physiological and molecular characterization of

Physiological and molecular

characterization of organophosphate

pesticide (profenofos and chlorpyrifos)

degrading bacterial strains

Hina Jabeen

Submitted in partial fulfillment of

requirement for the degree

Doctor of Philosophy

2015

Department of Biotechnology (NIBGE)

Pakistan Institute of Engineering and Applied Sciences

Nilore-45650 Islamabad, Pakistan

National Institute for Biotechnology and Genetic Engineering

P. O. BOX 577, JHANG ROAD, FAISALABAD.

(Affiliated with PIEAS, Islamabad)

Declaration of Originality

I hereby declare that the work accomplished in this thesis is the result of my own research

carried out in Environmental Biotechnology Division (EBD), NIBGE. This thesis has not

been published previously nor does it contain any material from the published resources that

can be considered as the violation of international copyright law.

Furthermore, I also declare that I am aware of the terms “copyright” and “plagiarism”, and if

any copyright violation was found out in this work, I will be held responsible of the

consequences of any such violation.

Signature: _______________

Name of the Student: Hina Jabeen

Registration No. 10-7-1-015-2008

Date: ______________

Place: NIBGE, Faisalabad

National Institute for Biotechnology and Genetic Engineering

P. O. BOX 577, JHANG ROAD, FAISALABAD.

(Affiliated with PIEAS, Islamabad)

Research Completion Certificate

Certified that the research work contained in this thesis entitled “Physiological and

molecular characterization of organophosphate pesticide (profenofos and

chlorpyrifos) degrading bacterial strains” has been carried out and completed by

“Hina Jabeen” under my supervision during her PhD studies in the subject of

Biotechnology.

____________________ _________________

Date Dr Samina Iqbal

Research Supervisor

Submitted Through

________________

Dr Shahid Mansoor

Director NIBGE

Certificate of Approval

This is to certify that the work contained in this thesis entitled “Physiological and

molecular characterization of organophosphate pesticide (profenofos and

chlorpyrifos) degrading bacterial strains”, carried out by “Hina Jabeen” in our

opinion is fully adequate, in scope and quality, for the degree of Doctor of Philosophy in

Biotechnology from Pakistan Institute of Engineering and Applied Sciences (PIEAS).

Approved by:

Internal Examiner/Supervisor:

Signature: _________________

Name: Dr. Samina Iqbal

External Examiner

Signature: _________________

Name: Dr Irshad Hussain

External Examiner:

Signature: _________________

Name: Dr Tahira Iqbal

Verified by:

Signature: _________________

Name: Dr. Shahid Mansoor

Head, Department of NIBGE (Biotechnology)

Stamp: _____________________

Dedicated to My beloved Parents, Bhaya,

And My grandfather

A spiritual companion who guided me

at all the steps of this task

Allah Bless his Soul!

Acknowledgements

Foremost, I am thankful to ALLAH ALMIGHTY, the only Creator, for blessing me with the

opportunity to seek knowledge and granting me success and determination through all the

phases of life. My deepest feelings of gratefulness are for HOLY PROPHET (PEACE BE

UPON HIM), the most loved one of Allah Almighty and a role model for humanity in all the

aspects forever. I would like to extend my gratitude to Higher Education Commission (HEC),

Pakistan for providing me the financial support for PhD studies and making my dreams come

true. My special gratitude for Dr Shahid Mansoor, Director NIBGE, for providing an excellent

environment for research and academics. Next I would like to pay my heartiest gratitude to

someone I revere as much, my supervisor, Dr Samina Iqbal, as she molded me as an

independent researcher through her continuous advices, systematic guidance, encouragement

and support. No words can suffice her role in my accomplishment.

I am greatly obliged to Dr Rebecca Parales, my foreign supervisor and Juan Parales,

foreign instructor for their cordial advices, endeavors and support during the tenure of IRSIP

fellowship (International Research Support Initiative Program, HEC) at University of

California, Davis, CA, USA. I am profoundly indebted to Head Environmental Biotechnology

Division, (SEBD) NIBGE Dr Qaiser Mehmood Khan for his kind attitude, guidance and

cooperation. I wish to express my sincere gratitude to Samina Anwar for her support, advices

and encouragement at all the points during my research which helped me to sum up my

determination and courage at many hopeless moments in research life. I am really grateful to

Dr Mohammad Afzal for his sincere advices and guidance during this research. I offer my

generous thanks to Dr. Sajjad Mirza for providing HPLC facility in his laboratory and for his

help in plant growth promoting analysis of bacteria, to Dr. Asma Imran for her cooperation

and support in the accomplishment of Confocal Laser Scanning Microsopic analysis to

complete a part of this research. I would like to say heartfelt thanks to Dr Mohammad

Ibrahim Rajoka for his valuable guidance in kinetic analysis of pesticides degradation.

My cordial gratitude to my friends Tanveer Majeed, Mariaum Zain, Muissa Fatima,

Saira Ali, and Maryam Zafar for their constant help and company, my lab fellows Sadiqa

Firdous, Fiaz Ahmad and Muhammad Asif Nadeem for their contributions which goes

beyond any measurable value and all my colleagues at NIBGE.

For any successful task, the support, faith and blessings of near and dear ones is a

must. I am totally indebted to my great grandparents, my beloved father and mother, my

brothers, sisters and bhabi, for their true love, patience, understanding and endurance during

the whole tenure of this journey. Their prayers traced me everywhere which were the source of

hope and success in this task. I cannot express my feelings for the everlasting love of my

mother and her prayers and I can never possibly repay. I can never forget to mention the name

of my beloved cousin & an affectionate companion, Saman Hina, for her great support,

encouragement and well wishes at all the points of this task.

Hina Jabeen

i

Table of Contents

Acknowledgement

Table of Contents................................................................................................................i

List of Figures..................................................................................................................vii

List of Tables....................................................................................................................xii

List of Abbreviations......................................................................................................xiii

Abstract............................................................................................................................xv

Chapter 1

Introduction and Review of Literature

1.1 Environmental xenobiotics ................................................................................... 1

1.2 Pesticides .............................................................................................................. 1

1.3 Pesticides: A necessary evil ................................................................................. 3

1.4 Organophosphate pesticides ................................................................................. 5

1.4.1 Mode of action and toxicity of organophosphate pesticides ......................... 8

1.4.2 Environmental impact and hazards of OP pesticides .................................. 11

1.5 Pesticide situation in Pakistan ............................................................................ 13

1.6 Bioremediation/Biodegradation ......................................................................... 15

1.7 Bioremediation/Biodegradation of pesticides .................................................... 15

1.8 Microbial degradation of organophosphate pesticides ....................................... 18

1.9 Factors affecting biodegradation of pesticides ................................................... 24

1.10 Organophosphate degrading enzymes ................................................................ 27

1.11 Bioremediation of organophosphate contaminated soil ..................................... 27

1.12 Plant microbe interaction for the remediation of pesticides/pollutants .............. 28

1.13 Objectives of the study ....................................................................................... 30

Chapter 2

Materials and Methods

2.1 Chemicals ........................................................................................................... 31

2.2 Bacterial strains used in the study ...................................................................... 31

2.3 Soil collection ..................................................................................................... 32

ii

2.4 Growth media ..................................................................................................... 32

2.4.1 Maintenance and preservation of the bacterial strains ................................ 32

2.5 Equipment used in the study .............................................................................. 33

2.6 Enrichment of profenofos and chlorpyrifos degrading bacterial strains ............ 33

2.7 Isolation of pesticide degrading bacterial strains ............................................... 33

2.8 Molecular characterization of bacterial isolates ................................................. 34

2.8.1 DNA Isolation ............................................................................................. 34

2.8.2 Preparation of Heat shock competent cells (C-cells) .................................. 35

2.8.3 Amplification of 16S rRNA gene from bacterial isolates ........................... 35

2.8.4 Agarose gel electrophoresis ........................................................................ 36

2.8.5 Ligation and cloning of the 16S rRNA gene .............................................. 36

2.8.6 Sequencing of 16S rRNA gene and bacterial identification ....................... 36

2.9 Morphological and biochemical characteristics of the bacterial isolates ........... 37

2.9.1 Morphological characterization .................................................................. 37

2.9.2 Physiology and Biochemical characterization ............................................ 37

2.9.2.1 Gram staining .......................................................................................... 37

2.9.2.2 Antibiotic resistance of isolates............................................................... 38

2.10 Inoculum preparation ......................................................................................... 38

2.11 Experimental set up for pesticide degradation studies ....................................... 39

2.11.1 Determination of the detection wavelength of the pesticides ..................... 39

2.11.2 Extraction of pesticide residues from liquid cultures ................................. 39

2.11.3 HPLC conditions for pesticide residual analyses........................................ 40

2.12 Soil microcosm studies (Pot Experiments) ........................................................ 40

2.12.1 Soil collection for microcosm experiments ................................................ 40

2.12.2 Determination of Maximum Water Holding Capacity (MWHC) of the

soil ............................................................................................................... 40

2.12.3 Preparation of pesticide-contaminated soil ................................................. 41

2.12.4 Extraction and analysis of pesticide residues from soil .............................. 41

2.12.5 Optimization of soil moisture on pesticide degradation ............................. 41

2.12.6 Optimization of inoculum density for pesticide degradation in soil ........... 42

2.13 Identification of pesticide metabolites ............................................................... 42

iii

2.14 Study of potential genes encoding hydrolases/oxygenases in pesticide degrading

bacterial strains ...................................................................................................... 43

2.14.1 Amplification of the OP degrading genes ................................................... 43

2.14.2 Agarose gel electrophoresis, cloning and sequencing of the amplified

gene ............................................................................................................. 43

Chapter 3

Isolation, characterization and degradation potential of profenofos


3.1 Introduction ...................................................................................................... 46

3.2 Materials and Methods .................................................................................... 49

3.2.1 Development of profenofos degrading bacterial consortium ...................... 49

3.2.2 Molecular identification of PFF degrading bacterial strains comprising the

consortium PBAC ....................................................................................... 50

3.2.3 Morphology and biochemical analysis of PFF degrading bacterial isolates50

3.2.4 Biodegradation of PFF by pure cultures and bacterial consortium PBAC . 50

3.2.5 Extraction and HPLC analysis of PFF residues .......................................... 50

3.2.6 Optimization of culture conditions for PFF degradation using Response

surface Methodology (RSM) ...................................................................... 51

3.2.7 PFF degradation by PBAC at different initial concentrations .................... 52

3.2.8 Soil microcosm studies for PFF degradation .............................................. 52

3.2.9 Identification of PFF metabolites................................................................ 52

3.2.10 Study of potential genes encoding OP hydrolases/oxygenases .................. 52

3.2.11 Biodegradation of other pesticides .............................................................. 53

3.2.12 Data analysis ............................................................................................... 53

3.3 Results ............................................................................................................... 54

3.3.1 Molecular identification of PFF degrading bacterial isolates ..................... 54

3.3.2 Morphological and biochemical characterization of PFF degrading bacterial

isolates......................................................................................................... 60

3.3.3 Antibiotic resistance assay .......................................................................... 63

3.3.4 Biodegradation of PFF in aqueous medium by pure cultures and PBAC .. 64

3.3.5 Biodegradation of PFF with different initial concentrations by PBAC ...... 67

3.3.6 Optimization of culture conditions for PFF degradation using RSM ......... 69

3.3.7 Response surface plots for PFF degradation ............................................... 75

3.3.8 Soil microcosm studies of PFF degradation ............................................... 80

iv

3.3.8.1 Optimization of soil moisture contents for PFF degradation .................. 80

3.3.8.2 Optimization of inoculum density for PFF degradation in sterilized soil 80

3.3.9 Identification of profenofos metabolites ..................................................... 84

3.3.10 Detection of OP degrading genes in PFF degrading bacterial strains (PF1-

PF4) ............................................................................................................. 96

3.3.11 Biodegradation of other pesticides by the bacterial consortium PBAC ..... 97

3.4 Discussion .......................................................................................................... 99

Chapter 4

Isolation, characterization and degradation potential of chlorpyrifos


4.1 Introduction

4.2 Materials and methods

4.2.1 Enrichment and isolation of CP degrading bacterial strains ..................... 108

4.2.2 Identification and characterization of selected strain HN3 ....................... 108

4.2.3 Experimental set up for CP degradation studies ....................................... 108

4.2.4 Extraction and analysis of CP residues ..................................................... 109

4.2.5 Turbidometric study to monitor the growth of the bacterial strain, HN3 . 109

4.2.6 Optimization of temperature and pH for biodegradation of CP by HN3 .. 109

4.2.7 CP degradation in minimal and complex media ....................................... 109

4.2.8 Kinetics of CP degradation by HN3 at different initial concentrations

of CP ......................................................................................................... 110

4.2.9 Biodegradation of TCP (primary metabolite of CP) ................................. 110

4.2.9.1 Biodegradation of TCP in minimal and complex media ....................... 110

4.2.9.2 Extraction and analysis of TCP residues ............................................... 110

4.2.9.3 Detection of chloride ions produced during CP and TCP degradation . 111

4.2.10 Soil microcosm studies of CP ................................................................... 111

4.2.11 Identification of CP metabolites ............................................................... 112

4.2.12 Study of potential genes encoding OP hydrolases/oxygenases ................ 112

4.2.13 Data Analysis ............................................................................................ 112

4.3 Results ............................................................................................................. 112

4.3.1 Isolation and selection of CP degrading bacterial strain ........................... 112

4.3.2 Molecular, morphological and biochemical identification of strain HN3 114

v

4.3.3 Antibiotic resistance assay ........................................................................ 114

4.3.4 Biodegradation of CP by Mesorhizobium sp. HN3 .................................. 118

4.3.4.1 Optimum temperature for the CP degradation by Mesorhizobium sp.

HN3 ....................................................................................................... 118

4.3.4.2 Optimization of pH for CP degradation by Mesorhizobium sp. HN3 ... 118

4.3.4.3 Biodegradation of CP at different initial concentrations ....................... 118

4.3.4.4 Kinetics of CP degradation and TCP accumulation and degradation

thereafter ................................................................................................. 123

4.3.4.5 Co-metabolic degradation of CP by Mesorhizobium sp. HN3 .............. 128

4.3.5 Biodegradation of TCP by Mesorhizobium sp. HN3 ................................ 130

4.3.5.1 Biodegradation of TCP at different initial concentrations .................... 130

4.3.5.2 Release of chloride ions in culture media containing CP and TCP ...... 130

4.3.6 Soil microcosm studies of CP ................................................................... 135

4.3.6.1 Optimization of soil moisture level for CP degradation in unsterilized

soil ......................................................................................................... 135

4.3.6.2 Biodegradation of CP in sterilized and unsterilized soil ....................... 135

4.3.6.3 Optimization of inoculum density for CP degradation in sterilized soil 136

4.3.7 Identification of Chlorpyrifos metabolites ................................................ 140

4.3.8 Detection of OP degrading genes in Mesorhizobium sp. HN3 ................. 155

4.4 Discussion ........................................................................................................ 157

Chapter 5

Bio-stimulation: Microbe Assisted Phytoremediation

5.1 Introduction .................................................................................................... 162

5.2 Materials and Methods .................................................................................. 163

5.2.1 Soil fortification with CP .......................................................................... 163

5.2.2 Bacterial strains used in the study ............................................................. 164

5.2.3 Plasmid used for transformation ............................................................... 164

5.2.4 Preparation of electrocompetent cells of Mesorhizobium sp. HN3 .......... 164

5.2.5 Electroporation of yfp gene into Mesorhizobium sp. HN3 ....................... 165

5.2.6 Experimental design.................................................................................. 165

5.2.7 Extraction and analysis of chlorpyrifos residues in the soil and plant ...... 166

vi

5.2.8 Detection and enumeration of the bacteria in the soil............................... 166

5.2.9 Root and shoot colonization by Mesorhizobium sp. HN3yfp ................... 167

5.2.10 Measurement of growth parameters.......................................................... 167

5.2.11 Phosphate solubilization ........................................................................... 167

5.2.12 Indoleacetic acid production ..................................................................... 168

5.2.13 Data analysis ............................................................................................. 168

5.3 Results ............................................................................................................. 169

5.3.1 Biodegradation of CP in the planted and un-planted soil ......................... 169

5.3.2 Chlorpyrifos uptake by plant .................................................................... 172

5.3.3 Colonization of Mesorhizobium sp. HN3 in soil ...................................... 173

5.3.4 Colonization of Mesorhizobium sp. HN3 in the roots and shoots ryegrass

173

5.3.5 Plant biomass......................................................................................... 178

5.3.6 Plant growth promoting properties of Mesorhizobium strain HN3 .......... 181

5.3.6.1 Indoleacetic acid production .............................................................. 181

5.3.6.2 Phosphate solubilization .................................................................... 181

5.4 Discussion ........................................................................................................ 182

Chapter 6

General Discussion.........................................................................................................186

Chapter 7

References ......................................................................................................................191

Appendices

Publications

vii

LIST OF FIGURES

Figure 1.1 General chemical structure of organophosphate pesticides 6

Figure 1.2 Normal Mode of action of acetycholinesterase in the absence of OP

compounds 10

Figure 1.3 Mechanism of action of OP compounds or inhibition of

acetylcholinesterase enzyme 10

Figure 1.4 Pesticides movement in the environment after their application 11

Figure 1.5 Trend of insecticide use in Pakistan 14

Figure 1.6 Hydrolysis of organophosphate by a bacterial phosphotriesterase enzyme 18

Figure 1.7 Plant microbe interaction for the remediation of pesticide contaminated

soil 29

Figure 3.1 Chemical structure of profenofos 46

Figure 3.2 Restriction Fragment Length Polymorphism of IGS gene from PFA-PFH 55

Figure 3.3 Neighbor joining tree showing the phyllogenetic relationship of strain PF1 56




Figure 3.7 Morphology of profenofos degrading pure bacterial isolates grown on LB-

agar medium 62

Figure 3.8 Degradation of profenofos and its metabolite BCP by pure bacterial

isolates and the consortium PBAC 65

Figure 3.9 Degradation of profenofos by the PBAC at different initial concentrations of

profenofos in MSM 68

Figure 3.10 The parity plot of PFF degradation (%) 72

Figure 3.11 Coutour and Response surface plots for profenofos degradation (%) as a

result of interaction of pH and temperature at constant inoculum size 76

Figure 3.12 Contour and response surface plots for profenofos degradation (%) as a

result of interaction of pH and inoculum size at constant temperature 77

Figure 3.13 Contour and response surface plots for profenofos degradation (%) as a

result of interaction of temperature and inoculum size at constant pH 78

viii

Figure 3.14 Optimization Ramp for profenofos degradation 79

Figure 3.15 Degradation (% ) of profenofos by the bacterial consortium PBAC in the

sterilized soil at different moisture levels 82

Figure 3.16 Degradation (% ) of profenofos by the bacterial consortium PBAC in the

sterilized soil at different inoculum sizes 83

Figure 3.17 Total ion chromatogram (TIC) of the extract of the spent culture medium

containing profenofos at zero time 86

Figure 3.18 Mass spectrum of of profenofos 87


containing profenofos harvested after 24 h 88


containing profenofos harvested after 72 h. 89


containing CP harvested after 96 h 90

Figure 3.22 Mass spectrum of 4-bromo-2-chlorophenol (BCP), the hydrolysis

products of profenofos 91

Figure 3.23 Mass spectrum of O- ethyl-S-propyl-O-hydrogen phosphorothioate

(EPHP), the hydrolysis products of profenofos 92

Figure 3.24 Mass spectrum ethylene glycol, a proposed ring cleavage product of

BCP 93

Figure 3.25 Mass spectrum of 4-bromo-2-chlorophenyl ethyl propyl phosphate

(BCPEPP) 94

Figure 3.26 Proposed biodegradation pathway of Profenofos by the bacterial

consortium 95

Figure 3.27 Amplification of opdA gene encoding an OP hydrolase (OPAA) in PFF

degrading bacterial strains (PF1-PF2) 96

Figure 3.28 Degradation of PFF and other pesticides by bacterial consortium 98

Figure 4.1 Chemical structure of chlorpyrifos 103

Figure 4.2 Degradation (%) of chlorpyrifos by 8 selected bacterial strains 113

Figure 4.3 Mesorhizobium sp. HN3 grown on LB-agar plate after 48 h of incubation 115

Figure 4.4 Scanning Electron Microscopy image of Mesorhizobium sp. HN3 115

ix

Figure 4.5 UPGMA tree showing the phyllogenetic relationship of strain HN3

116

Figure 4.6 Degradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 as a sole

source of carbon and energy at different incubation temperatures 120


source of carbon and energy at different initial pH 121

Figure 4.8 Biodegradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 at different

initial concentrations as a sole source of carbon and energy 122

Figure 4.9 Kinetics of CP degradation at 37C by Mesorhizobium sp. HN3 at different

initial concentrations 124

Figure 4.10 First order kinetics of chlorpyrifos degradation in MSM at different initial

concentrations 126

Figure 4.11 Co-metabolic degradation of chlorpyrifos by Mesorhizobium sp. HN3 128

Figure 4.12 Biodegradation of chlorpyrifos by Mesorhizobium sp. HN3 in nitrogen

free medium 129

Figure 4.13 Biodegradation (%) of TCP by Mesorhizobium sp. HN3 as a sole source

of carbon and energy at different initial concentrations 131

Figure 4.14 Co-metabolic degradation of TCP by Mesorhizobium sp. HN3 132

Figure 4.15 Analysis of chloride ions produced as a result of chlorpyrifos

degradation in a chloride free medium 133

Figure 4.16 Analysis of chloride ions produced as a result of TCP degradation in a

chloride free medium 134

Figure 4.17 Biodegradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 in the

unsterilized soil at different soil moistures 137


unsterilized and sterilized soil 138


sterilized soil at different inoculums densities 139


containing CP at zero time 142

Figure 4.21 Mass spectrum of chlorpyrifos (CP) 143

x


containing CP harvested at 48 h (2 days)

144

Figure 4.23 Mass spectrum of 3,5,6-trichloro-2-pyridinol 145

Figure 4.24 Mass spectrum of Diethylthiophosphate (DETP) 146

Figure 4.25 Mass spectrum of 3,5,6 trichloro-2-methoxypyridine (TMP) 147


containing CP harvested after 72 h (3 days). 148

Figure 4.27 Mass spectrum of 3,5-dichloropyridine 149

Figure 4.28 Mass spectrum of 3-chloro-2-pyridinol 150

Figure 4.29 Mass spectrum of 3,5-trichloro-2-methoxypyridine 151


containing CP harvested after 120 h (5 days) of incubation. 152

Figure 4.31 Mass spectrum of chromatogram of maleamic acid 153

Figure 4.32 Predicted biodegradation pathway of chlorpyrifos 154

Figure 4.33 Amplification of opdA gene encoding an OP hydrolase (OPAA) in CP

degrading Mesorhizobium sp. HN3 156

Figure 4.34 Amplification pcaH gene encoding an protocatechuate dioxygenase in CP

degrading Mesorhizobium sp. HN3 156

Figure 5.1 Degradation of CP and accumulation & subsequent disappearance of TCP

by ryegrass (Lolium multiflorum) and Mesorhizobium sp. 170

Figure 5.2 CLSM images (10X) of time course colonization process of Lolium

multiflorum (ryegrass) roots by Mesorhizobium sp. HN3yfp after 15 days of

inoculation

174



inoculation

175



inoculation

176

Figure 5.5 A comparison of shoot lengths and root lengths among different ryegrass

plant treatments 179

xi

Figure 5.6 Qualitative test of IAA production by Mesorhizobium sp. HN3 using Ferric

chloride

181

Figure 5.7 Qualitative test of phosphate solubilization by Mesorhizobium sp. HN3 on

Pikoviskaya medium 181

xii

List of Tables

Table 1.1 Some commonly used organophosphate pesticides and their metabolites 7

Table 1.2 Microorganisms isolated for the biodegradation of organophosphorus pesticides 21

Table 2.1 Bacterial strains used in the study 32

Table 2.2 Antibiotics used in the study 38

Table 2.3 Sequences of the previously reported primers used in this study 44

Table 2.4 Primer sequences designed by aligning the already reported

organophosphate degrading genes 45

Table 3.1 Experimental ranges and levels of independent variables 51

Table 3.2 Percent (%) similarity of profenofos degrading bacterial strains with reported

16SrRNA gene sequences in the GenBank 54

Table 3.3 Biochemical characteristics of profenofos degrading bacteria 61

Table 3.4 Response of profenofos degrading bacterial strains to different antibiotics 63

Table 3.5 Degradation kinetics of profenofos by pure and mixed cultures 66

Table 3.6 The 23 factorial and central composite design for experiment 71

Table 3.7 The CCD matrix showing actual values (%) along with the experimental values

of PFF degradation 73

Table 3.8 Analysis of Variance (ANOVA) for the response (% degradation of PFF) 74

Table 3.9 Different metabolites of profenofos and their detail 85

Table 4.1 Previously reported chlorpyrifos and TCP degrading bacterial strains 106

Tabl Table 4.2 Biochemical and morphological characteristics of Mesorhizobium sp. HN3 117

Table 4.3 Kinetic parameters for chlorpyrifos degradation and product (TCP) formation 125

Table Table 4.4 First order kinetics parameters for chlorpyrifos degradation by Mesorhizobium

sp. HN3 in liquid medium (MSM) 127

Table 5.1 First order kinetics parameters of chlorpyrifos degradation in planted and un-

planted soils 171

Table 5.2 CP uptake and accumulation in roots and shoots of ryegrass 172

Table 5.3 Colonization of Mesorhizobium sp. HN3yfp in planted (ryegrass) and un-planted

soil 177

Table 5.4 Effect of chlorpyrifos on plant growth parameters 180

xiii

LIST OF ABBREVIATIONS

2,4-D 2,4-Dichlorophenoxyacetic acid

AcH Acetylcholine

AChE Acetylcholinesterase enzyme

ASS Antibiotic sensitivity sulphonamide agar

CFU Colony Forming Units

CLSM Confocal Laser Scanning Microscope

CP Chlorpyrifos

DDT Dichloro-diphenyl-trichloro-ethane

DETP diethyl thiophosphoric acid

ECDs Endocrine disruptors

EPA Environmental Protection Agency

EPHP O- ethyl-S-propyl-O-hydrogen phosphorothioate

GC-MS Gas Chromatography Mass Spectrometer

HPLC High Performance Liquid Chromatography

IAA Indoleacetic acid

mpd Methyl parathion degrading

MPH Methyl parathion hydrolase

MS Mass Spectrum

MSM Minimal salt medium

NFM Nitrogen Free Medium

OC Organochlorine

OP Organophosphorus

xiv

OPAA Organophosphorus acid anhydrolase

opd Organophosphate degrading

opdA Organophosphorus acid anhydrolase gene

OPH Organophosphorus hydrolase

pcaH Protocatechuate hydrolyzing (gene)

PFF Profenfos

PGPR Plant growth promoting Rhizobia

POPs Persistent organic pollutants

P-solubilization Phosphate solubilization

SEM Scanning Electron Microscopy

TCP 3,5,6 trichloro-2-pyridinol

TMP 3,5,6 trichloro-2-methoxypyridine

WHO World Health Organization

xv

ABSTRACT

Organophosphate pesticides (OPs) are the synthetic chemicals that have broad

applications in agriculture for controlling different kinds of pests such as insects and

weeds etc. They poison the insects and mammals by paralyzing their central nervous

system which is linked to many acute and long term health disorders. Two of the most

widely used and broad-spectrum OP pesticides are the chlorpyrifos (CP) and profenofos

(PFF) which are used for protecting various crops against serious insect pests. However,

continuous and indiscriminate use of these pesticides is of great concern due to their

serious impacts and hazards on the environment and humans. Remediation of these toxic

pesticides and related contaminants using microorganisms having the right metabolic

pathways seems to be the most effective technology. Objectives of this study were to

isolate and characterize bacterial strains capable of complete degradation of CP, PFF and

their toxic metabolites, optimize culture conditions that govern degradation of these

compounds by the isolated bacteria and investigate the pathways of degradation.

A chlorpyrifos degrading bacterial strain, Mesorhizobium sp. HN3 was isolated and

characterized. Time course shake flask experiments and kinetic analysis revealed high

efficiency of Mesorhizobium sp. HN3 for CP degradation up to 300 mg/L at range of at a

broad range of culture conditions. Importantly, HN3 also degraded 3,5,6 trichloro-2-

pyridinol (TCP), a more toxic and persistent metabolite of CP. Further, enhanced CP

degradation in soil was achieved by the combined use of Mesorhizobium sp. HN3 and

ryegrass (Lolium multiflorum). Moreover, a yfp-tagged variant of Mesorhizobium sp.

HN3 (HN3yfp) was used to study the colonization of this strain in the rhizosphere and

endosphere of ryegrass. The strain HN3yfp proficiently colonized the rhizosphere & roots

of ryegrass, removed CP and TCP residues uptaken by the plant thus enhanced plant

growth.

For PFF degradation, a bacterial consortium PBAC, consisting of Achromobacter

xylosoxidans, Pseudomonas aeruginosa, Bacillus sp. and Citrobacter koseri, was

isolated. PBAC was capable of degrading PFF and its toxic hydrolysis product 4-bromo-

2-chlorophenol (BCP). The efficacy of PFF degradation was modeled by central

composite design (CCD) based on response surface methodology (RSM). The

simultaneous effects of three test interacting factors on the PFF degradation (%) were

xvi

monitored and conditions were optimized for maximum degradation of PFF. Gas

Chromatography Mass Spectrometry (GC-MS) analysis of CP and PFF provided plenty

of information regarding their metabolites and hence biodegradation pathways of the two

pesticides were predicted successfully. The detection of dehalogenation and ring cleavage

metabolites of the pesticides indicated the complete degradation of the toxic pesticides.

The overall study indicates that CP degrading Mesorhizobium sp. HN3 and PFF

degrading bacterial consortium PBAC are the promising candidates for the remediation

of OP contaminated sites. Further, the study provides insight into the fate and

biodegradation pathways of the two pesticides.

Validity of the study is that fate of TCP or BCP have seldom been addressed. Rather,

previous reports emphasis on the parent compound degradation. But the degradation of

the metabolites is more important due to the fact that OP pesticides degrade to their

metabolites soon after they reach soil. Metabolites are usually more toxic and persistent

than the parent compounds. Moreover, to best of our knowledge this is the first study

involving the elaborately designed optimization experiments for profenofos degradation

by a diverse bacterial consortium. Also, degradation of BCP by the microbial

communities has not already been reported.

Chapter 1 Introduction and Review of Literature

1

Chapter 1

Introduction and Review of Literature

1.1 Environmental xenobiotics

The term xenobiotic originated from the Greek word xeno which means “foreign”

and biotic means “pertaining to life” hence xenobiotics are generally referred to as

compounds that are foreign to living bodies. Xenobiotics are the synthetic chemicals that

were developed in the last century with the major purpose of benefit to mankind. Such

synthetic compounds have been employed as explosives, pesticides, dyes, solvents and

refrigerants in industrial, urban and agricultural applications. However, an ever mounting

production of synthetic chemicals is attributed largely to fast growing global

industrialization and new innovations in synthetic chemistry. Generally the term

xenobiotic is used with the context of pollutants or contaminants that are harmful for the

life. This is because the discharge and accumulation of these noxious compounds into the

environment have become enormous (Duong et al., 1997). Millions of kilograms of toxic

chemicals are discharged worldwide in water, air and soils as reported by Third World

Network (TWN) reports. These toxic chemicals cause environmental problems thus

disturbing the original balance in nature. Although, they enter the environment at very

low residual concentrations but their consequences are aggravated due to bio-

accumulation and bio-concentration leading to the overall disturbed quality of the

environment. Although, scientists all over the world are trying to develop strategies to

reduce the indiscriminate use of these pollutants so as to overcome the problems

associated with them, however, their words are not well given attention and many

substances are still in use without considering their unfavorable impact (Shukla et al.,

2010).

1.2 Pesticides

Among all xenobiotics, pesticides are the most widely used all over the world.

Pests are defined differently depending on different habitats and the roles they play in

different locations. For example, in rural areas, pests include all the species of insects,

weeds, bird, mites, slugs, rodents, snails and arthropods that cause damage to the crops.


2

For urban inhabitants, pests include all kinds of mosquitoes, flies, insects, bacteria and

viruses that can be vectors for diseases, cause disturbance & unhygienic conditions in the

environment and endanger health and comfort. The term pesticide, in particular describes

a group of chemical substances that modify natural processes of living organisms deemed

to be pests, whether these are insects, mold or fungi, weeds or deleterious plants.

Environmental Protection Agency (EPA) which is the primary regulator of the pesticide

use, defines pesticide more elaborately as: “any substance or mixture of substances

intended for preventing, destroying, repelling or mitigating any pest.”

Pesticides are classified on the basis of their physical properties, chemical

structure, target organism and mode of action. Based on the target organism, pesticides

are named as insecticides (used to kill the insect pests of crops and also mosquitoes, flies

and the insect vectors for human diseases), herbicides (intended to mitigate the unwanted

plants), fungicides (to kill fungi), avicides (to kill bird pests) and acaricides (to get rid of

tick and mites). Classification of pesticides based on chemical structure is usually more

preferred by the scientists because mode of action, toxicity and characteristics of

pesticides depend on the chemical structure. Major chemical families of pesticides

include organochlorines (OCs), organophosphates (OPs), carbamates and pyrethroids.

Organochlorines are the organic compounds with several chlorine atoms. Owing to

the presence of chlorine atoms, they are highly persistent and non biodegradable hence

stay in the environment for long period of time. Their mode of action includes the

disruption of sodium or potassium balance of nerve fibers thus continuously transmitting

the nerve impulses. Commonly known OC pesticides are DDT, eldrin and endosulfan..

Organophosphate pesticides are the organic phosphorus containing substances

which control the target pests by irreversibly inhibiting the acetyl cholinesterase (AChE)

enzyme (Clark, 2006). Currently, organophosphates are the most widely used insecticides

as they effectively control the serious insect pests and are less persistent compared to

organochlorines. Commonly used OP pesticides include chlorpyrifos, diazinon,

profenofos, parathion and triazophos.

Carbamates are the derivatives of carbamic acid and they also act as cholinesterase

inhibitors but they cause reversible and less severe inhibition compared to that caused by

OPs. Some of the carbamates include carbaryl, carbofuran and aldicarb. Pyrethroids are


3

analogs of the naturally occurring insecticides found in the pyrethrum extracted from the

chrysanthemum flowers. Being highly effective against insect pests at low concentrations,

they are also getting popularity for crop protection.

1.3 Pesticides: A necessary evil

Pesticides are very old existing since around 1000 BC when ancient Chinese

started using sulphur for the control of fungi and bacteria. After that, arsenic and honey

were used as a mixture to kill insect pests. Arsenic compounds were used as herbicides as

well as insecticide in late 1800’s. By the late 1900s, a compound called Paris green

(copper acetoarsenite) became a tool for farmers in USA to get rid of the agricultural

pests (Lah, 2011). These primitive forms of pesticides paved the way for developing

advanced and effective formulations of pesticides in the hope of providing more benefits

to mankind.

Use of advanced synthetic chemicals which include, herbicides, insecticides and

fertilizers (collectively called agrochemicals) started in late 1930s and early 1940s to

enhance crop quality and yield. It was the time when world population was increasing

tremendously more than the availability of resources. The population reached 6.0 billion

in the year 2000, and is projected to increase to approximately 8.0 billion by the year

2025. Hence there developed an immense need of improving the quality and quantity of

the food to support everyday growing population. Pesticide popularity soared after the

insecticidal properties of many organochlorine compounds including aldrin, DDT

(dichloro-diphenyl-trichloro-ethane), endrin and dieldrin were discovered. Being

inexpensive and effective; these products were widely used with DDT being the most

popular owing to its strong and broad-spectrum activity. These chemicals controlled a

great variety of sucking and chewing insects as well as aphids, spiders, mites, and other

pests that attack important crops like sugarcane, cotton, tobacco, peanuts, many fruits and

vegetables. Moreover, these chemicals helped in controlling many life threatening

diseases such as yellow fever, malaria and typhus by killing the insect vectors responsible

for the spread of these diseases. In this way many insect control programs were able to

save millions of lives. Hence, the use of pesticides both in the field of public health and


4

crop protection increased and they emerged as a necessary tool for economical pest

management in modern agriculture.

Unluckily, most of the synthetic pesticides, predominantly the chlorinated

compounds are highly persistent in the environment causing toxicity to all life forms

including humans. The adverse impact of these toxic chemicals can extend from the point

of production or application to a large-scale and even to global level. Chemical pesticides

can harm agricultural workers who do not follow the safety guidelines such as proper

protective dressing. Usually, human exposure to pesticides occurs through ingestion

(accidentally or intentionally eating the contaminated food) or inhalation (though air

contaminated with pesticide residues as a result of spray on crops).

Public concerns regarding pesticide use appeared after Rachel Carson’s book,

“Silent Spring” was published in 1962, which addressed the toxicity and hazards

associated with DDT and other related pesticides. The book helped in developing

awareness in the people about the persistence of pesticides in the environment and

subsequent contamination of food chain. It described about the decline of bird population

due to thinning of egg shells and other toxic effects in birds. Further, it has been reported

extensively that organochlorine pesticides accumulate in the fatty tissues of animals and

cause chronic health effects, such as cancer, genetic, teratogenic and neurological effects

(Chaudhry and Chapalamadugu, 1991; Dua et al., 2002; Vaccari et al., 2006).

Organochlorines were also classified in persistent organic pollutants (POPs) that are

environmentally persistent, highly toxic and tend to bioaccumulate (Yu et al., 2006).

