Review of Literature - INFLIBNETshodhganga.inflibnet.ac.in/bitstream/10603/13612/12/12...Review of Literature 14 to cis-1, 2 DCE (dichloroethene) and synthesizes energy via electron

Review of Literature

9

2. Review of Literature

Chlorinated organic molecules constitute the largest single group of

compounds on the list of priority pollutants compiled by the U.S Environmental

Protection Agency (U.S. EPA 1986). They are extraneously added into the

environment in large quantities as a result of their widespread use as

herbicides, insecticides, fungicides, solvents, hydraulic and heat-transfer

fluids, plasticizers, cleaning agents, fumigants, aerosol propellants, gasoline

additives, degreasers, and intermediates for chemical synthesis. The ability of

chlorinated compounds to impart toxicity, bioconcentrate, and persist and

consequently their ubiquitous distribution in the biosphere has caused public

concern over their possible effects on the quality of life (Fetzner and Lingens

1994). A list of synthetic chlorinated compounds and their use is given in

Table 2.1 (Rossberg et al., 1986; Anonymous 1993; Muller et al., 1986; Leng

1986). Some chlorinated compounds also occur naturally in the environment,

although in lower concentrations. For example, many different genera of wood

rotting fungi produce chlorinated anisyl metabolites in their natural

environments.These chloroanisyl-derivative-producing fungi are widespread in

nature. A ubiquitous production of chloroanisyl metabolites under natural

conditions was proposed by de Jong et al., (1994). More than 130 chlorine-

containing compounds have been isolated from higher plants and ferns. Many

compounds are chlorohydrins, which are isolated along with their related

epoxides (Engvild 1986).

As is true for many organic compounds, the turnover of chlorinated

molecules in the environment is largely determined by their susceptibility to

biotransformation by microorganisms (Dagley 1975). Many of the chloro-

organics that are not degraded by bacteria and fungi have the potential to

persist in the environment and express their toxicity over extended periods of

time (Hutzinger and Veerlamp 1981). Thus, identification and application of

novel organisms that use chlorinated pollutants for growth have become an

important area of research today. Further, process optimization for

biodegradation of these hazardous chemicals requires an understanding of

microorganisms involved in the degradation, their nutrient requirements, the


10

biochemistry of their mediated reactions, and why they promote these

reactions.

2.1 Utilization of chlorinated compounds by microorganisms The biological destruction of toxic and hazardous chemicals is also

based on the principles that support all ecosystems. These principles involve

the circulation, transformation, assimilation of energy and matter (Cookson

1995). Microorganisms convert complex organic compounds, via their central

metabolic routes, to CO2 or other simple organic compounds. The oxidation

yields energy and reducing equivalents that are used for conversion of a part

of the intermediates to cell mass (assimilation), enabling growth of the

organisms that carry out the degradation process (Janssen et al., 1989).

Degradation of compounds of natural origin is usually easy to achieve, and

organisms that bring about their degradation can be easily isolated from their

natural environments. However, in general, compounds having a structure

that is different from naturally occurring compounds (xenobiotics, most of

which are toxic and hazardous) are more difficult to degrade (Hutzinger and

Veerlamp 1981). Nevertheless, in the recent past, an array of microorganisms

has been identified that use xenobiotics such as chlorinated alkanes,

chlorinated halohydrins, polychlorinated biphenyls, and chlorobenzenes for

their survival.

Table 2.1: Major Chlorinated Hydrocarbons (HC) and Their Applications

Chlorinated HC Major uses Chloromethanes

Monochloromethane Production of silicones, tetramethyllead,

methylcellulose, other methylation

reactions

Dichloromethane Degreasing agent, paint remover, pressure

mediator in aerosols; extract technology

Trichloromethane Production of monochloro-difluoromethane

(for the production of tetrafluoroethene,


11

which is used for the manufacture of

Hostaflon and Teflon), extractant for

pharmaceutical products

Chloroethanes

Monochloroethane Production of tetraethyllead, production of

ethylcellulose; ethylating agent for fine

chemical production, solvent for extracting

processes

1,1- Dichloroethane Feedstock for the production of

1,1,1-trichloroethane

1, 2-Dichloroethane Production of vinyl chloride production of

chlorinated solvents such as 1,1,1-trichloro-

ethane and tri- and etrachloroethane,

synthesis of diethylenediamines

1,1,1-Trichloroethane Dry cleaning, vapor degreasing, solvent for

adhesives and metal cutting fluids; textile

processing

1,1,2-Trichloroethane Intermediate for production of 1,1,1-

trichloroethane and 1,1-dichloroethane

Chloroethenes Monochloroethene Production of polyvinyl chloride (PVC),

production of vinyl chloride chlorinated

solvents, primarily 1,1,1-trichloroethane

Trichloroethene Solvent for vapor degreasing in the metal

industry and for dry cleaning, extraction

solvent, solvents in formulations for

rubbers, elastomers and industrial paints

Tetrachloroethene Solvent for dry cleaning, metal degreasing,

textile finishing, dyeing, extraction

processes, intermediate for the production

of trichloroacetic acid and some

fluorocarbons


12

2-Chloro-1, 2-butadiene Starting monomer for polychloroprene

(Chloroprene) rubber

Chlorinated paraffins Plasticizers in PVC; flameproofing agents

in rubber textiles, plastics,H2O repellent and

not preventive agents; elastic sealing

compounds paints & varnishes;Metal

working agents (cutting oils); leather

Finishing

Chlorinated aromatic HC

Monochlorobenzene Production of nitrophenol, nitroanisole,

chloroaniline, phenylenediamine for the

manufacture of dyes, crop protection

products, pharmaceuticals and rubber

chemicals

1,2-Dichlorobenzene Production of 1,2-dichloro-4-nitrobenzene

for the production of dyes and pesticides;

production of disinfectants, room

deodorants

1,4-Dichlorobenzene Production of disinfectants, room

deodorants, moth control agent;

production of insecticides; production

of 2,5-dichloronitrobenzene for the

manufacture of dyes, production of

polyphenylenesulfide-based plastics

Chlorinated toluenes Hydrolysis of cresol, solvent for dyes;

precursors for dyes, pharmaceuticals,

pesticides, preservatives and disinfectants

Chlorophenols Preparation of agricultural chemical

herbicides etc,

Chlorophenonyalkanoic

Acids Herbicides

Side-chain chlorinated


13

aromatic HC

Chloromethylbenzene

(benzylchloride) Production of plasticizer, benzyl alcohol,

phenylacetic acid, quarternary ammonium

salts, benzyl esters, triphenylmethane

dyes, dibenzyl disulfide, benzylphenol,

benzylamines

Dichloromethylbenzene

(benzalchloride) production of benzaldehyde

Trichloromethylbenzene

(benzotrichloride) Production of benzoylchloride; Production of

pesticides; UV stabilizers and dyes

Pesticides, herbicides

and fungicides For seed treatment, for treatment of

diseases of plants,animals, and humans

2.2 Energy Metabolism

Several bacterial strains have been isolated that utilize chlorinated

compounds for synthesis of energy. Many have been shown to couple

reductive dechorination to energy metabolism (Holliger and Schumacher

1994; Holliger and Schraa 1994; Wolhfarth and Diekert 1997). Desulfomonile

tiedjei uses H2 or formate as an electron donor and 3-chlorobenzoate as a

terminal electron acceptor in a respiratory process (De Weerd et al., 1990;

Dolfing 1990; Mohn and Tiedje 1991). Chemo-osmotic coupling of reductive

dechlorination and ATP synthesis has been demonstrated in bacterium DCB-

1 (Mohn and Tiedje 1991). This organism can biosynthesize ATP by coupling

hydrogen oxidation to reduction of the C–Cl bond of 3-chlorobenzoate. Using

acetate or fumarate as electron donor, the isolate CP-1 grows via reductive

dechlorination of chlorophenol (CP) (Cole et al., 1994). Holliger et al., (1993)

have studied a highly purified enrichment culture that couples dechlorination

of tetrachloroethene (TeCE) to growth. They demonstrated that PER-K23

catalyzes transformation of PCE (perchloroethene) via TCE (trichloroethene)


14

to cis-1, 2 DCE (dichloroethene) and synthesizes energy via electron

transport phosphorylation (Holliger et al., (1993). Bradley and Chapelle (1998)

studied aerobic microbial mineralization of DCE as sole carbon substrate.

Methylobacterium, Methylophilus, and Hyphomicrobium are aerobic bacteria

capable of growth with DCM (dichloromethane) as sole source of energy and

carbon (Leisinger et al., 1993). Susanna et al., (1993) have demonstrated

DCM as sole “C”source for an acetogenic mixed culture.

Desulfitobacterium chlororespirans gains energy from the reductive

ortho-dechlorination of 3-chloro-4-hydroxy benzoate and 2, 3-di- and

polychloro-substituted phenols (Loeffler et al., 1996). Neumann et al., (1994)

unambiguously demonstrated coupling of reductive dechlorination to

respiratory growth in Desulfitobacterium multivorans.

However, utilization of chlorinated compounds is not based purely on

energy metabolism. Although traditionally it was believed that organisms must

obtain energy from an organic compound by degrading it, now it has been

shown that organisms growing at the expense of one substrate can also

transform a different substrate that is not associated with that organism’s

energy production, “C” assimilation, or any other growth process. This mode

of activity is called “cometabolism” (Cookson 1995).

2.3 Cometabolism

Cometabolism is defined as the degradation of a compound only in the

presence of another organic material that serves as the primary energy

source (McCarty 1987). A number of laboratory studies have demonstrated

that several chlorinated hydrocarbons are transformed cometabolically by

bacteria that degrade the chlorine unsubstituted aliphatic and/or aromatic

hydrocarbons (Ensley 1991; Vogel et al., 1987). Several studies on

chlorinated solvents undergoing fortuitous dechlorination by microorganisms

growing on other electron donors and acceptors have also been documented

(Semprini 1997). Nitrosomonas europea can cometabolize dichloromethane

(DCM), trichloromethane (TCM), 1,1,2-trichloroethane,1,1,1-trichloroethane,

and 1,2,3-trichloropropane while utilizing ammonia as the primary substrate

(Vannelli et al., 1991). Several bacteria capable of oxidizing toluene,methane,


15

and ammonia can cometabolize TCE, DCE, and vinyl chloride (VC) (Nelson et

al., 1986). Pseudomonas cepacia G4 is one such organism that uses toluene

and can degrade TCE cometabolically (Krumme et al., 1993). Reductive

dechlorination or reduction of TeCM (tetrachloromethane) by Escherichia coli

K12 under fumarate respiring conditions and by a denitrifying strain

Pseudomonas KC are cometabolic processes that are mediated by electron

carriers of the respiratory electron transport chain (Criddle et al., 1990; Dybas

et al., 1995). It has been observed that in some bacteria if nonhalogenated

diphenylmethane is added as a primary substrate, the chlorinated substituted

form is degraded by cometabolism (McCarty 1987). Hage et al., (2001) have

reported Pseudomonas strain DCA1 could cometabolize a broad range of

chlorinated methanes, ethanes, propanes, and ethenes using chloroacetic

acid as cosubstrate. Phenol-oxidizing microorganisms have been shown to

effectively transform cis- and trans-DCE and TCE in laboratory as well as in

situ field studies (Hopkins et al., 1993). Alcaligenes denitrificans and

Rhodococcus erythropolis can cometabolize TCE, DCE, and VC. A

Xanthobacter has been reported to degrade TCE, VC, cis- and trans-1,2-

DCE, 1,3-DCP (dichlorophenol), and 2,3-DCP cometabolically (Ensign et al.,

1968).

The phenomenon of cometabolism has been attributed to the

production of broad-specificity enzymes. Both the primary substrate and the

chlorinated compound compete for the same enzyme (McCarty 1987). It has

been reported that several oxygen-catalyzed dehalogenation reactions of

chlorinated methanes, ethanes, and ethylenes are due to multifunctional

enzymes with broad specificity or involve enzymes from aromatic degradative

pathways (Leng 1986). For a cometabolic mode, the degradation rate of the

target chlorinated compound is dependent on the electron flow from the

primary substrate.

2.4 Aerobic Degradation

During aerobic degradation of chlorinated compounds by

microorganisms, molecular oxygen serves as the electron acceptor. Several

chloroaliphatic compounds have been shown to be degraded aerobically. A


16

number of studies have demonstrated that microorganisms degrade DCE

under aerobic conditions (Bradley and Chapelle 1998; Hopkins and McCarty

1995; Bielefeldt et al., 1995; Bradley et al., 1998). Lee et al., (2000) observed

sustained degradation of TCE in a suspended growth reactor by an

Actinomycetes enrichment culture. Aerobic mineralization has been well

documented for chlorobenzenes with up to four chlorine substituents in

microcosms and by pure cultures (De Bont et al., 1986; Haigler et al., 1988;

Marinucci and Bartha 1979). Several of the chlorobenzenes (containing one,

two, three, or four chlorine substituents) could be biotransformed only under

aerobic conditions and were unstable in the absence of molecular oxygen

(Van der Meer et al., 1987). It has been reported that 4-chlorophenol (4-CP)

can be partially or completely degraded aerobically by several bacteria,

including Pseudomonas (Knackmus and Hellwig 1978), Alcaligenes (Hill et al.,

1996), Rhodococcus, Azotobacter (Wiesir et al., 1994) etc. Richard and

Michael (Lamar et al., 1990) studied degradation of pentachlorophenol (PCP)

by Phanerochaete spp., and studied its sensitivity to the compound.

Microbes play an essential role in the bioconversion and total

breakdown of pesticides. Among the microbial communities, bacteria and

fungi are the major degraders of pesticides. Yeasts, microalgae and protozoa

are less frequently encountered in the degradation process. Microbes

responsible for the degradation of various pesticides have been described in

Table 2.2. Among bacteria, Pseudomonads are considered to be the most

efficient group in bioremediation. Table 2.3 describes the bioconversion of

xenobiotics affected by Pseudomonads.

Table 2.2: Microorganisms responsible for pesticide degradation

Pesticide Microorganism Reference

Chlorophenoxy acids

2,4-D Alcaligenes

eutrophus

Don and Pemberton

(1981)


17

Alcaligenes

xylosoxidans

Gunulan and

Fournieer (1993)

Flavobacterium

spp.,. 50001

Chaudhry and

Huang(1988)

Pseudomonas

putida Lillis et al (1983)

Pseudomonas

cepacia Kilbane et al (1982)

Comamonas spp.,. Bulinski and

Nakatsu (1998)

2,4,5-T Pseudomonas

cepacia Karns et. al. (1982)

DPA Flavobacterium

spp.,.

Horvath et. al.

(1990)

Mecoprop Sphingomonas

herbicidivorans MH Zipper et al (1966)

Mecocarp Alcaligenes

denitrificans Tett et. al. (1997)

Organochlorines

Aerobacter

aerogenes Wedemeyer (1966)

Alcaligenes

eutrophus A5

Nadeau et. al.

(1994)

Agrobacterium

tumefaciens Johnson et al (1967)

Arthrobacter spp.,. Patil et. al. (1967)


18

DDT

DDT

Bacillus cereus Johnson et. al.

(1967)

Bacillus cooagulans Langlois et. al.

(1970)

Bacillus megaterium Plimmer et. al.

(1968)

Bacillus subtilis Johnson et. al.

(1967)

Clostridium

pasteurianum

Johnson et. al.

(1967)

Clostridium

michiganense Johnson et al (1967)

Enterobacter

aerogenes

Langlois et. al.

(1970)

Erwinia amylovora Johnson et. al.

(1967)

Escherichia coli Langlois et. al.

(1970)

Hydrogenomonas

spp.,.

Focht and

Alexander (1970)

Klebsiella

pneumonia Wedemeyer (1966)

Kurthia zapfii Johnson et. al.

(1967)

Micrococcus spp.,. Plimmer et. al.

(1968)

Nocardiai spp.,. Chacko et. al.

(1996)


19

Pseudomonas

aeruginosa DT-Ct1

Bidlan and

Manonmani (2002)

Pseudomonas

aeruginosa DT-Ct2

Bidlan and

Manonmani (2002)

Pseudomonas

fluorescens DT-2

Bidlan and

Manonmani (2002)

Serratia

marcescens

Mendel and Walton

(1966)

Serratia

marcescens DT-1P

Bidlan and

Manonmani (2002)

Streptomyces

annomoneus

Chacko et. al.

(1996)

Streptomyces

aureofacians

Chacko et. al.

(1996)

Streptomyces

viridochromogens

Chacko et. al.

(1996)

Xanthomonas spp.,. Johnson et. al.

(1967)

Phanerochaete

chrysosporium

(fungus)

Bumpus and Aust

(1987)

Trichoderma viride

(fungus)

Matsumura and

Boush (1968)

Aerobacter

aerogenes

Mecksongsee and

Guthrie (1965)

Bacillus cereus Mecksongsee and

Guthrie (1965)


20

-HCH

-HCH

Bacillus

megaaterium

Mecksongsee and

Guthrie (1965)

Citrobacter freundii Jagnow et. al.

(1977)

Clostridium rectum Jagnow et. al.

(1977)

Escherichia coli Francis et. al.

(1975)

Pseudomonas

fluorescens

Mecksongsee and

Guthrie (1965)

Pseudomonas

putida

Benzet and

Matsumara (1973)

Pseudomonas

paucimobilis

Bachmann et. al.

(1988)

Pseudomonas

spp.,. Sahu et. al. (1990)

Anabaena spp.,.

(Cyanobacteria)

Kurtiz and Wolk

(1995)

Nostocellipssosun

(Cyanobacterium)

Kurtiz and Wolk

(1995)

Phaenrochaete

chrysosporium

(fungus)

Mougin et. al. (1996)

Trametesversicolor

(fungus)

Singh and Kuhad

(1999a)

Phanerochaete

sordida (fungus)

Singh and Kuhad

(1999b)


21

Cyathus bulleri

(Fungus)

Singh and Kuhad

(1999b)

Organophosphates

Parathion

Flavobacterium

spp.,

Sethunathan and

Yoshida (1973)

Pseudomonas

aeruginosa

Gibson and Brown

(1974)

Pseudomonas

diminuta Serdar et. al. (1982)

Pseudomonas

melophthara

Boush and

Matsumura (1967)

Pseudomonas

stutzeri

Doughton and Hsieh

(1967)

Phorate

Rhizobium japonium Mich and Dahm

(1970)

Rhizobium melioloti Mich and Dahm

(1970)

Streptomyces

lividans Steiert et. al. (1989)

Bacillus megaterium La Partourel and

Wright (1976)

Carbamates

Pseudomonas

cepacia

Venkateswarlu et.

al. (1980)


22

Carbaryl

Pseudomonas

melophthora

Bousch and

Matsumura (1967)

Pseudomonas

aeruginosa

Chapalamadugu

and Chaudhry

(1993)

Gliocladium roseum

(Fungi)

Liu and Bollog

(1971)

Aspergillus flavus

(Fungi)

Bollog and Liu

(1972)

Aspergillus terreus

(Fungi)

Bollog and Liu

(1972)

Culcitalna spp.,.

(Fungi) Sikka et. al. (1975)

Halosphaeria spp.,.

(Fungi) Sikka et. al. (1975)

Fusarium solani

(Fungi)

Bollog and Liu

(1972)

Rhizopus spp.,.

(Fungi)

Bollog and Liu

(1972)

Penicillium spp.,

(Fungi)

Bollog and Liu

(1972)

Carbofuran

Achromobacter

spp., WMIII Karns et. al. (1986)

Arthrobacter spp., Ramanand et. al.

(1988)

Flavobacterium

spp.,.

Chaudhry and Ali

(1988)


23

Pseudomonas

cepacia

Venkateswarlu et.

al. (1980)

Pseudomonas

stutzeri

Mohapatra and

Awasthi (1977)

Bacillus pumilis Mohapatra and

Awasthi (1977)

s-Triazines

Pseudomonas spp., Cook and Hutter

(1981)

Klebsiella

pheumoniae

Cook and Hutter

(1981)

Rhodococcus

corallinus

Cook and Hutter

(1981)

Rhizobium spp.,

PATR

Bauguard et. al.

I(1997)

Phanerochaete

chrysosporium

(Fungus)

Mougine et. al.

(1994)

Source: Singh et al.,1999.

Table 2.3: Bioconversion of xenobiotics effected by Pseudomonads

Mode of action Species

Hydrolysis of carbaryl, dichlorphos, diazinon,

parathion

Hydrolysis of parathion

Ps.melophthora

Ps.stutzeri

Dehalogenation of halide acetate Ps. Species

Total dehalogenation of DDT, aromatic ring cleavage Ps.aeruginosa

Total degradation of 3- chlorobenzoate Ps.putida


24

Oxidative dehalogenation of lindane Ps.putida

Reduction of nitro group in 4,6- dinitro-q- cresol Ps. Species

Total degradation of 2,4,5- T Ps.sepacia

Degradation of toluene, xylene, styrene, α-

methylstyrene

Ps.putida

Ps.aeruginosa

Source: Golovleva et al., 1990.

Several microorganisms are known to degrade DDT anaerobically

(Wedemeyer 1966). The primary metabolic mechanism that was studied was

the reductive dechlorination of DDT, with the formation of DDD (1,1-dichloro-

2,2-bis(4-chlorophenyl)ethane or dichlorodiphenyldichloroethane) (Kallman

and Andrews 1963; Barker and Morrison 1964). This degradation was later

determined to be microbial and Proteus vulgaris was isolated (Barker et al.,

1965) which could degrade DDT mainly to DDD. Nadeau et al., (1994) has

reported the aerobic degradation of DDT via 4- chlorobenzoic acid by

Alkaligenes eutrophus A5. Cell free extracts of Escherichia coli, Klebsiella

pneumoniae and Enterobacter aerogenes dechlorinated p,pˈ-DDT to DDE

anaerobically (Singh et al., 1999). Bumpus and Aust (1987) reported the

degradation of DDT by the white rot fungus, Phanerochaete chrysosporium.

Chacko et. al. (1966) isolated numerous actinomycetes (Nocardia spp.,

Streptomyces aureofaciens, Streptomyces cinnamoneus, Streptomyces

viridochromogenes) from soil, which readily degraded DDT to DDD. These

organisms however, required another carbon source to facilitate degradation.

