bioremediation of organophosphate contaminated

BIOREMEDIATION OF ORGANOPHOSPHATE

CONTAMINATED AGRICULTURAL SOILS

USING INDIGENOUS BACTERIA

By

Samina Ambreen

Fatima Jinnah Women University

Rawalpindi, Pakistan

2020

BIOREMEDIATION OF ORGANOPHOSPHATE

CONTAMINATED AGRICULTURAL SOILS USING

INDIGENOUS BACTERIA

By

Samina Ambreen

Registration No: 2014-Ph.D-Env.Sci-004

A dissertation submitted to the

Fatima Jinnah Women University

Rawalpindi.

In partial fulfilment of the requirement for THE DEGREE OF DOCTOR OF PHILOSOPHY IN

ENVIRONMENTAL SCIENCES

August 2020

Author’s Declaration

I Samina Ambreen, hereby state that my PhD thesis titled “Bioremediation of

Organophosphate contaminated agricultural soils using indigenous bacteria” is

my own work and has not been submitted previously by me for taking any degree

from Fatima Jinnah Women University, Rawalpindi, or anywhere else in the

country/world. At any time, if my statement is found to be incorrect even after my

graduate, the university has the right to withdraw my Ph.D degree.

Samina Ambreen

Reg. No: 2014-Ph.D-Env.Sci-004

Plagiarism Undertaking

I solemnly declare that research work presented in the thesis entitled

“Bioremediation of Organophosphate contaminated agricultural soils using

indigenous bacteria” is solely my research work with no significant contribution

from any other person. Small contribution/help wherever taken has been duly

acknowledged and that complete thesis has been written by me. I understand the zero

tolerance policy of the HEC and Fatima Jinnah Women University, Rawalpindi

towards plagiarism. Therefore, I as an Author of the above titled thesis declare that no

portion of my thesis has been plagiarized and any material used as reference is

properly referred/ cited.

I undertake that if I am found guilty of any formal plagiarism in the above titled thesis

at any time even after award of Ph.D degree, the university reserves the rights to

withdraw/revoke my Ph.D degree and that HEC and the university has the right to

publish my name on the HEC/university website on which names of students are

placed who submitted plagiarized thesis.

Student/Author Signature: _________________

Name: Samina Ambreen

CERTIFICATE

It is certified that this Ph.D dissertation entitled “Bioremediation of

Organophosphate contaminated agricultural soils using indigenous bacteria”

submitted by Samina Ambreen, Reg No: 2014-Ph.D-Env.Sci-004 is an original work

and has not been presented for a degree wholly or partially in any other university. All

sources, references and literature used or excerpted during elaboration of this work

are properly cited and mentioned the due source.

Prof. Dr. Azra Yasmin

Supervisor

Table of Contents

Contents Page No

List of Abbreviations and Acronyms i

List of Figures v

List of Tables xvi

Acknowledgements xix

Summary xx

Chapter 1: Introduction 1

1.1 Pesticides in Pakistan 3

1.2 Organophosphate pesticides 4

1.2.1 The toxicity of OP compounds and inhibition of Acetyl

Cholinestrase (AChE) 7

1.3 Chlorpyrifos (CPF) 9

1.3.1 Mode of Action 10

1.3.2 Acute toxicity of CPF to non-target organisms 10

1.3.3 Microbial Degradation of CPF 12

1.3.4 Metabolic products of CPF 13

1.4 Triazophos (TAP) 14

1.4.1 Biodegradation of TAP 15

1.4.2 Metabolites of Triazophos 16

1.5 Dimethoate (DM) 16

1.5.1 Toxicological effects of Dimethoate 17

1.5.2 Environmental fate of Dimethoate 18

1.5.3 Microbial degradation of DM 18

1.5.4 Metabolites of Dimethoate 20

1.6 Microbial organophosphate degrading enzymes 20

1.7 Modern Analytical methods for the analyses of Pesticides 22

1.8 Aims and Objectives 22

Chapter 2: Materials and Methods

2.1 Preparation of solutions and media

24

24

2.2 Sampling Site and Soil Sampling 32

2.3 Physicochemical Analysis of Soil 32

2.3.1 Measurement of the soil pH 32

2.3.2 Electrical Conductivity (EC) 32

2.3.3 Soil organic matter 33

2.3.4 Soil texture 33

2.3.5 Soil moisture Content 34

2.4 Isolation of bacteria from the soil samples 34

2.4.1 Colony Counting (CFU/ml) 35

2.4.2 Single colony streaking (bacterial culture purification) 35

2.5 Screening for OP pesticide tolerance 35

2.6 Screening for 3, 5, 6-Trichloropyridinol (TCP) tolerance 35

2.7 Morphological characterization of OP pesticide degrading

bacterial isolates

36

2.7.1 Colony morphology 36

2.7.2 Cell morphology 36

2.7.2.1 Gram’s staining 36

2.7.2.2 Capsule staining 37

2.7.2.3 Spore staining 37

2.7.2.4 Motility test 38

2.8 Biochemical characterization 38

2.8.1 Oxidation Fermentation Test 38

2.8.2 Oxidase Test 39

2.8.3 Catalase Test 40

2.8.4 Other biochemical/enzyme Tests 40

2.9 Physiological Characterization 41

2.9.1 pH Optimization for Bacterial Growth 41

2.9.2 Effect of temperature on the growth of bacterial isolates 41

2.10 Heavy Metal Resistance Profile 42

2.11 Tolerance against different organic pollutants 43

2.12 Bacterial Inoculum and Consortium Preparation 43

2.13 Analysis of OP pesticide degradation using UV-VIS

Spectrophotometer

44

2.14 Qualitative and Quantitative Analysis for OP pesticide

degradation using GC-MS and HPLC

44

2.14.1 Extraction of pesticide residues from M-9 Culture Broth 44

2.14.2 Extraction of Trichloropyridinol (TCP) residues from M-

9 Culture Broth

45

2.14.3 Extraction from soil slurry for OP pesticide degradation

analysis

46

2.14.4 Extraction from soil microcosm for OP pesticide

degradation analysis

46

2.14.5 GC-MS conditions used for the analyses of CPF, TCP,

TAP and DM biotransformation

47

2.14.6 HPLC conditions used for the analyses of CPF, TCP,

TAP and DM degradation

47

2.15 Optimization for OP pesticide biodegradation 49

2.15.1 pH Optimization for OP pesticide biodegradation 49

2.15.2 Temperature Optimization for OP pesticide

biodegradation

50

2.15.3 Effect of Shaking and Static conditions for OP pesticide

biodegradation

50

2.16 Molecular Studies 50

2.16.1 DNA Extraction 50

2.16.2 Polymerase Chain Reaction (PCR) 51

2.16.3 Gel Electrophoresis 51

2.16.4. Reagents for DNA Extraction 52

2.17 Enzyme studies for Organophosphorus phosphatase (OPP)

Enzyme

53

2.17.1 Screening for phosphate Solubilization potential of

bacterial isolates

53

2.17.2 Screening of Extracellular Organophosphorus

phosphatase (OPP) and Enzyme Assay

54

2.17.3 Screening of intracellular Organophosphorus

phosphatase (OPP)

55

2.17.4 Factors effecting the production of Organophosphorus

phosphatase (OPP) Enzyme

55

2.17.4a Effect of pH on OPP production 55

2.17.4b Effect of temperature on Organophosphorus

phosphatase (OPP) production

56

2.17.4c Effect of incubation time on Organophosphorus

phosphatase (OPP) enzyme production

56

2.17.5 Factors affecting Organophosphorus phosphatase

enzyme Activity

56

2.17.5a Effect of temperature on Organophosphorus

phosphatase enzyme Activity

56

2.17.5b Effect of chemicals on Organophosphorus


56

2.17.5c Effect of metals on Organophosphorus phosphatase

enzyme Activity

57

2.17.5d Effect of Substrate concentration on

Organophosphorus phosphatase enzyme Activity

57

2.17.5e Effect of incubation period on Organophosphorus


57

2.17.6 Metal bioprecipitation by Organophosphorus phosphatase

enzyme

57

2.17.7 Substrate specificity determination of Organophosphorus

phosphatase enzyme against OP pesticides (CPF, TAP and DM)

58

RESULTS Chapter 3: Isolation, characterization and screening

of organophosphate degrading soil bacterial isolates

59

3.1 Soil Sampling and Physical and Chemical properties of Soil 61

3.2 Isolation and purification of soil bacteria 62

3.3 Screening experiments against OP pesticides (Chlorpyrifos,

Triazophos and Dimethoate

62

3.3.1 Screening against Chlorpyrifos 63

3.3.2 Screening against Triazophos 63

3.3.3 Screening against Dimethoate 63

3.2.4 Screening experiments with Trichloropyridinol (TCP)

(Major metabolite of CPF)

64

3.2 Morphological characterization of OP degrading bacterial

isolates

65

3.3 Biochemical characterization 66

3.4 Physiological characterization 68

3.4.1 Screening Experiments against Heavy metals 68

3.4.2 Screening Experiments against other Organic Pollutants 68

3.4.3 Effect of pH on the growth of bacterial isolates 68

3.4.4 Temperature optimization for the growth of bacterial

isolates

69

3.5 Quantitative analysis of Chlorpyrifos degradation by bacterial

isolates

70

3.6 Genetic Analyses 73

3.6.1 Extraction of genomic DNA 73

3.6.2 PCR 73

3.6.3 Bacterial strain Identification 73

Chapter 4: Bacterial degradation/transformation of

Chlorpyrifos and its metabolite 3, 5, 6-Trichloropyridinol

76

4.1 Optimization of environmental parameters (pH, temperature,

shaking and static incubation conditions) for CPF degradation

78

4.1.1 Growth pattern and optimization for CPF degradation at

different pHs by bacterial isolates and their consortia

78

4.1.2 Effect of temperature on growth and CPF degradation by

bacterial isolates and their consortia

80

4.1.3 Effect of shaking versus static conditions on growth and

CPF degradation by four isolates (MB490, MB497, MB498 and

MB504) and seven consortia (A, B, C, D, E, F, and G) after 24hrs of

incubation

82

4.1.4 Biotransformation and degradation of Chlorpyrifos in the

presence of TAP

83

4.2 Degradation of Chlorpyrifos by the bacterial isolates and their

consortia in M-9 broth

85

4.3 Degradation of Chlorpyrifos (CPF) by the bacterial isolates and

their consortia in the soil slurry

88

4.4 Degradation of Chlorpyrifos (CPF) by the 4 isolates and the 7

consortia in the soil microcosm

91

4.5 Degradation and transformation experiments with 3, 5, 6-

trichloropyridinol (TCP)

94

4.5.1 Effect of incubation period and concentration of TCP on

the growth (OD600) and degradation of Trichloropyridinol (TCP)

by the 4 isolates in the bacterial culture broth

94

4.6 Quantitative analysis through HPLC to determine

biodegradation of Chlorpyrifos (CPF) and its metabolite 3, 5, 6-

Trichloropyridinol (TCP)

97

4.6.1 Effect of pH on CPF degradation by isolates and

consortia

97-108

4.6.2 Effect of temperature on CPF degradation by isolates and

consortia

108-119

4.6.3 Degradation of 3, 5, 6-Trichloropyridinol (14mg/l) by

four isolates after 72hrs incubation

119

4.7 Qualitative analysis through GCMS to detect metabolites of

Chlorpyrifos (CPF) by the 4 isolates and their consortia in the M-9

broth, soil slurry and soil microcosm

121

4.8 Biotransformation of 3, 5, 6 Trichloropyridinol (TCP) by the 4

bacterial isolates in M-9 broth as analyzed by GCMS

128

4.9 Proposed pathway for 3, 5, 6-Trichloropyridinol (TCP)

degradation by bacterial isolates (MB490, MB497, MB498 and

MB504)

131

4.10 Proposed metabolic pathway of Chlorpyrifos degradation by

bacterial isolates (MB490, MB497, MB498 and MB504) and their

consortia A, B, C, D, E, F and G

132

Chapter 5: Bacterial degradation and transformation of

Triazophos (TAP)

135

5.1 Optimization of environmental parameters for TAP

biodegradation

137

5.2 Bacterial growth/ degradation of TAP in the presence of

Chlorpyrifos

140

5.3 Effect of incubation period on bacterial growth and %

degradation of TAP in M-9 medium

141

5.4 Degradation of Triazophos by the bacterial isolates and their

consortia in soil slurry

143

5.5 Degradation of Triazophos by bacterial strains and their

consortia in the soil microcosm

145

5.6 Quantitative analysis through HPLC for biodegradation of TAP 147

5.6.1 Effect of pH on TAP degradation 147-158

5.6.2 Effect of temperature on TAP degradation 158-169

5.7 Qualitative analysis through GCMS to detect metabolites of OP

Pesticide Triazophos (TAP) by bacterial isolates and their consortia

in M-9 broth, soil slurry and soil microcosm

169

5.8 Proposed Metabolic Pathway for the biotransformation of TAP

by bacterial isolates MB490, MB497, MB498 and MB504

174

Chapter 6:

Biodegradation and biotransformation of OP pesticide

Dimethoate (DM)

178

6.1 Optimization of environmental conditions for Dimethoate

biodegradation

179


degradation of DM in M-9 broth

183


degradation of DM in the soil slurry

186


degradation of DM in the soil microcosm

187

6.5 Quantitative analysis through HPLC for Dimethoate

biodegradation

189

6.5.1 Effect of pH on DM degradation 189-193

6.5.2 Effect of temperature on DM degradation

6.5.3 DM degradation in soil microcosm by four isolate after 9

days of incubation

193-197

197


Dimethoate (DM) by the 4 isolates and their consortia in the M-9


199

6.7 Proposed Metabolic Pathway for the Transformation of

Dimethoate by Bacteria

204

Chapter 7: Organophosphorus phosphatase (OPP) enzyme

studies

207

7.1 Screening for phosphate solubilization potential of bacterial

isolates

209

7.2 Screening of extracellular Organophosphorus phosphatase (OPP)

production in bacterial isolates (MB490, MB497, MB498 and

MB504)

210

7.3 Screening of intracellular Organophosphorus phosphatase (OPP)

in bacterial isolates (MB490, MB497, MB498 and MB504)

210

7.4 Factors affecting the production of extracellular

Organophosphorus Phosphatase (OPP)

7.4a Effect of pH on extracellular OPP production by bacterial

isolates (MB490, MB497, MB498 and MB504)

211

211

7.4b Effect of temperature on extracellular OPP production by

bacterial isolates (MB490, MB497, MB498 and MB504)

212

7.4c Effect of incubation time on OPP production by bacterial


213

7.5 Factors affecting Organophosphorus Phosphatase (OPP) Activity 214

7.5a Effect of temperature on OPP activity and stability 215

7.5b Effect of substrate (p-NPP) concentration on OPP activity 216

7.5c Effect of incubation time on OPP activity 218

7.6 Study of stimulatory or inhibitory effect of chemicals (SDS,

EDTA, metals) on OPP Activity

220

7.6.1 Effect of Sodium dodecyl sulphate (SDS) on OPP

Activity

220

7.6.2 Effect of EDTA on OPP Activity 223

7.6.3 Effect of metals on OPP Activity 225

7.7 Metal bioprecipitation by Organophosphorus Phosphatase (OPP) 227

7.7.1 Bioprecipitation of Nickel (Ni++) by OPP 228

7.7.2 Bioprecipitation of Manganese (Mn++) by OPP 229

7.7.3 Bioprecipitation of Chromium (Cr++) by OPP 231

7.7.4 Bioprecipitation of Cadmium (Cd++) by OPP 232

7.8 Substrate specificity of acidic and alkaline OPP for CPF, TAP

and DM

234

Chapter 8: Discussion 240

i

List of Abbreviations and Acronyms

% Percent

°C Degree centigrade

DDT Dichlorodiphenyltrichloroethane

WHO World Health Organization

ACh Acetylcholine

AChE Acetylcholinesterase

DDT Dichlorodiphenyltrichloroethane

HCH Hexachlorocyclohexane

OP Organophosphate

WHO World Health Organization

USA United States of America

ADHD Attention deficit hyperactivity disorder

LD Lethal Dose

LC Lethal Concentration

UV Ultra Violet

CPF Chlorpyrifos

TAP Triazophos

DM Dimethoate

Hg Mercury

mg/ml Milligram/milliliter

g/cm3 Gram/cubic centimeter

DNA Deoxyribose Nucleic Acid

rRNA Ribosomal ribose nucleic Acid

DETP Diethyl thiophosphate

TCP 3, 5, 6-trichloropyridinol

mg/l Milligram/liter

P Phosphorus

O Oxygen

mg/kg Milligram/kilogram

ii

USDA United States Department of Agriculture

OPAA Organophosphate acid anhydrolase

OPH Organophosphate hydrolase

MPH methyl parathion hydrolase

OpdA Organophosphate degrading enzyme extracted from

Agrobacterium radiobacter

ADPase Aryldialkylphosphatase

OPP Organophosphorus Phosphatase

A-OPH Aspergillus derived Organophosphate hydrolase

P-OPH Penicillium derived Organophosphate hydrolase

PTE Phosphotriestrases

PLL Phosphotriesterase‐Like‐Lactonase

MS Mass Spectrometry

GCMS Gas Chromatography Mass Spectrometry

HPLC High Performance Liquid Chromatography

M-9 Minimal salts medium

EC of pesticide Emulsifiable concentrate of pesticide

EC of soil Electrical conductivity of soil

Na2HPO4 Di Sodium Hydrogen Phosphate

lb/inch2 Pounds per square inch

μl Microliter

µg Microgram

µg ml-1 Microgram per milliliter

rpm Revolutions per minute

pH Potential Hydrogen

HCl Hydrochloric Acid

Ni Nickel

Cr Chromium

Mn Manganese

Fe Iron

Cd Cadmium

iii

Cl Chlorine

Cu Copper

S Sulphur

Zn Zinc

Pb Lead

Co Cobalt

CFU Colony forming units

H2O2 Hydrogen peroxide

API Analytical Profile Index

LDC Lysine decarboxylase

H2S hydrogen sulfide production

ADH Arginine dihydrolase

ODC Ornithine decarboxylase

URE Urea hydrolysis

TDA Tryptophan deaminase test

VP Voges Proskauer

EM Eosin Methylene Blue

OD Optical Density

CIT Citrate Utilization

GEL Gelatine Liquefaction

Vis Visible

Consortium A MB490+ MB498

Consortium B MB490+ MB497

Consortium C MB490+ MB504

Consortium D MB497+ MB498

Consortium E MB498+ MB504

Consortium F MB497+ MB504

Consortium G MB490+MB497+ MB498+MB504

TCD Thermal Conductivity Detector

EI Electron impact

eV Electron volt

iv

min minutes

Hrs Hours

PCR Polymerase Chain Reaction

µM Micromolar

TaqDNA polymerase DNA Polymerase derived from Thermus aquaticus

dNTP Deoxyribonucleotide triphosphate

TBE buffer Tris/Borate/EDTA buffer

kb Kilo bases

Tris HCl (hydroxymethyl)aminomethane/ tromethamine

hydrochloride

STE buffer Sodium Chloride-Tris-EDTA

TE buffer Tris-EDTA buffer

mM millimolar

NBRIP National Botanical Research Institute’s phosphate

PSI Phosphate solubilization index

p-NP p-nitrophenol

p-NPP p-nitrophenol phosphate

nm nanometer

1N 1Normal

SDS Sodium dodecyl sulphate

EDTA Ethylene diamine tetraacetic acid

ALP Alkaline phosphatases

Vmax Maximum velocity

> More than

SBs slurry bioreactors

µS/cm micro-Siemens per centimeter

CIAP Calf Intestinal Alkaline Phosphatase

PhoN Non-specific Phosphatase

v

List of Figures

Figure No. Caption Page No.

1.1 Fate of pesticides in the environment 2

1.2 Different sources of pesticide pollution 2

1.3 Chemical structure of an organophosphate compound 4

1.4 Chemical structure of a). Tetrachlorvinphos. b). Dichlorvos. c).

Paraoxon.

5

1.5 Chemical structure of a). Chlorpyrifos. b). Methyl parathion. c).

Triazophos. d). Diazinon.

6

1.6 Chemical structures of: a). Dimethoate b). Malathion c). Phosmet

d). Oxydemeton methyl.

6

1.7 Diagram representing the possible routes of OP pesticides entering

humans and other animals

8

1.8 The chemical structures of triazophos and its main metabolite 1-

phenyl-3-hydroxy-1, 2, 4-triazole

15

1.9 Chemical structure of Dimethoate 17

2.1 Graphical scheme of methodology 31

2.2 Standard curve for p-Nitrophenol 55

3.1 Effect of different pH on the growth of bacterial isolates MB490,

MB497, MB498 and MB504.

70

3.2 Effect of different temperatures on the growth of bacterial isolates

MB490, MB497, MB498 and MB504

70

3.3 Percent degradation of Chlorpyrifos (800mg/l) by bacterial isolate

a) MB490. b) MB497. c) MB498. d) MB504 after 24, 48 and 72hrs

of incubation, using UV-Vis spectrophotometer.

72

3.4 DNA bands of 1). MB490, 2). MB498, 3). MB497 and 4). MB504

in the 2nd, 3rd, 4th and 5th well respectively, along with DNA ladder

in the 1st well.

73

vi

3.5 Evolutionary relationships of Pseudomonas kilonensis MB490. 74

3.6 Evolutionary relationships of Bacillus thuringiensis sp. MB497. 74

3.7 Evolutionary relationships of Pseudomonas kilonensis MB498. 74

3.8 Evolutionary relationships of Pseudomonas sp. MB504. 75

4.1 Growth and % degradation of CPF by four isolates (MB490,

MB497, MB498 and MB504) at different pHs after 24hrs. Error

bars represent standard errors for values of three sample replicates.

80

4.2 Growth and % degradation of CPF by 7 consortia (A, B, C, D, E,

F and) G) at different pHs after 24hrs.

80

4.3 Effect of temperature on growth and degradation of CPF by four

isolates (MB490, MB497, MB498 and MB504) after 24hrs.

81

4.4 Effect of temperature on growth and degradation of CPF by 7

consortia (A, B, C, D, E, F and) G) after 24hrs.

82

4.5 Effect of shaking versus static conditions on growth and %

degradation of CPF by four isolates (MB490, MB497, MB498 and

MB504) after 24hrs.

83

4.6 Effect of shaking versus static conditions on growth and %

degradation of CPF by 7 consortia (A, B, C, D, E, F and G) after

24hrs.

84

4.7 Effect of presence of TAP on the degradation of Chlorpyrifos

(CPF) and growth (OD) by the 4 isolates (MB490, MB497, MB498

and MB504) along with the consortium G (mixture of all four

bacterial isolates) in M-9 broth after 24hrs incubation.

84

4.8 Effect of incubation period on the degradation of Chlorpyrifos and

growth of the 4 isolates (MB490, MB497, MB498 and MB504) in

the M-9 broth.

88

4.9 Effect of incubation period on the biodegradation of Chlorpyrifos

(CPF) by the 7 consortia (A, B, C, D, E, F and G) of bacterial

isolates in the M-9 bacterial culture broth.

88

vii

4.10 Effect of incubation period on percentage degradation of

Chlorpyrifos (CPF) by the 4 isolates (MB490, MB497, MB498 and

MB504) in the soil slurry

90

4.11 Effect of incubation period on percentage degradation of

Chlorpyrifos (CPF) by 7 consortia (A, B, C, D, E, F and G) in soil

slurry.

90

4.12 Effect of incubation period on degradation of Chlorpyrifos (CPF)

by the 4 isolates (MB490, MB497, MB498 and MB504) in the soil

microcosm

93

4.13 Effect of incubation period on degradation of CPF by 7 consortia

(A, B, C, D, E, F and G) in soil microcosm

94

4.14 Effect of incubation period on growth (OD600) and degradation of

3, 5, 6-Trichloropyridinol (TCP) by the 4 isolates (MB490,

MB497, MB498, MB504) in the M-9 broth as analyzed by HPLC

96

4.15 Effect of incubation period on growth (OD600) and degradation of

3, 5, 6- Trichloropyridinol (TCP) by the 4 isolates (MB490,

MB497, MB498, MB504) in the M-9 broth as analyzed by HPLC

97

4.16 Effect of pH on degradation of CPF (RT= 5.4min) by MB490. (a)

Control, (b) MB490 at pH 6 (c) pH 7 (d) pH 8.

98

4.17 Effect of pH on degradation of CPF (RT= 5.4min) by MB497. (a)


99

4.18 Effect of pH on degradation of CPF (RT= 5.4) by MB498. (a)


100

4.19 Effect of pH on degradation of CPF (RT= 5.4) by MB504. (a)

Control, (b) MB504 at pH 6, (c) pH 7, (d) pH 8.

101

4.20 Effect of pH on degradation of CPF (RT= 5.4min) by Consortium

A. (a) Control, (b) Consortium A at pH 6 (c) pH 7 (d) pH 8.

102

4.21 Effect of pH on degradation of CPF (RT= 5.4min) by consortium

B. (a) Control, (b) Consortium B at pH 6 (c) pH 7 (d) pH 8

102

viii


C. (a) Control, (b) Consortium C, at pH 6 (c) pH 7 (d) pH 8.

104


D. (a) Control, (b) Consortium D, at pH 6 (c) pH 7 (d) pH 8.

105


E. (a) Control, (b) Consortium E, at pH 6 (c) pH 7 (d) pH 8.

106


F. (a) Control, (b) Consortium F, at pH 6 (c) pH 7 (d) pH 8. TCP

peak is prominent (RT= 1.7min).

107


G. (a) Control, (b) Consortium G, at pH 6 (c) pH 7 (d) pH 8.

108

4.27 Effect of temperature on degradation of CPF (RT= 5.4min) by

MB490. (a) Control, (b) MB490, at 25°C(c) 30°C (d) 37°C.

109



110

4.29 Effect of temperature on degradation of CPF (RT= 5.4) by MB498.

(a) Control, (b) MB498, at 25°C(c) 30°C (d) 37°C. A single and

prominent peak of TCP can be noted at 1.77 min at 37°C.

111

4.30 Effect of temperature on degradation of CPF (RT = 5.4min) by


112

4.31 Effect of temperature on degradation of CPF (RT= 5.4) by

consortium A. (a) Control, (b) consortium A, at 25°C (c) 30°C (d)

37°C.

113


consortium B. (a) Control, (b) consortium B, at 25°C(c) 30°C (d)

37°C.

114


consortium C. (a) Control, (b) consortium C, at 25°C(c) 30°C (d)

37°C.

115

ix


consortium D. (a) Control, (b) consortium D, at 25°C(c) 30°C (d)

37°C.

116


consortium E. (a) Control, (b) consortium E, at 25°C(c) 30°C (d)

37°C.

117


consortium F. (a) Control, (b) consortium F, at 25°C(c) 30°C. Note

peak for TCP at RT = 1.77min. (d) 37°C.

118


consortium G. (a) Control, (b) consortium G, at 25°C(c) 30°C (d)

37°C.

119

4.38 (a) HPLC chromatogram of standard TCP. Degradation of 14mg/l

3, 5, 6-Trichloropyridinol (TCP) by (b). MB490, c). MB497, d).

MB498, e). MB504 after 72hrs incubation.

120

4.39 (A). GCMS Chromatogram of Chlorpyrifos standard (RT =

22minutes). (B). GCMS Chromatogram of 3, 5, 6

Trichloropyridinol (TCP) standard with retention time (RT) = 12.8

minutes.

122

4.40 (A). Mass spectra of Chlorpyrifos standard (m/z: 20-320). (B).

Mass spectra of Chlorpyrifos degraded by bacteria (m/z: 20-260).

123

4.41 GCMS Chromatograms of Chlorpyrifos degraded by: (A). MB490

(B). MB497 (C). MB498 after 3 days incubation in M-9 broth

125

4.42 GCMS Chromatograms of Chlorpyrifos degraded by MB504 after

3 days incubation in M-9 broth.

126

4.43 GCMS Chromatogram of Chlorpyrifos degraded by: (a).

Consortium A (b). Consortium B (c). Consortium C (d).

Consortium D (e). Consortium E (f). Consortium F (g).

Consortium G after 3 days incubation in soil microcosm.

127

4.44 (A). GCMS chromatogram of TCP standard (RT = 12.8min). (B).

Mass spectra of standard TCP.

129

x

4.45 GCMS chromatogram (A) and Mass spectrum (B) of TCP

degraded by MB490. GCMS chromatogram of TCP degraded by

(C). MB497 after 72hrs in M-9 broth

130

4.46 GCMS chromatogram of TCP degraded by (A). MB498 (B).

MB504 after 72hrs in M-9 broth

131

4.47 Proposed pathway for 3, 5, 6-Trichloropyridinol (TCP)

degradation by bacterial isolates.

132

4.48 Proposed metabolic pathway of Chlorpyrifos degradation by

bacteria.

134

5.1 Optical density and % degradation of TAP by four isolates

(MB490, MB497, MB498 and MB504) at different pH after 24 hrs.

138

5.2 Optical density and % degradation of TAP by 7 consortia (A, B, C,

D, E, F, and G) at different pH after 24 hrs.

139

5.3 Optical density and % degradation of TAP by four isolates

(MB490, MB497, MB498 and MB504) at different temperatures

after 24 hrs.

139

5.4 Optical density and % degradation of TAP by 7 consortia (A, B, C,

D, E, F, and G) at different temperature after 24 hrs.

139

5.5 Effect of shaking and static conditions on growth and TAP

degradation by bacterial isolates (MB490, MB497, MB498 and

MB504).

140

5.6 Effect of shaking and static conditions on growth and TAP

degradation by bacterial consortia (A, B, C, D, E, F and G).

140

5.7 Effect of presence of CPF on the growth (OD600) and degradation

of Triazophos by the 4 isolates (MB490, MB497, MB498, MB504)

and the consortium G

141

5.8 Effect of incubation period on bacterial growth and % degradation

of TAP by bacterial isolates (MB490, MB497, MB498, MB504) in

M-9 broth.

142

xi

5.9 Effect of incubation period on the growth and degradation of

Triazophos by the bacterial consortia (A, B, C, D, E, F, and G) in

M-9 broth

143

5.10 Effect of incubation period on degradation of Triazophos by the

bacterial isolates (MB490, MB497, MB498, and MB504) in the

soil slurry

144

5.11 Effect of incubation period on degradation of Triazophos by the 7

consortia (A, B, C, D, E, F and G in the soil slurry

144

5.12 Effect of incubation period on degradation of Triazophos by

bacterial isolates (MB490, MB497, MB498, and MB504) in the

soil microcosm

146

5.13 Effect of incubation period on degradation of Triazophos by the

bacterial consortia (A, B, C, D, E, F and G) in the soil microcosm

146

5.14 Effect of pH on degradation of TAP (RT= 2.4) by MB490. (a)


148

5.15 Effect of pH on degradation of TAP (RT= 2.4) by MB497. (a)


149

5.16 Effect of pH on degradation of TAP (RT = 2.4min) by MB498. (a)


150

5.17 Effect of pH on degradation of TAP (RT= 2.4min) by MB504. (a)


151

5.18 Effect of pH on degradation of TAP (RT= 2.4) by consortium A.

(a) Control, (b) consortium A, at pH 6, (c) pH 7, (d) pH 8.

152

5.19 Effect of pH on degradation of TAP (RT= 2.4) by consortium B.

(a) Control, (b) consortium B, at pH 6, (c) pH 7, (d) pH 8.

153

5.20 Effect of pH on degradation of TAP (RT= 2.4) by consortium C.

(a) Control, (b) consortium C, at pH 6, (c) pH 7, (d) pH 8.

154

xii

5.21 Effect of pH on degradation of TAP (RT= 2.4) by consortium D.

(a) Control, (b) consortium D, at pH 6, (c) pH 7, (d) pH 8.

155

5.22 Effect of pH on degradation of TAP (RT= 2.4) by consortium E.

(a) Control, (b) consortium E, at pH 6, (c) pH 7, (d) pH 8.

156

5.23 Effect of pH on degradation of TAP (RT= 2.4) by consortium F.

(a) Control, (b) consortium F, at pH 6, (c) pH 7, (d) pH 8.

157

5.24 Effect of pH on degradation of TAP (RT= 2.4) by consortium G.

(a) Control, (b) consortium G, at pH 6, (c) pH 7, (d) pH 8.

158

5.25 Effect of temperature on degradation of TAP (RT= 2.4) by MB490.

(a) Control, (b) MB490 at 25°C, (c) 30°C, (d) 37°C.

159



160



161

5.28 Effect of temperature on degradation of TAP (RT = 2.4) by

MB504. (a) Control, (b) MB504 at 25°C, (c) 30°C, (d) 37°C.

162

5.29 Effect of temperature on degradation of TAP (RT= 2.4) by

consortium A. (a) Control, (b) consortium A at 25°C, (c) 30°C, (d)

37°C.

163


consortium B. (a) Control, (b) consortium B at 25°C, (c) 30°C, (d)

37°C.

164


consortium C. (a) Control, (b) consortium C at 25°C, (c) 30°C, (d)

37°C.

165


consortium D. (a) Control, (b) consortium D at 25°C, (c) 30°C, (d)

37°C.

166

xiii


consortium E. (a) Control, (b) consortium E at 25°C, (c) 30°C, (d)

37°C.

167


consortium F. (a) Control, (b) consortium F at 25°C, (c) 30°C, (d)

37°C.

168


consortium G. (a) Control, (b) consortium G at 25°C, (c) 30°C, (d)

37°C.

169

5.36 (a). GCMS chromatogram of Triazophos standard (RT= 27.8min)

(b). GCMS mass spectrum of Triazophos standard.

170

5.37 GCMS chromatogram of Triazophos in the sample treated with (a)

MB490 (b) MB497 after 3 days of incubation in M-9 broth.

171

5.38 GCMS chromatogram of Triazophos in the sample treated with (a)


172

5.39 Proposed metabolic pathway for the biotransformation of TAP by

bacterial isolates MB490, MB497, MB498 and MB504.

176

6.1 Effect of pH on growth (OD600nm) and % degradation of

Dimethoate by four isolates (MB490, MB497, MB498 and

MB504) after 24 hrs.

181

6.2 Effect of temperature on growth (OD600nm) and % degradation of

Dimethoate by four isolates (MB490, MB497, MB498 and

MB504) after 24 hrs.

181

6.3 Effect of pH on growth (OD600nm) and % degradation of

Dimethoate 7 consortia (A, B, C, D, E, F, and G) after 24 hrs.

182

6.4 Effect of temperature on growth (OD600nm) and % degradation of

Dimethoate by 7 consortia (A, B, C, D, E, F, and G) after 24 hrs.

182

6.5 Effect of Shaking versus static conditions on growth and %

degradation of Dimethoate by four isolates (MB490, MB497,

MB498 and MB504) after 24hrs.

183

xiv

6.6 Effect of Shaking versus static conditions on growth and %

degradation of Dimethoate by seven consortia (A, B, C, D, E, F,

and G) after 24hrs.

183

6.7 Effect of incubation period on growth and degradation of

Dimethoate (DM) by the 4 isolates (MB490, MB497, MB498,

MB504) in the M-9 broth

185

6.8 Effect of incubation period on growth and degradation of

Dimethoate (DM) by seven consortia (A, B, C, D, E, F, and G) of

bacterial isolates in the M-9 broth

185

6.9 Effect of incubation period on degradation of Dimethoate (DM) by

the 4 bacterial isolates (MB490, MB497, MB498 and MB504) in

the soil slurry

186


the 7 consortia (A, B, C, D, E, F, and G) in the soil slurry

187


the bacterial isolates (MB490, MB497, MB498 and MB504) in the

soil microcosm

188


the consortia (A, B, C, D, E, F, and G) in the soil microcosm

189

6.13 a) HPLC chromatogram of Dimethoate control. b). HPLC

chromatogram of Dimethoate degraded by MB490 in M-9 broth

after24hrs at pH 6. c). pH 7. d). pH 9.

190



after24hrs at pH 6. c). pH 7. d). pH 9.

191

6.15. a) HPLC chromatogram of Dimethoate control. b). HPLC


after 24hrs at pH 6. c). pH 7. d). pH 9.

192



after 24hrs at pH 6. c). pH 7. d). pH 9.

193

xv



after 24hrs at 25°C. c). 30°C. d). 37°C.

194




195




196




197

6.21 a) HPLC chromatogram of Dimethoate control. HPLC

chromatogram of Dimethoate degraded by: b). MB490. c).

MB497. d). MB498. e).MB504 in soil microcosm after 9 days of

incubation.

198

6.22 (a). GCMS chromatogram of Dimethoate standard (RT =

17.7min). (b). Mass spectrum of Dimethoate standard.

200

6.23 GCMS chromatogram of Dimethoate (RT = 17.7min) degraded by:

A). MB490. B). MB497. C). MB498. D). MB504 in soil

microcosm after 3 days of incubation.101

202

6.24 GCMS chromatogram of Dimethoate (RT = 17.7min) degraded by

a). Consortium A, b). Consortium B, c). Consortium C, d).

Consortium D, e). Consortium E, f). Consortium F, g). Consortium

G, h). Mass spectrum of Dimethoate degraded by bacteria in soil

microcosm after 9 days of incubation.

203

6.26 Proposed metabolic pathway for the transformation of Dimethoate

by bacteria.

205

7.1 Extracellular production of Organophosphorus-phosphatase (OPP)

by bacterial isolates (MB490, MB497, MB498 and MB504) at

405nm.

211

xvi

7.2 Intracellular production of Organophosphorus-phosphatase (OPP)

by bacterial isolates (MB490, MB497, MB498 and MB504)

measured at 405nm.

211

7.3 Organophosphorus-phosphatase production by bacterial isolates

(MB490, MB497, MB498 and MB504) at different pHs

212

7.4 Organophosphorus phosphatase production by bacterial isolates

(MB490, MB497, MB498 and MB504) at different temperatures.

213

7.5 Effect of incubation time on Organophosphorus-phosphatase

production by bacterial isolates (MB490, MB497, MB498 and

MB504).

213

7.6 Effect of temperature on activity of Organophosphorus-

phosphatase produced by bacterial isolates (a). MB490, (b).

MB497.

215

7.7 Effect of temperature on activity of Organophosphorus-

phosphatase produced by bacterial isolates (a). MB498 (b).

MB504.

216

7.8 Effect of substrate (p-NPP) concentration on activity of

Organophosphorus-phosphatase produced by bacterial isolates (a).

MB490 (b). MB497.

217

7.9 Effect of substrate (p-NPP) concentration on activity of

Organophosphorus-phosphatase produced by bacterial isolates (a).

MB498 (b). MB504.

218

7.10 Effect of incubation period on activity of Organophosphorus-


MB497.

219

7.11 Effect of incubation period on activity of Organophosphorus-


MB504.

220

7.12 Effect of SDS on activity of Organophosphorus-phosphatase

produced by bacterial isolates (a). MB490 (b). MB497 (c). MB498

(d). MB504

222

xvii

7.13 Effect of EDTA on activity of Organophosphorus-phosphatase


(d).MB504.

224

7.14 Effect of metals on activity of Organophosphorus-phosphatase


(d).MB504.

226

7.15 Effect of incubation period on Metal bioprecipitation of Ni++

(1000ppm) by OPP produced by a). MB490. b). MB497. c).

MB498. d). MB504.

229

7.16 Effect of Incubation period on Metal bioprecipitation of Mn++


MB498. d). MB504.

230

7.17 Effect of Incubation period on Metal bioprecipitation of Cr++


MB498. d). MB504.

232

7.18 Effect of Incubation period on Metal bioprecipitation of Cd++


MB498. d). MB504.

233

7.19 Degradation of OP pesticides by acidic and alkaline OPP as

analyzed by HPLC after 30min of incubation. a). MB490. b).

MB497, c). MB498. d). MB504.

237

7.20 HPLC chromatograms showing degradation of 50mg/l of TAP (RT

= 2.4min) by alkaline OPP as analyzed by HPLC after 30min of

incubation. a). Control b). MB490. c).MB497. d). MB498. e).

MB504.

238

8.1 Comparison of degradation of CPF, TAP and DM by MB490

(Pseudomonas kilonensis), MB497 (Bacillus thuringiensis),

MB498 (Pseudomonas kilonensis), and MB504 (Pseudomonas

sp.) after 9 days of incubation. a). M-9 broth b). Soil slurry c). Soil

microcosm.

245

xviii

8.2 Comparison of degradation of 3, 5, 6 Trichloropyridinol (TCP) at

two different concentrations (14mg/l and 28mg/l) by bacterial

isolates MB490 (Pseudomonas kilonensis), MB497 (Bacillus

thuringiensis), MB498 (Pseudomonas kilonensis) and MB504

(Pseudomonas sp.).

246

8.3 Comparison of degradation of TAP by pure strains (MB490,

MB497, MB498 and MB504) with their respective consortia A, B,

C, D, E, F and G in M-9 broth after 9 days.

251



C, D, E, F and G in soil slurry after 9 days.

252



C, D, E, F and G in soil microcosm after 9 days.

253

8.6 Comparison of degradation of DM by pure strains (MB490,


C, D, E, F and G in M-9 broth after 9 days.

255



C, D, E, F and G in soil slurry after 9 days.

256



C, D, E, F and G in soil microcosm after 9 days.

257

xix

List of Tables

Table No Title Page No.

1.1 Physical and chemical properties of Chlorpyrifos, Triazophos and

Dimethoate

11

2.1 Nutrient Broth 24

2.2 Nutrient Agar Medium 24

2.3 M-9 Medium 25

2.4 Oxidase Test Reagent 25

2.5 Oxidation Fermentation Medium 26

2.6 Crystal Violet Solution 26

2.7 Safranin Solution 26

2.8 Iodine Solution 26

2.9 40% CuSO4 Solution 26

2.10 0.5% Malachite Green Solution 26

2.11 Hydrogen Peroxide Solution 26

2.12 National Botanical Research Institute’s phosphate growth broth

(NBRIP)

27

2.13 0.1M Sodium acetate Buffer 27

2.14 0.1M Tris HCl Buffer 27

2.15 0.1M Sodium Phosphate Buffer 27

2.16 p-Nitrophenyl Phosphate buffer mixture (pNPP) 28

2.17 p-Nitro phenol (p-NP) 28

2.18 1N NaOH 28

2.19 1N HCl 28

2.20 Ni+2 solution 29

2.21 Cr+6 solution 29

2.22 Mn+2 solution 29

2.23 Cd+2 solution 29

2.24 Cu+2 solution 29

2.25 Zn+2 solution 30

xx

2.26 Pb+2 solution 30

2.27 Co+2 solution 30

2.28 Fe+3solution 30

2.29 Naphthalene solution 30

2.30 Consortia made by 4 bacterial isolates (MB490, MB497, MB498,

and MB504).

43

2.31 The GCMS conditions used for the detection of metabolites of

Chlorpyrifos, Triazophos and Dimethoate.

48

2.32 HPLC conditions used for the analyses of Chlorpyrifos,

Triazophos and Dimethoate degradation.

49

2.33 10X TBE Buffer 52

2.34 STE Buffer 52

2.35 TE Buffer 52

2.36 1M Tris-HCl 53

2.37 Universal 16s RNA gene amplification primers 53

2.38 PCR amplification conditions 53

3.1 Physical and Chemical properties of Soil. 62

3.2 CFU/ml of Bacterial isolates 62

3.3 Comparison of maximum tolerance of bacterial isolates against

three OP pesticides (CPF, TAP and DM).

64

3.4 Effect of different concentrations of Trichloropyridinol (TCP) on

Growth and color of bacterial isolates grown on M-9 medium.

65

3.5 Colony morphology of Orgnophosphate degrading bacterial

isolates.

65

3.6 Cell morphology of organophosphate degrading bacterial isolates. 66

3.7 Results of different biochemical tests 67

3.8 Maximum tolerance of bacterial isolates against heavy metals. 69

3.9 Maximum tolerance of bacterial isolates against organic

pollutants/chemicals.

70

xxi

3.10 Percentage degradation of Chlorpyrifos (800mg/l) by bacterial

isolates as analyzed by UV-Vis Spectrophotometer.

71

4.1 Bacterial Population in soil slurry spiked with 200mg/l of CPF 91

4.2 Bacterial Population in soil microcosm spiked with CPF

(200mg/kg).

93

4.3 Biotransformation of Chlorpyrifos by the 4 bacteria in M-9 broth,

soil slurry and soil microcosm as analyzed by GCMS.

124

4.4 Biotransformation of 3, 5, 6-Trichloropyridinol (TCP) by the 4

bacteria in M-9 broth as analyzed by GCMS.

128

5.1 Bacterial Population in soil slurry spiked with 200mg/l of TAP 144

5.2 Bacterial Population in soil microcosm spiked with 200mg/kg of

TAP.

146

5.3 Biotransformation of Triazophos by bacterial isolates and their

consortia in M-9 broth, soil slurry and soil microcosm.

173

6.1 Bacterial Population in soil slurry spiked with 200mg/l of DM 187

6.2 Bacterial Population in soil microcosm spiked with 200mg/kg of

DM

189

6.3 Biotransformation of Dimethoate (DM) by the 4 isolates (MB490,

MB497, MB498, and MB504) and the consortia of these 4 isolates

in the M-9 broth, soil slurry and soil microcosm.

201

7.1 Screening for Phosphate solubilization potential of bacterial

isolates (MB490, MB497, MB498 and MB504).

209

8.1 Main compounds identified in the GCMS chromatogram of

Dimethoate degraded by bacterial isolates (MB490, MB497,

MB498 and MB504) and their consortia.

260

xxii

Dedication

My Ph.D thesis is dedicated to my Loving Parents, whose unconditional love

and support along with their prayers enabled me to complete my Ph.D.

xxiii

Acknowledgements

All praises and thanks to Almighty Allah for giving me strength and courage to complete my thesis.

Modest respects to Holy Prophet Muhammad (PBUH), Who is persistently a guidance for entire

human kind.

I would like to express my sincere gratitude to my advisor Prof. Dr. Azra Yasmin for the

continuous support of my Ph.D study and related research work, for her patience, motivation, and

immense knowledge. Her guidance helped me in all aspects of my research and writing of this

thesis. She proved best advisor and mentor for my Ph.D study. Special thanks to Mrs. Irum Asif,

Mr. Joseph Gill, Mr. Riaz and Mrs. Komal for their technical assistance during my research in

their Analytical lab. My heartiest thanks to all lab fellows especially Anila, Satara, Uruj, Tabeer,

Ayesha and Monaza for their tolerance and cooperation during my research work.

Last but not least, I am especially thankful to my parents, brothers and sisters for their love,

prayers, patience and endless support throughout my life. I specially acknowledge my husband

and my kids who facilitated me and provided me a peaceful environment at home to complete this

study.

Samina Ambreen

xxiv

Summary

Over use of pesticides including organophosphates has resulted in the contamination of soil, water

and food resources, thus harming non-target organisms via food chain. The removal of pesticides

and other pollutants from soil and water has become the point of interest for many research

workers since last few decades. Multiple varieties of microbes have been reported that have

potential to degrade pesticides. Therefore present study was aimed to isolate indigenous

organophosphate (OP) degrading soil bacteria from various agricultural soils of district Mianwali,

Punjab, Pakistan. Among a large number of bacterial isolates obtained, fifteen isolates (MB490

to MB505) were selected and purified based on their discrete morphological characters and their

resistance against three OP pesticides Chlorpyrifos (CPF), Triazophos (TAP) and Dimethoate

(DM). Four best isolates (MB490, MB497, MB498 and MB504) were selected for further

analyses. Strain MB497 was the most tolerant for CPF (8g/l), followed by MB490 and MB498

(6g/l) and then MB504 (0.8g/l). Strains MB490, MB497 and MB498 were tolerant up to 4g/l of

TAP and 0.4g/l of DM, whereas MB504 was least tolerant for TAP (2g/l) and DM (0.22g/l).

Isolate MB497 was Gram positive, while the other three were Gram negative. All were rods and

facultative anaerobes. All the isolates were able to grow under wide range of temperature (25-

42◦C) and pH (5-11), and were characterized as neutrophile (MB490), slightly acidophile

(MB504) and moderately alkaliphile (MB497 and MB498).

All of the isolates showed resistance against multiple organic and inorganic pollutants including

heavy metals (Pb, Mn, Zn, Fe, Cr, Cu, Ni) and aromatic hydrocarbons (Benzene, Toluene, Xylene,

Biphenyl and Naphthalene). Biochemical analysis of these isolates revealed the presence of

important enzymes like nitrate reductase, oxidase and catalase needed for biodegradation. Four

selected strains were identified as Pseudomonas kilonensis MB490, Bacillus thuringiensis

MB497, Pseudomonas kilonensis MB498 and Pseudomonas sp. MB504 on the basis of 16s rRNA

analysis. These strains and their seven consortia (A, B, C, D, E, F and G) efficiently degraded

40.01 - 99.99% of initial 200mg/l of CPF, TAP and DM at different pHs (6, 7, and 8) and at

different incubation temperatures (25, 30 and 37◦C). There was significant degradation of CPF

(up to 99%), TAP (88.4 – 99.9%) and DM (77.67 to 99%) in M-9 broth, soil slurry and soil

microcosm for most of the isolates/consortia within 9 days. Negligible pesticide degradation was

observed in the control samples without bacterial inoculum. The bacterial isolates were also found

xxv

capable of degrading 3, 5, 6-Trichloropyridinol (28mg/l) up to 90.57% within 72hrs. GCMS

analysis of CPF degradation revealed the formation of 3, 5, 6 Trichloropyridinol (TCP) along

with [(3, 5, 6-trichloro-2-pyridinyl) oxy] acetic acid; Diethyl thiophosphate (DETP);

phosphorothioic acid and Diisopropyl methanephosphonate. TCP was further metabolized

into 1-methyl-2-pyrrolidine ethanol; 2-Ethoxy-4, 4, 5, 5-tetramethyloxazoline; 3-(2, 4, 5-

Trichlorophenoxy)-1-propyne and p-Propyl phenol.

While 7 novel metabolites of TAP were identified according to NIST library i.e 1, 2, 4-Triazole-

4—amine, N-(2-Thienylmethyl); Benzene sulfonic acid hydrazide; Benzene sulfonic acid

methyl ester; 4H-1,2,4-Triazole-4-benzenesulfonamide; 4, 5 dihydro-N-(O-toyl)-3-furamide;

Ethyl 4-phenyldiazenylbenzoate and Dibutyl methanephosphonate. Dimethoate was

metabolized into 5 products i.e Methyl diethanol amine; Phosphonothioic acid propyl-O, S-

dimethyl ester; O, O, O- Trimethyl thiophosphate; Omethoate and Aspartylglycine ethyl

ester.