According to Stockholm Convention on POPs, nine out of the twelve POPs belong to

organochlorines. They are also designated as endocrine disruptors (ECDs) (Tsai, 2010).

World Health Organization (WHO) has classified pesticides toxicity effects in classes

ranging from class Ia to class III. Class Ia includes highly toxic pesticides and class III

includes slightly toxic pesticides (Copplestone, 1988). Most of the class Ia pesticides are

still in use in the developing countries (Bull, 1982).

Since DDT and other OC pesticides were banned by 1970s owing to their

persistence, toxicity and health effects, organophosphate and carbamate pesticides gained

popularity. Organophosphate pesticides had already been synthesized as potential warfare

agents by the Germans for use in World War II but they were kept secret. However, they


5

were re-purposed as insecticides after the organochlorine pesticides. Organophosphates

and carbamate pesticides became widespread in agriculture as they are effective against a

broad range of insects pests. Also being less persistent and less toxic, OPs were

considered environmentally safe compared to OCs. Moreover, being more biodegradable,

they were perceived as an absolute alternative to the persistent organochlorine pesticides

(Hansen et al., 1983; Coats et al., 1989).

“Silent Spring” also redirected the research towards more environment friendly

and natural types of pesticides. In this regard pyrethroids were introduced as they were

synthesized from pyrethrins that are the natural poisons for the insects. Synthetic

pyrethroids also target the nervous system of pests. Recent literature further indicates that

pesticide exposure is widespread and presents potential risks to humans, especially to

susceptible populations such as pregnant women and children (Whyatt et al., 2004 and

Eskenazi et al., 2004). Potential adverse effects of pesticide exposure to children’s health,

including reproductive outcomes, childhood cancers, neurobehavioral toxicity, and

endocrine disruption have been well studied (Garry, 2004).

1.4 Organophosphate pesticides

Organophosphorus compounds are ubiquitous and constitute a large group of

chemical agents (Kamanyire and Karalliedde, 2004). Synthetic organophosphorus (OP)

compounds are being used throughout the world as additives of pesticides, petroleum and

plasticizers and cover more than 38% of world pesticides used (Singh, 2008). Regarding

the chemical structure, they are the esters of phosphoric acids and generally consist of a

central phosphorus atom attached with a double bond to either oxygen (P=O) or sulfur

(P=S), two organic side chains (R1 and R2), and an additional side chain that behaves as

the leaving group (X) and belongs to a variety of substituted aromatic, heterocyclic or

aliphatic groups such as cyanide, thiocyanate, halide, phosphate, phenoxy, thiophenoxy,

or carboxylate group (Figure 1.1). Leaving group imparts variations in the properties of

different OP compounds. Insecticidal properties of the OP compounds depend largely on

the R1 and R2. Organophosphates with methyl or ethyl as R1=R2 are very effective

insecticides compared to those containing propyl or isopropyl groups (Fukuto, 1990).

Usually the “R” group is bonded to the phosphorus atom in different ways. Generally, an


6

oxygen or sulfur atom is present between the R group and phosphorus atom and the

structures are known as phosphate or phosphothioate respectively (Karpouzas and Singh,

2006). Some OPs contain R1 bonded to phosphorus atom through a direct bond and R2 is

bonded to phosphorus atom by a sulfur atom (thion phosphonates) or an oxygen atom

(phosphonates).

Names and chemical structures of some of the commonly used OP pesticides and their

respective hydrolytic metabolites are given in Table 1.1.

Figure 1.1 General chemical structure of organophosphate pesticides

P

O

R2(O,S)

R1(O,S)(S)

(O,S)

X


7

Table 1.1: Some commonly used organophosphate pesticides and their

metabolites

Compound Chemical structure Metabolite Chemical structure

Chlorpyrifos [O, O diethyl- O (3, 5, 6

trichloro-2-pyridyl

phosphorothioate)]

3,5,6-trichloro-2-

pyridinol

Profenofos O-4-bromo-2-chlorophenyl O-

ethyl S-propyl phosphorothioate

4-bromo-2-

chlorophenol

Br

Cl

HO

Methyl parathion O,O-dimethyl-O-p-nitrophenyl

phosphorothioate

4-nitrophenol

Diazinon O,O-Diethyl O-[4-methyl-6-

(propan-2-yl)pyrimidin-2-yl]

2-isopropyl-4-

methyl-6-

hydroxypyrimidine N

N

CH

CH3

CH3

HO

Triazophos 0, O-diethyl o-(1-phenyl-1H-1,

2, 4-triazol-3-yl)

phosphorothioate

N

N

N

O

P

OS

O

1-phenyl-1,2,4-

triazole-3-ol N

N

N

OH

Cl

N

Cl

O

Cl

P

S

O

O Cl

NCl

OH

Cl

Br

Cl

OP

O O

S

NOP

O

O

OS

O-

N

N

CH

CH3

CH3

OP

S

O

O

O

H

NO2


8

1.4.1 Mode of action and toxicity of organophosphate pesticides

Organophosphate insecticides are known to inhibit the acetyl cholinesterase

(AChE) which is a potent enzyme responsible for the normal nervous coordination in

insects, humans and other organisms (Yair et al., 2008). AChE is present in peripheral,

central and autonomic nervous system and regulates the level of acetylcholine which is

an important neurotransmitter and transmits the nerve impulses within the brain and

different parts of body. Normally, after impulse transmission, acetylcholine binds to the

active site of AChE due to its chemical structure complementarity to the active site of the

enzyme (Figure 1.2). AChE then hydrolyzes the acetylcholine to acetyl CoA and choline

to avoid accumulation of the neurotransmitter and overstimulation of muscles (Fukuto,

1990; Karpouzas and Singh, 2006).

OPs mimic the structure of acetylcholine hence easily bind to AChE thereby

inhibiting the normal activity of the AChE. Binding of OPs to active site of the enzyme

alters its structure and function and acetylcholine cannot bind to the enzyme due to

competitive inhibition (Figure 1.3). AChE inhibition leads to the accumulation of the

acetylcholine in the brain, at synapses, ganglia in the autonomic nervous system, skeletal

neuromuscular junctions, adrenal medulla and some in the sympathetic nervous system

(Chandrasekara and Pathiratne, 2005). Ultimate effects of the AChE inhibition include

headache, convulsion, disturbed breathing, paralysis and death of pests. (Ragnarsdottir,

2000).

Besides the target pests, OPs are known to adversely affect or kill other organisms

including ecologically important insects like beetles, bees, and wasps as well as aquatic

organisms e.g. protozoans, fish and tadpoles and higher animals including humans.

(Colosio et al., 2009; Jokanovic and Prostran, 2009). Routes of entry of

organophosphates to human body generally involve oral (intentional or unintentional

ingestion), respiratory (through inhalation) or dermal (through skin contact) exposure.

Dermal and respiratory exposure can also occur accidently and both are the efficient

entry routes for the pesticides (Khan et al., 2010). The inception and severity of OP

poisoning is usually determined by the exposure route, dose and physicochemical

properties of the pesticide e.g. lipid or tissue solubility and rate of

metabolism/degradation (Fukuto, 1990). Moreover, OP exposure is related to broad-


9

spectrum effects including a variety of nerve disorders and interruption of many vital

functions in higher animal and humans (Eaton et al., 2008; Ghisari and Bonefeld-

Jorgensen, 2005; Middlemore-Risher et al., 2010; Ventura et al., 2012).

OP poisoning results in a number of health effects which have been ascribed to be

the consequence of AChE inhibition. However, it was suggested that inactivation of

AChE by itself cannot account for all the adverse health effects, hence there may be some

other mechanisms responsible for OP toxicity within the organism’s body which

contribute to the overall effect (Zaki, 1982). It is now clear that, although inhibition of

AChE is a key mechanism of OPs, however, the disruption of other enzymes, individual

susceptibility and the direct exposure of tissues to OPs are also important factors to be

considered. They are very toxic to vertebrates and many reports indicate that they are

more toxic than organochlorines (Khanna et al., 1995).


10

Acetylcholine

Acetylcholine

Figure 1.3 Mechanism of action of OP compounds or inhibition of acetylcholinesterase enzyme

Figure 1.2 Normal mode of action of acetylcholinesterase in the absence of OP


11

1.4.2 Environmental impact and hazards of OP pesticides

Environment is polluted by the excessive and continuous use of pesticides. Once

these chemicals are applied on crops, they are distributed in the environment by winds,

water, by being washed off crops with rain water into soil and water reservoirs. The

contaminated soil and water reach the river systems which finally end up in the sea

(Figure 1.4). Sometimes pesticides break down to more toxic and persistent metabolites

which pollute the air, water, soil and ultimately all the ecosystems as well as their natural

biota. A huge number of environmental problems such as pollution of air, water and

terrestrial ecosystems, hazardous effects on different organisms, and disturbances in

biogeochemical cycles can be attributed to the excessive applications and large scale

synthesis of toxic chemicals.

Figure 1.4 Pesticide movement in environment after their applications


12

Approximately, 10% of the applied pesticides succeed in reaching the target site

or pests while the rest is just dispersed into soil, air and water. Consequently, they are

routinely found in non-target areas of the environment and vital non target species. The

residues and metabolites of pesticides tend to accumulate in soil at very unacceptable

levels (Shalaby and Abdou, 2010; Ortiz-Hernández, 2011). Eventual impact of the soil

contamination of pesticides is the decline of soil biodiversity. The small organisms are

essential to ecosystems because they are important in maintaining the structure and

function of natural ecosystems (Pal et al., 2005). Chlorpyrifos and glyphosate are

reported to affect earthworm and predatory arthropods respectively (Reinecke and

Reinecke, 2007; Muangphra et al., 2012).

Soil erosion and water runoff are the major ways of pesticide contamination of

aquatic ecosystems. Soluble pesticides can easily leach into lakes, streams and rivers.

Organophosphate pesticides have been shown to strongly affect the aquatic species of

vertebrates and invertebrates thus destroying natural biota (Beketov and Leiss, 2008;

DeLorenzo et al., 2001). Dichlorvos, a very toxic OP pesticide, has been reported to

cause toxicity in fish through impaired metabolism and death (Mir et al., 2012; Das,

2013). Mammals and birds are hurt by drinking the pesticide contaminated water or

eating fish and aquatic animals with toxic residues accumulated in their bodies. Such kind

of animals appears as good “indicator species”. Lethal impacts on non target life from

direct exposure include very severe kind of malfunctioning of body functions leading to

death. Indirect poisoning leads to long-term effects such as reduced growth, short

survival, and disturbed reproductive rate.

Food commodities become contaminated by direct applications of the OP

pesticides on crops. Bio-augmentation of these pollutants through food chains and food

web leads to the toxic impact on the non target organisms (Grzelak et al., 2012). OPs are

degraded as soon as they reach soil, however, degradation metabolites are often more

persistent which continue to accumulate in the environment and ultimately enter food

chain, animal and human tissues. Therefore, OPs proved to be much more toxic and

hazardous than were originally considered. For example, malathion has been reported to

affect the aquatic food web (Relyea and Hoverman, 2008).


13

1.5 Pesticide situation in Pakistan

Pesticide use in Pakistan started decades ago in the form of chemicals to control

the pests. The consumption of pesticides increased in the country from 250 Metric Tons

(MT) in 1950 to 670 MT in 1980s when their sales and distributions were transferred to

private sector (Tariq et al., 2007). According to FAO, (2002) 2.5 million tons/year

pesticides are being used worldwide including Pakistan. Pakistan is one of the developing

countries who are using pesticides at high rate in the hope of sustaining the ever

increasing population of the country. One of the reasons of high pesticide consumption is

that pesticide companies motivated the farmers to use higher dose of pesticides than the

recommended dose. Moreover, higher rate of illiteracy and lack of knowledge in the crop

growers also contributed to haphazard use of pesticides in Pakistan. Insecticides are the

most widely used (almost 85%) of all the pesticides in Pakistan. The consumption of

insecticides was highest in the year 2003-2005 followed by a decline. However, the rate

of their consumption again jumped in the year 2008 onward with a fluctuating rate of

consumption (Economic survey of Pakistan, 2012-2013) (Figure 1.5).

In Pakistan, cotton crop is under the extensive insecticide applications. The reason

behind the extreme dependence of cotton on the insecticides is that cotton was attacked

severely by the insect pests in 1995 to 1998 that reduced the overall yield and quality of

the cotton leading to severe economic loss to the country. Hence pesticides were applied

to cotton with the hope to recover the loss (Tariq, 2005; Mazari, 2005). Farmers were

attracted towards the apparent benefit and outcomes of insecticides and continued

excitedly to apply them in excessive amounts on cotton and other crops. But in this way,

these chemicals became a disaster for the country leading towards the contamination of

drinking water, food, fruits, milk, vegetables, fish and meat (Baig et al., 2009). Many

reports are present regarding the pesticide residues in water and food in Pakistan (Masud

and Farhat, 1985; Cheema and Shah, 1987; Parveen et al., 1996). According to a report

(1984) from National Institute for Health, (NIH) Islamabad, Pakistan, insecticide residues

were found in poultry, dairy products and vegetable oils. The insecticides found were

DDT, Methyl parathion, malathion and dieldrin. Commonly found insecticide residues in

water and food stuff (cotton seed oil) include organophosphates e.g. chlorpyrifos,

monocrotophos and methamidophos. (Parveen et al., 1996). Pakistani soils are also


14

reported to contain pesticides residues. Soil and groundwater contamination reports are

available from different parts of the country (Bano et al., 1991).

Figure 1.5 Trend of insecticide use in Pakistan


15

1.6 Bioremediation/Biodegradation

A number of conventional methods had been in use to eliminate or reduce the

ever increasing contamination of the planet. These methods include landfill,

incineration, and cap & contain methods and complete destruction of the pollutants by

the use of UV oxidation and chemicals decomposition (Parasad et al., 2012). However,

major drawback associated with the conventional methods is the release of toxic

intermediates into the environment (Debarati et al., 2005). Additionally, these methods

are expensive, laborious and environment unfriendly especially in extensive agricultural

areas (Jain et al., 2005).

Keeping in view the drawbacks of conventional methods, some eco-friendly

option needs to be adapted that renders the destruction and elimination of pollutants

without producing harmful substances in the environment. This aim was achieved by

exploiting the naturally existing biological properties of microorganisms and the process

is called bioremediation. ‘‘Bioremediation is a process that utilizes the metabolic

potential of living organisms such as plants, bacteria or fungi to clean up contaminated

environments, to detoxify, degrade or remove environmental pollutants”.

Bioremediation uses biological or metabolic processes of the microorganisms.

The importance of the microorganisms in bioremediation process lies in their diversity;

ubiquity and metabolic flexibility which make them use diverse ecological conditions.

Many microorganisms may grow in diverse media because of their excellent capacity of

adaptation and mutation. Furthermore, microorganisms have been found to possess

tremendous potential to acquire capacities of xenobiotic degradation when exposed to

these xenobiotics for long periods. Microorganisms can survive in almost any kind of

the environmental conditions if an appropriate source of energy and carbon is available.

1.7 Bioremediation/Biodegradation of pesticides

The extensive and widespread use of synthetic pesticides has led to a significant

effort to develop modern technologies for the elimination or reduction of these

contaminants from the environment. “Bioremediation” is a promising approach which

exploits the ability of microbial organisms (bacteria or fungi) to degrade or eliminate

toxic chemicals from the environment. This is the most effective, economical and


16

environment-friendly strategy to date which is based on environmental biotechnology

field. Environmental biotechnology is an old field that introduced the composting and the

wastewater treatment systems. But the advanced and recent accomplishments of this

research area include the bioremediation for the safe cleanup of contaminated

environment.

Attention has been focused on the isolation of bacteria with the capability of

degrading two types of compounds due to their widespread environmental problems; the

petroleum hydrocarbons; and chlorinated compounds including the pesticides. Literature

indicates that there are mainly three objectives of microbial isolations for pesticide

degradation; first is to get insights into the mechanisms of the inherent microbial

metabolism; second, to determine the mechanisms of gene/enzyme evolution and third is

to apply these microbes for the detoxification of the contaminated sites (Singh and

Walker, 2006).

A complete knowledge regarding ecological, physiological, biochemical and

molecular aspects of microorganisms has a significant role in analyzing their potential for

bioremediation (Mishra et al., 2001). Moreover, an understanding of evolutionary history

of microbes is also important in getting insights into their degradation capabilities.

Microorganisms play major roles in determining the environmental fate of chemical

pesticides because they are used as source of nitrogen, carbon or other nutrients as well as

energy. Pesticides are the recalcitrant compounds and may be resistant to complete

biodegradation in some cases because microorganisms are not adapted to utilize these

chemicals. However, frequent and repeated applications of a certain pesticide without the

crop rotation often results in the development of pesticide degrading capabilities in the

indigenous soil microorganisms, a phenomenon known as “enhanced biodegradation”

(Walker and Suett, 1986; Zhang and Bennett, 2005). Wider implications of enhanced

biodegradation were observed in crop fields in 1980s. Many reports are available that

demonstrate the use of native microorganisms in contaminated soil and sediment for the

degradation of pesticides (Walker and Suett, 1986). First report of enhanced

biodegradation of pesticides by microorganisms came in 1971 (Sethunathan, 1971).

Following the discovery of environmental persistence, a number of bacterial strains have

been isolated that are capable of degrading DDT (Nadeau et al., 1994; Bidlan and


17

Manonmani, 2002; Blasco et al., 2008; Fang et al., 2010). In the similar way, discovery

of toxic effect of other pesticides resulted in the unearthing of their degrading organisms.

By now, many microorganisms possessing pesticide utilization capabilities have been

isolated. These isolates include fungi and bacteria that can utilize the pesticides as a

source of essential nutrients e.g. carbon, phosphorus or nitrogen (Singh and Walker,

2006). Certain rhizospheric organisms were also reported to take part in degradation of

pesticide contaminants. Degradation studies in a rhizosphere system suggest that rather

than a single microorganism, a diverse microbial community is involved in the enhanced

degradation of chemical pollutants. Rhizospheric microorganisms found at

bioremediation sites mostly include Ectomycorrhizal fungi, Bacillus sp., Pseudomonas

sp. and Trichoderma sp. (Singh et al., 2002). However, after fifty years of research on the

microbial degradation, detailed knowledge about degradation pathways is available for

only a limited number of pesticides (Gomez, et al., 2007). A detailed knowledge about

the micro-organism and its environment is required that gives an insight into the

physiology, ecology, biochemistry and molecular aspects of that organism before

employing it for the remediation of a contaminated site (Iranzo et al., 2001).

The biochemical or enzymatic basis, an important aspect of microbial degradation

of pesticides, has been given considerable attention. The microbial metabolism of

pesticides is catalyzed by various extracellular or intracellular microbial enzymes

attacking a pesticide. These enzymes may be peroxidases, oxygenases or hydrolytic

enzymes. The attacked pesticide serves as a carbon, nitrogen, phosphorus, energy source

or as final electron acceptor. Sometime, a physiologically useful primary substrate (easy

to degrade) for example glucose induces the production of enzymes which can then

modify the molecular structure of another compound making it predisposed to microbial

enzymes (Luo et al., 2008). This process is called co-metabolic degradation and it takes

place in conditions when attack by the microbial enzymes is not strong enough to degrade

the pesticide. Microbial degradation therefore, presents a promising approach for the

removal of pesticide contaminants from the environment. Bacterial biochemical systems

transform the pesticides and modify their structure and toxicological properties leading to

complete conversion of the toxic chemical into inoffensive inorganic end products

(Racke et al., 1990; Singh, 2009).


18

Complete biodegradation eventually results in the mineralization of xenobiotic

compounds to CO2 and water. Many of the reactions involved in co-metabolism of

pesticides including oxidation-reduction, de-halogenation, ring-cleavage and hydrolysis,

occur simultaneously (Hazen, 1997). This transformation can lead to complete

detoxification, breakdown of products, which may be further attacked by other microbial

groups.

1.8 Microbial degradation of organophosphate pesticides

OP compounds contain the ester bond in the form of phospho (P=O) or

phophothio esters (P=S). The ester structure makes these compounds susceptible to

hydrolysis which is considered to be the major step in detoxification or degradation of the

OP pesticides as it removes the toxicity inducing leaving group (Figure 1.6). Further,

complete mineralization following the hydrolysis occurs through important reactions

such as alkylation, dealkylation, oxidation, reduction or ring cleavage (Singh et al.,

1999).

Figure 1.6 Hydrolysis of organophosphate pesticide by a bacterial phosphotriesterase

enzyme

Oxidation is usually uncommon in OPs (Lal, 1982). As described in the previous

section, enhanced biodegradation seems to be very significant in detoxifying the

repeatedly used pesticides without the introduction of degrading microbial communities

from other soils. This has been reported in many OP pesticides including isofenofos,

ethofos and feminofos. However, sometimes, soil micro-biota shows same degradation

tendencies towards different pesticides belonging to the same chemical family. This kind

P

O

R2(O,S)

R1(O,S)(S)

(O,S)

X

P

O

R2(O,S)

R1(O,S)(S)

OH

H(O,S)X

+

Bacterial

phosphotriesterase

Organophosphate pesticide


19

of trend is known as “cross adaptation” which is common in carbamates and other groups

of pesticides. Such pesticides are usually structural analogs of each other in terms of their

chemistry. However, within organophosphate pesticide group only limited trend of cross

adaptation has been observed (Racke and Coats, 1988) and more recently this trend has

been reported between chlorpyrifos and fenamifos (Singh et al., 2005). Further, cross

adaptation between organophosphates and pentachlorophenols is evident from a recent

study carried out by Fuentes et al., (2013) who reported that a single culture of

Streptomyces was capable of degrading chlorpyrifos and pentachlorophenol. This might

be due to the fact that characteristic microbial enzymes recognize the chemicals with

similar structure and bonds hence catalyze the degradation. The positive impact of the

cross adaptation of enhanced biodegradation is that microorganisms isolated for one

pesticide can be used for degrading or remediating the other pesticide for which no

microbial isolates are known. This aspect of cross adaptation has been extensively used

for organophosphate pesticides because mostly, two or more pesticides are applied

concurrently for crop protection which often leads to a mixed contamination of pesticide

residues in the soil environment. In such situations cross adaptation works very well and

helps in remediation of mixed pesticide residues.

A large number of bacterial species have been isolated till recent years which can

degrade these compounds in aqueous medium as well as in soils (Table 1.2). Degradation

of the OPs as a sole carbon source as well as co-metabolically is evident. Parathion (O,O-

diethyl-O-p-nitrophenyl phosphorothioate) is one of the widespread and most toxic

insecticides. The microbial degradation of parathion has received extensive attention

among the other OP pesticides. The first organophosphorus degrading bacterium,

Flavobacterium sp. that could degrade parathion and diazinon was isolated and reported

by Sethunathan and Yoshida, (1973). Further, Siddaramappa et al., (1973) isolated a

Pseudomonas sp. capable of hydrolyzing parathion as well as its hydrolysis product, p-

nitrophenol, as a carbon and nitrogen source. Mineralization of parathion has been

reported where it has been used as a source of carbon (Rani and Lalithakumari, 1994) or

source of phosphorus (Rosenberg and Alexander, 1979). Moreover, a pathway of

parathion degradation was also studies by Munnecke and Hsieh, (1976).


20

Hydrolysis of OP pesticides significantly reduces their toxicity towards

mammalian organisms, but sometimes hydrolysis products are more toxic compared to

the parent compound hence their degradation is relatively more important. However,

mostly the detoxification is the major concern of pesticide degradation; therefore further

metabolism of the degradation products is rarely studied. This can be explained by the

microbial degradation of chlorpyrifos which is one of the most extensively used OP

pesticides. Initially, Racke et al., (1990) reported that chlorpyrifos was resistant to the

phenomenon of enhanced degradation. It was suggested that production and

accumulation of first toxic metabolite, 3,5,6-trichloro-2-pyridinol (TCP), which has

antimicrobial properties, inhibits the proliferation of CP degrading microbes thereby

resisting the enhanced degradation of the pesticide (Racke et al., 1990).Toxicity of TCP

can be attributed to the existence of three chloride residues on the aromatic ring which

render it antimicrobial becoming more toxic and more persistent to microbial

degradation.


21

Table 1.2: Microorganisms isolated for the biodegradation of organophosphate

pesticides [a modification of table described by Singh and Walker, (2006)]

Organophosphate

pesticides

Microorganisms

References

Chlorpyrifos

Fungi

Phanerochaete chrysosporium

Hypholoma fasciculare

Coriolus versicolor

Stereum hirsutum

Trichoderma harzianum

Penicillium brevicompactum

Bacteria

Enterobacter sp.

Flavobacterium sp. ATCC27551

Bacillus pumilus C2A1

Stenotrophomonas maltophilia

Pseudomonas putida NII 1117,

Klebsiella sp., NII 1118,

Pseudomonas stutzeri NII 1119,

Pseudomonas aeruginosa NII 1120

Gordonia sp JAAS1

Sphingobacterium sp. JAS3

Bumpus et al., (1993)

Bending et al., (2002)

Omar, (1998)

Singh et al., (2004)

Mallick et al., (1999)

Anwar et al., (2009)

Dubey and Fulekar, (2012)

Sasikala et al., (2012)

Abraham et al., (2013)

Abraham and Silambrasam, (2013)

Parathion

Flavobacterium sp. ATCC27551

Pseudomonas diminuta

Arthrobacter spp.

Bacillus spp.

Agrobacterium radiobacter

Xanthomonas sp.

Suthantthan and Yoshida, (1973)

Serdar et al., (1982)

Nelson et al., (1982)

Horne et al., (2002)

Rosenberg and Alexander, (1979)


22

Continued from previous page

Table 1.2: Microorganisms isolated for the biodegradation of organophosphate pesticides [a

modification of table described by Singh and Walker, (2006)]

Organophosphate

pesticides

Microorganisms

References

Methyl parathion

Pseudomonas sp.

Bacillus sp.

Plesiomonas sp. M6

Pseudomonas putida

Pseudomonas sp. A3

Pseudomonas sp. WBC-3

Flavobacterium balustinum

Chaudry et al., (1988)

Sharmila et al., (1989)

Zhongli et al., (2001)

Rani and Lalitha-kumari, (1994)

Zhongli et al., (2002)

Chen et al., (2002)

Somara and Siddavattam, (1995)

Coumaphos

Nocardiodes simplex NRRL B24074

Agrobacterium radiobacter P230

Pseudomonas monteilli

Flavobacterium sp.

Mulbry, (2000)



Adhya et al., (1981)

Triazophos Bacillus Tap-1

Klebsiella sp. E6

Diaphorobacter sp.TPD-1

Diaphorobacter sp. GS-1

Ochrobactrum sp. mp-4

Roseomonas rhizosphaerae

Tang and You, (2012)

Wang et al., (2005)

Yang et al., (2011)

Liang et al., 2011

Dai et al., 2005

Chen et al., 2014

Profenofos Pseudomonas putida and

Burkholderia gladioli

Pseudomonas aeruginosa OW

Malghani et al., (2009a)

Malghani et al., (2009b)


23

In recent years, another aspect of microbial degradation was highlighted by the

use of mixed microbial communities for the remediation of the soils contaminated with

chlorpyrifos. A variety of chlorpyrifos degrading microorganisms isolated from diverse

environments (including highly contaminated sites) have been used as the mixed cultures

(Chishti and Arshad, 2013: Bhagobaty et al., 2007).

Another organophosphate pesticide, monocrotophos has many environmental and

health concerns associated with its high solubility in water and potential toxicity towards

aquatic organisms which in turn makes it important to investigate its detoxification.

Several bacterial strains have been isolated from monocrotophos treated soils. These

include Clavibacter michiganense sp. and Pseudomonas aeruginosa (Singh and Singh,

2003) which were isolated from soil and could utilize monocrotophos as a phosphorus

source. Also few fungal isolates were found to degrade monocrotophos such as

Aspergillus oryzae (Bhalerao and Puranik, 2009). However, it could not utilize

monocrotophos as a carbon source). Further, a Pseudomonas mendocina was reported as

the most capable monocrotophos degrader with the plasmid based degrading capability

(Bhadbhade et al., 2002).

Fenitrothion, a broadly used insecticide (Hayatsu et al., 2000) was found to be

degraded by Burkholderia sp. strain NF100 as a source of carbon with the involvement of

two plasmids. The first plasmid, pNF2, was found to be involved in the hydrolysis of

fenitrothion to a metabolite, 3-methyl- 4-nitrophenol with subsequent removal of the nitro

group to form methyl hydroquinone. Second plasmid, pNF2, further metabolized the

hydroquinone (Hayatsu et al., 2000).

Biodegradation of diazinon has also been given attention and various bacterial

isolates were reported for its remediation. Initially, two strains of Arthrobacter sp. and

two Pseudomonas spp were isolated which utilized diazinon as a source of carbon and

energy (Rosenberg & Alexander, 1979). However, recently, more reports are available

which throw light on diazinon degrading microorganisms (Cycon et al., 2009; Cycon et

al., 2013). Dimethoate degradation has been evidenced by a plasmid based gene from a

P. aeruginosa strain (Deshpande et al., 2001). A novel dimethoate degrading enzyme was

purified and characterized from a fungal strain, Aspergillus niger (Liu et al., 2001). This


24

enzyme could hydrolyze almost all of the compounds with phosphorothio linkage e.g.

fermothion and malathion but not the compounds with the phosphate linkage.

Ethoprophos and cadusafos are also two of the toxic OP pesticides. A

Pseudomonas putida strain was isolated with the potential to utilize ethoprophos as a sole

source of carbon (Karpouzas and Walker, (2000). Moreover, a Sphingomonas

paucimobilis and Flavobacterium sp. have also been reported by Karpouzas et al., (2005)

which can metabolize cadusafos. Similarly, several species of bacteria were isolated from

different environments which degrade OP pesticides in laboratory cultures and in soils

(Singh et al., 1999).

The metabolic regulation of microorganisms depends very strongly on effects of

organophosphate insecticides or pesticide left on particular organisms. For example, a

parathion degrading bacterium utilized parathion as a source of carbon and hydrolyzed it

to the diethyl phosphorothionate product but it could not metabolize it further, even in

the medium free of sulfur or phosphorus. Similarly, a variety of isolates have been

recognized that could use a pesticide as a sole source of phosphorus but not as a source

of carbon (Rosenberg & Alexander, 1979). A bacterial consortium was isolated that

degraded and used diethylthiophosphoric acid as a carbon source but failed to utilize it

as a source of sulfur or phosphorus. A possible explanation of difference in bacterial

metabolic behavior was provided earlier by Kertesz et al., (1994) who suggested that the

microbial enrichment conditions can potentially be crucial in the strain selection not

only with the most wanted degradative enzyme systems but also with specific regulation

mechanisms for the biodegradation pathways.

1.9 Factors affecting biodegradation of pesticides

Microbial degradation has been distinguished as an efficient strategy for

degradation of pesticides in soil which forms the basis for different strategies of

bioremediation and bioaugmentation. Consequently, conditions which support microbial

activity and growth in soil generally enhance the metabolic degradation of pesticides

(Gavrilescu, 2005). These environmental conditions are very crucial in the proliferation

and survival of microorganisms and also affect the chemical stability of the pesticide. In

soil, pesticide metabolism is influenced by abiotic and biotic factors which work in

succession and collaboration with one another. Completion of any bioremediation


25

process depends on several environmental factors such as chemical and physical

characteristics of the pesticide or substrate (hydrophilicity and solubility), availability of

nutrients, pH, temperature and biotic factors such as inoculum density of the degrading

microbiota (Karpouzas and Walker, 2000). It was suggested that the characteristics,

number of microbial organisms and the metabolic activity are the determining factors

regarding the biodegradation feasibility for pesticide remediation (Racke et al., 1996).

Effect of different environmental conditions on bioremediation of fenamiphos and

chlorpyrifos in soil and water to study the bioremediation was reported earlier (Singh et

al., 2006).

The rate of degradation of different pesticides depends largely on the soil pH.

This is because microbial enzymes are largely affected by the pH and their activity is

reduced or increased as a function of change in pH. Moreover, pH is also important in

maintaining the stability of the chemical substances (pesticides in this case). An

increased degradation of atrazine was observed by a Pseudomonas sp. in the soil

containing higher organic matter and low pH (Kontchou and Gschwind, 1995) whereas

high soil pH was reported to have enhancing effect on chlorpyrifos degradation.

Pesticide concentration has also been reported to affect bioremediation process.

For any xenobiotic compound, the concentration serves as a limiting factor and above a

certain level of concentration, the process of remediation becomes indispensable and this

level is regarded as “remediation trigger level.” When pesticide is applied at normal

agricultural rates, almost 99% of applied pesticide may be removed over the course of a

growing season. Unfortunately, even when present in soil at very low levels, many

recalcitrant pesticides often spread around and reach water resources because they are

not degradable by the microbial activities.

The soil temperature is one of the most significant factors which have a profound

effect on microbial activity and consequently affect the overall process of microbial

degradation of pesticides. Generally, at very low or very high soil temperature, the

efficiency of microbial degradation of soil contaminants is reduced. For an effective

bioremediation process an optimum temperature is required which is toleratable by the

degrading microbiota. This is again due to the dependency of the bacterial enzymes on

the temperature which require a suitable temperature to work properly.


26

Several researchers have demonstrated that the degradation of pesticides in soil

can be boosted by inoculation with appropriate population of the microorganisms.

Comeau et al., (1993) suggested that inoculum level of 106

to108

CFU/g of soil was

suitable for the pesticide removal in a contaminated site. However in another study it

was discovered that a lower inoculum size of an Agrobacterium sp. was adequate to

rapidly degrade atrazine (Struthers et al., 1998).

Bioremediation techniques attempt to increase the rate of naturally occurring

biodegradation processes by optimizing the degradation conditions (Semple et al., 2001).

Without suitable environmental conditions in a contaminated area, process of

bioremediation can be inhibited even if microbial populations are available for

biodegradation of a particular contaminant. In such cases, addition of nutrients manually

to the contaminated places, a process known as biostimulation or the addition of more

microbes (bioaugmentation) have been found effective in increasing the bioremediation

(Sasek, 2003; Mrozik et al., 2010). Although biostimulation may have poor

reproducibility and can be dependent on the characteristics of microbial populations but

still it is an efficient bioremediation strategy as it replaces the shortage of carbon and

nitrogen, O2, acid or bases for pH optimization, and sometimes water or specific

substrates to induce specific enzymes (Margesin et al., 2000). Bioaugmentation is

another efficient option. which introduces the exotic bacterial species to the site in

question. But a major advantage of bioaugmentation is that a full knowledge of process

and its conditions is mandatory as the different bacterial species are to be introduced. In

this case, the success of bioremediation is mainly dependent on the microbial

proliferation capability, competition of the introduced species and the bioavailability of

the pesticide (Gavrilescue, 2005). Bioavailability here means the acquirement of a

compound and its degradation to harmless products.

Sometimes microorganisms degrade an easily available carbon source and acquire

energy and enzyme induction for the degradation of xenobiotic compound. This kind of

metabolism is known as co-metabolism. Usually, biodegradation and co-metabolic

degradation occur at a time in soil. Bioremediation has been emerged as cheaper and

effective piece of technology in the recent era compared to parallel chemical and physical


27

methods of remediation. Moreover, this strategy has a greater potential to cope with the

contaminated groundwater and soil ecosystems.

1.10 Organophosphate degrading enzymes

Microorganisms have been equipped with the extensive enzyme systems that can

catalyze the degradation of toxic OP pesticides. Most important of all OP degrading

enzymes are the hydrolases which are also called phosphotriesterases or esterases. These

enzymes can be constitutive or inducible and catalyze the hydrolysis of OPs during

detoxification. Very first time reported enzyme catalyzing OP degradation was

organophosphate hydrolase (OPH) that was isolated from Flavobacterium sp. and

Pseudomonas strains (Serdar et al., 1982; Somara and Siddavattam, 1995). After this

discovery, OP hydrolases and genes encoding these OP hydrolases were reported from

different bacterial or fungal genera. Mostly the OP degrading genes including opd and

mpd have been reported to be plasmid borne which encode organophosphate degrading

hydrolase (OPH) and methyl parathion enzyme hydrolase (MPH) respectively (Singh and

Walker, 2006). However, another enzyme organophosphorus acid anhydrolase (OPAA)

was identified in Agrobacterium radiobacter (Horne et al., 2002) and Alteromonas sp.

(Cheng et al., 1996). OPAA is encoded by opdA gene and has been found to have

chromosomal origin. From time to time various genes have been isolated and reported for

degradation of OPs in various organisms such as Ochrobactrum, Pseudaminobacter and

Burkholderia (Zhang et al., 2005; Goda et al., 2010). However, a very less extent of

similarity is present among the genes. There is a considerable need for further

investigation of OP degrading genes and enzymes. Few other OP degrading enzymes

include dehydrogenases, mono-oxygenases, dioxygenases.

1.11 Bioremediation of organophosphate contaminated soil

As mentioned in previous sections, excessive use of pesticides, their

accumulation and persistence in soil leads to the disastrous results every year (Singh and

Walker, 2006). Although OPs are easily biodegradable and less persistent than

organochlorines but the problem is associated with the quick biodegradation of the OP

pesticides in the soil to the degradation metabolites which are usually more toxic and

persistent than the parent compound hence ultimately their residues persist in the soil. In


28

North Carolina chlorpyrifos residues were observed in 16 houses even long time after its

applications for insect control (Wright et al., 1994). Higher levels of chlorpyrifos and

TCP have been found in many soil samples by many researchers. In Pakistan, many

reports emphasis on the detection of pesticide residues in soil (Anwar et al., 2012)

Bioremediation has been proved to be a safer and gainful technology for the

treatment of pesticide contaminated soils as it converts the toxic pesticides into

undisruptive inorganic products (Cappello et al., 2007; Yang et al., 2005; Nawaz et al.,

2011). Numerous reports of bioremediation of CP contaminated or CP spiked soils are

available to date. A chlorpyrifos degrading Enterobacter strain B-14 was introduced by

Singh et al., 2004 into a CP contaminated soil containing a small indigenous population

of CP degrading bacteria. The strain B-14 efficiently remediated the contaminated soil.

