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.
-------
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.
-------
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.
-------
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
-------
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
-------
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
-------
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
-------
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
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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.
32
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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.
33
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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
36
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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.
37
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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.
38
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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.
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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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
COOH
3-CHLOROBENZOIC ACID
COOH
OH
3 CHLOROCATECHOL
COOK
OOH
COOH
2-CHLOflOMUCONIC ACID
\ •
COOH
OOH
Cl
OOH
OH
Cl
2-CHLOROCATECHOL
| COOH
3-CHLOROMUCONIC ACID
COOH
3.5-DICHLOROBENZOIC ACID
t
COOH
,OH
1H
3.5-DICHLOROCATECHOL
COOH
COOH
2.4-DICHLOROMUCONIC ACID
COOH
COOH
MUCONOLACTONE
2-CHLOROMUCONOLACTONE
COOH
MALEYLACETIC ACID
OOH
COOH
3-KETOADIPIC ACID,
[ COOH
2-CHLOHOMALEYLACETIC ACID
ODOH
Cl
2-CHLORO KETOADIPIC ACID
ACETIC ACID
2-CHLOHOSUCCINIC ACID
FUMARICACID ^*^ ^"^-Cr
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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OJ
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cu
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co
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CT
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e hydrolase.
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3
m
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conate isomet
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mes: (a) benzo
CD
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tabolism of
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cn
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3-CHLORO- 3-CHLOROCATECHOL
DHB
HOOC
3-CHLORO-cis,cis-
MUCONIC ACID
0
4-CHLORO-2-
HYDROXY-
PHENOXYACETIC
ACID
3-CHLOROBENZOIC ACID
.a
OH
5-CHLORO-DHB 4-CHLOROCATECHOL
2-CHLORO-CIS.CIS-
MUCONIC ACID
COOH
S
C=0
CH2
COOH
MALEYLACETIC
ACID
CIS-4-CARBOXYMETHYLENEBUT-
2-EN-4-OLIDE
-------
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
-------
COOH
•B S.H 2 g-
a 5 ~ " i
SfiSfS
* « o g o 4-CHLOROBENZOIC ACID
3- -* 3 o ±»
i||f|
w Jo ro « o-
COOH
OH
-<
o ^,
pi
is
<« o
CD
-t=.
8
10 3
B -» — Q.
M M 3 Q)
S m|i
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=; <3'
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Is?
15S
•< lo CD
|||
® to w
l£&
Q. ii m o
=r •£. - 2 S
° £ § ° ~
^ Q) z; * •<
(D
5- •<.
s
=•"'
DO
33s
5T 9L »'
COOH
COOH
5-CHLOROSALICYLIC
ACID
Cl
7-CARBOXY-4-
7-CARBOXY-4-CHLORO- CHLORO-
2-KETO-HEPT-3.5- 2-KETO-HEPT-3.5-
DIENOIC ACID 4,7-LACTONE
MALEYLPYRUVIC ACID
H20
COOH
r-o
CH3
PYRUVIC
ACID
COOH
, rM —
CH
COOH
FUMARIC
ACID
H2O
FUI\
COOH
FUMARYLPYRUVIC
ACID
COOH COOH
C=0 + CH
CH3 CH
COOH
PYRUVIC MALEIC
ACID ACID
O- ro i
-------
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
-------
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
-------
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
-------
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.
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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
-------
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
-------
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
-------
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
-------
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
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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
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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
^Y^ci
OH
TCHQ
PROPOSED
INTERMEDIATE
^j
J? OH
' CHQ
OH
Clvivci
T/^T — b.
i ^w'j "
spontaneous ^f^
chemical OH
^reaction 2,6-DCHQ
X 0
CIYUTOH
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
-------
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
-------
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
-------
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
-------
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
-------
f
i
> CD
f!
if
I 2.
31
a>
s
«
•O
COOH
COOH
Cl
COOH.
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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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
-------
ID
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-------
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
-------
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
-------
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-
FORMYLANILIDE
3,4-DICHLORO
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3,4-
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3,4-DICHLOROMUCONIC ACID
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3-CHLOROLEVULINIC ACID
3-CHLORO-
L 4-KETOADIPIC ACID
3-CHLOROBUTENOLIDE
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CH2
COOH
SUCCINIC
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HCCI
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2-CHLOROSUCCINIC
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ACETIC
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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
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-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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3
oi
8. S
. QJ
3 §•
3 S1
33 3
§ °
o a
(0 D
O
3
w'
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
-------
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
-------
I
T3
to
c
a\
ai
CD O
B
P
o
Q.
3'
CO
o1
i
CHLORDIMEFORM
4'-CHLORO-2'-METHYL-
MALONANILIC ACID
NO2
V
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.
128
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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
-------
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
-------
COOH
o ^-
CO
to
-------
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
-------
s
3
01
2,4,4'-TRICHLOROBIPHENYL
o> «.
•o ^
CD "!
a. Z
=? o
o 3
3 2
£ 1
S =-
g 1
CD S
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O) 2
?
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
-------
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
-------
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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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.
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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
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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
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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
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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
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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.
159
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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.
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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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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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
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
-------
Kef.
484
485
486
487
488
489
Page
IIS, 116
144
74
115, 117
115, 125
144
264
-------
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
-------
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
-------
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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