microbial degradation of organophosphorus compounds

Microbial degradationoforganophosphorus compoundsBrajesh K. Singh1 & Allan Walker2

1Environmental Sciences, Macaulay Institute, Craigiebuckler, Aberdeen and 2Horticulture Research International, Wellesbourne, Warwick, UK

Correspondence: Brajesh Singh,

Environmental Sciences, Macaulay Institute,

Craigiebuckler, Aberdeen, AB15 8QH, UK.

Tel.: 144 1224 498200; fax: 44 1224

498207; e-mail: [email protected]

Received 16 June 2005; revised 24 November

2005; accepted 6 January 2006.

First published online April 2006.

doi:10.1111/j.1574-6976.2006.00018.x

Editor: Alexander Boronin

Keywords

organophosphorus compounds; microbial

degradation; metabolic pathways; detoxifying

enzymes; genetic basis; biotechnological

aspects.

Abstract

Synthetic organophosphorus compounds are used as pesticides, plasticizers, air

fuel ingredients and chemical warfare agents. Organophosphorus compounds are

the most widely used insecticides, accounting for an estimated 34% of world-wide

insecticide sales. Contamination of soil from pesticides as a result of their bulk

handling at the farmyard or following application in the field or accidental release

may lead occasionally to contamination of surface and ground water. Several

reports suggest that a wide range of water and terrestrial ecosystems may be

contaminated with organophosphorus compounds. These compounds possess

high mammalian toxicity and it is therefore essential to remove them from the

environments. In addition, about 200 000 metric tons of nerve (chemical warfare)

agents have to be destroyed world-wide under Chemical Weapons Convention

(1993). Bioremediation can offer an efficient and cheap option for decontamina-

tion of polluted ecosystems and destruction of nerve agents. The first micro-

organism that could degrade organophosphorus compounds was isolated in 1973

and identified as Flavobacterium sp. Since then several bacterial and a few fungal

species have been isolated which can degrade a wide range of organophosphorus

compounds in liquid cultures and soil systems. The biochemistry of organopho-

sphorus compound degradation by most of the bacteria seems to be identical, in

which a structurally similar enzyme called organophosphate hydrolase or phos-

photriesterase catalyzes the first step of the degradation. organophosphate hydro-

lase encoding gene opd (organophosphate degrading) gene has been isolated from

geographically different regions and taxonomically different species. This gene has

been sequenced, cloned in different organisms, and altered for better activity and

stability. Recently, genes with similar function but different sequences have also

been isolated and characterized. Engineered microorganisms have been tested for

their ability to degrade different organophosphorus pollutants, including nerve

agents. In this article, we review and propose pathways for degradation of some

organophosphorus compounds by microorganisms. Isolation, characterization,

utilization and manipulation of the major detoxifying enzymes and the molecular

basis of degradation are discussed. The major achievements and technological

advancements towards bioremediation of organophosphorus compounds, limita-

tions of available technologies and future challenge are also discussed.

Introduction

The excessive use of natural resources and large scale

synthesis of xenobiotic compounds have generated a num-

ber of environmental problems such as contamination of air,

water and terrestrial ecosystems, harmful effects on different

biota, and disruption of biogeochemical cycling. At the

present time, the most widely used pesticides belong to the

organophosphorus group. The first organophosphorus in-

secticide, tetraethyl pyrophosphate, was developed and used

in 1937 (Dragun et al., 1984). At the same time, two

chemical warfare agents (also called nerve agents), Tabun

and Sarin, were developed and produced. Later, several

other organophosphorus pesticides were developed and

commercialized. These pesticides are widely used world-

wide to control agricultural and household pests. Overall,

organophosphorus compounds account for �38% of total

pesticides used globally (Post, 1998). In the USA alone over

40 million kilos of organophosphorus are applied annually

(Mulchandani et al., 1999a; EPA, 2004). Glyphosate and

FEMS Microbiol Rev 30 (2006) 428–471c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

chlorpyrifos are the most widely used in the US and account

for 20% and 11% of total pesticide use, respectively (EPA,

2004). Organophosphorus compound poisoning is a world-

wide health problem with around 3 million poisonings

and 200 000 deaths annually (Karalliedde & Senanayake,

1999; Sogorb et al., 2004). The compounds have been

implicated in several nerve and muscular diseases in human

beings. Their acute adverse effects have been discussed by

Colborn et al. (1996) and Ragnarsdottir (2000). Immuno-

toxicity of organophosphorus compounds towards human

beings and wild-life has been reviewed by Galloway &

Handy (2003).

Continuous and excessive use of organophosphorus

compounds has led to the contamination of several ecosys-

tems in different parts of the world (EPA, 1995; McConnell

et al., 1999; Cisar & Snyder, 2000; Tse et al., 2004). For an

example, surveys revealed that 100% of sampled catchments

in Scotland and 75% of sampled aquatic sites in Wales were

contaminated with organophosphorus compounds used in

sheep dips (Boucard et al., 2004). Several organophosphorus

compounds are used on animals for the control of body

pests as several of them are fat soluble and can thus enter the

body readily through the skin and potentially find their way

into meat and milk (MAFF/HSE, 1995). Contamination of

grains, vegetables and fruits with organophosphorus com-

pounds is also well documented (Pesticide Trust 1996;

National Consumer Council 1998). Another potential and

more dangerous source of organophosphorus contamina-

tion comes from chemical warfare agents. About 200 000

tons of extremely toxic organophosphorus chemical warfare

agents such as Sarin, Soman, and VX were manufactured

and are stored. As required by the Chemical Weapon

Convention (CWC) 1993, these stocks must be destroyed

within 10 years of ratification by the member states. Use of

micro-organisms in detoxification decontamination of or-

ganophosphorus compounds is considered a viable and

environment friendly approach.

The available literature on the microbial degradation of

xenobiotics indicates that most studies have considered

three aspects:

(1) The fundamental basis of biodegradation.

(2) Evolution and transfer of such activities among micro-

organisms.

(3) Bioremediation techniques to detoxify contaminated

environments (Singh et al., 1999).

However, the use of micro-organisms for bioremediation

requires an understanding of all physiological, microbiolo-

gical, ecological, biochemical and molecular aspects in-

volved in pollutant transformation (Iranzo et al., 2001).

There are two types of xenobiotics that cause environ-

mental concerns: (1) compounds that are persistent and

therefore provide long exposure to non-target organisms

such as lindane and DDT, and (2) compounds that are

biodegradable but mobile in soil and are toxic and therefore

have the potential to pollute ground water, such as carbo-

furan. Extensive and repeated use of the same pesticide

without any crop or pesticide rotation for a number of years

has occasionally resulted in unexpected failures to control

the target organisms. It has been demonstrated that a

fraction of the soil biota can develop the ability rapidly to

degrade certain soil-applied pesticides. This phenomenon

has been described as enhanced or accelerated biodegrada-

tion (Walker & Suett, 1986). The first evidence of biodegra-

dation of pesticide affecting its efficacy was reported in 1971

(Sethunathan, 1971). However, it was not until the early to

mid 1980s that the wider implication of enhanced bio-

degradation became observable in the field (Walker & Suett,

1986) and since then this phenomenon has been reported

for several other pesticides such as isofenphos (Chapman

et al., 1986), fenamiphos (Stiriling et al., 1992) and etho-

prophos (Karpouzas et al., 1999).

The practical significance of enhanced bio-degradation

depends on a number of interactive factors like the use of the

pesticides (soil or foliage applied), the frequency of use, the

interval between successive applications and the stability of

the active microflora without the presence of pesticides

(Kaufman et al., 1985). Recently, soil pH has been impli-

cated as a factor in enhanced degradation of atrazine in

different soils (Houot et al., 2000). This hypothesis has been

supported by recent reports of high enzymatic activity

(Acosta-Martinez & Tabatabai, 2000) and higher bacterial

activity at higher soil pH (Vidali, 2001). Sims et al. (2002)

suggested that soil pH may influence the rate of degradation

by affecting the uptake of the herbicide by soil micro-

organisms. The problem of enhanced bio-degradation be-

came more acute, following the observation that a pesticide

can be degraded rapidly in soil from a field to which it had

never been applied before but which had been exposed to a

pesticide from the same chemical group (Prakash et al.,

1996). This phenomenon is known as cross-adaptation.

Cross-adaptation of enhanced biodegradation has been

reported within many groups of pesticide, such as the

carbamates (Morel-Chevillet et al., 1996), dicarboximides

(Mitchell & Cain, 1996) and isothiocyanates (Warton et al.,

2002). On the other hand, only limited cross-adaptation for

enhanced biodegradation within the organophosphorus

class has been reported (Racke & Coats, 1988; Singh et al.,

2005). Cross-adaptation within groups is unpredictable and

may occur only in one direction. The positive side of this

problem is that micro-organisms isolated for degradation of

one compound can be used for bioremediation of other

compounds for which no known degrading microbial

system is known. This aspect is well established for organo-

phosphorus compounds where a parathion-degrading bac-

terium was able to degrade a wide range of other structurally

similar compounds including chemical warfare agents.

FEMS Microbiol Rev 30 (2006) 428–471 c� 2006 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

429Microbial degradation of organophosphorus compounds

Isolation of pesticide degrading microorganisms is impor-

tant for three main reasons:

(1) To determine the mechanism of the intrinsic process of

microbial metabolism.

(2) To understand the mechanisms of gene/enzyme evolu-

tion.

(3) To use these microbes for the detoxification and decon-

tamination of polluted aquatic and terrestrial environ-

ments (bioremediation).

Several microorganisms have been isolated which are able

to utilize pesticides as a source of energy. There are some

examples of fungi including Trametes hirsutus, Phanero-

chaete chrysosporium, Phanerochaete sordia and Cyathus

bulleri that are able to degrade lindane and other pesticides

(Singh & Kuhad, 1999, 2000; Singh et al., 1999). However,

most evidence suggests that soil bacteria are the principal

components responsible for enhanced bio-degradation

(Walker & Roberts, 1993). Several pure bacterial isolates

with the ability to use specific pesticides as a sole source of

carbon, nitrogen or phosphorus have been isolated (Singh

et al., 1999, 2000).

On numerous occasions, mixed bacterial cultures with

pesticide degradation ability are isolated but their individual

components are unable to utilize the chemical as an energy

source when purified (Shelton & Somich, 1988; Mandel-

baum et al., 1993; De Souza et al., 1993; Roberts et al., 1993);

an example is the organophosphorus nematicide fenami-

phos (Ou & Thomas, 1994; Singh et al., 2003b). Several

other studies failed to obtain micro-organisms capable of

growing on specific chemicals. However, this failure does

not exclude biological involvement in degradation and

could be attributed to the selection and composition of the

liquid media under artificial environments, strains requiring

special growth factors, or a major role of non-culturable

microorganisms (Walker & Roberts, 1993). A recent report

of growing previously non-culturable bacteria in the labora-

tory with a simulated natural environment (Kaeberlein

et al., 2002) may lead to isolation and characterization of

several new chemical-degrading bacteria.

The main aim of this article is to review the metabolic

pathways involved in organophosphorus compound degrada-

tion. Our understanding of the molecular basis of organopho-

sphorus degradation has progressed dramatically in recent

years. Additional information has become available by gen-

ome sequencing of several microorganisms and advancement

in molecular techniques. There is growing interest in devel-

oping biotechnological methods for clean up of contaminated

water and soil with organophosphorus compounds and to aid

in the destruction of large amounts of nerve agents. In this

article we also critically review recent biotechnological ad-

vancements in the development of bio-catalysts and bio-

sensors for organophosphorus compounds and their possible

application in bioremediation of contaminated ecosystems.

Chemistry and toxicology oforganophosphorus compounds

Most organophosphorus compounds are ester or thiol

derivatives of phosphoric, phosphonic or phosphoramidic

acid. Their general formula is presented in Fig. 1. R1 and R2

are mainly the aryl or alkyl group, which can be directly

attached to a phosphorus atom (phosphinates) or via

oxygen (phosphates) or a sulphur atom (phosphothioates).

In some cases, R1 is directly bonded with phosphorus and R2

with an oxygen or sulfur atom (phosphonates or thion

phosphonates, respectively). At least one of these two groups

is attached with un-, mono- or di-substituted amino groups

in phosphoramidates. The X group can be diverse and may

belong to a wide range of aliphatic, aromatic or heterocyclic

groups. The X group is also known as a leaving group

because on hydrolysis of the ester bond it is released from

phosphorus (Fig. 1) (Sogorb & Vilanova, 2002).

The mode of action of organophosphorus compounds

includes inhibition of neurotransmitter acetylcholine break-

down. Acetylcholine is required for the transmission of

nerve impulses in the brain, skeletal muscles and other areas

(Toole & Toole, 1995). However, after the transmission of

the impulse, the acetylcholine must be hydrolyzed to avoid

overstimulating or overwhelming the nervous system. This

breakdown of the acetylcholine is catalyzed by an enzyme

called acetylcholine esterase. Acetylcholine esterase converts

acetylcholine into choline and acetyl CoA by binding the

substrate at its active site at serine 203 to form an enzyme

substrate complex. Further reactions involve release of cho-

line from the complex and then rapid reaction of acylated

enzymes with water to produce acetic acid and the

Fig. 1. General formula of organophosphorus compounds and major

pathway of degradation.


430 B.K. Singh & A. Walker

regenerated acetylcholine esterase. It has been estimated that

one enzyme can hydrolyze 300 000 molecules of acetylcho-

line every minute (Ragnarsdottir, 2000).

Organophosphorus compounds inhibit the normal activ-

ity of the acetylcholine esterase by covalent bonding to the

enzyme, thereby changing its structure and function. They

bind to the serine 203 amino acid active site of acetylcholine

esterase. The leaving group binds to the positive hydrogen of

His 447 and breaks off the phosphate, leaving the enzyme

phosphorylated. The regeneration of phosphorylated acet-

ylcholine esterase is very slow and may take hours or days,

resulting in accumulation of acetylcholine at the synapses.

Nerves are then overstimulated and jammed (Manahan,

1992). This inhibition causes convulsion, paralysis and

finally death for insects and mammals (Ragnarsdottir,

2000).

Microbial degradation oforganophosphorus compounds

Use of organochlorine pesticides such as dichloro-diphenyl-

trichloroethane (DDT), lindane, etc., has been reduced

drastically in developed countries due to their long persis-

tence, tendency towards bioaccumulation and potential

toxicity towards non-target organisms. This group of com-

pounds has been replaced by the less persistent and more

effective organophosphorus compounds. However, most of

the organophosphorus compounds possess high mamma-

lian toxicity. Among the organophosphorus compounds,

glyphosate, chlorpyrifos, parathion, methyl parathion, dia-

zinon, coumaphos, monocrotophos, fenamiphos and pho-

rate have been used extensively and their efficacy and

environmental fate have been studied in detail. The chemical

and physical properties of some of these compounds are

listed in Table 1. The phosphorus is usually present either as

a phosphate ester or as a phosphonate. Being esters they

have many sites which are vulnerable to hydrolysis. The

principal reactions involved are hydrolysis, oxidation, alky-

lation and dealkylation (Singh et al., 1999). Microbial

degradation through hydrolysis of P-O-alkyl and P-O-aryl

bonds is considered the most significant step in detoxifica-

tion (Fig. 1). Both co-metabolic and bio-mineralization of

organophosphorus compounds by isolated bacteria have

been reported. A list of micro-organisms capable of degrad-

ing these compounds is presented in Table 2.

Hydrolysis of organophosphorus compounds leads to a

reduction in their mammalian toxicity by several orders of

magnitude. Since most of the research has been directed

towards detoxification, studies on the further metabolism of

the phosphorus containing products have not been exten-

sive. Hypothetical phospho-ester hydrolysis steps can be

postulated, yielding mono-ester and finally inorganic phos-

phate, but this pathway has not been specifically studied.

Analogous phospho-monoesterase and diesterase, which

degrade methyl and dimethyl phosphate, respectively, have

been reported in Klebsiella aerogenes (Wolfenden & Spence,

1967) and are produced only in the absence of inorganic

phosphate from the growth medium. The final enzyme in

the postulated degradative pathway is bacterial alkaline

phosphatase, which can hydrolyze simple monoalkyl phos-

phates and is also regulated by the level of phosphate

available to the cell (Wolfenden & Spence, 1967). A similar

mechanism of metabolism has been reported for phospho-

nates (Kertesz et al., 1994a). The way in which metabolism is

regulated depends very strongly on what role the organo-

phosphorus compound plays for the particular organisms

studied. Most often these compounds are used to supply

only a single element (carbon, phosphorus or sulfur) and

the relevant gene cannot be expressed as a response to

starvation for another of these elements (Kertesz et al.,

1994a). For example, a strain of Pseudomonas stutzeri

isolated to utilize parathion as a carbon source released the

diethylphosphorothioanate products quantitatively and

could not metabolize them further, even when alternative

source of phosphorus or sulfur were removed (Daughton &

Hsieh, 1977). Similarly, a variety of isolates that could use

phosphorothionate and phosphorodithionate pesticides as a

sole source of phosphorus were unable to utilize these

compounds as a source of carbon (Rosenberg & Alexander,

1979). Shelton (1988) isolated a consortium that could use

diethylthiophosphoric acid as a carbon source but was

unable to utilize it as a source of phosphorus or sulfur.

