STUDIES ON BIODEGRADATION OF GLYPHOSATE HERBICIDE B Y
BACTERIAL SPECIES ISOLATED FROM RICE FIELDS
BY
OKPALA, GLORIA NGOZI
PG/M.Sc./06/41700
DEPARTMENT OF MICROBIOLOGY
UNIVERSITY OF NIGERIA, NSUKKA
SEPTEMBER, 2010.
i
TITLE PAGE
STUDIES ON BIODEGRADATION OF GLYPHOSATE HERBICIDE B Y
BACTERIAL SPECIES ISOLATED FROM RICE FIELDS
BY
OKPALA, GLORIA NGOZI
PG/M.Sc./06/41700
A Dissertation Submitted to the Department of Microbiology
in the Faculty of Biological Sciences In Partial Fulfillment of the Requirement For
The Award Of A Master’s Degree (M.Sc.) in Environmental Microbiology
Supervisor: DR. A. N. MONEKE
SEPTEMBER, 2010
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CERTIFICATION
Miss Gloria Ngozi Okpala a postgraduate student in the Department of
Microbiology, majoring in Environmental Microbiology has satisfactorily completed the
requirements of course work and research for the degree of Master of Science (M.Sc) in
Environmental Microbiology. The work embodied in her Dissertation is original and has
not been submitted in part or full for either diploma or degree of this or any other
University.
Dr. (Mrs.) I. M. Ezeonu Dr. A. N. Moneke Head, Supervisor Department of Microbiology Department of Microbiology University of Nigeria, Nsukka University of Nigeria, Nsukka.
iii
DEDICATION
This work is dedicated to my father, Mr. Cyril Okpala, for the uncompromising moral
philosophy guiding his life, my mother for her nurturing and my late grandmother Mrs.
Florence Agwuncha for believing in me and encouraging me to reach beyond the skies.
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ACKNOWLEDGEMENT
I thank God for His innumerable blessings in my life, most especially for good
health and grace to complete this programme. I acknowledge with deep sense of
indebtedness and appreciation, my supervisor Dr A.N. Moneke for his unflinching
support, directives, advice and supervision of this work.
I also owe a great debt of gratitude to Prof. C. U. Iroegbu who has always had my
welfare at heart from my undergraduate days in the University. He made sure I lacked
nothing throughout my course of study. I wish to especially thank Dr. C.U. Anyanwu of
Microbiology Department, for his patience in listening to my questions and proffering
solutions to them. He was always there, ever ready, to guide and get me back on track any
time I derailed.
. My thanks also go to Dr. E. A. Eze, for his support and encouragement
throughout my research work. My gratitude goes to the following postgraduate lecturers of
the department for their various contributions towards the actualization of this work; Dr.
(Mrs.) I. M. Ezeonu, Prof J.O. Ugwuanyi, Prof J. C. Ogbonna, Dr. Arinze Okoli, Mr.
Nwokoro, Mr. C. N. Eze, Mr. Nnamchi and Mr. I. V. Chigor.
I will remain ever grateful to my parents Mr. and Mrs. C. Okpala for their care and
encouragement. I am equally indebted to my cousin Mr. C. Ejeagwu for the immense
financial support he gave me during the course of this programme. I am grateful to my
brothers and sisters Lillian, Emmanuel, Rosemary and David for their encouragements
during this programme. May the Lord preserve and bless you all.
I also want to thank my room-mates, friends and coursemates whose contributions
are immeasurable. These include; Rev. Sr. Eyisi, Ken, Madam Chioma, Ogechi, Ifedi,
Phidelia, Ogoo, Francis, Okwy, Nkechi, Attai, Bonny, Uche, Ogugua, Victor and others,
you are all wonderful people.
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ABSTRACT
Nineteen bacterial strains were obtained from rice field soil samples and on further
culturing on glyphosate agar medium, seven of the isolates showed consistent growth. The
seven bacterial isolates were identifed as Acetobacter sp., Escherichia sp., Pseudomonas
fluorescens, Azotobacter sp., Pseudomonas sp., Pseudomonas cepacia. and Alcaligenes sp.
Microbial growth during the study was monitored by measuring the optical density at 660
nm. Pseudomonas fluorescens and Acetobacter sp. utilized glyphosate most significantly
(P < 0.05) when compared to the other isolates and were therefore used for further
biodegradation studies. Evaluation of the effect of glyphosate as carbon and/or phosphorus
source on the growth of the isolates showed that there was significant (P < 0.05) growth of
both isolates in the medium containing glucose as carbon source and glyphosate as
phosphorus source. Evaluation of the effects of different concentrations of glyphosate on
the growth of the isolates showed significant (P < 0.05) increase in growth at the lower
concentration (7.2 mg/ml - 25 mg/ml). Although the isolated bacteria were able to tolerate
up to 250 mg/ml of glyphosate, a significant (P < 0.05) reduction in growth of both
isolates was observed as the concentration of glyphosate increased from 100-250 mg/ml.
Growth in the culture medium with different nitrogen sources (0.2%), showed that organic
nitrogen sources (Peptone, tryptone and yeast extract) enhanced growth most significantly
(P < 0.05) when compared with inorganic sources (NH4Cl and (NH4)2SO4) of nitrogen.
Peptone as an alternative nitrogen source yielded the highest growth. Acetobacter sp.
utilized glyphosate significantly (P < 0.05) at pH of 5.0 and 8.0, while Pseudomonas
fluorescens grew significantly (P < 0.05) at pH 4.0. Evaluation of the effects of heavy
metals on growth of the isolates in the presence of glyphosate revealed that lead at 500
µg/ml significantly (P < 0.05) enhanced the growth of Acetobacter sp. while zinc at 500
µg/ml also significantly (P < 0.05) stimulated the growth of Pseudomonas
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fluorescens.This study showed that Pseudomonas fluorescens and Acetobacter spp
exhibited a high capacity to efficiently degrade glyphosate under the environmental
conditions studied. Thus, they can be exploited for biodegradation of this compound and
also should be checked for their ability to degrade other organophosphates.
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TABLE OF CONTENTS
Title Page --------------------------------------------------------------------------------------i
Certification -----------------------------------------------------------------------------------ii
Dedication -----------------------------------------------------------------------------------iii
Acknowledgement ---------------------------------------------------------------------------iv
Abstract ----------------------------------------------------------------------------------------v
Table of Contents ----------------------------------------------------------------------------vii
List of Tables ---------------------------------------------------------------------------------xiii
List of Figures --------------------------------------------------------------------------------xiv
CHAPTER ONE
1.0 Introduction … … … … … … … … 1
1.2 The identified problems … … … … … … 3
1.3 Objectives of the Study … … … … … … 3
1.4.0 Literature Review … … … … … … … 4
1.4.1 What are Pesticides? … … … … … … … 4
1.4.2 Pesticides and pollution … … … … … … 5
1.4.3 Characteristics of an ideal pesticide … … … … … 6
1.4.4 Concerns on the use of pesticides … … … … … 6
1.4.5.0 Nature and types of organophosphate pesticides … … … 8
1.4.5.1 Organophosphate toxicity … … … … … … … 17
1.4.6 Glyphosate herbicide … … … … … … … 18
1.4.6.1 Properties of glyphosate … … … … … … 19
1.4.6.2 Mode of action of glyphosate … … … … … … 28
1.4.6.3 Persistence of glyphosate in the environment … … … … 31
1.4.7.0 Glyphosate toxicity … … … … … … … 34
1.4.7.1 Toxicity and effects on experimental animals and in vitro systems … 34
viii
1.4.7.1.1 Sub chronic toxicity in laboratory animals … … … … 35
1.4.7.1.2 Chronic toxicity in laboratory animals …… … … … 35
1.4.7.2 Acute toxicity in humans … … … … … … 35
1.4.7.3 Reproductive effects . … … … … … … 36
1.4.7.4 Mutagenicity … … … … … … … 36
1.4.7.5 Ecological effects of glyphosate … … … … … 37
1.4.7.5.1 Effect of glyphosate on non target animals … … … … 37
1.4.7.5.2 Effects on non target plants … … … … … … 38
1.4.8 Glyphosate degradation … … … … … … 40
1.4.8.1 Abiotic degradation … … … … … … … 42
1.4.8.1.1 Hydrolytic cleavage … … … … … … … 42
1.4.8.1.2 Photodegradation … … … … … … … 42
1.4.8.2 Biodegradation … … … … … … … … 43
CHAPTER TWO
2.0 Materials and Methods … … … … … … 47
2.1 Isolation and characterization of glyphosate degrading bacteria from
rice field soil samples … … … … … … ... 47
2.1.1 Collection of soil samples … … … … … … 47
2.1.2 Estimation of soil moisture content … … … … 49
2.1.3 Isolation medium … … … … … … … 49
2.1.4 Isolation of glyphosate utilizing bacteria … … … … 50
2.1.5 Storage … … … … … … … … … 50
2.2 Inoculum preparation and standardization … … … … 50
2.2.1 Glyphosate utilization patterns of the different isolates … … … 51
2.3 Identification and characterization of Isolates … … … … 51
2.3.1 Microscopic examination of cell morphology … … … … 51
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2.3.1.1 Gram stain … … … … … … … … 51
2.3.2 Biochemical tests … … … … … … … 52
2.3.2.1 Catalase test … … … … … … … … 52
2.3.2.2 Oxidase test … … … … … … … … 52
2.3.2.3 Sulfide-indole-motility (SIM) screening test …. … … … 53
2.3.2.4 Starch hydrolysis test … … … … … … … 54
2.3.2.5 Nitrate reduction … … … … … … … 54
2.3.2.6 Gelatin hydrolysis … … … … … … … 54
2.3.2.7 Growth on Kligler iron agar … … … … … … 55
2.3.2.8 Sugar fermentation test … … … … … … … 55
2.3.2.9 Citrate utilization … … … … … … … 56
2.4 Biodegradation study in liquid medium … … … … 56
2.4.1 Inoculum preparation and standardization … … … … 56
2.4.2 Preparation of turbidity standard … … … … 57
2.4.3 Growth kinetics of Pseudomonas fluorescens and Acetobacter sp
in glyphosate … … … … … … … … 58
2.4.4 Comparative role of glyphosate as carbon or phosphorus source … 58
2.4.5 Effects of different concentration of glyphosate on the growth of
the Isolates …. …. … … … … … … 59
2.4.6 Effects of nitrogen supplements on growth of the isolate and
Utilization of glyphosate … … … … … … 59
2.4.7 Effects of Medium pH on Growth of the Isolate and
Utilization of Glyphosate … … … … … … 60
2.4.8 Effect of heavy metals on glyphosate degradation … … … 60
2.4.8.1 Comparative effects of heavy metals on Pseudomonas fluorescens and
Acetobacter sp… … … … … … … … 60
x
2.4.8.2 Effect of different concentrations of heavy metals … … … 60
2.5 Colorimetric determination of glyphosate … … … … 61
2.5.1 Preparation of Molybdenum (V)-Molybdenum (VI) reagent … … 61
2.5.2 Preparation of standard curve … … … … … … 61
2.5.3 Determination of residual glyphosate … … … … … 62
2.6 Statistical analysis … … … … … … … 62
CHAPTER THREE
3.0 Results … … … … … … … … 63
3.1 Soil moisture content … … … … … … … 63
3.2 Isolation of glyphosate degrading bacteria from soil … … … 63
3.3 Glyphosate utilization pattern by the different isolates … … … 65
3.4 Growth kinetics of Pseudomonas fluorescens and Acetobacter sp in
glyphosate … … … … … … … … 65
3.5 Effect of glyphosate as carbon and/or phosphorus source on the growth
of the isolates …. … … … … … … … 68
3.5.0 Comparative role of glyphosate as carbon or phosphorus source … 68
3.5.1 Growth kinetics of Acetobacter sp and Pseudomonas fluorescens
on glyphosate as carbon or phosphorus source … … … … 68
3.6 Effect of nitrogen supplementation on microbial utilization of glyphosate 73
3.6.0 Comparative effect of nitrogen supplementation on the growth
Acetobacter sp and Pseudomonas fluorescens in the presence
of glyphosate … … … … … … … … 73
3.6.1 Growth kinetics of Acetobacter sp on the nitrogen supplements … 73
3.6.2 Growth kinetics of Pseudomonas fluorescens on the nitrogen
Supplements … … … … … … … … 74
xi
3.7 Effects of different glyphosate concentrations on the growth of
Acetobacter sp and P. fluorescens … … … … … 78
3.8 Effect of heavy metals on microbial utilization of glyphosate … … 82
3.8.0 Comparative effects of heavy metals on Acetobacter sp and
Pseudomonas fluorescens … … … … … … 82
3.8.1 Effects of different concentrations of cadmium on the growth of
Acetobacter sp … … … … … … … … 82
3.8.2 Effects of different concentration of chromium on the growth of
Acetobacter sp … … … … … … … … 83
3.8.3 Effects of different concentrations of lead on the growth of
Acetobacter sp … … … … … … … … 83
3.8.4 Effects of different concentrations of zinc on the growth of
Acetobacter sp … … … … … … … … 83
3.8.5 Effects of different concentrations of cadmium on the growth
of Pseudomonas fluorescens … … … … … … 83
3.8.6 Effects of different concentrations of chromium on the
growth of Pseudomonas fluorescens … … … … … 84
3.8.7 Effects of different concentrations of lead on the growth of
P. fluorescens … … … … … … … … 84
3.8.8 Effects of different concentrations of zinc on the growth of P. fluorescens 85
3.9 Effect of pH on microbial degradation of glyphosate … … … 95
3.9.1 Effect of pH on the utilization of glyphosate by Acetobacter sp
and P. fluorescens … … … … … … … 95
3.9.2 Effect of pH on the growth of Acetobacter sp and P. fluorescens and
degradation of glyphosate … … … … … … … 95
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CHAPTER FOUR
Discussion … … … … … … … … … 121
References … … … … … … … … … 129
APPENDIX I … … … … … … … … 142
APPENDIX II … … … … … … … … … 145
APPENDIX III … … … … … … … … 146
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List of Tables
Table 1: Physical properties of glyphosate (IPCS, 1994) … … … 23
Table 2: Some pesticides with their representative Koc values … … 32
Table 3: State and location from where the soil samples were collected … 48
Table 4: Moisture content of the different soil samples … … … 64
xiv
List of Figures
Figure 1: Pathway for the degradation of parathion … … … … 11 Figure 2: Pathway for the degradation of Diazinon … … … … 13 Figure 3: Pathway for the degradation of dichlorovos … … … 15 Figure 4: Glyphosate Structure … … … … … … … 20 Figure 5: Ionic species of glyphosate and their pka values … … … 22 Figure 6: Glyphosate proton dissociation reactions … … … … 25 Figure 7: Formation of N,N'-diphosphonomethyl-2,5-piperazinedione … 27 Figure 8: Shikimic acid pathway and the inhibition by glyphosate in plants and
microorganisms. ... … … … … … … 30
Figure 9: Abiotic and biodegradation processes for glyphosate degradation in the environment … … … … … … … 41 Figure 10: Degradation routes of glyphosate in soil … … … … 46 Figure 11: Screening of the isolates for glyphosate utilization … … 66 Figure 12: Growth kinetics of Acetobacter sp and P. fluorescens on glyphosate 67 Figure 13: Comparative effect of glyphosate as carbon and/or phosphorus source 70 Figure 14: Growth kinetics of Acetobacter sp in glyphosate as carbon or
phosphorus source … … … … …… … … 71
Figure 15: Growth kinetics of Pseudomonas fluorescens in glyphosate as carbon or phosphorus source … … … … … 72
Figure 16: Comparative effect of nitrogen supplementation on the growth Acetobacter sp and Pseudomonas fluorescens in the presence of
glyphosate … … … … … … … … 75 Figure 17: Growth kinetics of Acetobacter sp on the nitrogen supplements
in the presence of glyphosate … … … … … 76 Figure 18: Growth kinetics of Pseudomonas fluorescens on the
nitrogen supplements in the presence of glyphosate … … 77 Figure 19: Effects of the different concentrations of glyphosate on the
growth of Acetobacter sp and Pseudomonas fluorescens … … 79
xv
Figure 20: Growth kinetics of Acetobacter sp on the different concentrations of glyphosate … … … … … … 80
Figure 21: Growth kinetics Pseudomonas fluorescens on the different concentrations of glyphosate … … … … … 81
Figure 22: Comparative effects of heavy metals on Acetobacter sp and
P. fluorescens … … … … … … … 86
Figure 23: Effects of different concentrations of cadmium on the growth of Acetobacter sp … … … … … … … 87
Figure 24: Effects of different concentrations of chromium on the growth of Acetobacter sp … … … … … … … 88 Figure 25: Effects of different concentrations of lead on the growth of
Acetobacter sp … … … … … … … 89 Figure 26: Effects of different concentrations of zinc on the growth of Acetobacter sp … … … … … … … … 90 Figure 27: Effects of different concentrations of cadmium on the growth of
Pseudomonas fluorescens … … … … … … 91
Figure 28: Effects of different concentrations of chromium on the growth of Pseudomonas fluorescens … … … … … … 92 Figure 29: Effect of different concentrations of lead on the growth of
Pseudomonas fluorescens … … … … … … 93
Figure 30: Effects of different concentrations of zinc on the growth of Pseudomonas fluorescens … … … … … … 94
Figure 31: Effect of pH 4 on the utilization of glyphosate by Acetobacter sp 97 Figure 32: Effect of pH 5 on the utilization of glyphosate by Acetobacter sp 98 Figure 33: Effect of pH 6 on the utilization of glyphosate by Acetobacter sp 99 Figure 34: Effect of pH 7 on the utilization of glyphosate by Acetobacter sp 100 Figure 35: Effect of pH 8 on the utilization of glyphosate by Acetobacter sp 101 Figure 36: Effect of pH 9 on the utilization of glyphosate by Acetobacter sp 102 Figure 37: Effect of pH 4 on growth of Acetobacter sp and degradation of
glyphosate … … … … … … … … 103 Figure 38: Effect of pH 5 on growth of Acetobacter sp and degradation of
glyphosate … … … … … … … … 104
xvi
Figure 39: Effect of pH 6 on growth of Acetobacter sp and degradation of
glyphosate … … … … … … … … 105 Figure 40: Effect of pH 7 on growth of Acetobacter sp and degradation of
glyphosate … … … … … … … … 106 Figure 41: Effect of pH 8 on growth of Acetobacter sp and degradation of
glyphosate … … … … … … … … 107 Figure 42: Effect of pH 9 on growth of Acetobacter sp and degradation of
glyphosate … … … … … … … … 108 Figure 43: Effect of pH 4 on the utilization of Glyphosate by P. fluorescens 109 Figure 44: Effect of pH 5 on the utilization of Glyphosate by P. fluorescens 110 Figure 45: Effect of pH 6 on the utilization of Glyphosate by P. fluorescens 111 Figure 46: Effect of pH 7 on the utilization of Glyphosate by P. fluorescens 112 Figure 47: Effect of pH 8 on the utilization of Glyphosate by P. fluorescens 113 Figure 48: Effect of pH 9 on the utilization of Glyphosate by P. fluorescens 114 Figure 49: Effect of pH 4 on growth of Pseudomonas fluorescens and degradation
of glyphosate … … … … … … … 115 Figure 50: Effect of pH 5 on growth of P. fluorescens and degradation
of glyphosate … … … … … … … 116 Figure 51: Effect of pH 6 on growth of P. fluorescens and degradation of
glyphosate … … … … … … … … 117 Figure 52: Effect of pH 7 on growth of P. fluorescens and degradation of
glyphosate … … … … … … … … 118 Figure 53: Effect of pH 8 on growth of P. fluorescens and degradation of
glyphosate … … … … … … … … 119 Fig. 54: Effect of pH 9 on growth of Pseudomonas fluorescens and degradation of
glyphosate … … … … … … … … 120
1
CHAPTER ONE
1.0 INTRODUCTION
The use of pesticides in agriculture has increased over 50-fold since 1950. About
2.5 million tons of industrial pesticides are used each year to secure the food supply for the
growing global population (Miller and Tyler, 2002). In the tropics, intensive agriculture
has led to higher pesticide utilization. (Racke et al., 1997).
