, De Maio A, Fiorentino A, Previtera L (1994) Progesterone bioconversion by microalgal cultures. Phytochem 37: 1269–1272. Pridham JB (1964) The phenol glucosylation reaction in the plant kingdom. Phytochemistry 3: 493–497. Quigley MA, Cavaletto JF, Gardner WS (1989) Lipid composition related to size and maturity of the amphipod Pontoporeia hoyi. J Great Lakes Res 15: 601–610.

Bioconcentration, Bioaccumulation, and Metabolism

125

Rajendran N, Venugopalan VK (1991) Bioconcentration of endosulfan in different body tissues of estuarine organisms under sublethal exposure. Bull Environ Contam Toxicol 46: 151–158. Rakotondravelo ML, Anderson TD, Charlton RE, Zhu KY (2006) Sublethal effects of three pesticides on activities of selected target and detoxification enzymes in the aquatic midge, Chironomus tentans (Diptera: Chironomidae). Arch Environ Contam Toxicol 51: 360–366. Rao VVSN, Lal R (1987) Uptake and metabolism of insecticides by blue-green algae Anabaena and Aulosira fertilissima. Microbios Lett 36: 143–147. Rathore DS, Kumar A, Kumar HD (1993) Lipid profiles of three microalgae. Phykos 32: 1–8. Ravi V, Balakumar T (1998) Biodegradation of the C–P bond in glyphosate by the cyanobacterium Anabaena variabilis L. J Sci Ind Res 57: 790–794. Reichert WL, Le Eberhart BT, Varanasi U (1985) Exposure of two species of deposit-feeding amphipods to sediment-associated [3 H]benzo[a]pyrene: Uptake, metabolism and covalent binding to tissue macromolecules. Aquat Toxicol 6: 45–56. Reinert RE (1972) Accumulation of dieldrin in an alga (Scenedesmus obliquus), Daphnia magna and the guppy (Poecilia reticulate). J Fish Res Bd Canada 29: 1413–1418. Renberg L, Tarkpea M, Lindén E (1978) The use of the bivalve Mytilus edulis as a test organism for bioconcentration studies. I. Designing a continuous-flow system and its application to some organochlorine compounds. Ecotoxicol Environ Saf 9: 171–178. Rewitz KF, Styrishave B, Løbner-Olesen, Andersen O (2006) Marine invertebrate cytochrome P450: Emerging insights from vertebrate and insect analogies. Comp Biochem Physiol 143C: 363–381. Rice CP, Sikka HC (1973) Uptake and metabolism of DDT by six species of marine algae. J Agric Food Chem 21: 148–152. Richardson GM, Qadri SU, Jessiman B (1983) Acute toxicity, uptake and clearance of aminocarb by the aquatic isopod, Caecidolea racovitzai racovitzai. Ecotoxicol Environ Saf 7: 552–557. Richter S, Nagel R (2007) Bioconcentration, biomagnification and metabolism of 14 C-terbutryn and 14 C-benzo[a]pyrene in Gammarus fossarum and Asellus aquaticus. Chemosphere 66: 603–610. Roghair CJ, Buijze A, Yedema E, Hermans JLM (1992) Toxicity and toxicokinetics for benthic organisms: I: Toxicity and BCF values of chlorobenzenes to Chironomus riparius in water only experiments. RIVM Report 719101002, Rijksinstituut voor Volksgezondheid en Milieu RIVM, http://www.rivm.nl/bibliotheek/rapporten/719101002.html . Rose FL, McIntire CD (1970) Accumulation of dieldrin by benthic algae in laboratory streams. Hydrobiologia 35: 481–493. Roy S, Hänninen O (1994) Pentachlorophenol: Uptake/elimination kinetics and metabolism in an aquatic plant, Eichhornia crassipes. Environ Toxicol Chem 13: 763–773. Rust AJ, Burgess RM, Brownawell BJ, McRlroy AE (2004) Relationship between metabolism and bioaccumulation of benzo[a]pyrene in benthic invertebrates. Environ Toxicol Chem 23: 2587–2593. Sabali¯unas D, Lazutka J, Sabali¯uniene I, Södergren A (1998) Use of semipermeable membrane devices for studying effects of organic pollutants: Comparison of pesticide uptake by semipermeable membrane devices and mussels. Environ Toxicol Chem 17: 1815–1824. Safonova E, Kvitko K, Kuschk P, Möder M, Reisser W (2005) Biodegradation of phenanthrene by the green alga Scenedesmus obliquus ES-55. Eng Life Sci 5: 234–239. Sanborn HR, Malins DC (1980) The disposition of aromatic hydrocarbons in adult spot shrimp (Pandalus platyceros) and the formation of metabolites of naphthalene in adult and larval spot shrimp. Xenobiotica 10: 193–200. Sanborn JR, Metcalf RL, Bruce WN, Lu PY (1976) The fate of chlordane and toxaphene in a terrestrial-aquatic model ecosystem. Environ Entmol 5: 533–538. Sanborn JR, Yu CC (1973) The fate of dieldrin in a small ecosystem. Bull Environ Contam Toxicol 10: 340–346. Sanders HO, Hunn JB (1982) Toxicity, bioconcentration and depuration of the herbicide Bolero 8EC in freshwater invertebrates and fish. Bull Jpn Soc Sci Fish 48: 1139–1143.

126

T. Katagi

Sathe MC, Shrotri RV, Raghu K, Murthy NBK (2005) Observation on accumulation and depuration of fenthion in various tissues of marine edible clam, Marcia haintina (Lamarck). J Ecophysiol Occup Health 5: 33–36. Saxena DM, Lal R, Reddy BVP (1982) DDT uptake and metabolism in Blepharisma intermedium. Acta Protozool 21: 173–175. Schauberger CW, Wildman RB (1977) Accumulation of aldrin and dieldrin by blue-green algae and related effects on photosynthetic pigments. Bull Environ Contam Toxicol 17: 534–541. Schell Jr JD, James MO (1989) Glucose and sulfate conjugation of phenolic compounds by the spiny lobster (Panulirus argus). J Biochem Toxicol 4: 133–138. Schimmel SC, Garnas RL, Patrick Jr JM, Moore JC (1983) Acute toxicity, bioconcentration, and persistence of AC 222,705, benthiocarb, chlorpyrifos, fenvalerate, methyl parathion, and permethrin in the estuarine environment. J Agric Food Chem 31: 104–113. Schimmel SC, Patrick Jr JM, Forester J (1977) Uptake and toxicity of toxaphene in several estuarine organisms. Arch Environ Contam Toxicol 5: 353–367. Schlenk D (2005) Pesticide biotransformation in fish. In: Biochemistry and Molecular Biology of Fishes. Environmental Toxicology. Mommsen TP, Moon TW (eds) Elsevier B.V., Amsterdam vol 6, Chapter 6, pp 171–190. Schlenk D, Buhler DR (1988) Cytochrome P450 and phase II activities in the gumboot chiton Cryptochiton stelleri. Aquat Toxicol 13: 167–182. Schlenk D, Buhler DR (1989a) Determination of multiple forms of cytochrome P-450 in microsomes from the digestive gland of Cryptochiton stelleri. Biochem Biophys Res Commun 163: 476–480. Schlenk D, Buhler DR (1989b) Xenobiotic biotransformation in the Pacific oyster (Crassostrea Gigas). Comp Biochem Physiol 94C: 469–475. Schmidt K, Pflugmacher S, Staaks GBO, Steunberg CEW (2006) The influence of tributyltin chloride and polychlorinated biphenylas on swimming behavior, body growth, reproduction, and activity of biotransformation enzymes in Daphnia magna. J Freshwater Ecol 21: 109–120. Schoeny R, Cody T, Warshawsky D, Radike M (1988) Metabolism of mutagenic polycyclic aromatic hydrocarbons by photosynthetic algal species. Mutat Res 197: 289–302. Schooley DA, Quinstad GB (1979) Metabolism of insect growth regulators in aquatic organisms. In: Pesticide and Xenobiotic Metabolism in Aquatic Organisms. Khan MAQ, Lech JJ, Menn JJ (eds) ACS Symp Ser 99, American Chemical Society, Washington, DC, Chapter 10, pp 161–176. Schramm KW, Behechti A, Beck B, Kettrup A (1998) Influence of an aquatic humic acid on the bioconcentration of selected compounds in Daphnia magna. Ecotoxicol Environ Saf 41: 73–76. Schrenk C, Pflugmacher S, Brüggemann R, Sandermann Jr H, Steinberg CEW, Kettrup A (1998) Glutathione S-transferase activity in aquatic macrophytes with emphasis on habitat dependence. Ecotoxicol Environ Saf 40: 226–233. Schuler LJ, Wheeler M, Bailer AJ, Lydy MJ (2003) Toxicokinetics of sediment-sorbed benzo[a]pyrene and hexachlorobiphenyl using the freshwater invertebrates Hyalella azteca, Chironomus tentans and Lumbriculus variegatus. Environ Toxicol Chem 22: 439–449. Schuphan I (1974) Zum metabolismus von phenylharnstoffen. III. Metabolismus von monolinuronO-methyl 14 C in Chlorella pyrenodoisa. Chemosphere 3: 131–134. Schuth CK, Isensee AR, Woolson EA, Kearney PC (1974) Distribution of 14 C and arsenic derived from [14 C]cacodylic acid in an aquatic ecosystem. J Agric Food Chem 22: 999–1003. Schuytema GS, Krawczyk DF, Griffis WL, Nebeker AV, Robideaux ML (1990) Hexachlorobenzene uptake by fathead minnows and macroinvertebrates in recirculating sediment/water systems. Arch Environ Contam Toxicol 19: 1–9. Schuytema GS, Krawczyk DF, Griffis WL, Nebeker AV, Robideaux ML, Brownawell BJ, Westall JC (1988) Comparative uptake of hexachlorobenzene by fathead minnows, amphipods and oligochaete worms from water and sediment. Environ Toxicol Chem 7: 1035–1045. Semple KT, Cain RB (1996) Biodegradation of phenols by the alga Ochromonas danica. Arch Environ Microbiol 62: 1265–1273.