Kertesz et al. (1994a) explained possible underlying reasons

for this phenomenon. They suggested that the conditions

under which environmental isolates enriched were crucial in

selecting for strains not only with the desired degradative

enzyme systems but also with specific regulation mechan-

isms for the degradation pathways.

Table 1. History, toxicity and half-life of some organophosphorus

pesticides

Name Type

Year of

introduction

Mammalian

LD50

(mg kg�1)

Half-life

soil

(days)

Chlorpyrifos Insecticide 1965 135–163 10–120

Parathion Insecticide 1947 2–10 30–180

Methyl parathion Insecticide 1949 3–30 25–130

Glyphosate Herbicide 1971 3530–5600 30–174

Coumaphos Acaricide 1952 16–41 24–1400

Fenamiphos Nematicide 1967 6–10 28–90

Monocrotophos Insecticide 1965 18–20 40–60

Dicrotophos Insecticide 1965 15–22 45–60

Diazinon Insecticide 1953 80–300 11–21

Dimethoate Insecticide 1955 160–387 2–41

Fenitrothion Insecticide 1959 1700 12–28

Ethoprophos Nematicide 1966 146–170 3–30



Table 2. Microorganisms isolated for the degradation of organophosphorus compounds

Compound Microorganisms Mode of degradation Reference

Chlorpyrifos Bacteria

Enterobacter sp. Catabolic (C, P) Singh et al. (2003c)

Flavobacterium sp. ATCC27551 Co-metabolic Mallick et al. (1999)

Pseudomonas diminuta Co-metabolic Serdar et al. (1982)

Micrococcus sp. Co-metabolic Guha et al. (1997)

Fungi

Phanerochaete chrysosporium Catabolic (C) Bumpus et al. (1993)

Hypholama fascicularae ND Bending et al. (2002)

Coriolus versicolor ND Bending et al. (2002)

Aspergillus sp. Catabolic (P) Obojska et al. (2002)

Trichoderma harzianum Catabolic (P) Omar (1998)

Pencillium brevicompactum Catabolic (P) Omar (1998)

Parathion Bacteria

Flavobacterium sp. ATCC27551 Co-metabolic Sethunathan & Yoshida (1973)


Pseudomonas stutzeri Co-metabolic Daughton & Hsieh (1977)

Arthrobacter spp. Co-metabolic Nelson et al. (1982)

Agrobacterium radiobacter Co-metabolic Horne et al. (2002b)

Bacillus spp. Co-metabolic Nelson et al. (1982)

Pseudomonas sp. Catabolic (C, N) Siddaramappa et al. (1973)

Pseudomonas spp. Catabolic (P) Rosenberg & Alexander (1979)

Arthrobacter sp. Catabolic (C) Nelson et al. (1982)

Xanthomonas sp. Catabolic (C) Rosenberg & Alexander (1979)

Methyl parathion

Pseudomonas sp. Co-metabolic Chaudry et al. (1988)

Bacillus sp. Co-metabolic Sharmila et al. (1989)

Plesimonas spM6 Co-metabolic Zhongli et al. (2001)

Pseudomonas putida Catabolic (C) Rani & Lalitha-kumari (1994)

Pseudomonas sp. A3 Catabolic (C, N) Zhongli et al. (2002)

Pseudomonas sp. WBC Catabolic (C, N) Yali et al. (2002)

Flavobacterium balustinum Catabolic (C) Somara & Siddavattam (1995)

Glyphosate Bacteria

Pseudomonas ssp. Catabolic (P) Kertesz et al. (1994a)

Alcaligene sp. Catabolic (P) Tolbot et al. (1984)

Bacillus megaterium 2BLW Catabolic (P) Quinn et al. (1989)

Rhizobium sp. Catabolic (P) Liu et al. (1991)

Agrobacterium sp. Catabolic (P) Wacket et al. (1987)

Arthrobacter sp. GLP Catabolic (P) Pipke et al. (1987)

Arthrobacter atrocyaneus Catabolic (P) Pike & Amrhein (1988)

Geobacillus caldoxylosilyticus T20 Catabolic (P) Obojska et al. (2002)

Flavobacterium sp. Catabolic (P) Balthazor & Hallas (1986)

Fungi

Penicillium citrium Co-metabolic Pothuluri et al. (1998)

Pencillium natatum catabolic (P) Pothuluri et al. (1992)

Penicillium chrysogenum Catabolic (N) Klimek et al. (2001)

Trichoderma viridae Catabolic (P) Zboinska et al. (1992b)

Scopulariopsis spand Catabolic (P) Zboinska et al. (1992b)

Aspergillus niger Catabolic (P) Zboinska et al. (1992b)

Alternaria alternata Catabolic (N) Lipok et al. (2003)

Coumaphos

Nocardiodes simplex NRRL B24074 Co-metabolic Mulbry (2000)

Agrobacterium radiobacter P230 Co-metabolic Horne et al. (2002b)

Pseudomonas monteilli Co-metabolic Horne et al. (2002c)

Flavobacterium sp. Co-metabolic Adhya et al. (1981)


Nocardia strain B-1 Catabolic (C) Mulbry (1992)



Chlorpyrifos

Chlorpyrifos (O,O-diethyl O-(3,5,6-trichloro-2-pyridyl)

phosphorothioate) is one of the most widely used insecti-

cides effective against a broad spectrum of insect pests of

economically important crops. It is effective by contact,

ingestion and vapour action but is not systemically active. It

is used for the control of mosquitoes (larvae and adults),

flies, various soil and many foliar crop pests and household

pests. It is also used for ectoparasite control on cattle and

sheep. It has low solubility in water (2 mg L�1) but is readily

soluble in most organic solvents. It has a high soil sorption

co-efficient (Racke, 1993) and is stable under normal storage

conditions. Chlorpyrifos is defined as a moderately toxic

compound having acute oral LD50; 135–163 mg kg�1 for rat

and 500 mg kg�1 for guinea pig.

The environmental fate of chlorpyrifos has been studied

extensively. Degradation in soil involves both chemical

hydrolysis and microbial activity. The half-life of chlorpyr-

ifos in soil varies from 10 to 120 days (Getzin, 1981; Racke

et al., 1988) with 3,5,6-trichloro-2-pyridinol (TCP) as the

major degradation product. This large variation in half-life

has been attributed to different environmental factors, the

most important of which are soil pH, temperature, moisture

content, organic carbon content and pesticide formulation

(Getzin, 1981a, b; Chapman & Chapman, 1986). Initially,

the high rate of chlorpyrifos degradation in soils with

alkaline pH was attributed to chemical hydrolysis. Later,

Racke et al. (1996) concluded that the relationship between

high soil pH and chemical hydrolysis was weak and that

other factors like soil silt content might be important in

determining environmental fate.

Unlike other organophosphorus compounds, chlorpyri-

fos has been reported to be resistant to the phenomenon of

enhanced degradation (Racke et al., 1990). There have been

no reports of enhanced degradation of chlorpyrifos since its

first use in 1965 until recently. It was suggested that the

accumulation of TCP, which has anti-microbial properties,

acts as a buffer in the soil and prevents the proliferation of

chlorpyrifos degrading microorganisms (Racke et al., 1990).

However, Robertson et al. (1998) suggested that chemical

hydrolysis of chlorpyrifos and enhanced degradation of TCP

can result in loss of efficacy of the insecticide against

termites in sugar cane fields in Australia. Attempts to

introduce enhanced degradation in the laboratory or in the

field by repeated application have failed (Racke et al., 1990;

Mallick et al., 1999).

In recent experiments, we found that the degradation of

chlorpyrifos was very slow in acidic soils but that the rate of

degradation increased considerably with an increase in soil

pH. However, in 90 days of incubation, there was no

difference between soils in release of 14CO2 from the

pyridine ring despite the large differences in degradation

rate. Repeated applications of chlorpyrifos did not affect

Monocrotophos

Pseudomonas spp. Catabolic (C) Bhadbhade et al. (2002b)

Bacillus spp. Catabolic (C) Rangaswamy & Venkateswaralu (1992)

Arthrobacter spp. Catabolic (C) Bhadbhade et al. (2002b)

Pseudomonas mendocina Catabolic (C) Bhadbhade et al. (2002a)

Bacillus megaterium Catabolic (C) Bhadbhade et al. (2002b)

Arthrobacter atrocyaneus Catabolic (C) Bhadbhade et al. (2002b)

Pseudomonas aeruginosa F10B Catabolic (P) Singh & Singh (2003)

Clavibacter michiganense SBL11 Catabolic (P) Singh & Singh (2003)

Fenitrothion

Flavobacterium sp. Co-metabolic Adhya et al. (1981)

Arthrobacter aurescenes TW17 Catabolic (C) Ohshiro et al. (1996)

Burkholderia sp. NF100 Catabolic (C) Hayatsu et al. (2000)

Diazinon

Flavobacterium sp. Catabolic (P) Sethunathan & Yoshida (1973)

Pseudomonas spp. Co-metabolic Rosenberg & Alexander (1979)

Arthrobacter spp. Co-metabolic Barik et al. (1979)

Chemical warfare agents

G Agent Pseudomonas diminuta Co-metabolic Mulbry & Rainina (1998)

Altermonas spp. Co-metabolic DeFrank et al. (1993)

V Agent Pseudomonas diminuta Co-metabolic Mulbry & Rainina (1998)

Pleurotus ostreatus (fungus) Co-metabolic Yang et al. (1990)

Symbol in brackets after mode of degradation represents the type of nutrient that the pesticide provides to degrading microorganisms. C, carbon; N,

nitrogen; P, phosphorus.



either the degradation rate or degradation kinetics, suggest-

ing that repeated treatment did not result in enhanced

degradation. Fumigation of soil samples completely inhib-

ited hydrolysis of chlorpyrifos, suggesting an involvement of

soil micro-organisms (Singh et al., 2003c). Chlorpyrifos has

been reported previously to be resistant to enhanced degra-

dation. Given the tremendous adaptability of the soil

microbial community for degradation of a wide variety of

synthetic compounds, Racke et al. (1990) cited three possi-

ble reasons why a specific pesticide might not be susceptible

to enhanced degradation. One possibility is an inability of

the microflora to initiate degradation of the parent pesticide

easily. This may be due to factors such as steric hindrance of

enzymes by functional groups, electronic stability against

hydrolysis or lack of weak links in the molecule (Alexander,

1965; Niemi et al., 1987). The pesticide may also be unavail-

able for uptake and degradation by soil microorganisms due

to strong sorption to organic surfaces in the soil (Orgam

et al., 1985). However, these reasons cannot explain the

present results because chlorpyrifos is rapidly hydrolyzed by

the soil bacterial community in alkaline soils. The second

possibility is that the soil environmental conditions may in

some way inhibit the development or expression of en-

hanced degradation. This also cannot explain the present

results because repeated treatment of the same soil samples

resulted in enhanced degradation of fenamiphos (Singh

et al., 2003b). A third possibility is that the soil micro-

organisms cannot beneficially catabolize pesticide metabo-

lites. In these circumstances co-metabolism may occur (e.g.

hydrolysis of parent pesticides), but the microbial metabo-

lism of the degradation products is not possible. This is the

case with such relatively recalcitrant pesticides as DDT and

alachlor, which are converted to products that are them-

selves quite resistant to further metabolism (Tiedje &

Hagedorn, 1975). From our experiments we concluded that

in high pH soils, the microbial community transforms

chlorpyrifos co-metabolically into TCP. However, TCP con-

tains three chlorine atoms on the pyridinol ring. To break

this ring, chlorine atoms have to be removed (Feng et al.,

1997), and free chlorine has toxic effects on the micro-

organisms. Thus TCP metabolism may be toxic to micro-

organisms. Similar results were obtained by Price et al.

(2001) in a field where degradation of chlorpyrifos was

strongly related with soil pH but degradation was mediated

by soil micro-organisms. Later, Singh et al. (2003c) sug-

gested that chlorpyrifos is degraded by non-specific and

non-inducible enzyme systems produced in high pH soils.

This suggests that chlorpyrifos is co-metabolically hydro-

lyzed to TCP and that because the TCP has toxic effects,

normally enhanced degradation does not occur. Although

Shelton & Doherty (1997) in their model proposed a

significant role of bioavailability in degradation of xenobio-

tics, the toxic effect of TCP seems to be a realistic explana-

tion of its resistance to enhanced degradation because TCP

has high water solubility and therefore is bioavailable for the

degradation. However, repeated treatment with chlorpyrifos

over many years in an Australian soil resulted in develop-

ment of some opportunist microorganisms with the cap-

ability to use the toxic compound as has been reported with

organochlorine compounds (Robertson et al., 1998; Singh

et al., 2000). This adaptation can provide them with a

competitive advantage over other microbes in terms of

sources of energy. Further studies found higher copy num-

bers of opd (organophosphate degrading) gene in higher pH

soils (Singh et al., 2003a, c).

In most cases described to date, the aerobic bacteria tend

to transform chlorpyrifos by hydrolysis to produce

diethylthiophosphoric acid (DETP) and TCP, which in turn

accumulate in the culture medium without further metabo-

lism. This transformation reaction removes chlorpyrifos and

its mammalian toxicity but yields compounds that are not

metabolized by the microorganisms that produce them

(Richins et al., 1997; Mallick et al., 1999; Horne et al.,

2002b; Wang et al., 2002b).

Chlorpyrifos has been reported to be degraded co-meta-

bolically in liquid media by Flavobacterium sp. and Pseudo-

monas diminuta, which were initially isolated from a

diazinon treated field and by parathion enrichment, respec-

tively (Sethunathan & Yoshida, 1973; Serdar et al., 1982).

However, these microbes do not utilize chlorpyrifos as a

source of carbon. A Micrococcus sp. was isolated from a

malathion enriched soil which was later reported to degrade

chlorpyrifos in liquid media (Guha et al., 1997). We have

isolated an Enterobacter sp. from a soil from Australia

showing enhanced degradation of chlorpyrifos. This bacter-

ium degrades chlorpyrifos to DETP and TCP and utilizes

DETP as a source of carbon and phosphorus (Singh et al.,

2003c, 2004). Cook et al. (1978a) isolated several bacteria

from sewage sludge that were able to use dialkylthiopho-

sphonic acid as a sole source of phosphorus. One of these

organisms, Pseudomonas acidovorans, was able to use DETP

as a sole source of sulfur (Cook et al., 1980). Another

significant observation was the utilization of organopho-

sphorus insecticides as a source of phosphorus by Entero-

bacter sp. (Singh et al., 2003c, 2004). Sethunathan & Yoshida

(1973) isolated a Flavobacterium sp. that could use diazinon

as a source of carbon. However, Flavobacterium was not able

to use other organophosphorus pesticides as a source of

either phosphorus or carbon. Similarly, a variety of isolates

that could use phosphorothionate or phosphorodithionate

compounds as a sole source of phosphorus were unable to

degrade these compounds as a source of carbon (Rosenberg

& Alexander, 1979). Shelton (1988) isolated a consortium

that could use DETP as a carbon source but was unable to

degrade it when presented as source of phosphorus or sulfur.

It is believed that the conditions under which environmental



isolates are enriched are crucial in selecting for strains not

only with the desired degradative enzymes systems, but also

with the specific regulation mechanisms for the degradation

pathways (Kertesz et al., 1994a).

Studies on further metabolism and identification of

intermediate products of the phosphorus containing pro-

ducts have not been extensive. The postulated pathway steps

include hydrolysis, yielding monoester and finally inorganic

phosphate (Fig. 2). Bacterial phosphodiesterase has been

purified from a wide range of organisms including Escher-

ichia coli (Imamura et al., 1996), Haemophilus influenzae

(Macfadyen et al., 1988), and Burkholderia caryophylli

PG2982 (Dotson et al., 1996). The phosphodiesterase from

the first two bacteria are similar in sequence and both

moderate intracellular cyclic AMP levels. However, the

phosphodiesterase from B. caryophilli has a different se-

quence from that in the first two bacteria (Dotson et al.,

1996). This enzyme could not be assigned a clear function

but was thought to play a role in xenobiotic degradation

pathways because it degraded glycerol glyphosate. However,

until recently no phosphodiesterase had been isolated or

characterized which could utilize xenobiotic degradation

products such as diethyl phosphate and diethyl phospho-

nate. A novel phosphodiesterase was isolated and cloned

from Delftia acidovorans which has both mono- and di-

esterase activity (Tehara & Keasling, 2003). This enzyme

allows D. acidovorans to use diethyl phosphonate as a sole

source of phosphorus under phosphorus limiting condi-

tions. The final enzyme in the postulated degradative path-

way is alkaline phosphatase, which can hydrolyze simple

monoalkyl phosphates (Neidhardt et al., 1996).

Since only one bacterium has been isolated so far which

can degrade TCP in liquid medium, little literature is

available on microbial metabolism of TCP. Feng et al.