Organophosphates including glyphosate account for half of the pesticides used
worldwide with glyphosate based formulations such as Roundup, Accord, Touchdown
consisting the commonest types used for agricultural purposes.
Glyphosate (N-phosphonomethylglycine) is a weak organic acid comprising a
glycine moiety and a phosphonomethyl moiety. It is a broad spectrum, non-selective
herbicide used in the control and/or killing of grasses, herbaceous plants, including deep
rooted perennial weeds, brush, some broad-leaf trees and some shrubs (United State
Department of Agriculture (USDA), 2000; Cox, 2000). It can be used in no-till agriculture,
to prepare fields before planting, during crop development and after crop harvest (USDA,
2000).
Glyphosate inhibits an enzyme found in plants and bacteria that is essential for the
synthesis of amino acids which are building blocks of plant protein (WHO, 2004;
International Programme on Chemical Safety (IPCS), 1994). Inhibition of this enzyme is
via the shikimic acid pathway in which the enzyme, 5-enolpyruvylshikimic acid-3-
phosphate synthase (that catalyses the condensation of shikimic acid and
phosphoenolpyruvate to form chorismate an intermediate in phenylalanine, tyrosine and
tryptophan biosynthesis) is inhibited (Zablotowicz and Reddy 2004; Steinrucken and
Amrhein, 1980).
2
Pesticides are used extensively for agricultural purposes and have been found to
pollute virtually every lake, pond, river and stream in areas where they are used due to run
off and leaching (Amoros et al., 2007). This has severe environmental implications as less
than 1% of the pesticides used actually reach the target organisms. The rest end up
polluting the air, soil, water, plants and animals (Centre for Disease Control (CDC), 2003).
In a study conducted by Johnston (1986) it was observed that the use of pesticides
decreased the biodiversity of the soil. The massive use and biocide activity of pesticides
increased the probability of negative impacts on non-target organisms such as aquatic
biota and soil microorganisms (Amoros et al., 2007; DeLorenzo et al., 2001).
Glyphosate on its own may be relatively harmless to humans. It is, however,
formulated with surfactants such as POEA (Polyoxy-ethyleneamine) which is more toxic
than glyphosate alone. Also 2-4-dichlorophenoxy-acetic acid increases the toxicity of
glyphosate as it is used by most farmers to spike glyphosate in order to boost its efficacy
(Cox, 2000).
On application, glyphosate remains unchanged in the soil for varying lengths of
time, depending on soil texture and organic matter content (Penaloza-Vazquez et al.,
1995). Various processes have been described for the removal of pesticides from soil and
water. The removal/degradation process could be by chemical or microbiological process.
Chemical degradation occurs through reactions such as photolysis, hydrolysis, oxidation
and reduction (Andreu and Pico, 2004). These processes are not effective in the removal
of glyphosate and other pesticides because the bonds present in these pesticides such as
the carbon-phosphorus bond in glyphosate are highly stable to these reactions. Microbial
degradation utilizes microorganisms that have the ability to breakdown pesticides by
cleaving the bonds C-P, C-N present in glyphosate (Jacob et al., 1988).
3
In most Nigerian rice fields, the pesticides of choice are glyphosate formulations. It
is therefore necessary to isolate and characterize indigenous bacterial population capable
of degrading glyphosate herbicide as a veritable alternative to the use of chemical
degradation processes. This work aims at providing an environmentally friendly solution
to the problems associated with glyphosate pesticide application. It is also aimed at
isolating glyphosate-utilizing bacteria and using the isolates to study degradation of
glyphosate in vitro.
1.2 THE IDENTIFIED PROBLEMS
These include the following:
1) Pollution of both surface and groundwater by pesticides (glyphosate)
2) Toxicity of pesticides (glyphosate) to non-target organisms
3) Persistence of glyphosate and other pesticide in the environment, long after application.
1.3 OBJECTIVES OF THE STUDY
1) To isolate and characterize glyphosate-degrading bacteria from some rice fields
(Omor, Omasi, Adani and Abakaliki).
2) To carry out biodegradation of glyphosate in vitro using the microbial isolates
3) To determine the effects of the following factors on the biodegradation of
glyphosate in liquid medium:
a) Heavy metals; b) pH; c) Nitrogen supplementation; d) Carbon source;
e) Concentration of glyphosate
4
1.4.0 LITERATURE REVIEW
1.4.1 What are Pesticides?
The United States Environmental Protection Agency (US EPA, 2006) defines
pesticide as “any substance or mixture of substances intended for preventing, destroying,
repelling, or lessening the damage of any pest. A pesticide may be a chemical substance,
biological agent (such as a virus or bacteria), antimicrobial, disinfectant or device used
against pests including insects, plant pathogens, weeds, molluscs, birds, mammals, fish,
nematodes (roundworms) and microbes that compete with humans for food, destroy
properties, spread or are vectors for diseases or are nuisance. Many pesticides are
poisonous to humans.
The Australian Pesticides and Veterinary Medicines Authority (APVMA) (2004)
defines pesticide as any substance or a mixture of substances used to destroy or alter the
life cycle of any pest. In the APVMA pesticides Act of 1999, it was stated that a pesticide
is a substance or mixture of substances that is represented, imported, manufactured,
supplied or used as a means of directly or indirectly:
a) Destroying, stupefying, repelling, inhibiting the feeding of, or preventing infestation by
or attacks of, any pest in relation to a plant, a place or a thing; or b) Destroying a plant; or
c) Modifying the physiology of a plant or pest so as to alter its natural development,
productivity, quality or reproductive capacity; or d) Modifying an effect of another
agricultural product; or e) Attracting a pest for the purpose of destroying it.
In the light of the above definitions, it can be said that ‘pesticide’ is a broad term,
covering a range of products that are used to control pests. A pest is an organism that is
considered harmful or inconveniencing (APVMA, 2004).
There are several types of pesticides which include;
5
Bactericides - These destroy, suppress or prevent the spread of bacteria. Fungicide -
These are used to control, destroy; render ineffective or regulate the effect of a fungus.
Herbicide - Used in destroying or controlling the spread of a weed or unwanted
vegetation. Insecticides - Used for the control of insects which could be ovicides,
larvicides or adulticides. Miticides - Used in the control of mites. Rodenticides - These
are pesticides used specifically for controlling rodents such as mice and rats. Molluscides
- Used in the control of slugs and snails. Nematicides - Used in the control of nematodes
(APVMA, 2004)
1.4.2 PESTICIDES AND POLLUTION
The use of pesticides has increased 50-fold since 1950 and 2.5 million tonnes of
industrial pesticides are currently being used each year (Miller and Tyler, 2002)
worldwide to secure the food supply of the growing global population. In the tropical
regions of the world, agricultural intensification has led to higher pesticide consumption
(Racke et al; 1997). Pesticides are equally used in household as well as in silvicultural
(cultivation, and management of forest trees) applications.
Pesticides have been found to pollute virtually every lake, river and stream due to
runoff and leaching. This has severe environmental implications as less than 1% of
pesticides used actually reached the target organisms and the rest end up polluting the air,
soil or water. From the study conducted by Johnston (1986), the use of pesticides
decreases biodiversity of the soil. Not using them results in higher soil quality and higher
water retention due to more life in the soil
Pesticides found in aquatic systems are detrimental to many aquatic organisms and
they have been shown to be highly lethal to amphibians and implicated in acute health
problems such as abdominal pains, headaches etc. Many studies have indicated that
6
exposure to pesticides is also associated with long-term health problems such as
respiratory problems, memory disorders etc (Talbot et al., 1999).
1.4.3 CHARACTERISTICS OF AN IDEAL PESTICIDE
An ideal pesticide is one that is environmentally friendly and should possess the
following characteristics;
A) It should be highly specific to the target pest.
B) It should have a short residence time in the environment.
C) It should have harmless breakdown products
D) It should not have any long-term health effects on people.
1.4.4 CONCERNS ON THE USE OF PESTICIDES
Pesticides are of serious concern for human health because many are lipid soluble
and as such they accumulate in our fatty tissues through a process called
bioaccumulation. This is the accumulation of toxic substance in an organism especially in
an organism that forms part of the food chain. Persistent pesticides do not readily cycle
through the ecosystem, because they are not broken down readily. As a result, a persistent
pesticide undergoes bioaccumulation to a much higher degree than a pesticide that is
quickly broken down. Bioaccumulation is then followed by biomagnification or
bioamplification. At higher concentrations, many pesticides are toxic to organisms other
than their intended target. Although pesticides are used in controlled amounts, the levels of
the pesticide can increase as it travels up through the food chain. This process is referred
to as bioamplification/biomagnification. For example, pesticides that contain chlorine,
such as Dichlorodiphenyl trichloroethane (DDT) and Dieldrin, are soluble in fat but not in
water. As a result, these toxins cannot be excreted from the body in urine or sweat. They
7
therefore accumulate in the fatty tissues of animals. When the pesticide is in the
environment, it will enter the bodies of animals that are low in the food chain. Though
there is only a small amount of the toxin in each of the prey animals that a secondary
consumer eats, the amount of the toxin in this secondary consumers body will be larger
because each predator eats many preys. When the secondary consumer’s is eaten, the
higher-level predator gets all of its toxins, plus those of all the other preys it has eaten. At
each stage of the food chain the concentration becomes greater. Hence, the higher the
trophic level, the greater the concentration of toxins.
Furthermore, because pesticides are used to kill target organisms due to their
neurological or reproductive toxicity, they also have many similar deleterious effects in
humans. Recognized health effects of pesticides include cancer, developmental effects,
reproductive effects, endocrine disruption, immunotoxicity, neruotoxicity, and toxicity to
various organ systems such as skin or sense organs, cardiovascular or blood, stomach,
intestines, liver, kidney and respiratory systems. The United States Environmental
Protection Agency (US EPA), (1990) stated that direct exposure to these pesticides can be
harmful particularly to foetuses, infants and children since many of them have adverse
effects on the immune system at very low doses. More insidious are indirect but
cumulative exposures to outdoor and indoor air, dust, food and drinking water. The routes
of exposure contribute to our bodies’ pesticide burden. In a study by the Centre for
Disease Control (CDC) in 2003, 9283 people were tested and the average person had 13
pesticides in his or her body. Most worrisome, the CDC data show that children have
nearly twice that of adults of chlorpyrifos metabolite, an organophosphate pesticide.
Pesticides also present many ecological challenges. These ecological effects are
often considered to be an early warning indicator of potential human health impacts.
Pesticides in the environment can kill organisms, cause cancer, and produce lesions in fish
8
and wildlife, suppress the immune system, cause reproductive failures and can also cause
physiological birth defects such as deformed beaks in birds or malformed reproductive
organs as observed in alligators exposed to DDT. Also, they can cause dramatic decline in
biodiversity in areas that used to be filled with various species community (US EPA,
1990).
1.4.5.0 NATURE AND TYPES OF ORGANOPHOSPHATE PESTICIDES
Organophosphonates are a class of organic compounds characterized by the
presence of one or more carbon-phosphorus (C-P) bonds. The C-P bond is resistant to
chemical hydrolysis, thermal decomposition, and photolysis. Organophosphonates, which
are widely used as pesticides, lubricant additives, flame retardants, plasticizers, corrosion
inhibitors, and drugs, are potent biocides. The most conspicuous examples include the
popular herbicides glyphosate and phosphinothricin; ethyl- and phenylphosphonate
derivatives commonly used as insecticides; Fyrol 76, an oligomer of vinylphosphonate-
methylphosphonate representative of flame retardant; polyaminopolyphosphonic acids,
widely used as corrosion inhibitors; bisphosphonates, which have an application for the
treatment of bone mineralization disorders; the antibiotics alafosfalin and
phosphonomycin; and cyclic esters of aromatic bisphosphonates used as polymer additives
(Zboinska et al., 1992).
Organophosphates are a group of highly toxic compounds that are used extensively
as agricultural and domestic pesticides (Costa, 1998). They are used to control a wide
variety of insect pests, weeds, and disease transmitting vectors (Chaudhry, 1994).
In the United States alone, organophosphate pesticides account for about half of
the insecticides used. Approximately 60 million pounds of organophosphates are applied
9
annually; non agricultural uses accounts for about 17 million pounds per year (US, EPA,
2001; CDC, 2001).
Organophosphates were developed to replace halogenated pesticides such as
dichlorodiphenyl trichloroethane (DDT) which were banned in 1979 along with other
chlorinated pesticides because halogenated pesticides are recalcitrant in nature and are
susceptible to biomagnification. The toxicity, mutagenicity and carcinogenicity have
raised public health concerns (Alexander, 1981). Although they are safer than the older
pesticides, the organophosphates are not without their problems. First, they break down
quickly in the soil, and so must be applied to crops more frequently. Second, these new
chemicals are not selective. Since the nerve action of larger animals are very similar,
organophosphates are capable of killing mammals, birds, reptiles, amphibians, and fish,
thereby resulting in unintended changes to the food web which becomes difficult to
predict. Third, animals that have died or been weakened by the toxin put other animals that
eat them at risk through bioamplification, (Thomson, 1989). The organophosphates are
comparatively short lived under most environmental conditions. They are rapidly
metabolized or excreted by most animals (Eto, 1974) and do not bioaccummulate in the
food chain.
Some types of organophosphates include:
a) Parathion:- Parathion and its methyl analog methyl parathion (O,O-dimethyl O-P-
nitrophenyl phosphorothioate). It is of the class phosphorothioate which are
organophosphate pesticides that are hydrolytically stable in aqueous solutions, and
biodegradable at various rates (MacRae, 1989). It is a potent neurotoxic pesticide
that kills bugs by disrupting vital transmitters in their nervous systems. Thus,
parathion has been widely used for controlling insects of agricultural and public
health importance (Mateen et al., 1994; Cook and Hutter, 1981). Although
10
parathions are considered to be less persistent, these pesticides and their
metabolites, particularly p-nitro phenols have caused environmental pollution
(Mateen et al., 1994). ρ-nitro phenol imparts odor problems in water.
Parathion is degraded in soils and water by three pathways, all of which require
acclimatization for complete expression (Figure 1):
1. Hydrolysis to ρ -nitrophenol and diethylthiophosphoric acid usually occurs in
aerobic soils. ρ-Nitrophenol is often degraded by reduction to ρ-aminophenol.
2. Reduction to aminoparathion, which is then hydrolyzed to p-aminophenol and
diethylthiophosphoric acid under low oxygen levels (microaerophilic) in
anaerobic environments.
3. Small amounts of oxidation to paraoxon. This is the main mammalian metabolic
pathway. Paraxon is further degraded to nitrophenol and diethyl phosphoric acid in
the environment. The p-nitrophenol is metabolized aerobically by mono-
oxygenation ortho to the hydroxyl, followed by loss of the nitroso group and
cleavage of the catechol. Alternately, the nitroso group is cleaved by an oxygenase
attack forming hydroquinone, which is further hydroxylated and cleaved. p-
Nitrophenol also degrades in flooded soils with the release of NO2 and CO2.
Parathion is degraded by a bacterium, Flavobacterium sp, an algae Chlorella
pyrenoidosa, and a fungus Penicillium waksmani. Degradation occurs in aerobic
and flooded (anaerobic) soils, often involving consortia of microorganisms.
11
Fig 1: Pathway for the degradation of parathion, indicating the pathways mentioned
above by the numbers 1, 2 and 3. (MacRae, 1989)
12
b) Diazinon:- This is another example of a phosphorothioate. Diazinon (O,O-
dimethyl 0-2-isopropyl-4-methyl-6-pyrimidyl phosphorothioate) is used as a soil
and foliar insecticide and is effective against a broad range of insect pests of crops
and ornamental plants (Mateen et al., 1994). It provides a good residual treatment
for control of fleas in barns and also used in household sprays and dusts for ants
and cockroach control (McEwen and Stephenson, 1979). It is a neurotoxin that
causes headaches, dizziness, nausea, blurred vision, impaired memory and motor
skills. Ecological effects include bird and fish mortality, interference with nitrogen
fixing soil organisms, and reduction in the numbers of eggs laid by birds. The
potential toxicity of diazinon to birds has brought about its suspension as an
insecticide of choice on golf courses (Anonymous, 1990). Little is known about the
degradation of diazinon, but Flavobacterium spp. ATCC 27551 was originally
isolated as a diazinon degrading bacterium. It was characterized with respect to
parathion hydrolysis (Mateen et al., 1994). Diazinon is hydrolyzed to diethyl
phosphorothioate and 2-isopropyl-4-methyl-6-hydroxypyrimidine, leading to
complete mineralization, often requiring consortia. Figure 2 depicts the pathway
for the degradation of diazinon
13
Fig 2: Pathway for the degradation of Diazinon (Aislabie and Lloyd-Jones, 1995).
14
c) Isofenphos:- Isofenphos (O-ethyl-O-2-isopropoxycarbonyl)phenyl] N-isopropyl
phosphoro-amidothiate] is a systemic nematicide (Fest and Schmidt, 1983).
Isofenphos was degraded more rapidly in soils with history of the insecticide than
in unexposed soils (Racke and Coasta, 1988). Soils with enhanced isofenphos
degradation, contained an adapted population of soil microorganisms. Two
bacterial isolates an Arthrobacter sp. and a Pseudomonas sp. obtained from
adapted cultures, metabolized the pesticide in pure culture (Racke and Coasta,
1988).
d) Dichlorovos:- Dichlorovos is under the phosphate class (2,2-dichlorovinyl O,O-
dimethyl phosphate). It is used extensively in vaponastrips, (a preparation in which
insecticides are impregnated in resin and volatilized at fairly uniform rates) to give
control of household pests, especially fleas (McEwen and Stephenson, 1979).
Dichlorovos is also effective against ectoparasites and is used in flea collars for
dogs and cats and a number of veterinary applications. A likely degradative
pathway is shown in Figure 3.
15
Fig 3: Pathway for the degradation of dichlorovos (Aislabie and Lloyd-Jones, 1995).
16
e) Coumaphos:- [O,O-diethyl O-(3-chloro-4-methyl-2-oxo-2H-1-benzo-pyran-7-yl)
phosphorothioate] is used as an acaricide for the control of the southern cattle tick
(Boophilus microplus) and the cattle tick (Boophilus annulatus). Coumphos was
used by the Animal and Plant Health Inspection Service (APHIS). Since the half
life of coumaphos in the soil and water is about 300-days (Kearney et al., 1986),
safe and effective methods for disposal of coumaphos waste are required.
f) Methidathion: - [S-(S-methoxy-2-oxo-1,3,4-thiadiazol-3-(2H)-yl)methyl]O,O-
dimethyl phosphorodithioate] is used for the control of insects on alfalfa, cotton
and fruit crops. It is also used in greenhouses, mainly for rose cultures against
thysanopterae and lepidopterae, and in vegetable nurseries (Mateen et al., 1994).
g) Fenitrothion: - (O,O-dimethyl-O-(4-nitro-m-tolyl)-phosphorothioate) is a contact
insecticide and a selective acaricide of low ovicidal properties (Spencer, 1981). It
is considered a cholinesterase inhibitor (Agrochemical Handbook, 1983).
Fenitrothion is effective against a wide range of pests such as penetrating, chewing
and sucking insect pests (e.g. coffee leaf miners, locust, rice stem borers, wheat
bugs, flower beetles) found on cereals, cotton, orchard fruits, vegetables and
forests. It may also be used as a fly, mosquito and cockroach residual contact spray
for farms and public health programs (Meister, 1994). Fenitrothion is also effective
against household insects and all of the nuisance insects listed by the World Health
Organization (Worthing, 1987). Fenitrothion is a non systemic and a non persistent
organophosphate.
h) Glyphosate: - it has been extensively used for insect and weed control and it is also
a broad spectrum pesticide.
17
1.4.5.1 ORGANOPHOSPHATE TOXICITY
Organophosphate toxicity is due to the ability of these compounds to inhibit
acetylcholinesterase at cholinergic junctions of the nervous system. These junctions
include postganglionic parasympathetic neuroeffector junctions (sites of muscarinic
activity), autonomic ganglia and the neuromuscular junctions (sites of nicotinic activity)
and certain synapses in the central nervous system. Acetylcholine is the neurohumoral
mediator at these junctions. Since acetylcholinesterase is the enzyme that degrades
acetylcholine following stimulation of a nerve, its inhibition allows acetylcholine to
accumulate and result in initial excessive stimulation followed by depression (Macmullan,
2006).
Some compounds have a direct effect on the inhibition of acetylcholinesterase
while others such as parathion are converted in the liver to metabolites which inhibit
acetylcholinesterase. In addition to the anti-acetylcholinesterase activity of these
compounds, Mipafox causes demyelination in peripheral nerves, causing lesions which
resemble those due to thiamine deficiency. Many of these compounds are excreted in milk
and are able to cross placental membranes causing toxicity in offspring (US. EPA, 2001).
Organophosphate compounds vary greatly in their toxic capabilities and have the
advantage over other types of insecticides in that they produce little or no tissue residues.
All have a cumulative effect with chronic exposure causing progressive inhibition of
cholinesterase. Liquid organophosphates are absorbed readily by all routes, although
malathion, which is the least toxic of these chemicals, is only slightly absorbed through
the skin (CDC, 2001).
18
1.4.6 GLYPHOSATE HERBICIDE
Glyphosate was first reported as a herbicide in 1971. Three related products are
now manufactured under the name glyphosate: glyphosate-isopropylammonium and
glyphosate-sesquisodium patented by Monsanto, and glyphosate-trimesium
(trimethylsulfonium), patented by Zeneca (Franz, et al., 1997). However, it does not affect
the nervous system in the same way as other organophosphate pesticides, and is not a
cholinesterase inhibitor. It inhibits 5-enolpyruvylshikimate-3-phosphate synthase which is
needed for the synthesis of three essential amino acids (US EPA, 2001; Cox, 1995).
Glyphosate (N-[phosphonomethyl]glycine) is a broad spectrum non-selective post-
emergence herbicide that was first used in the early 1970’s (Battaglin et al., 2002).
Glyphosate product sales are worth $1,200 million a year. In the US, glyphosate was used
on about 12 to 25 million acres annually in the 1980s. In the UK, it was used on almost
800,000 acres in 1994. It is broad spectrum in action. It is used to control a great variety of
annual, biennial, and perennial grasses, broad leafed weeds and woody shrubs (WHO,
1994; Cox, 2000) through direct contact with the leaves from where it is translocated to
the root of plants. It is used in fruit orchards, vineyards, conifer plantations and many
plantation crops (e.g. coffee, tea, bananas); in pre-crop, post-weed emergence in a wide
range of crops (including soybean, cereals, vegetables and cotton); on non-crop areas (e.g.
road shoulders and rights of way); in cereal stubble; forestry; gardening and horticulture.
Other uses of the salts of glyphosate are in growth regulation in peanuts and in sugarcane
to regulate growth and speed fruit ripening. Glyphosate can be used throughout the
growing season. It is widely used in no-till agriculture to prepare fields for planting,
controlling of weeds during crop development, or controlling perennial weeds after crop
harvest (United State Department of Agriculture (USDA), 2000). It has high activity when
applied to foliage (WHO, 2004). Glyphosate is used worldwide in both agriculture and
19
non-agricultural areas all over the world (IPCS, 1994; Woodburn, 2000). The major
commercial formulations of glyphosate include Roundup, Touchdown, Poledo, Accord
(US-EPA, 1993).
1.4.6.1 PROPERTIES OF GLYPHOSATE
Glyphosate is a weak organic acid that contains a glycine moiety as shown in
Figure 4. It is amphoteric and may exist as different ionic species, depending on the actual
pH. Its structural formula is as follows
20
Fig 4: Glyphosate Structure (IPCS, 1994)
21
Pure glyphosate has a relative molecular mass of 169.07. It is a white, odourless
crystal. It has a melting point of 1850C and decomposes at 1870C producing toxic fumes
including nitrogen oxides and phosphorous oxides. Other properties of glyphosate are
shown in Table 1. It has three functional groups; phosphonic acid, carboxylic acid and
secondary amine. It is a very stable compound that undergoes most reactions that
phosphonic acid, carboxylic acid or secondary amine will undergo. The most important
reactions glyphosate undergoes are esterification amination, dehydration, N-alkylation, N-
acylation, N-sulfonylation and the formation of acyl and phosphonyl halides. The
following are the pKa values of different ionic species of glyphosate (Figure 5):
22
Fig 5: Ionic species of glyphosate and their pka Values (Sprankle et al., 1975)
23
Table 1: Physical properties of Glyphosate (IPCS, 1994)
PROPERTY STATE REMARK
Physical state a Crystalline Powder
Relative molecular mass 169.07
Colour White
Odour None
Melting point b 184.50C Decomposition at 1870C
Boiling point Not applicable
Specific gravity (Density) c 1.704 200C
Vapour pressure <1x10-5Pa 250C
Solubility in water b, e 10-100mg/litre 200C
Octanol-water Partition
Coefficient (Log Kow) d
-2.8
Surface tension 0.072N/m 0.5%(w/v) at approximate
(Sprankle et al., 1975)
pKa values d, f <2, 2.6, 5.6, 10.6
Molar absorptivity c 0.086 litre/mol per cm At 295nm
Flammability d Not flammable
Explosiveness d Not explosive
pH d 2.5 1% solution
Key:
a) Data provided by Monsanto Ltd. b) Purity 96%. c) Purity 100%. d) Purity not
reported. e) Pure glyphosate has been reported to have a water solubility of 11-
600mg/litre at 250C. f) Free acid.
24
Pure glyphosate is slightly soluble in water (12 g/litre at 250C) and it is practically
insoluble in most organic solvents due to its high polarity in ethanol, acetone and benzene.
The alkali-metal and amine salts are readily soluble in water. This is thought to be as a
result of the strong and very extensive system of hydrogen bonding within the glyphosate
crystal lattice. The pKa values for glyphosate in aqueous solution correspond to the proton
dissociation reactions shown below. The values for pK1, pK2 and pK3 are 2.27, 5.58 and
10.25 respectively.
25
Fig 6: Glyphosate proton dissociation reactions (Franz, et al., 1997)
26
The fluids within plants, generally operates within very narrow pH ranges. Phloem
sap has a pH of 8-8.5; and xylem and apoplastic solutions, pH of 5-6. This means that
glyphosate exists in the apoplast mainly in its monoanionic form and it is translocated
about the plant mainly in its dianionic form. Glyphosate formulations are stable for
extended periods below 600C (IPCS, 1994)
It can be seen that glyphosate is an acid that can be converted to just about any salt via
its reaction with the appropriate base. For example metal salts can be obtained by reaction
of glyphosate with aqueous bicarbonate, carbonate and hydroxide solution. Glyphosate is
amphoteric in nature and so will dissolve in strong acids to produce salts with negative
pKa values and crystalline hemisalts have, also, been isolated. Glyphosate has pKa values
which show that it dissociates in three stages and hence, capable of forming mono-, di-
and tri- salts. The stability constants of the Cu, Zn, Ca and Mg complexes of glyphosate
have been reported.
When glyphosate is heated at 200-230oC it softens and then resolidifies. N,N'-
diphosphonomethyl-2,5-piperazinedione is formed. This occurs via a dehydration reaction.
The 2,5-piperazinedione formed is very water soluble and is thermally stable to 316oC.
Glyphosate can be regenerated by refluxing the 2,5-piperazinediones with strong mineral
acids such as hydrobromic acid (Figure 7).
27
Fig 7: Formation of N,N'-diphosphonomethyl-2,5-piperazinedione (IPCS, 1994).
28
Glyphosate has the reactivity typical of many secondary alpha-amino acids, but its
low solubility in aqueous solutions and organic solvents makes some transformations quite
difficult.
Glyphosate is a polar compound known for its adsorption to iron and aluminum
oxides and clay (Busse et al., 2001; Morillo et al., 1997). It is a weak acid that donates its
hydrogen ion to another compound. Thus when formulated into a commercial product, the
hydrogen ion on the parent weak acid is replaced with a different salt (ion).
1.4.6.2 MODE OF ACTION OF GLYPHOSATE
Glyphosate is very effective in killing all plant types including grasses, perennials
and woody plants. It is able to carry out this function by being absorbed into the plant
mainly through its leaves and also through soft stalk tissues. It is then transported
throughout the plant where it acts on various enzyme systems, by inhibiting amino acid
metabolism in what is known as the shikimic acid pathway. Plants treated with
glyphosate die slowly over a period of days or weeks and no part of the plant survives
because of the chemical transportation.
The glyphosate herbicide inhibits the synthesis of aromatic amino acid
(tryptophan, phenylalanine and tyrosine) in plants and microorganisms (Zablotowicz and
Reddy, 2004; Fisher et al., 1986; Arhens, 1994).
The mechanism of action of glyphosate is unique because unlike other
organophosphate pesticides, glyphosate specifically inhibits the enzyme 5-
enolpyruvylshikimic acid-3-phosphate synthase (Steinrucken and Amrhein, 1980), which
catalyses the condensation of shikimic acid and phosphoenolpyruvate (Figure 8).
Inhibition of the shikimic acid pathway by glyphosate results in the accumulation of
29
shikimic acid and/or certain hydroxybenzoic acid such as Protocatechuic and/or gallic acid
in sensitive plant species (Becerril et al., 1989).
The amino acids inhibited are essential for the growth and survival of plants. Two
of the three aromatic amino acids are essential amino acids in human diet because humans,
like other higher animals, lack the shikimic acid pathway and thus cannot synthesize the
amino acids. They therefore rely on their foods to provide these compounds. Glyphosate
can affect other enzymes not connected with the shikimic acid pathway. In sugarcane, it
reduces the activity of one of the enzymes involved in sugar metabolism (Su, 1992). It
also inhibits a major detoxification enzyme in plants (plant cytochrome P-450) which is
involved in the detoxification of some herbicides.
30
Erythrose-4-P + Phosphoenolpyruvate (PEP)
DAHP
Gallate Dehydroquinate
Protocatehuate Dehydroshikimate
3-oxoadipate Shikimate
TCA Cycle Shikimate-3-phosphate
Glyphosate -----------
5-enolpyruvylshikimate-3-Phosphate Synthase
Aromatic Amino acids Chorismate
Prephenate
Fig 8: Shikimic acid pathway and the inhibition by glyphosate in plants and
microorganisms.
Key:
Stripped arrows ( ) indicate the overall effects of glyphosate inhibition of 5-
enolpyruvylshikimic-3-phosphate synthase (ESPS) and pathways for accumulation of
hydroxybenzoic acids.
(Moorman et al., 1992).
31
The toxic effects of glyphosate may be attributed to the following:
• The inability of the organism to synthesize the needed aromatic amino acids
• An energy drain on the organism resulting from adenosine triphosphate and
phosphoenolpyruvate spent in the accumulation of shikimate, 3-deoxy-D-arabino-
heptulose-7-phosphate (DAPH), and hydroxybenzoic acids.
1.4.6.3 PERSISTENCE OF GLYPHOSATE IN THE ENVIRONMENT
The persistence of glyphosate in the environment is variable. It is strongly bound
to clay particles and is considered moderately persistent in soils. Glyphosate readily
binds to cations (Fe2+, Fe3+ and Al3+) that are adsorbed to soils (Carlisle and Trevors,
1988). This binding is unlike other organic compounds that primarily adsorb to
organic matter in soils. Studies show that the soil-binding potential of glyphosate is
stronger than that of any other herbicide. A ratio known as “soil adsorption
coefficient” (Koc) measures the soil-binding capacity of chemical compounds, with
higher numbers meaning greater adsorption of the compound to the soil.
Table two (2) shows representative Koc values for some herbicides, (US EPA,
1993)
32
Table 2: Some pesticides with their representative Koc values (USEPA, 1993)
Active Ingredient Koc Value
2,4-Dichlorophenol 109
Alachlor 170
Metolachlor 200
Trifluralin 7,000
Glyphosate 24,000
Pendimethalin 24,300
33
Glyphosate is also adsorbed by hydrous oxides (Glass, 1981; McConnell and
Hossner, 1985). The role of adsorption on the glyphosate bioavailability and
biodegradation is not completely conclusive. Viega et al. (2001) stated that the
adsorbed glyphosate is more persistent in the soil because its degradation is slower
than that of the free herbicide.
Glyphosate binding is similar to phosphate binding, and it is possible that
phosphate accumulation in soils could reduce the capacity for glyphosate binding
(Gimsing et al., 2004). Glyphosate adsorption correlates with the amount of vacant
phosphate sorption sites and may occur through binding of the phosphonic acid moiety
(Arhens, 1994).
The active ingredients in some herbicides are volatile, meaning that they can move
as vapours to non-target areas after application. This can result in unintended
consequences on sensitive plant species outside the treated area. Several laboratory
studies show that glyphosate has a very low vapour pressure, suggesting that loss to
the atmosphere from treated surfaces will be small (Battaglin et al., 2005; Giesy et al.,
2000).
Although glyphosate is not recommended for direct application in soil, a
significant amount may reach the soil during early-season or preplant applications
(Haney et al., 2000). Glyphosate has a half life of 2 -197 days in the soil and that of its
breakdown product ranges from 76-240 days. Biodegradation of glyphosate in the soil
is dependent on the amount of herbicide available to soil microorganisms and this
depends on various factors including available nutrients, pH, temperature and soil
type.
In aquatic environments, the half life of glyphosate and its breakdown product,
aminomethylphosphonic acid (AMPA) is reported to range from 7 to 14 days (Giesy et
34
al. 2000). Monsanto (2003) and Giesy et al (2000) suggested that the occurrence and
persistence of glyphosate in surface run-off would be similar. This is because the
physical and chemical properties of glyphosate suggest a low probability that it will
runoff from fields, persist in surface water, or leach through soils to ground water.
1.4.7.0 GLYPHOSATE TOXICITY
1.4.7.1 Toxicity and Effects on Experimental Animals and In Vitro systems.
The acute toxicity of glyphosate is very low (Monsanto, 1992). Glyphosate acute
oral median dose (that is the dose that causes death in 50 percent of test animals; LD50)
in rats is greater than 4,230mg/kg of body weight (WHO, 1996; IPCS, 1994; Cox,
2000). The low acute toxicity of Glyphosate can be attributed to its biochemical mode
of action through a metabolic pathway in plants called the shikimic acid pathway
which does not exist in animals (Carlisle and Trevors, 1988).
It has been reported that technical grade glyphosate (98.7% purity) fed to
laboratory mice in diet dose levels of 0%, 0.5%, 1.0%, or 5.0% revealed an increase in
weight of several organs (liver, brain, heart and kidney). Also, growth retardation was
observed at dose levels of 5% (IPCS, 1994; WHO, 2004).
Glyphosate can also disrupt the functions of enzymes in animals. In rats, it was
found to decrease the activity of some detoxification enzymes when injected into the
abdomen (Cox, 1995). It is markedly more toxic by intraperitoneal route than by other
routes (IPCS, 1994).
Commercial glyphosate herbicides are more acutely toxic than pure glyphosate
(Martinez and Brown, 1991; Agriculture Canada, 1991). Glyphosate containing
products are more toxic via inhalation than orally (IPCS, 1994). Inhalation of Roundup
by rats caused gasping, congested eyes, reduced activity and body weight loss
35
(Agriculture Canada, 1991). It also caused lung damage when given intravenously to
dogs; and increased the ability of the heart muscle to contract causing cardiac
depression (Tai, 1990).
1.4.7.1.1 Sub chronic Toxicity in Laboratory Animals
Medium term studies on rats and mice carried out by the National Toxicology
Program (NTP) (1992) showed that salivary gland lesions occurred in all doses (200-
3400mg/Kg). Blood levels of potassium and phosphorus in rats also increased in all
doses tested (60-1600mg/Kg). Glyphosate containing products are more toxic than
glyphosate in sub chronic tests. In a seven day study with calves, 790mg/Kg per day of
Roundup caused pneumonia, and death in one-third of the animals tested. At lower
doses decreased food intake and diarrhoea were observed (WHO, 1994).
1.4.7.1.2 Chronic Toxicity in Laboratory Animals
Few effects were observed at all but the lowest dose tested. Glyphosate caused
inflammation of the stomach lining in both male and female mice (US.EPA, 1993).
Growth retardation, hepatocyte hypertrophy and excessive cell division in the urinary
bladder in male mice were also observed at 30-1000mg/Kg (IPCS, 1994).
1.4.7.2 Acute Toxicity in Humans
Doctors in Japan first exposed the acute hazards of surfactants in glyphosate
products in a study of 56 cases of Roundup poisoning, mostly resulting from suicides
or attempted suicides, which included nine fatalities. Rough estimates of the amounts
ingested in lethal cases varied from 85 to 200ml (corresponding to 30 to 70g of
Glyphosate acid) (Sawada et al., 1988). However, larger amounts (up to 500ml) were
reported to have been ingested by patients with mild or moderate symptoms (WHO,
1996). A variety of symptoms have been noted from acute poisoning incidents. They
include; intestinal pain, vomiting, excess fluid in the lungs, pneumonia, lung
36
dysfunction, low blood pressure, damage to the larynx and destruction of the red blood
cells ( Talbot et al., 1991; Temple and Smith, 1992).
1.4.7.3 Reproductive Effects
Glyphosate exposure has been linked to reproductive problems in humans. In a
study conducted in Canada, it was found that in fathers that use glyphosate there was
an increase in miscarriages and premature births (Savitz, 1997). In rats, glyphosate
reduced sperm counts at the two highest doses tested. In male rabbits, glyphosate at
doses of one-tenth and one-hundredth of the LD50 increased the frequency of abnormal
and dead sperms (Welsh, et al., 2000).
1.4.7.4 Mutagenicity
Glyphosate and glyphosate-containing products have been reported to be
mutagenic. However, it is to be noted that glyphosate-containing products are more
potent mutagens than glyphosate itself. This could be as a result of surfactant and other
substances used in the formulation of commercial products (Bolognesi et al., 1997).
In fruit flies, Roundup was found to increase the frequency of sex-linked, recessive
mutations. These mutations are only visible in males (Cox, 2000). In another study of
human lymphocytes an increase in the frequency of sister chromatid exchanges
following exposure to the lowest dose of Roundup tested was observed (Bolognesi et
al., 1997). In mice injected with Roundup, the frequency of DNA adducts (the binding
to genetic material of reactive molecules that lead to mutations) in the liver and kidney
increased at all three doses tested (Peluso et al., 1998).
37
1.4.7.5 ECOLOGICAL EFFECTS OF GLYPHOSATE
1.4.7.5.1 Effect of Glyphosate on Non Target Animals
A) Beneficial Insects: - Glyphosate treatment has reduced populations of beneficial
insects, birds, and small mammals by destroying vegetations on which they depend for
food and shelter (Cox, 2000). These beneficial insects are insects that kill other species
that are agricultural pests. The International Organization for Biological Control
(IOBC) found that exposure to freshly dried Roundup killed over 50 percent of 3
species of beneficial insects; a parasitic wasp, lacewing and a lady bug (Burst, 1990).
B) Other Insects: - Roundup treatment of a field caused an 89% decline in the
number of herbivorous plant eating insects because of the destruction of the vegetation
on which they live and feed. These insects serve as food resources for birds and insect
eating small mammals (Santillo et al., 1989). Glyphosate and glyphosate-containing
products kill a variety of other arthropods. In one laboratory study, over 50 percent of
test populations of a beneficial predatory mite were killed by exposure to Roundup
(Cox, 2000). Also in a similar study, Roundup exposure caused a decrease in survival
and a decrease in body weight of woodlice. These arthropods are important in humus
production and soil aeration (Cox, 2000). Roundup and some other formulations of
glyphosate are slightly toxic to earthworms with 14-day No-observed-effect
concentration values of 500 and 158mg product per kilogram dry weight respectively.
Increase in maturity time and increase in mortality was also observed (IPCS, 1994).
C) Fish:- Glyphosate is relatively non-toxic to fish, with a 24-96 hour LC50
(concentration of a chemical calculated to kill 50% of test animals) values ranging
from approximately 10ppm in acidic water (pH 6) to >200ppm in alkaline water
(USDA, 2000; Gardner and Grue, 1996). Other commercial products that contain
38
glyphosate are acutely toxic to fish (Cox, 2000). They have intermediate toxicity due
to the surfactants present in them.
Acute toxicities of glyphosate vary widely; median lethal concentrations i.e.
concentrations killing 50 percent of a population test animals as have been stated
above have been reported depending on the species of fish and test conditions.
Roundup toxicity increases with water temperature. In both rainbow trout and blue
gills, toxicity was shown to double between 70C and 170C (Abdelghani et al., 1997).
Treatments along river banks (riparian) with glyphosate, causes water temperature to
increase for several years because the herbicide kills shading vegetation (Holtby,
1989). Glyphosate also causes difficulty in breathing in trouts and gill damage in carp
(USDA, 2000).
D) Birds: - Glyphosate has indirect impacts on birds. Because it kills plants, its use
can create a dramatic change in the structure of the plant community. This affects bird
populations, since birds depend on the plants for food, shelter, and nest support (Cox,
2000).
1.4.7.5.2 Effects on Non Target Plants
Glyphosate as stated earlier is a broad spectrum herbicide which has potent acutely
toxic effects on most plant species. There are also other kinds of serious effects on
endangered species, reduced seed quality, reduction in the ability to fix nitrogen,
increased susceptibility to plant disease and reduction in the activity of mycorrhizal
fungi (Cox, 2000).
• Endangered Species and Seed quality:- Most plants are susceptible to
glyphosate. It can have a serious impact on endangered plant species. Sub
lethal treatment of cotton with Roundup “severely affects seed germination and
39
vigour. Seed germination was reduced between 24 and 84 percent and seedling
weight was reduced between 19 and 83 percent (Locke et al., 1995).
• Nitrogen Fixation:- Cultures of soil bacteria have shown the effects of
glyphosate on nitrogen fixation, denitrification and nitrification (Cox, 2000).
The processes of nitrogen fixation and nitrification are carried out by bacteria
which can be found in soil and in nodules on roots of legumes and other plants
(Atlas and Bartha, 1997; Raina, et al., 2000; Hutchinson, 1995). From studies
showing effects of glyphosate on nitrogen fixation, it was observed that at
concentrations corresponding to typical application rates, glyphosate reduced
by 70 percent the number of nitrogen fixing nodules on clover planted 120
days after treatment. The authors could not conclude whether the reduction was
due to direct effect of glyphosate on the bacteria, or on plant process that
support nitrogen fixation (Eberbach and Douglas, 1983). A similar study
showed that a concentration of glyphosate reduced by 20 percent nitrogen-
fixation by soil bacteria (Santos and Flore, 1995). Most of the studies
mentioned above were carried out in the laboratory and the results stated
above have corroborated in field experiments.
• Mycorrhizal Fungi: - Laboratory studies have shown that Roundup is toxic to
mycorrhizal fungi. Mycorrhizal fungi are beneficial fungi that live in and
around plant roots (Raina et al., 2000). They help plants absorb nutrients and
water and can protect them from cold and drought (Cox, 2000). In orchids,
treatment with glyphosate changed the mutually beneficial interaction between
the orchid and its mycorrhizae into a parasitic interaction which does not
benefit the plant (Bayne et al., 1995).
40
1.4.8 GLYPHOSATE DEGRADATION
Various processes have been described for the removal of pesticides from soil and
water. The removal/degradation process could be by chemical or microbiological process.
Chemical degradation occurs through reactions such as photolysis, hydrolysis, oxidation
and reduction (Andreu and Pico, 2004). These processes are not effective in the removal
of glyphosate and other pesticides because the bonds present in these pesticides such as
the carbon-phosphorus bond in glyphosate are highly stable to these reactions.
41
Fig 9: Abiotic and Biodegradation Processes for glyphosate degradation in the
environment (IPCS, 1994)
42
1.4.8.1 Abiotic degradation
1.4.8.1.1 Hydrolytic cleavage
Hydrolysis of glyphosate in sterile buffers is very slow. After 32 days less than 6.3%
of the applied glyphosate was recovered as aminomethylphosphonate (AMPA), after
applying 14C-glyphosate at rates of 25 and 250 mg/litre to aqueous buffer solutions of pH
3, 6 and 9 (Monsanto, 1978b). These tests were performed at both 5 and 35 °C.
1.4.8.1.2 Photodegradation
Photochemical degradation in water may occur under laboratory and field
conditions, depending mainly on the type of light source. In sterile aqueous buffers of pH
5, 7, and 9, less than 1% of the applied dose was degraded (photodecomposition of 14C-
phosphonomethyl-labelled glyphosate) during 29-31 days, when exposed to sunlight
(IPCS, 1994).
Lund-Hoie & Friestad (1986) in IPCS 1994) exposed Roundup to several light
sources under different conditions. When exposed to UV light (lambda = 254 nm) under
laboratory conditions, concentrations of 1 and 2000 mg active ingredient/litre in deionized
water showed DT50 values of 4 and 14 days, respectively. When exposed to sunlight under
field conditions 1 mg active ingredient/litre in polluted water without sediment showed a
much slower decomposition (DT50 > 63 days). This was possibly due to pollution
preventing adequate UV penetration in the water. Polluted water with sediments showed a
rapid dissipation from water, probably due to adsorption onto the sediments. In another
field experiment 2 and 100 mg active ingredient/litre in deionized or polluted water
without sediment showed DT50 values of < 28 days, when exposed to sunlight. At the low
concentration the dissipation in polluted water was more rapid than in deionized water. In
the dark no dissipation occurred.
43
In laboratory experiments 1 mg/litre of glyphosate in sterilized natural and deionized
water showed DT50 values of 4 to > 14 days when exposed to artificial light (350-450 nm)
in photoreactors without sediment (Monsanto, 1978a). In these experiments Ca2+ acted as
a photosensitizing agent.
Photodegradation by sunlight of glyphosate applied to a soil appeared to be an
insignificant route of dissipation (PTRL Inc., 1989). In this study, 14C-glyphosate mixed
with unlabelled glyphosate was exposed for 31 days to natural sunlight, after application
to a sandy loam at a rate of 4.5 kg active ingredient/ha. Extrapolated DT50 values that were
based on first-order kinetics were 90 days in the sunlight and 96 days in the dark,
indicating no substantial degradation due to photolysis. The temperature of the soil surface
was 22-23 °C. Under unnatural light conditions glyphosate appeared not to be
photodegraded substantially (Monsanto, 1972; Rueppel et al., 1977; Monsanto, 1978a).
1.4.8.2 Biodegradation
A number of bacteria have been found that degrade phosphonates (compounds that
contain a carbon to phosphorus bond), including glyphosate (N-phosphonomethyl glycine)
which is a potent, widely used broad spectrum herbicide. The earliest studies of bacterial
metabolism of glyphosate were performed with mixed bacterial cultures of soil-water
mixtures to simulate the ecological fate of glyphosate in soil (Nomura and Hilton, 1977;
Ruepple et al., 1977; Sprankle et al., 1975). Studies of glyphosate degrading bacteria have
involved selection for, and isolation of pure bacterial strains with enhanced or novel
detoxification capabilities for potential uses in biotechnology industry.
Microorganisms known for their ability to degrade glyphosate in soil and water
include Pseudomonas sp strain LBr (Jacob et al, 1988), Arthrobacter atrocyaneus (Pipke
et al., 1987), certain members of the Rhizobiaceae (Liu et al., 1991), including
44
Binorhizobium melitoli, Rhizobium trifolii, Rhizobium leguminosarurm, Agrobacterium
rhizogenes, Agrobacterium tumefaciens. Others include Bacillus subtillis, Escherichia coli
(Fischer et al., 1986; Quang, 1988), Bacillus cereus (Rosenberg and Nauze, 1967),
Flavobacterium sp (Balthazor and Hallas, 1986), Spirulina spp. (Lipok et al., 2007).
Bacteria degrade glyphosate via two general pathways leading to the intermediate
production of either glycine or aminomethylphosphonate (AMPA). Microorganisms
known to degrade glyphosate by way of glycine include Pseudomonas sp. strain PG2982
which breaks down glyphosate by utilizing it as its source of phosphorus by breaking
down the carbon-phosphorus bond. This results in the release of a phosphate group and a
molecule of sarcosine. The sarcosine is cleaved by a sarcosine-oxidizing enzyme
(sarcosine oxidase-dehydrogenase) present in Pseudomonas sp. strain PG2982 to glycine
and formaldehyde, and then the glycine produced is used for the biosynthesis of proteins
and purine bases (Shinabarger and Braymer, 1986).
Jacob et al. (1988) in their work reported that a Pseudomonas strain LBr is able to
degrade glyphosate via both pathways stated above. They also pointed out that the
bacterium was able to completely degrade glyphosate at concentrations as high as 19mM.
In another experiment carried out by Jacob et al. (1985), using solid state NMR in
the determination of glyphosate metabolism, in a Pseudomonas sp, it was reported that the
bacterium cleaved glyphosate directly to glycine. The phosphonomethyl carbon of the
glyphosate molecule enters into the tetrahydrofolate (THF) directed pathway of single-
carbon transfers, and the phosphate group, the only source of phosphorus for the
bacterium is used for growth.
A second group of bacteria, represented by a Flavobacterium sp strain GD1
(Balthazor and Hallas, 1986), and some other mixed bacterial strains are able to degrade
glyphosate by cleaving its carboxymethyl C-2 carbon-nitrogen bond to produce AMPA
45
(Jacob et al., 1988; Nomura and Hilton, 1977). AMPA is then cleaved by other
microorganisms with the ability to metabolize C-P bonds. Thus the complete elimination
of glyphosate from the soil, results from co-metabolism. Some of the AMPA generated in
this way can be further metabolized providing phosphorus for growth.
46
Fig 10: Degradation routes of glyphosate in soil (Liu et al., 1991)
47
CHAPTER TWO
2.0 MATERIALS AND METHODS
2.1 Isolation and Characterization of Glyphosate Degrading Bacteria from Rice
Field Soil Samples
2.1.1 Collection of Soil Samples
Soil samples were obtained from four rice fields located at Adani in Enugu, Omor
and Omasi in Anambra and Abakaliki in Ebonyi states, all in Southeastern Nigeria. These
rice fields are known to have been previously exposed to glyphosate-based formulation
(Roundup®) for long periods of time. Soil samples were collected with a sterile scoop
from a depth of 0-15 cm from three different sites in each of the four locations. Soil
samples from each site were thoroughly mixed and placed in sterile polyethylene bags.
They were taken immediately to the laboratory and stored at 40C before use within 72 h
(Kassem and Nannipieri, 1995).
48
Table 3: State and location from where the soil samples were collected
State Location Code
Anambra Omor
Omasi
OMR
OMS
Enugu Adani ADA
Ebonyi Abakaliki AKL
49
2.1.2 Estimation of soil moisture content
The moisture content of each soil sample was measured according to the method
described by Pansu and Gautheyrou (2003).The weighing bottles (tare) were dried for 2 h
at 105°C, allowed to cool in a desiccator and the weight of the tare (m0) was taken with
the lid placed underneath. Five grammes of air-dried soil (fine earth sieved through a 2
mm mesh) was placed in the tare box and the new weight taken as m1. The weighing
bottles with their flat caps underneath were placed in a ventilated drying oven for 4 h at
105°C (the air exit must be open and the drying oven should not be overloaded). The
bottles with the soil samples were cooled in a desiccator and the weight was measured as
m2 (all the lids of the series contained in the desiccator were closed to avoid moisture
input). The opened weighing bottles were again placed in the drying oven for 1 h at 105°C
and weighed under the same conditions; this process was continued till a constant weight
was attained.
% moisture content at 1050C = m1 – m2
m1 – m0
2.1.3 Isolation medium
A modified mineral salts medium (MSM) of Dworkin and Foster (1958) consisting
of (g/l) (NH4)2SO4, 0.375; MgSO4, 0.075; CaC03, 0.03; FeSO4.7H2O, 0.001; H3BO3,
0.000001, MnSO4, 0.000001, yeast extract, 0.0053 was used.. Phosphate buffer was
replaced by tris buffer (6.05 g/L) and pH adjusted to 7.0. All glasswares were washed with
1 N HCl and thoroughly rinsed with deionized water to remove contaminating phosphate
before use. The medium was autoclaved at 1210C and 15 psi for 15 min prior to the
addition of the filter sterilized Roundup® (isopropylamine salt of glyphosate) and glucose
(1.0 g/l) autoclaved at 1100C and 10 psi as carbon source.
100 x
50
2.1.4 Isolation of glyphosate utilizing bacteria
The soil samples were air-dried and sieved using a 2 mm mesh. Five gram of each
soil sample was suspended in 250-ml Erlenmeyer flask containing a mixture of 50-ml of
mineral salts medium and 1 ml of Roundup® (7.2 mg/ml of glyphosate). This
concentration was used because it is equivalent to the field application rate). The flasks
were incubated on a rotary shaker (Gallenkamp, England) at 120 rpm for 7 days at 300C.
The above steps were repeated by taking 1 ml of sample from each broth culture and
transferring to fresh enrichment medium followed by incubation as described for 7 days.
Isolation was done using the spread plate method on the solid mineral salts medium
described above with added glyphosate. The plates were incubated at 300C for 5 days.
Morphologically distinct colonies were isolated and were repeatedly sub-cultured on
nutrient agar (Fluka). Identity of the isolates was affirmed after characterization by
standard bacteriological methods (Holt et al., 1994; Cheesbrough, 1984).
2.1.5 Storage
Pure cultures were maintained on nutrient agar slants and on slants of enrichment
medium containing 15 g/L agar and stored at 40C and they were routinely sub-cultured on
the same media.
2.2 Inoculum Preparation and standardization
Inocula used for the study were prepared by inoculating isolates into nutrient broth
and incubated at 300C for 24 h. Using sterile normal saline, the cells from the above
cultures were re-suspended to a 0.5 McFarland nephelometer standard (optical density of
0.17 at 660 nm).
51
2.2.1 Glyphosate utilization patterns of the different isolates
A 1.0 ml portion of each isolate was inoculated into 150-ml of the screening medium
(contained in a 500-ml flask) which is the isolation medium without yeast extract. It
contained 3-ml of roundup (7.2 mg/ml of glyphosate). The flasks were incubated on a
rotary shaker (Gallenkamp, England) at 120 rpm for 180 h at 300C. The ability of each
isolate to utilize glyphosate was measured based on the turbidity of the medium at 660nm
using a spectrophotometer (Spectronic 20, USA).
2.3 Identification and characterization of Isolates
The isolates from the screening procedure were identified by conventional
microbiological and biochemical procedures according to the WHO Manual for the
Laboratory Identification and Antimicrobial Susceptibility Testing of Bacterial Pathogens
of Public Health Importance in the Developing World (WHO, 2003). Microscopic
examination of cell morphology, biochemical and substrate utilization tests were used to
characterize and identify the organisms. The following media were used in the
identification of the isolates; nutrient broth, nutrient agar, sulphide indole motility agar
(Oxoid), Kligler iron agar (Lab M), urea agar base (Oxoid), Simon’s citrate agar. All
culture media except where otherwise indicated, were products of Fluka laboratories
2.3.1 Microscopic Examination of Cell Morphology
2.3.1.1 Gram Stain
A representative of each distinct colony type was Gram stained according to the
method described in Prescott et al. (2002). A smear of each isolate was prepared on a
clean grease free glass slide with the aid of a wire loop by putting a drop of sterile saline
and heat fixed. The smear was then flooded with solutions of crystal violet (comprising
52
10g of crystal violet, 100ml ethanol; 1% of ammonium oxaloacetate) and was allowed to
stand for 30 s and then rinsed with water. It was covered with Gram’s (Lugols) iodine (2
g potassium iodide in 300 ml distilled water plus 1 g iodine crystals) for 1 min and rinsed
with water, acetone alcohol (95% ethanol and/or isopropanol-acetone mixture (3:1 v/v))
was used to decolorize the cell and then rinsed immediately with water. Finally, safranin
was used in counter staining the smear for 30 s, rinsed with water, air dried and observed
under light microscope using oil immersion objectives lens (x 100).
2.3.2 Biochemical Tests
2.3.2.1 Catalase Test
On clean grease free slide, a few drops of 3% v/v hydrogen peroxide (H2O2) was
dropped and the isolates emulsified on the slide. A vigorous evolution of oxygen bubbles
or effervescence is indicative of a positive result. This test shows the ability of the bacteria
to produce catalase enzyme that breaks down H2O2 (which is usually toxic to
microorganisms) to water and oxygen (Wistreich, 1997).
2H2O2 2H2O + O2
2.3.2.2 Oxidase Test
This test is used to detect the presence of cytochrome C (Wiestreich, 1997). It
demonstrates the presence in certain bacteria of oxidase enzyme which catalyses the
transport of electrons between donors in the bacteria and a redox dye (tetramethyl-para-
phenylenediamine hydrochloride). This dye differentiates colonies of oxidase-producing
bacteria from those not producing the enzymes.
The test was carried out by soaking filter papers in the reagent prepared by
dissolving 1 g of dye in 10 ml of deionized water. The isolates were then smeared on the
53
filter papers. Positive result is observed by the appearance of blue or deep purple
colouration within one min of application. It confirms the presence of the oxidase enzyme.
An absence of colour indicates a negative result.
2.3.2.3 Sulfide-indole-motility (SIM) screening test
Sulfide-indole-motility medium (SIM) is a commercially available combination
medium that combines three tests in a single tube: hydrogen sulfide (H2S) production,
indole production, and motility.
SIM medium was prepared from the dehydrated medium according to
manufacturer’s instruction. Escherichia coli, which is indole positive, H2S negative and
motility positive was used as control. The SIM medium were inoculated with a straight
inoculating needle, by making a single stab about 1–2 cm down into the medium and
incubating overnight at 37°C, according to the protocol in the District Laboratory Practice
in Tropical Countries (Cheesbrough, 2004). The surface of the motility agar was dry when
used, since moisture is known to cause a non-motile organism to grow down the sides of
the agar creating a haze of growth and appearing to be motile. The motility reaction in
SIM was indicated by the presence of diffuse growth (appearing as clouding of the
medium) away from the line of inoculation. The organisms that did not grow out from the
line of inoculation were recorded as non-motile. As in Kligler iron agar, H2S production
was indicated by blackening of the medium. Indole production was tested by adding 0.5
ml of Kovac’s reagent to the tube. Indole positive organisms produced red colour at the
top part of the test tube after shaking.
54
2.3.2.4 Starch Hydrolysis Test
This test as described by Wiestreich (1997) demonstrates the production of
amylase by the isolates. Plates of nutrient agar containing 0.2% soluble starch was
inoculated with the isolates and incubated for 5 days at 300C. The plates after incubating
were flooded with Lugol’s iodine solution. If blue black colouration is observed then the
starch in the medium has not been hydrolyzed. However, a clear colourless zone indicates
hydrolysis.
2.3.2.5 Nitrate Reduction
This is used to show the ability of the isolate to reduce nitrate to nitrite with the aid
of the enzyme nitrate reductase according to the method of Wiestreich, (1997). Nitrate
broth comprising 1 g of KNO3 in 1000 ml of nutrient broth was inoculated with isolates
and incubated at 300C for five days. Exactly 1.9 ml of nitrite reagent A, followed by 1.0
ml of reagent B will be added after incubation. A deep red colour showed the presence of
nitrite, thus indicating that nitrate has been reduced. Lack of colour change indicates a
negative result. Reagent A is prepared by dissolving 8 g of sulphanilic acid in one litre of
5 N acetic acid, while reagent B contained 5 g of naphthylamine in one litre of 5 N acetic
acid.
2.3.2.6 Gelatin Hydrolysis
This medium was used for the detection of proteolysis which indicated liquefaction
of gelatin due to the production of gelatin hydrolysing enzymes as described by
Wiestreich, (1997). The medium was prepared by dissolving 6.49 g of gelatin and 5 g
nutrient agar completely in 50 ml of deionized water and then dispensed into sterile
containers in 10 ml aliquots, before being autoclaved. After allowing to cool, the isolates
55
were inoculated by stabbing into the containers and thereafter incubate at 300C for 48 h.
After incubation, the tubes were further cooled to 220C to see if the gelatin will liquefy or
not. A positive result was observed if liquefaction of gelatin occurs at 220C.
2.3.2.7 Growth on Kligler iron agar
This test was carried out to determine the ability of isolates to ferment glucose and
lactose with the production of acid. In most situations, presumptive identification was
based on the reaction of the isolate on Kligler iron agar (KIA). Colonies were carefully
picked from agar media by selecting one discrete colony. Extreme care was taken to avoid
picking up contaminants that might be present on the surface of the agar by making sure
that the inoculating needle did not go through the colony to touch the surface of the plate
as describe in the protocol in the District Laboratory Practice in Tropical Countries
(Cheesbrough, 2004). Tubes of KIA were inoculated by stabbing the butt and streaking the
surface of the slant. The caps of the tubes were loosened before incubation. After
incubation for 24 hours at 37°C, the KIA slants were observed for reactions and the result
recorded. Yellowing of the butt indicated glucose fermentation; yellowing of slant
indicated lactose fermentation; while reddening of the slant indicated inability of the
organism to ferment lactose. Gas production was indicated by air bubbles, cracks or
displacement of the medium. Hydrogen sulphide production was indicated by blackening
of the medium.
2.3.2.8 Sugar Fermentation Test
This test was carried out to determine the ability of isolates to metabolize sugar
with the production of acid. The sugars prepared and used for the test include mannitol,
sorbitol, inositol, xylose and inulin. One percent Peptone water was used as the base
56
medium for fermentation. To a 100 ml solution, 0.1 g methyl red was added as indicator.
The peptone water with the indicator was sterilized at 1210C for 15 mins. The sugars (1.0
g each) to be tested was sterilized separately by autoclaving at 1100C for 10 mins and
allowed to cool before adding to the peptone water. The medium was then inoculated with
pure isolates using sterile loops; and afterwards incubated at 300C for 48 h. A change in
colour from red to yellow indicated acid production.
2.3.2.9 Citrate utilization
The citrate utilization test is a selective test for bacteria that have the ability to
consume citrate as their sole source of carbon and ammonium as sole nitrogen source. The
test carried out in this study was the method using Simmon’s citrate agar. Slopes of the
medium in bijou bottles were prepared according to the manufacturer’s instructions. Using
a sterile straight wire, the slope was streaked first with a saline suspension of the organism
and the butt-stabbed, according to the protocol in the District Laboratory Practice in
Tropical Countries (Cheesbrough, 2004). The bottles were then incubated at 35˚C for 48
hours. Bacteria that metabolized citrate produced an acid end product that changed the
colour of the medium blue and were recorded as positive. Those that remained green were
recorded as negative.
2.4 BIODEGRADATION STUDY IN LIQUID MEDIUM
2.4.1 Inoculum Preparation and Standardization
Inocula for this study were prepared by inoculating isolates into nutrient broth and
incubating at 300C for 24 h. Using sterile normal saline, the cells from the above cultures
were re-suspended to a 0.5 (optical density of 0.17 at 660 nm) McFarland nephelometer
standard. This comparison was made easy by viewing the tube against a sheet of white
57
paper on which sharp black lines were drawn. The turbidity standard was agitated on a
vortex mixer immediately prior to use. The turbidity of the bacterial suspension was
adjusted to the proper density (0.5 McFarland turbidity standards) by adding sterile broth
or adding more bacterial cells.
2.4.2 Preparation of Turbidity Standard
This standard was prepared as described by Baron et al. (1990). McFarland
turbidity standard (0.5) was prepared by adding 0.5 ml of a 1.175% (w/v) barium chloride
dihydrate (BaCl2.H20) solution to 99.5 ml of 1% (v/v) sulfuric acid (H2SO4). The turbidity
standard was then aliquoted into test tubes identical to those used to prepare the inoculum
suspension. The McFarland turbidity standard tubes were properly sealed to prevent
evaporation and stored in a black polyethylene bag in the dark at room temperature. Fresh
tubes were prepared on weekly basis or any time reduction in volume was noticed. (The
tube was marked to indicate the level of liquid, and checked before use to be sure that
evaporation did not occur). Before each use, the tube containing the turbidity standard was
shaken well, so that the fine white precipitate of barium sulfate was mixed in the tube. The
composition of McFarland turbidity standards and the corresponding densities of bacteria
(cfu/ml) are presented in Appendix II. The accuracy of the density of a prepared
McFarland turbidity standard was confirmed by using a spectrophotometer with a 1-cm
light path; for the 0.5 McFarland turbidity standard, the absorbance at a wavelength of 660
nm was 0.17.
2.4.3 Growth kinetics of Pseudomonas fluorescens and Acetobacter sp in glyphosate
Erlenmeyer flasks (500-ml) containing 150-ml of the sterile screening medium
(already described) was prepared and 3 ml of roundup (containing 7.2 mg/ml of
58
glyphosate) was added to each flask. 1-ml of inoculum (0.5 Macfarland standard) of each
selected isolate (P. fluorescens and Acetobacter sp) was used to inoculate each flask
(experiments were carried out in 3 replicates). The two isolates used were selected based
on their utilization patterns. The overall medium was incubated at 300C for 192 h on a
shaker at 120 rpm. A 5-ml volume of the culture medium was collected from each flask at
twelve hourly intervals and assayed for growth by measuring the optical density at 660 nm
using a spectrophotometer.
2.4.4 Comparative role of glyphosate as carbon or phosphorus source
The screening medium (150-ml) was prepared as earlier described and 3.0 ml
filter-sterilized roundup (containing 7.2 mg/ml of glyphosate) was added as phosphorus or
carbon source. When used as carbon source, denoted by Gly and Pi, the medium consisted
of the following (g/L): (NH4)2SO4, 0.375; MgSO4, 0.075; CaC03, 0.03; FeSO4.7H2O, 0.001;
H3BO3, 0.000001; MnSO4, 0.000001; NaHPO4, 6.0 and KH2PO4, 2.0. When used as
carbon and phosphorus source, denoted by (Glyphosate), the medium consisted of the
following (g/L): (NH4)2SO4, 0.375; MgSO4, 0.075; CaC03, 0.03; FeSO4.7H2O, 0.001;
H3BO3, 0.000001; MnSO4, 0.000001; tris buffer, 6.05g. When used as phosphorus source
denoted by (Gly and Glu), the medium consisted of the following (g/L): (NH4)2SO4, 0.375;
MgSO4, 0.075; CaC03, 0.03; FeSO4.7H2O, 0.001; H3BO3, 0.000001; MnSO4, 0.000001;
glucose, 1.0; tris buffer 6.05g. The media were incubated at 300C for 120 h on a shaker at
120 rpm. A 5-ml volume of the culture medium was collected from each flask at 12-h
intervals and assayed for growth by measuring the optical density at 660 nm using a
spectrophotometer.
59
2.4.5 Effects of different Concentrations of Glyphosate on the Growth of the
Isolates
Aliquots (1 ml) of 24-h old bacterial cultures (0.5 MacFarland standard) grown in
nutrient broth were inoculated into 300-ml Erlenmeyer flasks containing 150 ml of MSM
supplemented with various concentrations of glyphosate (25, 50, 100 and 250 mg/ml) to
test their ability to degrade the supplemental substrate (herbicide). A control was
maintained with MSM supplemented with 7.2 mg/ml of glyphosate. Bacterial growth was
monitored by increase in cell number immediately after inoculation at 0 h and at every 12
h interval till 108 h of incubation. Bacterial inoculum (5 ml) was drawn at regular intervals
from the test and control cultures and optical density taken at 660nm.
2.4.6 Effects of Nitrogen Supplements on Growth of the Isolate and Utilization of
Glyphosate
The effects of three organic nitrogen sources (yeast extract, peptone and tryptone)
and two inorganic nitrogen sources (NH4Cl and (NH4)2SO4) on the growth of the isolates
in glyphosate were studied. The screening medium was prepared as earlier described
without the supplements and dispensed in 150ml portions into 500 ml Erlenmeyer flasks.
Peptone, tryptone, yeast extract and NH4Cl each at o.2% (w/v) were added to different
flasks containing the medium in triplicates. The medium with (NH4)2SO4 served as the
control. After autoclaving at 1210C for 15mins, 3 ml of roundup (containing 7.2 mg/ml of
glyphosate) was added as before into each flask, including the controls. Each set of
triplicate flasks (15) was then inoculated with aliquots (1 ml) of 24-h old bacterial cultures
(0.5 MacFarland standard) grown in nutrient broth and incubated as earlier described.
Thereafter, 5 ml samples were aseptically collected from each flask and assayed for
60
growth (optical density at 660 nm) at zero (0) hour and subsequently at twelve hour
intervals.
2.4.7 Effects of Medium pH on Growth of the Isolate and Utilization of Glyphosate
The screening medium was prepared as earlier described and dispensed in 150 ml
portions into 500 ml Erlenmeyer flasks. The pH of each flask was adjusted to the
following levels 4.0, 5.0, 6.0, 7.0, 8.0, 9.0 with 1 M HCl or 0.1 M NaOH, using a pH
meter. Sterile glyphosate (7.2 mg/ml) was added to each replicate flask and also freshly
prepared inoculum of the isolates was used in inoculating the flasks and growth monitored
for 120 hours. Inoculated flasks were incubated on a shaker as described earlier. Samples
(5 ml) were aseptically withdrawn at 12 h intervals and used to assay for growth by
measuring the optical density at 660 nm.
2.4.8 Effect of heavy metals on glyphosate degradation
2.4.8.1 Comparative effects of heavy metals on Pseudomonas fluorescens and
Acetobacter sp growth
Salts of the selected heavy metals ions Pb2+, Cd2+, Zn2+ and Cr2+ at a concentration
of 25-µg/ml were added to 150-ml of the mineral salts medium. The medium was
autoclaved prior to the addition of 1-ml of the filter-sterilized Roundup® (7.2-mg/ml of
glyphosate). One millilitre aliquot of each isolate was used in inoculating the media and
incubation was carried out as described above.
2.4.8.2 Effect of different concentrations of heavy metals
The medium for the study was prepared as described earlier, and was autoclaved
prior to the addition of the filter-sterilized Roundup®. The medium was supplemented
61
with the following heavy metal ions Pb2+, Cd2+, Zn2+ and Cr2+ at concentrations of 50, 100
and 500 µg/ml of their salts. Filter-sterilized glyphosate (7.2 mg/ml) was added to each
replicate flask and also freshly prepared inoculum of the isolates was used in inoculating
the flasks and growth monitored for 120 hours. Medium without heavy metals was used as
control. Inoculated flasks were incubated at 300C on a rotary shaker (Gallenkamp,
England) at 120 rpm. Samples (5 ml) were aseptically withdrawn at 12-hourly intervals
and used to assay for growth by measuring the optical density with a spectrophotometer
(Spectronic 20, USA) at 660 nm.
2.5 Colorimetric Determination of Glyphosate
2.5.1 Preparation of Molybdenum (V)-Molybdenum (VI) reagent
The Molybdenum (V)-Molybdenum (VI) reagent was prepared by dissolving 17.5
g of (NH4)6Mo7O24.4H2O in 200ml of 6N HCl this gave Mo(VI). Zinc metal (1.5 g) was
added to the solution and allowed to dissolve completely. This reduced part of the Mo(VI)
to Mo(V) and a change in colour from light yellow to brown was observed. To the
resultant solution, 100 ml of concentrated HCl was added slowly after which 200 ml of
concentrated H2SO4 was equally added slowly in an ice bath. The solution was then
diluted to a ratio of 1:1 with deionized water to give a green Mo(V)-Mo(VI) reagent.
2.5.2 Preparation of standard curve
A standard curve was prepared by adding 2 ml aliquots of Mo(V)-Mo(VI) reagent
to 0.5-5.0 ml aliquots of 50 µg/ml potassium dibasic phosphate solution in 25-mL
volumetric flasks. The standards were then diluted to 22 ml with distilled water and
allowed to stand in a 1000C water bath for 20 min. The absorbance of the
phosphomolybdate heteropoly blue complex was measured against water using a
62
spectrophotometer at an absorbance of 830nm. A straight line was obtained when quantity
of orthophosphate phosphorus was plotted against absorbance.
2.5.3 Determination of residual glyphosate
This method was carried out as described by Glass (1981). This procedure was
used in the determination of the residual glyphosate. Five millitre of culture medium was
withdrawn every 12 or 24 hours and centrifuged at 10,000 rpm for 10 mins after which 2
ml of the supernatant was used. To the 2 ml of supernatant, 1 ml of 30% H2O2 was added
and the resultant solution was boiled at moderate rate to dryness. It was then allowed to
cool and 20 ml of 0.25 M HCl was added to it to help re-dissolve the residue. Two
millilitre of the Mo(V)-Mo(VI) reagent described above was added to 2 ml of the re-
dissolved residue above and then diluted to 22 ml with distilled water. This was then
allowed to stand in a water bath for 20 mins at 1000C in order for the heteropoly blue
complex to develop. Absorbance was read at 830 nm using water as a blank. The quantity
of glyphosate in each sample was calculated by multiplying the number of micrograms of
orthophosphate phosphorus measured in solution times the factor 5.46. This factor was
derived on the assumption that 100% of the organic phosphorus in glyphosate was
converted to the orthophosphate.
2.6 STATISTICAL ANALYSIS
Graphs and tables were used for data presentation. Treatment effects on the growth of the
bacterial isolates at the different time periods were tested by using two way ANOVA with
replications (P < 0.05). The statistical package used was GENSTAT.
63
CHAPTER THREE
RESULTS
3.1 SOIL MOISTURE CONTENT
Table 4 shows the result of the moisture content of the rice field soil samples The
soil sample designated OMS had the highest moisture content of 18.20% , with AKL
having the least (12.88%).
3.2 ISOLATION OF GLYPHOSATE DEGRADING BACTERIA FROM SOIL
The preliminary studies with glyphosate as carbon or phosphorus source showed
that a total of twelve bacterial isolates were able to grow in the presence of glyphosate as
sole phosphorus source, while seven were able to grow in the medium containing
glyphosate as carbon source (Table 5).
On further sub-culturing, seven isolates (Acetobacter spp, Escherichia spp,
Pseudomonas fluorescens, Azotobacter sp, Alcaligenes sp, Pseudomonas cepacia, and
Pseudomonas spp) consistently grew on the MSM enriched with glyphosate as carbon or
phosphorus source. Those isolated from mineral salt medium containing glyphosate as
sole phosphorus source were:
i. Acetobacter sp - G ADA3
ii. Escherichia sp - G AKL2
iii. Pseudomonas fluorescens - G AKL5
iv. Azotobacter sp - G OMR1
v. Alcaligenes sp - G OMS1
Those isolated from mineral salt medium containing glyphosate as sole carbon source and
inorganic phosphate as phosphorus source (GPi) were tentatively identified as;
i. Pseudomonas cepacia - GPi AKL1; ii. Pseudomonas sp - GPi OMS2
64
Table 4: Moisture content of the different soil samples
Soil Sample Soil Moisture Content (%)
ADA 16.26
AKL 12.88
OMR 14.68
OMS 18.20
65
3.3 GLYPHOSATE UTILIZATION PATTERN BY THE DIFFERENT
ISOLATES
The seven bacterial isolates were screened for glyphosate utilization by measuring
their growth at 660 nm. Of the five (5) bacterial isolates grown on the medium containing
glyphosate as sole phosphorus source, Pseudomonas fluorescens most significantly (P <
0.05) utilized glyphosate (mean OD 0.1268). This was followed by Acetobacter sp,
Azotobacter sp and Alcaligenes sp (mean OD 0.1069, 0.0858 and 0.0841), respectively.
Escherichia sp did not show any appreciable growth as seen in Figure 11. The growth of
Pseudomonas sp and Pseudomonas cepacia were non-significant (P > 0.05) with mean
values of 0.0034 and 0.00438, respectively.
3.4 Growth kinetics of Pseudomonas fluorescens and Acetobacter sp in glyphosate
The growth kinetics of Pseudomonas fluorescens and Acetobacter sp were further
monitored over time at 660 nm, using the MSM enriched with glyphosate as sole
phosphorus source. Their growth was both significant (P < 0.001), But that of
Pseudomonas fluorescens was significantly (P < 0.05) higher than that of Acetobacter sp
as shown in figure 12
66
Fig 11: Screening of the isolates for glyphosate utilization
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0 12 24 36 48 60 72 84 96 108 120 132 144 156 168 180Time (Hours)
Opt
ical
Den
sity
(66
0nm
)
Acetobacter sp Escherichia sp Pseudomonas fluorescens Azotobacter sp
Alcaligenes sp Pseudomonas sp Pseudomonas sp
67
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0 12 24 36 48 60 72 84 96 108 120 132 144 156 168 180 192
Time (Hours)
Op
tica
l D
ensi
ty (
660
nm
)
Acetobacter sp Pseudomonas fluorescens
Figure 12: Growth kinetics of Acetobacter sp and Pseudomonas fluorescens on
glyphosate.
68
3.5 EFFECT OF GLYPHOSATE AS CARBON AND/OR PHOSPHORUS
SOURCE ON THE GROWTH OF THE ISOLATES
3.5.0 Comparative role of glyphosate as carbon or phosphorus source
The ability of glyphosate to serve as carbon source, phosphorus source, carbon and
phosphorus source was monitored. Optical density measurement at 660 nm was used to
monitor increase in cell numbers. The growth of Acetobacter sp was non-significantly (P <
0.05) higher in the Glucose and Glyphosate (Gly and Glu) medium (mean OD value =
0.0907) when compared to Pseudomonas fluorescens (mean OD value = 0.09003) (Figure
13). On the medium with glyphosate as both carbon and phosphorus source, the growth of
Pseudomonas fluorescens was significantly higher when compared to Acetobacter sp as
shown in Figure 13.
3.5.1 Growth kinetics of Acetobacter sp and Pseudomonas fluorescens on glyphosate
as carbon or phosphorus source
The growth kinetics of the isolates in the different carbon sources showed that their
was progressive increase in growth of the isolates when glyphosate was used as a
phosphorus source and glucose as carbon source. The growth of Acetobacter sp after 24 h
incubation on Glu and Gly medium was more significant (P < 0.05) with a mean OD value
of 0.0933 when compared with the growth on the GPi, glyphosate and the control (Figure
14). Also, the growth of Acetobacter sp on the GPi medium peaked after the 36 h with
mean OD value of 0.02733, which was significantly (P<0.05) higher than growth on the
medium containing glyphosate and the control. After the 36 h of incubation, the growth on
the Glu and Gly medium consistently increased till the end of the monitoring (120 h). The
69
growth of Pseudomonas fluorescens in the Glu and Gly medium followed a similar pattern
as that of Acetobacter sp. as shown in Figure 15
70
Fig 13: Comparative effect of glyphosate as carbon and/or phosphorus source
on growth of Acetobacter sp and Pseudomonas fluorescens
0.00000
0.02000
0.04000
0.06000
0.08000
0.10000
0.12000
0.14000
Glucose Gly and Glu Glyphosate Gly and Pi
Carbon source
Op
tica
l D
ensi
ty (
660n
m)
Acetobacter sp Pseudomonas fluorescens
71
Fig 14: Growth kinetics of Acetobacter sp in glyphosate as carbon or phosphorus
source
-0.02000
0.00000
0.02000
0.04000
0.06000
0.08000
0.10000
0.12000
0.14000
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Abs
orba
nce
(660
nm)
Glucose Glyphosate Gly and Glu Gly and Pi
72
Fig 15: Growth kinetics of Pseudomonas fluorescens in glyphosate as carbon or
phosphorus source
0.00000
0.020000.04000
0.060000.08000
0.10000
0.120000.14000
0.16000
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Ab
sorb
ance
(66
0nm
)
Glucose Glyphosate Gly and Glu Gpi
73
3.6 EFFECT OF NITROGEN SUPPLEMENTATION ON MICROBIAL
UTILIZATION OF GLYPHOSATE
3.6.0 Comparative effect of nitrogen supplementation on the growth of Acetobacter
sp and Pseudomonas fluorescens in the presence of glyphosate
Addition of the different organic and inorganic sources of nitrogen (0.2%)
(Peptone, Tryptone, Yeast extract, NH4Cl and (NH4)2SO4) to the mineral salt medium
yielded a progressive increase in the growth of the isolates. The medium containing
peptone gave the most significant (P < 0.05) growth. This was followed closely by that on
yeast extract and peptone. The growth of the isolate on the inorganic nitrogen sources
yielded the least growth. However, its growth in the medium containing NH4Cl was
significant (P < 0.05) when compared with the control-(NH4)2SO4). The growth of
Pseudomonas fluorescens in the medium containing peptone as organic nitrogen source
was significantly (P < 0.05) higher when compared with its growth in the other media
containing other organic nitrogen sources. The growth of the Pseudomonas fluorescens in
the control was significantly (P < 0.05) higher, when compared with its growth in the
medium containing NH4Cl as inorganic nitrogen source (Fig. 16).
3.6.1 Growth kinetics of Acetobacter sp on the nitrogen supplements
After 24 h of incubation, a progressive increase in growth of the isolate was
observed in all the media with peptone medium giving the most significant (p < 0.05)
growth (mean OD = 0.2150). This consistent increase in growth (Figure 16) continued in
all the media until after 156th h when a peak in growth in the medium containing yeast
extract as nitrogen source was noted. This was the highest growth observed with a mean
74
value of 0.440 and it was significant (P < 0.05), when compared to the growth in the other
media (Fig 17).
3.6.2 Growth kinetics of Pseudomonas fluorescens on the nitrogen supplements
The growth of Pseudomonas fluorescens in the different nitrogen sources increased
progressively with the medium containing peptone having the most significant (P < 0.05)
growth (Mean OD = 0.240) after 24 h. After 132 h, the growth in the medium containing
yeast extract increased above that in the peptone medium. The highest increase in the
growth of Pseudomonas fluorescens in the mineral salt medium containing yeast extract
was observed after 168 h (mean OD = 0.4100) as shown in Figure 18.
75
0
0.05
0.1
0.15
0.2
0.25
0.3
0.35
Tryptone Yeast extract Peptone NH Cl (NH ) SO
Nitrogen source
Ab
sorb
ance
(66
0nm
)
Acetobacter spp Pseudomonas fluorescens
Fig 16: Comparative effect of nitrogen supplementation on the growth Acetobacter sp
and Pseudomonas fluorescens in the presence of glyphosate
76
0.0000.0500.1000.1500.2000.2500.3000.3500.4000.4500.500
0 12 24 36 48 60 72 84 96 108 120 132 144 156 168
Time (Hours)
Op
tica
l D
ensi
ty (
660n
m)
Tryptone Yeast Extract Peptone NH Cl (NH ) SO4
Fig 17: Growth kinetics of Acetobacter sp on the nitrogen supplements in the
presence of glyphosate
77
0.000
0.100
0.200
0.300
0.400
0.500
0.600
0 12 24 36 48 60 72 84 96 108 120 132 144 156 168
Time (Hours)
Op
tica
l D
ensi
ty (
660n
m)
Tryptone Yeast Extract Peptone NH Cl (NH ) SO4
Fig. 18: Growth kinetics of Pseudomonas fluorescens on the nitrogen supplements in
the presence of glyphosate
78
3.7 EFFECTS OF DIFFERENT GLYPHOSATE CONCENTRATIONS ON THE
GROWTH OF ACETOBACTER SP AND PSEUDOMONAS FLUORESCENS
The growth of Acetobacter sp and Pseudomonas fluorescens in different
concentrations of glyphosate gave an inverse result as shown in Figure 19. As the
concentration of glyphosate increased there was a corresponding decrease in the growth of
the isolates. The highest growth was observed in the control (7.2 mg/ml) which contained
the least concentration of glyphosate. The growth kinetics of Acetobacter sp and
Pseudomonas fluorescens in different concentrations of glyphosate, gave a progressive
decrease as is seen in Figure 20 and 21. A 12 h lag phase was observed at concentrations
of 7.2 mg/ml, 25 mg/ml and 50 mg/ml, 100mg/ml in both isolates.
79
0.0000
0.0200
0.0400
0.0600
0.0800
0.1000
0.1200
0.1400
0.1600
Op
tica
l d
ensi
ty (
660
nm
)
7.2mg/ml 25mg/ml 50mg/ml 100mg/ml 250mg/ml
Acetobacter sp Pseudomonas fluorescens
Fig 19: Effects of the different concentrations of glyphosate on the growth of
Acetobacter sp and Pseudomonas fluorescens
80
Fig 20: Growth kinetics of Acetobacter sp on the different concentrations of
glyphosate
0.000
0.050
0.100
0.150
0.200
0.250
0 12 24 36 48 60 72 84 96 108
Time (hours)
Ab
sorb
ance
(66
0nm
)
7.2mg/ml 25mg/ml 50mg/ml 100mg/ml 250mg/ml
C
81
Fig 21: Growth kinetics Pseudomonas fluorescens on the different concentrations of
glyphosate
0.0000
0.0500
0.1000
0.1500
0.2000
0 12 24 36 48 60 72 84 96 108Time (Hours)
Ab
sorb
ance
(66
0nm
)
7.2mg/ml 25mg/ml 50mg/ml 100mg/ml 250mg/ml
82
3.8 EFFECT OF HEAVY METALS ON MICROBIAL UTILIZATION OF
GLYPHOSATE
3.8.0 Comparative effects of heavy metals on Acetobacter sp and Pseudomonas
fluorescens
Figure 22 depicts the effect of the different heavy metals on the growth of
Acetobacter sp and Pseudomonas fluorescens. For Acetobacter sp maximum growth was
observed in the medium containing lead (mean OD = 0.1434). This was highly significant
(P < 0.05) when compared to the growth of Acetobacter sp in the presence of the other
heavy metals. The growth of Acetobacter sp in the medium containing zinc was also
significantly (P <0.05) higher when also compared with its growth in cadmium (0.1143)
and chromium (0.0783), which gave the least growth. The growth of Pseudomonas
fluorescens in the medium containing zinc gave the most significant (P < 0.05) growth
(mean OD = 0.1338). This was closely followed by lead, cadmium and chromium (mean
OD = 0.1184, 0.1004 and 0.09927), respectively.
3.8.1 Effects of different concentrations of cadmium on the growth of Acetobacter
sp
As shown in Figure 23, the growth of Acetobacter sp in the medium containing
500 µg/ml of cadmium was significantly (P < 0.05) higher when compared with its growth
at the other concentrations used. A 12 h lag period was observed at concentrations of 50
and 100 µg/ml before any appreciable growth was observed after the 36 h incubation. The
highest (peak) growth was seen in the medium containing 100 µg/ml after 120 h. This
increase was significant (P < 0.05) when compared with others after 120 h incubation.
83
3.8.2 Effects of different concentration of chromium on the growth of Acetobacter
sp
The growth of the isolate at the different concentration of chromium followed a
similar pattern as shown in Figure 24. A 12 h time lag was observed in the medium
containing 50 and 100 µg/ml of chromium. Maximum (peak) growth was observed in the
medium containing 50 µg/ml of chromium after 96 h of incubation (mean OD = 0.170)
and it was significant when compared to others
3.8.3 Effects of different concentrations of lead on the growth of Acetobacter sp
The growth of Acetobacter sp increased progressively after a 12 h lag period at the
different concentrations (Figure 25). The growth at a concentration of 500 µg/ml of lead
gave the most significant (P < 0.05) yield, with maximum growth observed after the 72 h
(mean OD = 0.2900)
3.8.4 Effects of different concentrations of zinc on the growth of Acetobacter sp
The addition of zinc at a concentration of 500 µg/ml gave the most significant (P <
0.05) yield (Figure 26). Maximum growth was observed after 96 h with a mean OD of
0.3250. After 12 h of incubation, a lag period was observed with the three different
concentrations before any appreciable increase in growth
3.8.5 Effects of different concentrations of cadmium on the growth of Pseudomonas
fluorescens
Appreciable growth of Pseudomonas fluorescens in the medium containing
cadmium (at different concentrations) was only observed after 24 h (50 µg/ml) and 36 h
(100 and 500 µg/ml) of incubation. Figure 27 shows a progressive increase in the growth
84
of Pseudomonas fluorescens at the different concentrations of cadmium. Its growth in the
medium containing 500 µg/ml concentration was significantly higher (P < 0.05) when
compared with others. Peak growth at this concentration was seen after 96 h incubation
(mean OD = 0.2350).
3.8.6 Effects of different concentrations of chromium on the growth of
Pseudomonas fluorescens
In Figure 28, a consistent increase in growth of Pseudomonas fluorescens was
observed in the media containing 50 and 100 µg/ml concentrations of chromium all
through 120 h of monitoring. Maximum growth was recorded in the medium containing
100 µg/ml of chromium after 96 h incubation (mean OD = 0.180) and was significant (P <
0.05) when compared with other concentrations at that time. The growth of the isolate in
the medium containing 500 µg/ml had its peak growth after 36 h (mean OD of 0.130) and
thereafter, there was a steady decline.
3.8.7 Effects of different concentrations of Lead on the growth of Pseudomonas
fluorescens
The growth pattern of Pseudomonas fluorescens in the medium containing
lead at different concentrations was similar (Figure 29). However, after 72 h incubation, a
significant (P < 0.05) increase in growth was observed in the medium containing 500
µg/ml ((mean OD = 0.210). Also, after 108 h incubation another growth peak was
observed in the medium containing 50 µg/ml (mean OD = 0.2130).
85
3.8.8 Effects of different concentrations of zinc on the growth of Pseudomonas
fluorescens
The addition of 500 µg/ml of zinc to the glyphosate mineral salt medium yielded
the most significant (P < 0.05) growth when compared with its growth at 50 and 100
µg/ml concentrations. A growth peak was recorded after 96 h incubation (mean OD =
0.335) in this medium (Figure 30).
86
0.00000.02000.04000.06000.08000.10000.12000.14000.1600
Opt
ical
Den
sity
(660
nm
)
Lead Cadmium Zinc Chromium
Heavy Metals
Acetobacter sp Pseudomonas fluorescens
Fig 22: Comparative effects of heavy metals on Acetobacter sp and Pseudomonas
fluorescens
87
Fig 23: Effects of different concentrations of cadmium on the growth of
Acetobacter sp
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l D
ensi
ty (
660n
m)
50µg/ml 100µg/ml 500µg/ml Metal Free
88
Fig 24: Effects of different concentrations of chromium on the growth of
Acetobacter sp
0.000
0.050
0.100
0.150
0.200
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l D
ensi
ty (
660n
m)
50µg/ml 100µg/ml 500µg/ml Metal Free
89
Fig 25: Effects of different concentrations of lead on the growth of Acetobacter sp
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0.350
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l d
ensi
ty (
660n
m)
50µg/ml 100µg/ml 500µg/ml Metal Free
90
Fig 26: Effects of different concentrations of zinc on the growth of Acetobacter sp
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0.350
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l D
ensi
ty (
660n
m)
50µg/ml 100µg/ml 500µg/ml Metal Free
91
Fig 27: Effects of different concentrations of Cadmium on the growth of
Pseudomonas fluorescens
0.0000
0.0500
0.1000
0.1500
0.2000
0.2500
0.3000
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l d
ensi
ty (
660n
m)
50 µg/ml 100 µg/ml 500 µg/ml Metal free
92
Fig 28: Effects of different concentrations of chromium on the growth of Pseudomonas
fluorescens
0.0000
0.0500
0.1000
0.1500
0.2000
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l d
ensi
ty (
660n
m)
50 µg/ml 100 µg/ml 500 µg/ml Metal free
93
Fig 29: Effect of different concentrations of lead on the growth of Pseudomonas
fluorescens
0.000
0.050
0.100
0.150
0.200
0.250
0.300
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Ab
sorb
ance
(66
0nm
)
50µg/ml 100µg/ml 500µg/ml Metal free
94
Fig 30: Effects of different concentrations of zinc on the growth of Pseudomonas
fluorescens
0.00000.0500
0.10000.1500
0.20000.2500
0.30000.3500
0.4000
0 12 24 36 48 60 72 84 96 108 120
Time (Hours)
Op
tica
l d
ensi
ty (
660n
m)
50 µg/ml 100 µg/ml 500 µg/ml Metal free
95
3.9 Effect of pH on microbial degradation of glyphosate
3.9.1 Effect of pH on the utilization of glyphosate by Acetobacter sp and P.
fluorescens
The effects of pH on the utilization of glyphosate by Acetobacter sp were
evaluated by adjusting the initial pH of the medium to different pH values (4, 5, 6, 7, 8 and
9). The growth of Acetobacter sp at pH 5, 6, 7, 8 and 9 preceded with a very short lag
phase, while at pH 4 appreciable growth was only observed after a 24-h lag phase (figures
31, 32, 33, 34, 35, 36). The growth of the isolate at pH 8 yielded the most significant (P <
0.05) growth when compared with others. Although this growth was not significant (P <
0.05) when compared to its growth at pH 5, the mean OD was higher at pH 8 than at 5.
The resultant pH in the medium starting with pH 4 was 5.0. For medium pH 5, the
resultant pH was between 5.0-5.1, for pH 6 it was 5.0-5.1, pH 7 (5.1-5.3). pH 8 had a
resultant pH of between 5.7-5.9 and then at pH 9 the resultant pH was 6.8-7.7.
A 24-h lag phase was seen in the growth of P. fluorescens at medium pH 4 and 9,
while pH 5, 6, 7 and 8 had a 12-h lag phase. a similar drop in pH as was observed for
Acetobacter sp at medium pH 6, 7, 8 and 9. The resultant pH at medium pH 4 also
increased to 5.0 as the isolate utilized/metabolised glyphosate (Figure 43, 44, 45, 46, 47
and 48).
3.9.2 Effect of pH on the growth of Acetobacter sp and P. fluorescens and
degradation of glyphosate
The effect of pH on the growth of Acetobacter sp and degradation of glyphosate
showed (figures 37, 38, 39, 40, 41 and 42) that as the growth of the isolate increased, the
amount of glyphosate in the medium decreased with time. Medium pH 8 had the highest
amount of glyphosate degraded, while pH 9 had the least amount of glyphosate degraded
96
(Figures 41 and 42). A similar trend was observed for the effect of pH on growth of P.
fluorescens and degradation of glyphosate as shown in figures (49, 50, 51, 52, 53 and 54)
97
1
2
3
4
5
6
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0.0000.0200.0400.0600.0800.1000.1200.140
OD
(66
0nm
)
pH Cell Density
Fig. 31: Effect of pH 4 on the utilization of Glyphosate by Acetobacter sp
98
4.944.96
4.985
5.025.04
5.065.08
5.15.12
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0.0000.0200.0400.0600.0800.1000.1200.1400.1600.1800.200
OD
(66
0nm
)
Resultant pH Cell Density
Fig. 32: Effect of pH 5 on the utilization of Glyphosate by Acetobacter sp
99
4.44.64.8
55.25.45.65.8
66.2
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0.000
0.050
0.100
0.150
0.200
OD
(66
0nm
)
pH Cell Density
Fig. 33: Effect of pH 6 on the utilization of Glyphosate by Acetobacter sp
100
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
pH Cell Density
Fig. 34: Effect of pH 7 on the utilization of Glyphosate by Acetobacter sp
101
0
2
4
6
8
10
0 12 24 36 48 60 72 84 96 108 120Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
pH Cell Density
Fig. 35: Effect of pH 8 on the utilization of Glyphosate by Acetobacter sp
102
0
2
4
6
8
10
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
(pH
)
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
pH Cell Density
Fig. 36: Effect of pH 9 on the utilization of Glyphosate by Acetobacter sp
103
pH4
Time (h)
Res
idu
al g
lyp
ho
sate
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 1200.000
0.050
0.100
0.150
OD
(66
0nm
)
Residual Glyphosate Cell Density
pH4
Time (h)
Res
idu
al g
lyp
ho
sate
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 1200.000
0.050
0.100
0.150
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 37: Effect of pH 4 on growth of Acetobacter sp and degradation of glyphosate
104
pH 5
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
idu
al g
lyp
ho
sate
0.000
0.050
0.100
0.150
0.200
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 38: Effect of pH 5 on growth of Acetobacter sp and degradation of glyphosate
105
Time (h)
Res
idu
al G
lyp
ho
sate pH6
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0.000
0.050
0.100
0.150
0.200
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
al G
lyp
ho
sate pH6
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0.000
0.050
0.100
0.150
0.200
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 39: Effect of pH 6 on growth of Acetobacter sp and degradation of glyphosate
106
Time (h)
Res
idu
al G
lyp
ho
sate pH7
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
al G
lyp
ho
sate pH7
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 40: Effect of pH 7 on growth of Acetobacter sp and degradation of glyphosate
107
Time (h)
Res
idu
al g
lyp
ho
sate
pH8
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
al g
lyp
ho
sate
pH8
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 41: Effect of pH 8 on growth of Acetobacter sp and degradation of glyphosate
108
Time (h)
Res
idu
al G
lyp
ho
sate
pH9
0.00
1.00
2.00
3.00
4.00
5.00
6.00
12 24 36 48 60 72 84 96 108 120
0.000
0.050
0.100
0.150
0.200
0.250
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
al G
lyp
ho
sate
pH9
0.00
1.00
2.00
3.00
4.00
5.00
6.00
12 24 36 48 60 72 84 96 108 120
0.000
0.050
0.100
0.150
0.200
0.250
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 42: Effect of pH 9 on growth of Acetobacter sp and degradation of glyphosate
109
pH4
0.01.02.03.04.0
5.0
6.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
00.050.1
0.15
0.2
0.25
0.3
OD
(66
0nm
)
Resultant pH Cell Density
pH4
0.01.02.03.04.0
5.0
6.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
00.050.1
0.15
0.2
0.25
0.3
OD
(66
0nm
)
Resultant pH Cell Density
Fig. 43: Effect of pH 4 on the utilization of Glyphosate by Pseudomonas fluorescens
110
pH Cell density
Res
ult
ant
pH pH 5
4.5
5.0
5.5
6.0
6.5
0 12 24 36 48 60 72 84 96 108 120
Time (h)
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
pH Cell density
Res
ult
ant
pH pH 5
4.5
5.0
5.5
6.0
6.5
0 12 24 36 48 60 72 84 96 108 120
Time (h)
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)pH 5
4.5
5.0
5.5
6.0
6.5
0 12 24 36 48 60 72 84 96 108 120
Time (h)
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Fig. 44: Effect of pH 5 on the utilization of Glyphosate by Pseudomonas fluorescens
111
pH 6
pH
4.5
5.0
5.5
6.0
6.5
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Cell density
pH 6
pH
4.5
5.0
5.5
6.0
6.5
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Cell density
Fig. 45: Effect of pH 6 on the utilization of Glyphosate by Pseudomonas fluorescens
112
pH7
4.0
5.0
6.0
7.0
8.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
pH Cell Density
pH7
4.0
5.0
6.0
7.0
8.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
OD
(66
0nm
)
pH Cell Density
Fig. 46: Effect of pH 7 on the utilization of Glyphosate by Pseudomonas fluorescens
113
pH8
0.0
2.0
4.0
6.0
8.0
10.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
pH Cell Density
pH8
0.0
2.0
4.0
6.0
8.0
10.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
pH Cell Density
Fig. 47: Effect of pH 8 on the utilization of Glyphosate by Pseudomonas fluorescens
114
pH9
0.0
2.0
4.0
6.0
8.0
10.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
OD
(66
0nm
)
pH Cell Density
pH9
0.0
2.0
4.0
6.0
8.0
10.0
0 12 24 36 48 60 72 84 96 108 120
Time (h)
Res
ult
ant
pH
0
0.05
0.1
0.15
OD
(66
0nm
)
pH Cell Density
Fig. 48: Effect of pH 9 on the utilization of Glyphosate by Pseudomonas fluorescens
115
Time (h)Residual Glyphosate Cell Density
pH4
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
0.25
0.3
Res
idu
al g
lyp
ho
sate
OD
(66
0nm
)
Time (h)Residual Glyphosate Cell Density
pH4
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
0.25
0.3
Res
idu
al g
lyp
ho
sate
OD
(66
0nm
)
Fig. 49: Effect of pH 4 on growth of Pseudomonas fluorescens and degradation of
glyphosate
116
OD
(66
0nm
)
Time (h)
Res
idu
alg
lyp
ho
sate
pH5
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
Residual Glyphosate Cell density
OD
(66
0nm
)
Time (h)
Res
idu
alg
lyp
ho
sate
pH5
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
Residual Glyphosate Cell density
Fig. 50: Effect of pH 5 on growth of Pseudomonas fluorescens and degradation of
glyphosate
117
Time (h)
Res
idu
alG
lyp
ho
sate
pH6
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 1200
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
alG
lyp
ho
sate
Res
idu
alG
lyp
ho
sate
pH6
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 1200
0.05
0.1
0.15
0.2
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 51: Effect of pH 6 on growth of Pseudomonas fluorescens and degradation of
glyphosate
118
Time (h)
Res
idu
al G
lyp
ho
sate pH7
0
2
4
6
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
al G
lyp
ho
sate pH7
0
2
4
6
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 52: Effect of pH 7 on growth of Pseudomonas fluorescens and degradation of
glyphosate
119
Time (h)
Res
idu
al G
lyp
ho
sate pH8
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
Residual Glyphosate Cell Density
Time (h)
Res
idu
al G
lyp
ho
sate pH8
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
0.2
0.25
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 53: Effect of pH 8 on growth of Pseudomonas fluorescens and degradation of
glyphosate
120
Time (h)
Res
idu
al G
lyp
ho
sate pH9
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
OD
(66
0nm
)
Residual Glyphosate Cell DensityTime (h)
Res
idu
al G
lyp
ho
sate pH9
0
2
4
6
8
0 12 24 36 48 60 72 84 96 108 120
0
0.05
0.1
0.15
OD
(66
0nm
)
Residual Glyphosate Cell Density
Fig. 54: Effect of pH 9 on growth of Pseudomonas fluorescens and degradation of
glyphosate
121
CHAPTER FOUR
DISCUSSION
Nineteen bacterial isolates were initially isolated from rice field soil samples. On
further sub-culturing on solid media enriched with glyphosate, only seven showed the
capacity to grow in the presence of the herbicide. The seven bacterial isolates were
identified as Acetobacter sp, Escherichia sp, Pseudomonas fluorescens, Azotobacter sp,
Arthrobacter sp, Pseudomonas cepacia and Alcaligenes sp, respectively. Studies by Busse
et al., (2001) showed that culturable bacteria and fungi are usually reduced in number or
eliminated when extracted from soil or grown on solid media containing glyphosate. The
results in this study which showed a reduction in the number of bacterial species grown on
the glyphosate solid medium are consistent with the report of Busse et al. (2001). Toxicity
of the artificial media is expected based on the mode of action of glyphosate (inability of
the organism to synthesize the needed aromatic amino acids). Other studies have found
similar reductions in population counts when glyphosate is added to culture media (Quinn
et al., 1988; Santos and Flores, 1995; Kryzsko-Lupicka and Orlik, 1997). Unlike the
response in artificial media, no toxicity was expressed when glyphosate was added to soil
in laboratory bioassays (Busse et al., 2001).
The identified bacterial species have been previously isolated from other soil
samples (Zboinska et al., 1992, Franz et al., 1997). Of the seven identified bacterial
species, two (Acetobacter sp. and Pseudomonas fluorescens) were selected for further
utilization studies based on their short lag phase and rapid utilization of glyphosate (Figure
1). Many species of Pseudomonas have been used extensively in the degradation/or
metabolism of glyphosate (Jacob et al., 1988; Shinabarger and Braymer, 1984; Jacob and
Kishore, 1987; Talbot et al., 1984. But Zboinska et al. (1992) reported that Pseudomonas
fluorescens could not utilize glyphosate contrary to our result. The strain of Pseudomonas
122
fluorescens used for this study was not only able to utilize glyphosate but was also able to
thrive at high concentrations of the herbicide. The use of Acetobacter sp in the degradation
or metabolism of glyphosate has not been reported. Both organisms used in this study
showed appreciable growth in the culture media containing glyphosate as sole phosphorus
source. The difference in the growth of the isolates in the medium is indicative of the
differences between the organisms in tolerating the herbicide. The effective utilization of
glyphosate by the selected isolates is indicated by the rapid growth with little lag phases.
In addition, Pseudomonas fluorescens attained maximum growth and peaked after 132 h
incubation, while Acetobacter sp achieved maximum growth and peaked after 72 h
incubation. There have been several reports on the ability of microorganisms including
some Pseudomonas sp to effectively utilize glyphosate by naturally synthesizing
appropriate enzymes or as a result of genetic mutation (Jacob et al., 1988; Shinabarger,
and Braymer, 1984; Jacob and Kishore, 1987). But so far, there has been no report on the
ability of Pseudomonas fluorescens to utilize glyphosate as sole phosphorus or carbon
source. The high capacity of these two organisms to utilize this herbicide in vitro could be
attributed to their previous contact with the herbicide in the soil (rice fields) from where
they were isolated. It is also possible that the organisms have undergone genetic mutation
leading to the adaptability of the organisms to their microenvironment.
The testing of the comparative role of glyphosate as carbon or phosphorus source
for the two isolates showed that glyphosate serves as a better phosphorus source for the
two isolates. Many bacterial isolates have been reported to utilize glyphosate as a
phosphorus source (Liu et al., 1991; Balthazor and Hallas, 1986; Dick and Quinn, 1995).
Of the two isolates, Pseudomonas fluorescens demonstrated a better capacity to utilize
glyphosate as carbon and phosphorus source. Glyphosate served as a better phosphorus
source for the Acetobacter sp than as a carbon source. In both organisms, the presence of
123
inorganic phosphate in the Gpi medium affected the effective uptake of glyphosate. This is
because inorganic phosphate (Pi) has been reported to suppress the genes coding for the C-
P layse system and thus make them unable to metabolise glyphosate (Liu et al., 1991). Our
results in this study agree with this report
The results in this study showed that supplementing the MSM-glyphosate with
organic nitrogen sources increased the growth of the isolates when compared to the
control (NH4)2SO4 and NH4Cl which are the inorganic nitrogen sources tested. This could
be explained by the fact that organic nitrogen sources are ready-made, and therefore
considerably easier for the isolates to utilize than their inorganic counterparts in the culture
medium, which must be processed in some way before it could be utilized by the
organisms (Acetobacter sp and Pseudomonas fluorescens). This may explain why the
organisms utilized the organic nitrogen sources in preference to the inorganic control
((NH4)2SO4), which already was a part of the mineral salts medium formulated for its
isolation. Since nitrogen is usually not a part of the precursor metabolites or even
intermediate building blocks in most bacterial biosynthetic pathways, there is the need to
provide it externally, either through inorganic or organic sources. Thus nitrogen is
considered a limiting nutrient in most bacterial growth experiments (Prescott et al., 2002;
Annuar et al., 2008; Nnamchi, 2004). This no doubt explains why the addition of peptone,
yeast extract, and tryptone to glyphosate containing mineral salts medium yielded more
bacterial growth than the control and NH4Cl. Organic nitrogen sources that contain
aromatic amino acids help to support protein synthesis and overcome the inhibition of the
shikimic acid pathway in bacteria grown in the presence of glyphosate (Busse et al.,
2001). The result showed that peptone serves as a better organic nitrogen source followed
by yeast extract and tryptone. The growth of Pseudomonas fluorescens in glyphosate
medium containing yeast extract increased more than that in the peptone medium, after
124
132 h and maximum growth was observed after 168 h. This same pattern was observed for
Acetobacter sp but in this case, the peak growth was observed after 156 h. This suggests
that the nitrogen in the yeast extract is slowly released for use by the isolates and would be
better for use in biodegradation of glyphosate.
Carlisle and Trevors (1988) observed that glyphosate can either stimulate or inhibit
soil microorganisms depending on the soil type or herbicide concentration. The results in
this study showed that increase in glyphosate concentration led to a concomitant decrease
in the growth of the isolates. Amoros et al., (2007), while studying the effects of roundup
at different glyphosate concentrations (50 and 100 mg/l) observed an increase in
Aeromonas counts at these concentrations in contrast to the control which contained no
glyphosate at all. In our study, after 24 h at 300C, high cell density of both isolates was
recorded at glyphosate concentrations of 7.2 - 50 mg/ml. However, at higher
concentrations of glyphosate (100 and 250 mg/ml), the cell density was very low
compared with the control. Even though a severe decline in growth of the organisms
occurred at high concentrations (100 and 250 mg/ml), they were still able to tolerate up to
250 mg/ml of glyphosate. A possible explanation may be the presence of novel
degradative systems in the organisms. The growth kinetics of both isolates in increasing
concentrations of glyphosate followed a similar pattern with a lag phase of about 12 h and
steady increase in growth. After 84 h of incubation the growth of Pseudomonas
fluorescens in medium containing 25 mg/ml of glyphosate increased significantly (P <
0.05) when compared with its growth in the medium with 7.2 mg/ml of glyphosate till the
end of the monitoring at 108 h.
Gram negative bacteria showed higher tolerance to heavy metals than their gram
positive counterparts due to their higher level of intrinsic metal resistance (Ahmad et al.,
2005). This difference is based on the chemical composition of their cell wall. The two
125
bacterial species used are gram negative organisms thus supporting their ability to tolerate
high concentrations of the heavy metals. Noghabi et al. (2007) reported the high capability
of heavy metals bioaccumulation by Gram negative bacteria. Many bacterial-resistance
systems for toxic metals are plasmid- encoded (Silver, 1996; Jankowska et al., 2006),
however, some are chromosomal (Gupta et al., 1999).
Heavy metals have been reported to stimulate microbial growth (Gikas, 2007;
Gikas, et al., 2009). Kools et al., (2005) reported that the presence of heavy metals
increased the rate of glyphosate utilization. They proposed that this could be because
metal-glyphosate complexes are transported more efficiently across microbial cell walls
than the sole compound. This supports the findings of this study. We observed that the
addition of lead (Pb), cadmium (Cd), and zinc (Zn) to the glyphosate medium used in
growing the Acetobacter sp increased the rate of glyphosate utilization as indicated by the
increase in the growth of the organisms. None of the metals used completely inhibited the
growth of the organism. The organisms grew very well in the presence of lead at all the
concentrations (50, 100, 500 µg/ml) used. This was closely followed by zinc, cadmium
and chromium. The highest growth of the organisms was observed at the highest
concentration of the heavy metal (500 µg/ml) with the exception of chromium. Although
chromium serves as an essential trace metal, overexposure or very high concentrations of
it has cytotoxic and genotoxic effects (cell death, cell transformation and mutation)
(Carmago et al., 2005).
The growth of Pseudomonas fluorescens was more enhanced in the presence of
zinc (zinc is an essential element for the normal activity of DNA polymerase and protein
synthesis) in the glyphosate medium. This was followed by lead, cadmium and chromium
respectively. