wEPA
           United States
           Environmental Protection
           Agency
               Hazardous Waste Engineering
               Research Laboratory
               Cincinnati OH 45268
Research and Development    EPA/600/2-86/090

Microbial
Decomposition of
Chlorinated
Aromatic
Compounds

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                                    EPA/600/2-86/090
                                       September 1986
     MICROBIAL DECOMPOSITION
             OF CHLORINATED
        AROMATIC COMPOUNDS
        Melissa L. Rochkind and James W. Blackburn

                    IT Corporation
               Knoxville, Tennessee 37923
                         and
                     Gary S. Sayler
              The University of Tennessee
               Knoxville, Tennessee 37916
                Contract No. 68-03-3074

               Technical Project Monitors
               P.R. Sferra and J.A. Glaser
            Alternative Technologies Division
    Hazardous Waste Engineering Research Laboratory
                 Cincinnati, Ohio 45268
HAZARDOUS WASTE ENGINEERING RESEARCH LABORATORY
        OFFICE OF RESEARCH AND DEVELOPMENT
       U.S. ENVIRONMENTAL PROTECTION AGENCY
                CINCINNATI, OHIO 45268

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                          DISCLAIMER
  The information in this document has been funded wholly or in part by the United
States Environmental  Protection Agency  under  Contract  68-03-3074 to IT
Corporation. It has been subject to the Agency's peer and administrative review, and
it has been approved for publication as an EPA document. Mention of trade names
or commercial products does not constitute endorsement or recommendation for
use.

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                             FOREWORD

  Today's rapidly developing and changing technologies and industrial products
and practices frequently carry with them the increased generation of solid and
hazardous wastes. These materials, if improperly dealt with, can threaten both public
health and the environment. Abandoned waste sites and accidental releases of toxic
and hazardous substances to the environment also have important environmental
and  public  health implications.  The Hazardous Waste  Engineering Research
Laboratory assists in providing an authoritative and defensible engineering basis for
assessing and solving these problems. Its products support the policies, programs and
regulations of the Environmental  Protection Agency, the permitting and other
responsibilities of State and local governments and the needs of both large and small
business in handling their wastes responsibly and economically.
  This report is a compendium describing the current level of understanding of
chlorinated aromatic compound decomposition by microbiological pathways. The
halogenated aromatic compounds  are one  of the most persistent collections of
chemicals contaminating the environment. The persistent nature of these chemicals is
attributable to the inability of the environment to cleanse  itself of these contami-
nants. Since microbiological communities  are  fundamental participants in the
detoxification chain, the environment generally does not have microorganisms
capable of degrading the halogenated aromatic compounds. This report specifically
identifies microorganisms capable of degrading many of the halogenated organic
species. In many cases, the substrate is tracked through a decomposition pathway to
end product. Many factors contribute to the biodecomposition of a given chemical;
among the most important are: the chemical nature of the  substrate molecule and
substituents,  substrate concentration, environmental parameters, nutrient  and
growth factor availability, and the presence of organisms capable of degrading the
substrate.  However, insufficient information is  currently  available to assess the
potential biodegradability of a substrate based  on information generated under
different environmental conditions.
                                                   David G. Stephan
                                                        Director
                                              Hazardous Waste Engineering
                                                  Research Laboratory
                                   in

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                              ABSTRACT

   This report was initiated because of a need to bring together a review of the
literature pertaining to microbial metabolism of chlorinated aromatic compounds.
The information gathered here is extensive although not exhaustive. Most attention
has been given to reports of bacterial, fungal, and cyanobacterial pathways of
substrate degradation where metabolites or end products have been identified.
Studies which report data on metabolites arising from incubation of the substrate
with mixed cultures or environmental samples and  studies which show disap-
pearance of the compound have also been evaluated  and included.
  In addition to separate chapters on each class of chlorinated aromatic compounds,
reviews of microbial physiology, genetics, and methods of biodegradation assess-
ment are included.  One chapter reviews biodegradation of these compounds in
scaled-up processes.
  The  potential  biodegradation  pathways  for  all  classes of chloroaromatic
compounds have been brought together into an overview diagram.
  The review indicates that many factors are involved in assessing the biodegrad-
ability of a compound including the nature of the molecule, substrate concentration,
environmental parameters, availability of nutrients and growth factors, and presence
of degradative microorganisms. Not enough information is presently available to
permit extrapolation from one environment to another or to utilize data on a similar
compound to assess the biodegradability potential of a given substrate.
  This report includes an illustrated list of compounds, a glossary, a reference list, an
additional bibliography, citation index, and a subject index.
                                    IV

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                              CONTENTS

Foreword  	   iii
Abstract  	   iv
Figures  	  viii
Tables  	   xi
Abbreviations and Symbols  	  xii
Acknowledgments 	  xiii

      1. Introduction	   1

      2. Conclusions  	   4

      3. Overview of Microbial Physiology  	   6
        Microbial cell structure	   7
        Growth requirements  	  10
       The cell growth cycle	  11
        Population growth  	  11
       Continuous culture  	  12
       Cell death  	  13
        Pure and mixed culture metabolism  	  13
        Substrate uptake and transport	  13
       Enzymes  	  14
        Metabolic energy production  	  16
       3-Ketoadipic acid pathway	  18
       Plasmids  	  19

      4. Cellular Gene Coding and Genetic Technologies  	  20
       Structure and function of DNA  	  20
       Transcription  	  23
       Translation  	  23
       Mutagenic events   	  25
       Current biochemical tools for genetic manipulation  	  28
       Mechanical shearing of DNA  	  31
       Cloning vehicles  	  31
       Methods of manipulating  DNA  	  34
       Mapping of restriction endonuclease recognition sites	  35
       Identification of DNA sequences within fragments	  36
       Expression of prokaryotic genes in foreign hosts	  36
       Gene cloning in yeasts	  36
       Expression of eukaryotic genes in a prokaryotic host  	  36

     5. Methods of Biodegradation Assessment	  38
       Chemical analytical techniques  	  41
       Analyses of metabolic activity	  43
       Parameters for pure culture studies	  44

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  6. Metabolism of Nonchlorinated Aromatic Compounds	     . .
   Chemistry of benzene and substituted benzenes  	     ..
   Microbial attack on benzene structures	     ,_
   Attack on aromatic structures by cyanobacteria	     ^_
   Ring fission of dihydroxy aromatic compounds by bacteria	     ~.
   Attack of aromatic structures by eukaryotes	
   Degradation of dihydroxylated aromatic compounds by
     yeasts and fungi	     -Jl
   Summary  	     "'

  7. Chlorobenzoic Acids	     ""
   Bacterial metabolism and chlorobenzoic acids	     69
   Algal metabolism of chlorobenzoic acids  	     74
   Fungal metabolism of chlorobenzoic acids  	     77
   Metabolism of chlorobenzoic acids in soils and by consortia  	     77
   Reductive dechlorination  	     78
   Summary   	     80

  8. Chlorobenzenes   	     81
   Microbial metabolism of chlorobenzenes	     81
   Metabolism of chlorobenzenes by microbial communities	     86
   Summary  	     87

 9. Chlorophenols	     88
   Bacterial metabolism of chlorophenols  	     88
   Metabolism of chlorophenols by fungi   	     91
   Metabolism of chlorophenols by mixed microbial cultures	     91
   Summary  	     93

10. Pentachlorophenol	     94
   Bacterial metabolism of PCP	     94
   Fungal metabolism of PCP	     96
   Disappearance of PCP in environmental samples	     96
   Summary  	     98

11. Chlorophenoxy and Chlorophenyl Herbicides 	    99
   2,4-D	    99
   MCPA  	   104
   2,4,5-T	   104
   4-Chlorophenoxyacetic acid  	   106
   Other phenoxy herbicides  	   106
   Fungal metabolism of phenoxy herbicides  	   109
   Metabolism of phenoxy herbicides in soils  	   Ill
   Chlorophenyl herbicides  	   113
   Summary  	   114

12. Phenylamide and Miscellaneous Herbicides	   115
   Bacterial metabolism of chlorinated anilines	   115
   Fungal metabolism of chlorinated anilines  	   118
   Metabolism of chlorinated anilines in soils  	   118
   Metabolism of urea herbicides   	   120
   Metabolism of chlorinated phenyl carbamate herbicides	   122
   Metabolism of acyl anilide herbicides 	   123
   Miscellaneous pesticides  	   126
   Summary  	   128

                               vi

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    13. Chlorinated Biphenyls  	  129
      Microbial metabolism of PCBs	  129
      Metabolism of PCBs by mixed microbial cultures	  135
      Summary  	  136

    14. DDT and Related Compounds	  138
      Bacterial metabolism of DDT	  138
      Fungal metabolism of DDT 	  142
      Fungal metabolism of other compounds	  143
      Persistence and degradation of DDT in the environment 	  143
      Summary  	  144

    15. Chlorinated Dioxins and Dibenzofurans  	  146
      Microbial metabolism of dioxins and furans	  146
      Dioxin persistence and degradation in  soils	  149
      Summary  	  149

    16. Biodegradation of Chlorinated Aromatic Compounds in
         Scaled-up Biological Water Related Treatment Processes	  152
      Introduction  	  152
      Pentachlorophenol  	  154
      Chlorinated biphenyls	  156
      Dichlorophenol	  162
      Trichlorocarbanilide	  164
      Dichlorobenzene	  165
      Combined studies on several classes  of chloroaromatics 	  166
       Summary  	  170

    17. Overview of Microbiological Decomposition of
         Chlorinated Aromatic Compounds	   171

References	  176
Bibliography  	  207
Appendix: Illustrated list of compounds 	  234
Glossary  	  252
Citation Index	  261
Organism Index  	  265
Subject Index  	  267
                                  vn

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                               FIGURES

Number                                                              Pase

     1     Cellular organization  	    °
     2     Cyanobacterial vegetative cell  	    1
     3     Cell wall structures	    8
     4     Structure of membranes	    9
     5     Bacterial growth curve 	   11
     6     Regulation of enzyme synthesis  	   15
     7     Tricarboxylic acid cycle 	   17
     8     The 3-Ketoadipic acid pathway in bacteria and fungi	   18
     9     Hydrogen bonding of adenine with thymine and guanine with
          cytosine 	   21
    10     The DNA double helix	.'	   22
    11     Synthesis of nucleic acids and proteins	   24
    12     Transfer and recombination of genes during bacterial
          conjugation  	   27
    13     Genetic recombination during viral transduction of bacterial
          genes into a recipient cell 	   28
    14     Integration of materials balance and mineralization
          approaches in biodegradation assessment  	   40
    15     Common names and conventional nomenclature for substituted
          benzenes  	   46
    16     Oxidation of aromatic molecules by bacteria	   48
    17     Pathways for the bacterial metabolism of toluene  	   49
    18     Pathways for the metabolism of benzoic acid   	    50
    19     Metabolism of biphenyl by P. putida and Beijerinckia sp	   52
    20     Mechanism of  bacterial attack on naphthalene, anthracene,
          and phenanthrene	   53
    21     Pathway of phenanthrene metabolism by Pseudomonas sp	   54
    22     Pathway of naphthalene metabolism by Pseudomonas spp	   55
    23     Pathway of anthracene metabolism by Pseudomonas spp	   56
    24     Pathways of naphthalene metabolism  by Oscillatoria sp.,
          strain JCM	   58
    25     Ortho- and mete-cleavage pathways of catechol metabolism
          by bacteria  	   59
    26     Ortho- and meta-cleavage pathways of protocatechuic acid
          metabolism by bacteria  	   60
    27     Pathways of gentisic acid and  homogentisic acid
          metabolism by bacteria  	    62
    28     Divergent pathways for the metabolism of benzoic acid,
          p-hydroxybenzoic acid, and m-hydroxybenzoic acid by
          P. testosteroni and P. acidovorans	    63

                                                               (continued)

                                  viii

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Number                 FIGURES (continued)                    Page

   29    Formation of catechol from benzene in fungi, yeasts,
         and mammals	  64
   30    Pathway of naphthalene metabolism by C. elegans  	  65
   31    Metabolism of aromatic compounds by T. cutaneum	  66
   32    Formation of 3,5-dichlorocatechol from 2,3,6-trichloro-
         benzoic acid by Brevibacterium sp	  70
   33    Pathways of metabolism of chlorobenzoates by
         Pseudomonas sp. WR912	  72
   34    Metabolism of 3-chlorobenzoic acid by Pseudomonas sp. B13 ...  73
   35    Metabolism of 4-chlorobenzoic acid and 3,5-dichloro-
         benzoic acid by Pseudomonas sp. B13 transconjugants  	  75
   36    Metabolism of 5-chlorosalicylic acid by B. brevis	  76
   37    Representative pathway for the reductive dechlorination
         of chlorobenzoic acids by anaerobic microbial consortia  	  79
   38    Primary metabolic reductive dechlorination of gamma-
         hexachlorocyclohexane by anaerobic microorganisms 	  79
   39    Pathway of chlorobenzene mineralization by
         bacterial strain WR1306	  82
   40    Pathways of metabolism of 4-chloronitrobenzene by
         Rhodosporidium sp	  84
   41    Metabolism of pentachloronitrobenzene by F. oxysporum
         and 2,4-dichloro-l -nitrobenzene by M.javanicus AHU6010  ....  85
   42    Methylation of chlorophenols by Arthrobacter spp	  89
   43    Cometabolism of chlorocatechols via catechol 1,6-oxygenase
         by resting cell suspension of Achromobacter sp	  90
   44    Action of aromatic and chloroaromatic enzymes from
         P. putida B13 and P. putida derivative strains  	  92
   45    Proposed pathway for pentachlorophenol (PCP) metabolism
         by the bacterial culture KC-3 and by Pseudomonas sp	  95
   46    Bacterial metabolism of 2,4-D  	  102
   47    Metabolism of 4-chlorocatechol by Arthrobacter sp	103
   48    Pathway of MCPA metabolism by Pseudomonas sp.
         NCIB 9340	  105
   49    Pathway of 4-chlorophenoxyacetic acid metabolism by
         a soil pseudomonad  	  107
   50    Pathway for the metabolism of 4-(2,4-dichlorophenoxy)butyric
         acid by Flavobacterium sp	  108
   51    Metabolism of 2,4-D and MCPA by A. niger  	  110
   52    Pathways of 3,4-dichloroaniline metabolism by P. putida  	116
   53    Metabolism of 4-chloroaniline by microorganisms	  119
   54    Metabolism of solan by microorganisms  	  124
   55    Metabolism of chlordimeform in soils  	  127
   56    Microbial degradation of 4-chlorobiphenyl  	  131
   57    Pathway of 2,4,4'-trichlorobiphenyl metabolism by
         Acinetobacter sp. P6  	134
   58    Reductive dechlorination of DDT and dehydrochlorination
         of DDT 	                                139
                                                              (continued)
                                  IX

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Number                 FIGURES (continued)                     Page

    59    Metabolism of DDT by bacteria  	   141
    60    Oxidation of dibenzo-p-dioxin by Pseudomonas sp.
         NCIB 9816 	   147
    61    Oxidation of dibenzofuran by Beijerinckia sp.
         and C. elegans  	   150
    62    Chlorinated aromatic compounds metabolized to
         chlorophenols   	   172
    63    Chlorinated aromatic compounds metabolized to
         chloroanilines   	   173
    64    Metabolism of chlorocatechols, chlorobenzoic acids, and
         chlorosalicylic acids	   174

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                                TABLES

Number                                                               Page

     I    Recognition sites for restriction endonucleases	   29
     2    Materials balance and mineralization approaches to
          biodegradation assessment	   39
     3    Comparison of metabolites formed by eukaryotes and
          mammalian systems  	   68
     4    Microorganisms that metabolize phenoxy acids	  100
     5    Metabolism of chlorinated biphenyl compounds by
          Alcaligenes sp. Y42 and Acinetobacter sp	  132
     6    Chemical factors influencing organic biodegradability	  153
     7    Biological factors influencing organic biodegradability  	  153
     8    Experimental factors influencing organic biodegradability	  154
     9    The effect of pentachlorophenol purity on disappearance
          in continuous systems	  155
    10    Disappearance of commercial chlorobiphenyl mixtures  	  157
    11    Distribution of Kaneclor 500 in activated sludge semi-
          continuous systems  	  158
    12    Distribution of 2,3',4,5'-tetrachlorobiphenyl in a sediment-
          water-air-model system  	  160
    13    Distribution of 2,2',4,4',5,5'-hexachlorobiphenyl in sediment-
          water-air-model system  	  160
    14    Distribution of 2,2',3,3',4,4',5,6'-octachlorobiphenyl in a
          sediment-water-air model system	  161
    15    Fate estimates of 2,4-dichlorophenol removal from a lab activated
          sludge system using proposed equations	  164
    16    Fate of trichlorocarbanilide in lab-scale activated sludge
          systems	  165
    17    Classes of chloroaromatics studied in several experimental
          studies  	  166
    18    Biokinetic results from 2,4-D and chlorobenzoic acids	  168
    19    Fate of several chloroaromatics in a lab-scale activated sludge
          system  	  168
    20    Mass removal of chloroaromatic  compounds in a full-scale
          wastewater treatment plant  	  169
                                XI

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              ABBREVIATIONS AND SYMBOLS
ABBREVIATIONS

BOD         — biochemical oxygen demand
COD         — chemical oxygen demand
Hc            — Henry's law constant
HRT         — hydraulic residence time
MCRT        — mean cell residence time
MLSS        — mixed liquor suspended solids
MLVSS       — mixed liquor volatile suspended solids
ppb           — parts per billion (jug/ 1)
ppm          — parts per million (//g/ ml or mg/1)

 Abbreviations of herbicides are noted at first mention and in index.
SYMBOLS

a
b
Ca
C|
Cs
fL
Kb

Kf
Kow
Ks
Ksl
n
Qair/ V
REMb

REMe
REMS

REMst

X
Y
 m
              — empirical constant in Langmuir isotherm
              — empirical constant in Langmuir isotherm
              — concentration of substrate in the mixed liquor
              — concentration of substrate in the liquid phase
              — concentration of substrate in the solid phase
              — weight fraction of lipid-like compounds in the biomass
              — a biological disappearance rate constant, first order in substrate
                concentration
              — empirical constant for the Freundlich isotherm equation
              — octanol-water partition coefficient
              — Monod half-saturation constant
              — first order stripping rate constant
              — empirical constant in Freundlich isotherm equation
              — ratio of the air flow rate to the hydraulic reactor volume
              — percent removal of the substrate from the system by the
                biotransformation fate mechanism
              — percent removal of the substrate from the system in the effluent
              — percent removal of the substrate from the system by sorption on
                biomass
              — percent removal of the substrate from the system  by the
                stripping fate mechanism
              — concentration of biomass in the mixed liquor
              — yield coefficient
              — specific growth rate
              — maximum specific growth rate

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                      ACKNOWLEDGMENTS

   The authors would like to thank  Drs.  P.R.  Sferra and John A. Glaser, the
Technical Program Monitors (EPA, HWERL, Cincinnati), for their helpful support
and assistance throughout this project. We would also like to acknowledge Mr.
David R. Watkins of the same organization for his considerable help in the planning
and inception phases of this project.
  Mrs. Kim Truong of IT Corporation provided valuable assistance in reviewing the
manuscript and assisting  in the production of  this report.  Her  contribution  is
gratefully appreciated.
  Finally, Dr. David T. Gibson, Director of the Center for Applied Microbiology,
University of Texas at Austin, Dr. John C. Loper,  Department of Microbiology and
Molecular Genetics, University of Cincinnati College of Medicine, and Dr. Christen
J. Hurst, Health Effects Research Laboratory, U.S.E.P.A., Cincinnati, reviewed this
document and offered technical perspective on the  contents of this work. The authors
believe these comments improve this work and are indebted to Drs. Gibson, Loper,
and Hurst for their cooperation.
                                    xiu

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                              SECTION 1

                          INTRODUCTION

  The first synthesized organochloride compound, ethyl chloride, was prepared in
about 1440, but large scale synthesis of industrially important chlorinated organics,
including chlorinated aromatic compounds, occurred during only the past few
decades (214a). In general, the chlorinated  aromatics of industrial synthesis  or
byproducts thereof represent one class of xenobiotic recalcitrant compounds. These
compounds have few or no naturally occurring structural analogs and are persistent
or resistant to both biological and abiotic degradation. Many of the chlorinated
aromatics share similar physical chemical properties of low water solubility and high
Kow (octanol-water partition coefficients) which suggest lipophilicity or bioaccumu-
lation potential.
  Properties such as persistence, bioaccumulation, and demonstrable chronic and
acute toxicity to human and nonhuman animal populations cause immediate
concern  related to their environmental  health effects  and their potential  for
ecosystem perturbations on long- and short-term exposure. These concerns cause the
frequent appearance of these chemicals on EPA priority pollutant lists and have led
to extensive research on their fate in the environment and their potential  for
microbiological transformation to less hazardous molecules.
  The term biodegradation has had many different meanings. The Biodegradation
Task Force, Safety of Chemicals Committee,  Brussels (299) has  defined biode-
gradation as the molecular degradation of an organic substance resulting from the
complex action of living organisms. A  substance is said to be biodegraded to  an
environmentally acceptable extent when environmentally undesirable properties are
lost.  Loss of some characteristic function or property  of the substance  by
biodegradation may be referred to as biological transformation. In this text, we have
attempted to restrict use of the term biodegradation in favor  of more specific
terminology. In this respect, biotransformation refers to any alteration of an organic
molecule by organisms, and mineralization means the transformation of an organic
molecule to its inorganic component  parts with release of halide, CO2, and/or
methane.
  The potential for microbial transformation of chlorinated aromatic compounds is
related to the two fundamental roles of heterotrophic microorganisms in the global
ecosystem. Both roles relate to the central concept of microbial decomposition of
organic matter to release stored energy in the organic molecules (whether natural or
anthropogenic in  origin)  and to return essential  nutrients, such  as CO2,  to
biogeospheric nutrient pools. The first has thermodynamic implications while the
second relates to elemental and nutrient cycling. While these simplified generaliza-
tions apply to most naturally produced  organic matter, certain aromatic polymers
such as lignin are persistent in the environment.  Factors contributing to persistence
of organic compounds have been discussed previously (92a) and can be summarized
as insolubility, large molecular size or polymeric nature, toxicity, and anthropogenic
origin. Although most or all of these factors are important for various chlorinated
aromatics, the anthropogenic origin of most chlorinated aromatics is critical for the

                                     1

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prediction of the rate of elimination of the compound from the environment and its
eventual fate.
   In general, microbial decomposition or biodegradative capabilities have coeyolved
with the synthesis of organic matter by plants  and animals over the millema.
However, in the case of many chlorinated organics of industrial synthesis, 40 to 50
years is an unlikely time frame to expect evolution of enzyme systems capable of
decomposition of such compounds. Yet evidence has accumulated that some of these
compounds can be biologically  transformed and extensively biodegraded by a
diversity of heterotrophic microorganisms.
   Such evidence has arisen primarily from studies using pure and mixed microbial
cultures and from lesser-controlled environmental fate experiments. This evidence
has promoted much additional research on the molecular mechanism of biodegra-
dation, which in turn has permitted studies leading to increased knowledge of more
detailed aspects of biodegradation itself. Currently, major questions exist as to the
rate at which biodegradation occurs in various environments and the potential for
kinetic  prediction of  pollutant fate  based  on  laboratory and  environmental
observations. Research needs directed at this major question have caused a renewed
focus on individual microbial populations that may be specifically responsible for
biodegradation of a narrow spectrum of chlorinated aromatic substrates, and  on
physical/chemical  environmental parameters that may modulate both the  popu-
lation and their catabolic activity. Developments in these areas have led to the
relatively recent detection  of bacterial strains harboring extrachromosomal DNA
(plasmids) that genetically encodes enzymes which mediate the biodegradation of
specific groups of aromatic and chlorinated aromatic compounds. Coupled with the
availability of new molecular genetic techniques, such as DNA probe technology, it
has become feasible to plan research to examine  catabolic gene maintenance and
transfer in natural populations and to detect specific biodegradative microorganisms
in the environment. Such development will lead to greater predictive capabilities on
the long-term persistence  of selected chlorinated aromatic pollutants, and will
provide for insight in the evolution and reassortment of genes responsible for
biodegradation. With these genetic techniques there is  a potential therefore to
enhance biodegradative capacity among natural populations.
   This  report begins  with three sections that provide an overview of microbial
physiology, genetic information transfer and processing, technologies for gene
manipulation, and methods of biodegradation assessment. These principles are
applicable to microorganisms in general and are specifically relevant to  metabolism
of chlorinated  aromatic  compounds.  Readers  wishing review of  these basic
microbiological concepts should read these chapters before turning to subsequent
sections.
  The next section includes a review of pathways of metabolism of non-chlorinated
aromatic compounds from which the chlorinated  pollutants are  derived. The
microbial metabolism of most of these compounds has been studied extensively and
these data form the basis for an understanding of the biotransformation pathways of
the more complex chlorinated aromatic molecules.
  The following nine sections discuss the microbial metabolism of the chlorobenzoic
acids, chlorobenzenes, chlorophenols, pentachlorophenol, chlorophenoxy and
chlorophenyl herbicides, chlorinated biphenyls, DDT and related compounds, and
chlorinated dioxins and dibenzofurans. These compound classes represent all of the
major classes of chlorinated  aromatic molecules with environmental pollution
potential.
  Each chapter details the information available on metabolism of these compounds
by pure cultures or consortia of bacteria, cyanobacteria, and fungi. Emphasis has
been placed on studies in which metabolites have been identified and, where possible

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complete or partial pathways have been reported. Studies reporting evolution of CO2
and chloride release have been reviewed and are discussed as well. The disappearance
of these pollutants in soils, water, sewage, and other environments has also been
noted, although the body of literature relating to herbicide disappearance is so
extensive that it has only been summarized and has not been reviewed as completely
as the other topics. The fate of these compounds in scaled-up biological treatment
processes is the subject of the next section.
  The final chapter summarizes the information reviewed here and draws together
the pathways of metabolism of chlorinated aromatic molecules into an overview
diagram indicating the potential for biodegradation of a compound under optimum
conditions. This is an idealized representation, as few  environmental situations
would comprise all the factors  necessary for  complete biotransformation and/or
mineralization of these recalcitrant compounds.
   This report includes a list of citations and a bibliography of additional references,
including several excellent  reviews on the biodegradation of various classes of
organic compounds. All of the compounds noted in this report, including metabo-
lites, are registered in an illustrated alphabetical list in the appendix. A citation index
and a subject index are also included. Finally, a glossary of many of the scientific
terms that appear in the text is appended in order to aid the reader.
  This document is intended to be a general reference for environmental decision-
makers who are interested in the fate of chlorinated aromatic compounds with
respect to microbial activity. It is also meant to provide a resource for scientists and
engineers involved in environmental predictive fate assessments. In addition, this
review is designed to be a continuing resource for environmental microbiologists
interested in the areas of metabolic pathways of chlorinated aromatic compound
dissimilation and biodegradative fate of these potential pollutants.
  The organization of this document into specific chapters and associated review
sections is intended to facilitate its use by these diverse groups of people. This is an
extensive although not exhaustive review of the literature pertaining to biodegra-
dative fate  of these classes of chlorinated aromatic compounds.

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                               SECTION 2
                            CONCLUSIONS

 1.   The biodegradability potential of a given compound depends on many factors.
     Chemical determinants include the ionic state of the compound, the number,
     types, and position of substituents, and  the  general form  of the molecule.
     Environmental parameters affecting microbial metabolism include pH, tempera-
     ture, redox state, moisture, reactor configuration, kinetics, and system turbu-
     lence  considerations, and  interference by competitive processes such  as
     sorption, stripping, and photodegradation. Other factors involved in microbial
     growth and metabolism include availability of nutrients and growth factors,
     concentration of substrate, competitive interference by other substrates, and
     formation of toxic metabolites.

 2.   Many microorganisms require a period of acclimation before biodegradation
     occurs. Once a population is acclimated, it sometimes retains its predisposition
     to metabolize the substrate, and subsequent additions of substrate are metabo-
     lized after a shorter  lag or no lag period.

 3.    Some compounds are mineralized by consortia of microorganisms in cases
     where no single  species  has been shown to  be capable of that process.

4.    Fungi and bacteria metabolize most compounds  by different  biochemical
     pathways.

 5.    In general, chlorophenoxy herbicides and chlorobenzenes can be metabolized to
     chlorophenols and then to chlorocatechols. Phenylamide herbicides and other
     compounds  with  nitrogen-containing substituents are decomposed to chloro-
     anilines, which can be oxidized to chlorocatechols and a variety  of products.
     Chlorocatechols may be  converted by  a variety of different mechanisms to
     nonchlorinated ring cleavage products. Chlorobenzoic acids may be trans-
     formed by three  different routes  to ring cleavage products: (1) through a
     substituted catechol,  (2)  through chlorosalicylic acid, or (3) anaerobically
     through benzoic  acid to  methane and carbon  dioxide.

6.    The above conclusions represent general pathways for substrate dissimilation.
     Prediction of the biodegradability of a particular compound on which no data
     exists, based on data about similar types of compounds, can be loosely made.
     For instance, the degree of chlorination  affects  the rate  and extent of
     metabolism  of PCBs  and other compounds.

7.    Experiments in the laboratory or in a particular environment cannot be readily
     extrapolated  to  other environments, because of the need  to consider the
     parameters affecting the biodegradability potential listed above.

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8.    Small-scale lab tests are essential in order to perform properly controlled
     experiments to assess the effects of many of the above-mentioned biodegrad-
     ation factors. Experimental design necessary to achieve the highly controlled
     results  desired in small-scale testing nearly always requires compromise of
     factors needed to  apply results to  larger-scale systems.

9.   Scale-up testing in larger-scale systems designed to simulate the  desired full-
     scale application (wastewater treatment system, land farming, environmental
     scenario, etc.) is required to collect data (often  kinetic data)  allowing the
     prediction of process performance and comparison among full-scale systems.

10.  Many of the available scaled-up studies focus on substrate disappearance or
     removal from the given feedstream without quantification of abiotic fates or
     adequate regard for calculation of biokinetic rate  constants.

11.  Additional  emphasis  should be given to establishing consistent  scale-up
     methodology and to implementing additional work for generation of reliable
     scale-up data on the biological treatment of chlorinated aromatic compounds.

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                              SECTION 3
          OVERVIEW OF MICROBIAL PHYSIOLOGY

  A study of the specific pathway of degradation of a compound by an organism
necessarily is limited to specific biochemical reactants,  products, and reactions
occurring within the cell. Much of the content of this document is concerned with just
such features. When genetics of the pathways are discussed, the relationship to the
total cell is even  more remote. Therefore,  it is important to begin with a firm
understanding of the physiology of the microbial cell, its structure, requirements for
growth and survival, and relationship to its environment. While  the cell is often
described as a microscopic biochemical reactor, the activities of the cell are intimately
connected to and shaped by its external environment. Information presented in this
chapter is based on several general references, which may be consulted for further
details (4, 107, 276).
  The primary physical difference between bacteria and fungi is the presence of a
membrane surrounding the  DNA  material. The enclosed structure is  called the
nucleus. Cells containing a membrane-bound nucleus are referred  to as eukaryotic
cells. The DNA of bacteria and cyanobacteria is contained in a diffuse region without
a surrounding membrane called the nucleoid and these forms are called prokaryotes
(Figure 1). Although it is tempting to consider bacteria as primitive compared to
eukaryotes, the  complexity of their biochemical reactions, and their regulatory and
adaptive  mechanisms, preclude such a label.
 Nucleolus
                                   Chromosomes

                                   Nuclear
                                   membrane
                                   Mitochondrion
                                   Endoplasmic
                                   reticulum
                                           Vacuole

                                        Mesosomes

                                         Ribosomes
                                         Inclusions
Chromosome
  (nucleoid)
Cell wall
Plasma
membrane
      Figure 1.   Cellular organization. A - Typical eukaryotic animal cell. B - Typical
                        prokaryotic rod-shaped bacterium.
                                 Reference 4.

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MICROBIAL CELL STRUCTURE
  Most bacterial cells average about 1 to 2 microns in length and are rod shaped.
Among all bacteria, however, cell size ranges from one tenth to 100 times the average
bacteria size. Although most  of the bacteria found  in the environment are rod
shaped, some water isolates are shaped like commas or spirals, and many bacteria,
especially pathogenic species, are spherical (cocci). Other bacteria take unique shapes
and forms as well.
  The  shape of the  cell  is conferred by a rigid cell wall composed mostly  of
peptidoglycan, lipid, lipopolysaccharide, and protein (Figure 2). There are charged
polymers within the cell wall which assist in the uptake of ions and some nutrients.
The wall also acts as a molecular sieve which prevents entry of some large molecules
and prohibits loss of proteins,  i.e.,  enzymes, from within the cell.
                Cyanophycin
                    granule
              Cell membrane
               Carboxysome

              Polyphosphate
                     granule

                   Thylakoid

                Nucleoplasm

             70S Ribosomes
                    Glycogen
                    granules

                  Gas vesicle
 Figure 2.  Schematic diagram of cyanobacterial vegetative cell. (Insert) Enlarged view
 of cell envelope, showing  outer membrane and peptidoglycan wall layers and cell
 membrane.
                             Reference unknown.

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  Bacteria and cyanobacteria can be divided into two groups based on cell wall
structure and composition. Classically the bacteria have been differentiated into
Gram positive and Gram negative groups according to a staining procedure called
the Gram stain. Electron microsopic techniques have shown differences in the form
of the cell walls between the two types of bacteria. The Gram positive cell wall is
composed of a single dense layer of peptidoglycan. Embedded in the peptidoglycan
matrix are polysaccharides and teichoic acids. The cell wall is closely associated with
the cytoplasmic membrane which has a double-track appearance with a central
transparent layer. The Gram negative cell wall is more complex. The outermost layer
is a wavy, double-track membrane which differs in chemical composition and in
function from  the cytoplasmic membrane. This layer is composed of lipopoly-
saccharides, phospholipids, and proteins.  Internal to the outer membrane is a thin
rigid layer of peptidoglycan. Between the cytoplasmic membrane and the outer cell
wall membrane lies the periplasmic space containing enzymes (Figure 3). In contrast,
the fungi have cell walls composed mainly of polysaccharides. The particular types of
polysaccharides are characteristic of the taxonomic group of the fungi.
               Gram positive
               Gram negative
                                 peptidoglycan
                                 and teichoic acid
                                cytoplasmic membrane
                                              Slipopolysaccharide
                                              lipoprotein
                                              lipid, etc.
                                peptidoglycan
                                cytoplasmic membrane
        Figure 3.  Cell wall structures seen in thin-section electron microscopy.
   A - Diagrammatic representation of the Gram-positive wall; B - of the Gram-negative
                 wall. The location of wall components is indicated.
                               Reference 107.

  Some bacterial cells are motile, and of these the most common mechanism is by use
of flagella, hairlike helical structures several times the length of the cell. Some genera
possess only 1  or 2 flagella, while in other genera the flagella are present over the
entire cell surface. The flagella rotate to propel  the cell through the water. Some
bacteria, including some cyanobacteria, move in  a characteristic gliding motion by
flexing the cell wall against a surface in a manner similar to inchworrn movement.
  Some bacteria have the ability to attach to solid surfaces. In a few genera this may
be accomplished by hairlike pili or by structures  called holdfasts. In most bacteria
attachment occurs by a capsule or slime layer composed of organic polymers, mostly
polysaccharides. After initial contact with the surface, the cell synthesizes polymers
which bridge the gap and attach firmly to the surface. It then may become impossible
to remove the cell without destroying it. When  the cell divides, the nonattached
portion can move, but the new cell arising from the attached portion of the cell
remains in place.

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  Other functions of the capsule or slime layer include protection of the cell from
such conditions as desiccation.  In many pathogenic bacteria, the presence of the
capsule affords protection against white blood cells and antibodies. The capsule
seems also to serve as storage sites for excess nutrients or wastes.
  In freshwater environments, the cell contains a higher concentration of salts than
the  surrounding medium. The cell would expand and lyse without the protection
against osmotic shock afforded by the cell wall together with the cell membrane. The
cytoplasmic membrane is internal to the cell wall and has additional functions
(Figure 4). It is selectively permeable and often facilitates movement of a substrate
into or out of the cell against a concentration gradient. For other substrates transport
is almost  completely prevented. The rate of transport can be  specific for  the
particular substrate, and  two substrates very closely related structurally can have
very different transport rates. Some substances enter or leave by passive diffusion.
The cytoplasmic  membrane also maintains  the osmotic gradient, is the site of
enzymes involved  with cell wall synthesis, and is the site of oxidative metabolism and
energy conversions. More complex constructions of the cytoplasmic membranes are
found in specialized groups of bacteria such as the cyanobacteria and the methane-
utilizing bacteria. The cyanobacteria contain internal membrane structures called
thylakoids which  contain the photosynthetic apparatus.

                                   Proteins
           Hydrophilic  groups
           on phospholipid
Lipid bilayer
      Figure 4.  Structure of membranes, a diagrammatic representation. The lipid
                    molecules are probably in constant motion.
                               Reference 107.

  In fungi, cell growth occurs only at the tip of the hypha, and the plasma membrane
below the tip contains a large number of membrane-bound vesicles which may hold
the enzymes and cell wall precursors needed for cell growth. In addition, the fungal
plasma  membrane  is involved  with  osmotic regulation and nutrient  uptake.
Functions such as oxidative metabolism are reserved to certain membrane-bound
organelles which are absent in bacteria.
  All cells contain chromosomal DN A; in bacteria it is circular and double stranded,
resembling a helical ladder. Bacteria also contain extrachromosomal DNA called
plasmids which code for  auxiliary functions in the cell, such  as  resistance to
antibiotics and  heavy metals and ability to metabolize some organic compounds.
Fungi contain a number of linear chromosomes. The DNA contains the code which
guides the structure and metabolism of the cell. Specific features in the functioning of
DNA will be discussed in a later section.

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  Eukaryotic cells contain mitochondria, which are organelles bounded by a double
membrane. These function in ATP generation and the oxidative metabolism of
substrates, activities carried out in bacteria by the cytoplasmic membrane. There are
other specialized structures within some bacterial or fungal cells which function in
storage of excess nutrients or gaseous products. The Gram-positive genera Bacillus
and Clostridium form spores when exposed to unfavorable conditions. The spores
are extremely resistant  to heat, desiccation, radiation,  acids, and chemical disin-
fectants,  yet when exposed to favorable conditions will germinate and form a
vegetative cell within hours.

GROWTH REQUIREMENTS
  Bacteria of one type or another have been found in all environments and under all
conditions with the possible 'exception of pure vacuum.  A specific bacterial  species
may grow  under a wide variety of conditions or it  may have very exacting
requirements for cell growth.
  Certain nutrients are required by all cells. Carbon is most important and those cells
that obtain it from organic substrates are referred to as heterotrophs. Autotrophs can
fix carbon from carbon dioxide. A few specialized groups can utilize other substrates;
methylotrophs, for example, can oxidize methane at aerobic/anaerobic interfaces.
Other essential nutrients include phosphorus, usually derived from phosphates, and
nitrogen, usually obtained from nitrate or ammonia. These three elements are the
most common nutrients that limit growth. Other necessary growth factors include
sulfur, magnesium, potassium, calcium, and other metallic elements. While some
bacteria synthesize all their required vitamins and growth factors, other bacteria
must obtain some from the environment. Water is also a specific requirement in
cellular metabolism. The bacterial cell is composed of about 80% water, and water is
both the solvent  and a specific cofactor in many biochemical reactions.
  An important physical parameter for growth is temperature.  An increase in
temperature may inactivate enzymes or may be lethal to the cells, while a decrease in
temperature may simply inhibit growth. Cellular enzyme activity is also governed
partially by the ambient temperature. Upon warming, the cells may resume normal
cellular  function. An individual microbial species usually has  a minimum, an
optimum, and  a maximum temperature for growth.  Those  that grow best at
temperatures below 20°C are  called psychrophiles.  Mesophiles grow from about
15°C to about 45°C, and most bacteria are grouped into this category. Thermophiles
grow at temperatures above 50°C. These names permit categorization of a situation
which in reality  represents a gradation of microbial tolerances for temperatures
ranging from the arctic  environment to thermal springs.
  The oxygen requirements of microorganisms vary considerably. Obligate aerobes
grow only in the presence of air and use aerobic respiration to obtain energy. Obligate
anaerobes grow only in  the absence of air. The sensitivity of anaerobes to molecular
oxygen is due  to lack  of enzymes which render the superoxide free radical ion
harmless through reduction. Facultative anaerobes will grow  in the presence or
absence  of air using alternate chemical electron acceptors such as O  or NO .
Microaerophiles have a narrow range of tolerance for their gaseous environment and
require a reduced air environment or in some cases an increased proportion of carbon
dioxide.
  Bacteria also respond to changes in pH of the medium. Most bacteria (neutro-
philes) grow best at  neutral  pH  (pH 7).  However,  acid-producing bacteria
(acidophiles) grow very well at lower pH values and strains adapted to alkaline
environments (basophiles) grow at pH of 8 or 9. Fungi often tolerate extremes of pH
better than bacteria. Halophilic bacteria require high salt concentrations, while other
species are salt-tolerant although high salt concentrations are not mandatory for

                                    10

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survival. Organisms have been isolated from the deep ocean. Some strains survive
both at the deep  ocean pressure and at atmospheric  pressure and are  called
barotolerant. Other deep ocean strains have only been kept alive by bringing them to
the surface in pressurized vessels, and these bacteria are said to be barophilic. Many
common strains of bacteria found at atmospheric pressure are killed when placed in a
high-pressure environment.

THE CELL GROWTH CYCLE

  The bacterial cell normally grows until it reaches a certain size, at which time it
divides by binary fission into two identical  daughter cells. During this time the
bacterial DN A replicates and is partitioned to opposite sides of the growing cell. The
cell wall and cytoplasmic membrane divide the cell in 1 of 2 ways depending on the
particular genus.
  In some genera the elongated cell pinches in equatorially until 2 cells are formed. In
other genera a double cytoplasmic membrane is formed  in the middle of the cell
followed by synthesis of a double cell wall. When wall construction is complete the 2
daughter cells separate. Not all cells divide in synchrony,  so in a culture of cells all
stages of the growth cycle are represented.
  Fungi grow by elongation at the tip of the  hypha. Many move into a yeast stage
during which the cell divides by budding. An outgrowth appears at some point on the
cell surface and  grows until it is  almost the size of the mother cell. The cellular
organelles including DNA are replicated and a copy is partitioned into the daughter
cell which eventually is walled off. The mother cell does not increase greatly in size
during cell division.

POPULATION GROWTH
  Laboratory studies with pure cultures of bacteria traditionally have demonstrated
exponential growth in batch culture, in which all essential nutrients are present in
excess and growth parameters are optimal.  In this situation,  population growth
follows a characteristic cycle which begins with the lag phase of growth, during which
the cells are adapting to the new environment (Figure 5). Enzyme synthesis induced
    ffi
    a
    s»
    O
    3
    z
    O>
    O
                                 Time, Hours

Figure 5.  Bacterial growth curve. A, lag phase; B, logarithmic phase; C, stationary
              phase; D, decline phase; E, surviving population.
                                     11

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 by contact with a new substrate occurs during this phase. Cells that are preadapted to
 the substrate or the growth conditions experience a shortened lag phase or no lag
 phase at all. Following the lag phase is a period of unrestricted multiplication called
 the log or exponential growth phase,  so named because of the binary fission process
 of cell division. The rate of growth (cell division) depends upon the composition of
 the growth medium and the environmental parameters,  and under optimal condi-
 tions a cell may divide every 15 minutes. When a nutrient becomes limiting or when
 inhibitory or toxic products accumulate, the cell enters the stationary growth phase.
 The individual cell is still  viable although not replicating. This phase is manifested
 within the total culture by an equivalence between the number of cells produced by
 cell division and the number of cells dying; thus, there is no net change in the number
 of cells in the population.  When the number of cells dying becomes greater than the
 number of cells being formed, the death phase ensues. The population eventually
 stabilizes at a  constant  low number of surviving cells.  Because  this cycle is
 characterized by  an abundance of nutrients,  it is rarely  seen in  the  natural
 environment.
   In cultures where (1) cells are able to proliferate, (2) there is an absence of
 inhibitors, (3) there is a homogeneous mixture of cells and nutrients, and (4) the
 substrate is the limiting factor in growth, cellular growth  can be related  mathe-
 matically with the disappearance of substrate. In this case, Monod kinetics apply as
 in equation 1 :


                                                                          (1)

                           dX              CaX
                           dt
   The instantaneous change in cell concentration over time, dX/dt, is equal to the
 maximum specific growth rate, times a fraction including the substrate concentration
 in the mixed liquor, Ca, the cell concentration, X, and the Monod half-saturation
 constant, Ks.
   The half-saturation constant is equal to the substrate concentration at which the
-specific growth rate is one-half /um (in batch culture this situation occurs at the end of
 the experimental growth phase. Typically the constants /zm and Ks are determined in
 batch culture tests using linearized graphical plots of the reciprocals of the measured
 cellular growth and substrate concentrations. Alternatively, they may be determined
 in a series of continuous culture tests.

CONTINUOUS CULTURE

  Techniques for continuous cultures have  been  developed  to provide a constant
environment for microbial  growth. The physical factors of temperature,  pH, O2
concentration, etc., are well controlled, and nutrients can be  supplied at controlled
rates coupled with removal of potentially toxic waste materials. Population growth,
therefore, occurs at a constant rate which in some systems can be varied by changing
the availability of a  nutrient. Studies conducted in continuous systems such as the
chemostat  and the  recycling  fermentor have helped  to  establish  the  energy
requirements of cells under growth or maintenance (survival) conditions in addition
to exploring the response of cells to various types of nutrient or other growth factor
limitations. Maintenance of a cell population under steady-state conditions with
selective pressure, such as a nonutilizable carbon source, may enable mutants with

                                      12

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capability to metabolize the substrate to be generated and then grow to sufficient
population levels to be recovered.

CELL DEATH

  The most widespread measure of cell death is loss of reproductive capability.
However, the medium used to detect survivors may not be adequate to demonstrate
cell division, although the cell may be viable and capable of reproduction in another
environment. The problem of defining cell death has not yet been resolved.

PURE AND MIXED CULTURE METABOLISM

  Populations in which all the cells are of the same species are considered to be a pure
culture. Except for the ongoing process of mutation, discussed in a later section, the
process of cell reproduction by binary fission with replication of genetic information
ensures that the pure culture will express essentially the same properties. The
population will be nearly homogeneous in its ability to metabolize a substrate.
  In some cases, a substrate may be metabolized only partially by a particular species
and a product may accumulate. In a parallel situation, another species may be able to
metabolize that product further, although the second species may lack enzymes
needed to metabolize the parent substrate. By themselves, neither species could
mineralize the substrate of interest.  However, a mixed culture of the two organisms
might act in concert with one species, mineralizing  the product resulting from
metabolism of the substrate by the other species.  A  consortium of more than two
species may be required to mineralize a substrate and the effective species may be
bacteria, fungi, or a mixture of the two.
  Cometabolism refers to the fortuitous metabolism of a compound while the cell
obtains its carbon and energy from another source (273). Such metabolism may be
partial or complete and depends upon enzymes already active in the cell.

SUBSTRATE UPTAKE AND TRANSPORT
  Some motile bacteria possess the ability to move along the concentration gradient
of a specific compound in its environment. This phenomenon, called chemotaxis,
may permit these strains to scavenge some nutrients more efficiently or to move away
from toxic or inhibitory compounds.
  Some bacteria are able to utilize as carbon and energy sources substrates which are
too large to enter the cell. These strains secrete hydrolytic enzymes into the culture
medium which break the high molecular weight compounds (such as proteins, starch,
or cellulose) into smaller components which can enter the cell.
  The transport of other substrates (lower molecular weight) into cells depends on a
number of interrelated factors. The substrate must  be able to pass the complex
cytoplasmic  membrane which is composed of a hydrophobic zone  surrounded on
both sides by hydrophilic layers. Some lipid-soluble substances can pass across this
zone by free diffusion which is dependent on the difference in substrate concentration
inside and outside of the membrane. The rate  of uptake by this mechanism,  called
passive transport, depends upon the size and charge of the substrate.
  Within the cytoplasmic membranes are proteins which couple substrate transport
to an energy-yielding process. Called active transport, this mechanism is the route of
entry for most substrates and ions, and the concentration within the cell protoplasm
can be much greater than the concentration outside the cell. The proteins involved in
active transport can be very specific for a particular substrate to the exclusion of
structurally related analogs.
                                    13

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ENZYMES
  Enzymes are proteins which are the most efficient known catalysts for biochemical
processes. They serve to increase the rate of a reaction, often causing a reaction to
occur under physiological  conditions which  otherwise could  only occur under
extremes of pH, temperature, or concentration.
  Classically, enzymes were grouped into  9 categories based on function. These
names are still used frequently. (1) Dehydrogenases mediate the  loss of a hydrogen
ion from a substrate  with  the acceptor being other than molecular oxygen.  (2)
Oxidases catalyze loss of a hydrogen ion with molecular oxygen as the acceptor. (3)
Kinases transfer a phosphate group from ATP or other nucleoside triphosphate to
the substrate. (4) Phosphatases mediate the hydrolytic cleavage of phosphate esters.
(5) Mutases catalyze transfer of a functional group between two positions in the same
molecule. (6) Synthetases mediate condensation of two separate molecules coupled
with cleavage of ATP. (7) Decarboxylases achieve decarboxylation of the substrate.
(8) Thiokinases catalyze the ATP-dependent formation of thiol esters. (9) Carbo-
xylases catalyze the ATP-dependerit addition of carbon dioxide to the acceptor
substrate.
  These categories have been replaced by 6 classes of enzymes in a formal system
developed by the International Enzyme Commission (39). (1) Oxidoreductases act on
the CH-OH group of a substrate, requiring NAD+ or NADP+ as the  hydrogen
acceptor. This category includes  dehydrogenases and oxidases.  (2) Transferases
catalyze the transfer of an  intact  group of atoms, such as methyl or phosphorus
containing groups from a donor to an acceptor molecule. Kinases and mutases are
included in this group. (3) Hydrolases, including phosphatases, mediate the transfer
of chemical groups to water. (4) Lyases, such as decarboxylases, catalyze the addition
of groups to substrates containing double  bonds, or the removal of groups from
substrates to yield products with double bonds. (5) Isomerases catalyze a change in
the atomic configuration of a molecule without a change in the number or kind of
atoms. (6) Ligases are involved in the formation of a product  resulting from the
condensation of 2 different molecules coupled with the breaking of a pyrophosphate
linkage in ATP. This class includes synthetases, thiokinases and carboxylases.
  Most enzymes are notably specific in their actions, catalyzing the reaction of a
particular substrate, but having no activity against a very closely structurally related
substrate. Some enzymes, however, act on many related compounds. These enzymes
act on a specific structural component of different substrates. Enzyme specificity is
related to two features of the substrate. First is the specific chemical structure which
is attacked by the enzyme. Second, the substrate must also contain a binding group
which binds to the enzyme in such a way as to permit optimal association of the
susceptible structure with the enzyme. The active site on the enzyme is the area
containing both the binding site and the catalytic site, and the  three-dimensional
configuration of the complex resembles a "lock and key" relationship.
  The activity of enzymes can  be inhibited either  irreversibly  or reversibly.
Irreversible inhibitors destroy or bind to a functional group on an enzyme which is
necessary for its catalytic activity. Reversible inhibitors may be either competitive or
noncompetitive.  A competitive inhibitor has a similar structure to that of the
substrate and therefore can be bound by the enzyme. However, the enzyme has no
activity against the inhibitor. Since the inhibitor and the substrate compete for the
binding sites of enzymes, the  action of a  competitive inhibitor can  be partially
reversed by increasing the concentration of substrate. Noncompetitive  inhibitors
bind to the enzyme in an area other than the binding site, and in so doing alter the
catalytic site so as to make it inactive. The affected site is often called the regulatory
site of the enzyme and is reversibly occupied by the inhibitor. Lowered concen-
trations of the inhibitor increase the activity of the enzyme. When the inhibitor is a

                                    14

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direct product of a series of reactions involving the enzyme, the regulatory process is
called feedback inhibition and is an important cellular mechanism for regulating
metabolic processes such  that energy is not wasted on production of unnecessary
metabolites.  Enzymes with a regulatory site,  called allosteric  enzymes, can be
stimulated as well  as inhibited  by specific effector molecules which bind to the
regulatory site. The effector molecule may be the substrate itself, signalling the
enzyme to initiate the metabolic pathway. Often only one key enzyme in a pathway is
regulated; the activity of the rest of the enzymes is limited by the availability of their
specific substrate.
  Enzymes may also be controlled at the level of enzyme synthesis. A reduction in the
amount of enzyme would reduce the total enzymic activity. The genetic system for
synthesis of enzymes consists of several parts (Figure 6). This general model shows
several structural genes which code for the enzyme proteins. More than one enzyme
may be part of a system. In addition, each system contains one control gene coding
  Regulatory Gene
                  Control Region
                                               Structural Genes
                Promoter   Operator
                 Site   i   Site
                                                                         mRNA
                    Figure 6.  Regulation of enzyme synthesis.
                          Adapted from Reference 276.
                                      15

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for a protein, called the represser, which binds specifically to a control site called the
operator region. The represser protein binds to the operator region in the absence of
an inducer molecule, and the entire enzyme system is inactive. When the inducer is
present it binds with the one represser protein and the complex has reduced affinity
for the operator region. The operator region then complexes with RNA polymerase
at the adjacent promoter region and transcription of the structural genes is initiated.
When the concentration of the inducer molecule falls below a critical point, the
operator region is again blocked by the represser protein and enzyme synthesis
ceases.
  Very long or branched metabolic pathways, in which an intermediate substrate
may be directed to alternative pathways, are regulated by sequential induction, in
which sections of the pathway are under separate regulatory control. The product of
one series of steps acts as the inducer for the next several steps. This prevents the cell
from wasting energy on unnecessary or unproductive metabolic processes.
  Gene systems for a particular function which are grouped in one place physically
on the genome are rare in eukaryotes. Genes for a particular metabolic pathway are
more likely to be scattered over many chromosomes. However, regulatory genes still
function in a similar fashion at separate control sites. Some bacteria (prokaryotes)
also have systems in which the genes are  scattered along the chromosome.
  A more general type of control is called catabolite repression, in which the control
protein binds to operator sites  of many enzyme systems. This  permits  a favored
substrate to be utilized preferentially before other substrates are metabolized. As
long as the substrate of choice is present,  other substrates are not metabolized, even
though they may also be present. When the concentration of the favored substrate is
reduced, enzyme systems for metabolism of the other substrates are induced. The
favored pathway is more efficient and therefore costs the cell less energy.
  Another consideration in the effectiveness of enzyme activity  is the physical
location of the enzyme with respect to the substrate. In eukaryotes the enzyme may be
enclosed within membrane-bound organelles. In  prokaryotes an enzyme may be
enclosed within the periplastic  space between the cell wall and the cytoplasmic
membrane, while the substrate may be extracellular or intracellular.
  All  of the factors regulating enzyme activity may act in concert. Control of
metabolic processes is finely tuned to the nutritional opportunities available, so that
the cell acts in the most energy-efficient manner possible.

METABOLIC ENERGY PRODUCTION

  All microorganisms need a source of energy for maintenance of cell viability and
growth. The manner in which energy is obtained varies, and bacteria can be classified
according to the source of their energy requirement. Phototrophs such as cyano-
bacteria use light directly in a photosynthetic process. Chemotrophs oxidize organic
or inorganic compounds. A chemotroph  which can derive its carbon requirements
from carbon dioxide is a lithotroph, while a  chemotroph which utilizes organic
carbon is known as an organotroph.
  The most common method of gaining energy is through oxidation reactions, which
are normally coupled  to the formation of ATP and other high energy molecules.
Many different kinds of substrates can be oxidized, but eventually the substrates are
modified to  metabolites  which can enter one  of only a few pathways for carbon
dissimilation. These pathways can be divided into two categories, fermentation
pathways in which organic compounds serve as  both the electron donor and electron
acceptor, and respiration pathways in which oxygen or an inorganic compound or
ion serves as the terminal electron acceptor.
  Aerobes utilize a respiratory pathway known as the  tricarboxylic  acid cycle
(Figure 7). For each mole of glucose converted to acetyl-CoA which completes the

                                     16

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                  o
                  II
                  C-S-CoA

                ACETYL-CA
                                                 CoA-SH+H+

COO^ ^
?=° H20
CH2
coo-
OXALOACETIC ACID
f
coo-
HO-CH
i
CH2
•
COO

L-MALIC ACID
It
coo-
CH
HC
COQ-
/ coo-

HO-C-COQ-
9H« V\
coo- \\
CITRIC ACID \ »
1 \
coo-
CH2
c-coo-
II
CH

COQ-

cis-ACONITIC ACID
i i
COQ-
CH2
HC-COO-
HO-CH
coo-
ISOCITRIC ACID
 FUMARIC ACID
COO"

CH2 ,

CH2

COQ-
                                    coo-
                                              CoA-SH
                           f
                         CoA-SH
             SUCCINICACID
  CH2

  C-S-CoA   C02

  0

SUCCINYL-CoA oc _ KETOGLUTARIC ACID
                       Figure 7.  Tricarboxylic acid cycle.
                         Adapted from Reference 276.
cycle, 38 moles of ATP are generated. The intermediates in the cycle are precursors to
important cell macromolecules and may be utilized to fulfill other needs. Other
metabolic reactions act to replace the intermediates in order to maintain functioning
of the cycle.
  Under anaerobic conditions some facultatively anaerobic bacteria utilize anaer-
obic respiration. This is an oxidative process utilizing the same pathway for substrate
degradation  as  aerobic respiration,  except that nitrate  or another inorganic
compound is substituted for oxygen as the terminal electron acceptor.
                                     17

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3-KETOADIPIC ACID PATHWAY

  One of the maj or pathways for the degradation of aromatic compounds in bacteria
and fungi is the 3-ketoadipic acid  pathway (Figure 8).  The primary  aromatic
substrate is  converted to either catechol or protocatechuic acid, each  of which
undergoes several catabolic reactions in two separate but parallel pathways, until
they converge to three common intermediates, 3-ketoadipic acid enol-lactone, 3-
ketoadipic acid and 3-ketoadipyl-CoA, which is cleaved to form succinic acid and
acetyl-CoA. These two end products enter the tricarboxylic acid cycle. The pathway
                                                      PROTOCATECHUIC ACID
  cis.cis-MUCONICACID
         I cycloisomerase

        T
           :=o
(+)- MUCONOLACTONE
                      4-CARBOXYMUCONOLACTONE
                                                  3-CARBOXY-cis, cis-MUCONIC ACID
                      isomerase    . ,
                                                    3-CARBOXYMUCONOLACTONE
                    3—KETOADIPIC ACID ENOL—LACTONE
                          3-KETOADIPIC ACID
                                                                SUCCINIC ACID
u V^ C -SCoA-CoA-SH
k^Cig
9Ha 3-KETOADIPYL-CoA
C = 0
SCoA
ArETvL_r~A -^

thiolase
	 4
COOH
CHZ
CH2
C=0
SCoA
to- sunn
       Figure 8.  The 3-ketoadipic acid pathway in bacteria (path I) and fungi (path II).
                       Adapted from References 66a, 93, 408.
                                      18

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is strictly aerobic and regulated exclusively by control of enzyme synthesis. Although
the chemistry of the pathway is the same in all bacteria, the pathway is regulated
differently in different groups. Details of this pathway are found in the chapter on
parent compounds.
  The 3-ketoadipic  acid pathway is  one of  the best studied cellular metabolic
processes. The chemistry of the pathway has been elucidated (144,238,289,340,341,
342, 409),  the  enzymes isolated  and  their amino acid  composition and other
properties identified (123, 307, 336, 337, 345, 348, 396, 477, 478, 479, 480), and
microbial regulation of the pathway described (335, 338, 339).  A comprehensive
review of the pathway has been published (408).