(1997) isolated a Pseudomonas sp. which can mineralize

TCP in liquid medium. Later the same group, on the basis of

combined experiments with photolysis and microbial de-

gradation, suggested that TCP was metabolized by a Pseu-

domonas sp. by a reductive dechlorination pathway (Feng

et al., 1998). In this pathway, TCP is first reductively

dechlorinated into chlorodihydro-2-pyridone, which is

further dechlorinated to tetra-hydro-2-pyridone. Ring clea-

vage of this compound resulted in formation of maleamide

semialdehyde, which is metabolized to water, carbon diox-

ide, and ammonium ions. Microbial degradation of analo-

gous compounds such as pyridine and hydroxypyridine has

been researched and reviewed extensively (Shukla, 1984;

Sims & O’Loughlin, 1989; Kaiser et al., 1996). Several micro-

organisms were reported to degrade hydroxypyridine (Kai-

ser et al., 1996). Cain et al. (1974) reported that 2- or 3-

hydroxypyridine was oxidized to 2,5-dihydroxypyridine and

production of maleamic acid occurred later through ring

cleavage. Oxygen atoms used to transform 4-hydroxypyr-

idine via 3,4-dihydroxypyridine were derived from water

molecules by hydroxypyridine hydrolase (Watson et al.,

1974). It is likely that TCP is metabolized in a similar

manner as one of the metabolites of TCP was identified to

have similar structure to 2-hydroxypyridine.

Fungal mineralization of chlorpyrifos by Phanerochaete

chrysosporium was reported by Bumpus et al. (1993).

Chlorpyrifos was hydrolyzed and then the pyridinyl ring

underwent cleavage before being converted to carbon diox-

ide and water. Degradation of chlorpyrifos in ‘biobed’

composting substrate by two other white-rot fungi, Hypho-

loma fascicularae and Coriolus versicolor, was observed

(Bending et al., 2002). Degradation of a wide range of

xenobiotic compounds by white-rot fungi is well documen-

ted (Kuhad et al., 1997; Singh & Kuhad, 1999, 2000; Singh

et al., 1999). These organisms have been reported to degrade

several persistent aromatic compounds by ring cleavage

(Armenante et al., 1994; Reddy & Gold, 2000). The multi-

step pathway of pentachlorphenol degradation by the white-

rot fungus Phanerochaete chrysosporium is initiated by lignin

peroxidase and manganese peroxidase, producing tetra-

chloro-1-4-benzoquinone, which is further metabolized by

two parallel but cross-linked pathways. The tetrachloroben-

zoquinone is reduced to tetrachlorodihydroxybenzene,

which can undergo four successive dechlorinations to pro-

duce 1,4-hydroquinone. This is then hydroxylated to pro-

duce the final aromatic metabolite, 1,2,4-trihydroxybenzene.

Alternatively the tetrachlorobenzoquinone converts to

2,3,5-trichlorotrihydroxybenzene, which undergoes succes-

sive reductive dechlorination to produce 1,2,4-trihydroxy-

benzene. At several points, hydroxylation reaction converts

chlorinated dihydroxybenzene to chlorinated trihydroxy-

benzene, linking two pathways. The 1,2,4-trihydroxyben-

zene is ring cleaved to produce CO2 and water (Reddy &

Gold, 2000). Mineralization of TCP by white-rot fungi is

possible via reductive de-chlorination. White-rot fungi have

been reported previously to use this transformation step to

degrade other chlorinated compounds such as pentachlor-

ophenol (Aiken & Logan, 1996) and hexachlorocyclohexane

(Mougin et al., 1996; Singh & Kuhad, 1999, 2000). Degrada-

tion of several polychlorinated compounds by white-rot

fungi suggests that they produce a range of isoenzymes with

a wide range of substrate specificity. Several species of

Aspergillus, Trichoderma harzianum and Penicillium brevi-

compactum were reported to utilize chlorpyrifos as sources

of phosphorus and sulfur (Omar, 1998) (Table 1). On the

basis of the above discussion, the authors propose possible

pathways for microbial degradation of chlorpyrifos (Fig. 2).

Parathion

Parathion (O,O-diethyl-O-p-nitrophenyl phosphorothio-

ate) is one of the most toxic insecticides registered with the



US Environmental Protection Agency (EPA). Extreme toxi-

city with ease of exposure has resulted in numerous human

and non-target species deaths in several developing coun-

tries (McConnell et al., 1999). The microbial degradation of

parathion has received extensive attention among the orga-

nophosphorus compounds because of its widespread use

and the ready detection of its hydrolytic product (p-nitro-

phenol). Parathion is rapidly degraded in biologically active

Fig. 2. Proposed pathways for chlorpyrifos

degradation by microorganisms. The scheme is

based on articles cited in the text. When the

conversion of one compound to another is

believed to occur through a series of inter

mediates, the steps are indicated by dotted

arrows. DETP, diethylthiophosphate; TCP,

trichloropyridinol.



soil. A proportional increase in the bacterial population in

soils was observed with an increase in the concentration of

parathion added (Nelson, 1982). Flooded soil conditions

favoured hydrolysis of parathion and release of 14CO2 from

ring labelled parathion in the rhizosphere of rice seedlings

(Reddy & Sethunathan, 1983).

Several species of bacteria have been isolated either from

parathion enrichment or other organophosphate enriched

environments, which can hydrolyze parathion (Table 2)

(Munnecke et al., 1982; Kertesz et al., 1994a; Racke et al.,

1996). Both mineralization, where parathion was used as a

source of carbon (Munnecke & Hsieh, 1976; Rani & Lalitha-

kumari, 1994) or phosphorus (Rosenberg & Alexander,

1979), and co-metabolic hydrolysis (Serdar et al., 1982;

Horne et al., 2002b) have been reported. Sethunathan &

Yoshida (1973) isolated the first organophosphorus degrad-

ing bacterium, Flavobacterium sp., that could degrade para-

thion and diazinon. Siddaramappa et al. (1973) isolated a

Pseudomonas sp. that was able to hydrolyze parathion and

utilize the hydrolysis product p-nitrophenol as a carbon or

nitrogen source. Later, P. stutzeri was isolated, which can

hydrolyze parathion although p-nitrophenol was metabo-

lized by a separate bacterium (Daughton & Hsieh, 1977).

Rosenberg & Alexander (1979) isolated two Pseudomonas

ssp. that were able to hydrolyze a number of organopho-

sphorus compounds including parathion, and to use the

ionic cleavage products as a sole source of phosphorus.

Several species of Bacillus and Arthrobacter have been

isolated that were capable of hydrolyzing parathion; one of

the Arthrobacter strains was also able to utilize p-nitrophenol

as a sole source of carbon (Nelson, 1982). A Pseudomonas sp.

and a Xanthomonas sp. were isolated which can hydrolyze

parathion and can further metabolize p-nitrophenol (Tche-

let et al., 1993). A Moraxella sp. can use p-nitrophenol as the

sole source of carbon and nitrogen (Spain & Gibson, 1991).

This bacterium degrades p-nitrophenol to p-benzoquinone

using the enzyme p-nitrophenol monooxygenase. p-Benzo-

quinone is transformed to hydroquinone by a reductase

(Spain & Gibson, 1991). Candida parapsilosis has been

reported to produce hydroquinine 1,2-dioxygenase, which

converts hydroquinone to cis,trans-4-hydroxymuconic

semialdehyde. This is then metabolized to maleylacetate by

semialdehyde dehydrogenase. Maleylacetate is converted to

3-oxoadipate by a reductase, which is finally metabolized to

intermediary metabolites of the tricarboxylic acid (TCA)

cycle (Carnett, 2002). A Pseudomonas putida strain was

found to metabolize p-nitrophenol to hydroquinone and

1,2,4-benzenetriol, which was further cleaved by benzene-

triol oxygenase to maleylacetate (Rani & Lalitha-kumari,

1994). A similar pathway of p-nitrophenol degradation was

reported in Pseudomonas cepacia that can utilize p-nitro-

phenol as a source of carbon and nitrogen (Prakash et al.,

1996).

A different pathway of degradation was reported in

Arthrobacter sp. strain JS443 and Arthrobacter protophormiae

RHJ100 where p-nitrophenol was mineralized via p-nitroca-

techol. Nitrocatechol is converted to 1,2,4-benzenetriol by

benzotriol dehydrogenase, which in turn is directly con-

verted to maleylacetate by benzotriol dioxygenase (Jain et al.,

1994; Bhushan et al., 2000a; Chauhan et al., 2000). Recently,

a consortium of two Pseudomonas ssp. (strains S1 and S2)

was isolated which can also metabolize p-nitrophenol via

p-nitrocatechol (Qureshi & Purohit, 2002). The analogous

compound 3-methyl-4-nitrophenol has also been reported

to be metabolized by Ralstonia sp. via catechol formation

(Bhushan et al., 2000b). A Nocardia sp. was reported to

produce p-nitrophenol-2-hydroxylase, which catalyzes trans-

formation of p-nitrophenol to p-nitrocatechol (Mitra &

Vaidyanathan, 1984). A mono-oxygenase from a Moraxella

sp. that releases nitrite from p-nitrophenol has been partially

purified (Spain & Gibson, 1991). A soluble nitrophenol

oxygenase was purified from P. putida B2 that converts

ortho-nitrophenol to catechol and nitrite (Zeyer & Kocher,

1988). A novel monooxygenase was characterized from

Bacillus sphaericus that catalyzes the first two steps of the

degradation of p-nitrophenol via p-nitrocatechol and benzo-

triol. This enzyme consists of two components, a reductase

and oxygenase, and catalyzes two sequential mono-oxygena-

tion reactions that convert p-nitrophenol to benzotriol. The

first reaction converts p-nitrophenol to p-nitrocatechol and

the second removes the nitro group (Kadiyala & Spain,

1998). A pentachlorophenol degrading Sphingomonas

sp. UG30 was found to degrade p-nitrophenol. A pentachloro-

phenol-monooxygenase was purified from this bacterium

that can catalyze the hydroxylation of p-nitrocatechol to

benzotriol (Leung et al., 1999). A hydroxyquinol (benzo-

triol) ring cleavage dioxygenase was isolated and character-

ized from p-nitrophenol degrading Arthrobacter sp. strain

JS443. The gene encoding this dioxygenase (npdB) was

found to be in the same gene cluster as reductase (npdA1)

and oxygenase (npdA2) components of the p-nitrophenol

mono-oxygenase, maleylacetate reductase (npdC), and a

regulatory protein (npdR) (Zylstra et al., 2000; Parales

et al., 2002). Rhodococcus strain PN1 and Rhodococcus

erythropolis HL PM-1, which degrade 2,4-dinitrophenol

and p-nitrophenol, were reported to contain an npd gene

cluster including npdC (encoding hydride transferase I),

npdG (encoding the NADPH-dependent F420 reductase)

and npdI (encoding hydride transferase II). It was observed

that npdG and npdI genes have the same function as the

homologous genes (Heiss et al., 2003). Recently, a novel gene

called orf243 was reported from Flavobacterium sp. orf243

which is transposon based and is linked with the opd gene

(Siddavattam et al., 2003). This gene encodes a protein with

homology to a family of aromatic compound hydrolases and

is able to degrade p-nitrophenol.



Although in most of the studies on microbial degradation

of parathion, the first reaction was hydrolysis of the phos-

photriester bond, there have been reports of different

degradation pathways. In one study, degradation of para-

thion by a mixed culture and a Bacillus sp. (Sharmila et al.,

1989) was shown to occur by reduction of the nitrogroup

that was later hydrolyzed to p-aminophenol. Another report

of conversion of parathion to paraoxon before hydrolysis of

phosphotriester bond was reported in a mixed bacterial

culture (Tomlin, 2000).

Studies on the degradation of methyl parathion (O,O-

dimethyl-O-p-nitrophenyl phosphorothioate) have also

been reported. Methyl and ethyl parathion have identical

chemical structures except for the ethyl groups of the P

chain of parathion, which are replaced by methyl groups as

evident by the name of the compound. A Pseudomonas sp.

was isolated that can co-metabolically degrade methyl para-

thion (Chaudry et al., 1988). Rani & Lalitha-kumari (1994)

isolated P. putida that could hydrolyze methyl parathion and

utilize p-nitrophenol as a source of energy. A Bacillus sp. was

reported to degrade methyl parathion by both hydrolysis

and nitro group reduction (Sharmila et al., 1989). Utiliza-

tion of methyl parathion by Flavobacterium balustinum as

the sole source of carbon was observed earlier (Somara &

Siddavattam, 1995). In this bacterium the opd gene was

found to be linked with a novel gene involved in degradation

of p-nitrophenol (Siddavattam et al., 2003). Degradation of

methyl parathion by a Pseudomonas sp. in soil and on

sodium alginate beads was reported (Ramanathan & La-

lithakumari, 1996). Co-metabolic degradation of methyl

parathion by Plesimonas sp. strain M6 was observed (Zhon-

gli et al., 2001) which was mediated by a novel degrading

gene. They also isolated Pseudomonas sp. A3 which can

utilize p-nitrophenol as sole source of carbon and nitrogen.

This isolate can also utilize a series of aromatic compounds

as a sole source of carbon (Zhongli et al., 2002). Another

strain of Pseudomonas sp. WBC was isolated from polluted

soils around a Chinese pesticide factory. The isolate was

capable of complete degradation of methyl parathion and

could utilize it as sole source of carbon and nitrogen (Yali

et al., 2002). The hydrolysis product of methyl parathion is

also p-nitrophenol, for which the degradation pathways

have already been described. The different proposed path-

ways of parathion and methyl degradation are presented

in Fig. 3.

Glyphosate

Glyphosate (N-(phosphonomethyl) glycine) is a globally

used broad-spectrum herbicide. It is a representative of the

phosphonic acid group of compounds, which is character-

ized by a direct carbon to phosphorus (C–P) bond. The C–P

linkage is chemically and thermally very stable and renders

the molecule much more resistant to non-biological degra-

dation in the environment than its analogues with O-P

linkage (Hayes et al., 2000). Mode of action of glyphosate

includes inhibition of the plant enzyme 5-enol-pyruvyl-

shikimate-3-phosphate synthase, which catalyzes synthesis

of aromatic amino acids (Fisher et al., 1984; Cole, 1985).

Glyphosate is moderately persistent with a half-life of

30–170 days (Tomlin, 2000). Microbial degradation is

considered to be the most important of the transformation

processes controlling its persistence in soil (Araujo et al.,

2003). It was observed that mineralization of glyphosate is

related to both the activity and biomass of soil micro-

organisms (Wiren-Lehr et al., 1997). Microbial degradation

of glyphosate produces the major metabolite aminomethyl

phosphonic acid and ultimately leads to the production of

CO2, phosphate and water (Forlani et al., 1999; Araujo

et al., 2003). Several species of bacteria have been isolated

from previously treated and untreated environments, which

can degrade glyphosate either co-metabolically or as a

source of phosphorus. There has been no report of the

utilization of glyphosate as a source of carbon or nitrogen

(Dick & Quinn, 1995). Several species of Pseudomonas have

been isolated which can degrade glyphosate (Moore et al.,

1983; Tolbot et al., 1984; Jacob et al., 1988; Quinn et al.,

1989). Similarly, a Flavobacterium sp. (Balthazor & Hallas,

1986), an Alcaligenes sp. (Tolbot et al., 1984), Bacillus

megaterium strain 2BLW (Quinn et al., 1989), several species

of Rhizobium (Liu et al., 1991), three species of Agrobacter-

ium (Wacket et al., 1987; Liu et al., 1991) and an Arthro-

bacter sp. (Pipke et al., 1987) have also been reported to

degrade this herbicide (Table 2).

Three different pathways for C–P bond cleavage have

been reported for the use of phosphonate as a source of

phosphorus for growth.

The phosphonatase pathway is involved in degra-

dation of alpha carbon substituted phosphonates, which are

primarily naturally occurring phosphonates such as

2-aminoethylphosphonates that have been reported in

Bacillus cereus (Lee et al., 1992b), and Pseudomonas aeru-

ginosa (Lacoste et al., 1993), Salmonella typhimurium and

several other organisms (Jiang et al., 1995). In a two-step

process, this pathway leads to the cleavage of the C–P bond

by a hydrolysis reaction requiring an adjacent carbonyl

group. 2-Aminoethylphosphonate is converted to phosp-

honoacetaldehyde by a specific transaminase, which is

further degraded to acetaldehyde by phosphonatase.

The C–P lyase pathway is involved in the cleavage of

both substituted and unsubstituted phosphonates such as

methylphosphonates (Lee et al., 1992b).The phospho-

noacetate hydrolase pathway specifically degrades phospho-

noacetate and appears to have evolved for phosphonate use

as a carbon source. This enzyme catalyzes the hydrolysis of

phosphonoacetate ;to acetate and inorganic phosphonates



via metal cation-assisted P–C bond cleavage (McMullan &

Quinn, 1994; McGrath et al., 1995). Glyphosate has been

found to be degraded by the second of these pathways.