Bioconcentration, Bioaccumulation, and Metabolism

127

Semple KT, Cain RB, Schmidt S (1999) Biodegradation of aromatic compounds by microalgae. FEMS Microbiol Lett 170: 291–300. Serrano R, Hernández F, López FJ, Peña JB (1997a) Bioconcentration and depuration of chlorpyrifos in the marine mollusk Mytilus edulis. Arch Environ Contam Toxicol 33: 47–52. Serrano R, Hernández F, Peña JB, Dosda V, Canales J (1995) Toxicity and bioconcentration of selected organophosphorus pesticides in Mytilus galloprovincialis and Venus gallina. Arch Environ Contam Toxicol 29: 284–290. Serrano R, López FJ, Hernández F, Peña JB (1997b) Bioconcentration of chlorpyrifos, chlorfenvinphos and methidathion in Mytilus galloprovincialis. Bull Environ Contam Toxicol 59: 968–975. Serrano R, López FJ, Roig-Navarro A, Hernández F (1997c) Automated sample clean-up and fractionation of chlorpyrifos, chlorpyrifos-methyl and metabolites in mussels using normalphase liquid chromatography. J Chromatogr A 778: 151–160. Sethunathan N, Megharaj M, Chen ZL, Williams BD, Lewis G, Naidu R (2004) Algal degradation of a known endocrine disrupting insecticide α-endosulfan, and its metabolite, endosulfan sulfate, in liquid medium and soil. J Agric Food Chem 52: 3030–3035. Sharma HA, Barber JT, Ensley HE, Polito MA (1997) A comparison of the toxicity and metabolism of phenol and chlorinated phenols by Lemna gibba, with special reference to 2,4,5-trichlorophenol. Environ Toxicol Chem 16: 346–350. Shaw GR, Connell DW (1984) Physicochemical properties controlling polychlorinated biphenyl (PCB) concentrations in aquatic organisms. Environ Sci Technol 18: 18–23. Shaw GR, Connell DW (1987) Comparative kinetics for bioaccumulation of polychlorinated biphenyls by the polychaete (Capitella capitata) and fish (Mugil cephalus). Ecotoxicol Environ Saf 13: 84–91. Shaw B, Hopke PK (1975) The dynamics of diquat in a model eco-system. Environ Lett 8: 325–335. Shofer SL, Tjeerdema RS (1993) Comparative disposition and biotransformation of pentachlorophenol in the oyster (Crassostrea gigas) and abalone (Haliotis fulgens). Pestic Biochem Physiol 46: 85–95. Sikka HC, Lynch RS, Lindenberger M (1974) Uptake and metabolism of dichlobenil by emersed aquatic plants. J Agric Food Chem 22: 230–234. Sims JG, Steevens JA (2008) The role of metabolism in the toxicity of 2,4,6-trinitrotoluene and its degradation products to the aquatic amphipod Hyalella Azteca. Ecotoxicol Environ Saf 70: 38–46. Skoglund RS, Stange K, Swackhamer DL (1996) A kinetics model for predicting the accumulation of PCBs in phytoplankton. Environ Sci Technol 30: 2113–2120. Snyder MJ (2000) Cytochrome P450 enzymes in aquatic invertebrates: Recent advances and future directions. Aquat Toxicol 48: 529–547. Södergren A, Svensson B (1973) Uptake and accumulation of DDT and PCB by Ephemera danica (Ephemeroptera) in continuous-flow systems. Bull Environ Contam Toxicol 9: 345–350. Soeder CJ, Hegewald E, Kneifel H (1987) Green microalgae can use naphthalenesulfonic acid as sources of sulfur. Arch Microbiol 148: 260–263. Solé M, Livingstone DR (2005) Components of the cytochrome P450-dependent monooxygenase system and “‘NADPH-independent benzo[a]pyrene hydroxylase” activity in a wide range of marine invertebrate species. Comp Biochem Physiol 141C: 20–31. Southworth GR, Beauchamp JJ, Schmieder PK (1978) Bioaccumulation potential of polycyclic aromatic hydrocarbons in Daphnia pulex. Water Res 12: 973–977. Spacie A, Hamelink JL (1982) Alternative models for describing the bioconcentration of organics in fish. Environ Toxicol Chem 1: 309–320. Spehar RL, Tanner DK, Gibson JH (1982) Effects of kelthane and pydrin on early life stage of fathead minnows (Pimephales promelas) and amphipods (Hyalella azteca). In: Aquatic Toxicology and Hazard Assessment: Fifth Conference, Pearson RB, Foster RB, Bishop WE (eds) ASTM STP 766, ASTM, West Conshohocken, pp 234–244.

128

T. Katagi

Spehar RL, Tanner DK, Nordling BR (1983) Toxicity of the synthetic pyrethroids, permethrin and AC 222,705 and their accumulation in early life stages of fathead minnows and snails. Aquat Toxicol 3: 171–182. Standley LJ (1997) Effect of sedimentary organic matter composition on the partitioning and bioavailability of dieldrin to the oligochaete Lumbriculus variegates. Environ Sci Technol 31: 2577–2583. Stange K, Swackhamer DL (1994) Factors affecting phytoplankton species-specific differences in accumulation of 40 polychlorinated biphenyls (PCBs). Environ Toxicol Chem 13: 1849–1860. Stehly GR, Landrum PF, Henry MG, Klemm C (1990) Toxicokinetics of PAHs in Hexageniza. Environ Toxicol Chem 9: 169–174. Stenersen J, Kobro S, Bjerke M, Arend U (1987) Glutathione transferases in aquatic and terrestrial animals from nine phyla. Comp Biochem Physiol 86C: 73–82. Stief P, Eller G (2006) The gut microenvironment of sediment-dwelling Chironomus plumosus larvae as characterized with O2 , pH, and redox microsensors. J Comp Physiol B 176: 673–683. Sturm A, Hansen PD (1999) Altered cholinesterase and monooxygenase levels in Daphnia magna and Chironomus riparius exposed to environmental pollutants. Ecotoxicol Environ Saf 42: 9–15. Subramanian G, Sekar S, Sampoornam S (1994) Biodegradation and utilization of organophosphorus pesticides by cyanobacteria. Int Biodeterior Biodegradation 33: 129–143. Sukhija PS, Singh M, Bhatia IS (1979) Lipid composition and fatty acid synthesis in the blue green alga Anabaena doliolum. Biochem Physiol Pflanzen 174: 685–690. Sundaram KMS (1995) Distribution, persistence, and fate of mexacarbate in the aquatic environment of a mixed-wood boreal forest. J Environ Sci Health B30: 651–683. Sundaram KMS, Szeto SY (1979) A study on the lethal toxicity of aminocarb to freshwater crayfish and its in vitro metabolism. J Environ Sci Health B14: 589–602. Suora KE, Fisher SW (1986) Effect of pH on the environmental fate of [14 C] aldicarb in an aquatic microcosm. Ecotoxicol Environ Saf 11: 81–90. Sushchick NN, Gladyshev MI, Moskvichova AV, Makhutova ON, Kalachova GS (2003) Comparison of fatty acid composition in major lipid classes of the dominant benthic invertebrates of the Yenisei river. Comp Biochem Physiol B 134: 111–122. Swackhamer DL, Skoglund RS (1993) Bioaccumulation of PCBs by algae: Kinetics versus equilibrium. Environ Toxicol Chem 12: 831–838. Sweeney RA (1969) Metabolism of lindane by unicellular algae. Proceedings of the 12th Conference on Great Lakes Research, International Associationforf Great Lakes Research, pp 98–102. Takimoto Y, Ohshima M, Miyamoto J (1987a) Comparative metabolism of fenitrothion in aquatic organisms. II. Metabolism in the freshwater snails, Cipangopaludina japonica and Physa acuta. Ecotoxicol Environ Saf 13: 118–125. Takimoto Y, Ohshima M, Miyamoto J (1987b) Comparative metabolism of fenitrothion in aquatic organisms. III. Metabolism in the crustaceans, Daphnia pulex and Palaemon paucidens. Ecotoxicol Environ Saf 13: 126–134. Tang JX, Hoagland KD, Siegfried BD (1998a) Uptake and bioaccumulation of atrazine by selected freshwater algae. Environ Toxicol Chem 17: 1085–1090. Tang JX, Siegfried BD (1996) Bioconcentration and uptake of a pyrethroid and organophosphate insecticide by selected aquatic insects. Bull Environ Contam Toxicol 57: 993–998. Tang JX, Siegfried BD, Hoagland KD (1998b) Glutathione-S-transferase and in vitro metabolism of atrazine in freshwater algae. Pestic Biochem Physiol 59: 155–161. Tantawy MM, Braumann T, Grimme LH (1984) Uptake and metabolism of the phenylpyridazinone herbicide metflurazon during the bleaching and regeneration process of the green alga, Chlorella fusca. Pestic Biochem Physiol 22: 224–231. Tatsuzawa H, Takizawa E, Wada M, Yamamoto Y (1996) Fatty acid and lipid composition of the acidophilic green alga Chlamydimonas sp. J Phycol 32: 598–601.

Bioconcentration, Bioaccumulation, and Metabolism

129

Teisseire H, Vernet G (2001) Effects of the fungicide folpet on the activities of antioxidative enzymes in duckweed (Lemna minor). Pestic Biochem Physiol 69: 112–117. Thies F, Backhaus T, Bossmann B, Grimme LH (1996) Xenobiotic biotransformation in unicellular green algae. Plant Physiol 112: 361–370. Thies F, Grimme LH (1995) O-dealkylation of coumarin and resorufin ethers by unicellular green algae: Kinetic properties of Chlorella fusca and Chlorella sorokiniana. Arch Microbiol 164: 203–211. Thomann RV, Komlos J (1999) Model of biota-sediment accumulation factor for polycyclic aromatic hydrocarbons. Environ Toxicol Chem 18: 1060–1068. Thomas TM, Seaman DE (1968) Translocation studies with endothal-14 C in Potamogeton nodosus Poir. Weed Res 8: 321–326. Thompson Jr GA (1996) Lipids and membrane function in green algae. Biochim Biophys Acta 1302: 17–45. Thompson HM (1999) Esterases as markers of exposure to organophosphates and carbamates. Ecotoxicology 8: 369–384. Thybaud E, Caquet T (1991) Uptake and elimination of lindane by Lymnaea palustris (Mollusca: gastropoda): A pharmacokinetic approach. Ecotoxicol Environ Saf 21: 365–376. Thybaud E, Le Bras S (1988) Absorption and elimination of lindane by Asellus aquaticus (Crustacea, Isopoda). Bull Environ Contam Toxicol 40: 731–735. Tjeerdema RS, Crosby DG (1992) Disposition and biotransformation of pentachlorophenol in the red abalone (Haliotis rufescens). Xenobiotica 22: 681–690. Todd SJ, Cain RB, Schmidt S (2002) Biotransformation of naphthalene and diaryl ethers by green algae. Biodegradation 13: 229–238. Tront JM, Saunders FM (2006) Role of plant activity and contaminant speciation in aquatic plant assimilation of 2,4,5-trichlorophenol. Chemosphere 64: 400–407. Tront JM, Saunders FM (2007) Sequestrations of a fluorinated analog of 2,4-dichlorophenol and metabolic products by L. minor as evidenced by 19 F NMR. Environ Pollut 145: 708–714. Tsorbatzoudi E, Vockel D, Korte F (1976) Beiträge zur ökologischer Chemie Cx1. Metabolismus von buturon-14 C in algen. Chemosphere 5: 49–52. Tsuge S, Kazano H, Tomizawa C (1976) Some devices in a model ecosystem for volatile compounds and its application to carbaryl and p,p -DDT. J Pestic Sci 1: 307–311. Turgut C (2005) Uptake and modeling of pesticides by roots and shoots of parrot feather (Myriophyllum aquaticum). Environ Sci Pollut Res Int 12: 342–346. Tweedy BG, Loeppky C, Ross JA (1970) Metabolism of 3-(p-bromophenyl)-1-methylurea (metobromuron) by selected soil microorganisms. J Agric Food Chem 18: 851–853. Ueda T, Sadakane S, Yamaoka K, Nakagawa M, Ishida M (1988) Bioaccumulation and metabolic fate of pyrazolate and its hydrolysates in guppies and short-necked clams. J Pestic Sci 13: 85–92. Ullrich WR, Ullrich-Eberius CI, Köcher H (1990) Uptake of glufosinate and concomitant membrane potential changes in Lemna gibba G1. Pestic Biochem Physiol 37: 1–11. Uno S, Shiraishi H, Hatakeyama S, Otsuki A (1997) Uptake and depuration kinetics and BCFs of several pesticides in three species of shellfish (Corbicula leana, Corbicula japonica and Cipangopludina chinensis): Comparison between field and laboratory experiment. Aquat Toxicol 39: 23–43. Uno S, Shiraishi H, Hatakeyama S, Otsuki A, Koyama J (2001) Accumulative characteristics of pesticide residues in organs of bivalves (Anodonta woodiana and Corbicula leana) under natural conditions. Arch Environ Contam Toxicol 40: 35–47. USEPA (2007) ECOTOX User Guide: ECOTOXicology Database System. Version 4.0. United States Environmental Protection Agency, Washington, DC, USA. http:/www.epa.gov/ecotox/. R USEPA (2008) Estimation Programs Interface SuiteTM for Microsoft Windows, v3.20. United States Environmental Protection Agency, Washington, DC, USA. http://www.epa.gov/oppt/ exposure/pubs/episuite.htm.

