REVIEW OF LITERATURE

2.1 Pesticides

The term “pesticide” refers to chemical substances intended to be biologically

harmful to pests (weeds, insects, mould or fungi) and interfere with the normal

biological processes of living organisms. The basic aim of developing different

pesticides is to obtain higher yield from agricultural fields. Thus, are designed to be

persistent in environment to achieve effective control of pests over long period of

time. The pesticides are classified into five main classes (Table 2.1).

Table 2.1: Classification of pesticides on the basis of their mode of application

Pesticides Examples

Insecticides

Organophosphates Chlorpyrifos, Parathione

Carbamate Esters Primicarb, Aldicarb

Pyrethroids Fenvelerate, Cypermithrin

Organochlorines DDT, Cyclohexane

Botanical Insecticides Scilliroside, Strychnine

Herbicides

Chlorophenoxy compounds 2,4-dichlorophenoxy acetic acid

Bipyridyl derivatives Paraquat, Diquat

Rodenticides

Metal Phosphide Zinc Phosphide, Magnesium Phosphide

Organofluorine Sodium fluoroacetic acid, Gliftor

Thiourea Promurit, Thiosemicarbazide

Anticoagulants 4-Hydroxycoumarine, Indian dione

Fungicides

Hexachlorobenzene Anticaril

Organomercurials Phenylmercuric acetate

Phenolic compounds Pentachlorophenol, 2,4-Dinitrophenols

Phthalimides Thiochlorfenphim, Folpet, Captan

Dithiocarbamates Amobam, Asomate, Azithiram

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Fumigants

Phosphine Bedfont EC80

Brominated compounds Ethylene di-bromide, Methyl bromide

Dibromochloropropane Fumazone, Nembrom

The populations of insects are more in number as compared to other living

organisms and are known to cause extensive damage at food production to storage

levels form domestic front to industrial units. The role of insecticides is crucial to

control and kill insects and is used in very large amount per year world wide. Aktar et

al. 2009 reported that among the total consumption of pesticides in India the major

share is of insecticides (76%) followed by fungicide (13%), herbicides (10%) and

others (1%). The higher consumption of insecticides may be attributed to higher

hatching rate of insects in warm humid and tropical climate which provides favorable

breeding environment.

On the basis of their chemical properties insecticides may be classified as:

2.2 Classification of pesticides

2.2.1 Inorganic compounds

These insecticides are used both at domestic front and agricultural fields

against beetles, cockroaches and moths. These are low molecular weight compounds

and do not contain carbon in their chemical composition. Few commonly used

inorganic compounds are paris green, lead arsenate, sodium fluoride etc.

2.2.2 Thiocyanates

Thiocyanates such as lethane and thanite are used to tackle public health

problems to control mosquitoes, flies and bed bugs. These insecticides are found in

photoprocessing, electroplating, and chemical-fertilizer wastewaters (Goncalves et al.

1998). Thiocyanates are toxic to humans and animals resulting in problems like

irritability, hallucination, convulsions and nervousness because of their strong

tendency to bind to proteins and as non-competitive inhibitors of enzymes in central

nervous system (Lewis, 1992; Wood et al. 1998).

2.2.3 Organochlorines

Organochlorine (OC) insecticides include well known compounds such as

dichlorodiphenyltrichloroethane (DDT), heptachlor, hexachlorobenzene etc. which

were introduced in 1940s and extensively used during the 1950s–1970s in food and

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5

non-food crops such as corn, wheat and tobacco. They can be classified into four

categories: dichlorodiphenylethanes (e.g., DDT), cyclodienes (e.g., heptachlor),

chlorinated benzenes (e.g., hexachlorobenzene) and cyclohexanes (e.g.,

hexachlorocyclohexane). Dichlorodiphenylethanes and cyclodienes were used as

agricultural insecticides whereas chlorinated benzenes and cyclohexanes are used as

fungicides and antimicrobials. Organochlorine pesticides vary in their chemical

structures and mechanisms of toxicity. These are lipophilic compounds and keeps on

accumulating in the higher trophic levels and their concentration magnifies along with

the food chain in fatty tissues of the body (Poon et al. 2005). The chemicals affect the

humans and animals health due to their ability to interact with endocrine system

(Munozde-Toro et al. 2006). The compounds may get transferred from nursing

mothers to offspring via breast milk (Munozde-Toro et al. 2006).

The exposure of hexachlorobenzene to humans leads to formation and

accumulation of heme precursors in the body by interfering with normal synthesis of

heme which may cause hyperpigmentation in skin, weakness, arithritis etc. The

exposure of hexachlorocyclohexane on the other hand blocks inhibitory

neurotransmitters in the central nervous system causing seizure and death. There have

been reports of blood dyscrasias anemia, leucopenia after high exposure of HCH

(Morgan et al. 1980; Rugman et al. 1990). The exposure of humans and animals to

these chemicals occurs through food of animal origin, dust, and soil (Cruz et al.

2003).

2.2.4 Carbamates

In early 1950s carbamate were introduced as pesticides and are being used in

pest control due to short lifetime, effectiveness and broad spectrum of biological

activity. These are used for treatment of seeds, soil and crops for controling insects

e.g. leaf monors, cockroaches, ants, scale insects, mealy bugs and whiteflies.

Carbamates include three subgroups: 1) N-methylcarbamate ester of phenols, e.g.

methiocarb and propoxur 2) N-methyl- and N-dimethylcarbamate esters of

heterocyclic phenols e.g. primicarb and carbofuran 3) Oxime derivatives of aldehydes

e.g. aldicarb. Carbamate pesticides are related to organophosphates by their mode of

action but carbamate pesticides are less dangerous as the dose required producing

minimum poisoning symptoms and mortality in humans is higher for carbamate

compounds than for organophosphorus compounds (Goldberg et al. 1963; Vandekar

et al. 1971).

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2.2.5 Pyrethroids

Pyrethroids are synthesized from pyrethrins of pyrethrum (the oleo-resin

extract of dried Chrysanthemum flower) and include cypermithrin, fenvelerate,

deltamethrin, bifenthrin etc. These insecticides are applied in agricultural fields to kill

insects causing damage to crops (e.g., alfalfa, cotton, lettuce) and orchards (e.g.,

almonds, pistachios, and peaches) and are used in the home in pet sprays and

shampoos to remove lice/ticks. In natural pyrethroids, the attachment of alcohol to

dichlorovinyl derivative of the cyclopropanecarboxylic acid moiety gave less toxic

and low persistence property to these pesticides. However, attachement of α-cyano

group to the 3-phenoxybenzylalcohol moiety enhances the toxicity of synthetic

pyrethroids (Litchfield 1983). The chrysanthemic and pyrethroic acids of pyrethrins

are strongly lipophilic and interfere with ionic conductance of nervous system of

insects by prolonging the sodium current to paralyze them (Reigart and Roberts

1999). These are less toxic to mammals due to their rapid biodegradation by

mammalian liver enzymes, while insects are more susceptible to these chemicals due

to absence of such enzymatic system (Reigart and Roberts 1999).

2.2.6 Organophosphorus pesticides

There are number of organic phosphorus compounds which were synthesized

around 1800s but organophosphate (OP) based pesticides were used first in 1937

(Dragun et al. 1984). The first commercialized OP insecticide Bladan, containing

TEPP (tetraethyl pyrophosphate) was synthesized by German chemist Gerhard

Schrader, who later synthesized insecticide parathion in 1944 (Gallo and Lawryk

1991). Thus, their development as insecticides took place in the late 1930s and early

1940s.

2.3 Structure of organophosphorus pesticides

The general structure of OP pesticide is represented as:

R2

R1 P

O/S

X

• R1 and R2 are the aryl or alkyl groups which either directly bonded to

phosphorous (P) as in case of phosphinates or through oxygen (phosphates) or

sulphur (phosphothioates) atoms.

