Dehalogenation: Microbial Processes and Environmental Applications

DEHALOGENATION Microbial Processes and Environmental Applications

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DEHALOGENATION Microbial Processes and Environmental Applications

edited by

Max M. Häggblom and

Ingeborg D. Bossert Rutgers University U.S.A.

KLUWER ACADEMIC PUBLISHERS NEW YORK, BOSTON, DORDRECHT, LONDON, MOSCOW

eBook ISBN: Print ISBN:

0-306-48011-5 1-4020-7406-9

©2004 Kluwer Academic Publishers New York, Boston, Dordrecht, London, Moscow Print ©2003 Kluwer Academic Publishers Dordrecht All rights reserved No part of this eBook may be reproduced or transmitted in any form or by any means, electronic, mechanical, recording, or otherwise, without written consent from the Publisher Created in the United States of America Visit Kluwer Online at: and Kluwer's eBookstore at:

http://kluweronline.com http://ebooks.kluweronline.com

CONTENTS CONTRIBUTORS

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PREFACE Max M. Häggblom and Ingeborg D. Bossert

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PART I. INTRODUCTION

1. Halogenated Organic Compounds – A Global Perspective Max M. Häggblom and Ingeborg D. Bossert

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PART II. MICROBIAL PROCESSES

2. Microbial Ecology of Dehalogenation Ingeborg D. Bossert, Max M. Häggblom, and L.Y. Young

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3. Diversity of Dechlorinating Bacteria Frank E. Löffler, James R. Cole, Kirsti M. Ritalahti, and James M. Tiedje

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4. Thermodynamic Considerations for Dehalogenation Jan Dolfing

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5. Dehalogenation by Anaerobic Bacteria Christof Holliger, Christophe Regeard, and Gabriele Diekert 6. Biodegradation of Chlorinated Compounds by White Rot Fungi James A. Field

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PART III. BIOCHEMISTRY AND CHEMISTRY

7. Bacterial Growth on Halogenated Aliphatic Hydrocarbons: Genetics and Biochemistry Dick B. Janssen, Jantien E. Oppentocht, and Gerrit J. Poelarends 8. Aromatic Dehalogenases: Insights into Structures, Mechanisms, and Evolutionary Origins Shelley D. Copley

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227

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9. Abiotic Dehalogenation by Metals Lisa A. Totten and Nada M. Assaf-Anid

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PART IV. ENVIRONMENTAL FATE AND APPLICATIONS 10. Bioavailability of Organohalides Kyoungphile Nam and Jerome J. Kukor

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11. Biotransformation of Halogenated Pesticides Dennis D. Focht

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12. Biodegradation of Atmospheric Halocarbons Ronald S. Oremland

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13. Dechlorination of Sediment Dioxins: Catalysts, Mechanisms, and Implications for Remedial Strategies and Dioxin Cycling Cyndee L. Gruden, Q. Shiang Fu, Andrei L. Barkovskii, Iris D. Albrecht, Mary M. Lynam, and Peter Adriaens

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14. Redox Conditions and the Reductive/Oxidative Biodegradation of Chlorinated Ethenes in Groundwater Systems Francis H. Chapelle and Paul M. Bradley

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15. Microcosms for Site-Specific Evaluation of Enhanced Biological Reductive Dehalogenation Donna E. Fennell and James M. Gossett

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16. Chlorinated Organic Contaminants from Mechanical Wood Processing and Their Bioremediation M. Minna Laine, Minna K. Männistö, Mirja S. Salkinoja-Salonen, and Jaakko A. Puhakka

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17. Polychlorinated Biphenyls in Aquatic Sediments: Environmental Fate and Outlook for Biological Treatment Donna L. Bedard

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PART V. SUMMARY 18. Environmental Dehalogenation – Problems and Recommendations Charles E. Castro

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INDEX

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CONTRIBUTORS Peter Adriaens, Environmental and Water Resources Engineering, The University of Michigan, 180 EWRE Bldg., 1351 Beal St., Ann Arbor, Ml 48109-2125, USA [email protected] Iris D. Albrecht, Environmental and Water Resources Engineering, The University of Michigan, Ann Arbor, Ml 48109-2125, USA Nada Assaf-Anid, Chemical Engineering Department, Manhattan College, Riverdale, NY 10471, USA [email protected] Andrei L. Barkovskii, Georgia College and State University, Milledgeville, GA 31061, USA [email protected] Donna L. Bedard, Department of Biology, Rensselaer Polytechnic Institute, MRC 236, 110 8th Street, Troy, NY 12180-3590, USA [email protected] Ingeborg D. Bossert, Department of Biochemistry and Microbiology, Cook College, Rutgers, The State University of New Jersey, 76 Lipman Drive, New Brunswick, NJ 08901-8525, USA [email protected] Paul M. Bradley, Water Resources Division, U.S. Geological Survey, 720 Gracern Road, Suite 129, Columbia, SC 29210, USA [email protected] Charles E. Castro, Chemical and Environmental Consulting, 1090 Madison Place, Laguna Beach, CA 92651, USA [email protected] Francis H. Chapelle, Water Resources Division, U.S. Geological Survey, 720 Gracern Road, Suite 129, Columbia, SC 29210, USA [email protected] James R. Cole, Center for Microbial Ecology, Michigan State University, East Lansing, Ml 48824-1325, USA [email protected] Shelley D. Copley, Department of Chemistry and Biochemistry and Department of Molecular, Cellular, and Developmental Biology, University of Colorado, Boulder, CO 80309-0215, USA [email protected] Gabriele Diekert, Lehrstuhl für Angewandte und Oekologische Mikrobiologie, Philosophenweg 12, D-07743 Jena, Germany [email protected] Jan Dolfing, Alterra, Wageningen University and Research Centre, P.O. Box 47, 6700 AA Wageningen, The Netherlands [email protected]

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Donna E. Fennell, Department of Environmental Sciences, Cook College, Rutgers, The State University of New Jersey, 14 College Farm Road, New Brunswick, NJ 08901-8851, USA [email protected] James A. Field, Department of Chemical and Environmental Engineering, University of Arizona, P.O. Box 210011, Tucson, AZ 85721-0011, USA [email protected] Dennis D. Focht, Department of Plant Pathology, 2317 Webber Hall, University of California, Riverside, CA 92521, USA [email protected] Q. Shiang Fu, Department of Civil and Environmental Engineering, Stanford University, Palo Alto, CA 94305, USA [email protected] James M. Gossett, Department of Civil and Environmental Engineering, Cornell University, 319 Hollister Hall, Ithaca, NY 14853, USA [email protected] Cyndee L. Gruden, Environmental and Water Resources Engineering, The University of Michigan, Ann Arbor, Ml 48109-2125, USA [email protected] Max M. Häggblom, Department of Biochemistry and Microbiology and Biotechnology Center for Agriculture and the Environment, Cook College, Rutgers, The State University of New Jersey, 76 Lipman Drive, New Brunswick, NJ 08901-8525, USA [email protected] Christof Holliger, DGR-IGE/Laboratory for Environmental Biotechnology, Swiss Federal Institute of Technology (EPFL), CH-B Ecublens, 1015 Lausanne, Switzerland [email protected] Dick B. Janssen, Department of Biochemistry, University of Groningen, Nijenborgh 4, Groningen, 9747AG, The Netherlands [email protected] Jerome J. Kukor, Biotechnology Center for Agriculture and the Environment and Department of Environmental Sciences, Cook College, Rutgers, The State University of New Jersey, 59 Dudley Road, New Brunswick, NJ 08901-8520, USA [email protected] M. Minna Laine, Science Support/Biosciences, CSC-Scientific Computing Ltd., P.O. Box 405, FIN-02101 Espoo, Finland [email protected] Frank E. Löffler, School of Civil and Environmental Engineering, Georgia Institute of Technology, 200 Bobby Dodd Way, 202 DEEL, Atlanta, GA 30332-0512, USA [email protected] Mary M. Lynam, School of Public Health, The University of Michigan, Ann Arbor, 48109 Ml, USA [email protected] viii

Minna K. Männistö, Arctic Microbiology Research Consortium, Metla Rovaniemi Research Station, P.O. Box 16, 96301 Rovaniemi, Finland [email protected] Kyoungphile Nam, School of Civil, Urban & Geosystem Engineering, Seoul National University, San 56-1, Shinlim-dong, Kwanak-gu, Seoul 151-742, The Republic of Korea [email protected] Jantien E. Oppentocht, Department of Biochemistry, University of Groningen, Nijenborgh 4, Groningen, 9747AG, The Netherlands Ronald S. Oremland, U.S. Geological Survey, 345 Middlefield Rd., Menlo Park, CA 94025, USA [email protected] Gerrit J. Poelarends, Department of Biochemistry, University of Groningen, Nijenborgh 4, Groningen, 9747AG, The Netherlands Jaakko A. Puhakka, Institute of Environmental Engineering and Biotechnology, Tampere University of Technology, P.O. Box 541, FIN33101 Tampere, Finland [email protected] Christophe Regeard, Laboratory for Environmental Biotechnology, Swiss Federal Institute of Technology (EPFL), CH-B Ecublens, 1015 Lausanne, Switzerland Kirsti M. Ritalahti, School of Civil and Environmental Engineering, Georgia Institute of Technology, 200 Bobby Dodd Way, 202 DEEL, Atlanta, GA 30332-0512, USA Mirja S. Salkinoja-Salonen, Department of Applied Chemistry and Microbiology, University of Helsinki, P.O. Box 56, FIN-00014 University of Helsinki, Finland [email protected] James M. Tiedje, Center for Microbial Ecology, Michigan State University, 540 Plant and Soil Science Building, East Lansing, Ml 48824-1325, USA [email protected] Lisa A. Totten, Department of Environmental Sciences, Rutgers, The State University of New Jersey, 14 College Farm Road, New Brunswick, NJ 08901-8851, USA [email protected] L.Y. Young, Biotechnology Center for Agriculture and the Environment and Department of Environmental Sciences, Rutgers, The State University of New Jersey, 59 Dudley Road, New Brunswick, NJ 08901-8520, USA [email protected]

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PREFACE Halogenated organic compounds constitute one of the largest groups of environmental chemicals. The industrial production of new halogenated organic compounds has increased throughout the last century peaking in the 1960s, and continuing in widespread use today. Organohalides are integral to a variety of industrial applications, including use as solvents, degreasing agents, biocides, pharmaceuticals, plasticizers, hydraulic and heat transfer fluids, and intermediates for chemical synthesis, to name a few. It is important to recognize the beneficial aspects of halogenated organic compounds, as well as their potentially deleterious impact on the environment and health. Recognition ofthe adverse environmental effects of many types of organohalide compounds has led to efforts to reduce or eliminate the most problematic ones. Although organohalide compounds are typically considered to be anthropogenic industrial compounds, they have their counterpart in several thousands of natural biogenic and geogenic organohalides, representing most classes of organic chemicals. Natural sources account for a significant portion of the global organohalogen budget. This volume authored by recognized experts in the field provides a current perspective on how both natural and synthetic organohalides are formed and degraded, and how these processes are incorporated into a global halogen cycle. The focus is on microbial processes, since these play a major role both in the production and degradation, i.e., cycling of halogenated organic compounds in the environment. This book is organized into five parts. Part I, Introduction, provides a global perspective on the issues of organohalides and their fate in the environment. The unique physicochemical properties that also impart resistance to both chemical and biological degradation is one of the qualities that has made many organohalides useful in industrial applications, but it is also the reason for many of the environmental problems related to the use of these compounds. Part II, Microbial Processes provides an overview of the diversity of dehalogenating microorganisms, and brings together our current understanding of microbially mediated processes for the biodegradation of organohalides. A critical step in the degradation of organohalides is cleavage of the carbon-halogen bond, and microorganisms have evolved a variety of metabolic strategies for dehalogenation, which include oxidation, reduction, substitution, intramolecular substitution, dehydrohalogenation, hydration, and methyl transfer reactions. The list of organohalides that can be utilized by microbes continues to increase dramatically, as do the number of dehalogenating microorganisms that have been identified and characterized. The chapters in this section of the book examine the ecology and diversity of dehalogenating bacteria, archaea, and fungi, including the thermodynamics of microbial dehalogenation. Part III, Biochemistry and Chemistry explores the biochemistry and genetics of the dehalogenating micoorganisms. The chapters provide an overview of the range of biologically-mediated dehalogenation mechanisms. In addition, some of the abiotic processes involved in dehalogenation are examined. Part IV, Environmental Fate and Applications takes a more detailed look at some of the problematic organohalides, such as pesticides, chlorofluorocarbons, chlorinated solvents, polychlorinated dibenzo-p-dioxins and polychlorinated biphenyls. The chapters

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in this section discuss the fate of these compounds in the environment and explore strategies for the development of bioremediation technologies for organohalidecontaminated sites. Part V, Summary provides a look at future research needs for understanding the role of organohalides in the environment and their influence upon human health. Detailed information on biodegradation and biotransformation mechanisms for a variety of organohalides and on the microorganisms mediating these processes has greatly increased our understanding of the cycling and fate of these unique and widespread compounds in our environment. We believe that this volume will serve as a comprehensive reference source for information on microbial degradation of halogenated organic compounds, in particular, as a resource on the microbial processes and applications of dehalogenation. We thank our colleagues for their efforts and excellent contributions, and for their patience during the editing process. We are grateful to Melinda Paul, Editor at Kluwer Academic Publishers, for interest in this project and cooperation in seeing this volume to fruition. Max M. Häggblom Ingeborg D. Bossert

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PART I. INTRODUCTION

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Chapter 1 HALOGENATED ORGANIC COMPOUNDS A GLOBAL PERSPECTIVE MAX M. HÄGGBLOM 1,2 AND INGEBORG D. BOSSERT 1 1

Department of Biochemistry and Microbiology and 2 Biotechnology Center for Agriculture and the Environment, Rutgers University, New Brunswick NJ, USA

1. INTRODUCTION Halogenated organic compounds constitute one of the largest groups of environmental chemicals. Their use and misuse in industry and agriculture represent a large entry of these chemicals into the environment, resulting in widespread dissemination and oftentimes undesirable conditions, i.e., environmental contamination. It is important to recognize the beneficial aspects of halogenated organic compounds, as well as their potentially deleterious impact on the environment. While it is recognized that a large number of synthetic organohalides arise from anthropogenic activity, it is equally important to note that “naturally produced” organohalides abound in nature and have been present on earth for eons. In a discussion of the global cycling of halogenated organic compounds, it is necessary to consider all aspects of this diverse and biologically challenging group of compounds, including their production, biodegradation, assimilation, integration (e.g., sorption and coupling to organic matter), and also their persistence in the environment. Microorganisms impact each of these processes, and therefore play an essential role in the global cycling of organohalides. This chapter provides an overview of each of these aspects of global cycling, and the ultimate fate of organohalides in the environment. In many respects, the chemistry of halogenated organic compounds is due to the unique physicochemical properties of their halogen substituent (F, Cl, Br, or I). At the start of the series, the carbon-fluorine bond is very strong with high polarity. With increasing molecular weight of the halogen, carbon-halogen bond energies decrease markedly, i.e., F > Cl > Br > I. Other characteristics, such as the electron-withdrawing effect of the halogen substituent impact chemical reactivity of the molecule and its heat transfer and dielectric characteristics (e.g., polychlorinated phenols; polychlorinated biphenyls, PCBs). The physical size and shape of the halogen substituent may also affect reactivity, due to steric constraints and may also hinder uptake into cells and enzymatic attack during biodegradation. In addition, the halogen moiety of an organic compound Dehalogenation: Microbial Processes and Environmental Applications, pages 3-29 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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generally reduces its water solubility and conversely increases lipid solubility. The biological consequence of increased lipophilicity may be reduced biodegradation due to decreased bioavailability, and/or biomagnification in the food chain as the nondegraded haloorganic compounds sequester in the fatty tissues of higher animals. Finally, the halogen substituent and its potential organohalide metabolites may alter, oftentimes increasing, the inherent toxicity of the molecule. The latter characteristic is apparent both in synthetic, xenobiotic (foreign to the biosphere) compounds, such as toxic dibenzo-p-dioxins, chlorophenols, PCBs, and halogenated aliphatic compounds (tetrachloroethene, PCE; trichloroethene, TCE; vinyl chloride, VC), as well as in biologically produced natural products, such as antibiotics (e.g., chlortetracycline, chloramphenicol, drosophilin, bromopyrroles) and other antibiologicals, produced by both micro- and macro- organisms for protection against competition and predation. The discovery of chlorine and other halogens, and the elucidation of their unique chemistry was followed by their synthesis and large-scale industrial production and application (see 162). The scale of production (past and present) of these organohalide compounds has had direct implications for their occurrence and fate in the global environment. Organohalides are integral to a variety of applications, including use as solvents, degreasing agents, biocides, Pharmaceuticals, plasticizers, hydraulic and heat transfer fluids, intermediates for chemical synthesis, and numerous other industrial functions. Other halogenated compounds are produced as by-products during combustion, chlorine bleaching of pulp, or disinfection of water and wastewater. As a result, many halogenated organic compounds, including aliphatic, aromatic and heterocyclic derivatives, have been produced and used in vast quantities over the last 50 to 80 years. The majority of these compounds are chlorinated, but brominated, fluorinated and iodinated compounds also have industrial applications. Historically, with increasing use, a number of the negative properties often assoicated with organohalide compounds had become widely evident in the 1960s. With the growing use of industrial chemicals, in particular organohalides, and their oftentimes indiscriminate dissemination in the environment, concern increased over the potential adverse effects of organohalides. This was brought to public attention in 1962 by Rachel Carson in her seminal work, “Silent Spring” (19). Further evidence for the widespread occurrence of organohalides has emerged from the development of improved sampling and analytical methods with greater sensitivity, in particular the invention of the electron capture detector for gas chromatography by James Lovelock in 1961 (104). With increased detection sensitivities, industrial organohalides have been increasingly found as trace contaminants broadly disseminated in the environment. The initial recognition of the persistence and potential toxicity of different organohalides, and their propensity to bioaccumulate in the food chain, among other adverse effects, has led to extensive efforts to replace halogenated chemicals today. In cases where the unique physicochemical properties and/or economics of halogenated organic compounds are difficult or impossible to match with non-halogenated alternatives, greater care is exercised in their use and disposal. Current and improving regulatory guidelines and mandates will help to further control the potential adverse effects of organohalides in the environment. Introduction of industrial halogenated compounds into the environment occurs through terrestrial, aquatic and atmospheric discharges. Therefore, their impact is on all

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major environmental compartments, i.e., soils sediments, water, and air. Depending on their ultimate fate, organohalides may be degraded to harmless byproducts, or they may exert harmful effects through toxicity, biomagnification, and/or persistence in the environment. Their harmful impact on the biota may be direct, i.e., toxicity, or indirect, such as by destruction of the protective ozone layer in the stratosphere by atmospheric halocarbons. Owing in part to their often xenobiotic origin and persistent character, many industrial organohalides are resistant to biodegradation, and therefore accumulate and exert their harmful effects in the environment. Within the past decade, as greater focus and research efforts have been placed on determining the fate of these compounds, a more comprehensive understanding of the environmental fate and impact of halogenated organic compounds has been gained. Although organohalide compounds were initially considered anthropogenic (i.e., man-made or synthetic) contaminants with limited or no natural counterparts and thus inherently problematic, this line of thinking is contrary to the scientific evidence. A number of natural biogenic and geogenic sources have now been identified and are further discussed in section 2 of this chapter. Biogenic producers of organohalides have been found in both marine and terrestrial environments and include bacteria, fungi, plants, sponges, insects and mammals (49, 61, 62, 126, 155, 182). Geogenic sources of organohalides include forest fires and other pyrogenic processes, including volcanic emissions (80,86). For example, organochlorine compounds, including polychlorinated phenols, dibenzo-p-dioxins and dibenzofurans, are produced during combustion of organic material containing inorganic chloride (79, 143). There is now an increasing understanding of how natural organohalides are formed and degraded in the environment and how anthropogenic organohalides are incorporated into a halogen cycle. A brief overview of this halogen cycle, as depicted in Figure 1.1, illustrates how biotic and abiotic processes contribute to the overall fate of halogens in the environment. Of particular interest, especially in the context of this text, is the

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extensive role that microorganisms play in the biogeochemical cycling of halogens, and the movement of halogenated organic compounds through the global carbon cycle. These interrelated and often complementary processes provide an effective means for reducing the environmental load of harmful organohalides. Microorganisms are the major mediators for the cycling of halogenated organic compounds in the environment. It is therefore essential to fully delineate and understand the diverse microbial roles in biodegradation and biotransformation processes. The nature of the carbon-halogen bond is a key factor, that combined with other structural properties of the molecule, will affect the reactivity of organohalides. Halogens are strongly electronegative; the electron-withdrawing effect of halogen substituents increases electrophilicity of the carbon atom. The carbon-halogen bond strength decreases with increasing molecular weight. For example, the carbon-fluorine bond energy is among the highest found in natural compounds, and is also more polar than carbon-chlorine and carbon-bromine bonds. The ionic radius of the halogen substiutent also affects metabolism of the organohalide substrate. For example, the bulkiness on bromine and iodine substituents may in some cases limit metabolism. The combination of physicochemical properties can therefore greatly influence bioavailability and the metabolic pathways for biodegradation and biotransformation (see Chapter 2). Extensive progress that has been made in the last few years towards a better understanding of the mechanisms of microbial action on halogenated compounds, in both oxic and anoxic environments. These are summarized in the remaining chapters of this book.

2. NATURAL SOURCES OF ORGANOHALIDES 2.1. Biogenic Sources A great variety of halogenated compounds are produced naturally. Since they are part of the natural carbon cycle, halogenated organic compounds should not be considered solely as anthropogenic contaminants. Natural organohalides were originally considered to be biosynthetic oddities of nature; only 30 organohalide compounds had been identified by 1968 (60). However, over 3,000 naturally-produced organohalides have been identified to date, in almost all classes of organic chemicals (60, 61). Brominated and chlorinated organohalides are produced most abundantly, iodinated compounds are found less frequently, while fluorinated metabolites are very rare (even though fluorine is the most abundant halogen found in the earth’s crust) (92, 133). The organisms that produce organohalides represent a diverse group, and encompass both eukaryotic and prokaryotic kingdoms. These organisms include bacteria, fungi, lichens, marine sponges, worms, insects, and mammals (49, 60-63, 126, 182, 183). A few examples are listed in Table 1.1 and highlighted below. Not surprisingly, the saline marine environment is an incredibly rich source of naturally occurring halogenated compounds. Organohalides are produced by a diversity of marine organisms, including mollusks (10), algae (142), polychaetes (3,54), jellyfish (189), and sponges (57, 151). Marine macroalgae, e.g., kelp, produce a wide range of volatile chlorinated, brominated and iodinated hydrocarbons, the most abundant of which is bromoform (59). A number of sponge species in the phylum Porifera, in particular Aplysina sp., have been shown to produce an amazing variety of brominated

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metabolites, including bromoindoles, bomophenols, polybrominated diphenyl ethers, and even brominated dibenzo-p-dioxins (43, 62, 128, 129, 171, 173). Bromine-containing metabolites can account for up to 7 to 12 percent of the sponge dry weight (171). These compounds may serve as a chemical defense against predators and inhibit biofouling. In addition, Aplysina sponges harbor large amounts of bacteria which can amount to 40% of the biomass of the animal (73), and it has been hypothesized that some of the organobromine compounds may in fact be synthesized by bacteria associated with the

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sponge. Chlorinated sponge metabolites are relatively rare (49). Acorn worms (phylum Hemichordata) represent another group of marine animals with an incredible capacity to produce organobromine compounds. 2,6-Dibromophenol, 2,4,6-tribromophenol and brompyrroles are produced by various species, apparently as defensive agents against predation (24, 54, 55, 93, 94, 192). The concentration of brominated phenols may reach several hundered per liter in the burrow lining of these worms (93). Another interesting example of biologically-mediated production of halogenated phenols, in this case by a non-marine land animal, is the production of 2,5dichlorophenol, a compound that is emitted in the defensive froth of the grasshopper Romalea microptera (46). Halogenated phenols do not necessarily always serve as an inhibitor or deterant. The Lone-Star tick and other tick species, produce 2,6dichlorophenol as an attractant, in this case, as a sex pheromone (15, 16, 112). Fungi have also been shown to produce a variety of chlorinated organic compounds in terrestrial environments (see Chapter 6). The basidiomycetes have a widespread capacity for organohalogen biosynthesis, in particular chlorinated aromatic and aliphatic compounds (32-34). For example, chlorinated anisyl metabolites are produced in situ by wood and forest litter degrading fungi, and have been found at concentrations of 75 mg per kg of the wood or litter substrate (33). In addition to the individual compounds thus far identified, a large pool of halogenated organic material, defined analytically as the sum parameter of organically bound halide in the form of adsorbable organic halogen (AOX) or extractable organic halogen (EOX), has been detected. Recent estimates suggest that up to 3 g of AOX per kg of mycelium (dry weight) may be produced in some fungal species (34), with some strains producing organic halogens up to 3% of their biomass dry weight (185). Verhagen et al. (185) have calculated that 2 kg AOX per hectare would be produced annually in a peat bog with a Hypholoma mycelial biomass of 80 kg per hectare. Naturally-formed chlorinated phenolics may further be transformed by fungal peroxidases to yield dimeric compounds such as chlorinated phenoxyphenols, dibenzop-dioxin and dibenzofurans (119,130,132,166). Laccase from the fungus Rhizoctonia practicola has been shown to transform 2,4-dichlorophenol to chlorodiphenyl ethers (119), Similarly, polychlorinated dibenzo-p-dioxins and dibenzofurans are formed from 2,4,5- and 3,4,5-trichlorophenol by peroxidase-catalyzed reactions (131,166), indicating that biogenic formation of these highly toxic compounds is possible (see also Chapter 16). The natural formation of chlorinated dibenzo-p-dioxin and dibenzofurans has been detected in soil of a Douglas fir forest (74). A recent study suggests that vinyl chloride is formed in terrestrial environments during oxidative degradation of organic matter by a variety of abiotic and biotic processes (90). Combined, these findings explain how organohalogens (AOX and EOX) have been widely found in a variety of “unpolluted” environments, including groundwater, surface water, and soil (8, 9, 56).

2.2. Geogenic Sources The geological origins of halogenated organic compounds are generally associated with high temperature and pressure, such as found in volcanic eruptions and forest fires. Under these extreme conditions, inorganic sources of halogen salts, in particular chloride, may combine with organic molecules to form organohalides. A range of

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halogenated organic compounds have been detected in lava gas, including chlorinated, fluorinated, brominated, and iodinated compounds (60,80,86). HCl and HF likely react with organic compounds to produce organohalogens. Chlorinated methanes, tetrachloroethane, trichloroethene, and chlorobenzene are some of the most abundant organohalogen species that have been detected in lava gas samples (86). However, in the case of chlorofluorocarbons (CFCs), it is estimated that emissions from volcanoes are negligible compared to the anthropogenic CFC burden. Halogenated organic compounds are similarly generated during combustion of organic material in the presence of chloride and other halides, e.g., burning of fresh wood or incineration of municipal solid waste (1, 88, 137, 168). It has been estimated that approximately 20 to 50 thousand tons of bromomethane are produced by burning biomass, which corresponds to approximately 30% of the stratospheric bromine budget (62). In comparison, marine emissions of bromomoethane are estimated at over 50 thousand tons a year, and that of bromoform is estimated at 1 to 2 million tons a year (62). Chloromethane is the most abundant volatile organohalogen in the atmosphere, with estimates of natural (mainly biogenic) emissions at 3 to 8 million tons a year, a value two orders of magnitude more than that estimated from anthropogenic sources (34). In addition to halogenated aliphatic compounds, halogenated aromatics are also generated during combustion. For example, terra- and pentachlorophenols are generated during combustion of fresh wood (1). During combustion, organohalides that are formed may volatilize or sorb onto fly ash, resulting in widespread airborne distribution. For example, chlorophenols most likely produced during forest fires are ubiquitously distributed in remote pristine areas, and have been found in lake sediments located far from industrial sites (149). Pentachlorophenol has been found in sediment layers older than 50 years, dating to before industrial production, and presumably originating from forest fires. Polychlorinated dibenzo-p-dioxins can be produced from halogenated precursors (such as chlorophenols) by surface-catalyzed combustion reactions (79, 88, 168). Their formation may also proceed from chlorinated phenols via biologically mediated peroxidase reactions as discussed in Section 2.1.

3. ANTHROPOGENIC SOURCES OF ORGANOHALIDES The industrial production of organohalogen compounds has increased throughout the last century. During this time period, industrial haloorganics have been found increasingly as contaminants in the environment, especially with improved methods for detection and quantifitation. The industrial production of new halogenated organic compounds peaked in the 1960s, and their widespread use continues to grow today. For example, world chlorine consumption in 1987 totaled 34.5 million tons, with over 60% used in organic synthesis (162). This has increased to over 48 million tons in 1996, and is increasing at an estimated rate of 3.4% per year. Of the many haloorganic productsproduced, polyvinyl chloride (PVC), a widely used polymer in plastic production, represents the largest single end-use of chlorine. PVC manufacture accounted for 36% of global chlorine consumption in 1995, and is projected to reach 50% by 2005 (162). Published values (22) for the production of some main industrial organohalides are listed in Table 1.2. The two PVC pre-cursors, 1,2-dichloroethane

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(ethylene dichloride) and chloroethene (vinyl chloride) comprise the top two organohalogen compounds produced industrially, with a combined worldwide annual production of over 20 million tons. Of these two major products, 1,2-dichloroethane production in the United States has grown steadily over the last decade to approximately 10 million tons in 2000 (23). As shown in Table 1.2, the top chlorinated aromatic compounds are chlorobenzenes, used as solvents, intermediates, and biocides. The annual production (estimated in 1982) of chlorobenzene was 600 million kg, in addition to over 200 million kg of di- to hexachlorobenzenes (141). Other industrial compounds include a wide variety of chlorinated hydrocarbons (such as chloromethanes, chloroethanes, chloroethenes and chlorobenzenes), chlorofluorocarbons, and halogenated phenolics, representing products with diverse end uses inlcuding solvents, degreasers, fumigants, biocides, dielectric fluids, flame retardants and chemical intermediates.

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3.1. Chlorine Disinfection and Bleaching Water disinfection using chlorine accounts for about 6% of the global chlorine production (162). Since its introduction in 1908, chlorination of drinking water has brought about a dramatic reduction in the occurrence of water-borne diseases. Aside from its beneficial use as a disinfectant, chlorination can lead to the formation of a variety of chlorinated byproducts, especially trihalomethanes, such as chloromethane and chloroform. These undesirable byproducts arise from reactions with humic and fulvic acids that are present in the water prior to treatment, and have been associated with mutagenicity of drinking water (162). Similarly, a wide range of chlorinated compounds, including both low- and highmolecular weight material, is formed during the chlorine bleaching of pulp for paper manufacture (100). Several hundred chlorinated compounds have been identified in bleach plant effluents, ranging from simple compounds such chlorinated aliphatic acids and phenolics, to a poorly defined class known as “chlorolignin” (85, 100, 114, 138, 165). Significant levels of chlorinated dioxins and dibenzofurans have also been found (167). Modifications in the bleaching process over the last decade have substantially reduced the formation of these harmful compounds. Although the discharge of organochlorine compounds into the environment persists, it occurs at significantly lower levels.

3.2. Solvents Chlorinated solvents are necessary for many industrial applications. Although current practices follow stringent guidelines regarding their use and disposal, past practices were often haphazard at best and as a result, halogenated solvents are a major contaminant in many environments, especially the subsurface. PCE and TCE, and their metabolites, dichloroethene (DCE) and VC, are among the more common halogenated contaminants. These and related compounds, such as chloroform (CF) and carbon tetrachloride (CT), are excellent degreasing agents and have been widely used in cleaning operations, ranging from aviation mechanics to residential dry-cleaning. Accidental spills and leaks, as well as past indiscriminate disposal practices, have resulted in their spread throughout the environment. Once released into the environment, the physical properties of these halogenated compounds often render them fairly resistant to biodegradation. Typical physicochemical characteristics, such as a high specific density, poor water solubility, and low vapor pressure, increase mobility to further disseminate halogenated contaminants throughout the environment, including the subsurface and atmosphere. These same characteristics can also affect the inherent biodegradability of a compound, and thereby provide a two-fold impact on the biological fate of halogenated solvents in the environment, in that they partition into environmental compartments which are less hospitable to microorganisms, and may also be less supportive of microbial activity (biodegradation).

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3.3. Biocides Biocides, by definition, are toxic to life. Of the most widely-used biocides, e.g., pesticides, many are halogenated compounds. The majority of these compounds are chlorinated, but brominated and fluorinated organic pesticides are also in use. These compounds, which have been designed and are used for their biocidal properties, are deliberately introduced into the environment, and therefore are of a priori environmental concern. Whereas some non-target haloorganics inadvertently exert unwanted and often deleterious effects in a particular environment, other haloorganics, such as bacteriocides, herbicides, insecticides, and other types of microbicides and pesticides, may be intentionally applied to control unwanted populations, including pathogens, weeds, and insect pests, respectively. Their effect may be broad or very targeted to specific biota, therefore their uncontrolled release may have serious impact on the environment. The use of synthetic pesticides is less than a century old, yet the impact of pesticides on the environment is far-reaching. Perhaps the best-known pesticide, DDT (1,1 -bis[pchlorophenyl]-2,2,2-trichloroethane) has been in use since 1942 as an agricultural and domestic insecticide, with cumulative world production estimated at up to 3 million tons (25). Likewise, toxaphene is an insecticide used since 1948 with cumulative world production estimated at 1.3 million tons. Relative to the application rates of these insecticides, their production numbers are large, and represent a significant impact on the environment. There has been a marked increase in both the number and amounts of different pesticides applied in the last 40 years. For example, the annual use of pesticides on croplands and pastures in the United States increased from less than 90 million kg (based on active ingredient) in 1964, to an estimated 300 million kg in 1993 (11, 101). The total annual use of different pesticides in the United States in agricultural and nonagricultural settings is approximately 500 million kg (11). Contamination of groundwater and surface waters by pesticides has become a major concern throughout the world. Because pesticides are deliberately introduced to the environment, it is important to understand the consequences and potential adverse effects of such environmental contamination. Recognition of the adverse environmental effects has provided impetus for replacing problematic halogenated pesticides with compounds of lesser environmental concern, which are either less- or non-halogenated. These are generally more biodegradable and persist for a shorter period of time in the environment. 4. EVIRONMENTAL FATE The introduction of oranohalide compounds into the environment may occur through direct release, industrial emissions, or by indirect and non-industrial sources. Accurate data on the quantities of organohalides released into the environment are relatively scarce (162). Their eventual fate in the environment will depend on their behavior and mobililty in terrestrial, aquatic and atmospheric compartments. Volatilization of halogenated compounds increases their distribution into the atmosphere, where secondary effects, such as ozone-depletion, take on a global impact of haloorganics on the environment. The global distribution of many persistent haloorganic pollutants, for example DDT, PCB, dioxins, and halogenated flame retardants, is driven by a process

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known as the “global distillation effect”. This climatic phenomenon helps to distribute organohalide contaminants through volatilization and condensation mechanisms on a global scale, resulting in a greater accumulation at colder higher latitudes (4, 81, 106, 154, 162). Not only volatile compounds (e.g., chlorofluorocarbons), but also semivolatile compounds, such as DDT, PCBs, and chlorinated dioxins are transported in the atmosphere, both in the gaseous phase and adsorbed onto particles. The polar regions are environmental sinks for these contaminants through a combination of geographical and meteorological phenomena (12, 35). Many halogenated organic compounds of industrial origin can manifest harmful effects on the environment by acute or chronic toxicity to the exposed biota, biomagnification, and/or persistence. Some chlorinated industrial chemicals, including heat exchanger fluids (PCBs) or solvents such as TCE and CF, do not have intended uses in the environment-at-large (in contrast to biocides), but rather are released though accidental spills and leaks or through poor management practices. Flame retardants are yet another class of haloorganic compounds which, owing to their widespread use, present potential harm to the environment. Other potentially toxic organohalogens, such as polychlorinated dibenzo-p-dioxins, are inadvertent byproducts from industrial combustion sources or bleaching of pulp for paper manufacture. Recognition of the adverse environmental effects of many of the various organohalide compounds (e.g., CFCs, PCBs, dioxins, persistent pesticides) has led to efforts to reduce or eliminate the most problematic ones. International agreements (Montreal Protocol in 1987 and subsequent amendments) have recently been negotiated for the phase-out of ozone-depleting chemicals, in particular, selected volatile halohydrocarbons, i.e., chlorohydrocarbons (see Chapter 12). The Stockholm Convention signed in 2001 (161) is a global treaty to protect human health and the environment from persistent organic pollutants (POPs). POPs are defined as chemicals that remain intact in the environment for long periods, become widely distributed geographically, accumulate in the fatty tissue of living organisms, and are toxic to humans and wildlife. The twelve compounds that are considered particularly hazardous and covered by this treaty are all organochlorine compounds, produced for use as pesticides or industrial chemicals, or are unintended byproducts of combustion and industrial manufacturing (Table 1.3). Resistance to both chemical and biological degradation is one of the qualities that has made many organohalides useful in industrial applications, but it is also the reason for many of the environmental problems related to the use of these compounds. The persistence of organohalides in the environment varies from days to several decades or centuries (3), depending on the chemical structure and the prevailing environmental conditions, and can play a major role in their overall global impact.

5. BIODEGRADATION OF ORGANOHALIDES – MICROBIAL MEDIATORS OF THE HALOGEN CYCLE Microbial degradation is one of the key factors that determine the ultimate fate of organohalides in the environment. The work of Kluyver and van Niel (see 96) was central in advancing the concept of the pivotal role of microoganisms in the biodegradation of organic material. From their early studies, it was recognized that all

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naturally-occurring organic compounds can be biodegraded in some fashion by the diversity of microorganisms found on earth. This concept was further expanded by Martin Alexander (2), with his “principle of microbial infallibility”, to articulate the remarkable adaptability of microorganisms and their ability to biodegrade synthetic compounds, often novel structures not found in nature. Microbial evolution over the last 3.6 billion years, has provided ample time for the evolution of metabolic pathways necessary for the biodegradation of most halogenated compounds. Biodegradation of anthropogenic chemicals is achieved when these compounds can feed into similar and relevant enzymatic pathways that are already present for the biodegradation of natural compounds (31). The current evidence supports and furthers this early thinking, by helping to define mechanisms such as mutation and gene transfer that develop new pathways for biodegrading the xenobiotic contaminants introduced by humans in the last century (177, 178). These are discussed in greater detail in Chapters 7 and 8. A critical step in the degradation of organohalides is cleavage of the carbon-halogen bond. Microbial degradation requires the presence of enzymes that cleave this bond under physiological conditions. Based on the long-standing natural biogenic and geogenic processes that form halogenated organic compounds, it is therefore not surprising that many recent organohalides which are synthetically-derived have been shown to be biodegradable under some environmental conditions. The importance of microbial degradation of organohalides is reflected in the large number of excellent reviews published over the last 20 years (27, 47, 50, 51, 64, 66, 69, 72, 76, 83, 97, 102, 122, 144, 145, 156, 163, 177, 186, 187). In fact, den Dooren de Jong (36) first

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demonstrated in 1926 that brominated aliphatic acids could be used as carbon sources by bacteria (referenced in 31, 139, 157). Further evidence indicates that the biodegradation of anthropogenic halogenated organic compounds has basis in two phenomena: 1) the fortuitous degradation of structures analogous or similar to naturallyoccuring compounds, using existing pathways for naturally-occurring organohalides, or 2) a selective genetic transfer, amalgamation or mutation, so that new pathways evolve from existing ones. It should be noted that these processes are not exclusive of one another, but rather provide complimentary means for establishing new and efficient metabolic pathways for the biodegradation of new and novel halogenated organic compounds. The growing list of organohalides that can be utilized by microbes continues to increase dramatically, as can be found in the remaining chapters of this book. Microorganisms have evolved a variety of metabolic strategies for cleaving the carbon-halogen bond. The range of biologically-mediated dehalogenation reactions and processes are summarized in a number of key reviews (47, 50, 51, 64, 72, 76, 83, 122, 144, 163, 186, 177, 187). Examples of currently recognized dehalogenation mechanisms, substrates, and their mediating organsims are presented in Table 1.4. These biodegradation mechanisms include oxidation, reduction, substitution, intramolecular substitution, dehydrohalogenation, hydration, and methyl transfer reactions. In addition to specific enzymes that catalyze dehalogenation (i.e., dehalogenases), the carbonhalogen bond can also be cleaved spontaneously via production of unstable metabolites, or by fortuitous reactions. Oxidative dehalogenations are catalyzed by mono- and dioxygenases, in reactions that incorporate either one or two oxygen atoms from into the substrate. Oxidative dehalogenation is thus restricted to oxic environments. Unlike oxidative dehalogenation, which is restricted to aerobes operating in containing environments, other dehalogenation mechanisms generally operate under a wide range of redox conditions, from aerobic to highly reduced anaerobic conditions. Depending on the prevailing redox environment and availability of electron acceptors and donors, halogenated compounds offer a wide variety of substrate types that may either accept or donate electrons to microbially mediated biochemical reactions. Dehalogenation reactions comprise different strategies, where organohalides serve either as electron donors (and carbon sources) or electron acceptors, or undergo exchange reactions, as follows: 1) the organohalide serves as a carbon and energy source and dehalogenation occurs in order to break down the carbon backbone, 2) the organohalide serves as an alternate electron acceptor for anaerobic respiration, termed (de)halorespiration, 3) dehalogenation occurs as a detoxification mechanism, or 4) the organohalide is dehalogenated through fortuitous reactions that do not yield any benefit to the organism. Structural features of a particular compound are central in determining which dehalogenation mechanisms are possible for the overall biodegradation of the compound. Metabolic strategies for degradation vary between types of halogenated organic compounds (e.g., aliphatic vs. aromatic), as well as redox environments, (e.g., aerobic vs. anaerobic) and the availability of electron donors and acceptors. A large variety of microbes have evolved that utilize these different strategies to degrade organohalides. The following two sections provide a brief overview of aerobic and anaerobic microorganisms, and their abilities to metabolize halogenated organic compounds.

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5.1. Dehalogenation Strategies for Aerobic Microorganisms Numerous aerobic bacteria and fungi use halogenated compounds as a carbon and energy source for growth. Initial attack on the carbon skeleton is typical for the degradation of aliphatic and aromatic organohalides by aerobic microorganisms, followed by dehalogenation during subsequent metabolism. In some cases, only partial biotransformation or cooxidation occurs. Under those circumstances, the organohalide may be cooxidized, or cometabolized, in the presence of an additional substrate that provides growth and energy for the microorganism. Aerobes have also adapted some of mechanisms as a means to degrade organohalides, including reductive and hydrolytic dehalogenation. The dehalogenation reactions involved in the degradation of aliphatic halides are diverse (see Table 1.4 and Chapter 7). Hydrolytic and substitutive dehalogenase enzymes are commonly found in many types of organohalide-degrading microorgansims, and have been studied in detail (51, 72, 83, 156, 157). Oxygen incorporation during metabolism of aliphatic halides can result in the formation of corresponding epoxides or halohydrins, which spontaneously decompose with the elimination of halide. This is typically a fortuitous, or cometabolic (cooxidative), transformation that does not benefit the organism. The transformation of trichloroethene and other haloethenes, for example, is mediated by several mono- and dioxygenases (e.g., methane monooxygenase, ammonia monooxygenase, phenol hydroxylase, toluene monooxygenases, and toluene dioxygenase), that exhibit broad substrate range (82, 134, 170, 181, 188, 193). For halogenated aromatic compounds, two main degradation strategies are used to cleave the halogen-carbon bond: 1) dehalogenation as an initial step in degradation via reductive, hydrolytic, or oxygenolytic mechanisms, or 2) dehalogenation of an aliphatic intermediate after cleavage of the aromatic ring. In some organisms, initial dioxygenase attack yields a halogenated cis-dihydrodiol, with spontaneous elimination of halide during reductive rearomatization, to yield anon-halogenated catechol product. For example, two-component dioxygenases in Pseudomonas strains catalyze the dehalogenation of 2-halobenzoates (18, 48, 52, 53, 118) via dehalogenation from the dihydrodiol intermediate. Following a different strategy, most mono- and dichlorinated aromatic compounds (e.g., phenols, benzoic acids, benzenes) are degraded by initial dioxygenase attack without dehalogenation to yield the corresponding halocatechols (for reviews, see 64, 97, 144, 177, 178). Halogenated catechols are thus central intermediates in the aerobic degradation of many mono- and dihalogenated aromatic compounds. Dehalogenation proceeds after oxidative ring cleavage, either through the formation of unstable metabolic intermediates (e.g., during lactonization and isomerization reactions), or by the action of specific enzymes. The ring-cleavage step is critical, since meta-cleavage typically produces acylhalides as “suicide” or dead-end metabolites, while orthocleavage leads to a productive biodegradation route (97). One of the most extensively investigated strains is Pseudomonas sp. B13, first isolated by Dorn et al. (42) for its ability to utilize 3-chlorobenzoate. The biochemistry and genetics of the chlorocatechol pathways have been well characterized (see for reviews 144, 177, 178). In contrast, degradation of many polychlorinated aromatics, such as pentachlorophenol (PCP), by aerobic bacteria is initiated by oxygenase attack to yield

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chlorohydroquinones. The aromatic ring is typically cleaved only after oxidative, hydrolytic, or reductive removal of all or most of the halogens (64). In Sphingobium chlorophenolicum (formerly Sphingomonas chlorophenolica and Flavobacterium sp.), a soluble flavoprotein monooxygenase (PCP monooxygenase) catalyzes a NADPHdependent dechlorination to yield tetrachloro-p-hydroquinone (29, 136, 190; see also Chapter 8). In contrast, transformation of PCP to tetrachloro-p-hydroquoinone by Mycobacterium chlorophenolicum and M. fortuitum is thought to be catalyzed by a membrane-bound cytochrome enzyme (5, 174, 175). Fungi and algae also play a role in the cycling of halogenated organic compounds, most notably in the degradation of halogenated products and also in their production (see Chapter 6). White rot and other fungi produce strong and non-selective extracellular oxidative (ligninonlytic) enzymes that catalyze the fortuitous transformation of a variety of organohalides. These enzymes catalyze one-electron oxidation reactions with the formation of highly reactive radicals. Here, the degradation of organohalides is cometabolic and other substrates are needed for carbon and energy.

5.2. Dehalogenation Strategies for Anaerobic Microorganisms Reductive dehalogenation is considered to be the predominant process in the anaerobic transformation of halogenated compounds. Although it is also a strategy used by aerobic microorganisms, reductive dehalogenation of both aliphatic and aromatic organohalides is frequently observed in anaerobic environments. In addition to potentially serving as a carbon source (or cosubstrate), organohalides function as terminal electron acceptors in an anaerobic respiration process, termed dehalorespiration or halorespiration (see Chapters 3, 4 and 5). Studies on the fate of pentachlorophenol in anaerobic rice paddy soils led Ide et al. (78) in 1972 to be among the first to suggest the process of anaerobic dechlorination. In other studies, microbially-mediated reductive dehalogenation of chlorobenzoates was conclusively demonstrated in anaerobic sewage sludge and lake sediments (77, 164). Following these early studies, the enrichment of a 3-chlorobenzoate-utilizing culture eventually led to the isolation of an anaerobic dehalogenating bacterium, Desulfomonile tiedjei (37, 153), and the first demonstration that dehalogenation yielded energy that could be conserved for growth (40, 41, 120, 121; see Chapter 4). Several other anaerobic bacteria that respire by dehalogenation of organohalides have since been isolated and characterized, and are discussed more extensively in Chapters 3 and 5. This metabolic capability appears to be distributed in diverse bacterial lineages, including (but not limited to: Desulfomonile, Desulfitobacterium, Dehalobacter, Dehalococcoides, Desulfovibrio and Dehalospirillum. During their anaerobic respiration and growth, halogenated compounds such as halobenzoates, halophenols, and chloroethenes serve as electron acceptors, and hydrogen and formate are widely used as electron donors. A variety of anaerobic bacteria also use halogenated compounds as carbon sources for growth, in a wide range of electron-accepting environments. For example, the homoacetogenic bacteria, Acetobacterium dehalogenans and Dehalobacterium formcoaceticum, can utilize chloromethane and dichloromethane as carbon and energy sources (105, 169). It is now known that several types of anaerobic bacteria catalyze cometabolic reductive dehalogenations involving tetrapyrrole cofactors, flavoprotein-

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flavins, and ferredoxins (see Chapters 5 and 9). In addition, the anaerobic utilization of chlorinated aliphatic compounds has also been demonstrated under different redox conditions, with oxidation of dichloroethene and vinyl chloride coupled to Fe(III)-, Mn(IV)-, sulfate-, or carbonate-reduction (see Chapter 14). Degradation of halogenated aromatic compounds generally proceeds via reductive dehalogenation, with further degradation of the carbon skeleton after complete dehalogenation has occurred. The anaerobic mineralization of halobenzoates and halophenols coupled to nitrate-, sulfate-, and Fe(III)-reduction has also been demonstrated (58, 68, 70, 89, 98, 117, 123, 150, 184). In fact, several chloro-, bromo-, and fluorobenzoate-utilizing denitrifying strains, representative of different groups within the Proteobacteria, (e.g., Acidovorax, Azoarcus, Bradyrhizobium, Ensifer, Mesorhizobium, Ochrobactrum, Paracoccus, Pseudomonas, and Thauera), have been isolated. Their presence in many soils and sediments, indicates a wide distribution of dehalogenation processes under denitrifying conditions (71, 158, 159, 184). In another group of bacteria, phototrophic Rhodospirillum and Rhodopseudomonas species have been shown to grow on halogenated aliphatic acids and on 3-chlorobenzoate (44, 87, 113, 179). The mechanisms involved in the anaerobic degradation of organohalides by these phototrophic organisms is still poorly understood.

6. CONCLUDING REMARKS The following chapters bring together our current understanding of the microbial processes of organohalide degradation, including their biochemistry and genetics. Detailed information now available on biodegradation and biotransformation mechanisms for a variety of organohalides and on the microorganisms mediating these processes has greatly increased our understanding of the cycling and fate of these unique and widespread compounds in our environment. While the focus is on microbial degradation processes, it is also important to recognize the contribution of microorganisms to the overall production of organohalides. In addition to substrate mineralization, biodegradation processes influence the mobility of the organohalide chemical in the subsurface and surface soils and waters, and also change the toxicity and bioavailability of the compound. These aspects are covered more extensively in the subsequent chapters of this book. Knowledge gained from a better understanding of the role that microorganisms play in the biodegradation and removal of harmful halogenated compounds from our environment will provide a strong basis for evaluating potential hazards to human and environmental health, and for effective implementation of bioremediation technologies for the cleanup of contaminated sites. The future provides exciting possibilities for development of technologies for bioremediation of organohalide-contaminated sites polluted by past practices. REFERENCES 1.

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44. Egland PG, Gibson J & Harwood CS (2001) Reductive, coenzyme A-mediated pathway for 3-chlorobenzoate degradation in the phototrophic bacterium Rhodopseudomonas palustris. Appl. Environ. Microbiol. 67:13961399 45. Ehrlich J, Bartz QR, Smith RM, Joslyn DA & Burkholder PR (1947) Chloromycetin, a new antibiotic from a soil actinomycete. Science 106:417 46. Eisner T, Hendry LB, Peakall DB & Meinwald J (1971) 2,5-Dichlorophenol (from ingested herbicide?) in defensive secretion of grasshopper. Science 172:277-278 47. El Fantroussi S, Naveau H & Agathos SN (1998) Anaerobic dechlorinating bacteria. Biotechnol. Prog. 14:167-188 48. Engesser KH & Schulte P (1989) Degradation of 2-bromo-, 2-chloro- and 2-fluorobenzoate by Pseudomonas putida CLB 250. FEMS Microbiol. Lett. 60:143-148 49. Faulkner DJ (1980) Natural organohalogen compounds. In: Hutzinger O (Ed) Handbook of Environmental Chemistry, Vol 1, Part A. Anthropogenic Compounds (pp 229-254). Springer-Verlag, Berlin 50. Fetzner S (1998) Bacterial dehalogenation. Appl. Microbiol. Biotechnol. 50:633-657 51. Fetzner S & Lingens F (1994) Bacterial dehalogenases: Biochemistry, genetics, and biotechnological applications. Microbiol. Rev. 58:641-685 52. Fetzner S, Müller R & Lingens F (1989)

Degradation 53.

54.

55.

56.

57.

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PART II. MICROBIAL PROCESSES

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Chapter 2 MICROBIAL ECOLOGY OF DEHALOGENATION INGEBORG D. BOSSERT 1, MAX M. HÄGGBLOM 1,2 AND L. Y. YOUNG 2,3 1

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Department of Biochemistry and Microbiology, Biotechnology Center for Agriculture and the Environment, and 3 Department of Environmental Sciences, Rutgers University, New Brunswick NJ, USA

1. INTRODUCTION The great diversity of microorganisms and their versatile metabolism mirrors the wide range of substrates that are used for microbial growth and the natural environments in which they are found. Many types of microorganisms are able to adapt and metabolize chemically different and often relatively new industrial (xenobiotic) compounds. Halogenated organic compounds fall into these substrate categories, in that they are often novel and/or relatively new types compounds of industrial (anthropogenic) origin, yet they also include natural products from a variety of biogenic and geogenic sources (see Chapter 1). Paralleling their diverse composition and occurrence, organohalide substrates can be biodegraded by microorganisms in many different types of environments, reflecting a variety of metabolic strategies. Given this metabolic diversity, the fundamental commonality of metabolic reactions is remarkable, irrespective of substrate and organism. A unifying principle for most metabolic functions, whether they arise from prokaryotic or eukaryotic organisms, relies on two phenomena: 1) electron transfer (energy production) and 2) carbon metabolism (growth). Electron (hydrogen) transport through a series of oxidation-reduction reactions, ultimately producing ATP, provides the energy flow for biochemical and biological function. In this process, a compound serving as the electron donor is oxidized, giving up its electrons (protons) to an acceptor compound, which becomes chemically reduced. Both inorganic and organic compounds, including organohalides, can serve in these functions. Organohalide substrates can be further biodegraded, serving as a carbon source for growth. Dehalogenation reactions are often an integral part of the catabolic pathway, and together with anabolic (biosynthetic) reactions, play a most important role in the global cycling of organohalides.

Dehalogenation: Microbial Processes and Environmental Applications, pages 33-52 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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2. OVERVIEW OF THE ROLE OF ENVIRONMENT ON MICROBIAL DEHALOGENATION The microbial ecology of dehalogenation follows an overall metabolic scheme where halogenated compounds (naturally-occurring or anthropogenic) serve as electron acceptors (reduction) through dehalogenation, or as electron donors (oxidation), providing carbon for growth through degradation of the carbon “backbone”. In addition to energy-yielding, coupled redox reactions, other enzyme-mediated mechanisms, such as hydrolysis or hydrogenation, may account for dehalogenation of a substrate. Cometabolism, a phenomenon where the substrate is fortuitously dehalogenated in the presence of another growth substrate, is another microbial means for degrading organohalides, in particular those which are more biologically recalcitrant. Spontaneous dehalogenation reactions also occur in the environment through abiotic mechanisms, generally through chemical reduction by redox coupled reactions, as with oxidized metal species (see Chapter 9). Reactive metals may be present as organic species, e.g., complexed porphyrins, or as inorganic species within the mineral matrix of soils. All of the aforementioned dehalogenation processes largely define the role that microorganisms play in the ecology of a particular site, and how a particular environment will impact the fate of organohalide compounds. This has direct application to the bioremediation of sites contaminated with haloorganic compounds. By manipulating the environmental characteristics of a site, e.g., through fertilization, pH control, change in redox/electron acceptor, or addition of a secondary substrate (electron donor or cometabolite), the rate and extent of biodegradation (and dehalogenation) can be optimized. Table 2.1 provides a summary of factors affecting the biodegradation of halogenated organic compounds. These serve as guidelines for improving activity in soils, sediments, and groundwater. The information pertains primarily to soils and sediments since many organohalides are sequestered in this environmental compartment, owing to their relative water insolubility, density, and partitioning characteristics (e.g., non-aqueous phase liquids, NAPLs; 7, 70; see Chapter 10). Over the past decade, as more information on the fate of organohalides in the environment has become available, some general trends have emerged. Figure 2.1 (modified from 2,92) depicts the relationship between type of halogenated substrate and its fate in the environment (i.e., biodegradation and phase compartmentalization), as defined by its degree of halogenation (polychlorinated vs. monochlorinated compounds) and the prevailing redox conditions (i.e., terminal electron accepting processes). Although exceptions exist and may become more common as more information on dehalogenation is obtained, the schematic provides a good framework on expected dechlorination activity in a given environment with a given substrate. The actual biochemical mechanisms and the type of microbial community mediating dehalogenation reactions depend primarily on environmental constraints, as defined by the following factors: 1) redox environment, 2) substrate type and availability, and 3) electron acceptor type and availability (see Chapter 3). Conversely, these environmental factors themselves are impacted by microbial activity that can further alter the physicochemical characteristics of the environment. Other environmental factors, such as extremes in salinity, pH and temperature, and/or the presence of toxic or potentially inhibitory compounds, may play secondary roles in determining microbial

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community structure and activity. The influence and interaction of all these parameters underscores the dynamic flux and interdependence within a microbial community and the environment, thereby defining its microbial ecology. The purpose of this chapter is to provide an overview on the interrelatedness of the many types of microorganisms, their metabolism, and the interactions that occur in and with their environment, as it relates to the fate of organohalides.

3. TYPES OF DEHALOGENATING MICROORGANISMS AND THEIR ECOLOGICAL ROLES Within a given environment, the metabolic reactions that occur are due to the inherent physiological capabilities of the microbial community, the thermodynamic constraints of the prevailing redox conditions and the availability of external electron acceptors, and the availability and intrinsic biodegradability of the (halogenated) substrate. In general, microbial dehalogenation strategies comprise oxidation, elimination, reduction, and substitution (e.g., hydrolysis, methyl transfer) reactions, where organohalides serve either as electron donors or acceptors, or undergo exchange reactions. For a survey of biologically-mediated dehalogenation reactions see Chapter 1. Halogenated organic compounds offer a wide variety of substrate types, naturallyoccurring and anthropogenic, that can either accept or give up electrons to microorganisms and thus contribute to the energy flow of microbial communities, and ultimately, to the global cycling of halogens. Dehalogenation reactions occur in aerobic, facultative, and anaerobic organisms spanning three phylogenetic kingdoms (Bacteria, Archaea, and Eukarya), and are an integral part of cometabolic or energy-yielding metabolic processes (for comprehensive reviews, see 22, 25, 71). Many types of microorganisms with different metabolic strategies exist over this broad redox spectrum, helping to define the ecology of a

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particular environment, from aerated, oxidized to highly reduced, anaerobic strata. Figure 2.2 depicts examples of some of the types of organohalide-degrading microorganisms that are active in different redox zones and their ecological relationship to one another. The dehalogenation capabilities of aerobic and anaerobic microorganisms often overlap, underscoring their complementary interactions in and with their environment. Aerobes such as dehalogenating pseudomonads, mycobacteria and methylotrophs occupy the most oxidized zones, requiring to serve as reactant (oxygenase mediated attack) or as an electron acceptor for metabolism, although not necessarily for dehalogenation of halogenated substrates. Oxygenolytic, hydrolytic and reductive dehalogenation mechanisms are commonly found in aerobic bacteria. Facultative, denitrifying microorganisms that dehalogenate, such as Pseudomonas stutzeri and Thauera chlorobenzoica, may compete or overlap at the periphery of aerobic zones if nitrate is present. Dehalorespirers, strict anaerobes that gain energy from electron transfer to organohalides, such as the recently isolated and characterized Dehalococcoides ethenogenes and Dehalobacter restrictus, generally occupy a distinct niche in the middle of the redox range. As depicted in the figure, the dehalorespiring microorganisms are flanked by iron- and sulfate-reducing bacteria, at the positive and negative ends of their redox range, respectively. At the far end of the redox spectrum, usually at deeper depths, acetogens and methanogens occupy the most reduced anoxic strata, often competing for the same electron donor and electron acceptor The thermodynamics of uptake and utilization, however, favor methanogenesis over acetogenesis, all other environmental factors being equal. Note that fermenters, which produce both of these substrates during fermentation, are usually present in close proximity. The hydrogen produced by this group of microorganisms also serves as an electron source for sulfate-reducers and other dehalogenating microorganisms that operate higher on the redox scale. In effect, or hydrogen-donating substrates, serve as the conduit for energy flow through the microbial community (see Section 4.2).

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Through syntrophic associations and/or cross-feeding phenomena, hydrogen transfer spans the entire redox environment. The prevailing electron accepting processes that participate in dehalogenation (Figure 2.2) are determined not only by the availability of electron accepting species, but also by the concentration of (or other hydrogen donor) in the environment (see Chapter 4). Table 2.2 and Figure 2.3 illustrate how the available hydrogen pool and threshold concentrations help to determine prevailing species and energy flow. Recent studies by Löffler et al. (49) provide clear evidence for the role of threshold concentrations in determination of an electron accepting process, and have shown that thermodynamic constraints, i.e., uptake and energy yield, favor utilization by halorespirers in the presence of haloorganic substrates (contaminants). As more information is gained about dehalorespirers, a unique and thermodynamically efficient group of microorganisms, a clearer understanding of their potential for the bioremediation of organohalide-contaminated environments will ensue.

4. ENVIRONMENTAL FACTORS AFFECTING MICROBIAL ACTIVITY 4.1. Effect of Substrate Biodegradability, Bioavailability, and Toxicity on Dehalogenation The intrinsic characteristics of an organohalide substrate, as determined by its biodegradability, bioavailabililty, and toxicity to degrading microorganisms, will ultimately determine its biological fate in the environment (see also Chapters 11 and 12). Halogen substituents generally reduce the biodegradability of an organic substrate in one or both of two ways, by changing the physical or chemical attributes of the substrate. For

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example, biodegradability of a compound may be reduced by the large molecular size of its one or more halogen substituents that can sterically hinder enzymatic attack. Halogen substituents may also impact chemical stability, i.e., reactivity, of a compound. Within the periodic class of halogen substituents (F, Cl, Br, or I), halogen-carbon bond strengths and bond dissociation energies decrease with increasing molecular weight (31, 93), so that fluorinated compounds tend to be less chemically labile (more stable) than iodinated ones. For aromatic compounds, the electron-withdrawing characteristics of a halogen substituent help to stabilize the aromatic ring, making it less susceptible to attack. Multiple halogen substitution, however, may cause ring destabilization by decreasing electron density (82), as do carboxy, hydroxy, and cyano groups (45). In addition to affecting the intrinsic biodegradability of a compound, the extent of halogenation may also affect bioavailabililty, and indirectly impact its biodegradability. Within an analagous set of substates, a greater degree of halogenation decreases the water solubility and increases sorption potential (e.g., partition coefficient, With some exception, most hydrophobic substrates are taken up and utilized by cells in the aqueous phase (discussed in 9, 60), therefore substrate availability to the degrading microorganism is reduced with increasing halogenation. In some cases where the substrate is toxic to cells, e.g., trichloroethene (TCE) and polychlorinated phenols, this reduced bioavailability might prove advantageous to the health and overall biological activity in a given environment. Many halogenated compounds exhibit some toxicity, although this can vary considerably between even similar substrates and isomers. Toxicity is manifest in a variety of ways, including solvation of cell structures, e.g., dissolution of the cell membrane, metabolic (enzyme) inhibition, or decoupling of electron transport. Bioavailability is also affected by partitioning between different environmental compartments, e.g., gas, liquid and solid matrices (7, 89). For volatile substrates, the vapor pressure of a compound, or Henry’s constant, H, will impact the concentration and availability of a substrate to the degrading microorganisms. In general, vapor pressure decreases with increasing degree of halogenation, so that within a class of compounds, less-halogenated analogues will exhibit increased volatility: chloromethane methylene chloride chloroform carbon tetrachloride or as in another series, vinyl chloride > dichloroethene > trichloroethene > tetrachloroethene. Partitioning onto solid surfaces will also depend greatly on the nature of the solid phase, i.e., hydrophilic (mineral) or hydrophobic (humus and natural organic) material. In general, increased halogenation of a substrate increases hydrophobicity and decreases water solubility, so that substrates partition more readily onto hydrophobic, e.g., organic, surfaces. Some organohalides degrade rapidly at low concentrations, but are persistent at higher concentrations, often due to bacteriostatic or toxic effects of the substrate, e.g., 3-chloro- or 3-fluorobenzoate (82). A similar phenomenon also has been reported for chlorophenols and related compounds (6, 51, 62). Conversely, some organohalides exhibit lower threshold concentrations for degradation, e.g., 4-amino-3,5dichlorobenzoate (82), suggesting that bioavailability may be the bottleneck for biodegradation. Under such circumstances, the amount of substrate available to the degrading microorganisms is below the concentration limit for uptake and bioavailability, perhaps due to poor water solubility or unfavorable partitioning.

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Threshold concentrations for biodegradation have been observed for a variety of halogenated compounds (3), including apparent thresholds for the anaerobic dechlorination of PCBs (1, 26, 69, 72). For this group of compounds, the evidence suggests that the enhanced dechlorination rates observed at higher PCB concentrations are due to increased bioavailability. Biodegradation of harmful organohalides provides a means for detoxification in the environment (for examples, see Chapters 13 to 17). Unless biodegradation is complete, however, some partially biodegraded (dehalogenated) metabolites may be even more harmful than the parent compound, both with regard to intrinsic toxicity as well as mobility in the environment. For example, vinyl chloride (VC) is a partially dehalogenated analog of tetrachloroethene (PCE) and TCE dechlorination often encountered in the field as a result of natural attenuation or engineered remediation processes and also in laboratory microcosms (18, 24, 58, 77, 78, 84; see Chapters 14 and 15). VC is a volatile, toxic metabolic product and a known carcinogen. Alternatively, in aerobic environments TCE may undergo partial degradation to chloral or dichloroacetate, both also toxic products through the activity of (methane)monooxygenases (93). Microbial food chains, comprised of two or more types of microorganisms, are often required to biodegrade (and detoxify) recalcitrant organohalides. By combining the metabolic activities of several microorganisms, difficult substrates may be completely mineralized. Complete biodegradation of a highly halogenated substrate may require dehalogenation by one species, followed by further degradation of the carbon “backbone” by a second species. Whereas the first metabolic step in this example may produce energy for the dehalogenating microorganism, it does not provide carbon for growth. This illustrates the role of secondary substrates which are often required, either as carbon source and/or hydrogen source for the biodegradation (dehalogenation) of highly halogenated organic compounds, e.g., PCE, TCE, PCBs. The role of an auxiliary substrate in this case should not be confused with cometabolism, or a fortuitous cooxidation of substrate where no benefit is derived for the mediating organism. Cosubstrates are often required for anaerobic dehalogenation processes. Their role may be multi-fold, to provide reducing power, co-factors, supplementary carbon and energy to the complex microbial communities often associated with the biodegradation of highly chlorinated compounds under anaerobic conditions.

4.2. Effect of Electron Acceptors on Dehalogenation and Electron Flow The environmental factors that often define microbial community structure and activity are generally not discrete entities, but are dynamic, both with respect to time and locale. For example, in a soil representative of a multiphasic system (gas, liquid, and solid phases), the redox environment is best represented as a continuum, ranging from most oxidized at the surface to highly reduced conditions. In an oxic environment, aerobic microorganisms predominate, utilizing as a reactant and an electron acceptor for respiratory processes. With depth, a decreasing redox potential ensues as conditions become more reduced, due in large part to microbial activity and a hindered physical exchange of atmospheric gases. becomes depleted and concentrations increase. At the lower depths, alternative electron acceptors, especially

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inorganic species such as and are present as dissolved species or as part of the surrounding mineral matrix, and serve as electron acceptors in support of respiratory processes mediated by facultative and strict anaerobic microorganisms (29). Other, relatively minor mineral species also support microbiological activity in soils by serving as alternate electron acceptors in soils and sediments. For example, selenate and arsenate have been reported to serve as terminal electron acceptors in support of biodegradation activity in soil and sediment enrichment cultures (67, 81), although not specifically attributed to dehalogenation processes. In addition, organic species including organohalides, may serve as electron acceptors during fermentative, as well as respiratory processes (49; see Chapters 3,4 and 5). The use of organohalides as electron acceptors, (as well as electron donors), has important implications for the fate of organohalides in the environment and the cleanup of contaminated sites, in particular in less accessible areas which may be restricted to natural attenuation processes. The presence and absence of respiratory electron acceptors will have a direct impact on the activity of the microbial community, including the biodegradation of organohalides. Dehalogenating microorganisms requiring molecular oxygen for initial attack (via oxygenases) or for respiration will naturally be inactive in anoxic environments. In the absence of oxygen, organic compounds including organohalides may be degraded by mixed microbial communities using alternate electron acceptors. This includes the use of organohalide as the electron acceptor for respiration, especially in environments rich in halogenated substrates. A number of halogenated organic compounds can be degraded under different electron accepting conditions and their complete oxidation to can be coupled to processes such as denitrification, Fe(III)reduction, sulfate reduction and methanogenesis (10, 12-14, 27, 28, 32, 34-36, 41,42, 61,65,48,75). Several studies have shown that nitrate and sulfate, in particular, impede the enrichment of dehalogenating populations, or that interspecific competition for electron donors inhibits anaerobic reductive dechlorination (5,30,33,40,44,46,52,63,75,82, 86,87). Sulfate-reducing bacteria may out-compete dehalogenating microorganisms for hydrogen, but these interactions appear to be more complex (86). For example, in studies with Desulfomonile tiedjei DCB-1, thiosulfate and sulfite competed with 3chlorobenzoate for reducing equivalents and their reduction was favored over dechlorination (16, 17, 86). Recently, some anaerobic dehalogenating bacteria that are not inhibited by sulfate have been isolated (11, 83). For many soil-type environments, a continuous redox spectrum can be envisioned, where stratification of substrate (electron donor) and electron acceptors occurs in relation to the redox gradient and availability of electron acceptor. Competition for hydrogen (or other electron donor) is key in determining the succession of microorganisms and redox processes. A variety of compounds, including organohalides and soil minerals, may serve either as carbon source and/or electron donor for energy flow through the community, or as external electron acceptors for respiratory processes. This is at best a simplistic representation of actual field conditions, since a soil or sediment environment seldom is a uniformly mixed system, and is likely to have pockets of heterogeneity. This heterogeneity is reflected in redox environment as well, with low redox anoxic centers (microniches) existing within areas of higher redox surroundings.

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The graduated continuum of changing redox and substrates is a dynamic one, as defined by partition equilibria, biological activity (i.e., biodegradation), and physical mixing (e.g., via air or water flow, the burrowing activity of invertebrates, or engineered intervention) within the environmental matrix (89). Of particular importance, and arguably the underlying basis of the relationships that define the microbial ecology of an environment, is the flow of electrons through the system. Energy flow generally follows a decreasing trend in redox, from most oxidized to least oxidized systems as depicted in Figure 2.3. This parallels a decreasing trend in energy yield as the electron accepting process becomes more reduced. With ensuing changes in the redox environment, different microbial populations predominate, so that electron flow is not necessarily intraspecific, but often passes from one microbial population to another, from aerobic to facultative to anaerobic species. Electron transfer generally occurs through changes in the oxidation state of metabolic substrates and products, but in many anoxic environments, is the prevailing electron donor and vehicle for interspecies electron transfer between respiratory and fermentative species in the anaerobic food chain (Figure 2.3; 50). In environments with multiple available electron acceptors, the concentration of can determine which electron accepting process will predominate (15, 49, 50). The dynamics of production and consumption, i.e., flow in a microbial food web, have been clearly detailed in an anaerobic 3-chlorobenzoate degrading methanogenic consortium first described by Shelton and Tiedje (79). This stable consortium, consisting of a dechlorinating bacterium (Desulfomonile tiedjei DCB-1), a hydrogenproducing benzoate degrader, and a methanogen illustrates the synergistic interactions that provide thermodynamically favorable conditions for complete degradation of the 3-chlorobenzoate substrate (19,20,79). As described, benzoate degradation to acetate, and provided the reducing power for reductive dechlorination. One third of the benzoate-derived hydrogen was recycled through reductive dechlorination with the remainder being consumed by the methanogen. This type of cross-feeding of energyyielding substrates (as well as additional growth factors such as vitamins) appears to be a common phenomenon in anaerobic dehalogenating communities.

4.3. Matrix Mineralogy Matrix mineralogy implies a solid structure, therefore the discussion here focuses primarily on soils and sediments. It should be noted, however, that minerals in the aqueous phase, as in marine and freshwater environments, can significantly impact microbial activity (at water-rock interfaces and soluble phases) and overall ecology. For detailed information on soil mineralogy and its associated geomicrobial processes, the reader is referred to Ehrlich (21). The soil matrix, in particular its mineral component, may affect the microbial ecology of dehalogenation in a variety of ways: 1) through physical microbe-mineral interactions, i.e., mobility and sorption; 2) through substratematrix interactions, i.e., sorption that can alter substrate bioavailability; and 3) by providing alternate electron acceptors for microbial activity in environments, i.e., the subsurface. Most soils and sediments contain varying combinations of both organic and mineral components. Within the inorganic mineral matrix of a soil, a wide variety of minerals

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provide structure and ionic definition, e.g., carbonates, silicates, and oxides. In general, these structural components have a net negative charge and help to determine the cation exchange capacity of a soil. The cation exchange capacity, together with the porosity and fine structure of a soil, will affect how well microorganisms (which generally carry a net negative charge) can move through the matrix. Metallic ions, such as and further define the mineral nature of a soil, as well as its ability to sorb and/or intercalate substrates and microorganisms, by affecting its net charge, as well as its mineral (clay) structure. In this very complex environment, both carbonate and other ionic mineral components may interact with soil microorganisms and become altered, physically and chemically, through microbial activity. Mineral species in the soil matrix, in particular Fe(III) and Mn(IV), can serve as electron acceptors during the biodegradation of organohalides. In some cases, the reduced species, for example, may further the process and directly interact with the organohalide substrates by catalyzing abiotic reductive dehalogenation reactions (see below). During the biodegradation of organohalides, the production of organic and/or inorganic acids (e.g., HCl) may solubilize minerals, mobilizing and/or making them more bioavailable. It should be noted that most soils exhibit a large buffering capacity, due primarily to the mineral (i.e., carbonate and clay matrix), as well as the organic fraction (humus and natural organic material), providing a natural buffer to acid production. Other environmentally important electron acceptors include carbonate, sulfate, and nitrate, as noted earlier. These anions may be naturally present as part of the soil mineralogy and microbial activity, but they may also occur as a result of man-made intervention, e.g., fertilization. Carbonate is one of the most abundant mineral species on earth, serving as a major carbon sink in both terrestrial and marine environments. It also serves as an electron acceptor for methanogens and acetogens in highly reduced environments. Its speciation in the environment, i.e., equilibration between carbonate and bicarbonate is dependent on a number of environmental factors, including pH, temperature, and pressure. These environmental factors often determine the bioavailability of the electron acceptor to carbonate-reducing, i.e., methanogenic and acetogenic, bacteria. Sulfate is produced biologically in aerobic zones from elemental sulfur and S-containing minerals, by the action of sulfur-oxidizing bacteria. It is also one of the predominant anions found in marine habitats, approximately 28 mM in seawater. Nitrate can be formed from ammonia-oxidizing bacteria or by urea hydrolysis, but one of its major sources in the environment comes from surface run-off from agricultural practices such as fertilization. Whereas carbonate exhibits a very complex behavior in soils, the latter two anions are generally quite mobile in most soils and sediment. By their ability to serve as electron acceptors during anaerobic respiration, all three play an important role in defining the microbial ecology of the anaerobic soil environment, including the support of microbial mediators of organohalide degradation reactions. Beyond supporting microbial activity by providing alternate electron acceptors and increasing buffering capacity, the mineral component of many soils, in particular iron, can contribute biologically, as well as mediate the abiotic dehalogenation of organohalide contaminants (90; see also Chapter 9). For example, oxidized iron mineral species, in the presence of an electron donor, can serve as an electron acceptor for iron-

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reducing bacteria. During this process, the oxidized iron is reduced and becomes solubilized. Reduced iron species can catalyze an abiotic reductive dehalogenation of halogenated organic substrates (23, 57). An extention of this concept can be applied to the field for the bioremediation of sites contaminated with organohalides, such as PCE and TCE. In soils naturally high in iron an organic substrate (electron donor) is added to the contaminated area, and iron-reducing bacteria (naturally-occurring or augmented) reduce the to thereby producing a soluble catalyst for dehalogenation of the target contaminant. The catalytic properties of iron, in particular zero-valent iron, also have been applied to engineered remediation strategies for the in-situ treatment of soils and groundwater contaminated with a variety of halogenated solvents, including PCE, TCE, and chlorinated methanes (59, 73, 74, 76, 85). In the cited studies, permeable reactive barriers of zero-valent iron were placed downstream or within a contaminant plume to intercept and treat contaminants as they pass through with groundwater flow. Other reduced metals, copper, zinc or aluminum, have been successfully tested in the laboratory, but have not yet been applied to the field, in large part due to economic reasons. By several mechanisms, all of which are not yet fully understood, the reactive surfaces of zero-valent iron catalyze reductive dehalogenation of the target contaminants. The potential role of microorganisms in the engineered permeable reactive barriers environment also is not clear, but evidence for qualitiative changes in bacterial populations upstream and downstream of the iron barrier has been reported (90). In their pilot-scale field study at an industrial site with chlorinated solvent contaminantion, the researchers observed greater numbers of sulfate-reducing bacteria within the barrier, and a shift towards obligate anaerobes downstream, in response to generated during abiotic dehalogenation by zero-valent iron. The influence of changing environmental parameters, in this case, redox and electron donor, again underscores the important role that environment plays in defining the microbial ecology of a site and its dehalogenation processes.

4.4. Other Environmental Determinants A number of other environmental factors also play a role in determining the microbial ecology of a site, and for purposes of discussion here, its dehalogenation potential. Environmental effects may be physicochemical or biological, or both. Temperature can modify the phase separation or compartmentalization of a substrate, and therefore also its bioavailability, i.e., higher temperatures may favor a gaseous state, while low temperatures result in a liquid or solid phase, depending on the vapor pressure, and/or melting point of a particular substrate. Temperature can also determine rates of microbial activity, both with respect to rate and type. A general rule of thumb for temperature influence on the rate of reaction within a given biological range is represented by the quotient, or a doubling of reaction rate for each 10 °C rise in temperature. Temperature may also exert a selective effect on anaerobic dehalogenation activity, whereby certain temperature ranges elicit specific dechlorination patterns, based on the distinct activity ranges of different members in acclimated, mixed cultures. This has been observed for reductive dechlorination of PCBs in anaerobic sediments (97-99).

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The optimum temperature range for most microorganisms falls within the mesophilic range, reflecting the environment where they are found. However, isolates from extreme environments, such as thermal pools or polar regions, often exhibit optimum ranges outside of the norm. For example, diverse psychrotolerant, pentachlorophenol-degrading bacteria have been found in a cold chlorophenol-contaminated aquifer (54,55). Aerboic degradation of TCE and PCBs has also been observed at low temperatures (64,66,96), and psychrotolerant PCB-degrading bacteria have been isolated (56). Biological temperature constraints have also been a consideration in the development of biological treatment protocols. For example, a fluidized bed reactor for on-site treatment of groundwater contaminated with chlorophenols was successfully acclimated to the low temperatures required for operation (38, 39; see also Chapter 16). As with temperature, the hydrogen reactivity, i.e., pH, of a given environment also can have profound effects on both biological and physicochemical components. Extremes in pH can directly affect cellular physiology, and may select for specially adapted microorganisms that function well in either highly acidic, or in very alkaline environments. pH may also directly affect the speciation of substrates and minerals, which in turn impacts microbial activity. Control of pH is especially critical for environments where halogenated substrates predominate. In the absence of adequate buffering capacity, acid production, e.g., HCl, from the biodegradation of halogenated substrates may significantly change prevailing pH conditions. Together with pH, redox conditions in large part determine mineral speciation and ambient concentrations, which in turn help to define community and metabolic processes that predominate in a given environment. A wide range in environmental conditions support biogeochemically relevent microbial communities and activity, e.g., dehalogenation (21). Subsequent changes in redox potential (reduction of electron acceptor) and pH (acid production from dechlorination), as a function of microbial activity, can change the type of microbial community that predominates in a given environment. Other environmental factors which do not directly support or define the biodegradation of organohalides may impact overall activity, not as supporters of activity, but rather as inhibitors. High salt concentrations (salinity) generally depress microbial activity, and can therefore inhibit biodegradation of organohalides. Extreme halophilic microorganisms which require high salt concentrations (> 15%) have not yet been implicated in the dehalogenation of organohalides. Other types of inhibitory compounds, such as heavy metals or other potentially toxic materials, can negatively impact overall microbial activity (47,68,94,95). In some cases, the negative impact of environmental extremes can be mitigated by adjusting other environmental factors, and thereby minimize impact on exposed microbial populations. For example, a change in redox can alter heavy metals speciation to a less toxic form, or an increase in prevailing pH may precipitate undesirable metals and reduce their mobility, thereby localizing any detrimental effects. These examples further underscore the many interrelationships between substrate, microorganism, and environment, and how interdependent they are on one another.

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5. SUMMARY In reviewing the physicochemical factors and biological activities that describe the microbial ecology of an environment, in particular how it pertains to the global cycling of halogens, many elements are evident. A unifying principle in these often complex systems can perhaps be found in the energy flow through the system. On a small scale, this can be seen in the microbial communities which support and mediate dehalogenation. Hydrogen transfer, through syntrophic and cometabolic microbial associations, together with its impact on the environmental matrix, e.g., redox and mineral speciation, provides a commonality to the cycling of nutrients in the environment, including halogenated substrates. A better understanding can serve as a basis for exploiting these processes for the better good, in particular for the cleanup of contaminated environments, but also for better management of naturally-occurring processes. REFERENCES 1. Abramowicz DA, Brennan MJ, Van Dort HM & Gallagher EL(1993) Factors influencing the rate of polychlorinated biphenyl dechlorination in Hudson River sediments. Environ. Sci. Technol. 27:1125-1131 2. Adriaens P & Vogel T (1995) Biological treatment of chlorinated organics. In: Young LY & Cerniglia CE (Eds) Microbial Transformation and Degradation of Toxic Organic Chemicals (pp 435-486) Wiley-Liss, New York 3. Alexander M (1985) Biodegradation of organic chemicals. Environ. Sci. Technol. 18:106-112 4. Alexander M (1981) Biodegradation of chemicals of environmental concern. Science 211:132-138 5. Allard A-S, Hynning P-Å, Remberger M & Neilson AH (1992) Role of sulfate concentration in dechlorination of 3,4,5-trichlorocatechol by stable enrichment cultures grown with coumarin and flavanone glycones and aglycones. Appl. Environ. Microbiol. 58:961-968 6. Apajalahti JHA & Salkinoja-Salonen MS (1984) Absorption of pentachlorophenol (PCP) by bark chips and its role in microbial PCP degradation. Microb. Ecol. 10:359-367 7. Arands R, Bossert I, Kosson D, Lederman P, Lyman W, Massry I, Schaefer C & Tekrony M (1999) Mobility and Degradation of Organic Contaminants in Subsurface Environments: Update for Phase I. EPA report 600/2-91/053 8. Bartha R & Atlas R (1998) Microbial Ecology: Fundamentals and Applications. Benjamin Cummings Publ. Co. Menlo Park, CA 9. Bossert I & Compeau G (1995) Petroleum hydrocarbon contamination in soil. In: Young LY & Cerniglia CE (Eds) Microbial

Transformation and Degradation of Toxic Organic Chemicals (pp 77-125) Wiley-Liss, New York 10. Boyle AW, Knight VK, Häggblom MM & Young LY (1999) Transformation of 2,4dichlorophenoxyacetic acid in four different marine and estuarine sediments: effects of sulfate, hydrogen and acetate on dehalogenation and side-chain cleavage. FEMS Microbiol. Ecol. 29:105-113 11. Boyle AW, Phelps CD & Young LY (1999) Isolation from estuarine sediments of a Desulfovibrio strain which can grow on lactate coupled to the reductive dehalogenation of 2,4,6-tribromophenol. Appl. Environ. Microbiol. 65:1133-1140 12. Bradley PM, Chapelle FH (1996) Anaerobic mineralization of vinyl chloride in Fe(III)reducing aquifer sediments. Environ. Sci. Technol. 30:2084-2086 13. Bradley PM & Chapelle F (1998) Microbial Mineralization of VC and DCE under different terminal electron accepting conditions. Anaerobe 4:81-87 14. Bradley PM & Chapelle FH (2000) Aerobic microbial mineralization of dichloroethene as sole carbon substrate. Environ. Sci. Technol. 34:221-223 15. Chapelle FH, Haack SK, Adriaens P, Henry MA & Bradley PM (1996) Comparison of and measurements for delineating redox processes in a contaminated aquifer. Environ. Sci. Technol. 30:3565-3569 16. DeWeerd KA, Concannon F & Suflita JM (1991) Relationship between hydrogen consumption, dehalogenation, and the reduction

MICROBIAL ECOLOGY OF DEHALOGENATION of sulfur oxyanions by Desulfomonile tiedjei. Appl. Environ. Microbiol. 57:1929-1934 17. DeWeerd KA & Suflita JM (1990) Anaerobic aryl reductive dehalogenation of halobenzoates by cell extracts of "Desulfomonile tiedjei". Appl. Environ, Microbiol. 56:2999-3005 18. DiStefano TD, Gossett JM & Zinder SH (1992) Hydrogen as an electron donor for dechlorination of tetrachloroethene by an anaerobic mixed culture. Appl. Environ. Microbiol. 58:3622-3629 19. Dolfing J & Tiedje JM (1986) Hydrogen cycling in a three-tiered food web growing on the methanogenic conversion of 3-chlorobenzoate. FEMS Microbiol. Ecol. 38:293-298 20. Dolfing J & Tiedje JM (1991) Kinetics of two complementary hydrogen sink reactions in a defined 3-chlorobenzoate degrading methanogenic co-culture. FEMS Microbiol. Ecol. 86:25-32 21. Ehrlich, HL (1996) Geomicrobiology, Third Edition. Marcel Dekker, Inc., New York 22. El Fantroussi S, Naveau H & Agathos SN (1998) Anaerobic dechlorinating bacteria. Biotechnol. Prog. 14:167-188 23. Erbs M, Hansen HCB & Olsen CE (1999) Reductive dechlorination of carbon tetrachloride using iron(II) iron(III)hydroxide sulfate (green rust). Environ. Sci. Technol. 33:307-311 24. Fennell DE, Gossett JM & Zinder SH (1997) Comparison of butyric acid, ethanol, lactic acid, and propionic acid as hydrogen donors for the reductive dechlorination of tetrachloroethene. Environ. Sci. Technol. 31:918-926 25. Fetzner S (1998) Bacterial dehalogenation. Appl. Microbiol. Biotechnol. 50:633-657 26. Fish KM (1996) Influence of Aroclor 1242 concentration on polychlorinated biphenyl biotransformations in Hudson River test tube microcosms. Appl. Environ. Microbiol. 62:3014-3016 27. Genthner BRS, Price WA II & Pritehard PH (1989) Anaerobic degradation of chloroaromatic compounds in aquatic sediments under a variety of enrichment conditions. Appl. Environ. Microbiol. 55:1466-1471 28. Genthner BRS, Price WA II & Pritchard PH (1989) Characterization of anaerobic dechlorinating consortia derived from aquatic sediments. Appl. Environ. Microbiol. 55:14721476 29. Ghiorse WC, Wilson JT (1988) Microbial ecology of the terrestrial subsurface. Adv. Appl. Microbiol. 33:107-172 30. Gibson SA & Suflita JM (1986) Extrapolation of biodegradation results to groundwater aquifers: reductive dehalogenation of aromatic

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50 43. Kohring G-W, Rogers JE & Wiegel J (1989) Anaerobic biodegradation of 2,4-dichlorophenol in freshwater lake sediments at different temperatures. Appl. Environ. Microbiol. 55:348353 44. Kohring G-W, Zhang X & Wiegel J (1989) Anaerobic dechlorination of 2,4-dichlorophenol in freshwater sediments in the presence of sulfate. Appl. Environ. Microbiol. 55:2735-2737 45. Kuhn EP & Suflita JM (1989) Dehalogenation of pesticides by anaerobic microorganisms in soils and groundwater A review. In: Sawhney BL, Brown K (Eds) Reactions and Movements of Organic Chemicals in Soils (pp 111 -180) Soil Sci. Soc. of America and American Soc. of Agronomy, Madison, WI 46. Kuhn EP, Townsend GT & Suflita JM (1990) Effect ofsulfate and organic carbon supplements on reductive dehalogenation of chloroanilines in anaerobic aquifer slurries. Appl. Environ. Microbiol. 56:2630-2637 47. Kuo C-W & Genthner BRS (1996) Effect of added heavy metal ions on biotransformation and biodegradation of 2-chlorophenol and 3chlorobenzoate in anaerobic bacterial consortia. Appl. Environ. Microbiol. 62:2317-2323 48. Liu S-M, Kuo C-E & Hsu T-B (1996) Reductive dechlorination of chlorophenols and pentachlorophenol in anoxic estuarine sediments. Chemosphere 32:1287-1300 49. Löffler FE, Tiedje JM & Sanford RA (1999) Fraction of electrons consumed in electron acceptor reduction and hydrogen thresholds as indicators of halorespiratory physiology. Appl. Environ. Microbiol. 65:4049-4056 50. Lovley DR, Chapelle FH & Woodward JC (1994) Use of dissolved concentrations to determine distribution of microbially catalyzed reactions in anoxic groundwater. Environ. Sci. Technol. 28:1205-1210 51. Madsen T & Aamand J (1992) Anaerobic transformation and toxicity of trichlorophenols in a stable enrichment culture. Appl Environ Microbiol 58:557-561 52. Madsen T & Aamand J (1991) Effects of sulfuroxy anions on degradation of pentachlorophenol by a methanogenic enrichment culture. Appl. Environ. Microbiol. 57:2453-2458 53. Männisto MK & Puhakka JA (2002) Psychrotolerant and microaerophilic bacteria in boreal groundwater. FEMS Microbiol. Ecol. 41:9-16 54. Männisto MK, Tiirola MA & Puhakka JA (2001) Degradation of 2,3,4,6-tetrachlorophenol at low temperature and low dioxygen concentrations by phylogenetically different

BOSSERT ET AL. groundwater and bioreactor bacteria. Biodegradation 12:291-301 55. Männistö MK, Tiirola MA, Salkinoja-Salonen MS, Kulomaa MS & Puhakka JA (1999) Diversity of chlorophenol-degrading bacteria isolated from contaminated boreal groundwater. Arch. Microbiol. 171:189-197 56. Master ER, Mohn WW (1998) Psychrotolerant bacteria isolated from arctic soil that degrade polychlorinated biphenyls at low temperatures. Appl. Environ. Microbiol. 64:4823-4829 57. Matheson LJ & Tratnyek PG (1994) Reductive dehalogenation of chlorinated methanes by iron metal. Environ. Sci. Technol. 28:2045-2053 58. McCarty PL (1993) In situ bioremediation of chlorinated solvents. Curr. Opin. Biotechnol. 4:323-330 59. McNab WW & Ruiz R (2000) In-situ destruction of chlorinated hydrocarbons in groundwater using catalytic reductive dehalogenation in a reactive well: testing and operational experiences. Environ. Sci. Technol. 34:149-153 60. Miller R (1994) Surfactant-enhanced bioavailability of slightly soluble organic compounds. In: Skipper H (Ed) Bioremediation – Science and Applications. Soil Science Society of American Publications, Madison, WI 61. Milligan PW & Häggblom MM (2000) Anaerobic biodegradation of halogenated pesticides: Influence of alternate electron acceptors. In: Bollag J-M& Stotzky G (Eds) Soil Biochemistry, (pp 1-34) Marcel Dekker, New York 62. Mohn WW & Kennedy KJ (1992) Reductive dehalogenation of chlorophenols by Desulfomonile tiedjei DCB-1. Appl. Environ. Microbiol. 58:1367-1370 63. Mohn WW & Tiedje JM (1992) Microbial reductive dehalogenation. Microbiol. Rev. 56:482-507 64. Mohn WW, Westerberg K, Cullen WR, Reimer KJ (1997) Aerobic biodegradation of biphenyl and polychlorinated biphenyls by arctic soil microorganisms. Appl. Environ. Microbiol. 63:3378-3384 65. Monserrate E & Häggblom MM (1997) Dehalogenation and biodegradation of phenols and benzoic acids under iron-reducing, sulfidogenic, and methanogenic conditions. Appl. Environ. Microbiol. 63:3911-3915 66. Moran BN & Hickey WJ (1997) Trichloroethylene biodegradation by mesophilic and psychrophilic ammonia oxidizers and methanotrophs in groundwater microcosms. Appl. Environ. Microbiol. 63:3866-3871 67. Oremland RS, Newman DK, Kail BW & Stolz

MICROBIAL ECOLOGY OF DEHALOGENATION JF (2001) Bacterial respiration of arsenate and its significance in the environment. In: Frankenberger WT Jr (Ed) Environmental Chemistry of Arsenic (pp 273-295) Marcel Dekker, New York 68. Pardue JH, Kongara S & Jones WJ (1996) Effect of cadmium on reductive dechlorination of trichloroaniline. Environ. Toxicol. Chem. 15:1083-1088 69. Quensen JF III, Tiedje JM & Boyd SA (1988) Reductive dechlorination of polychlorinated biphenyls by anaerobic microorganisms from sediments. Science 242:752-754 70. Ramaswami A & Luthy R (1997) Measuring and modeling physicochemical limitations to bioavailability and biodegradation. In: Hurst C, Knudsen G, McInerney M, Stetzenbach L & Walter M (Eds) Manual of Environmental Microbiology (pp 721-729) ASM Press, Washington, DC 71. Reineke W (2001) Aerobic and anaerobic biodegradation potentials of microorganisms. In: Beek B (Ed) Handbook of Environmental Chemistry, Vol 2, Part K. Biodegradation and Persistence (pp 1-161). Springer-Verlag, Berlin 72. Rhee G-Y, Bush B, Bethoney CM, DeNucci A, Oh H-M & Sokol RC (1993) Reductive dechlorination of Aroclor 1242 in anaerobic sediments: pattern, rate and concentration dependence. Environ. Toxicol. Chem. 12:10251032 73. Richardson JP & Nicklow JW (2002) In situ permeable reactive barriers for groundwater contamination. Soil Sediment Contam. 11:241268 74. Ritter K, Odziemkowski MS & Gillham RW (2002) An in situ study of the role of surface films on granular iron in the permeable iron wall technology. J. Contam. Hydrol. 55:97-111 75. Sanford RA & Tiedje JM (1997) Chlorophenol dechlorination and subsequent degradation in denitrifying microcosms fed low concentrations of nitrate. Biodegradation 7:425-434 76. Scherer MM, Balko BA, Gallagher DA & Tratnyek PG (1998) Correlation analysis of rate constants for dechlorination by zero-valent iron. Environ. Sci. Technol. 32:3026-3033 77. Semprini L, Hopkins GD, McCarty PL & Roberts PV (1992) In-situ transformation of carbon tetrachloride and other halogenated compounds resulting from biostimulation under anoxic conditions. Environ. Sci. Technol. 26:2454-2461 78. Semprini L, Kitanidis PK, Kampbell DH & Wilson JT (1995) Anaerobic transformation of chlorinated aliphatic hydrocarbons in a sand aquifer based on spatial chemical distributions.

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79. Shelton DR & Tiedje JM (1984) Isolation and partial characterization of bacteria in an anaerobic consortium that mineralizes 3chlorobenzoic acid. Appl. Environ. Microbiol. 48:840-848 80. Smith RL (1997) Subsurface and landfills. In: Hurst C, Knudsen G, McInerney M, Stetzenbach L & Walter M (Eds) Manual of Environmental Microbiology (pp 577-585) ASM Press, Washington, DC 81. Stolz JF & Oremland RS (1999) Bacterial respiration of arsenic and selenium. FEMS Microbiol. Rev. 23:615-627 82. Suflita JM & Townsend GT (1995) The microbial ecology and physiology of aryl dehalogenation reactions and implications for biorermediation In: Young LY & Cerniglia CE (Eds) Microbial Transformation and Degradation of Toxic Organic Chemicals (pp 243-268) Wiley-Liss, New York 83. Sun B, Cole JR & Tiedje JM (2001) Desulfomonile limimaris sp. nov., an anaerobic dehalogenating bacterium from marine sediments. Int. J. Syst. Evol. Microbiol. 51:365-371 84. Tandol V, DiStefano TD, Bowser PA, Gossett JM & Zinder SH (1994) Reductive dehalogenation of chlorinated ethenes and halogenated ethanes by a high-rate anaerobic enrichment culture. Environ. Sci. Technol. 28: 973-979 85. Tratnyek, PG, Scherer MM, Deng B & Hu S (2001) Effects of natural organic matter, anthropogenic surfactants, and model quinones on the reduction of contaminants by zero-valent iron. Water Research 25:4435-4443 86. Townsend GT & Suflita JM (1997) Influence of sulfur oxyanions on reductive dehalogenation activities in Desulfomonile tiedjei. Appl. Environ. Microbiol. 63:3594-3599 87. Townsend GT, Ramanand K & Suflita JM (1997) Reductive dehalogenation and mineralization of 3-chlorobenzoate in the presence of sulfate by microorganisms from a methanogenic aquifer. Appl. Environ. Microbiol. 63:2785-2791 88. Valo R, Apajalahti J & Salkinoja-Salonen M (1985) Studies on the physiology of microbial degradation of pentachlorophenol. Appl. Microbiol. Biotechnol. 21:313-319 89. Venkatramen S, Schuring JR, Boland TM, Bossert ID & Kosson D (1998) Application of pneumatic fracturing to enhance in-situ bioremediation. J. Soil Contam. 7:143-162 90. Vogan JL, Focht RM, Clark DK & Graham SL (1999) Performance evaluation of a permeable

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Chapter 3 DIVERSITY OF DECHLORINATING BACTERIA FRANK E. LÖFFLER 1, JAMES R. COLE 2, KIRSTI M. RITALAHTI 1, AND JAMES M. TIEDJE 2 1 School of Civil and Environmental Engineering, Georgia Institute of Technology, Atlanta, GA, USA 2 Center for Microbial Ecology, Michigan State University, East Lansing, Ml, USA

1. CHLORINATED COMPOUNDS IN THE ENVIRONMENT Chlorinated compounds are ubiquitous environmental pollutants due to their extensive use in industry, agriculture, and private households. Despite efforts to replace chlorinated chemicals by compounds that are of lesser environmental concern, large amounts of chlorinated chemicals are still produced for a variety of applications (44). In the early 1970s, the potential danger that chloroorganic chemicals pose to the environment and human health was recognized and consequently, production and handling operations were optimized and restricted to minimize human exposure, as well as to improve environmental quality. At the time, it was believed that the chemically synthesized chloroorganic compounds had no naturally occurring counterparts, and were therefore termed “xenobiotics” (from the greek, xenos = foreign, and bios = life). Research over the last two decades, however, has demonstrated that most, if not all, chloroorganic pollutants are also produced naturally, and are not “foreign to life”. Hence, the term xenobiotic is misleading, and should only be used to indicate that anthropogenic (from the greek, anthropos=people, and genesis = creation) activity has changed the concentration of a chloroorganic compound in certain environments. Significant amounts of chloroorganic compounds are of biogenic and geogenic origin (7, 28, 59-62, 76, 77, 146). Marine organisms, in particular seaweeds, sponges, ascidians, soft corals, and algae, commonly produce a variety of secondary metabolites, many of which are halogenated compounds (62, 72, 73). The function of these compounds for the producing organisms is largely unknown, however, in some cases haloorganic compounds may play a role as feeding deterrents (72,73). Abrahamsson et al. (1) demonstrated the biogenic formation of tetrachloroethene (PCE) and trichloroethene (TCE) by different species of temperate, subtropical, and tropical marine micro- and macro- algae. Other algal species have been shown to produce halogenated propanes (63), and cyanobacterial mats were implied to be a major source of a variety of halogenated aliphatic compounds (161). The natural global production of Dehalogenation: Microbial Processes and Environmental Applications, pages 53-87 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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chloromethane in marine systems has been estimated at 5 million tons per year, exceeding the anthropogenic production 200-fold (70). The natural production of halogenated compounds has also been observed in terrestrial habitats. A recent report demonstrated the natural formation of chloroform and chlorodibromomethane in soil (77). Enormous quantities of an array of chlorinated phenolic compounds are produced naturally, several of which are on the United States Environmental Protection Agency (USEPA) priority pollutants list. Swedish peat bogs alone are estimated to contain 400,000 tons of chlorophenols produced by biological processes. De Jong et al. (28) demonstrated that white rot fungi produce high concentrations of chlorinated anisyl alcohols and aldehydes (up to 75 mg per kg of wood), which are precursors for the biogenic formation of chlorophenols. Recent research has demonstrated that even compounds which were viewed as typical anthropogenic chemicals only a few years ago can be produced naturally, e.g., through the pyrolysis of organic material in the presence of inorganic chloride. An estimated 60 kg of polychlorinated dibenzo-p-dioxins (PCDDs) and dibenzofurans (PCDFs) are formed annually in Canadian forest fires (168). For comparison, the Seveso accident in Italy in 1976, with its well-documented impact on human health and on the environment, released “only” between 0.1 and 20 kg of 2,3,7,8-tetrachlorodibenzodioxin. Fires have occurred on earth since land plants evolved 350 to 400 million years ago, and it is clear that many chloroorganic compounds have been natural components of our environment for a long time. In addition to combustion processes, volcanic emissions have been shown to contain significant amounts of a variety of chloroorganic compounds, including chlorinated alkanes (e.g., chlorinated propanes), chlorinated alkenes (e.g., PCE and TCE), chlorinated benzoates, and polychlorinated biphenyls (82, 145, 177). These natural chloroorganic compounds have been discharged into this planet’s biosphere over a period of billions of years, likely dating back to the very beginning of life on earth. In fact, the concentrations of chlorinated compounds in areas of high volcanic activity on early earth were probably elevated because no biological consumption, i.e., biodegradation, was yet occurring. Just like the cycling of other elements such as carbon, nitrogen, iron, or sulfur, the biogeochemical cycling of chlorine depends on microbial communities that degrade chlorinated compounds. The balance between production and consumption of chlorinated compounds maintains low, steady-state concentrations of chlorinated compounds in the environment. Such environments are called pristine. Anthropogenic influences have significantly changed the concentrations of certain chlorinated compounds in today’s environment, and the natural balance between production and consumption has been disturbed. Substantial efforts currently focus on understanding better the ecology and physiology of microorganisms that degrade haloorganic compounds, and exploiting their capabilities in bioremediation. The focus of this chapter is on the types of bacteria that transform chloroorganic compounds, a major group of halogenated pollutants with the most data available on their environmental cycling by dechlorinating bacteria. It should be noted, however, that other halogenated compounds are also of environmental concern, in particular fluorinated chemicals (e.g., fluorinated surfactants such as perfluorooctanyl sulfonate [PFOS]) (55, 88, 94). In summary, numerous animals, plants, algae, fungi, and bacteria, as well as combustion processes and volcanic activity, are responsible for the production

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of chloroorganic compounds. These processes occur in both terrestrial and marine environments, and their global production is significant.

2. EVOLUTION OF DECHLORINATING BACTERIA Chlorinated compounds were present in the environment prior to their anthropogenic production and release. Before human intervention, chloroorganic compounds did not accumulate in the environment because microorganisms had evolved strategies to benefit from the presence of these compounds, possibly in the earliest stages of microbial evolution. Chloroorganic aliphatic and aromatic compounds represent feedstocks in microbial food webs, serving as carbon and energy sources, or as terminal electron acceptors (chloridogenesis, see below). For the first 2 billion years on earth, oxygen was not available, and oxygenolytic dechlorination could not occur until oxygen-producing photosynthesis evolved and an oxygen-rich atmosphere developed. Early evolution was restricted to pathways that did not involve oxygen as a cofactor, and used electron acceptors other than oxygen. Based on thermodynamic considerations (see Chapter 4 by J. Dolfing), complete oxidation of chlorinated compounds coupled to the reduction of nitrate, ferric iron, sulfate, and other alternate electron acceptors, can be an energetically favorable process. However, not all theoretically feasible physiologies have actually been observed in the bacterial world (33). Novel enrichment strategies have helped to explore how thermodynamic considerations can predict the existence of yet unidentified physiological properties of microbes in nature, and indicate that a greater diversity of dechlorinating bacteria awaits discovery. Considering the long evolutionary time microbes have had to develop strategies to benefit from chloroorganic compounds, it is not surprising that dechlorinators are found in numerous groups of Gram-positive and Gram-negative bacteria. A number of bacteria that obtain energy for growth from the consumption of a chlorinated compound have been recovered over the last two decades from contaminated and uncontaminated (pristine) sites (44, 45, 133). These findings suggest that nature has indeed provided the appropriate conditions for long-term microbial adaptation and for the evolution of enzyme systems specific for chloroorganic compounds.

3. DECHLORINATION MECHANISMS Microorganisms have evolved different strategies to remove the chlorine substituent and to benefit from chlorinated compounds. Commonly observed mechanisms that are exploited by metabolically dechlorinating microorganisms to remove the chlorine substituent from an organic compound are depicted in Figure 3.1. Oxygenolytic dechlorination is restricted to aerobic conditions because oxygen is a required cofactor in catalysis. Hydrolytic and reductive dechlorination can occur under aerobic and anerobic conditions, although reductive dechlorination is most often observed in strict anaerobic environments. Representative of oxygenolytic dechlorination pathways are the degradation of 2chlorobenzoate (2-CBA) by Burkholderia cepacia strain 2CBS and by Pseudomonas sp. strain 273 (46,200). Oxygenolytic dechlorination can be mediated by diooxygenases or monooxygenases. The 2-CBA dechlorinase is a dioxygenase that

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incorporates both of the oxygen atoms from molecular oxygen into the product, whereas the 1,10-dichlorodecane dechlorinase is a monooxygenase incorporating only one oxygen atom into the product and reducing the other one to water. Chlorine substituents can also be replaced by hydroxyl groups derived from water. A well studied example is the hydrolytic dechlorination of 1,2-dichloroethane to 2chloroethanol by Xanthobacter autotrophicus GJ10 (83). Hydrolytic dechlorination is not limited to aliphatic hydrocarbons, and also occurs with aromatic compounds. The initial step in the degradation of 4-CBA by Pseudomonas sp. strain CBS3 involves an enzyme system that hydrolytically removes the chlorine substituent yielding 4hydroxybenzoate (96, 111, 163). Reductive dechlorination is the replacement of a chlorine substituent by hydrogen. Many highly chlorinated environmental pollutants that are resistant to oxygenolytic and hydrolytic dechlorination can be reductively dechlorinated (see chloridogenesis below). The first well-documented example of this type of reaction was the reductive dechlorination of 3-CBA to benzoate by Desulfomonile tiedjei DCB-1 (30, 179). A special case of a reductive dechlorination reaction, known as vicinal reduction or dichloroelimination, can occur when aliphatic chloroorganic compounds have chlorines located on adjacent saturated carbon atoms. An example is the reductive dechlorination of 1,2-dichloropropane to propene, which was observed in anaerobic mixed cultures (109). Specific examples of each of the commonly observed dechlorination reactions are shown in Figure 3.2. Common to all biologically-mediated dechlorination reactions is the release of chloride and protons. Hence, dechlorination is an acidifying process, and may result in a pH decrease in environments with low buffering capacity.

4. STRATEGIES OF MICROBIAL DECHLORINATION Bacterial metabolism is based on redox reactions, and bacteria have evolved strategies to oxidize or reduce chloroorganic compounds by using them as electron donors or electron acceptors, respectively (Figure 3.3). Bacteria benefit from chlorinated aromatic and aliphatic compounds in two ways: (A) by using them as a source of carbon and energy in the presence of a suitable electron acceptor (e.g., oxygen), thereby

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oxidizing the carbon skeleton completely to carbon dioxide, and (B) by using them as a terminal electron acceptor for energy generation. Diverse groups of bacteria completely oxidize chloroorganic compounds to generate carbon and energy, and many pure bacterial cultures have been isolated using a

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chloroorganic compound as the sole substrate under aerobic conditions (44, 45). The different microbial strategies, the evolution of different degradation pathways, and the dissemination of the genes encoding for these degradation pathways in the natural environment, is appropriately illustrated using chlorobenzoates, the commonly used herbicide 2,4-dichlorophenoxyacetate, and pentachlorophenol as examples.

4.1. Chlorinated Compounds as Carbon and Energy Sources

4.1.1. Degradation under Aerobic Conditions Chlorobenzoates A variety of bacteria, including Pseudomonas, Burkholderia, Sphingomonas, Alcaligenes, Micrococcus and Arthrobacter species, grow with chlorobenzoates as the sole source of carbon under aerobic conditions (44, 45). During growth, the chlorine substituent(s) can be removed in the initial enzymatic attack on the halogenated compound prior to the cleavage of the aromatic ring system (Figure 3.4). Most bacteria that utilize 4-CBA as a growth substrate possess the fcb genes encoding for the 4-CBA dechlorinase (consisting of 4-chlorobenzoate coenzyme A ligase, 4-chlorobenzoylcoenzyme A dechlorinase, and 4-hydroxybenzoyl-coenzyme A thioesterase) (25, 110, 111, 163). The chlorine substituent is removed in the initial degradation step from the aromatic ring system, yielding a non-chlorinated aromatic product, 4-hydroxybenzoate, which is then further degraded by standard degradation pathways (174).

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The chlorine substituent can also be removed from the aromatic ring system oxygenolytically, e.g., by the attack of dioxygenase enzyme systems on the chlorinesubstituted carbon. The resulting chlorohydrodiols are unstable and spontaneously eliminate HC1 to form the corresponding catechols. For instance, the cba operon encoding a 3,4-(4,5)-dioxygenase found in Alcaligenes sp. strain BR60 can result in initial dechlorination of 3-CBA and 4-CBA yielding protocatechuate (136). The 2-CBA 1,2-dioxygenases from Burkholderia cepacia 2-CBS and Pseudomonas putida P111 catalyze the dihydroxylation of 2-CBA in 1,2-position with concomitant release of chloride and carbon dioxide, yielding catechol (14, 64). On the other hand, CBA degradation pathways with chlorocatechols as intermediates also exist. Here, the chlorine substituent is released from aliphatic intermediates after cleavage of the aromatic ring system. Multiple pathways for CBA degradation have been found in a single organism, e.g., Pseudomonas putida P111. This strain degrades 3-CBA and 4CBA via chlorocatechols (removal of the chlorine substituent after ring cleavage), whereas the chlorine substituent from 2-CBA is removed in an initial dioxygenolytic attack to yield catechol (14). There is evidence that even more pathways for CBA degradation exist. For example, Alcaligenes sp. strain L6 metabolizes 3-CBA via gentisate (100). In a recent study, less than 10% of the isolates obtained with 3-CBA or 4-CBA as the growth substrate hybridized to gene probes targeting sequences encoding the known pathways (144). This suggests that other, as yet unknown genes may be involved in CBA dechlorination. Although Table 3.1 is not a complete list of the numerous isolates obtained with 4-CBA, it becomes obvious that 4-CBA degradation pathways are distributed among several bacterial groups. Gram-positive bacteria and different subdivisions of the Proteobacteria have similar pathways, which suggests lateral transfer of the gene cassettes coding for 4-CBA degradation between phylogenetically distant populations. A basis for the genetic transfer and relatedness of divergent strains is shown through a number of recent studies that have demonstrated that the operons with gene cassettes involved in CBA degradation can be found on the chromosome or transmissible elements, e.g., plasmids or transposons. For instance, the fcb genes encoding for the hydrolytic 4-CBA dehalogenase were found on plasmid pSS70 in Alcaligenes sp. strain ALP83 (104), but were located on the chromosome in other strains, such as Pseudomonas sp. DJ-12 (17). A detailed sequence analysis of the fcb genes constituting the 4-CBA dehalogenase revealed that they may have evolved from genes encoding for proteins involved in preexisting pathways (39). The high degree of sequence similarity among the genes coding for the 4-CBA-CoA ligase and the 4-chlorobenzoylCoA dehalogenase suggest that different bacteria share common ancestoral genes, and that the fcb genes did not evolve separately, but may have been spread by horizontal (lateral) gene transfer. Another example of horizontal gene transfer is the cbaAB operon, encoding a 3-CBA 3,4-[4,5]-dioxygenase that is carried on plasmid pBRC60, which was found in Alcaligenes sp. strain BR60. Lateral transfer of pBRC60 to species of both the and of the Proteobacteria was demonstrated in microcosm experiments (51). Further investigation of this system demonstrated that the genes specifying 3-CBA degradation are located on the catabolic transposon Tn5271 (31, 135, 203).