130

T. Katagi

Väisänen MVT, Mackenzie PI, Hänninen OOP (1983) UDP glucosyltransferase and its kinetic fluorimetric assay. Eur J Biochem 130: 141–145. Valentine JP, Bingham SW (1974) Influence of several algae on 2,4-D residues in water. Weed Sci 22: 358–363. Van der Linde A, Hendriks AJ, Sijm DTHM (2001) Estimating biotransformation rate constants of organic chemicals from modeled and measured elimination rates. Chemosphere 44: 423–435. Van Hattum B, Cid Montanes JF (1999) Toxicokinetics and bioconcentration of polycyclic aromatic hydrocarbons in freshwater isopods. Environ Sci Technol 33: 2409–2417. Van Hattum B, Curto MJ, Cid Montanes JF (1998) Polycyclic aromatic hydrocarbons in freshwater isopods and field-partitioning between abiotic phases. Arch Environ Contam Toxicol 35: 257–267. Vance BD, Drummond W (1969) Biological concentration of pesticides by algae. J Am Water Works Assoc 61: 360–362. Vance BD, Maki AW (1976) Bioconcentration of dibrom by Stigeoclonium pachydermum. Bull Environ Contam Toxicol 15: 601–607. Vanderford M, Shaanks JV, Hughes JB (1997) Phytotransformation of trinitrotoluene (TNT) and distribution of metabolic products in Myriophyllum aquaticum. Biotechnol Lett 19: 277–280. Varanasi U, Reichert WL, Stein JE, Brown DW, Sanborn HR (1985) Bioavailability and biotransformation of aromatic hydrocarbons in benthic organisms exposed to sediment from an urban estuary. Environ Sci Technol 19: 836–841. Varó I, Serrano R, Pitarch E, Amat F, López FJ, Navarro JC (2000) Toxicity and bioconcentration of chlorpyrifos in aquatic organisms: Artemia parthenogenetica (Crustacea), Gambusia affinis and Aphanius iberus (Pisces). Bull Environ Contam Toxicol 65: 623–630. Veith GC, DeFoe DL, Bergstedt BV (1979) Measuring and estimating the bioconcentration factor of chemicals in fish. J Fish Res Board Can 36: 1040–1048 Vidal ML, Narbonne JF (2000) Characterization of glutathione S-transferase activity in the Asiatic clam Corbicula fluminea. Bull Environ Contam Toxicol 64: 455–462. Vidal ML, Rouimi P, Debrauwer L, Narbonne JF (2002) Purification and characterization of glutathione S-transferases from the freshwater clam Corbicula fluminea (Müller). Comp Biochem Physiol 131C: 477–489. Vioque-Fernández A, Alves de Almeida E, López-Barea J (2007) Esterases as pesticide biomarkers in crayfish (Procambarus clarkii, Crustacea): Tissue distribution, sensitivity to model compound and recovery from inactivation. Comp Biochem Physiol 145C: 404–412. Virtanen MT, Hattula ML (1982) The fate of 2,4,6-trichlorophenol in an aquatic continuous-flow system. Chemosphere 11: 641–649. Vose JR, Cheng JY, Antia NJ, Towers HN (1971) The catabolic fission of the aromatic ring of phenylalanine by marine planktonic algae. Can J Bot 49: 259–261. Wandiga SO, Ongeri DMK, Mbuvi L, Lalah JO, Jumba IO (2002) Accumulation, distribution of 14 C-1,1,1-trichloro-2,2-bis-(p-chlorophenyl)ethane (p,p -DDT) residues in a model tropical marine ecosystem. Environ Technol 23: 1285–1292. Wang JS, Simpson KL (1996) Accumulation and depuration of DDTs in the food chain from Artemia to brook trout (Salvelinus fontinalis). Bull Environ Contam Toxicol 56: 888–895. Wang WH, Lay JP (1989) Fate and effects of salicylic acid compounds in freshwater systems. Ecotoxicol Environ Saf 17: 308–316. Wang X, Harada S, Watanabe M, Kashikawa H, Geyer HJ (1996) Modelling the bioconcentration of hydrophobic organic chemicals in aquatic organisms. Chemosphere 32: 1783–1793. Wang YS, Hwang KL, Hsieh YN, Chen YL (1992a) Fate of the herbicide naproanilide in a rice paddy model ecosystem. J Pestic Sci 17: 161–167. Wang YS, Jaw CG, Tang HC, Lin TS, Chen YL (1992b) Accumulation and release of herbicides butachlor, thiobencarb and chlomethoxyfen by fish, clam and shrimp. Bull Environ Contam Toxicol 48: 474–480. Warner NA, Wong CS (2006) The freshwater invertebrate Mysis relicta can eliminate chiral organochlorine compounds enantioselectively. Environ Sci Technol 40: 4158–4164.

Bioconcentration, Bioaccumulation, and Metabolism

131

Warshawsky D, Keenan TH, Reilman R, Cody TE, Radike MJ (1990) Conjugation of benzo[a]pyrene by freshwater green alga Selenastrum capricornutum. Chem-Biol Interact 74: 93–105. Warshawsky D, Radike MJ, Jayasimhulu K, Cody T (1988) Metabolism of benzo[a]pyrene by a dioxygenase system of the freshwater green alga Selenastrum capricornutum. Biochem Biophys Res Commun 152: 540–544. Watanabe S, Watanabe S, Ito K (1985) Accumulation and excretion of herbicides in various tissues of mussel. J Food Hyg Soc Jpn 26: 496–499. Watanabe S, Watanabe S, Ito K (1987) Reduction metabolites of the herbicide chlornitrofen (CNP) in mussel. J Food Hyg Soc Jpn 28: 277–280. Watanabe Y (2000) High-valent intermediates. In: The Porphyrin Handbook. Biochemistry and Binding: Activation of Small Molecules. Kadish KM, Smith KM, Guilard R (eds) Academic Press, New York, vol 4, Chapter 30, pp 97–117. Weinberger P, Greenhalgh R (1985) The sorptive capacity of an aquatic macrophyte for the pesticide aminocarb. J Environ Sci Health B 20: 263–273. Weinberger P, Greenhalgh R, Moody RP, Boulton B (1982) Fate of fenitrothion in aquatic microcosms and the role of aquatic plants. Environ Sci Technol 16: 470–473. Weiner JA, DeLorenzo ME, Fulton MH (2004) Relationship between uptake capacity and differential toxicity of the herbicide atrazine in selected microalgal species. Aquat Toxicol 68: 121–128. Wheelock CE, Shan G, Ottea J (2005) Overview of carboxylesterases and their role in the metabolism of insecticides. J Pestic Sci 30: 75–83. Whitten BK, Goodnight CJ (1966) The comparative chemical composition of two aquatic oligochaetes. Comp Biochem Physiol 17: 1205–1207. Wiegand C, Pehkonen S, Akkanen J, Penttinen OP, Kukkonen JVK (2007) Bioaccumulation of paraquat by Lumbriculus variegatus in the presence of dissolved natural organic matter and impact on energy costs, biotransformation and antioxidative enzymes. Chemosphere 66: 558–566. Wilbrink M, Groot EJ, Jansen R, de Vries Y, Vermeulen NPE (1991b) Occurrence of a cytochrome P-450-containing mixed-function oxidase system in the pond snail, Lymnaea stagnalis. Xenobiotica 21: 223–233. Wilbrink M, van de Merbel NC, Vermeulen NP (1991a) Glutathione-S-transferase activity in the digestive gland of the pond snail, Lymnaea stagnalis. Comp Biochem Physiol 99C: 185–189. Wildi E, Nagel R, Steinberg CEW (1994) Effects of pH on the bioconcentration of pyrene in the larval midge, Chironomus riparius. Water Res 28: 2553–2559. Williamson KC, Shofer SL, Tjeerdema RS (1995) Toxicokinetics and biotransformation of p-nitrophenol in the black turban snail (Tegula funebralis). Aquat Toxicol 33: 113–123. Wofford HW, Wilsey CD, Neff GS, Giam CS, Neff JM (1981) Bioaccumulation and metabolism of phthalate esters by oysters, brown shrimp, and sheephead minnows. Ecotoxicol Environ Saf 5: 202–210. Wolf SD, Lassiter RR, Wooten SE (1991) Predicting chemical accumulation in shoots of aquatic plants. Environ Toxicol Chem 10: 665–680. Wolfe NL, Hoehamer CF (2003) Enzymes used by plants and microorganisms to detoxify organic compounds. In: Phytoremediation: Transformation and Control of Contaminants. McCutcheon SC, Schnoor JL (eds) John Wiley & Sons, Inc., Chichester, Chapter 5, pp 159–187. Wood BJB (1974) Fatty acids and saponifiable lipids. In: Algal Physiology and Biochemistry. Stewart WDP (ed) Botanical Monographs vol 10, Blackwell Scientific Publications, Oxford, Chapter 8, pp 236–265. Woodburn KB, Hansen SC, Roth GA, Strauss K (2003) The bioconcentration and metabolism of chlorpyrifos by the eastern oyster, Crassostrea virginica. Environ Toxicol Chem 22: 276–284. Wright SJL, Maule A (1982) Transformation of the herbicide propanil and chlorpropham by microalgae. Pestic Sci 13: 253–256.