Two different pathways of glyphosate degradation are

presented in Fig. 4. Arthrobacter sp. GLP-1 and Pseudomonas

sp. PG2982 degrade glyphosate by initial cleavage of the C–P

bond, resulting in the production of sarcosine (N-methyl-

glycine) by C–P lyase activity (Moore et al., 1983; Shinabar-

ger & Braymer, 1984; Pipke et al., 1987; Liu et al., 1991; Dick

& Quinn, 1995). Rhizobium meliloti has also been reported

to degrade glyphosate by this pathway but, unlike other

bacteria, it has only one C–P lyase, which is able to degrade a

wide range of phosphonates (Park & Hausinger, 1995). The

sarcosine formed is further degraded to the amino acid

glycine and a C1-unit, which is incorporated into purines,

and the amino acids serine, cysteine, methionine and

histidine (Pipke et al., 1987). The second pathway involves

the conversion of glyphosate to aminomethylphosphonic

acid (AMPA) by the loss of a C2 unit. This compound is then

dephosphorylated by C–P lyase and further broken down by

subsequent steps to methylamine and formaldehyde (Pike &

Amrhein, 1988; Lerbs et al., 1990). An identical pathway has

Fig. 3. Different pathways of parathion and

methyl parathion degradation by microorgan-

isms. When the conversion of one compound to

another is believed to occur through a series of

intermediates, the steps are indicated by dotted

arrows. DATP, dialkylthiophosphate.



been observed in Arthrobacter atrocyaneus (Pike & Amrhein,

1988) and Flavobacterium sp. (Balthazor & Hallas, 1986;

Pipke et al., 1987). Recently, a thermophile, Geobacillus

caldoxylosilyticus T20 was isolated from a central heating

system which also degrades glyphosate by this pathway,

utilizing the compound as a sole source of phosphorus

(Obojska et al., 2002). A halophilic bacterium, Chromohalo-

bacter marismortui, isolated from soil beneath a road

gritting salt pile was capable of utilizing several organopho-

sphonates including aminomethyl phosphonic acid as a

source of phosphorus (Hayes et al., 2000). Utilization of

aminoalkylphosphonates as a source of nitrogen by different

bacterial isolates has been reported (McMullan & Quinn,

1994; Ternana & McMullan, 2000). Pseudomonas fluorescens

was reported to utilize a diverse range of organophospho-

nates as sources of carbon, nitrogen and phosphorus

(Zboinska et al., 1992a). A strain of Kluyveromyces fragilis

has been shown to utilize AMPA as a source of nitrogen

(Ternana & McMullan, 2000). Strains of Streptomyces were

also reported to degrade and utilize several organopho-

sphonate compounds as sources of carbon and nitrogen.

These strains were capable of degrading glyphosate in

phosphate-free media via C–P bond cleavage accompanied

by sarcosine formation (Obojska et al., 1999). Streptomyces

morookaensis DSM 40565 could degrade aminoalkylpho-

sphonate as a sole source of nitrogen and phosphorus

(Obojska & Lejczak, 2003). Alkyl amines are intermediate

degradation products for several xenobiotics such as carbo-

furan, atrazine, and monocrotophos and have been reported

to serve as a source of energy for different micro-organisms

(Strong et al., 2002). Use of methylamine as a source of

carbon is widespread in nature (Hanson & Hanson, 1996;

Trabue et al., 2001).

Fungi play an important role in degradation of xenobio-

tics and biospheres (Pothuluri et al., 1998, 1992) including

glyphosate. Probably the first fungal degradation of glypho-

sate by Penicillium citrinum was reported by Zboinska et al.

(1992b). Penicillium notatum can utilize the herbicide as a

source of phosphorus and can degrade it by the amino-

methyl phosphonic acid pathway (Bujacz et al., 1995).

Strains of Trichoderma harzianum, Scopulariopsis spand and

Aspergillus niger were able to degrade glyphosate and

aminomethyl phosphonic acid in the laboratory (Krzysko-

Lupicka et al., 1997). The first report of utilization of

glyphosate as a source of nitrogen by a microorganism was

reported for Penicillium chrysogenum (Klimek et al., 2001).

The fungal cells were found to lack detectable nitrogen

reductase activity and therefore this isolate seemed to lack

Fig. 4. Pathways of microbial degradation for

glyphosate. AMPA, aminomethyl phosphonic

acid.



the ability to convert nitrate to ammonium. Recently,

Alternaria alternata, a plant pathogen, was found to utilize

glyphosate as a source of nitrogen (Lipok et al., 2003).

The above observations suggest that glyphosate is de-

graded by several soil microorganisms, and different steps of

the degradation involve different microorganisms which

utilize different degradation products as different sources of

energy. The possible pathways of glyphosate degradation are

presented in Fig. 4.

Coumaphos

Coumaphos (O,O-diethyl-O-(3-chloro-4-methyl-2-oxo-

2H-1-benzo-pyran-7-yl) phosphorothioate) is used as an

acaricide for the control of cattle ticks. It is widely used by

different government agencies for tick eradication and

quarantine purposes. The primary tool used in the eradica-

tion programme is a series of dipping vats placed at border

crossing points. The cattle are induced to jump into the deep

end of the vat, resulting in their complete immersion in

coumaphos. They then swim the length of the vat and climb

out to other end. There are around 42 vats in the USA alone

and each vat contains about 15 000 L of coumaphos suspen-

sion at the rate of 1600 mg L�1 (42% active ingredient, a.i.)

(Shelton & Somich, 1988; Mulbry et al., 1998). The vats are

cleaned and recharged every 2 years to keep the concentra-

tion of acaricide at a desirable level. These operations

generate approximately 460 000 L of concentrated insecti-

cide waste yearly in USA alone (Mulbry et al., 1996). A

similar programme within Mexico is thought to produce a

much larger volume. Coumaphos is comparatively persis-

tent in soil, with a half-life of about 300 days (Kearney et al.,

1986) and it possesses a very high mammalian toxicity.

Because of these characteristics, it requires a safe and

effective method for disposal. Rapid degradation of couma-

phos was observed in several cattle-dipping vats, resulting in

loss of efficacy against cattle ticks (Shelton & Karns, 1988).

Under aerobic conditions, experiments with radiolabelled

coumaphos demonstrated that the aromatic portion of the

molecule is susceptible to mineralization by bacteria in

problematic vat dips (loss of efficacy). Three morphologi-

cally distinct bacteria (designated B-1, B-2 and B-3) that

could metabolize coumaphos were isolated from a problem

vat dip (Shelton & Somich, 1988). All these bacteria hydro-

lyzed coumaphos to DETP and chlorferon. Chlorferon was

further metabolized by B-1 and B-2 to a-chloro-b-methyl-

2,3,4-trihydroxy-trans-cinnamic acid (CMTC). Further ex-

periments demonstrated that B-1 was capable of mineraliz-

ing and incorporating the aromatic portion of the

coumaphos molecule into biomass, but this was inhibited

by the accumulation of metabolites that was due apparently

to the inefficient metabolism of a chlorinated intermediate.

Combination of B-1 with another organism from the vat,

designated strain B-4, which metabolized these inhibitory

products, yielded a stable two-member consortium able to

grow at the expense of coumaphos (Shelton & Haperman-

Somich, 1991). No further study on the degradation path-

way or metabolite identification has been carried out.

Ralstonia sp. LD35 has been reported to degrade an

analogous compound, 3,4-dihydroxycinnamic acid via ben-

zoic acid (Gioia et al., 2001). A similar breakdown pathway

for the propenoic side chain of substituted cinnamic acid

molecule, p-coumaric acid, has been observed in Pseudomo-

nas sp. (Tse et al., 2004) and Acinetobacter strains (Delneri

et al., 1995). These bacteria use p-coumaric acid as the

source of carbon. In the first step, they convert p-coumaric

acid into p-hydroxybenzoic acid which is then transformed

to protocatechuic acid and integrated to the TCA cycle via

the b-ketodipate pathway. Many bacteria degrade substi-

tuted cinnamic acid by decarboxylation of side chains.

Enzymes and genes responsible for such degradation have

been purified and characterized (Degrassi et al., 1995;

Barthelmebs et al., 2000). Streptomyces setonii (Sutherland

et al., 1983) and Rhodopseudomonas palustris (Harwood &

Gibson, 1988) have been shown to degrade cinnamic and 4-

coumaric acids to their corresponding benzoic acid deriva-

tives. Several other bacteria follow the same pathway for

degradation of substituted cinnamic acids. Monooxygenase

and dioxygenase catalyze the formation of the 2-, 3-, and 4-

hydroxy derivatives as substituted acid and/or substituted

catechol (Peng et al., 2003).

The b-oxidation pathway has been proposed for the

degradation of substituted cinnamic acids by Pseudomonas

putida (Zenk et al., 1980). This pathway, which is analogous

to the b-oxidation of fatty acids, is thought to include

thiolytic cleavage of 4-hydroxy-3-methoxy-b-ketopropinyl-

CoA to yield acetyl CoA and vanillyl CoA, which is catalyzed

by b-ketoacyl CoA thiolase. The pathway subsequently leads

to ring fission and requires several co-factors including ATP,

CoA and NAD1 (Zenk et al., 1980). Under anaerobic

conditions, coumaphos undergoes reductive dechlorination

to form potasan (Mulbry et al., 1998).

Nocardia sp. strain B-1 was reported to degrade couma-

phos by a different gene enzyme system to the known opd

gene (Mulbry, 1992). Another microorganism, Nocardiodes

simplex NRRL B-24074, was found to have a distinct

enzymes system for coumaphos degradation (Mulbry,

2000). Horne et al. (2002b) isolated an Agrobacterium

radiobacter P230 capable of hydrolyzing coumaphos from

an enrichment culture containing organophosphorus as the

sole source of phosphorus. This bacterium degrades couma-

phos by hydrolysis of the phosphotriester bond. Pseudomo-

nas monteilli was isolated which can hydrolyze coumaphos

as well as its oxo analogue coroxon but it can utilize only

coroxon as a sole source of phosphorus, not coumaphos or

its hydrolysis product DETP. This bacterium degrades



coumaphos and diazinon but not parathion (Horne et al.,

2002a). Coumaphos is degraded by the other microorgan-

isms like Flavobacterium sp. (Sethunathan & Yoshida, 1973),

P. diminuta (Serdar et al., 1982), and Enterobacter sp. B-14

(Singh et al., 2004), which were isolated for their ability to

degrade other organophosphorus compounds. This obser-

vation suggests that these microorganisms produce several

isoenzymes or broad-specificity enzymes that can degrade a

range of organophosphorus compounds. The proposed

pathway of microbial degradation of coumaphos is shown

in Fig. 5.

Fenamiphos

Fenamiphos (ethyl 4-methylthio-m-tolyl isopropylpho-

sphoramidate) is an organophosphorate used extensively

for the control of soil nematodes. It is systemic, active

against ecto- and endo-parasitic, cyst forming and root-

knot nematodes, and is recommended for application at

5–20 kg a.i.ha-1. Its solubility at room temperature is

700 mg L�1 water. The acute oral LD50 is 15.3–19.4

mg kg�1 for rats, 10 mg kg�1 for dogs and 75–100 mg kg�1

for guinea pigs (Tomlin, 2000).

Fig. 5. Proposed pathways for microbial

degradation of coumaphos. The scheme is based

on articles cited in the text. When the conver

sion of one compound to another is believed

to occur through a series of intermediates, the

steps are indicated by dotted arrows. DETP,

diethylthiophosphate; CMTC, chloromethyl

trihydroxy cinnamic acid.



Although, there have been reports of enhanced degrada-

tion of fenamiphos, the mechanism of degradation has

received little attention. Fenamiphos is oxidized rapidly to

fenamiphos sulfoxide (FSO) which in turn is oxidized to

fenamiphos sulfone (FSO2). As FSO and FSO2, have nema-

ticidal activity and toxicity similar to fenamiphos (Waggoner

& Khasawinah, 1974), degradation and persistence studies

usually include estimation of total toxic residue, which is the

combination of the two oxidation products along with

parent compound. The half-life in soil for fenamiphos and

its metabolites (total toxic residues) varies from 30 days to 90

days (Johnson, 1998). More rapid rates of degradation in soil

repeatedly treated with the fenamiphos in the laboratory

have been reported (Chung & Ou, 1996) and enhanced

degradation of fenamiphos in the field has been observed in

many countries (Stiriling et al., 1992; Smelt et al., 1996;

Meghraj et al., 1999). It was suggested that 3–4 years were

necessary before the accelerated degradation of fenamiphos

declined in a sandy soil in a temperate region (Ou, 1991).

Fenamiphos rapidly disappears from both enhanced and

non-enhanced soils but FSO2 is rarely formed in enhanced

soils (Ou, 1991). This suggests that enhanced bio-degrada-

tion of fenamiphos total toxic residue was due to an increase

in the disappearance rate of FSO in soil samples collected

from field sites treated one or two consecutive times with

fenamiphos (Davis et al., 1993). In a recent study of soil

samples from a field in the UK, which had similar physical

characteristics except for soil pH, the degradation rate of

fenamiphos increased with the increase in pH. Repeated

application of fenamiphos slowed down the rate of degrada-

tion in acidic soils, and in the neutral pH soil, three

consecutive treatments did not result in the development of

enhanced degradation of fenamiphos. However, in the two

alkaline soils, a second treatment with fenamiphos led to

enhanced degradation (Singh et al., 2003b). Chung & Ou

(1996) have tried to shed light on the mechanism of

fenamiphos degradation in soils that showed enhanced

degradation. They reported that fenamiphos is degraded

into FSO which in turn is rapidly degraded into FSO-

phenol, which is subsequently mineralized into CO2. There-

fore in enhanced soil, degradation of fenamiphos (total toxic

residue) is rapid because it misses one step, FSO to FSO2. In

enhanced UK soils, fenamiphos was rapidly oxidized to FSO,

which in turn, was quickly degraded. The major fenamiphos

metabolites identified were FSO and FSO-phenol. No FSO2

was detected in the enhanced soil samples (Singh et al.,

2003b). However, in two Australian soils, a different me-

chanism of fenamiphos degradation was observed where the

nematicide was directly converted to fenamiphos phenol,

suggesting that the first oxidation step was replaced by

hydrolysis (Singh et al., 2003b).

Ou & Thomas (1994) isolated the first microbial con-

sortium with six different bacterial species that degraded

fenamiphos in liquid culture. A pure culture of Brevibacter-

ium sp. MM1 was isolated which hydrolyzed fenamiphos

and its hydrolysis products but did not utilize these chemi-

cals as energy sources (Megharaj et al., 2003). Two different

consortia from Australian soils, made up of five and four

different bacterial strains, were isolated [B. K. Singh, un-

published]. Both consortia could utilize fenamiphos as sole

sources of carbon and nitrogen. In contrast to the con-

sortium isolated by Ou & Thomas (1994), the two Austra-

lian consortia (CRF and BEP) did not require any

supplementary nutrient source for fenamiphos degradation

and were active in liquid media in the absence of mineral

surfaces (Singh et al., 2003b). These microbial systems were

found to mineralize fenamiphos or its oxidative metabolites

by hydrolysis as a first step. The hydrolytic product fenami-

phos phenol, FSO-phenol or fenamiphos sulfone phenol

(FSO2-OH) can be further degraded by desulfonation. Three

modes of desulfonation are reported for aromatic sulfo-

nates: desulfonation (a) before, (b) during or (c) after ring

cleavage (Kertesz et al., 1994a). Mode (a) is considered to be

most common pathway of desulfonation in the environ-

ment. In this pathway, the target compound is oxygenated

by a multi-component oxygenase, yielding an unstable

sulfono cis-diol, which then spontaneously re-aromatizes

to the corresponding catechol with the loss of sulfite. An

enzyme which catalyzes this reaction in toluene sulfonate

and benzene sulfonate has been isolated from an Alcaligenes

sp. (Thurnheer et al., 1986, 1990). In Pseudomonas putida

S-313, a broad-spectrum monooxygenolytic sulfonatase

catalyzes the conversion of sulfonate to a phenol with

incorporation of one oxygen atom from molecular oxygen

(Kertesz et al., 1994b). Alcaligenes sp. strain O-1 is reported

to contain two different desulfonative pathways where the

initial desulfonation is catalyzed by different dioxygenase

enzyme systems. One enzyme system can degrade 2-amino-

benzenesulfonate, benzene sulfonate and 4-toluene sulfo-

nate but the other one can degrade only the last two

compounds (Junker et al., 1994). Hydrogenophaga palleronii

S1 has been reported to degrade 4-carbo-4-sulfoazobenzene

by the 4-sulfocatechol pathway via the formation of 4-

aminobenzenesulfate (Vickers, 2002). Another proposed

pathway is transformation of toluene sulfonate to hydroxy

toluene by toluenesulfonate monooxygenase. Pseudomonas

putida strain S-313 catalyzes toluene sulfonate desulfona-

tion, which can serve as its sole source of sulfur and leaves 4-

hydroxytoluene unmetabolized. However 4-hydroxy toluene

is a metabolite that is readily catabolized by other bacteria

via the toluene pathway (Eisenmaan & McLeish, 2002).

Another toluene sulfonate degrading bacterium, Coma-

monas testosteroni T-2, was found to contain a degrading

gene on a plasmid (Hooper et al., 1990). Simple alkane

sulfonates are utilized by Pseudomonas sp. as a carbon source

where crude cell extract catalyzes the oxidation of the



a-carbon atom of alkanesulfonate to an aldehyde bisulfite

adduct. This adduct then degrades to produce the corre-

sponding aldehyde and sulfite. The substrate range for this

reaction has been reported to be relatively broad where

hydroxy-, methyl-, and alkenyl-substituted compounds are

all transformed (Thysse & Wanders, 1974). Degradation of

alkylsulfate proceeds via initial hydrolysis of the sulfate ester

linkage and subsequent oxidation of the released alkanol

(Kertesz et al., 1994a). Pseudomonas sp. C12B and a strain of

Comamonas terrigena were reported to utilize a range of

alkylsulfates as a source of carbon (Payne & Faisal, 1963;

Fitzgerald et al., 1977). Five different alkylsulfatases were

characterized from Pseudomonas sp. C12B and two from C.

terrigena (Dodgson et al., 1982). On the basis of the above

studies, we propose the microbial degradation pathways for

fenamiphos as presented in Fig. 6.