Similarly, a Sphingomonas sp. has been reported to degrade about 98% chlorpyrifos in

10 days in CP spiked soil (Li et al., 2007). Profenofos has been reported as less

persistent in soil. Environmental fate studies of profenofos show that it is not very

mobile in soil and the route of degradation is hydrolysis that needs microbial processes.

However, first degradation metabolite, 4-bromo-2-chlorophenol is more persistent and

toxic than profenofos itself hence its remediation needs attention.

1.12 Plant microbe interaction for the remediation of pesticides/pollutants

Phytoremediation is a low-cost and economical technology that employs plants for the

cleanup of the environment contaminated with the pollutants (Pilon-Smits, 2005; Wang

et al., 2008). However, one of the few limitations of the phytoremediation is the

sensitivity of the plants to pollutants which adversely affect the plants in terms of growth

and physiology hence pollutant tolerant plants are also found to exhibit reduced growth

which consequently results in reduced effectiveness of pollutant remediation by the plant

(Gaskin et al., 2008; Weyens et al., 2009). Impaired germination and growth of rice

seeds and seedlings was observed as a result of imdacloprid applications (Stevens et al.,

2008). Moreover, alteration in the nitrogen metabolism and growth of Vigna radiata (L.)

in response to chlorpyrifos has also been documented (Parween et al., 2011). This

limitation has been compensated by the microbially-assisted phyto-remediation which

involves the use of plants and microbes together for the toxic pesticide remediation.

Plant-microbe interaction for pollutant remediation helps both the partners. Plant root


29

exudates stimulate microbial growth and enhance the pollutant degradation properties of

the microbes. However, Plants might get profit from bacteria (and other microbes)

because plant associated rhizospheric and endophytic bacteria have been reported to

degrade the toxic pollutants in the contaminated soil and potentially improve the

phytoremediation (Afzal et al., 2012). Moreover, rhizobacteria and endophytic bacteria

have been reported to increase plant growth by different beneficial biochemical

processes such as mineral solubilization, phytohormone production and nitrogen fixation

thus alleviating the stress induced by the pollutant on the plant (Glick, 2010). Endophytic

bacteria reduce phytotoxicity by producing pollutant degrading enzymes inside plant

tissues thereby increasing the plant resistance to contaminants and ultimately improving

the plant growth (Tang et a.,l 2010; Fernández et a.l 2011).

The mechanism of plant interaction with microbes is given in Figure 1.7. The inoculation

of beneficial microbes to plants offers a feasible and inexpensive alternative technology

to clean up pesticide contaminated sites. However, little is known concerning the

potential of plant growth promoting rhizo and endophytic bacteria in the

phytoremediation of pesticides.

Figure 1.7 Plant-microbe interactions for remediation of pesticide contaminated soil


30

1.13 Objectives of the study

Profenofos (PFF) and chlorpyrifos (CP) are among the commonly used OP

insecticides. They are biodegradable in soil but their degradation metabolites, being

aromatic in nature, are highly toxic and persistent. However, bacterial strains capable of

degrading these toxic metabolites are scarce. Therefore, the present study was designed to

isolate and characterize bacterial strains capable of complete degradation of PFF and CP

as well as their metabolite, 3,5,6 trichloro-2-pyridinol (TCP) and 4-bromo-2-

chlorophenol (BCP) respectively.

A well optimized bioremediation of pesticides is the basis of successful

remediation of the particular pesticide. Hence, study was planned to optimize culture

conditions of isolated bacterial strains that govern CP and PFF degradation, the kinetics

of CP and PFF biodegradation, accumulation and utilization of their toxic metabolites

and the governing constants thereafter. As, these parameters vary depending on bacterial

strains and concentration/nature of pollutant, a clear understanding of the biodegradation

kinetics of pesticides would determine suitability of the bacterial strain for in situ

bioremediation.

A little is known about the biochemical pathways of the two pesticides. It is

important to investigate, what metabolites are produced by the degradation of these

pesticides and whether they are further degraded or accumulate in the culture medium?

To answer these questions, biodegradation products of CP and PFF were studied and

identified by GC-MS analyses. Moreover, a trial was made to track genes involved in the

degradation of the two pesticides.

Chapter2 Materials and Method

31

Chapter 2

Materials and Methods

1.14 Chemicals

Analytical grade chlorpyrifos (CP, 98.6%) was obtained from Dr Ehren Stofer

GmbH (Germany). 3,5,6-trichloro-2-pyridinol (TCP), the major hydrolysis product of

CP, analytical grade profenofos (PFF, 95%) and its hydrolysis metabolite 4-bromo-2-

chlorophenol (BCP) were purchased from Chem Service (West Chester, Pennsylvania,

USA). Technical grade CP (95%) and PFF (92%) were purchased from Pak China

Chemicals Lahore. Dichloromethane (DCM), n-hexane, methanol, acetone and

acetonitrile (HPLC grade) were procured from Merk (Frankfurter, Germany). N, O-Bis

(trimethylsilyl) trifluoroacetamide (BSTFA) kit was purchased from Supelco, Bellefonte,

Pennsylvania, USA. All other chemicals (used in preparation of growth and minimal

media) were purchased from Merk, Sigma or Eldrich. Reagents used in molecular

techniques such as restriction enzymes, PCR and cloning reagents were purchased from

Fermentas.

1.15 Bacterial strains used in the study

Profenofos and chlorpyrifos degrading bacterial strains used in the study were

isolated indigenously (Table 2.1). Escherichia coli (E. coli) DH5 carrying pk18 vector

and Pseudomonas diminuta were kindly provided by Dr. Rebecca Parales, Department of

Microbiology, University of California, Davis, California, United States of America.

Escherichia coli DH5α carrying a broad host range ampicillin resistant (AmpR) plasmid

pBBRIMCS-4 harboring yfp gene was obtained from NBRC, NIBGE, Faisalabad,

Pakistan.


32

Table 2.1: Bacterial strains used in the study

1.16 Soil collection

Soil samples were collected from fields of Ayub Agricultural Research Institute

(AARI) and NIBGE, Faisalabad, Pakistan. The soil samples had a long history of

organophosphate (OP) insecticides applications. Hence extensive use of the pesticide

made this soil vital for the isolation of selective pesticide degrading microbes.

1.17 Growth media

Growth media used in the study were Luria Bertani (LB), Minimal Salt Medium

(MSM), Focht solution, Pikovskaya medium, Yeast Extract Mannitol (YEM), Nitrogen

free medium, and Chloride free medium. Compositions of all growth media are given in

appendices.

2.4.1 Maintenance and preservation of the bacterial strains

Profenofos and chlorpyrifos degrading bacterial strains/consortia were maintained in

MSM supplemented with respective pesticide. Glycerol stocks (50% v/v) were prepared

by aseptically mixing bacterial cultures grown in LB broth and the 50% glycerol. The

stocks were preserved at -80°C for months. PFF degrading bacterial isolates were

maintained separately as well as in mixed culture in glycerol stocks.

Name of the strain Source Origin

Mesorhizobium sp. HN3 OP contaminated soil Pakistan, Isolated in the current study

Achromobacter xylosoxidans PF1 OP contaminated soil Pakistan, Isolated in the current study

Pseudomonas aeruginosa PF2 OP contaminated soil Pakistan, Isolated in the current study

Bacillus sp. PF3 OP contaminated soil Pakistan, Isolated in the current study

Citrobacter koseri PF4 OP contaminated soil Pakistan, Isolated in the current study


33

2.5 Equipment used in the study

Most of the facilities and equipments used were available at NIBGE. These

included High Performance Liquid Chromatograph (HPLC) Varian Pro Star 325, UV VIS

detector; Spectrophotometer; Eppendorf centrifuge machines (models 5424 and 5816);

Biorad Thermocycler; Stereomicroscope; Rotary shaker; Light microscope; Olympus

Confocal Laser Scanning Microscope (CLSM) and Scanning Electron Microscope

(SEM). Agilent Gas Chromatograph Mass Spectrometry was availed at UC Davis, USA.

DNA sequencing was done using commercial sequencing facility of Macrogen, South

Korea, until otherwise mentioned.

2.6 Enrichment of profenofos and chlorpyrifos degrading bacterial strains

Enrichment culture technique was used to isolate pesticide degrading bacterial

strains from soil using MSM as enrichment medium. For this purpose, soil (10g) was

added to 250 ml Erlenmeyer flasks containing 100 ml sterile MSM (in duplicate)

supplemented with 50 and 100 mg/l (individually) of respective pesticide

(profenofos/chlorpyrifos). These flasks were incubated in a rotary shaker at 37°C and 100

rpm. Two weeks following the incubation, 5 ml suspension from each replicate was

transferred to flasks containing fresh MSM supplemented with 50 & 100 mg/l of the

respective pesticide. Similarly, four transfers were carried out by sub culturing 5 ml

inoculum into fresh MSM containing respective pesticide each time. In this way

enrichment cultures containing diverse number of pesticide degrading bacteria were

developed.

2.7 Isolation of pesticide degrading bacterial strains

The enrichment cultures were used for isolations of pure, pesticide degrading

bacterial strains. Two weeks following the last transfer, the enrichment cultures were

serially diluted (10-1

to 10-7

) in 0.9% saline solution. Selected dilutions 10-5

, 10-6

and 10-7

of each culture were used to spread (100 µL) in triplicates on solid LB agar plates

containing 100 mg/l respective pesticide. Following two days of incubation,

morphologically distinct bacterial colonies were separated and purified by repeated

streaking on LB agar plates.


34

Once all the isolates were purified by repetitive sub culturing, their degradation potential

was investigated in terms of their growth on MSM agar plates containing respective

pesticide as the source of carbon and energy. Moreover, the pesticide degradation

efficiency of the isolates was also tested in MSM broth containing respective pesticide as

the only source of carbon (and incubated at 37°C). Turbidometric method described by

Jyothi et al., (2012) was employed for monitoring the growth of the degrading bacterial

cultures in MSM broth containing respective pesticide as a carbon source. Cell dry mass

was determined for the bacterial cultures having an OD600nm of 1.0 and was used as

standard for calculating cell dry mass of all the culture samples.

Liquid cultures were harvested after five days of incubation and the pesticide

residues were extracted using dichloromethane and analyzed by quantifying the residual

concentrations of respective pesticide on HPLC. Best degrading bacterial isolates were

characterized and used for further degradation studies.

2.8 Molecular characterization of bacterial isolates

2.8.1 DNA Isolation

Genomic DNA was extracted from bacterial isolates by CTAB (Cetyl Tri-methyl

Ammonium Bromide) Method (Mateen, 1998) as follow:

Bacterial cells were grown in MSM containing glucose at 37C overnight and harvested

by centrifugation at 8000 rpm.

Cells were re-suspended in 5 ml T.E buffer and 20 mg lysozyme was added to this

suspension.

Suspended cells were incubated at 37C for 5 min.

500 µl of 10% SDS, 25 µl proteinase K (25 mg/ml) and 3 µl RNAase were added to the

incubated suspension.

The contents were mixed thoroughly and incubated at 37C for 10 min.

After incubation, 0.9 ml 5M NaCl and 0.75 ml NaCl/CTAB were added, mixed thoroughly

and incubated at 65C for 20 min.

The protein contents were extracted (twice) with an equal volume of Phenol-

chloroform-isoamyl alcohol (25:24:1) and centrifuged for 10 min at 8000 rpm and 4C to

separate the organic and aqueous phase (containing DNA).


35

Aqueous supernatant was separated in fresh falcon tube and 0.6 volume of isopropanol

was added and mixed thoroughly until a white DNA pellet precipitated out of solution.

The solution containing DNA was stored at -20°C overnight.

The solution containing DNA was centrifuged at 8000 rpm for 5 min and supernatant

was discarded.

The DNA pellet was washed with 70% ethanol (twice) and dried at room temperature.

Dried DNA pellet was re-suspended in 100 µl T.E and saved at -20°C until used.

2.8.2 Preparation of Heat shock competent cells (C-cells)

E. coli cells were grown overnight in 5 ml LB at 37°C with constant shaking.

0.25 ml of the overnight grown culture was inoculated into 50 ml LB and grown at 37°C

with constant shaking until the culture reached an OD600nm of approximately 0.4.

The culture was poured into cold Oakridge tubes and centrifuged for 5 min at 4,500

rpm.

The supernatant was discarded removing as much liquid as possible.

GENTLY each cell pellet was re-suspended in 10 ml COLD Frozen Storage Buffer (FSB,

Appendix 4), combined them into one tube and incubated on ice for 15 to 60 min, and

pelleted by centrifugation at 45,00 rpm for 5 min.

Cell pellet was gently re-suspended in 4 ml of cold FSB.

Pipetted 200 ul aliquots into COLD 75 12 mm polypropylene snap cap tubes and

stored the competent cells at –70˚C.

2.8.3 Amplification of 16S rRNA gene from bacterial isolates

16S rRNA gene of all bacterial isolates was amplified using forward primer FD1,

5-AGAGTTTGATCCTGGCTCAG-3 (E. coli bases 8–27) and reverse primer RP1, 5-

ACGGHTACC TTGTTACGACTT-3 (E. coli bases 1507–1492) (Wilson et al., 1990).

PCR reactions were performed in 50 µl reaction volumes containing 1 µl of Taq DNA

polymerase (2.5 U/µl), 5 µl of 10×PCR reaction buffer, 2 µl of each of the primers (10

µM), 2 µl dNTPs mixture (10 mM) and 38 µl of sterile distilled water. 2 µl of each

individual bacterial strain was used as template. Thermal cycler (Biorad) was used for

amplification from DNA which was programmed as: pre-heat treatment at 94C for 1 min


36

followed by annealing at 52C for 1 min, and extension at 72C for 1.5 min and all the

three steps were repeated for 30 cycles.

2.8.4 Agarose gel electrophoresis

The amplified PCR products were analysed on 1 % agarose gel stained with

ethidium bromide (100 μg/ml). Amplified PCR products were mixed with 6X loading

dye (bromophenol blue) and loaded on the agraose gel immersed in 0.5X TAE buffer.

Samples were electrophoresed at 60-100 volts for about 1 to 1.5 h followed by the

examination of gel under UV transilluminator for analyzing the PCR products.

2.8.5 Ligation and cloning of the 16S rRNA gene

The 16S rRNA gene amplicons were ligated into TA cloning vector (PCR Product

cloning Kit, Fermentas) and the ligation mixtures were kept at 16°C overnight. The

following day, ligation mixtures were transformed into E. coli TOP10 competent cells by

heat shock methodology. Plasmids were isolated from E. coli using mini prep kit

(Fermentas) according to the manufacturer’s instructions and restricted with EcoRI

(restriction enzyme, Fermentas) to confirm the product size of the insert.

2.8.6 Sequencing of 16S rRNA gene and bacterial identification

The inserts were sequenced by using M-13 primers. The 16S rRNA gene

sequences were compared to the already known nucleotide sequences using BLAST

search algorithm (http://www.ncbi.nlm.nih.gov/BLAST) and molecular evolutionary

relationships were accomplished using MEGA 5.0 version with the Kimura two-

parameter model and the neighbor-joining algorithm (Saitou and Nei, 1987). The

sequences of the isolates obtained were submitted in GenBank.

2.9 Morphological and biochemical characteristics of the bacterial isolates

2.9.1 Morphological characterization

Bacterial isolates were examined for colony morphology (size, margins and

surface texture), cell morphology, pigmentation and motility as per the standard

procedures given by Barthalomew and Mittewer, (1950).

http://www.ncbi.nlm.nih.gov/BLAST


37

2.9.2 Physiology and Biochemical characterization

A series of studies were conducted to find out the biochemical and physiological

characteristics of isolated bacterial strains. These characteristics include utilization of

different carbon sources (glucose, sucrose), enzymatic properties (oxidase, arginine).

Biochemical and physiological tests of the isolates were carried out using QTS-24

(Quick Test Strip) kit developed by Defense Science and Technology Organization

Laboratories (DESTO), Karachi, Pakistan. According to the manufacturer instructions,

the kit is based on dehydrated substrates contained in microcupules for different

enzymatic and assimilation reactions. Further, gram staining and antibiotic assays were

performed as described in the following sections:

2.9.2.1 Gram staining

The compositions of all the solutions used in Gram’s staining (the crystal violet

solution, iodine solution and safranin solution) are described in Appendix 5.

Following protocol was adapted for gram staining of bacterial isolates:

Bacterial strains were grown on LB agar and incubated overnight at 37C.

A single colony of each freshly grown culture (on LB agar plates) was picked with a

sterile loop and mixed in a drop of saline (0.9%) on a glass slide to make a thin

smear.

The slide was air dried, heat fixed and stained with primary stain, the crystal violet

solution for one min.

The smear was washed with distilled water and flooded with iodine solution for one

min.

Iodine solution was washed off with water and de-colorized with ethanol (70%) for

one and a half minute.

Again slide was washed with distilled water and stained with secondary stain,

safranin solution for one min.

Finally the slide was washed with distilled water, air dried and observed under light

microscope.


38

2.9.2.2 Antibiotic resistance of isolates

For this purpose, 1-2 ml of a freshly grown bacterial cultures (in LB) were spread

(100 ml) aseptically onto the antibiotic sensitivity sulphonamide agar (ASS; Merk,

Germany) plates until the bacterial cultures were completely absorbed to the agar.

The intrinsic antibiotic resistance pattern of pesticide degrading bacterial isolates

was determined by disc diffusion method (Valverde et al., 2005) using ready to use

antibiotic discs (Bioanalyse, Turkey). The antibiotics used are given in Table 2.2.

Table 2.2 Antibiotics used in the study

No. Antibiotic Abbreviation Disk concentration

1 Chloramphenicol C 30µg

2 Ampicilin AM 10µg

3 Rifampicin RA 5µg

4 Streptomycin S 10µg

5 Carbenicillin PY 100µg

6 Erythromycin E 15µg

7 Gentamycin CN 10µg

8 Kanamycin K 30µg

9 Nalidixic Acid NA 30µg

10 Tetracycline TE 1.25µg

2.10 Inoculum preparation

The chlorpyrifos and profenofos degrading bacterial isolates were grown

aerobically in LB medium containing 100 mg/l respective pesticide in Erlenmeyer flasks

with constant shaking in a rotary shaker at 37°C. The overnight grown bacterial culture

was harvested centrifuged at 4600×g for 10 to 15 min and used for inoculum preparation

following Anwar et al., (2009). The cell pellet was washed with 0.9% saline solution

(Appendix 6) and re-suspended in the same to get an OD600nm of 0.8. Colony forming

units (CFU/ml) were determined by dilution plate count technique. This suspension (2%

v/v) was used as inoculum in biodegradation experiments until otherwise described.


39

2.11 Experimental set up for pesticide degradation studies

Pesticide degradation studies were carried out in 250 ml Erlenmeyer flasks

containing 50 ml MSM supplemented with 100 mg/l respective pesticide and 2%

inoculum of respective pesticide degrading bacterium under various culture conditions as

described in respective sections. The flasks were incubated at 37°C (or at other

temperatures as per the requirement of experiment) and 100 rpm in rotary shaker for 10

days. For all the treatments, un-inoculated flasks served as controls and all the

experiments were performed in triplicates. Samples were periodically harvested for

analyzing the growth rates and residues of pesticide and its metabolites.

2.11.1 Determination of the detection wavelength of the pesticides

For a good chromatographic detection of PFF and CP (and others pesticides used

in Chapter 3), the wavelength, at which they display maximum absorption of UV

radiation, was determined spectrophotometrically. The optimum wavelength called

“λmax” was obtained by recording the UV absorption spectra of all the pesticides used in

this study. Moreover, for the confirmation, the λmax was evaluated using HPLC and

comparing the absorbance of the pesticides at about three different wavelengths.

2.11.2 Extraction of pesticide residues from liquid cultures

Pesticide residues were extracted using equal volumes of water immiscible solvent

(mostly dichloromethane, DCM). Culture samples (20 ml) were recovered from flasks after every

24 h and centrifuged at 4600×g for 10 min to obtain cell free medium. Pesticide residues were

extracted from supernatant using equal volume of dichloromethane (DCM) twice. Organic layer

of DCM was aspirated, pooled and evaporated using rotavapor at 37°C. The dried residues were

dissolved in HPLC grade acetonitrile (1 ml), and filtered through flouropore TM filter membrane

(0.45 μm FH) to remove any particles. Samples were finally diluted (if required) and analyzed by

High Performance Liquid Chromatography (HPLC).

2.11.3 HPLC conditions for pesticide residual analyses

Quantitative analyses of pesticides were carried out by High Performance Liquid

Chromatography (HPLC) equipped with ODS2 C18 reversed-phase column. The gradient


40

mobile phase consisting of acetonitrile:water: acetic acid (20:80 to 80:20, Appendix 7) was

programmed at the flow rate of l ml/min. Retention time for the pesticide and its

metabolites were deduced by running the respective standards solutions (Anwar et al.,

2009). Concentration of pesticides in the culture medium and that in the unioculated

control samples was calculated using unit method.

2.12 Soil microcosm studies (Pot Experiments)

2.12.1 Soil collection for microcosm experiments

For pesticide degradation studies in soil microcosm, soil was collected randomly

from 0-25cm depth from an area in Faisalabad, Pakistan where no pesticides had been

applied. The samples were pooled, brought to the laboratory in polyethylene bags and

kept in refrigerator at 4°C to maintain the biological activity of the soil microbes until

used.

2.12.2 Determination of Maximum Water Holding Capacity (MWHC) of the soil

Oven dried soil samples (100g) were taken in funnel in triplicates. The funnels

having short length rubber tubing at their mouths with a clamp were used for this

purpose. Cotton swabs of equal weight were placed in the funnel at the top of the stem.

Water was transferred into soil slowly until it was saturated. As soon as first drop of the

water escaped from the funnel, water addition was stopped and all extra water was

allowed to come out of the funnel. After about half an hour, volume of the water run

away through the soil in the beaker below was measured. This volume of water retained

by the soil was used for calculating MWHC of soil.

2.12.3 Preparation of pesticide-contaminated soil

In order to conduct microcosm experiments, pesticide stock solutions were used

for spiking the soil in their respective experiments according to the procedure described

by Brinch et al., (2002) and is described below:

The pesticide in the form of acetonitrile solution was added to sand (25% of the

total quantity of dry soil), mixed thoroughly and left closed for few hours. The solvent

was volatilized under fume-hood and the spiked soil was mixed with rest of experimental


41

non-contaminated soil. A metal spatula was used to mix sand and soil so as to mix the

pesticide homogeneously in the total soil and to obtain final required concentrations of

the pesticide for respective experiments. In this way heterogeneity of the soil is reduced

to maximum. The spiked soil was saved at room temperature.

2.12.4 Extraction and analysis of pesticide residues from soil

For pesticide extraction from soil, 5g soil was sampled from each pot of

experiment. The soil samples were transferred to a 20 ml glass tube. Acetonitrile (5 ml)

was added to each of the soil samples and the tubes were sealed. The sealed tubes were

vortexed for 3 to 5 min and placed in a sonication bath for 10 min. The tubes containing

the extracts and soil materials were centrifuged at 4600×g for about 15 min. The soil

pellet was discarded and the supernatant was separated containing the pesticide residues

from each sample was filtered through fluorropore ™ membrane filters (0.5 µm) and

stored at 4°C until analyzed through HPLC.

2.12.5 Optimization of soil moisture on pesticide degradation

The microcosm studies were carried out with soil contaminated with 50 mg/kg

pesticide in plastic pots. Triplicate soil samples (100 g) containing pesticide were

inoculated with respective pesticide degrading bacterial strains (2x109 CFU/g). Varying

soil moisture contents were adjusted and maintained at 20%, 40%, 60% and 80% of the

MWHC by adding sterile MSM. Un-inoculated soil samples served as controls. Pesticide

residues were extracted after every ten days and analysis was carried out for about 40

days.

2.12.6 Optimization of inoculum density for pesticide degradation in soil

Pesticide contaminated soil was inoculated with varying inoculum densities

prepared by a dilution series using overnight grown bacterial culture in nutrient broth.

Triplicate sterilized soil samples (100 g) at 40% MWHC of the soil, containing 50mg/kg

pesticide and were inoculated with pesticide degrading bacteria to achieve varying cell

densities (CFU/g) fresh weights and incubated at room temperature. Sampling and


42

extraction of pesticide residues were carried out after every ten days and the experiment

was carried out for 40 days.

2.13 Identification of pesticide metabolites

Samples containing residues of chlorpyrifos and profenofos along with their

degradation metabolites, periodically obtained from culture flasks were extracted with

solvent (as described in previous sections) and derivatized with N, O-Bis-(trimethylsilyl)

-trifluoroacetamide (BSTFA, derivatizing reagent for Gas Chromatography) using

BSTFA kit according to the protocol as instructed by manufacturer (Supelco).

GC-MS analyses were performed with an Agilent 6890N Gas Chromatograph

equipped with a Supleco Equity-1 capillary column (30 m by 250 µm and 25 µm film

thickness), an auto-injector (7683 series), and an Agilent 5973 network mass selective

detector (Agilent Technologies, Palo Alto, Calif.). Helium was used as the carrier gas

with a constant flow rate of 0.5 ml/min. The injector and transfer lines were 220 and

300°C respectively. The chromatography program was as follows: total run time was 33

min; Initial temperature of column was 70°C, a temperature increase of 10°C/min and

final heating to 240°C. The ionization voltage and electron multiplier settings were 70eV

and 1,294 V, respectively. Product identities were confirmed by a comparison of

retention times and MS fragmentation profiles to authentic chemical standards of the

respective pesticides.

2.14 Study of potential genes encoding hydrolases/oxygenases in pesticide degrading

bacterial strains

2.14.1 Amplification of the OP degrading genes

Primers were designed and synthesized to amplify the potential genes encoding

hydrolases, hydroxylases or oxygenases from PFF and CP degrading bacterial isolates.

Primers already reported for OP hydrolase genes (opd, mpd & opdA) and oxygenases of

different aromatic compounds were selected from previously published work as

mentioned in Table 2.3. Some primers were also designed (Table 2.4) by retrieving

known sequences of OP hydrolase or oxygenase genes which have already been

submitted in the GenBank. These sequences were aligned using ClustalW software,


43

conserved regions of the sequences were selected and tested in silico using the Primer3

program available online (http://frodo.wi.mit.edu/cgi-bin/primer3/primer3.www.cgi).

PCR reactions were performed in 25 µl reaction volumes containing 0.5 µl of Taq

DNA polymerase (2.5 U/µl), 2.5 µl of 10×PCR reaction buffer, 0.75 µl of each of the

primers (10 µM), 1 µl dNTPs mixture (10 mM), 1 µl of the bacterial genomic DNA

(as the template) and 18.5 µl of sterile distilled water. The PCR amplification

protocol was: denaturation at 94C for 1 min, annealing at 52-65C (depending on

melting temperatures of primer pairs) for 30 seconds and extension at 72C for 1 min,

and all the three steps were repeated for 35 cycles.

2.14.2 Agarose gel electrophoresis, cloning and sequencing of the amplified gene

The amplicons of expected size obtained were analyzed on agarose gel (1 to

1.5%) depending on the size of the PCR product. Same protocol was followed as

described in Section 2.8.4 for 16S rRNA gene.

The amplified PCR products were purified using PCR purification kit (Fermentas),

ligated to pk18 vector and transformed in E. coli DH5 following heat shock method.

Plasmids were isolated from transformants using mini prep kit (Fermentas). Isolated

plasmid was digested with required restriction enzyme to confirm the product sizes of the

amplicons. The plasmids thus obtained were sequenced and the sequences obtained were

analyzed using the BLASTN search program of the GenBank database in National Center

for Biotechnology Information, NCBI.

http://frodo.wi.mit.edu/cgi-bin/primer3/primer3.www.cgi


44

Table 2.3: Sequences of the previously reported primers used in this study

Primer Sequence

Product

size Reference

Gene/enzyme recognized

by the primer

OpdA-F1

OpdA-R1

5’GCACTGCAGATGCAAACGAGAAGAGATGCACTT-3

5- GTCGAATTCTCATCGTTCGGTATCTTGACGGGG-3’

1155bp Sharaf et al., 2006

OPDA enzyme, a natural

variant of OPH enzyme

OpdA-F2

OpdA-R2

5'GCGATGTTCCGGTAACCACTCACA3'

5'GCAACACTCTCAGAGGGACGAAGG3'

412bp Ali et al., 2011 OPDA enzyme, a natural

variant of OPH enzyme

Opd-F1

Opd-R1

5'GCAAGGGTTGTGCTCAAGTCTGC 3'

5'GACCAATAAACTGACGTCGCGAC 3'

327bp Ali et al., 2011 Organophosphate Hydrolase

(OPH)

Mpd-F1

Mpd-R1

5'GAAAAGCAGGTCGACGAGATCTAC3'

5'ACCTTTGACGACCGAGTAGTTCAC3'

547bp Ali et al., 2011

Methyl parathion hydrolase

(MPH)

NAH-F

NAH-R

5' CAAAA(A/G)CACCTGATT(C/T)ATGG 3'

5' (C/T)(A/G)CG(A/G)G(C/G)GACTTCTTTCAA 3'

377bp Baldwin et al., 2003

Naphthalene dioxygenase

TOD-F

TOD-R

5' ACCGATGA(A/G)GA(C/T)CTGTACC 3'

5' CTTCGGTC(A/C)AGTAGCTGGTG 3' 757bp Baldwin et al., 2003

Toluene dioxygenase

TOL-F

TOL-R

5' TGAGGCTGAAACTTTACGTAGA 3'

5' CTCACCTGGAGTTGCGTAC 3' 475bp Baldwin et al., 2003

Toluene monoxygenase

BPH1-F

BPH1-R

5' GGACGTGATGCTCGA(C/T)CGC 3'

5'TGTT(C/G)GG(C/T)ACGTT(A/C)AGGCCCAT 3'


Biophenyl dioxygenase

BPH2-F

BPH2-R

5' GACGCCCGCCCCTATATGGA 3'

5' AGCCGACGTTGCCAGGAAAAT3'


Biophenyl dioxygenase

PHE-F

PHE-R

5' GTGCTGAC(C/G)AA(C/T)CTG(C/T)TGTTC 3'

5' CGCCAGAACCA(C/T)TT(A/G)TC 3' 206bp Baldwin et al., 2003

Phenol hydroxylase

PCAHF

PCAHR

5' GAGRTSTGGCARGCSAA[Y] 3'

5' CCG[Y]SSAGCACGATGTC 3' 390bp Azhari et al., 2007

Protocatechuate 1,2,

dioxygenase


45

Table 2.4: Primer sequences designed by aligning the already reported

organophosphate degrading genes

Primer Sequence Product size Gene/enzyme recognized

by the primer

Opd-F2

Opd-R2

5' AGGTGGTGTTCCGGTAACCACT 3'

5' GGACTGAGCGCCTTGATCAAGA 3' 276bp Organophosphate

degrading hydrolase

Opd-F3

Opd-R3

5' ACAGTGTTCCGGTAACCACTCACAC 3'

5' CCGGCTCAAATCGTCAGTATCATCG 3' 474bp Organophosphate

degrading hydrolase

Opd-F4

Opd-R4

5' AAAGCGGCTGGCGTGCGAACGAT 3'

5' GAGGTTCACGCGATCCATCACGT 3'

691bp

Organophosphate

degrading hydrolase

OpdUpF

OpdDnR

5' GACAGGATTCTTGCGTGCTTGGC 3'

5' CAACAACCCGAACAGCCAGTCATTC 3' 752bp Organophosphate

degrading hydrolase

mpdUpF

mpdDnR

5' GAAACAAGCTGGTGCTGGTGGACAC 3'

5' CATAGTATCAGGTCGCCGAGCAGG 3' 530 bp Methyl Parathion

degrading hydrolase

Chapter 4 Biodegradation of profenofos

46

Chapter 3

Isolation, characterization and degradation potential of profenofos


1.20 Introduction

Profenofos (PFF) [(O-4-bromo-2-chlorophenyl)-O-ethyl-S-propylphophorothioate]

is a phosphorothioate organophosphate insecticide (Figure 3.1) which is a broad spectrum,

non-systemic and foliar insecticide. It is also used as acaricide i.e. to kill mites on a wide

range of crops including cotton, maize, sugar beet, soya beans, potatoes, vegetables and

tobacco (Reddy and Rao, 2008). It was developed for controlling insect pests that were

resistant to chlorpyrifos and other OPs (Worthing and Hance, 1991). It is one of the widely

used pesticides in Pakistan (Ismail et al., 2009) and other countries e.g. India, Australia,

Korea and USA (Rao et al., 2003; Kumar and Chapman, 2001; Min and Cha, 2000;

Dadson et al., 2013). One reason for the extensive use of PFF is a deceptive view of its

short half life in soil but it has been recognized as highly persistent and toxic at even low

concentrations (Zhao et al., 2008).

Figure 3.1 Chemical structure of profenofos

According to World Health Organization (WHO) this compound has been

classified as a moderately hazardous (Toxicity class II) pesticide (Abass et al., 2007;

Malghani et al., 2009a). Many reports reveal the toxicity of PFF to non target aquatic and

terrestrial organisms such as fish & earthworms (Kavitha and Rao, 2009; Liu et al., 2012).

Genotoxic effects of PFF on fish have also been reported (Prabhavathy et al., 2006;

Pandey et al., 2011). Moreover, it interferes with the biochemical activity in non target

insects, birds, animals and humans (Rao et al., 2003, McDaniel et al., 2004; Abass et al.,

2007). Generally, like other OP pesticides, PFF also exhibits acute neurotoxicity by


47

inhibiting acetyl cholinesterase enzyme (Costa, 2006). Other health effects include the

induced oxidant stress and consequent nephrotoxicity at high doses (Lin, et al., 2003).

Due to the toxicity concerns of PFF, remediation of this pesticide from

contaminated soils and water needs serious attention. In this context microbial degradation

is considered to be a potential strategy for the remediation of pesticide residues from

contaminated sites (Watanabe, 2001). Considerable attention has been given towards the

naturally existing bacterial isolates bestowed with pesticide degrading capacities. Natural

degraders endow the environment with the prospect of both eco-friendly and in situ

detoxification. Moreover, they exhibit efficient degradation in a wide range of

environmental conditions (Abe et al., 2011; Latifi et al., 2012).

Regarding the microbial degradation of PFF, limited number of bacterial strains

have been reported to date which include Pseudomonas putida & Burkholderia gladioli

(Malghani et al., 2009a), Pseudomonas aeruginosa (Malghani et al., 2009b) and Bacillus

subtilis (Salunkhe et al., 2013). These strains were found to hydrolyze PFF to produce 4-

bromo-2-chlorophenol (BCP). Further degradation of BCP by the isolated bacteria has not

been reported yet. BCP has also been proved to be toxic and a specific and sensitive

exposure biomarker for PFF (Dadson et al., 2013).

In addition to pure bacterial isolates, bacterial consortia obtained from highly

contaminated sites have also been found very efficient in removing the contaminants at

high concentrations (Krishna and Philip, 2008). While isolating the bacteria using

enrichment culture technique, highly efficient bacterial consortia are obtained for the

degradation of the pesticides. Sometimes, single pure isolates are not able to withstand

stress induced by the toxic pesticide metabolites; hence complete degradation of the

pesticide is resisted. However, bacterial consortium, a composite of some suitable bacterial

strains, helps in complete transformation of a pesticide into potentially harmless end

products. This can be attributed to the mutual activity of constitutive bacterial strains

which work in harmony with each other to overcome the toxic effects of pesticide.

The fate of pesticides in soil depends on both biotic and abiotic factors such as

physical and chemical characteristics of the substrate, nutrients status, pH, pesticides

characteristics (hydrophilicity, level of solubility), temperature and biotic factors such as

inoculum density (Karpouzas and Walker, 2000). These factors collaborate and harmonize


48

one another in the microenvironment to affect the process of bioremediation. Racke et al.,

(1990) suggested that the composition and size of soil microbial populations as well as the

status of metabolic activity are the determining factors regarding the biodegradation

feasibility as a remediation option. Bioremediation techniques endeavor to speed up the

naturally occurring biodegradation process by optimizing the conditions under which it

occurs. Although suitable microbial populations may be available for biodegradation of a

given contaminant, environmental conditions may limit or even inhibit this process in

many contaminated areas. In such type of cases, bio-stimulation of the degrading potential

of native microbial populations and/or the bioaugmentation of selected degrading

microorganisms to contaminated soil have been found efficient at enhancing pesticide

metabolism (Margesin et al., 2000; Sasek, 2003).

Generally, studies regarding the optimization of pesticide degradation involve one

factor at a time. But in the recent years, these have been found to be laborious and might

also lead to misinterpretations of the results. Therefore, a statistically developed model or

set of experiments, response surface methodology (RSM) was employed. RSM,

supported by the software, a statistical design of experiments, is a pragmatic modelization

technique derived for the estimation of the relationship of a set of controlled experimental

variables and the observed results. This method was developed to create a set of designed

experiments to find an optimum response of different variables (Box and Wilson, 1951).

However, central composite design based on response surface methodology (RSM) has

been found to effectively cope with most of the limitations encountered in the

optimization of pollutant degradation processes (Sridevi et al., 2011).

Although biodegradation of profenofos has been reported, such studies are scarce

or do not exist as yet for agricultural soils in Pakistan. Its extensive use and high

persistence determines its presence in agricultural lands where it is applied repeatedly. The

environmental concern of profenofos or its metabolites, therefore, has prompted to pay

attention towards the biodegradation of this pesticide. It has become increasingly

important to isolate microorganisms indigenously that are capable of degrading profenofos

and it would greatly hamper the problems associated with the agricultural and non-

agricultural use of PFF.


49

Here we aim to achieve scientific understanding of the metabolism of profenofos

and its degradation products by indigenously isolated bacterial strains/consortia in liquid

media and contaminated soil. Moreover, Central Composite Design (CCD) based on RSM

was employed for the optimization of various culture conditions for biodegradation of PFF

by the bacterial consortium. “CCD is a standard design of RSM and is well suited for

fitting process optimization.” This design is quite efficient in reducing the number of

experiments and gaining desired results and generating useful information about a

process.” This study can be an important asset in the area of developing processes for the

enhanced biodegradation/bioremediation of profenofos contaminated soils, sediments and

groundwater which would provide valid information for the environmental risk assessment

related to pesticide profenofos and attracting the attention of environmental scientists

towards this pesticide which has been neglected yet from the bioremediation point of view.

1.21 Materials and Methods

1.21.1 Development of profenofos degrading bacterial consortium

Soil samples were collected from cotton fields with prior history of PFF

applications and used for the isolation of PFF degrading bacteria. Repeated enrichment

culture technique was employed for the development of a bacterial consortium (Lakshmi

et al., 2009) using 250 ml Erlenmeyer flasks containing soil (10g), 100 ml MSM

supplemented with 1 ml/l Focht trace element solution (Appendix 8) and 50 and 100 mg/l

PFF as described in Chapter 2 Section 2.6. However, after three successive transfers into

the fresh MSM containing PFF a consortium named as PBAC was developed with PFF

degrading capability. For isolation of bacterial strains the consortium was serially diluted

and plated on MSM agar medium containing 100 mg/l PFF. The colonies presenting

visually distinct morphology were purified by repeated streaking and eight distinct

colonies were obtained which were named as PF-A, PF-B, PF-C, PF-D, PF-E, PF-F, PF-

G and PF-H. Growth of all the isolated strains and the consortium (PBAC) in MSM broth

with PFF was monitored spectrophotometerically in terms of optical density (OD) at 600

nm.


50

1.21.2 Molecular identification of PFF degrading bacterial strains comprising the

consortium PBAC

Ribosomal Intergenic Spacer Analysis (RISA) is a method of microbial

community analysis which provides means of comparing specie-specific sequences of

microbes in a community. Therefore, 16S-23S intergenic spacer region of all the eight

isolates (PF-A to PF-H) was amplified using primers IGS-forward (5′-

TGCGGCTGGATCACCTCCT-3′) and IGS-reverse (5′- GGCTGCTTCTAAGCCAAC-

3′) as described earlier (Yousaf et al., 2010). PCR products (10 μl) were digested with

EcoRI & HindIII (10U/µl, Fermentas) and the resulting DNA fragments were separated

by agarose gel (1% w/v) electrophoresis in TAE buffer to carry out the Restriction

Fragment Length Polymorphism (RFLP) analysis of 16S-23S IGS of the bacterial strains

in the consortium. Molecular identification was confirmed by 16S rRNA gene analysis

and evolutionary relationships of the isolates were studied as described in Chapter 2

Section 2.8. The 16Sr RNA gene sequences of the isolates were submitted in GenBank.

1.21.3 Morphology and biochemical analysis of PFF degrading bacterial isolates

Profenofos degrading bacterial strains were characterized by the standard

morphological and biochemical methods as described earlier (Chapter 2 Section 2.9).

1.21.4 Biodegradation of PFF by pure cultures and bacterial consortium PBAC

Degradation experiments with PFF as a sole source of carbon were carried out in

Erlenmeyer flasks containing 50 ml MSM (pH 7.0) supplemented with 100 mg/l PFF,

inoculated with pure isolates or the consortium (PBAC) to give final culture density of

0.6 g/l. Flasks were incubated at 37°C in a rotary shaker at 100 rpm. Un-inoculated flasks

served as controls and all the experiments were performed in three replicate.

1.21.5 Extraction and HPLC analysis of PFF residues

Cultures were harvested after regular intervals (24 h) and PFF residues were

extracted using equal volumes of dichloromethane and analyzed using HPLC as

described in Chapter 2 Section 2.11.1 - 2.11.3. A mixture of acetonitrile and water

(80:20) was used as mobile phase. However, PFF and its degradation product were


51

detected and quantified at a wavelength of 275 nm. Retention time for PFF and 4-bromo-

2-chlorophenol were 7.5 and 2.7 min respectively.

1.21.6 Optimization of culture conditions for PFF degradation using Response

surface Methodology (RSM)

The preliminary studies showed that pH, temperature and inoculum size (as

explained in Section 3.2.6) significantly affect the degradation of PFF by the consortium

PBAC. To optimize the interactive effects of three factors on PFF degradation, response

surface methodology (RSM) was employed. The analyzed response was the degradation of

PFF after three days of incubation. Using Design Expert software (trial version 8, Stat-

Ease, Inc., MN, USA), a 23 full factorial central composite design (CCD) with total 20

runs was employed. pH, temperature and inoculum density are the three experimental

factors. The symbols and levels of the three variables are given in Table 3.1. General

experimental setup (Chapter 2 Section 2.11) was followed to perform all 20 experiments

with modifications according to the variable ranges. Samples were harvested, extracted

and analyzed by HPLC as described above (Section 3.2.5).

Table 3.1: Experimental ranges and levels of independent variables

Independent variables Symbols Coded levels (Range)

Low (-1) Centre (0) High (+1)

pH of medium X1 6.0 7.0 8.0

Incubation temperature (°C) X2 30 37 40

Inoculum size of the culture (g/l) X3 0.2 0.4 0.6

X1, X2 and X3 are the short notations for the independent variables pH, temperature and inoculums size respectively.

These are the culture conditions which affect the rate of degradation of pesticide in liquid culture (and in soil)

In the system involving three variables, a mathematical relationship of the

response (PFF % degradation) of these variables was approximated by the quadratic

polynomial equation given below:

Y= a0 + a1X1 + a2X2 + a3X3 + a12X1X2 + a13X1X3 +

a23X2X3 + a11X12

+ a22X22 + a33X3

2 Eq. (1)


52

Where Y is the predicted response value, a0 is the constant, a1, a2 and a3 are the linear

coefficients; a12, a13 and a23 are the cross product coefficients; a11, a22 and a33 are the

quadratic coefficients.

1.21.7 PFF degradation by PBAC at different initial concentrations

Degradation experiments with various initial PFF concentrations (50–300 mg/l)

as the sole carbon source were performed in 250 ml Erlenmeyer flasks containing 50 ml

sterile MSM using PBAC as inoculum. Profenofos was introduced in the form of

acetonitrile solution. The cultures were incubated at 37°C and 100 rpm in a rotary

shaker for 192 h (8 days). Each treatment was set in triplicates with un-inoculated

samples as control. The extraction and analysis of PFF residues was carried out to

assess the potential of PBAC to tolerate and degrade PFF.

1.21.8 Soil microcosm studies for PFF degradation

Biodegradation of profenofos in soil (50 mg/kg, dry weight) was studied at

different soil moisture levels (20%, 40% 45% and 60%) as described in Section 2.12.

Moreover, PFF degradation was also studied in sterilized soil containing 50 mg/kg PFF

and different inoculum densities (1.6×105

, 1.6×10

6 and 1.6×10

7 and 1.6×10

8 CFU/g) of

bacterial consortium PBAC.

1.21.9 Identification of PFF metabolites

Samples containing residues of PFF and its metabolites obtained from culture

flasks were extracted with n-hexane-acetone (80:20) following Malghani et al., (2009a)

and dehydrated with anhydrous sodium sulphate. The solvent was evaporated with

rotavapor, dried residues were dissolved in dried acetonitrile, derivatized and analyzed by

Agilent Gas Chromatograph as described in Chapter 2, Section 2.13.

1.21.10Study of potential genes encoding OP hydrolases/oxygenases

Amplification of genes encoding organophosphate degrading hydrolases and

mono- or di-oxygenases in PFF degrading bacterial consortium PBAC was carried out

using primers mentioned in Table 2.4 and 2.5 as described in Chapter 2, Section 2.14.


53

1.21.11Biodegradation of other pesticides

Degradation experiments were carried out using various pesticides including

organophosphates (chlorpyrifos, diazinon, methyl-parathion, triazophos and

imidacloprid) and pyrethroids (cypermethrin) at 50 mg/l concentration as sole source

of carbon to evaluate the degradation spectrum of the PFF degrading bacterial

consortium PBAC. Minimal salt medium supplemented with each pesticide

individually was inoculated with PBAC. Cultures were harvested after 3 days of

incubation and pesticides residues were extracted. Cypermethrin was extracted two

times with n-hexane, separated layer of n-hexane containing pesticide was

evaporated and the dried residues were dissolved in methanol. A mobile phase

containing methanol and water was used for analyzing cypermethrin residues using

HPLC. All other pesticides used in the experiment, were extracted and analyzed as

described in section 3.2.5. However, detection wavelengths (λmax) were measured for

all other pesticides as described in Chapter 2, Section 2.12.1.

1.21.12Data analysis

Degradation rates and percent degradation were calculated as described in El-

Helow et al., (2013). Using following formulae:

(1). Degradation (%) = [initial profenofos concentration − residual profenofos

concentration]/initial pesticide concentration) × 100.

(2). Degradation rate (mg/h) = profenofos biodegradation (mg/l)/ time (h).

Kinetic model was determined by plotting log profenofos residues against time.

Degradation rate constant (k, 1/h) and half-life in days (DT50) were determined using Eq.

(2) and Eq. (3) as described in Cycon et al., (2009) and Jabeen et al., (2014).

Ct = C0e-kt

Eq. (2)

T1/2 = ln (2) / k Eq. (3)

Ct and C0 indicate the pesticide concentration at time “t” and time “zero”

respectively. Statistical analyses were performed on three replicates of data obtained from

all treatments. The significance of differences were treated statistically by one, two or


54

three way ANOVA and evaluated by post hoc comparison of means using Tukey’s test in

Statistica 6.0 software.

1.22 Results

1.22.1 Molecular identification of PFF degrading bacterial isolates

Following RFLP analysis of 16S to 23S IGS amplicons it was found that the

bacterial consortium PBAC was comprised of four bacterial strains based on the

restriction pattern. Restriction patterns of isolates B, D, G and H were found to be similar

(Figure 3.2A & B). Similarly, the restriction patterns of isolates E and F were similar

using both the enzymes indicating that they might be the same strains. However, isolates

A and C showed unique restriction patterns. Further, the sequence similarity deduced

from the 16S rRNA gene analysis and database comparison was in correspondence with

RFLP results. The isolates were identified as Achromobacter xylosoxidans PF1,

Pseudomonas aeruginosa PF2, Bacillus sp. PF3 and Citrobacter koseri PF4. Accession

numbers of PF1, PF2 PF3 and PF4 are KF201649, KF207917, KF207918 and KJ561165

(Table 3.2). Phyllogenetic relationship of the four bacterial strains showed that PF1, PF2,

PF3 and PF4 were placed with the well supported branches of Achromobacter

xylosoxidans, Pseudomonas aeruginosa, Bacillus sp. and Citrobacter koseri respectively

(Figure 3.3-3.6).

Table 3.2: Percent (%) similarity of profenofos degrading bacterial strains with

reported 16SrRNA gene sequences in the GenBank alongwith the GenBank

accession numbers

Bacterial Isolate Similarity with %

Similarity

PF1 (KF201649) Achromobacter xylosoxidans strain J2 (GU014534.1) 99%

PF2 (KF207917) Pseudomonas aeruginosa strain JKCM-H-2B (LC010673.1) 100%

PF3 (KF207918) Bacillus sp. strain 2DT (LM655315.1) 99%

PF4 (KJ561165) Citrobacter koseri isolate URMITE (LK054630.1) 99%

Numbers in parentheses indicate the GenBank accession numbers


55

Figure 3.2 Restriction Fragment Length Polymorphism of IGS gene from PF-A to PF-H

(Lane 1-8) restricted with A) EcoR1 and B) HindIII. PF-A displays a unique restriction

pattern (named as PF1); restriction pattern of PF-B, PF-D, PF-G and PF-H, are similar

(named as PF2); PF-C also displays a unique pattern (named as PF3); restriction patterns

of PF-E and PF-F are similar (named as PF4). M indicates the 1Kb Marker.

A

B


56

Figure 3.3 Neighbor joining tree showing the phyllogenetic relationship of strain PF1

with the related species based on the 16S rRNA gene sequences. Bootstrap values that are

expressed as the percentages of 1000 replications are shown at the nodes of the branches.

Achromobacter spanius LMG 5911 025686.1

Achromobacter piechaudii EY3860 (027186.1)

Achromobacter xylosoxidans A8 (074754.1)

Achromobacter insolitus LMG 6003 (025685.1)

Achromobacter denitrificans DSM 30026 (042021.1)

Achromobacter ruhlandii EY3918 (027197.1)

Achromobacter xylosoxidans Hugh 2838(044925.1)

Achromobacter xylosoxidans PF1 (KF201649.1)

Bordetella avium ATCC 35086 (041769.1)

Bordetella hinzii LMG 13501 (027537.1)

Bordetella pertussis CS (103933.1)

Bordetella parapertussis 522 (025950.1)

Bordetella petrii Se-1111R (025369.1)

Bordetella petrii DSM 12804 (074291.1)

Herbaspirillum huttiense subsp. putei 7-2 (028656.1)

Azohydromonas lata IAM 12599 (041244.1)

Burkholderia andropogonis LMG 2129 (104960.1)

Burkholderia soli GP25-8 (043872.1)

Burkholderia multivorans Struelens (029358.1)

Burkholderia latens R-5630 (042632.1)100

82

100

80

100

93

42

100 76

69

95

96

94

68

62

67

84

0.01


57




Pseudomonas_peli_R-20805_16S_(042451)

Pseudomonas_guineae_M8_(042607

Pseudomonas_cuatrocienegasensis_1N_(044569)

Pseudomonas_borbori_strain_R-20821(042450)

Pseudomonas_fulva_12-X_strain_(074659)

Pseudomonas_marincola_KMM_3042_(041592)

Pseudomonas_alcaliphila_AL15-21_(024734)_

Pseudomonas_psychrotolerans_C36_(042191)

Pseudomonas_stutzeri_A1501_(074829

Pseudomonas_stutzeri_ATCC_17588__LMG_11199_(103934)

Pseudomonas_stutzeri_ATCC_17588__LMG_11199_(041715)

Pseudomonas_otitidis_MCC10330_(043289)

Pseudomonas_resinovorans_NBRC_106553_(103921)

Pseudomonas_resinovorans_LMG_2274_(026534)_

Pseudomonas_aeruginosa_DSM_(026078)

Pseudomonas_aeruginosa_PF2_(KF207917)

Pseudomonas_aeruginosa_PAO1_(074828)

Pseudomonas_azotifigens_6H33b_16S_(041247)

Pseudomonas_denitrificans_ATCC_13867_(102805)

Pseudomonas_knackmussii_B13_16S_(041702)

Pseudomonas_delhiensis_RLD-1(04373184

100

100

83

80

100

100

54

29

60

86

73

37

48

39

86

36

0.005


58




Bacillus pocheonensis Gsoil 420 (041377.1)

Bacillus ginsengisoli DCY53 (109068.1)

Bacillus niacini IFO15566 (024695.1)

Bacillus vireti R-15447 (025590.1)

Bacillus circulans ATCC 4513(104566.1)

Bacillus siralis 171544 (028709.1)

Bacillus purgationiresistens DS22 (108492.1)

Bacillus firmus IAM 12464 (025842.1)

Bacillus flexus IFO15715 (024691.1)

Bacillus megaterium IAM 13418 (043401.1)

Bacillus megaterium QM B1551 (074290.1)

Bacillus sp. PF3 (KF207918.1)

Bacillus endophyticus 2DT (025122.1)

Bacillus humi LMG 22167(025626.1)

Bacillus niabensis 4T19 (043334.1)

Bacillus seohaeanensis BH724 (043083.1)

Bacillus aquimaris TF-12 (025241.1)

Bacillus isabeliae CVS-8(042619.1)

Bacillus carboniphilus JCM9731 (024690.1)

Bacillus sporothermodurans M215 (026010.1)

Bacillus shack letonii LMG 18435 (025373.1)

100

97

100

88

45

95

84

44

59

38

75

93

51

68

52

41

21

67

0.005


59




Salmonella enterica subsp. enterica serovar Paratyphi A str. AKU_12601 (074935.1)

Salmonella enterica subsp. enterica serovar Paratyphi A str. ATCC 9150 (074934.1)

Salmonella enterica subsp. enterica serovar Paratyphi C RKS4594 (074899.1)

Salmonella enterica subsp. enterica serovar Choleraesuis str. SC-B67 (074800.1)

Salmonella enterica subsp. houtenae DSM 9221 (044371.1)

Salmonella enterica subsp. salamae DSM 9220 (044372.1)

Salmonella enterica subsp. enterica serovar Typhi str. Ty2 (074799.1)

Salmonella enterica subsp. enterica serovar Typhimurium str. LT2 (074910.1)

Salmonella bongori NCTC 12419 (074888.1|)

Citrobacter farmeri CDC 2991-81 (024861.1)

Citrobacter koseri ATCC BAA-895 (102823.1)

Citrobactor koseri PF5

Citrobacter koseri CDC-8132-86 (104890.1)

Shigella flexneri 2a str. 301 (074882.1)

Shigella flexneri ATCC 29903 (026331.1)

Shigella sonnei CECT 4887 (104826.1)

Shigella boydii Sb227 Sb227 (074893.1)

Shigella boydii P288 (104901.1)98

97

59

100

52

85

9485

5227

49

72

83

89

86

0.002

Citrobactor koseri PF4


60

1.22.2 Morphological and biochemical characterization of PFF degrading bacterial

isolates

Morphological and biochemical characterization of all eight isolates is described

in Table 3.3. Based on the comparison with Bergey’s Manual of Determinative

Bacteriology following observations were recorded:

PF-A was found to resemble to Achromobacter.

PF-B, PF-D, PF-G and PF-H resembled to Pseudomonas sp.

PF-E and PF-F resembled to Citrobacter

PF-C resembled to Bacillus.

Hence these results were in harmony with the molecular identification of the bacterial

consortium. Morphology of the four strains in shown in Figure 3.7.


61

Table 3.3: Biochemical characteristics of profenofos degrading bacteria

Bacterial strain Morphology Biochemical characters

Colony

Morphology

Cell

morphology Gram’ s Staining

Sucrose/glucose

/Maltose

Oxidase/Orthinin

/arginine

PF-A

Round,

Smooth,

margin, shiny,

small, convex,

Rod

Motile

Gram negative -/+/- +/-/-

PF-B

Oval, Irregular

margin,

medium size,

shiny, green ,

flat,

Rod shaped

Motile

Gram negative +/+/+ +/-/+

PF-C

Off-white to

white, regular,

opaque

Short rods

Motile Gram positive +/+/- -/-/+

PF-D

Oval, Irregular

margin,

medium size,

shiny, green ,

flat,

Rod shaped

Motile


PF-E

Round,

smooth,

larger,

convex,

creamy color

Rods, Motile Gram negative +/-/- -/+/+

PF-F

Round,

smooth,

larger,

convex,

creamy color

Rods, Motile Gram negative +/-/- -/+/+

PF-G

Oval, Irregular

margin,

medium size,

shiny, green ,

flat,

Rod shaped

Motile


PF-H

Oval, Irregular

margin,

medium size,

shiny, green ,

flat,

Rod shaped

Motile



62

Figure 3.7 Profenofos degrading pure bacterial strains grown on LB agar medium


63

1.22.3 Antibiotic resistance assay

Response of all four PFF degrading bacterial strains to different antibiotics is

presented in Table 3.4.

Table 3.4: Response of profenofos degrading bacterial strains to different

antibiotics

Bacterial strains

PF1 PF2 PF3 PF4

Antibiotics Symbols

Kanamycin K Sensitive Resistant Sensitive Sensitive

Rifamycn RA Resistant Resistant Sensitive Resistant

Tetracycline TE Resistant Resistant Sensitive Sensitive

Chloramphenicol C Resistant Resistant Sensitive Sensitive

Nalidixic acid NA Resistant Resistant Sensitive Sensitive

Streptomycin S Resistant Resistant Sensitive Resistant

Erythromycin E Resistant Resistant Sensitive Sensitive

Gentamycin CN Resistant Sensitive Sensitive Sensitive

Carbenicillin PY Resistant Resistant Resistant Resistant

Ampicillin AM Resistant Sensitive Resistant Sensitive


64

1.22.4 Biodegradation of PFF in aqueous medium by pure cultures and PBAC

A comparative analysis of the degradation abilities of the consortium PBAC and

that of the pure bacterial cultures was carried out (Figure 3.8A). After 24 h of inoculation,

PBAC metabolized 37% of the added PFF while pure cultures of A. xylosoxidans PF1, P.

aeruginosa PF2, Bacillus sp. PF3 and C. koseri PF4 metabolized 9%, 12%, 10% and 7%

of added PFF respectively. PFF degradation by the PBAC significantly increased (P <

0.05) with incubation time, reaching 100% PFF within 72 h. However, none of the pure

cultures could significantly metabolize PFF to an extent as high as the consortium could.

The accumulation and subsequent degradation of 4-bromo-2-chlorophenol (BCP),

the major hydrolysis product of PFF was also monitored (Figure 3.8B). Maximum

amount of BCP was observed in the cultures containing PBAC where 20 mg/l BCP was

found after 48 h which was in line with the degradation of PFF. The concentration of

BCP decreased subsequently as the experiment proceeded. However, the scenario was

different in case of the individual bacterial cultures. It was found that after 72 h of

incubation 8, 9.7, 7.8 and 4 mg/l BCP was produced in the cultures inoculated with PF1,

PF2, PF3 and PF4 respectively. Further, the concentration of BCP was decreased to 7,

4.3, and 5.6 mg/l in the cultures containing PF1, PF2 and PF3 respectively. Noticeably,

no observable degradation of BCP by the strain PF4 was found as it continued to increase

until the end of experiment (120 h). Hence, degradation of both PFF and BCP was

significantly higher in cultures containing consortium PBAC. No significant degradation

was observed in un-inoculated controls.

The degradation followed the first order reaction as a straight line was produced

by plotting the log (ln) values (Ct/C0) of PFF residues against respective hours. The data

was interpreted statistically for the calculation of regression equation and first order

kinetic parameters (Table 3.5). Regression coefficient (R2) ranging from 0.8 to 0.998

indicated a good fit. The kinetic constant k (day-1

) was significantly higher for the PBAC

than that for the pure cultures. Similarly, the observed half lives of PFF in the cultures

containing pure PF1, PF2, PF3 and PF4 were significantly higher than that for the

cultures containing PBAC. An un-inoculated control showed a half life of 38.5 days.

Based on these observations the PBAC containing all four bacterial isolates was

considered for further PFF degradation studies.


65

Figure 3.8 Degradation (%) of profenofos and its metabolite BCP by pure

bacterial isolates and the bacterial consortium PBAC: A) Degradation (%) of PFF

(100 mg/l) and B) accumulation and degradation of BCP as a result of PFF

degradation by PBAC () and pure isolates PF1 (), PF2 (), PF3 (), PF4 ()

and control (). Each value is the mean of three replicates and error bars show

the standard error.

B

A


66

Table 3.5: Degradation kinetics of profenofos by pure cultures and consortium

PBAC

Treatments

MSM+PFF (100 mg/l) +

bacterial culture

Regression equation k (d-1

) t1/2 (Days) R2

A. xylosoxidans PF1 ln(Ct/C0)= 4.598-0.100x 0.100±0.003 6.9 0.964

P.aeruginosa PF2 ln(Ct/C0)= 4.623-0.151x 0.151±0.009 4.6 0.998

Bacillus sp. PF3 ln(Ct/C0)= 4.611-0.120x 0.120±0.009 5.8 0.958

C. koseri PF4 ln(Ct/C0)= 4.602-0.080x 0.080±0.006 8.7 0.989

Bacterial consortium

(PBAC) ln(Ct/C0)= 5.497-1.444x 1.444±0.003 0.5 0.970

Un-inoculated control ln(Ct/C0)= 4.601-0.018x 0.018±0.001 38.5 0.860


67

1.22.5 Biodegradation of PFF with different initial concentrations by PBAC

Dynamic curves of PFF degradation and subsequent accumulation and

degradation of BCP at different initial concentrations of PFF (Figure 3.9) indicate a

higher rate of PFF degradation at its lower initial concentrations. At 25, 50 and 100 mg/l,

added PFF completely disappeared within 24, 36 and 72 h respectively (Figure 3.9 A, B

and C). However, rate of PFF degradation slowed down at relatively higher

concentrations (Figure 3.9 D and E) as indicated by 95% and 80% degradation at 200 and

300 mg/l initial PFF concentration respectively.

A concomitant accumulation and degradation of BCP was observed at different

initial concentrations of PFF. As described in Section 3.3.4, in the cultures where higher

PFF was degraded, higher BCP concentration was observed. However, BCP production

decreased with increasing the initial PFF concentration which was in line with the slow

rate of PFF degradation at higher concentrations. After 24 h of incubation, 8 and 7 mg/l

BCP was observed at 25 and 50 mg/l PFF which was disappeared after 24 and 48 h

respectively. At 100 mg/l PFF, 22 mg/l BCP was observed after 48 h which decreased to

11 and 7 mg/l after 72 and 96 h respectively and completely disappeared after 120 h of

experiment.

Nevertheless, at higher concentration of PFF (200 and 300 mg/l), BCP

concentration continued to increase at a slow rate concomitantly with PFF degradation

and small degradation of BCP was observed at higher concentrations of PFF. These

results indicate that accumulation of higher concentrations of PFF and BCP caused

inhibitory effect on microbial growth and activity hence degradation of PFF and

subsequently the BCP is reduced.


68

Figure 3.9 Degradation of profenofos by the PBAC at different initial concentrations of

profenofos as a sole source of carbon and energy in MSM at 37°C and pH 7.0, (A) 25

mg/l, (B) 50 mg/l, (C) 100 mg/l, (D) 200 mg/l and (E) 300 mg/l indicating disappearance

of PFF (), accumulation and degradation of BCP () & un-inoculated control ().

Values are the Means of three replicates and error bars show the standard error.


69

1.22.6 Optimization of culture conditions for PFF degradation using RSM

Based on the CCD, 20 experiments were performed consisting of 8 full factorial

points, 6 central and 6 axial points located at the central and the extreme levels with 6

centre points designated as replications (Table 3.6). By applying the multiple linear

regression analysis on the experimental data, a polynomial quadratic equation was found

to represent the % PFF degradation as given by the following mathematical expression:

Y= – 417.17 + 47.20X1 + 14.37X2 + 346.25X3 + 0.61X1X2

– 27.33X1X3+1.77X2X3 – 3.76X12

–0.28X22

–192X32

(4)

X1, X2 and X3 denote three independent variables i.e. pH, incubation temperature

and inoculums size (of the culture) respectively. The negative and positive signs of the

regression coefficients indicate the antagonistic and synergistic effects of each variable

respectively. In this case an antagonistic effect associated with X13, X12, X2

2and X3

2

while a synergistic effect associated with X1, X2, X3, X1X2 and X2X3 can be concluded

from the regression equation.

The predicted values of PFF degradation (%) using Eq. (4) along with the

experimental values are given in Table 3.7. The parity plot (Figure 3.10) indicates a

satisfactory correlation (R2= 0.968) between the predicted and the actual (observed)

response values of % PFF degradation. R2 is the determination coefficient which is used

to measure the goodness of fit for the model. Value of R2 closer to 1 is an indication of

the stronger model and good prediction of the response. Hence a good value of the R2

(0.968) in this case is the evidence of the best fit of the model which shows a good

correlation between predicted and observed response.

The results of the quadratic response surface model fitting in the form of analysis

of variance (ANOVA) are given in Table 3.8. It is required to test the significance and

adequacy of the model. Moreover, the Fisher variance ratio (F-statistics) is the valid

measure of how well the variables describe the variation in the data about its mean. The

greater F-value indicates that the factors adequately explain the variation in the data and

the estimated factors are real. Hence the current model is highly significant as indicated

by the Fisher’s F-test (Fmodel= 31) and very low p-value (p>F= 0.0001). Moreover, p-

values and the F- statistics were calculated for the coefficient of each term (Table 3.8).


70

This implies that the main effects of pH, temperature and inoculum size were highly

significant as indicated by their p-values. This indicates that the three factors can act as

the limiting factor and small increase in values can alter the degradation rate of PFF. The

interaction effect of pH & temperature and pH & inoculum size were found to be very

significant (p<0.05) while interaction effect of temperature & inoculum size was

insignificant (p>0.05). However, the p-value of the model was 0.0001 implying that the

model is significant.


71

Table 3.6: The 23 factorial and central composite design for experiment

Run no. Order no. Variables in coded levels Comment

X1 (pH) X2 (incubation

Temperature)

X3 (Inoculum size)

1 5 -1 -1 -1 Full factorial

2 11 1 -1 -1 Full factorial

3 19 -1 1 -1 Full factorial

4 13 1 1 -1 Full factorial

5 7 -1 -1 1 Full factorial

6 14 1 -1 1 Full factorial

7 6 -1 1 1 Full factorial

8 15 1 1 1 Full factorial

9 4 -2 0 0 Axial

10 16 2 0 0 Axial

11 3 0 -2 0 Axial

12 1 0 2 0 Axial

13 10 0 0 -2 Axial

14 12 0 0 2 Axial

15 9 0 0 0 Center

16 18 0 0 0 Center

17 20 0 0 0 Center

18 17 0 0 0 Center

19 8 0 0 0 Center

20 2 0 0 0 Center


72

Figure 3.10 The parity plot of PFF degradation (%)


73

Table 3.7: The CCD matrix showing actual values (%) along with the experimental

values of PFF degradation

Standard no. Variables in un-coded levels Response (PFF degradation %)

X1(pH) X2 (Incubation temperature) X3 (Inoculum size) Actual Predicted

1 6.0 30 0.2 56.00 54.00

2 8.5 30 0.2 68.00 69.00

3 6.0 40 0.2 38.00 38.84

4 8.5 40 0.2 68.00 68.66

5 6.0 30 0.6 85.00 87.52

6 8.5 30 0.6 73.10 74.57

7 6.0 40 0.6 75.00 78.63

8 8.5 40 0.6 74.24 81.13

9 5.15 37 0.4 63.00 61.46

10 9.43 37 0.4 85.00 80.84

11 7.0 26 0.4 75.12 73.61

12 7.0 43 0.4 69.21 65.64

13 7.0 37 0.06 45.00 45.98

14 7.0 37 0.74 93.00 86.32

15 7.0 37 0.4 87.60 87.80

16 7.0 37 0.4 88.34 87.80

17 7.0 37 0.4 87.70 87.80

18 7.0 37 0.4 88.10 87.80

19 7.0 37 0.4 87.90 87.80

20 7.0 37 0.4 86.80 87.80


74

Table 3.8: Analysis of Variance (ANOVA) for the response (% degradation of PFF)

Source Degree of freedom F-value p-value

Prob>F

Model 9 31.25 < 0.0001*

X1 1 15.42 0.0028*

X2 1 5.19 0.0459*

X3 1 31.93 <0.0001*

X1X2 1 8.43 0.0157*

X1X3 1 24.74 0.0006*

X2X3 1 1.78 0.2122

X1X1 1 32.18 0.0002*

X2X2 1 40.15 <0.0001*

X3X3 1 10.30 <0.0001*

Residual 10

Lack of fit 5 105.59 <0.0001*

*Significant (p<0.05)

Adjusted R square = 0.9657

Predicted R square= 0.8578

Adeqate precision= 17.813


75

1.22.7 Response surface plots for PFF degradation

To better understand the relationship between the response (% degradation of

PFF) and the experimental variables (X1, X2 & X3) two dimensional surface plots were

analyzed. Figure 3.11 indicates that at optimum inoculum size, an increase in temperature

with pH up to an optimum point resulted in the increased % PFF degradation. However,

the trend is reversed with further increase in pH and temperature.

Figure 3.12 indicates the maximum % degradation of PFF at high inoculum size

and an optimum pH value. Hence effect of inoculum size was dependent on pH value. A

similar trend was observed by the inoculum size and temperature interaction (Figure

3.13). Hence effect of inoculum size increase on % degradation of PFF was dependent on

pH and temperature.

The predicted optimized % degradation of PFF was found to be 93.39% (~ 94%)

based on software at optimized values of variables i.e. pH 6.83, temperature 34.59°C (~

35°C) and inoculum size 0.59 g/l. Experiments were performed with the optimized values

deduced by software (Figure 3.14) to confirm the validity of the applied model and hence

94% degradation was obtained. This value is very close to the predicted % PFF

degradation (93.39%~94%) indicating the adequacy of the obtained model for PFF

degradation.


76

Figure 3.11 Contour plot (A) and Response surface plot (B) for profenofos

degradation (%) as a result of interaction of pH and temperature at constant

inoculum size (0.59 g/l) and 100 mg/l initial concentration of PFF.


77

Figure 3.12 Contour plot (A) and response surface plot (B) for profenofos

degradation (%) as a result of interaction of pH and inoculum size at constant

temperature (35°C) and 100 mg/l initial concentration of PFF.


78

Figure 3.13 Contour plot (A) and response surface plot (B) for profenofos

degradation (%) as a result of interaction of temperature and inoculum size at

constant pH (6.83) and 100 mg/l initial concentration of PFF.


79

Figure 3.14 Optimization ramp for profenofos degradation


80

1.22.8 Soil microcosm studies of PFF degradation

1.22.8.1 Optimization of soil moisture contents for PFF degradation

Biodegradation of PFF was found to be influenced significantly (P<0.05) by the

soil moisture contents. Data indicated that with increasing from 20 to 40% of soil

moisture contents (MC), PFF degradation increased from 20% to 31% of the added PFF

(50 mg/kg) after 10 days of experiment (Figure 3.15). After 40 days of experiment, the

56.5% and 92% PFF was degraded at 20% and 40% MC. However, increasing the MC to

45%, further increased PFF degradation and all of the added PFF was degraded by the

end of experiment. Further increase in soil moisture resulted in the retardation of PFF

degradation process as it is obvious from the data at 60% MWHC where only 48%

degradation was achieved after 40 days of experiment.

Effect of soil moisture contents on BCP accumulation (as a result of PFF

degradation by the consortium PBAC) and subsequent degradation was also monitored

and data showed that accumulation of BCP was increased in soils with 20 and 60% MCs

compared to that in the soil with 40 and 45% MWHC. Hence 45 % MC of soil was taken

as optimum for PFF and BCP degradation in soil by the PBAC. Further soil experiments

were performed at 45% soil MC.

1.22.8.2 Optimization of inoculum density for PFF degradation in sterilized soil

Soil was sterilized to clearly identify the efficiency of bacterial strains of the

PBAC in soil and optimize its inoculums size for the efficient PFF degradation. Data

showed that after 10 days of experiment, 37% of added PFF was degraded in soil

containing 1.6×108 CFU/g soil which completely disappeared by the 40 days of

experiment. Soils containing 1.6×105

, 1.6×10

6 and 1.6×10

7 CFU/g soil showed a slower

rate of PFF disappearance compared to that at 1.6×108. However, extending the

incubation period might lead to the complete degradation even at lower inoculums sizes.

BCP accumulation and degradation was also dependent on inoculum size and

minimum accumulation and maximum degradation of BCP was found at highest

inoculum density compared to that at lower ones (Figure 3.16). It was observed that

accumulation of BCP was increased to 5, 9, 10 and 8 mg/kg at 1.6×105

, 1.6×10

6 and

1.6×107 and 1.6×10

8 CFU/g respectively after 10 days of incubation. By the end of


81

experiment, no BCP was observed in the soil samples at 1.6×108

CFU/g soil inoculum

size. However at lower inoculums sizes, BCP tended to accumulate with slow rate of

degradation in the soil until the end of experiment which might be attributed to the fact

that lower inoculum size does not produce sufficient enzymatic activity to degrade the

PFF metabolite.


82

Figure 3.15 Degradation (%) of profenofos by bacterial consortium PBAC in the

sterilized soil at different moisture contents (MC) of soil; 20% (), 40% (),

45% () and 60% (), un-inoculated controls at 20% (), 40% (), 45 () and

60% () Solid lines indicate % profenofos degradation and dashed lines with

same marker indicate BCP accumulation and degradation at respective moisture

levels. Values are the means of three replicates and error bars represent the

standard error.


83

Figure 3.16 Degradation (%) of profenofos by bacterial consortium PBAC in the

sterilized soil at different inoculums densities: 1.6105 (), 1.610

6 (),

1.6107

(), 1.6108

() and un-inoculated control ().

Solid lines indicate

PFF degradation (%) and dashed lines with same marker indicate BCP

accumulation and subsequent degradation (mg/kg) at respective inoculum

densities. Values are the means of three replicates and error bars represent the

standard error.


84

1.22.9 Identification of profenofos metabolites

Time course analysis of the PFF degradation was carried out to get a clear idea of

production and subsequent degradation PFF metabolites using GC-MS as mentioned in

Materials and Methods.

Different metabolites were observed based on different retention times (RT) in

samples harvested at different time intervals. The sample at zero time showed a peak at

17.79 min as the major peak (peak 1) corresponding to PFF (Figure 3.17). Mass spectrum

of this peak was identical to that of authentic PFF standard showing molecular ion peak

with m/z 373 (Figure 3.18). Peak 1 was found to reduce in abundance in the

chromatograms of sample extracts obtained after 24 h and some new peaks (2,3,4,5 and

6) appeared at 10.37, 9.58, 8.25, 13.02 and 16.83 minutes respectively (Figure 3.19). All

of these peaks started decreasing in abundance as found in the TIC of samples harvested

after 72 h (Figure 3.20). Interestingly, in the TIC of samples obtained after 96 h (Figure

3.21), peak 2 again rose up compared to that in the previous TIC. Furthermore, two new

peaks appeared (7 and 8) at 7.63 and 16.99 min

Metabolites of PFF were identified on the basis of mass spectrum analysis. The

peaks 2 and 3 with m/z 208 and 183 appeared in the form of trimethylsilyl (TMS)

derivatives with m/z 280 and 241 respectively (Figure 3.22-3.23). These were identified

as 4-bromo-2-chlorophenol (BCP) and O- ethyl-S-propyl-O-hydrogen phosphorothioate

(EPHP). Peak 7 observed in the TIC of samples obtained after 96 h was identified to be

ethylene glycol with m/z 62 that was present in the form of TMS derivative with m/z 208

(Figure 3.24). Other peaks (4, 5, and 8) corresponded to some unknown metabolites. Peak

6 corresponding to m/z 356 was identified to be 4-bromo-2-chlorophenyl ethyl propyl

phosphate (BCPEPP) (Figure 3.25). The BCPEPP indicates the replacement of sulphur

by oxygen forming profenofos oxon. The formation of 4-bromo-2-cholorophenol

(molecular weight 208) proves the breaking of ester bond linkage of the parent compound

by the bacterium. Further disappearance of hydrolysis products in the subsequent samples

indicates their degradation to smaller products. Hence the bacterial consortium PBAC is

efficient enough to degrade profenofos and its metabolites. Based on these observations a

degradation pathway of PFF was predicted (Figure 3.26). Table 3.9 presents the summary

of identified metabolites and their detail.


85

Table 3.9: Different metabolites of profenofos and their detail

Name Adduct

m/z

Retention

time (RT

min)

m/z of original

compound

4-bromo-2-chlorophenol (BCP) 280 10.35 208

O-ethyl-S-propyl-hydrogen

phosphate 256 9.58 183

Ethylene glycol 208 7.63 62

4-bromo-2-chlorophenyl ethyl

propyl phosphate (BCPEPP) - 16.83 356


86

At zero time

Figure 3.17 Total ion chromatogram (TIC) of the extract of the spent culture

medium containing profenofos at zero time showing the only peak corresponding

to profenofos at 17.79 min as the most abundant or major peak. No metabolite

peaks were found in this TIC.

Retention Time


87

Profenofos

Figure 3.18 Mass spectrum of profenofos

Br

Cl

OP

S

OO


88

After 24 h


medium containing profenofos harvested after 24 h. Peak 1 is reduced in

abundance while new peaks 2, 3, 4 5 and 6 appeared at different retention times.

Retention Time


89

After 72 h


medium containing profenofos harvested after 72 h. Peak 1 is reduced further

compared to that in previous TIC. Peaks 2, 3, 4 and 6 decreased in abundance

while peak 5 disappeared.

Retention Time


90

After 96 h


medium containing PFF harvested after 96 h. Abundance of peak 1 is reduced

very much, of peak 2 increased compared to previous TICs. Other peaks

disappeared with the appearance of two new peaks 7 and 8 which were not found

in previous TICs.

Retention Time


91

4-bromo-2-chlorophenol

Figure 3.22 Mass spectrum of 4-bromo-2-chlorophenol (BCP), the major

hydrolysis product of profenofos, formed in the culture medium after 1

day of incubation.

OH

Cl

Br


92

O- ethyl-S-propyl-O-hydrogen phosphorothioate (EPHP)

Figure 3.23 Mass spectrum of O- ethyl-S-propyl-O-hydrogen

phosphorothioate (EPHP), the second hydrolysis products of profenofos,

formed in the culture medium after 1 day of incubation.

P

OO•

OS


93

Ethylene glycol

Figure 3.24 Mass spectrum of ethylene glycol, a proposed ring cleavage

product of BCP, formed in the culture medium after 3 days of incubation.


94

4-bromo-2-chlorophenyl ethyl propyl phosphate (BCPEPP)

Figure 3.25 Mass spectrum of 4-bromo-2-chlorophenyl ethyl propyl

phosphate (BCPEPP)


95

Figure 3.26 Proposed biodegradation pathway of profenofos by the bacterial consortium

PBAC: 1) Profenofos is hydrolyzed to EPHP and a halogenated metabolite, 4-bromo-2-

chloropheno (BCP); 2) Debromination of BCP; 3) Hydroxyl radical attack on 2-

chlorophenol to produce hydroquinone; 4) Conversion of hydroquinone to benzoquinone;

5) Ring cleavage of quinone and (proposed) formation of ethylene glycol (OTMS); 6)

Production of ethylene glycol


96

1.22.10Detection of OP degrading genes in PFF degrading bacterial strains (PF1-

PF4)

PCR was carried out with opd, opdA, mpd and oxygenase primers (Table 2.3 &

2.4). Chapter 2, Section 2.15, Potentially opdA gene was amplified from genomic DNA

of the profenofos degrading bacterial isolates by using primers described in Sharaf et al.,

(2006) (Chapter 2, Table 2.3). PCR product of expected size i.e. 1155 bp was obtained

which was confirmed by agrarose gel electrophoresis (Figure 3.27). The variants of opdA

in different bacteria encode enzyme (organophosphate hydrolase, OPH) capable of

hydrolyzing a variety of organophosphate pesticides.

Although potential opdA with expected product sizes were obtained, further quest

into the opdA gene sequence analysis revealed no definitive success because the

sequences of the potential opdA genes (amplified in this study) did not match any of the

previously known hydrolase gene sequences reported in the GenBank.

Figure 3.27 Amplification of opdA gene potentially encoding an OP hydrolase

(OPAA) in PFF degrading bacterial strains PF1, PF2, PF3 & PF4 (Lanes 3, 4, 5 and

6 respectively). M indicates 1 Kb marker.


97

1.22.11Biodegradation of other pesticides by the bacterial consortium PBAC

Degradation capability of the PBAC was studied by inoculating the consortium

PBAC in minimal media containing different pesticide substrates as sole source of carbon.

Medium containing PFF served as a positive control for biodegradation activity of PBAC

and PFF was degraded completely within 3 days. All other pesticides were detected at their

respective detection wavelength (λmax) that were found to be 275 nm, 290 nm, 245 nm, 280

nm, 254 nm, 270 nm and 254 nm for profenofos, chlorpyrifos, triazophos, methyl

parathion, diazinon, imidacloprid and cypermethrin respectively.

Among all substrates (other than PFF), significantly higher degradation (69.3%) of

cypermethrin was observed and lowest degradation was observed for imidacloprid.

Interestingly, similar extent of degradation of all organophosphates tested (other than PFF)

was observed. Chlorpyrifos, methyl parathion, triazophos and diazinon were degraded to

55, 59, 57.3 and 57.7% within 3 days of incubation (Figure 3.28). Hence the PFF

degrading PBAC had a broad spectrum of degradation. This is the first report describing

the biodegradation affinity of PFF degrading microorganisms for diverse class of

chemicals.


98

PFF= Profenofos CP= Chlorpyrifos IMD= Imidacloprid

CYP= Cypermethrin TR= Trizophos M.P=Methyl parathion

Dia= Diazinon

C = indicates un-inoculated control for the respective pesticide

Figure 3.28 Degradation of PFF and other pesticides (50 mg/l) as a sole source of

carbon by PBAC (after 3 days of incubation). Data are the means of three

replicates and error bars show the standard error. Values with different letters are

significantly different statistically (p<0.05, Tukey’s test).


99

1.23 Discussion

In the present study, complete degradation of PFF and its hydrolysis metabolite,

BCP by a bacterial consortium was observed. In contrast, individual strains viz.

Achromobacter xylosoxidan, Pseudomonas aeruginosa, Bacillus sp. and Citrobacter

koseri degraded PFF and BCP inefficiently and the extent of degradation was

significantly lower compared to that by the PBAC. Effectiveness of bacterial consortium

has been attributed to the combined and collaborative activity of all the bacterial isolates

that constitute a consortium and help one another to withstand the stressed conditions

produced due to the generation of toxic pollutant metabolites (Nestler, 2001). Our

findings are in line with the above statement as bacterial consortium displayed the

complete degradation of PFF and its toxic metabolite, BCP. However, within the same

incubation period, incomplete degradation of PFF was obtained by pure isolates.

Moreover, capability of PFF degrading consortium to degrade high concentration

of PFF (200 mg/l) and its toxic metabolite, BCP might be regarded as the result of the

synergistic effect of different bacterial members of the PBAC. Malghani et al., (2009b)

reported PFF degradation by a pure P. aeruginosa at lower PFF concentration. Similar

enhanced and effective degradation of many recalcitrant pollutants has been reported by

the pesticide-degrading bacterial consortia enriched from various crop soils (Sørensen et

al., 2002, 2008). Some reports of bacterial consortia include those degrading 4-

nitrophenol (Laha and Petrova, 1998), endosulfan (Awasthi et al., 2000), chlorpyrifos

(Lakshmi et al., 2009) and methyl parathion & chlorpyrifos simultaneously (Pino and

Penuela, 2011).

The bacterial consortium PBAC was found to embrace the bacterial strains which

belong to metabolically active and diverse genera such as Bacillus and Pseudomonas are

known to be metabolically very active genera capable of degrading a variety of

organophosphates including the diazinon, chlorpyrifos, methyl parathion and other

chemicals (Ghassempour et al., 2002; Lakshami et al., 2008; Anwar et al., 2009).

Literature survey reveals that no report is available for the PFF degradation by

Achromobacter xylosoxidans and Citrobacter koseri yet both are important candidates for

the biodegradation of many other compounds. A. xylosoxidans has been reported to

degrade nitro compounds such as p. nitrophenol (Wan et al., 2007) and organochlorines


100

(Singh and Singh, 2011). C. koseri is a facultative anaerobe and belongs to

Enterobacteriaceae which is a large family consisting of gram negative harmless

symbiotic as well as many human pathogenic bacteria. Interestingly, most of them have

been helpful in biodegradation of OP and other pollutants (Singh et al., 2004; Cycon et

al., 2013; Ghanem et al., 2007).

Previous studies showed inoculum size, temperature and pH to be significant

factors, affecting biodegradation of pollutants (Awasthi, 2000; Diez, 2010; Wolski et al.,

2005). This may be due to the environmental stresses such as high concentrations of the

pollutant, variations in pH and temperature which may be responsible for retarding or

enhancing the growth of the degrading bacteria. Temperature has been reported as an

important factor to affect the degradation of hazardous chemical compounds (Zhao et al.,

2008). Therefore, in the present study such parameters were optimized for the

degradation of PFF and regarding the effect of pesticide concentration,

In contrast to the conventional “single factor at a time experiments”, we report the

interaction and simultaneous effect of various factors; pH, temperature and initial

inoculum size on PFF degradation following RSM which greatly reduced the time,

explained the effect of different factors in a more pronounced way as well as described

the interactive effect of different variables on PFF degradation. The PFF degrading

consortium was capable of degrading PFF at all the tested pH, temperature and inoculums

sizes which can be attributed to the combined activities of the bacterial strains in the

consortium. However, RSM model helped to get maximum degradation at a set of

optimized range of conditions.

Increasing the inoculums size, no doubt, increased the degradation of PFF but this

effect was in turn dependent of pH and temperature. At optimum pH and temperature,

high population of degrading bacteria can degrade the pollutant more quickly and

efficiently. Effect of temperature and pH on PFF degradation is evident from Ali and

Badawy, 1982 who demonstrated remarkable effect of temperature of on PFF

degradation. Contrary to Malghani et al., (2009b) who described that lower pH was more

supportive for PFF degradation than the higher pH; our study revealed that PBAC was

capable of exhibiting higher PFF degradation at higher pH. It might be possible that some


101

key enzymes of the current bacterial strains involved in PFF degradation would be active

at neutral to high pH.

The current study was also extended to optimize PFF degradation in soil. For such

studies bioaugmentation of selected species of microorganism has proved an excellent

approach (Gentry et al., 2004). Successful remediation of pesticide contaminated soils by

the use of bacteria has been reported extensively for parathion (Barles et al., 1979),

ethoprophos (Karpouzas et al., 2005) and chlorpyrifos (Lakshami et al., 2008). The

optimization of bioremediation processes in soil depends on many factors such as the soil

properties (moisture contents, soil pH) and the survival and population of the degrading

cultures. Optimum soil moisture plays important role in the remediation or removal of the

pollutants from the soil (Johnson et al., 1998).

In a soil remediation system, inoculum size plays a key role for degrading the

organic pollutants (Labana et al., 2005). High initial density of bacteria could compensate

for the initial population decline, degrade the toxic pollutant and multiply (Comeau et al.,

1993). The present study was consistent with the previous findings in that longer lag

phase was observed at low initial inoculum densities as compared to that at higher

inoculum densities. These results signify the importance of optimizing the inoculums

density while studying pollutant degradation.

Previous studies regarding the PFF degradation described the formation of BCP

through ester bond breakage but no further metabolites were investigated. In this study an

attempt was made to predict the biodegradation pathway of PFF. Being in line with

previous studies, hydrolysis was found to be the first step in degradation of PFF

producing 4-bromo-2-chlorophenol (BCP) and O- ethyl-S-propyl-O-hydrogen

phosphorothioate (EPHP). However, our study provided the insights into the further

degradation of BCP accompanied by ring cleavage. Identification of ethylene glycol

helped to predict a pathway for PFF degradation. BCP was predicted to undergo

debromination followed by the hydroxyl radical attack which resulted in various oxidized

products (catechol, quinones). Ethylene glycol was assumed to be produced through ring

cleavage of quinone (Hong et al., 2003). The quinone is not phenolic hence TMS

derivative was not possible. Therefore being unstable, quinone was not detected

throughout the course of experiment. However, we strongly assume the generation of


102

ethylene glycol through further cleavage of the quinone in this study. This is the first

report of biodegradation of BCP by profenofos degrading bacteria and predicted

biodegradation pathway of profenfos. These results put forward the candidacy of PFF

degrading consortium for remediation of PFF contaminated sites. However, the full

degradation pathway and mineralization of intermediates require further investigation.

The PFF degrading bacterial consortium proved a good candidate for

biodegradation of other organophosphate pesticides and pyrethroids as it efficiently

degraded cypermethrin and other tested OP pesticides (diazinon, methyl parathion,

imidacloprid) This aspect reveals the diverse metabolic and enzymatic activities of the

bacterial strains comprising the consortium which help in inducing each other’ s enzyme

systems for utilizing different pollutants and hence allow the consortium to quickly adapt

to pesticide contaminated environments. Therefore, we conclude that the isolated

bacterial consortium can potentially be used for the bioremediation of not only

organophosphate pesticides but also pyrethroid contaminated sites.

Potentially, organophosphate degrading gene, opdA was amplified in the PFF

degrading bacterial strains. This gene encodes organophosphate hydrolase which

hydrolyzes many OP pesticides. There are many reports that emphasize the involvement

of opdA gene in OP degrading organisms (Horne et al., 2002; Sharaf et al., 2006).

However, no report is yet available for the existence of opdA in PFF degrading bacteria.

In the present study, we hypothesized that potentially, opdA like gene could be involved

in the hydrolysis of PFF to 4-bromo-2-chlorophenol (BCP) and O- ethyl-S-propyl-O-

hydrogen phosphorothioate (EPHP). However, the sequence of the amplicon regarded as

probable “opdA gene” did not match with any of the reported OP degrading genes in the

GenBank. This aspect of the study needs further research and analysis to identify the

gene(s)/enzymes involved in the hydrolysis of PFF.

Chapter 4 Biodegradation of chlorpyrifos

103

Chapter 4

Isolation, characterization and degradation potential of chlorpyrifos


4.1 Introduction

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

(CP), is one of the widely used, toxic and broad spectrum organophosphate (OP)

insecticides. Chemical structure is shown in Figure 4.1. It is an important ingredient of

various household formulations which are effective against termites, bees, flies and

mosquitoes (Bicker et al., 2005; Mohan et al., 2007). Chorpyrifos is also used extensively

against agricultural insect pests of a variety of vital crops such as cotton, cereals,

vegetables since many years (Fang et al., 2006; Wang et al., 2007). Although, many OP’s

including CP were initially regarded as less persistent and toxic, there is escalating

concern that these pesticides or there metabolites are highly persistent in the environment

as well as toxic and hence lead to undesirable health issues (Ragnarsdottir 2000; Alavanja

et al., 2013).

Figure 4.1 Chemical structure of chlorpyrifos

A consequence of continuous domestic and agricultural use of CP is widespread

contamination of environment leading to serious damage to non-target organisms and

ecosystems (Rovedatti et al., 2001; Anderson and Hunta, 2003; Vogel et al., 2008; Farag

et al., 2010). In the environment, CP is converted to TCP, a persistent metabolite which is

resistant to biotic and abiotic degradation owing to the presence of three chloride residues

on the N-aromatic ring (Racke et al., 1996; Robertson et al., 1998; Singh et al., 2003;


104

Chishti and Arshad, 2013). Moreover, TCP has higher water solubility as compared to the

parent compound; hence it leaches to the water bodies causing widespread contamination

of aquatic environments (Xu et al., 2008; Grzelak et al., 2012; Watts, 2012). CP and TCP

toxicity has been linked to broad-spectrum effects including neurological disorders,

developmental disorders, autoimmune disorders and interruption of many vital functions

in higher animals and humans (Sogorb et al., 2004; Mehta et al., 2008; Alavanja and

Bonner, 2012; Ventura et al., 2012; Estevan et al., 2013). However, TCP contributes

more than CP to pollute the environment due to its antimicrobial nature and hence

leaches to and persists in the water bodies (Vogel et al., 2008; Xu et al., 2008).

All these concerns imply that elimination of both CP and TCP from the

environment to alleviate their hazardous effects is imperative. Conventionally, many

approaches, including chemical treatment, photodecomposition and incineration can be

applied for the remediation of contaminants (Olexsey and Parker, 2006), however, most

of them are expensive, environmental unfriendly and not applicable for contamination at

low concentration. Bioremediation approaches which mainly involve microorganisms or

plants with the right metabolic pathways seem to be the most feasible technology for

remediation of CP, TCP and related contaminants (Thengodkar and Sivakami, 2010).

Contrary to the earlier findings that TCP, being highly toxic, inhibits the

proliferation of CP degrading microorganisms in soil (Racke et al., 1990), many bacterial

strains capable of degrading CP and TCP as a sole source of carbon and energy have been

isolated and characterized during recent years (Feng et al., 1997; Lu et al., 2013). Most of

them can utilize CP co-metabolically. A review of CP and TCP biodegradation studies

has been presented by Maya et al., (2011). A modified and updated review of CP and

TCP degrading microorganisms during last decade has been given in Table 4.1. Recently

reported CP and TCP degrading bacterial strains include Bacillus cereus (Liu et al.,

2012), a Stenotrophomonas maltophilia strain MHF ENV20 (Dubey and Fulekar, 2012)

and Cupriavidus sp. DT-1 (Lu et al., 2013).

It has also been generally recognized that microbial degradation of CP may be

affected by many biotic and abiotic factors such as tolerance to initial pesticide

concentration, microbial population, optimum growth temperatures and optimum pH. It

has been well established that tolerance to these environmental factors vary from one


105

microorganism to another (Karpouzas and Walker, 2000; Sharma, 2012; Singh et al.,

2003; Chishti et al., 2013). Either biotic or abiotic, degradation of CP in the environment

results in the production of TCP and diethyl thiophosphoric acid (DETP) (Racke et al.,

1996; Robertson et al., 1998; Singh et al., 2003 Chishti et al., 2013). Recently, more

attention has been given to metabolism of CP indicating TCP degradation and few de-

chlorination products of TCP have been reported by researchers few years back.

Complete metabolism of CP and its metabolites has not yet been reported.

In the present study, Mesorhizobium sp. HN3 a novel CP degrading bacterial

strain also capable of degrading TCP under different culture conditions was isolated and

characterized. The kinetics of CP biodegradation including accumulation and utilization

of TCP and the governing constants thereafter were also determined, as these parameters

vary depending upon bacterial strains and concentration/nature of pollutants, a clear

understanding of the biodegradation kinetics of CP and TCP is prerequisite for in situ

bioremediation. Moreover, degradation products of CP and TCP were also identified.


106

Table 4.1: Previously reported chlorpyrifos and TCP degrading bacteria

No.of

reports

Chlorpyrifos degrading

bacteria

Degradation efficiency (%) of CP

& TCP References

1. Enterobacter strain B-14 100% of 250 mg/l CP in 2 days Singh et al., 2004

2. Alcaligenes faecalis DSP3 i)100% of 100 mg/l CP in 10 days

ii) 100% of 100 mg/l TCP in 10

days

Yang et al., 2005

3. Agrobacterium tumefaciens

ASM-5 50% of 100 mg/l CP in 14 days

Sharaf et al.,

2006

4. Stenotrophomonas sp. YC-1 100% of 100 mg/l CP in 24 h Yang et al., 2006

5. Sphingomonas sp. i) 100% of 100 mg/l CP in 24 h

ii).100% of 20 mg/l TCP in 2 days Li et al., 2007

6. Pseudomonas fluorescens,

Brucella melitensis, Bacillus

subtilis, Bacillus cereus,

Klebsiella species, Serratia

marcescens and Pseudomonas

aeruginosa

i) 75–87% of 50 mg/l CP in 20 days

by Pseudomonas aeruginosa

ii) 67% of 2.5 mg/l TCP in 12 h

only by Pseudomonas aeruginosa

Lakshami et al.,

2008

7. Sphingomonassp.Dsp-2

Stenotrophomonas sp Dsp-4,

Brevimundus sp.Dsp-7

Bacillus sp Dsp-6

100% of 100 mg/l CP in 24 h

Li et al., 2008

8. Paracoccus sp. TRP i) 100% of 50 mg/l CP in 4 days

ii) 100% of 50 mg/l TCP in 4 days Xu et al., 2008

9. Bacillus firmus 50% of 25 mg/l CP in 20 h Sabdono, 2008

10. Pseudomonas aeruginosa 52% of 50-75 mg/l CP Fulekar and

Geetha, 2008

11. Bacillus pumilus C2A1 i) 89% of 1000 mg/l CP in 15 days

ii) 100 % of 300 mg/l TCP in 8

days

Anwar et al.,

2009

12. Pseudomonas sp. 84% of 10 mg/l CP in 120 h Singh et al., 2009

13. Leuconostoc mesenteroides

WCP907, Lactobacillus

brevis WCP902,

Lactobacillus plantarum

WCP931, and Lactobacillus

sakei WCP904.

100% of 30 mg/l CP after 8 days

Cho et al., 2009

14. Bacillus licheniformis ZHU-1 99% of 100 mg/l CP in 10 days Zhu et al., 2010


107

Continued from previous page

Table 4.1: Previously reported chlorpyrifos and TCP degrading bacteria

15. Four Pseudomonas spp. Two

Agrobacterium spp. One

Bacillus sp.

i) 76-84% of 100 mg/l CP in 10days

by four Pseudomonas spp.

ii) 87.5-90% of 90 mg/l in 10 days 10

days by Pseudomonas spp.

iii) 62.7 & 64% of 75 mg/l CP in 10

days by two Agrobacterium spp

iv) 76.8 and 77.5% of 60 mg/l TCP

by Agrobacterium spp

v) 52% of 50 mg/l CP by Bacillus sp.

vi) 79.5% of 45 mg/l TCP by

Bacillus sp.

Maya et al.,

2011

16. Acinetobacter sp,

Pseudomonas

putida, Bacillus sp,

Pseudomonas aeruginosa,

Citrobacter freundii,

Stenotrophomonas sp,

Flavobacterium sp,

Proteus vulgaris,

Pseudomonas sp,

Acinetobacter sp, Klebsiella

sp and Proteus sp.

i) 39% of 150 mg/l CP after 5 days

ii) 100% of 150 mg/l CP after 5 days

in the presence of glucose

Pino and

Peñuela, 2011

17. Stenotrophomonas

maltophilia MHF ENV20

100% of 50 mg/l CP in 2 days Dubey and

Fulekar, 2012

18. Bacillus cereus 78.85% of <150 mg/l CP in 5 days Liu et al., 2012

19. Streptomyces sp. AC5 and

Streptomyces sp. AC7 90% of 25 and 50 mg/l CP after 24 h

Briceno et al.,

2012

20. Pseudomonas putida NII

1117,

Klebsiella sp NII 1118,

Pseudomonas stutzeri NII

1119,

Pseudomonas aeruginosa NII

1120

70.84% of 500 mg/l of CP by

consortium after 30 days in soil

Sasikala et al.,

2012

21. Cupriavidus sp. DT-1 i) 100% of 100 mg/l CP in 6 h

ii) 100% of 100 mg/l TCP in 6 h Lu et al., 2013

22. Cupriavidus pauculus P2 100% of 100 mg/l TCP in 10 h Cao et al., 2012

23.

24. Alcaligenes sp JAS1.

300 mg/l of CP in 12 h

Silambarasan

and Abraham,

2013


108

4.2 Materials and methods

4.2.1 Enrichment and isolation of CP degrading bacterial strains

Three different agricultural soil samples were collected from fields with previous

history of OP (including CP) applications. Chlorpyrifos degrading bacteria were isolated

by enrichment culture technique following Anwar et al., (2009) as described in Chapter

2, Section 2.6-2.7.

Once all the isolates were purified by repetitive sub culturing, their growth and

CP degradation potential was monitored on MSM agar plates containing CP (100 mg/l)

as the only source of carbon and energy and incubated at 37°C until growth appeared.

Growth and CP degradation potential of the isolates were also observed in liquid

cultures (MSM) supplemented with 100 mg/l CP as the only source of carbon and energy.

Degradation potential of all the isolates was determined and compared by quantifying the

residual concentration of chlorpyrifos using HPLC. On the basis of degradation

capability, a bacterial isolate named as HN3 was selected for further CP degradation

studies.

4.2.2 Identification and characterization of selected strain HN3

Total genomic DNA of the isolate HN3 was extracted. Molecular identification

was carried out by 16S rRNA gene analysis and evolutionary relationships of the isolate

were studied as described in Chapter 2 Section 2.8. The 16S rRNA gene sequence of the

isolate was submitted in GenBank.

Biochemical and morphological characterization were carried out as described in

Chapter 2 Section 2.9. Moreover, morphology was also studied using Scanning Electron

Microscopy (SEM).

4.2.3 Experimental set up for CP degradation studies

Inoculum of strain HN3 was prepared and biodegradation experiments were

performed as described in Chapter 2 Section 2.10-2.11. Chlorpyrifos degradation studies

were carried out in 250 ml Erlenmeyer flasks containing 50 ml MSM supplemented with

2% HN3 inoculum and 100 mg/l CP under various culture conditions as described in

respective Sections. The flasks were incubated at 37°C and 100 rpm in rotary shaker


109

unless otherwise mentioned. For all the treatments, un-inoculated flasks served as

controls and all the experiments were performed in triplicates. Samples of the liquid

medium were periodically removed for analyzing the growth rates and the residues of CP

and TCP.

4.2.4 Extraction and analysis of CP residues

Extraction and analysis of CP residues were carried out following the method

described in Chapter 2 Section 2.11.1-2.11.3. CP and TCP were detected at a wavelength

of 290 nm. Retention time for CP and TCP were 8 and 4.47 min respectively.

4.2.5 Turbidometric study to monitor the growth of the bacterial strain, HN3

A Turbidometric method as described in Jyothi et al., (2012) was employed for

monitoring the growth of the strain HN3 wherever required throughout the course of this

study. Increase in turbidity of the culture was due to the growth of the HN3 by utilizing

the pesticide (Maya et al., 2011). Cell dry mass was determined for the Mesorhizobium

culture having an OD600nm of 1.0 and was used as standard for calculating cell dry mass

for all the cultures (of HN3) of different optical densities.

4.2.6 Optimization of temperature and pH for biodegradation of CP by HN3

To optimize temperature for degradation of CP by strain HN3, culture flasks

containing MSM (pH 7.0) were incubated at 30, 37 and 40°C in rotary shaker at 100 rpm.

Degradation capacity of CP by Mesorhizobium sp. HN3 was also monitored at different

initial pH i.e. acidic (6.0), basic (8.0) and neutral (7.0). MSM with different pH

(maintaining buffered conditions) was prepared as described by Anwar et al., (2009),

supplemented with CP (100 mg/l), inoculated with HN3 and incubated at 37°C.

4.2.7 CP degradation in minimal and complex media

To explore the degrading potential of Mesorhizobium sp. HN3 at a range of initial

pesticide concentrations, MSM containing different initial concentrations of CP i.e., 10 to

300 mg/l as sole source of carbon and energy was used.


110

To investigate if HN3 can utilize CP as a sole source of carbon and nitrogen, a

nitrogen free medium (Appendix 9) was prepared and used. The medium was

supplemented with 100 mg/l CP that served as a source of carbon and nitrogen.

To determine if Mesorhizobium sp. HN3 can degrade CP in the presence of easily

available carbon source, MSM was supplemented with CP (100 mg/l) and glucose (1 g/l).

4.2.8 Kinetics of CP degradation by HN3 at different initial concentrations of CP

Biodegradation of CP by strain HN3 was investigated at different initial

concentrations (50, 100, 200, 300 and 400 mg/l) and kinetic parameters were determined

as described by Pirt, (1975). Rates of pesticide degradation (Qs), metabolite production

(QP) and cell mass productivity (Qx) were determined by calculating the slope in their

respective plots versus time (h). Product yields (Yp/s) and cell mass yield (Yx/s) were

determined by dP/dS and dX/dS where dP, dS and dX are the changes in concentrations of

product, pesticide and the bacterial cell mass respectively per unit time. Specific growth

rate (), the growth of a bacterial cell per unit cell per unit time, was determined by

plotting the ln(X/X0) versus time, where X0 and X are the initial cell mass (g/l) and cell

mass (g/l) at time ‘t’ respectively (calculated during exponential phase at different time

intervals). Specific productivity (qp) and specific rate of CP degradation (qs) were

multiple of and Yp/x and Yx/s.

4.2.9 Biodegradation of TCP (primary metabolite of CP)

4.2.9.1 Biodegradation of TCP in minimal and complex media

MSM supplemented with different initial concentrations of TCP (25, 50, 100, 200

and 300 mg/l) was inoculated with Mesorhizobium sp. HN3 and incubated at 37°C and

100 rpm in rotary shaker. Degradation of TCP was also investigated in the presence of

easily available carbon source, by adding glucose (1 g/l) along with the TCP (100 mg/l)

in the culture medium.

4.2.9.2 Extraction and analysis of TCP residues

Extraction and analysis of TCP residues were performed as described in this

Chapter Section 4.2.4.


111

4.2.9.3 Detection of chloride ions produced during CP and TCP degradation

To investigate the production of chloride ions as a result of CP and TCP

degradation, a chloride assay described by Cao et al., (2012) was used with some

modifications to determine the concentration of chloride ions released due to degradation

of CP and TCP. For this purpose, MSM was replaced by a chloride free medium

(Appendix 10). Two initial concentrations of CP and TCP in (independent experiments)

i.e., 100 and 200 mg/l were employed (in triplicates) to study accumulation of chloride

ions in the media.

Mesorhizobium was inoculated in chloride free medium containing 100 mg/l and

200 mg/l CP separately (in duplicates). Un-inoculated flasks served as controls. After

three days of incubation at 37oC and 100 rpm, samples from all replicates were harvested

and centrifuged. During each sampling, 20 ml culture was centrifuged at 12,000 rpm for

15 minutes and the supernatant was treated to estimate the chloride ions concentration in

the culture media. Similarly the chloride ion concentration was also determined for the

cultures containing 100 and 200 mg/l TCP.

Supernatant (10 ml) of each sample was taken in Erlenmeyer flasks. A 0.5 ml of

potassium dichromate (K2CrO4) was added to each sample and titrated with 0.0141 molar

solution of silver nitrate until a reddish brown color appeared. Chloride ion was

calculated according to the following formula:

C (mg/l) = 24.99225 (V1-V2)/V1*1000

Where C is the concentration of chloride ions, V1 and V2 represent the titration volume

of sample and of control respectively.

4.2.10 Soil microcosm studies of CP

Biodegradation of chlorpyrifos was studied in soil at different moisture levels and

inoculum densities as described in Section 2.12. Moreover, a comparison of CP

biodegradation was carried out in sterilized and unsterilized soil to test the efficacy of

Mesorhizobium sp. HN3.


112

4.2.11 Identification of CP metabolites

Samples containing residues of CP and its metabolites, periodically obtained from

culture flasks were extracted with dichloromethane and derivatized with N, O-Bis-

(trimethylsilyl) -trifluoroacetamide (BSTFA, a derivatizing reagent for Gas

Chromatography) and analyzed by gas chromatography-mass spectrometry (GC-MS) as

described in Chapter 2 Section 2.13.

4.2.12 Study of potential genes encoding OP hydrolases/oxygenases

Amplification and analyses of opd, mpd, opdA encoding hydrolases and different

oxygenase genes encoding mono- or di-oxygenases in Mesorhizobium sp. HN3 were

carried out using primers mentioned in Tables 2.4 and 2.5 in Chapter 2, Section 2.14.

4.2.13 Data Analysis

Statistical analyses were performed on three replicates of data obtained from all

treatments. The significance of differences were treated statistically by one, two or three

way ANOVA and evaluated by post hoc comparison of means using Tukey’s test in

Statistica 6.0 software.

Kinetic model of CP degradation was determined by plotting log CP residues

against time. A straight line was obtained for all test concentrations of CP (50- 400 mg/l),

following first order kinetics model. Therefore, degradation rate constant (k, h-1

) and half-

life (T1/2) in days were determined using Eq. (2) and Eq. (3) as described in Chapter 3

Section 3.2.12.

Ct = C0e-kt

Eq. (2)

T1/2 = ln (2) / k Eq. (3)

4.3 Results

4.3.1 Isolation and selection of CP degrading bacterial strain

Following enriched culture technique sixteen bacterial isolates were obtained

initially. These isolates were streaked individually and purified on LB agar plates

containing 100 mg/l CP. Eight of the sixteen bacterial isolates were found to grow on


113

MSM agar as well as in MSM broth containing 100 mg/l CP as a sole source of carbon

and energy (Figure 4.2). One isolate HN3, showing complete degradation of CP (100

mg/l) within 5 days of incubation was considered to be the most efficient. Chlorpyrifos

degradation by this strain was further investigated under a range of culture conditions

and in soil.

Figure 4.2 Degradation (%) of chlorpyrifos as a sole source of carbon and energy in the

MSM by 8 selected bacterial strains: HN1 (), HN2 (), HN3 (), HN4 (), HN5

(), HN6 (), HN7 () and HN8 () and in un-inoculated control () at 37C and 7.0

pH. Values are the means of three replicates and error bars represent standard error.


114

4.3.2 Molecular, morphological and biochemical identification of strain HN3

Morphological and biochemical characters of isolate HN3 were compared to those

described by Jarvis et al., (1997) which confirmed strain HN3 to be a Mesorhizobium sp

(Table 4.2). Morphology is apparent from Figure 4.3 and 4.4. The 16S rRNA gene

sequence of the strain HN3 showed 99% identity with 16S rRNA gene sequence of

Mesorhizobium sp. STM 4018, GenBank accession number. EF100516.1 and it was

grouped in a well-supported branch with various Mesorhizobium spp. (Figure 4.5).

GenBank accession number of the Mesorhizobium sp. HN3 16s rRNA gene is JN119831.

4.3.3 Antibiotic resistance assay

Antibiotic assay showed that Mesorhizobium sp. HN3 was resistant to kanamycin,

nalidixic acid, streptomycin, rifamycin, tetracycline, and carbenicillin while it showed

sensitivity towards gentamycin, erythromycin and chloramphenicol and ampicilin.