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Interestingly, virtually identical cbaAB operons appear to be distributed globally, indicating that transposition events may occur frequently, and may be the driving force behind the horizontal dissemination of catabolic genes cassettes (134). Similarly, genes encoding for chlorobenzene, chloroacetate, and chlorobiphenyl degradation were found

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on transposon-like elements (92, 173, 193). The clc genes encoding the chlorocatechol ortho ring cleavage pathway in Pseudomonas putida P111 were found on the chromosome, whereas the CBA 1,2-dioxygenase present in the same strain was shown to be encoded on plasmid pPB 111 (14). Interestingly, in a variant strain ofP111, the clc operon was translocated into a plasmid (14). Recent work with Pseudomonas sp. strain B13 demonstrated that the clc element includes sequences encoding for integrase functions, emphasizing that mobile DNA elements may play an important role in the horizontal transfer of genes encoding for catabolic pathways (192). Another example that stresses the relevance of horizontal gene transfer is found in haloalkane-degrading bacteria. Rhodococcus rhodochrous NCIMB13064, Pseudomonas pavonaceae 170, and Mycobacterium sp. strain GP1 are phylogenetically distant organisms, however, they share a highly conserved haloalkane dehalogenase gene (dhaA), which allows these strains to degrade 1-chlorobutane, 1,3-dichlorobutene, and 1,2-dibromoethane, respectively (147). Flanking regions encoding the dhaA gene contained sequences encoding for integrase function. Other examples of catabolic pathways for chlorinated compounds that are encoded on transmissible elements are shown in Table 3.2.

2,4-Dichlorophenoxyacetate (2,4-D) The catabolism of the broad-spectrum herbicide 2,4-D is another example that demonstrates the evolution and dissemination of degradation pathways. 2,4-D has been widely studied since its initial use during the 1940s for agriculture, land management and urban needs. Shortly after 2,4-D is applied to target plants, it is readily degraded by the combined efforts of microbial communities and numerous individual bacterial populations found in soil (40, 50, 85). Interesting discoveries about the organization, regulation and assembly of2,4-D catabolic pathways have been made in recent decades. In this section, the diversity of 2,4-D-degrading bacteria is summarized, along with evidence for the evolution and transfer of2,4-D degradation pathways between bacterial populations (69, 117, 124, 137, 184). The bacteria that degrade 2,4-D span several bacterial divisions and many bacterial genera. The first pure cultures of2,4-D-degrading bacteria were isolated from soils, and grouped among the high G+C Gram-positive bacteria. Examples included an Arthrobacter sp. (116) and a Corynebacterium sp. (151). As more 2,4-D-degrading bacteria were isolated throughout the 1980s and 1990s, an interesting trend was seen. A large number of representative isolates were found among the of the Proteobacteria, and far fewer populations were characterized from other bacterial groups (37, 50). Pseudomonas, Alcaligenes, Ralstonia and Burkholderia are that can expand their metabolic range by using genes on broad host range plasmids. Interestingly, many of the 2,4-D-degrading populations are also closely related to chlorobenzoate (CBA)- and chloroatechol-degrading strains. Often the 2,4-D-degrading populations, like Pseudomonas sp. 965 and Ralstonia eutropha JMP 134, have the ability to degrade 3-CBA or other chlorinated compounds (38,85). In addition to the frequently encountered Sphingomonas species and haloalkalophilic bacteria Flavobacteria (Bacteroides-CytophagaFelexibaxter group) and Nocardioides (high G+C Gram-positive group) were also described to degrade 2,4-D. Table 3.3 shows selected examples of the wide variety of

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bacteria that are known to completely mineralize 2,4-D under aerobic conditions. Many were obtained from agricultural soils and sites with prior exposure to the herbicide. The bacteria that utilize 2,4-D possess pathways that are encoded by genes residing on chromosomes or on mobile genetic elements such as plasmids, transposons and insertion elements. Mobile genetic elements are powerful means of disseminating favorable combinations of genes among bacteria. They allow for inherited properties to be passed onto offspring, as well as across population boundaries. The mobile elements provide a means of assembling combinations of genes that developed in different bacterial lineages. For example, the proteobacterial populations that degrade 2,4-D often possess this trait because of a mobile genetic element that originated in another bacterial line. Over time, the genes evolve in the new host, and the modified genes may again find their way to another population. Slightly different variations of the genes and pathways become widely distributed among the populations that benefit from the new gene combinations. The mobile genetic elements, then, act as reservoirs for gene cassettes involved in utilizing 2,4-D and other chlorinated aromatic compounds. The ability to utilize chlorinated aromatic compounds like 2,4-D is a cumulative ability, created by stacking together mobile gene cassettes, each specifying a particular function. Initial examination of the diversity of 2,4-D degradative genes took advantage of the tfd gene sequences obtained from plasmid pJP4 of Ralstonia eutropha. Plasmid pJP4 carries five of the crucial genes encoding for enzymes involved in the degradation

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of 2,4-D, as well as the regulatory elements. As increasing numbers of tfd genes were analyzed, unexpected diversity in gene sequences and organizational patterns emerged from each newly discovered population. Also, it appeared that contemporary genes of the 2,4-D pathway diverged from ancient common ancestors, which can be taken as evidence that substrates resembling 2,4-D have been utilized for a large span of bacterial history. In Figure 3.5, the mosaic pattern of gene assembly found among some representative 2,4-D-degrading isolates is illustrated, and when known the location of the genes, whether plasmid-borne or chromosomal, is indicated. Among the Proteobacteria, the degradative genes might be integrated into the chromosome or on plasmids, and sometimes a portion of the pathway can exist in either location. As opposed to the diverse genetic organization of the other genera, e.g., the sphingomonads, tend to carry their 2,4-D degradation genes (spa genes) on the chromosome (85). The initial 2,4-D degradation step on plasmid pJP4 is encoded by the gene tfdA. An dioxygenase removes the acetyl group from 2,4-D to form dichlorophenol and glyoxylate (48, 49, 89, 90). Genes that resemble the tfdA gene of plasmid pJP4 have been found on other plasmids such as pEMTl, pEMT3, and pEST4001, as well as in multiple bacterial populations (e.g., Ralstonia, Pseudomonas). There are also at least two different lineages of tfdA genes. While many bacteria have tfdA genes that are nearly identical to those of plasmid pJP4, some Burkholderia strains utilize alternative tfdA genes that retain similar activity toward 2,4-D (50, 181). The Burkholderia genes are not limited to the chromosome, and can move around on mobile genetic elements as well. Other plasmids contain only a single gene of the 2,4-D degradative pathway, indicating that independent elements are maintained as genetic reservoirs that can supplement existing degradation pathways (185). As suggested by Figure 3.5, not all bacteria that degrade 2,4-D have a tfdA gene that can be detected with the available gene probes, which implies even greater diversity than described here. In the second degradation step, a hydroxyl group is added to the aromatic ring by way of the gene product of tfdB. Analogous to the diversity in tfdA genes, there is considerable diversity in tfdB genes. Different tfdB genes can be found in combination with the same tfdA gene (e.g., on plasmids pEMT3 and p JP4), or the same tfdB gene can function with different tfdA genes. In some strains, like Ralstonia eutropha JMP134, evolution of tfdA has occurred in parallel with tfdB, while in other populations both genes appear to have evolved independently, and were assembled more recently. Even greater diversity is seen in the genes of the tfdCDEF operon responsible for ring cleavage and subsequent transformation reactions, including the first dechlorination step. These genes are closely related to the chlorocatechol (clc) degradative genes described previously. The tfd and clc genes can even share the same regulatory elements, which often transfer together with the catabolic genes (107). Sequence level variation as well as the possible origins and evolution of 2,4-D catabolic genes and pathways are compared in detail in an article by Valleyes et al, (189). Clearly, great variation exists in the tfd genes of 2,4-D-degrading bacteria. And moreover, any individual organism can contain an independent assortment of tfd genes. Lateral gene transfer is responsible for shuffling 2,4-D-degrading gene cassettes as they ride across species boundaries on catabolic plasmids. Ka et al. found a 2,4-D catabolic plasmid pKA2 that integrated into the chromosome of Alcaligenes paradoxus (86). The

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plasmid could excise from the chromosome to again exist freely as a transmissible plasmid. Lateral transfer of genes through recombination events was also shown with the well-studied plasmid pJP4 that has an insertion element (ISp JP4) in the regulatory region of the tfdCDEF operon. The ISpJP4 element was able to transpose and move tfd genes in laboratory experiments (107, 108). It was also found that Burkholderia cepacia CSV90 harboring plasmid pMABl, recruited the chlorocatechol-degrading genes (tfdCDEF) using the same insertion element that appears on plasmid pJP4 (9, 189). Furthermore, the 2,4-D genes on plasmid pJP4 appear to have experienced a duplication event, and since the two sets of tfd genes are different, it provides indirect evidence that new pathways may arise by subsequent duplication and modification of preexisting pathways (54, 103). Why is there so much diversity in the 2,4-D degradative genes? Possibly, it is because different pathways evolved to degrade the assortment of naturally occurring chlorinated phenolic products. Compelling evidence supports the divergence of tfd genes from ancient origins, with subsequent rounds of assembly and evolution. Each of the independent gene cassettes can be recruited in order to assemble a complete and functional 2,4-D degradation pathway. Further evolution of the duplicated genes leads to divergence of catabolic genes. Independently evolved degradation pathways assemble by way of insertion elements or catabolic plasmids that transfer laterally to other microbial populations. Consequently, different organisms possess a mosaic of 2,4-D degradation genes, often arising by recombination of genes from divergent sources. In addition to detailed genetic characterization, biochemical and immunological studies of dechlorinating enzyme systems provide additional evidence for lateral gene transfer. For instance, the comparison of dichloromethane dechlorinases from different aerobic methylotrophs capable of using dichloromethane as a growth substrate revealed high similarities among dechlorinating enzyme systems. Again, these observations suggest that the structural genes for the dichloromethane dechlorinases were laterally transferred among methylotrophic populations and have not evolved separately (97). In summary, three relevant mechanisms for lateral transfer of catabolic (dechlorinating) genes or gene cassettes have been identified: conjugative plasmids, transposons, and integrons (178, 187, 203). The genetic environments of most catabolic genes involved in dechlorination and degradation of halogenated compounds are largely unknown. Progress in microbial genomics will reveal whether dissemination of catabolic genes among microbial populations via transmissible elements is a major driving force behind the diversity of dechlorinating populations, or if separate evolution is a more common theme. Pentachlorophenol

Pentachlorophenol (PCP) has been widely used as a wood preservative for decades, and is a major environmental pollutant. Despite its toxicity and high degree of chlorination, several aerobic PCP-degrading bacteria were isolated (Table 3.4). The enzymes and genes involved in the PCP degradation pathways have only recently been elucidated, or are still awaiting isolation and complete characterization. The aerobic PCP degradation pathway in some species involves oxygenolytic, reductive, and hydrolytic dechlorination reactions prior to ring cleavage (Figure 3.6).

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Interestingly, gene sequences encoding for the monooxygenase catalyzing the initial hydroxylation step were detected in bacteria that failed to degrade PCP (157). Further studies on the pathway and the progress in bacterial genomics will help to explain the evolution and the recruitment of genes for assembling this complex pathway. It will be interesting to see if similar patterns emerge for the PCP degradation pathways as were found for the tfd genes. PCP can also be degraded under strict anaerobic conditions, which will be discussed below.

4.1.2. Non-Oxygen Respiring Dechlorinating Organisms The ability to utilize chlorinated compounds as a source of carbon and energy is not limited to aerobic bacteria. Recent research demonstrates that pure cultures of denitrifying bacteria are also capable of completely oxidizing halogenated aromatic compounds, including chlorobenzoates. Several pure cultures of Proteobacteria belonging to the genera Thauera Ochrobactrum and Pseudomonas were isolated with 3-CBA providing the sole source of carbon and energy under denitrifying conditions (66, 172). Using the same starting materials, degradation of 4-CBA under denitrifying conditions was also observed, although no pure cultures could be obtained. Interestingly, degradation of 2-CBA was not detected under denitrifying conditions (172). Despite extensive efforts, enrichments to obtain denitrifying chlorophenol degraders were unsuccessful. Anaerobic conditions merely stimulated reductively dechlorinating populations, even though this process is energetically less favorable than complete oxidation coupled to denitrification (160). The complete mineralization of chlorinated compounds under iron(III)-reducing, sulfidogenic, or methanogenic conditions has also been observed (11,12,65). However, no pure cultures have been obtained so far, and it is unclear whether a single population is sufficient, or if a consortium is necessary to completely oxidize chloroorganic compounds under the specified redox conditions. Circumstantial evidence suggests the involvement of multiple populations, because reductive dehalogenation precedes the complete oxidation of the remaining carbon skeleton (65). The ability of phototrophic bacteria to use chloroorganic compounds under anoxic conditions is not well explored but might be widely distributed. The type strains of the purple nonsulfur species Rhodospirillum rubrum, Rhodospirillum photometricum, and Rhodopseudomonas palustris grow phototrophically on a variety oftwo- and three-carbon halocarboxylates, including chloroacetate, 2-chloropropionate,

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and 3-chloropropionate. In each case, degradation was initiated by a reductive dechlorination step (128). Different Rhodopseudomonas palustris strains degraded 3-CBA with stoichiometric release of chloride under anaerobic conditions when incubated in light (43,194). The chlorine substituent was released in a reductive process from 3-chlorobenzoyl-CoA rather than 3-CBA (43). Specialized acetogenic populations use dichloromethane and chloromethane as sole sources of carbon and energy. Dehalobacterium formicoaceticum strain DMC belongs to the Bacillus/Clostridium subphylum of the low G+C Gram-positive bacteria, and ferments dichloromethane to acetate and formate. Dichloromethane has the same carbon oxidation state as formaldehyde, and can feed directly into the acetogenic pathway. Strain DCM is highly specific for dichloromethane as a substrate and grows best in association with a formate-consuming population (118). Acetobacterium dehalogenans (strain MC, DSM 11527), a low G+C Gram-positive methylotrophic homoacetogen, grows with chloromethane as the sole source of carbon and energy and produces acetate as the main fermentation product (129, 130, 186, 195). Acetobacterium dehalogenans, however, can not use dichloromethane as a substrate, and chloroform and carbon tetrachloride inhibited growth with chloromethane.

4.2. Chlorinated Organic Compounds as Electron Acceptors (Chloridogenesis ) Another strategy commonly observed in anaerobic environments is reductive dechlorination. Here, certain groups of anaerobic bacteria take advantage of chlorinated compounds by using them as favorable terminal electron acceptors for the oxidation of available substrates (i.e., electron donors). Under anaerobic conditions the availability of suitable terminal electron acceptors for energy generation often limits bacterial growth and the degradation of organic compounds. In such environments bacteria not only compete for oxidizable substrates as carbon sources, but also for favorable terminal electron acceptors. Reductive dechlorination is an electron consuming process, and the energy available from reductive dechlorination reactions is substantial, making chlorinated compounds good electron acceptors under anaerobic conditions (see Chapter 4) (33-36, 68). Desulfomonile tiedjei strain DCB-1, a was the first organism obtained in pure culture that derived all of its energy required for growth from the reductive dechlorination of 3-CBA to benzoate (Figure 3.7) (30, 169, 179). The process of reductive dechlorination as a respiratory electron accepting reaction

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was originally designated chlororespiration (113, 127, 159, 171), but the term dechlororespiration (3, 79, 126) is found in literature as well. Both terms describe exactly the same process although the names indicate that one is the opposite of the other. The term respiration implies a change in redox state, however, the chloride ion that is liberated in (de)chlororespiration is neither oxidized nor reduced, and therefore both terms are misleading. In addition, the term chlororespiration is often confused with the respiratory electron transfer system in plant chloroplasts (140). For clarification, we propose the term chloridogenesis to describe reductive dechlorination processes that are linked to energy conservation, and organisms with this capability are termed chloridogens. Figure 3.8 illustrates the key physiology of a chloridogenic population. It was several years after the isolation of Desulfomonile tiedjei that additional chloridogens were isolated, possibly because enrichment procedures with chloroorganic compounds as the only available electron acceptors were not consequently pursued.

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Since the mid 1990’s the number of isolates capable of using chlorinated compounds as metabolic electron acceptors steadily increased, and it is likely that the number of new isolates will continue to rise (Figure 3.9). Chloridogens are distributed among the bacterial domain, and are found in different lineages. The phylogenetic relationships of known chloridogenic bacteria are shown in Figure 3.10. Many chlorinated compounds are found on the USEPA list of priority pollutants. Of particular environmental concern are polychlorinated compounds such as chlorinated solvents, chlorinated phenols, chlorinated benzenes, chlorinated biphenyls, and chlorinated pesticides, Due to the oxidized nature of polychlorinated compounds, the electrophilic attack of oxygenase enzyme systems is hindered, and many highly chlorinated chemicals are not efficiently metabolized under aerobic conditions. Consequently, the discovery of chloridogenesis initiated intense research focusing on anaerobic populations that use chlorinated compounds as respiratory electron acceptors under anaerobic conditions. Several isolates using (poly)chlorinated priority pollutants as metabolic electron acceptors have been recently obtained.

4.2.1. Chlorophenols as Electron Acceptors Desulfitobacteria belong to the Bacillus / Clostridium group within the low G+C Gram-positive bacteria. Members of the genus Desulfitobacterium are commonly

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isolated with chlorophenolic compounds or PCE as electron acceptors. With one exception (190), all isolates are capable of chloridogenic growth with at least one chlorinated compound. Despite physiological similarities, strain differences exist and the dechlorinating abilities differ among members of the genus. Table 3.5 lists different desulfitobacteria, as well as the chlorinated electron acceptor that was used for each isolation procedure. The genus was designated Desulfitobacterium because the first isolate, Desulfitobacterium dehalogenans, grows with sulfite, but not with sulfate, as a terminal electron acceptor (188). Other isolates are also able to use sulfite as an electron acceptor, and only one strain, the PCE-to-cis-l,2-dichloroethene (cis-DCE)-dechlorinating strain PCE-S, failed to grow with sulfite (79). Desulfitobacteria are nutritionally versatile with regard to electron donor and acceptor utilization. Ortho-, meta-, and para-dechlorination of chlorophenolic compounds have been observed with desulfitobacteria, however, strain-specific differences exist, and the available isolates can be distinguished by the spectrum of chloroorganic compounds used to support growth. Only a few isolates, including Desulfitobacterium strain PCE1 and Desulfitobacterium frappieri strain TCP-A, are capable of complete dechlorination to phenol (13, 53). Some strains use chlorophenolic compounds and chloroethenes as electron acceptors, while others use only chlorophenolic compounds or only chloroethenes (see Table 3.5). There is evidence that different enzyme systems are responsible for chloroaryl and chloroalkyl reduction, and that the reductive dechlorinases are specifically induced by the respective substrates (52,132). The PCE-dechlorinating Desulfitobacterium isolates produce TCE or cis-DCE as end products, and none of the available pure cultures can dechlorinate beyond cisDCE. Desulfitobacteria were found in diverse anaerobic environments and are probably widely distributed in nature. They are robust organisms, relatively easy to grow under laboratory conditions, and endospores have been reported in at least some strains. These properties are desired for use in bioremediation approaches. Interesting to note is the occurrence of multiple 16S rRNA operons in Desulfitobacterium strains, e.g., extended 16S rDNA sequences resulting in modified secondary structures were observed (10,41). Enrichment with monochlorophenols, e.g., 2-chlorophenol, typically results in the isolation of other groups of chloridogens, such as Desulfovibrio dechloracetivorans SF3, a marine isolate, or strains of Anaeromyxobacter dehalogenans (24,115,158,180). The anaerobic myxobacteria are unique chloridogens because they are facultative anaerobes, and use acetate as well as hydrogen as electron donors for reductive dechlorination (158). Reductive dechlorination of 3-chlorophenol has also been observed in methanogenic consortia (84, 115). Anoxic degradation of phenol can occur under varying redox conditions, and hence, the complete anaerobic degradation of polychlorinated phenols is possible (75).

4.2.2. Polychlorinated Biphenyls (PCBs) as Electron Acceptors Highly chlorinated biphenyls resist aerobic degradation but can be reductively dechlorinated under anaerobic conditions. Similar to the observations made with polychlorinated phenols, diverse PCB dechlorination patterns have been seen in microcosms studies (8, 183, 197). Circumstantial experimental evidence suggests that PCBs are used as terminal electron acceptors by chloridogenic bacteria. The isolation

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of pure cultures, however, has proven to be very challenging, and none are available to date. PCBs have a very low aqueous solubility, and hence, growth of PCB-respiring bacteria is limited by electron acceptor availability. Establishing sediment-free cultures was a difficult hurdle to overcome but has been recently accomplished (26,201). A 16S rDNA-based community analysis of a 2,3,5,6-tetrachlorobiphenyl-dechlorinating, sediment-free culture revealed the presence of a Dehalococcoides species (80). This is an interesting observation because the three known Dehalococcoides isolates are obligate hydrogenotrophic chloridogens, and one can speculate about the involvement of a Dehalococcoides population in PCB dechlorination. This hypothesis was recently substantiated using a 16S rDNA-based approach, and a green non-sulfur bacterium (e.g., a Dehalococcoides population) was identified to catalyze double-flanked chlorine removal from 2,3,4,5-pentachlorophenol (202). Desulfitobacterium dehalogenans reductively dechlorinated hydroxylated PCBs, however, dechlorination was not seen with PCBs lacking the hydroxyl group (198). Nevertheless, this is a very encouraging finding, as it demonstrates that PCB dechlorination is possible with pure bacterial cultures.

4.2.3. Chlorinated Solvents as Electron Acceptors Chlorinated solvents are among the most abundant groundwater pollutants, many of which are not degraded efficiently under aerobic conditions. Of particular environmental concern are chlorinated ethenes. Considerable efforts over the last decade resulted in the isolation of bacteria that can reductively dechlorinate chlorinated ethenes. Table 3.6 lists chloridogenic bacteria that use chloroethenes as electron acceptors. Several isolates that dechlorinate PCE to TCE or cis-DCE were obtained from contaminated and pristine sites. These isolates belong to different phylogenetic groups, including the Gram-positive bacteria and different subdivisions of the Proteobacteria. The PCE-to-cis-DCE-dechlorinating enzyme systems and/or the encoding gene sequences have been characterized from Dehalospirillum multivorans (138, 139), Dehalobacter restrictus (165), Desulfitobacterium sp. strain PCE-S (132), Clostridium bifermentans (143), and Dehalococcoides ethenogenes (119,120). Common to all PCE reductases investigated so far are iron-sulfur clusters and a corrinoid cofactor, which are involved in the catalytic cycle of the dechlorination reaction. Molecular analysis of the gene sequences encoding for different reductive dehalogenases suggests that these enzymes constitute a novel class of redox enzymes (171). The evolution of genes coding for the reductive dehalogenases is currently unknown. Once additional sequences become available, it will be very interesting to see if the reductive dechlorinase genes have deep evolutionary branches, and how they have been disseminated among reductively dechlorinating populations. Dehalococcoides ethenogenes is a fascinating bacterium which was isolated at Cornell University (126). This unusual bacterium is not affiliated with any known bacterial lineage, and appears to be most closely related to the green non-sulfur bacteria. This organism has an extremely limited substrate range, and can only be grown with hydrogen as the electron donor and chloroethenes (PCE, TCE, cis-DCE, and 1,1 -DCE) or 1,2-dichloroethane as electron acceptors. Vinyl chloride (VC) and trans-DCE do not support growth but VC is dechlorinated cometabolically to ethene at low rates.

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Nevertheless, Dehalococcoides ethenogenes was the first isolate that completely dechlorinated PCE to the non-toxic end product ethene (125). Another Dehalococcoides species, designated strain FL2, was isolated from a TCE-to-ethene-dechlorinating mixed culture (114). Dehalococcoides. sp. strain FL2 lacks the ability to dechlorinate PCE at high rates, but dechlorinates TCE, cis-DCE, and trans-DCE rapidly to VC and ethene (74). Like with Dehalococcoides ethenogenes, the final dechlorination step to ethene occurrs at a much lower rate. Hence, current efforts focus on populations that use VC as electron acceptor and exhibit high dechlorination rates. Circumstantial evidence strongly suggests that VC-respiring populations exist. For instance, several highly enriched VC-dechlorinating cultures consumed to threshold concentrations below 0.4 ppmv. In contrast, the same cultures that were not fed with VC did not consume to concentrations below 100 ppmv, indicating that VC supported a terminal electron accepting process (115). An exponential increase in VC dechlorination rates was also observed, which provides additional evidence that VC reduction to ethene is linked to growth of the dechlorinating population(s) (115,153). The 16S rDNA-based community analysis of VC-dechlorinating cultures that were repeatedly transferred to mineral salts medium with VC as the only available electron acceptor revealed the presence of multiple Dehalococcoides species (74, 99). Interestingly, the VCdechlorinating cultures failed to dechlorinate PCE or TCE, suggesting that these Dehalococcoides species are different from Dehalococcoides ethenogenes and Dehalococcoides sp. strain FL2 (47). Recently, a third Dehalococcoides isolate, strain CBDB1, was described. This organism was isolated using trichlorobenzenes as electron acceptors (3). These reported observations suggest that Dehalococcoides species are diverse, exhibiting different substrate specificities, and might be involved in the reductive dechlorination of a variety of chlorinated compounds. It should be noted that the three Dehalococcoides populations available as pure cultures have no regular peptidoglycan-containing cell wall, and are resistant to ampicillin and vancomycin. Interestingly, several 16S rDNA sequences affiliated with the Dehalococcoides cluster were retrieved from diverse habitats, including a deep subsurface environment (18), an oligotrophic open ocean environment (56), a contaminated aquifer (32), Baltimore Harbor sediment (80), anaerobic sludge from a lagoon of a distillery (57), freshwater sediment from the Saale River (196), and the Obsidian Pool (a hot spring in Yellowstone National Park) (81). Only speculation is possible regarding the physiologies of the populations represented by the environmental clones affiliated with the Dehalococcoides cluster. It is important to note, however, that the clones pNl-52, SAR202, and SAR307 represent open ocean bacterioplankton (56). As mentioned earlier, extensive amounts of halogenated compounds are constantly produced in the oceans, and it is tempting to hypothesize that these uncultured populations play a relevant role in the biogeochemical cycling of chloroorganic compounds in marine environments.

4.3. Electron Donors for Chloridogens Most known bacterial populations with the ability to use a chlorinated compound as a terminal electron acceptor utilize hydrogen as electron donor. Some dechlorinating isolates are very versatile with regard to the electron donors used for reductive dechlorination, e.g., desulfitobacteria, Dehalospirillum multivorans, Clostridium

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bifermentans, etc. can couple reductive dechlorination of PCE to the oxidation of a variety of electron donors, including hydrogen, lactate, pyruvate. None of these organisms, however, can use acetate as an electron donor for reductive dechlorination. In contrast, PCE-dechlorinating Desulfuromonas species oxidize acetate but are unable to couple reductive dechlorination to hydrogen oxidation. Unique with regard to electron donor utilization are the chlorophenol-dechlorinating Anaeromyxobacter dehalogenans strains, which can use both hydrogen and acetate as a source of reducing equivalents (158). Dehalococcoides and Dehalobacter species are the least metabolically versatile groups of chloridogens and depend on molecular hydrogen as their electron donor. Desulfovibrio dechloracetivorans SF3 uses a variety of electron acceptors, including sulfate, sulfite, thiosulfate, nitrate, and 2-chlorophenol. Interestingly, acetate support the reductive dechlorination of 2-chlorophenol, but strain SF3 failed to couple the reduction of any of the other physiological electron acceptors to acetate oxidation. These observations suggest that the spectrum of electron donor utilization in strain SF3 depends on the terminal electron accepting process (180). Acetate and hydrogen are the major degradation products of organic matter in anaerobic environments. In the absence of alternate electron acceptors, hydrogen is typically consumed by methanogens for the reduction of carbon dioxide to methane, and acetate can be fermented to methane and carbon dioxide by acetoclastic methanogens. Hydrogenotrophic chloridogens are excellent competitors for molecular hydrogen, and can use hydrogen at concentrations that are too low to support methanogenic activity (115,205). Similar observations have been made with acetotrophic PCE-dechlorinating Desulfuromonas species that reduced the acetate concentration below the level that supported acetoclastic methanogenesis in Methanosarcina barkeri (Sanford & Löffler, unpublished). An intriguing observation was made with Trichlorobacter thiogenes (29). This organism grows with trichloroacetate as electron acceptor, converting it to dichloroacetate in the presence of acetate and sulfide. Surprisingly, acetate was not the direct electron source for reductive dechlorination. Trichlorobacter thiogenes uses sulfide as the source of reducing equivalents for dechlorination, thereby forming sulfur, which occurs as intracellular granules. Sulfur reduction to sulfide, in turn, is coupled to acetate oxidation regenerating sulfide, the electron donor for reductive dechlorination. Trichlorobacter thiogenes belongs to the of the Proteobacteria, and acetate oxidation coupled to sulfur reduction is a common feature of closely related Geobacter and Desulfuromonas species. The PCE dechlorinator Desulfuromonas michiganensis strain BB1 reduces sulfur to sulfide in the presence of acetate, however, has failed to dechlorinate with sulfide as the electron donor (Löffler, unpublished). Sulfide, the end product of sulfate reduction, is frequently found in anaerobic environments, and it remains to be seen how common the ability to use sulfide as an electron donor for reductive dechlorination is in nature.

5. CONCLUDING REMARKS Dechlorinating bacteria are members of the different subdivisions of the Proteobacteria, the low and high G+C Gram-positive bacteria, the Cytophaga, and the green non-sulfur bacteria. Figure 3.11 shows the number of isolates of each phylogenetic

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affiliation that is currently (www.cme.msu.edu/BSD).

listed in the Biodegradative

Strain

Database

Several phylogenetic branches within the domain Bacteria currently contain no recognized dechlorinating populations, and to date no dechlorinators are described in the domain Archaea. Considering the minute fraction of microorganisms that has been so far cultured (4), and the fact that the dechlorinating abilities ofavailable cultures have not been comprehensively explored, it is likely that a large number of new dechlorinating species await discovery. Innovative culture techniques and enrichment conditions are needed to obtain novel cultures and isolates, Microbial genomics will provide a basis to decipher the evolution of dechlorinase genes, and shed light on the mechanisms of how these genes are disseminated among bacteria within the same phylogenetic group and beyond. This information will help to exploit dechlorinating microorganisms in environmental biotechnology, e.g., bioremediation, biotransformations, and to more comprehensively understand the function and ecology of microorganisms in the natural cycling of chlorine. REFERENCES 1.

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Chapter 4 THERMODYNAMIC CONSIDERATIONS FOR DEHALOGENATION JAN DOLFING Alterra, Wageningen University and Research Centre, Wageningen, The Netherlands

1. INTRODUCTION Insight into the energetics of a microbial process helps to understand the logic behind the process and the ecology of the organisms that catalyze it. The present chapter focuses on the energetics of dehalogenation reactions, using thermodynamics as a tool. Thermodynamics gives us insight into the amount of energy that is available from a dehalogenation reaction, or how far from equilibrium that reaction is. Thus, thermodynamics is used to predict whether an organism can in principle obtain energy for growth from catalyzing a certain reaction. Thermodynamics does not give insight into how much energy an organism actually captures, or how much it has to invest to capture this energy. It does help to explain why and which fortuitous reactions (40) take place. For example, when the system is out of equilibrium an organism can fortuitously help to move the system towards equilibrium. Fortuitous in this context means that organisms catalyzing the reaction do not obtain energy from it. The second application of thermodynamics is as a tool to assist in formulating a framework to rationalize dehalogenation pathways. Here, an analogy to the global biogeochemical redox concept can be drawn, whereby is preferentially used as electron acceptor, then nitrate, then sulfate, and finally carbonate. Microorganisms will follow a similar redox sequence also on a more subtle scale, and preferentially catalyze those reactions that have the highest energy yield. Reasoning along these lines, one can predict the likelihood of certain pathways associated with specific environmental conditions, in particular, prevailing redox conditions and the type and availability of electron acceptor. In the past two decades we have learned that: (i) most halogenated compounds can be dehalogenated under anaerobic conditions, (ii) the type of reaction involved is generally (but not always, see below) a hydrogenolysis, and (iii) in many cases, the microorganisms that mediate the reaction obtain energy for growth (14; see Chapters 3 and 5). Dehalogenation via hydrogenolysis is a reductive process, which requires reducing equivalents for the reaction to proceed. The energetic value of reducing equivalents depends on the redox potential of the environment, and is highest under aerobic conditions, i.e., with as electron acceptor. Based on these assumptions, it is Dehalogenation: Microbial Processes and Environmental Applications, pages 89-114 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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logical that reductive dehalogenation is not often observed under oxic conditions. There is, however no a priori (thermodynamic) reason why reductive dechlorination could not be possible in the presence of The objective of the present chapter is to provide a thermodynamic framework to rationalize these observations, and to discuss the potential ecophysiological consequences within this framework, including the hypothesis that anaerobic microorganisms exist that benefit from dechlorination via mechanisms other than hydrogenolysis. Brief attention will also be paid to the question of why highly halogenated compounds, like terra- (per-) and trichloroethylene, do not appear to be used by aerobic microorganisms as a sole source of carbon and energy.

2. CALCULATION METHODS FOR Gibbs free energy values, or better, values for changes in Gibbs free energy ( values), provide a useful tool to evaluate whether a reaction is exergonic of endergonic, i.e., whether or not an organism can obtain energy for growth from catalyzing such a reaction (37). The use of changes in Gibbs free energy values has a long and fruitful tradition in microbial ecology. Bryant et al., for example, in their landmark paper on Methanobacillus omelianskii (4), used thermodynamics to point out why interspecies hydrogen transfer (25) was the mechanism behind the growth of this syntrophic co-culture. Anaerobic microbiology subsequently benefited from the excellent review as well as comprehensive tables of values of physiological reactions listed by Thauer et al. (37). In conjunction with the increasing interest in the degradation of halogenated compounds in the last two decades, values have been estimated and tabulated for a wide array of both aromatic and aliphatic halogenated compounds (15, 17, 23, 24). For actual values, the reader is referred to the primary literature. The procedures to calculate values for microbial reactions are well described in easily accessible texts such as, “Brock Biology of Microorganisms” (30). Briefly, calculations of the changes in Gibbs free energy of a system are made according to the equation:

The naut sign (°) indicates that the calculations are made for standard conditions, i.e., concentrations of 1 molar, temperature of 25°C and a partial pressure of 1 atmosphere for gases. For evaluations under actual physiological conditions, this is not really relevant. In biological systems, equilibrium between gas and liquid phases is assumed. For example, at a partial pressure of 1 atm is in equilibrium with an aqueous concentration of . This is taken into consideration in the calculation of the value for (See Box 4.1 for details). Deviation from equilibrium is calculated according to the Nernst equation, because the change in Gibbs free energy values of a reaction are directly linked to its equilibrium constant. Chemists prefer the value to remain as simply derived (“clean”) as possible; e.g., for a compound that is normally a gas at 25°C, should be given for the gaseous state of the compound. However, for proper evaluation of the in microbiological systems, it is more straightforward to directly measure and calculate for the aqueous phase, even when the conversion of the value from the gaseous to the aqueous phase introduces

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Box 4.1 The amount of free energy available from a reaction depends on the Gibbs free energies of formation of substrates and products, as given by the relationship (substrates). is the increment in free energy for the reaction under standard conditions. For biological systems the conventional standard conditions are 25°C and a pressure of 1 atm (101 kPa). In aqueous solutions, the standard condition of all solutes is 1 mol/kg activity, that of water is the pure liquid. Example: reductive dechlorination of 1,1,2,2-tetrachloroethane to 1,1,2-trichloroethane is given by the reaction:

The values for tetrachlorethane, and trichloroethane in the gaseous state the –85.48, 0, and -77.4 kJ/mol, those for and in the aqueous phase are 0 and -131.3 kJ/mol respectively. Thus

It is illuminating to notice that the values for 1,1,2,2-tetrachloroethane and 1,1,2trichloroethane in the aqueous phase are different from those in the gaseous phase: -88.9kJ/mol for 1,1,2,2-tetrachloroethane and –77.6 kJ/mol for 1,1,2-trichloroethane respectively. Consequently, the of the above reaction with 1,1,2,2-tetra-and 1,1,2trichloroethane in the aqueous phase at concentrations (activities) of 1M is: Thus, the outcome of calculations is affected by the choice of the standard conditions (hence the naut sign in Conversion of free energy values from the gaseous phase to the aqeous phase are made with Henry’s constant: where R is the universal gas constant (8.314 J/K·mol), T is the temperature (K) and H is Henry’s constant in At 25°C (298K), The Henry’s constants for 1, 1, 2, 2-tetra– and 1, 1, 2-trichloroethane are and respectively. Hence, the above difference is 5.7·log0.91 - 5.7·log0.25 = 3.2 kJ/mol, based on results between the Gibbs free energy calculations with standard conditions, gaseous vs. aqueous phase. Under environmentally relevant conditions, the concentrations of substrates and products are not 1 mol/kg and the partial pressures are not 1 atm. This is considered in values. For a hypothetical reaction values are calculated by using the mass equation:

Thus, if in the above example the hydrogen partial pressure is atm rather than 1 atm, the value becomes: in these calculations is a substrate, hence the value. This example underscores that at low substrate concentrations values can be significantly higher than values, while at low product concentrations values can be significantly lower. The degree to which the values are affected depend on the concentrations of the substrates and products involved.

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Box 4.1 (continued) Under biological conditions the hydrogen concentration is usually closer to rather than This is considered in values. As we have seen above, the value for is 0. The correction for pH=7 follows from the mass (Nernst) equation: Thus, in calculations, is corrected by -39.9 kJ for every mol of involved, with the sign of the correction depending on whether is a substrate or a product. The same logic can be applied to

Under standard conditions (1M concentrations, i.e., Thus, at It may be useful to point out that the sum of these two values (i.e., the values for and for each at pH=7) equals –237.2 kJ/mol, the value for This is in full agreement with the rule that the combined with the fact Gibbs free energy values are related to equilibrium constant K via the equation:

With this relationship in mind, it is instructive to point out that the difference in values in the earlier example on dechlorination of 1,1,2,2-tetrachloroethane with standard conditions set for either the gaseous of the liquid phase are in fact academic. Introduction of Henry’s constant into that example implied the assumption of full equilibrium. Under equilibrium conditions, the organisms catalyzing the reaction of interest cannot obtain different amounts of energy, depending on where and how the reactants are measured. These effects are implicitly compensated for when the equilibrium constant K is defined. The direct relationship between and K makes it easy to appreciate some of the salient characteristics of i.e., the information of whether a reaction yields energy (is exergonic) or costs energy (is endergonic). This intuitive logic also has the advantage of using or values rather than using redox potentials ( values) for evaluating and comparing the energetics ofredox reactions. Redox potentials are defined at pH =7, i.e., at a redox couple between and of 0. 41 V per 2 electrons. Thus the value of 0.41 V is the starting point for evaluations based on the redox scale, rather than the value of zero on scale. The conversion between the two scales is straightforward and given by the equation:

where is the number of electrons involved, F is the Faraday constant (96.48 kJ/V) and is the difference in potentials. As pointed out in student texts, e.g., Brock’s Biology of Microorganisms, the couple has a potential of –0.41 and the pair has a potential of +0.82, so the potential difference is 1.23 V. Because two electrons are involved this is equivalent to a free energy yield of 237.34 kJ/mol. In the opposite direction: based on the redox potential for the dehalogenation redox couple for the reaction:

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additional uncertainty into the numerical value of (15). In biochemistry, the conventional way to evaluate the tendency of a compound to donate or accept electrons is by means of the redox, or reduction, potential (expressed in volts). For the present chapter, I prefer the use of values. This convention is more intuitive since it gives direct information about whether a reaction is endergonic or exergonic, or which of a suite of reactions is the energetically most favorable one. Where necessary, values can be converted into redox potentials, or conversely with the formula (see Box 4.1, and 14, 30 or 37 for details).

3. REDUCTIVE DEHALOGENATION Values for the of reductive dechlorination (hydrogenolysis) of halogenated compounds range from -130 to -180 kJ/mol per halogen removed, corresponding to a redox potential of +260 to +480 mV (15, 17, 23, 24) (Figure 4.1). This makes halogenated compounds excellent electron acceptors, with redox potentials comparable to the redox couple of This is substantially higher than the values for sulfate and bicarbonate and suggests that dehalogenating organisms will out-compete sulfate reducers or methanogens for reducing equivalents when these are rate limiting. Based on these considerations, it follows that reductive dehalogenation will be rare, but not impossible, under aerobic conditions. This is indeed in agreement with practical experience (1, 38; Chapter 3).

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3.1. Effect of Position of the Substituent The amount of energy released upon hydrogenolysis depends on the nature and the position of the halogen substituent removed. Figure 4.2 illustrates this for halobenzoates. The amount of energy released increases in order with increasing atomic number of the halogen substituent: fluoro
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3.2. Dehalogenation Pathways Estimates of the Gibbs free energy of formation for chlorinated compounds have an associated uncertainty value of 2.5 kJ/mol for compounds in the gaseous phase (15). For compounds in the aqueous phase, there is additional uncertainty, depending on the reliability of the data for vapor pressure, solubility and, where applicable, for the pK value of the reactants. The total uncertainty is then about 5.5 kJ/mol. This accuracy makes it possible to formulate and test hypotheses about the logic behind the prevalence of certain dehalogenation pathways in the environment. When differences in Gibbs free energy yield indicate the potential for competing dechlorination steps to occur, in general, microorganisms will choose the pathway that yields the most energy. If the difference in free energy between alternative dechlorination reactions is more than 5.5 kJ/mol, analysis of the thermodynamic values can predict which pathway will occur. This assumes that the presumed differences in the change in Gibbs free energy can be effectively harnessed by the organisms involved. The rationale for this hypothesis is, as indicated in the introduction, the (presumed) analogy to the preferential use of inorganic electron acceptors in microbial ecosystems. Here, the electron acceptor with the highest redox potential, i.e., the electron acceptor giving the highest energy yield, is used preferentially, followed by those of decreasing redox potential, e.g., The hypothesis has been tested for the transformation of pentachlorobenzene. Dechlorination of pentachlorobenzene to 1,2,3,4tetrachlorobenzene, 1,2,3,5- tetrachlorobenzene, or 1,2,4,5- tetrachlorobenzene yields 161, 168, or 163 kJ/mol respectively. Thus, microorganisms would be expected to preferentially catalyze dechlorination of pentachlorobenzene to 1,2,3,5-

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tetrachlorobenzene, and this is indeed what is generally observed in the environment. Similarly, microorganisms predominantly convert 1,2,3,5-tetrachlorobenzene to 1,3,5trichlorobenzene, in a step that yields substantially more energy than conversion of 1,2,3,5- tetrachlorobenzene to either 1,2,3- trichlorobenzene or 1,2,4- trichlorobenzene (16). The hypothesis has now been evaluated in various environments, and has needed to be rejected on several occasions. One of the reasons for this may be that in many, if not all environments contaminated with halogenated compounds, these compounds do not occur alone, but rather in mixtures. Thus, the microorganisms that evolve or predominate to degrade this mixture may be influenced by a contaminant history that selects for other, perhaps less thermodynamically favorable dehalogenation pathways. This may be due to the steric preferences of enzymes already present that are adapted to cover novel compounds. Most anaerobic cultures show initial ortho-dechlorination of pentachlorophenol (PCP), which is at odds with the thermodynamic prediction made in Section 3.1, but agrees with a model where the bond charge of the carbon-halogen bond predicts the degradation pathway ofhalogenated aromatics (6). However, the fact that pathways can vary implies that it is apparently not possible to predict with absolute certainty the dechlorination patterns in anaerobic microbial habitats (14).

3.3. Accuracy of the Estimates In experiments conducted to determine the pathway of vitamin reductive dechlorination of aqueous 2,3,4,5,6-pentachlorobiphenyl, Woods et al. (43) found that theoretical product distributions based on free energies of formation agreed with the observed product distributions. The authors used the premise that the value Box 4.2

The regiospecificity of a reductive dechlorination reaction of a compound can be evaluated based upon data for that compound and for its potential dechlorination products. Theoretical product distributions are estimated by using the relationshipbetween and given in Box 1 . The standard free energy for the reaction under consideration is estimated from the reductive dechlorination half reaction At equilibrium:

At 25°C, this equation can be expressed as:

Writing down the dechlorination reaction and expressing substrates and products as a function ofK makes it easy to see, for example, that the distribution of dechlorination products for a system with three potential products can be estimated with the following expression:

where

is the fraction of the products represented by product

(after reference 43).

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of an equilibrium reaction is directly related to the equilibrium constant of that reaction, via the relationship where is the value for the reductive dechlorination half reaction. This equation can be used to calculate equilibrium constants for each parent compound and its potential products (see Box 4.2 for details). The product yields thus predicted are rather sensitive to small differences in Gibbs free energy values between potential products (Figure 4.4), and hence to the accuracy of their estimates. The accuracy of the data for polychlorinated biphenyls (PCBs) was estimated at better than 3.2 kJ/mol. Errors of ~3 kJ/mol would, however, have a significant impact on the the resulting product distribution, as in the experiments described by Woods et al. (43). Nevertheless, the prediction of which products would

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predominate was folly confirmed by the observed product distributions. In fact, the quantification of predicted and observed product distributions matched quite well (Figure 4.4B). From this good match, it appears that the accuracy of the data are indeed better than 3 kJ/mol. Given these results, it is tempting to conclude that the correlation between predicted product distributions and values can be used to fine tune the estimates of values. An example of such an approach is presented in Figure 4.4D, which shows the correlation between observed and measured dechlorination patterns for the dechlorination of tetrachlorobiphenyls to trichlorobiphenyls. Here, two small adjustments, of 1.5 to 2 kJ/mol, improve the correlation between the estimated and the observed product ratios for the tetrachlorobiphenyls, as analyzed by Woods et al. (43) Interestingly, these corrections were for both compounds with substituents at the 3- and 5-positions. This suggests that an additional interaction occurs with these substituents that was not (fully) accounted for in the original calculations. The vitamin system developed by Woods provides a powerful tool, not only to study both biological and abiological reductive dechlorination processes (8, 35, 42, 43), but also to refine existing estimates for values of halogenated organics. The above discussion centers on the agreement between model predictions and experimental results of short-term experiments, but results from long-term experiments are also convincing. Figure 4.5 depicts this agreement in a plot of observed versus predicted distributions. For example, the best fit of the data for terra-, tri-, and dichlorobiphenyls, but without the monochlorobiphenyl congeners, shows a slope of 1.0, with an value of 0.95. This was done because the measured data showed no 2- and 4-

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chlorobiphenyl, which is illogical unless one assumes that the reaction time had been too short to achieve a balanced formation of monochlorobiphenyls. Figure 4.5 illustrates that the data for the monochlorobiphenyls were true outliers, while the correlation between model predictions and experimental results for the other congeners was excellent.

4. DEHALOGENATION OF ALIPHATIC HYDROCARBONS AND PESTICIDES 4.1. Aliphatic compounds The values for the reductive dehalogenation of halogenated aliphatic compounds fall into the same range as those for aromatics. For both types of compounds,

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microorganisms are known to grow dependent on the hydrogenolysis of the carbonchlorine bond (see Chapter 3). The amount of energy released upon defluorination of chlorofluorocarbons is less than the amount available from dechlorination (Figure 4.6). Because C-C1 bonds generally provide a greater net energy gain than C-F bonds, the preferred transformation route of chlorofluorocarbons under anaerobic conditions is dechlorination rather than defluorination (22, 33, 36). The amount of energy available per dechlorination step is fairly constant. It is therefore not clear why reductive dechlorination of highly chlorinated aliphatics, such as tetrachloroethene, appears easier and proceeds faster than dechlorination of less chlorinated ones, such as vinyl chloride. This is a problem in many environments since vinyl chloride is a toxic intermediate that sometimes tends to accumulate as a “quasi-endproduct” of the anaerobic transformation of tetra- and trichloroethene.

4.2. Dehydrodehalogenation The reductive dehalogenation reaction discussed so far is essentially a hydrogenolysis. This, however, is not the only type of reductive dehalogenation reaction observed in nature. In another type of vicinal reduction, known as dihaloelimination or dehydrodehalogenation, two halogen substituents are removed from adjacent (vicinal) carbon atoms, while the aliphatic C-C bond is converted into an unsaturated C=C bond (Figure 4.7). Due to the electron configuration of the aromatic ring, dehydrodehalogenation does not occur on aromatic compounds. The important difference between dehydrodehalogenation and “normal” hydrogenolytic dehalogenation reactions is that, in the former two mol of halogens are removed per mol of while

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only one mol of halogen is removed in the hydrogenolytic type of reductive dehalogenation. This has consequences for the energy balance of the reaction. For example, approximately 160 kJ/mol is available per hydrogenolytic dechlorination step of chlorinated ethanes (i.e., one mol halogen per mol whereas this amount increases to 240 kJ/mol for dichloroelimination of chlorinated ethanes (Table 4.1). Under conditions where is limiting, i.e., in environments where concentrations are very low, or with competing populations, dihaloelimination is expected to prevail over hydrogenolysis. This line of reasoning suggests that organisms which use the dehydrodehalogenation route are more likely to be found under nitrate- or sulfatereducing conditions, rather than under methanogenic conditions. A practical implication ofthis hypothesized phenomenon is that it may be possible to steer the degradation route of halogenated ethanes away from the formation of vinyl chloride by adding an easily available source of reducing equivalents (12, 13). The observation that diluting a lactatefed methanogenic, 1,1,2,2-tetrachloroethane-transforming batch culture resulted in the formation of more 1,1,2-trichloroethane and less dichloroethene (5) is in line with the latter hypothesis, but is not conclusive evidence. This attractive hypothesis needs to be tested further both in cultures and in the environment.

4.3. Pesticides For many exotic compounds including most halogenated pesticides, values are not readily available in the literature. This is not always a problem; reductive dechlorination of all classes of halogenated compounds, aromatic as well as aliphatic, appears to be strongly exergonic. There is no reason why this should be different for other types of organic compounds, such as pesticides. In some cases, it may be of

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interest to have a more precise estimate of the of a particular dehalogenation reaction. In those cases, the group contribution method outlined by Mavrovouniotis (31, 32) offers a solution. Group contribution methods are widely used to estimate numerical values of thermodynamic properties of organic compounds. To estimate the property for a particular compound, one views the compound as composed of functional groups, and sums the values contributed by each group to the overall value of the property. Mavrovounitis did not present data for chloro substituents, but following his approach it has been possible to give values for aliphatic compounds which are in the order of 0-8 kcal/mol (17). A similar approach for aromatic compounds yields values of ~10 kcal/mol for chloro substituents, which is, however, only a first approximation. As discussed previously, the thermochemistry of a chloro substiruent is strongly affected by the presence of other substituents. Thus, if it is necessary to estimate the of an unknown compound, the best procedure is to use the group contribution method, with reference to the of a halogenated compound in which the halogen is surrounded by substituents similar to those present in the compound of interest (17).

5. HALOGENATED COMPOUNDS AS ELECTRON ACCEPTORS: HALORESPIRATION The amount of energy available from reductive dehalogenation is large enough to permit growth of microorganisms and they indeed are able to do so. This is expected from an energetic point of view, but fortuitous dehalogenation with no energetic benefit to the organism may also occur. For example, vitamin catalyzes reductive dehalogenation reactions. This cofactor is abundantly present in methanogens, and therefore fortuitous dehalogenation in methanogenic systems readily occurs (e.g., 40). The fact that microorganisms can harness large amounts of energy from dechlorination, raises a number of questions, including: how efficient are these organisms, what is their growth rate and growth yield, and what is their affinity for halogenated compounds and for It is hypothesized that these parameters are related to the amount of energy intrinsically available, and that the limiting (threshold) concentration of decreases with increasing energy availability. The same also follows for steady state concentrations in such systems. All of these considerations are based on dehalogenation via halorespiration (26). Two thermodynamic indicators are currently used to determine if dehalogenating microorganisms possess a halorespiratory physiology. Specifically, an evaluation of (i) the hydrogen threshold concentration, and (ii) the fraction of electrons consumed in electron acceptor reduction, can help to determine the metabolic mode used for dehalogenation. The hydrogen threshold can only be used for halorespirers growing with as a source of electrons, and cannot be used for organisms that grow on acetate. Furthermore, it has not yet been specified what the threshold is for microorganisms that catalyze reductive dechlorination reactions without obtaining energy for growth, such as methanogens catalyzing dechlorination of tetrachloroethene. Another method used to identify the activity of chlororespiring organisms is to determine The term is the fraction of electrons used for energy formation (i.e., the fraction of electrons released during oxidation of the electron donor that are directed toward reduction of the terminal electron acceptor). The fraction of electrons derived from the electron donor which is incorporated into biomass is the synthesis fraction

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with With chlororespirers, is in the range of 0.63 to 0.7 in pure cultures growing with halogenated compounds as electron acceptors. A minor problem here is that reductive dechlorination may continue in the stationary phase, which can affect the outcome of Thus, should be measured only during the growth phase of pure cultures. A problem encountered with mixed cultures is that a substantial portion of the electrons can flow to other electron acceptors. In such cases, it will be difficult to accurately assess the extent of electron flow to the presumed halorespirers. The values of chlororespiring cultures are considerably lower than those of sulfidogenic cultures (0.84 to 0.94) or methanogenic cultures (0.95), but higher than those of aerobic cultures (0.57) or denitrifying cultures (0.54). This comparison is based on the utilization of the same electron donor, i.e., acetate (28). Energetically, this makes sense since for chlororespirers is between the values for sulfate reduction and denitrification. Halorespirers often face a dilemma in that they generally function at the beginning of a food chain, although they require the reducing equivalents provided by others (further down the food chain). In a very simplified system, this unidirectional dependency may lead to metabolic bottlenecks. However, once reducing equivalents have been used by the dehalogenating organisms, the dehalogenated products are unlocked for consumption by successive organisms in the chain, assuming that dehalogenation is required before other metabolic processes can take place (14). This simplified picture may be typical for some microbial, e.g., methanogenic, communities, but in other, perhaps more complicated, multimember ecosystems this is less likely to occur (9). The available evidence suggests schemes whereby dehalogenators can harness more energy than just from the dechlorination reaction alone. Thus, 3-chlorobenzoate can theoretically be fermented to benzoate and acetate in a pure culture. Alternatively, a system containing a 3-chlorobenzoate dechlorinator plus a benzoate degrader could grow together without the need for a methanogen to remove the electrons from the degradation of benzoate, since dehalogenation (halorespiration) will help to pull the reaction by removing electrons. For example, methanogen- and sulfate- free medium inoculated with a co-culture of Desulfomonile tiedjei plus a benzoate degrader indeed showed growth as indicated by an increase in and depletion of 3-chlorobenzoate (Dolfing, unpublished).

5.1. Thermodynamic and Physiological Threshold Concentrations The for reductive dechlorination is higher than the for methanogenesis. Based on these thermodynamic considerations, it follows that chlorinated compounds are favored as electron acceptors by microorganisms under methanogenic conditions. In addition, halogenated compounds are generally also good electron acceptors under sulfate reducing conditions. For nitrate reducing conditions, the results are less clear-cut. The mechanism by which dechlorinating microorganisms outperform their competitors in mixed culture is thought to be due, at least in part, to prevailing concentrations. The energetics of the terminal electron accepting process controls the minimum concentration that can be consumed. This concentration is also known as the “threshold”. There is strong evidence in support of this, and recent reports indicate that it also applies to halorespiration (28). This is somewhat surprising, because the concept is based primarily on values, not on values. In other words, little

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attention has been paid to product concentrations and their effect on the actual values. Another flaw in many of the discussions of this concept is that the terms “steady state concentration” and “threshold” are used interchangeably, yet they are not synonymous. It is clear that halorespirers can operate at concentrations below the hydrogen uptake threshold for methanogens (28,44), but it should be kept in mind that in many dehalogenating ecosystems only a part of the electrons will be consumed by dehalogenating organisms. The partial pressure, therefore, cannot be used to evaluate whether dehalogenation occurs in a certain system. For example, in a methanogenic system, it is possible that roughly 90% of the electrons flow (as ) to the dehalogenators and only 10% to the methanogens. Here, because 10% actually flows to the methanogens, it follows that the hydrogen concentration is above the hydrogen threshold for methanogens. The reason that the dechlorinators do not consume all in this hypothetical example could be due to a number of factors, including a lack of halogenated substrate or not enough active biomass, i.e., compositional differences in the community strcuture. The of a reaction needs to be less than zero, in order to enable microorganisms to obtain energy for growth. Hence, it is possible to estimate a minimum substrate concentration below which reactions will not sustain growth. In anaerobic ecosystems, the limiting energy substrate is frequently dependent on the availability of reducing equivalents. All things being equal, and if a number of different types of electron acceptor is available, hydrogen concentrations generally will dictate which electron acceptor is used by microorganisms to sustain growth. Each electron acceptor has its own optimal range of hydrogen concentrations. Below a particular range, the microorganisms using the corresponding electron acceptor cannot obtain energy; above it another electron acceptor will divert part of the electron flow. There are indications that due to physiological reasons, the threshold is not when but when is approximately -20 kJ/mol (34). It appears that microorganisms need a certain amount of electron pressure before they can actually start harnessing energy. This small amount does not affect the idea of the minimum threshold. It only “moves the scale” by ~20 kJ/mol. However, these “first” 20 kJ/mol of a reaction are not lost. Microorganisms working with a reaction that yields 30 kJ/mol can harness all of this energy (although not with 100% efficiency), and not just the 10 kJ/mol above the threshold of 20 kJ/mol. Recent evidence however indicates that anaerobic microbial metabolism can proceed close to thermodynamic limits, even though the energy remaining when metabolism stops is highly variable (26). Strict application of the threshold concept yields realistic values for sulfate reduction and methanogenesis, but not for dechlorination. For example, thermodynamic considerations indicate a threshold (equilibrium) concentration of roughly for the dechlorination of chlorinated ethenes. In practice, this is unrealistic because microorganisms cannot metabolize at meaningful rates at such low concentrations. Even the premise that organisms need at least one third of an ATP, i.e., ~18 kJ/mol does not help; this would result in an ~1000 fold increase in the threshold for dechlorination, which is still unrealistically low. Thus, the threshold as well as its concentration in dechlorinating ecosystems appears to be under kinetic rather than thermodynamic control. Recently, a very useful extension of the threshold concept has been developed. It has

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been argued (20) that product inhibition can be quantified through a simple thermodynamic function, where The term is the amount of energy available from substrate conversion. The term is the value below which the organisms cannot obtain enough energy from the substrate for the reaction to be beneficial. Note that the values for are between 1 and 0. The factor is used as a correction on the concentrations of substrates and products in MichaelisMenten-like kinetics. In many, if not all, acetogenic degradation reactions (7), these values, and their factors are strongly dependent on the concentrations of acetate and The factor of dechlorination reactions will also depend on the concentration of although in this case is a substrate rather than a product. Recently, Liu et al. (27), in a study on bacterial reduction of goethite, reported that the observed match between theoretical (model) and experimental results hinged on the inclusion of the factor in their model.

5.2. Competition with Other Electron Acceptors The effect of substrate and product concentration on the of a reaction (see Box 1 for details) may also be impacted by relatively high local concentrations of alternative electron acceptors (other than chlorinated compounds), such as nitrate and sulfate, so as to affect their competitiveness. Relatively high concentrations of a particular electron acceptor can make these reactions thermodynamically more favorable, especially because the concentration of both substrate and product affects High concentrations per se do not necessarily result in a high For example, during nitrate reduction to nitrite, the ratio of nitrate to nitrite determines the similarly for sulfate reduction, it is the ratio of sulfate to sulfide. Therefore, the of a reaction is essentially unaffected, as long as the concentrations of nitrate and nitrite, or sulfate and sulfide, are within the same concentration range. Whereas with nitrate this is usually the case, the situation can vary with sulfate. When sulfate is in the millimolar range, as it frequently is, and sulfide is in the micromolar or nanomolar range, the will be 3 times (sulfide in the micromolar range), or even 6 times (sulfide in the nanomolar range) more beneficial than under standard conditions, i.e., when equimolar concentrations of both oxidized and reduced species of electron acceptor are present. For dechlorination, the concentration of chlorinated substrates and lesser-chlorinated products is also an issue. One of the characteristics of chlorinated compounds is that they are rather hydrophobic and have a tendency to sorb to surfaces, such as sediment (14). Actual concentrations in aqueous solution can thus be quite low. The hydrophobicity of chlorinated compounds decreases with decreasing degree of halogenation. The lesser-chlorinated products have a smaller tendency to sorb to the sediment matrix. Thus, if a significant part of the parent compound has been dechlorinated (to lesser-chlorinated products), it is likely that the concentration of the dechlorinated product in the pore-water of the sediment becomes considerably greater than that of the parent compound. Conceptually, there is therefore reason to expect a thermodynamic auto-inhibition of dechlorination in sediments. Quantitatively, however, this is unlikely, since the differences in sorption characteristics between chlorinated parent compound and the lesser-chlorinated, or even dechlorinated, daughter product, are not that great. Even an 100-fold concentration difference has an effect of only 2 x

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5.7=11.4 kJ/mol, which is less than 10% of the values of > 130 kJ/mol per reductive dechlorination under standard conditions, or 130 - 5·5.7 = ~110 kJ/mol for dechlorination at hydrogen partial pressures of 1 Pa rather than 1 atm (100 kPa). Because reductive dechlorination is a step by step process, such a hypothetical “desorption induced thermodynamic auto-inhibition” does not also play a significant role in the dechlorination of highly chlorinated compounds, such as PCBs.

6. AEROBIC DEGRADATION OF HALOGENATED COMPOUNDS The presence of a halogen substituent makes a compound more oxidized than its non-halogenated analog. The thermochemical characteristics of halogens are such that halogenation decreases the amount of energy available to organisms which grow at the expense of these compounds under aerobic conditions, i.e., with as electron acceptor. The amount of energy available from mineralization of a halogenated compound with serving as final electron acceptor decreases with the degree of halogenation of the compound, while the amount of energy available to methanogenic consortia increases with the degree of halogenation of a compound (Figure 4.8). Interestingly, the amount of energy available from the aerobic mineralization of highly halogenated compounds, such as tri- and tetrachloroethene, is theoretically more than sufficient to sustain growth. Thermodynamically, there is no reason why aerobic growth with these compounds as electron donor has never been observed. With trichloroethene, only cometabolic degradation has been observed while tetrachloroethene appears to be completely recalcitrant under aerobic conditions (19).

6.1. Hydrolytic versus Reductive Dehalogenation Hydrolytic dehalogenation is an exergonic reaction, but is energetically less favorable than reductive dehalogenation. For a compound such as 2-chlorobenzoate, reductive dechlorination with as electron donor source has a of-145.5 kJ/mol, whereas hydrolytic dehalogenation of 2-chlorobenzoate to 2-hydroxybenzoate yields only 78.3 kJ/mol. Likewise, under standard conditions (pH=7; present at a partial pressure of 100 kPa), reductive dehalogenation of 2-chlorobenzoate is significantly (67.1 kJ/mol) more favorable than hydrolytic dehalogenation. This also holds for environmentally more realistic concentrations of 1 Pa. Under these conditions, the for reductive dechlorination becomes 5 times 5.7, or 28.5 kJ/mol higher, providing an energy yield of –145.4 + 28.5, or -116.9 kJ/mol. Similar calculations for more highly chlorinated compounds cannot be made currently due to a lack of thermodynamic data, in particular the free energy of formation for most chlorinated benzoates and chlorohydroxybenzoates. For a hydrolytic dechlorination of chlorobenzenes, such a calculation can be made and the outcome (presented in Figure 4.9) indicates that reductive dechlorination of chlorinated benzenes yields approximately 73 kJ/mol more Gibbs free energy than hydrolytic dehalogenation. The above calculations indicate that under aerobic conditions, where reducing equivalents are expensive, and to a lesser extent under nitrate reducing conditions, microorganisms may tend to use hydrolytic rather than reductive dehalogenation. It is

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not necessarily because this dechlorination mechanism yields the most energy, but simply because competition for electrons is fierce and alternative dechlorination mechanisms are feasible and not overly expensive. Based on this information, onemust be cautious in assuming that the first step in the anaerobic degradation pathway of halogenated compounds is always a reductive dechlorination step (8).

7. ENVIRONMENTAL CONSTRAINTS The focus of this chapter is on the thermodynamic range in which microorganisms can obtain energy for growth from halogenated substrates. It should be emphasized, however, that other factors will also influence the energetics, and ultimately the in situ reaction. Many halogenated compounds, especially the more chlorinated ones, are rather hydrophobic and will strongly sorb to the environmental matrix, e.g., soil or sediment (14). Actual concentrations that microorganisms encounter in the aqueous phase will often be orders of magnitude lower than the total concentration. Under these conditions, sorption/desorption processes, diffusion, and low concentrations will strongly affect bioavailability of halogenated substrates, and therefore also the energetics of the reactions. In this context, it is prudent to stress that dechlorination will increase the

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bioavailability of less-chlorinated daughter products. The thermodynamic calculations made here are equilibrium calculations and they generally assume equimolar concentrations. However, at extremely low concentrations it is likely that sufficient fluxes cannot be maintained, not so much because insufficient energy is derived from the molecule, but rather because the diffusion rate becomes limiting.

7.1. Reductive Debromination It is well established that the addition of specific PCB congeners will stimulate, i.e., “prime”, dechlorination in PCB-contaminated sediments (41; see Chapter 17). Following a similar strategy, and combined with the knowledge that reductive dehalogenation of brominated benzoates yields slightly more energy than reductive dehalogenation of their corresponding chlorinated analogs, analog enrichment techniques with polybrominated biphenyls (PBBs), have been employed to stimulate PCB biodegradation in PCB-contaminated sediments. These attempts have been successful (2). An evaluation of the energetics of reductive debromination versus reductive dechlorination of halogenated biphenyls has helped to explain the observed phenomenon. By estimating the Gibbs free energy of formation data for PBBs with the same methods previously used to estimate values for PCBs (Dolfing and Harrison, unpublished), it was possible to estimate the values for reductive debromination for a series of brominated biphenyls. The outcome, as presented in Figure 4.10, shows that reductive debromination of PBBs is energetically slightly more favorable (~10 kJ/mol) than reductive dechlorination of the corresponding PCBs. However, no

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differences have been observed for dehalogenation at the ortho position of compounds with no halogens at any of the other ortho positions.

8. FERMENTATION Degradation of organic compounds is conceptually a series of dehydrogenations and decarboxylations. For example, an organic compound such as ethene undergoes mineralization to and in an endergonic reaction that only becomes exergonic at low hydrogen concentrations. The presence ofchloro substituents changes this picture; mineralization of chlorinated ethenes to and is an exergonic reaction, even under the standard conditions of 1 atm The energy content of the chloro substituent of chlorinated aliphatics is such that mineralization becomes more favorable with increasing degree ofchlorination (13). The formation of implies that the energetics of these hypothetical mineralization reactions depends strongly on the partial pressure of Since more is formed from the complete mineralization of the lesser chlorinated congeners, these compounds would be expected to be more sensitive to changes in concentration. In addition, sensitivity to changes in concentration during biodegradation of the lesser chlorinated congeners is less than that of more chlorinated congeners. It is tempting to speculate that organisms may exist that are able to obtain energy for growth from such mineralization reactions. The most likely habitat for these organisms would be in environments where the partial pressure is very low, i.e., environments where iron, nitrate or sulfate serves as final electron acceptor. Furthermore, the large amount of energy available to microorganisms that can catalyze these reactions makes this metabolic (dechlorination) strategy especially attractive for highly chlorinated compounds that are generally present at very low concentrations due to of their low solubility in water. A low aqueous solubility of may impede the flow of such compounds towards the microorganisms that degrade them. Conversely, hydrophobic cells may actually sorb compounds, thereby increasing ambient concentrations. From an energetic point of view, it is rather surprising that polychlorinated compounds are mostly degraded by more than one organism in what resembles a “food chain”, but what in fact more aptly can be referred to as a “waste chain” (18). It is not one organism being eaten by another, as in the classical food chain, but rather the "waste" of one organism being consumed by the next organism in the chain. It is tempting to hypothesize that the wellknown division of labor, and hence of energy profits, between dehalogenators, acetogens, and methanogens suggests fairly evolved system, whereby each organism has its own specialized role in the ultimate degradation of a complex, and often recalcitrant, substrate. The aforementioned sequential dehalogenation, consisting of a series of metabolic steps carried out by different organisms, can be viewed as a sign of further specialization towards a climax structure. On the other hand, it seems equally valid to argue that it is inefficient to have two dehalogenators do what one could probably do as well alone. The availability of microorganisms that are able to carry out only the initial two dehalogenation steps of tetrachloroethylene dechlorination, and of microorganisms that can dehalogenate tetrachloroethylene completely to ethene, suggests that it is now possible to study these questions experimentally.

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9. ENVIRONMENTAL APPLICATIONS Thermodynamic logic may help to design in situ bioremediation schemes. For example, Lorah and Olsen (29) have described a site where transformation of a chlorinated ethane proceeds via dehydrodehalogenation, giving rise to vinyl chloride which can accumulate as a carcinogenic intermediate. Thermodynamics would predict that addition of an easily available source of reducing equivalents would favor a route via hydrogenolysis (12). The addition of reducing equivalents, i.e., a secondary substrate, is problematic in that it may support competition between dehalogenators and other organisms for a commonly metabolized source of energy. Based on thermodynamics, dehalogenator populations will outcompete methanogens for (28). However, methanogens are relatively abundant in many anaerobic environments, and although dechlorinators may have a lower uptake threshold, their numbers are not sufficient to reduce concentrations to levels below the threshold for uptake by methanogens. So, whereas the dechlorinators are thermodynamically more capable of utilizing under nonlimiting conditions, a considerable fraction of the reducing equivalents will flow to methane (44). In order to circumvent this possibility for practical applications, it has been shown in the laboratory that yeast extract, rather than acetate, can serve as a good source of reducing equivalents for dehalogenation (11,45). Another viable strategy may be the addition of a source of reducing equivalents that is degraded rather slowly. Propionate has been suggested as such a source, since propionate degradation in methanogenic ecosystems is only sustainable at low concentrations (21).

10. FINAL REMARKS As we gain a better understanding of the role of thermodynamic principles in biological systems, it becomes clear that thermodynamics helps to rationalize biogeochemical processes, especially the use of inorganic electron acceptors (46). For organic compounds, the applicability is less clear-cut. In the present chapter, it is argued that chlorinated compounds are excellent electron acceptors, possessing redox potentials similar to the nitrate/nitrite couple. To what extent microorganisms can use these compounds for the generation of metabolic energy remains to be seen. Growth yields of halorespirers are considerable, but not as high as those of organisms that use nitrate as an electron acceptor. Apparently, the metabolic efficiency of the dehalogenating organisms leaves room for improvement. Although there is reason to believe that some halogenated compounds are synthesized in nature (10), many of these compounds are considered to be xenobiotic. They often occur at relatively high concentrations in the environment, especially in the last century with increased industrialization and pollution. Hence, it seems reasonable to assume that the organisms which metabolize them still have not evolved optimally towards their substrate use. This may explain why dechlorinating organisms do not always behave in a way that would seem logical from a thermodynamic point of view, and suggests that the organisms have not yet evolved fully, since the appropriate selection pressure has not yet been applied long enough. Another point is the presence of other compounds in the environment, and the genetic background of the bacteria involved in their degradation. When xenobiotic compounds

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are introduced, potential degraders do not start from scratch, but adapt existing enzymes and pathways. This may explain why theoretical predictions are frequently not confirmed by environmental observations. A further complicating factor may be the availability of the compound in the environment. Many halogenated compounds strongly sorb to solid surfaces, e.g., the soil matrix, which makes them poorly available for biodegradation. Low concentrations and low diffusion rates may preclude the development of optimally efficient degradation routes. For some chlorinated aliphatic compounds, dehydrodehalogenation is energetically more efficient than hydrogenolysis and would be expected to dominate in environments where reducing equivalents are expensive. Especially in environments where the supply of reducing equivalents fluctuates, it would be beneficial for organisms to ferment halogenated compounds. The carbon-halogen bond can then serve as internal electron acceptor, while electrons are generated from the dehydrogenation of the dehalogenated product. This is essentially what happens is in a methanogenic consortia, but could theoretically also be performed by a single organism. Obviously, this line of reasoning is very speculative but there is one indication that it may not be too far-fetched, as Bradley and Chapelle have recently observed that chloroethene (vinyl chloride) can be fermented to acetate and ethene (3). REFERENCES 1.

2.

3.

4.

5.

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Apajalahti JHA & Salkinoja-Salonen MS (1987) Complete dechlorination of tetrachloroh y d r o q u i n o n e by cell extracts of pentachlorophenol-induced Rhodococcus chlorophenolicus. J. Bacteriol. 169:5125-5130 Bedard DL, Van Dort H & DeWeerd KA (1998) Brominated biphenyls prime extensive microbial reductive dehalogenation of Aroclor 1260 in Housatonic River sediment. Appl. Environ. Microbiol. 64:1786-1795 Bradley PM & Chapelle FH (1999) Methane as a product of chloroethene biodegradation under methanogenic conditions. Environ. Sci. Technol. 33:653-656 Bryant MP, Wolin EA, Wolin MJ & Wolfe RS (1967) Methanobacillus omelianskii, a symbiotic association of two species of bacteria. Arch. Microbiol. 59:20-31 Chen C, Puhakka JA & Ferguson JF (1996) Transformations of 1,1,2,2-tetrachloroethane under methanogenic conditions. Environ. Sci. Technol. 30:542-547 Cozza CL & Woods SL (1992) Reductive dehalogenation pathways of substituted benzenes: A correlation with electronic properties. Biodegradation 2:265-278 Dolfing J (1988) Acetogenesis. In: Zehnder AJB (Ed) Biology of Anaerobic Microorganisms (pp 417-469). John Wiley & Sons, Inc, New York Dolfing J (1995) Regiospecificity of chlorophenol reductive dechlorination by vitamin

Appl. Environ. Microbiol. 61:2450 Dolfing J (1996) degradation of monochlorinated and nonchlorinated aromatic compounds under iron-reducing conditions. Appl. Environ. Microbiol. 62:3554-3556 10. Dolfing J (1998) Halogenation of aromatic compounds: Thermodynamic, mechanistic and ecological aspects. FEMS Microbiol. Ecol. 167:271-274 11. Dolfing J (1999) Comment on "Competition for hydrogen within a chlorinated solvent dehalogenating anaerobic mixed culture". Environ. Sci. Technol. 33:2127 12. Dolfing J (1999) Comment on "Degradation of 1,1,2,2-tetrachloroethane in a freshwater tidal wetland: Field and laboratory evidence". Environ. Sci. Technol. 33:2680 13. Dolfing J (2000) Energetics of anaerobic degradation pathways of chlorinated aliphatic compounds. Microb. Ecol. 40:2-7 14. Dolfing J & Beurskens JEM (1995) The microbial logic and environmental significance of reductive dehalogenation. Adv. Microb. Ecol. 14:143-206 15. Dolfing J & Harrison BK (1992) The Gibbs free energy of formation of halogenated aromatic compounds and their potential role as electron acceptors in anaerobic environments. Environ. Sci. Technol. 26:2213-2218 16. Dolfing J & Harrison BK (1993) Redox and reduction potentials as parameters to predict the 9.

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Brock Biology of Microorganisms, Eight degradation pathway of chlorinated benzenes in anaerobic environments. FEMS Microbiol Ecol Edition. Prentice Hall International, Inc. 13:23-30 31. Mavrovouniotis ML (1990) Group contributions 17. Dolfing J & Janssen DB (1994) Estimates of for estimating standard Gibbs energies of Gibbs free energies of formation of chlorinated formation of biochemical compounds in aqueous solution. Biotechnol. Bioeng. 36:1070-1082 aliphatic compounds. Biodegradation 5:21-28 18. Dolfing J & Prins RA (1996) Methanogenic 32. Mavrovouniotis ML (1991) Estimation of standard Gibbs energy changes in 'food chains'. ASM News 62:117-118 biotransformations. J. Biol. Chem. 19. Dolfing J, van den Wijngaard AJ & Janssen DB 266:14440-14445 (1993) Microbiological aspects of the removal of chlorinated hydrocarbons from air. 33. Oremland RS, Lonergan DJ, Culbertson CW & Lovley DR (1996) Microbial degradation of Biodegradation 4:261-282 hydrochlorofluorocarbons and 20. Fennell DE & Gossett JM (1998) Modeling the in soils and sediments. Appl. production of and competition for hydrogen in a Environ. Microbiol. 62:1818-1821 dechlorinating culture. Environ. Sci. Technol. 34. Schink B (1997) Energetics of syntrophic 32:2450-2460 cooperation in methanogenic degradation. 21. Fennell DE, Gossett JM & Zinder SH (1997) Microbiol. Mol. Biol. Rev. 61:262-280 Comparison of butyric acid, ethanol, lactic acid, and propionic acid as hydrogen donors for the 35. Smith MH & Woods SL (1994) Regiospecificity of chlorophenol reductive dechlorination by reductive dechlorination of tetrachloroethene. vitamin Appl. Environ. Microbiol. Environ. Sci. Technol. 31:918-926 60:4107-4110 22. Hageman KJ, Istok JD, Field JA, Buscheck TE & Semprini L (2001) In situ anaerobic 36. Sonier DN, Duran NL & Smith GB (1994) Dechlorination of trichlorofluoromethane transformation of trichlorofluoroethene in (CFC-11) by sulfate-reducing bacteria from an trichloroethene contaminated groundwater aquifer contaminated with halogenated aliphatic Environ. Sci. Technol. 35:1729-1735 compounds. Appl. Environ. Microbiol. 23. Holmes DA, Harrison BK & Dolfing J (1993) Estimation of Gibbs free energies of formation 60:4567-4572 of polychlorinated biphenyls. Environ. Sci. 37. Thauer RK, Jungermann K & Decker K (1977) Energy conservation in chemotrophic anaerobic Technol. 27:725-731 bacteria. Bacteriol. Rev. 41:100-180 24. Huang C-L, Harrison BK, Madura J & Dolfing J (1996) Thermodynamic prediction of 38. Van den Tweel WJJ, Kok J & de Bont JAM (1987) Reductive dechlorination of dehalogenation pathways for PCDDs. Environ. 2,4-dichlorobenzoate to 4-chlorobenzoate and Toxicol. Chem. 15:824-836 hydrolytic dehalogenation of 4-chloro-, 25. Ianotti EL, Kafkewitz P, Wolin MJ & Bryant 4-bromo-, and 4-iodobenzoate by Alcaligenes MP (1973) Glucose fermentation products of denitrificans NTB-1. Appl. Environ. Microbiol. Ruminococcus albus grown in continuous culture with Vibrio succinogenes: Changes 53:810-815 caused by interspecies transfer of hydrogen. J. 39. van Eekert MHA (1999) Transformation of Bacteriol. 114:1231-1240 chlorinated compounds by methanogenic granular sludge. PhD Thesis, Wageningen 26. Jackson BE & McInerney MJ (2002) Anaerobic Agricultural University, Wageningen, The microbial metabolism can proceed close to Netherlands thermodynamic limits. Nature 415:454-456 27. Liu C, Kota S, Zachara JM, Fredrickson JK & 40. van Eekert MHA, Stams AJM, Field JA & Schraa G (1999) Gratuitous dechlorination of Brinkman CK (2001) Kinetic analysis of the chloroethanes by methanogenic granular sludge. bacterial reduction of goethite. Environ. Sci. Appl Microbiol Biotechnol 51:46-52 Technol. 35:2482-2490 28. Löffler FE, Tiedje JM & Sanford RA (1999) 41. Van Dort HM, Smullen LA, May RJ & Bedard DL (1997) Priming microbial metaFraction of electrons consumed in electron dechlorination of polychlorinated biphenyls that acceptor reduction and hydrogen thresholds as have persisted in Housatonic River sediments indicators of halorespiratory physiology. Appl. for decades. Environ. Sci. Technol. Environ. Microbiol. 64:4049-4056 29. Lorah MM & Olsen LD (1999) Degradation of 31:3300-3307 1,1,2,2-tetrachloroethane in a tidal freshwater 42. Woods SL&Smith MH (1995) Regiospecificity of chlorophenol reductive dechlorination by wetland: Field and laboratory evidence. Environ. vitamin Appl. Environ. Microbiol. Sci. Technol. 33:227-234 61:2450-2451 30. Madigan MT, Martinko JM & Parker J (1997)

114 43. Woods SL, Trobaugh DJ & Carter KJ (1999) Polychlorinated biphenyl reductive dechlorination by vitamin Thermodynamics and regiospecificity. Environ. Sci. Technol. 33:857-863 44. Yang Y & McCarty PL (1998) Competition for hydrogen within a chlorinated solvent dehalogenating anaerobic mixed culture. Environ. Sci. Technol. 32:3591-3597

DOLFING 45. Yang Y & McCarty PL (1999) Response to "Comment on 'Competition for hydrogen within a chlorinated solvent dehalogenating anaerobic mixed culture'". Environ. Sci. Technol. 33:2128 46. Zehnder AJB & Stumm W (1988) Geochemistry and biogeochemistry of anaerobic habitats. In: Zehnder AJB (Ed) Biology of Anaerobic Microorganisms (pp 1 -38). John Wiley & Sons, Inc, New York

Chapter 5 DEHALOGENATION BY ANAEROBIC BACTERIA CHRISTOF HOLLIGER 1, CHRISTOPHE REGEARD1, AND GABRIELE DIEKERT 2 1

Laboratory for Environmental Biotechnology, Swiss Federal Institute of Technology (EPFL), Lausanne, Switzerland 2 Lehrstuhl für Angewandte und Ökologische Mikrobiologie, Friedrich-Schiller-Universität, Jena, Germany

1. INTRODUCTION Degradation of halogenated compounds under anoxic conditions was first studied in the 1950s and 1960s when the fate of halogenated pesticides in agricultural soils was investigated (2, 54, 58, 70). Only 15 to 20 years later, the anaerobic degradation of halogenated compounds has become a matter of special concern due to the almost ubiquitous presence of chlorinated compounds as pollutants of groundwater, soils, and sediments. Highly chlorinated compounds that resist aerobic degradation, such as tetrachloroethene (PCE) and polychlorinated biphenyls (PCBs), are transformed under anoxic conditions by reductive reactions (8, 119, 121). Such reductive dehalogenations may be a threat to living organisms if compounds more toxic than the parent compound are formed, such as carcinogenic vinyl chloride (VC) from PCE (157) and 2,3,7,8,tetrachlorodibenzo-p-dioxins from polychlorinated dibenzo-p-dioxins (4). However, reductive dehalogenation reactions also have a large potential for application in treatment processes for materials contaminated with halogenated compounds, such as industrial wastes, soils, sediments, waste- and groundwater. Aerobically persistent polyhalogenated compounds can be transformed by anaerobic mechanisms into harmless compounds, or into halogenated compounds that are further degradable by aerobic microorganisms (59). This potential has already found its application in different bioremediation projects (160). Reductive dehalogenation under anoxic conditions has been reported for many different compounds, such as chlorobenzoates, chlorophenols, chlorobenzenes, chloromethanes, chloroethanes, and chloroethenes. Although abiotic processes might be involved in some of the reductive transformations observed in environmental samples, recent evidence has been presented to show that the majority of these reactions are biologically catalyzed. Several reviews are available that summarize the knowledge of reductive dehalogenations catalyzed by mixed and pure cultures (39, 61, 78, 105). Long Dehalogenation: Microbial Processes and Environmental Applications, pages 115-157 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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acclimation periods, substrate specificity, high dehalogenation rates, and the possibility to enrich for the dehalogenation activity by subcultivation in media containing a selective organic or inorganic electron donor indicate that many of the reductive dehalogenations in the environment are catalyzed by specific bacteria (60). Despite the strong evidence for the involvement of biological processes in reductive dehalogenations, a biological activity can be unambiguously assigned only to a few of the reductive dehalogenation reactions observed in environmental samples. The availability of pure cultures catalyzing reductive dehalogenations allows for more detailed investigations on the metabolic function of these types of reactions. In addition to reductive dehalogenation, mineralization of halogenated compounds to carbon dioxide and chloride has also been observed in environmental samples under anoxic conditions. For chlorinated aromatic compounds such as chlorobenzoates and chlorophenols, mineralization only occurs after complete reductive dechlorination to benzoate and phenol (97, 122, 142). The reductive dehalogenation reactions appear to be catalyzed by specific bacteria, rather than by the benzoate- and phenol-oxidizing organisms (34). Carbon dioxide production has been reported for and chlorinated compounds, such as carbon tetrachloride (8, 9), chloroform (8, 9), 1,2-dichloroethane (8), 1,1,1-trichloroethane (8, 9, 158), tetrachloro- (157) and dichloro- ethene (12), and VC (11, 12, 16). The latter two seem to be used directly as a carbon and energy source by anaerobic bacteria, without initial reductive dehalogenations. Recently, two strictly anaerobic bacteria have been isolated that are able to grow on chlorinated methanes as sole energy sources (87, 146). In other chapters of this book, the biodiversity of anaerobic dehalogenating bacteria is discussed in detail (see Chapters 2 and 3), and some of the physiological principles are introduced (see Chapters 3 and 4). Here, the different physiological roles that halogenated compounds can have in the metabolism of anaerobic bacteria is described. Also, a detailed discussion of the physiology of the anaerobic bacteria isolated in pure culture and their ability to degrade halogenated compounds, as well as the biochemistry and the genetics of the implicated dehalogenation reactions is presented.

2. METABOLIC FUNCTION OF THE DEHALOGENATION REACTION Halogenated compounds can serve in three different metabolic functions in anaerobic bacteria: i) as carbon or energy source or both, ii) as substrate for cometabolic activity, and iii) as terminal electron acceptor in an anaerobic respiration process. All of these metabolic functions have been identified in different anaerobic bacteria. Acetobacterium dehalogenans strain MC (146) and Dehalobacterium formicoaceticum strain DMC (87) can utilize the compounds methyl chloride or dichloromethane as carbon and energy source, respectively. Both bacteria are homoacetogenic organisms, and are the only pure cultures of anaerobic bacteria which so far have been isolated that use an aliphatic halogenated compound as a carbon source for growth. The physiology and biochemistry of the degradation of these two chloromethanes will be discussed in Section 3 of this chapter. Indications for the anaerobic utilization of the compounds dichloroethene (DCE) and VC as carbon and energy source have been observed in microcosms with compounds (13). For example, formation of was found to occur in different redox environments, such

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as under Mn(IV)-, Fe(III)-, and (methanogenic) conditions (14). acetate formation in microcosms under methanogenic conditions, in the presence of the specific methanogenesis inhibitor, 2-bromoethanesulfonate (BES), indicates that oxidative acetogenesis may be the initial step in the net oxidation of VC to (15). Only under Mn(IV)-reducing conditions does DCE appear to be used directly without initial reductive dechlorination to VC, as was observed under Fe(III)-, and reducing conditions (17). With one exception, aromatic chlorinated compounds have been demonstrated to serve as carbon and energy source, but only after complete dechlorination (34, 97). For the degradation of 3-chlorobenzoate by a mixed culture under methanogenic conditions, it has been shown that hydrogen produced by benzoateoxidizing bacteria serves as electron donor for 3-chlorobenzoate-dechlorination (34). The only reported exception is a recently isolated denitrifying bacterium that utilizes 3chlorobenzoate as carbon and energy source. In this case, degradation activity was inducible and degradation was specific to the position of the halogen substituent (55). In addition to microcosm studies, work with pure cultures of anaerobic bacteria has shown that a broad range of bacteria are capable of dehalogenation. Reductive dehalogenation reactions are a common mechanism catalyzed by these bacteria, but often occur at very low rates. Other transformation reactions, however, have been observed leading to more oxidized products. An overview of the work with pure cultures of fermentative bacteria, iron- and sulfate-reducers, homoacetogens, methanogens, and others is presented in Section 4 of this chapter. The redox potential of the redox couples involved in dechlorination, R-C1/R-H, lies between +250 and +600 mV and therefore, chlorinated compounds are thermodynamically favored as electron acceptors. Based on these considerations, bacteria have been enriched and isolated in pure culture that are capable of coupling the oxidation of an organic or inorganic electron donor, to the reduction of a chlorinated compound in an anaerobic respiration process. The increasing number of such dehalogenating bacteria isolated during the past decade, as well as numerous detailed studies of their metabolism, biochemistry, and genetics, has helped to establish the general principles of this anaerobic respiration process (see Section 5).

3. DECHLORINATION OF CHLORINATED METHANES VIA CORRINOIDDEPENDENT ALKYL TRANSFER Several aerobic and anaerobic bacteria have been reported to metabolize chlorinated methanes (for a review, see reference 79). In most of the studies performed with anaerobes, dechlorination of these compounds proved to be a very slow reductive dehalogenation carried out by cofactors, such as vitamin derivatives or coenzyme of methanogenic bacteria (36, 74, 75, 139). In the last decade, two strictly anaerobic bacteria have been isolated that are able to grow on chlorinated methanes as sole energy sources. The substrates utilized were methyl chloride for Acetobacterium dehalogenans and dichloromethane for Dehalobacterium formicoaceticum. Both organisms belong to the homoacetogenic bacteria, which share the ability to form acetate as a major end product from compounds such as or methyl moieties.

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3.1. Anaerobic Dechlorination of Methyl Chloride Acetobacterium dehalogenans has been isolated with methyl chloride as energy source (146). The organism converts methyl chloride to acetate in its energy metabolism according to the following equation:

Growth occurs with a doubling time of about 30 h with 2-3% methyl chloride in the gas phase. In addition to methyl chloride, a variety of other substrates including, CO, sugars, and methoxylated aromatic compounds are also utilized by A. dehalogenans. The ability to metabolize methyl chloride appears to be inducible; do not dechlorinate the substrate. For a review on the natural origin of methyl chloride and on its global cycle and bioconversion, see (56). According to studies performed with crude extracts of the organism (95), the initial step of methyl chloride dehalogenation is the transfer of the methyl group to tetrahydrofolate. Hence, the biotransformation of chlorinated methanes as growth substrates occurs via a methyl transfer reaction, rather than a reductive dechlorination. Methyl tetrahydrofolate participates in divergent pathways: it is oxidized to or it combines with coenzyme A and carbon monoxide to form acetyl-CoA. Carbon monoxide is formed via reduction of with the reducing equivalents derived from methyl group oxidation. Acetyl-CoA is then either converted to acetate during energy metabolism, or to cell carbon in anabolism. A summary of the pathway of methyl chloride conversion to acetate is depicted in Figure 5.1. The methyl transfer from methyl chloride to tetrahydrofolate can be assayed spectrophotometrically in crude extracts by a newly developed coupled test, in which

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methyl tetrahydrofolate as the reaction product is quantified by its oxidation via methylene tetrahydrofolate to methenyl tetrahydrofolate (96). Little is known about the chloromethane dehalogenase, which has not yet been purified due to its oxygen sensitivity and to the complex composition of this multi-enzyme system. The methyl chloride dehalogenase of A. dehalogenans appears to consist of at least 3 to 4 separate proteins, one of which carries a corrinoid cofactor as the primary acceptor of the methyl group (Messmer et al., unpublished data). The corrinoid protein has been purified to apparent homogeneity. The methyl chloride dehalogenase enzyme system appears to be analogous to Odemethylases purified from the same organism (40, 71-73). The latter enzyme system mediates the transfer of methyl groups derived from different methoxylated aromatic compounds to tetrahydrofolate. The reaction mechanism ofthe O-demethylase is shown in Figure 5.2. The O-demethylases consist of four distinct protein components. The first component, designated methyl transferase I (MT I), catalyzes the transfer of the methyl group from the substrate to a second component, a corrinoid protein in its super-reduced Co(I)-state. The third component is a methyl transferase II (MT II), which mediates the transfer of the methyl group to tetrahydrofolate. The super-reduced corrinoid protein is known to undergo autoxidation due to its low redox potential. Since the corrinoid has to be reduced to the Co(I)-state to accept the methyl group, a repair mechanism is required in case of autoxidation of the corrinoid protein. This repair mechanism, which requires a fourth protein component (activating enzyme, or AE) was shown to be dependent on an electron donor and substoichiometric amounts of ATP, both for the Odemethylases (40, 71) and the methyl chloride dehalogenase (95). For the latter enzyme, the electrons are provided by in the presence of hydrogenase, whereas the former system (O-demethylase) can be replaced by titanium(III) citrate. Reduction of the corrinoid protein of the dehalogenase with an artificial electron donor is difficult, and might contribute to its extreme oxygen sensitivity. The mechanism of reactivation of the corrinoid protein is not yet understood.

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Evidence is available that methyl chloride dehalogenase, as well as O-demethylase are inducible enzyme systems, that are formed during incubation with the corresponding growth substrates. In addition, cells induced with methoxylated aromatics are also able to slowly convert methyl chloride in vitro (F. Kaufmann, unpublished results), which suggests that methylation of the O-demethylase corrinoid protein by methyl chloride may occur either enzymatically or abiotically. It should be noted, however, that the corrinoid proteins of the O-demethylases and of the methyl chloride dehalogenase are different with respect to their N-terminal ammo acid sequence (M. Messmer et al., unpublished).

3.2. Aerobic Dechlorination of Methyl Chloride Surprisingly, several aerobic bacteria have been reported to possess a similar pathway of methyl chloride conversion (28, 155, 156). However, the enzyme systems of these bacteria are less complex than the strictly anaerobic system described above. For example, in reported studies where chloromethane served as a growth substrate for Methylobacterium sp. strain CM4 (155), the methyl chloride dehalogenase was inducible. Mutants lacking the ability to grow with methyl chloride were still able to utilize methanol, methylamine, and other reduced compounds, indicating that different enzyme systems were responsible for conversion of the different methyl substrates (155, 156). Studies on the enzyme activities involved in methyl chloride conversion by these bacteria led to the conclusion that the methyl group is first transferred to a bound corrinoid cofactor by an enzyme designated CmuA. Subsequently, a second enzyme, CmuB, mediates the methyl transfer from the corrinoid protein to tetrahydrofolate. The gene sequences encoding CmuA and CmuB appear to exhibit some similarity to the methanogenic methyl transferase systems (156). Although methyl chloride dehalogenase activity is stimulated by titanium(III) citrate, but without a strict requirement for this artificial electron donor, ATP or other potentially reactivating cofactors, such as GTP or S-adenosyl-methionine, are not required. This is, in contrast to the anaerobic enzyme system described above. CmuA has been purified and characterized (T. Leisinger, personal communication), and the 68 kDa protein was shown to contain the corrinoid cofactor. Therefore, it is assumed that in this organism, the putative methyl transferase I and the corrinoid protein involved in methyl chloride dehalogenation of A. dehalogenans are replaced by a single polypeptide in strain CM4. CmuB (methyl transferase II) has recently been purified and characterized (140, 141). The homodimeric enzyme has an apparent molecular mass of 33 kDa per subunit. It mediates the methyl transfer from methylcobalamin, which could serve as a substitute for the methylated CmuA, to tetrahydrofolate. Tetrahydromethanopterin cannot replace tetrahydrofolate as methyl acceptor. Another aerobic, facultatively methylotrophic bacterium which clusters with Rhizobium spp. and is designated strain CC495, has recently been isolated with methyl chloride as sole carbon source (28). Higher growth yields are observed with methyl chloride than with methylamine as growth substrate. Growth on methyl chloride strictly depends on the presence of cyanocobalamin. Methyl chloride conversion was inducible, since methylamine-grown cells did not convert methyl chloride. Hydrogen sulfide was assumed to be the physiological methyl acceptor, since a methanethiol oxidase could be detected in cell extracts. Cell extracts mediated the transfer of the methyl group

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of methyl chloride, methyl bromide, or methyl iodide to a variety of methyl acceptors such as and (in order of decreasing efficacy). Hence, the methyl halide dehalogenase of strain CC495 could act as a transhalogenation system. The monomeric 67 kDa methyl transferase of strain CC495 has been purified and characterized (28). The enzyme contains one mol of corrinoid per mol of enzyme. Metals other than cobalt, such as zinc or iron, were not detectable in the purified protein. The enzyme was activated by dithiothreitol (DTT) and methyl chloride; Ti(III) could not replace DTT and S-adenosylmethionine could not replace methyl chloride. As isolated, the enzyme exhibited spectral properties suggesting the corrinoid component to be present in the cob(II)alamin form. Upon activation of the enzyme with DTT and methyl chloride, the methylated cobalamin was formed. Under these conditions, DTT was able to completely reduce the corrinoid to its super-reduced oxidation state. It appears that the aerobic, corrinoid-dependent methyl halide dehalogenases are less susceptible to oxidation and can more easily be reduced, indicating that the redox potential of the aerobic corrinoids is more positive than those found in strictly anaerobic systems. Together with the lower complexity of the aerobic systems, as compared to the anaerobic enzymes, this has proved helpful for the elucidation of the reaction mechanism of the corrinoid-dependent methyl halide dehalogenation.

3.3. Anaerobic Dechlorination of Dichloromethane A strictly anaerobic homoacetogen, Dehalobacterium formicoaceticum, has been recently isolated with dichloromethane as energy source (87). The organism converts dichloromethane to formate and acetate (2:1) as end products of the fermentation (Figure 5.1). Cell extracts of the organism mediate the formation of methylene tetrahydrofolate from dichloromethane and tetrahydrofolate (88). The reaction is dependent on the presence of substoichiometric amounts of ATP, methyl viologen as artificial electron carrier, and hydrogen as electron donor. The reaction is inhibited by 1-iodopropane in a lightly-reversible manner, suggesting the involvement of a corrinoid in the superreduced oxidation state. Hence, the system appears to resemble the strictly anaerobic methyl chloride dehalogenase of A. dehalogenans, described in Section 3.1. Surprisingly, no intermediary formation of methyl chloride, methyl tetrahydrofolate, or other free- or tetrahydrofolate- bound compounds was observed. In contrast to the methyl chloride dehalogenation, dichloromethane conversion requires two cleavages of a carbon-halogen bond. The mechanism of methylene tetrahydrofolate formation from dichloromethane and tetrahydrofolate is not yet understood. Although experimental evidence for this is still lacking, chloromethyl-cobalamin and 5-chloromethyl tetrahydrofolate are implicated to be potential intermediates in the reaction.

4. COMETABOLIC ANAEROBIC REDUCTIVE DECHLORINATION 4.1. Bacteria Catalyzing Cometabolic Reductive Dechlorination In the late 1970s, degradation studies of hexachlorocyclohexane transformation with a number of pure cultures showed that reductive dechlorination of certain compounds is quite widespread among bacteria. Numerous studies with facultative and strictly

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anaerobic bacteria have subsequently been carried out, mainly with and chlorinated compounds such as tetra- and trichloromethane, di- and trichloroethanes, and tetrachloroethene (Table 5.1; reference 39). The broad spectrum of bacteria with the ability to reductively dehalogenate aliphatic compounds suggests that alkyl reductive dehalogenation is a cometabolic activity of many of these bacterial strains. There is only one report of cometabolic reductive dehalogenation activity with aromatic chlorinated compounds, namely the reductive dechlorination of 1,2,4-trichlorobenzene by Staphylococcus epidermidis (147). The involvement of cometabolism in reductive dechlorination reactions that had been observed in environments such as aquifers and sediments is difficult to assess. The reductive dechlorination activity observed in unadapted methanogenic granular sludge (152-154) and the isolation of clostridia (23, 47), methanogens (20), and even facultative anaerobes (129) with quite high dechlorination activities, indicate that the importance of such cometabolic processes should not be underestimated.

4.2. Biochemistry of Cometabolic Dechlorination by Anaerobic Bacteria Several enzyme systems appear to be involved in cometabolic alkyl reductive dehalogenation reactions. They include protein-bound tetrapyrrole cofactors (iron(II) porphyrins, corrinoids, or factor flavoprotein-flavin complexes, and ferredoxins. Although reductive dehalogenation activity using the free forms of cofactors, such as iron(II) porphyrins, corrinoids, and coenzyme has been demonstrated (48, 51, 65, 74, 75), it is most probably the enzyme-bound form of the cofactors that catalyses the reductive dehalogenation activity observed with whole cells. The reducing equivalents, i.e., electron donors, required for enzyme activity are generated by cellular metabolism and thus cometabolic reductive dechlorinations should be designated as biological rather than abiotic reactions. The heme protein, cytochrome isolated from Pseudomonas putida grown on camphor as energy and carbon source, is able to reductively dechlorinate trichloronitromethane (21). Evidence has been presented that indeed the cytochrome is responsible for the dechlorination observed with whole cells. The ironreducing bacterium Shewanella putrefaciens reductively dechlorinates tetrachloromethane stoichiometrically to trichloromethane (120). Indications have been obtained that c-type cytochromes produced during microaerophilic growth are involved in the reductive dechlorination reaction observed in cell suspension experiments. Attempts to link the reductive dehalogenation activity found in homoacetogens and methanogens with corrinoid-containing enzyme systems has not lead to conclusive results. In some cases, strong indications for a link between reductive dechlorination and corrinoid-containing enzymes have been obtained, whereas other studies have indicated the absence of such a link. Homoacetogenic bacteria reductively dechlorinate tetrachloroethene and tetrachloromethane. The photoreversible inhibition of the reductive dechlorination of tetrachloroethene by 1-iodopropane in cell-free extracts of the homoacetogen, Sporomusa ovata, suggests the involvement of corrinoids (145). On the other hand, only a poor correlation between corrinoid content and rates of tetrachloromethane dechlorination by different protein fractions of another homoacetogen, Acetobacterium woodii, has been found (139). It is noteworthy that the

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transformation of tetrachloromethane by acetogens only partially leads to the formation of reductive dechlorination products. Carbon monoxide and carbon dioxide, as well as non-volatile compounds, have been observed in whole cell experiments. Although the formation of the latter products can be explained by a two-electron reduction to a carbenoid that hydrolyzes to form carbon monoxide, it is possible that as yet unidentified enzyme systems are involved in the tetrachloromethane transformation by homoacetogens. The dechlorination mechanism of PCE by Clostridium bifermentans strain DPH-1, recently isolated from a contaminated site (23), has also been characterized in cell extracts. Photoreversible inhibition of PCE dechlorination by 1-iodopropane in titanium(III) citrate reduced cell extracts, and inhibition by cyanide indicate that a corrinoid enzyme is involved in the dechlorination reaction (22). A 70 kDa dimeric PCE reductive dehalogenase, with a specific activity of 1.2 nkat/mg protein and a of 12 has been isolated from cell extracts of C. bifermentans strain DPH-1 (118). The photoreversible inhibition by 1-iodopropane indicates the presence of a corrinoidenzyme. However, cobalt analysis, or cyanolysis to extract the corrinoid from the enzyme, to prove the presence of a corrinoid have not yet been performed. Methanogens reductively dechlorinate compounds such as terra- and trichloromethane, 1,2-dichloroethane, and tetrachloroethene. The purified corrinoidcontaining carbon monoxide dehydrogenase of Methanosarcina thermophila, an acetoclastic methanogen, has been reported to dechlorinate trichloroethene, mainly to cis-l,2-dichloroethene and to traces of trans-1,2-dichloroethene, vinyl chloride, and ethene (68). The absence of inhibition by the corrinoid inhibitor, 1-iodopropane, reported for cell-free extracts of Methanobacterium thermoautotrophicum on the other hand, has suggested that corrinoid-containing enzymes might not be involved in the reductive dechlorination of 1,2-dichloroethane, a reaction catalyzed by many different methanogens (66). Rather, the specific inhibitor approach, using 2bromopropanesulfonate with cell-free extracts of Methanobacterium thermoautotrophicum indicated that the reductive dechlorination of 1,2dichloroethane by methanogens was catalyzed by the coenzyme enzyme, methyl-coenzyme M reductase (66). 2-Bromopropanesulfonate almost completely inhibited 1,2-dichloroethane dechlorination in cell-free extracts, whereas 1 -iodopropane had no effect. The reductive dechlorinating activity of purified methyl-coenzyme M reductase further supports the possible role methyl-coenzyme M reductase in the reductive dechlorination of 1,2-dichloroethane by methanogens. The involvement of flavoprotein-flavin complexes in reductive dehalogenation reactions has been observed with Escherichia coli and Pseudomonas putida. Membrane fractions of E. coli cells reductively dechlorinated l,l,l-trichloro-2,2-bis(pchlorophenyl)ethane (DDT) under anaerobic conditions to l,l-dichloro-2,2-bis(pchlorophenyl)ethane (DDD), in the presence of reduced FAD (46). In anaerobic incubations of a 6-10 kDa flavoprotein isolated from Ps. putida, DDT was dechlorinated when flavin cofactors such as FAD, riboflavins, or FMN were added to the reaction mixture (41).

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5. DEHALORESPIRATION BY ANAEROBIC BACTERIA At present, there are numerous bacterial pure cultures that are able to utilize a chlorinated compound as terminal electron acceptor during anaerobic respiration. The majority of the isolates utilize chlorophenolic compounds, tetrachloroethene, or both, whereas some isolates utilize chlorobenzoate, bromophenols, trichloroacetate, and chlorobenzenes (Table 5.2). All of the known dehalorespiring microorganisms are bacteria, and fall into four distinct phylogenetic branches, namely the Gram-positives with a low G+C content, the and and the chloroflexi. A more detailed discussion of the phylogeny and diversity of dehalorespiring bacteria is given in Chapter 3 of this book. In this section, we focus on the physiology (electron transport, and donors and acceptors used, nutritional requirements), the bioenergetics (growth yields, generation of a proton-motive force, composition of the respiration chain), the biochemistry (properties of the reductive dehalogenases, reaction mechanisms), and the genetics (characteristics of the genes and operons, regulation of gene expression) of dehalorespiring bacteria (DRBs).

5.1. Physiology of Dehalorespiring Bacteria (DRB)

5.1.1. Electron Donors Used by DRB Electron donors utilized by many DRB are pyruvate, lactate, hydrogen, and formate (Table 5.2). With the latter two substrates, it has been possible to show unambiguously that halogenated compounds can serve as terminal electron acceptors in an anaerobic respiration process, as was first demonstrated for Desulfomonile tiedjei, which utilizes 3-chlorobenzoate as electron acceptor (35,102). Some isolates utilize acetate as electron donor, but not hydrogen (31, 77, 144). Other organic compounds that are used by some DRB as a carbon and energy source include fermentation products such as ethanol, propionate, butyrate, and crotonate. Some DRB are also able to ferment pyruvate. Hydrogen and formate are two electron donors widely utilized by the majority of DRB. Three isolates, Dehalobacter restrictus PER-K23, Dehahcoccoides ethenogenes strain 195 and Dehalococcoides sp. strain CBDB1, are even restricted to the utilization of hydrogen. Hydrogen and formate play an important role in syntrophic interactions, i.e., electron transport, between bacteria oxidizing fermentation products such as propionate, butyrate, and ethanol, and bacteria utilizing these two substrates (124). Based on thermodynamic considerations, DRB should out-compete hydrogenotrophic sulfate-reducers and methanogens for the hydrogen produced by syntrophic bacteria. This has actually been confirmed by several studies with mixed bacterial cultures (3, 44, 45, 81, 131, 164). Under conditions where hydrogen is slowly produced at low levels, reductive dechlorination generally occurs in favor of methanogenesis (see Chapter 4).

5.1.2. Electron Acceptors Used by DRB The range of halogenated compounds that are used as electron acceptors in anaerobic respiration processes comprises three major groups, e.g., chlorobenzoates, halogenated phenolic compounds, and chloroethenes (Table 5.2). One isolate utilizes trichloroacetate

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and one utilizes chlorobenzenes as electron acceptor. Some DRB, such as Desulfomonile tiedjei and Desulfitobacterium dehalogenans, use a certain group of chlorinated compounds as electron acceptor but also fortuitously dechlorinate others, presumably in a cometabolic reaction (26). The majority of DRB that utilize chlorophenolic compounds belong to the genus Desulfitobacterium. DRB dechlorinate chlorophenolic compounds primarily at the ortho position. However, some strains are able to dechlorinate at the ortho and meta positions, and one strain, Desulfitobacterium frappieri strain PCP-1, dechlorinates at the ortho, meta, and the para position. The dechlorination ofpolychlorinated phenolic compounds is often incomplete, resulting in the accumulation of tri-, di-, and monochlorophenols. Among the PCE-dechlorinating DRB, only Dehalococcoides ethenogenes performs a complete dechlorination to ethene. However, the last dechlorination step from VC to ethene is a cometabolic transformation and not linked to anaerobic respiration (93). Other PCE-dechlorinating DRB, such as Desulfitobacterium sp. strains PCE-S and TCE1, Dehalobacter restrictus, Desulfuromonas chloroethenica TT4B, and Dehalospirillum multivorans, only partially dechlorinate PCE, producing primarily cisDCE. Desulfitobacterium sp. strain PCE1, a member of the genus from which a remarkably high number of DRB has been isolated, dechlorinates PCE only to TCE. However, this strain can also utilize chlorophenolic compounds as electron acceptors, a property also found for the PCE-dechlorinating Desulfitobacterium sp. strain PCE-S (52). It has been shown recently that Desulfitobacterium sp. strain PCE 1 possesses two different reductive dehalogenases, one specific for PCE and the other for chlorophenolic compounds (150). This is probably also true for Desulfitobacterium sp. strain PCE-S, since the purified PCE reductive dehalogenase of this organism is not able to dechlorinate chlorinated phenols, which are reduced in growing cultures. Trichlorobacter thiogenes, a bacterium isolated from an anaerobic soil enrichment, utilizes acetate as carbon and energy source and dechlorinates trichloroacetate to dichloroacetate with a stoichiometry of four mol chloride produced per mol of acetate oxidized to (31). It is proposed that a sulfur-sulfide redox cycle is involved in this process. Dehalococcoides sp. strain CBDB1 is the only bacterium isolated in pure culture so far that dechlorinates chlorobenzenes in an anaerobic respiration process, although indications for the existence of such bacteria had been obtained quite some time ago (63). This strain is phylogenetically closely related to Dehalococcoides ethenogenes, but cannot use PCE. Most DRB are able to use electron acceptors other than chlorinated compounds (Table 5.2). Three known exceptions are Dehalobacter restrictus and the two Dehalococcoides strains 195 and CBDB1. This is very surprising since the chlorinated compounds utilized by these strains, although possibly produced naturally in certain specific environments (53), probably do not accumulate naturally in significant amounts to guarantee the survival of these bacteria. Nitrate is another exogenous electron acceptor that is used by a considerable number of DRB, and is incompletely reduced to nitrite. Sulfur oxyanions, such as sulfite and thiosulfate, but not necessarily sulfate, are also used, as by the genus Desulfitobacterium. Sulfite and sometimes also thiosulfate have an inhibitory effect on the dechlorination activity of DRB (Table 5.2). Since sulfite also often inhibits reductive dehalogenases directly (Table 5.3), it is not known whether the inhibition of dechlorination in whole

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cells is due to a preferential use of this electron acceptor, or due to a direct inhibition of the dechlorinating enzyme. Dechlorination by Desulfitobacterium dehalogenans cultures is not inhibited in the presence of nitrate, sulfite, or thiosulfate, whereas relative activities of 78, 39, and 65% have been observed upon addition of nitrate, sulfite, and thiosulfate to cell extracts, respectively (150). This suggests that the reductive dehalogenase of Desulfitobacterium dehalogenans is protected against these compounds in intact cells. However, a competition for reducing equivalents between the fumarate reductase and the reductive dehalogenase has been proposed (150). This is based on the observation that the dechlorination rate was only 85% upon addition of fumarate to a growing culture, compared to controls without fumarate present. Moreover, fumarate did not inhibit the reductive dehalogenase in cell extracts. PCE dechlorination by growing cultures of Dehalospirillum multivorans was almost completey inhibited in the presence of fumarate, whereas fumarate had only little effect on the dechlorinating activity of

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resting cells (107). This indicates that fumarate may be preferentially used as electron acceptor. A similar effect has been observed with the electron acceptor polysulfide. Bacteria that use a broad range of electron acceptors must also develop a regulation strategy to determine which electron acceptor to use. In several cases, it has been shown that dehalogenation activity is induced by the chlorinated electron acceptor (99, 112, 137). In Desulfitobacterium dehalogenans, evidence has been obtained that regulation takes place at the level of gene transcription (137) but more studies with other DRB are necessary to draw general conclusions. Indications for the induction of different reductive dehalogenase enzyme systems have been reported for Desulfitobacterium frappieri strain PCP-1 and Desulfitobacterium sp. strain PCE1. Strain PCP-1, possesses one inducible reductive dehalogenase system for ortho-dechlorination of chlorophenols and one for meta-dechlorination. Desulfitobacterium sp. strain PCE1 grown with 3chloro-4-hydroxyphenoxyacetate (Cl-OHPA) as electron acceptor did not dechlorinate PCE, whereas chlorophenolic compounds were not dechlorinated by cells grown with PCE (50,150). Whether electron acceptors such as nitrate, fumarate, sulfate, sulfite, and thiosulfate have a regulatory effect on the genetic level of anaerobic dechlorination is not yet known.