132

T. Katagi

Wright SJL, Stainthorpe AF, Downs JD (1977) Interactions of the herbicide propanil and a metabolite 3,4-dichloroaniline with blue-green algae. Acta Phytopathol Acad Sci Hung 12: 51–60. Yadav DV, Agarwal HC, Pillai MKK (1978) Uptake, metabolism and excretion of DDT by the fresh water snail, Vivipara heliciformis. Bull Environ Contam Toxicol 15: 300–306. Yamato Y, Kiyonaga M, Watanabe T (1983) Comparative bioaccumulation and elimination of HCH isomers in short-necked clam (Venerupis japonica) and guppy (Poecilia reticulate). Bull Environ Contam Toxicol 31: 352–359. Yang S, Wu RSS, Kong RYC (2002) Biodegradation and enzymatic responses in the marine diatom Skeletonema costaum upon exposure to 2,4-dichlorophenol. Aquat Toxicol 59: 191–200. Yang W, Spurlock F, Liu W, Gan J (2006) Effects of dissolved organic matter on permethrin bioavailability to daphnia species. J Agric Food Chem 54: 3967–3972. Yockim RS, Isensee AR, Jones GE (1978) Distribution and toxicity of TCDD and 2,4,5-T in an aquatic model ecosystem. Chemosphere 7: 215–220. Yockim RS, Isensee AR, Walker EA (1980) Behavior of trifluralin in aquatic model ecosystem. Bull Environ Contam Toxicol 24: 134–141. Yoshida T, Sako Y, Uchida A, Kakutani T, Arakawa O, Noguchi T, Ishida Y (2002) Purification and characterization of sulfotransferase specific to O-22 of 11-hydroxy saxitoxin from the toxic dinoflagellate Gymnodinium catenatum (dinophyceae). Fish Sci 68: 634–642. You J, Landrum PF, Lydy MJ (2006) Comparison of chemical approaches for assessing bioavailability of sediment-associated contaminants. Environ Sci Technol 40: 6348–6353. Yu CC, Booth GM, Hansen DJ, Larsen JR (1974a) Fate of bux insecticide in a model ecosystem. Environ Entomol 3: 975–977. Yu CC, Booth GM, Hansen DJ, Larsen JR (1974b) Fate of carbofuran in a model ecosystem. J Agric Food Chem 22: 431–434. Yu CC, Booth GM, Hansen DJ, Larsen JR (1975b) Fate of pyrazon in a model ecosystem. J Agric Food Chem 23: 309–311. Yu CC, Booth GM, Hansen DJ, Larsen JR (1975c) Fate of alachlor and propachlor in a model ecosystem. J Agric Food Chem 23: 877–879. Yu CC, Booth GM, Hansen DJ, Larsen JR (1975d) Fate of triazine herbicide cyanazine in a model ecosystem. J Agric Food Chem 23: 1014–1015. Yu CC, Hansen DJ, Booth GM (1975a) Fatae of dicamba in a model ecosyetm. Bull Environ Contam Toxicol 13: 280–283. Yu CC, Sanborn JR (1975) The fate of parathion in a model ecosystem. Bull Environ Contam Toxicol 13: 543–550. Yuen WK, Ho JW (2001) Purification and characterization of multiple glutathione S-transferase isozymes from Chironomidae larvae. Comp Biochem Physiol 129A: 631–640. Yu-yun T, Thumm W, Jobelium-Korte M, Attar A, Freitag D, Kettrup A (1993) Fate of two phenylbenzoylurea insecticides in an algae culture system (Scenedesmus subspicatus). Chemosphere 26: 955–962. Zablotowicz RM, Schrader KK, Locks MA (1988) Algal transformation of flumeturon and atrazine by N-dealkylation. J Environ Sci Health B33: 511–528. Zaroogian GE, Heltshe JF, Johnson M (1985) Estimation of bioconcentration in marine species using structure-activity models. Environ Toxicol Chem 4: 3–12.

Electron Transfer as a Potential Cause of Diacetyl Toxicity in Popcorn Lung Disease Peter Kovacic and Andrew L. Cooksy

Contents Introduction . . . . . . . . . . . . . . . . . . . . . Diacetyl Toxicity . . . . . . . . . . . . . . . . . . . 2.1 Electron Transfer . . . . . . . . . . . . . . . . 2.2 Metabolism . . . . . . . . . . . . . . . . . . . 3 A Unifying Mechanism for Pulmonary Toxicity . . . . . 4 Diacetyl: Evidence to Support an ET Mechanism of Action 5 Other Relevant α-Dicarbonyls . . . . . . . . . . . . . 5.1 Ethanol Metabolites . . . . . . . . . . . . . . . 5.2 Bacterial Cell Signaling Agents . . . . . . . . . . 5.3 Other Toxins . . . . . . . . . . . . . . . . . . 5.4 Protein Derivatives . . . . . . . . . . . . . . . 5.5 Advanced Glycation End Products (AGEs) . . . . . 5.6 Other Bioactive α-Dicarbonyls . . . . . . . . . . 6 Other Physiological Effects . . . . . . . . . . . . . . . 7 Summary . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . .

1 2

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

. . . . . . . . . . . . . . . .

133 135 135 136 136 136 140 140 140 142 142 142 143 144 145 146

1 Introduction Bronchiolitis obliterans, a serious lung disease, arises in microwave popcorn plant workers who are exposed to butter-flavored volatiles, particularly diacetyl. Findings indicate that chronic inhalation of butter-flavoring volatiles compromises lung epithelial barrier function (Fedan et al. 2006).

P. Kovacic (B) Department of Chemistry and Biochemistry, San Diego State University, San Diego, CA, 92182-1030, USA e-mail: [email protected] D.M. Whitacre (ed.), Reviews of Environmental Contamination and Toxicology Volume 204, Reviews of Environmental Contamination and Toxicology 204, C Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-4419-1440-8_2, 

133

134

P. Kovacic and A.L. Cooksy

A considerable amount of information is available on the toxicity of artificial butter-flavoring agents and their associated constituents (Kreiss 2007; ILS 2007). One inhalation toxicity study indicated necrosis of tracheal and nasal epithelium. Necropsy results revealed general pulmonary tissue congestion, focal hyperemia, atelectasis, bloody edema, bronchial edema, and altered thorax in rats that expired after 4-h diacetyl exposures. Histopathological examination indicated the presence of emphysema and hyperemia, as well as necrosis in kidney proximal tubules and swelling of hepatocytes. Subacute exposure to diacetyl caused death, necrotizing rhinitis, necrotizing laryngitis, and bronchitis (Morgan et al. 2008). The foregoing pattern of injury replicates features of human injury observed among workers engaged in processing and packaging of popcorn. Inhalation of diacetyl induced epithelial necrosis and inflammation in the nose, larynx, trachea, and bronchi (Hubbs et al. 2008), also produced were edema, hemorrhage, and ultrastructural changes in the tracheal epithelium. Although the threat of lung disease is most serious among plant workers, one investigation suggests that diacetyl, used as a butter substitute in cooking, puts professional cooks at risk as well (Schneider 2007). Thus far, however, few kitchen-related injuries from diacetyl exposure have been identified. One article describes occupational injury from exposure to diacetyl as an avoidable tragedy caused by corporate and regulatory negligence (Egilman et al. 2007). Because considerable evidence is accumulating concerning diacetyl toxicity, major popcorn makers have finally and recently stopped using the compound (Hanson 2009). The preponderance of bioactive substances or their metabolites incorporates ET (electron transfer) functionalities which we believe play an important role in many physiological responses. Some chemical functionalities that are engaged in ET include quinones (or phenolic precursors), metal complexes (or complexors), aromatic nitro compounds (or reduced hydroxylamine and nitroso derivatives), and conjugated imines (or iminium species). In vivo redox cycling with oxygen of substances that retain ET functionalities may occur and, thereby, may produce oxidative stress (OS) through generation of reactive oxygen species (ROS), such as hydrogen peroxide, hydroperoxides, alkyl peroxides, and diverse radicals (hydroxyl, alkoxyl, hydroperoxyl, and superoxide). The mechanism of the reaction that occurs between electrons and oxygen produces superoxide, which acts as a precursor for hydrogen peroxide and other ROS. In some cases, ET results in interference with normal electrical effects, e.g., in respiration or neurochemistry. In addition, ET, ROS, and OS have been increasingly implicated in the mode of action of drugs and toxins. The α-dicarbonyl group of diacetyl is a functional moiety that has ET effects, and Niufar et al. (2002) are among the few authors who have addressed this aspect. On preparing this review, we found that there have been increasing numbers of reports that address the bioactivity of diacetyl and its connection to ET. Therefore, it is our goal in this review to propose and explain a novel mechanism for how diacetyl may express its toxic action, along with the electrochemical, metabolic, and cell signaling evidence that support our thesis.

Electron Transfer as a Potential Cause of Diacetyl Toxicity

135

2 Diacetyl Toxicity 2.1 Electron Transfer The literature is incomplete on the question of how diacetyl manifests its toxic action. Notwithstanding, diacetyl and other α-dicarbonyls have been implicated to act through mechanisms that involve ET and possibly ROS and OS, and, in this section, we will summarize the cogent literature on that topic. Evidence has been presented in various reviews that support involvement of ET-ROS-OS in the toxic action of several categories of chemical substances: therapeutics (Kovacic and Becvar 2000), anticancer agents (Kovacic and Osuna 2000), carcinogens (Kovacic and Jacintho 2001), and toxins (Kovacic et al. 2005; Kovacic and Somanathan 2006a; Kovacic et al. 2002; Poli et al. 1989; Kovacic and Thurn 2005; Kovacic and Somanathan 2005; Kovacic and Cooksy 2005a; Kovacic and Somanathan 2008; Halliwell and Gutteridge 1999). Sometimes, however, no evidence is found for toxic action that derives from participation of ROS or OS. For example, there are no substantial data to support the involvement of ROS in the metabolism of zolpidem (Ambien), although ET may participate (Kovacic and Somanathan 2009a). The toxicity of zolpidem is quite low (Berson et al. 2001), when compared with several other drugs that produce effects through the involvement of ROS (Kovacic et al. 2005; Kovacic and Somanathan 2006a; Kovacic et al. 2002; Poli et al. 1989; Kovacic and Thurn 2005; Kovacic and Somanathan 2005; Kovacic and Cooksy 2005a; Kovacic and Somanathan 2008; Halliwell and Gutteridge 1999). Another feature of zolpidem is that it does not deplete GSH (glutathione), which suggests that the molecule only induces low levels of OS (Berson et al. 2001). Zolpidem may not be alone. There are other drugs that engage in ET reactions, but do so without appreciable formation of ROS. For example, amsacrine, a 9-anilinoacridine derivative and anticancer agent, appears to exert its cytotoxic action by poisoning of cellular topoisomerase enzymes (Baguley et al. 2003). Evidence indicates, however, that the drug may act as an electron donor in ET reactions, perhaps in a way that involves DNA. Other substances, 5,6dimethylxanthenone-4-acetic acid (DMXAA) and flavone-8-acetic acid (FAA), also appear to be capable of ET as pyrylium-type species (Kovacic 2005), but do not produce effects through ROS and OS. DMXAA is an anticancer agent and exhibits tumor antivascular activity. Administration of DMXAA or FAA in cultured marine splenocytes results in the synthesis of tumor necrosis factor (TNF), in addition to various cytokines, chemokines, and transcription factors. Because little is known about the effect of diacetyl on ROS production, it may be that this α-dicarbonyl produces its effects mainly by ET. On the other hand, a recent review provides extensive evidence for the participation of ET-ROS for the vast majority of pulmonary toxins (Kovacic and Somanathan 2009b), which diacetyl certainly is. More work is needed to elucidate the degree to which ROS are involved in diacetyl toxicity.

136

P. Kovacic and A.L. Cooksy

2.2 Metabolism Although metabolites often play important roles in therapeutic action and toxicity, evidence suggests that, in the case of diacetyl, the parent toxin is the active agent. In rats, labeled diacetyl was rapidly metabolized and excreted, mainly as carbon dioxide (ILS Inc. 2007). Acetoin may also play a role in lung toxicity as a precursor of diacetyl. Metabolism was mainly by oxidation to diketone at low concentrations, but by reduction to diol at higher levels. Alternatively, since carbonyls condense readily with protein primary amino acid groups, the imine derivative may be physiologically active, as discussed below.