Other organophosphorus pesticides

Several other organophosphorus compounds have been

used extensively for pest control. Diazinon, monocrotophos,

malathion, dimethoate, etc., are being used world-wide.

Several species of bacteria have been isolated and character-

ized that can degrade these compounds in liquid medium

and soils (Table 2).

Fig. 6. Proposed pathways for fenamiphos

degradation by microorganisms. The scheme

is based on articles cited in the text. FSO,

fenamiphos sulfoxide; FSO2, fenamiphos

sulfone; FSO-phenol, fenamiphos sulfoxide

phenol; FSO2-OH, fenamiphos sulfone phenol;

F phenol, fenamiphos phenol.



Monocrotophos ((3-hydroxy-N-methyl-cis-crotonamide)

dimethyl phosphate) is widely used to control aphids, leaf

hoppers, mites and other foliage pests. It has been classified

as extremely hazardous, with an LD50 value of 20 mg kg�1

for mammals. The half-life of monocrotophos in soil was

reported to be 40–60 days (Tomlin, 2000). Monocrotophos

is easily soluble in water and therefore has potential to

contaminate ground water. Together with its high mamma-

lian toxicity, these characteristics make monocrotophos an

ideal compound for decontamination and detoxification.

Rangaswamy & Venkateswaralu (1992) isolated a monocro-

tophos degrading Bacillus sp. from previously treated soil.

Megharaj et al. (1987) isolated monocrotophos degrading

algae from soil. Two different algae, Aulosira fertilissima

ARM 68 and Nostoc muscorum ARM 221, were found to

utilize monocrotophos as a sole source of phosphorus

(Subramanian et al., 1994). Pseudomonas aeruginosa F10B

and Clavibacter michiganense ssp. insidiosum SBL 11 were

isolated from soil. These bacteria can utilize monocrotophos

as a phosphorus source but not as a carbon source (Singh &

Singh, 2003). Two species of Pseudomonas, three species of

Bacillus and three species of Arthrobacter were isolated from

soils, which can utilize monocrotophos as a sole source of

carbon (Table 2). Further studies demonstrated that Pseu-

domonas mendocina is the most efficient monocrotophos

degrader among the isolated bacteria and its degrading

capability is plasmid based (Bhadbhade et al., 2002a). The

same group isolated another 17 bacterial isolates from

previously exposed soils which can mineralize monocroto-

phos in liquid culture (Bhadbhade et al., 2002b). The two

most versatile degraders, Bacillus megaterium and A. atro-

cyaneus, were chosen for further studies on the biochemical

mechanisms and pathways of monocrotophos degrada-

tion. Phosphatase activities were observed in both cul-

tures, and it was suggested that the phosphates identified

may be mono- and dimethyl phosphates (Bhadbhade et al.,

2002b). Dimethyl- and monomethyl phosphates were

involved as intermediates in monocrotophos degradation

in plants and animals (Menzer & Cassida, 1965; Muck,

1994). Another intermediate identified during monocroto-

phos degradation was methylamine, produced by an esterase

enzyme. This esterase could be an amidase capable of

selecting amides as substrates since esterases sometimes

attack the amide bond (Hassal, 1990). Similar pathways of

degradation were reported for dicrotophos, which is first

demethylated to monocrotophos and then further degraded

to methyl amine (Eto, 1974). As with most of the other

organophosphorus compounds, the first degradation step

of monocrotophos should involve hydrolysis, which

could produce N-methyl acetoacetamide and dimethyl

phosphate (Beynon et al., 1973). Further degradation

of N-methyl acetoacetamide produced valeric acid in A.

atrocyaneus and acetic acid in B. megaterium (Bhadbhade

et al., 2002b). Acetic acid is the key intermediate of the

glycolytic pathway in microorganisms. The pathway of

dicrotophos- and monocrotophos degradation is shown in

Fig. 7.

Degradation of fenitrothion (O,O-dimethyl O-4-nitro-m-

tolyl phosphorothioate), a widely used insecticide, by Bur-

kholderia sp. strain NF100 was reported (Hayatsu et al.,

2000). This strain utilized fenitrothion as a source of carbon

with the help of two plasmids. The first plasmid (pNF2) was

found to catalyze the hydrolysis of fenitrothion to 3-methyl-

4-nitrophenol. The nitro group from this compound

was oxidatively removed to form methylhydroquinone,

which was further metabolized by the second plasmid

(pNF2) (Hayatsu et al., 2000). This bacterium was also

found to degrade p-nitrophenol as a source of energy.

Methylhydroquinone may be degraded by ring fission as

one of the two methods described for p-nitrophenol

degradation in the section dealings with parathion. p-

Nitrophenol degrading Ralstonia sp. SJ98 was reported to

have chemotaxis towards 3-methyl-4-nitrophenol and to

utilize it as a source of carbon. This strain degrades 3-

methyl-4-nitrophenol by the formation of catechol (Bhush-

an et al., 2000b).

Microbial degradation of various other organopho-

sphorus compounds has been documented. Diazinon de-

gradation by a Flavobacterium sp. was reported in 1973

(Sethunathan & Yoshida, 1973). Two Pseudomonas spp.

isolated from sewage sludge were found to degrade diazinon

in a culture medium (Rosenberg & Alexander, 1979). Two

strains of Arthrobacter sp. were reported to hydrolyze

diazinon (Barik et al., 1979). Dimethoate degradation was

reported to be carried out by a plasmid based gene of

P. aeruginosa MCMB-427 (Deshpande et al., 2001). A novel

dimethoate degrading enzyme was purified and character-

ized from a strain of the fungus Aspergillus niger. This

enzyme was found to degrade all compounds containing

P–S linkage like malathion and fermothion but not com-

pounds with the P–O linkage (Liu et al., 2001).

Utilization of ethoprophos as a sole source of carbon by P.

putida has been observed (Karpouzas et al., 2000). Isolation

and metabolism of cadusafos by Sphingomonas paucimobilis

and Flavobacterium sp. have been reported recently (Kar-

pouzas et al., 2005). Similarly, several species of bacteria

were isolated from different environments which degrade

organophosphorus compounds in laboratory cultures and

in soils (Singh et al., 1999). Microorganisms isolated from

enrichment of one organophosphorus compound can de-

grade other structurally similar compounds. For example,

Flavobacterium sp. and P. diminuta were isolated by diazinon

and parathion enrichment but they can degrade a wide

range of other organophosphorus compounds such couma-

phos, methyl parathion, chlorpyrifos and nerve agents

(Adhya et al., 1981; Singh et al., 1999).



Chemical warfare agents

Among lethal chemical warfare agents, the nerve agents have

played a dominant role since the Second World War. Nerve

agents acquired their name because they affect the function-

ing of nerve impulses like other organophosphorus com-

pounds. The nerve agents are a group of particularly toxic

warfare agents. There are five major substances that are

classified as nerve agents and they can be divided into two

main groups:

(1) G agents, including Tabun (GA), Sarin (GB), Soman

(GD) and cyclohexyl methylphosphonofluoridate

(commonly referred to as cyclosarin or GF).

(2) V agent, represented by VX.

G agents are usually non-persistent volatile liquids

whereas VX is highly persistent, non-volatile and much

more active than any of the G agents. Physical and chemical

properties of these nerve agents are listed in Table 3. Munro

et al. (1994) described the acute and chronic toxicity of

nerve agents. In another review by this group, they listed the

different sources, fate and toxicity of degradation products

of chemical warfare agents (Munro et al., 1999).

It is estimated that about 30 000 tons in USA and about

200 000 tons nerve agents globally have to be destroyed

under the Chemical Weapons Convention (CWC), 1993. As

of 30 January 2002, 175 states have made CWC commit-

ments. CWC bans the use of chemical weapons but more

significantly also bans their development, production,

stockpiling and transfer and requires that all existing stocks

be destroyed by the member states within 10 years of

ratification. Other known chemical warfare agents concen-

trations include Japanese chemical weapons munitions

abandoned in China in 1945, and an estimated 100 000 tons

of German chemical weapons munitions that were dumped

O

O

O

C

H3C

CH3

CH3

C

Dicrotophos

H

C NP

O

O

O

C

H3C

CH3

CH3

CH3

C

MCP

H H

C NP

H3CO

H3CO

O

C C C N

HH

OOH

HOP

Dimethyl phosphate N-Methyl acetoacetamide

Methylamine

H3CO

H3CO

O

OH

Phosphoric acidPhosphate HCHO

Valeric acid Acetic acid

CH3(Ch2)3COOH CH3 COOH

PHO

HO

CO3

O

O C CH

COOH

NH4+

PH3CO

CH3NH2H3CO

H3C

H3CO

H3CO

Formaldehyde

C1 metabolic cycleFig. 7. Pathways for microbial degradation

of dicrotophos and monocrotophos (MCP) .



into the Baltic Sea at the end of World War II. Prior to 1969,

the US army disposed of chemical weapons by open pit

burning, evaporative atmospheric dilution, burial and pla-

cement of munitions in concrete coffins for ocean dumping.

In the 1970s, alkaline hydrolysis replaced the above methods

for destroying nerve agents. Later, due to the resistance of

GB to alkaline hydrolysis, the incineration method was

adopted for destroying all groups of chemical weapons.

However, due to strong opposition to incineration by

environmentalist and local populations, this method of

chemical warfare agents destruction was stalled in the USA

and republics of the former Soviet Union. Consequently,

there is a need to find alternative remediation methods that

can provide an environmentally safe and economically

viable solution. In this section, we review the environmental

fate of important nerve agents and the pathways of micro-

bial degradation.

Tabun (GA)

It is believed that Tabun (name given by its inventor), or GA

(ethyl N,N-dimethylphosphoroamidocyanidate), was the

first nerve agent ever discovered. It was manufactured in

1937 in Germany, although large scale production and

stockpiling started in 1942 (Robinson, 1967). GA is the

American denomination of Tabun. It enters the body mainly

through the respiratory tract and the primary action is on

the respiratory system. It can also cause vision impairment

through its anti-acetylcholine esterase activity. GA has a

high water solubility but is also readily soluble in organic

solvents and can therefore easily penetrate skin (Munro

et al., 1999).

GA contains a cyanide group and is subject to hydrolysis.

Under neutral and acidic conditions, the first step, which is

very rapid, includes formation of O-ethyl N,N-dimethyl

amidophosphoric acid and hydrogen cyanide. The subse-

quent hydrolytic step, which is comparatively slow, is

hydrolysis of O-ethyl N,N-dimethyl amidophosphoric acid

to dimethylphosphoramidate and then finally to phosphoric

acid. Under acidic conditions, hydrolysis to ethylphosphor-

ylcyanide and dimethylamine occurs. The final product of

all pathways is phosphoric acid. Several different metabolites

were identified in soil exposed to GA. D’Agostino & Provost

(1992) identified 16 different compounds from a soil con-

taminated with GA. However, several of them were impu-

rities and some were degradation products.

The chemical structure of GA suggests that it contains

several possible microbial degradation sites. The initial steps

are potentially O-dealkylation and C-dealkylation, nitrile

hydrolysis and N-dealkylation (Morrill et al., 1985). No

specific microorganism has been isolated for exclusive GA

degradation from natural environments but P. diminuta

isolated for degradation of other organophosphorus com-

pounds can degrade several chemical warfare agents includ-

ing GA (Mulbry & Rainina, 1998). DeFrank et al. (1993)

isolated several strains of Alteromonas that can effectively

degrade all G nerve agents. As with other organophosphorus

compounds, the complete degradation of GA is likely to

produce phosphoric acid. Several bacterial species have been

reported to cleave C–P bonds. The mechanism and asso-

ciated micro-organisms have been described in detail under

the section dealing with glyphosate.

Several intermediate metabolites of GA have been identi-

fied from soils that include dimethylamine and triethyl

phosphate (Sanches et al., 1993; Verschueren, 1996), and

diethyl dimethylphosphoramidate (Munro et al., 1999).

These compounds are readily biodegradable (Verschueren,

1996). Degradation and utilization of alkylamine as a source

of energy is widespread in natural environments as dis-

cussed for glyphosate degradation. On the basis of the above

details, a proposed pathway for GA degradation is presented

in Fig. 8.

GB

GB (isopropyl methylphosphonofluoridate) is a highly toxic

nerve agent first produced in Germany in 1937 (Bakshi et al.,

2000). The term Sarin is an acronym of its discoverers

(Gerhard Schrader, Ambros Rudriger and Van der Linde).

Immediate death from exposure occurs because of respira-

tory tract failure (Rickett et al., 1986). Other routes of

exposure include the gastro-intestinal tract and skin absorp-

tion (Spruit et al., 2000). GB was implicated in terrorist

attacks in 1994 and 1995 in Japan, which caused death and

Table 3. Chemical, physical and biological properties of some organophosphorus chemical warfare agents

Name

First made

(Year)

Vapour pressure

(mmHg)

Volatility

(mg m�3)

Solubility in

water (g L�1)

Lethal dose

Breathing

(mg min�1 m�3)

Skin

(mg individual-1)

Tabun (GA) 1936 0.07 600 98 150–400 1000–1700

Sarin (GB) 1938 2.9 17 000 00 75–100 1000–1700

Soman (GD) 1944 0.3 3900 21 35–50 50–100

VX 1952 0.0007 10 30 10 6–10



injured many people (Abu-Qare & Abou-donia, 2002).

People exposed to GB during the incident in Japan reported

symptoms such as darkness of vision, ocular pain, dyspnoea

and headache. A review on GB effects on health is available

elsewhere (Abu-Qare & Abou-donia, 2002). GB is non-

persistent, volatile and completely soluble in water and

subject to acid and alkaline hydrolysis (Munro et al., 1999).

Like other xenobiotics, the fate of GB in soil includes

biotic and abiotic degradation, evaporation and leaching.

More than 90% of GB added to soil was reported to be

degraded within 5 days (Small, 1984); however, degradation

is comparatively slow at low temperature (Morrill et al.,

1985; Sanches et al., 1993). As discussed for GA, several

bacteria have been reported to degrade G agents including

GB. The major metabolites identified for GB degradation

are isopropylmethylphosphonic acid (IMPA) and methyl

phosphonic acid (MPA) (Mulbry & Rainina, 1998; Munro

et al., 1999). Chemically, IMPA is extremely stable and is

predicted to have a half-life of over 1900 years (Rosenblatt

et al., 1975). IMPA is relatively resistant to bacterial degra-

dation. However, two bacterial species, P. testosteroni and

Pseudomonas melophthora have been reported to degrade

Fig. 8. Possible pathways for microbial degrada-

tion of GA. The scheme is based on articles cited

in the text.



IMPA to MPA (Daughton et al., 1979). These bacteria

metabolize IMPA via cleavage of the C–P bonds to methane

and inorganic phosphorus compounds. Zhang et al. (1999)

reported biotransformation of IMPA by a microbial con-

sortium. Four mixed cultures were acclimated to IMPA. Two

of these cultures, namely APG and SX microorganisms, used

IMPA as the sole source of phosphorus. The intermediate

metabolites were identified as MPA and inorganic phos-

phates. Although attempts to use IMPA as a source of

carbon to support microbial growth were not successful, in

a bioreactor 85 mg L�1 IMPA was decreased to non-detect-

able level within 60–75 h. MPA is also susceptible to C–P

lyase producing bacteria (Zhang et al., 1999). Use of MPA as

a source of phosphorus by a P. putida has been observed

(Cook et al., 1978b). Several other bacteria were reported to

possess C–P lyases and they have been described for

glyphosate degradation. The microbial degradation pathway

for GB is presented in Fig. 9.

GD

Soman, or GD (pinacolyl methylphosphonofluoridate), is

structurally similar to GB. GD was the given identifier of

Soman post war (American denomination, GC was already

in medical use) when the information relating to Soman was

recovered by old Soviet Union in 1949. Its volatility is

intermediate between GA and GB. It is less water soluble

and more lipid soluble than the other two G agents, which

results in more rapid skin penetration and greater toxicity

(Munro et al., 1999). Like other G agents, GD is subject to

hydrolysis but the rate of hydrolysis is five times slower than

GA (Hambrook et al., 1971). The first step in hydrolysis is

fluoride removal to form pinacolyl methylphosphonic acid

(PMPA), which is then slowly degraded to MPA and

pinacolyl alcohol (Kingery & Allen, 1995). No data were

found on the fate of PMPA or pincolyl alcohol in the

environment. It is assumed that, like alkyl methylphospho-

nic acid, PMPA is probably resistant to degradation (Munro

et al., 1999). However, like other G agents, GD is hydrolyzed

by P. diminuta and several strains of Alteromonas (DeFrank

et al., 1993). Similarly, IMPA degrading consortia were able

to degrade PMPA as a sole source of phosphorus. In

successive batch experiments using immobilized cells,

PMPA level decreased from 164 mg L�1 to below the detec-

tion limit within 60 h (Zhang et al., 1999). The proposed

microbial degradation pathway is presented in Fig. 9.

VX

O-ethyl-S[2-(di-isopropylamino) ethyl] methylphospho-

nothioate was first discovered by British scientists. Later,

the US produced it in large quantities under code name VX.

It is a moderately persistent nerve agent characterized by a

P–S bond and, therefore, it belongs to the phosphorothio-

lates group. It is less volatile than G agents and does not

evaporate easily (Munro et al., 1999). VX is soluble in water

(30 g L�1 at 25 1C) and is relatively resistant to hydrolysis.

However, at acidic and extreme alkaline pH, cleavage of the

P–S bond predominates, resulting in formation of ethyl

methylphosphonic acid (EMPA) and diisopropylethyl mer-

captoamine (DIEM). The latter can be oxidized to bis (2-

diisopropylaminoethyl) disulfide (BIAEDS) (Yang et al.,

1993). At neutral and alkaline pH, the common pathway of

hydrolysis includes cleavage of C–O bonds to ethanol and S-

(2-diisopropyl aminoethyl) methyl phosphonothioate

(DIAEMP). The half-life of VX in water at pH 7 and 25 1C

is 17–42 days (Clark, 1989). Laboratory and field studies on

the fate of nerve agents demonstrated that loss is due to a

Fig. 9. Microbial degradation pathway for GB and GD. IMPA, isopropyl-

methyl phosphonic acid; PMPA, pinacolylmethyl phosphonic acid; MPA,

methyl phosphonic acid.



combination of abiotic and biotic processes such as evapora-

tion, hydrolysis and microbial degradation. According to one

study, 90% of added VX was lost from soil in 15 days (Small,

1984). Diethyl methyl phosphonate and BIAEDS were ex-

tracted when VX was added to soil (Sanches et al., 1993). In

other studies, EMPA and DIEM were found to be major

metabolites in soils (Kaaijk & Frijlink, 1977; Omar, 1998).

Further degradation of EMPA resulted in formation of MPA

(Omar, 1998). EMPA can be used as a phosphorus source for

natural microbial systems (Cook et al., 1978b; Mulbry &

Rainina, 1998). Diethyl dimethylpyrophosphonate, diisopro-

pylaminoethanol (DIPAE), diisopropylamine (DIPA) ethyl-

methylphosphonothioic acid (EMPTA) were reported as

other possible intermediate metabolites (Munro et al., 1999).

VX is resistant to microbial hydrolysis. It cannot be

hydrolyzed by any of the strains of Alteromonas isolated for

the degradation of G agents. OPH of P. diminuta is found to

be active against VX and Russian VX but its activity is less

than 0.1% toward VX as compared with parathion. However,

site-specific mutagenesis in OPH resulted to a 33% increased

activity against VX (Gopal et al., 2000). It was noted that VX

could be rapidly degraded by chemical oxidation of the P–S

bond using various peroxides (Yang et al., 1993) and mono-

magnesium perphthalate (Amitai et al., 1998). Oxidative

hydrolysis of VX produces EMPA and dialkylaminoethane-

sulfonate as compared with the corresponding alkylthion

hydrolytic product formed via the hydrolysis pathway. EMPA

has been reported to be degraded as a source of phosphorus

by two glyphosate-degrading bacteria, Burkholderia caryo-

philli and Pseudomonas testosteronis (Elashvili & DeFrank,

2001). A partially purified enzyme from B. caryophilli

bacterium has shown a broad specificity towards neutralized

nerve agents, including GF, GB, GD, VX and Russian VX

(Elashvili & DeFrank, 2001).

Oxidative hydrolysis of VX by the enzyme laccase from a

white-rot fungus, Pleurotus ostreatus was observed. The

mechanism of such degradation is not fully understood. It

was suggested that the sulfur atom is oxidized followed by

cleavage of the P–S bond (Yang et al., 1990). The nitrogen

atom at the b position to the carbon bound to the sulfur

atom was assumed to play an important role in the

enzymatic reaction. One suggested pathway is the formation

of an N-oxide intermediate in the N,N dialkyl aminoethyl

moiety at alkaline pH that may affect the cleavage of the P–S

bond. Cleavage of the S–C bond may also occur forming O-

ethyl methyl phosphorothoic acid and 2-diisopropylami-

noethanol (Yang et al., 1990). The proposed pathways of VX

degradation are shown in Fig. 10.

Detoxifying enzymes

Microbial enzymes that can hydrolyze organophosphorus

compounds have been identified and characterized from

different microbial species. The hydrolysis of organopho-

sphorus compounds leads to a decrease in mammalian

toxicity by several order of magnitudes and therefore this

step is also called detoxification. An excellent review on the

role of bacterial enzymes in detoxification of organopho-

sphorus nerve agents has been published recently (Raushel,

2002). Consequently, in the present article, we only review

the characteristics, improvement and utility of a few of the

most extensively studied organophosphorus hydrolyzing

enzymes. Several bacterial and fungal isolates with novel

enzyme/gene systems are reported (Table 4). However,

despite the apparent diversity of the enzyme systems, most

studies of organophosphorus degrading enzymes have fo-

cused on organophosphorus hydrolase (OPH) and organo-

phosphorus acid anhydrolase (OPAA), which are among the

most extensively studied enzymes in the biological sciences.

Organophosphorus hydrolase has been isolated from

several bacteria (Serdar et al., 1982; Mulbry & Karns, 1989a;

Singh et al., 1999). Among bacterial enzymes, OPH from P.

diminuta has the widest range of substrate specificity and,

therefore, has received most attention. OPH is a dimer of

two identical subunits containing 336 amino acid residues

(Dumas et al., 1989) that folds into a (ab)8-barrel motif

(Gerlt & Raushel, 2003). Each subunit contains a binuclear

zinc situated at the C-terminal portion. The two zinc atoms

are separated by about 3.4 A and linked to the protein

through the side chain of His 55, His 57, His 201, His 230,

Asp 301 and a carboxylated Lys 169. Both the Lys 169 and

the water molecule (or hydroxide ion) act to bridge the two

zinc ions together (Benning et al., 2001). A schematic

diagram of the structure of the binuclear metal centre within

the active site of OPH is presented in Fig. 11. It has a

molecular weight of 72 kDa.

Organophosphorus compounds bind to the binuclear

metal centre within the active site via co-ordination of the

phosphoryl oxygen to the b-metal ion. This interaction

weakens the binding of the linking hydroxide to the b-metal.

The metal-oxygen interaction polarizes the phosphoryl

oxygen bond and creates a more electrophilic phosphorus

centre. Subsequent nucleophilic attack by the bound hydro-

xide is assisted by proton abstraction from Asp 301. The

hydroxide attacks the phosphorus centre, resulting in weak-

ening of the bond to the leaving group (Raushel, 2002). A

working model for the OPH reaction mechanism is shown

in Fig. 12. In summary, the role of one metal ion in the

active site of OPH is to increase the electrophilicity of the

phosphorus centre through co-ordination with the non-

ester oxygen atom of the substrate, whereas the second metal

ion acts as a promoter of the attacking nucleophile (Efr-

menko & Sergeeva, 2001). However, questions regarding the

mechanisms of catalytic activity remained unanswered. The

pKa value of the bridging solvent is not known; it is believed

to be determined by variation of the kinetic parameters with



pH. In addition, it has still to be resolved whether the two

Zn ions within the active site have distinct functions or

whether they act in tandem (Raushel, 2002).

Organophosphorus hydrolase has a wide range of sub-

strate specificity. It hydrolyzes P–O, P–F, and P–S bonds to

different extents (Table 4). The lowest specificity is for the

P–S bond. However, the enzyme does not catalyze the

cleavage of carbonyl groups such as those found in p-

nitrophenyl acetate. Similarly, organophosphate diesters are

very poor substrates. The Kcat and VKm values for the

H3C

H3C

O

PS

O

PS NH5C2O

HO

O

PO

O

PS OH

HO

HO

O

POHHO

HO

O

POH

H3C

H5C2OO

O

SHO

CH2

VX

HS

CH2

DIEM

EMPA

BIAEDS

DIPAE

DIPA

HOH2C

H5C2O

CO2 + H2O NH4+ + CO2 + H2O

H3C

EMPTA

HOC2H4

sulfurous acid

phosphoric acid

DIAEMP

CH (CH3)2

CH (CH3)2

NCH2 CH2

CH (CH3)2

CH (CH3)2

CH (CH3)2

CH (CH3)2

CH3CH2OHEthanol

NCH2 CH2

CH (CH3)2

CH (CH3)2

N

N

NH

CH2

CH (CH3)2

CH (CH3)2

CH (CH3)2

CH (CH3)2

NCH2 CH2

CH (CH3)2

CH (CH3)2

C2H5OH

Fig. 10. Proposed pathways for VX degradation by microorganisms. The scheme is based on articles cited in the text. When the conversion of one

compound to another is believed to occur through a series of intermediates, the steps are indicated by dotted arrows. DIAEMP, diisopropyl aminoethyl

methyl phosphonothioate; DIEM, diisopropylethyl mercaptoamine; EMPA, ethylmethyl phosphonic acid; BIAEDS, bis-diisopropyl aminomethyl disulfide;

DIAPAE, diisopropyl amino ethanol; DIPA, diisopropyl amine; EMPTA, ethylmethyl phosphonothioic acid.



hydrolysis of paraxon by the enzyme have been determined

to be nearly 104 s�1 and 108 M�1 s�1, respectively (Omburo

et al., 1992).

The effects of metal substitution on the catalytic activity

of OPH were studied by removing the native metal (Zn)

from purified OPH and reconstitution with a series of

divalent cations which include Co, Cd, Cu, Fe, Mn and Ni

(Omburo et al., 1992; Di Sioudi et al., 1999; Benning et al.,

2001). Further enzymatic assays showed that Co21 had the

greatest activity against paraoxon (Omburo et al., 1992). It

was suggested that divalent cations increased the activity of

enzyme by assisting folding of expressed enzyme in the

medium (Manavathi et al., 2005). Site specific mutagenesis

was used to substitute the original histidinyl residues at

positions 254 and 257. Of these mutant enzymes, H254R

(histidine at 254 was replaced with arginine) and H257L

(histidine at 257 was substituted with leucine) demonstrated

a four- to five-fold higher catalytic activity against the P–S

bond (VX and demeton-S). Other mutants also showed

higher activity against Soman and other nerve agents (Di

Sioudi et al., 1999).

The hydrolysis of asymmetric organophosphorus com-

pounds catalyzed by OPH is stereoselective (Lewis et al.,

1988; Chae et al., 1994). For example, OPH degrades the Sp

isomer of EPN 21 times faster than the Rp isomer. Similarly

Sp isomers of acephate and methamidophos are catalyzed

preferentially by OPH (Hong & Raushel, 1999). To achieve a

practical solution to nerve agent contamination, the enzyme

should be able to degrade the racemic mixture, as both

isomers are usually present in compounds such as Soman,

GB and VX. This aim was achieved by rational modification

to the substrate binding activities. The size and shape of

these binding subsites were remolded through a rational

restructuring via site-directed modification (Wu et al., 2001;

Raushel, 2002). Preferential degradation of the Sp isomer of

the EPN by OPH presumably arises because the bulkier

phenyl substituent is better accommodated in the large

subsite and the ethyl group within the small subsite of OPH

for the Sp–enantiomer. To increase the degradation of the

Rp-enantiomer, the small subsite was expanded. This was

achieved by the substitution of Phe132, Ser308 and Ile106

to glycine and/or alanine. With these mutants, the

Table 4. Organophosphorus degrading microbial enzymes

Enzyme Origin MW Structure

Bond cleavage

P–O P–F P–S P–C

Bacterial

OPH Pseudomonas diminuta 72 Dimer 1 1 1 1

OPAA Alteromonas spp. 50–60 Monomer 1 1 � 1

OPDA A. radiobacter 70 Dimer 1 1 1 1

ADPase Nocardia sp. 43 Monomer 1 ND ND �AMPP Escherichia coli 52 Tetramer 1 ND ND ND

HOCA Pseudomonas monteilli 19 Monomer 1 ND � ND

SC-OPH SC strain 67 Tetramer 1 ND ND �NS-OPH Nocardiodes simplex 45 Monomer 1 ND ND ND

PEH Burkholderia caryophilli 58 Tetramer � � � 1

C–P lyase Pseudomonas spp. 200 ND � � � 1

Phosphonatase Bacillus cereus 37 Dimer � � � 1

Fungal

A-OPH Aspergillus niger 67 Monomer � ND 1 ND

P-OPH Penicillium lilacinum 60 Monomer 1 ND 1 ND

Laccase Pleurotus ostreatus ND ND � � 1 ND

1, positive activity; � , no activity; ND, not determined; MW, molecular weight.

Fig. 11. Representation of the structure of the binuclear centre within

the active site of the bacterial organophosphorus hydrolase (reproduced

with permission from Benning et al., 2001).



stereoselectivity for the Sp and Rp enantiomers for EPN

decreased from 21 : 1 to 1 : 1.3 (Chen-Goodspeed et al.,

2001; Wu et al., 2001).

The course of evolution of OPH in bacteria remains an

unresolved question. Organophosphorus compounds were

released as pesticides after World War II and it is difficult to

understand how these OPH catalytic activities appeared so

quickly. One suggestion is that this enzyme was already

present in the environment before the application of orga-

nophosphorus pesticides. This hypothesis has gained im-

portance following isolation of similar OPH from

phylogenetically and geographically different micro-organ-

isms (Mulbry et al., 1986; Somara et al., 2002). The recent

finding of OPH encoding genes in a field soil which has

never been exposed to this group of pesticides supports this

hypothesis (Singh et al., 2003c). Another possibility is that

this enzyme has evolved new substrate specificity from pre-

existing enzymes as it has been shown that OPH (phospho-

triesterase) could acquire phosphodiesterase activity by

alteration of only one amino acid (Shim et al., 1998). Urease

has been found to have carbamylated lysine as a bridging

ligand with binuclear Ni at the active site (Park & Hausinger,

1995). The binuclear centre of urease and OPH was found to

be remarkably similar. However, the chemical nature of the

active sites of these enzymes is quite different (Raushel,

2002). A larger group of enzymes with similar active site

architecture has been identified (Holm & Sander, 1997).

Interestingly this superfamily also includes atrazine chlor-

ohydrolase.

A similar enzyme, OPDA, has been isolated from A.

radiobacter and was found to have 90% homology to OPH

at the amino acid level and a very similar overall secondary

structure (Horne et al., 2002b; Yang et al., 2003). Despite

these similarities, the two enzymes have different substrate

specificities. There is about a 30-sequence difference be-

tween OPH and OPDA. The largest group consists of 19

residues at the C-terminus. In addition, two regions with

significant difference in OPDA from OPH are a large pocket

at the active site and another in the region of the protein that

is responsible for binding phenyl ethanol (an inhibitor) in

OPH. Apart from the sequence difference, the water struc-

ture in this region differs in the two enzymes. There are two

water molecules that form a network of hydrogen bonds in

OPDA. The equivalent residues in OPH could not form the

same hydrogen bonds but were found to be stabilized by the

presence of phenyl ethanol. These differences at the active

site of OPDA likely to give it a preference for substrates that

have shorter alkyl substituents. Further studies using site-

specific mutagenesis in OPH gave a series of mutants that

had activities similar to those of OPDA. Yang et al. (2003)

argued that this alteration in the active site gave substrate

specificity and represented the progressive natural evolution

of the enzyme from OPH to OPDA. However, the observa-

tion that alterations of amino acids in the active site

increases the activity of enzymes emphasizes that enzyme

efficiency depends on several factors and that evolution can

take place in many ways.

Another enzyme that has received considerable attention

recently, for detoxification of organophosphorus nerve

agents is OPAA. A highly active OPAA from Alteromonas

undina was isolated and purified and is composed of a single

polypeptide with molecular weight 53 kDa (Cheng et al.,

1993). However, another OPAA isolated from Alteromonas

sp. JD6.5 is composed of 517 amino acids with molecular

weight of 60 kDa. However, one from Alteromonas halo-

planktis contains 440 amino acids (Cheng et al., 1996, 1997)

with molecular weight 50 kDa. The 10 kDa difference be-

tween OPAAs of these two Alteromonas spp was found to be

Fig. 12. Working model for the catalytic me-

chanism for hydrolysis of organophosphorus

nerve agents by organophosphorus hydrolase

(reproduced with permission from Raushel,

2002).



due to the presence of an extended C-terminal region in the

JD 6.5 enzyme (DeFrank & White, 2002). The three-dimen-

sional structure of this enzyme is not yet known. It possesses

low catalytic activity against P–O but high activity against

P–F bonds. OPAA also displays stereoselectivity towards the

chiral phosphorus centre by displaying preference for the

Rp-enantiomers. It also exhibits an additional preference for

the stereochemical configuration at the chiral carbon centre

of the Soman analogue (Hill et al., 2001). This observation

was confirmed by a recent study, which found that OPAAs

along with wild-type phosphotriesterase catalyze preferen-

tially the hydrolysis of (1) GF isomer in a racemic mixture

(Harvey et al., 2005). However, OPAAs from different

species of Alteromonas have demonstrated wide variation in

catalytic activity, with the highest activity observed with the

enzyme obtained from Alteromonas sp. J.D.6.5 (DeFrank &

White, 2002). OPAA from Alteromonas sp. JD6.5 has high

degree of homology at amino acid level with E. coli X-Pro

dipeptidase (48%) and E. coli aminopeptidase P (31%).

Further molecular and biochemical analyses of OPAA have

established that this enzyme is a prolidase; a type of

dipeptidase cleaving dipeptide bond with a prolyl residue at

the carboxyl terminus (Cheng et al., 1999). Although the

native function of OPAA is not yet known, it has been

suggested to play an important role in cellular dipeptide

metabolism because all OPAAs were found to have activity

against several dipeptides (DeFrank & White, 2002). Mole-

cular modelling studies with Soman and Leu-Pro revealed

that the three-dimensional structure and electrostatic den-

sity maps of the two are nearly identical (DeFrank & White,

2002). This explains why several dipeptidase enzymes have

catalytic activity against organophosphorus compounds.