PLASMIDS

  Genes coding  for vital functions  of the bacterial cell  are located  on the
chromosome and are passed to every daughter cell.  However, some metabolic
processes, while not essential, confer considerable advantages on cells with those
capabilities. The genes for these processes are coded for on plasmids, circular strands
of DNA which can replicate autonomously. Plasmids can be passed from cell tb cell
as well as being replicated in the progeny; thus, an entire population can  quickly
acquire the  specific characteristic. Traits which are often coded for on plasmids
include the ability to metabolize unusual substrates including many aromatic
compounds, resistance to antibiotics, and ability to survive in the presence of heavy
metals. Presence of a specific plasmid is often a guide to the metabolic capability of
that cell. Regulation of enzymes coded for on plasmids is similar to that discussed
earlier, and catabolic repression  may  be effective  across both plasmid  and
chromosomal DN A. Thus, a substrate which could be metabolized by two pathways,
one on the plasmid and one on the chromosome, may be metabolized by one pathway
preferentially while the other is repressed. A pathway for mineralization may involve
some steps  coded for on the plasmid and others coded for on the chromosome.
  Since plasmids may be considered potential vehicles for genetic reassortment and
transfer, they  may also be viewed as mediators of evolution  of biodegradative
capabilities  within microbial populations. At the population level, the development
of DNA probes labeled with 32P or fluorescent reagents can permit detection and
monitoring  of specific catabolic genes.  Such applications would likely utilize colony
hybridization  techniques  to  directly  probe for  complementary target DNA in
individual microbial colonies. Information derived from such experiments would
allow measuring the selective pressure required to maintain catabolic genes in the
natural population. In addition, the survival and transfer of novel catabolic genes
originating  from  recombinant DNA  technologies can  also be tracked in the
environment. Such information can be useful in the finer detailed prediction of the
kinetics of biodegradation and the likelihood of utilizing genetically engineered
microorganisms to degrade specific chlorinated aromatic pollutants.
                                     19

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                              SECTION 4

           CELLULAR GENE CODING AND GENETIC

                          TECHNOLOGIES

STRUCTURE AND FUNCTION OF DNA
   Deoxyribonucleic acid (DNA) consists of 4 kinds of deoxyribonucleotides linked
together in a specific sequence. DNA is usually double-stranded. Of the four kinds of
ribonucleoside bases, there are two subsets of hydrogen bonded pairings, adenine
with thymine and guanine with cytosine. RNA (ribonucleic acid) is also composed of
4 types of bases, but with uracil substituted for thymine. Each nucleoside base is
joined to the carbon-1 of a pentose sugar (deoxyribose in DNA, ribose in RN A). A
phosphate molecule is joined through  an ester linkage to the carbon-5 and  the
resulting molecule is known as a nucleotide. The nucleotides are joined by linking a
hydroxyl group on the carbon-3 of a pentose to the phosphate group on the carbon-5
of another pentose to form a phosphodiester bridge. The pentose carbons are primed
(" ' ") in order to distinguish them from carbons in the bases,'thus the two ends of
single-strand DNA are known as the 5'-phosphate end and the 3'-hydroxyl end. DNA
from different species has characteristic relative amounts of the 4 nucleotides and this
property has  served  to help identify unknown species  of bacteria  and  establish
evolutionary relationships among the species (Figure 9, Figure 10).
  Double-stranded DNA consists of two strands in which the 5'-end of one strand is
paired with the 3'-end of the opposite strand. The two strands are complementary —
anadenine on one strand always pairs through hydrogen bonding with a thymine on
the other strand, as does guanine with cytosine.
   Each set of three bases along a strand is called a  codon and codes for a specific
message, usually formation of an amino acid.  Some triplet sets of bases are stop
messages while others are nonsense codons and lead to  premature termination of
message reading. There is some redundancy in the triplet codes. Several triplets code
for the same amino acid, so a change in one base may not cause a functional change in
the message. Groups of triplets code for sequences  of amino  acids which become
proteins  after  some modifications.  The DNA  segment  which codes for a single
sequence of amino acids is known as a gene. The products of several genes may
combine to form a protein.  The total genetic material of a cell is called the genome,
consisting of the chromosome and in some cases plasmids. In eukaryotic cells the
chromosome includes some proteins also.
   Since bases are read in groups of three, the deletion or addition of one or two bases
will cause a shift in the reading frame such that the DNA is no longer read correctly.
This alteration is called a frame-shift mutation and may or may not be lethal to the
cell.
   Hydrogen bonding as well as hydrophobic interactions of the molecules free DNA
to take on a highly structured configuration resembling a double helix. Physical and
chemical perturbations such as heat or acidity cause denaturation or  unwinding of
the DNA,  eventually leading to separation  of the two strands.  The  covalent
molecules joining the bases  within each strand are not broken. When the physical or
                                    20

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                        \ g   // 5  6 NN           / 6  5 \\
                         N-C4      1N	H-N1      4C-H
                       £      \ 3  2 /            \ 2_3p./

                                     H         O        ,$~
                           adenine                thymine
                                             H-/       H
                           "C4     1N-H	 N1      4C-H
                              ^ 3  2 /             2  3  '
                               N  =C             C  - N
                                     >H	
                                    H
                           guanine                 cytosine
Figure 9.  The pairing of adenine with thymine and guanine with cytosine by hydrogen
bonding. The symbol—dR—represents the deoxyribose moieties ol the sugar-phosphate
      backbones o1 the double helix. Hydrogen bonds are shown as dotted lines.
                                       21

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  Figure 10.   Schematic representation of the DNA double helix. The outer ribbons
  represent the two deoxyribosephosphate strands. The parallel lines between them
 represent the pairs of purine and pyrimidine bases held together by hydrogen bonds.
Specific examples of such bonding are shown in the center section, each dot between
  the pairs of bases representing a single hydrogen bond. The direction of the arrows
  correspond to the 3' to 5' direction of the phosphodiester bonds between adjacent
  molecules of 2'deoxyribose. After J. Mandelstam and K. McQuillen, Biochemistry of
                Bacterial Growth, 2nd ed. New York: Wiley, 1973.
Reference: Stanier, K.Y., E.A. Adelberg, and J. Ingraham, The Microbial World, Prentice
                     Hall, Inc., Englewood Cliffs, N.J.  1976.
                                     22

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chemical stress is removed, the two strands will anneal spontaneously. When
denatured DNA from two different sources is mixed, a certain percentage of DNA
from the separate sources will anneal to form a DNA-DNA hybrid. The amount of
hybridization depends on the percentage of complementary sections between the
DNA. Strands of RNA can also combine with DNA to form RNA-DNA_hybrids.
  The chromosomal DNA of E. coli consists of about 4 x 106 base pairs, has a
molecular weight  of about 2.6 x 105, and is about 1400 ftm long (the E. coli cell is
about 2 /urn long).  Eukaryotic cells contain from 10 to 600 times as much DNA as E.
coli  cells. Eukaryotic DNA is organized into several linear chromosomes, each
carrying a unique  set of genes as well as proteins.
   Segments of DNA in eukaryotic cells are repeated many times, while prokaryotes
usually lack repetitive sequences. Some regions of eukaryotic DNA are characterized
by inverted repetitions of base sequences which may be a few to a thousand base pairs
long. Eukaryotic genes also contain segments which do not code for an amino acid
and are not translated. These intervening sequences or introns have been found in all
eukaryotic genes yet examined. Their function is unclear. They have been postulated
to contain regulatory signals or to separate the genes into smaller units which can be
readily recombined into new genes.
   The definition  of what constitutes a gene  has modified  over the years as the
biochemical exploration of the cell has become more detailed. A gene classically has
been taken to mean the genetic material which specifies a single trait. More recently,
portions of the DNA have been classified as structural genes if they code for a single
polypeptide (a portion of a protein) or for a specific type of RNA, or as regulatory
sequences if they function to mark the beginning and end of structural genes or start
or terminate transcription.
   There are two functions of the DNA molecule. The first function is to serve as a
template for its own  replication.  Enzymes separate the two strands, add new
complementary bases to each intact strand, and ligate the bases. Since the original
two  strands were complementary,  the  two  new complete DNA molecules  are
identical, each containing one old strand and one newly synthesized complementary
strand.

TRANSCRIPTION
   The process of converting the information  coded by DNA into RNA is called
transcription and  is the second function of DNA (Figure  11). Only portions of the
chromosome which code for a specific sequence or sequences of required genes is
transcribed at any given time. A single strand of RNA complementary to the DNA
strand is generated. Most of the RNA so formed is called messenger RNA (mRNA),
which codes for the amino acids which comprise  the polypeptides of the proteins.
Other sections generated are transfer RN As (tRNA), ribosomal RN As (rRN A), and
regulatory sequences.
   The mRNA serves to carry the genetic message from the DNA in the nucleus or
nucleolus to the ribosome, a collection of proteins and RNA units which is the site of
protein synthesis in the cytoplasm. A single mRNA molecule contains the message
needed to code for one or several polypeptides as well as a leader region and
intergenic spacer regions which are not translated.

TRANSLATION
  The process of protein biosynthesis according to the code carried by the mRNA is
called translation  and  takes place at the ribosomes in the cytoplasm. There  are
specific transfer RNAs that recognize each triplet codon which codes for an amino
acid. The tRNA molecule has receptor sites for both the mRNA chain and specific


                                    23

-------
   CO
   i
 3  =?
 3  »
 3  
-------
amino acids, and serves as an adapter to bring the appropriate amino acid in close
proximity to the developing polypeptide. Enzymes then attach the amino acid to the
chain and remove the tRNA. When a termination codon is read, biosynthesis stops
and the chain is released from the ribosome. The polypeptide finally is subjected to
post-translational modification of some of its amino acids and undergoes folding
into  its  characteristic three-dimensional  shape,  which  renders the  molecule
biologically active.
   Ribosomes are composed of ribosomal RNA and proteins. The rRNAs have a
specific three-dimensional structure and serve as a framework for the binding of the
polypeptide  subunits. The  ribosomal proteins are postulated to function  in the
process of synthesizing the polypeptide chain.
   The process of translation is repetitive and a single mRNA molecule can be read
simultaneously by several ribosomes spaced closely along the length of the molecule.
In bacteria, translation of mRNA begins while the molecule is still being transcribed
from the DNA.  Thus,  in prokaryotes these two processes  are closely  linked. The
prokaryotic  mRNA is  quickly degraded by nucleases,  so that efficient regulatory
control over protein synthesis is maintained.

MUTAGENIC EVENTS
   Most traits of bacteria are conservative and are reproduced in each generation.
However, like all living organisms, bacteria may undergo mutations in which the
genetic message  is altered. If the alteration is lethal for the  cell, the message is not
passed on because the cell dies.  Other mutations may not result  in a change in
expression of the message, since there is some redundancy built into DNA codes. In
some cases,  an alteration in the DNA code may lead to an alteration in the cell's
metabolism. Some mutations allow the cell to survive at unfavorable temperatures or
in the presence of potentially toxic compounds. In other cases, the cell acquires the
ability to use previously unsuitable substrates. Most mutations are either lethal or
place the cell at an environmental disadvantage.
   Mutations occur randomly, on the order of approximately one in one million cells
for a given characteristic: Some agents, including ultraviolet  light, some kinds of
radiation, and some chemical agents, cause increased mutagenesis. These mutations
are characterized by being randomly distributed across the DNA. Mutations can also
be selected by applying selective pressure to a population.  For instance,  in the
presence of an unfavorable environment, only those cells which have mutated in such
a way as to adapt to the environmental situation will survive.
   Cells have powerful mechanisms for excising mutations from DNA. There are
enzymes which recognize specific types of  mutations  and replace them with the
correct message. Thus, the number of mutations passed to  progeny cells may be a
small fraction of the total number of mutagenic events sustained by the cell. In the
case  of massive  mutations, such  as radiation damage, the  cell sets into motion a
complex series of steps designed to foster cell replication at the expense of almost
every other cellular function. The resultant cells are usually heavily damaged and do
not function normally.  The survival rate of these cells is very low.
   Classical genetic techniques are  based  on manipulation  of whole  cells and
environments  to select and induce desired  mutations. One  common induction
procedure is to expose a population of cells to a broadly acting mutagen and then
place the surviving cells in  the desired environment. Some mutations which have
occurred in the genetic region of interest may enable those cells to grow or express the
desired trait. A disadvantage of this method is that multiple  mutations may have
occurred in other genetic regions of the cell which may change the properties of the
cell in unknown ways. Another method  of obtaining  mutations is to expose the


                                     25

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population directly to the selective property. Only cells which can adapt will grow.
However, such adaptation may take weeks, months or longer before the altered
population is large enough to be observed.
   Some types of genetic alterations in the cell occur as normal cellular events.
Exogenous DN A can enter the cell by a variety of mechanisms and once inside can
recombine with the chromosomal DN A. This process, called genetic recombination,
can result in addition of new genetic information or the substitution of homologous
DNA sequences. One method of genetic recombination is the transfer of exogenous
DNA into a recipient bacterial cell. This process is called transformation and is the
only direct evidence for DNA being the genetic material. Only a minority of recipient
cells is competent at any given time to receive the DNA. Once the DNA has entered
the cell, it may find its homologous region on the chromosome, recombine, and
become a permanent part of the host chromosome, or it may recircularize into an
autonomous plasmid which replicates and is passed to daughter cells along with the
chromosome.
   Conjugation is a method which permits the entry of large segments of DNA from a
donor cell into a recipient (Figure 12). Direct cell-to-cell contact is required between
the donor cell, which possesses a  particular plasmid-encoded mating appendage
through which the DNA passes, and the recipient cell which lacks the appendage.
Upon contact, the donor cell is stimulated to begin replication of the plasmid and the
copy is threaded through the conjugation bridge  to the  recipient.  The donor
chromosome itself can be transferred to the recipient cell if the plasmid has integrated
into the chromosome. As long as  contact  can be maintained, transfer of genetic
material continues. The transfer of genetic material always begins at the plasmid
origin of replication. Therefore, by separating the cells at specific time intervals and
noting  which  traits  have been transferred,  mapping  of the  genes along the
chromosome can be achieved.
   Transduction is the term given to transfer of genetic material by bacteriophages
which are viruses specific for bacteria (Figure 13). During the packaging of viral
DNA into phage heads in the lytic cycle, some portions of bacterial DNA will be
incorporated instead. When these particles are expelled from the cell and infect
another cell, the bacterial DNA is released into the new cell to recombine with
homologous host DNA.
   In eukaryotic cells both parental chromosomes contribute genes to the daughter
chromosome. Both parental chromosomes undergo cleavage at homologous points
and segments of the chromosomes are exchanged. The new combinations of genes
are spliced together and passed to the progeny cells.
   Some segments of DNA are highly mobile and can leave their original position in
the chromosome to be inserted elsewhere. Each end of these transposable elements or
transposons contains short DNA pieces called insertion sequences. The insertion
sequences are recognized by specific enzymes which catalyze their insertion into new
sites on the chromosome or plasmid.
    In  recent years  techniques  have been developed  which permit the direct
manipulation of specific genes. Many of these techniques are now well established
and are being applied to solve specific problems. Descriptions of the fundamental
techniques and methods  follow. Additional information  can be obtained from
references which served as the basis for this chapter (276, 294, 334). In particular,
reference 294 contains details of the methods discussed here.
                                    26

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F incorporated
into chromosome
  Segment of
  (+) DMA
  inserted into
  (-) DNA
                                        Chromosome
                                        Sex factor F
                                        (a plasmid)
                                        (+) cell
                                      Conjugation
                                        (-) cell
Deleted
portion of DNA
of (-) cell

(-) cell
  Recombinant cell, now containing genes from (+) cell
Figure 12.  Transfer and recombination of genes during bacterial conjugation. The
DNA of the (+) cell \s replicated by the rolling-circle process, and the resulting single
            strand containing F is introduced into the (-) cell.
                       Reference 276.
                           27

-------
                 Viral DNA
                 Transducible
                    genes
                                        Bacterial
                                        chromosome
                                         Donor
                                         bacterium
                              LYSIS
                 Viral DNA
                 with genes
                 from donor
                 cell
 Transducible
 genes of donor
 bacterium now
 carried by phage
Chromosome
                                         Acceptor
                                         cell

                                          Transduced
                                          gene is
                                          incorporated
                                          into the
                                          chromosome
                                          of the
                                          acceptor
                                          cell
                Figure 13.  Genetic recombination during viral trans-
                   duction of bacterial genes into a recipient cell.
                              Reference 276.

CURRENT BIOCHEMICAL TOOLS FOR GENETIC

MANIPULATION

  A series of prokaryotic enzymes  has been isolated which can be used to cleave
DNA and then splice different pieces together to form new strands. These enzymes
are now being utilized in the laboratory.
                                 28

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Restriction Endonucleases

   Restriction  endonucleuses  are enzymes which specifically recognize certain
sequences  within double-stranded DNA. These sequences are usually 4 to  6
nucleotides long with  a two-fold axis  of symmetry. Examples  of restriction
endonuclease recognition sequences are  shown in Table 1.  A particular tetra-
nucleotide recognition site might arise once in every 46  (40%) pairs, assuming
random distribution of DNA base pairs. The endonucleases  cleave the DNA
molecules either at the axis of symmetry, yielding blunt double-stranded ends, or at
positions offset  from the center, giving fragments of DNA with one protruding
single-stranded end known as "stieky"ends. DNA from different sources acted on by
the same restriction endonuclease will produce complementary termini. In some
cases, different  restriction endonucleases with different recognition sequences will
produce complementary termini as well. These ends can join with complementary
ends on a different fragment to form new molecules.

                 TABLE 1. RECOGNITION  SITES FOR
                  RESTRICTION ENDONUCLEASES*
    Enzyme
  Recognition
   sequence
          Termini
  Bam HI
  EcoRI
 Haelll
 3'  5'

GGATCCt
CCTAGG
      I

GAATTC
CTTAAG

  I
GGCC
CCGG
G
CCTAG
G
CTTAA
GG
CC
GATCC
     G
AATTC
     G
    CC
    GG
 Hindlll
AAGCTT
TTCGAA
A
TTCGA
AGCTT
     A
 Mbol
GATC
CTAG
                                              xx
                                              xxCTAG
                GATCxx
                     XX
 Pstl
      i
CTGCAG
GACGTC
CTGCA
G
     G
ACGTC
 Thai
CGCG
GCGC
CG
GC
    CG
    GC
*Reference 294.
'Arrows indicate site of cleavage. (A) Adenine. (C) Cytosme, (G) Guanme. (T)
 Thymine.

-------
Deoxyribonuclease
  Deoxyribonuclease (DNase) is an enzyme that cleaves double-stranded or single-
stranded DNA randomly, yielding fragments with 5'-phosphate termini. Depending
on the conditions of the reaction, either the double strand of DNA is cleaved at
approximately the same site or each strand is cleaved independently. Under certain
conditions  this enzyme creates nicks  in double-stranded DNA which does  not
destroy the unity of the molecule. The concentration of DNase in the solution  will
affect the extent of nicking. Fragments of DNA containing regions of interest can be
inserted into  other molecules. Nicked regions  on  DNA permit insertion of
nucleotides. The order of insertion can be controlled to permit creation of a defined
DNA strand, and the nucleotides can be radiolabeled to permit tracking of the
constructed strain during other manipulations.

Polymerases
  Polymerases are enzymes that add nucleotides to the 3'-hydroxyl terminus created
when the double-stranded DNA molecule is nicked by DNase. The enzyme  can also
remove nucleotides from the 5'-phosphate ends. Both of these processes acting at the
same time result in  movement of the nick  along the intact strand of DNA (nick
translation). If the nucleotides being added are radioactive, such as with  32P,  the
labeled DNA can be prepared with a high specific activity. Normally the replacement
nucleotides are distributed uniformly along the DNA molecule, since the nicks occur
randomly throughout the DNA.

Reverse Transcriptase
   This enzyme, also known as RNA-dependent DNA polymerase, uses mRNA as
the template for transcription to form double-stranded DNA. Single-stranded DNA
or RNA can also be utilized by this enzyme  to make probes for use in hybridization
experiments.  Initiation of the  action of reverse transcriptase requires a short DNA
primer sequence  base-paired to  the template.  Primers can be generated  by
exhaustively  digesting DNA  and retrieving the fragments. Since the fragments
represent random portions of DNA, some fraction of them will bind to the template
and  can be used as primers. The discovery that eukaryotic mRNA contains  multiple
adenylate bases at the 3'-end allows construction of complementary polymer thymine
residues at that site, which then will act as a primer.
   The RNA template possessing a primer is then mixed with solutions of the 4
nucleotides. Usually only one nucleotide is radioactively labeled. Reverse transcrip-
tase then catalyzes synthesis of the complementary DNA  (cDNA). Following
completion  of the reaction, the RNA  strand  can be  selectively degraded.
Complementary DNA to be  used as hybridization probes are retained in single-
stranded form. However, the cDNA can form a hairpin loop at its 5'-end which acts
as a self primer for synthesis of the complementary strand, or a second primer can be
added to the  cDNA to initiate synthesis, resulting in double-stranded cDNA.

Ligases

  T4 DNA ligase links together complementary fragments of double-stranded DNA
by forming a phosphodiester bond between adjacent 3'-hydroxy and 5'-phosphate
ends. The ends may be either sticky (one strand of the double-stranded molecule
extends beyond the other) or  blunt (both strands  end at the same place). T4 RNA
ligase  joins single-stranded RNA or  DNA. These  enzymes  can  also  catalyze
circularization of DNA molecules if the concentration of DNA in solution is low.

                                     30

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Transferases

  The enzyme, terminal deoxynucleotide transferee adds deoxynucleotides to the
3'-hydroxyl end of DNA. By adding homopolymer  sequences  of one type of
nucleotide  to one set  of DNA fragments, and a series of complementary homo-
polymer nucleotides to a second set of DNA molecules, the two populations can be
joined by their newly formed complementary ends.

Methylases

   These enzymes add a methyl group to particular nucleotides. Some restriction
endonucleases will fail to recognize a sequence which differs only by addition of a
methyl group. This is an important component of cellular defense systems, in which
host DNA is methylated to protect against host restriction endonucleases which will
attack foreign (nonmethylated) DNA.

MECHANICAL SHEARING OF DNA
   Double-stranded DNA can be broken by the shearing forces present in solutions.
Very small fragments  (approximately 300 base pairs in length) can be obtained by
subjecting the solution to  sonication with ultrasound.  Larger fragments of about
8,000 base pairs result  from stirring the solution at high speed in a blender. The DNA
molecule is sheared randomly along its length, producing fragments with short
single-stranded ends.

CLONING VEHICLES
   In order for  a fragment of DNA to be replicated, it must contain a specific
sequence called an origin  of replication. Plasmids and prokaryotic chromosomes
each usually contain one origin of replication. DNA which possesses an origin of
replication is called a replicon. If a fragment of DNA in a cell cannot replicate, it will
be diluted out of the population after several generations. Therefore, DNA fragments
of interest must be attached to replicons, called vectors or cloning vehicles, before
insertion into the cell. Replicons which are not native to the host cell  may not be
functional  after insertion.  The combination of the replicon and the foreign DNA
fragment  creates a hybrid molecule often called a chimera. The  process  of
constructing a hybrid  DNA molecule is known by several names, including genetic
engineering or gene manipulation,  to acknowledge the potential for creating new
combinations of genes, and gene cloning or molecular cloning because this method
allows amplification of the chimera via growth of the host population of organisms,
each carrying the identical piece of genetic information. The DNA can be extracted
from the new population and the chimeras recovered.

Plasmids
  Plasmids are stable, extrachromosomal, circular double-stranded DNA replicons
which are inheritable,  but are also dispensable. Under constant selective pressure the
plasmids will be replicated in the daughter cells, but  when not essential for  cell
function many plasmids are lost from the cell. Plasmids contain from 1,000 to
200,000 base pairs. Conjugative plasmids carry a set of genes that promotes bacterial
conjugation; nonconjugative plasmids lack these genes. The term relaxed plasmids
refers to plasmids which are present as multiple copies (10 to 200 copies) within a
single cell, while stringent plasmids are limited to 1 to 3 copies per cell. Generally,
plasmids of relatively high molecular weight are conjugative and stringent, while low
molecular weight plasmids are nonconjugative and are present in multiple numbers.

                                     31

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If cellular protein synthesis is blocked, chromosomal and stringent plasmid
replication ceases, while relaxed plasmids continue to replicate and can increase their
copy number to several thousand per cell. Therefore, relaxed plasmids are most
useful for molecular cloning processes. Some plasmids ("promiscuous plasmids")
can be transferred into a wide range of Gram-negative bacteria. These plasmids are
potentially useful in transferring genetic information to diverse bacterial hosts. Some
plasmids are incompatible with others and  cannot coexist within the  same cell.
Plasmids have been  grouped into incompatibility classes on  the basis  of mutual
incompatibility.
   Plasmids useful as cloning vectors generally are small and under relaxed control.
They carry an easily  selectable marker (such as antibiotic resistance) which allows
identification of transformants which have acquired the plasmid. These plasmids
also contain a single recognition sequence for a given restriction endonuclease which
permits insertion of DNA into a region of the plasmid not essential for replication. A
restriction site located within the marker genes will inactivate the gene when foreign
DNA is inserted, providing a tracer for successful DNA insertion.
   Some plasmids have been modified to include polylinkers, segments of DNA that
contain closely  spaced recognition  sites for several  restriction endonucleases.
Generally, plasmids so modified are small and lack natural restriction sites. Use of
small plasmids is advantageous in that they are less likely to be damaged physically
during handling. Small plasmids also tend to generate higher copy numbers.
   Construction of plasmid vectors involves cleaving both foreign DNA and plasmid
DNA with the same restriction endonuclease to form complementary ends. Both
types of DNA are mixed and are ligated. In some of the resulting molecules the
foreign DNA will be ligated to the plasmid DNA, and a circular recombinant plasmid
recovered. Use of a restriction site within a marker gene simplifies the process of
detecting recombinant plasmids. The inserted DNA inactivates the gene. Plasmids
which  recircularize without insertion of foreign  DNA will  express the marker
characteristic and can be rejected during the screening process.
   Recircularization  of plasmid DNA can be minimized using a procedure called
directional cloning. This method takes advantage of  the fact that most plasmid
vectors carry single recognition sites for more than one restriction enzyme.  A plasmid
is digested with two such endonucleases. The larger fragment is separated and ligated
with foreign DNA containing ends complementary with the  two dissimilar ends
generated by the two restriction enzymes. The plasmid fragment itself does  not
contain complementary ends and therefore will not circularize.
   Another  method  of preventing recircularization involves  treating  the linear
plasmid  DNA with  alkaline phosphatase. This enzyme removes the 5'-terminal
phosphates. Ligation  requires both  a 3'-hydroxyl  and a 5'-phosphate end. The
foreign DNA combines with the treated plasmid DNA  to create a circular molecule
with a single nick on each strand where the phosphates have been removed. This open
circular molecule can be inserted into cells much more efficiently than linear DNA
and so most of the transformants will carry recombinant plasmids.


Bacteriophages

   Bacteriophages (phages) are viruses which attack bacteria. The most extensively
studied phage is bacteriophage 1 which contains  a double-stranded linear DNA
molecule about 50,000 base pairs long with single-stranded complementary sticky
ends. The intact phage consists of the genomic material surrounded by a protein coat
with a protruding tail. The tail attaches to the bacterial cell and injects only the DNA
into the cell. One of two processes of replication, either lysogeny or the lytic cycle, can
be initiated within the cell.

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   During lysogeny, the viral DNA is integrated into the host chromosome and is
replicated and transmitted to progeny cells along with the host chromosome. This
can happen indefinitely. At some point the lysogenic cell is triggered to begin the
second process of replication, the lytic cycle. Alternatively, upon infection the viral
DNA can initiate the lytic cycle immediately.
   The lytic cycle begins with viral adsorption and DNA penetration. These steps
require specific conditions and the phages are host specific. After entry into the host
cell, the linear viral DNA  circularizes via its  complementary sticky  ends and
replicates as an independent molecule. Copies of the DN A are continuously made.
Transcription of the molecule is initiated soon after replication begins. One of the
earliest  proteins produced is a regulatory element which acts to prevent defensive
activities of the host which might otherwise prevent further transcription. During the
late phase of transcription, proteins involved in assembly of the head and tail and cell
lysis are produced. As many as 200 copies of a phage can be replicated within a single
host cell.
   During the assembly phase of the lytic cycle, a linear copy of the DNA becomes
coiled into a phage prehead. When the head is filled, an additional protein attaches
and locks the DNA into the head. The head finally attaches to the preassembled tail
unit to form a complete phage  particle. Progeny phage  particles are released in a
single burst when the host cell lyses. Each phage is then able to  infect another cell.

Bacteriophage 1 Vectors

   About one-third of the phage 1 genome is nonessential for virus replication and can
be replaced by foreign DNA so that the total length of the genome is conserved.
Although phage  1 contains several  recognition sites  for each of the restriction
endonucleases ot interest, derivatives of phage 1 have been developed which no longer
carry restriction sites in critical areas of the genome, but carry only 1 or 1 such sites in
nonessential regions. The phage 1 thus  has been manipulated to become a  useful
cloning  vector while still retaining its infective and lytic properties.

Cosmid Vectors
   Cosmids are  constructed vectors designed  for cloning large  fragments  of
eukaryotic DNA.  They consist of a drug-resistance marker, a plasmid origin of
replication, one or more restriction sites,  and the ligated sticky end of phage 1 (the cos
site). They are very small, so that large amounts of foreign DNA can be added to the
molecule. The complete cosmid  DNA is packaged into a bacteriophage coat which
mediates  its injection into the host  cell.  Inside  the  cell the cos  site allows
circularization and the plasmid origin of replication initiates replication.

Single-stranded Bacteriophage Vectors
   Bacteriophages containing single-stranded DNA replicate in a different manner
from  phage 1. After penetration, the single-stranded form is converted to double-
stranded DNA which can be isolated and used  as a cloning vector. The double-
stranded form replicates until 100 to 200 copies are made. Then DNA replication
shifts  to produce large amounts of only one of the two DNA strands. Single strands
are incorporated into the phage coats and the mature phage particles are continually
extruded from the cell without lysing the host cell.  The single-stranded DNA thus
produced can be recovered  and used as a template for DNA sequencing  or for
generating DNA probes.
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METHODS OF MANIPULATING DNA

Isolation and Purification of Plasmid DNA

  Plasmids inserted into cells can be amplified by growing the bacteria to high cell
yields in the presence of an antibiotic providing selective pressure from the plasmid.
The cells can be harvested by centrifugation and then lysed by several methods. Lysis
procedures include boiling, treatment with alkali, and treatment with the surfactant
sodium dodecyl sulfate. Lysozyme is added to help break apart the cell walls. The
treated solution is centrifuged to remove DNA from other cellular material.
   The DNA preparation contains chromosomal DNA as well as plasmid  DNA.
These two types of DNA can be separated by taking advantage of several differences
in the properties of chromosomal and plasmid DNA. For some applications, the
crude DNA preparation from as little as 10 ml of culture may be used successfully.
After treatment by one of the lysis procedures, plasmid DNA is recovered from cells
in intact circular form, while chromosomal DNA generally is extracted in short linear
pieces. When plasmid DNA of high purity is required, centrifugation at very high
speeds in a solution of cesium chloride and ethidium bromide will separate the DNA
according to density.  The  linear chromosomal DNA takes up  more  of the
intercalating agent ethidium bromide than the plasmid  DNA. The  chromosomal
DNA is physically stretched more than the plasmid DNA, and therefore will be a less
dense molecule. When subjected to ultracentrifugation, the two types of DNA form
narrow bands in separate regions of the centrifuge tube. Contaminating protein will
form a third band, and RNA will form a pellet. The ethidium bromide can be
removed after this step or it can be retained during subsequent procedures.

Isolation and Purification of Bacteriophage X DNA

   Bacteriophages which are lysogenic can be recovered by inducing the bacterial
culture  to begin  the lytic cycle.  One method useful  for  phages  containing a
temperature-sensitive represser is to raise the temperature of the culture briefly.
   Phages which are not lysogenic may be amplified by infecting the host bacterial
culture with a low number of phages. Much of the bacterial culture will replicate for
several generations, increasing the number of host cells, before successive rounds of
the lytic cycle infect the entire culture.
    Cell  debris remaining  after completion  of the lytic cycle  is removed  by
centrifugation.  The remaining solution can be  subjected  to  density gradient
ultracentrifugation, after which the intact bacteriophage particles  appear as a thin
band. Crude bacteriophage preparations useful for many purposes can be obtained
without density gradient ultracentrifugation.
   DNA can be recovered from the phage particles by treatment with  a solution of a
protein-digesting enzyme and sodium dodecyl sulfate. The protein components can
be extracted into phenol and removed by centrifugation.

Separation and Purification of DNA Fragments

   Gel electrophoresis is a sensitive method for resolving mixtures of DNA. Samples
containing DNA are loaded onto a slab of agarose or polyacrylamide gel. The gel is
submerged into a buffer solution of nearly the same electrical resistance and a current
is applied. Various types of DNA—linear, nicked circular, and closed circular—will
migrate through the gel at different rates, depending upon the molecular size of the
DNA, the concentration of agarose or polyacrylamide, the applied current, and the
DNA base composition and temperature. Under some conditions single strands of
DNA can be separated. The DNA is stained with the fluorescent dye ethidium
bromide and can be  detected directly. Bands of DNA can be cut from the gel and the
DNA recovered.
                                      34

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   The DNA is then packaged into a vector and transformed into a bacterial culture
for amplification. Colonies containing plasmids are grown on nitrocellulose or nylon
filters seated on agar petri plates, while bacteriophage plaques are formed in a lawn of
indicator bacteria on agar media and then eluted into a liquid suspension which is
stable indefinitely.

Identification of Recombinant Clones

   In situ hybridization of bacterial colonies or bacteriophage plaques on agar media
is  rapid  and can efficiently  screen large numbers of potential clones.  Colony
hybridization involves lysing the colonies on the nitrocellulose or nylon filter and
then fixing the DNA to the filter in situ. Bacteriophage plaques are transferred to the
nitrocellulose filters following plaque formation. Filters are placed in contact with
plaques on the agar and then removed. Some portion of the viruses in the plaques will
be removed with the filter. The DNA is then fixed, in some cases by heat treatment.
The DNA probe of choice is labeled with "P and the hybridization reaction between
the probe and  the fixed DNA carried out.

Hybridization of Probes to Immobilized DNA

    Hybridization  reactions  are governed  by such factors  as solvent used,
temperature, length  of  hybridization, concentration and specific activity  of the
32P-labeled probe or density of fluorescent probe,  and washing procedures after
hybridization. Prior to hybridization, the filters are treated with one of a number of
compounds  to saturate  sites on the filter with  nonspecific affinity for  single- or
double-stranded DNA, in order to ensure that the probe DNA will not bind directly
to the filter. The 12P-labeled probe DNA is denatured (double-stranded molecule
separated into its single-strand components) and  added to  the filters.  During
incubation, the single strands of the probe DNA will join to complementary DNA
strands on the filter. Following hybridization the filters are washed thoroughly to
remove unbound DNA and dried. An autoradiograph is made by placing the filters in
contact with X-ray film. Following development, radioactive signals representing
positive homology of the probe with plasmid  or phage DNA can be correlated with
colonies or plaques on  the agar plates.  Those colonies  can be retrieved and the
recombinant DNA contained therein amplified.
   Other procedures employ  DNA probes with  induced fluorescence  and/or
antigenic properties for  use with fluorescent  antibodies to detect positive hybrids.
Both methods have the sensitivity to detect as little as 1 picogram of DNA or as few as
10,000 copies of a single gene.

MAPPING OF RESTRICTION ENDONUCLEASE RECOGNITION

SITES
   The order of bases on a strand of DNA can  be determined using a number of
techniques; usually more than one is required to obtain a detailed map. By cleaving
DNA with restriction endonucleases having known recognition sites, the presence
and number of such sequences can be resolved.
   The relative positions of dissimilar endonuclease  recognition sequences can be
determined by first labeling one end of linear DNA with radioactive nucleotides to
obtain a reference point. Digestion of separate  aliquots  of the  DNA by different
restriction enzymes  is  followed  by gel  electrophoresis and  autoradiography.
Resulting fragments of different lengths define the distance of each recognition site
from the labeled end, and the relative distances between each site can be determined
as well.
                                      35

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   Small fragments of DNA can be mapped using an exonuclease which digests the
DNA from each end by single nucleotides. Samples of the digestion reaction are
withdrawn at time intervals and are treated with restriction endonucleases. As the
DNA digestion progresses, the recognition sites disappear in specific order related to
their position along the molecule.

IDENTIFICATION OF DNA SEQUENCES WITHIN FRAGMENTS

   The Southern transfer technique is an effective method for identifying particular
sequences of DNA. Fragments of DNA created by one of the previously described
methods are separated by size on agarose gels using electrophoresis. The DNA within
the gel is stained with ethidium bromide and denatured (strands separated) in situ.
The DNA is then eluted from the gel directly onto nitrocellulose filters by placing the
gel on absorbent paper whose edges are trailing in a salt solution. The filter is placed
on the gel and more absorbent paper placed above the filter.  Wicking action will
cause the DNA to migrate from the gel to the filter with the relative positions of the
fragments intact. The filter is treated with 32P-labeled probe DNA of known base
composition and then washed well. Fragments containing complementary bands will
hybridize to the probe and can be visualized after autoradiography.

EXPRESSION OF PROKARYOTIC GENES IN  FOREIGN HOSTS

   Most genetic engineering studies have used £. colias the host with introduction of
either E. coli genes or genes from other prokaryotes or  eukaryotes. Other studies
have involved introduction of genes from one  strain to another strain of the same
species. Little research has been conducted on expression of E.  coli genes into other
hosts. Some studies have noted that E.  coli genes cloned into a B. subtilis host were
not expressed, although the genes themselves when recovered and inserted back into
E. co/i were functional (334). This has been attributed to differences in the specificity
of the RNA polymerases of the two hosts. There are differences as well between the
translation mechanisms of E. coli and B. subtilis. Thus, 6. subtilis genes function in
E. coli but the reverse is not true.

GENE CLONING IN YEASTS

   The yeast Saccharomyces cerevisiae has received the most attention with respect
to application of genetic engineering techniques. This species contains a plasmid
which replicates with high copy number although it has  no known function (334).
Fragments of yeast DNA as well as an E. coli plasmid vector have been cloned into
this plasmid and the recombinant molecule transforms yeast with high frequency and
replicates in both E. coli and yeast. Some yeast genes replicate autonomously and
these can be used to construct vectors which transform yeasts with high efficiency.

EXPRESSION OF EUKARYOTIC GENES IN A PROKARYOTIC
HOST
   Genes from the fungi S. cerevisiae and Neurospora crassa have been cloned into
and expressed in E. coli(334). However, many  other genes from eukaryotic sources
have been cloned into prokaryotic hosts but have not been expressed. This has been
explained in part by the differences in mechanism of expression of the genes (protein
synthesis).  The steps involved in synthesis  of a functional  protein  include
transcription of the  DNA,  translation of the  mRNA,  and post-translational
modification of the newly-formed polypeptide. Transcription and translation require
a promoter or a binding site recognizable by the host RNA polymerase. Further, the
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protein produced is often subject to rapid degradation unless it is protected by the
modified amino acids or three-dimensional configuration of the native proteins.
Even if all of these components are present, genes which are expressed in a foreign
host may not necessarily give rise to a stable gene product.
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                              SECTION 5

     METHODS OF BIODEGRADATION ASSESSMENT

  A decade ago biodegradation was measured at a worst case level by substrate
disappearance.  Many reports  in  the  area  of environmental  microbiology  and
wastewater engineering documenting biodegradation were accompanied by gas
chromatographic analysis showing the net loss over time of a parent compound. In
some cases, even visual disappearance of insoluble crystalline organics, such as
naphthalene, was used as a gross measure of biodegradation. Most often, abiotic or
sterile control samples were used in such assays of biodegradation. However, little
insight was  developed into nonmetabolic interactions  among organisms  and
pollutant substrates and the measured biodegradation. In the area of wastewater
treatment, BOD or COD removal measured by comparing influent and effluent
concentrations  was  used  as  measures  of  pollutant biodegradation, with the
assumption that recalcitrant organics had essentially the same fate as labile organics
in wastewater treatment.
  Such approaches to measuring biodegradation have  been  replaced by more
stringent parameters to give accurate  estimates of microbial catabolic potential
under laboratory conditions and to determine more accurately the environmental
fate of chlorinated aromatic pollutants. The most stringent criteria for accurately
estimating biodegradation include mass or  material  balance approaches  and
mineralization  approaches. In actual  practice, both approaches are frequently
integrated to give  the best  estimate  and predictive capability of  determining
biodegradative  fate. In either  instance,  the  use of radiolabeled (primarily  I4C)
substrates complements the approach,  especially at environmentally realistic, low
concentrations. These trace concentrations  may  make  conventional analytical
approaches more difficult and/or expensive. The approaches are summarized in
Table 2.
  Laboratory assays for biodegradation, using labeled or unlabeled  compounds,
require a determination of physical processes which contribute to the overall loss of
the substrate. Accurate  material  balances are determined by an accounting of
substrate loaded onto biomass and suspended particulates,  aqueous phase substrate,
residual substrate sorbed to glassware and reaction vessels,  and volatilized substrate.
Assuming efficient recovery for each component phase, the difference between input
and accounted-for residual and comparison to abiotic control samples should give a
reasonable approximation of  true biodegradation.  However, even  under  these
circumstances,  biodegradation may be poorly understood if aqueous phase and
cellular-associated substrate is transformed to polar oxidized products. In such cases
I4C analysis  without conventional analysis (HPLC) may  underestimate biodegra-
dation, or conventional analysis may fail to detect transformation products that in
themselves are resistant to further microbial degradation. Joint conventional and I4C
analysis (where available) provide excellent material balance analyses for biodegrada-
tion of parent substrate and accumulated metabolic products.
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  TABLE 2. MATERIALS BALANCE AND MINERALIZATION APPROACHES
                  TO BIODEGRADATION ASSESSMENT
    Biodegradation approach	Process examined
Materials balances                Recovery of parent substrate
                                  Recovery of radiolabeled parent
                                    substrate and metabolic products
Mineralization                    Production of carbon dioxide,
                                    methane, or their carbon
                                    radiolabeled congeners from the
                                    parent substrate
                                  Release of substituents groups,
                                    e.g. chloride or bromide ion
  Mineralization assays  as a measure of total  biodegradation (oxidation  or
reduction to terminal decomposition products) have enjoyed utility as unambiguous
measures  of  biodegradation. In the event that non-MC-labeled substrates are
employed, mineralization assays must include an absolute materials balance for the
system. Such approaches have been used to study anaerobic biodegradation resulting
in methane (CH4) production as the terminal mineralization product (417).
  In more general  practice, CO2 production  is the most common measure of
pollutant mineralization.  With the commercial  availability or custom synthesis of
l4C-labeled organic pollutants, measurements of I4CO2 production indicating both
the extent and rates  of substrate degradation have become common practice.
  In cases involved with biodegradation of aromatic  and chlorinated aromatic
pollutants, the parent substrate is generally chosen with aromatic ring-labeled I4C
atoms. Production of I4CO2 during biodegradation is therefore the result of aromatic
ring oxidation and cleavage, representing virtually total destruction of the parent
aromatic molecule and associated bioactive properties  that are  of initial concern
from environmental health and ecological perspectives. In alternative mineralization
approaches where l4C-parent substrate may be available, the release of halogen ions,
generally Cl- or Br, from  the aromatic ring during ring oxidation and cleavage is a
good measure of biodegradation. However,  if the goal is to determine terminal
decomposition, care must be taken to assure that halogen ion release follows ring
cleavage  rather than preliminary reductive or oxidative  dehalogenation of the
aromatic ring.
  An integrated flow  diagram (Figure  14) describes biodegradation assessment.
Carbon-radiolabeled parent substrate is added to a reaction system containing the
microbiological  population  of interest. In  a time course fashion, samples are
withdrawn or replicate samples are sacrificed. At each time point, material balances
for the parent substrate and I4C are prepared using a combination of conventional
procedures (most conveniently analyzed by HPLC) and liquid scintillation analysis
of radioactive decay of I4C. Mineralized products such as  I4CO2 or I4CH4 can be
collected and analyzed by liquid scintillation analysis and confirmed by conventional
analytical methods.  Where  specific identification and confirmation  are required,
mass spectrometry (MS) or GC/MS can be employed for isolated products.
  The  resulting data, when compared to appropriate abiotic controls to accom-
modate nonspecific  sorption, volatilization or stripping, and photolytic or other
abiotic degradation mechanisms, provide the information to determine the potential

                                    39

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                       I  14C Parent Substrate]
1
Biodegr
| Assay
I
Conventional Analysis
Residual Substrate (GC/HPLC)
I
Bioti'aiibfoi iiialioii Pioducls (HPLC) •*
i
Identification (GC/MS)
Ba14COi Precipitation + IR/14CO, „
Gc/'4CH4 ~~

adation
System I
Liquid Scintillation
•^ 14C Residual Substrate

Products
Mineralization
Products
                 Time Course Determination/Kinetics
            Material Balance for Parent Substrate and '4C
          Comparison to Biologically Inhibited Control Sample
Figure 14.  Integration of materials balance and mineralization approaches in
                      biodegradation assessment.
                                  40

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and extent of degradation. A relative rank, in terms of rates of degradation, may be
assigned if multiple comparisons are being made.
  Biodegradation  assessment  may  be divided into  two broad  categories,  one
comprising studies of the environment or laboratory studies which simulate the
environment, and  a  second category  which includes pure culture  studies  under
defined environmental conditions. Environmental studies yield  information  on
disappearance or movement of the substrate. Pure culture systems permit studies at
the molecular level, including information about specific enzyme systems and gene
involvement in the manifestation of degradative capability. Studies of degradation
must include both mechanisms of induction and the mode of action of the enzymes.
However, results from laboratory studies are not necessarily an indication of results
to be  expected  in  the  environment. These  studies must  be  correlated  with
environmental conditions for an assessment of in situ biodegradation.
  Primary degradation or biotransformation is considered to be disappearance of
the substrate, without consideration of metabolite formation or mineralization.
While  primary degradation is important evidence to assess  the  potential biode-
gradability of the molecule, only knowledge of the metabolites formed  or of complete
mineralization will enable  confirmation of the  biodegradability of the substrate
under the specific environmental conditions used.
  The  biodegradation of a chlorinated compound is considered complete when the
chloride ion is returned to its  mineral state (HC1 or CF) and the  carbon skeleton
converted to cellular products (83).  Appearance of the chloride ion  is most
conveniently measured by using an ion-selective electrode.
  Disappearance of the substrate may be due to a number of factors in addition to
biodegradation. These include photolytic decomposition, volatilization, chemical
degradation, and sorption and  irreversible binding to soils, clays, or organic matter
including cells. 1 n addition to their separate effects, these factors may work in concert
with the biota to degrade the substrate. In studies of biological degradation, these
factors must be controlled, eliminated, or  accounted for.

CHEMICAL ANALYTICAL TECHNIQUES
  The ability to quantify the amount of substrate in a given experimental system is of
prime  importance. To this end,  sophisticated  analytical techniques  have  been
developed which  allow the unambiguous identification  of  the  substrate  or its
metabolites.
  Gas-liquid  partition chromatography  (GC) is  a  separation technique which
combines  high sensitivity, accuracy, and repeatability. Low concentrations of the
sample are required. The sample may be solid, liquid, or gaseous as long as the
sample can be volatilized at the operating temperature of the instrument. Samples
which are insufficiently volatile can sometimes be derivatized and converted to a
more volatile  ether or ester compound.
  The  sample to be analyzed is injected along with a gas which carries the sample
along a column packed with inert particles coated with a liquid. The solutes in the
sample are distributed between the liquid  and gas phases according  to the relative
solubility of the solute in the liquid. Solutes of lower solubility or high volatility move
through the column  at faster rates.  As the bands exit from the column they are
recorded as roughly symmetrical peaks with retention times related to their relative
partition coefficients. These can be compared with the retention times of standard
materials. This does  not constitute absolute proof of identity, however, as two or
more solutes can elute from the column with the same retention time.
  The solutes separated by GC can be analyzed directly by a mass spectrometer (MS)
for determination of the molecular structures of the compounds (GC/ MS). In gas
                                    41

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chromatography-mass spectrometry, the separated sample effluent from the GC is
highly ionized and accelerated in a vacuum, causing it to fragment into smaller ions.
These ions are separated according to their mass-to-charge ratio (m/e) and then
collected according to their relative abundance in the sample. The fragmentation
pattern  results from the molecular structure and is characteristic of the type of
compound. The mass  spectrum of the compound can be used to recreate its
molecular structure and often it can be compared with mass spectra in computerized
libraries for quick identification of unknown compounds.
  The use of GC/ MS has become widespread because of the potential unequivocal
identification and quantification of metabolites and residual substrate. Materials
present in minute quantities can be identified within a mixture of other materials.
  Additional information regarding the types of bonds between functional groups
can be obtained through infrared spectroscopy. The different types of bonds absorb
at specific frequencies when infrared radiation is passed through the molecule, and
the resulting spectrum is unique to the particular material. Infrared spectra are most
useful when used in conjunction with other methods of compound identification and
when looking for specific functional groups.
  Liquid  chromatography  can be an effective technique for the  separation of
materials  of either lower volatility or ionic structure.  The sample, in a solvent, is
passed through a column containing  an absorbing material dispersed on an inert
support. The solutes partition between  the liquid  and the absorbing material
according to  their  relative affinities and solubilities.  Each solute elutes from the
column at characteristic time intervals.  The great variety of both solvent  and
stationary phases makes this  a very versatile and sensitive technique for both
separation and identification of compounds by comparison with known standards.
Increased resolution can be obtained  by decreasing the  size of the particles  in the
stationary phase. This formerly required high pressures to achieve the desired flow
rates, leading to the term high pressure liquid chromatography (HPLC). Newer
developments have reduced the required pressure and  this technique is now widely
referred to as high performance liquid chromatography. Solutes eluted from a liquid
chromatography column  can be  used for  other applications  including  mass
spectrometry.
  A convenient way to monitor a substrate, subjected to biodegradation testing, is to
use a radiolabeled compound. When the exposure of a compound to biodegrading
agents is halted, the residual quantity of the compound can be measured by counting
the radioactive emission of the  solution  in a  liquid  scintillation counter.  If the
compound is uniformly labeled (all atoms of the given element are radioactive) then
the metabolites separated  by  liquid  chromatography can be examined for
radioactivity and a determination of the fate of the original substrate can be  made.
This  identifies  the  metabolites  arising from  the substrate  and eliminates  any
ambiguity arising from  the presence  of metabolites which may have arisen from
sources other than the substrate of interest.
  Radioactive-carbon (I4C) labeled compounds are particularly useful in mineral-
ization  experiments in  which carbon dioxide  is evolved. Mineralization  of the
substrate  will result  in radioactive carbon dioxide which can be quantified. The
advantage of this method is  that it  is  extremely specific  and gives unequivocal
evidence of the extent of mineralization for the  radiolabeled compound. However,
few compounds are routinely available in labeled form and custom synthesis is
expensive. Stringent regulations govern the  use and disposal of radioactive
compounds.
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ANALYSES OF METABOLIC ACTIVITY
  Determination ot the fate of a substrate in a biological community gives specific
evidence of metabolic activity. However, several nonspecific methods of analysis are
available which measure the  metabolic  activity  of the microorganisms.  If the
substrate of interest  is the only available  source of a required nutrient or energy,
metabolic activity is considered to be directly related to the presence of the substrate.
Lack of metabolic activity is considered to be evidence of inability of the microbial
population to utilize the substrate as long as all other essential nutrients and growth
factors are present. These methods do not give information about the metabolic
products arising from substrate utilization.
  The biochemical oxygen demand (BOD) is a measure of the oxygen required to
biochemically oxidize all the carbonaceous and nitrogenous matter in a sample. This
test  is subject  to many  variations in the  methods  of sample  collection  and
measurement of data, and results are comparable only between tests performed by
the same protocol. The BOD test is most useful for sewage and other samples with
high organic content. The test is not very sensitive as a measure of the bio-oxidation
of recalcitrant substrates.
  A more sensitive measurement of oxygen uptake, as a substrate is utilized,  can be
obtained with such manometric devices as a Warburg respirometer. The procedure is
based on the ideal gas law which states that at constant temperature and constant gas
volume a change in the amount of a gas can be measured by the change in its pressure.
The utilization of a substrate  by  microorganisms usually involves utilization of
oxygen and production of carbon dioxide. If the carbon dioxide is absorbed in alkali
the only change in gas volume or pressure will  be uptake of oxygen. Both the rate and
amount of oxygen uptake can be determined by this method. Measurements may be
taken at specified time intervals and a graph of  oxygen uptake vs. time can be
constructed. When the graph indicates a straight line function with time, the enzyme
systems are usually considered to be saturated with respect to substrate, although
exceptions  exist and in  some  cases higher  levels of substrate  may result in an
increased rate. Determination of the amount of oxygen in moles taken up per mole of
substrate yields information on the completeness of substrate oxidation.
  Alternatively, carbon dioxide evolution as  measured by trapping in alkali gives a
measure of the complete mineralization  of a compound supplied as the sole source of
carbon. 11 the substrate contains radioactively-labeled carbon, the liquid scintillation
counter can be used to measure carbon dioxide evolution as captured in the alkali
trap.
  Warburg respirometry  has been  used to develop the technique of simultaneous
adaptation (407) for determination  of the involvement of specific compounds in the
pathway of substrate metabolism.  Simultaneous adaptation  is based on the  theory
that cells adapted to metabolize  the primary substrate will  also be adapted to
metabolize all the intermediates of the pathway, but will not attack other substrates.
In respirometry tests, this  is seen as immediate uptake of oxygen after addition of the
substrate or its metabolites. When other  substrates are introduced, no uptake or
uptake only after a lag period is observed. The only prerequisite for this system is that
the enzymes be largely adaptive, such that they are not induced until the specific
substrate is present. Therefore,  a limitation to this test is in\ olvement of nonspecific
enzymes.
  Assays have been developed for determining the presence of specific enzyme in a
solution or culture. The assays are usually designed so thai a substrate which is acted
on  directly and  specifically by the  enzyme of interest is made available.  Enzyme
activity is monitored by measuring loss of the substrate or appearance of a product by
spectrophotometric, chemical or chromatographic methods.  These  assays are most

                                     43

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useful for enzymes which are specific for a single substrate. Some enzyme assays
require that a cell-free culture filtrate be prepared, while others work with intact cells.
  Enrichment cultures are widely used to search for organisms with degradative
ability. The substrate of interest is supplied as the growth-limiting nutrient in a
culture medium to which a mixed culture of microorganisms is added. The substrate
most commonly is the sole source of carbon, and the inoculum may include sewage
sludge, sediment samples, or river or ocean water. An inoculum is often selected from
environments  thought to be contaminated with the substrate of interest. Only
organisms with ability to degrade the substrate will be able to grow in the culture
medium and eventually will become the dominant population. These cells then can be
recovered and isolated from the other cells added in the original inoculum. Evidence
of substrate utilization is obtained indirectly by measuring growth of the population
of the degradative organism, indicating incorporation of the substrate into cellular
material. Further tests, usually with the isolated culture, are required to determine if
the substrate is completely mineralized or whether intermediate metabolites remain.
As some bacteria can grow in the absence of specifically added carbon, control
experiments must be performed to ensure that  the substrate is necessary for growth.

PARAMETERS  FOR PURE CULTURE STUDIES

  Bacteria used for pure culture studies may be selected from enrichment cultures.
Identification of these isolates permits the results of such studies to be analyzed in the
context of other such studies. Investigations of specific enzymatic or genetic features
of degradative bacteria are more easily integrated with other studies when reference
bacteria  are used. These reference bacteria represent  isolates which have been
identified and then deposited in culture libraries such as the American Type Culture
Collection (ATCC) or the National Collection of Industrial Bacteria (NCIB).
Bacteria  registered with such libraries are always indicated by a reference number
which permits other researchers to obtain the same strain for subsequent studies.
However, bacteria frequently mutate during repeated subculturing in a laboratory,
and a strain studied for a length of time may no longer resemble the original culture.
For this  reason,  the  conditions under which a strain is  maintained should be
reported. Of particular importance is the  frequency with which bacteria lose their
degradative capability when removed from the substrate of interest. Such cultures
must be maintained on media containing the substrate.
  Some general parameters for biodegradation in solution are:

  •  The concentration of the substrate is an important consideration in all studies
     of biodegradation. The capability of bacteria to degrade substrates supplied at
     trace levels may be very different from the response to high concentrations.

  •  Chemicals used  in formulating culture media should be of the highest purity
     possible, particularly when the contaminating chemical may be implicated in
     the degradative strategy of the organism.

  •  Parameters such as  pH,  temperature,  and  dissolved  oxygen should be
     monitored,  as fluctuations may affect not only the metabolic activities of the
     bacteria but also the chemical nature of the substrate.

  •  As  newer techniques of analysis become available, broad studies comprising
     many of the techniques discussed here become feasible. Knowledge of both the
     environmental degradative behavior of  the bacterial culture and its genetic
     structure can lead to manipulation of the culture toward increased degradative
     capability.
                                    44

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                              SECTION 6

     METABOLISM OF NONCHLORINATED AROMATIC

                            COMPOUNDS
CHEMISTRY OF BENZENE AND SUBSTITUTED BENZENES

  The basic chemical structure of most aromatic compounds  is the benzene ring.
Benzene (molecular formula C6H6) is  the  simplest six-membered aromatic
carbocycle. The entire molecule is structurally flat, i.e., the six carbon atoms and
attached hydrogens lie in the same plane. The six aromatic electrons are delocalized
and thereby confer the stability which is inherent to aromatic structures. The stability
of benzene refers to the availability of the aromatic electrons for bonding. When the
reactivities  of aromatic  and  aliphatic carbocycles are compared under identical
conditions, the aromatic  systems are found to be less reactive, hence more stable.
   The  naming of substituted benzenes  can  be  quite confusing  due to several
completely different sets  of nomenclature rules. For instance, there is a set of trivial
names such as toluene, phenol, and aniline. These trivial names are enigmatic to the
casual observer since there are no rules, only a historical selection. Positional isomers
have at least two nomenclature systems. The terms ortho, meta, and para have been
used to identify the position of the substituents attached to the benzene ring. Finally,
the International  Union for  Pure  and Applied Chemistry  offers a numerical
description for positional isomers (Figure 15). To show the interrelationships of these
systems, catechol, for example, is the trivial name  for 1,2-dihydroxy benzene and a
more cumbersome ortAo-hydroxy phenol.
   Positional location of substituents (ortho, meta, para or  1,2; 1,3; 1,4) is important
to the overall reactivity of an aromatic molecule. A substituent on the benzene ring
substantially influences the mode of reaction for  a given chemical  or biochemical
system, i.e., the where and how of the attack by another reactive molecule. Common
substituents attached to chlorinated aromatic molecules, available common articles
of commerce, are: the hydroxyl (-OH), amino (-NH2), methyl (-CH3), and phenyl
(-C6H5)  groups which can  render  the molecule more  reactive to electrophilic
("electron-loving")  reaction  conditions. Additional halogen substituents  serve to
deactivate the aromatic ring for electrophilic attack. Each of the cited substituent
groups has a directing influence on subsequent  electrophilic substitution which
occurs mainly at the ortho and para positions (316).
   The chemical nature and structural position of a substituent on the benzene ring
affects the mode and ease of microbial attack on the compounds. The biological
"recalcitrance"  of  a  molecule, i.e., the resistance of a  molecule to microbial
degradation, is a direct consequence of the chemical nature and structural position of
a substituent on the aromatic ring (2).  Mention throughout  this work of the
recalcitrance of a compound will refer to biological activity rather than chemical or
photooxidative processes.

MICROBIAL ATTACK ON BENZENE STRUCTURES
  The first step in oxidative microbial attack on benzene involves hydroxylation of
the ring. This is accomplished by different mechanisms in prokaryotes (bacteria) and

                                   45

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     Cl
    (6)
       OH
      (O)
CHLOROBENZENE   PHENOL
CH3
                          TOLUENE
                                        COOH
                                     BENZOIC
                                       ACID
  NH2
(Q)
  S03H
Co)
                                       ANILINE   BENZENESULFONIC
                                                    ACID
  Cl
Co]
                                  Cl
                                 [Q]
                                      Cl
     o-DICHLOROBENZENE
  1,2~DICHLOROBENZENE
                           m-DICHLOROBENZENE
                                                  p-DICHLOROBENZENE
               1,3-DICHLOROBENZENE  1,4-DICHLOROBENZENE
    NAPHTHALENE
                               ANTHRACENE
             PHENANTHRENE
Figure 15.  Common names and conventional nomenclature for substituted benzenes.
                                   46

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eukaryotes (fungi, mammals, etc.). Bacteria employ a dioxygenase which incor-
porates two atoms of oxygen from an oxygen molecule simultaneously into the ring
(Figure 16) (15, 175, 205). Molecular oxygen is a required substrate for this enzyme
(84). The resultant intermediate compound is a c;s-l,2-dihydroxy-l,2-dihydro-
benzene which loses two hydrogens to become catechol (1,2-dihydroxybenzene), a
process mediated by the enzyme  cj's-benzene glycol  dehydrogenase (14).  The
stereospecific nature ("cis") of the intermediate compound was identified in 1968
(181) and confirmed in 1970 (178) for Pseudomonas putida, and since then has been
shown to be true for all bacterial species studied (14, 176, 177, 205).
  Substituted  benzenes similarly  are  oxidized to  c/'s-dihydrodiols (substituted
catechols) by bacteria. Some species oxidize the substituent before hydroxylation of
the aromatic ring while  others attack the ring yielding a substituted catechol. For
example, toluene is attacked by P. aeruginosa with oxidation of the methyl group to
benzyl alcohol, benzaldehyde and benzoic acid,  followed by ring hydroxylation to
form catechol  (254,  331), while Achromobacter spp. and  Pseudomonas spp.,
including P. putida, hydroxylate toluene directly to form 3-methylcatechol (Figure
17) (82, 175, 176, 179, 182, 332). Other alkylbenzenes are also subject to these two
types of oxidative degradation,  either  oxidation of  the aromatic ring to form an
alkylcatechol or oxidation of the alkyl  substituent to form an aromatic carboxylic
acid which is dihydroxylated to catechol (84, 177, 177a,  181).
  Benzoic acid is metabolized by a number of different pathways, depending on the
bacterial strain. Alcaligenes  eutrophus oxidizes benzoic acid to catechol. This
mechanism in  A. eutrophus is by way  of the 3,5-cyclohexadiene-l,2-diol-l-
carboxylic acid intermediate (74, 368) catalyzed by benzoic acid 1,2-dioxygenase, a
two-protein enzyme  composed  of  NADH-cytochrome c reductase and another
protein (475). The intermediate is converted to catechol  by a single protein (Figure
18). The fluorescent pseudomonad group also oxidizes benzoic acid to catechol (410,
467). In contrast, the acidovorans pseudomonad group monohydroxylates benzoic
acid to m-hydroxybenzoic  acid and  subsequently to either gentisic  acid  (P.
acidovorans) by m-hydroxybenzoic acid 6-hydroxylase  or protocatechuic acid (P.
testosteroni) by m-hydroxybenzoic acid 4-hydroxylase (467). Other Pseudomonas
spp. monohydroxylate benzoic acid to p-hydroxybenzoic acid by utilizing benzoate
4-hydroxylase, and metabolize this intermediate further to protocatechuic acid by
the enzyme p-hydroxybenzoate 3-hydroxylase (Figure 9) (82).
  Pseudomonas PN-1 metabolizes  benzoic acid, p-hydroxybenzoic acid and m-
hydroxybenzoic acid to protocatechuic  acid aerobically.  However, under anaerobic
conditions  of nitrate respiration both protocatechuic acid and m-hydroxybenzoic
acid are metabolized through p-hydroxybenzoic acid to  benzoic acid. The mode of
attack resulting in ring cleavage has not been elucidated (430, 431).
  Members of Streptomyces spp. metabolize benzoic acid via catechol, m-hydroxy-
benzoic acid via gentisic acid, and p-hydroxybenzoic acid via protocatechuic acid
(421). Two pathways have been shown in facultatively thermophilic Bacillusspp. In
one species, benzoic acid, m-hydroxybenzoic acid and p-hydroxybenzoic acid are all
metabolized via gentisic acid (64).  The conversion  of  p-hydroxybenzoic acid to
gentisic acid by this organism requires an "NIH Shift" of the carboxyl group. In
another species, benzoic  acid is metabolized through salicylic acid to 2,3-dihydroxy-
benzoic acid (406). The latter pathway is  analogous to that shown for Azotobactersp.
(450).
  The phototrophic bacterium  Rhodopseudomonas palustris is unable to use
benzoic acid as a substrate for aerobic growth (191), although early work postulated
an aerobic  pathway via protocatechuic acid and catechol (357). The organism can
metabolize p-hydroxybenzoic acid  aerobically  via the protocatechuic acid path
(125).  Under  anaerobic photosynthetic conditions,  however, benzoic acid  is

                                     47

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                  HYPOTHETICAL
                   DIOXETANE
                                   crs-1. 2-DIHYDRO-
                                1,2-DIHYDROXYBENZENE
                                               ..._. NADH2
                                               NAD   A  X^OH
                                                        01
                                                             OH
                                                      CATECHOL
 ANILINE
                            CATECHOL
                            CATECHOL
                                     NAD   NADH"  [O
                                ^H
                                .OH

                                '^
                      cis-2,3-DIHYDRO-
                   2;3-DIHYDROXYBIPHENYL
            OH


           SOH

2,3-DIHYDROXYBIPHENYL
         Figure 16.  Oxidation of aromatic molecules by bacteria.
Adapted from References 14, 16, 63, 71a, 141, 176, 178, 183, 196, 287, 326.
                                 48

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                 BENZALDEHYDE
                                                                  OH
                                                                      OH
          cis-TOLUENE
          DIHYDRODIOL
                                                                      OH

                                                          3-METHYLCATECHOL
Figure 17.  Pathways for the bacterial metabolism of toluene. Pathway (a)P. putida
    mt-2; (b) P. aeruginosa; (c) P. putida; Pseudomonas sp.; Achromobacter sp.
            Adapted from References (a) 471 a; (b) 254; (c) 92, 179,
                                    49

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    5    3
    I'D «
    o  w  c
    sT  c  3
    . CD  ^
     & 3
If
•a  o
       3  T)
       m  a
      . S-  3-
      .•3  §
f  f°^«
1  s  is °
I  IIIf
      5' 3
      .—. 01
     i-
w  x  
   O OJ 3 (b
ssut
•   1^11
   w Q. — C
    t» 7 H,
    s   1
                                 COOH
                                                   COOH
H
S
NAD+
DIENE-
a ^
X^
NADH
a-m
COOH
IL-OH
rDROXY-
OH
a ^fnT01"
^+(0)
C02
CATECHOL
                   /-
                 NADH NAD+
                                          1.2-DIOL-1-             /3-KETOCARBOXYLIC
                                          CARBOXYLIC ACID          ACID
0
                 COOH
                                    COOH
                        X. b
                        S ^>V
                                          O,
                                                       COOH
                                                                         COOH
              BENZOICNADP NADPH
                ACID
                NADPH  NADP
             OH
     4-HYDROXYBENZOIC
            ACID
PROTOCATECHUIC
      ACID
 /3-CARBOXY-
c/s,c/s-MUCONIC
     ACID
                                                                                            COOH
/3-KETOADIPIC
    ACID
                            CYCLOHEX-1-ENE-
                           1-CARBOXYLIC ACID
                                                2-HYDROXY-
                                               CYCLOHEXANE
                                              CARBOXYLIC ACID
                                                       2-KETO-
                                                     CYCLOHEXANE
                                                   CARBOXYLIC ACID
                                                                                           PIMELIC ACID

-------
metabolized reductively topimelicacid viacyclohexanol(Figure 18)(124,125,191).
Early decarboxylation does not occur. The enzymes involved are thought to be
reductases such as ferredoxin coupled to the light-induced electron transport system

  Rhodococcus sp. strain AN-117  utilizes aniline as a sole source of carbon and
energy and metabolizes it exclusively by conversion to and ortho cleavage of catechol
by inducible enzymes (235). In contrast, strain SB3, thought to be a pseudomonad,
utilizes a constitutive  meta cleavage pyrocatechase and hydroxymuconic semial-
dehyde dehydrogenase. However, aniline degradation occurs only when the cells are
grown on aniline,  indicating the presence of another inducible enzyme. Aniline-
grown resting cells of Frateuria sp. ANA-18 oxidize aniline without a lag and oxidize
catechol at a faster rate than aniline (9). Metabolites resulting from incubation of a
cell-free extract with aniline include cis, c/s-muconic acid, beta-ketoadipic acid and
ammonia.
  A mutant strain of a Nocardia sp. has been shown to convert aniline to catechol via
simultaneous dioxygenation (16). This pathway is partially corroborated by data
indicating that a Pseudomonas sp. grown on aniline oxidizes catechol rapidly,
4-aminophenol moderately quickly, 2-aminophenol slowly, and phenol not at all
(449). However, only half  the  ammonia theoretically  expected  from  the  direct
formation of catechol from aniline was recovered. This discrepancy has not been
resolved. P. multivoransstrain AN1 growing on aniline was simultaneously adapted
to oxidize catechol but not phenol or 2-aminophenol (199). Transient formation of a
catechol was noted indicating replacement of the amine group with a hydroxyl.
  Biphenyl (a benzene-substituted benzene) is dihydroxylated by Beijerinckia sp., P.
putida, and an unidentified Gram-negative bacterium to c;s-2,3-dihydro-2,3-dihy-
droxybiphenyl and subsequently to 2,3-dihydroxybiphenyl (Figure 19) (183, 287).
Phenol is oxidized directly to catechol via phenol hydroxylase(63,141,196,326).
  Polynuclear aromatic hydrocarbons  are made up of fused aromatic  rings. The
three simplest compounds, naphthalene,  anthracene,  and  phenanthrene,  are
metabolized by bacteria to  form c/s-dihydrodiols by the same mechanism as that
shown for benzene (Figure 20) (72, 227). A mutant strain of Beijerinckia sp. (strain
B836), as well as P. putida strain  199, forms (+)-cj's-l,2,-dihydroxy-l,2-dihydro-
anthracene from anthracene (226). These two organisms also convert phenanthrene to
(+)-cj's-3,4,-dihydroxy-3,4-dihydrophenanthrene. A minor product formed is cis-l ,2-
dihydroxy-l,2-dihydrophenanthrene and there were no other  diols  recovered
during these experiments.  Pseudomonads  also oxidize phenanthrene  to  3,4-
dihydroxyphenanthrene (Figure 21) and then to 1,2-dihydroxynaphthalene which is
metabolized via the naphthalene pathway (Figure 22) (145, 267, 282, 373).
  Naphthalene is  metabolized by  pseudomonads  through  c/s-l,2-dihydro- 1,2-
dihydroxynaphthalene, 1,2-dihydroxynaphthalene, salicylaldehyde, salicylic acid,
and catechol (106). The enzyme which catalyzes the conversion of the 1,2-dihydro-
1,2-dihydroxynaphthalene to 1,2-dihydroxynaphthalene is c/s-naphthalenedihydro-
diol dehydrogenase (224).  Its  proposed mechanism  is stepwise, the first step
bacterial-enzyme catalyzed and the second step a nonenzymatic enolization (224).
  Anthracene is oxidized to 1,2-dihydroxyanthracene by a  naphthalene-grown
Pseudomonas sp.  (151, 374).  This intermediate is metabolized  further to  2,3-
dihydroxynaphthalene which follows an unknown degradative pathway through
salicylic acid (Figure 23) (145).
  In all of the above examples with the exception of phenol, bacterial attack on the
benzene ring proceeds  via a dioxygenase with formation of a c/s-dihydroxybenzene.
This mechanism is consistent with almost all bacterial oxidations of all substituted
benzenes studied to date. The phenol hydroxylase is a monooxygenase but results in
an ort/jodihydroxylated molecule.

                                     51

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                                          COOH
                                         BENZOICACID
                                       CH2  COOH    CH3  COOH
                         _0        2-HYDROXYPENTA-
                          COOH  2,4-DIENOIC ACID
                            OH
     2-HYDROXY-6-OXO-6-PHENYLHEXA-
                  2,4-DIENOIC ACID
                                          4-HYDROXY-2-OXOVALERIC ACID
                 "OH  \b
BIPHENYL
    2,3-DIHYDROXYBIPHENYL
                                                  COOH
                                         PHENYLPYRUVIC ACID
       2-HYDROXY-3-PHENYLMUCONIC SEMIALDEHYDE
  COOH
CHO
      Figure 19.  Metabolism of biphenyl by (a) P. putida and (b) Beijerinckia sp.
             Adapted from References (a)70, 71; (b)183, 287; (c)287.
                                   52

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              cis-1,2-DIHYDRO-1,2-
            DIHYDROXYNAPHTHALENE
                       1,2-DIHYDROXYNAPHTHALENE
                                      1,2-DIHYDROXY ANTHRACENE
  cis-1,2-DIHYDRO-1,2-
DIHYDROXY ANTHRACENE
               cis-3,4-DIHYDRO-3,4-    3,4-DIHYDROXYPHENANTHRENE
            DIHYDROXYPHENANTHRENE
Figure 20.  Mechanism of bacterial attack on naphthalene, anthracene, and
                         phenanthrene.
                Adapted from References 145, 224.
                             53

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                          H0?%
 PHENANTHRENE      cis-3,4-DIHYDRO-3,
               4-DIHYDROXYPHENANTHRENE
                                3,4-DIHYDROXYPHENANTHRENE
                                                OH
                                          HOOO
 pyrilium cation
 in strongly
 acidic solutions
hemiacetal form
in neutral and weakly
acidic solutions
     cis-4-(1 -HYDROXY-
      NAPHTH-2-YU-2-
   OXOBUT- 3-ENOIC ACID,
    unstable anion in alkali

              i
              '— **
CHO
                                                        7,8-BENZOCOUMARIN
 1-HYDROXY-2-
 NAPHTHOIC ACID
   1-HYDROXY-2-      CH3-C— COOH  hypothetical intermediate
   NAPHTHALDEHYDE      Q
                     PYRUVIC ACID
                        naphthalene degradation
1,2-DIHYDROXYNAPHTHALENE
 Figure 21.  Pathway of phenanthrene metabolism by Pseudomonas sp. Dashed lines
                       indicate hypothetical pathways.
                        Adapted from Reference 145.
                                     54

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                                                  OH
    NAPHTHALENE
                            cjs-1.2-DIHYDRO-1.2-       1,2-NAPHTHOQUINONE
                          DIHYDROXYNAPHTHALENE        (nonbiological)
                                     1,2-DIHYDROXYNAPHTHALENE

                   1,2-dihydroxynaphthalene oxygenase
                                            quinoid form

                         nonbiological transformation I
   2-CARBOXYBENZO-
   PYRILIUM
   cation in strongly acidic solutions
            cis-o-HYDROXYBENZALPYRUVIC ACID
        hemiacetal in neutral or weakly acidic solutions
                              salicylate
                              hydroxylase
                                                  -°T-T(oj
                                         COUMARALDEHYDE COUMARIN
                                         anionic form in alkali
            caiechol2,3-
            oxygenase
                                       •-COOH
2-HYDROXYMUCONIC  CATEtHOL            SALICYLALDEHYDE
SEMIALDEHYDE                  SALICYLIC
                                  ACID
                                                 salicylaldehyde
                                                 dehydrogenase
                                       "OH
                          °.. COOH
                                                                 OH
    T
CH3C-COOH
    0
PYRUVIC ACID
                                                         4-HYDROXY-4-0-
                                                        HYDROXYPHENYL-
                                                       2-OXOBUTYRICACID
Figure 22.  Pathway of naphthalene metabolism by Pseudomonas spp. Dashed lines
                       indicate hypothetical pathways.
                        Adapted from Reference 106.
                                    55

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                  cis-1,2-DIHYDRO-1,2-
                DIHYDROXYANTHRACENE
                                       1,2-DIHYDROXYANTHRACENE
                COOH

               '
  pyrium cation in  ""  hemiacelal form in  cis-4-(2-HYDROXY-  6,7-BENZOCOUMARIN
   strongly acidic     neutral and weakly   NAPHTH-3-YL)-2-
     solutions         acidic solutions  OXOBUT-3- ENOIC ACID
                                        anion in alkali
                OH
       Colo
             ""COOH
      2-HYDROXY-3-
    NAPHTHOIC ACID  NAPHTHALEDHYDE
                                 CH3C-COOH
OH
                               PYRUVIC ACID

                              	^- degradation through salicylate
2,3-DIHYDROXYNAPHTHALENE
     Figure 23.  Pathway of anthracene metabolism by Pseudomonas spp. Dashed lines
                            indicate hypothetical pathways.
                            Adapted from Reference 145.
                                       56

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ATTACK ON AROMATIC STRUCTURES BY CYANOBACTERIA
  The cyanobacteria have not been studied as extensively with regard to aromatic
degradation. An investigation of the degradation of naphthalene by the cyano-
bacterium  Oscillatoria sp.  revealed  the  presence of  1-naphthol as the major
metabolite (Figure 24); 57% of the 1-naphthol formed involved the incorporation of
molecular oxygen (76a, 78a). Small amounts  of CJS-l,2-dihydroxy-l,2-dihydro-
naphthalene were recovered which readily dehydrated to form 1-naphthol. Trans-
l,2-dihydroxy-l,2-dihydronaphthalene was not recovered. There are three possible
pathways which suggest a mechanism for attack of cyanobacteria on  the aromatic
ring: (a)  similar to that for mammalian systems (path I, Figure 24), (b) unique to
photosynthetic organisms (path II), or (c) similar to that for heterotrophic bacteria
(path III) (78a).

RING FISSION OF DIHYDROXY AROMATIC COMPOUNDS BY
BACTERIA

   Bacteria can open an aromatic ring containing two hydroxyl groups if the groups
are located ortho (adjacent) or para (opposite) to each other. If the groups are ortho,
ring cleavage can occur either between the two (ortho cleavage) or next to one group
(meta cleavage). The choice of which pathway is induced depends partly on the
substrate and  partly  on the genetic constitution of the particular bacterial species.
Both pathways can be induced independently of each other. An organism may utilize
one pathway preferentially although it contains the enzymes for both pathways. For
example, P. putida  (arvilla) mt-2 metabolizes catechol via  the  meta  pathway,
although it contains the enzymes for both the meta and  ortho pathways (323).
   The ortho cleavage pathway of catechol leads to formation of 3-ketoadipic acid
(Figure 25). Substituted catechols can be metabolized by an analogous path until
either the substituent is expelled or the compound formed along the pathway cannot
be metabolized further. Thus, the ortho pathway for protocatechuic acid dissimi-
lation  converges  with  that  of catechol at 3-ketoadipate enol-lactone  after the
carboxyl group  is expelled  (Figure 26) (66, 84, 104).  This latter compound  is
converted to 3-ketoadipic acid, which picks up coenzyme-A from succinyl-CoA to
form the intermediate 3-ketoadipyl-CoA. Cleavage of this compound  results in
acetyl-CoA and succinyl-CoA which in turn  exchanges its coenzyme-A  with 3-
ketoadipyl-CoA  leaving  one molecule  of succinic acid (408). These aliphatic
molecules enter the cell's tricarboxylic acid cycle.
   The meta cleavage  pathway (Figure 25) results in the formation of pyruvic acid and
acetaldehyde from catechol (102, 103, 180, 330). Substituted catechols usually form
pyruvic acid and  another aldehyde or an acid (Figure 26). The substrate 3-m_ethyl_
catechol yields acetate while 4-methyl catechol yields formic acid (27).  The non-
fluorescent pseudomonad group metabolizes protocatechuate by the meta pathway,
utilizing protocatechuate 4,5-oxygenase with subsequent production of oxaloacetic
acid and pyruvic acid. Meta cleavage of protocatechuic acid  in Bacillus spp.  is
catalyzed by protocatechuate 2,3-oxygenase and  yields pyruvic acid and acetal-
dehyde via the 2-hydroxymuconic semialdehyde intermediate (95). Meta cleavage
degradation of catechol results in two possible routes of metabolism as demonstrated
in Azotobactersp. and P. putida NCIB 10015 (382,383), the major route involving an
NAD* dependent dehydrogenase and resulting in formation of 4-oxalocrotonic acid
and the minor route not requiring NAD' but rather employing a hydrolase to form
2-oxopent-4-enoic acid directly. The paths converge at this step. Degradation of
3-methylcatechol  is constrained to follow the hydrolase pathway only, as the NAD*
dependent path requires the presence of an aldehyde group on the molecule. Other

                                     57

-------
              NAPHTHALENE 1,2-OXIDE
 [OJOJ  -
NAPHTHALENE
1-NAPHTHOL
            H   OH
4-HYDROXY-1 -TETRALONE
     cis-1,2-DIHYDRO-l ,2-DIHYDROXYNAPHTHALENE
   Figure 24.  Pathways for naphthalene metabolism by Oscillatoria sp., strain JCM.
  (I) Metabolism via 1,2-oxide; (II) light-dependent direct hydroxylation of naphthalene;
                        (III) metabolism via dihydrodiol.
                         Adapted from Reference 78a.
                                     58

-------
           ORTHO
          CATECHOL
                ,c-o
                c-o
                  OH
     cis, cis-MUCONIC ACID
                                               META
                                   2-HYDROXYMUCONIC SEMIALDEHYDE
                                      x©
      +1-MUCONOLACTONE

              '
                  -OH
                            4-OXALOCROTONIC ACID
 3-KETOADIPATE ENOL-LACTONE

              T   nH
                  OH
       3-KETOADIPIC ACID
             ^OH
  3-KETOADIPYL CoA
          {  COOH
    CH3      CH2
    C=0   +  CH2
    SCoA    COOH
ACETYL-CoA  SUCCINIC ACID
                             2-OXOPENT-4-ENOIC ACID (ENOL FORM)



                                           CH3 cf
                                     4-HYDROXY-2-OXOVALERIC ACID
                                              CH3
                                            HC=0

                                     ACETALDEHYDE  PYRUVIC ACID
COOH
C=0
CH3
Figure 25.  Ortho- and mefa-cleavage pathways of catechol metabolism by bacteria.
   Adapted from References (1) 27, 103; (2) 382, 382a, 383, 468a; (3) 340, 408.
                                  59

-------
          QRTHO

           ,XX^OH
          IP!
     HOOCy^OH
   PROTOCATECHUIC ACID
            T   nn
                                            META
                                        HOOC'
                                      PROTOCATECHUIC ACID
3-CARBOXY-cjS.cis-MUCONIC ACID
                                 2-HYDROXY-4-CARBOXYMUCONIC
                                         SEMIALDEHYDE
                                             H     ^
     HOOC
 4-CARBOXYMUCONOLACTONE
              c-o
                              2-HYDROXY-   2-HYDROXY-4-     2-0X0-4-
                               MUCONIC CARBOXYMUCONIC CARBOXYPENT-
                             SEMIAL.DEHYDE    A9ID        4-ENOIC ACID

                                                                 *0
              —OH
S-KETOADIPATE'ENOL-LACTONE
                                           HO   0
      3-KETOADIPIC ACID
             T   oui
                          2-HYDROXYMUCONIC
                                 ACID

                                .,, T ^0
                                       2-OXO-4-HYDROXY-
                                    4-CARBOXYMUCONIC ACID
                                                     2-OXO-4-HYDROXY-
                                                  CARBOXYPENTANOIC ACID
      3-KETOADIPYL-CoA
                                             COOH
                                             C=0
                                             COOH
                                  |       OXALO ACETIC
                             4-HYDROXY-     ACID
                          2-KETOVALERIC ACID
       ?H3
       C=0
                   COOH
                   CH,
   ACETYL-CoA
                   COOH
                SUCCINIC ACID
      CH3
     HC=0
ACETALDEHYDE
COOH
C=0
                                          PYRUVIC ACID
 Figure 26.  Ortho- and mefa-cleavage pathways of protocathechuate metabolism by
                  bacteria. (1) Bacillus sp., (II) P. testosteroni.
                  Adapted from References 66a, 95, 340, 408.
                                    60

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compounds  which undergo  meta cleavage include naphthalene, 2,3-dihydroxy-
phenylpropionic acid, 2,3-dihydroxybenzoic acid, and homoprotocatechuic acid
(84, 106).
  Compounds that contain two hydroxyl  groups located opposite to each other
(para substitution), such as homogentisic acid and gentisic acid, are usually cleaved
at the bond between one hydroxyl and the  adjacent side chain, leading to fumaric
acid and acetoacetic acid or fumaric acid and pyruvic acid, respectively (Figure 27)
(84).
  Metabolism of benzoic acid by P. testosteroni and P. acidovorans follow divergent
pathways  with  a different hydroxylase  being induced in  each one (Figure 28),
resulting in meta and para cleavage pathways, respectively. P. testosteroni produces
two moles of pyruvic acid and one of formic acid, while P. acidovorans induces the
gentisic acid pathway and produces one mole each of fumaric acid and pyruvic acid
(467).
  Biphenyl hydroxylated at the carbon-2 and carbon-3 positions may be cleaved in
the meta position in two ways. An unidentified Gram-negative organism and a
mutant strain of Beijerinckia sp. metabolize this compound by cleavage between the
carbon-3 and carbon-4 positions to yield 2-hydroxy-3-phenylmuconic semialdehyde
and subsequently phenylpyruvic acid (183, 287). However,  P putida cleaves 2,3-
dihydroxybiphenyl between the carbon-1 and carbon-2 positions to form 2-hydroxy-
6-oxo-6-phenylhexa-2,4-dienoic acid and then benzoic acid (70, 71).
  The dihydroxy  fused ring compounds are cleaved initially by meta cleavage
between the angular carbon and the adjacent hydroxyl. Hydroxynaphthalene  is
cleaved to form c/s-o-hydroxybenzalpyruvic acid and subsequently salicylaldehyde,
salicylic acid, and catechol (Figure 13). Catechol may be degraded by ortho or meta
cleavage (106, 224, 255,267, 353). In some bacteria, phenanthrene is dihydroxylated
to ds-4-(l-hydroxy-naphth-2-yl)-2-oxobut-3-enoic  acid, 1-hydro xy-2-naphthalde-
hyde, l-hydroxy-2-naphthoic acid, and 1,2-dihydroxynaphthalene (Figure 12) (145,
151, 267, 373). Subsequent steps follow the pathway through salicylic acid. Other
bacteria, including fluorescent and nonfluorescent pseudomonad groups, vibrios,
and Aeromonas spp., metabolize phenanthrene to l-hydroxy-2-naphthoic acid,
2-carboxybenzaldehyde, o-phthalic acid,  and protocatechuic  acid,  which may
undergo ortho  or meta cleavage  (255, 256). The  first ring cleavage product of
1,2-dihydroxyanthracene is  cj's-4-(2-hydroxynaphth-3-yl)-2-oxobut-3-enoic acid
which is degraded to 2-hydroxy-3-naphthaldehyde, 2-hydroxy-3-naphthoic acid and
2,3-dihydroxy-naphthalene (Figure 14) (145, 151, 267, 374). Degradation of 2,3-
dihydroxynaphthalene continues  through salicylic acid by  an unknown pathway
(145).

ATTACK OF AROMATIC STRUCTURES  BY
EUKARYOTES
  The fate of aromatic-organic substances in mammalian systems has been studied
extensively, both in vivo (injecting animals directly and recovering metabolites in
body fluids or tissues) and using extracts  of liver (or other) cells which contain the
enzymes active  in degradation of compounds (the  microsomal  enzymes). The
mechanisms by  which fungi  and yeasts degrade  aromatic compounds have been
shown  to be analogous to that of mammalian systems (404).
  In contrast to bacteria, fungi utilize a  monooxygenase which incorporates one
atom of molecular oxygen into the benzene ring while converting the other to water.
The resulting intermediate is an epoxide, which undergoes hydration with water to
form a rrans-l,2-dihydroxy-l,2-dihydro  intermediate  and  subsequently a  trans-
dihydroxy compound (Figure 29) (175). Alternatively, the epoxide can isomerize to
form phenols (177). Both of these mechanisms operate in the attack on naphthalene

                                     61

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         OH
             COOH
         OH
   GENTISIC ACID
                                                  OH
                                                       CH,COOH
           OH
  HOMOGENTISIC ACID
        COOH COOH
        0
MALEYLPYRUVIC ACID
 HOOC
              COOH
    PYRUVIC ACID
             g<^,
             -0

           0
MALEYLACETOACETIC ACID


                CH2COOH
                                           HOOC

                                       FUMARYLACETOACETIC ACID
L_r i M w v i\^ r^i^i i_
COOH
C=0
CH3


111 " COOH "*
CH
CH
COOH
^
cc
F
OH
CH2
9=o
CH3
                         FUMARIC ACID
                                           ACETOACETIC ACID
Figure 27.  Pathways of gentisic acid and homogentisic acid metabolism by bacteria.
                 Adapted from References 84, 84a, 266a.
                                62

-------
                                    COOH
                                  (o
                              BENZOIC ACID
                        Benzoate
                        3-hydroxylase
                                         £-HYDROXYBENZOIC ACID
                                     fHydroxybenzoate  |
                                     •hydroxylase    1»

COOH   m-Hydroxybenzoate  COOH  m-Hydroxybenzoate  COOH
        6-hydroxylase
                                           4-hydroxylase
    HOOC
    FUMARIC ACID
           +
     PYRUVIC ACID
                         m-HYDROXYBENZOIC ACID
                                                    PROTOCATECHUIC ACID

                                                Protocatechuate
                                                4,5-oxygenase
                                                OHC    .
                                                 HOOC^OH
                                                    It

                                               2 PYRUVIC ACID

                                                FORMIC ACID
Figure 28.  Divergent pathways for the metabolism of benzoic acid, p-hydroxybenzoic acid, and
 m-hydroxybenzoic acid by P. testosteroni(so\\d arrows) and P. acidovorans(broken arrows).
                           Adapted from Reference 467.
                                      63

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  BENZENE   BENZENE 1,2-OXIDE                    CATECHOL

               trans-1,2-DIHYDRO-1,2-DIHYDROXYBENZENE

    Figure 29.  Formation of catechol from benzene in fungi, yeasts, and mammals.

by Cunninghamella elegans in which the primary metabolite is 1-naphthol (Figure
30) (76). Anthracene is oxidized by C. elegans predominantly to trans-1,2-dihydroxy-
1,2-dihydroanthracene with formation of 1-anthryl sulfate (sulfate conjugation of
1-anthrol) as well (73). Other unidentified metabolites are also produced.  Other
compounds which have a hydroxyl group added during fungal metabolism include
acetanilide, aniline, anisole, benzene, benzoic acid, biphenyl, and toluene (404). In
some cases the position of the hydroxyl is variable or more than one hydroxyl group
is added  to the  ring.  For example,  C.  elegans hydroxylates biphenyl  to 2-
dihydroxybiphenyl,  3-hydroxybiphenyl,  4-hydroxybiphenyl,  2,4'-dihydroxybi-
phenyl, and 4,4'-dihydroxybiphenyl (116).

DEGRADATION OF DIHYDROXYLATED AROMATIC

COMPOUNDS BY YEASTS AND FUNGI
  In general, fungi and  yeasts lack  many  of  the  ring fission  dioxygenases
characteristic of bacteria  (5, 274).  In most  fungi  and yeasts, catechol and
hydroxyquinol are cleaved only by the ortho mechanism, utilizing 1,2-dioxygenases
only (5, 327, 395). Phenol is metabolized through catechol by the ortho pathway
(395). Aspergillus niger converts benzoic acid to benzaldehyde (359). A single strain
of Penicillium sp. only  has been found  to utilize the  mera-fission pathway (67).
However, certain catabolic enzymes of the yeasts have broader substrate specificities
than the equivalent bacterial enzymes.  In addition, a third  hydroxyl group can be
introduced into the  aromatic  ring (5).  Thus, although the  yeast Trichosporon
cutaneum  lacks dioxygenases for protocatechuic acid, gentisic acid, and homopro-
tocatechuic acid, it can metabolize these substrates by means of NADH-dependent
hydroxylases (Figure 31).
  Methoxylated  aromatic  compounds are demethylated  and converted  to  the
corresponding hydroxybenzoic acids by microfungi (42) but  are reduced to their
corresponding aldehydes or alcohols by the wood-rotting basidiomycete Polystictus
versicolor(\47). The metabolism of protocatechuic acid through 3-carboxymuconic
acid and 3-carboxymuconolactone to 3-ketoadipate by Neurospora crassa is typical
of many fungi (67,188). Protocatechuic acid is formed from p-hydroxybenzoic acid
or p-methoxybenzoic acid. The protocatechuic acid 3,4-oxidase of Rhodotorula
mucilaginosa was used  to identify the first metabolite of  protocatechuic acid as
3-carboxy-cis, c/s-muconic acid (67). Fungi contain a lactonizing enzyme  which
converts this compound to 3-carboxymuconolactone. This is in contrast to bacteria,
which  form  4-carboxymuconolactone (93). The  product  of 3-carboxymucono-
lactone degradation is 3-ketoadipic acid.
  Some groups of fungi form catechol from protocatechuic acid (67) via catechol
1,2-oxygenase. Further degradation leads to cys,CK-muconic acid and (+)-mucono-
lactone via  a  ds,c/s-muconic acid-lactonizing enzyme and a  muconolactone.
Eventually 3-ketoadipic acid is formed and subsequently metabolized to succinic
acid and acetyl-CoA. The catechol pathway is similar to that of bacteria (67). Fungal

                                    64

-------
        (ojgj
      NAPHTHALENE
  NAPHTHALENE 1,2-OXIDE
trans-1,2-DIHYDRO-
1,2-DIHYDROXYNAPHTHALENE
        [oioj
 1,2-DIHYDROXYNAPHTHALENE
   1,2-NAPHTHOQUINONE
1,4-DIHYDROXYNAPHTHALENE

             Q

        [oT


  1,4-NAPHTHOQUINONE
        Figure 30.  Pathway of naphthalene metabolism by C. elegans.
                    Adapted from Reference 76.
                           65

-------
(o)
:oic

 I
  CH2COOH

Co]
                                                           CH2COOH
                                                          to)
                         BENZOIC ACID         OH       PHENYLACETIC ACID
                                 4-HYDROXYPHENYLACETIC ACID
  \
  CH2COOH

fol
                                                           CH2COOH
                     4-HYDROXYBENZOICACID    ?   3-HYDROXYPHENYLACETIC
                                  HOMOPROTOCATECHUIC ACID  V


                                                           CH2COOH

                                                          (o)

                     PROTOCATECHUIC ACID     i       HOMOGENTISIC ACID
          OOH
                       c°2
    GENTISIC ACID
                              OH
                        HYDROXYQUINOL
                                         HO
                                                       HO
                                                             H2COOH

                                                             °
                                                             COOH
                       MALEYLACETOACETIC
                              ACID
COOH
                COOH

              Co
   ACID
\
      CATECHOL
jn
^OH
VOH
1OXY-
ACID

JkCOQH
^COOH
MALEYLACETIC
ACID
1
T
O""OH 	 *X*S
:OOH [ COOH
C°°H
lUCDNIC
ACID

3-KETOADIPIC
ACID
KREBS rvri F -*-
COOH
T °
CH2
COOH
OXALOACETIC
ACID
COOH -*~~
CH2
CH3
ACETOACETIC
ACID
)
>0
HO'^Xs*^ COOH
MALEYLPYRUVIC
ACMCl
MUIU
A
^^
COOH
i
COOH
FUMARIC ACID
I
J
     Figure 31.  Metabolism of aromatic compounds by 7". cutaneum.
                     Adapted from Reference 5.
                               66

-------
metabolism  of 1,2-dihydroxynaphthalene or  1-naphthol leads to 1,2-naphtho-
quinone or 1,4-naphthoquinone, respectively (75,76,78). These pathways are similar
to those of mammalian microsomal extracts and are due to the cytochrome P-450
present in some fungi as well as in mammals (75). Monohydroxylated biphenyl
compounds are hydroxylated further to various dihydroxy compounds (184a). These
transformations are similar to those of mammals (184a, 404).

SUMMARY

  Bacteria and eukaryotes differ fundamentally in the mechanism  of primary
oxidation of aromatic compounds. Bacteria usually add two atoms of molecular
oxygen from the same atmospheric oxygen molecule using a dioxygenase enzyme.
The mechanism of the oxidative addition results  in a c;'s-l,2-dihydrodiol inter-
mediate. In a few cases, such as when the aromatic ring is already monooxygenated
(as for phenol), a hydroxylase  (a monooxygenase enzyme) is utilized. The ortho-
dihydrodiol  molecules are subject to ring cleavage by either of two mechanisms.
Ortho cleavage enzymes such as catechol 1,2-oxygenase open the ring between the
adjacent hydroxyl groups. The molecule subsequently is metabolized via cis, cis-
muconic acid and 3-ketoadipic acid to acetyl-Co A and succinic acid. In contrast, the
meta cleavage pathway opens the ring adjacent to one hydroxyl group using enzymes
such  as  catechol  2,3-oxygenase. The intermediate compounds of this pathway
include 3-hydroxymuconic acid, and the end products include pyruvic acid and an
aldehyde. Compounds such as benzoic acid initially may be monohydroxylated in
the meta position. A second hydroxylase may then form a para-dihydroxylated
molecule which is  metabolized via the gentisic acid pathway to fumaric acid and
pyruvic acid. An alternative pathway of benzoic acid dissimilation attributable to
bacteria  is demonstrated in anaerobic reductive metabolism by R. palustris which
forms pimelic acid via the cyclohexanol intermediate.
  Fused-ring compounds are ort/jo-dihydroxylated at the positions adjacent to the
angular carbon, and are cleaved by the meta pathway.
  Cyanobacteria appear to attack aromatic  structures by a mechanism similar to
that of heterotrophic bacteria, which results in a c/s-dihydrodiol intermediate.
However, the exact mechanism has not been elucidated. Fungi and other eukaryotes
attack benzene structures by monooxygenases which incorporate one atom of
molecular oxygen, forming an epoxide.  Subsequently the epoxide may undergo
hydration with water to form a tra.ns-l,2-dihydrodiol intermediate. Alternatively, a
monohydroxylated compound may be formed.
  The ortho mechanism is most commonly  utilized by fungi for ring fission. The
protocatechuic acid pathway in fungi differs from that of bacteria in that 3-carboxy-
c/s,c/s-muconic acid is lactonized to the 3-lactone in fungi and the 4-lactone in
bacteria. Fungi lack many of the substrate-specific dioxygenases characteristic of
bacteria, but some of their catabolic enzymes  have broader substrate specificities
than the  equivalent bacterial enzymes. A third hydroxyl group may be added to the
ring by fungi to facilitate metabolism of a molecule. The similarity in metabolism by
fungi and mammals of many aromatic compounds  has been demonstrated by
comparison  of the resultant metabolites (Table 3) (404). Thus, fungi may serve as
models for many mammalian metabolic mechanisms.
                                     67

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       TABLE 3. COMPARISON OF METABOLITES FORMED BY
            EUKARYOTES AND MAMMALIAN SYSTEMS*
                                    Mammalian metabolites
Substrate
Aniline
Fungal metabolites In vitro
Acetanilide, 4-Hydroxy-
2-hydroxy- aniline
aetanilide, and
4-hydroxyaniline
In vivo
Acetanilide,
2-, 3-, and 4-
hydroxyaniline
Anisole
2- and 4-
 Hydroxy
 anisole,
 phenol
2- and 4-
 Hydroxy
 anisole,
 phenol
2- and 4-
 Hydroxy
 anisole
Benzene
Benzoic acid
Biphenyl
Chlorobenzene
Naphthalene
Toluene
Phenol
2- and 4-
Hydroxy-
benzoic acid,
3,4-dihydroxy-
benzoic acid
2- and 4-
Hydroxy-
biphenyl, 4,4'-
dihydroxybiphenyl
2- and 4-
Hydroxy-
chlorobenzene
1-and2-
Hydroxy-
naphthalene
2- and 4-
Hydroxy
toluene
Phenol
3-Hydroxy-
benzoic acid
2- and 4-
Hydroxy-
biphenyl
2-, 3-, and 4-
Hydroxychloro-
benzene
1-and2-
Hydroxy-
naphthalene
2- and 4-
Hydroxy
toluene, benzyl
alcohol
Phenol
2-, 3-, and 4-
hydroxybenzoic
acid
4-Hydroxy-,
3,4-dihydroxy-
and 4,4'-dihydroxy
biphenyl
2-, 3-, and 4-
Hydroxy-
benzene
1-and2-
Hydroxy-
naphthalene
Benzoic acid and
conjugates

 * Adapted from reference 404.
                               68

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                              SECTION 7

                     CHLOROBENZOIC ACIDS
  Chlorobenzoic acids are introduced into the environment as degradative products
of polychlorinated biphenyls (1, 334a) and herbicides as well as in direct application
as herbicides (41). For example, the  multisubstituted  herbicide 2,3,6-trichloro-
benzoic acid is a growth regulator similar in function to 2,4-dichlorophenoxyacetic
acid (208).

BACTERIAL METABOLISM AND CHLOROBENZOIC ACIDS
  The degradation of 2,3,6-trichlorobenzoic acid (Figure 32)  has been investigated
using resting cell suspensions of Brevibacterium sp. grown on benzoic acid (208,210).
The major resulting product is 3,5-dichlorocatechol which appears with stoichio-
metric release of one mole of chloride and one mole of CO2 per mole of herbicide
metabolized (208). The initial oxidation of 2,3,6-trichlorobenzoic acid takes place at
the unsubstituted carbon-4 position. This is followed by a one-step oxidation-
dechlorination at the adjacent chlorinated carbon.  The pathway thus proceeds
through 2,3,6-trichloro-4-hydroxybenzoic acid to 2,3,5-trichlorophenol and subse-
quently to 3,5-dichlorocatechol (Figure 32). The dichlorocatechol accumulates in the
medium  and  is toxic  to the Brevibacterium sp. cells.  However, resting cell
suspensions of Achromobacter sp. grown on benzoic acid will cleave 3,5-dichloro-
catechol by the meta pathway to form 2-hydroxy-3,5-dichloromuconic semialdehyde
(210). This metabolite accumulates and is toxic to the Achromobacter sp. cells.
  Several species of bacteria have  been shown to metabolize 3-chloro- and 4-
chlorobenzoic  acid. Resting cell suspensions of Arthrobacter sp. grown on  benzoic
acid  oxidize 3-chlorobenzoic acid to 4-chlorocatechol, which is not inhibitory to
growth or oxygen  uptake (210, 211). Pseudomonas aeruginosa strain  B23
accumulates 3-chlorocatechol from the metabolism of 3-chlorobenzoic acid (216).
Acinetobacter calcoaceticus strain Bs5 grown on succinic acid or pyruvic acid will
cometabolize 3-chlorobenzoic acid to both 3-chloro- and 4-chlorocatechol, which
accumulate and are toxic (362). When mixtures of chlorocatechols can be formed,
3-chlorocatechol is the major metabolite (261, 362).  Meta cleavage of 3-chloro-
catechol results in an acylhalide which acts as an acylating agent and inactivates the
meta pyrocatechase (catechol-cleaving enzyme) irreversibly, resulting in the lethal
accumulation of catechols (261). Inefficient ortho cleavage will also result in the
accumulation of chlorocatechols. Meta cleavage of 4-chlorocatechol yields 5-chloro-
2-hydroxymuconic semialdehyde. Corresponding chlorocatechols are also formed
from 3-chlorobenzoic acid and 4-chlorobenzoic acid by Azotobactersp. grown on
benzoic acid (414) and by Pseudomonas sp. WR912 (195).
  Cells  grown on chlorinated  compounds  including 3-chlorobenzoic acid  are
induced to produce high levels of pyrocatechase II, which has high activity against
chlorocatechols as compared to catechols  (118). Cells grown on nonchlorinated
substrates express only  pyrocatechase I, which does not function in chlorocatechol
oxygenation. Pyrocatechases I and II are separate catechol 1,2-dioxygenases (118).
Pyrocatechase I is involved in the degradation of catechol via the 3-ketoadipic acid
pathway (117). Pyrocatechase II is similar to the Brevibacterium spp. pyrocatechase

                                     69

-------
         S
         w
         10
    £
         o_
         CO
    o  & o
^   all
0   S  1°
                                               CO OH
       u>
       -
         o

         JO
         CO
         O)
2,3,6-TRICHLORO-
  BENZOIC ACID
   2,3,6-TRICHLORO-
4-HYDROXYBENZOIC ACID
       OH
2,3,5-TRICHLORO-
    PHENOL
                                               3,5-DICHLOROCATECHOL
         o
         o
         ff
         §
         o
         Q)
         g
         CL

-------
(321) and is unusual in its broad substrate specificity (118). The ability to cleave
chlorocatechols, which are toxic, appears to be a crucial factor in the ability of
microorganisms to degrade chloroaromatic compounds (118).
  In cells of Pseudomonas sp. WR912, pyrocatechases I and  11 are both induced
when the growth substrate is unsubstituted benzoic acid (195). This organism can use
benzoic acid, 3-chloro-, 4-chloro-, and 3,5-dichlorobenzoic acids as sole sources of
carbon and energy. Each is metabolized to the corresponding chlorocatechol, which
undergoes ortho cleavage to form the chlorinated muconic acid. The muconic acid in
each case is cycloisomerized with coincident or subsequent stoichiometric elimi-
nation of the chloride ion (Figure 33). Because of the wide range of substrates, the
benzoic acid 1,2-dioxygenase of Pseudomonas sp. WR912 is characterized as being
of low substrate specificity and also not stereospecific, similar to the corresponding
enzyme of P. putida mt-2 (195).
   In contrast, the benzoic acid 1,2-dioxygenase of Pseudomonas sp. B13 (Figure 34)
shows  narrow  substrate  specificity, as this organism metabolizes only 3-chloro-
benzoic acid (195). The isomer 2-chlorobenzoic acid does not induce oxygen uptake,
and 4-chlorobenzoic acid is  oxidized  only at very low reaction rates (363). Cells
grown on 3-chlorobenzoic acid are adapted simultaneously to metabolize  benzoic
acid, but the reverse is not true (119). The enzymes of the 3-keto-adipic acid pathway
are induced in cells grown on benzoic acid, and the chlorinated catechols accumulate.
In contrast, resting cell suspensions of cultures grown  on 3-chlorobenzoic  acid
produce both  3-chloro-  and  5-chlorodihydroxybenzoic acid in  almost equal
quantities. A branched pathway thus exists for metabolism of 3-chlorobenzoic acid
by Pseudomonas sp. B13 (Figure 34). Along one branch 3-chlorodihydroxybenzoic
acid is  metabolized  to 3-chlorocatechol and  along a  parallel branch 5-chloro-
dihydroxybenzoic acid is converted to 4-chlorocatechol.  The common  enzyme
involved, 3,5-cyclohexadiene-l,2-diol-l-carboxylic acid  dehydrogenase, has the
same relative activity in both benzoic acid-grown and 3-chlorobenzoic acid-grown
cells (365). The chlorocatechols are metabolized to muconic acids by pyrocatechases
which are induced only in  3-chlorobenzoic acid-grown  cells. The muconate
cycloisomerase II enzyme which acts on the muconic  acids to perform cyclo-
isomerization has much higher activity in  3-chlorobenzoic acid-grown cells (119).
Combined dechlorination and regeneration of the diene system is a spontaneous
secondary reaction (386). P.  putida  strain 87 isolated from soils treated with
pesticides also  contains two pyrocatechases,  one specific for  the nonchlorinated
catechol and the other specific for  chlorinated catechols  (187). The former  is
controlled by chromosomal genes and the latter is plasmid mediated. Chloromuconic
acid was detected upon incubation of this strain with 3-chlorobenzoic acid.
   The mutant  strain  Alcaligenes eutrophus B9 also produces 3-chloro- and 5-
chlorodihydroxybenzoic  acid from -cooxidation of 3-chlorobenzoic acid (367). A
strain of P. putida, harboring a plasmid termed pAC25, degrades 3-chlorobenzoic
acid via 3-chlorocatechol and 3-chloromuconic acid  (86). Chloride is released and
maleylacetic  acid rather than  3-ketoadipic acid is  produced. This  pathway  is
analogous to one path of 3-chlorobenzoic acid metabolism by Pseudomonas sp. B13.
   Four strains  of Pseudomonas spp. which utilize 3-chlorobenzoic acid as the sole
source of carbon and energy for growth were isolated from sewage which had been
enriched with the substrate (194). One isolate studied in detail, Pseudomonas sp.
strain HI, resembles Pseudomonas sp. B13 in metabolism of 3-chlorobenzoic acid
and benzoic acid, and therefore seems to possess both  pyrocatechases. A black
pigment, resulting from oxidation and polymerization of unmetabolized catechols,
forms when Pseudomonassp. strain H1 is incubated with both 3-chlorobenzoic acid
and benzoic acid without prior  adaptation to 3-chlorobenzoic acid.  In contrast,
another species isolated from the sewage enrichment culture metabolizes catechols
and chlorocatechols rapidly enough to prevent occurrence of the black pigment.

                                      71

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     COOH
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2-CHLOflOMUCONIC ACID

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                                         COOH
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                                                                  3.5-DICHLOROCATECHOL
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  Figure 33.  Pathways of metabolism of chlorobenzoic acids by Pseudomonas sp.
           WR912. Compounds in brackets are hypothetical intermediates.
                            Adapted from Reference 195.
                                         72

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  A hybrid strain has been developed which combines the ortho cleavage pathway of
Pseudomonas sp. B13 with the relatively nonspecific toluate 1,2-dioxygenase of P.
putida mt-2 (364). This derivative of Pseudomonas sp. B13  acquired the TOL
plasmid from P. putida mt-2.  In the resulting cells, both 4-chlorobenzoic acid and
3,5-dichlorobenzoic acid as well as 3-chlorobenzoic acid are metabolized (Figure 35).
The enzyme, toluate 1,2-dioxygenase from the genes on the plasmid, is induced and
slightly modified to result in increased turnover of the chlorinated compound used as
the selective substrate. Dihydrodihydroxybenzoic acid dehydrogenases from both
plasmid and chromosomal sources are induced. In addition, ortho pyrocatechases 1
and II are induced, but not the unproductive meta pyrocatechase.
  A  Bacillus sp. grown on benzoic acid uses a unique  pathway to cometabolize
3-chlorobenzoic acid to 5-chloro-2,3-dihydroxybenzoic  acid via 5-chlorosalicylic
acid (406). Another unique pathway involving enzymatic rather than spontaneous
elimination  of  chloride ion was demonstrated in Bacillus  brevis isolated from
polluted river water (96). This  organism utilizes 5-chloro-2-hydroxybenzoic acid (5-
chlorosalicylic acid) as a sole carbon and energy source. The first step in metabolism
is cleavage between the carbon-1  and carbon-2  by a specific 5-chlorosalicylate
1,2-dioxygenase. This enzyme requires a  halogen (except iodine) or a methoxyl
group (but not  a hydroxyl) at the carbon-5 position. Only one hydroxyl group is
present  on  the molecule.  After   loss of the  chloride ion  and  formation of
maleylpyruvic  acid, metabolism continues along the steps of the gentisic acid
pathway (Figure 36).
  A  novel pathway for 3-chlorobenzoic acid  metabolism in which  the chloride is
replaced by a  hydroxyl  group in the first step has  been  demonstrated in  a
Pseudomonas sp.  isolated from  soil (231). This organism utilizes 3-chlorobenzoic
acid as a sole source of carbon for growth and metabolizes it to 3-hydroxybenzoic
acid and subsequently to 2,5-dihydroxybenzoic acid.
  Similarly, an  Arthrobactersp.  growing on 4-chlorobenzoic acid as the sole source
of carbon produces 4-hydroxybenzoic  acid and 3,4-dihydroxybenzoic acid (proto-
catechuic acid)  (380). A strain of Arthrobacter globiformis also utilizes 4-chloro-
benzoic acid as the sole carbon source and  metabolizes it via 4-hydroxybenzoic acid
and protocatechuic acid, with release  of chloride (486). Direct replacement of the
chloride by the hydroxyl group precludes  formation  of the potentially toxic
chlorocatechol. Pseudomonas sp. strain CBS 3 also utilizes 4-chlorobenzoic acid as
the sole source of carbon for growth (257) by the same pathway. The enzymes
induced by growth with this substrate have been identified as 4-chlorobenzoate-4-
hydroxylase, 4-hydroxybenzoate-3-hydroxylase,  and protocatechuate-3,4-dioxy-
genase. The first enzyme was not induced by growth with 4-hydroxybenzoic acid or
any of several other chlorinated  and nonchlorinated substrates. The mechanism of
action of this enzyme has not been elucidated.  Protocatechuic acid was metabolized
by the 3-ketoadipic acid pathway following ortho cleavage.

ALGAL METABOLISM OF CHLOROBENZOIC ACIDS

  The only reference to algal metabolism of chlorobenzoic acids obtained involves a
monoalgal culture of Chlamydomonas sp. strain A2 isolated from sewage (221). The
nonaxenic culture (bacteria present) transforms 4-chloro-3,5-dinitrobenzoic acid to
3-hydroxymuconic semialdehyde, indicating  a meta-cleavage pathway. Approxi-
mately 20% chloride release was reported. Since the culture contained bacteria along
with the algae, dechlorination might have been due to the action of the algae or  it
might have been a bacterial process with the algae providing fixed carbon or growth
factors.
                                     74

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-------
FUNGAL METABOLISM OF CHLOROBENZOIC ACIDS

   Aspergillus niger cultures utilized both 2-chloro- and 3-chlorobenzoic acid as sole
sources of carbon and energy (390). Protocatechuic acid and 4-hydroxybenzoic acid
were isolated from both samples. Cells grown on these substrates oxidize all four
compounds as well as benzoic acid more  rapidly than do  cells grown on glucose
Adapted cultures dechlorinate either substrate, while glucose grown cells do not have
this capability. Dehalogenating activity was also noted in the cell-free extracts of
cultures grown on 2-chlorobenzoic acid.

 METABOLISM OF CHLOROBENZOIC ACIDS IN SOILS AND BY
 CONSORTIA

  Under cometabolic conditions with glucose as the additional carbon source, a
sewage plant effluent inoculum metabolized 3-chlorobenzoic acid with production of
3-chlorocatechol (212). Upon continued incubation this metabolite disappeared with
concomitant appearance of 2-hydroxy-3-chloromuconic semialdehyde. After a 29-
day period  of no discernible metabolism,  the semialdehyde was metabolized with
appearance of  stoichiometric  amounts  of  inorganic  chloride.  There was  no
additional increase in cell numbers due to  the presence of chlorobenzoic acid until
degradation of the semialdehyde occurred.
  Diluted wastewater sludge supernatant fluid mediated disappearance of 16 mg/1
3-chlorobenzoic  acid within 14 days, although no degradation of 2- or 4-chloro-
benzoic acid was seen after 25  days (193). Readaption of the sludge inocula to
3-chlorobenzoic acid greatly reduced the time required for metabolism of both 3- and
4-chlorobenzoic acid. Soil suspension also  did not degrade  4-chlorobenzoic acid in
25 days, although the  other 2  substrates  were  metabolized within 14 days.
Application of 2,3,6-trichlorobenzoic acid to  soil resulted  in 30% chloride release
within one month (113). No intermediate metabolites were detected.
  A  sewage  microcosm  enriched with  chlorinated  benzoic  acids  resulted in
development of  a  consortium of Gram-negative motile rods and Gram-positive
pleomorphic rods which could utilize as sole carbon and energy sources benzoic acid,
2-chloro-, 3-chloro-, 4-chloro- and 3,4-dichlorobenzoic acids but not 2,4-dichloro- or
2,3,6-trichlorobenzoic acid (114). Addition of biodegradable benzoic acids did not
lead to decomposition of any of the substrates.  Degradation of 3-chlorobenzoic acid
led to formation of  both  4-chlorocatechol and  5-chlorosalicylic acid,  the latter
compounds disappearing from solution. Formation of both metabolites indicates
two separate pathways of metabolism within the consortium.
  Pronamide [3,5-dichloro-N-(l,l-dimethyl-2-propynyl)benzamide] is an herbicide
used for weed control on crops of lettuce and  alfalfa and other legumes (153).
Pronamide was metabolized in soils to 14CO2 from both l4C(carbonyl)- and l4C(ring)-
labeled  substrate. In addition, seven other metabolites were found, none of which
was dechlorinated. The potential metabolite 3,5-dichlorobenzoic acid was also
metabolized with 80% of  i"C(carbonyl)-  and 50% of "
-------
recovered as 14CO2, and metabolites which were not radioactive were not identified.
Dicamba applied at a rate of 1.1 kg/ ha to a silty clay with high organic content almost
entirely disappeared within two weeks (399a). At the same application rate onto
sandy loam and heavy clay, there was less than 10% substrate remaining after 4 weeks
when the moisture content was high, although residues were still detected after 6
weeks in low-moisture soils. After 4 weeks, over 90% of the substrate applied to
sterile  soils was  recovered. Studies  with  l4C(carboxyl)-  and I4C(ring)-labeled
dicamba revealed that 18% was released as I4CO2 in 6 weeks and 45% in 17 weeks
from 14C(carboxyl)-dicamba, and 9% in 6 weeks from 14C(ring)-dicamba (398). The
only metabolite which could be recovered was 3,6-dichlorosalicylic acid.

REDUCTIVE DECHLORINATION

  The chlorobenzoic acids have served as model substrates for the elucidation of
anaerobic reductive dechlorination by consortia of bacteria from anaerobic sediment
or sludge environments. This pathway for dechlorination of aromatic compounds
involves removal of the aryl halide from the aromatic ring (Figure 37) (417). A
consortium resulting from enrichment with 3-chlorobenzoic acid mineralizes this
substrate through benzole acid to methane and CO2. The substrate 4-amino-3,5-
dichlorobenzoic acid is converted to 4-amino-3-chlorobenzoic acid by replacement
of one chlorine atom with  a hydrogen atom. No chloride shift takes place. Neither
2-chloro- nor 4-chlorobenzoic acid was metabolized in experiments lasting for one
year of  incubation (206). In multi-substituted compounds, the meta substituent is
utilized  preferentially. Thus, 2,5-dichlorobenzoic acid is reduced to 2-chlorobenzoic
acid, 3,4-dichlorobenzoic acid  to 4-chlorobenzoic acid, and  2,3,6-trichlorobenzoic
acid to  2,6-dichlorobenzoic acid (417). In these experiments, persistence was not
correlated with the number of halogens present on the molecule.
  Although benzoic acid was always an intermediate in anaerobic mineralization of
the chlorobenzoic acids, acclimation to the chlorinated substrate did not result in
acclimation to benzoic acid (206). This phenomenon was  explored further with the
consortium acclimated to 3-chlorobenzoic acid. Metabolism of 3,5-dichlorobenzoic
acid to 3-chlorobenzoic acid proceeded with accumulation of 3-chlorobenzoic acid
until the parent  substrate concentration fell to a low  level.  Only then was 3-
chlorobenzoic acid metabolized to benzoic acid followed by production of methane
and CO2. Similarly, 4-amino-3,5-dichlorobenzoic acid was metabolized to 4-amino-
3-chlorobenzoic acid which accumulated until the concentration of parent substrate
decreased to a low level, after which the intermediate metabolite was converted to
4-aminobenzoic acid which was not degraded further. These events were postulated
to be due to competitive  substrate inhibition, in which one enzyme involved in
multiple steps of a degradative pathway acts only on the parent compound until its
concentration falls below a threshold level (418). Under these conditions, bacteria
from environments receiving several structurally related chemicals may metabolize
substrates selectively due to competitive substrate inhibition.
  The consortium could be acclimated to degrade 4-amino-3,5-dichlorobenzoic acid
to its metabolites, even though this substrate was not used as a sole carbon and energy
source  and  was  not mineralized (206).  Aliquots of the  substrate  added after
acclimation were metabolized without a lag.  Partial dechlorination of several
compounds by sewage  microflora acclimated  to  nonchlorinated products was
reported (221). Substrates  attacked include 2-chlorotoluene, 3-chlorobenzoic acid,
4-chloro-2,5-dinitrobenzoic acid, 4-chloroaniline, 4-chlorobiphenyl, 4-chlororesor-
cinol and 4-chlorobenzonitrile. From 2-47% of the chlorine from these substrates was
removed within 2 days. The substrate 4-chloro-3,5-dinitrobenzoic acid was 13 to 45%
dechlorinated in sewage but was only 13 to 20% dechlorinatea within 20 days by
single isolates of bacteria from the sewage.

                                     78

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   3,5-DICHLORO-        3-CHLORO-          BENZOIC         METHANE +
   BENZOIC ACID       BENZOIC ACID           ACID        CARBON DIOXIDE
  Figure 37.  Representative pathway for the reductive dechlorination of chlorobenzoic
                     acids by anaerobic microbial consortia.
                      Adapted from References 206, 417.

  Reductive  dechlorination of  pentachlorophenol has  been demonstrated  in
anaerobic soils (217, 266, 320). Resultant products include isomers of tetrachloro-
phenols,  trichlorophenols, dichlorophenols and 3-chlorophenol. The methylated
chloroanisole analogues  of these isomers have  been detected as well.  These
investigations led to the conclusion that the chloride ions ortho and para to the
hydroxyl group are utilized preferentially (217). Pentachlorophenol metabolism is
discussed further in a later section. In contrast, the reductive dechlorination of DDT
to DDD described in another section involves the alkyl chlorides but not the
chlorides attached to the ring (107, 189a, 202, 234). This has been demonstrated in
yeasts as well as bacteria and in pure cultures as well as consortia. Lindane (gamma-
hexachlorocyclohexane) is a  nonaromatic molecule  which is also  reductively
dechlorinated (Figure 38). The major metabolite is gamma-tetrachlorocyclohexene,
followed by benzene, monochlorobenzene, and small amounts of tri-and tetrachloro-
benzenes (192).
              Cl                          Cl
     HEXACHLOROCYCLOHEXANE   X-TETRACHLOROCYCLOHEXENE
  Figure 38.   Primary metabolic reductive dechlorination of y-hexachlorocyclohexane by
                          anaerobic microorganisms.
                       Adapted from References 192, 267.
                                     79

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SUMMARY

  The chlorobenzoic acids can be metabolized to several different intermediate
products in bacteria. The most common mechanism results in the conversion of the
chlorobenzoic acids to chlorocatechols.  If the meta cleavage pathway is the only
route induced in the bacteria, then  chlorocatechols are metabolized usually to
chloromuconic semialdehydes, which are not metabolized further. In addition, the
meta cleavage product of 3-chlorocatechol inactivates  the  meta pyrocatechase,
causing an accumulation of the toxic chlorocatechols. This pathway does not lead to
mineralization of the chlorinated substrate, and ultimately results in cell death and
release of chlorinated intermediates  into  the environment. In contrast, ortho
cleavage  of  chlorocatechols  is a successful pathway  for chlorobenzoic acid
mineralization. The  ortho pyrocatechase results in the formation of chlorinated
muconic acids, which are cycloisomerized with coincident or subsequent elimination
of the chloride  ion.  After release of the  chloride ion, the compound  is fully
metabolized by established cellular mechanisms.
  One block to utilization of chlorinated aromatic compounds in organisms
expressing the ortho pyrocatechase is specificity of the enzyme required for the first
oxygenation step. The hybrid strain constructed from Pseudomonas sp. B13 and P.
putida mt-2 incorporates  the relatively nonspecific oxygenase of P. putida mt-2,
carried  on the TOL plasmid, into   Pseudomonas sp.  B13 which metabolizes
chlorinated compounds via the ortho pathway.  The hybrid strain is  capable of
mineralizing a wider range of chlorobenzoic acids than either parent.
  Another  pathway which has been  discovered replaces the chloride ion by a
hydroxyl group directly, resulting in a nonchlorinated hydroxybenzoic acid which
can be metabolized by established cellular mechanisms. A third series of metabolic
pathways results in  hydroxylation of the chlorobenzoic acid  without loss of prior
substituents. A specific and unique enzyme opens the ring and an enzyme has been
identified which specifically dechlorinated the resulting aliphatic acid molecule. Data
on algal and fungal pathways for metabolism of chlorobenzoic acids are lacking. The
relaxed substrate specificities of fungi suggest that these organisms may be of prime
importance in metabolism of the chlorobenzoic acids.
                                     80

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                              SECTION 8

                        CHLOROBENZENES

MICROBIAL METABOLISM OF CHLOROBENZENES

  Chlorobenzenes are used as industrial solvents and diluents for polychlorinated
biphenyl compounds  (PCBs) and  thus  have a complementary distribution  as
pollutants in the environment, including capacitor and transformer storage and
disposal (295). They are solvents for paints and appear as byproducts in the textile
dyeing industry and in  other industries.  Pentachloronitrobenzene is  used as a
fungicide for seeds and soils.
  A chemostat seeded with a mixture of soil and sewage samples was used to enrich
for an organism capable of utilizing chlorobenzene as a sole growth substrate (366).
After nine months an unidentified bacterium, strain WR1306, was isolated which
degrades chlorobenzene with stoichiometric release of chloride ion. Detection of the
enzymes c/s-l,2-dihydroxycyclohexa-3,5-diene (NAD+)-oxidoreductase, catechol
1,2-dioxygenase, muconate  cycloisomerase, 4-carboxymethylenebut-2-en-4-dlide
hydrolase and NADH-dependent maleylacetate reductase, and isolation  of the
metabolite 3-chlorocatechol, enabled construction of a pathway for chlorobenzene
dissimilation (Figure 39). The proposed pathway is similar to that demonstrated for
the metabolism of other nonphenolic benzene compounds such as 3-chlorobenzoic
acid. Chlorobenzene is converted to 3-chlorocatechol which is cleaved by the ortho
pathway. The substituted muconic acid  thus formed is  cycloisomerized with
coincident or subsequent elimination of chloride. The nonchlorinated intermediate is
metabolized to 3-oxoadipic acid  which enters the cell's tricarboxylic acid cycle.
  Attempts to find other strains of bacteria capable of using chlorobenzene as a sole
source of carbon and energy for growth have been hampered by the lipophilicity of
the compound (366). Strain WR1306, although capable of growth on chlorobenzene,
was inhibited by high concentrations of the compound. Accumulation of the toxic
metabolite 3-chlorocatechol is lethal for cells which have not evolved a mechanism
for efficient metabolism of this compound. The mechanism of catechol cleavage must
be by the ortho rather than the meta pathway for production of metabolites useful in
cell biosynthesis (discussed further in section on chlorobenzoates).
  Resting cells of P. putida grown on toluene oxidize chlorobenzene and to a lesser
extent all three isbmers of chlorotoluene (182). Cometabolic growth of P. putida on
toluene and chlorobenzene results in formation of 3-chlorocatechol. Cells grown on
toluene  and 4-chlorotoluene  metabolize the  latter through  c/s-4-chloro-2,3-di-
hydroxy-1 -methylcyclohexa-4,6-diene to 4-chloro-2,3-dihydroxy-1 -methylbenzene.
  Bacteria  were isolated which  utilize  1-chloronaphthalene for growth (45la).
Chloronaphthalene  diol (8-chloro-l,2-dihydro-l,2-dihydroxynaphthalene) and  3-
chlorosalicylic acid were recovered as metabolites.
  An alternative pathway has been developed which proposes chlorophenols  as
intermediates in the degradation of chlorobenzenes (19). However, the formation of
phenolic products from the dihydrodiol metabolite may occur spontaneously under
                                    81

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-------
mild acid conditions as well as enzymatically (366). Confirmation of this pathway
requires isolation of the enzymes involved.
  Pure cultures of bacteria isolated from pond water and pond sediment samples
metabolize dichlobenil (2,6-dichlorobenzonitrile) to 2,6-dichlorobenzamide, 2,6-
dichlorobenzoic  acid and several other metabolites  which  appeared in trace
quantities (131,  132). An Arthrobacter sp. which was grown  on  benzonitrile
metabolized more  than 70% of l4C-dichlobenil to 2,6-dichlorobenzamide and an
additional 20% to other metabolites within 6 days (314).
  The degradation of chloronitrobenzenes  by fungi  has  been studied and  the
mechanism is believed to result in detoxification of the fungicides (94). The yeast
Rhodosporidium sp., when grown in a complex nutrient medium containing 4-
chloronitrobenzene,  produces  several  metabolites.  This information  enabled  a
branched pathway to be proposed (Figure 40) (94). The common early steps of
4-chloronitrobenzene metabolism involve sequential reduction of the substrate to
form 4-chloronitrosobenzene and subsequently 4-chlorophenylhydroxylamine. This
product may be metabolized by two mechanisms. The main  pathway is further
reduction of the hydroxylamine to 4-chloroaniline, followed by acetylation to
produce 4-chloroacetanilide, the major metabolite. This product accumulates in the
culture medium. An alternative mechanism involves a shift of the hydroxyl group
from nitrogen to carbon (called a Bamberger rearrangement) resulting in conversion
of the hydroxylamine to 4-hydroxyaniline and 2-amino-5-chlorophenol. Formation
of 4-hydroxyaniline involves loss of the chloride ion and subsequent acetylation
results in formation of 4-hydroxyacetanilide.
  Pentachloronitrobenzene is metabolized to pentachloroaniline by Streptomyces
aureofaciens, Rhizoctonia solani, Fusarium oxysporum and many other genera of
fungi and actinomycetes (80, 322). In addition, F. oxysporum also metabolizes the
substrate to  pentachlorothioanisole  (Figure 41)  (322).  The  introduction of  a
methylthio group was also noted in the  metabolism of 2,4-dichloro-l-nitrobenzene
by Mucorjavanicus AHU6010 (Figure 41) (428). The source of the sulfur atom has
not been established. The metabolite may be formed by  secondary degradation of a
glutathione degradation product, as proposed for rhesus monkey metabolism, or the
methylthio group may be transferred from S-adenosylmethionine as demonstrated
by cells of Mycobacterium sp. (428).
  Metabolism of l,4-dichloro-2,5-dimethoxybenzene (chloroneb) to 2,5-dichloro-4-
methoxyphenol by R. solaniis a detoxification mechanism which results in tolerance
by the organism to at least twice the concentration of metabolite as product (204).
This conversion occurred only at a high ratio  of mycelia to growth medium.
Sclerotium rolfsii and Saccharomyces pastorianus did  not metabolize chloroneb.
Neurospora crassa  converts chloroneb to an unidentified product.
  In  a broad study of 23 species, conversion of chloroneb to 2,5-dichloro-4-
methoxyphenol occurred only in cultures actively growing in nutrient medium (468).
The most active species was a Fusarium sp. which demethylated 60 to 80% of a 5 ppm
solution within 10 days. Thirteen other  species demethylated the fungicide as well,
although R. solani neither grew in the presence  of nor demethylated chloroneb.
Fusarium sp. also converted the metabolite back to chloroneb at the rate of 4% in 7
days. Eleven other species performed the same transformation. Demethylation of
2,5-dichloro-4-methoxyphenol to form 2,5-dichlorohydroquinone occurred in four
species, but at  concentrations of 10% or  less of the applied 2,5-dichloro-4-
methoxyphenol.
                                    83

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-------
METABOLISM OF CHLOROBENZENES BY MICROBIAL

COMMUNITIES
  The chlorobenzenes are largely volatile and attempts to study their biodegradation
have been hampered by disappearance of the substrates. A mesocosm experiment
using tanks  of seawater amended with mixtures of volatile organic compounds
showed that at 3  to 7°C  both 1,4-dichlorobenzene  and 1,2,4-trichlorobenzene
disappeared  at rates explainable by volatilization (446). At warmer temperatures (20
to 22°C), the rate of disappearance was much more rapid, indicating biodegradation
by the planktonic and microbial communities.
  Application of 1,2,3- and 1,2,4-trichlorobenzene at a rate of 50 mg/g soil resulted
in I4CO2 evolution of more than 10% after several weeks (295). Addition of high levels
of organic matter increased only 1,2,3-trichlorobenzene mineralization. Extracts of
soil dosed with 1,2,3-trichlorobenzene yielded 2,3- and 2,6-dichlorophenol, and
3,4,5-trichlorophenol, while 1,2,4-trichlorobenzene samples yielded 2,4-, 2,5-, and
3,4-dichlorophenol.
  A mixed population of soil bacteria precultured on benzene metabolized benzene
and chlorobenzenes (20 to 200 mg/1) to chlorophenols (19). Mono- through tetra-
chlorobenzenes were monohydroxylated at a  position ortho to the chloride. No
phenol was detected in media containing pentachlorobenzene. Diphenyls eventually
were detected in the media.
  A granular activated carbon column was seeded with a mixed culture of bacteria
(primary sewage) and supplied with acetate as a carbon source (52). A biofilm was
formed which after acclimation metabolized more than 90% of a 10 to 30 mg/1
solution each of chlorobenzene, 1,2-di-, 1,4-di-, and 1,2,4-trichlorobenzene. Partial
disappearance of  1,3-dichlorobenzene was noted. Studies with 14C-l,4-dichloro-
benzene confirmed that these substrates were mineralized to I4CO2.
  Hexachlorobenzene was applied to zoysia plots at a rate resulting in 6 ppm in the
upper 2 cm  of the soil (28). The bulk of the material was volatilized with 24%
remaining after 29 days and 3.4% after  19 months. The remaining material was
unchanged substrate. The original application resulted in 0.11 ppm in the 2 to 4 cm
layer which did not change during the course of the experiment.
  Several experiments concerning the metabolism of dichlobenil in soils have shown
that the major metabolite is 2,6-dichlorobenzamide (88, 314, 314a, 442a). Other
metabolites appeared in trace quantities and could not be identified. After 61 days'
incubation of the substrate l4C-labeled in the nitrile group, only trace amounts of
I4CO2 or volatile l4C-compounds could be recovered. More than 85% of the substrate
added at 1 ppm remained unaltered.
  Formation of 2,6-dichlorobenzamide in a pond water and sediment system was
followed by a decrease in its concentration, indicating further transformation (314).
Carbonyl-l4C-labeled 2,6-dichlorobenzamide was metabolized with 5.6% converted
to I4CO2 after 40 days and 28% recovered as metabolites.
  Application of 1  mg/1 dichlobenil to a farm pond resulted in initial sorption of the
herbicide to  the soil with subsequent disappearance from both water and soil (371).
Less than  10% remained 90 days after treatment. Soils which had  been pretreated
with 2,6-dichloro-4-nitroaniline showed evolution of I4CO2 when treated  with this
radiolabeled fungicide (189). No I4CO2 evolution was noted in soils which had not
been pretreated. Some unidentified metabolites were also seen. A pure culture of rod
shaped bacteria was isolated which also converted the fungicide to  I4CO2.
  Greenhouse soils treated with pentachloronitrobenzene were analyzed for the
presence of  metabolites (108). Products recovered included pentachloroaniline,
pentachlorothioanisole, and  tetrachloronitrobenzene.  Hexachlorobenzene and
                                    86

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pentachlorobenzene were detected but are known 'to be  present as impurities in
technical grade pentachloronitrobenzene.
  Extracts of soils amended with 1,000 ppm pentachloronitrobenzene revealed the
presence of pentachloroaniline but  no polychloroazobenzenes (62). Soils treated
periodically for 11  years  still showed  residual  pentachloronitrobenzene and  the
technical grade impurities tetrachloronitrobenzene, pentachlorobenzene, hexachloro-
benzene, pentachloroaniline and methylthiopentachlorobenzene when analyzed 1 to
5 years later (29).
  Anaerobic flooded or moist Hagerstown silty clay loam  was treated with 10 ppm
pentachloronitrobenzene (319). After 40 days' incubation there was no evolution of
14CO2 and only slight volatilization of the unchanged substrate. The main metabolite
formed was pentachloroanisole, and lesser amounts of pentachlorothioanisole and
pentachlorophenol were also detected.
  Chloroneb (14C-ring labeled) was applied to soil plots at the rate of 2 lb/ acre (370).
Another layer of soil was applied to the plots. After 12 months 40% of the original
activity was recovered,  of which 90% was unchanged  substrate  and 10% an
unidentified metabolite.

SUMMARY

  Chlorobenzenes  containing  less  than five chlorines  can be  mineralized by
acclimated populations under permissive conditions.  High concentrations of these
compounds are toxic to the bacteria. Most of the information regarding metabolism
of the chlorobenzenes has come from studies with soil or  mixed culture consortia.
There is little information available on pathways of metabolism by pure cultures.
  A single study indicates that hexachlorobenzene  is not metabolized in soils.
Pentachlorobenzene was not oxidized  in a sole study, although under anaerobic
conditions pentachloronitrobenzene is converted to several metabolites.
  The available evidence indicates that chlorocatechols or chlorophenols are the
primary  degradation  products of chlorobenzenes.   These metabolites can be
metabolized by mechanisms discussed in the appropriate sections.
                                     87

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                              SECTION 9

                        CHLOROPHENOLS

  The chlorophenols are used extensively as antifungal agents and are often applied
as a preservative to freshly sawn lumber. They have found some use as herbicides and
in food processing plants to control mold (99). Chlorophenols are also common
degradation products of chlorophenoxy herbicides. The wood shavings from lumber
processes have been used for litter in chicken houses and contain high levels of these
chlorophenols, especially 2,3,4,6-tetrachlorophenol  and pentachlorophenol (101).
The chlorophenols degrade to volatile chloroanisoles via methylation of the oxygen
atom and the resulting compounds have been implicated in the "musty taint" of
chicken eggs and meat (101).

BACTERIAL METABOLISM OF CHLOROPHENOLS

  Studies with Arthrobacter spp. have confirmed methylation as a mechanism for
chlorophenol metabolism.  Conversion of  2,4,6-trichlorophenol  to  2,4,6-
trichloroanisole  has been demonstrated.  Methylation is  also the predominant
reaction  in the  conversion  of guaiacols (o-methoxyphenol)  to  veratroles (1,2-
dimethoxybenzene) by Arthrobacter spp. (325). For example, 4-chloroguaiacol,
4,5-dichloroguaiacol, and 3,4,5-trichloroguaiacol are converted to the correspond-
ing veratroles by dense cell suspensions of cultures grown on hydroxybenzoic acid
(Figure 42). Low concentrations of catechols have also been found in the culture
medium.  An exception to this mechanism has been demonstrated in the metabolism
by Arthrobacter sp. strain 1395 of 3,4,5-trichloroguaiacol to 3,4,5-trichlorosyringol,
which requires  hydroxylation and subsequent methylation  of the previously
unsubstituted carbon-6 (Figure 42). This latter compound is resistant to further
degradation by this species.
  An alternative mechanism results  in the  formation of chlorocatechols  from
chlorophenols. Cells of a Nocardia sp. grown on phenol metabolize 2-chlorophenol
to 3-chlorocatechol, 3-chlorophenol to  4-chlorocatechol, and 4-chlorophenol to
4-chlorocatechol (406).  Similarly, phenol-grown Pseudomonas  sp. B13 or
Alcaligenes eutrophus cells metabolize 2-chlorophenol to 3-chlorocatechol and 4-
chlorophenol to 4-chlorocatechol  (262).  Pseudomonas sp.  B13  can utilize 4-
chlorophenol as the sole source of carbon and energy, and with this substrate can
cometabolize 2-chlorophenol and 3-chlorophenol completely without accumulation
of metabolites (262). A phenylcarbamate-degrading  Arthrobacter sp. also metabo-
lizes  4-chlorophenol to 4-chlorocatechol (466a). While the  same initial enzyme is
used for the first step in both phenol and 4-chlorophenol degradation, phenol grown
cells  contain a  muconate-lactonizing enzyme  which has little activity for 3-
chloromuconic acid, the metabolite of 4-chlorophenol (118).
  Resting cell suspensions of Achromobacter sp. metabolize 4-chlorocatechol to
4-chloro-2-hydroxymuconic semialdehyde and 3,5-dichlorocatechol to 3,5-dichloro-
2-hydroxymuconic semialdehyde via a unique catechol 1,6-oxygenase which differs
from the more common catechol 2,3-oxygenase (Figure 43) (209). Neither product is
metabolized further. Pseudomonas putida metabolizes 4-chlorophenol to 4-chloro-
catechol,  and then employs the meta cleavage enzyme catechol 2,3-dioxygenase to

-------
                OCH,
   4-CHLOROGUAIACOL
                                  4-CHLOROVERATROLE
                OCH,
 4,5-DICHLOROGUAIACOL
 4,5-DICHLOROVERATROLE
               OCH,
3,4,5-TRICHLOROGUAIACOL
3,4,5-TRICHLOROVERATROLE
                                3,4,5-TRICHLOROSYRINGOL
  PENTACHLOROPHENOL
  PENTACHLOROANISOLE
       Figure 42.  Methylation of chlorophenols by Arthrobacterspp.
                  Adapted from Reference 325.
                           89

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3-CHLOROBENZOIC
      ACID
4-CHLOROCATECHOL
     4-CHLORO-
2-HYDROXYMUCONIC
   SEMIALDEHYDE
 2,6,6-TRICHLORO-
   BENZOIC ACID
   3,5-DICHLORO-
     CATECHOL
  3,5-DICHLORO-2-
 HYDROXYMUCONIC
  SEMIALDEHYDE
a = catechol 1,6-oxygenase
Figure 43.  Cometabolism of chlorocatechols via catechol 1,6-oxygenase by resting
                cell suspensions of Achromobacter sp.
                   Adapted from Reference 209.
                             90

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produce 2-hydroxy-5-chloromuconic semialdehyde, which accumulates to 10% of
the starting substrate. Free chloride amounting to 85% of the substrate is recovered,
although a pathway for liberation of the chloride has not been elucidated (333).
  Two species of bacteria were utilized to produce a genetically constructed strain
with altered ability to metabolize aromatic compounds (388). Pseudomonas sp. B13,
with ability to metabolize  chlorophenols, and Alcaligenes sp. A7, which degrades
phenol by the meta path and has no activity against chlorophenols, were combined to
produce a mutant (designated A7-2) which utilizes phenol by the ort/iopath and also
metabolizes  2-, 3-,  and 4-chlorophenol as well  as  3-chlorobenzoic acid. Three
enzymes  were isolated, pyrocatechase II and cycloisomerase II,  which have high
activity for chlorinated substrates, and a third enzyme which functions exclusively in
the chloroaromatic  pathway to perform a dehalogenating cycloisomerization of
chloromuconic acids (Figure 44).
  The 2,4,5-trichlorophenoxyacetic acid-degrading strain of P. cepacia AC1100 can
dechlorinate  a wide variety of chlorophenols (237). Resting cell suspensions can
dechlorinate  within 3 hours at 0.1  mM substrate concentration, the following
chlorophenols: 2,3-, 2,4-, and 2,5-dichlorophenol, 2,3,4- and 2,4,5-trichlorophenol,
2,3,4,6- and 2,3,5,6-tetrachlorophenol and pentachlorophenol. The strain has less
activity against 2,4,6-trichlorophenol and 2,3,4,5-tetrachlorophenol and metabolizes
2,6- and 3,5-dichlorophenol and 2,3,5-, 2,3,6-, and 3,4,5-trichlorophenol poorly.

METABOLISM OF CHLOROPHENOLS BY FUNGI

  Fungal metabolism of chlorophenols  often involves methylation in a  manner
analogous to that of bacteria (172). A study of 116 fungal isolates from chicken house
litter revealed that 59% produce 2,3,4,6-tetrachloroanisole from 2,3,4,6-tetrachloro-
phenol at conversion efficiencies of from 1 to 83%. Flask cultures in this study were
sealed and incubated for five days, so the transformation may have occurred under
aerobic or anaerobic conditions. The  fungi demonstrating this ability include
Aspergillusspp., Paecilomycesspp., Penicilliumspp., and Scopulariopsisspp. (100,
101, 172). Some strains metabolize 2,3,4,6-tetrachlorophenol without formation of
the anisole, suggesting an alternate mechanism for chlorophenol metabolism (172).
The  yeast Rhodotorula glutinis grown on phenol converts 3-chlorophenol to 4-
chlorocatechol (448).
  There is  little additional information available on  fungal metabolism of
chlorophenols, although there is evidence to indicate that a Penicillium sp. produces
2,4-dichlorophenol as a natural metabolite (8).

METABOLISM OF CHLOROPHENOLS BY MIXED MICROBIAL

CULTURES
  Soil populations exhibited enhanced rates of metabolism of 2-chlorophenol after
prior acclimation by soil  percolation (447). Following an initial  decrease  in 4-
chlorophenol concentration during  soil percolation, however, additional appli-
cations of that substrate were not  metabolized. In other experiments, soil inocula
mediated the complete disappearance of 4-chlorophenol within 25 days, although
neither 2- nor 3-chlorophenol was degraded during that time (193).
  Wastewater sludge  supernatant liquid  required 14 to 25 days  for complete
disappearance of 16 mg/1  2-chlorophenol and 3-chlorophenol, although 4-chloro-
phenol disappeared within  14 days (193). Disappearance of 1 mg/1 2- and 4-
chlorophenol was faster in  polluted acclimated or nonacclimated river water than in
diluted sewage inocula experiments (139). The substrates were not degraded by the
sewage microorganisms after 25 days,  while less  than 15 days was required for
diappearance of the substrates from river water.

                                    91

-------
       OH

      [o]
         Pyrocatechase I
          O-
          :
 COOH
  COOH
         Cycloisomerase 1
 COOH

.0'
        COOH
         C=0
         Hydrolase 1
3-KETOADIPIC ACID --
                           OH

                          [Ol
                                                              OH
                               OH
                                          Pyrocatechase I
COOH
 COOH
                                                              Cl
 ,COOH
[I   COOH
U  jJ
                                          Cycloisomerase II
                              C,
                                          I
                                    COOH   HOOC
                                     ,'C=0
                                            Hydrolase II
                             --*--MALEYLACETIC ACID
                    TRICARBOXYLIC
                     ACID CYCLE
                                                              Cl
 Figure 44.  Action of aromatic and chloroaromatic enzymes from P. putida B13 and
                        P. putida derivative strains.

                       Adapted from Reference 261.
                                  92

-------
  An acclimated sludge culture exposed to 100 mg/1 substrate was able to metabolize
the following compounds within 5 days with chloride release as noted (218): 2-, 3-, or
4-chlorophenol,  100%; 2,4-dichlorophenol, 100%; 2,5-dichlorophenol,  16%; 2,4,6-
tnchlorophenol,  75%; and sodium pentachlorophenolate, 0%.
  The fate of the monochlorophenols when incubated anaerobically with a 10%
municipal sewage sludge inoculum was  determined (54). At a concentration of 50
mg/1,2-chlorophenol required 3 weeks, 3-chlorophenol 7 weeks, and 4-chlorophenol
16 weeks for complete disappearance. Mineralization was monitored by measuring
net methane production, and results indicated nearly complete mineralization of
2-chlorophenol. Methane was produced  from 3-chlorophenol after a lag period of 5
weeks, and 4-chlorophenol was  not mineralized.  During the degradation of 2-
chlorophenol, phenol was recovered as the initial metabolite, followed by methane
production. This is consistent with other studies in which dechlorination was shown
to be the initial step in the reductive metabolism of 3-chlorobenzoic acids (417).
  Fresh undiluted sludge samples also reductively dechlorinated several dichloro-
phenols with removal  of the ortho  chloride,  such  that 2,6-dichlorophenol was
converted to 2-chlorophenol, 2,3- and 2,5-dichlorophenol to 3-chlorophenol, and
2,4-dichlorophenol  to  4-chlorophenol  (53).  Two  isomers  which lack an ortho
substituent, 3,4- and 3,5-dichlorophenol, were not metabolized during the 6 weeks of
the experiment.
  Undiluted sludge samples were acclimated to the monochlorophenols by repeated
inoculations with 20 mg/1 substrate, until  the cultures could metabolize 25 mg/1
within 1 week (53). Each acclimated sludge culture was then challenged with a 20
mg/1  solution of other chlorophenols. Cultures  acclimated  to 2-chlorophenol
metabolized both 2- and 4-chlorophenol at equal rates and 2,4-dichlorophenol
somewhat more slowly. However, 3-, 2,3-di-, 2,5-di-, and 2,6-dichlorophenol were
not metabolized. Sludge inocula acclimated to 3-chlorophenol also metabolized
4-chlorophenol, and 3,4- and 3,5-dichlorophenol but  not 2-chlorophenol or 2,3- or
2,5-dichlorophenol. Acclimation  to 4-chlorophenol also permitted metabolism of
3-chlorophenol and at much slower  rates 2-chlorophenol and  2,4- and 3,4-
dichlorophenol as well. The population acclimated to 4-chlorophenol seemed to have
a broader substrate range than the other acclimated populations. Incubation of
2- and 4-chlorophenol-acclimated sludge inocula with uniformly ring-'4C-labeled
2-and 4-chlorophenol  and  2,4-dichlorophenol showed that in all cases nearly
complete mineralization of the substrates to l4C-methane and I4CO2 occurred.
  Experiments in which tainted  litter was incubated with sawdust, pentachloro-
phenol, and 2,3,4,6-tetrachlorophenol, showed nearly  quantitative conversion of the
latter substrate to 2,3,4,6-tetrachloroanisole(101). There was virtually no conversion
in the absence of the litter inoculum. Pentachlorophenol was 50% converted to
pentachloroanisole after 29 days.
  Aspergillus sydowi, Scopulariopsis brevicaulis, and  a Penicillium sp. were isolated
from the litter and each species was also found to be capable of the above substrate
conversions.

SUMMARY
  Many of the chlorophenol compounds have been shown to be metabolizable by
pure cultures or mixed natural populations of microorganisms both aerobically and
anaerobically. In most cases complete mineralization occurs.  However, the rates of
disappearance of the isomers vary widely, depending upon degree of acclimation of
the population and other environmental  factors. Mixed cultures seem to be required
for complete mineralization of the chlorophenols.
                                    93

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                             SECTION 10

                     PENTACHLOROPHENOL

  Pentachlorophenol (PCP) is widely used in a variety of agricultural and industrial
applications as a fungicide, bactericide, insecticide, herbicide, and molluscicide (36,
97,  466). It is most widely used in the  United States and elsewhere for wood
preservation,  both for newly cut timber and for slime control in pulp and paper
production. PCP is usually used as a 5% solution in petroleum solvents or as the
water-soluble  sodium or potassium salt.

BACTERIAL METABOLISM OF PCP

  In spite of its use as a fungicide and bactericide, PCP is metabolizable by a variety
of microorganisms. Reports of decomposition of PCP in rice paddy soil and other
soils or aquatic environments (90,  217, 253, 266, 282, 320, 354) were followed by
experiments with consortia and pure cultures of bacteria which demonstrated
chloride release and '4CO2 formation from labeled PCP (89,354,369,411,423,459).
However,  few  studies have  identified metabolites  arising from pure culture
metabolism of PCP.
  Cultures of Pseudomonas spp. produce both tetrachlorocatechol and tetrachloro-
hydroquinone from PCP (Figure 45) (423,424).  These are metabolized rapidly soon
after they are produced. There is no evidence of methylation  of PCP to form
pentachloroanisole. Amino acid analyses with hydrolysates of bacterial cells indicate
incorporation of I4C derived  from PCP into  the cell constituents (423). Penta-
chlorophenol  is metabolized by Arthrobacterspp. to pentachloroanisole at levels of
less than 0.5% conversion at approximately  44 mM substrate concentration (325).
  A bacterium identified as Mycobacterium sp. which cannot use PCP as a growth
substrate methylates PCP to pentachlaoroanisole (424, 425). Further methylations
by washed cell suspensions of this culture result  in the formation of tetrachloro-1,2-
dimethoxybenzene and tetrachloro-1,4-dimethoxybenzene. Additional metabolites
include tetrachlorocatechol, tetrachlorohydroquinone,  tetrachloro-2-methoxyphe-
nol  and tetrachloro-4-methoxyphenol. The formation  of these products indicates
that the main metabolite is the methylated derivative of PCP, but in addition, PCP is
hydroxylated  in the ortho or para positions and subsequently methylated at these
positions. As pentachloroanisole is less toxic to the bacteria than PCP, methylation
is suggested as a detoxification mechanism (424). The methylation of PCP was also
demonstrated in cell-free systems  of Mycobacterium  sp. (422). The  mechanism
appears to involve the enzymes that transfer the methyl group from S-adenosyl-
methionine to the hydroxyl groups of these  substrates.
  A saprophytic soil corynebacterium was isolated which utilizes PCP as a sole
source of carbon and energy for growth (369, 89). By measuring I4CO2 evolution, the
conversion rate was calculated to be 10 mg PCP per mg of dry cell weight per hour
(90). Cells  of this isolate, referred  to as KC3, when  grown on  PCP also show
immediate uptake, as measured by ultraviolet spectrophotometry, of a wide variety
of chlorophenol isomers, including 2,3,5-, 2,3,6-, and 2,4,6-trichlorophenol, 2,3,4,6-,
and 2,3,5,6-tetrachlorophenol and  pentachlorophenol (89). Uptake of 3,4,5-tri-

                                   94

-------
                       3
                       5
    PCP
                    o
                  TeCBQ
OH
xk^ci
._ _ frvT — ^

OH
Clv,xlvv,CI
TOT
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OH
TCHQ
PROPOSED
INTERMEDIATE
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' CHQ



OH

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i ^w'j "
spontaneous ^f^
chemical OH
^reaction 2,6-DCHQ
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ci-^r^ci


0
TCHBQ

slow metabolism
with ring fission






rapid metabolism
with ring fission



incomplete chloride
release; ring fission
questionable


Tetrachlorocatechol
                      TeCHQ
                      TeCBQ
                      TCHQ
                      TCHBQ
                      CHQ
                      DCHQ
= tetrachlorohydroquinone
= tetrachlorobenzoquinone
= trichlorohydroquinone
= trichlorohydroxybenzoquinone
= chlorohydroquinone
= dichlorohydroquinone
Figure 45.   Proposed pathway for pentachlorophenol (PCP) metabolism by the bacterial
                       culture KC-3 and by Pseudomonas sp.
                        Adapted from References 369, 423.
                                      95

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chlorophenol and 2,3,4,5-tetrachlorophenol is delayed. While the para and meta
monochlorophenols are oxidized, as measured by manometric techniques, the ortho
isomer is oxidized poorly and phenol itself not at all (89). Chloride is not released to
an appreciable degree from any  of the monochlorophenols. In general, release of
chloride is greatest from the 2,6-substituted di-, tri-, and tetrachlorophenols. Isomers
with chloride substitutions in other positions are less well attacked by culture KC3.
  Substantial investigations into  the metabolism of PCP by the KC3 isolate failed to
show accumulation of metabolites in the medium. However, mutants were developed
which failed to grow in  a PCP-minimal salts  medium (369). One of these mutants,
designated  ER-47,  converts PCP primarily to 2,6-dichlorohydroquinone. KC3
parent cells adapted to PCP release chloride from 2,6-dichlorohydroquinone rapidly
and without a lag. A trace of monochlorohydroquinone also appears but its role in
the pathway of biological degradation is uncertain, as it is only slowly attacked by
parent KC3 cells.  A second mutant,  ER-7,  accumulates  several metabolites,
including tetrachlorohydroquinone, tetrachlorobenzoquinone, and trichlorohydroxy-
benzoquinone (Figure 36). These three products are converted rapidly and spon-
taneously from the hydroquinone through the benzoquinone to the more stable
hydroxybenzoquinone. The latter product is metabolized by KC3 but dechlorination
is not complete and the ring is not ruptured. Tetrachlorohydroquinone is rapidly
metabolized to 2,6-dichlorohydroquinone but this metabolic transformation must
compete with the rapid spontaneous transformation to the trichlorohydroxybenzo-
quinone.
  Another series of experiments  explored the metabolism of sodium pentachloro-
phenate by a wide variety of bacteria metabolically active for phenols, chlorophenols
or chlorobenzenes (379). Metabolites were identified by detecting acetyl derivatives
using combined gas chromatography and mass spectrometry. With few exceptions
metabolites  occurred in concentrations  of less  than  one percent of the starting
material. Reported metabolites  included PCP-acetate, pentachloroanisole, tetra-
chloroanisoles, tetrachlorophenols, and tetrachlorodihydroxybenzenes.

FUNGAL METABOLISM OF PCP

  The role of fungi in  detoxifying PCP has been  studied to  some degree. Fungi
associated with PCP-treated wood reduce PCP to a less toxic metabolite (122, 288).
Three Trichoderma spp. metabolized sodium pentachlorophenate (Na-PCP) within
2 weeks in a malt extract medium as well as on wood treated with Na-PCP (98).
Pentachloroanisole was detected  in the culture medium of T. virgatum after 5 days'
incubation at levels corresponding to 10-20% of the starting Na-PCP. It is unclear
whether this is an  integral step  of the pathway or whether methylation is a side
reaction (98).
  During  a comparison  of  the growth of several  species of fungi  on PCP,
Trichoderma spp.  were the only ones which reduced PCP  levels  after 12 days'
incubation at 5 to  10 mg/1 concentration (98).  Fungi inactive against PCP were
Cephaloascus fragrans, C. pilifera, Graphium  spp., and Penicillium sp. Another
experiment showed that Trichoderma viride and Coniophora puteana reduced the
concentration of PCP in treated wood blocks, although C. puteana was much more
sensitive to PCP in liquid culture (441). It was postulated that the  presence of an
alternative substrate  of wood or the binding capacity of PCP to  wood reduced
exposure of the fungus to below the toxic level, thus permitting metabolism of PCP.

DISAPPEARANCE OF PCP IN ENVIRONMENTAL SAMPLES

  The standard procedure of applying PCP in a carrier solvent to wood products has
complicated subsequent analyses of disappearance and biodegradability. A carrier

                                    96

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which is too volatile will carry PCP with it as it evaporates (277). A carrier which
retains liquidity at ambient  temperatures  will bleed from the wood until  an
equilibrium is established. PCP in solution will be carried along, reducing the final
concentration in the wood  and increasing the amount reaching the surrounding
environment.
  Extraction and analysis procedures are also subject to error, including incomplete
extraction due to poor choice of extracting solvent, and use of procedures which
extract pure PCP but not polymerized molecules, which may be present in technical
grade PCP at levels as high  as 18 percent (277).
  Degradation of PCP with release of chloride and CO2 has been demonstrated in a
number of environments. In a waste stream continuously contaminated with PCP,
acclimation occurred after 3 weeks (354). The microflora, particularly the attached
bacteria, metabolized  up to 0.43 ppm influent concentration. Pure cultures were
isolated from the waste stream which were capable of mineralizing 100 mg/1 PCP in
90 hr with almost complete  chloride release.
  A soil perfusion apparatus using rice paddy soil effected disappearance  of PCP
with more than 90% liberation of chloride (459). A Pseudomonas sp. isolated from
the enrichment culture degraded 40 mg/1 PCP in 10 days with complete chloride
release.
  PCP added to moist garden soil at 150 to 200 mg/1 soil-water concentration was
25% metabolized after 12 days when the experiment was conducted using outdoor
shaded test plots (128). When a culture of Arthrobactersp. ATCC 33790 was mixed
into the soil, about 85% of the  PCP disappeared during the same time. Under
laboratory conditions addition of the bacterial culture reduced the half-life for PCP
disappearance from 12 - 14 days to 1 day. This Arthrobactersp. utilizes PCP as the
sole source of carbon and energy with complete release of chloride (130).
  Comparisons of aerobic  and anaerobic metabolism of PCP have shown that
aerobic metabolism is much more efficient (282). Enrichment cultures established in
fermentors fed with 2 mg/1 PCP revealed a half-life of 0.36  days under  aerobic
conditions and 192 days under anaerobic conditions. Addition of glucose or 4-
chlorophenol as cometabolic substrates depressed the rate of PCP metabolism.
  Soils treated with 10 mg/1  PCP and incubated for 24 days under aerobic conditions
revealed considerable loss of 14C-labeled material from the system (320). Of 59% total
recovered material, 51% was identified as pentachloroanisole. Volatile products and
CO2 were not measured. In the same system maintained under anaerobic conditions,
7% of the material was converted to  metabolites and no 14CO2 was detected.
Metabolites included  about 5% pentachloroanisole and lesser amounts of 2,3,6-
trichlorophenol, 2,3,4,5-tetrachlorophenol and 2,3,5,6-tetrachlorophenol.
  PCP  applied  to flooded paddy  soil, simulating  anaerobic conditions,  was
metabolized after 3 weeks  to the following products: 3-chlorophenol, 3,4-, and
3,5-dichlorophenol, 2,3,5-,  and  2,4,5-trichlorophenol, and  2,3,4,5-,  2,3,4,6-, and
2,3,5,6-tetrachloroanisole(217).
  The rate of PCP metabolism in 11  soils was found to be related to the  organic
matter content of the soils (266). Degradation products included a mixture of tri- and
tetrachlorophenols.
  A major PCP spill on the Mississippi River Gulf Outlet left PCP levels as high as
1.60 mg/g in the sediment (109). At 18 months there was no detectable PCP in the
sediment. Studies arising from the spill indicated that the degradation rates increased
with increasing sediment redox potential. Maximum degradation occurred at pH 8 at
+500 mV. Less degradation  occurred at pH 9 and at pH less than 8.
  PCP-degrading bacteria have been isolated both from polluted sites and from sites
not  known to be contaminated with PCP (411).  An enrichment consortium
established under continuous culture conditions became adapted to metabolize 500

                                    97

-------
mg/1PCP. Arthrobacter sp. strain NC was isolated from the culture and metabolized
100 mg/1PCP until the pH decreased to 6.15. Upon adjustment to pH 7.1 the residual
PCP was metabolized.  The strain was capable of growth at pH  6.0 upon other
substrates. Other experiments showed a correlation of toxicity with the acid form of
PCP.

SUMMARY

  In summary, there is evidence  that  PCP is attacked by bacteria and fungi
cometabolically or as sole source for growth with release of chloride and CO2. The
pathway involves  dechlorination and hydroxylation either ortho or para  to the
phenolic hydroxyl group, forming a catechol or a quinone, respectively. However,
the mechanism of this process is not understood and the enzymes involved have not
been isolated. Further, the steps of the pathway leading to carbon incorporation into
cell contents and CO2 formation have not been elucidated. In fungi, methylation has
been detected as a prominent metabolic process, but its role in PCP degradation has
not been established.
                                   98

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                              SECTION 11
 CHLOROPHENOXY AND CHLOROPHENYL HERBICIDES

   Compounds  with an arylcarboxylic acid parent structure have plant growth-
regulating  properties.  They produce  physiological effects such as morphogenic
abnormalities, promote the rooting of cuttings, and aid in setting fruit in the absence
of pollination (445).
   There are three principal chlorine-substituted phenoxyacetic acids used widely as
herbicides, (2,4-dichlorophenoxy)acetic acid (2,4-D), (2,4,5-trichlorophenoxy)acetic
acid (2,4,5-T),  and  (4-chloro-2-methylphenoxy)acetic  acid  (MCPA). They are
selective against broadleaved  weeds and woody broadleaved  plants  and are
commonly used in lawns, grass pastures, and cereal crops (283). Other phenoxy-
alkanoic acids  are useful in controlling weed species which are resistant to the
phenoxyacetic acids. The phenoxybutyric acid herbicides have very low toxicity to
plant species such as legumes which are damaged by exposure to phenqxyacetic
acids. After application, they are activated by target plants (weeds) which /3-oxidize
them to their corresponding toxic phenoxyacetic acids. Other herbicides in this class
include the phenoxyethyl esters, which are applied when deep soil penetration is
required or in noncrop areas.
   The phenoxyalkyl acid herbicides are detoxified in soils and aquatic environments
due to microbial action (13, 60, 111, 283, 343, 376, 377).  Bacteria and fungi which
metabolize the herbicides have been isolated from soils (Table 4). The products of
microbial metabolism may be phytotoxic or they may result in inactivation of the
herbicide (290,292). These products may be similar to those formed as a consequence
of plant metabolism, such as 2,4-dichlorophenol from 2,4-D and 2,4,5-trichloro-
phenol from 2,4,5-T. There is evidence that 2,4,5-trichlorophenol, rather than 2,4,5-
T, is the  active agent which damages  the  plants (283).  Some processes of
microorganisms prevent activation of the herbicides rather than actually detoxifying
them (3). For this reason the pathways by which  microorganisms metabolize the
phenoxy herbicides are of importance in determining the choice of herbicide for a
given application.

2,4-D

   The herbicide most extensively studied has been 2,4-D. Bacteria including
Pseudomonas sp., Arthrobacter  sp., Achromobacter sp., Mycoplana sp., and
Flavobacterium peregrinum, cleave the molecule at the ether linkage between the
oxygen and the aliphatic side chain to form glyoxylic acid and 2,4-dichlorophenol
(Figure 46) (31, 32, 40, 152, 285, 286, 412,  433, 434, 451). The latter compound is
metabolized to  3,5-dichlorocatechol, a's,cjs-2,4-dichloromuconic acid, 2-chloro-4-
carboxymethylene but-2-enolide and 2-chloromaleylacetic acid (46, 393).  Chloro-
maleylacetic acid is degraded further to succinic acid via 2-chloro-4-ketoadipic acid
and chlorosuccinic  acid.  The entire pathway has also been  demonstrated using
cell-free extracts of Arthrobacter sp. (40, 44, 126,  142, 393, 434). Cleavage of the
ether-oxygen bond in phenoxyacetic acid to form the phenol has been proven to

                                    99

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occur between the aliphatic side chain and the ether-oxygen in experiments with
Arthrobacter sp. using 1802 (198).
                TABLE 4. MICROORGANISMS THAT
                   METABOLIZE PHENOXY ACIDS
      Phenoxy acid
      Organism
                             References
2,4-D
Achromobacter sp.
Arthrobacter sp.
                      31,32,412,414
                      45, 126,285,434,
                      44, 46, 284, 286
Arthrobacter globiformis 283
Corynebacterium sp.    375
Flavobacterium
  peregrinum
Mycoplana sp.
Nocardia sp.
2,6-D

2-Chlorophenoxyacetic
  acid
Pseudomonas sp.

Sporocytophaga
  congregate
  (F. aquatile)
Streptomyces
  viridochromogenes

Achromobacter sp.
Achromobacter sp.
Arthrobacter sp.

F. peregrinum
                                            405a, 411 a, 412, 414
                                            451
                                            283
                                            146, 169, 170,171, 142,
                                            152, 171a
                                            225
                                            51a
                      31,32
                      31,32,412,414
                      45, 284, 285, 286,
                      44, 46, 125, 434
                      412,414
4-Chlorophenoxyacetic
  acid
Pseudomonas sp.
Achromobacter sp.
Arthrobacter sp.

F. peregrinum

Mycoplana sp.
Nocardia sp.
Pseudomonas sp.
                      142, 146, 152
                      31,32,412,414
                      45, 284, 285, 286

                      412,414

                      451
                      283
                      126, 142,143,146,152,
                      169,170, 171,171a
                                                         (continued)
                               100

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 TABLE 4. (continued)


 MCPA                 Achromobacter sp.        31,32,412,414
                         Arthrobacter sp.          45, 284, 285, 286,
                                                  44, 46, 126, 434
                         F. peregrinum            45
                         Mycoplanasp.            451
                         Pseudomonas sp.         142, 146, 152,169, 170,
                                                  171,171a

 2,4,5-T                 Achromobacter sp.        31,32
                         Brevibacterium sp.        207
                         Mycoplanasp.            451
                         S. viridochromogenes     51 a
  The  enzymes mediating the degradation of 2,4-D are relatively nonspecific.
Oxygen and  a reduced  pyridine nucleotide (NADH  or  NADPH) are required,
indicating that the enzyme(s) may be a mixed function oxidase (44). The broad
specificity of the enzymes is reflected in the findings that Pseudomonas sp. cells
grown  on 2,4-D also metabolize 4-hydroxyphenoxyacetic acid and phenoxyacetic
acid (285). Arthrobacter sp. cell-free extracts grown on 2,4-D also oxidize MCPA,
2,4-dichlorophenol, 4-chloro-2-methylphenol and 3,5-dichlorophenol.  Neither 6-
hydroxy-2,4-dichlorophenol  nor 2,4-dichloroanisole is oxidized, indicating that
neither is an  intermediate in 2,4-D metabolism  (284). The enzyme extracts also
convert 2-chlorophenoxyacetic acid to 2-chlorophenol and 4-chlorophenoxyacetic
acid to 4-chlorophenol which  subsequently forms 4-chlorocatechol. Catechol is
converted to cjs,c/s-muconic acid and 4-chlorocatechol to cjs,cis-3-chloromuconic
acid. Chloride is released from 3-chloromuconic acid to form 4-carboxymethylene
but-2-enolide, maleylacetic acid and subsequently succinic acid (Figure 47) (40, 44,
434). This pathway is analogous to that demonstrated for 3,5-dichlorocatechol
during 2,4-D degradation.
  A Corynebacterium sp. isolated from soil by enrichment culture metabolized
2,4-D with nearly complete chloride release after 48 hours (375). An application rate
to soil of 3,000 ppm was metabolized but neither growth nor metabolism was noted
upon application of 3,500 ppm. No metabolites  were seen during the incubation
period.
  A bacterial strain tentatively  identified as  F. peregrinum was isolated from
enrichment culture with 2,4-D in soil (414). This strain metabolized 100 ppm 2,4-D in
25 days and upon addition of 0.1 % yeast extract metabolized 0.1% 2,4-D in 12 to 16
days. Chloride release was estimated at 70% of that in 2,4-D within 39 days.
                                   101

-------
OCH2COOH
      COOH
    HCCI
      CH2
      C=0
      CH2
      COOH
                            glyoxylic acid
                 OH

               0-CHCOOH
                          4
                                              hydroxy-
                                              malonic
                                            semialdehyde
                                        CHO

                                        COOH

                                        C02
                                                                  COOH
                                   2,4-dichlorophenol   3,5-dichlorocatechol
                                     COOH.
     C\    POOH
NADH  Y  COOH

          Q    2-chloro-4-carboxymethylene
    2-chloromaleyl      but-2-enolide
     acetic acid
 CI^COOH
	T   COOH
                                                     V
                                                       Cl
                                            cjs,cjs-2,4-dichloromucanic
                                                      acid
    2-chloro-
 4-ketoadipic acid
 CHjCOSCoA    9OOH
	*- HCCI
ACETYL-CoA    £HZ
               COOH
                                        COOH
                                        CH2
                                        
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  f
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                                   COOH
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       Cl                 Cl
4-CHLOROCATECHOL  cis,cis-3-CHLORO-
                    MUCONIC ACID
        O
        ii
        ,C                ^^\j
                  	^ (|   COOH
                         V
                          o
4-CARBOXYMETHYLENE-   MALEYL-
    BUT-2-ENOLIDE      ACETIC ACID
                                                           COOH
                                                           CH2
                                                           9H2
                                                           COOH

                                                          SUCCINIC
                                                            ACID

-------
  Flavobacterium aquatile metabolized 0.01% 2,4-D in sterile soil, nonsterile soil,
and on solid agar plates, but not in a soil extract medium or on semisolid agar (225).
MCPA was not metabolized in sterile soil. In similar experiments, Corynebacterium
sp. metabolized both herbicides in sterile soil and on solid agar.
  A number of bacteria were isolated by enrichment culture from sewage or soil
amended with 2,4-D or 2,4,5-T (376). None utilized either substrate as the sole source
of carbon. Forty-one of 52 strains cometabolized 2,4-D only while 19 strains utilized
both 2,4-D and 2,4,5-T. Experiments with these 19 isolates incubated with 2,4,5-T in
nutrient medium showed that 12 isolates produced chloride ion and 8 produced a
phenolic compound with or without concomitant production of free chloride.

MCPA

  The  metabolism of MCPA has been studied extensively in cultures of Pseudo-
monas sp. NCIB 9340 and Arthrobactersp. (45,46,169,170,412) as well as cell-free
extracts of Arthrobactersp. (40). Initial attack on the molecule results in oxidative
cleavage of the ether linkage to form a phenol and glyoxylic acid (Figure 48) (169).
The phenol thus formed is 4-chloro-2-methylphenol (5-chloro-o-cresol). In Pseudo-
monas sp. NCIB 9340 this product is metabolized to 5-chloro-3-methylcatechol and
then to  c/s,cjs-4-chloro-2-methylmuconic  acid. The chloride ion  is  lost upon
lactonization  by dehydrochlorination to form 4-carboxymethylene-2-methyl-2,3-
butenolide and subsequently 4-hydroxy-2-methylmuconic acid (170).  Formation of
the two double bonds of the lactone is an unusual feature in the metabolism  of
aromatic  compounds by bacteria.
  The  three enzymes  mediating the  conversion of 5-chloro-3-methylcatechol  to
4-hydroxy-2-methylmuconic acid have been isolated (171). These enzymes, respon-
sible for ring cleavage, lactonization, and delactonization, confirm that lactonization
and dehalogenation is a one-step process.
  The enzymes which attack MCPA are also relatively nonspecific. Oxygen as well
as NADH or NADPH are required for enzymatic activity. F. peregrinum cells grown
in the presence of MCPA are induced to oxidize 2,4-D as detected by manometric
techniques, although MCPA is not metabolized (413).  A strain thought to be an
Achromobactersp., isolated from enrichment culture with MCPA, metabolized 50
mg/1  MCPA  or  2,4-D with addition of  0.05% yeast extract in  4 days (414).
Experiments showed a faster rate of oxygen uptake with 2,4-D than MCPA although
the organism was cultured on MCPA. Nonacclimated cultures showed no oxygen
uptake with either MCPA or 2,4-D.

2,4,5-T

  The tri-chlorinated phenoxy herbicide, 2,4,5-T, has proven much more difficult to
degrade. The utilization of 2,4,5-T by a Pseudomonas sp. appears not to be plasmid
encoded,  unlike 2,4-D and MCPA metabolism (154). The herbicide is metabolized to
2,4,5-trichlorophenol by both  P. fluorescens and P. cepacia  AC1100 (249, 377).
Brevibacterium sp. when grown on benzoic acid converts 2,4,5-T  to 3,5-dichloro-
catechol without a lag, indicating removal of one chloride ion (207).  This latter
compound can be metabolized  (Figure 46).
  Pseudomonas cepacia AC1100 is a strain modified in the laboratory which utilizes
2,4,5-T as the sole source of carbon and energy for growth (249). More than 97% of a
1 g/1 solution was degraded within 6 days with stoichiometric chloride release. In 2
hr, resting cells promoted 50% disappearance of 2,4,5-T although only 15% chloride
release was evident. Within 24  hours, there was complete substrate disappearance
with 94% chloride release. Resting cells also mediated release of more than 80% of the
chloride  from 2,4,5-trichlorophenol, 2,3,4,6-tetrachlorophenol and pentachloro-

                                    104

-------
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phenol. Additionally, resting cells were also shown by oxygen electrode deter-
mination to oxidize 2,4-D at relatively high rates but not (2,4-5-trichloro-phenoxy)
propionic acid. Phenoxyacetic acid was oxidized at low rates only. When inoculated
into soil P. cepacia AC1100 mediated 95% chloride release of a 1 mg/ g application of
2,4,5-T within 1 week (85). The optimum conditions were 25% moisture content at
30°C.

4-CHLOROPHENOXYACETIC ACID

  The degradation of 4-chlorophenoxyacetic acid by a soil pseudomonad proceeds
through 4-chloro-2-hydroxyphenoxyacetic acid to 4-chlorocatechol (Figure 49)
(143). This organism also metabolizes 4-chlorocatechol to cjs,c/s-3-chloromuconic
acid. While direct evidence for subsequent steps in the pathway was not obtained, the
culture medium did contain a lactone which is analogous to that described for
Arthrobactersp. The pseudomonad was not induced to grow on 4-chlorophenol and
this product was not found in the culture medium, indicating that this compound is
not an intermediate in 4-chlorophenoxyacetic acid metabolism.
  An unidentified gram-negative organism isolated from soil also metabolizes 4-
chlorophenoxyacetic acid through 4-chloro-2-hydroxyphenoxyacetic acid and 4-
chlorocatechol, as determined by simultaneous adaptation experiments (146). The
same  technique was used  to determine that  Achromobacter sp.  grown  on  4-
chlorophenoxyacetic acid immediately oxidizes 4-chloro-2-hydroxyphenoxyacetic
acid, 4-chlorocatechol, and catechol, but not 4-chlorophenol (414). The first step of
4-chlorophenoxyacetic acid metabolism in these pathways is hydroxylation of the
ring, followed by ether cleavage. This is in contrast to Arthrobacter sp. metabolism of
several chlorophenoxy compounds in which cleavage of the ether  linkage is the
primary step yielding 4-chlorophenol  (284).