3 A Unifying Mechanism for Pulmonary Toxicity The lung is a major target for toxicity. The atmosphere is replete with natural and manmade agents that may be toxic by the inhalation route. Some prominent toxic chemicals, or associated categories that manifest their action on the lung, include industrial materials (asbestos, sand, silicates, etc.), particulates from mining and combustion, agricultural chemicals, cigarette smoke, ozone, nitrogen oxides, and miscellaneous types that include diacetyl. Although pulmonary toxins may manifest their actions through many modes of action, we have recently postulated that oxidative processes (ET, ROS, and OS) are often strong contributors to pulmonary damage (Kovacic and Somanathan 2009b). We regard these aforementioned oxidative processes to form the basis of a unifying mechanism that may well describe how pulmonary toxins often produce their effects (Kovacic and Somanathan 2009b). The underlying concept behind this unifying mechanism is that the vast majority of toxins or their metabolites possess ET functionalities, which may undergo redox cycling and, thereby, generate ROS that are capable of injuring various cellular constituents of the lung or other organs by oxidative attack. Such attack is commonly associated with lipid peroxidation and oxidation of genetic materials that may produce DNA strand cleavage and production of 8-hydroxy-D-guanosine (8-OHD G). Antioxidants (AOs), sourced both naturally in fruits and vegetables, as well as synthetic ones may provide protection from these adverse effects. However, oxidative reactions in tissues are often accompanied by depletion of natural AOs, which further exacerbates toxicity or susceptibility thereto. The mechanistic framework described above is also applicable to some more prominent pulmonary illnesses, such as asthma, COPD (chronic obstructive pulmonary disease), and cancer. We propose that because diacetyl possesses an ET moiety and is a pulmonary toxin, it is plausible that it acts by oxidative processes known for other pulmonary toxins.

4 Diacetyl: Evidence to Support an ET Mechanism of Action Electrochemistry can provide valuable insight on mechanisms by which chemicals may produce their effects, because it deals with the energetics of electron uptake, which is relevant to how ET functionalities react in vivo. If, in a biosystem reaction,

Electron Transfer as a Potential Cause of Diacetyl Toxicity

137

Fig. 1 Diacetyl diimine

the reduction potential is measured and found to be more positive than –0.5 V, the possibility exists for ET. This value for diacetyl (at pH 5) is –0.37 V; for diimine derivatives (Fig. 1) this value is –0.45 to –0.49 V (Niufar et al. 2002). Thus, these key substances are within the electrochemical range that may sustain ET reactions in vivo, possibly with the formation of ROS. The imines represent models of counterparts generated by the reaction of diacetyl with the primary amino groups of proteins. Because ET processes are catalytic, only small amounts of the agents would be needed to induce significant physiological effects. This thesis is supported by the observation that α-dicarbonyl has an affinity for electrons (Jacobs 1986); in fact, the detailed nature of the resultant radical anion has been elucidated (Russell 1968). Valuable additional insight may be gained from evaluating electroreduction data from α-dicarbonyl compounds other than diacetyl (Kovacic 2006). For example, methylglyoxal (CH3 COCHO) possesses a reduction potential in cyclic voltammetry of –0.18 V in the presence of strong acids (Montoya et al. 1993). From polarography, the range of values for the reduction potential (E1/2 ) of this substance was –0.34 to –0.46 V for pH 2.5–5 (Mellado and Montoya 1994a). This range clearly supports the view that these α-dicarbonyl compounds may result in ET reactions in biological systems. It is interesting that methylglyoxal is a prominent member of a host of products that arise from oxidation of sugars, lipids, and amino acids and induces various toxic responses. Wu (2005) and Du et al. (2001) report that some ROS species are generated in such toxic responses, and AOs, if present, offer benefits. The electrochemical characteristics of methylglyoxal, and the degree to which it forms imines from protein interaction, may be important to its biochemical behavior. Unfortunately, research on this aspect of its mode of action has largely been neglected (Choudhary et al. 1997). The electron uptake efficiency of any ET agent is strongly correlated to its electron affinity (EA). Electron affinity is expressed as the enthalpy change ( H) for the process A + e– → A – . The electron uptake is determined by the free energy ( G) for this electron attachment, which is related to the EA at any given temperature (T) by G = H – T S = EA – T S ( S = entropy change). However, because S for this process is roughly independent of the electron acceptor (A), the free energy for the electron uptake is determined almost entirely by the electron affinity. Electron affinities for many organic electron acceptors were obtained by

138

P. Kovacic and A.L. Cooksy

directly measuring electron absorption coefficients in electron-rich environments, which yield a relative electron uptake constant (KA ). The process requires a direct measurement of the anion/neutral concentration ratio of anthracene to standardize the results through determining an absolute equilibrium constant (KA ) (Briegleb 1964) (See Eq. (1) below). EA(A) = RT ln KA = RT[ ln KA (A) + ln KA (anthracene) − ln KA (anthracene)]. (1) These measurements of EA are more direct and precise than methods based on the Huckel orbital energies or on relative electron transfer efficiencies that use a common donor. The EA determined by this method for diacetyl is 0.60 eV. It is illuminating to compare this value for diacetyl with those of prominent ET functionalities, such as quinone and aromatic nitro compounds, and to consider the effect of structural differences. The EA value for diacetyl is within the range of those for quinones (0.54–0.64 eV) and dinitrophenol (0.59 eV) (Briegleb 1964). The strongest acceptors in the quinone category are 2,3-dichloro-5,6-dicyano- p-benzoquinone, 2,6-dinitrobenzoquinone, tetracyano- p-benzoquinone, 2,3-dicyanobenzoquinone, and bis(cyanomethylene)quinone. EAs of unsubstituted quinones are as follows: 9,10-anthraquinone < 1,4-naphthoquinone ≈ 9,10-phenanthrenequinone ≈ 1,2-naphthoquinone ≈ p-benzoquinone < o-benzoquinone < 1,8-diphenoquinone. Nitroaromatics are relatively good electron acceptors: 2,4,7-trinitrofluorenone > 1,3,5-trinitrobenzene ≈ 1,4-dinitrobenzene > nitrobenzene. It is of particular interest that molecules with two adjacent electrophilic carbonyl groups (e.g., diacetyl, ethyl pyruvate, and dimethyloxaloacetate) are relatively good electron acceptors, and, in this regard, are comparable to trinitrobenzene. These findings represent favorable delocalization of the derived radical anion over the conjugated α-dicarbonyl system and anion stabilization by electronegative oxygen. By comparison, the absence of conjugation in acetylacetone results in a low EA value (0.34 eV). Diacetyl has long been used to stabilize electrons in biological systems. Diacetyl has been used to function in a fashion similar to oxygen in enhancing the response of bacterial spores to x-rays (Tallentire et al. 1968). Diacetyl, bound to an active site in vivo, may well be present in a hydrogen-bonded complex involving acidic agents in proteins that could be in the form of alcohols, phenols, carboxylic acids, or RNH3 + . The favorable influence of H-bonding on electron uptake and ET is indicated by the following data for reduction potentials of diacetyl vs. pH: −0.2 V, pH 1.5; −0.3 V, pH 5; −0.5 V, pH 7 (Niufar et al. 2002). Electrostatics constitutes those effects determined by the distribution of fixed charges in a system, such as the interactions between two polar groups of a protein. The application of electrostatics to biology and medicinal chemistry has been neglected, which is unfortunate because the literature discloses extensive involvement of electrostatics in living systems. Electrostatics is involved in living systems at a fundamental level, because electromagnetic forces are primarily responsible for the structure of matter (from atoms to more complex substances). Electrical

Electron Transfer as a Potential Cause of Diacetyl Toxicity

139

forces also appear to be a vital factor in the early evolution of life (Gagliardi 2006). Electrostatic force, a component of electromagnetic interaction, has played a dominant role in the dynamics of cell division and other biochemical processes in primitive cells, with the involvement persisting in modern, highly evolved eukaryotic cells. An electrostatic mechanism is believed to play a role in receptor–ligand activity, and therefore in receptor docking (Kovacic et al. 2007a). A fundamental characteristic of receptor–ligand activity entails a molecular electrostatic potential (MEP) that is associated with ions and dipoles, and diacetyl carbonyl groups have an appreciable dipole character. The exact role of MEP is speculative, but the fields produced may function as conduits for electrons and radicals. Such fields may also affect the energetics associated with the ET process. Electrical fields are also known to influence electron spin (Nowack et al. 2007). Phosphorylation and sulfation play essential roles in cell signaling. The MEP associated with the phosphate and sulfate anions is a key element in linking communication among cells and may also favorably affect the energetics involved (Kovacic et al. 2007b). With this background, it is reasonable to rationalize that electrostatic interactions are involved in the mechanism by which diacetyl produces its effects. We provide some lines of evidence below that support the involvement of electrostatics in ET and in the action of α-dicarbonyls, with emphasis on energetics (Kovacic 2008). Electrostatic effects have been studied in the plant kingdom. Ishikita and Knapp (2006) reported strong support for electrostatic action that energetically facilitates electron migration in the photosynthetic process. In one bacterial photosynthetic reaction, pheophytin is transferred to a primary quinone (QA), followed by passage to a secondary quinone (QB). A non-heme iron complex is situated midway between the two Qs. Removal of the iron complex causes a 15-fold decrease in ET rate from pheophytin to QA. Computational data support the conclusion of a predominant role for Fe electrostatics in relation to a favorable energetic effect on forward ET. Hence, the electrostatic field of Fe can be regarded as a force that facilitates efficient electron migration. MEP is a significant force in DNA function, which has not attracted appreciable attention. The MEP results from the presence of repeating phosphate links in the DNA chain, along with associated anions. One review addresses the influence of electrostatics on the chemical behavior of DNA (Kovacic and Wakelin 2001). This article addresses the impact of MEP on ligand binding to DNA, guanine oxidation, axial charge transport, hopping termination, and reactions with charged and uncharged ROS. In the Hofmeister series, a system designed to classify ions according to their ability to affect water structure, ammonium-type cations and sulfate or phosphate anions are the most active, which is generally in line with their involvement in cell signaling and certain other biological processes (Kovacic 2008). The evidence that diacetyl and its imine derivatives act through mechanisms that involve ET is based on the following: ease of electron uptake, including having an appropriate reduction potential; stability of the resultant radical anion that

140

P. Kovacic and A.L. Cooksy

permits time for biological interaction; ability to interact with proteins to form conjugated imines (which may serve to anchor the agent at the receptor and illicit a physiological response); similarity to other bioactive α-dicarbonyls in both electrochemical character and in vivo action; and, finally, the propensity of diacetyl to cause pulmonary damage similar to that caused by selected other pulmonary toxins.

5 Other Relevant α-Dicarbonyls The literature contains articles that address appreciable numbers of other α-dicarbonyls which are structurally related to diacetyl. The chemical and physiological properties of these other α-dicarbonyls, discussed in the following sections, provide additional support for the mechanistic theme presented for diacetyl.

5.1 Ethanol Metabolites Diacetyl is involved in the metabolism of ethanol, which involvement has been fairly well delineated (Kovacic and Cooksy 2005b; Kovacic and Somanathan 2006b). As ethanol is metabolized, there is an increase in ROS and an initial generation of hydroxyethyl radicals. A key intermediate in the metabolism of ethanol is acetaldehyde, which is converted to acetic acid. Many investigators consider the acetaldehyde to be the main toxin from ethanol consumption. Other reactions that damage tissues as ethanol is metabolized entail condensation with protein and formation of ROS during the oxidation of acetaldehyde to acetic acid. Oddly, despite their toxicity and the problems emanating from alcohol addiction, scant attention has been paid to the products of acetaldehyde’s side reactions, which mostly contain four carbons. Most of these side reactions result in the production of diacetyl, acetoin, and 2,3-butanediol. Acetaldehyde reacts with pyruvate to form acetoin, after which it is oxidized to α-dicarbonyl. One hypothesis that has been advanced is that the side products may be responsible for “hangovers” (Otsuka et al. 1996).