5.1.3. Nutritional Requirements Many DRB are often cultured with yeast extract in the medium, but it is not certain whether the DRB really depend on the growth factors added with yeast extract. Some strains, such as Desulfomonile tiedjei DCB-1, Dehalobacter restrictus strain PER-K23, and Dehalococcoides ethenogenes 195 were only isolated upon addition of rumen fluid, fermented yeast extract, or anaerobic digester sludge supernatant, to their respective culture media (33, 64, 94). For two of the strains, the precise need of growth factors has been studied in detail. Desulfomonile tiedjei requires the vitamins thiamine, nicotinamide, and lipoic acid. In addition, hemin and an unusual amendment of 1,4naphthoquinone is required by this bacterium (33). Two halophiles, Desulfomonile liminaris strain DCB-M and strain DCB-F, have been enriched on media with similar growth factors as for Desulfomonile tiedjei, although it is not known whether the Desulfomonile liminaris strains really depend on the presence of these growth factors (144). Growth of Dehalobacter restrictus strain PER-K23 depends on the addition of two vitamins, thiamine and cyanocobalamin, and three amino acids, arginine, histidine, and threonine (67). For the isolation of Dehalobacter restrictus strain TEA, 5% (v/v) of filter-sterilized spent medium from a fixed-bed anaerobic reactor was added as source of growth factors (162). The exact nutritional requirements of this strain, however, have not been determined.

5.2. Bioenergetics of Dehalorespiration

5.2.1. Growth Yields of DRB The coupling of hydrogen and formate oxidation to reductive dechlorination implies energy conservation via electron transport phosphorylation. The free energy available from the dechlorination of PCE to DCE with hydrogen as electron donor is 189

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kJ per mol This is enough energy to yield approximately 2.5 ATP, assuming an energy requirement of 70 kJ per mol for the formation of 1 ATP, out of 1 ADP and 1 (125). For the dechlorination of 3-chlorobenzoate, the available free energy is somewhat lower (137 kJ per mol H2), but it would still be sufficient to yield approximately 2 ATP. Growth yields of DRB with hydrogen or formate as electron donor range from 1.2 to 2.1 g protein per mol chloride released (Table 5.3). Two exceptions are Desulfitobacterium frappieri strain TCE1 and Dehalococcoides ethenogenes, for which growth yields of 3.2 to 3.9, and 4.8 g protein per mol chloride released have been reported, respectively (50, 94). DRB growing with hydrogen as electron donor need acetate as a carbon source. Theoretically, 10 g of cell material can be formed per mol ofATP when acetate is the carbon source (138). Estimating the yields in cell dry weight from the reported protein yields, with the assumption that the protein content is equal to 60% of the cell dry weight, one obtains a range of 2 to maximally 8 g cell dry weight per mol chloride released. This indicates that the anaerobic respiration with chlorinated compounds generally yields less than 1 ATP per chloride released. Numerous DRB can ferment pyruvate, and it is less clear whether the reductive dechlorination reaction is coupled to electron transport phosphorylation if pyruvate serves as carbon and energy source in the presence of a chlorinated compound serving as electron acceptor. Two studies have investigated in more detail the differences between the growth yields on pyruvate with and without a chlorinated compound. In the absence of3-chloro-4-hydroxyphenyl acetate (Cl-OHPA), a growth yield of 13.4 to 14.2 g protein per mol acetate produced was found for Desulfitobacterium dehalogenans (85, 150). Pyruvate was fermented to equimolar amounts of lactate and acetate in a HEPESbuffered medium (85), whereas a ratio between lactate and acetate of 1:2 was found when cultivated in a bicarbonate-buffered medium (150). The additional acetate produced is probably the product of reduction to acetate. On the other hand, the growth yields obtained on pyruvate plus Cl-OHPA were quite different. In the first study, the growth yield on pyruvate was found to be twice as high in the presence of Cl-OHPA [25.4 g protein per mol acetate; reference (85)]. In the other study, no increase in growth yield per mol acetate produced was found upon addition of the chlorinated electron acceptor[12.8 g protein per mol acetate (150)]. In both studies, it has been shown that pyruvate is stoichiometrically oxidized to acetate and carbon dioxide, that one mol of Cl-OHPA is dechlorinated per mol pyruvate oxidized, and that lactate is not produced. During the oxidation of pyruvate to acetate, one ATP can be formed as a result ofsubstrate-level phosphorylation, and 13.5 g cell biomass can be formed per mol ATP when pyruvate is the carbon source (138). Based on this reasoning, the second study indicates that no proton motive force is created upon electron transfer from pyruvate to Cl-OHPA, whereas the first study suggests that additional ATP is formed during reductive dechlorination, presumably by electron transport phosphorylation.

5.2.2 Generation of a Proton Motive Force by Dehalorespiration Experiments with uncouplers, protonophores, and oxidant pulses in cell suspensions of Desulfomonile tiedjei DCB-1 have indicated chemiosmotic coupling of dechlorination and ATP synthesis (104). Oxidant pulse experiments, performed by adding limiting amounts of 3-chlorobenzoate to cell suspensions of Desulfomonile tiedjei that

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were saturated with as electron donor, resulted in an acidification of the cell suspension with a ratio of 1.05 (104). This proton liberation was significantly reduced in the presence of the uncouplers 2,4-dinitrophenol or pentachlorophenol. A very similar ratio of 1.25 has been reported for the PCE-dechlorinating bacterium Dehalobacter restrictus PER-K23 (127). This proton liberation was inhibited by the protonophore carbonylcyanide m-chlorophenylhydrazone (CCCP), but PCE and TCE were still reductively dechlorinated. A similar effect that has also been observed for Desulfomonile tiedjei. Dechlorination by Dehalospirillum multivorans, on the other hand, has been shown to be inhibited by ionophores in cell suspensions but not in cell extracts. This indicates that the bacterium needs a membrane potential and/or a pH gradient for the in vivo reaction.

5.2.3. Topology of the Dehalorespiration Chain Electron transport phosphorylation requires an association of the mediating proteins with the cytoplasmic membrane. The localization and arrangement of the two major enzymes involved in hydrogen- or formate-dependent dehalorespiration have been investigated for six DRB. The majority of the reductive dehalogenases described so far have been found in the membrane fraction, and only the use of detergents allowed their solubilization and purification (Table 5.4). One exception is the PCE reductive dehalogenase of Dehalospirillum multivorans that has always been recovered in the soluble fraction. Based on DNA sequence data, it has been proposed that reductive dehalogenases may be anchored on the membrane by a hydrophobic protein RdhB, a product of the gene rdhB which is located in the same operon as the gene rdhA that encodes the reductive dehalogenase (see Section 5.4.2). In all cases where also the electron donating part of the respiration chain has been investigated, a certain degree of membrane association of the hydrogenase or formate dehydrogenase has been reported (82, 90, 99, 100, 127, 150). In addition, it has been shown by the use of the membrane-impermeable artificial electron acceptor methyl viologen and the membrane-impermeable hydrogenase inhibitor that the hydrogenases and formate dehydrogenases are facing the outside of the cytoplasmic membrane. The same approach was applied for reductive dehalogenases, with reduced methyl viologen as electron donor, and results indicated that reductive dehalogenases are facing the inside of the cytoplasmic membrane due to the 2 to 10-fold increase in dechlorinating activity upon permeabilization of the cells (99, 100, 127, 150). For Desulfitobacterium dehalogenans, this increase in activity was rather low, and it has been reported for Dehalococcoides ethenogenes that four-fold higher activities with whole cells have been found with methyl viologen as electron donor compared with hydrogen (113). It is not known from the latter study whether a permeabilization of the cells causes an increase of the activity of the reductive dehalogenase. Nevertheless, it cannot be ruled out that the reductive dehalogenase can also face the outside of the cytoplasmic membrane. For the reductive dehalogenase of Dehalospirillum multivorans which has always been found in the soluble fraction, it has been shown with spheroplast experiments that the enzyme is not present in the periplasm, but in the cytoplasm (107).

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5.2.4. Other Components of the Dehalorespiration Chain A s in any respiratory system, electron transport from the electron donating member to the electron accepting terminal reductase, e.g., the reductive dehalogenase, has to occur in the cytoplasmic membrane, during which a proton gradient is possibly generated. Hypothesized components that take part in the electron transfer within the membranes of DRB are cytochromes and quinones. Membrane-bound c-type cytochromes have been found in Desulfomonile tiedjei (83), Desulfitobacterium hafniense (24), Desulfitobacterium dehalogenans (150), and Desulfuromonas chloroethenica (76), whereas a soluble one is present in Dehalospirillum multivorans (126). Indications for the involvement of cytochrome c in electron transfer have been obtained with Desulfitobacterium dehalogenans, where the cytochrome has been shown to be reduced when formate was added as electron donor and re-oxidized upon addition of Cl-OHPA (150). The cytochrome c of Desulfomonile tiedjei is co-induced with dechlorination activity and is only present in dechlorinating cells (83). Cytochromes b are present in membranes of Dehalospirillum multivorans (126), Dehalobacter restrictus (67), and Desulfitobacterium dehalogenans (150). Indications for an electron transfer function of these cytochromes during dehalorespiration have been found for Desulfitobaterium dehalogenans (150) and for Dehalobacter restrictus (Schumacher & Holliger, unpublished), by recording difference spectra of cell fractions in the presence of a physiological electron donor, before and after the addition of an electron accepting chlorinated compound. Menaquinones have been detected in membranes of Dehalospirillum multivorans (126), Dehalobacter restrictus (127), and Desulfitobacterium dehalogenans (150). Redox difference spectra of membrane fractions and experiments with the menaquinone analog, 2,3-dimethyl-l,4-naphthoquinone (DMN), and the inhibitor, 2-n-heptyl-4hydroxyquinoline-N-oxide (HOQNO), have provided evidence for the involvement of menaquinone in the electron transfer in Dehalobacter restrictus (127). An electron transfer component between menaquinone and the PCE reductive dehalogenase has been proposed, based on experiments with reduced DMN (61). DMNH2 could act as electron donor for the PCE reductive dehalogenase in membrane fractions, but not in membrane extracts and not for the purified enzyme. This indicates that the respiration chain has to be intact, and that menaquinone is not the direct electron donor of the PCE reductive dehalogenase. Indications for menaquinone-mediated electron transfer during dehalorespiration have also been obtained for Desulfitobacterium dehalogenans, by determining the ratio of reduced and oxidized menaquinone extracted under different conditions from cells grown with formate and Cl-OHPA (150). A yet undefined quinone has been isolated from Desulfomonile tiedjei, and the inhibition of dechlorination by HOQNO indicates that a quinone is involved in the electron transfer to the chlorinated compound (82). HOQNO did not inhibit the dechlorination by whole cells of Dehalospirillum multivorans and Dehalococcoides ethenogenes (99, 114), whereas the presence of quinones has been shown for the first organism, it is not known whether any are present in the latter.

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5.2.5. Proposed Schemes of the Dehalorespiration Chain Based on the experimental evidence concerning the topology of the enzymes involved in dehalorespiration and the components that appear to be involved in the electron transfer from the electron donating enzyme to the terminal reductase, two generalized schemes of the dehalorespiration chain are proposed (Figure 5.3). In both schemes, hydrogenases are the electron donating enzymes, but they can be replaced by formate dehydrogenases, since these enzymes are also facing the outside of the cytoplasmic membrane. Scheme A depicts a proposed respiration chain based in large part on the available evidence. The proposed localization of the hydrogenase and the reductive dehalogenase enzymes yields a proton gradient of per mol consumed by a scalar process. In Dehalobacter restrictus PER-K23, it has been shown that menaquinone is not acting as

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a proton pump (127). It can, however, not be excluded that quinones in other DRB contribute to the creation of proton motive force (dashed arrow in the figure). Since it is not known what other components are involved in electron transfer, proton translocation across the membrane during another step of this process is also conceivable (pointed arrow). The two protons produced by the scalar process of hydrogen oxidation correspond to two thirds of an ATP, assuming that three protons cross the membrane per ATP formed (125). This is in agreement with the low cell yields of DRB, and indicates that not even one mol ATP is gained per mol chloride removed. It also corresponds with the ratios found for Desulfomonile tiedjei DCB-1 and Dehalobacter restrictus PER-K23 (104, 127). Scheme B is a quite speculative representation of the dehalorespiration chain. All reductive dehalogenases sequenced so far share a signal sequence with a twin arginine motif (see section 5.4.1). This signal sequence is cleaved off during post-translational processing. These signal peptides are thought to play a key role in the maturation and translocation of periplasmic proteins by the recently described Twin Arginine Translocation (TAT) system (6). Therefore, reductive dehalogenases facing the outside of the cytoplasmic membrane cannot be excluded. In such a respiration chain, proton translocation during electron transfer has to occur, since the scalar proton production by hydrogen oxidation is compensated by the reductive dehalogenation reaction.

5.3. Biochemistry of Reductive Dehalogenases

5.3.1. Major Properties of Reductive Dehalogenases The reductive dehalogenases so far studied catalyze the in vitro reduction of chlorinated compounds with the artificial electron donor, methyl viologen -446 mV; according to the following equation:

This simple reaction allows for spectrophotometric quantification of the reaction and was essential for the purification of the dehalogenases. Due to the low redox potential of the reduced MV, this reaction is measured in the absence of molecular oxygen. The characteristics of purified reductive dehalogenases are summarized in Table 5.4. The 3-chlorobenzoate reductive dehalogenase of Desulfomonile tiedjei was the first that has been characterized biochemically (112). This reductive dehalogenase has several properties that distinguish it from the other reductive dehalogenases described. It is a heterodimer, with a small subunit that might contain a heme as cofactor. It has a very low specific activity (0.3 nkat/mg protein) and is insensitive to oxygen. All other biochemically characterized reductive dehalogenases have been isolated as monomeric enzymes. SDS gel analysis has revealed the presence of one subunit, with molecular masses between 48-65 kDa. First indications for the involvement of a corrinoid in reductive dehalogenation have been obtained with photoreversible inhibition by 1-iodopropane (100, 108, 127). For some of the purified reductive dehalogenases, determination of the cobalt content, and extraction of corrinoids by cyanolysis, have unambiguously shown that corrinoids are a cofactor of these enzymes, with a

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stoichiometry of 1 mol corrinoid per mol of catalytic unit (25, 101, 109, 128, 151). Electron paramagnetic resonance (EPR) spectroscopy with the reductive dehalogenases of Dehalobacter restrictus (128) and Desulfitobacterium dehalogenans (151) have revealed that the Co(I/II) and the Co(II/III) have relatively high redox potentials of-350 mV and >150 mV, respectively, compared with other corrinoid enzymes. Furthermore, the EPR studies have shown another unusual feature, in that the corrinoid in the Co(II)state is present in the base-off configuration. Analysis of the iron and acid-labile sulfide content of the purified enzymes has indicated the presence of iron-sulfur clusters. The PCE reductive dehalogenase of Dehalobacter restrictus contains two [4Fe-4S] clusters with rather low redox potentials of as shown by EPR spectroscopy (128). In another microorganism, Desulfitobacterium dehalogenans, the ortho-chlorophenol reductive dehalogenase contains one [4Fe-4S] cluster and one [3Fe-4S] cluster with redox potentials of and respectively (151). The identification of a ferredoxin-like and a truncated Fe-S cluster binding motif in the sequence of the reductive dehalogenase gene of Desulfitobacterium dehalogenans supports the presence of these two prosthetic groups (151; see also section 5.4.1). Three chloroethene and three chlorophenol reductive dehalogenases have N-terminal sequences with high homologies, and they also have quite similar molecular masses of 59-65 kDa and 47-48 kDa, respectively (Table 5.5). The N-termini of the rest of the three reductive dehalogenases do not have any homologies with each other and with the other six enzymes, and their molecular masses are either similar to the one (60 kDa) or the other group (48 kDa).

5.3.2. Reaction Mechanism Dehalogenases

of Corrinoid-Dependent Reductive

Low-potential electron donors (< -360 mV) are required for reductive dechlorination (99), yet the standard redox potential of the couple R-C1/R-H is usually > 200 mV. This suggests that corrinoids, which are required for PCE reductive dehalogenase activity, are present in an enzyme-bound Co(I)-state. Two feasible reaction mechanisms for reductive dechlorination of chlorinated compounds have been proposed: i) an addition of the Co(I) corrinoid to one of the carbons and subsequent of the chlorine substituent (163), or ii) a dissociative one-electron transfer from the Co(I) corrinoid to the chlorinated compound upon formation of a radical, which combines with a H-radical after elimination of a chloride anion (128). Evidence for a one-electron transfer is available from EPR data of the purified PCE reductive dehalogenase of Dehalobacter restrictus PER-K23 (128), and the chlorophenol reductive dehalogenase of Desulfitobacterium dehalogenans (151). In the reported studies, the completely reduced enzymes react very rapidly with the chlorinated substrate, yielding the base-off form of the corrinoid in the Co(II)-state. A radical signal has not been observed until now with EPR, but it is possible that the radical exists in an enzyme- or corrinoid-bound state, rather than in a free form. However, it cannot be excluded that the Co(III)-state, possibly formed in the course of the reaction, is immediately reduced to the Co(II)-state, since reducing conditions were required for these experiments.

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The PCE reductive dehalogenases of Dehalospirillum multivorans and Desulfitobacterium sp. strain PCE-S also enzymatically dechlorinate chlorinated propenes (100). 2,3-Dichloropropene and 1,1,3-trichloropropene were dechlorinated to 2-chloropropene and 1,2-dichloropropene, respectively (111). This reported elimination of the chlorine substituent from the hybridized C-1 carbon, rather than from the hybridized carbon, argues against an addition at C-1 and an elimination at the C-2 position, since this mechanism would yield vinyl chloride plus methyl chloride as the reaction products. Trans-l,3-dichloropropene has been shown to dechlorinate to a mixture of trans-1-chloropropene, cis-1-chloropropene, and 3-chloropropene. The formation of these three chloropropene isomers is in accordance with a dechlorination mechanism involving a radical intermediate (111). A study with the free corrinoids in reaction mixtures with titanium(III)-citrate or

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titanium(III)-NTA as reducing agents also indicates that a dissociative one-electron transfer mechanism is involved in the dechlorination of PCE and TCE (51). These results have been corroborated by a study employing radical traps, labeling experiments, and stopped-flow spectroscopy (130). A tentative scheme of a radical mechanism involved in reductive dechlorination is outlined in Figure 5.4. From all of the available experimental data , it appears that the corrinoid undergoes a change in its redox status in the catalytic cycle of the reaction. Hence, reductive dechlorination represents a new type of corrinoid-dependent reaction. The radical formed upon reduction by the Co(I) corrinoid appears to react with the simultaneously formed Co(II) corrinoid, yielding an alkyl-Co(III) corrinoid; the alkyl moiety gets protonated and is cleaved off. The remaining Co(III) corrinoid is reduced by the two iron-sulfur centers of the enzyme to the Co(I)-state. Another possibility is that the radical formed during the reaction gets reduced by one of the two iron-sulfur centers, whereas the other is reducing the Co(II) corrinoid to the Co(I)-state.

5.4. Genetics of Reductive Dehalogenases

5.4.1. Molecular Characteristics of Reductive Dehalogenases The gene sequences of different reductive dehalogenases (rdh) have been obtained by cloning and sequencing with reversed genetic approaches, using the N-terminal amino acid sequences of the purified proteins (89, 110, 136,151; Table 5.5) and from complete genome sequences of two DRB, Dehalococcoides ethenogenes strain 195 (www.tigr.org) and Desulfitobacterium hafniense strain DCB-2 (www.jgi.doe.gov). Their availability has enabled the elucidation of common features and differences among this novel class of corrinoid enzymes (136). Alignment of the complete amino acid sequence (Table 5.6) reveals a high degree of homology between chlorophenol reductive dehalogenases (60-99% similarity, and a low degree of homology between chloroethene reductive dehalogenases (27-32% similarity) and between chlorophenol reductive dehalogenases (26-36% similarity). This is in agreement with the alignment of the Nterminal sequences showing a quite similar grouping (Table 5.5). As another means of characterizing the genetic relationships between specific dehalogenase enzymes, Figure 5.5 shows the alignment of the amino acid sequences in the N-teminal and C-terminal part of the gene rdhA. Here, all sequences of the gene clearly show the presence of two conserved Fe/S cluster binding motifs. With the exception of the TCE reductive dehalogenase from Dehalococcoides ethenogenes (TceA), the first expected cysteine residue of the second cluster is in all cases replaced by a glycine residue, compared with normal bacterial ferredoxins (19). This is in agreement with the Fe/S clusters detected by EPR spectroscopy with the purified enzyme of Desulfitobacterium dehalogenans, showing one [4Fe-4S] cluster and one [3Fe-4S] cluster. The sequence of the TceA dehalogenase of Dehalococcoides ethenogenes indicates the presence of two [4Fe-4S] clusters, as has been shown for the purified reductive dehalogenase of Dehalobacter restrictus PER-K23 (128). Unfortunately, nothing is known about the content of iron and acid-labile sulfide of the enzyme from Dehalococcoides ethenogenes. In addition, the sequence of the reductive dehalogenase gene of Dehalobacter restrictus PER-K23 is not available.

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Despite the rather high degree of homology between the C-terminal regions of the reductive dehalogenases, they all lack the consensus sequence for binding of a corrinoid that has been identified in methylcobalamin-dependent methyltransferases and mutases (84). Since the EPR measurement with reductive dehalogenases clearly have indicated the presence of the base-off form of the Co(II) corrinoid in the enzyme, this finding is

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not surprising. Highly conserved residues of special interest are various histidines and tryptophans that might, in analogy, be involved with hydrolytic dehalogenases (30) and dichloromethane dehalogenases (91), in catalysis by binding the corrinoid cofactor, or by stabilizing the leaving halide (136). All rdhA gene sequences show that the enzyme RdhA is produced as a pre-protein containing a twin arginine signal sequence (RRXFXK), that is characteristic for extracytoplasmic proteins that bind different complex redox cofactors (Figure 5.5). This leader sequence is normally found in proteins which are exported to the periplasmic space by a twin arginine translocation (TAT) system (6), and is in contradiction with the topology of reductive dehalogenases since they most probably orient towards the cytoplasm. The dimethylsulfoxide reductase of Escherichia coli is one example of a cytoplasmic protein not exported to the periplasm that also contains the typical twin arginine leader sequence This protein contains a small hydrophobic subunit that prevents the enzyme from being exported (159). The product of the gene rdhB, a hydrophobic protein (see section 5.4.2), may fulfill a similar role in the reductive dehalogenases (110).

5.4.2. The Reductive Dehalogenase Gene Cluster and its Regulation The rdhA genes that code for catalytically active reductive dehalogenases have always been found to be linked with an open reading frame, referred to as rdhB, encoding a small hydrophobic protein that contains two to three transmembrane helices. This gene was first described in Dehalospirillum multivorans, and it has been proposed that it serves a double function, anchoring the RdhA protein on the membrane and preventing the export of the RdhA protein by the TAT system (110). This close linkage of an open reading frame encoding a small hydrophobic protein with the rdhA gene has also been found in all putative reductive dehalogenase gene clusters found in the genomes of Dehalococcoides ethenogenes (at least 17 homologues) and Desulfitobacterium hafniense (at least 5 homologues) (134, 136). The co-transcription of both genes rdhA and rdhB, as shown for Dehalospirillum multivorans (110) and Desulfitobacterium dehalogenans (137) by the Reverse Transcriptase-Polymerase Chain Reaction (RT-PCR), suggests a functional linkage of both gene products. All reductive dehalogenases have been purified as monomers, and the product of the gene rdhB never has been co-purified with the catalytic unit. Despite all of the indications on a genetic level, the role of RdhB remains to be confirmed on the protein level. The molecular characterization of a 11.5-kb genomic fragment containing the orthochlorophenol reductive dehalogenase-encoding cprAB genes of Desulfitobacterium dehalogenans has revealed the presence of eight designated genes with the order cprTKZEBACD (137). All of the genes have the same polarity, except cprT. The deduced products of the different genes indicated that cprC and cprK belong to the NirI/NosR- and CRP-FNR families of transcription regulatory proteins, respectively, where cprD and cprE are possibly encoding molecular chaperons of the GroEL type, and that cprT appears to encode a homologue of the trigger factor folding catalysts. No function has been assigned to the predicted product of cprZ. Transcription of cprAB increases 15-fold upon addition of Cl-OHPA, with a concomitant induction of dechlorination activity. Binding motifs that somewhat resemble the FNR box have been

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identified in the cprT-cprK and the cprK-cprZ intergenic region, as well as in the promoter of cprB, suggesting a role of CprK in the expression of the cprTKZEBACD genes (137). Based on these and other results obtained by the characterization of dehalorespiration deficient mutants (135), a model for the regulation and functional assembly of the ortho-chlorophenol reductive dehalogenase of Desulfitobacterium dehalogenans has been proposed. It consists of a histidine kinase that senses the chlorinated compound, and a response regulator that transfers the signal from the histidine kinase to constitutively produced CprK. The latter protein subsequently induces the transcription of the cprTKZEBACD genes (132). Nothing is known about the gene clusters and their regulation in other DRB, but analyses of the genomes of Desulfitobacterium hafniense and Dehalococcoides etheneogenes have indicated similarities of gene clusters present in the neighborhood of putative reductive dehalogenase genes to the gene cluster identified in Desulfitobacterium dehalogenans (134).

6. CONCLUSIONS Halogenated compounds developed by the chemical industry during the last few decades, and used as intermediates for chemical synthesis and in many other industrial applications, are metabolized by a wide spectrum of anaerobic bacteria. Metabolism occurs in three different ways as demonstrated by a variety of substrates. Chloro- and dichloromethane have been shown to serve as carbon and energy sources for specialized homoacetogenic bacteria, and 3-chlorobenzoate has been found to be a substrate for a denitrifying bacterium. Many different anaerobic bacteria cometabolically transform halogenated compounds via reductive dehalogenation reactions, while others utilize the halogenated compounds as terminal electron acceptors in an anaerobic respiration process. Corrinoids seem to play a key role in all different types of anaerobic metabolism of halogenated compounds. Biochemical studies performed on the mechanisms of chloromethane metabolism have shown a high resemblance to the homoacetogenic metabolism of methoxylated aromatic compounds. The initial step of methyl chloride dehalogenation is the transfer of the methyl group to tetrahydrofolate. Here, the biotransformation of chlorinated methanes as growth substrates occurs via a methyl-transfer reaction, rather than a reductive dechlorination. Subsequent steps in the pathway are the same as in acetate production from the methyl-tetrahydrofolate derived from methoxylated aromatic compounds. The methyl chloride dehalogenase contains a corrinoid protein, which is involved in the methyl-transfer. Cometabolic reductive dehalogenation has been described for many different groups of bacteria, including facultative anaerobes, fermentative bacteria, methanogens, and nitrate-, iron-, and sulfoxy anion-reducers. The reductive dechlorination activity observed in unadapted methanogenic granular sludge, and the subsequent isolation of clostridia, methanogens, and even facultative anaerobes with reltively high dechlorination activities, indicate that the importance of such cometabolic processes should not be underestimated. Enzyme systems so far recognized for their involvement in the cometabolic reductive dehalogenations, comprise protein-bound tetrapyrrole cofactors such as iron(II) porphyrins, corrinoids, and factor flavoprotein-flavin

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complexes, and ferredoxins. The enzyme-bound forms of the cofactors appear to catalyze the reductive dehalogenation reactions in whole cells, yet the supply of reducing equivalents to these enzymes depends on overall metabolic activity. This supports the premise that cometabolic reductive dechlorinations should be recognized as biological, rather than abiotic reactions. Dehalorespiring bacteria belong to four different phylogenetic groups. Primarily, they use fermentation products, such as hydrogen, formate, pyruvate, and lactate as electron donors. With a few exceptions, they can also use inorganic and organic compounds as electron acceptors. Rather simple respiratory chains appear to be involved, which use the energy gained from exergonic dechlorination reactions quite inefficiently. The majority of reductive dehalogenases purified so far are corrinoidcontaining enzymes that constitute a new class of corrinoid-enzymes, due to the electron transfer that they catalyze. There is some evidence that a dissociative one-electron transfer is involved, but additional research is required to fully understand the reaction mechanism. The gene cloning and gene inactivation systems recently developed for the dehalorespiring bacterium, Desulfitobacterium dehalogenans (133, 135), provide necessary tools for the elucidation of structure-function relations and the indentification of residues involved in catalysis. Comparison of apparent and putative reductive dehalogenase gene sequences has revealed the presence of a twin-arginine leader sequence that is cleaved off from the pre-protein. Also, the presence of binding motifs for two iron-sulfur clusters has been determined. In addition, an open reading frame encoding a hydrophobic protein has always been found in the vicinity of the reductive dehalogenase gene rdhA. This rdhB gene is co-transcribed with rdhA, and is proposed to function as a membrane-anchor of RdhA. The hypothesis remains to be validated at the protein level. It has also been shown that the production of the reductive dehalogenase is regulated on the transcription level. The increasing amount of sequence information on this novel anaerobic respiration process not only helps to unravel fundamental questions, but also provides a better means to exploit the potential of dehalorespiring bacteria for bioremediation of organohalide-contaminated sites. The development of dehalorespiration-specific molecular tools will enable the determination of whether a certain reductive dehalogenation potential is present at a specific site, and whether it is expressed under certain conditions. The future will tell whether transfer of this knowledge has occurred and helped to clean up the sites that we have polluted in the past.

Acknowledgments The research of the group of C.H. is supported by grants from the Swiss National Science Foundation and the Swiss Federal Office for Education and Science. The research group of G.D. is supported by grants from the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie. The authors thank Anke Neumann for critically reading the manuscript.

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DEHALOGENATION BY ANAEROBIC BACTERIA 371 144. Sun B, Cole JR., Sanford RA & Tiedje JM (2000) Isolation and characterization of Desulfovibrio dechloracetivorans sp. nov., a marine dechlorinating bacterium growing by coupling the oxidation of acetate to the reductive dechlorination of 2-chlorophenol. Appl. Environ. Microbiol. 66:2408-2413 145. Terzenbach DP & Blaut M (1994) Transformation of tetrachloroethylene to trichloroethylene by homoacetogenic bacteria. FEMS Microbiol. Lett. 123:213-218 146. Traunecker J, Preuss A & Diekert G (1991) Isolation and characterization of a methyl chloride utilizing, strictly anaerobic bacterium. Arch. Microbiol. 156:416-421 147. Tsuchiya T & Yamaha T (1984) Reductive dechlorination of 1,2,4-trichlorobenzene by Staphylococcus epidermidis isolated from intestinal contents of rats. Agric. Biol. Chem. 48:1545-1550 148. Utkin I, Woese C & Wiegel J (1994) Isolation and characterization of Desulfitobacterium dehalogenans gen. nov., sp. nov., an anaerobic bacterium which reductively dechlorinates chlorophenolic compounds. Int. J. Syst. Bacteriol. 44:612-619 149. Utkin I, Dalton DD & Wiegel J (1995) Specificity of reductive dehalogenation of substituted ortho- chlorophenols by Desulfitobacterium dehalogenans J W/IU-DC1. Appl. Environ. Microbiol. 61:346-351 150. van de Pas BA (2000) Biochemistry and physiology of h a l o r e s p i r a t i o n by Desulfitobacterium dehalogenans. Ph.D. thesis. University of Wageningen, The Netherlands. 151. van de Pas BA, Smidt H, Hagen WR, van der Oost J, Schraa O, Stams AJM & de Vos WM (1999) P u r i f i c a t i o n and molecular characterization of ortho-chlorophenol reductive dehalogenase, a key enzyme of halorespiration in Desulfitobacterium dehalogenans. J. Biol. Chem. 274:20287-20292 152. van Eekert MHA, Stams AJM, Field JA & Schraa G (1999) Gratuitous dechlorination of chloroethanes by methanogenic granular sludge. Appl. Microbiol. Biotechnol. 51:46-52 153. van Eekert MHA, Schröder TJ, Stams AJM, Schraa G & Field JA (1998) Degradation and fate of carbon tetrachloride in unadapted

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methanogenic granular sludge. Appl. Environ. Microbiol. 64:2350-2356 154. van Eekert MHA, Schröder TJ, van Rhee A, Stams AJM, Schraa G & Field JA (2001) Constitutive dechlorination of chlorinated ethenes by a methanol degrading methanogenic consortium. Biores. Technol. 77:163-170 155. Vannelli T, Studer A, Kertesz M & Leisinger T (1998) Chloromethane metabolism by Methylobacterium sp. strain CM4. Appl. Environ. Microbiol. 64:1933-1936 156. Vannelli T, Messmer M, Studer A,Vuilleumier S & Leisinger T (1999) A corrinoid-dependent catabolic pathway for growth of a Methylobacterium strain with chloromethane. Proc. Natl. Acad. Sci. USA 96:4615-4620 157. Vogel TM & McCarty PL (1985) Biotransformation of tetrachloroethylene to trichloroethylene, dichloroethylene, vinyl chloride, and carbon dioxide under methanogenic conditions. Appl. Environ. Microbiol. 49:1080-1083 158. Vogel TM & McCarty PL (1987) Abiotic and biotic transformations of 1,1,1-trichloroethane under methanogenic conditions. Environ. Sci. Technol. 21:1208-1213 159. Weiner JH, Bilous PT, Shaw GM, Lubitz SP, Thomas GH, Cole JA & Turner RJ (1998) A novel and ubiquitous system for membrane targeting and secretion of cofactor-containing proteins. Cell 93:93-101 160. Wickramanayake GB, Gavaskar AR, Alleman BC & Magar VSE (Eds) (2000) Bioremediation and phytoremediation of chlorinated and reclacitrant compounds. Battelle Press, Columbus, Ohio. 161. Wiegel J, Zhang X & Wu Q (1999) Anaerobic dehalogenation of hydroxylated polychlorinated biphenyls by Desulfitobacterium dehalogenans. Appl. Environ. Microbiol. 65:2217-2221 162. Wild A, Hermann R & Leisinger T (1996) Isolation of an anaerobic bacterium which reductively dechlorinates tetrachloroethene and trichloroethene. Biodegradation 7:507-511 163. Wohlfarth G & Diekert G (1997) Anaerobic dehalogenases. Curr. Op. Biotechnol. 8:290-295 164. Yang Y & McCarty PL (1998) Competition for hydrogen within a chlorinated solvent dehalogenating anaerobic mixed culture. Environ. Sci. Technol. 32:3591-3597

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Chapter 6 BIODEGRADATION OF CHLORINATED COMPOUNDS BY WHITE ROT FUNGI JAMES A. FIELD Department of Chemical and Environmental Engineering, University of Arizona, Tucson, AZ, USA

1. WHITE ROT FUNGI AS LIGNIN DEGRADERS Lignin is the second most abundant polymer (after cellulose) in the biosphere, accounting for 20 to 35% of the dry weight of wood. Lignin is a complex high molecular weight aromatic polymer formed from the random condensation of three different phenylpropanoid precursors, p-coumaryl alcohol, coniferyl alcohol and sinapyl alcohol (4-hydroxy-, 3-methoxy-4-hydroxy-, and 3,5-dimethoxy-4-hydroxy-phenylpropanol, respectively). Lignin behaves as a glue providing strength to fibers, serves as a hydrophobic impermeable seal across cell walls, and as a barrier to microbial attack (34). The hydrophobic and irregular structure of lignin renders the polymer inaccessible to hydrolytic enzymes. The only mechanism that can account for the initial attack of lignin is a non-specific extracellular oxidative process (79). White rot fungi constitute the most important group of microorganisms responsible for the biodegradation of nature's most complex polymer, lignin (39, 79, 136). White rot fungi are, for the most part, higher fungi with macroscopic fruiting bodies (e.g., mushrooms, conches etc.), belonging to the order Basidiomycetes. These organisms play a vital role in earth's global ecology, by secreting extracellular ligninolytic enzymes to bring about an unique oxidative erosion of the complex aromatic structure of lignin. The lignin polymer generally does not serve as a sole source of carbon and energy, rather it is cometabolized by white rot fungi utilizing carbohydrates or other readily assimilatable compounds. The physiological purpose of lignin degradation is to provide white rot fungi with better access to hemicellulose and cellulose in wood, which are the true primary substrates of the fungi. Ligninolytic metabolism is usually triggered by nutrient limitation, typically either nitrogen or carbon starvation (79, 109). The main components of the ligninolytic enzyme system are shown in Table 6.1. The most common lignin degrading enzymes are two closely related peroxidases, manganese peroxidase (MnP) and lignin peroxidase (LiP), which are glycosylated extracellular proteins with iron-bearing heme groups. As the name suggests, the peroxidases require as an electron acceptor in order to oxidize substrates (39, 53, 159). The main difference in the enzymes is in their binding sites. MnP has a specific manganese binding Dehalogenation: Microbial Processes and Environmental Applications, pages 159-204 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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site and MnP almost exclusively oxidizes manganese. Oxidized manganese or in turn causes a chemical oxidation of phenolic lignin (52). LiP can oxidize nonphenolic compounds (methoxybenzenes or R-O-phenyl moeities in lignin). LiP producing fungi biosynthesize the metabolite veratryl alcohol (3,4-dimethoxybenzene), which works in conjunction with LiP as a redox mediator to facilitate the oxidation of other compounds. In addition, laccases are copper-containing extracellular proteins that utilize as an electron acceptor to support substrate oxidation (6, 39). Laccases cause the oxidation of phenolic compounds. A variety of extracellular oxidases are also produced to supply the ligninolytic system with extracellular as shown in Table 6.1. The substrates for the oxidases include simple organic acids, aldehydes, and cellobiose produced during the degradation of wood. Many white rot fungi also produce biosynthetic aryl alcohols, including 3,5dichloro-p-anisyl alcohols which serve as recyclable substrates of aryl alcohol oxidases (AAO) (37, 40). Oxalic acid is an additional metabolite of white rot fungi that is implicated in the lignin degrading system (44, 83). Oxalic and other organic acid metabolites can complex oxidized manganese, assisting in the function of MnP (53). They are also precursors to radicals that eventually can generate reduced oxygen species, such as (77, 82). The main reaction mechanism of ligninolytic enzymes can be characterized as a oneelectron oxidation of aromatic moeities, as illustrated for lignin in Figure 6.1 (145, 159).

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The resulting cation radicals react spontaneously with oxygen, either from water or as elemental dioxygen, resulting in the formation of oxidized products. The presence of elemental oxygen is required for extensive oxidation (159). Due to their capacity to oxidize and modify lignin, white rot fungi and their oxidative enzymes are being considered as biocatalysts to develop clean technologies in the paper and pulp industry (4, 106,121). In addition to its natural substrate, lignin, a wide variety of aromatic priority pollutants are also susceptible to the non-specific oxidizing reactions caused by ligninolytic enzymes. Consequently, this has motivated research and development in the application of white rot fungi to bioremediation techniques for the degradation of persistent environmental pollutants (49, 111, 130). Included here is the use of white rot fungi for the degradation of organochlorine pollutants. It should be noted that extracellular ligninolytic enzymes are not always responsible for the organochlorine degradation, and it is therefore necessary to consider their use on a case by case basis, following adequate evaluation of their efficacy. Aside from their capacity to degrade chlorinated compounds, many white rot fungi are also capable of biosynthesizing chlorinated compounds. Therefore, before reviewing organohalogen degradation, a short overview of organochlorine production by these fungi will be addressed.

2. ORGANOCHLORINE PRODUCTION BY WHITE ROT FUNGI Organoholagens have both anthropogenic and natural sources. Recent surveys indicate that more than 1500 natural-occurring organochlorine compounds are known (56). The ecologically important group of Basidiomycetes are responsible for a

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significant production of the terrestrial sources. This group includes white rot fungi that produce significant quantities of de novo chlorinated metabolites. The predominant organochlorine metabolites produced by white rot fungi are chloromethane, chlorinated anisyl metabolites, chlorinated phenylpropanoid metabolites, and chlorinated hydroquinone methyl ethers (38, 50, 61). Typical examples of these metabolites are illustrated in Figure 6.2. Fungi are estimated to emit 160,000 tons of chloromethane to the atmosphere each year. The most prominent contributors are white rot fungi from the genus Phellinus, accounting for 86% of the fungal input (61,178). Commonly occurring white rot fungi from the genus Hypholoma produce the organochlorine metabolite, 3,5dichloro-p-anisyl alcohol, equivalent to 3% of their dry biomass weight (174). Average field-measured concentrations of this metabolite in wood and litter colonized by these fungi are (40). Presently, there is no clear evidence for a chlorinating enzyme from white rot fungi. The formation of chloromethane is assumed to result from a halide methyltransferase. Labelled methionine is incorporated into chloromethane when provided to Phellinus pomaceus, suggesting that S-adenosylmethionine (SAM) is the direct precursor. A SAMdependent methyl transferase isolated from the cell membranes of the fungus was able to methylate chloride, but the for chloride was very high (300 mM) and thus could not account for physiological chloromethane formation (144). The affinity for chloride may not be adequately represented by the disturbed conformation of the purified enzyme. The chlorinating enzymes involved in the formation of chloroaromatic metabolites are even more elusivé. Chloroperoxidase activity in Hypholoma spp. was

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extensively monitored, both in extracellular culture broth and in cell-free extracts of mycelium when the fungus was actively producing organohalogens. However, no activity was detected (160) suggesting that alternative means of chlorination are probably involved. Recently, evidence has been reported for novel and dependent halogenating enzymes in bacteria (173). The biosynthetic precurors of the chlorinated anisyl metabolites and chlorinated phenylpropanoid metabolites are better understood. Evidence based on labeling experiments indicates that the aromatic amino acid, phenylalanine, is the common precursor for these metabolites (110, 148). The scope of organohalogen metabolites production by some white rot Basidiomycetes indicates that their synthesis is purposeful. Indeed, several physiological functions have been proposed for the organohalogen metabolites. These physiological functions include antibiotic properties, metabolites involved in lignin degradation, and synthons for biosynthesis. An important physiological function attributed to organohalogens is their antibiotic properties. The antibiotic properties of drosophilin A were described, along with the first characterization of its metabolite (74). The antibiotic properties of chlorinated metabolites from Basidiomycetes have previously been reviewed (50). Further evidence of antifungal activity of chlorinated anisyl metabolites, supplied at 50 to has been reported (165). Apart from their antibiotic properties, chlorinated metabolites are also involved in lignin degradation, either as substrates for extracellular production or as redox mediators. Most white rot fungi that produce chlorinated anisyl metabolites concurrently produce extracellular aryl alcohol oxidases (AAO). This group of enzymes is responsible for the oxidation of aryl alcohols to aryl aldehydes, at the expense of reduction to (37). The chlorinated anisyl alcohols are much better substrates for AAO compared to their structurally similar, non-chlorinated secondary metabolite counterparts, veratryl alcohol and p-anisyl alcohol. Aryl aldehydes produced from the extracellular oxidation are recycled to the corresponding alcohols by intracellular NADPH-dependent aryl alcohol dehydrogenases. This process generates a physiological cycle for continuous production. Both veratryl alcohol and p-anisyl alcohol are substrates of lignin peroxidase, whereas the chloro group of chlorinated anisyl metabolite (CAM) alcohols renders the aryl alcohol resistant to lignin peroxidase, conserving the molecule for the producing redox cycle (37). The fungal metabolite 2-chloro-1,4-dimethoxybenzene (2CDMB) has been found to act as a redox mediator for LiP in the same fashion as veratryl alcohol. The presence of sub-stoichiometric concentrations of 2CDMB enable lignin peroxidase to oxidize various substrates which otherwise are not oxidized very well or not at all, replacing the function ofveratryl alcohol (162-164). The evidence indicates that 2CDMB is oxidized to a stable cation radical and the cation radical oxidizes the terminal substrate. The occurrence of the cation radical has been demonstrated with an electron paramagnetic resonance spectrometer (164). Additionally, the presence of terminal substrates was shown to decrease the conversion of 2CDMB by lignin peroxidase, which would be expected if the cation radical was reduced back to 2CDMB upon reaction with the terminal susbstrate (162). Finally, organochlorines can function as synthons in biosynthesis. Chloromethane has been shown to be readily utilized by several white rot fungi as a methyl donor for the biosynthesis of veratryl alcohol. Laboratory studies have shown that deuterium-

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labeled chloromethane was incorporated as methoxy groups in veratryl alcohol. This compound, as was already mentioned, is an important secondary metabolite of white rot fungi (61, 62). Additionally, chloromethane is the source of methyl groups in the formation of methyl ester metabolites, such as benzoate methyl ester, from the genus Phellinus (61).

3. BIODEGRADATION OF ORGANOCHLORINES BY WHITE ROT FUNGI White rot fungi are capable of degrading a wide range of chlorinated priority pollutants. Interest in their use for bioremediation was initiated due to the non-specific lignin degrading enzyme system these fungi possess. The extracellular ligninolytic system is well known for its oxidative reactions (79, 159). However, the same enzyme system can generate radicals from organic acids, which can also behave as powerful reducing agents (13, 30, 31, 77, 134). Thus, the extracellular enzymes are responsible for both oxidation and reduction reactions. Also, white rot fungi have many of the same intracellular xenobiotic degrading enzymes found in many other fungi, such as monooxygenase activity from the cytochrome system (86, 101, 114). The ligninolytic system in the most well-studied model white rot fungus, Phanerochaete chrysosporium, is strongly regulated by nutrient nitrogen. Expression of the ligninolytic enzymes is triggered by nutrient-nitrogen, i.e., bioavailable nitrogen, limitation (79). Thus, a common method of determining the dependence of chlorinated pollutant degradation on the ligninolytic enzymes is to compare degradation in lownitrogen and high-nitrogen culture media. Also, manganese induces expression of MnP (39, 108), therefore a high-manganese culture medium can be used to determine the importance of MnP in a particular degradation scheme.

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Degradation of chlorinated priority pollutants by white rot fungi is always due to cometabolism. In most of the experiments covered in this chapter, the primary substrates are simple sugars (e.g., glucose) or polysaccharides, either in a purified form such as cellulose or as natural lignocellulosic substrates (e.g., wood chips, straw). Among researchers of white rot fungi, these primary substrates are referred to as the "cosubstrate". Therefore, unlike bacteria, white rot fungi cannot be enriched on organohalogen pollutants.

3.1. Biodegradation Of Simple Chlorinated Alkanes and Alkenes The white rot fungus, Phanerochaete chrysosporium, is known to degrade a variety of chlorinated solvents, such as trichloroethene (TCE) and carbon tetrachloride, as shown in Table 6.2 TCE is extensively mineralized to however the two studies which have documented this finding, have conflicting results with respect to the nutrient nitrogen regime required. Khindharia et al. (78) found that ligninolytic conditions at low nitrogen were required for extensive mineralization of both TCE and carbon tetrachloride, suggesting the involvement of ligninolytic enzymes. In contrast, Yadav et al. (179) found that nutrient-rich malt extract culture medium was required for extensive mineralization of TCE, indicating that the involvement of ligninolytic enzymes was not likely. In any case, the research group of Aust and coworkers has demonstrated that lignin peroxidase incubated in vitro with the physiological metabolites, veratryl alcohol and oxalate, caused reduction of the chlorinated solvents (78, 147). The incubation results in the oxidation of oxalic acid to and formate anion radical. The formate anion radical causes the reductive dehalogenation of the chlorinated solvents, such as the conversion of carbon tetrachloride to chloroform radical, as shown in Figure 6.3. In a similar fashion, cellobiose dehydrogenase (CDH) can cause the reduction of

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ferric iron to ferrous iron, which can generate hydroxyl radicals that can oxidize oxalate to formate anion radicals. CDH incubated with Fe(III), oxalate and hydrogen peroxide was shown to reductively dehalogenate bromotrichloromethane to form the chloroform radical (24). Similar degradation patterns have been observed with carbon tetrachloride, when the formate anion radical was generated by photolysis of the Fe(III)-oxalate

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complex (69). Although chloromethane is produced by some white rot fungi (see Section 2 on organochlorine biosynthesis), chloromethane can also be utilized by nonchloromethane producing white rot fungi. The evidence of its utilization is the conversion of deuterium-labeled chloromethane to the methoxy groups of the secondary metabolite, veratryl alcohol in P. chrysosporium (62).

3.2. Biodegradation of Chlorinated Heterocyclic Insecticides The evidence for biodegradation of chlorinated heterocyclic insecticides by white rot fungi is shown in Table 6.3. Among these compounds, significant mineralization has only been observed for lindane and chlordane. The mineralization of labeled dieldrin and aldrin is so low that the mineralization could potentially be due to radiolabeled impurities. The only confirmed bioconversion of these latter compounds is the formation of dieldrin resulting from the epoxidation of a double bond between two carbons in aldrin (76). This reaction, however, does not result in any dechlorination of the substrates. Also, endosulfan metabolism is limited to the oxidation of a sulfur-containing side chain, without any dechlorination (85). Lindane is degraded by many different white rot fungal strains. Aside from three dechlorinated metabolites have been identified and these are: tetrachlorocyclohexene, tetrachlorocyclohexene epoxide; tetrachlorocyclohexenol (116, 150, 151). Tetrachlorocyclohexene was also a substrate for P. chrysosporium, yielding tetrachlorocylohexanol as a metabolite (116). Lindane metabolism is not associated with the production of ligninolytic enzymes, and ligninolytic enzymes do not directly biotransform lindane. Rather, the lindane-degrading activity of P. chrysosporium can be inactivated by 1-aminobenzotriazole (a specific cytochrome inhibitor) (116). This evidence, together with the product spectrum of lindane metabolism, suggests the involvement of monooxygenase activity of cytochromes as the initiating enzymes for lindane degradation by white rot fungi. Laboratory experiments have demonstrated that lindane biodegradation in soil by the natural microbiota can be increased by inoculating the soil with P. chrysosporium (115). In the cited study, the mineralization of lindane increased from 21.6% to 49.1%, due to inoculation of P. chrysosporium. Moreover, the white rot fungus did not significantly mineralize lindane in sterile soil (only 0.7%), so that the combined activity of soil microbiota and the white rot fungus accounted for the extensive mineralization.

3.3. Biodegradation of a Chlorinated Organophosphorus Insecticide Chlorpyrifos is the only chlorinated organophosphorus pesticide which has been tested for biodegradation by a white rot fungus. This insecticide contains an aromatic pyridine moiety, bearing three chloro-groups. In one study, P. chrysosporium mineralized labeled chlorpyrifos by 28% (Table 6.3). Results further demonstrated that the chlorinated pyridinyl ring of chlorpyrifos undergoes cleavage during biodegradation by the fungus (21).

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3.4. Biodegradation of a Chlorinated Triazine Herbicide, Atrazine Several studies have investigated the biodegradation of the chlorinated triazine herbicide, atrazine (Table 6.4). Two white rot fungi, P. chrysosporium and Pleurotus pulmonarius, were shown to convert atrazine to several N-dealkylated metabolites (99, 102, 114). These metabolites are desethylatrazine, desisopropylatrazine, and desethyldesisopropylatrazine. Also, one hydroxylated alkyl side chain metabolite, hydroxyisopropylatrazine, was identified by the activity of Pleurotus pulmonarius (99, 102).

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Since all of these metabolites still contain the chloro group, the white rot fungi were not able to dechlorinate atrazine. The observed bioconversions were not caused by purified ligninolytic peroxidases (114). The conversion was localized in mycelium that contained significant amounts of microsomal cytochrome activity. Specific inhibitors, either piperonyl-butoxide or 1-aminobenzotriazole, effectively inhibited the atrazine biotransformation (99, 114). Manganese supplements increased atrazine biotransformation in Pleurotus pulmonarius, but this observation does not necessarily implicate the involvement of MnP, since cytochrome activity also increased in response to manganese (99). In several experiments, the combined activity of the natural microbiota and white rot fungi has been evaluated in non-sterile soils or in non-sterile spent straw from Pleurotus mushroom cultivation. In both cases, there was insignificant conversion of -labeled atrazine to (46, 64, 100). Instead, a large fraction of the label was converted to non-extractable, bound residues (64, 100).

3.5. Biodegradation of Chloroacetanilide Herbicides Chloroacetanilide herbicides are phenyl-N-alkylated structures with chloro-groups on the aliphatic side chains. Evidence for their biodegradation by white rot fungi is shown in Table 6.4. Two white rot fungal strains (Ceriporiopsis subvermispora and Phlebia tremellosa) were found to mineralize -labeled alachlor to by 12 to 14%, after 122 days of incubation on malt extract medium supplemented with wood (48). The chlorine moiety of metachlor was mineralized by 30% to in16 days by P. chrysosporium (97). Seven dechlorinated metabolites were identified, which together accounted for 28% of the initial The three major metabolites (metabolites D, G and I) are shown in Figure 6.4.

3.6. Biodegradation of Chlorophenoxyacetate Herbicides Chlorophenoxyacetate herbicides are phenyl-O-alkylated structures with chlorogroups on the ring structure. Both of the widely used members of this herbicide group, 2,4-dichlorophenoxyacetate (2,4-D) and 2,4,5-trichlorophenoxyacetate (2,4,5-T), are extensively mineralizedby white rot fungi, as shown in Table 6.4. Based on experiments in a high nitrogen medium, in which labeled 2,4-D and 2,4,5-T were mineralized by P. chrysosporium, it was concluded that chlorophenoxyacetate herbicide degradation

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does not depend on the ligninolytic enzymes (43, 181, 182). This hypothesis was confirmed further by observing robust mineralization of 2,4-D and 2,4,5-T with a peroxidase-negative mutant of P. chrysosporium (181, 182). However, studies with Dichomitus squalens indicate that, while the mineralization of the side chain (acetoxy group) is not dependent on ligninolytic metabolism, the mineralization of the 2,4,5-T ring is dependent on ligninolytic metabolism (135). The latter activity (ring metabolism) was greatest in supplemented cultures, which provides strong evidence for the induction of MnP by and points to the involvement of MnP in ring degradation. Similarly, the intermediate 2,4-dichlorophenol was identified as a metabolite from 2,4-D metabolism (135). The involvement of MnP in the initial oxidation of 2,4dichlorophenols has been confirmed in another study (171). A known product of 2,4dichlorophenol oxidation by MnP, 2-chloro-p-benzoquinone, was also identified during 2,4-D metabolism by Dichomitus squalens (135). A unique feature of 2,4-D degradation by Dichomitus squalens was the formation of a xyloside from the 2,4-dichlorophenol intermediate (135). In these studies, crude cell extracts of the mycelium had activity that could cleave the gylocisidic bond. The xylosylation mechanism is suggested to be a detoxification strategy, allowing the fungus to accumulate intermediates without deleterious effects. Additionally, the xyloside is a protecting group that prevents the polymerization of the chlorophenol intermediates. A scheme indicating the general pathway of 2,4-D degradation by Dichomitus squalens is shown in Figure 6.5.

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3.7. Biodegradation of Chlorinated Benzenes, Polychlorinated Biphenyls (PCBs) and Diphenyl Ethers The white rot fungus, P. chrysosporium has also been shown to partially mineralize radiolabeled mono- and di-chlorobenzenes to (Table 6.5). The metabolism of chlorobenzene by P. chrysosporium was found to be very limited in a low nitrogen medium, and greatly improved in high nitrogen, indicating that ligninolytic enzymes were most likely not involved (183). Another, more highly chlorinated compound, hexachlorobenzene, has been shown to be partially eliminated from soil by a Lentinus isolate (104), but no information on mechanism was provided. Polychlorinated biphenyls (PCBs) are also metabolized by various white rot fungi (Table 6.5). As a general rule, the extent of PCB mineralization by white rot fungi decreases with increasing chlorine number. For example, the low levels of mineralization reported for hexa- and tetrachlorobiphenyls (0.4 to 1.4%, respectively), do not provide very conclusive evidence for their mineralization, since radiolabeled impurities could account for the low levels of observed mineralization. Only in one study, where very low concentrations of 2,4,2',4'-terachlorobiphenyl were utilized, did mineralization increase to more demonstrable levels (9.6%) (166). Radiolabeled tri-, di-, and mono- chlorinated biphenyls are more extensively mineralized to with values as high as 11 to 16% being reported (14, 42, 166). Similar trends have also been observed with technical PCB mixtures. For example, P. chrysosporium decreased PCB concentrations of Aroclor 1242, 1254, and 1260 (42, 54 and 60% chlorinated) by 60.9, 30.5, and 17.6%, respectively (180). Congeners of lower chlorine number were degraded more extensively compared to those of higher chlorine number during the remediation of the technical mix Delor103 by Pleurotus ostreatus (84). Bioavailability has been also found to be important. For example, mineralization of 2,4’,5-trichlorobiphenyl by Trametes versicolor significantly increased from 7% to 19% by supplementing the culture with the surfactant Triton-X-100 (15). In some cases of the more highly chlorinated PCBs, such as terra-, penta-, and hexa- chlorinated congeners, white rot fungi appear to play a role in their degradation (14, 180, 186), although the mechanism of removal is not clear. A large fraction of the PCB-label is found bound to mycelium, which may result from either covalent bonding or simple phase-partitioning of unaltered PCBs (42, 45, 166). Little information is available on the types of metabolites formed during the degradation of PCB by white rot fungi. Two metabolites, 4-chlorobenzoic acid and 4chlorobenzyl alcohol have been observed during the degradation of 4,4’dichlorobiphenyl by P. chrysosporium (42), however, there is little evidence for any involvement of ligninolytic enzymes in PCB degradation by white rot fungi. In general, PCB degradation proceeds under high nutrient nitrogen conditions (45, 81, 180). Severe repression of PCB degradation would be expected if ligninolytic metabolism was involved, at least in the case of P. chrysosporium, due to its requirement for nitrogenpoor conditions. In the screening of various white rot fungi, no correlation has been observed between ligninolytic enzyme titers and PCB mineralization (166). While chlorinated biphenyls were not subject to oxidation by ligninolytic enzymes, hydroxylated chlorinated biphenyls are readily oxidized by laccase (146). The products of the oxidation are dimers from the coupling of two hydroxybiphenyls, either by C-C or C-O-C bonds. At least one of the dimers formed from 3-chloro-4-hydroxy-biphenyl

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and 5-chloro-2-hydroybiphenyl was completely dechlorinated, indicating the occurrence of dehalogenation during the laccase-mediated coupling reaction. In addition to PCB, related chlorodiphenyl ether compounds are also subject to biotransformation by white rot fungi (Table 6.5). In one study, Triclosan (2,4,4’trichloro-2’-hydroxydiphenylether), an antimicrobial compound used in soaps and deodorants, was either cleaved to 2,4-dichlorophenol or xylosylated and glucosylated by Trametes versicolor, and additionally methylated by Pycnoporus cinnabarinus in the hydroxy position (68). However, none of these metabolites resulted from dechlorination. A similar compound, 4-chlorodiphenyl ether, was shown to be metabolized to several hydroxylated metabolites by Trametes versicolor, including metabolites where the nonhalogenated ring is cleaved, as well as 4-chlorophenol (67). Also, no dechlorinated products were observed with this diphenyl ether. The metabolism of 4-chlorobiphenyl ether has been attributed to cytochrome activity, based on metabolic inhibition of by 1-aminobenzotriazole. In this study, extracellular fluids with laccase and MnP activity were not able to convert the unhydroxylated diphenyl ethers (67).

3.8. Biodegradation of Chlorinated bis(Phenyl)Ethanes The insecticide 1,1,1 -trichloro-2,2-bis(p-chlorophenyl)ethane, (DDT), is a persistent environmental pollutant that has been a major concern due to its uptake in the food chain. White rot fungi have been shown to biotransform and partially mineralize DDT (Table 6.6). The extent of DDT mineralization by cultures of various white rot fungi has been reported to range from 5.3 to 13.5%, after 30 days in liquid cultures with glucose as the primary substrate in the medium (20). The highest levels of mineralization were observed with P. chrysosporium. When the culture conditions were optimized utilizing cellulose as the primary substrate instead of glucose, the extent of DDT mineralization reached 32% after 60 days (47). Also, high levels of mineralization (30%) were achieved by reducing the DDT concentration in the medium to 0.01 mg/1 (47). Results from these studies clearly indicate the potential of ligninolytic fungi to significantly biodegrade During the degradation of DDT by P. chrysosporium, several intermediates have been identified (Table 6.6), which indicate metabolism at the or position of the ethane group. Only one metabolite, l,l-dichloro-2,2-bis(4chlorophenyl)ethane (DDD), is formed immediately during primary metabolism. Several oxidized intermediates, 2,2,2-trichloro-1,1-bis(4-chlorophenyl)ethanol (dicofol), 2,2dichloro-1,1-bis(4-chlorophenyl)ethanol (FW-152), and 4,4’-dichlorobenzophenone (DBP) are formed, but only after a 6 day lag period (20, 23). In addition to the aforementioned intermediates, another dechlorinated biotransformation product, 1,1dichloro-2,2-bis(4-chlorophenyl)ethylene (DDE) has been identified during the degradation of DDT by P. chrysosporium, in a soil with corn as the primary substrate (47). The formation of DDE also has been observed in autoclaved cultures, indicating that its formation may be due to abiotic processes (73). These intermediates emerge transiently and disappear again, indicating that they are metabolized further. This indication was further proven for dicofol and DDE. When supplied as labeled compounds to P. chrysosporium, they were directly mineralized to (Table 6.6). The mineralization of DDT and DDE by P. chrysosporium requires ligninolytic culture conditions, created by utilizing a nitrogen-limited medium (20, 22, 23).

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Conversely, mineralization of DDT is insignificant when culture conditions are not nitrogen-limited. Likewise, the conversion of DDT to oxidized intermediates, such as dicofol, FW-152, and DBP occurrs only in nitrogen-limited medium, and only after a lag-phase of several days which corresponds to the onset of ligninolytic enzyme production (20). Other reports further indicate that the addition of LiP inhibitors to P. chrysosporium cultures inhibits DDT mineralization (11). While the data seem to infer the involvement of LiP in the degradation of DDT, there is no report of purified LiP directly oxidizing DDT (12). An indirect mechanism involving LiP might be implicated, such as the dechlorination of chlorinated solvents by LiP incubated together with

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mediators veratryl alcohol and oxalic acid (78). It also has been suggested that DDT and dicofol have suitable reduction potentials, so as to be reductively dechlorinated by the formate anion radical produced by the LiP/veratryl alcohol/oxalic acid system. This may account for the formation of DDD from DDT, or the formation of FW-152 from dicofol (78). While the suggestion may be valid for the conversion of dicofol to FW-152, the suggestion is not consistent with the conversion of DDT to DDD, prior to the onset of lignin metabolism. In contrast to DDT mineralization, DDT biotransformation is not affected by nitrogen nutrients. The biological removal of DDT by P. chrysosporium has been observed in nitrogen-sufficient medium, where no LiP is produced (73,80), This has led one author to conclude that DDT metabolism was not due to LiP activity (80). However, in another report, the conversion of DDT to DDD was recognized as a non-ligninolytic conversion (20). Thus, the occurrence of DDT metabolism under culture conditions in which no LiP is formed does not disprove the involvement of LiP in the oxidation of DDT to The proposed pathway DDT metabolism by white rot fungi is shown in Figure 6.6. DDT is either initially dechlorinated by reductive dechlorination to DDD, or by abiotic dehydrochlorination to DDE. A more predominant initial pathway is through direct hydroxylation of DDT to form dicofol. The bond of dicofol can be cleaved abiotically under mild chemical conditions to DBP, or it may be reductively dechlorinated by the fungus to yield FW-152. Also, a hydroxylation of DDD can yield FW-152. The FW-152 metabolite can be further oxidized to DBP in several steps, involving dechlorination and oxidation of the dichloromethyl group to carboxylic acid that is decarboxylated. Finally, DBP is the last metabolite prior to ring cleavage and mineralization (20, 22). The methoxylated analogue of DDT, 1,1,1 -trichloro-2,2-bis(4-methoxyphenyl)ethane (methoxychlor), is more readily mineralized by P. chrysosporium, compared to DDT. Reports indicate that up to 54% of is mineralized to (Table 6.6). Similar to DDT metabolism, biotransformation products that are hydroxylated on dechlorinated on or exhibiting both modifications, have been observed (57). Moreover, these studies have shown that three biotransformation products are readily mineralized by P. chrysosporium, indicating that they are true intermediates of methoxychlor biodegradation to (57).

3.9. Biodegradation of Chlorinated Phenols The conversion of chlorinated phenols by white rot fungi has been extensively studied. Unlike many of the chlorinated compounds considered so far, chlorinated phenols are truly direct substrates for ligninolytic enzymes of white rot fungi. Many other types of enzymes from white rot fungi are also capable of transforming chlorinated phenols. Tables 6.7 through 6.10 summarize the literature data for the biodegradation penta-, terra-, tri-, di-, and mono-chlorinated phenols, respectively. In whole culture, chlorinated phenols are extensively mineralized to and chloride by white rot fungi. The extent of (PCP) mineralization PCP-organochlorine to chloride has been reported to be around 40% (Table 6.7). The extent of mineralization to has been reported upwards to 70%, and the

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mineralization of various isomers to has been reported to be approximately 60%, and the mineralization of organochlorine in 2,4,6-trichlorophenol to chloride has been reported to be nearly 100% (Table 6.8). The only dichlorophenol tested to date has been and it was mineralized 50% to (Table 6.9). Another isomer, 3,4-dichlorophenol was tested for mineralization of organochlorine, and showed a 15% release as chloride (Table 6.9). The only mineralization result available for monochlorinated phenols is the 94% conversion of the organochlorine in 2-chlorophenol to chloride (Table 6.10). There are number of different mechanisms involved in the biotransformation and mineralization of chlorinated phenols by white rot fungi. These can be categorized according to the enzymes or biochemical conversions involved. The most important mechanisms are as follows: 1) oxidation by ligninolytic enzymes; 2) methylation (and glycosylation); 3) benzoquinone reduction; 4) hydroxylation; and 5) reductive dehalogenation.

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3.9.1. Oxidation of Chlorinated Phenols by Ligninolytic Enzymes The importance of ligninolytic enzymes in chlorophenol degradation by the white rot fungus, P. chrysosporium is reflected by the great impact of nitrogen-limited culture conditions. Nitrogen-limitation is generally found to greatly improve chlorophenol removal, as well as its mineralization (112, 131, 133, 171). However, some white rot fungi that are less nitrogen-regulated, degrade chlorophenols quite well in nitrogensufficient culture conditions (95, 118). The best evidence for the involvement of ligninolytic enzymes is their ability to directly oxidize chlorinated phenols. Tables 6.7 to 6.10 show examples of the direct oxidation of a wide variety of chlorinated phenols by LiP, MnP, manganese independent peroxidase (MIP), and laccase. The chlorinated phenols susceptible to ligninolytic enzyme oxidation range from penta- to monochlorinated phenols. Since oxidized manganese species are responsible for the terminal reactions carried out by MnP, the oxidation of chlorophenols by or are indicative of the capacity of MnP. The data summarized in Tables 6.8 through 6.10 indicate that these oxidized manganese species also are implicated in the oxidation of tri- to monochlorinated phenols. Products from the oxidation of chlorinated phenols by ligninolytic enzymes vary from the formation of quinones to polymerized products (Table 6.7). The classic reaction of chlorinated phenols with ligninolytic enzymes results in the formation of 1,4benzoquinone, where the chlorine in the 4-position is dechlorinated (60, 70). The formation of chlorinated 1,4-benzoquinones from the oxidation of penta-, tri-, and dichlorophenols, as well as 2-chlorophenol, by ligninolytic enzymes and oxidized

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manganese species is evident from many of the examples presented in Tables 6.7 to 6.10. Additionally, the formation of 1,4-benzoquinone from 4-chlorophenol by and the formation of 2-methoxy- 1,4-benzoquinone from 4-chloroveratrole by LiP have been reported (Table 6.10). Evidence for the 4-dechlorination is also evident from the release of chloride when 4-substituted chlorophenols are incubated with ligninolytic enzymes or oxidized manganese species (Tables 6.7-6.10). In some cases, the release of chloride is found to be stoichiometric with dechlorination of the 4-substituted chlorogroup (29, 60, 139). Based on labeling experiments with the 4-dechlorination mechanism has been shown to be due to a nucleophilic addition of water to a cyclohexadienone cation intermediate, as shown in Figure 6.7 (70, 72). Alternatively, it has been suggested that the 4-dechlorinated chloro-1,4-benzoquinone products are artifacts of the chemical decomposition of chlorophenolic dimers in organic solvent (often used to extract the the chlorinated benzoquinones for analysis). For example, the pentachlorophenyl dimer, 2,3,4,5,6-pentachloro-4-pentachlorophenoxy-2,5cyclohexadienone (PPCHD in Figure 6.8), formed by the oxidation of PCP by a plant peroxidase, has been shown to decompose chemically to tetrachloro-1,4-benzoquinone upon exposure to organic solvents (75). While this artifact cannot be ignored, the numerous studies detecting the release of chloride, where no extracting organic solvents have been utilized, would indicate that the direct formation of the 1,4-benzoquinones represents true products of ligninolytic enzymes (Tables 6.7-6.10). In one study, the direct oxidation of by MnP resulted in 30% mineralization to indicating that in some cases extensive oxidation can be achieved by ligninolytic enzymes (66). During the oxidation of chlorophenols by ligninolytic enzymes, polymers from the coupling of chlorophenol radicals are readily formed (3, 35, 63, 98, 152). The chlorophenols in the polymerized products are either joined by direct C-C bonds, or more commonly by ether bridges(C-O-C) (63, 98, 157, 175). Examples of the types of coupling products formed are illustrated in several dimers for which the structures have been elucidated (Figure 6.8). The tendency for polymerization decreases with increasing chlorine substitution (35); but even the most chlorinated species, such as PCP and drosophilin A (tetrachloro-4-methoxyphenol), can become polymerized (140,161,170). During treatment of chlorinated phenols by white rot fungi in soils, there is a tendency for chlorophenols to become covalently linked with humus. Labeling experiments have demonstrated the incorporation of PCP into different humus fractions. In soil culture experiments with various white rot fungi, 16 to 44% of was recovered as radioactivity in the fulvic acid, humic acid and humin fractions (141). In another study, the polymerization of 2,4-dichlorophenol by a plant peroxidase was carried out in the

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presence of humic acid, and experiments revealed covalent linkages of the chlorophenol with the humus (63). The formation of chlorophenols bound to humus can also be explained by the copolymerization of chlorophenols with nonhalogenated phenolics (e.g., catechol, guaiacol, ferulic acid, coniferyl alcohol, syringic acid) by ligninolytic enzymes (17, 26, 123, 138, 140). During the treatment of PCP in soils, a large fraction ofthe chlorophenol becomes copolymerized and incorporated into humus (bound residue), and helps to explain why mineralization in soil is often far less than that achieved in liquid cultures (Table 6.7).

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3.9.2. Methylation (and Glycosylation) of Chlorophenols Another major biotransformation observed during the metabolism of chlorophenols is the methylation of the phenolic hydroxy-groups. The conversion of PCP to pentachloroanisole (PCA) has been reported in many studies (Table 6.7). Also, the conversion of 3,4- and 2,4- dichlorophenols to their respective anisoles has been reported (Table 6.9). PCA does not appear to be a dead-end metabolite since it is mineralized by white rot fungi (Table 6.7). Methylation is also observed during the degradation of chlorinated-1,4-hydroquinones. P. chrysosporium converts tetrachloro1,4-hydroquinone to drosophilin A (DA, tetrachloro-4-methoxyphenol) and drosophilin A methyl ether (DAME, tetrachloro-4-dimethoxybenzene) and similar conversions have also been observed with trichloro-, dichloro- and monochloro- 1,4-hydroquinones, as shown in Tables 6.8, 6.9, and 6.10. Likewise, P. chrysosporium has been reported to convert chlorocatechol to chloroveratrole (Table 6.10). Methylation does not necessarily have to be the initial biotransformation step. For example, 3,4-dichlorophenol is converted to 4,5-dichloroguaiacol and 4,5-dichloroveratrole (Table 6.9), which results from hydroxylation followed by methylation (41). Also, 2,4-dichlorophenol is converted to 2CDMB (Table 6.9), a major product from oxidative 4-dechlorination, followed by 1,4-benzoquinone reduction and subsequent methylation of the resulting 2-chloro-l,4hydroquinone (171). The purpose of methylation is not fully understood, but methylation can serve to detoxify chlorophenols (88), as well as protect chlorophenols from oxidative coupling. Also, methylation is a key biotransformation step in the biosynthesis of important secondary metabolites of white rot fungi, such as the formation of veratryl alcohol (61) and chlorinated anisyl metabolites (110). Thus, methylation of chlorophenols can also be viewed as a fortuitous reaction that occurs when these compounds are accidentally utilized as precursors for secondary metabolite biosynthesis (110). The final step in the biosynthesis of veratryl alcohol involves methylation by Sadenosylmethionine dependent methyl transferases (SAM-MT), or chloromethanedependent methyl transferases (61). From white rot fungi, only SAM-MT have been isolated and characterized. A SAM-MT isolated from P. chrysosporium has been shown to readily convert 2,4-dichlorophenol to 2,4-dichloroanisole (33). Aside from methylation, chlorophenolic hydroxy-groups can also be glycosylated by sugar moeities. Dichomitus squalens has been shown to convert 2,4-dichlorophenol to xylosylated 2,4-dichlorophenol (135), illustrated in Figure 6.5. In a similar fashion, the free hydroxy-group of triclosan (2,4,4’-trichloro-2’-hydroxydiphenyl ether) is xylosylated and glucosylated by Trametes versicolor and Pycnoporous cinnabarinus (68).

3.9.3. Benzoquinone Reduction during Chlorophenol Degradation The oxidation of chlorophenols and their methylated biotransformation products by ligninolytic enzymes results in the formation of benzoquinone intermediates, especially chlorinated 1,4-benzoquinones. For continued degradation, these benzoquinone intermediates must be reduced to their corresponding hydroquinones. The hydroquinones are then subject to all of the other major biotransformations of chlorophenols (oxidation by ligninolytic enzymes, methylation, glycosylation,

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hydroxylation and reductive dehalogenation). The reduction of chlorinated 1,4benzoquinones is clearly an intracellular process carried out by the mycelium. Studies with washed mycelial preparations indicate a ready and nearly stoichiometric conversion of chlorinated 1,4-benzoquinones to their corresponding hydroquinones (71,131,133). Only marginal reduction appears to occur in the extracellular culture broth (71, 131). Intracellular enzymes responsible for the reduction of 1,4-benzoquinones are also known to occur in white rot fungi. A benzoquinone reductase (BQR) from the mycelium of P. chrysosporium has been isolated and characterized (18, 19). The BQR is a flavin mononucleotide-containing protein that is dependent on NADH or NADPH as an electron donor. However, the chemical reaction between NADH and various chlorinated 1,4-benzoquinones (e.g., 2-chloro-, 2,6-dichloro- and tetrachloro-) is so rapid that it suggests that the enzyme may not even be required to mediate their reduction by the mycelium (18). A cell free extract of P. chrysosporium, filtered through an ultrafiltration membrane to remove proteins, readily reduced 2-chloro-1,4-benzoquinone to 2-chloro1,4-hydroquinone (18).

3.9.4. Hydroxylation of Chlorophenols While hydroxy-groups can be introduced into chlorophenols through the combined activity of ligninolytic enzymes to yield benzoquinones, followed by benzoquinone reduction, there are also intracellular enzymes responsible for hydroxylating chlorophenols. The direct evidence for intracellular enzymes involved in the hydroxylation of chlorophenols is the conversion of 3,4-dichlorophenol to 4,5dichlorocatechol, observed in cell free extracts (CFE) prepared from mycelium of P. chrysosporium. According to one study, hydroxylase activity in the CFE is dependent on the presence of NADPH (41). The hydroxylase enzyme has been partially purified and found to be a dimeric protein. The only other aromatic hydroxylase purified from a white rot fungus, Pleurotus pulmonarius, is a benzo[a]pyrene hydroxylase, which is a microsomal cytochrome Otherwise, the involvement of hydroxylases are inferred from the conversion of chlorinated-1,4-hydroquinone substrates to hydroxylated metabolites by whole cultures of P. chrysosporium (131, 133).

3.9.5. Reductive Dehalogenation of Chlorophenols Because white rot fungi have generally been recognized for their ability to oxidize pollutants, the discovery that such fungi mediate reductive dehalogenation of chlorinated 1,4-hydroquinone substrates was surprising. This phenomenon was first observed during studies that unraveled the pathways used by P. chrysosporium to degrade 2,4,6trichlorophenol and pentachlorophenol (131, 133). Examples of these reductive dehalogenations by whole cultures are shown for various chlorinated 1,4hydroquninones in Tables 6.8 through 6.10. Reductive dehalogenation is not an exclusive capacity of P. chrysosporium, since several different white rot fungi have been shown to metabolize 2-chloro-l,4,5-trihydroxybenzene to 1,4,5-trihydroxybenzene (Table 6.10). Recently, the enzymes involved in the reductive dehalogenation have been isolated and characterized (132, 134). The dehalogenation is carried out by a twocomponent enzyme system involving the formation and reduction of a glutathione

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conjugate of the chlorinated 1,4-hydroquinone substrate. The first component of the membrane-bound enzyme complex is a glutathione transferase (GST), which displaces a chloro-group and forms a glutathionyl conjugate of the dechlorinated 1,4hydroqunione. The glutathionyl conjugate is then reduced by a soluble enzyme known as glutathionyl conjugate reductase (GSCR). The reducing equivalents for the GSCR are provided by glutathione. A scheme illustrating the two enzymatic steps leading to the reductive dehalogenation of tetrachloro-1,4-hydroquinone is shown in Figure 6.9.

3.9.6. Pathways of Chlorophenol Degradation There are several pathways proposed for the degradation of chlorophenols by white rot fungi. In one case, chlorophenol degradation can be envisioned as cycles of extracellular ligninolytic oxidation, followed by intracellular reduction and methylation. The ligninolytic enzymes generate chlorinated benzoquinones, with chloro-groups that are displaced by the nucleophilic addition of Benzoquinone reduction by mycelium can generate hydroquinones which are substrates for LiP, MnP and laccases. Full methylation of polychlorinated hydroquinones to polychlorinated 1,4dimethoxybenzenes can temporarily protect chlorophenols from ligninolytic oxidation (and thus polymerization) and perhaps detoxify them. However, the lower chlorinated hydroquinone, 2CDMB, is a substrate of LiP and thus enters directly into the cycling. As the cycling proceeds, more and more hydroxy groups are introduced into the molecule, via the nucleophilic addition of Eventually, tetrahydroxybenzene is formed which is then subject to ring cleavage. This metabolic pattern has been proposed for 2,4-dichlorophenol and 2,4,5-trichlorophenol degradation by P. chrysosporium (71, 171). A scheme depicting the proposed pathway for 2,4-dichlorophenol is shown in Figure 6.10. A second type of pathway is rationalized in which ligninolytic enzymes are only involved in the initial oxidation of the chlorophenol, generating a chlorinated 1,4benzoquinone intermediate dechlorinated in the 4-position. Thereafter, the degradation is postulated to be intracellular. The chlorinated 1,4-benzoquinone is converted to its corresponding chlorinated-1,4-hydroquinone by reduction. The chlorohydroquinone is than further decomposed by a network of hydroxylation or reductive dechlorination reactions. The hydroxylations are most likely carried out by an intracellular NADPH dependent hydroxylase (perhaps a monooxygenase). The reductive dehalogenations are carried out by GST and GSCR system. This pathway has been proposed for 2,4,6-

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trichlorophenol and pentachlorophenol (131, 133). A scheme depicting the proposed pathway for 2,4,6-trichlorophenol is shown in Figure 6.11.

3.10. Biodegradation of Chlorinated Dioxins Chlorinated dibenzo-p-dioxins and dibenzofurans are also found to be susceptible to degradation by the activity of white rot fungi (Table 6.11). The most important evidence is the biologically mediated elimination of chlorinated dioxins from white rot fungal cultures (156, 172). When 2,7-dichlorodibenzo-p-dioxin (25 mM) was incubated with P. chrysosporium under ligninolytic conditions (low N), 50% of the compound was removed in 4 weeks; little removal occurred in non-lignonolytic cultures (high N) (172). The only intermediates detected were 4-chlorocatechol and trihydroxybenzene (Table 6.11). LiP has been shown to directly oxidize the chlorinated dioxins, 2-chlorodibenzop-dioxin and 2,7-dichlorodibenzo-p-dioxin (59, 172). In these studies, chloride, 2chloro-1,2-benzoquinone, and 2-hydroxy-1,4-benzoquinone were the products identified from the LiP catalyzed oxidation of 2,7-dichlorodibenzo-p-dioxin (Table 6.11). The results indicate that LiP is able to cleave the dioxin structure. Label experiments with indicate that molecular oxygen is not the source of O in quinone cleavage products, leading to the hypothesis that O is incorporated by nucleophilic additions of water. The initiating reaction is most likely the formation of a cation radical, which has been demonstrated for the non-chlorinated dibenzo-p-dioxin (59). Extensive degradation of a mixture of 10 dioxin compounds (tetrachloro- to octochloro- dibenzo-p-dioxins and dibenzofurans) has been observed in experiments with either P. chrysosporium or P. sordida cultured in low nitrogen medium for 14 days

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compared to heat killed controls (156). In the study, the percent degradation ranged from approximately 40% (tetrachloro-) to 76% (hexachloro-) for dibenzo-p-dioxins, and from 45% (tetrachloro-) to 70% (hexachloro-) for dibenzofurans. During the metabolism of tetrachloro- and octochloro-dibenzo-p-dioxins, dichloro- and tetrachloro- catechols were identified as intermediates (Table 6.11). The mineralization of 2,3,7,8tetrachlorodibenzo-p-dioxin to has also been demonstrated, albeit the percent mineralization was low (Table 6.11). The dependence of degradation on low nitrogen and the direct oxidation by LiP, clearly indicates the involvement of ligninolytic enzymes in dioxin biodegradation by P. chrysosporium. Quinone products of the LiP-catalyzed cleavage of dioxin are reduced by the fungal mycelium yielding phenols and chlorinated catechols, which are further metabolized in accordance with the pathways available for chlorophenols (172).

3.11. Biodegradation of Chloroanilines Chlorinated anilines are important xenobiotic residues released during the metabolism of phenylurea, N-phenylcarbamates and acylanilide herbicides. Both 4chloroaniline and 3,4-dichloroaniline are extensively mineralized by P. chrysosporium (Table 6.12). Efficient mineralization of chloroanilines depends on adequate aeration, uptake and utilization in culture flasks with large head-space volumes together with

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nitrogen-limited conditions (143). Several studies have demonstrated that chloroanilines covalently bound to lignin in plant cell walls are also mineralized by P. chrysosporium (7, 105). Ligninolytic enzymes can directly oxidize chloroanilines, readily forming oligomers (Table 6.12). Examples of oligomeric structures that have been identified are shown in Figure 6.12. Some structures completely conserve the chlorine, such as the dimer, 4,4'-dichloroazobenzene. Other structures, such as trimers and tetramers, indicate that at least some of the original chloroaniline moieties are dechlorinated during the polymerization process. Some of the oligomers observed in vitro have also been found in vivo (25). The polymerization of chloroanilines is suggested to account for the formation of bound residue in soil humus. To explore this possibility, copolymerization of chloroaniline with phenolic compounds has been shown to result in the formation of humic polymers with incorporated chloroanilines (123, 158). Aside from polymerization reactions, products conjugated with organic acids have also been observed as metabolites during degradation of chloroanilines by P. chrysosporium. These include chloroanilines that are N-conjugated with succinic and acids (Table 6.12). The initial congugation is proposed to be a spontaneous reaction between A and 3,4-dichloroaniline, forming (34DC1-PKGA) (143). 34DC1PKGA is suggested to be the precursor of N-(3,4-dichlorophenyl)succinimide. The latter is formed after cleavage of 34DC1-PKGA by Conjugation is thought to be an important detoxification mechanism. By protecting anilines from polymerizatiom, the fungus is protected from toxic oligomerization products (143). The tendency for chloroanilines to undergo oxidative coupling in the presence of ligninolytic enzymes is extremely high. Protective groups are therefore necessary to afford the high levels of chloroaniline mineralization observed with P. chrysosporium. The biodegradation pathway is thus best described as a competition between polymerization and conjugation. The polymerization route ultimately leads to humic polymers; although, even chloroanilines bound to humus may eventually be subject to partial mineralization by white rot fungi (58). The formation of conjugates appears to be a prerequisite for direct mineralization of chloroanilines. An overall scheme depicting chloroaniline degradation by white rot fungi is shown in Figure 6.13.

3.12. Biodegradation of Chlorolignin During elemental chlorine bleaching and, to a lesser extent, during chlorine dioxide bleaching of pulp, lignin becomes chlorinated, is subsequently extracted by alkali, and is discharged in bleachery effluents. The resulting chlorolignin is responsible for a major portion of the adsorbable organic halogen (AOX) load at a pulp mill (129, 153, 154). White rot fungi have been investigated for their potential application in the treatment of chlorolignin in bleachery effluents, both for the removal of color and AOX. Various white rot fungi have been shown to remove 31 to 62% of the organohalogens in bleachery effluents (Table 6.13). In some cases, the evidence for dehalogenation is further supported by an increase in the inorganic chloride content (124, 125). The removal of organohalogens is associated with effluent decolorization, as well as a decrease in molecular weight of the lignin in some cases (51, 94, 117). In order to sustain high levels of organohalogen removal, glucose must be added at relatively high

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concentrations (125, 177). A number of observations have indicated the involvement of ligninolytic enzymes in dehalogenation. First, catalase added to cultures causes a white rot fungus to loose its ability to remove AOX, indicating involvement of peroxidases (94). Second, the dehalogenation is associated with the onset of ligninolytic enzyme production (92, 184). Third, MnP appears to directly depolymerize chlorolignin (87). Two types of technology have been developed for treatment of chlorine bleachery effluents, using white rot fungi. One technology, known as Mycelial Color Removal (MyCoR), utilizes P. chrysosporium immobilized onto rotating biological contactors (RBC) (125, 128, 184). Two distinct phases are involved in the MyCoR system. First, there is a primary growth phase to establish fungal mycelium on the RBC discs. Second, there is a secondary stage where chlorolignin degradation occurs. This phase is initiated with the appearance of ligninolytic enzymes triggered by nutrient limitation. During the secondary phase, from 44 to 62% of organohalogens are removed, with hydraulic retention times of 1 to 2 days (124, 184). Another technology is known as the MYCOPOR process, which is basically an unsubmerged down-flow trickle filter with counter-current aeration (107). P. chrysosporium is immobilized onto polyurethane foam blocks. In this system, from 45 to 55% organohalogen removal is obtained at a hydraulic retention time of 12 h. The MYCOPOR system has been operated for up to two years under non-sterile conditions. The best results for organohalogen removal from bleachery effluents have been obtained with sequential treatment systems. For example a sequence

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involving MyCoR as the first stage followed by anaerobic treatment, resulted in 65% AOX removal (128).

4. CONCLUSIONS White rot fungi play an important role in the global ecology by degrading complex lignin polymers. Their ability to degrade lignin is attributed to their unique ability to produce extracellular oxidative enzymes (ligninolytic enzymes). These enzymes cause the random oxidation of aromatic compounds, including priority pollutants. Their ability to also fortuitously dechlorinate a wide array of chlorinated organics has resulted in their use for bioremediation technologies. The white rot fungi, in particular, with their wide ranging metabolic capabilities, have served to motivate a growing body of research and development of these fungi for bioremediation. This chapter reviews the potential of white rot fungi to degrade specific chlorinated priority pollutants. First, evidence is presented in support that some white rot fungi are actually producers of organochlorine compounds. The most important organochlorine compounds produced are methyl ethers of chlorophenols (e.g., 3,5-dichloro-p-anisyl alcohol) and chloromethane. Some of the methyl ethers of chlorophenols are secondary metabolites which support lignin degradation, whereas others are antibiotic compounds. In addition, chloromethanes can serve as synthons or methyl donors during secondary metabolite biosynthesis. Second, evidence is presented which indicates that a wide variety of organochlorine pollutants are cometabolically degraded by white rot fungi (summarized in Table 6.14). Organochlorine compounds in most of the categories listed exhibit at least 10% mineralization of the parent compound, indicating that a significant and complete level of biodegradation is achieved by white rot fungi. Since the main motivation for considering white rot fungi for biodegradation was due to their ligninolytic enzymes, it is noteworthy that direct oxidation by ligninolytic enzymes is only truly established for four of the thirteen compound categories evaluated (chlorophenols, dioxins, cloroanilines and chlorolignin). The actual mechanism of dechlorination is generally considered to be nucleophilic substitution of water on a cation radical of the substrate, thereby displacing a chloro- group with a hydroxy-group. Ligninolytic enzymes are also most likely involved in the degradation of chlorophenoxyacetates, where chlorophenols are generated as intermediates after side chain hydrolysis. The ligninolytic enzymes are also indirectly implicated in the reduction of polychlorinated ethenes and methanes, through the formation of reducing radicals with organic acids. When considering both direct and indirect effects, the involvement of ligninolytic enzymes in the degradation of chlorinated organics has been established in only six of the compound categories. As expected with ligninolytic enzymes, a low nitrogen medium is required to induce their activity, thereby stimulating degradation in these cases (Table 6.14). In addition to the dechlorination reactions mediated by ligninolytic enzymes, free radical production and subsequent polymerization reactions by these same enzymes have had a profound effect on the fate of chlorinated organics in the environment. For example, chlorophenols and chloroanilines are highly susceptible to polymerization reactions by ligninolytic enzymes and readily combine with the organic soil fraction, to form covalently bound humic residues. This is commonly reported during white rot fungal bioremediation of

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these pollutants. Ligninolytic enzymes have no demonstrated involvement in the degradation of the remaining seven categories of compounds discussed here. Instead, monooxygenase activity, generally related to cytochromes, has been established in a few cases. For example, the use of selective inhibitors has established the involvement of cytochrome in the degradation of lindane, atrazine, and chlorodiphenyl ether by white rot fungi. Also, hydroxylases (monooxygenase) have been implicated in chlorophenol degradation. The observed profile of metabolic intermediates from chloroacetinilide and DDT biotransformations reflects monooxygenase (hydroxylase) activity. The enzymes implicated in PCB biodegradation by white rot fungi are still not identified.

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Although white rot fungi are best known for their ability to oxidize pollutants, the recent discovery of a reductive dehalogenase from Phanerochaete chrysosporium is remarkable (132, 134). The enzyme is involved in dechlorinating chloro-1,4hydrqoquinone intermediates generated during chlorophenol degradation. The reductive dehalogenase is a two-component enzyme. The first component is a glutathione transferase that produces a dechlorinated glutathione-hydroquinone conjugate. The second component is a glutathione-conjugate reductase, which reduces the conjugate to hydroquinone and glutathione. Finally, based on isotope labeling experiments, the use of chloromethane as a synthon in the biosynthesis of methyoxylated secondary metabolites has been proven, implying that a methyltransferase is involved (32). However, no methyltransferase utilizing chloromethane as a methyl donor has of yet been isolated from white rot fungi. In conclusion, white rot fungi have multiple enzymatic strategies to dechlorinate chlorinated pollutants, utilizing either ligninolytic enzymes, monooxygenases (including cytochrome glutathione-dependent reductive dehalogenases and methyl transferase. REFERENCES 27:173-181 1. Aiken BS & Logan BE (1996) Degradation of pentachlorophenol by the white rot fungus 9. Armenante PM, Lewandowski G & Haq IU Phanerochaete chrysosporium grown in (1992) Mineralization of 2-chlorophenol by P. ammonium lignosulphonate media. chrysosporium using different reactor designs. Biodegradation 7:175-182 Hazard. Waste Hazard. Mater. 9:213-229 2. Aislabie JM, Richards NK & Boul HL (1997) 10. Armenante PM, Pal N & Lewandowski G (1994) Role of mycelium and extracellular protein in Microbial degradation of DDT and its residues A review. N. Z. J. Agric. Res. 40:269-282 the biodegradation of 2,4,6-trichlorophenol by Phanerochaete chrysosporium. Appl. Environ. 3. Aitken MD, Massey IJ, Chen TP & Heck PE (1994) Characterization of reaction-products Microbiol. 60:1711-1718 from the enzyme-catalyzed oxidation of phenolic 11. Aust SD (1990) Degradation of environmentalpollutants by Phanerochaete chrysosporium. pollutants. Water Res. 28:1879-1889 Microb. Ecol. 20:197-209 4. Akhtar M, Blanchette RA & Kirk. TK (1997) Fungal delignification and biomechanical 12. Barr DP & Aust SD (1994) Mechanisms whitepulping of wood. In: Scheper T (Ed) Advances rot fungi use to degrade pollutants. Environ. Sci. in Biochemical Engineering / Biotechnology, Technol. 28:A78-A87 (pp 159-195). Springer-Verlag, Berlin 13. Barr DP, Shah MM, Grover TA & Aust SD (1992) Production of hydroxyl radical by lignin 5. Alleman BC, Logan BE & Gilbertson RL (1995) Degradation of pentachlorophenol by fixed films peroxidase from Phanerochaete chrysosporium. of white-rot fungi in rotating tube bioreactors. Arch. Biochem. Biophys. 298:480-485 Water Res. 29:61-67 14. Beaudette LA, Davies S, Fedorak PM, Ward OP & Pickard MA (1998) Comparison of gas 6. Archibald FS, Bourbonnais R, Jurasek L, Paice chromatography and mineralization experiments MG & Reid ID (1997) Kraft pulp bleaching and for measuring loss of selected polychlorinated delignification by Trametes versicolor. J. biphenyl congeners in cultures of white rot Biotechnol. 53:215-236 7. Arjmand M & Sandermann H (1985) fungi. Appl. Environ. Microbiol. 64:2020-2025 Mineralization of chloroaniline lignin conjugates 15. Beaudette LA, Ward OP, Pickard MA & and of free chloroanilines by the white rot Fedorak PM (2000) Low surfactant concentration increases fungal mineralization of fungus Phanerochaete chrysosporium. J. Agric. a polychlorinated biphenyl congener but has no Food Chem. 33:1055-1060 effect on overall metabolism. Lett. Appl. 8. Arjmand M & Sandermann HJ (1987) N(chlorophenyl)-succinimides: A novel metabolite Microbiol. 30:155-160 class isolated from Phanerochaete 16. Bergbauer M, Eggert C & Kraepelin G (1991) Degradation of chlorinated lignin compounds in chrysosporium. Pesticide Biochem. Physiol.

198 a bleach plant effluent by the white-rot fungus Trametes versicolor. Appl. Microbiol. Biotechnol. 35:105-109 17. Bollag JM, Liu SY & Minard RD (1980) Crosslinkage ofphenolic humus constituents and 2,4dichlorophenol. Soil Sci. Soc. Am. J. 44:52-56 18. Brock BJ & Gold MH (1996) 1,4-Benzoquinone reductase from the basidiomycete Phanerochaete chrysosporium: Spectral and kinetic analysis. Arch. Biochem. Biophys. 331:31-40 19. Brock BJ, Rieble S & Gold MH (1995) Purification and characterization of a 1,4benzoquinone reductase from the basidiomycete Phanerochaete chrysosporium. Appl. Environ. Microbiol 61:3076-3081 20. Bumpus JA & Aust SD (1987) Biodegradation of DDT [1,1,1 -trichloro-2,2-bis(4- chlorophenyl) ethane] by the white rot fungus Phanerochaete chrysosporium. Appl. Environ. Microbiol. 53:2001-2008 21. Bumpus JA, Kakar SN & Coleman RD (1993) Fungal degradation of organophosphorous insecticides. Appl. Biochem. Biotechnol. 39:715-726 22. Bumpus JA, Powers RH & Sun T (1993) Biodegradation of DDE (1,1 -dichloro-2,2-bis(4chlorophenyl)ethene) by Phanerochaete chrysosporium. Mycol. Res. 97:95-98 23. Bumpus JA, Tien M, Wright D & Aust SD (1985) Oxidation of persistent environmental pollutants by a white rot fungus (Phanerochaete chrysosporium). Science 228:1434-1436 24. Cameron MD & Aust SD (1999) Degradation of chemicals by reactive radicals produced by cellobiose dehydrogenase from Phanerochaete chrysosporium. Arch. Biochem. Biophys. 367:115-121 25. Chang CW & Bumpus JA (1993) Oligomers of 4-chloroaniline are intermediates formed during its biodegradation by Phanerochaete chrysosporium. FEMS Microbiol. Lett.107:337342 26. Cho NS, Rogalski J, Jaszek M, Luterek J, Wojtas-Wasilewska M, Malarczyk E, FinkBoots M & Leonowicz A (1999) Effect of coniferyl alcohol addition on removal of chlorophenols from water effluent by fungal laccase. J. Wood Sci. 45:174-178 27. Choi SH, Moon SH, Lee JS & Gu MB (2000) Biodegradation of 2,4,5-tichlorophenol using cell-free culture broths of Phanerochaete chrysosporium. J. Microbiol. Biotechnol. 10:759-763 28. Chung N, Kang GY, Kim GH, Lee IS & Bang WG (2001) Effect of nutrient nitrogen on the degradation of pentachlorophenol by white rot

FIELD fungus, Phanerochaete chrysosporium. J. Microbiol. Biotechnol. 11:704-708 29. Chung NH & Aust SD (1995) Veratryl alcoholmediated i n d i r e c t oxidation of pentachlorophenol by lignin peroxidase. Arch. Biochem. Biophys. 322:143-148 30. Chung NH, Shah MM, Grover TA & Aust SD (1993) Reductive activity of a manganesedependent peroxidase from Phanerochaete chrysosporium. Arch. Biochem. Biophys. 306:70-75 31. Collins PJ, Dobson ADW & Field JA (1998) R e d u c t i o n of the 2 , 2 ' - a z i n o b i s ( 3 ethylbenzthiazoline-6- sulfonale) cation radical by physiological organic acids in the absence and presence of manganese. Appl. Environ. Microbiol. 64:2026-2031 32. Coulter C, Hamilton JTG & Harper DB (1993) Evidence for the existence of independent chloromethane-utilizing and S-adenosylmethionine-utilizing systems for methylation in Phanerochaete chrysosporium. Appl. Environ. Microbiol. 59:1461-1466 33. Coulter C, Kennedy JT, McRoberts WC & Harper DB (1993) Purification and properties of an S-adenosylmethionine - 2,4-disubstituted phenol O-methyltransferase from Phanerochaete chrysosporium. Appl. Environ. Microbiol. 59:706-711 34. Dean JFD & E. EK (1992) Biotechnological modifications of lignin structure and composition in forest trees. Holzforschung 46:135-147 35. Dec J & Bollag JM (1990) Detoxification of substituted phenols by oxidoreductive enzymes through polymerization reactions. Arch. Environ. Contam. Toxicol. 19:543-550 36. Dec J & Bollag JM (1994) Dehalogenation of chlorinated phenols during oxidative coupling. Environ. Sci. Tech. 28:484-490 37. de Jong E, Cazemier AE, Field JA & DeBont JAM (1994) Physiological role of chlorinated aryl alcohols biosynthesized de-novo by the white-rot fungus Bjerkandera sp. strain Bos55. Appl. Environ. Microbiol. 60:271-277 38. de Jong E & Field JA (1997) Sulfur tuft and turkey tail: Biosynthesis and biodegradation of organohalogens by basidiomycetes. Annu. Rev. Microbiol. 51:375-414 39. de Jong E, Field JA & de Bont JAM (1994) Aryl alcohols in the physiology of ligninolytic fungi. FEMS Microbiol. Rev. 13:153-188 40. de Jong E, Field JA, Spinnler HE, Wijnberg J & de Bont JAM (1994) Significant biogenesis of chlorinated aromatics by fungi in natural environments. Appl. Environ. Microbiol. 60:264-270

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PART III. BIOCHEMISTRY AND CHEMISTRY

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Chapter 7 BACTERIAL GROWTH ON HALOGENATED ALIPHATIC HYDROCARBONS: GENETICS AND BIOCHEMISTRY DICK B. JANSSEN, JANTIEN E. OPPENTOCHT AND GERRIT J. POELARENDS Biochemical Laboratory, Groningen Biomolecular Sciences and Biotechnology Institute, University of Groningen, Groningen, The Netherlands

1. INTRODUCTION Many synthetically produced halogenated aliphatic compounds are xenobiotic chemicals in the sense that they do not naturally occur on earth at biologically significant concentrations. Nevertheless, various microorganisms have been isolated that possess the capacity to grow at the expense of these compounds, and to use them as a carbon and energy source under aerobic conditions. This raises a number of interesting questions concerning the mechanisms of dehalogenation, and the evolution and distribution of dehalogenase-encoding genes. For example, to what extent were the biochemical pathways for the mineralization of such xenobiotic halogenated chemicals preexisting before their large-scale anthropogenic introduction into the environment? What adaptation events at the genetic level occurred after the release of the xenobiotics? What are the origins of enzymes that cleave carbon-halogen bonds? Did similar mechanisms evolve in different organisms, at different sites, or were genes and organisms distributed from one site to another? Over the last few decades, several classes of reactions for the dehalogenation of xenobiotic compounds have been found, and in some cases the enzymatic mechanisms have been unraveled to the atomic level by x-ray crystallographic studies. At the same time, genetic studies have revealed much about the diversity, similarities, and possible distribution mechanisms of dehalogenase genes. This review focuses on the catabolic pathways for the aerobic mineralization of halogenated aliphatics, including dehalogenation reactions and issues of adaptation and distribution, with an emphasis on events that may have occurred after introduction of these xenobiotics into the environment. Bacterial growth on halogenated compounds requires the presence of enzymes that are capable of cleaving carbon-halogen bonds. The mechanisms and diversity of some Dehalogenation: Microbial Processes and Environmental Applications, pages 207-226 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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of these enzymes have been studied in detail. Furthermore, genetic analysis has yielded insight into the evolutionary relationships of dehalogenase genes and their distribution. This allows one to understand genetic events that have contributed to the recruitment of catabolic pathways for synthetic compounds. In this chapter, the most important principles are illustrated with enzymes and genes involved in the degradation of xenobiotic organohalogens such as dichloromethane, 1,2-dichloroethane, 1,2dibromoethane, and 1,3-dichloropropylene.

2. AEROBIC DEGRADATION OF HALOALIPHATICS Most halogenated aliphatic compounds that occur as environmental pollutants should, from a thermodynamic point of view, be suitable growth substrates for aerobic bacteria (7). Indeed, a large number of aerobic bacterial cultures that utilize haloaliphatics with a low degree of halogenation (1-3 halogens per molecule) have been found. Their generation times may be in the range of 2 to 10 hours, which makes their cultivation for physiological studies convenient. Even some highly chlorinated compounds can serve as a growth substrate, e.g., dichloromethane, trichloroacetate, and hexachlorocyclohexane (Figure 7.1). Enrichment trials and degradation studies have also shown that such bacterial cultures can only be obtained for a restricted number of haloaliphatics, whereas many other chlorinated compounds appear to be refractory to degradation. Recalcitrance may either be due to the lack of an enzyme that can catalyze the first step, often a dehalogenation reaction, or to the lack of a productive metabolic pathway. If dehalogenation is not the first step, there is a high risk of formation of toxic intermediates which may prohibit the use of a compound as growth substrate. Examples include the difficulties of obtaining growth on substrates such as 1,2-dibromoethane and chloroethenes. Modified pathways following different metabolic strategies may be used to prevent the formation of toxic intermediates. For example, the conversion of 1,2dichloroethane and 1,2-dibromoethane starts with a hydrolytic cleavage by a haloalkane dehalogenase for both compounds, resulting in their corresponding alcohols. Whereas 1,2-dichloroethane-degrading Xanthobacter strains further oxidize the 2-chloroethanol product by an alcohol dehydrogenase, the 2-bromoethanol formed from 1,2dibromoethane in a 1,2-dibromoethane-utilizing Mycobacterium is directly dehalogenated by a halohydrin lyase to epoxyethane. In this way, the accumulation of the highly toxic intermediate bromoacetaldehyde is prevented (42). Some of the most important pathways for the aerobic degradation of haloaliphatics, and the role of dehalogenases are presented in Figure 7.1. It appears that the actual dehalogenation reactions are very diverse. Dehalogenation may proceed, for example, by substitution (e.g., with water), elimination of HCl, or by reduced cofactor-dependent oxidative conversion. Hydrolytic dehalogenases have received considerable attention, and x-ray structures of some of these enzymes have been determined from which detailed mechanistic insight has been obtained.