Soil fungi not only produced DDD and small amounts of dicofol (4,4’-dichloro-

a-(trichloromethyl) benzhydrol), but some variants could produce DDA (bis(4-

chlorophenyl)acetic acid) or DDE (1,1-dichloro-2,2-bis(4-chlorophenyl)ethene)

exclusively (Matsumura and Boush 1968). Wedemeyer (1967) reported

dehalogenation of DDT to various metabolites under anaerobic conditions by

Aerobacter aerogenes. DDD was obtained under both aerobic as well as

anaerobic conditions when DDT was incubated with Aerobacter aerogenes

(Mendel et al., 1967; Wedemeyer 1967). Escherichia coli dechlorinated 50%


25

of DDT to DDE when grown in various broths or skimmed milk (Langlois

1967). Under aerobic conditions the major product of DDT metabolism, in

Bacillus cereus, B. coagulans, B. subtilis, was DDD while DDMU (1-chloro-

2,2-bis(4-chlorophenyl)ethylene), DDMS (1-chloro-2,2-bis(4-

chlorophenyl)ethane), DDNU (2,2-bis4-chlorophenyl)ethane), DDOH (2,2-

bis(4-chlorophenyl)ethanol), DDA and DBP (4,4’-dichlorobenzo phenone)

were in trace amounts and were found under anaerobic conditions (Langlois

et al., 1970). Hydrogenomonas spp., yielded DDD, DDMS, DDMU, DBH (4,4’-

dichlorobenzhydrol), DDM (bis(4-chlorophenyl)methane) and DDA (Focht and

Alexander 1970). DDD was further degraded through dechlorination,

dehydrochlorination and decarboxylation to DBP or to a more reduced form

DDM.

2.5 Anaerobic Degradation

In the anaerobic mode of degradation the electron acceptor is a

molecule other than O2. This could be NO-3, SO4

2- , Fe3+, H+, S, fumarate,

trimethylamineoxide, an organic compound, or CO2 (Cookson 1995). The

term “dehalorespiration” has been coined for anaerobic bacteria that couple

the reductive dehalogenation of chlorinated aliphatic and aromatic compounds

to ATP synthesis via an electron transport chain (Wolhfarth and Diekert 1997).

Reductive dechlorination or reductive dehydrogenolysis is a common

biotransformation pathway for chloroaliphatics containing one or two carbon

atoms, under methanogenic conditions (Semprini 1997). Chen et al., (1996)

studied the biotic transformation of TeCE under methanogenic conditions. A

strictly anaerobic homoacetogenic bacterium and an uncharacterized

anaerobic mixed culture were shown to use chloromethane as a ‘C’ and

energy source (Susanna et al., 1993). Most of the chlorinated aromatic

compounds and several pesticides are known to be best degraded under

anaerobic conditions. Ramanand et al., (1993) have reported rapid

degradation of chlorinated benzenes and toluenes under methanogenic

conditions. Several chlorinated aromatic compounds have been shown to be

degraded under methanogenic conditions. These include 2,4,5-

trichlorophenoxyacetate, 3-chlorobenzoate, 2,4-dichlorophenol, 4-


26

chlorophenol, 2,3,6-trichlorobenzoate, and dichlorobenzoates (Gibson and

Sulfita 1990; Zang and Wiegel 1990). Jiangzhong et al., (2003) have reported

complete detoxification of VC by an anaerobic enrichment culture, which they

later identified as Dehalococcoides spp.,. Buser and Muller (1995) have

studied degradation of pesticide hexachlorocyclohexane (HCH) and its

isomers in sewage sludge under anaerobic conditions. Studies by Holscher et

al., (2003) showed anaerobic reductive dechlorination of chlorobenzene

congeners in cell extracts of Dehalococcoides strain CBDB1. Chlorophenol

degradation coupled to SO42− reduction has been documented by Haggblom

and Young (Haggblom and Young 1990). They suggested that degradation of

chlorinated aromatic compounds not only takes place under sulfate-reducing

conditions but is in fact coupled to sulphate reduction (Castro and Belser

1990). Vargas et al., (2001) have given an account of anaerobic

dechlorination of chlorinated dioxins in estuarine sediments.

2.6 Sequential Degradation

Although degradation of chlorinated aliphatic and aromatic compounds

has been reported both under aerobic and anaerobic conditions, sequential

use of these processes always has an advantage over using them individually

for complete mineralization of heavily chlorinated compounds. It is generally

implied that aerobic microbes often fail to metabolize heavily chlorinated

compounds. For example, several bacteria capable of oxidizing TCE, DCE,

and VC by using nonspecific enzymes cannot oxidize TeCE by any of these

enzyme systems (Nelson et al., 1988; Wackett and Gibson 1988; Tsien et al.,

1989; Vannelli et al., 1990). Aerobic bacteria that rapidly biodegrade

monochlorinated benzenes are usually unable to degrade heavily chlorinated

benzene compounds (Zang and Wiegel 1990). Similarly, increased resistance

of chloroalkenes to biological reductive dechlorination has been observed in

anaerobic reactors and anaerobic freshwater microcosms (Bouwer and

McCarty 1983; Parsons et al., 1984). Therefore, it has been suggested that

detoxification and complete mineralization of chlorinated wastes can be easily

achieved by using a sequential treatment process, that is, anaerobic followed

by aerobic treatment. For instance, the fungicide HCB (hexachlorobenzene)


27

and polychlorinated biphenyl (PCB) undergo reductive dechlorination in

anaerobic environments (Tsien et al., 1989; Vannelli et al., 1990; Fathepure et

al., 1988). The products are congeners bearing fewer chlorine substituents,

which are more susceptible to biodegradation by aerobic bacteria (Spain and

Nishino 1987; Brown et al., 1987). A sequential treatment step will ensure

total mineralization of these chlorinated toxic compounds.

2.7 Role of Electron Donors in Dechlorination

Reductive dehalogenation reaction, whether catalyzed by a transition

metal, bacterial cofactors, or an enzyme, requires two electrons. Therefore, a

source of electrons must be available for the reaction to take place (Bhaskara

et al., 1998). The source of electrons (or electron donor) for a dechlorination

reaction is usually a reduced substrate provided for microbial growth. Nies

and Timothy (Nies and Vogel 1990) studied the effects of different organic

substrates on the ability of anaerobic sediment enrichment to reductively

dechlorinate polychlorinated biphenyls. They used acetate, acetone,

methanol, and glucose and found that the relative rates of dechlorination were

in the order methanol > glucose > acetone > acetate fed cultures (Nies and

Vogel 1990). De Bruin et al., (1995) observed biological reductive

dechlorination of TeCE to ethane with lactate as the electron donor (Gibson

and Sulfita 1990) observed that addition of butyrate, propionate, ethanol, or

acetate increased not only the rate of dehalogenation of

trichlorophenoxyacetic acid but also the extent of its degradation. Hydrogen,

formate, ethanol, propionate, or acetate can serve as the source of reducing

equivalents required for dechlorination in the bacteria Desulomonile tiedje

(Dolfing 1990; Mohn and Tiedje 1991). A similar observation was seen in case

of trichlorophenol (TCP) degradation, where yeast extract was the preferred

primary substrate and resulted in complete degradation of the target

compound within 3 days (Madsen and Aamand 1992). With peptone and

casamino acid, complete transformation was observed only after 6–7 days

(Galli and McCarty 1989). Studies by Holliger et al., (1992) showed that HCB,

all three isomers of TeCE, 1,2,3-TCB (trichlorobenzene), and 1,2,4-TCB were

reductively dechlorinated by enrichment culture in the presence of lactate,


28

glucose, ethanol, or isopropanol as electron donors. Lactate, ethanol, and H2

appeared to be the best substrates. Moreover, dechlorinating activity could

not be maintained when electron donor was not added (Holliger et al., 1992).

Gibson and Sewell (Gibson and Sewell 1992) observed that lactate,

propionate, crotonate, butyrate, and ethanol stimulated dechlorination activity

of TeCE in methanogenic slurries made with aquifer solids. Acetate,

methanol, and isopropanol did not support dehalogenation (Gibson and

Sewell 1992). For bacteria like the Nitrosomonas spp., capable of degrading

several chlorinated aliphatic compounds, ammonia served as the electron

donor. A study demonstrated that dehalogenation of DCE in a contaminated

soil required fatty acids and alcohols as electron donors. Supporting evidence

was also given to show that the dechlorination process stops once the

electron donor is depleted (Villarante et al., 2001). Smatlak et al., (1996)

observed that PCE dechlorination rates decreased significantly at lower H2

concentrations, which was added as an electron donor in the experiment.

Dechorination of PCP was enhanced by the addition of glucose to a UASB

reactor fed with PCP and phenol (Hendriksen et al., 1992).

2.8 Role of Electron Acceptors in Dechlorination

All energy-yielding reactions are oxidation–reduction reactions. The

reduction reaction that is, the reaction involving the electron acceptor,

establishes the metabolism mode. Microbes preferentially utilize electron

acceptors that provide the maximum free energy during respiration (Stumm

and Morgan 1981). Among the common electron acceptors used by

microorganisms, O2 typically provides the maximum free energy during

electron transfer, followed by nitrate, Mn(IV),Fe(III), SO42−, and CO2 (Cobb

and Bouwer 1991). Cobb and Bouwer (1991) used a mixture of primary

electron acceptors like O2, nitrate, and sulfate for the transformation of 1,1,1-

TCE, TeCE, and chlorinated benzenes, and suggested sulfate to be an

important primary acceptor. Experimental studies with a biofilm using a single

electron acceptor showed that halogenated aliphatic compounds such as

TCE, chloroform, and others could be transformed under methanogenic and

sulfate reducing conditions (Bouwer and McCarty 1983). Chlorinated


29

compounds are stronger oxidants than nitrate (Vogel, et al., 1987). On the

basis of such thermodynamic considerations, chlorinated hydrocarbons have

been shown to act as terminal electron acceptors in a respiratory process

(Dolfing and Harrison 1992; Holliger et al., 1988).

Cupples (2003) observed growth of a Halococcoides-like organism on

VC and cis dichloroethene as electron acceptor. Dehalococcoides

ethenogens strain 195 completely dechlorinates PCE to ethene using H2 as

electron donor and PCE as the electron acceptor (Maymo-Gatell 1997). A

study by Holliger et al., (1993) revealed that a highly purified enrichment

culture could use only PCE or TCE as electron acceptor and O2, NO3−, NO−

2,

SO42−, SO3

2−, S2O32−, S, or CO2 could not replace PCE or TCE as electron

acceptor. Even organic electron acceptors such as acetoin, acetol, dimethyl

sulfoxide, fumarate, and trimethylamine N-oxide were not utilized by the

organisms (Holliger et al., 1993). PCE as an electron acceptor was used by

an acetate-oxidizing anaerobic bacteria identified as Desulfomonas

michiganensis spp., nov (Sung et al., 2003).

2.9 Role of Transition Metal Cofactors in Dechlorination

Transition metal cofactors can mediate nonspecific reactions with

hydrophobic chlorinated pollutants that gain entry into bacterial cells by

partitioning through membranes (Gantzer and Wackett 1991). There are two

different classes of transition metal cofactors found in bacteria that grow

under anaerobic conditions (Wackett et al., 1989). In the first type, the metal is

coordinated by a stable macrocyclic ligand system, which in turn can be

bound by proteins. In the second type, metal(s) is (are) directly coordinated to

protein ligands. Both type of redox-active centers display great versatility in

their biological functions (Gantzer and Wackett 1991). The cobalt-containing

cobalamins and the iron coenzyme hematin (II) show catalytic activities in

addition to their biological role as electron carriers (Hogenkamp 1975;

Hambright 1975). Iron–S clusters, which also function in electron transfer, are

now implicated as key participants in several enzyme-catalyzed hydrolytic

reactions (Krone et al., 1989). Gantzer and Wackett (1991) noted that

bacterial transition metal coenzymes vitamin B12 (Co), coenzyme F430 (Ni),


30

and hematin (Fe) catalyzed the reductive dechlorination of polychlorinated

ethylenes and benzenes, whereas the electron-transfer proteins four-iron

ferridoxin, two-iron ferrodoxin, and azurin (Cu) did not. Cobalamins, coezyme

F430, and hematin have recently been shown to dehalogenate chlorinated

methanes in the presence of a reductant (Krone et al., 1989). Carbon

tetrachloride (CT) degradation rates increased linearly with higher intracellular

vitamin B12 content (Zon et al., 2000). In many cases, the microbial

transformation of CT is considered to be closely related to the presence of

microbial cofactors such as porphinoids and corrinoids (Villarante et al.,

2001). Corrinoids such as aquocobalamin or methylcobalamin catalyze the

reduction of tetrachloromethane or trichloromonofluoromethane with titanium

(III) citrate or with dithiothreitol as electron donors (Van Eckert et al., 1998).

Klecka and Gonsior (1984) observed transformation of CT, chloroform, and

1,1,1-tetrachloroethane by iron porphyrins with sulfide as the reductant. More

recently, zero-valent iron has also been reported to catalyze reductive

dechlorination reactions at extremely high rates (Lu et al., 2004).

2.10 Enzymes involved in dechlorination

Microorganisms have evolved a diverse potential to transform and

degrade halogenated organic compounds. They produce an array of enzymes

that bring about dehalogenation and degradation of both aliphatic and

chloroaromatics compounds. The reactions catalyzed by such enzymes can

be broadly classified as follows:

Reaction Enzymes

Oxidative dehalogenation Mono- or dioxygenases

Dehydrohalogenation Dehydrohalogenases

Substitutive dehalogenation Halidohydrolases

Dechlorination via methyl transfer Methyltransferases

Reductive dehalogenation Dehydrohalogenases


31

2.10.1 Oxidative Dehalogenation

Oxidative dechlorination of aliphatic chlorinated compounds is a result

of mono- or dioxygenase enzymes that function via metabolic or cometabolic

reactions. The chlorinated hydrocarbon competes along with the growth

substrate of the organism for the active site of the oxygenase enzyme. The

organisms, however, are not known to benefit from the cometabolic processes

(Fetzner 1998). The initial step in the aerobic transformation of chlorinated

alkenes is generally assumed to be an epoxidation of the carbon–carbon

double bond (Hartmans et al., 1989). The subsequent metabolism of the

reactive haloepoxides is not known in detail, but extensive dehalogenation is

frequently observed (Klecka and Gonsior 1984). An example of this kind of

dehalogenation is by methane monooxygenases (MMO), which is thought to

catalyze the conversion of haloalkenes such as TCE to its epoxide, which

subsequently undergoes isomerization or hydrolysis. The reaction is

represented by (Fetzner 1998) (Fig.2.1).

Fig. 2.1: Oxidative dehalogenation reaction

where (a) is trichloroethene, (b) methane monooxygenase (MMO), and

(c)epoxide.

A high degree of specificity of this enzyme toward TCE was observed

in Methylosinus trichosporium OB3b (Fetzner and Lingens 1994; Fox et al.,

1990). There probably are different mechanisms of TCE oxidation by

oxygenases. Microbial oxidation of TCE has been reported to be catalyzed by

toluene 2,3-dioxygenase (Li and Wackett 1992; Wackett and Householder

1989; Zylstra et al., 1989), toluene2-monooxygenase (Folsom et al 1990;

Nelson et al., 1987; Shields et al., 1989; Shields et al., 1991), toluene 4-


32

monooxygenase (Winter et al., 1989), phenol hydroxylase (Harker and Kin

1990) and 2,4-dichlorophenol hydroxylase and propane monooxygenase

(Wackett et al., 1989) and the involvement of separate dioxygenases was

noted from data on plasmid profiles and DNA hybridization in Pseudomonas

putida (Lu et al., 2004) which utilizes a broad range of mono-, di-, and

trichlorinated benzoates (Brenner et al., 1993). Similar to methane

monooxygenase, ammoniamonooxygenase oxidizes not only TCE but a

variety of n-chlorinated alkenes (Leng 1986). Oxygenolytic dehalogenation of

haloaromatic compounds is either catalyzed by specific oxygenases or occurs

during a conversion, catalyzed by the enzyme for the corresponding

unsubstituted substrate.

Two-component dioxygenases such as 4-chlorophenyl acetate, 3,4-

dioxygenase, and 2-halobenzoate 1,2-dioxygenase preferentially catalyze

chloroaromatic compounds (Markus et al., 1986; Schweizer et al., 1987). In

the degradation of 1,2,4,5-tetrachlorobenzene by Pseudomonas strain PS14,

an initial 5,6-dioxygenating attack is followed by spontaneous elimination of

HCl during rearomatization of the dehydrodiol, yielding 3,4,6-trichlorocatechol

(Sander et al., 1991).

Dioxygenolytic dechlorination of 2, 2-dichlorobiphenyl, 2, 3-

dichlorobiphenyl, and 2, 5, 2-trichlorobiphenyl at the ortho position is

catalyzed by biphenyl 2,3-dioxygenase of Pseudomonas strain LB400

(Haddock et al., 1995). All these dioxygenases have been proposed to

catalyze the formation of cis-diols, which spontaneously rearomatize with

concomitant Cl- elimination, yielding a catechol product. In the first step of

PCP degradation by Sphingomonas chlorophenolica ATCC 39723, a soluble

flavoprotein monooxygenase catalyzes its NADPH-dependent conversion to

tetrachloro p-hydroquinone (Xun et al., 1991; Xun et al., 1992).

2.10.2 Dehydrohalogenation

This type of dechlorinating process eliminates HCl from the haloorganic

substrate, leading to the formation of a double bond. Dehydrohalogenation

occurs during the mineralization of insecticide γ –HCH by Sphingomonas

paucimobilis UT26 (Nagata et al., 1993). The elimination of HCl from both γ -


33

HCH and an intermediate metabolite γ –pentachlorocyclohexene is catalyzed

by a dehydrochlorinase designated LinA (Imai et al., 1991). The enzyme

catalyzes the release of three chloride ions per molecule of γ -HCH, but its

substrate specificity is narrow. α-HCH, γ -HCH, δ-HCH, α-

pentachlorocyclohexene and γ -pentachlorocyclohexene are the only

substrates converted. It has been suggested that Lin A catalyzes the

stereoselective dehydrochlorination of HCH with a trans and diaxial pair of

hydrogen and chloride. Two other dehydrochlorinase enzymes have also

been described, namely, glutathione-dependent DDT dehydrochlorinase from

the housefly and the 3-chloro-D-alanine dehydrochlorinase from P. putida,

which requires pyridoxal 5-phosphate (Nagata et al., 1993).

Dehydrohalogenases are also involved in the ortho cleavage of

chlorocatechols, which results in chlorinated cis-muconates, which are

cycloisomerized to diene lactones (Vollmer et al., 1994).

2.10.3 Substitutive Dehalogenation

Substitutive dehalogenation of chlorinated compounds takes place by

three different processes:

1. Hydrolytic processes.

2. Thiolytic processes.

3. Intramolecular substitution reactions.

2.10.3.1 Hydrolytic processes

Hydrolytic dehalogenation of several heterocylic, aromatic, and

aliphatic compounds has been reported (Fetzner and Lingens 1994; Hardman

1991; Leisinger and Bader 1993; Janssen et al., 1994; Slater et al., 1995;

1997). These reactions are catalyzed by halidohydrolases. Hydrolytic

dechlorination of haloalkanes was first found with the haloalkane

dehalogenase from the nitrogen-fixing hydrogen bacterium Xanthobacter

autotrophicus GJ10. Because of the presence of two halidohydrolases, strain

GJ10 is capable of rapid utilization of 1,2-dichloroethane. Both these

dehalogenases in X. autotrophicus are synthesized constitutively (Janssen et

al., 1985). These enzymes have a broad specificity and catalyze the


34

dehalogentation of more than 24 haloaliphatic compounds. The haloalkane

dehalogenase gene dhl A has been cloned and sequenced (Janssen et al.,

1989). One haloalkane halidohydrolase encoding gene is present in the

plasmid (designated pXAU1), while the gene encoding the second enzyme, 2-

haloalkanoic acid halidohydrolase, is located on the chromosome.

Nucleophilic displacement with H2O was suggested as the mechanism

of halide release (Tardif et al., 1991). Asp-24 is the nucleophilic residue

attacking the substrate. It is assumed that the covalent intermediate is an

ester, which must be subsequently cleared by water molecule, releasing the

alcohol (Franken et al., 1991). The hydrolytic dechlorination reaction of 1,2-

dichloroethane in Xanthobacter autotrophicus GJ10 is given as follows

(Fetzner 1998) (Fig. 2.2).

Fig. 2.2: Hydrolytic dehalogenation reaction

(I) 1,2-Dichloroethane; DhlA, haloalkane dehalogenase.

(II) 2-Chloroethanol; MoX, alcohol dehydrogenase.

(III) Chloroacetaldehyde; Ald, aldehyde dehydrogenase.

(IV) Chloroacetate; Dhl B, 2-haloacid dehalogenase.

(V) Glycolate; PQQ, pyrroquinoline quinone.

Since delocalization of the pi electrons considerably stabilizes the

aromatic ring system, it was previously thought unlikely that bacteria have

evolved enzymes for the direct hydrolysis of the aromatic carbon–halogen


35

bond. Deethylsimazine, a monohydroxylated s-triazine derivative, has

considerable aromatic character, but in contrast to the benzenoid ring,

delocalization of the pi electrons is not complete. Hydrolytic removal of

substituents has been described for various s-triazines (Knackmuss 1981).

Cook and Hutter (1986) have shown that two isofunctional but different

enzyme fractions from Rhodococcus corallinus NRRLB-15444R hydrolytically

dechlorinated diethylsimazine to N-ethylamine. No cofactors were required for

dechlorination. This hydrolytic substitution at the aromatic ring is chemically

feasible because of the low electron density at the ring carbon atoms (Cook

and Hutter 1986). An example for hydrolytic dehalogenation reaction is the

conversion of 4-chlorobenzoate to 4-hydroxybenzoate. This reaction requires

three enzymes, namely 4-chlorobenzoate coenzyme A (CoA) ligase, 4-

chlorobenzoyl-CoA dehalogenase, and 4-hydroxybenzoyl CoA thioesterase.

This conversion has been shown to be catalyzed by a number of bacterial

strain belonging to the genera Pseudomonas, Arthrobacter, Acinetobacter,

Alcaligenes, Nocardia, and Corynebacterum (Brunner et al., 1980). In the

conversion of 4-chlorobenzoate to 4-hydroxybenzoate by Pseudomonas strain

CBS3, Loffler and Muller (1991) identified 4-chlorobenzoyl CoA as an

intermediate in the dehalogenation reaction and proposed the reaction

mechanism. In the first step, a 4-chlorobenzoate lyase catalyzes the

adenylation of the carboxy group followed by displacement of the AMP, a thiol

group from CoA, leading to the formation of the thioester 4-chlorobenzoyl

CoA. The formation of the CoA ester activates the substituent in the para

position for a nucleophilic attack and enables the substitution of the chlorine

by a hydroxyl group from H2O, catalyzed by dehalogenase (Loffler and Muller

1991). The reaction can be represented as Fig. 2.3.


36

Fig. 2.3: Hydrolytic dehalogenation reaction

where (I) is 4-chlorobenzoate CoA ligase (II) is 4-chlorobenzoyl CoA

dehalogenase and (III) is 4-hydroxybenzoyl CoA thioesterase.

2.10.3.2 Thiolytic processes

The thiolytic substitutive dehalogenation process is catalyzed by

glutathione S-transferase enzymes. This process has been extensively

studied in methylotrophic bacteria. Dechlorination of dichloromethane by

facultative methylotrophic bacteria is catalyzed by inducible glutathione S-

transferases. Dichloromethane is converted to formaldehyde and inorganic

chloride with S-chloromethylgutathione as intermediate and the formaldehyde

so formed is a central metabolite of methylotrophic growth (Fetzner 1998).

Pseudomonas strains, Hyphomicrobium strains, and several

Methylobacterium spp., strains have been shown to contain these enzymes

(Loffler and Muller 1991; Galli and McCarty 1989; Kohler et al., 1986; Kohler-

Staub and Leisinger 1985). The reaction has been described as follows (Fig.

2.4).


37

Fig. 2.4: Thiolytic substitutive dehalogenation reaction

where (I) is dichloromethane dehalogenase (II) is formaldehyde dehalogenase

and (III) is formate dehydrogenase.

2.10.3.3 Intramolecular substitution reactions

These reactions are catalyzed by halohydrin–hydrogen halide lyases,

also called halohydrin epoxidases. They were first discovered by Castro and

Bartnicki (1968) and Bartnicki and Castro (1969) in 1968 from a 2,3-dibromo-

1-propanol utilizing Flavobacterium spp., They constitute a unique group of

dehalogenating enzymes (Castro and Bartnicki 1968; Bartnicki and Castro

1969). In 1989, Van den Wijngaard et al., (1989) reported the degradation of

epichlorohydrin and halohydrins by Pseudomonas strain AD1, Arthrobacter

strain AD2, and Coryneform strain AD3. Halohydrin dehalogenase from strain

AD2 converted C-2 and C-3 chloroalcohols and was active with chloroacetone

and 1, 3-dichloroacetone as well, yielding epoxides as products. Neither

cofactors nor O2 was required for the dehalogenation. Thus, the reaction

mechanism was thought to proceed via intramolecular substitution (Van den

Wijngaard et al., 1989; Kesai et al., 1990). The reaction did not require any

cosubstrate, and purified haloalcohol dehalogenase from AD2 showed no

immunological cross-reactions with haloalkane or 2-haloacid halidohydrolases

(Van den Wijngaard et al., 1991).


38

2.10.4 Dechlorination via Methyl Transfer

A chloromethane dehalogenase, which is inducible by chloromethane,

transfers the methyl group of its substrate onto tetrahydrofolate, producing

methyltetrahydrofolate and chloride. The further metabolism of

methyltetrahydrofolate to acetate proceeds via the reactions of the acetyl CoA

pathway (MeBmer et al., 1993; MeBmer et al., 1996) Dehalobacterium

formicoaceticum, which utilizes dichloromethane as sole energy source,

ferments DCM to acetate and formate in a molar ratio of 1:2 (Knackmuss

1981). Cell extracts in the presence of tetrahydrofolate, ATP, methyl viologen,

and H2 were found to convert DCM to methylene tetrahydrofolate. DCM is

assumed to react with a reduced Co(I) corrinoid, forming chloride and

chloromethyl-Co(III) corrinoid, which acted as a donor for a methyltransferase,

generating chloromethyltetrahydrofolate. The latter spontaneously rearranged

to yield chloride and N5, N10-methylenetetrahydrofolate (Magli et al., 1998).

2.10.5 Reductive Dehalogenation

Reductive dehalogenation is a two-electron-transfer reaction that

involves the release of the halogen as a halogenide ion and its replacement

by hydrogen. The mechanisms of reductive dehalogenation of haloaliphatic

compounds is not fully understood, although there are a number of reports on

the metabolism of halogenated aliphatic hydrocarbons under methanogenic,

sulfate-reducing, and denitrifying conditions (Barrio-Lage et al., 1986; Belay

and Daniels 1987; Bouwer and McCarty 1985; Di Stefano et al., 1991; Egli et

al., 1988; 1990; 1989; Freedman and Gosett 1989; Krone and Thauer 1992;

Lewis and Crawford 1993; Mikesell and Boyd 1990; Pavlostathis and Zhuang

1991; Tatara et al., 1993). For the strictly anaerobic methanogens, Fathepure

and Boyd (1988) presented a scheme linking reductive dechlorination to

methanogenisis. In this scheme they proposed that the chlorinated ethylenes

serve as electron acceptors. Clostridium strain TCAIIB isolated from a

methanogenic mixed culture was found to reductively dechlorinate 1,1,1-

trichloroethane to 1,1-dichloroethane and dechlorination of

tetrachloromethane to tri- and dichloromethane (Kobayashi and Rittmann

1982). There is evidence for reductive dehalogenation under methanogenic,


39

sulfidogenic, and even denitrifying conditions of a number of haloaromatics

such as chlorobenzenes, chlorotoluenes, chlorobenzoates, 2,4

dichlorobenzoate, a number of chlorinated phenols, tri- and

tetrachlorocatechols, di-, tri-, and tetrachloroanilines, 2,4,5-trichlorophenoxy

aceticacid and polychlorinated biphenyls. In the reductive dechlorination

mechanism, a reduced organic substrate or H2 might be the source of both

the reducing power and the protons (Leng 1986).

Biotransformation of many halogenated pesticides has been known to

involve reductive dehalogenation. A list of halogenated pesticides (most of

which are chlorinated) and anthropogenic compounds undergoing reductive

dehalogenation was presented by Kobayashi and Rittmann (1982) and Mohn

and Tiedje (1992). Desulfomonile tiedjei DCB-1 reductively dechlorinates 3-

chlorobenzoate, meta-substituted dichlorobenzoates, chlorophenols, and

tetrachloroethylene (Linkfield and Tiedje 1990; De Weerd and Sulfita 1990;

Mohn and Kennedy 1992; Fathepure 1987) Clostridium rectum S-17, C.

sphenoides, several Bacillus strains, and members of the family

Enterobacteriaceae are involved in reductive dechlorination of lindane

(Jagnow et al., 1977; Haider 1979; Mac Rae et al., 1969; Heritage Mackar

1977; Ohisa et al., 1982; Ohisa and Yamaguchi 1978; Ohisa et al., 1980). A

metabolic pathway of DDT dechlorination by Aerobacter aerogenes involving

reductive and dehydrochlorination steps, yielding 4,4’-dichlorobenzophenone,

was proposed by Wedemeyer (1967). Dicamba, after demethylation, was

reductively dechlorinated to 6-chlorosalicylate by an anaerobic consortium

(Taraban et al., 1993).

2.11 Evidence of Enzyme-mediated Degradation of Xenobiotic Compounds The application of fungi for the cleanup of contaminated soil first came

to attention in the mid-1980s when the white rot fungus Phanerochaete

chrysosporium was shown to metabolize a range of organic environmental

contaminants (Canet et al., 2001; Trejo-Hernandez et al., 2001). Later, this

ability was demonstrated for other white rot fungi, including Trametes

versicolor and Pleurotus ostreatus (Ghani et al., 1996). Xenobiotics have


40

been shown to share at least one of many sub-structures (e.g., combination of

functional groups) present in the lignin molecule (Gadd 2001). It has been

shown that both laccase (LAC) and peroxidises co-metabolize these

compounds with lignin through similar oxidative mechanisms (Han et al.,

2004; Gadd 2001; Pointing 2001), Determining the activity of LAC in soil

inoculated with white rot species provides a measure of the colonizing ability

of the fungus and can be used to monitor the bioremediation of numerous soil

contaminants, among them triazine pesticides (Fragoeiro and Magan 2005;

2008; Pointing 2001). Novotný et al., (2004) measured LAC activity to

demonstrate the correlation between its production and the degradation of

polycyclic aromatic hydrocarbons (PAHs) by several strains of white rot fungi

in both liquid culture and soil. The applications of fungi for biodegradation of

xenobiotics were found to be related to the production of LACs, Mn-

peroxidase or (less frequently) ligninperoxidases, both alone or in

combination, which has been corroborated by other studies (Fragoeiro and

Magan 2005; 2008; Pointing 2001; Mswaka and Magan 1999). LAC activity in

the biodegradation of xenobiotic compounds with lignin-like structures has

already attracted considerable interest (Tuor et al., 1995), and its

biodegradative effects on different contaminants have been exhaustively

studied. Specifically, LAC enzyme is a copper-containing phenoloxidase

involved in the degradation of lignin (Radtke et al., 1994), and it oxidizes

phenol and phenolic lignin sub-structures (Valli et al., 1992). The catabolic

role of fungal LAC in lignin biodegradation is not well understood (Tuor et al.,

1995; Hestbjerg et al., 2003), but there have been some successful instances

of this enzyme performing decontamination. Complete removal of benzene

and toluene was observed with the involvement of LAC (Demir 2004). Han et

al., (2004) studied the degradation of phenanthrene by T. versicolor and

purified its LAC. Valli et al., (1992) demonstrated the mineralization of 2,7-

dichlorobenzenop-dioxin by P. chrysosporium and the purified LiPs and MnPs

were capable of mineralization in a multistep pathway. Esposito et al., (1998)

showed that different actinomycetes were able to degrade diuron in soil using

MnPs.


41

2.12 Plants and their associated enzymes as decontaminating agents

An appealing alternative for overcoming some of the drawbacks related

to the use of enzymes in in situ remediation of polluted environments is

phytoremediation. Phytoremediation is the in situ use of plants, their

enzymatic system, their roots and associated microorganisms to degrade,

contain or render harmless pollutants present in different environmental

systems (soil, sediments, groundwater, and air). With respect to their direct

roles in remediation processes, plants may utilize different mechanisms to

efficiently remove both organic and inorganic pollutants from a polluted

environment: a) rhizofiltration; b) absorption; c) concentration and precipitation

of heavy metals by roots; d) phytoextraction, i.e. extraction and accumulation

of pollutants in plant tissues including roots and leaves; e) phytodegradation

i.e. degradation of complex organic molecules in CO2 and H2O and their

incorporation in plant tissues f) rhizodegradation or plant assisted

bioremediation i.e. stimulation of microbial and fungal degradation by the

release of root enzymes and exudates in the rhizosphere; and g)

phytostabilitation, i.e. adsorption and precipitation of pollutants (mainly

metals) with a consequent reduction of their mobility. An interesting

phenomenon is the synergic interaction between plants and microorganisms

that specifically occurs in the soil environment influenced by plant roots, or

rhizosphere. Since plants may be deficient in catabolic pathways for the

complete degradation of pollutants compared with microorganisms, research

efforts have been devoted to engineer plants with genes that can confer them

additional and enhanced degradation abilities. The efficacy of

phytoremediation can be directly enhanced by overexpressing the genes

involved in metabolism, uptake, or transport of specific pollutants in plants.

Moreover, suitable genes may be expressed in roots to enhance the

rhizodegradation of highly recalcitrant pollutants (Abhilash et al., 2009).

Several transgenic plants enriched with genes from humans, microbes,

plants and animals have been produced and have shown enhanced abilities

of metabolizing several xenobiotics. For instance, human and mammalian

(e.g. rat, mouse, rabbit) CYP450 isoenzymes (CYP1, CYP3) genes have

been inserted in Nicotiana tabaccum, Solanum tuberosum, Oryza sativa or


42

Arabidopsis thaliana and the modified plants have shown either herbicide

resistance (e.g. tolerance towards atrazine, simazine) or enhanced

metabolization of xenobiotics (herbicides or volatile halogenated

hydrocarbons) and their subsequent removal from contaminated soil and

groundwater (Abhilash et al., 2009).

Another most promising approach to enhancing phytoremediation

ability is the production of transgenic plants secreting enzymes for the

rhizoremediation of xenobiotics (Abhilash et al., 2009). In these plants

xenobiotics degrading genes have been inserted in their root system and

therefore plants have achieved the capability of secreting degrading enzymes

into the rhizosphere. This method has the unquestionable advantage that

pollutants have not been taken up by plants to be degraded; instead, the

secreted enzymes can degrade the pollutants in the rhizospheric zone (Fig.

2.5). Additional rhizosphere effects may contribute to enhance pollutant

degradation. Microbial density, diversity and/or metabolic activity may

increase because of the release of plant root exudates, mucigel and root

lysates. In addition, the physical and chemical properties of the contaminated

soil can be increased by plants as well as by the contact between the root-

associated microorganisms and the soil contaminants (Fig. 2.6). However, the

use of plants alone can present some limitations. Recently, application of

plant growth-promoting rhizobacteria (PGPR), i.e. bacteria capable of

promoting plant growth by colonizing the plant root has received much

attention for their use in bioremediation of polluted soils (Zhuang et al., 2007).

Several examples of bioremediation of inorganic and organic contaminants by

PGPR are now available. Various bacteria associated with plants like wheat,

alfalfa, tall fescue, Brassica juncea, Indian mustard, canola and others have

been successfully used in the bioremediation of crude oil, PAHs, total

petroleum hydrocarbons, TCE, PCBs and lead, zinc, nichel, cadmium

(Zhuang et al., 2007). Therefore, phytoremediation in conjunction with

rhizospheric microbes may provide sustainable, eco-friendly and efficient

rhizoremediation processes for contaminated ecosystems.


43

Fig. 2.5: Enzymatic and microbial activities responsible for the enhanced remediation in rhizospheric zone (Abhilash et al., 2009)

Fig. 2.6: Overview of the enzymology of biological remediation (Whiteley and Lee, 2006)


44

2.13 Genes Mediating Xenobiotic Degradation in bacteria Many xenobiotic compounds are degraded by microorganisms,

primarily bacteria and fungi (Lal 1982; Lal and Saxena 1982). Usually fungi

are adept in degrading complex and large biomolecules like lignin, whereas

bacteria are proficient in catabolising nonpolymeric, mononuclear aromatic

compounds. Numerous bacterial strains, primarily Pseudomonads, have been

isolated on a wide range of environment contaminated with aromatic

compounds. As early as 1924, de Jong listed 80 different organic compounds

that were degradable by Pseudomonas alone (De Jong 1994). Later Kluyver

(1931) listed 78 compounds that could serve as carbon and energy sources

for the growth of a strain of P. putida. However, the credit of extending our

knowledge of the genus in their major survey of the nutritional capabilities of

representative strains of the fluorescent and nonfluorescent Pseudomonads

goes to Stanier et al., (1966) Since then, this bacterium was found to degrade

a wide array of aromatic compounds, ranging from benzene to benzo (pyrene)

(Gibson et al., 1990; Zylstra and Gibson 1991). Apart from Pseudomonas, the

other bacterial strains known to degrade aromatic compounds include species

of the genus Alcaligenes, Acinetobacter, Arthrobacter, Corynebacterium,

Rhodococcus, and Nocardia (Gibson et al., 1990; Zylstra and Gibson 1991;

Cain 1981; Asturias and Timmis 1993). The degradation pathways and the

genetic mechanisms operative in Pseudomonads are predominantly known

for aromatic compounds and are not fully clear for the degradation of

pesticides or other compounds with complicated structures (except for a few

such compounds as 2,4D, 2,4,5T etc.). With the advancement in recombinant

DNA technology, the understanding of genetic mechanisms and the genetic

manipulations of catabolic genes have mainly emerged from Pseudomonads.

However, in recent years certain bacteria other than Pseudomonads have

also been explored for understanding the genetic mechanism of degradation

(Cain 1981). The commonly used bacteria other than the Pseudomonads

comprise mainly the Gram-positive bacteria such as Rhodococcus globerulus

P6 for biphenyl degradation (Asturias and Timmis 1993). Nocardia spp., for

phenol degradation (Asturias and Timmis 1993) and a few Gram-negative


45

bacteria, such as Sphingomonas paucimobilis for the degradation of HCH and

its isomers (Imai et al., 1991). Degradation of xenobiotics are mediated by a

panoply of enzymatic machinery, which consists of hydroxylating,

dehalogenating, dehydrogenating, and hydrolyzing systems along with the

further complete cleavage systems. The importance of various degradative

enzymes, primarily the oxygenases of the hydroxylating systems in

degradation, prompted microbiologists to manipulate the catabolic genes of

majority of these enzymes to augment the catabolic range and efficiency of

the bacterial strains (Harayama et al., 1992) This has facilitated a greater

understanding of the subject vis-à-vis adding improved methods to the

repertoire of knowledge on conferring catabolic superiority to the concerned

bacterial strains. The progress in this field over the last 2 decades has been

rapid. Generally, it has been found that the pathways of degradation could not

always undergo smoothly, or sometimes do not even start because of the

presence of one or the other bottlenecks or constraints in degradation.

The biological enzymes, isolated chiefly from microorganisms, are

capable of breaking this resonance stability by adding dioxygen to the ring,

which is so important for the operation of Earth’s carbon cycle (Dagley 1986).

All these processes are under the manifestation of several enzymes that have

evolved as individual module during the process of evolution (Van der Meer et

al., 1992; Timmis et al., 1994). The substitution of the aromatic nucleus has

often resulted in the slowing down of the disruption of resonance stability by

the catabolic enzymes. In other terms, substitution of aromatics contributes

differently to the available energy content of the molecule. For instance, one

chlorine atom reduces the energy content of organic substances by

decreasing the available electrons by one, and consequently is reflected in a

reduced growth efficiency of such compounds (Müller and Babel 1994). As a

result, highly chlorinated compounds such as DDT, HCH, PCP, etc. do not

allow bacteria to grow proficiently when any of these compounds are used as

the sole source of carbon and energy. Although aerobic and anaerobic both

mechanisms contribute significantly to the process of decontamination of the

environment, the former method is preferred from environmental point of view,

because this method is fast and substantive when compared with the latter


46

mode. Furthermore, the initial introduction of oxygen into a hydrocarbon by

hydration (anaerobic process) is thermodynamically highly unfavourable

(Dagley 1986). As a result, oxidative cleavage of the aromatic ring is more

prevalent in the biosphere.

Microorganisms channel a compound to the intermediates of Kreb’s

cycle through a known series of dihydroxylated intermediates, such as

catechol, protocatechuate, gentisate, homoprotocatechuate, homogentisate,

or other derivatives (Fig. 2.7). However, in the case of compounds with

complex structures, such as HCH, DDT, PCP, and others, the modes of

aerobic ring cleavages have been different. All these compounds are first

converted into less chlorinated intermediates, similar to that of monochloro- or

dichlorobenzoates, and then are subjected to enzymatic transformations by

the microorganisms (Fig. 2.8).


47

Fig. 2.7: Generalized mechanism of degradation of aromatic hydrocarbons. Formation of catechol, protocatechuate, or gentisate has been predominant in the degradation of aromatic hydrocarbons


48

Fig. 2.8: Degradation of hexachlorocyclohexane (HCH), DDT, and

Pentachlorophenol (PCP) by different bacteria. Formation of

chlorinated products could be noted


49

2.14 Catabolic enzymes of degradation pathways Catabolic enzymes have broadly been grouped into peripheral [or

upper pathway (Carrington et al., 1994; Khanna et al., 1998) and ring-

cleavage or lower pathway (Carrington et al., 1994; Khanna et al., 1998)]

enzymes. The ring cleavage enzymes from a variety of microbes exhibit

significant functional similarity. The peripheral enzymes, however, convert a

xenobiotic compound into metabolites, which are degradable. Peripheral

enzymes thus are the ones that recognize and convert pollutants into

degradable molecules. These are the enzymes for which the products initially

act as substrates, and thus these have to be tailored to suit the chemo- and

region specificities of a variety of xenobiotics. The product of these enzyme

catalyzed reactions are called central metabolites, such as catechol,

gentisate, protocatechate, or their derivatives. Peripheral enzymes, thus

assumes much significance as regards the degradation of a variety of

xenobiotics. Some such important enzymes are:

2.14.1 Peripheral Enzymes

Main enzymes of this group are:

2.14.1.1 Aromatic Ring-Oxygenases

These enzymes add molecular dioxygen into the aromatic ring and

need cofactors such as NADH, NADPH during this process. These

dioxygenases play a significant role in the bacterial catabolism of naturally

occurring and xenobiotic compounds. By catalyzing the incorporation of two

hydroxyl groups into the aromatic ring, dioxygenases increase the reactivity of

these compounds, making them susceptible to enzymatic ring fission

reactions. A number of highly chlorinated compounds (including numerous

polychlorinated biphenyl congeners) are resistant to aerobic biodegradation

because of the inability of bacterial dioxygenases to accept them as

substrates. Therefore, it is important to develop a greater understanding of

dioxygenase structure and to identify the factors that influence congener


50

specificity. Depending on the structure and components, these enzymes can

be divided into following subgroups, viz.

2.14.1.1.1 Multicomponent Dioxygenase

The enzyme complex is generally formed from three different

components, a terminal oxygenase (also called iron-sulfur protein or

hydroxylase protein), which consists of two different subunits ( and β), a

ferredoxin, and an NADPH-ferredoxin reductase. These multicomponent

proteins form short electron transport chains with flavins and iron-sulfur

clusters as redox components (Fig. 2.9). The initial component of the chain is

a flavoprotein that oxidizes reduced pyridine nucleotides and passes the

electrons to the terminal dioxygenases via a ferredoxin electron carrier. The

ferredoxin and dioxygenase contained [2Fe-2S] redox centers also known as

Rieske type iron-sulfur center, which is either associated with the oxygenase

itself or as part of a small electron transport protein and is involved in electron

transport. The latter protein contains the active site for the incorporation of

oxygen into the aromatic substrate (Hoefer et al., 1993). In contrast to the

plant-type ferredoxins, which have four symmetrically placed cystein sulfurs

coordinating to the [2Fe-2S] core of the center, recent studies have

unequivocally established that two of the ligands to the Rieske [2Fe-2S]

center of these dioxygenases were histidine nitrogens, which coordinate to

the ferrous ion site of the spin-coupled [Fe2+(S=2), Fe3+(S=5\2)] pair of the

reduced cluster (Gurbiel et al., 1989). Highly conserved cystein-histidine pairs

separated by 16 or 17 amino acids were present in Rieske proteins of the

biphenyl and toluene dioxygenases. On the basis of the comparison of the

deduced amino acid (AA) sequences of PCB degradation enzymes BphB,

BphC, and BphD, the dioxygenases of PCB have revealed the presence of

catalytically important amino acid residues and the functions of such residues

were also studied (Gurbiel et al., 1989). In them, the acidic amino acid at

position 18 (or19) C-terminal to the invariant Gly19 (of short-chain alcohol

dehydrogenases) is not absolutely required for their functions in different

enzymes. Among the biphenyl degrading strains of Pseudomonas spp.,

LB400 and KF707, the biphenyl dioxygenases exhibit dramatic differences in


51

PCB substrate range despite nearly identical amino acid sequences (Haddock

et al., 1993). This is mainly because of the amino acid differences within a

141 amino acid region in the large subunit of the terminal dioxygenase. The

remainder of the proteins are essentially identical, lacking even silent

nucleotide changes in their sequences. This implies that such proteins have

diverged recently and as such the difference in amino acid is prevalent in only

a small region. In a recent report, Hugo et al., (2000) have characterized the

xylT gene product, a component of the xylene catabolic pathway of

Pseudomonas putida mt-2, as a novel [2Fe-2S] ferredoxin that specifically

reactivates oxygen-reactivated catechol 2, 3 dioxygenase (XylE). Their study

provides evidence for a subgroup of [2Fe-2S] ferredoxins with distinct

biochemical properties whose specific function is to reactivate intrinsically

labile extradiol ring cleavage dioxygenases involved in the catabolism of

various aromatic hydrocarbons (Hugo et al., 2000).


52

Fig. 2.9: Catabolic enzymes designated as peripheral or upper

pathway enzymes with their basic structure


53

The common examples of this group of dioxygenases are the benzene

dioxygenase from P. putida, benzoate dioxygenase from P. putida, toluene

dioxygenase from P. putida F1, biphenyl dioxygenase from P.

pseudoalcaligenes, and from Pseudomonas spp., strain LB400 (Haddock et

al., 1993; Erickson and Mondello 1992; Zylstra et al., 1988). The three-

component dioxygenases, which have not been characterized on the DNA

sequence level, include naphthalene dioxygenase from plasmid NAH7,

biphenyl dioxygenase from P. paucimobilis Q1, and chlorobenzene

dioxygenases from Pseudomonas spp., strain P51 (Yen and Serdar 1988;

Taira et al., 1988; Erickson and Mondello 1992; Zylstra et al., 1988). There

are also two-component di-oxygenases, such as toluate dioxygenase from P.

putida, benzoate dioxygenase from Acinetobacter calcoaceticus, in which the

electron transfer function is fulfilled by a single protein possessing a

ferredoxin-like structure in its C-terminal region (Van der Meer et al., 1991).

Fetzner et al., (1992) have isolated a novel two-component 2-halobenzoate 1,

2 dioxygenase from Pseudomonas cepacia 2CBS, which has activity toward

ortho substituents of chlorobenzoates. Another two component dioxygenase,

4-chlorophenylacetate 3,4 dioxygenase from Pseudomonas spp., strain

CBS3, shows dehalogenation activity. These enzymes are members of the

short chain alcohol dehydrogenase family (Neidle et al., 1991).

Of all multicomponent dioxygenases characterized so far, toluene

dioxygenase has been found to be the most versatile and hence the best-

studied catabolic enzyme. It has the remarkable power of catalyzing a wide

range of substrates and produces optically pure hydroxylated products

(Zylstra and Gibson 1991). Toluene dioxygenase can also function as

monooxygenase when it oxidizes the benzylic carbon atom of indan to yield (-

)1(R)-indanol and indene to (-)-cis- (1S,2R)-dyhydroxyindan and (+)-(1S)-

indenol 50. Toluene dioxygenase has been critical in the degradation of

trichloroethylene also (Furukawa et al., 1993; Bellard et al., 1983).


54

2.14.1.1.2 Multicomponent Monooxygenase

Phenol hydroxylase is a multicomponent monooxygenase that

transforms phenol to catechol. Some dioxygenases such as toluene

dioxygenase also function as monooxygenases. A few dioxygenases, like that

of toluene and naphthalene, have significant homology in the amino acid

sequences of the ferredoxin components with the toluene 4-monooxygenase

from P. mendocina KR1 (Yen et al., 1991) while other of its components do

not show any similarity to the dioxygenases. However, similarity exists in the

three components of toluene 4-monooxygenase to phenol hydroxylase from

Pseudomonas spp., strain CF600 (Nordlund et al., 1993; Powlowski and

Shingler 1990). These multicomponent monooxygenases are structurally

related to methane monooxygenases (Powlowski and Shingler 1990). The

existence of six polypeptides has been found to be involved in the activity of

hydroxylase in the initial conversion of phenol into catechol in the

Pseudomonas spp., strain CF 600 (Powlowski and Shingler 1990). However,

only the five polypeptide products have been found to be required for in vitro

activity of this multicomponent enzyme (Ballou 1982). The multicomponent

nature of phenol hydroxylase has been intriguing, because in general, mono-

hydroxylated ring structures such as phenol are oxygenated by single

component flavoprotein monooxygenases (Ballou 1982; Bertoni et al., 1998;

Weijer et al., 1982).

2.14.1.1.3 Single Component Monooxygenase

Various single component hydroxylases and monooxygenases have

been reported and were found to share conserved domains. Salicylate

hydroxylase NahG, encoded on the NAH7 plasmid, was shown to be 25%

homologous in amino acid sequence to p-hydroxybenzoate hydroxylase from

P. fluorescens. (Weijer et al., 1982). Similarities have also been reported in

the salicylate hydroxylase and phenol hydroxylase (Kivisaar et al., 1991).

2.14.2 Dehalogenase

These key enzymes catalyze dehalogenation of aromatic hydrocarbons

by cleaving the carbon-halogen bond. The haloacid dehalogenases differ with


55

respect to their substrate specificities, electrophoretic mobilities, and inhibition

by sulfhydryl-blocking agents (Weightman et al., 1982). Based on substrate

range, reaction type, and gene sequences, the dehalogenating enzymes are

classified in different groups, including hydrolytic dehalogenases, glutathione

transferases, monooxygenases, and hydratases (Janssen et al., 1994). A

hydration type of dehalogenation reaction has been proposed for aromatic

compounds and aliphatic acrylic acids (Babbitt et al., 1992; Chang et al.,

1992; Scholten et al., 1991; Hartmans et al., 1991; Vlieg JET and Janssen

1992). The best evidences of hydratase-type reaction comes from the studies

of 4-chlorobenzoate degradation studies of Pseudomonas CBS3 and an

Arthrobacter 4CB1 (Müller et al., 1984; Crooks and Copley 1994) 4-

Chlorobenzoyl coenzyme A dehalogenase from Arthrobacter spp., strain

4CB1 (previously known as Acinetobacter spp., strain 4CB1), which is a

bacterium isolated from PCB-containing soil, was found to be a homotetramer

consisting of 33 kDa subunits with an isoelectric point of 6.1 (Crooks and

Copley 1994; Perkins et al., 1990). The enzyme is able to dehalogenate

substrates bearing fluorine, chlorine, bromine, and iodine in the 4-position,

although the rate of dehalogenation of 4-fluorobenzoyl CoA is quite slow

(Perkins et al., 1990). While three polypeptides with sizes of 57, 30 and 16

kDa were investigated to consist of the 4-chlorobenzoate dehalogenase

activity in Pseudomonas spp., strain CBS3 (Scholten et al., 1991). This

activity was proposed to be the sum of the individual activities of a 4-

chlorobenzoate: CoA ligase, a chlorobenzoate: CoA dehalogenase existing as

a heterodimer of 57- and 30-kDa components, respectively, and a 16-kDa 4-

hydroxybenzoate:CoA thioesterase (Scholten et al., 1991). Some

oxygenases, such as 2-chlorobenzoate and 4-chlorophenoxyacetate

dioxygenases and pentachlorophenol monooxygenase, have also been

implicated in the dehalogenation of their substrates. (Van der Meer et al.,

1991; Crooks and Copley 1993). The dehalogenation reaction is believed to

be a nucleophilic aromatic substitution in which chloride substituent is

replaced by a hydroxyl group derived from water (Perkins et al., 1990). An

unusual enzymic dehalogenation reaction, these are intrinsically difficult

reactions and take place in nonenzymic systems only under special


56

circumstances or very extreme conditions (Weightman and Slater 1980).

Dehalogenases with different substrate specificity and thermal stabilities were

isolated from Pseudomonas putida PP3 and were found to be distantly related

to each other, which is in contrast to the original contention about it (Nagata et

al., 1993). A dehydrochlorinase activity functionally in the dehalogenation of -

hexachlorocyclohexane (BHC) to 1,2,4-trichlorobenzene via -

pentachlorocyclohexene was isolated from P. paucimobilis UT26. Degradation

assays of halogenated compounds by purified dechlorinase (linA) showed that

the substrate specificity of linA is very narrow. Another dehalogenase,

encoded by linB, has been found to show similarities to hydrolytic

dehalogenase, dhlA, when their amino acid sequences were deduced and

were compared (Persson et al., 1991; Neidle et al., 1992). 1-chlorobutane

(C4), 2-chlorobutane, and 1-chlorodecane (C10) have been found to be the

substrates for this dehalogenase (linB), which suggested that this

dehalogenase may be a member of the haloalkane dehalogenase family with

broadrange specificity for substrates (Persson et al., 1991), playing a key role

in the hydrolytic dehalogenations of halogenated aliphatic compounds (Van

der Meer et al., 1991).

2.14.3 Dehydrogenase

Dehydrogenases are members of shortchain alcohol family, which

have their N-terminal similar to known adenosinetriphosphate-binding motifs

of NAD+-binding domains (Wierenga et al., 1986). On the basis of the known

three-dimensional structures of five proteins out of the 15 or 20 family

member dehydrogenases containing such motifs, an anionic side-chain close

to the C-terminal end of ß--ß fold of dehydrogenases has been suggested.

This anionic side chain functions as a hydrogen bond acceptor for 2'-OH

group of adenosine moiety of NAD+, but acts unfavourably with the 2'-

phosphate group of NADP+ (Irie et al., 1987). Gene sequence homology is

found to be nearly 96 or 99% in the dehydrogenase coding regions of different

PCB-degrading strains for which genetic sequence data are available. The

substrates of BphB, TodD, BnzE, and EntA, dihydrodiol dehydrogenases of

biphenyls, toluene, benzene, and benzoate degradation, respectively, differ


57

only in their substituents at one of their carbon atoms next to the dihydrodiol

carbons (Irie et al., 1987; Neidle et al., 1991). Based on sequence

comparison, polychlorinated biphenyl (PCB) degrading dihydrodiol

dehydrogenase share about 61% amino acid with Tod (toluene dihydrodiol

dehydrogenase) and BnzE (benzene dihydrodiol dehydrogenase),while only

28% with EntA (benzoate dihydrodiol dehydrogenase). BenD of Acinetobacter

calcoaceticus and XylL of P. putida, two dehydrogenases acting on the

product of 1,2-dihydroxylation of benzoate and sharing 58% identical amino

acid between them, were found to be about equally related to BphB and EntA

(28 to 32%) (Neidle et al., 1991; Zylstra et al., 1988; Irie et al., 1987; Nakai et

al., 1983; Franklin et al., 1981).

2.15 Aromatic Ring-Cleavage Dioxygenase

These enzymes incorporate two atoms of dioxygen into aromatic

substrates, and aromatic ring is cleaved. This reaction does not require any

external reductant, such as NAD or NADPH or others. Based on the cleavage

of the aromatic ring, they are classified into two types as follows.

2.15.1. Extradiol Enzymes

The extradiol ring-cleavage dioxygenases (EDOs) seem to form a

superfamily of enzymes that catalyze meta cleavage of catechols. The best-

characterized EDO is catechol 2, 3,-dioxygenase ( C23O), encoded by xylE

gene (Ghosal et al., 1987) which is located on TOL plasmid, PWWO (Ghosal

et al., 1987). This enzyme consits of four identical subunits of 32 kDa and

contains one catalytically essential Fe(II) ion per subunit (Harayama and

Rekik 1989; Kimbara et al., 1989). The substrate range of this enzyme is

relatively broad: this enzyme oxidizes 3-methyl, 3-ethyl, 4-methyl, and 4-

chlorocatechol.3-chloro and 4-ethycatechol, in contrast, are not efficiently

oxidized by this enzyme. Other dioxygenases of this superfamily include

catechol 2,3-dioxygenase,encoded by the nahH gene (Ghosal et al., 1987) on

a NAH7 plasmid, 1,2 dihydroxynaphthalene dioxygenase encoded by nahC

gene on a NAH7 plasmid (Ghosal et al., 1987) and 2,3-dihydroxybi-phenyl

dioxygenase (BphCs) from four different Pseudomonas strains, such as P.


58

pseudoalcaligenes KF 707, P. putida KF 715, P. paucimobilis Q1, and

Pseudomonas spp., strain KKS102 (Kimbara et al., 1989). The EDOs derived

from seven different Pseudomonas strains were expressed in Escherichia

coli, and the substrate specificities were investigated for a variety of catecholic

compounds. The dioxygenases from Pseudomonas pseudoalcaligenes KF707

showed a very narrow substrate range, while the dioxygenase from pWWO

showed a relaxed substrate range. Thus, the seven EDOs from various

Pseudomonas strains are highly diversified in terms of substrate specificity

(Hirose et al., 1994). Catechol 2,3-dioxygenase from A. eutrophus has a

primary sequence quite different from other C23Os of this superfamily. In a

recently sequenced trihydroxybiphenyl extradiol dioxygenase, all the six

candidates viz. One Tyr, one Glu, and four His for Fe2+ coordination were

conserved.

2.15.1.1 Protocatechuate 4, 5-Dioxygenase

This EDO catalyzes extradiol cleavage of protocatechuate. The

enzyme consists of an equal number of two different subunits, and ß, 18

and 34 kDa, respectively, and its quaternary structure may be (ß)2 Fe2

(Arciero and Lipscomb 1986). The amino acid sequences of the subunits of

protocatechuate 4,5-dioxygenase differ from C2, 3O. However, the ß-subunit

of this enzyme resembles that of A. eutrophus C2, 3O (Kimbara et al., 1989).

Investigation of the Fe2+ environment of this enzyme from C. testosterone

using EPR spectroscopy revealed that electron delocalization in the ternary

complex, enzyme-Fe (II)-O-O, of a hypothetical reaction sequence is

assumed to polarize dioxygen, thus preparing the distal oxygen atom for

nucleophilic attack on the aromatic ring of the substrates. The iron peroxy-

substrate intermediate, enzyme-Fe (II)-O-O-S, thus produced initiates a

sequence of reaction resulting in the ring fission of the substrate (Arciero and

Lipscomb 1986). Homoprotocatechuate dioxygenase is another class of EDO.

Its amino acid sequence indicates that it constitutes a discrete class among

EDOs (Roper and Cooper 1990).


59

2.15.1.2. Intradiol Enzymes

This group of enzymes consists of catechol 1,2 dioxygenases and

protocatechuate 3,4 dioxygenases. Both these enzymes contain Fe2+ as

cofactors and contain a nonheme, noniron sulfur Fe3+ as a prosthetic group

(Ghosal et al., 1987; Hirose et al., 1994). Usually, C1, 2O from many bacteria

consist of nonidentical - and ß-subunits, (ß-Fe3+), whereas in some

bacterial strains C1,2O consist of a single polypeptide chain (-Fe3+). Nakai

et al., 1983 have reported a Pseudomonas species, which produces two types

of C1, 2O polypeptide subunits, and ß, resulting in the presence of three

isozymes, , ß, and ßß, in the same bacterium. Chlorocatechol 1, 2-

dioxygenase (Clc-C12O) is another class of intradiol enzyme, characterized

by broad substrate specificity.It degrades both catechol as well as

chlorocatechol, while C12O is not able to catalyze chlorocatechols (Van der

Meer et al., 1992).

2.15.1.2.1 Protocatechuate 3, 4 Dioxygenase

This enzyme catalyzes ortho cleavage of protocatechuate to yield ß-

cis-cis-muconate and has been characterized in a number of microorganisms,

including various Pseudomonas strains, Acinetobacter calcoaceticus,

Nocardia spp., Etc (Sterjiades and Pelmont 1989). Protocatechuate

dioxygenases (Pcases) thus far characterized contain equal number of two

different subunits, and ß, and form different quaternary structures of (ß)n (n

= 3-12) (Kimbara et al., 1989). Two alternative forms of PCase have also

been found in Moraxella spp., which were induced as a result of growth of the

bacterium on two different compounds, such as protocatechuate and guaicol

or other phenolic compounds (Sterjiades and Pelmont 1989). However, the

basic structures of such PCases are similar. The similarities in the primary

sequences of C12O and in the and ß subunits of protocatechuate 3, 4-

dioxygenases indicate their origin from a common ancestor (Dercora et al.,

1999).


60

2.15.1.2.2 Gentisate 1, 2 Dioxygenase

Many xenobiotic compounds, such as hydroxybenzoates, salicylates,

naphthalene disulfonate, etc. are degraded through the formation of gentisate

and the enzyme catalyzing ring-cleavage is gentisate 1,2-dioxygenase.

Enzyme, purified from Pseudomonas acidovorans and C. testosteroni,

consists of a single polypeptide of about 40 kDa with a quaternary structure of

(Fe2+). This enzyme contains Fe2+ as cofactor (Harpel and Lipscomb 1990).

Recently, gentisate 1, 2 dioxygenase purified and characterized from two

different species of Pseudomonas reveal significant differences in the first 23

amino acid residues. However, both of these exhibited wide substrate

specificity toward alkyl and halogenated gentisate analogues (Feng et al.,

1999).

2.16 Catabolic genes and their manipulations

The degradation genes of xenobiotics are either located on

chromosomes or on plasmids and sometimes partially on both. These genes

are usually clustered. The plasmid-coded pathways of degradation have

special advantage in that plasmid being a flexible genetic unit that can easily

move into an organism by the natural conjugation process and the entire

population can acquire the trait governed by a plasmid in a reasonable span

of time. Thus, the horizontal gene transfer in a population occurs through

plasmid, and also some new pathways of metabolism can evolve in this

process. However, this is based on the types of genes encoded on plasmids,

their mode of replication, and their ability to promote their own natural transfer

(Hooper et al., 1989) These attributes of a plasmid are referred as the

‘backbone’ of the plasmid (Burlage et al., 1990) Usually homology exists in

the backbone inside or outside the catabolic genes of the degradative

plasmids, for example, homologies exist in the catabolic genes of plasmids

TOL, NAH, CAM, OCT, etc. Burlage et al., (Burlage et al., 1990; Smets et al.,

1993) have described the role of homologous plasmid backbone of pJP4,

pAC24, pSS50, and pBR60 in their replication and transfer functions. Indeed,

direct conjugal transfer of naturally occurring or engineered plasmids has

resulted in the development of bacteria that possess novel biodegradative


61

capabilities. Continuous cocultivations of different organisms having unique

degradative genes on plasmids could lead to the rearrangements of genetic

information within a single organism that manifests new catabolic functions

not shared by or derivable from the separate starter strains (Fernandez-

Astorga et al., 1992).

The kinetic events controlling conjugal plasmid transfer must influence

their contribution to community adaptation and environmental changes.

Factors affecting plasmid transfer in mating experiments like transconjugant

concentration, transconjugant-to-donor ratio, and transconjugant-torecipient

ratio allow comparisons within one study only, because they depend on the

variables like cell densities, population ratios,and period of incubation

(Fernandez-Astorga et al., 1992). Thus, intrinsic parameters describing

plasmid transfer kinetics independent of such factors are needed. The

encounters between plasmid-harboring and plasmid less strains are assumed

to occur at random with a frequency jointly proportional to both population

densities and a fraction of these encounters result in transmission of the

plasmid, thus envisaging a mass action approach (Fernandez-Astorga et al.,

1992; Kinkle et al., 1993). There is considerably less quantitative data on

conjugal transfer of catabolic plasmids, since most of this type of work has

centered around the medically important plasmids (Fernandez-Astorga et al.,

1992; Kinkle et al., 1993). It is only recently that catabolic plasmids have

gained some importance for this study. Once this is studied extensively, some

control strategies for enhancing the dergradative capability of the microbial

community could be worked out. Smets et al. (Fernandez-Astorga et al.,

1992) on the basis of preliminary analysis suggested that the transfer rates of

the TOL plasmid are large enough to maintain it in a dense microbial

population without applying selection pressure. The transfer of plasmid pJP4,

a plasmid coding 2,4-dichloro-phenoxyacetic acid degradation and some

accessory functions between populations of Bradyrhizobia in nonsterile soil

has also been possible (Yen and Gunsalus 1982). It was shown that it could

be transferred to some specific strains only, revealing the fact that this

plasmid is transfer selective to only certain of its hosts. It was found that the

choice of donor microorganism might be a key factor to consider for


62

bioaugmentation efforts. In addition, the establishment of an array of stable

indigenous plasmid hosts at sites with potential for reexposure or long-term

contamination may be particularly useful (Yen and Gunsalus 1982).

TOL is one of the best-studied catabolic plasmids as regards the

genetics and enzymatics of degradation. It bears the degradative genes for

toluene, xylene, and benzoate. The original TOL plasmid, pWWO from

Pseudomonas putida mt-2, is 117 kb in size and belongs to incompatibility

group P-9. Its various derivatives have been found mediating degradation of

compounds like naphthalene, salicylate, catechol, phenol, biphenyl, etc.

(Chatterjee and Chakrabarty 1982; Kivisaar et al., 1990; Lloyd-Jones et al.,

1994; Schmidt 1987). Although TOL predominantly occurs in Pseudomonas

spp., a similar plasmid has been reported in Alcaligenes.( Burlage et al., 1989;

Tsuda and Iino 1988). The basic reason behind the finding of a good number

of TOL derivatives is supposed to be due to the presence of a 56-kb

transposable region on it, which endows transposase genes also (Tsuda et

al., 1989; Romine et al., 1999). About a 40-kb region on plasmid has been

found to be involved in the catabolic functions. Very little is known about the

TOL plasmid, aside from its catabolic region (Burlage et al., 1989). The

location of the genes for replication and conjugal transfer have been mapped

only roughly, and little is known about either process. Recently, a 184-kb

catabolic plasmid has been reported from Sphingomonas aromaticivorans

F199, which has genes for integration and excision events with chromosome

and has many homologous catabolic genes on it (Nurk et al., 1991).

2.16.1 Mechanism of Catabolic Gene Action

An important structural feature of catabolic genes is that they are

generally organized in one or more operon(s), which contribute to the different

reaction(s) for the catabolism of the xenobiotic compound(s). Thus, based on

the number of operons, catabolic genes can be grouped as follows.

2.16.1.1 Single Operon Genes

This group includes such catabolic genes that possess only one

operon. Examples to this group are phenol, biphenyls, etc. (Fig. 2.10). The


63

degradation in case of both of these compounds have been reported to be

mediated by plasmids and chromosomes, although in case of phenol,

degradation genes have predominantly been located on plasmids (Shingler et

al., 1992; Herrmann et al., 1995; Khan and Walia 1990). However, in case of

the latter compound, that is, biphenyls, very few plasmids, such as pSS50,

pWW100, and few others, have been reported, and most of the studies

pertaining to its degradative genes are based on the reports on chromosomal

genes (Kosono et al., 1997). The location of three out of seven genes

involved in biphenyl degradation were found to occur on plasmids in

Rhodococcus erythropolis TA 421 (Assinder and Williams 1990). The

degradation of phenol has been suggested to be governed by multiplasmid

system in Pseudomonas spp., strain EST1001, while in P. putida strain H and

Pseudomonas spp., strain CF600, by plasmid pPGH1 and pSVI, respectively

(Herrmann et al., 1995; Khan and Walia 1990). Because phenol has been

found to be degraded by catechol formation, there exist two mechanisms of

catechol ring fission, that is, ortho and meta. The genetics of ortho pathway of

phenol degradation is little understood. The plasmid-mediated genes of ortho

pathway were found as pheA- and pheB encoding phenol monooxygenase

and catecholdioxygenase, respectively. These two genes have been

sequenced and have been found to possess similarities with the catabolic

operons of chlorocatechol (Clc), catechol (cat), pJP4 genes tfdA and tfdB

(Lloyd-Jones et al., 1994; Shingler et al., 1992). The plasmid isolated by

Herrmann et al., (1995) pPGH1 from P. putida strain H encodes the

degradation of phenol and also some of the methylated derivatives through

the meta pathway. The catabolic genes for the complete degradation span

about 16 kb and consist of a single operon in P. putida strain H. Contrary to

these reports, the degradative genes of phenol have been supposed to be

located on chromosomes, but this has yet to be ascertained. The operons of

phenol and biphenyls have been named as phe and bph, respectively.

However, Khan and Walia (1990) designated the biphenyl operons as cbp.

They even doubt the single operon organization of biphenyl genes, because

(1) the structural genes, cbpCD, alone consisted of an independent operon,

and (2) two types of cbpC genes have been found to exist, which specified


64

two different enzymes, one with a broad substrate range and another with

narrow substrate specificity. This observation is corroborated by the finding

that four groups exist within the PCB-degrading bacteria. Based on the

genetic and immunological characteristics, PCB degraders have been

classified into four groups (Taira et al., 1988). These groups are first,

possessing the bph operon very similar to that of P. pseudoalcaligenes KF

707, the second, having homologous bph operon, but different restriction map

profiles from the operon of KF 707; and the third, group showing weak

homology with KF 707 bph operon; the fourth group, including P. paucimobilis

Q1 showing no genetic or immunological homologies with the KF 707 operon.

The presence of an extra 3 kb DNA lying 30 kb downstream of the bphC

gene, termed bphX, in the strain KF 707 supports this notion (Khan and Walia

1990).


65

Fig. 2.10: Structural genes degrading few pesticides


66

2.16.1.2 Double Operon Genes

This group comprises such catabolic genes that possess two operons

for their complete degradative functions. The structural genes falling in this

category include the genes for xylene/toluene, naphthalene, phenanathrene,

1,2,4-trichlorobenzene degradation (Fig. 2.10). The most versatile gene of this

group has been found to be located on plasmid TOL, which is invariably the

most thoroughly characterized catabolic plasmid. This plasmid has been

reported from a number of Pseudomonas strains degrading a number of

compounds, such as biphenyls, (Lloyd-Jones, et al., 1994), phenol (Kivisaar

et al., 1990) and naphthalene (Yen and Gunsalus 1982). The two operons

encoded by the TOL plasmid, which was discovered from the Pseudomonas

strain degrading xylene and toluene (Assinder and Williams 1990) are

designated as the upper and lower operons (Assinder and Williams 1990).

These two operons are responsible for the degradation of xylene/toluene to

benzoate and later into the intermediates of the Kreb’s cycle. The degradation

of toluene/xylene to benzoate/toluate respectively, is encoded by xylCAB,

while the lower pathway starting from benzoate or toluate and culminating into

acetaldehyde and pyruvate is encoded by genes xylDLEGFJKIH (Assinder

and Williams 1990).

2.16.1.3 Multiple Operon Genes

There are few examples of catabolic genes that have been organized

into more than two operons (Fig. 2.11). Of which three operon organizations

have been found for the genes of dinitrotoluene dioxygenase, 2,4-D

degradation in Alcaligenes eutrophus JMP 134 and benzoate degradation

from A. calcoaceticus (Suen and Spain 1993; Harayama et al., 1986; Harker

et al., 1989). The 2,4 D degradation has been demonstrated to occur due to

the presence of the plasmid pJP4 in Alcaligenes, for which even chromosomal

genes are essential. The catabolic genes of plasmid pJP4 have been mapped

by transposon mutagenesis (Don et al., 1985). The plasmid pJP4, isolated

from A. eutrophus JMP 134, is 80 kb size and has a broad host range.

Several restiction maps of this and other similar plasmids have been


67

published (Don and Pemberton 1981; 1985; Ghosal et al., 1985a). Plasmid

pJP4 carries the gene essential for the degradation of 3-CBA, mercury

resistance as well as 2,4-dichlorophenoxy acetate (2,4-D) degradation (Don

and Pemberton 1985). The metabolic pathways for 3-CBA and 2,4-D

degradation utilize enzymes common to the degradation of chlorocatechol to

chloromaleyl acetate (Dorn and Knackmuss 1978; Evans et al., 1971; Kukor

et al., 1989). The gene clusters involved are tfdA and tfdB encoding TFD

monooxygenase (32 kDa) and 2, 4 dichlorophenol (TFP) hydroxylase (65

kDa), respectively, and convert chlorophenoxyacetate to chlorocatechol (Dorn

and Knackmuss 1978). Genes for the degradation of 2,4-D to 2-chloromaleyl

acetate are plasmid pJP4 mediated and have been sequenced, while genes

encoding degradation of 2-chloromaleyl acetate are located on the

chromosomes (Dorn and Knackmuss 1978). The ability or inability of a

microorganism possessing initial catabolic genes to degrade 2,4-D completely

depends on the presence of the complementary enzymes encoded by

chromosomal genes (Kukor et al., 1989).

2.16.1.4 Transposon-Mediated Genes

The existence of catabolic genes on transposons has been known

since the discovery of the ability of the TOL operons to be mobile. In fact, this

transposon dependent mobility was accounted to be present due to a Tn4652

mobile genetic element on the TOL plasmid. This transposon has been

suggested to belong to the family of Tn-3 transposons (Tsuda et al., 1989;

Romine et al., 1999). Similar mobile regions within the catabolic genes have

been reported later for a number of compounds, such as phenol (Kasak et al.,

1993), chloro-benzoates (Nakatsu et al., 1991), chlorobenzene (Van der Meer

et al., 1991), biphenyls (Springael et al., 1993) and 2,4,5-

trichlorophenoxyacetate (2,4,5 T) (Haughland et al., 1990). Another

transposon bearing the catabolic genes of 3-CBA degradation was known and

was designated Tn5271, which has been regarded as composite class I

element with a flanking region of class II insertion sequences (Nakatsu et al.,

1991). A composite transposon, Tn5280, has been known to act in the

degradation of chlorobenzene, although its origin is still unclear. A. eutrophus


68

strain A5 was reported to possess a 59-kb transposon, on which the complete

set of genes for the conversion of BP and 4CBP were located (Haughland et

al., 1990). Similar genes were also found to be present in some other

bacterial strains (Sylvestre et al., 1985). The degradative genes of a common

herbicide 2,4,5-trichlorophenoxyacetic acid (2,4,5-T) have been investigated

less extensively and most of the information pertaining to this degradation has

been accumulated by using reductive (anaerobic) sediments (Sangodkar et

al., 1988; Kilbane et al., 1982). Little is known about the precise gene location

and function in degradation of 2, 4, 5-T (Chaudhary and Chapalamadugu

1991). Two insertion elements, RS1100, redesignated as IS931 and IS932,

have been reported in the Pseudomonas spp., strain AC1100, which have

roles in the degradation of 2,4,5-T. Amino acid sequence homology with BenA

and XylY from toluate 1,2 dioxygenase of A. calcoaceticus and XylX and XylY

from toluate 1,2 dioxygenase of P. putida also existed in the two catabolic

genes (tftA1 and tftA2) present on transposons mediating the early

transformation of 2,4,5-T (Danganan et al., 1994; Schell 1993).

2.17 Regulation of catabolic gene action

A number of factors have been found to influence the expression of

catabolic genes. These factors include the structure of the genes, enzymes,

substrates, and the metabolites. In fact, the overall interactions of all these

together results in the onset of process of degradation. Usually, degradation

at substrate or metabolite level is supposed to be coordinately regulated,

while at the gene level it is subject to such a control by a set of structural

genes, the products of which are regulatory. These products are proteins and

are designated as LysR family of regulators (Coco et al., 1993; 1994). Several

LysR type of regulators have been found for compounds such as

chlorobenzoates, phenol, benzoate, and others, which consist of single

operon in their catabolic genes (Parsek et al., 1994; Henikoff et al., 1988;

Aldrich et al., 1987). In benzoic acids, the genes catB and catC, encoding the

first two reactions of catechol catabolism after cis-cis muconate fromation in

ortho pathway, are coordinately controlled and have been found to be closely

linked on the chromosome, but catA (gene for C12O) is separated from the


69

catBC operon (Wheelis and Ornston 1972; Rothmel et al., 1990). The

regulatory gene catR maps upstream of catBC operon and it is transcribed

divergently from the operon (Rothmel et al., 1990). The binding of the catR to

the regulatory region that includes its own promoter leads to both auto

regulation and the activation of catBC genes (Rothmel et al., 1990). CatR

binds to the catR-catBC promoter control region in both the presence or

absence of the inducer cis-cis-muconate, but activates the catBC operon only

in the presence of the inducer (Rothmel et al., 1990). Studies have also

demonstrated that cis-cis-muconate allows CatR to bind to another site of the

catBC promoter region, thereby favoring the formation of the transcription

initiation complex of the catBC promoter (Aldrich et al., 1987). Similarly, the

expression of the biodegradation pathway for 3-chlorocatechol in

Pseudomonas putida, which is encoded by the clcABD operon, has been

shown to require the divergently transcribed lysR type regulatory gene clcR

for activation (Parsek et al., 1994) After cloning and sequencing of the clcR

genes, it is revealed that ClcR inducibly activates the clcABD operon and

represses its own transcription (Parsek et al., 1994) Although similarities

among the genes and regulatory proteins of several ortho-cleavage pathway

operons are thought to point to a shared ancestry, the extent of cross-binding

and cross-tack among LysR family members of regulators is yet unclear

(Parsek et al., 1994) Similar controlling element, tfdR, was found to regulate

the synthesis of TFD monooxygenase (tfdA gene product) in case of 2,4-D

degradation in A.eutrophus (Kaphammer et al., 1990) It also regulated the

tfdCDEF operon, and not tfdB, the other operon. Recently in Rhodococcus

opacus 1CP, the presence of CatR regulatory protein resembling members of

the PopR family of IclR type regulatory protein has been found (Eulberg et al.,

1998). However, in the case of toluene/xylene, where two operons encode the

degradative functions, there are two regulatory genes, xylR and xylS for the

upper and lower operons, respectively (Harayama et al., 1986). These

operons are transcribed from physically close but functionally divergent

promoters (Inouye et al., 1983; 1985). Although only a fraction of xylR gene

has been sequenced (Spooner et al., 1986; Mermod et al., 1987) the size of

the xylR protein has been estimated to be 68kDa. XylS protein of 36.5 kDa


70

size has its gene completely sequenced (Mermod et al., 1987; Inouye et al.,

1986; Ramos et al., 1987).

The control of the upper and lower pathways of TOL and of the

regulatory genes is still not fully clear (Kinkle et al., 1993). However,

generalities have been made about the induction of the pathways and roles of

the regulatory proteins. For instance, substrates for the upper pathway

enzymes, such as toluene or m-methyl benzyl alcohol, are activators of the

pathway in the presence of xylR (Burlage et al., 1989; Ramos et al., 1987). In

a similar manner, m-toluate is both a substrate and an inducer for the lower

pathway in conjunction with the xylS gene product. However, upper pathway

substrates can activate the lower pathway if both xylR and xylS are present.

The genetic mechanisms of these inductions have been investigated and

found that the substrate (m-xylene) itself is an inducer of the first operon,

designated as OP1, resulting in the synthesis of the products (m-toluate) in

large amounts. The latter products act as inducers for the other operon,

designated as OP2, by binding to the XylS protein to enhance the activation of

OP2 as an inducer (Inouye et al., 1990; Jeffery et al., 1992; Cowles et al.,

2000).

XylR activated promoter of OP1 and xylS gene share sequence

similarity to the nitrogen- regulated (ntr) and the nitrogen fixation promoters

(Ramos et al., 1987). The ntrA gene of P. putida has been cloned and was

found to be required to be intact for the activation of OP1 and xylS genes

(Jeffery et al., 1992). Jeffery et al., (1992) have reported a substitute for xylS

regulatory gene, which is designated benR. This new regulatory gene,

originally found in a benzoate-degrading Pseudomonas, has been used to

activate the lower pathway operon in some Pseudomonas strains, viz., P.

aeruginosa PAO1 and P. putida mt-2 and PRS 2000. The two originally

different operons, benR and xylS, thus imply evolutionary relationships

between them (Jeffery et al., 1992). Pseudomonas putida converts benzoate

to catechol using two enzymes that are encoded on the chromosome and

whose expression is induced by benzoate. Benzoate also binds to the

regulator XylS to induce expression of the TOL (toluene degradation) plasmid-

encoded meta pathway operon for benzoate and methylbenzoate


71

degradation. Finally, benzoate represses the ability of P. putida to transport 4-

hydroxybenzoate (4-HBA) by preventing transcription of pcaK, the gene

encoding the 4-HBA permease. Cowles et al., (2000) have identified a gene,

benR, as a regulator of benzoate, methylbenzoate, and 4-HBA degradation. A

benR mutant isolated by random transposon mutagenesis was unable to grow

on benzoate. The deduced amino acid sequence of BenR showed high

similarity (62% identity) to the sequence of XylS, a member of the AraC family

of regulators (Jeffery et al., 1992; Cowles et al., 2000). An additional seven

genes located adjacent to benR were inferred to be involved in benzoate

degradation based on their deduced amino acid sequences. The benABC

genes likely encode benzoate dioxygenase and benD likely encodes 2-hydro-

1, 2-dihydroxybenzoate dehydrogenase. BenK and benF were assigned

functions as a benzoate permease and porin, respectively. The possible

function of a final gene, benE, is not known. BenR activated the expression of

a benA-lacZ reporter fusion in response to benzoate. It also activated

expression of a meta cleavage operon promoter-lacZ fusion inserted in an E.

coli chromosome (Cowles et al., 2000). Third, benR was required for

benzoate-mediated repression of pcaK-lacZ fusion expression. The benA

promoter region contains a direct repeat sequence that matches the XylS

binding site previously defined for the meta cleavage operon promoter. It is

likely that BenR binds to the promoter region of chromosomal benzoate

degradation genes and plasmid-encoded methylbenzoate degradation genes

to activate gene expression in response to benzoate. The action of BenR (the

protein encoded by benR gene) in repressing 4-HBA uptake is probably

indirect (Cowles et al., 2000).


72

Fig. 2.11: Regulation of catabolic gene action. The function of catabolic

operons are under the regulation of LysR family of regulators.

Substrates for the upper pathway enzymes are activators of the pathway

in the presence of LysR type of regulators. This substrate is also an

inducer for the lower pathway in conjunction with the upper pathway

gene product


73

2.18 Conclusion

With the reports of enormous pollution of our environment by the

release of synthetic compounds, bacterial degradation for the

decontamination of polluted sites and detoxification of the compounds itself

have assumed significant importance. Numerous bacterial strains are being

isolated and the mechanism of degradation of xenobiotic is being studied. A

general view regarding the pathway of degradation entails that there are only

three options, such as catechol, gentisate, or protocatechuate, available to the

bacterial species for the degradation of concerned compound when it is

subject to such incubation. There are also examples of formation of products

other than these three. For instance, the degradation of benzoate by a new

pathway, that is, benzoyl Co-A, has also been reported, which is an

uncommon pathway of benzoate degradation. Varied mechanisms have been

adopted by the microorganisms for the degradation of organic compounds.

The enzymatic machinery and the genetic system of these organisms, and the

flexibility manifested have been suggested to be largely due to location of

majority of the catabolic genes on the plasmids and transposons. For

example, the catabolic genes of chlorobenzoates, chlorobenzene,

chlorobiphenyls, benzoate, xylene, etc. have been found to reside on

transposons. The presence of two transposons on TOL plasmid is probably

the only fact behind the occurrence of a number of TOL derivatives from

bacterial strains growing on various xenobiotic compounds. As a result, there

exists usually homology in the catabolic genes of various aromatic

compounds. To explore the possibilities of construction of genetically

engineered or altered microorganisms through the applications of random

mutations, a basic understanding of their degradative enzymes, mainly the

peripheral enzymes, catabolic genes, and operons involved in the act, is

necessary. Efforts are on to develop monooxygenases and dioxygenase with

overlapping novel regio- and chemo specificities. Directed enzyme evolution

in combination with recombinant DNA technology is being exploited to

broaden the substrate specificity. Ultimately, bioremediation in all probability

will be carried out at the field level by a cell rather than an individual enzyme.

Thus, novel metabolite pathways have to be engineered in a particular strain.


74

As our understanding of the enzymes and genes involved increases, novel

pathways could be engineered by introducing genes into a particular strain.

Alternatively, a mixture of microbes that can coexist together and carry out

different parts of the pathway separately could also be exploited. Although

several bacterial strains have been isolated on a wide range of aromatic

hydrocarbons, the search for better and better strains is still on. By better, we

mean strains that degrade a compound and its analogues completely.

However, the main limitations in microbial degradation are the incomplete

degradation of the xenobiotic compound, inhibition of the bacterial growth or

catabolic enzymes by intermediates of the pathway, and nonselective

induction of some of pathway enzyme(s) leading to the production of dead-

end metabolites, etc. For a better understanding of the molecular processes

involved in degrading these xenobiotics in their microbes, the microbial

ecology of the contaminated sites are some of the areas that need intensive

investigation. In order to achieve bioremediation, these constraints have to be

overcome. Genetic manipulation of the microorganism by recombinant DNA

technology holds promise. No bacterial strains have yet been released to

degrade the pollutants for the bioremediation of the contaminated field sites

on a large scale. However, several in situ trials have been conducted with a

few bacterial strains, mainly Pseudomonas. The isolation of novel strains and

construction of novel bacterial genotypes for degradation of pollutants are

essential steps for the efficient decontamination of polluted sites. The

availability of data banks on degradative pathways, their enzymes and genes,

and the efforts to predict novel pathways in silico and their subsequent

utilization in combination with recombinant DNA technology would greatly help

in ameliorating environmental pollution.


75

References: Abhilash PC, Jamil S, Singh N, 2009, Transgenic plants for enhanced

biodegradation and phytoremediation of organic xenobiotics,

Biotechnol. Adv., 27: 474-488.

Anonymous, 1993, Facts and Fig.s for the chemical industry, Chem. Eng.

News, 17 (June 28), 38–46.

Aldrich TL, Frantz B, Gill JF, Kilbane JJ, Chakrabarty AM, 1987, Cloning and

complete nucleotide sequence determination of the catB gene

encoding cis-cis-muconate lactonizing enzyme, Gene, 52: 185.

Arciero DM and Lipscomb JD, 1986, Binding of the 17O-labeled substrate and

inhibitors to protocatechuate 4, 5 dioxygenase-nitrosyl complex, J. Biol.

Chem., 261: 2170.

Assinder SJ and Williams PA, 1990, The TOL plasmids: determinants of the

catabolism of toluene and the xylenes, Adv. Microbiol. Phys., 31: 1.

Asturias JA and Timmis KN, 1993, Three different 2,3-dihydroxybiphenyl-1,2-

dioxygenase genes in the Gram-positive polychlorobiphenyl degrading

bacterium Rhodococcus globerulus P6, J. Bacteriol., 175: 4631.

Babbitt PC, Kenyon GL, Martin BM, Charest H, Sylvestre M, 1992, Ancestry

of the 4-chlorobenzoate dehalogenase: analysis of amino acid

sequence identities among families of acyl:adenyl ligases, enoyl-CoA

hydratases/isomerases, and acetyl-CoA thioesterases, Biochemistry

31, 5594.

Ballou D, 1982, Flavoprotein monooxygenases. In: Massey, V. and Williams,

C.H. (Eds.) Flavin and Flavoproteins, New York, Elsevier, North

Holland Publishing, pp. 301.

Barker PS and Morrison FO, 1964, Breakdown of DDT to DDD in mouse

tissue, Canadian Journal of Zoology, 42: 324- 325.

Barker PS, Morrison FO, Whitaker RS, 1965, Conversion of DDT to DDD by

Proteus vulgaris- A bacterium isolated from the intestinal flora of a

mouse, Nature, 205: 621- 622.

Barrio-Lage G, Parsons FZ, Nassar RS, Lorenzo PA, 1986, Sequential

dehalogenation of chlorinated ethenes, Environ. Sci. Technol., 20: 96-

99.


76

Bartnicki EW and Castro CE, 1969, Biodehalogenation: The pathway for

transhalogenation and the stereochemistry of epoxide formation from

halohydrins, Biochemistry, 8: 4677-4680.

Belay N and Daniels L, 1987, Production of ethane, ethylene and acetylene

from halogenated hydrocarbons by methanogenic bacteria, Appl.

Environ. Microbiol., 53: 1604–1610.

Bellard DGH, Curtis A, Shirley A, Taylor SC, 1983, A biotech route to

polyphenylene, J. Chem. Soc. Chem. Commun., 954.

Bertoni G, Martino M, Galli E, Barbieri P, 1998, Analysis of the gene cluster

encoding toluene/o-xylene monooxygenase from Pseudomonas

stutzeri OX1. Appl. Environ.Microbiol. 64: 3626.

Bhaskara B, David SH, Ferguson JF, Magar VS, Puhakka JA, 1998, Toxic

chlorinated compounds: Fate and biodegradation in anaerobic

digestors, Water Environment Research Foundation.

Bielefeldt AR, Stensel HD, Strand SE, 1995, Cometabolic degradation of

TCE and DCE without intermediate toxicity, J. Environ. Eng., 791-

797.

Bouwer EJ and McCarty PL, 1983, Transformations of 1 and 2 carbon

halogenated aliphatic organic compounds under methanogenic

conditions, Appl. Environ. Microbiol., 45: 1286-1294.

Bouwer EJ and McCarty PL, 1985, Ethylene dibromide transformation under

methanogenic conditions, Appl. Environ Microbiol., 50: 527-528.

Bradley PM and Chapelle FH, 1998, Effect of contaminant concentration on

aerobic microbial mineralization of DCE and VC in stream bed

sediment, Environ. Sci. Technol., 32: 553-557.

Bradley PM, Landmeyer JE, Dinicola RS, 1998, Anaerobic oxidation of (1,2-

14C) dichloroethene, Appl. Environ. Technol., 64: 1560-1562.

Brenner V, Hernandez BS, Focht DD, 1993, Variation in chlorobenzoate

catabolism by Pseudomonas putida P111 as a consequence of genetic

alterations, Appl. Environ. Microbiol., 59: 2790–2794.

Brown JF, Bedard DL, Brennan MJ, Carnahan JC, Feng H, Wagner RE, 1987,

Polychlorinated biphenyl dechlorination in aquatic sediments,

Science, 236: 709–712.


77

Brunner W, Staub D, Leisinger T, 1980, Bacterial degradation of

dichloromethane, Appl. Environ. Microbiol., 40: 950–958.

Bumpus JA, Aust SD, 1987, Biodegradation of DDT (1,1,1-trichloro-2,2-bis(4-

chlorophenyl)ethane) by a white rot fungus Phanerochaete

chrysosporium, Applied and Environmental Microbiology, 53: 2001- 08.

Burlage RS, Hooper SW, Sayler GS, 1989, The TOL (pWWO) catabolic

plasmid, Appl. Environ. Microbiol., 55: 1323.

Burlage RS, Bemis LA, Layton AC, Sayler GS, Larimer F, 1990,

Comparative genetic organization of incompatibility group P

degradative plasmids, J. Bacteriol., 172: 6818.

Buser HR, Muller MD, 1995, Isomer and enantioselective degradation of hexa

chlorocyclohexane isomers in sewage sludge under anaerobic

conditions, Environ.Sci. Technol., 29: 664–612.

Cain RB, 1981, Regulation of aromatic and hydroaromatic catabolic pathways

innocardioform actinomycetes. Zentralbl.Bakteriol., Hyg. Abt. 1 Orig. C,

1: 335.

Canet R, Birnstingl JG, Malcolm DG, Lopez-Real JM, Beck AJ, 2001,

Biodegradation of polycyclic aromatic hydrocarbons (PAHs) by native

microflora and combinations of white-rot fungi in a coal-tar

contaminated soil, Bioresour Technol, 76: 113-7.

Carrington B, Lowe A, Shaw LE, Williams PA, 1994, The lower pathway

operon for benzoate catabolism in biphenyl utilizing Pseudomonas

spp., Strain JC and the nucleotide sequence of the bphE gene for

catechol 2,3 dioxygenase, Microbiology, 140: 409.

Castro CE, Bartinicki EW, 1968, Biodegradation, The epoxidation of

halohydrins, epoxide opening and transhalogenation by a

Flavobacterium spp., Biochemistry, 7: 3213–3218.

Castro CE and Belser NO, 1990, Biodehalogenation: Oxidative and reductive

metabolism of 1,1,2-trichloroethane by Pseudomonas putida

biogeneration of vinyl chloride, Environ. Toxicol. Chem., 9: 707–714.

Chacko CL, Lockwood JL, Zabik M, 1966, Chlorinated hydrocarbon pesticides

degradation by microbes, Science, 154: 893- 895.


78

Chang KH, Liang PH, Beck W, Scholten JD, 1992, Isolation and

characterization of the three polypeptide components of 4-

chlorobenzoate dehalogenase from Pseudomonas spp., strain CBS-3.

Biochemistry, 31: 5605.

Chatterjee DK and Chakrabarty AM, 1982, Genetic rearrangements in

plasmids specifying total degradation of chlorinated benzoic acids,

Mol. Gen. Genet., 188: 279.

Chaudhary GR and Chapalamadugu S, 1991, Biodegradation of

halogenated organic compounds, Microbiol. Rev., 55: 59.

Chen C, Puhakka JA, Ferguson JF, 1996, Biotic transformation of

tetrachloroacetylene with 1,1,2-trichloroethane, trans-1,2-

dichloroethene and cis-1,2-dichloroethene as products, Environ. Sci.

Technol., 30: 542–547.

Cobb GD and Bouwer EJ, 1991, Effects of electron acceptors on halogenated

organic compound biotransformations in a biofilm column, Environ. Sci.

Technol., 25: 1068–1074.

Coco WM, Rothmel RK, Henikoff S, Chakrabarty AM, 1993, Nucleotide

sequence and initial functional characterization of the clcR gene

encoding a LysR family activator of the clcABD chlorocatechol

operon in Pseudomonas putida, J. Bacteriol., 175: 417.

Coco WM, Parsek MR, Chakrabarty AM, 1994, Purification of the LysR

family regulator, ClcR, and its interaction with the Pseudomonas putida

clcABD chlorocatechol operon promoter, J. Bacteriol., 176: 5530.

Cole JR, Caserelli AL, Mohn WW, Tiedje JM, 1994, Isolation and

characterization of a novel bacterium growing via reductive

dehalogenation of 2- chlorophenol, Appl. Environ. Microbiol., 60:

3536–3542.

Cook AM and Hutter R, 1986, Ring dechlorination of diethylsimazine by

hydrolases from Rhodococcus corallinus, FEMS. Microbiol. Lett., 34:

335–338.

Cookson JT Jr, 1995, In: Bioremediation Engineering: Design and Application,

New York: McGraw-Hill.


79

Cowles CE, Nichols NN, Harwood C S, 2000, BenR, a XylS homologue,

regulates three different pathways of aromatic acid degradation in

Pseudomonas putida, J. Bacteriol., 182: 6339.

Criddle CS, De Witt JT, McCarty PL, 1990, Reductive dehalogenation of

carbon tetrachloride by Escherichia coli K-12, Appl. Environ. Microbiol.,

56, 3247–3254.

Crooks GP and Copley D, 1993, A surprising effect of leaving group on the

nucleophilic aromatic substitution reaction catalyzed by 4-

chlorobenzoyl-CoA- dehalogenase, J. Am. Chem. Soc., 115: 6422.

Crooks GP and Copley SD, 1994, Purification and characterization of 4-

chlorobenzoyl-CoA dehalogenase from Arthrobacter spp., strain 4-

CB1, Biochemistry, 33: 11645.

Cupples AM, Spormann AM, McCarty PL, 2003, Growth of a

Dehalococcoides like microorganism on VC and cis-dichloroethene as

electron acceptors as determined by competitive PCR, Appl. Environ.

Microbiol., 69: 953–959.

Dagley S, 1975, In: Essays in Biochemistry, Campbell, P.N., and Aldridge,

W.N., eds., London: Academic Press, 11: 81–138.

Dagley S, 1986, Biochemistry of aromatic hydrocarbon degradation. In:

Sokatch, J.R. and Ornston, L.N. (Ed.), The Bacteria, vol. X, Academic

Press, pp. 527.

Danganan CE, Ye RW, Daubaras DL, Xun L, Chakrabarty AM, 1994,

Nucleotide sequence and functional analysis of the genes encoding

2,4,5- trichlorophenoxyacetic acid oxygenase in Pseudomonas

cepacia AC 1100, Appl. Environ. Microbiol., 60: 4100.

De Bont JAM, Vorage MJAW, Hartmans S, Van den Tweel WJJ, 1986,

Microbial degradation of 1,3-dichlorobenzene, Appl. Environ.

Microbiol., 52: 677–680.

De Bruin WP, Kotterman MJJ, Posthumus MA, Schraa G, Zehnder AJB,

1995, Complete biological reductive transformation of

tetrachloroethene to ethane, Appl. Environ. Micrbiol., 58: 1996–2000.


80

De Jong E, Field JA, Spinnler HE, Wijnberg JBPA, de Bont JNM, 1994,

Significant biogenesis of chlorinated aromatics by fungi in natural

environments, Appl. Environ. Microbiol., 60: 264–270.

Demir G, 2004, Degradation of toluene and benzene by Trametes versicolor.

J Environ Biol, 25: 19-25.

Dercora K Vrb, Balaz S, Klasinc L,Peijnenburg WJGM, Sabljic A, Slanina J,

1999, Degradation process in the environment IV Pac int.

Conference. Pubronic, Goatia, 24–28 May 1998, Chemophore 38:

1391.

De Weerd KA, Sulfita JM, 1990, Anaerobic aryl reductive dehalogenation of

halobenzoates by cell extracts of Desulfomonile tiedje, Appl. Environ.

Microbiol., 56: 2999–3005.

Di Stefano TD, Gosett JM, Zinder SH, 1991, Reductive dechlorination of

high concentration of tetrachloroethene to ethane by an anaerobic

enrichment culture in the absence of methanogenisis, Appl. Environ.

Microbiol., 57: 2287– 2292.

Dolfing J, 1990, Reductive dechlorination of 3-chlorobenzoate is coupled to

ATP production and growth in an anaerobic bacterium, strain DCB-1,

Arch. Microbiol., 153: 264–266.

Dolfing J, Harrison BK, 1992, Gibbs free energy of formation of halogenated

aromatic compounds and their potential role as electron acceptors in

anaerobic in environments, Environ. Sci. Technol., 26: 2213–2218.

Don RH and Pemberton JM, 1981, Properties of six pesticide degrading

plasmids isolated from Alcaligenes paradoxus and Alcaligenes

eutrophus, J. Bacteriol.,145: 681.

Don RH, Weightman AJ, Knackmuss HJ, Timmis KN, 1985, Transposon

mutagenesis and cloning analysis of the pathways for degradation of

2,4-dichlorophenoxyacetic acid and 3-chlorobenzoate in

Alcaligenes eutrophus JMP134 (pJP4), J.Bacteriol., 161: 85.

Dorn E and Knackmuss HJ, 1978, Chemical structure and biodegradability of

halogenated aromatic compounds. Two catechol 1,2-dioxygenases

from a 3-chlorobenzoate-grown Pseudomonad, Biochem. J., 174: 73.


81

Dybas MJ, Tatara GM, Criddle CS, 1995, Localization and characterization

of the carbon tetrachloride transformation activity of Pseudomonas

spp., Strain KC, Appl. Environ. Microbiol., 61, 758–762.

Egli C, Tschan T, Scholtz R, Cook AM, Leisinger T, 1988, Transformation of

Tetrachloromethane to dichloromethane and CO2 by Acetobacterium

woodii, Appl. Environ. Microbiol., 54: 2819–2824.

Egli C, Thuer M, Suter D, Cook AM, Leisingen T, 1989, Monochloro- and

dichloroacetic acids and carbon and energy sources for a stable,

methanogenic mixed culture, Arch. Microbial., 152: 218–223.

Egli C, Stromeyer S, Cook AM, Leisinger T, 1990, Transformation of tetra-

and trichloromethane to carbon dioxide by anaerobic bacteria is a non-

enzymic process, FEMS. Microbiol. Lett., 68: 207–212.

Engvild KC, 1986, Chlorine-containing natural compounds in higher plants,

Phytochemistry, 25: 781–791.

Ensign SA, Hyman MR, Arp DJ, 1968, Cometabolic degradation of

chlorinated alkenes by alkene monoxygenase in a propylene grown

Xanthobacter strain, Appl. Environ. Microbiol., 53: 3038–3046.

Ensley BD, 1991, Biochemical diversity of TCE methabolism, Annu. Rev.

Microbiol., 45: 283–299.

Erickson BD and Mondello FJ, 1992, Nucleotide sequencing and

transcriptional mapping of the genes encoding biphenyl

dioxygenase, a multicomponent polychlorinated biphenyldegrading

enzyme in Pseudomonas strain LB 400, J. Bacteriol., 174: 2903.

Esposito E, Paulillo SM, Manfio GP, 1998, Biodegradation of the herbicide

diuron in soil by indigenous actinomycetes. Chemosphere, 37: 541-8.

Eulberg D, Schlomann M, Alderson G, 1998, in M. Goodfellow (Ed.) Biology of

Exploitable Bacteria in the Genus Rhodococcus. Antonio van

Leeuwenhoek. 74: 71.

Evans WC, Smith BSW, Fernley HN, Davies JI, 1971, Bacterial

metabolism of 2,4-dichlorophenoxyacetate, Biochem. J, 122: 543.

Fathepure BZ, Nengu JP, Boyd SA, 1987, Anaerobic bacteria that

dechlorinate perchloroethene, Appl. Environ. Microbiol., 53: 2671–

2674.


82

Fathepure BZ, Tiedje JM, Boyd SA, 1988, Reductive dechlorination of

hexachlorobenzene to tri and dichlorobenzenes in anaerobic sewage

sludge, Appl. Environ. Microbiol., 54: 327–330.

Fathepure BZ and Boyd SA, 1988, Dependence of tetrachloroethylene

dechlorination on methanogenic substrate consumption by

Methanosarcina sp strain DCM, Appl. Environ. Microbiol., 54: 2979–

2980.

Feng V, Khoo HE, Poh CL, 1999, Purification and Characterization of

Gentisate 1,2-Dioxygenases from Pseudomonas alcaligenes NCIB

9867 and Pseudomonas putida NCIB 9869, Appl. Environ. Microbiol.,

165: 946.

Fernandez-Astorga A, Muela A, Cisterna R, Iriberri J, Barcina I, 1992, Biotic

and abiotic factors affecting plasmid transfer in Escherichia coli strains,

Appl. Environ. Microbiol., 58: 392.

Fetzner S, Muller, R., and Lingens, F.Purification and some properties of 2-

halobenzoate-1,2-dioxygenase, a two component enzyme system from

Pseudomonas cepacia 2CBS. J. Bacteriol. 174, 279, 1992.

Fetzner S and Lingens F, 1994, Bacterial dehalogenases: Biochemistry,

genetics and biotechnological applications, Microbiol. Rev., 58:

641–685.

Fetzner S, 1998, Bacterial dehalogenation, Appl. Microbiol. Biotechnol., 50:

633–657.

Focht DD and Alexander M, 1970, DDT metabolites and analogues: ring

fission by Hydrogenomonas, Science, 170: 91- 92.

Folsom BR, Chapman PJ, Pritchard PH, 1990, Phenol and trichloroethylene

degradation by Pseudomonas cepacia G4: Kinetics and interaction

between substrates, Appl. Environ. Microbiol., 56: 1279–1285.

Fox BG, Borneman JG, Wackett LP, Lipscomb JD, 1990, Haloalkene

oxidation by the soluble methane monooxygenase from Methylosinus

trichosporium OB3b: Mechanistic and environmental implications,

Biochemistry, 29: 6419–6427.

Fragoeiro S and Magan N, 2005, Enzymatic activity, osmotic stress and

degradation of pesticide mixtures in soil extract liquid broth inoculated


83

with Phanerochaete chrysosporium and Trametes versicolor, Environ

Microbiol., 7: 348-55.

Fragoeiro S and Magan N, 2008, Impact of Trametes versicolor and

Phanerochaete crysosporium on differential breakdown of pesticide

mixtures in soil microcosms at two water potentials and associated

respiration and enzyme activity, Int Biodeterior Biodegrad., 62: 376-83.

Franken SM, Rozeboom HJ, Kalk KH, Dijkstra BW, 1991, Crystal structure

of haloalkane dehalogenase an enzyme to detoxify halogenated

alkanes, EMBO J., 10: 1297–1302.

Franklin FCH, Bagdasarian M, Bagdasarian MM, Timmis KN, 1981, Molecular

and functional analysis of the TOL plasmid pWWO from

Pseudomonas putida and cloning of genes for the entire regulated

aromatic ring-cleavage pathway, Proc. Natl. Acad. Sci. U.S.A., 78:

7458.

Freedman DL and Gosett JM, 1989, Biological reductive dechlorination of

tetrachloroethylene and trichloroethylene to ethylene under

methanogenic conditions, Appl. Environ. Microbiol., 55: 2144–2157.

Furukawa K, Hirose J, Suyama A, Zaika T, Hayashida S, 1993, Gene

components responsible for discrete subtrate-specificity in the

metabolism of biphenyl (bph operon) and toluene (tod operon), J.

Bacteiol., 175: 5224.

Gadd GM, 2001, Fungi in bioremediation, Cambridge: Cambridge University

Press.

Galli R and McCarty PL, 1989, Biotransformation of 1,1,1-trichloroethane,

trichloromethane and tetrachloromethane by Clostridium spp., Appl.

Environ.Microbiol., 55: 837–844.

Gantzer CJ and Wackett LP, 1991, Reductive dechlorination catalyzed by

bacterial transition metal coenzymes, Environ. Sci. Technol., 25: 715–

722.

Ghani A, Wardle DA, Rahman A, Lauren DR, 1996, Interactions between

14C-labelled atrazine and the soil microbial biomass in relation to

herbicide degradation. Biol Fertil Soils, 21: 17-22.


84

Ghosal D, You IS, Chatterjee DK, Chakrabarty AM, 1985a, Microbial

degradation of halogenated compounds. Science (Washington, D.C.),

228: 135.

Ghosal D, You IS, Chatterjee DK, Chakrabarty AM, 1987, Nucleotide

sequence and expression of gene nahH of plas mid NAH7 and

homology with gene xylE of TOL pWWO, Gene, 55: 19.

Gibson SA and Sulfita JM, 1990, Anaerobic biodegradation of 2,4-5-

trichlorophenoxyacetic acid in samples from a methanogenic aquifer:

Stimulated by short chain organic acids and alcohols, Appl. Environ.

Microbiol., 56: 1825–1832.

Gibson DT, Zylstra GJ, Chauhan S, 1990, Biotransformations catalyzed by

toluene dioxygenase from Pseudomonas putida F1 In: Silver et al.

(Ed.) Pseudomonas: Biotransformations, Pathogenesis, and Evolving

Biotechnology. American Society for Microbiology, Washington, D.C.,

p.132.

Gibson SA and Sewell GW, 1992, Stimulation of reductive dechlorination of

tetrachloroethene in anaerobic aquifer microcosms by addition of short

chain organic acids or alcohols, Appl. Environ. Microbiol., 58: 1392–

1393.

Golovleva LA, Aharonson N, Greenhalgh R, Sethunathan N, Vonk JW, 1990,

The role and limitations of microorganisms in the conversion of

xenobiotics, Pure and Applied Chemistry, 62(2): 351- 364.

Gurbiel RJ, Batie CJ, Sivaraja M, True AE, Fee JA, Hoffman BM, Ballou DP,

1989, Electron nuclear double resonance spectroscopy of 15N-

enriched phthalate dioxygenase from Pseudomonas cepacia proves

that two histidines are co-ordinated to [2Fe-2S] Rieske-type clusters,

Biochemistry, 28: 4861.

Haddock JD, Nadim LM, Gibson DT, 1993, Oxidation of biphenyl by a

multicomponent enzyme system from Pseudomonas spp., strain

LB400, J. Bacteriol., 175: 395.

Haddock JD, Horton JR, Gibson DT, 1995, Dihydroxylation dechlorination of

chlorinated biphenyls by purified biphenyl 2,3-dioxygenase from

Pseudomonas spp., strain CB400, J. Bacteriol., 177: 20–26.


85

Hage JC and Kiestra, 2001, Hartmans Cometabolic degradation of chlorinated

hydrocarbons by Pseudomonas spp., strain DCA-1, Appl. Microbiol.

Biotechnol., 57: 548–554.

Haggblom MM and Young LY, 1990, Chlorophenol degradation coupled to

sulphate reduction, Appl. Environ. Microbiol., 56: 3255–3260.

Haider K, 1979, Degradation and metabolism of lindane and other

hexachlorocyclohexane isomers by anaerobic and aerobic soil micro-

organisms, Z.Naturforsch.Sect. C., 34: 1066–1069.

Haigler BE, Nishino SF, Spain JC, 1988, Degradation of 1,2-dichlorobenzene

by Pseudomonas spp., Appl. Environ. Microbiol., 54: 294–301.

Hambright P, 1975, In: Porphyrins and Metalloporphyrins, Smith, K.M., eds.,

Amsterdam: Elsevier Scientific, 233–278.

Han MJ, Choi HT, Song HG, 2004, Degradation of phenanthrene by Trametes

versicolor and its laccase, J Microbiol, 42: 94-8.

Harayama S, Rekik M, Timmis KN, 1986, Genetic analysis of a relaxed

substrate- specificity aromatic ring dioxygenase, toluate 1,2

dioxygenase, encoded by TOL plasmid pWWO of Pseudomonas

putida, Mol. Gen. Genet., 202: 226.

Harayama S and Rekik M, 1989, Bacterial aromatic ring cleavage enzymes

are classified into different gene families, J. Biol. Chem., 264:

15328.

Harayama S, Kok M, Neidle EL, 1992, Functional and evolutionary

relationships among diverse oxygenases, Annu. Rev. Microbiol., 46:

565.

Hardman DJ, 1991, Biotransformation of halogenated compounds, Crit.

Rev.Biotechnol., 11: 1–40.

Harker AR, Olsen RH, Seidler RJ, 1989, Phenoxyacetic acid degradation by

the 2,4-dichlorophenoxyacetic acid (TFD) pathway of plasmid pJP4:

mapping and characterization of the TFD regulatory gene, tfdR, J.

Bacteriol., 171: 314.

Harker AR and Kin Y, 1990, Trichloroethylene degradation by two

independent aromatic-degrading pathways in Alcaligenes eutrophus J

MP134, Appl. Environ. Microbiol., 56: 1179–1181.


86

Harpel MR and Lipscomb JD, 1990, Gentisate 1, 2 dioxygenase from

Pseudomonas: Purification, characterization and comparison of the

enzymes from Pseudomonas testosterone and Pseudomonas

acidovorans, J. Biol. Chem., 265: 6301.

Hartmans S, de Bont JAM, Harder W, 1989, Microbial metabolism of short

chain unsaturated hydrocarbons, FEMS., Microbiol. Rev., 63: 235–264.

Hartmans S, Janssen MW, van der Werf MJ, deBont JAM, 1991, Bacterial

metabolism of 3-chloroacrylic acid, J. Gen. Microbiol., 137: 2025.

Haughland RA, Sangodkar UMX, Chakrabarty AM, 1990, Repeated

sequences including RS 1100 from Pseudomonas cepacia AC 1100

function as IS elements, Mol. Gen. Genet, 220: 222.

Hendriksen HV, Larsen S, Ahring BK, 1992, Influence of a supplemental

carbon source on anaerobic dechlorination of pentachlorophenol in a

granular sludge, Appl. Environ. Microbiol., 58: 365–370.

Henikoff S, Haughn GW, Calvo JM, Wallace JC, 1988, A large family of

bacterial activator proteins, Proc. Natl. Acad. Sci. U.S.A., 85: 6602.

Heritage AD and Mackar IC, 1977, Degradation of lindane by cell free

preparations of Clostridium sphenoides, Appl. Environ. Microbiol., 34:

222–224.

Herrmann H, Müller C, Schmidt I, Mahnke J, Petrschka L, Hahnke K, 1995,

Localization and organization of phenol degradation genes of

Pseudomonas putida strain, H. Mol. Gen. Genet., 247: 240.

Hestbjerg H, Willumsen PA, Christensen M, Andersen O, Jacobsen CS, 2003,

Bioaugmentation of tar-contaminated soils under field conditions using

Pleurotus ostreatus refuse from commercial mushroom production.

Environ Toxicol Chem, 22: 692-8.

Hill GA, Milne BJ, Nawrocki PA, 1996, Cometabolic degradation of 4-

chlorophenol by Alcaligenes eutrophus, Microbiol., Biotechnol., 46:

163–168.

Hirose J, Kimura N, Suyama A, Kobayashi A, Hayashida S, Furukawa K,

1994, Functional and structural relationship of various extradiol

aromatic ringcleavage dioxygenases of Pseudomonas origin, FEMS

Microbiol. Lett., 118: 273.


87

Hoefer B, Eltis LD, Dowling DN, Timmis KN, 1993, Genetic analysis of a

Pseudomonas locus encoding a pathway for biphenyl/polychlorinated

biphenyl degradation. Gene, 130: 47.

Hogenkamp HPC, 1975, In: Cobalamin: Biochemistry and Pathophysiology,

Babior, B.M., eds., New York: John Wiley & Sons, 21–75.

Holliger C, Schraa G, Stams AJM, Zehnder AJB, 1988, In: Tilche, A., and

Rozzi, A., eds., Poster papers of the V International Symposium on

Anaerobic Digestion, Monduzzi Editre, Bologna, Italy, 847–850.

Holliger C, Schraa G, Stams AJM, Zehnder AJB, 1992, Enrichment and

properties of an anaerobic mixed culture reductively dechlorinating

1,2,3- trichlorobenzene to 1,3-dichlorobenzene, Appl. Environ.

Microbiol., 58: 1638–1644.

Holliger C, Schraa G, Stams AJM, Zehnder AJB, 1993, A highly purified

enrichment culture couples the reductive dechlorination of

tetrachloroethene to growth, Appl. Environ. Microbiol., 59: 2991–2997.

Holliger C and Schumacher W, 1994, Reductive dehalogenation as a

respiratory process, Antonie van Leuwenhock, 66: 239–246.

Holliger C, Schraa G, 1994, Physiological meaning and potential for

application of reductive dechlorination by anaerobic bacteria, FEMS

Microbiol Rev., 15: 297–305.

Holscher T, Gorisch H, Adrian L, 2003, Anaerobic reductive dehalogenation of

chlorobenzene congeners in all extracts of Dehalococcoides spp.,

strain CBDB1, Appl. Environ. Microbiol., 69: 2999–3001.

Hooper SW, Dockendorff TC, Sayler GS, 1989, Characteristics and

restriction analysis of the 4-chlorobiphenyl catabolic plasmid,

pSS50., Appl. Environ. Microbiol., 55: 1286.

Hopkins GD, Semprini L, McCarty PL, 1993, Microcosm and in situ field

studies of enhanced biotransformation of trichloroethylene by phenol

oxidizing micro-organisms, Appl. Environ. Microbiol, 59: 2277–2285.

Hopkins GD and McCarty PL, 1995, Field evaluation of in-situ aerobic

cometabolism of trichloroethylene and 3-dichloroethylene isomers

using phenol and toluene as the primary substrates, Environ. Sci.

Technol., 29:1628–1637.


88

Hugo N, Meyer IC, Armengaud IJ, Gaillard J, Timmis KN, Jouanneau Y, 2000,

Characterization of three XylT-Like [2Fe-2S] ferredoxins

associated with catabolism of cresols or naphthalene: Evidence for

their involvement in catechol dioxygenase reactivation, J. Bacteriol.

182: 5580.

Hutzinger O and Veerlamp W, 1981, In: Microbial Degradation of Xenobiotic

and Recalcitrant Compounds, Leisinger, T., Cook, A., Hutter, R., and

Nuesch, J., eds., London: Academic Press 3–45.

Imai R, Nagata Y, Fukuda M, Takagi M, Yano K, 1991, Molecular cloning of

a Pseudomonas paucimobilis gene encoding a 17-kilodalton

polypeptide that eliminates HCl molecules from γ -HCH, J. Bacteriol.

173: 6811–6819.

Inouye S, Nakazawa A, Nakazawa T, 1983, Molecular cloning of regulatory

gene xylR and operator-promoter regions of the xylABC and xylDEGF

operons of the TOL plasmid, J. Bacteriol., 155: 1192.

Inouye S, Nakazawa A, Nakazawa T, 1986, Nucleotide sequence of the

regulatory gene xylS on the Pseudomonas putida TOL plasmid and

identification of the protein product, Gene, 44: 235.

Inouye S, Nakazawa A, Nakazawa T, 1985, Determination of the

transcription initiation site and identification of the protein product of the

regulatory gene xylR for xyl operons on the TOL plasmid, J. Bacteriol.,

163:863.

Inouye S, Kimoto M, Nakazawa A, Nakazawa T, 1990, Presence of flagella

in Pseudomonas putida is dependent on the ntrA (rpoN) gene, Mol.

Gen. Genet., 221: 295.

Irie S, Doi S, Yorifuji T, Takagi M, Yano K, 1987, Nucleotide sequencing

and characterization of the genes encoding benzene oxidation

enzymes of Pseudomonas putida, J. Bacteriol., 169: 5174.

Jagnow G, Haider K, Ellwardt PC, 1977, Anaerobic dechlorination and

degradation of HCH isomers by anaerobic and facultative anaerobic

bacteria, Arch.Microbiol., 115: 285–292.


89

Janssen DB, Scheper A, Dijkhuizen L, Witholt B, 1985, Degradation of

halogenated aliphatic compounds by Xanthobacter autotrophicus

GJ10, Appl. Environ Microbiol., 49: 673–677.

Janssen DB, Pries F, Van der Ploeng J, Kazenier B, Terpstra P, Witholt B,

1989, Cloning of 1,2-dichloroethane degradation gene of Xanthobacter

autotrophicus GJ10 and expression and sequencing of the dhl A gene,

J. Bacteriol., 171: 6791–6799.

Janssen DB, Oldenhius R, Van den Wijngaard AJ, 1989, Degradation of

xenobiotic compounds by microorganisms, VDI Berichte, 735: 25–39.

Janssen DB, Pries F, van der Ploeg JR, 1994, Genetics and biochemistry of

dehalogenating enzymes, Annu. Rev. Microbiol., 48: 163–191.

Jeffery WH, Cuskey SM, Chapman PJ, Resnick S, Olsen RH, 1992,

Characterization of Pseudomonas putida mutants unable to catabolize

benzoate: cloning and characterization of Pseudomonas genes

involved in benzoate catabolism and isolation of a chromosomal DNA

fragment able to substitute for XylS in activation of the TOL lower

pathway promoter, J. Bacteriol., 174: 4986.

Jiangzhong HC, Ritalahti KM, Aiello MR, Loeffler FE, 2003, Complete

detoxification of VC by an anaerobic environment culture and

identification of the reductively dechlorinating population as a

Dehalococcoides species, Appl. Environ. Microbiol., 69: 996–1003.

Kallman BJ and Andrews AK, 1963, Reductive dechlorination of DDT to DDD

by yeast, Science, 141: 1050- 1051.

Kaphammer B, Kukor JJ, Olsen RH, 1990, Regulation of tfdCDEF by tfdR of

the 2,4-dichlorophenoxyacetic acid degradation plasmid pJP4. J.

Bacteriol. 172: 2280.

Kasak L, Hörak R, Nurk A, Talvik K, Kivisaar M, 1993, Regulation of the

catechol 1,2-dioxygenase and phenol monooxygenases encoding

pheBA operon in Pseudomonas putida PaW85, J. Bacteriol., 175:

8038.

Kesai N, Tsujimura K, Unoura K, Suzuki T, 1990, Degradation of 2,3-dichloro-

1-propanol by a Pseudomonas spp., Agric Biol Chem. 54, 3185–3190.


90

Khan A and Walia S, 1990, Identification and localization of 3-phenylcatechol

dioxygenase and 2-hydroxy-6-oxo-6-phenyl-hexa-2,4-dienoate

hydrolase genes of Pseudomonas putida and expression in

Escherichia coli, Appl. Environ. Microbiol., 56: 956.

Khanna S, Lal B, Chandra R, Rajam S, Bajpai U, 1998, Enzymes and

bioremediation, In Enzyme Chemistry: Impact and applications.

Suckling, C.J., Gibson, C.L.,and Pitt, A.K., Eds., Blackie Academic and

Professional, Chapman and Hall, London, p.240.

Kilbane JJ, Chatterjee DK, Karns JS, Kellog ST, Chakrabarty AM, 1982,

Biodegradation of 2,4,5-trichlorophenoxyacetic acid by a pure culture of

Pseudomonas cepacia., Appl. Environ. Microbiol., 44: 72.

Kimbara K, Hashimoto T, Fukuda M, Koana T, Takagi M, Oishi M, Yano K,

1989, Cloning and sequencing of two tandem genes involved in

degradation of 2,3-dihydroxybiphenyl into benzoic acid in the

polychlorinated biphenyl degrading soil bacterium Pseudomonas spp.,

strain KKS102, J.Bacteriol., 1171: 2740.

Kinkle BK, Sadowsky MI, Schmidt EL, Koskinen WC, 1993, Plasmids pJP4

and r68.45 can be transferred between populations of Bradyrhizobia in

nonsterile soil, Appl. Environ. Microbiol., 59: 1762.

Kivisaar M, Hörak R, Kasak L, Heinaru A, Habicht J, 1990, Selection of

independent plasmids determining phenol degradation in

Pseudomonas putida and the cloning and expression of genes

encoding phenol monooxygenases and catechol 1,2-dioxygenase,

Plasmid, 24: 25.

Kivisaar M, Kasak L, Nurk A, 1991, Sequence of the plasmid-encoded

catechol 1,2-dioxygenase expressing gene, pheB, of phenol-degrading

Pseudomonas spp., strain EST 1001, Gene, 98: 15.

Klecka GM and Gonsior SJ, 1984, Reductive dechlorination of chlorinated

methanes and ethanes by reduced iron(II) porphyrins, Chemosphere

13, 391-402, 1984.

Kluyver AJ, 1931, The chemical activities of microorganisms. Univ. of London

Press, London.


91

Knackmus HJ and Hellwig M, 1978, Utilization and co-oxidation of chlorinated

phenols by Pseudomonas spp., B13, Arch. Microbiol., 117: 1–7.

Knackmuss HJ, 1981, Degradation of halogeneted & sulfonated

hydrocarbons, FEMS Symp., 12: 189–212.

Kobayashi H and Rittmann BE, 1982, Microbial removal of hazardous organic

compounds, Environ. Sci. Technol., 16: 170A–183A.

Kohler-Staub D and Leisinger T, 1985, Dichloromethane dehalogenase of

Hyphomicrobium spp., strain DM2, J. Bacteriol., 162: 676–681.

Kohler-Staub D, Hartmans S, Galli R, Suter F, Leisinger T, 1986, Evidence

for identical dichloromethane dehalogenases in different methylotrophic

bacteria, J. Gen. Microbiol., 132: 2837–2843.

Kosono S, Maeda M, Fuji F, Arai H, Kudo T, 1997, Three of the seven

bphC genes of Rhodococcus erythropolisTA421, isolated from a

termite ecosystem, are located on a indigenous plasmid associated

with biphenyl degradation, Appl. Environ. Microbiol., 63: 3282.

Krone U, Thauer RK, Hogenkamp HPC, 1989, Reductive dehalogenation

of chlorinated Cl-hydrocarbons mediated by corrinoids, Biochemistry

28, 4908–4914.

Krone VE and Thauer RK, 1992, Dehalogenation of trichlorofluronethane

(CFC- 11) by Methanosarcina barkeri, FEMS Microbiol. Lett., 90, 201–

204.

Krumme ML, Timmis KN, Dwyer DF, 1993, Degradation of trichloroethylene

by Pseudomonas cepacia G4 and the constitutive mutant strain

G45223 PR 1 in aquifer microcosms, Appl. Environ. Microbiol. 59,

2746–2749.

Kukor J, Olson R, Siak JS, 1989, Recruitment of a chromosomally encoded

maleylacetate reductase for degradation of 2,4-dichlorophenoxyacetic

acid by the plasmid pJP4, J. Bacteriol., 171: 3385.

Lal R, 1982, Accumulation, metabolism and effects of organophosphorous

insecticides on microorganisms, Adv. Appl. Microbiol., 28: 149.

Lal R and Saxena DM, 1982, Accumulation, metabolism and effects of

organochlorine insecticides on microorganisms, Microbiol. Rev., 46: 95.


92

Lamar RT, Larsen MJ, Kirk JK, 1990, Sensitivity to and degradation of

pentachlorophenol by Phanerochate spp., Appl. Environ. Microbiol.

Langlois BE, 1967, Predictive dechlorination of DDT by Escherichia coli.

Journal of Dairy Science, 50: 1168- 1170.

Langlois BE, Collins JA, Sides KG, 1970. Some factors effecting degradation

of organochlorine pesticides by bacteria, Journal of Dairy Science, 53:

1671- 1675.

Lloyd-Jones G, deJong C, Ogden RC, Duetz WA, Williams PA, 1994,

Recombination of the bph(biphenyl) catabolic genes from plasmid

pWW100 and their deletion during growth on benzoate, Appl. Environ.

Microbiol., 60: 691.

Lee SB, Strand SE, Stensel HD, 2000, Sustained degradation of

trichloroethylene in a suspended growth gas treatment reactor by an

Actinomycetes, Environ. Sci. Technol., 34: 3261.

Leisinger T, Roche S La, Bader R, Appert MS, Stromeyer SB, Cook AM,

1993, In: Microbial growth on chlorine compounds, Murrel, C., and

Kelly, D.P. eds., Andover, UK: Intercept.

Leisinger T, Bader R, 1993, Microbial Dehalogenation of synthetic

organohalogen compounds: Hydrolytic dehalogenases, Chimia 47,

116–121.

Leng ML, 1986, Chlorophenoxyalkanoic acids. In: Gerhartz, W., eds.,

Ullmann’s Encyclopedia of Industrial Chemistry, Weinheim: VCH A7,

9–16.

Lewis TA and Crawford RL, 1993, Physiological factors affecting carbon

tetrachloride dehalogenation by the denitrifying bacterium

Pseudomonas spp., strain KC, Appl. Environ. Microbiol., 59:

1635– 1641.

Li S and Wackett LP, 1992, Trichloroethylene oxidation by toluene

dioxygenase, Biochem. Biophys. Res. Commun., 185: 443–451.

Linkfield TG and Tiedje JM, 1990, Characterization of the requirements and

substrates for reductive dehalogenation by strain DCB-1, J. Ind.

Microbiol., 5: 9–16.


93

Loeffler FE, Sanford RA, Tiedje JM, 1996, Initial characterization of a

reductive dehalogenase from Desulfitobacterium chlrorespirans Co 23,

Appl. Environ. Microbiol., 62: 3809–3813.

Loffler F and Muller R, 1991, Identification of 4-chlorobenzoyl coenzyme A as

intermediate in the dehalogenation catalyzed by 4-chlorobenzoate

dehalogenase from Pseudomonas spp., CBS3, FEBS Lett. 290: 224–

226.

Lu MC, Anotai J, Liao CH, Ting WP, 2004, Dechlorination of

hexachlorobenzene by zero valent iron, Pract. Period. of Haz. Toxic

Radioact. Waste Manage., 8: 136–140.

Mac Rae IC, Raghu K, Bautista EM, 1969, Anaerobic degradation of the

insecticide lindane by Clostridium spp., Nature (Lond.) 221, 859–860.

Madsen T and Aamand J, 1992, Anaerobic transformation and toxicity of

trichlorophenols in a stable enrichment culture, Appl. Environ.

Microbiol., 58: 577–561.

Magli A, Messmer M, Keisinger T, 1998, Metabolism of dichloromethane by

the strict anaerobic Dehalobacterium formicoaceticum, Appl. Environ.

Microbiol., 64: 646–650.

Marinucci AC and Bartha R, 1979, Biodegradation of 1,2,3- and 1,2,4-

trichlorobenzene in soil and in liquid enrichment culture, Appl.

Environ. Microbial., 38: 811–817.

Markus A, Krekel D, Singens F, 1986, Purification and some properties of

compound A of the 4-chlorophenylacetate 3,4-dioxygenase from

Pseudomonas species strain CBSE, J. Biol. Chem., 261:12883–12888.

Matsumura F and Bousch GM, 1968, Degradation of insecticides by a soil

fungus Trichoderma viride, Journal of Economic Entomolology, 69:

610-612.

Maymo-Gatell XM, Chien YT, Gosett JM, Zinder SH, 1997, Isolation of a

bacterium that reductively dechlorinates tetrachloroethene to ethene,

Science, 276: 1568–1571.

McCarty PL, 1987, Bioengineering issues related to in-situ remediation of

contaminated soils and groundwater, In: Environmental Biotechnology,

Omenn, G.S. eds., New York: Plenum Press, 143–162.


94

MeBmer M, Wohlfarth G, Dickert G, 1993, Methyl chloride metabolism of the

strictly anaerobic, methyl chloride-utilizing homoacetogen strain MC,


MeBmer M, Reinhardt S, Wohlfarth G, Diekert G, 1996, Studies on methyl

chloride dehalogenase and o-demethylase in cell extracts of the

homoacetogen strain MC based on a newly developed coupled

enzyme assay, Arch. Microbiol., 165: 18–25.

Mendel JL, Klein AK, Chen JT, Walton MS, 1967, Metabolism of DDT and

some other chlorinated organic compounds by Aerobacter aerogenes,

Journal of Association Official Analytical Chemists, 50: 897-903.

Mermod N, Ramos JL, Bairoch A, Timmis KN, 1987, The xylS gene

positive regulator of TOL plasmid pWWO: identification, sequence

analysis and overproduction leading to constitutive expression of meta

cleavage operon. Mol. Gen. Genet., 207: 349.

Mikesell MD, Boyd SA, 1990, Dechlorination of chloroform by Methanosarcina

strains, Appl. Environ Microbiol., 56: 1198–1201.

Mohn WW and Tiedje JM, 1991, Evidence for chemiosmotic coupling of

reductive dechlorination and ATP synthesis in Desulfomonile tiedje,


Mohn WW and Tiedje JM, 1992, Microbial reductive dehalogenation,

Microbiol. Rev., 56: 482–507.

Mohn WW and Kennedy KJ, 1992, Reductive dehalogenation of

chlorophenols by Desulfomonile tiedje DCB-1, Appl. Environ.

Microbiol., 58: 1367–1370.

Mswaka AY and Magan N, 1999, Temperature and water potential relations of

tropical Trametes and other wood-decay fungi from the indigenous

forests of Zimbabwe. Mycol Res., 103: 1309-17.

Muller F and Caillard L, 1986, Chlorophenols, In: Gerhartz, W., eds.,

Ullmann’s Encyclopedia of Industrial Chemistry, Weinheim: VCH 7,

1–8.

Müller R, Thiele J, Klages U, Lingens F, 1984, Incorporation of (18O) water

into 4-hydroxybenzoic acid in the reaction of 4-chlorobenzoate


95

dehalogenase from Pseudomonas spec. CBS3, Biochem. Biophys.

Res. Commun, 124: 178.

Müller RH and Babel W, 1994, Phenol and its derivatives as heterotrophic

substrates for microbial growth-an energetic approach, Appl.

Microbiol. Technol., 42: 446.

Nadeau LJ, Menn FM, Breen A, Sayler GS, 1994, Aerobic degradation of

1,1,1- trichloro-2,2-bis(4-chlorophenyl)ethane (DDT) by Alcaligenes

eutrophus A5, Applied and Environmental Microbiology, 60: 51- 55.

Nagata Y, Hatta T, Imai R, Kimbara K, Fukuda M, Iano K, Takagi M, 1993,

Purification & characterization of γ HCH dehydrochlorinase (Lin A) from

Pseudomonas paucimobilis, Biosci. Biotechnol. Biochem., 57: 1582–

1583.

Nagata Y, Nariya T, Ohtomo R, Fukuda M, Yano K, Takagi M, 1993,

Cloning and sequencing of a dehalogenase gene encoding an enzyme

with hydrolase activity involved in the degradation of _-

hexachlorocyclohexane in Pseudomonas paucimobilis, J.Bacteriol.,

175: 6403.

Nakai C, Kagimiyama H, Nozaki M, Nakazawa T, Inouye S, 1983, Complete

nucleotide sequence of the metapyrocatechase gene on the TOL

plasmid of Pseudomonas putida mt-2, J. Biol. Chem., 258: 2923.

Nakatsu CJ, Singh R, Straus N, Wyndham C, 1991, Chlorobenzoate

catabolic transposon Tn 5271 is a composite class I element with

flanking class II insertion sequences. Proc. Natl. Acad. Sci. U.S.A. 88:

8312.

Neidle EL, Hartnett C, Ornston LN, Bairoch A, Rekik M, Harayama S, 1991,

Nucleotide sequences of the Acinetobacter calcoaceticus benABC

genes for benzoate 1,2-dioxygenase reveal evolutionary relationships

among multicomponent oxygenases, J. Bacteriol., 173: 5385.

Neidle EL, Hartnett C, Ornston NL, Bairoch A, Rekik M, Harayama S, 1992,

Cisdiol dehydrogenases encoded by the TOL pWWO plasmid

xylL gene and the Acinetobacter calcoaceticus chromosomal benD

gene are members of the short-chain alcohol dehydrogenase

superfamily, Eur. J. Biochem., 204: 113.


96

Nelson MJK, Montgomery SO, O’Neill EJ, Pritchard PH, 1986, Aerobic

metabolism of trichloroethylene by a bacterial isolate, Appl. Environ.

Microbiol., 52: 383–384.

Nelson MJK, Montgomery SO, Mahaffey WR, Pritchard PH, 1987,

Biodegradation of trichloroethylene and involved of an aromatic

biodegradative pathway, Appl. Environ Microbiol., 53: 949–954.

Nelson MJK, Montgomery SO, Pritchard PH, 1988, Chloroethylene

metabolism by micro-organisms that degrade aromatic compounds,

Appl. Environ Microbiol., 54: 604–606.

Neumann A, Scholtz MH, Diekert G, 1994, Tetrachloroethene metabolism of

Dehalospirillum multivorans, Arch Microbiol., 162: 295–301.

Nies L and Vogel TM, 1990, Effects of organic substrates on dechlorination of

aroclor 1242 in anaerobic sediments, Appl. Environ. Microbiol., 56:

2612– 2617.

Nordlund I, Powlowski J, Hagström A, Shingler V, 1993, Conservation of

regulatory and structural genes for a multi-component phenol

hydroxylase within phenol catabolising bacteria that utilize meta-

cleavage pathway, J. Gen. Microbiol., 139: 2695.

Novotný C, Svobodová K, Erbanová P, Cajthaml T, Kasinath A, Lang E,

Šašek V, 2004, Ligninolytic fungi in bioremediation: extracellular

enzyme production and degradation rate, Soil Biol Biochem., 36: 1545-

51.

Nurk A, Kasak L, Kivisaar M, 1991, Sequence of the gene (pheA) encoding

phenol monooxygenase from Pseudomonas spp., EST1001:

expression in Escherichia coli and Pseudomonas putida, Gene, 102:

13.

Ohisa N and Yamaguchi M, 1978, Gamma BHC degradation accompanied by

growth of Clostridium rectum isolated from paddy field soil, Agric. Biol.

Chem., 42: 1819–1823.

Ohisa N, Yamaguchi M, Kurihara N, 1980, Lindane degradation of cell-free

extracts of Clostridium rectum, Arch. Microbiol., 125: 221–225.

Ohisa N, Kurihara N, Nakajima M, 1982, ATP synthesis associated with the

conversion of HCH related compounds, Arch. Microbiol., 131, 330–333.


97

Parsek MR, Ye RW, Pun P, Chakrabarty AM, 1994, Critical nucleotides in the

interaction of a LysR-type regulator with its target promoter region:

catBC promoter activation by CatR., J. Biol. Chem., 269: 11279.

Parsons F, Wood PR, DeMarco JJ, 1984, Transformation of tetrachloroethene

and trichloroethene in microcosms and ground water, J. Am. Water

Works Asso., 76: 56–59.

Pavlostathis SG and Zhuang P, 1991, Transformation of trichloroethylene by

sulfate-reducing cultures enriched from a contaminated subsurface

soil, Appl. Microbiol. Biotechnol., 36: 416–420.

Perkins EJ, Gordon MP, Caceres O, Lurquin PF, 1990, Organization and

sequence analysis of the 2,4-dichlorophenol hydroxylase and

dichlorocatechol oxidative operons of plasmid pJP4, J Bacteriol.,

172: 2351.

Persson B, Krook M, Jornvall H, 1991, Characteristics of short-chain alcohol

dehydrogenases and related enzymes, Eur. J. Biochem., 200: 537.

Pointing SB, 2001, Feasibility of bioremediation by white-rot fungi.Appl

Microbiol Biotechnol, 57: 20-33.

Powlowski J and Shingler V, 1990, In vitro analysis of polypeptide

requirements of multicomponent phenol hydroxylase from

Pseudomonas spp., strain CF600, J. Bacteriol., 172: 6834.

Radtke C, Cook WS, Anderson A, 1994, Factors affecting antagonism of the

growth of Phanerochaete chrysosporium by bacteria isolated from

soils. Appl Microbiol Biotechnol., 41: 274-80.

Ramanand K, Balba MT, Duffy J, 1993, Reductive dehalogenation of

chlorinated benzenes and toluenes under methanogenic conditions,


Romine MF, Stillwell LC, Wong KK,Thurston SJ, Sisk EC, Sensen C,

Gaasterland T, Fredrickson JK, Saffer JD, 1999, Complete Sequence

of a 184-Kilobase Catabolic Plasmid from Sphingomonas

aromaticivorans F199, J. Bacteriol., 181: 1585.

Ramos JL, Mermod N, Timmis KN, 1987, Regulatory circuits controlling

transcription of TOL plasmid operon encoding meta-cleavage pathway


98

for degradation of alkylbenzoates by Pseudomonas, Mol. Microbiol., 1:

293.

Roper DI and Cooper RA, 1990, Subcloning and nucleotide sequence of the

3,4- dihydroxyphenyl acetate (homoprotocatechuate) 2,3-

dioxygenase gene from Escherichia coli C., FEBS Lett., 275: 53.

Rossberg M, Lendle W, Togel A, Dreher EL, Langer E, Rossaerts H,

Kleinschmidt P, Streck H, Beck U, Lipper KA, Torkelson TR, Loser E,

Beutel KK, 1986, Chlorinated hydrocarbons. In: Gerhartz W., ed.

Ullmann’s Encyclopedia of Industrial Chemistry, Weinheim: VCH 6,

233–239.

Rothmel RK, Aldrich TL, Houghton JE, Coco WM, Ornston LN, Chakrabarty

AM, 1990, Nucleotide sequencing and characterization of

Pseudomonas putida catR: a positive regulator of the catBC operon is

a member of the LysR family, J.Bacteriol, 172: 922.

Sander P, Wittich RM, Fortnagel P, Wilkes H, Francke W, 1991,

Degradation of 1,2,4-trichloro and 1,2,4,5-tetrachlorobenzene by

Pseudomonas strains, Appl.Environ. Microbiol., 57: 1430–1440.

Sangodkar UMX, Chapman PJ, Chakrabarty AM, 1988, Cloning, physical

mapping and expression of chromosomal genes specifying degradation

of the herbicide 2,4,5-T by Pseudomonas cepacia AC 1100. Gene 71,

267.

Schell MA, 1993, Molecular biology of the LysR family of transcriptional

regulators, Ann. Rev. Microbiol., 47: 597.

Schmidt E, 1987, Response of a chlorophenol degrading mixed culture to

changing loads of phenol, chlorophenol and cresols. Appl. Microbiol.

Biotechnol., 27: 94.

Scholten JD, Chang KH, Babbitt PC, Charest H, Sylvestre M, Dunaway MD,

1991, Novel enzymic hydrolytic dehalogenation of a chlorinated

aromatic, Science, 253: 182.

Schweizer D, Markus A, Seez M, Ruf H, Lingens F, 1987, Purification &

some properties of component B of the 4-chlorophenylacetate 3-4

dioxygenase from Pseudomonas species strain CBS3, J. Biol Chem.,

262: 9340–9346.


99

Semprini L, 1997, Strategies for the aerobic cometabolism of chlorinated

solvent, Curr. Opin. Biotechnol., 8: 296–308.

Shields MS, Montgomery SO, Chapman PJ, Cuskey SM, Pritchard PH, 1989,

Novel pathway of toluene catabolism in the trichloroethylene-degrading

bacterium G4, Appl. Environ. Microbiol., 55: 1624–1629.

Shields MS, Montgomery SO, Cuskey SM, Chapman PJ, Pritchard PH, 1991,

Mutants of Pseudomonas cepacia G4 defective in catabolism of

aromatic compounds & trichloro ethylene, Appl. Environ. Microbiol. 57,

1935–1941

Shingler V, Powlowski J, Marklund U, 1992, Nucleotide sequence and

functional analysis of the complete phenol/3,4-dimethylphenol catabolic

pathway of Pseudomonas spp., strain CF600, J. Bacteriol., 174: 711.

Singh BK, Kuhad RC, Singh A, Lal R, Tripathi KK, 1999, Biochemical and

molecular basis of pesticide degradation by microorganisms, Critical

Reviews in Biotechnology, 19(3): 197- 225.

Slater JH, Bull AT, Hardman DJ, 1995, Microbial dehalogenation,

Biodegradation, 6: 181–189.

Slater JH, Bull AT, Hardman DJ, 1997, Microbial dehalogenation of

halogenated alkanoic acids alcohols and alkanes, Adv. Microb Physiol.,

38:133–176.

Smatlak CR, Gosett JM, Zinder SH, 1996, Comparitive kinetics of hydrogen

utilization for reductive dechlorination of tetrachloroethene and

methanogenisis in an anaerobic enrichment culture, Environ. Sci.

Technol., 30: 2850–2856.

Smets BF, Rittmann BE, Stahl DA, 1993, The specific growth rate of

Pseudomonas putida PAW1influences the conjugal transfer rate of the

TOL plasmid, Appl. Environ. Microbiol., 59: 3430.

Spain JC, Nishino SF, 1987, Degradation of 1,4-dichlorobenzene by

Pseudomonas spp., Appl. Environ. Microbiol., 448: 840–848.

Spooner RA, Lindsay K, Franklin FCH, 1986, Genetic, functional and

sequence analysis of the xylR and xylS regulatory genes of the TOL

plasmid pWWO, J. Gen. Microbiol., 132: 1347.


100

Springael D, Kreps S, Mergeay M, 1993, Identification of a catabolic

transposon, Tn 4371, carrying biphenyl and 4-chlorobiphenyl

degradation genes in Alcaligenes eutrophus A5, J Bacteriol., 175:

1674.

Stanier RY, Palleroni NJ, Doudoroff M, 1966, The aerobic Pseudomonads:

A taxonomic study, J. Gen. Microbiol., 43: 159–271.

Sterjiades R, Pelmont J, 1989, Occurrence of two different forms of

protocatechuate 3,4-dioxygenase in a Moraxella spp., Appl. Environ

Microbiol. ,55: 340.

Stumm W and Morgan JJ, 1981, Aquatic Chemistry, 2nd ed., New York:

Wiley. Suen WC, Spain JC, 1993, Cloning and characterization of

Pseudomonas spp., strain DNT genes for 2,4-dinitrotoluene

degradation, J. Bacteriol., 175: 1831.

Sung Y, Ritalahti KM, Sanford RA, Urbance JW, Flynn ST, Tiedje JM,

Loeffler FE, 2003, Characterization of two tetrachloroethene reducing

acetateoxidixing anaerobic bacteria and their description as

Desulfuromonas michiganensis spp., nov., Appl. Environ. Microbiol.,

69: 2964–2974.

Susanna A, Stromeyer SB, Hermann R, Cook AM, Leisinger T, 1993,

Dichloromethane as the sole carbon source for an acetogenic mixed

culture and isolation of a fermentative degrading bacterium, Appl.


Sylvestre M, Masse R, Ayotte C, Messier F, Fauteux J, 1985, Total

biodegradation of 4-chlorobiphenyl (4CBP) by a twomembered

bacterial culture. Appl. Microbiol. Biotechnol., 21: 191.

Taira K, Hayase N, Arimura N, Yamashita S, Miyazaki T, Furukawa K, 1988,

Cloning and nucleotide sequences of the 2,3-dihydroxybiphenyl

dioxygenase gene from the PCB degrading strain of Pseudomonas

paucimobilis Q1, Biochemistry, 27: 3990.

Taraban RH, Derry DF, Berry DA, Walker HL, 1993, Degradation of Dicamba

by an anaerobic consortium enriched from wetland soil, Appl.



101

Tardif G, Greer CW, Labbe D, Lau PCK, 1991, Involvement of a large

plasmid in the degradation of 1,2-dichloroethane by Xanthobacter

autotrophicus, Appl. Environ. Microbiol., 57: 1853–1857.

Tatara GM, Dybas MJ, Criddle CS, 1993, Effects of medium and trace

metals on kinetics of carbon tetrachloride transformation by

Pseudomonas spp., strain KC, Appl. Environ. Microbiol., 59: 2126–

2131.

Timmis KN, Steffan RJ, Unterman R, 1994, Designing microorganisms for the

treatment of toxic wastes, Ann. Rev. Microbiol., 48: 525.

Trejo-Hernandez MR, Lopez-Munguia A, Ramirez RQ, 2001, Residual

compost of Agaricus bisporus as a source of crude laccase for enzymic

oxidation of phenolic compounds, Process Biochem., 36: 635-9.

Tsien HC, Brusseau GA, Hanson RS, Wackett LP, 1989, Biodegradation of

trichloroethylene by Methanosinus trichosporium OB3b, Appl. Environ.

Microbiol., 55: 3155–3161.

Tsuda M and Iino T, 1988, Identification and characterization of Tn4653, a

transposon covering the toluene transposon Tn4651 on TOL plasmid

pWWO. MOl. Gen. Genet., 213: 72.

Tsuda M, Minegishi KI, Iino T, 1989, Toluene transposons Tn4651 and

Tn4653 are class II transposons, J. Bacteriol., 171: 1386.

Tuor U, Winterhalter K, Fiechter A, 1995, Enzymes of white-rot fungi involved

in lignin degradation and ecological determinants for wood decay, J

Biotechnol., 41: 1-17.

U. S. EPA .1 Leisinger, T., and Brunner, W. In: Biotechnology, Rehm, H.J.,

Reed, G., eds., VCH Verlagsgesellschaft: Weinheim 8, 475–513, 1986.

Valli K, Wariish H, Gold MH, 1992, Degradation of 2,7-dichlorodibenzo-p-

dioxin by the lignin degrading basidiomycete Phanerochaete

chrysosporium, J Bacteriol., 174: 2131-7.

Vannelli T, Logan M, Arciero DM, Hopper AB, 1990, Degradation of

halogenated aliphatic compounds by ammonia-oxidizing bacterium

Nitrosomonas europia, Appl., Environ. Microbiol., 56: 1169–1171.

Vannelli T, Chapman P, Hooper AB, 1991, Degradation of halogenated

aliphatic compounds by the ammonia-oxidizing bacteria Nitrosomonas


102

europia. In: EPA Symposium on Bioremediation of Hazardous

Wastes, Falls Church, VA.

Van den Wijngaard AJ, Janssen DB, Witholt B, 1989, Degradation of

epichlorohydrin and halohydrins by bacterial cultures isolated from

freshwater sediments, J. Gen. Microbiol., 135: 2199–2208.

Van den Wijngaard AJ, Reuvekamp PT, Janssen DB, 1991, Purification and

characterization of haloalcohol dehalogenese from Arthrobacter spp.,

strain AD2, J. Bacteriol., 173: 124–129.

Van der Meer JR, Roelofsen W, Schraa G, Zehnder AJB, 1987,

Degradation of low concentrations of dichlorobenzenes and 1,2,4-

trichlorobenzene by Pseudomonas spp., strain P51 in non-sterile soil

columns, FEMS Microbiol. Ecol., 45: 333–341.

Van der Meer JR, Zehnder AJB, deVos WM, 1991, Identification of a novel

composite transposable element, Tn 5280, carrying chlorobenzene

dioxygenase genes of Pseudomonas spp., strain P51, J. Bacteriol.,

173: 7077.

Van der Meer JR, deVos WM, Harayama S, Zehnder AJB, 1992, Molecular

mechanisms of genetic adaptation to xenobiotic compounds, Microbiol.

Rev., 56: 677.

Van Eckert MHA, Schroder TJ, Stams AJM, Schraa G, Field JA, 1998,

Degradation and fate of carbon tetrachloride in unadapted

methanogenic generator sludge, Appl. Environ. Microbiol., 64: 2350–

2358.

Vargas C, Fennell DE, Haggblom MM, 2001, Anaerobic reductive

dechlorination of chlorinated dioxins in estuarine sediments, Appl.

Microbiol. Biotechnol., 57: 786–790.

Villarante NR, Armenante PM, Quibuyen TAO, Favak F, Kafkewitz D, 2001,

Dehalogenation of DCE in a contaminated soil: Fatty acids &

alcohols as e donors and an apparent requirement for

tetrachloroethene, Appl. Microbiol.Biotech., 55: 239–247.

Vlieg JET VH, Janssen DB, 1992, Bacterial degradation of 3-chloroacrylic

acid and the characterization of cis- and trans-specific

dehalogenases. Biodegradation, 2: 139.


103

Vogel TM, Criddle CS, McCarty PL, 1987, Transformation of halogenated

aliphatic compounds, Environ. Sci. Technol., 21: 722–736.

Vollmer MD, Fischer P, Knackmuss HJ, Schlomann M, 1994, Inability of

muconate cycloisomerases to cause dehalogenation during conversion

of 2- chloro-cis, cis-muconate, J. Bacteriol., 176: 4366–4375.

Wackett LP and Gibson DT, 1988, Degradation of trichloroethylene by toluene

dioxygenase in whole cell studies with Pseudomonas putida F1, Appl.


Wackett LP, Orme-Johnson WH, Walsh CT, 1989, In: Metal Ions and

Bacteria, Beveridge, T.J., and Doyle, R.J., eds., John Wiley and Sons,

165–206.

Wackett LP and Householder SR, 1989, Toxicity of trichlorothylene to

Pseudomonas putida F1, Appl. Environ. Microbiol. 55, 2723–2725.

Wedemeyer G, 1966, Dechlorination of DDT by Aerobacter aerogenes.

Science, 152: 647- 649.

Wedemeyer G, 1967, Dechlorination of 1,1,1-Trichloro-2,2-bis(p-

chlorophenyl)ethane by Aerobacter aerogenes, Applied Microbiology.,

15: 569- 574.

Weightman AJ and Slater H, 1980, Selection of Pseudomonas putida strains

with elevated dehalogenase activities by continuous culture growth on

chlorinated alkanoic acids, J. Gen. Microbiol., 121: 187.

Weightman AJ, Weightman AL, Slater JH, 1982, Stereospecificity of 2-

monochloropropionate dehalogenation by the two dehalogenases of

Pseudomonas putida PP3: evidence for two different dehalogenation

mechanisms, J. Gen. Microbiol., 128: 1755.

Weijer WJ, Hofsteenge J, Vereijken JM, Jekel PA, Beintema JJ, 1982,

Primary structure of p-hydroxybenzoate hydroxylase from

Pseudomonas fluorescens, Biochem.Biophys. Acta., 704: 385.

Wheelis ML and Ornston LN, 1972, Genetic control of enzyme induction in ß-

ketoadipate pathway of Pseudomonas putida: deletion mapping of cat

mutations, J. Bacteriol., 109: 790.


104

Wierenga RK, Terpstra P, Hol WGJ, 1986, Prediction of the occurrence of the

ADP-binding ß-_-ß fold in proteins, using an amino acid sequence

fingerprint, J. Mol. Biol., 187: 101.

Wiesir M, Eberspacher J, Vogler B, Lingens F, 1994, Metabolism of 4-

chlorophenol by Azotobacter spp., GP1: Structure of the meta cleavage

product of 4-chlorocatechnol, FEMS Microbiol. Lett., 166: 73–78.

Winter RB, Yen KM, Ensley BD, 1989, Efficient degradation of

trichloroethylene by a recombinant Escherichia coli, Biotechnology , 7:

282–285.

Wolhfarth G and Diekert G, 1997, Anaerobic dehalogenases, Curr. Opin.

Biotechnol., 8: 290–295.

Xun L and Orser CS, 1991, Purfication and properties of pentachlorophenol

hydroxylase, a flavoprotein from Flavobacterium spp., strain ATCC

39723, J. Bacteriol., 174: 4447–4453.

Xun L, Topp E, Orser CS, 1992, Diverse substrate range of a Flavobacterium

pentachlorophenol hydroxylase and reaction stoichiometries, J.

Bacteriol., 174: 2898–2902 1992.

Yen KM and Gunsalus IC, 1982, Plasmid gene organization:

naphthalene/salicylate oxidation, Proc. Natl. Acad. Sci. U.S.A., 79: 874.

Yen KM and Serdar CM, 1988, Genetics of naphthalene catabolism in

Pseudomonads, Crit. Rev. Microbiol., 15: 247.

Yen KM, Karl MR, Blatt LM, Simon MJ, Winter RB, Fausset PR, Lu HS,

Harcourt AA, Chen, KK, 1991, Cloning and characterization of a

Pseudomonas mendocina KR1 gene cluster encoding toluene-4-

monooxygenase, J. Bacteriol., 173: 5315.

Zang X and Wiegel J, 1990, Sequential anaerobic degradation of 2,4-

dichlorophenol in freshwater sediments, Appl. Environ., Microbiol.,

56: 1119–1127.

Zhuang X, Chen J, Shim H, Bai Z, 2007, New advances in plant growth-

promoting rhizobacteria for bioremediation, Environ. Int., 33: 406-413.

Zon S, Stensel HD, Ferguson JF, 2000, CCl4 degradation: Effect of

microbial growth substrate and Vit B14 content, Environ. Sci. Technol.,

34, 1751–1757.


105

Zylstra GJ, McCombie WR, Gibson DT, Finette BA, 1988, Toluene

degradation by Pseudomonas putida F1: genetic organization of the

tod operon, Appl. Environ. Microbiol., 54: 1498.

Zylstra GJ, Wackett LP, Gibson DT, 1989, Trichloroethylene degradation by

Escherichia coli containing the cloned Pseudomonas putida

dioxygease genes, Appl. Environ. Microbiol., 55: 3162–3166.

Zylstra GJ, McCombie WR, Gibson DT, Finette BA, 1988, Toluene

degradation by Pseudomonas putida F1: genetic organization of the

tod operon, Appl. Environ. Microbiol., 54: 1498.

Zylstra GJ and Gibson DT, 1991, Aromatic Hydrocarbon Degradation: a

Molecular Approach. In: Setlow, J.K. (Ed.) Genetic Engineering,

Plenum Press, New York, p.203.

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Review of Literature - INFLIBNETshodhganga.inflibnet.ac.in/bitstream/10603/13612/12/12...Review of Literature 14 to cis-1, 2 DCE (dichloroethene) and synthesizes energy via electron