5.2 Bacterial Cell Signaling Agents The α-dicarbonyl functionality is also present in 4,5-dihydroxy-2,3-pentanedione (DPD)(Fig. 2), an important bacterial cell signaling molecule whose activity may be related to that of diacetyl. Cell signaling has attracted attention in higher organisms for some time and has recently been reported to occur in bacteria. If cell signaling is disrupted, it may have substantive impact on living systems. Cell signaling may be important for several reasons, including its role in quorum sensing. It is known that chemical exposure may affect cell signaling. DPD acts as an autoinducer, one of the trigger molecules in quorum sensing. In vivo DPD is in equilibrium with a furanone and a furanosyl-borate diester (AI-2). In one hypothesis, ET and ROS are involved in cell signaling in higher organisms and are also involved in biological conduits, relays, and electrical effects (Kovacic and Pozos 2006).

Electron Transfer as a Potential Cause of Diacetyl Toxicity

141

Fig. 2 4,5-Dihydroxy-2,3pentanedione (DPD)

As ET agents, diacetyl or its imine derivatives can serve as a model for DPD, since diacetyl possesses a reduction potential amenable to ET in the biological domain. Hence, it is conceivable that DPD and its imine derivatives also may be involved in ET-ROS processes (Kovacic 2007). The presence of hydroxyl groups should facilitate ET by DPD vs. diacetyl, as shown in Fig. 3; these hydroxyl groups stabilize the anionic character of the oxygen molecules via bonding with the partially acidic hydrogens of the alcohol groups. A 2007 article focuses on 3hydroxytridecane-4-one (CAI-1), an α-hydroxyketone, as a bacterial signaling agent (Higgins et al. 2007). In this chapter, it is pointed out that the α-hydroxyketone acetoin is oxidized in vivo to diacetyl. Similar metabolism may occur with CAI1 to produce an ET α-dicarbonyl that is capable of playing a bacterial cell communication role.

Fig. 3 Hydrogen-bonded radical anion of DPD

142

P. Kovacic and A.L. Cooksy

5.3 Other Toxins Exposure to reactive α-dicarbonyl species, of which diacetyl is a member, is known to induce ROS and OS. In addition, one recent report casts light on cellular damage induced by reactive carbonyl species (RCS) (Wondrak et al. 2002). Such toxic entities, mainly in the α-dicarbonyl class, are generated by oxidation of sugars, lipids, and amino acids. Among the more prominent examples are glyoxal, methylglyoxal, and α-dicarbonyls such as 3-deoxyosones from monosaccharides. This category can exert its adverse effects by increasing OS, thereby generating a vicious cycle of ROS and RCS formation. Lately, α-dicarbonyls have attracted appreciable attention as key reactive intermediates in toxicity and involvement in neurodegenerative diseases, for example, Alzheimer’s and amyotrophic lateral sclerosis, involving the Maillard reaction, which yields AGEs. As discussed in another section, imines from condensation with protein primary amino groups may be other important participants in the induction of such disease.

5.4 Protein Derivatives Protein cross-linking via the Maillard reaction has been the subject of much scrutiny. A study was made on the impact of this process on enzyme action (Miller and Gerrard 2005). Protein glycation can generate α-dicarbonyls (see below). Protein function following glycation was examined after treatment of ribonuclease A with various α-dicarbonyl compounds, including diacetyl, which induced cross-linking and impaired activity. The effects of two Maillard reaction inhibitors, namely aminoguanidine and 3,5-dimethylpyrazole-1-carboxamidine, were assessed, along with a parallel measurement of the effect on enzyme activity. Evidence indicates that prevention of protein cross-linking does not necessarily preserve enzyme activity. Thus, the data point to the involvement of another mode of action. We suggest that electrochemical properties, including ET, may be important factors in this alternative mode of action. In a related investigation, odorous products were formed from reactions of diacetyl with amino acids involving the Maillard and Strecker reaction (PripasNicolau et al. 2000). In the presence of cysteine, many of the derivatives formed were heterocyclic in nature.

5.5 Advanced Glycation End Products (AGEs) Among AGEs are α-dicarbonyls that result from protein–carbohydrate interactions. Accumulated evidence supports the hypothesis that there is mutual enhancement between glycation and oxidation and entails closely linked processes (Traverso et al. 1997). Glucose auto-oxidation plays an essential role in non-enzymatic glycation

Electron Transfer as a Potential Cause of Diacetyl Toxicity

143

of proteins. Glucose and glycated collagen show a catalytic role in lipid peroxidation. Early glycation products can be transformed into AGEs by oxidative processes involving ROS. Glyoxal and methylglyoxal products result from oxidative degradation of glucose. Condensation of the carbonyl functions, present in these compounds, with primary amino groups of protein occurs and has been described in the literature (Walker and Feather 1983). Non-degraded α-carbonyls such as deoxyosone, glucosome, and 3-deoxyglucose are also formed and may display potential for ET and formation of ET imines as well.

5.6 Other Bioactive α-Dicarbonyls Cyclohexane-1,2-dione is an antiviral agent (Tiffany et al. 1957). Diacetyl monoxime, an imine analog, has attracted attention because of its physiological activity, including its effects on cardiac muscle tension, neuromuscular transmission, action potential, and ion currents (Schlichter et al. 1992). This molecule also displays some activity as an antidote for nerve gas (Kovacic 2003). Since the protonated form exhibits a reduction potential of −0.43 V (Niufar et al. 2002), some of its bioresponses may reflect ET-ROS activity. o-Quinones represent a special class of α-dicarbonyl compounds that possess reduction potentials amenable to ET with the capacity to produce ROS. Various members of this group are physiologically active as (for example) anti-infective agents (Kovacic and Becvar 2000) or anticancer drugs (Kovacic and Osuna 2000) and, thereby, emulate the profile of substances that fit the ET-ROS-OS mechanistic framework. A new structural class of antibiotics, represented by marinopyrroles (Fig. 4), was recently reported (Hughes et al. 2008). These antibiotics were obtained from actinomycetes that inhabit ocean sediments. The agents possess potent antibiotic activities

Fig. 4 Marinopyrrole with α-dicarbonyl conjugation

144

P. Kovacic and A.L. Cooksy

against methicillin-resistant Staphylococcus aureus. Extensive evidence has been presented to support ET–ROS–OS involvement in the action of anti-infective agents (Kovacic and Becvar 2000). This evidence has been buttressed by research on a common cell death mechanism for antibiotics that involves ROS (Kohanski et al. 2007). Inspection of the marinopyrrole structure (Fig. 4) reveals the presence of a highly conjugated α-dicarbonyl analog, obtained from conjugation by aromatic nuclei, namely pyrrole and benzenoid. Hence, the electron affinic nature of the moiety should be appreciably greater than for diacetyl. The reduction potential of carbonyls is enhanced by hydrogen bonding, which increases electron attraction and stabilizes the resultant radical anion. It is interesting that the phenolic hydroxyl groups in Fig. 4 are in position to coordinate with the carbonyls, with resultant enhancement of the potential for electron uptake. A relevant feature of carbonyls is the ease by which they condense with protein primary amino groups to yield imines. As previously mentioned, conjugated imines and iminiums are ET agents and may play important physiological roles in organisms. Two aromatic oximes, zinviroxime (Fig. 5a) and enviroxime (Fig. 5b), have been shown to be potent inhibitors of rhinovirus multiplication in vivo (Kovacic et al 1990). The protonated, iminium-like forms displayed reduction potentials in the range amenable to ET in the biodomain. A similar situation may pertain to the structure in Fig. 4.

Fig. 5 Zinviroxime (syn) and enviroxime (anti)

These reports show that various α-dicarbonyl derivatives are electron affinic and possess reduction potentials favorable for in vivoET. In addition, the compounds display diverse physiological activities that are compatible with the ET–ROS–OS theme.

6 Other Physiological Effects When tested for genotoxicity, diacetyl generally exhibited mutagenic activity in Salmonella typhimurium (ILS Inc. 2007). ROS are believed to be involved in

Electron Transfer as a Potential Cause of Diacetyl Toxicity

145

the induction of genotoxicity (Kovacic and Jacintho 2001). Sulfite, a reducing or inactivating agent toward ROS, countered the mutagenicity effect of diacetyl (ILS Inc. 2007). When diacetyl was tested in a sister chromatid exchange (SCE) assay, positive results were recorded in Chinese hamster ovary cells. The reducing agent bisulfite significantly diminished the frequency of SCEs and proportion of endoreduplicated cells. Diacetyl completely inhibited cell growth in sarcoma cells at high doses, but only moderately at lower levels. Mutagenesis is believed to reflect the formation of adducts between α-dicarbonyls and guanine residues of DNA, giving rise to conjugated imines that possess ET characteristics and are capable of modifying DNA (Mellado and Montoya 1994b). ET and ROS action may at times apply to both mutagenesis and carcinogenesis, since these processes are closely linked (Kovacic and Jacintho 2001). Diacetyl has activating and deactivating influences on a number of enzymes and metabolic processes. Diacetyl is known to have increased ornithine decarboxylase activity and DNA synthesis. Moreover, diacetyl inactivated estradiol 17-α-dehydrogenase in the human placenta under ultraviolet light. Imines and oximes of α-dicarbonyl compounds are good complexing agents for heavier metals. Since metal complexes can undergo ET and generate ROS (Kovacic et al. 2005; Kovacic and Somanathan 2006a; Kovacic et al. 2002; Poli et al. 1989; Kovacic and Thurn 2005; Kovacic and Somanathan 2005; Kovacic and Cooksy 2005a; Kovacic and Somanathan 2008; Halliwell and Gutteridge 1999), it is plausible that these complexes possess physiological activity.

7 Summary Diacetyl, a butter-flavoring component, has recently attracted scientific and media attention because it has been implicated as an agent that induces popcorn lung disease in exposed plant workers. This disease, officially referred to as bronchiolitis obliterans, entails exposure-induced compromise to the lung’s epithelial barrier function. In this review, we present a novel molecular mechanism (electron transfer, ET) designed to explain how diacetyl and its imine derivatives might interact to produce lung damage. We relate the fact that diacetyl and related compounds possess reduction potentials amenable to electron transfer (ET) in vivo. The electrochemical nature of these toxicants can potentially disrupt normal ET processes, generate reactive oxygen species (ROS), and participate in cell signaling events. Condensation of diacetyl with protein may also play a role in the toxicity caused by this compound. ET is a common feature of toxic substances, usually involving their metabolites which can operate per se or through reactions that generate ROS and oxidative stress (OS). Examples of agents capable of ET are quinone and metal compounds, aromatic nitro compounds, and iminium salts. Among compounds that generate ET, the α-dicarbonyl ET class, of which diacetyl is a member, is much less studied. This review emphasizes diacetyl as an agent that acts through oxidative processes to cause its effects. However, we also treat related substances that appear to act by a similar mechanism. This mechanism forms a theoretical framework capable of

146

P. Kovacic and A.L. Cooksy

describing the mechanism by which diacetyl may induce its effects and is in accord with various physiological activities displayed by other α-dicarbonyl substances. Examples of substances that may act by mechanisms similar to that displayed by diacetyl include cyclohexane-1,2-dione, marinopyrroles, reactive carbonyl species, the bacterial signaling agent DPD, and advanced glycation end products. Acknowledgments Editorial assistance by Angelica Ruiz and Thelma Chavez is appreciated. ALC is partly supported through grant CHE-0719575 of the National Science Foundation.