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• In phosphonates R1 is directly bonded to P but R2 is bonded to oxygen. In

phosphonothioates R1 is directly bonded to P but R2 is bonded to sulphur

atom.

• In phosphoramidates both or atleast one R group is attached to –NH2.

• X is the ‘‘leaving group’’ (as it gets displaced upon hydrolysis of OP).

• Oxygen and sulphur (O/S) are directly attached to P by a double bond.

Based on their chemical structure different types of OPs were produced and

used for control of insects as shown in Table 2.2.

Table 2.2: Organophosphate pesticides (OPs) based on their chemical structure:

Types of OPs Examples

Phosphate Dichlorvos, Chlorfenvinphos

Phosphonate Trichlorfon

Phosphorothioate Chlorpyrifos, Chlorpyrifos methyl,

Diazinon, Parathion

Phosphorothiolate Demeton-S-methyl, Omethoate,

Profenofos

Phosphorodithioate Dimethoat, Disulfoton, Malathion,

Thiometon

Phosphonothioate EPN (O-ethyl O-(4-nitrophenyl)

phenylphosphonothioate)

Phosphorothioamidate

Phosphoramidothionate)

Isofenphos, Propetamphos

Phosphorothioamidate

(Phosphoramidothiolate)

Methamidophos

Phosphonofluoridate Sarin

2.4 Use of organophosphorus pesticides

Organophosphate pesticides are widely used in variety of cereal crops, oil

seed crops, vegetable crops and fruit crops for effective control of insect such as

termites, jassids, aphids, white fly, leaf hoppers etc. They are also used in some

veterinary and human medicines to remove parasitic insects, in various public hygiene

products generally for the control of cockroaches and termites (Racke et al. 1994).

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8

Among them chlorpyrifos (O, O-diethyl-O-3,5,6 trichloro-2-pyridyl

phosphorothioate) is being extensively used in developing countries like India where

it was the fourth highest consumed pesticide after monocrotophos, acephate and

endosulfan, in the year 2000 (Ansaruddin, 2003).

2.5 Toxicity of organophosphorus pesticides

The persistent organochlorine pesticides were widely used around the world

before the 1970s. Initially OPs were considered as safe alternative to organochlorines

but over the years due to their inordinate use their accumulation and exposure lead to

acute toxicity to non-target organisms. There are reports of high mammalian toxicity

due to OPs resulted in three million poisonings and 200,000 deaths annually

(Karalliedde and Senanayak 1999; Sogorb et al. 2004). OP’s have been cited as

potential cause of many diseases as given in Table 2.3.

Table 2.3: Diseases due to excessive use of OP’s:

Disease’s References

Miscarriage Ballentyne & Marrs, 1992

Asthma Hodgson & Smith, 1992

Polyneuropathy Lotti et al. 1993; Mc Conell et al. 1994

Cancerous lymphomas Newcombe et al. 1994

Chronic neurological sequelae Steenland et al. 1994

Immune dysfunction Newcombe et al. 1994

Saku disease Dementia, 1994

Parkinson’s disease Mars, 1995

Sensory neuropathy Stephans et al. 1995

Peripheral neuropathy, Intermediate

syndrome

Marrs, 1995

Psychiatric disorder Stephan et al. 1995

Myalgic encephalomyelitis or Chronic

fatigue syndrome

Behan, 1996

BSE (bovine spongiform encephalopathy) King 1996; Gordan et al. 1998

CJD (Creutzfeldt-Jakob disease) King, 1996

Motor neuron disease and Multiple

sclerosis

Purdey, 1996

Delayed psycho-neurodegenerative Purdey, 1996

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syndrome

Gulf War syndrome Hom et al. 1997; Jamal, 1998

Mental retardation Weiss, 1997

Induced hypothermia Gordan et al. 1998

The accumulation of chlorpyrifos in soil has many adverse effects as at higher

concentrations (10–300 mg Kg-1) it results in lowering the number of di-nitrogen-

fixing bacteria as well as total bacterial population (Martinez et al. 1992). This leads

to decrease in nitrogen and phosphorus content of soil. There have been reports of

delayed seedling emergence, fruit deformities and abnormal cell division upon

prolonged exposure to chlorpyrifos (NCAP, 2000; Sardar and Kole 2005).

Although, solubility of chlorpyrifos is less in water even then its toxicity is

prevalent in aquatic ecosystem. In case of fish and aquatic invertebrates, chlorpyrifos

is found to be moderately to highly toxic (US Environmental Protection Agency,

2002). Van Wijngaarden et al. (2005) reported in microcosm’s studies that

cladocerans (small crustaceans, commonly called water fleas), other zooplankton and

phytoplankton are adversely affected when they were exposed to chlorpyrifos.

2.5.1 Mode of action of organophosphorus pesticides

These pesticides act primarily at the synapses, altering the regulation of the

transmission of the signal from one cell to the next by inhibiting the enzyme acetyl

cholinesterase (AChE). This enzyme normally rapidly deactivates/hydrolyse

acetylcholine, a major neurotransmitter in animals into choline and acetylCoA to

prevent over stimulation of nerves. Organophosphorus compounds inhibit the normal

activity of the acetylcholinesterase by covalent bonding to the enzyme, thereby

changing its structure and function. They bind to the amino acid serine 203 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 acetylcholine esterase is very slow and may take

hours or days, resulting in accumulation of acetylcholine at the synapses, leading to

over stimulated and jammed nerves (Manahan, 1992). This inhibition causes

convulsion, paralysis and finally death for insects and mammals (Ragnarsdottir,

2000).

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The organophosphate pesticides can mimic hormones and acts as endocrine

disrupters (Prakash et al. 1992). Some OP’s have been reported to be mutagenic,

causing chromosomal abberations and other genetic toxicity to humans (Kiraly et al.

1979; Flessel et al. 1993).

2.5.2 Bioaccumulation of organophosphorus pesticides

The bioaccumulation is the primary cause of toxicity of pesticides as their

higher concentrations in biological systems leads to major health problems as listed in

Table 2.3. The persistence of OP in soil has also been related to the organic matter,

clay content and iron and/or aluminiumoxy(hydro)oxide content of the soil (Weber,

1972). These have higher affinity to absorb/adsorb the pesticides and act as a sink for

such hydrophobic compounds and discharge into other phases included living

organisms and plants (Boonsaner et al. 2002). Parathion, an organophosphate

pesticide, has been found to persist in soil for more than 16 year (Stewart et al. 1971).

The bioaccumulation of OPs is primarily related to their molecular weight and

aqueous phase solubility as cited in Table 2.4.

There are reports regarding their bioaccumulation in living systems ranging

from blue green algae to higher systems. Chlorpyrifos due to their very low aqueous

phase solubility (2ppm) had the highest bioaccumulation factor in the blue-green

algae, aquatic plants, gold fish, mosquito fish and Mytilus galloprovincialis, a

Mediterranean mussel (Spacie and Hamelink, 1985; Lal et al. 1987). Thus, being at

the top of the food chain humans indirectly gets much adversely affected by

bioaccumulation of chlorpyrifos and other pollutants in aquatic fauna (Serrano et al.

1997).

Table 2.4: List of commonly used organophosphate pesticides along with their

chemical structure, molecular weight and aqueous phase solubility.

Organophosphate

pesticides

Chemical structure Mol. Wt/Solubility

in water at 25º C

Monocrotophos

223.2/

100 g l-1

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Chlorpyrifos

350.6/

2 mg l-1

Diazinon

304.35/

40 mg l-1

Parathion (R=CH2CH3 X=H)

Methyl parathion (R=CH3

X=H)

291.3/

12.4 mg l-1 and

263.2/

55-60 mg l-1

Phorate

260.38/

50 mg l-1

Dimethoate

229.28/

25 g l-1

As evident from the Table 2.4 chlorpyrifos has least water solubility and presence of

three chlorine atoms attached to its ring make it more resistant to microbial

degradation. Due to these structural features the limiting rate of degradation results in

its accumulation in environment (Volkering et al. 1998; Angelova and Schamander

1999)

2.6 Fate of organophosphate pesticides

The improper storage facilities and handling of insecticides in different parts

of world prone to higher pest attack generally result in pollution of environment

Alemayehu, 2004; Biratu, 2005). The leakage of corroded pesticide containers in open

or inappropriate stores and their burning add to pollution world wide risking the

human and environmental health (Curtis and Olsen, 2004; Minh et al. 2006; Misra

and Pandey, 2005). Furthermore, empty pesticide containers at domestic front for

storage of fuel, food and water causing direct exposure to human populations (Dalvie

and London, 2001; Dalvie et al. 2006). The disposal of empty pesticide containers

near or into irrigation canals and streams is also a normal exercise for farmers

(Damalas et al. 2008).

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Pesticides undergo various changes in environment including their adsorption

transmission and degradation depending on the physicochemical nature of the

pesticide and the soil (Redondo et al. 1997). The predominant processes involved in

transformation of such molecules is mediated by microbes (Vink and Van der Zee,

1997) followed by photolysis or photodegradation and chemical transformations

(Roberts, 1998; Stangroom et al. 2000).

Thus, generally fate of pesticide involves both biological and non-biological

agents.

2.6.1 Abiotic transfer/transformation of OP pesticides

The abiotic processes involved either their transport where parent compound

remains unchanged and simply transferred from one matrix to another depends on the

physicochemical properties of pesticides itself (Stangroom et al. 2000) e.g.

volatilization, leaching (Laabs et al. 2000), runoff (Moore et al. 2002), absorption and

adsorption of pesticides (Yu et al. 2006) or by abiotic transformations by

photodegradation (Walia et al. 1988) and chemical hydrolysis (Liu et al. 2001).

Volatilization can transform pesticides from liquid or solid into gaseous form

depending on temperature of the surface and air currents in that area (Yates et al.

2002; Haith et al. 2002). The run off/soil erosion causes movement of pesticides

either on their dissolution in the water or through attachment with soil particles or

sediments and add a fairly large part (e.g., 5–30% for chlorpyrifos) of the overall

mass transfer from ground soil to surface water aquifers (Bailey et al. 1974; Richards

and Baker, 1993; Wood and Stark, 2002; Luo and Zhang 2009). Leaching also plays

role in transmission of higher water soluble pesticides to ground water through

fractures, root holes and earthworm burrows in earth crust (Stagnitti et al. 1994;

Magri and Haith, 2009). Pesticides can also be absorbed/adsorption by plants and

soils depend on their water solubility, soil characteristics and properties of pesticide

itself (Gan et al. 1996; Trapp, 2000; Davis et al. 2002; Paranychianakis et al. 2006;

Johnson et al. 2007).

2.6.1.1 Photodegradation

Photodegradation is one of the abiotic routes of degradation for many

organophosphate pesticides including certain pyrethroids and urea pesticides

(Stangroom et al. 2000). The reports indicated that even after the photodegradation of

primary toxic compound the toxic intermediates and products persists for longer time

periods (Stangroom et al. 2000; Zamy et al. 2004). The organophosphate pesticides

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may undergo photodegradation by direct photolysis under different sources of light

including sunlight, mercury and Xenon lamps. The absorption maxima of 240–310

nm in UV region is mandatory to exhibit photodegradation of OPs, which can be

further enhanced by the presence of oxygen and humic substances acting as natural

sensitizers (Pehkonen and Zhang, 2002). Photolytic studies of OP by irradiation

revealed that their degradation leads to formation of more toxic oxons (Zamy et al.

2004). The photodegradation of chlorpyrifos results in formation of chlorpyrifos-oxon

as a single major product and longer irradiation is required for complete removal of

toxic degradation products (Kralj 2007). While in case of azinphos-methyl, malathion

and malaoxon two toxic photoproducts, i.e., phosphorodithioic O,O,S-trimethyl ester

and phosphorothioic O,O,S-trimethyl ester, having the potential to inhibit

acetylcholine esterase were identified.

2.6.1.2 Chemical degradation

The chemical degradation and microbial degradation is difficult to distinguish

as both the process goes side by side. Further the physical properties of the soil also

plays integral part, as the clay content in soil leads to increase in surface area which

enhances hydrolytic conversion (Yaron, 1978). Smalling and Aelion in 2006 studied

that in different phases of estuarine sediments chemical processes involving sorption

and hydroxylation brings up 30 to 70% removal of atrazine and its metabolites.

Some of the pesticides are hydrolysed under acidic conditions and others are

hydrolysed under basic conditions in the soil. Thus, pH plays an important role in

hydrolysis of pesticides depending on the nature of the pesticide. According to a study

by Huang et al. (2000) the hydrolysis of chlorpyrifos is enhanced under basic

conditions. The effluent from different industries is used in many parts of the world

for ferti-irrigation. However this may increase dissolve organic matter content and

alter the pH of the soil resulting in chemical hydrolsis of pesticides (Stevenson, 1982;

Muller et al. 2007).

2.6.2 Biotic transformation of OP pesticides

Xenobiotic compounds like organophosphate pesticides are man made

compounds and were not previously present in nature. Consequently the natural

microflora does not have potential to metabolize these pesticides due to lack of

enzyme and proper transport processes. But over the year due to excessive use of

xenobiotic compounds microbes have evolved the new degradation pathways

resulting in accelerated degradation of such compounds (Seffernick and Wackett,

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2001; Johnson and Spain, 2003). For instance Seffernick et al. in 2001 suggests that

enzyme Atrazine chlorohydrolase (AtzA) responsible for dechlorination of herbicide

atrazine under strong selective pressure was evolved from enzyme melamine de-

aminase (TriA) and is 98% similar to the TriA. The accelerated bioremediation under

natural conditions is also helped by transfer of genes among different microbial

cultures by transformation, transduction and conjugation (Ghigo 2001 and Fux 2005).

Chlorpyrifos, an OP compound has large variation in half life from less than

60 days to more than 100 days attributed to difference in formulations, moisture and

organic carbon content of the soil, climatic condition, soil pH, native microbial

community and availability of desired genes among these microbial populations to

degrade chlorpyrifos (Getzin, 1981; Chapman & Chapman, 1986; Singh et al. 2002).

The most common metabolic pathway of chlorpyrifos degradation in soil involves an

initial hydrolytic cleavage of the P–O–C bond leading to the formation of

diethylthiophosphoric acid (DETP) and 3,5,6-trichloro-2-pyridinol (TCP).

The resistance of chlorpyrifos to enhanced microbial degradation may be

attributed to accumulation of TCP, which is known to have anti-microbial activity that

may hinder the proliferation of natural microbial diversity including chlorpyrifos

degraders (Racke et al. 1990). The studies suggest that TCP is more soluble in water

as compared to its parent compound thus may degrades faster depending upon

metabolic potential of the microbial populations of the polluted site (Barrett et al.

2000; Caceres et al. 2007).

The microbial diversity at the contaminated site is constituted by diverse

group of microscopic organisms like bacteria, fungi, viruses, protozoa and algae.

Among them bacteria, fungi and to some extent algae are the main contributors to

degradation of pesticides.

N

Cl Cl

ClHO

+

diethylthiophosphate

3,5,6-trichloro-2-pyridinol (TCP)

Hydrolysis

P

O

CH2

CH3

S

C

H2

H3C O OHN

Cl Cl

Cl

OP

O

CH2

CH3

S

C

H2

H3C O

Chlorpyrifos 0,0-diethyl 0-(3,5,6-trichloro-2-pyridinyl)

phosphorothioate

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Microbial transformation can be mainly achieved by following three different

mechanisms:

• Biodegradation

• Co-metabolism and

• Bioaccumulation

Bacteria are dominantly involved in accelerated biodegradation of pesticides

(Racke and Coats, 1990). The bacterial strains from different taxonomic groups with

potential to degrade the organophosphorus insecticides have been reported (Yasouri,

2006; Li et al. 2008). While in case of fungus along with normal degradation process

co-metabolic process of OP degradation by lignolytic or cytochrome 450 associated

enzymes is also prevalent (Fernando and Aust, 1994; Yadav et al. 2003). On the other

hand reports in literature indicating that algae and blue green algae are mainly

involves in binding, adsorption and bioaccumulation of organophosphate pesticides

during their mesocosm and microcosm studies (Lal and Lal 1987; Laabs et al. 2007;

Pablo et al. 2009).

2.6.2.1 Bacterial degradation of OPs

In literature there are many reports regarding screening and isolation of

microbes capable of degrading pollutants under laboratory conditions. However, their

use at the contaminated sites under field scale application has not been successful

(Pilon-Smits 2005; Dua et al. 2002; Kuiper et al. 2004). The reasons behind this

include the competition faced from the natural microflora and microfauna of the soil,

suboptimal nutrition or nutritional deficiency leading to low microbial growth, non

availability or less bioavailability of the pollutant desired to be degraded and the

growth inhibitory concentration of pollutant itself (Kuiper et al. 2004; Dillewijn et al.

2007).

The other major factor is selective consumption of substrates by microbes as it

has been observed that chlorpyrifos and diazinon hydrolyzed 30–1000 times slower

than paraoxon as latter was more favorable substrate of organophosphate hydrolase

(Dumas et al. 1989). The reason for such selective substrate consumption may be due

to lack of transport processes for uptake of these pollutants (Dumas et al. 1989; Chen

and Georgiou, 2002). The direct expression of OPH at the surface of E.coli resulted in

80% increase in degradation efficiency of OPs (Richins et al. 1997) so as to overcome

this limitation of availability. Racke and Coats (1987) reported Pseudomonas sp.

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capable of degrading isofenphos was not able to degrade other organophosphorus

insecticides. On the other hand, there are reports that organophosphorus hydrolase

(OPH), present in many microbes of diverse groups have the ability to hydrolyze

broad range of organophosphates by cleaving the P–O, P–F and P–S bonds (Ang et al.

2005).

Further, the studies suggest that the higher concentration of pesticides was

always found to be detrimental for microbial growth by inhibiting respiration and

enzymatic activities of soil microflora. Rangaswamy et al. (1994) reported that OPs

like monocrotophos and quinalphos and synthetic pyrethroids viz. cypermethrin and

fenvalerate at concentration of more than 5 kg ha−1 have a deleterious effect on

dehydrogenase and protease activities in soil. Pozo et al. (1995) also reported that

presence of organophosphate pesticides may leads to increase or decrease in microbial

biomass in the soil depending on the ability of soil microflora to degrade/tolerate OPs.

Diazinon exhibited an reversible repressive effect on the urease-producing microbial

inhabitants of soil (Ingram et al. 2005). Shan et al. (2006) reported the suppressed

growth of bacterial, fungal, and actinomycete populations in the presence of

chlorpyrifos (10 mg kg-1). Vischetti et al. (2007) reported reduction in soil microbial

biomass in an Italian soil field by 25% and 50% after chlorpyrifos treatment at 10 mg

kg-1and 50 mg kg

-1, respectively.

Inspite of these limitations, microbial degradation of organophosphate

pesticides is an important process responsible for their biotic degradation in

environment (Felsot, 1989). There are reports regarding ability of microbes to degrade

pesticides co-metabolically or as source of carbon, nitrogen and phosphorous.

Sethunathan and Yoshida (1973) reported Flavobacterium sp. having the ability to

degrade chlorpyrifos in liquid medium by cometabolism. Similarly, Serdar et al. in

1982 isolated Pseudomonas diminuta degrading chlorpyrifos co-metabolically rather

than as a source of carbon On the other hand, Ohshiro et al. (1996) reported that

Arthrobacter sp. strain B- 5 can use chlorpyrifos as a substrate rather than a co

metabolite. Singh et al. (2003) isolated six chlorpyrifos degrading bacteria capable of

degrading chlorpyrifos in both liquid medium and soil. Dutta et al. (2004) observed

increase in net microbial biomass carbon (MBC) in chlorpyrifos treated soils as

compared to the control containing no chlorpyrifos. Singh et al. (2004) also reported

degradation of chlorpyrifos by pure culture Enterobacter sp. B-14 in liquid as well as

in soils. Wang et al. (2006) reported degradation of chlorpyrifos by pure culture of

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Bacillus laterosporus DSP. Li et al. (2008) reported isolation of chlorpyrifos-

degrading bacterial strains Dsp-2, Dsp-4, Dsp-6 and Dsp-7 identified as

Sphingomonas sp., Stenotrophomonas sp., Bacillus sp. and Brevundimonas sp.

respectively and few other strains distinguished as members of Pseudomonas sp. from

chlorpyrifos-contaminated samples.

There are many chlorpyrifos degraders reported but very few degraders are

known to degrade the compound at higher rates. Mallick et al. (1999) reported

complete degradation of 10 mg l-1 of chlorpyrifos in the mineral salts medium by

Flavobacterium sp. ATCC 27551 and Arthrobacter sp. within 24 h and 48 h

respectively. As described earlier, the major limitation in the process of chlorpyrifos

degradation is the formation an anti-microbial compound 3,5,6-trichloro-2-pyridinol

(TCP) which may also affect the growth of chlorpyrifos-transforming microorganisms

(Racke et al. 1990). A report by Racke and Coats (1990) indicate that transformation

of 30 mg kg-1 of chlorpyrifos in the soil resulted in production of TCP which repress

the proliferation of microbes introduced into the soil. The accelerated degradation of

chlorpyrifos was observed either due to the ability of degraders to tolerate TCP or

their potential to mineralize TCP efficiently at a rate higher rapidly than the rate of its

formation in the medium. There are reports regarding degradation of both chlorpyrifos

and TCP in aqueous phase (Feng et al. 1998; Mallick et al. 1999; Horne et al. 2002;

Bondarenko et al. 2004). A Stenotrophomonas sp. isolated by Yang et al. (2006) was

found to be a degrader of both chlorpyrifos and TCP. On the other hand, Singh et al.

(2004) isolated Enterobacter sp. capable of degrading chlorpyrifos was not able to

degrade TCP but utilize diethylthiophosphate as carbon and phosphorus source. The

Enterobacter species in this case showed the tolerance against TCP even at higher

concentrations (150 mg l-1), which might be the reason of effective chlorpyrifos

degradation.

2.6.2.2 Fungal degradation of OPs

There are not many reports of OP degradation by fungal species as compared

to those by their bacterial counterparts. Furthermore the organophosphates

degradation rate by fungal isolates was found to be slower. Jones and Hastings (1981)

reported 95% to 98% degradation of 50 ppm chlorpyrifos by a group of forest fungi

namely Trichoderma harzianum, Penicillium vermiculatum, and Mucor sp. after 28

days of incubation along with accumulation of its metabolite TCP. Bumpus et al.

(1993) reported a fungal strain Phanerochaete chrysosporium able to mineralize only

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18

26.6% of added chlorpyrifos after 18 days of incubation. Bending et al. (2002)

reported Hypholoma fasciculare and Coriolus versicolor degraded chlorpyrifos in soil

bio-bed after 42 days. Studies had also reported the chlorpyrifos degradation in soil by

Aspergillus sp., Trichoderma sp. (Liu et al. 2003) and Fusarium sp. (Wang et al.

2005).

There are reports regarding fungal degradation of other OP compounds.

Mostafa et al. (1972) studied the transformation of malathion by Aspergillus niger,

Penicillium notatum and Rhizoctonia solani. The conversion of malathion to

malaoxon (AChE inhibitor) by Rhizoctonia solani was reported. However, A. niger

and P. notatum transformed malathion to malathion monoacid and malathion diacid.

Rao and Sethunathan (1974) isolated Penicillium waksmani from soil having the

ability to degrade large amounts of parathion to aminoparathion and two unidentified

polar metabolites. Hasan (1999) reported few fungal species having ability to degrade

various organophosphate compounds and reported Aspergillus sydowii capable of

using dimethoate as a sole source of phosphorus. Liu et al. 2001 isolated a fungal

strain, A. niger ZHY256 from sewage having the potential to hydrolyze P–S–C bonds

characteristics of dimethoate, malathion and formothion. Kim et al. (2005) observed

that a fungal cutinase produced by Fusarium oxysporum f. sp. pisi metabolize

malathion to its monacid and diacid derivatives.

Zboinska et al. (1992) reported fungal strain Penicillium citrinum capble of

metabolizing organophosphate pesticide glyphosate. Krzysko-Lupicka et al. 1997 also

reported five fungal strains namely Trichoderma viride, T. harzianum, Scopulariopsis

sp., Alternaria sp. and A. niger, isolated from soil, capable of utilizing glyphosate via

AMPA (amino methyl phosphonic acid) pathway as a source of phosphorous. Further,

Lipok et al. (2003) isolated four fungal strains Penicillium janthinellum, Penicillium

simplicissimum, Mucor sp. and Alternaria alternata from carrot seeds utilizing

glyphosate as a phosphorus source.

2.6.2.3 Algal degradation of OPs

There are only few algal species that have been reported for their ability to

degrade/bioaccumulate OPs as compared to bacterial and fungal isolates. Zuckerman

et al. (1970) isolated an alga, Chlorella pyrenoidosa having ability to metabolize

parathion to aminoparathion. Megharaj et al. (1987) reported degradation of

monocrotophos by algae, Chlorella vulgaris and Scenedesmus bijugatus.

Subramanian et al. (1994) reported two filamentous cyanobacteria, Aulosira

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19

fertilissima ARM 68 and Nostoc muscorum ARM 221 using malathion and

monocrotophos as a phosphorus source even in the presence of additional inorganic

phosphorus.

Cyanobacteria are known to absorb phosphorus in surplus than required and

also reported to absorb OPs as a phosphorous source (Stewart and Alexander, 1971).

Lal and Lal, (1987) also reported bioaccumulation of chlorpyrifos in blue-green algae.

Similarly, Mesocosm studies by Pablo et al. (2009) described that algae plays

important role in binding of chlorpyrifos along with the organic matters in the soil.

2.6.2.4 Degradation of OPs by microbial consortia

Most of the lab scale microcosm based in-situ bioremediation studies

involving addition of pure cultures to polluted soil are prone to problem because of

poor survival or low activity of these cultures in the natural environmental conditions.

Munnecke and Hsieh (1974) reported a mixed bacterial consortium consisting of

Pseudomonas sp., Xanthomonas sp., Azotomonas sp. and a Brevibacterium sp.

capable of hydrolysing 50 mg l-1 of

parathion. The biodegradation of pesticides, such

as 4-nitrophenol (Laha & Petrova 1998), endosulfan (Awasthi et al. 2000), 1,3-

dichloropropene (Ou et al. 2001) and diazonin degradation (Cycon et al. 2009) by a

microbial consortium has been reported.

The entire pesticide degradation pathways generally may not be present in

individual species. However, different components of microbial consortia can work in

concerted manner to acheive effeciant degradation of these compounds (Macek et al.

2000; Kuiper et al. 2004; Chaudhry et al. 2005). The rhizosphere soil contains 10–100

times more microbes than un-vegetated soil due to presence of plant exudates such as

sugars, organic acids, and larger organic compounds in the soil (Lynch 1990; Kumar

et al. 2006). However, there are certain factors those can interfere with the enrichment

process for development of degradative consortium including (a) the complex

molecular and structural features of the degrading compound that may limit its

degradability e.g. polyhalogenated compounds (Wackett et al. 1994), (b) natural

dominance of a non-productive metabolic pathway (Oh and Bartha 1997), (c) low

frequency of an essential degradative gene (Shapir et al. 1998), (d) poor

bioavailability e.g. polycyclic aromatic hydrocarbons (Bastiaens et al. 2000) and (c)

production of recalcitrant intermediates (Van Hylckama Vlieg and Janssen 2001). The

problem may be overcome by developing genetically engineered microbial strains or

by developing an efficient consortium from natural degraders. The metabolic

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20

synergism between different microbial species encourages the biodegradation of

recalcitrant molecules via aerobic and anaerobic reactions. Gilbert et al. 2003

developed a consortium comprised of two engineered strains, Escherichia coli SD2

with plasmids encoding a gene for parathion hydrolase and Pseudomonas putida

KT2440 having pSB337 plasmid contained a p-nitrophenol-inducible operon

encoding the genes for p-nitrophenol mineralization, to hydrolyze 500 µM of

organophosphate insecticide parathion without the accumulation of p-nitrophenol in

suspended culture.

Vidya Lakshmi et al. 2008 developed a microbial consortium consisting of

Pseudomonas fluorescence, Brucella melitensis, Bacillus subtilis, Bacillus cereus,

Klebsiella sp., Serratia marcescens and Pseudomonas aeruginosa supported 75–87%

degradation of chlorpyrifos after 20 days of incubation.

Immobilization of the pure cultures and consortium may help to improve

bioremediation potential as immobilized cells have prolonged microbial cell viability

ranging from weeks to months and improved capacity to tolerate higher

concentrations of pollutants (Richins et al. 2000; Chen and Georgiou 2002).

Karamanev et al. 1998 immobilized bacterial consortium in alginate beads and on

tezontle (a porous igneous rock) by biofilm for the removal of a pesticide mixture

form liquid medium composed of methyl-parathion and tetrachlorvinphos.

2.7 Factors affecting degradation of OPs

The most important parameters involved in pesticide biodegradation are

pesticide concentration, inoculum size, pH, temperature, and its bioavailability

(Karpouzas and Walker, 2000; Singh et al. 2006).

2.7.1 Effect of substrate concentration

The chlorpyrifos concentration found in the upper 10 mm of the soil sediment

after its application was observed to be the highest (Brock et al. 1992). Cink and

Coats (1993) observed that after the use of agricultural application rate of 10 mg kg-1

of chlorpyrifos 5% of chlorpyrifos persisted in the soil. However, at termite

infestation sites where chlorpyrifos might be used at higher level almost 57% of

appended chlorpyrifos remained in soil.

The ability of microbes to degrade a pollutant depends on the available

concentration of polluting chemicals, as high concentrations are usually toxic for

microbial degraders and low concentrations may not be able to induce the enzymes

involved in degradation (Block et al. 1993; Morra, 1996). Menon et al. (2005)

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21

reported delayed dehydrogenase activity in the soil after chlorpyrifos application at

0.20 µg g-1 indicated inhibition of microbial growth at the polluted site. A similar

observation was reported by Pandey and Singh (2004) where a dose of 4 L/hm2

chlorpyrifos showed a short-term inhibitory effect on the total microbial population.

Shan et al. (2006) also indicated that the application of chlorpyrifos lead to decrease

in bacterial, fungal and actinomycete populations with increasing chlorpyrifos

concentration (2, 4, and 10 mg kg-1) in the soil. Hua et al. (2009) reported that soil

ammended with chlorpyrifos at the initial level of 4, 8, and 12 mg kg-1, the

chlorpyrifos was degraded after 35 days with average half-live of 14.3, 16.7, and 18.0

respectively. The initial inhibition of soil microbial communities was followed by

recolonization of soil microbial communities after two weeks.

2.7.2 Effect of inoculum size

Ramadan et al. 1990 observed that at low inoculum levels,

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22

of chlorpyrifos degradation in alkaline soils was due to chemical hydrolysis. In

general, higher the pH higher is the rate of hydrolysis of OP pesticides) which may be

due to higher copy numbers of opd (organophosphate degrading) gene (Sparks, 1989;

Singh et al. 2003; Singh et al. 2003).

Wang et al. (2006) had also reported that chlorpyrifos degradation rate by B.

laterosporus DSP was increased with increase in pH from 7.0 to 9.0. Al-Qurainy and

Abdel-Megeed 2009 observed the effect of pH on two OP pesticides malathion and

dimethoate. They reported complete degradation of malathion and dimethoate by

Pseudomonas frederiksbergensis at pH 7.0 after 6 days of incubation with half lives

accounted by 3 and 2.3 days respectively. However, when the medium pH was set at

8.0, biodegradation began on the first day and complete degradation was observed

after 3 days. Wang et al. (2005) reported that the biodegradation rates of chlorpyrifos

in the pH range 6.5–9.0 by Fusarium LK. ex Fx. WZ-I were higher.

2.7.4 Effect of solubility/bioavailability

Many researchers have reported that high organic matter content of the soil

leads to a absorption of pesticide to soil particles resulting in lower bioavailability of

organophosphorus pesticides and hence decreases their degradation rate (Barriuso et

al. 1992; Weber and Huang, 1996; Karpouzas and Walker, 2000; Nelson et al. 2000;

Ben-Hur et al. 2003). Knuth and Heinis (1992) and Brock et al. (1992) also reported

high absorption of chlorpyrifos to sediments in static aquatic systems sediments due

to the hydrophobic nature of aquatic sediments and lower solubility of chlorpyrifos in

aqueous phase. Similarly, Civilini, 1994 reported that it is easy to remove more water

soluble lighter hydrophilic compounds than heavier hydro-phobic PAHs. In general

pesticides having low water solubility are less susceptible to accelerated degradation

due to their limited dissolution rates (Alexander, 1999).

The fate and behavior of pesticide in the environment is determined by its

solubility, half-life and partitioning coefficients (Neitsch et al. 2005). The

hydrophobic compounds become non-available to microbial degraders as these

compounds get entrapped in nanopores of the solid phase of organic matter (Arbeli

and Fuentes 2007). However, many researchers used chemical surfactant or

biosurfactants produced by microorganisms to desorb chemical compounds from soil

organic matter so as to make them bioavailable for their consumption (Aronstein et al.

1991; Brown and Jaffe, 2006; Zhou and Zhu, 2008).

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23

2.8 Biosurfactants

The biosurfactant molecules are composed of a hydrophobic and a hydrophilic

moiety. Hydrophobic end of the molecule is comprised of a long-chain fatty acids,

hydroxy fatty acids or α-alkyl-β-hydroxy fatty acids while the hydrophilic part can be

a carbohydrate phosphate, cyclic peptide, amino acid, carboxylic acid or alcohol.

Biosurfactants are broadly grouped as:

• Glycolipids (Rhamnolipids, Sphorolipids, Trehalolipids etc.)

• Lipopeptides (surfactin, viscosin, lichenysin etc.)

• Phospholipids

• Fatty acids/Neutral lipids (corynomicolic acids)

• Polymeric surfactants (emulsan, liposan, alsan etc.)

• Particulate compounds (vesicles, whole microbial cells)

Most of biosurfactant are either anionic or neutral while few of these

containing amine groups are cationic. The surfactant-producing microorganisms

include:

Table 2.5: Different types of biosurfactants produced by microorganisms

Types of Biosurfactants Microorganisms

Glycolipids Pseudomonas aeruginosa, , Candida tropicalis

Phospholipids Corynebacterium, Nocardia and Rhodococcus spp.

Lipopeptides Bacillus licheniformis

Lipopolysacchrides Acinitobacter calcoaceticus

Sophorolipids Torulopsis spp.

Corynemycolic acids Corynebacterium spp.

Trehalose and sucrose lipids Arthrobacter paraffineus

Particulate biosurfactants Arthrobacter radioresistens

Jarvis and Johnson (1949) reported that two L-rhamnose molecules linked to a

chain of β-hydroxydecanoyl β -hydroxydecanoate were produced from Pseudomonas

aeruginosa. They were classified in two major groups: the monorhamnolipids, which

contain one unit of rhamnose and two of β-hydroxydecanoic acid (Rha-C10-C10) and

the di-rhamnolipids, which contain two units of rhamnose linked to two units of

hydroxydecanoic acid (Rha-Rha-C10-C10). The rhamnolipid biosynthesis is catalyzed

by two rhamnosyl-transfer reactions catalyzed by rhamnosyl-transferase Rt1 and Rt2

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24

where β-hydroxydecanoyl β–hydroxydecanoate or mono-rhamnolipid act as a

rhamnosyl recipient while deoxythymidine diphosphate (dTDP)-L-rhamnose acts as

the rhamnosyl donor (Burger et al. 1966). A total of 28 different homologues, of

rhamnolipid with acyl chains varying from C8–C14 and branched sugar moieties have

been reported till date (Mulligan, 2005 and Sober´on-Chavez et al. 2005). Bacterial

cultures mostly Pseudomonas sp. are known to produce glycolipids including

rhamnolipids that are involved in degradation of polyaromatic hydrocarbons (Arino et

al. 1996). The rhamnolipid biosurfactants are extensively studied for solubilization

and bioremediation of many hydrophobic environmentally toxic compounds

(Monteiro et al. 2007).

2.8.1 Methods for screening of biosurfactant producers

Due to diversity of surface active molecules produced by microbes a range of

screening methods have been developed including:

• Blood cell lysis agar method first time reported by Bernheimer and Avigad

1970 for screening of Surfactin produced by Bacillus subtilis. The effective

biosurfactant production leads to haemolysis of blood cells forming a clear

zone around the microbial colony (Carillo et al. 1996). However its not so

reliable method as certain other haemolysin microbial products may also

disrupt the cell membrane (Seigmund and Wagner 1991).

• Cetyltrimethylammonium bromide-methylene blue (CTAB-MB) plate assay is

specific only for detection of anionic biosurfactants e.g. rhamnolipid,

cellobioselipid, sophorolipids etc. The cultures producing anionic surfactant

form insoluble ionic pairs with cationic CTAB to give a dark blue halo around

their colony (Seigmund and Wagner 1991).

• Surface activity measurement of biosurfactant by surface tension/interfacial

tension reduction of growth medium by the Ring method (Magaritis et al.

1979; Persson and Molin 1987). It is the most reliable and standard method to

detect the biosurfactant production by the microbes (Willumsen and Karlson

1997; Makkar and Cameotra 1999; Joshi et al. 2008).

• Drop collapse method where a drop of liquid containing biosurfactant will

spread over the surface of oil due to reduction in its surface tension and ability

to bridge between water and oil molecules due to the amphipathic nature of

biosurfactant (Bodour and Maier 1998; Morikawa et al. 2000).

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25

• Acid precipitation method can be used for the separation of biosurfactant

produced in the growth medium by lowering its pH as biosurfactants have the

property to precipitate at or below pH 2.0 (Mukherjee et al. 2009).

• Thin layer chromatography technique involves either direct application of

bacterial biomass on to the TLC plate to develop with suitable solvent system

(Matsuyama et al. 1991) or extraction of the growth medium with solvents and

then spotted on to the TLC plate to develop e.g. by Zhang and Miller 1997 for

detection of mono and di-rhamnolipids.

Effective surfactants have low CMC or critical micellar concentration values. The

CMC is the minimum amount of a surfactant required to induce micelles formation.

The CMCs of the biosurfactants range from 1 to 200 mg l-1 (Lang and Wagner, 1987).

Surfactants those can lower the surface tension of water from 72 mN m-1 to 35 mN m

-

1 are considered as good surfactants (Mulligan 2005). The rhamnolipid surfactant

produced by P. aeruginosa can lower the surface tension of water to 26 mN m-1

(Syldatk et al. 1985). Thus, biosurfactants at low concentrations can be used for

mobilization of soil adsorbed hydrophobic contaminants (Tsomides et al. 1995).

Further the emulsification potential of a surfactant depends on its hydrophilic and

lipophilic balance (HLB) which directly related to the length of the hydrocarbon chain

(Schramm et al. 2003). Surfactants with HLB 3 or less have hydrophobic property

while those have HLB value more than 11 are hydrophilic and can used for

solubilizing the hydrophobic compounds (Sabatini et al. 1995).

Microbially produced surfactants have the ability to modulate the cell surface

properties of their producers which regulate the attachment and detachment of

microbial cells from a surface (Rosenberg 1993). As the biosurfactant produced by

Acinetobacter sp. reduces its cell surface hydrophobicity whereas, rhamnolipid

increase the hydrophobicity of P.areuginosa cell surface that may allow uptake of

hydrophobic pollutants by microbial cells (Rosenberg and Rosenberg 1983; Zhang

and Miller 1994).

Pseudomonas aeruginosa can produce rhamnolipids from wide range of

substrates such as alkanes, pyruvate, fructose, succinate, citrate, glycerol, mannitol,

glucose and olive oil (Robert et al. 1989). Yield of the biosurfactant also depend on

the fermentor design, nutrients and physico-chemical conditions used (Mulligan and

Gibbs, 1993). The high production costs for biosurfactants currently prohibits their

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26

large-scale utilization, which can be reduced by exploring biosurfactant producing

strains capable of using a wide range of cheap substrates. Patel and Desai 1997

reported that Pseudomonas aeruginosa GS3 produced rhamnolipid biosurfactant with

0.25 g l-1 of rhamnose concentration using 7% (w/v) molasses and 0.5% (w/v)

cornsteep liquor as the primary carbon and nitrogen source. Similarly, Haba et al.

(2000) reported production of 8.0 g l-1 of rhamnolipid biosurfactant from 20 g l

-1 of

canola oil refinery waste by Pseudomonas sp.

2.9 Role of surfactants in solubility and bioremediation of pesticides

Surfactants are amphiphilic molecules consisting of a hydrophilic tail and a

hydrophobic head (Banat et al. 2000). Thus, surfactants at concentrations above the

CMC may increase the solubility of organic pollutants by its partitioning at the

hydrophobic core of the surfactant micelles (Di Cesare and Smith, 1994). Surfactants

are of main point of interest to researchers in recent years for bioremediation as they

can enhance the solubility and bioavailability of non-aqueous phase soluble

hydrophobic compounds in aqueous phase (Brown and Jaffe, 2006; Zhou and Zhu,

2008).

Surfactant can enhance the process of microbial degradation of HOCs by three

ways i) the surfactants addition may alter the hydrophobicity of cell which allows the

direct contact of the microbial cells and the pollutant molecules (Tang et al. 1998). ii)

The microbial cell membrane fuse with the micelles formed by the surfactants

containing HOC molecules (Schippers et al. 2000). iii) The bacteria can directly use

the surfactant solubilized HOCs from solid phase of the soil into the aqueous phase

(Kim et al. 2001).

2.9.1 Chemical surfactants

The ability of the surfactants to increase desorption of HOCs from the soil

particles and increase their apparent aqueous solubility resulting in improved

bioavailability to microbes for their bioremediation has been reported (Lopes et al.

1995; Mata-Sandoval et al. 2001). Both anionic and nonionic surfactants are used for

bioremediation of land polluted with oils and hydrocarbons (Haigh 1996). It was

observed that surfactant enhanced solubilization enhances biodegradation of

polyaromatic hydrocarbons and phenanthrene (Boonchan et al. 1998; Guha and Jaffe

1996). Microbial cells membrane up to a certain extent fuse with the micelles to take

up the pollutant (Miller and Bartha 1989; Edwards et al. 1991). Usually synthetic

surfactants are used as mixtures as they act better in mixtures than individual

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27

components (Bruheim et al. 1999). Chiou 2002 observed that aqueous phase partition

of pesticides can be enhanced by surfactant polyethylene lauryl ether (C12E10) as

observed by fluorescence anisotropy method. Further it was observed that surfactants

facilitate the uptake of HOCs by plants as it increases their aqueous phase solubility

(Li et al. 2001; Gao et al. 2004; Gao et al. 2006).

Although use of chemical surfactants for bioremediation purposes is well

known but there are certain drawbacks for their use. Chemical surfactants are less

biodegradable, toxic for indigenous microbial populations of soil and usually their

high concentrations are required to mobilize non-aqueous phase soluble compounds.

Pinto and Moore 2000 in soil slurries studies observed that 156 g l-1 of Tween 80

(10,000 times the CMC) was required to mobilize 70% of high molecular weight

polyaromatic hydrocarbons from contaminated soil. So microbially produced

surfactants produced by diverse range of prokaryotic and eukaryotic microorganisms

can be used as a safe and effective alternative (Van-Dyke et al. 1993; Mata-Sandoval

et al. 2001).

2.9.2 Biosurfactants

The applications of biosurfactant have been picking the pace in their use in

pollution removal including pesticides (Banat et al. 2000), oil (Ron and Rosenberg

2002) and polyaromatic hydrocarbons (Cameotra and Bollag 2003) by increasing their

solubility and hence availability towards microbial degradation.

Van Dyke 1993 isolated Pseudomonas aeruginosa having ability to produce

rhamnolipid biosurfactants with ability to remove 25-70 % hydrocarbons from sandy-

loam soil and 40-80 % from silt-loam soil when used at the concentration of 5 g l-1.

There are many reports in the literature regarding the use of biosurfactant in enhanced

biodegradation of polyaromatic hydrocarbons (Tiehm 1994; Churchill 1995). Kim et

al. (2001) also showed that with addition of nonionic surfactants biodegradation of

PAHs could be enhanced.

Microbial surfactants are more useful than synthetic ones because of their low

toxicity and high biodegradability (Zajic et al. 1997). Kanga et al. 1997 isolated

Rhodococcus species 413A capable of producing a glycolipid which was observed to

be 50% less toxic than a synthetic surfactant Tween 80 in naphthalene solubilization

tests. A biosurfactant produced by P. aeruginosa was compared with Marlon A-350 a

synthetic surfactant widely used in the industry and it was observed that biosurfactant

was comparatively less toxic and mutagenic (Flasz et al. 1998). Further there is less

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28

need for product purity and in-situ production of microbial surfactant by indigenous

or introduced microbial populations is also possible (Ivshina et al. 1998; 2001).

Awasthi et al. 1999 showed enhanced microbial degradation of endosulfan a

hydrophobic organic compound by 30–45% in presence of biosurfactant produced by

Bacillus subtilis MTCC1427. Mata-Sandoval et al. (2001) studied the biodegradation

of the three pesticides trifluralin, atrazine and coumaphos in aqueous phase in

presence of rhamnolipid and Triton X-100. The biodegradation of atrazine decreased

in presence of both the surfactants but trifluralin biodegradation was enhanced while

coumaphos biodegradation increased in presence of rhamnolipid but declined when

Triton X-100 concentration was used above its CMC. In soil slurries with increase in

concentration of rhamnolipid removal of coumaphos increased. Conte et al. (2005)

observed that humic acid solution can be used as a natural surfactant for 90%

desorption of polyaromatic hydrocarbons and thiophenes from soils which was

comparable to that of the synthetic surfactants (SDS and Triton X-100).

The use of biosurfactants to improve bioavailability of toxicants in soils and

other environments is an attractive option because of their biodegradability (Herman

et al. 1995). Makkar and Cameotra 1997 studied the effect of surfactin produced by B.

subtilis on biodegradation of the hydrophobic pesticide endosulfan. Zhang et al. 1997

studied the effect of two different types of rhamnolipid biosurfactants on the

dissolution, bioavailability and biodegradation of phenanthrene. It was observed that

with addition of biosurfactants, solubility and hence degradation of phenanthrene was

increased as compared to the control without biosurfactants. Page et al. (1999)

reported that the mass transfer of PAHs into the aqueous phase was increased up to

35-fold more effectively by biosurfactant produced by Rhodococcus strain H13-A

than the synthetic surfactant Tween 80. Robinson et al. 1996 used 4 g l-1 Rhamnolipid

R1 biosurfactant produced by P. aeruginosa to mineralized 4,4- chlorobiphenyl and

observed that there was 213 times more mineralization of the added polychlorinated

biphenyls as compared to the control without biosurfactant. Similarly Fiebig et al.

1997 has shown that in the presence of a glycolipids (GL-K12) biosurfactant from

Pseudomonas cepacia enhanced degradation of Arochlor 1242 by mixed cultures.

Warranaphon et al. 2008 isolated a biosurfactant-producing bacterium Burkholderia

cenocepacia BSP3 with high CMC value of 316 mg l-1 but very low surface tension

(25 mN M-1) and have the ability to emulsify methyl parathion, ethyl parathion and

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29

trifluralin hence having the potential to use for bioremediation of pesticide-

contaminated soil.

There are many additional potential applications of biosurfactant includes

cosmetic and soap formulations, foods, and dermal/transdermal drug delivery systems

(Itoh 1987; Brown 1991). Studies revealed that many biosurfactants can be applied

for accelerated biodegradation of hydrocarbons in soils contaminated by oil spillage

near beaches, enhanced oil recovery, crude oil drilling lubricants, food processing

industry and health care (Harvey et al. 1990; Fiechter 1992; Providenti et al. 1995;

Desai and Banat 1997; Cameotra and Makkar 1998).

On the contrary there are reports (Hisatsuka et al. 1971) that rhamnolipid

produced by Pseudomonas aeruginosa failed to stimulate degradation of hydrophobic

compounds by other strains or by mixed cultures which are not known to produce

biosurfactants. Similarly, degradation of hexadecane by rhamnolipid-producing

organisms is stimulated to a greater extent by rhamnolipid rather than by any other

biosurfactant (Itoh et al. 1972). The surfactants at concentrations above the CMC

inhibit adhesion of bacteria to the surfaces of droplets of liquid hydrocarbons and thus

inhibit biodegradation (Ortega-Calvo and Alexander. 1994). Thus there is need to

optimize the effective concentration of biosurfactant in bioremediation protocols.

2.10 Molecular basis of organophosphate pesticide degradation

Two enzymes namely phosphotriesterase also known as organophosphate

hydrolase (OPH) and organophosphorus acid anhydrolase (OPAA) capable of

degrading organophosphorus pesticides encoded by opd and opaA genes respectively.

The enzyme organophosphate hydrolase (OPH) is extensively studied enzyme due to

its ability to degrade a wide range of OP compounds.

2.10.1 Phosphotriesterase

Phosphotriesterase from Pseudomonas diminuta is an efficient metalloenzyme

that hydrolyses a variety of organophosphorus nerve agents. The phosphotriesterase

was first identified in Flavobacterium sp. from Philippine rice patty samples

(Sethunathan and Yoshida 1973). Munnecke 1976 isolated strain P. diminuta through

its ability to hydrolyze parathion having enzyme phosphotriesterase. In both cases

organophosphate-degrading genes (opd) encoding the active enzymes were localized

on extra chromosomal plasmids. The opd (organophosphate degrading) gene encoding

OPH was first isolated from P. diminuta and was reported to be present on a 66 kb

plasmid, pCMS1 (Serdar et al. 1982). Serdar (1989) cloned the opd gene from

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30

Pseudomonas sp. into Escherichia coli. Similarly the gene has been subcloned into

Streptomyces sp. (Steiert 1989) and insect cells (Dumas 1990). This strategy led to a

variant that maintains the specific activities of wild-type phosphotriesterase, but

expresses functionally at higher levels to accquire better degradation efficiency.

Phosphotriesterase has a wide range of substrate specificities with ability to

hydrolyze P–O, P–S, P–F and P–CN bonds with highest activity against the P–O

linkage and least specificity for the P–S bond (Efremenko and Sergeeva, 2001). The

phosphotriesterase from Pseudomonas diminuta is a highly efficient zinc

metalloenzyme carrying out hydrolysis of variety of organophosphorus nerve agents

(Donarski et al. 1989). Two metal ions are vital for maximal catalytic activity of the

enzyme (Dumas et al. 1989; Omburo et al. 1992) which was shown by X-ray

crystallography to have a two binuclear metal center embedded within a cluster of

histidine residues (Vanhooke et al. 1996; Benning et al. 2001).

Omburo et al. 1992 also isolated an opd gene encoding a 40 kDa homodimer

parathion hydrolase, which contains divalent zinc ions as a cofactor. Horne et al.

(2002) also suggest that phosphotrieseterase is a 384-amino-acid protein with a

molecular mass of approximately 35 kDa when it is cleaved from its signal peptide.

The two native Zn2+ ions of this enzyme can be substituted with either Co

2+, Ni

2+,

Cd2+, or Mn

2+ with/without the restoration of catalytic activity. Recent findings have

shown that two metal atoms are closely associated and the water molecule that attacks

the phosphoryl center is bound directly to the binuclear metal center (Benning et al.

1995; Vanhooke et al. 1996).

2.10.2 Organophosphorus acid anhydrolase (OPAA)

The natural function of the OPAA is not known but has been proposed that it

is a dipeptidase that catalyses dipeptide with a proline residue at the C-terminus

(Cheng et al. 1996). The gene (opaA) was isolated from Alteromonas sp. JD6.5

encodes OPAA a single peptide with molecular weight of 60 kDa (Cheng et al. 1996).

There was no sequence similarity between opd and opaA but functionally they show

similarity to act against OPs.

OPAA was mostly isolated from strains of Alteromonas with higher levels of

activity and a broad range of substrate specificity against OPs preferably against sarin

and soman (Cheng et al. 1993; Hill et al. 2000). However, it does not have any

activity against P-S bond. OPAA has lower efficiency against paraoxon as comparied

Review

31

to OPH. Maximum activity of OPAA was reported in the presence of Mn2+ and Co

2+

(DeFrank and White, 2002).

There are restriction governing uses of genetically modified organisms for “in-

situ” bioremediations protocols. Thus it is important to understsnd the conditions

which can keep in optimum expression of enzyme in microbial isolates resulting in

efficient degradation of target molecules.