All bacterial strains were capable of phosphate solubilization and could produce substantial

amounts of extracellular, acidic, neutral and alkaline organophosphorous phosphatases (OPP).

Mainly, there was more production of acidic and alkaline phosphatases as compared to neutral

phosphatases. Among three types of OPP, Pseudomonas kilonensis MB490 produced maximum

amount of neutral OPP, Bacillus thuringiensis MB497 produced maximum alkaline OPP, whereas

Pseudomonas sp. MB504 showed highest production of acidic OPP. The highest OPP production

was observed at pH 11 by all isolates. The maximum OPP enzyme production was exhibited at

50°C (MB490, MB497 and MB498) and at 45°C (MB504). Generally, OPP production was

reduced by all isolates after 48hrs. Though OPP enzyme activity and stability was maximum at

37⁰C, but remained still active even at highest given temperature (70⁰C) by all isolates. Largely,

acidic, neutral and alkaline OPP activity was inhibited by SDS and EDTA especially at higher

concentrations, whereas, Zn++, Cu++ and Cd++ significantly enhanced OPP activity. All three OPPs

exhibited 86-100% bioprecipitation of selected metals (Ni, Mn, Cr and Cd). Alkaline OPP showed

more degradation (80 to 99%) of given 50mg/l of three OP pesticides (CPF, TAP and DM), within

30min of incubation as compared to acidic OPP that was able to degrade up to 40 to 80%.

xxvi

Present research revealed that these indigenous bacteria (Pseudomonas kilonensis MB490,

Bacillus thuringiensis MB497, Pseudomonas kilonensis MB498 and Pseudomonas sp. MB504)

exhibited significant tolerance and degradation of multiple organophosphate pesticides (CPF,

TAP and DM) along with their unique aspect to degrade and transform recalcitrant 3, 5, 6

Trichloropyridinol (metabolite of CPF) which was further supported by the identification of many

known as well as novel metabolites formed by these isolates during degradation process. These

strains were resistant to a variety of metals and organic pollutants. Presence of acidic, neutral and

alkaline OPP and other enzymes (nitroreductase, oxidase and catalase) involved in degradation

process further support the possible use of these indigenous bacteria for bioremediation of OP

pesticide contaminated soils of Pakistan.

1

Chapter 1

Introduction

Pesticides are the organic compounds which are synthesized and used to control pest

either by killing or slowing down their growth. Pesticides include herbicides,

insecticides, fungicides, and nematocides. These are being used extensively in modern

age agriculture to increase food production for huge human population. The extensive

and continuous use of man-made pesticides leads to environmental pollution which is

enhanced by other diffused sources like unintentional spills, left over pesticides in the

drums or vessels, mishandling of application tools and also the undesirable application

methods (Diez, 2010). Pesticide contamination affects the quality of air, soils, food and

water resources, ultimately harming non-intended organisms including humans and

inducing pesticide resistance in pests by inborn mutations (Gavrilescu et al., 2015;

Aceves-Diez et al., 2015).

Biological, chemical and physical agents/factors regulate the steadiness as well as

movement of pesticides in the environment. Most of the pesticides are obstinate, as they

persevere for a longer time in soils and deposits, ultimately entering the food chain

(Ortiz-Hernandez et al., 2013). Though quantity of pesticides which are applied

annually reaches millions of tons, yet merely a minor portion of these pesticides targets

the intended organisms, while the major part is wasted reaching the soil, water and

atmosphere harming humans and other non-subjective species (Gill et al., 2014; Lewis

et al., 2016).

Traditional methods to deactivate pesticides mainly depend on chemical treatment,

incineration and landfilling. These methods have many downsides including the release

of large amounts of acids and alkali as secondary waste product (in case of landfills),

leaching of pesticides into surrounding soil and ground water supplies, and toxic

contamination of the environment as a result of incineration. As opposed to the old and

outdated methods, bioremediation using microorganisms or plants (phytoremediation)

is now considered as an alternative and smooth, potentially applicable, active, cheaper

and an eco-friendly method (Liu et al., 2016; Rayu et al., 2017). The microorganisms

are the chief biological mediators that can remove and degrade waste materials, in order

to allow their recycling in the surroundings (Parte et al., 2017).

2

Figure 1.1 Fate of pesticides in the environment (Source: Ortiz-Hernandez et al., 2013).

Figure1.2 Different sources of pesticide pollution (Source: Diez, 2010).

Soil bioremediation includes methods like phytoremediation, bioaugmentation,

biostimulation and enzymatic bioremediation (Adams et al., 2015). A bacterial

consortium had ability to degrade Profenofos up to 93.39% (Jabeen et al., 2015).

Various isolates (Bacillus alkalinitrilicus, Brevibacterium sp., Bacillus sp.,

3

Pseudomonas putida F1, Rhizobium sp.) had been reported to breakdown imidacloprid

(Sharma et al., 2014; Sabourmoghaddam et al., 2015).

1.1 Pesticides in Pakistan

The pesticides have been imported in Pakistan since 1954. The pesticide business was

given to the private sector in 1980. This resulted in a high rise of pesticide consumption

reaching to 5519 metric tons in 1992. Total 85% of the pesticides consist of insecticides

used mostly on cotton crop (Jabbar and Mallick, 1994). The pesticides in Pakistan are

regularized/controlled by the Agricultural Pesticide Rules, 1973 with respect to their

production/ import, registration and use etc. The Department of Plant Protection

working under government of Pakistan is responsible for the registration of pesticides.

All the agricultural workers especially the spray men and female cotton pickers are at

greater risks to pesticide poisoning. The pesticides are mostly handled without

appropriate precautionary measures as a result of lack of awareness. Pesticide remains

have been detected in many food and feed items sometimes exceeding the WHO

recommended limits. Likewise, crop soils and groundwater at various sites have

been found contaminated with the pesticide residues, ultimately reaching the human

bodies. There are only limited studies available in Pakistan on the human exposure to

pesticides. Moreover, these are only limited to organochlorine and organophosphates.

In Quetta and Karachi, organochlorine like DDT and HCH have been detected in the

blood and fat tissues of human population (Jabbar and Mallick, 1994). Similarly, the

activities of blood choline-esterase have been found reduced to dangerously low levels

in the spray men and cotton pickers in cotton growing area of Multan. Khan et al. (2015)

studied risks related to pesticide use among farmers from cotton belt of Punjab,

Pakistan. According to their survey, 4875 kg of pesticides/year was being used by the

farmers including 55% moderately risky and 23% extremely dangerous. There were 10-

11 average pesticide applications in each growing season. Most of the farmers were

unaware or less informed about pesticide risk and safe use of pesticides. So, there must

be alternative safer measures of integrated pest management (involving natural pest

control and biological control) and decrease in dependence on man-made pesticides.

Pesticide residues (chlorpyrifos, fenvalerate, dimethoate, methyl parathion,

fenitrothion, cypermethrin etc) were present in various fruit samples and in apple,

these were found above the permissible limits (Anwar et al., 2011). Chlorpyriphos

4

followed by Endosulfan and Dimethoate were the mostly observed pesticides in soil

samples of various cotton fields in Sindh (Anwar et al., 2012). Similarly, high

concentrations of DDTs and HCHs were noticed at pesticide dumping sites and adjacent

areas in Hyderabad, posing potential risk to living organisms, safety of food crops and

ultimately to human health (Alamdar et al., 2014).

1.2 Organophosphate pesticides

Organophosphate (OP) pesticides are man-made esters, thiol derivatives, or amides, of

phosphoric, phosphonic, phosphonothioic or phosphorothioic acids. In OP pesticides

there is a fundamental phosphorus atom, which is doubly bonded either with oxygen to

form oxon pesticides, or with sulfur to give thion pesticides (figure 1.3).

Figure. 1.3 Chemical structure of an organophosphate compound.

The R1, R2 and X groups which are attached to the central phosphorus atom

(pentavalent) by a single-bond, are different for Oxons and thions. R1 and R2 are

commonly alkyl (ethyl or methyl) groups, whereas X is the unstable leaving acyl

residue indicating the main metabolite of a certain OP compound. Many

organophosphates are resistant to chemical, thermal and photolytic breakdown due to

the stability and inertness of C-P bond as compared with their analogues having O-P,

S-P or N-P linkages which are more volatile (Kazemi et al., 2012). Organophosphate

pesticides (OPs) are major group of pesticides being used widely since World War II

to control agricultural pests, disease vectors, public sanitation, and domestic pests. They

have also been used as chemical warfare agents because of their potential to inactivate

acetyl cholinesterase enzyme in insects, humans and many other animals, leading to

nerve toxicity (Gupta, 2015). There are 3 types of Organophosphate pesticides (Silva

et al., 2013).

1. Phosphotriesters

2. Thiophosphotriesters

5

3. phosphorothiolesters

Phosphotriesters have a centeral phosphate attached with three O-linked groups. Their

main reperesentatives are Tetrachlorvinphos, Dichlorvos, Paraoxon (figure 1.4).

a. b.

.

c.

Figure 1.4 Chemical structure of a). Tetrachlorvinphos. b). Dichlorvos. c). Paraoxon.

Thiophosphotriesters contain sulfur instead of the phosphoryl oxygen (Silva et al, 2013;

Bigley and Raushel, 2013). They include Chlorpyrifos, Methyl parathion, Triazophos

and Diazinon (figure 1.5). In phosphorothiol esters, atleast one of the ester oxygen is

replaced by sulfur. They have members like Dimethoate, Malathion, Phosmet,

Oxydemeton methyl (figure 1.6). OP pesticides have replaced the highly toxic

organochlorine pesticides and include the most extensively used insecticides that are

currently available (Abhilash and Singh, 2009). OP pesticides are also greatly toxic

because of their potential to inhibit acetylcholinesterase (AChE) irreversibly thus

seriously effecting the nervous system of man and other exposed organisms (Gonzalez-

Alzaga et al., 2014). These compounds are commercially much successful and are main

part of agrichemicals, being used as an integral element in the modern agriculture

worldwide. According to a research report, 19 % (1.6 billion kg) of total world use of

glyphosate (8.6 billion kg) during last few decades has been utilized in the USA only.

Glyphosate use has increased to 15 times worldwide especially after introduction of

genetically modified glyphosate-tolerant crops in 1996 (Benbrook, 2016).

Dichlorvos Tetrachlorvinphos

Paraoxon

6

a. b.

c. d.

Figure 1.5. Chemical structure of a). Chlorpyrifos. b). Methyl parathion. c).

Triazophos. d). Diazinon.

a. b.

c. d.

Figure. 1.6 Chemical structures of: a). Dimethoate b). Malathion c). Phosmet d).

Oxydemeton methyl.

Diazinon

Triazophos

Methyl parathion Chlorpyrifos

Dimethoate Malathion

Phosmet Oxydemeton methyl

7

Even though OP compounds are considered harmless as compared to organochlorines,

yet they are still extremely neurotoxic for humans (Martin-Reina et al., 2017).

Pesticides may enter the living body by inhalation, ingestion, and skin penetration

(Garcia-Garcia et al., 2016). In the modern times, there are 140 types of pesticides as

well as plant growth regulators belonging to organophosphates being used worldwide.

Moreover it has been assessed that 1.5 thousand or more types of organophosphates

had been manufactured during the last century (Kang et al., 2006). Though they may

be very beneficial as insecticides, their exhaustive and haphazard use has resulted in

acute and chronic environmental threats and health issues (Suratman et al., 2015). The

inappropriate handling and improper storage of OP pesticides may give rise to severe

environmental hazards at both waste disposal sites especially near to cultivated areas as

well as at OP manufacturing factories resulting in leaching into soil/water resources.

These environmental risks and health hazards due to obsolete pesticides have the

potential to affect several countries (Matthews, 2015).

1.2.1 The toxicity of OP compounds and inhibition of Acetyl Cholinestrase (AChE)

OP pesticides are strong inhibitors of cholinesterase enzymes that function as

neurotransmitters. These enzymes include acetylcholinesterase, butylcholinesterase,

and pseudocholinesterase. These enzymes are inhibited as they bind to the OP

compound. This inhibition results in the accumulation of neurotransmitter acetylcholine

at neuron gaps (O'Brien, 2016). Under normal conditions, the Acetyl Cholinestrase

(AChE) catalyzes the hydrolysis reaction at the neuron synapsis and this reaction

depends on a serine residue present in the active site of enzyme reacting with the

carbonyl group in Acetylcholine (Ach) neurotransmitter compound. But when

organophosphates are present, the serine residue is phosphorylated quickly so that the

phosphorylated acetyl cholinesterase is inactivated (Santos et al., 2007). OP pesticides

are the most widely observed residues within contaminated foodstuff samples. These

include methamidophos, Chlorpyrifos and acephate as the major active components

which are causing food contamination (Silva et al., 2013; Xu et al., 2015).

Each year thousands of humans experience OPs toxicity all over the world. OP

pesticides along with some other chemicals have been declared as developmental

neurotoxicant since 2006 (Grandjean and Landrigan, 2014). It was reported that there

8

is more chance of cancer development in farm children exposed to pesticide-treated

agricultural fields (How et al., 2014). It had been observed that children affected by OP

pesticides were much expected to be diagnosed with attention deficit hyperactivity

disorder (ADHD) (Bouchard et al., 2010). Human exposure has been due to the

frequent use of OP pesticides in agriculture and their residues present in vegetables,

fruits, drinking water resources dairy and poultry products (Kim et al., 2017). Once in

the environment, pesticides are regulated by a number of biological, chemical and

physical routes. Though many OP pesticides can be degraded by microbial or

environmental methods, some of the pesticides may either be used up by living

organisms, or they might leach into ground water. When a pesticide reaches the ground

water, it can persist there as such for a significant period of time. As no sunlight

approaches in ground water, pesticide degradation is reduced, enhancing their possible

threats for human health/environment (Vermeire et al., 2003). The potential entry ways

of OP pesticides to humans or other biota in the environment are shown in figure1.7

Figure 1.7 Diagram representing the possible routes of OP pesticides entering humans

and other animals (Source: Vermeire et al., 2003).

9

Toxicity of OP compounds interacts with any given biological system depending upon

dose and is expressed as lethal dose (LD) which is the dose required to kill 50% of the

given animal species (LD50). These LD50 values are usually expressed as amount per

unit weight (mg/kg). There are both living as well as non-living factors responsible for

pesticide degradation in the natural environment. Yet microbes are the major

contributors for pesticide removal in various environmental compartments (Moorman,

2018). The studies on microbial degradation are helpful to develop strategies for

decontamination of pesticides. Biodegradation has become an important method to

remove organic impurities due to its cheap cost and less negative after-effects like

devastation of native animal and plant organisms as against other remediation

techniques (Verma et al., 2014). Recently isolation of indigenous bacteria that are able

to transform OP pesticides has been a focus for many scientists as these bacteria offer

a technique for in situ detoxification of pesticides and is also an environment friendly

method (Tang et al., 2017). The degradation of the pesticide deposits in the soil by the

natural local microflora is a much slower process which may be accelerated by

externally inoculating the soil with more competent/effective degrading microbes by

the practice of bioaugmentation (Cycon et al., 2013). Cyanobacteria have also been

reported to degrade OP pesticides. Malathion was removed by Nostoc muscorum up to

91% (Ibrahim et al., 2014). A bacterial strain Ochrobactrum sp. HZM was reported to

utilize quinalphos, profenofos, parathion‐methyl and chlorpyrifos as energy sources

(Talwar et al., 2014). Ortiz-Hernandez et al. (2003) revealed tetrachlorvinphos (TCV)

degradation by bacterial co-culture. Bacillus aryabhattai could degrade 70% parathion

within 120 hrs in mineral broth (Pailan et al., 2015). It has been observed that certain

native microorganisms living in polluted environments for a long period of time have

become adapted to these contaminants due to genetic evolution. These locations are the

most suitable ecological places for the isolation of OP degrading strains (Ortiz-

Hernandez et al., 2013). Hossain et al. (2015) reported complete removal of 20 mg/l

CPF by three bacterial strains within 8 -10 days.

1.3 Chlorpyrifos (CPF)

Chlorpyrifos is an important organophosphate pesticide used to control a large variety

of agricultural, veterinary and domestic pests (Das and Adhya, 2015). Generally half-

life of Chlorpyrifos in soil ranges from 60 to 120 days. Yet it may be as low as 2 weeks

to as long as more than one year, due to differences in the formulation, rate of

10

application, soil type, and the environmental conditions like soil type, pH, temperature

etc. (Racke et al., 1994; Williams et al., 2014). Soil bound Chlorpyrifos may undergo

break down by chemical hydrolysis, dechlorination, UV light and soil microbes

(Williams et al., 2014). Chlorpyrifos was reported to be immobile in certain soils like

clay, clay loam and peat (Halimah et al., 2016). Chlorpyrifos binding to soil particles

is very strong. It is less water soluble, thus has low mobility through soil. While its

metabolite TCP loosely binds to soil particles and is considerably more leachable in

soils (John and Shaike, 2015; Jaiswal et al., 2017). The widespread usage of

chlorpyrifos has led to pollution of soils in addition to water resources causing great

harm to non-target species (Solomon et al., 2014). The CPF contamination has been

noticed up to approximately 24 kilometres from the place of its application (Bhagobaty

et al., 2007). CPF being an OP pesticide has the potential to cause diseases like

Alzheimer’s and chronic fatigue (Peris-Sampedro et al., 2015). Neurobehavioural,

cardiovascular, and the respiratory health problems may have a link to low level

exposure to CPF due to its high acute toxicity resulting in increased public concern

about the extensive use of Chlorpyrifos in agriculture having potential human health

risks (Deeba et al., 2017; Dominah et al., 2017). Among organophosphates,

Chlorpyrifos is the much comprehensively studied pesticide. It interrupts neurons

duplication and maturation, axon production and neural function resulting in

developmental defects associated with experimental animals (used as models of

developmental CPF action) as well as in children highly exposed to CPF present in the

environment (De Felice et al., 2016). Among CPF breakdown products both CPF oxon

and trichloropyridinol have been reported as less effective to inhibit DNA synthesis in

neurons as compared with the parent compound (Wu et al., 2017). Physical and

chemical Properties of CPF are given in Table 1.1.

1.3.1 Mode of Action

CPF acts in a similar way for both intended and non-intended organisms (Roberts and

Reigart, 2013). Chlorpyrifos inhibits the cholinesterase enzyme, which results in

buildup of Acetylcholine in the synaptic gap causing over-reactivity of the neurons,

leading to neurological toxicity which ultimately may kill the CPF exposed organism.

Potentially lethal symptoms include diarrhea, neck muscle weakness, and respiratory

depression (Jacquet et al., 2016). In addition to cholinesterase, many other

11

biomolecules are probable targets for chlorpyrifos poisoning including interruption of

macromolecular (nucleic acids and proteins) synthesis, cytotoxicity and also

interactions with several enzymes. In humans, symptoms of CPF severe toxicity include

pain in head, weakness, vomiting, dizziness, muscle jerking, increased sweating and

salivation, happen as a result of about 50% decrease in cholinesterase activity (Eaton et

al., 2008). In case of mammalion non target organisms, CPF interrupts many other

enzymes, like A-esterases and carboxylesterases that occur in many mammalian

systems, though the function of these enzymes is not well understood (Karanth and

Pope, 2000).

1.3.2 Acute toxicity of CPF to non-target organisms

In general, LD50 is expressed as mg (milligrams) of the chemical per kg (kilogram) of

body weight, while LC50 is mostly expressed as weight (e.g mg) of the chemical per

volume (e.g liter) of the medium (air or water) to which the organism is exposed

(Adamson, 2016).

Table 1.1 Physical and chemical properties of Chlorpyrifos, Triazophos and

Dimethoate (Occupational Health Services, 1991; Meister, 1992; Marrs, 2002; Tomlin,

2009; Mackay et al., 2014).

Properties Pesticides

Chlorpyrifos Triazophos Dimethoate

Chemical formula C9H11Cl3NO3PS C12H16N3O3PS C5H12NO3PS2

Molecular weight

(g/mol)

350.6 313.312 229.26

Physical

appearance

Colorless/white

crystalline solid,

mild mercaptan

like smell

yellowish brown

oil

A colorless,

crystalline solid,

camphor-like

smell

Melting point 42°C 32-41°C 45-52°C

Boiling point 160 °C

Evaporates before

boiling

107°C at 0.05

mm Hg

12

Solubility 30-40 mg/l at 20°C

in water/ soluble

in many organic

solvents

39 mg/l in water at

20°C/ soluble in

various organic

solvents

25 mg/ml in

water at 21◦C.

Soluble in many

organic solvents

Vapor Pressure 2.02 × 10−5 mm Hg

(25 °C)

2.9 x 10-6 mm Hg

(30°C)

8.5 x 10-6 mm

Hg at 25°C

Density (g/cm3) 1.4 1.25 1.3

For Chemicals, when the LD50/LC50 is small, their toxicity is considered high and when

its value is larger, they are considered safe for health. Yet, the LD50/LC50 does not

indicate any long-lasting effects (like cancer, inborn defects or reproductive infertility)

resulting from longstanding exposure that usually occur at much lower concentrations

than those that cause death (Christensen et al., 2009).

Chlorpyrifos is highly toxic to birds, honey bees and mammals, freshwater fish, and

other aquatic organisms (Tomlin, 2009; Giesy et al., 2014). Different studies have

shown that newly born and the young ones are more vulnerable to harmful effects of

chlorpyrifos exposure as compared to adults at levels below those causing ChE

inhibition (Smegal, 2000: Grandjean and Landrigan, 2014). Research reports indicate

severe neurobehavioral effects, including effects on rat neuronal cell development and

DNA synthesis in experimental rat models (Kim et al., 2017).

1.3.3 Microbial Degradation of CPF

The degradation of CPF has been much less investigated inspite of their extensive use

(Kanekar et al., 2004). The CPF contains chloride radical in its structure, which makes

them less soluble in water. The biodegradation of CPF depends upon lipid (e.g.

rhamnolipids) producing microbes where lipid biosurfactants first dissolve and then

degrade CPF. Likewise, removal of chloride ion from CPF is carried out easily by the

microorganisms which have halogenase enzyme (Kanekar et al., 2004). Research

13

studies indicate that CPF is broken down to DETP and TCP by hydrolysis during

biodegradation by bacteria. Yet limited studies have been conducted to search for the

metabolism and degradation products of CPF (Xu et al., 2007). The degradation by

microorganisms is now understood to be the basic tool which determines the

environmental outcome and conduct of Chlorpyrifos (Kulshrestha and Kumari, 2011;

Ortiz-Hernandez et al., 2013). Research studies on microbial degradation are vital in

developing strategies for bioremediation to detoxify Chlorpyrifos and other pesticides

(Chen et al., 2011). Bioremediation is the biological process that involves using living

organisms or their products like enzymes to change a toxic/harmful material to a non-

toxic substance or to achieve the original condition (before contamination) of the

contaminated environment (Singh, 2009). During the last decade, bioremediation has

been established as a safe, cost effective, easily handled and smooth method to clear-

out polluted environments (Gao et al., 2012). Many different chemicals have been

removed effectively from soils and aquatic environments using degrading microbes

(Alvarenga et al., 2015).

The Chlorpyrifos degrading bacteria have been sequestered from CPF contaminated

agricultural land deposits, industrial effluents and waste water in the recent years (Li et

al., 2007; Latifi et al., 2012; Supreeth and Raju, 2016; Rayu et al., 2017). In alkaline

soils, Chlorpyrifos is briskly hydrolyzed by the soil bacteria (Singh and Walker, 2006).

Strain Burkholderia cepacia KR100 could hydrolyze chlorpyrifos-methyl to 3, 5, 6-

trichloro-2-pyridinol (TCP) and was also capable of degrading chlorpyrifos (Kim and

Ahn, 2009). Strains Ralstonia sp. T6 and Cupriavidus sp. DT-1 were capable of

metabolizing both CPF and TCP (Li et al., 2010; Lu et al., 2013). Similarly, a

cyanobacterium Synechocystis sp. PUPCCC 64 could metabolize CPF in to 3, 5, 6-

trichloro-2-pyridinol (Singh et al., 2011). Microbes were important factor in

degradation of Chlorpyrifos in non-sterilized soil slurries of paddy soils (Das and

Adhya, 2015). It was revealed in a study that Penicillium sp., Bacillus sp. and

Streptomyces thermocarboxydus were capable of degrading Chlorpyrifos in mineral

broth in the order: bacteria > fungi > actinomycetes (Eissa et al., 2014). A consortium

consisting of bacterial and fungal isolates (Cellulomonas fimi and Phanerochaete

chrysosporium) was able to completely remove CPF (50 mg/l) within 16 hrs, while

fungus alone took 6 days to degrade 50mg/l of Chlorpyrifos (Barathidasan et al., 2014).

14

A bacterial strain Acinetobacter calcoaceticus D10 could degrade 100 mg/l of CPF up

to 60% within 4 days (Zhao et al., 2014). It was revealed in a study that isolate Bacillus

safensis exhibited highest CPF degeadation followed by Bacillus subtilis and then

Bacillus cereus respectively (Ishag et al., 2016). Biodegradation is considered as the

most active route for the elimination of Chlorpyrifos even on the glacier surface

(Ambrosini et al., 2017).

1.3.4 Metabolic products of CPF

Chlorpyrifos metabolic pathway depends upon the various microorganisms involved in

the biotransformation and also the environmental conditions. It may result in four major

metabolites i.e 3, 5, 6-trichloropyridinol, chlorpyrifos oxon, Diethyl phosphate and

Diethyl thiophosphate (Sanchez-Hernandez et al., 2018). The CPF degradation

pathway leads to the formation of two unstable metabolites (chlorpyrifos oxon and

desethyl chlorpyrifos). Further, these metabolites are hydrolyzed to form the major

metabolite TCP, which is ultimately broken down to trichloro methoxypyridinol/carbon

dioxide (Racke, 1993). It has been previously reported that TCP is persistent in the soil

having t1/2 from 42-49 days in top soils and 64 -117 days in subsurface soils (Racke,

1993; Baskara et al., 2003). But in acidic top soils, the t1/2 of TCP has been reported

from 10 to 325 days (Racke, 1993). The major metabolites of CPF formed in soils,

plants and animals are identical to each other. These metabolites (diethyl phosphates

and 3, 5, 6-trichloropyridinol) are produced by hydrolysis/oxidative dealkylation of

CPF (Das and Adhya, 2015; Li et al., 2017).

So, briefly a great variety of microorganisms have been explored that are able to break

down CPF. Future studies must be sought for the complete mineralisation of

Chlorpyrifos as some of the intermediate metabolites of CPF degradation like TCP are

more insistent towards degradation by microbes and has also antimicrobial action.

Moreover, there should be enhanced research for co-culture of two or more

microorganisms, in order to achieve complete degradation of both chlorpyrifos and

TCP.

15

1.4 Triazophos (TAP)

Triazophos (TAP) has been an effective alternate to highly toxic organophosphate

pesticides, and applied worldwide on crops (maize, cotton, vegetables and paddy rice

etc.) for the last 40-50 years (Lin and Dongxing, 2005). Triazophos is being used with

wide-range and random effects as insecticide and miticide with a certain degree of

nematicidal effects. It exhibits tremendous preventive effects for many pests of the

major crops (grain, cotton and vegetables), such as stem borers, rice plant hoppers,

spider mites, cotton bollworms, lepidoptera and nematodes etc. (Aungpradit et al.,

2007). But its extensive use has resulted in environmental contamination by the TAP.

There are reports of higher concentrations of TAP in food, soil and water resources

(Zhong et al., 2006; Cherukuri et al., 2015; Bhamore et al., 2016). Average t1/2 of

Triazophos in different plant parts of wheat is 5.22 days, whereas in soil, it is 7.93 days

(Li et al., 2008). TAP may be harmful to beneficial organisms as it is inhibitor of

enzyme acetyl cholinesterase and neurotoxicant, thus possibly posing hazard to

mankind. It can also show genotoxicity, damaging immune genes and membrane

structural proteins of non- intended organisms (Zhong et al., 2005; Xiao et al., 2010;

Zhu et al., 2014; Wu et al., 2018). Ghaffar et al. (2014) conducted a research in Pakistan

to analyze the changes in blood composition caused by oral intake of TAP in birds

(male quail). They reported several medical symptoms like messy feathers, trembles,

watery stools, saliva secretion, less count of crowing and mating at high TAP

concentrations (6 and 8mg/kg/day) in treated Quails. Mahboob et al. (2015) studied the

severe effects of Triazophos in Cirrhinus mrigala baby fish in Pakistan and reported

critical toxic stress (choking, bottom dwelling, inconsistent swimming, exhaustion and

swigging before death). Therefore, it is need of the day to increase public concern about

its biological and ecological effects and to search for an efficient/suitable technique to

remove TAP residue. Physical and chemical properties of TAP are given in Table 1.1.

1.4.1 Biodegradation of TAP

Cheng et al. (2007) reported the 41–55% removal of Triazophos by Canna indica

(Linn.) in a water culture system. Research on biodegradation helps in developing

strategies to clear up pollutants by microorganisms (Hayatsu et al. 2000). Only few

triazophos degrading bacteria have been obtained from various sites (Wang et al., 2005;

16

Yang et al., 2011; Gu et al., 2014). Liang et al. (2011) revealed greater adsorption

tendency and microbial degradation of Triazophos along with chlorpyrifos, 1-phenyl-

3-hydroxy-1, 2, 4-triazole and 3, 5, 6-trichloro-2-pyridinol within paddy soil. Wu et al.,

(2016) reported the positive role of microbes in removal of TAP from polluted

wetlands. A bacterial isolate Roseomonas rhizosphaerae sp. nov. YW11T was reported

to be capable of degrading TAP (Chen et al., 2014). Isolate Pseudomonas stutzeri YC-

YH1 was capable of degrading Triazophos and other OP pesticides (Shi et al., 2015).

Burkholderia sp. SZL-1 was reported to hydrolyze Triazophos into 1-phenyl-3-

hydroxy-1, 2, 4-triazole (Zhang et al., 2016).

Figure 1.8 The chemical structures of triazophos and its main metabolite 1-phenyl-3-

hydroxy-1, 2, 4-triazole.

1.4.2 Metabolites of Triazophos

The well-known intermediate of TAP degradation is 1-phenyl-3-hydroxy-1, 2, 4-

triazole, which is formed by the hydrolytic break down of P–O ester bond in triazophos

molecule (Dai et al., 2007; Liang et al., 2011). Lin and Dongxing (2005) identified 4

main degradation products of TAP (monoethyl phosphorothioic acid, O, O diethyl

phosphorothioic acid, phosphorothioic acid and 1-phenyl-3-hydroxy-1, 2, 4-triazole).

Much earlier Schwalbe-Fehl and Schmidt (1986) working on degradation pathway of

TAP in dogs and rats suggested 3 metabolites of TAP i.e 1-phenyl-3-hydroxy-(1H)-1,

2, 4-triazole and its two conjugation products with glucuronide and sulfate. TAP oxon

and Phenylsemicarbazine has also been reported as metabolite of TAP in soil (Bock,

1975). Triazophos is one of major organophosphate pesticides being used worldwide

but there are only few reports on microbial degradation and metabolites of TAP as given

above. There is still no research done on TAP microbial degradation in Pakistan, though

it is being used here extensively contributing to TAP contamination in land and water

bodies. The main objective behind the present research work is to evaluate the potential

17

of indigenous bacteria to degrade TAP and thus to contribute to bioremediation of

agricultural soils contaminated with TAP.

1.5 Dimethoate (DM)

Dimethoate is an organophosphate pesticide containing an amide group with a systemic

action. DM has been used extensively all over the world since 1956 to kill a wide variety

of agricultural insects, though it is highly toxic for non-subjective organisms

(Lozowicka et al., 2016). Dimethoate is an organophosphate insecticide but is also

included in carbamate pesticides because of the presence of an amide group in its

molecule (Li et al., 2010). The widespread use of amide OP pesticides has contaminated

natural resources and food items. Amide pesticides are hazardous for human well-being

and can also harm vegetation if applied inappropriately (Tomlin, 2009; Aslanturk and

Çelik, 2016). The dimethoate and its remains have been found in farm products, land

and even milk products (Kotinagu and Krishnaiah, 2015; Hou et al., 2017; Han et al.,

2017). Dimethoate is harmful to various organisms including plants, birds and aquatic

animals. It badly affects photosynthesis and growth in plants, while inhibits brain

enzymes in birds, whereas swimming behavior of aquatic organisms is changed (Scoy

et al., 2016). Dimethoate has a half-life of 12 days under alkaline conditions

(Deshpande et al., 2004). It decomposes at higher (above 96◦C) temperatures

(Andreozzi et al., 1999). In soil without biodegradation and at low temperature (25◦C),

half-life of dimethoate may be increased to 206 days (Hassal, 1990). Physical and

chemical properties of Dimethoate are shown in Table 1.1.

Figure 1.9 Chemical structure of Dimethoate

1.5.1 Toxicological effects of Dimethoate

Dimethoate is considerably toxic by all means of intake i.e ingestion, inhalation and

dermal absorption (Damalas and Koutroubas, 2016). Similar to other

organophosphates, it freely enters body by the skin and lungs. Persons having

18

respiratory syndromes, recently exposed to cholinesterase inhibitors, already reduced

cholinesterase production, or having liver diseases may be highly threatened by

exposure to DM. Dimethoate toxicity may be boosted by elevated temperatures or

visible/UV light exposure of DM (Occupational Health Services, 1991). Severe eye

irritation has been reported to occur in industrial workers manufacturing Dimethoate,

firefighters and farmers when exposed to fumes of dimethoate (Mattos and Lorripa,

1982; Pham et al., 2016). In an experimental study, human lymphocytes were damaged

by dimethoate in a dose dependent manner (Soni et al., 2016). Being an

organophosphate insecticides, dimethoate is a cholinesterase inhibitor and is extremely

toxic by all means of exposure. Inhalation of DM results in coughing, runny nose, chest

pain, short breathing. Similarly, contact with Dimethoate through skin may give rise to

local sweating and spontaneous muscle tightening while eye contact would cause

blurred vision, pain and tears/ bleeding. The systemic effects may start immediately or

after up to 12 hrs of exposure by any means. Higher level of toxicity will damage the

central nervous system, thus producing incoordination, loss of reflexes, speech

difficulty, lethargy, jolting, tongue/eyelids trembling, and ultimately paralyzing the

body and leading to death (Occupational Health Services, 1991). For adult humans,

cholinesterase inhibition was observed at 30 mg/day or higher dosage (Hayes and Laws

1990). The continued exposure as well as severe exposure to Dimethoate may result in

the similar effects.

Dimethoate was found as carcinogenic, mutagenic teratogenic in cats and rats (Hayes,

1982; Hallenbeck and Cunningham-Burns, 1985). Dimethoate appeared to be less toxic

for animals having fast DM metabolism and larger weight ratios of liver to body (Hayes

and Laws, 1990). Dimethoate is more toxic to birds than mammals as birds cannot

metabolize dimethoate as quickly as mammals (Hudson et al., 1984; Meister, 1992).

Dimethoate (500 mg/kg) caused significant mortality within 24 hrs in rabbits and

symptoms included congestion, hemorrhages, and extensive degeneration in vital

organs of liver, kidneys, and brain (Baba et al., 2015).

1.5.2 Environmental fate of Dimethoate

Dimethoate is rapidly biodegraded in waste water treatment tanks. It under goes

significant leaching as it is highly soluble in water and is weakly adsorbed to soil

19

particles. There is hydrolytic degradation of DM, particularly under high pH condition.

Its evaporation occurs in dry soil surfaces. 23 to 40% of applied Dimethoate have been

reportedly lost by evaporation. Significant DM biodegradation (77%) has been reported

in a non-autoclaved soil within 14 days (Howard, 1991; USDA, 1990). T ½ of

dimethoate varies from 4 -122 days in soil. The rate of dimethoate break down increases

in moist soils and DM is readily degraded by soil microbes (Howard, 1991). There is

no Dimethoate adsorption to deposits or scattered particles under water bodies. It is

expressively hydrolyzed, specifically in waters with basic pH. It does not undergo

photolytic breakdown and is not evaporated significantly from surfaces of water bodies.

Its half-life was 8 days in fresh water body (river) and removal was probably as a result

of microbial and/or chemical degradation (Anyusheva et al., 2016). The half-lives of

dimethoate in purple flowering stalk, celery, Chinese kale and soil were reported to be

5.9–6.5, 3.5–5.4, 3.8–5.1, 3.4–3.6 days respectively (Chen et al., 2018).

1.5.3 Microbial degradation of Dimethoate

There has been a significant role for microorganisms to degrade or detoxify amide/OP

pesticides. Moreover metabolic pathways of pesticides by microbes have also been

revealed (Hashimoto et al., 2002; Li, et al., 2010; Pandey et al., 2010; Zhang et al.,

2011). Pseudomonas sutzeri was reported to tolerate dimethoate up to 900ppm and

completely removed dimethoate at pH8.5 and 30°C within72 hrs of treatment (Jadhav,

2014). There have been some reports for bacterial degradation of dimethoate from

different sites (Deshpande et al. 2001; DebMandal et al. 2002 and 2005). DebMandal

et al. (2008) reported four unidentified potential metabolites of dimethoate formed by

Bacillus licheniformis and Pseudomonas aeruginosa. Being a member of carbamate

pesticides, DM is slightly less readily degraded as compared with other common

organophosphates (Kanekar et al., 2004). The stability of dimethoate in nature depends

upon pH, temperature and type of the medium (Megeed and El-Nakieb, 2008). Liu et

al. (2001) reported 71.82% degradation of dimethoate by Pseudomonas stutzeri within

72 hrs. Pseudomonas putida and Bacillus pumulus could degrade DM up to 88 and 92%

respectively after72 hrs of incubation (Madhuri, 2014). 100 mg/l of dimethoate was

completely removed under 6 hrs of incubation in liquid culture by Paracoccus sp. (Li

et al., 2010). The concept of Effective Microorganisms (EM) utilizes different types of

microorganisms including bacteria, fungi, actinomyces and cyanobacteria. This unique

20

approach of the Effective Microorganisms was utilized by Megeed and El-Nakieb

(2008) to study the degradation of dimethoate in broth using the microbial consortia

that could tolerate and completely degrade dimethoate up to 120 mg/l within 3 days.

Strain Raoultella sp. could remove dimethoate up to 75% co-metabolically (Liang et

al., 2009).

Generally, microbial degradation of pesticides follows two types of mechanisms. The

first type is catabolism that utilizes both individual isolates as well as consortia that are

able to utilize precise pesticides as a sole source of carbon (Barton et al., 2004). The

individual components of mixed microbial cultures are not able to utilize chemicals as

a source of carbon/energy, so they are defined independently (Shelton and Somich,

1988). The second type is co-metabolism that indicates the biotransformation of a non-

growth substance only in the existence of an obligatory substrate needed for growth

(Liang et al., 2009). In this case, there is mutual use of enzymes for transformation of

the secondary substance as well as for utilizing primary growth substance. Optimization

of co-metabolism can be achieved by upholding the appropriate ratio of primary to

secondary substrates (Claus et al., 2007). Shinde et al. (2015) revealed that isolate

Pseudomonas spp.A1113 could degrade Dimethoate up to 19%. Chen et al. (2016)

reported the expression of cloned gene (dmhA) belonging to Sphingomonas, responsible

for dimethoate amidohydrolase in Escherichia coli BL21, which could transform DM

into two metabolites (methylamine and dimethoate carboxylic acid). Two bacterial

isolates Acinetobacter sp. Yj2 and Bacillus sp. Yj3 were able to degrade Dimethoate

due to the action of both acid and alkaline phosphatases having key roles in degrading

organophosphates (Yang et al., 2016). Begum et al. (2016) revealed the isolation of B.

pumilus from agricultural soil of Tamil Nadu which was very effective in DM

degradation. Co-culture of bacterial isolates Enterobacter cloacae, Panteoa sp,

Pseudomonas putida were reported to play important role in DM degradation in

Tunisian soil having t1/2 of 2.39 days for dimethoate in non-sterilized soil as compared

to 3.3 days in sterilized soil (Salem et al., 2016). Cellulolytic bacterial isolates

Pseudomonas putida and Bacillus pumulus could degrade dimethoate up to 88 and 92%

respectively within72 hrs of incubation (Madhuri, 2014).

21

1.5.4 Metabolites of Dimethoate

Li et al (2010) identified many metabolites of dimethoate (O,O,S-trimethyl

thiophosphorothioate , dimethoate carboxylic acid, 2-(hydroxyl (methoxy)

phosphorylthio)acetic acid, and O-methyl O,S-dihydrogen phosphorothioate etc,)

formed by Paracoccus sp. thus suggesting a metabolic pathway for DM. The

degradation of dimethoate even in mammals and plants has been reported (Gandhi et

al., 2015; Meghesan-Breja et al., 2017). Though Omethoate (dimethoate Oxon) is

considered as most common metabolite of dimethoate, but in plants, Dimethoate

carboxylic acid, Omethoate carboxylic acid and des-O-methyl-dimethoate carboxylic

acid were also reported as metabolites of DM in plants (Meghesan-Breja et al., 2017).

1.6 Microbial organophosphate degrading enzymes

Among the OP degrading enzymes, earliest isolated is phosphotriesterase (PTE). This

enzyme is capable of hydrolyzing variety of OP compounds (Serdar et al., 1989; Singh

et al., 2006). Another name for this enzyme is organophosphorus hydrolase (OPH) and

it shows great catalytic activity to hydrolyze a large number of organophosphates by

breaking P-S and P-O linkages. Enzymes mainly derived from microbes, like OPH,

OPAA (OP acid anhydrolases) and methyl parathion hydrolase (MPH), have been

proved as strong mediators for OP removal (Schenk et al., 2016). Ortiz-Hernandez et

al. (2003) tested a number of organophosphate pesticides with phosphotriesterase

isolated from Flavobacterium sp. They noted that some chemical structures of OP (like

TCV) were not responsive towards the enzyme activity. Gao et al. (2014) revealed that

immobilized OpdA was very effective to remove methyl parathion from solution. The

OP compounds detoxify initially by hydrolysis, which is considered a vital step as they

become more exposed for further microbial breakdown (Pinto et al., 2017). There have

been extensive studies to understand the process of hydrolysis and also its kinetic

features (Ortiz-Hernandez et al., 2003). Among OP hydrolases OpdA extracted from

Agrobacterium radiobacter is much effective enzyme (Selleck et al., 2017). The OP

hydrolase is capable of detoxifying OP-polluted environments (Ortiz-Hernandez et al.,

2011). Though there is much diversity of enzyme systems, but OPH and OPAA have

been focus of most studies of organophosphorus degrading enzymes.

22

OPDA has similarity to OPH in general secondary structure, yet these two enzymes

differ in substrate specificities due to difference in active site structure and thus OPDA

prefers for substrates having smaller alkyl substituents. It has been suggested that OPH

naturally evolved to OPDA by mutation in active site (Yang, 2003). There is another

important enzyme OPAA (organophosphorus acid anhydrolase), isolated from different

species of Alteromonas (Cheng et al., 1997). OPAA is less responsive for P–O but more

for P–F linkage (Hill et al., 2001). This enzyme has been found in both prokaryotes and

various eukaryotes thus suggesting its primitive origin (Cheng et al., 1999). Some other

organophosphorus degrading enzymes like parathion hydrolases,

aryldialkylphosphatase (ADPase), phosphotriesterases have been reported which are

structurally and functionally different (Mulbry and Karns, 1989a; Mulbry, 1992;

Mulbry, 2000; Horne et al., 2002). There have been reports of only a few

organophosphate hydrolyzing enzymes isolated from fungi including laccase (phenol

oxidase), A-OPH, P-OPH (Amitai, 1998; Liu et al., 2001; Liu et al., 2004). These

enzymes that have capability to immobilise or degrade organophosphate residues, may

be applied in bioreactors to decontaminate polluted water, after post-harvest

purification of farm /animal products to decrease or remove pesticide levels. Recently

Phosphotriesterase‐Like‐Lactonase (PLL) enzymes have been extracted from

thermophilic/extremophilic bacteria/hyperthermophilic archaea having extraordinary

thermal stability and are much promising for detoxification of OPs/nerve agents

(Manco et al., 2018).

1.7 Modern Analytical methods for the analyses of Pesticides

Multiple approaches like enzymatic analyses along with colorimetric methods have

been explored for the detection of organophosphates in different samples (Turner et al.,

2015). Chromatographic methods assisted with various detectors and spectroscopy

along with immunoassays/enzyme biosensors using cholinesterase activity inhibition

have been used for detecting pesticides and other pollutants (Samuels and Obare, 2011;

Tuzimski and Sherma. 2015; Zhang et al., 2016). Recently, Mass spectrometry (MS)

methods are being successfully applied for the qualitative and quantitative analysis of

several OP pesticides. These methods have extremely low detection limits (part per

billion and part per trillion) and also have the selective detection of analytes in multi-

residue samples (Wu et al., 2015). The mass spectrometry procedures are mostly joined

23

with separation methods like gas chromatography/high performance liquid

chromatography (Prapamontol et al., 2014).

GC-MS has been a most commonly used method for OP detection in any given sample

since the last decade. It has been utilized for different sample types like biological,

agricultural, environmental and foodstuffs (Fontana et al., 2010; Prapamontol et al.,

2014; Xu et al., 2015; Tette et al., 2016). The sensitive and selective determination of

OP analytes is possible through GC-MS analysis by using different combinations of

extraction procedures, ionization sources, analyzers and retention time.

1.8 Aims and Objectives

Present study is aimed to screen the indigenous isolates for their tolerance against OP

pesticides and also evaluate their degradation potential by analyzing their metabolites

both in liquid culture and in soil microcosm in order to use them for bioremediation of

contaminated agricultural soils in Pakistan. The novelty of this work lies in the fact that

up to my knowledge, it is first time that three important OP pesticides i.e Chlorpyrifos,

Triazophos and Dimethoate have been analyzed together in one study for degradation

by indigenous soil bacteria in Pakistan. In this study, the bacterial degradation of

Dimethoate and Triazophos is being studied for the first time in Pakistan. In order to

devise a strategy for bioremediation of OP contaminated agricultural soils, multiple

experiments in series were carried out with the objectives given below:

1. Characterization of organophosphate pesticide degrading indigenous bacterial

isolates. This includes morphological, biochemical, physiological and

molecular characterization.

2. Screening of organophosphate pesticide degrading bacteria against different

pollutants (organic and inorganic).

3. Optimization of the environmental conditions (pH, temperature/substrates) for

OP degradation.

4. To analyze the effect of OP pesticides (both separately and in mixture) on

degradation potential of OP degrading bacteria and their consortia in M-9 broth,

soil slurry and agricultural soil (microcosm).

5. Detection and identification of the metabolites produced by OP degrading

bacteria in culture media, soil slurry and in agricultural soil (microcosm).

6. Analysis/Assaying of OP degrading enzymes.

24

Chapter 2

Materials and Methods

All the solutions and media being used were prepared in distilled water and autoclaved

at 121°C and 15 lb/inch2 for 15 minutes for complete sterilization. Similarly, glassware

and plasticware were washed accurately and also oven dried to sterilize completely

before use. Moreover, all the research work was done under complete sterilized

conditions to avoid any contamination. All experiments related to screening,

optimization, quantitative and qualitative analyses etc. were performed in triplicate to

obtain more authentic results.

For this study, commercial-grade Organophosphate pesticides Chlorpyrifos,

Triazophos and Dimethoate (40% EC) were used supplied by Four Brothers Agri

services, Pakistan. The chemical composition of different chemicals along with media

used in this study is given below.

2.1 Preparation of solutions and media

Table 2.1 Nutrient Broth

S. No Ingredients g L-1

1 Peptone 5.0

2 Meat extract 3.0

3 NaCl 5.0

Table 2.2 Nutrient Agar Medium

S. No Ingredients g L-1

1 Peptone 5.0

2 Meat extract 3.0

3 NaCl 8.0

4 Agar agar 12.0

pH 7 ± 0.2

25

Table 2.3 M-9 Medium

S.

No

Ingredients g/l

1 Na2HPO4 6

2 KH2PO4 3

3 NaCl 0.5

4 NH4Cl 1.0

5 Agar 15

pH 7.4 ± 0.2

Among these ingredients in the media bottles, 1ml of 0.1M CaCl2 solution, 10 ml of

20% glucose solution (if required), 1ml of 1M MgSO4 solution and 5 g/l of Casein

hydrolysate were added after autoclaving of rest of the media, stirred and then poured

into the pre-autoclaved petri plates.

Table 2.4 Oxidase Test Reagent

S. No Ingredients g100 ml-1

1 NNNʹNʹ-Tetramethyl-p-Phenylenediamine

dihydrochloride

1.0 g

2 Distilled water 100 ml

Table 2.5 Oxidation Fermentation Medium

S. No Ingredients g m L-1

1 Peptone water 2.0

2 Sodium chloride 5.0

3 Dipotassium hydrogen phosphate 1.0

4 Bromothymol blue 0.03

5 Glucose 10.0

6 Agar agar 8.0

pH 7.1 ± 0.2

26

Table 2.6 Crystal Violet Solution

S. No Ingredients g 500 ml-1

1 Crystal violet 10.0

2 Ammonium oxalate 4.0

3 20% Ethanol 500ml

Table 2.7 Safranin Solution


1 Safranin 2.5

2 Ethanol 100ml

Table 2.8 Iodine Solution


1 Iodine 1.0

2 Potassium iodide 2.0

3 Ethanol 125ml

Table 2.9 40% CuSO4 Solution


1 CuSO4 40.0

2 Distilled water 100ml

Table 2.10 0.5% Malachite Green Solution


1 Malachite green 0.5

2 Distilled water 100ml

Table 2.11 Hydrogen Peroxide Solution

S. No Ingredients v/v

1 Hydrogen peroxide 3.0ml

2 Distilled water 7.0ml

27

Table 2.12 National Botanical Research Institute’s phosphate growth broth

(NBRIP) (Nautiyal, 1999).

S. No Ingredients g/l

1 Glucose 10

2 KCl 0.2

3 Ca3(PO4)2 5

4 MgCl2. 6H2O 5

5 MgSO4.7H2O 0.25

6 (NH4)2SO4, 0.1

Table 2.13 0.1M Sodium acetate Buffer (Weinberg and Zusman, 1990)

S. No Ingredients g/100ml

1 Sodium acetate 0.82

2 Acetic acid Up to pH adjustment

pH 5.2

Table 2.14 0.1M Tris HCl Buffer (Plisova et al., 2005)


1 Trizma base 1.2114

2 HCl Adjust pH

pH 9.5

Table 2.15 0.1M Sodium Phosphate Buffer


1 Sodium Phosphate Monobasic 1.09

2 Sodium Phosphate dibasic dehydrated 0.31

pH 7

28

Table 2.16 p-Nitrophenyl Phosphate buffer mixture (pNPP)

S. No Ingredients Amount

1 pNPP 0.6 mg

2 Sodium Phosphate Buffer (for neutral

Phosphatase enzyme assay)

1ml

3 Sodium acetate Buffer (for acidic

Phosphatase enzyme assay)

1 ml

4 Tris HCl Buffer (for alkaline Phosphatase

enzyme assay)

1 ml

For phosphatase enzyme essay, p-Nitrophenyl Phosphate (pNPP) was used as substrate

and was stored at 4°C along with the working stocks. Depending upon the type of

enzyme essay, i.e acidic, neutral or alkaline, buffer solutions were used accordingly.

Table 2.17 p-Nitro phenol (p-NP)

S. No Ingredients µg/ml

1 p-Nitro phenol 800

2 Methanol

As per

requirement

p-Nitro phenol (pNP) solution was used in standard curve preparation.

Table 2.18 1N NaOH


1 NaOH 0.4

2 Water 10 ml

Table 2.19 1N HCl


1 HCl 0.36

2 Water 10 ml

29

Table 2.20 Ni+2 solution


1 NiCl2.6H2O 2

2 Water 20 ml

Table 2.21 Cr+6 solution


1 K2Cr2O7 2

2 Water 20 ml

Table 2.22 Mn+2 solution


1 MnCl2 2

2 Water 20 ml

Table 2.23 Cd+2 solution


1 CdCl2 2

2 Water 20 ml

Table 2.24 Cu+2 solution


1 CuSO4 2

2 Water 20 ml

30

Table 2.25 Zn+2 solution


1 ZnSO4 2

2 Water 20

Table 2.26 Pb+2 solution


1 Pb(NO3)2 2

2 Water 20

Table 2.27 Co+2 solution


1 CoCl2 2

2 Water 20

Table 2.28 Fe+3solution


1 FeCl3 2

2 Water 20

Table 2.29 Naphthalene solution


1 Naphthalene 0.5

2 Ethanol 10

31

Figure 2.1. Graphical scheme of methodology

Soil Sampling and Physicochemical analysis

of Soil

Isolation of bacteria from soil

Purification of bacteria Morphological, Physiological and

biochemical characterization of bacterial isolates

Molecular Characterization of bacterial isolates

Screening of bacterial isolates against CPF, TAP,

DM and TCP on M-9 medium

Optimization of biodegradation of CPF,

TAP and DM

HPLC analysis of pesticide degradation by

Bacteria.

Organophosphorus Phosphatase (OPP)

Assay

Detection of Metabolite of pesticides formed

during Biodegradation using GCMS

32

2.2 Sampling Site and Soil Sampling

Soil samples used in the current study, were collected from three different cotton and

wheat fields with previous 20 years history of treatment with organophosphate (OP)

pesticides frequently, at Dera Saleemabad, Mochh, District Mianwali, Punjab, Pakistan.

The OP pesticides are mostly used to protect cotton and wheat crops from insects and

pests in this area. Generally, the climate in this part of southern Punjab is very hot and

dry during summers, while moderately cold in winters. For sample collection, the sandy

loam soil samples were collected in triplicate in polythene bags from 0-12 inches depth

in the early hours of morning. The collected soil had been treated with the OP pesticide

two weeks earlier, before the sampling. During sampling, separate and sterilized

trowels were used to collect each sample in order to reduce chance of external

contamination. Collected soil samples were then brought to laboratory immediately to

isolate bacteria, where these soil samples were sieved after being partially air dried

overnight before making suspension for isolation of bacteria.

2.3 Physicochemical Analysis of Soil

2.3.1 Measurement of the soil pH

The pH of soil samples was measured with the help of BMS Neomet pH meter (Thomas,

1996). For this purpose, 20 g of soil sample was suspended in 40 ml distilled water (1:2

ratio) followed by stirring intermittently using glass rod for half an hour and then

was left for an hour. The pH meter electrode was dipped into the soil suspension and

pH was recorded. The soil pH indicates the acidity and alkalinity of the soil and is a

very important soil property as it is deeply related with the nutrients accessibility, soil

microbial activity along with physical state of soil.

2.3.2 Electrical Conductivity (EC)

Electrical conductivity (EC) is another very important property of soil which gives ion

contents of soil solution indicating the soluble salt content of soil and thus current

carrying capacity of soil. The electrical conductivity (EC) of soil samples was

measured using a digital electrical conductivity meter (Rayment and Higginson,1992;

Wagh et al., 2013) and for this 20 g soil was mixed and stirred in 40 ml distilled

water for half an hour and kept undisturbed for 30 minutes so that soluble salts may

33

be dissolved completely and the soil is settled down and then filtered. Then conductivity

cell was dipped in soil suspension filtrate to note the EC value.

2.3.3 Soil organic matter

Soil organic matter consists of the remnants of plant material, roots and soil organisms

at various levels of degradation as well as synthesis. It also varies in composition.

Although soil organic matter is small in amounts yet it has a main impact on nutrient

availability, moisture retention, soil aggregation and biological activity. The Weight

Loss on Ignition method (Davies, 1974) was used to estimate soil organic matter and it

measures the weight loss from a dry soil sample when heated to high temperatures

(360oC). Finely ground soil (10 g) was taken in a crucible oven dried at 105◦C for two

hours and then weighed. It was heated again in a pre-heated oven at 360◦C for two hours

and then cooled to 150oC and weighed again. The weight loss occurring at this

temperature was then interrelated to oxidizable organic carbon using the following

formula:

% Organic carbon = (Soil weight at 105oC-weight at 360 oC) × 100

Soil Weight at 105◦C

2.3.4 Soil texture

Soil texture analysis refers to the measurement of the proportions of the different sizes

of the soil particles. There are three major groups of soil particles i.e sand (2.0-0.05

mm), silt (0.05-0.002 mm) and clay (< 0.002 mm). Soil texture effects the rate of

organic matter breakdown. The higher clay content in soils usually indicate a higher

organic matter content, as a result of slower rate of decomposition of organic matter.

The soil texture was determined with the help of Siever or Oscillator using standard

methods (Kettler et al., 2001) depending upon fractionation of different particle size of

soil and then calculating the percentage of each fraction against total weight of soil

sample used. Finally soil texture was determined by putting/comparing percentage of

each particle size i.e sand, silt and clay in Texture Triangle Chart.

34

2.3.5 Soil moisture Content

The soil water content was measured using analytical balance according to the standard

method (Wagh et al., 2013). For this purpose, fresh weight of soil along with crucible

dish was recorded. It was kept in the oven at 105 ◦C for overnight and then dry weight

of soil sample along with crucible dish was noted. The weight of the dish was then

subtracted from both dry and fresh weight of soil. The % water content of soil was

calculated by the following formula:

% water content of soil = Soil dry weight × 100

Soil fresh weight

2.4 Isolation of bacteria from the soil samples

In order to make soil suspension, 1 gm of each of the three soil samples was suspended

in 10 ml of autoclaved distilled water in a sterilized universal bottle, separately using a

vortex. The resultant soil suspensions were labelled as sample suspension (direct

sample) and were left on bench for 2-3 hours before use. This direct suspension was

used to make further dilutions of 1/10, 1/100, and 1/1000. In order to make 1/10

dilution, 100 µl of sample suspension was added to 1 ml of autoclaved distilled water

in an eppendorf and labelled as 1/10. Next, 100 µl of this 1/10 sample was mixed with

1 ml of autoclaved distilled water in an eppendorf and then labelled as 1/100. Likewise,

100µl of this 1/100 sample was taken and added to 1 ml of autoclaved distilled water

to get 1/1000 dilution.

For the initial isolation of bacterial population, nutrient agar medium (Table 2.2) was

prepared. Then 10 µl of direct as well as diluted samples (1/10, 1/100, 1/1000) was

taken and poured in the center of different agar plates labeled as direct, 1/10, 1/100,and

1/1000, respectively. Next, these soil suspensions were spread using glass spreader over

the whole surface of agar medium in the petri plate. This spreading was continued untill

the drying of liquid on the agar surface was achieved. Then the plates after being

covered with lids were incubated at 37◦C for 24-48 hours and different bacterial

colonies having different morphology were observed after incubation period.

35

2.4.1 Colony Counting (CFU/ml)

Colony counting is a measure of viable cells where a colony represents an aggregate of

cells derived from a single cell. CFU is applied to determine the number of active living

bacterial cells in a sample per ml or per gram. In order to determine the colony forming

units (CFU) of bacterial isolates in 1 g of soil, following formula was used:

CFU/g of soil ꞊ no. of colonies×1/dilution

2.4.2 Single colony streaking (bacterial culture purification)

For the purification of bacterial isolates, single colony streaking method was used,

where morphologically distinct bacterial colonies after selection, were picked and

purified by streaking on nutrient agar plates. This streaking method was continued until

a pure colony was obtained.

2.5 Screening for OP pesticide tolerance

In order to determine their resistance or tolerance for organophosphate pesticides, the

bacterial isolates MB490, MB497, MB498 and MB504 were grown on Minimal

medium M-9 (Table.2.3) (with and without 20% glucose) supplemented with different

concentrations of OP pesticides Chlorpyrifos, Triazophos and Dimethoate. The stock

solution of commercial grade Chlorpyrifos was made by dissolving 1 ml of OP in 10

ml of autoclaved distilled water and different concentrations of this stock solution (0.1,

0.2, 0.3, 0.4…….6%) were provided in the M-9 medium (Table 2.3), which were easily

tolerated by the isolates with very good growth. Next, these isolates were tested against

different concentrations i.e 0.1 to 20% (0.04 g/l to 8 g/l) of concentrated pure

commercial grade (EC 400 g/l) OP pesticides (Chlorpyrifos, Dimethoate and

Triazophos) supplemented to M-9 medium to check bacterial growth.

2.6 Screening for 3, 5, 6-Trichloropyridinol (TCP) tolerance

Trichloropyridinol is the major metabolite of Chlorpyrifos, and is known for its

antimicrobial effects (Racke et al., 1988). The isolation of CPF degrading bacteria had

been difficult in the past mainly due to killing of these bacteria by TCP. Therefore, the

four isolates MB490, MB497, MB498 and MB504 were tested for their tolerance

36

against TCP (Sigma Aldrich) by growing them on M-9 medium supplemented with 14

and 28 mg/l of TCP. Their growth was checked after 24 hrs incubation at 37°C.

2.7 Morphological characterization of OP pesticide degrading bacterial isolates

After screening, the morphological and biochemical characterization of purified

bacterial isolates was carried out.

2.7.1 Colony morphology

The visual and microscopic studies were conducted to observe and record colony

morphology of the bacterial isolates including shape, color, margins, elevation, texture

and size of colonies.

2.7.2 Cell morphology

Different tests like Gram’s staining, capsule staining, spore staining and motility test of

bacterial isolates were carried out to study the cell morphology.

2.7.2.1 Gram’s staining

Gram’s staining is a technique used to differentiate bacteria into two main group i.e

Gram-positive and Gram-negative, on the basis of cell wall composition. It was

introduced in 1884 by Hans Christian Gram as a differential staining method to study

the phenotypic characters of bacteria and their taxonomic grouping. The cell walls have

thick layers of peptidoglycan (90% of cell wall) in case of Gram-positive bacteria.

These bacteria retain the crystal violet-iodine complex during the staining process and

so, these are stained purple. While, Gram-negative bacteria have thin layers of

peptidoglycan (10% of wall) and higher lipid content in their cell walls and hence are

stained pink.

During Gram’s staining the first step is the smear preparation where a drop of distilled

water was put on a clean glass slide. Next, a small amount of fresh 24 hours old bacterial

culture was mixed well with the drop of water so that the smear was spread uniformly

on the slide surface, followed by air drying. Then, bacterial smear was heat fixed by

passing across the flame twice or thrice and stained with crystal violet solution (Table

2.6) for 1 minute followed by washing with distilled water. Next step was flooding with

37

iodine solution (Table 2.8) for 1 minute and washing with distilled water. The smear

was then dipped in 70% ethanol for 5-6 minutes, in order to remove extra stain.

Afterwards, the smear was air dried and stained with Safranin (Table 2.7) for 30-45

seconds. It was then washed with distilled water and air dried. Finally, a drop of oil

immersion was put on the smear and covered by a cover slip and observed under the

microscope using oil immersion lens.

2.7.2.2 Capsule staining

Many different types of bacteria both Gram-positive and Gram-negative, may possess

an outer layer called the glycocalyx that is referred to as a capsule if it is tightly bound

and remains attached to cells. Whereas the loosely bound layers are known as slime

layers. These bacterial capsules are composed of high molecular weight

polysaccharides and/or polypeptides along with polyalcohol and polyamines. Capsules

give protection against dehydration and assist in adherence of cells to surfaces and with

other bacteria thus forming biofilms.

The first step for the capsule staining is the smear preparation by mixing the 24 hours

fresh bacterial culture in a drop of distilled water. Next, the smear was air dried and

stained with 1% aqueous crystal violet solution (Table 2.6) for 1 minute. Then, it was

washed well with 40% copper sulfate solution (Table 2.9). After washing with copper

sulfate, the slide was instantly dried with the help of filter paper to avoid crystallization

of copper sulfate solution after its direct contact with air. Ultimately, a drop of oil

immersion was put on the slide and was covered with a cover slip. These slides were

then observed under the microscope (100X) in order to identify the shiny layer of

capsule in dim light.

2.7.2.3 Spore staining

The spores are formed in many genera of bacteria like Bacillus and Clostridium, not for

their reproduction, but as a mean of survival. The sporulation is stimulated by stress

conditions like shortage of nutrients or unfavorable environmental conditions. Spores

are produced in bacteria as a thick protective wall and are called endospores. While the

rest of the cell is known as vegetative cell.

38

In case of spore staining, 72-96 hours old bacterial cultures were used. The smear was

made by mixing bacterial culture in a drop of distilled water on the slide. Smear was

air dried, heat fixed and the malachite green solution (Table 2.10) was put continuously

on the slide over steam for 15-20 minutes. Then, the slide was washed with distilled

water followed by staining with safranin solution (Table 2.7) for 30-45 seconds. Stained

smear was washed again with distilled water, air dried and a drop of oil immersion was

put on the slide before covering with the cover slip. Slide was observed using the

microscope (100X) where the spores appear green after staining and the vegetative cells

give pink appearance.

2.7.2.4 Motility test

Bacterial motility usually depends on the presence of flagella or axial filaments. The

flagella are thin, protoplasmic, whip-like extensions of the cell. For motility test, a tiny

drop of water was placed on cover slip to prepare a hanging drop culture. Then, the

cover slip was inverted on a cavity slide, so as the bottom of drop did not touch the

slide itself. Next a drop of oil immersion was placed on the slide and the slide was then

observed in weak light using oil immersion lens.

2.8 Biochemical characterization

2.8.1 Oxidation Fermentation Test

Some bacteria, utilize glucose or other carbohydrates by oxidative pathway (aerobic

routes), while others are able to ferment sugars anaerobically (fermentation reaction).

There also may be facultative anaerobes capable of using both aerobic and anaerobic

pathways. In case of Oxidation Fermentation test, the carbohydrate source is the

glucose, whereas bromothymol blue is used as an indicator. If the acid is produced only

in the open tube (without agar plug), it indicates that the organism is an oxidizer. While

in case of a fermenter, acid production will be both in the agar-covered tube and in the

open tube. The peptone in the medium may be utilized by some aerobic bacteria to form

ammonia resulting in appearance of blue color in the upper part of the open tube due to

alkaline conditions.

For oxidation fermentation (OF) test, four test tubes were used for each isolate while

four test tubes were labeled as control. Oxidation fermentation medium (Table 2.5) was

39

dissolved carefully in distilled water near boiling. Four ml of this medium was poured

in each test tube and then autoclaved. The fresh bacterial culture was used to streak all

test tubes after solidification except the control, while two of the inoculated test tubes

for each strain were sealed with 4 ml of molten agar to create anaerobic conditions.

Also, the two test tubes of control were plugged with agar and other two were left

unplugged. All the test tubes were incubated at 37°C for 24-48 hours and then the

results were recorded. The change in color from bluish green to yellow in the unplugged

test tubes indicated the oxidative pathway of isolates. While change in color of the

medium and the gas production in both open and plugged tubes indicated the

fermentative pathway of carbohydrates by the bacteria.

2.8.2 Oxidase Test

The oxidase test which is a biochemical reaction, is carried out to detect the presence

of enzyme cytochrome oxidase (indophenol oxidase). During this test, the colorless

reagent is reduced to an oxidized colored product by the organism having cytochrome

oxidase enzyme. This enzyme cytochrome oxidase takes part in the electron transport

chain reactions of bacterial respiration, to catalyze the oxidation of cytochrome c and

reduces oxygen to give water. For this oxidase test, tetra-methyl-p-phenylene diamine

dihydro chloride reagent is utilized as an artificial electron donor for cytochrome c.

There is a change in the reagent from colorless to a dark blue or purple compound,

indophenol blue, when it is oxidized by cytochrome C.

For the oxidase test, 1% Oxidase reagent (Table 2.4) was prepared in autoclaved

distilled water and this solution was kept in dark bottle and stored in refrigerator for

maximum a week. Next step was to saturate the autoclaved filter paper strips in the

oxidase reagent followed by air drying. A little amount of fresh bacterial culture was

taken with the help of inoculating loop and was rubbed against the filter paper strips

previously treated with the reagent. Then any change in color of strips was observed.

The isolates were oxidase positive, if the color changed to purple within 5-10 seconds

and when the color changed to purple after 60-90 seconds, the organism was considered

weak positive. The organism was considered negative, if the color did not change

during more than 2 minutes.

40

2.8.3 Catalase Test

In order to survive, organisms utilize defense mechanisms that help them to restore or

escape the oxidative breakdown by hydrogen peroxide (H2O2). For this purpose, the

enzyme catalase is produced by some bacteria to help in the cellular detoxification. The

catalase test is carried out for detection of the enzyme catalase in bacteria. This enzyme

named catalase counteracts the toxic effects of hydrogen peroxide. Its concentration is

especially higher in the pathogenic bacteria. The breakdown of hydrogen peroxide to

water and oxygen is catalyzed by catalase. This reaction is indicated in the form of

quick bubble formation. For the catalase test conduction, a little amount of bacterial

culture was put on a clean glass slide with the help of sterilized loop. Then 50 μl of

hydrogen peroxide (Table 2.11) was added to the culture. The catalase-positive

organism was confirmed by the formation of bubbles or foam.

2.8.4 Other biochemical/enzyme Tests

The API-20E System was used to characterize Enterobacteriaceae and other non-

fastidious Gram-negative bacteria. For API strip test, 0.8% saline solution was made

first, 7 ml of which was poured in each test tube for a bacterial isolate, and then test

tubes were sealed and autoclaved. After then, they were kept to cool down at room

temperature. A loopful of fresh bacterial culture (24 hours old) was put in the saline

solution in the test tube and was vortexed to get uniform bacterial suspension. The base

of incubation tray consisting of wells was filled with autoclaved distilled water to

provide humid conditions. Next, the strip having the dehydrated substrates was put in

the incubation tray. The bacterial culture prepared in saline solution was poured in to

each cup containing the dehydrated substrates. Five cups {Lysine decarboxylase

(LDC), hydrogen sulfide production (H2S), Arginine dihydrolase (ADH), Ornithine

decarboxylase (ODC), and urea hydrolysis (URE)} were half filled with bacterial

culture solution, whereas mineral oil was used to fill other half to provide anaerobic

conditions. The incubation tray was covered with the lid and then incubated for 18-24

hours at 37°C. After the incubation period, the strips were observed, interpreted and

recorded consequently. Specific reagents were also added for some of the tests such as

James reagent for Indole test, NIT 1 and NIT 2 reagents for Nitrate reduction test in

glucose tube, TDA reagent for Tryptophan deaminase test, and VP 1 and VP 2 reagents

41

for Voges Proskauer test. In the case of no red or orange color within the glucose tube,

small quantity of zinc dust was added and observed after five minutes for color change.

The nitrate reduction to molecular nitrogen was indicated by the appearance of red color

on addition of zinc dust while appearance of yellow color indicated a positive result.

2.9 Physiological Characterization

2.9.1 pH Optimization for Bacterial Growth

Different bacteria have specific optimal pH range for their best growth. The growth

reduction can occur, in case the pH of the environment varies from the optimal range.

There are many bacterial species that grow best at neutral or slightly alkaline pH.

Likewise, some bacteria grow best at low pH like 4, whereas others may grow at higher

pH as 11. The potential of bacteria to survive beyond their optimal pH range, depends

on their ability to tolerate or self-adjust changes in the environmental pH. Some bacteria

are naturally able to acclimatize to pH changes and are classified as acidophiles,

neutrophiles or alkaliphiles.

The effect of pH was studied by growing all the isolates in nutrient broth (Table 2.1) at

pH 5, 6, 7, 8, 9, 10 and 11 on a shaker at 37◦C. Earlier, four millilitre of this broth was

poured in each test tube, cotton plugged and autoclaved. For preparing bacterial

suspension, inoculum from fresh bacterial culture was mixed in 1ml autoclaved distilled

water in eppendorf and then 25 μl of bacterial suspension was added to each test tube

containing the sterilized nutrient broth which were then kept in the shaking incubator

(150 rpm) at 37°C for 24 hours. After the incubation period, the optical density (OD)

was measured using UV/VIS spectrophotometer (BMS UV-160) at 600 nm.

2.9.2 Effect of temperature on the growth of bacterial isolates

Like pH, temperature is very important factor, for bacterial growth. Bacteria have a

wide range of temperature for their growth. Psychrophilic bacteria are able to grow at

low temperature. Whereas, thermophilic bacteria can grow at higher temperatures.

However, most of the bacteria are mesophilic and can grow at temperatures between

5◦C to 63◦C (growth zone). They are having an optimum temperature of about 37 ◦C for

their growth.

42

The effect of temperature on bacterial growth was studied by growing all the bacterial

isolates (MB490, MB497, MB498 and MB504) at various temperatures (25◦C, 30◦C,

37◦C and 42◦C) in the nutrient broth at pH 7. For this purpose, 4 ml of nutrient broth

was taken in each test tube. After being cotton plugged, tubes were autoclaved.

Bacterial suspension was made using 24 hrs fresh bacterial culture for each isolate using

a sterilized loop and mixed with 1 ml of autoclaved distilled water in each eppendorf.

For each bacterial isolate, five test tubes having nutrient broth were taken and 25 μl of

bacterial suspension was added to each tube as inoculum, except the control test tubes.

Test tubes were incubated at the selective temperature for 24 hrs. After incubation

period, bacterial growth was estimated by measuring optical density (OD) using

UV/VIS spectrophotometer at 600 nm.

2.10 Heavy Metal Resistance Profile

The metals having density above 5 g/cm3 are called heavy metals and are essential trace

elements for living organisms. Though they play significant role in important

biochemical reactions but in higher concentration they lead to formation of complex

chemical compounds with toxic effects. Many bacteria are resistant to heavy metals due

to their inheritance and the genes responsible for metal resistance are related to plasmids

and transposon. As a result of natural and industrial processes, greater concentrations

of heavy metals are accumulated in the microbial environment. The microbial

organisms have evolved different methods for metal toleration like complexation, metal

ion reduction, or effluxion.

The isolates (MB490, MB497, MB498 and MB504) were tested for their heavy metal

resistance using various metals like Ni+2(NiCl2), Cr+6(K2Cr2O7), Mn+2(MnCl2),

Cd+2(CdCl2), Cu+2(CuSO4), Zn+2(ZnSO4), Pb+2[Pb(NO3)2], Co+2(CoCl2) and

Fe+3(FeCl3) (Table 2.20, 2.21, 2.22, 2.23, 2.24, 2.25, 2.26, 2.27, and 2.28 respectively).

Different concentrations of these metals ranging from 50-3000 μg/ml were

supplemented to the autoclaved M-9 medium and poured in Petri plates. It was allowed

to solidify and then, it was streaked with isolates. The Petri plates were kept in the

incubator at 37◦C for 24 hrs. After incubation period, bacterial growth and their color

was checked and noted.

43

2.11 Tolerance against different organic pollutants

Due to increasing industrialization, these organic pollutants are constantly

contaminating our soil and water resources Therefore, bacterial isolates MB490,

MB497, MB498 and MB504 were also tested for their tolerance against various

concentrations ranging from 0.1% to 6% of different organic pollutants (Benzene,

Toluene, Xylene, Aniline, Biphenyl and Naphthalene).

2.12 Bacterial Inoculum and Consortium Preparation

Potential bacterial isolates MB490, MB497, MB498 and MB504 were cultured in 30

ml of nutrient broth containing respective pesticides (200 mg/l) at 37ºC for 24 h to

obtain an OD600 of 0.6 and CFU/ml/g of 106 (Kumar et al., 2008; Sasikala et al., 2012).

During all the culture experiments, 500 µl of 24 hr old bacterial inoculum was used for

individual isolates. In case of biodegradation experiments in m-9 broth, viability of the

inoculated bacterial strains was determined in terms of OD600 by using UV-VIS

Spectrophotometer at various intervals of incubation in triplicate. Whereas, in case of

biodegradation experiments in soil slurry and soil microcosm, total viable counts of the

inoculated bacteria were determined by plate counts via serial dilution method at

different intervals (0, 3, 6 and 9 days) during the experiment (Salam et al., 2015). For

developing a consortium, the compatibility of individual bacterial isolates was checked

by streaking them together in combination of two and four isolates on nutrient agar

plate (Sasikala et al., 2012). Following consortia were developed.

Table 2.30 Consortia made by 4 bacterial isolates (MB490, MB497, MB498, and

MB504).

S. No Consortium Bacterial isolates

1 A MB490+ MB498

2 B MB490+ MB497

3 C MB490+ MB504

4 D MB497+ MB498

5 E MB498+ MB504

6 F MB497+ MB504

7 G MB490+MB497+ MB498+MB504

44

2.13 Analysis of OP pesticide degradation using UV-VIS Spectrophotometer

Initially, the degradation of Chlorpyrifos was studied by growing isolates MB490,

MB497, MB498 and MB504 in M-9 broth in test tubes supplemented with 800 mg/l

Chlorpyrifos as the sole source of carbon and nitrogen. Then, the test tubes including

control (without inoculum), were incubated at 120 rpm on rotary shaker at 37◦C. The

contents of the test tubes were centrifuged at 3500 rpm for 12 minutes after every 24

hrs interval up to 72 hrs. Next, the supernatant was analyzed by UV-VIS

Spectrophotometer (BMS UV-160) to analyze the Chlorpyrifos degradation by the four

isolates. The percentage of the compound degraded was found by the formula:

Percentage of degradation = (Xa–Xb) /Xa × 100

Where, Xa = absorbance of compound before degradation

Xb = absorbance at the same wavelength after degradation (Rokade and Mali,

2013).

2.14 Qualitative and Quantitative Analysis for OP pesticide degradation using

GC-MS and HPLC

A more elaborate and advanced analysis for OP pesticide degradation was conducted

using GC-MS (Shimadzu QP5050), and HPLC (Shimadzu LC-20AT). For this

purpose, bacterial isolates MB490, MB497, MB498 and MB504 and their different

consortia were grown stepwise in M-9 broth, soil slurry and soil microcosm

respectively. Extraction method for each is given below.

2.14.1 Extraction of pesticide residues from M-9 Culture Broth

Bacterial strains MB490, MB497, MB498 and MB504 and their different consortia

were grown in 30 ml of M-9 broth in 100 ml flasks supplemented with 200 mg/l of OP

pesticides CPF, TAP and DM separately and combined (CPF and TAP) in certain cases.

The culture flasks along with controls (without any inoculum) were incubated at 120

rpm in a shaker for 9 days. After incubation period of 3 days, 6 days and 9 days, 4 ml

of sample was taken from each flask using micropipette under sterile conditions in a 10

ml falcon tube. Its absorbance/optical density (OD) was observed at 600 nm to estimate

bacterial growth (Kale et al., 1989; Negi et al., 2014) followed by centrifugation at

45

3500 rpm for 20 minutes. The supernatant was taken in another falcon and ethyl acetate

was added to it in 1꞉1 ratio, and dried with 5 g of anhydrous Na2SO4 to absorb moisture

and to concentrate the extract. The sample was shaken well and left for 30 minutes

(Rokade and Mali, 2013; Pawar and Mali, 2014). Then 1.5 ml of upper organic layer

was collected in an Eppendorf and stored in biorefrigerater at -20◦C for few days (if

delay is necessary) untill used for GC-MS analyses etc. Just before GC-MS analyses,

1ml of extract was taken in a falcon and mixed with 9 ml of methanol to obtain 1: 9

ratio. Next, 1.5 ml of this mixture was filtered using Sartorius Ministart sterile syringe

filters (0.45 µm) to remove any particles and put in the GC vials for GC-MS analyses.

Each experiment was conducted in triplicate.

2.14.2 Extraction of 3, 5, 6-Trichloropyridinol (TCP) residues from M-9 Culture

Broth

Bacterial strains MB490, MB497, MB498 and MB504 were tested for their potential to

degrade and transform TCP, the major metabolite of CPF. For this purpose, they were

grown in 10 ml of M-9 broth separately in two sets of test tubes supplemented with 14

mg/l and 28 mg/l of Trichloropyridinol. The test tubes along with controls (without any

inoculum) were incubated at 120 rpm in a shaker for 24, 48 and 72 hrs. 4 ml of sample

was taken from each test tube after incubation period of 24, 48 and 72 hrs, with the help

of micropipette under sterile conditions in a 10 ml falcon tube. Its absorbance/optical

density (OD) was observed at 600 nm to determine bacterial growth (Anwar et al.,

2009) followed by centrifugation at 3500 rpm for 20 minutes. The supernatant was

shifted to another falcon and ethyl acetate was added to it in 1꞉1 ratio. It was dried with

5 g of anhydrous Na2SO4 to absorb moisture. After 30 minutes of shaking well, 1.5 ml

of upper organic layer was collected in an Eppendorf and stored in biorefrigerater at -

20◦C for few days until used for GC-MS and HPLC analyses. GC-MS and HPLC

instrumental conditions used were same for both CPF and TCP with slight

modifications (Brzak et al., 1998; Li et al., 2014).

Just prior to GC-MS analyses, 1 ml of extract was taken in a falcon and mixed with 9

ml of methanol to obtain 1:9 ratio. Next, 1.5 ml of this mixture was filtered using

Sartorius Ministart sterile syringe filters (0.45 µm) to remove any particles and put in

the GC vials for GC-MS analyses. For HPLC analyses, about 1 ml of the sample was

46

filtered through 0.45 µm syringe filter and put into the sample vial. Then a set volume

of the sample was injected into the column of HPLC system.

2.14.3 Extraction from soil slurry for OP pesticide degradation analysis

The biodegradation of OP pesticides by MB490, MB497, MB498 and MB504 and by

their consortium was also tested in soil slurry. For this purpose, 50 g soil was suspended

in 30 ml M-9 broth in 250 ml flasks to achieve 60% soil water content and autoclaved.

After then, this soil suspension was spiked with 200 mg of OP pesticide (CPF, TAP or

DM) and 500 µl of bacterial inoculum and incubated at 37◦C for 9 days. After 3 days,

6 days and 9 days, 3 ml of soil slurry was taken from each flask using a micro pipette

in a falcon tube and 10 ml of ethyl acetate was added to it. This suspension was filtered

through Butchner funnel with a cotton pad and filterate was centrifuged at 3500 rpm

for 20 minutes. The supernatant was shifted to another falcon containing 5 g of

anhydrous Na2SO4 and kept for 30 minutes. Then, 1 ml of upper organic layer was

taken in an Eppendorf and stored at -20◦C in biomedical freezer for further analyses by

GCMS. Before GC-MS analyses, 1ml of extract was mixed with analytical grade

methanol in 1: 9 ratio. Next, 1.5 ml of this extract solution in methanol was syringe

filtered using Sartorius Ministart sterile syringe filters (0.45 µm) and added to GC vials

being used for GCMS analyses (Polese et al., 2002; EL-Bestawy et al., 2014). Each

experiment was conducted in triplicate.

2.14.4 Extraction from soil microcosm for OP pesticide degradation analysis

For OP pesticide degradation analysis in soil microcosm, 50 g of agricultural soil was

finely ground, sterilized and then added to autoclaved glass Petri plates. These 50 g soil

samples were spiked with 200 mg/kg of OP pesticides (CPF, TAP and DM) separately

and 500 µl of 24 hrs old bacterial inoculum (106- 107cfu/g of soil, suspended in 10 ml

M-9 broth as fertilizer) (Ueno et al., 2006) was mixed well to obtain uniform

concentration of pesticide. Additionally, 10 ml of autoclaved distilled water was also

added to soil samples in Petri plates to maintain 40% of water-holding capacity (Singh

et al., 2004; Hong et al., 2007). In order to compare bacterial degrading efficiency,

similar process was carried out with sterilized as well as unsterilized soil sample

(without inoculum) as control. Finally these soil samples were incubated at 37°C under

dark conditions to avoid photo oxidation of pesticides. Autoclaved distilled water was

47

added periodically to overcome the loss of water by evaporation (Hong et al., 2007;

Sasikala et al, 2012; Akbar and Sultan, 2016). The experiment was conducted in

triplicate to authenticate the results. For extraction of pesticide residues, 10 g of soil

was taken from all the petriplates separately on 0, 3, 6 and 9th day of incubation in a 50

ml universal falcon and shaken well with 10 ml of Ethyl acetate along with 5 g of

anhydrous Na2SO4 and then kept for 2 hrs. This mixture was then filtered through

Buchner funnel with cotton pad followed by centrifugation of filtrate at 3500 rpm for

20 minutes. The supernatant was microfiltered and 1 ml of filterate was taken in

eppendorfs and stored at -20°C for few days (if necessary otherwise used freshly) till

further analyses. For GC-MS analyses, 1ml of extract was mixed with analytical grade

methanol in 1: 9 ratio. Then 1.5 ml of this extract sample in methanol was syringe

filtered using Sartorius Ministart sterile syringe filters (.45 µm) and added to GC vials

being used for GCMS analyses.

2.14.5 GC-MS conditions used for the analyses of CPF, TCP, TAP and DM

biotransformation

GC-MS analyses of OP pesticides was done using QP5050 GC-MS equipped with a

DB-5 MS capillary column (30 m × 0.25 mm ×0.25 mm) and helium as carrier gas (100

ml/min) with Thermal Conductivity Detector (TCD) with splitless injection system.

The GCMS conditions used for the detection of intermediate products of CPF/TCP,

TAP and DM (formed during the degradation) are given in Table 2.31.

2.14.6 HPLC conditions used for the analyses of CPF, TCP, TAP and DM

degradation

The degradation of Chlorpyrifos, 3, 5, 6 Trichloropyridinol, Triazophos and

Dimethoate by four isolates MB490, MB497, MB498 and MB504 was also analyzed

using High-performance liquid chromatography (HPLC) LC-20AT equipped with a

UV-VISible detector (SPD-20A) and a C18 column (0.46 x 15 cm). For this purpose,

sample extract with ethyl acetate after being filtered by syringe filters (0.45 µm) was

directly analyzed by HPLC according to conditions given in Table 2.32 for each

pesticide respectively. Each experiment was conducted in triplicate. The HPLC

conditions used are given in Table 2.32. The percentage degradation of OP

pesticide/TCP was calculated using the equation:

48

B% = (Ca – Cb) ÷ Ca ×100 Where,

B = Pesticide degradation. Cb = the concentration of OP pesticide (mg/l) or (mg/kg)

residues left in the medium containing OP pesticide degrading microbial isolates. Ca =

the initial concentration of OP pesticide (mg/l) or (mg/kg) supplemented to the medium.

(Eissa et al., 2014).

Table. 2.31 The GCMS conditions used for the detection of metabolites of

Chlorpyrifos, Triazophos and Dimethoate.

GC-MS parameters OP pesticides

CPF TAP DM

Injection initial

temperature

Interface temperature

Control mode

Column inlet pressure

Column flow

Linear velocity

Split ratio

Total flow

Solvent cut time

Detector gain mode

Sampling rate

Injection volume

Electron impact (EI) using

SIM mode

Oven temperature program

Protocol followed with

modification

200◦C

250◦C

Split

60.6 KPa

1 ml/min

36.6 cm/sec

47

50 ml/min

2 min

Relative

0.50 sec

1 𝜇l

70 eV

Initially

maintained at

70∘C for 0 min

and then

increased to

250∘C at a rate

of 8∘C per min

and hold at

250∘C for 5

minutes.

(Reddy et al.,

2012)

200◦C

250◦C

Split

56.7 KPa

1 ml/min

36.5 cm/sec

48

50 ml/min

2 min

Relative

0.50 sec

1 𝜇l

70 eV

Initially

maintained at

70∘C for 0 min

and then

increased to

250∘C at a rate

of 10∘C per min

and hold at

250∘C for 5

minutes.

(Wang et al.,

2005)

220◦C

250◦C

Split

52.8 KPa

1 ml/min

36.3 cm/sec

48

50 ml/min

3 min

Relative

0.50 sec

1 𝜇l

70 eV

Initially

maintained at

50∘C for 1.5 min

and then

increased to

250∘C at a rate

of 10∘C per min

and hold at

250∘C for 5

minutes.

(Li et al., 2010)

49

During HPLC analysis, the concentration of OP pesticide (mg/l) or (mg/kg) was

calculated by comparing peak areas in the chromatogram of sample with that of

peak area of the standard chromatogram. Concentration of OP in sample (mg/kg) =

Peak area of chromatogram of sample ÷ Peak area of chromatogram of standard OP

compound × concentration of standard OP compound (Bishnoi et al., 2009).

Table 2.32 HPLC conditions used for the analyses of Chlorpyrifos, Triazophos and

Dimethoate degradation.

HPLC parameters OP pesticides

CPF TAP DM

Isocratic mobile phase

Detection wavelength

injection volume

flow rate

Oven temperature

Protocol followed with

modification

Acetonitrile and

water (70:30)

280 nm

10 μl

1.0 ml/ min

40°C

(Alvarenga et al.,

2015)

Acetonitrile and

water (80:20)

270 nm

10 μl

1.0 ml/ min

40°C

(Rani and Dhiraj,

2015)

Acetonitrile and

water (60:40)

221 nm

15 μl

1.0 ml/ min

30°C

(Bagyalakshmi et

al., 2011)

2.15 Optimization for OP pesticide biodegradation

2.15.1 pH Optimization for OP pesticide biodegradation

In order to study the effect of different pHs on the biodegradation of OP pesticides, 4

bacterial isolates and their consortium were grown in 30 ml M-9 broth supplemented

with 200 mg/l of OP pesticides at different pHs (6,7, and 8) in 100 ml flasks. 500 µl

inoculum was used for each isolate. All the flasks along with un-inoculated controls

were incubated in shaker incubator (150 rpm) at 37°C for 72 hrs. After incubation

period of 24, 48 and 72 hrs, 4 ml of sample was taken from each flask followed by the

method of extraction described earlier in section 2.15.1 and then OP pesticide

degradation in samples at different pH was analyzed using HPLC.

50

2.15.2 Temperature Optimization for OP pesticide biodegradation

The effect of different temperatures on the degradation of OP pesticides was studied by

growing four bacterial isolates (MB490, MB497, MB498 and MB504) and their

consortia in 30 ml of M-9 medium with pH 7 supplemented with 200 mg/l of OP

pesticide and 500µl of bacterial inoculum in 100 ml flasks at different temperatures (25,

30 and 37◦C) under static conditions for 72 hrs. For each temperature, control flask

without inoculum was also incubated. Extraction was carried out after 24, 48 and 72

hrs of incubation as described earlier in section 2.12.1. Degradation of OP pesticide in

each sample was analyzed using HPLC and % degradation was calculated using the

equation 2.6.

2.15.3 Effect of Shaking and Static conditions for OP pesticide biodegradation

The effect of shaking and static conditions on OP pesticide biodegradation was also

tested by growing four isolates (MB490, MB497, MB498 and MB504) and their

consortia in M-9 broth supplemented with different OP pesticides at 37°C and at pH 7

under shaking and static conditions.

2.16 Molecular Studies

For the identification of the isolates, 16S rRNA Gene Sequencing analysis was used.

2.16.1 DNA Extraction

For extracting the total genomic DNA from each isolate (MB490, MB497, MB498 and

MB504), a loop full of 24 hrs fresh bacterial culture was picked and mixed well with

1ml of autoclaved distilled water in an eppendorf. The cell suspension was centrifuged

at 13000 g for 2 minutes. The supernatant was discarded and pellet was washed twice

with 400 µl of STE buffer (Table 2.34). It was centrifuged at 13000 g for 2 minutes and

pellet was re-suspended in 200 µl of TE buffer (Table 2.35). Then, 100 µl of Tris

saturated phenol (Table 2.21) with pH 8 was added to this mixture and mixed well by

inverting gently for 1 minute. It was centrifuged at 13000 g for 5 minutes at 4◦C so as

to obtain 2 layers. Next, 160 µl of upper phase was picked and transferred to new

Eppendorf. 40 µl of TE buffer and 100 µl of chloroform was added to it and mixed well

by inverting gently until a white interface was observed between two layers. It was

51

centrifuged again at 13000 g for 5 minutes. The upper phase containing pure DNA was

shifted to another eppendorf and was confirmed by Gel electrophoresis.

2.16.2 Polymerase Chain Reaction (PCR)

The colony PCR was performed for the amplification of 16S rRNA gene. For this

purpose a 25 µl PCR reaction was established in the thermo cycler (96 universal

Gradient Peq Star, Peq Lab UK). Earlier, 0.4 µl each of forward and reverse primers

(0.5 µM), 12.5 µl of commercial master mix (Go Green Mastermix, Promega) and 8.7

µl PCR water were mixed together in a sterilized PCR tube. Then, 22 µl of this PCR

mix was taken in another sterile PCR tube for each isolate and 3µl of genomic DNA

was mixed with it to obtain final volume of 25 µl. All the reaction steps were done on

ice. Negative and positive controls were also incorporated. Go Green Mastermix

contains Taq DNA polymerase, dNTPs, MgCl2 and reaction buffers with best possible

concentrations. During agarose gel electrophoresis of PCR products, loading dye was

also integrated. Universal primers 27F and 1492R (Table 2.37) were used in the PCR

reaction to give a product size of nearly 1500 base pairs. The reaction conditions which

were used for the PCR are given in the Table 2.38. The blank without DNA was used

as control.

2.16.3 Gel Electrophoresis

Agarose gel (1%) prepared in 1X TBE Buffer (Table 2.33) and stained with ethidium

bromide was used to analyze PCR amplified samples. Gel was prepared by dissolving

0.5 g of Agarose (Oxoid) in 50 ml of 1X TBE Buffer after being heated in the

microwave up to boiling. It was cooled down to about 60˚C. Then a final concentration

of 0.5 µg/mL was achieved by adding 2.5 µl of ethidium bromide. Further, the gel was

poured vigilantly into the gel casting tray fitted with a comb and then it was solidified

in 20-30 min at room temperature. The combs were detached cautiously from the gel

and then, they were placed horizontally into the electrophoresis unit. For the running

medium, 1 X TBE buffer was used and poured into the electrophoresis chamber so that

to cover the gel. 3µl of each PCR amplified product was loaded in each well. In the first

well, 2 µL of 1 kb DNA ladder (Invitrogen) was loaded. The apparatus was run using a

current of 80 V for 30-45 min after the lid and power leads being placed on it. The

bubbles were given off the electrodes confirming the current flow. After giving enough

52

time, DNA bands were observed under UV light. Dolphin gel documentation system

(Wealtec, USA) was used to photograph DNA bands.

2.16.4. Reagents for DNA Extraction

In order to make 10X TBE buffer, 54 g of Tris HCl, 27.5 g boric acid and 4.65 g of

disodium EDTA were dissolved in distilled water and made up to a final volume of 1

L. It was stored at room temperature and diluted to 1X concentration before using for

agarose gel electrophoresis.

Table 2.33 10X TBE Buffer

Ingredients Amount

Autoclaved distilled water 1 L

Boric acid 27.5 g

Tris HCL 54 g

Disodium EDTA 4.65 g

Table 2.34 STE Buffer

Ingredients Amount

Autoclaved distilled water 200 ml

1 mM NaCl 0.011688 g

10 mM Tris HCL 0.03152 g

1 mM EDTA 0.074448 g

Table 2.35 TE Buffer

Ingredients Amount

Autoclaved distilled water 200 ml

10mM Tris HCL 0.03152 g

1mM EDTA 0.011688 g

53

Table 2.36 1M Tris-HCl

Ingredients Amount

Tris-HCl 157.64 g

Distilled water 1 L

pH 8.0

Table 2.37 Universal 16s RNA gene amplification primers

Primers Sequences

27F AGAGTTTGATCCTGGCTCAG

1492R TACGGCTACCTTGTTACGACTT

Table 2.38. PCR amplification conditions

2.17 Enzyme studies for Organophosphorus phosphatase (OPP) Enzyme

2.17.1 Screening for phosphate Solubilization potential of bacterial isolates

The four bacterial strains MB490, MB497, MB498 and MB504 were grown in National

Botanical Research Institute’s phosphate (NBRIP) medium (Table 2.12) containing

calcium phosphate as a substrate and supplemented with 1.5% Agar-agar for an agar

assay of phosphate solubilizing soil bacteria (Nautiyal, 1999). All four strains per plate

were stabbed with the help of sterile toothpicks in triplicate. After 10 days of incubation

at 37°C, the capability of the bacteria to solubilize insoluble phosphate was

demonstrated by the solubilization index (Premono et al., 1996) as given below.

Phosphate solubilization index (PSI) = (Colony diameter + Halo zone)/colony diameter

1 Cycle 33 Cycles 1 Cycle

Initial

Denaturation

Denaturation Annealing Extension Final extension

Temp.

(˚C)

Time

(Min)

Temp.

(˚C)

Time

(Sec)

Temp.

(˚C)

Time

(Sec)

Temp.

(˚C)

Time

(Min)

Temp.

(˚C)

Time

(Min)

94 5 94 40 56 45 72 1.8 72 6

54

2.17.2 Screening of Extracellular Organophosphorus phosphatase (OPP) and

Enzyme Assay

The fresh bacterial cultures (500 µl) of four isolates MB490, MB497, MB498 and

MB504 were grown seperately in 30 ml NBRIP broth to the late logarithmic phase for

three days. The cells were centrifuged at 3,000 rpm for 10 minutes and were harvested

and pelleted. The supernatant was used for the production of extracellular

Organophosphorus phosphatase (OPP) (acidic, neutral and alkaline) by using p-

nitrophenyl phosphate (p-NPP) as colorless substrate (Table 2.16), which turns into

yellow end product p-nitrophenol after hydrolysis by OPP to cleave the phosphate

linkage (Burns, 1982; Hernandez et al., 1995). In order to measure acidic, neutral and

alkaline OPP activity, buffer substrate mixtures were prepared by dissolving 0.6 mg of

p-nitrophenol phosphate /ml of respective buffer i.e for acidic OPP, 0.1M sodium

acetate buffer (Table 2.13) for neutral OPP, 0.1M sodium phosphate buffer (Table 2.15)

and for alkaline OPP, 0.1M Tris HCl buffer (Table 2.14) were used to dissolve

substrate. Then in 3 ml of crude enzyme extract, 1 ml of respective buffer substrate (p-

nitrophenyl phosphate) was added. The incubation of mixture was carried out for 20

min at 37°C. The reaction was terminated by adding 1ml of 1N NaOH (Table 2.18) to

increase pH of the reaction mixture. Both p-NPP and p-Np are colorless at acidic and

neutral pH but p-nitrophenol formed turns yellow at alkaline pH and was measured in

a UV-VIS Spectrophotometer (BMS UV-160) at 405 nm (Harishankar et al., 2013).

The amount of enzyme liberating 1nmol of p-nitrophenol per minute at 37°C is

equal to one unit (U) of OPP activity (Zhang et al., 2016). The concentration of OPP

produced was calculated using standard curve of p-Nitrophenol (p-NP, Sigma) (Table

2.17) with the help of serial dilutions of p-NP (1-10 µg/l).

55

Figure. 2.2 Standard curve for p-Nitrophenol.

2.17.3 Screening of intracellular Organophosphorus phosphatase (OPP)

The resuspension of cell pellet was made in 1ml of phosphate buffer with pH 7 and

silica/glass beads were used for disrupting cells. The centrifugation of the lysate was

carried out at 30,000 g at 4°C for 30 min to remove all cell debris and the

supernatant was used as enzyme source for intracellular OPP activity (Chaudhry et

al.,1988). The 3 ml of this supernatant enzyme extract was mixed with substrate buffer

mixture (1 ml) after various time intervals and incubated for 30 min. The reaction was

stopped by adding 1ml of 1N NaOH. All the experiments were performed in triplicates.

2.17.4 Factors effecting the production of Organophosphorus phosphatase (OPP)

Enzyme

2.17.4a Effect of pH on OPP production

The effect of pH on enzyme production was studied by inoculating24 hrs fresh bacterial

culture (50 µl) in 5 ml of NBRIP broth at pH range of 6, 7, 8, 9, 10 and 11 along with

substrate p-NPP and then incubated for 24 hrs. Initially, the pH of the medium was

adjusted by 1N HCl (Table 2.19) or 1N NaOH before autoclaving. After incubation, the

0.033

0.231

0.7920.893

1.021

1.2131.311

1.4321.53

1.65

2.025

2.52y = 0.1889x - 0.0069

R² = 0.9468

0

0.5

1

1.5

2

2.5

3

0 5 10 15

O.D

(4

05

nm

)

p-Nitrophenol (nmol)

56

sample cultures were centrifuged at 3000 rpm for 10 min and supernatant was used for

enzyme assay.

2.17.4b Effect of temperature on Organophosphorus phosphatase (OPP)

production

The effect of temperature on OPP enzyme production was checked by taking 5ml of

NBRIP broth in test tubes inoculated with 50 µl of 24 hrs fresh bacterial cultures and

then incubating at different temperatures (37, 45, 50 and 60°C) in a rotary shaker for

24 hrs. The enzyme was extracted by centrifugation at 3000 rpm for 10 min.

Supernatant was used to analyze the enzyme production by UV-VIS spectrophotometer

at 405 nm.

2.17.4c Effect of incubation time on Organophosphorus phosphatase (OPP)

enzyme production

In order to study the effect of incubation time on the production of acidic, neutral and

alkaline organophosphate phosphatase enzyme, the NBRIP broth inoculated with 24

hrs fresh bacterial cultures and supplemented with substrate buffer mixture was

incubated at 37°C in a rotary shaker at 120 rpm for 24, 48 and72 hrs. The enzyme was

extracted after incubation, centrifuged and supernatant was used for acidic, neutral and

alkaline organophosphate phosphatase enzyme assay as described above.

2.17.5 Factors affecting Organophosphorus phosphatase enzyme Activity

2.17.5a Effect of temperature on Organophosphorus phosphatase enzyme Activity

For temperature optimization, OPP activity, was assayed at 37, 45, 50, 60 and 70°C,

while pH was maintained according to the acidic, neutral or alkaline nature of enzyme.

The relative residual enzyme activity was measured using UV-VIS Spectrophotometer

at 405 nm as before immediately.

2.17.5b Effect of chemicals on Organophosphorus phosphatase enzyme Activity

The effect of different concentrations of SDS and EDTA on OPP activity was studied

using the supernatant enzyme extract mixed with substrate buffer and then adding SDS

and EDTA (2.5, 5 and 7.5%) separately to it. This enzyme reaction mixture was

57

incubated for 30 min at 37°C and then enzyme activity was assayed/analyzed using

UV-VIS Spectrophotometer at 405 nm. Control was without these chemicals.

2.17.5c Effect of metals on Organophosphorus phosphatase enzyme Activity

In order to study the effect of different metals on the activity of acidic, neutral and

alkaline OPP, 2.5% of stock solutions of different metals like ZnSO4, CuSO4 and CdCl2

(Table 2.27, 2.26 and 2.25, respectively) were added separately to enzyme substrate

mixture and was incubated at 37°C for 30 min while control was without metals. The

enzyme activity was analyzed using UV-VIS Spectrophotometer at 405 nm as

mentioned before.

2.17.5d Effect of Substrate concentration on Organophosphorus phosphatase

enzyme Activity

In order to study the effect of substrate concentration on the activity of OPP, different

substrate concentrations of 0.06, 0.6, 0.8 and 1.1% were used in the supernatant enzyme

extract of acidic, neutral and alkaline phosphatases and incubated for 30 min. Then

enzyme activity was observed and calculated with the help of UV-VIS

Spectrophotometer at 405nm.

2.17.5e Effect of incubation period on Organophosphorus phosphatase enzyme

Activity

Similarly enzyme activity at different incubation times of 30, 50, 70 and 90 minutes

was observed using p-NPP as substrate in the supernatant enzyme extract. The enzyme

activity determined with the help of UV-VIS Spectrophotometer at 405 nm.

2.17.6 Metal bioprecipitation by Organophosphorus phosphatase enzyme

The bioprecipitation potential of acidic, neutral and alkaline phosphatases was studied

for different heavy metals i.e Ni+2, Cr+6, Mn+2 and Cd+2. For this purpose, 30 ml NBRIP

broth was inoculated with 24 hrs fresh bacterial culture, and incubated for 3 days at

37°C at 150 rpm. Then, the bacterial culture broth was centrifuged at 8000 rpm for 10

minutes and supernatant (extracellular OPP phosphatase) was collected and used for

metal bioprecipitation study. The reaction mixture was prepared by using buffer

substrate mixture and extracellular phosphatase extract (supernatant) supplemented

58

with 1000 ppm of respective metal ion stock solutions of Ni+2 (NiCl2), Cr+6 (K2Cr2O7),

Mn+2 (MnCl2) and Cd+2 (CdCl2) (Table 2.20, 2.21, 2.22 and 2.23 respectively) at

different incubation times of 60 min, 120 min and 180 min. The amount of

bioprecipitated metal was measured by quantifying the reduction in the concentration

of respective metal content in the supernatant before and after incubation using Atomic

Absorption Spectrometer (AAS Shimadzu AA 7000), with the help of following

equation:

X = (A-B)/A×100 (Chaudhary et al., 2013)

Where

X = % of metal precipitation

A = Initial concentration of metal in the aliquot

B = Final concentration of metal in the aliquot i.e the supernatant.

2.17.7 Substrate specificity determination of Organophosphorus phosphatase

enzyme against OP pesticides (CPF, TAP and DM)

Finally the substrate specificity of OPP against different organophosphorus insecticides

(Chlorpyrifos, Dimethoate and Triazophos) was determined by measuring the OPP

activity against 50 mg/l of respective pesticides using HPLC and metabolites formed

were analyzed using GCMS (Liang et al., 2005; Gao et al., 2012).

59

RESULTS

Chapter 3

Isolation, characterization and screening of organophosphate

degrading soil bacterial isolates

Overview

Organophosphates are among the main group of pesticides being used extensively

worldwide to control pests as well as other domestic and veterinary insects. Their over

use has led to the contamination of soil, water and food resources. These become

harmful for non-target organisms by entering the food chain and causing

biomagnification. On the other hand, a large number of microbes have been reported

that are capable of degrading pesticides. Therefore present study was carried out to

isolate indigenous organophosphate (OP) degrading soil bacteria from different

agricultural soils of district Mianwali, Punjab, Pakistan. For this purpose, a large

number of bacterial isolates were obtained from the soil of three different wheat and

cotton fields at 6-12 inch depth. Diverse bacterial colonies were obtained by spread

plate method (105 CFU/g of soil). Out of them, fifteen bacterial isolates were selected

based on their distinct morphological characters and their tolerance against the OP

pesticides Chlorpyrifos (CPF), Triazophos (TAP) and Dimethoate (DM). After

purification on nutrient agar medium, they were named as MB490 to MB505. Among

them, four best isolates (MB490, MB497, MB498 and MB504) were selected for

further analyses. In case of CPF, MB497 was seen most tolerant as it could tolerate up

to 8 g/l of CPF, while MB490 and MB498 could tolerate up to 6 g/l and MB504 up to

0.8 g/l of CPF concentration. For TAP, MB490, MB497 and MB498 were tolerant up

to 4 g/l, while MB504 could grow up to 2 g/l of TAP. Similarly, for Dimethoate (DM),

MB504 was least tolerant up to 0.22 g/l, while MB490, MB497 and MB498 showed

growth up to 0.4 g/l of Dimethoate. MB504 showed the highest biodegradation of

63.56 % followed by MB490 with 63.19% .While MB497 and MB498 could degrade

41.57% and 37.97% respectively of Chlorpyrifos (0.8 g/l) after 72 hrs of incubation.

60

Isolate MB497 was Gram positive, while the other three were Gram negative. All were

rods and facultative anaerobes. All the isolates were mesophiles growing best at 30 to

42◦C. While on the basis of pH, these were characterized as neutrophile (MB490),

slightly acidophile (MB504), and moderately alkaliphile (MB497 and MB498). All of

the isolates showed multiple heavy metal resistance.The four isolates were positive for

important enzymes like Nitrate Reductase, Oxidase and Catalase etc. needed for

biodegradation. On the basis of all above mentioned facts, it can be suggested that these

isolated bacteria have good potential for degradation of OP pesticides and can be

utilized for bioremediation of OP pesticide contaminated soils.

Background

There are many reports that indigenous microbes from soil have potential for pesticide

degradation. The degradation of ipconazole (triazole fungicide) by soil

microorganisms (bacteria, Actinomycetes and fungi) was reported by Eizuka et al.

(2003). Similarly, the degradation of a chlorpyrifos contaminated soil using indigenous

mixed microorganisms in a slurry bioreactor at 3000 μg/g, 6000 μg/g and 12000 μg/g

was studied by Mohan et al. (2004) where they observed that 91, 82 and 14% of

chlorpyrifos was respectively degraded after 72 hrs. Singh (2008) reported the potential

of an isolated inborn fungal strain from corn field soil to degrade 44% of atrazine

within 20 days and utilize atrazine as a source of nitrogen. The isolation and

characterization of a fungal strain was mentioned by Yu et al. (2006) having potential

of degrading more than 80% of Chlorpyrifos and using it as a sole source of carbon

and energy. Mostly pesticides are applied simultaneously or in sequence for crop

protection, which results in a collective contamination of pesticide residues in the soil

environment (Diez, 2010). The biodegradation of lindane by a native bacterial

consortium was studied by Pesce and Wunderlin (2004) and they reported the

degradation of lindane by B. thiooxidans and S. Paucimobilis within 3 days of aerobic

incubation. Chu et al. 2008 investigated the impact of chlorothalonil on degradation of

chlorpyrifos and its effects on microbial populations in the soil. They found that

Chlorpyrifos degradation was not effected significantly by the presence of

chlorothalonil but there was increased inhibitory effect of chlorpyrifos on soil

microorganisms. Chirnside et al. (2007) reported the isolation of an indigenous

microbial consortium from polluted soils having a potential to degrade both atrazine

61

and alachlor. According to them, the consortium revealed a unique degradation pattern

in which atrazine degradation was dependent on alachlor degradation.

There have been many recent studies for isolation and characterization of various

Actinomycetes able to degrade pesticides (Benimeli et al., 2003). The lindane

biodegradation capability of Streptomyces sp. M7 in soil samples was studied by

Benimeli et al. (2008). They worked out effects of pesticide on maize plants grown in

lindane-polluted soil formerly inoculated with Streptomyces sp. M7. The researchers

found that Streptomyces sp. improved the biomass and concurrently reduced lindane

residues. According to them, the activity and growth of this strain was not inhibited by

natural soil microbial flora and by high pesticide concentration.

On the basis of above studies, current research was conducted to isolate, screen and

characterize the indigenous soil bacteria and to optimize different parameters for the

bacterial growth and OP pesticide biodegradation. The Detail of experimental results

with optimized parameters has been given in this chapter.

Results and Discussion

3.1 Soil Sampling and Physical and Chemical properties of Soil

For the isolation of OP degrading bacteria, soil sample collected was analyzed for its

physical and chemical properties as explained in section 2.2 of chapter 2 (Materials

and Methods). Soil pH was found to be 6.21, soil EC 143.3 µS/cm and soil organic

matter was 3.15%. Soil texture was to be sandy loam with 5% moisture. (Table 3.1).

Soils with EC below 0.4 mS/cm (400 µS/cm) are considered slightly or non-saline,

wheras soils having EC more than 0.8 mS/cm are considered highly saline (Wagh et

al., 2013). The agricultural soil used for bacterial isolation under current study was

found non-saline having EC value of 143.3µS/cm (0.1433 mS/cm).

62

Table 3.1 Physical and Chemical properties of Soil.

S. No. Physical and Chemical

properties of Soil

Remarks

1 pH 6.21

2 EC 143.3 µS/cm

3 Organic matter 3.15 %

4 Texture Sandy loam

5 Moisture 5%

3.2 Isolation and purification of soil bacteria

During present study, several colonies of bacterial isolates (106 to 107 CFU/g of soil)

were obtained through serial dilution and spread plate method from the soil of three

different organophosphate (OP) pesticide treated wheat and cotton fields at 6-12 inch

depth from district Mianwali. Among these diverse colonies, ultimately four bacterial

isolates were selected on the basis of their distinct morphological characters and the

maximum tolerance against the OP pesticides (Chlorpyrifos, Triazophos and

Dimethoate). They were purified on nutrient agar medium and were named as MB490,

MB497, MB498 and MB504 (Table 3.2).

Table 3.2 CFU/ml of Bacterial isolates

S. No. Bacterial isolates CFU/ml

1 MB490 9.4×107

2 MB497 3.25×106

3 MB498 3.92×107

4 MB504 1.63×107

3.3 Screening experiments against OP pesticides (Chlorpyrifos, Triazophos and

Dimethoate

The isolated bacteria MB490, MB497, MB498, and MB504 were grown on M-9

medium with and without glucose supplemented with various concentrations (0.04 g/l

to 8 g/l) of commercial samples of OP pesticides (CPF, TAP and DM). All four

63

isolates, (MB490, MB497, MB498, and MB504) showed very good growth even

without glucose, therefore indicating their ability to utilize pesticide as a sole source

of carbon. There have been several reports for biodegradation of many OP pesticides

(Zeinat et al. 2008; Megeed and El-Nakieb, 2008; Latifi et al., 2012).

3.3.1 Screening against Chlorpyrifos

Among the four isolates, MB497 was seen most tolerant as it could tolerate up to 8 g/l

of CPF. MB504 was the least tolerant (only up to 0.8 g/l), while MB490 and MB498

could tolerate up to 6 g/l of CPF (Table 3.3). Klebsiella sp. was capable of tolerating

upto 17.3 g/l of Chlorpyrifos (Ghanem et al., 2007). Current findings are comparable

to that reported by Ajaz et al. (2005) where Aeromonas, Pseudomonas and Klebsiella

could tolerate 4 g/l, 2 g/l and 8 g/l of CPF respectively.

3.3.2 Screening against Triazophos

When grown on M-9 medium supplemented with different concentrations of

triazophos, the isolates MB490, MB497 and MB498 were most tolerant up to 4 g/l of

TAP, while MB504 could grow up to 2 g/l of TAP (Table 3.3). Bacillus TAP1 strain

isolated from waste water sludge was reported to tolerate TAP up to 100 mg/l (Tang

and You, 2012). Similarly many other bacterial isolates capable of degrading TAP have

been reported like Ochrobactrum sp.mp-4 and Klebsiella sp. E6 (Dai et al., 2005;

Wang et al., 2005).

3.3.3 Screening against Dimethoate

Isolate MB504 was least tolerant for Dimethoate (up to 0.22 g/l), whereas the isolates

MB490, MB497 and MB498 showed growth up to 0.4 g/l of Dimethoate (Table3.3).

Megeed and El-Nakieb (2008) reported a microbial consortium that was able to tolerate

DM up to 120 mg/l. Raoultella sp. X1 was able to degrade DM co-metabolically

(Liang et al. 2009). Three bacterial isolates Brevundimonas sp., Bacillus sp. and

Klebsiella oxytoca were reported by Deshpande et al. (2004) to tolerate Dimethoate up

to 1000 mg/l. Similarly Proteus vulgaris and Bacillus licheniformis could degrade

Dimethoate up to 0.005 mg/ml and 3.5 mg/ml respectively (Debmandal et al., 2002

and 2005).

64

Table 3.3 Comparison of maximum tolerance of bacterial isolates against three

OP pesticides (CPF, TAP and DM).

Bacterial

Isolates

Maximum OP Pesticide tolerance (g/l)

CPF TAP DM

MB490 6 4 0.4

MB497 8 4 0.4

MB498 6 4 0.4

MB504 0.8 2 0.22

3.2.4 Screening experiments with Trichloropyridinol (TCP) (Major metabolite of

CPF)

The bacterial isolates (MB490, MB497, MB498 and MB504) were tested for their

tolerance to Trichloropyridinol (TCP) by growing them on M-9 medium supplemented

with different concentrations (14 and 28 mg/l) of TCP. All isolates exhibited very good

growth for both 14 mg/l and 28 mg/l of TCP. MB490 and MB504 appeared pale, while

MB497 and MB498 appeared as whitish and yellow growth respectively as given in

Table 3.4. TCP is known for its antimicrobial effects (Racke et al., 1988). So there are

only few reports of bacteria having ability to tolerate and degrade both CPF and its

metabolite TCP (Anwar et al., 2009; Liang et al., 2011; Lu et al. 2013). So, the bacterial

isolates MB490, MB497, MB498 and MB504 in the present study proved to be tolerant

for both CPF and TCP indicating their stronger bioremediation potential for CPF

contaminated sites.

Table 3.4. Effect of different concentrations of Trichloropyridinol (TCP) on

Growth and color of bacterial isolates grown on M-9 medium.

Bacterial

Isolates

Concentrations of Trichloropyridinol (mg/l)

14 28

MB490 ++P ++P

MB497 ++W ++W

MB498 ++ Y ++ Y

MB504 ++ P ++ P

65

++ = very good growth, P = Pale yellow, W = Whitish, Y= Yellowish

3.2 Morphological characterization of OP degrading bacterial isolates

The colonial shape, color, size, margins, texture and elevation were considered for the

morphological characterization of the bacterial isolates. The colony morphology was

examined both visually and microscopically. Isolates MB490, MB498, MB504 were

off-white, while MB497 produced white colored colonies. Colonies were circular with

smooth margins (MB490 and MB498), filamentous margins (MB497) and undulate

margins (MB504). All four isolates showed convex elevation and were having sticky

texture. The colony size ranged from 1mm to 4mm (Table 3.5).

All isolates (except MB497) were Gram negative and rod shaped (Bacilli). Rods were

arranged as single, pairs, chains or clusters. All isolated strains were non-capsulated

and non- spore forming (Table3.6).

Table 3.5 Colony morphology of Organophosphate degrading bacterial isolates.

Bacterial

Isolates

Visual

colour

Color

under

microscope

Shape

Size

(mm) Margin Elevation Texture

MB490 OW LY circular 1-1.5 Smooth convex Sticky

MB497 W GrB circular 2-4 Filamentous convex Sticky

MB498 OW GY circular 1-2 Smooth convex Sticky

MB504 OW GrB circular 1-2 Undulate convex Sticky

OW =off white, GY = golden yellow, GrB = grayish brown, LW=light yellow.

Table 3.6 Cell morphology of organophosphate degrading bacterial isolates.

Bacterial

Isolates

Gram

staining

Shape Arrangement Motility Capsule

staining

Spore

staining

MB490 G –ve Bacilli Pairs, groups - - -

MB497 G +ve Bacilli Pairs, chains + - -

MB498 G –ve Bacilli Pairs, groups - - -

MB504 G –ve Bacilli pairs, groups + - -

66

+꞊ Motile/Capsulated/Spore forming, _ = Non-motile/Non-capsulated/Non-spore

forming

3.3 Biochemical characterization

All isolates were facultative anaerobes as they were able to grow under aerobic as well

as anaerobic conditions (Table 3.7). Isolates MB490, MB498 and MB504 were able to

produce acid only under aerobic conditions thus indicating oxidative pathway, while

MB497 showed acid production under both aerobic and anaerobic conditions thus

indicating both oxidative and fermentive capability of the strains. No isolate could

show Gas production under both aerobic and anaerobic conditions.

All four isolates showed positive test for enzyme oxidase, catalase, and nitrate

reductase (Table 3.7). When other biochemical tests were performed, isolates MB490,

MB497, MB498 and MB504 showed negative test for Orthonitrophenyl- βD-

Galactopyranosidase, H2S production, Indole production, Mannitol

oxidation/fermentation, Inositol, D-Sorbitol, L-Rhamnose and Amygdalin tests.

MB497 and MB504 gave negative results for Lysine decarboxylase, Ornithine

decarboxylase, and Urease. Whereas all isolates were positive for Nitrate reduction

(NIT), Tryptophane deaminase (TDA), Citrate utilization (CIT) and Arginine

dihydrolase (ADH). Only MB490 was negative for Gelatinase (GEL) test, while

positive for Urease and Arabinose. In case of different sugars, all isolates were positive

for glucose, while all were negative for Mannitol, Inositol, Sorbitol, L-Rhamnose, and

Amygdalin sugars (Table.3.7).

When grown on MacConkey’s agar medium, only MB490 and MB498 exhibited

pinkish growth color and decolorization of the medium, thus showing lactose

fermentation, while MB504 showed transparent growth with non-lactose fermentation

(Table 3.7). While in case of Eosin Methylene Blue (EMB) agar medium, MB490,

MB498 and MB504 showed good growth on EMB agar medium with color variation,

thus supporting their gram negative character. The isolates MB490 and MB498 showed

light purple growth with pinkish spot and no decolorization of the medium thus

indicating weak lactose fermentation (Table 3.7) while MB497 and MB504 showed

non-lactose fermentation.

67

Table 3.7 Results of different biochemical tests

Biochemical tests Bacterial Isolates

MB490 MB497 MB498 MB504

Orthonitrophenyl-βD

Galactopyranosidase

- - - -

Arginine dihydrolase + + + +

Lysine decarboxylase + + - -

Ornithine decarboxylase + - - -

Citrate utilization + + + +

H2S production - - - -

Urease + - - -

Tryptophane deaminase + + + +

Indole production - - - -

Voges Proskauer - - - -

Gelatinase - + + +

Glucose + + + +

Mannitol - - - -

Inositol - - - -

Sorbitol - - - -

L-Rhamnose - - - -

Saccharose + - - +

Melibiose - - - +

Amygdalin - - - -

Arabinose + - - -

Nitrate reductase + + + +

Catalase + + + +

Oxidase + + + +

Oxidation Fermentation

Test (Aerobic)

+A +A +A +A

Oxidation Fermentation

Test (Anaerobic)

+ +A + +

68

Growth on MacConkey’s

Agar

+ pink - + pink +

transparent

Growth on EMB Agar + Light

Purple

+ pink + Light

Purple

+ Dark

purple

+ = Positive result, - = Negative result, A= Acid production

3.4 Physiological characterization

3.4.1 Screening Experiments against Heavy metals

Heavy metals are defined as group of metals having density more than 5 g/cm3. These

occur naturally as well as due to human activities as a contaminant in ecosystems. They

ultimately result in bioaccumulation in food chains. There have been many reports of

bacterial isolates capable of tolerating heavy metals (Castro-Silva et al. 2003; Silva et

al. 2007). All the four isolates could tolerate lead as [Pb(NO3)2] up to 2000 µg/ml,

showing good growth. The most tolerant to Manganese (MnCl2) was MB497 which

could grow up to 3000 µg/ml of Mn, while isolates MB504, MB498 and MB490 could

grow up to 1000, 500 and 300 µg/ml of Mn respectively (Table 3.8). On the other hand

all isolates were very sensitive to Cadmium (CdSO4). Isolates could grow in the

presence of different concentrations of Ni, Fe, Cu and Zn. Begum and Aundhati (2016)

reported Pseudomonas sp R2- KJ461965 that was capable of both heavy metal

tolerance and organophosphate degradation.

3.4.2 Screening Experiments against other Organic Pollutants

The isolates were checked for their tolerance against different organic pollutants other

than pesticides. All isolates MB490, MB497, MB498 and MB504 were able to tolerate

up to 5% of Benzene, Toluene, Xylene, Biphenyl and Naphthalene both with and

without glucose whereas they were very sensitive to Aniline showing no growth at all

concentrations tested (Table 3.9).

3.4.3 Effect of pH on the growth of bacterial isolates

All the isolates exhibited a wider pH range. At pH 7, MB490 showed its optimum

growth and thus was neutrophile. Isolates MB504 showed best growth at pH 6 thus

69

showing acidophilic character, while optimum pH for MB497 and MB498 was at pH

8 and pH 9 respectively. All the isolates were observed with decreasing trend of growth

towards alkaline pH 10 and 11 (Figure 3.1)

3.4.4 Temperature optimization for the growth of bacterial isolates

All isolates showed a wide temperature range for their growth. Isolates MB490 and

MB504 were growing optimally at 37◦C (Figure 3.2). While MB498 and MB497

showed their optimum growth at 30◦C and 42◦C respectively.

Table 3.8 Maximum tolerance of bacterial isolates against heavy metals.

Bacterial

isolates

Maximum Tolerance against heavy metals concentration(µg/ml)

NiCl2 FeCl3 CoCl2 K2Cr2O7 CuSO4 ZnSO4 PbNO3 CdSO4 MnCl2

MB 490 50 100 - - 100 - 2000 - 300

MB497 300 50 50 50 100 200 2000 - 3000

MB498 50 100 - 50 100 200 2000 - 500

MB504 100 50 - - 100 200 2000 - 1000

Table 3.9 Maximum tolerance of bacterial isolates against organic

pollutants/chemicals.

Bacterial

isolates

Maximum Tolerance against organic chemicals concentration (%)

Benzene Toluene Xylene Aniline Biphenyl Naphthalene

MB490 5 5 5 - 5 5

MB497 5 5 5 - 5 5

MB498 5 5 5 - 5 5

MB504 5 5 5 - 5 5

70

Figure 3.1 Effect of different pH on the growth of bacterial isolates MB490, MB497,

MB498 and MB504.

Figure 3.2 Effect of different temperatures on the growth of bacterial isolates MB490,

MB497, MB498 and MB504.

3.5 Quantitative analysis of Chlorpyrifos degradation by bacterial isolates

The four isolates were analyzed by UV-VIS Spectrophotometer for CPF degradation

potential within three days of incubation using initial CPF concentration of 0.8 g/l.

Isolates MB504 showed the highest biodegradation of CPF upto 63.56% followed by

MB490 with 63.19% CPF degradation after 72 hrs. While lowest degradation of

37.97% was found in MB498. MB497 could degrade 41.57% of Chlorpyrifos (Table

3.10).

All the isolates exhibited a prominent trend of increasing rate of biodegradation with

the passage of time from 24 hrs to 72 hrs (Figure 3.3 a, b, c & d). The Chlorpyrifos

showed maximum wave length of 299 nm before degradation. Pseudomonas putida

0.1

0.2

0.3

0.4

0.5

0.6

0.7

0.8

0.9

25◦C 30◦C 37◦C 42◦C

OD

60

0n

m

Temperature

MB490

MB497

MB498

MB504

0.1

0.3

0.5

0.7

0.9

1.1

1.3

1.5

5 6 7 8 9 10 11

pHs

MB490 MB497

MB498 MB504

71

could degrade CPF up to 76% when grown at initial CPF concentration of 1%

(Vijayalakshmi and Usha, 2012). Pseudomonas desmolyticum NCIM 2112 was

reported to degrade 98% of 10 mg/l in 6 days (Rokade and Mali, 2013).

Table 3.10 Percentage degradation of Chlorpyrifos (800 mg/l) by bacterial

isolates as analyzed by UV-VIS Spectrophotometer.

Bacterial

Isolates Percentage CPF degradation after

24 hrs 48 hrs 72 hrs

MB 490 11.91 40.73 63.19

MB497 19.16 23.79 41.56

MB498 24.92 24.29 37.97

MB504 28.95 53.27 63.56

The current study revealed an effective involvement of soil bacteria in the

bioremediation of Organophosphate pesticides by tolerating high OP concentrations

and also by their degradation capabilities to utilize these pesticides as a sole source of

carbon. Further studies were carried out in order to understand mechanisms of

biotransformation and to identify different metabolites of OP pesticides used, which

have been formed during biodegradation process.

72

a.

b.

c.

d.

Figure 3.3 Percent degradation of Chlorpyrifos (800 mg/l) by bacterial isolate a)

MB490. b) MB497. c) MB498. d) MB504 after 24, 48 and72 hrs of incubation, using

UV-VIS spectrophotometer.

Ab

sorb

ance

A

bso

rban

ce

Ab

sorb

ance

A

bso

rban

ce

73

3.6 Genetic Analyses

3.6.1 Extraction of genomic DNA

The 16S rRNA Gene Sequencing analysis was used to identify the isolated strains. The

DNA of each isolate was extracted using lysing buffer as given in methodology. Then

the extracted DNA was run through gel electrophoresis in order to confirm the presence

and size of DNA. Next, the DNA bands were observed under UV light and were

photographed using Dolphin gel documentation system (Wealtec, USA).

Figure 3.4 DNA bands of 1). MB490, 2). MB498, 3). MB497 and 4). MB504 in the

2nd, 3rd, 4th and 5th well respectively, along with DNA ladder in the 1st well.

3.6.2 PCR

The amplified PCR products were submitted to Macrogen for 16S rRNA sequence

based identification of MB490, MB497, MB498 and MB504.

3.6.3 Bacterial strain Identification

Selected isolates were sent for sequencing from Macrogen Inc., for characterization on

the basis of 16s rRNA analysis. Strains were identified as Pseudomonas kilonensis

MB490, Bacillus thuringiensis MB497, Pseudomonas kilonensis MB498 and

Pseudomonas sp MB504. Following 16s rRNA gene sequence accession numbers have

been assigned by NCBI to these four strains:

1 2 3 4 5

74

1. MG685888 for MB490 (Pseudomonas kilonensis)

2. KP886829 for MB497 (Bacillus thuringiensis)

3. MG685889 for MB498 (Pseudomonas kilonensis)

4. KP886830 for MB504 (Pseudomonas sp.)

The phylogenetic trees of MB490, MB497, MB498 and MB504 were made using

MEGA6 software. The evolutionary history of the four isolates was inferred using the

maximum likelihood method (Tamura et al. 2013) on the basis of maximum similarity

(figurrespectively).

Figure 3.5. Evolutionary relationships of Pseudomonas kilonensis MB490.

Figure 3.6 Evolutionary relationships of Bacillus thuringiensis sp. MB497.

Figure 3.7 Evolutionary relationships of Pseudomonas kilonensis MB498.

75

Figure 3.8. Evolutionary relationships of Pseudomonas sp. MB504.

Conclusion

Soil microflora has a great tendency for remediation of contaminated soils to protect

human health, environment and non-target organisms. The soil microbes are the most

eco-friendly, cost effective and reliable resources for decontamination of

organophosphate pesticides. In the present study, based on the morphological,

biochemical and physiological characterization, all the four isolates (Pseudomonas

kilonensis MB490, Bacillus thuringiensis sp. MB497, Pseudomonas kilonensis MB498

and Pseudomonas sp. MB504) were found facultative anaerobes, capable of growing

over a wide range of pH and temperature. Furthermore, all of them exhibited significant

tolerance and biodegradation of organophosphate pesticides. Therefore, these isolates

can be applied for detoxification of pesticide residues from agricultural soils in

Pakistan, due to well acclimatization to local environmental conditions (like

temperature and pH). Besides, enzymatic and metabolic studies would lead to more

efficient decontamination of pesticide pollutants from the environment

76

Chapter 4

Bacterial degradation/transformation of Chlorpyrifos and its

metabolite 3, 5, 6-Trichloropyridinol

Overview

Four isolates (MB490, MB497, MB498 and MB504) and their seven consortia (A, B,

C, D, E, F and G) were optimized for Chlorpyrifos (CPF) degradation (analyzed by

HPLC)/ growth at 600 nm (using UV-VIS Spectrophotometer) at different pHs (6, 7,

and 8) and at different incubation temperatures (25, 30 and 37◦C) for 24 hrs incubation.

These isolates and their consortia exhibited their potential to degrade CPF considerably

under wide range of pH and temperatures with 67.22 to 99.63% degradation of initial

200 mg/l of CPF. The MB490 showed best CPF degradation at pH 6, while MB497,

MB498 and MB504 were best at alkaline pH 8. The consortia B, C, and F were best at

pH 7, whereas consortia A, D, E and G were best CPF degraders at pH 8. The isolates

MB490 and MB498 exhibited maximum degradation of CPF at 37◦C as compared to

the MB497 and MB504, which were best at 30◦C. Consortia A, B, D and E showed best

CPF degradation at 37◦C while consortia C, F and G were outstanding at 30◦C. More

CPF degradation with less growth under static conditions was observed as compared to

shaking conditions. Degradation of CPF was studied in M-9 culture broth, soil slurry

and soil microcosm under different incubation periods. There was almost 99% CPF

degradation in M-9 broth, soil slurry and soil microcosm for most of the isolates and

consortia within 9 days. It was noticed that degradation was most rapid during first

three days of incubation followed by little increase in degradation up to 9 days. There

was a little bit enhanced CPF degradation and increased biomass in the presence of

TAP in the bacterial culture. There was negligible CPF degradation in the control

samples without bacterial inoculum. The bacterial isolates were also found capable of

degrading 3, 5, 6 Trichloropyridinol (28 mg/l) up to 90.57% within72 hrs. During

HPLC analysis, the retention times for CPF and TCP were determined to be 5.4 minutes

and 1.77 minutes respectively, while through GCMS, the detection time was 22 and

12.8 minutes respectively.

77

GCMS analysis of bacterial isolates in M-9 broth, soil slurry and soil microcosm at

different intervals revealed the formation of 3, 5, 6 Trichloropyridinol (TCP) along with

[(3, 5, 6-trichloro-2-pyridinyl) oxy] acetic acid, Diethyl thiophosphate (DETP),

phosphorothioic acid and DiIsopropyl methanephosphonate. TCP was further

metabolized into 1-methyl-2-pyrrolidine ethanol, 2-Ethoxy-4, 4, 5, 5-

tetramethyloxazoline, 3-(2, 4, 5-Trichlorophenoxy)-1-propyne and p-Propyl phenol,

thus proving that TCP was mineralized by bacterial isolates and consumed as a carbon

source of energy.

Background

The organophosphorus pesticides (OPs) are extremely toxic and used worldwide

including Pakistan for pest control. This extensive use has resulted in grave concerns

for food safety and environmental contamination. Many scientists have reported

different bacterial strains capable of degrading organophosphates e.g Pseudomonas

aeruginosa could degrade methamidophos, acephate, dimethoate, methyl parathion and

malathion (Ramu and Seetharaman, 2013), while Serratia sp. was capable of degrading

methidathion (Li et al., 2013). Similarly three soil bacterial isolates (Pseudomonas,

Agrobacterium and Bacillus) were able to degrade CPF (Maya et al., 2011). Isolate

Serratia marcescens could remove chlorpyrifos in addition to diazinon, fenitrothion,

and parathion (Cycon et al., 2013). Bacterial degradation of OPs, may be affected by

various factors like temperature, pH, soil moisture and also the period of pesticides

existence within soils before bacterial treatment. Singh and Walker (2006) reported that

more is the aging of Chlorpyrifos, greater is its persistence in soil. Xu et al. (2009)

revealed in a research work that Stenotrophomonas sp. PF32 could degrade up to 99%

each of fenthion, methyl parathion (in24 hrs), fenitrothion (in 18 hrs), phoxim (after

30 hrs), up to 97% of TAP and CPF (in 42 hrs and 48 hrs respectively) with initial

concentration of 100 mg/l each. It was reported that Stenotrophomonas maltophilia was

capable of degrading CPF, its metabolite TCP and DETP (Dubia and Fulekar, 2012).

Likewise, Stenotrophomonas sp. YC-1 had potential of completely removing CPF (100

mg/l) in24 hrs but could not consume its metabolite TCP (Yang et al., 2006). So, on the

basis of above mentioned studies, it was decided to isolate some indigenous soil

bacteria to test their potential to degrade Chlorpyrifos and also its major metabolite

Trichloropyridinol which is known to have antimicrobial effect. For this purpose four

78

bacterial strains (Pseudomonas kilonensis MB490, Bacillus thuringiensis MB497,

Pseudomonas kilonensis MB498 and Pseudomonas sp. MB504) and their seven

consortia (A, B, C, D, E, F, and G) were analyzed by HPLC for degradation studies

under different conditions and by GCMS to identify degradation products of CPF and

3, 5, 6 Trichloropyridinol (TCP). The results are discussed below in detail.


4.1 Optimization of environmental parameters (pH, temperature, shaking and

static incubation conditions) for CPF degradation

Four isolates (MB490, MB497, MB498 and MB504) and their seven consortia (A, B,

C, D, E, F, and G) were tested to optimize for Chlorpyrifos (CPF) degradation at

different pHs (6, 7, and 8) under shaking conditions and at different incubation

temperatures (25, 30 and 37◦C) under static conditions for 24 hrs. The growth of isolates

and consortia was also observed under different conditions at 600 nm using UV-VIS

Spectrophotometer. All isolates and their consortia exhibited significant degradation

under all conditions ranging from 67.22 to 99.50% as analyzed by HPLC.

4.1.1 Growth pattern and optimization for CPF degradation at different pHs by

bacterial isolates (MB490, MB497, MB498 and MB504) and their consortia (A, B,

C, D, E, F, and G)

The MB490 showed best CPF degradation (97%) at acidic pH (pH 6), while MB497,

MB498 and MB504 were best at alkaline pH (pH 8) with 99.39, 92.91 and 98.87%

degradation respectively (figure 4.1). All consortia showed excellent degradation

(83.74 to 99.5%) at all given pHs (figure 4.2). However, with the minor difference,

consortia B, C, and F were best at neutral pH i.e 7, while consortia A, D, E and G

exhibited maximum degradation under alkaline conditions (pH 8) indicating

involvement of alkaline phosphatases or some other similar degrading enzymes. The

bacterial growth (OD600 nm) monitored in all isolates and consortia, mostly indicated

an abrupt correlation with % degradation at the given pH range. In some cases, little

bacterial growth exhibited highest degradation of CPF (figure 4.1 & 4.2), indicating

79

involvement of all bacterial machinery and energy for the production of OP degrading

enzymes rather than cell division (Hett and Rubin, 2008).

On the other hand, only 2 to 9% CPF degradation was observed in the control at all

given pH and temperatures. Thus, a strong role of bacterial isolates in CPF degradation

was indicated. According to Singh et al. (2003), highest CPF degradation was exhibited

by an Enterobactor sp. at alkaline pH as compared to slower degradation at acidic pH.

Likewise, Anwar et al. (2009) reported greatest degradation of CPF (40 mg/l) at higher

pH of 8.5 by Bacillus pumilus strain C2A1. The results in the present study, where three

of the four isolates (MB497, MB498 and MB504) showed maximum CPF degradation

at pH 8 are in accordance with these previous studies, indicating that some vital

enzyme(s) involved in Chlorpyrifos degradation may have their optimum enzymatic

activity at high pH.

Abraham et al. (2014) reported the complete degradation of 300 mg/l of CPF in 24 hrs

by a consortium comprising of ten isolates in a laboratory scale fermenter which is

comparable to the present study. In another study by John et al. (2014), it was revealed

that the consortium C2 consisting of three isolates could degrade 25 ppm of CPF up to

95.38% in three days of treatment period. Complete degradation of 150 mg/l of CPF in

120 hrs was achieved by a consortium comprising 12 bacterial isolates (Pino and

Penuela, 2011). In the current study, the consortia consisted of only 2 isolates except

consortium G which was comprised of 4 isolates. Their response regarding CPF

degradation was very unique and extraordinary as they degraded more than 95% of

initial 200 mg/l CPF (except consortium G which degraded 83.74 to 98.43% CPF) at

all given pH after 24 hrs.

80

Figure 4.1. Growth and % degradation of CPF by four isolates (MB490, MB497,

MB498 and MB504) at different pHs after 24 hrs. Error bars represent standard errors

for values of three sample replicates.

Figure 4.2 Growth and % degradation of CPF by 7 consortia (A, B, C, D, E, F and) G)

at different pHs after 24 hrs. Error bars represent standard errors for values of three

sample replicates.

4.1.2 Effect of temperature on growth and CPF degradation by bacterial isolates

and their consortia

Bacterial isolates exhibited considerable CPF degradation ranging from 83.74-99.48%

at different temperatures (25, 30, 37◦C) with various growth trends. Isolates MB490 and

MB498 were found best CPF degraders at 37◦C, whereas MB497 and MB504 showed

maximum degradation at 30◦C (figure 4.3). Consortia exhibited significant CPF

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

pH6 pH7 pH8 pH6 pH7 pH8 pH6 pH7 pH8 pH6 pH7 pH8

% D

egra

dat

ion

OD

60

0n

m

% Degradation growth

MB490 MB497 MB498 MB504

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

6 7 8 6 7 8 6 7 8 6 7 8 6 7 8 6 7 8 6 7 8

A B C D E F G%

Deg

rad

atio

n

OD

60

0n

m

pHs% degradation Growth

81

degradation (90.88 to 99.63%) at different temperatures with optimum growth.

Consortia A, B, D and E showed best CPF degradation at 37◦C while consortia C, F and

G were outstanding at 30◦C (figure 4.4). In case of control, only 3. 6 and 9% CPF

degradation was observed at 25, 30, 37◦C respectively thus confirming the role of

bacterial isolates in pesticide degradation. According to Vijayalakshmi and Usha

(2012), strain Pseudomonas putida was able to achieve maximum (76%) degradation

of given 2% CPF at pH 7 and at 35°C. Singh et al. (2004) isolated Enterobacter sp.

from Australian soil that degraded Chlorpyrifos most rapidly at temperature 35°C. It

was reported that the isolated strains (Agrobacterium and Enterobacter spp.,

Pseudomonas spp. and Bacillus cereus) degraded Chlorpyrifos most efficiently at

optimal temperature of 30°C and pH 7 (Chishti and Arshad, 2012; Awad et al., 2011;

Liu, et al., 2012). Consortium C5 was able to degrade up to 90.02% CPF (125 ppm) at

optimum 30 ºC and pH 7 after 8 days of incubation (John et al., 2016).

Figure 4.3 Effect of temperature on growth and degradation of CPF by four isolates

(MB490, MB497, MB498 and MB504) after 24 hrs. Error bars represent standard errors

for values of three sample replicates.

10

30

50

70

90

110

0

0.5

1

1.5

2

25

°C

30

°C

37

°C

25

°C

30

°C

37

°C

25

°C

30

°C

37

°C

25

°C

30

°C

37

°C

MB 490 MB497 MB498 MB504%

Deg

rad

atio

n

OD

60

0n

m

Temperatures% Degradation growth

82

Figure 4.4 Effect of temperature on growth and degradation of CPF by 7 consortia

(A, B, C, D, E, F and) G) after 24 hrs. Error bars represent standard errors for values

of three sample replicates.

4.1.3 Effect of shaking versus static conditions on growth and CPF degradation by

four isolates (MB490, MB497, MB498 and MB504) and seven consortia (A, B, C,

D, E, F, and G) after 24 hrs of incubation

There was significant variation in biodegradation of CPF and growth of bacterial

isolates under shaking (aerobic) and static culture conditions after 24 hrs of incubation.

Generally there was little bit more CPF degradation by the bacterial isolates (96 to

99.36%) and their consortia (98.05 to 99.63%) but less growth under static than shaking

conditions, though more than 90% CPF degradation was noted under both conditions

(figures 4.5 & 4.6). These results are partly in contrast to Chishti and Arshad, (2012),

who revealed greater degradation of Chlorpyrifos (87−92%) under shaking/aeration

conditions, by four isolated strains after 18 days as compared to lesser 50−56%

degradation under static condition. However, according to them, strain SWLC2

exhibited highest degradation of Chlorpyrifos under shaking incubation which is in

accordance with present study. Similar to present study, they observed more bacterial

growth and higher abiotic degradation under shaking than under static conditions.

Pseudomonas putida was reported to have lesser degradation efficiency at higher speed

of shaking condition which was explained to be due to less contact between the

pesticide and the culture (Vijayalakshmi and Usha, 2012).

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

A B C D E F G

% D

egra

dat

ion

OD

60

0n

m

Temperatures

% degradation Growth

83

4.1.4 Biotransformation and degradation of Chlorpyrifos in the presence of TAP

by bacterial isolates and consortium G

Four bacterial isolates and consortium G (MB490+MB497+MB498+MB504) were

grown in M-9 broth supplemented with 200 mg/l of CPF and TAP each to analyze

quantitatively using HPLC, after 24 hrs of incubation under 120 rpm shaking at 37°C

and pH 7. All the isolates showed significant results regarding CPF degradation under

both conditions of CPF alone and in the presence of TAP. The CPF biodegradation and

growth of bacteria was little enhanced in the presence of TAP (figure 4.7). The reason

may be that bacteria were using both pesticides as their energy source increasing their

biomass and ultimately having more degrading enzymes to degrade both pesticides.

While there was only 2% CPF degradation in control under both condition of CPF alone

and CPF + TAP. There was 91.35 to 97.50 % CPF degradation by isolates and

consortium G after 24 hrs, when only 200 mg/l of CPF was supplied. Whereas in the

presence of both TAP and CPF, bacterial isolates and consortium G exhibited higher

(93 to 98%) CPF degradation (figure 4.7).

Figure. 4.5 Effect of shaking versus static conditions on growth and % degradation of

CPF by four isolates (MB490, MB497, MB498 and MB504) after 24 hrs. Error bars

represent standard errors for values of three sample replicates.

80

85

90

95

100

105

00.5

11.5

22.5

33.5

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4

Shaking conditions Static conditions

% D

egra

dat

ion

of

CP

F

OD

60

0n

m


84

Figure 4.6 Effect of shaking versus static conditions on growth and % degradation of

CPF by 7 consortia (A, B, C, D, E, F and G) after 24 hrs. Error bars represent standard

errors for values of three sample replicates.

Figure 4.7 Effect of presence of TAP on the degradation of Chlorpyrifos (CPF) and

growth (OD) by the 4 isolates (MB490, MB497, MB498 and MB504) along with the

consortium G (mixture of all four bacterial isolates) in M-9 broth after 24 hrs incubation

(initial concentration of CPF and TAP used = 200 mg/l). Error bars represent standard


There are other reports regarding degradation of mixtures of pesticides. Fuentes et al.

(2013) studied the degradation of mixture of CPF and Pentachlorophenol (PCP) by the

combined cultures and revealed that two immobilized consortia, first composed of 2

species of Streptomyces (AC5-AC7) and second consisting of six Streptomyces strains

80

85

90

95

100

105

0

0.5

1

1.5

2

2.5

3

3.5

4

A B C D E F G A B C D E F G

Shaking Conditions Static Conditions

% D

egra

dat

ion

of

CP

F

OD

60

0n

m

% Degradation Growth

80

85

90

95

100

0

0.5

1

1.5

2

2.5

3

3.5

4

MB490 MB497 MB498 MB504 Consortium G%

CP

F d

egra

dat

ion

OD

60

0n

m

% degradation of CPF alone % CPF degradation in presence of TAP

OD of isolates in presence of CPF alone OD of isolates in presence of CPF + TAP

85

(A2-A5-A11-M7-AC5-AC7) could degrade up to 48.64 and 42.90%, of 1.66 mg/l of

CPF and PCP respectively. Yanez-Ocampo et al. (2009) studied the consumption of 25

mg/l each of methyl-parathion (MP) and tetrachlorvinphos (TCF), by a bacterial co-

culture immobilized in three different ways (suspended, alginate immobilized and

tezontle-immobilized) and found 41, 72 and 66% removal for MP and 53, 65 and 47%

for TCF in three types of cultures respectively. A consortium consisting of many

bacterial species could degrade methyl parathion up to 72% and CPF up to 39% in a

combination of both pesticides with 150 mg/l each in 120 hrs, though it had completely

consumed each pesticide when used separately (Pino and Pinuela, 2011). The present

study is different and unique in the sense that both CPF and TAP in a mixture were

degraded independently to each other with very astonishing results, rather TAP

showing little enhancing effect on CPF degradation possibly due to fortification of

degrading microbial population.

4.2 Degradation of Chlorpyrifos by the bacterial isolates and their consortia in M-

9 broth

When bacterial isolates and their consortia (A, B, C, D, E, F and G) were grown in M-

9 broth supplemented with 200 mg/l of CPF and analyzed quantitatively using HPLC,

after 1, 3, 6 and 9 days of incubation, they all showed very good results regarding CPF

degradation and bacterial growth monitored by UV-VIS spectrophotometer (BMS UV-

160). Strains MB490, MB497, MB498 and MB504 consumed up to 99.90% of the

spiked CPF (200 mg/l) reaching maximum after 9 days of incubation (figure 4.8). These

results are very unique and excellent as compared to those reported by other researchers

especially with respect to time period required to achieve maximum degradation.

Chishti and Arshad (2012) reported 86, 81 and 89% degradation of Chlorpyrifos (100

mg/l) by the strains SGB2, SWLC1 and SWLH2 in 18 days, respectively. The bacterial

strain Pseudomonas WW5 was reported to degrade 94% of supplied CPF (400 mg/l) in

18 days (Farhan et al., 2012). Similarly two isolated strains of Streptomyces AC5 and

AC7 could degrade 90% of only 50 mg/l CPF after 24 hrs incubation (Briceno et al.,

2012). Feng et al. (2017) revealed more than 90% degradation of initial 20 mg/l CPF

in 12 days by a plant-derived bacterium Sphingomonas sp. Akbar and Sultan (2016)

reported 84.4 and 78.6% degradation of initial 100mg/l CPF in 10 days by the two

86

isolates. Hamsavathani et al. (2017) reported 30.78 to 82.06% CPF degradation of 0.5%

initial amount in 2 weeks by the 3 isolates. In the current study, CPF degradation rate

was most rapid during initial 3 days due to log phase of bacteria in accordance with the

previous reports (Hossain et al., 2015; Ishag et al., 2016).

Consortia A, B, C, D, E, F and G showed maximum (>99 %) CPF degradation from

day 3 to 9 as no significant increase was noticed. After 9 days, minimum (99.61%) CPF

degradation was exhibited by consortium A and maximum (99.95%) degradation by

consortium E was observed (figure 4.9). On contrary, much less degradation was

observed in the control with only 2, 3, 5 and 7% CPF degradation after 1, 3, 6 and 9

days of incubation respectively. Recently bioremediation research have been focused

more on using microbial consortia as they were considered to be more effective than

single culture methods due to their synergistic effects. So, it is obvious that ability of

consortia to metabolize CPF to use it as a nutrient for their growth, is a growth linked

process. Earlier, Singh et al. (2003) as well as Chishti and Arshad (2012) had reported

growth linked degradation of CPF by bacterial isolates but in the current study, the CPF

degradation efficiency of the respective consortia is extraordinary and excellent as

compared to previous studies. Despite, there are many reports of isolation of many

Chlorpyrifos degrading microorganisms from various polluted sources, yet there are

very few reports on consortial biodegradation of Chlorpyrifos (Fulekar and Geetha,

2008; Pino and Penuela, 2011; Sasikala et al., 2012). Sasikala et al. (2012) reported

70% degradation of 200 mg/l CPF by a bacterial co-culture composed of three bacterial

strains after 30 days of incubation. A consortium consisting of multiple microorganisms

could consume 150 mg/l of Chlorpyrifos in 120 hrs (Pino and Pinuela, 2011). The

consortium C5 consisting of 3 species could degrade up to 90% of 125 ppm

Chlorpyrifos within 8 days (John et al., 2016). Therefore, all the consortia in the current

study have best potential to remove Chlorpyrifos from contaminated sites and are most

suitable to be used for

Ivashina (1986) studied chlorpyrifos degradation by co-culture of Trichoderma sp. and

Bacillus sp. Shaker et al. (1988) reported 72-83% CPF degradation after 96 hrs by two

lactic acid bacterial strains. Chishti and Arshad (2012) studied the growth linked

biodegradation of chlorpyrifos by mixed culture (Agrobacterium and Enterobacter sp).

A co-culture consisting of a bacterium and a fungus could mineralize given CPF (100

87

mg/l) completely in 24 hrs (Xu et al., 2007). Likewise, Yang et al. (2005) revealed 40%

degradation of CPF (100 mg/l) within two days of incubation by an isolated bacterium.

Strain Paracoccus sp. demonstrated complete mineralization of CPF (50 mg/l) within

4 days (Xu et al., 2008). It was reported that a variety of bacterial isolates (Bacillus

subtilis, Bacillus cereus, Klebsiella, Brucella melitensis species, Pseudomonas sp.)

could degrade 46-72% of CPF in a culture broth within 20 days (Lakshmi et al., 2008).

Similarly, Ajaz et al. (2012) stated that Pseudomonas putida MAS-1 exhibited about

95% degradation of 200 mg/l CPF in 3 days in minimal medium.

In preset study, the isolates showed rapid degradation in minimal medium without

glucose because they have no other option of carbon source, thus these isolates had to

depend upon the Chlorpyrifos for their carbon and energy needs. Maximum growth

activity for all isolates and their consortia was noticed after first three days of incubation

which was followed by little decrease up to 9th day indicating either toxicity of

metabolites formed during CPF degradation thus killing the bacterial population or

scarcity of carbon source as almost all the CPF have been degraded (figure 4.8 and 4.9).

Moreover, the less availability of dissolved oxygen (DO), may be another reason for

lagging phase as increase in the organic load might result in the decreased DO

concentration (Corbitt, 1998). Moreover, the bacterial cells present in log phase during

rapid biodegradation achieved within initial three days also indicated that substrate

transformation would be at its maximum (Gray, 1989; Jilani, 2013). These results

indicated a strong association between bacterial growth activity and degradation

processes of CPF just like that reported by Chishti and Arshad (2012).

In the present study, 200 mg/l of CPF was almost completely mineralized in maximum

9 days, which indicated the incredible potential of 4 bacterial isolates and their consortia

to degrade and mineralize CPF and utilize resulting carbon and phosphorus for their

energy source. So, these isolates and their consortia may be considered as best

candidates to be applied for bioremediation of CPF contaminated places. Moreover,

these studies also revealed that these strains may have alkaline Phosphatase activity as

this enzyme is a phosphomonoestrase playing a very vital role in the CPF degradation

metabolism by hydrolysis of O-P bonds to give off phosphorus along with ethanol

which can be assimilated by bacteria (Singh and Walker, 2006).

88

Figure 4.8 Effect of incubation period on the degradation of Chlorpyrifos and growth

of the 4 isolates (MB490, MB497, MB498 and MB504) in the M-9 broth. Error bars


Figure 4.9 Effect of incubation period on the biodegradation of Chlorpyrifos (CPF) by

the 7 consortia (A, B, C, D, E, F and G) of bacterial isolates in the M-9 bacterial culture

broth (initial concentration of CPF used = 200 mg/l). Error bars represent standard


4.3 Degradation of Chlorpyrifos (CPF) by the bacterial isolates and their consortia

in the soil slurry

The bacterial isolates and their consortia were grown in soil slurry supplemented with

200 mg/l of CPF for up to 9 days and CPF degradation by them was analyzed using

HPLC after 3, 6 and 9 days of shaking incubation at pH 7 and 37°C. All the four

bacterial isolates showed excellent results of CPF degradation in soil slurry reaching

maximum degradation (>99%) during 9 days of incubation (figure 4.10). On the other

10

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hand, there was only 3, 5 and 7% CPF degradation in the control after 3, 6 and 9 days

of incubation respectively. Bacterial growth was monitored in terms of CFU/ml of soil

slurry after 3, 6 and 9 days. It was observed that bacteria were in log phase up to 3 days

followed by stationary phase up to 9th day due to lack of space and nutrients after almost

complete utilization of given pesticide (Table 4.1).

Similarly, all the consortia A, B, C, D, E, F and G exhibited outstanding results and

almost completely (>99%) removed given CPF in soil slurry after 9 days of incubation

(figure 4.11). Recently, soil slurry bioreactors (SB) have been used as an effective in

situ and ex situ technology that target the bioremediation of challenging sites, like those

having soils rich in clay and organic matter, having recalcitrant and toxic pollutants

exhibiting unusual behavior for degradation. These slurry bioreactors can achieve

enhanced and rapid treatment of polluted soils under controlled environmental

conditions assisted by biostimulation (nutrient supply of N, P and organic carbon

source), additional inocula (bioaugmentation), or by using surfactants or inducing

biosurfactant production inside the SB to increase the availability of pollutants (Robles-

Gonzalez et al., 2008). The CPF degradation results observed during present study are

much better than those given by Kumar (2011) who studied the Chlorpyrifos

degradation in soil slurry by four bacterial monocultures (RCC-2, GCC-1, GCC-3 and

JCC-3) and two bacterial co-cultures (GCE345 and GCC134). He reported monoculture

RCC-2 to be most effective having 21, 37, 54 and 77% degradation of 20 mg/l

Chlorpyrifos in 5, 10, 15 and 30 days of incubation respectively while the consortium

GCC134 was best with 24, 38, 56 and 85% Chlorpyrifos degradation in 5, 10, 15 and

30 days of incubation, respectively. Note that in his results, spiked amount of CPF was

only 20 mg/l and maximum degradation of 77 and 85% by monoculture and mixed

culture respectively was achieved in 30 days which is much longer period than in the

current study.

90

Figure 4.10 Effect of incubation period on percentage degradation of Chlorpyrifos

(CPF) by the 4 isolates (MB490, MB497, MB498 and MB504) in the soil slurry (initial

concentration of CPF used = 200 mg/l). Error bars represent standard errors for values


Figure 4.11. Effect of incubation period on percentage degradation of Chlorpyrifos

(CPF) by 7 consortia (A, B, C, D, E, F and G) in soil slurry (initial concentration of

CPF used = 200 mg/l). Error bars represent standard errors for values of three sample

replicates.

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91

Table 4.1 Bacterial Population in soil slurry spiked with 200 mg/l of CPF

Bacterial

isolates

CFU/ml of soil slurry


MB490 1.5×106 ± 0.1 7.2×106 ± 0.1 5.2×106 ± 0.1 3.2×106 ± 0.1

MB497 1.4×106 ±0.05 8.5×106 ± 0.1 6.5×106 ± 0.1 4.2×106 ± 0.1

MB498 1.7×106 ± 0.1 7.4×106 ± 0.05 4.4×106 ± 0.05 2.4×106 ± 0.05

MB504 1.6×106 ± 0.1 7.1×106 ± 0.05 4.1×106 ± 0.05 3.2×106 ± 0.05

Each value is the mean of three replicates; CFU = colony forming unit. ± values of

standard error.

4.4 Degradation of Chlorpyrifos (CPF) by the 4 isolates and the 7 consortia in the

soil microcosm

The bacterial isolates and their consortia were inoculated in soil microcosm in Petri

plates having 50 g of soil with 40% water holding capacity supplemented with 200

mg/kg of CPF and incubated under dark conditions for up to 9 days. CPF

biodegradation was analyzed using HPLC after 3, 6 and 9 days of incubation. All the

bacterial isolates showed a gradually increasing trend of CPF degradation with

increasing period of incubation in soil microcosm ranging from minimum (68.5%) after

3 days to maximum (99% or above) after 9 days of incubation (figure 4.12). Moreover,

the CPF degradation was most rapid during initial 3 days of incubation, then it slowed

down gradually. It was observed that after 9 days, among the four isolates, MB490

exhibited highest CPF degradation (99.8%). Bacterial population of each isolate in the

soil microcosm was monitored at different intervals of 0, 3, 6 and 9 days by counting

CFU through serial dilutions and spread plate method as given in Table 4.2. It was

observed that CPF degradation was growth linked as microbial population also

increased most rapidly during first 3 days, then it slowed down and ultimately decreased

until 9th day, probably due to lack of nutrition.

In general, the CPF degradation rates by the bacterial isolates and consortia followed a

biphasic model with an initial faster rate in the first phase of degradation followed by a

second phase at a slower rate. There are previous reports regarding biphasic

92

biodegradation for many other pesticides (Rigas et al., 2007; Shaer et al., 2013;

Abdurruhman et al., 2015; Ishag et al., 2016).

On the other hand, only 10, 12 and 15% CPF degradation was observed in the control

unsterilized soil and 3, 5 and 7% CPF degradation in the control sterilized soil without

any bacteria after 3, 6 and 9 days of incubation respectively. Three bacterial strains

Serratia liquefaciens, Serratia marcescens and Pseudomonas sp. and their consortium

exhibited efficient degradation of 100 mg/kg soil of diazinon in sterilized soil with a

rate constant of 0.032–0.085/day, while half-life for diazinon was observed from 11.5

to 24.5 day (Cycon et al., 2009). Earlier, it had been revealed that high organic matter

content in soil resulted in reduced bioavailability of organophosphorus pesticides and

their degradation (Karpouzas and Walker, 2000), but on the opposite, Singh et al.

(2006) proved that the organic matter had no major effect on fenamiphos and

chlorpyrifos removal by bacteria. Farhan et al. (2017) revealed 69, 60 and 45%

degradation of CPF at different concentrations of 40, 60 and 80 mg/Kg respectively by

Pseudomonas sp. in soil after 21 days. They also reported bacterial lag phase at higher

CPF concentration of 60-80 mg/Kg.

Surprisingly, all the consortia exhibited exceptional results for CPF degradation in soil

microcosm with minimum 98.60% CPF degradation (consortium A) to maximum

99.99% (consortium B) degradation after 9 days of incubation (figure 4.13).

93

Figure 4.12. Effect of incubation period on degradation of Chlorpyrifos (CPF) by the

4 isolates (MB490, MB497, MB498 and MB504) in the soil microcosm (initial

concentration of CPF used=200 mg/kg). Error bars represent standard errors for values


Table 4.2 Bacterial Population in soil microcosm spiked with CPF (200mg/kg)

Bacterial

isolates

CFU/g soil


MB490 1.8×106 ± 0.1 7.5×106 ± 0.05 4.2×106 ± 0.1 2.4×106 ± 0.1

MB497 1.2×106 ± 0.05 8.5×106 ± 0.1 5.1×106 ± 0.05 3.3×106 ± 0.05

MB498 1.5×106 ± 0.1 8.4×106 ± 0.05 4.3×106 ± 0.1 3.2×106 ± 0.1

MB504 1.4×106 ± 0.05 9.6×106 ± 0.1 4.5×106 ± 0.05 2.4×106 ± 0.05


standard error

0

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40

60

80

100

120

MB490 MB497 MB498 MB504 Control soil Unsterilizedsoil

% D

egra

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F

Incubation time


94

Figure 4.13 Effect of incubation period on degradation of CPF by 7 consortia (A, B,

C, D, E, F and G) in soil microcosm (initial concentration of CPF used=200 mg/kg).

Error bars represent standard errors for values of three sample replicates.

4.5 Degradation and transformation experiments with 3, 5, 6-trichloropyridinol

(TCP)

Compound 3, 5, 6-trichloropyridinol (TCP) is one of the main metabolites of CPF. It is

considered recalcitrant and resistant to microbial degradation due to antimicrobial

effects (Racke et al., 1988). Only few bacteria were reported earlier capable of

degrading CPF as well as TCP (Liang et al., 2011; Lu et al. 2013). So, in the present

study, four bacterial isolates (MB490, MB497, MB498 and MB504) were tested for

their potential to degrade TCP. Results are discussed below.

4.5.1 Effect of incubation period and concentration of TCP on the growth (OD600)

and degradation of Trichloropyridinol (TCP) by the 4 isolates in the bacterial

culture broth

When TCP concentration used was 14 mg/l, all the four bacterial isolates (MB490,

MB497, MB498 and MB504) showed excellent results for TCP degradation with

maximum degradation achieved within 72 hrs of incubation. Strain MB490 was most

efficient as it exhibited maximum (98%) degradation of TCP followed by MB504

(97.90%), MB497 (96.20%) and MB498 (93.80%) after 72 hrs with least growth (figure

4.14). There was only 2, 3 and 5% TCP degradation in control without inoculum. All

results are mean values of three replicates.

0

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95

On the other hand, when initial concentration of TCP was increased to 28 mg/l, all the

isolates exhibited gradual increase in TCP degradation with increasing incubation time

from 24 hrs to 72 hrs. Strain MB497 exhibited highest TCP degradation of 90.57%,

followed by MB504 (83.07%), while isolates MB490 and MB498 could degrade up to

67.53 and 60.39% TCP after 72 hrs (4.15). In contrast, they showed gradual decreasing

trend in their growth from 24 hrs to 72 hrs. The reason for highest growth and lowest

degradation during 24 hrs may be that bacteria were growing rapidly to produce more

enzymes for the degradation of higher concentration of TCP. During maximum

degradation, bacterial growth decreased rapidly due to different factors like

accumulation of toxic metabolites of TCP or some other toxic secondary secretion of

bacteria themselves. The TCP utilization by all the 4 strains was confirmed by HPLC

analysis (figure 4.38a, b, c, d and e).

Previously, it has been suggested that CPF biodegradation is resisted due to the

accumulation of its main metabolite 3, 5, 6-trichloro-2-pyridinol (TCP). TCP had been

declared as a persistent and leachable pollutant by the US Environmental Protection

Agency (Armbrust, 2001). It has somewhat higher antimicrobial effects on microbes,

which prevents its own degradation (Caceres et al., 2007; Cao et al., 2012). The

resistance of TCP towards degradation might be due to the presence of three chlorine

atoms in the pyridinol ring of TCP structure which must be eliminated in order to break

the ring (Feng et al., 1997).

There are only few reports of TCP degradation due to its antimicrobial effects. Present

results are in consistence with those reported by Chen et al. (2012), which revealed

89% degradation of CPF and 93.5% degradation of TCP (50 mg/l each) by fungal strain

Cladosporium cladosporioides Hu-01 within 1 day and their complete disappearance

after 5 and 6 days of incubation respectively.

A bacterial strain, Cupriavidus sp. DT-1, was also reported by Lu et al. (2013) that was

able to degrade both Chlorpyrifos (100 mg/kg) and 3,5,6-trichloro-2-pyridinol (50

mg/kg) up to 100 and 94% respectively in 30 days of incubation in soil. The strain

Bacillus pumilus C2A1 was reported to degrade up to 90% of initial 300 mg/l of TCP

after 8 days of incubation and up to 73, 83 and 87% of the spiked Chlorpyrifos (100,

200 and 300 mg/l respectively) in broth within 10 days of incubation (Anwar et al.,

96

2009). Similarly, Yang et al. (2005) reported degradation of both Chlorpyrifos and TCP

(100 mg/l) by Alcaligenes faecalis. It was reported that Pseudomonas sp. could degrade

only TCP but not Chlorpyrifos in broth culture (Feng et al., 1997). Similarly, three

isolated strains (Xanthomonassp. 4R3-M1, Pseudomonas sp. 4H1-M3,

and Rhizobium sp. 4H1-M1) were capable of degrading 10 mg/l of both CPF and its

primary metabolite, 3, 5, 6-trichloro-2-pyridinol (Rayu et al., 2017). Nevertheless, in

the current studies, both CPF and TCP were degraded by the four isolates which is an

exceptional outcome.

Figure 4.14. Effect of incubation period on growth (OD600) and degradation of 3, 5, 6-

Trichloropyridinol (TCP) by the 4 isolates (MB490, MB497, MB498, MB504) in the

M-9 broth as analyzed by HPLC (initial concentration of TCP used = 14 mg/l). Error


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Figure 4.15 Effect of incubation period on growth (OD600) and degradation of 3, 5, 6-

Trichloropyridinol (TCP) by the 4 isolates (MB490, MB497, MB498, MB504) in the

M-9 broth as analyzed by HPLC (initial concentration of TCP used= 28 mg/l). Error


4.6 Quantitative analysis through HPLC to determine biodegradation

of Chlorpyrifos (CPF) and its metabolite 3, 5, 6-Trichloropyridinol

(TCP)

4.6.1 Effect of pH on CPF degradation by isolates and consortia

4.6.1a Effect of pH on CPF degradation by isolate MB490

After HPLC analysis, it was observed that in case of MB490, there was a very sharp

decrease in peak area at pH 6 (figure 4.16b) as compared to control (figure 4.16a),

showing highest CPF degradation. While there was less decrease in peak area (figure

4.16d) and lowest degradation at pH 8 after 24 hrs of incubation. At pH 7, there was

considerable decrease in peak area of CPF (RT= 5.4 min), thus indicating considerable

degradation as given in (figure 4.16c). There are some new peaks appearing at retention

time of 2.2 min at pH 7 and pH 8.

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a. b.

c. d.

Figure. 4.16 Effect of pH on degradation of CPF (RT= 5.4 min) by MB490. (a)


4.6.1b Effect of pH on CPF degradation by isolate MB497

It was observed that at pH 8, MB497 showed much reduction in CPF peak area (figure.

4.17d) with maximum CPF degradation, while, it exhibited almost equally decreased

CPF peaks at pH 6 and pH 7, indicting considerable but equal degradation of CPF at

both pH 6 and pH 7 as shown in figure. 4.17c & d.

Retention time (min)

Ab

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Retention time (min) Retention time (min)

99

a. b.

c. d.

Figure. 4.17 Effect of pH on degradation of CPF (RT= 5.4 min) by MB497. (a)


4.6.1c Effect of pH on CPF degradation by isolate MB498

In case of MB498, maximum reduction in CPF peak area was observed at pH 8 (figure

4.18d) thus indicating highest CPF degradation, whereas least reduction of peak area

and thus least degradation of CPF was observed at pH 6 (figure 4.18b). While, at pH 7,

reduction in peak area was moderate showing considerable degradation as shown in

figure 4.18c.


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a. b.

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Figure. 4.18 Effect of pH on degradation of CPF (RT= 5.4) by MB498. (a) Control,

(b) MB498 at pH 6 (c) pH 7 (d) pH 8.

4.6.1d Effect of pH on CPF degradation by isolate MB504

Strain MB504 showed highest reduction in CPF peak area (figure 4.19d) and thus

highest degradation at pH 8, followed by considerable decrease in peak area at pH 6

and then at pH 7, respectively. It seemed as pH had no significant effect on CPF

degradation in case of MB504 (figure 4.19). It exhibited excellent degradation of CPF

at all given pH range.



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a. b.

c. d.

Figure. 4.19 Effect of pH on degradation of CPF (RT= 5.4) by MB504. (a) Control,

(b) MB504 at pH 6, (c) pH 7, (d) pH 8.

4.6.1e Effect of pH on CPF degradation by consortium A

Consortium A, exhibited sharpest decrease in CPF peak area (RT = 5.4 min) and thus

greatest degradation at pH 8 along with new prominent peaks depicting some unknown

metabolites at different retention times (4.20b). It showed considerable reduction in

peak area at both pH 6 and pH 7 with very noticeable CPF degradation as illustrated in

figure 4.20b & c respectively.

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a. b.

c. d.

Figure. 4.20 Effect of pH on degradation of CPF (RT= 5.4 min) by Consortium A. (a)

Control, (b) Consortium A at pH 6 (c) pH 7 (d) pH 8.

4.6.1e Effect of pH on CPF degradation by consortium B

In case of consortium B, it was observed that very considerable reduction in peak area

of CPF occurred at all three pH (6, 7, 8) respectively with almost equal CPF degradation

as shown in figure 4.21b, c & d. Many new peaks can also be seen indicating metabolite

formation during CPF transformation reactions, where peak at retention time of 1.7 min

is present at all pH 6, 7 and 8 indicating formation of 3, 5, 6-Trichloropyridinol (TCP).

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a. b.

c. d

Figure. 4.21 Effect of pH on degradation of CPF (RT= 5.4 min) by consortium B. (a)

Control, (b) Consortium B at pH 6 (c) pH 7 (d) pH 8.

4.6.1f Effect of pH on CPF degradation by consortium C

In case of consortium C, CPF peak almost diminished at pH 7 (figure 4.22 c) showing

nearly complete degradation of CPF while at pH 6 and pH 8 it showed remarkable

decrease in peak area depicting excellent degradation as illustrated in figure 4.22 b & d

respectively. At all pH range, many secondary peaks of metabolites at different

retention times can be observed including TCP at RT of 1.7 min. The consortium C

proved very efficient degrader of CPF at the given pH range of 6, 7 and 8.

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a. b.

c. d.

Figure. 4.22 Effect of pH on degradation of CPF (RT= 5.4 min) by consortium C. (a)

Control, (b) Consortium C, at pH 6 (c) pH 7 (d) pH 8.

4.6.1g Effect of pH on CPF degradation by consortium D

The consortium D showed greatest decrease in CPF peak area at pH 8 depicting highest

degradation. There was very noticeable reduction in peak area at pH 6 and pH 7 (figure

4.23b and c respectively). The peak for 3, 5, 6-Trichloropyridinol is prominent in figure

4.23b & d with RT of 1.7 min. So, the consortium D proved excellent degrader of CPF

at wide pH range.

Ab

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a. b.

c. d.

Figure. 4.23 Effect of pH on degradation of CPF (RT= 5.4 min) by Consortium D. (a)

Control, (b) Consortium D, at pH 6 (c) pH 7 (d) pH 8.

4.6.1h Effect of pH on CPF degradation by consortium E

Likewise in case of consortium E, highest reduction in peak area was observed at pH 8.

While at pH 6 and pH 7, there was very prominent decrease in peak area for CPF

showing outstanding results for CPF degradation as illustrated in figure 4.24b & c

respectively. The metabolites are indicated at different retention times in figure 4.24 b,

c & d.

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a. b.

c. d.

Figure. 4.24 Effect of pH on degradation of CPF (RT= 5.4 min) by consortium E. (a)

Control, (b) Consortium E, at pH 6 (c) pH 7 (d) pH 8.

4.6.1i Effect of pH on CPF degradation by consortium F

Consortium F exhibited much reduction in peak area for CPF (RT = 5.4 min) at pH 7

(figure 4.25 c). Whereas at pH 6 and pH 8, peak area decreased almost equally but

significantly, indicating tremendous CPF degradation (figure 4.25b and d respectively).

Different peaks for metabolites appeared at all pH, where TCP is identifiable at RT =

1.7 min.

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a. b.

c. d.

Figure. 4.25 Effect of pH on degradation of CPF (RT= 5.4 min) by Consortium F. (a)

Control, (b) Consortium F, at pH 6 (c) pH 7 (d) pH 8. TCP peak is prominent (RT= 1.7

min).

4.6.1j Effect of pH on CPF degradation by consortium G

In case of consortium G (MB490+MB497+MB498+MB504), it was observed that

highest decrease in CPF peak area occurred at pH 8 (figure 4.26 d). There was least

reduction in peak area at pH 6, while at pH 7, considerable decrease in peak area was

noticed as exhibited in figure 4.26 b & c. Different new peaks for metabolites are

prominent in figure 4.26c & d).

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a. b.

c. d.

Figure. 4.26 Effect of pH on degradation of CPF (RT= 5.4 min) by Consortium G. (a)

Control, (b) Consortium G, at pH 6 (c) pH 7 (d) pH 8.

4.6.2 Effect of temperature on CPF degradation by isolates and

consortia

4.6.2a Effect of temperature on CPF degradation by isolate MB490

HPLC analysis was conducted to determine the effect of temperature on CPF

degradation by the bacterial isolates and their consortia. Isolate MB490, exhibited

maximum reduction of CPF peak at 37◦C and also highest CPF degradation. While

minimum peak reduction with least CPF degradation at 25◦C and considerable peak

reduction at 30◦C was observed (figure 4.27 b & c). There was prominent peak for

metabolite TCP (RT = 1.77 ± 0.2 min) at 25◦C and 37◦C, though it was also present at

30◦C as a very low peak.

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a. b.

c. d.

Figure. 4.27 Effect of temperature on degradation of CPF (RT= 5.4 min) by MB490.

(a) Control, (b) MB490, at 25°C(c) 30°C (d) 37°C.

4.6.2b Effect of temperature on CPF degradation by isolate MB497

Strain MB497 showed maximum peak reduction of CPF (RT = 5.4) at 30◦C (figure

4.28c). There was also considerable reduction in CPF peak at 25◦C and 37◦C (figure

4.28 b and d). TCP peak (RT = 1.77 min) was conspicuous at 30◦C and 37◦C while at

25◦C, it was diminished may be due to its utilization by the MB497 as energy source.

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a. b.

c. d.

Figure. 4.28 Effect of temperature on degradation of CPF (RT= 5.4 min) by MB497.


4.6.2c Effect of temperature on CPF degradation by isolate MB498

In case of MB498, CPF peak was almost completely diminished at 37◦C with a single

and prominent peak for metabolite TCP having RT = 1.7 min (figure 4.29 c). There was

considerable reduction in peak at 25 and 30◦C (figure 4.29 b and c).

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a. b.

c. d.

Figure. 4.29 Effect of temperature on degradation of CPF (RT= 5.4) by MB498. (a)

Control, (b) MB498, at 25°C(c) 30°C (d) 37°C. A single and prominent peak of TCP

can be noted at 1.77 min at 37°C.

4.6.2d Effect of temperature on CPF degradation by isolate MB504.

Strain MB504 exhibited very outstanding reduction in CPF peak at 30◦C, though there

was considerable decrease in peak at 25◦C and 37◦C as illustrated in figure 4.30 b & d

respectively. Many different peaks also appeared at all three temperatures including

TCP peak (RT= 1.7±0.2 min).

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a. b.

c. d.

Figure. 4.30 Effect of temperature on degradation of CPF (RT = 5.4 min) by MB504.


4.6.2e Effect of temperature on CPF degradation by consortium A

The consortium A, exhibited almost diminished CPF peak at 37◦C (figure 4.31 d) while

least reduction in peak occurred at 25◦C (4.31 b). There was very noticeable decrease

in peak area at 30◦C (4.31 c). There appeared a new prominent peak at RT = 7.1, 7.2

and 6.75 min at all the temperatures (25, 30, 37◦C), indicating formation of some new

metabolite by the consortium A. In figure 4.31 d at 37◦C, a small peak for TCP at RT =

1.77 min was noticed and its absence at other temperatures could be due to its rapid

mineralization by the bacteria in the culture as has been confirmed in experiments with

TCP degradation by the HPLC chromatograms given in figure 4.38.

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a. b.

c. d.

Figure. 4.31 Effect of temperature on degradation of CPF (RT= 5.4) by consortium

A. (a) Control, (b) consortium A, at 25°C (c) 30°C (d) 37°C. Note peak for TCP at RT

= 1.77 min.

4.6.2f Effect of temperature on CPF degradation by consortium B

In case of consortium B, there was almost complete reduction of CPF peak area

indicating a remarkable CPF degradation at 25, 30 and 37◦C along with appearance of

new and prominent peaks with retention time above 6 and 7 min, depicting some

unknown metabolites. It seemed as temperature had no effect on CPF degradation by

consortium B (figure 4.32b, c and d).

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a. b.

c. d.

Figure. 4.32 Effect of temperature on degradation of CPF (RT = 5.4 min) by

consortium B. (a) Control, (b) consortium B, at 25°C(c) 30°C (d) 37°C.

4.6.2g Effect of temperature on CPF degradation by consortium C

Likewise, the consortium C also followed the same trend as consortium B and exhibited

nearly complete reduction of CPF peak area along with appearance of same new

metabolites at all three temperatures (25, 30 and 37◦C) as illustrated in figure 4.33 b, c

& d respectively.

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a. b.

c. d.


consortium C. (a) Control, (b) consortium C, at 25°C(c) 30°C (d) 37°C.

4.6.2h Effect of temperature on CPF degradation by consortium D

Consortium D showed maximum reduction in CPF peak area at 37◦C, followed by

decrease in peak area at 25◦C while there was also considerable peak area reduction at

30◦C (figure 4.34 d, b, and c respectively). The unknown metabolites at RT of more

than 7 min were prominent at all three temperatures.

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a. b.

c. d.

Figure. 4.34 Effect of temperature on degradation of CPF (RT= 5.4 min) by consortium

D. (a) Control, (b) consortium D, at 25°C(c) 30°C (d) 37°C.

4.6.2i Effect of temperature on CPF degradation by consortium E

Consortium E showed the same trend with almost same tremendous reduction in CPF

peak area at 25, 30 and 37◦C with appearance of new peaks at retention time of 7 to

8.5min (figure 4.35 b, c and d).

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a. b.

c. d.


E. (a) Control, (b) consortium E, at 25°C(c) 30°C (d) 37°C.

4.6.2j Effect of temperature on CPF degradation by consortium F

Consortium F behaved similarly and exhibited almost complete reduction of CPF peak

area at 25, 30 and 37◦C thus indicating excellent results for CPF degradation at all

temperatures along with presence of same peaks of metabolites as described in above

consortia (figure 4.36 b, c and d).

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a. b.

c. d.


consortium F. (a) Control, (b) consortium F, at 25°C(c) 30°C. Note peak for TCP at RT

= 1.77 min. (d) 37°C.

4.6.2k Effect of temperature on CPF degradation by consortium G

Same was the case with consortium G, having almost complete diminishing of CPF

peak area at all three given temperatures (25, 30 and 37◦C) with the formation of same

metabolites (figure 4.37 b, c and d).

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a. b.

c. d.


G. (a) Control, (b) consortium G, at 25°C(c) 30°C (d) 37°C.

4.6.3 Degradation of 3, 5, 6-Trichloropyridinol (14mg/l) by four isolates after72

hrs incubation

In case of all isolates, there was great reduction in TCP peak area and thus maximum

degradation of TCP (14 mg/l) after 72 hrs of incubation. Some new peaks with different

retention times can also be seen in the chromatogram indicating the new metabolites

formed by TCP degradation (figure 4.38b, c, d, e).

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a. b.

c. d.

e.

Figure 4.38 (a) HPLC chromatogram of standard TCP. Degradation of 14mg/l 3, 5, 6-

Trichloropyridinol (TCP) by (b). MB490, c). MB497, d). MB498, e). MB504 after 72

hrs incubation.

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4.7 Qualitative analysis through GCMS to detect metabolites of Chlorpyrifos

(CPF) by the 4 isolates and their consortia in the M-9 broth, soil slurry and soil

microcosm

The intermediate byproducts formed during biodegradation of CPF were identified by

comparing their mass spectra either to that of National Institute of Standards and

Technology (NIST) library or to the mass spectra observed for standards. When

bacterial isolates and their 7 consortia were analyzed for CPF metabolite formation

using GCMS, many peaks appeared in the chromatogram at different retention times.

As a whole, five metabolites of CPF i.e 2-Hydroxy-3, 5, 6-trichloropyridine (TCP),

[(3,5, 6-trichloro-2-pyridinyl)oxy] acetic acid (observed only in MB490), Diethyl

thiophosphate (DETP), phosphorothioic acid and Diisopropyl methanephosphonate

(observed only in samples with MB497, MB498 and MB504), were detected in all the

samples of given media (M-9 broth, soil slurry and soil microcosm) after 3, 6 and 9

days of incubation with the help of NIST library (Table 4.3 and figures 4.40, 4.41, 4.42,

4.43). When GCMS spectral data was compared with similar data reported in literature

for degradation of CPF (Díaz-Cruz and Barcelo, 2006), it was determined that peaks at

m/z 258 or 255 correspond to the removal of two ethylene molecules from the parent

molecule of CPF, at m/z 314 respectively. The peak at m/z of 27, indicated one ethylene

molecule removed from side chain of parent molecule. The breaking of the P O bond

is indicated at m/z 97 corresponding to the molecule H3PO4 released from the parent

ion. The peak at m/z 81 indicated release of an oxygen from H3PO4 to give H3PO3.

Whereas, the peak at m/z 197 indicated the formation of the stable and main metabolite

TCP by complete removal of the side chain. Diethyl thiophosphate (DETP) was

detected at m/z values of either 169, 170, or 171, while phosphorothioic acid at m/z

value of 109.

The m/z peaks of some metabolites in the mass spectrum may appear higher than their

mass values by +1,+2 or even +3 due to the fact that a considerable portion of isotopes

of Hydrogen (2H), Carbon (13C), Oxygen (18O) and Nitrogen (15N) is mostly present in

chemical compounds in nature. The appearance of m/z peak for a certain compound at

a mass which is less than its mass value by 1 (M+-1), or 2 (M+-2) etc. may be due to

loss of hydrogen ions during fragmentation process (Reddy et al., 2012). All isolates

122

and their consortia showed similar major fragments of Chlorpyrifos having almost

identical spectra. So, the present results are in complete agreement with previous

reports where TCP and DETP have been reported and detected as major metabolites of

CPF (Feng, 1998; Yang et al., 2005; Kim and Ahn, 2009; Li et al., 2010; Ajaz et al.,

2012; Alvarenga et al., 2015; Rayu et al., 2017). GCMS chromatograms for consortia

in soil microcosm after 3 days of incubation are shown in figure 4.43 respectively,

where TCP is clearly indicated at retention time of 12.8 minutes and CPF at 22 minutes.

It is worthy to mention that formation of 3, 5, 6 Trichloropyridinol during CPF

biodegradation was also confirmed during HPLC analysis with a peak appearing at

retention time of 1.77 min (figures 4.26, 4.27, 4.28, 4.29 and 4.36). The absence of TCP

peak in some CPF degradation chromatograms by HPLC may be due to its rapid

mineralization and consumption by the bacterial isolates.

A.

B.

Figure 4.39 (A). GCMS Chromatogram of Chlorpyrifos standard (RT = 22 minutes).

(B). GCMS Chromatogram of 3, 5, 6 Trichloropyridinol (TCP) standard with retention

time (RT) = 12.8 minutes.


CPF standard

RT = 22 min

TCP standard.

RT =12.8 min


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Figure 4.40 (A). Mass spectra of Chlorpyrifos standard (m/z: 20-320). (B). Mass spectra

of Chlorpyrifos degraded by bacteria (m/z: 20-260).

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Table 4.3: Biotransformation of Chlorpyrifos by the 4 bacteria in M-9 broth, soil

slurry and soil microcosm as analyzed by GCMS.

CPF and its metabolites detected by GCMS

Compound Name Chemical structure Retention

Time

(minutes)

m/z

Chlorpyrifos

22 314, 97, 197,

199

3, 5, 6-Trichloropyridinol

(TCP)

12.8

197, 198, 199

[(3,5, 6-trichloro-2-

pyridinyl)oxy] acetic acid

(detected only in samples with

MB490)

4.4 255

Diethyl thiophosphate (DETP)

17 169, 170

Diisopropyl

methanephosphonate (detected

only in samples with MB497,

MB498 and MB504 )

18 81, 97, 134

Phosphorothioic acid

8 109

125

A.

B.

C.

Figure 4.41 GCMS Chromatograms of Chlorpyrifos degraded by: (A). MB490 (B).

MB497 (C). MB498 after 3 days incubation in M-9 broth. Peak a (RT = 4.4 min) = [(3,

5, 6-trichloro-2-pyridinyl) oxy] acetic acid detected only in samples with MB490. Peak

b (RT = 8 min) = phosphorothioic acid. Peak c (RT =12.8 min) = TCP. Peak d (RT =

17 min) = Diethyl thiophosphate (DETP). Peak e (RT = 18 min) = Diisopropyl

methanephosphonate, detected only in samples with MB497, MB498 and MB504.

Peak f (RT = 22 min) = Chlorpyrifos.

b f

a d

d b e

f

b c e f

c

c

c

d




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Figure 4.42 GCMS Chromatograms of Chlorpyrifos degraded by MB504 after 3 days

incubation in M-9 broth. Peak b (RT = 8 min) = phosphorothioic acid. Peak c (RT =12.8

min) = TCP. Peak d (RT = 17 min) = Diethyl thiophosphate (DETP). Peak e (RT = 18

min) = Diisopropyl methanephosphonate, detected only in samples with MB497,

MB498 and MB504. Peak f (RT = 22 min) = Chlorpyrifos.

b d e f c


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a. b.

c. d.

E. e. f.

g.

Figure 4.43 GCMS Chromatogram of Chlorpyrifos degraded by: (a). Consortium A

(b). Consortium B (c). Consortium C (d). Consortium D (e). Consortium E (f).

Consortium F (g). Consortium G after 3 days incubation in soil microcosm. Note the

peak for CPF at 22 min and for TCP at12.8 min. All other peaks and metabolites are

almost similar to those detected in case of bacterial isolate except peak at 4.4 min for

[(3, 5, 6-trichloro-2-pyridinyl) oxy] acetic acid is absent in all consortia.





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4.8 Biotransformation of 3, 5, 6-Trichloropyridinol (TCP) by the 4 bacterial

isolates in M-9 broth as analyzed by GCMS

All four bacterial isolates were tested for their potential to transform TCP, main

metabolite of CPF in M-9 broth supplemented with TCP and analyzed by GCMS after

24, 48 and 72 hrs. It was noticed that all exhibited almost same pattern of peaks in the

chromatogram and they formed same type of metabolites as identified by the NIST

library after 24, 48 and 72 hrs of incubation i.e 1-methyl-2-pyrrolidine ethanol, p-

Propyl phenol, 2-Ethoxy-4, 4, 5, 5-tetramethyloxazoline and 3-(2, 4, 5-

Trichlorophenoxy)-1-propyne (Table 4.4 & figures 4.45, 4.46). Moreover, it can be

seen in the chromatograms that peak for TCP (RT = 12.8 minutes), were highly reduced

indicating transformation and mineralization of TCP by the isolates.

Table 4.4: Biotransformation of 3, 5, 6-Trichloropyridinol (TCP) by the 4 bacteria

in M-9 broth as analyzed by GCMS.

TCP and its metabolites detected by GCMS

Compound Name Chemical structure Retention

Time

(min)

m/z

3, 5, 6-Trichloropyridinol

(TCP)

12.8

197, 198, 199

1-methyl-2-pyrrolidine ethanol.

7

129

p-Propyl phenol.

13

136

2-Ethoxy-4, 4, 4, 5, 5-

tetramethyloxazoline.

13.6 171

3-(2, 4, 5-Trichlorophenoxy)-1-

propyne.

15.9 234

129

A.

B.

Figure 4.44. (A). GCMS chromatogram of TCP standard (RT =12.8 min). (B). Mass

spectra of standard TCP.


m/z values

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A.

B.

C.

Figure 4.45. GCMS chromatogram (A) and Mass spectrum (B) of TCP degraded by

MB490. GCMS chromatogram of TCP degraded by (C). MB497 after 72 hrs in M-9

broth with following identified peaks: b. 3, 5, 6-Trichloropyridinol (RT=12.8 min), c.

p-Propyl phenol (RT= 13 min). d. 2-Ethoxy-4, 4, 4, 5, 5-tetramethyloxazoline (RT=

13.6 min). e. 3-(2, 4, 5-Trichlorophenoxy)-1-propyne (RT= 15.9 min).

b e

b

c

d e


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A.

B.

Figure 4.46. GCMS chromatogram of TCP degraded by (A). MB498 (B). MB504 after

72 hrs in M-9 broth with following identified peaks: a. 1-methyl-2-pyrrolidine ethanol

(RT= 7 min) detected only in MB504, b. 3, 5, 6-Trichloropyridinol (RT = 12.8 min), c.

p-Propyl phenol (RT= 13 min). d. 2-Ethoxy-4, 4, 4, 5, 5-tetramethyloxazoline (RT=

13.6 min). e. 3-(2, 4, 5-Trichlorophenoxy)-1-propyne (RT = 15.9 min). Note that peak

for TCP (RT =12.8 min) in C (MB504) has been completely diminished indicating its

complete mineralization.

4.9 Proposed pathway for 3, 5, 6-Trichloropyridinol (TCP) degradation by

bacterial isolates (MB490, MB497, MB498 and MB504)

On the basis of above findings by GCMS analysis, the metabolic pathway for TCP can

be suggested as illustrated in figure 4.47. According to this pathway, TCP underwent

two different and parallel reactions catalyzed by two different enzymes depending upon

bacterial strains. In the 1st reaction chlorine atoms were removed from TCP with the

help of dehalogenase resulting in the formation of 1-methyl-2-pyrrolidine ethanol,

detected only in strain MB504, which was further oxidized by oxidase enzyme giving

rise to 2-Ethoxy-4, 4, 5, 5-tetramethyloxazoline followed by removal of nitrogen by

b a c

. d e

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132

deaminase to give p-Propyl phenol. Similarly in the second parallel reaction in other

three strains, TCP was first converted to 3-(2, 4, 5-Trichlorophenoxy)-1-propyne by

deaminase, followed by removal of three chlorine atoms with the help of dehalogenase

enzyme to form p-Propyl phenol which was ultimately mineralized into CO2 and H2O

(figure 4.46). Li et al. (2010) studied 3, 5, 6-trichloro-2-pyridinol degradation by

Ralstonia sp. strain T6 and identified two metabolites namely 3, 6-dihydroxypyridine-

2, 5-dione and 5-amino-2, 4, 5-trioxopentanoic acid. Many intermediates including 5,

6-DCP, dihydroxypyridine along with pyrrol structures substituted with carboxylic

groups were formed during photocatalytic degradation of TCP (Zabar et al., 2016).

Figure 4.47 Proposed pathway for 3, 5, 6-Trichloropyridinol (TCP) degradation by

bacterial isolates. The metabolite in red box (1-methyl-2-pyrrolidine ethanol) was

observed specifically in samples with MB504.

4.10 Proposed metabolic pathway of Chlorpyrifos degradation by bacterial

isolates (MB490, MB497, MB498 and MB504) and their consortia A, B, C, D, E, F

and G

Based on above mentioned findings by GCMS analysis, a metabolic pathway can be

suggested (figure 4.48) for Chlorpyrifos degradation by bacterial isolates (MB490,

MB497, MB498 and MB504) and their consortia A, B, C, D, E, F and G. The first step

3, 5, 6-trichloropyridinol

1-methyl-2-pyrrolidine ethanol

2-Ethoxy-4, 4, 5, 5-tetramethyloxazoline

3-(2, 4, 5-Trichlorophenoxy)-1-propyne

p-Propyl phenol

Mineralization

Deaminase

Dehalogenase

Dehalogenase

Oxidase

Deaminase

133

in this pathway possibly involved the hydrolysis of CPF into 2-Hydroxy-3, 5, 6-

trichloropyridine (TCP) and Diethylthiophosphate (DETP) by organophosphate

phosphatase (OPP). Then TCP followed two independent and parallel pathways

depending upon bacterial strains, probably involving different enzymes like

dehalogenase, oxidase and deaminase respectively giving rise to two products:

1. 1-methyl-2-pyrrolidine ethanol (formed only by strain MB504).

2. 3-(2, 4, 5-Trichlorophenoxy)-1-propyne (formed in all four strains).

But in case of MB490, TCP first underwent conjugation with acetic acid to form [(3, 5,

6-trichloro-2-pyridinyl) oxy] acetic acid, which was dehalogenated to give 1-methyl-2-

pyrrolidine ethanol. Then 1-methyl-2-pyrrolidine ethanol was converted to 2-Ethoxy-

4, 4, 5, 5-tetramethyloxazoline after oxidation by oxidase. Next step involved

dechlorination from 3-(2, 4, 5-Trichlorophenoxy)-1-propyne and deamination from 2-

Ethoxy-4, 4, 5, 5-tetramethyloxazoline to convert both of them into p-Propyl phenol

which was ultimately mineralized to CO2 and H2O via Krebs cycle. Similarly DETP

followed two pathways. In case of MB497, MB498 and MB504, it was first converted

to Diisopropyl methanephosphonate by alkylation which was then dealkylated to give

Phosphorothioic acid. While in MB490, it was directly dealkylated to form

Phosphorothioic acid which underwent mineralization providing carbon and

phosphorus for bacterial growth (figure 4.48). The formation of TCP during CPF

degradation has been revealed in many studies (Das and Adhya, 2015; Aceves-Diez et

al., 2015; Pradeep and Subbaiah, 2015; Rosbero and Camacho, 2017). Liu et al. (2016)

reported formation of 3, 5, 6-trichloro-2-pyridinol and diethyl phosphate by engineered

Pseudomonas putida. Recently 5, 6-DCP, dihydroxypyridine along with TCP were

detected during CPF degradation by integrated recirculating constructed wetlands

including microbial community (Tang et al., 2017).

134

Figure 4.48. Proposed metabolic pathway of Chlorpyrifos degradation by bacteria.

Metabolite in green box was specific for MB490 and metabolite in yellow box was

noticed in MB497, MB498, and MB504.

Conclusion

Four bacterial strains (Pseudomonas kilonensis MB490, Bacillus thuringiensis MB497,

Pseudomonas kilonensis MB498 and Pseudomonas sp. MB504) and their consortia

proved very efficient for CPF degradation and biotransformation. These strains also

showed the potential to degrade and mineralize TCP. So, keeping in view the prevailing

situation of extensive use of OP pesticides resulting in contamination of agricultural

soil and water resources, these isolated bacterial strains can be applied for the

bioremediation of CPF contaminated places.

Chlorpyrifos

Diethylthiophosphate

Phosphorothioic acid

2-Hydroxy-3,5,6-trichloropyridine

1-methyl-2-pyrrolidine ethanol

2-Ethoxy-4, 4, 5, 5-tetramethyloxazoline

3-(2, 4, 5-Trichlorophenoxy)-1-propyne

p-Propyl phenol

Mineralization

Organophosphate phosphatase

Dehalogenase

Deaminase

Dehalogenase

Oxidase

Deamination

Dealkylation

[(3, 5, 6-trichloro-2-

pyridinyl)oxy] acetic

acid

Diisopropyl

methanephos

phonate

Alkylation

conjugation

135

Chapter 5

Bacterial degradation and transformation of Triazophos (TAP)

Overview

Four bacterial isolates (MB490, MB497, MB498 and MB504) as well as their seven

consortia (A, B, C, D, E, F, and G) were examined for Triazophos (TAP) degradation

and bacterial growth (OD 600nm) at different pHs (6, 7, and 8) and temperatures (25,

30 and 37◦C) after 24 hrs incubation. The Triazophos degradation was also studied in

M-9 broth, soil slurry and soil microcosm after incubating for 1, 3, 6 and 9 days using

HPLC and GCMS for quantitative and qualitative analysis. All the bacterial isolates,

exhibited considerable TAP degradation (70.92 to 86.88%) at all pHs. Strains MB490,

MB497 and MB498 were best at pH 7 while MB504 showed almost equal and

maximum degradation at both pH 6 and pH 8. Strains MB497 and MB498 exhibited

maximum TAP degradation at 25°C, whereas MB490 and MB504 showed best results

at 37°C. Similarly, all the consortia exhibited excellent results for TAP degradation

(91.04 to 99.99%) at all pHs. Consortia A, B, C, F and G showed best degradation at

pH 8, while D and E could degrade maximally at pH 7. Consortia A, B, C, F and G

showed best degradation at 37°C (93.99 to 99.99%), while consortia D and E were

most active at 30 (99.99%) and 25°C (99.40%). Generally both individual isolates and

consortia exhibited more TAP degradation when grown statically. Presence of CPF

improved both TAP degradation and bacterial growth. In M-9 broth, there was 88.4 -

99% TAP degradation, while 98.67 - 99.90% degradation in soil slurry and 90.89 to

99.99% in soil microcosm was observed by bacterial isolates and consortia after 9 days.

During HPLC analysis, retention time (RT) for TAP standard was determined to be 2.4

min, while in GCMS analysis, it increased to 27.8 min. GCMS analysis revealed 7

unique and novel metabolites of TAP according to NIST library i.e 1, 2, 4-Triazole-4—

amine, N-(2-Thienylmethyl), Benzene sulfonic acid hydrazide, Benzene sulfonic acid

methyl ester, 4H-1,2,4-Triazole-4-benzenesulfonamide, 4, 5 dihydro-N-(O-toyl)-3-

furamide, Ethyl 4-phenyldiazenylbenzoate and Dibutyl methanephosphonate. In

general, results revealed the potential of these strains for the degradation/remediation

of TAP contaminated agricultural soils.

136

Background

Triazophos (TAP) is an important OP pesticide which is being extensively used to

protect agricultural crops (paddy rice, cotton, maize and vegetables) from pests

worldwide. There are reports of TAP contamination in soil deposits, water bodies and

food items (Alexis et al., 2001; Li et al., 2008; Ghaffar et al., 2014). As a result of

greater chemical and photochemical stability, TAP has been considered potentially

harmful to aquatic organisms. Zhu et al. (2014) reported the toxic effects of TAP on

the early development of a fish (Gobiocypris rarus) including deformity, slower heart

rate, decreased body length, altered enzymatic functions and changed mRNA levels at

lower concentration (0.05 mg/l for embryos and 0.01 mg/l for larvae). Sharma et al.

(2015) studied harmful effects of triazophos in female Wistar rats and revealed

significant alteration of estrous cycle along with altered activity of various enzymes in

the ovary of treated rats. Triazophos was among most frequently detected pesticides

during 2010-2011 in food samples tested in India (Malik, 2014).

In view of current situation of ever increasing pesticide contamination in environment

and its harmful effects on non-target organisms including humans, bioremediation

using microbes, plants and other organisms has been considered as a less time

consuming, result oriented, economical and ecologically safe technique for clearing up

pollutants from environment. Effective degradation of TAP along with ethoprophos and

fenamiphos by two fungal strains (Fusarium oxysporum and Aspergillus flavus)

isolated from sandy soil was reported (Thabit and El-Naggar, 2014). Bacterial strain

Stenotrophomonas G1 was capable of degrading TAP (50 mg/l) up to 34% during 24

hrs (Deng et al., 2015). Similarly, there had been other reports for the degradation of

TAP by microbes (Dai et al., 2007; Yang et al., 2011; Tang and You, 2012).

The current study was conducted with the objective to investigate the bacterial

degradation of TAP in order to determine the potential of these indigenous bacterial

isolates (MB490, MB497, MB498 and MB504) and their consortia in order to possibly

use them for the remediation of TAP contaminated agricultural soils. For this purpose,

different parameters affecting the growth and degradation activities of the isolate in M-

9 broth, soil slurry as well as in soil microcosm were studied. GCMS analysis was

137

conducted to identify biodegradation products of TAP and to propose a degradation

pathway of TAP by the bacterial isolates.


5.1 Optimization of environmental parameters for TAP biodegradation

Four bacterial isolates as well as their seven consortia were optimized for Triazophos

degradation and bacterial growth (OD600 nm) at different pHs (6, 7, 8), temperatures

(25, 30, 37◦C) and different incubation period (1, 3, 6 and 9 days). In case of bacterial

isolates, MB490, MB497, MB498 were best TAP degraders at pH 7 while MB504

performed best TAP degradation at pH 6, while TAP degradation varied from 70.92 to

86.88% (figure 5.1). Consortia A, B, C, F and G exhibited highest TAP degradation at

pH 8 while consortia D and E were best at pH 7. Similarly, all the consortia exhibited

excellent results for TAP degradation ranging from 91.04-99.99% (figure 5.2).

Optimum temperature for TAP degradation was 37◦C (MB490, MB504) and 25◦C

(MB497, MB498). Percentage degradation ranged from 59.70 to 99.70% for different

isolates (figure 5.3). It was reported by Tang and You, (2012) that the optimal pH and

temperature for the degradation of TAP by Bacillus sp. TAP-1 were 6.5 to 8 and 32 °C,

respectively. Zhang et al. (2016) revealed that complete removal of 100 mg/l triazophos

by Burkholderia sp. in 5hrs at 25°C and pH 8 as optimal temperature and pH for

triazophos degradation. Consortia A, B, C, F and G had their maximum TAP

degradation at 37◦C, while consortia D and E were best at 30◦C and 25◦C respectively.

Degradation percentage ranged from 66.70% to 99.99% in various cases (figure 5.4).

There was only 1.3, 2 and 3% TAP degradation in the control (without any bacterial

inoculum) at 25, 30 and 37◦C respectively, thus endorsing the contribution of bacterial

isolates in pesticide degradation. Thus they showed their great potential to degrade TAP

under a wide range of pH and temperature.

Bacterial isolates generally exhibited more TAP degradation under static conditions

(figure 5.5), probably due to immobile bacterial cells having more contact with the

dissolved pesticide, or less available dissolved oxygen. Consortia showed a mixed

behavior, as some (consortia D and E) were best degraders under shaking conditions,

while others (consortia A, C and G) showed best results under both conditions.

138

Consortia B and F responded better under static conditions (figure 5.6). There are

reports proving that immobilized cells are much more tolerant to harmful chemicals,

due to which immobilized systems of microbial cells have been predominantly effective

for the removal of pesticide contamination (Ha et al., 2008; Ortiz-Hernandez et al.,

2013). There was little bit more abiotic degradation (5%) under aerobic (shaking)

conditions and only 3% under static conditions probably due to more abiotic oxidation

reactions occurring under agitation.

Figure 5.1. Optical density and % degradation of TAP by four isolates (MB490,

MB497, MB498 and MB504) at different pH after 24 hrs. Error bars represent standard


Figure 5.2. Optical density and % degradation of TAP by 7 consortia (A, B, C, D, E,

F, and G) at different pH after 24 hrs. Error bars represent standard errors for values


10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

3.5

4


MB490 MB497 MB498 MB504

% D

egra

dat

ion

OD

60

0n

m


10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

6 7 8 6 7 8 6 7 8 6 7 8 6 7 8 6 7 8 6 7 8

A B C D E F G

% D

egra

dat

ion

of

TAP

OD

60

0n

m

pHs% degradation Growth

139

Figure 5.3. Optical density and % degradation of TAP by four isolates (MB490,

MB497, MB498 and MB504) at different temperatures after 24 hrs. Error bars represent

standard errors for values of three sample replicates.

Figure 5.4. Optical density and % degradation of TAP by 7 consortia (A, B, C, D, E,

F, and G) at different temperature after 24 hrs. Error bars represent standard errors for

values of three sample replicates.

0

20

40

60

80

100

0

0.5

1

1.5

2

2.5

3

25

°C

30

°C

37

°C

25

°C

30

°C

37

°C

25

°C

30

°C

37

°C

25

°C

30

°C

37

°C

MB490 MB497 MB498 MB504

% D

egra

dat

ion

of

TAP

OD

60

0n

m


0

20

40

60

80

100

0

0.5

1

1.5

2

2.5

3

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

25

°C3

0°C

37

°C

A B C D E F G%

Deg

rad

atio

n o

f TA

P

OD

60

0n

m

% degradation Growth

140

Figure 5.5 Effect of shaking and static conditions on growth and TAP degradation by

bacterial isolates (MB490, MB497, MB498 and MB504). Error bars represent standard


Figure 5.6 Effect of shaking and static conditions on growth and TAP degradation by

bacterial consortia (A, B, C, D, E, F and G). Error bars represent standard errors for


5.2 Bacterial growth/ degradation of TAP in the presence of Chlorpyrifos

Four bacterial isolates ((MB490, MB497, MB498, MB504) and consortium G were

cultured in M-9 broth supplemented with 200 mg/l of TAP alone and also with mixture

of TAP and CPF (200 mg/l each). TAP degradation was quantitatively analyzed using

HPLC, after 24 hrs of incubation. The bacterial growth was also checked using UV-

VIS spectrophotometer. It was observed that presence of CPF positively enhanced TAP

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4


% D

egra

dat

ion

of

TAP

OD

60

0n

m

%Degradation Growth

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3


Shaking conditions Static conditions%

Deg

rad

atio

n

OD

60

0n

m

Incubation conditions


141

degradation as well as bacterial growth as compared to TAP alone. TAP degradation

was increased from 77.75-86.88 (TAP alone) to 91.4-99.6% (CPF + TAP) by bacterial

isolates and consortium G. (figure 5.7). This enhancing effect of CPF was correlated

with increased bacterial growth due to availability of more nutrients by

degradation/mineralization of both TAP and CPF by the bacteria. Abiotic degradation

of TAP was almost unaffected in the presence of CPF.

Figure 5.7. Effect of presence of CPF on the growth (OD600) and degradation of

Triazophos by the 4 isolates (MB490, MB497, MB498, MB504) and the consortium G

(initial concentration of TAP and CPF used = 200 mg/l each) in the M-9 broth. Error

bars represent standard errors for values of three replicates.

5.3 Effect of incubation period on bacterial growth and % degradation of TAP in

M-9 medium

The effect of incubation period (1, 3, 6 and 9 days) on the rate of TAP degradation was

studied by growing four isolates (MB490, MB497, MB498 and MB504) and their

consortia (A, B, C, D, E, F and G) in M-9 broth supplemented with 200 mg/l of TAP.

Analysis was done by HPLC for TAP degradation and by UV-VIS spectrophotometer

for growth periodically. Individual bacterial strains showed 88.4 to 95.8% TAP

degradation, while consortia performed better with more than 99% degradation after 9

days with very good growth (figures 5.8 and 5.9).

Maximum rate of TAP degradation and bacterial growth was achieved after first day of

incubation followed by slow increase up to 9th day (figures 5.8 and 5.9). The microbial

10

30

50

70

90

110

0

1

2

3

MB490 MB497 MB498 MB504 Consortium G % D

egra

dat

ion

of

TAP

OD

60

0n

m

% degradation of TAP alone

% degradation of TAP in presence of CPF

Growth of isolates in presence of TAP alone

Growth of isolates in presence of CPF + TAP

142

growth activity and chemical degradation processes of TAP were proved to be closely

linked. There was only 2, 3, 5 and 7% TAP degradation observed in the control having

no bacterial inoculum after 1, 3, 6 and 9 days respectively. Tang and You (2012)

reported that Bacillus sp. TAP-1, could degrade up to 98.5% of 100 mg/l TAP in the

medium after 5 days of incubation. Stenotrophomonas sp. PF32 could degrade 97% of

initial100 mg/l triazophos in 42 hrs and 97% of 100 mg/l Chlorpyrifos in 48 hrs (Xu et

al., 2009). Deng et al. (2015) isolated a bacterium Stenotrophomonas sp. G1 from

sludge that could degrade 34% of 50 mg/l of Triazophos in 24 hrs. Present study results

are much significant as compared to those reported earlier. Jabeen et al. (2015) reported

a consortium PBAC that could degrade Triazophos (50 mg/l) up to 57.7% within 3 days

of incubation.

Figure 5.8. Effect of incubation period on bacterial growth and % degradation of TAP

by bacterial isolates (MB490, MB497, MB498, MB504) in M-9 broth. The error bars

represent standard error from the mean of three replicates.

1030507090110

00.5

11.5

22.5

3

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4


% D

egra

dat

ion

of

TAP

OD

60

0n

m

Incubation period


143

Figure 5.9 Effect of incubation period on the growth and degradation of Triazophos by

the bacterial consortia (A, B, C, D, E, F, and G) in M-9 broth (initial concentration of

TAP used=200 mg/l). Error bars represent standard error from the mean of three

replicates.

5.4 Degradation of Triazophos by the bacterial isolates and their consortia in soil

slurry

The TAP degradation by bacterial isolates and their consortia in soil slurry

supplemented with 200 mg/l of TAP was analyzed using HPLC after 3, 6 and 9 days of

incubation. Maximum TAP degradation (98.67-99.90% and 92.60-99.90%) was

achieved by individual bacterial strains and their consortia respectively in soil slurry

after 9 days of incubation (figure 5.10 and 5.11). Moreover, TAP degradation was most

rapid during first 3 days of incubation in collaboration with bacterial population at peak

growth (Table 5.1), after which it increased slowly reaching highest degradation after

9 days in soil slurry (figure 5.10 and 5.11). Stationary phase of bacterial growth was

observed during 3rd to 9th day indicated by decreased CFU of surviving bacterial cells

(Table 5.2). On contrary, only 2, 4 and 6% TAP degradation in the control was observed

after 3, 6 and 9 days of incubation respectively.

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3



% D

egra

dat

ion

of

TAP

OD

60

0n

m

Incubation period% Degradation Growth

144

Table 5.1 Bacterial Population in soil slurry spiked with 200 mg/l of TAP

Bacterial isolates CFU/ml soil


MB490 1.9×106 ± 0.1 6.1×106 ± 0.05 3.5×106 ± 0.1 2.8×106 ± 0.1

MB497 1.3×106 ± 0.05 5.7×106 ± 0.1 4.1×106 ± 0.05 2.5×106 ± 0.05

MB498 1.4×106 ± 0.1 6.4×106 ± 0.05 3.6×106 ± 0.1 2.4×106 ± 0.1

MB504 1.3×106 ± 0.05

6.6×106 ± 0.1 4.6×106 ± 0.05 2.2×106 ± 0.05


standard error.

Figure 5.10 Effect of incubation period on degradation of Triazophos by the bacterial

isolates (MB490, MB497, MB498, and MB504) in the soil slurry (initial concentration

of TAP used=200 mg/l). Error bars represent standard errors for values of three

replicates.

Figure. 5.11 Effect of incubation period on degradation of Triazophos by the 7 consortia (A,

B, C, D, E, F and G in the soil slurry (initial concentration of TAP used=200 mg/l).

Error bars represent standard errors for values of three sample replicates.

10

30

50

70

90

110

MB490 MB497 MB498 MB504

% D

egra

dat

ion

Incubation period


10

30

50

70

90

110

A B C D E F G

% D

egra

dat

ion

of

TAP

Incubation period (days)


145

5.5 Degradation of Triazophos by bacterial strains and their consortia in the soil

microcosm

Triazophos biodegradation in soil microcosm by the bacterial isolates and their

consortia was analyzed using HPLC after 3, 6 and 9 days of incubation. For this

purpose, they were inoculated in petri plates having 50 g of soil with 40% water holding

capacity supplemented with 200 mg/kg of TAP and incubated under dark conditions

for up to 9 days. There was observed a gradually increasing trend of TAP degradation

by all bacterial isolates (MB490, MB497, MB498 and MB504) with increasing period

of incubation in soil microcosm ranging from minimum 92.72 (MB498) to maximum

95% (MB504) after 9 days of incubation (figure 5.12). Furthermore, the rate of TAP

degradation i.e amount of TAP degraded per day was observed to be most rapid during

first 3 days of incubation linked with log phase of bacterial growth in terms of CFU/g

of soil (Table 5.2), then it slowed down gradually. In present study, results of TAP

degradation are much better than that reported by Liang et al. (2011), where

Diaphorobacter sp. GS-1 could degrade 95.38% of TAP (50 mg/kg) in paddy soil

microcosm after 21 days of incubation. On contrary, there was only 7, 10 and 16% TAP

degradation in the unsterilized soil and 3, 5 and 8% degradation in the control (sterilized

soil) without any bacteria after 3, 6 and 9 days of incubation respectively. Thus proving

the positive contribution of isolated bacteria in TAP degradation.

All the consortia exhibited remarkable TAP degradation in soil microcosm with

minimum 90.89% (consortium B) to maximum 99.99% (consortium A and D) after 9

days of incubation. It was observed that maximum degradation rate had been achieved

during first 3 days of incubation mainly due to co-metabolic activities of consortia

members in log phase. Then, there was slow increase in degradation from 3rd to 9th day

along with stationery phase of bacteria (figure 5.13).

146

Figure 5.12 Effect of incubation period on degradation of Triazophos by bacterial isolates

(MB490, MB497, MB498, and MB504) in the soil microcosm (initial concentration of TAP

used = 200 mg/kg). The error bars represent standard error from the mean of three replicates.

Table 5.2 Bacterial Population in soil microcosm spiked with 200 mg/kg of TAP

Bacterial

isolates

CFU/g soil


MB490 1.6×106 ± 0.1 6.5×106 ± 0.05 3.5×106 ± 0.1 2.1×106 ± 0.1

MB497 1.3×106 ± 0.05 5.5×106 ± 0.1 3.1×106 ± 0.05 2.5×106 ± 0.05

MB498 1.4×106 ± 0.1 6.2×106 ± 0.05 3.4×106 ± 0.1 2.4×106 ± 0.1

MB504 1.5×106 ± 0.05 5.6×106 ± 0.1 4.1×106 ± 0.05 2.6×106 ± 0.05


standard error.

Figure 5.13. Effect of incubation period on degradation of Triazophos by the bacterial

consortia (A, B, C, D, E, F and G) in the soil microcosm (initial concentration of TAP

used = 200 mg/l). The error bars represent standard error from the mean of three

replicates.

0

20

40

60

80

100

MB490 MB497 MB498 MB504 Control Unsterilizedsoil

% D

egra

dat

ion

of

TAP


0

20

40

60

80

100

120

A B C D E F G

% D

egra

dat

ion

of

TAP

Incubation period (days)


147

5.6 Quantitative analysis through HPLC for biodegradation of TAP

5.6.1 Effect of pH on TAP degradation

5.6.1a Effect of pH on TAP degradation by isolate MB490

When TAP degradation by MB490 at different pH was analyzed by HPLC, it was observed

that there was a very sharp decrease in peak area of TAP (RT = 2.4 min) at pH 7 (figure

5.14c) as compared to control (figure 5.14a), corresponding to maximum TAP degradation

at pH 7. Whereas, there was least decrease in peak area (figure 5.14b) and lowest TAP

degradation at pH 6 after 24 hrs of incubation. While, there was considerable decrease in

peak area of TAP at pH 8, thereby indicating considerable degradation as given in (figure

5.14d). Some new peaks appeared at retention time of 1.3, 1.5 and 1.9 min at pH 6, pH 7 and

pH 8 (figure 5.14b, c and d respectively).

148

a. b.

c. d.

Figure 5.14 Effect of pH on degradation of TAP (RT= 2.4 min) by MB490. (a)


5.6.1b Effect of pH on TAP degradation by isolate MB497

In case of MB497, there was much reduction in TAP peak area (RT = 2.4 min) at pH 7

with maximum TAP degradation (figure. 5.15c). Almost equally decreased TAP peaks

were noticed at pH 6 and pH 8, thus showing considerable but equal degradation of

TAP at both pH 6 and pH 8 as shown in figure. 5.15b & d respectively.

Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)



Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

149

a. b.

c. d.

Figure. 5.15 Effect of pH on degradation of TAP (RT= 2.4) by MB497. (a) Control,

(b) MB497 at pH 6, (c) pH 7, (d) pH 8.

5.6.1c Effect of pH on TAP degradation by isolate MB498

For MB498, there was maximum decrease in TAP peak area at pH 7 thus showing

maximum TAP degradation, while there was least decrease in peak area at pH 6

followed by decrease at pH 8 as illustrated in figure 5.16c, b, & d respectively.



Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

150

a. b.

c. d.

Figure. 5.16 Effect of pH on degradation of TAP (RT = 2.4 min) by MB498. (a)


5.6.1d Effect of pH on TAP degradation by isolate MB504

There was maximum and nearly equal decrease in peak area of TAP at pH 6 and pH 8

(figure 5.16b and d). There was also considerable decrease in peak area at pH 7 (figure

5.17c).



Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

Ab

sorb

ance

(m

V)

151

a. b.

c. d.

Figure. 5.17 Effect of pH on degradation of TAP (RT= 2.4 min) by MB504. (a)


5.6.1e Effect of pH on TAP degradation by consortium A

There was little more decrease in peak area of TAP at pH 8 in case of consortium

indicating higher TAP degradation as compared to that at pH 7 and 6 as shown in figure

5.18d, c & b respectively. Many new peaks can be seen in the chromatograms at pH 6,

7 and 8 denoting some unknown metabolites.



Ab

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(m

V)

Ab

sorb

ance

(m

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a. b.

c. d.

Figure. 5.18 Effect of pH on degradation of TAP (RT= 2.4 min) by consortium A. (a)

Control, (b) consortium A, at pH 6, (c) pH 7, (d) pH 8.

5.6.1f Effect of pH on TAP degradation by consortium B

In case of consortium B, maximum decrease in TAP peak area was observed at pH 8

(figure 5.19d), followed by pH 6 and 7 as presented in figure 5.19b & c respectively,

thus it could degrade TAP considerably at all given pH after 24 hrs.



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a. b. .

c. d.

Figure. 5.19 Effect of pH on degradation of TAP (RT= 2.4) by consortium B. (a)

Control, (b) consortium B, at pH 6, (c) pH 7, (d) pH 8.

5.6.1g Effect of pH on TAP degradation by consortium C

Consortium C demonstrated highest loss of peak area at pH 8 with remarkable decrease

in peak area and thus TAP degradation at pH 7 and pH 6 as shown in figure 5.20 d, c

& b respectively.



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a. b.

c. d.

Figure. 5.20 Effect of pH on degradation of TAP (RT= 2.4) by consortium C. (a)

Control, (b) consortium C, at pH 6, (c) pH 7, (d) pH 8.

5.6.1h Effect of pH on TAP degradation by consortium D

The peak area for TAP decreased greatly at pH 7 (figure 5.21c) for consortium D as it

could degrade TAP to highest value. While at pH 6 and pH 8, it could show conspicuous

degradation with remarkable loss of peak area of TAP as given in figure 5.21b & d,

respectively.



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a. b.

c. d.

Figure 5.21 Effect of pH on degradation of TAP (RT= 2.4) by consortium D. (a)

Control, (b) consortium D, at pH 6, (c) pH 7, (d) pH 8.

5.6.1i Effect of pH on TAP degradation by consortium E

In case of consortium E, maximum TAP degradation with greatest loss of TAP peak

area at pH 7 (figure 5.22c). While there was remarkable decrease in peak area at pH 6

and pH 8, with considerable TAP degradation (figure 5.22b and d respectively).



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a. b.

c. d.

Figure. 5.22 Effect of pH on degradation of TAP (RT= 2.4) by consortium E. (a)

Control, (b) consortium E, at pH 6, (c) pH 7, (d) pH 8.

5.6.1j Effect of pH on TAP degradation by consortium F

Consortium F could degrade maximum TAP with highest decrease in TAP peak area at

pH 8 (figure 5.23d) followed by pH 6 and pH 7 respectively with considerable

degradation (figure 5.23b and c).



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a. b.

c. d.

Figure. 5.23 Effect of pH on degradation of TAP (RT= 2.4) by consortium F. (a)

Control, (b) consortium F, at pH 6, (c) pH 7, (d) pH 8.

5.6.1k Effect of pH on TAP degradation by consortium G

Consortium G (MB490+MB497+MB498+MB504) exhibited highest TAP degradation

and peak area almost diminished at pH 8 and pH 7 with the appearance of a prominent

peak at RT = 1.9 min indicating some unknown metabolite (figure 5.24d and c

respectively). While, it showed minimum degradation and least decrease of TAP peak

area at pH 6 with many new peaks (figure 5.24b).



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a. b.

c. d.

Figure. 5.24 Effect of pH on degradation of TAP (RT= 2.4) by consortium G. (a)

Control, (b) consortium G, at pH 6, (c) pH 7, (d) pH 8.

5.6.2 Effect of temperature on TAP degradation

5.6.2a Effect of temperature on TAP degradation by isolate MB490

TAP degradation by bacterial isolate MB490 was analyzed by HPLC at different

temperatures i.e 25, 30 and 37◦C after 24 hrs. Maximum decrease was observed in TAP

peak area at 37◦C (figure 5.25d) while it showed minimum but almost equal decrease

in peak area at 25◦C and 30◦C as illustrated in figure 5.25b & c respectively. Some new

peaks of degradation products especially at RT of 1.8 and 1.9 min are prominent in

chromatograms at 30◦C and 37◦C.



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a. b.

c. d.

Figure. 5.25 Effect of temperature on degradation of TAP (RT= 2.4 min) by MB490.


5.6.2b Effect of temperature on TAP degradation by isolate MB497

Strain MB497 could exhibit maximum decrease in TAP peak area and thus maximum

degradation at 25◦C with a new remarkable peak at 1.8 min denoting some unidentified

metabolite of TAP (figure 5.26b). While loss of peak area was almost equal at 30◦C and

37◦C (figure 5.26c and d).



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a. b.

c. d.

Figure. 5.26 Effect of temperature on degradation of TAP (RT= 2.4) by MB497. (a)

Control, (b) MB497 at 25°C, (c) 30°C, (d) 37°C.

5.6.2c Effect of temperature on TAP degradation by isolate MB498.

In case of MB498, complete disappearance of TAP peak at RT of 2.4 min indicated

excellent and highest TAP degradation at 25◦C, with appearance of a new peak at 1.8

min (figure 5.27b). Whereas, there was minimum decrease in TAP peak area at 30◦C

followed by that at 37◦C as shown in figure 5.27c & d respectively.



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a. b.

c d.

Figure. 5.27 Effect of temperature on degradation of TAP (RT= 2.4 min) by MB498.


5.6.2d Effect of temperature on TAP degradation by isolate MB504

Strain MB504 could show maximum decline of TAP peak area at 37◦C (figure 5.28d),

while minimum decrease occurred at 30◦C. There was moderate lessening of peak area

at 25◦C. Some small new peaks were also observed at all given temperatures (figure

5.28b, c and d).



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a. b.

c. d.

Figure. 5.28 Effect of temperature on degradation of TAP (RT = 2.4 min) by MB504.


5.6.2e Effect of temperature on TAP degradation by consortium A

TAP peak completely vanished in case of consortium A at 37◦C, while least decline in

peak area was observed at 25◦C followed by 30◦C as shown in figure 5.29d, b & c

respectively.



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a. b.

c. d.

Figure. 5.29 Effect of temperature on degradation of TAP (RT= 2.4 min) by consortium

A. (a) Control, (b) consortium A at 25°C, (c) 30°C, (d) 37°C.

5.6.2f Effect of temperature on TAP degradation by consortium B

Consortium B exhibited best TAP degradation at 37◦C with more or less complete

decline of TAP peak area (figure 5.30d), while minimum decrease in peak area occurred

at 30◦C followed by 25◦C with many new peaks appearing after degradation (figure

5.30c and b respectively).

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a. b.

c. d.

Figure. 5.30 Effect of temperature on degradation of TAP (RT= 2.4) by consortium B.

(a) Control, (b) consortium B at 25°C, (c) 30°C, (d) 37°C.

5.6.2g Effect of temperature on TAP degradation by consortium C

Likewise, consortium C demonstrated maximum reduction of TAP peak area at 37◦C

(figure 5.31d) while minimum decline in peak area was observed at 25◦C followed by

30◦C (figure 5.31b and c respectively). Many new peaks can be seen in figure 5.31b, c

& d.

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a. b.

c d.

Figure. 5.31 Effect of temperature on degradation of TAP (RT= 2.4) by consortium C.

(a) Control, (b) consortium C at 25°C, (c) 30°C, (d) 37°C.

5.6.2h Effect of temperature on TAP degradation by consortium D

Consortium D showed almost diminished peak area of TAP at 30◦C (figure

5.32c).While it exhibited minimum decline in TAP peak area at 25◦C and moderate

reduction at 37◦C as shown in figure 5.32b & d respectively.

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a. b.

c. d.


D. (a) Control, (b) consortium D at 25°C, (c) 30°C, (d) 37°C.

5.6.2i Effect of temperature on TAP degradation by consortium E

Consortium E showed maximum decrease in TAP peak area at 25◦C. While it exhibited

minimum reduction at 30◦C and moderate decline at 37◦C (figure 5.33c and d

respectively) along with many new peaks appearing after degradation.

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a. b.

c. d.


E. (a) Control, (b) consortium E at 25°C, (c) 30°C, (d) 37°C.

5.6.2j Effect of temperature on TAP degradation by consortium F

Consortium F exhibited almost equal decrease in TAP peak area at 37, 25 and 30◦C

showing excellent TAP degradation of 99.99, 99.74 and 99.73% respectively. It seemed

as temperature had no effect on TAP degradation by consortium F (figure5.34).

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a. b.

c. d.

Figure. 5.34 Effect of temperature on degradation of TAP (RT= 2.4) by consortium F.

(a) Control, (b) consortium F at 25°C, (c) 30°C, (d) 37°C.

5.6.2k Effect of temperature on TAP degradation by consortium G

Consortium G showed maximum reduction of TAP peak area at 37◦C and minimum

decline at 30◦C, while there was also remarkable peak reduction at 25◦C (figure5.35d, c

and b respectively).

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a. b.

c. d.


G. (a) Control, (b) consortium G at 25°C, (c) 30°C, (d) 37°C.

5.7 Qualitative analysis through GCMS to detect metabolites of OP Pesticide

Triazophos (TAP) by bacterial isolates and their consortia in M-9 broth, soil slurry

and soil microcosm

The GCMS analysis of the bacterial cultures (M-9, soil slurry and soil microcosm)

treated with Triazophos (200 mg/l/kg) was performed after 3, 6 and 9 days of

incubation. During all experiments, as a whole there were total 8 major peaks identified

in the chromatograms for MB490, MB497, MB498 and MB504 and their consortia by

comparing the mass spectra with those of the reference compounds in the data systems

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of NIST library. Following compounds were identified (figures 5.37, 5.38 & Table 5.3):

1, 2, 4-Triazole-4—amine, N-(2-Thienylmethyl), Benzene sulfonic acid hydrazide,

Benzene sulfonic acid, methyl ester. 4H-1, 2, 4-Triazole-4-benzenesulfonamide, 4, 5

dihydro-N-(O-toyl)-3-furamide, Ethyl 4-phenyldiazenylbenzoate, Dibutyl

methanephosphonate and parent compound Triazophos.

The metabolites observed in present study are quite different and novel from those

reported earlier due to photo-fenton degradation (Lin et al., 2004), hydrolytic

degradation (Rani et al., 2001; Lin et al., 2004), photocatalytic degradation (Aungpradit

et al., 2007), microbial and soil degradation (Wang et al., 2005; Bajeer et al., 2016).

Similar to current study, glucuronide and sulfate conjugates of 1-phenyl-3-hydroxy-

(1H)-1, 2, 4-triazole were detected in the urine of rats and dogs (Schwalbe-Fehl and

Schmidt, 1986). This suggested the presence of same metabolizing enzymes in both

bacterial isolates and in higher animals like rats and dogs etc.

a.

b.

Figure. 5.36 (a). GCMS chromatogram of Triazophos standard (RT= 27.8 min) (b).

GCMS mass spectrum of Triazophos standard.

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e A

bu

nd

ance


m/z values

Triazophos

RT= 27.8 min

171

a.

b.

Figure. 5.37 GCMS chromatogram of Triazophos in the sample treated with (a)


Peak 1 (RT = 11.5min) = 1, 2, 4-Triazole-4—amine, N-(2-Thienylmethyl).

Peak 2 (RT = 15.2min) =. Benzene sulfonic acid hydrazide

Peak 3 (RT = 17.2min) = Benzene sulfonic acid, methyl ester.

Peak 4 (RT = 18.2min) = 4H-1, 2, 4-Triazole-4-benzenesulfonamide.

Peak 5 (RT = 21.1min) = 4, 5 dihydro-N-(O-toyl)-3-furamide.

Peak 6 (RT = 21.5min) = Ethyl 4-phenyldiazenylbenzoate (absent in samples with

MB498)

Peak 7 (RT = 24.2min) = Dibutyl methanephosphonate.

Peak 8 (RT = 27.8 min) = Triazophos.

1 3 4 5 2 7 6 8



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a.

b.

Figure. 5.38 GCMS chromatogram of Triazophos in the sample treated with (a)


Peak1 (RT = 11.5 min) = 1, 2, 4-Triazole-4—amine, N-(2-Thienylmethyl).

Peak2 (RT = 15.2 min) =. Benzene sulfonic acid hydrazide

Peak3 (RT = 17.2 min) = Benzene sulfonic acid, methyl ester.

Peak4 (RT = 18.2 min) = 4H-1, 2, 4-Triazole-4-benzenesulfonamide.

Peak5 (RT = 21.1 min) = 4, 5 dihydro-N-(O-toyl)-3-furamide.

Peak6 (RT = 21.5 min) = Ethyl 4-phenyldiazenylbenzoate (absent in samples with

MB498)

Peak7 (RT = 24.2 min) = Dibutyl methanephosphonate.

Peak8 (RT = 27.8 min) = Triazophos. (This peak could not be detected in case of

MB498 may be due to complete consumption).

1 2

3

4

5 7



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1

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Table 5.3 Biotransformation of Triazophos by bacterial isolates and their

consortia in M-9 broth, soil slurry and soil microcosm.

S. No. Triazophos and its metabolites detected by GCMS

Compound Name Structure Retention

Time

(minutes)

1 Triazophos

27.8

2 4H-1, 2, 4-Triazole-4-

benzenesulfonamide.

18.2

3 Benzene sulfonic acid

hydrazide

15.2

4 1,2,4-Triazole-4—

amine,N-(2-

Thienylmethyl)

11.5

5 4,5 dihydro-N-(O-toyl)-

3-furamide.

21.1

6 Ethyl 4-

phenyldiazenylbenzoate

(observed only in

MB490, MB497,

MB504)

21.5

7

Benzene sulfonic acid,

methyl ester.

17.2

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8 Dibutyl

methanephosphonate

24.2

5.8 Proposed Metabolic Pathway for the biotransformation of TAP by bacterial

isolates MB490, MB497, MB498 and MB504

In the current study, the first step involved in the transformation of Triazophos is the

hydrolytic breakage of the P–O ester bond of Triazophos (figure 5.39) by

organophosphorus phosphatase (OPP) to form 1-phenyl-3-hydroxy-1, 2, 4-triazole (one

of most commonly reported metabolites of TAP though not detected in this form but in

conjugated form in the current study) and Diethylthiophosphate, the most common

initial intermediate of TAP (Wang et al., 2005), though not detected in present study.

It is important to note that Dibutyl methanephosphonate detected at retention time of

24.2 minutes may be formed by alkylation and desulfulrization of Diethylthiophosphate

(figure 5.39). Next 1-phenyl-3-hydroxy-1, 2, 4-triazole is conjugated either with

sulfonamide or thiophene by to form 4H-1, 2, 4-Triazole-4-benzenesulfonamide and 1,

2, 4-Triazole-4—amine, N-(2-Thienylmethyl) respectively by two parallel reactions.

The former of the two products is hydrolyzed/oxidized to Benzene sulfonic acid

hydrazide which is further hydrolyzed in to Benzene sulfonic acid, methyl ester and

hydrazine (not detected in current study). While 1, 2, 4 -Triazole-4—amine, N-(2-

Thienylmethyl) follows two pathways depending upon bacterial strains. Firstly, it is

hydrolyzed to Ethyl 4-phenyldiazenylbenzoate (only in three strains MB490, MB497

and MB504) which ultimately undergoes mineralization. Secondly in general, it is

oxidized to 4, 5 dihydro-N-(O-toyl)-3- furamide followed by desulfurization and

deamination. Further, 4, 5 dihydro-N-(O-toyl)-3- furamide and Benzene sulfonic acid,

methyl ester may undergo mineralization along with Hydrazine and Dibutyl

methanephosphonate. So, most of the products identified in this pathway of TAP

degradation are novel and has been reported first time.

The metabolism of pesticides in bacteria and fungi may involve three phases using

intracellular/extracellular enzymes like hydrolytic enzymes, oxygenases and

peroxidases etc. The phase I may transform the parent compound via oxidation,

reduction, or hydrolysis reactions to give a product that is more water-soluble and

commonly less toxic than the parent compound. In the second phase, a pesticide or its

175

metabolite may conjugate to a sugar (glucoronidation) or amino acid (amination), thus

increasing the water solubility and reducing toxicity as compared to the parent

pesticide. In the third phase, metabolites of phase II may be converted to non-toxic

secondary conjugates (Eerd et al., 2003; Ortiz-Hernandez et al., 2011).

Wang et al. (2005) reported the formation of O, O diethyl phosphorothioic acid and 1-

phenyl-3-hydroxy-1, 2, 4-triazole, as metabolites of TAP biodegradation, by Klebsiella

sp. E6 as a result of TAP hydrolysis. Then Yang et al. (2011) studied almost complete

degradation of 50 mg/l of TAP and its intermediate, 1-phenyl-3-hydroxy-1, 2, 4-triazole

by Diaphorobacter sp. TPD-1 in 24 and 56 hrs, respectively. They could identify three

metabolites i.e 1-phenyl-3-hydroxy-1, 2, 4-triazole, O, O-diethyl phosphorothioic acid

and 1-formyl-2-phenyldiazene. They further proposed that 1-formyl-2-phenyldiazene

is converted to 2-phenylhydrazinecarboxylic acid by addition of one water molecule

followed by decarboxylation of the 2-phenylhydrazinecarboxylic acid to form

phenylhydrazine. In the current study, metabolites Ethyl 4-phenyldiazenylbenzoate and

Benzene sulfonic acid hydrazide are very close to 2-phenylhydrazinecarboxylic acid

previously reported by Yang et al. (2011). Zhang et al. (2016) could only detect 1-

phenyl-3-hydroxy-1,2,4-triazole as initial metabolite during degradation of TAP by

Burkholderia sp. Wu et al. (2016) studied the metabolism of 1-phenyl-3-hydroxy-1, 2,

4-triazole by Shinella sp. NJUST26, and identified three main metabolites i.e 1, 2-

dihydro-3H-1,2,4-triazol-3-one, semicarbazide and urea. In the present study,

hydrazide compound is closely related to semicarbazide which is derivative of urea and

hydrazine (Schirmann and Bourdauducq, 2002). Jawale and Gogate, (2018) studied

the abiotic TAP degradation based on ultrasound technique and identified some

metabolites like 3Z Undecene-5,7,10-triynoicacid, 1-(2-Hydroxyethyl)-2-

hydroxymethyl-5-nitroimidazole, Acetylisoniazid, Swietenine,

Dihydrodeoxystreptomycin and Crocetin which seem very different from biotic and

specially microbial metabolites.

176

Figure 5.39. Proposed metabolic pathway for the biotransformation of TAP by

bacterial isolates MB490, MB497, MB498 and MB504. Metabolite in green box (Ethyl

4-phenyldiazenylbenzoate) was absent in MB498 and produced only by MB490,

MB497 and MB504 and their consortia.

Conclusion

The bioremediation may be considered as an effective alternative to the traditional

treatment methods of Triazophos contamination by application of microbial

metabolism. Present study proved that four isolates (Pseudomonas kilonensis MB490,

Bacillus thuringiensis MB497, Pseudomonas kilonensis MB498 and Pseudomonas sp.

Triazophos

Diethyl thiophosphate

Dibutyl methanephosphonate.

4H-1, 2, 4-Triazole

4H-1, 2, 4-Triazole-4-benzenesulfonamide

Benzene sulfonic acid hydrazide

Hydrazine Benzene sulfonic acid, methyl ester

1,2,4-Triazole-4—amine, N-(2-

Thienylmethyl)

4, 5 dihydro-N-(O-toyl)-3- furamide

Mineralization

Organophosphate Phosphatase

Hydrolysis

Conjugation

Hydrolysis

and

oxidation

Des

ulf

uri

zati

on,

dea

min

atio

n a

nd

Glu

coro

nid

atio

n Ethyl 4-

phenyldiazenyl

benzoate Hydrolysis

177

MB504) and their consortia were capable of degrading and transforming TAP into

simple products. Moreover, novel metabolites (1, 2, 4-Triazole-4—amine, N-(2-

Thienylmethyl), Benzene sulfonic acid hydrazide, Benzene sulfonic acid, methyl ester.

4H-1, 2, 4-Triazole-4-benzenesulfonamide, 4, 5 dihydro-N-(O-toyl)-3-furamide, Ethyl

4-phenyldiazenylbenzoate, Dibutyl methanephosphonate) of TAP were produced by

these bacterial strains indicating presence of some unique genes/enzymes. Therefore

these isolates would be valuable for the investigation of new bioremediation approaches

for TAP contaminated sites like agricultural soils and water bodies.

178

Chapter 6

Biodegradation and biotransformation of OP pesticide Dimethoate

(DM)

Overview

Four isolates and their seven consortia were analyzed by HPLC for Dimethoate (DM)

degradation alongwith their growth being monitored at 600 nm (using UV-VIS

Spectrophotometer) at different pHs (6, 7, and 8) and incubation temperatures (25, 30

and 37◦C) for 24 hrs. The isolates and their consortia exhibited good potential to degrade

DM (200 mg/l) under wider range of pH and temperatures with 40.01 to 77.69%

degradation. The MB490 and MB497 proved best DM degraders at all pHs (6, 7 and 8)

thus showing that DM degradation by these isolates was pH independent. Strains

MB498 and MB504 showed best DM degradation at pH 7. All consortia showed

maximum DM degradation in alkaline pH 8. Optimum temperature for DM degradation

by the isolates and their consortia ranged from 25- 30◦C. More DM degradation under

shaking conditions with more growth was exhibited by all isolates and consortia.

Isolates and their consortia showed 80.1 to 99.90% DM degradation in M-9 broth within

9 days, while it ranged from 77.67 to 93.06% in soil slurry and 77.89 to 89.99% in soil

microcosm. The GCMS analysis of bacterial isolates in M-9 broth, soil slurry and soil

microcosm at different intervals revealed that DM (RT = 17.7 min) was metabolized

into 5 products i.e Methyl diethanol amine, Phosphonothioic acid propyl-O, S-dimethyl

ester, O, O, O- Trimethyl thiophosphate, Omethoate and Aspartylglycine ethyl ester.

Background

Dimethoate is an organophosphate systemic insecticide used to control various insect

pests like Acari, Aphididae, Aleyrodidae, Lepidoptera etc. in cereals, fruits and

vegetables. It is also used for the control of domestic and veterinary insects (Hayes and

Laws, 1990). Dimethoate is toxic to mammals and is declared as a possible carcinogenic

agent in humans, as it caused tumors in exposed mice (US EPA, 2006). Being an

organophosphate, dimethoate is an acetyl cholinesterase inhibitor, and thus is a

neurotoxic agent (Tomlin, 2009). Dimethoate is depleted from the soil by the process

179

of leaching, evaporation and biodegradation with half-life ranging from 4-16 days

(Megeed and El-Nakieb, 2008). It is hydrolyzed to O, O-dimethyl dithiophosphoric acid

in plants and animals (Megeed and Mlilo, 2014). Dimethoate belongs to carbamate

group of organophosphate and thus is not easily degraded. The stability of dimethoate

in the environment depends upon pH, temperature and the type of medium. In the

environment, biotic degradation and metabolic reactions by microbes play a very

important role not only for the removal of the original pesticides, but also for

transformation of pesticide, thus changing their properties and affecting their spread in

various environmental sections (Megeed and El-Nakieb, 2008). Only few bacteria

having potential to degrade Dimethoate have been reported (Liu et al., 2001; Jiang et

al., 2007; DebMandal et al., 2008). The two DM degrading enzymes have been isolated

from Aspergillus niger and Klebsiella sp. (Liu et al., 2001; Jiang et al., 2007). In

Pakistan, Dimethoate and other OP pesticides are extensively being used to control

pests. This has resulted in contamination of agricultural soils, water resources and even

food crops. Earlier, there have been no reports on DM degrading bacteria in Pakistan.

This is the first study in Pakistan about isolation and characterization of indigenous soil

bacteria in order to evaluate their potential to tolerate and degrade DM under local

conditions and to identify the metabolites of Dimethoate formed so as to understand the

transformation pathway of DM in these selected isolates.


6.1 Optimization of environmental conditions for Dimethoate biodegradation

Multiple experiments were conducted to optimize the four isolates and their seven

consortia for Dimethoate (DM) degradation at different pH (6, 7, and 8) under shaking

conditions and at different incubation temperatures (25, 30 and 37◦C) under static

conditions within 24 hrs. The isolates and consortia were also monitored for their

growth under different conditions at 600 nm using UV-VIS Spectrophotometer. The

isolates and their consortia exhibited 40.01 to 77.69% DM degradation under wider

range of pH and temperatures. Strains MB490 and MB497 were best DM degraders at

all given pHs i.e 6, 7 and 8 indicating pH had no effect on DM degradation by these

isolates. Strain MB498 was best DM degrader at pH 7 and 8, while MB504 showed

maximum degradation at pH 7 (figure 6.1). In case of consortia, maximum DM

180

degradation was observed at alkaline pH8 (figure 6.3), thus indicating contribution of

alkaline phosphatases or other pH dependent degrading enzymes. Earlier Megeed and

El-Nakieb (2008) reported that acidic pH range support biotransformation of DM which

is in contrast to current study results.

Bacterial strains MB490 and MB497 exhibited maximum DM degradation both at 25

and 30◦C (figure 6.2). While MB498 and MB504 were best degraders of DM at 25◦C.

Consortia A and C exhibited maximum DM degradation at 25◦C, while consortia B, D,

E, F and G showed highest degradation at 30◦C with good growth (figure 6.4). Liang et

al. (2009) isolated a bacterium Raoultella sp. that could degrade DM at 30◦C and pH

6–8. Maximum abiotic DM degradation (2%) was observed at pH 7 and at 37◦C in the

control, thus demonstrating a vital contribution of bacterial isolates in DM degradation.

In some cases, bacterial growth pattern was not in accordance with their rate of

degradation, as least bacterial growth exhibited highest DM degradation and vice versa

(figure 6.1, 6.2, 6.3 & 6.4). This growth pattern may either be due to death of bacteria

by toxic metabolites formed during DM degradation or utilization of all bacterial

machinery and energy for the production of OP degrading enzymes rather than cell

division (Yale et al., 2017). All isolates and consortia exhibited better DM degradation

(49.41- 64.99%) under shaking conditions with more growth after 24 hrs in contrast to

static conditions of incubation (31.87-50.87%), which may be due to more aeration

supporting aerobic bacterial growth and producing more degrading enzymes as well as

shaking conditions, increase solubility and availability of pesticide as depicted (figures

6.5 & 6.6).

181

Figure 6.1 Effect of pH on growth (OD600 nm) and % degradation of Dimethoate by

four isolates (MB490, MB497, MB498 and MB504) after 24 hrs. Error bars represent

standard errors for values of three replicates.

Figure 6.2 Effect of temperature on growth (OD600 nm) and % degradation of

Dimethoate by four isolates (MB490, MB497, MB498 and MB504) after 24 hrs. Error

bars represent standard error for values of three replicates.

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3


MB490 MB497 MB498 MB504

% D

egra

dat

ion

of

DM

OD

60

0n

m

pHs


10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

25 30 37 25 30 37 25 30 37 25 30 37

MB490 MB497 MB498 MB504

%d

egra

dat

ion

of

DM

OD

60

0n

m

Temperatures (°C)


182

Figure 6.3 Effect of pH on growth (OD600 nm) and % degradation of Dimethoate 7

consortia (A, B, C, D, E, F, and G) after 24 hrs. Error bars represent standard errors for

values of three replicates.

Figure 6.4 Effect of temperature on growth (OD600 nm) and % degradation of

Dimethoate by 7 consortia (A, B, C, D, E, F, and G) after 24 hrs. Error bars represent

standard errors for values of three replicates.

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

6 7 8 6 7 8 6 7 8 6 7 8 6 7 8 6 7 8 6 7 8

A B C D E F G

% D

egra

dat

ion

OD

60

0n

m

pHs


10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

25

30

37

25

30

37

25

30

37

25

30

37

25

30

37

25

30

37

25

30

37

A B C D E F G%

Deg

rad

atio

n o

f D

M

OD

60

0n

m

Temperatures °C


183

Figure 6.5 Effect of Shaking versus static conditions on growth and % degradation of

Dimethoate by four isolates (MB490, MB497, MB498 and MB504) after 24 hrs. Error

bars represent standard errors for values of three replicates.

Figure 6.6 Effect of Shaking versus static conditions on growth and % degradation of

Dimethoate by seven consortia (A, B, C, D, E, F, and G) after 24 hrs. Error bars

represent standard errors for values of three replicates.

6.2 Effect of incubation period on bacterial growth and % degradation of DM in

M-9 broth

Isolates MB490, MB497, MB498, MB504 and their consortia (A, B, C, D, E, F and G)

were grown in M-9 broth supplemented with 200 mg/l of DM and analyzed

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3

MB490 MB497 MB498 MB504 MB490 MB497 MB498 MB504

Shaking conditions static conditions

%D

M D

egra

dat

ion

OD

60

0n

m

%Degradation Growth

10

30

50

70

90

110

0

0.5

1

1.5

2

2.5

3



% D

egra

dat

ion

of

DM

OD

60

0n

m


184

quantitatively using HPLC, after 1, 3, 6 and 9 days of incubation. All the strains and

their consortia showed very efficient DM degradation increasing with increase in

incubation period reaching its maximum (80.1-92% for isolates and almost 99% for

consortia) after 9 days of incubation (figure 6.7 & 6.8). After 6 days, the bacterial

growth declined up to 9 days, due to death of bacterial cells resulting from nutritional

competition and toxic products of degradation. The rate of degradation seemed

maximum during initial 24 hrs which may be linked to log phase of bacterial growth

(Gray, 1989; Jilani, 2013). The decrease in degradation rate after 6 and 9 days may be

related to pH change in the medium as a result of metabolite production, which may

denature the degrading and other vital enzymes. Siddique et al. (2003) suggested that

microbial strains considerably reduced the pH of growth medium from 7.2 to 3.2 during

15 days of incubation. Li et al. (2010) isolated Paracoccus sp. strain Lgjj-3 from waste

water that could completely degrade 100 mg/l of DM in broth culture within 6 hrs.

There are reports that Dimethoate can be biodegraded up to about 50% during 3 to 5

days in soil and river water (Thapar et al., 1995). Liang et al. (2009) reported that isolate

Raoultella sp. X1could degrade only 27% of DM in 10 days, when DM (200 mg/l) was

provided as only source of carbon and nitrogen. Madhuri (2014) reported that

Pseudomonas sp. and Bacillus sp. could degrade Dimethoate (0.24 mg/ml) up to 88 and

92% after 72 hrs which is in agreement to present study results.

Moreover, bacterial isolates and their consortia were able to metabolize DM and use it

as a nutrient for their growth in a growth linked process. The toxic compounds need a

minimum level of cell density for their significant biodegradation. This strategy applies

to both pure bacterial cultures as well as microbial consortia (Tharakan and Gordon,

1999). Further, they had tendency for rapid DM degradation in minimal medium

without glucose so that these isolates had to depend upon the DM for their carbon and

energy needs. Similarly, Megeed and El-Nakieb (2008) reported the complete removal

of 120 mg/ml of Dimethoate from aqueous medium by effective microbial consortia

within 3 days. On the other hand, DM degradation in the control was negligible with

only 7, 8, 9.6 and 11% degradation after 1, 3, 6 and 9 days of incubation respectively.

Therefore, in the current study, all the isolates and consortia have great potential to

eradicate DM from contaminated sites and can be applied for bioremediation.

185

Similar to CPF and TAP degradation, the biodegradation of DM followed a biphasic

model where more rapid rate of removal was observed in the first phase i.e up to 3 days

of incubation followed by slower rate in the second phase up to 9 days. Same case was

reported by Abdurruhman et al. (2015) for degradation of mixture of Atrazine and

Pendimethalin by Pseudomonas pickettii where maximum degradation had been done

during 1st three days of incubation thus attaining the equilibrium. The current study

results indicated a strong correlation between microbial growth and degradation

processes.

Figure 6.7 Effect of incubation period on growth and degradation of Dimethoate (DM)

by the 4 isolates (MB490, MB497, MB498, MB504) in the M-9 broth (initial

concentration of Dimethoate used = 200 mg/l). Error bars represent standard errors for


Figure 6.8. Effect of incubation period on growth and degradation of Dimethoate (DM)

by seven consortia (A, B, C, D, E, F, and G) of bacterial isolates in the M-9 broth (initial

concentration of Dimethoate used = 200 mg/l). Error bars represent standard errors for


1030507090110

00.5

11.5

22.5

3

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4

MB

49

0

MB

49

7

MB

49

8

MB

50

4


% D

egra

dat

ion

of

DM

OD

60

0n

m

Incubation period% Degradation growth

10

30

50

70

90

110

00.5

11.5

22.5

3


1 day 3 days 6 days 9 days % D

egra

dat

ion

of

DM

OD

60

0n

m

Incubation period


186

6.3 Effect of incubation period on bacterial growth and % degradation of DM in

the soil slurry

The bacterial isolates and their consortia were cultured in soil slurry supplemented with

200 mg/l of DM for up to 9 days to study effect of incubation period on DM degradation

using HPLC after 3, 6 and 9 days of shaking at pH 7 and 37°C. Maximum DM

degradation by bacterial isolates (80% by MB504 to 91.8% by MB498) and their

consortia (77.67% by consortium D to 93.06% by consortium F) was achieved after 9

days of incubation in soil slurry (figure 6.9 & 6.10). The rate of degradation was rapid

in initial 3 days followed by decrease in degradation rate in next phase up to 9th day due

to different limiting factors like death of bacteria or/due to toxic metabolites or due to

competition for space and nutrients. DM degradation was biphasic and positively

correlated with growth of bacteria in terms of CFU/ml of soil slurry (Table 6.1) as most

rapid degradation occurred during log phase of bacterial growth (initial 3 days)

followed by stationary phase (decreased CFU of viable bacterial cells) where rate of

bacterial cell division was equal to rate of bacterial death. On the contrary, there was

only 3, 6 and 11% DM degradation in the control after 3, 6 and 9 days of incubation

respectively. Ishag et al. (2016) also reported the Dimethoate degradation by bacteria

to be biphasic with respect to given time period.

Figure 6.9 Effect of incubation period on degradation of Dimethoate (DM) by the 4 bacterial

isolates (MB490, MB497, MB498 and MB504) in the soil slurry (initial concentration of

DM used = 200 mg/l). Error bars represent standard errors for values of three replicates.

0

20

40

60

80

100

MB490 MB497 MB498 MB504

% D

egra

dat

ion

of

DM

Incubation time


187

Table 6.1 Bacterial Population in soil slurry spiked with 200 mg/l of DM

Bacterial

isolates

CFU/ml of soil slurry


MB490 1.6×106 ± 0.05 5.7×106 ± 0.05 4.5×106 ± 0.05 3.5×106 ± 0.1

MB497 1.5×106 ± 0.1 5.5×106 ± 0.1 3.5×106 ± 0.05 2.5×106± 0.05

MB498 1.7×106 ± 0.1 5.4×106 ± 0.05 4.4×106 ± 0.1 2.4×106 ± 0.05

MB504 1.4×106 ± 0.05

5.6×106 ± 0.1 3.6×106 ± 0.05 2.6×106 ± 0.1

Each value is the mean of three replicates; CFU = colony forming unit; ± values of

standard error.

Figure 6.10 Effect of incubation period on degradation of Dimethoate (DM) by the 7

consortia (A, B, C, D, E, F, and G) in the soil slurry (initial concentration of DM used = 200

mg/l). Error bars represent standard errors for values of three replicates.

6.4 Effect of incubation period on bacterial growth and % degradation of DM in the

soil microcosm

In order to study effect of incubation period on degradation of DM in soil microcosm,

bacterial isolates and their consortia were inoculated in 50 g of soil with 40% water holding

capacity supplemented with 200 mg/kg of DM in Petri plates and incubated under dark

conditions for up to 9 days. DM biodegradation was analyzed using HPLC after 3, 6 and 9

0

20

40

60

80

100

A B C D E F G Control% D

egra

dat

ion

of

DM

Incubation Time


188

days of incubation. DM degradation gradually reached maximum by bacterial isolates

(78.80 to 82.1%) and their consortia (77.89 to 89.99%) in soil microcosm within 9 days

(figure 6.11 & 6.12). The growth and survival of inoculated bacteria in soil microcosm was

checked by calculating CFU via serial dilution and spread plate method at 0, 3, 6 and 9 days

interval as given in Table 6.2. Generally, the DM degradation by the bacterial isolates and

consortia followed a biphasic model starting with faster and increasing rate in the first phase

of degradation up to 3 days due to bacterial log phase having plenty of nutrients followed

by a second phase of decreasing rate due to stationary phase of bacteria (indicated by

decreased CFU of viable cells in Table 6.2) till 9th day as a result of toxicity of some

metabolites and scarcity of nutrients under limited space of soil microcosm. This stationary

phase of bacteria in the presence of toxic metabolites corresponds to acclimation period that

prepares them to activate the production of degradative enzymes for metabolites (Jilani,

2013). Previously, biphasic biodegradation has also been reported for other pesticides (Rigas

et al., 2007; Ishag et al., 2016). On the other hand, there was only 4, 6 and 7% DM

degradation in the control sterilized soil (without any bacterial inoculum) and 6, 8 and 10%

DM degradation in the control unsterilized soil after 3, 6 and 9 days of incubation

respectively. Previously Dimethoate degradation by fungus Aspergillus niger was reported

by Liu et al. (2001). There are few earlier reports of Dimethoate degradation using bacterial

isolates (Deshpande et al., 2001; DebMandal et al., 2008).

Figure 6.11 Effect of incubation period on degradation of Dimethoate (DM) by the bacterial

isolates (MB490, MB497, MB498 and MB504) in the soil microcosm (initial concentration

of DM used = 200mg/kg of soil). Error bars represent standard errors for values of three

replicates.

0

20

40

60

80

100

MB490 MB497 MB498 MB504 Control soil Unsterilizedsoil

% D

egra

dat

ion

of

DM

Incubation time


189

Figure 6.12 Effect of incubation period on degradation of Dimethoate (DM) by the

consortia (A, B, C, D, E, F, and G) in the soil microcosm (initial concentration of DM

used = 200 mg/kg of soil). Error bars represent standard errors for values of three

replicates.

Table 6.2 Bacterial Population in soil microcosm spiked with 200 mg/kg of DM

Bacterial

isolates

CFU/g soil


MB490 1.5×106 ± 0.1 6.5×106 ± 0.05 3.2×106 ± 0.1 2.8×106 ± 0.1

MB497 1.4×106 ± 0.05 6.5×106 ± 0.1 4.2×106 ± 0.05 3.2×106 ± 0.05

MB498 1.8×106 ± 0.1 6.4×106 ± 0.05 4.1×106 ± 0.1 3.2×106 ± 0.1

MB504 1.6×106 ± 0.05

5.6×106 ± 0.1 3.2×106 ± 0.05 2.5×106 ± 0.1

Each value is the mean of three replicates; CFU = colony forming unit; ± values of standard

error.

6.5 Quantitative analysis through HPLC for Dimethoate biodegradation

6.5.1 Effect of pH on DM degradation

6.5.1a Effect of pH on DM degradation by isolate MB490

The MB490 exhibited almost equal peak reduction in all given pHs (6, 7, 8) at retention time

of 2.9 minutes, thus indicating its efficiency to degrade Dimethoate under wider range of

pH after 24 hrs in M-9 broth (figure 6.1

0

20

40

60

80

100

% D

egra

dat

ion

of

DM

Incubation time


190

a. b.

c d.

Figure 6.13. a) HPLC chromatogram of Dimethoate control. b). HPLC chromatogram

of Dimethoate degraded by MB490 in M-9 broth after24hrs at pH 6. c). pH 7. d). pH 9.

6.5.1b Effect of pH on DM degradation by isolate MB497

In case of MB497, DM peak (RT = 2.9 min) is almost equally reduced in pH 6, 7 and

8 as compared to control, thus indicating equal degradation at all given pH as given in

figure 6.14.



Ab

sorb

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(m

AU

)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

191

a. b.

c. d.


of Dimethoate degraded by MB497 in M-9 broth after24hrs at pH 6. c). pH 7. d). pH 9.

6.5.1b Effect of pH on DM degradation by isolate MB498

Isolate MB498 showed same and little more decrease of DM peak (RT = 2.9 min) at

pH 7 and 8 as compared to pH 6 as given in figure 6.15. There are some other peaks at

RT = 1.8 and 1.17 min indicating formation of metabolites.



Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

192

a. b.

c. d.


of Dimethoate degraded by MB498 in M-9 broth after 24 hrs at pH 6. c). pH 7. d). pH

9.

6.5.1d. Effect of pH on DM degradation by isolate MB504

Strain MB504 demonstrated maximum peak reduction of DM at pH 7 followed by pH

6 and pH 8 (figure 6.16).



Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

193

a. b.

c. d.


of Dimethoate degraded by MB504 in M-9 broth after 24 hrs at pH 6. c). pH 7. d). pH

9.

6.5.2 Effect of temperature on DM degradation

6.5.2a Effect of temperature on DM degradation by isolate MB490

Strain MB490 demonstrated nearly equal and sharp DM peak reduction at 25 and 30°C,

showing best DM degradation at these two temperatures (figure 6.17b and c) followed

by considerable peak decrease at 37°C (figure 6.17d).



Ab

sorb

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(m

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(m

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)

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(m

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)

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sorb

ance

(m

AU

)

194

a. b.

c. d.


of Dimethoate degraded by MB490 in M-9 broth after 24 hrs at 25°C. c). 30°C. d).

37°C.

6.5.2b Effect of temperature on DM degradation by isolate MB497

Similarly in case of MB497, there was great and equal depletion of DM peak at 25 and

30°C followed by peak reduction at 37°C corresponding to their DM degradation at

these temperatures (figure 6.18b, c, d).



Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

) A

bso

rban

ce (

mA

U)

Ab

sorb

ance

(m

AU

)

195

a. b.

c. d.



37°C.

6.5.2c Effect of temperature on DM degradation by isolate MB498

Likewise MB498 was more efficient degrader of DM at 25°C, thus showing significant

reduction of DM peak at this temperature (figure 6.19b). There was less reduction in

peak at 30°C along with metabolite peaks prominent at 1.3, 1.9 and 2.8 min (figure

6.19c).



Ab

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(m

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Ab

sorb

ance

(m

AU

)

Ab

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(m

AU

)

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(m

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)

196

a. b.

c. d.



37°C.

6.5.2d Effect of temperature on DM degradation by isolate MB504

In case of MB504, a sharp decrease in DM peak (RT = 2.9 min) was noticed at 25°C

followed by 37 and 30°C (figure 6.20b, c, d) corresponding to maximum degradation

at 25°C and minimum at 30°C.



Ab

sorb

ance

(m

AU

)

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(m

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)

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(m

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)

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(m

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197

a. b.

c. d.



37°C.

6.5.3 DM degradation in soil microcosm by four isolate (MB490, MB497, MB498

and MB504) after 9 days of incubation

All the isolates demonstrated very sharp and prominent decline of DM peak (RT = 2.9

min) after 9 days of incubation, indicating maximum depletion of DM by bacteria along

with metabolite peaks appearing at retention time of 1.17 and 1.8 min (figure 6.21b, c,

d & e).



Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

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)

Ab

sorb

ance

(m

AU

)

Ab

sorb

ance

(m

AU

)

198

a. b.

c. d.

e.

Figure 6.21. a) HPLC chromatogram of Dimethoate control. HPLC chromatogram of

Dimethoate degraded by: b). MB490. c). MB497. d). MB498. e).MB504 in soil

microcosm after 9 days of incubation




Ab

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(m

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(m

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(m

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(m

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199


Dimethoate (DM) by the 4 isolates and their consortia in the M-9


The bacterial cultures (M-9, soil slurry and soil microcosm) treated with Dimethoate

(200 mg/l/kg) were analyzed by GCMS after 3, 6 and 9 days of incubation. In all

bacterial samples, total 6 main peaks were identified in the chromatograms for MB490,

MB497, MB498 and MB504 and their consortia with the help of NIST library, USA.

These compounds were identified with a resemblance percentage above 90% as follows

(figures 6.23, 6.24 & 6.25): Dimethoate, Methyl diethanol amine, Phosphonothioic

acid, propyl-O, S-dimethyl ester, O, O, O- Trimethyl thiophosphate, Omethoate and

Aspartylglycine ethyl ester (detected only in samples with MB490) (Table 6.3). The

control medium without bacteria incubated for 9 days produced no detectable

compound, other than dimethoate. Earlier in a study. seven metabolites of dimethoate

degradation, including dimethoate carboxylic acid, 2-(hydroxyl (methoxy)

phosphorylthio)acetic acid, O,O,S-trimethyl thiophosphorothioate, O-methyl O,S-

dihydrogen phosphorothioate, phosphorothioic O,O,S-acid, O,O,S-trimethyl

phosphorothiate and O,O,O-trimethyl phosphoric ester, were formed by Paracoccus sp.

strain Lgjj-3 (Li et al., 2010).

200

a.

b.

Figure 6.22 (a). GCMS chromatogram of Dimethoate standard (RT = 17.7 min). (b).

Mass spectrum of Dimethoate standard.


m/z values

Rel

ativ

e A

bu

nd

ance

In

ten

sity

Dimethoate

(standard)

RT= 17.7 min

201

Table 6.3 Biotransformation of Dimethoate (DM) by the 4 isolates (MB490,

MB497, MB498, and MB504) and the consortia of these 4 isolates in the M-9 broth,

soil slurry and soil microcosm.

Dimethoate and its metabolites detected by GCMS

S. no Name of

compound

Chemical structure Retention Time

(minutes)

m/z

1 Dimethoate

17.7 47, 87,

93, 125

2 Methyl diethanol

amine.

15.9 125

(M++H)

3 Phosphonothioic

acid,propyl-O,S-

dimethyl ester.

4.5

47, 79,

126

4 O, O, O- Trimethyl

thiophosphate

5

93,

126,

156

5 Omethoate

9.78

213.15

6, 110,

79

6 Aspartylglycine

ethyl ester

14.46 88

202

A.

B.

C.

D.

Figure 6.23 GCMS chromatogram of Dimethoate (RT = 17.7 min) degraded by: A).

MB490. B). MB497. C). MB498. D). MB504 in soil microcosm after 3 days of

incubation. Peaks in the chromatograms are identified as :a). Dimethoate (RT = 17.7

min). b). Methyl diethanol amine (RT= 15.9 min). c). Aspartylglycine ethyl ester (RT

= 14.46 min) detected in MB490 only. d). Omethoate (RT = 9.78min). e). O, O, O-

Trimethyl thiophosphate (RT = 5 min). f). Phosphonothioic acid,propyl-O,S-dimethyl

ester (RT = 4.5 min).

c

a

d e f

a

b d e f

b

203

a. b.

c. d.

e. f.

g. h.

Figure 6.24. GCMS chromatogram of Dimethoate (RT = 17.7 min) degraded by a).

Consortium A, b). Consortium B, c). Consortium C, d). Consortium D, e). Consortium

E, f). Consortium F, g). Consortium G, h). Mass spectrum of Dimethoate degraded by

bacteria in soil microcosm after 9 days of incubation. Peaks in the chromatograms are

identified as :a). Dimethoate (RT = 17.7 min). b). Methyl diethanol amine (RT= 15.9

min). d). Omethoate (RT = 9.78 min). e). O, O, O- Trimethyl thiophosphate (RT = 5

min). f). Phosphonothioic acid, propyl-O, S-dimethyl ester (RT = 4.5 min). While peak

c for Aspartylglycine ethyl ester (RT = 14.46 min) disappeared in these chromatograms.

a b

. e

c

d f




m/z values (40-220) Retention time (min)

Rel

ativ

e A

bu

nd

ance

Inte

nsi

ty

Inte

nsi

ty

Inte

nsi

ty

Inte

nsi

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Inte

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Inte

nsi

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Inte

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204

6.7 Proposed Metabolic Pathway for the Transformation of Dimethoate by

Bacteria

On the basis of above findings, metabolic pathway of Dimethoate transformed and

mineralized by bacteria can be suggested as given in figure 6.26. According to this

model, Dimethoate first oxidized to Omethoate which then followed two pathways. In

first pathway, (generally observed in all isolates and their consortia), it is first

hydrolyzed to Phosphonothioic acid, propyl-O, S-dimethyl ester and Methyl diethanol

amine by phosphatase and/or amidase enzyme. Methyl diethanol amine is further

mineralized by bacteria as a source of carbon and nitrogen. In second pathway observed

only in MB490, Methyl diethanol amine is further converted to Aspartylglycine ethyl

ester by amination which becomes assimilated by bacteria. Phosphonothioic acid,

propyl-O, S-dimethyl ester is very unstable and is oxidized to O, O, O-Trimethyl

thiophosphate which undergoes desulphurization and dephosphorylation by

phosphatase and is ultimately mineralized providing carbon, sulphur, phosphorus and

nitrogen to bacteria. Paracoccus sp. strain FLN-7 was able to use dimethoate as a

carbon source for growth by assimilating the small aliphatic compounds like

methylamine derived as a result of hydrolysis of DM via Arylamidase (Zhang et al.,

2012).

In a degradation pathway observed in certain microbes, dimethoate was degraded by

the hydrolysis of the phosphotriester bond (POS) by OPH which belongs to the

amidohydrolase superfamily that are capable of hydrolyzing multiple bonds like POO,

POF, and POS in OP pesticides (Horne et al., 2002). Another degradation pathway for

dimethoate involves reduction of the P=S linkage by an aldo-keto reductase (Jiang et

al., 2007). DebMandal et al. (2008) reported many unknown metabolites of Dimethoate

formed during bacterial degradation. Moreover, the metabolic pathway of DM has also

been studied in mammals (Franca and Emanuela, 2007). Photocatalytic oxidation of

DM mediated by TiO2 was studied by Evgenidou et al. (2006). Similar to current study

results, O, O, S-trimethyl thiophosphorothioate was oxidized to O, O, O-trimethyl

phosphoric ester during three different processes i.e thermal, photochemical and

microbial degradation (Andreozzi et al., 1999; Evgenidou et al., 2006; Li et al., 2010).

The enzymes involve in the biodegradation of organophosphate pesticides include

phosphatase, esterase, hydrolase and oxygenase (Kanekar et al., 2004). The role of

205

eseterases especially carboxylamidases has been reported in the degradation of

dimethoate thus releasing methylamine (Hassal, 1990). Recently, Chen et al., (2016)

revealed the formation of Dimethoate carboxylic acid and Methylamine by

amidohydrolase in Sphingomonas sp. Omethoate was detected as only metabolite of

Dimethoate in pesticide treated cucumber fruit (Geng et al., 2018).

Figure 6.26. Proposed metabolic pathway for the transformation of Dimethoate

by bacteria. Metabolite in purple box (Aspartylglycine ethyl ester) was detected

only in MB490, while in green box (Methyl diethanol Amine) was detected in

all isolates and their consortia.

Conclusion

There are very few reports about biodegradation and transformation study of

Dimethoate in the world and none in Pakistan. The current study is the first report of its

kind in Pakistan about DM showing potential of indigenous bacteria to degrade DM

considerably and metabolize it into simpler products. Some novel metabolites of DM

have been detected by GCMS like Methyl diethanol amine and Aspartylglycine ethyl

ester and a metabolic pathway has been proposed in the current study which will

definitely help to understand fate of DM degraded by the bacterial cells as well as to

Dimethoate

Omethoate

Phosphonothioic acid,propyl-O,S-dimethyl ester.

O,O,O- Trimethyl thiophosphate

Methyl diethanol Amine

Aspartylglycine ethyl ester

Mineralization

Phosphatase

Amination Oxidation

Oxidation

Hydrolysis by Carboxyl Amidase

Phosphatase

206

study the enzymes and genes involved as a future perspective. Therefore these four

bacterial strains MB490 (Pseudomonas kilonensis), MB497 (Bacillus thuringiensis),

MB498 (Pseudomonas kilonensis) and MB504 (Pseudomonas sp.) can be

recommended as strong candidate for application to bioremediate DM contaminated

agricultural soils and water resources.

207

Chapter 7

Organophosphorus phosphatase (OPP) enzyme studies

Overview

Four bacterial strains MB490 (Pseudomonas kilonensis), MB497 (Bacillus

thuringiensis), MB498 (Pseudomonas kilonensis) and MB504 (Pseudomonas sp.) were

studied for the production of intracellular and extracellular organophosphorus

phosphatase enzyme (OPP). All bacterial strains were found positive for phosphate

solubilization and were producing significant amount of extracellular, acidic, neutral

and alkaline phosphatases. Largely, there was more production of acidic and alkaline

phosphatases as compared to neutral phosphatase. For three types of OPP,

Pseudomonas kilonensis MB490 showed maximum production of neutral OPP though

less than alkaline OPP. Bacillus thuringiensis MB497 exhibited maximum production

of alkaline OPP, whereas Pseudomonas sp. MB504 showed highest production of

acidic OPP. The strain Pseudomonas kilonensis MB498 could produce equal and

significant quantity of acidic and alkaline phosphatases. The highest OPP production

was observed at pH 11 by all isolates. Among four strains, Pseudomonas sp. MB504

showed best production of OPP from pH 6-11 (especially at pH 10 and 11). The

maximum OPP enzyme production was exhibited at 50°C by MB490, MB497 and

MB498, whereas at 45°C by MB504. All strains showed minimum enzyme production

at 37°C. In general, OPP production was decreased by all isolates after 48 hrs.

The OPP enzyme activity and stability was maximum at 37⁰C followed by decrease at

higher temperatures but remained still active even at highest given temperature (70⁰C)

by all isolates. Generally, SDS and EDTA proved inhibitors for acidic, neutral and

alkaline OPP activity especially at higher concentrations, while, Zn++, Cu++ and Cd++

greatly enhanced OPP activity. There was 86-100% bioprecipitation of selected metals

(Ni, Mn, Cr and Cd) by all three OPPs. Acidic OPP exhibited 40 to 80% degradation

of given 50 mg/l of three OP pesticides (CPF, TAP and DM), after 30 min of incubation,

while alkaline OPP showed more degradation (80 to 99%).

208

Back ground

The conventional methods used for the removal of OP compounds include physical and

chemical treatment, incineration and deep ocean dumping which are both costly and

pose environmental threats such as release of toxic chemicals (Richins et al., 2000).

Therefore, recently research is focused on new techniques like enzymatic detoxification

in order to remove the OP pesticide residues, which is cheap as well as environmental

friendly (Cheng et al., 1999). It is need of the day to study new enzymes and

biocatalysts that are stable over a wide range of temperature, pH, salts etc. and that do

not represent an environmental hazard in order to find a better solution for degrading

OP compounds. These enzyme characteristics are important in order to determine the

potential use of enzymes in future applications.

La Nauze (1970) isolated the first enzyme capable of degrading phosphonates, from B.

cereus and named as 2-phosphonoacetaldehyde hydrolase (phosphonatase). Its optimal

activity was observed at pH 8 and it is similar to alkaline phosphatase in many respects

except having narrow substrate specificity. According to Kononova and Nesmeyanova,

(2002), phosphonatase could not degrade phosphomonoesters. Yet it is able to degrade

variety of phosphonates like glyphosate and has been isolated from many bacterial

species (Baker et al., 1998). Mazur (1946) isolated the first OP-degrading enzymes

(DFPases and sarinases) from rabbit and human tissue extracts that were able to

hydrolyze diisopropylfluorophosphate (DFP) and Sarin (nerve agents). Later on, in

1992, these were included among Phosphoric triester hydrolases by the Nomenclature

Committee of the International Union of Biochemistry and Molecular Biology (Hill et

al., 2001). Among OP-degrading enzymes, the organophosphorus hydrolases (OPH)

also known as phosphotriesterases isolated from Pseudomonas diminuta and OPAA

from Alteromonas sp. strain JD6.5 are the well-characterized enzymes (Theriot and

Grunden, 2011).

Usually, enzymes exhibit great sensitivity to pH changes. The enzyme activity can be

affected by pH in various ways. The ionization of the enzyme substrate complex can be

altered by pH or, the ionization of various groups within enzyme molecule may be

changed, thus affecting the enzyme-substrate affinity. Moreover, it may affect the

ionization of the substrate itself, thus changing its affinity to the enzyme. Further,

protein structure of enzyme may be denatured at extreme pH (Palmer, 1995).

209

The degradation potential of bacterial isolates (MB490, MB497, MB498 and MB504)

against three OP pesticides (CPF, TAP and DM) has been confirmed on the basis of

HPLC and GCMS analyses (results discussed in chapter 4, 5 and 6 respectively along

with the detection of their metabolites), thus strongly indicating the presence of OP

degrading phosphatases (involved in cleavage of P-O-C bond of these pesticides) in the

four bacterial isolates. Thus an enzyme assay was carried out to check the potential of

these isolates to produce acidic, neutral and alkaline organophosphorus phosphatases

(OPPs). Further the isolated crude extracts of OPP enzymes were optimized for their

production, activity and stability against different factors like pH, temperature,

substrate concentrations, incubation time and different metals. The bioprecipitation

potential of OPP was also tested against different metals. Finally, the acidic and alkaline

OPP were cross checked for their substrate specificity against CPF, TAP and DM.

Results are discussed below in detail.


7.1 Screening for phosphate solubilization potential of bacterial isolates

The bacterial isolates (MB490, MB497, MB498 and MB504) were grown on National

Botanical Research Institute’s phosphate (NBRIP) medium supplemented with calcium

phosphate as a substrate. All the isolates exhibited positive results by forming clear

zones of phosphate solubilization (Table 7.1). These zones were measured using

phosphate solubilization index (Premono et al., 1996). Islam et al. (2007) reported

varying levels of phosphate solubilizing activity in 6 isolates using National Botanical

Research Institute’s phosphate medium in both agar plate and broth assays. Generally,

the greater intensity of dissolved phosphate indicates greater phosphatase production

and there is positive correlation between the two (Sakurai et al., 2008).

Table 7.1 Screening for Phosphate solubilization potential of bacterial isolates

(MB490, MB497, MB498 and MB504).

Incubation

period

diameter (mm) of Halo zone

MB490 MB497 MB498 MB504

Day 3 14 10 24 26

Day 6 15 12 25 28

Day 9 16 13 26 30

210

7.2 Screening of extracellular Organophosphorus phosphatase (OPP) production

in bacterial isolates (MB490, MB497, MB498 and MB504)

Generally, there was less production of neutral phosphatase as compared to acidic and

alkaline phosphatases. The strains MB490 and MB497 exhibited maximum production

of alkaline phosphatase (ALP) followed by acidic phosphatase, whereas MB504

produced more acidic than alkaline phosphatase. Isolate MB498 could produce equal

amount of acidic and alkaline phosphatases (figure 7.1). The bacterial requirements for

inorganic phosphorus (Pi) are satisfied by the phosphorus removal from

organophosphate and phosphonate sources (Kriakov et al., 2003). ALPs are considered

the most promising dephosphorylating enzymes in the periplasmic space thus providing

the Pi cell demand (Manoil et al., 1990; Kriakov et al., 2003).

7.3 Screening of intracellular Organophosphorus phosphatase (OPP) in bacterial


All the four isolates were also tested for production of intracellular acidic, neutral and

alkaline OPP. It was noticed that the intracellular OPP production and activity was

much reduced as compared to extracellular phosphatases (figure 7.2), so further enzyme

assay was conducted with only extracellular phosphatases due to their better

production. Pesticide biodegradation is catalyzed by either extracellular or intracellular

enzymes (Tang and You, 2012). Among the intracellular phosphatases, MB490

produced more acidic OPP followed by alkaline and neutral OPP respectively i.e acidic

> alkaline > neutral, while MB497 and MB498 showed more production of alkaline

OPP followed by acidic and neutral respectively (Alkaline >acidic>neutral). In case of

MB504, production of OPP followed the sequence neutral > alkaline > acidic.

211

Figure 7.1 Extracellular production of Organophosphorus-phosphatase (OPP) by

bacterial isolates (MB490, MB497, MB498 and MB504) at 405 nm. Error bars


Figure 7.2 Intracellular production of Organophosphorus-phosphatase (OPP) by

bacterial isolates (MB490, MB497, MB498 and MB504) measured at 405 nm. Error


7.4 Factors affecting the production of extracellular Organophosphorus

Phosphatase (OPP)

7.4a Effect of pH on extracellular OPP production by bacterial isolates (MB490,

MB497, MB498 and MB504)

The four bacterial isolates were grown in NBRIP broth under variable pH range (6, 7,

8, 9 10 and 11) for 24 hrs and analyzed for OPP production. It was observed that OPP

production was highest at pH 11 by all the four isolates. OPP enzyme production

increased with increasing pH from 6 to 11. Strain MB504 was best among the four

isolates with respect to enzyme production at all pH especially at pH 10 and 11 (figure

7.3). Naturally, alkaline phosphatases require about neutral pH environment for

production, yet these exhibit maximum catalytic activity at pH 8 or above (Rina et al.,

0

10

20

30

MB490 MB497 MB498 MB504

Enzy

me

acti

vity

(u

/ml)

Acidic Phosphatase Neutral Phosphatase Alkaline Phosphatase

0

1

2

3

4

MB490 MB497 MB498 MB504Enzy

me

acti

vity

(u

/ml)

Acidic Phosphatase Neutral Phosphatase Alkaline Phosphatase

212

2000). The alkaline phosphatase isolated from Rhizobium demonstrated a wide range

of pH optimum (6.8-11.8) for the enzymatic reaction with p-NPP having maximum

activity at pH 9.8 (kumar et al., 2008). Bacterial alkaline phosphatases are reported to

have broader pH range which can go beyond physiological pH range when assayed with

p-NPP (Mori et al., 1999). The ALP isolated from Pyrococcus abyssi was reported as

most alkaline working best at pH 11 (Zappa et al., 2001). In the current study maximum

OPP production was observed at pH 11 by MB504, indicating its alkaline behavior.

Figure 7.3 Organophosphorus-phosphatase production by bacterial isolates (MB490,

MB497, MB498 and MB504) at different pHs. Error bars represent standard errors for


7.4b Effect of temperature on extracellular OPP production by bacterial isolates

(MB490, MB497, MB498 and MB504)

In order to study the effect of temperature on the production of organophosphorus

phosphatase, four isolates were grown in NBRIP broth at different temperatures (37,

45, 50 and 60°C) in a rotary shaker for 24 hrs. Among them MB490, MB497 and

MB498 exhibited maximum OPP enzyme production at 50°C, whereas MB504 showed

highest enzyme production at 45°C (figure 7.4). There was minimum enzyme

production at 37°C by all the isolates. Shah et al. (2008) reported an alkaline

phosphatase isolated from Bacillus subtilis KIBGE-HAS, which exhibited maximum

activity at pH 8 and 37°C.

0

10

20

30

40

50

pH 6 pH 7 pH 8 pH 9 pH 10 pH 11Enzy

me

acti

vity

(u

/ml)

MB490 MB497 MB498 MB504

213

Figure 7.4 Organophosphorus phosphatase production by bacterial isolates (MB490,

MB497, MB498 and MB504) at different temperatures. Error bars represent standard


7.4c Effect of incubation time on OPP production by bacterial isolates (MB490,

MB497, MB498 and MB504)

In order to study effect of incubation period on the production of acidic, neutral and

alkaline phosphatases, the bacterial isolates (MB490, MB497, MB498 and MB504)

were cultured in NBRIP broth for different time intervals (24, 48 and 72 hrs).

Figure 7.5 Effect of incubation time on Organophosphorus-phosphatase production by

bacterial isolates (MB490, MB497, MB498 and MB504). Error bars represent standard


Acidic and neutral OPP production after 24 hrs, followed the pattern as MB498>

MB497>MB490>MB504, while in case of alkaline OPP production was as

MB504>MB498> MB497> MB490 (figure 7.5). After 48 hrs, acidic, neutral and

alkaline OPP production decreased by all the isolates, whereas after 72 hrs incubation,

010203040506070

37⁰C 45⁰C 50⁰C 60⁰CEn

zym

e ac

tivi

ty (

u/m

l)

Temperatures

MB490 MB497 MB498 MB504

0123456

Acidic Neutral Alkaline Acidic Neutral Alkaline Acidic Neutral Alkaline


OP

P a

ctiv

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(u/m

l)

Incubation Time

MB490 MB497 MB498 MB504

214

it increased a little, especially alkaline OPP production was maximum by MB498.

Mahesh et al. (2015) reported the optimum production of extracellular alkaline OPP

derived from Proteus mirabilis M54 at pH 7-8 and 35°C for 28 hrs incubation time.

The classification of phosphatases into acid, neutral and alkaline phosphatase is based

on the optimum pH required for their catalytic activity (Shahbazkia et al., 2010). Their

production has been reported by prokaryotes as well as eukaryotes and are recognized

to transform insoluble organic phosphorous into soluble and available phosphorous

during their active cellular metabolism (Eshanpour and Amini, 2003; Amlabu et al.,

2009).

7.5 Factors affecting Organophosphorus Phosphatase (OPP) Activity

7.5a Effect of temperature on Organophosphorus phosphatase enzyme activity

and stability

There was considerable enzyme activity and stability observed over a wide range of

temperatures (37, 45, 50, 60 and 70⁰C) as discussed in detail below (figures 7.6 and

7.7). There was decreased enzyme activity by all isolates at higher temperatures but

still active even at 70⁰C. Previously, it was reported that Alkaline phosphatases (ALP)

from the mesophilic Escherichia coli and Pyrococcus abyssi showed activity at 80 and

105°C respectively (Janeway et al., 1993; Zappa et al., 2001). Behera et al. (2017)

studied alkaline phosphatase activity isolated from phosphate solubilizing Alcaligenes

faecalis and reported maximum activity at pH 9.0, at 45 °C and substrate concentration

of 1.75 mg/ml. Shah et al. (2008) reported that the enzyme activity of alkaline

phosphatase was maximum at 37°C and decreased at higher temperatures. The kinetic

energy of the protein molecules in enzyme structure is increased at high temperatures,

breaking the bonds between the active amino acids, thus enzyme activity is lost (Bryan

and Keith, 1994).

7.5a.1 Pseudomonas kilonensis MB490

For MB490, maximum acidic OPP activity was found at 60⁰C, while both neutral and

alkaline OPP activities were highest at 37⁰C (figure 7.6a).

http://www.scialert.net/asci/result.php?searchin=Keywords&cat=&ascicat=ALL&Submit=Search&keyword=Escherichia+coli

215

7.5a.2 Bacillus thuringiensis MB497

In case of MB497, highest acidic and alkaline OPP activity was noticed at 37⁰C while

neutral OPP was most active at 50⁰C (figure 7.6b).

7.5a.3 Pseudomonas kilonensis MB498

Likewise, for MB498, acidic and neutral OPP showed its maximum activity at 37⁰C,

while alkaline OPP was most active at 70⁰C (figure 7.7a).

7.5a.4 Pseudomonas sp. MB504

For MB504, maximum activity of acidic and alkaline OPP was noticed at 37⁰C,

whereas for neutral OPP it was observed at 60⁰C (figure 7.7b).

a.

b.

Figure 7.6 Effect of temperature on activity of Organophosphorus-phosphatase

produced by bacterial isolates (a). MB490, (b). MB497. Error bars represent standard


0

6

12

18

24

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Temperatures (°C)

216

a.

b.

Figure 7.7 Effect of temperature on activity of Organophosphorus-phosphatase

produced by bacterial isolates (a). MB498 (b). MB504. Error bars represent standard


7.5b Effect of substrate (p-NPP) concentration on OPP activity

The activity of OPP was measured at different concentration of substrate (p-NPP) i.e

0.06, 0.6, 0.8, 1.1%. Detailed account of OPP activity at different concentrations of p-

NPP for different isolates is given below:

7.5b.1 Strain MB490

In case of MB490, maximum acidic OPP activity was found at 0.6% p-NPP, while

neutral and alkaline OPP activity was observed at 1.1 and 0.8% respectively (figure

7.8a). It was reported that alkaline phosphatase exhibits greater substrate specificity for

p-nitro phenyl phosphate (Zappa et al., 2001; Boulanger and Kantrowitz, 2003). Shah

et al. (2008) also revealed that the alkaline phosphatase activity was increased with the

increase in substrate concentration.

0

6

12

18

24

Aci

dic

Ne

utr

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alin

e

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37 45 50 60 70

OP

P a

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(u/m

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Temperatures °C

0

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24

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37 45 50 60 70

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Temperatures (°C)

217

7.5b.2 Strain MB497

For MB497, highest acidic OPP activity was measured at 0.6%, while neutral and

alkaline were most active at 0.8 and 0.06% respectively (figure 7.8b).

7.5b.3 Strain MB498

Likewise, for MB498, acidic OPP showed its maximum activity at 0.06%, while neutral

and alkaline OPP were most active at 0.6 and 0.8% respectively (figure 7.9a).

7.5b.4 Strain MB504

In case of MB504, maximum activity for acidic OPP was noted at 0.06%, whereas for

neutral and alkaline OPP it was observed at 1.1 and 0.8% respectively (figure7.9b).

a.

b.

Figure 7.8 Effect of substrate (p-NPP) concentration on activity of Organophosphorus-

phosphatase produced by bacterial isolates (a). MB490 (b). MB497. Error bars


010203040

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218

a.

b.

Figure 7.9 Effect of substrate (p-NPP) concentration on activity of Organophosphorus-

phosphatase produced by bacterial isolates (a). MB498 (b). MB504. Error bars


7.5c Effect of incubation time on OPP activity

Detailed account of OPP activity at different incubation times (50, 70, 90 min) for

different isolates is given below:

7.5c.1 Strain MB490

In case of MB490, maximum acidic OPP activity was found after 50 min of incubation

time, whereas highest neutral and alkaline OPP activity was observed after 90 min

(figure 7.10a).

7.5c.2 Strain MB497

For isolate MB497, highest acidic, neutral and alkaline OPP activity was noticed after

70min of incubation period (figure 7.10b).

0

10

20

30

40

Aci

dic

Ne

utr

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e

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0.06 0.6 0.8 1.1O

PP

act

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y (u

/ml)


0

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0.06 0.6 0.8 1.1

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219

7.5c.3 Strain MB498

Likewise, for MB498, acidic OPP showed its maximum activity after 90 min, while

neutral and alkaline OPP were most active after 70 min of incubation (figure 7.11a).

7.5c.4 Strain MB504

In case of MB504, maximum activity for acidic OPP was noted after 50 min, whereas

for neutral and alkaline OPP it was observed after 70 and 90 min of incubation

respectively (figure 7.11b). It was revealed by Shah et al. (2008) that enzyme activity

of alkaline phosphatase was markedly decreased with increase in incubation time. This

may be due to the thermal sensitive nature of the enzyme with respect to time and

increased time period enhances the temperature of reaction mixture, which may break

the bonds between the amino acids (Hulett-cowling and Campbell, 1971).

a.

b.

Figure 7.10 Effect of incubation period on activity of Organophosphorus-phosphatase



0

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40

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a.

b.

Figure 7.11 Effect of incubation period on activity of Organophosphorus-phosphatase



7.6 Study of stimulatory or inhibitory effect of chemicals (SDS, EDTA, metals) on

OPP Activity

7.6.1 Effect of Sodium dodecyl sulphate (SDS) on OPP Activity

The effect of different concentrations (2.5, 5 and 7.5%) of SDS on OPP activity for

selected isolates (MB490, MB497, MB498 and MB504) was also checked and detail is

given below:

7.6.1a Strain MB490

In case of MB490, acidic and alkaline OPP activity was found to decrease gradually

thus inhibited at all given concentrations (2.5, 5 and 7.5%) after 30min of incubation

time, whereas neutral OPP activity was enhanced up to 2.5% SDS followed by

inhibition at higher SDS concentrations (figure 7.12a).

0

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221

7.6.1b Strain MB497

For isolate MB497, acidic OPP activity was noticed to increase at all given

concentrations of SDS after 30min of incubation period (figure 7.12b), while neutral

OPP activity was enhanced up to 2.5% followed by inhibition at higher concentrations

of SDS. Alkaline OPP activity remained almost unaffected by SDS at all concentration.

7.6.1c Strain MB498

For MB498, acidic and neutral OPP activity was increased by 2.5% SDS followed by

inhibition at higher concentrations, while alkaline OPP activity was enhanced by 2.5%

SDS, after which it remained unaffected by higher SDS concentrations of 5 and 7.5%

(figure 7.12c).

7.6.1d Strain MB504

In case of MB504, acidic and neutral OPP activity was inhibited at all given SDS

concentrations, whereas alkaline OPP activity was increased a little bit at 2.5% SDS

but then it remained unaffected at higher concentrations (figure 7.12d).

222

a.

b.

c.

d.

Figure 7.12 Effect of SDS on activity of Organophosphorus-phosphatase produced by

bacterial isolates (a). MB490 (b). MB497 (c). MB498 (d). MB504. Error bars represent

standard errors for values of three sample replicates.

0

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223

7.6.2 Effect of EDTA on OPP Activity

The study was conducted to check the effect of different concentrations (2.5, 5 and

7.5%) of EDTA on OPP activity for selected isolates (MB490, MB497, MB498 and

MB504) and detail is given below (figures 7.13).

7.6.2a Strain MB490

In case of MB490, acidic OPP activity remained unaffected at 2.5% of EDTA but

inhibited at higher concentrations (5 and 7.5%) after 30min of incubation time, whereas

neutral and alkaline OPP activity was inhibited and decreased at all given

concentrations of EDTA (figure 7.13a).

7.6.2b Strain MB497

For MB497, acidic, neutral and alkaline OPP activities were inhibited by all

concentrations of EDTA (2.5, 5 and 7.5%) after 30min of incubation period (figure

7.13b).

7.6.2c Strain MB498

Likewise in case of MB498, acidic, neutral and alkaline OPP activities were decreased

and inhibited at all concentrations (2.5, 5 and 7.5%) of EDTA (figure 7.13c).

7.6.2d Strain MB504

In case of MB504, acidic OPP activity was promoted by 2.5 and 5% of SDS followed

by inhibition at higher concentration, whereas neutral and alkaline OPP activities were

inhibited at all given concentrations of EDTA (figure 7.13d).

224

a.

b.

c.

d.

Figure 7.13 Effect of EDTA on activity of Organophosphorus-phosphatase produced

by bacterial isolates (a). MB490 (b). MB497 (c). MB498 (d).MB504. Error bars


01020304050

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225

7.6.3 Effect of metals on OPP Activity

The inhibitory/stimulatory effect of metals on the activity of acidic, neutral and alkaline

OPP produced by the bacterial isolates was studied by using 100 µl (2.5%) of selected

metals Zn++ (ZnSO4), Cu++ (CuSO4) and Cd++ (CdSO4), with the enzyme extract and

substrate mixture and incubated at 37⁰C for 30 min along with the control without

metal. It was noticed (after UV-VIS Spectrophotometer analysis at 405 nm) that acidic,

neutral and alkaline OPP activity from all isolates was greatly enhanced by all the

metals used. Alkaline OPP activity was stimulated more in the presence of Cu++ in all

the isolates except MB490. Likewise, neutral OPP activity was promoted by Cd++ in all

the isolates, while acidic OPP activity in all isolates was enhanced only to a limited

extent by all the metals (figures 7.14). Alkaline phosphatases are known to be metalo-

dependent enzymes requiring Zn and Mg as essential bivalent cations for their activity

and stability (Mori et al., 1999). Alnuaimi et al. (2012) reported that Hg2+, Cu2+, and

Cd2+ inhibited E. coli ALP, whereas Co2+ had little promoter effect on ALP and divalent

alkaline earth metals like Ca2+ and Mg2+ activated the enzyme.

226

a.

b.

c.

d.

Figure 7.14 Effect of metals on activity of Organophosphorus-phosphatase produced

by bacterial isolates (a). MB490 (b). MB497 (c). MB498 (d). MB504. Error bars


01020304050

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227

7.7 Metal bioprecipitation by Organophosphorus Phosphatase (OPP)

The acidic, neutral and alkaline phosphatases were checked for their bioprecipitation

potential for different heavy metals (Ni+2, Cr+6, Mn+2 and Cd+2). For this purpose, the

reaction mixture was prepared by using buffer substrate mixture and extracellular

phosphatase extract (supernatant) supplemented with 1000 ppm of respective metal ion

stock solutions of Ni+2 (NiCl2), Cr+6 (K2Cr2O7), Mn+2 (MnCl2) and Cd+2 (CdCl2) at

different incubation times of 60, 120 and 180 minutes. The free metal ions are

complexed with inorganic phosphate (Pi) produced by catalytic breakdown of pNPP as

metal-phosphate that can be measured by estimating the decrease in the free metal ion

concentration present in the supernatant of reaction mixture with the help of the Atomic

Absorption Spectrometer (AAS Shimadzu AA 7000). At present, progressive

industrialization has resulted in a serious problem environmental metal pollution and

major challenge is to recover desired metals from such waste metal residues. There

must be effective immobilization of metals to prevent contamination of groundwater

(Lieser, 1995). Currently, bioprecipitation of heavy metals as metal phosphates by

enzymatic methods is gaining much interest, as metals can be recovered from very low

concentrations as compared to chemical techniques (Lokhande et al., 2001). There are

many reports of effective bioprecipitation of metals, like uranium and cadmium, with

the application of acid phosphatase extracted from various bacterial

strains(Citrobacter sp., Bacillus sp., Rahnella sp., Pseudomonas sp.

and Salmonella sp.) in an acidic-to-neutral pH range (Macaskie et al., 2000; Renninger

et al., 2004; Shelobolina et al., 2004; Martinez et al., 2007). A radio-resistant

bacterium Deinococcus radiodurans R1 was genetically engineered to secrete a

nonspecific acid phosphatase, PhoN in order to biorecover uranium from acidic or

neutral wastes (Appukuttan et al., 2006). CIAP mediated bioremediation (white

biotechnology) is a novel technique that is eco-friendly and cost effective as compared

to the traditional chemical technologies used for the bioremediation of heavy metal

contaminated soils and industrial effluents (Chaudhuri et al., 2013).

228

7.7.1 Bioprecipitation of Nickel (Ni++) by OPP

7.7.1a Strain MB490

In case of MB490, maximum Ni++ bioprecipitation (97, 96 and 96%) was observed after

60min of incubation by all three types (acidic, neutral and alkaline) of OPP respectively

(figure7.15a).

7.7.1b Strain MB497

For MB497, highest Ni++ bioprecipitation, (98, 98 and 94%) occurred after 60 and 120

min followed by decrease after 180min incubation by acidic, neutral and alkaline OPP

respectively (figure7.15b).

7.7.1c Strain MB498

Maximum Ni++ bioprecipitation, (100%) was achieved after 60min by acidic and neutral

OPP, while it was observed after 120 min by alkaline OPP in case of isolate MB498

(figure7.15c).

7.7.1d Strain MB504

Similarly, in MB504, acidic and neutral OPP exhibited maximum Ni++ bioprecipitation

(99%) after 120 min, while highest 93% Ni++ bioprecipitation was achieved by alkaline

OPP after 180 min of incubation (figure7.15d).

229

a. b.

c. d.

Figure 7.15 Effect of incubation period on Metal bioprecipitation of Ni++ (1000 ppm)

by OPP produced by a). MB490. b). MB497. c). MB498. d). MB504. Error bars


7.7.2 Bioprecipitation of Manganese (Mn++) by OPP

7.7.2a Strain MB490

In case of MB490, acidic OPP achieved maximum Mn++ bioprecipitation (99%) after

120 min, while neutral and alkaline OPP obtained their maximum (100 and 97%) after

180 min of incubation respectively (figure7.16a).

7.7.2b Strain MB497

For MB497, highest Mn++ bioprecipitation by acidic OPP (99%) occurred after 120 min

while neutral and alkaline OPP achieved their maximum value (93%) after 60 min

(figure7.16b).

020406080

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230

7.7.2c Strain MB498

Similar was the case with MB498, where maximum Mn++ bioprecipitation, (99%) was

achieved after 120 min by acidic OPP, while 93% was observed after 60 min by neutral

and alkaline OPP (figure7.16c).

7.7.2d Strain MB504

Similarly, in MB504, acidic OPP exhibited maximum Mn++ bioprecipitation (99%)

after 120 min, while highest 93 and 97% Mn++ bioprecipitation was achieved by neutral

and alkaline OPP respectively after 60 min of incubation (figure7.16d).

a. b.

c. d.

Figure 7.16 Effect of Incubation period on Metal bioprecipitation of Mn++ (1000 ppm)



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231

7.7.3 Bioprecipitation of Chromium (Cr++) by OPP

7.7.3a Strain MB490

In case of acidic OPP from MB490, Cr++ bioprecipitation (86%) remained same after

60, 120, and 180 min, while neutral and alkaline OPP obtained their maximum (86 and

89%) after 60 and 120 min of incubation respectively (figure7.17a).

7.7.3b Strain MB497

In case of MB497, equal but considerable (86%) Cr++ bioprecipitation by acidic OPP

occurred at all given incubation times while maximum value (90 and 87%) was

observed for neutral and alkaline OPP after 60 and 120 min respectively (figure7.17b).

7.7.3c Strain MB498

Similarly, for MB498 maximum Cr++ bioprecipitation, (87 and 86%) was achieved after

180 min by acidic and neutral OPP respectively, while 88% was observed after 120min

by alkaline OPP (figure7.17c).

7.7.3d Strain MB504

In isolate MB504, acidic OPP exhibited maximum Cr++ bioprecipitation (87%) after

180 min, while highest 89 and 88% Cr++ bioprecipitation was achieved by neutral and

alkaline OPP respectively after 120 min of incubation (figure7.17d).

232

a. b.

c. d.

Figure 7.17 Effect of Incubation period on Metal bioprecipitation of Cr++ (1000 ppm)



7.7.4 Bioprecipitation of Cadmium (Cd++) by OPP

7.7.4a Strain MB490

In case of MB490, all three types of OPP (acidic, neutral and alkaline), achieved

maximum Cd++ bioprecipitation (99-100%) after 60min of incubation (figure7.18a).

The current study results are in agreement with those reported by Chaudhuri et al.

(2013) who investigated the removal of heavy metals (Cd 2+, Ni 2+, Co 2+ and Cr 3+/6+)

from single ion solutions by calf intestinal alkaline phosphatase (CIAP) enzyme and

reported highest (80.99%) removal of Cd2+ (initial 250 ppm) followed by Ni 2+

(64.78%) > Cr 3+ (46.15%) > Co 2+ (36.47%) > Cr 6+ (32.33%). They further revealed

greater metal precipitation at pH 11 and at 250 ppm of initial metal concentration as

compared with 1000 ppm.

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7.7.4b Strain MB497

For MB497, highest Cd++ bioprecipitation by acidic, neutral and alkaline OPP (99-

100%) occurred after 60min (figure7.18b).

7.7.4c Strain MB498

Similarly for MB498, maximum Cd++ bioprecipitation, (99-100%) was achieved after

60min by all three types (acidic, neutral and alkaline) of OPP (figure7.18c).

7.7.4d Strain MB504

Likewise, in case of MB504, acidic, neutral and alkaline OPP exhibited maximum Cd++

bioprecipitation (99-100%) after 60 min of incubation (figure7.18d). Nilgiriwala et al.

(2008) studied bioprecipitation of uranium by extracellular alkaline phosphatase,

secreted by recombinant Escherichia coli strain EK4 that was able to precipitate more

than 90% of initial uranium in less than 2 hrs from alkaline solutions.

a. b.

c. d.

Figure 7.18 Effect of Incubation period on Metal bioprecipitation of Cd++ (1000 ppm)



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Incubation Time (min)

234

7.8 Substrate specificity of acidic and alkaline OPP for CPF, TAP and DM

Substrate specificity against different organophosphorus insecticides (Chlorpyrifos,

Dimethoate and Triazophos) was determined by measuring the acidic and alkaline OPP

activity (as neutral OPP production was negligible by the isolates) against 50 mg/l of

respective pesticides after 30 min of incubation at 37°C followed by HPLC and GCMS

analyses (Liang et al., 2005; Gao et al., 2012). Recently, the degradation of pesticides

by using enzymes is being considered as a more effective technology for

bioremediations (Cycon et al., 2013). As phosphatase enzyme hydrolyzes its substrate

and cleaves a phosphoric acid monoester into a phosphate ion and an alcohol by the

addition of water, thus it belongs to hydrolases (Liberti et al., 2012).

Previously, there are many reports of enzymatic degradation of OP pesticides.

Thengodkar and Sivakami (2010) reported 50 and 85% degradation of 10 ppm CPF by

intracellular alkaline phosphatase (extracted from Spirulina sp.) after 1 and 2hrs of

incubation respectively. Similarly, Tang and You (2012) revealed 84.4%, degradation

of TAP by intracellular cell extracts of Bacillus sp. TAP-1 respectively after 1hr of

incubation, and suggested that the Triazophos hydrolase expression was mainly

intracellular and constitutive. Gothwal et al. (2014) extracted and purified

organophosphorus hydrolase (OPH) from Brevundimonas diminuta capable of

degrading methyl parathion up to 500 μM with Vmax of 50 μM/min indicating a

hyperbolic relationship. They further reported the maximum activity of cell free OPH

at optimum pH 7.5 and at 35°C with incubation time of 8 min.

7.7.1 Strain MB490

In case of OPP extracted from MB490, there was less degradation of CPF, TAP and

DM (47, 43 and 40% respectively) by acidic OPP as compared to alkaline OPP i.e 98.8,

94.58 and 90% degradation of CPF, TAP and DM respectively (figure 7.19a).