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3. CLASSIFICATION AND MECHANISTIC ASPECTS OF DEHALOGENASES Traditionally, dehalogenases have been classified according to their substrate range. Such a classification scheme places haloacid dehalogenases, haloacetate dehalogenases, and haloalkane dehalogenases in different groups. We think this classification should be abandoned, since more recent information obtained from sequence analysis has shown that dehalogenases can be ordered into different homology groups, corresponding to the structural folds and catalytic mechanisms of the enzymes, rather than their observed substrate range. For example, fluoroacetate dehalogenase from a Moraxella strain (DehH1) (23) appears to be structurally more similar to several haloalkane dehalogenases which are also fold enzymes, than to most haloacid dehalogenases or haloacetate dehalogenases. Furthermore, enzymes that convert chloroacetate can be divided in different groups, some using a covalent mechanism whereas others not forming a covalent intermediate. In this case, a classification scheme according to substrate range provides little information on the evolution and distribution of enzymes and metabolic pathways. An overview of dehalogenase types, their relationship to other enzymes of the same class, and their genes and accession numbers, is given in Table 7.1. The first well-characterized dehalogenase, with respect to structure and mechanism, was that of Xanthobacter autotrophicus GJ10, an organism isolated in 1983 in The Netherlands (19). The enzyme, DhlA, catalyzes dehalogenation of a variety of short chain haloalkanes. The reaction mechanism of DhlA is characterized by covalent catalysis, with an Asp-His-Asp catalytic triad similar to that found in the classical chymotrypsin-like serine proteases as the most salient feature (Figure 7.2). In the first step, there is a nucleophilic displacement in which the carbon atom of the substrate to which the halogen atom is bound is attacked by the nucleophile aspartate residue (Asp), thereby forming a covalent alkyl-enzyme ester intermediate (63). In the second step, the histidine base residue (His), assisted by the second Asp which acts as a charge-relay residue, activates a water molecule that hydrolyzes the ester intermediate by attacking the carbonyl carbon atom of the enzyme. The most important difference between this mechanism and that of serine proteases is that the first and second nucleophilic substitutions occur on different carbon atoms in the dehalogenase enzyme, whereas in proteases they occur on the same carbon. In agreement with this, the carbonyl function required for hydrolysis of the covalent intermediate, is supplied by the dehalogenase enzyme, rather than by the substrate as in serine hydrolases (44). In addition to the catalytic triad, the haloalkane dehalogenase posesses two tryptophan (Trp) residues that are involved in leaving group stabilization. Thus, the enzyme is really evolved to carry out a dehalogenation reaction, and it is not a general hydrolase that fortuitously also dechlorinates. Other haloalkane dehalogenases have been found in various Rhodococcus strains (26, 41). The DhaA enzyme of strain NCIMB13064 has a different substrate range than DhlA. The former enzyme also converts long-chain chloroalkanes, but its activity with 1,2-dichloroethane is very low. Both DhlA and DhaA enzymes belong to the hydrolase fold superfamily, a group to which also epoxide hydrolases belong. Other enzymes belonging to this group include DehHl from Moraxella sp. strain B, a haloacetate dehalogenase which converts both fluoro- and chloroacetate, and LinB, a

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halidohydrolase involved in the degradation of in Sphingomonas paucimobilis (Figure 7.1). Group classification is based on sequence similarity and use of a similar hydrolytic mechanism with involvement of a catalytic triad (17, 30, 31). In DhaA and LinB, the catalytic triad consists of Asp-His-Glu, with the glutamate residue (Glu) acting as the charge-relay acid rather than an Asp as in DhlA of X. autotrophicus (17, 38). In these enzymes, the catalytic Glu is located after strand 6, which is a different position than that of the catalytic Asp of DhlA, which is located behind strand 7 (6). It has been shown that the position ofthis charge relay residue can be moved in DhlA from behind strand 7 to a position behind strand 6 (24). Haloacid dehalogenases (HAD) are classified into at least two groups, which are evolutionarily unrelated (15). The HAD-type haloacid dehalogenases (group II) are the best characterized enzymes. These have a three-dimensional structure which is different from that found in haloalkane dehalogenases, but hydrolysis also proceeds via covalent catalysis (Figure 7.2). This class ofdehalogenases has a topological fold which is similar

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to the one found in several phosphatases, and the conserved nucleophilic Asp residue involved in the displacement of the halogen is located close to the N-terminus. DhlB of X. autotrophicus GJ10 and L-DEX of Pseudomonas sp. YL are examples where x-ray structures have been determined (16, 29, 46). These group II enzymes usually convert L-2-chloropropionic acid, but not D-2-chloropropionic acid, always with inversion of product configuration. Haloacetate dehalogenase DehH2 from Moraxella sp. B also belongs to this group, although substrate usage by this enzyme is non-stereospecific, illustrating that stereoselectivity and substrate range are not well correlated with structural and mechanistic features (23). Besides covalent catalysis, non-covalent catalysis has also been proposed for haloacid dehalogenases. There are indications that with the dehalogenase DL-DEX from Pseudomonas sp. strain 113, hydrolysis does not proceed via formation of a covalent ester intermediate, but the hydroxyl group of water directly attacks the of 2-haloalkanoic acid to displace the halogen atom (37). This protein belongs to a group of homologous haloacid dehalogenases (group I) that includes enzymes which convert both L-2-chloropropionic acid and D-2-chloropropionic acid, as well as enzymes which convert only D-2-chloropropionic acid with inversion of product configuration (2, 15). The group also includes a dehalogenase enzyme involved in the conversion of trihaloacetate to carbon monoxide (52). Other hydrolytic dehalogenases with activity towards chloroacetate and chloropropionates probably exist, but sequence data and structural information are too scarce to allow adequate classification (15). Halohydrin lyases (also called haloalcohol dehalogenases) provide another dehalogenation mechanism that uses intramolecular substitution, as is found in Corynebacterium sp. N-1074. These proteins share partial sequence similarity with proteins of the short-chain reductase/dehydrogenase (SDR) family. Such an enzyme is involved in the degradation of l,3-dichloro-2-propanol, where two halohydrin-lyase mediated dehalogenation steps yield an intermediary epoxide (Figure 7.1) (34, 35). Different halohydrin lyases in Corynebacterium sp. N-1074 have been described, which display different enantioselectivities. For example, HheA acts non-stereospecifically, while HheB is enantioselective, converting mostly the (R)-enantiomer from a racemic substrate mixture. The HheA and HheB enzymes show no significant structural similarity, except in the carboxyl-terminal region (65). Possibly, they share similar catalytic mechanisms but have major differences in substrate recognition domains. Dehalogenation by dichloromethane dehalogenase (DcmA) of Methylobacterium sp. DM4 and Methylophilus sp. DM11 involves a thiolytic substitution reaction (Figure 7.1). DcmA is an inducible enzyme enabling bacterial growth on dichloromethane as the sole carbon and energy source, All known dichloromethane dehalogenases belong to the gluthatione S-transferase super family The enzymatic mechanism involves a nucleophilic displacement of halogen (Cl) by glutathione, rather than by a hydroxyl or carboxylate oxygen, producing a halomethylthioether intermediate which is converted to formaldehyde (3). Two DcmA gene types have been found which are not directly related to each other. Comparison of strain DM11 and strain DM4 dichloromethane dehalogenase sequences revealed 56% identity at the protein level, thus indicating an ancient divergence of the two enzymes (1). Also structural differences between the two enzymes exist. The Methylobacterium sp. DM4 enzyme has a subunit molecular mass of 33 kDa and an structure, while the Methylophilus sp. DM11 enzyme has a subunit

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mass of 34 kDa and an structure (25, 27). Chloromethane has been shown to be degraded and utilized via formate by the facultative methylotrophic bacterium CC495 under aerobic conditions (4). The dehalogenating enzyme appeared to be an inducible 67 kDa methyltransferase with a corrinoid-bound cobalt atom. In the reaction, the methyl group of chloromethane appears to be transferred to a sulfhydryl acceptor yielding methanethiol, which can be converted to formaldehyde by a methanethiol oxidase (Figure 7.1G). The methyltransferase enzyme shows mechanistic similarities to cobalamin-dependent methionine synthase from Escherichia coli. Alternatively, in Methylobacterium sp. the methyltransferase becomes methylated and subsequently donates the methyl group to tetrahydrofolate in a reaction catalyzed by another methyltransferase (61). This second methyltransferase was recently purified and accepted methylcobalamin as the methyl donor (53). Further metabolism then proceeds via a pterine-dependent pathway. Chloroacrylic acid dehalogenase of Pseudomonas pavonaceae is the second dehalogenating enzyme in the degradation pathway of 1,3-dichloropropene (43) (Figure 7.1C). A formal hydration reaction at the C=C bond of 3-chloroacrylic acid takes place, yielding the unstable intermediate 3-chloro-3-hydroxypropanoic acid, from which HCl is eliminated to give malonate semialdehyde (13). In a coryneform strain, two different enzymes are present, which convert either cis- or trans-3-chloroacrylic acid (60). The cis-3-chloroacrylic acid dehalogenase is a 38 kDa trimeric protein, while the trans-3chloroacrylic acid dehalogenase consists of 7.4 and 8.7 kDa subunits. In the first two steps in the degradation pathway of Sphingomonas paucimobilis (Figure 7.IE), a dehydrohalogenation reaction catalysed by hexachlorocyclohexane dehydrochlorinase (LinA) eliminates two biaxial HCl pairs in two sequential steps, leading to the formation of double bonds in the substrate (33, 55). LinA is a homotetramer consisting of 16.5 kDa subunits. The linA gene shows no homology to other known dehalogenase genes (18). However, the involvement in catalysis of a histidine residue (His73) as a base that abstracts a proton from the substrate, facilitated by Asp25, was proposed on the basis of similarity to a hydratase and isomerase with known x-ray structures (55). How the enzyme interacts with the halogen is at present unknown. The product formed by LinA, 1,3,4,6-tetrachloro-1,4cyclohexadiene, is converted by the halidohydrolase LinB, an enzyme of the fold superfamily, as discussed above. Vinyl chloride, a compound that is produced on a large scale by industry and is also biologically formed during the reductive breakdown of chlorinated ethenes, can be degraded oxidatively. Many organisms have been described that degrade vinyl chloride cometabolically during utilization of other compounds, but so far only two strains, Mycobacterium aurum L1 and Pseudomonas aeruginosa MF1, have been isolated which grow on vinyl chloride as the sole carbon and energy source (12, 62). Vinyl chloride breakdown in M. aurum L1 was shown to be mediated by an alkene monooxygenase, yielding chlorooxirane, which is presumably further degraded by an epoxide dehydrogenase (11). In this pathway, the monooxygenase-mediated conversion is not the dehalogenating reaction. In the oxygenolytic degradation of chloromethane and 1,2-dichloroethane, the first degradation step is most likely catalyzed by a dehalogenating monooxygenase. Chloromethane serves as a sole carbon and energy source for growth of

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Hyphomicrobium MC1 (14). Degradation is possibly catalyzed by methyl chloride monooxygenase, yielding formaldehyde. Recently, a Pseudomonas strain (DCA1) was described that grows aerobically on 1,2-dichloroethane as the sole carabon source (Figure 7.1). The first step of the 1,2-dichloroethane degradation pathway is thought to be mediated by a monooxygenase because the reaction is dependent of oxygen and NADH (10), In the proposed pathway, oxidation of 1,2-dichloroethane yields chloroacetaldehyde, which is most likely further degraded to glycolate. This is analogous to the 1,2-dichloroethane degradation pathway described in X. autotrophicus GJ10. At the moment, no DNA sequences or information on protein structure or reaction mechanisms of the monooxygenase proteins involved in the dehalogenation reactions are available.

4. RECENT EVOLUTION OF HALOALKANE DEHALOGENASE The evolution of enzymes and catabolic pathways for the degradation of synthetic compounds has received considerable attention. This has often been based on the notion that the degradation of xenobiotics presents a real challenge for microorganisms in polluted environments. Often, however, the evolution of catabolic pathways for xenobiotics is discussed in the context of systems that have only distant evolutionary relationships. It seems unlikely that such distant similarities are related to, or reflect evolutionary processes that occurred over the last 100 years, as might occur in response to the release of synthetic chemicals. If we want to understand how microorganisms present in the natural environment adapt to the introduction of industrial pollutants into the environment, it is important to focus on evolutionary events which have occurred recently. In fact, very little is known about the genetic changes that have occurred during the exposure and adaptation of microorganisms to industrial chemicals. Even though an organism degrades a chlorinated (xenobiotic) compound, it does not mean that a new catabolic pathway is required, since numerous organohalogens are naturally produced. Different views have been presented with respect to the degree of mutations that occur during the evolution of new enzymes. A spectacular hypothesis has been proposed by Ohno (39), who suggests that the nylon oligomer hydrolase gene of Flavobacterium sp. K172 had evolved by a shift in reading frame, which occurred in a very ancient gene that was composed of primordial oligomeric repeats and encoded an arginine-rich protein of completely unrelated function. This view supports that a completely new protein arose due to exposure and adaptation to a xenobiotic chemical. However, gene sequences homologous to the oliogomer hydrolase can also be found in other bacteria, including organisms with no known history of nylon oligomer degradation, such as P. aeruginosa PAO. Furthermore, it is impossible to rule out that a very similar hydrolase exists that is involved in cleavage of amide bonds found in natural compounds. We support the view that new catabolic activities evolve by a limited number of mutations in genes that encode proteins with very similar function. A natural enzyme for which a recent adaptation process has been proposed is the haloalkane dehalogenase from X. autotrophicus GJ10 (45). Several lines of evidence indicate that this dehalogenase is especially adapted to convert haloalkanes, and has undergone the following evolutionary changes during adaptation to synthetic 1,2dichloroethane:

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The nucleophile consists of aspartate and two tryptophan residues which are involved in stabilization of the leaving group. This indicates that the active site of the enzyme is specifically designed to catalyze dehalogenation reactions. Although the dehalogenase is evolutionarily related to fold enzymes, it most probably was already a dehalogenase before industrial pollution started. Because active dehalogenases have been found in other organisms (e.g., Mycobacterium), not selected for growth on chlorinated compounds (21), it is likely that the enzyme preexisted, or was a dehalogenase for some other naturally-occurring compounds prior to being challenged with industrial 1,2-dichloroethane. The enzyme actually has evolved from a more primitive dehalogenase by recent genetic adaptation events. Short direct sequence repeats are present in the cap domain, i.e., a 15 bp perfect repeat and a 9 bp imperfect (one substitution) repeat. It is unlikely that these repeats are of ancient evolutionary origin, since it is difficult to envision that the presence of short repeated sequences would be needed for proper enzyme function. Instead, we propose that they are the result of recent mutations (less than 100 years ago), which adapted the enzyme to 1,2-dichloroethane. This could be achieved by starting from a dehalogenase that was active with an unknown natural halogenated compound. Experimental work supports the role of spontaneous mutants has been investigated. The mutations selected are all localized in the cap domain, and included repeats similar to of those identified in the wild-type enzyme (45). This 1,2-dichloroethane dehalogenase is probably the only dehalogenase for which there is some evidence that it has undergone mutations during adaptation to a synthetic organohalogen. Although speculation exists, there is little proof for recent adaptations to halogenated substrates of, for example, glutathione transferases and enoyl hydratases. There certainly is an evolutionary relationship between these latter enzymes and the dehalogenases, but the time of branching may have occurred very long ago, and may not be related to adaptation to the relatively recent presence of xenobiotics. Again, it should be noted that many natural organohalogens exist, and that haloaromatics are particularly abundant, e.g., in decomposing wood (see also Chapter 1 and 6). Features in the DNA sequence which are the result of fortuitous mutations and serve no useful function should be regarded as a sign of recent genetic change, as opposed to those evolutionary changes that are the result of continued selection and optimization of enzyme activity (42). A striking example of how the sequence of a gene that has undergone recent mutations may be influenced by the mutagenic event is provided by the haloalkane dehalogenase gene of Mycobacterium sp. GP1 (Figure 7.3). The gene of this organism is fused in frame to a segment of the hheB gene, encoding haloalcohol dehalogenase, resulting in an extension of the dhaA open reading frame by 14 codons. Since this hheB part of is identical to the 3' end of an intact hheB gene that is located approximately 2.6 kb further downstream, it seems likely that a duplication of the hheB gene occurred prior to acquisition and insertion of the dhaA gene. The organization and sequence of these catabolic genes as we see it now, is in part the result of fortuitous genetic events that yielded a suitable activity which probably holds for other systems as well. Moreover, it does not appear to be the result of extensive evolutionary optimization.

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5. REGULATION OF DEHALOGENASE ACTIVITY A dehalogenase enzyme may be acquired by a microorganism and altered to convert a novel compound, but this only contributes to a functional catabolic pathway if the protein is produced when it is needed. Acquisition of a regulatory system for gene expression is a fundamental second step in the evolution of an efficient new catabolic pathway. In addition to the catabolic enzyme, it requires a second protein molecule that can recognize and bind the target compound. The expression of various dehalogenases is indeed regulated by the presence of substrate, which has been identified for enzymes converting halocarboxylic acids, chloroalkanes, chloroacrylic acids, chloroalcohols, 4-chlorobenzoate, and pentachlorophenol. Such inducible expression suggests that the catabolic pathways are well developed. Apparently, there has been enough time to evolve a regulatory system. A number of regulatory systems have been characterized in more detail. In Pseudomonas putida PP3, the regulatory protein DehRl responds to inducers such as 2-monochloropropionate and monochloroacetate, and activates transcription of the

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haloalkanoic acid dehalogenase gene dehI (56-58). Transcription is mediated by an alternative RNA polymerase containing the factor. In dichloromethane-utilizing methylotrophs, the negative regulator DcmR controls expression of dichloromethane dehalogenase DcmA (27, 28, 48). The plasmid-located haloalkane dehalogenase gene (dhaA) in various haloalkane-utilizing strains of Rhodococcus erythropolis is regulated by the product of the adjacent dhaR gene (41). The DhaR protein belongs to the TetR family of transcriptional repressor-type regulators and responds to 1-chlorobutane and several other 1-halo-n-alkanes (5, 40). It is likely that the regulatory proteins for these enzymes were designed originally to respond to natural organohalogens, and that they have been modified during the development of degradative pathways for the respective halogenated xenobiotics. In other cases, a regulatory system is lacking or it is not functional, which may be seen as a sign of evolutionary primitivity. For example, in X. autotrophicus GJ10 (Figure 7.1), the haloalkane dehalogenase gene (dhlA) is expressed constitutively and addition of 1,2-dichloroethane does not enhance expression. However, an open reading frame that is located directly upstream of the dehalogenase gene encodes a putative regulatory protein that shares low but significant sequence similarity with members of the TetR family of repressor-type regulators (20). In addition, the promoter region of the haloalkane dehalogenase gene contains two copies of the palindromic sequence TAGGTCNNNNGACCTA, which may serve as binding sites for the putative repressor. These observations suggest that normal transcriptional regulation of the dhlA gene (or its ancestral form) has been relaxed to allow expression in the presence of 1,2-dichloroethane, which is not recognized by the regulator. Another example in which alteration of a regulatory protein played a role in the ability to utilize a new carbon source is found in Mycobacterium sp. strain GP1 (40, 42). This bacterium is capable of growth on 1,2-dibromoethane (Figure 7.1) via a haloalkane dehalogenase (DhaAf), for which 1,2-dibromoethane is a substrate but not an inducer. A deletion of 12 nucleotides has occurred in the regulatory dhaR gene, which in other strains encodes a repressor. Although this deletion is in frame, it inactivates the DhaR protein, leading to constitutive expression of the dehalogenase.

6. MOBILIZATION AND DISTRIBUTION OF DEHALOGENASE GENES 6.1. Plasmids, Transposons and Insertion Elements Various studies on the evolution of catabolic pathways for haloaliphatics have provided evidence that gene transfer plays an important role. Important aspects are: Dehalogenating systems are often encoded on self-transmissible plasmids. This has been found for several haloalkane dehalogenases, such as the dhlA genes of Xanthobacter and Ancylobacter strains, the dhaA genes of Rhodococcus strains, and the dcm genes of dichloromethane-degrading bacteria (9, 26, 41, 48, 54). Gene clusters encompassing dehalogenases are often associated with insertion elements and located in a transposon-like structure, as has been found in chloropropionate, dichloromethane, and chloroalkane-utilizing organisms. For example, the highly conserved dichloromethane degradative gene (dcm) region in several phylogenetically distinct methylotrophic bacteria is associated with three

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distinct insertion elements (48). Another example is insertion element IS1071, first detected in a gene region involved in chlorobenzoate metabolism. The chlorobenzoate catabolic genes (cbaABC) of Alcaligenes sp. strain BR60 have been found to be encoded on a composite transposon, designated Tn5271, that is flanked by two copies of this 3.2 kb insertion element (IS1071) (64). Subsequently, the same insertion element was shown to flank the haloacetate dehalogenase gene dehH2 on plasmid pUOl in Moraxella sp. strain B (23, 64), the haloalkane dehalogenase gene dhaA on the chromosome in Pseudomonas pavonaceae 170 (40), the aniline degradative genes on a plasmid of Pseudomonas putida UCC22 (8), and presumably also the p-sulfobenzoate degradative genes (psbAC) on plasmids of Comamonas testosteroni (22). IS1071 thus seems to be responsible for mobilizing catabolic genes to the chromosome or a plasmid in different host organisms. How an insertion element may transpose itself to a dehalogenase gene and mobilize this gene into a plasmid has been demonstrated by studies on the adaptation of X. autotrophicus GJ10 to toxic concentrations of monobromoacetate (59). Bromoacetate can be hydrolyzed by haloacid dehalogenase (DhlB), but substrate concentrations above 5 mM are toxic to strain GJ10. Mutants which are able to grow in the presence of higher concentrations of bromoacetate were found to overexpress haloacid dehalogenase, and it appears that this resistance is accompanied by the incorporation of an unlinked insertion element (IS1247) to an upstream site adjacent to the chromosomally located dhlB gene. This insertion causes increased expression of haloacid dehalogenase. Furthermore, one-ended transposition of IS1247, together with dhlB, can lead to insertion of a copy of the dhlB gene into a plasmid present in the host. A dehalogenase gene thus may become activated by an insertion element, and the combination of a single insertion element and dehalogenase gene can transpose to another replicon. Similar evolutionary changes have been observed earlier by Senior and coworkers (49). Their continuous culture selection experiments have revealed that during a period of thousands of hours, one member of a seven-membered microbial community capable of utilizing the herbicide 2,2-dichloropropionic acid (Dalapon) acquired the ability to grow on Dalapon through the production of two dehalogenases. Apparently, cryptic dehalogenase genes in the original strain became activated during prolonged selection, most likely through the action of insertion elements (50). Several years later, Thomas et al. (56, 57) showed that the dehalogenase gene dehI of P. putida PP3 indeed is located on a transposable element, designated DEH.

6.2. Do DNA Integrases Play a Role in the Acquisition of Dehalogenase Genes? Acquisition of foreign DNA by horizontal gene transfer requires integration into a replicon that is stably maintained in the recipient microorganism. Both in P. pavonaceae 170 and in Mycobacterium sp. strain GP1 a gene encoding a putative site-specific recombinase (intP in strain 170 and intM in strain GP1), is present directly upstream of the recruited DNA segment harboring the haloalkane dehalogenase gene (Figure 7.3) (40). The intP and intM gene products share significant sequence similarity with members of the integrase (Int) family of site-specific recombinases, and both harbor the conserved, catalytically important, tetrad R-H-R-Y of the Int family. These putative

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integrase proteins probably mediate the insertion of the dehalogenase genes into the genome, although their activity remains to be established experimentally, The finding of putative integrase genes next to the dehalogenase genes further reinforces our hypothesis that these DNA segments were acquired by horizontal transmission. Integrase-mediated gene acquistion has previously been associated with a class of genetic elements called integrons (51). Integrons harbor a gene for a site-specific DNA integrase, which can mediate the incorporation of one or more foreign genes in a specific site (the recombination or core site), directly upstream of the integrase gene. Thus far, integrons have been implicated in the acquisition of antibiotic resistance genes by various bacterial species.

6.3. Global Distribution of a Gene Cluster Encoding Haloalkane Catabolism In some cases, organisms possessing identical dehalogenase genes have been found in very different geographical areas, For example, Gram-negative methylotrophs containing identical or nearly identical dhlA or dcmA genes have been isolated from widely separated sites in Europe. Furthermore, we have found that haloalkane-utilizing Rhodococcus strains isolated from contaminated sites in Europe, Japan, and the United States possess identical haloalkane dehalogenase (dhaA) genes (41). The same gene is also present in the 1,3-dichloropropylene degrader P. pavonaceae 170 and, in slightly modified form, in the 1,2-dibromoethane degrader Mycobacterium sp. GP1. Thus, transfer of the dhaA gene from a Rhodococcus strain to a Gram-negative organism may have occurred. Apparently, the capacity to use haloalkanes as a carbon source has become widespread due to the recent and global distribution of a single catabolic gene cluster. In all strains analyzed, the gene cluster is localized on plasmids or associated with insertion elements, suggesting a role for these mobile elements in gene transfer. Such observations are difficult to interpret without considering long-distance distribution mechanisms for microorganisms. Despite growing concern about biological invasions and emergent diseases, not much is known about the long-distance distribution mechanisms for microorganisms. Recently, Ruiz et al. (47) have suggested that the global movement of ballast water in ships creates a long-distance dispersal mechanism for human pathogens. Similarly, the global movement of seed and agriculture food products (e.g., potatoes, fruits and vegetables) may have contributed to the global spread of organohalogen-degrading microorganisms. Agricultural products are often treated with halogenated fumigants and/or are harvested from soils treated with fumigants, such as the soil disinfectants 1,3-dichloropropene and methyl bromide (see Chapter 12). This may have created ecological niches were dehalogenase genes proliferate, and thereby increased their occurrence and distribution.

7. WHY ARE SOME COMPOUNDS STILL RECALCITRANT? Several important chlorinated aliphatics that occur as environmental pollutants have so far not been found not to serve as a growth substrate for bacterial cultures under aerobic conditions. This includes compounds such as 1,1-dichloroethene, 1,2dichloropropane, 1,2,3-trichloropropane, the 1,2-dichloroethenes, trichloroethanes, and

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trichloroethene. For all these compounds, some form of degradation under aerobic conditions has been demonstrated. Thus, 1,1 -dichloroethane may be slowly oxidized by monooxygenases, 1,2-dichloropropane and 1,2,3-trichloropropane are (poor) dehalogenase substrates, and the dichloroethenes can be converted to their epoxides by monooxygenase activity. Rates of transformation are very low, however, or the products of the initial conversions are toxic, as with the chloroethene epoxides. There are no fundamental reasons that prohibit the use of these compounds as a substrate for aerobic growth. Their current recalcitrance may well be a fortuitous situation, and with ongoing evolution and adaptation, new organisms may appear with time. Of course, the time needed to evolve new metabolic activities is of great practical importance. Since evolution processes can be accelerated under laboratory conditions, it is entirely possible that directed evolution techniques will contribute to obtaining organisms with enhanced catabolic potential. REFERENCES 1. Bader R & Leisinger T (1994) Isolation and

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characterization of the Methylophilus sp. strain DM11 gene encoding dichloromethane dehalogenase/glutathione S-transferase. J. Bacteriol. 176:3466-3473 Barth PT, Bolton L & Thomson JC (1992) Cloning and partial sequencing of an operon encoding two Pseudomonas putida haloalkanoate dehalogenases of opposite stereospecificity. J. Bacteriol. 174:2612-2619 Blocki FA, Logan MS, Baoli C & Wackett LP (1994) Reaction of rat liver glutathione S-transferases and bacterial dichloromethane dehalogenase with dihalomethanes. J. Biol. Chem. 269:8826-30 Coulter C, Hamilton JT, McRoberts WC, Kulakov L, Larkin MJ & Harper DB (1999) Halomethane:bisulfide/halide ion methyltransferase, an unusual corrinoid enzyme of

environmental significance isolated from an aerobic methylotroph using chloromethane as the sole carbon source. Appl. Environ. Microbiol. 65:4301-4312 5. Curragh H, Flynn O, Larkin MJ, Stafford TM, Hamilton JTG & Harper DB (1994) Haloalkane degradation and assimilation by Rhodococcus rhodochrous NCIMB13064. Microbiology 140:1433-1442 6. Damborsky J & Koca J (1999) Analysis of the reaction mechanism and substrate specificity of haloalkane dehalogenases by sequential and structural comparisons. Protein Eng. 12:989-998 7. Dolfing J & Janssen DB (1994) Estimation of the Gibbs free energies of formation of chlorinated aliphatic compounds. Biodegradation 5:21-28 8. Fukomori F & Saint CP (1997) Nucleotide sequences and regulation analysis of genes

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involved in conversion of aniline to catechol in Pseudomonas putida UCC22 (pTDN1). J. Bacteriol. 179: 399-408 Gälli R & Leisinger T (1988) Plasmid analysis and cloning of the dichloromethane-utilization genes of Methylobacterium sp. DM4. J. Gen.icrobiol. 134:943-952 Hage JC & Hartmans S (1999) Monooxygenasemediated 1,2-dichloroethane degradation by Pseudomonas sp. strain DCA1. Appl. Environ. Microbiol. 65:2466 Hartmans S & De Bont JA (1992) Aerobic vinyl chloride metabolism in Mycobacterium aurum L1. Appl. Environ. Microbiol. 58:1220-1226 Hartmans S, De Bont JAM, Tramper J & Luyben KCAM (1985) Bacterial degradation of vinyl chloride. Biotechnol. Lett. 7:383-388 Hartmans S, Jansen MW, van der Werf MJ & de Bont JA (1991) Bacterial metabolism of 3-chloroacrylic acid. J. Gen. Microbiol. 137:2025-2032 Hartmans S, Schmuckle A, Cook A & Leisinger T (1986) Methyl chloride: Naturally occurring toxicant and substrate. J. Gen. Microbiol. 132:1139-1142 Hill KE, Marchesi JR & Weightman AJ (1999) Investigation of two evolutionarily unrelated halocarboxylic acid dehalogenase gene families. J. Bacteriol. 181:2535-2547 Hisano T, Hata Y, Fujii T, Liu JQ, Kurihara T, Esaki N, Soda K (1996) Crystal structure of L-2haloacid dehalogenase from Pseudomonas sp. YL. An alpha/beta hydrolase structure that is different from the alpha/beta hydrolase fold. J Biol Chem. 271:20322-30. Hynkova K, Nagata Y, Takagi M & Damborsky J (1999) Identification of the

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from Sphingomonas paucimobilis UT26. FEBS Lett. 446:177-181 18. Imai R, Nagata Y, Fukuda M, Takagi M & Yano K (1991) Molecular cloning of a Pseudomonas paucimobilis gene encoding a 17-kilodalton polypeptide that eliminates HC1 molecules from J. Bacteriol. 173:6811-6819 19. Janssen DB, Scheper A, Dijkhuizen L & Witholt B (1985) Degradation of halogenated aliphatic compounds by Xanthobacter autotrophicus GJ10. Appl. Environ. Microbiol. 49:673-677 20. Janssen DB, Pries F, van der Ploeg J, Kazemier B, Terpstra P & Witholt B (1989) Cloning of 1,2-dichloroethane degradation genes of Xanthobacter autotrophicus GJ10 and expression and sequencing of the dhlA gene. J. Bacteriol. 171:6791-6799 21. Jesenska A, Sedlacek I & Damborsky J (2000) Dehalogenation of haloalkanes by Mycobacterium tuberculosis H37Rv and other mycobacteria. Appl. Environ. Microbiol. 66:219-222 22. Junker F & Cook AM (1997) Conjugative plasmids and the degradation of arylsulfonates in Comamonas testosteroni. Appl. Environ. Microbiol. 63:2403-2410 23. Kawasaki H, Tsuda K, Matsushita I & Tonomura K (1992) Lack of homology between two haloacetate dehalogenase genes encoded on a plasmid from Moraxella sp. strain B. J. Gen. Microbiol. 138:1317-1323 24. Krooshof GH, Kwant EM, Damborsky J, Koca J & Janssen DB (1997) Repositioning the catalytic triad aspartic acid of haloalkane dehalogenase: Effects on stability, kinetics, and structure. Biochemistry 36:9571-9580 25. Kohler-Staub D, Hartmans S, Gälli R, Suter F & Leisinger T (1986) Evidence for identical dichloromethane dehalogenases in different methylotrophic bacteria. J. Gen. Microbiol. 132:2837-2843 26. Kulakova AN, Larkin MJ & Kulakov LA (1997) The plasmid-located haloalkane dehalogenase gene from Rhodococcus rhodochrous NCIMB 13064. Microbiology 143:109-115 27. La Roche SD & Leisinger T (1990) Sequence analysis and expression of the bacterial dichloromethane dehalogenase structural gene, a member of the glutathione S-transferase supergene family. J. Bacteriol. 172:164-171 28. La Roche SD & Leisinger T (1991) Identification of dcmR, the regulatory gene governing expression of dichloromethane dehalogenase in Methylobacterium sp. strain DM4. J. Bacteriot. 173:6714-6721 29. Li YF, Hata Y, Fujii T, Hisano T, Nishihara M, Kurihara T & Esaki N (1998) Crystal structures

JANSSEN ET AL. of reaction intermediates of L-2-haloacid dehalogenase and implications for the reaction mechanism. J Biol Chem. 273:15035-44 30. Liu JQ, Kurihara T, Ichiyama S, Miyagi M, Tsunasawa S, Kawasaki H, Soda K & Esaki N (1998) Reaction mechanism of fluoroacetate dehalogenase from Moraxella sp. B. J. Biol. Chem. 273:30897-30902 31. Marek J, Vevodova J, Smatanova IK, Nagata Y, Svensson LA, Newman J, Takagi M & Damborsky J (2000) Crystal structure of the haloalkane dehalogenase from Sphingomonas paucimobilis UT26. Biochemistry 39:1408214086 32. Nagata Y, Futamura A, Miyauchi K & Takagi M (1999) Two different types of dehalogenases, LinA and LinB, involved in cyclohexane degradation in Sphingomonas paucimobilis UT26 are localized in the periplasmic space without molecular processing. J. Bacteriol. 181:5409-5413 33. Nagata Y, Hatta T, Imai R, Kimbara K, Fukuda M, Yano K & Takagi M (1993) Purification and characterization of dehydrochlorinase (LinA) from Pseudomonas paucimobilis. Biosci. Biotechnol. Biochem. 57:1582-1583 34. Nakamura T, Nagasawa T, Yu F, Watanabe I & Yamada H (1992) Resolution and some properties of enzymes involved in enantioselective transformation of 1,3-dichloro2-propanol to (R)-3-chloro-l,2-propanediol by Corynebacterium sp. strain N-1074. J. Bacteriol. 174:7613-7619 35. Nakamura T, Nagasawa T, Yu F, Watanabe I & Yamada H (1994) Characterization of a novel enantioselective halohydrin hydrogen-halidelyase. Appl. Environ. Microbiol. 60:1297-1301 36. Nardi-Dei V, Kurihara T, Park C, Esaki N & Soda K (1997) Bacterial D,L-2-haloacid dehalogenase from Pseudomonas sp. strain 113: Gene cloning and structural comparison with Dand L-2-haloacid dehalogenases. J. Bacteriol. 79:4232-4238 37 . Nardi-Dei V, Kurihara T, Park C, Miyagi M, Tsunasawa S, Soda K & Esaki N (1999) D,L-2Haloacid dehalogenase from Pseudomonas sp. 113 is a new class of dehalogenase catalyzing hydrolytic dehalogenation not involving enzyme-substrate ester intermediate. J. Biol. Chem. 274:20977-20981 38. Newman J, Peat TS, Richard R, Kan L, Swanson PE, Affholter JA, Holmes IH, Schindler JF, Unkefer CJ & Terwilliger TC (1999) Haloalkane dehalogenases: Structure of a Rhodococcus enzyme. Biochemistry 38:16105-16114 39. Ohno S (1984) Birth of a unique enzyme from an alternative reading frame of the preexisted,

BACTERIAL GROWTH ON HALOGENATED ALIPHATIC HYDROCARBONS 225 internally repetitious coding sequence. Proc. 51. Stokes HW & Hall RM (1989) A novel family of Natl. Acad. Sci. USA 81:2421-2425 potentially mobile DNA elements encoding site40. Poelarends GJ, Kulakov LA, Larkin MJ, van specific gene-integration functions: Integrons. Mol. Microbiol. 3:1669-1683 Hylckama Vlieg JET & Janssen DB (2000) Roles of horizontal gene transfer and gene 52. Stringfellow JM, Cairns SS, Cornish A & integration in evolution of 1,3-dichloropropeneCooper RA (1997) Haloalkanoate dehalogenase and 1,2-dibromoethane-degradative pathways. II (DehE) of a Rhizobium sp. - Molecular J. Bacteriol. 182:2191-2199 analysis of the gene and formation of carbon monoxide from trihaloacetate by the enzyme. 41. Poelarends GJ, Zandstra M, Bosma T, Kulakov LA, Larkin MJ, Marchen JR, Weightman AJ & Eur. J. Biochem. 250:789-793. Janssen DB (2000) Haloalkane-utilizing 53. Studer A, Vuilleumier S & Leisinger T (1999) Rhodococcus strains isolated from Properties of the methylcobalamin: geographically distinct locations possess a methyltransferase involved in chloromethane utilization by Methylobacterium sp. strain CM4. highly conserved gene cluster encoding haloalkane catabolism. J. Bacteriol. 182: 2725Eur. J. Biochem. 264:242-249 54. Tardif G, Greer CW, Labbe D & Lau PC (1991) 2731 Involvement of a large plasmid in the 42. Poelarends GJ, van Hylckama Vlieg JET, degradation of 1,2-dichloroethane by Marchesi JR, Freitas dos Santos LM & Janssen Xanthobacter autotrophicus. Appl. Environ. DB (1999) Degradation of 1,2-dibromoethane Microbiol. 57:1855-1857 by Mycobacterium sp. strain GP1. J. Bacteriol. 181:2050-2058 55. Trantirek L, Hynkova K, Nagata Y, Murzin A, Ansorgova A, Sklenar V & Damborsky J (2001) 43 . Poelarends GJ, Wilkens M, Larkin MJ, van Reaction mechanism and stereochemistry of Elsas JD & Janssen, DB (1998) Degradation of dehydrochlorinase 1,3-dichloropropene by Pseudomonas cichorii LinA. J. Biol. Chem. 276:7734-7740 170. Appl. Environ. Microbiol. 64 2931-2936 44 . Pries F, Kingma J, Pentenga M, van Pouderoyen 56. Thomas AW, Slater JH & Weightman AJ (1992) The dehalogenase gene dehI from Pseudomonas G, Jeronimus-Stratingh CM, Bruins AP & putida PP3 is carried on an unusual mobile Janssen DB (1994) Site-directed mutagenesis genetic element designated DEH. J. Bacteriol. and oxygen isotope incorporation studies of the 174:1932-1940 nucleophilic aspartate of haloalkane 57. Thomas AW, Topping AW, Slater JH & dehalogenase. Biochemistry 33:1242-1247 Weightman AJ (1992) Localization and 45. Pries F, van den Wijngaard AJ, Bos R, functional analysis of structural and regulatory Pentenga M & Janssen DB (1994) The role of dehalogenase genes carried on DEH from spontaneous cap domain mutations in Pseudomonas putida PP3. J. Bacteriol. 174: haloalkane dehalogenase specificity and 1941-1947 evolution, J. Biol. Chem. 269:17490-17494 46. Ridder IS, Rozeboom HJ, Kalk KH & Dijkstra 58. Topping AW, Thomas AW, Slater JH & Weightman AJ (1995) The nucleotide sequence BW (1999) Crystal structures of intermediates in of a transposable haloalkanoic acid the dehalogenation of haloalkanoates by L-2dehalogenase regulatory gene (dehRI) from haloacid dehalogenase. J. Biol. Chem. Pseudomonas putida strain PP3 and its 274:30672-30678 relationship with activators. 47. Ruiz GM, Rawlings TK, Dobbs FC, Drake LA, Mullady T, Huq A & Colwell RR (2000) Global Biodegradation 6:247-255 spread of microorganisms by ships. Nature 59. Van der Ploeg J, Willemsen M, van Hall G & Janssen DB (1995) Adaptation of Xanthobacter 408:49-50 autotrophicus GJ10 to bromoacetate due to 48. Schmid-Appert M, Zoller K, Traber H, activation and mobilization of the haloacetate Vuilleumier S & Leisinger T (1997) Association dehalogenase gene by insertion element IS 1247. of newly discovered IS elements with the J. Bacteriol. 177:1348-56 dichloromethane utilization genes of methylotrophic bacteria. Microbiology 60. Van Hylckama Vlieg JET & Janssen DB (1992) Bacterial degradation of 3-chloroacrylic acid 143:2557-2567 and the characterization of cis- and trans49. Senior E, Bull AT & Slater JH (1976) Enzyme specific dehalogenases. Biodegradation 2:139evolution in a microbial community growing on the herbicide Dalapon. Nature 263: 476-479. 150 50. Slater JH, Weightman AJ & Hall BG (1985) 61. Vannelli T, Messmer M, Studer A, Vuilleumier S & Leisinger T (1999) A corrinoid-dependent Dehalogenase genes of Pseudomonas putida catabolic pathway for growth of a PP3 on chromosomally located transposable Methylobacterium strain with chloromethane. elements. Mol. Biol. Evol. 2: 557-567

226 Proc. Natl. Acad. Sci. USA 96:4615-20 62. Verce MF, Ulrich RL & Freedman DL (2000) Characterization of an isolate that uses vinyl chloride as a growth substrate under aerobic conditions. Appl. Environ. Microbiol. 66:35353542 63. Verschueren KH, Seljee F, Rozeboom HJ, Kalk KH & Dijkstra BW (1993) Crystallographic analysis of the catalytic mechanism of haloalkane dehalogenase. Nature 363:693-698

JANSSEN ET AL. 64. Wyndham RC, Cashore AE, Nakatsu CH & Peel MC (1994) Catabolic transposons. Biodegradation 5:323-342 65. Yu F, Nakamura T, Mizunashi W & Watanabe I (1994) Cloning of two halohydrin halogenhalide lyase genes from Corynebacterium sp. strain N-1074 and structural comparison of the genes and gene products. Biosci. Biotechnol. Biochem. 58:1451-1457

Chapter 8 AROMATIC DEHALOGENASES: INSIGHTS INTO STRUCTURES, MECHANISMS, AND EVOLUTIONARY ORIGINS SHELLEY D. COPLEY Department of Molecular, Cellular, and Developmental Biology, University of Colorado at Boulder, Boulder, CO, USA

1. INTRODUCTION Chlorinated aromatic compounds are among the most significant environmental contaminants introduced into the environment by human activities. Pesticides such as atrazine, 2,4-D (2,4-dichlorophenoxyacetic acid), and pentachlorophenol are introduced intentionally, while compounds such as PCBs have been released accidentally or by poor disposal practices in the past. Many of these novel compounds are xenobiotic (foreign to life), and therefore microorganisms may not have enzymes capable of transforming them. As expected, many chlorinated aromatic xenobiotics are quite resistant to biodegradation. However, microorganisms capable of degrading such compounds have often been isolated from contaminated sites, suggesting that microorganisms can adapt to the presence of novel compounds by evolving new metabolic pathways that allow them either to take advantage of a new nutrient source or to destroy a toxin. The novel enzymes that participate in newly evolved pathways may be acquired by horizontal transfer from another organism, or by recruitment of a pre-existing cellular protein that has at least a modest ability to catalyze the newly needed reaction. Given sufficient time and selective pressure, genetic changes can occur that will improve the function of these recruited proteins. It is, of course, not clear what length of time is “sufficient”. However, anthropogenic chlorinated aromatics have been present in the environment for less than a century. That period, an eye-blink in the nearly 4 billion-year history of life on earth, is apparently enough to get the process started, but is unlikely to be long enough to generate enzymes of their usual level of sophistication in terms of regulation and catalytic proficiency. Biodegradation of chlorinated aromatic compounds requires removal of chlorine substituents either before or after the aromatic ring is cleaved, although for highly chlorinated compounds such as pentachlorophenol and some PCBs, at least some of the chlorines must be removed before ring cleavage. The aromatic dehalogenases that accomplish the difficult task of removing chlorines from aromatic rings are the subject Dehalogenation: Microbial Processes and Environmental Applications, pages 227-259 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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of this chapter. Three types of aromatic dehalogenases have been identified (Figure 8.1). Hydrolytic dehalogenases catalyze the replacement of a chlorine substituent with a hydroxyl group derived from water. Reductive dehalogenases catalyze the replacement of a chlorine substituent with hydrogen, a reaction that results in reduction of the aromatic ring and concomitant oxidation of a cosubstrate. Figure 8.1 depicts tetrachlorohydroquinone dehalogenase, which uses glutathione as a reductant (84), but other reductive dehalogenases use different reductants. 2,4-Dichlorobenzoyl CoA dehalogenase from Corynebacterium sepedonicum KZ-4 uses NADPH as a reductant (62), and 3chlorobenzoate dehalogenase from Desulfomonile tiedjei DCB-1 is a membrane-bound heme enzyme whose physiological reductant is unknown (53). Finally, oxygenolytic dehalogenases catalyze the replacement of a chlorine substituent with a hydroxyl group derived from Some of these enzymes are dioxygenases that add both oxygen atoms of to the aromatic ring, while others are monooxygenases that add one oxygen of to the aromatic ring and reduce the other to water. The monooxygenase class includes both flavin monooxygenases, which act only upon substrates containing a hydroxyl or amino group, and enzymes, which are less particular. The wealth of genomic and structural data that has been acquired in the last decade now allows us to examine the functions and origins of aromatic dehalogenases in a way that was previously impossible. Knowledge of the amino acid sequence of an enzyme in most cases allows its assignment to a particular superfamily, a group of enzymes derived by divergent evolution from a common progenitor. Proteins in a superfamily share a common structural fold, although they often catalyze very dissimilar transformations. Before the genomic era, there would have been little reason to suspect that many enzymes in a superfamily might be related to each other at all. However, the developing field of genomic enzymology demonstrates that a common structural fold that provides a particular catalytic ability can be elaborated by the addition of other catalytic groups to generate a variety of enzymes capable of catalyzing diverse reactions. Several of the aromatic dehalogenases to be discussed here have been assigned to superfamilies. These assignments allow prediction of structure, potential catalytic residues, and mechanistic features, as well as provide insight into the types of enzymes that may have given rise to novel dehalogenases in response to selective pressure from anthropogenic compounds. One of the most interesting questions regarding the evolution of aromatic dehalogenases is when these enzymes originated. This is an issue of great importance with respect to environmental quality, because microorganisms play a critical role in the degradation of the thousands of chemicals released by agriculture, industry, and household use. In the case of xenobiotic compounds, the ability of microorganisms to evolve new enzymes and metabolic pathways may be the limiting factor in the removal of these compounds from the environment. Many studies of microbial metabolism have shown that microbes, when faced with an “eat this or die” scenario, are often capable of finding a way to survive. Thus, it is certainly possible that new enzymes have evolved during the past few decades. However, in some cases, dehalogenases may have evolved long ago to dehalogenate chlorinated aromatic natural products. A dehalogenase involved in degradation of a natural product might be well-suited for biodegradation of a chlorinated anthropogenic compound, or at least provide a reasonable starting place.

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The question of when dehalogenases originated may never be answered with certainty. However, there are genetic and functional features of at least some aromatic dehalogenases that are consistent with a recent origin. This chapter will review the existing information about selected members of the three classes of aromatic dehalogenases, chosen because there is a substantial body of genetic, mechanistic, and/or structural information that informs an enlightened discussion in the context of the family or superfamily to which the enzyme belongs. The following questions will be emphasized: 1. How do the enzymes catalyze the very difficult reactions required to remove chlorine substituents from an aromatic ring? 2. Where have the enzymes come from? 3. When were they recruited to serve as aromatic dehalogenases? 4. How well are they serving their function in the degradation of anthropogenic chlorinated aromatic compounds?

2. HYDROLYTIC DEHALOGENASES 2.1. 4-Chlorobenzoyl CoA Dehalogenase 4-Chlorobenzoyl CoA dehalogenase is the best understood aromatic dehalogenase. A crystal structure of the enzyme from Pseudomonas sp. strain CBS3 is available (9), and many years of mechanistic and spectroscopic studies have provided detailed insights into the means by which its active site promotes this very difficult chemical reaction. Furthermore, the relationship of this enzyme to its superfamily and the special adaptations to the superfamily active site that allow this particular transformation to occur are now well understood. 4-Chlorobenzoyl CoA dehalogenase catalyzes the hydrolytic dechlorination of 4chlorobenzoyl CoA during the degradation of 4-chlorobenzoate by a number of soil bacteria, including Pseudomonas sp. strain CBS3 (42, 66), Arthrobacter sp. strain SU (65) and Arthrobacter sp. strain 4-CB1 (formerly designated Acinetobacter sp. strain 4CB1) (17). 4-Chlorobenzoate is a product formed during degradation of certain PCBs. Arthrobacter sp. strain 4-CB1 was isolated from PCB-contaminated soil (1), but, interestingly, Pseudomonas sp. CBS3 (40) was not. The pathway for degradation of 4chlorobenzoate (Figure 8.2) begins with the conversion of 4-chlorobenzoate to 4chlorobenzoyl CoA in a reaction catalyzed by 4-chlorobenzoate:Coenzyme A ligase. Subsequently, 4-chlorobenzoyl CoA dehalogenase replaces the chlorine at the 4-position with a hydroxyl group derived from The CoA thioester is hydrolyzed in the third step. The transient formation of a CoA thioester which is hydrolyzed only one step after its formation is rather odd, since the formation of the high-energy thioester bond requires the hydrolysis of an ATP molecule, and this might appear to be a waste of chemical energy. Formation of CoA thioesters is not required for degradation of all chlorobenzoates–for example, 3-chlorobenzoate is degraded without formation of CoA thioesters (34). In contrast, certain benzoates are in anaerobes converted to CoA thioesters at the beginning of the pathway and the thioester is left in place, apparently for the entire degradation pathway (27). In the case of 4-chlorobenzoate degradation, the CoA thioester is specifically required for effective dehalogenation. The reasons for this

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will become apparent in the following discussion of the structure and mechanism of the dehalogenase. Since the dehalogenation reaction can not occur on 4-chlorobenzoate itself, the expenditure of one molecule of ATP is warranted because it allows the molecule to be degraded and used as a source of carbon and energy. 4-Chlorobenzoyl CoA dehalogenase is a member of the 2-enoyl-CoA hydratase/isomerase superfamily (79). The superfamily is named for the functions of the prototypical members of the superfamily, although there is considerable diversity among the reactions catalyzed by other members. The common catalytic strategy among the members of the superfamily is the polarization of the thioester carbonyl in order to facilitate a reaction involving development of negative charge in the carbonyl. Such reactions include nucleophilic attack at the carbonyl itself or at a position in conjugation with the carbonyl, proton abstraction from the position, or carbon-carbon bond cleavage between the and carbons. In some members of the superfamily, additional steps have been incorporated before or after the step involving delocalization of the negative charge into the carbonyl. Figure 8.3 shows examples of reactions catalyzed by several members of the superfamily. Structures are now available for four superfamily members, 4-chlorobenzoyl CoA dehalogenase (9), 2-enoyl CoA hydratase (crotonase) (28), CoA isomerase (52), and methylmalonyl CoA decarboxylase (7). These proteins are made up of monomers with a common structural fold (Figure 8.4), although they differ in quaternary structure and, in the case of methylmalonyl CoA decarboxylase, in the disposition of the final two that form the C-terminal domain (see Figure 8.4c). The active sites in this superfamily are specialized to provide an oxyanion hole for binding and polarization of the carbonyl group of the thioester substrate, as well as a CoA binding pocket. The range of catalytic abilities found in the superfamily is made possible by alterations in an expandable binding pocket which accommodates the acyl group, and variations in catalytic groups positioned on several segments of the protein which line the active site (79). The mechanism of 4-chlorobenzoyl CoA dehalogenase is shown in Figure 8.5a (9, 18, 85). It begins with attack of the Asp 145 residue upon 4-chlorobenzoyl CoA, followed by expulsion of chloride to form an aryl-enzyme intermediate. Subsequently,

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the aryl-enzyme intermediate is hydrolyzed via attack of a water molecule. His90 acts as a general base to remove a proton from water, and of Trpl37 appears to stabilize the oxyanion resulting from attack of water via a hydrogen bonding interaction. The presumed tetrahedral intermediate decomposes to give 4-hydroxybenzoate and to regenerate the free enzyme. The mechanistic and structural information described above provides insight into some intriguing aspects of this reaction. First of all, the replacement of chlorine on an aromatic ring by a hydroxyl group is an extremely difficult reaction. Nucleophilic aromatic substitution reactions that proceed by direct attack of the nucleophile inevitably

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require multiple strongly electron-withdrawing substituents (such as nitro groups) to stabilize the negative charge resulting from nucleophilic attack. Pseudomonas sp. CBS3 successfully overcomes this difficulty using several ingenious strategies. First, 4chlorobenzoate is converted to 4-chlorobenzoyl CoA prior to the dehalogenation reaction because thioesters are more strongly electron-withdrawing than are carboxylates. Second, the oxyanion hole at the active site strongly polarizes the carbonyl of the substrate by interactions with the backbone amides of Gly 114 and Phe64 and with the helix dipole originating from residues 114-121 (9). This polarization has been demonstrated by Raman and NMR studies of complexes of the enzyme with product or

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substrate analogues. The C=O stretching frequency of the substrate analogue 4methylbenzoyl CoA is shifted from in aqueous solution to in the active site. This polarization is equivalent to that which would be provided by hydrogen bonds worth (14). Following attack of the nucleophile, the same interactions stabilize the Meisenheimer complex intermediate in which the negative charge is largely delocalized into the carbonyl. Third, the efficacy of this polarization is likely to be increased by the hydrophobic active site, which creates a low dielectric environment in which electrostatic interactions are strengthened. Fourth, the enzyme positions the nucleophilic carboxylate in an appropriate position to attack the ring, thus lowering the for the reaction, and probably also promotes the nucleophilicity of the carboxylate by removing intervening water molecules that might hinder nucleophilic attack. Finally, the need for a second energetically-demanding nucleophilic aromatic substitution reaction to generate the product is avoided by the use of a carboxylate as a nucleophile; a simple ester hydrolysis suffices to form the product and regenerate the free enzyme. Of these strategies, the extremely important polarization of the carbonyl is provided by the generic superfamily oxyanion hole. The carboxylate nucleophile (Asp145) and the residues that assist in the ester hydrolysis (His90 and Trp137) are specific adaptations of the generic superfamily active site to allow catalysis of this

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particular reaction. For purposes of comparison, Figure 8.5b shows the mechanism of 2-enoyl CoA hydratase (crotonase), which againutilizes the generic superfamily strategy of polarization of the carbonyl via a structurally identical oxyanion hole, but utilizes a different set of catalytic residues (Glu164 and Glu 144) to catalyze the syn addition of water across a double bond. These two reactions are excellent examples of how an active site with a particular capability (in this case, the polarization of a carbonyl in an oxyanion hole) can be tailored by the addition of different catalytic groups, to allow catalysis of quite different overall transformations. The substrate specificity of the enzyme is easily understood in the context of the structure and mechanism discussed above. Although the enzyme can efficiently remove chlorine, bromine, or iodine from the 4-position, it cannot remove halogens from either the 2- or 3-position (41). In order to utilize the carbonyl of the thioester as an electron sink, nucleophilic attack must occur at either the 4- or the 2- position. This mechanistic strategy could not be used to dehalogenate 3-chlorobenzoate, and, indeed, 3chlorobenzoate is degraded in aerobes by a different series of reactions in which the aromatic ring is cleaved before the chlorine is removed (34). Furthermore, a nucleophile positioned to attack position 4 would not be able to reach position 2. In theory, hydrolytic dehalogenation at the 2-position should be possible from an electronic standpoint, but it would require further adaptation of the active site by movement of the catalytic nucleophile into a different position. A final interesting aspect of this enzyme is that its rate is rather slow: under steady state conditions is It appears from pre-steady state kinetic studies that

the initial nucleophilic aromatic substitution reaction, which might be expected to be the most difficult part of the reaction, is not rate-limiting. The rate constant for the nucleophilic attack of Asp 145 upon the substrate is and the rate constant for the loss of chloride is The rate of hydrolysis ofthe aryl-enzyme intermediate is quite slow, with a rate constant of The rate is actually limited by the slow rate of product release (D. Dunaway-Mariano, personal communication). It is especially interesting that the ester hydrolysis step is so slow. The intrinsic difficulty ofthe reaction is probably not the cause. Many enzymes are known to catalyze ester hydrolysis with substantially higher rates. For example, acetylcholinesterase cleaves its substrate to acetate and choline with a turnover number ofgreater than (4). The explanation may lie in the requirement for Asp 145 to serve as a nucleophile in the first step, and as an electrophile in the ester hydrolysis step. It may not be possible for Asp145 to “multitask” with optimum effectiveness in these two different roles (D. Dunaway-Mariano, personal communication). Enhancement of the electrophilicity of the carbonyl of Asp 145 by interaction with an oxyanion hole in order to facilitate the ester hydrolysis would decrease its nucleophilicity and diminish its ability to attack 4-chlorobenzoyl CoA in what is intrinsically a more difficult reaction.

2.2 Atrazine Chlorohydrolase Atrazine chlorohydrolase catalyzes the hydrolytic dehalogenation of atrazine, a chlorinated heteroaromatic herbicide used to control broad leaf weeds in corn and sorghum fields. This enzyme catalyzes the first step in the degradation of atrazine by Pseudomonas sp. ADP (Figure 8.6) (21). Subsequent steps, which remove the N-

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alkylamine substituents, are similar hydrolytic reactions (11, 63). Atrazine is believed to be a xenobiotic compound, and therefore atrazine chlorohydrolase has presumably been recruited recently to catalyze the dehalogenation reaction. There are numerous heteroaromatic natural products – most notably purines and pyrimidines – and enzymes that catalyze substitution reactions on such compounds are known. Thus, there is a considerable repertoire of enzymes capable of catalyzng this type of transformation, and the ability to degrade atrazine likely requires a rather simple adjustment in the active site to accommodate the peculiar shape of atrazine, as well as possible adaptations to facilitate the departure of the chloride leaving group. Although a number of microorganisms are capable of completely degrading atrazine, in natural ecosystems it is likely that consortia will be involved in atrazine degradation. Wackett and co-workers have analyzed the degradation of atrazine by a four-membered bacterial consortium enriched from agricultural soil that had been treated with atrazine for 15 years (20). One member of the consortium, Clavibacter michiganensis ATZ1, converted atrazine to N-ethylammelide, while a second member, Pseudomonas sp. strain CN1, converted N-ethylammelide to cyanuric acid and then further metabolized the cyanuric acid. The role of the additional two unidentified members may have been to metabolize the alkylamines released from atrazine, which could otherwise lead to a harmful increase in the pH of the medium. It is interesting that the order of steps in the pathway carried out by the consortium is different from that in Pseudomonas strain ADP. All of the steps involve essentially the same type of chemical reaction, and it apparently does not matter which substituent is removed first. Thus, the order probably simply reflects the chance occurrence of different adjustments to a similar active site pocket to accommodate different substituents on the heteroaromatic ring. The mechanism, structure and evolutionary origin of this enzyme have not been investigated as thoroughly as those of 4-chlorobenzoyl CoA dehalogenase. However, the superfamily approach provides important insights into each of these issues. Wackett and coworkers recently reported that atrazine chlorohydrolase (AtzA), and AtzB and AtzC as well, are members of the amidohydrolase superfamily (63). Again, the superfamily is named for the activity of some of the prototypical members, but a range of chemical transformations is catalyzed by members of the superfamily. This superfamily was identified by Holm and Sander in 1997 (36) by comparison of the structures of urease, phosphotriesterase, and adenosine deaminase. The characteristic superfamily fold

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comprises a common barrel domain, consisting of eight alternating strands and alpha helices, with the strands forming a compact core surrounded by the helices. A metal-binding site is formed by several residues at the C-terminal ends of four of the strands (see Figure 8.7). The residues involved in the metal-binding site are few in number and are widely separated in the primary sequence, so it would have been difficult to recognize this superfamily from sequence data in the absence of structural information. In some members of the superfamily, such as adenosine deaminase, a single metal is bound in the active site pocket, while in others, such as urease and phosphotriesterase, two metal ions are bound with the assistance of a carbamoylated lysine that bridges the two metal centers. The primary catalytic activity that is supported by this active site structure appears to be the activation of a metal-bound hydroxide for nucleophilic attack upon a substrate. A nucleophilic hydroxide situated at such a metal center can be used to attack a variety of substrates. Thus, the specificity of the enzyme will be dictated by the shape of the active site, and by additional interactions that can stabilize the transition state resulting from hydroxide attack and which depend on the particular characteristics of the substrate. Some of these interactions involve the active-site metal ions. In phosphotriesterase, the more solvent-exposed zinc ion coordinates the phosphoryl oxygen of the substrate, thus activating the phosphorous for attack by the metal-bound hydroxide (8, 61). In urease, both metals are believed to participate in substrate binding in order to polarize the substrate (13, 58). Reactions catalyzed by several members of the superfamily are shown in Figure 8.8.

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The transformation catalyzed by atrazine chlorohydrolase is strikingly similar to that catalyzed by adenosine deaminase. In both cases, the enzymes catalyze the substitution of a hydroxyl group for a substituent on a heteroaromatic ring. The mechanism of adenosine deaminase (Figure 8.9) is well understood, particularly because of a crystal structure of the enzyme in complex with 6-hydroxyl-1,6-dihydropurine ribonucleoside, an analogue of the intermediate that would normally be formed by attack of the metalbound hydroxide on adenosine (Figure 8.10) (78). The mechanism begins with attack of the zinc-bound hydroxide at the C-6 position of adenosine. Either concomitant with or after hydroxide attack, N-6 is protonated by Glu217. Subsequently, elimination of in either a stepwise or concerted process, generates the product. Adenosine deaminase uses three primary strategies to accelerate the deamination reaction. First, binding of the substrates at the active site prior to reaction significantly decreases the

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unfavorable for the hydration reaction, which is nearly -10.5 kcal/mol for the nonenzymatic reaction (39). Second, the nucleophilicity of the attacking water molecule is increased substantially by its interactions with the active site zinc, as well as with Asp295, which is proposed to act as a general base. Third, Glu217 provides a proton to the N-l position of adenosine; subsequently, the carboxylate of Glu217 forms a hydrogen bond to the protonated N-1 which is estimated by Raman spectroscopy to be worth 4-10 kcal/mol (25). The mechanistic strategy used by adenosine deaminase should certainly be applicable to other substitution reactions involving different leaving groups on heteroaromatic rings. A likely mechanism for atrazine chlorohydrolase is shown in Figure 8.9b. Support for this proposed mechanism is shown in Figure 8.11, which showsa motif found in a set of proteins including atrazine chlorohydrolase and several homologues of unknown function, as well as adenosine, guanine and cytosine deaminases. Highlighted in grey are the histidine residue corresponding to His214 of adenosine deaminase, which is one of the ligands to the active site zinc, and the glutamate residue corresponding to Glu217, which is the critical active site general acid that protonates N-1 of adenosine. The conservation of this Glu in this set of proteins is consistent with the hypothesis that they may all function in a manner similar to that of adenosine deaminase. Homologues ofadenosine deaminase, including atrazine chlorohydrolases and other nucleobase deaminases, are found in diverse organisms from all three kingdoms of life, suggesting that catalysis of a nucleophilic substitution reaction on a heteroaromatic substrate is an ancient catalytic strategy built into the versatile barrel protein scaffold. The adaptation of the active site to bind different substrates while retaining the basic catalytic machinery has apparently allowed diversification to provide enzymes capable of catalyzing many different transformations. AtzB and AtzC, the two additional enzymes that remove substituents from atrazine, are two examples of the utilization of

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this catalytic, strategy using substrates that differ only slightly in structure in atrazine. Since atrazine is a xenobiotic compound but is closely related to natural products, it is interesting to consider when and how atrazine chlorohydrolase arose. Bacteria capable of mineralizing atrazine were not reported until 1993 (45), although atrazine had been in use since 1959. Thus, atrazine chlorohydrolase may have arisen over the relatively short period of time since 1959, although it is impossible to be sure. However, it is certainly clear that a few genes involved in atrazine degradation have spread globally in a very short period of time. Wackett and coworkers have analyzed the sequences of atzA, atzB and atzC genes from several different atrazine-degrading bacteria, including strains of Pseudomonas, Alcaligenes, Ralstonia, and Agrobacterium (22). Despite the fact that these strains represent different bacterial genera and were isolated from locations scattered all over the world, the genes are 99-100% identical. This astonishing finding can only be explained by horizontal transfer of these genes; independent evolution in such diverse species and locations could not have resulted in such a high degree of identity. Thus, it appears that the release of over 40 billion pounds

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of atrazine in the past few decades has provided strong selective pressure for the evolution of atrazine-catabolizing enzymes and their rapid spread in the environment. Although we have come to expect rapid dispersal of antibiotic resistance genes, the extraordinary rapidity of the dispersal of these degradative genes is intriguing and profoundly important. The rapid spread of the atrazine catabolism genes among bacteria is likely due to their location on mobile genetic elements that are readily transferred between bacteria. In Pseudomonas strain ADP, atzA, atzB and atzC are encoded on a 96kb self-transmissable plasmid (pADP-1) (23). Notably, atzA, atzB and atzC are physically distinct and not encoded by an operon. pADP-1 can be transferred easily from Pseudomonas strain ADP to E. coli, as well as several Gram-negative soil bacteria. However, the spread ofthe atrazine catabolism genes is clearly not due to simple transfer of pADP-1, since the genes are found on plasmids of different sizes in several atrazinedegrading bacteria (L. Wackett, personal communication), Furthermore, atzA is flanked by insertion sequences in several of these cases, suggesting that transposable elements may be involved in the movement of these genes (47).

3. A REDUCTIVE DEHALOGENASE: TETRACHLOROHYDROQUINONE DEHALOGENASE Tetrachlorohydroquinone (TCHQ) dehalogenase catalyzes the reductive dehalogenation of TCHQ and trichlorohydroquinone (TriCHQ) during the degradation of pentachlorophenol (PCP) by the Gram-negative soil bacterium Sphingomonas chlorophenolica (Figure 8.12) (82). The reducing equivalents for each step are provided by two molecules of glutathione, which are oxidized to glutathione disulfide (84). TCHQ dehalogenase is a particularly intriguing enzyme for two reasons. First, PCP was first introduced as a pesticide in 1936 (15). Thus, it is likely that the pathway for degradation of PCP has been assembled quite recently, and some of the enzymes, particularly those in the early stages of the pathway, may have been recently recruited to serve new functions in this evolving pathway. Secondly, reductive aromatic dehalogenation reactions are uncommon in aerobic microorganisms. Anaerobic bacteria are known to catalyze a variety of reductive dehalogenation reactions, but they generally employ transition metal cofactors; TCHQ dehalogenase has no transition metal cofactors (51). Thus, the origin of TCHQ dehalogenase is a particularly intriguing puzzle. TCHQ dehalogenase is a member of the glutathione S-transferase (GST) superfamily (51, 54). The generic reaction promoted by enzymes in this superfamily is the nucleophilic attack of glutathione upon an electrophilic substrate to form a glutathione conjugate (Figure 8.13a). Many of the enzymes in this superfamily, particularly those involved in detoxification reactions, catalyze just this simple reaction. However, in a few cases, additional steps have been added before and/or after the nucleophilic attack of glutathione to facilitate a more complex transformation. Examples of enzymes in the superfamily that catalyze such reactions are shown in Figure 8.13 b-f. (Note that in the case of dichloromethane dehalogenase presented in Figure 8.13c, the second reaction following the typical GST step is probably non-enzymatic). A crystal structure of TCHQ dehalogenase is not yet available, but its membership in the GST superfamily allows a reasonable prediction of its structure. Crystal structures have been solved for members of several classes of the superfamily, and the overall

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structural fold is strikingly similar throughout the superfamily (5). A structure of the monomer of the human theta class GST is shown in Figure 8.14, in order to demonstrate the typical structural features of the GSTs. Most GSTs are dimers, although TCHQ dehalogenase is not (48). The monomers contain two domains. The N-terminal domain provides most of the binding contacts for glutathione, while the C-terminal domain provides most of the binding contacts for the electrophilic substrate. Much of the catalytic power of the GSTs comes from their ability to lower the of glutathione in the active site from 9 to 6.5 - 7, thus potentiating its nucleophilicity. The thiolate of the glutathione is stabilized in the active site in most classes by a hydrogen bonding interaction with the hydroxyl group of a nearby Ser or Tyr (5). TCHQ dehalogenase is one of the members of the superfamily in which additional steps before and after the typical GST step allow catalysis of a more complex transformation. The current working model for the mechanism of TCHQ dehalogenase is shown in Figure 8.15 (S. Copley, unpublished). The typical GST step is boxed. The steps preceding the GST step can be considered “substrate preparation” steps that convert the relatively non-electrophilic TCHQ to the very electrophilic tetrachlorobenzoquinone. This conversion is postulated to occur via an initial ketonization step, followed by 1,4-elimination of HCl. Nucleophilic attack of glutathione upon the electrophilic tetrachlorobenzoquinone results in formation of a glutathione conjugate in a step characteristic of GSTs. The subsequent steps convert the glutathione conjugate, which would be the product of most GST reactions, further into a reduced

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product. Cys13 attacks the sulfur of the glutathionyl substituent, releasing the reduced product, TriCHQ, and forming a mixed disulfide between Cys13 and glutathione. In the final step, a thiol-disulfide exchange reaction with a second molecule of glutathione regenerates the free enzyme and forms glutathione disulfide. The steps involving Cys13

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in the later part of the reaction are well characterized (50). At this point, the initial steps in the model are consistent with all of the data, but not uniquely so, and investigations of the mechanism are continuing. Recently it was recognized that TCHQ dehalogenase is a member of the zeta class of the GST superfamily (10). This class is distinguished primarily by a highly conserved and distinctive region in the N-terminal domain in the active site region (Figure 8.16). Three members of this class are known maleylacetoacetate isomerases, enzymes that catalyze the glutathione-dependent isomerization of a double bond from cis to trans during the degradation of phenylalanine and tyrosine in many, but not all, species. The reaction catalyzed by maleylacetoacetate isomerase is shown in Figure 8.13d. Several bacterial zeta class GST genes are also likely to encode maleylacetoacetate isomerases based upon their proximity to another gene involved in tyrosine degradation. The zeta

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class also includes a bacterial maleylpyruvate isomerase that is involved in a similar transformation during degradation of benzoate. The striking similarity between the active site regions ofTCHQ dehalogenase and these isomerases prompted Anandarajah and coworkers to assay the dehalogenase for isomerase activity (2). Maleylacetone, a substrate analogue that is more stable than maleylacetoacetate because it lacks the beta carboxylate group that tends to undergo decarboxylation, was used for these assays. TCHQ dehalogenase does indeed have considerable maleylacetone isomerase activity, nearly equivalent to that of a bona fide bacterial maleylacetoacetate isomerase from Vibrio sp. 01(70,71). The for both enzymes is about and the for TCHQ dehalogenase is only four-fold lower than that for the Vibrio sp. MAA isomerase Furthermore, both activities occur at the same active site, since mutation of both Ser11 and Cysl3 affects both reactions. This finding is particularly important for two reasons. First, it is astounding that the same active site can catalyze such different reactions. This is an unusual and possibly unprecedented level ofcatalytic promiscuity. Experiments are underway to elucidate how the active site participates in both reactions. Insights into this question will greatly enhance our understanding ofhow the generic chemistry provided by the superfamily fold can be elaborated to allow catalysis of new and different reactions. Second, the high level of isomerase activity is consistent with the possibility that TCHQ dehalogenase was recruited from a preexisting isomerase (ofeither the maleylacetoacetate or maleylpyruvate variety) that may have had some dehalogenase activity as a consequence of the particular arrangement of catalytic groups in its active site. The recruitment of a glutathione-dependent double bond isomerase to serve as a reductive dehalogenase illustrates the important point that novel enzymes can arise from sources we might never suspect based upon the chemical reactions catalyzed. In many cases, novel enzymes involved in xenobiotic degradation are likely to have arisen from broad-specificity enzymes that catalyze the same type of chemical reaction on multiple substrates, and may be modified to favor a novel substrate. In the case of TCHQ dehalogenase, a suitable reductive dehalogenase precursor was likely not available, since reductive aromatic dehalogenases are uncommon in aerobes. Thus, the reductive dehalogenase required to carry out the critical dehalogenation steps was apparently recruited from a different source, one that would not have been suspected based upon the type of transformation catalyzed by these glutathione-dependent double bond isomerases. Clearly, nature will capitalize on anything that works, and the “modular” design of the GST superfamily active site is well–suited for modifications. The regions involved in binding glutathione and optimizing its nucleophilicity can be held constant, while other regions involved in binding the electrophilic substrate and providing additional catalytic groups can be varied without affecting the nucleophile. The finding that TCHQ dehalogenase likely originated from a glutathione-dependent double-bond isomerase brings up additional interesting questions regarding the evolution of the dehalogenase. First, why does the enzyme retain such a high level of isomerase activity? The isomerase activity is not required for tyrosine degradation, since the bacterium has another maleylacetoacetate isomerase that is induced in response to tyrosine (2). It also seems unlikely that efficient isomerase activity would be an accidental consequence of an active site that was truly specialized to provide reductive dehalogenation. The most likely explanation appears to be that the enzyme has not yet

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evolved to be an optimal dehalogenase, either because of lack of time or lack of sufficient selective pressure. A second interesting question focuses on the quality of function of the dehalogenase. If the enzyme is indeed in the process of evolving, then it might not be expected to be an optimal catalyst. Certain properties of the enzyme are indeed rather unusual in terms of our usual expectations for bacterial metabolic enzymes. The enzyme is subject to profound inhibition by its aromatic substrate (S. Copley, unpublished). The substrate inhibition appears to arise because an enzyme with one glutathione binding site (which is typical of GSTs) has been recruited to catalyze a reaction requiring two equivalents of glutathione. As shown in Figure 8.17, the aromatic substrate and the second glutathione, which lacks its own binding site, appear to compete for access to the active site at the stage of the mixed-disulfide between glutathione and Cysl3. Binding of the aromatic substrate at this stage sequesters the enzyme as a dead-end complex. Although it is unusual for an enzyme to neglect to bind one of its substrates, and the substrate inhibition is quite severe, these apparently sub-optimal characteristics do not appear to have any biological consequences. The first step in the pathway, the conversion of PCP to TCHQ, is so slow that only very low levels of the aromatic substrate occur inside the cell (49), so that substrate inhibition should be minimal in vivo. Furthermore, there is no selective pressure to improve the performance of the dehalogenase because the first step in the pathway is rate-limiting. Finally, as with the other dehalogenases discussed above, it would be interesting to know whether the recruitment of an isomerase to serve as a dehalogenase occurred during the last century in response to the introduction of a xenobiotic compound, or whether it occurred long ago in response to some unidentified chlorinated aromatic natural product. The dehalogenase is clearly not a highly evolved enzyme. The high

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level of residual isomerase activity and the profound substrate inhibition clearly suggest a “work in progress”. Furthermore, TCHQ dehalogenase expression is not induced in response to pentachlorophenol in conjunction with the first and third enzymes in the pathway (54, 56, 81). This finding suggests that the regulatory apparatus is also in an immature stage. These catalytic and regulatory defects are certainly consistent with a recent recruitment of the isomerase to serve as a dehalogenase, but the possibility that the recruitment occurred long ago but has not been under sufficient selective pressure to improve the regulation and function of the enzyme cannot be ruled out.

4. AN OXYGENOLYTIC DEHALOGENASE: PENTACHLOROPHENOL HYDROXYLASE PCP hydroxylase catalyzes the first step in the degradation of PCP in Sphingomonas chlorophenolica (Figure 8.12), the conversion of PCP to TCHQ. The reaction requires and two equivalents of NADPH (83). The enzyme contains a flavin cofactor. Notably, the Gram-positive microorganism Mycobacterium chlorophenolicum also converts PCP to TCHQ, but utilizes a monooxygenase rather than a flavindependent monooxygenase (35, 74, 75). As mentioned above, PCP is not known to occur in nature, so it is likely that PCP hydroxylase has been recently recruited to catalyze this hydroxylation reaction. However, phenols are ubiquitous natural products, and naturally occurring chlorinated phenols containing up to four chlorines have been identified (72). Many microorganisms are capable of degrading such compounds, so it is reasonable to expect that the genetic information for a catalytically promiscuous enzyme capable of dealing with PCP might have been readily available. PCP hydroxylase is not very closely related to any protein in the database. It is only 27% identical to the most closely related protein, an oxygenase involved in biosynthesis of the antitumor polyketide antibiotic mithramycin in Streptomyces argillaceus. However, its sequence and cofactor requirements indicate that it belongs to a very divergent family of FAD-dependent monooxygenases that hydroxylate phenols. Crystal structures are available for two of these proteins, phenol hydroxylase from Trichosporon cutaneum (29) and p-hydroxybenzoate hydroxylase from Pseudomonas fluorescens (32, 68,69). Phenol hydroxylase and p-hydroxybenzoate hydroxylase have strikingly similar structures. Three hundred residues can be superimposed with an r.m.s. deviation of only 1.9 Å, even though the sequence identity is only 20% for the structurally aligned residues (29). The key structural differences are the presence of a third domain in phenol hydroxylase which adopts a thioredoxin-like fold and is involved in dimer-dimer interactions that hold the tetramer together, and a lid consisting of residues 170-210 in phenol hydroxylase that sequesters the active site from solvent during the catalytic cycle. The structure of phenol hydroxylase, which is the closest structurally characterized relative of PCP hydroxylase, is shown in Figure 8.18. Like phenol hydroxylase, PCP hydroxylase contains a C-terminal region that is not found in p-hydroxybenzoate hydroxylase, but this C-terminal region bears no resemblance to any sequence in the database, including that of phenol hydroxylase. Of course, this lack of detectable homology does not preclude a similar structural fold, and it will be interesting to discover whether this domain in PCP hydroxylase also adopts a thioredoxin-like fold and is involved in quaternary structure interactions.

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Several of the enzymes in this family, including p-hydroxybenzoate hydroxylase, phenol hydroxylase, anthranilate hydroxylase, and melilotate hydroxylase, have been characterized (76), and functions for others have been suggested by genetic analyses. Figure 8.19 shows a number of reactions catalyzed by members of this family of proteins. These enzymes replace a hydrogen ortho or para to a hydroxyl group (or, rarely, an amino group), with another hydroxyl group derived from Several, including PCP hydroxylase, will replace halogen, nitro, and amino groups, as well as hydrogen (59, 73, 83). Since phenols are both abundant and diverse in nature, it is not surprising that an active site capable of hydroxylating a phenol has diverged to allow binding and transformation of many different substrates. These versatile enzymes serve a variety of functions in soil microorganisms. They are found in bacterial pathways for degradation of many phenols, including some chlorinated phenols. In streptomycetes, they are involved in the synthesis of a wide variety of antibiotics, including urdamycin (31), mithramycin (43), tetracenomycin (24), landomycin (77) mitomycins (46), and chlortetracycline (19). (Note that in some of these cases, the involvement of a flavin hydroxylase is inferred from sequence homology, and the function of the putative hydroxylase has not been demonstrated.) Conversely, recent evidence suggests that a flavin monooxygenase in Rhodococcus equi may hydroxylate rifampin, leading to resistance to this clinically important antibiotic (3). The mechanisms of flavin monooxygenases, and p-hydroxybenzoate hydroxylase in particular, have been studied in great detail. The model for the mechanism of PCP hydroxylase shown in Figure 8.20 is based directly upon this large body of mechanistic and kinetic work. The reaction is proposed to begin with binding of the substrate to the enzyme bearing an oxidized flavin at the active site (step 1). Once the substrate is bound, the flavin can be reduced by NADPH (step 2). The reduced flavin then reacts with to form a C(4a)hydroperoxyflavin (step 3). The hydroxylation step occurs via nucleophilic attack of the substrate upon the C(4a)hydroperoxyflavin (step 4). Subsequently, the hydroxylated substrate eliminates HCl, and a proton is transferred to

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the flavin to form the C(4a)hydroxyflavin intermediate (step 5). The tetrachlorobenzoquinone formed is reduced non-enzymatically by NADPH. The C(4a)hydroxyflavin eliminates water to form the oxidized flavin, thus completing the catalytic cycle (step 6). One of the critical aspects of catalysis in flavin monooxygenases is the control ofthe reactivity of the C(4a)hydroperoxyflavin, which can undergo reaction with solvent to form oxidized flavin (thus wasting an equivalent of NADPH) and hydrogen peroxide. In order to prevent this side reaction, the oxidized flavin in phenol hydroxylase and p-hydroxybenzoate hydroxylase is prevented from reacting with NADPH to form the reduced flavin until substrate is present at the active site (26, 38). Consequently, reaction of the reduced flavin with to form the C(4a)hydroperoxyflavin occurs only when substrate is present. Furthermore, in both enzymes, the C(4a)hydroperoxyflavin is sequestered from solvent so that it can only undergo reaction with the phenol at the active site, although the timing of the movement ofthe flavin from the “out” to the “in” conformation may be different in these two cases (29, 57). Whether PCP hydroxylase also protects the C(4a)hydroperoxyflavin is not known, but it is clear from the sequence that PCP hydroxylase lacks the lid region that is found in phenol hydroxylase. PCP hydroxylase may catalyze the rate-limiting step for the degradation of PCP in S. chlorophenolica (49). Levels of PCP, TCHQ, TriCHQ and DCHQ in S. chlorophenolica Strain RA-2 have been measured in cells that were actively metabolizing The cells accumulated very high levels of PCP, undoubtedly primarily in the membranes. In contrast, the levels of the chlorinated hydroquinones

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were very much lower. These data suggest that the conversion of PCP to TCHQ is the rate-determining step for PCP biodegradation in vivo. Either mass transport of PCP from the membrane into the cytoplasm or the actual chemical conversion of PCP to TCHQ might limit the rate. If the chemical conversion of PCP to TCHQ is rate-limiting, the poor catalytic performance of the enzyme may be due to one or more of three factors. First, the reaction may be intrinsically more difficult than typical phenol hydroxylation reactions. This possibility is difficult to assess at this point. The multiple electron-withdrawing chlorines should make the ring less nucleophilic than other phenols. On the other hand, the multiple chlorine substituents also lower the pKa of the hydroxyl group to 3.5, so that the proton does not have to be removed in order to hydroxylate the ring. Second, the enzyme may be rather ineffective because of its broad substrate specificity. PCP hydroxylase is able to turn over a broad range of substrates bearing variable numbers of halogen substituents, and can replace nitro, amino, cyano, and hydrogen substituents, as well as all of the halogens, with a hydroxyl group (83). The broad substrate specificity implies a capacious active site with relatively relaxed binding interactions. Since enzymes gain much of their catalytic power by immobilizing their substrates, thus decreasing the entropy of activation and orienting the substrate optimally with respect to catalytic groups, it is to be expected that a non-specific enzyme would be rather slow. The third possibility is that the enzyme is in the process of evolving to become a specific PCP hydroxylase, and has not yet had time to optimize this particular activity. The gene encoding PCP hydroxylase is part of an operon that also encodes the third enzyme in the pathway, DCHQ dioxygenase (PcpA), a putative regulatory protein (PcpR), and a protein of unknown function (PcpD) (55). Expression of PCP hydroxylase and DCHQ dioxygenase is induced by PCP. Clearly, these two proteins are functionally linked. Preliminary data suggest that DCHQ is the best substrate for DCHQ dioxygenase (80). These findings suggest that these enzymes may have originally been designed to degrade a naturally occurring phenol such as 2,6-dichlorophenol (the sex pheromone of the female Lone Star tick) (72). A model for the “patchwork” assembly of the PCP degradation pathway from components of two pre-existing pathways has recently been proposed (Figure 8.21) (16). A dichlorophenol hydroxylase may have been recruited to catalyze the chemically similar hydroxylation of the more highly substituted phenol, PCP. It appears (see above) that a reductive dehalogenase recruited from a glutathione-dependent double bond isomerase was patched in next to convert TCHQ to TriCHQ and then to DCHQ. At this point, the pathway may have tied back into the pre-existing pathway. Thus, PCP hydroxylase may be another example of a relatively easy type of recruitment of a preexisting enzyme to accommodate a substrate with a different shape and consequently somewhat different electronic properties, but utilizing the original catalytic machinery to carry out a comparable chemical transformation.

5. CONCLUSIONS Based upon the examples of aromatic dehalogenases discussed here, some principles regarding the relationship between structural features and the mechanism of dehalogenation can be deduced. Hydrolytic dehalogenation reactions require the presence of a group in either the ortho or para position that can act as an electron sink.

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In the case of 4-chlorobenzoyl CoA dehalogenase, this group is the thioester, whose electron-withdrawing capabilities are enhanced by the oxyanion hole provided by the enzyme. In the case of atrazine, the electron sink is provided by the N-1 nitrogen of the heteroaromatic ring, which can be protonated, nicely neutralizing the negative charge created by attack of the nucleophilic hydroxide on the ring. In the absence of a strategy such as these for accommodating the negative charge formed by attack of the nucleophile, hydrolytic dehalogenation is not expected. The structural requirements for reductive dehalogenation may be more varied; sufficient mechanistic information is not yet available for many reductive dehalogenases. However, mechanistic studies of TCHQ dehalogenase suggest that reductive dehalogenation using glutathione requires two hydroxyl groups in ortho or para positions. Studies of dehalogenation of chlorinated compounds such as PCBs by anaerobes, which appear to use transition metal cofactors, demonstrate that more highly chlorinated congeners are more readily dehalogenated (12). Such enzymes were not discussed here because there is little structural, mechanistic or genetic information, largely due to the difficulties in obtaining pure cultures of these organisms. However, this result is consistent with the expected mechanism for the reaction. This mechanism begins with initial electron transfer to the aromatic ring, followed by

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expulsion of chloride and further reaction of the resulting radical. Since compounds with more electron-withdrawing chlorine substituents are more easily reduced, they are more susceptible to reductive dehalogenation by transition-metal containing enzymes that operate by the mechanism. Aerobic microorganisms commonly use in the initial attack upon aromatic rings, and such reactions can result in dehalogenation. Flavin monooxygenases such as PCP hydroxylase require a hydroxyl substituent (or occasionally an amino substituent) in order to push electrons toward the C(4a)hydroperoxyflavin at the active site. This electron-donating group can be either ortho or para to the position of hydroxylation. The position of hydroxylation is determined by the orientation of the substrate in the active site. Similar interactions can facilitate hydroxylations carried out by monooxygenases. However, enzymes create a more powerful hydroxylating intermediate that is capable of hydroxylating inactivated positions in a wide variety of substrates (33), so dehalogenation reactions might not be limited to rings bearing hydroxyl or amino substituents. Finally, dioxygenase enzymes are also capable of attacking aromatic compounds that lack hydroxyl or amino substituents, as long as there are not too many electron-withdrawing chlorine substituents present on the ring. This collection of information on dehalogenases involved in xenobiotic degradation also allows some generalizations to be made about the potential for recruitment of an enzyme to serve a new function in the degradation of a novel compound. Three scenarios can be envisioned. First, it may be possible for a microorganism to recruit an enzyme that already catalyzes a certain type of transformation to carry out a similar reaction on a novel substrate. PCP hydroxylase and atrazine chlorohydrolase are likely examples of this strategy. Second, it may be possible to take advantage of a secondary reaction catalyzed by a catalytically promiscuous enzyme as a starting place for the evolution of a new enzymatic activity. This secondary activity need not be obviously related to the primary activity, although it will take advantage of some or all of the catalytic residues present at the active site of the enzyme. TCHQ dehalogenase is a particularly striking example of this strategy, as it appears to have evolved from an enzyme that catalyzed a very different reaction, the glutathione-dependent isomerization of a double bond. Finally, a third possibility is that there is no enzyme available at all to initiate the degradation of a xenobiotic compound. In such a case, the compound will not be biodegradable. In summary, a perusal ofthe catalytic strategies involved in aromatic dehalogenation provides some insights into the prospects for biodegradation of anthropogenic chlorinated aromatic compounds. In theory, nearly all chlorinated aromatic compounds should be susceptible to attack by one of the three classes of aromatic dehalogenases. The efficacy of biodegradation may be limited by the time it takes to evolve an enzyme with reasonable catalytic effectiveness, and this could take a long time, indeed. In addition, the biodegradation of such compounds may be inefficient because of the limited bioavailability of chlorinated aromatic compounds, which tend to be hydrophobic and therefore partition onto soils. Finally, the type of chemistry that is needed to dehalogenate a chlorinated aromatic compound is not always consistent with the environmental compartment in which the compound is found. For example, highly chlorinated compounds are most readily dechlorinated by transition metal enzymes found in anaerobes, but are difficult for aerobic microorganisms to attack. Conversely, compounds with only one or two chlorines are more readily attacked by aerobic

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microorganisms utilizing monooxygenases or dioxygenases. Thus, highly chlorinated aromatic compounds would be expected to persist in aerobic environments, and lesser chlorinated aromatic compounds to persist in anaerobic environments. REFERENCES atzB gene of Pseudomonas sp. strain ADP encodes the second enzyme of a novel atrazine DD (1989) Bacterial dehalogenation of degradation pathway. Appl. Environ. Microbiol. chlorobenzoates and coculture biodegradation of 63:916-923 4,4'-dichlorobiphenyl. Appl. Environ. Microbiol. 12. Brown JF, Bedard DL, Brennan MJ, Carnahan 55:887-892 JC, Feng H & Wagner RE (1987) Anandarajah K, Kiefer PM & Copley SD (2000) Polychlorinated biphenyl dechlorination in Recruitment of a double bond isomerase to serve aquatic sediments. Science 236:709-712 as a reductive dehalogenase during biodegradation of pentachlorophenol. 13. Ciurli S, Benini S, Rypniewski WR, Wilson KS, Miletti S & Mangani S (1999) Structural Biochemistry 39:5303-5311 properties of the nickel ions in urease: Novel Andersen SJ, Kuan S, Gawan B & Dabbs ER insights into the catalytic and inhibition (1997) Monooxygenase-like sequence of a mechanisms. Coordination Chem. Rev. 190Rhodococcus equi gene conferring increased 192:331-355 resistance to rifampin by inactivating this antibiotic. Antimicrobial Agents Chemother. 14. Clarkson J, Tonge PJ, Taylor KL, DunawayMariano D & Carey PR (1997) Raman study of 41:218-221 the polarizing forces promoting catalysis in Anglister L, Stiles JR & Salpeter MM (1994) 4-chlorobenzoate dehalogenase. Biochemistry Acetylcholinesterase density and turnover number at frog neuromuscular junctions, with 36:10192-10199 modeling of their role in synaptic function. 15. Cline RE, Hill RH, Phillips DL, & Needham LL Neuron 12:783-794 (1989) Pentachlorophenol measurements in Armstrong RN (997) Structure, catalytic body fluids of people in log homes and mechanism, and evolution of the glutathione workplaces. Arch. Env. Contam. Toxicol. transferases. Chem. Res. Toxicol. 10:2-18 18:475-481 Beaty NB & Ballou DP (1981) The oxidative 16. Copley SD (2000) Evolution of a metabolic half-reaction of liver microsomal FADpathway for degradation of a toxic xenobiotic: containing monooxygenase. J. Biol. Chem. The patchwork approach. Trends Biochem. Sci. 25:261-265 256:4619-4625 Benning MM, Haller T, Gerlt JA & Holden HM 17. Copley SD & Crooks GP (1992) Enzymic (2000) New reactions in the crotonase dehalogenation of 4-chlorobenzoyl coenzyme A superfamily: Structure of methylmalonyl CoA in Acinetobacter sp. strain 4-CB1. Appl. decarboxylase from Escherichia coli. Environ. Microbiol. 58:1385-1387 18. Crooks GP, Xu L, Barkley RM & Copley SD Biochemistry 39:4630-4639. Benning MM, Hong S-B, Raushel FM & Holden (1995) Exploration of possible mechanisms for HM (2000) The binding of substrate analogs to 4-chlorobenzoyl CoA dehalogenase: Evidence phosphotriesterase. J. Biol. Chem. 275:30556for an aryl-enzyme intermediate. J. Am. Chem. 30560 Soc. 117:10791-10798 Benning MM, Taylor KL, Liu R-Q, Yang G, 19. Dairi T, Nakano T, Aisaka K, Katsumata R & Xiang H, Wesenberg G, Dunaway-Mariano D & Hasegawa M (1995) Cloning and nucleotide Holden HM (1996) Structure of 4-chlorobenzoyl sequence of the gene responsible for chlorination coenzyme A dehalogenase determined to 1.8 Å of tetracycline. Biosci. Biotechnol. Biochem. resolution: An enzyme catalyst generated via 59:1099-1106 adaptive mutation. Biochemistry 35:8103-8109 2 0. de Souza ML, Newcombe D, Alvey S, Crowley Board PG, Baker RT, Chelvanayagam G, & DE, Hay A, Sadowsky MJ & Wackett LP (1998) Jermiin LS (1997) Zeta, a novel class of Molecular basis of a bacterial consortium: glutathione transferases in a range of species Interspecies catabolism of atrazine. Appl. from plants to humans. Biochem. J. 328:929Environ. Microbiol. 64:178-184 935 21. de Souza ML, Sadowsky MJ & Wackett LP Boundy-Mills K, de Souza ML, Mandelbaum (1996) Atrazine chlorohydrolase from RM, Wackett LP & Sadowsky MJ (1997) The Pseudomonas sp. strain ADP: Gene sequence,

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258 studies. J. Biol. Chem. 268:4134-4144 45. Mandelbaum RT, Wackett LP & Allan DL (1993) Mineralization of the s-triazine ring of atrazine by soil bacteria. Appl. Environ. Microbiol. 59:1695-1701 46. Mao Y, Varoglu M & Sherman DH (1999) Molecular characterization and analysis of the biosynthetic gene cluster for the antitumor antibiotic mitomycin C from Streptomyces lavendulae NRRL 2564. Chem. and Biol. 6: 251-263 47. Martinez B, Tomkins J, Wackett, LP, Wing, R & Sadowsky, MJ (2001) Complete nucleotide sequence and organization of the atrazine catabolic plasmid pADP-1 from Pseudomonas sp. Strain ADP. J. Bacteriol. 183:5684-5697 48. McCarthy DL (1998) Studies of tetrachlorohydroquinone dehalogenase. Ph.D. Thesis, University of Colorado at Boulder, 1998 49. McCarthy DL, Claude A & Copley SD (1997) In vivo levels of chlorinated hydroquinones in a pentachlorophenol-degrading bacterium. Appl. Environ. Microbiol. 63:1883-1888 50. McCarthy DL, Louie DF & Copley SD (1997) Identification of a covalent intermediate between glutathione and cysteine13 during catalysis by tetrachlorohydroquinone dehalogenase. J. Am. Chem. Soc. 119:11337-11338 51. McCarthy DL, Navarrete S, Willett WS, Babbitt PC & Copley SD (1996) Exploration of the relationship between tetrachlorohydroquinone dehalogenase and the glutathione S-transferase superfamily. Biochemistry 35:14634-14642 52. Modis Y, Filppula SA, Novikov DK, Norledge B, Hiltunen JK & Wierenga RK (1998) The crystal structure of dienoyl-CoA isomerase at 1.5 A resolution reveals the importance of aspartate and glutamate sidechains for catalysis. Structure 6: 957-970 53. Ni S, Fredrickson JK & Xun L (1995) Purification and characterization of a novel 3-chlorobenzoate reductive dehalogenase from the cytoplasmic membrane of Desulfomonile tiedjei DCB-1. J. Bacteriol. 177: 5135-5139 54. Orser CS, Dutton J, Lange C, Jablonski P, Xun L & Hargis M (1993) Characterization of a Flavobacterium glutathione-S-transferase gene involved in reductive dehalogenation. J. Bacteriol. 175: 2640-2644 55. Orser CS & Lange CC (1994) Molecular analysis of pentachlorophenol degradation. Biodegradation 5: 277-288 56. Orser C, Lange CC, Xun L, Zahrt TC & Schneider BJ. (1993) Cloning, sequence analysis, and expression of the Flavobacterium pentachlorophenol-4-monooxygenase gene in Escherichia coli. J. Bacteriol. 175: 411-416 57. Palfey BA, Ballou DP & Massey V (1997)

COPLEY Flavin conformational changes in the catalytic cycle of p-hydroxybenzoate hydroxylase substituted with 6-azido- and 6-aminoflavin adenine dinucleotide. Biochemistry 36: 1571315723 58. Pearson MA, Park I.-S, Schaller RA, Michel LO, Karplus PA & Hausinger RP (2000) Kinetic and structural characterization of urease active site variants. Biochemistry 39: 8575-8584 59. Peelen S, Rietjens IMCM, Boersma MG, Vervoort J (1995) Conversion of phenol derivatives to hydroxylated products by phenol hydoxylase from Trichosporon cutaneum: A comparison of regioselectivity and rate of conversion with calculated molecular orbital substrate characteristics. Eur. J. Biochem. 227: 284-291 60. Powlowski J, Ballou DP & Massey V (1990) Studies of the oxidative half-reaction of anthranilate hydroxylase (deaminating) with native and modified substrates. J. Biol. Chem. 265:4969-4975 61. Raushel FM & Holden HM (1998) Phosphotriesterase: An enzyme in search of its natural substrate. Adv. Enzymol. Rel. Areas Mol. Biol. 74:51-93 62. Romanov V & Hausinger RP (1996) NADPHdependent reductive ortho dehalogenation of 2,4-dichlorobenzoic acid in Corynebacterium sepedonicum KZ-4 and coryneform bacterium strain NTB-1 via 2,4-dichlorobenzoyl coenzyme A. J. Bacteriol. 178: 2656-2661 63. Sadowsky MJ, Tong Z, de Souza ML & Wackett LP (1998) AtzC is a new member of the amidohydrolase protein superfamily and is homologous to other atrazine-metabolizing enzymes. J. Bacteriol. 180: 152-158 64. Sayle RA, Milner-White EJ (1995) RasMol: Biomolecular graphics for all. Trends Biochem. Sci. 20(Sept): 374-376 65. Schmitz A, Gartemann K-H, Fiedler J, Grund E & Eichenlaub R (1992) Cloning and sequence analysis of genes for dehalogenation of 4-chlorobenzoate from Arthrobacter sp. strain SU. Appl. Environ. Microbiol. 58:4068-4071 66. Scholten JD, Chang K-H, Babbitt PC, Charest H, Sylvestre M & Dunaway-Mariano D (1991) Novel enzymic hydrolytic dehalogenation of a chlorinated aromatic. Science 253: 182-185 67. Schopfer LM & Massey V (1980) Kinetic and mechanistic studies on the oxidation of the melilotate hydroxylase-2-OH-cinnamate complex by molecular oxygen. J. Biol. Chem. 255:5355-5363 68. Schreuder HA, Mattevi A, Obmolova G, Kalk KH, Hol WGJ, van der Bolt FJT & van Berkel WJH (1994) Crystal structures of wild-type phydroxybenzoate hydroxylase complexed with 4-

AROMATIC DEHALOGENASES aminobenzoate, 2,4-dihydroxybenzoate, and 2-hydroxy-4-aminobenzoate and of the Tyr222Ala mutant complexed with 2-hydroxy4-aminobenzoate: Evidence for a proton channel and a new binding mode of the flavin ring. Biochemistry 33: 10161-10170 69. Schreuder HA, Prick PAJ, Wierenga RK, Vriend G, Wilson KS, Hol WGJ & Drenth J (1989) Crystal structure of the p-hydroxybenzoate hydroxylase-substrate complex refined at 1.9 Å resolution. Analysis of the enzyme-substrate and enzyme-product complexes. J. Mol. Biol. 208: 679-696 70. Seltzer S (1973) Purification and properties of maleylacetone cis-trans isomerase from Vibrio 01. J. Biol. Chem. 248: 215-222 71. Seltzer S & Lin M (1979) Maleylacetone cistrans-isomerase. Mechanism of the interaction of coenzyme glutathione and substrate maleylacetone in the presence and absence of enzyme. J. Am. Chem. Soc. 101: 3091-3097 72. Siuda JF & DeBernardis JF (1973) Naturally occurring halogenated organic compounds. Lloydia 36: 107-143 73. Suzuki K, Gomi T, Kaidoh T& Itagaki E (1991) Hydroxylation of o-halogenophenol and onitrophenol by salicylate hydroxylase. J. Biochem. 109:348-353 74. Uotila JS, Kitunen VH, Saastamoinen T, Coote T, Häggblom MM & Salkinoja-Salonen MS (1992) Characterization of aromatic dehalogenases of Mycobacterium fortuitum CG-2. J. Bacteriol. 174: 5669-5675 75. Uotila JS, Salkinoja-Salonen MS & Apajalahti JHA (1991) D e c h l o r i n a t i o n of pentachlorophenol by membrane bound enzymes of Rhodococcus chlorophenolicus PCP-1. Biodegradation. 2: 25-31 76. van Berkel WJH & Müller F (1990) Flavindependent monooxygenases with special reference to p-hydroxybenzoate hydroxylase. In: Chemistry and Biochemistry of Flavoenzymes. Müller F (Ed) CRC Press. Boca Raton, 1990; Vol. II.

259 77. Westrich L, Domann S, Faust B, Bedford D, Hopwood DA & Bechtold A (1999) Cloning and characterization of a gene cluster from Streptomyces cyanogenus S136 probably involved in landomycin biosynthesis. FEMS Microbiol. Lett. 170: 381-387 78. Wilson DK, Rudolph FB & Quiocho FA (1991) Atomic structure of adenosine deaminase complexed with a transition-state analog: Understanding catalysis and immunodeficiency mutations. Science 252:1278-1284 79. Xiang H, Luo L, Taylor KL & DunawayMariano D (1999) Interchange of catalytic activity within the 2-enoyl-coenzyme A hydratase/isomerase superfamily based on a common active site template. Biochemistry 38:7638-7652 80. Xun L (1999) Studies of 2,6-Dichlorohydroquinone Dioxygenase and 4-Chlorobenzoyl CoA Dehalogenase. Ph.D. Thesis, University of Colorado at Boulder. 81. Xun L & Orser CS (1991) Purification of a Flavobacterium pentachlorophenol-induced periplasmic protein (PcpA) and nucleotide sequence of the corresponding gene (pcpA). J. Bacteriol. 173: 2920-2926 82. Xun L, Topp E & Orser CS (1992) Purification and characterization of a tetrachloro-phydroquinone reductive dehalogenase from a Flavobacterium sp. J. Bacteriol. 174:8003-8007 83. Xun L, Topp E & Orser C (1992) Diverse substrate range of a Flavobacterium pentachlorophenol hydroxylase and reaction stoichiometries. J. Bacteriol. 174: 2898-2902 84. Xun L, Topp E & Orser CS (1992) Glutathione is the reducing agent for the reductive dehalogenation of tetrachloro-p-hydroquinone by extracts from a Flavobacterium sp. Biochem. Biophys. Res. Commun. 182: 361-366 85. Yang G, Liang P-H & Dunaway-Mariano D (1994) Evidence for nucleophilic catalysis in the aromatic substitution reaction catalyzed by (4-chlorobenzoyl)coenzyme A dehalogenase. Biochemistry 33: 8527-8531

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Chapter 9 ABIOTIC DEHALOGENATION BY METALS LISA A. TOTTEN1 AND NADA M. ASSAF-ANID2 1

2

Department of Environmental Sciences, Rutgers University, New Brunswick NJ, USA Chemical Engineering Department, Manhattan College, Riverdale, NY, USA

1. BACKGROUND Zero-valent metals have received much recent attention in the scientific community because they can serve as a fixed source of electrons for the reductive dehalogenation of organic contaminants, such as carbon tetrachloride (CT) and perchloroethene (PCE) in groundwater. Zero-valent metal remediation technology has already been tested at several sites in the United States (50, 72, 89, 94, 102, 140). It typically involves the installation of a permeable barrier containing granular iron (Fe°), which transects the natural flow path of the local groundwater. The encounter between the iron granules and the groundwater contaminants results in iron oxidation (corrosion) and diminished downgradient contaminant concentrations through reductive dechlorination. This type of passive remediation system provides several significant advantages over conventional pump-and-treat methods. It virtually eliminates daily operation and maintenance costs, and because all remediation activities take place below ground, it is aesthetically acceptable and allows the property at the remediation site to be used for other purposes, thereby significantly enhancing its value. Although the reactions of alkyl halides with zero-valent metals have long been recognized in the field of corrosion chemistry (4, 5, 124), only recently have engineers and scientists begun to realize the potential of metals for degrading these compounds (126, 127). Interest in this field began when Reynolds et al. (106) conducted a series of experiments to determine whether different materials used to monitor groundwater may introduce sampling bias. Stainless steel, aluminum, and galvanized steel all reacted with at least some of the five test compounds: bromoform; 1,1,1-trichloroethane (TCA); 1,1,2,2-tetrachloroethane (TeCA); hexachloroethane (HCA); and PCE. Since this initial publication on the subject, dozens of papers dealing with remediation of alkyl halide contaminants by zero-valent metals have appeared in refereed journals. Most of this work has centered on reductions promoted by granular iron metal because the iron species which dissolve into groundwater during corrosion of the Fe(0) are non-toxic and therefore of no concern. Some work has investigated Zn(0), which is thermodynamically a stronger reductant than Fe(0), and a few other metals (Cu, Al and others), including bimetallic systems that usually consist of Fe and Pd, Pt, Cu, Ag, or Ni. Dehalogenation: Microbial Processes and Environmental Applications, pages 261-287 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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2. SCOPE A substantial amount of research has been performed to date (see Table 9.1 for a summary of research findings from 1990 to present). All of the tested chlorinated ethenes react with various metals, including iron, zinc, tin, palladium (with amendment), as well as bimetallic reductants (2, 8, 23, 27, 83, 97, 116, 117, 125). Aliphatic halides containing 3 or more chlorines are typically reactive with iron and zinc (13, 43, 51, 55, 62, 80, 86, 116, 117). Granular zinc is generally a stronger reductant (27), and is thus capable of dehalogenating dichloromethane (DCM) and the dichloroethanes (7, 17). It is not clear whether zinc reacts with chloroethane or chloromethane, but it has been shown to react with hexachlorobenzene (9). Schreier and Reinhard (116) demonstrated that granular manganese dehalogenates PCE, 1,1,1 -TCA, and 1,1-dichloroethene (DCE). The ability of granular iron to dechlorinate various substrates cannot be predicted a priori from the reduction potential of the substrate. The Fe(II)/Fe(0) couple has a redox potential of -0.447 V, suggesting that it is capable of reducing even monohalogenated methanes and ethanes, which have positive two-electron reduction potentials (130). However, aliphatic halides containing two or fewer chlorine atoms are often unreactive with granular iron, although they may react readily with zinc or bimetallic reductants. It is somewhat surprising that DCM is largely unreactive with granular iron, while the chlorinated ethenes react rapidly with iron, given that the oneelectron reduction potential for dichloromethane is -0.428 V (standard hydrogen electrode), while the most favorable among the chloroethenes is -0.598 V for tetrachloroethene (130). Granular iron also dehalogenates aromatic chlorides under some conditions, including polychlorinated biphenyls (PCBs) and chlorinated benzenes. Wang and Zhang (133) have demonstrated that nanoscale (1-100 nm) iron particles slowly dehalogenate PCBs, resulting in approximately 25% reduction in overall PCB concentration in 17 hours. Others (28, 138, 139) have demonstrated reductive dehalogenation of PCBs in water by granular iron at high temperature and pressure. Addition of palladium to the iron results in a dramatic increase in reactivity; PCBs may be completely dehalogenated to biphenyl at ambient temperature by Pd/Fe, in a timeframe of minutes to hours (56, 133). Similarly, subcolloidal iron/silver particles dechlorinate benzenes containing 3 to 6 chlorines, while unamended iron is largely unreactive (137). Some chlorophenols are dehalogenated by iron (71) or palladinized iron (54). Vicinal dibromides are dehalogenated by iron, zinc, copper, and aluminum (105, 119, 129). Bromoform is reduced by iron, zinc, tin, and aluminum (135). Iron and zinc dehalogenate monobromo compounds, including vinyl bromide, bromoethane, and others (105, 128). Given that iron and zinc dehalogenate monobromides, they are likely capable of dehalogenating other brominated species with more favorable redox potentials, such as fluorobromomethanes and -ethanes (halons). Other chlorinated organics are dehalogenated by granular iron, including haloacetic acids (64), and herbicides (38, 40, 47, 48, 91, 110). Granular iron is also effective at reducing nitroaromatic compounds (1, 37, 66, 120), azo dyes (24, 36, 93), and other Nsubstituted organics (47, 49, 59, 96). Transition metals, including uranium and chromium, are reduced by granular iron to insoluble (and therefore immobile) forms (44,

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57, 100, 101). Granular iron has, for example, been used to remediate groundwater contaminated with Cr(VI) by reducing it to insoluble Cr(III) (12, 103, 104).

3. DEHALOGENATION PRODUCTS For saturated and unsaturated aliphatic polyhalides, reductive dehalogenation may proceed via two possible pathways (Table 9.2): hydrogenolysis (replacement of halogen by hydrogen), or elimination in which two halogens are eliminated from the substrate resulting in an increase in bonding order (when halogens are vicinal), or a carbene (when halogens are geminal). Most metals also promote other transformation reactions, including hydrogenation of double and triple bonds. All the above reactions can lead to complicated pathways (e.g., Figure 9.1). The thermodynamics of reductive elimination tend to be more favorable than those of hydrogenolysis, due largely to the production of two heavily solvated chloride (or bromide) ions (130). Thus, it is not surprising that many vicinal dihalides react primarily via (see Table 9.2.). Because carbenes are stabilized by halogen substituents (63), tetrahalomethanes are more likely to undergo reduction via yielding a dihalocarbene than trihalomethanes, which react via to form a monohalocarbene. Thus, it would appear that chloroform (CF) reacts solely via hydrogenolysis to DCM. Although the DCM product is unreactive with granular iron under most conditions (86), it may react with zinc to give chloromethane and methane (11, 16, 17). Although carbene intermediates have not been directly detected as products of CT reduction by granular metals in water, their role as intermediates in the reaction has been invoked to explain the complete dechlorination of CT (as indicated by production of chloride or methane) often observed with zero-valent metals such as iron, zinc, and tin (13, 16). The proposed pathways for reduction of CT by granular metals is shown in Figure 9.2. Reaction of CT

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with granular iron, zinc, magnesium, and tin may therefore occur by both hydrogenolysis and pathways to yield chloroform and CO, respectively (16, 86). Chlorinated ethenes have been shown to react via elimination with granular iron and zinc (6, 8, 107). The percent of the reaction proceeding via elimination in the presence of zinc is between 15% (for PCE) and 95% (for trans-DCE), and is related to the difference in redox potential between the hydrogenolysis and reactions (6). When granular iron serves as the reductant, the pathway dominates, accounting for 85-99% of products for all of the vicinal polychloroethenes (8). Liu et al. (82) noted that chlorinated ethenes react via similar pathways during electrolysis on a porous nickel cathode. This elimination pathway is advantageous in that each elimination step allows the loss of two halogens leading to full dehalogenation while circumventing the production of generally toxic intermediates. The final products of reduction of chlorinated ethenes by granular iron, granular zinc, and the nickel cathode are completely dehalogenated hydrocarbons such as acetylene, ethene and ethane, and also some compounds formed in the presence of iron (2, 8, 23, 97, 116). The hydrocarbons may result in part from the reduction of or carbide carbon residual in the iron (34). Deng et al. (34) demonstrated that reaction of

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trichloroethene (TCE) with granular iron produces hydrocarbons. It is striking that although factors such as solution pH and surface condition of the iron dramatically effect the kinetics of transformation (as described below), the distribution of reduced products remains remarkably constant over a wide pH range. Products ofthe reaction also appear to be the same despite a variety of species in solution (both organic and inorganic), in both column and batch reactors, and in the presence of both acidwashed and unwashed iron. While reactions of chlorinated ethenes have been investigated in much detail, relatively little work has been done on the reactions of chlorinated ethanes with zerovalent metals. A few studies have been conducted with granular iron (43, 51, 112, 116). Arnold and Roberts (7) noted that in the presence of granular zinc, most vicinal polychloroethanes react exclusively via reductive including HCA, both TeCAs, 1,1,2-TCA, and 1,2-dichloroethane (1,2-DCA). Pentachloroethane reacted mostly via reductive (93%), but also underwent some dehydrohalogenation (hydrolysis) to TCE. Reductive elimination from chlorinated ethanes yields chlorinated ethenes which then react via the pathways described above to result in dehalogenated hydrocarbon products. Given that, for chlorinated ethenes is more extensive with granular iron than zinc, it is likely that chlorinated ethanes also react almost exclusively via elimination pathways with granular iron. The products of these reactions have not been investigated in any detail. Liu et al. (82) investigated electrolytic reduction of chloroethanes with a porous Ni cathode and noted that vicinal polychloroethanes reacted primarily via reductive No lesser-chlorinated ethanes were detected as products. For polychloroalkanes which do not have a pair of vicinal halogens, such as CT,

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1,1,1-TCA and 1,1-DCA, reductive is not possible. Fennelly and Roberts (43) investigated reduction of 1,1,1-TCA and 1,1-DCA by granular zinc. In their study, 1,1,1-TCA yielded approximately 75% ethane and 25% 1,1-DCA. The 1,1-DCA product reacted slowly with granular zinc to form ethane. In contrast, 1,1,1-TCA reacted with granular iron to form primarily 1,1-DCA (~67%), and lesser amounts of ethane, cis-2butene, ethene, and 2-butyne. The 1,1-DCA product reacted slowly with iron to form primarily ethene and ethane. Plating the iron surface with nickel or copper resulted in a similar product distribution as granular iron from reduction of 1,1-DCA and 1,1,1TCA, but with decreased 1,1-DCA yields. In the presence of copper-plated granular iron, 1,1,1,-TCA reacted to form traces of 1,1-DCE, most likely by dehydrohalogenation. In all cases, 1,1-DCA reacted too slowly with the granular metal to be an intermediate for ethane or ethene formation, and chloroethane was not detected as a reaction product. Based on the distribution of products, Fennelly and Roberts (43) hypothesized that 1,1,1-TCA and 1,1-DCA undergo reductive toform either a free carbene or organometallic carbenoid intermediate in the presence of each of these metals/bimetals. This mechanism satisfactorily explains the formation of hydrocarbon products without the formation of lesser chlorinated ethane intermediates. The products observed by Gregory et al. (55) for the reduction of 1,1,1-TCA in a column containing steel wool are consistent with such a mechanism, as are the products of electrolysis of 1,1,1-TCA and 1,1-DCA on a porous nickel cathode reported by Liu et al. (82). In contrast, in the presence of granular aluminum 1,1,1,-TCA was reported to undergo dehydrohalogenation to yield significant amounts of 1,1-DCE (5). A great deal of research has been conducted on the reactions of CT and other chlorinated methanes with zero-valent metals. Criddle and McCarty (31) studied electrolysis of CT on a silver electrode and hypothesized that reduction occurred via two competing pathways: hydrogenolysis to CF, and to form the dichlorocarbene. The latter product resulted in the production of CO and formate. Liu et al. (82) reported that electrolytic reduction of CT produces similar products (CF, DCM, and methane, but no chloromethane) regardless of the cathode material (Ag, Al, Au, Cu, Fe, Ni, Pd, or Zn). With the nickel cathode, ethane, ethene, and propane were detected as products. Mass balances in this study were typically 90% or better. In studies with granular iron, CT has been shown to be typically reduced (dehalogenated) to CF, although the reported yields ofCF vary widely and mass balances are usually incomplete (13, 55, 62, 80, 86). Matheson and Tratnyek (86) reported that CT was dehalogenated to CF (70%), and CF was further dehalogenated to DCM (50%), which was unreactive. Lien and Zhang (80) reported reduction of CT by granular iron via CF to DCM (71%), methane (25%), and traces of chloromethane. DCM appears to be reduced too slowly to account for the production ofmethane in this system. Use ofnanoscale iron increased the yield of methane and decreased production of DCM, but chloromethane was not detected as an intermediate (80). Balko and Tratnyek (13) hypothesized that conduction band electrons may be the reactive species in the zero-valent iron system. They reacted CT with iron under photoirradiation to increase the production of conduction band electrons in the oxide which coats the metal. The amount of CF produced decreased from ~30% to ~40% and the overall rate of reaction increased slightly. The researchers hypothesized that the remaining mass consisted of CO and other products of dichlorocarbene hydrolysis.

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Application of ultrasound (65), besides increasing the reactivity of granular iron, does change the reaction pathways for CT substantially, resulting in the production of coupling products, such as HCA and pentachloroethane (PCA). These products arise from free-radical chemistry initiated by sonolysis (65). Lipczynska-Kochany et al. (81) also reported that CF is the primary product of CT reduction by unamended granular iron, but in the presence of FeS, or In this study, CF was not detected as a product. Metallic Pd/Fe also produces traces of coupling products (PCE and TCE) from reduction of CT, as well as promoting dehalogenation of DCM to methane, but not chloromethane (80). Relatively little research has been conducted to investigate the reductive dehalogenation of brominated compounds by zero-valent metals. Warren et al. (135) observed reduction of bromoform to dibromomethane with iron, zinc, tin, and aluminum. In the presence of granular zinc or iron, methane was also a product of bromoform reduction. Rajagopal and Burris (105) investigated the reactions of a few brominated compounds with granular iron. In their study, 1,2-dibromoethane reacted via to form ethene exclusively (i.e., vinyl bromide and bromoethane were not intermediates in ethene production). Vinyl bromide reacts to form ethene, while bromoethane undergoes both hydrogenolysis to ethane and dehydrohalogenation to ethene. Other researchers (128, 129) have utilized alkyl bromides to investigate the mechanism of dehalogenation promoted by granular iron, zinc, aluminum, and copper. With all of these metals, threo- and erythro-2,3-dibromopentane underwent stereospecific reductive to >95% production of cis- and trans-2-pentene, respectively. In addition, D,L-stilbene dibromide was reduced to a mixture of cis- and trans-stilbenes. No hydrogenolysis products were observed in these studies. Monobromides (6-bromo-1-hexene and bromomethylcyclopropane) have also been investigated for use as “radical clocks”, i.e., species which yield characteristic rearrangement products if reactions proceed via single electron transfer pathways, and which can also in some cases allow an estimate of the pseudo first-order rate constant associated with reduction of the intermediate radicals (128). In the presence of zinc, both radical clocks were reduced to the expected (hydrogenolysis) products: 1-hexene and methylcyclopentane (from 6-bromo-1-hexene), and methylcyclopropane and 1-butene (from bromomethylcyclopropane), although hydrolysis was also significant. In contrast to this simple product distribution, reduction of the radical clocks by granular iron was more complicated. Reaction of both radical clocks with granular iron gave rise to several by-products, including 1,5-hexadiene, coupling products such as dodecane, and isomerized olefins such as 2-hexene from 1-hexene, which are characteristic of the decomposition of organoiron species. The formation of such organometallic intermediates may explain the formation of rearrangement products, suggesting that a radical intermediate are not necessarily produced in this system. Alternatively, the low yields of rearranged products suggest that any radical intermediate formed is very shortlived.

4. KINETICS Reactions of alkyl halides with granular zinc appear to obey a simple pseudo firstorder kinetic model in the absence of mass transfer limitations (6, 135), making the

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measurement of intrinsic reaction rate constants a relatively easy process. Arnold et al. (6, 7) have demonstrated that the reactivity of vinyl and ethyl chlorides with granular zinc increases with their one- and two- electron reduction potentials. This relationship suggests that electron transfer, i.e., reaction at the metal surface, is the rate-limiting step of the reaction. The exceptions to this relationship are the heavily chlorinated ethanes, whose reactions with zinc appear to be mass-transfer limited (7). In contrast, Liu et al. (82) observed that, while rate constants for the reaction of chlorinated ethanes on a porous nickel cathode displayed good correlations with both values and C-C1 bond dissociation energies (a surrogate for the correlation did not extend to chlorinated ethenes. These results suggest that the olefins react with the cathode via a different mechanism. In general, the reaction of chlorinated methanes, ethanes, and ethenes at iron surfaces appears to be rate-limiting, i.e., the mass transfer or desorption of products is not one of the rate-limiting processes in the iron system. This conclusion is based in part on the observed activation energies for reduction of chlorinated compounds by granular iron. Diffusion-controlled reactions are expected to have low activation energies typically less than 21 kJ/mol, while chemical processes generally display higher values (76). Sivavec and Horney (121) reported values in the range of 15 to 18 kJ/mol for the reductive dehalogenation of chlorinated ethenes by commercial iron powders, suggesting a mass-transfer limited process. More recently, however, Scherer et al. (111) reported values of 55.9 ± 12.0 and 40.5 ± 4.1 kJ/mol for reactions of CT and hexachloroethane with granular iron, respectively. Similarly, Su and Puls (125) reported values ranging from 25.8 to 81.8 kJ/mol for reduction of TCE on many types of granular iron. Rajagopal and Burris (105) measured to be 50 kJ/mol for reaction of 1,2-dibromoethane with cast iron, and Deng et al (35) found the for reaction of vinyl chloride (VC) with granular iron to be 41.6 ± 2.0 kJ/mol. The dependence of reaction rates on the surface condition of the iron is further evidence that the rate-limiting step for reductive dehalogenation involves reaction at the iron surface (111). Another line of evidence which suggests that reaction at the iron surface is ratelimiting is the reported intra- (parent-parent) and inter- (parent-daughter) species competition for reactive sites (8, 69, 136). Unlike the reactions of chlorinated ethenes with zinc (6), reaction of many chlorinated solvents with iron metal cannot be described by a simple pseudo first-order kinetic model. Johnson et al. (68) noted that most of the variation in the reported rate constants for reactions of alkyl halides with granular iron can be explained by normalizing the experimental data to the iron surface area. Others (8, 69, 136) have, however, observed that rate constants for the reaction of many halides, even when normalized to surface area, vary significantly depending on the initial ratio of substrate to metal surface area. Based on this information, the reaction rate data must be modeled as a process involving reaction of the chlorinated compound at a limited number of reactive sites on the metal surface. Arnold and Roberts (8) likewise observed that during reaction with granular iron, most chlorinated ethenes compete both with themselves and with any daughter products for the reactive sites on the metal. Because of these complicated kinetics, the intrinsic reactivity of an alkyl halide with granular iron can only be determined by measuring the bulk disappearance rate of the compound and its daughter products at a variety of initial alkyl halide concentrations. The data must then be modeled to account for intra- and inter- species competition for reactive sites.

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The exception to this model is VC, which appears either to react via a mechanism that is not site-limited or to have a very low affinity for sites, such that intraspecies competition is not observed at the concentration of VC typically employed (8, 35). Scherer et al. (112) developed a linear free-energy relationship (LFER) for reactions of chlorinated methanes, ethanes, and ethenes with zero-valent iron by using published reaction rate constants and values. This analysis revealed a direct relationship between faster reduction kinetics and increasing values (Figure 9.3). The researchers relied on rate constants derived from bulk disappearance kinetics and the values calculated by Curtis (32) and Roberts et al. (107) to develop their correlations. The research of others (8, 69) has demonstrated that rate constants obtained in this way may not accurately portray the intrinsic reactivity of alkyl halides with granular iron due to intra- and inter-species competition for reactive sites. Also, the sorption of alkyl halides to unreactive sites consisting of exposed graphite inclusions on the iron surface can cause rapid initial declines in aqueous concentrations of the reactants which might be mistaken for reaction (18, 19, 23). Finally, it should be noted that Totten and Roberts (130) have demonstrated that the values calculated by Curtis may be in error by as much as 0.4 V. Using the values calculated by Totten and Roberts (130) and the

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kinetic data compiled by Scherer et al. (112) indicates that reduction kinetics become faster as values increase. Using rate constants derived from a model of the reactions of chlorinated ethenes that accounts for intra- and inter- species competition for a limited number of reactive sites on the iron metal surface, Arnold and Roberts (8) have derived a log k vs. correlation (Figure 9.4) that resulted in a negative slope, indicating that the second-order rate constants for the reactions of these compounds with granular iron decreased with increasing The greater reactivity of ethenes containing fewer chlorine substituents with iron has also been observed by others (41, 42, 136). The kinetics of reductive dehalogenation for aliphatic chlorides such as CT and chlorinated ethanes by granular iron have not been analyzed via a model that accounts for inter- and intra-species competition, so it is not known whether they will also display decreasing reaction rates with increasing reduction potentials. The apparent discrepancy between the conclusions of these two groups of researchers is puzzling, but it may arise in part from the different data sets considered. Arnold and Roberts (8) included only chlorinated ethenes in their correlation, while Scherer et al. (112) included chlorinated methanes, ethanes, and ethenes in their analysis. Studies have shown that reaction rates tend to decrease with increasing pH (50, 52, 86, 92, 116, 135). Acid washing or polishing of the iron surface can increase reaction rates, at least initially (86, 109, 111, 125). Sulfur species, chloride, and surfactants can

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increase the reactivity of granular iron (69, 54, 61, 77, 81), while oxygen and anionic organic ligands inhibit the reaction (62, 69).

5. MECHANISTIC ISSUES The first attempt to understand the mechanisms of metal-catalyzed abiotic dehalogenation reactions was reported by Matheson and Tratnyek (86), who investigated the reaction of CT with granular iron. They focused on identifying the exact species responsible for the reduction reaction, and on quantifying the effects of changing pH and iron surface characteristics. These researchers hypothesized that, although iron metal is the initial source of the electrons, three species may be responsible for the reductive dehalogenation observed: iron metal itself, [Fe(0)]; Fe(II) generated from the corrosion of the iron metal and sorbed via an inner- or outer-sphere complex to the iron oxide coating the metal; or nascent hydrogen, also produced via the reduction of protons (Figure 9.5). Assigning possible reductants into one of these three classes is perhaps an oversimplification of the complexities inherent in the granular iron system. Dihydrogen would require a suitable catalyst, (assumably Fe(0) or an Fe(II) species), to be an effective reductant under these conditions, Scherer et al. (113) hypothesized that electron transfer may occur via three pathways in the presence of granular iron (Figure 9.6). First, electrons may be transferred from bare metal Fe(0) to the electron acceptor Fe(II) oxide in corrosion pits on the metal surface acting as the conduction band that transfers electrons to the substrate. Changes in the granular iron surface induced by reactions with alkyl halides have been investigated via scanning electron microscopy, surface profilometry and atomic force microscopy (53). These studies showed that pitting corrosion does occur, raising the possibility that the galvanic cells induced via pitting may be responsible for much of the dehalogenating activity of the iron. The results did not, however, rule out the possibility that hydrogen gas generated in the pits acts as a reductant, with surface defects acting as catalysts (53). Further research has provided clues about the possible role of hydrogen species in reductive dehalogenation promoted by granular iron. Scherer et al. (111) and Li and Farrell (78) observed that negligible amounts of hydrogen are produced during reduction of CT at an oxide-free iron electrode, and therefore concluded that hydrogen species are unimportant in the reduction of CT. Liu et al. (82) similarly noted that electrode efficiency for the reductive dehalogenation of CT is inversely related to the current density for hydrogen evolution for a range of metal cathodes (i.e., cathodes which produce large amounts of hydrogen are less efficient at reducing CT). The strong correlation between dissolved Fe(II) and CF produced from CT dehalogenation (67, 111) has also been cited as evidence that Fe(0) or Fe(II), but not hydrogen, is the dominant reductant in the granular iron system (115). These arguments all involve CT. Li and Farrell (78) have used several lines of evidence, including the fact that the applied potential at an iron electrode has little effect on the rate of TCE reduction, to conclude that TCE is reduced indirectly by hydrogen species at iron electrodes. Reduction of TCE by an iron electrode likely proceeds overwhelmingly via to chloroacetylene (8), and subsequent reaction steps must then involve hydrogenation of double bonds, a reaction which is presumably mediated by nascent hydrogen. It remains possible that nascent hydrogen also plays a significant role in

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hydrogenolysis reactions under some conditions. Totten (128) has also identified a role for Fe-H species in determining the product distribution arising from reduction of 6bromo-1-hexene. In this system, the Fe-H species are thought to catalyze the isomerization of olefins and elimination reactions which convert 6-bromo-1 hexene to 1,5-hexadiene. The respective roles of Fe(0) and Fe(II) species in the granular iron system are very difficult to distinguish. Fe(II)-containing minerals, including green rust (39) and

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magnetite (87, 88) can dehalogenate CT. Moreover, the products of CT reduction by magnetite are essentially the same as those produced with granular iron (87). Magnetite also dehalogenates TCE (122). Several studies have demonstrated that magnetite is the predominant mineral phase formed during corrosion of granular iron (14, 15, 95). In addition to being a strong reductant itself, magnetite is also a good conductor (29), and

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appears to facilitate reductive dehalogenation by allowing the transfer of electrons from the underlying Fe(0) to the alkyl halide at the magnetite surface. Fe(II) adsorbed to goethite, nontronite, lepidocrocite, hematite and siderite reduces HCA and CT (3, 45, 118). Charlet et al. (26) note that the measured by Orth and Gillham (97) from reduction of TCE and PCE by granular iron is closer to that of the redox couple for oxidation of Fe(II)sorbed to but is not similar to the Fe(0)/Fe(II) or the redox couples. This observation led Charlet et al. (26) to conclude that the dehalogenation reaction is mediated by Fe(II) adsorbed to the mineral surface, with Fe(0) acting as a source of Fe(II). Iron sulfide minerals also promote reductive dehalogenation. CT and HCA are dehalogenated by pyrite and marcasite, as well as biotite and vermiculite, in the presence of dissolved hydrogen sulfide (73, 74, 75). It has also been shown that FeS dehalogenates CT (10), TCE and PCE (21, 61), as well as many other polyhaloalkanes (20, 22). It is interesting to note that TCE is dehalogenated more rapidly than PCE by FeS (21), because the same trend in reactivity is seen with granular iron (8,41). As with granular iron, reductive elimination appears to be the dominant pathway for dehalogenation of the chlorinated ethenes by FeS. However, FeS does not appear to be capable of hydrogenating double bonds, resulting in the accumulation of acetylene as the ultimate reaction product (21). FeS also dehalogenates ethanes containing 3 to 6 chlorines, with rate constants demonstrating a strong positive correlation with values (22). Addition of sulfur compounds, such as to granular iron enhances its reactivity with TCE (61, 81). All commercially available granular iron contains at least traces of sulfur, allowing FeS to be formed as a product of the reaction of the metal with water. Hassan (61) has theorized that in the granular iron system, FeS catalyzes the reaction of alkyl halides with hydrogen, with iron serving primarily as a source of hydrogen gas. This interpretation, if correct, does not rule out an important role of Fe(0) and nascent hydrogen in the hydrogenation of double bonds. Johnson et al. (69) have also investigated the role of the oxide coating in the granular iron system on the rate of CT reduction by studying the effects of both redox active and redox inactive ligands and anions. Both types of ligands inhibited the reaction. The authors hypothesized that these ligands block CT from interacting with Fe(II) on the passive oxide coating of the iron metal, thereby slowing the reaction. However, the researchers also concluded that the slower reaction rates of CT, TCE, and nitrobenzenes with magnetite and Fe(II) adsorbed to goethite (relative to granular iron) might be due to one of two factors (69). Either reaction within corrosion pits, presumably with Fe(0), is an important process, or the Fe(II) generated in the granular iron system is produced more efficiently or is more reactive than the Fe(II) that may be introduced from solution. Amonette et al. (3) have noted that Fe(II) adsorbed to goethite reduces CT to CF exclusively, which suggests that further reduction of CF to DCM requires the presence of granular iron. Similarly, biogenic magnetite does not reduce 6-bromo-1 -hexene, while granular iron does (128). The products of the reduction of CT on an oxide-free iron electrode are the same as those observed in the granular iron system (82), which might suggest that Fe(0) acts as the reductant via pitting corrosion. Several lines of evidence suggest that organometallic species are formed during reduction of alkyl halides by granular metals. There is ample evidence in the organic synthesis literature to suggest that both iron and zinc are likely to form organometallic

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intermediates during reduction of alkyl halides (25, 30, 70, 85, 90, 131). In addition, product distributions suggest that geminal polyhalides, such as CT and 1,1,1-TCA, are reduced in the presence of granular iron to hydrocarbons without mono- and dihalide intermediates, a process which could occur via an organometallic intermediate. Boronina et al. (16) suggested that an organometallic carbenoid could account for the production of CO from reductive dehalogenation of CT by zinc. Fennelly and Roberts (43) also suggested that 1,1,1-TCA is fully dehalogenated by iron via a carbenoid intermediate. The production of 1,5-hexadiene and coupling products during reduction of 6-bromo-1 hexene by iron suggests a organometallic intermediate (128). Coupling products might also arise from organometallic intermediates. Sivavec and Horney (121) hypothesized that products resulting from chlorinated ethene reduction by iron arise from the coupling of intermediate radicals. Others (128, 129) have demonstrated that any radicals produced during reduction of alkyl halides by zinc or iron persist for a very short time and do not build up to concentrations where radical-radical coupling becomes an important process. Reduction of chlorinated ethenes and 1,1,1trichloroethane by Fe(0) results in the formation of to hydrocarbons (2, 8, 23, 43, 97, 116). Arnold and Roberts (8) noted that such hydrocarbons are produced during the reduction of chlorinated ethenes only when acetylene is formed as an intermediate. Acetylene is known to undergo radical polymerization on metal surfaces (99). In contrast, others have hypothesized that the production of longer carbon chains from the reduction of alkyl halides by granular iron might arise from organometallic intermediates (16, 17, 43, 128). Kinetic evidence also supports the formation of organometallic intermediates. Arnold and Roberts (8) have suggested that the decrease in rate constants with increasing oneand two-electron reduction potentials is evidence that the formation of a organometallic intermediate is the rate-limiting step in the reaction of chlorinated ethenes with iron (Figure 9.7). In addition, they noted that the isotope effect for reduction of chlorinated ethenes with granular iron is 1.017-1.022 (123), which is large enough to be considered a primary isotope effect involving a change in bonding at carbon. Because of the inverse relationship between reactivity and reduction potentials, breakage of the C-Cl bond does not appear to be the rate-limiting step. Thus, formation of a C-metal bond may be responsible for the observed kinetic isotope effect (8). However, the formation of an organometallic intermediate does not imply anything about the active reductant in the iron system [Fe(0) or Fe(II)].

6. SUMMARY A great deal of sometimes conflicting information exists regarding the roles of Fe(0), Fe(II), and hydrogen species in the granular iron system. More research is needed to identify the active reductant(s) in this system. It should be noted that different compounds may react via different mechanisms and with different reductants in the highly complex granular iron system. It has been clearly demonstrated that CT, which possesses very favorable one- and two-electron reduction potentials (0.085 and 0.673 V, respectively) (130), is reduced by Fe(II) sorbed to mineral surfaces, by magnetite, by conduction-band electrons, and by Fe(0) itself. It is reasonable to assume that all of these processes are occurring (at widely varying rates) in the granular iron system. In contrast,

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a compound such as trans-DCE, with less favorable reduction potentials (130), might be unreactive with weaker reductants and might react solely with Fe(0). In addition, the chlorinated ethenes are thought to react with granular iron via interaction of the double bond with Fe, resulting in the formation of an organometallic intermediate, a pathway which is unavailable to the chlorinated methanes and ethanes.

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It is interesting to note the differences observed between the reaction of alkyl halides with granular iron and with zinc. In the presence of zinc, reductive dehalogenation reactions do not display saturation kinetics, and do not yield coupling products, but do increase in rate with increasing one- and two-electron reduction potentials. In contrast, reactions with iron display saturation kinetics, a decrease in rate with increasing redox potentials for the chlorinated ethenes, and frequently yield coupling products. These differences may indicate that the mechanisms of alkyl halide reaction with iron and zinc are not the same. In this respect, it is important to note that because zinc has only two stable oxidation states [Zn(0) and Zn(II)], cannot promote reductive dehalogenation reactions. Zero-valent metals have been demonstrated to be effective and efficient reagents for the destruction of a wide range of halogenated organics in water. A great deal of research has investigated the long-term performance of in situ barriers, with emphasis on understanding how groundwater composition, precipitation of mineral phases, passivation, and microbial activity can influence reactivity (42, 46, 58, 79, 84, 94, 98, 108, 113, 114, 115, 132, 134). A better understanding of the mechanism through which zero-valent metals reduce organohalides will aid in the design ofmore effective systems for in situ treatment of contaminated groundwater. Specifically, understanding the factors that drive reductive dehalogenation of CT toward the production of fully dehalogenated species via a carbene intermediate, and away from the production of chloroform, would allow the design oftreatment systems that could effectively remediate groundwater contaminated with CT (and other organohalide candidates), without producing toxic and recalcitrant intermediates. Further research should also focus on developing effective and non-toxic catalysts for the reductive dehalogenation of aromatic compounds such as PCBs. REFERENCES 1.

Agrawal A & Tratnyek PG (1996) Reduction of nitro aromatic compounds by zero-valent iron metal. Environ. Sci. Technol. 30:153-160 2. Allen-King RM, Halket RM & Burris DR (1997) Reductive transformation and sorption of cis- and trans-1,2- dichloroethene in a metallic iron-water system. Environ. Toxicol. Chem. 16:424-429 3. Amonette JE, Workman DJ, Kennedy DW, Fruchter JS & Gorby YA (2000) Dechlorination of carbon tetrachloride by Fe(II) associated with goethite. Environ. Sci. Technol. 34:4606-4613 4. Archer WL & Simpson EL (1977) Chemical p r o f i l e of p o l y c h l o r o e t h a n e s and polychloroalkenes. Ind. Eng. Chem. Prod. Res. Dev. 16:158-162 5. Archer WL (1982) Aluminum-1,1,1trichloroethane. Reactions and inhibition. Ind. Eng. Chem. Prod. Res. Dev. 21:670-672 6. Arnold WA & Roberts AL (1998) Pathways of chlorinated ethylene and chlorinated acetylene reaction with Zn(0). Environ. Sci. Technol. 32:3017-3025

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Arnold WA, Ball WP & Roberts AL (1999) Polychlorinated ethane reaction with zero-valent zinc: Pathways and rate control. J. Contam. Hydrol. 40:183-200 Arnold WA & Roberts AL (2000) Pathways and kinetics of chlorinated ethylene and chlorinated acetylene reaction with Fe(0) particles. Environ. Sci. Technol. 34:1794-1805 Assaf-Anid N & Nies L (1995) Reductive dechlorination of aliphatic and aromaticcompounds by zero-valent metals. Abstracts of Papers of the American Chemical Society 209:219-ENVR Assaf-Anid N & Lin K-Y (2002) Carbon tetrachloride reduction by and FeS with vitamin as organic amendment. ACSE J. Environ. Eng. 128:94-99 Assaf-Anid N, Anid PJ, Häggblom M, Nies L & Vogel TM. Reductive dechlorination of chlorinated biphenyls: Limitations and perspectives. in prep. Astrup T, Stipp SLS & Christensen TH (2000) Immobilization of chromate from coal fly ash

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283 Chemosphere 38:565-571

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PART IV. ENVIRONMENTAL FATE AND APPLICATIONS

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Chapter 10 BIOAVAILABILITY OF ORGANOHALIDES KYOUNGPHILE NAM1 AND JEROME J. KUKOR2 1

School of Civil, Urban and Geosystem Engineering, Seoul National University, Seoul, The Republic of Korea 2 Biotechnology Center for Agriculture and the Environment and Department of Environmental Sciences, Rutgers University, New Brunswick, NJ, USA

1. INTRODUCTION The extensive use of halogenated organic compounds (HOCs) in dyes, pesticides, fire retardants and solvents has resulted in widespread environmental contamination, either by broad use (in the case of certain pesticides that do not degrade rapidly and remain as persistent environmental contaminants), or by improper disposal or other unintentional releases. In addition to these anthropogenic sources of HOCs, certain marine invertebrates and algae, and some terrestrial fungi and bacteria, biosynthesize a variety of halogenated compounds which can also contribute to the overall input of these compounds into the environment (26). Regardless of their origin, certain HOCs can pose potential hazards that either arise directly from acute or chronic toxicity, or from longterm effects, due to the tendency of HOCs to bioaccumulate and bioconcentrate. In general, removal of halogen substituents tends to reduce or eliminate toxic effects, and microorganisms capable of mineralizing such compounds have been reported (13, 29). However, many HOCs seem to be resistant to microbial attack, and as the number of halogen substituents on a compound increases, the compound becomes less biodegradable (18). This is significant since many of the anthropogenic HOCs are multiply-halogenated (75). Biodegradation is one of the major mechanisms by which HOCs are transformed, immobilized, or mineralized in the environment (2). A clear understanding of the major processes that affect the interactions between HOCs and the microorganisms which mediate biodegradation in the environment is thus important for determining persistence of HOCs, for predicting in situ transformation rates, and for developing site remediation programs. This chapter focuses on some of the major factors governing bioavailability of HOCs in the environment, with emphasis on how bioavailability constraints affect biodegradation. Emphasis will be placed on some of the major classes of HOCs for which bioavailability studies have been performed, e.g., pesticides (DDT [1,1,1trichloro-2,2-bis[p-chlorophenyl]ethane],aldrin[1,2,3,4,10,10-hexachloro-1,4,4a,5,8,8ahexahydro-exo-1,4-endo-5,8-dimethanonaphthalene], dieldrin [1,2,3,4,10,10hexachloro-6,7-epoxy-1,4,4a,5,6,7,8,8a-endo,exo-1,4,5,8-dimethanonaphthalene], Dehalogenation: Microbial Processes and Environmental Applications, pages 291-302 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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lindane [1,2,3,4,5,6-hexachlorocyclohexane], chlordane [1,2,4,5,7,8,8-octachloro2,3,3a,4,7,7a-hexahydro-4,7-methanoindane], 2,4-D [2,4-dichlorophenoxyacetic acid], and atrazine [2-chloro-4-ethylamino-6-isopropyl-l,3,5-triazine]), chlorinated solvents (PCE [perchloroethene] and TCE [trichloroethene]), and polychlorinated biphenyls (PCBs). Many of the fundamental processes that affect the bioavailability of hydrophobic compounds in complex environmental matrices (e.g., soil, aquifers and sediments) have not been investigated with a focus on HOCs as the model contaminants. Instead, a major focus of research has been on other environmentally persistent contaminants, such as polynuclear aromatic hydrocarbons (PAHs). We will also make reference to such studies for purposes of illustration of fundamental principles and mechanisms. 2. BIODEGRADABILITY AND BIOAVAILABILITY Biodegradation is a (largely) microbially-mediated process that can result in partial transformation or complete mineralization of organic compounds. The biodegradative process can contribute to a reduction of, or complete elimination of, anthropogenic organic contaminants in the environment. This is achieved either through specific enzymatic mechanisms that accommodate the target compound, or through fortuitous transformations from which the biodegradative population may, or may not, gain any benefit in terms of carbon or energy for growth. For successful biodegradation of a target compound in a particular environment, one or more microorganisms capable of degrading the compound of interest must be present in the environment containing the compound. However, the mere presence of a microorganism capable of biodegrading the compound, although necessary, is not sufficient for biodegradation to occur. Additional necessary components are accessibility of the compound to the microorganism and favorable conditions in the local environment for the biodegradative activities of the microorganism (4). It has often been observed that biodegradation of an organic compound is limited by environmental constraints, even when microorganisms competent to degrade the compound are present in the same environment with the compound. This limitation on intrinsic biodegradability is described by the term, “bioavailability.” In this context, bioavailability describes the degree of availability or accessibility of a compound to microorganisms under a particular set of environmental conditions. It is important to note that bioavailability is operationally defined and is context dependent. Bioavailability is not an inherent property of a compound or of a microorganism. Bioavailability is extremely important, in that bioavailability constraints frequently account for the environmental persistence of organic compounds that are inherently biodegradable, and that might otherwise be assumed to be readily metabolized as a result of inappropriately designed treatability studies (4). Given microorganisms with the appropriate catabolic potential in an environmental matrix (such as soil, aquifers and sediments), the inherent biodegradability ofa compound does not change. However, the bioavailability of the compound can be altered as a result of interactions between the compound, the microorganisms, and the characteristics of the matrix materials. The possible interactions that can occur among these components are illustrated in Figure 10.1.

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3. BIOAVAILABILITY OF ORGANIC CONTAMINANTS IN THE ENVIRONMENT The fact that a compound in aqueous solution may be readily utilized by bacteria does not guarantee that the compound will be similarly biodegraded by the same bacteria in a complex environmental matrix, such as soils, aquifer or sediment solids, which often are associated with natural organic matter. Determinants of the bioavailability of organic compounds in such complex environments depend on a variety of factors. These include the physical state (85), aqueous solubility (79), and chemical structure (8) of the compound; the extent of sorption of the compound to environmental matrix materials (66); the composition, structure and size of matrix aggregates (52, 71); the moisture content of the environmental matrix (73); and the types of microorganisms present (27). The physico-chemical properties of organic compounds play a major role in determining their bioavailability. One measure that is frequently used to assess the extent to which a compound will partition into an aqueous vs. an organic phase in multiphasic systems is the octanol-water partition coefficient This coefficient serves as an indicator of the tendency of a compound to partition into natural organic matter, which is thought to behave as an organic solvent. Organic compounds, when introduced into natural environments, tend to sorb to clay minerals, to natural organic matter (e.g., humic substances), or to other complex mineral/organic solids. This sorption can be defined as any accumulation of organic compounds at a surface, or within the matrix of an aggregate or particle. A considerable amount of research has been done to elucidate the basic mechanisms of sorption, and two general models are current in the literature: hydrophobic sorption, and partitioning (15, 34). Hydrophobic sorption refers to the accumulation of hydrophobic organic compounds on hydrophobic surfaces. In this model, there are sites available for interaction between the organic compound and the surface of the sorbent. In the partitioning model, on the other hand, the interaction between the organic compound and the sorbent is described as a continuous process in which the hydrophobic compound enters into the three-dimensional structure of the

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sorbent (e.g., natural organic matter) in a gradual manner. In fact, the two processes may not be separable and may operate simultaneously in the natural environment (64). The initial phase of the process may involve accumulation of hydrophobic compounds at hydrophobic attachment sites on the sorbent, followed by a second phase where the sorbed compound may partition into the matrix of the sorbent. It is widely accepted that this second phase sorption results in the formation of sorbed molecules that are resistant to desorption and biological degradation, and even to chemical extraction. A detailed review of the mechanisms involved in this type of multiphase partitioning is beyond the scope of this chapter and may be found elsewhere (45, 64). Regardless of the relative contribution of the two mechanisms, it is clear that sorption of organic compounds has a profound impact on their bioavailability to microorganisms. Organic compounds introduced into a complex environmental matrix such as soil, aquifer or sediment materials, are likely to be less bioavailable than the same compounds in solution, presumably due to the sorption of these compounds. One of the most important sorbents of hydrophobic compounds in nature is natural organic matter (15), and reduction in bioavailability of hydrophobic compounds is directly related to organic matter content of the matrix. This reduced bioavailability is often accompanied by a reduction in apparent toxicity of a compound (80, 82). Peterson et al. (59), in a study of how soil properties affected toxicity, determined values for DDT towards Drosophila melanogaster. These researchers concluded that DDT may present little biological hazard when present in soils rich in organic matter. Other studies have also reported an inverse relationship between toxicity (or biodegradation) and extent of sorption for a variety of organic compounds, including chlorinated insecticides and PAHs (33, 47). Although not extensively studied, there is some evidence that the quality of natural organic matter may influence bioavailability by affecting the partitioning process of organic compounds (25, 41). A study (77) on several chlorinated pesticides (alachlor, metolachlor, propachlor) suggested that values decreased as the ratio of oxygenplus-nitrogen to carbon (O+N:C) of a sorbent increased.

4. DEGRADATION OF SORBED COMPOUNDS Sorption of organic compounds often results in the formation of desorption-resistant fractions. This process reduces the overall rate of release of the compound from the sorbent, and thereby decreases bioavailability and results in increased environmental persistence. Studies with (7, 68) in soil have shown that mass transfer – not intrinsic microbial biodegradative activity – is in most cases the critical factor limiting biodegradation of this compound. Reduced bioavailability of organic contaminants in soil is the result of reduction in mass transfer of the contaminant from the sorbent to the degradative microorganisms. Consistent with this, several lines of evidence indicate that organic compounds are available to bacteria only when they are present in the aqueous phase. Ogram et al. (55), by use of mathematical models and experimental observation, have shown that only soluble 2,4-D is available to degradation by a model bacterium. Similar results were obtained for the degradation of fatty acids by marine bacteria (31). From these and comparable studies, it appears that the bioavailability of sorbed compounds is less than that of compounds in solution, and it

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appears that the desorption rate can control bioavailability of the sorbed compounds (69). In contrast, although sorption often results in a decline in bioavailability, other lines of evidence suggest that at least some part of the sorbed fraction can be available for biodegradation by bacteria (43). This sorbed fraction can be available for microbial utilization owing, in part, to microbial processes that facilitate desorption of sorbed compounds (86). Remberger et al. (67) have provided evidence that even tightly bound residues are accessible to biological transformation. In some instances, bacteria may act on sorbed compounds without the need for prior desorption of the compounds (11, 20, 28, 74). Differences in microbial metabolic activities have also been found to affect bioavailability. Harms and Zehnder (32) reported that the rate at which 3chlorodibenzofuran became available to model bacteria was influenced by the specific metabolic activity of the organisms, as well as the tendency of the organisms to adhere to the matrix in which the 3-chlorodibenzofuran was sorbed. Guerin and Boyd (27) found that sorbed naphthalene was less available than naphthalene in aqueous solution for several bacterial strains tested, however the magnitude of the sorption effect was highly organism-specific. Interestingly, other researchers have found that a variety of factors that affect overall microbial metabolism, including methods of inoculum preparation, can influence the results obtained in bioavailability studies (46, 70).

5. FORMATION OF DESORPTION-RESISTANT FRACTIONS Sorption of organic compounds to complex environmental matrices such as soil, aquifer solids and sediments has been well established, largely from research done on the environmental persistence of halogenated pesticides such as 2,4-D and picloram (40). Prior to the 1980s, tests on the sorption of organic compounds were routinely conducted over a 24- to 72-hour time interval, based largely on the assumption that sorption was a rapid process. However, in 1980 Karickhoff (37) presented evidence that sorption may be a slow process, extending from days to weeks. Subsequent to this study, two-compartment sorption models were developed (38) that consisted of two domains: a rapid sorption component (S1), and a much slower sorption component (S2). Organic compounds sorbed during the S1 phase were easily extracted from the sorbent by relatively mild extraction conditions, and these sorbed components were readily bioavailable. However, compounds or fractions in the S2 sorption phase were resistant to solvent extraction as well as to microbial utilization. This biphasic sorption behavior, often called nonequilibrium sorption, may result in the formation of desorption-resistant fractions, which may account for the environmental persistence of some organic compounds. A conceptual diagram illustrating initial sorption, formation of desorptionresistant fractions, and the biological consequences of these processes, is presented in Figure 10.2. Numerous laboratory studies with historically contaminated soils, as well as environmental samples to which organic compounds have been intentionally added, are supportive of a model in which desorption-resistant fractions develop as a consequence of irreversible sorption. A number of studies have shown that a fraction of sorbed organic compounds is difficult to remove from the sorbent matrix, and this fraction has

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been termed the “irreversible” or “resistant” fraction (60). DiToro and Horzempa (21) have provided evidence that hexachlorobiphenyl sorbed onto sediments has both a reversible and an irreversible fraction, and similar conclusions have been drawn from sorption studies with PCB congeners (17) and PAHs (23, 36). Biodegradation studies that have been conducted on sorbed organic compounds have demonstrated that at least part of the desorption-resistant fraction is available to microorganisms, however a significant part of this fraction appears not to be utilized in biologically meaningful time periods (24, 84). Investigations into the mechanistic basis of nonequilibrium sorption have been conducted using several halogenated and non-halogenated compounds. The formation of slowly reversible sorbed fractions in soil has been observed for halogenated alkanes and alkenes, including PCE, TCE, and 1,2-dibromoethane (EDB) (61). These compounds are volatile, have weak to moderate equilibrium sorption tendencies, and do not contain strongly interacting functional groups that could form strong chemical bonds with soil matrix materials. This suggests that strong chemical reactivity is not required for the formation of slowly reversible and desorption-resistant fractions. In addition to information obtained from chemical reactivity considerations, several studies have been done on the kinetics of the process. Pavlostathis and Jaglal (57) observed an initially fast rate ofdesorption, followed by a second-phase slower rate of desorption for soil samples from a TCE-contaminated site. Moreover, the extent of TCE desorption decreased with increasing residence time of the compound in soil (58). In a 300-day study with picloram (51), the amount of the compound readily released from soil decreased appreciably with time. Connaughton et al. (19) investigated the rate of release of freshly-added and 30year aged naphthalene and found that increased residence time allowed the PAH to be sorbed to compartments in soil that exhibited very slow desorption kinetics. Similar observations were reported by Pignatello and Huang (63), who examined the reversibility of sorption of atrazine and metolachlor (2-chloro-N-[2-ethyl-5methylphenyl]-N-[2-methoxy-1-methylethyl]-acetamine) in a field soil. Their results suggested that the soil may contain a large fraction of slowly reversible HOC

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contaminants, and that the size of this fraction increases with time. Although most studies on nonequilibrium sorption processes have focused on chemicals that, based on their physico-chemical properties, would be predicted to behave in a strongly hydrophobic manner, it should be noted that more polar compounds can also exhibit irreversible sorption behavior. Formation of desorption-resistant fractions has been reported for PCBs (12, 17, 21) TCE (57, 58), PCE and EDB (61), as well as PAHs (23, 26), and benzene and toluene (1).

6. “AGING” AS AN EXPLANATION FOR REDUCTION IN BIOAVAILABILITY A growing body of evidence has been accumulated in support of the argument that the bioavailability of some organic compounds, as well as the ease of extractability of these compounds, diminishes as the residence time of the compounds in complex environmental matrices such as soil, sediment, and aquifers increases. A considerable amount of this supporting evidence comes from long-term monitoring studies of the persistence of HOCs applied to agricultural fields. For example, in a long-term study of DDT persistence, it was found that there was detectable disappearance of the compound during the first 10 years after application, but little change in disappearance rates could be detected in subsequent years (54). A graphic representation of the disappearance kinetics reveals a hockey stick-shaped curve. A similarly shaped disappearance curve has been observed for HOC pesticides such as aldrin, dieldrin, lindane, and chlordane in soil (14, 44, 54, 56). This time-dependent, inverse bioavailability/extractability process has been termed “aging”, or sequestration (3). A study with field soil contaminated with EDB showed that aging altered the bioavailability of the compound to indigenous microorganisms (76). Freshly added EDB could be totally destroyed by the indigenous microbial population within a few weeks, whereas very little of the EDB that had been present in the soil for at least 19 years was transformed in the 30 days of the experiment. Similar observations have been reported for PAHs (35, 78). Aging does not include reactions that alter the structure of the parent molecule, such as polymerization or covalent bonding to matrix materials. Therefore, aged compounds should be readily biodegraded if the sequestered fractions are extracted from a matrix (soil, sediment, or aquifer solids) and added back to the same matrix. This, in fact, was demonstrated experimentally for PAHs by Weissenfels et al. (82). The time-dependent reduction in bioavailability that has been observed for microorganisms has been observed for multicellular organisms as well. Scribner et al. (72) showed that aged simazine (2-chloro-4,6-bis[ethylamino]-S-triazine) residues were less bioavailable to sugarbeet roots, whereas unaged simazine was able to slowly desorb into the soil solution. The bioavailability of sediment-sorbed PAHs to the amphipod Diporeia sp. and to the oligochaete Lumbriculus variegatus also decreased with increasing length of contact time between the sediment and the contaminant (30, 42). Similar findings have been obtained with earthworms (39) and plants (9). One of the consequences of the aging or sequestration process is that, for some compounds, a decrease in bioavailability also appears to correlate with an apparent decrease in toxicity. It has been found that the of DDT to fruit flies increases (i.e., the toxicity decreases) as the compound resides in soil for increasing periods of time (59). It should also be noted that reduction in bioavailability due to aging or sequestration

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may not occur, or may not have an appreciable effect, for all compounds in all environments. For example, both freshly added, as well as historically aged PCDDs (polychlorinated dibenzo-p-dioxins) were equally bioavailable for indigenous microbes carrying out reductive dechlorination processes (6).

7. MECHANISMS OF “AGING” Since aging, or sequestration, is believed to be the product of a very slow process of sorption within the matrix of the sorbent material, the mechanisms that underlie aging would be expected to be closely linked to the processes described for nonequilibrium sorption. The general view is that organic compounds are taken up rapidly via sorption from the bulk aqueous phase onto external sorption sites on the sorbent, and then move slowly into internal remote sites, or sites that are less accessible with time to solvent extraction or to microbial uptake, or both. This physical sequestration of organic compounds may occur by association with the polymeric structure of the matrix organic matter (a process that is referred to as intraorganic matter diffusion), or by diffusion into tortuous pores located within the matrix of a microporous particle (a process that is referred to as retarded intraparticle diffusion). Brusseau and Rao (10), in reviewing the available data prior to 1990, concluded that there was considerable support for intraorganic matter diffusion as a mechanism to explain the nonequilibrium sorption of hydrophobic organic compounds. This model envisions organic compounds as partitioning into natural organic matter (16, 22). An implicit assumption of the model is the three-dimensional structure of natural organic matter. In support of this, a number of experimental and modeling studies have provided evidence that natural soil organic matter is comprised of a three-dimensional network of polymeric chains with a relatively open and flexible structure perforated with voids (12, 40, 48, 83). The second proposed mechanism for sequestration of organic compounds – the retarded intraparticle diffusion model – envisions aqueous-phase diffusion ofa compound within micropores ofsorbents, with diffusion ofthe compound in pore water being retarded by sorption onto pore walls (87). The finding that bioavailability, or desorption of environmentally persistent HOCs, such as aliphatic halocarbons, 1,2,4,5-tetrachlorobenzene, PCE, and EDB, were enhanced by pulverization provides support for the intraparticle diffusion model (5, 62, 76). It is likely that both mechanisms may operate simultaneously, and their relative significance may depend on the characteristics of the sorbates and the sorbents. It is also possible that sorbed compounds may diffuse into organic matter-coated micropores where the two mechanisms would work together. In this scenario, sorbed compounds may undergo extensive sorption/desorption processes inside micropores. Indeed, a sorption model having a rapid component governed by partitioning, and a slow component controlled by micropore diffusion, provides a good fit to data generated from the elution of freshly added and aged atrazine and metolachlor from a fine sandy loam (65). Such a mechanism would be expected if micropores are coated with organic matter, and existing evidence indicates that a significant amount of natural organic matter resides within pores of less than 10 nm in diameter in soils and sediments (49, 50). In a model system using synthetic beads, microporous sorbents having hydrophobic surfaces showed a significant decrease in bioavailability of the test compound, phenanthrene, to bacteria.

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In contrast, nonporous hydrophobic beads and microporous hydrophilic beads did not affect bioavailability as drastically (53). More recently, two additional models have been developed to account for the complex nature of sorbent/sorbate interactions vis-à-vis the role of soil organic matter in the non-equilibrium sorption of hydrophobic organic compounds. These are the dualmode sorption model (89), and the distributed reactivity model (81). Both models originate from the same observations of different competitive effects in the sorption of nonpolar organic compounds (in this case, atrazine and its analogs). The dual-mode sorption model proposes that soil organic matter has two types of sorption sites: a partition domain and an adsorption domain. Adsorption sites are believed to reside internal to the soil organic matter matrix, and are proposed to be responsible for most of the slow and apparently irreversible sorption (88). In contrast, the distributed reactivity model proposes three different sorption sites: an exposed inorganic mineral domain (domain I), an amorphous and swollen organic matter domain (domain II), and a condensed and highly crystalline organic matter domain (domain III). In this model, domain II-associated processes are associated with most of the observed partitioning of HOCs, whereas domain III-associated processes account for slow and nonlinear sorption that results in the formation of desorption-resistant fractions in soil.

8. SUMMARY From the brief overview and analysis provided in this chapter, it is clear that bioavailability is an extremely important criterion when considering the environmental fate and persistence of HOCs. Because of the hydrophobic nature of many of the compounds in this class, their sorption onto, and sequestration into, natural organic matter matrices will more often than not govern and limit the potential intrinsic biodegradation of many of these compounds. Moreover, these same sorption and sequestration processes that affect biodegradation by microorganisms can also affect the bioavailability of these compounds to multicellular organisms. This, in turn, can have a profound impact on assessment of apparent toxicity, as well as the impact of long-term exposure to contaminated environmental matrices. REFERENCES Ahlert WK & Uchrin CG (1990) Rapid and secondary sorption of benzene and toluene by two aquifer solids. J. Hazard. Mater. 23:317-333 2. Alexander M (1981) Biodegradation of chemicals of environmental concern. Science 211:132-138 3. Alexander M (1995) How toxic are toxic chemicals in soil? Environ. Sci. Technol. 29:2713-2717 4. Alexander M (1999) Biodegradation and Bioremediation, 2d ed. Academic Press, San Diego, CA 5. Ball WP & Roberts PV (1991) Long-term sorption of halogenated organic chemicals by aquifer materials. 2. Intraparticle diffusion. 1.

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NAM AND KUKOR pond sediment. Science 214:790-792 23. Fu G, Kan AT & Tomson MB (1994) Adsorption and desorption hysteresis of PAHs in surface sediment. Environ. Toxicol. Chem. 13:1559-1567 24. Fu MH, Mayton H & Alexander M (1994) Desorption and biodegradation of sorbed styrene in soil and aquifer soilds. Environ. Toxicol. Chem. 13:749-753 25. Garbarini DR & Lion LW (1986) Influence of the nature of soil organics on the sorption of toluene and trichloroethylene. Environ. Sci. Technol. 20:1263-1269 26. Gribble GW (1994) The natural production of chlorinated compounds. Environ. Sci. Technol. 28:310A-319A 27. Guerin WF & Boyd SA (1992) Differential bioavailability of soil-sorbed naphthalene to two bacterial species. Appl. Environ. Microbiol. 58:1142-1152 28. Guha S & Jaffé PR (1996) Bioavailability of hydrophobic compounds partitioned into the micellar phase of nonionic surfactants. Environ. Sci. Technol. 30:1382-1391 29. Hardman DJ (1991) Biotransformation of halogenated compounds. Crit. Rev. Biotechnol. 11:1-40 30. Harkey GA, van Hoof PL & Landrum PF (1995) Bioavailability of polycyclic aromatic hydrocarbons from a historically contaminated sediment core. Environ. Toxicol. Chem. 14:1551-1560 31. Harmansson M & Marshall KC (1985) Utilization of surface localized substrate by non-adhesive marine bacteria. Microb. Ecol. 11:91-105 32. Harms H & Zehnder AJB (1995) Bioavailability of sorbed 3-chlorodibenzofuran. Appl. Environ. Microbiol. 61:27-33 33. Harris CR (1966) Influence of soil type on the activity of insecticides in soil. J. Econ. Entomol. 59:1221-1225 34. Hassett JJ & Banwart WL (1989) The sorption of nonpolar organics by soils and sediments. In: Sawhney BL, Brown K (Eds) Reactions and Movement of Organic Chemicals in Soils, (pp 31-45). Soil Science Society of America, Madison, WI 35. Hatzinger PB & Alexander M (1995) Effect of aging of chemicals in soil on their biodegradability and extractability. Environ. Sci. Technol. 29:537-545 36. Kan AT, Fu G & Tomson MB (1994) Adsorption/desorption hysteresis in organic pollutants and soil/sediment interaction. Environ. Sci. Technol. 28:859-867 37. Karickhoff SW (1980) Sorption kinetics of

BIOAVAILABILITY OF ORGANOHALIDES hydrophobic pollutants in natural sediments. In: Baker RA (Ed) Contaminants and Sedimants: Analysis, Chemistry, Biology. Vol 2, (pp 193-205). Ann Arbor Science Publishers, Ann Arbor, MI 38. Karickhoff SW & Morris KR (1985) Sorption dynamics of hydrophobic pollutants in sediment suspensions. Environ. Toxicol. Chem. 4:469-479 39. Kelsey JW & Alexander M (1997) Declining bioavailability and inappropriate estimation of risk of persistent compounds. Environ. Toxicol. Chem. 16:582-585 40. Khan SU (1973) Equilibrium and kinetics studies of the adsorption of 2,4-D and picloram on humic acid. Can. J. Soil Sci. 53:429-434 41. Kile DE, Chiou CT, Zhou H, Li H & Xu O (1995) Partition of nonpolar organic pollutants from water to soil and sediment organic matters. Environ. Sci. Technol. 29:1401-1406 42. Landrum PF, Eadie BJ & Faust WR (1992) Variation in the bioavailability of polycyclic aromatic hydrocarbons to the amphipod Diporeia (spp.) with sediment aging. Environ. Toxicol. Chem. 11:1197-1208 43. Laor Y, Strom PF & Farmer WJ (1996) The effect of sorption on phenanthrene bioavailability. J. Biotechnol. 51:227-234 44. Lichtenstein EP, DePew LJ, Eshbaugh EL & Sleesman JP (1960) Persistence of DDT, aldrin, and lindane in some midwestern soils. J. Econ. Entomol. 53:136-142 45. Luthy RG, Aiken GR, Brusseau ML, Cunningham SD, Gschwend PM, Pignatello JJ, Reinhard M, Traina SJ, Weber WJ Jr & Westall JC (1997) Sequestration of hydrophobic organic contaminants by geosorbents. Environ. Sci. Technol. 31:3341-3347 46. Madsen EL, Mann CL & Bilotta SE (1996) Oxygen limitations and aging as explanations for the field persistence of naphthalene in coal tar-contaminated surface sediments. Environ. Toxicol. Chem. 15:1876-1882 47. Manilal VB & Alexander M (1991) Factors affecting the microbial degradation of phenanthrene in soil. Appl. Microbiol. Biotechnol. 35:401-405 48. Maurice PA & Namjesnik-Dejanovic K (1999) Aggregate structure of sorbed humic substances observed in aqueous solution. Environ. Sci. Technol. 33:1538-1541 49. Mayer LM (1994) Relationships between mineral surfaces and organic carbon concentrations in soils and sediments. Chem. Geol. 114:347-363 50. Mayer LM (1994) Surface area control of organic accumulation in continental shelf

301 sediments. Geochim. Cosmochim. Acta 58:1271-1284 51. McCall PJ & Agin GL (1985) Desorption kinetics of picloram as affected by residence time in the soil. Environ. Toxicol. Chem. 4:37-44 52. Mott SC, Groenevelt PH & Voroney RP (1990, Biodegradation of a gas oil applied to aggregates of different sizes. J. Environ. Qual. 19:257-260 53. Nam K & Alexander M (1998) Role of nanoporosity and h y d r o p h o b i c i t y in sequestration and bioavailability: Tests with model solids. Environ. Sci. Technol. 32:71-74 54. Nash RG & Woolson EA (1967) Persistence of chlorinated hydrocarbon insecticides in soils. Science 157:924-927 55. Ogram AV, Jessup RE, Ou LT & Rao PSC (1985) Effects of sorption on biological degradation rates of (2, 4 - d i c h l o r o phenoxy)acetic acids in soils. Appl. Environ. Microbiol. 49:582-587 56. Onsager JA, Rusk HW & Butler LI (1970) Residues of aldrin, dieldrin, chlordane, and DDT in soil and sugarbeets. J. Econ. Entomol. 63:1143-1146 57. Pavlostathis SG & Jaglal K (1991) Desorptive behavior of trichloroethylene in contaminated soil. Environ. Sci. Technol. 25:274-279 58. Pavlostathis SG & Mathavan GN (1992) Desorption kinetics of selected volatile organic compounds from field contaminated soils. Environ. Sci. Technol. 26:532-538 59. Peterson JR, Adams Jr. RS & Cutkomp LK (1971) Soil properties influencing DDT bioactivity. Soil. Sci. Soc. Am. Proc. 35:71-78 60. Pignatello JJ (1989) Sorption dynamics of organic compounds in soils and sediments. In: Sawhney BL, Brown K (Eds) Reactions and Movement of Organic Chemicals in Soils, (pp 45-80). Soil Science Society of America, Madison, WI 61. Pignatello JJ (1990) Slowly reversible sorption of aliphatic halocarbons in soils. I. Formation of residual fractions. Environ. Toxicol. Chem. 9:1107-1115 62. Pignatello JJ (1990) Slowly reversible sorption ofaliphatic halocarbons in soils. II. Mechanistic aspects. Environ. Toxicol. Chem. 9:1117-1126 63. Pignatello JJ & Huang LQ (1991) Sorptive reversibility of atrazine and metolachlor residues in field soil samples. J. Environ. Qual. 20:222-228 64. Pignatello JJ & Xing B (1996) Mechanisms of slow sorption of organic chemicals to natural particles. Environ. Sci. Technol. 30:1-10 65. Pignatello JJ, Ferrandino FJ & Huang LQ (1993) Elution of aged and freshly added herbicides

302 from a soil. Environ. Sci. Technol. 27:1563-1571 66. Rao PSC & Davidson JM (1980) Estimation of pesticide retention and transformation parameters required in nonpopint source pollution models. In: Overcash MR, Davidson JM (Eds) Environmental Impact of Nonpoint Source Pollution, (pp 23-67). Ann Arbor Science Publishers, Ann Arbor, MI 67. Remberger M, Allard A-S &Neilson AH (1986) Biotransformation of chloroguaiacols and chloroveratroles in sediments. Appl. Environ. Microbiol. 51:552-558 68. Rijnaarts HHM, Bachmann A, Jumelet JC & Zehnder AJB (1990) Effect of desorption and intraparticle mass transfer on the aerobic biomineralization of hexachlorocyclohexane in a contaminated calcareous soil. Environ. Sci. Technol. 24:1349-1354 69. Robinson KG, Farmer WS & Novak JT (1990) Availability of sorbed toluene in soils for biodegradation by acclimated bacteria. Wat. Res. 24:345-350 70. Sandoli RL, Ghiorse WC & Madsen EL (1996) Regulation of microbial phenanthrene mineralization in sediment samples by sorbent-sorbate contact time, inocula and gamma irradiation-induced sterilization artifacts. Environ. Toxicol. Chem. 15:1901-1907 71. Scow KM & Alexander M (1992) Effect of diffusion on the kinetics of biodegradation: Experimental results with synthetic aggregates. Soil Sci. Soc. Am. J. 41:340-342 72. Scribner SL, Benzing TR, Sun S & Boyd SA (1992) Desorption and bioavailability of aged simazine residues in soil from a continuous corn field. J. Environ. Qual. 21:115-120 73. Shelton DR & Parkin TB (1991) Effect of moisture on sorption and biodegradation of carbofuran in soil. J. Agric. Food Chem. 39:2063-2068 74. Shimp RJ & Young RL (1988) Availability of organic chemicals for biodegradation in settled bottom sediments. Ectoxicol. Environ. Safety 15:31-45 75. Slater JH, Bull AT & Hardman DJ (1995) Microbial dehalogenation. Biodegradation 6:181-189 76. Steinberg SM, Pignatello JJ & Sawhney BL (1987) Persistence of 1,2-dibromoethane in soils: Entrapment in intraparticle micropores. Environ. Sci. Technol. 21:1201-1208 77. Torrents A, Jayasundera S & Schmidt WJ (1997) Influence of the polarity of organic matter on the sorption of acetamide pesticides. J. Agric. Food Chem. 45:3320-3325 78. Varanasi U, Reichert WL, Stein JE, Brown DW

NAM AND KUKOR & 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 79. Volkering F, Breure AM, Strekenburg A & van Andel JG (1992) Microbial degradation of polycyclic aromatic hydrocarbons: effect of substrate availability on bacterial growth kinetics. Appl. Microbiol. Biotechnol. 40:535-540 80. Weber JB, Best JA & Gonese JU (1993) Bioavailability and bioactivity ofsorbed organic chemicals. In: Luxmore RJ, Peterson GA (Eds) Sorption and Degradation of Pesticides and Organic Chemicals in Soil, (pp 153-196). Soil Science Society of America, Madison, WI 81. Weber, Jr. WJ & Huang W (1996) A distributed reactivity model for sorption by soils and sediments. 4. Intraparticle heterogeneity and phase-distribution relationships under nonequilibrium conditions. Environ. Sci. Technol. 30:881-888 82. Weissenfels WD, Klewer H & LanghoffJ (1992) Adsorption of polycyclic aromatic hydrocarbons (PAHs) by soil particles: influence on biodegradability and biotoxicity. Appl. Microbiol. Biotechnol. 36:689-696 83. Wershaw RL (1989) Application of a membrane model to the sorptive interactions of humic substances. Environ. Health Perspect. 83:191-203 84. White JC & Alexander M (1996) Reduced biodegradabilityofdesorption-resistant fractions of polycyclic aromatic hydrocarbons in soil and aquifer solids. Environ. Toxicol. Chem. 15:1973-1978 85. Wodzinski RS & Coyle JE (1974) Physical state of phenanthrene for utilization by bacteria. Appl. Microbiol. 27:1081-1084 86. Wszolek PC & Alexander M (1979) Effect of desorption rate on the biodegradation of N-alkylamines bound to clay. J. Agric. Food Chem. 27:410-414 87. Wu S & Gschwend PM (1986) Sorption kinetics of hydrophobic organic compounds to natural sediments and soils. Environ. Sci. Technol. 20:717-725 88. Xing B & Pignatello JJ (1996) Time-dependent isotherm shape of organic compounds in soil organic matter: Implications for sorption mechanism. Environ. Toxicol. Chem. 15:1282-1288 89. Xing B, Pignatello JJ & Gigliotti B (1996) Competitive sorption between atrazine and other organic compounds in soils and model sorbents. Environ. Sci. Technol. 30:2432-2440

Chapter 11 BIOTRANSFORMATION OF HALOGENATED PESTICIDES DENNIS D. FOCHT Department of Plant Pathology, University of California, Riverside, CA, USA

1. HISTORICAL ASPECTS OF PESTICIDE USAGE 1.1. Evolution of Modern Agriculture Three scientific discoveries, directly intertwined to each other, led to the increased usage and design of synthetic chemicals prior to the end of the World War II (WWII). The first was the introduction of a remarkable new insecticide DDT (dichlorodiphenyltrichloroethane, or 1,1-bis[p-chlorophenyl]-2,2,2-trichloroethane) in the 1930s. Ironically, this chemical had languished on the shelf since 1874 when it was synthesized as the topic of a doctoral thesis in 1874 by Zeidler (39). It was hailed as the ultimate insecticide because it appeared to have no injurious effects on mammals. The U.S. Army used DDT to combat body lice during WWII by direct application of a 5% mixture in talcum powder; similar mixtures were commonly used with household pets and farmyard animals during this time. The second related event in agriculture occurred in 1940 with the Haber-Bosch synthesis of ammonia from atmospheric nitrogen and hydrogen. This process effected the greatest increase ever achieved in the production of food and fiber. The dramatic shift from crop rotation to continuous mono-cropping created a potential paradise for the development of insect and weed pests. Nevertheless, the production of high crop yields, through use of pesticides and fertilizers, was one of the marvels of agricultural science that went unchallenged for two decades. Increased demand for food and fiber would have diminished were it not for increased life expectancies and population growth (the third event). The establishment of public health standards and the discovery of antibiotics during the first halfofthe 20th century, greatly reduced mortality from common bacterial infections. Increased life expectancy and standard of living also causes demand for more consumer goods. As an industrialized society generates products to meet these demands for consumer goods and services, it generates waste by-products in the manufacture of them. Many of the industrial synthesis procedures use halogenated compounds as coupling reagents and catalysts. This can be seen by the development of novel organochlorine compounds, which reached their highest level in the early 1960s (Figure 11.1). Dehalogenation: Microbial Processes and Environmental Applications, pages 303-322 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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1.2. The Silent Spring DDT and the chlorinated hydrocarbon insecticides were hailed as the elixir of modern agriculture until the publication of the book “Silent Spring” (15). Rachel Carson brought to the attention of the public and the scientific community that the wonders of the green revolution were not without cost. Adverse side effects were observed with unintended targets, particularly birds of prey. Because of low water solubility and high adsorption affinity to organic matter, chlorinated hydrocarbon insecticides became concentrated in sediments that deposit in surrounding streams and rivers from agricultural run-off during rainfall or irrigation. Biomagnification of these insecticides occurred in each successively higher consumer of the food chain, from bacteria, protozoans, insects, and finally to the larger animals at the top. Aerial dissemination and deposition had also contributed to DDT and related insecticides being distributed throughout the global ecosystem, as noted by its accumulation in tissues of polar bears and penguins (61). Pesticide contamination of groundwater soon became another issue in the 1980s. Normally, contaminant levels are below 1 ppm (part per million). Nevertheless, acceptable levels below ppb (part per billion) have been set for groundwater contamination of pesticides that are acutely toxic. Groundwater contamination is a more serious problem with those compounds that are more water soluble (Tables 11.1, 11.2, and 11.3), than those that are confined in the soil matrix near the surface. Nevertheless, immobile compounds (e.g., DDT), and biodegradable ones (e.g., malathion), have been

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commonly found in groundwater (4). This probably occurs as a result of transport through large channels in soil, or between well casings where pesticide application machinery was commonly washed over 20 years ago. The accepted Federal standard in the United States of 5 ppm DDT (and its metabolites) in cropland soil is frequently exceeded in many areas throughout the southwestern part of the country (36).

2. CHEMICAL CONFIGURATION AND BIOTRANSFORMATION 2.1. Microbial Infallibility Throughout most of the 20th century, it had been assumed a priori that all carbonaceous compounds would be processed through the geochemical carbon cycle.

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Gale (25) went so far to state that there were microorganisms in nature somewhere that could degrade any conceivable molecule that might be synthesized by organic chemists. During this long time, the idea that soil, through the action of its microbiota, could serve as a great cleansing agent, was never challenged despite the continued synthesis and usage of novel organic compounds. Xenobiotic (derived from the Greek xenos, meaning foreign) compounds are those that are foreign to biology. In one of the earliest studies on the fate of pesticides, Martin (48) reported that neither 2,4-D (2,4-dichlorophenoxy acetic acid) nor DDT had any

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injurious effects on the soil microbiota at concentrations two orders ofmagnitude higher than what was used in agriculture (1 ppt). However, no consideration was given to whether or not these compounds decomposed in the environment. The earliest work on biodegradation of synthetic organic compounds was with the herbicide 2,4-D, (5, 68). Alexander (1) reviewed the state of the art regarding biodegradation, which consisted mostly of hydrocarbon metabolism and these few studies on 2,4-D, and he recognized

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that studies were needed to determine if pesticides were readily degraded in soil. Alexander (2) also questioned Gale's concept of microbial infallibility, and asked whether or not there may be some compounds that are inherently recalcitrant to microbial attack. He suggested that it might be expecting too much of microorganisms, which evolved their metabolic capabilities over a billion years of evolutionary development, to evolve enzymes able to metabolize this new array of synthetic substrates, in a span of merely decades.

2.1. Genomics and Catabolic Enzymes Among all organochlorine pesticides that have been used since WWII, there is a unanimous consensus that the chlorinated hydrocarbon insecticides are the most resistant to microbial decomposition. Nevertheless, microorganisms, plants and animals synthesize many natural organochlorine compounds (see Chapter 1) (18). Thus, the concept that synthetic compounds are inherently unfamiliar to microorganisms and their long evolutionary history is a questionable one. Moreover, this concept is inconsistent with the fundamental basis of cometabolism, a phenomenon whereby enzymes, which serve as catalysts for natural products, fortuitously metabolize many synthetic compounds (or other natural compounds). For example, dehydrogenases that reduce PCB (polychlorinated biphenyl) diols to the corresponding catechols have very broad substrate specificity and show similar sequence homologies to oxidoreductases of polyols, sterols, and antibiotics (6). The addition of biphenyl to soil has been shown to greatly affect aerobic biodegradation of PCBs by providing a growth substrate for indigenous biphenylutilizing bacteria (13, 23, 30). However, biphenyl is a carcinogen and a listed priority pollutant by the United States Environmental Protection Agency (USEPA), which makes its use limited. Moreover, it is not very effective for stimulating transformation of the more highly chlorinated PCB congeners that comprise Aroclor 1254 (24). Flavinoids (21) and terpenes (27, 34) are natural substrates that have been shown to enhance biodegradation of Aroclor 1242. It has been suggested that terpenes, rather than biphenyls, might be the natural substrates for which the enzymes of the biphenyl pathway function (34). As genomic databases expand, it should be possible to determine which natural substrates are involved in promoting the transcription and synthesis of enzymes that act upon a specific xenobiotic chemical. Contaminated soil could then be treated by adding an innocuous substrate that enhances growth of microorganisms involved in biodegradation of the target chemical.

3. BIOAVAILABILITY 3.1. Chlorine Substituents: The Major Factor in Recalcitrance? Although it is generally accepted that chlorinated compounds are less biodegradable than their non-chlorinated analogs, chlorine substitution per se can not alone account for persistence. For example, pentachlorophenol (Table 11.1) has the highest extent of chlorine substitution per carbon atom than any of the organochlorine insecticides shown in Table 11.2, yet it is utilized as a growth substrate by different species ofbacteria (16,

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29, 59). A similar comparison of aryl chlorine substitution reveals that 2,4-D contains two chlorines per aromatic ring, while DDT contains only one (Tables 11.1 and 11.2). While bacterial cultures able to utilize 2,4-D as a sole carbon source are relatively common, no isolate has been rigorously proven to do the same with DDT or any of the insecticides shown in Table 11.2. DDT and the chlorinated hydrocarbon insecticides persist for decades in soil, while 2,4-D is gone within the same growing season of its application. Clearly, extent of aryl chlorine substitution can not be the sole factor that effects the persistence of chlorinated hydrocarbon insecticides in the environment. The common features of PAHs (polyaromatic hydrocarbons), which have no chlorine substituents, and chlorinated hydrocarbon insecticides, are their persistence in the environment, their low water solubility, and their sorption to soil (see Chapter 10). The solubility of these compounds is directly proportional to their rates of biotransformation. This has practical implication for laboratory studies as well, in that their respective water solubilities determine the ease with which enrichment cultures that utilize them as sole carbon sources can be obtained. The water solubilities of naphthalene, biphenyl, phenanthrene, pyrene, and chrysene are 31, 7, 1.1, 0.13, and 0.002 mg/L, respectively. Naphthalene- and biphenyl-utilizers are readily obtained from soil, and in some cases may utilize the same enzymes for metabolism (43). Although bacteria that utilize phenanthrene (3-membered ring), pyrene (4-membered ring), and chrysene as growth substrates have been reported (33, 62), they are less readily isolated than naphthalene/biphenyl-degraders. Similarly, water soluble herbicides and fungicides, such as the chlorophenols and chlorobenzoates are far more biodegradable than the nonsoluble chlorinated insecticides (Tables 11.1 – 11.3). The problem of bioavailability of PCBs led to a novel idea advanced by Lajoie et al. (44), in the genetic construction ofa PCB-degrading bacterium containing a cloned gene for surfactant production. Soil inoculated with this bacterium had markedly lower concentrations of PCBs than controls (without inoculation) or inoculated with the wild type strain, which did not contain the gene encoding surfactant utilization. The question of how microorganisms metabolize insoluble compounds has not been resolved. In some cases, common soil polymers such as lignin and cellulose are hydrolyzed by extracellular enzymes secreted by soil microorganisms, breaking the larger molecules into smaller, soluble units that can be transported into microbial cells and further metabolized.

3.2. Aged Residues in Soil Although surfactants may enhance biodegradation ofinsoluble compounds added to soil in experimental studies, their effectiveness in treating aged residues in the field is less successful. A certain proportion of aged pesticide is entrapped in soil. Steinberg et al. (63) extracted soil periodically over 200 days, and continued to recover progressively diminishing quantities of EDB (ethylene dibromide) that had been a soil contaminant many years prior to analysis. They concluded that equilibrium between "free" and sorbed or bound contaminant existed in soil, and that extractions shifted the equilibrium continuously in the direction of the free state. The two primary mechanisms accounting for slow release ofchemicals from soil are sorption to, and diffusion into organic matter, and entrapment in the clay mineral pores. More detail on these processes can be found in Chapter 10.

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The effect of entrapment can be seen in Figure 11.2, providing a composite illustration (3) of several field studies (46, 54, 55). In all three cases, there is an initial period of relatively rapid biotransformation, followed by a plateau, in which further disappearance is minimal. Note also that the fraction of "bound" or sorbed DDT is proportionally greater with higher concentrations.

4. SPATIAL VARIABILITY Laboratory investigations designed to measure rates of pesticide degradation characteristically involve the addition (spiking) of known quantities to soil. Although this facilitates sampling and analysis, it is difficult to extrapolate laboratory-derived data to the field, where spatial variability is the rule, rather than the exception. Few soil parameters can be found which follow a normal arithmetic distribution, and most tend to be log-normally distributed. An example of this variability is shown with soil concentrations of dieldrin from a golf course (Figure 11.3), in which the pesticide was discontinued more than 25 years prior to sampling. The consequence of a log normal distribution means that three sample replicates, as used with normal distributions, are unreliable in making valid statistical comparisons between different treatments. Many of the discrepancies reported for rates of pesticide degradation in the field are a result of spatial variability. Aside from the statistical difficulties in assessing remediation, spatial variability of

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concentrations may influence the rates of degradation due to kinetic effects. Uniform degradation rates in the field can not be assumed to occur throughout the soil matrix. For example, "hot spots" (areas of high concentration) of DDT contamination are still found in agricultural soils, throughout the San Joaquin Valley of California, more than 25 years after discontinuation oftheir use. These hot spots are found near sites where mixing and washing of spraying equipment occurred. Such areas will continue to have much higher levels of persistent pesticides (sometimes exceeding 100 mg/kg), than what is normally found in crop land soils (less than 5 mg/kg) (36) throughout the southwestern United States.

5. PERSISTENT METABOLITES 5.1. Biodegradation vs. Biotransformation Because pesticide concentrations are regulated by governmental standards, it has been customary and convenient to measure biodegradation by disappearance. In this sense, the word biodegradation is close to its etymological derivation (Latin degradare, meaning to "reduce in rank"). However, disappearance of a compound raises the question of what it disappears to, and whether or not the products that are formed are “reduced in rank”, to compounds less toxic than the original material. Thus, the term

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"biotransformation" is more appropriate in referring to a limited number ofone or more reactions that are carried out by microorganisms. A more environmentally conscious definition of biodegradation is the ultimate biological destruction to innocuous products. Mineralization, the complete destruction of a compound to its inorganic mineral forms, is one way of producing innocuous

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products. For example, mineralization of DDT would produce carbon dioxide, water, and chloride. However, complete mineralization of the most biodegradable compounds (e.g., glucose) never occurs because some of it is incorporated into microbial biomass, which subsequently dies and becomes part of the amorphous soil organic matter, referred to as humus. In the environment, the metabolites may be toxic, or moderately recalcitrant to other soil microorganisms, and persist considerably longer than the compound from which they originated. In other cases, they may become polymerized by the action of peroxidases and free radical formation, and become indistinguishable as part of the soil humus. There is no general rule that the accumulation of degradation products is environmentally problematic or innocuous, because each compound must be considered on a case by case basis.

5.2. Total DDT Residues in the Environment The most persistent and environmentally problematic biotransformation product in the environment is DDE (dichlorodiphenyldichloroethylene, or 1,1 -bis [p-chlorophenyl] 2,2,-dichloroethene), which results from the dehydrodehalogenation of DDT by all life forms, including mammals (Figure 11.4). Reference to DDT concentrations in the environment now means total DDT (TDDT), which includes DDE and another major anaerobic breakdown product DDD (dichlorodiphenyldichloroethane, or 1,1-bis[pchlorophenyl]-2,2,-dichloroethane). DDE accounts for more than 70-90% of pesticide residues in soil that remain 25 years after DDT was banned in by the United States government. Soils in the southwestern United States have characteristically higher ratios of DDE/DDT, as a result of slow conversion to DDE over the years. Data obtained from field plots in 1969, 1973, and 1989 in the southwestern United States indicate approximate half-lives for DDT and DDE of 10 to 20 years. A field having the highest original concentration of DDT decreased from 60 to 42 to 22 ppm, while DDE actually increased, from 3.3 to 4.5 to 7.0 ppm through the respective years noted above (47). Ratios of DDE to DDT have been used occasionally to determine if DDT has been used illegally after its ban in 1972. DDE has diametrically opposite effects on insects and birds. The production of DDE is a detoxification mechanism by which insects develop resistance to DDT, while in birds, DDE is a potent inducer of steroid hydroxylase, which disrupts calcium metabolism and leads to thin egg shell development (50). DDE is also a potent androgen receptor in mammals (41). Lately, it has been recognized that, although DDT concentrations in wild birds in the southwestern United States have been decreasing, the levels of DDE have been increasing (36). The natural transformation of DDT to DDE in soil may actually result in greater risks to birds that nest in transient winter ponds, created from agricultural run-off, during the rainy season in California (41).

5.3. Effects of Temperature on Biotransformation of DDT, DDE, and DDD Temperature affects the biotransformation rate of DDT and other chlorinated hydrocarbon insecticides the same as it does with other substrates. Soil organic matter is more abundant in humid, cool climates than in dry, hot climates because of greater plant productivity and slower microbial breakdown of humus in the former. Therefore,

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the 30-year persistence of DDT in forest soils of Maine (20), compared with a 39% disappearance over a four-week period in agricultural soils of Colorado (28), is consistent with the basic principles of biological rate processes. Many of the developing countries in the tropics still use DDT on a large scale. Although this has raised objections by environmentally conscious people, TDDT does not present the long- term persistence problem common to temperate climates. Lalah et al. (45) found that total residues of disappeared rapidly to 11% of the added amount in less than 6 months, with 8.5% being mineralized, in field studies conducted in Kenya. DDE was the primary transient intermediate, and very little DDD was produced. They concluded that DDT is not a persistent pesticide that is likely to accumulate in tropical soils. This conclusion is supported by dozens offield studies from the tropics, published in this same special volume (42). It remains unclear, however, if losses of TDDT are due to more rapid biodegradation in tropical soils as a result of higher mean temperatures, or to greater vapor losses and global transport to the Northern Hemisphere.

5.4. Bacterial Metabolism of DDT, DDE, and DDD Until recently, the biodegradation of DDT was thought to occur through a series of coupled, anaerobic-aerobic biotransformations (22, 66, 67), as advanced by Pfaender and Alexander (56). They suggested that these diametrically opposed, and often spatially separated, redox conditions might account for the persistence of DDT in the environment. For example, anaerobic dehalogenation of PCBs, which were not degraded aerobically, was later demonstrated by a number of researchers (12, 57) (see Chapter 17). These studies suggested that recalcitrant chlorinated hydrocarbons might be slowly decomposed innature, where aerobic and anaerobic zones such as sediments are in close proximity. A very recent study (58) has shown that DDE was reductively dehalogenated to DDMU (1-chloro-2,2-bis(p-chlorophenyl)ethene) in sediments off the Palos Verde shelf near Los Angeles harbor (Figure 11.4). Prior to this study, the fate of DDE in the environment was unknown. Nadeau et al. (52, 53) made the first definitive observation that DDT was metabolized aerobically to p-chlorobenzoate by a biphenyl-utilizing bacterium, Ralstonia eutropha A5 (formerly Alcaligenes eutrophus A5). This was followed in studies by Hay and Focht, who showed that DDE and DDD were also oxidized aerobically through the biphenyl pathway to p-chlorobenzoate, respectively by Pseudomonas acidovorans M3GY (31), and Ralstonia eutrophua A5 (32). Some alternate variations of the DDE and DDD aerobic pathways were noted (Figure 11.4), particularly the formation of p-chloroacetophenone, a secondary metabolite common to the biphenyl pathway (9, 10). The arrows from anaerobic dehalogenation products in Figure 11.4 indicate that these compounds undergo metabolism through the same pathway shown for DDE.

5.5. Coupled Anaerobic-Aerobic Processes: From Fungicide to Herbicide For many years, mercurial-based fungicides had been used in Japan for the treatment of rice blast disease. Shortly after the discovery of chronic and severe mercurial poisoning in residents near Miyamato Bay in 1956, all mercurial fungicides were phased

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out of usage. The most popular replacement was pentachlorobenzyl alcohol (PCBA), which was used extensively throughout the 1960s. Within a decade, phytotoxic effects became noticed in lowland (continuously flooded) rice paddies. The cause of this problem was shown by Ishida (38) to be caused by microbial conversion of PCBA to the herbicide 2,3,6-trichlorobenzoate. Ishida correctly postulated that PCBA was oxidized by aerobic microorganisms in the oxygen rich sediments at the surface, where it was recalcitrant to further aerobic attack. It then diffused to the anaerobic zone below, where it was reductively dehalogenated to 2,3,6-trichlorobenzoic acid (2,3,6-TCB), a herbicide (Figure 11.5). As this problem did not occur in upland paddies, which are subjected to drying cycles when the land lies fallow, draining the fields periodically solved the lowland rice paddy problem. The mechanism postulated by Ishida was later demonstrated by Gerritse and Gottschal (26), who established a reactor system consisting of anaerobic and aerobic consortia of bacteria, which effected complete mineralization of 2,3,6-TCB. The anaerobes grew within the anoxic volume of vermiculite particle and dehalogenated 2,3,6-TCB to 2,5-DCB. 2,5-DCB diffused to the aerobic zone outside, where it was utilized as a growth substrate by Pseudomonas aeruginosa JB2 (35).

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5.5. Formation of Tetrachloroazobenzene Dimers from Chloroaniline Herbicides The first example of a stable xenobiotic intermediate was observed within the class of acyl-aniline and phenyl urea herbicides (8). It can be readily noted from the structure of these compounds (Figure 11.6), that the amide and ureide bonds should be readily hydrolyzed by microorganisms in soil. However, the subsequent formation of 3,4dichloroaniline gave rise to a compound that was not readily metabolized. Consequently, this product accumulated and was oxidized by soil microorganisms to aryl hydroxylamine and aryl nitroso compounds, both of which are highly reactive chemically, and dimerize to give the azo product shown. The half-life of this compound has been estimated to be about 6 months to a year in contrast to a half-life of one or two weeks with the herbicide (7).

5.6. Formation and Biodegradation of Chlorinated Humus By definition of the term “biodegradation”, substrates do not have to necessarily be completely degraded, or mineralized; conversion of incomplete biodegradation products to soil humus represents a natural, stabilized, and innocuous fate. TNT (2,3,6trinitrotoluene) is an example of a compound that is not mineralized, but is polymerized into biomass and organic polymers (14). The process occurs by reduction of the aryl nitro groups to amino substituents, which make the aniline ring susceptible to free radical formation by the peroxidases and laccases of soil microorganisms (60). Similar reactions occur with 2,4-dichlorophenol, the initial breakdown product of 2,4-D (2,4dichlorophenoxy acetic acid), which, if not further degraded, may be incorporated and

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stabilized into the soil humus (11, 19). Free radical addition can also involve polymerization of chloroaromatic molecules with phenolic acids from lignin as well as other biological constituents, namely polysaccharides and proteins, as shown in the schematic, Figure 11.7. The question of whether or not chlorinated humus is environmentally objective needs to be put in the perspective of whether or not these compounds are of foreign origin. As already mentioned here and in Chapter 1, a vast array of natural organochlorine compounds including antibiotics, exist in the environment. Perhaps most revealing of

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all is the observation that chlorinated phenols can be synthesized by common soil fungi at concentrations of 75 mg/kg wood (17, 18). The following statement of the cited authors should help to dispel the negative mystique about xenobiotic compounds: "The widespread ability among common fungi to produce large amounts of chlorinated aromatic compounds in the environment makes us conclude that these kinds of compounds can no longer be considered to originate mainly from anthropogenic sources." Given that chlorinated humus is synthesized naturally, in the absence of xenobiotic compounds, and that their metabolites are benign, is chlorinated humus more stable than non-chlorinated humus? This question was addressed using model polymers synthesized from catechol, 4-chlorocatechol, and 4,5-dichlorocatechol (64). The model polymers were synthesized by fungal peroxidases in a manner identical to the process as it occurs in nature. evolution was monitored periodically from four different soils. Only the cumulative results at the end of the experiment are shown in Figure 11.8. In all four soils, the rate of decomposition was greatest with the dichlorinated model polymer (4,5-dichlorocatechol), and least with the non-chlorinated polymer (catechol). However, the authors found no significant differences between mineralization rates of the three catechol monomers. That more chlorinated humus is mineralized to a greater extent than non-chlorinated humus may appear contrary to the central dogma, namely that biodegradation is inversely proportional to the extent of chlorine substitution. However, the reasons for stability ofhumus in soil are more complex. Infra red spectra ofhumus characteristically show no strong C-H scissoring vibrations in the 690-890 cm region, always typical of aromatic monomers, because of extensive cross-linkage and substitution ofthe aromatic units. Thus, if a chlorine is not removed during free radical addition (Figure 11.7), then that carbon atom will not be available for cross-linking to impart greater structural stability. Thus, the greater rate of decomposition with chlorinated humic polymers (Figure 11.8) is most likely due to fewer aryl ether and C-C linkages.

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6. HISTORIC PERSPECTIVE: SOILS AND ACCELERATED BIODEGRADATION The flip side to the enhancement of biodegradation of persistent chemicals is the awareness that many of the carbamate insecticides and thiocarbamate herbicides, long used in agriculture, eventually were observed to rapidly disappear in the mid 1980s (40). This was initially surprising, as these pesticides were usually effective through the early part ofthe growing season, and generally required a single application. It is tempting to speculate that the continued used ofa single herbicide in the field for many years is akin to a laboratory enrichment culture that microbiologists use for selecting organisms able to utilize a specific compound as a sole carbon source. Although the scales ofsize differ, the principle may be the same in that, over time and exposure, selective populations of microorganisms are selected for, which metabolize, i.e., biodegrade, the target compound. It must be noted, however, that inmany cases, enrichment cultures fail, most notably with attempts to isolate DDT or PCB utilizers, because two or more organisms are involved in the catabolic pathway, and these may require very divergent, e.g., aerobic vs. anaerobic, conditions for growth and activity. The isolation of bacteria able to utilize multi-chlorinated aromatic compounds as growth substrates in the last decade has been astounding. In almost all cases, these cultures may take a year or more to isolate, and invariably come from polluted (and therefore also selective) environments. It is difficult to rigorously prove the source of isolation, or that the prolonged incubation time was the result of a rare event (mutation, genetic exchange). Yet, this occurrence has a parallel in the widespread dissemination of antibiotic-resistant bacteria, as first documented by Watanabe some 20 years after the use of antibiotics in medicine (65). Though it is true that microorganisms represent the evolution of a billion or more years, it does not follow that their response to immediate and constant selection pressure is on a geological time scale. To the contrary, the evolution of novel catabolic genotypes is all around us. REFERENCES 1. Alexander M (1961) Introduction to Soil Microbiology. John Wiley and Sons, New York 2. Alexander M (1965) Biodegradation: Problems of molecular recalcitrance and microbial fallability. Adv. Appl. Microbiol. 7:35-76 3. Alexander M (1999) Biodegradation and Bioremediation. Academic Press, San Diego 4. Anonymous (1986) Ground water quality in California: A review of scientific and technical issues. ESE Report No. 86-61. UCLA Environmental Science and Engineering Program 5. Audus LJ (1952) The decomposition of 2,4dichlorophenoxyacetic acid and 2-methyl-4chlorophenoxyacetic acid in the soil. J. Sci. Food Agr. 3:268-274 6. Baker ME (1990) Sequence similarity between Pseudomonas dihydrodioldehydrogenase, part of the gene cluster that metabolizes PCBs, and

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S-C (2001) Mechanisms involving aerobic Microbiol. 53:1103-1112 biodegradation ofPCBs in nature. In: Agathos S 11. Bollag J-M & Liu S-Y (1985) Copolymerization & Reineke W (Eds) Focus on Biotechnology Vol of halogenated phenols and syringic acid. 3A. Biotechnology for the Environment: Pesticide Biochemistry and Physiology 23:261Strategy and Fundamentals. Kluwer Academic 272 Publishers, BV Dordrecht, The Netherlands. 12. Brown JF Jr, Wagner RE, Bedard DL, Brennan MJ, Carnahan JC, May RJ & Tofflemire TJ 25. Gale EF (1952) The Chemical Activities of Bacteria. Academic Press, London (1984) PCB transformations in upper Hudson 26. Gerritse J & Gottschal JC (1992) Mineralization sediments. Northeast. Environ. Sci. 3:167-179 of the herbicide 2,3,6-trichlorobenzoic acid by a 13. Brunner W, Sutherland FH & Focht DD (1985) co-culture of anaerobic and aerobic bacteria. Enhanced biodegradation of polychlorinated FEMS Microbiol. Ecol. 101:89-98 biphenyls in soil by analog enrichment and bacterial inoculation. J. Environ. Qual. 14:324- 27. Gilbert ES & Crowley DE (1997) Plant compounds that induce polychlorinated biphenyl 328 biodegradation by Arthrobacter sp. strain B1B. 14. Carpenter DF, McCormick NG, Cornell JH & Appl. Environ. Microbiol. 63:1933-1938 Kaplan AM (1978) Microbial transformation of 2,4,6-trinitrotoluene in an activated 28. Guenzi WD & Beard WE (1967) Anaerobic biodegradation of DDT to DDD in soil. Science sludge system. Appl. Environ. Microbiol. 156:1116-1117 35:949-954 15. Carson R(l962) Silent Spring. Houghton Miflin 29. Häggblom MM, Nohynek LJ & SalkinojaSalonen MS (1988) Degradation and OCo., Boston methylation ofchlorinated phenolic compounds 16. Crawford RL & Mohn WW (1985) by Rhodococcus and Mycobacterium strains. Microbiological removal of pentachlorophenol Appl. Environ. Microbiol. 54:3043-3052 from soil using a Flavobacterium. Enzyme 30. Harkness MR,McDermott JB,Abramowicz DA, Microb. Technol. 7:617-620 17. De Jong E, Cazemie AE, Field JA & De Bont Salvo JJ, Flanagan WP, Stephens ML, Mondello FJ, May RJ, Lobos JH, Carroll KM, M.J. B, JAM (1994) Physiological role of chlorinated Bracco AA, Fish KM, Warner GL, Wilson PR, aryl alcohols biosynthesized de novo by the Dietrich DK, Lin DT, Morgan CB & Gately WL white rot fungus Bjerkandera sp. strain BOS55. (1993) In-situ stimulation of aerobic PCB Appl. Environ. Microbiol. 60:271-277 biodegradation in Hudson River sediments. 18. De Jong E, Field JA, Spinnler HE, Wijnberg Science 259:503-507 JBPA & De Bont JAM (1994) Significant biogenesis of chlorinated aromatics by fungi in 31. Hay AG & Focht DD (1998) Cometabolism of 1,1-dichloro-2,2-bis(4-chlorophenyl)ethyleneby natural environments. Appl. Environ. Microbiol. 60:264-270 Pseudomonas acidovorans M3GY grown on biphenyl. Appl. Environ. Microbiol. 64:214119. Dec J & Bollag J-M (1994) Dehalogenation of chlorinated phenols during binding to humus. 2146 In: Anderson TA & Coats JR (Eds) 32. Hay AG & Focht DD (2000) Transformation of Bioremediation through Rhizosphere 1,1 -dichloro-2,2-(4-chlorophenyl)ethane (DDD) Technology (pp 102-111) American Chemical by Ralstonia eutropha A5. FEMS Microbiol. Society, Washington, DC Ecol. 31:249-253 20. Dimond JB, Belyea GY, Kadunce RE, Getchell 33. Heitkamp MA & Cerniglia CE (1989) AS & Blease JA (1970) DDT residues in robins Polycyclic aromatic hydrocarbon degradation by and earthworms associated with contaminated a Mycobacterium sp. in microcosms containing forest soils. Can. Entomol. 102:1122-1130 sediment and water from a pristine ecosystem. 21. Donnelly PK, Hegde RS & Fletcher JS (1994) Appl. Environ. Microbiol. 55:1968-1973 Growth of PCB-degrading bacteria on 34. Hernandez. BS, Koh S-C, Chial M & Focht DD compounds from photosynthetic plants. (1997) Terpene-utilizing isolates and their Chemosphere 28:981-988 relevance to enhanced biotransformation of 22. Focht DD & Alexander M (1971) Aerobic polychlorinated b i p h e n y l s i n soil. cometabolism of DDT analogues by Biodegradation 8:153-158 Hydrogenomonas sp. J. Agric. Food Chem. 35. Hickey WJ & Focht DD (1990) Degradation of 19:20-22 mono-, di-, and trihalogenated benzoic acids by 23. Focht DD & Brunner W (1985) Kinetics of Pseudomonas aeruginosa JB2. Appl. Environ. biphenyl and polychlorinated biphenyl Microbiol. 56:3842-3850 metabolism in soil. Appl. Environ. Microbiol. 36. Hitch RK & Day HR (1992) Unusual 50:1058-1063 persistence of DDT in some western USA soils. 24. Focht DD, McCullar MV, Searles DB & Koh Bull. Environ. Contam. Toxicol. 48:259-264

BIOTRANSFORMATION OF HALOGENATED PESTICIDES 37. Hutzinger O & Veerkamp W (1981) Xenobiotic chemicals with pollution potential. In: Leisinger T, Cook AM, Hütter R& Nüsch J (Eds) Microbial Degradation of Xenobiotic and Recalcitrant Compounds (pp 3-45) Academic Press, New York 38. Ishida M (1972) Phytotoxic metabolites of pentachlorobenzyl alcohol. In: Matsumura F, Bousch GM & Misato T (Eds) Environmental Toxicology of Pesticides (pp 281-299) Academic Press, New York 39. Jensen S (1972) The PCB story. Ambio 1:123131 40. Kaufman DD, Katen Y, Edwards DS & Jordon EJ (1985) Microbial adaptation and metabolism of pesticides. In: Hilton JL (Ed) Agricultural Chemicals of the Future (pp 437-451) BARC Symposium 8. Rowman and Allanheld, Totawa, NJ 41. Kelce WR, Stone CR, Laws SC, Gray LE, Kemppainen JA & Wilson EM (1995) Persistent DDT metabolite p,p'-DDE is a potent androgen receptor antagonist. Nature 375:581-585 42. Khan SH (1994) Special Issue on DDT in the Tropics: Appraisal of overall programme accomplishments. J. Environ. Sci. Health Part B-Pesticides Food Contamin. Agricult. Wastes 29:205-226 43. Kuhm AE, Stolz A & Knackmuss H-J (1991) Metabolism of naphthalene by the biphenyldegrading bacterium Pseudomonas paucimobilis Q1. Biodegradation 2:115-120 44. Lajoie CA, Layton AC & Sayler GS (1994) Cometabolic oxidation of polychlorinated biphenyls in soil with a surfactant-based field application vector. Appl. Environ. Microbiol. 60:2826-2833 45. Lalah JO, Acholla FV & Wandiga SO (1994) Fate of in Kenyan tropical soils. J. Environ. Sci. Health Part B-Pesticides Food Contam. Agricult. Wastes 29:57-64 46. Lichtenstein EP, DePew LJ & Quirk AV (1960) Persistence of DDT, aldrin, and lindane in some midwestern soils. J. Econ. Entomol. 53:136-142 47. Martijn A, Bakker H & Schreuder RH (1993) Soil persistence of DDT, dieldrin, and lindane over a long period. Bull. Environ. Contam. Toxicol. 51:178-184 48. Martin JP (1946) The hormone weed killer 2,4D. Calif. Citrograph 31:248, 264-265 49. Martin JP & Focht DD (1977) Biological properties of soil. In: Elliot LF & Stevenson FJ (Eds) Soils for Management of Organic Wastes and Waste Waters (pp 115-169) Soil Science Society of America, Madison, WI 50. Metcalf RL & Fukuto TR (1968) The comparative toxicity of DDT and analogues to susceptible and resistant houseflies and

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mosquitos. Bull. Wld. Hlth. Org. 38:633-647 51. Montgomery JH (1993) Agrochemicals Desk Reference: Environmental Data. Lewis Publishers, Boca Raton 52. Nadeau LJ, Menn FM, Breen A & Sayler GS (1994) Aerobic degradation of 1,1,1,-trichloro2,2-bis(4-chlorophenyl)ethane (DDT) by Alcaligenes eutrophus A5. Appl. Environ. Microbiol. 60:51-55 53. Nadeau LJ, Sayler GS & Spain JC (1998) Oxidation of l,l, l-trichloro-2,2-bis(4chlorophenyl)ethane (DDT) by Alcaligenes eutrophus A5. Arch. Microbiol. 171:44-49 54. Nash RG & Woolson EA (1967) Persistence of chlorinated hydrocarbon insecticides in soil. Science 157:924-927 55. Onsager JA, Rusk HW & Butler LI (1970) Residues of aldrin, dieldrin, chlordane, and DDT in soil and sugarbeets. J, Econ. Entomol. 63:1143-1146 56. Pfaender FK & Alexander M (1972) Extensive microbial degradation ofDDT in vitro and DDT metabolism by natural communities. J. Agric. Food Chem. 20:842-846 57. Quensen III, JF, Tiedje JM & Boyd SA (1988) Reductive dechlorination of polychlorinated biphenyls by anaerobic microorganisms from sediments. Science 242:752-754 58. Quensen III, JF, Tiedje JM, Jain MK & Mueller SA (2001) Factors controlling the rate of DDE dechlorination to DDMU in Palos Verdes margin sediments under anaerobic conditions. Environ. Sci. Technol. 35:286-291 59. Radehaus PM & Schmidt SK (1992) Characterization of a novel Pseudomonas sp. that mineralizes high concentrations of pentachlorophenol. Appl. Environ. Microbiol. 58:2879-2885 60. Rieger P-G & Knackmuss H-J (1995) Basic knowledge and perspectives on biodegradation of 2,4,6-trinitrotoluene and related aromatic compounds in contaminated soil. In: Spain JC (Ed), Biodegradation of Nitroaromatic Compounds (pp 1 -18) Plenum Press, New York 61. Riseborough RW, Reiche P, Peakall DB, Hersman SG & Kirven MN (1968) Polychlorinated biphenyls in the global ecosystem. Nature 220:1098-1102 62. Sanseverino J, Applegate BM, King JMH & Sayler GS (1993) Plasmid-mediated mineralization of naphthalene, phenanthrene, and anthracene. Appl. Environ. Microbiol. 59:1931-1937 63. Steinberg SM, Pignatello JJ & Sawhney BL (1987) Persistence of 1,2-dibromoethane in soils: Entrapment in intraparticle micropores. Environ. Sci. Technol. 21:1201-1208 64. Stott DE, Martin JP, Focht DD & Haider K

322 (1982) Biodegradation, stabilization in humus, and incorporation into soil biomass of 2,4-D and chlorocatechol carbons. Soil Sci. Soc. Am. J. 47:66-70 65. Watanabe T (1963) Infective heredity of multiple drug resistance in bacteria. Bacteriol. Rev. 27:87-115 66. Wedemeyer G (1967) Biodegradation of dichlorodiphenyltrichloroethane:Intermediates in dichlorodiphenylacetic acid metabolism by

FOCHT Aerobacter aerogenes. Appl. Environ. Microbiol. 15:1494-1495 67. Wedemeyer G (1967) Dechlorination of 1,1,1trichloro-2,2-bis(p-chlorophenyl)ethane by Aerobacter aerogenes. Appl. Environ. Microbiol. 15:569-574 68. Whiteside JS & Alexander M (1960) Measurement of microbiological effects of herbicides. Weeds 8:204-213

Chapter 12 BIODEGRADATION OF ATMOSPHERIC HALOCARBONS RONALD S. OREMLAND U.S. Geological Survey, Menlo Park, CA, USA

1. INTRODUCTION Two major environmental challenges for the twenty-first century concern the earth’s atmosphere: global warming of the troposphere, and the integrity of the stratospheric ozone layer. The presence of volatile, long-lived halocarbons in the atmosphere has contributed significantly to both problems. For this reason, international agreements have been forged (e.g., the Montreal Protocol) in order to eliminate or severely constrain anthropogenic releases ofselected volatile halocarbons. However, as we shall see in this chapter, some of these halocarbons have biogenic as well as anthropogenic sources, a facet that complicates our understanding of their global budgets. Warming of the troposphere drives the phenomenon of global climate change. Certain volatile halocarbons in the atmosphere are “radiatively-active”, in that they absorb infrared energy released from the earth’s surface that was originally derived from the warming effect of incident sunlight. Thus, certain halocarbons, in addition to gases like and contribute to the heat capacity of the troposphere. An index of the extent to which an individual radiative trace gas contributes to this phenomenon is termed its global warming potential, or GWP. All GWP values are set relative to which has a GWP of 1.0. Similarly, the contribution a gas makes to the degradation of the stratospheric ozone layer is termed its ozone depletion potential, or ODP, set relative to CFC-11 that has an ODP of 1.0. The calculation of GWP and ODP for a given halocarbon involves a number of factors. These include its ability to absorb infrared energy of a given wavelength (“IR cross section”), how many and what type of halogens are attached to it, its reactivity with atmospheric oxidants, its susceptibility to photolysis, and its global budget with respect to sources and sinks. It is in this last term, namely global sinks, that microorganisms make their contribution by carrying out degradation reactions. These degradation reactions can occur in proximity to sites of biogenic formation of the gas in question, thereby intercepting and diminishing release to the atmosphere of the volatile halocarbon. In addition, some degradation reactions occur at the ambient concentrations that these gases attain when they enter soils, sediments, and natural waters from the atmosphere. In some cases, (e.g., methyl Dehalogenation: Microbial Processes and Environmental Applications, pages 323-345 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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bromide), these reactions are of sufficient magnitude to affect the global biogeochemical budgets of the compound.

1.1. Global Atmospheric Considerations A simple schematic of the global atmospheric cycles for halocarbon trace gases is given in Figure 12.1. The concentration of a trace gas in the troposphere is termed its “mixing ratio,” or mole fraction (in parts per billion or parts per trillion by volume), that in effect represents a quasi steady-state balance between its global sources and sinks. The mixing ratio for a given trace gas can increase, decrease, or fluctuate with time, depending upon temporal changes between the relative strengths of sources and sinks. The time scale component varies quickly on a local basis, seasonally on a regional basis, over decades in the case of atmospheric trends, and even much longer time scales years) for regional or global climatic changes. Very long-term global records of trends in tropospheric trace gas concentrations can be determined by extraction of gases frozen in ice cores retrieved from polar glaciers. At any given instant in time, mixing ratios may vary spatially, either narrowly on a local basis, or broadly with latitude. In the latter case, concentration disparities between the northern and southern hemispheres are commonly observed for anthropogenic trace gases, a factor caused by the larger land mass and greater human population of the northern vs. southern hemispheres. Volatile halocarbons enter the troposphere from natural and/or anthropogenic sources. Their time of residence within the troposphere is a critical factor in determining ODP and GWP. Very reactive gases, such as iodomethane, have values of measured in days or even hours,

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while non-reactive gases like chlorofluorocarbons (CFCs) have as long as centuries in duration. Some examples of the for selected tropospheric trace gases are given in Table 12.1. The major sink for reactive trace gases is oxidation with hyroxyl radicals (·OH), the cleansing agent of the troposphere. Hydroxyl radicals are powerful oxidants that arise from the photochemical reaction of ozone with water vapor. The tropospheric mixing ratio of ·OH cannot be directly determined analytically with precision because it is too reactive and short-lived, although it can be inferred (see below). Oxidation of reactive trace gases is initiated by ·OH, and subsequently leads to oxidized products such as CO or as occurs in the case of or to partially oxidized products in the case of hydrochlorofluorocarbons (HCFCs) or hydrofluorocarbons (HFCs). Partially oxidized products are removed from the troposphere by precipitation, either in “dry” form as in attachment (sorption) to particles, or “wet” as dissolved in rainwater. These tropospheric outfall oxidation products in turn enter the earth’s terrestrial and aquatic biomes. Obviously, the longer its tropospheric residence time, the more a gas will absorb infrared energy and contribute to warming of the troposphere. Likewise, the longer a non-reactive gas hangs around in the troposphere, the better are its chances ofeventually breaking through the physical boundary imposed by the tropopause, and being wafted up to the stratosphere. The stratosphere is the major sink for CFCs, where they undergo photolysis, resulting in the slow release of Cl and other halogen atoms. It is halogen atoms, especially Cl and Br, which cause the destruction of as was first noted by Molina and Rowland (45):

The affinity ofbromine for is approximately a 50-fold greater than the affinity ofCl, which is the reason the Montreal Protocol calls for severe limitations on the use of substances like methyl bromide and halons (bromine-containing fire retardants). This greater reactivity partially offsets its much lower abundance (~400-fold) than chlorine:

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The products of reactions 1 and 2 can recombine in a catalytic manner to regenerate the original reactant halogen atoms and accelerate the overall destruction of which further magnifies the effect of free bromine (40):

The efficacy of agreements like the Montreal Protocol can be discerned by monitoring the tropospheric mixing ratio of anthropogenic halocarbons with time and latitude. Thus, a clear decline in the abundance of methyl chloroform 1,1,1trichloroethane; “TCA”) in the troposphere was noted starting in 1991 for stations in the northern hemisphere, and in 1992 for stations in the southern hemisphere (Figure 12.2). Northern hemispheric stations exhibited higher mixing ratios than those of the southern hemisphere because most of the is produced and utilized in the north. Because the major sink for methyl chloroform is its oxidation with ·OH, the kinetics of this reaction have been used to indirectly deduce the ·OH concentration of the troposphere, a value which has been revised upward by 20% (54). The decline in methyl chloroform is rapid because it has a of about 5 years (Table 12.1). For the non-reactive chlorofluorocarbons (e.g., CFC-11 and CFC-12), and halons like H-1301 which have ofseveral decades or longer, the immediate effect of the Montreal Protocol has been to slow their rate of growth in the troposphere, rather than to cause their overall decline (12). Cumulatively, the effect of the Montreal Protocol appears to have led to a tropospheric decline of halogen abundance derived from anthropogenic halocarbons (46). This tropospheric decline was predicted to be followed by a stratospheric decline starting close to the turn of the century, year 2000.

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2. MICROBIOLOGICAL SINKS 2.1 Analytical Aspects The following discussion is meant to illustrate some of the experimental approaches taken, as well as the technical limitations of determining if microorganisms comprise a significant sink for atmospheric halocarbons. Whereas many studies on the transformation and fate of halogenated organics in the environment focus on relatively high concentrations in the milieu studied, and are often attributable to a point source of pollution, this is not the case for atmospheric halocarbons, which are present at sub-ppb to a few ppt mixing ratios, or roughly 4 to 5 orders of magnitude below that of atmospheric methane (approximately 1.75 ppm). These substances can be detected when dissolved in natural waters roughly in equilibrium with the atmosphere (oceans, lakes, rivers, groundwaters), which results in their concentrations being in the low picomolar range. Often, technical experimental difficulties are related to the selective sensitivity of the analytical instrumentation. For example, the electron capture detectors (ECD) of the gas chromatographs routinely employed in these studies have different sensitivities for the detection of halogens, where I > Br > Cl > F, and halocarbons with multiple halogens like Cl or Br give stronger responses than those with fewer halogens, or those with a proportional abundance of F. Thus, when a reductive dechlorination reaction takes place at near-ambient mixing ratios, it is often difficult to detect the product. In general, the ECD is more sensitive to CFCs or methyl bromide, than to HCFCs or methyl chloride. Some key questions are therefore not so easy to resolve because of these technical limitations. For example, do microbes degrade CFCs and HCFCs at ambient tropospheric mixing ratios of these gases, and if so, what are the products? Can the activity observed with whole cells or extracts at high concentrations (e.g., approximately 1,000 ppm) be extended down to sub-ppb levels? If no consumption of a gas by microbes is observed on exposure to high concentrations (approximately 1,000 ppm), is it possible that the gas inhibits the central metabolism of the microbe, and that its consumption may actually proceed at low concentrations? What possible benefit would a microorganism attain by having a capacity to degrade halocarbons at ppb or ppt concentrations, a situation that would require expenditure of more energy to maintain the necessary constitutive enzymes than could be derived by oxidation of the substrate? How can the degradation of halocarbons observed in laboratory studies with cultures, soils, or sediments be verified with field results? Finally, how can these field results be scaled up to determine if the global microbial sink for a halocarbon is of sufficient magnitude to affect its

2.2. Degradation of Chlorofluorocarbons, Hydrochlorofluorocarbons, and Volatile Chlorinated Hydrocarbons Chlorofluorocarbons (CFCs), because of their great utility as heat exchangers and their relatively inert chemical properties, have been widely used since the 1930s and released into the troposphere. Indeed, because CFCs have permeated the earth’s surface and have established equilibrium with its surface waters, they are commonly employed as tracers of water movement for both oceanic and groundwater systems. The central

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tenet of this application is that CFCs are conservative tracers and do not undergo significant chemical or biological degradation along their flow paths. While CFCs have proven resistant to aerobic biodegradation, they are susceptible to reductive dehalogenation (see below). This calls into question their efficacy as tracers of water mass flow through anoxic environments. The first hint that CFCs might be subject to biodegradation was attained during the course of gas flux chamber incubations over termite mounds and rice paddy soils. Both CFC-11 and CFC-12 were noted to decrease with time, to levels below the ambient mixing ratios present in the initially entrapped air, but the mechanism for these removals was not pursued (26). Reasoning that both of these environments (termite hindguts and paddy soil) have important functioning, anaerobic ecosystems, Lovley and Woodward (35) were able to demonstrate microbial consumption, at sub-ppb levels, of these two CFCs in anaerobic sediments (Figure 12.3), and with cells of Clostridium pasteurianum. In addition, CFC-11 was more susceptible to degradation

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than was CFC-12, and no consumption occurred under aerobic conditions. Reductive dehalogenation of chlorinated solvents had been previously noted with Methanosarcina barkerii, in a reaction mediated by coenzyme (30). Subsequently, Krone and Thauer (31) reported M. barkerii to reductively dechlorinate CFC-11 with as the electron donor, forming HCFC-21 and fluoride as products. The authors also found that CFC-11 fully inhibited methanogenesis by cells under the incubation conditions (approximately 1,000 ppm), but they speculated that at the prevailing atmospheric mixing ratios (roughly 0.2 ppb), this inhibition is unlikely to occur. Bacteria in groundwater samples were found to reductively dechlorinate ppb levels of CFC-11 with stoichiometric recovery of HCFC-21 as the product (60). The authors concluded from amendment experiments that this reaction was achieved by acetate-oxidizing sulfate-reducers, but not those that utilize lactate or by nitraterespirers. Inhibition experiments with molybdate reinforced this interpretation, and the lack of inhibition by 2-bromoethanesulfonate indicated that the phenomenon was not attributable to methanogens. Hydrochlorofluorocarbons (HCFCs) are currently being employed as interim freon replacement compounds because they are more reactive than CFCs, and therefore have much lower values for (approximately 10 to 15 years). As we have seen above, HCFCs might also arise locally from the reductive defluorination of CFCs by methanogens in anoxic environments. In experiments with anoxic sediments (Figure 12.4) and aerobic soils (Figure 12.5), HCFC-21 was shown to undergo further degradation, although the products could not be discerned using ECD analysis at these low concentrations. Under aerobic conditions, methanotrophic soils can oxidize HCFC21, and because HCFC-21 is a methane analogue, its presence partially inhibited methane oxidation (Figure 12.6). HCFC-21 and HCFC-22 were subsequently shown to be irreversible inhibitors ofboth the soluble and particulate forms ofmethane mono-oxygenase in Methylococcus capsulatus, and when applied at low concentrations, these substrates were co-oxidized with methane (39). In addition, both of these HCFCs partially inhibited the oxidation of methanol to indicating that they

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are also potentially disruptive in other facets of the methane oxidation pathway (e.g., methanol dehydrogenase). A number of other HCFCs can undergo co-oxidation by methanotrophs, including HCFC-142b (1-chloro-1,1-difluoroethane), HCFC-141 b (1,1 dichloro-1-fluoroethane), and HCFC-123 (2,2-dichloro-1,1,1-trifluoroethane), as well as selected HFCs like 134a (1,1,1,2-tetrafluoroethane) (4, 9, 61). Methanotrophs, however, cannot oxidize CFCs (R. Oremland, unpublished data). Information gained from field studies has reinforced the above results from laboratory experiments. The anoxic Gotland Basin of the Baltic Sea has been shown to be highly depleted inboth CFC-11 and carbontetrachloride relative to the aerated surface waters, indicating a bacterial water-column removal process is operative (32). Similarly, CFC-11 and (but not CFC-12) were removed, with depth, in vertical profiles ofthe anoxic waters of Saanich Inlet (Figure 12.7), a well-studied episodically anoxic fjord offshore of British Columbia, Canada (33). Several types of brominated and chlorinated methanes and ethanes have been observed to be removed from the anoxic waters of the Black Sea, presumably due to microbiologically mediated reactions (62). Happell and Wallace (17) investigated halocarbon profiles in short (approximately 2 m), aerobic surface soil cores taken on Long Island, New York. These cores received significant episodic input of halocarbons that were dissolved in rainfall. However, significant removal of and methyl chloroform (TCA) were noted some time after rain events, while there was no apparent removal of CFCs under these aerobic conditions, implying that a selective bacterial sink existed in the soil (Figure 12.8). The authors extrapolated their data to estimate the significance of live soils as a global sink for these volatile halocarbons. Surprisingly, aerobic soils were found to account for an average of 40% and 5%, for and TCA respectively, of the previously established global removal rates from the atmosphere. This soil sink estimate for removal of from the atmosphere was in good agreement with the earlier estimate of Khalil and Rasmussen (26). Since decay of tropospheric TCA is used as an indirect index of ·OH

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abundance (54), an additional global biological sink for TCA causes a modest downward revision of the estimated ·OH abundance. As was seen in the previous section, accurate ·OH concentrations are critical in determining the oxidation kinetics of a number of tropospheric halocarbons. In summary, a number of laboratory and field experimental observations on CFCs, HCFCs, and TCA have shown that natural populations of microorganisms can degrade these substances by aerobic or anaerobic attack, and that the reactions occur at environmentally-relevant (near-ambient) concentrations. These reactions can ultimately contribute to removal of certain halocarbons from the atmosphere, although the global significance ofprocesses occurring in anoxic environments may be small. In the case of volatile chlorinated hydrocarbons, the impact of aerobic soils appears to be quite significant. For CFCs and HCFCs, the importance ofthese reactions as a global sink has yet to be estimated with any degree of certainty, but it is unlikely that they are of comparable magnitude as that of oxidation by ·OH (HCFCs) or with escape to the stratosphere (CFCs). Nonetheless, better assessments ofglobal biological sinks for CFCs and HCFCs may affect future estimates of the values for these materials in the atmosphere. In addition, certain HCFCs are predicted to contribute to the precipitation oftrifluoroacetate from the troposphere, a subject ofcurrent environmental concern (see Section 2.3).

2.3. Degradation of Hydrofluorocarbons Hydrofluorocarbons (HFCs) represent the new class of freons meant to replace CFCs. HFCs have estimated values of approximately 15 years, and therefore some will survive oxidation in the troposphere and be transported to the stratosphere. Fluorinated moieties are generally not thought to cause ozone depletion,

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although this is still a matter of ongoing debate (28). HFCs (e.g., HFC-134a) undergo substantial chemical attack in the troposphere with a series of complex oxidations initiated by hydroxyl radicals. The products of these reactions lead to the formation of trifluoroacetyl fluoride (or trifluoroacetyl chloride in the case ofHCFC-123 and HCFC124). These trifluoroacetyl moieties hydrolyse within water droplets and form the highly stable molecule, trifluoroacetate (TFA) (20, 68). Because TFA is refractory to both chemical and biological degradation, it is expected to accumulate in the environment and the living biota with increased usage of HFCs and HCFCs, as these replace CFCs. Certain environments, such as evaporative vernal pools and coastal wetlands, are predicted to quickly reach dissolved TFA concentrations greater than from the combined effects of precipitation, evaporative concentration, and the absence of chemical or microbiological sinks (64). Recently, dissolved TFA concentrations as high as ~90 nM were detected in seasonal wetlands (3). The consequences of such an accumulation are unforeseen. The review by Key et al. (25) gives the reader a more complete discussion of the potential manifestations of this problem. The concept that TFA is a molecule highly recalcitrant to chemical or biological degradation underwent serious challenge by the results ofVisscher et al. (67). Using labeled TFA, these workers discovered that anaerobic saltmarsh and freshwater sediments were able to completely degrade low micromolar levels ofapplied TFA within incubation periods of 1 to 4 weeks, to either (under sulfidogenic conditions) or (under methanogenic conditions). Degradation proceeded by a series of sequential defluorinations, which resulted in the transient accumulation of difluoroacetate, monofluoroacetate, and acetate (Figure 12.9). The methyl group of the acetate was subsequently oxidized to (under sulfidogenic conditions), or cleaved and reduced to (under acetoclastic methanogenic conditions). With aerobic conditions, TFA was cleaved into fluoroform and No degradation was observed under nitratereducing conditions, although after the nitrate was depleted, TFA underwent reductive

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defluorination as had been previously seen (52). Not long after these observations were reported, another group at Michigan State University observed that denitrifying cultures of Azoarcus tolulyticus could decarboxylate TFA as a cometabolic product of toluene degradation (25). Collectively, these results underscored the idea that TFA may be readily susceptible to biodegradation by several different microbial pathways. The facility with which TFA was degraded in these experiments was quite unexpected, as were the novel observations of its reductive defluorination. Aside from the reassuring notion that TFA was easily biodegradable, some of the results also gave rise to a few reasons for concern. For example, the production of fluoroform (trifluoromethane, or TFM), observed in the aerobic incubations, generated some stir because this gas has a long a known large GWP (approximately 2100), and an unknown ODP (29). Because on a global basis, aerobic soils have far more surface area in contact with the atmosphere than do anaerobic sediments, TFM might wind up being the major TFA degradation product of natural systems, rather than methane or carbon dioxide. Also, unlike atmospheric methane, TFM is quite resistant to oxidation by soil microbes, indicating the general lack of a significant biospheric sink for TFM removal (28). Atmospheric TFM concentrations appear to have been increasing since 1978 (48), but not all of the TFA in the environment is attributable to HFC and HCFC breakdown products (24). The appearance of monofluoroacetate as an intermediate was also a cause for concern because this substance is a potent metabolic inhibitor of the tricarboxylic acid cycle, as well as many other biochemical pathways that utilize acetate for anabolism or catabolism. However, production of organofluorides in nature, including monofluoroacetate, can also be attributed to biological sources like Streptomyces cattleya (15). It would seem therefore, that microbial pathways for the dehalogenation of low molecular weight fluorinated substances are common in nature, although this may not extend to perfluorinated compounds like TFA. An analogous compound, chlorodifluoroacetate (CDFA), was recently detected in rainwater as a possible photolysis degradation product of atmospheric CFC-113 (trifluorotrichloroethane). CDFA is also thought to be resistant to biodegradation (38). The concerns about TFA were subsequently allayed by the subsequent research on its degradation in soils and sediments. Unfortunately, follow-up experiments failed to reproduce the earlier results (52). Despite repeated attempts using the same and also new biological materials, no significant loss of was noted, no defluorination products were ever detected, and no gaseous products accumulated. Curiously, the TFA decarboxylation results with A. tolulyticus could also not be repeated (C. Griddle, personal communication). This odd state of affairs is where matters stood for several years, with two independent groups of scientists shaking their heads over these bizarre observations. It seems that TFA degradation by microbes, while not a chimera, nonetheless has elusive, gossamery qualities that make it difficult for a researcher to grasp and hold. Sequential defluorination of TFA under methanogenic conditions has been recently reported during prolonged (90 week) experiments with an anaerobic bioreactor (27). While it is now clear that TFA is not readily biodegraded by microorganisms, the fact that its biodegradation has been observed certainly merits further investigation. The widespread industrial use of a number of volatile and nonvolatile perfluorinated compounds, coupled with their release into the environment,

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beckons the need to re-examine this problem (25). If experimental success is realized, it is likely that eventual isolation of the mechanisms for reductive defluorination will be of great scientific interest and have practical applications for the remediation of perfluorinated wastes. The reader is referred to the reviews of Harper and O’Hagan (17) and O’Hagan and Harper (47) for details on natural sources of fluorinated compounds in the environment.

2.4. Degradation of Methyl Halides Methyl halides, especially methyl chloride (MeCl) and methyl bromide (MeBr), represent the largest sources of halogen atoms to the stratosphere, and are thus of the greatest importance with regard to ozone degradation. However, unlike CFCs, HCFCs, and HFCs, methyl halides in the atmosphere arise from a mixture of anthropogenic and biological sources (2, 34), plus other “natural” sources that include emissions from biomass burning (37), and even a very minor source from volcanoes (24). The global biological sources of these two gases greatly exceed their anthropogenic ones. Biogenic sources of methyl halides include their production by fungi, marine phytoplankton, salt marsh macrophytes, and macroalgae (37, 56, 69,71). This topic has been the subject of a recent comprehensive review by Harper (18). Transhalogenation reactions, both chemical and biological, also occur between these two gases (and with methyl iodide), making their source identification more complicated (19). Anthropogenic sources of MeBr to the atmosphere arise mostly from its use as a biocide, primarily as a preplanting soil fumigant for highly valued specialty crops like strawberries, flowers, and certain vegetables. MeCl is used for a variety of applications including refrigeration and as a chemical methylating agent. MeCl and MeBr can be degraded in anaerobic environments. MeCl will support the growth of certain homoacetogenic bacteria (57). Dechlorination of MeCl by homoacetogen strain MC involves transfer of the methyl group to a folate, with subsequent formation of a methyl corrinoid, coupled with the fixation of to produce a de novo synthesis of acetate (40, 42):

In environments containing free sulfide, MeCl is susceptible to nucleophilic attack and will form methane thiol (MeSH) in a biologically accelerated reaction (1). MeBr is even more susceptible than MeCl to chemical nucleophilic substitution with sulfide, forming both MeSH and dimethylsulfide (DMS) (50):

Because DMS, as well as MeSH, can fuel both methanogenesis and sulfate-reduction, there is a microbiological link to this chemical pathway. This can be best illustrated from the pathway of degradation occurring within a sulfide-rich microbial mat (Table 12.2).

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Aerobic degradation of MeCl and MeBr can be achieved via the monooxygenases of methanotrophs and nitrifying bacteria (5, 55). Because methyl halides are molecular analogues of methane and ammonium, they are cometabolized and act as competitive inhibitors of methane monooxygenase (MMO) and ammonia monooxygenase (AMO) (11, 39, 49, 50). However, methyl halides do not support the growth of these microbes. Methanotrophic soils readily were shown to oxidize MeBr, and because significant inhibition of MeBr oxidation (72%) was achieved with methyl fluoride, therefore most of the observed MeBr oxidizing activity was attributed to methane-oxidizing bacteria (51). In aquatic environments, bacteria readily oxidized MeBr in freshwater, estuarine, marine, and hypersaline/alkaline systems (Figure 12.10). The oxidation of MeBr in a freshwater lake was entirely inhibited by methyl fluoride, and this inhibitor also achieved 82% inhibition of dibromomethane oxidation. The clear participation of methanotrophs and/or nitrifiers was therefore indicated for most, if not all, of the measured MeBr oxidation (13). In contrast to the above studies which directly implicated the involvement of monooxygenases, MeBr oxidation in marine and hypersaline systems has been shown to display much different patterns of transformation. The aforementioned results implicated the involvement of methylotrophs that normally grow on the breakdown products of common osmolytes, such as trimethylamine (from glycine betaine) and dimethylsulfide (from dimethylsulfoniopropionate). Thus, oxidation of in hypersaline and alkaline Mono Lake proved insensitive to a number of monooxygenase inhibitors (e.g., methyl fluoride, allyl sulfide, acetylene). In addition, oxidation was stimulated by trimethylamine but not by dimethylamine or glucose (6). Similarly, methyl fluoride did not inhibit MeBr oxidation in seawater and had little effect in estuarine waters (13). Oxidation of in seawater was unaffected by additions of unlabeled methanol or MeCl, was partially inhibited by MeBr (due to isotope dilution), and was stimulated by trimethylamine. Collectively, these results implicated that MeBr was degraded by a cometabolic phenomenon, most probably carried out by trimethylamine-utilizing methylotrophs. Hoeft and coworkers (22) recently demonstrated MeBr cometabolism in several strains of newly isolated marine methylotrophic bacteria, but noted their oxidation activity to be linked to dimethylsulfide rather than trimethylamine. Miller et al. (44) investigated the oxidation of MeBr in two fumigated strawberry fields in California. They found that in situ oxidation was highest during fumigation events, but they could not attribute the observed activity to a mono-oxygenase because neither methane-oxidation nor ammonium-oxidation could be detected in soil samples. Furthermore, although incubated soils from these sites were readily capable of MeBr oxidation (Figure 12.11), cometabolic linkage to a number of possible substrates (e.g., methylated amines, formate, methanol, ethanol, glucose, ammonia, methane) could not be demonstrated. Further experimentation with soils demonstrated enhanced MeBr degradation occurred with repeated additions of MeBr (Figure 12.12), leading to the conclusion that microorganisms were using this substance as a growth-supporting substrate. Using a mineral salts medium with MeCl as a substrate, Doronina et al. (10) reported on the isolation of a soil bacterium (eventually identified as Methylobacterium CM4) that was a facultative methylotroph. A similar strategy was employed to isolate a MeBr-oxidizing aerobe from fumigated soils. This bacterium, designated strain IMB-1,

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proved capable of achieving growth on MeBr in a mineral salts medium (Figure 12.13). Subsequent investigations have determined that strain IMB-1 is a facultative methylotroph able to grow on MeI, MeBr, MeCl, and methylated amines, as well as on substrates like acetate, pyruvate, and glucose. No growth occurs with methane, methyl fluoride, methanol, or formate (7). Growth on MeI, MeBr, and MeCl probably goes through a single enzyme system because all substrates can serve as competitive inhibitors of methyl halide degradation (57). In addition, methyl fluoride did not inhibit oxidation of MeBr. Chloropicrin (trichloro-nitro-methane) is inhibitory to the growth of this organism, as well as to its ability to oxidize MeBr. This is significant because chloropicrin (“tear gas”) usually comprises one third of the MeBr-fumigation mixtures applied to agricultural soils, and therefore probably restricts the amount of MeBr oxidation mediated by the resident microbiota. Addition of IMB-1 cells to soils greatly speeded their ability to consume MeBr. This suggests that artificially enhanced soil populations of this organism may be employed to constrain or eliminate outflux of MeBr to the atmosphere during fumigation operations, provided that the chloropicrin problem can be overcome. Moreover, the pathway of MeBr oxidation could be induced by exposure of glucose-grown cells of IMB-1 to MeBr (57). This observation facilitates the mass culture of IMB-1, and opens the door for its possible use as a MeBr-absorbing and -degrading “bioreactor” during field fumigations. Based on its 16s rRNA gene sequence, strain IMB-1 is a member of the aligning near the clade of Rhizobium, although it does not fix nitrogen (7). Currently there exist in culture five aerobic members of the that can grow on one or more methyl halides (Figure 12.14). These include the closely related strains IMB-1 and CC495, the latter isolated from leaf-litter with MeCl as its substrate (8). Both of these strains belong to the genus Aminobacter. A marine organism, strain

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MB2 was isolated from a California tidepool with MeBr as its substrate, and has been named Leisingera methylohalidivorans (58). It grows on only a few substances: MeI, MeCl, MeBr, methionine, and glycine betaine. Strain CC495, as well as Hyphomicrobium chloromethanicum CM2 and Methylobacterium chloromethanicum CM4, are facultative methylotrophs that can grow on MeCl; they can oxidize, but cannot grow on MeBr or MeI. The pathway for methyl halide oxidation initially follows a corrinoid-mediated methyltransferase reaction followed by transfer to a folate, rather than one initiated by a monooxygenase or a hydroxylase (8, 65, 66). The methyltransferase genes of Methylobacterium strain CM4 have significant sequence similarity to methyltransferases of methanogens (66). Similar methyltransferase genes have been detected in strain IMB1 (70). Methyl halide oxidation by strains IMB-1, MB2, and CC495 also achieves very large isotope discrimination factors (Figure 12.15), equivalent to the enrichments observed with methanogenic Archaea (44). These results with stable C isotopes may provide information to be used in constraining the global atmospheric budgets for MeBr and MeCl, as has been done for methane. Soils represent significant global sinks for atmospheric MeBr, consuming about 42 grams/year, or about 30% of the atmospheric burden (59). Together with the tropospheric ·OH and a net oceanic sink, the for MeBr is currently estimated to be approximately 0.7 years. The soil sink has been shown to be bacterial in its nature, unrelated to methane-oxidation, yet able to consume ambient mixing ratios (roughly 10 ppt) of MeBr (21, 59). Recent studies (14) with strains IMB-1 and MB2 have demonstrated that they can oxidize MeBr at these ambient tropospheric mixing ratios (Figure 12.16).

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3. SUMMARY Diverse aerobic and anaerobic microorganisms have the ability to degrade selected atmospheric halocarbons. In many cases, these degradative reactions are operative at the ambient tropospheric mixing ratios of these substances. This will either result in their complete destruction, or in the accumulation of non-degradable (and perhaps toxic) intermediates within aquatic and terrestrial ecosystems. The significance of these microbiological sinks with respect to the global biogeochemical budgets of halocarbons has not been accurately assessed. However, preliminary results suggest that they depend on the particular compound in question. Thus, biodegradation appears to be a minor global sink for certain CFCs while for methyl halides it is a major one. A challenge for future research, therefore, is not only to define novel pathways and microorganisms, but also to determine if these reactions truly represent important biogeochemical sinks. REFERENCES 1.

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OREMLAND Elkins JW (1996) Decline in the tropospheric abundance of halogen from halocarbons: Implications for stratospheric ozone depletion. Science 272:1318-1322 47. O’Hagan DO & Harper DB (1999) Fluorinecontaining natural products. J. Fluorine Chem. 100:127-133 48. Oram DE, Sturges WT, Penkett SA, McCulloch A & Fraser PJ (1998) Growth of fluoroform in the background atmosphere. Geophy. Res. Lett. 25:35-38 49. Oremland RS & Culbertson CW (1992) Evaluation of methyl fluoride and dimethyl ether as inhibitors of aerobic methane oxidation. Appl. Environ. Microbiol. 58:2983-2992 50. Oremland RS, Miller, LG & Strohmaier FE (1994) Degradation of methyl bromide in anaerobic sediments. Environ. Sci. Technol. 28:514-520 51. Oremland RS, Miller LG, Culbertson CW, Connell TL & Jahnke L (1994) Degradation of methyl bromide by methanotrophic bacteria in cell suspensions and soils. Appl. Environ. Microbiol. 60:3640-3646 52. Oremland RS, Matheson L, Guidetti J & Visscher PT (1995) Summary of research results on bacterial degradation of trifluoroacetate (TFA), November, 1994 - May, 1995. USGS Open-File Rpt. 95-422, Menlo Park, CA 53. Oremland RS, Lonergan DJ, Culbertson CW & Lovley DR (1996) Microbial degradation of hydrochlorolfluorocarbons and in soils and sediments. Appl. Environ. Microbiol. 62:1818-1821 54. Prinn RG, Weiss RF, Miller BR, Huang J, Alyea FN, Cunnold DM, Fraser PJ, Hartley DE & Simmonds PG (1995) Atmospheric trends and lifetime of and global OH concentrations. Science 269:187-192 55. Rasche ME, Hyman HR & Arp DJ (1990) Biodegradation of halogenated hydrocarbon fumigants by nitrifying bacteria. Appl. Environ. Microbiol. 56:2568-2571 56. Rhew RC, Miller BR & Weiss RF (2000) Natural methyl bromide and methyl chloride emissions from coastal salt marshes. Nature 403:292 -295 57. Schaefer JK & Oremland RS (1999) Oxidation of methyl halides by the facultative methylotroph strain IMB-1. Appl. Environ. Microbiol. 65:5035-5041 58. Schaefer JK, Goodwin KD, McDonald I, Murrell JC & Oremland RS. Leisingera methylohalidivorans gen. nov., sp. nov., a marine methylotroph that grows on methyl halides. Int. J. System. Evol. Microbiol. 52:851859

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Chapter 13 DECHLORINATION OF SEDIMENT DIOXINS: CATALYSTS, MECHANISMS, AND IMPLICATIONS FOR REMEDIAL STRATEGIES AND DIOXIN CYCLING CYNDEE L. GRUDEN 1, Q. SHIANG FU 2, ANDREI L. BARKOVSKII 3, IRIS D. ALBRECHT 1, MARY M. LYNAM 4, AND PETER ADRIAENS 1 1

Environmental and Water Resources Engineering, University of Michigan, Ann Arbor, Ml, USA Department of Civil and Environmental Engineering, Stanford University, Palo Alto, CA, USA Georgia College and State University, Milledgeville, GA, USA 4 School of Public Health, University of Michigan, Ann Arbor, Ml, USA 2

3

1. Introduction Polychlorinated dibenzo-p-dioxins (PCDD) are among the most hazardous environmental pollutants. The United States Environmental Protection Agency (USEPA) has stated that dioxins should be regulated as probable carcinogens and that these compounds pose a concern at any concentration level in the environment (34). The laterally substituted congeners (2,3,7,8-substituted) are considered to exhibit the highest toxicity, due to their molecular planarity and stability, and their capacity for bioaccumulation. Despite the regulatory emphasis on 2,3,7,8-substituted PCDD, dioxins are typically found as complex mixtures that are globally distributed in the environment. Prior to the late 1970s, it was believed that dioxin contamination was attributable solely to chemical manufacturing processes (e.g., chlorophenol and herbicide synthesis). It was later determined that PCDD are ubiquitous environmental contaminants generated by natural and anthropogenic processes and combustion(26), and have even been found in a million-year-old ball clay (37). Aerosols and particulate material (e.g., fly ash) are recognized as a major means of PCDD transport through the atmosphere, toward deposition in the ultimate environmental sinks, predominantly soils and sediments. These combined processes and pathways have resulted in dioxin concentrations in sediments, ranging from picograms to nanograms per kilogram for and nanograms to micrograms per kilogram, for all 2,3,7,8-substituted congeners For comparison, these concentrations are three to five orders of magnitude below sediment concentrations of polychlorinated biphenyls (PCBs) (16). In studies on Dehalogenation: Microbial Processes and Environmental Applications, pages 347-372 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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the fate of dioxins, it has been observed that the corresponding homologue profiles, or chlorination patterns, from source and sink are distinct. The multitude of PCDD sources (104), and environmental transformation reactions (32), may be responsible for the limited successes in establishing source-sink correlations for dioxin residues. Recent peer-reviewed literature has confirmed the potential for dechlorination of environmentally significant PCDD concentrations in sediments via biotic and abiotic transformation reactions (1, 3, 6, 12, 41, 43). River and lake sediments are very reduced beneath the surficial layers, which render them ideal habitats for anaerobic microbial communities. Moreover, functional groups from hurnic acids, heavy metals, biogenic factors, and reduced mineral surfaces constitute potentially significant electron shuttles in reduced environments. Characterization of microbial activities, in combination with geochemical indicators, may provide a method for predicting PCDD dechlorination in a given sediment environment. In addition, this information provides a scientific rationale for the interpretation of observed dechlorination patterns in environmental sinks, and also a potential strategy for the remediation of environmental concentrations of laterally substituted PCDD congeners.

2. DISPOSITION OF DIOXINS IN THE ENVIRONMENT 2.1. Physical and Chemical Characteristics of Dioxins Dioxins are a complex group of compounds consisting of two benzene rings joined by ether bonds. There are eight carbon positions available for chlorination, resulting in 8 homologue (isomer) groups. Depending upon the chlorination pattern, 75 dioxin congeners can theoretically be formed (Table 13.1). The different substitution patterns determine the chemical properties of the PCDD congeners, which, in turn, affect their

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environmental fate. Shiu et al. (97) endeavored to provide a reliable database of PCDD properties based on the peer-reviewed literature (90, 92), and from thermodynamic equations based on fundamental molecular properties. Other researchers have relied on prediction methods for instances when accurate experimental data were unavailable or extremely difficult to obtain (50, 55, 61, 91). Due to the aromaticity, molecular symmetry, and low polarity of PCDD, the congeners are considered hydrophobic. This is supported by published chemical characteristics; their octanol-water partition coefficient and aqueous solubility (25°C) range from 5.30 to 11.16, and 417 to respectively (Table 13.2). The increases with increasing molar volume in a near linear fashion, and aqueous solubility decreases slightly with each addition of a chlorine substituent, comparable to PCBs (69). However, the Henry’s Law constant was reported to range from 0.1 and for PCDD, values that are significantly lower than those reported for PCBs by Burkhard et al. (22). The decline in H per sequential chlorine added is a result of the concomitant decrease in both vapor pressure and solubility. Due to their hydrophobic and non-volatile nature, dioxins predominantly partition into sediments and soils as well as biological tissue (e.g., fish, plants). The laterallysubstituted congeners (2,3,7,8-substituted) are considered to exhibit the most toxicity, due to their molecular planarity and their ability to bind to biological receptors. A toxicity equivalency procedure, which involves assigning individual toxicity equivalency

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factors (TEFs) to the 2,3,7,8-substituted congeners, was developed for risk assessment (70, 102) (Table 13.3). Toxic equivalency (Teq) can be calculated by multiplying the observed concentration in a given environment by the assigned TEF. 2.2. Energetics of Dioxin Reactivity The intrinsic thermodynamic properties of compounds serve as descriptors for their reactive potential in environmental systems. Molecular calculations may provide information to predict, or to interpret observed degradation/dechlorination pathways, assuming no enzymatic or microbial preferences for specific substituent patterns are inferred. In the case of reductive dechlorination of aryl halides, good correlations have been obtained between predicted pathways based on molecular redox potentials, or Gibbs free energy of formation, and experimental observations (24, 80, 101). Huang et al (50) estimated the Gibbs free energy for the reductive dechlorination of chloro-pdioxins and their redox potential, based on predicted thermochemical values (31, 62, 63). Under standard conditions, the amount of Gibbs free energy available from dehalogenation ranges from -137 to -180 kJ/mol, depending on the position of the removed substituent and the parent molecule. These thermochemical properties provide a basis for indicating the (biochemical) potential for microbial dechlorination. The redox potentials range from 295 to 516 mV, similar to other chlorinated compounds (30). Dechlorination of distinct substituents on the same congener may yield different amounts of energy, indicating the potential for preferred dechlorination pathways. With increasing frequency, calculations of HOMO (Highest Occupied Molecular Orbital) – LUMO (Lowest Unoccupied Molecular Orbital) gaps are used to describe molecular reactivity. The HOMO-LUMO Gap is an indicator of stability in a molecule; a larger gap indicates a decrease in the probability of further reaction. More highly chlorinated dioxin congeners will likely be more susceptible to further reaction due to a smaller HOMO-LUMO Gap (Figure 13.1) (60). Isomers within the same homologue group may exhibit differing reactivities based on their substitution pattern. Whereas

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should, due to its smaller HOMO-LUMO gap, exhibit significant reactivity, the apparent environmental persistence of may be due to factors other than molecular descriptors. Under reducing conditions, these predictions are consistent with those obtained for other halogenated aromatic compounds, such as chlorobenzenes and polychlorinated biphenyls (PCBs), where higher rates of dechlorination were observed with increasing chlorine content (2, 7). 2.3. Sources, Sinks, and Distribution of Dioxins in the Environment Dioxins have never been intentionally manufactured (except during production of analytical standards), but are formed during a number of processes, including: (i) synthesis of chlorophenol and chlorinated herbicides (e.g., 2,4-D and 2,4,5-T), (ii) wood treatment and preservation with pentachlorophenol, (iii) pulp and paper bleaching, and (iv) incineration processes (23, 49, 75, 88). The environmental burden of PCDD in waterways and sediments is dependent upon the point sources of discharge and the nature of their origin. Congener-specific analysis helps to differentiate signature patterns for certain point sources or process characteristics (Table 13.4). Diffuse land-based sources, such as domestic burning of coal/wood, are the principal supply of to the atmosphere (57). Incineration processes have been implicated in de novo (e.g., from elemental carbon) or precursor-based (e.g., chlorobenzenes, chlorophenols) generation of dioxins on fly ash. Data collected worldwide in 1995 demonstrates a direct correlation ( with a probability greater than 99%) between PCDD release and annual emissions (10). Dioxin emissions from chemical manufacturing are difficult to quantify as these include direct discharge and accidental release events, and the contamination remains localized. Atmospheric deposition and industrial discharges have resulted in total dioxin concentrations on the order of nanograms to micrograms per kilogram in urban environments (9). The dominant

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homologue group in all environmental compartments is . The 2,3,7,8-substituted congeners constitute a minor fraction of the and total PCDD homologue groups in incineration-dominated patterns. As PCDD are very poorly soluble in water and have relatively high partition coefficients, they do not stay in the aqueous phase, but strongly sorb to particles. Thus, sediments of rivers, lakes, and oceans are the ultimate sinks for water-borne dioxins (39, 45,87). Research reports indicate that particle or aerosol-associated PCDD are subjected to UV-mediated dechlorination reactions in the atmosphere, prior to wet or dry deposition in topsoils and sediments (54, 56). Furthermore, sediment analyses from a site impacted only by atmospheric deposition (Siskiwit Lake) suggest the photochemical synthesis of from pentachlorophenol (10). In general, wet deposition processes, i.e., precipitation, that scavenge highly chlorinated PCDD that are largely particlebound, may result in the enrichment of these congeners in terrestrial sinks (26). The fate of PCDD in the environment will depend upon their fugacity in each medium, and determination of fugacity ratios may assist in determining the direction of exchange. Adriaens et al. (3) determined in laboratory microcosms that the net exchange of PCDD was in the direction of sediments (fugacity ratios <1). On the contrary, Lohman et al (57) discovered net volatilization of PCDD from the Hudson River Estuary water column, and Harner et al. (47) suggested soil-to-air PCDD transfer

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contributing to atmospheric burdens when PCDD air concentrations are low. The highly chlorinated dioxin congeners are non-volatile compounds with an extremely low vapor pressure (91) and Henry’s Law constant (Table 13.2), however volatilization from water and sediments, in some instances, may merit quantification (97).

2.4. Trends of Dioxin Sediment Burdens and Source-Sink Correlations Due to the abundance and complexity of PCDD source-sink data, multivariate data analysis has been used to define similarities and differences among samples. Pattern recognition techniques such as Principal Component Analysis (PCA) have been used to establish trends from composite PCDD distribution data. PCA involves threedimensional expression of data while preserving the variance and relativity among the samples. Figure 13.2 includes a location map and a PCA plot for 2,3,7,8-substituted dioxin residue patterns in international industrial waterways (103). Principal component scores and loadings for each sample were calculated from the Eigen vectors of the scaled data matrix. Each plane represents a principal component that was calculated to factor or group the samples. Sediment samples are clustered according to similarities and differences among isomeric patterns. This method, in combination with hierarchical cluster analysis, has been used to characterize PCDD residues, and to differentiate sources of these compounds inthe environment. Analyses of dioxin distribution data indicate that the corresponding homologue profiles of PCDD from source and sink diverge (104). As previously described, initial environmental PCDD patterns are characteristic for the process that generated them. Incineration source profiles exhibit a more uniform distribution of homologues than profiles generated from chemical synthesis, which are composed of primarily 2,3,7,8substituted congeners, depending on the contribution of point and diffuse sources. Sink profiles, on the other hand, are dominated by either highly chlorinated PCDD, (e.g., and ), or by 2,3,7,8-substituted congeners. A principal components loadings plot of normalized 2,3,7,8-substituted PCDD can be used to provide information on which congeners in this homologue group best explain the differences between sources and sink samples. Highly chlorinated congeners, (e.g., and ), and particularly the 2,3,7,8-substituted isomers, are responsible for the observed differences. The discrepancy between source and sink residue patterns may be attributable to the diversity of PCDD sources or to environmental transformation reactions that occur prior to PCDD deposition.

3. DIOXIN REACTIVITY IN SEDIMENTS Extensive field observations have demonstrated that sediments can be considered a reactive system, in which microbial and abiotic attenuation (including dechlorination) processes may alter PCDD source profiles, ultimately resulting in the release of lesserchlorinated congeners. In order to better understand, and to develop fundamental information to predict the fate of PCDD in environmental sediments, a comprehensive study of dechlorination processes in field samples was undertaken by the Fundamental and Applied Microbiology for the Environment (FAME) laboratory at the University of Michigan. The overarching objectives of the research performed by FAME were to, (i)

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interpret the potential for dioxin reactivity in sediments, (ii) determine the yield and extent of dechlorination, (iii) assess dominant dechlorination pathways and kinetics, (iv) evaluate the fate of 2,3,7,8-substituted residues and the potential accumulation of , and (v) investigate the potential for stimulating sediment dechlorination kinetics, based on organic and inorganic electron donors and priming compounds. The freshwater and estuarine sediments sampled for this work were collected from a variety of locations in the United States including the Hudson River, Fort Edward, New York; American Creosote Works Aquifer, Pensacola, Florida; Cherokee Pond, Athens, Georgia; Paletta Creek, San Diego Bay, California; and Lower Passaic River, New Jersey. Results were dependent upon the amendment applied, and the contamination history and source of the sediments used. Most of the research performed focused on the Passaic River Dioxin Superfund Site. The Passaic River Estuary is an enclosed estuary formed by the confluence of the tidal Passaic River and Hackensack River in the north, and the Arthur Kill and the Van Kull in the south. Nearby metropolitan areas include New York City and Newark, New Jersey. Previous research indicated that the ecology of the Newark Bay and the lower Passaic River has been adversely impacted by century-long anthropogenic activities (65, 67, 68), including bleaching effluents, 2,4,5-T manufacturing, and atmospheric deposition via incineration. Evidence from sediment analyses has suggested the primary source to be the Diamond Alkali Site, an industrial site where pest and weed killers were manufactured during the 1950s and 1960s (20, 86, 100). Dioxin patterns from this pesticide manufacturing process are dominated by the congener, and concentrations up to 21,000 ppt were found in sediments proximal to the site (20). Source-sink correlations are indeterminate; this location has been impacted by many point and diffuse sources, including atmospheric deposition, which contributed to elevated and concentrations. Passaic River sediment cores and saline bottom river water were collected for a prolonged series of laboratory studies. An experimental array summarizing the research performed during the last decade is provided in Table 13.5; detailed experimental protocols were included in previous publications (1, 3, 5, 7, 8, 12, 13, 41-43). Varied biogeochemical conditions (freshwater, estuarine, marine), as well as amendments of dissolved organic matter (DOM) (e.g., humic acid, polymaleic acid, 3,4dihydroxybenzoate, catechol, and resorcinol), a bulk electron donor (titanium citrate), hydrogen gas, organic acids, and a potential priming compound (2-monobromodioxin) served as variable parameters for the study. 3.1. Sediments and Their Potential for Dioxin Dechlorination The initial step in the anaerobic degradation of most chlorinated aromatic compounds is reductive dechlorination (2, 16, 99). This reaction results from electron transfer between a reduced organic or inorganic electron donor and a chlorinated compound, such as PCDD. Previous dioxin transformation studies incorporated single congeners and soil- or sediment- derived pure and mixed cultures (3,5,11,12,18). Only enrichments derived fromsediments demonstrated dechlorination activity, but the reason for this is not yet clear.

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3.1.1. Humic Substances Contributing to Abiotic Dechlorination Anaerobic sediments contain aquatic humic substances, heavy metals, biogenic factors, and mineral surfaces; these constitute a significant reducing capacity. Aquatic humic substances are organic acids of biological origin found in all natural waters. They are ubiquitous due to their diverse origins and pathways of formation, as well as their recalcitrance to geochemical and microbial degradation (66). Humic materials are categorized based on their solubility: humic acids are soluble at pH above 2, fulvic acids are soluble at any pH, and humins are insoluble. Sources of aquatic humus include terrestrial (e.g., plant) organic material and microbial biomass. Photosynthetic aquatic microorganisms are the main source of organic matter in the oceans, while terrestrial contributions constitute a small fraction (less than 5%). Humic substances marine environments (comprising less than 15% of carbon in marine dissolved organic matter), are primarily composed of unsubstituted alkyl carbons, and are low in aromatic carbon relative to humic substances from freshwater environments (17). The physical and chemical properties of aquatic humic substances, which exhibit temporal and spatial variation, are not yet well characterized. Soil humic substances, composed of humic and

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fulvic acids, appear to have higher aromaticity than aquatic humics, where paraffinic and carbohydrate-like structures dominate. However, the NMR spectral features of paraffinic structures in terrestrial deposits (peat, soils) are very similar to those of aquatic sedimentary humic acids, characterized by highly branched and cross-linked macromolecular structures attributed to microbial and algal residues, suggesting similar origins (48). Previous research suggests that the quinone-hydroquinone functional groups of humic materials are responsible for their associated redox activities (94). Quinone moieties act as electron acceptors during microbial reduction of humic substances (95). Current information further indicates that quinones may be the primary site for abiotic humic-mediated electron transfer, from reduced sulfur and iron species to nitroaromatic and chlorinated compounds (25, 94). In light of the importance of the quinonic functionality during electron transfer, and the evidence for abiotic humic constituentmediated dechlorination of dioxins, humics may contribute significantly to dioxin dechlorination.

3.1.2. Biogeochemical Indicators for Respiratory Activity Microbial diversity in estuarine and marine environments is impacted by salinity gradients, high sulfate concentrations, and availability and competition for electron donors. Salinity gradients between 5 to 20 parts per thousand have been reported to exert a negative effect on aerobic heterotrophic (77, 83), methanogenic (27), and anaerobic (74, 76) respiratory activities. In estuarine and marine environments, a high sulfate content (20 to 30 mM) results in the dominance of sulfate reduction (23, 98), but denitrification (19, 74), methanogenesis (27, 76), iron reduction, and fermentation (76) have also been observed. A fundamental difference in fatty acid consumption in low and high sulfate environments has been observed (72). Under freshwater conditions, fermentative processes appear to be responsible for fatty acid consumption, while a coupling between organic acid consumption and sulfate reduction occurred under sulfate-rich conditions (59). Growing evidence indicates that hydrogen is a key electron donor used in the dehalogenation of lesser-chlorinated organics, and organic electron donors (e.g., fatty acids) serve primarily as precursors to hydrogen generated during fermentation (29,35). This observation suggests that dehalogenating microorganisms occupy an anaerobic niche similar to hydrogen-utilizing methanogens, homoacetogens and sulfidogens, and will compete for hydrogen (64). Under freshwater conditions, discrete dissolved hydrogen concentration ranges correlate with specific dominant terminal electron accepting processes (e.g., 7 to 20 nM for methanogenesis; 1 to 5 nM for sulfatereduction; 0.8 to 1.0 nM for iron-reduction). Likewise, ambient hydrogen concentrations in saline environments (less than 6 nM) suggest a predominance of sulfate-reduction. Yang and McCarty (107) have reoprted on studies of benzoate-acclimated, dehalogenating mixed cultures with organic acids, where benzoate metabolism produced hydrogen at higher levels and faster rates than propionate. In these experiments, methanogenesis was stimulated in the former (benzoate-fed), and dechlorination in the latter (propionate-fed) cultures. An optimum hydrogen concentration (2 to 11 nM) resulted in reductive dechlorination without competition from methanogens. A similar

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mechanism may be operative during PCDD dechlorination. Therefore, reductive dechlorination rates and pathways in freshwater sediments will likely differ from those prevailing in estuarine or marine environments, underscoring the importance of the inclusion of biogeochemical parameters in this research.

3.2. Dechlorination Catalysis in Biotic and Abiotic Model Systems

3.2.1. Dechlorination Yields Based on literature reports on the reactivity of structurally similar compounds (e.g., PCBs) in sediments, dechlorination reactions are presumably the dominant pathway in the fate of these compounds under reducing conditions (7). Published studies from our laboratory on the fate of PCDD in anaerobic sediments have indicated, surprisingly, that dechlorination products accounted for a minor fraction (less than 20%) of the total reactivity, based on the disappearance of parent compound (7, 8, 12, 41, 43). The remainder of products observed, but not quantified or identified, provided evidence of transchlorination (migration of chlorine from PCDD to organic matter), or polymerization. Estuarine (12) and freshwater (5,18) sediment-derived microorganisms, humic acid constituents (12,40), biogenic macromolecules, and heavy metals (1) have been found to catalyze dechlorination activities. Microbially mediated yields were strongly dependent on the prevailing biogeochemical conditions; estuarine conditions resulted in the highest relative yield (normalized to parent compound concentration) (Figure 13.3). It should be noted that in these studies, freshwater samples were spiked with while the estuarine and marine samples received (both isomers). The yields of the DOM-mediated (monomer and polymer) reactions (2 to 20%) were similar in magnitude to those observed for microbial cells alone (6.5%). DOM-mediated yields normalized to the available phenolic acidity indicated differences in dechlorination efficiencies between humics (42). These results suggest that electron shuttling from an electron donor to an acceptor may also depend on structural differences in humic materials, such as percent aromaticity (95) and hydrophobic interactions with the helical cavity of the polymers (96).

3.2.2. Dechlorination Pathways Two distinct pathways have been observed during an investigation of PCB dechlorination (107). Microorganisms which survived heat and ethanol treatments preferentially removed meta chlorines, while non-thermo- and non-ethanol-resistant subpopulations mainly contributed to the observed para-dechlorination activity. Similar results were obtained using PCDD-spiked microbial communities eluted from estuarine Passaic River sediment cores. Here, dechlorination proceeded via two concurrent pathways :peri-dechlorination (removal of a 1,4,6,9-chlorine) that resulted in transient formation of and mixed peri-lateral dechlorination for non-2,3,7,8substituted congeners (Figure 13.4). Results suggested that peri-lateral dechlorination represented the result of microbially mediated activity, while the mixed peri-lateral pathway represented combined biotic and abiotic activities (12).

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The magnitude of the 2,3,7,8-PCDD to non-2,3,7,8-PCDD ratio reflects the dominance of the peri-dechlorination (ratio greater than 1), or mixed peri-lateral dechlorination (ratio less than 1). In this study, the majority of samples (biotic and abiotic) resulted in a ratio less than 1, indicating an overall preference for the mixed peri-lateral (2,3,7,8) dechlorination pathway. Thermodynamic calculations determined that the most favorable pathway to occur was at the most positively charged carbon atom in the ring, typically a lateral carbon (2,3,7, or 8 position) (60). Production of 2,3,7,8increased with decreasing salinity. The magnitude ofthe 2,3,7,8-PCDD to non2,3,7,8-PCDD ratio was significantly higher in the presence of DOM, suggesting a shift in the reaction towards peri-dechlorination (Figure 13.3). This strong fractionation ratio in DOM or DOM-microbial incubations is similar to that observed in sediment systems (7), where only the production of was observed. Further dechlorination of 2,3,7,8- or isomers to has been demonstrated in spiked (11, 18) and historically contaminated (12) environmental systems containing microorganisms. These were naturally-occurring (11, 12) and

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enriched or hexachlorobenzene-acclimated cultures (18). In general, the endpoint of microbially-mediated dechlorination was whereas the organic matter-mediated reactions ceased at (41,42). Therefore, abiotic or biotic dechlorination processes may be detected based on isomer-specific ‘fingerprinting’ ofthe dechlorination pattern. These findings also suggest that biotic and abiotic transformation processes in sediments may account for the distinction between PCDD source and sink patterns (Figure 13.5).

3.3. Effects of Chemical inducers on Microbial PCDD Dechlorination Activity

3.3.1. Structurally-Analogous Compounds as Primers for Dechlorination PCB dechlorination in historically contaminated freshwater sediments has been stimulated using structurally analogous compounds, such as chloro-and bromobiphenyls, and bromobenzoates (14-16). These compounds appear to act as a preferred dechlorination substrate and electron acceptor, effectively “priming” dechlorination (1416). Similar results were obtained upon amendment with a structurally analogous compound (2-monobromodioxin, or MBrDD) to sediment-free estuarine incubations spiked with PCDD and historically contaminated sediments. Primer addition resulted in a moderate increase in relative yield (Figure 13.6a) (43), and a corresponding preference for the mixed peri-lateral dechlorination pathway (ratio less than1) (Figure 13.6b). Molecular analyses (denaturing gradient gel electrophoresis) revealed significant shifts in community structure as a result of the MBrDD amendments, independent of the

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spiked dioxin congeners and biogeochemical conditions. This result suggests that changes in dechlorination patterns and yields were due to the priming (or analogue enrichment) of populations with distinct dechlorinating capabilities. However, the responsible populations have not been identified to date.

3.3.2. Hydrogen-Enhanced Microbial Dechlorination Activity There are a limited number of studies describing the use of hydrogen to stimulate anaerobic dechlorination. Organic acids (which ferment to hydrogen) have stimulated reductive dechlorination in a benzoate-acclimated dehalogenating mixed culture (106). Also, Newell et al. (73) reported on the complete dechlorination of chlorinated solvents to ethene in aquifer material amended with hydrogen gas. No mechanistic information or correlations to respiratory activity were inferred. Hydrogen amendments in freshwater and estuarine sediments contaminated with dioxins resulted in extensive dechlorination

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of to at the expense of a decrease in production. In addition, dechlorination occurred at accelerated (200-fold) rates (6, 7). Because anaerobic inorganic electron acceptors are generally found in excess in sediments and respiratory activities coexist, Kerner (53), and Postma and Jakobson (81), have argued that key metabolites, such as hydrogen and acetate that are produced from fermentation, may be rate-limiting for microbially-mediated reduction reactions in sediments. These findings suggest hydrogen amendment as a potential strategy for stimulating PCDD dechlorination to without the accumulation of in contaminated sediments.

3.4. Dechlorination Kinetics in Biotic, Abiotic, and Sediment Systems Since disappearance of PCDD congeners is not limited to dechlorination reactions, dechlorination kinetics are generally based on product appearance. Productionrates have been accurately determined, based on isomer-specific analysis. However, the calculated dechlorination rates are uncertain due to a lack of commercially available and isomers for product identification. These lesser-chlorinated congeners may be derived from the dechlorination of any isomer. Production rates of the various dioxin homologue group model systems generally decrease with subsequent dechlorination. This observation corresponds to thermodynamic predictions, in that both the Gibbs free energy for dechlorination and the HOMO-LUMO gap generally decrease with decreasing levels of chlorination (60). Production rates of lesser-chlorinated congeners due to dechlorination range from 0.05 to 0.90 nanograms per day (ng/d) in sediment-free systems and 0.01 to 17.29 ng/d in sediment systems with varied amendments (7). In abiotic reactions, the time-dependent appearance of lesser-chlorinated products as a function of incubation conditions shows an increase in all congener groups with time, whereas dechlorination in cell-mediated incubations has resulted in the sequential production of lesser-chlorinated congeners (41). No information is available on the concentration-dependent dechlorination of single dioxin congeners, but studies with PCBs indicated that the sequential removal of chlorine atoms prevailed over the concentration range studied (16). The addition ofDOM (e.g., 3,4-dihydroxybenzoic acid [DHBA], Aldrich humic acid, polymaleic acid) generally resulted in an increase in the production rate of (40). A 3,4-DHBA amendment to abiotic systems resulted in increased production rates of congeners (10 times higher), when compared to cell-mediated reactions. The presence of an electron donor (titanium citrate) resulted in an order of magnitude increase in the production rates of all PCDD congeners measured. In sediment systems amended with organic acid and MBrDD, the 1,2,3,4,6,7,8and isomers comprised a 50 to 70% disappearance of an indication that dechlorination reactions of PCDD transformation predominate (7) (Figure 13.7). Amendment with MBrDD resulted in enhanced dechlorination (transformation rate = 0.13 ng/d), and a corresponding increase in the production rate of (0.84 ng/d). From these studies, the production rates for dechlorinated products from farexceeded those from in organic acid amended systems, suggesting that dechlorination may be rate-limiting. To the contrary, the

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rate of production in the hydrogen-amended system was two orders of magnitude higher than the combined disappearance of other measured congeners. The relative rates of production from and dechlorination to are of particular interest due to toxicity concerns (Figure 13.7). Except in the cases of spiked cells under freshwater conditions and hydrogen-stimulated sediments, the isomer is likely to accumulate or remain at steady state concentrations during dechlorination reactions in sediments ( and dominated residues). Extrapolation of the rates at which these dechlorination and detoxification reactions are likely to occur in the environment is a difficult task. Some assumptions need to be made with respect to the dominant catalysts of dechlorination activity in sediments, the bioavailability of the contaminant, and the electron fluxes in reduced sediments. Based on the assumptions that, (i) the dechlorination reaction is cometabolic, with an approximate ratio of 1:1000, chlorine removal to methane/sulfate generation, from Barkovskii and Adriaens (12) and Fu et al. (42), and (ii) the reaction is coupled to either methanogenesis or sulfate-reduction, sediment dioxin reaction rates would be on the order of 5 to 10 years per chlorine removed. This corresponds well to half-lives reported for dioxins (approximately 2 to 10 years in aquatic sediments) in the literature (61).

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4. IMPLICATIONS FOR NATURAL ATTENUATION AND REMEDIAL STRATEGIES 4.1. Natural Dioxin Cycling Studies of PCDD distribution in archived and contemporary sediment and soil samples (from e.g., Lake Ketelmeer, The Netherlands and Broadbalk, England), including sites primarily impacted by atmospheric deposition rather than anthropogenic activity, indicate a preferential accumulation of (18). Atmospheric deposition residues are dominated by and (9), which suggests a discrepancy between source and sink profiles, thereby indicating processes other than emissions may have impacted the sink patterns. Beurskens et al. (18) concluded that there was evidence of post-depositional, microbially mediated dechlorination of some of the higher chlorinated PCDDs. The peer-reviewed literature provides abundant evidence for the dechlorination of PCDD at environmentally relevant concentrations. Dechlorinating activities were catalyzed by estuarine (7, 12) and freshwater (5, 18) sediment-derived microorganisms, humic acids (13,41), biogenic macromolecules, and heavy metals (1). Although natural attenuation of dioxins was not directly measured in the series of studies presented here, it is apparent that dioxins can undergo extensive transformation reactions in sediments, with dechlorination processes representing up to 20% of total reactivity. For example, the accumulation of during the incubation of PCDDspiked and historically contaminated sediments correspond to profiles observed at the air-water interface of the Passaic River; fugacity calculations have implied a net flux from the water body, indicating that sediments may serve as a PCDD source (57). Also, some core samples have shown evidence for the presence of but the source of this isomer was not determined (8). Moreover, a decrease in and and the production of (25 mol%) were observed in sediment cores incubated in the presence of hydrogen gas, where the original PCDD residues did not include Microbially-mediated dechlorination of historical residues to has also been observed in earlier studies (12). These results suggest that sediment reactivity may be a natural attenuation mechanism. 4.2. Dioxin-lmpacted Sediment Management Strategies The cleanup of PCDD-contaminated sediments poses some of the most challenging issues for site remediation in regional and national freshwater and estuarine coastal environments, due to point-source releases and diffusive contaminant sources. An estimated 14 to 28 million cubic yards (11 to 22 million cubic meters) of contaminated sediment must be managed annually (71). Whereas volumes of contaminated sediments tend to be large, contaminant concentrations are low, such that application of expensive conventional control technologies. For example, incineration or disposal in a secure landfill, cannot be justified. When removal is necessary (e.g., to maintain navigational channels) or feasible, on-land treatment or disposal options may be ineffective or costprohibitive. Under such circumstances, natural or enhanced recovery methods may provide an ideal alternative for environmental cleanup and the reduction of human and

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ecological risks from sediment contamination. Natural recovery includes naturallyoccurring environmental processes, such as sorption, sequestration and biodegradation (36). Enhanced recovery processes are those that involve manipulation strategies designed to stimulate natural fate pathways. Both in situ and ex situ sediment remediation techniques have been developed and some (mainly ex situ) are extensively applied in practice (93). Ex situ applications for toxicity reduction typically involve a volume reduction technique, such as particle separation to separate sand from contaminated fines (84), followed by chemical, thermal (89), or biological (36) treatment methods that can be contained in landfills, landfarms, or reactors. In situ applications are limited to a few pilot demonstrations of enhanced bioremediation, with application to either the aerobic (46), or anaerobic (16) degradation of PCBs and PAHs (33), or immobilization and chemical treatment (71). Biostimulation of PCB dechlorination reactions has been achieved by adding and mixing brominated biphenyl analogues and nutrients in sediments. Alternatively, powdered products containing oxygen carriers, nutrients, and bacteria have been spread over surface waters and allowed to sink to the bottom, to stimulate in situ PAH degradation (36). Experience with in situ bioremediation technologies, in land-based soils or sediments and groundwater systems, have limited potential for transfer to freshwater or marine sediments due to the dearth of knowledge on the degradative capabilities of microbial populations in these environments. Evidence from the groundwater literature and our previous sediment research, however, indicates a role for hydrogen as a suitable driving force for dechlorination reactions in these environments. Due to competition for hydrogen in anaerobic environments, differences in hydrogen concentrations at equilibrium, resulting from fermentation and direct additions, are expected to affect the extent and kinetics ofdechlorination. For example, complete fatty acid turnover and low residual hydrogen concentrations have been observed under saline conditions (3 to 17 ppt). In these studies, metabolic endpoints did not infer any correlation between respiratory and dechlorination activity (43), but our previous research has demonstrated extensive reduction of to without the accumulation of in hydrogen-amended sediment systems (7). A conceptual illustration ofcontainment, disposal, and natural recovery technologies that incorporate our current understanding of biogeochemical effects on dioxin dechlorination is shown in Figure 13.8 (modified from reference 71). Natural recovery involves leaving the contaminated sediments in place, allowing on-going aquatic processes to contain (e.g., burial by cleaner sediments), destroy, or otherwise reduce the bioavailability of the contaminants. The applicability of these processes is generally limited in sediment beds offlowing streams, high-energy coastal zones, or sites requiring navigational dredging. Due to the unknown potential for contaminant destruction, this has only been applied to a few cases (51,105). Even though natural in situ dechlorination processes in sediments have been demonstrated for compounds such as PCBs (16,21,40), and DDT (85), it is unclear whether these processes are sufficiently effective for risk mitigation. A central limitation to the decision-making framework for (monitored and verifiable) natural recovery is the lack of knowledge on natural transformation rates vs. chemical release rates from the sediment bed (108). In particular, the latter has been the main motivation behind in situ or ex situ capping, where an isolating material cover (e.g., sand, clay, uncontaminated

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sediment, geosynthetic materials) is placed over contaminated sediments (78, 79). Capping has become one of the few accepted management techniques for contaminated sediments, despite a dearth of information on long-term chemical fluxes into the overlying water column, verification methods for cap thickness, and available monitoring data on cap integrity. From a regulatory perspective, capping is not currently considered a permanent solution, but rather a “preferred remedy status”, reflecting technologies geared at toxicity reduction (71). Indeed, capped sediments may lend themselves for further innovative technology applications, such as nutrients and gas (hydrogen) amendments to stimulate biodegradation (Figure 13.8).

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Containment of Contaminated Sediments (pp 289-296) American Society of Testing and Materials, ASTM STP 1293, Philadelphia 80. Peijnenburg WJGM, t’Hart MJ, den Hollander HA, van de Meent D, Verboom HH & Wolfe NL (1992) QSARs for predicting reductive transformation rate constants of halogenated aromatic hydrocarbons in anoxic sediment systems. Environ. Toxicol. Chem. 11:310-314 81. Postma D & Jakobsen R (1996) Redox zonation: Equilibrium constraints on the reduction interface. Geochim. Cosmochim. Acta 60:3169-3175 82. Praeger TH, Messur SD & DiFiore RP (1996) Remediation of PCB-containing sediments using surface water diversion ‘dry excavation’: A case study. Water Sci.Technol. 33:239-245 83. Prieur D, Troussellier M, Romana A, Chamroux S, Mevel G & Baleux B (1987) Evolution of bacterial communities in the Gironde estuary (France) according to a salinity gradient Estuar. Coast. Shelf Sci. 24:95-108 84. Pruijn MF, van Winden SC & Wevers HHAG (1998) From characterization of sediment to full scale treatment by wet particle separation. Water Sci.Technol. 37:189-198 85. Quensen JF, Mueller SA, Jain MK & Tiedje JM (1998) Reductive dechlorination of DDE to DDMU in marine sediment microcosms. Science 280:722-724 86. Rappe C, Berquist P-A, Kjeller L-O, Swanson SE, Belton T, Rupper B, Lockwood K & Kahn PC (1991) Levels and patterns of PCDD and PCDF in fish, crabs, and lobsters from Newark Bay and New York Bight. Chemosphere 22:239266 87. Rappe C, Andersson R, Berquist P-A, Brohede C, Hansson M, Kjeller L-O, Lindstrom G, Marklund S, Nygren M, Swanson SE, Tysklind M & Wiberg K (1987) Overview on environmental fate of chlorinated dioxins and dibenzofurans: Sources, levels and isomeric patterns in various matrices. Chemosphere 16:1603-1618 88. Rappe C, Choudhary G & Keith LH (1986) Chlorinated Dioxins and Dibenzofurans in Perspective. Lewis Publishers, Inc. Chelsea, MI 89. Rienks J (1998) Comparison of results for chemical and thermal treatment of contaminated dredged sediments. Water Sci. Technol. 37:355362 90. Rordorf BF (1986) Thermal properties of dioxins, furans, and related compounds. Chemosphere 15:1325-1332 91. Rordorf BF (1987) Prediction of vapor pressure, boiling points, and enthalpies of fusion for twenty-nine halogenated dibenzo-p-dioxins.

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372 Thermochim. Acta 112:117-122 92. Rordorf BF (1989) Prediction of vapor pressures, boiling points and enthalpies of fusion for twenty-nine halogenated dibenzo-p-dioxins and fifty-five dibenzofurans by a vapor pressure correlation method. Chemosphere 18:783-788 93. Rulkens WH, Tichy R & Grotenhuis JTC (1998) Remediation of polluted soil and sediment: Perspectives and failures. Water Sci. Technol. 37:27-35 94. Schwarzenbach RP, Stierly R, Lanz K & Keyer J (1990) Quinone and iron porphyrin mediated reduction of nitroaromatic compounds in homogeneous aqueous solution. Environ. Sci. Technol. 24:1566-1574 95. Scott DT, McKnight DM, Blunt-Harris EL, Kolesar SE & Lovely DR (1998) Quinone moieties act as electron acceptors in the reduction of humic substances by humicsreducing microorganisms. Environ. Sci. Technol. 198:2984-2989 96. Sein LT, Varnum JM & Jensen SA (1999) Conformational modeling of a new building block of humic acid: Approaches to the lowest energy conformer. Environ. Sci. Technol. 33: 546-552 97. Shiu WY, Doucette W, Gobas F, Andren A & Mackay D (1988) Physical-chemical properties of chlorinated dibenzo-p-dioxins. Environ. Sci. Technol. 22: 651-658 98. Skyring GW (1987) Sulfate reduction in coastal ecosystems. Geomicrobiol. J. 5:295-374 99. Suflita JM & Townsend GT (1995) The microbial ecology and physiology of aryl dehalogenation reactions and implications for bioremediation. In: Young LY & Cerniglia C (Eds) Mircrobiological Transformation and Degradation of Toxic Organic Chemicals (pp 243-268) Wiley-Liss, New York, NY 100. Tong HY, Monson SJ, Gross ML, Bopp RF, Simpson HJ, Deck BL & Moser FC (1990) Analysis of dated sediment samples from the Newark Bay area for selected PCDD/Fs.

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Chapter 14 REDOX CONDITIONS AND THE REDUCTIVE/OXIDATIVE BIODEGRADATION OF CHLORINATED ETHENES IN GROUNDWATER SYSTEMS FRANCIS H. CHAPELLE AND PAUL M. BRADLEY Microbial Studies Group, U.S. Geological Survey, Columbia, SC, USA

1. INTRODUCTION Chlorinated ethenes are subject to a variety of microbial degradation processes that include reductive dechlorination (26), aerobic oxidation (12, 16), anaerobic oxidation (3), and aerobic cometabolism (21, 28). Because of this metabolic variety, assessing the relative efficiency of chlorinated ethene biodegradation in groundwater systems is complex. The conditions under which these biodegradative processes occur have now been well-documented, and it is often possible to assess the potential contribution of particular degradation processes at a given site. The purpose of this chapter is to illustrate how the documentation and assessment of ambient redox conditions can help to determine reductive and oxidative biodegradation processes occurring in groundwater systems.

1.1. Reductive and Oxidative Mechanisms for Chlorinated Ethene Biodegradation In the early 1980s a number of reports described the accumulation of daughter products from highly chlorinated ethene biotransformation, such as cis-dichloroethene (cis-DCE) and vinyl chloride (VC), in anaerobic groundwater systems. At first, this phenomenon was considered to be cometabolic in nature, with reduction of the primary chlorinated contaminants occurring due to the gratuitous interaction of reduced microbial cofactors with the chlorinated ethenes. Later, it became evident that some microorganisms could use chlorinated ethenes as electron acceptors in their respiration. The consensus today is that microbial reductive dechlorination processes are ubiquitous in anaerobic, chloroethene-contaminated aquifers, but that the extent of dechlorination is highly variable from site to site (2, 10, 15, 20, 21, 26). Lightly chlorinated ethenes such DCE and VC are also subject to oxidative, as well Dehalogenation: Microbial Processes and Environmental Applications, pages 373-384 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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as reductive biodegradation processes. Rapid microbial degradation of VC, including mineralization, has been observed in laboratory cultures and aquifer samples under aerobic conditions (3, 12). Moreover, under aerobic conditions, VC can be used as a sole carbon source for growth and metabolism (16). DCE has also been shown to oxidize under aerobic conditions in liquid culture, and this activity may support microbial growth (7; J.M. Gossett, personal communication). Because the production ofDCE and VC generally occurs by reductive dechlorination ofmore highly chlorinated ethenes under anaerobic conditions, the aerobic oxidation of these less chlorinated compounds is often limited in groundwater systems. However, where anaerobic conditions that produce DCE and VC grade to less reduced, more oxic conditions, as often occurs on the fringes of a contaminant plume, aerobic oxidation of these compounds can be significant. An oxygen source added to groundwater systems provides a strategy to stimulate the degradation of the less chlorinated analogs, i.e., VC and DCE. Anaerobic oxidation of DCE and VC is a process that has only been recognized recently (3). Given the early emphasis on reductive dechlorination under anaerobic conditions as a principal degradation pathway, the fact that these compounds also oxidize anaerobically was unexpected. In a series of papers, it was shown that VC, and to a lesser extent DCE, could be oxidized to carbon dioxide under Fe(III) reducing conditions (3-5). The fact that Fe(III) reduction could support the anaerobic oxidation of VC was initially attributed to Fe(III) being a fairly strong oxidant, and may thereby catalyze abiotic transformations. Further investigation with substrates showed that a portion of both and was mineralized to under a variety of

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anoxic conditions which represent a broad redox spectrum, including Fe(III)-reducing conditions, sulfate-reducing conditions, and even methanogenic conditions (Figure 14.1). In addition to inorganic electron acceptors, it was shown that organic compounds, such as humic acids could serve as an electron acceptor for VC and DCE oxidation (4). Clearly, the simple explanation that a strong oxidant such as Fe(III) was solely responsible for anaerobic oxidation could not account for observed oxidation under methanogenic conditions (7). Studies reporting that was an intermediate of mineralization under methanogenic conditions (8,9; Figure 14.2) suggested that the actual degradation of VC may not be directly mediated by any particular terminal electron accepting process, but rather by fermentative, acetogenic microorganisms. This suggestion was later confirmed by the observed accumulation of from when methanogenesis and reductive dechlorination were blocked by addition of BES to the microcosms (9). Acetogens are known to metabolize a wide variety of organic compounds while producing acetate as a waste product (14, 22). The conceptual model proposed in Figure 14.3 depicts transformation of VC to acetate by acetogenic populations; the acetate product is readily oxidized to by a wide variety of terminal electron accepting processes. The model is consistent with the observation that the rate of anoxic VC mineralization decreases with the oxidizing potential of the electron acceptor. Because Fe(III)-reducing bacteria have a higher affinity for acetate than methanogens, they maintain lower concentrations of acetate. This, in turn, allows acetogenic microorganisms to produce acetate (their waste product) at a faster rate. This in turn, is a possible explanation for the faster production observed under progressively

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more oxidizing conditions (Figure 14.1). The observed presence of acetate as an intermediate in VC mineralization does not rule out the possibility that VC can also be directly oxidized to A direct oxidation would also be consistent with the pattern shown in Figure 14.1. It is now clear that VC can be degraded by both reductive dechlorination and by acetogenic fermentation under anaerobic conditions. Because the efficiency ofthese processes varies with redox conditions, it is clear that the assessment of ambient redox conditions is key to understanding the behavior of chlorinated ethenes in groundwater systems.

1.2. Degradation of Chlorinated Ethenes in Groundwater Systems Although microbial degradation processes of chlorinated ethenes are complex, they are to some extent predictable. Under highly reducing conditions, e.g., acetogenesis or methanogenesis, all chlorinated ethenes can serve as electron acceptors in microbial metabolism. In this case, microorganisms reduce chlorinated ethenes by transferring electrons to them. Under other redox conditions, microorganisms can use less chlorinated ethenes such as VC as electron donors, resulting in a net oxidation of the substrate. Under aerobic conditions, this appears to be a direct oxidation process. However, under anoxic conditions, oxidation can proceed with acetogenic microorganisms (and possibly other fermentative microorganisms) playing an intermediate role (Figure 14.3). Under anoxic conditions, highly chlorinated ethenes such as tetra- or perchloroethene (PCE) and trichloroethene (TCE) are subject to reductive dechlorination according to the sequence:

Here, the efficiency of dechlorination varies under different redox conditions and degree of chlorination. Dechlorination of PCE and TCE to DCE occurs under mildly reducing nitrate- or Fe(III)-reducing conditions, whereas efficient transformation of DCE to VC,

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or VC to ethene requires the more strongly reducing conditions of methanogenesis or sulfate reduction (25). Results from laboratory studies (13) indicate that reductive dechlorination can be driven by molecular hydrogen. If this process is widespread in nature, it follows that the efficiency of reductive dechlorination is related to the availability of Concentrations of in groundwater systems are controlled by ambient microbial terminal electron-accepting processes (TEAPs). Under anaerobic conditions, is continuously produced by fermentative microorganisms metabolizing natural or anthropogenic organic matter. The generated in situ can be used by respiratory microorganisms that most commonly use Fe(III), sulfate, or as terminal electron acceptors. Significantly, each of these TEAPs has a different affinity for uptake (18). reduction (methanogenesis) has the lowest affinity, and steady-state concentrations in methanogenic aquifers are therefore relatively high (approximately 10 nM). Sulfate reduction has a slightly greater affinity for than methanogenesis, and is characterized by lower concentrations (1-4 nM). Fe(III) reduction and denitrification have even greater affinitites for (0.2-0.8 nM and <0.10 nM, respectively), and are characterized by progressively lower steady-state concentrations (Table 14.1). Since chlororespiration becomes hydrogen-limited at concentrations characteristic of Mn(IV)-reduction and Fe(III) reduction, it is less efficient under these conditions than under methanogenic or sulfate-reducing conditions where concentrations are characteristically higher. In most groundwater systems, biodegradation of chlorinated ethenes proceeds via both reductive and oxidative pathways. In fact, overall biodegradation in groundwater systems is favored by the transition from relatively reducing to relatively oxidizing conditions. Under reducing conditions, PCE and TCE may be transformed to DCE and VC. Then, as groundwater migrates to zones dominated by Fe(III) reduction or oxygen reduction, DCE and VC can be further oxidized to carbon dioxide:

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The efficiency with which chlorinated ethenes are completely degraded, therefore, depends on the prevailing redox conditions in an aquifer, and on the redox gradient that chlorinated ethenes are exposed to along particular groundwater flowpaths. Thus, an

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important component to evaluating the biodegradation of chlorinated ethenes in groundwater systems is an assessment of ambient redox conditions.

1.3. Assessment of Redox Conditions in Groundwater Systems Identifying the distribution of TEAPs in groundwater systems is often difficult. This is in large part due to the inherently heterogeneous lithologic and geochemical nature of groundwater systems (11). Nevertheless, because particular TEAPs require the presence of certain electron acceptors e.g., sulfate or Fe(III), and because particular TEAPs produce characteristic final products, e.g., methane from methanogenesis, sulfide from sulfate reduction, or Fe(II) from Fe(III) reduction, the concentration trends of potential electron acceptors and final products through time and/or space can be used to deduce ongoing TEAPs. Similarly, concentrations of intermediate products such as can provide insight into the spatial and temporal distribution of TEAPs. A schematic diagram showing the logic behind this methodology is shown in Figure 14.4. Using this flowchart as a guide, if, for example, sulfate concentrations are observed to decrease with time or along a flowpath segment of a ground-water system, sulfide concentrations are observed to increase, and hydrogen concentrations are observed to be in the range characteristic of sulfate reduction, the occurrence of sulfate reduction can be identified in that portion of the aquifer with a high degree of confidence. In practice, however, this methodology often encounters uncertainties. For example, if methane, sulfide, and Fe(II) concentrations are observed to increase along a flowpath segment, or with time, it can be concluded that methanogenesis, sulfate reduction, and Fe(III) reduction are ongoing (1). Because methane, sulfide, and Fe(II) are actively transported by flowing ground water, it is often difficult to determine the spatial location of each redox zone. If methane is present in ground water at a particular well location, it cannot be concluded that methanogenesis is occurring within the screened intervals of the well. Rather, it can only be concluded that methanogenesis is occurring upgradient of the well. In these cases, concentrations of , continuously cycled by microorganisms and thus not subject to significant advective transport, may more accurately identify redox zones. However, it is always possible that wells can be screened across different redox zones, leading to intermediate, non-diagnostic concentrations of Moreover, redox zones shift with time and space in response to recharge events (27), the availability of organic matter, or other environmental factors. Because of this observed variability, the term “predominant TEAPs”, rather than “exclusive TEAPs”, is appropriate and must always be considered when following the methodology outlined in Figure 14.4.

2. REDOX CONDITIONS AND THE BIODEGRADATION OF CHLORINATED ETHENES IN GROUNDWATER SYSTEMS The influence of differing redox conditions on the biodegradation behavior of chlorinated ethenes can be illustrated by observing conditions at two different sites. One site, Naval Air Station Cecil Field, in Florida, is characterized by relatively low oxidizing conditions in the contaminant source area, followed by progressively more oxidizing conditions downgradient. The other site, Naval Submarine Base Kings Bay,

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in Georgia, is characterized by reducing conditions in the source area, followed by more oxidizing conditions downgradient. Because of these differing conditions, the biodegradation behavior of chlorinated ethenes is much different at each of these sites, even though their hydrology is quite similar. Concentrations of chlorinated ethenes, and concentrations of redox-sensitive water chemistry parameters for the Cecil Field site are shown in Figure 14.5. The plume is characterized by relatively high concentrations of TCE in and near the source area, that persist at least 400 meters downgradient. As groundwater flows along the hydrologic gradient, concentrations of iron (~6 mg/L) are relatively high and (~0.5 nM) is in the range characteristic of Fe(III) reduction and chlororespiration (Table 14.1). In addition,

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sulfate concentrations decrease and concentrations of sulfide increase slightly, indicating that sulfate-reduction captures part of the electron flow in this part of the system. Beyond 400 meters along the flowpath, sulfide concentrations increase sharply, and concentrations rise into the range characteristic of sulfate reduction. This indicates that sulfate reduction becomes the predominant electron-accepting process downgradient. The data from Cecil Field indicate that reductive dechlorination of TCE occurs under the Fe(III)-reducing conditions that are predominant near the source area, resulting in the formation of cis-DCE. Significantly, however, VC, the dechlorination product of cis-

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DCE is not present. This indicates that either VC is not being formed, or that it is degrading faster than it is being produced so that it does not accumulate in solution. Downgradient, where sulfate reduction is more in evidence, contaminant concentrations decrease below measurable levels. The relatively recalcitrant behavior of chlorinated ethenes observed at the Cecil Field site (Figure 14.5) is much different than is observed in other hydrologic systems. For example, at the Kings Bay site (Figure 14.6), concentrations of the highly chlorinated ethenes PCE and TCE degrade rapidly and drop below measurable levels by 80 meters downgradient. In the source area, concentrations of sulfate are relatively low, concentrations of sulfide are relatively high, and concentrations are in the range characteristic of sulfate reduction. These observations indicate that sulfate reduction sequesters the majority of electron flow in the source area. Downgradient, however, concentrations of sulfate increase, concentrations of sulfide decrease, concentrations of iron remain relatively high, and concentrations are in the range characteristic of Fe(III) reduction and chlororespiration (Table 14.1). The data from the Kings Bay site indicate that reductive dechlorination of PCE and TCE occurs rapidly under the sulfatereducing conditions that predominate at the contaminant source area. Downgradient, the equally rapid decrease in cis-DCE and VC indicates rapid biodegradation under the more oxidizing conditions. The hydrology of the Cecil Field and Kings Bay sites is similar, both being aquifers of marginal marine origin, of similar age, and both with groundwater flow rates on the order of 0.16 meters per day. The behavior of chlorinated ethenes, however, is markedly different between the sites. At Cecil Field, high concentrations of TCE extend at least 400 meters downgradient. In contrast, at Kings Bay, PCE and TCE do not extend beyond 100 meters downgradient. The observed difference in the biodegradation profiles for chlorinated ethenes at these two sites reflects a succession of redox processes. At Kings Bay, efficient PCE and TCE biodegradation reflects the higher efficiency of reductive dechlorination, at concentrations characteristic of sulfate reduction (15) near the source area, followed by the oxidation of VC under Fe(III)reducing conditions downgradient. In contrast, the relatively low concentrations, which are characteristic of Fe(III) reduction, result in lower rates of reductive dechlorination near the source area at the Cecil Field site, and the relative persistence of cis-DCE in much of this aquifer.

3. CONCLUSIONS Chlorinated ethenes are subject to a variety of biodegradation processes including chlororespiration, cometabolic reductive dechlorination, fermentative acetogenesis,aerobic and anaerobic oxidation, and aerobic cometabolism. Due to this metabolic diversity, the behavior of chlorinated ethenes in groundwater systems is often puzzling. The rate and extent of these biodegradation processes depends on predominant redox conditions and on the succession of predominant redox conditions. Thus, an assessment of ambient redox processes can help to explain the fate of chlorinated ethenes in different hydrologic systems. By careful evaluation of the redox conditions in groundwater systems, a clearer understanding of the biodegradation and natural attenuation of chlorinated ethenes in the environment can be achieved.

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REFERENCES 1. Baedecker MJ, Cozzarelli IM, Eganhouse RP, Siegel DI & Bennett PC (1993) Crude oil in a shallow sand and gravel aquifer, III. Biogeochemical reactions and mass balance modeling in anoxic ground water. Appl. Geochem. 8:569-586 2. Bouwer EJ (1994) Bioremediation of chlorinated solvents using alternate electron acceptors. In: Norris RD et al. (Eds) Handbook of Bioremediation (pp 149-175). Lewis Publ., Boca Raton 3. Bradley PM & Chapelle FH (1996) Anaerobic mineralization of vinyl chloride in Fe(III)reducing, aquifer sediments. Environ. Sci. Technol. 30:2084-2086 4. Bradley PM & Chapelle FH (1998) Microbial mineralization of VC and DCE under different terminal electron accepting conditions. Anaerobe 4:81-87 5. Bradley PM, Chapelle FH & Lovley DR (1998) Humic acids as electron acceptors for anaerobic microbial oxidation of vinyl chloride and dichloroethene. Appl. Environ. Microbiol. 64:3102-3105 6. Bradley PM, Landmeyer JE & Dinicola RS (1998) Anaerobic oxidation of [1,2-14C] dichloroethene under Mn(IV)-reducing conditions. Appl. Environ. Microbiol. 64:15601562 7. Bradley PM & Chapelle FH (1999) Methane as a product of chloroethene biodegradation under methanogenic conditions. Environ. Sci. Technol. 33:653-656 8. Bradley PM & Chapelle FH (1999) Role for acetotrophic methanogens in methanogenic biodegradation of vinyl chloride. Environ. Sci. Technol. 33:3473-3476 9. Bradley PM & Chapelle FH (2000) Acetogenic microbial degradation of vinyl chloride. Environ. Sci. Technol. 34:2761-2763 10. Chapelle FH (1996) Identifying redox conditions

(1991) Reductive dechlorination of high concentrations of tetrachloroethene to ethene by an anaerobic enrichment culture in the absence of methanogenesis. Appl. Environ. Microbiol. 57:2287-2292 14. Drake HL (1994) Acetogenesis, acetogenic bacteria, and the acetyl-CoA “Wood/Ljungdahl” pathway: Past and current perspectives. In: Drake HL (Ed.) Acetogenesis (pp 3-62) Chapman and Hall, New York 15. Gossett JM & Zinder SH (1996) Microbiological aspects relevant to natural attenuation of chlorinated ethenes. In: Symposium on Natural Attenuation of Chlorinated Organics in Ground Water (pp 10-13) EPA/540/R-96/509

16. Hartmans S (1995) Microbial degradation of vinyl chloride. In: Singh VP (Ed.) Biotransformations: Microbial Degradation of Health Risk Compounds (pp 239-248) Elsevier Science, Amsterdam 17. Löffler FE, Tiedje JM & Sanford RA (1999) Fraction of electrons consumed in electron acceptor reduction and hydrogen thresholds as indicators of halorespiratory physiology. Appl. Environ. Microbiol. 65:4049-4056 18. Lovley DR & Goodwin S (1988) Hydrogen concentrations as an indicator of the predominant terminal electron-accepting reactions in aquatic sediments. Geochim.Cosmochim. Acta 52:2993-3003 19. Maymo-Gatell X, Chien Y-T, Gossett JM & Zinder S H (1997) Isolation of a bacterium that reductively dechlorinates tetrachloroethene to ethene. Science. 276:1568-1571 20. McCarty PL (1996) Biotic and abiotic transformations of chlorinated solvents in groundwater. In: Symposium on Natural Attenuation of Chlorinated Organics in Ground Water (pp 5-9) EPA/540/R-96/509 21. McCarty PL & Semprini L (1994) Groundwater treatment for chlorinated solvents. In: Norris RD et al. (Eds.) Handbook of Bioremediation (pp that favor the natural attenuation of chlorinated 87-116). Lewis Publ., Boca Raton ethenes in contaminated groundwater systems. In: Symposium on Natural Attenuation of 22. Schink B (1994) Diversity, ecology, and isolation of acetogenic bacteria. In: Drake Chlorinated Organics in Ground Water (pp 17HL(Ed.) Acetogenesis (pp 197-235) Chapman & 20) EPA/540/R-96/509 Hall, New York 11. Cozzarelli IM, Herman JS, Baedecker MJ & Fischer JM (1999) Geochemical heterogeneity of 23. Vogel TM & McCarty PL (1985) Biotransformation of tetrachloroethylene to a gasoline-contaminated aquifer. J Contam. trichloroethylene, dichloroethylene, vinyl Hydrol. 40:261-284 chloride, and carbon dioxide under 12. Davis JW & Carpenter CL (1990) Aerobic methanogenic conditions. Appl. Environ. biodegradation of vinyl chloride in groundwater Microbiol. 49:1080-1083 samples. Appl. Environ. Microbiol. 56:387024. Vogel TM, Criddle CS & McCarty PL (1987) 3880 Transformation of halogenated aliphatic 13. DiStefano, T D, Gossett, J M & Zinder SH

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CHAPELLE AND BRADLEY accepting processes in a petroleumhydrocarboncontaminated aquifer and the significance for contaminant biodegradation. Water Resources Res 30:1561-1570 27. Wilson JT & Wilson BH (1985) Biotransformation of trichloroethylene in soil. Appl. Environ. Microbiol. 49:242-243

Chapter 15 MICROCOSMS FOR SITE-SPECIFIC EVALUATION OF ENHANCED BIOLOGICAL REDUCTIVE DEHALOGENATION DONNA E. FENNELL1 AND JAMES M. GOSSETT2 1 2

Department of Environmental Sciences, Rutgers University, New Brunswick NJ, USA School of Civil and Environmental Engineering, Cornell University, Ithaca, NY, USA

1. INTRODUCTION: FACTORS INFLUENCING THE PROBABLE SUCCESS OR FAILURE OF REDUCTIVE DEHALOGENATION SYSTEMS Enhancement of in situ anaerobic biological dehalogenation processes is accomplished by providing an exogenous electron donor to stimulate the indigenous dechlorinating microbiota to use chloroethenes or other halogenated compounds as electron acceptors (Figure 15.1). This coupled oxidation-reduction scheme may appear to be a straightforward process. However, the subsurface is a complex environment with regard to its hydrology, chemistry, and microbiology, and each candidate site must undergo a thorough evaluation prior to remedial efforts. Application of this relatively new bioremediation technology to sites that are poorly characterized will inevitably produce inconsistent outcomes and result in wasted funds. The link between a clear understanding of the microbiology of organohalide-contaminated aquifers, and the likelihood of successful application of biological cleanup methods, is still relatively new, and characterization methods are still being developed. As we have seen in prior chapters, an overall understanding of the environmental significance of dehalorespiring bacteria is increasing at a remarkable rate. Molecular level detection of dehalorespiring bacteria has demonstrated their wide distribution, both in pristine and contaminated sediments and aquifers (28, 30, 58). Bioaugmentation of contaminated aquifers with cultures containing dechlorinating bacteria has also been demonstrated successfully (45). Despite these advances, use of the technology for remediation is still limited by many factors—chief among them is the inherent heterogeneity of the chemical, hydrological, and biological components of the subsurface. These heterogeneities make all aspects of site characterization and remediation challenging, and in some cases, with our current technology, next to impossible (14, 102). Within this context of uncertainty, we have a significant stake in understanding how native dechlorinating bacteria can be stimulated and manipulated in order to meet Dehalogenation: Microbial Processes and Environmental Applications, pages 385-420 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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remediation goals. This chapter focuses on the use of laboratory microcosm studies to characterize native microbial populations, prior to initiating pilot- or full-scale chloroethene dehalogenation systems. It addresses how microcosm results may be used to assess the likely response of the microbial community to in situ enhancement in the field, and how such results may be used to determine parameters that are useful in modeling or designing field-scale systems. While the focus is on the chloroethenes, many of the ideas that are presented are applicable to other halogenated contaminants. A microcosm study may provide answers to the following questions: Are native dechlorinating bacteria capable of complete dechlorination of the original chloroethene contaminants to ethene (ETH), or do partially chlorinated daughter products accumulate under reduced conditions? Which electron donor is most effective for stimulating reductive dechlorination? What is the fermentation pathway of the electron donor? Do other terminal electron-accepting processes control the effectiveness of the electron donor addition? How much electron donor should be added?

1.1. Site Hydrogeological and Chemical Evaluation A number of protocols and screening tools describe the proper steps required for general site characterization (96), for assessing natural attenuation potential (91,99), or for determining if enhanced reductive dehalogenation is an appropriate technology for a site (71). This section addresses some aquifer and groundwater qualities that are desirable for the application of enhanced reductive dechlorination, and thus for choosing candidate sites for microcosm studies. The time, effort, and expense involved in running microcosm studies clearly dictate that the locations for testing must be carefully chosen according to the best and most current site data. Candidate areas within the site should be further characterized through monitoring and sampling, from additional well

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installations or from existing wells, if necessary. The microcosm study should be as expansive as possible, including studies with sediment/groundwater mixtures from a number of promising locations, and/or creating composite sediment core samples from multiple locations.

1.1.1. Hydrogeological Requirements Candidate sites for enhanced biological reductive dechlorination must have physical characteristics that enable the establishment and control of a flow scheme needed to distribute electron donor within the subsurface. Low hydraulic conductivities, less than cm per sec), can retard electron donor movement in the aquifer. In addition, a widely fluctuating water table; extreme depth to groundwater; and site heterogeneities, such as significant stratigraphy or buried objects, would cause substantial shortcircuiting during imposition of a flow scheme, and may disqualify a site (71). Even with such limitations, enhanced reductive dehalogenation has been successfully implemented in difficult situations, such as fractured basalt aquifers (89, 90), and in peat layers overlying aquifers (87).

1.1.2. Temperature Requirements Because enhanced reductive dehalogenation relies on microbial activity, a groundwater temperature of greater than 10 °C to 15 °C is desirable (71). Upper temperature limits are generally not of concern since most groundwater systems do not exceed temperatures between 20 and 30 °C. Dehalogenation has been demonstrated at temperatures of 35 °C (34), and even up to 65 °C (52). The primary limitation of low temperature is its slowing effect on microbial metabolism. An extended period of remediation, as required due to low biological reaction rates, may allow the contaminant plume to spread unacceptably. However, low ambient temperatures are not necessarily an impediment to the application of dechlorination technology, especially if other site parameters are promising. One of the earliest incidences of intrinsic in situ tetrachloroethene (PCE) dechlorination to ETH was documented at a solvent transfer facility site in northern Toronto, Canada, with a groundwater temperature of 10 °C (65). It is crucial to conduct microcosm studies at a similar temperature range to that encountered in the field. Oftentimes, this will require extended incubation periods for microcosm studies conducted at temperatures representative of the subsurface.

1.1.3. pH and Alkalinity Requirements In general, aquifers with pH values less than 5 or greater than 9 are not considered good candidates for reductive dechlorination (71,99). Anaerobic systems which support dehalogenation generally operate best between pH 6 and 8. This is not surprising since biological systems generally function within these pH limitations. This is further supported by most laboratory studies, which indicate a pH optimum near neutral for dehalogenating bacteria. For example, in one series of experiments with an enrichment culture known to contain Dehalococcoides, dechlorination of PCE was four-fold slower at pH 6 and two-fold slower at pH 8, than at pH 7 (105).

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Due to the formation of acids during fermentation of the electron donor, adequate alkalinity, or a sufficient buffering capacity, is required to prevent a pH drop and possible slowing of the dechlorination rate. Acidification (pH drop to as low as 5.5) and inhibition of PCE dechlorination was observed in a large-scale sandbox aquifer seeded with Dehalospirillum multivorans and a mixed DCE-dechlorinating culture, with ethanol as an electron donor (20). Buffer addition alleviated this problem to some extent. Aside from biological considerations, regulatory requirements also dictate groundwater pH limits. In general, groundwater alkalinity of less than 1 g per L (71) is considered insufficient for enhanced remediation. Buffer addition to adjust the pH to near neutral and maintain it during system operation should be added to the enhancement regimen. It is also conceivable that site organisms may be adapted to the ambient pH. If possible, microcosm studies should be performed both at ambient pH and with a buffered neutral pH to determine the best site approach.

1.1.4. Presence of Co-Contaminants Toxic organic co-contaminants, such as chloroform and cyanide (16), or toxic heavy metals may inhibit dehalogenating bacteria and other community members. Heavy metal toxicity can be lessened by the onset of reduced conditions if the site geochemistry supports the formation of sulfides that precipitate or sequester free metals. However, any pH drop associated with enhancement of reduced systems may in turn mobilize heavy metals in the soil—thus another potential reason for buffering the system. Ironically, other co-contaminants, such as petroleum hydrocarbons or organic solvents, such as acetone or methanol, may actually be beneficial since they provide a co-substrate, and their degradation produces reduced conditions and simpler organic acids, that may act as electron donor substrates to support reductive dechlorination (24, 65,85). Indeed, the presence of enough of these types of co-contaminants may make the site a better candidate for natural attenuation than enhanced remediation (99).

1.2. Site Microbiological Evaluation 1.2.1. Presence of Indigenous Dehalogenating Bacteria Examination of historical site data is the first step to assess whether indigenous dechlorinating bacteria are present in a chlorinated solvent plume. For example, if an aquifer originally contaminated with trichloroethene (TCE), shows the presence of dechlorinated metabolites, such as dichloroethene (DCE) isomers, vinyl chloride (VC), and ETH, these metabolites are indicative of microbial dechlorination activity. Site peculiarities may, however, prevent an easy analysis of the problem. Site data are often collected over a large grid with widely spaced sampling points or times. In a very heterogenous matrix, flow patterns in an aquifer may shift over time, and the dispersion of daughter products may occur. Additionally, secondary microbial processes may result in the oxidation of the daughter products, either in anoxic or aerobic portions of a chloroethene plume (12, 13, 97). Moreover, it is also difficult to determine whether dechlorination activity is on-going, or has occurred (possibly many years) in the past. In some cases, a finer grid of monitoring wells and more periodic sampling may be

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required to clarify the situation. In addition to using field data to identify promising locations for obtaining groundwater and sediments for microcosm studies, analysis of site material for specific dehalogenating bacteria using molecular methods may provide useful information during initial screening activities (see Chapter 3). While it is currently difficult to correlate positive PCR results with an actual abundance or activity of dechlorinating bacteria, such information provides some indication whether local conditions are (or at least were) favorable for supporting dechlorinating populations. In the absence of available sediment cores, groundwater or sediment fines obtained from existing monitoring wells (23) can also be tested using sensitive, nested PCR techniques (30,58). Such testing, however, supposes that the current dehalogenating bacterial isolates (the only ones for whom genetic information is available) represent the full spectrum of dehalogenating bacteria. This is probably not the case, but as our base of knowledge increases, such techniques will become increasingly useful.

1.2.2. Presence of Microbial Mediators or Competitors of Dehalogenation A second question regarding the candidacy of a site for bioremediation, is the presence and ability of microbes to support a dehalogenating microbial community. Conversely, it is also necessary to evaluate whether competing processes are present or may occur. To this end, electron donors must be identified that will be degraded by the supporting microbial populations and effectively stimulate dechlorination. Also, the pathway(s) of electron donor degradation and other important competing terminal electron-accepting processes should be identified. Groundwater geochemistry and in situ hydrogen measurements can be used to delineate the dominant terminal electronaccepting processes at different points in a plume (18,60,61). This type of information can be used to determine the potential fate of an electron donor in the subsurface, e.g., how much donor will be used directly for competing processes such as sulfate- or iron (Ill)-reduction during the course of the bioremediation effort. Just as it is possible via molecular methods to detect specific dechlorinating bacteria, it is also possible to detect supporting community members, such as fermentative syntrophic bacteria, sulfatereducing species, and others (27 ,43) that may be important in the bioremediation process.

2. USE OF LABORATORY MICROCOSMS TO ASSESS DEHALOGENATION POTENTIAL OF THE INDIGENOUS MICROBIAL COMMUNITY 2.1. Usefulness of Microcosm Studies Microcosm studies provide physiological evidence of the likely response of the subsurface microbial community to the addition of specific electron donors, and allow for a mass balance determination on the chloroethene contaminants and the added electron donor(s). The amount of electron donor necessary for dechlorination and how much is used by competing electron-accepting processes, such as sulfate reduction and methanogenesis, can be determined. By following a variety of fermentation products it is also possible to determine the pathway of electron donor fermentation. Enriching the

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system also increases relevant populations and allows more ready detection through molecular methods.

2.1.1. Competition for Electron Donor During a microcosm study, the number of electron donor candidates that can be tested for their ability to stimulate reductive dechlorination is limited only by time and money. Potential candidates include simple organic acids to complex substrates such as compost leachate. The use of simpler electron donors makes it easier to quantify results, and provides an overall better understanding of the processes involved. If more complex substrates are to be tested, simpler electron donors should also be included in the protocol. In the end, economic, regulatory, and technical considerations, in addition to microcosm results, will determine the electron donor for full-scale application. Hydrogen is an important electron donor for several known dehalogenating species. Dehalococcoides ethenogenes uses only molecular hydrogen as an electron donor for reductive dechlorination of PCE to ethene (66, 67). A closely related organism has been isolated which dechlorinates chlorinated benzenes with hydrogen as an electron donor (2), and organisms that phylogenetically group very closely with Dehalococcoides have been widely detected using 16S rDNA molecular methods (28, 30, 45, 58, 78, and Chapter 3). One study has shown a correlation at many contaminated sites between the presence of 16S rDNA sequences closely related to Dehalococcoides, with concomitant reductive dechlorination of PCE or TCE past cis-DCE (28). Evidence has also been presented for the existence of other hydrogen-utilizing bacteria that dechlorinate DCEand VC to produce ethene (33, 82). Hydrogenotrophic dechlorinating bacteria that can convert the chloroethenes to ethene are of great environmental and engineering significance because through their activity, the chloroethenes may be completely detoxified. Other dehalogenating bacteria, e.g., Dehalobacter restrictus (limited to molecular hydrogen), Dehalospirillum multivorans and Desulfomonile tiedjei (able to utilize molecular hydrogen as well as other organic compounds), and Desulfitobacterium and Desulfuromonas strains (able to use acetate or other organic compounds), are unable to dechlorinate the DCE daughter product. Although these organisms are also environmentally important, their activity constitutes only the first step in dechlorination and detoxification of PCE and TCE. In cases where DCE accumulates, indicating a block in the pathway, strategies such as bioaugmentation, anoxic or aerobic mineralization, or aerobic co-metabolism, may be used. The addition of the appropriate electron donor may have a two-fold impact for enhancing in situ dechlorination. The addition of electron donor not only stimulates the activity of dechlorinating organisms, but also the activity of other organisms that utilize the electron donor or products of its degradation. This response could be beneficial—e.g., increasing the activity of syntrophic hydrogen-producing organisms, or detrimental—e.g., increasing the growth of methanogens through the addition of methanogenic substrates like methanol. A simplified schematic of the electron flow that exists in complex subsurface communities is shown in Figure 15.2. Focus is on the use of hydrogen by dechlorinating bacteria that can transform the chloroethenes to ethene. However, other dechlorinating

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bacteria may produce DCE while using acetate or the added electron donor. The diagram depicts the various electron acceptors that may be present in the subsurface, including the chloroethenes that are the target of remediation. The diagram does not show oxygen or nitrate, however these electron acceptors may be important initially or under certain circumstances. The effect of adding exogenous organic substrates (donors) may be multi-fold: 1. Support sulfate or iron (III) reduction, or in the case of methanol or acetate, be directly used for methanogenesis; 2. Stimulate production of hydrogen and acetate by proton-reducing syntrophic organisms or other fermenters; Channel the resulting hydrogen to dechlorination, methanogenesis, 3. acetogenesis, sulfate reduction, or iron (III) reduction; and 4. Provide acetate as an electron donor for sulfate reduction, iron (III) reduction, acetotrophic methanogenesis, and dechlorination to DCE. Use of supplied reducing equivalents for processes other than dechlorination results in a requirement for more electron donor than is stoichiometrically necessary for complete chloroethene dechlorination. This condition should be avoided for a number of reasons. First, electron donor costs can be a significant portion of the total cost of a field remediation effort, and the use of electron donor for purposes other than dechlorination becomes a wasted resource. Second, unnecessary stimulation of biomass production of non-target microorganisms causes well and aquifer biofouling (10) and rapid depletion of supplied electron donor near sites of injection, clogging and exacerbating problems of distributing electron donor effectively throughout the desired remediation zone. Excessive bubble formation during methanogenesis can significantly diminish the hydraulic conductivity of an aquifer (10). Finally, as with any scenario

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where a significant mass of organic matter is subject to anaerobic conditions, excessive methanogenesis may result in unsafe, explosive levels of methane (5,15,42) that could result in safety concerns during remedial work. Although such a scenario during enhanced reductive dechlorination would be extreme, this possibility should be considered and monitored.

2.1.2. Minimizing Competition for Electron Donor In laboratory cultures operated at non-inhibitory levels of PCE, it has been shown that providing reducing equivalents for dechlorination through the addition of excess methanol as organic electron donor resulted in the growth of more methanogenic biomass vs. dechlorinators. The initially perceived solution to this problem of competition (addition of higher methanol concentrations to account for an increasing competitive demand for it) eventually resulted in failure of dechlorination due to a displacement of dechlorinators by methanogens (92). Addition of non-methanogenic substrates such as organic acids that are fermented directly to hydrogen is one strategy for suppressing this type of direct methanogenic competition. Hydrogenotrophic dechlorinating bacteria have a lower threshold for hydrogen than methanogens and can out-compete hydrogenotrophic methanogens for hydrogen at low hydrogen levels (8, 32, 59, 88, 104). Thus, the addition of electron donors such as fatty acids that produce hydrogen only at low levels (low donors), is a technique that can be used to channel hydrogen to dechlorination and away from methanogenesis (32, 104). Competition by sulfate-reducing organisms is a more complex problem (Figure 15.2). Sulfate-reducing bacteria comprise a diverse group of organisms that can utilize many different organic compounds in addition to hydrogen (44). Hydrogen levels of 115 nM have been found to support sulfate reduction in a variety of sediments and laboratory cultures (61, 62, 68, 81). The hydrogen threshold for dechlorinators is somewhat, but not substantially below that of sulfate-reducing bacteria (8, 59, 68, 88, 103,104). Nonetheless, absence of dehalogenation under sulfate-reducing conditions is sometimes observed. The lack of dechlorination under sulfate-reducing conditions may be the result of: 1) direct competition for supplied organic electron donor by sulfatereducers; 2) inhibition by sulfate, thiosulfate, sulfite, or sulfide ions of enzymes involved in dehalogenation; or 3) selective use of sulfate, thiosulfate, or sulfite as the terminal electron acceptor, instead of chlorinated compounds within the same organism (26, 38, 64). Furthermore, in a high-sulfate environment, the initial dechlorinator population may be low compared to that of the sulfate-reducing bacteria. When an electron donor is initially added, the sulfate-reducing population may keep hydrogen near the threshold for sulfate reduction. Although both dehalogenation and sulfate reduction can occur simultaneously under these conditions, the stimulatory effect of added donor upon the dechlorinator population may be barely detectable at such a low hydrogen level, When sulfate is eventually depleted, hydrogen may increase transiently and stimulate more rapid growth of the dechlorinator population, which would then decrease the hydrogen concentration to the hydrogen threshold of dechlorinators. Therefore, initial reduction of sulfate may proceed comparatively rapidly, while dechlorination may appear to lag. Interestingly, upon depletion of sulfate, some sulfate-reducing bacteria have the ability

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to convert to fermentative growth (on lactate, for example) and live as a syntrophic partner to hydrogenotrophic dechlorinators (37). The question of competition between dechlorinating and nitrate- or iron (III)reducing bacteria has not yet been addressed to a great extent in chloroethene-degrading systems. However, energetic considerations would appear to place dechlorination between these two processes in terms of hydrogen threshold range (59,103). Debromination of bromophenols and bromobenzoates has been shown to occur in iron (III)-rich enrichments (70) and dechlorination of PCE has been demonstrated to occur by both biotic and abiotic mechanisms under iron(III)-reducing conditions at a lakeaquifer interface (1). In a microcosm study of TCE dechlorination in groundwater from a fractured dolomite aquifer with bulk iron(III)-reducing conditions, TCE dechlorination was observed only in microcosms prepared under conditions intended to support methanogenesis and was not observed in the presence of iron (III) oxide (103). For this and similar sites, electron donor addition will eventually result in the depletion of Fe(III) or sulfate, and a microcosm study can provide quantitative assessment of the excess electron donor needed to fulfill these requirements (see Section 3).

2.1.3. Electron Donor Fermentation Pathways In addition to being unable to assess which electron donor will effectively stimulate dechlorination, and whether a electron donor will be fermented at all at a site, we also cannot tell prior to microcosm testing what fermentation pathway the electron donor will follow. Figure 15.3 shows a number of possible electron donor fermentation pathways. The fermentation pathway may help to shift hydrogen availability by converting a more rapidly degraded, higher-hydrogen-producing substrate such as lactate to propionate, a more slowly degraded compound that produces lower hydrogen levels. The fermentation of lactate to propionate may proceed through one of two different fermentation pathways that may (25) or may not (47, 55, 98) result in the formation of hydrogen. In one fullscale system, the fermentation of lactate to propionate apparently occurred without the production of hydrogen, and dechlorination did not proceed until the resulting propionate was fermented to hydrogen (89). Similarly, the fermentation of benzoate through butyrate (see section 2.3.2) and fermentation of ethanol through propionate (84) have also been observed in sediments and cultures.

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2.1.4. Nutrients Along with electron donors, nitrogen, phosphorous, and trace nutrients (e.g., trace metals and vitamins), may be important for reductive dechlorination in the subsurface. For oligotrophic freshwater aquifers with low organic matter and microbial populations, these aspects may be particularly important. Some dehalogenating bacteria have required growth factors beyond the usual nitrogen and phosphorus. For example, Dehalococcoides ethenogenes strain 195 requires vitamin and other, as-yet-unknown factors (67). During enhanced dechlorination, nutritional requirements may be met by directly adding specific growth factors along with the electron donor, or may be produced in situ by the supporting microbial community. Thus, choice of electron donor can directly affect the nutritional status of the dehalogenating population. A microcosm study will demonstrate which electron donor yields the most rapid, efficient dechlorination with the shortest lag period. While not all of the difference between electron donors can easily be ascribed to nutritional status, this effect would be a part of the overall observed difference. Addition of trace nutrients via yeast extract should be done only at low levels, since this rich carbon source is notorious for producing unwanted biomass. Another means for minimizing unwanted biomass production is to separate, temporally or physically, dual nutrient requirements, so that field injection of the electron donor and nutrient source occurs in separate wells or through time-separated pulse inputs to avoid biofouling (76). A typical laboratory protocol used to address these issues is presented in Table 15.1, and includes replications of lactate-amended bottles, with and without yeast extract or vitamin or both.

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2.2. Microcosm Protocols A number of protocols are available that describe microcosm preparation and various techniques for working anaerobically and aseptically with sediments and groundwater (36,63,71,108). Likewise, many reports are available that describe specific techniques that have been used for microcosm studies (see for example, 1, 9, 35, 39, 40, 41, 49, 50, 54, 58, 65, 69, 72, 86, 101, 103). Microcosm studies are sometimes performed destructively so as to minimize (sub)sampling error (57 ,65, 101). This method drastically increases the number of replicate bottles that are periodically sacrificed for analysis during a study, and risks the danger of running out of bottles before the desired activity has been documented. Non-destructive microcosm procedures are briefly described in this section.

2.2.1 Collection of Material Aquifer and subsurface characteristics, previous site experience, price, and availability of equipment will dictate what sediment collection technology is utilized at a particular site (51). Depending upon the site and the material to be sampled, aquifer solids may be obtained at relatively shallow depths through use of a hand auger or slide hammer. For more difficult material and deeper sampling, core collection may be accomplished via direct-push methods such as Geoprobe®, or, drilling with a hollowstem auger and use of split spoons or continuous samplers to collect sediment at discrete depths. For aquifer material that is not amenable to coring, sediment may be obtained by pumping sediment-groundwater slurry from a temporary well. Regardless of the collection methods, cross-contamination between coring locations, laboratory strains and exposure to air should be avoided. Samples may be temporarily stored at 4 °C, and then used as soon as possible to construct the microcosms, since the microbial population will potentially change even at this low temperature.

2.2.2. Preparation of Composite Samples and Microcosms Preparation of samples representative of a site requires special and stringent procedures. These are explained in detail to provide a better understanding of some of the difficulties encountered with microcosm studies. For anoxic subsurface samples, microcosms are generally prepared in a standard anaerobic chamber or in separate, disposable glovebags to prevent oxygen contamination. Use of disposable glovebags helps prevent microbial cross-contamination. Alcohol should be used minimally for disinfection of surfaces in these closed environments because the fumes can potentially contaminate sediments and serve as an electron donor. Hydrogen in the anaerobic chamber atmosphere will contribute to the electron donor pool unless it is subsequently removed by purging. When cores are available from several sampling locations at a site, a decision must be made as to the value of retaining information concerning spatial variation in dechlorination potential. Separate microcosm studies can be run on material from each core; however, that is not likely to be economically feasible. In most cases, representative cores are grouped together to create a composite and then sampled for

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initial analyses such as moisture content, pH, alkalinity, and molecular or microbial analyses. The composite sediment may be distributed gravimetrically or volumetrically to sterile serum bottles capped with Teflon®-lined septa to minimize sorptive loss of chloroethenes. Groundwater may be amended with a redox indicator (e.g., resazurin) and buffer, if desired, to achieve pH values between 6 and 8 and to provide adequate buffering for acid formation due to electron donor fermentation and dechlorination. The pH, alkalinity, and required buffer amendment may be empirically determined using a few test microcosms. Sediment/groundwater ratios and volumes should be adequate to minimize disturbance of the system by sample removal and ensure adequate material to cany through a several-hundred-day incubation (30). After distribution of the material, the bottles may be randomly used in a protocol such as the one shown in Table 15.1 (71).

2.2.3. Microcosm Amendments After setup, background concentrations of chloroethene, methane, hydrogen, and cocontaminants must be determined. Initial chloroethene concentrations should: 1) more or less reflect that found at the site; 2) present no analytical difficulties; 3) be high enough to allow clear delineation between activity and normal analytical variability; and 4) not be so high as to potentially form a separate phase (DNAPL) or pose toxicity problems. The latter concerns are dependent on the aqueous solubility of substrate. If necessary, filter-sterilized, anoxic PCE or TCE (or whatever substrate is appropriate for the microcosm study) is added to boost the level over what was observed upon set-up and to allow clear delineation of dechlorination activity. The prepared microcosms are shaken overnight at the incubation temperature to equilibrate the added PCE or TCE with the sediment and groundwater, and then analyzed to determine the initial concentrations of chloroethenes. The microcosms are then amended with electron donors and/or nutrients (for example as in Table 15.1).

2.2.4. Microcosm Operation Inverted microcosms should be incubated in the dark with gentle or intermittent agitation at the prevailing groundwater temperature. At regular intervals, the microcosms are monitored and re-amended with the appropriate substrate when parent chloroethenes (PCE or TCE) or electron donor has been depleted. Microcosms may be sampled non-destructively by withdrawing small headspace and liquid samples. Volatile substrates and products, such as chloroethenes, ethane, methane, and hydrogen are analyzed using gas chromatography (29, 32, 34). For gas sampling, headspace samples are removed aseptically and anoxically using a sterile gas-tight syringe. For liquid analysis, groundwater (0.25-ml) samples are removed from microcosms using a sterile, disposable 1 -ml plastic syringe. Smaller sized needles should be used whenever possible to avoid septum coring and loss of mass balance. Sacrificial sampling is preferred for assessment of acid-volatile sulfide (AVS) formation (4), which allows quantification not only of sulfides in uncomplexed soluble form, but also sulfides that may have precipitated after their formation. (Other

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techniques—such as headspace analysis, ion-chromatographic analysis of soluble samples, or sulfide-ion-specific electrodes—will not usefully assess total sulfide).

2.3 Microcosm Case Studies As a part of a protocol development project (71), microcosm studies were conducted for several TCE-contaminated sites. Laboratory results were linked to small-scale fieldtest results in order to determine if the microcosm studies predicted the same response to enhancement that was observed in the field-scale tests (3). To illustrate some of the differences between the microbial ecology of TCE-contaminated sites, three examples are presented: 1. A location within a TCE-contaminated aquifer at Cape Canaveral Air Station (CCAS) Florida, with low daughter-product concentrations; 2. A location within the same CCAS plume, with higher daughter-product concentrations; and 3. A TCE-contaminated aquifer at the former Naval Air Station at Alameda Point (AP), California.

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The CCAS site is a shallow plume of TCE, DCE isomers, and VC, in a relatively homogeneous aquifer consisting of coarse to fine sands with shell deposits that may exist as distinct layers or lenses (74). Sediment and groundwater samples used for a microcosm study were taken from 12 to 15 ft below ground surface. The AP location is a TCE, DCE isomer, and VC plume in a homogeneous, sandy clay aquifer. The AP environment is high in sulfate and is estuarine in nature. Depth of groundwater and sediment recovery was 15 to 17 ft below ground surface. There were no other known significant contaminants in either plume. Table 15.2 shows the groundwater geochemistry of the three locations. AP exhibits one order of magnitude greater sulfate concentration than CCAS. Aside from different TCE and daughter product concentrations and a slightly higher methane concentration, there was no discernable difference between the two CCAS sites which were only about 10m apart (30). The CCAS microcosms had a high buffering capacity from the seashell (carbonate) deposits in the aquifer solids, and no additional buffer was required, whereas AP microcosms were buffered with sodium bicarbonate.

2.3.1. Dehalogenation In samples from one CCAS location (W9), dechlorination was observed in only two of the microcosms and only cis-1,2-DCE was produced, whereas dechlorination of TCE was observed in all microcosms from the other site (W6) (Figure 15.4). The pattern of rapid dechlorination of TCE to VC and then slower dechlorination of VC to ethene, is very typical of that observed with D. ethenogenes, strain 195 (67). Polymerase chain reaction (PCR) 16S rDNA analyses of the sediments from W6 indicated the presence of an organism very similar to D. ethenogenes. However, PCR analyses at location W9 that exhibited a different dechlorination pattern, showed very faint or no detection of the organism (30). With the exception of daughter product profiles and slight differences in methane production, the groundwater characteristics of these two locations were very

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similar, yet the dechlorinating bacterial populations were clearly different, whether assessed by the physiological study or PCR analyses (30). The heterogeneous distribution of dechlorinating activity at CCAS within a relatively close proximity highlights a potential weakness in extrapolating the results of microcosms derived from small samples to predict overall field responses. As illustrated in the above example, different microbial populations may exist within discrete locations at a site. It is likely that the indigenous dechlorinating organisms could be distributed to inactive locations via extensive groundwater pumping that would most likely accompany in situ bioremediation. Thus, a “positive” result at one location (as described for W6) suggests that the site is amenable to engineered bioremediation, provided the system installation encompasses the active location. Alone, the negative result both in dechlorination activity and PCR analyses seen in W9 would suggest that field-scale efforts at the site would be very risky. One can not overemphasize the need for careful and comprehensive analyses of site data—"up-front"—and further characterization of locations evidencing higher daughter product concentrations when data are spotty. In microcosms from the AP site, the pattern typically observed was TCE dechlorination to ETH, with only transient accumulation of cis-1,2-DCE and VC (Figure 15.5). The rate of complete conversion to ethene was dependent upon the electron donor added. Butyrate, the electron donor ultimately chosen for field-testing, gave the most rapid ethene production. However, propionate, yeast extract and some lactate-amended bottles also showed complete conversion to ethene. The sudden onset of ethene production shows a different pattern than that observed in the CCAS sediments, and suggests the presence of very different dechlorinating populations. This is supported by PCR analyses that detected D. ethenogenes in only a few of the microcosms prepared for this site (17).

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2.3.2. Electron Donor Degradation and Fermentation Pathway The fermentation pathway for electron donors was also shown to differ between the CCAS and AP sites, demonstrated in Figures 15.3, 15.6 and 15.7. These charts present a mass balance for each compound based on electron equivalents [Equivalents are based on complete conversion to for the electron donors, and on equivalents needed for reduction ofthe electron acceptors]. At the CCAS site, lactate was fermented through propionate, particularly during re-spiking of electron donor (Figure 15.6). This is the effect depicted in the schematic of Figure 15.3, where addition of a rapidly degraded electron donor produces a slowly degraded electron donor. Data from the microcosm studies demonstrate fermentation ofbenzoate through a pathway that includes significant butyrate accumulation (Figure 15.7). In the high sulfate environment at AP, electron donors were often immediately consumed without the concomitant appearance of fermentation products (Figure 15.8). AP sediment did not show as much, or as consistent formation ofpropionate from lactate as the CCAS sediments. Butyrate fermentation was slower than lactate or propionate (Figure 15.9). As butyrate eventually disappeared, acetate did not accumulate, but was used apparently by sulfate reduction. The fermentation ofbenzoate was observed in only one of the triplicate bottles from Alameda Point.

2.3.3. Competition for Electron Donor Further observation of the equivalents balance of electron donor added and the measured reduction products (Figures 15.6 to 15.9) for each location allows a quick

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determination of the fate of the electron donor. For CCAS W6, the bulk of hydrogen formed from electron donor was channeled to dechlorination. Methanogenesis was observed in some bottles (Figure 15.7), while sulfate (1-2 mM) was not a significant sink for the applied electron donor. Most of the donor equivalents added were recovered at the end of the study as acetate. The equivalents balance for AP shows a much different scenario (Figures 15.8 and 15.9), with rapid loss of added organic electron donors, presumably through concomitant reduction of sulfate. Methanogenesis was not observed in the Alameda Point microcosms. Thus, sulfate reduction and dechlorination were the primary terminal electron-accepting processes. Note that, the sulfide that was recovered at the end of the study through sacrificial AVS analyses did not completely close the equivalence balance. Some of the "missing" equivalents would have been channeled to biomass synthesis, neglected in this balance construction. Results from the case studies show significant site-specific differences that could only have been discerned through microcosm studies. While two other lines of evidence were available for these sites, i.e., site data and PCR analyses for a specific dechlorinating bacterial species, only the microcosm studies clearly showed the differences in electron donor usage, electron donor fermentation pathways, and extent of dechlorination.

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3. COMPUTATION OF SITE-SPECIFIC ELECTRON DONOR DOSAGES USING MICROCOSM RESULTS In principle, site analyses and microcosm results can be used to design a bioremediation plan that often includes enhancement through the injection of alternative electron donors. One strategy might be to satisfy the competitive demand for reducing equivalents from such activities as methanogenesis and sulfate reduction. Results from microcosm studies can be used to assess electron donor fermentation pathways and the fraction of reducing equivalents that will be channeled to dechlorination. A more elegant approach is potentially available, in that microcosm results can be used to develop sitespecific inputs to comprehensive contaminant transport/fate models, allowing quantitative model estimates of dynamic response to alternative enhancement strategies. This approach is described in Section 4. In the absence of more sophisticated modeling approaches, a crude, but rational injection formulation may be derived as follows. First the volume of water and the mass of the soil in the treatment zone is calculated:

where: = volume of water within the zone (m3) = total effective volume of affected treatment zone (m3) = porosity (void-volume fraction) (generally, varies between 0.2 and 0.4)

where: = mass of dry soil solids within the zone (kg) = bulk mass density of solids within zone (g per cm3 of bulk volume) (generally can be assumed approximately 1.8 g/cm3)

Estimate the hydraulic residence time within the zone:

where: = estimated hydraulic residence time within the zone — i.e., the time it takes for a conservative tracer to pass through the zone under treatment conditions (days), = estimated flow rate through the zone (including injected water) (m3/d);

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Determine the retardation factor for the chloroethenes:

where: = retardation factor for chloroethenes in the zone = the sorption distribution coefficient for chloroethenes (cm3/g). While values of FR can vary considerably from site to site, depending on organic-carbon content of the soil, a reasonable default value would be around 5 for chloroethenes.

Calculate the electron donor demand for the groundwater and soil phases:

where: = electron donor demand from contributions in the groundwater phase (mol of donor per of groundwater); = electron donor demand from alternative electron acceptors within the groundwater phase (mol of donor per of groundwater). This value can be estimated from stoichiometric equations for oxidation/reduction ofdonor/acceptor couples, groundwater analyses of electron acceptors, and adjusted for results observed in microcosm studies (i.e., some alternative electron acceptors may not have been significantly reduced and can therefore be ignored at the site); = electron donor demand from reductive dechlorination of dissolved chloroethenes (mol of donor per of groundwater). Values are corrected for estimated methanogenic competition for or through direct methanogenic consumption of the supplied donor. can be estimated from groundwater analyses of chloroethenes, and then inflated through use of a safety factor, for methanogenic competitive demand (Equation 15.6). It is expected that may vary from 1 (no methanogenesis from and no direct methanogenic use ofsupplied donor) to 20, depending upon the donor selected and site-specific conditions of microbial ecology. Thus:

where the concentrations of the chloroethenes are expressed in mol per groundwater, and is the number of moles of donor required to yield one mole of in fermentation (e.g., in the case of butyrate

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where: = electron donor demand from contributions within the soil phase (mol of donor per kg soil solids) = electron donor demand from alternative electron acceptors within the soil phase (mol of donor per kg of soil solids). This can be estimated from soil analysis using measured particulate nitrates, sulfates, and Fe(III) per kg dry soil. Bioavailability, however, may not be representative of the total values. The microcosm results may be usefully employed here to estimate the concentrations of bioavailable alternative electron acceptors. Note, however, that one should correct for the fact that microcosms employ a matrix that is 33% solids and 67% groundwater (wt/wt), whereas the in situ aquifer is perhaps 85% solids and 15% water (wt/wt). = electron donor demand from reductive dechlorination of chloroethenes sorbed to the aquifer solids within the test plot (mol of donor per kg of soil solids). The fraction of soluble chloroethenes (of the total within the treatment zone) can be estimated as the reciprocal of the retardation factor, FR (Equation 15.4). Thus, by extension of what was earlier estimated for dechlorination of soluble species,

From the information derived from the preceding calculations, a two-phased strategy is recommended for electron donor dosing. Phase I Based on calculations, Phase I will utilize a higher dosing rate than Phase II, and last for one hydraulic retention time During this period, sufficient electron donor is required to meet the estimated demand from all soil-phase sources within the zone (chloroethenes and alternative electron acceptors), plus the demand from all groundwater entering the zone during Phase I. The objective is to eliminate competing electron acceptors as quickly as possible, thereby establishing fermentation/ methanogenic conditions as quickly as possible. Furthermore, it is recommended that the electron donor administered during Phase I include some readily degradable substrate (e.g., yeast extract or ethanol or lactate), as well as less-readily degradable substrate intended for later use in Phase II (to promote growth of organisms that use such substrates).

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Alternatively, the dose may be expressed as a concentration to be achieved in the aquifer at the injection point:

[Note: In instances where a high demand for electron donor exists due to high levels of particulate forms of alternative electron acceptors, the above-described strategy can potentially result in inhibitory (or otherwise unacceptably high) concentrations of electron donor. In such cases, Phase I may be extended beyond a single retention time, reducing the dosage rate and concentration. For example, if Phase I were extended to two retention times, then one would substitute and in Equations 15.9 and 15.10, for and respectively.] Phase II Phase II is the treatment period extending beyond Phase I, to the end of the treatment period. It is assumed that the particulate-based demand from alternative electron acceptors has been met in Phase I. However, it cannot be assumed that dechlorination activity in Phase I will completely biodegrade and remove sorbed chloroethenes. A conservative dosing strategy is therefore recommended for Phase II which targets sorbed chloroethenes (at their estimated, original levels), and also accomodates the demand for electron donor arising from influent groundwater (chloroethenes and alternative electron acceptors). Furthermore, the dose rate is selected to theoretically meet the demand from sorbed chloroethenes within one hydraulic retention time.

The assumption for Phase II, i.e., that dosing attempts to dechlorinate the entire sorbed inventory of chloroethenes in a single retention time, might seem excessive. However, it is always possible to alter dosing rates during Phase II to reflect results from ongoing monitoring. If, for example, biofouling becomes an issue, donor could be withheld and the biomass formed would then be available as a slow-release donor. Donor selected for Phase II should be slowly utilized substrate(s) (or be fermented to such desirable substrates), to minimize methanogenic competition. The choice of donor also minimizes biofouling. Choice for field application should be guided by the microcosm results and ultimate final adjustments made in the field.

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4. DETERMINATION OF SITE-SPECIFIC MODEL PARAMETERS USING MICROCOSM RESULTS A more elegant approach for designing an in situ enhancement strategy employs a conceptually based computational model of dechlorination and competition for electron donor with full integration into a subsurface, contaminant-transport model. This type of model allows designers to explore alternative dosing strategies, including alternative dosing locations, flows, and extraction wells. Microcosm data can be useful in estimating site-specific parameters for such models. A primary application for microcosm studies has been to evaluate chloroethene transformation rates for natural attenuation applications (101). The purpose is to determine in situ dechlorination rates driven by naturally occurring organic matter or cocontaminants such as aromatic hydrocarbons. Dechlorination rates obtained in the laboratory are expressed as first-order rate constants or half-lives, and these values are extrapolated to the field via incorporation into fate and transport models to project plume movement. This approach assumes that the half-lives of target contaminants and all of the factors that control observed transformation of the contaminant, i.e., bacterial populations, dominant bacterial species, nutrient status, concentrations of competitive terminal electron acceptors and the concentration of electron donor (Figure 15.2), will remain constant over the lifetime of the project. For some systems, e.g., landfill plumes with a high or continuous input of organic matter and relatively low levels of chloroethene contaminants, this approach may be valid. However, for many sites, and especially for enhanced systems, these assumptions simply are not valid.

4.1. Models for Reductive Dechlorination An improved model of dechlorination, including the incorporation site-specific parameters is an area deserving further research and development. Many of the models for contaminant degradation in groundwater were originally developed for petroleum hydrocarbons (99). These compounds serve as electron donors and are mineralized by heterotrophic organisms that use oxygen or nitrate as electron acceptors. In contrast, chlorinated ethenes generally serve as electron acceptors in anoxic environments, and may be dechlorinated by only a few highly specialized organisms that use a limited number of electron donors, in competition with other bacteria. Several models are available for application to reductive dechlorination. The widely used modeling package RT3D (21, 93) includes several modules for biokinetic expressions, and also a kinetic package for reductive dechlorination. BIOPLUME III (79) and UTCHEM (11) incorporate Monod kinetics for describing dechlorination. For another site model for reductive dechlorination, chloroethene and electron-donor degradation kinetics, equations for the conversion of an applied donor (methanol) to end products (acetate) thought to be used by dechlorinators, and the competitive inhibition between PCE and TCE were incorporated (95). One- and two-dimensional, finite-element models have been used to describe the effect of hydraulic conductivity on microbial activity during enhanced reductive dechlorination (19). The biological component of this type of model is based on Monod kinetics, and describes a separate biomass population for each dechlorination step. The

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kinetic parameters incorporated here have been taken from experimental results with Dehalospirillum multivorans and a mixed DCE-dechlorinating culture. In addition to chloroethene, an unspecified electron donor (e.g., hydrogen or organic donors) is included in the kinetic expression for chloroethene degradation. A separate biomass component is used to describe the competitive degradation of the electron donor at a constant, first-order decay rate. From these models it was concluded that the delivery of the electron donor to the aquifer is the limiting factor for enhanced dechlorination, even when the competitive use of the donor is set at relatively conservative values. A comprehensive dechlorination model based upon the International Association on Water Quality framework for modeling activated sludge has been described (7). This model incorporates dechlorination and competitive processes, and has a flexible framework that is amenable for adding other processes. Competitive inhibition between the chloroethene compounds is included and inhibition parameters have been added to show the effect of end product buildup during electron donor fermentation. Hexachlorobenzene dechlorination in aquifer sediment enrichments in the presence of excess electron donor and biomass has been modeled using Michaelis-Menten kinetics (75). For this application, it was speculated that the dechlorination was cometabolic in nature. Even so, the fit of the Michaelis-Menten model to the data appeared to be superior to pseudo-first-order rate equations. Monod kinetics have been used to describe reductive dechlorination in a system that was apparently dominated by two separate dechlorinating populations: one converting TCE to DCE, and one converting DCE to VC and ETH. The model expressions include components for both electron donor and chloroethenes, and incorporate competitive inhibition between the different chloroethenes. A comprehensive list of kinetic parameters was developed (46, 80). In addition to adequately describing the complexity of dechlorination, the transformation of electron donors and competition for electron donor and evolved hydrogen must also be adequately described. Hydrogen is produced from fatty acids and alcohols under conditions that involve highly complex interactions of syntrophic, obligate-proton-reducing bacteria (83). Kinetics and thermodynamics govern the oxidation of these compounds. The coupling of syntrophic organisms (hydrogenproducers) and dechlorinators (hydrogen-utilizers) mirrors the well-studied interactions between syntrophic organisms and methanogens. Several models have been described that incorporate mathematical expressions for limiting the rate of electron donor degradation when product formation begins to thermodynamically limit the reaction, as occurs with syntrophic organisms (6, 48, 53, 56, 77). Building on the principles of these closely coupled syntrophic reactions, we have developed a model that incorporates the biokinetic expressions which describes closely coupled syntrophic associations of hydrogen-producing and hydrogen-utilizing microorganisms (29, 31). The model equations that are shown were designed to allow simulation of cultures in 160-ml serum bottles (and therefore be used to extract values of model parameters from microcosm studies). The basic biokinetic structure has also been incorporated into a generalized fate and transport model suitable for field-scale application (100). The expression for electron donor fermentation is the most unusual aspect of the model and is shown in Equation 15.13.

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where: =

total amount of electron donor in the bottle (µmol);

= time (hr); = maximum specific rate of electron donor degradation (µmol mg ); = electron donor-fermenting biomass in the bottle (mg VSS); =

half-velocity coefficient for the donor (µM);

=

electron donor concentration (µM); and

= hypothetical electron donor concentration that, under the instantaneous culture conditions, would result in given the concentrations of all the other reactants and products at that instant. S* is the “equilibrium” concentration of S, and it is related to The term refers to some marginally negative free energy that the organisms must have available to live and grow (83) (Equation 15.14).

S is the actual substrate concentration at a particular time and is related to the free energy available from the fermentation at that point in time (Equation 15.15).

Subtracting and simplifying,

where:

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(S–S*) in Equation 15.13 is replaced by as per Equation 15.16, with calculated using Equation 15.17, giving Equation 15.18, the form used in the implementation of the model.

is a measure of the distance of the reaction from thermodynamic equilibrium. If the electron donor concentration is high relative to the concentrations of the products of the reaction, hydrogen and acetate, the driving force is high, then approaches one and the fermentation reaction is limited primarily by intrinsic kinetics. After the electron donor concentration has decreased and hydrogen and acetate have increased, the driving force is lessened, the value of approaches zero and the fermentation is limited primarily by thermodynamics. While these expressions complicate the model substantially and increase the number of calculations per time step, they must be included for an accurate description of electron donor degradation, the resulting hydrogen levels and subsequent stimulation of either reductive dechlorination or competing processes such as methanogenesis or sulfate reduction. Equations for utilization in dechlorination, methanogenesis, sulfate reduction, and acetogenesis are shown in Table 15.3. Note that the equation parameters units are the same as those described for Equation 15.13. For each process, the key component is the dependence of the substrate transformation rate upon both electron donor (e.g., hydrogen) and electron acceptor concentration, and the hydrogen threshold value for each process. Other equations may be added as needed, for example to show competitive use of added organic donors directly by sulfate-reducers.

4.2. Site-Specific Model Parameters With more descriptive models it is possible to gain a better representation of dechlorination processes in the complex subsurface environment, and therefore more accurately predict and design remediation processes. Field-scale modeling of processes to stimulate in situ dechlorination requires site-specific information for model parameters, including biokinetic data and the biomass of dechlorinating, methanogenic, sulfate-reducing, and electron-donor-degrading populations present at the site. Information obtained from microcosm studies may be fit with a biological model to extract these parameters. Clearly, obtaining these values is not easy; there are two possible approaches. In the first method, microcosm data may be evaluated to provide a conceptual model that describes pathways, organism-types and terminal electron-accepting processes that are likely to occur. Molecular methods, dechlorination patterns, and fermentation pathways can be used to determine the likely dominant populations and pathways at a given site. Adaptation of literature values of relevant kinetic parameters can be used to reflect in situ temperatures (see for example, 31,46, 80). The model is then employed with these parameters, to fit the microcosm data by inversely determining the initial biomass for each relevant population. This method is demonstrated in Section 4.2.4.

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An alternative approach is to determine the initial biomass and types of populations through a counting method such as quantitative PCR or most-probable-number analysis. The model may be fit to the microcosm data by manipulating the biokinetic parameters to determine site- and population-specific parameters.

4.2.1. Determination of Site-Specific Indigenous Biomass Concentrations Using Microcosm Data Sets Here, a proof-of-concept demonstration is provided using our existing model (29, 31) to extract initial and final biomass population distributions from a microcosm data set. The thermodynamic parameters and kinetic parameters of the model have been modified for application to the CCAS sediment/groundwater microcosms previously described. Model inputs, such as pH, ionic strength, sulfate concentration, and bicarbonate alkalinity were modified to reflect the groundwater properties at CCAS (Table 15.2). Biokinetic modules for sulfate-reducing bacteria and homoacetogenic bacteria were added. Relevant kinetic parameters used for the simulation are shown in Table 15.4. The primary constraints for estimating the initial biomass for each physiological group required that the total biomass be within the same order of magnitude of the biomass estimated from the actual cell counts for the CCAS sediment (Table 15.2; 17). Additionally, each replicate microcosm data set was fit using the same initial biomass

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distribution, even though under different circumstances, certain biomass groups would be inactive and simply undergo decay. Thermodynamic parameters were modified to reflect environmental temperatures (24 °C) (29,31). An analysis of the free energy available to butyrate oxidizers in the microcosms was performed to determine and was found to be approximately –5 kJ per mol, as opposed to almost –20 kJ per mol; the latter value has often been observed (31,83). It is likely that the bulk concentrations of acetate and hydrogen that were measured in the microcosms did not reflect the actual concentrations in the pore spaces and microenvironments within the unmixed sediment in the microcosms. Therefore, the assumed equilibrium between all the phases of the microcosm may not be valid. The microcosm system was assumed to be at equilibrium with respect to transfer of volatile components between the gaseous and liquid phases. pH, bicarbonate alkalinity, and ionic strength remained constant during the course of study. Results of a simulation for the CCAS site, butyrate-fed microcosm that also received 20 mg per 1 yeast extract are shown in Figure 15.10. Yeast extract added an additional per bottle per addition. This source of reducing equivalents was simulated as if added as butyrate The initial biomass values used for simulation and the final biomass values obtained after 387 days of simulation are shown in Table 15.5.

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It should be noted that although methane was produced beginning at Day 110, the initial rate of methanogenesis was not sustained. If methane production was from acetate, it is likely that the rate of production would have been sustained or increased, rather than leveling off after Day 170. The decrease in rates appears to be connected to decreasing hydrogen levels. It is therefore likely that the bulk of methane production was from hydrogenotrophic methanogenesis. Addition of acetoclastic methanogens to the model simulation, even at an extremely low initial biomass level, eventually resulted in a very rapid acetotroph bloom with a corresponding exponential increase in methane that was not observed in the microcosm. Inclusion of a homoacetogen pool allowed better simulation of acetate levels and hydrogen consumption. The biomass per bottle values obtained via the model (Table 15.5) should be adjusted to reflect in situ conditions. Assumptions must be made about the distribution coefficient for biomass between sediment and groundwater phases and also take into account any difference between the sediment/water ratios in the microcosms and the in situ situations they purport to represent. Many groundwater models express concentrations (of various parameters) in mg (or moles) per unit volume of groundwater. Thus, the total amount of biomass (mg) associated with one liter of groundwater, including all of the solids associated with that 1 liter of groundwater—i.e., the aquifer volume containing one liter of water—should be obtained. Thus, obtaining useful initial biomass values for groundwater models would require some manipulation of the per bottle values obtained from the biokinetic model. This type of data fitting obviously involves some degree of uncertainty. Sensitivity analysis can help to determine which parameters are most influential in the overall trends of the data set (100). It is also important to assess which aspect of the data set is most important in terms of prediction. For example, is it most important to use biomass numbers that most accurately fit the TCE time course? Or since VC is a more important regulatory target, should we try to most accurately fit the VC time course? These types of considerations are crucial in the successful use of modeling to extract quantitative site parameters, which in turn will improve the applicability of the model for real-site remediation. The approach to be taken is likely to be dependent upon the purpose of modeling in any specific instance.

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5. CONCLUSIONS Microcosm studies are currently one of the best tools available for the assessment of a site-specific biodehalogenation potential. Physiological and quantitative studies conducted in laboratory microcosms can be combined with molecular tools to determine which dechlorinating bacteria and supporting community members are present, and can also help to develop rate information for the design of field implementation. Until existing technologies for rapid, inexpensive detection of specific microbial populations and the active expression of important genes are improved, laboratory microcosm studies continue to be a vital tool for assessing bioremediation potential in the field and providing a design basis for implementation. Small-scale field tests or in situ microcosms are currently the only alternative to laboratory microcosm studies. Ultimately, for full-scale site remediation, reliable, flexible models are needed which can aid in design and management of these complex systems. The determination of relevant, site-specific parameters for input into the models is another important potential use for laboratory microcosm studies.

Acknowledgments The authors gratefully acknowledge the financial support of the Environmental Security Technology Certification Program of the Department of Defense and the Air Force Research Laboratory, Tyndall Air Force Base, FL. REFERENCES 1. Adriaens P, Lendvay JM, McCormick ML & Dean SM (1997) Biogeochemistry and dechlorination potential at the St. Joseph aquifer-Lake Michigan interface. In: Alleman BC & Leeson A (Eds) The Fourth International In Situ and Onsite Bioremediation Symposium. Vol 3 (pp 173-178) Battelle Press, Columbus, OH 2. Adrian L, Szewzyk U, Wecke J & Gortsch H (2000) Bacterial dehalorespiration with chlorinated benzenes. Nature 408:580-583 3. Alleman, BE, Morse, JM, Snyder, F, Ackert, L, Sewell, G, Gossett, JM & Fennell, DE (2000) Reductive anaerobic biological in situ treatment technology (RABITT) treatability test: Results from Cape Canaveral Air Station. Presented at the Second International Conference on Remediation of Chlorinated and Recalcitrant Compounds. Monterey, CA, May 22-25 4. Allen HE, Fu G, Boothman W, DiToro DM & Mahony JD (1991) Determination of acid volatile sulfide and selected simultaneously extractable metals in sediment. U.S.EPA 821/R91-100. US Environmental Protection Agency, U.S. Government Printing Office: Washington, DC

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depends on the redox potential of the terminal NAPL flow and biodegradation model. In: Reddi electron acceptor. Arch. Microbiol. 149:350-357 LN (Ed) Non-Aqueous Phase Liquids (NAPLs) in Subsurface Environments: Assessment and 23. Cox S (1999) personal communication Remediation (pp 478-489) ASCE, Washington, 24. Cozzarelli IM, Herman JS & Baedecker MJ (1995) Fate of microbial metabolites of DC hydrocarbons in a coastal plain aquifer: the role 12. Bradley PM & Chapelle FH (1996) Anaerobic of electron acceptors. Environ. Sci. Technol. mineralization of vinyl chloride in Fe(III)29:458-469 reducing aquifer sediments. Environ. Sci. 25. Delwiche EA, Pestka JJ & Tortorello ML (1985) Technol. 30:2084-2086 The Veillonellae: Gram-negative cocci with a 13. Bradley PM & Chapelle FH (2000) Acetogenic unique physiology. Ann. Rev. Microbiol. microbial degradation of vinyl chloride. 39:175-193 Environ. Sci. Technol. 34:2761-2763 14. Brockman FJ & Murray CJ (1997) Subsurface 26. DeWeerd KA, Concannon F & Suflita JM (1991) Relationship between hydrogen microbiological heterogeneity: Current consumption, dehalogenation, and the reduction knowledge, descriptive approaches, and of sulfur oxyanions by Desulfomonile tiedjei. applications. FEMS Microbiol. Rev. 20:231-247 Appl. Environ. Microbiol. 57:1929-1934 15. Brown, D, Burns, DA, Bell, M & Lee, MD (2001) Biologically-enhanced reductive 27. Dojka MA, Hugenholtz P, Haack SK & Pace NR (1998) Microbial diversity in a hydrocarbondechlorination. In: Magar V, Fennell D, Alleman and chlorinated solvent-contaminated aquifer B, Morse JL & Leeson A (Eds) Anaerobic undergoing intrinsic bioremediation. Appl. Degradation of Chlorinated Solvents Vol 6, Environ. Microbiol. 64:3869-3877 Battelle Press, Columbus, OH 16. Carney A (1996) M.S. Thesis, Cornell 28. Elberson MA, Tabinowski JA, Ebersole RC, Ellis DE & Hendrickson ER (2000) Detection University, Ithaca, NY and characterization of Dehalococcoides 17. Carroll AB, Fennell DE, Anguish TW, Gossett ethenogenes' 16S rRNA sequences found in JM & Zinder SH (2000) Molecular groundwater and soils contaminated with PCE characterization of TCE-contaminated sites. and TCE. Presented at the 100th General Presented at the 100th General Meeting of the American Society for Microbiology. Abstract QMeeting of the American Society for Microbiology. Abstract Q-125. Los Angeles, 129. Los Angeles, CA, May 21-25, 2000 CA, May 21-25, 2000 18. Chapelle FH (1997) Identifying redox conditions that favor the natural attenuation of chlorinated 29. Fennell DE (1998) PhD Thesis, Cornell University, Ithaca, NY ethenes in contaminated ground-water systems In: Proceedings of the Symposium on Natural 30. Fennell DE, Carroll AB, Gossett JM & Zinder SH (2001) Assessment of indigenous reductive Attenuation of Chlorinated Organics in Ground Water (Dallas, TX, 1996), EPA/540/R-97/504; dechlorinating potential at a TCE-contaminated US Environmental Protection Agency, (pp 19site using microcosms, polymerase chain 22), U.S. Government Printing Office: reaction analysis, and site data. Environ. Sci. Washington, DC Technol. 35:1830-1839 19. Cirpka O (1995) Influence of hydraulic aquifer 31. Fennell DE & Gossett JM (1998) Modeling the properties on reductive dechlorination of production of and competition for hydrogen in a tetrachloroethene. In: Hinchee RE, Leeson A & dechlorinating culture. Environ. Sci. Technol. Semprini L (Eds) Third International In Situ and 32:2450-2460 On-Site Bioreclamation Symposium Vol. 3 (pp 32. Fennell DE, Gossett JM & Zinder SH (1997) 25-34) Battelle Press, Columbus, OH Comparison of butyric acid, ethanol, lactic acid, 20. Cirpka OA, Windfuhr C, Bisch G, Granzow S, and propionic acid as hydrogen donors for the Scholz-Muramatsu H & Kobus H (1999) reductive dechlorination of tetrachloroethene. Microbial reductive dechlorination in a largeEnviron. Sci. Technol. 31:918-926 scale sandbox model. J. Environ. Eng. 125:861- 33. Flynn SJ, Loffler FE & Tiedje JM (2000) 870 Microbial community changes associated with a 21. Clement TP, Sun Y, Hooker BS & Petersen JN shift from reductive dechlorination of PCE to (1998) Modeling multispecies reactive transport reductive dechlorination of cis-DCE and VC. in groundwater. Ground Wat. Mon. Remed. Environ. Sci. Technol. 34:1056-1061 18:79-92 34. Freedman DL & Gossett JM (1989) Biological 22. Cord-Ruwisch R, Seitz H-J & Conrad R (1988) reductive dechlorination of tetrachloroethylene The capacity of hydrogenotrophic anaerobic and trichloroethylene to ethylene under bacteria to compete for traces of hydrogen methanogenic conditions. Appl. Environ.

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35. Gao J, Skeen RS, Hooker BS & Quesenberry RD (1997) Effects of several electron donors on tetrachloroethene dechlorination in anaerobic soil microcosms. Wat. Res. 31:2479-2486 36. Gerhardt P, Murry R, Wood W & Krieg N (1994) Methods For General and Molecular Bacteriology; ASM: Washington, DC 37. Gerritse J, Alphenaar A & Gottschal JC (1998) Ecophysiology and application of dechlorinating anaerobes. In: Wilson TE (Ed) Water Resources and the Urban Environment--98. Proceedings of the 1998 National Conference on Environmental Engineering (pp 227-232) ASCE, Reston VA 38. Gerritse J, Drzyzga O, Kloetstra G, Keijmel M, Wiersum LP, Hutson R, Collins MD & Gottschal JC (1999) Influence of different electron donors and acceptors on dehalorespiration of tetrachloroethene by Desulfitobacterium frappieri TCE1. Appl. Environ. Microbiol. 65:5212-5221 39. Gibson SA, Roberson DS, Russell HH & Sewell GW (1994) Effects of three concentrations of mixed fatty acids on dechlorination of tetrachloroethene in aquifer microcosms. Environ. Toxicol. Chem. 13:453-460 40. Gibson SA & Sewell GW (1992) Stimulation of reductive dechlorination of tetrachloroethene in anaerobic aquifer microcosms by addition of short-chain organic acids or alcohols. Appl. Environ. Microbiol. 58:1392-1393 41. Gibson SA & Suflita JM (1986) Extrapolation of biodegradation results to groundwater aquifers: reductive dehalogenation of aromatic compounds. Appl. Environ. Microbiol. 52:681688 42. Hackley KC, Liu CL & Trainor D (1999) Isotopic identification of the source of methane in subsurface sediments of an area surrounded by waste disposal facilities. Appl. Geochem. 14:119-131 43. Hansen KH, Ahring BK & Raskin L (1999) Quantification of syntrophic fatty bacteria in a mesophilic biogas reactor by oligonucleotide probe hybridization. Appl. Environ. Microbiol. 65:4767-4774 44. Hansen, TA (1993) Carbon metabolism of the sulfate-reducing bacteria. In: Odom JM & Singleton R, Jr. (Eds) The Sulfate-Reducing Bacteria: Contemporay Perspectives, (pp 21 -40) Springer-Verlag, New York 45. Harkness MR, Bracco AA, Brennan, Jr. MJ, Deweerd KA & Spivack, JL (1999) Use of bioaugmentation to stimulate complete reductive dechlorination of trichloroethene in Dover soil columns. Environ. Sci. Technol. 33:1100-1109 46. Haston ZC & McCarty PL (1999) Chlorinated

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FENNELL AND GOSSETT 20:67-73 70. Monserrate E & Häggblom MM (1997) Dehalogenation and biodegradation of brominated phenols and benzoic acids under iron-reducing, sulfidogenic, and methanogenic conditions. Appl. Environ. Microbiol. 63:39113915 71. Morse JL, Alleman BC, Gossett JM, Zinder SH, Fennell DE, Sewell GW & Vogel CM (1997) A treatability test for evaluating the potential applicability of the reductive anaerobic biological in situ treatment technology (RABITT) to remediate chloroethenes. A Draft Technical Protocol developed for the E n v i r o n m e n t a l Security Technology Certification Program, Department of Defense, U.S. Government Printing Office: Washington, DC 72. Odom JM, Tabinowski J, Lee MD & Fathepure BZ (1995) Anaerobic biodegradation of chlorinated solvents: Comparative laboratory study of aquifer microcosms. In: Hinchee RE, Leeson A & Semprini L (Eds) Bioremediation of Chlorinated Solvents, (pp 17-24) Battelle Press, Columbus, OH 73. Ohtsubo S, Demizu K, Kohno S, Miura I, Ogawa T & Fukuda H (1992) Comparison of acetate utilization among strains of an aceticlastic methanogen, Methanothrix soehngenii. Appl. Environ. Micro. 58:703-705 74. Parsons Engineering Science (1997) RCRA Facility Investigation Report For Facility 1381 SWMU 21 (DP-32) Volume 1 of 2. January. 75. Pavlostathis SG & Prytula MT (2000) Kinetics of the sequential microbial reductive dechlorination of hexachlorobenzene. Environ. Sci. Technol. 34:4001-4009 76. Peyton BM, Truex MJ, Skeen RS & Hooker BS (1995) Design of an in situ carbon tetrachloride bioremediation system. In: Hinchee RE, Leeson A & Semprini L (Eds) Third International In Situ and On-Site Bioremediation Symposium Vol 3 (pp 111-116) Battelle Press, Columbus, OH 77. Powell GE (1984) Equalisation of specific growth rates for syntrophic associations in batch culture. J. Chem. Tech. Biotechnol. 34B:97-100 78. Pulliam-Holloman TR, Elberson MA, Cutter LA, May HD & Sowers KR (1998) Characterization of a defined 2,3,5,6-tetrachlorobiphenyl-orthodechlorinating microbial community by comparative sequence analysis of genes encoding for 16S rRNA. Appl. Environ. Microbiol. 64:359-367 79. Rifai HS, Newell CJ, Miller RN, Taffinder S & Rounsaville M (1995) Simulation of natural attenuation with multiple electron acceptors In:

MICROCOSMS FOR ASSESSMENT OF BIODEHALOGENATION POTENTIAL Ward, H. (Ed) Intrinsic Bioremediation, Vol 1 (pp 53-59) Battelle Press, Columbus, OH 80. Rittmann BE & McCarty PL (2001) Environmental Biotechnology: Principles and Applications, McGraw-Hill Higher Education, New York 81. Robinson JA & Tiedje JM (1984) Competition between sulfate-reducing and methanogenic bacteria for under resting and growing conditions. Arch. Microbiol. 137:26-32 82. Rosner BM, McCarty PL & Spormann AM (1997) In vitro studies on reductive dechlorination of vinyl chloride dehalogenation by an anaerobic mixed culture. Appl. Environ. Microbiol. 63:4139-4144 83. Schink B & Friedrich M (1994) Energetics of syntrophic fatty acid degradation. FEMS Microbiol. Rev. 15:85-94 84. Schink B, Kremer DR & Hansen TA (1987) Pathway of propionate formation from ethanol in Pelobacter propionicus. Arch. Microbiol. 147:321-327 85. Sewell GW & Gibson SA (1991) Stimulation of the reductive dechlorination of tetrachloroethene in anaerobic aquifer microcosms by the addition of toluene. Envir. Sci. Technol. 25:982-984 86. Skubal KL, Haack SK, Forney, LJ & Adriaens P (1999) Effects of dynamic redox zonation on the potential for natural attenuation of trichloroethylene at a fire-training-impacted aquifer. Phys. Chem. Earth (B). 24:517-527 87. Slenders H, Bosma T, Gerritse J, Borger A, Baartmans R, Hofstra S, de Sain H, Hetterschijt R, te Street C & de Kreuk H (1999) Bioremediation of chlorinated solvents in peat and natural attenuation of plume. In: Leeson A & Alleman BC (Eds) The Fifth International In Situ and Onsite Bioremediation Symposium Vol 2 (pp 141-146) Battelle Press, Columbus, OH 88. Smatlak CR, Gossett JM & Zinder SH (1996) Comparative kinetics of hydrogen utilization for reductive dechlorination of tetrachloroethene and methanogenesis in a anaerobic enrichment culture. Environ. Sci. Technol. 30:2850-2858 89. Sorenson KS, Jr. (2000) PhD Thesis; University of Idaho, Moscow, ID 90. Sorenson KS, Jr., Peterson LN & Ely RL (1999) Enhanced reductive dechlorination of TCE in a basalt aquifer. In: Leeson A & Alleman BC (Eds) The Fifth International In Situ and On-site Bioremediation Symposium, Vol 2 (pp 147-155) Battelle Press, Columbus, OH 91. Stiber NA, Pantazidou M & Small MJ (1999) Expert system methodology for evaluating reductive dechlorination at TCE sites. Environ. Sci. Technol. 33:3012-3020 92. Stover MA (1993) MS Thesis; Cornell

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93. Sun Y, Petersen JN, Clement TP & Hooker BS (1996) A modular model for simulating natural attenuation of chlorinated organics in saturated ground-water aquifers. In: Proceedings of the Symposium on Natural Attenuation of Chlorinated Organics in Ground Water (Dallas, TX, 1996), EPA/540/R-97/504; US Environmental Protection Agency, (pp 170), U.S. Government Printing Office: Washington, DC 94. Tandoi V, DiStefano TD, Bowser PA, Gossett JM & Zinder SH (1994) Reductive dehalogenation of chlorinated ethenes and halogenated ethanes by a high-rate anaerobic enrichment culture. Environ. Sci. Technol. 28:973-979 95. Tonnaer H, Alphenaar A, de Wit H, Grutters M, Spuij F, Gerritse J & Gottschal JC (1997) Modelling of anaerobic dechlorination of chloroethenes for in situ bioremediation. In: Alleman, BC & Leeson, A (Eds) The Fourth International In Situ and On-Site Bioremediation Symposium, Vol 5, (pp 591596) Battelle Press, Columbus, OH 96. USEPA (1991) Site characterization for subsurface remediation. EPA/625/4-91/026; US Environmental Protection Agency, U.S. Government Printing Office: Washington, DC 97. Verce MF, Ulrich RL & Freedman DL (2000) Characterization of an isolate that uses vinyl chloride as a growth substrate under aerobic conditions. Appl. Environ. Microbiol. 66:35353542 98. de Vries W, van Wyck-Kapteyn MC & Stouthamer AH (1973) Generation of ATP during cytochrome-linked anaerobic electron transport in propionic acid bacteria. J. Gen. Microbiol. 76:31-41 99. Wiedemeier TH, Swanson MA, Moutoux DE, Gordon EK, Wilson JT, Wilson BH, Kampbell DH, Hansen JE, Haas P & Chapelle FH (1996) Technical protocol for evaluating natural attenuation of chlorinated solvents in groundwater; EPA/600/R-98/128; US Environmental Protection Agency, U.S. Government Printing Office: Washington, DC 100. Willis M, Shoemaker C, Gossett J & Fennell D (1999) Applications of a competitive hydrogenotrophic biological dechlorination transport model for groundwater remediation. In: Leeson A & Alleman BC (Eds) The Fifth International In Situ and On-Site Bioremediation Symposium, Vol 2 (pp 27-33) Battelle Press, Columbus, OH 101. Wilson, BH, Wilson, JT & Luce, D (1997) Design and interpretation of microcosm studies

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Chapter 16 CHLORINATED ORGANIC CONTAMINANTS FROM MECHANICAL WOOD PROCESSING AND THEIR BIOREMEDIATION M. MINNA LAINE 1, MINNA K. MÄNNISTÖ 2, MIRJA S. SALKINOJA-SALONEN 3 AND JAAKKO A. PUHAKKA2 1

Applied Environmental Biotechnology Unit, Finnish Environment Institute, Helsinki, Finland Institute of Environmental Engineering and Biotechnology, Tampere University of Technology, Tampere, Finland 3 Department of Applied Chemistry and Microbiology, University of Helsinki, Finland 2

1. CHLOROPHENOLIC CHEMICALS USED IN TIMBER AND LUMBER TREATMENT Among the man-made chemicals, chlorophenols have been most widely used for impregnation or short-term preservation of wood and wooden products (46, 90, 141). In Finland, approximately 30,000 tons of chlorophenols, mostly with the trade name “Ky-5”, were used by the timber industry to impregnate wood until 1988. The Ky-5 preservative consisted mainly of 2,3,4,6-tetrachlorophenol (TeCP), 2,4,6-trichlorophenol (TCP) and pentachlorophenol (PCP). It also had dimeric impurities of polychlorinated dibenzodioxins (PCDDs) and polychlorinated dibenzofurans (52). This preparation was used in sawmills for timber protection, either by the dipping or spraying of an 1 to 5% alkaline chlorophenol solution. Due to work practices and accidental releases, the soil, surface, and ground water adjacent to the sawmills was often contaminated. The use of chlorophenolic wood preservatives has been reviewed by Kitunen (52).

1.1. Physico-Chemical Characteristics of Chlorophenols Chlorinated phenols are weak acids and thus exist as neutral phenols or dissociated phenoxide ions in the environment. The occurrence of the protonated and unprotonated species depends on the ambient pH in relation to the dissociation constant. For example, when the the neutral form predominates and when the the ionized form predominates. Thus, at neutral pH, PCP occurs mainly as its ionized form, whereas TCP is present in both forms at roughly equal proportions. The properties and environmental fate of the neutral and ionized forms differ markedly. Ionization affects Dehalogenation: Microbial Processes and Environmental Applications, pages 421–442 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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the solubility and sorption and therefore the mobility of polychlorophenols. With an increase in pH from 4 to 8.5, the predominant form of PCP changes from the protonated to unprotonated form. Under these conditions, sorption decreases by three- fold whereas solubility increases by three orders of magnitude (117).

1.2. Impurities of Chlorophenol Preparations Chlorophenol-contaminated sawmill soil in Finland is often also contaminated with polychlorinated dibenzo-p-dioxins and dibenzofurans (PCDD/Fs), as well as with polychlorinated phenoxyphenols (PCPPs), polychlorinated diphenyl ethers, and polychlorinated biphenyls (PCBs). These compounds are all present as impurities in the technical wood preservative Ky-5 that was produced and used in Finland and in other Nordic countries for many years (44, 45, 51, 91, 105). Chlorinated dioxins and other dimeric compounds may also be biologically generated in contaminated soil. For example, white rot fungi may form PCDD/Fs from chlorophenols using peroxidase enzymes (99), or from chlorinated anisyl compounds that they produce de novo (22; see Chapter 6).

2. CONTAMINATION OF SAWMILL ENVIRONMENTS 2.1. Scale of the Problem Nearly 800 former Finnish sawmill sites are contaminated with chlorophenols, PCDD/Fs, and phenoxyphenols originating from the commercial chlorophenol mixture Ky-5 (129). By the year 1997, 56 of these sites had been remediated, at least 36 of them by composting (136). The treatment of timber by Ky-5 was done either by dipping the timber in 1-2% sodium chlorophenolate solution, or by spraying the timber with the solution in a spraying hood (52). The most common practice was the dipping treatment after which the preservative was allowed to drip to the ground (52). Chlorophenols have, therefore, spread to the environment during normal sawmill operations. Accidental and intentional discharges of chlorophenol storing tanks have also occurred and as a result, severe contamination from soil around several sawmills has been reported (51,53,130). Due to their higher lipophilicity, the dimeric Ky-5 impurities (e.g., PCDDs and PCDFs) are retained in the topmost 5 cm of soil, whereas chlorophenols contaminate soil at great depth and leach down to the groundwater (51). As a consequence, groundwater and surface waters have been seriously contaminated around several sawmills (51, 64).

2.2 Environmental Fate of Contamination Chlorophenols are recalcitrant compounds that have been used for decades to impregnate wood, and many residues can be found in the environment long after the use of the chlorophenols has been discontinued. Chlorophenols are incorporated into soil to form a so-called “bound residue”, or non-extractable fraction, principally by two mechanisms: sorption and oxidative coupling (15). These processes affect bioavailability (see Chapter 10). Sorption can be physical sorption on surfaces of the solid material

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(adsorption) or partitioning into the solid material (absorption) (4), whereas oxidative coupling involves covalent bonding to form dimers, oligomers or polymerized fraction, and is usually mediated by enzymatic reactions in soil (15). Sorption of chemicals in soil appears to occur in two stages: an initially rapid stage followed by a slow stage (69). The slow sorption can be defined as “aging”: the aged or desorption-resistant fraction of the chemicals in soil may result from a slow diffusion of these molecules within the matrix of soil organic matter, or small pores of soil aggregates (4, 36). Covalent bonds in polymerization reactions are formed with chemicals that have functional groups similar to humic monomers (14). It has been demonstrated that a large part of the PCP present in contaminated soil is covalently bound to the soil organic matter, primarily to humic acid, by the action of white-rot fungi (108, 109). Oxidoreductive enzymes such as manganese peroxidase, lignin peroxidase and laccase are involved in copolymerization reactions of chlorophenols and ferulic acid, a precursor of humic acid (23, 35, 108). Inorganic chemicals, such as birnessite (manganous manganite, 23) and clay minerals can also catalyze oxidative reactions (4, 135). The nature of soil organic matter influences the extent of sorption and desorption of chlorophenols in soil. Benoit et al. (13) studied sorption of 4-chlorophenol (4-CP) and 2,4-dichlorophenol (2,4-DCP) to wheat straw, chestnut wood, Kraft lignin, composted straw and soil humic acid, and found that the sorption of 4-CP and 2,4-DCP on lignin and composted straw was greater than on humic acid, but desorption from humified materials was less than from fresh organic materials, indicating weaker interactions with the latter. The role of bio tic and abiotic reactions was further studied (12) during straw composting, and it was found that the formation of the bound residue, or immobilization of chlorophenols, in composted straw was mainly due to biological activity. Schäfer and Sandermann (112) have suggested that cell cultures of wheat metabolized PCP to tetrachlorocatechol, which in turn was either conjugated or incorporated into the lignin cell wall. Peat has also been shown to sorb PCP irreversibly from wastewater (137).

2.3. Public Health and Ecotoxicological Concerns The fate and potential hazard of organic halogen compounds in contaminated soil are affected by (bio)transformation reactions of the chemicals in soil. The compounds may volatilize, become mineralized or transformed by microbes. Chlorophenols may be hydroxylated, methoxylated or polymerized by soil microbes at sawmill sites. Hence, the chlorophenols in nature are prone to O-methylation to the corresponding chloroanisoles (39-41, 88, 122), dimerization or polymerization (8, 14, 61, 99, 138), as well as other metabolic alterations (15, 34). These biotransformation products may cause ecotoxicological risks and impact on the soil environment. For example, methylation of chlorophenols produces chlorinated anisoles which are more lipophilic and become more susceptible to bioaccumulation. Since chlorophenols are relatively water-soluble, they may leach into the groundwater and thus cause a threat to humans and the environment. Toxicologically important impurities of wood preservatives, e.g., PCDD/Fs, are less mobile since they adsorb to the organic matter in soil (2). Food is the major overall source of PCDD/Fs exposure for humans (128). Chickens and cows have been found to accumulate

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PCDD/Fs from contaminated soil, even at concentrations as low as 45 ppt, measured as international 2,3,7,8-TCDD toxic equivalents, I-TEQ (equivalent to as total PCDD/F concentration) (118). The main route of PCDD/F exposure from contaminated soil to humans is through skin contact or inhalation of dust particles. The release and dissemination of PCDD/Fs on airborne particles has been monitored during chlorophenol composting (60). In the cited study, the PCDD/Fs concentrations were determined as a function of particle size. The congener distribution of PCDD/Fs was similar in the collected air particle fractions as to that of the compost windrows, and the concentrations of the airborne PCDD/Fs were 1000-fold higher than the previously reported atmospheric background values (60). The average human inhales approximately of air per day (72). Composts were usually turned six times during the summer period. Every mixing lasted approximately 2 hours, but a working day at a sawmill lasts for 8 hours. Thus, workers in the sawmill area inhale approximately of air during that time. Based on these assumptions a cumulative dose of particles containing PCDDs and PCDFs as calculated per average daily intake was between 0.1 and 4.8 ng I-TEQ, during mixing of the compost windrows in one summer season. That dose equals the amount of PCDDs and PCDFs ingested by eating 1 kg of Finnish lake fish, such as pike, which contains 0.2 to 0.5 ng of PCDDs and PCDFs as I-TEQ per kg, or by eating 100 g of Baltic salmon (134). For a 70 kg person the daily intake will be between 1.4 to 69 pg This maximum value in summer season is about 20 times higher when compared to the acceptable daily intake (ADI) given by the Nordic countries, 35 pg It should be noted, however ADI value is given for the lifetime exposure and not for the occupational exposure only.

3. MICROBIOLOGY OF CONTAMINATED SITES Although several Gram-positive and Gram-negative polychlorophenol-degrading bacteria have been isolated from contaminated soils, relatively few studies have assessed total microbial diversity at polychlorophenol contaminated sites. Polychlorinated phenols may significantly alter the composition of microbial communities in contaminated soil and groundwater. As potent broad-scale biocides, chlorophenols present at relatively low concentrations may reduce microbial biomass (18). Tolerance to polychlorophenols may vary greatly among different microorganisms. For example, Gram-positive bacteria are generally more sensitive to PCP than Gram-negatives (78, 110,144). High doses of PCP in soil have been reported to increase the share of Gramnegative bacteria, while the numbers of Gram-positives decrease (110). Chaudri et al. (18) reported a 91% decrease in soil biomass content, 6 months after application of 50 mg/kg of PCP. A much higher dose of PCP, 200 mg/kg over a similar timeframe, reduced microbial biomass only by 22%. The authors suggested that low PCP concentrations may have eliminated bacteria that are more sensitive to polychlorophenols, whereas high concentrations may have selected for PCP-tolerant bacteria that could grow and degrade PCP with much less competition (18). Among Gram-negative bacteria, sphingomonads appear remarkably tolerant to PCP (104, 106, 144). Moreover, several species of this group can degrade PCP and other polychlorinated phenols as sole carbon sources (24, 76, 78, 93, 94). The taxonomy of

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the genus Sphingomonas was recently revised by Takeuchi et al. (124) and several species have been transferred to the new genera Novosphingobium, Sphingobium and Sphingopyxis (124). Consequently, polychlorophenol-degrading sphingomonads have been (re)classified as species of Sphingobium and Novosphingobium (124, 126). Their degradation ability, together with a high tolerance to the chlorinated organics, may lead to the selection of these types of microorganisms at contaminated sites. Beaulieu et al. (11) have reported on the predominance of Sphingomonas species in PCP-contaminated soil. In their study of PCP-enriched soil, 96% of the examined clones generated from PCR-amplified 16S rRNA genes were related to the genus Sphingomonas. This indicates that PCP contamination may result in a decrease of phylogenetic diversity and the selective enrichment of PCP-degraders, in particular sphingomonads, in soil. A number of studies have shown adaptation of microorganisms to the existing conditions in contaminant plumes (70; for review, see 32). In environments contaminated with chlorinated organic compounds such as PCP, the contaminant may serve as a selective pressure, and support specific microbial populations which either utilize or detoxify the organic contaminant. A change in numbers or a shift in the diversity of the microbial community may ensue. This can be observed as increased numbers of total bacteria, increased numbers of contaminant-degrading bacteria, and/or increased degradation rates of the contaminant in the area of impact, as compared to sediments obtained from an uncontaminated reference site (133). Very few studies have been made, however, in chlorophenol-contaminated sites. Bacterial abundance and diversity of a chlorophenol-contaminated aquifer in Southern Finland was investigated by Männistö et al. (76, 78). In this study, up to cells/ml and CFU/ml were enumerated from groundwater samples obtained within the area of contamination, as well as surrounding aquifer. No correlation was found between the extent of chlorophenol contamination and the number of microorganisms in various groundwater wells along the aquifer (78). A total of 190 bacterial strains isolated from inside and outside the chlorophenol plume were characterized. These isolates represented a large spectrum of bacterial taxa, as indicated by a cluster analysis based on fatty acid compositions of the isolates. The proportion of chlorophenol-degrading bacteria was significantly higher among isolates from inside the chlorophenol plume (57%), than among isolates originating from outside the plume (7%) (78). This indicates a selective enrichment of chlorophenol-degraders, as a consequence of chlorophenol contamination that had been ongoing at the site for at least 25 years (64). Furthermore, the 17 chlorophenol-degrading isolates obtained from inside the plume comprised a diverse group of bacteria, as indicated by whole cell fatty acid compositions and partial 16S rRNA gene sequences (78). Based on phylogenetic analyses, the 17 chlorophenol degraders represented 5 different phylogenetic groups: Grampositive bacteria with high G+C, and the Cytophaga/Flexibacter/Bacteroides group.

4. BIODEGRADATION STRATEGIES Several bacteria and fungi can mineralize chlorophenols (for review, see 37; Chapters 3 and 6). The mineralization of chlorophenols may proceed aerobically (6,21, 114; for review, see 38) or via reductive dehalogenation (43, 83; for review, see 85). However, reductive dehalogenation does not always lead to complete mineralization;

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instead, metabolic products may accumulate (19,116). Several chlorophenol-degrading microorganisms, including both Gram-negative and Gram-positive bacteria, have been isolated and their properties have been carefully investigated in the laboratory (see reviews 5, 27, 37, 38, 79, 85, 132). Many Gram-negative bacteria have a common pathway to degrade PCP and other highly chlorinated phenols. The pathway involves an initial oxygenolytic dechlorination of PCP to tetrachloro-p-hydroquinone by PCP-4-monooxygenase. It has been suggested that the PCP monooxygenase has evolved to accommodate PCP from a previously existing flavin monooxygenase (21). This dehalogenase ofmonooxygenase type and the corresponding pcpB gene share homology among Sphingomonas chlorophenolica (formerly Flavobacterium sp.) ATCC 39723 (97), Sphingomonas sp. RA2 (104), Arthrobacter sp. ATCC 33790 (111), and Pseudomonas sp. strain SR3 (106), as well as other Pseudomonas spp. (67). According to studies of Ederer et al. (24), the homology of pcpB among these four PCP-degrading microorganisms is exceptionally high and due to additional culture characteristics it was suggested that all of these strains be reclassified into the genus Sphingomonas of the (50, 94). Moreover, all four strains were members of the same species, Sphingomonas chlorophenolica, which was reclassified as Sphingobium chlorophenolicum (124). The sequencing of new degrading genes from environmental isolates and clones has shown that pcpB –type of genes are also found in novel bacterial strains. All of these bacteria, primarly identified as sphingomonads, had been isolated from PCP-contaminated soil, water, or enrichment cultures. In conclusion, most of the Gram-negative bacteria able to degrade PCP appear to belong to the genus previously called Sphingomonas, and they have very similar degradation pathways, enzymes, and genes to do so. In addition, Männistö et al. (78) have isolated and characterized 17 TeCP degrading bacteria from contaminated boreal groundwater, that were members of and and the Cytophaga/Flexibacter/Bacteroides group. Several polychlorophenol degraders isolated from the contaminated groundwater were members of genera other than the genus Sphingomonas, including Pseudomonas, Ralstonia, Caulobacter, Flavobacterium and Herbaspirillum (77, 78). Chlorinated hydroquinones are the initial metabolites during degradation of PCP and other polychlorinated phenols by many Gram-positive chlorophenol-mineralizing strains, including Mycobacterium chlorophenolicum (7, 42), Mycobacterium fortuitum (41, 92), and Streptomyces rochei 303 (31). In contrast to Gram-negative bacteria, these Grampositive bacteria have different PCP-hydroxylating enzymes (37, 50) and the further degradation of chlorinated hydroquinone follows a different pathway (38). For less chlorinated phenols, bacteria have yet different degradation pathways, including the oxidation of mono- or dichlorophenols to corresponding chlorocatechols with dehalogenation occurring after ring cleavage (38). White rot fungi have been reported to degrade a wide range of lignocellulosic and xenobiotic compounds (143). The fungal metabolism of PCP by Phanerochaete chrysosporium may form either pentachloroanisole (61) or 2,3,5,6,-tetrachloro-2,5cyclohexadiene-l,4-dione (84). However, it has been shown that one or more bacterial species are associated and coexist with Phanerochaete chrysosporium (115). These bacteria are not eliminated from the fungus by treatment with either heat, antibiotics, ozone, or sodium hypochlorite (115). One ofthese bacteria, Agrobacterium radiobacter,

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appears to also synergistically associate with other bacteria to degrade PCP, and has been isolated together with Pseudomonas sp. and Flavobacterium gleum from a mixed PCP-degrading culture (147). Since other bacteria associated with fungi are efficient degraders of aromatic compounds (115), it can not be disregarded that degradation of xenobiotic compounds by Phanerochaete chrysosporium may in fact be due to a symbiotic (synergistic) relation between bacteria and the fungus.

5. EFFECTS OF ENVIRONMENTAL CONDITIONS ON BIODEGRADATION Degradation rates of chlorophenols are usually higher in the laboratory than in the field, since laboratory-scale incubations are conducted generally under more favorable and controlled conditions (see also Chapter 15). In fact, the oftentimes poor survival of laboratory cultures inoculated into contaminated soils may limit their use for effective bioremediation in the field. Bioavailability may also play an important role in the biodegradation of contaminants (see Chapter 10). To promote bioremediation through increased bioavailability of chlorophenol contaminants, as well as the survival of inoculated pure cultures, bacteria immobilized in physical supports, such as polyurethane (16, 96), sodium alginate (71), or K-carrageenan (17), has been shown to improve degradation of chlorophenols, at least on a laboratory scale. It is often assumed that low temperatures decrease biodegradation rates significantly, and therefore make the bioremediation of organic compounds non-feasible in cold climates. Bacterial degradation of polychlorophenols has been primarily been studied in the laboratory at temperatures higher than 20°C using mesophilic organisms (for reviews, see 101, 102). In general, biodegradation rates at low temperatures can be estimated from literature values determined at 20 to 30°C, by assuming that the degradation rate is decreased by 50% for each 10°C decrease in temperature. In one series of studies, polychlorophenol degradation has been detected down to 4°C in fluidized-bed enrichments (47, 81). As expected, large decreases in degradation rates due to lowering temperature have been observed (81, 145). An unusually strong effect of temperature on chlorophenol degradation in fluidized-bed reactors was reported at temperatures ranging from4 to 16.5°C (81). In these studies, the highest degradation rate was observed at the highest temperature (16.5°C), and a 10°C decrease in temperature generally decreased chlorophenol degradation rates by a factor of seven. When assessing temperature effects on microbial activity, it is important to recognize that some microorganisms exhibit different temperature optima depending on substrate. For example, Wittmann et al. (145) studied the well-known PCP degraders, Sphingobium chlorophenolicum (formerly Sphingomonas chlorophenolica) RA2 and Mycobacterium chlorophenolicum PCP-1, which have optimum growth temperatures on glucose at 33 °C and 30°C, respectively. The optimum temperature for PCP degradation by S. chlorophenolicum was similar to its optimum growth temperature on glucose, but M. chlorophenolicum exhibited highest PCP-degrading activity at 41°C, which was 10°C higher than its optimum temperature for growth. In contrast, the effect of temperature on chlorophenol degradation by cold-adapted groundwater isolates has been shown to be different (75, 77). Although the optimum growth temperatures of these isolates, Nocardioides sp. K44 and Sphingomonas spp. 74, and Sphingomonas spp. MT1 [reclassified as Novosphingobium sp. (126)], were 25°C, 25°C, and 23°C, respectively

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(75), 2,3,4,6-TeCP was degraded faster at 8°C than at room temperature by all of these isolates, indicating a much colder temperature optimum for chlorophenol degradation (77).

6. REMEDIATION OF CHLOROPHENOL-CONTAMINATED SOIL USING WINDROW COMPOSTING Composting is an accelerated biological process for effectively degrading organic compounds in soil. Chlorophenols can be effectively degraded by composting. During a three-year, full-scale composting operation of contaminated sawmill soil, at least 93% of the chlorophenols contaminants were removed in all compost windrows containing a mixture of contaminated soil and straw compost or bark chips (57). Biodegradation of chlorophenols was efficient and fast, irrespective of the addition of a chlorophenoldegrading inoculum. In general, it was observed that indigenous soil microorganisms degraded chlorophenols under suitable conditions (59). Complete mineralization of chlorinated contaminants should always be the goal of the bioremediation process, since partially degraded metabolites may be more toxic or mobile (water soluble) than the parent contaminants. Composting in biopiles, i.e., windrow composting, is very simple and cost-effective method. It is important, however, to maintain appropriate environmental conditions for effective microbial mineralization of chlorinated pollutants in soil. A special bench or pilot-scale treatability evaluation needs to be performed to demonstrate the feasibility of the process before full-scale implementation. Before construction of a full-scale treatment system, it is necessary to assess the available nutrients and the activity of the pollutant-degrading microbial population. Laboratory studies usually provide the required information, but it is important to recognize that laboratory scale experiments are generally conducted under more favorable and controlled conditions. For example, a constant room temperature above 20°C improves chlorophenol degradation in the laboratory, but is difficult to maintain during outdoor composting. Bioremediation should be performed so that it does not cause further contamination of the environment or adverse health effects. Thus, contaminated soil is excavated and placed in a specially prepared treatment area. During the excavation, workers should be protected against dust. The windrows are constructed on a plastic bed, such as a high density polyethylene (HDPE) liner, which has an inclination towards separate underdrainage. After the contaminated soil has been placed on the liner, the plastic bed is covered with an insulation layer of bark. Alternatively, windrows can be constructed on a clay foundation surrounded by a berm. In some cases, native topsoil can act as a permeable barrier to contaminant migration. The leachate from the treatment system is recovered and recycled through the irrigation system, thus eliminating the need for disposal of potentially contaminated water. Use of a vacuum source to draw air through the treatment system facilitates control and treatment of volatile compounds in the offgas, if required. The best solution is to perform composting in a closed environment (e.g., tent or building in a special composting facility), both for control of leachates, and to overcome adverse weather effects. During the actual construction of a windrow, the soil is homogenized, often with a screened crusher, to obtain relatively homogeneous dispersal of the contaminated

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materials, and to remove stones and other non-reactive materials prior to composting. The compost windrows contain chlorophenol-contaminated soil, nutrients and lime, and additional amendments. The supplementary admendments often include inoculants of mixed microbial populations, nutrient-rich material (such as composted straw) with bacterial consortium (56), or pure cultures of pollutant-degraders, depending on the soil type and the degree of contamination. Bulking agents such as straw compost (55, 56) and bark chips, that contain high amounts of easily available carbon sources and active microbial populations can enhances chlorophenol degradation in contaminated soil, as well as improve overall microbial functionality and activity (Figure 16.1 A). The ratio of the materials is roughly 2 parts of contaminated soil to 1 part of bulking agent by volume. Nitrogen and phosphorus nutrients are often applied as commercial fertilizers. Each inoculant is mixed with contaminated soil, pH is adjusted to near neutral with fine granular lime, nutrients are added in the form of a nitrogen-rich commercial fertilizer, and bark chips (fresh softwood) as supporting and aerating material are added to enhance microbial degradation. The moisture content of the compost windrows is adjusted to about 26%, and is maintained by watering when necessary. After all the materials are piled in a sandwich-type, the windrows are properly mixed, sampled, and covered with liner. Oxygen supply is needed for aerobic degradation of contaminants in compost windrows. Composting can be performed in dynamic or static windrows. The dynamic style of windrows can be mechanically aerated and mixed with several different tools: a mechanical clamshell, a windrow turner (mixing drum machine), or by using an excavator which turns the windrows by moving the compost soil 5 m along the windrow length is opposing directions (Figure 16.1B). In a static style of windrow, air is circulated through the compost via a perforated pipe placed within the compost bed. In the case of static biopiles, the spacing of aeration pipes, adjustment to an optimal air flow rate, and vacuum removal of off-gases need to be determined on the basis of soil permeability and the anticipated size of the treatment system. Static windrows are better in preventing the spread of airborne contaminants from the biopiles, whereas mixing of the dynamic windrows provides better mixing and improves the close contact of degrading microbes to contaminants. The composting operation can be evaluated indirectly by monitoring microbial activity in the windrows. This is accomplished by analyzing the soil air composition in the compost windrows. Before each mixing, soil off-gas, collected in perforated tubes installed inside the windrows is analyzed to determine how much oxygen is consumed, as well as the amounts and water produced during the most active phases of contaminant, e.g., chlorophenol, degradation (56). Lower levels of microbial activity are reflected by a drop in temperature, relatively high levels, and low and water concentrations, in the pore spaces of the compost windrow (56). With elevated microbial activity, a reverse profile is observed, i.e., low but high and water levels. In the case of chlorinated contaminants such as chlorophenol, that release Cl- ions during biodegradation, the formation of HC1 lowers the pH of the compost windrows, from near neutral to slightly acidic (56). Ambient pH may also be lowered during composting, by the catabolism of secondary carbon sources to organic acids and Augmentation with chlorophenol-degrading pure cultures can significantly improve chlorophenol degradation in soil, provided that mineralization occurs. The augmentation

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of a windrow compost with a chlorophenol-mineralizing strain of Mycobacterium chlorophenolicum, resulted in chlorophenol removal rates of 0.8 and 9.5 mg (kg dry where initial contamination levels ranged from 10 to and from 18 to respectively (37). Removal rates in field experiments with the white rot fungi, Phanerochaete chrysosporium and Phanerochaete sordida,

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were from 4.3 to with contamination levels from 74 to 1010 mg In these studies, mineralization was not monitored, but up to 15% methylation of PCP with accumulation of pentachloroanisole was observed (62). According to earlier laboratory studies by Lamar and his coworkers (63), Phanerochaete spp. mineralized PCP to a small extent (2 to 12% per month, or 0.1 to 0.4 % per day) in liquid culture, but in a soil environment, most of the observed PCP depletion by Phanerochaete spp. was due to methylation of the parent compound, or conversion to non-extractable compounds. Relatively good degradation rates are also achieved by amendment with mixed culture. In pilot-scale composting studies by Valo and Salkinoja-Salonen (129), using a chlorophenol-degrading enrichment culture, the removal rate of chlorophenols was 2.2 mg with initial concentrations from 20 to In studies by Mahaffey and Sanford (73) using a mixed, chlorophenol-degrading culture in slurry bioreactors, the chlorophenol removal rate was at a contamination level ranging from 0.5 to 100 mg chlorophenols In both of these cited cases, the laboratory studies confirmed that the chlorophenol removal was due to mineralization (73, 129). In a solid-phase treatment of soil in the field, Seech et al. (113) observed a chlorophenol removal rate of in soil inoculated with Pseudomonas resinovorans, with contamination levels from 6 to 680 mg The inoculated strain mineralized 12% of radiolabelled PCP in 48 hours (6% per day) in the laboratory. When the contaminated soil itself has a sufficient number of chlorophenol-degrading (mineralizing) microorganisms, further inoculation will not enhance degradation. For example, in a pilot-scale study of composting with an indigenous consortium of chlorophenol-degraders (56), the chlorophenol removal rate was about 0.6 mg (kg dry at low contamination levels, and from 2.2 to at high contamination levels, (from about 50 to 1800 mg (kg dry In parallel laboratory tests, the mineralization of radiolabeled PCP was 60% per month, corresponding to 2% per day, assuming first order decay (56). In addition to determining contaminant concentrations and soil respiration profiles, there are several parameters that can be regularly monitored during composting to assess the efficacy of the operation. Monitoring of both physical and microbiological determinants is needed to ensure optimal composting conditions. Although composting is a cost-effective bioremediation method, the comprehensive additional sampling and analysis required for maintaining optimal operating conditions, can contribute significantly to overall costs. In addition to the previously mentioned soil respiration profiles, a number of other environmental determinants can be used to evaluate composting activity. For example, microbial status can be determined by conventional enumeration of microorganisms, on selective and general culture media. Microbial activity and community structure can be further assessed by establishing a substrate utilization pattern for a large range of substrates (59), or by using gene probes. The temperature gradient inside the windrows can be measured using a 2-m long temperature probe. The average temperature in the compost windrows rises to 36 to 40 °C, which is well above the ambient day temperature (56). During the winter, the degradation of chlorophenols may stop if the outdoor biopiles cannot keep the temperature above zero.

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In general, the amounts of total phosphorus and total nitrogen remain the same during the composting, although the amounts of bioavailable nutrients, e.g., soluble phosphorus and ammonium (and nitrate), are rapidly used by the microbes (59). Based on this observation, the analysis of total nutrient concentrations is not an informative parameter to follow during the composting process. Measurements of bioavailable nutrients, e.g., utilization of ammonium, nitrate and soluble P, provide a tool to better evaluate the microbial status during the composting. In the studies of Laine and coworkers (58), the toxicity of contaminants and their metabolites, as assessed by luminescent bacteria tests, was observed to decrease during composting; degree of toxicity followed the chlorophenol concentrations in the compost windrows. The threshold value for chlorophenol toxicity appeared to be 200 mg total chlorophenols per kg dry weight. Based on these results, the toxicity tests provided a fast turnaround, and also a promising tool for assessing toxicity changes in the chlorophenolcontaminated soil. However, the toxicity screening was not sensitive enough to detect chlorophenol concentrations that meet the remediation criteria for the clean-up of contaminated soil, which in this case was (dry wt) of total chlorophenols. Moreover, in assessing the environmental impact of soil contamination or of bioremediation products, a set of different trophic level biotests should be used, in order to see, for example, the toxic effect of PCDD/Fs remaining in contaminated soil. Some of the drawbacks of composting technology include the need for excavation, transportation, and treatment of the contaminated soil at a special composting site. Only biodegradable contaminants can be treated. For example, large amounts of highly chlorinated dibenzo-p-dioxins and dibenzofurans (PCDD/Fs) originating from the wood preservative Ky-5 were present in composted soil in mg per kg concentrations even after treatment (58). The congener composition of PCDD/Fs resembled that of the original wood preservative, indicating that PCDD/Fs were neither formed nor degraded during the composting process (58).

7. REMEDIATION OFCHLOROPHENOL-CONTAMINATEDGROUNDWATER Historically, the treatment of contaminated groundwater follows a general scheme. One of the earliest and still most widely used methods for the cleanup of contaminated groundwater entails an on-site “pump-and-treat” system for clean-up. The major limitation of such a system is often due to the difficulty of bringing, i.e., pumping, the contaminated groundwater to the surface for treatment (for review, see 66). Treatment at the surface is generally either by air stripping and recovery, or by biological treatment. Many organic contaminants are by nature hydrophobic, and tend to partition with the subsurface solids, thereby reducing their mobility (and recovery) in the aqueous phase that is recovered for remediation. In these cases, the volume of water to be pumped for treatment may be ten-fold greater than the volume of contaminated groundwater. Often, groundwater pumping is also needed for containment of contaminants. In these cases, the on-site treatment is a useful unit process in aquifer remediation. During recent years, the research and development of aquifer cleanup systems have focused on in situ and especially on intrinsic bioremediation. This general development is also seen in the bioremediation of chlorophenol contaminated groundwater.

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7.1. On-Site Bioremediation Strategies Owing to their solubility characteristics in an aqueous environment, i.e., groundwater, the sorption tendency of chlorophenols is relatively weak as compared to other groundwater contaminants such as hydrocarbons. Despite their relative increased solubility and mobility, the recovery and cleanup of a chlorophenol-contaminated aquifer by “pump and treat” technology is still impacted by limited chlorophenol solubility in the recoverable aqueous phase. Over the years, numerous studies of on-site chlorophenol bioremediation have been reported, but only a few have resulted in actual application (for reviews, see 37, 101, 102). This review summarizes only known, fullscale treatments. One of the first successful operations using a bioreactor on the surface, was the BioTrol system designed for PCP treatment (119). This high-rate system, with a chlorophenol load of 365 mg PCP/l/d, relied on inoculation with a Sphingomonas (formerly Flavobacterium) strain and achieved removal efficiencies of 96%; approximately 1 mg PCP/1 remained in the treated effluent. Another high-rate system was developed for cold-temperature applications in Finland (47, 48, 74, 81, 82). This fluidized-bed process was developed to treat water contaminated with a mixture of 2,4,6trichlorophenol, 2,3,4,6-tetrachlorophenols, and pentachlorophenol. The system functions at 8°C with a hydraulic retention time of 2.5 hours, and has been in operation for over 5 years. During periods of steady operation, its performance is characterized by over 99% chlorophenol removal and effluent concentrations ranging from 0.002 to 0.1 mg/1. Chlorophenol-contaminated groundwater often contains other chemicals used for wood treatment. Bioremediation systems have been used for groundwater containing multiple contaminants. Groundwater contaminated with a mixture of PCP and polycyclic aromatic hydrocarbons (PAH) has been treated in a two-stage system consisting of a continuously stirred tank reactor for PAHs, followed by PCP treatment on a fill-anddraw batch system (87). Another application for PAH/PCP groundwater uses up-flow aerobic fixed-film reactors (100). These multiple contaminant treatment systems are characterized by long treatment times and partial removal of chlorophenols.

7.2. In Situ Bioremediation In situ bioremediation technology aims to improve natural biodegradation processes by enhancing bioavailability of the contaminant and the increasing activity of the indigenous microbiota. In some case soil or groundwater may also be augmented with specific contaminant-degrading microorganisms, including pure cultures specifically selected for their ability to degrade the target contaminant(s) (89); genetically modified cultures that have been manipulated to enhance their degradation ability (54, 107); or selective enrichment cultures developed from the contaminated site (9, 98, 139). Bioaugmentation, i.e., inoculation with contaminant-degrading microorganisms may not always be successful, because the introduced microorganisms may not be able to compete with the indigenous microbiota and survive in the target environment for a prolonged time (68). In most cases, the degradation ability of the indigenous microbiota from a site is sufficient, and may be stimulated by improving the ambient environmental

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conditions. In situ bioremediation may include supplementation of one or more of the following: an electron acceptor, nutrients, electron donor (for anaerobic degradation) or primary substrate for cometabolic degradation (26, 80). The design of the in situ bioremediation system is highly site specific. In situ bioremediation technologies have been successfully applied for groundwater contaminated with petroleum compounds and chlorinated solvents (reviewed by 25,26). In most cases, aerobic biodegradation is faster and more extensive, and therefore most in situ remediation projects are engineered to use oxygen as the terminal electron acceptor. It should be noted, however, that nitrate can also serve as an electron acceptor to facultative aerobes in less oxidized environments. In contrast to the bioremediation of petroleum-contaminated soils, bioremediation of groundwater and subsurfaces contaminated with chlorinated organics generally follow a different strategy (see Chapters 14 and 15). As discussed in Chapter 3, reductive dechlorination serves as a major mechanism for the biodegradation of chlorinated organics, especially those with a high degree of chlorination, such as tetrachloroethene and trichloroethene. For these types of compounds, anaerobic, or even mixed, anaerobic/aerobic treatment schemes have been developed to adequately treat those compounds which may be less biodegradable under aerobic conditions (26). Chlorophenols generally are degradable under aerobic conditions, and therefore most in situ treatment schemes attempt overcome any potential oxygen limitations as might occur in contaminated subsurface environments. Due to the low solubility of oxygen, distribution of oxygen to the subsurface is a challenge. Oxygen may be added to the saturated subsurface physically by aeration, or chemically using oxygen releasing compounds such as hydrogen peroxide or magnesium peroxide (131). Hydrogen peroxide decomposes rapidly in soil and may be released prematurely to the atmosphere. Depending on the groundwater chemistry and soil type, the addition of hydrogen peroxide may also result in the formation of mineral precipitates that reduce hydraulic conductivity and thus water movement (142). Hydrogen peroxide is toxic at high concentrations and may inhibit biodegradation at concentrations greater than 100200 ppm (131). Magnesium oxide is used in so-called oxygen releasing compounds, where oxygen is released more slowly. Low concentrations of nutrients, particularly phosphorus, may limit biodegradation in groundwater and nutrient addition may be a key for enhanced biodegradation. In situ bioremediation is considered applicable for aquifers with hydraulic cm/s (125). In highly impermeable aquifers, the groundwater flow may be too slow for efficient distribution of electron acceptors, nutrients or other supplements (25). When applicable, in situ bioremediation is generally less costly than other remediation techniques, and transforms contaminants into harmless endproducts. If no natural enrichment of chlorophenol degraders in aquifers takes place, the possibilities for biological treatment in situ are limited. The study of Männistö et al. (76, 78) showed that a long-term chlorophenol contamination had resulted in development of an abundant and diverse chlorophenol degrading microbiota in a boreal aquifer. From pilot-scale experiments, the in situ activation of chlorophenol degradation in this case required only an oxygen supply (65). This indicated significant in situ bioremediation potential of the contaminated aquifer. The actual application of increasing the supply may, however, be limited by simultaneous iron oxidation and precipitation which may

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result in aquifer clogging (65). In another example, an amendment-strategy with degrader microbes was proposed as one possibility for the cleanup of a chlorophenol-contaminated aquifer (120). A PCP degrading Shingomonas (Flavobacteriuni) culture was encapsulated into microspheres of alginate and showed enhanced survivability and long-term PCP biodegradation in microcosms (121). Bioaugmentation may have potential especially in recently contaminated aquifer cleanup where intrinsic contaminant degraders have not yet been naturally enriched. The first in situ PCP remediation experiences were gained in Libby, Montana. (100,146). Pumping of oxygen and nutrient supplemented water to the aquifer resulted in PCP degradation as indicated by decreasing PCP concentrations as well as oxygen and nutrient consumption in the groundwater. To date, in situ bioremediation of chlorophenols has a limited number of applications, but the potential for this technology has the same promise as for other aerobically biodegradable contaminants.

8. FUTURE CHALLENGES (PCDDs & PCDFs) Polychlorinated dibenzo-p-dioxins and dibenzofurans (PCDD/Fs) are some of the most toxic man-made compounds in the world. The toxic mechanism of PCDD/Fs and related compounds is based on their binding to an Ah-receptor found in higher organisms. The Ah-receptor-mediated mechanism of action, however, is only one of the several ones suggested as the toxic mechanism of PCDD/Fs (140). Some studies indicate that chlorophenols, rather than the PCDDs and PCDFs, are the main cause for detrimental health effects in contaminated sawmill areas (134). Exposure to wood preservatives containing PCP, and a correlation of blood PCP levels to gynecological and endocrine dysfunction in women have been reported (29). PCP may, therefore, play a role in infertility. Removal of chlorophenols from a contaminated sawmill soil by composting, for example, was shown to reduce the risk for substantial spreading of the chlorophenol pollution (58, 59), but the problem with PCDD/Fs remains. PCDDs and PCDFs are less mobile than chlorophenols since they are not soluble in water and adsorb stronger to the organic matter in soil (2). However, they may spread from the soil via air particles (see section 2.3. “Public Health and Ecotoxicological Concerns” above). This needs to be kept in mind when planning any remediation actions for contaminated soil. Although non-chlorinated dibenzo-p-dioxin and dibenzofuran biodegrade to some extent (28,49, 86, 143), only a few reports on biological breakdown or dehalogenation of PCDD/Fs exist (1, 3, 10, 30, 33, 123, 127). Bioremediation of PCDD/Fs is hindered by their low availability to microorganisms and their strong sorption to soil. Chlorophenol-contaminated soil often contains PCDD/Fs as inpurities, therefore it seems impossible to clean sawmill soil completely using biological treatment only. This is especially true for aerobic processes. How much of PCDD/Fs can be degraded through sequential anaerobic-aerobic treatments (e.g., lowering redox potential after aerobic treatment), or by phytoremediation, remains to be seen. Clearly, anaerobic microorganisms seem to be far more promising candidates for degrading highly chlorinated dibenzo-p-dioxins and dibenzofurans (see Chapter 13), especially in situations where time is not a limiting factor in the cleanup process.

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CHLORINATED PHENOLIC CONTAMINANTS IN SAWMILL ENVIRONMENTS Pentachlorophenol degradation by microencapsulated flavobacteria and their enhanced survival for in situ aquifer bioremediation. In: Hinchee RE, Anderson DB, Metting FB & Sayles GD (Eds) Applied Biotechnology for Site Remediation (pp 422427). Lewis Publishers, Boca Raton, Florida 122. Suzuki T(l 983) Methylation and hydroxylation of pentachlorophenol by Mycobacterium sp. isolated from soil. J. Pesticide Sci. 8:419-428 123. Takada S, Nakamura M, Matsueda T, Kondo R & Sakai K (1996) Degradation of polychlorinated dibenzo-p-dioxins and polychlorinated dibenzofurans by the white rot fungus Phanerochaete sordida YK-624. Appl. Environ. Microbiol. 62:4323-4328 124. Takeuchi M, Hamana K & Hiraishi A (2001) Proposal of the genus Sphingomonas sensu stricto and three new genera, Sphingobium, Novosphingobium and Sphingopyxis, on the basis of phylogenetic and chemotaxonomic analyses. Int. J. Syst. Evol. Microbiol. 51:1405-1417 125. Thomas JM & Ward CH (1989) In situ restoration of organic contaminants in the subsurface. Environ. Sci. Technol. 23:760-766 126. Tiirola MA, Männistö MK, Puhakka JA, Kulomaa MS (2002) Isolation and characterization of Novosphingobium sp. strain MT1, a dominant polychlorophenol-degrading strain in a groundwater bioremediation system. Appl. Environ. Microbiol. 68: 173-180 127. Toussaint M, van der Steen JMD, Slot PC, de Wolf J, Beurskens JEM & Parsons JR (1995) Reductive dechlorination of chlorinated dibenzo-p-dioxins by a bacterial consortium isolated from Lake Ketelmeer sediment (pp 423-424). Proceedings of The Fifth International FZK/TNO Conference on Contaminated Soil, Maastricht, the Netherlands, 30 October-3 November 1995. Kluwer Academic Publishers, The Netherlands 128. Travis CC & Hattemer-Frey HA (1991) Human exposure to dioxin. Sci. Total Environ. 104:97127 129. Valo R & Salkinoja-Salonen M (1986) Bioreclamation of chlorophenol-contaminated soil by composting. Appl. Microbiol. Biotechnol. 25:68-75 130. Valo R, Kitunen V, Salkinoja-Salonen M & Räisänen S (1984) Chlorinated phenols as contaminants of soil and water in the vicinity of two Finnish sawmills. Chemosphere 13:835844 131. van Cauwenverghe L & Roote DS (1998) In situ bioremediation. Technology overview report TO-98-01. Groundwater remediation

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technologies analysis center. 132. van der Meer JR, de Vos WM, Harayama S & Zehnder AJB (1992) Molecular mechanisms of genetic adaptation to xenobiotic compounds. Microbiol. Rev. 56:677-694 133. van der Meer JR, Werlen C, Nishino SF & Spain JC (1998) Evolution of a pathway for chlorobenzene metabolism leads to natural attenuation in contaminated groundwater. Appl. Environ. Microbiol. 64:4185-4193 134. Vartiainen T, Lampi P, Tuomisto JT & Tuomisto J (1995a) Polychlorodibenzo-p-dioxin and polychlorodibenzofuran concentrations in human fat samples in a village after pollution of drinking water with chlorophenols. Chemosphere 30:1429-1438 135. Verstraete W & Devliegher W (1996) Formation of non-bioavailable organic residues in soil: Perspectives for site remediation. Biodegradation 7:471-485 136. Viitasaari S & Mikkola M (1995) Kloorifenolien saastuttamien saha-alueiden kunnostuksen nykytila. Vesi- ja ympäristöhallituksen monistesarja, nro 636. (In Finnish). 137. Viraraghavan T & Tanjore S (1994) Removal of pentachlorophenol from wastewater using peat. Hazardous Waste & Hazardous Materials. 11:423-433 138. Warith MA, Fernandes L & La Forge F (1993) Adsorption of pentachlorophenol on organic soil. Hazardous Waste & Hazardous Materials 10:13-25 139. Weber WJ & Corseuil HX (1994) Inoculation of contaminated subsurface soils with enriched indigenous microbes to enhance bioremediation rates. Wat. Res. 28:1407-1414 140. Webster T & Commoner B (1994) Overview. The Dioxin Debate. In: Schecter A (Ed) Dioxins and health, (pp 1-50). Plenum Press, New York 141. Wilkinson JG (1981) Industrial Timber Preservation. Associated Business Press, London 142. Wilson JT, Armstrong JM & Rafai HS (1994) A full-scale field demonstration of the use of hydrogen peroxide for in situ bioremediation of an aviation gasoline-contaminated aquifer. In: Flathman PE, Jerger DE & Exner JH (Eds) Bioremediation field experience (pp 333-359). Lewis Publishers, Boca Raton, Florida 143. Wittich R-M (1996) Biodegradability of xenobiotic organic compounds depends on their chemical structure and efficiently controlled, and productive biochemical reaction mechanisms. In: Peijnenburg WJGM & Damborský J (Eds) Biodegradability Prediction, pp 7-16. Kluwer Academic Publishers, The

442 Netherlands 144. Wittich R-M, Wilkes H, Sinnwell V, Francke W & Fortnagel P (1992) Metabolism of dibenzo-pdioxin by Sphingomonas sp. strain RW1. Appl. Environ. Microbiol. 58:1005-1010 145. Wittmann C, Zeng A-P & Deckwer W-D (1998) Physiological characterization and cultivation strategies of the pentachlorophenol-degrading bacteria Sphingomonas chlorophenolica RA2 and Mycobacterium chlorophenolicum PCP-1. J. Industr. Microbiol. Biotechnol. 21:315-321

LAINE ET AL. 146. Woodward & Clyde Consultants & Champion International (1995) Annual Operations Report for the Upper Aquifer. Submitted to USEPA Region VIII, Helena, Montana Office. February 28, 1995 147. Yu J & Ward O (1996) Investigation of the biodegradation of pentachlorophenol by the predominant bacterial strains in a mixed culture. Int. Biodeterioration & Biodegradation 37:181-187

Chapter 17 POLYCHLORINATED BIPHENYLS IN AQUATIC SEDIMENTS: ENVIRONMENTAL FATE AND OUTLOOK FOR BIOLOGICAL TREATMENT DONNA L. BEDARD Department of Biology, Rensselaer Polytechnic Institute, Troy, NY, USA

1. INTRODUCTION Polychlorinated biphenyls, PCBs, were widely used from 1929 through 1978 for multiple industrial applications: hydraulic fluid, dielectric fluid in capacitors and transformers, solvent extenders, and more. Approximately 1.4 billion pounds of PCBs were manufactured, and several hundred million pounds were released to the environment (59), most of which are now in aquatic sediments (78). PCBs are hydrophobic, hence they adsorb strongly to organic matter, especially fatty tissue. As a consequence, they accumulate in biota and biomagnify in the food chain. Because PCBs are potential human carcinogens and are potentially toxic to both humans and wildlife, cleanup of PCB-contaminated sediments is a national priority in the United States and has been a hotly debated issue at many sites. Dredging is the solution most often proposed by regulators to remediate PCB-contaminated sediments, but dredging simply moves the PCB-contaminated sediment which still requires treatment or disposal. Usually the contaminated dredgings are deposited in a specially constructed landfill approved for PCBs. Finding a site where such a landfill will be accepted by residents and landowners can be a major problem because of the perceived risks associated with transport of the contaminated sediments to the landfill and seepage of PCBs from the landfill. Most likely no dechlorination or degradation of PCBs will occur in landfills (32). This means that dredged landfilled PCBs may create new problems in the future if the landfills should fail. There are also situations where inaccessibility, or the sheer volume of contaminated sediment may preclude dredging. And, of course, the impact of dredging on the ecosystem is a major consideration. Given all of the above, there is a real need for the development of cost-effective and environmentally compatible methods of PCB bioremediation in situ. The purpose of this chapter is to give a brief overview of PCB bioremediation, and to point out challenges that need to be addressed. (For more detailed dechlorination and degradation, to highlight promising leads that are Dehalogenation: Microbial Processes and Environmental Applications, pages 443-465 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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relevant to reviews of microbial PCB dechlorination and degradation the reader is referred to references 7, 42, 98, and 101).

2. BENEFITS OF PCB DECHLORINATION Over the past 15 years it has become clear that microbially mediated PCB dechlorination in aquatic sediments is widespread, and that this process has significant implications for both risk assessment and remediation strategies. Dechlorination reduces the “dioxin-like” toxicity of PCB mixtures, because it preferentially targets and removes meta and para chlorines (7, 85, 87). Figure 17.1 illustrates the structure, numbering scheme, and naming of PCBs. Hence dechlorination converts “dioxin-like” PCB congeners (i.e., those with two para and at least two meta chlorines, but no more than one ortho chlorine, ref. 89) into less chlorinated congeners that have no “dioxin-like” toxicity. Microbial dechlorination also reduces PCB bioaccumulation in vertebrates by converting PCB congeners that have long half-lives in vertebrate tissue into congeners with much shorter half-lives (6, 24). If carried far enough, microbial dechlorination transforms highly chlorinated PCB congeners to mono- through trichlorobiphenyl congeners that can be oxidized by aerobic biphenyl-degrading bacteria which are typically found in soils and sediments (12, 42, 98). These bacteria use dioxygenases to hydroxylate PCBs, thus initiating a four step “upper pathway” which ultimately breaks open one of the rings (2, 50) to generate chlorobenzoic acid and a 5-carbon fragment that is further metabolized to acetyl CoA and enters the tricarboxylic acid cycle. Chlorobenzoic acids can also be mineralized by some bacteria. Thus, sequential anaerobic/aerobic microbial processes have the potential to completely destroy PCBs. Recent experimental evidence indicates that it may be possible to promote or accelerate PCB dechlorination in situ (see Section 7.1). Based on the available evidence, microbial dechlorination may become an attractive future option for remedial action at some sites.

3. PCB DECHLORINATION IN THE ENVIRONMENT AND PCB ANALYSIS PCB dechlorination in aquatic sediments has been reported worldwide, in both freshwater and estuarine sediments (for review, see references 7 and 101). Table 17.1

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lists many locations that show strong evidence of microbial dechlorination of PCBs in situ, or that have been shown to contain PCB-dechlorinating microorganisms. Quensen and Tiedje (82, 83) give an excellent description of how to determine whether in situ PCB dechlorination has occurred at any given site, and how to determine if PCBdechlorinating microorganisms are present. At the heart of these analyses is an accurate congener-specific analysis of PCBs. This is a difficult analysis because there are 209 different PCB congeners. Commercial PCB mixtures such as Aroclors typically contain

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60-80 PCB congeners. (Aroclor is the tradename for commercial PCB mixtures manufactured in the United States. For more information about commercial PCB mixtures, see reference 7.) Furthermore, microbial dechlorination can generate additional congeners that are not present in significant amounts in the commercial mixtures (6,43,46), Recently, several major advances have made accurate PCB analysis easier. The first is that all 209 PCB congeners are now commercially available; both individually from AccuStandard (New Haven, CT) and Ultra Scientific (North Kingstown, RI), and in a set of 11 mixtures from AccuStandard. The second is the publication by Frame (44) of the relative retention times of all 209 congeners on 17 different capillary GC column phases. Use of the congener mixtures and relative retention time tables published by Frame greatly facilitates accurate congener assignments on almost any GC column. The third is the publication of the quantitative congener distribution for all of the major Aroclor mixtures (45, 46). Frame and his colleagues insist that their tables of congener distribution are only semi-quantitative, but these tables still provide the best quantitation available.

4. THE DIVERSITY OF MICROBIAL PCB DECHLORINATION ACTIVITIES To date, our progress in understanding microbial PCB dechlorination has been limited, because no pure cultures of PCB-dechlorinating microorganisms are yet available. However, even in the absence of pure cultures, we have been able to distinguish eight different PCB-dechlorinating activities on the basis of congener selectivity, targeted chlorines, and dechlorination products. These activities, also called dechlorination processes, each refer to a distinct set of dechlorination reactions which exhibit a particular pattern of congener selectivity and chlorophenyl reactivity. Hence, each dechlorination process can be described by a set or series of reactions that indicates which congeners are substrates, which chlorines will be removed from those congeners, and the order in which they will be removed. The eight dechlorination processes described to date have been given letter names and are briefly summarized in Table 17.2. For more complete descriptions the reader is referred to additional references (7, 15,107). Various investigators have proposed that each of these distinct PCB dechlorination activities is expressed by a different microbial population. The most commonly observed dechlorination is the removal of a meta or para chlorine flanked by two other chlorines, e.g., the meta chlorine from a 234-, 2345-, or the para chlorine from a 2345-chlorophenyl group. The next most common is the removal of singly flanked para chlorines (Dechlorination Processes H, H', and P), or meta chlorines (Dechlorination Process N). Unfortunately, the flanked meta and para dechlorination activities do not complement each other, but instead compete for many of the same substrates. Processes M, Q, and LP each have the rare ability to remove unflanked chlorines. This enables these dechlorination activities to target the terminal products of Dechlorination Processes H, H', N, and P. For example, Processes LP and Q can remove the unflanked para chlorines from the terminal products of Process N dechlorination, and Process M can remove the unflanked meta chlorines from the terminal products of Process P or H Dechlorination (Figure 17.2). However, dechlorination Processes M, Q, and LP appear to have impaired activity against PCBs with six or more chlorines.

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The implications are that two or more dechlorination processes with complementary activities will achieve more extensive dechlorination than any single dechlorination activity can accomplish by itself. Fortunately, complementary activities do occur in at

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least some sites. The clearest example is the upper Hudson River. The extensive meta and para dechlorination of PCBs that has occurred there is likely the result of the combined activities of Dechlorination Processes H, M, and Q (7, 26, 40, 84). Another example is the St. Lawrence River (31). All of the major microbial PCB dechlorination processes that have been described to date target meta or para chlorines, but ortho dechlorination of a few PCB congeners has been observed in the laboratory. Microorganisms from the Woods Pond section of the Housatonic River (Lenox, MA), from Silver Lake (Pittsfield, MA), and from Baltimore Harbor can all remove ortho chlorines from a few PCB congeners, e.g., 246-CB, 2356-CB, 235-CB (17, 34, 99, 102, 106). However, most of these congeners are at best minor constituents of Aroclors (43, 46). There is neither evidence of ortho dechlorination of the Aroclor 1260 in Housatonic River sediments, nor can such an activity be stimulated in these sediments (107; Van Dort and Bedard, unpublished data). A small amount of ortho dechlorination has been observed when Aroclor 1260 was incubated with a microbial enrichment culture that exclusively removed ortho chlorines from several PCB congeners (109). However, even with this enrichment, the primary dechlorination activity against Aroclor 1260 was meta dechlorination (Process N), and only a few congeners were dechlorinated from the ortho position.

5. THE NATURE OF PCB DEHALOGENATORS PCBs have only been manufactured since 1929. How is it then that the ability to dechlorinate PCBs is so widespread, and that there are so many different microbial dechlorination activities? Because of the specificity of the PCB dechlorination activities that have been described, the general assumption is that PCB dechlorination is enzymatic. But how did this capacity evolve? Seventy years is a very short time in the evolutionary scale. Most likely, PCB-dehalogenating enzymes evolved for some other purpose, perhaps for dehalogenation of naturally produced halogenated organics. More than 2000 different naturally-occurring halogenated compounds have been described (51, see Chapter 1). These include many halogenated phenols, phenolic ethers, pyrroles, indoles, heterocycles, and aromatics. In fact, several research groups have concluded that forest and brush fires are the major source of chlorinated dioxins and dibenzofurans in the environment (51). Many of the naturally occurring halogenated compounds are toxic and others have strong antibiotic activity. Perhaps dehalogenating enzymes developed by fortuitious mutations of pre-existing enzymes, that enabled microorganisms to dehalogenate one or more of these natural halogenated organics and in turn gave them a selective advantage in certain environments. Over time, the ability to halorespire, i.e., to use halogenated compounds as electron acceptors, evolved. More recently, exposure to high concentrations of PCBs in contaminated sediments for several decades may have led to further modification and improvement of PCB-dehalogenating enzymes and electron transfer mechanisms. The reader is also referred to Chapters 5 and 8 for further discussion on the evolution of enzymatic mechanisms for dechlorination. If, as hypothesized, PCBs act as electron acceptors, it should be possible to selectively enrich for PCB-dechlorinating microorganisms by incubating sediment with a high concentration of halogenated biphenyls. When sediment from the Woods Pond section of the Housatonic River was incubated with a high concentration of

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2,6-dibromobiphenyl, the dibromobiphenyl was completely dehalogenated to biphenyl (13). At the same time, the number of PCB-dechlorinating microorganisms in the sediment increased up to a 1000-fold from cells per g sediment (108). This finding provides strong support for the hypothesis that PCBdechlorinating organisms obtain energy for growth from dehalogenation, presumably by using halogenated aromatics as electron acceptors. As of this writing, repeated attempts by several laboratories to isolate PCB dechlorinators have not yet succeeded. It is widely assumed that this is at least in part because PCB dechlorination in sediments requires a consortium of several mutually dependent microorganisms. The results of PCB-dechlorinating enrichments treated in various ways to promote or inhibit the growth of particular groups of prokaryotes suggest that the microorganisms responsible for PCB Dechlorination Processes H, M, and Q may be methanogens, spore-forming sulfate reducers, and non-spore-forming sulfate reducers, respectively (111-114). Another intriguing possibility is that a novel group of organisms, Dehalococcoides, may be capable of dechlorinating PCBs. At the moment, only two examples of this genus have been isolated; both grow exclusively by halorespiration. Dehalococcoides ethenogenes dehalogenates tetrachloroethylene and trichloroethylene (PCE and TCE, respectively) (68,69), but the second, strain CBDB1, removes flanked and doubly flanked chlorines from tri- and tetrachlorobenzenes (1). It is not difficult to imagine that this same organism may also dechlorinate PCBs, and experiments to test this are in progress (L. Adrian, personal communication). In the meantime, several stable, highly enriched sediment-free consortia of PCB-dechlorinating microorganisms have been developed (34, 110). This means that the isolation of PCBdechlorinating bacteria may soon become a reality. The ability to monitor changes in microbial populations during enrichment of PCB dechlorinating microorganisms, by characterizing the changes in the distribution of their 16S rRNA gene sequences using denaturing gradient gel electrophoresis (DGGE; 74), or similar techniques, should facilitate our understanding of the microorganisms involved in PCB dechlorination and their isolation. Several laboratories, including ours, are now using this approach.

6. CRITICAL FACTORS AFFECTING THE RATE AND EXTENT OF PCB DECHLORINATION Multiple factors influence PCB dechlorination activity at any given site. The potential for in situ dechlorination is determined by which PCB-dechlorinating microorganisms are present, and by their substrate range and specificity. The PCB congener distribution is also a major factor. The abundant pentachlorobiphenyls of Aroclors 1248 and 1254 are susceptible to dechlorination by most microbial dechlorination activities (Table 17.2). However, the substantial proportions of heptathrough nonachlorobiphenyls in Aroclors 1260 and 1268 make these PCB mixtures more difficult to dechlorinate. This is probably a result of 1) steric hindrance to the dehalogenases caused by increased chlorine substitution and 2) greater insolubility of the PCBs with seven or more chlorines. Conversely, the PCBs in Aroclors 1242 and 1016 are primarily trichlorobiphenyls and have fewer flanked chlorines. Hence, dechlorination processes that can target only flanked chlorines will have minimal impact

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on these Aroclors. Finally, as the meta and para chlorines are depleted, the PCB mixture will approach an undechlorinatable residue, composed of PCB congeners that cannot be dechlorinated by any of the known dechlorination processes. These include 2-CB, 26-CB, 2-2-CB, 26-2-CB, and 26-26-CB. Even if PCB dechlorinating microorganisms are present and have the proper specificity to dechlorinate the PCB mixture associated with a particular sediment, their impact on the PCBs may still be very limited or not even detectable. The PCB concentration will have an effect on the potential rate of dechlorination, as will temperature, pH, and the availability of electron donors and electron acceptors. (For detailed review of these topics, see references 7 and 101). Furthermore, the PCB dechlorinators at a given site may not be able to effectively compete with the other indigenous microorganisms for electron donors. Alternatively, if PCB dechlorinators can use alternate electron acceptors that are more favorable (e.g., sulfate, nitrate, iron), they may not dechlorinate the PCBs until the more favorable electron acceptors are exhausted. Co-contaminants may also affect PCB dechlorination. For example, PCBs will partition into organics and oil, hence these will affect bioavailability (115). It is also possible that heavy metals or pesticides may be toxic to some PCB dechlorinators (115). Quensen and Tiedje (82, 83) give methods for determining whether PCB dechlorinating microorganisms are present at any given site, and whether the PCBs in the sediment are accessible to dechlorination. They also give examples of the results of applying these methods to the Sheboygan River (Wisconsin), the River Raisin (Michigan) and the Saginaw River (Michigan). For example, comparison of the PCB congener distribution profile in the sediment of the River Raisin in Michigan with that ofthe probable original source PCBs, a mixture ofAroclors 1242 and 1248, shows very modest decreases of the more highly chlorinated congeners and corresponding increases in less chlorinated congeners (83). The clearest indications of dechlorination are elevated levels of 24-3-CB and 25-3-CB. These particular products are likely the products of flanked para dechlorination of 24-34-CB and 25-34-CB, respectively. Laboratory assays showed that dechlorinating microorganisms were present, and that the PCBs in the sediment were bioavailable and could be dechlorinated by the indigenous microorganisms under optimal conditions (83). Furthermore, the rate of dechlorination could be accelerated by “priming” with a halogenated substrate (see below). Hence this site could be a candidate for bioremediation. In contrast, laboratory assays showed that sediment from the Saginaw River does not harbor PCB-dechlorinating microorganisms and does not support PCB dechlorination, even when inoculated with PCBdechlorinating microorganisms from another site. This site, therefore, is not likely to be a good candidate for bioremediation.

7. TREATMENT STRATEGIES: PROBLEMS AND POTENTIAL SOLUTIONS 7.1. Activating or Accelerating Anaerobic Dechlorination Two approaches have been used to try to activate or accelerate PCB dechlorination: biostimulation (adding some compound to stimulate the indigenous organisms) and bioaugmentation (adding organisms). For the latter approach, the organisms may be enriched from the same site or may be from a different source.

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7.1.1. Biostimulation Sometimes, even though PCB-dechlorinating microorganisms are present at a site (as evidenced by dechlorination assays conducted in the laboratory), the site shows little or no evidence of in situ dechlorination activity. It may be possible to stimulate the activity of the indigenous PCB-dechlorinating microorganisms in some sediments. Information from research studies has provided several insights on how this might be done. Early studies focused on carbon amendments or addition of electron acceptors to promote the growth of specific types of prokaryotes, but these failed to impact the overall extent of dechlorination, as measured in chlorine loss (7). More recently, two very different approaches have shown considerable promise. The first, “priming”, uses a halogenated aromatic substrate to stimulate PCB dechlorination by the indigenous microbial community. This process was pioneered in our laboratory (8, 14, 100), based on the hypothesis that high concentrations of an appropriate dehalogenation substrate will selectively promote the growth of dehalogenating microorganisms that use the compound as an electron acceptor. The enhanced population of dehalogenators will then dechlorinate the PCBs in the sediment. Our priming studies used sediment from the Housatonic River (Lenox, MA), which is contaminated with Aroclor 1260 (predominantly hexa- through nonachlorobiphenyls). The PCBs in this sediment show clear evidence of past dechlorination (6) and laboratory experiments confirm that PCBdechlorinating organisms are still present (8, 15, 100, 107), yet no dechlorination could be detected in unamended sediment incubated anaerobically for more than a year (11, 107). However, a variety of halogenated aromatics have been identified that can specifically stimulate Process N or Process P dechlorination in this sediment (13, 36, 100). Certain brominated biphenyl congeners have proven to be highly effective substrates for stimulating Process N in Housatonic River sediments. For example, 2,6dibromobiphenyl (26-BB) primes rapid dechlorination of hexa- through nonachlorobiphenyls, converting them to mainly ortho, para-substituted tetrachlorobiphenyls (13). Furthermore, 26-BB will prime PCB dechlorination in this sediment at temperatures as low as 8°C (108). From 1992 to 1995 a field test was conducted in the Woods Pond stretch of the Housatonic River (Lenox, MA) to determine if priming PCB dechlorination in situ was possible. The experiment was carried out at ambient temperatures in 6-foot diameter caissons driven into the sediment. The results were very promising. A single addition of 26-BB activated the indigenous PCB-dechlorinating bacteria and stimulated a 62% decrease in sediment-associated PCBs with six or more chlorines in just three months, and a 74% decrease in a year (11). The dechlorination activity was Process N, and the products were primarily ortho-, and para-substituted tetrachlorobiphenyls, especially 24-24-CB and 24-26-CB. This result clearly demonstrates that it is indeed possible to activate large-scale PCB dechlorination in situ. A second dechlorination activity native to the Housatonic River, Process LP, targets the products of Process N dechlorination and converts them to di- and trichlorobiphenyls that are aerobically degradable (92, 93). In the laboratory, highly dechlorinated PCBs resulting from Process N dechlorination were used to foster, i.e., “prime” LP dechlorination. The results were very promising: most of the products of Process N dechlorination were further dechlorinated to di- and tri-chlorobiphenyls. In fact, 42% of

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the total PCBs were converted to 2-2-CB, 26-CB, 25-2-CB, and 26-2-CB (92, Bedard & Smullen, unpublished data). Based on the results of these studies, even highly chlorinated PCBs can be biologically treated. The major challenge now is to identify environmentally compatible and cost-effective priming substrates. A second promising approach to biostimulation involves the addition of ferrous sulfate, an inexpensive innocuous compound, to promote nearly complete meta and para dechlorination ofAroclor 1242 (114). A series ofexperiments have provided persuasive evidence that the microorganisms responsible for dechlorination processes M and Q in Hudson River sediment are sulfate-reducing bacteria (114). These dechlorination processes are not active in the presence of sulfate, but are enhanced after the sulfate has been exhausted, provided that ferrous iron or another divalent metal is present to precipitate sulfide and thus reduce its toxicity. Apparently, the sulfate stimulates the growth of the bacteria responsible for Dechlorination Processes M and Q, and these bacteria use the PCBs as an alternate electron acceptor when the sulfate is exhausted. The experiments on which these findings were based were carried out with sediment spiked with Aroclor 1242, and incubated in anaerobic mineral medium. If future experiments carried out in actual contaminated river sediment under simulated field conditions show similar results, this finding could be a major breakthrough for PCB remediation. As envisioned, treatment would require only 10 mM, or approximately 10.6 pounds of ferrous sulfate per ton of sediment (114).

7.1.2. Bioaugmentation Another potential alternative to bioremediation is augmentation with PCBdechlorinating organisms enriched from the same site, or from another site. If there are no PCB dechlorinators present at a given site, it might be possible to introduce PCB dechlorinators enriched from another site. Of course this would only be possible if the microbial ecology and biogeochemistry of the site to be treated were suitable (similar temperature, pH, electron acceptors and donors, organic components, mineral composition, and sediment composition) and if co-contaminants do not preclude dechlorination. There have been several laboratory attempts to stimulate PCB dechlorination in sediments from the Housatonic River by inoculating with PCB-dechlorinating enrichments from the same site. Unfortunately these have had little or no impact (15, 104). Bioaugmentation with a stable anaerobic consortium with PCB-dechlorinating ability derived from other sources is another possible approach. Natarajan and colleagues developed such a consortium in the form of methanogenic granules (77). The granules are predominantly composed of syntrophic, hydrolytic, acetogenic, fermentative, and methanogenic bacteria, and have the ability to remove ortho, meta, and para chlorines from 23456-pentachlorobiphenyl (76, 77). These granules were added to PCB-contaminated (Aroclors 1242 and 1248) sediment from the River Raisin, in bench-scale experiments conducted in an anaerobic upflow reactor fed 0.01 % poplar wood to sustain the granules (76). Unamended sediment showed very modest but detectable dechlorination. In contrast, the sediment treated with the dechlorinating consortium (granules) showed strong decreases of tri- through pentachlorobiphenyls, and the accumulation of primarily ortho-substituted mono- and di-chlorobiphenyls.

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Moreover, dechlorination occurred at 12 °C, as well as at 22 to 24 °C. This approach offers considerable promise for remediation, but only if it can be adapted to work with a much smaller inoculum of the bacterial consortium. In the bench-scale experiments, the volume of granules applied was 10% of the volume of the sediment. This raises concerns about the practical and economic feasibility of such an approach.

7.2. Aerobic Biodegradation of PCBs

7.2.1. Evidence of PCB Degradation In Situ Anaerobic PCB dechlorination detoxifies PCBs and decreases their persistence in vertebrates (24), but it does not destroy the PCBs – it converts them to predominantly ortho-substituted, less chlorinated congeners. Therefore, a total solution for PCB bioremediation is usually envisioned as a two stage process: PCB dechlorination, an anaerobic process, followed by the aerobic degradation via the biphenyl (bph) degradation pathway. This pathway (Figure 17.3) converts biphenyl to benzoic acid and 2-hydroxy-penta-2,4-dienoic acid in four steps. The latter compound is further metabolized to acetyl-CoA and then enters the tricarboxylic acid (TCA) cycle (56, 57, 61). The biphenyl degradation pathway is very common in soil and sediment microorganisms (for review, see references 42 and 98). Biphenyl degraders have even been isolated from termite guts, suggesting that the bph pathway is involved in lignin degradation (33). The bph pathway in many biphenyl-degrading bacteria can metabolize PCBs, albeit the PCB substrate specificity in different bacteria varies considerably (12). It was originally proposed that PCBs would be metabolized to chlorobenzoic acid (2), but even as early as 1979 it was recognized that hydroxylated metabolites and the metacleavage product accumulated from some PCB congeners (50). Several lines of evidence indicate that aerobic degradation of the dechlorinated PCBs in the upper Hudson River can and does occur in situ. First, fourteen different strains of PCB-degrading bacteria were isolated from the upper Hudson River in the region below Hudson Falls (73; all strains whose designation begins with an H described in reference 12). A fifteenth strain, Ralstonia eutropha H850 (formerly Alcaligenes eutrophus), was

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isolated from river dredge spoils in the same area (16). Second, a field test conducted in the upper Hudson River in 1991 clearly demonstrated that the mono-, di-, and trichlorobiphenyls in the sediment, the products of PCB dechlorination in situ, were degraded by indigenous bacteria (53). Chlorobenzoic acids briefly accumulated and then were further degraded. Nutrient supplements (ammonia-N, phosphate, biphenyl, and hydrogen peroxide as a source of oxygen) accelerated the degradation, but simply stirring the sediment also helped, presumably because of increased aeration. However, bioaugmentation with Ralstonia eutropha H850 had no effect. This was surprising since H850 has broad PCB congener specificity. Moreover, because it was isolated from river dredge spoils near the site of the field test (9, 16), it was expected that this organism would thrive when reintroduced to the sediment and would increase the efficiency of PCB degradation (7). Third, the isolation of PCB-degrading bacteria from the Hudson River, and the demonstration that PCB degradation occurred during the Hudson River Field Test (53), prove that microorganisms indigenous to the Hudson River are able to degrade the dechlorinated PCBs in the river. However, these observations do not prove that PCB degradation actually occurs in situ. Nevertheless, three additional observations do provide convincing evidence that PCB degradation occurs in situ. 1) Flanagan and May (41), using a sensitive GC/MS technique, identified the expected chlorobenzoic acid products and several hydroxylated metabolites of dechlorinated PCBs in undisturbed sediment cores taken from the region of PCB contamination/dechlorination in the upper Hudson River. 2) Fish and Principe (40) demonstrated anaerobic PCB dechlorination followed by aerobic degradation in submerged test-tube microcosms of Hudson River sediment. PCB dechlorination occurred in the anaerobic zone and in the surface sediment. This was followed by aerobic degradation of the dechlorinated PCBs in the surface sediments (0-4 mm). No nutrients were provided and the only source of oxygen was from contact of the overlying circulating water in the tank where the microcosms were submerged. 3) Williams and May (103) demonstrated that the indigenous microorganisms in the aerobic zone of Hudson River sediment (0-15 mm) could oxidize the dechlorinated PCBs associated with the sediment when statically incubated at 2 to 5 °C. PCB degradation was detectable at six weeks and was extensive by five months. Furthermore, the degradation showed the same congener specificity as the PCB-degrading bacteria isolated from the Hudson River. Together, these observations indicate that the aerobic degradation of dechlorinated PCBs can and does occur in the upper Hudson River, perhaps even during the winter months. The problem of PCB degradation in the environment is not solved, however. Some of the products of PCB dechlorination, especially 26-CB, 26-2-CB, 26-3-CB, and 26-4-CB are recalcitrant, and 26-26-CB completely resists degradation. It is assumed that the two ortho chlorines on the 2,6-chlorophenyl ring sterically hinder the access of the 2,3-biphenyl dioxygenase to both rings. In addition, other congeners, such as 2-4-CB, 24-4-CB, and 25-4-CB are often not metabolized all the way to chlorobenzoic acids and metabolism of chlorobenzoic acids by pathways not optimized for chlorinated aromatics can generate metabolites that arrest chlorobenzoate metabolism (4) or inhibit PCB degradation (21, 55). Some of these issues are being addressed by various genetic engineering approaches (see Section 7.2.2).

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7.2.2. Genetic Engineering to Expand PCB Congener Specificity of PCBDegrading Bacteria The genes encoding the biphenyl/PCB (bph) degradation pathway have been cloned and sequenced for a variety of PCB-degrading bacteria (98). In particular, a great deal has been learned about the sequences controlling the substrate range and position of attack from the study of two PCB-degrading bacteria isolated from opposite sides of the world: Burkholderia cepacia (formerly Pseudomonas sp.) LB400, isolated from a landfill in Moreau, NY (23), and Pseudomonas pseudoalcaligenes KF707, isolated near a biphenyl manufacturing plant in Kitakyushu, Japan (48). Both the genetic organization and the amino acid (aa) sequence of the bph genes in these two organisms are nearly identical (61), yet the PCB congener substrate range of the two organisms is markedly different. For example, LB400 degrades 17 out of 19 PCB congeners in two test mixtures, but KF707 degrades only 7 of these (73). LB400 preferentially attacks 2- or 2,5-chlorophenyl rings, but KF707 preferentially attacks 4-chlorophenyl rings (61). The first enzyme in the bph pathway, biphenyl dioxygenase (BP DOX) is an ironsulfur protein composed of four subunits, each encoded by a separate gene. These are the large subunit (458 aa), the small subunit (213 aa), ferredoxin (109 aa) and ferredoxin reductase (408 aa). The amino acid sequence of the large subunit of the biphenyl dioxygenase differs by less than 5%, or 20 amino acids out of 459, in LB400 and KF707, and the remaining three subunits are identical in 99.3 to 100% of the amino acid sequence. A series of elegant experiments by Mondello et al. (37, 38, 73) and by Furukawa et al. (47, 48, 61, 97) resulted in cloning and sequencing the bph operon for LB400 and KF707, respectively. Site-specific mutagenesis was then used to generate chimeric biphenyl dioxygenases in order to determine which of the amino acids that differ in the two strains are responsible for the differences in congener specificity. These experiments determined that all of the key differences are in the C-terminal region of the large subunit. Mondello et al. (73) found that changing Asn 376 (LB400) to Thr 376 (KF707) in the LB400 BP DOX resulted in loss of most of the ability to attack di-ortho substituted congeners and 2,5-chlorophenyl rings and in narrower substrate specificity. Furukawa and colleagues (61) showed that the reverse change, i.e., changing Thr 376 to Asn in the KF707 BP DOX, expanded the range of PCB substrates and conferred the ability to attack a 2,5-chlorophenyl ring at carbons 3,4. Mondello’s group (73) also showed that changing four other amino acids in the LB400 BP DOX expands the ability to attack di-para substituted congeners such as 4-4-CB. In particular, changing Phe 336 (LB400) to Ile 336 (KF707), or Asn 338 (LB400) to Thr 338 (KF707), enhanced the ability of the LB400 BP DOX to degrade 4-4-CB. Two groups (29,49, 63) have used gene shuffling between KF707 and LB400 to create variant BP DOX enzymes with increased PCB substrate range. However, more work is needed in this area; none of the variants expanded the PCB substrate range beyond that of the two parental strains combined. An optimal two-stage anaerobic/aerobic biodegradation scheme requires that the BP DOX substrate range include 2,6-substituted congeners: 26-CB, 26-2-CB, 26-3CB, 26-4-CB, 24-26-CB, and 26-26-CB, because these are often significant terminal dechlorination products. This may be an area in which gene shuffling could lead to a break-through, since no organisms that can readily degrade these congeners have been found.

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7.2.3. The Intermediate Steps: From Dihydrodiol to Chlorobenzoic Acid The biphenyl dioxygenase is only the first enzyme in a multi-step pathway, and it has become quite clear that other enzymes in the bph pathway are far less tolerant of chlorines than BP DOX. The result is that many congeners that are attacked by BP DOX are not efficiently metabolized to chlorobenzoic acid, but instead accumulate as chlorodihydrodiols, chlorodihydroxybiphenyls, or chlorinated mata-cleavage products (Figure 17.3; references 5, 30, 50, 66, 90, 91). Many congeners containing 2chlorophenyl rings are efficiently metabolized to chlorobenzoic acids by ortho-directed strains such as Burkholderia cepacia LB400, Ralstonia eutropha H850, and Rhodococcus sp. strain RHA1 (5,66). This is apparently because the BP DOX of these strains attacks at the ortho chlorine, generating an unstable diol intermediate which spontaneously loses the ortho chlorine to form a dihydroxybiphenyl (Figure 17.4; reference 52). The same is apparently true for 2,4-chlorophenyl rings. Fortunately, this means that many of the common dechlorination products of Aroclors 1242 and 1248 (e.g., 2-CB, 2-2-CB, 2-3-CB, 2-4-CB, 24-2-CB, 24-4-CB, and 25-2-CB) are readily degradable to chlorobenzoic acids by these organisms. On the other hand, the attack on a 2,5-chlorophenyl ring occurs at position 3,4 and results in a dead-end dihydrodiol (Figure 17.4; references 52, 75). Attack on a 4-chlorophenyl ring occurs at carbons 2 and 3, but often leads to the accumulation of a yellow meta-cleavage product which can inhibit growth and metabolism (5, 66, 91). Therefore, engineering the BP DOX alone does not ensure effective metabolism of all PCBs to chlorobenzoic acids. It will likely be necessary to engineer other enzymes in the upper bph pathway as well.

7.2.4. Metabolism of Chlorobenzoic Acids PCB-degrading bacteria do not generally have the ability to efficiently metabolize chlorobenzoic acids. As a result, chlorobenzoic acids accumulate, and can impair growth and activity of the PCB degraders. Many natural populations can metabolize chlorobenzoic acids to chlorocatechols, but few have the specialized enzymes needed to mineralize the chlorocatechols. Bartels et al. (4) showed that if 3-chlorobenzoic acid is metabolized to 3-chlorocatechol, meta-cleavage will produce a reactive acyl chloride which will irreversibly inactivate the meta-cleavage enzyme. Havel and Reineke (55) reported an unidentified toxic metabolite of 4-chlorobenzoic acid produced by natural microflora that resulted in rapid death of their PCB-degrading strain when it was inoculated into non-sterile soil. More recently, Blasco et al. (21) showed that metabolism of 4-chlorobenzoic acid can lead to the production of protoanemonin, a potent antibiotic which is toxic to many microorganisms including Burkholderia cepacia LB400. Many investigators have sought to combine the bph pathway and chlorobenzoic acid (CBA) degradation pathways into a single microorganism, in order to circumvent these problems and develop bacteria that will grow on PCBs which are chlorinated on both rings. However, in contrast to PCB-degrading bacteria which have a broad substrate range, CBA-degrading bacteria can usually degrade only one isomer of CBA. Most 4CBA-degrading bacteria use a hydrolytic dehalogenase to convert 4-CBA to 4-hydroxybenzoic acid, which is then readily mineralized (96). On the other hand,

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degradation of 2-CBA proceeds by 1,2-dioxygenase attack at the ortho chlorine resulting in oxygenolytic dehalogenation similar to that shown for PCBs in Figure 17.4 (96). Degradation of 3-CBA and the various dichlorobenzoic acids each require other specialized enzymes. Consequently, engineering a microorganism that can completely degrade the desired range of PCBs plus their CBA products is a daunting task, requiring the assembly and coordinated function of multiple pathways. Several PCB/CBA-degrading stains have been constructed by moving bph genes into CBA-degrading strains (58, 90). More recently, investigators have moved CBAdegradation pathways into PCB-degrading microorganisms. The first of these efforts

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developed two recombinant strains of Comomonas testosteroni VP44 that grow on monochlorobiphenyls without accumulation of CBA; one grows on 2-CB, and the other on 4-CB (58). However, neither of these recombinants could grow on PCBs that are chlorinated on both rings, because the BP DOX in the parent strain cannot attack a chlorinated ring. A second effort in the same laboratory (90) developed a recombinant by inserting the operon encoding the 4-CBA hydrolytic dehalogenase pathway fcb into a PCB-degrading strain with broad PCB congener specificity, Rhodococcus sp. strain RHA1 (91). This strain is robust and has the ability to degrade several other aromatic compounds such as benzene, toluene, ethylbenzene, and o-xylene (54), which may be co-contaminants at some sites. The inserted operon functioned and was stable in the recombinant and enabled growth on 4-CB without accumulation of 4-CBA. The recombinant also degraded 2-4-CB, a significant PCB dechlorination product, without accumulation of CBA. However, the strain could only grow on 4-CB if the bph pathway was not fully induced. When the bph pathway was fully induced; the 4-CB was apparently processed to 4-CBA more rapidly than the fcb pathway could break it down. This resulted in a backup of products and accumulation of the yellow meta-cleavage product which in turn inhibited growth and PCB degradation (90, 91). Unfortunately, if the bph pathway is not fully induced, the recombinant will not degrade the full spectrum of PCBs that its parent strain degrades. More engineering will be necessary to better coordinate the bph and fcb pathways in this recombinant, and then to incorporate additional CBA degradation pathways.

8. FUTURE DIRECTIONS AND NEEDS The anaerobic microbial dechlorination of PCBs, perhaps combined with aerobic degradation, holds much promise for the in situ detoxication and/or remediation of contaminated aquatic sediments. In order to realize this potential, much work is still needed. The key challenge for improved PCB dechlorination is to develop economical and environmentally compatible methods for activating the appropriate PCBdechlorinating organisms (either indigenous or introduced) in situ. Identification, isolation, and characterization of the microorganisms responsible for the various PCB dechlorination processes is crucial. However, it will not be sufficient to understand these organisms in axenic cultures in defined media. The very fact that these organisms have been so difficult to isolate testifies to their close interdependence on other organisms, and it is crucial that we understand these interactions. It is also critical to understand how PCB-dechlorinating microorganisms interact with their environment. For example, why is evidence for LP dechlorination in situ restricted to a very short stretch of the Housatonic River (10, Bedard et al., unpublished data)? Moreover, it is intriguing that the microorganisms responsible for LP dechlorination are present about 7 miles downstream in Woods Pond, as demonstrated by their activity in laboratory experiments (15,92), but the PCB congener distribution in multiple samples from Woods Pond show no evidence of LP dechlorination in situ at this location (10, Bedard and Smullen, unpublished data). Understanding why they are active in one site and not another, i.e., understanding their ecology, will lead to insights on how to promote PCB dechlorination in situ. Aerobic degradation of terminal dechlorination products, primarily ortho-substituted

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PCBs, will be required to completely eliminate PCBs. A great deal is already known about PCB degradation in many different bacteria, yet significant challenges remain. Perhaps the most obvious is the problem of supplying a steady supply of oxygen in sufficient quantity to maintain full function of the dioxygenases in the bph pathway. This will be much more difficult in some sites than others. It will be particularly difficult in highly methanogenic sediments with high organic content and low water flow, such as the Woods Pond stretch of the Housatonic River. In addition to the challenges discussed in the preceding sections, there is a particular need to discover or engineer a biphenyl dioxygenase that will not be impaired by chlorines in positions 2,6 on the opposite ring. This would enable degradation of 26-CB, 26-2-CB, 26-3-CB, and 26-4-CB, all of which are significant terminal dechlorination products and all of which are recalcitrant. There is also a need to discover or engineer enzymes with broader specificity for chlorinated substrates for the remaining steps in the upper bph pathway, specifically dihydrodiol dehydrogenase, 2,3-dihydroxybiphenyl dioxygenase, and 2-hydroxy-6-oxo-6-phenylhexa-2,4-dienoic acid hydrolase (HOPDA hydrolase). This last enzyme in particular is a bottleneck for many congeners. Incorporating CBA-degradation pathways into PCB-degrading organisms will also help, but their regulation must be coordinated with that of the upper bph pathway. Excellent pathways are already available for 2-CBA and 4-CBA. Additional CBA pathways that will be most helpful are those for 24-CBA, 25-CBA, and 26-CBA. Clearly there are many challenges, but none that are insurmountable. Acknowledgments

The author acknowledges grant support from National Science Foundation Award 0077837 and from GE Corporate Environmental Programs. REFERENCES 1. Adrian L, Szewzyk U, Wecke J & Görisch H 2. 3.

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database. Fresenius J. Anal. Chem. 357:701-713 Rhee G-Y (2001) Microbial PCB dechlorination in dredged sediments and the effect of moisture. 45. Frame GM (1997) A collaborative study of 209 PCB congeners and 6 Aroclors on 20 different Chemosphere 43:1119-1126 HRGC columns: 2. Semi-quantitive Aroclor 33. Chung S-Y, Maeda M, Song E, Horikoshi K & congener distributions. Fresenius J. Anal. Chem. Kudo T (1994) A Gram-positive polychlorinated 357:714-722 biphenyl-degrading bacterium, Rhodococcus erythropolis strain TA421, isolated from a 46. Frame GM, Cochran JW & Bøwadt SS (1996) Complete PCB congener distributions for 17 termite ecosystem. Biosci. Biotechnol. Biochem. Aroclor mixtures determined by 3 HRGC 58:2111-2113 systems optimized for comprehensive, 34. Cutter L, Sowers KR & May HD (1998) quantitative, congener-specific analysis. J. High Microbial dechlorination of 2,3,5,6Resolut. Chromatogr. 19:657-668 tetrachlorobiphenyl under anaerobic conditions in the absence of soil or sediment. Appl. 47. Furukawa K, Hayase N, Kazunari T & Tomizuka N (1989) Molecular relationship of Environ. Microbiol. 4:2966-2969 chromosomal genes encoding biphenyl/ 35. David M, Brondyk LM & Sonzogni WC (1994) polychlorinated biphenyl catabolism: Some soil Distribution of PCB congeners in Sheboygan bacteria possess a highly conserved bph operon. River (Wisconsin) Sediments. J. Great Lakes J. Bacteriol. 171:5467-5472 Res. 20:510-522 36. DeWeerd KA & Bedard DL (1999) Use of 48. Furukawa K & Miyazaki T (1986) Cloning of gene cluster encoding biphenyl and halogenated benzoates and other halogenated chlorobiphenyl degradation in Pseudomonas aromatic compounds to stimulate the microbial dechlorination of PCBs. Environ. Sci. Technol. pseudoalcaligenes. J. Bacteriol. 166:392-398 33:2057-2063 49. Furukawa K, Nishi A, Watanabe T, Suyama A & 37. Erickson BD & Mondello FJ (1992) Nucleotide Kimura N (1998) Engineering microorganisms capable of efficient degradation of chlorinated sequencing and transcriptional mapping of the environmental pollutants. Rev. Toxicol. 2:179genes encoding biphenyl dioxygenase, a multicomponent polychorinated-biphenyl187 degrading enzyme in Pseudomonas strain 50. Furukawa K, Tomizuka N & Kamibayashi A LB400. J. Bacteriol. 174:2903-2912 (1979) Effect of chlorine substitution on the 38. Erickson BD & Mondello FJ (1993) Enhanced bacterial metabolism of various polychlorinated biphenyls. Appl. Environ. Microbiol. 38:301biodegradation of polychlorinated biphenyls after site-directed mutagenesis of a biphenyl 310 dioxygenase gene. Appl. Environ. Microbiol. 51. Gribble GW (1994) The natural production of 59:3858-3862 chlorinated compounds. Environ. Sci. Technol. 39. Farley KJ, Germann GG & Elzerman AW (1994) 28:310A-319A Differnetial weathering of PCB congeners in 52. Haddock JD, Horton JR & Gibson (1995) Lake Hartwell, South Carolina. In: Baker LA Dihydroxylation and dechlorination of (Ed) Environmental Chemistry of Lakes and chlorinated biphenyls by purified biphenyl 2,3Reservoirs, (pp 575-600). American Chemical dioxygenase from Pseudomonas sp. strain Society, Washington, D.C. LB400. J. Bacteriol. 177:20-26 40. Fish KM & Principe JM (1994) 53. Harkness MR, McDermott JB, Abramowicz DA, Biotransformations of Aroclor 1242 in Hudson Salvo JJ, Flanagan WP, Stephens ML, Mondello River test tube microcosms. Appl. Environ. FJ, May RJ, Lobos JH, Carroll KM, Brennan Microbiol. 60:4289-4296 MJ, Bracco AA, Fish KM, Warner GL, Wilson 41. Flanagan WP & May RJ (1993) Metabolite PR, Dietrich DK, Lin DT, Morgan CB & Gately detection as evidence for naturally occurring WL (1993) In situ stimulation of aerobic PCB aerobic PCB biodegradation in Hudson River biodegradation in Hudson River sediments. sediments. Environ. Sci. Technol. 27:2207-2212 Science 259:503-507 42. Focht DD (1993) Microbial degradation of 54. Hatta T, Shimada T, Yoshihara T, Yamada A, chlorinated biphenyls. In Bollag J-M & Stotzky Masai E, Fukuda M & Kiyohara H (1998) metaG (Eds): Soil Biochemistry Vol 8. New York: Fission product hydrolases from a strong PCB Marcel Dekker, Inc., pp 341-407 degrader Rhodococcus sp. RHA1. J. Ferment. 43. Frame G (1997) Congener-specific PCB Bioeng. 85:174-179 analysis. Analyt. Chem. 69:468A-478A 55. Havel J & Reineke W (1992) Degradation of 44. Frame GM (1997) A collaborative study of 209 Aroclor 1221 and survival of strains in soil PCB congeners and 6 Aroclors on 20 different microcosms. Appl. Microbiol. Biotechnol. HRGC columns: 1. Retention and coelution 38:129-134

PCBs IN AQUATIC SEDIMENTS 56. Hofer B, Backhaus S & Timmis KN (1994) The biphenyl/polychlorinated biphenyl-degradation locus (bph) of Pseudomonas sp. LB400 encodes four additional metabolic enzymes. Gene 144:916 57. Hofer B, Eltis D, Dowling DN & Timmis KN (1993) Genetic analysis of a Pseudomonas locus encoding a pathway for biphenyl/ polychlorinated biphenyl degradation. Gene 130:47-55 58. Hrywna Y, Tsoi TV, Maltseva OV, Quensen JF III & Tiedje JM (1999) Construction and characterization of two recombinant bacteria that grow on ortho- and para-substituted chlorobiphenyls. Appl. Environ. Microbiol. 65:2163-2169 59. Hutzinger O & Veerkamp W (1981) Xenobiotic chemicals with pollution potential. In Leisinger T, Hutter R, Cook AM & Nuesch J (Eds): Microbial Degradation of Xenobiotics and Recalcitrant Compounds. New York: Academic Press, p 3-45. 60. Imamoglu I, Li K, Christensen ER & McMullin J (2000) Transformations of PCBs in sediments of Fox River, Wisconsin. Proceedings of the 43rd Conference on Great Lakes Research (IAGLR 2000), May 22-26, 2000, Cornwall, Ontario 61. Kimura N, Nishi A, Goto M & Furukawa K (1997) Functional analyses of a variety of chimeric dioxygenases constructed from two biphenyl dioxygenases that are similar structurally but different functionally. J. Bacteriol. 179:3936-3943 62. Kuipers B, Cullen WR & Mohn WW (1999) Reductive dechlorination of nonachlorobiphenyls and selected octachlorobiphenyls by microbial enrichment cultures. Environ. Sci. Technol. 33:3579-3585 63. Kumamaru T, Suenaga H, Mitsuoka M, Watanabe M & Furukawa K (1998) Enhanced degradation of polychlorinated biphenyls by directed evolution of biphenyl dioxygenase. Nature Biotechnol. 16:663-666 64. Lake JL, Pruell RJ & Osterman FA (1991) Dechlorination of polychlorinated biphenyls in sediments of New Bedford Harbor. In: Baker RA (Ed) Organic Substances and Sediments in Water, (pp 173-197). Lewis Publishers Inc., Chelsea, MI 65. Lake JL, Pruell RJ & Osterman FA (1992) An examination of dechlorination processes and pathways in New Bedford Harbor sediments. Marine Environ. Res. 33:31-47 66. Maltseva OV, Tsoi TV, Quensen JF III, Fukuda M & Tiedje JM (1999) Degradation of anaerobic reductive dechlorination products of Aroclor

463 1242 by four aerobic bacteria. Biodegradation 10:363-371 67. Maruya KA, Lee RF, Frischer MF, Kostka JE, King JK & Saunders FM (1999) Biotransformation of co-occurring PCBs and mercury by sulfate reducing bacteria in estuarine sediments. SETAC 20th Annual Meeting, November 14-18. Philadelphia, PA 68. Maymó-Gatell X, Anguish T & Zinder SH (1999) Reductive dechlorination of chlorinated ethenes and 1,2-dichloroethane by ”Dehalococcoides ethenogenes” 195. Appl. Environ. Microbiol. 65:3108-3113 69. Maymó-Gatell X, Nijenhuis I & Zinder SH (2001) Reductive dechlorination of cis-1,2dichloroethene and vinyl chloride by “Dehalococcoides ethenogenes." Environ. Sci. Technol. 35:516-521 70. McLaughlin D (1994) Natural and Induced Transformations of PCBs in sediments. Ph.D. Thesis. Department of Land Resources, University of Wisconsin-Madison 71. McNulty AK (1997) In Situ Anaerobic Dechlorination of Polychlorinated Biphenyls in Hudson River Sediments. MS Thesis. Department of Earth and Environmental Sciences, Rensselaer Polytechnic Institute, Troy, NY 72. Mohn WW & Tiedje JM (1992) Microbial reductive dehalogenation. Microbiol. Rev. 56:482-507 73. Mondello FJ, Turcich MP, Lobos JH & Erickson BD (1997) Identification and modification of biphenyl dioxygenase sequences that determine the specifity of polychlorinated biphenyl degradation. Appl. Environ. Microbiol. 63:30963103 74. Muyzer G & Smalla K (1998) MiniReview: Application of denaturing gradient gel electrophoresis (DGGE) and temperature gradient gel electrophoresis (TGGE) in microbial ecology. Antonie van Leeuwenhoek 73:127-141 75. Nadim L, Schocken MJ, Higson FJ, Gibson, DT, Bedard DL, Bopp LH & Mondello FJ (1987) Bacterial oxidation of polychlorinated biphenyls, In: Proceedings of the 13th Annual Research Symposium on Land Disposal, Remedial Action, Incineration, and Treatment of Hazardous Waste. EPA/600/9-87/015, (pp 395-402). US Environmental Protection Agency, Cincinnati 76. Natarajan MR, Nye L, Wu WM, Wang H & Jain MK. (1997) Reductive dechlorination of PCBcontaminated Raisin River sediments by anaerobic microbial granules. Biotech. Bioeng. 55:182-190 77. Natarajan MR, Wu WM, Nye L, Wang H,

464 Bhatangar L & Jain MK (1996) Dechlorination of polychlorinated biphenyl congeners by an anaerobic microbial consortium. Appl. Microbiol. Biotechnol. 46:673-677 78. NRC (National Research Council) (1979) Polychlorinated Biphenyls. National Academy of Sciences, Washington, D.C. 79. Øfjord GD, Puhakka JA & Ferguson JF (1994) Reductive dechlorination of Aroclor 1254 by marine sediment cultures. Environ. Sci. Technol. 28:2286-2294 80. Pakdeesusuk, U, Lee, CM, Coates, JT & Freedman, DC (2001) Reductive dechlorination of polychlorinated biphenyls in Twelve Mile Creek/Lake Hartwell watershed. American Chemical Society, Division of Environmental Chemistry, Preprints of Extended Abstracts, 221st ACS National Meeting, San Diego, CA, April 1-5, 2001,Vol. 41, No. 1, Paper #246, pp 334-337 81. Palekar L, Wiegel J, Maruya K & Kostka J (2000) Dechlorination patterns of PCBs by an anaerobic consortium of estuarine sediment microorganisms. Abstracts, 100th General Meeting, American Society for Microbiology. Q-148, p 574 82. Quensen JF III & Tiedje JM (1997) Methods for evaluation of PCB dechlorination in sediments. In: Sheehan D (Ed) Methods in Biotechnology Vol. 2: Bioremediation Protocols, (pp 241 -256). Humana Press Inc., New Jersey 83. Quensen JF III & Tiedje JM (1997) Evaluation of PCB dechlorination in sediments. In: Sheehan D (Ed) Methods in Biotechnology Vol. 2: Bioremediation Protocols, (pp 257-273). Humana Press Inc., New Jersey 84. Quensen JF III, Boyd SA & Tiedje JM (1990) Dechlorination of four commercial polychlorinated biphenyl mixtures (Aroclors)by anaerobic microorganisms from sediments. Appl. Environ. Microbiol. 56:2360-2369 85. Quensen JF III, Mousa MA, Boyd SA, Sanderson JT, Froese KL & Giesy JP (1998) Reduction of receptor mediated activity of PCB mixtures due to anaerobic microbial dechlorination. Environ. Toxicol. Chem. 17:806-813 86. Quensen JF III, Tiedje JM & Boyd SA (1988) Reductive dechlorination of polychlorinated biphenyls by anaerobic microorganisms from sediments. Science 242:752-754 87. Quensen JF III, Tiedje JM, Boyd SA, Enke C, Lopshire R, Giesy JP, Mora M, Crawford R & Tillit D (1992) Evaluation of the suitability of reductive dechlorination for the bioremediation of PCB-contaminated soils and sediments. In: I n t e r n a t i o n a l Symposium on Soil

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465 107. Wu Q, Bedard DL & Wiegel J (1997) Temperature determines the pattern of anaerobic microbial dechlorination of Aroclor 1260 primed by 2,3,4,6-tetrachlorobiphenyl in Woods Pond sediment. Appl. Environ. Microbiol. 63:4818-4825 108. Wu Q, Bedard DL & Wiegel J (1999) 2,6Dibromobiphenyl primes extensive dechlorination of Aroclor 1260 in contaminated sediment at 8 to 30°C by stimulating the growth of anaerobic PCB-dehalogenating microorganisms. Environ. Sci. Technol. 33:595602 109. Wu Q, Sowers KR& May HD (1998) Microbial reductive dechlorination of Aroclor 1260 in anaerobic slurries of estuarine sediments. Appl. Environ. Microbiol. 64:1052-1058 110. Wu Q, Sowers KR & May HD (2000) Establishment of a polychlorinated biphenyldechlorinating microbial consortium, specific for doubly flanked chlorines, in a defined sediment-free medium. Appl. Environ. Microbiol. 66:49-53 111. Ye D, Quensen JF III, Tiedje JM & Boyd SA (1992) Anaerobic dechlorination of polychlorinated biphenyls (Aroclor 1242) by pasteurized and ethanol-treated microorganisms from sediments. Appl. Environ. Microbiol. 58:1110-111 112. Ye D, Quensen JF III, Tiedje JM & Boyd SA (1995) Evidence for para dechlorination of polychlorobiphenyls by methanogenic bacteria. Appl. Environ. Microbiol. 61:2166-2171 113. Ye D, Quensen JF III, Tiedje JM & Boyd SA (1999) 2-Bromoethanesulfonate, sulfate, molybdate, and ethanesulfonate inhibit anaerobic dechlorination of polychlorinated biphenyls by pasteurized microorganisms. Appl. Environ. Microbiol. 65:327-329 114.ZwiernikMJ, Quensen JF III & Boyd SA(1998) amendments stimulate extensive anaerobic PCB dechlorination. Environ. Sci. Technol. 32:3360-3365 115. Zwiernik MJ, Quensen JF III & Boyd SA (1999) Residual petroleum in sediments reduces the bioavailability and rate of reductive dechlorination of Aroclor 1242. Environ. Sci. Technol. 33:3574-3578

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PART V. SUMMARY

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Chapter 18 ENVIRONMENTAL DEHALOGENATION PROBLEMS AND RECOMMENDATIONS CHARLES E. CASTRO Chemical & Environmental Consulting, Laguna Beach, CA, USA

1. INTRODUCTION The wide array of microbial activities associated with halogen cycling that are presented in earlier chapters gives the reader a good idea of the current state of the field of microbial dehalogenation research. Are there additional areas that should be explored, and what needs to be done to put these endeavors in context with current and developing environmental concerns? This chapter attempts to address these questions. The chapter begins with six recommendations. These are followed by a more cohesive presentation of the role of haloorganics in the environment, their influence upon human health, and the influence their continued study may have upon understanding the underlying chemistry and dynamic nature of the environment. (1) Probing the reactivity of the environment is essential. “Site Reactivity Probes” (SRPs) now exist that can determine the reactivity of any terrestrial site by direct analysis using nuclear magnetic resonance (NMR) techniques. Oxidative, reductive, and substitutive capacities can be measured in-situ and their rates assessed. Rates measured by a reactivity probe can be taken as a maximum for that site to degrade or transform a target contaminant or xenobiotic. The probes should be generally employed in evaluating any site. (2)

The oceans must be explored for their dehalogenation potential.

The oceans are the great repositories of runoff waters from the earth surface, and are now known to be the source of a vast array of haloorganics (for a summary, see Chapter 1). The synthesis of these compounds is primarily associated with kelp and algae. It appears that chloro- and bromoperoxidases, iron porphyrin-containing enzymes, and some enzymes with vanadium centers are involved. However, the study of the enzymes present in the life forms that dwell in the oceans is in its infancy. A reasonable question to ask is: If the ocean is a vast resource for the synthesis of haloorganics, might it not

Dehalogenation: Microbial Processes and Environmental Applications, pages 469-479 Edited by M.M. Häggblom and I.D. Bossert, Kluwer Academic Publishers, 2003.

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also be a main source of destruction of these compounds? Is there a reasonable cycle of synthesis and destruction of haloorganics in the ocean itself? It would seem this must be so. For example, low valent iron and vanadium in the same enzymes may serve as powerful reductants.

(3) The dehalogenation ability of plants merits much study. Only a few examples of biodehalogenation by plants can be found in the literature, though “phytoremediation” studies for the cleanup of halogenated organic contaminants are in progress. For example, it is clear from our early work (unpublished), that orange seedlings can degrade alkyl halides. A much-increased effort in this area of research is necessary. For a summary of fungal dechlorinations, see Chapter 6.

(4)

Studies of abiotic processes important to halogen cycling, including the nature and mechanisms of these transformations need to be examined.

The chemical reactivities of geological minerals, humic acids, and other earth constituents are largely uncharted, making this field ripe for investigation. Minerals containing iron, copper, nickel, manganese, and cobalt should be prime targets for study. These elements in various valence states and complexes have well established dehalogenating potential. See Chapter 9 for an analysis.

(5) Studies with organic halides can be used as models for predicting the rate and fate of other environmental transformations. Properly chosen organic halides release halide ion regardless of the nature of the transformation. Hence, good rate data can be obtained. SRPs are an example of this general principal, but little is known about the degree to which substances differ in their in situ rates of transformation at a given site. Structure-activity relationships need to be designed based upon the reality that the inherent reactivity of the site controls the rates of conversion and the fate of substances present. Polarity, size, and shape are main structural characteristics for any substance.

(6) More realistic assessments of the hazards associated with the trace quantities of organic halides detectable in the environment need to be made. After all, haloorganics are natural products and have been present on earth for millennia. All life forms have developed in their presence. Moreover, this treatise demonstrates, with great force, that microbes have an enormous capacity to dehalogenate haloorganics and render them non-toxic. Mammals, plants, and nearly all life forms have this capacity. As improved sampling and analytical technologies continue to decrease levels of detection, much attention has been given to the trace levels of haloorganic contaminants. Some of the current regulations need rethinking. A methyl bromide standard for toxicity is proposed.

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2 . THE INFLUENCE OF DEHALOGENATION STUDIES UPON AN UNDERSTANDING OF THE ENVIRONMENT AND HUMAN HEALTH 2.1. The Reactivity of the Environment Although it may seem obvious, it is not widely understood that the reactivity of the environment itself is the governing factor that determines the rate of conversion and the fate of haloorganics and all other substances on earth. Moreover, the reactivity of any segment of the terrestrial environment may vary widely with time, season, weather, and any other change it may undergone. Thus, one might expect the microbiological composition and population at any ground site to be in continual flux. A good example of the variability that may be observed is evidenced by studies of hexadecane, naphthalene, and phenanthrene metabolism at different sites and times along the coast following the Exxon Valdez oil spill (31). A vast array of environmental dehalogenations by microbes has been reported to date. For leading references see Chapters 2 through 8, and 11 through 17, and reference (3). In contrast there is a paucity of examples for corresponding abiotic processes (21, 22), although this potential bares much further scrutiny (Chapter 9). At this time, it would seem that the dominant mode of transformation of haloorganics in the terrestrial environment is biotic in nature, and this capacity for transformation resides in the microorganisms, principally bacteria, that inhabit the earth. Plants, however, may also be very important in dehalogenation processes, but an understanding of their role is only beginning to emerge (25, 26). With any given site or environmental segment then, it is clear that the transformations wrought by it must be a function of the combined chemical, microbiological, and plant species present. The nature of this complex matrix is almost impossible to ascertain, but its composite reactivity can be measured experimentally (6). This measurement is essential for predicting the rate and fate of haloorganics and other xenobiotics. Given that abiotic dehalogenation in the terrestrial environment is rare or slow compared to microbial processes, it is reasonable to assume that biodehalogenation processes will dominate. However, rates of dehalogenation for whole bacterial cells, responsible enzymes, and corresponding active site mechanistic chemistry establish that the rate limiting step in the process is never the chemistry at the active site. For an amplification and salient data, see reference (3). For example, the rates of dehalogenation of polyhalomethanes by Methylosinus trichosporium OB-3b vary only mildly and do not follow the bond dissociation energies of the corresponding C-H bonds (1). In this case, the dehalogenation chemically follows the insertion of an oxygen atom into the C-H bond by the enzyme methane monooxygenase, and a large isotope effect has been observed for C-H insertion by this enzyme (28). Substitution of deuterium for hydrogen in the polyhalomethanes, however, had no effect upon the rates of oxidative dehalogenation of these substrates by resting cells. Another startling example is vested in the unusually high reactivity of nickel(I)isobacteriochlorins to alkyl halides as compared to the very slow reductions by the anaerobe Methanobacter thermoautotrophicum (MTM) that contains the nickel corphin cofactor Nickel(I)octaethylisobacteriochlorin is a good model for cofactor It reduces a wide array of alkyl halides with astonishing speed. Indeed, it is the

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fastest nucleophile on record. The containing organism, M. thermoautotrophicum, possesses the same broad capacity for dehalogenation as this active site model, but the reactions are exceedingly slow and do not vary with substrate structure. Thus, while the products and overall chemistry of the two systems are the same (5), the speeds of reaction differ by a factor of Moreover, the vast difference in rates with substrate structure noted with the isobacteriochlorin in homogeneous solution is not observed with the organism. It is ironic that this organism with the greatest capacity for rapid reduction, is the slowest we have observed. Clearly, the chemistry at the active site within the organism is not rate-limiting. The mild or lack of variation of the organismal rates with structure here, in a reductive process, is similar to that observed in the oxidative process noted above with the soil methylotroph (M. trichosporium OB-3b), but the rates are slower. The data suggest that the microbial rates are dependent upon the organism and not the mechanism or the structure of the substrate. For a summary of anaerobic reduction reactions by bacteria, see Chapter 5. More elaborate studies (8) of the nature and rates of organohalide reduction by Pseudomonas putida (strain Ppg-786), the responsible enzyme, cytochrome in its native cell and in homogeneous solution, are documented, including the corresponding chemistry of the active site of the iron porphyrin in homogeneous amide solvent. Both the active site heme and the enzyme reduce the halides in an overall second-order process that is first-order with respect to the heme component or enzyme, and first-order with respect to organic halide substrate. The heme in homogeneous solution generally reduces halides much more quickly than the enzyme because of a steric effect. The larger structures have difficulty penetrating the enzyme protein to get to the iron moiety. In addition, the enzyme in its native cell reacts more slowly than the enzyme inhomogeneous solution. Direct spectral observation of the oxidation of the heme by haloorganics within whole cell suspensions established that the rate law for the dehalogenation is independent of the enzyme. This represents a direct experimental verification that the rate of dehalogenation by whole cells does not reflect the chemistry at the active site within the cell. That is, the rate-limiting step in microbial dehalogenation is not the chemistry that occurs directly at the active site. Thus, overall biotransformation rates cannot be predicted by factors or relationships drawn solely from chemical reactivity and thermodynamics. Indeed, it may be more accurately portrayed as a composite of permeation factors for the halide. These entail permeation of the cell, getting to the dehalogenating enzyme within the cell, and getting through the tertiary structure of the protein to the active site in the enzyme. This latter steric effect at the enzyme level is exemplified by the inertness of DDT with P. putida. DDT is readily reduced to DDD by heme in homogeneous solution (2, 9, 33). however, does not react with DDT because the substrate cannot penetrate the relatively small pocket in which the heme is ensconced. Thus, although DDT may penetrate P. putida Ppg-786, and the cells have the capacity to dehalogenate it, they do not. This case is also a good example of kinetic rather than thermodynamic control of reactivity, and is typical of microbial dehalogenations. The studies noted above with M. trichosporium OB-3b and M. thermoautotrophicum MTM were conducted with small halides such that any steric inhibition of rates at the enzyme or cofactor level would be small. Hence, nearly the same rates (within a factor of 4) are obtained for all substrates with the same organism. Taken together, the

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results with these three organisms and a variety of substrates suggest that all relatively small organic halides, to a first approximation, will be converted at the same rate with a given organism. Extrapolating this thinking to the “environment” means that a small molecule of medium polarity, such that it can permeate lipid, protein and water and is capable of oxidation, reduction or substitution (hydrolysis), can indicate the transformation capacity of a particular site, if products from it are characteristic of the process, and are easily discerned and measured. In a sense, in addition to the facets of microbial permeation, any substrate in the environment must find a reactive site that can transform it to be degraded. That is, it must get to the chemically reactive site(s) whether they are within microorganisms, plant roots, or represent natural chemical constituents of a particular soil or site. Thus, rates for oxidation, reduction, or substitution at any site assessed by a “Site Reactivity Probe” as having the properties noted above can be taken to represent the maximum rate of conversion by that site at that time for any xenobiotic whether it contains halogen or not. For such an assessment, three probes have been designed so that minimum sample handling and work up are required. They are acid (CA), (CAM), (CCN). The details of the methodology for their use and effectiveness as Site Reactivity Probes (SRPs) for soil, lake water and sediments, coastal marine water and sediments, and sludge has demonstrated a large capacity for the environment to transform xenobiotics (3). A simple incubation of the sample with the probe in water, followed by centrifugation/filtration and NMR analysis of products has measured the various reactivities of these sites (6). Unless a transformation by a given site is purely chemical in nature, for example aqueous hydrolysis, structure-activity relationships based upon physical organic chemical principles (18, 24) have no relevance, and even here environmental permeation factors could dominate. The same can be said for QSAR modeling based upon physical properties or hypothetical considerations of the xenobiotic substrate under study (10, 11, 20, 23). These models must be calibrated to the reactivity of the site by experimentallyderived information, and formulated to reflect the dominant controlling factors of the environment in which the transformation takes place. In short, there is no such thing as a specific environmental degradation rate for a specific substance. These reactivities are site-specific. Realizing that it is the environmental site that controls the rate of transformation, and that this reactivity can be measured, should lead to a routine use of SRPs in planning and assessing remediation strategies, pesticide use, or the influence of any change upon a the reactivity of a site. Because of steric constraints, rigid larger molecules will react more slowly than these probes, as will very non-polar or very highly polar charged substances, because of lessened permeation capacity. Attempts to assess their respective transformation rates using the SRPs will require a structure reactivity equation that considers molecular size and shape, and polarity. For example, the rate of transformation of substance X maybe estimated using a relationship of the form:

where is the rate for the reactivity probe to be oxidized (assuming in this case, the site has been demonstrated to oxidize the probe), and is the corresponding rate for

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substance X. An octanol/water partition coefficient closest to 1 is ideal in that it represents a molecule of medium polarity soluble in a broad range of matrices. The may be taken as a function of the logarithm of the for X such that any deviation from a of 0 results in a lessened rate of transformation. The function:

will do this nicely, wherein

is the absolute value of log

The term f (size and shape) can be formulated arbitrarily using the solvent diameter of benzene (5.2 Ångstroms) as a standard for which steric inhibition at an active site is minimal. In this case, the function:

where

is the minimum diameter of the structure X, in Ångstroms.

Thus, overall rates of transformation for unknown substances may be estimated from the experimental rate of the reactivity probe by the equation:

Clearly, this equation is speculative, but it is based upon known parameters that effect environmental dehalogenation processes. The reactivity of any environmental segment must be measured and adjusted for the polarity of X and its molecular size, before any estimates of degradation rates of X are possible. However, in order for this experimental, size and shape, and polarity approach to be effective, much work is in order. At this time, though favorable examples exist (3), there are not enough rate measurements for defined transformations by single organisms under constant conditions to verify that these assumptions are broadly true or not. Thus, additional work with pure cultures under resting cell conditions is essential. Moreover, there are few examples of actual environmental rates at a specific site for a variety of substances. Work of this kind is also needed. In sum, environmental structure/activity relationships are meaningless unless they are calibrated to the reactivity of the environmental site. 2.2 Human Health Haloorganic substances that serve as ‘environmental drugs’ can be greatly beneficial to human health. Indeed, mosquito abatement by DDT , and the consequent control of malaria in the world could rank this haloorganic compound as the environmental equivalent of ‘penicillin’ in saving human lives. However, the presence of DDT and other haloorganics in the environment and their potential deleterious effects upon human health and other organisms is the basis of much concern. Moreover, federal and private support for work in the environmental dehalogenation area is in large measure based upon this and similar concerns. Given the widespread natural generation and presence

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of haloorganics on earth, along with new knowledge of environmental reactivity, environmental dehalogenation, and human metabolism, it is reasonable to ask whether trace quantities of haloorganics in the environment pose any health risk at all. Some particularly striking observations of naturally-occurring haloorganics lend weight to the notion that man and the rest of the earth have been living with these substances for a very long time. For example, an ester of 4- bromoacetoacetate has been isolated recently from the cerebrospinal fluid of normal humans (14). It is thought to have a physiological function. Another example is shown in the tetraiodophenolic diphenylether linkage to alanine to produce thyroxine, a critical regulator of the thyroid gland. Also, the ketone, 1-bromo-3-chloroacetone, an oxidative metabolite of 1,2dibromo-3-chloropropane (DBCP) (19), has been isolated from the red alga, Asparagopsis taxiformis, along with 100 chloro-, bromo-, iodo- and mixed polyhalogenated ketones, alcohols, amides and carboxylic acids (12-13) The alga is a favorite in the diet of Hawaiians. Ethylene dibromide (EDB) is now known to be synthesized by giant kelp of the Antarctic, and it has also been detected as one of the simple alkyl halide constituents emanating from crushed rocks, shales and minerals (12). Freons, trichlorethylene (TCE), and a variety of fluorochloroalkenes and alkanes are now well established constituents of the gases emitted from volcanoes and sulfataras (12). Polyhalo-N-methylbipyrroles, similar in structure to polychlorinated biphenyls (PCBs), have been identified as natural components of sea bird eggs (15). Finally, ancient groundwater samples and soil itself have been shown to contain haloorganics (12). Both chloroform and trichloroacetic acid have been found to be biogenerated in soil (12). Haloorganics are also generated in spruce forest soil (27). The major enzyme thought to be responsible for the syntheses of haloorganics in the environment is haloperoxidase. It is a heme-containing peroxidase (17), and represents a beautiful example of the capacity of nature to use the same reactant, an iron porphyrin or heme, for both the synthesis and destruction of haloorganics. Similarly, hemoglobin, myoglobin, the cytochromes peroxidase and any hemeprotein in the ferrous state that allows access to iron (a protein in the G conformation) can reduce a broad range of haloorganics (3). Moreover, in higher valence states of iron, the and peroxidases have the capacity to oxidize them. The ubiquity of hemes and hemeproteins on earth and in man is no accident. They are responsible for the respiratory cycle on this planet, but they are also intimately involved in the halogen cycle as well. The reader is referred to Chapter 1 for a review of the halogen cycle and to the excellent reviews of Gribble (12-14) for a description of naturally-occurring organic halides and their abundance. Mammals can rapidly detoxify haloorganics, even very toxic ones. A good example is the sequential reduction of the war gas chloropicrin (trichloronitromethane) to nitromethane by mice, using a glutathione dependent pathway. This dechlorination can occur in the liver cytosol and microsomes (30). The same reductive sequence occurs with the soil bacterium P. putida via the enzyme cytochrome and it is quite rapid (7). Given the massive production of haloorganics by natural sources, and an equally great capacity for the transformation of them by natural means, it is the author’s opinion that current regulations and practices relating to these compounds are unnecessarily restrictive. Their toxicity to man and the environment is greatly exaggerated. Moreover, the contribution to the natural haloorganic pool by human activity is small. For example,

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it is possible, with reasonable accuracy to estimate the amount of methyl bromide used and produced by man because production records are kept. However, this substance is produced in enormous quantities by ocean algae and kelp as well as many other natural sources, and it is difficult to estimate the total natural production. Moreover, methyl bromide is readily degraded by soil methylotrophs (1) and marine nitrifiers (29). It can hydrolyze abiotically at reasonable speed, and the chemical hydrolysis rate is enhanced by light (4). However, methyl bromide is a gas at room temperature, and consequently some of the substance, used as a preplant soil fumigant in agriculture (principally with strawberries in California), may escape to the upper atmosphere. On the basis that it will contribute to ozone depletion, methyl bromide is scheduled for elimination from use by the United States Environmental Protection Agency (USEPA). Methyl bromide is a toxic, odorless, gas, and protection of applicators requires appropriate safety mandated standards. In light of the natural production and destruction of this substance, however, its elimination from use based upon ozone depletion makes no sense. The trace quantities of pesticides detected in groundwater have also fostered what oftentimes seem to be illogical and unnecessarily low values for tolerances. Are organic halides or substances at 1 ppb and less truly deleterious to anyone or anything? Consider the Threshold Limit Values (TLV) established for some well-known toxic gases presented in Table 18.1. These values represent concentrations in air that workers may be exposed to day after day without adverse effects. As noted above, methyl bromide, the third listing in the table, is a toxic gas to humans. Indeed, it is toxic to most organisms by virtue of its strong alkylating capacity and excellent permeation characteristics. It can also bind to heme iron (32). Thus, the substance was commonly used as a house fumigant for termites, a sterilant for nuts and grains, as well as a preplant soil sterilant. Clearly, however, humans have a greater tolerance for it than they do for nitrogen dioxide, chlorine, or ozone. Its low molecular weight, established chemistry and reactivity, and its physical characteristics suggest methyl bromide as an alkyl halide

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standard for human toxicity. That is, all haloorganics will be less toxic to humans than methyl bromide is. For example, methyl bromide is a much better alkylating agent than EDB (ethylene dibromide) or DBCP (l,2-dibromo-3-chloropropane). It also possesses much better permeation characteristics. It is soluble in media of widely varying polarity. Hence, it can alkylate nucleic acids in a cell nucleus more quickly and be able to induce cancer via this pathway more readily than EDB or DBCP. Indeed, methyl bromide has a great capacity to interfere with all bioprocesses in which nucleophilic sites may be important. This includes all proteins where alkylation may alter overall conformation and effectiveness. Using the TLV for methyl bromide (mol. wt. 94.94) in air from Table 18.1, a conservative estimation of the tolerable exposure for an eight hour period can be made. For this calculation it is assumed that one breathes in 0.5 L with each breathing cycle, at 12 times per minute. Thus the number of liters breathed in is: 0.5 · (12) · (60) · (8) = 2,880 L for an 8 hour day. Taking the density of air at 300 °K (80 °F) at 1.16 g/L, the number of grams of methyl bromide inhaled at 5000 ppb is:

On the other hand, assume one drinks 5 L of water per day that contains the legal maximum for DBCP (0.2 ppb, taken from the USEPA web site, 2000). The number of moles of DBCP (mol. wt. 236.3) taken in would be:

This would suggest that DBCP is or approximately 40,000 times more toxic than methyl bromide! This is of course not true, and it is preposterous. If the methyl bromide TLV were used for water, that is assuming DBCP were at most as toxic as methyl bromide is, then, using the above values, the level would be set at:

To be sure, there are other modes of intoxication by haloorganics that do not entail alkylation or bond-breaking processes. For example, oxidation at key metal centers, or simply binding intact to specific receptor sites may occur. Indeed, binding at specific sites is a dominant mode of drug action and an important basis for drug design. The tetrachlorodioxins and polychlorobiphenyls may function in this way. However, can this binding be more drastic than a ubiquitous irreversible alkylation by methyl bromide? It is hard to see a coherent reasoning behind current drinking water and other environmental standards set by the government for haloorganics. It is time they were reexamined. These low, and legally enforceable standards, enhance the public hysteria about “chemicals” and “pesticides” in the environment, and they contribute to the large expenditure of funds associated with legal disputes and mitigation procedures.

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3. FUTURE WORK Continued studies of haloorganics in the environment and all life forms can generally open the door to an understanding of the dynamic nature of all structures on earth. The halogen of the carbon-halogen bond contained in organic halides, can function as a reporter group for kinetic analyses of fragmentation, or chemical and biochemical conversion. More accurate rates for these processes are needed. In addition, microbiologists, botanists, and environmental scientists generally need to collaborate more closely with chemists, in order to establish the fundamental nature, scope, and speed of these reactions. Structures of reactants and products must be established for each bond making-bond breaking process. Labeled substrates should be employed in these studies to demonstrate unambiguously the products observed are derived from the substrate. Moreover, the ocean, the life forms within it, and in its sediments need to be explored for naturally-occurring dehalogenation processes. After all, this vast expanse of organic halide generation undoubtedly has great capacity for dehalogenation as well. Plants, minerals, man, and other microorganisms also require much further study. Placing dehalogenation in the proper framework can also cast a more reasonable view of “chemicals” in the environment, and a better understanding of the dynamics of all substances on earth. No single thing abides, but all things flow. Fragment to fragment clings and thus they grow,

Until we know and name them. Then by degrees they change, and are no more the things we know. Lucretius, 50 B. C. (from De Rerum Natura)

REFERENCES 1. Bartnicki EW & Castro CE (1994) Biodehalogenation: Rapid oxidative metabolism of mono- and polyhalomethanes by Methylosinus trichosporium OB-3b. Environ. Toxicol. Chem. 13:241-245 2. Castro CE (1964) The rapid oxidation of iron(II) porphyrins by alkyl halides: A possible mode of intoxification of organisms by alkyl halides. J Am. Chem. Soc. 86:2310 3. Castro CE (1998) Environmental Dehalogenation-Chemistry and Mechanism. In: Ware G (Ed) Reviews of Environmental Contamination and Toxicology 155 (pp 1-171). Springer-Verlag, New York 4. Castro CE & Belser NO (1981) Photohydrolysis ofmethyl bromide and chloropicrin. J Agr. Food Chem. 29:1005-1008 5. Castro CE, Helvenston MC & Belser NO (1994) Biodehalogenation, reductive dehalogenation by Methanobacterium thermoautotrophicum:

6.

7.

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Comparison with nickel(I)octaethylisobacteriochlorin anion. An F-430 model. Environ. Toxicol. Chem. 13:429-433 Castro CE, O'Shea SK, Wang W & Bartnicki EW (1996) 13 C-NMR reactivity probes for the environment. Environ. Sci. Technol. 30:1185-1191 Castro CE, Wade RS & Belser NO (1983) Biodehalogenation: The metabolism of chloropicrin by Pseudomonas sp. J. Agr. Food Chem. 31:1184-1187 Castro CE, Wade RS & Belser NO (1985) Biodehalogenation: Reactions of cytochrome P-450 with polyhalomethanes. Biochemistry 24:204-210 Castro CE & Wade RS (1985) N-Alkenylporphyrins from the reaction of DDT with chloroiron(III) tetraphenylporphine and iron powder. J. Org Chem. 50:5342-5351 Eriksson L, Jonsson J & Tysklind M (1995)

ENVIRONMENTAL DEHALOGENATION - PROBLEMS AND RECOMMENDATIONS Multivariate QSRB modeling of biodehalogenation: Half-lives of halogenated aliphatic compounds. Environ Toxicol. Chem. 14:209-207 11. Eriksson L (1997) Cluster-based design in environmental QSAR. Quant Struct-Act. Relat. 16:383-390 12. Gribble GW (1998) Naturally-occurring organohalogen compounds. Acc. Chem. Res. 31:141-142 13. Gribble GW (1988) Chlorinated compounds in the biosphere. Natural Production. In: Meyers RA (Ed) Encyclopedia of Environmnental Analysis and Remediation, (pp 972-1035). John Wiley and Sons Inc., New York 14. Gribble GW ( 2000) The natural production of organobromine compounds. Environ. Sci. Pollut. Res. 7: 37-49 15. Gribble GW, Blank DH & Jasinski JB (1999) Synthesis and identification of two halogenated bipyrroles present in sea bird eggs. Chem. Comm. 1999:2195-2196 16. Helvenston MC & Castro CE (1992) Nickel(I)octaethylisobacteriochlorin anion an exceptional nucleophile. Reduction and coupling of alkyl halides by anionic and radical processes. A Model for Factor J. Am. Chem. Soc. 114:8490-8496 17. Hewson WD & Hager LP (1979) Peroxidases, catalases and chloroperoxidases. In: Dolphin D (Ed) The Porphyrins Vol. VII, Chapter 6 (pp 295-332). Academic Press, New York 18. Klump G (1982) Reactivity in Organic Chemistry, John Wiley and Sons, New York 19. Koe GS & Vilker VL (1993) Dehalogenation by cytochrome Effect of oxygen level on the decomposition of 1,2-dibromo-3-chloropropane. Biotechnol. Prog. 9:608-614 20. Kompare B (1998) Estimating environmental pollution by xenobiotic chemicals using QSAR (QSRB) models based on artificial intellegence. Water Sci. Technol. 37:9-18 21. Kriegman-King MR & Reinhard M (1992) Transformation of carbon tetrachloride in the presence of sulfide, biotite, and vermiculite: Environ Sci. Technol. 26:2198-2206 22. Kriegmann-King MR & Reinhard M (1994) Transformation of carbon tetrachloride by pyrite

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in aqueous solution. Environ. Sci. Technol. 28:692-700 23. Lohman PH (1999) Qualitative and quantitative procedures for health-risk assessment. Mut. Res. 428:237-254 24. March J (1992) Advanced Organic Chemistry. Reactions, mechanisms, and structure. John Wiley and Sons Inc. New York 25. Newman LA, Strand SE, Choe N, Duffy J, Ekuan G, Ruszaj M, Shurtleff BB, Wilmoth J, Heilman P & Gordon MP (1997) Uptake and biotransformation of trichloroethylene by hybrid poplars. Environ. Sci. Technol. 31:1062-1067 26. Newman LA, Doty SL, Gery KL, Heilman PE, Muiznieks I, Shang TQ, Siemieniec ST, Strand SE, Wang X, Wilson AM & Gordon MP (1998) Phytoremediation of organic contaminants: a review of phytoremediation research at the University of Washington. J. Soil Contam. 7:531-542 27. Öberg GO & Grøn C (1998) Sources of organic halogen in spruce forest soil. Environ Sci. Technol. 32:153-1579 28. Pilkington SJ & Dalton H (1990) Soluble methane monooxygenase from Methylococcus capsulatas Bath. In: Lindstrom (Ed) Methods in Enzymology, Vol 188 (pp 181-190). Academic Press, New York 29. Rasche ML Hyman M & Arp DJ (1990) Biodegradation of halogenated hydrocarbon fumigants by nitrifying bacteria. Appl. Environ. Microbiol. 56:2568-2571 30. Sparks SE, Quistad GB & Casida JE (1997) Chloropicrin: Reaction with biological thiols and metabolism in mice. Chem. Res. Toxicol. 10:1001-1007 31. Sugai SF, Lindstrom JE & Braddock FF (1997) Environmental influences on the microbial degradation of Exxon Valdez Oil in the shorelines of Prince William Sound, Alaska. Environ. Sci. Technol. 31:1564-1572 32. Wade RS & Castro CE (1985) Halomethane adducts of iron(II)porphyrins. Inorg. Chem. 24:2862-2863 33. Wade RS & Castro CE (1973) Oxidation of iron(II) porphyrins by alkyl halides. J. Am. Chem. Soc. 95:226-230

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481

INDEX abiotic dehalogenation, 34, 261-282, 358364, 470, alkyl halides, 261-272 formation of organometallic intermediates, 279-280 kinetics, 272-276, 363-365 pathways, 269, 361 reaction mechanisms, 268, 276-281 Acetobacterium dehalogenans, 20, ecological role, 38 fermentation of chloromethane, 69 growth on methyl chloride, 116-121 Acetobacterium woodii, cometabolic reductive dehalogenation, 122 ecological role, 38 acetogens, ecological role, 38-39 reductive dehalogenation, 69, 124-125 role in VC degradation, 375-376 Acinetobacter sp., growth on chlorobenzoates, 60, 230 aerobes, ecological role, 38-39 aerobic metabolism, 19-20, 54-55, 58-68, 208, biocides, 61-66, 167-170 haloaliphatic compounds, 19, 165, 207 haloaromatic compounds, 58-61 halophenolics, 19,67,71, 177-190,425427 thermodynamics, 106-108 (see also oxygenase-mediated dehalogenation) aged residues, 297-299, 309-310, (see also sorption) Agrobacterium radiobacter, synergistic PCP degradation, 426 alachlor, abiotic dehalogenation, 266 biodegradation by fungi, 168 Alcaligenes denitrificans, growth on 4-CB, 60 Alcaligenes eutrophus, 2,4-D degradation, 62-63 Alcaligenes paradoxus, 2,4-D degradation, 63 Alcaligenes sp., 62, growth on chlorobenzoates, 58, 60

Alcaligenes xylosoxidans, 2,4-D degradation, 63 aldrin, bioavailability, 291 biotransformation, 166, 306 production and use, 14 aliphatic hydrodrocarbons, halogenated, 10-11, 165, aerobic degradation, 209-212 anaerobic dechlorination, 74-76 degradation by fungi, 165 natural production, 6, 54 Aminobacter sp., growth on methyl halides, 339 analog enrichment, enhanced PCDD degradation, 361-363 using PBBs, 109, 449-450, 452-453 anaerobes, ecological role, 38-39 anaerobic metabolism, 20-21, 68-77, 115157, 355, oxidation of DCE, VC, 373-374 respiration, 20, 55, 69, 89, (see also dehalorespiration; electron acceptors) Anaeromyxobacter dehalogenans, as chloridogen, 71 anthropogenic compounds, 3, 9, 14, 33-35, 53, 161,351,443, (see also xenobiotic) antibiotics, chlorinated, 163,457, (see also chloramphenicol; drosophilin A) AOX (adsorbable organic halogen), 8, 192, 194 aquatic sediments, PCB dechlorination, 444-445 PCDD dechlorination, 355 aquifers, cleanup of TCE-contaminated sites, 397402 (see also aquatic sediments; groundwater) Aroclors, 171, 308, 446, 449-451, (see also polychlorinated biphenyls) aromatic compounds, halogenated, aerobic degradation, 19, 227 anaerobic dechlorination, 20, 71, 115 degradation by fungi, 169 natural production, 6, 54 aromatic dehalogenases, 227-256

482

Arthrobacter globiformis, 62, growth on 4-CB, 60 Arthrobacter oxydans, growth on 4-CB, 60 Arthrobacter sp., 2,4-D degradation , 61 dichloropropanol mineralization, 208 growth on chlorobenzoates, 58, 60, 230 atmospheric gases, 323-325, global warming potential (GWP), 323 mixing ratio, 324 ozone depletion potential (ODP), 323 troposphere residence time 324, 332 atrazine, abiotic dehalogenation, 266 bioavailability, 292 biotransformation, 168, 307 dechlorination, 235-242 sorption in soil, 296 atrazine chlorohydrolase, 235-242 Azoarcus tolulyticus, TFA cometabolism, 334 Basidiomycetes, biodegradation, 164-197,430-431 organochlorine production, 161-164 (see also white rot fungi) benzoquinones, chlorinated, biodegradation by fungi, 180,183 reduction during chlorophenol degradation, 186-187 bioaugmentation, 385, 429, 433, 453-454 bioavailability, 40-41,291-299, aging phenomenon in soil, 297-299, 422 definition, 292 desorption-resistant fractions, 294-297, 422 effect on mineralization, 171 environmental determinants, 293-294 impact on toxicity, 294 polymerization (oxidative coupling),423 solubility of substrate, 309 sorption, 294-299, 423 biocides, 10,12, (see pesticides) biodegradability, effect of bioavailability, 291-299 effect of halogen substituent, 40-41, 450 effect of substrate concentration, 37, 41 intrinsic (definition), 292 solubility of substrate, 305-309, 450

INDEX

biodegradation, 14, assessment and optimization, 389-394 constraints, 108-109, 450 coupled anaerobic-aerobic processes, 312,314-315,373-382, 454 environmental determinants, 36-37,4647, 427, 450-451 factors, 34, 36-37 mechanisms, 15-18, 55, 164, 207 biogeochemical cycling of organohalides, 6, 33, 54, 324, 469, dioxin dechlorination, 366-367 redox concept, 89 role of microorganisms, 54 biokinetic constants, 412, (see also modeling) bioremediation, cleanup of contaminated sites, 21, 365366, 397-402, 428-435 co-contaminants, 388 composting, 428-432 electron donors, 390-393 in situ, 46, 282, 291, 366, 433, 451-456 immobilized cells, 427 microcosm studies, 389-402 natural attenuation (intrinsic), 365, 387 nutrient addition, 36-37,394 site-specific evaluation, 385-415 thermodynamic basis, 111 two-stage process (anaerobic/aerobic), 373-386, 454 biostimulation, 385-386, 451-453 biotransformation, 311-313, function of substrate solubility, 305-309 (see also biodegradation) biphenyl, degrading bacteria, 444 bph pathway, 454-458 1,1-bis(p-chlorophenyl)-2,2-dichloroethene, (see DDE) 1,1 -bis(p-chlorophenyl)-2,2,2-trichloroethane, (see DDT) Bjerkandera adusta, PCP biodegradation, 179 bjerkanderol, 162 Bradyrhizobium sp., 2,4-D degradation, 63 brominated aliphatic compounds, abiotic dehalogenation, 272 bromobenzoic acid, 7

INDEX

bromoethane (BA), anaerobic dechlorination, 122 bromoform, abiotic dehalogenation, 261, 266 bromomethanes, 7, 332-341 bromoperoxidase, 469, (see also haloperoxidase) bromopyrroles, 7 bromotrichloromethane, dehalogenation, 164 Burkholderia anthropogonis, 2,4-D degradation, 63 Burkholderia cepacia, 2-CB biodegradation, 55, 57 dechlorination of PCB metabolites, 457 growth on 4-CB, 60 haloaliphatic dehalogenase, 212 Burkholderia mallei, 2,4-D degradation, 63 Burkholderia sp., 2,4-D degradation, 63 growth on chlorobenzoates, 58, 60 carbonate, as electron acceptor, 45 carbon tetrachloride (CT), abiotic dechlorination, 263-264, 270271, 276 anaerobic dechlorination, 122 anaerobic mineralization, 116, 164 dechlorination mechanism, 165 mineralization by Phanerochaete chrysosporium, 164 reaction with lignin peroxidase, 165 removal from the environment, 331 catabolic mobile elements, 61-63 catabolic pathways, 208, evolution, 217-219 CB, (see chlorobenzoate) Ceriporiopsis subvermispora, biodegradation of chlorinated pesticides, 168 biotransformation of chlorophenols, 183 CFCs, (see chlorofluorocarbons; halocarbons) CFC-11 (trichlorofluoromethane), 328-329 CFC-113 (trichlorotrifluoroethane), 334 CFC-12 (dichlorodifluoromethane), 328 chloral, formation during TCE oxidation, 42 chloramphenicol, 7

483

chlordane, bioavailability, 292 biodegradation by fungi, 166 biotransformation, 306 production and use, 14 chloridogenesis, 20, 55, 69-77, (see also dehalorespiration) chloridogens, 70, (see also dehalorespirers) chlorinated anisyl compounds. (see chloroanisoles) chlorinated diphenyl ethers, (see chlorodiphenyl ethers) chlorinated ethenes, (see chloroethenes; see specific compounds) chlorinated humus, 316, formation from pesticide residues, 317 natural synthesis, 318 chlorinated organophosphorus pesticides, biodegradation by fungi, 166-168 chlorinated phenols, (see chlorophenols) chlorinated phenylpropanoid compounds, fungal metabolites, 162 chlorinated solvents, (see solvents; see also tetrachloroethene; trichloroethene) chlorine, (see halogen) chlorine disinfection, 11 chloroacetanilides, biodegradation by fungi, 168-169 chloroacetate, genes encoding degradation, 60 chloroanilines, biodegradation by fungi, 191-192 chloroanisoles, biodegradation by fungi, 179 bioformation, 423 biotransformation, 426 fungal metabolites, 162, 423 chlorobenzenes, biodegradation by fungi, 171-172 genes encoding degradation, 60 reductive dehalogenation, 115 2-chlorobenzoate (2-CB; 2CBA), oxygenolytic dehalogenation, 55 3-chlorobenzoate (3-CB; 3CBA), biodegradation, 16, 20, 68-69, 117 dechlorination by Desulfomonile tiedjei, 20,43,44,56,69, 126 utilization by Pseudomonas sp., 19 4-chlorobenzoate (4-CB; 4CBA), metabolic pathway, 231 utilization by Pseudomonas sp., 56, 230

484

chlorobenzoates, aerobic growth substrate, 58 genes encoding degradation, 60 production of toxic metabolites, 457 reductive dehalogenation, 115 4-chlorobenzoyl CoA dehalogenase, 230-235 chlorobiphenyl, (see polychlorinated biphenyls) 1 -chlorobutane, 61 3-chlorodibenzofuran, bioavailability, 295 chlorodibromomethane, production in soil, 54 chlorodifluoromethane, (see HCFC-22) 2-chloro-1,4-dimethoxybenzene (2DCDMB), redox mediator in fungal metabolism, 163 chlorodiphenyl ethers, 422, biodegradation by fungi, 171 -174 chloroethanes, energetics of reductive dechlorination, 101 production and use, 10 reductive dehalogenation, 115 chloroethene, (see vinyl chloride) chloroethenes, 373-382, cleanup of contaminated sites, 385-415 dechlorination pathway, 269, 386 degradation in groundwater, 376-377 fermentation, 110 mineralization under anaerobic conditions, 116 reductive dehalogenation, 115 reductive/oxidative biodegradation, 373382 (see also specific compounds) chloroform (CF), abiotic dechlorination, 263-264 anaerobic dechlorination, 122 anaerobic mineralization, 116 dechlorination by fungi, 164 production and use, 10, 11, 54 chlorofluorocarbons (CFCs), 325, degradation, 327-332 history and use, 327-328 properties, 326-329 (see also halocarbons; halomethanes) chloroheterocyclic pesticides, biodegradation by fungi, 166-167 (see also specific compounds) chlorohydrodiols,

INDEX

formation, 59 chlorohydroquinones, dehalogenation, 16, 242-249 formation, 20 chlorolignin, 11, biodegradation by fungi, 192-193 (see also lignin) chloromethanes, 335-341, oxygenolytic degradation, 216 production and use, 7, 10, 54 production by fungi, 162 utilization by acetogens, 69, 116 reductive dehalogenation, 115, 125 (see also methyl chloride) chloroorganic compounds, 53, (see also organohalides) chloroperoxidase, 469, (see also haloperoxidase) chlorophenol reductive dehalogenase, 136, 142-143, 188, cpr genes, 148-149 chlorophenols, as electron acceptors, 71-73 bioavailability, 422-423 biodegradation by fungi, 177-190 biotransformation by fungi, 186-187 impurities, 421-422, (see also polychlorinated dibenzo-p-dioxins; polychlorinated dibenzofurans) methylation, 186, 423, 426 natural production, 54, 318 pathways for biodegradation, 188-190 polymerization, 422-423 reductive dehalogenation, 115, 132, 187188 soil contamination, 421-423 chlororespiration, 70, 379, (see also dehalorespiration) chlorophenoxyacetates, biodegradation by fungi, 168-170 (see also 2,4-dichlorophenoxyacetate; 2,4,5-trichlorophenoxyacetate) chloropicrin, (see trichloronitromethane) chlorpyrifos, biodegradation by fungi, 166 Chrysosporium lignorum, dichloroaniline mineralization, 191 dieldrin biodegradation, 166 Citrobacter freundii, cometabolic reductive dehalogenation, 123

INDEX

Clostridium bifermentans, as chloridogen (dehalorespirer), 71 cometabolic reductive dehalogenation, 123, 125 ecological role, 38 PCE dechlorination, 74 Clostridium pasteurianum, utilization of CFCs, 328 Clostridium rectum, cometabolic reductive dehalogenation, 123 Clostridium thermoaceticum, cometabolic reductive dehalogenation, 122 CoA dehalogenases, 230-235, genes, 59 CoA thioesters, role in dehalogenation reactions, 230 cobalamin, (see also vitamin abiotic dehalogenation, 263-265 coenzyme role in dechlorination, 124-125,263265,471 cofactor mediated dehalogenation, 124-125, 263-265, 471, (see also cobalamin; corrinoids; tetrahydrofolate; coenzyme cometabolism, 19, 123-125, 132, 308, aerobic, 159, 195 alkyl reductive dehalogenation, 124-125 composting, 428 congeners, dioxins (PCDDs), 365 PCBs, 444 selectivity in dechlorination, 446-449 conservative tracers, 328 cooxidation, 329-332, (see also cometabolism) corrinoids, reaction mechanism for reductive dehalogenases, 142-145 role in dehalogenation, 117-125, 335, 341 corrosion, role in abiotic dehalogenation, 279 Corynebacterium sepedonicum, 2,4-D degradation, 61 growth on 4-CB, 60 Corynebacterium sp. dichloropropanol mineralization, 208 haloaliphatic dehalogenases, 211-212

485

cosubstrates, role in anaerobic dehalogenation, 42 CT, (see carbon tetrachloride) Cyathus bulleri, biotransformation of lindane, 166 cytochrome role in dechlorination, 124, 473 white rot fungi, 164, 196 cytochromes, role in electron transfer, 139-140 cytochromes c (and c-type), electron transfer in Desulfomonile tiedjei, 139 Cytophaga sp., growth on PCP, 66 2,4-D, (see 2,4-dichlorophenoxyacetate) dalapon, 307 DCE, (see dichloroethene) DDD (dichlorodiphenyldichloroethane, or 1,1 -dichloro-2,2-bis[4-chlorophenyl] ethane), abiotic dechlorination, 266 biodegradation, 314 metabolite of DDT biodegradation, 174 DDE (dichlorodiphenyldichloroethylene, or 1,1 -dichloro-2,2-bis[4-chlorophenyl] ethene, or 1,l-bis[p-chlorophenyl]2,2-dichloroethene), 174, 472, abiotic dechlorination, 266 biodegradation, 314 environmental effects, 313 DDT (dichlorodiphenyltrichloroethane, or 1,1,1 -trichloro-2,2-bis[4-chlorophenyl]ethane), or l,l-bis[pchlorophenyl]-2,2,2-trichloroethane abiotic dechlorination, 266 aerobic degradation, 314 anaerobic dechlorination, 125 bioavailability, 291 biodegradation, 174-177, 314 dechlorination by heme, 472 environmental fate, 13, 313-314 history and use, 12, 14, 303-305 metabolites, 174 pathways, 176, 306, 312 debromination, reductive, 109-110 (see also dehalogenation) dechlorinases, 55, 4-CB dechlorinase, 58

486

dechlorinases (continued), dehydrodechlorinase, 216 dichloromethane dechlorinase, 67 dechlorination, mechanisms, 55 (see also dehalogenation) dechlororespiration, 70, (see chlororespiration; dehalorespiration) degree of chlorination, effect on biodegradability, 35, 94, 171, 291, 308-309, 444, 450 effect on polymerization reactions, 184, 363 energetics of dechlorination, 110, 363, 376-377 impact on environmental fate, 35, 41, 443-445 PCB toxicity, 444 solubility, 450 Dehalobacter restrictus, as chloridogen, 71, 126 ecological role, 38-39 growth yield, 133 PCE dechlorination, 74, 390 PCE/TCE reductive dehalogenase, 137, 142 phylogeny and properties, 129 Dehalobacterium formicoaceticum, 20, anaerobic metabolism of dichloromethane, 69, 116-117, 121 ecological role, 38 Dehalococcoides ethenogenes, cofactor requirements, 394 ecological role, 38-39 electron donors for dehalorespiration, 126, 390 growth yield, 133 PCE dechlorination, 74, 132 PCE reductive dehalogenase, 136 phylogeny and properties, 131 potential role in PCB dechlorination, 450 TCE reductive dehalogenase, 137 Dehalococcoides sp., as chloridogen, 71 effect of pH on dechlorination, 387 dehalogenases, 15, aliphatic dehalogenases, 61, 210-213 amino acids at catalytic sites, 145-148, 456 aromatic dehalogenases, 227-256 atrazine chlorohydrolase, 235-242

INDEX

biochemistry of reductive dehalogenases, 136-137, 141-145 biphenyl dioxygenase, 456-458 catalysis, 214-217, 228, 231-253 catalytic triads, 213 3-CB reductive dehalogenase, 136, 229 characterization and classification, 136137, 213-217, 227-256 4-chlorobenzoyl CoA dehalogenase, 228, 230-235 chlorophenol reductive dehalogenase, 188 4-chlorophenylacetate dioxygenase, 228 2,4-dichlorobenzoyl CoA dehalogenase, 229 evolution, 217-220, 227-256 gene products, 213-215 genetic engineering, 456 genetic organization, 219 genetics, 145-149, 219-222, (see also genes; genetic transfer) group classification, 208-217, 227-256 haloacid dehalogenase, 208-215 halohydrin lyase, 209 homology groups, 213, 229, (see also enzyme superfamily) methyl halide dehalogenases, 120-121 methyl transferases, 162, 216, 341 pentachlorophenol hydroxylase, 249-253 pentachlorophenol monooxygenase, 228 reaction mechanisms, 142-145, 213-217, 228, 230-256 reductive dehalogenase, 145-149 regulation, 219-220 sequence analysis in classification, 213, 229, 230-256 tetrachlorohydroquinone dehalogenase, 228, 242-249 (see also dechlorinases; dehalogenation; reductive dehalogenases) dehalogenation, by plants, 470 effect of chloro substitution, 35, 94, 110, 350, 444 environmental constraints, 37-38, 40, 108-109, 450 pathway determination using Gibbs free energy, 95-96 pathways, 15, 16, 95-96, 208, 231, 236 role in metabolism, 116-117

INDEX dehalogenation activity, assessment and optimization, 389-394, (see also microcosms) global distribution of activity, 324-326, 386,445,449 dehalogenation mechanisms, 15-18, 35, 55, 207, 227, 473, dehydrodehalogenation, 18,100 dihaloelimination, 56, 211, 268-271 hydration, 18, 212 hydrogenolysis, 89-90, 268, 272 hydrolysis, 17, 55-57, 106, 179, 208, 210-211, 228, 230-242 methyl transfer, 18, 117-121, 179, 211 oxygenolysis, oxidation, 17,19-20, 39, 43, 55-57, 165, 179, 208, 211, 228, 249-253, 373-376 reduction, 16, 20, 55-56, 106, 115-116, 179, 228, 242-249, 268, 355, 373376 thiolysis, 17, 188, 215, 228 242-249 substitution, 211 vicinal reduction, 56, 100 dehalorespiration, 20, 40, 126-141, ATP production, 135 biogeochemical indicators, 357-358 electron acceptors, 69, 93, 102, 126-134, 379, 450 electron donors, 126 generation of proton motive force, 135, 138 hydrogen limitation, 377 hypothesized scheme for electron transfer, 140 topology of respiratory chain, 138 dehalorespiring bacteria (DRB), 71, ecological role, 38-39 global distribution, 385 growth yields, 133-135 molecular probes, 390 nutritional requirements, 134 phylogenetic groups, 126-134 physiology, 126-134 role in bioremediation, 385-386 Dehalospirillum multivorans, as chloridogen, 71 ecological role, 38 growth yield, 133 PCE dechlorination, 74, 390 PCE/TCE reductive dehalogenase, 137, 142

487

phylogeny and properties, 131 denitrification, degradation of organohalides, 38, 68 denitrifying bacteria, chlorobenzoate degradation, 38, 68 ecological role, 38 examples of dehalogenators, 39, 68 Desulfitobacterium chlororespirans, growth yield, 133 phylogeny and properties, 127 Desulfitobacterium dehalogenans, o-chlorophenol reductive dehalogenase, 137, 142 ecological role, 38 fortuitous cometabolism, 132 growth yield, 133 phylogeny and properties, 127 reductive dehalogenase, 133 Desulfitobacterium frappieri, growth yield, 133 PCE/TCE reductive dehalogenase, 137 phylogeny and properties, 127-129 Desulfitobacterium hafniense, o-chlorophenol reductive dehalogenase, 137 phylogeny and properties, 127 Desulfitobacterium sp., as chloridogen, 71-72 PCE dechlorination, 74, 390 Desulfobacterium autotrophicum, cometabolic reductive dehalogenation, 122 Desulfomonile sp., as chloridogen, 71 Desulfomonile limimaris, growth yield, 133 phylogeny and properties, 129 Desulfomonile tiedjei, anaerobic respiration, 71 biodegradation of 3-CB, 20, 43, 56, 57, 126 3-CB reductive dehalogenase, 137, 141 ecological role, 38 electron transfer by cytochromes, 139 fortuitous cometabolism, 132 growth yield, 133 phylogeny and properties, 129 Desulfovibrio dechloroacetivorans, as chloridogen, 71 ecological role, 38 growth yield, 133

488

Desulfovibrio dechloroacetivorans (continued), phylogeny and properties, 130 Desulfuromonas chloroethenica, ecological role, 38 growth yield, 133 phylogeny and properties, 130 Desulfuromonas sp., 71, 390 detoxification via dehalogenation, 15, 475 dibenzo-p-dioxins, chlorinated, (see polychlorinated dibenzo-p-dioxins) dibenzofurans, chlorinated, (see polychlorinated dibenzofurans) l,2-dibromo-3-chloropropane, 266, 475 l,2-dibromoethane(DBA; EDB), anaerobic dechlorination, 122 bioavailability, 296 mineralization by Mycobacterium, 208 natural production, 473 dibromoethene (DBE), 475, anaerobic dechlorination, 122 1,2-dibromomethane, genes for degradation, 61 dibromophenols, 7 3,4-dichloraniline, 316 dichloroalkanes, abiotic dechlorination, 265 biodegradation by fungi, 172 dichloroanisyl alcohol, 7 dichlorobenzenes, oxygenolytic dehalogenation, 55 2,4-dichlorobenzoate, dehalogenation, 16 dichloroelimination, 56, (see also dehalogenation mechanisms) 1,3-dichlorobutene, genes for degradation, 61 1,1 -dichloro-2,2-bis(4-chlorophenyl)ethane, (see DDD) 1,1 -dichloro-2,2-bis(4-chlorophenyl)ethene (see DDE) dichlorodifluoromethane (see CFC-12) dichlorodiphenyldichloroethane, (see D D D ) dichlorodiphenyldichloroethylene, (see DDE) 1,2-dichloroethane (DCA; ethylene chloride), 10, abiotic dehalogenation, 265, 270-271 anaerobic dechlorination, 122, 125 anaerobic mineralization, 116

INDEX

catabolic pathway, 208 degradation by Xanthobacter, 56, 208 evolution of degradation pathway, 217219 oxygenolytic degradation, 216 dichloroethane dehalogenase, enzyme evolution, 218 dichloroethene (DCE), abiotic dehalogenation, 262-263 aerobic degradation, 374 dehydrohalogenation product of TCA, 271 formation during PCE/TCE degradation, 11, 74, 101, 129-132, 269, 373, 386 in groundwater, 373 metabolism, 74, 101 mineralization under anoxie conditions, 374-375 dichlorofluoromethane, (see HCFC-21) dichloromethane (DCM), abiotic dehalogenation, 263-264 dechlorination, 164 dehalogenase, 215 growth substrate for anaerobic growth, 116, 121 mineralization by methylotrophs, 208 dichlorophenols, biodegradation by fungi, 182 electron acceptor for isolation, 72 glycosylation by fungi, 186 natural source, 7 2,4-dichlorophenoxyacetate (2,4-D), 61-67, 306-308, biodegradation by bacteria, 61 -64 biodegradation by fungi, 168 humified metabolites, 316-317 pathway of Dichomitus squalens, 170 1,3-dichloropropanol, aerobic mineralization, 208 1,3-dichloropropene, mineralization by Pseudomonas pavonaceae, 208 Dichomitus squalens, biodegradation of chlorinated pesticides, 168 biotransformation of chlorophenols, 182183 glycosylation reactions, 186 dicofol, biodegradation by fungi, 175

INDEX

dieldrin, bioavailability, 291 biodegradation by fungi, 166 production and use, 14 dioxins, chlorinated, (see polychlorinated dibenzo-p-dioxins) dioxygenases, dehalogenation, 17, 19, 55, 59, 444 (see also dehalogenases) diuron, 307 DRB, (see dehalorespiring bacteria) drosophilin A, 7, 162, 180, 184 electron accepting processes, comparison, 38-40, 376-377 competition, 43, 105-106, 357, 376-377 degree of chlorination, 376-377 determination of predominant pathways, 378 ecological role, 38, 43, 375-376 energetics, 95 field assessment, 389 impact on community structure, 43, 102105, 357, 376-377 redox conditions, 376-377 regulation, 133 role of 39,40,377,391 electron acceptors, 21, 89, 93, 127-131, 376377 carbonate, 39, 43, 93, 105, 117, 357, 374, 376-377 iron, 43, 117, 357, 374,377 manganese, 43, 117, 377 nitrate, 43, 68, 93, 103, 105, 132, 357, 377 organohalides, 20, 43, 55, 69-77, 93, 102-103, 126-134, 376, 449-450, (see also dehalorespiration) oxygen 15, 39, 106-108 sulfate, 43, 93, 105, 117, 357, 374, 377 electron donors,127-131, 357, acetate, 73, 77, 375-376 competition for, 390-393 fatty acids, 357, 392-394 for chloridogenesis (dehalorespiration), 76-77, 126 formate, 126 39, 74, 76, 126, 329, 357, 390, 392 substrates for dehalogenation, 127-131 electron flux (see also electron transfer), 33, dechlorination in reduced sediments, 364

489

role of and formate, 126, 357, 391 electron transfer, 140-141, coupling, 357 dehalorespiratory chain, 135-140 energy production during phosphorylation, 135, 138 role of cytochromes and quinones, 139 abiotic dehalogenation, 268-272, (see also dehalogenation mechanisms) endosulfan, biodegradation by fungi, 166 endrin, production and use, 14 energetics, dechlorination, 89-112 dechlorination vs. debromination, 109 degree of chlorination, 110 dehydrodehalogenation, 100-101 fermentation, 110 consumption, 103-105 prediction of product yields, 97 terminal electron accepting processes, 103-106 (see also thermodynamics) energy conservation, 3-CB dechlorination, 20, 69 in DRB, 70, 134-135 proton motive force, 139 proton pump, 141 via proton gradient, 135, 140-141 energy flow, coupling, 357 electron-accepting processes, 40 food chain, 44, 110 function of redox, 39, 44 role of dehalogenators in food chain, 103 role of 39-40, 44 energy yield (from dehalogenation), 89-114, comparison between halomethanes, 99 dechlorination of chloroethanes, 101 dechlorination of pesticides, 101 dechlorination vs. debromination, 109 and fractions, 102-103 thermodynamic predictions, 96 Enterobacter aerogenes, cometabolic reductive dehalogenation, 123 Enterobacter agglomerans, cometabolic reductive dehalogenation, 123

490

Enterobacter cloacae, cometabolic reductive dehalogenation, 123 Enterobacter sp., as chloridogen, 71 environment, constraints, 36-37, 108-109 determinants for biodegradation, 36-37, 46-47 effect on dehalogenation, 471 enzymes, (see dehalogenases) enzyme superfamily, aminohydrolase, 236-238 2-enoyl-CoA-hydratase/isomerase, 231 235 glutathione-S-transferase, 215, 242-249 fold, 213-215 short-chain reductase/dehydrogenase (SDR), 215 EOX (extractable organic halogen), 8 Escherichia coli, cometabolic reductive dehalogenation, 122-123, 125 ethene (ETH), dechlorination product of PCE, TCE, 386 ethylene chloride, (see dichloroethane) ethylene dibromide (EDB), 473 evolution, catalytic pathways, 14, 217, 227-256 2,4-D degradation, 67 dehalogenases, 217-219, 227-256 dehalogenating bacteria, 55 dehalogenation mechanisms, 55-56, 6269, 449 dissemination of degrading elements, 6269 enzyme regulation, 219-220 haloalkane dehalogenase, 217-219 role of microbial adaptation, 55 (see also genetic transfer) electron fraction used for energy production, 102-103 electron fraction for biosynthesis, 102-103 fatty acids, substrates for production, 392-394, 401-402 Fe(III), (see iron) Fe(0), (see zero-valent iron) fermentation,

INDEX

hydrogen production, 390-394, 401 pathways, 393 (see also electron donors) fermenters, ecological role, 38-39, 376-377 role in VC degradation, 375-376 Flavobacterium gleum, 427 Flavobacterium sp., 2,4-D degradation, 63 fluoroacetate, 7 fluoroform (trifluoromethane, TFM), atmospheric characteristics, 334 TFA metabolite, 334 fluorooleic acid, 7 fluorothreonine, 7 food chain, dehalogenating consortia, 44, 110 energy (electron) flow, 103, 377 role in biodegradation and detoxification, 42 “waste chain”, 110 fortuitous dehalogenation, 15, 19, 132, energy production, 102-103 freons, 332, 475, (see also chlorofluorocarbons; hydrochlorofluorocarbons; hydrofluorocarbons; specific compounds) fugacity ratio (of PCDDs), 352 fungi, dehalogenating, (see white rot fungi) fungicides, chlorinated, biotransformation, 305 gene cassettes, 59, (see also genetic transfer; genes) gene evolution, 217-221, gene clusters, 220 horizontal transfer, 241 genes, bph genes, 456, 458 cbaAB operon, 60, 220 elc genes, 65 cryptic genes, 220 2,4-D catabolism, 62-69 dem gene, 220 dehalogenases, 148-149 dehH2 gene, 220 dehI gene, 220 dhaA gene, 61,220 dhlA gene, 220 dhlB gene, 220 encoding dehalogenation, 59-61

INDEX genes (continued), evolution, 217-222 fcb genes, 59 global distribution, 222 methyl transferases, 341 mobile elements, 220-221 rdh genes, 145-148 regulation, 148-149, 219-220 sequences for dehalogenating enzymes, 120, 145, 219 tfd genes, 64-67 genetic engineering, PCB congener specificity, 456-459 genetic transfer, 59, 2,4-D degrading elements, 62-64 insertion sequences, 220-221 mobilization and distribution, 59, 220221 plasmids, 59, 62, 220-222 transposons, 59, 64, 220-221 (see also horizontal transfer) Gibbs free energy, calculation, 90-94 estimation by group contribution method, 102 of formation, 90 predicting product distribution, 97, 363 global distillation effect, 13 global warming potential (GWP), 323 glutathione-S-transferase, 215, (see also enzyme superfamily) granular iron, biotic dechlorination pathways, 270 groundwater, anaerobic/aerobic system, 374 chloroethene contamination, 373 degradation of chloroethenes, 376-377 hydrogen 377, 379 redox gradients, 374, 379 remediation, 379-382 haloalkane dehalogenase, evolution, 217 genetic organization, 219 (see also dehalogenases) halobenzoates, effect of substituent position on dehalogenation, 94 (see also chlorobenzoates), halocarbons, atmosphere, 323-327

491 biodegradation, 323-342 effect of ambient concentrations on degradation, 332 reaction mechanisms, 325 halocatechols, formation during aromatic dehalogenation, 19 halogen, discovery and production, 4 physicochemical properties, 3-4, 6 substituents, 94 halogenated aliphatic compounds, abiotic dehalogenation, 262-272 halogenated organic compounds, (see organohalides) halogenated pollutants, 54 halogenation, de novo synthesis, 161-164 haloperoxidase, 475 halogen cycle, 5-6, 54, 324, 469, microbiological sinks, 6, 13-15, 34, 55, 327-342 (see also biogeochemical cycling of organohalides) halohydrins, epoxide formation of, 19 halomethanes, 327-331 inhibitors of methanogenesis, 328-329 Halomonas sp., 2,4-D degradation, 63 halons, 326, (see also halocarbons) haloorganic compounds, (see organohalides) haloperoxidase, 475 halorespiration, (see dehalorespiration) HCFCs, (see halocarbons; hydrochlorofluorocarbons) HCFC-21 (dichlorofluoromethane), 329, (see also halomethanes; hydrochloro-fluorocarbons) HCFC-22 (chlorodifluoromethane), 329, (see also halomethanes; hydrochloro-fluorocarbons) heptachlor, production and use, 14 Herbaspirillum, 426 herbicides, biodegradation by fungi, 168 biotransformation, 307 dimerization of metabolites, 316 (see also specific compounds)

492

hexachlorobenzene (HCB), biodegradation by fungi, 172 production and use, 14 modeling aquifer dechlorination, 408 (HCH; lindane), anaerobic dechlorination, 122 bioavailability, 292 biodegradation by fungi, 166-167 biotransformation, 306 mineralization by Sphingomonas paucimobilis, 208 hexachloroethane (HCA), abiotic dehalogenation, 261 HFCs, (see hydrofluorocarbons) homoacetogens, anaerobic metabolism of chloromethanes, 116-121 cometabolic reductive dehalogenation, 123-125 growth on methyl chloride, 335 HOMO-LOMO gap, 350, effect of degree of chlorination, 363 horizontal gene transfer, gene casettes, 59 mechanisms, 67 role of DNA integrases, 221-222 hydraulic conductivity, (see site assessment) hydrochlorofluorocarbons (HCFCs), 329331, as biotransformation products, 329 cooxidation by methanotrophs, 331 degradation, 327-331 reactivity, 329 (see also halocarbons) hydrofluorocarbons (HFCs), 325, 331, 475, degradation, 332-335 (see also halocarbons) hydrogen abiotic dehalogenation with metals, 263266 competition between methanogens and sulfidogens, 39, 77, 102-105, 126, 357, 392-393 electron donor for dehalogenation, 7677, 357, 391 in groundwater, 377, 389 partial pressure, 104 stimulation of PCDD dechlorination, 361-363 uptake and utilization, 39, 377 hydrogenotrophic dehalogenators, 392, (see

INDEX

also DRB) hydrogen threshold concentration, 39, 102105, 377, 392-393 hydrogen transfer, 40, energy flow in microbial community, 44, 391 hydrolytic dehalogenation, 17, 19, 55-57, 106, 208, 210-211, 228, 230-242 hydroquinones, chlorinated, biodegradation by fungi, 181,183 methylation by fungi, 186 Hyphomicrobium chloromethanicum, growth on methyl chloride, 341 Hyphomicrobium sp., growth on chloromethane, 217 haloaliphatic dehalogenase, 212 Hypholoma sp., synthesis of organochlorides, 162-163 insecticides, biodegradation by fungi, 167 biotransformation, 306 (see also pesticides; specific compounds) in situ dechlorination, 445, indicators, 451 in situ remediation, addition of electron donor, 390-392 assessment of microbial activity, 388 competition for electron donor, 390-393 microcosms for field design, 385-415 model development, 407-415 permeable barriers, 46, 261 site hydrological and physicochemical evaluation, 386-388 treatment strategies for anaerobic dechlorination, 451-454 (see also bioremediation; site assessment) intrinsic bioremediation, (see bioremediation) iodobutanes, 7 iodomethanes, 7 1-iodopropane, dehalogenase inhibitor, 141 inhibition of dechlorination, 125 Ionotus dryophilus, PCP dechlorination, 178 iron, abiotic dehalogenation, 262-271, 276280

INDEX

iron (continued), as electron acceptor, 45-46 porphyrins mediating dehalogenation, 124, 472 used in bioremedation, 46, 261 (see also zero-valent iron) iron minerals, role in abiotic dehalogenation, 278-279 iron-reducing bacteria, ecological role, 38 iron-reducing conditions, mineralization of DCE, VC, 374 laccase, 160 lateral gene transfer, 59, (see horizontal gene transfer) Leisingera methylohalidivorans, growth on methyl halides, 341 Lentinula edodes, biotransformation of PCA, 179 Lentinus sp., biodegradation, 172 lignin, 159 lignin degraders, role in dehalogenation, 159-204 ligninolytic enzymes, 160, biodegradation of chlorinated phenols, 181-186 components, 159-160 oxidation of chloroanilines, 191-192 products of reaction, 178-184 reaction mechanisms, 161, 184 role in dehalogenation, 20, 174-179 substrate polymerization, 184-185 substrates, 161, 178-185 lignin peroxidase (LiP), 160-161, reaction mechanism with CT, 165 role in PCP degradation, 177-179 lindane, (see low donors, 392, 406 manganese, as electron acceptor in soil, 45 manganese peroxidase (MnP), 160-161, role in 2,4-D degradation, 170 role in PCP degradation, 177-178 matrix mineralogy of soil, 44-46, composition, 45 impact on microbial ecology, 44 menaquinones, (see quinones) metabolic strategies, 15, 19-21, 33-35, 56-

493

77, 425-426, (see also dehalogenation mechanisms) metabolism, (see aerobic metabolism; anaerobic metabolism) metals for abiotic dehalogenation, 262-282, 358, bimetal combinations, 263, 265 catalytic mechanisms, 276-280 methane monooxygenase, 10, 42 Methanobacterium thermoautotrophicum, cometabolic reductive dehalogenation, 122, 471 ecological role, 38 methanogens, cometabolic reductive dehalogenation, 122-125, 329 competition with DRB, 39, 126, 105106, 377, 393 ecological role, 38-39 possible PCB dechlorinators, 450 methanogenesis, (see methanogens) Methanosarcina barkeri, cometabolic reductive dehalogenation, 122 ecological role, 38 reductive dehalogenation of CFCs, 329 Methanosarcina thermophila, TCE dechlorination, 125 methanotrophs, (see methylotrophs) methoxychlor, biodegradation by fungi, 175 biotransformation pathways, 306 methylation, of chlorophenols, 186, 423, 426 methyl bromide, 335-342, degradation in sediments, 336 soil as global sink, 341 toxicity, 475 use as soil fumigant, 335 (see also bromomethanes; methyl halides) methyl chloride, aerobic metabolism, 120-121 anaerobic dechlorination, 118-120 degradation, 335-342 dehalogenase, 120 (see also chloromethanes; methyl halides), methyl chloroform, 326, 331, (see also trichloroethane) methyl fluoride, 337, (see methyl halides)

494

methyl halides, dechlorination pathway, 341 degradation in the environment, 335-341 natural sources, 335 role in ozone depletion, 335 (see also bromomethanes; chloromethanes; halomethanes) Methylobacterium chloromethanicum, 38, growth on methyl chloride, 341 Methylobacterium sp., degradation of chloromethanes, 120, 208 growth on methyl chloride, 337 haloaliphatic dehalogenase, 211-212 Methylophilus sp., haloaliphatic dehalogenase, 211 mineralization of dichloromethane, 208 Methylosinus trichosporium, dehalogenation of halomethanes, 471 methylotrophs, cooxidation of organohalides, 329-332 degradation of methyl halides, 208, 211212, 337 growth on methyl halides, 338-339 phylogenetic classification, 340 role in dehalogenation, 39, 120-121 methyl transfer, role in methyl chloride metabolism, 118120, 341 metolachlor, biochlorination by fungi, 168 pathway by Phanerochaete chrysosporium, 169 sorption in soil, 296 microbial ecology, 33-48 microbial infallibility, 14, 305-308 microbial mediators, 38 role in dehalogenation, 13, 35 (see also biodegradation; dehalorespiring bacteria) microbial production of organohalides, 6, 55, 161-164, 218, 473 Micrococcus sp., growth on chlorobenzoates, 58 microcosms, case study of TCE contamination, 397402 determination of site-specific model parameters, 403-414 optimization of dehalogenation potential, 389-394 protocols, 395-397

INDEX

use in field design, 389-402 mineralization, definition, 312-313 (see also biodegradation) mirex, production and use, 14 Mn(IV), (see manganese) modeling in situ bioremedation, biokinetic expressions, 408-409 BIOPLUME III, 407 development of transport/fate models, 403-414 kinetic parameters for dechlorination activity 408 reductive dechlorination, 407-410 site-specific parameters, 410-411 use in field design of bioremediation, 407-414 UTCHEM, 407 molecular probes, for DRB, 390, 398, 450 2-monobromodioxin (MBrDD), primer for PCDD dechlorination, 361 monofluoroacetate, metabolic inhibitor, 334 natural production, 334 monooxygenases, ammonia monooxygenase, 19, 337 dehalogenation, 20, 55, 68, 337 methane monooxygenase, 10, 19, 42, 337,471 methyl halides as competitive inhibitors, 337 (see also dehalogenases; methylotrophs) Montreal Protocol, 13, 323, 325-326 Moraxella sp., 62, haloaliphatic dehalogenases, 210 multiphase partitioning, 41, 294 mycelial color removal (MyCoR) system, 194 mycobacteria, role in dehalogenation, 39 Mycobacterium aurum, growth on VC, 216 Mycobacterium chlorophenolicum, 20, dehalogenation, 20, 249 ecological role, 38 PCP degradation, 66, 426 Mycobacterium fortuitum, 20, 426 Mycobacterium sp., haloalkane degradation, 61, 208

INDEX

MYCOPOR treatment process, 194 NAPLs (non-aquaeous phase liquids), 34 natural organic matter (see organic matter) natural production of organohalides, 5-9, 5355, 161-164, (see also organohalides) nitrate, as electron acceptor in soils, 45 (see also denitrifying bacteria; electron acceptors) nitrate reduction, (see denitrification; denitrifying bacteria) Nocardioides simplex, 2,4-D degradation, 63 Novosphingobium, 425, (see Sphingobium) nutrients, enhancement of dechlorination activity, 36-39, 394-396 Ochrobactrum sp., growth on 3-CB, 68 organic matter, 41, 294, 423, impact on bioavailability, 294, 298 production of chlorinated NOM, 316318 organochlorine, synthesis by white rot fungi, 161-164 synthons for biosynthesis, 163-164 (see also organohalides) organofluorides, 334, (see also organohalides) organohalides, as terminal electron acceptor, 69-77 biodegradation by fungi, 164-197 biogenesis, 5-8, 53-54 fate in the environment, 4-5, 13, 110, 295-297, 323-327, 471 genes encoding degradation, 60, (see also genes) geogenesis, 8, 54 industrial, anthropogenic sources, 9-12, 53 mineralization under anoxic conditions, 116 natural production, 5-9, 53-55, 161-164, 218, 473 physicochemical characteristics, 3-4, 6, 41, 110, 292, 308-309, 324-327, 348-351, 421-422 production and use, 4-5,10-12, 14, 161-

495

164, 303-304, 351-353, 443, 473 pyrogenesis (forest fires; volcanic emissions) 9, 54, 473 organophosphorus pesticides, chlorinated, biodegradation by fungi, 166 oxygenase-mediated dehalogenation, 15, 1920, 39, 55, 249, (see also dioxygenases; monooxygenases) oxygenolytic dehalogenation, (see dehalogenation mechanisms; oxygenasemediated dehalogenation) ozone, depletion by halohydrocarbons, 13, 332333 depletion potential (ODP), 323 Panus tigrinus, biotransformation of chlorophenols, 180, 183 PCBA, (see pentachlorobenzyl alcohol) PCBs, (see polychlorinated biphenyls) PCDDs, (see polychlorinated dibenzo-pdioxins) PCDFs, (see polychlorinated dibenzofurans), PCE, (see tetrachloroethene) PCP, (see pentachlorophenol) pentachloroanisole (PCA), 426, biotransformation by fungi, 179 (see also chloroanisoles) pentachlorobenzyl alcohol (PCBA), biotransformation, 314-3150 (see also herbicides) pentachlorophenol (PCP), aerobic biodegradation, 16, 20, 67-68, 177-179, 242, 426 as electron acceptor, 72 dehalorespiration, 127-131 O-methylation, 423 pathways for metabolism, 94, 305 toxicity, 424 wood treatment, 351 (see also chlorophenols) perchloroethene (see tetrachloroethene) perchloroethylene (see tetrachloroethene) perfluorooctanyl sulfonates (PFOs), 54 permeable reactive barriers, 46, 261 peroxidases, 159-161, 165, 170, 176, polymerization of chloroaromatic residues, 317 (see also ligninolytic enzymes)

496

persistent metabolites, (see recalcitrance) pesticides, bioavailability, 292 biodegradation, 166, 174 biotransformation, 303-319 chloracetanilides, 168, (see alachlor; metolachlor) chlorinated atrazines, 168, (see atrazine) chlorinated organophosphorus compounds, 166-168 chloroheterocyclic compounds, 166-167, (see aldrin; chlordane; dieldrin; lindane) chlorophenoxyacetates, 168, (see also 2,4-dichlorophenoxyacetate; 2,4,5trichlorophenoxyacetate) coupled anaerobic/aerobic processes, 312-315 energetics of dehalogenation, 102 production and use, 12, 303-305 sorption in soil, 296-297 triazines, 168, (see also atrazine) (see also fungicides; herbicides; insecticides) pH, effect on dechlorination, 387 impact on redox, 47 requirements for bioremediation, 36-37, 387-388 speciation of metals, 47 Phanerochaete chrysosporium, biodegradation of chlorinated organics, 164, 166, 168, 171-174, 175, 177192, 425-427 biotransformation of PCDDs, 190-191 ligninolytic system, 164, 181-186 methylation reactions, 186, 426 mycelial color removal (MyCoR), 194 MYCOPOR treatment process, 194 reductive dehalogenase, 188 Phanerochaete sordida, biotransformation of lindane, 166 biotransformation of PCDDs, 188-191 PCP biodegradation, 178, 426 Phellinus pomaceus, synthesis of chloromethane, 162 Phellinus weirii, DDT biodegradation, 175 Phellinus sp., synthesis of organohalides, 161-162 phenylpropanoid compounds, chlorinated,

INDEX

de novo synthesis, 162 Phlebia strigoso-zonata, DDT biodegradation, 175 Phlebia tremellosa, biodegradation of chlorinated pesticides, 168, 175 photoirradiation, with iron, 271 phototrophic dehalogenators, 68 ecological role, 38 phylogenetic analysis of DRB, affiliation of dechlorinating isolates, 78, 127-131 Gram-positive bacteria, 60, 127-129, green non-sulfur, 131 129-130 131 phylogenetic classification, methyl halide-degrading methylotrophs, 340 picloram, sorption in soil, 296 plasmids for dehalogenation, 59, 64, (see also genetic transfer) Pleurotus ostreatus, biodegradation of chlorinated pesticides, 175, 179 biodegradation of PCBs, 171-172 Pleurotus pulmonarius, biotransformation of chlorinated pesticides, 168 polybrominated biphenyls (PBBs), stimulation of PCB dechlorination, 109, 449-450 polybrominated dibenzodioxins (PBDDs), 7 polychlorinated biphenyls (PCBs), 443-460, abiotic dehalogenation, 265 aerobic biodegradation, 454-455 as electron acceptor, 73-74, 449-450 bioavailability, 292, 296, 450 biodegradation by fungi, 171-174 biodegradation, two-stage process, 454 congener-specific analysis, 445-446 dechlorination, 16, 444-451, 453, 457459 determination of product ratios, 98 dredging, 443 enhanced degradation, 452 genetic engineering, 456-460 in situ biodegradation, 454 pathways, 448, 454-459

INDEX

polychlorinated biphenyls (continued), production and use, 14, 54, 443 structure and nomenclature, 444 (see also Aroclors) polychlorinated dibenzo-p-dioxins (PCDDs), 347-368 bioavailability, 435 biodegradation by fungi, 190-191 cometabolism, 364 congener profiles, 365 dechlorination kinetics, 363-365 dechlorination pathways, 358-361 dechlorination yields, 359 inducers of dechlorination, 361-363 natural production, 54, 351 reactivity in sediments, 353-357, 363 reductive dehalogenation, 358 remediation, 365-368 source-sink correlations, 351-354, 365 toxicity, 13 transchlorination reactions, 351 polychlorinated dibenzofurans (PCDFs), bioavailability, 295, 435 natural production, 54 polymerase chain reaction (PCR), correlation to dechlorination activity, 398-399 (see also molecular probes) polymerization of chlorophenols, 422-423 polyvinyl chloride (PVC), 9 POPS (persistent organic pollutants), 13-14 propanil, 307 Proteus mirabilis, cometabolic reductive dehalogenation, 123 proton motive force, energy production, 135 proton pump, 141 pseudomonads, role in dehalogenation, 39 (see also individual species) Pseudomonas acidovorans, DDT biotransformation, 314 Pseudomonas aeruginosa, growth on dichlorobenzoate, 315 growth on VC, 216 Pseudomonas cepacia, (see Burkholderia cepacia) Pseudomonas pavonaceae, 61 dichloropropene mineralization, 208 Pseudomonas putida,

497

cometabolic reductive dehalogenation, 122, 125 DDT dehalogenation by cytochrome 472 ecological role, 38 growth on 4-CB, 60 haloaliphatic dehalogenase, 211 Pseudomonas recinovorans, chlorophenol degradation, 431 Pseudomonas sp., dechlorination mechanism, 231, 236 degradation of dichloroalkanes, 55, 57, 208 growth on chlorobenzoates, 58, 60, 68, 230 haloaliphatic dehalogenases, 210-212 haloaromatic dehalogenases, 230-242 mobile elements for 2,4-D degradation, 62 PCP degradation, 66, 426 Pseudomonas stutzeri, 38-39 PVC, (see polyvinyl chloride) Pycnoporus cinnabarinus, biotransformation of chlorodiphenyl ether, 173 biotransformation of chlorophenols, 183 glycosylation of chlorodiphenyl ethers, 186 QSAR modeling, 473 quinones, role in electron transfer, 139-140 radical clocks, 272 Ralstonia eutropha, 2,4-D degradation, 61 DDT biotransformation, 314 dechlorination of PCB metabolites, 457 ecological role, 38 recalcitrance, biodegradation vs. biotransformation, 311-313 compounds resisting degradation, 222223, 305-309 redox, competition for electron donor, 39, 375376 coupled reactions, 34, 58, 117 groundwater, 375-376 microbial ecology, 39, 375 processes for biodegradation, 373-382

498

redox (continued), soil, 43 spectrum, 38 redox conditions, assessment in groundwater, 379, 382 impact on electron accepting processes, 377-382 support for dehalogenation, 21, 376, 379-382 redox potential, role in energetics, 89-111, 376-379 reducing equivalents, 391, 403, (see also electron donors; hydrogen) reductive dechlorination, (see reductive dehalogenation) reductive dehalogenase genes, 145-149 regulation, 148-149 reductive dehalogenases (RDase), amino acid sequences, 145-148 corrinoid-dependent reaction, 145, (see also methyl transfer) genetics, 145-149 reaction mechanism, 142-145 (see also dehalogenases) reductive dehalogenation, 15, 34, 56, 89, 115-116, aerobes, 19, 54-77 aerobic conditions, 90 anaerobes, 20-21, 54-77 cometabolism, 122-126 phototrophs, 68-69 regiospecificity of reaction, 96 role of tetrapyrrole cofactors, 20 thermodynamic calculations, 93-95 remediation, dioxins, 366-367 ex situ applications, 366, 428, 432 (see also bioremediation; in situ remediation) respiration, role of chloroorganics as electron acceptors, 69-70 (see dehalorespiration; see also electron acceptors) Rhodococcus erythopolis, haloaliphatic dehalogenase, 210 Rhodococcus rhodochrous, 2,4-D degradation, 61-62, Rhodococcus sp., dechlorination of PCB metabolites, 457 Rhodoferax fermentans,

INDEX

2,4-D degradation, 63 Rhodopseudomonas palustris, ecological role, 38 rice paddy soils, dehalogenation activity, 20, 314-315 salinity, effect on biodegradation, 46, 357 SDR enzyme family, short-chain reductase/dehydrogenase, 215, (see also enzyme superfamily) sediments, (see soils and sediments) Serratia marcescens, cometabolic reductive dehalogenation, 123 Shewanella putrefaciens, cometabolic reductive dehalogenation, 122, 124 simazine, 307 site assessment, microbiological activity, 388-389 physicochemical characteristics, 387388 site reactivity probes, 465, 473 soils and sediments, aging phenomenon, 297-299, 422 bioremediation, 21, 365-367, 397-402, 428-435 global sink for methyl bromide, 341 heterogeneity and variability in sampling, 310-311, 385-386 inoculation, 167, (see also bioaugmentation) mineral matrix, 43-46 multiphasic system, 42 natural organic matter, 294 redox and electron acceptors, 36, 38, 4246 sorption phenomena, 295-297, 422 solubility, 305-309, effect of chlorination on PCBs, 450 solvents, halogenated, 10-12, as electron acceptors, 74 bioavailability, 292 biodegradation (see also specific compounds) sorption kinetics, 298-296 sorption of organohalides, aging phenomenon in soil, 297-299, 422 effect on biodegradation, 294-299 multiphase partitioning, 294-297

INDEX

fractions, 295 Sphingobium chlorophenolicum, 20, degradation of PCP, 66, 68, 249, 425427 ecological role, 38 reductive dechlorination, 242 Sphingomonas chlorophenolica, (see Sphingobium chlorophenolicum) Sphingomonas paucimobilis, 2,4-D degradation, 63 growth on 4-CB, 60 haloaliphatic dehalogenase, 210, 212 mineralization of hexane, 208 Sphingomonas sp., 2,4-D degradation, 63 growth on chlorobenzoates, 58 growth on PCP, 66 Staphylococcus epidermidis, cometabolic reductive dehalogenation, 123 Streptomyces cattleya, synthesis of monofluoroacetate, 334 Streptomyces rochei, PCP degradation, 426 suicide metabolites, formation during meta-cleavage, 19, 457 sulfate, soil and marine environments, 45 sulfate-reducing bacteria (sulfidogens), competition with DRB, 126 ecological role, 38 possible PCB dechlorinators, 450 sulfate-reduction (sulfidogenesis), 38 competition with dehalorespiration, 105 (see also electron acceptors) synthons for biosynthesis, 163-164 2,4,5-T, (see 2,4,5-trichlorophenoxyacetate) TAT system, 148, (see also twin arginine translocation) TCE, (see trichloroethene) TEAPs, terminal electron accepting processes, (see electron accepting processes) temperature, effect on biodegradation, 36, 46-47 effect on DDT biodegradation, 313-314 physicochemical effects, 46 quotient, 46 requirements for bioremediation, 387

499

tetrachloroazobenzene, 316 2,3,7,8-tetrachlorodibenzodioxin, (see polychlorinated dibenzo-p-dioxins) 1,1,2,2-tetrachloroethane, abiotic dehalogenation, 261, 265 tetrachloroethene (PCE; perchloroethylene; tetrachloroethylene), abiotic dehalogenation, 261-263, 279, 281 as electron acceptor, 72-73, bioavailability, 291, 296 dechlorination pathway, 376, 386, 391 dehalorespiration, 127-132 production and use, 10, 53 reductive dehalogenation, 16, 122, 125, 376 tetrachloroethylene, (see tetrachloroethene) tetrachlorohydroquinone (TCHQ), 242-249, 426 tetrachlorophenol, 421, (see also chlorophenols) tetrahydrofolate, role in dehalogenation, 118-120, TFA (see trifluoroacetate) TFM (trifluoromethane; see fluoroform) Thauera chlorobenzoica, 38-39 Thauera sp., growth on 3-CB, 68 thermodynamic indicators of dehalorespiration, 102 thermodynamics of dechlorination, 89-112, competition for 103 -105 considerations for bioremediation, 111 threshold concentrations, thermodynamic and physiological aspects, 103-105 thyroxine, 7 topology of dehalorespiratory chain, 138 TOX, total organic halogen, 194 toxaphene, production and use, 14 toxicity of organohalides, 41 Trametes hirsutus, biotransformation of lindane, 166 PCB mineralization, 171 PCP biodegradation, 178 Trametes versicolor, biodegradation of chlorophenols, 180, 182-183 biodegradation of organohalides, 172173, 178

500

Trametes versicolor (continued), DDT biodegradation, 175 glycosylation of chlorodiphenyl ethers, 186 trametol, 162 transhalogenation reactions, 335, (see also polychlorinated dibenzo-p-dioxins) transposons, 59, (see also genetic transfer) tribromophenols, 7 trichloroacetate, 307 as electron acceptor, 77, 127-131 Trichlorobacter thiogenes, as chloridogen (dehalorespirer), 71 phylogeny and properties, 130 1,2,4-trichlorobenzene (TBA), anaerobic dechlorination, 122 2,3,6-trichlorobenzoate (TCB), 307, formation, 315 mineralization, 315 1,1,1 -trichloroethane (TCA), abiotic dehalogenation, 261, 264-265, 270-271 anaerobic dechlorination, 122 anaerobic mineralization, 116 atmospheric concentration, 326 trichloroethene (TCE; trichloroethylene), 11, abiotic dehalogenation, 262-263, 276278 bioavailability, 292, 296 bioremediation of contaminated aquifer, 397-402 dechlorination pathway, 376, 386, 391 dehalorespiration, 127-131 mineralization by Phanerochaete chrysosporium,164 oxygenase-mediated dehalogenation, 19, 42 production and use, 53 reductive dehalogenation, 16, 42, 376 trichloroethylene, (see trichloroethene) trichlorofluoromethane, (see CFC-11) trichloromethane, (see chloroform) trichloronitromethane, dechlorination, 124, 473 trichlorophenols (TCP), as electron acceptor, 72-73 biodegradation by fungi, 180 dehalogenation, 16 2,4,5-trichlorophenol (TCP), 180, (see also trichlorophenols)

INDEX 2,4,6-trichlorophenol (TCP), 180, 421, (see also trichlorophenols) 2,4,5-trichlorophenoxyacetate (2,4,5-T), 307, dioxin byproducts, 351 mineralization by fungi, 168 trichlorotrifluoroethane (see CFC-113) triclosan, glycosylation by fungi, 186 (see also chlorodiphenyl ethers) trifluoroacetate (TFA), biodegradation, 333-334 formation from halocarbons, 333 trifluoromethane (TFM; see fluoroform) twin arginine translocation (TAT), role in electron transfer, 141, 148 Variovorax sp., 2,4-D degradation, 63 VC, (see vinyl chloride) vicinal reduction, 56, (see dehalogenation mechanisms) vinyl chloride (VC), abiotic dehalogenation, 262, 273 aerobic degradation, 216, 374 as electron acceptor, 76, 101 dechlorination pathway, 376, 386 dechlorination via acetogenesis, 117 formation during PCE, TCE degradation, 42, 74, 101, 373 in groundwater, 373 mineralization under anoxic conditions, 374-375 production and use, 10 toxicity, 42 vitamin cofactor requirement for dehalogenation, 263-265, 394 reductive dehalogenation reactions, 102, (see also cobalamin) volatile organohalides, degradation, 327-331 environmental effects, 12-13, waste chain, (see food chain) white rot fungi, biodegradation of chlorophenols, 177190, 426, 430-431 biodegradation of PCBs, 171-174 biodegradation of PCDDs, 190-191 dechlorination of chlorolignin, 192-193

INDEX

white rot fungi (continued), de novo synthesis of organohalides, 162164 ligninolytic enzymes, 177-190 methylation and glycosylation, 186 organochlorine production, 161-164 pathway for DDT biodegradation, 176 polymerization reactions, 184-185 treatment of bleachery effluents, 192 (see also ligninolytic enzymes, Phanerochaete chrysosporium) Xanthobacter autotrophicus, dechlorination of chloroalkanes, 56-57 ecological role, 38 haloaliphatic dehalogenases, 210 mineralization of dichloroethane, 208

501

xenobiotic compounds, 4, 11, 33, 53, 164, 207, 469, definition, 306-308 evolution of metabolic pathways, 207, 217-219, 227-256 residues, 191-192 zero-valent iron, abiotic dehalogenation, 261-282 dehalogenation pathways, 269 in situ treatment, 46,282 kinetics, 274 use in bioremedation, 282 (see also granular iron; permeable reactive barriers) zinc, abiotic dehalogenation, 268-272