References Baguley BC, Wakelin LPG, Jacintho JD, Kovacic P (2003) Mechanisms of action of DNA intercalating acridine-based drugs: How important are contributions from electron transfer and oxidative stress. Curr Med Chem 24:2643–2649. Berson A, Descatoire V, Sutton A, Fau D, Maulny B, Vadrot N, Feldmann G, Berthon B, Torkjmann T, Pessayre D (2001) Toxicity of alpidem, a peripheral benzodiazepine receptor ligand, but not zolpidem, in rat hepatocytes: Role of mitochondrial permeability transition and metabolic activation. J Pharmacol Exp Ther 299:793–800. Briegleb G (1964) Electron affinity of organic molecules. Angew Chem Int Edit 3:617–632. Choudhary D, Chandra D, Kale RK (1997) Influence of methylglyoxal on antioxidant enzymes and oxidative damage. Toxicol Lett 93:141–152. Du J, Suzuki H, Nagase F, Akhand A.A, Ma XY, Yokoyama T, Miyata T, Nakashima I, (2001) Superoxide-mediated early oxidation and activation of ASKI are important for initiating methylglyoxal-induced apoptosis process. Free Rad Biol Med 31:469–478. Egilman D, Mailloux C, Valentin C (2007) Popcorn-worker lung caused by corporate and regulatory negligence: An avoidable tragedy. Int J Occup Environ Health 13:85–98. Fedan JS, Dowdy J.A, Fedan KB, Hubbs AF (2006) Popcorn worker s lung: In vitro exposure to diacetyl, an ingredient in microwave popcorn butter flavoring, increases reactivity to methacholine. Toxicol Appl Pharmacol 215:17–22. Gagliardi LG (2006) Electrostatic considerations in nuclear envelope breakdown and reassembly. J Electrostat 64:843–849. Halliwell B, Gutteridge JMC (1999) Free Radicals in Biology and Medicine, Oxford University Press, New York pp 1–897. Hanson DG (2009) OSHA withdraws diacetyl proposal. Chem Eng News 87:23. Higgins DA, Pomianek ME, Kraml CM, Taylor RK, Semmelhack MF, Bassler BL (2007) The major Vibrio cholerae autoinducer and its role in virulence factor production against oxidative stress. Nature 450:883–886. Hubbs AF, Goldsmith WT, Kashon ML, Frazer D, Mercer RR, Battelli LA, Kullman GJ, Schwegler-Berry D, Friend S, Castranova V (2008) Respiratory toxicologic pathology of inhaled diacetyl in Sprague-Dawley rats. Toxicol Pathol 36:330–344. Hughes CC, Prieto-Davo A, Jensen PR, Fenical W (2008) The marinopyrroles, antibiotics of an unprecedented structure class from a marine Streptomyces sp.Org Lett 10:629–631. ILS (2007) Integrated Laboratory Systems Inc: Chemical Information Review Document for Artificial Butter Flavoring and Constituents Diacetyl [CAS No. 431-03-08] and Acetoin [CAS No. 513-86-0], http://ntp.niehs.nih.gov/ntp/htdocs/Chem_Background/ ExSumPdf/Artificial_butter_flavoring.pdf 2007. Ishikita H, Knapp EW (2006) Electrostatic role of the non-heme iron complex in bacterial photosynthetic reaction center. FEBS Lett 580:4567–4570. Jacobs GP (1986) Non-nitro radiation sensitizers. Int J Radiat Biol Relat Stud Phys Chem Med 49:887–890.

Electron Transfer as a Potential Cause of Diacetyl Toxicity

147

Kohanski MA, Dwyer DJ, Hayete B, Lawrence CA, Collins J (2007) A common mechanism of cellular death induced by bactericidal antibiotics. J Cell 130:797–810. Kovacic P (2003) Mechanism of organophosphates (nerve gases and pesticides) and antidotes: Electron transfer and oxidative stress. Curr Med Chem 10:2705–2710. Kovacic P (2005) Fundamental, electron transfer mechanism by pyrylium-type ions for the anticancer drugs 5,6-dimethylxanthenone-4-acetic acid (DMXAA) and flavone-8-acetic acid (FAA). Curr Med Chem-Anti-Cancer Agents 5:501–506. Kovacic P (2006) Novel electrochemical approach to enhanced toxicity of 4 oxo-2-nonenal vs. 4-hydroxy-2-nonenal (role of imine): Oxidative stress and therapuetic modalities. Med Hypotheses 67:151–156. Kovacic P (2007) Unifying mechanism for bacterial cell signaling (4,5-dihydroxy-2,3-petanedione, lactones and oligopeptides) electron transfer and reactive oxygen species. Practical medical features. Med Hypotheses 69:1105–1110. Kovacic P (2008) Bioelectrostatics: Review of widespread importance in biochemistry. J Electrostat 66:124–129. Kovacic P, Becvar LE (2000) Mode of action of anti-infective agents: Focus on oxidative stress and electron transfer. Curr Pharmaceut Des 6:143–167. Kovacic P, Cooksy AL (2005a) Unifying mechanism for toxicity and addiction by abused drugs: Electron transfer and reactive oxygen species. Med Hypotheses 64:357–366. Kovacic P, Cooksy AL (2005b) Role of diacetyl metabolite in alcohol toxicity and addiction via electron transfer and oxidative stress. Arch Toxicol 79:123–128. Kovacic P, Draskovich CD, Pozos RS (2007b) Unifying electrostatic mechanism for phosphates and sulfates in cell signaling. J Recept Signal Transduct 27:433–443. Kovacic P, Jacintho JD (2001) Mechanisms of carcinogenesis: Focus on oxidative stress and electron transfer. Curr Med Chem 8:773–796. Kovacic P, Kassel MA, Popp WJ, Feinberg BA (1990) Mechanisms of antiviral action: Focus on electron transfer and oxidative stress. J Biopharmaceut Sci 1:315–330. Kovacic P, Osuna JA (2000) Mechanisms of anti-cancer agents: Emphasis on oxidative stress and electron transfer. Curr Pharmaceut Des 6:277–309. Kovacic P, Pozos RS, Somanathan R, Shangari R, O Brien P (2005) Mechanism of mitochondrial uncouplers, inhibitors, and toxins: Focus on electron transfer, free radicals, and structure-activity relationships. J Curr Med Chem 5:2601–2623. Kovacic P, Pozos RS (2006) Cell signaling (mechanism and reproductive toxicity): Redox chains, radicals, electrons, relays, conduit, electrochemistry, and other medical implications. Birth Defects Res Part C 78:333–344. Kovacic P, Pozos RS, Draskovich CD (2007a) Unifying electrostatic mechanism for receptorligand activity. J Recept Signal Transduct 27:411–432. Kovacic P, Sacman A, Wu-Weis M (2002) ephrotoxins: Widespread role of oxidative stress and electron transfer. Curr Med Chem 9:823–847. Kovacic P, Somanathan R (2005) Neurotoxicity: The broad framework of electron transfer, oxidative stress and protection by antioxidants. Curr Med Chem-CNS Agents 5:249–258. Kovacic P, Somanathan R (2006a) Mechanism of teratogenesis: Electron transfer, reactive oxygen species, and antioxidants. Birth Defects Res Part C 78:308–344. Kovacic P, Somanathan R (2006b) In: New Research on Alcohol Abuse and Alcoholism. Brozner EY (ed), Nova Science, New York, pp 51–58. Kovacic P, Somanathan R (2008) Ototoxicity and noise trauma, electron transfer, reactive oxygen, species, cell signaling, electrical effects, and protection by antioxidants: Practical medical aspects. Med Hypotheses 70:914–923. Kovacic P, Somanathan R (2009a) Zolipidem, a clinical hypnotic that affects electronic transfer, alters synaptic activity through potential GABA receptors in the nervous system without significant free radical generation. Oxid Med Cell Longev 2: 116–121. Kovacic P, Somanathan R (2009b) Pulmonary toxicity and environmental contamination: Radicals, electron transfer and protection by antioxidants. Rev Environ Contam Toxicol 201:41–69.

148

P. Kovacic and A.L. Cooksy

Kovacic P, Thurn LA (2005) Cardiovascular toxicity from the perspective of oxidative stress, electron transfer, and prevention by antioxidants. Curr Vasc Pharmacol 3:107–118. Kovacic P, Wakelin LPG (2001) DNA molecular electrostatic potential, novel perspectives for the mechanisms of action of anticancer drugs involving electron transfer and oxidative stress. Anti-Canc Drug Des 16:1–10. Kreiss K (2007) Flavoring-related bronchiolitis obliterans Curr Opin Allergy Clin Immnol 7: 162–167. Mellado JMR, Montoya MR (1994a) CEC mechanisms in the electroreduction of α-dicarbonyl compounds on mercury electrodes. J Electroanal Chem 365:71–78. Mellado JMR, Montoya MR (1994b) Correlations between chemical reactivity and mutagenic activity against S. typhimurium mechanism TA100 for dicarbonyl compounds as proof of mutagenic mechanism. Mutat Res 304:261–264. Miller AG, Gerrard JA (2005) Assessment of protein function following crosslinking by dicarbonyl. Ann N Y Acad Sci 1043:195–200. Montoya MR, Zon MA, Mellado JMR (1993) Investigation of the reduction of 1,2cyclohexanedione and methylglyoxal on mercury electrodes under pure kinetic conditions by linear sweep voltammetry. J Electroanal Chem 353:217–224. Morgan DL, Flake GP, Kirby PJ, Palmer SM (2008) Respiratory toxicity of diacetyl in C57BL/6 mice. Toxicol Sci 103:169–180. Niufar NN, Haycock FL, Wesemann JL, MacStay JA, Heasley VL, Kovacic P (2002) Reduction potentials of conjugated aliphatic ketones, oximes and imines: Correlation with structure and bioactivity. Rev Chem Soc Mex 46:307–312. Nowack KC, Koppens FHL, Nazarov YV, Vandersypen LMK (2007) Coherent control of a single electron spin with electric fields. Science 318:1430–1433. Otsuka M, Mine T, Ohuchi K, Ohmori S (1996) A detoxification route for acetaldehyde, metabolism of diacetyl, acetoin and 2,3-butanediol in liver homogenate perfused liver of rats. J Biochem (Tokyo) 119:246–251. Poli G, Cheeseman KH, Dianzani MU, Slater TF (1989) Free Radicals in the Pathogenesis of Liver Injury, Pergamon, New York, pp 1–330. Pripis-Nicolau L, de Revel G, Bertrand A, Maujean A (2000) Formation of flavor components by the reaction of amino acids and carbonyl compounds in mild conditions. J Agric Food Chem 48:3761–3766. Russell GA (1968) In: Radical Ions. Interscience. Kaiser ET, Kevan N (eds), New York, pp 91–106. Schlichter LC, Pahapill PA, Chung I (1992) Dual action of 2,3-bulanedione monoxime (BDM) on K+ currents in human T lymphocytes. J Pharmacol Exp Ther 261:438–446. Schneider A (2007) Seattle Post-Intelligencer, Dec: 21: p. A.1. Tallentire A, Schiller NL, Powers EL (1968) 2,3-Butanedione, an electron-stabilizing compound, as a modifier of sensitivity of Bacillus megateriumspores to X-rays. Int J Radiat Biol 14: 397–402. Tiffany BD, Wright JB, Moffett RB, Heinzelmann RV, Strube RE, Aspergren BD, Lincoln EH, White JL (1957) Antiviral compounds, aliphatic glyoxals, hydroxyldehydes and related compounds. J Am Chem Soc 79:1682–1687. Traverso N, Manini S, Cottalasso D, Odetti P, Marinari UM, Pronzato MA (1997) Mutual interaction between glycation and oxidation during non-enzymatic protein modification. Biochim Biophys Acta 1336:409–418. Walker GR, Feather MS (1983) The Maillard Reaction in Food and Nutrition, ACS Symposium Series. 215:ACS, Washington, DC, p 36. Wondrak GT, Cervantes-Laurean D, Roberts MG, Qrasem JG, Kim M, Jacobson EL, Jacobson MK (2002) Identification of dicarbonyl scavengers for cellular protection against oxidative stress. Biochem Pharmacol 63:361–373. Wu L (2005) The pro-oxidant role of methylglyoxal in mesenteric artery smooth muscle cells. Can J Physiol Pharmacol 83:63–68.

Index

A Absorption & elimination, bioconcentration theory, 19 Acyltransferases, aquatic organisms, 79 Advanced glycation end products, α-dicarbonyl species, 142 Age-dependence, pesticide bioconcentration, 14 Aquatic organism acyltransferases, 79 conjugation reactions (diag.), 74 glutathione-S-transferases, 75 growth, bioconcentration effects, 14 hydrophobicity effects, 8 metabolism organo- & organo-phosphorus pesticides, 85 pesticide bioaccumulation, 1 ff. bioconcentration, 1 ff. metabolism, 1 ff. Aquatic species BCF (bioconcentration factor), pesticide clearance times, 23 BCF vs. log Kow (diag.), 41 bioconcentration humic acid effects, 18 lipid-content effects, 9 organochlorine pesticides (table), 24 organophosphorus pesticides (table), 30 pesticides (table), 35 pyrethroid insecticides (table), 34 bioconcentration theory, 19 chemical metabolism (table), 80 differences, pesticide metabolism, 90 lipid content (table), 10 lipid content variability, 11 metabolism

glucose & glucuronic acid transferases, 72 organochlorine pesticides (table), 86 pesticides (table), 67 pesticides, 89 pesticide bioaccumulation (table), 43 biota-sediment accumulation factors (table), 46 metabolic pathways (table), 53 pesticide metabolism enzymes, 54 esterases, 63 regulatory testing, 2 Aquatic system, chemical flow (diag.), 3 B Bacterial cell signaling, chemical exposure, 140 BAF (bioaccumulation), controlling factors, 42 BCF (bioconcentration factor) defined, 4 effect pesticide class, 42 pesticide clearance times, 23 physico-chemical effects (table), 5 vs. log Kow , pesticides in aquatic species (diag.), 41 vs. log Kow , pesticides in fish (diag.), 7 Bioaccumulation description, 2 dietary route, 47 ecological effects, 45 effects, dissolved organic carbon, 47 flow in aquatic system (diag.), 3 metabolic effects, 48 of pesticides, 1 ff. aquatic species (table), 43

D.M. Whitacre (ed.), Reviews of Environmental Contamination and Toxicology Volume 204, Reviews of Environmental Contamination and Toxicology 204, C Springer Science+Business Media, LLC 2010 DOI 10.1007/978-1-4419-1440-8, 

149

150 Bioaccumulation (cont.) theory, 49 sediment effects, 44 Bioaccumulation factor (BAF), controlling factors, 42 Bioactive α-dicarbonyls, examples, 143 Bioconcentrated pesticides, organ distribution, 15 Bioconcentration in aquatic species organochlorine pesticides (table), 24 organophosphorus pesticides (table), 30 pesticides (table), 35 pyrethroid insecticides (table), 34 controlling factors, 3 description, 2 dissolved organic carbon effects, 18 effects biological factors, 14 humic acids, 18 environmental effects, 16 flow in aquatic system (diag.), 3 illumination effects, 16 isomerism effects, 9 lipid-content effects, 9 metabolism effects, 14 pesticides, 1 ff. pH-dependency, 17 potential, test regulations, 2 theory, 19 absorption & elimination, 19 Bioconcentration factor (BCF), defined, 4 Biological factors, bioconcentration effects, 14 Biomagnification, description, 2 Biota-sediment accumulation factors (BSAF) for pesticides, aquatic species (table), 46 Bio-uptake by aquatic species, metabolic effects, 51 Bronchiolitis oblitherans, lung disease, 133 C Carbamate pesticides, chemical structures (illus.), 102 Cell signaling chemical exposure, 140 electron transfer & reactive oxygen species, 140 Chemicals exposure, cell signaling effects, 140 flow, aquatic system (diag.), 3 hydrophobicity, BCF effects, 8 metabolism, aquatic species (table), 80

Index physico-chemical effects on BCF (table), 5 uptake, aquatic organism growth effects, 14 (various), structures (illus.), 99 Conjugation reactions, aquatic organisms (diag.), 74 Cytochrome P450, reaction scheme (diag.), 54 D Diacetyl connection, ethanol metabolites, 140 electron affinity, 138 enzyme activator, 145 genotoxicity, 144 mechanism of action, 136, 139 mechanism of action, electron transfer, 139 molecular electrostatic potential, 139 and other α-dicarbonyls, protein glycation, 142 ROS (reactive oxygen species) production, 135 structure (illus.), 137 toxic effects, metabolite vs. parent, 136 toxicity caused by electron transfer, 133 ff., 135 lung effects, 134 oxidative effects, 134 α-Dicarbonyls bioactive examples, 143 diacetyl relationship, 137 species advanced glycation end products, 142 reactive oxygen & carbonyls, 142 Dietary route, bioaccumulation, 47 Dissolved organic carbon bioaccumulation effects, 47 bioconcentration effects, 18 E Ecological factors, bioaccumulation effects, 45 Electrochemistry, toxicity connection, 137 Electron affinity, diacetyl relationship, 138 Electron transfer diacetyl toxicity, 133 ff., 135 oxidative stress, 134 physiological effects, 134 redox cycling, 136 Electrostatics, action in living systems, 138 Environmental factors, lipid-content effects, 13 Environmental fate, test regulations, 2 Enzyme activator, diacetyl, 145

Index inhibitors, glutathione-S-transferases, 78 -reaction scheme, cytochrome P450 (diag.), 54 role, pesticide metabolism, 52 Enzymology glutathione-S-transferases, 77 pesticide metabolism (illus.), 57 Esterases aquatic species pesticide metabolism, 63 substrates & inhibitors (illus.), 57 Ethanol metabolites, diacetyl connection, 140 F Florey-Higgens theory, chemical interactions, 8 G Genotoxicity, diacetyl, 144 Glucose transferases, pesticide metabolism, 72 Glucuronic acid transferases, pesticide metabolism, 72 Glutathione-S-transferase aquatic organisms, 75 enzymology, 77 inhibitors, 78 substrates (illus.), 57

151 Metabolic effects, aquatic species pesticide uptake, 51 Metabolic pathways, pesticides (table), 53 Metabolism in aquatic species organophosphate insecticides (table), 64 PAHs (table), 80 of chemicals, aquatic species (table), 80 diacetyl, 136 effects, bioaccumulation, 48 of organochlorine pesticides, aquatic species (table), 86 pesticides & other chemicals, 79 of pesticides aquatic species, 51, 89 aquatic species (table), 67 enzymology (illus.), 57 pesticides, 1 ff. polycyclic aromatic hydrocarbons, 79 Microcosm studies, pesticides, 96 Model ecosystems, pesticide studies (table), 93 Model ecosystem studies, pesticides, 92 Molecular electrostatic potential, diacetyl, 139 Mutagenesis, diacetyl, 145

L Light effects, bioconcentration, 16 Lipid content aquatic species (table), 10 in aquatic species, bioconcentration effects, 9 effects, environmental factors, 13 Lung disease, Bronchiolitis oblitherans, 133 pathology, diacetyl effects, 134

O Organochlorine pesticide bioconcentration, aquatic species (table), 24 chemical structures (illus.), 100 metabolism aquatic organisms (table), 86 aquatic species, 85 Organophosphate insecticide metabolism, aquatic species (table), 64 Organophosphorus pesticide aquatic species metabolism, 85 bioconcentration, aquatic species (table), 30 chemical structures (illus.), 101 Oxidases pesticide metabolism, 54 substrates, inducers & inhibitors (illus.), 57 Oxidative processes possible unifying toxic mechanism, 136 pulmonary toxicity mechanism, 136 Oxidative stress electron transfer, 134 toxic action, 135

M Mechanism of action, diacetyl, 136 Mesocosm studies, pesticides, 96

P PAH (polycyclic aromatic hydrocarbon) chemical structures (illus.), 99

H Humic acids, bioconcentration effects, 18 Hydrophobicity, BCF effects, 8 I Inducers, oxidases (illus.), 57 Inhibitors esterases (illus.), 57 oxidases (illus.), 57 Insecticide metabolism in aquatic species, organophosphates (table), 64 Isomerism, bioconcentration effects, 9

152 PAH (polycyclic aromatic hydrocarbon) (cont.) metabolism, 79 aquatic organisms (table), 80 Pesticides and chemicals, metabolism, 79 and other chemicals, structures (illus.), 102 in aquatic species, BCF vs. log Kow (diag.), 41 bioaccumulation aquatic organisms, 1 ff. aquatic species, (table), 43 theory, 49 bioconcentration age dependence, 14 aquatic organisms, 1 ff. aquatic species (table), 35 environmental effects, 16 biota-sediment accumulation factors, aquatic species (table), 46 chemical structures (illus.), 103 class, BCF effect, 42 clearance times, BCF effects, 23 elimination, species variability, 40 factors affecting bioconcentration, 3 in fish, BCF vs. log Kow (diag.), 7 metabolic pathways, aquatic species (table), 53 metabolism aquatic organisms, 1 ff. aquatic species, 51, 89 aquatic species (table), 67 aquatic species differences, 90 in aquatic species, esterases, 63 bioconcentration effects, 14 enzyme role, 52 enzymology (illus.), 57 oxidases, 54 sulfotransferases, 73 model ecosystem studies, 92 physico-chemical effects on BCF (table), 5 studies micro- & mesocosms, 96 model ecosystems (table), 93 uptake, aquatic organism growth effects, 14 PH-dependency, bioconcentration, 17 Popcorn lung disease

Index diacetyl toxicity, 133 ff. electron transfer, 133 ff. Popcorn plant workers, Bronchiolitis oblitherans, 133 Protein glycation, diacetyl & other α-dicarbonyls, 142 Pulmonary toxicity, unifying mechanism proposal, 136 Pyrethroid insecticide bioconcentration, aquatic species (table), 34 R Reactive oxygen species (ROS) electron transfer, 134 toxic action, 135 Redox cycling, electron transfer, 136 Regulatory testing, bioconcentration, 2 S Sediment effects, bioaccumulation, 44 Species variability, pesticide elimination, 40 Substrates esterases (illus.), 57 glutathione-S-transferase (illus.), 57 oxidases (illus.), 57 Sulfotransferases, pesticide metabolism, 73 T Temperature effects, bioconcentration, 16 Theoretical approach, pesticide bioaccumulation, 49 Tissue distribution, bioconcentrated pesticides, 15 Toxic action, electron transfer & oxidative mechanisms, 135 Toxicity electrochemical interactions, 137 from oxidative mechanisms, examples, 135 U Urea pesticides, chemical structures (illus.), 102 W Water chemistry effects, bioconcentration, 16