Determination of the crystal structure of OPAA may con-

firm this hypothesis. Both prokaryotes and several eukar-

yotes have been found to possess this enzyme (Mazur, 1946;

Hoskin et al., 1999), which suggests that OPAA is not a

newly evolved enzyme (Cheng et al., 1999). In a recent study,

aminopeptidase P (AMPP) was found to catalyze the

hydrolysis of a wide range of organophosphate triesters.

AMPP belongs to the family of protein-specific peptidases

and catalyzes the cleavage of amino-terminal X-Pro peptide

bonds (Jao et al., 2004). This enzyme possesses many

similarities with OPAA, such as two divalent (Mn21) ions,

which are critical for maximal activity of AMPP. Replace-

ment of the Mn21 ion with other divalent ions except Co21

resulted in the loss of catalytic activity (Jao et al., 2004).

AMPP is a tetramer with each sub-unit composed of a ‘pita

bread’ fold of the C-terminus domain (Wilce et al., 1998).

The active site of AMPP is located at the C-terminal portion

of the b-sheet with the two manganese ions separated by

33 A. The two ions are co-ordinated by Asp 260, Asp 271,

His 354, Glu 383, Glu 406 and two water molecules. A water

molecule or hydroxide ion bridging between the two metal

ions is believed to be strongly activated, and acts as the

nucleophile in the attack on the scissile peptide bond

Xaa–Pro. A single amino acid mutation with hydrophobic

side chains such as R153W, R153L, R370L increased the

hydrolysis rates towards most of the organophosphorus

substrates compared to the wild-type enzyme (Jao et al.,

2004). This result suggests that the further protein engineer-

ing of AMPP may significantly enhance the cleavage of P–O

bond in a variety of organophosphorus compounds.

Other structurally and functionally different organopho-

sphorus degrading enzymes have been reported. Three

unique parathion hydrolases were isolated, purified and

characterized from gram-negative bacterial isolates. One

cytosolic hydrolase described as an ADPase (aryldialkylpho-

sphatase) from Nocardia sp. strain B-1 was composed of a

single sub-unit of approximately 43 kDa (Mulbry, 1992).

Another hydrolase from strain SC was membrane bound

and is composed of four identical sub-units of 67 kDa.

While having some common features such as constitutive

production and similar temperature optima around 40 1C,

the substrate specificity and structure of these enzymes

differ one from another, and also from the other known

OPHs (Mulbry & Karns, 1989a). A unique phosphotriester-

ase has been characterized from Nocardioides simplex NRRL

B-24074. The purified enzyme is monomeric, has a native

molecular weight of 45 kDa, is constitutively expressed and

located in the cytoplasm. This enzyme is quite distinct with

respect to its activity towards different substrates and also in

its stimulation or inhibition by divalent cations and dithio-

threitol (Mulbry, 2000). Another novel phosphotriesterase

HocA (hydrolysis of caroxon) was isolated from P. monteilli

(Horne et al., 2002c). This enzyme was required by the host

for phosphate metabolism and was suggested to be evolved

from phosphodi- or mono-esterase. HocA (19 kDa) does

not require a metal ion for its catalytic activity but was

reported to be less efficient at hydrolyzing organopho-

sphorus compounds than other reported microbial phos-

photriesterases (Horne et al., 2002c). HocA is not a

metalloenzyme and its activity is controlled by the presence

of phosphate in the medium.

There have been a number of reports on isolation and

purification of organophosphorus hydrolyzing enzymes

from pure isolates or from mixed cultures of bacteria. Only

a few organophosphorus hydrolyzing enzymes have been

reported from fungi. Degradation of phosphorothiolates by

a broad-spectrum fungal enzyme, laccase (phenol oxidase)

from a white-rot fungus P. ostreatus was reported (Amitai

et al., 1998). This is a significant observation as this enzyme

attacks P–S bond, which is comparatively resistant to OPH

and OPAA cleavage. Laccase was observed to be capable of

complete and rapid degradation of VX and Russian VX

(Amitai et al., 1998). Several white-rot fungi are capable of

organophosphorus degradation (Table 2), and it will be



interesting to know if the degradation capability of all white-

rot fungi towards organophosphorus compounds is

mediated by the presence of laccase, or whether different

fungi possess different enzyme systems. A novel organopho-

sphorus degrading fungal enzyme (A-OPH) was isolated

from A. niger ZHY256 that could hydrolyze a range of P–S

bonds containing organophosphorus compounds (Liu et al.,

2001). This 67-kDa enzyme has found to have optimal pH at

7 with thiol and sulfhydryl groups in the active catalytic site.

A-OPH does not require divalent cations for activation;

however, Cu21 was found to activate its activity. Another

novel organophosphorus-hydrolyzing enzyme was purified

from Penicillium lilacinum BP303. Interestingly, this peni-

cillium OPH (P-OPH) was found to degrade various

organophosphorus compounds by cleaving both P–O and

P–S linkages (Liu et al., 2004). The molecular mass of P-

OPH is 60 kDa with optimal activity at pH 7.5. The purified

enzyme was reported to be a member of a cysteine hydrolase

group and similar to A-OPH and OPAA. Despite having

several similar structural components, P-OPH is different

from OPH, OPAA and A-OPH in its catalytic activity. P-

OPH degraded all organophosphorus compounds with P–O

and P–S linkage, whereas OPH in Flavobacterium sp. only

attacks P–O bond and A-OPH splits only P–S linkage.

The first reported enzyme able to degrade the phospho-

nates, 2-phosphonoacetaldehyde hydrolase (phosphona-

tase), was isolated from B. cereus (La Nauze et al., 1970).

The isolated and purified phosphonatase showed optimal

activity at pH 8, required Mg21 for its activity, and was

inhibited by sulfhydryl reagents (La Nauze et al., 1970).

Phosphonatase resembles alkaline phosphatase in many

properties but has narrow substrate specificity. Phosphona-

tase does not degrade phosphomonoesters and is not a

metalloenzyme (Kononova & Nesmeyanova, 2002). This

enzyme has been reported from several bacterial species

and can degrade a range of phosphonates including glypho-

sate (Baker et al., 1998). Further analysis suggests that

phosphonatases belong to a new family of hydrolase having

a high conservative aspartate residue in their active site to

which the phosphoryl group from a lysine residue of the

enzyme is transferred (Baker et al., 1998). Phosphonatase is

a homodimer of 33–37 kDa subunits, and its active site is

mainly comprised of polar amino acid residues, which

suggests that phosphonatase may have a common origin to

the NAD dependent superfamily of dehalogenase, phospho-

tase and phosphomutase (Kononova & Nesmeyanova,

2002). Another interesting enzyme that can degrade phos-

phonates is C–P lyase. There is one report suggesting partial

purification of this enzyme from Pseudomonas sp. GLC11

(Selvapandiyan & Bhatnagar, 1994). The molecular mass of

this enzyme was reported to be approximately 200 kDa and

it was found to be localized in the periplasmic space of

bacteria. However, subsequent reports suggest that C–P

lyase manifests its activity only in cells and has never been

reliably found in cell-free extracts (Kononova & Nesmeya-

nova, 2002). This obstacle considerably limits the possibility

of understanding the mode of catalytic action of C–P lyase.

A good review on the proposed mechanism of phosphonates

degradation by C–P lyase on the basis of computer model-

ling and gene structures is available (Kononova & Nesmeya-

nova, 2002). In brief, the action is initiated by the generation

of a phosphonyl radical. Subsequent cleavage of this reactive

intermediate would lead to metaphosphate and alkyl moi-

eties as the corresponding alkenes. Abstraction of hydrogen

by an alkyl radical would yield the corresponding alkanes as

products, which is a specific feature of phosphonate degra-

dation.

Several microbial isolates have been reported to have

further novel enzyme/gene systems but most of these were

not isolated or purified such as C–P lyase (Kertesz et al.,

1994a), methyl parathion hydrolase (Zhongli et al., 2001)

and chlorpyrifos degrading enzyme (Singh et al., 2004).

Most of the enzymatic studies were carried out to improve

the catalytic activity of OPH and OPAA by protein or

genetic engineering. Nonetheless, a few novel enzymes have

been purified recently and these enzymes differed in mole-

cular mass, substrate specificity and, sensitivity to chemicals

(Table 4). Improvements of known enzymes should con-

tinue but this trend of characterizing new and diverse

enzymes from prokaryotes and eukaryotes needs to be

sustained to find the best bioremedial enzymes with optimal

activity in detergents and in the presence of metal ions, and

which have a broad pH and temperature optima. Discovery

of diverse microbial enzymes will also facilitate understand-

ing of the evolutionary structure–function relationship of

organophosphorus-degrading enzymes.

Genetic basis of organophosphorusdegradation

The first described organophosphorus degrading (opd) gene

was found in P. diminuta, and was shown to be present on a

plasmid (Serdar et al., 1982). The plasmid size was 66 kb and

was termed pCMS1. By cloning into different plasmids and

into the broad range cloning vector, it was shown that a

1.5 kb BamHI fragment with single restriction sites for SalI,

PstI and XhoI encoded this enzyme (Serder & Gibson, 1985).

Mulbry et al. (1987) found that the opd gene from Flavo-

bacterium sp. strain ATCC 27551 was encoded on a 43-kb

plasmid (pPDL2) and had a similar restriction map to the

opd gene from P. diminuta. Southern hybridization experi-

ments demonstrated that the opd gene from the two bacteria

possessed significant homology. This finding of homologous

genes on two non-homologous plasmids from two phylo-

genetically and temporally different bacteria isolated from

different geographical regions suggests that the gene may be



a mobile genetic element or transposon (Mulbry et al.,

1987). Sequencing of the opd gene proved that the gene

from both bacteria had identical sequences (Harper et al.,

1988). Later, the nucleotide sequence of the opd gene from P.

diminuta was determined and a single open read frame

located (Serder et al., 1989). The opd gene from Flavobacter-

ium sp. consists of 1693 base pairs with one open reading

frame (Mulbry & Karns, 1989b). The opd gene has been

cloned into various bacterial strains (Serder & Gibson,

1985), actinomycetes (Steiert et al., 1989), fungi (Xu et al.,

1996) and insect cells (Dumas et al., 1989). Several other

bacteria have opd genes with almost identical nucleotide

sequences (Chaudry et al., 1988; Somara et al., 2002). Horne

et al. (2002b) reported a similar opd gene from A. radio-

bacter P230 isolated in Australia for coumaphos degrada-

tion, which was chromosome based. This gene, called opdA,

was approximately 88% identical at the nucleotide level to

opd (Horne et al., 2002b). Sequencing of the whole genome

revealed the presence of opd-like genes in Mycobacterium

tuberculosis (Philipp et al., 1996) and E. coli (Blattner et al.,

1997), which supports the hypothesis that the opd gene may

be transposon based. However, this evidence has been

provided only recently (Siddavattam et al., 2003) where a

complete sequence of a region of plasmid pPDL2 from

Flavobacterium sp. is reported, which has identical restric-

tion patterns to opd containing plasmid pCMS1 of P.

diminuta (Fig. 13). The opd gene was found to be flanked

by an insertion sequence, ISF1sp1 (encoding a complete

istAB operon), which is a member of the IS21 family, and

downstream by a Tn3-like element (tnpA and tnpR) encod-

ing a transposase and a resolvase. It was also observed that

adjacent to opd, but transcribed in the opposite direction, is

an open reading frame (orf243) which encodes a polypep-

tide of 27 kDa that plays a role in the degradation of p-

nitrophenol (the major degradation product of parathion

and methylparathion). A 2.5-kb region upstream of the opd

gene contains two ORFs transcribed in the same direction as

opd, which have significant homology to the IstA and IstB

genes. Similarly, Horne et al. (2003) reported that opdA in A.

radiobacter P230 is transposable. A tnpA gene was found

upstream of the opdA. The two genes are flanked by

insertion sequences which resembles Tn 610 transposon

from Mycobacterium fortuitum. Two additional putative

ORFs separate opdA and tnpA, and the deduced translation

products show similarity to two proteins encoded on the

Geobacillus stearothermophilus IS5376 (Horne et al., 2003).

These observations of linkage of the opd/opdA genes to IS

elements and transposase genes supports the idea that the

widespread distribution of the opd gene could be due to its

lateral transfer by a combination of transposition and

plasmid transfer (Siddavattam et al., 2003).

The evolutionary origin of the opd gene is presently not

known. However, it has been argued that the closest homo-

logues of the IstA, IstB, and tnpR genes are all found in

strains of Agrobacterium tumefaciens and an opd gene has

recently been reported from a strain of A. radiobacter

(Horne et al., 2002b). These similarities could indicate an

evolutionary origin for these genes in Agrobacterium (Sid-

davattam et al., 2003). Another hypothesis is that the gene

was present in the environment long before organopho-

sphorus compounds were commercialized. The presence of

genes similar to opd in several bacteria that have never been

exposed to this group of compounds also supports this

argument (Philipp et al., 1996; Blattner et al., 1997; Richins

et al., 1997). Recently, higher copy numbers of the opd gene

were observed in higher pH soils (Singh et al., 2003a, c).

Fig. 13. Map of the sequence region from pPDL2 showing eight ORFs identified together with the location of the IR sequences that flank ISF1sp1 (&).

The extent of the plasmid subclones (pWWM1079, pWWM44 and pSM1) used as a starting material for sequencing is indicated and the region

identified previously as homologous to plasmid pCMS1 of Pseudomonas diminuta is indicated by a dashed arrow. The complete opd gene cluster (on

pSM2) is included. BamH1, EcoRI, HindIII, and PstI sites in the sequence are indicated by B, E, H and P, respectively. pWWM1079 is a recombinant

plasmid that contains an entire conserved region of opd gene cluster located downstream of the opd gene, pWWM44 is a recombinant plasmid that

contains an entire conserved region of opd gene cluster located upstream of the opd gene. Isolation and religation of the larger HindIII fragment from

pWWM1079 gave a plasmid (pSM1) carrying part of orf243 with tnpA and tnpR. The conserved region upstream of opd was isolated on a 5.2-kb HindIII

fragment from plasmid pWWM44 and cloned into pSM1 in the correct orientation to reconstitute the entire region, thereby giving pSM2 (reproduced

and adapted with permission from Siddavattam et al. (2003)).



This observation has considerable significance because this

field had not been exposed to organophosphorus com-

pounds. It is worth mentioning that OPH has optimal

activity at higher pH. It is possible that opd or its ancestor

has some important function to play at high pH. It is

unlikely that such gene/enzyme systems evolved to protect

against anticholinesterase compounds since bacteria do not

contain acetylcholinesterases but their widespread distribu-

tion suggests that the gene/enzyme system does serve an

important function. One suggested function is the role of an

opd-like gene in phosphate metabolism (Horne et al.,

2002b). These genes may have evolved from pre-existing

mono-phosphatase or phosphodiesterase as it has been

shown that OPH (phosphotriesterase) could acquire phos-

phodiesterase activity by the change of only one amino acid

(Shim et al., 1998). Isolation, characterization and cloning

of a novel phosphodiesterase (which degrades organopho-

sphate xenobiotics) gene from Delftia acidovorans is strong

evidence to support this hypothesis (Tehara & Keasling,

2003). This gene shows sequence similarity to cyclic AMP

(cAMP) phosphodiesterase and cyclic nucleotide phospho-

diesterases and exhibits activity on cAMP in vivo when the

gene is expressed in E. coli, suggesting that it may have

evolved from a common ancestor of the cAMP gene and

may regulate cAMP levels in bacterial cells (Tehara &

Keasling, 2003). Further evidence in the form of the gene

structure and function relationships is required to reach to a

definitive conclusion.

Another organophosphorus degrading gene which has

received considerable attention is opaA, first isolated and

cloned from Alteromonas sp. JD6.5 (Cheng et al., 1996,

1997). In spite of functional similarity with the opd gene,

no sequence homology was found between them. One ORF

of 1552 nucleotides was identified that codes for OPAA. The

OPAA enzyme has amino acid sequence similarity with that

of E. coli AMPP and human prolidase. It is believed that

opaA and the prolidase gene may have evolved from the

same ancestral gene and may play a role in bacterial peptide

metabolism (Cheng et al., 1996). However, the role of pepP

gene (encodes for AMPP) in organophosphorus degrada-

tion has also been reported (Jao et al., 2004).

Genes for phosphonate degradation have received con-

siderable attention due to their commercial exploitation in

genetically modified crops. Seventeen open reading frames

(phnA to phnQ) were reported to be involved in phospho-

nate uptake and degradation by E. coli (Chen et al., 1990).

Further study suggested that the genes phn ABQ were not

involved in degradation and that the phn operon therefore

consisted of 14 genes which are transcribed from a single

promoter preceding the phnC gene. On the basis of sequence

analysis and comparison with known motifs, reading frames

phnCDE have been proposed to form a phosphonate trans-

port complex, whereas phnF and phnO may be involved in

regulation. Two genes (phnNP) are not required for phos-

phonate use and may encode accessory proteins for the C–P

lyase. The remaining seven genes (phnG-M) were suggested

to be involved in the C–P lyase complex itself (Metcalf &

Wanner, 1991). Parker et al. (1999) reported the presence of

a homologous part of the E. coli phn gene cluster in

Sinorhizobium meliloti. By cloning, phnGHIJK genes were

identified in S. meliloti. However, several genes from phn

cluster of E. coli were not detected in S. meliloti, despite the

fact that S. meliloti appeared to have a broader substrate

specificity. This observation suggests that not all genes in

phn cluster may be required for phosphonate metabolism or

that these genes are functionally redundant in S. meliloti.