115

Figure 4.3 Mesorhizobium sp. HN3 grown on LB-agar plate after 48 h of incubation

Figure 4.4 Scanning Electron Microscopy Image of Mesorhizobium sp. HN3


116

Figure 4.5 UPGMA tree showing the phyllogenetic relationship of strain HN3 with the

related species based on the 16S rRNA gene sequences. Bootstrap values that are


Mesorhizobium_metallidurans_(NR_042685.1)

Mesorhizobium_tarimense_(NR_044051.1)

Mesorhizobium_gobiense_(NR_044052.1|)

Mesorhizobium_tianshanense_(NR_024880.1)

Mesorhizobium_mediterraneum_(NR_042483.1)

Mesorhizobium_temperatum_(NR_025253.1)

Mesorhizobium_huakuii_(NR_043390.1)

Mesorhizobium_amorphae(NR_024879.1)

Mesorhizobium_loti_(NR_074162.1)

Mesorhizobium_opportunistum_(NR_074209.1)

Mesorhizobium_plurifarium_(NR_026426.1)

Mesorhizobium_thiogangeticum_(NR_042358.1)

Mesorhizobium_sp.(HQ836166.1)

HN3_(JN119831.1)

Mesorhizobium_sp.(EF100516.1)


Sinorhizobium_terangae_(NR_044842.1)

Sinorhizobium_kostiense_(NR_042484.1)

Sinorhizobium_saheli(NR_026096.1)

98

75

96

95

71

56

67

32

100

76

40

100

71

99

0.0000.0050.0100.015


117

Table 4.2: Biochemical and morphological characteristics of Mesorhizobium sp. HN3

Biochemical Tests Colony and cell

Morphology

Test Reaction with

Mesorhizobium sp. HN3 Result Colony shape Round

Sucrose Yellow green + Size Small

Maltose Orange - Surface Shiny

Innocitol Brown + Odour Plant

like

Mannose Yellow green + Elevation Flat

Raffinose Yellow green + Margins Smooth

O-nitrophenyl B-

galactopyraoside (ONPG)

Yellow + Cell motility Motile

Cell shape Short

rods

Lysin decarboxylase Orange _ Gram’ s

reaction

Gram

positive

Arginin dihydrolase Light green +

Ornithine decarboxylase Green _

Tryptophane deaminase Yellow _

H2S Off white _

Urea Pink +

Melonic acid Brown +

ADON Green _


118

4.3.4 Biodegradation of CP by Mesorhizobium sp. HN3

4.3.4.1 Optimum temperature for the CP degradation by Mesorhizobium sp. HN3

Data indicating the effect of temperature on the biodegradation of CP by

Mesorhizobium sp. HN3 is shown in Figure 4.6. In the presence of 100 mg/l CP, 33 and

18% of the added pesticide was degraded after 24 h of incubation at 37 and 40°C which

was significantly higher compared to that at 30°C where only 5% degradation was

observed. After 5 days of incubation, complete CP degradation was observed at 37°C

compared to 85% and 55% at 40 and 30C respectively. At 30°C complete degradation

was achieved after 10 days of incubation. Results indicate that the temperature

significantly affected the rate of CP degradation.

4.3.4.2 Optimization of pH for CP degradation by Mesorhizobium sp. HN3

Efficient degradation of CP was achieved at all the three initial pH tested. At 100

mg/l initial concentration, at pH 7.0, the entire added CP was degraded after 5 days of

incubation (Figure 4.7). Degradation was relatively slow at alkaline pH (8.0) whereby

100% degradation was achieved after 7 days of incubation and a further decline in the

degradation was observed at acidic pH (6.0) as 100% degradation was observed after 8

days of incubation. Among the three pH conditions tested for CP degradation by

Mesorhizobium sp. HN3, pH 7.0 was found to be optimum.

4.3.4.3 Biodegradation of CP at different initial concentrations

Mesorhizobium sp. HN3 was able to degrade CP efficiently up to 400 mg/l initial

concentration in MSM whereby degradation was achieved in concentrations dependent

manner. Chlorpyrifos degradation and bacterial biomass production at different initial CP

concentrations after 3 days of incubation are presented in Figure 4.8A. In the cultures

containing 50 and 100 mg/l CP, 100 and 85 % degradation was achieved respectively

whereas at 200, 300 and 400 mg/l initial concentrations, 45%, 33% and 15% CP was

degraded respectively. After three days of incubation, cell biomass (g/l) was highest at

100 mg/l initial CP concentration and declined gradually at concentrations beyond this.

As depicted in Figure 4.8B, specific rate of CP degradation was dependent on initial

concentration with an increase in specific degradation rate at lower initial CP


119

concentrations (50-100 mg/l) and a decline in the rate at higher initial CP concentrations.

Mesorhizobium sp. HN3 could tolerate higher CP concentration, with delayed

degradation i.e., 40% and 30% respectively after 16 days of incubation at 1000 and 1200

mg/l CP (data not shown). Complete degradation of CP was achieved after 5, 7, 9 and 10

days at 100, 200, 300 and 400 mg/l respectively.


120

Figure 4.6 Degradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 as a sole source

of carbon and energy at different incubation temperatures; 30C (), 37C () and 40C

(). Dashed lines show un-inoculated controls; 30C (), 37C () and 40C ().

Values are the means of three replicates and error bars represent standard error.


121


source of carbon and energy at different initial pH; 6.0 (), 7.0 () and 8.0 ().

Dashed lines indicate un-inoculated controls at pH 6.0 (), 7.0 () and 8.0 ().

Values are the means of three replicates and error bars represent the standard

error.


122

Figure 4.8 A) Degradation of chlorpyrifos by Mesorhizobium sp. HN3 at different initial

concentrations of chlorpyrifos in MSM as a sole source of carbon and energy after 3 days

of incubation () and consequent growth of Mesorhizobium sp. HN3 (). Values are the

means of three replicates and error bars represent the standard error. B) Effect of initial

CP concentrations on specific degradation rate of CP (, qs).


123

4.3.4.4 Kinetics of CP degradation and TCP accumulation and degradation

thereafter

Effect of initial concentrations of CP on kinetic parameters viz. specific

degradation rate (qs), specific growth rate (µ),substrate (CP) consumption variables (Qs,

Qx, Yx/s, qs) and product (TCP) formation parameters (Qp, Yp/s, qp) are presented in Table

4.3. The CP consumption and TCP production parameters were high at lower CP

concentrations i.e., (50-100 mg/l) followed by a decline at higher concentrations (200-

400 mg/l). Rate of TCP accumulation (QP) increased with the increase in CP

concentrations up to 200 mg/l with a gradual decrease at higher concentrations. TCP

yield (Yp/s) and qp were highest at 100 mg/l initial concentration and decreased at higher

concentrations. As illustrated in Figure 4.9, at 50 and 100 mg/l, maximum concentration

of TCP was observed after 48 hrs of incubation and all of the TCP produced as a result of

CP hydrolysis was degraded after 96 and 144 hrs respectively. At higher concentrations,

TCP was detected in the culture media even after 240 hrs of incubation which might be

due to the continuous production and slow degradation. Lag phase of bacterial growth

was extended with an increase in initial CP concentration beyond 100 mg/l.

Figure 4.10 shows the fitting results of the kinetic model based on the

experimental data of CP degradation. The degradation followed the first order reaction as

a straight line was produced by plotting the ln values (Ct/C0) of CP residues against

respective hrs. The residue data were therefore, interpreted statistically for the calculation

of regression equation and first order kinetic parameters (Table 4.4). Regression

coefficient indicating the degradation rate further supported the findings that the CP

persistence increases with increasing initial concentration.


124

Figure 4.9 Kinetics of CP degradation at 37C by Mesorhizobium sp. HN3 as a sole

source of carbon and energy at different initial concentrations; a) 50 mg/l, b) 100 mg/l,

c) 200 mg/l and d) 300 mg/l showing residual CP concentration (), TCP

concentration () and cell biomass of Mesorhizobium sp. HN3 () in the culture

media. Values are the means of three replicates and error bars represent the standard

error.


125

Table 4.3: Kinetic parameters* for chlorpyrifos degradation and product (TCP)

formation thereafter by Mesorhizobium sp. HN3 in liquid cultures containing different

initial concentrations of the pesticide

Each value is the means of three replicates± standard errors. All the values differ from

each other significantly at p< 0.05

*Kinetic parameters:

µ (/h): specific growth rate

Qs: mg substrate consumed /l/h

Qx: mg cell mass produced /l/h

Yx/s: mg cells /mg substrate utilized

qs: mg substrate consumed /mg cells /h

Qp: mg TCP produced /l/h

Yp/s: mg TCP produced /mg substrate consumed

qp: mg TCP produced /mg cells /h

Initial CP

concentration

(mg/l)

Substrate utilization parameters

µ

(/h)

Qs

(mg/l/h)

Qx

(mg/l/h)

Yx/s

(mg/mg/h)

qs

(mg/mg/h)

50 0.090±0.000 1.15±0.01 1.52±0.01 1.24±0.005 0.080±0.001

100 0.280±0.005 1.43±0.02 4.60±0.05 1.70±0.040 0.164±0.002

200 0.058±0.000 1.30±0.01 2.88±0.00 0.78±0.003 0.075±0.001

300 0.023±0.000 0.72+0.00 2.56+0.03 0.45±0.001 0.050±0.000

400 0.025±0.000 0.26+0.00 2.21+0.01 0.35±0.003 0.045±0.001

Product formation parameters

QP

(mg/l/h)

YP/S

(mg/mg/h)

qp

(mg/mg/h)

50 1.15±0.010 0.40±0.005 0.015±0.040

100 2.24±0.020 0.81±0.010 0.073±0.030

200 2.67±0.010 0.50±0.020 0.013±0.010

300 1.08±0.030 0.23±0.010 0.005±0.000

400 0.99±0.001 0.17±0.000 0.002±0.000


126

Figure 4.10 First order kinetics of chlorpyrifos degradation in MSM at different initial

concentrations; 50 mg/l (), 100 mg/l (), 200 mg/l (), 300 mg/l () and 400 mg/l

().


127

Table 4.4: First order kinetics parameters for chlorpyrifos degradation by Mesorhizobium

sp. HN3 in liquid cultures containing different initial concentrations of the pesticide

CP Concentration

(mg/l) Rate constant (h

-1)

t1/2

(Days) R

2

Regression

equation

50 0.025 1.16 0.917 3.853-0.025x

100 0.023 1.26 0.977 4.699-0.023x

200 0.015 1.93 0.933 5.557-0.015x

300 0.014 2.06 0.911 6.161-0.014x

400 0.012 2.41 0.900 6.451-0.012x


128

4.3.4.5 Co-metabolic degradation of CP by Mesorhizobium sp. HN3

Co-metabolic degradation of CP by Mesorhizobium sp. HN3 was monitored by

comparing the pesticide degradation by adding glucose in the culture medium. Higher CP

degradation was observed in the cultures containing glucose as compared to those

without glucose. After 2 days of incubation, 75% and 59% CP was degraded in the

cultures with and without glucose respectively. After 4 days of incubation no CP residues

were recovered in cultures containing glucose while 9% CP was still present in cultures

without glucose that also vanished completely by the 5th

day (Figure 4.11).

Figure 4.11 Co-metabolic degradation of chlorpyrifos by Mesorhizobium sp. HN3.

Degradation of chlorpyrifos by Mesorhizobium sp. HN3 in MSM containing chlorpyrifos

and added glucose () and in MSM containing chlorpyrifos as a sole source of carbon

and energy () with respective un-inoculated controls () and ().Values are the means

of three replicates and error bars represent the standard error.


129

Degradation of CP as a sole source of carbon and nitrogen by HN3

In nitrogen free medium, efficient degradation of CP by Mesorhizobium sp. HN3

was achieved as 100% of the added CP was degraded within 6 days of incubation. By the

7th

day of incubation no CP could be recovered from the cultures (Figure 4.12).

Figure 4.12 Degradation of chlorpyrifos (100 mg/l) by Mesorhizobium sp. HN3 in

nitrogen free medium (), bacterial biomass () and un-inoculated control ().Values

are the means of three replicates and error bars represent the standard error.


130

4.3.5 Biodegradation of TCP by Mesorhizobium sp. HN3

4.3.5.1 Biodegradation of TCP at different initial concentrations

Chlorpyrifos degrading bacterium, Mesorhizobium sp. HN3 was found to degrade

TCP, a more toxic chlorinated metabolite of CP. At different initial concentrations, TCP

was degraded in concentration dependent manner similar to the observations recorded for

CP degradation. At 25, 50, 100 and 200 mg/l initial concentrations of TCP complete

degradation was observed after 3, 5, 6 and 9 days (of incubation) respectively. However,

with increasing concentrations of TCP up to 300 mg/l biodegradation rate was relatively

slower with 88% degradation after 10 days of incubation (Figure 4.13). In another

experiment, addition of glucose in the medium containing TCP at 100 mg/l resulted in

enhanced biodegradation as it shortened the duration of TCP utilization from 6 to 4 days

of incubation (Figure 4. 14).

4.3.5.2 Release of chloride ions in culture media containing CP and TCP

Release of chloride ions in the culture media as a result of biodegradation of CP

and TCP was investigated to understand dehalogenation potential of Mesorhizobium sp.

HN3. The calculated equimolar concentrations of chloride ions in 100 and 200 mg/l CP is

30.4 and 60 mg/l respectively and is equivalent to the total of three chloride ions from

one molecule of CP. Hence in our study, estimated amount of chloride ions in the culture

medium (containing 100 and 200 mg/l CP) was 20.1 and 40 mg/l after three and four

days respectively. These chloride concentrations are corresponding to the release of two

chloride ions in the culture media from one molecule of CP. As the experiment

proceeded, concentration of chloride ions in the above mentioned media increased to

almost 30.4 and 60 mg/l after 7 and 9 days respectively showing the release of all three

chloride ions (Figure 4.15).

In case of TCP, calculated equimolar concentrations of chloride ions in 100 and

200 mg/l TCP is equal to 53.7 and 106 mg/l that corresponds to total three chloride ion.

Hence during the TCP degradation by Mesorhizobium sp. HN3, chloride ions

concentration reached almost 53.7 and 106 mg/l in the media with 100 and 200 mg/l

initial TCP after 7 and 10 days respectively indicating the release of total three chloride

ions (Figure 4.16).


131

Figure 4.13 Degradation (%) of TCP by Mesorhizobium sp. HN3 as a sole source of

carbon and energy at different initial concentrations: 25 mg/l (), 50 mg/l (), 100 mg/l

(), 200 mg/l (), 300 mg/l () and un-inoculated control (). Values are the means of

three replicates and error bars represent the standard error.


132

Figure 4.14 Co-metabolic degradation of TCP. Degradation of TCP by Mesorhizobium

sp. HN3 in MSM containing TCP and added glucose (), in MSM containing TCP as a

sole source of carbon and energy () and respective un-inoculated controls () and

().Values are the means of three replicates and error bars represent the standard error.


133

Figure 4.15 Analysis of chloride ions produced as a result of chlorpyrifos degradation in

a chloride free medium. Biodegradation and release of chloride ions is shown side by

side. Biodegradation of chlorpyrifos in the chloride free culture medium containing CP

100 mg/l (), CP 200 mg/l (), concentration of chloride ions released at CP 100 mg/l

() and CP 200 mg/l ().Values are the means of three replicates and error bars represent

the standard error.


134

Figure 4.16 Analysis of chloride ions produced as a result of TCP degradation in a

chloride free medium. Biodegradation and release of chloride ions is shown side by side.

Biodegradation of TCP in the chloride free culture medium containing TCP 100 mg/l ()

and 200 mg/l (). Concentration of chloride ions released at CP 100 mg/l () and CP

200 mg/l ().Values are the means of three replicates and error bars represent the

standard error.


135

4.3.6 Soil microcosm studies of CP

4.3.6.1 Optimization of soil moisture level for CP degradation in unsterilized soil

Biodegradation of CP was found to be affected significantly by the soil moisture

contents. Data showed that as the soil moisture was increased from 20% to 40% of

maximum water holding capacity (MWHC) of soil, CP degradation was increased from

18% to 55% of the added CP (50 mg/l) after 10 days of experiment (Figure 4.17).

However, in the soils with higher MCs i.e. 60% and 80%, about 21% and 13% of the

added CP was respectively left in the soil extracts after 10 days of experiment. At the end

of experiment (40 days), 59%, 100%, 69% and 37% of the added CP was degraded in soil

with 20%, 40%, 60% and 80% MCs respectively.

In inoculated soil, effect of soil moisture contents on TCP accumulation (as a

result of CP degradation) and degradation was also monitored and data showed that

accumulation of TCP was higher in soils with 20%, 60% and 80% MCs compared to the

soil with 40% MC. As with CP, TCP degradation was also supported at 40% MC of soil.

Hence 40% MC of soil was considered to be the optimized moisture level for CP and

TCP degradation by Mesorhizobium sp. HN3 and further soil experiments were

performed at 40% soil MC.

4.3.6.2 Biodegradation of CP in sterilized and unsterilized soil

Degradation efficiency of Mesorhizobium sp. HN3 (2x109 CFU/ g of soil) in

sterilized and un-sterilized soil was studied. The treatments to be compared included

followings:

1. Un-sterilized and un-inoculated soil (control-1, T1)

2. Un-sterilized soil inoculated with Mesorhizobium sp. HN3 (T2)

3. Sterilized and un-inoculated soil (control-2, T3)

4. Sterilized soil inoculated with Mesorhizobium sp. HN3 (T4)

The comparative analyses of all four treatments showed that HN3 not only degraded CP

in the sterilized soil but it also enhanced the degradation in the un-sterilized soil.

After 10 days of experiment, only 19% CP was degraded in the un-sterilized &

un-inoculated soil (T1) followed by a non significant increase in further CP degradation

(only 38% at the end of experiment). In un-sterilized soil containing HN3 (T2) and


136

sterilized soil containing HN3 (T4), 58% and 50% CP was degraded respectively,

followed by the complete degradation of CP at the end of experiment. No CP residues

were detected in the samples harvested at the end of experiment (Figure 4.18).

A similar trend was observed in case of TCP degradation. In un-sterilized control

soil (T1), more TCP was accumulated than that in the un-sterilized containing HN3 (T2)

and sterilized soil containing HN3 (T4). Moreover, very small CP degradation and

negligible TCP residues were observed in the sterilized and un-inoculated soil (T3).

We can conclude that Mesorhizobium HN3 is an efficient tool in the

bioremediation of CP contaminated soil where it can work equally well alone as well as

in collaboration with the existing indigenous microbial communities of soil.

4.3.6.3 Optimization of inoculum density for CP degradation in sterilized soil

Soil was sterilized to clearly identify the efficiency of HN3 strain and optimize its

inoculums size for the efficient CP degradation. Data indicated that after 20 days of

experiments, in soil containing 2×109 CFU/g soil, 75% of the added CP was degraded

which was completely disappeared by the 30 days of experiment. Soils containing 2×105

and 2×107 CFU/g soil, showed a slower rate of CP disappearance compared to that at

2×109.

TCP accumulation and degradation was also dependent on inoculum size and

minimum accumulation and maximum degradation of TCP was found at highest

inoculum density compared to that at lower ones (Figure 4.19).


137

Figure 4.17 Degradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 in the

unsterilized soil at different soil moistures: 20% (), 40% (), 60% () and 80% ().

Solid lines indicate CP degradation (%) and dashed lines with same marker indicate TCP

accumulation and degradation (mg/kg) at respective moisture level. Un-inoculated

controls are indicated by 20% (), 40% (), 60% () and 80% (). Values are the

means of three replicates and error bars represent the standard error.


138

Figure 4.18 Degradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 in the

unsterilized and sterilized soil. Un-sterilized control soil (un-inoculated) (), Un-

sterilized soil inoculated with HN3 (), Sterilized control soil (un-inoculated) (),

Sterilized soil inoculated with HN3 (). Solid lines indicate CP degradation (%) and

dashed lines with same marker indicate TCP accumulation and degradation (mg/kg) at

respective moisture level. Values are the means of three replicates and error bars

represent the standard error.


139

Figure 4.19 Degradation (%) of chlorpyrifos by Mesorhizobium sp. HN3 in the sterilized

soil at different inoculums densities: 2109 (), 210

7 () and 2 10

5 () and un-

inoculated control (). Solid lines indicate CP degradation (%) and dashed lines with

same marker indicate TCP accumulation and subsequent degradation (mg/kg) at

respective inoculum densities. Values are the means of three replicates and error bars



140

4.3.7 Identification of Chlorpyrifos metabolites

Biodegradation of CP was confirmed by GC-MS analysis of the extracts

obtained from the spent culture media containing CP as a source of carbon and energy.

Metabolites of CP were identified on the basis of mass spectrum analysis. Presence of CP

was indicated at a retention time of 16.02 min and the peak corresponding to CP was

designated as peak 1 (Figure 4.20). Mass spectrum of this peak was identical to that of

authentic CP standard showing molecular ion peak with m/z 351 (Figure 4.21). The time

course GC-MS analysis of bacterial culture extracts containing CP and its metabolites

showed that peak corresponding to CP disappeared as the metabolism proceeded and in

the meanwhile some new peaks appeared at different retention times. Hence in the TIC of

48 h extracts five peaks designated as 2, 3, 4, 5 and 6 appeared at the RT of 10.23, 11.39,

7.72, 7.30 and 18.98 min (Figure 4.22).

Mass spectra of peaks 2, 3 and 4 were further analyzed. Base peaks at m/z 197

and 169 were identified to be the hydrolysis products TCP (peak 2) and DETP (peak 3)

respectively. While peak at m/z 210 was identified to be 3,5,6, trichloro-2-

methoxypyridine, TMP (peak 4). TCP was found to be the predominant metabolite in the

initially harvested culture extracts. TMP indicated the O-methylation of TCP. Mass

spectra of TCP, DETP and TMP are shown in Figure 4.23-4.25. The metabolite DETP,

being unstable was not found persistently and disappeared subsequently.

In the TIC of samples extracted after 72 h, peak corresponding to TCP increased

further while TMP peak disappeared completely. Peaks 5 and 6 further reduced and a

new peak 7 appeared at RT 14.89 (Figure 4.26). Analysis of mass spectra of peaks 5, 6

and 7 showed them to be the dechlorination products of TCP and TMP and they were

identified as viz 3,5 dichloropyridine, 3-chloro-2-pyridinol and 3,5-trichloro-2-

methoxypyridine corresponding to m/z 147, 129 and 179 respectively (Figure 4.27, 4.28

and 4.29). Appearance of these three metabolites strengthens the degradation of TCP and

TMP through reductive dechlorination as their structure indicates the removal of one or

two chlorine atoms from TCP and TMP structure.

In the TIC of 120 h (5 days) extracts, a new peak 8 appeared which was not

found in the TICs of previous sample extracts (Figure 4.30). Peak corresponding to CP

disappeared completely and TCP peak was reduced compared to that in the previous day


141

culture TICs. Further, peak 8 corresponding to m/z 117 was identified to be maleamic

acid with a chemical structure indicating a ring cleavage product (Figure 4.31). Hence,

the appearance of a ring cleavage product further supports the capability of

Mesorhizobium sp. HN3 for efficient degradation of CP and its toxic metabolites

containing aromatic rings. On the basis of the above findings, we predicted pathway for

the CP biodegradation by Mesorhizobium sp. HN3 as shown in Figure 4.32.


142

At zero time


containing CP at zero time showing the only peak (peak 1) corresponding to CP at 16.02

min as the most abundant or major peak. No metabolite peaks were found in this TIC.

Retention Time

Peak 1


143

Chlorpyrifos

Figure 4.21 Mass spectrum of chlorpyrifos (CP)

N

Cl

Cl

Cl

O

PO

SO

CH3

CH3


144

After 48 h (2 days)


medium containing CP harvested after 48 h (2 days). Peak 1 is reduced in

abundance while new peaks 2, 3, 4 5 and 6 appeared at different retention times.

Retention Time

Retention Time


145

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

Figure 4.23 Mass spectrum of 3,5,6-trichloro-2-pyridinol (TCP)


146

Diethylthiophosphate (DETP)

Figure 4.24 Mass spectrum of Diethylthiophosphate (DETP)


147

3,5,6-trichloro-2-methoxypyridine (TMP)

Figure 4.25 Mass spectrum of 3,5,6 trichloro-2-methoxypyridine (TMP)


148

After 72 h (3 days of incubation)


medium containing CP harvested after 72 h (3 days). Abundance of peak 1 is

reduced further, of peak 2 increased, of 3, 5 and 6 decreased and peak 4 (which

was present in the previous TIC) is totally disappeared in this. Peak 7 is a new

peak which was not found in previous TICs.

Retention Time


149

3,5-dichloropyridine

Figure 4.27 Mass spectrum of 3,5-dichloropyridine


150

3-chloro-2-pyridinol

Figure 4.28 Mass spectrum of 3-chloro-2-pyridinol


151

3,5-dichloro-2-methoxy Pyridine

Figure 4.29 Mass spectrum of 3,5-trichloro-2-methoxypyridine


152

After120 h (5 days of incubation)


containing CP harvested after 120 h (5 days) of incubation. Abundance of peak 1 is

reduced almost to its complete, of peak 2 decreased, of 3 and 6 decreased and peak 5

(which was present in the previous TIC) is totally disappeared in this TIC. Peak 7 is

larger relative to previous TIC and a new peak, 8 is present at 13.62 that was not found in

previous TICs.

Retention Time


153

Maleamic acid (Ring cleavage product)

Figure 4.31 Mass spectrum of maleamic acid

C

COOH

O

NH2


154

Figure 4.32 Predicted biodegradation pathway of chlorpyrifos


155

4.3.8 Detection of OP degrading genes in Mesorhizobium sp. HN3

Various primers were used for amplifying different OP hydrolases and oxygenases

(Tables 2.3 & 2.4). However, only opdA and protocatechuate (pcaH) genes were

amplified from genomic DNA of CP degrading Mesorhizobium sp. HN3 by using primers

described in Sharaf et al., (2006) and Azhari et al., (2007). PCR products of expected

sizes i.e. 1155 and 390 bp were obtained for opdA and pcaH gene respectively (Figure

4.33 and 4.34). As in case of PFF degrading bacterial isolates (Chapter 3), potential opdA

gene PCR product of expected size was obtained, but further sequence analysis revealed

no similarity with the sequences of any of the previously known hydrolase gene

sequences reported in the GenBank.

However, the sequence of the pcaH gene was found to be 81% identical to the beta

subunit of protocatechuate 3,4- dioxygenase gene of Cupriavidus necator striain

(Accession number JMP131).


156

Figure 4.33 Amplification of opdA gene encoding an OP hydrolase (OPAA) in CP

degrading Mesorhizobium sp. HN3 (Lanes 1 & 2). M indicates the 1Kb marker.

Figure 4.34 Amplification pcaH gene encoding an protocatechuate dioxygenase in CP

degrading Mesorhizobium sp. HN3 (Lanes 1). M indicates the 1Kb marker.


157

4.4 Discussion

This study describes the isolation and characterization of a novel bacterial strain

Mesorhizobium sp. HN3 capable of complete degradation of chlorpyrifos, a chlorinated

organophosphate pesticide. To date many bacterial strains capable of CP degradation

have been reported including a few PGPRs i.e. Pseudomonas sp. (Fulekar and Geetha,

2008), Bacillus sp. (Zhu et al., 2010), Klebseilla sp. (Ghanem et al., 2007). However,

PGPR belonging to Rhizobia group have never been reported to degrade pesticides and

related contaminants.

Initially, the isolation of CP degrading bacteria was considered difficult

potentially due to the accumulation of TCP (an antimicrobial metabolite of CP) in the

culture media and in soil which hindered the enhanced degradation of CP (Racke et al.,

1990). However, in recent years due to the long exposure to CP, microorganisms might

have developed capabilities not only to survive in its presence but also to degrade it

(Singh et al., 2003; Chen et al., 2012). Hence CP as well as TCP degradation potential of

some bacterial isolates has been reported (Maya et al., 2011; Briceno et al., 2012; Chishti

and Arshad, 2013).

Notably, temperature and pH of the soil and water greatly affect the pesticide

degrading activity of the microorganisms (Goda et al. 2010). Different bacterial species

have been reported to show different optimal temperatures for CP degradation (Li et al.

2007; Lu et al., 2013). However, strain HN3 performed well at a range of temperatures

i.e. 30-40°C.

Concerning the effect of pH on CP degradation, the study indicated that all of the

tested pH supported the biodegradation of CP by Mesorhizobium sp. HN3. However, an

efficient degradation of CP was observed at neutral (7.0) and basic (8.0). This was in

contrast to the findings of Racke et al. (1996) who exclaimed that high pH had a co-

relation with hydrolysis of CP in soil and Singh et al. (2003) who demonstrated that high

(basic) pH support microbial hydrolysis of CP.

Mesorhizobium sp. HN3 degraded CP over a wide range of initial concentrations

in contrast with previous reports showing inhibition of bacterial growth and CP

degradation rates at concentrations < 300 mg/l (Singh et al. 2004; Singh et al. 2006; Li et

al. 2007). Kinetic parameters deduced for CP consumption and TCP and bacterial


158

biomass production give a good picture of TCP production as a consequence of CP

degradation. High rate of TCP production and complete TCP removal at relatively lower

initial CP concentrations might be because HN3 would have more easily adapted to lower

TCP concentrations accumulated in the culture media as a result of CP degradation.

Further, higher specific growth rate (µ), cell mass formation rate (QX) and the substrate

uptake rate (QS) at 50-100 mg/l initial CP concentration indicated a remarkable impact of

initial concentration on CP degradation rate. CP degradation by Mesorhizobium sp. HN3

followed first order kinetics with a dramatic decrease in degradation rate with increasing

initial concentration. A similar trend for CP degradation have been reported earlier

(Dubey and Fulekar, 2012).

Efficient degradation of CP by Mesorhizobium sp. HN3 was observed in the

presence and absence of glucose, whereby degradation was higher in former case

indicating co-metabolism of CP. There are few reports showing simultaneous utilization

of CP and glucose by bacterial isolates including Actinobacteria (Briceno et al., 2012)

and Paracoccus sp. TRP (Xu et al., 2008). The present studies showed that

Mesorhizobium sp. HN3 degrades CP better in the presence of an easily available carbon

source and supports the idea that co-metabolism enhances the degradation of recalcitrant

compounds by enhancing the growth of bacteria which in turn enhances utilization of

xenobiotics (De Schrijver and Mot, 1999).

Mesorhizobium sp. HN3 inoculated in the medium deprived of inorganic nitrogen

was capable to scavenge the nitrogen from the pyridine ring of the chlorpyrifos. The lag

phase of the HN3 was relatively longer (48 h) potentially due to the inavailability of the

nutrients and it took time to acclimate to the stressed culture condition after which the

growth started rapidly. As the only nitrogen source available for growth of HN3 in the

culture medium was that in the pyridine ring of CP, the results indicate ring cleavage to

make the nitrogen available for growth. Use of CP as a source of nitrogen by

Acremonium sp. strain GFRC-1 (a fungal isolate) has been reported (Kulshrestha and

Kumari, 2010). However, such data for the bacterial isolates is meagerly available.

Mesorhizobium sp HN3 was also capable of degrading TCP at high

concentrations when provided as a sole source of carbon in MSM. More recently, Lu et

al., (2013) isolated a bacterial strain, DT-1 that can degrade TCP efficiently but


159

increasing the TCP concentration beyond 50 mg/l subsequent decrease in growth was

observed followed by inhibition 100 mg/l. In contrast to this, Mesorhizobium sp. HN3

appears to be more efficient as it can degrade both CP and TCP even at higher

concentrations.

CP and its metabolite TCP contain three chloride atoms on the aromatic ring thus

behaving like an organochlorine and posing a great threat to the environment. For this

reasons, it is important to know the fate of chloride ions during CP and TCP degradation.

Investigation of chloride ions release during biodegradation of CP and TCP further

supported the complete degradation of CP and TCP by Mesorhizobium sp. HN3 whereby

degradation was correlated with the rhythm of chloride ions release in the culture

medium. According to Feng et al., (1998), the TCP mineralization proceeds through

reductive dechlorination under anaerobic conditions. HN3 proves to be one of the best CP

as well as TCP degrading organisms to date that can not only hydrolyze CP but also

release chloride ions and scavenge nitrogen from the aromatic ring of TCP.

CP degradation by Mesorhizobium sp. HN3 was also investigated in soil. A

number of reports indicate the CP remediation in soil by microbial isolates such as

parathion (Barles et al. 1979), ethoprophos (Karpouzas et al., 2005) and chlorpyrifos

(Lakshami et al., 2008). The optimization of bioremediation processes depends on many

factors as the soil properties (moisture contents, soil pH) and the survival and population

of the degrading cultures (Duquenne, et al., 1996). The present study was conducted to

optimize the soil moisture levels and inoculums density of the degrading bacterial

consortium for efficient degradation of CP. Soil moisture contents are very important for

the effective remediation of pesticide contaminated soil as they are one of the key factors

which affect the proliferation of the degrading cultures (Van Veen et al., 1997). The

optimum soil moisture for CP degradation by the bacterial consortium was found to be

40% and an increase or decrease in this moisture produced undesirable results. The

present study was consistent with the previous findings in that longer lag phase was

observed at low initial inoculum density compared to that at higher inoculum densities.

However, inoculation of HN3 in the soil resulted in maximum pesticide degradation both

in sterilized as well as unsterilized soil. The degradation of CP by Mesorhizobium sp.

HN3 was higher in un-sterilized soil than that in sterilized soil. Higher degrading activity


160

of HN3 in the un-sterilized soil can be attributed to the presence of native microbial

communities, some of which might be capable of CP degradation. Therefore, inoculation

of HN3 further enhanced the degradation of chlorpyrifos in un-sterilized soil. Whereas in

sterilized soil, no indigenous microbial population was present therefore, HN3 was alone

to cope with CP contamination. These results signify the importance of optimizing the

inoculums density while studying pollutant degradation.

Mass spectrometric analysis of CP degradation revealed that first step of the

metabolic pathway is hydrolysis of O-P ester linkage to produce TCP and DETP. As

mentioned earlier that samples extracted from the culture media were derivatized with

BSTFA, the trimethyl-silyl portion (m/z 73) attaches to hydroxyl group of TCP and

DETP giving aggregates with m/z 270.619 ~ 271 and 241 for TCP and DETP

respectively. Most of the CP degrading bacteria reported earlier could hydrolyze CP to

TCP by the hydrolysis of O-P bond thus leaving phosphorus atoms to be utilized as a

source of phosphorus by the microorganisms (Singh et al., 2004; Singh and walker 2006;

Chen et al., 2012; Lu et al., 2013). TCP and DETP were not persistently found during the

course of experiments and disappeared subsequently. Singh et al., (2004) suggested that

DETP is utilized as a carbon and energy source by microorganisms for further

degradation of TCP. Moreover, strain HN3 was also capable to grow in phosphorus free

medium containing CP as the only P source (data not shown) indicating utilization of

DETP as P source. Importantly, detection of methylation product of TCP that is TMP is

another discovery about the potential of the strain HN3. Furthermore, subsequent

disappearance of TMP obviously supports degradation of TMP as well.

Identification of dechlorination products viz 3,5 dichloropyridine, 3-chloro-2-

pyridinol and 3,5-trichloro-2-methoxypyridine confirms the degradation of TCP and

TMP because as their structures indicate the removal of chlorine atoms from them.

Further appearance of a ring cleavage product, maleamic acid is a strong evidence of

breakage of pyridine ring and consequently TCP degradation. Moreover, the mass

spectrum of CP contains three peaks corresponding to m/z 314, 258 and 286 were

observed. According to Reddy et al., (2012) peak with m/z 314 corresponds to [M-HCl]

and that with 286 and 258 are obtained after removal of one and two ethylene molecules

form m/z 314 respectively. These findings enabled us to predict a pathway for the CP


161

biodegradation which can be a very good addition to the existing repertoire of knowledge

about biodegradation of chlorpyrifos. Concerning the investigation of degrading genes in

Mesorhizobium sp. HN3, we could only amplify the opdA gene. The variants of opdA

gene have been reported in different bacteria capable of hydrolyzing a variety of

organophosphate pesticides (Horne et al., 2002; Sharaf et al., 2006). However,

identification of protocatechuate dioxygenase (PCD) encoding gene (pcaH) indicate the

capability of the strain HN3 to degrade aromatic compounds as PCD is the key enzyme

involved in the β-keto adipate pathway which is the main pathway for the degradation of

aromatic compounds (Azhari et al., 2007). Therefore, identification of pcaH in the

chlorpyrifos degrading HN3 strain indicates the efficacy of the strain in breaking

aromatic pyridine ring thus degrading the pesticide to harmless products.

Further, genetic mechanism involved in the hydrolysis of chlorpyrifos degrading

bacteria could not be determined conclusively hence it leaves a speculation of some novel

genetic degradation pathway.

Chapter 5 Bio-stimulation

162

Chapter 5

Bio-stimulation: Microbe Assisted Phytoremediation

5.1 Introduction

CP has low water solubility (2 mg/l) and strong affinity for organic matter and

soil particles, hence, its residues remain in the environment for undefined period of time

(DeLorenzo, 2001; Gavrilescu, 2005). The absorption and translocation of CP residues by

wheat and oil seed rape roots and other crop systems have been reported (Wang et al.,

2007) and its residues have also been detected in vegetables, cattle meat and fruits

(Parveen et al., 2004; Mohammad et al., 2010). Exposure and uptake of CP may

adversely affect the plant in terms of its growth and physiology such as delayed

emergence of seedlings (Sinclair et al., 1992) and deformities in fruits (Beck et al.,

1991). The problems associated with CP pose serious concerns regarding the health of

environment as well as inhabitants of the environment which in turn arouse need to pay

utmost attention towards remediation of CP in eco-friendly and cheaper way described

earlier in the context of bioremediation.

Phytoremediation, use of plants for the detoxification of pollutants in the

environment, is accomplishing a significant level of public attention and becoming a

rapidly expanding field owing to its ‘green’ approach, (Aken et al., 2010; Huang et al.,

2011). In this regard, biovailavility and phytotoxicity of the contaminant and

biodegradation/biotransformation capability of the plant are important parameters to be

considered (Wang et al., 2008; Abhilash et al., 2013). However, one of the limitations is

that pollutants adversely affect the plant growth resulting in reduced biomass which in

turn could influence the phytoremediation process (Gaskin et al., 2008; Weyens et al.,

2009). This limitation has been compensated by the combined use of plants and

microorganisms for the remediation of contaminated sites (Khan et al., 2012; Segura et

al., 2009). Term rhizoremediation has been employed for microbe assisted

phytoremediation which involves mutual interactions of plant and rhizospheric

microorganisms exhibiting contaminant degradation activities. More recently, enhanced

pollutant removal by plants in combination with bacterial endophytes capable of

degrading a certain contaminant has been demonstrated as a promising approach to


163

alleviate contaminant induced stress on the plant and increase remediation efficiency

(Kang et al., 2012; Ho et al., 2013).

In some cases, it has been demonstrated that plants can stimulate the pesticide

degradation by microbial communities (Yu et al., 2003; Sun et al., 2004).

Phytoremediation of CP and involvement of associated microbes is also limited to few

studies (Moore et al., 2002; Lee et al., 2012; Dubey and Fulekar, 2012). The present

study describes the inoculation of a chlorpyrifos degrading Mesorhizobium sp. HN3 in

the rhizosphere of ryegrass for enhancing the degradation of the pesticide. Ryegrass is a

good choice to exploit plant-microbe interactions for degradation of contaminants

because of its extensive root system that helps in improving the growth of microbes in its

rhizosphere and in turn the remediation potential of the system is enhanced (Korade and

Fulekar, 2009a). Mesorhizobium sp. HN3 is a CP degrading bacterium isolated and

characterized in our research group (Jabeen et al., 2014). It belongs to plant growth

promoting rhizobia and survives in the plant rhizosphere as well as in the bulk soil.

Moreover, its ability to live as plant endophyte was exploited for the degradation and

removal of CP residues accumulated in plant roots and shoots hence rendering it a good

candidate for the detoxification of CP.

5.2 Materials and Methods

5.2.1 Soil fortification with CP

The experimental soil was without background contamination of pesticides.

Technical grade CP (5% stock solution in acetonitrile) was used to spike 20 g sand and

mixed with 25 % of the experimental soil. The solvent (acetonitrile) was allowed to

disperse and evaporate completely at room temperature for about 24 hours. The spiked

soil was mixed with the rest of the experimental soil to obtain a final concentration of 50

mg/kg (w/w). The experimental soil was the mixture of soil and sand (for aeration,

drainage and storage capacity of water and nutrients).

5.2.2 Bacterial strains used in the study

Mesorhizobium sp. HN3 was employed in this study. E. coli DH5α carrying a

broad host range ampicillin resistant (AmpR) plasmid pBBRIMCS-4 (Kovach et al.,


164

1995) containing yfp (yellow fluorescent protein) cassette was obtained from National

Institute for Biotechnology and Genetic Engineering, NIBGE, Biotechnology Resource

Centre (NBRC), Faisalabad.

5.2.3 Plasmid used for transformation

Plasmid DNA (AmpR, plasmid pBBRIMCS-4 containing yfp cassette) was

isolated from E.coli srain DH5 following Mini-prep standard protocol. The

composition of the solutions used for mini-prep is described in Appendix 11.

E. coli was grown overnight in LB broth containing ampicillin (50 mg/ml) in a

rotary shaker at 37°C and 100 rpm with constant shaking. Culture (5 ml) was transferred

into eppendorff tubes, centrifuged at 8000 rpm for one minute and supernatant was

discarded. The pellet was resuspended in 100 µl ice-cold Solution 1 followed by the

addition of 150 µl Solution 2. The suspension was incubated at room temperature for 5

min and immediately after incubation, 200 µl Solution 3 was added and incubated on ice

for 10 min. The chilled suspension was centrifuged for 5 min at 8000 rpm and the

supernatant was transferred to fresh micro centrifuge tubes.

1 ml of absolute ethanol was added to the supernatant and incubated for 20 min at

-20°C and centrifuged at 14000 rpm for 5 min. Supernatant was discarded and the pellet

was washed twice with 70% ethanol. The washed pellet was dried and re-suspended in 30

µl TE buffer (Tris-EDTA buffer). The plasmid DNA was separated through

electrophoresis using 1% agarose gel.

5.2.4 Preparation of electrocompetent cells of Mesorhizobium sp. HN3

Electrocompetent cells of Mesorhizobium sp. HN3 were prepared following Wu

et al., (2010). 48 h grown culture of Mesorhizobium sp. HN3 was harvested and 5 ml

was used to inoculate a secondary culture in 500 ml flask containing 250 ml LB medium

which was again incubated in rotary shaker at 37°C until the cell density reached

OD600nm=0.6. The culture was transferred aseptically into ice cold 45 ml falcon tubes,

incubated on ice for 30 minutes and centrifuged at 6000 rpm for 10 minutes at 4°C. The

pellet was re-suspended in 45 ml sterile cold 10% glycerol and centrifuged. Cells were

re-suspended in 25 ml 10 % glycerol and centrifuged followed by the re-suspension in 18


165

ml cold 10% glycerol and centrifugation. Finally the cells were re-suspended in 1 ml

filter sterilized cold 10 % (v/v) glycerol and cell aliquots of 200 µl were prepared and

stored at -80ºC.

5.2.5 Electroporation of yfp gene into Mesorhizobium sp. HN3

Frozen electro-competent cells were allowed to thaw on ice. Plasmid DNA

pBBRIMCS-4 conatining yfp was introduced into the Mesorhizobium sp. HN3 by

electroporation following Shahid et al., (2012). For this purpose, an electroporator, Gene

Probe set at a R5 (129 ohm) resistor, 25 µF capacitor and 12.5 KV/cm field strength, was

used. Desired pulse length was 5-6 msec. Mixture of HN3 cells (200 µl) and plasmid

DNA (1 µl = 0.1 µg) was transferred into the electroporation cuvette. Following a short

electric pulse, 1 ml LB medium was mixed gently to the transformation mixture; the

whole mixture was transferred into a fresh 1.5 ml eppendorff tube and incubated at 37°C

for 45 minutes with constant shaking.

The transformed cells were screened on LB-ampicillin (50 µg/µl) agar plates and

the transformed colonies were confirmed under Confocal Laser Scanning Microscope

(CLSM) at 530 nm. The transformed bacterium was named as Mesorhizobium sp.

HN3yfp. Inoculum of HN3yfp (10

7 CFU/ml) was prepared as described in Chapter 2

Section 2.10 and used for CP remediation studies.

5.2.6 Experimental design

Ryegrass (RG, Lolium multiflorum var Taurus) previously reported to tolerate CP

(Korade and Fulekar, 2009a; Ahmad et al., 2012) was used in these studies. For the

experiment, the plastic pots (1.5 kg soil each) were filled with agricultural soil spiked

with CP (50 mg/kg). The study included following treatments:

1. CP contaminated soil (Soil+CP)

2. CP contaminated soil inoculated with HN3yfp (Soil+CP+HN3yfp).

3. Ryegrass planted in un-contaminated soil (Soil+RG)

4. Ryegrass planted in CP contaminated soil (Soil+RG+CP)

5. Ryegrass planted in CP contaminated soil and inoculated with HN3yfp

(Soil+RG+CP+HN3yfp)


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The seeds of the Lolium multiflorum were surface sterilized, soaked in bacterial

suspension (except for the un-inoculated treatments) and finally sown in each pot (150

seeds/pot) containing CP spiked soil. For the inoculated treatments, the soil was mixed

with 50 ml bacterial suspension (3.3×105 CFU/g of soil) and with 0.85% NaCl for the un-

inoculated control treatments before sowing. The plants were grown in green house at a

temperature of 25±2°C with 16 h light and 8 h dark. The pots were watered when needed

for 45 days. One week following seed germination, seedlings were counted, poor

emerging were removed and 100 plants were maintained per pot. Plants were harvested

after 15, 30 and 45 days of experiment and shoots were cut 2 cm above ground, roots

were separated from the bulk soil and the soil from each pot was mixed thoroughly to get

homogenized samples for CP residue analysis. CP concentration in the rhizosphere of

ryegrass, bulk soil and within plant tissues, plant growth parameters, bacterial population

in soil and roots were observed. Whole experiment was performed in triplicates.

5.2.7 Extraction and analysis of chlorpyrifos residues in the soil and plant

CP and TCP residues were extracted from rhizospheric (planted) and bulk (un-

planted) soil following the protocol described in chapter 2 Section 2.12.4 for extraction of

soil from the soil. CP residues were extracted from residues from roots and shoots

following Ahmad et al., (2012). The protocol is described in Appendix 12. The

estimation of CP and TCP in soil, root and shoot samples was carried out using HPLC as

described in Chapter 4 Sections 4.2.4.

5.2.8 Detection and enumeration of the bacteria in the soil

Rhizospheric soil was collected by removing soil adhered to roots and suspended

in 10 ml 0.85 % saline solution. The suspension was agitated for 1h at 37°C, the soil

particles were allowed to settle down and 10 fold dilutions were prepared. Same

procedure was adopted for bulk soil. By spreading these dilutions on LB-CP-ampicillin

(50 µg/ml) agar plates and incubating, HN3yfp colonies were counted. Ampicillin

resistance is the result of an enzyme, beta lactamase which breaks down the ampicillin.

Ampicillin resistant gene usually serves as a useful selectable marker and bacteria that

are subjected to introduce a foreign DNA into a cell are grown on the medium containing


167

ampicillin. The bacterial colonies which successfully grow on the ampicillin containing

medium are taken up and selected for the expression of the introduced DNA/gene. The

colonies morphologically similar to Mesorhizobium sp. HN3yfp were further confirmed

by restriction fragment length polymorphism (RFLP) analysis of 16S-23S rDNA

intergenic spacer region (IGS) using genomic DNA of the randomly selected bacterial

colonies as described earlier (Chapter 3 Section 3.2.2).

5.2.9 Root and shoot colonization by Mesorhizobium sp. HN3yfp

For observing colonization of Mesorhizobium sp. HN3yfp in the plant tissues, 15,

30 and 45 days old roots and shoots of the ryegrass were washed separately with sterile

distilled water as soon as they were harvested. The washed root and shoot samples were

examined under CLSM (Olympus Fluoview Version 1.3) at 4X and 10X magnifications

for observing the colonization of Mesorhizobium sp. HN3yfp within ryegrass root hairs,

root and shoot surfaces and inside the tissues.

1. Measurement of growth parameters

Plant growth parameters measured were length and weights (fresh and dry) of root

and shoot at each harvest. Elongation of roots and shoots was measured using a ruler.

Fresh weights were measured by directly weighing the freshly separated and cleaned

roots and shoots with a physical balance. For assessing the dry weights, the roots and

shoots were dried in oven at 65°C for 8-12 hours until a constant weight was achieved.

5.2.11 Phosphate solubilization

Capability of the chlorpyrifos degrading isolate HN3 to solubilize phosphate was

also explored to check its potential in plant growth promotion. For this purpose,

Pikovskaia (1948) medium was prepared with Pikovskaia’s agar and tri-calcium

phosphate as insoluble phosphate. Overnight grown bacterial culture of LB media (10 µl)

was spotted onto the plates containing the above media. The plates were incubated at

37°C until a clear zone appeared around the culture spots. This clear zone was considered

as a sign of positive result for phosphate solubilization.


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5.2.12 Indoleacetic acid production

A colorimetric assay was performed for indoleacetic acid (IAA) production from

Mesorhizobium sp. HN3 (Gordon and Weber, 1951). For this purpose a fully grown

bacterial culture (100 µl) was transferred into eppendorf vials containing LB (200 µl)

supplemented with 100 mg/l tryptophane (as a precursor of IAA) in three replicates.

Eppendorf vials were incubated at 30°C without shaking. The incubated cultures were

mixed with Salkowski reagent (Appendix 13) in 96-well microtitre plate. IAA standard

(100 µl) was mixed with the same volume of Salkowski reagent. The isolates were

observed instantly for the development of pink, purple or purplish pink color after mixing

for 30 minutes. Quantitative estimation of IAA was carried out following Tien et al.,

(1979). Bacterial culture was grown for 7 days in LB- broth supplemented with

tryptophan (100 mg/l). Cells were harvested at 8000 rpm, culture supernatant was

acidified with hydrochloric acid (to get a pH 2.8) and extracted twice with an equal

volume of ethyl acetate. The extracts were collected, dried, re-suspended in ethanol and

analyzed on HPLC.

5.2.13 Data analysis

Statistical analyses of plant biomass and CP & TCP degradation in soil, roots and

shoots were performed on three replicates of data obtained from all treatments. Standard

error and the significance of differences were treated statistically by the ANOVA and

evaluated by post hoc comparison of means using Tukey’s test in Statistica 6.0 software.

The degradation rate constants and theoretical half-life values (DT50) of CP

degradation in soil and were determined using algorithms Ct/C0= e-kt

and from the linear

regression equation between ln(Ct/C0) and time as already described in previous chapters.


169

5.3 Results

5.3.1 Biodegradation of CP in the planted and un-planted soil

The residual concentration of the CP was determined to assess the effect of

ryegrass and Mesorhizobium sp. HN3yfp partnership on CP degradation in contaminated

soil. CP degradation and TCP accumulation and removal were compared between three

treatment soils i.e., planted (un-inoculated), un-planted (inoculated), and

planted+inoculated soil (Figures 5.1A & B). After 15 days of sowing, 22% of the added

CP was degraded in the planted soil (un-inoculated) whereas in the inoculated (un-

planted) soil, 36% degradation was achieved. On the other hand when the planted soil

was inoculated with Mesorhizobium sp. HN3yfp, 44% of the total applied pesticide was

degraded. At the end of the experiment (45 days), the CP degradation was 79% and 91%

in the planted (un-inoculated) and inoculated (un-planted) soils respectively whereas

complete degradation was achieved in the planted+inoculated soil as no residual pesticide

was detected in the soil samples (planted+inoculated soil) . Hence inoculation of

Mesorhizobium sp. HN3 in the planted soils significantly enhanced the removal of CP as

compared to un-inoculated soils. Furthermore, kinetic analysis (Table 5.1) showed that

higher degradation rate and lower half-life values of CP were achieved in the plant-

bacterial system as compared to those containing plants or bacteria solely.

Regarding the accumulation and degradation of TCP, at the first harvest (15 days

of the experiment) 9 and 10 mg/kg TCP was observed in the solely planted and solely

inoculated soils respectively which continued to increase upto 30 days and then declined

towards the end of experiment leaving 11 and 7 mg/kg TCP. Whereas, higher TCP

concentration (15 mg/kg) was found in the planted+inoculated soil at the first harvest

which further indicate higher CP degradation in this treatment. However, it decreased

gradually and more rapidly as compared to other treatments leaving only 3 mg/kg after 45

days.


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Figure 5.1 Degradation of CP (A) and accumulation and subsequent disappearance of

TCP (B) as a result of CP hydrolysis by ryegrass (Lolium multiflorum) and

Mesorhizobium sp. HN3 in different treatment soils at different time ntervals. Each value

is the mean of three replicates with error bars representing the standard error

B

A


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Table 5.1: First order kinetics parameters of chlorpyrifos degradation in planted

and un-planted soils.

Where, k = First order rate constant for degradation

C0 = Initial CP concentration

Ct = CP concentration at time (t)

t1/2 = Half-life of CP

R2 = Regression coefficient

Treatments Regression equation R2 k (Day

-1) t1/2(Days)

Control ln(Ct/C0)= -0.0042x+3.915 0.995 0.0042 165

Planted ln(Ct/C0)= -0.0353x+4.038 0.963 0.0353 20.5

Inoculated ln(Ct/C0)= -0.0532x+4.094 0.956 0.0532 13

Planted+inoculated ln(Ct/C0)= -0.0858x+4.291 0.929 0.0858 8


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5.3.2 Chlorpyrifos uptake by plant

Residual concentrations of chlorpyrifos in vegetative parts (root and shoot) of

ryegrass indicating uptake of the pesticides by the plant are presented in Table 5.2. Plants

vegetated in un-inoculated soil exhibited significantly higher concentration of CP in roots

and shoots as compared to plants vegetated in HN3yfp inoculated soil.

Table 5.2: CP uptake and accumulation in roots and shoots of ryegrass.

Values are the means of three replicates for CP uptake and followed by the standard error

(in the parentheses). For control no pesticide was added to the soil.

Treatments CP uptake (µg/g plant dry mass)

Roots Shoots

50 mg/kg

Control

(Un-inoculated) 0.41 (0.03) 0.22 (0.025)

Inoculated

(HN3yfp

) 0.13 (0.04) Not found


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5.3.3 Colonization of Mesorhizobium sp. HN3 in soil

In general, Mesorhizobium sp. HN3yfp population declined with time in the un-

planted or bulk soil. However, in the planted or rhizospheric soil, the inoculant

proliferated above the initially inoculated level and reached 106 CFU/g (Table 5.3).

However, the population declined towards the end of experiment. In sum, total

population of bacteria was higher in the rhizospheric soil than in the bulk soil throughout

the course of experiment.

5.3.4 Colonization of Mesorhizobium sp. HN3 in the roots and shoots ryegrass

Use of fluorescently (yfp) tagged Mesorhizobium sp. HN3 and CLSM enabled to

demonstrate that HN3yfp actively colonized the ryegrass roots. The entire colonization

process i.e. starting from the attachment of bacteria with root hairs and lateral roots to

invasion into the internal tissues of the roots was observed. At each harvest, fluorescence

denoted the presence of bacterial aggregates on root hairs and the junctions between the

primary and lateral root surfaces indicating the possible points of entry into the ryegrass

roots. Maximum colonization was observed on the lateral root surfaces, root tips and the

zone of elongation and differentiation (Figure 5.2-5.4). Fluorescence was not detected in

un-inoculated control. These results confirmed the colonization of Mesorhizobium sp.

HN3 inside ryegrass roots rendering it active endophyte


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Figure 5.2 CLSM images of Mesorhizobium sp. HN3yfp colonization in ryegrass roots

after 15 days of inoculation. (A) In vitro grown cells at 100X (B) time course

colonization process of Lolium multiflorum (ryegrass) roots by yfp-tagged

Mesorhizobium sp. HN3 to show bacterial colonization on root tip surfaces, (C) inside the

lateral roots and root hairs. Arrows indicate the aggregates of bacterial cells (bright

green) attached inside or outside the roots surfaces, the root tip and root hairs.


175


multiflorum (ryegrass) roots by Mesorhizobium sp. HN3yfp after 30 days of inoculation

to show bacterial colonization on lateral roots (A, C), inside the roots (B) and inside the

root tip and root hairs (D). Arrows indicate the aggregates of bacterial cells (bright green)

attached inside or outside the roots surfaces, the root tip and root hairs


176


multiflorum (ryegrass) roots by Mesorhizobium sp. HN3yfp after 45 days of inoculation

to show bacterial colonization inside the root hairs (A), inside and outside the root

surfaces and root tip (B,C,D). Arrows indicate the aggregates of bacterial cells (bright

green) attached inside or outside the roots surfaces, the root tip and root hairs.


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Table 5.3: Colonization of Mesorhizobium sp. HN3yfp in planted (ryegrass) and un-

planted soil

Each value is a mean of three replicates.

Treatments CFU/g soil

Zero day 15 Days 30 Days 45 Days

50 mg/kg

Control (inoculated with HN3yfp)

3.3105

2.1105

1.4104

4.9103

Ryegrass and inoculated (HN3yfp) 3.3105 8.510

5 6.610

6 2.110

5


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5.3.5 Plant biomass

Contamination of soil with CP drastically decreased plant growth as indicated by

various growth parameters of plant such as root length (RL), shoot length (SL), root &

shoot fresh weights (RFW & SFW) and root & shoot dry weights (RDW & SDW). A

significant decline was observed in RL and SL of ryegrass planted in the CP

contaminated soil, however, both the parameters increased in HN3yfp inoculated soil.

Similarly a significant decrease in the RFW and SFW was observed in the CP

contaminated soil as compared to un-contaminated soil. When CP contaminated soil was

inoculated with HN3yfp significantly (P<0.05) higher RFW and SFW were observed

(Table 5.4). Root and shoot dry weights corresponded with the RFW and SFW of

ryegrass in all the treatments. Difference in the root and shoot length of different

treatment plants is shown in Figure 5.5.


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Figure 5.5 A comparison of shoot lengths (A) and root lengths (B) among

different treatments of experiment: control ryegrass plant (a), ryegrass plant

grown in CP contaminated soil (b) and ryegrass plants grown in CP contaminated

soil inoculated with Mesorhizobium sp. HN3 (c).

a c b A

c b a B


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Table 5.4: Effect of chlorpyrifos on plant growth parameters {root length (RL), shoot

length (SL), root fresh weight (RFW), shoot fresh weight (SFW), root dry weight (RDW)

and shoot dry weight (SDW)}

Each value is mean of three replicates. Means in the same column followed by the

different letters are significantly different at a 5 % level of significance. Standard error of

three replicates is presented in parenthesis.

Sampling

days Treatments RL (cm) SL (cm) RFW (g) SFW (g) RDW (g) SDW (g)

15 Ryegrass only

8.20e(0.1) 8.39e(0.39) 3.00f(0.05) 3.70f(0.06) 0.52ef(0.03) 0.65e(0.02)

CP+ Ryegrass

7.75e(0.25) 7.50e(0.50) 2.56f (0.170) 2.87f (0.09) 0.45e(0.01) 0.51e(0.03)

CP+HN3 yfp +Ryegrass

9.87de(0.07) 9.59e(0.185) 3.32ef (0.31) 4.32f (0.03) 0.62f (0.04) 0.81f (0.04)

30 Ryegrass only 11.50cd(0.5) 17.5d(0.50) 7.44de(0.26) 15.00e(0.50) 1.23e(0.00) 2.63de(0.11)

CP+ Ryegrass 10.25b(0.25) 14.65c(0.35) 6.67cd(0.45) 13.50d(0.50) 1.19cd(0.20) 2.41de(0.01)

CP+HN3yfp+Ryegrass 15.25b(0.95) 19.90bc(0.90) 9.38c(0.56) 20.00d(1.00) 1.76d(0.15) 3.76d(1.00)

45 Ryegrass only 16.60bc(0.5) 30.00b(2.0) 17.15a(1.15) 30.00c(0.50) 3.01b(0.44) 5.27c(0.40)

CP+ Ryegrass 13.27a(1.23) 25.50a(0.50) 14.05b(0.25) 26.96b(0.15) 2.51a(0.05) 4.82b(0.50)

CP+HN3 yfp +Ryegrass 20.83a(0.47) 34.50a(0.50) 18.62a(0.19) 35.75a(0.45) 3.49c(0.15) 6.72a(1.00)


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5.3.6 Plant growth promoting properties of Mesorhizobium strain HN3

5.3.6.1 Indoleacetic acid production

Indoleacetic acid (IAA) production by the Mesorhizobium strain HN3 was recorded upto

5.34 pp. Moreover, colorimetric method indicated the color change in the medium from

colorless to purplish pink which was very close to the color of the standard solution

(Figure 5.6).

5.3.6.2 Phosphate solubilization

A clear zone was formed by solubilizing tricalcium phosphate on the Pikoviskaya

medium plates inoculated with Mesorhizobium sp HN3 which indicates the high

capability of the strain HN3 to solubilize phosphate (Figure 5.7).

Ability of Mesorhizobium sp. HN3 to solubilize phosphate and IAA production

along with CP degrading potential demonstrate its high ability to promote plant growth

which would render this strain a very good tool in plant growth promotion in the polluted

soils.

Clear zone of Phosphate -

solubilization

Clear zone of P-solubilization

Figure 5.7 Qualitative test of phosphate

solubilization by Mesorhizobium sp. HN3 on

Pikoviskaya medium

Figure 5.6 Qualitative test of IAA production by

Mesorhizobium sp. HN3 using Ferric chloride.

A). Standard solution for IAA. B). IAA

production by Mesorhizobium sp. HN3


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5.4 Discussion

Bioremediation of toxic pollutants employing bacteria that possess pollutant

removing capability towards a given contaminant is generally considered as a safe and

advantageous technology. However, sometimes bacteria cannot successfully thrive in the

contaminated sites. In such cases bio-stimulation i.e. use of nutrients and electron

acceptors might help bacteria to survive/multiply and hence enhance bioremediation

process. Although, phytoremediation (use of green plants for the pollutants remediation)

is emerging as a promising technology due to its low cost and minimal environmental

disturbances, however, use of plants along with microorganisms (also referred to as

phyto-stimulation) is a contemporary approach for enhancing the removal of

environmental contaminants (Segura et al., 2009; Weyens et al., 2009).

Inoculation of organic pollutant degrading bacteria in the plant rhizosphere has

recently been reported as a useful strategy for bioremediation (McGuinness and Dowling,

2009). Among other plants, ryegrass (Lolium multiflorum) has been revealed as a good

candidate for such studies because of its extensive root system and capability to support

the microorganisms in its rhizosphere which in turn enhance pollutant degradation.

In the current study suitability of plant-bacteria partnership using a CP degrading

Mesorhizobium sp. HN3yfp and ryegrass plant for the remediation of CP contaminated

soil was investigated. A comparative study of CP degradation in the rhizospheric and

bulk soil (planted and un-planted soil respectively) was carried out. Lower rates of CP

and TCP degradation were observed in soils solely planted with ryegrass or solely

inoculated with HN3yfp whereas the rates increased to maximum following the combined

use of the two. Higher rate of bacterial degradation of CP in the rhizospheric soil as

compared to that in the bulk soil displays the importance of plant root exudates (vitamins,

amino acids and other nutrients) that can nourish microbes in the rhizosphere and also

induce biochemical pathways which in turn display the enhanced CP degradation. It has

already been reported that even though many endophytic bacteria are capable of living in

bulk soil, their pollutant degradation efficiency is higher in the rhizospheric soil because

they need plant exudates to boost up the pollutant degrading enzyme activities (Leigh et

al., 2002).


183

Relatively low concentrations of CP in the roots of the inoculated plants were

observed as compared to the un-inoculated plants. Very interestingly, in the presence of

HN3yfp, no CP was detected in the shoots as compared to the shoots of un-inoculated

plants. A possible explanation for this finding could be the extensive colonization of the

Mesorhizobium sp. inside the roots of ryegrass which would be responsible for maximum

degradation of CP accumulated within the roots and hence no CP could move towards

shoots.

Colonization studies of HN3yfp in the soil and inside the plant revealed that

HN3yfp colonized both in the rhizosphere and inside the roots of ryegrass which is not

surprising because most of the endophytes have been reported as capable of surviving

outside their host plant (Di Fiori and Del Gallo, 1995). Higher proliferation of HN3yfp in

the rhizospheric soil as compared to that in the bulk soil can be attributed to the fact that

plant roots release exudates that can nourish microbes in the rhizosphere (Leigh et al.,

2002). Moreover, root exudates also stimulate chemo tactic movements in bacterial cells.

Further, a decline in the rhizospheric population of the inoculant at the end of the

experiment could mean that HN3yfp tended to move into the plant roots from rhizosphere

and built up its population inside the plant. It also supports the previously established

view that endophytic pollutant degrading bacteria get more abundant inside the plant

tissues in contaminated sites (Siciliano et al., 2001) and that endophytic bacteria existing

in the rhizosphere make their path towards internal plant parts (Sturz et al., 2000; Prieto

et al., 2011).

The inoculants, HN3yfp was found densely populated on the root surface and

inside roots including zone of differentiation and elongation, inside the tip, the root hairs

and other root parts. Although, bacterial colonization inside root hairs has been scarcely

reported, in the present studies, ryegrass root hairs were intensely colonized by HN3yfp.

Root hairs can be regarded as the possible entry points for the inoculants. Shoot

colonization of the inoculants was not observed which is in harmony with the fact that

high densities of endophytes are generally observed in the roots and decrease from stem

to the leaves (Prieto et al 2011; Moore et al., 2006; Weyens et al., 2012).

Although, numerous reports indicate the role of rhizospheric bacteria in

improving the phytoremediation of pollutants (Gerhardt et al., 2009; Afzal et al., 2011),


184

effective use of endophytic bacteria to improve phytoremediation of organic

contaminants is a relatively new concept (Germaine et al., 2006; Ho et al., 2012, 2013).

The current results compliment to some recent findings that phytoremediation of

aromatic contaminants can be enhanced by inoculation of functional endophytic bacterial

species.

Root and shoot growth decreased in the CP contaminated soil as compared to the

un-contaminated soil which might be attributed to the toxic effects of CP and TCP.

Pesticides are known to inhibit the plant growth by disrupting the synthesis of DNA and

proteins in the cells (Sinclair et al., 1992; Pereira et al., 2010). Reduction in the seed

germination of ryegrass with increasing the CP concentration has been reported

previously (Korade and Fulekar, 2009b) and attributed to different kinds of impairments

associated with the cell division apparatus. It was further explained that the CP induced

toxicity in cell division stages ultimately induces abnormalities in the seedlings hence

seed germination is adversely affected.

An increase in the plant growth parameters in contaminated soil in the presence of

inoculant, Mesorhizobium sp. HN3yfp can be explained by the fact that the inoculant is an

efficient degrader of CP and TCP. We believe that due to degradation of CP and TCP by

the HN3yfp, toxic effects of the pesticide were alleviated and hence root and shoot

growth was not hindered. Microorganisms (rhizospheric or endophytic) are known to

play a central role in conferring tolerance to plant against specific xenobiotics (Siciliano

et al., 2001). Moreover, HN3yfp belongs to PGPR owing to IAA production and P-

solubilization activity and hence can impart its role in enhancing ryegrass growth.

Proliferation of the bacterium in the rhizosphere and within the roots of ryegrass also

supports the fact that HN3yfp might have enhanced the plant growth owing to its plant

growth promoting activities. Our findings are in line with the previous studies showing

the importance of PGPR inoculation in relieving the contaminant stress in the plant

environment (Glick, 2003).

Here we conclude that Mesorhizobium sp. HN3 used in the present study is

interestingly an example of facultative endophyte that can also colonize the rhizosphere

and efficiently degrades CP in the rhizosphere and within the roots. A few endophytes of

Lolium multiflorum have been reported that settle in its roots and shoots and assist in


185

degrading hydrocarbons (Yousaf et al., 2010), Mesorhizobium sp. as a root endophyte of

this plant has never been reported. Moreover, colonization of the pesticide degrading

endophytic bacteria inside the ryegrass roots has not been reported earlier.

Mesorhizobium sp. HN3 hindered the translocation of the pesticide inside plant parts and

hence can potentially be employed for controlling CP movement into plant tissues.

Chapter 6 General Discussion

186

Chapter 6

General discussion

Aim of thesis

The pesticides entered into the environment as the beneficial tools to cope with

agricultural problems. They resulted in a great revolution in agricultural products as they

control the insect vector of different diseases, pests of plants and crops thus improving

the crop protection. However, extensive applications of toxic chemical pesticides played

havoc with human and other life forms as their residues persist in the environment for the

variable period of time. These toxic pesticide residues significantly disturb the ecosystem

by affecting the non target micro & macro flora and fauna. Initially organochlorines

(OCs) were introduced and they were found effective for pest control but appeared to be

very much persistent as well as toxic for non target aquatic and terrestrial life and hence

had a negative impact on the environment. To combat this problem, OPs, carbamates and

pyrethroids were introduced. They were considered less persistent and less toxic than

OCs. However, later on, the researchers found that OP, carbamates and pyrethroids also

had the toxic impact on disturbing the ecological balance and health of humans and

animals (Reviewed in chapter 1).

In the recent years, the use of organophosphate pesticides and their metabolites

have been recognized as an emerging worldwide problem and their impacts are now

becoming a subject of wider scientific and the social interest. Previous reports have

shown that microorganisms are the key players that work in the diverse range of the

environment and can survive against environmental fluctuations. Repeated applications

of a same pesticide at a particular site help microorganisms adapt capabilities to utilize

the pesticide as a source of carbon and energy. This phenomenon is called “enhanced

biodegradation” which has already been discussed in chapter 1. Hence this capability of

the microorganism is exploited for the benefit of the mankind, wildlife and also for

reducing the xenobiotic stress on the environment.

Therefore, the exploitation of bacterial potential to degrade pesticides was a main

gizmo of this study. Microbial degradation studies were established to investigate the

complete degradation of two model organophosphate pesticides, profenofos and

chlorpyrifos (Chapter 3, 4) by indigenously isolated bacterial strains. Various


187

environmental factors were optimized for maximum degradation of the two pesticides.

An attempt was made to elucidate the biochemical pathway for both the pesticides. The

main purpose of our study was to achieve systematic understanding of the metabolism of

pesticides by microorganisms and to develop methods for accelerating these metabolic

processes for the bioremediation of contaminated soils and groundwater. Moreover, it

was desired 1) to gain scientific understanding of in situ bioremediation by performing

laboratory and field research on biotransformation and biodegradation of pesticides, and

2) accelerating in situ biodegradation by bio augmentation and bio stimulation processes.

Main findings

The bacterial consortium PBAC

While screening the bacterial strains for the degradation of profenofos, an

efficient bacterial consortium PBAC was obtained. PBAC was found to degrade

profenofos, containing halogenated aromatic moiety, within a week upto 300 mg/L. Its

ability was also checked to remediate the profenofos contaminated soil at lab scale and it

worked remarkably to degrade profenofos as well as its toxic metabolite, BCP which is

rarely reported previously. Its potential was also studied to degrade other pesticides of the

same chemical class and others (Pyrethroids) as well. Surprisingly, it was capable of

degrading all the tested pesticide which is a significant aspect of this bacterial

consortium. Hence this consortium will prove to be a potential candidate for the removal

of organophosphate pesticides from the liquid culture, soil and sediments.

Response surface methodology (RSM) was employed for the optimization of

different environmental factors for degradation of profenofos by PBAC. It was found to

be a useful tool to elucidate the different culture conditions as it enabled us to study three

variables at a time with changing two variables and keeping one constant thus developing

a polynomial equation. This equation can predict % degradation of profenofos for

different levels of variables and can also be helpful for the future researchers and

scientists to predict % degradation for profenofos. RSM is a three-dimentional (3-D)

graphical representation of the data which looks beautiful enough to capture the attention

of viewers and develop an immediate understanding of the trend of the response (%

biodegradation in this case). Another significant aspect of this strategy is that it shifted


188

the trend of laborious and tedious one-factor-at a time approach towards the study of

multiple factors at a time. In this way it is a time effective approach with maximum

understanding of interaction of multiple variants on the response.

Mesorhizobium sp. HN3, a novel strain

Another key to the thesis was the achievement of a novel chlorpyrifos degrading

Mesorhizobium sp. HN3 which is a rhizobial strain (dwelling in the rhizo-sphere of the

plant). The novelty of the strain lies in the fact that it was a first report of a rhizobial

degradation of xenobiotics, importantly the pesticides. It presented a high capability of

chlorpyrifos degradation at high concentrations. The studies to optimize different culture

conditions depicted the marvellous efficacy of the Mesorhizobium sp. HN3 to tolerate

wider range of the media pH, temperature, inoculum density and different initial pesticide

concentrations. Kinetic analysis of the CP degradation by Mesorhizobium sp. HN3

revealed that this bacterium was helpful to degrade not only the parent compound but its

subsequent toxic hydrolysis metabolite, TCP which had long been known to be a main

culprit causing resistance to CP degradation (Racke et al., 1990). Luckily this strain was

useful to utilize TCP as a carbon source in the presence /or absence of a readily available

carbon source, glucose. This happened to be another important finding of this study.

Bio stimulation of Mesorhizobium sp. HN3 by ryegrass

Mesorhizobium sp. HN3 being a rhizobial strain, was believed (also investigated

in this study) to harbour plant growth promoting properties (PGPR) such as phosphate

solubilisation, indoleacetic acid (IAA) production. These characteristics of PGPR

bacteria are known to help in improving plant growth. Keeping in view these PGPR

properties of the strain (its ability to reside in the rhizosphere of the plant) and its

potential to degrade chlorpyrifos, a microbe assisted phytoremdiation system was

developed using yfp-tagged variant of the strain HN3. The aim of this technology was to

assess the enhanced degradation of chlorpyrifos in the rhizosphere and different parts of

the model plant (ryegrass). yfp tagging of the Mesorhizobium sp. HN3 enabled us to get

visual proof (as confirmed by the CLSM) of the colonization of the HN3 inside the

ryegrass roots. Further, extraction of CP residues from ryegrass roots and shoots and


189

HPLC analysis provided the evidence of Mesorhizobium sp. HN3 being an efficient

isolate to alleviate the stress on the plant induced by the chlorpyrifos (chapter 5). This

was a remarkable on site preliminary application of the Mesorhizobium sp. HN3. In this

way it appeared to be an exclusively new approach for remediation of pesticide

contaminated soil.

Metabolic pathways

During the bioremediation process, it is important to investigate the fate of

xenobiotics in the environment. To achieve this goal, it was important to elucidate the

biochemical pathways of the two pesticides in question (profenofos and chlorpyrifos) by

their respective isolates. GC-MS was employed for metabolites analysis in a time course

fashion. Successfully both the pesticides were found to be degraded to potentially

harmless products (results discussed in chapter 3 and 4). This finding enabled us to

demonstrate the fate of the two pesticides which can enable the future researchers to

investigate the genes/enzymes involved in the different steps/chemical reactions like

hydrolysis, dehalogenation, oxygenation and ring cleavage of the xenobiotics.

However, in this study, we attempted to find genes involved in different chemical

reactions during the biodegradation of chlorpyrifos and profenofos. Unfortunately no

success was achieved in this area of the study. Although, the primers were designed

based on the previously reported OP degrading genes but their sequences did not show

homology to previously reported OP degrading gene sequences present in the NCBI

database. This indicates that the present isolates might have some novel genes which

need further investigation.

Future recommendations specific to this thesis

1. Microbe assisted phyto-remediation system comprising profenofos degrading

consortium PBAC and ryegrass as model plant can be employed to check the

efficacy of the PBAC for the degradation of profenofos in a way as

determined for the remediation of chlorpyrifos by Mesorhizobium sp. HN3.

Moreover, this strategy can also be applied using other grasses and crops such

as wheat, cotton and rice where the two pesticides are commonly sprayed.


190

2. On site remediation of profenofos and chlorpyrifos using bacterial consortium

PBAC and Mesorhizobium sp. HN3 respectively in the contaminated soils

(micro plots, field trials) under different environmental conditions can be

carried out. Using this technology, it will be possible to get rid of

contaminating pesticides and hence improving soil quality without any

environmental risks. The process will also be beneficial to farmers interested

in organic farming as it will make it possible to get rid of the pesticide

residues in their lands and hence the products. This is an important step to

meet the quality standards and increasing the export of agricultural and other

products by minimizing the pesticide residues.

3. Identification of unknown metabolites of the chlorpyrifos and profenofos

should be given attention as in the current study, in the total ion

chromatograms of culture extracts of the two pesticides; some peaks were

observed which remained unknown. The investigation of these unknown

metabolites would provide an insight into the metabolic pathways of the

pesticides through GCMS, LCMS or MS/MS.

4. As the genes and enzymes involved in the degradation remained un-identified,

hence study can be planned to solely throw the light on the investigation of

OP degrading genes and enzymes of the isolates of this study. Identification of

genes would help in identifying the enzymes. Based on the concept of

enhanced biodegradation, a comparison can be made between induced and

non induced bacterial cultures using protein expression assays.

Chapter 7 References

191

Chapter 7

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Appendices

Appendix 1

Minimal Salt Media (MSM) preparation

Component Composition (g/l)

Na2 H PO4 5.8 g/l

KH2PO4 3.0 g/l

NaCl 0.5 g/l

NH4Cl 1.0 g/l

MgSO4.7H2O 0.25 g/l

pH 6.8-7.00

Appendix 2

Luria Bertani medium

Appendix 3

Preparation of standard stock solutions

Standard stock solutions of chlorpyrifos, TCP, profenofos and BCP (1%) were prepared

in acetonitrile/methanol. The stocks were mixed well, filter sterilized and stored at 4°C

until used. Stock solutions of antibiotics (ampicillin, kanamycin), IPTG and X-gal etc

were prepared and stored at -20°C until used.


Trypton 10 g/l

Yeast extract 5 g/l

NaCl 5 g/l

pH 7.2±0.2

Appendices

Appendix 4

Frozen Storage Buffer (FSB)

Before preparing FSB, prepare 1M KCO2CH3:

Dissolve 9.8 g KCO2CH3 into 80 ml water. Adjust pH to 7.5 with KOH. Make

up final volume to 100 ml with water. Autoclave and store at 4˚C.

So the composition of FSB is as follow

Dispense into bottles and autoclave. Store at 4˚C.

Appendix 5

Solutions for Gram’s reaction

Appendix 5.1

Crystal violet solution

Component Amount to be used for 400 ml

Crystal violet 10g

Ammonium oxalate 4 g

Ethanol 100 ml

Component Amount to be used for 1 litre

KCl 7.4 g

CaCl2-2H2O 7.5 g

Glycerol 100 ml

1M KCO2CH3 10 ml

pH 6.2

Appendices

Appendix 5.2

Iodine solution


Iodine 1g

Potassium iodide 2 g

Ethanol 10 ml

Appendix 5.3

Safranin solution


safranin 2.5g

Ethanol 10 ml

Appendix 6

Saline solution (0.9%)

Appendix 7

Mobil phase for used HPLC


Acetonitrile 800 ml

Water(deionized) 200 ml

Acetic acid 2.5 ml

Sonicated and used

for the HPLC.

Component Amount to be used

Distilled water 100 ml

NaCl 0.9 g

Appendices

Appendix 8

Focht solution

The Focht trace element solution (Malghani et al., 2009) contains:

Component Composition (mg/L)

MnSO4.H2O 169 mg/l

ZnSO4.7H2O 288 mg/l

CuSO4.5H2O 250 mg/l

NiSO4.6H2O 26 mg/l

CoSO4 28 mg/l

NaMoO4.2H2O 24 mg/l

pH 7.2±0.2

Appendix 9

Nitrogen free medium (NFM)


Na2 H PO4 5.8 g/l

KH2PO4 3.0 g/l

NaCl 0.5 g/l

MgSO4.7H2O 0.25 g/l

pH 6.8-7.00

Appendix 10

Chloride free medium (CFM)

Component Composition g/l

Distilled water 1L

Na2 H SO4 5.8 g/l

KH2SO4 3.0 g/l

MgSO4.7H2O 0.25 g/l

pH 6.8-7.00

Appendices

Appendix 11

Solutions for standard Miniprep protocol

Solution 1 (Suspension Buffer)

Component Concentration

Tris (pH 8.0) 50 mM

EDTA 10 mM

RNAase A 100 mM

Solution 2


NaOH 200 mM

(SDS) 10%

Solution 3 (pH 4.8-5.0)


Potassium acetate 3.0 mM

Glacial acetic acid 11.5 ml/l

Appendix 12

Extraction of pesticide residues from roots and shoots

Column preparation

Took a glass column with stopper at its one end

Placed the glass wool at the base of the column above the stopper and fill it with

charcoal

Conditioned the charcoal with solvent (acetone)

Poured the sample into the column and let it to be filtered under gravity

Washed the column with acetone than with distilled water thrice to remove any

remnants of the first sample

Appendices

Sample preparation and purification

Took 5 g sample of shoots/roots

Dipped into the liquid nitrogen and macerate into the piston Morton to obtain cell

extract

Mixed the cell extract with acetone

Passed this extract through the column to remove the cell debris

Saved the filtrate

Extracted the CP with dichloromethane

The solvent, dichloromethane was allowed to evaporate

Dissolved the residues of CP in acetonitrile and filtered for analysis on HPLC

Appendix 13

Salkowski reagent


Distilled water 50 ml

Ferric chloride0.5M 1 ml

Sulphuric acid

with (Specific gravity

1.84)

30 ml

Publications

Jabeen, H., Iqbal, S., Anwar, S. 2014. Biodegradation of chlorpyrifos and 3, 5, 6-

trichloro-2-pyridinol by a novel rhizobial strain Mesorhizobium sp. HN3. Water

and Environment Journal (Published Online on 7th

March, 2014)

Biodegradation of chlorpyrifos and 3, 5, 6-trichloro-2-pyridinolby a novel rhizobial strain Mesorhizobium sp. HN3Hina Jabeen1,2, Samina Iqbal1,2 & Samina Anwar1

1Soil and Environmental Biotechnology Division, National Institute for Biotechnology and Genetic Engineering (NIBGE), Faisalabad, Pakistan and 2Pakistan

Institute of Engineering and Applied Sciences (PIEAS), Islamabad, Pakistan

Keywords3,5,6-trichloro-2-methoxypyridine;

3,5,6-trichloro-2-pyridinol; biodegradation;

chlorpyrifos; Mesorhizobium sp.

CorrespondenceSamina Iqbal, Soil and Environmental

Biotechnology Division, National Institute for

Biotechnology and Genetic Engineering

(NIBGE), PO Box 577, Jhang Road, Faisalabad

38000, Pakistan. Email: [email protected]

doi:10.1111/wej.12081

Abstract

A chlorpyrifos (CP) and 3,5,6-trichloro-2-pyridinol (TCP) degrading bacterial strain,Mesorhizobium sp. HN3, was isolated and characterized. Mesorhizobium sp. HN3degraded CP efficiently up to 400 mg/L initial concentration at wide range of tem-peratures (30–40°C) and pH (6.0–8.0). However, optimal degradation of CP wasachieved at 37°C and neutral pH (7.0) at an initial inoculum density 2 × 107 colonyforming unit/mL of culture medium. Kinetic parameters for CP degradation byMesorhizobium sp. HN3 were estimated at different initial concentrations. Culturesexhibited significant variation (P ≤ 0.05) in the specific growth rate (μ), cell massformation rate (QX) and the substrate uptake rate (QS) during degradation of CP. Thevalues of kinetic parameters increased up to 100 mg/L CP and decreased at higherconcentration. Investigation of degradation metabolites indicated that CP is con-verted to diethylthiophosphate and TCP that leads to the formation of 3,5,6-trichloro-2-methoxypyridine.

Introduction

Chlorpyrifos [O,O-diethyl-O (3,5,6-trichloro-2-pyridyl phos-phorothioate)] (CP) is a moderately toxic and broad spectrumorganophosphate (OP) insecticide. It is an important ingredi-ent of common household formulations that are effectiveagainst mosquitoes, termites, bees, flies, etc. (Bicker et al.2005; Mohan et al. 2007). CP is also extensively used in agri-culture to kill the insect pests of a variety of crops such ascereals, cotton, fruits and vegetables since many years (Fanget al. 2006; Wang et al. 2007). Although many OPs includingCP were initially regarded as less persistent and toxic, there isescalating concern that these pesticides or their metabolitesare highly persistent in the environment as well as toxic andhence lead to undesirable health issues (Ragnarsdottir 2000;Alavanja et al. 2013).

A consequence of continuous domestic and agriculturaluse of CP is a widespread contamination of environmentleading to serious damage to nontarget organisms and eco-systems (Rovedatti et al. 2001; Anderson & Hunta 2003; Vogelet al. 2008). In the environment, CP is converted to 3,5,6-trichloro-2-pyridinol (TCP), a persistent metabolite that isresistant to biotic and abiotic degradation owing to the pres-ence of three chloride residues on the N-aromatic ring (Rackeet al. 1996; Robertson et al. 1998; Singh et al. 2003; Chishti &Arshad 2013). Moreover, TCP has higher water solubility ascompared with the parent compound; hence, it leaches to the

water bodies causing widespread contamination of aquaticenvironments (Vogel et al. 2008; Xu et al. 2008; Grzelak et al.2012; Watts 2012). CP and TCP toxicity has been linked tobroad-spectrum effects including neurological disorders,developmental disorders, autoimmune disorders and inter-ruption of many vital functions in higher animals and humans(Sogorb et al. 2004; Mehta et al. 2008; Alavanja & Bonner2012; Ventura et al. 2012; Estevan et al. 2013).

All these concerns imply that removal of both CP and TCPfrom the environment to alleviate their hazardous effects isimperative. A number of approaches including chemical treat-ment, photodecomposition and incineration can be appliedfor the remediation of contaminants (Olexsey & Parker 2006);however, most of them are expensive, environmentally unfa-vourable and not applicable for diffused contamination at lowconcentration. Use of microorganisms having the right meta-bolic pathways seems to be the most feasible technology forremediation of CP, TCP and related contaminants (Thengodkar& Sivakami 2010; Singh et al. 2011).

Bacterial strains capable of degrading CP and TCP as a solesource of carbon and energy as well as cometabolically havebeen isolated and characterized during recent years. Asummary of biodegradation studies of CP and TCP has beenreviewed by Maya et al. (2011). Recently reported CP degrad-ing bacterial strains include Bacillus cereus (Liu et al. 2012),Stenotrophomonas maltophilia strain MHF ENV20 (Dubey &Fulekar 2012) and Cupriavidus sp. DT-1 (Lu et al. 2013). It has

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also been generally recognized that microbial degradation ofCP can be affected by many biotic and abiotic factors andtolerance to initial pesticide concentration, microbial popula-tion, optimum growth temperatures and optimum pH varyfor different microorganisms (Singh et al. 2003; Anwar et al.2009; Sharma 2012). The objectives of the present studywere to isolate and characterize bacterial strain capable ofcomplete degradation of CP and TCP, optimize culture condi-tions that govern CP degradation by the isolate and investi-gate the pathway of degradation. The kinetics of CPbiodegradation, accumulation and utilization of TCP, and thegoverning constants thereafter were also determined. As,these parameters vary depending on bacterial strains andconcentration/nature of pollutant, a clear understanding ofthe biodegradation kinetics of CP and TCP would determinesuitability of the bacterial strain for in situ bioremediation.

Materials and methods

Chemicals

Analytical standard and technical grade CP were purchasedfrom Dr Ehren Stofer GmbH (Germany) and Chem Service(Web Chester), respectively. Dichloromethane (DCM) andhigh performance liquid chromatography (HPLC) grade sol-vents were purchased from Merck. N,O-Bis (trimethylsilyl)trifluoroacetamide (BSTFA) kit was purchased from Supelco,Bellefonte, PA, USA.

Enrichment, isolation and selection of CPdegrading bacterial strains

Three different agricultural soil samples were collected fromfields where CP had been applied frequently. CP degrading bac-teria were isolated by enrichment culture technique followingAnwar et al. (2009). Initially, CP utilization by the isolates wasmonitored on minimal salt medium (MSM) agar plates contain-ing CP (100 mg/L). Growth and CP degradation potential of theisolates was also observed in liquid cultures (MSM) sup-plemented with 100 mg/L CP as the only source of carbon andenergy. A bacterial isolate, HN3 was found most efficient for CPutilization and hence selected for further studies.

Identification of isolate HN3

Total genomic DNA of the Mesorhizobium sp. HN3 was iso-lated, and 16S rRNA gene was amplified using universalprimers FD1 (5'-AGAGTTTGATCCTGGCTCAG-3'; Escherichiacoli bases 8-27) and RP1 (5'-ACGGHTACCTTGTTTACGACTT-3';E. coli bases 1507-1492) (Wilson et al. 1990). Polymerasechain reaction product was cloned into TA cloning vector(pTZ57R) and sequenced. Biochemical and physiologicaltests of the strain HN3 were carried out using QTS-24 kitdeveloped by Defense Science and Technology Organization

Laboratories, Karachi, Pakistan according to the manufac-turer instructions.

Inoculum preparation of HN3 forbiodegradation studies

The overnight grown culture of strain HN3 in Lauria Bertanimedium containing 100 mg/L CP was harvested and centri-fuged at 4600 g for 10 min. The cell pellet was washed with0.9% normal saline and suspended in the same solution toobtain an optical density of 0.8 at 600 nm (OD600nm). Dilutionplate count technique was used to determine the colonyforming units/mL, and 2% of this suspension was used asinoculum in CP biodegradation experiments until otherwisestated.

Experimental set-up for CP degradation studiesby HN3

CP degradation studies were carried out in 250-mL Erlen-meyer flasks containing 50-mL MSM supplemented with 2%HN3 inoculum and 100 mg/L CP under various culture condi-tions as described in respective sections. The flasks wereincubated at 37°C and 100 rpm in rotary shaker for 10 days.For all the treatments, uninoculated flasks served as controls,and all the experiments were performed in triplicate. Sampleswere periodically harvested for analysing the growth rates andresidues of CP and TCP. Extraction and HPLC analysis of pesti-cide residues was carried out as described in Anwar et al.(2009). Biodegradation was estimated by comparing theremoval of CP in samples and controls over time. Turbi-dometric method described by Jyothi et al. (2012) wasemployed for monitoring the growth of the Mesorhizobiumsp. HN3. Cell dry mass was determined for the Mesorhizobiumculture having an OD600nm of 1.0 and was used as standard forcalculating cell dry mass of samples.

Optimization of temperature and pH forbiodegradation of CP by HN3

To optimize temperature for degradation of CP by strain HN3,culture flasks containing MSM (pH 7.0) were incubated at 30,37 and 40°C in rotary shaker at 100 rpm. Degradation capac-ity of CP by Mesorhizobium sp. HN3 was monitored in MSMwith different initial pH, that is, acidic (6.0), basic (8.0) andneutral (7.0) prepared according to Anwar et al. (2009).

Kinetics of CP degradation by HN3 at differentinitial concentrations of CP

Biodegradation of CP by strain HN3 was investigated at differ-ent initial concentrations (50, 100, 200, 300 and 400 mg/L) andkinetic parameters were determined as described by Pirt

Biodegradation of CP and TCP H. Jabeen et al.

2 Water and Environment Journal (2014) © 2014 CIWEM.

(1975). Rates of pesticide degradation (Qs), metabolite pro-duction (QP) and cell mass productivity (Qx) were determinedby calculating the slope in their respective plots versus time(h). Product yields (Yp/s) and cell mass yield (Yx/s) were deter-mined by dP/dS and dX/dS where dP, dS and dX are thechanges in concentrations of product, pesticide and the bac-terial cell mass, respectively, per unit time. Specific growthrate (μ) was determined by plotting the ln(X/X0) versus time,where X0 and X are the initial cell mass (g/L) and cell mass (g/L)at time ‘t’, respectively (calculated during exponential phaseat different time intervals). Specific productivity (qp) and spe-cific rate of CP degradation (qs) were multiple of μ and Yp/x andYx/s.

Kinetic model was determined by plotting log CP residuesagainst time. A straight line was obtained for all CP concen-trations (50–400 mg/L), following first-order kinetics model.Therefore, degradation rate constant (k, h−1) and half-life (T1/2)in days were determined using Eqs (1) and (2) as described inDubey and Fulekar (2012).

C C ett= × −

0k (1)

T1 2 2= ( )ln k (2)

Identification of CP metabolites

Samples containing residues of CP and its metabolitesobtained from culture flasks were extracted with DCM andderivatized with BSTFA [CF3C=NSi(CH3)3OSi(CH3)3] usingBSTFA kit according to the protocol. Gas chromatography-mass spectrometry (GC-MS) analyses were performed withan Agilent 6890N gas chromatograph, equipped with aSupleco Equity-1 capillary column (30 m by 250 μm and25 μm film thickness), an auto-injector (7683 series) and anAgilent 5973 network mass selective detector (Agilent Tech-nologies, Palo Alto, CA, USA).

Helium was used as the carrier gas with a constant flowrate of 0.5 mL/min. The injector and transfer lines were 220and 300°C, respectively. The chromatography program wasas follows: total run time 33 min, initial temperature ofcolumn 70°C, a temperature increase of 10°C/min and finalheating to 240°C. The ionization voltage and electron multi-plier settings were 70 eV and 1294 V, respectively. Metabo-lites were identified by comparison of retention time (RT) andMS fragmentation profile of the metabolites to those ofauthentic standards.

Statistical analysis

All of the experiments were performed in triplicate. Meansand the standard deviations were determined using StatisticAnalysis System (SAS 9.0) software packages.

Results

Isolation and selection of CP degradingbacterial strain

Sixteen bacterial isolates capable to grow on MSM agarplates containing CP were obtained initially. Eight of theseisolates utilized CP (100 mg/L) as a sole source of carbon andenergy in MSM broth. One of the isolates, HN3, showing com-plete degradation of CP (100 mg/L) within 5 days of incuba-tion was recognized as most efficient and employed forfurther CP degradation studies.

Molecular, morphological and biochemicalidentification of strain HN3

The 16S rRNA gene sequence of the strain HN3, Gene bankaccession number JN119831, showed > 97% identity with cor-responding sequences of Mesorhizobium spp. and wasgrouped in a well-supported branch with various Mesor-hizobium spp. (Fig. 1). Morphological and biochemical charac-ters of isolate HN3 were compared with those described byJarvis et al. (1997) that also confirmed strain HN3 to be aMesorhizobium sp.

Biodegradation of CP by Mesorhizobiumsp. HN3

Optimum temperature for the degradation of CPby Mesorhizobium sp. HN3

Data indicating the effect of temperature on biodegradationof CP by Mesorhizobium sp. HN3 is shown in Fig. 2. In thepresence of 100 mg/L CP, 33 and 18% of the added pesticidewas degraded after 24 h of incubation at 37 and 40°C thatwas significantly higher as compared with that at 30°C whereonly 5% degradation was observed. After 5 days of incuba-tion, complete CP degradation was observed at 37°C as com-pared with 85 and 55% at 40 and 30°C, respectively, indicatinga significant effect of temperature on CP degradation rate.

Optimization of pH for CP degradation byMesorhizobium sp. HN3

Efficient degradation of CP was achieved at all the three initialpH levels tested (Fig. 3). At 100 mg/L initial concentration, atpH 7.0, the entire added CP was degraded after 5 days ofincubation (Fig. 3). Degradation was relatively slow at alkalinepH (8.0), whereby 100% degradation was achieved after 7days of incubation and a further decline in the degradationwas observed at acidic pH (6.0) as 100% degradation wasobserved after 8 days of incubation. Among the three pHconditions tested for CP degradation by Mesorhizobium sp.HN3, pH 7.0 was found to be optimum.

H. Jabeen et al. Biodegradation of CP and TCP


Mesorhizobium_metallidurans_(NR_042685.1)

Mesorhizobium_tarimense_(NR_044051.1)

Mesorhizobium_gobiense_(NR_044052.1|)

Mesorhizobium_tianshanense_(NR_024880.1)

Mesorhizobium_mediterraneum_(NR_042483.1)

Mesorhizobium_temperatum_(NR_025253.1)

Mesorhizobium_huakuii_(NR_043390.1)

Mesorhizobium_amorphae(NR_024879.1)

Mesorhizobium_loti_(NR_074162.1)

Mesorhizobium_opportunistum_(NR_074209.1)

Mesorhizobium_plurifarium_(NR_026426.1)

Mesorhizobium_thiogangeticum_(NR_042358.1)


HN3_(JN119831.1)

Mesorhizobium_sp.(EF100516.1)


Sinorhizobium_terangae_(NR_044842.1)

Sinorhizobium_kostiense_(NR_042484.1)

Sinorhizobium_saheli(NR_026096.1)

98

75

96

95

7156

67

32

100

76

40

100

71

99

0.0000.0050.0100.015

Fig. 1. Unweighted pair group mean average tree showing the phylogenetic relationship of strain HN3 with the related species based on the 16S rRNA

gene sequences. Bootstrap values that are expressed as the percentages of 1000 replications are shown at the nodes of the branches.

CP

degr

adat

ion

(%)

Time in days

0102030405060708090

100

0 1 2 3 4 5 6 7 8 9 10

Fig. 2. Degradation of chlorpyrifos (CP) by Mesorhizobium sp. HN3 as a

sole source of carbon and energy at different incubation temperatures;

30 (◆), 37 (■) and 40°C (▲). Dashed lines show uninoculated controls; 30

(◇), 37 (□) and 40°C (△). Values are the means of three replicates and

error bars represent standard error.

Time in days

CP

degr

adat

ion

(%)

0102030405060708090

100

0 1 2 3 4 5 6 7 8

Fig. 3. Degradation of chlorpyrifos (CP) by Mesorhizobium sp. HN3 as a

sole source of carbon and energy at different initial pH; 6.0 (◆), 7.0 (■)

and 8.0 (▲). Dashed lines show uninoculated controls at pH 6.0 (◇), 7.0

(□) and 8.0 (△). Values are the means of three replicates, and error bars




Biodegradation of CP at differentinitial concentrations

Mesorhizobium sp. HN3 was able to degrade CP efficiently upto 400 mg/L initial concentration in MSM, whereby degrada-tion was achieved in concentration-dependent manner. CPdegradation and bacterial biomass production at differentinitial CP concentrations after 3 days of incubation are pre-sented in Fig. 4(a). In the cultures containing 50 and 100 mg/LCP, 100 and 85% degradation was achieved, respectively,whereas at 200, 300 and 400 mg/L initial concentrations, 45,33 and 15% CP was degraded, respectively. After 3 days ofincubation, cell biomass (g/L) was highest at 100 mg/L initialCP concentration and declined gradually at concentrationsbeyond this. As depicted in Fig. 4(b), specific rate of CP deg-radation was dependent on initial concentration with anincrease in specific degradation rate at lower initial CP con-centrations (50–100 mg/L) and a decline in the rate at higherinitial CP concentrations. Mesorhizobium sp. HN3 could tol-erate higher CP concentration, with delayed degradation,that is, 40 and 30%, respectively, after 16 days of incubation at1000 and 1200 mg/L CP (data not shown). Complete degra-dation of CP was achieved after 5, 7, 9 and 10 days at 100,200, 300 and 400 mg/L, respectively.

Kinetics of CP degradation and TCP accumulationand degradation thereafter

Effect of initial concentrations of CP on kinetic parametersviz. specific degradation rate (qs), specific growth rate (μ) sub-strate (CP) consumption variables (Qs, Qx, Yx/s) and product(TCP) formation parameters (Qp, Yp/s, qp) are presented in

Table 1. The CP consumption and TCP production parameterswere high at lower CP concentrations (50–100 mg/L) followedby a decline at higher concentrations (200–400 mg/L). Rate ofTCP accumulation (QTCP) increased with the increase in CPconcentrations up to 200 mg/L with a gradual decrease athigher concentrations. TCP yield (Yp/s) and qp were highest at100 mg/L initial concentration and decreased at higher con-centrations. As illustrated in Fig. 5, at 50 and 100 mg/L,maximum concentration of TCP was observed after 48 h ofincubation, and all of the TCP produced as a result of CPhydrolysis was degraded after 96 and 144 h, respectively. Athigher concentrations, TCP was detected in the culturemedia even after 240 h of incubation that might be due to thecontinuous production and slow degradation. Lag phase ofbacterial growth was extended with an increase in initial CPconcentration beyond 100 mg/L.

Figure 6 shows the fitting results of the kinetic modelbased on the experimental data of CP degradation. The deg-radation followed the first-order reaction as a straight linewas produced by plotting the ln values (Ct/C0) of CP residuesagainst respective hours. The residue data were thereforeinterpreted statistically for the calculation of regression equa-tion and first-order kinetic parameters (Table 2). Regressioncoefficient indicating the degradation rate further supportedthe findings that the CP persistence increases with increasinginitial concentration.

Identification of CP metabolites andpathway prediction

Metabolites of CP were identified by GC-MS analysis ofsamples obtained from culture media containing CP as a

Initial CP concentration (mg/L)

CP

degr

adat

ion

(%)

Cel

l Bio

mas

s (dr

y w

t. g

/L)

0

0.05

0.1

0.15

0.2

0

20

40

60

80

100

50 100 200 300 400

a


Spec

ific

CP

degr

adat

ion

rate

(h- 1

)

0

0.02

0.04

0.06

0.08

0.1

0.12

0.14

0.16

0.18

0 50 100 150 200 250 300 350 400

b

Fig. 4. (a) Degradation of chlorpyrifos (CP) by Mesorhizobium sp. HN3 at different initial concentrations as a sole source of carbon and energy after 3 days

of incubation (●) and consequent growth of Mesorhizobium sp. HN3 (■). Values are the means of three replicates and error bars represent the standard

error. (b) Effect of initial CP concentrations on specific degradation rate of CP (◆, qs).



Table 1 Kinetic parametersa for chlorpyrifos (CP) degradation and product [3,5,6-trichloro-2-pyridinol (TCP)] formation thereafter by Mesorhizobium sp.

HN3 in liquid cultures containing different initial concentrations of the pesticide


Substrate utilization parameters

μ (/h) Qs (mg/L/h) Qx (mg/L/h) Yx/s (mg/mg/h) qs (mg/mg/h)

50 0.090 ± 0.000 1.15 ± 0.01 1.52 ± 0.01 1.24 ± 0.005 0.080 ± 0.001

100 0.280 ± 0.005 1.43 ± 0.02 4.60 ± 0.05 1.70 ± 0.040 0.164 ± 0.002

200 0.058 ± 0.000 1.30 ± 0.01 2.88 ± 0.00 0.78 ± 0.003 0.075 ± 0.001

300 0.023 ± 0.000 0.72 + 0.00 2.56 + 0.03 0.45 ± 0.001 0.050 ± 0.000

400 0.025 ± 0.000 0.26 + 0.00 2.21 + 0.01 0.35 ± 0.003 0.045 ± 0.001

Product formation parameters

QP (mg/L/h) YP/S (mg/mg/h) qp (mg/mg/h)

50 1.15 ± 0.010 0.40 ± 0.005 0.015 ± 0.040

100 2.24 ± 0.020 0.81 ± 0.010 0.073 ± 0.030

200 2.67 ± 0.010 0.50 ± 0.020 0.013 ± 0.010

300 1.08 ± 0.030 0.23 ± 0.010 0.005 ± 0.000

400 0.99 ± 0.001 0.17 ± 0.000 0.002 ± 0.000

Each value is the means of three replicates ± standard errors. All the values differ from each other significantly at P < 0.05.aKinetic parameters: μ (/h), specific growth rate; Qs, mg substrate consumed/L/h; Qx, mg cell mass produced/L/h; Yx/s, mg cells/mg substrate utilized; qs,

mg substrate consumed /mg cells /h; Qp, mg TCP produced/L/h; Yp/s, mg TCP produced/mg substrate consumed; qp, mg TCP produced /mg cells/h.

CP/

TC

P co

ncen

trat

ion

and

cell

biom

ass (

mg

/L)

Time in hours

a

c

b

d

Fig. 5. Kinetics of chlorpyrifos (CP) degradation at 37°C by Mesorhizobium sp. HN3 as a sole source of carbon and energy at different initial concentra-

tions; (a) 50, (b) 100, (c) 200 and (d) 300 mg/L showing residual CP concentration (△), 3,5,6-trichloro-2-pyridinol (TCP) concentration (○) and cell biomass

of Mesorhizobium sp. HN3 (●) in the culture media. Values are the means of three replicates, and error bars represent the standard error.



source of carbon and energy. In the total ion chromatogram(TIC), CP was indicated at an RT of 16.02 min. Mass spectrumof this peak was identical to that of authentic CP standardshowing molecular ion peak with an m/z value of 351. In theTIC of samples obtained after 3 days of incubation, peak cor-responding to CP disappeared as the metabolism proceeded,and some new peaks appeared at different RTs. These peakswere identified as (a) TCP at RT of 10.23 min, (b)diethylthiophosphate (DETP) at RT of 11.39 min and (c) 3,5,6-trichloro-2-methoxypyridine (TMP) at RT of 7.72 min (Fig. 7).As mentioned earlier, the extracted samples were derivatizedwith BSTFA, trimethylsilyl group (m/z 73) attached to thehydroxyl group of TCP and DETP. The derivatives thus hadmasses of ∼271 and 241 for TCP and DETP, respectively, dem-onstrating a mass of 198 for TCP and 169 for DETP. Peakscorresponding to these three metabolites were not detectedin TIC of cultures by the end of experiments. The resultsindicate that Mesorhizobium sp. HN3 efficiently hydrolysedCP to TCP and DETP. TCP was converted to TMP, whichwas further degraded as indicated by appearance ofdechlorination and ring cleavage metabolites in the subse-quent cultures (data not shown). On the basis of the previousfindings, a pathway was predicted for CP biodegradation byMesorhizobium sp. (Fig. 8).

Discussion

The present study describes the isolation and characteriza-tion of a novel bacterial strain Mesorhizobium sp. HN3capable of complete degradation of CP, a chlorinated OP pes-ticide. To date, many bacterial strains capable of CP and TCPdegradation have been reported (Chishti & Arshad 2013;Chishti et al. 2013) including a few PGPRs, that is,Pseudomonas sp. (Fulekar & Geetha 2008), Bacillus sp.(Zhu et al. 2010), Flavobacterium sp. (Mallick et al. 1999),Klebseilla sp. (Ghanem et al. 2007), etc. However, PGPRbelonging to Rhizobia group have rarely been reported todegrade pesticides and related contaminants.

Notably, temperature and pH of the soil and water greatlyaffect the efficiency of the microorganisms to degrade pesti-cides (Goda et al. 2010). Different bacterial species havebeen reported to show different optimal temperatures for CPdegradation (Li et al. 2007; Lu et al. 2013). However, strainHN3 performed well at a range of temperatures, that is,30–40°C. Although, neutral pH was found to support the bio-degradation of CP by Mesorhizobium sp. HN3, efficient deg-radation was also achieved at acidic (6.0) and basic (8.0) pH.This was in contrast with the findings of Racke et al. (1996)who exclaimed that high pH had a co-relation with hydrolysis

R² = 0.917 R² = 0.977R² = 0.933

R² = 0.911

R² = 0.900

00.5

11.5

22.5

33.5

44.5

55.5

66.5

7

0 24 48 72 96 120 144 168 192 216 240

lnR

esid

ues o

f CP

Time in hours

Fig. 6. First-order kinetics of chlorpyrifos (CP)

degradation in minimal salt medium at differ-

ent initial concentrations; 50 (◆), 100 (■), 200

(▲), 300 (✗) and 400 mg/L (●).

Table 2 First-order kinetics parameters for chlorpyrifos degradation by Mesorhizobium sp. HN3 in liquid cultures containing different initial

concentrations of the pesticide

CP concentration (mg/L) Rate constant (h−1) T1/2 (days) R2 Regression equation

50 0.025 1.16 0.972 3.853 − 0.025x

100 0.023 1.26 0.977 4.699 − 0.023x

200 0.015 1.93 0.933 5.557 − 0.015x

300 0.014 2.06 0.911 6.161 − 0.014x

400 0.012 2.41 0.900 6.451 − 0.012x



of CP in soil and Singh et al. (2003) who demonstrated thathigh (basic) pH support microbial hydrolysis of CP.

Mesorhizobium sp. HN3 degraded CP over a wide range ofinitial concentrations (50–400 mg/L) in contrast with previousreports showing inhibition of bacterial growth and CP degra-dation rates at concentrations < 400 mg/L (Singh et al. 2004,2006; Li et al. 2007). Higher specific growth rate (μ), cell massformation rate (QX) and the substrate uptake rate (QS) at

50–100 mg/L initial CP concentration indicated a remarkableimpact of initial concentration on CP degradation rate. CPdegradation by Mesorhizobium sp. HN3 followed first-orderkinetics with a dramatic decrease in degradation rate withincreasing initial concentration. A similar trend for CP degra-dation has been reported earlier (Dubey & Fulekar 2012).

High rate of TCP production and complete TCP removal atrelatively lower initial CP concentrations (50–100 mg/L) wasobserved, which might be attributed to the easy adaptation ofHN3 to low TCP concentrations in the culture media accumu-lated as a result of CP degradation. Delay in complete disap-pearance of CP and TCP at higher concentrations might be dueto the antimicrobial/inhibitory effect of relatively higher con-centration of TCP produced towards Mesorhizobium sp. HN3.Generally, the lag phase of bacterial growth and CP degrada-tion was extended with the increase in initial CP concentra-tion. These results are consistent with the previous reportsthat reveal the extended lag phase at higher CP concentra-tions (Singh et al. 2006; Anwar et al. 2009; Chen et al. 2012).

Mass spectrometric identification of the metabolites pro-duced during CP degradation indicated that first step of themetabolic pathway is hydrolysis of O-P ester linkage toproduce TCP and DETP. However, TCP and DETP were notpersistently found during the course of experiments and dis-appeared subsequently. Singh et al. (2004) suggested thatDETP is utilized as a carbon and energy source by microor-ganisms for further degradation of TCP. Strain HN3 was alsocapable to grow in phosphorus-free medium containing CP asthe only P source (data not shown) indicating utilization ofDETP as P source as well. Importantly, O-methylation of TCPto produce TMP and its subsequent disappearance wasobserved, which substantiates that TMP was furtherdegraded. Biodegradation of TCP is a crucial part in the reme-diation of CP contaminated sites. If left accumulated, TCP willaffect the beneficial microbial communities in the soilbecause of its antimicrobial properties. These findings makestrain HN3 an efficient tool for remediation of CP, TCP andrelated contaminants.

Conclusions

(1) Mesorhizobium sp. strain HN3 is capable of degrading CPat a wide range of initial concentrations, temperatures andpH.(2) HN3 degrades CP through P-O-C bond hydrolysis produc-ing DETP and TCP that leads to the formation of TMP.(3) TCP and TMP produced as a result of CP metabolism arefurther degraded.(4) Kinetic parameters indicate that fairly faster CP and TCPdegradation by Mesorhizobium sp. HN3 can be obtained atrelatively lower concentrations.(5) Mesorhizobium sp. HN3 can potentially be used for thebiodegradation and bioremediation of CP and TCP.

─Si (CH3)3

m/z 197 m/z 73.19 m/z 271

NCl OH

ClCl

NCl O

ClCl

Si(CH3)3

a

m/z 210

c

N

Cl

OCH3

Cl

Cl

m/z 169 m/z 73.19 m/z 241

PS

HO

O

OCH3

CH3─Si (CH3)3 P

S

O

O

OCH3

CH3

Si(CH3)3

b

Fig. 7. (a) Mass spectra of 3,5,6-trichloro-2-pyridinol; (b) diethylt-

hiophosphate and (c) 3,5,6-trichloro-2-methoxypyridine formed in the

culture medium after 3 days of incubation corresponding to peaks

obtained at retention time of 7.72, 10.23 and 11.39 min, respectively, in

the total ion chromatogram.



Acknowledgements

The present research was financially supported by HigherEducation Commission (HEC), Pakistan. Dr. Sajjad Mirza andDr. Ghulam Rasool, National Institute for Biotechnology andGenetic Engineering, are greatly acknowledged for grantingaccess to HPLC facility. Authors are grateful to Professor Dr.Mohammad Ibrahim Rajoka for guidance in kinetic analysis ofdata.

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Chlorpyrifos(CP)

Diethylthiophosphoricacid (DETP)

3,5,6-trichloro-2-methoxypyridine

(TMP)

O-methylation

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

Hydrolysis

dechlorination and ring cleavage

Further breakdown

NCl

OP

O

CH3

S OCH3Cl

Cl

PS

HO

O

OCH3

CH3NCl OH

ClCl

N

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Fig. 8. Proposed pathway for biodegradation

of chlorpyrifos by Mesorhizobium sp. HN3 in

liquid cultures.



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