OTHER PHENOXY HERBICIDES

  The metabolism of phenoxy herbicides with longer aliphatic side chains has also
been studied. Two mechanisms appear to mediate degradation of these compounds.
The primary  mechanism is /3-oxidation, a mechanism common to plants (283).
Evidence for /3-oxidation comes from observations with cultures of Flavobacterium
sp. which were grown on 4-(2,4-dichlorophenoxy)butyric acid (4-(2,4-D)B) and then
tested for products  arising from oxidation of higher carbon-number homologs.
Phenols were detected in all cases but more so with  compounds containing an odd
number of carbons in the side chain (291). When the side chain contained an odd
number of carbons,  primarily 2,4-dichlorophenol was recovered, while 2,4-D was
recovered from metabolism of compounds with an even number of carbons in the
side chain.  Extracts of these cultures also contained free aliphatic acids  which
indicates ether cleavage, a second mechanism of phenoxy acid degradation similar to
that shown for other herbicides (290,292). This organism when grown on 4-(2,4-D)B
also oxidizes  (as determined  by manometric techniques) 3-(2,4-D)propionic acid,
4-(4-chlorophenoxy)butyric acid, 4-(4-methyl-2-chlorophenoxy)butyric acid, 2,4-
dichlorophenol, and 4-chlorocatechol but not 2,4-D (60, 292). The failure to oxidize
2,4-D argues  against /3-oxidation as the controlling  mechanism for these degra-
dations as the product of /3-oxidation of 4-(2,4-D)B would  be  2,4-D (292). The
enzymes involved in these oxidations are adaptive rather than  constitutive. The
oxidation of these substrates led to the proposal of a pathway for the degradation of
4(2,4-D)B through 2,4-dichlorophenol to 4-chlorocatechol with loss of one chloride
ion and subsequent metabolism of 4-chlorocatechol by established pathways (Figure
50). The side chain is cleaved initially at the ether linkage and then is metabolized by
/3-oxidation (292).

                                   106

-------
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4-CHLOROCATECHOL  /3-CHLOROMUCONIC

                        ACID
                  terminal
                  respiratory
                  cycle
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                                                                        9
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                                            . n ,  .        yr -CARBOXYM1THYLENE-
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                            2,4-DICHLOROPHENOL  4-CHLOROCATECHOL
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-------
  Two species of Nocardia, N. opaca strain Tl6 and Nocardia sp. strain P2 have been
shown to use ^-oxidation for degradation of phenoxy acid homologs above the acetic
congener (461).  The corresponding chlorinated  phenols  are generated from (3-
chlorophenoxy)propionic and (4-chlorophenoxy)propionic acids. The six-carbon
homolog (2-chlorophenoxy)caproic acid is  metabolized to (2-chlorophenoxy)bu-
tyric acid and (4-methyl-2-chlorophenoxy)caproic acid is metabolized to (4-methyl-
2-chlorophenoxy)butyric  acid.  Similarly, (2,4-dichlorophenoxy)caproic acid is
metabolized  to  (2,4-dichlorophenoxy)butyric  acid. Metabolism of 4-(4-chloro-
phenoxy)butyric acid to 3-hydroxy-4-(4-chlorophenoxy)butyric acid is followed by
metabolic conversion to (4-chlorophenoxy)propionic acid.  A similar pathway is
followed by 4-(3-chlorophenoxy)butyric acid (462). These studies  with strain T,6
showed that 3-hydroxy acid intermediates appear during the metabolism of all the
o-arylobutyric acids.
  An alternative mechanism has been noted  in N. coeliaca (432). Although /3-
oxidation is operative in this organism, cr-oxidation operates in the metabolism of
compounds with 10 or 11 side-chain carbons (phenoxydecanoic acid and phenoxy-
undecanoic acid).  This process, demonstrated for nonchlorinated molecules,
involves two enzymes: (a) a peroxidase catalyzing peroxidative decarboxylation of
the fatty acid to yield CO2  and the fatty aldehyde with one less carbon, and (b) a
dehydrogenase catalyzing oxidation of the aldehyde to the corresponding acid.
  The salt of phenoxy compound, sodium 2-(2,4-dichlorophenoxy)ethyl sulfate, is
widely used in commercial formulations and  is metabolized to 2-(2,4-dichloro-
phenoxy)ethanol by both P. putida FLA and cell-free filtrates of Bacillus cereus var.
mycoides (279, 443). 2,4-D eventually appears in the B.  cereus  cell-free filtrates.
Metabolism by P. putida does not result in production of 2,4-D. The enzyme of P.
putida which breaks the oxygen-sulfur bond does not require prior activation (279).
This is a novel alkylsulfatase, as other microbial alkylsulfatases break the chain at the
carbon-oxygen bond.

FUNGAL METABOLISM OF PHENOXY HERBICIDES

  Studies on fungal metabolism of phenoxy compounds have included studies with
Aspergillus niger. Hydroxylation is a major mechanism, although not all vacant ring
sites are hydroxylated. Thus, (2,4-dichloro-5-hydroxyphenoxy)acetic acid is the
major metabolite  of 2,4-D metabolism (Figure 51).  A  minor  metabolite, (2,5-
dichloro-4-hydroxyphenoxy)acetic  acid, appears as  the result of a novel hydroxyl-
chloride replacement and chloride shift (148). The latter compound also is the only
compound  formed from metabolism of 2,5-D(148, 149). The hydroxyl-chloride shift
is similar to  that seen  in 2,4-D metabolism by many plants,  in which the major
metabolite  is (2,5-dichloro-4-hydroxyphenoxy)acetic acid and the minor metabolite
is (2,3-dichloro-4-hydroxyphenoxy)acetic acid. Both  metabolites require a hydroxyl-
chloride shift (283).
  Hendersonula toruloidea metabolizes 2,4-D with production of I4CO2 (470). In 8
weeks, 28.8% of (carbon-I)-l4C 2,4-D and  2.8% of ring-l4C 2,4-D were released.
Stachybotrys atra produced only 3% of (carbon-1 )-l4C 2,4-D as I4CO2 after 8 weeks.
  Phytophthora megasperma var. sojae metabolized 10 mg/1 4-(2,4-D)B with 45%
disappearance in 21 days, but no production of 2,4-D was noted (402). The organism
also did not metabolize  2,4-D, suggesting that /3-oxidation is  not  a primary
mechanism in the metabolism of 4-(2,4-D)B.
  MCPA is metabolized to (4-chloro-5-hydroxy-2-methylphenoxy)acetic acid by A.
niger(\49). The metabolism of 2- or 4-chlorophenoxyacetic acid by fungi does not
result in ring cleavage, in contrast to the activity of bacteria (150).  Metabolism of
(4-chlorophenoxy)acetic acid yields compounds hydroxylated  in the 2- or 3-
                                    109

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ID
C
                  OCHoCOOH
       (2,4-DICHLOROPHENOXY)
            ACETIC ACID
        OCH2COOH
 (2,4-DICHLORO-5-
HYDROXYPHENOXY)-
   ACETIC ACID
  major metabolite
        OCH2COOH
        OH
 (2,5-DICHLORO-4-
HYDROXYPHENOXY)-
   ACETICACID
  minor metabolite
t
           H,C
                 OCH2COOH
        OCHoCOOH
                                 (4-CHLORO-5-HYDROXY-
                                  2-METHYLPHENOXY)-
                                      ACETIC ACID

-------
positions, the latter a novel product. Similarly, (2-chlorophenoxy)acetic acid yields
compounds hydroxylated in the 4- or 5-positions, the latter also a novel product.
Minor products include 2-chloro-3-hydroxy and 2-chloro-6-hydroxy acids.
  The microorganisms present in soil  treated repeatedly with  herbicides were
isolated  and identified (436).  Bacteria capable of  metabolizing 2,4-D include
Arthrobacter sp., Bacillus sp.,  Pseudomonas sp., and Sarcina sp. Fungi include
Penicillium megasporum and another Penicillium sp. Bacteria which can metabolize
MCPA include Arthrobacter sp., Corynebacteriumsp., and Pseudomonas sp., and
fungi include  Fusarium  culmorum, Mucor sp., Penicillium sp., Zygorhynchus
moelleri  and four Verticillium spp.  Of these,  two bacteria and five fungi  also
metabolize 2,4-D.

METABOLISM OF PHENOXY HERBICIDES IN SOILS
  A sample of Philippine soil was treated with 2,4,5-T for 4 months, after which
2,4,5-trichlorophenol  was recovered  (377). A  mixture of microorganisms  was
removed and incubated with 2,4,5-T. Loss of 10% of substrate was recorded with
liberation of 8% of the initial radioactivity of the uniformly ring-labeled substrate as
I4CO2 in 25 days. The major metabolite was 2,4,5-trichlorophenol, which was readily
metabolized with about  75% of the chloride in this metabolite liberated as free
chloride. About 40% of this  compound was released as 14CO2 in 25 days. Products
arising from incubation of the mixed culture include 3,5-dichlorocatechol, cis,cis-
2,4-dichloromuconic acid, 2-chloro-4-carboxymethylene but-2-enolide, chlorosuc-
cinic acid, succinic acid, and 4-chlorocatechol. These products with the exception of
4-chlorocatechol, are  all found in the 2,4-D  degradative pathway subsequent to
3,5-dichlorocatechol (Figure 46).
  The fate of uniformly ring-'"C-labeled 2,4-D and 2,4,5-T was explored  in 6
different soils (304). Metabolites of 2,4,5-T included 2,4-5-trichlorophenol and 2,4-5-
trichloroanisole, while no metabolites were detected after 2,4-D incubation. About
20 to 35% of the substrates were recovered from the humic and fulvic acids and humin
fractions, indicating formation of polymeric humic substances of 2,4-D and 2,4,5-T
mediated by additional hydroxyl groups on the rings. Depending on the soil, up to
83% of 2,4-D and 71% of 2,4,5-T applied at 1 ppm concentration was converted to
i4CO2in  150 days.
  Diclorfop-methyl,  (i)-methyl 2-[4-(2,4-dichlorophenoxy)phenoxyl]  propionic
acid, undergoes rapid hydrolysis of the ester bond in field soils (297). At 1 ppm the
resultant diclorfop is  rapidly metabolized in aerobic soils with isolation of two
metabolites when the l4C-label is the chlorinated ring and one metabolite when the
label is in  the nonchlorinated ring. The  ubiquitous  metabolite was identified as
4-(2,4-dichlorophenoxy)phenol. Other experiments indicated  that intermediates
include dichlorfop acid with subsequent decarboxylation to form phenyl ether (397).
In 25 weeks, 25 to 35% of each type of labeled substrate was converted to I4CO2 (397).
In anaerobic soils diclorfop persists with no evolution of CO2 and formation of only
trace amounts of a metabolite.
  The herbicide 2-(2,4-dichlorophenoxy)ethanol is often applied to soils in the inert
form sodium 2-(2,4-dichlorophenoxy)ethyl sulfate (443). In sterile soils conversion to
the  active form occurs only at pH 3 to 4, while in nonsterile soils conversion takes
place at pH 3 to 7, within 45 minutes after application. Thus, in soils with pH greater
than 4, herbicide activation is thought to be biologically mediated.
  The isopropyl, n-butyl, and  isooctyl esters  of 2,4,5-T, the n-butyl  ester of (2,4-
dichlorophenoxy)butyric acid,  and the isooctyl ester of  (2,4-dichlorophenoxy)-
propionic acid were applied to 4 Saskatchewan soils at 4 ppm concentration (400). In
moist soils there was nearly  complete conversion of the substrate to the free acids

                                    111

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within 24 hours, with the exception of the isooctyl ester of (2,4-dichlorophenoxy)-
butyric acid, which was completely converted in 72 hours. In air-dried soils there was
very little loss of ester in the same time.
  The isooctyl ester or dimethylamine  salt of 2,4-D was  applied to soils at a
concentration of either 1.6 or 16 ppm(469). After 58 days 60 to 80% of the ring- or
carboxyl-labeled substrate was released as I4CO2, while 1% was recovered as 2,4-D.
Eighty percent of the labeled substrate in the runoff water was recovered as I4CO2
after  5 weeks, with an additional 3% more recovered in the next 5 weeks; the
remaining material was not 2,4-D.
  The primary effluent of municipal sewage  was added to a nutrient medium
containing 2,4-D (376). Within 7 days almost  all the substrate disappeared. In a
similar test phenoxyacetic acid disappeared within 12 days. Subsequent additions of
either of  these  substrates resulted  in  metabolism  without a lag period.  No
disappearance  of 2,4,5-T was noted after 60 days.
  Incubation of Maahas clay with medium containing these herbicides resulted in
90% disappearance of 2,4-D in 14 days on initial application, with 3 days required for
75% disappearance of additional applications of substrate (376). Phenoxyacetic acid
required  16 days  for initial  disappearance,  and subsequent applications were
metabolized in 4 days. Evolution of I4CO2 began from 7 to 60 days after herbicide
application, and after 4 months from 5.2 to 34% was recovered depending on the soil.
  After 12 weeks of incubation in sandy loam, 71 to 84% of either ring-'4C- or
(carbon-l)-l4C- or (carbon-2)-l4C 2,4,-D  was released  as I4CO2 (470). The concen-
tration of substrate in the soil was not given.
  Several South Vietnamese soil and  mud samples were treated with carboxyl-l4C
labeled 2,4,5-T (65). Two samples were thought to be treated previously with a 50:50
mixture of 2,4-D and 2,4,5-T, while two samples were thought to be uncontaminated.
At 1 ppm concentration, almost 70%  was evolved as I4CO2 in 49 days. At 15 ppm,
three  soils converted 70 to 80% of the substrate  to I4CO2 in 168 days, while  one
sample, thought to be previously uncontaminated, evolved more than 95% of the
material as I4CO2.
  In moist Philippine soils (upland conditions), 20 ppm 2,4-D disappeared more
rapidly than when applied to flooded soils, but after 6 weeks the concentration of
2,4-D was similar in both moist and flooded soils (482). The same results were
reported in the disappearance of 10 ppm 2,4,5-T from one of the soils. However, in
another soil 2,4,5-T remained  in flooded soils  for a 4  week lag before undergoing
rapid and complete disappearance in the  next 4 weeks, while in the  moist soils
gradual disappearance was noted with about  40% remaining after 12 weeks.  No
disappearance of 2,4-D or 2,4,5-T was noted in sterile control samples after 12 weeks'
incubation.
  The persistence of 2,4-D and MCPA in soils following repeated applications was
measured (436).  Ten weeks were required  for disappearance of 2,4-D upon first
application. The herbicide disappeared after 7 weeks upon second application in the
second year, and required only 4 weeks for disappearance after 18 years of repeated
applications. MCPA required 20 weeks for disappearance in the first year, 10 weeks
in the second year, and 7 weeks after 18 years. Soils pretreated with either herbicide
showed accelerated 2,4-D disappearance after  18 years but  not after one year of
herbicide application. Enhancement of MCPA disappearance was noted after either
one year or  18 years  of pretreatment  with  either  herbicide. The numbers of
degradative bacteria or fungi were not significantly different after 0, 1 or 18 years of
pretreatment.

  A seed bioassay (mustard or cress) showed that 2,4-D disappears  faster than
MCPA, and 2,4,5-T much slower than either, from both a light clay soil and a sandy
loam (414). The first application of 2,4-D to soil at 55 ppm concentration required 14

                                    112

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days for disappearance of herbicidal  activity, while 7 days were  required for
disappearance of the second  application at 120  ppm and 4 days upon third
application of 200 ppm.
  A soybean bioassay also indicated that 2,4,5-T lasted much longer than 2,4-D or
MCPA (111). The rate of application had no effect at application rates from 5 to 20
Ib/acre. The rate of disappearance of herbicide increased with increasing temper-
ature and increasing moisture.  Although MCPA is subject to  degradation  by
photolysis, experiments with  1  mg/1 MCPA incubated in rice water in the dark
showed disappearance of 75% in 6 days, as opposed to  15% disappearance due to
photolysis alone (405).
  Application of bifenox, methyl 5-(2,4-dichlorophenoxy)-2-nitrobenzoate, to a
greenhouse soil mix at a rate equivalent to 1.7 kg/ ha showed that after 313 days 78%
of the benzoate ring-MC-labeled  material and 67% of phenoxy ring-"C-labeled
material was bound to the soil (275). After 7 days  following initial  application very
little additional bifenox disappeared, although only 20 to 26% was bound to the soils.
The metabolites, which were identified by thin layer chromatography, included the
acid of bifenox, 5-(2,4-dichlorophenoxy)-2-nitrobenzoic acid, nitrofen (2,4-dichloro-
phenoxy-4-nitrophenyl ether), 5-(2,4-dichlorophenoxy)anthranilic acid, and other
unidentified compounds. These metabolites were also found as degradation products
in plants grown in bifenox treated soil.

CHLOROPHENYL HERBICIDES

  Pseudomonassp. strain CBS 3 utilizes 4-chlorophenylacetic acid as the sole source
of  carbon and  energy  (258).  Initially-formed  metabolites include 4-chloro-3-
hydroxyphenylacetic  acid, 3-chloro-4-hydroxyphenylacetic acid  and 4-chloro-2-
hydroxyphenylacetic acid. This strain, however, cannot grow on 3-chloro-4-hydroxy
or 4-chloro-3-hydroxyphenylacetic acid (296). Metabolism of 4-chloro-2-hydroxy-
phenylacetic acid results in formation of 4-chloro-2,3-dihydroxyphenylacetic acid.
This is not the primary pathway of substrate metabolism. Upon further incubatjon
3,4-dihydroxyphenylacetic acid  (homoprotocatechuate) appears, indicating direct
removal  and replacement of the chloride  before ring cleavage. Homoprotocate-
chuate is metabolized to homogentisic acid (2,5-dihyroxyphenylacetic acid) and then
to the meta cleavage product  maleylacetoacetate resulting from  the  action of
homogentisate 1,2-dioxygenase. An Arthrobactersp. similarly produces 4-chloro-3-
hydroxyphenylacetic acid  from 4-chlorophenylacetate,  along with  an additional
unidentified metabolite (110).
  The herbicide chlorofenprop-methyl [2-chloro-3-(4-chlorophenyl) propionic acid
methyl ester] is readily metabolized (264). In mixed cultures of soil microorganisms,
4-chlorocinnamic acid and 4-chlorobenzoic acid have been identified as metabolites.
The latter product has been recovered from soil amended with the herbicide. Two
strains, thought to be a Flavobacterium sp. and a  Brevibacterium sp., were isolated
from the soil and found to convert 4-chlorocinnamic acid to 4-chlorobenzoic acid.
Two Arthrobacter spp.  have been shown to grow on 4-chlorobenzoic as the sole
source of carbon and energy. Thus, a consortia or  a mixed soil population would be
able to mineralize chlorofenprop.

  A chlorophenyl insecticide known as SD 8280 [2-chloro-l-(2,4-dichlorophenyl)-
vinyl dimethyl phosphate] was studied with respect to its degradation in soils (372).
The major  products  were  2,4-dichlorobenzoic acid  and  l-(2,4-dichlorophenyl)-
ethanol.  Lesser amounts of 2-chloro-l-(2,4-dichlorophenyl)vinyl methyl hydrogen
phosphate and 2',4'-dichloroacetophenone were also formed. Other products were
noted but not identified, although they were shown not to be 2,4-dichlorophenol or


                                    113

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2,4-dichlorobenzyl alcohol. None of the recovered metabolites represent alterations
to the chlorinated ring of the molecule.

SUMMARY

  Most of the chlorophenoxy herbicides appear to be biodegradable to CO2 and free
chloride under the right conditions. These results have been shown for both pure
cultures and in soils containing mixed populations. Adaptation of the cultures to the
substrate is required and  results in faster disappearance of the compound.  The
persistence of some compounds in some environments indicates that under some
conditions  these herbicides could  be  considered to be  recalcitrant compounds.
Seasonal variations in herbicide degradation have also been noted (460).
  Once a population has become adapted to metabolize a substrate, however,  that
capability persists for long periods of time. Thus, yearly applications of an herbicide
are sufficient to maintain a degradative population in soil. A population adapted to
metabolize a particular substrate is often also adapted to metabolize other related
compounds. This has particular application where crop rotation is accompanied by
usage of different herbicides.
                                    114

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                             SECTION 12
   PHENYLAMIDE AND MISCELLANEOUS HERBICIDES

  The  phenylamide herbicides include the groups of phenyl ureas,  N-phenyl-
carbamates, and acylanilides. Each takes the general form R-NH-CO-X where R is a
halogenated or nonhalogenated aromatic hydrocarbon (239, 174). In urea herbi-
cides, X is an amino group with methyl, alkyl, or methoxy substituents. Carbamates
have the form whereby X is an alkoxy group. In acylanilide herbicides, X is an alkyl
group.  In many cases these herbicides are degraded to substituted anilines (305).
  The  urea herbicides are specific  inhibitors of photosynthesis,  but can have  a
selective effect because of their low water solubility and low mobility in soil. They can
be applied to kill shallow-rooted weeds while having no  effect on deeper-rooted
plants of interest. In the late 1940s, a large number of substituted urea compounds
were compared with 2,4-D for herbicidal activity and were found worthy of further
development. Originally used as industrial weed killers, they have more recently been
used in agricultural  applications (174). The carbamates comprise a wide range of
active compounds which, depending on the chemical substituents, are used for such
purposes as herbicides, insecticides, and medicinals (186a). As insecticides,  the
carbamates inhibit the action of acetylcholinesterase, an enzyme required for proper
neurotransmitter substance functioning. The degree of fit between the inhibitor and
the enzyme  lends selectivity  to the activity of the  carbamates. The  herbicidal
carbamates have  also  been used  since the 1940s. Amide herbicides (substituted
anilides)  were developed in the 1950s and are used for such crops as corn and rice.

BACTERIAL METABOLISM OF CHLORINATED ANILINES
  The degradation of such herbicides as monuron, diuron, linuron, and propanil
results  in formation of 3,4-dichloroaniline. A strain of P. putida has been isolated
which mineralizes 3,4-dichloroaniline in the presence of aniline (483). The rate of
mineralization was enhanced by  increasing the concentration of aniline. In  the
presence  of 500 mg/1 propionanilide, as much as 50% of ring labeled chloroaniline
(added at 10 to 60 mg/1 concentration) was metabolized to 14CO2 within 2 weeks,
accompanied by some chloride release (484). A pathway for 3,4-dichloroaniline was
proposed which involves ortho cleavage through 4,5-dichlorocatechol and further
metabolism to succinic acid (Figure 52). This pathway is analogous to that shown for
aniline. Dichloroaniline is also metabolized by a different  mechanism to dichloro-
formylanilide (245, 484).
  Another Pseudomonas sp., strain G, mineralizes 3,4-dichloroaniline to CO2 when
grown  in the presence  of 4-chloroaniline  (488).  In 9 days, 15%  of 0.5 mM
dichloroaniline was converted to CO2.
  Studies with 4-chloroaniline have shown that this metabolite is utilized as a sole
source of carbon and nitrogen by Pseudomonassp. strain G (487). After 10 days, 64%
of a 2.5 ppm solution of the substrate was released as I4CO2. Ammonium cation,
rather than nitrate or nitrite anions, accumulates suggesting that the amino group is
removed directly without oxidation. Other l4C-labeled products accumulate but are
                                   115

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         3,4-DICHLORO-
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-------
not incorporated into the cell biomass. Resting cells of this strain grown on 4-
chloroaniline  also  oxidize  aniline,  catechol,  and  4-chlorocatechol but not 4-
chloronitrobenzene or 4-chlorophenol, indicating  dioxygenase attack on  the
molecule (487). The meta-cleavage metabolite 2-hydroxy-5-chloromuconic semi-
aldehyde also  accumulates in the medium. This strain also utilizes 2-chloroaniline
and 3-chloroaniline as sole sources of carbon and nitrogen. P. multivorans strain An
1, when grown with aniline present in the medium, converts 2-chloroaniline to
3-chlorocatechol, and 3-chloroaniline and 4-chloroaniline both to 4-chlorocatechol,
which is metabolized subsequently to CO2 and cell constituents (361).
  A strain of Alcaligenes faecalis utilizes 3-chloroaniline or 4-chloroaniline under
cooxidative  conditions with sodium  acetate  or sodium pyruvate (419).  Both
chloroanilines are oxidized to 4-chlorocatechol, which undergoes meta cleavage to
5-chloro-2-hydroxymuconic  semialdehyde and then to  2-chloro-4-oxalocrotonic
acid. Similarly, an  unidentified isolate which utilizes 4-chloro-aniline as the sole
carbon source demonstrated, via Warburg respirometry,  oxidation of 4-chloro-
catechol without a lag (56).
  The metabolism of 4-chloroaniline by Paracoccus sp. under  both aerobic and
anaerobic conditions was investigated (43).  Transformation was faster under
anaerobic conditions with 100% of a 20 mg/1 solution converted within 2 days to a
volatile  product  plus several other metabolites. The volatile  product was not
identified but was shown not  to be CO2. In the same period, 75% of the substrate was
utilized aerobically, although the aerobic population was  larger than the anaerobic
population.  The  major product  of aerobic  metabolism  is 4-chloroacetanilide,
although several other products are formed as well. Paracoccus sp. also transforms
2-, 3-, 4-chloroaniline, 2,3-, 2,4-, 2,5-, and 3,4-dichloroaniline both aerobically and
anaerobically, with more complete primary degradation occurring under anaerobic
conditions.
  A study of  the anaerobic  conversion of 4-chloroaniline by the  same  organism
showed pH-dependent formation of products (310). In a medium containing nitrate,
80% of a 100 ppm solution was transformed within 48 hours to the condensation
product l,3-bis(p-chlorophenyl)triazine. However, sterile anaerobic solutions at pH
of 5 to 6 also yielded this product, although no transformation took place at pH 7. It
was postulated that the conversion of nitrate to nitrite and decrease in pH were the
primary effects of bacterial metabolism, while triazine formation was a nonbiological
secondary effect. Paracoccus sp. also formed small amounts of 4-chloroacetanilide
during incubation with 4-chloroaniline.
  A wide variety  of both Gram-positive and Gram-negative bacteria were isolated
from soil which had been enriched with 4-chloroaniline (134). The most active species
was identified  as Bacillus firmus. This species could not use the substrate as the sole
source of carbon, but when grown with ethanol transformed 4-chloroaniline to
4-chloroacetanilide  as the  main  metabolite,  with  lesser  amounts  of  4-chloro-
propionanilide also produced. Two other products were identified as 7-chloro-2-
amino-3H-phenoxyazine-3-oneand7-chloro-2-amino-3H-3-hydroxyphenoxyazine,
and were postulated to result from spontaneous condensation of 4-chloroaniline with
subsequent  hydroxylation.  The  acylation of the  substrate described here is
considered to be a detoxification process in microorganisms (134).
  Aniline-grown  resting cells of Rhodococcus sp. An 117 convert 2-chloro-  and
3-chloroaniline to  3-chloro- and  4-chlorocatechol,  respectively.  An additional
product identified as 2-chloromuconic acid results from  metabolism of 2-chloro-
aniline.  Cometabolism  of 3-chloroaniline  in  the presence of 18O2 resulted in
production  of another product  which was  identified  as the g-lactone of 3-
hydroxymuconic  acid, formed by incorporation of two molecules of oxygen (223).
Appearance of this product was associated with disappearance of 4-chloro-catechol.

                                     117

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This suggests that  dechlorination  is associated  with lactonization of  the  3-
chloromuconic acid, a mechanism shown previously in pseudomonads. When fresh
cultures were supplied with benzoate plus 2- or 3-chloroaniline as the sole source of
nitrogen, growth of cells and disappearance of both substrates occurred (222). The
growth yield  was similar to that obtained when NH4NO3 was utilized as the sole
nitrogen source. No  growth occurred when  4-chloroaniline was supplied  as the
nitrogen source.
  The  fungicide  2,6-dichloro-4-nitroaniline  is  metabolized by  many  bacteria
including E. coli and P. cepacia (442). The first step is reduction of the nitro group to
an amine,  forming 2,6-dichloro-p-phenylenediamine.  This compound is then
acetylated to produce  4-amino-3,5-dichloroacetanilide. The first  reductive  step
occurs much faster under anaerobic conditions than under aerobic conditions.

FUNGAL METABOLISM OF  CHLORINATED ANILINES

  Fungi as well as bacteria produce peroxidases  which are responsible for the
polymerization of chloroanilines. Species with this capability include Geotrichum
candidum L-3 and Aspergillus sp.  (47, 271).  The substrate 3,4-dichloroaniline is
converted to 3,3',4,4'-tetrachloroazobenzene by G. candidum L-3 and Aspergillussp.
(271) and is converted to 3,3',4,4'-tetrachloro-azoxybenzene in Fusarium oxysporum
cultures (243). However, this latter reaction varies with the culture conditions and the
azoxy condensation product has not been found in soils.
  The fungal metabolism of 4-chloroaniline follows several pathways (Figure 53). A
major pathway is N-hydroxylation such as is demonstrated by F. oxysporum (242).
This species metabolizes 4-chloroaniline to 4-chlorophenylhydroxylamine which is
subsequently  converted to  4-chloronitrosobenzene  and  4-chloronitro-benzene.
Condensation products which appear include both 4,4'-dichloroazo-benzene and
4,4'-dichloroazoxybenzene. In addition, 4-chloroacetanilide appears as an acylation
reaction  and  may undergo hydrolysis to yield 4-chloroaniline.  Free chloride  is
produced as the result of some of these reactions.
  The culture filtrate of G. candidum, as well as the purified fungal enzymes
peroxidase and aniline oxidase, converts 4-chloroaniline to several condensation
products including 4,4'-dichloroazobenzene and 4-chloro-4'-(4-chloroanilino)-axo-
benzene  (49). These reactions have also  been demonstrated with  horseradish
peroxidase.  Streptomyces  sp. also  formylates 4-chloroaniline  with  resultant
production of 4-chloroformylanilide as well as 4-chloroacetanilide and at least two
other metabolites (381).
  Ring hydroxylation is a mechanism demonstrated by F. oxysporum which  results
in metabolism of 3-chloroaniline to 2-amino-4-chlorophenol and 4-chloroaniline  to
2-amino-5-chlorophenol (Figure 53) (155). These molecules are hydroxylated in the
ortho position.  The  aminophenols  are  relatively  unstable and can undergo
condensation and polymerization reactions, although 2-amino-4-chlorophenol has
been detected in soil as well  as in  the pure culture studies reported above.  The
ort/jo-substituted substrate 2-chloroaniline is not hydroxylated by F. oxysporum.

METABOLISM OF CHLORINATED ANILINES IN SOILS

  There is evidence that in soils 3,4-dichloroaniline is slowly mineralized. Radio-
carbon-labeled humic-bound material is mineralized in some soils (14CO2 pro-
duction) at about the same rate as the average soil organic matter polymer (384). The
addition of aniline to soils enhances mineralization of both free and humic-bound
3,4-dichloroaniline (483). This was attributed to selection by the aniline analogue for
chloroaniline-degradative populations as well as the induction by aniline of the
common metabolic pathway.

                                    118

-------
                                 -NHCOCH
4-CHLOROANILINE
                       4-CHLOROACETANILIDE
4-CHLOROPHENYL-
 HYDROXYLAMINE
                    2-AMINO-5-CHLOROPHENOL
4-CHLORONITROSO
  BENZENE
                        4,4'-DICHLOROAZOBENZENE
                                                2-ACETAMIDE-5-
                                                CHLOROPHENOL
 Cl
 4-CHLORONITRO-
  BENZENE
                         CKO)~N=N-(O)-C|
                       4,4'-DICHLOROAZOXYBENZENE
                    4-CHLORO-4'-(CHLOROANILINO)-AZOBENZENE



      Figure 53.  Metabolism of 4-chloroaniline by microorganisms.

               Adapted from References 49, 155, 242.
                               119

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  Application of  10  ppm ring-labeled i4C-3,4-dichloroaniline to a rice paddy
ecosystem yielded a total recovery (extractable plus nonextractable) of almost 69%
(219). Less than 4% of the material applied to soil was recovered in the water or rice
plants. CO2 evolution was not determined. Extraction of the soil yielded 3.4% of the
original material as dichloroaniline, 4% as tetrachloro-azobenzene, and 44.7% as
polar material.
  The degradation of aniline, 4-chloroaniline,  and 3,4-dichloroaniline  in  four
different soils was compared (420). At 1 ppm application rate, 16 to 26% aniline was
converted to CO2 after 10 weeks, while after 16 weeks 12 to 27% chloroaniline and 4
to 12% dichloroaniline were mineralized. In soils 4-chloroaniline can be converted to
4-chlorophenylhydroxylamine via biological mechanisms (48). This compound then
undergoes condensation with 4-chloroaniline to form 4,4'-dichloroazobenzene in a
nonbiological reaction.
  Condensation of two molecules of 3,4-dichloroaniline forms 3,3',4,4'-tetrachloro-
azobenzene which  is relatively persistent in the environment (51). This reaction has
been demonstrated in Nixon sandy loam (11, 23, 24) as well as in other soils (22,48,
51, 245). The conversion of chloroanilines to chloroazobenzenes has been shown to
occur by a peroxidase mechanism (25). A mixture of substituted anilines, hydrogen
peroxide,  and peroxidase  resulted  in  the formation  of chloroazobenzene  (51).
Bacteria including Bacillus sp., Arthrobacter sp., and Pseudomonas sp. exhibit
peroxidase activity (271). A pathway  has been proposed which includes trans-
formation of the hypothetical 3,4-dichloroanilidyl molecules to 3,4-dichlorophenyl-
hydroxylamine. Two of these molecules are condensed to the hypothetical 3,3',4,4'-
tetrachlorohydrazobenzene which is  converted to 3,3',4-4'-tetrachloroazobenzene
(51). However, studies in which dichloroaniline was applied to herbicide-treated soils
did not show formation of tetrachloroazobenzene, suggesting that dichloroaniline is
not the prime precursor  for  tetrachloroazobenzene  (30). Also,  production  of
tetrachloroazobenzene in soils incubated with propanil was not correlated with the
quantity of  peroxidase-producing microorganisms recovered from the soil  (58).
Cell-free perioxidase  was found  only rarely in  the soil samples. Recovery  of
peroxidase increased  upon amendment  of  the  soils  with nutrient sources and
additionally upon sonification of the samples, which may have released cell-bound or
intracellular enzymes. Addition of proteose-peptone decreased recovery of peroxi-
dase-producing organisms but increased soil peroxidase  recovery.  However, this
peroxidase did not form  tetrachloroazobenzene from dichloroaniline.  Another
condensation product detected as a humic-bound residue in soils treated  with
propanil is 4-(3,4-dichloroanilino)-3,3',4'-trichloroazobenzene (22).
  Warburg respirometry studies indicated that aniline-acclimated activated sludge
microflora oxidized 500 mg/12-chloroaniline at low rates over a 192-hour incubation
period (293). Oxidation of 4-chloroaniline occurred at slightly higher rates, and after
a 100-hour lag period  rapid oxidation of 3-chloroaniline took place.

METABOLISM OF UREA HERBICIDES

  A Fusarium sp.  utilizes  the larvacide diflubenzuron [l-(4-chlorophenyl)-3-(2,6-
difluorobenzoyl)urea]  as its sole source of carbon and energy (389). The pathway was
elucidated to include  initial formation  of 2,6-difluorobenzoic acid and 4-chloro-
phenylurea.  The latter compound is  metabolized to 4-chloroaniline, and then 4-
chloroacetanilide, followed by reductive dehalogenation to acetanilide and further
metabolism to cell constituents. Other fungi including Cephalospocium sp.,
Penicillium sp., and Rhodotorula sp., although unable to utilize diflubenzuron as a
sole carbon  source, metabolized the compound to 2,6-difluorobenzoic acid and
4-chlorophenylurea, indicating cleavage of the urea bridge.
                                    120

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  Evidence that the urea herbicides are degraded in the environment accrues from
studies which showed that such compounds as 3-(4-chlorophenyl)-l, 1 -dimethylurea,
(monuron) and 3-(3,4-dichlorophenyl)-l,l-dimethylurea (diuron), when applied at
rates of 1 to 2 lb/ acre annually in the eastern part of the United States, left no residual
phytotoxicity after 4 to 8 months (203). Higher application rates required longer
times for disappearance of phytotoxicity.
  Potential  mechanisms for disappearance include biological, leaching, volatili-
zation and chemical decomposition (203). Leaching is considered to be significant
only in porous soil or if there is a great amount of rainfall. The low vapor pressure
and aqueous solubilities of these herbicides make volatilization unlikely to be an
important mechanism. Photodecomposition may be a factor in dry areas in which the
herbicide remains on the soil surface, but these compounds are stable to chemical
decomposition  in aqueous  solutions. Biological studies revealed that the  rate of
herbicide inactivation is greater in nonsterile than in sterile soils. This was shown by
the amount of radiocarbon-labeled CO2 evolved  from soils  amended with
l4C(methyl)-labeled monuron, and a Pseudomonas sp. was isolated which oxidized
this substrate in Warburg respirometry studies (203).
  The  herbicide N'-(4-chlorophenoxy)phenyl-N,N-dimethylurea is metabolized
when placed  in contact with soils such as sandy loam or humus soil (173). Sorption of
the compound to the soils has an effect on the rate of degradation. Enriched bacterial
cultures derived from the soil samples also metabolized the herbicide by successive
demethylation to N'-(4-chlorophenoxy)phenyl-N-methylurea and subsequently to
N'-(4-chlorophenoxy)phenylurea. No appreciable amounts of CO2 were released.
Fungal isolates  of Penicillium sp. and Aspergillus sp. removed less than 50% of the
carbonyl group but did not transform the compound further. The metabolism of this
herbicide in  plants also follows successive demethylation leading to CO2 evolution.
  Diuron  is used for long-term  weed control in peach orchards, and has been
detected as long as 3 years after the last application in a field consisting of Fox loamy
sand (248).  The levels of  3,4-dichloroaniline were very low and  decreased  to
undetectable levels in 3 years, while the potential condensation product tetrachloro-
azobenzene was not detected. The decomposition of diuron is enhanced by changes
in environmental conditions that  favor the growth of microorganisms (306). Thus,
increasing the temperature of incubation from 10 to 30°C or adding organic matter to
the soils both increase the rate of diuron decomposition.  The rate of herbicide
inactivation  is much greater than the rate of CO2 evolution, and investigations
showed that  the loss of one methyl group decreases herbicidal activity by half while
loss of both methyl groups completely inactivates the molecule. The pathway of
diuron metabolism in soils was proposed to be 3-(3,4-dichlorophenyl)-l,l-dimethyl-
ureato 3-(3,4-dichlorophenyl)-l-methylurea, then loss of the second methyl group to
form 3-(3,4-di-chlorophenyl)urea, followed by hydrolysis of the urea to form the
aniline derivative 3,4-dichloroaniline, which accumulates as the major product (103).
  Microbial enrichment cultures  from pond water and pond sediment treated with
diuron revealed a similar pathway of degradation (131, 132). Three  additional
unidentified products were also detected. The fenrichment cultures included mixtures
of fungi and bacteria as well as consortia of bacteria. Some of the mixed cultures
converted  more than 90%  of the substrate to CO2 within 3  weeks. Of 20 single
isolates, however, only 3 could partially metabolize diuron after 4 weeks' incubation.
  Under anaerobic conditions reductive ring dechlorination of diuron occurs (12).
Enrichment  cultures from pond water and sediment incubated anaerobically rapidly
degraded diuron to 3-(3-chlorophenyl)-1,1 -dimethylurea in stoichiometric amounts.
This product was  not degraded  further and no other products were detected.
Repeated  additions of diuron resulted in rapid metabolism of the substrate.
                                    121
                                    AWBERC  LIBRARY U.S.  EPA

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   Chlortoluron [N-(3-chloro-4-methylphenyl)-N'-dimethylurea] has a half-life in a
 variety of soils of 4 to 6 weeks (403). The major degradation product is monomethyl
 chlortoluron.  However, the product  of subsequent  demethylation was  never
 detected. This may indicate that  cleavage of the molecule to form 3-chloro-4-
 methylaniline  follows monomethyl chlortoluron formation. Although the sub-
 stituted aniline was not detected, this product rapidly disappears from the soils when
 applied directly.
   Methoxy phenyl  urea herbicides have  quite high herbicidal selectivity and
 additionally are  not very persistent in soil after application (174). Metabolism of
 these compounds differs from that  of the dimethyl phenyl urea herbicides.
   Soils treated  with  3-(4-chlorophenyl)-l-methoxy-l-methylurea (monolinuron)
 yielded  a Bacillus sphaericus isolate which could cometabolize l4C(ureido)-mono-
 linuron to I4CO2 and 4-chloroaniline (452). Maximum degradation occurred after
 the end of logarithmic growth. The substituted aniline was not degraded by  the
 organism but was lost from the culture  through volatilization.  Linuron [3-(3,4-
 dichlorophenyl)-l-methoxy-l-methylurea] was  metabolized  to stoichiometric
 amounts of 3,4-dichloroaniline, while the dimethyl compounds monuron and diuron
 were not degraded. Cell-free extracts of B.  sphaericus also transformed the methoxy
 herbicides to substituted anilines with the aliphatic portion of the molecule degraded
 to CO2 plus another metabolite (453). The degradation product of linuron was
 identified as N,O-dimethylhydroxylamine (133). The cell-free extracts were less
 active against the dimethyl substrates monuron and diuron (453).
   The  soil  fungus  Cunninghamella  echinulata Thaxter degrades  linuron and
 monolinuron  through stable  hydroxymethyl  intermediates  (435). Linuron is
 metabolized to 3-(3,4-dichlorophenyl)-l-methoxy-l-hydroxymethylurea and subse-
 quently 3-(3,4-dichlorophenyl)-l-methoxyurea and 3-(3,4-dichlorophenyl)-l-methyl-
 urea. Disappearance of compounds such  as linuron from nonsterile, but not from
 sterile, soils has  been noted (120).  The degradation of monolinuron during waste
 composting was investigated (318). After three weeks of composting, N-methoxy-N'-
 4-chlorophenylurea  was present in trace amounts  and was the only metabolite
 detected.

 METABOLISM OF CHLORINATED PHENYL CARBAMATE
 HERBICIDES

   The phenyl carbamate herbicides (also known as carbanilates) are used to kill
 weeds on crop plants such as rice.  Substituted anilines arise from degradation of
 these compounds, as the primary mechanism of degradation seems to be hydrolysis
 of the ester linkage (239).
   The  cell-free enzyme  extract from  a  Pseudomonas sp. isolated from a soil
 enrichment culture was capable of converting several chlorophenylcarbamates to
 corresponding chlorinated anilines (244). The compounds tested included CIPC,
 [isopropyl-N-(3,4-dichlorophenyl)carbamate],sec-butyl-N-(3,4-dichlorophenyl)-car-
 bamate, a-carboisopropoxyethyl-N-(3-chlorophenyl)carbamate,  2-chloroethyl-N-
 (3-chlorophenyi)carbamate, 2-( 1 -chloropropyl)-N-(3-chlorophenyl)carbamate, 2-
 ethylhexyl-N-(3-chlorophenyl)carbamate  and  a-carbo-(2,4-
 dichlorophenoxyethoxy)-ethyl-N-(3-chlorophenyl)carbamate. In  contrast,  the
 enzyme preparation had no activity against 3-(4-chlorophenyl)-l,l-dimethylurea
 (monuron).
   Penicillium jenseni was isolated  from soil which  had been treated with barban
 [(3-chlorophenyl)carbamic acid 4-chloro-2-butyl ester] (472).  The mold did  not
utilize barban  as a  carbon source, although trace  amounts  of  3-chloroaniline
appeared after incubation.  Incubation of the mycelia with barban resulted in

                                   122

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production of large amounts of 3-chloroaniline, which was metabolized by the
mycelia suspensions without production of free chloride.
  A mixture of  bacteria and fungi isolated from soil enriched with the non-
chlorinated analogue isopropyl  phenylcarbamate (propham) metabolized a wide
range of herbicides to their corresponding chlorinated aniline products (305). These
included compounds ring-substituted with 3-chloro-, 4-chloro-, 2,4-dichloro-, 3,4-
dichloro-, and 2,4,5-trichloro- substituents. Microorganisms comprising the con-
sortium  included Mycobacterium sp., Arthrobacter sp.,  Corynebacterium sp.,
Fusarium sp., Nocardia sp., Streptomyces sp., Aspergillus sp., and Penicillium sp.
  A wide variety of fungal  isolates from treated soil metabolized swep [methyl
N-(3,4-dichlorophenyl)carbamate] to 3,4-dichloroaniline with formation of trace
amounts of 3,3',4,4'-tetrachloroazobenzene (240). The fungi included Aspergillus
ustus, A. versicolor, Fusarium oxysporum, F. solani, Penicillium chrisogenum, P.
nigulosum and Trichoderma viride. The isolates were most active on CIPC and only
slightly active against the phenylureas diuron and 3-(3,4-dichlorophenyl)-l-methyl-
urea. The  rate of  formation of 3-chloroaniline was the same  as the  rate of
disappearance of CIPC due to metabolism by Aspergillus fumigatus, indicating that
hydrolysis of the ester bond is the first step in degradation of this herbicide (471).
  Swep is metabolized to 3,4-dichloroaniline with formation of 3,3',4,4'-tetrachloro-
azobenzene in soils such as Nixon sandy loam (11). Soil microorganisms obtained
from muck soil metabolized 1240 mg/1 isopropyl-N-(3-chloro-phenyl)carbamate
(CIPC, chlorpropham) to 3-chloroaniline with complete chloride release within 13
days (241). A  similar pathway with complete dechlorination within 16 days was
observed in the metabolism of 2-chloroethyl-N-(3-chloro-phenyl)carbamate (CEPC).
The isolates lost degradative capability if they were maintained on nutrient agar for
several days but they could be readapted to use these herbicides as a sole source of
carbon.

METABOLISM OF ACYL ANILIDE HERBICIDES

  The substituted anilide herbicides are structurally related to the urea herbicides
and the carbanilates, and like them are degraded to substituted anilines.
  Strains ofPseudomonasstriataand Achromobactersp. metabolize N-(3-chloro-4-
methylphenyl)-2-methylpentanamide(3'-chloro-2-methyl-p-valero-toluidide, solan)
to 3-chloro-p-toluidine from which  chloride is released quantitatively (Figure 54)
(240). Azobenzene products were not detected in these experiments.
  Fungi from several genera, including Aspergillus spp., Fusarium spp., Penicillium
spp., and Trichoderma  sp.  also metabolize solan  to 3-chloro-p-toluidine with
chloride release (240). Cultures of A. niger metabolize solan to a product identified as
3'-chloro-4'-methylacetamlide (Figure 45) (455). Cell-free extracts converted solan to
the substituted aniline, but in growing cultures this product was rapidly acetylated to
effect detoxification and the  free  aniline could not be recovered.
  Strains of P. striata and Achromobacter sp. metabolize both  swep and N-(3,4-
dichlorophenyl)propionamide (propanil) to 3,4-dichloroaniline and small quantities
of 3,3',4,4'-tetrachloroazobenzene (240). Corynebacterium pseudodiphtheriticum
NCIB  10803 utilizes propanil as  the sole source of carbon and energy for growth
(186).  The resulting products are 3,4-dichloroaniline which accumulates  in the
medium, and the propionic acid moiety which is utilized for cell growth.
  A strain of F. solani was isolated which utilizes propanil as a sole source of carbon
and energy (269).  Dichloroaniline accumulates in the medium until it reaches toxic
levels.  The enzyme responsible for propanil hydrolysis to propionate and dichloro-
aniline was identified as an acylamidase (270). This enzyme is specific for molecules
with a short chain,  as it could not hydrolyze dicryl [N-(3,4-dichlorophenyl)metha-


                                      123

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                                   H3C-/OVNHCOCH3H7

                                      SOLAN
          3-CHLORQ-p-TOLUIDINE
                                                 3-CHLORO-4-METHYLACETANILIDE

-------
crylamide] or the six-carbon herbicide 2-methylpentanamide. Other fungi, including
Aspergillus ustus,  A. versicolor,  Fusarium  oxysporum,  F. solani, Penicillium
cnrysogenum, P. nigulosum and Trichoderma viride, also metabolize propanil to
3,4-dichloroaniline with formation of trace amounts of 3,3',4,4'-tetrachloroazo-
benzene (240).
   1 he interaction of Penicillium piscarium and Geotrichum candidum incubated
with propanil results in increased growth over either  alone  (50).  P.  piscarium
contains an acylamidase which converts propanil to dichloroaniline. G. candidum
cannot utilize propanil but contains a peroxidase which converts dichloroaniline to
the tetrachloroazobenzene. Each  fungus reduces  the toxic level  of the  other's
byproduct of metabolism, demonstrating a synergistic or mutualistic interaction.
  The yeast Pullularia pullulans and two Penicillium spp. were isolated and found to
utilize N-(3,4-dichlorophenyl)-2-methylpentanamide as a sole source of carbon and
energy, metabolizing the herbicide to dichloroaniline and 2-methyl-valeric acid
(392).  The enzyme was inducible  and differed in  level  of activity and substrate
specificity among the three  species. Cell-free extracts of one of the Penicillium spp.
hydrolyzed a wide variety of other  acyl anilides as well, but had no activity  against
diuron or CIPC.
  An unusual product, identified as N-(3,4-dichlorophenyl)-2-methyl-2,3-dihydroxy-
propionamide, was detected in the culture medium of a Rhizopusjaponicus culture
growing in the  presence of dicryl (456). This product results from the  double
hydroxylation of the ethylene double bond of dicryl, and was the only metabolite
detected. R. japonicus also hydroxylates  the side chain of N-(3,4-dichlorophenyl)-
pentanamide to  produce N-(3,4-dichlorophenyl)-3-hydroxy-2-methylpentanamide
(457). This mechanism results in detoxification of the herbicide.
  Since propanil is an inhibitor of photosynthesis it is a potential poison to the algae
as well (473). A study of the effect of propanil on cyanobacteria indicated depression
of photosynthesis but also showed conversion of propanil to the less toxic 3,4-
dichloroaniline in both axenic (bacteria-free) and contaminated cultures. The species
studied incorporated those found in flooded rice paddy fields and soil, and included
Anabaenacylindrica, A. variabilis, Nostocmuscorum, N. entophytum, Tolypothrix
tenuis, and Gloeocapsa alpicola.
  In soils propanil is converted to 3,4-dichloroaniline and the condensation product
3,3',4,4'-tetrachloroazobenzene (30, 87). Uniformly-labeled 14C-propanil applied to
soils was transformed to 3,4-dichloroaniline and the multiple condensation product
4-(3,4-dichloroanilino)-3,3',4'-trichloroazobenzene which accumulated to 2% of the
substrate (22). At high concentration (500 mg/1), the dichloroaniline was volatile,
while at lower concentrations  (5 to 10 mg/1) dichloroaniline and its condensation
product were humic-bound. The aliphatic portion is degraded to CO2. Studies with
l4C(carbonyl)-propanil revealed 70% conversion to I4CO2 within 25 days, while soils
amended with l4C(ring)-propanil yielded only 3% 14CO2 during the same time period
(87). Pseudomonas sp. strain G also converts propanil to 3,4-dichloroaniline (488).
  Propanil sprayed onto flooded rice plots was dissipated quickly and disappeared
within 24 hours (112). The major metabolite was 3,4-dichloroaniline which sorbed to
soils. Only a trace of 3,3',4,4'-tetrachloroazobenzene was detected, although dilution
caused by the flooding may have precluded condensation of the dichloroaniline.
  The  amount of tetrachloroazobenzene  formed in soils as a result of propanil or
3,4-dichloroaniline  application was  found to be highly variable (213). In  a
comparison of 9 soils, those  at a pH of 4.5 to 5.5 showed the most production.
Tetrachloroazobenzene  formation was not correlated  with the organic  matter
content of the soils, and air-dried soil samples showed 87 to 99% reduction in product
formation. More tetrachlorozobenzene was produced from direct applicaton of
                                    125

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 dichloroaniline than from molar equivalent application of propanil. In a contra-
 dictory study, however, more tetrachloroazobenzene was produced from propanil
 than from the equivalent application of 3,4-dichloroaniline (58).
   The degradation rates of herbicides in Nixon sandy loam have been correlated with
 the number of carbon atoms in the side chain (23). The four-carbon molecule dicryl is
 metabolized  at a slower rate than propanil, and the six-carbon herbicide N-(3,4-
 dichlorophenyl)-2-methylpentanamide is the most persistent of the three. Each is
 converted to 3,4-dichloroaniline, 3,3',4,4'-tetrachloroazobenzene  and  another
 metabolite.

 MISCELLANEOUS PESTICIDES
   Chlordimeform[N'-(4-chloro-e-methylphenyl)-N,N-dimethylmethanimideaniide]
 represents another class of formamide compounds used as insecticides. Chlor-
 dimeform is metabolized in  sandy loam to  several products within 90 days (220).
 These include 4-chloro-6-nitro-o-toluidine and 4-chloro-o-toluidine (Figure 46). The
 latter product is condensed  to form 4,4'-dichloro-2,2'-dimethylazobenzene. Other
 coupling products which were detected include N-(4-chloro-o-tolyl)-2-methyl-p-
 benzoquinone monoimine  and 2-(4-chloro-o-toluidino)-N-(4-chloro-o-tolyl)-6-
 methyl-p-benzoquinone monoimine. These are formed by a one-electron oxidation
 mediated by peroxidases, and appearance of these products is pH-dependent.
   A mixed population of soil microorganisms mediated a novel conjugation of the
 aniline  moiety of chlordimeform with malonic acid to form 4'-chloro-2'-methyl-
 malonanilic acid (Figure 55) (378). This mechanism was previously found only in
 plants as a means for detoxification of certain D-amino  acids.
   The herbicide N-( 1,1 -dimethylpropynyl)-3,5-dichlorobenzamide underwent exten-
sive metabolism of the side chain during 90 days'incubation in soils (481). However,
 no alteration of the chlorinated ring structure was noted. The cyanobacterium
 Oscillatoriasp. metabolized N'-(4-chloro-o-tolyl)-N,N-dimenthylformamidine with
 production of 14CO2  from  either tolyl-14C- or ring-14C-labeled  substrate (34).
 Extensive nonbiological degradation of this compound occurs but  no evolution of
 14CO2 in the absence of the algae was noted.
   Techlofthalam  [N-(2,3-dichlorophenyl)-3,4,5,6-tetrachlorophthalamic acid] is a
 bactericide used on rice plants (252). An analysis of its fate under flooded soil
 conditions analogous to those of rice paddy fields indicated that after 32 weeks most
 of the recoverable  radiolabeled material was  isolated  as two or more  products
 chlorinated in the tetrachlorophthalamic ring. Nine percent of the carboxyl-labeled
 material was converted to 14CO2.  Techlofthalam was  recovered as  a minor
 metabolite. No further transformations occurred during  the  32 weeks of the
 experiments.
   Metabolism of chlornethoxynil  (2,4-dichlorophenyl-3'-methoxy-4'-nitrophenyl
 ether) in flooded soil resulted in production of a number of compounds due to
 alteration of the molecule without loss of chloride (328).  Cleavage of the ether bond
 results in production of 2,4-dichlorophenol.
   Production of amino derivatives from other diphenyl ether herbicides was shown
 to be faster in flooded than in moist soils (329). A number of bacteria including those
 from the genera Bacillus, Pseudomonas, Enterobactera.nd Escherichiamediated this
 conversion, although disappearance of the  herbicides was noted in sterile soils as
 well.  The  herbicides tested  included  nitrofen  (2,4-dichlorophenyl 4'-nitrophenyl
 ether),  2,4,6-trichlorophenyl 4'-nitrophenyl ether, 2,4-dichloro-6-fluorophenyl 4'-
 nitrophenyl ether, and chlomethoxynil.
                                    126

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         CHLORDIMEFORM
         4'-CHLORO-2'-METHYL-
          MALONANILIC ACID
                                                                        NO2
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                                         4-CHLORO-o-TOLUIDINE
                                               CH3
 4,4'-DICHLORO-2,2'-
DIMETHYLAZOBENZENE
                                                         N02
                                             4-CHLORO-6-NITRO-
                                                o-TOLUIDINE
                                                                    2-(4-CHLORO-o.-TOLUIDINO)-
                                                                     N-(4-CHLORO-o.-TOLYL)-6-
                                                                     METHYL-p-BENZOQUINONE
                                                                           MONOIMINE
                                        N-(4-CHLORO-o-TOLYL)-2-
                                        METHYL-p-BENZOQUINONE
                                              MONOIMINE

-------
SUMMARY
  The herbicides described here were formulated with degree of persistence after
application as a prime consideration. All of these compounds undergo primary
degradation readily in soils and in a variety of microbial cultures. The resulting
products are chlorinated anilines and other metabolites arising from the aliphatic
side  chain. When the side chain is not chlorinated, it  is metabolized  to  cell
constituents and CO2. Little research  has been published regarding  the  fate of
chlorinated or other highly substituted side chain metabolites.
  The bulk of the herbicides remain as chlorinated anilines. There is some evidence
for volatilization of these compounds in arid zones if sorption to soils is delayed or if
the concentration of chloroanilines is very high. However, the chlorinated anilines
are readily and strongly bound to soil humic substances. Monosubstituted anilines,
particularly 3-chloroaniline and 4-chloroaniline, are utilizable as sole carbon sources
by some microorganisms and are metabolized in laboratory experiments. Dichloro-
aniline is toxic  to microorganisms  at low  concentrations and evidence  for  its
degradation is indirect. It is not clear how sorption to soils affects degradation of the
chloroanilines in the environment.
  Chlorinated  anilines are metabolized by  dioxygenases to  chlorocatechols.
Organisms which convert the chlorinated anilines to chlorocatechols and then also
metabolize the chlorocatechols  do  so by the meta pathway.  Limited evidence
suggests that the amine group is removed directly without oxidation.
  Under some conditions azobenzenes are formed from condensation of chloro-
anilines via peroxidases. The concentration of chloroanilines must be very high for
this mechanism to be operative in soils.
  Primary degradation of the urea herbicides occurs by successive demethylation of
the side chain. The side chain is then metabolized leaving the chloroaniline moiety.
Methoxy phenyl ureas are metabolized by a different mechanism, as microorganisms
active against these compounds have little activity against the dimethyl  herbicides.
  Reductive dechlorination to a limited  extent of the aromatic moiety has been
reported. However, the effectiveness of this mechanism in the environment has not
been investigated. Anaerobic utilization of 4-chloroaniline  takes place readily, but
the resultant volatile product or products have not been identified.
  The primary mechanism of degradation of the phenyl carbamate herbicides is
hydrolysis of the ester linkage to form chlorinated anilines. These compounds are
metabolized by a wide variety of fungi and bacteria as noted above.
  Hydrolysis of the acyl anilide herbicides results in formation of the  chlorinated
anilines with utilization of the aliphatic side chain for cell growth. An unusual
mechanism employed by R.japonicus results in hydrolysis of the side chains of dicryl
and  N-(3,4-dichlorophenyl)-2-methyl pentanamide. This modification results in
detoxification of the molecules.
  Studies with various species of algae have shown that these organisms metabolize
acyl anilide compounds with production of chlorinated anilines.
  Metabolism of these classes of herbicides is dependent upon factors which affect
microbial soil activity, including such factors as pH and soil composition which
affect the chemical state of the compound as well. These compounds are relatively
easily broken down  to  chlorinated  anilines and  side chain  metabolites. The
chlorinated anilines may be volatilized or bound to soils before or concomitantly
with microbial metabolism. It  is not  clear what effects these competing processes
have on  biodegradation of these molecules. There have been few  studies on the
persistence or metabolism of compounds arising from the  side chain, particularly
those containing chloride ions.
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                             SECTION 13

                   CHLORINATED BIPHENYLS

  Polychlorinated biphenyls (PCBs) have been used widely in industrial applications
because of their  thermal stability,  excellent dielectric (electrically insulating)
properties, and resistance to oxidation, acids, bases, and other chemical agents.
PCBs therefore have found use in capacitors and transformers as dielectric fluids, in
hydraulic  systems, gas turbines and vacuum pumps, and as fire retardants and
plasticizers (215). In 1971, however, Monsanto Company, the sole U.S. producer,
voluntarily restricted the use of PCBs to closed systems (capacitors, transformers,
vacuum pumps, gas-transmission turbines) and discontinued production entirely in
1978 (281). These applications use complex mixtures of PCBs marketed under the
trade names Aroclor (Monsanto Company, USA), Clophen (Germany), Phenoclor
and  Pyralene (France), Kaneclor and Santotherm (Japan), and Fenclor (Italy).
Askarels are synthetic mixtures of chlorinated biphenyls and trichlorobenzenes.
  The Aroclor products are denoted  by a four-digit number in which the first two
indicate the type of molecule ("12" indicates a chlorinated biphenyl) and the last two
digits indicate the weight percent chlorine. Aroclor 1254 consists  of chlorinated
biphenyls  with 54% by weight chlorine and on average 5 chlorines per molecule,
although it has been reported to contain 69 different chlorinated biphenyl molecules
(162). Similarly, Aroclor 1242 is 42% chlorine by weight and averages 3 chlorines per
molecule.  Aroclor 1016 also consists primarily of trichlorinated compounds but
contains fewer penta- and hexachlorinated molecules than Aroclor 1242. There are
210  possible  PCB compounds containing 0 to 10 chlorine atoms per biphenyl
molecule.  However, many of these  have  never been found in commercial PCB
mixtures.
  The PCBs  have been released into the  environment for  many years and are a
worldwide contaminant. They are lipophilic and sorb strongly to the lipids and fats of
animals including fish, mussels, and birds. PCBs also undergo biological magnifica-
tion  in such common aquatic invertebrates as daphnids, mosquito larvae, stoneflies
and crayfish. The concentration of PCBs in the invertebrates can be as high as 27,500
times that in  water (162). These invertebrates subsequently are eaten by fish and
birds, and bioaccumulation occurs at all levels of the food chain.