7.7.2 Strain MB497

Similarly, for MB497, there was less degradation of CPF (70%), TAP (65%) and DM

(60%) by acidic OPP as compared to that degraded by alkaline OPP (94.94, 92.5 and

80% degradation of CPF, TAP and DM respectively). However, acidic OPP seemed

more active in MB497 than in MB490 (figure 7.19b).

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7.7.3 Strain MB498

Organophosphorus phosphatase from MB498 exhibited the same trend as there was

more degradation of CPF (99.32%), TAP (95.5%) and DM (92%) by alkaline OPP than

by acidic OPP i.e 75, 70 and 65% respectively (figure 7.19c). Furthermore, acidic and

alkaline OPP from MB498 showed more active degradation of three pesticide than

MB490 and MB497.

7.7.4 Strain MB504

In case of MB504, there was less degradation of CPF (80%), TAP (75%) and DM (69%)

by acidic OPP than by alkaline OPP (98.12, 96.82 and 94% respectively). Among the

four isolates, acidic OPP extracted from MB504 proved most efficient against CPF,

TAP and DM (figure 7.19d). Alkaline OPP was more efficient in MB504 against CPF

than in MB497 but less than in MB498 and MB490, while it was most active in MB504

against TAP and DM as compared to other three isolates as shown in figure 7.19.

HPLC chromatograms for TAP degradation by alkaline OPP are shown in figure 7.20.

GCMS analyses of the enzymatic reaction mixtures with the three pesticides again

confirmed the formation of same metabolites as observed earlier with OPs treated

bacterial samples (chapters 4, 5 & 6).

Phosphatases belong to the Organophosphorus hydrolases (OPH) and play significant

role in the degradation of OPs alongwith MPH and OpdA. OPH (organophosphorus

hydrolase) is involved in the hydrolysis of organophosphates and has been much

studied (Chaudhry et al., 1988; Mulbry and Karns 1989a; Raushel, 2002; Pinto et al.,

2017). It was reported by Barik et al. (1984) that parathion-hydrolase could hydrolyze

parathion efficiently and was found to be constitutive in nature. Tang and You (2012)

revealed the triazophos-hydrolase activity of crude cell free extracts of strain TAP-1,

thus indicating the enzymatic degradation of triazophos. They further demonstrated that

enzyme involved are intracellular and constitutive.

The main bacterial pathways for degradation of OP pesticides involved the cleavage of

phosphorus ester bond (P-O), which ultimately results in the release of inorganic

phosphorus that is utilized by bacteria as a sole source of phosphorus (Horne.et al.,

2002). Generally phosphotriestrases (PTE) are the enzymes catalyzing breakdown of

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phospho-ester bond of OP pesticides (Singh and Walker, 2006).Phosphatases like OPP

are related to phosphotriestrases (PTE). There are many reports of PTE isolated from

various bacterial strains capable of catalyzing different OPs (Mulbry et al., 1986;

Chaudhry et al., 1988; Horne et al., 2002). Khalid et al. (2016) reported 44 and 78%

degradation of 100 ppm CPF by 25 and 50 µl of ALP respectively after 1hr of

incubation. The bacteria are not adversely affected by OPs as they lack the

acetylcholine esterase which is usually inhibited by OPs in higher organisms like

insects and mammals (Singh et al., 2009).

Zhang et al. (2014) reported enhanced degradation of five OP pesticides in skimmed

milk by use of ten lactic acid bacteria. Moreover, phosphatase production by the strains

was positively correlated to increased degradation of given pesticides thus playing

possible vital role during OP pesticide degradation.

Majumder and Das (2016) studied the effect of four OP pesticides (monocrotophos,

profenophos, quinalphos and triazophos) in relation to the growth and activities of

phosphate solubilizing microorganisms and availability of insoluble phosphates in soil.

The growth of phosphate solubilizing microorganisms was reported to be greatly

encouraged by 38.3% profenophos, whereas solubility of insoluble phosphates in soil

was highly enhanced by 20.8% monocrotophos. On the other hand, the acid and alkaline

phosphatase activities of the soil were expressively improved by the integration of the

pesticides in general, manifesting more availability of water soluble phosphorus in soil.

237

a. b.

c. d.

Figure 7.19 Degradation of OP pesticides by acidic and alkaline OPP as analyzed by

HPLC after 30min of incubation. a). MB490. b). MB497, c). MB498. d). MB504. Error


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a. b.

c. d.

e.

Figure 7.20 HPLC chromatograms showing degradation of 50 mg/l of TAP (RT = 2.4

min) by alkaline OPP as analyzed by HPLC after 30 min of incubation. a). Control b).

MB490. c).MB497. d). MB498. e). MB504.

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Conclusion

The four bacterial strains MB490 (Pseudomonas kilonensis), MB497 (Bacillus


proved to be phosphate solubilizing bacteria with the production of extracellular acidic,

neutral and alkaline phosphatases. The three types of OPP were further found active

and stable over a wide range of pH, temperature, substrate (p-NPP) concentrations,

metal ions and different chemicals like SDS and EDTA. The acidic, neutral and alkaline

OPP further exhibited great potential for bioprecipitation of different metals like Ni,

Mn, Cd and Cr. Finally, acidic and alkaline OPP showed a broad substrate specificity

against three OP pesticides (CPF, TAP and DM) with significant degradation and

metabolites formation as analyzed by HPLC and GCMS. Therefore, the phosphate

solubilizing potential and phosphatase production activity of the bacteria in the current

study, strongly recommend these four strains for their application in the bioremediation

of soil and water resources contaminated with OP pesticides and heavy metals. These

strains can even be used as biofertilizers due to their phosphate solubilization ability.

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Chapter 8

Discussion

Organophosphate pesticides are playing vital role in the control of domestic and

agricultural pests and insects, but their over use has adversely affected our environment.

Recently, the OP pesticides have been comprehensively studied with respect to their

chemistry, mechanism of action and classification (Carr et al., 2001; Caceres et al.,

2007; Raposo et al., 2010). There may be both acute and chronic effects when humans

are exposed to OP pesticides. The acute effects are definitely fatal due to direct

poisoning, while the long-term effects may harm the peripheral and central nervous

system in mammals due to inhibition of acetylcholinesterase. According to Costa

(2006), this phenomenon is medically termed as organophosphate-induced delayed

neuropathy (OPIDN). Furthermore, it is a known fact that OP pesticides may be

potential teratogens and carcinogens for human (Chambers et al., 2001; De Silva et al.,

2006).

Recently biotechnological approaches are being used to degrade recalcitrant pesticide

residues present in our environment including soil and water resources. These

bioremediation techniques depend upon both natural and cultivated organisms like

bacteria, fungi, cyanobacteria and even higher plants for the transformation of

pollutants and are considered more effective, economical and environment friendly as

compared to conventional physical and chemical methods (Singh and Walker, 2006).

There are variety of fungal and bacterial microbes found capable of degrading OP

compounds e.g among bacteria Enterobacter B-14, Alcaligens faecalis DSP3,

Stenotrophomonas sp.YC-1, Sphingomonas Dsp-2, Paracoccus sp. TRP, Bacillus

pumilus C2A1, Serratia sp. (Singh et al., 2004; Yang et al., 2005; Yang et al., 2006;

Xu et al., 2008; Anwar et al., 2009; Li et al., 2010; Li et al., 2013). Similarly fungal

OP degraders include Verticillium sp. DSP, Acremonium sp. GFRC-1 and

Cladosporium cladosporioides Hu-0 (Fang et al., 2008; Kulshrestha and Kumari, 2011;

Chen et al., 2012).

The microbes, especially the common inhabitant of polluted sites are bestowed upon

natural mechanisms like enzymes to consume and transform toxic chemicals including

pesticides in to non- toxic or less toxic forms. Bioremediation using bacteria has been

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more focused by researchers mainly due to their easy handling, rapid growth and

minimum nutritional requirements. Therefore, in the present study, OP pesticide

degrading bacteria were isolated from agricultural fields of Mianwali with a previous

history of repeated application of OP pesticides. The main objective was to check the

potential of these indigenous bacteria to degrade and transform three OP pesticides

(CPF, TAP and DM) and to evaluate their ability to be utilized for the bioremediation

of Pakistani agricultural soils which have been contaminated with OP pesticides.

Plate assay for pesticide tolerance of four isolates Pseudomonas kilonensis MB490,


MB504 revealed that MB497 was most tolerant for CPF up to 8 g/l, followed by MB490

and MB498 up to 6 g/l and then by MB504 up to 0.8 g/l of CPF. In case of TAP, MB490,

MB497 and MB498 showed good growth up to 4 g/l, whereas MB504 was least tolerant

up to 2 g/l of TAP. Dimethoate proved more toxic for isolates as MB490, MB497 and

MB498 showed growth only up to 0.4 g/l of Dimethoate, while MB504 was least

tolerant (up to 0.22 g/l) for DM. For three pesticides, these isolates showed tolerance

as CPF>TAP>DM and MB504 proved least tolerant among four isolates. For CPF,

tolerance of isolates followed the order MB497>MB490= MB498>MB504, while for

TAP and DM toleratance, the order was MB490=MB497=MB498>MB504. Previously

bacterial tolerance to different concentrations of various pesticides is well documented.

Azotobacter chroococcum was capable of tolerating carbofuran up to 5 ppm (Kale et

al., 1989). It was reported by Harishankar et al. (2013) that L. fermentum, L. lactis and

E. coli could tolerate CPF up to 1400 µg/ml while E. faecalis and L. plantarum showed

tolerance up to 400 and 100 µg/ml respectively. Similarly Shafiani and Malik (2003)

reveled that a Pseudomonas sp. was able to resist endosulfan, carbofuran and malathion

up to 800, 1600 and 1600 µg/ml respectively.

The four selected strains MB490 (Pseudomonas kilonensis), MB497 (Bacillus

thuringiensis), MB498 (Pseudomonas kilonensis) and MB504 (Pseudomonas sp.) out

of a number of isolates showed their growth as well as pesticides (CPF, TAP and DM)

degradation under a wide range of temperatures (25◦C-37◦C) and pH (6-9) which are

mostly prevalent under local environment of Pakistan. The heavy metals and organic

pollutants other than pesticides are also being continuously accumulated in the soil

sediments and water bodies as a result of different human activities like industrial

242

effluents or biochemical spills. The selected isolates were able to tolerate significant

concentration of different heavy metals like Ni, Cr, Cu, Zn, Pb and Mn (Table 3.11)

and also organic compounds such as Benzene, Toluene, Xylene, Aniline, Biphenyl and

Naphthalene (Table 3.12).

Wang et al. (2005) reported that isolate Klebsiella sp. E6 exhibited good growth at pH

7–8 under a broader temperature range of 32 to 37◦C. Furthermore, it was capable of

utilizing methanol and TAP as carbon source and nitrogen source respectively. In case

of CPF, there was 83.74 to 99.48% degradation at all given temperatures (25, 30 and

37◦C) and 67.22 to 99.39% at all given pH (6, 7, 8) by all the four isolates. Maximum

CPF degradation was observed at 30 and 37◦C and at pH 8 in most isolates, which

corresponds to prevailing average temperature and soil pH in Pakistan. The soils in

Pakistan are considered mostly dry and alkaline (with pH almost 8 or above) due to the

presence of high quantities of calcium carbonate and a lower amount of organic matter

(Khalid et al., 2012). It is suggested that pH of experimental biological systems must

be maintained within range of 6.5-8.0 (Tyagi, 1991; Tano-Debrah et al., 1999).

Similarly, Singh et al. (2003) reported more rapid Chlorpyrifos biodegradation in the

two alkaline soils having pH 7.7 and 8.4 as compared to acidic and neutral pH. The

degradation of fenitrothion which is a nitrophenolic pesticide, by Burkholderia sp.

FDS-1, was reported to be more rapid at 30 °C and at slightly alkaline pH (Hong et al.,

2007).

Present study results are in agreement with Anwar et al. (2009) who reported more

efficient (80%) CPF degradation by Bacillus pumilus isolated from Pakistani

agricultural soils at alkaline and neutral pH as compared to lesser (only 50%)

degradation at acidic pH. On the other hand, there was higher CPF degradation at acidic

pH by a fungus, Cladosporium cladosporioides Hu-01 as revealed by Chen et al.

(2012).

During modern age, treatment of soils and sediments in slurry bioreactors (SBs) under

controlled environmental conditions is being considered as one of the most significant

ways of in situ and ex situ bioremediation techniques (Mueller et al., 1991; Mohan et

al., 2006). The main purpose to use SBs is to determine the practicability and real

potential of a certain biological approach in the final refurbishment of a polluted soil or

243

place (Fava et al., 2004). Indeed, the rates of removal of a pollutant with in slurry

environment are determined chiefly by the degradation potential of the microorganisms

existing in the system (Cookson, 1995).

Soil microcosms are like the miniaturized ecosystems that make it possible for the

researchers to probe the effects of presence of selective recalcitrant on natural microbial

populations under precise conditions (Grenni et al., 2012). The laboratory microcosms

are considered as model ecosystem having a part of the natural environment like soil or

water (Benton et al., 2007; Drake and Kramer, 2012). These models possess natural

biotic communities which are retained, under controlled environmental conditions like

temperature, light, humidity etc. equivalent to natural ones.

When the four isolates MB490 (Pseudomonas kilonensis), MB497 (Bacillus


compared for their potential to degrade CPF, TAP and DM in M-9 broth, soil slurry and

soil microcosm with respect to time period, it was noticed that all of these exhibited

outstanding consumption of CPF, TAP and DM within 9 days of incubation (figure

8.1a, b, c). In M-9 broth, all isolates consumed three given pesticides in the order

CPF>TAP>DM with CPF degraded up to almost 100%, TAP 88.4-95.8% and DM 80.1-

92% within 9 days. In soil slurry, all the strains exhibited CPF and TAP degradation up

to 99% or above, while these could degrade DM from 80 to 91.8%. When grown in

soil microcosm, these isolates could degrade more CPF (99 to 99.8%) followed by TAP

(92.72 to 95%) and then DM (78.8 to 82.1%), which corresponds well with their

tolerance behavior against CPF, TAP and DM. So, it can be noticed that CPF proved

more easy and favorite substrate for the isolates in all the three given media (M-9 broth,

soil slurry and soil microcosm), followed by TAP and then DM. When degradation

performance of individual bacterial strains was compared in the given media for each

of three pesticides, it was observed that all four strains showed >99% degradation of

CPF in all media (figure 8.1). Present study results are in strong agreement with those

revealed by Cho et al. (2009) who reported 83.3% degradation of CPF (200 mg/l)

within 3 days and 100% degradation after 9 days by four strains of lactic acid bacteria

(Leuconostoc mesenteroides WCP907, Lactobacillus brevis WCP902, Lactobacillus

plantarum WCP931, and Lactobacillus sakei WCP904). These strains were also

capable of degrading Coumaphos, diazinon, parathion, and methylparathion. Abraham

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and Silambarasan (2013) studied CPF degradation in soil microcosm (100 g of soil)

with and without supplementing nutrients by Sphingobacterium sp. JAS3 and reported

complete degradation of CPF within 24 hrs in both soils along with complete

degradation of TCP (product of CPF degradation) achieved in 5 days of incubation.

During present study, four individual strains MB490 (Pseudomonas kilonensis),

MB497 (Bacillus thuringiensis), MB498 (Pseudomonas kilonensis) and MB504

(Pseudomonas sp.) showed > 99% degradation of initial 200 mg/l of CPF in sterilized

soil within just 9 days of incubation which can be compared with that reported by Akbar

and Sultan (2016) who stated that Achromobacter xylosoxidans (JCp4) and

Ochrobactrum sp. (FCp1) could degrade 200 mg/kg of initial CPF in both sterilized

and non-sterilized soils up to 93–100% but within 42 days. Stenotrophomonas sp. YC-

1 was reported to degrade 100 mg/l of CPF up to 100% within 24 hrs (Yang et al.,

2006), while 93.8% degradation of initial supplemented 5 mg/l CPF was demonstrated

by Synechocystis sp. PUPCCC 64 within 5 days (Singh et al., 2011). Similarly, it was

revealed by Singh et al. (2004) that Enterobacter strain B-14 could degrade CPF (25

mg/l) up to 40% in 48 hrs of treatment.

For TAP degradation in M-9 broth, the isolates followed the order

MB490>MB497>MB498>MB504, while in soil slurry these were degrading TAP as

MB490=MB504>MB498>MB497 and in soil microcosm these isolates consumed TAP

as MB504>MB497>MB490>MB498 but with very minor difference (figure 8.1). There

are only few earlier reports about TAP degradation by bacteria. Wang et al. (2005)

reported bacterial degradation of TAP for the first time by Klebsiella sp. E6 which was

able to consume 100 mg/l of TAP as well as its metabolite 1-phenyl-3-hydroxy-1, 2, 4-

triazole as a carbon and nitrogen source within 7 days. Similarly, Bacillus sp., TAP-1

was reported to degrade 100 mg/l of TAP up to 98.5% in the minimal liquid medium

after 5 days of incubation (Tang and You, 2012). Yang et al. (2011) reported the

complete removal of 50 mg/l of TAP and its metabolite 1-phenyl-3-hydroxy-1, 2, 4-

triazole (PHT) by Diaphorobacter sp. TPD-1in mineral salt broth within 24 and 56 hrs

respectively. Stenotrophomonas sp. PF32 and Stenotrophomonas sp. G1 could degrade

100 mg/l and 50 mg/l of triazophos up to 97 and 34% respectively within 42 and 24 hrs

respectively (Xu et al., 2009; Deng et al., 2015).

245

In case of DM degradation in M-9 broth, the bacterial strains exhibited their potential

as MB490>MB498>MB497>MB504, while in soil slurry these followed the sequence

as MB498>MB490>MB497>MB504, whereas in soil microcosm their DM degradation

capacities were in the order MB490>MB497>MB498>MB504 (figure 8.1). So, it can

be noticed that strain MB490 remained prominent in most cases consuming maximum

pesticide as compared to other three isolates.

a. b.

c.

Figure 8.1 Comparison of degradation of CPF, TAP and DM by MB490 (Pseudomonas

kilonensis), MB497 (Bacillus thuringiensis), MB498 (Pseudomonas kilonensis), and

MB504 (Pseudomonas sp.) after 9 days of incubation. a). M-9 broth b). Soil slurry c).

Soil microcosm. The error bars represent standard errors for values of three sample

replicates.

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One of the unique feature of the selected four bacterial isolates MB490 (Pseudomonas

kilonensis), MB497 (Bacillus thuringiensis), MB498 (Pseudomonas kilonensis) and

MB504 (Pseudomonas sp.) was their ability to consume and degrade primary

metabolite of CPF i.e 3, 5, 6 Trichloropyridinol (TCP), which previously had been

considered recalcitrant due to its antimicrobial effect (Armbrust, 2001; Caceres et al.,

2007). At lower concentration (14 mg/l) of TCP, the four bacterial isolates showed more

degradation (93.80 to 98%) of TCP as compared to higher (28 mg/l) concentration of

TCP i.e 60.39 to 90.57% within 72 hrs of incubation (Figure 8.2), which was observed

to be positively correlated with bacterial growth during this period. At 14 mg/l, TCP

was degraded by isolates in the sequence MB490>MB504>MB497>MB498, while at

28 mg/l, TCP degradation followed the order: MB497>MB504>MB490>MB498. On

the other hand, there was only negligible (2-5%) TCP degradation in control without

inoculum.

Figure 8.2 Comparison of degradation of 3, 5, 6 Trichloropyridinol (TCP) at two

different concentrations (14 mg/l and 28 mg/l) by bacterial isolates MB490

(Pseudomonas kilonensis), MB497 (Bacillus thuringiensis), MB498 (Pseudomonas

kilonensis) and MB504 (Pseudomonas sp.). The error bars represent standard errors for


In the past, only a small number of microorganisms have been reported which were

capable of TCP degradation along with its parent compound CPF e.g Alcaligenes

Faecalis, Bacillus pumilus C2A1, Cladosporium cladosporioides Hu-01, Cupriavidus

sp. DT-1, Xanthomonas sp. 4R3-M1 and Pseudomonas sp. 4H1-M3 (Yang et al., 2005;

Anwar et al., 2009; Chen et al., 2012; Lu et al., 2013; Rayu et al., 2017). Current study

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results of TCP degradation are comparable to those reported by Chen et al. (2012),

where 89% CPF and 93.5% of TCP (50 mg/l each) were decomposed by fungal strain

Cladosporium cladosporioides Hu-01 within 24 hrs followed by complete removal

after 5th and 6th day of incubation respectively. Abraham and Silambarasan (2013)

reported that 300 mg/l of CPF was completely degraded in mineral broth within 12 hrs

of incubation and its metabolite TCP (so produced) was also disappeared completely

by Sphingobacterium sp. JAS3 within 5 days of treatment, while transformation

product benzene, 1,3-bis(1,1-dimethylethyl) was detected by GCMS. Alcaligenes

faecalis DSP3 was able to degrade CPF and TCP (100 mg/l each) up to 100 and 93.5%

within 12 days (Yang et al., 2005). Paracoccus sp. TRP completely degraded 50 mg/l

of CPF and TCP in 4 days (Xu et al., 2008). Bacillus pumilus C2A1 was reported to

consume 1000 mg/l of CPF and 300 mg/l of TCP up to 89 and 90% within 15 days and

8 days respectively (Anwar et al., 2009). Similarly Mesorhizobium sp. HN3 could

completely utilize initially supplemented 50 mg/l of CPF and TCP (Jabeen et al., 2015).

Recently, Rayu et al. (2017) demonstrated that Xanthomonas sp. 4R3-M1 and

Pseudomonas sp were capable of completely utilizing 20 mg/l CPF and no TCP

detected as metabolite after 6 days of incubation, indicating its complete consumption.

The same strains were reported to degrade TCP when supplied externally

In the present study, three of the four selected bacterial isolates i.e MB490

(Pseudomonas kilonensis), MB498 (Pseudomonas kilonensis) and MB504

(Pseudomonas sp.) belong to the genus Pseudomonas as identified based on 16s rRNA

analysis and all of them proved very effective in degrading CPF, TCP, TAP and DM in

the given media. Gram negative rods belonging to Pseudomonads (like Pseudomonas

putida and P. fluorescens), are known to have the best degradative capability (Fritsche

and Hofrichter, 2005), mainly due to their potential to utilize metabolic protocols along

with catabolic enzymes (Houghton and Shanley, 1994). In many previous

investigations, Pseudomonas sp. have been proved capable of degrading OP-

compounds either co-metabolically or catabolically such as Pseudomonas diminuta-

MG (Serdar et al., 1982; Mulbry et al., 1986), Pseudomonas sp. (Choi et al., 2009),

Pseudomonas putida POXN01 (Iyer et al., 2013). Almost 50% CPF consumption of

initial 150 mg/l of CPF by pure strain of Pseudomonas kilonensis SRK1 was

demonstrated by Khalid et al. (2016a) at optimal conditions of pH 8, CFU (306 × 106),

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and in the presence of glucose as extra carbon source. In the current study, two strains

of Pseudomonas kilonensis MB490, Pseudomonas kilonensis MB498 were able to

remove 97 and 99.36% of initial 200 mg/l of CPF respectively at optimum temperature

of 37°C, CFU/ml of 106 and at pH 7 under static conditions in M-9 broth without any

additional carbon source in 24 hrs which indicates their comparatively greater potential

to degrade CPF without requiring any additional nutrients like glucose for the

expression of OP degrading enzymes in contradiction to Pseudomonas kilonensis SRK1

as reported by Khalid et al. (2016a).

The complete CPF degradation was reported in a culture consisted of combination of

the cell surface-expressed laccases and living bacterial cells of Pseudomonas putida

MB285 (Liu et al., 2016). On the other hand, CPF was partially degraded into TCP by

the purified cell free laccase. Recently, Pailan and Saha (2015) has isolated an OP

degrading chemotactic Pseudomonas BUR11 that is capable of utilizing parathion, CPF

as well as their major hydrolytic metabolites for its growth. It could degrade 200 ppm

of parathion up to 62% in 96 hrs at 37 ◦C.

In the current study, MB497 also proved to be efficient degrader of CPF, TAP and DM

in M-9 broth, soil slurry and soil microcosm within 9 days of incubation (figure 8.1). It

was identified as Bacillus thuringiensis by 16s rRNA analysis. There are many earlier

reports of Bacillus sp. as a degrader of OP pesticides. Bacillus thuringiensis was

isolated from agricultural wastewater in Egypt and was reported as capable of

degrading malathion (Zeinat et al., 2008). Similarly Bacillus pumilus C2A1, isolated

from agricultural soil at Faisalabad, Pakistan, revealed to be very effective as degrader

of CPF (Anwar et al., 2009). Recently a bacterial strain Bacillus subtilis Y242 isolated

from agricultural wastewater in Egypt was found highly effective in degrading 150 mg/l

Chlorpyrifos up to 95.12% in 48 hrs of incubation (El-Helow et al., 2013) which is

comparable to CPF degradation observed in present study.

During current investigations, bacterial inoculum of 106 CFU/g/ml was used. The

inoculum size has been considered as an important factor to achieve a successful

bioremediation of pesticide contaminated sites (Ramadan et al., 1990). It was

recommended to use inoculum size of 106–108 cells/g of soil for the bioremediation of

pesticide-contaminated places (Comeau, et al., 1993).

249

In previous findings, microbial consortia have been proven an efficient tool for

pesticide degradation because of the fact that a solitary bacterium might not have all

the enzymes required to remove whole or even majority of the organic compounds in a

polluted site. The consortia are composed of mixture of microbial communities so that

different genes and their enzymes having the maximum potential for biodegradation

can be used to degrade the composite mixtures of recalcitrant organic compounds often

present in contaminated locations (Fritsche and Hofrichter, 2005). Abraham et al.

(2014) reported the complete biodegradation of a mixture of two OP pesticides

(Chlorpyrifos, monocrotophos) and an organochlorine pesticide (endosulfan) by a

consortium consisting of 10 bacterial species in a fermenter–bioreactor system within

24 hrs of treatment. It was revealed by Ortiz-Hernandez and Sanchez-Salinas (2010)

that a bacterial consortium consisting of six bacterial strains was able to degrade

tetrachlorvinphos (TCV) up to 57% in 36 hrs. While only one individual strain A3 could

remove 49% TCV in mineral medium, other strains could utilize it only in rich medium.

Degradation potential of individual strains (MB490, MB497, MB498 and MB504) for

CPF, TAP and DM was compared with their respective consortia i.e A

(MB490+MB498), B (MB490+MB497), C (MB490+MB504), D (MB498+MB497), E

(MB498+MB504), F (MB497+MB504), and G (MB490+MB497+MB498+MB504) in

M-9 broth, soil slurry and soil microcosm after 9 days of incubation. It was observed

that all the consortia and individual pure strains were equally effective in degrading

CPF (up to 99% or more), in all given media after 9 days of incubation. Barathidasan

et al. (2014) used a consortium consisting of a bacterium Cellulomonas fimi (capable

of transforming CPF to TCP) and a fungal strain Phanerochaete chrysosporium that

could consume both CPF and TCP. The consortium was able to completely mineralize

50 mg/l of CPF within 16hrs which was otherwise possible in 6 days by the fungus

alone. Present investigation results are much better than those recently reported by

Akbar and Sultan (2016) that two bacterial isolates Achromobacter xylosoxidans JCp4

and Ochrobactrum sp. FCp1 could degrade up to 84.4% and 78.6% respectively of

initially supplemented 100 mg/l of CPF in a treatment period of 10 days. Recently a

consortium consisting of five strains (Pseudomonas kilonensis SRK1, Serratia

marcescens SRK2, Bacillus pumilus SRK4, Achromobacter xylosoxidans SRK5 and

Klebsiella sp. T13) was reported to consume 400 mg/l of CPF up to 98 % within

250

sequencing batch reactors at pH 7, 10 % inoculum and in 48 hrs of reaction time (Khalid

et al., 2016a).

Triazophos is one of the major OP pesticides being used extensively in Pakistan to

control pests in agriculture. It is the first time that TAP biodegradation by indigenous

bacterial isolates has been investigated in Pakistan. In case of TAP degradation in M-9

broth, it was noticed that pure strains MB490, MB497, MB498 and MB504 showed

less TAP degradation (88.4-95.8%) than their respective consortia (>99%) as shown in

figure 8.3. In soil slurry, strains MB490 MB497, MB498 and MB504 along with

consortium G exhibited more TAP degradation (almost 99%) than other consortia

(92.69 to 95.93%) as given in figure 8.4a, b, c & d). In soil microcosm, all strain

performed less (92.72-95%) TAP degradation than their consortia A, D, E, F and G

(97.17-99.9%). While consortia B and C showed reduced TAP degradation than

respective individual strains MB490, MB497 and MB504 in soil microcosm (figure

8.5d).

251

a. b.

c. d.

Figure 8.3 Comparison of degradation of TAP by pure strains (MB490, MB497,

MB498 and MB504) with their respective consortia A, B, C, D, E, F and G in M-9

broth after 9 days. The error bars represent standard errors for values of three sample

replicates.

20

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100

alone A B C G

% D

egra

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MB490

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MB497

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MB498

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MB504

252

a. b.

c. d.


MB498 and MB504) with their respective consortia A, B, C, D, E, F and G in soil slurry

after 9 days. The error bars represent standard errors for values of three sample

replicates.

Dimethoate (DM) is also an important OP pesticide having an insecticidal effect to kill

a large variety of insects like aphids, thrips, plant hoppers and whiteflies both

systemically as well as by contact (Hayes and Laws, 1990). Again, in Pakistan, there is

no previously reported research on DM biodegradation by bacteria.

0

20

40

60

80

100

alone A B C G

% D

egra

dat

ion

of

TAP

MB490

0

20

40

60

80

100

alone B D F G

% D

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of

TAP

MB497

0

20

40

60

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alone A D E G

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MB498

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% D

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MB504

253

a. b.

c. d.


MB498 and MB504) with their respective consortia A, B, C, D, E, F and G in soil

microcosm after 9 days. The error bars represent standard errors for values of three

sample replicates.

So, the present investigation is novel in the sense that DM biodegradation by indigenous

bacterial isolates has been focused for the first time in Pakistan along with identification

of its metabolites suggesting a novel and unique pathway for its transformation in

bacteria. For DM degradation in M-9 broth after 9 days, more (95-99.9%) DM

degradation was observed by consortia as compared to pure strains (80.1-92%) (figure

8.6), probably due to more availability and solubility of pesticide in liquid media as

well as synergistic effect of all consortial members to degrade DM. In case of soil

0

20

40

60

80

100

alone A B C G

% D

egra

dat

ion

of

TAP

MB490

0

20

40

60

80

100

alone B D F G

% D

egra

dat

ion

of

TAP

MB497

0

20

40

60

80

100

alone A D E G

% D

egra

dat

ion

of

TAP

MB498

0

20

40

60

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MB504

254

slurry, it was noticed that MB490 showed more degradation than consortia B, C and G

but little bit less than consortium A after 9 days (figure 8.7a), while isolate MB497

showed higher (88.24%) DM degradation than B, D and F but less than G (90%) as

illustrated in figure 8.7b. Strain MB498 could degrade more DM than its respective

consortia A, D and G but less than E after 9 days in soil slurry (figure 8.7c), while

MB504 degraded more DM than E but less than C, F and G (figure 8.7d). In soil

microcosm, DM degradation by MB490 was more than consortia B and C but less than

A and G after 9 days of incubation (figure 8.8a), while MB497 showed higher

degradation than B, D and E but less than G (figure 8.8b). Strain MB498 performed

better as compared to D but less than A, E and G (figure 8.8c), whereas MB504

degraded lesser DM than C, E, F and G (figure 8.8d). So, consortia E and G (almost

90%) degraded highest DM in soil microcosm among all consortia. There are previous

reports of DM degradation by Pseudomonas and Bacillus sp. Similar to present results,

Debmandal et al. (2011) reported degradation of 100 mg/l of DM in mineral salt

solution by Bacillus licheniformis, Pseudomonas aeruginosa, Aeromonas hydrophila,

Proteus mirabilis and Bacillus pumilus up to 100, 96, 83, 72 and 71% respectively

within 7 days. It was previously reported by Deshpande et al. (2001) that Pseudomonas

aeruginosa, Bacillus megaterium degraded Dimethoate up to >95% within 8 days of

incubation. Liang et al. (2009) demonstrated 75% removal of Dimethoate by Raoultella

sp. via co-metabolism.

255

a. b.

c. d.

Figure 8.6 Comparison of degradation of DM by pure strains (MB490, MB497, MB498

and MB504) with their respective consortia A, B, C, D, E, F and G in M-9 broth after

9 days. The error bars represent standard errors for values of three sample replicates.

0

20

40

60

80

100

alone A B C G

% D

M d

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MB490

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% D

M d

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MB497

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256

a. b.

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and MB504) with their respective consortia A, B, C, D, E, F and G in soil slurry after

9 days. The error bars represent standard errors for values of three sample replicates.

0

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a. b.

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and MB504) with their respective consortia A, B, C, D, E, F and G in soil microcosm

after 9 days. The error bars represent standard errors for values of three sample

replicates.

The biodegradation of mixture of CPF and TAP was also investigated by the four pure

isolates and their consortium G. It was observed that TAP and CPF both had an

enhancing effect on CPF and TAP degradation respectively as well as on bacterial

growth which may be due to the availability of more nutrients formed as result of

mineralization of both pesticides resulting in the production of more degrading

0

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alone A B C G

% D

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MB490

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% D

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MB497

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258

enzymes. It was reported by Deng et al. (2015) that a bacterial strain Stenotrophomonas

G1 had the ability to degrade 50 mg/l of multiple OP pesticides i.e methyl parathion,

methyl paraoxon, diazinon, and phoxim (100% each), parathion (95%), CPF (63%),

profenofos (38%) and triazophos (34%) within 24 hrs. This report is comparable to

current learning, where four bacterial strains MB490 (Pseudomonas kilonensis),

MB497 (Bacillus thuringiensis), MB498 (Pseudomonas kilonensis) and MB504

(Pseudomonas sp.) were capable of effectively degrading three given OP pesticides

(CPF, TAP and DM) separately as well as these strains degraded CPF and TAP in a

mixture very efficiently.

In current investigation, there was only 11.7 and 45.5% CPF degradation, 10 and 28%

TAP degradation, 12 and 20% DM degradation in non-inoculated sterile and non-

sterilized soil after 9 days of incubation, thus suggesting a strong role of bacterial

isolates MB490, MB497, MB498 and MB504 in CPF, TAP and DM biodegradation in

soil. So, the data obtained in the current investigation about the existence of

autochthonous bacterial communities bestowed with a natural remediation capability,

which can be utilized to develop site-specific tactics to achieve natural reduction of

pollution in soil and water resources as well as a potential application of selective

identified bacterial strains for the purpose of bioremediation.

GCMS analysis of the samples treated with bacterial isolates MB490, MB497, MB498

and MB504 and their consortia separately in all given media (M-9 broth, soil slurry and

soil microcosm) treated with CPF indicated the formation of 2-Hydroxy-3, 5, 6-

trichloropyridine (TCP), Diethyl thiophosphate (DETP) and phosphorothioic acid

in all bacterial samples. While two metabolites were novel and detected first time. [(3,

5, 6-trichloro-2-pyridinyl) oxy] acetic acid was noted specifically in samples with

MB490 and Diisopropyl methanephosphonate was detected in samples of MB497,

MB498 and MB504. TCP, DETP and phosphorothioic acid have been stated in many

previous reports (Reddy et al., 2012; Jabeen et al., 2015; Khalid et al., 2016; Rayu et

al., 2017).

It is important to be noted here that 3, 5, 6 Trichloropyridinol production during CPF

biodegradation was also verified by HPLC analysis. During GCMS analysis, retention

259

time (RT) for CPF and TCP standard was determined to be 22 and12.8 min respectively,

while by HPLC analysis it was determined to be 5.4 and 1.77 min respectively.

Another important feature of the present study is that the bacterial isolates (MB490,

MB497, MB498 and MB504) were capable of metabolizing and mineralizing TCP

(main metabolite of CPF) added externally in M-9 broth as analyzed by GCMS. All the

four isolates formed four metabolites (1-methyl-2-pyrrolidine ethanol, p-Propyl

phenol, 2-Ethoxy-4, 4, 5, 5-tetramethyloxazoline and 3-(2, 4, 5-Trichlorophenoxy)-

1-propyne), based on NIST library identification after 24, 48 and 72 hrs of incubation

as shown in in Table 4.4. Metabolic pathways for TCP and CPF and have been proposed

in the current study and represented in the figures 4.47 & 4.48 respectively.

During GCMS and HPLC analysis, retention time (RT) for TAP standard was

determined to be 27.8 min and 2.4 min respectively. The GCMS analysis of TAP

supplemented bacterial samples after 3, 6 and 9 days of incubation in given media

indicated the following 7 metabolites of TAP, which are very novel and have not been

reported elsewhere before current study:

1. 1, 2, 4-Triazole-4—amine, N-(2-Thienylmethyl).

2. 4-Benzyl-4, 5-dihydroisoxazole.

3. Benzene sulfonic acid, methyl ester.

4. 4H-1, 2, 4-Triazole-4-benzenesulfonamide.

5. 4, 5 dihydro-N-(O-toyl)-3-furamide.

6. Ethyl 4-phenyldiazenylbenzoate.

7. Dibutyl methanephosphonate.

So all the four bacterial strains MB490 (Pseudomonas kilonensis), MB497 (Bacillus

thuringiensis), MB498 (Pseudomonas kilonensis) and MB504 (Pseudomonas sp.) in

the current study have been proved to be very promising towards degradation and

metabolization of TAP and can be considered as suitable for practical remediation of

TAP contaminated agricultural soils and other resources. Ishag et al. (2016) have

demonstrated that initial 400 mg/l of CPF, malathion and DM were removed up to >87,

>80 and 76% respectively from the mineral media by consortium of three bacterial

260

strains (Bacillus safensis FO-36bT, Bacillus subtilis KCTC13429T, and Bacillus cereus

ATCC14579T).

There were 6 metabolites of Dimethoate identified in the GCMS chromatograms for

individual bacteria and their consortia in the given media (M-9, soil slurry and soil

microcosm) at 37°C and pH 7.4 ± 0.2 with the help of NIST library as given below in

Table 8.1:

Table 8.1 Main compounds identified in the GCMS chromatogram of Dimethoate

degraded by bacterial isolates (MB490, MB497, MB498 and MB504) and their

consortia.

S.no Identified compound Retention time

(minutes)

1 Dimethoate 17.7

2 Methyl diethanol amine 15.9

3 Phosphonothioic acid, propyl-O, S-dimethyl ester 4.8

4 O, O, O- Trimethyl thiophosphate 9.56

5 Omethoate 9.78

6 Aspartylglycine ethyl ester 14.46

Li et al. (2010) proposed the transformation pathway of DM by Paracoccus sp. Lgjj-3.

Among many other metabolites formed, they also reported formation of Methyl amine

by amidase cleavage from Dimethoate just like Methyl diethanol amine in present

study. This reaction is in agreement to those previously reported in mammals and

during photocatalytic oxidation (Evgenidou et al., 2006; Franca and Emanuela, 2007).

Metabolic pathway for DM transformation by bacterial isolates in the current study has

been illustrated in figure 6.26 in chapter 6.

Another important aspect of the present study is that all the four selected bacterial

strains MB490 (Pseudomonas kilonensis), MB497 (Bacillus thuringiensis), MB498

(Pseudomonas kilonensis) and MB504 (Pseudomonas sp.) were found to produce

substantial amount of all three types (acidic, neutral and alkaline) of extracellular

Organophosphorus Phosphatases (OPP). Among the four strain. Pseudomonas

261

kilonensis MB490 was observed to produce maximum amount of neutral OPP. While,

Bacillus thuringiensis MB497 displayed highest production of alkaline OPP,

Pseudomonas sp. MB504 showed highest production of acidic OPP, whereas

Pseudomonas kilonensis MB498 was found to produce significant amount of both

acidic and alkaline phosphatases. On the whole, acidic and alkaline OPP production

was greater than neutral OPP.

There had been previous findings of extracellular alkaline phosphatase production in

various microbes like Pyrococus abyssi, Streptomyces griseus, Bacillus subtilis

KIBGE-HAS (Zappa et al., 2001; Moura et al., 2001; Shah et al., 2008). Bacterial

extracellular alkaline phosphatase production is induced under low phosphate

availability thus indicating its involvement in phosphate breakdown (Combs Jr. et al.,

1979). Phosphatases are considered as subclass of Organophosphorus hydrolases due

to similarity of mechanism of hydrolytic action to breakdown organophosphates

(Gandhi and Chandra. 2012).

Moreover, phosphatase activity is considered to comprise of activities of

phosphodiesterase (PDE) and phosphomonoesterase (PME) which are two

corresponding though discrete enzymes. PDE is responsible for the hydrolysis of

intricate P containing organic compounds like nucleic acids and phospholipids to form

phosphomonoesters such as inositol phosphates and mononucleotides, which are then

decomposed by PME into orthophosphate to make them available for direct uptake by

plants and microbes (Rejmankova et al., 2011; Stone and Plante, 2014).

In the current findings, OPP production was maximum at pH 11 by all isolates. Gothwal

(2014) reported that OPH extracted from Brevundimonas diminuta was optimized at

pH 7.5, while for OPH NL01 isolated from Pseudomonas aeruginosa NL01, optimum

pH was reported to be at pH 8 (Najavand et al., 2012), and for His6-OPH extracted

from E. coli, it was found to be at pH 10.5 (Votchitseva et al., 2006).

Among four strains, Pseudomonas sp. MB504 was noticed for significant production

of OPP at wide pH range of pH 6 to 11 and at 45°C, while Pseudomonas kilonensis

MB490, Bacillus thuringiensis MB497, Pseudomonas kilonensis MB498 showed

noteworthy OPP production even at 50°C. Enzyme production was minimum by all

strains at 37°C, thus indicating that enzyme producing gene was highly expressed under

262

stress conditions of high pH and high temperature. Generally, the OPP (acidic, neutral

and alkaline) production was decreased by all isolates after 48 hrs of incubation.

As far as OPP enzyme activity and stability is concerned, it was significant over a wide

range of temperatures (37, 45, 50, 60 and 70⁰C), though it was maximum at 37⁰C and

remained still active even at highest temperature of 70⁰C by all isolates. Likewise,

generally the activity for acidic OPP was increased at lower substrate (p-NPP)

concentrations (0.06-0.6%), while for neutral and alkaline OPP it was enhanced at

higher (0.8-1.1%) substrate concentrations. Shah et al. (2008) reported an increase in

the alkaline phosphatase activity when the substrate concentration was increased up to

20 mM. Alkaline phosphatase (EC 3.1.3.1) exhibits optimal activity at alkaline pH and

is related with non-specific phosphomonoesterase activity hydrolyzing the

phosphomonoester to give inorganic phosphate and corresponding alcohol (Boulanger

and Kantrowitz, 2003). Alkaline phosphatase has been used as a vital tool in molecular

cloning and DNA sequencing and ELISA base kits (Shah et al., 2008).

In general, acidic neutral and alkaline OPP activity in all isolates was observed to be

increased with increase in incubation time up to 90 min of incubation. It is known that

phosphatase activity is affected by SDS (Rotenberg and Brautigan, 1987). Mostly,

enzyme activities are inhibited by SDS (Husberg et al., 2012). But in present study,

very diverse effects of SDS were observed on acidic, neutral and alkaline OPP derived

from four isolates. Acidic and neutral, alkaline OPP activity in MB490 and MB504 was

inhibited or remained unaffected by SDS at all given concentrations up to 300 µl, while

activity was enhanced or remained unaffected in most of the isolates at lower

concentrations. Generally, activity of acidic, neutral and alkaline OPP derived from all

isolates was inhibited by EDTA particularly at higher concentrations. Wojciechowski

et al. (2002) reported inhibitory effect of 0.1 mM EDTA on the activity of alkaline

phosphatase extracted from hyperthermophilic bacterium Thermotoga maritima.

Similarly activity of alkaline phosphatase from Saccharomyces cerevisiae was

inhibited by 5 mM of EDTA and 1 mM of vanadate (Fernandes et al., 2008).

It was noticed that divalent metal ions (Zn++, Cu++ and Cd++) increased the activity of

acidic, neutral and alkaline OPP derived from all isolates. Zinc (Zn) is known to be the

most commonly needed metal ion for alkaline phosphatase activity (Cho et al., 2007).

The activity of alkaline phosphatase derived from a hyperthermophilic archaeon

263

Pyrococcus abyssi was reported to be enhanced by the addition of Zn, Mg and Co

(Zappa et al., 2001).

There are international environmental rules and regulations to decrease the

concentration of heavy metals in the industrial effluents to acceptable levels before their

discharging into soil and water bodies. The precipitation of heavy metals mediated by

enzymes is an innovative and eco‐friendly technique for removal of deadly heavy

metals from different industrial wastes including dye, tannery, and electroplating

industries. Recently, phosphatase‐mediated bioremediation of toxic heavy metals has

been proved to play a very promising role in the treatment of municipal, industrial, and

nuclear wastewater (Chaudhuri et al., 2017). Phosphatase being a hydrolase enzyme, is

considered to have a very vital contribution in the phosphate breakdown of living

organisms through hydrolysis of organic phosphates and polyphosphates (Pasqualini et

al., 1992). So, in the current findings, all three types (acidic, neutral and alkaline) of

OPP showed their ability to bioprecipitate selected metals (Ni, Mn, Cr and Cd) up to

86-90%. Alkaline phosphatases are nonspecific reacting with a diverse types (both

natural and synthetic) of substrates and have been found in various organisms from

bacteria to higher mammals. They are capable of effectively hydrolyzing numerous

mono‐ and diesters (Chaudhuri et al., 2017). Kulkarni et al. (2016) revealed uranium

bio-immobilization by extracellular alkaline phosphatase. Alkaline phosphatase

extracted from Escherichia coli and calf‐intestine was reported to enhance

bioprecipitation of Cd2+, Ni 2+, Co 2+, Cr 3+ and Cr6+ from both single‐ion metal solutions

and industrial effluents (Chaudhuri et al., 2017). Martinez et al. (2007) reported 73-

95% precipitation of uranium by the intracellular phosphatase extracted from

Arthrobacter, Bacillus and Rahnella.

The acidic and alkaline OPP derived from all four isolates were proved to have a

broader substrate specificity against three OP pesticides (CPF, TAP and DM). Among

the two types, acidic OPP exhibited 40 to 80% degradation of CPF, TAP and DM at 50

mg/l substrate concentration within 30 min of reaction time, while alkaline OPP was

observed more active with 80 to 99% degradation. Enzymatic degradation of given

pesticides was also confirmed by GCMS analyses with the detection of same respective

metabolites of CPF, TAP and DM as discussed in previous chapters (4, 5 and 6).

264

Present results are in agreement with that reported by Thengodkar and Sivakami (2010)

where 80% of CPF was degraded by alkaline phosphatase extracted from

cyanobacterium Spirulina platensis within one hour of incubation along with detection

of TCP as metabolite. Zhang et al. (2014) studied the degradation of five OPs

(Chlorpyrifos, diazinon, fenitrothion, malathion and methyl parathion) by phosphatase

enzyme produced by ten strains of lactic acid bacteria (LAB) and reported that

phosphatase production by the isolated strains was the main factor involved for the

rapid degradation of OPs. Gothwal (2014) isolated OPH from Brevundimonas

diminuta, which was found very active against methyl parathion. Earlier, phosphatase

extracted from fungal strains (Aspergillus flavus, A. fumigatus, A. niger, A. sydowii, A.

terreus, Emericella nidulans, Fusarium oxysporum and Penicillium chrysogenum) was

reported to effectively degrade three OPs pyrazophos, lancer and malathion (Hasan,

1999). Islam et al. (2010) revealed that organophosphorus hydrolase (OpdB) extracted

from Lactobacillus brevis WCP902 effectively degraded chlorpyrifos, coumaphos,

diazinon, methylparathion, and parathion.

Zhang et al. (2016) reported the expression of a cloned gene for Triazophos hydrolase

from Burkholderia sp. SZL-1 which was found very active against TAP degradation.

Kumar (1996) revealed that microbial degradation of dimethoate depends upon

esterases and phosphatases. Megeed and El-Nakieb (2008) suggested degradation of

Dimethoate by phosphotriestrase.

Conclusion

The current research study revealed the microbial diversity of the agricultural soils of

Pakistan. Selected indigenous bacterial strains (Pseudomonas kilonensis MB490,


MB504) were able to tolerate high concentrations of OP pesticides (CPF, TAP and

DM), heavy metals (Pb, Ni, Cu, Zn, Mn, Cr, Fe), and organic pollutants (Benzene,

toluene, Xylene, Naphthalene, biphenyl).

These strains were capable of effectively degrading Chlorpyrifos, Triazophos and

Dimethoate under a wide range of pH (6, 7, 8), temperature (25, 30, 37°C) and other

incubation conditions. Moreover, another distinctive and rare feature of these strains

was their capability to tolerate and transform highly recalcitrant metabolite of

265

Chlorpyrifos, namely 3, 5, 6-Trichloropyridinol. GCMS analyses confirmed the

biotransformation of three given pesticides by the detection of many unique and novel

metabolites along with known byproducts.

In addition to have nitro reductase, oxygenase and peroxidase, all the selected strains

were found to produce significant amount of acidic, neutral and alkaline

organophosphorus phosphatase (OPP) enzyme which showed stability and activity

under a broad range of pH, temperature and metal ions. This OPP enzyme showed great

capability for metal bioprecipitation and a diverse substrate specificity against the three

OP pesticides (CPF, TAP and DM). On the basis of all these facts, the four indigenous

bacterial isolates can be suggested as very strong candidates for the practical application

to bioremediate OP pesticide contaminated agricultural soils of Pakistan under local

climatic conditions for which these are better adopted.

Future Perspective

The four selected strains in the current study possibly possess genes encoding

phosphatase and other enzymes needed for degradation of OP pesticides. Further

studies are required to explore these OP degrading genes involved in the regulation of

concerned enzymes vital for OP biodegradation like OPPs, Oxidases, Nitro reductase,

Amidase etc. in the given strains through gene based sequencing which could not be

achieved in the present study due to limitation of time and funds. This would lead to

genetic basis of biodegradation. Genes once traced could be used to improve OP

degrading potential of other bacteria which are currently non-degrading or less-

degrading through genetic transformation. Furthermore, detailed open field studies of

present bacterial strains must be conducted to evaluate their practical application under

open natural field conditions.

266

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