The molecular-genetic analyses suggest that the process of

phosphonate degradation involves a multi-component sys-

tem with constituents localized in the membrane and

periplasm. This may explain the failure by several research

groups to isolate and purify cell free C–P lyase despite

considerable effort. Expression of the genes for phosphonate

degradation is controlled by phosphorus supply to the cell

and they have been suggested to be the part of the pho

regulon on the basis of their similarity to promoter se-

quences (Mulbry & Karns, 1989b) and the requirement of

regulatory genes (Wacket et al., 1987). Two other novel

genes involved in the degradation and utilization of glypho-

sate, glpA and glpB, were isolated and sequenced from

Pseudomonas pseudomallei. The gene glpA (1260 bp long)

encodes an enzyme (phosphotransferase) of 420 amino

acids, which confers increased tolerance to glyphosate. The

gene glpB encodes a protein (309 amino acids long) with the

ability to break the N–C bond of glyphosate to yield

aminomethyl phosphonic acid (Penaloza-Vazquez et al.,

1995). Another gene involved in glyphosate metabolism,

pehA (encodes PEH), was cloned and sequenced from B.

caryophilli PG2982 (Dotson et al., 1996).

Several other genes with similar or identical function but

totally different nucleotide sequences have been reported

(Table 5). A methyl parathion degrading (mpd) gene was

isolated from Plesiomonas sp. strain M6 (Zhongli et al.,

2001). Sequencing and cloning of mpd revealed that this

gene is 1061 bp long and encodes a 35-kDa product. When

the mpd nucleotide sequence and predicted protein se-

quence were compared with those in the Genbank database,

no region of extensive DNA homology was observed. The

highest similarity with predicted protein sequence was

found to be 31% with beta-lactamase, suggesting significant

novelty of the gene-enzyme system. Another novel gene,

adpB (which encodes ADPase), for organophosphorus de-

gradation was obtained from Nocardia strain (Mulbry,

1992). This 1600 bp long gene does not share homology

with any of the other known genes involved in organopho-

sphorus degradation. Horne et al. (2002c) isolated and

cloned a gene called hocA (hydrolysis of caroxon) gene from



P. monteilli, consisting of 501 bp; it has a different sequence

from all other known organophosphate degrading genes.

The hocA gene encodes a 19-kDa protein that can degrade a

range of oxon and thion organophosphorus compounds.

Increased expression of hocA was observed from an integra-

tive hocA–lacZ fusion when the culture was grown in the

absence of phosphate, suggesting that it might be part of the

pho regulon. This structural diversity and functional simi-

larity suggests that these genes have evolved from different

ancestors, but that the structural similarity of xenobiotics to

natural compounds, and the constant mutation in the

bacterial genome and rapid doubling time play an impor-

tant role in the evolution of the gene/enzyme systems for

xenobiotic degradation.

Biotechnological advancements

Biotechnological aspects of the degradation of organopho-

sphorus compounds have received considerable attention

recently due partly to their high mammalian toxicity

and partly to the requirements of the CWC. Several com-

pounds need sensible detoxification and disposal techniques

because of their bulk usage, storage and widespread use.

Current methods for detoxifying these compounds mainly

rely on incineration and landfills. Incineration of chemical

warfare agents has received strong and sustained opposition

from the public and environmental groups because of

potentially toxic emissions. This process is also very costly,

as it requires considerable amounts of energy to reach the

high temperatures needed to destroy the pollutants. Land-

fills provide an adequate short term solution but leaching of

pollutants to ground water is a major source of concern.

Bioremediation with micro-organisms is therefore an at-

tractive alternative to these conventional techniques for

pollutant disposal.

Munneck (1976) first reported the potential use of para-

thion hydrolase producing bacteria for the detoxifi-

cation and disposal of organophosphorus compounds.

Later, successful use of OPH producing bacteria for com-

plete destruction of coumaphos in cattle-dip waste was

reported (Kearney et al., 1986; Karns et al., 1987). The use

of a consortium of microbes in a filter bioreactor for

destruction of coumaphos has been very successful. Two

units, each capable of treating 15 000 litres of waste

cattle-dip at a time, have been operational since 1996.

The US Department of Agriculture has been using these

units for treatment of coumaphos waste generated under its

cattle fever tick eradication programme (Mulbry et al.,

1998). The use of whole living cells for bioremediation

presents some difficulties such as delivery of fresh

inocula and nutrient composition. To avoid these difficul-

ties, the use of cell free OPH was carried out successfully

(Karns et al., 1998). It was observed that addition of non-

ionic detergents and cobalt salts increased the efficiency of

OPH in waste cattle-dips. Both native and recombinant

OPHs, immobilized on a nylon membrane, powder and

tubing (Caldwell & Raushel, 1991a), silica beads and glass

(Caldwell & Raushel, 1991b) have been used for the detox-

ification of organophosphorus compounds. OPH from P.

diminuta was immobilized based on the formation of non-

composite protein-silicone polymers and was a highly

active, stable and versatile biocatalyst for the liquid and gas

phase detoxification of organophosphorus compounds. It

was fabricated as monoliths, sheets, thick films, granulates

or monoporous foams (Gill & Ballesteros, 2000). OPH and

OPAA were also incorporated into an aqueous fire-fighting

Table 5. List of genes, their origin, vector, and gene products involved in degradation of organophosphorus compounds

Gene Organism(s) Location Encoded enzyme Reference

opd Pseudomonas diminuta Plasmid OPH Serder et al. (1989)

Flavobacterium sp. Plasmid OPH Mulbry et al. (1986)

Flavobacterium balustinum Plasmid OPH Somara & Siddavattam (1995)

Pseudomonas sp. Plasmid OPH Chaudry et al. (1988)

opaA Alteromonas sp. JD6.5 Chromosome OPAA Cheng et al. (1996)

Alteromonas haloplanktis Chromosome OPAA Cheng et al. (1997)

Alteromonas undina Chromosome OPAA Cheng et al. (1996)

opdA Agrobacterium radiobacter Chromosome OPDA Horne et al. (2002b)

hocA Pseudomonas monteilli Chromosome ND Horne et al. (2002c)

mpd Plesiomonas sp. Chromosome ND Zhongli et al. (2001)

adpB Nocardia sp. B-1 Chromosome ADPase Mulbry (1992)

PdeA Delftia acidovorans Chromosome Phospho diesterase Tehara & Keasling (2003)

PepA Escherichia coli Chromosome AMPP Jao et al. (2004)

Phn Escherichia coli Chromosome Phosphonatase Chen et al. (1990)

Sinorhizobium meliloti Chromosome Parker et al. (1999)

glp A&B Pseudomonas pseuodomallei Chromosome C–P lyase Penaloza-Vazquez et al. (1995)

pehA Burkholderia caryophilli Chromosome PEH Dotson et al. (1996)



foam which was active and stable (Cheng et al., 1999;

Raushel, 2002).

Recently, considerable efforts have gone into the develop-

ment of OPH biosensors to detect contamination of orga-

nophosphorus compounds. This bioanalytical technique

provides rapid, cost effective and in-field monitoring of

contaminants. The use of OPH in this technique has

advantages over acetylcholine esterase and enzyme-linked

immunoassays (ELISA). ELISA can be sensitive but like

most immunoassays, targets a single compound and re-

quires multisteps. Acetylcholine esterase based enzyme

inhibitions are well suited for screening applications; how-

ever, due to their irreversible inhibition by organopho-

sphorus compounds, they are not particularly well suited

for process control monitoring applications that require

rapid and repeated measurements (Chough et al., 2002).

Because OPH based biosensors respond to organopho-

sphorus compounds as substrates rather than as inhibitors,

they show considerable potential for applications that

require repetitive analysis (White & Harmon, 2005). Two

assay formats were used with OPH for biosensors: potentio-

metric measurement of local pH change (Mulchandani

et al., 1998, 1999b) and amperometric measurement of

electroactive enzyme products (Wang et al., 1999; Chough

et al., 2002). A dual amperometric and potentiometric flow-

injection biosensor detection system was developed recently

which used different physical transducers simultaneously in

connection with OPH. This enhanced the output and

allowed discrimination between various organophosphorus

compounds (Wang et al., 2002a). This biosensor, which

combines the advantages of both the amperometric device

and potentiometric detection, displays well-defined signals

from the oxidized leaving group and has been accomplished

with silicon-based pH sensitive electrolyte-insulation-semi-

conductor transducers (Wang et al., 2002a, 2003). While

offering a fast response, such enzyme biosensors have

limitations in terms of the number of samples that can be

handled and discriminated among organophosphorus com-

pounds. To overcome this problem, an on-chip enzymatic

assay for screening organophosphorus nerve agents, based

on pre-column reaction of OPH, electrophoretic separation

of the phosphonic acid products, and their contactless

conductivity detection has been developed (Wang et al.,

2004). On-chip enzymatic assays combine the selectivity and

amplification features of biocatalytic reactions with the

analytic features and versatility of microchip devices. This

new microsystem holds promise for field screening of

organophosphorus compounds with the advantages of

speed/warning, efficiency, portability, sample size and cost.

Information regarding the structure and function of

enzymes and pathways involved in biodegradation will

provide opportunities for improving enzyme activities.

Catalytic mechanism and enzyme properties can be ma-

nipulated by site-directed mutagenesis guided by computer-

modelled three-dimensional structure of enzymes (Cheng

et al., 1999). Site-directed mutagenesis was successfully used

to enhance the activity of OPH against racemic mixtures of

organophosphorus enantiomers. The size and shape of the

substrate binding subsites were remoulded through rational

restructuring via site-directed mutagenesis (Wu et al., 2001;

Raushel, 2002). However, rational design can fail sometimes

due to unexpected influences exerted by substituted amino

acids. Another limitation imposed by this rational approach

is that only a limited sequence space can be explored at one

time. Irrational approaches such as DNA shuffling, random

priming and staggered extension processes have been sug-

gested as preferable alternatives to direct the evolution of

enzymes (Cheng et al., 1999). DNA shuffling was success-

fully used to isolate an improved variant of opd cloned E.

coli, which can degrade methyl parathion 25 times faster

than the wild type (Cho et al., 2002). However use of

enzymes for detoxification of pesticides is not a cost-

effective process. It was expected that genetic engineering

would provide a means for cheaper production of microbial

enzymes. The OPH encoding gene opd has been cloned

under different promoters to increase the amount of OPH

produced by cloned bacteria (Cheng et al., 1999).

It was suggested that the use of growing or non-growing

whole cells immobilized onto supports could offer cheaper

and more effective options. The major problem associated

with whole cell bioreactors is mass transport limitation of

substrate across the cell membrane where OPH resides

(Mulchandani et al., 1999a). Uptake of organophosphorus

compounds as a rate limiting factor has been reported by

several groups (Hung & Liao, 1996; Elashvili et al., 1998).

This barrier of substrate transport can be overcome by

treating cells with permeabilizing agents such as EDTA and

DMSO. However, several enzymes are sensitive to such

treatment and immobilized viable cells cannot be subjected

to permeabilization (Mulchandani et al., 1999a). To over-

come this difficulty, OPH was successfully anchored and

displayed onto the surface of E. coli using the same

Lpp–OmpA fusion system used for beta-lactamase (Richins

et al., 1997). Whole cells with surface expressed OPH had

seven times higher activity than whole cells expressing

similar amounts of OPH intracellularly. A genetically en-

gineered E. coli expressing both OPH and cellulose-binding

domain on the cell surface was constructed, enabling the

simultaneous hydrolysis of organophosphorus nerve agents

and immobilization via specific adsorption to cellulose

(Wang et al., 2002b). OPH was expressed on the surface by

the use of a truncated ice-nucleation protein-fusion system

while the cellulose-binding domain was surface anchored by

the Lpp–OmpA fusion system.

Microorganisms that can completely degrade and miner-

alize whole molecules of organophosphorus compounds



have not yet been reported, but a diverse set of organisms

has been isolated that are capable of collectively mineralizing

these compounds. For example, some microbes can rapidly

hydrolyze parathion and methyl parathion and utilize DETP

as a source of carbon or phosphorus but quantitatively

produce p-nitrophenol as a by-product. However, other

bacteria can utilize p-nitrophenol as a source of energy. A

micro-organism engineered to complete mineralization of

organophosphorus compounds would avoid the generation

of toxic hydrolytic products. This goal was achieved by the

genetic engineering of p-nitrophenol mineralizing Moraxella

sp., which was transformed with the opd gene. The truncated

ice nucleation protein anchor was used to express the OPH

onto the surface of the bacterium, overcoming the potential

substrate uptake limitation (Shimazu et al., 2001). In

another study, Walker & Keasling (2002) engineered P.

putida KT 2442 to use parathion as a source of carbon and

energy. Two separate plasmids, one harbouring a native opd

gene (pAWW04) and another harbouring an operon encod-

ing enzymes for p-nitrophenol transformation to b-ketoa-

dipate (pSB337), were introduced into P. putida; the

plasmids enabled the bacterium to utilize 0.8 mM parathion

as a source of carbon. Recently, an E. coli expressing

phosphotriesterase from A. radiobacter (opdA) and glycer-

olphosphodiesterase from Enterobacter aerogenes (GpdQ),

which can use methyl parathion as a source of phosphorus,

was used to screen for mutants with enhanced activity. This

process of directed evolution produced a variant with

increased protein expression and increased activity against

organophosphorus (McLoughlin et al., 2005). The intro-

duction of all degradative genes into a single organism

allows for future optimization of gene expression and the

potentials to utilize further directed evolution to optimize

degradation rates and minimize the metabolic burden

placed on the cell.

Perspectives

Degradation of organophosphorus compounds has at-

tracted considerable attention because of their widespread

use as pesticides, their high mammalian toxicity, and the

CWC (1993). A large number of microorganisms have been

isolated and characterized that can degrade organopho-

sphorus compounds by mineralization or co-metabolism.

Some microorganisms can degrade several compounds and

some can degrade only one or few structurally similar

organophosphorus compounds. Because hydrolysis of orga-

nophosphorus compounds reduces mammalian toxicity by

several orders of magnitude, the environmental fate of

degradation products has not received much attention from

the scientific community. Complete pathways of parathion

and glyphosate degradation are known but the pathways for

several other organophosphorus compounds are not yet

fully understood. This area of research needs concerted

efforts as degradation products of several compounds are

pollutants and may have deleterious effects on the environ-

ment and non-target organisms.

Organophosphorus degrading enzyme OPH has been

characterized, its three-dimensional structure determined

and its catalytic activity elucidated. Site-specific mutagenesis

has been carried out successfully to increase the catalytic

activity against poor substrates, and to decrease the stereo-

selectivity of the enzyme. In addition to the potential

bioremedial use of microbes and enzymes for dealing with

organophosphorus contamination in the environment,

there has been considerable interest in the use of organo-

phosphorus degrading enzymes prophylactically and ther-

apeutically for organophosphorus poisonings (Sogorb et al.,

2004; Petrikovics et al., 1999, 2000a, b). Future areas of

research include increasing enzyme activity against poor

substrates and improving enzyme catalytic activities in

mixtures of chemicals.

Sequencing and structure determination of new proteins

will provide missing links to relate and elucidate evolution

mechanisms. Determining the three-dimensional structure

of OPAA would be a major boost for furthering studies on

the manipulation of enzymatic activity, which in turn may

help in developing efficient enzymatic destruction methods

for chemical warfare agents, as this enzyme has higher

catalytic activity towards G-agents.

The biodegradation of chemical warfare agents has re-

cently been a major area of research because of the urgency

to destroy all stocks by 2007. Recent efforts have provided

successful laboratory results. However, only a few isolated

microorganisms have the capacity to degrade chemical

warfare agents. A comprehensive screening of microbial

dipeptidase activity from different sources may provide

new gene/enzyme systems with higher activities against G-

agents or V-agents. For example, screening of anaerobic

microorganisms and extremophiles may be useful but

this so far has received little attention for organophos-

phorus compound degradation. Another challenge for

the scientific community is scaling-up of laboratory success

to the field because of differential behaviour of iso-

lated micro-organisms in the environment and also because

stockpiles of chemical warfare agents contain a mixture

of different chemical contaminants and degradation pro-

ducts. The application of genetic engineering and biochem-

ical techniques to improve and evolve natural

biodegradative capabilities will ultimately create strains

capable of degrading complex mixtures of compounds. For

example, micro-organisms were isolated which can utilise

several neutralized chemical warfare agents as a phosphorus

source but require addition of excess nitrogen and carbon

which are rate limiting. Introducing cells containing C–P

lyase activity in consortia or C–P lyase gene in degrading



microorganisms might accelerate the overall degradation

process.

Acknowledgements

Work in BKS laboratory is supported by a grant from the

Scottish Executive Environment and Rural Affairs Depart-

ment (SEERAD). We are grateful to Ms Pat Carnegie for

drawing chemical structures and pathways. We also thank

Drs Pete Millard, Charlie Shand, Colin Campbell (MI), Alun

Morgan, Gary Bending (HRI), and Michael Kertesz (Uni-

versity of Manchester) for helpful discussions.

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