MICROBIAL METABOLISM OF PCBs

  Most of the studies. on  microbial metabolism of PCBs  have explored the
biodegradability of the Aroclors in natural environments or in laboratories using
pure strains or mixed cultures. These studies have shown that PCBs containing fewer
than 5 chlorines per molecule are extensively degraded, while heavier molecules tend
to persist in the environment (26,91,163,167,220a, 281,394,438,474). These studies
are corroborated by environmental  analyses which indicate that  PCBs found in
weathered samples contain 5 or more chlorine atoms per molecule (162).
  Few studies have attempted  to elucidate the pathways of degradation of pure
compounds of a chlorinated biphenyl. Several studies have shown that 4-chlorobi-
phenyl can be metabolized to 4-chlorobenzoic acid, indicating hydroxylation, ring

                                    129

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cleavage, and degradation of the nonchlorinated ring of the molecule. This has been
demonstrated with soil bacteria, a sewage effluent isolate identified as Achrom-
obacter sp. pCB, an unidentified facultative anaerobe called strain B206, Acineto-
bactersp. P6, and Alcaligenes sp. Y42 (1, 163, 164, 333, 426). Acinetobacter sp. P6
can use 4-chlorobiphenyl as the sole source of carbon for growth (164).  Growth of
both Achromobacter sp. strain B 218 and Bacillus brevis  strain B  257 on 4-
chlorobiphenyl as the sole source of carbon generates the same metabolites (298).
The pathway of degradation involves formation of a 2,3-dihydroxy intermediate
with meta cleavage to form eventually 4-chlorobenzoic acid. Other metabolites were
isolated which represent successive oxidation and utilization of the aliphatic carbons
from the cleaved ring (Figure 56).
  The formation of chlorinated benzoic acids from chlorinated biphenyls is the most
common route of PCB degradation. Both Alcaligenes sp. Y42 and Acinetobacter sp.
P6 convert a large number of biphenyl compounds to the corresponding chlorinated
benzoic acids (Table 5) (163,  168). For several compounds  containing multiple
chlorines with one on the second ring, loss of that chlorine occurs in the formation of
the chlorobenzoic  acid. Studies with 14C-2,5,2'-trichlorobiphenyl  confirmed the
formation of l4C-2,5-dichlorobenzoic acid and a yellow intermediate by resting cell
suspensions of both Alcaligenes sp. Y42 grown on biphenyl and Acinetobacter sp. P6
grown   on 4-chlorobiphenyl (164). The  patterns  of metabolism  of PCBs by
Alcaligenes sp. Y42  and Acinetobacter sp. P6 are similar.  The general path of
degradation proceeds through meta cleavage compounds to chlorobenzoic acids
which accumulate during the metabolism of chlorobiphenyls. Metabolism of some
chlorinated biphenyls is blocked after production of the dihydroxy intermediate
(precursor to ring cleavage), while  for other compounds the mefa-cleavage inter-
mediate accumulates.
  Pseudomonas sp. strain 7509 grown on biphenyl metabolizes 2,4'-dichlorobi-
phenyl to two different monochlorobenzoates, indicating that both rings are capable
of being attacked (26a). An intermediate metabolite was identified as 2-hydroxy-6-
oxo-6-(chlorophenyl)chlorohexa-2,4-dienoic acid.
  The  degradation of 2,4,4'-trichlorobiphenyl by Acinetobacter sp. P6 grown on
4-chlorobiphenyl was studied in detail and a pathway was proposed involving meta
cleavage after 2',3'-hydroxylation (Figure 57) (166). Hydroxylation  occurs on the
ring with the  fewest chlorine substituents. The predominant metabolite is the meta
cleavage product, although small amounts of the dichlorobenzoic acid appear. The
occurrence of a yellow meta  cleavage product  and subsequent  production of
chlorobenzoates in the degradation of other PCBs suggests that this may be a general
pathway for most PCB metabolism  in bacteria.
  On the basis of these studies, several generalizations were made regarding the effect
of the  structure of the PCBs on  microbial degradation (163, 164,  165, 168).
Degradation  decreases as the number of chlorines per molecule increases.  Two
chlorines on the ortho positions of a single ring (i.e., 2,6-) or on both rings (i.e., 2,2'-)
inhibit degradation. PCBs with one unsubstituted ring are more readily metabolized
than PCBs with the same  number of  chlorines  on both rings. On PCBs  with
unequally substituted rings, the ring with fewer substitutions is  preferentially
cleaved. PCBs with a chlorine  on the 4'-position are metabolized to stable meta-
cleavage products.
  The occurrence of nitro-containing metabolites in extracts of media containing the
unidentified facultative anaerobe B 206 and 4-chlorobiphenyl was investigated (427).
When ammonium sulfate is added  to the culture medium as the nitrogen source,
4-chlorobiphenyl is metabolized to 2-hydroxy-4'-chlorobiphenyl and 4-hydroxy-4'-
chlorobiphenyl. However, when the nitrogen source is sodium nitrate, the metabo-
lites 2-hydroxy-nitro-4'-chlorobiphenyl  and 4-hydroxynitro-4'-chlorobiphenyl ap-


                                   130

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                                                                                                           COOH
o     ^-
CO
to

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        TABLE 5. METABOLISM OF CHLORINATED BIPHENYL
            COMPOUNDS BY ALCALIGENES SP Y42 AND
                      ACINETOBACTER SP. P6*
              Substrate
                                                 Products
 Substitutions on one ring:

 2-Chlorobiphenyl
 3-Chlorobiphenyl
 4-Chlorobiphenyl
 2,3-Dichlorobiphenyl
 2,4-Dichlorobiphenyl
 2,5-Dichlorobiphenyl
 2,6-Dichlorobiphenyl+

 3,4-Dichlorobiphenyl
 3,5-Dichlorobiphenyl
 2,3,4-Trichlorobiphenyl
 2,3,6-Trichlorobiphenyl+

 2,4,5-Trichlorobiphenyl
 2,4,6-Trichlorobiphenyl
2,3,4,5-Tetrachlorobiphenyl
2,3,5,6-Tetrachlorobiphenyl
2,3,4,5,6-Pentachlorobiphenyl

Substitutions on both rings:
2,2'-Dichlorobiphenyl
2,4'-Dichlorobiphenyl
3,3'-Dichlorobiphenyl
4,4'-Dichlorobiphenyl
2,4,4'-Trichlorobiphenyl

2,5,2'-Trichlorobiphenyl

2,5,3'-Trichlorobiphenyl
2-Chlorobenzoic acid
3-Chlorobenzoic acid
4-Chlorobenzoic acid
2,3-Dichlorobenzoic acid
2,4-Dichlorobenzoic acid
2,5-Dichlorobenzoic acid
2,5-Dichlorodihydroxybiphenyl,
 2,6-dichlorotrihydroxybiphenyl
3,4-Dichlorobenzoic acid
3,5-Dichlorobenzoic acid
2,3,4-Trichlorobenzoic acid
2,3,6-Trichlorodihydroxybiphenyl,
 2,3,6-dichlorotrihydroxybiphenyl
2,4,5-Trichlorobenzoic acid
2,4,6-Trichlorobenzoic acidf,
 2,4,6- trichlorodihydroxybiphenyl.f
 2,4,6-trichlorotrihydroxybiphenyl+
2,3,4,5-Tetrachlorobenzoic acid
None
None
2-Chlorobenzoic acid
2-Chlorobenzoic acid
3-Chlorobenzoic acid
4-Chlorobenzoic acid
2-Chlorobenzoic acid,
 2,4-dichlorobenzoic acid
2-Chlorobenzoic acid,
 2,5-dichlorobenzoic acid
3-Chlorobenzoic acid,
 2,5-dichlorobenzoic acid
                                                             (continued)
                                  132

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TABLE 5. (continued)
 2,5,4'-Trichlorobiphenyl

 3,4,2'-Trichlorobiphenyl
 2,3,2',3'-Tetrachlorobiphenyl

 2,3,2',5'-Tetrachlorobiphenyl

 2,4,2',4'-Tetrachlorobiphenyl+
 2,4,2',5'-Tetrachlorobiphenyl+
 2,4,3',4'-Tetrachlorobiphenyl+
 2,5,2',5'-Tetrachlorobiphenyl+

 2,5,3',4'-Tetrachlorobiphenyl+
 2,6,2',6'-Tetrachlorobiphenyl+
 3,4,3',4'-Tetrachlorobiphenyl+
 2,4,5,2',3'-Pentachlorobiphenyl+

 2,4,5,2',5'-Pentachlorobiphenyl+
2-Chlorobenzoic acid,
 2,5-dichlorobenzoic acid
2-Chlorobenzoic acid
2,3-Dichlorobenzoic acid, an
  unidentified dichloro compund
Dichlorobenzoic acid, an unidentified
  dichloro compound
2,4-Dichlorobenzoic acid
Dichlorobenzoic acid
Dichlorobenzoic acid
2,5,2',5'-Tetrachlorodihydroxyl-
 biphenyl
Dichlorobenzoic acid
None
3,4-Dichlorobenzoic acid
2,4,5-Trichlorobenzoic acid, an
 unidentified trichloro compound
2,4,5,2',3'-Tetrachlorodihydroxy-
 benzoic acid
   *Adaptedfrom reference 163, 164, 168.
   +Metabolism by Acinetobacter sp. P6 only.
   fMetabolism by Alcaligenes sp. Y42 only.

 pear. These metabolites were interpreted to result from a nonenzymatic reaction
 between an arene oxide intermediate and nitrate or nitrite anions. This organism
 subsequently accumulates 4-chlorobenzoic acid in the medium (426). The appear-
 ance of the monohydroxy intermediates suggests a rare monooxygenase mechanism
 for PCB degradation, although the phenylphenols may also be an artifact arising
 during the isolation procedure (7la).
   Acinetobacter sp. strain P6 resting cells were incubated for 4 hours with several
 Kaneclor PCB mixtures (167). Kaneclor KC200 (primarily dichlorobiphenyls) was
 metabolized  to  monochlorobenzoates.  Kaneclor  KC300  was  metabolized  to
 benzoates with 1 to 3 chlorines, dihydroxybiphenyls  with 2 to 4 chlorines, ring meta
 cleavage products with 2 to 3 chlorines, and many  other unidentified compounds
 with  2 chlorines. Kaneclor KC500 was scarcely  metabolized,  although some
 dihydroxy isomers were noted.
   Studies were conducted utilizing Alcaligenes sp. strain BM 2 which was isolated on
 diphenylmethane and known to metabolize dichlorinated biphenyls (474). A mixture
 of di- and trichlorinated biphenyls at 0.05% concentration (32% by weight) was 80%
 metabolized in 1 day and completely metabolized in 3 days. At 0.25% concentration,
 22% was metabolized in 1 day and 29% in 3 days. Under cometabolic conditions, a
 100 mg/1 PCB mixture of di-, tri-, and tetrachlorobiphenyls (41% by weight) was 70%
 metabolized in 2 days and 80% in 6 days. In a minimal medium with only a small
 quantity of carbon source, 30% was metabolized in  6 days. Most of the remaining
 substrate was tetrachlorobiphenyl.  The metabolites  included mono-  di-, and
 trichlorobenzoates, monohydroxychlorobiphenyl, 2-hydroxy-6-oxo-chlorophenyl-
 hexa-2, 4-dienoic acid, chlorobenzoylpropionic acid, chlorophenylacetic acid, and
 3-chlorophenyl-2-chloropropenoic acid (a substituted cinnamic acid).
                                     133

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s

3
01
                              2,4,4'-TRICHLOROBIPHENYL
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CD  "!
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=?  o
o  3
3  2

£  1

S  =-
g  1
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   cn
   •D
           Cl
      Cl     OH  OH


Cl~(^	Cr~cl


      Cl     OH  OH

ci-(o)	(O)-ci
           ci     OOHOH
                          Cl
ci-(o)-COOH
                              1-CHLORO-2,3-DIHYDROXY-4-(2,4-DICHLOROPHENYL)-HEXA-4,6-DIENE
                              2,4,'-TRICHLORO-2',3'-DIHYDROXYBIPHENYL
                              3-CHLORO-2-HYDROXY-6-OXO-6-(2,4-DICHLOROPHENYL)HEXA-2,

                              4-DIENOIC ACID
                              2,4-DICHLOROBENZOIC ACID

-------
  Pseudomonas sp. strain 7509 also formed mono- and dichlorobenzoic acids during
metabolism of Aroclor 1242 (26a). A  number  of nonchlorinated aromatic and
aliphatic compounds were isolated after 2 months'incubation of Aroclor 1242 with
several strains of bacteria isolated from lake water (233). No chlorinated metabolites
or oxidized derivatives of PCBs were detected.
  The disappearance of Aroclor  products  and individual PCB  isomers during
incubation with Nocardia sp. NCIB 10603  was monitored (26). The following
isomers were 60 to 100% metabolized within 2 weeks: 2,4'-di, 2,3-di, 3,4-di, 2,3,2'-tri,
2,3,4'-tri-  and  3,4,3'-trichlorobiphenyl, while  2,5,4'-trichlorobiphenyl was 60%
metabolized in 73 days. There was little  transformation of 2,4,6-tri-, 2,4,2',4'-tetra-,
or  2,4,6,2'-tetrachlorobiphenyl in  9 days, and  no degradation  of 4,4'-
dichlorobiphenyl was detected after 121 days. However, 4,4'-dichlorobiphenyl was
50% metabolized in 2 days when present as a component of Aroclor  1242. During 52
days' incubation Aroclor 1242 was 88% metabolized and  in  100 days was 95%
metabolized. Aroclor 1016 was 96% metabolized in 52 days.
  There are two reports in the literature concerning the degradation of PCBs by
fungi.  Rhizopus japonicus converts  4-chlorobiphenyl to 4-chloro-4'-hydroxybi-
phenyl and 4,4'-dichlorobiphenyl to an unidentified hydroxylated metabolite (454).
Cunninghamella echinulata Thaxter metabolizes 2,5-dichloro-4'-isopropylbiphenyl
by oxidation of the isopropyl group to form 2,5-dichloro-4'-biphenylcarboxylic acid
and by hydroxylation of the chlorine-substituted phenyl group (440).

METABOLISM OF PCBs BY MIXED MICROBIAL CULTURES
  A mixed microbial population  in lake water metabolized 2-chlorobiphenyl to
2-chlorobenzoic acid and chlorobenzoylformic acid (394). In contrast, 2,4'-dichloro-
biphenyl was not metabolized after 8 months' incubation.
  A mixed microbial culture derived  from river sediments was able to metabolize
4-chlorobiphenyl rapidly, with 99%  removal in 30  days (265). Acclimated and
nonacclimated cultures showed similar results. There was transitory formation of a
metabolite thought to be 2-hydroxy-6-oxo-6-(4-chlorophenyl)hexa-2,4-dienoic acid
as well as production of 4-chlorobenzoic acid. The substrate was MC-labeled on the
chlorinated ring only and production of 14CO2 was noted after 4-chlorobenzoic acid
formation. The substrates 2-chloro- and 3-chlorobiphenyl were also metabolized,
but 2-chlorobenzoic acid was not degraded further.
  A marine mixed microbial community metabolized all three monochlorinated
biphenyls (52). Metabolites were not identified.
  A river water die-away test demonstrated 50% removal of 1 to 100 mg/12-chloro-,
3-chloro-, or 4-chlorobiphenyl within 2 to 5 days (17). The compound was uniformly
14C-labeled in  the chlorinated  ring  and up to 50% of the  label appeared as
monochlorobenzoic acid and subsequently as 14CO2. No degradation  of 2,2',4,4'-
tetrachlorobiphenyl was noted after 98 days' incubation.
  A mixture of bacteria was  isolated by enrichment culture with garden soil using
benzene as the substrate (18). Several PCB isomers were incubated with the mixed
culture and benzene for up to 6 weeks and  the medium subsequently tested  for
presence of chlorobenzoate metabolites. Neither 2-chloro- nor 2,2'-dichlorobiphenyl
was metabolized  to  chlorobenzoic acids, although  4-chlorobiphenyl formed 4-
chlorobenzoic acid. The substrate 2,4,4'-trichlorobiphenyl formed copious amounts
of 4-chloro- and 2,4-dichlorobenzoic acid in the ratio 5:2. Loss of one chloride was
noted in the formation of 4-chlorobenzoic  acid from 2,4'-dichlorobiphenyl, 2,5-
dichlorobenzoic acid from 2,2',5-trichlorobiphenyl, 4-chlorobenzoic acid and 3,4-
dichlorobenzoic acid from 3,4,4'-trichlorobiphenyl, 4-chlorobenzoic acid and 2,4-
dichlorobenzoic acid from 2,4,4'-trichlorobiphenyl, and 2,3,4,5-tetrachlorobenzoic
acid from both 2,3,3',4,5-pentachlorobiphenyl and 2,3,4,,4',5-pentachlorobiphenyl.

                                    135

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  A soil plot was treated with 1 ppm 2,2'-dichlorobiphenyl (317). After one growing
season almost half the remaining material was unchanged substrate. About 9% were
soluble metabolites and almost 42% were unextractable residues. After 1 year, 74% of
the remaining  material were  unextractable residues. The metabolites  included
monohydroxy derivatives of the substrate as well as other products.
  Metabolism of 4,4'-dichlorobiphenyl by a mixed microbial culture obtained from
the filtrate of an activated sludge sample resulted in formation of 4,4'-dichloro-2,3-
hydroxybiphenyl  and 4-chlorobenzoic acid (439). Metabolism of this substrate is
repressed by the presence of alternative carbon sources. Under similar experimental
conditions, including an incubation time of 14 days,  there was no metabolism of
2,4,5'-tri-, 2,2',5,5'-tetra, 2,2',3,4,5'-penta-, 2,2',3,4,5,5'-hexa- and decachlorobi-
phenyl.
  Metabolism of 2,5,2',5'-tetrachlorobiphenyl and 2,5,2'-trichlorobiphenyl occurred
in seawater with production of a compound thought to be a lactone acid (68a). No
degradation was  noted during the incubation of the  tetrachlorobiphenyl with
anaerobic marsh mud during a 45-day incubation period.
  A microbial consortium obtained from activated sludge metabolized the isopropyl
group of 4-chloro-4'-isopropylbiphenyl to a hydroxyl substituent, forming 4-chloro-
4'-hydroxybiphenyl,  followed by formation of 4-chlorobenzoic acid (440). Inter-
mediates of the isopropyl metabolism pathway were identified. Addition of glucose
as an alternate carbon source repressed metabolism  of the chlorinated substrate.
  Biphenyl enrichment of both uncontaminated soils  and soils contaminated with
PCBs resulted in isolation of mixed cultures which were incubated with Aroclor 1242
(91). Cometabolism  with sodium acetate enhanced metabolism of all the PCBs
including the higher chlorinated molecules, although this phenomenon may be due to
the increased biomass resulting from growth on the simpler carbon source. Extensive
degradation of all  the  lower  chlorinated isomers was noted with  up to 68%
metabolism of the tetrachlorinated biphenyls in 15 days.
  A related study was conducted to determine the influence of inoculum concen-
tration on the aerobic bio-oxidation of 3,3'-dichlorobenzidine which is used in the
manufacture of azo dyes (57). The effluent from a domestic sewage treatment plant
was used as the inoculum and the substrate concentration was 20 mg/1. When present
as the sole carbon source, the substrate was not metabolized. However, the presence
of yeast extract in the medium promoted extensive disappearance of the substrate
within 28 days.  In another experiment, 2 mg/1 3,3'-dichlorobenzidine was added to
lake or reservoir water and incubated 14 days (10). Neither metabolites nor I4CO2 was
recovered after incubation. Increasing disappearance of substrate with time was
correlated with increasing biomass, which served as a sorbent for the substrate. The
supernatant fluid from settled activated sludge material served as the inoculum for
flasks containing  3,3'-dichlorobenzidine and additional carbon sources. After 4
repeated weekly subcultures to flasks of fresh media, no metabolites of the substrate
were recovered.

SUMMARY

  The limited number of studies on the degradation of specific chlorinated biphenyl
compounds by pure strains of bacteria has served to establish some general features
of PCB metabolism. A few strains of bacteria have been shown to mineralize some
chlorinated biphenyls. In most cases bacteria with the capability of degrading one
ring of a chlorinated biphenyl compound are unable  to  degrade the resulting
chlorinated benzoates. These compounds accumulate in pure cultures.  Evidence
exists for the  complete mineralization of chlorinated benzoates by other strains of
bacteria (discussed in Section 7  on chlorobenzoates), and mixed cultures of bacteria


                                    136

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have been shown to mineralize PCBs with 4 or fewer chlorines per molecule. More
heavily substituted PCBs appear to resist degradation  and accumulate even in
environments where lower PCBs are degraded.
  The mechanism of hydroxylation of PCBs by bacteria has not yet been elucidated,
nor have the enzymes mediating the steps in the proposed pathways been isolated.
Two pathways have  been proposed, the first analogous  to  the  pathway  of
degradation of biphenyl. Initial hydroxylation occurs in the 2,3-position of the less
substituted ring, followed  by meta cleavage  and subsequent degradation of the
aliphatic portion of the molecule to form substituted benzoic acids. Chlorines on the
aliphatic carbons are lost during this process.  However,  this  may not be the
mechanism for degradation of PCBs substituted in all the ortho positions. A second
pathway based on presence of a monooxygenase in bacteria has been proposed after
discovery of 4-hydroxy-4'-chlorobiphenyl in extracts of bacterial cultures incubated
with 4-chlorobiphenyl. More evidence corroborating  this mechanism needs to  be
obtained to determine how widespread this pathway is. Limited evidence on fungal
metabolism of PCBs indicates activity of a monooxygenase in a manner similar to
that shown in biphenyl metabolism.
                                   137

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                              SECTION 14
               DDT AND RELATED COMPOUNDS
  DDT,  l,l,l-trichloro-2,2-bis(p-chlorophenyl)ethene, received  its abbreviation
from the trivial name dichlorodiphenyltrichloroethane. This chlorinated aromatic is
one of the most persistent pesticides in the environment. Since it is lipophilic, it
readily accumulates in microorganisms and invertebrates and undergoes biomagni-
fication as fish and  birds higher in  the  food chain ingest DDT-contaminated
organisms (79,229,263,311). Widespread effects of DDT poisoning include eggshell
thinning and birth defects. Reviews of the literature pertaining to DDT metabolism
in microbial systems have been published in 1976 and 1980 (138, 228).

BACTERIAL METABOLISM OF DDT

  Due to the widespread incidence of toxicity demonstrated after DDT ingestion,
the intestinal flora of various animals became the focus for studies of the metabolism
of DDT. DDT is converted directly to ODD (l,l-dichloro-2,2-bisOchlorophenyl)-
ethane, also referred to as dichlorodiphenyldichloroethane (Figure 58). The reaction
involves removal of a chloride ion from the aliphatic portion of the molecule. This is a
reductive dechlorination reaction requiring anaerobic conditions, in contrast to the
oxidative pathways of metabolism of most other pesticides.
  The involvement of intestinal bacteria in DDT metabolism by mammals has been
demonstrated in experiments which showed that rats converted DDT to DDD when
fed by stomach tube, but not  when injected intraperitoneally (308). The coliform
bacteria  Escherichia coli  and Enterobacter aerogenes isolates from rat  feces
demonstrate this reaction as well. Microorganisms isolated from rat intestines which
convert DDT to  DDD include  Clostridium perfringens,  Streptococcus sp.,
Bacteroides  sp., E.  coli and other  coliforms,  yeasts, and to a  lesser extent
Lactobacillus sp. (55). Klebsiella pneumoniae also converts DDT to DDD (464).
DDE (l,l-bis(p-chlorophenyl)-2-chloroethylene) can be detected after 20 hours at
concentrations from  5 to 10% in cultures of Streptococcus sp.,  Bacteroides sp.,
Pseudomonas sp. and Lactobacillus sp. (the latter after a 72 hour incubation) (55).
However, DDE  has  been shown to  be produced  nonenzymatically as well as
enzymatically (464).
  Proteus vulgaris, isolated from the intestines of DDT-resistant mice, converts
DDT  to DDD  subsequently to  l,l-bis(p-chlorophenyl)-2-chloroethane  and
l,l-bis(p-chlorophenyl)ethane, representing three successive reductive dechlori-
nations (Figure 58) (20,21). DDE also appears in the medium. The excreta of stable
flies became the source of three bacteria, E. coli, Serratia marcescens, and a third
unidentified strain, which convert DDT to DDD (90%) and DDE (5%) after 24 to 72
hours anaerobically but not aerobically (415). Bacteria from bovine rumen fluid
convert 14C-DDT to 14C-DDD  (311). This same reaction was noted for DDT
incubated with water from Clear Lake, California and with reduced iron porphyrins
(hemoglobin or hematin) (311). The isomer o,p -DDT which constitutes 10-20% of
                                    138

-------
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-------
technical grade DDT is converted by rumen microorganisms to o,p -DDD at the
same rates as p,p'-DDT (161). This conversion occurs as well in E. aerogenes both
aerobically and anaerobically (309).
  The direct conversion of DDT to DDD without the DDE intermediate was
confirmed in E. aerogenes using deuterated DDT (355). The deuterium atom present
at the 2-position in DDT is retained in the product, indicating that the chlorine is
replaced by hydrogen (or hydride ion) directly without the intermediary species. The
membranes  of E. coli were shown to be the site of reductive dechlorination of this
species. The process required flavine-adenine dinucleotide (FAD) and anaerobic
conditions (160).
  Cell-free extracts of E. aerogenes also convert DDT to DDD. This activity is due to
reduced Fe(II) cytochrome oxidase (464).  More complete degradation of DDT
occurs in both whole-cell preparations and cell-free extracts of this organism (Figure
59)(463,465). Metabolism follows the pathway DDT - DDD - DDMU - DDMS -
DDNU - DDA -  DPM - DBH - DBF -, where the abbreviations represent the
compounds  as follows: (DDMU)  l-chloro-2,2-bis(p-chlorophenyl)ethylene;
(DDMS)  l-chloro-2,2-bis(p-chlorophenyl)ethane; (DDNU)  unsym-bis(p-chloro-
phenyl)ethylene; (DDA) 2,2-bis(p-chlorophenyl)acetate or more commonly dichloro-
diphenylacetate; (DPM) dichlorodiphenylmethane; (DBH)  dichlorobenzhydrol;
and (DBF) dichlorobenzophenone. The enzymatic conversion of DDT to DDE is a
dead-end side reaction.  DDA is the end product of vertebrate metabolism. The
conversion of DDA to DBF does not require anaerobic conditions. This pathway has
also been demonstrated in anaerobic cultures of E. coli (268).
  A single study reports that under aerobic conditions,  cultures of Bacillus cereus
metabolize DDT by this pathway within 7 days, although the use of screw-cap flasks
in these experiments may have allowed some anaerobiosis to develop. Cultures of E.
coli incubated aerobically for 24 hours with intermittent shaking converted DDT to
DDD (75%) and DDE (25%) (247).
  Environmental isolates also have the ability to convert DDT to DDD. Viable cells
of Bacillus megaterium convert DDT to DDD (201). Three hundred bacterial strains
from Lake Michigan each converted DDT to DDD and many converted I4C-DDD
to 14C-DDNS (l-bis(p-chlorophenyl)ethane (302). Bacteria isolated from marine
and brackish water and sediment converted 14C-DDT to water-soluble metabolites
(232). Forty-seven of 100 isolates effected 5 to 10% conversion while an additional 38
isolates converted less than 5% of the starting material. Twenty-five isolates did not
produce water-soluble metabolites, indirectly indicating that the presence of those
metabolites in the other cultures was biologically mediated. Twenty-three of 26 plant
pathogenic and saprophytic strains of bacteria representing nine genera, converted
DDT to DDD anaerobically (230). Eighteen bacterial strains, mostly Pseudomonas
spp. which previously had been shown to metabolize dieldrin, also degraded DDT to
DDD (349). In addition, 14 of these isolates produced  DDA and 10 produced a
dicofol-like compound.
  Pseudomonas aeruginosa 640x isolated from DDT-polluted soil of the Crimean
region was used to construct 2  derivatives (185). Strain BS816 carries a plasmid
encoding the genes which degrade naphthalene and salicylate by ortho cleavage, and
strain BS827 carries a plasmid which effects meta degradation. Both plasmids were
obtained from strains of P. putida. The parent P. aeruginosa and the two derivatives
all metabolize DDT with the formation of the same metabolites. Strain  BS816,
carrying the plasmid coding for ortho cleavage, degrades DDT most extensively,
converting 89% of the DDT to DDD,l,l-dichloro-2,2-bis(p-chlorophenyl)ethylene
(DDDE), phenylpropionic acid (PPA) and phenylacetic  acid (PAA).
  An organism identified as a Pseudomonas sp. was isolated by enrichment culture
for its ability to use diphenylethane as a sole source of carbon and energy for growth

                                    140

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(157). The ability of this organism to grow on several other metabolites of DDT was
tested. Diphenylethane is converted  to  2-phenylpropionic acid and metabolized
further. Diphenylmethane is metabolized with intermediate production of phenyl-
acetic acid and l-(p-chlorophenyl)-l-phenylethane is metabolized with the produc-
tion of 2-(p-chlorophenyl)proprionic acid as the only metabolite, indicating cleavage
of the unsubstituted ring. The DDE  analog l-(p-chlorophenyl)-l-phenylethene is
metabolized only to 2-(p-chlorophenyl)-2-propenoic acid. The substrate 1-p-chloro-
phenyl)-l-phenylethanol is converted to 2-(p-chlorophenyl)-2-hydroxypro-
pionic acid which is slowly metabolized further. During metabolism of this substrate
minor amounts of the nonchlorinated metabolite 2-hydroxy-2-phenylpropionic acid
are produced, indicating attack on the chlorinated ring. Each of the above substrates
served as the sole  source of carbon and energy for growth. The compound 2,2-
diphenylethanol is  not metabolized. Analogs of DDT which have chlorine substi-
tuents on both rings are not metabolized, and l,l-diphenyl-2,2,2-trichloroethane is
not metabolized. However, when diphenylethane is available in the medium, this
organism cometabolizes bis(p-chlorophenyl)methane to p-chlorophenylacetic acid
which accumulates (158). With the same cosubstrate, the organism cometabolizes
l,l-bis(p-chlorophenyl)ethane  to 2-(p-chlorophenyl)propionic acid with transient
appearance  of two hydroxylated  metabolites,  l-(p-chloro-o-hydroxyphenyl)-l(p-
chlorophenyl)ethane and  l-(p-chloro-m-hydroxyphenyl)-l-(p-chlorophenyl)ethane.
Accumulation of toxic chlorinated carboxylic acids may have inhibited metabolism
of the substrate. Compounds which are recalcitrant to cometabolic activity have
substitutions in the ethane or ethene  sections of their structures which may cause
steric hindrance.
  A large sampling study of oceanic and near-shore environments established that 35
of 95 isolates degraded  DDT to many of the metabolic  products  previously
identified, with DDD  the major metabolite (350). The only environmental samples
which failed to mediate DDT degradation were the oceanic water samples.

FUNGAL METABOLISM OF DDT

  The earliest studies involving microbial metabolism of DDT were conducted using
commercial yeast cakes (Saccharomyces cerevisiae) (234).  Reductive dechlorination
was demonstrated  by the  appearance in culture  media of I4C-DDD from I4C-DDT
labeled in the phenyl  group. DDE was shown  not to be  a necessary intermediate
metabolite.
  A study of 8 fungi incubated with DDT for 6 days did not reveal degradation (80).
However, in the same study 6  of 9 actinomycetes were found to convert DDT to
DDD. These 6 actinomycetes are Nocardia erythropolis, Streptomyces aureofaciens,
S. viridochromogenes, S. cinnamoneus, and with lesser efficiency S.  albus and S.
antibioticus. All cultures in this study were incubated aerobically with shaking.
Another study of microbial cultures with the capability to degrade dieldrin showed
that 2 Trichoderma viride strains could degrade DDT to DDD, a "dicofol-like"
metabolite, and DDA (349). The dicofol-like compound was subsequently identified
as l-bis(p-chlorophenyl)ethane (302). Earlier studies with several strains of T. viride
established differences in the metabolites produced by each strain (300).
  Shake  cultures  of  Mucor alternans  in  nutrient  media containing I4C-DDT
produced 3 hexane  soluble and 2 water soluble metabolites  within 2 to 4 days (7). The
total activity recovered was about equally divided between the two phases. The major
metabolite is water soluble. These  metabolites  were unidentified since the results
obtained from thin  layer chromatography were different from those for DDE, DDD,
DDA, DBF, dicofol, or l,l-bis(p-chlorophenyl)ethane. M. alternans converted 15%
of the DDT starting material to 3 unidentified  water soluble products in another


                                    142

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experiment as well (232). A comparison of these products with l,l-bis(p-chloro-
phenyl)acetic acid (DDA), PCPA, DBF, DBH and 2-chlorosuccinic acid failed to
reveal their identities. Attempts to reproduce this metabolic activity in the natural
environment were unsuccessful, as the addition of M. alternans spores to DDT-
treated soil failed to promote any degradation after  11 weeks' incubation (7).
  A sequential experiment was developed to study the interactive effects of bacteria
and fungi (156). Hydrogenomonas sp. was grown on dichlorodiphenylmethane or
p-chlorophenylacetic acid, both metabolic products of DDT. Hydrogenomonas sp.
cannot liberate free chloride from metabolism of these compounds. The culture
supernatant fluid was extracted and added to a basal salts solution, which became the
growth medium for a culture of Fusarium sp. Growth occurred  under anaerobic
conditions, and chloride was detected in the medium, indicating that the products of
Hydrogenomonas sp.  metabolism were degraded to CO2,  H2O,  and HC1 by
Fusarium sp. The ability to perform this mineralization decreased  if the 2 microbial
populations were incubated together.
  In another study of the interactive effect of other fungi on DDT degradation by M.
alternans, the addition of other fungal cultures or the cell-free spent media from some
cultures repressed the formation of water soluble metabolites (6). Other fungi,
including Aspergillus flavus. A.  fumigatus. A.  niger,  Fusarium  oxysporum,
Penicillium notatum. Rhizopus arrhizus and Trichoderma ciride, failed to produce
water soluble metabolites of DDT. However, water soluble 14C-products appeared
after incubation of UC-DDT with the excretory products retained in culture media
after growth of all of the above fungi including M. alternans, with the exception of R.
arrhizus. This discrepancy in the appearance of water soluble metabolites may be
attributed to sorption of degradation products by the mycelia or to further
degradation by cells to metabolites that are not water soluble.
  The path of DDT metabolism by Fusarium oxysporum has been established and
follows the route DDT - ODD - DDMU - DDHO - DDOH - DDA - DBF, with
DDE formed from DDT(135,136,136a). DDHO is the aldehyde intermediate which
is rapidly converted to DDOH and DDA. This path is similar to the described for
bacteria. The enzymes involved in DDT metabolism which  have been isolated
include DDT dehydrochlorinase  and those that decompose DDMU, DDA, and
DDOH (165a). DDT inhibits the fungal esterase while ODD strongly activates the
same enzyme (159). The net effect is enzyme activation which results in detoxification
of the molecule.

FUNGAL METABOLISM OF OTHER COMPOUNDS

  The acaricide chlorobenzilate (ethyl 4,4'-dichlorobenzilate)  14C-labeled in  the
aliphatic moiety was cometabolized by Rhodotorula gracilis with glucose as an
additional carbon source (312,313). Production of 14CO: was correlated with culture
growth. Metabolites included 4,4'-dichlorobenzilic acid, dichlorobenzophenone,
and other unidentified products. The same results were obtained in the metabolism of
chloropropylate (isopropyl 4,4'-dichlorobenzilate).  Alteration of the chlorinated
rings was not noted.

PERSISTENCE AND DEGRADATION OF  DDT IN THE

ENVIRONMENT

  DDT is converted to DDD by anaerobic but not aerobic sludge microorganisms
(202). The same results were found using Pawnee silt loam treated with '•'C-DDT.
Under anaerobic conditions DDD was recovered while under aerobic conditions
DDE was the only metabolite (190). Another study of anaerobic soil treated with


                                    143

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DDT reported formation of DDD and traces of other metabolites including DDE,
DDA, dicofol, 4-chlorobenzoate, DBF, and DBM (189a). Conversion of DDT to
DDD in flooded soil was faster when more organic matter was present (69).
  The conversion of DDT to DDD has been demonstrated in sterile as well as
nonsterile environments (59), and has been related to the redox potential (Eh) of the
soil. Sewage sludge samples sterilized in a variety of ways all resulted in conversion of
DDT to DDD if the Eh was sufficiently low (489). The rate of DDT degradation is
highest in soils with the lowest redox potentials, in the range of -90 to -250 mV (184).
Studies of DDT degradation in a variety of anaerobic and aerobic environments
using various carbon sources and various soils, have shown differences in the amount
of degradation and the efficiency of substrate and product recovery which preclude
extrapolation from laboratory to the environment or even from one experiment to
another (346, 347). Little conversion was found in moist anaerobic soil with Eh of
+350 mV or in flooded anaerobic soil with an Eh that dropped from +400 to +200 mV.
Flooded stirred soil (Eh = 0 m V) also showed little degradation. However, stirred
anaerobic soil treated with lime (Eh = -250 m V) and glass beads inoculated with
muck (Eh = -250 to -300 m V) mediated greater than 95% conversion of DDT (346).
  It appears that at  low Eh, DDT undergoes an irreversible redox type of reaction
with transient formation of a free radical before  conversion to DDD (184). The
reaction is thought to be mediated by reduced iron porphyrins, with cell metabolism
not being necessary (489).
  Numerous studies have been conducted with regard to the persistence of DDT and
its metabolites in environments treated with the insecticide (115,121, 278, 324, 344,
429,444,485). All of the studies have reported residues of DDT, DDE and DDD in
the environment at the time of sampling. The longest period of time  between last
application of DDT and  sampling was 17 years, and on the basis of these data
half-life numbers for DDT of 2.5 to 35 years have been reported (324). In general, the
amount of  these residues that remains in the soil  1 to 2 years after application is
similar to the amount recovered 9 years or longer after the first sampling.

SUMMARY

  Studies on metabolism of DDT by bacteria and  fungi have shown that reductive
dechlorination of the nonaromatic portion of the molecule is the necessary primary
step. The pathway of metabolism of DDT, first  described for E.  aerogenes but
subsequently confirmed in other bacteria and in fungi, describes a series of steps
requiring anaerobiosis (DDT to DDA) followed by a series of steps that may require
aerobic conditions (DDA to DBF). Although some studies report aerobic conversion
of DDT to metabolites, the oxygen tension was  not rigorously defined in these
experiments. All of these metabolites retain the chloride ions on both aromatic rings
of the molecule.
  The  effect  of  a concerted  attack  by several  microbial  species  has  been
demonstrated in a two-stage experiment in which Hydrogenomonas sp. was grown
on either of the DDT metabolized dichlorodiphenylmethane or p-chlorophenyl-
acetic acid.  The resulting filtered media became the growth substrate for cultures of
the fungus  Fusarium sp., which anaerobically  liberated free chloride, indicating
mineralization to CO2, H2O, and HC1. The nature of this pathway and the enzymes
involved have not yet been elucidated.
  Other studies which have shown differing efficiencies of DDT metabolism between
pure culture and consortia or sewage/soil studies,  indicate that degradation of this
compound  is highly dependent on environmental  factors including coexistence of
other organisms. DDT persists in the environment despite the presence of organisms
capable of metabolizing the compound to at least DDD. Of particular importance is


                                   144

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the redox potential of the environment. Only under highly reducing conditions can
the necessary first step of conversion of DDT to ODD be achieved. This reaction
does not require microbial mediation. Further steps in DDT degradation may
require environmental conditions and microbial activities which have not yet been
elucidated. DDT, DDD, and DDE are highly persistent in all environments treated
with DDT.
                                     145

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                             SECTION 15

      CHLORINATED DIOXINS AND DIBENZOFURANS

  The chlorinated dibenzo-p-dioxins and dibenzofurans are produced as byproducts
during the formation of many other  chemicals, including 2,4,5-T, hexachloro-
phene, pentachlorophenoi and other chlorinated phenols, and polychlorinated
biphenyls (360). Chlorinated dioxins have been found in the fly ash and flue gases
from municipal generators in Switzerland, presumably due to pyrolysis of chloro-
phenol salts, and the formation of chlorinated furans has been tied to the pyrolysis of
polychlorinated biphenyls and polychlorinated diphenyl ethers. These compounds
are used as heat exchange fluids and as hydraulic liquids. From 3 to 25% of the
polychlorinated biphenyls burned may  be converted to chlorinated dibenzofurans
(360). There is no known technical use for the chlorinated dibenzo-p-dioxins, of
which 75 congeners can exist, and the chlorinated dibenzofurans, of which there are
135 theoretical congeners (360). The positional isomers of the dioxins vary greatly in
their acute toxicity and biological activity, and the most potent isomer, 2,3,7,8-
tetrachlorodibenzo-p-dioxin (TCDD), is considered the most potent low-molecular-
weight toxin known (mean lethal dose in guinea pigs 0.6 mg/kg body weight) (387).
  Interest in these compounds was generated after an epidemic of "chick edema
factor" in 1957 due to 1,2,3,7,8,9-hexachlorodibenzo-p-dioxin that caused the death
of millions of broiler chickens, and an accident in a chemical plant in 1976 in Seveso,
Italy that released a cloud of toxic materials including TCDD to the surrounding
environment (259, 356). TCDD has also been shown  to cause  chick edema factor
(387). The extreme toxicity of the compound of major interest, TCDD, has focused
most research on this isomer.

MICROBIAL METABOLISM OF DIOXINS AND FURANS

  To date none of the chlorinated or nonchlorinated dioxins or furans have served as
a sole source of carbon or energy for growth by any microorganism in a wide range of
screening and enrichment experiments (214,  259, 260, 301). Pseudomonas sp. NCIB
9816, which can utilize naphthalene as a sole carbon source, can cometabolize the
nonchlorinated  molecule  dibenzo-p-dioxin when salicylic acid is present in the
growth medium (259). Studies to  determine  the products of cometabolism were
conducted with a  mutant, Pseudomonas sp. NCIB 9816 strain 11, which oxidizes
naphthalene only  to ds-l,2-dihydroxy-l,2-dihydronaphthalene. Dibenzo-p-dioxin
is cometabolized to 2 neutral products, identified as cjs-l,2-dihydroxy-l,2-dihydro-
dibenzo-p-dioxin  and 2-hydroxydibenzo-p-dioxin.  When  the first  product  is
incubated aerobically or anaerobically with cell extracts of the parent organism in a
medium  containing NAD+, a  third product is formed and  was identified as 1,2-
dihydroxydibenzo-p-dioxin  (Figure  60). This metabolite completely inhibits or
inactivates the enzyme 1,2-dihydroxynaphthalene oxygenase, which in the parent
splits the naphthalene ring in the analogous  pathway.
  Similarly, a  Beijerinckia sp. grown on  dibenzo-p-dioxin  and  succinic  acid
produces 1,2-dihydroxydibenzo-p-dioxin (260).  Cell  growth is inhibited after 4
                                   146

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hours. Cell extracts incubated with 1,2-dihydroxydibenzo-p-dioxin show a brief
initial rate of oxidation followed by a decline to the nonenzymatic rate. The rate of
oxidation was determined polarographically using an oxygen electrode to measure
oxygen consumption. Two oxygenases  were isolated  from the cell extract, 2,3-
dihydroxybiphenyl oxygenase which also oxidizes 1,2-dihydroxydibenzo-p-dioxin,
and catechol oxygenase which has no activity against the dioxin metabolite. Both
these oxygenases are inhibited when incubated with cell extracts.
  This Beijerinckia sp. utilizes biphenyl as a sole carbon  and energy source. When
grown on succinic  acid plus biphenyl, resting  cells oxidize  1-chloro- and  2-
chlorodibenzo-p-dioxin at a high rate. Dibenzo-p-dioxin  and the isomers 2,3-, 2,7-,
and 2,8-dichlorodibenzo-p-dioxin are oxidized at a lower rate, followed by  1,2,4-
trichlorodibenzo-p-dioxin (260).
  A mutant strain of this species called Beijerinckia sp. B8/36 was isolated which
metabolizes several aromatic hydrocarbons to cis-dihydrodiols (260). When grown
on  succinic acid plus dibenzo-p-dioxin, a neutral product is formed which was
identified as  cj's-l,2-dihydroxy-l,2-dihydrodibenzo-p-dioxin  and has identical
characteristics to the product  of Pseudomonas sp. NC1B 9816 metabolism. Also
appearing in the medium is the metabolite 2-hydroxydibenzo-p-dioxin. The mutant
Beijerinckia sp. B8/36  cometabolizes  1-chlor-  and 2-chlorodibenzo-p-dioxin  to
neutral products which appear to be ds-dihydrodiols, but no products appear after
cometabolism with 2,3-dichloro- or 2,7-dichlorodibenzo-p-dioxin.
  An unidentified  bacterium  was isolated from contaminated Seveso  soil and
incubated aerobically in a complex nutrient medium containing I4C-TCDD (214,
351, 352). After 54 weeks 2 polar metabolites appeared.  One was  isolated in very
small quantities and was not identified. The other was found to be a hydroxylated
derivative. This microbial metabolite also appears in a  culture of P. testosteroni
strain G1036 after  incubation for 36 weeks and in a culture composed of a mix of 6
bacteria from Seveso soil. The metabolite was postulated to be l-hydroxy-2,3,7,8-
TCDD, assuming no chlorine rearrangement took place.
  TCDD was metabolized by Bacillus  megaterium to several polar metabolites
(358). The most active cultures were incubated with 5 //g/1 TCDD introduced in an
ethyl acetate carrier, a solvent which increases cell permeability. When ethyl acetate
was the carrier and the amount of soybean extract in the medium was reduced,  as
much as  55% of the dioxin was recovered as polar metabolites. TCDD was also
converted in small amounts to a polar metabolite in farm soil which had been
incubated for 2 months. The quantity of this metabolite did not increase with time
after 2 months. Other soils similarly incubated failed to  produce any metabolites.
Two strains of bacteria which converted TCDD  to polar  metabolites were isolated
from the farm soil  samples.
  A large screening study examined 100 bacterial isolates for ability to metabolize
TCDD (301). These strains all had  shown previous ability to metabolize persistent
pesticides, but only 5 showed some ability to metabolize  TCDD as determined by
thin-layer chromatography. The product or products were not identified.
  Degradation of  TCDD by an extracellular laccase (p-diphenol:oxygen oxido-
reductase) produced by the fungus Polyporus versicolor was investigated (68). Crude
enzyme extracts incubated with TCDD under a variety of conditions failed to modify
the  substrate.
  There has been one report on the microbial metabolism of dibenzofuran (77). A
comparison was made of the cooxidation of this compound by Cunninghamella
elegans, Beijerinckia sp. and Beijerinckia sp. B8/ 36 discussed previously with regard
to dibenzo-p-dioxin metabolism. The mutant strain oxidizes dibenzofuran to a
mixture of 1,2-dihydroxy-l ,2-dihydrodibenzofuran and the unstable 2,3-dihydroxy-
2,3-dihydrodibenzofuran which under acidic conditions dehydrates to a mixture of

                                    148

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2-hydroxy- and 3-hydroxydibenzofuran (Figure 61). The fungal culture forms a
much more stable 2,3-dihydrodiol which yields  2-hydroxydibenzofuran and 3-
hydroxydibenzofuran  only when heated with acid, although the ratios  of the 2
products are similar after bacterial or fungal metabolism. These results are consistent
with the unstable bacterial metabolite being of the cis configuration and the stable
fungal metabolite arising from an epoxide to form a trans configuration. The fungal
culture also forms  a small amount of 2,3-dihydroxydibenzofuran, and the parent
Beijerinckia sp. in the presence of NAD+ forms 1,2-dihydroxy- and 2,3-dihydroxy-
dibenzofuran.

DIOXIN PERSISTENCE AND DEGRADATION IN SOILS
  Evaluation of the persistence of TCDD in soils is complicated by the strong
sorptive properties of TCDD, making recovery for analysis difficult (301, 352). In
addition,  artifacts  may arise during the exhaustive extraction and analytical
procedures involved. TCDD incubated with lake sediment for less than 1 hour and
then extracted with solvents and analyzed by thin layer chromatography and I4C-
radioactivity showed 6 to 7% conversion by metabolism to metabolites, indicating
the generation of artifacts during the procedure or the presence of impurities (458).
Analysis of a  commercial I4C-TCDD preparation revealed the presence of 7%
contaminants, including TriCDD, some anisole isomers, and some other compo-
nents (352). Upon incubation, TCDD became less easily extractable while the other
components were in comparison  readily extractable, leading to artificial enrichment
of the isomer during the analysis.  This could cause misinterpretation of experimental
results. Formation  of metabolites would be expected to increase with time unless
precluded by a toxicity threshold or by further metabolism.
  Finally, since TCDD is only present in the environment as a contaminant of other
chemicals, the analytical procedure must be able to measure TCDD at levels lower
than  a few parts  per million  (197).  Recovery of I4C-TCDD as  measured  by
combustion from soils receiving 1.78 ppm  TCDD were 52% after  1 year from
Hagerstown silty clay loam containing 2.5% organic matter, and 67% from Lakeland
loamy sand containing 0.9% organic matter (246). At an application rate of 17.8 ppm,
89% was recovered from Hagerstown loam and 73% from Lakeland sand. No
metabolites were detected. Little MCO2 was evolved from TCDD-treated soils during
10 weeks' incubation.  Extracts of soil treated with 2,7-dichlorodibenzo-p-dioxin
(DCDD) contained a major metabolite in addition to the parent substrate (246). The
metabolite was not identified. About  5% of the added radioactivity in a 0.7 ppm
application of DCDD to soils was evolved after 10 weeks. TCDD incubated in lake
water for 589 days was not altered (458). However, a lake water and sediment system
incubated for the same length of time produced metabolites amounting to 1 to 4% of
the original substrate. These products were  polar (water soluble) and some were
extractable and some were nonextractable in chloroform. The addition of nutrients
enhanced formation of metabolites.

SUMMARY
  The chlorinated and nonchlorinated dioxins and furans have not yet been shown
to be utilizable as a  sole carbon source for growth and  energy. The parent
nonchlorinated  dibenzo-p-dioxin can be hydroxylated by several bacteria. The
hydroxylated products accumulate and are recalcitrant to further oxidation. Some of
the mono-, di-, and trichlorinated isomers  are also  oxidized by some bacteria,
although at lower rates. TCDD is also hydroxylated by a few species of bacteria. Not
all of the metabolites arising from TCDD metabolism have been identified. The
                                    149

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         Beijerinckia sp
        cis-1,2-DIHYDROXY-
     1,2-DIHYDRODIBENZOFURAN
DIBENZOFURAN
 2,3-EPOXIDE
1,2-DIHYDROXY
DIBENZOFURAN
                                                       2-HYDROXY-
                                                     DIBENZOFURAN
                                                       3-HYDROXY-
                                                     DIBENZOFURAN
           cis-2,3-DIHYDROXY-
                                trans-2,3-DIHYDROXY-2,3-
       2,3-DIHYDRODIBENZOFURAN  DIHYDRODIBENZOFURAN
                    2,3-DIHYDROXYDIBENZOFURAN
Figure 61.   Oxidation of dibenzofuran by Beijerinckia sp. and C. elegans. Bracketed
compound is hypothetical intermediate. Dashed lines indicate postulated reactions.
                       Adapted from Reference 77.
                                 150

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dibenzofurans have been less well studied, although it has been demonstrated that the
nonchlorinated substrate can be hydroxylated by both bacteria and fungi. Studies of
the metabolism  of these compounds have been hampered by the difficulty of
extraction and product analysis, and by the extreme toxicity of TCDD, the isomer of
greatest interest.
                                     151

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                            SECTION 16

    BIODEGRADATION OF CHLORINATED AROMATIC

                           COMPOUNDS
       IN SCALED-UP BIOLOGICAL WATER RELATED

                   TREATMENT PROCESSES

INTRODUCTION
  The  previous sections have  demonstrated that a wide variety of chlorinated
aromatic compounds are subject to biodegradation by a diversity of pure and mixed
bacterial cultures. The significance of this information relates to the  perceived
potential for both environmental and wastewater biodegradation and/or detoxifi-
cation  of chlorinated aromatic pollutants. The extrapolation of such laboratory-
derived results to environmental degradation and waste treatment is imprecise due to
optimization, acclimation  and high cell  density cultures  employed in  most
biochemical  and  physiological studies, which are rarely if ever met in real world
biodegradation scenarios. In addition, real world complexity of the environmental
matrix in which biodegradation occurs frequently necessitates the use  of imperfect
measures of  biodegradation that cannot readily be correlated with those  used to
assess biodegradation in a laboratory environment.
  Much of the work assessing biodegradation potential has been done in small-scale
laboratory glassware with the hope that adequate comparisons might  be drawn to
full-scale environmental or treatment systems. Many factors may vary  between the
small-scale lab systems and the full-scale  systems. A  summary of  factors that
influence organic biodegradability  is presented in Tables 6, 7, and 8 (385). Small-
scale lab tests can assess the importance of many of these factors in well-designed,
controlled experiments. However, many issues of importance in relating small-scale
test results to full-scale process performance depend on interactions of the  various
individual factors and the rates of material and biomass changes. These often are
influenced by the physical design of the system and mass and energy transfer
considerations. The turbulence and mixing potential of the system are also of major
importance.
  For these reasons, environmental scientists and engineers have turned to larger
scale experiments to simulate the design and physical features of the full-scale system,
whether a treatment or an environmental system. In these systems, material and
energy kinetics can be measured, competitive abiotic processes can be  studied, and
these results can be linked to mathematical models to describe the system and allow
scale-up to full-scale systems with more certainty.
  This chapter focuses on scale-up studies to determine the biodegradability of
chlorinated aromatics reported in the literature. Much information is  available on
continuous wastewater treatment systems. This was emphasized in this  chapter over
the environmental microcosm work, since the focus of this work is on treatability as
                                  152

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             TABLE 6. CHEMICAL FACTORS INFLUENCING
              ORGANIC BIODEGRADABILITY IN WATER*
       Chemical factors
          Consequences
Substrate structural
  considerations
    Molecular weight or size
    Polymeric nature
    Aromaticity
    Halogen substitution
    Solubility
    Toxicity
    Xenobiotic origin
Environmental factors
    Dissolved oxygen

    Temperature
    pH
    Dissolved carbon

    Particulates, surfaces
    Light
    Nutrient and trace elements
Limited active transport
Extracellular metabolism required
Oxygen-requiring enzymes
Lack of dehalogenating enzymes
Competitive partitioning
Enzyme inhibition, cell damage
Evolution of new degradative
  pathways
02-sensitive and Oa-requiring
  enzymes
Mesophilic temperature optimum
Narrow pH optimum
Organic/pollutant complexes are
  concentration dependent for growth
Sorptive competition for substrate
Photochemical enhancement
Limitations on growth and enzyme
  synthesis
*Reference 385.
            TABLE 7. BIOLOGICAL FACTORS INFLUENCING
               ORGANIC BIODEGRADABILITY IN WATER*
        Biological factors
          Consequences
Enzyme ubiquity
Enzyme specificity

Plasmid encoded enzymes
Enzyme regulation
Competition
Habitat selection

Population regulation
Low frequency of degradative species
Analogous substrates not
  metabolized
Low frequency of degradative species
Repression of catabolic enzyme
  synthesis required acclimation
  or induction
Extinction or low density populations
Lack of establishment of degradative
  populations
Low population density~of
  degradative organisms
*Reference 385.
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            TABLE 8. EXPERIMENTAL FACTORS INFLUENCING
                     ORGANIC BIODEGRADABILITY*
      Experimental factors	Problems encountered	

Analytical method
    Substrate disappearance     Competing abiotic processes
    Biotransformation           Complex analysis
    Mineralization               Incomplete biochemical pathways

Scale up/down                  Comparability among reactor designs
                                     and effects on kinetics

Feedstock complexity
    Chemically/biologically      Poor simulation and predictability
      defined
    Complex waste/wastewaters Difficult interpretation


*Reference 385.

opposed to persistence in the environment. Little information was found on scale-up
studies in the soil matrix.

PENTACHLOROPHENOL
   Pentachlorophenol has been studied in several scaled-up systems. These include
studies in aerobic "fiber-wall" reactor systems (140) as well as more conventionally-
designed lab activated sludge systems. (37, 129, 315.)
   A lab-scale test with a continuous, aerobic, "fiber-wall" reactor was used to study
the bio-oxidation of PCP in  a synthetic  and in an  authentic wood-preserving
wastewater (140). In the synthetic case, the concentration of PCP in the feed was 20
mg/1, the COD was 300 mg/1, the acclimation period was  15 days, and  the
operational period was 30 days. Reagent grade, commercial grade, and improved
commercial grade pentachlorophenol were  used. The improved commercial  grade
had fewer impurities, including chlorodioxins. Table 9 shows a survey of the results
of these experiments. Other concentrations were  also tested. These data show a
general inhibition of the disappearance of PCP  in the commercial preparation
relative to the reagent and improved commercial grade preparations. Presumably,
this is related to impurities present, possibly chlorodioxins. Actual waste rivaled the
synthetic tests in PCP degradation performance. No  proof of mineralization or
estimation of other fate mechanisms was given in these tests.
   A series  of lab-scale  continuous-stirred tank  reactors (CSTR) was used to
determine the aerobic biodegradation of PCP in wastewater treatment applications
(315). The testing protocol included a phase where the inoculum was acclimated to
PCP over 90 days from  initial concentrations of  1 mg/1 to 20 mg/1 of PCP in a
"fiber-wall" reactor. This sludge was then introduced into continuous-stirred tank
reactors with no sludge or cell recycle. The hydraulic residence times (HRT) and the
mean cell residence times (MCRT) were, therefore, equal and ranged from 3.2 to 18.3
days. Data collected  included disappearance data on COD and PCP  as well as
reactor suspended solids  concentrations. Proof of mineralization of PCP by use of
MC-PCP was employed in related batch studies and fate information was collected on
sorption and stripping mechanisms in adjunct batch experiments. No confirmatory


                                   154

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            TABLE 9. THE EFFECT OF PENTACHLOROPHENOL
        PURITY ON DISAPPEARANCE IN CONTINUOUS SYSTEMS*
Parameter
                      Improved
           Commer-   commer
Reagent      cial        cial      Actual
 grade      grade     gradef    waste
FeedPCP(mg/l)
Feed COD (mg/l)
Hydraulic residence
  time (hr)
Effluent PCP (mg/l)
Effluent COD (mg/l)
Activated sludge initial
  pentachlorophenate
  degrading capacity (mg
  PCP g cell - 1 hr -)
    20
   515

     6
 0.53-0.7
15.8-45.6
  20
515

   6
  1.9
29.1
 0.4-0.49    0.11
  20
515

   6
 0.3
52.8
            0.4
 17.8
1336

    6
  0.2
 216
*Reference 140.
 98% pure PCP.
 Blend from four manufacturers, 75-85% pure.
tSame PCP concentration as  commercial grade, "substantially reduced"
 chlorodioxins.  Dow Chemical Co., Improved Commercial  Grade  Penta
 (XD-8108.00L).

specific analysis of radiolabeled intermediates in the effluent or in the biomass are
offered.
  Results from this study include a first order kinetic rate constant, (the maximum
specific growth rate,//m, divided by the Monod saturation constant, Ks) of 0.00171
yug'd' with a minimum attainable PCP CSTR reactor concentration of 27 fig/1.
Aqueous phase concentrations of PCP ranged between 51-293 (ig/\ in the reactor.
PCP had little effect on the removal of other COD in this study. The batch fate testing
indicated  that neither stripping nor  sorption were significant PCP  removal
mechanisms and  PCP was mineralized with some carbon incorporated into the
cellular material.
  A synthetic waste containing PCP was treated in a continuous lab-scale activated
sludge system consisting of a 6.25-1 mixed liquor vessel and a 1.66-1 external clarifier
(129). Air was added at the rate of 6.251/min through a sintered glass sparger. Sludge
was wasted at 15 min intervals throughout the experiment. HRTs ranged from 8.9 to
10.4 hours and the MCRT  was  maintained at 6.2 days. Parameters measured
included total solids, sludge volume index, PCP and reducing sugar concentrations
in the clarifier, and clarifier effluent turbidity.
  Screened wastewater treatment plant sludge was acclimated to PCP using a fill and
draw reactor. An Arthrobacter sp. strain (ATCC 33790) was also added to another
acclimation reactor with the effect of a lag period reduction to 1 -3 days as opposed to
over 6 days for the unamended sludge.
  Steady-state  operation of the activated sludge reactor achieved reductions in
concentration from 40 mg/l PCP to about 1 mg/l. Unsteady-state transient
conditions were studied by increasing the feed to 120 mg/l. Shock loading effects
                                  155

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were analyzed with the use of kinetic models. Systems where the Arthrobactersp. was
added continuously showed a considerably improved transient response to the shock
load than did the acclimated systems without addition of the strain.
  Continuous lab-scale activated sludge units, consisting of 11-1 aeration factors (air
sparged in at * 21/min) and 4-1 external clarifiers with partial sludge recycle were
challenged with 8.6 mg/1 of PCP (37). The feed stream consisted of a pulp mill foul
condensate with substantial amounts of non-PCP carbon, largely as methanol. Mean
cell residence times were 4.9 and 9.3 days and hydraulic residence times were 25.4 and
24.0  hours, respectively. PCP was added as U-I4C-PCP  and  label analysis and
specific PCP analysis were performed on the mixed liquor supernate as well as waste
biomass and offgas. Other operating conditions are reported as well.
  Unlike the other studies, the emphasis of this study was to develop information on
PCP sorption on biomass and, therefore, a biomass not acclimated to PCP was used.
BOD6 and TOC removals from the aqueous phase were 63 to 69% and 86 to 96%,
while PCP removals were 11.6 to 7.0%, depending on the reactor MCRT. Essentially,
all of the PCP removal was either sorbed to or soluble in the aqueous component of
the waste sludge. No significant biological transformation or mineralization was
evident and stripping of PCP was below the detection limit. Good accountability was
found for both labeled and unlabeled  PCP.
  Batch sorption tests were also incorporated and comparison of the batch data and
the continuous runs suggests that the data fits a Langmuir-type isotherm with an
apparent saturation of the biomass at an aqueous phase concentration of 2 mg/1
PCP. The  PCP data collected at concentrations less than the apparent saturation
concentration agree well with a proposed sorption equilibrium equation based on the
PCP octanol-water partition coefficient. An equation proposed for estimation of
stripping based on the Henry's law  constant for associated PCP was found to
exaggerate the amount stripped. However, use of the Henry's law constant for the
pentachlorophenate form was expected to show better agreement. No such Henry's
law constant is available.
  Consistent with information in earlier chapters,  PCP has been found to
biotransform in scaled-up systems  using acclimated biomass or systems amended
with known PCP degraders. No scaled-up  studies have  attempted  to elucidate
transformation products other than 14CO2, but complete mineralization is strongly
indicated  in  at  least  one study.  Inhibition of other compounds  (potentially
chlorodioxins) on the  biomass has been noted and, therefore, the waste matrix in
which the PCP resides may be important in determining the extent and rate of
biodegradation. Conventionally-designed activated sludge systems  operating with
biomass not acclimated to PCP or experiencing transient shock loads either fail to
achieve biotransformation of-PCP or do so at greatly reduced rates. In these cases,
PCP removal from the aqueous phases is poor and sorption to biomass (or other
suspended solids) is expected to be a significant removal mechanism.
  In summary, effective continous  PCP degradation appears to require a biomass
with specific PCP  degradative capability  (PCP degrader sub-population) as
evidenced  by the importance of acclimation and  a steady  and transient-free feed.
Systems allowing longer cell residence times may also have an advantage over designs
incorporating short cell residence time, although evidence here is not conclusive.

CHLORINATED BIPHENYLS

  Several studies have occurred in which various mixtures and/ or specific congeners
of chlorinated biphenyls (PCBs) have been tested for biodegradability in larger,
continuous experiments. These have  included  tests  on consortia  taken from
operating wastewater treatment plants (200, 236, 438), as well as tests using specific


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 organisms (280). Still other research has been completed focusing on the fates of
 PCBs in environmental microcosms (272).
   PCB commercial mixtures (Aroclors 1221,1016,1242,1254,andMCS 1043)were
 tested for biodegradation using a lab-scale activated sludge test (438). Because of the
 test protocol, evaluation  of the HRT and MCRT was not possible. The sludge
 inoculum was obtained from a municipal treatment plant and  was acclimated for
 several weeks on a synthetic feed composed of glucose, nutrient broth, and KH2PO4.
 PCBs were not included in the acclimation feedstream. Initial MLSS concentrations
 were adjusted to 2500 mg/1.  PCBs dissolved in ethanol were injected into  the
 reactors. Disappearance of PCBs from the mixed liquor was measured using a
 hexane extraction followed by specific GC-EC or UV analysis. PCB spikes into the
 mixed liquor were used to measure analytical recoveries of PCBs from the liquor
 containing biomass and supernate.
   Sorption on biomass was checked by sonic homogenization and extraction of the
 mixed liquor from an Aroclor 1016 run and  extraction in the standard manner.
 Comparison with results without  sonic homogenization showed similar PCB
 recoveries. No sorption checks were made with higher chlorinated PCBs, nor was
 proof of mineralization or biotransformation reported. Stripping was checked with
 Aroclor 1221, MCS  1043, and Aroclor  1016 by use of hexane  offgas scrubbers
 connected to the reactors. Stripping rates of 4.2, 6.1, and 3.6%, respectively, were
 reported for the above PCB mixtures.
   Results of these tests, equating disappearance with degradation, are presented in
 Table 10. A 48-hour cycle with an addition rate of 1 mg PCB over this interval was
 used.

                    TABLE  10.   DISAPPEARANCE OF
              COMMERCIAL CHLOROBIPHENYL MIXTURES*
Mixture


Biphenyl
Aroclor 1221
MCS 1043
Aroclor 1016
Aroclor 1 242
Aroclor 1254
Percent
chlorine
of mixture
0
21
30
4-1
42
54
Percent
disappearance
during test
100
81+6
56+16
33 ±14
26 ±16
15 + 38

"Reference 438.

  A lab  study of activated sludge challenged with carbon radiolabeled 2,5,4'-
trichlorobiphenyl and 2,4,6,2',4'-pentachlorobiphenyl was performed to determine
the fates of these compourids in biological processes (200). Municipal sewage sludge
was placed in an aerated glass column and the offgas was scrubbed in hexane and
toluene to recover stripped I4C-PCB.  No acclimation of the sludge to PCBs was
reported. Initial concentrations were  0.178 mg/kg and  0.231 mg/kg trichloro-
biphenyl and pentachlorobiphenyl, respectively.
  Metabolic byproducts of the trichlorobiphenyl were found in the sludge while
none were found in the aqueous phase. One percent of the l4C-trichlorobiphenyl was
estimated to undergo biodegradation. The degradation products were found to be

                                   157

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less volatile than the parent compound and, therefore, accumulated in the biomass.
No evidence of degradation of the pentachlorobiphenyl was found in this study.
  A lab-scale semi-continuous test using a "fill-and-draw" technique (236) was
performed using municipal sludge acclimated over 3 months on 1,5 and 10/ug/l of
Kanechlor 500 (a  PCB  product with  its  main component being pentachloro-
biphenyl). The units were aerated for 12 hours and then settled for 0.5 hours, after
which time the clear supernate and sludge in excess of 25% of the reactor volume was
wasted. The reactor was recharged with synthetic feed containing the PCBs, glucose,
sodium glutamate, and inorganic nutrients.
  The BOD of the feed was 320 mg/1. Specific PCB analysis in the sludge included
centrifugation of the sludge, digestion of the solids  with methanol-KOH, and
subsequent hexane extraction. The aqueous phase was extracted with hexane and
extracts were combined, water washed, dried  with anhydrous Na2SO4, cleaned up
with a Florisil column, concentrated, and analyzed by GC-EC. COD measurements
and respirometric measurements were also taken during the study.
  In batch respirometric tests, oxygen uptake of biomass with 1 and 5 /ug/1 PCBs
was stimulated relative to the control without PCBs. Semicontinuous reactors fed up
to 10//g/l PCBs experienced  high BOD removal efficiencies (98.6 to 99.1%). Major
removals of PCBs in  the semicontinuous reactors were found at all feed concen-
trations over the 12 hour aeration period. Equilibrium concentrations were achieved
during the first hour of the period. Table 11 shows the distribution of PCBs in the
semicontinuous reactors for sludge acclimated at 1, 5, and 10 ^g/1 PCB.

            TABLE  11. DISTRIBUTION OF KANECLOR  500 IN
          ACTIVATED SLUDGE SEMICONTINUOUS SYSTEMS*
Sludge PCB
acclimation
concentra-
tion
M9/I
1
5
10
Initial
PCB con-
centra-
tion
/"9/I
0.6
0.48
0.85
Removal
in
wasted
sludge
(%)
45.6
75.9
81.5

Remaining
in
effluent
(%)
31.3
15.0
12.1


Unac-
counted
(%)
23.1
9.1
6.4
"Reference 236.

  The majority of the PCBs charged to the system were removed with the wasted
sludge. The authors tested stripping of PCBs in their semicontinuous reactors at air
flow rates of 0.1 1/min of  air per 1 of mixed liquor and  found significant
disappearance (65%) after 20 h aeration. They conclude that stripping could account
for the PCB losses experienced in the semicontinuous runs. Finally, challenge of a
municipal anaerobic digester sludge with 31 jug/1 of PCBs (wet weight) incubated at
38°C for 40 days showed no disappearance of the starting PCB material. Resistance
to anaerobic biotransformation was concluded.
  A  lab-scale continuous aerobic  study of the degradation of Aroclor 1221 by a
Pseudomonas sp. strain 7509 culture is reported (280). Inocula were acclimated on a
feed  in which Aroclor 1221  was the sole carbon source. A fermentation vessel was
used as the reactor and raw sewage with a BOD5 of 140 to 170 mg/1, fortified with 20
mg/1 each of nitrogen (as NH4C1) and phosphorus (as KH2PO4+K2HPO4), was
                                   158

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amended with Aroclor 1221 at concentrations of 50 and 100 mg/1. The waste was fed
at rates between 13 and 91 ml / hr to the 14-1 reactor in which the agitator rate and the
dissolved oxygen concentration was monitored. The HRT ranged from 4.6 to 32 days
and was equal to the MCRT.
  The broth (or mixed liquor) was sampled periodically and acidified prior to hexane
extraction. GC-FID was used for disappearance analysis. No distinction can be made
between the compounds in the aqueous and biomass compartments of the mixed
liquor since the solids were not separated prior to analysis. Neither stripping nor
sorption  was measured and no proof of transformation or mineralization was
reported.
  At high HRTs (16 to 32 days), all of the Aroclor 1221 fed disappeared. At lower
HRTs (4.6 to  10.7 days), some of the specific congeners began to appear in the mixed
liquor indicating, according to the author, a preference of the organisms for certain
congeners. The implication is  that under  nonstressed  conditions, biphenyl,  2-
chlorobiphenyl, and 4-chlorobiphenyl will degrade readily while  2,2'-dichloro-
biphenyl, 2,4'-dichlorobiphenyl, and 4,4'-dichlorobiphenyl are  more recalcitrant.
2,4'-dichlorobiphenyl was found to build up as an indicator of the lower biodegra-
dation rates. Aroclors 1016 and 1254 were also tested with resulting accumulation of
all of the components. Switching from continuous  to batch operation indicated
disappearance of many of those congeners. Specific congeners were not identified in
the Aroclor 1016 and 1254 runs.
  Fates of 2,2',4,5'-tetrachloro-, 2,2',4;4',5,5'-hexachloro- and  2,2',3,3',4,4',5,6'-
octachlorobiphenyls were introduced into a model system that included sediment,
water, and air compartments (272). The model systems were fitted with a special gas
bubbling device to enable investigation of removal by jet drop entertainment, and
were operated in the dark. Anaerobic, sterile (bactericidal HgCl2), and aerobic biotic
systems with  and without macroinvertebrates were studied. The macroinvertebrates
were grown in separate vessels and then added to the model systems. The equivalent
of 7500 chironomids/m2 (Chironomus plumosus-type) and 25,000 tubificids/m2
(Tubifex tubifex) were added to the system. Gas flow to the system was estimated to
be from 0.00005 to 0.003 ml/min of gas per ml of water. Thus,  air flow to liquid
volume ratio  was from 30 to 2000 times less than that found in diffused air biological
treatment systems.  14C-PCB was used and no specific compound analysis was
performed on the parent compound or possible metabolites or conversion products.
Congeners recovered in each experiment averaged 68%,  40%, and 31% for tetra-,
hexa-, and octachlorobiphenyls, respectively. Distribution of PCBs in the various
tests are found in Tables  12, 13, and 14.
  Although this experiment was intended to simulate a natural ecosystem and clearly
differs from experiments on engineered systems, some conclusions are suggested that
may relate to engineered treatment systems. First, although the vast majority of the
congeners partition with the sediments  (90 to 99.9%),  the presence of  biomass
(especially the macroinvertebrates) was a determinative factor in the partitioning
related to dispersion of sediment particles and jet-drop entrainment. In an engineered
system, the  turbulence of the system may  override the turbulence from the
macroinvertebrates with the result of much greater suspended material  potentially
being available for adhesion to walls and other surfaces.  Of potentially greater
importance is increased jet-drop entrainment, because of higher suspended solids
concentrations and substantially higher air flow  to water volume ratios in the
engineered system. Truly aerosolized jet drops may be collected in scrubbers or filters
and therefore may be included in measures of stripping potential, even though the
substrate  is actually not  a vapor but a mechanically-carried liquid-solid droplet.
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Unfortunately, no literature addressing this potentially important removal mech-
anism in engineered systems is evident.
     TABLE 12. DISTRIBUTION OF 2,2',4,5'-TETRACHLOROBIPHENYL
             IN A SEDIMENT-WATER-AIR MODEL SYSTEM*


Compartment


Sediment
Dissolved in water
Particles in water
Macroinvertebrates
Glass walls
(particle adhesion)
Glass walls
(extractable)
Air filters
Vessel stoppers
Surface microlayers
Jet-drop impactors
Aerobic
with
macro-
inverte-
brates
97.7T
0.03
0.06
0.55
0.20

0.10

0.03
0.06
0.03
0.93
Aerobic
without
macro-
inverte-
brates
98.3T
0.06
0.03



0.02

0.01
0.08
0.01
1.48


Sterile An-
aerobic

99.8! 99.3t
0.02 0.03


0.02



0.02
0.04

0.18 0.04
•Reference 272.
fAII values as percent of recovered compound.
IN A SEDIMENT-WATER-AIR


Compartment


Sediment
Dissolved in water
Particles in water
Macroinvertebrates
Glass walls
(particle adhesion)
Glass walls
(extractable)
Air filters
Vessel stoppers
Surface microlayers
Jet-drop impactors
Aerobic
with
macro-
inverte-
brates
96. 1t
0.06
0.14
2.73
0.48

0.02

0.01

0.04
0.36
MODEL
Aerobic
without
macro-
inverte-
brates
99.9t



0.01





0.03
0.03
SYSTEM*


Sterile


98.9 1





0.17

0.02
0.02

0.10



An-
aerobic

99.9T
0.02


0.02

0.02



0.03
0.03
tAII values as percent of recovered compound.
                                160

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             TABLE 14. DISTRIBUTION OF 2,2',3,3',4,4',5,6'-
                      OCTACHLOROBIPHENYL IN A
                SEDIMENT-WATER-AIR MODEL SYSTEM*



Compartment


Sediment
Dissolved in water
Particles in water
Macroinvertebrates
Glass walls
(particle adhesion)
Glass wails
(extractable)
Air filters
Vessel stoppers
Surface microlayers
Jet-drop impactors
Aerobic
with
macro-
inverte-
brates
90.0 1
0.10
1.00
3.00
4.67

0.07

0.01
0.30
0.07
0.99
Aerobic
without
macro-
inverte-
brates
99.61
0.03
0.02

0.0





0.03
0.24


Sterile An-
aerobic

99.8t 99.4t
0.06 0.45
0.09 0.01



0.01


0.05
0.11
0.04
'Reference 272.
tAll values as percent of recovered compound.

  A single scaled-up study has been reported to date that conclusively demonstrates
PCB biodegradation. In this study, metabolic byproducts accounting for about 1%
of the  trichlorobiphenyl were  recovered.  No  other studies  present conclusive
evidence  for biodegradation. This is largely due to use of parent compound
disappearance  data for analysis  and lack of proof for biotransformation or
mineralization. No work is evident using labeled congeners to support an argument
for enzymatic  processes. Several  studies have shown major  potential for PCB
disappearance  related  to stripping and  to sorption on  biomass or other solids
(sediments). Analytical  difficulty in extractive recovery of the higher chlorinated
congeners from sediments suggests that in the absence of methods to digest the
biomass or to determine specific recoveries of the higher congeners in the biomass,
extractive analysis of the biomass or the total mixed liquor (biomass and supernate)
may not recover substantial portions of the parent compounds strongly sorbed to the
cellular or solids matrix. Spiking with a PCB mixture and complete extractive
recovery is not conclusive proof of recovery of PCBs from unknown biomass samples
unless no saturation effect of the biomass with sorbed PCBs exists and sufficient time
is  allowed for  the PCBs to achieve  equilibrium partitioning with the biomass.
Absence  of either condition would lead to more of the spiked sample  residing in the
aqueous  compartment and misleadingly high apparent recovery values. In other
words, special caution must be taken when equating PCB-spiked mixed liquor or
biomass recovery with actual sample PCB recovery.
  The strongest evidence of biodegradation. notwithstanding the above comments,
is the study by Liu (280) in which ratios of the specific congeners in Aroclor 1221
change depending on the operating conditions. It is difficult to  see how sorption or
stripping mechanism would vary as widely as reported for the same congeners under
different  operating conditions unless enzymatic processes are at work. Also  the PCB


                                   161

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mixture studied is comprised of congeners that, in other microbial tests (reported in
an earlier chapter), have been shown to undergo biotransformation.
  In general, further scaled-up studies on PCBs are needed that include analytical
protocols  to offer proof of biotransformation or  mineralization,  quantify the
sorption and stripping mechanisms, and study other potentially important mech-
anisms  such as jet-drop  entrainment  in engineered systems, before  engineered
biological processes may be considered  for PCB treatment.

DICHLOROPHENOL
  A lab-scale activated sludge reactor with sludge recycle (liquid volume  of aeration
tank, 3 1) was used to study the simultaneous biodegradation of 2,4-dichlorophenol
(DCP) and phenol  (33). The phenol concentrations ranged from  14.9 to 45.7 mg/1
and the DCP concentrations ranged from 52.4 to  121 mg/1. A 1:1 carbon ratio of
both substrates was desired. The sludge was previously acclimated to phenol and
then was acclimated to DCP by gradual replacement of the phenol in the feed with
DCP. The acclimation  process required 70 days to  complete. The reactors were
operated with HRT between 2.5 to 6.25 hr and MCRT of between 1.75 and 10.7 days.
  MLVSS concentrations ranged between 46 and 299 mg/1.  Analysis focused on
substrate disappearance with no determination of stripping or sorption. Chloride
analysis as a test of mineralization and COD determinations did not account for the
phenol or DCP disappearance, implying biotransformation of the substrates.
  Biokinetic rate constants  for the runs yielded disappearance rate constants of
0.00098 1 mg-'hr1 for phenol and 0.045 hr1 for DCP. The Monod  half-saturation
constant for DCP was 63 mg/1. Yield coefficients for phenol and DCP were 0.67 mg
VSS/mg phenol and 0.39 mg  VSS/mg DCP, respectively. A combined biomass
decay coefficient of 0.014 hr1 was presented.
  A mathematical rationale  leading  to deterministic estimates  of fates from
biological waste water treatment processes has been proposed (38). For continuous,
complete-mix,  activated sludge units where the biological disappearance  of the
parent compound  is  described  by a rate  equation  first  order  in substrate
concentration, and where sorption is occurring at concentrations below any potential
biomass saturation concentration,  the following equations are proposed for first
estimates of the percent substrate stripped, sorbed, and wasted in the waste biomass,
and biotransformed to another compound:
                        REMS =
                                                                        (2)
                                     1 + A + S + B
                       REMS,  =           S                            (3)
                                     1 + A + S + B
                        REMb  =    _	2	                      (4)
                                    1 + A + S + B
                                   162

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                        REMe =    	'                             (5)
                                     1 + A + S + B
                             A = HRT    X  K-"" fL                     (6)
                                        1000 PL MCRT
                 S =  HRT  K"  =  -2^- 3.71 X 10"'  (H,.)1045                 (7)
                             B  = HRT Kb                               (8)
 where:     REMS, REMS1, REMh, REMe are the percent removals of the substrate
           from the system  by the sorption, stripping, biological transformation,
           and effluent fate  mechanisms, respectively,
           HRT is the hydraulic residence time of the activated sludge system (hr),
           X is the concentration of biomass as  MLSS (mg/1),
           Kow is the substrate octanol-water partition coefficient (concentration in
           octanol/concentration in  water),
           f L is the fraction of lipids or lipophilic compounds in the biomass (weight
           fraction),
           PL is the mean density of  the lipophilic biomass compounds (g/1),
           MCRT is the mean cell residence time for the biomass in the systems (hr),
           Qa,r/V is the  ratio of the air flow rate into the system to the  system
           hydraulic volume (min '),
           Kb is a biological disappearance rate constant, first order in substrate
           concentration (hr"1).

  Equations 6 to 8 are discussed individually in separate papers (37, 385, 437).
  If assumptions are made such that the equations described above are applicable to
the  DCP experiment discussed earlier, (complete-mix system, Q,,,/V =0.1 min"1, Hc
of DCP = 13.4torr L mof1, Kowof DCP= 1202, etc.) then the data reported  can be
used to calculate Kb for each experimental series and the mechanism removals can be
estimated. Table 15 presents the results of this analysis.  It may be concluded from
this analysis that stripping can be a significant removal mechanism, especially in the
instance where  Kb is relatively slow. In run  4, 16% of the DCP removed from the
system (other than  in the effluent) was stripped. As the biotransformation rate
increased, the stripping potential was vastly reduced to less than 0.1% of the total
DCP removed (other than in the effluent) (Run 1). Sorption of DCP to the waste
sludge taken from the system is not a significant fate mechanism for DCP. However,
depending  upon the level of extracellular water  wasted with the biomass  (waste
sludge solids concentration),  more DCP could be lost in the waste sludge than that
shown in Table 15.
                                    163

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          TABLE 15. FATE ESTIMATES OF 2,4-DICHLOROPHENOL
           REMOVAL FROM A LAB ACTIVATED SLUDGE SYSTEM
                     USING PROPOSED EQUATIONS*
                                          Percent removal of DCP by
Calculated

Run
1 +
2+
3t
4t
MCRT
(hr)
257
42
109
20
HRT
(hr)
25
6.25
6.25
6.25
K"
(hr-i)
0.075
0.043
0.070
0.020

Sorption
0.008
0.67
0.25
0.78
Biotrans-

Stripping
0.034
2.0
1.8
2.2
forma-
tion
63.0
21.6
29.8
10.8

Effluent
33.6
76.8
68.1
86.2
 +Runs with phenol and 2,4-dichlorophenol at a ratio of 1 to 1 carbon from
  each substrate.
 tRuns with glucose and 2,4-dichlorophenol at a ratio of 1 to 1 carbon from
  each substrate.
  Finally, the first order biotransformation rate constants, Kb, are similar to values
reported  by Beltrame, et al. (33)  derived using  more conventional empirical
biokinetic rate constant methods (K> = 0.045 ± 0.005-'). Calculated rate constants
trorn  I able 10 show wider variance and  indicate strong relationships between
biological rates and the MCRT.
TRICHLOROCARBANILIDE
  Trichlorocarbanilide (TCC) was studied in both lab-scale batch flask tests and
continuous activated sludge  systems (184a). TCC  with  MC  label on  the 4-
                                                                (DCA)
  Sludge was obtained  from a municipal  wastewater treatment plant. In the
a 400C ± 1 af Ml'sV e,bn°rSS C°ncentration and * *™ rates were control ed
at 4000 figl\ as MLSS and 0.05 standard ft3hr of CO,-free air  Offgas was traooed
m an amme solution to recover HCO, The .«C content^ tne^SL^LSJ

stream0"1           2 reC°Very method 2°° ^ of TCC ^ added to the feed
  Batch flask tests showed that 90% of the theoretical "CO2 evolved from incubation
  Sorption of TCC to activated sludge was determined by contacting activated
                                 164

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      TABLE 16.   FATE OF TRICHLOROCARBANILIDE IN LAB-SCALE
                     ACTIVATED SLUDGE SYSTEMS*
           Substrate               Stream      % of Feed   % Recovered
14C-PCA-TCCt

14C-DCA-TCC§



Effluent
14C02
Activated
sludge
Effluent
14C02
Activated
sludge
3.2
56.1
34.1
30.3
25.9

35.2
93.4


91.4


 *Reference 184a.
 fTrichlorocarbanilide with a labeled 4-Chloroaniline ring.
 §Trichlorocarbanilide with a labeled dichloroaniline ring.

theoretical 14CO2 was evolved from the chloroaniline ring. Table 16 highlights the
fate of the radioactivity in the continuous tests.
  Although no  specific analysis of the sludge  was undertaken to determine the
chemical composition of I4C found in the sludge (i.e., parent compound, metabolites,
cellular material), such analysis was done on the effluent. Chloroaniline, dichloro-
aniline, aniline condensation products, and unknowns were found in addition to the
TCC parent compound.
  This study convincingly supports conclusions related to the biotransformation and
mineralization of TCC. These findings are consistent with those stated in an earlier
chapter  on chloroaniline herbicides even though this molecule's structure  varies
somewhat. In addition, this study is among the earliest work found that considered
major fate mechanisms and offers proof of biotransformation or mineralization in a
scaled-up  biological  wastewater system treating  chloroaromatic  compounds.
Unfortunately, lack of information on the MCRT limits the calculation of biokinetic
rate  constants  and the direct extrapolation  of these results to other design
configurations.

DICHLOROBENZENE

  A pilot scale activated sludge system was operated on a side-stream of sewage from
the city of Zurich, Switzerland (303). 1,4-Dichlorobenzene was present at all times in
the system  feed and it was  used as an indicator compound to determine the
nonbiological removal mechanisms of stripping and sorption on the biomass. The
major assumption made in this study was that 1,4-dichlorobenzene was conserved
and was not biotransformed at the operating conditions of the  study. The aeration
vessel volume was 11.3 to 15m3 and the HRTs ranged from 2.5 to 6.5 hr with MCRTs
ranging from 74 to 182 hr. Sorption and stripping mechanisms  were quantified but
proof of biotransformation (or the absence of) was not provided.
  The fates of DCB were reported to be 72% stripped, less than 3% sorbed on wasted
sludge, 10% in the effluent, and 15% unaccounted. Data presented in an earlier
section indicated that chlorobenzenes (except hexachlorobenzene) can be mineral-
ized but there is a lack of knowledge  on the biochemical pathways. The DCB not
accounted for in this study may be undergoing biotransformation but the variance in
the material balance related to analytical  shortcomings precludes  a definite

                                   165

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statement. A study discussed  later in this  chapter offers  evidence  of DCB
biotransformation.

COMBINED STUDIES ON SEVERAL CLASSES OF

CHLOROAROMATICS

Lab and Pilot Studies
  Several studies have occurred using a given experimental protocol on a variety of
chloroaromatic compounds. Table 17 shows the classes of chloroaromatics studied
for each of the these compounds.
  Benzoic acid, 2-chlorobenzoic acid, 3-chlorobenzoic acid, 4-chlorobenzoic acid,
2,4-dichlorobenzoic acid, 2,5-dichlorobenzoic acid, 2,6-dichlorobenzoic acid, 3,5-
dichlorobenzoic acid, phenoxyacetic acid, and 2,4-dichlorophenoxyacetic acid were
studied in lab-scale continuous flow reactors resembling chemostats (391). The HRT
and MCRT were equal in these reactors. The reactors were completely mixed by
aeration and used suspended biomass. The lag for acclimation of municipal biomass
to the specific substrates was determined. Kinetic disappearance data were collected
on the specific substrates and on dissolved  organic carbon (DOC) as well. Batch
testing also was performed to determine kinetic rate constants for comparison with
the continuous tests.  Proof of mineralization of the specific substrates was
determined by measurement of chloride ion release.
  The lag for acclimation of the initial sludge for the monochlorobenzoic acids was
in the range of 10 to  20 days.  The biomass began to show  acclimation  to 3,5-
dichlorobenzoic acid at about 20 days but the acclimation process was continued
through 100 days. 2,5-dichlorobenzoic acid became acclimated abruptly at 100 days.
2,4- and 2,6-dichlorobenzoic acids did not  show acclimation during this  testing
protocol.
  The author argues that long  term acclimation for some of the compounds is
evidence for genetic changes in the organisms as opposed to enzyme induction  or
population effects.  Maximum specific growth rates, /um, the Monod half-saturation
constant, Ks, and the yield coefficient, Y, are shown in Table 18.
   Good agreement was found between //m and Ks for the continuous flow reactors
and associated batch tests. The  MCRT of the systems were found to be strongly
related to to the effluent concentrations of the specific substrates. MCRTs of 3 to  15
days were required to achieve a 0.5 mg/1 effluent concentration of the monochloro-
benzoic acids while 6  to  50 days  were  needed to achieve  0.25 mg/1 effluent
concentrations.

         TABLE 17. CLASSES OF CHLOROAROMATICS STUDIED
                  IN SEVERAL EXPERIMENTAL STUDIES
                                               Reference
          Compound             61     127   250   251    391    416

Pentachlorophenol                 X            XX            X

Chlorinated Biphenyls
 Aroclor 1242                    X
 Aroclor1254                    X
                                                              (continued)

                                   166

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TABLE 17. (continued)

Chlorophenols
 2-Chlorophenol                  X
 2,4-Dichlorophenol               X      X     X     X
 2,4,6-Trichlorophenol             X

Chlorobenzenes
 Chlorobenzene                   X
 1,2-Dichlorobenzene             X      X     X     X
 1,3-Dichlorobenzene             X      X
 1,4-Dichlorobenzene             X      X
 1,2,4-Trichlorobenzene           X      X
 Hexachlorobenzene              X

Chlorobenzoic acids
 2-,3- and 4-chlorobenzoic acid                               X
 2,4-, 2,5-, 2,6-, and
    3,5-Dichlorobenzoic acid                                   X

Chlorophenoxy compounds
 2,4-Dichlorophenoxyacetic acid                              X
   Benzoic acid, 2-chlorobenzoic acid, 3-chlorobenzoic acid, 4-chlorobenzoic
 acid, 2,4-dichlorobenzoic acid, 2,5-dichlorobenzoic acid, 2,6-dichlorobenzoic
 acid, 3,5-dichlorobenzoic acid, phenoxyacetic acid, and 2,4-dichlorophenoxy-
 acetic acid were studied in  lab-scale continuous flow reactors resembling
 chemostats (391). The HRT and MCRT were  equal in these  reactors. The
 reactors were completely mixed by aeration and used suspended biomass. The
 lag  for acclimation  of municipal biomass to the specific substrates was
 determined. Kinetic disappearance data were collected on the specific
 substrates and on dissolved organic carbon (DOC) as well. Batch testing also
 was performed to determine kinetic rate constants for comparison with the
 continuous tests. Proof  of mineralization of the specific substrates was
 determined by measurement of chloride ion release.
   The substrates were mineralized to CO2 and cellular material but no fate
 measurements  were made on stripping or sorption mechanisms. Performance of
 systems with glucose and the 2,4-dichlorophenoxyacetic acid indicated no effect of
 glucose on 2,4-D disappearance but lower glucose disappearance rates related to
 2,4-D presence. However, no strong inhibition ortoxic effects of the substrates on the
 biomass were  seen  at feed  concentrations of 50 to 200  mg/1. The effluent
 concentration is, of course, much less than the feed concentrations.
   A major study on a variety of organics was undertaken (250,251,416). Continuous
 lab-scale activated sludge units were challenged with chlorophenols, chlorobenzenes,
 and pentachlorophenol as well as a number of other organics. These compounds
 were added to a synthetic "base mix" containing ethylene glycol, ethyl alcohol, acetic
 acid, glutamic  acid, glucose, phenol, and various inorganic nutrients. The chloro-
 aromatics were added such that the BOD5 achieved was ~ 250 mg/1. HRT was held to
 about 8 hours and the MCRT ranged between  43 and 146 hrs. Stripping was
 measured  by trapping the organics from the offgas on a solid sorbent and  all specific
 compound analysis generally followed  EPA analytical protocols. Methodology for


                                   167

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           TABLE 18.  BIOKINETIC RESULTS FOR 2,4-D AND
                       CHLOROBENZOIC ACIDS*
Compound
2,4-Dichlorophenoxyacetic acid
2-Chlorobenzoic acid
3-Chlorobenzoic acid
4-Chlorobenzoic acid
2,5-Dichlorobenzoic acid
3,5-Dichlorobenzoic acid
M.**
(day-')
2.3
1.0
0.6
1.2
0.6
0.05
Kst
(mg/l)
5.4
2.4
2.0
1.1
1.5
25.3
Ytt
(mg/mg)
0.14
0.22
0.14
0.25
0.16

 *Reference 391.
**Maximum specific growth rate.
 fMonod half-saturation constant.
ftYield coefficient.

         TABLE 19.  FATE OF SEVERAL CHLOROAROMATICS IN A
                LAB-SCALE ACTIVATED SLUDGE SYSTEM*
                                           Percent removed by
           Compound              Stripping     Sorption     Biotrans-
                                                             formation
2,4-Dichlorophenol
Pentachlorophenol
1 ,2-Dichlorobenzene
95.2
0.58 97.3
21.7 78.2
  *Reference 250, 251,416.

 sorption quantitation was not clear. No proof of mineralization or biotransfor-
 mation was offered. Acclimation to the compounds was allowed for 4 weeks before 2
 months of continuous data collection. The reactor volume was 31 for aeration and 3.3
 1 for an internal clarifier. Air flow to the reactors ranged from 2 to 3 1/min. Table 19
 presents fate data on specific chloroaromatics. These studies utilized an air flow-to-
 liquid volume ratio greater than 1 min~1 and the resultant data probably exaggerates
 the stripping mechanism. Disappearance attributed to biotransformation for DCP
 and PCP are in agreement  with earlier studies reviewed in this chapter. Biotrans-
 formation of 1,2-dichlorobenzene is in conflict with a study discussed earlier but is
 consistent with general predictions on the biodegradability of chlorobenzenes.

 Full-Scale Studies

   Many full-scale  biodegradation studies  on chloroaromatics  are  reported.
 However, only a few have attempted to quantify the abiotic fates of the compounds.
 Also, variation in the waste composition and flow and in other physical variables
 often makes it difficult to interpret the results. Two of the most significant examples
 of full-scale plant studies are discussed here.
    A study to  determine the  fates of priority  pollutants for 50 publicly  owned
  wastewater treatment plants has generated input-output data on several chlorinated
                                    168

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aromatics including 2-chlorophenol, 2,4-dichlorophenol, 2,4,6-trichlorophenol, pentachloro-
phenol,  chlorobenzene, 1,2-dichlorobenzene,  1,3-dichlorobenzene, 1,4-dichloro-
benzene, 1,2,4-trichlorobenzene,  hexachlorobenzene, Aroclor 1242, and Aroclor
1254 (61). This work was designed to allow statistical analysis on the occurrence and
fates of priority pollutants in the system feedstream, intermediate process streams,
system effluent, and waste sludge streams. The material that disappeared in the
process  was reported but since no air sampling was undertaken, no quantitative
attempt  was made to distinguish between stripping and biotransformation removal
mechanisms. EPA analytical protocols were used and often the compounds were
grouped into the "volatile", "acid extractable", and "base neutral" groupings arising
from the analytical workup. Sampling periods were for approximately 6 days with 24
hr composite samples,  and compounds were often at concentrations near the
analytical detection limit. Since the influent was often variable and the sampling
period was of the same order of magnitude as the plant's MCRT, material balance
data and conclusions must be viewed with caution.
  A related study was performed on a single publicly-owned wastewater treatment
plant  for a 30 day period (127). Over this period, variations in influent flow and
substrate concentrations could  be more  precisely  quantified  and conclusions
regarding compound fate could be made. Table 20 summarizes the removal data of

  TABLE 20. MASS REMOVAL OF CHLOROAROMAT1C COMPOUNDS IN A
             FULL-SCALE WASTEWATER TREATMENT PLANT*
                                           Percent removal inf
           Compound              Primary     Secondary     Overall
                                  treatment    treatment*}    treatment
2,4-Dichlorophenol
1 ,3-Dichlorobenzene
1 ,4-Dichlorobenzene
1 ,2,4-Trichlorobenzene
2
14
0
12
46
30
88
79
47
40
88
82
 'Reference 127.
 t Calculation includes (total mass accounted for in minus total mass accounted
  for out)/total mass accounted for in. Thus, compound in the aqueous effluent
  is combined with that found in the waste solids. Removal mechanisms here
  are biotransformation, stripping,  and other abiotic mechanisms  excluding
  sorption.
 §Based on activated sludge units alone.

 2,4-dichlorophenol. 1,3-dichlorobenzene, 1,4-dichlorobenzene. and 1,2.4-trichloro-
 benzene found in this study. The removal calculation sums the substrate entering the
 system from all streams and the substrate leaving in all streams. Thus, sorption and
 effluent removal mechanisms are not considered "removed" whereas biotransfor-
 mation. stripping and other abiotic mechanisms, excluding sorption. are  considered
 "removed"
  Full-scale plant data requires special planning and careful implementation to yield
 satisfactory data on biotic  and abiotic removal mechanisms. Conclusive material
 balances often  are  impeded  because  of  feed  variability  and  low  compound
 concentrations. Use of labeled compounds is expensive at large scale and questions
 related to the environmental release  of labeled compounds exist. Therefore,  the
                                   169

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capability of collecting data leading to proof of biotransformation or mineralization
is limited.

SUMMARY
  In  order  to  conclusively  establish  biodegradation of  chlorinated  aromatic
compounds in larger scale systems and to collect data that are of use in extrapolation
and system design, several factors must be included in the experimental design. These
include: (1) measurement or prediction of abiotic fate  processes, (2)  proof of
biotransformation or mineralization, and (3) suitable measurement and reporting of
important process variables relating to the calculation of biokinetic rates. Failure to
include these factors leads to inconclusive results on compound fates or the inability
to use the data for predictions on other (even similarly designed) systems.
  Calculation of biokinetic rate constants based only on "removal" leads to wide
variances in the rate constants for compounds with major abiotic fate tendencies and
precludes reliable scale-up and more direct comparison between systems. Only a few
scale-up studies are available on chlorinated aromatics in general, and only a subset
of these contain data suitable for drawing conclusions relative to biodegradation and
which allow comparison between systems. At times, coupling the results from several
studies may allow an enlightened judgment regarding biodegradation, but there is no
substitute for a single well-designed  study. In general, much additional work is
needed to generate reliable scale-up data for biological treatment  of chloroaromatic
compounds.
                                    170

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                             SECTION 17

OVERVIEW OF MICROBIOLOGICAL DECOMPOSITION OF

         CHLORINATED AROMATIC COMPOUNDS

  Most of  the studies reviewed here have explored the metabolism of a single
compound by a single organism. A few have reported the metabolites formed in soils
or by contrived microbial consortia. Taken together, however, these studies indicate
the potential ultimate fate of the chloroaromatic compounds.
  The  chlorophenoxy  herbicides and the chlorobenzenes can be metabolized to
chlorophenols (Figure  62). Chlorophenols may in turn be metabolized to chloro-
anisoles, but the most common route of biodegradation is to chlorocatechols.
Phenylamide herbicides and other compounds with nitrogen-containing substituents
are metabolized  to chloroanilines. The chloroanilines form a variety of products
including chlorocatechols (Figure 63). Other products represent alterations of the
aliphatic moiety.
  Chlorocatechols, in turn, may be metabolized by several different mechanisms to
nonchlorinated ring cleavage products (Figure 64). There are two main pathways.
One results from meta cleavage to form a chlorohydroxymuconic semialdehyde
which, after loss of the chloride, forms pyruvate and an  aldehyde. The second
pathway involves ortho cleavage to form /8-ketoadipic acid via chloromuconate. The
products of this pathway that are incorporated into cell constituents are succinate
and  acetyl-CoA.
  Chlorobenzoic acids may be metabolized by 3 routes (Figure 64), the first through
protocatechuic acid (a substituted catechol) to 3-ketoadipic acid. The second route
metabolizes chlorobenzoic acids through chlorosalicylic acid to maleylpyruvic acid.
Chloronaphthalenes are also metabolized through chlorosalicylic acid. Anaerobic
metabolism of chlorobenzoic acids involves reductive dechlorination to benzoate
followed by formation of CH4 and CO2. The ultimate products of each of these
metabolic pathways are either available for incorporation into cellular material or
represent ultimate mineralization.
  These pathways all represent  a general biochemical potential. Whether or not a
specific compound is actually metabolized to nonchlorinated products depends on
many factors. The compounds  which are most readily metabolized are the lower
chlorinated forms. However, some compounds with only one chlorine, for instance
2-chloroaniline, are not readily metabolized. Thus both the position and the number
of substituents are important in  determining the biodegradability of a molecule. It is
not only the number of chlorines but also the general form of the molecule itself
which  governs biodegradability. Compounds such as chlorodioxins and DDT are
not shown on these figures because they appear to be highly resistant to substantive
microbial attack. The  exact physicochemical features of any given molecule  that
govern its biodegradability remain to be elucidated.
  Even chlorinated molecules for which complete pathways  of mineralization have
been developed are not metabolized in all systems. Although a few compounds are
amenable to anaerobic biodegradation, most require the availability of molecular
oxygen for ring cleavage. Other environmental parameters  which may place

                                   171

-------
          -OCH3

CHLOROANISOLE
                              Cln
                           CHLOROPHENOL-*
                                                                 X3H

                                                                 'OH
                                                          Cln
                                                      CHLOROCATECHOL
                                                        (See Figure 55)
                                                          OCH2COOH
           CI-(O)-OCH2COOH
                   "Cl         CI-/O)-OCH2COO|-f

              2,4,5-T                 CH3       CHLOROPHENOXYACETIC ACID
                                 MCPA

Figure 62,  Chlorinated aromatic compounds metabolized to chlorophenols. This figure
presents possible pathways extrapolated from various studies. In actual environmental
  systems a given transformation may be  inhibited by a number of factors. Terminal
 compounds shown may be recalcitrant or insufficient research may exist on which to
              base a conclusion. Refer to text for further discussion.
                                  172

-------
  CI-(O)-NHCOCH2COOH ~*— CI-(O)-NCHN2

CHLOROMETHYL-             CHLORDIMEFORM
 MALONIC ACID
    UREA
  HERBICIDES
                                                                 ,,OH

                                                                 SOH
                                                            Cl
  Cl..
 ACYLANILIDE
  HERBICIDES
                          CHLOROHYDROXY-
                               ANILINE
                                                       CHLOROCATECHOL
                                                         (See Figure 55)
                                                              NHCOCH,
                         CHLORONITROBENZENE
                          -OCH3

                  Cln
                CHLOROANISOLE
                                                 CHLOROACETANILIDE
                                                               NHCHO


                                                 CHLOROFORMYLANILIDE


                                                     rr\
                                                          •N=
                                                    , s—v
                                                  Cln
                                                  CHLOROAZOBENZENE

                                                    SCH3
                                        CHLOROTHIOANISOLE
Figure 63.  Chlorinated aromatic compounds metabolized to chloroanilines. This figure
presents possible pathways extrapolated from various studies. In actual environmental
  systems a given transformation may be inhibited by a number of factors. Terminal
 compounds shown may be recalcitrant or insufficient research may exist on which to
              base a conclusion. Refer to text for further discussion.
                                  173

-------
g s  a c
3 ~  <» 2
     CO
S-51!.
° I §
  33
S 2. i
?. o  2
  o  S
ic
o o
»fl
     22
<5: S
C CD
CD 5)
   C?
  CT '
 ! » i
 S 3 CD
 10 Q. 3
 CD CO 
 CO CO Q,
 S O CO.
 3 S <
 O. 3 5
 S 3 |
 Q 5 |
     .  m
|  I
3  CO
Is
°l
«  3;
(0  O
    "S
            c o
            COOH
                              CH3
                              CH2
                              COOH
             PYRUVIC ACID PROPIONIC ACID
 COOH
 CH2
 CH2
 COOH

SUCCINIC
                      T
                                      ACID
                                              CH3
                                              COOH
                                            ACETIC ACID
                                                                     CH4 + C02
                          CHO
                CHLOROHYDROXY-
                MUCONIC SEMIALDEHYDE
                                      ^-KETOADIPIC ACID   PROTOCATECHUIC ACID
                                                                               COOH
                                       CHLOROMUCONIC
                                           ACID
                              Cl'n
                           CHLOROCATECHOL
                  HYDROXYBENZOIC   BENZOIC
                       ACID  V  > ACID
                               ^anaerobic
                                 ,COOH


                           Cln"
                       CHLOROBENZOIC ACID
                                                                                    CHLOROHYDROXY-
                                                                                      BENZOIC ACID
      0
      'COOH
      COOH
MALEYLPYRUVIC
    ACID
                                       \
                  CHLOROPHENOL
                   (See Fig 53)
                                         CHLOROANILINE
                                           (See Fig 54)
                                                                c-COOH
                                                                                  Cln    *Cln
                                                                              CHLORONAPHTHALENE
                                                                                                             COOH
                                                                                                             OCH3
                                                           CHLOROBENZOYL
                                                            FORMIC ACID

-------
restrictions on biological activity include  pH, temperature,  and moisture. Upon
exposure to the environment, the chemical state of the substrate may be altered to a
form resistant to microbial attack.
  The chemical itself may be degradable but the system may lack other nutrients
necessary for microbial activity.  Alternatively, other chemicals present may be
preferred substrates, preventing metabolism of the substrate of interest. Another
compound may also act to repress the activity of enzymes required for substrate
metabolism. Accumulation of toxic metabolites may also repress further metabolic
activity.
  Ample evidence exists that some chemicals require the activities of several different
groups of microorganisms for complete mineralization. Such consortia may not be
found in the system containing the substrate.  The  interactions of these  micro-
organisms pose  additional constraints  regarding production and utilization  of
potentially toxic metabolites as well as competition for nutrients and growth factors.
  Finally, the dynamics of  pollutant  appearance  in the system  is of critical
importance. Most microorganisms require a period of acclimation to the substrate
before metabolism  occurs.  During this period, the  substrate level must be high
enough to promote acclimation without being toxic or inhibitory. Prior exposure to
the compound helps to shorten the acclimation period. Such exposure to other
pollutants may also predispose the microbial  population to adaptation  to  the
substrate  of interest. Or such  acclimation may result  in destruction  of  micro-
organisms capable of substrate utilization in  favor of a population adapted  to a
different substrate, or in virtually complete destruction of the microbial flora.
  This review of microbiological decomposition of chlorinated aromatic compounds
indicates that while certain metabolic pathway generalizations exist, as reflected in
Figures 62, 63, and 64, biodegradation of the compounds shown is dependent on
many other variables and cannot  be assumed in any given biotic system. Data on
many of these variables are needed to  allow prediction of the metabolic  fate of
chlorinated aromatic compounds.
                                     175

-------
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   methylnaphthalene, salicylate, and benzoate by Pseudomonas PG: Regulation of tan-
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Williams, P.P. 1977. Metabolism of synthetic organic pesticides by anaerobic microorga-
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                                    232

-------
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                                    233

-------
               APPENDIX

ILLUSTRATED LIST OF COMPOUNDS
                              11.





2.





CH3
HC=0


ACETALDEHYDE

COOH
CH2
CO
CH3

ACETOA.CETICACID
3.







4.







CH3
SCoA


ACETYL-CoA

COOH
CH3
C-COOH
CH
COOH
CH3
Hl^NH2
COOH

ALANINE
7.
COOH
' |x^f'C'
J| 1
CIX^^NNH2

AMIBEN (chloramben,
3-amino-2,5-dichlorobenzoic

NH2
^L. OH
(TyT
^Y"*
Cl
2-AMINO-5-
CHLOROPHENOL
9.

NH2
(o)

,. ,
f*^\l
^^Cl
BARBAN [(3-chlorophenyl)-
carbamic acid 4-chloro-
2-butynyl ester]
12' 0
CH2-S-C-NC 2 4
l^\^
(OJ

Cl
BENTHIOCARB (S-4-
ackf) chlorobenzyl-N,N-
diethylthiolcarbamate)
13.
CHO

(O)
^^

BENZAtDEHYDE
14.


(o)

                                               ,CI
ACONITIC ACID




    NH2


  ^*CY\
 HC^N,C  xCH

       H


  ADENINE
                 ANILINE
                                   BENZENE
            10.
                              15.
               ANTHRACENE
         ,H


         ^H



BENZENE 1,2-OXIDE
                      234

-------
16.
                        21.
              SO3H


            (o)

       BENZENESULFONIC
            ACID
                                                          COOH
17.
       BIPHENYL


22.


 CI-(O\-NHCON
                                              26.
                                               4-CARBOXYMETHYLENEBUT-
                                               2-EN-4-OLIDE
                                              27.
                                                         0
                                                         n
                                        CH-CH3

                                        C=CH
      nc  ~™  ..A      BUTURON[3-(4-chlorophenyl)-  4-CARBOXYMETHYLENE-2-
  7,8-BENZOCOUMARIN   , .isobutynyM.me,hylurea]    METHYL-A-BUTENOLIDE
18.
                        23.
                                              28.
19.
COOH
[0]
BENZOIC
ACID
CH2OH
(0)
BENZYL
ALCOHOL

©c>COOH
2-CARBOXYBENZO-
PYRILIUM
24.
COOH
Jj COOH
7-CARBOXY-4-CHLORO-
2-KETO-HEPT-3.5-
COOH
1
r^
1 COOH
COOH
3-CARBOXY-
cis.cis-MUCONIC
ACID
29.
°*C
3-CARBOXYMUCONOL
20.
                        25.
       -OCH3 CI
                                                 30.
                                     COOH
                             CI
                                 .0-9=0
                                                          -07.0
                                                   HOOC  ^^  XH
                            7-CARBOXY-4-CHLORO-
BIFENOX [methyl 5-(2,4-dichloro-  2-KETO-HEPT-
phenoxy}-2-nitrobenzoate]        4,7-LACTONE         4-CARBOXYMUCONOLACTONE
                                235

-------
   31.
                           36.
                                                   41.
                 _OH
            :-c<
                  OH

CHLORFENPROP METHYL
[methyl 2-chloro-3-
(4-chlorophenyl) propionate]
                                                             COOC2H5
                                                     CHLOROBENZILATE
                                                     (ethyl 4,4'-dichlorobenzilate)
                                                   42.
CFNP (2,4-dichloro-6-fluorophenyl-
4'-mtrophenyl ether)
                           38.
                                                              COOH
                                                              O
     CHLORFENVINPHOS           3-CHLORO-
     2-chloro-1 -(2,4-dichlorophenyl)-  BENZOIC
     vinyl diethyl phosphate]         ACID
                         43.
              0                     COOH
            HNCCH3
                                  fol
CHLOMETHOXYNIL (2,4-dichloro-           fa
phenyl-3'-methoxv-4'-nitroPhenyl 4.CHLOROACETANILIDE   4-CHLOROBENZOIC ACID
ether)
   34.                       39'                     44'
 CHLORDIMEFORM[N-(4-chloro-
 o-tolyl)-N',N'-dimethylformamidine]  4-CHLOROANILINE
   35.
                            40.
              CH2COOH
             Cl

           Col
  CHLORFENAC (fenac,
  2,3,6-trichlorophenylacetic acid)   CHLOROBENZENE
                             4-CHLOROBIPHENYL
                                                   45.
                                                                 COOH
                                                          Cl
                         2-CHLORO-4-CARBOXY-
                         METHYLENE-BUT-2-ENOLIDE
                                     236

-------
  46.
                         51.
                                                56.
                       5-CHLOR°-3.5-CYCLOHEXADIENE 4-CHLORO-2-HYDROXY-
                       1,2-DIOL-l-CARBOXYLIC ACID    ACETANILIDE
                         52.
                                                57.
  48.
                         3-CHLORO-1,2-DIHYDROXY-  (4-CHLORO-5-HYDROXY-
                         CYCLOHEXA-3.5-DIENE      2-METHYLPHENOXY)-
                                                  ACETIC ACID
                                                       H02C
                                                      OH
    5-CHLORO-o-CRESOL  HEXA-4,6-DIENE
  49.                     54.
             COOH
                                                           Cl
                        1 -CHLORO-2.3-DIHYDROXY-
                        4-(2,4-DICHLOROPHENYL)-   4-CHLORO-2-HYDROXY-
                                                MUCONIC SEMIALDEHYDE
                                                59.
                                       ,OCH,
                                                        Cl
                                                 ci-(O
                                                             oc-
                                                                    Cl
3-CHLORO-3.5-CYCLOHEXADIENE
                                               3-CHLORO-2-HYDROXY-6-(2,
                                               4-DICHLOROPHENYL) HEXA-2,
o-v^i ii_wnw-o,u-u i v^i_wnc/vrtuici>j[i                    .
1,2-DIOL-l -CARBOXYLIC ACID   4-CHLOROGUAIACOL 4-
                                                         Ann
                                                         ACID
  50.
                          55.
                                                60.
 4-CHLORO-3.5-CYCLOHEXADIENE
 1,2-DIOL-l -CARBOXYLIC ACID
                          CHLOROHYDROQUINONE
                                                 4-CHLORO-2-HYDROXY-
                                                 PHENOXYACETIC ACID
                                  237

-------
   61.
                          66.
            COOH
              COOH
                                    71.

                          ^0(CH2)3COOH
                                              COOH
                                               COOH
                                                         Cl
                             MCPB [4-(4-CHI_ORO-
                             2-METHYLPHENOXY)
  2-CHLORO-4-KETOADIPIC ACID  BUTYRIC ACID]      3-CHLOROMUCONIC ACID
   62.
                         67.
        CI
-------
76.   ci
                CI
                        81.
                                              86.
       Co) 
-------
 91.
                             96.
                                                         101.
                                                                     COOH
    DDA
    [2,2-bis(p-CHLOROPHENYL)
    ACETIC ACID]
         H-C-H

 DDNU
 [unsym-bis(pp-CHLOROPHENYL|
 ETHYLENE]
97.
 92.
                                                           DICAMBA (3.6-DICHLORO-
                                                           o-ANISIC ACID)
         Cl-c-ci
            H
ODD
[1,1-DICHLORO-2,2-bis
(£-CHLOROPHENYL)ETHANE)

93.
            9
         H-C-OH
            H
  DDOH
  [2,2-bis(£-CHLOROPHENYD-
  ETHANOL]
                             98.
         CI-C-CI
   DDE
   [2,2-bis(p-CHLOROPHENYL)-
   1,1-DICHLOROETHYLENE]
94.
                                                        102.
                                                               DICHLOBENIL
                                                        (2,6-DICHLOROBENZONITRILE)
                                                        103.
DDMS
[1,1 -bis(p_-CHLOROPHENYL-
2-CHLOROETHANE)]

95.
                             DDT
                             [1,1,1 -TRICHLORO-2,2-bis-
                             |£-CHLOROPHENYL)ETHANE]
                            99.
                                       COOH

                                       L'°"OH
                               DHB
                               (3,5-CYCLOHEXADIENE-
                               1.2-DIOL-1-
                               CARBOXYLIC ACID)
                            100.
                              DICHLOFENTHION
                              [£,£-DIETHYL-£-
                              (2.4.DICHLOROPHENYL)-
                              PHOSPHOROTHIOATE]

                           104.
                                  Cl
                                                CH3
                             DICHLORFOP METHYL
                             [|±)-METHYL 2-[4-(2,4-DI-
                             CHLOROPHENOXY)PHENOXY]-
                             PROPIONATE]

                           105.
            Y
         H-C-CI
DDMU
[1-CHLORO-2,2-bis(£-CHLORO-
PHENYDETHYLENE]
                                  DIBENZO-p-DIOXIN
                                                           1,2-DICHLOROBENZENE
                                          240

-------
106.
                       111.
                                             116.
          Cl
         [oXci

   1,3-DICHLOROBENZENE
107.
                        3,5-DICHLOROCATECHOL


                        112.
                         (2.4-DICHLORO-5-
                         HYDROXYPHENOXY)-
                         ACETIC ACID
                                             117.
                        3,5-DICHLORO-3,5-CYCLO-
   1 4-DICHLOROBENZENE  HEXAD1ENE-1.2-DIOL-1-
                        CARBOXYLIC ACID
108.
                        113.
      2,4-DiCHLORO-
      BENZENEAMINE
                                     OCH,
4,5-DICHLOROGUAIACOL

114.
                                OH
                         (2.5-DICHLORO-4-
                         HYDROXYPHENOXY)-
                         ACETIC ACID
                      118.

                             if  COOH
                           riS^^^n
                           Cl       Cl

                      DICHLOROMUCONIC ACID
 109.
2,4-DICHLOROBENZOIC ACID  DICHLOROHYDROQUINONE   2,4-DICHLORO-
                                                  1-NITROBENZENE
 110.
                        115.
                                              120.
                               H02C
                              OHC   T
       3,5-DICHLORO-
       BENZOIC ACID
   3.5-DICHLORO-2-
   HYDROXYMUCONIC
   SEMIALDEHYDE
2,4-DICHLOROPHENOL
     (2,4-DCP)
                                241

-------
                       126-
 121.
  (24DICHLORO-       DIFLUBENZURON               cis-1 ,2-DIHYDRO-1 ,2-
  PHENOXY ACETIC ACID) [1 -(4-CHLOROPHENYL)-        DIHYDROXYNAPHTHALENE
                     3-(2,6-DIFLUOROBENZOYL)UREA]
122.    OCH2CH2CH2COOH 127.
            cl
                                            132.
4-(24-D)B                  cis 1,2-DIHYDRO-1,2-       trans-1,2-DIHYDRO-
4-(2]4-DICHLOROPHENOXY)  DIHYDROXYANTHRACENE 1,2-DIHYDROXYNAPHTHALENE
BUTYRIC ACID
123.     „,             128.                   133.

                                    sOH

                                    •°H

                           cis-1,2-DIHYDRO-
                        1,2-DIHYDROXYBENZENE
(2,4-DICHLOROPHENOXY)
ETHANOL

124.
                       129.
         OCH2COOH-0-S03NA
                                    OH
                                   
-------
  136.
                        141.
                                     OH
                                             146.
             "OH             ^*^                OH

  2,3-DIHYDROXYBIPHENYL 3,4-DIHYDROXYPHENANTHRENE  GENTISIC ACID

  137.                   142. Cl          Cl     147.
                             (0)H<0)
                  CHO

                  COOH
      1,2-DIHYDROXY-   DPM
     DIBENZO-p-DIOXIN  DICHLORODIPHENYLMETHANE   GLYOXYLIC ACID
  138.
                        143.
                                             148.
                                COOH
                                6H
                                
-------
151.
                      156.
                                           161.
                                    SCOOH
       Cl
HEXACHLOROPHENE      o-HYDROXYBENZALPYRUVIC
[2,2'-METHYLENE bis-      ACID
(3,4,6-TRICHLOROPHENOL)]
152.        OH         157-                   162.
             .CH2COOH            COOH
         [OJ
           OH

   HOMOGENTISIC ACID
   0.-HYDROXYBENZOIC
   ACID
153.
                       158.
                                           163.
  HOMOPROTOCATECHUIC
  ACID
 154.
  m-HYDROXYBENZOIC
  ACID
                       159.
                                            164.
 4-HYDROXYACETANILIDE  £-HYDROXYBENZOIC ACID

155.                  160.                  165.
           NH2                    o
                         HOOC
                                     OH
                                     •H
                                HO
                          6-HYDROXY-6-
                          (4'CHLOROPHENYL)-
                          HEXANOIC ACID

                               COOH
                                   OH
      cir
 2-HYDROXY-
 CYCLOHEXANE-
 CARBOXYLIC ACID
    1-HYDROXY-
    DIBENZO-p-
    DIOXIN
                            2-HYDROXY-
                            DIBENZO-p-
                            DIOXIN
                                                HOOC
                                   -OH
                                   OH
           OH
    4-HYDROXYANILINE
2-HYDROXY-4-CARBOXY-
MUCONIC SEMIALDEHYDE
2-HYDROXY-4-CARBOXY-
MUCONIC ACID
                                244

-------
 166.
             O COOH
                       171.
                                             176.
            OH
   4-HYDROXY-1-0-
   HYDROXYPHENYL-
   2-OXOBUTYRIC ACID
                                    COOH
167.
    o-HYDROXY-
    /3-KETOCARBOXYLIC
    ACID
 168-      CHO
        HCOH
          COOH

    HYDROXYMALONIC
    SEMIALDEHYDE
                       3-HYDROXYMUCONIC ACID
                       172.
                                -C>L
              OH

  2-HYDROXYMUCONIC
  SEMIALDEHYDE
173.
 169.
           COOH
             COOH
                CHO
   1-HYDROXY-2-
   NAPHTHALDEHYDE
                       174.
  4-HYDROXY-2-METHYL-
  MUCONIC ACID
170.
               COOH
                           1-HYDROXY-2-
                           NAPHTHOIC ACID
               'OH
                        175.
 2-HYDROXYMUCONIC ACID
                                  HO'V°H
                        2-HYDROXY-6-OXO-6-(4'-
                        CHLOROPHENYL) HEXA-2,
                        4-DIENOIC ACID
                                             177.
 2-HYDROXY-6-OXO-6-(4'-
 CHLOROPHENYL)-4-HEXENOIC
 ACID
 178.
   2-HYDROXY-5-OXO-5-(4'-
   CHLOROPHENYL-
   PENTANOIC ACID

 179.
          |^

            =0
              COOH
              "OH
2-HYDROXY-6-OXO-6-
PHENYLHEXA-2.4-DIENOIC ACID
 180.    CH2 COOH
                       cis-4-  -HYDROXY-
                       NAPHTH-2-YL -2-OXOBUT-
                       3-ENOIC ACID
                  2-HYDROXYPENTA-2.4-DIENOIC ACID
                      181.
                             CH3  COOH
                           HQ^—^*0

                    4-HYDROXY-2-OXOVALERIC ACID
                                 245

-------
 182.
       HO
             H2cooH    188
           193.
                   COOH
                   CH
                   CH
                   COOH
3-HYDROXYPHENYLACETIC ACID
                      3-KETOADIPATE ENOL-LACTONE   MALEIC ACID
 183-
            CH2COOH
                       189.
                                            194.

OH
4-HYDROXYPHENYLACETIC
184.





fol
Ix^J
][

T COOH
XCOOH
ACID
/J-KETOADIPICACID

190.
SCoA
Oi«y^c = o
r COOH
0

MALEYL-
ACETIC ACID

195.
COOH
|| ccCH2COOH
           VCHO
 2-HYDROXY-3-PHENYLMUCONIC
 SEMIALDEHYDE
,C = 0
    OH
185.



186.



OH
(oXoH
OH
HYDROXYQUINOL

0
Ou
H^OH
4-HYDROXY-1 -TETRALONE
187. COOH
CH2
HC-COOH
-C-CH
COOH

ISOCITRIC ACID
3-KETOADIPYL CoA
191.
COOH
^vyO
^ 	 s

2-KETO-
CYCLOHEXANE-
CARBOXYLIC ACID
192.
COOH
CH2
$0
COOH
2-KETOGLUTARIC ACID
MALEYLACETOACETIC AC
196.
-COOJ^COOH
lljc^o
O


MALEYLPYRUVIC ACID
197.
COOH
HO'9H
COOH
MALIC ACID
                               246

-------
198.
            >CH2COOH
                        203.
     H3C
MCPA
(4-CHLORO-2-METHYL-
PHENOXYACETIC ACID)
                            •o_
                                    ,
                                  C = 0

                                  C = 0
                           MUCONOLACTONE
                        204.
 199.
            cci3
METHOXYCHLOR
[2,2-bis (p-methoxyphenyl)-! ,1 ,1 -
trichloroethane]                NAPHTHALENE
  200.
                        205.
         CH2
                                              208.
                                               1,2-NAPHTHOQUINONE

                                             209.
                                                         O
                                            1,4-NAPHTHOQUINONE

                                           210.
                                                   Cl
           ,OH

           VOH

   3-METHYLCATECHOL   NAPHTHALENE 1,2-OXIDE

201.                     206.

                                  OH
            COOH
                                               Cl-
                                                       •o-
                                                               -NO,
                                                NITROFEN
                                                (2,4-DICHLOROPHENYL-
                                                4'-NITROPHENYL ETHER)
         ^COOH
         Y  COO
                                              211
                                                    COOH
                                                    C=0
                                                    CH2
                                                    COOH
 3-METHYLMALEYL ACETATE      1 -NAPHTHOL

  202.                    207.
           OH

   MUCONIC ACID
                            2-NAPHTHOL
                                               OXALOACETIC ACID

                                             212.

                                                  HO    0 n
                                                     N '/  O
                                                      a"  "
                                                         -OH
                                                         OH


                                              4-OXALOCROTONIC ACID
                                247

-------
 213.
                        218.
                                               223.
  214.
             COOH


           O

                       5-OXO-5-(4'-CHLOROPHENYL)- PCP
     3-OXOADIPIC ACID  PENTANOIC ACID            (PENTACHLOROPHENOL)

                         219.                   224.
      HOOC
                            HOOC
   2-OXO-4-CARBOXYPENT-   2-OXO-4-HYDROXY-4-
   4-ENOATE               CARBOXYMUCONIC ACID   PENTACHLOROANILINE

                                               225.
  215.
                         220.
                            HOOC
                           2-OXO-4-HYDROXY-
4-OXO-4-(4'-CHLOROPHENYL)-  CARBOXYPENTANOIC
BUTANOIC ACID

 216.                     221.
                                  ACID
 PENTACHLOROANISOLE

226.
                               CH2
 6-OXO-6-(4'-CHLOROPHENYL(-  2-OXOPENT-4-ENOIC  DrMT
 2-HYDROXYHEXANOIC ACID          ACID         PENTACHLOROIMITROBENZENE
  217.
                          222.
                                               227.
                HO 0
                            PCMC
2,5-OXO-5-(4'-CHLOROPHENYL)-  (4-CHLOROPHENYL
PENTANOIC ACID              N-METHYLCARBAMATE) PENTACHLOROTHIOANISOLE
                                  248

-------
228.
     PHENANTHRENE
229.
        PHENOL

230.
         CH2COOH

       Co]


   PHENYLACETIC ACID

231.
           COOH

  PHENYLPYRUVICACID
232.
         COOH

       p   COOH
                       233.
                                              238.
                         PROTOCATECHUIC ACID
                                                       900H
                                                       
-------
243.
                       248.
                   Cl
    Cl
TETRACHLOROCYCLOHEXENE
                           2,3,6-TRICHLORO-
                           BENZOIC ACID
 244.
                        249.
                                             253.
                                                OH
                                      2,3,6-TRICHLORO-
                                      4-HYDROXYBENZOIC ACID

                                     254.
       Cl"^f"CI
            OH         2,4,4'-TRICHLOROBIPHENYL
TETRACHLOROHYDROQUINONE
                        250.                  255.
            0
            ii
        HN'CvC-CH3
                                                TRICHLOROHYDROXY-
                                                BENZOQUINONE
245.
 246.
247.
   H
THYMINE


   CH3

  (o)

TOLUENE
                          2,4,4'-TRICHLORO-2',
                         3'-DIHYDROXYBIPHENYL
                                                  2,3,5-TRICHLORO-
                                                  PHENOL
                       251.
                                             256.
                                Cl             2,4,5-T
                      3,4,5-TRICHLOROGUAIACOL  (2,4,5-TRICHLORO-
                                               PHENOXYACETIC ACID)
                       252.
                                             257.
           CH3
               OH
              ^-OH
        cis-TOLUENE
       DIHYDRODIOL
                      TRICHLOROHYDROQUINONE
                                          2-(2,4,5-TRICHLOROPHENOXYPRO-
                                          PIONIC ACID
                                          (SILVEX)
                                 250

-------
  258.
3,4,5-TRICHLOROSYRINGOL
 259.
 3,4,5-TRICHLOROVERATROLE
 260.
         HN
 0
 II
'C"CH
,N,CH
 H
          URACIL
                                       261.
                              2,3,7,8,-TETRACHLORODIBENZOFURAN
                            2,3,7,8-TETRACHLORODIBENZO-P-DIOXIN
                             251

-------
                                GLOSSARY
Acidophile:  Organism that grows very well at an acidic pH.
Active transport:   Process in which substrate entry into a cell is coupled to an energy-
yielding process.
Adenine:   Purine base unit of a nucleoside.
Allosteric enzyme:   Enzyme that contains a regulatory site.
Anaerobic respiration:  Oxidative process similar to aerobic respiration but which
utilizes nitrate or another inorganic compound as the terminal electron acceptor.
ATP (adenosine triphosphate):   Ribonucleoside 5'-triphosphate that serves as a phos-
phate-group donor in the cell's energy cycle.
Autoradiograph:   Picture formed on film resulting from exposure to radioactive parti-
cles.
Autotroph:  Organism that obtains its energy from the oxidation of organic or inorganic
compounds.
Bacteriophage:   Virus that attacks bacteria.
Barophilic:  Requiring high barometric pressures for survival.
Barotolerant:  Ability to survive at a wide range of barometric pressures.
Basophile:  Organism that grows at alkaline pH.
Binary fission:   Process of cell replication whereby a single cell divides into two.
Budding:  Process of cell replication whereby a protruberance from a cell grows into
another cell.
Capsule:   Extracellular porysaccharide which accumulates around the cell and functions
in cell attachment, defense, and protection.
Carboxylase:   Enzyme that catalyzes the ATP-dependent addition of carbon dioxide to
the  acceptor substrate.
Catabolite repression:  Inhibition of enzyme activity by binding of a control protein to
the  operator site.
Cell wall:  Outer portion of bacterial cell that confers upon the cell its shape.
Chemostat:  An apparatus used for the continuous culture of microbial populations in a
steady state in which the growth rate is maintained by the substrate dilution rate.
Chemotaxis:  Movement in response to a specific chemical.
Chemotroph:  See autotroph.
Chimera:  Molecule consisting of a replicon and another fragment of DNA.
Chromosome:   A single large molecule of DNA that contains many genes.
Codon:  A group of three adjacent nucleotides that codes for an amino acid.
                                    252

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Cometabolism:  Process by which a substrate is metabolized by a cell while the cell
utilizes another substrate as its energy source.
Competency:  State in which a cell is able to undergo transformation.
Competitive inhibitor:   Chemical which has a similar structure to an enzyme substrate
and therefore binds to the enzyme, but which does not activate the enzyme.
Complete-mix reactor:  A reactor in which, at any instant in time, the concentration of
constituents is the same  at any point in the reactor. The effluent concentration of a
constituent is therefore also the same as the concentration in the reactor.
Conjugation:  Process of DNA transfer from one bacterial cell to another by direct cell
to cell contact.
Conjugative plasmid:   Plasmid that contains genes for bacterial conjugation.
Consortia:   Mixtures of different populations.
Continuous stirred tank reactor (CSTR):  A reactor in which mechanical agitation is
used to generate a complete-mix condition where, at any instant in time, the concentration
of constituents is the same at any point  in the reactor. The effluent concentration of a
constituent is therefore also the same as the concentration in the reactor.
Copy number:  The number of copies of a single plasmid present within a single cell.
Cosmid:   Constructed vector used for cloning large fragments of DNA.
Cytochrome P-450:  Protein that serves as an electron carrier in enzymatic hydroxyla-
tion reactions and can also transfer electrons to oxygen.
Cytoplasmic membrane:  The limiting boundary of the cell protoplasm, composed of
protein and phospholipid, that functions  in substrate transport, osmotic  regulation, cell
wall synthesis, oxidative metabolism and energy production.
Cytosine:   Pyrimidine base unit of a nucleoside.
Death phase:  Period  in which reduction in population occurs due to cell death.
Decarboxylase:  Enzyme that catalyzes decarboxylation of the substrate.
Dehydrogenase:  Enzyme that mediates the loss of a hydrogen ion from a substrate with
the acceptor being other than molecular oxygen.
Denaturation:  Process of separating double-stranded DNA into single strands.
Deoxyribonuclease:  Enzyme  that catalyzes  random Cleavage of double-stranded  or
single-stranded DNA.
Deoxyribonucleic acid (DNA):  A polynucleotide consisting of deoxyribonucleotide
units that serves as the carrier of genetic information.
Deterministic:  Use of knowledge of the causes of  a process to arrive at a prediction of
its performance.
Dioxygenase:  Enzyme that catalyzes the addition of two atoms of molecular oxygen to a
molecule.
Enzyme:   Protein which both lowers the energy of activation of and directs the metabolic
pathway taken by chemical reactions in an organism.
Eukaryote:  Organism characterized by having a nucleus surrounded by a membrane.
Exogenous DNA:  DNA molecule which  is  not an integral part of the cell genome.
Exonuclease:  Enzyme that removes single nucleotides from the end of a DNA mole-
cule.

                                     253

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Exponential phase (Log phase):  Growth phase during which cells divide by binary
fission.
Extracellular:   Outside of the outermost layer of a cell (cell membrane or cell wall).
Extracellular  water:  The  water in biomass "solids" residing between cells.  Com-
pounds residing in extracellular water are not included in measures of passive or active
cellular uptake.
Facultative anaerobe:  Organism that grows in the presence or absence of air.
Feedback inhibition:  Inhibition of an allosteric enzyme early in a metabolic pathway
by a later product of the pathway.
Fermentation pathway:   Metabolic pathway in which organic compounds serve as both
the electron  donor and the electron acceptor.
Fiber wall reactor:   A reactor in which the biomass is contained within a fibrous inner
cavity. The  aqueous solution and its dissolved constituent are thus permitted to pass
through the fiber well but the biomass (fixed film and suspended) is contained within the
inner cavity. This type of reactor obviates the need for biomass separation by settling and
recycle to the system.
Fill-and-draw reactor:   A mode of operating a reactor in which reactants and products
are added or removed over discrete time intervals but the reactions are allowed to proceed
continuously. Since concentrations of constituents change cyclically during the test,
results of this type of test may only approximate true continuous and steady-state tests.
Flagellum:  A hairlike organelle attached to a cell that functions in motility.
Freundlich isotherm:  An isotherm equation relating the equilibrium partitioning of a
compound between liquid and solid compartments (or phases). The equation is of the form
      C, =  IQC,1"1
where C, and C, are concentrations of the compound in solid and liquid compartments,
respectively, and Kf and n are empirical constants.
Gene:   A DNA segment that codes for a single polypeptide chain or RNA molecule.
Genetic  recombination:  Process of combining DNA  from different  sources into a
molecule.
Genome:  The entire group of genes of a cell.
Gram negative:  Term given to bacteria that lose the primary stain (crystal violet) of the
Gram staining  procedure upon exposure to alcohol or other decolorizing agent.
Gram positive:   Term given to bacteria that retain the primary stain (crystal violet) of
the Gram staining procedure upon exposure to alcohol or other decolorizing agent.
Gram stain:  Differential staining procedure in which  crystal  violet, Gram's iodine,
decolorizing agent such as alcohol, and safranin are sequentially applied to bacterial cells.
Most bacteria can be divided into two groups based on whether they  retain or lose the
primary stain (crystal violet) during the procedure. The response of bacterial cells to this
procedure has been linked to differences in the cell wall composition.
Guanine:   Purine base unit of a nucleoside.
Halophile:  Organism that requires high salt  concentrations for growth.
Henry's  law constant:   A  constant that describes the  equilibrium  partitioning of a
compound between liquid and gas compartments (or phases) at a given condition, where
                                     254

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the concentration of the compound in the liquid compartment is sufficiently low. Many
units are possible with this constant and caution should be taken in its use.
Heterotroph:  Organism that requires an organic form of carbon for energy.
Holdfast:  Appendage of some bacteria consisting of a fine stalk which may possess
adhesive material; functions in attachment.
Hybridization:  Process of joining two nucleotides.
Hydraulic residence time (HRT):   The time that the bulk aqueous phase resides in a
continuous reactor volume. This may be calculated as the ratio of the system's hydraulic
volume and the flow rate of the aqueous feed stream, assuming no change in fluid density
during the reaction.
Hydrolase:   Enzyme that mediates the transfer of a chemical group to water.
Hydrolysis:   Cleavage of a molecule by reaction with water.
Hydrophilic:  Water-loving; refers to polar molecules that associate with water.
Hydrophobic:  Water-hating; refers to nonpolar molecules that are insoluble in water.
Hypha:   A fungus thread.
Inducer:  A molecule that induces the activity of an enzyme.
Insertion sequence:   Segment of DNA occurring on either end of a transposon.
In situ:  In its original position.
Intron:  An intervening sequence in a gene that is transcribed but excised before transla-
tion of the gene.
In vitro:  In a test tube or beaker ("in glass").
In vivo:   In a living organism.
Irreversible inhibitor:   Chemical that destroys or binds to a functional group on an
enzyme, thereby preventing its catalytic activity.
Isomerase:   Enzyme that catalyzes a change in the atomic configuration of a molecule
without a change in the number or kind of atoms.
Jet drop entrainment:   A mechanism in which small particles may be launched into a
gaseous phase based on the collapsing of bubbles at the gas-liquid surface. Small droplets
of fluid originating at the base of the bubble are formed and accelerated as the top of the
bubble breaks and  the bottom of the bubble merges with the gas-liquid surface. These
drops may be aerosolized and thus carried with the gas phase.
3-Ketoadipate pathway:  Aerobic pathway of aromatic compound dissimilation in
which the end products are the tricarboxylic acid cycle intermediates succinate and acetyl-
CoA.
Kinase:   Enzyme that catalyzes transfer of a phosphate group from ATP or other
nucleoside triphosphate to the substrate.
Lag phase:   Growth phase during which adaptation to the environment occurs and no
increase in cell number is seen.
Langmuir isotherm:   An isotherm equation relating the equilibrium partitioning of a
component between liquid and solid compartments (or phases). The equation is of the
form:
      c      abCL
            1 + b CL
                                    255

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where Cs and CL are concentrations of the compound in solid and liquid compartments,
respectively, and a and b are empirical constants. This isotherm may be derived from
considerations of mass transport onto  surfaces with limited capacity to sorb the com-
pound.
Ligase:   Enzyme that catalyzes the formation of a product resulting from the con-
densation of two different molecules, coupled with the  cleavage of a pyrophosphate
linkage in ATP.
Lipophilic:   Refers to molecules that  associate with lipids.
Lithotrophs:   Bacteria that use inorganic compounds  as substrates for respiratory
metabolism.
Log phase:   See exponential phase.
Lyase:   Enzyme that catalyzes the addition of a chemical group to the double bond of a
substrate or the removal of a chemical group to form a double bond.
Lyse:  Breaking apart of the cell wall.
Lysogeny:  Infection of a bacterial cell by a virus during which the viral genome
becomes integrated into the cell genome, is repressed, and is  replicated with the cell
genome.
Lytic cycle:   Process by which a bacteriophage infects a cell, replicates, and is released
into the environment.
Macroinvertebrates:   Group of organisms that lack a backbone; in this context refers to
species that live in the  water.
Maximum specific growth rate (fxm):   The specific growth rate of biomass on a sub-
strate that is limiting growth as defined by the Monod equation:
                                   M- =
                                           Ks + Ca
where u, is the specific growth rate, |xm is the maximum growth rate measured, Ks is the
Monod half-saturation constant (K, = C0 at a growth rate of ^J2), and Cn is the con-
centration of the growth-limiting substrate.
Mean cell residence time (MCRT) :  The mean of the cellular time distribution descrip-
tive for the reactor configuration under consideration. This equals the hydraulic residence
time in complete-mix, suspended growth reactors that have no biomass separation and
recycle. This is a design variable that may be determined in suspended growth reactor
systems that have provision for biomass separation and recycle and is determined by
biokinetic rate constants and physical design in fixed film systems.
Mesocosm:  A constructed laboratory representation  of  an environment including
atmospheric, hydrospheric, and geospheric parts with associated flora and fauna.
Mesophile:   Organism that grows best at temperatures from 15°C  to about 45°C.
Messenger RNA (mRNA):   RNA molecule that serves to carry  the genetic message
from the DNA to the ribosome.
Meta position:   Position on an aromatic molecule separated from the point of reference
by one carbon position.
Methylase:   Enzyme that adds a methyl group to particular nucleotides.
                                    256

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Microaerophile:   Organism that has a narrow range of tolerance for its gaseous environ-
ment  and requires either a reduced air environment or, in some  cases, an increased
proportion of carbon dioxide.
Microcosm:   A small-scale version of a mesocosm. See mesocosm.
Mineralize:  Convert a molecule to inorganic ions and molecules.
Mitochondria:  Organelles in a eukaryotic cell which are the sites of oxidative metabo-
lism.
Mixed liquor:  The mixture of the aqueous phase and suspended biomass in the aeration
reactor of a biological treatment process.
Mixed liquor suspended solids (MLSS):  A measure of biomass in suspended biolog-
ical processes where solids are separated from the mixed liquor, are dried, and weights are
determined gravimetrically. Standard methods exist for this test.
Mixed liquor volatile suspended solids (MLVSS):  A measure of biomass in suspended
biological processes where solids are separated  from the mixed liquor, dried, heated to
remove volatile organics, and weights are determined gravimetrically. Standard methods
exist for the test.
Monod half-saturation constant (KJ:  A constant defined in the Monod equation. See
maximum specific growth rate.
Monooxygenase:  Enzyme that catalyzes the addition of one atom of molecular oxygen
to a molecule.
Mutant:  Cell in which the genome has undergone mutation.
Mutase:   Enzyme that catalyzes transfer of a functional group between two positions on
the same molecule.
Mutation:  Alteration of the genetic message.
NAD  , NADH:   Nicotinamide adenine dinucleotide,  a coenzyme which functions in
oxidation-reduction reactions as hydrogen and electron carriers.
NADP   , NADPH:   Nicotinamide adenine dinucleotide phosphate. Same function as
NAD; see NAD.
Neutrophile:  Organism that grows best at a neutral pH.
NIH shift:  Migration of a hydrogen atom from one carbon to the adjacent carbon on an
aromatic molecule.
Noncompetitive inhibitor:   Chemical that binds to an enzyme in an area other than the
binding site, thereby altering and inactivating the catalytic site.
Nucleoid:  The area in a prokaryotic  cell  that contains the chromosome and is  not
bounded by a membrane.
Nucleoside:  A compound composed of a purine or pyrimidine base covalently linked to
a pentose  sugar.
Nucleotide:  A nucleoside with a phosphate group attached to one of the pentose hydro-
xyl groups.
Nucleus:  The membrane-bound organelle in a eukaryotic cell that contains the chromo-
some.
Nick:  A breakage  in one strand of a double-stranded DNA molecule.
Obligate aerobe:  Organism that grows in the presence of air and uses aerobic respira-
tion to obtain energy.

                                    257

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Obligate anaerobe:   Organism that grows only in the absence of air.
Octanol-water partition coefficient (K.J:   A constant that describes the equilibrium
partitioning of a compound between equal volumes of n-octanol  and water at a given
temperature. Partitioning between other immiscible fluids and between other compart-
ments can be mathematically related to this constant.
Operator region:   Regulatory site of a gene.
Organelle:  A discrete portion of a cell, with a specific function.
Organotrophs:  Bacteria that use organic compounds as substrates for respiratory
metabolism.
Origin  of replication:   Sequence of DNA required for  replication of  the molecule.
Ortho position:  Position on an aromatic molecule adjacent to the point of reference.
Osmotic shock:  Sudden change in the solute  concentration of the environment sur-
rounding a cell.
Oxidase:   Enzyme that catalyzes loss of a hydrogen ion with molecular oxygen as the
acceptor.
Oxidoreductase:  Enzyme that mediates alterations of the CH-OH group of a substrate
and requires NAD   or NADP   as the hydrogen acceptor.
Para position:  Position on an aromatic molecule separated from the point of reference
by two carbon positions, effectively opposite to the point of reference.
Passive transport:   Process by which substrates enter a cell by free diffusion dependent
on the difference in substrate concentration inside and outside of the  cell.
Pathogenic:   Capable of causing disease.
Periplasmic space:   Area between the cytoplasmic membrane and the cell wall.
Phosphatase:   Enzyme that mediates the hydrolytic cleavage of phosphate esters.
Phototroph:  Organism that obtains its energy from light.
Phytotoxic:  Capable of inhibiting the growth of plants  or algae.
Plasmid:   Small circular DNA molecule that is extrachromosomal and replicates auton-
omously.
Pleomorphic:   Capable of changing shape.
Polylinker:  Segment of DNA that contains closely spaced recognition sites for several
restriction endonucleases.
Polymerase:   Enzyme that adds nucleotides to the 3'-hydroxyl terminus or removes
nucleotides from the 5'-phosphate terminus of nicked DNA.
Primary degradation:  The initial alteration of a compound.
Primer:  Short section of DNA required to be attached to mRNA to initiate the activity
of reverse transcriptase.
Prokaryote:   Organism characterized by lacking a nuclear membrane.
Protoplasm:  The cell within the cell wall.
Protoplast:   A viable cell that lacks a cell  wall.
Psychrophile:  Organism that grows best at temperatures  below 20°C.
Recycling fermentor:  An apparatus for the continuous culture  of microorganisms in a
steady state whereby the cells are returned to the culture vessel while medium and waste
materials are removed.

                                    258

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Regulatory sequence:   A DNA segment involved in regulating a gene.
Regulatory site:  Area on an enzyme reversibly occupied by a noncompetitive inhibitor.
Relaxed plasmid:   Plasmid which is present in a cell as multiple copies.
Replicon:  DNA molecule that contains an origin of replication.
Represser protein:  Protein that binds to the operator region of a gene and blocks its
transcription.
Respiration pathway:   Metabolic pathway in which oxygen or other inorganic com-
pound or ion serves as the terminal electron acceptor.
Resting cell:   A viable cell which is not actively growing or dividing.
Restriction endonuclease:   Enzyme that recognizes and cleaves specific sequences of
nucleotides within double-stranded DNA.
Reverse transcriptase (RNA-dependent DNA polymerase):   Enzyme that catalyzes
the formation of double-stranded DNA from the information on mRNA.
Reversible inhibitor:  Chemical that  binds to an enzyme but which may be removed
with resulting activation of the enzyme; see competitive inhibitor, noncompetitive inhibi-
tor.
Ribonucleic acid (RNA):  A polynucleotide composed of ribonucleotide units.
Ribosomal RNA (rRNA):   RNA molecule attached to the ribosome that serves as a
framework for the binding of the polypeptide subunits of a protein.
Ribosome:  Site of protein biosynthesis.
RNA-dependent DNA polymerase:   See reverse transcriptase.
RNA polymerase:  Enzyme that catalyzes the formation of RNA from the information
on DNA or RNA.
Semi-continuous reactors:  Reactors that are operated by adding or removing reactant
or products over discrete time intervals, but where the reactions  are allowed to proceed
continuously. See fill and draw reactor.
Sequential induction:   Control of a long metabolic pathway such that sections of the
pathway are under separate regulatory control and each section is induced by the product
of a  prior section.
Sludge  volume index (SVI):   A measure of settleability of suspended biomass that is
based on the volume of solids  settled from a mixed liquor over a  given time interval.
Standard methods exist for this test.
Stationary phase:   Growth phase during which no net change in cell numbers is seen;
number of cells generated and dying is equivalent.
Steady  state:   The condition  where properties of a system of any given point in the
system are the same over time.
Steric hindrance:  The inability of atoms or groups on a molecule to  rotate freely
because of mutual repelling due to van der Waals forces.
Sticky end:   Linear double-stranded DNA in which one strand extends beyond the other.
Stringent plasmid:  Plasmid  which is present in a cell in one or,  at the most, three
copies.
Structural gene:  A gene  that codes for a protein.
                                    259

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Suspended biomass:  The state of biomass growth where sufficient mechanical energy
is introduced by the cells or from external sources to favor free suspension of cells of floes
of biomass in the mixed liquor or broth and to avoid the formation of a fixed biofilm on
reactor surfaces.
Synthetase:  Enzyme that mediates condensation of two separate molecules coupled
with cleavage of ATP.
T4 DNA ligase:   Enzyme that links together complementary fragments of double-
stranded DNA.
T4 RNA ligase:  Enzyme that links together complementary fragments of single-
stranded DNA or RNA.
Taxonomy:  The science of arranging organisms into logical groups describing in detail
the basic taxonomic unit, the  species.
Terminal deoxynucleotidyl transferase:   Enzyme that adds deoxynucleotides to the
3'-hydroxyl end of DNA.
Thermophile:  Organism that grows at temperatures above 50°C.
Thiokinase:   Enzyme that catalyzes the ATP-dependent formation of thiol esters.
Thylakoid:  Internal membrane structure in cyanobacteria that contains the photo-
synthetic apparatus.
Thymine:   Pyrimidine base unit of a deoxyribonucleoside.
Transcription:  Process of converting information coded by DNA into RNA.
Transduction:   Bacteriophage-mediated transfer of genetic material into a cell.
Transferase:  Enzyme that catalyzes the transfer of an intact group of atoms from a
donor to an acceptor molecule.
Transfer RNA (tRNA):  RNA molecule that serves to bring a specific amino acid into
proximity with the developing polypeptide.
Transformation:   Process of transfer of exogenous DNA into a cell.
Translation:  Process of protein biosynthesis according to the code carried by the
mRNA.
Transposon:  A segment of DNA that can be moved from one area on a chromosome to
another.
Tricarboxylic acid cycle:  Respiration  pathway utilized by aerobic organisms.
Unsteady state:   The condition where properties of a system at any given point in the
system are changing over time.
Uracil:  Pyrimidine base unit of a ribonucleoside.
Vector:  A replicon to which a fragment of DNA  may be attached so that the fragment
may be replicated.
Vesicle:  Cavity filled with liquid  or gas.
Viable:  Capable of growing.
Yield coefficient (Y):  The  ratio  of the change in biomass concentration, X, and the
change  in substrate  concentration, Ca, over an interval of time. This is a measure of
biomass production for unit substrate removal.
                                   260

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                             CITATION  INDEX
Ref.    Page
  1     69, 130
  2     45
  3     99
  4     6
  5     64, 66
  6     143
  7     142, 143
 g     91
  9     51
10     136
II     123
12     121
13     99
14     47, 48
15     47, 48
16     48
17     135
18     135
19     81,86
20     138
21     138
22     120, 125
23     120, 126
24     120
25     120
26     129, 135
26a    130, 135
27     57, 59
28     86
29     86
30     120, 125
31     99, 100
32     99, 100
33     162, 164
34     126
36     94
37     154, 156,  163
38     162
39     19
40     99, 101, 104
41     69
42     64
43     117
44     99, 100, 101
45     100, 104
46     99, 100, 102-104
47     118
48     120
49     118, 119
                                            261
Ref.
50
51
5la
52
53
54
55
56
57
58
59
60
61
62
63
64
65
66
66a
67
68
68a
69
70
71
71a
72
73
74
75
76
76a
77
78
78a
79
80
82
83
84
84a
85
86
87
88
89
90
91
92
Page
125
125
100
86, 135
93
93
138
117
118, 136
120, 126
144
99, 106
167, 168
87
48, 51
47
112
57
18, 60
64
148
136
144
52, 61
52,61
48, 133
51
64
47
67
64, 65. 67
57
148, 150
67
52, 55. 58
138
83, 142
47. 50
41
47, 57. 61, 62
62
104
71
125
86
94. 96
94
129, 136
49
Ref.
92a
93
94
95
96
97
98
99
100
101
102
103
104
106
107
108
109
110
III
112
113
114
115
116
117
118
119
120
121
122
123
124
125
126
127
128
129
130
131
132
133
134
135
136
I36a
138
139
140
141
Page
1
18, 64
83, 84
57. 60
74, 76
94
96
88
91
88, 91, 93
57
57, 59, 121
57
51, 55, 61
6, 79
86
97
113
99, 113
125
77
77
144
64
69, 71
69, 71. 88
71, 73
122
144
96
19
47, 50, 51
50, 51
99, 100
167, 169
97
154. 155
96
83. 121
83, 121
122
117
143
143
143
138
91
154. 155
48. 51

-------
Kef.
142
143
144
145
146
147
148
149
150
151
152
153
154
155
156
157
158
159
160
161
162
163
164
165
166
167
168
169
170
171
17la
172
173
174
175
176
177
177a
178
179
180
181
182
183
184
I84a
185
ise'
I86a
187
188
189
189a
190
191
192
193
194
195
196
197
Page
99, 100, 102
100, 106, 107
19
51-54, 56, 61
100, 106
64
109
109
109
51,61
99, 100
77
104
118. 119
143
142
142
143
140
140
129
129, 130, 133
130, 133
130
134
130, 133
130, 133
100. 104. 105
100. 104
100. 104
100
91
121
115, 122
47,61
47,48
47, 61
47
47. 48
48. 49
57
47
47. 81
48. 51. 52, 61
144
67. 164, 165
140
123
115
71
64
86
78, 144
143
47. 50, 51
79
77, 91
71
69, 71, 72
48, 51
149
Kef.
198
199
200
201
202
203
204
205
206
207
208
209
210
211
212
213
214
214a
215
216
217
218
219
220
220a
221
222
223
224
225
226
227
228
229
230
231
232
233
234
235
236
237
238
239
240
241
242
243
244
245
246
247
248
249
250
251
252
253
254
255
256
Page
100
51


156, 157
140
79,
121
83
47
78,

143



79
100, 104
69,
70
88,90
69
69
77
125




146. 148
1
129
69
79,
93
120
126
129
74,
118
117
51,
100
51
51
138
138
140
74
140
135
79,
51
156
91
19
115
123
123
118
118
122
115
149



94, 97




78


53,61
. 104






, 143

142

, 158


, 122
, 125. 127

, 119


, 120

140
121
104
167
167
126
94
47,
61
61


, 168
, 168


49


262
Kef.
257
258
259
260
261
262
263
264
265
265a
266
266a
267
268
269
270
271
272
273
274
275
276
277
278
279
280
281
282
283
284
285
286
287
288
289
290
291
292
293
'294
295
296
297
298
299
300
301
302
303
304
305
306
307
308
309
310
311
312
313
314
3l4a
  Page
74
113
146,  147
146,  148
69,92
88
138
113
135
143
79, 94, 97
62
51, 61, 79
140
123
123
118,  120
157,  160, 161
18
64
113
7, 15, 17, 26
97
144
109
157,  158, 161
129
51,94,97
99, 100, 106, 109
100,  101, 106
99, 100, 101
99, 100
48, 51, 52. 61
96
19, 120
99, 106
99, 106
99, 106
120
26. 29
81, 86
113
111
130,  131
I
142
146,  148, 149
140,  142
165
111
115,  123
121
19
138
140
117
138,  140
143
143
83, 86
86

-------
Ref.
315
316
317
318
319
320
321
322
323
324
325
326
327
328
329
330
331
332
333
334
334a
335
336
337
338
339
340
341
342
343
344
345
346
347
348
349
350
351
352
353
354
355
356
357
358
359
360
361
362
363
364
365
366
367
368
369
370
371
372
373
Page
154
45
136
122
87
79, 94, 97
71
83, 85
57
144
88, 89, 94
48, 51
64
126
126
57
47
47
91, 130
26,36
69
19
19
19
19
19
19, 59, 60
19
19
99
144
19
144
144
19
140, 142
142
148
148, 149
61
94, 97
140
146
47
148
64
146
117
69
71
74
71
81, 82, 83
71
47, 50
94, 95. 96
87
86
113
51, 61
Ref.
374
375
376
377
378
379
380
381
382
382a
383
384
385
386
386a
387
388
389
390
391
392
393
394
395
396
397
398
399
399a
400
401
402
403
404
405
405a
406
407
408
409
410
411
41 la
412
413
414

415
416
417
418
419
420
421
422
423
424
425
426
427
Page
51, 61
100, 101
99, 104
99, 104, 111
126. 127
96
74
118
57, 59
59
57, 59
118
152-154, 163
71, 73
73
146
91
120
77
166, 167
125
99
129, 135
64
19
111
78
77
78
111
77
109
122
61, 64, 67, 68
113
100
47, 74, 81
43
18, 19, 57, 59, 60
19
47
94, 97
100
99, 100, 104
104
69, 100, 101. 104,
106, 112
138
166, 167, 168
39, 78, 93
78
117
120
47
94
94,95
94
94
130, 133
130
                          Ref.
                          428
                          429
                          430
                          431
                          432
                          433
                          434
                          435
                          436
                          437
                          438
                          439
                          440
                          441
                          442
                          442a
                          443
                          444
                          445
                          446
                          447
                          448
                          449
                          450
                          451
                          45 la
                          452
                          453
                          454
                          455
                          456
                          457
                          458
                          459
                          460
                          461
                          462
                          463
                          464
                          465
                          466
                          465a
                          467
                          468
                          468a
                          469
                          470
                          471
                          47 la
                          472
                          473
                          474
                          475
                          477
                          478
                          479
                          480
                          481
                          482
                          483
   Page
83,85
144
47
47
109
99, 102
99, 100, 101, 103
122
111, 112
163
129, 156, 157
136
135, 136
96
118
86
109, 111
144
99
86
91
91
51
47
99, 100
81
122
122
135
123, 127
125
125
149
94, 97
114
109
109
140, 141
75, 138, 140
140. 141
94
88
47, 61, 63
83
59
112
109, 112
123
49
121
125
129, 133
47
19
19
19
19
126
112
115, 118
263

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 Kef.
484
485
486
487
488
489
   Page
IIS,  116
144
74
115,  117
115,  125
144
                                       264

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                               ORGANISM INDEX
Achromobacter, 47, 69, 88, 99, 104,106, 123, 130
Acinelobacler, 69, 130,  133
  A. calcoacelicus, 69
Aeromonas, 61
Alcaligenes, 47,  71, 88,  117, 130, 133
  A. eutrophus, 47, 71, 88
Anabaena, 125
  A. cylindrica, 125
  A. variabilis, 125
Arthrobacter, 69, 74, 83, 88, 94, 97-101,
  104, 106,  111, 113, 120, 123,  155,  156
  A. globiforms, 74
Aspergillus, 64, 77, 91, 93, 109, 118,  121, 123,
  125, 143
  A.flavus, 143
  A. niger, 64, 77, 109,  123, 143
  A. sydowi, 93
  A. ustus,  123, 125
  A. versicolor,  123, 125
Azolohacter, 47, 57, 69
Bacillus, 10, 47,  57, 74,  109, 111, 117, 120, 122,
  126, 130, 140,  148
  B. brevis, 74,  130
  B. cereus, 109, 140
  B.firmus, 117
  B. megaterium, 140,  148
  B. subtilis, 36
Bacleriodes, 138
Beijerinckia, 51,61, 146, 148, 149
Brevibacterium,  69, 104, 113
Cephaloascus, 96
  C.fragrans, 96
  C. />ro, 96
Chlamydomonas, 74
Closlridium, 10, 138
  C. perfringens, 138
Coniophora, 96
Corynebacterium, 101,  104, 111, 123
Cunninghamella, 64, 122, 135, 148
  C. echinulala, 122, 135
  C. efetfa/M, 64, 148
Enterobacter, 126,  138
  £. aerogenes,  138, 140, 144
Escherichia, 126, 138
  £. fo«, 23, 36, 118,  138, 140
Flavobacterium, 101, 104, 106,  113
  /•". aquatile, 104
  F. peregrinum, 99, 101, 104
Frateuria, 51
Fusarium, 83,  111, 118,  120, 123, 125, 143, 144
  F. culmorum,  111
  F. oxysporum, 83, 118, 123, 125, 143
Geolrichum, 118, 125
  G. candidum,  118, 125
Gloeocapsa, 125
  G. alpicola,  125
Graphium, 96
Hendersonula, 109
  //. toruloidea, 109
Hydrogenomonas,  143, 144
Klebsiella,  138
  AC. pneumoniae,  138
Laciobacillus,  138
A/uror, 83, 111,  142
  A/, allernans,  142, 143
  AY. javanicus,  83
Myco.bacterium, 83, 84,  123
Mycoplana, 99
Neurospora, 36,  64, 83
  M craMa, 36,  64, 83
Nocardia, 51, 88, 109, 123,  135, 142
  M coeliaca,  109
  M erylhropolis,  142
  JV. opafa, 109
/Voj/oc, 125
  M enlophytum,  125
  M muscorum, 125
Oscillaloria, 57,  126
Paecilomyces,  91
Paracoccus, 117
Penicillium, 64, 91, 93, 96, 111, 120, 122,  123,
 125, 143
  P.jenseni, 122
  /*. megasporum, 111
  /". noiaium, 143
  /*. piscarium,  125
Phytophthora, 109
  /". megasperma,  109
Polystictus, 64
  /". versicolor, 64
Proteus,  138
  /". vulgaris,  138
Pseudomonas, 47,  51, 69, 71, 74, 80, 81, 88, 91,
 94,97,99, 101, 104, 111, 113, 115, 120-123, 125,
 126, 130, 135, 138, 140, 146, 148, 158
                                               265

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  P. acidovorans, 47, 61
  P. aeruginosa, 47,  69, 140
  P. cepacia, 91, 104, 106,  118
  P. multivorans, 51, 117
  P. nigulosum, 123, 125
  P. piscarium, 125
  P.pulida,41, 51, 57,61,71,74,80,81,83,88,
    109, 115, 140
  P. teslosteroni, 47, 61,  148
Pullularia,  125
  P.pullulans,  125
Rhizoctonia, 83
  R. solani, 83
Rhizopus, 125,  135,  143
  R. arrhizus, 143
  R.japonicus, 135
Rhodococcus, 51, 117
Rhodopseudomonas, 47
  fl. paluslris, 47, 67
Rhodosporidium, 83
Rhodotorula, 64, 91, 120,  143
  Rhodotorula glutinis, 91
Saccharomyces, 36, 83, 142
  S. cerevisiae, 36
  5. pasiorianus, 83
Sclerotium, 83
   S. TO//JH, 83
Scopulariopsis, 91, 93
   5. brevicaulis, 93
Serratia, 138
   5. marcescens,  138
Stachybolrys, 109
   5. fl(ra, 109
Streptococcus, 138
Srrcpfo/ncra, 47, 83, 118, 123, 142
   S. aureofaciem, 83, 142
   S. o/6uj, 142
   5. anlibioticus, 142
   5. viridochromogenes, 142
Tolypoihrix, 125
   r. renuu, 125
Trichoderma, 96,  123, 142, 143
   7". virgalum, 96
   r. wride, 96, 123,  125, 142,  143
Tricchosporon, 64
   7". cutaneum, 64
Verlicillium,  111
Zygorhynchus, 111
  Z. moelleri, 111
                                             266

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                                SUBJECT  INDEX
abiotic, 1, 5, 38, 39, 152, 168, 169
abiotic degradation, 1, 39
acetanilide, 64
acetoacctic acid, 61
acylanilides, 115
algal, 74, 80
algal metabolism of chlorobenzoates, 74
4-amino-3,5-dichlorobenzoic acid, 78
aminophenol, 51
anaerobic, 17, 39, 47, 67, 78, 79, 87, 91, 97, 111,
  117, 118, 121, 138, 140, 143, 144, 158, 159, 171
anaerobic respiration, 17
aniline, 51, 64, 115, 117, 118, 120-123, 126, 165
anisole, 64, 91, 149
anthracene, 51
Aroclor,  129, 135, 136, 157-159, 161, 168
Askarels, 129
barban,  123
benzene, 45,47,51,64,79,81,86,94 113,118,135
benzoic acid, 4, 47, 61, 64, 67, 69, 71, 74, 77,78,
  118, 130, 135-137, 171
benzonitrile, 83
bifenox, 113
biphenyl, 64, 67, 81, 129, 136, 137, 148, 159
 1,1-bis (p-chlorophenyl)acetic acid (DDA),
  142, 143
catabolite repression, 19 -
catechol,  4, 18, 47, 51, 61, 64, 67, 69, 71, 81, 88,
  98, 101, 106, 117,  148, 171
chemostat, 10, 81
chlomethyoxynil, 126
chlordimeform, 126
chlorinated biphenyl, 129,  130, 136
chloroaniline, 78, 83,  115, 117,  118,  120, 123,
  128, 164, 165, 171
chloroanisoles, 88
chloroazobenzene, 120
chlorobenzene, 2, 4, 81, 86, 87, 96, 165, 167,
  168, 171
chlorobenzilate, 143
chlorobenzoic acid, 2, 69, 71, 74, 77, 78, 80, 81,
  91, 129,  130, 133, 135, 136, 144, 166, 167
chlorobiphenyl, 78, 129, 130, 135, 137, 157-159
chlorocatechol, 69, 71, 77, 79, 81, 88, 91, 101,
  106, 111, 117, 128
4-chloro-3,5-dinitrobenzoic acid, 74, 78
chlorofenprop-methyl, 113
chloroneb, 83
chloronitrobenzene, 83
chlorophenol, 2, 4, 79, 83, 86-88, 91, 93, 94, 96,
 97, 101,  106, 117, 118, 146,  167, 168, 171
chlorophenoxyacetic acid, 101, 106, 109, 111
chlorophenoxypropionic acid,  109
chlorophenylacetic acid, 113
chlorophenylcarbamic acid, 122
chloropropylate, 143
chlororesorcinol, 78
chlorosalicylate, 74
chlorotoluene, 78
chlortoluron, 122
chromosome, 19, 20, 23-26, 34
CIPC, 122, 123. 125
Clophen, 129
cometabolism,  146, 148
consortia, 2, 4, 13, 78,  79, 87, 94, 113, 121, 144,
  156, 171, 175
cyanobacteria,  2, 6-8, 16, 57, 67, 125
cyclohexanol, 51, 67
cytochrome, 67, 140
2,4-D, 99, 101, 104, 106,  109, 111-113,  115, 167
4-2,4-DB, 106,  109
DBH (dichlorobenzhydrol), 140, 143
DBF (dichlorobenzophenone), 140, 143, 144
DDA (2,2-bis(p-chlorophenyl)acetate, 140,
  142-144
DDMS (l-chloro-2,2-bis(p-chlorophenyl)-
 ethane, 140
DDMU (l-chloro-2,2-bis(p-chlorophenyl)-
 ethylene, 140, 143
DDNU (unsym-bis(p-chlorophenyl)ethylene,
  140
DDT, 2, 79, 138, 140,  142-145, 171
decachlorobiphenyl, 136
dehydrogenase, 47, 51, 57, 69, 109
dibenzo-p-dioxin, 2, 146,  148,  149
dibenzofuran,  2, 146, 151
dicamba, 77, 78
dichlobenil, 83, 86
dichloroaniline, 115, 118,  120-123,  125, 126,
  128
dichlorobenzamide, 83, 86
dichlorobenzene, 86, 165,  168, 169
dichlorobenzidine, 136
dichlorobenzoic acid, 77, 78,  135, 136,  166
dichlorobenzophenone, 140, 143
dichlorobiphenyl, 133, 135, 136,  159
dichlorocatechol, 69, 88, 99, 101, 104, 111,  115
l,4-dichloro-2,5-dimethoxybenzene, 83
dichlorodiphenyldichloroethane, 138
dichlorodiphenylmethane, 140, 144
2,4-dichloro-6-fluorophenyl 4'-nitrophenyl
 ether,  122
                                              267

-------
3,5-dichloro-N-(l,I-dimethyl-2-propynyl)-
 benzamide, 77
2,6-dichloro-4-nitroaniline, 86,  118
dichlorophenol, 86, 91, 93, 97, 99, 101, 106, 113,
 126, 162,  164, 168, 169, 187, 192, 193
(2,4-dichlorophenoxy)acetic acid, 99,  166
(2,4-dichlorophenoxy)butyric acid. 111,  112
2-(2,4-dichlorophenoxy)ethanol, 109,  111
(2,4-dichIorophenoxy)propionic acid,  111
3-(3,4-dichlorophenyl)- 1-methylurea, 123
2,4-dichlorophenyl 4'-nitrophenyl ether,  126
dicryl, 123,  125,  126, 128
diflubenzuron, 120
dihydroxybenzoic acid, 47, 51,  74
dihydroxydibenzo-p-dioxin, 146, 148
dioxin, 2, 146, 148, 149
dioxygenase, 51, 67, 117
diphenylmethane, 133, 142
diuronTllS, 121,  122
DNA (deoxyribonucleic acid), 2, 7, 9, 20-36
DPM (dichlorodiphenylmethane), 140
enzyme, 2, 7, 8, 11,  12, 14, 15, 23, 29, 32, 34, 35,
 41, 43, 44,  47, 51, 57, 61, 64, 67, 69, 74, 78, 80,
 81, 83, 88,  91,94, 98, 101,  104, 109,  115, 118,
 120, 121, 123, 125, 137, 143, 144, 146, 148, 166,
 175
ethyl 4,4'-dichlorobenzilate,  143
feedback inhibition,  15
Fenclor,  129
fermentation, 16, 158
fumarate, 61, 67
fungal metabolism, 64, 91, 109, 118,  137, 149
fungal metabolism of phenoxy,  109
gentisic acid, 47, 61, 64, 67,  113
guaiacols, 88
Henry's law constant, 156
herbicides,  3, 4, 88, 99, 104, 106, 109, 111-115,
 120-123, 125, 16,  128, 165,  171
hexachlorobenzene, 86, 87,  165, 168
homogentisic acid, 61, 138
hydraulic residence time (HRT), 154,  155, 157,
 159, 162-164, 166,  168
hydrolase, 57, 81
hydrolysis. 111, 118, 121,  123, 128
hydroxybenzoic acid, 47
isomerase, 61
isopropyl dichlorobenzilate,  143
isopropy] phenylcarbamate,  122
Kaneclor, 129, 133, 158
3-ketoadipate, 18, 64
3-ketoadipic, 64, 67, 71, 74
ligase, 30
linuron,  115, 122
maximum specific growth rate,  10, 155
MCPA, 99, 101, 104, 109, 111-113
mean cell residence time,  154, 156, 163
methane, 1, 4, 8, 39, 78, 93
methylcatechol, 57
2-methylpentanamide, 123
microcosm. 77,  152
mixed liquor suspended solids (MLSS), 157,163,
  164
mixed liquor volatile suspended solids
  (MLVSS), 162
monolinuron, 122
monoxygenase,  51, 61, 67, 133,  137
monuron,  115, 121,  122
N-(2-chloro-4-methylphenyl)-N'-dimethylurea,
  122
N-(3,4-dichlorophenyl)methacrylamide, 123
N-(l,l-dimethylpropynyl)-3,5-dichloro-
  benzamide, 126
naphthalene, 38, 51, 57, 61, 140, 146
naphthol, 57, 64, 67
N1H Shift, 47
nitrofen, 113,  128
0-phthalic acid,  61
octachlorobiphenyl,  159
octanol-water partition coefficient, 1, 156,  163
oxidase, 101, 118,  140
pentachlorophenol (PCP), 2, 79, 87, 88, 94,
  96-98, 104, 146, 154-157
penta'chlorobenzene, 86, 87
pentachlorobiphenyl, 135,  136, 157, 158
pentachloronitrobenzene, 86, 87
phenanthrene, 51,61
Phenoclor, 129
phenol, 34, 51, 86, 88, 91, 93, 96, 99,  104, 111,
  162, 163,  168
phenoxyacetic acid, 99,  112,  166
phenoxyalkyl acid herbicides, 99
phenoxyethyl esters, 99
phenyl ureas, 115, 128
phenylamide, 115
phosphatase, 32
photolysis, 113
pimelic acid, 51, 67
plasmids, 2, 9, 20,  31, 32, 33, 34, 140
polychlorinated  biphenyls (PCB), 69,  81, 129,
  130, 133, 135-137, 146, 156-159, 161,  162
polymerase, 16, 30, 36
propanil, 115, 120, 123, 125,  126
protocatechuic acid,  23, 47, 57, 61, 64, 67,
 74, 77
Pyralene, 129
pyrocatechase, 69, 71, 74, 80, 91
pyruvic acid, 57, 61, 67, 117,  171
reductive dechlorination, 78,  79, 138,  144, 171
RNA (ribonucleic acid), 16, 20-25, 30, 36
salicylaldehyde,  51, 61
salicylic acid, 47, 51, 61,  146, 171
Santotherm, 129
SD8280, 113
sewage, 3, 43, 44, 71, 74, 77,  78, 81, 86, 91, 93,
  104, 112, 130, 136,  157,  158
sludge, 44, 77, 78, 91, 93,  120, 136, 143,
  144, 154-158, 162-168
sludge volume index, 155
                                              268

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sodium 2-(2,4-dichlorophenoxy)cthyl
 sulfate, 111
solan,  123
sorption, 4,  39, 86, 128,  143, 154-157, 159,
 161-163, 165, 167-169
stripping, 4, 155, 156,  158, 159, 161-163,  165,
 167-169
structure, 6-8, 14, 25, 41. 4A, 45-, 99,  &** L30,
 165
suspended biomass, 166
swep,  123
2,4,5-T, 99,  104,  106, 111-113, 146
tetrachloroazobenzene, 118,  123,  125, 126
tetrachlorobiphenyl, 133, 135
tetrachlorodibenzo-p-dioxin  (TCDD), 146,  148,
 149,  151
tetrachlorophenol, 88. 91, 94, 96, 97, 104
toluene, 47, 64, 81, 157
transferase, 31
tricarboxylic acid cycle, 16, 57, 81
trichlorobenzene, 86, 168,  169
trichlorobenzoic acid, 69, 78
trichlorobiphenyl, 130, 133, 135,  136, 161
trichlorocarbanilide  (TCC), 164,  165
l,l,l-trichloro-2,2-bis(p-chlorophenyl)ethane
 (DDT), 2, 79, 138,  140, 142-145
trichlorophenol, 69,  86, 94, 96, 97, 99, 104,
 111, 168
2,4,6-trichlorophenyl 4'-nilrophenyl ether,  126
unsym-bis(p-chlorophenyl)ethylene, 140
urea herbicides, 115, 121, 122,  128
volatilization, 39, 86, 87, 121, 122, 128
yield coefficient,  166
                                               269
               
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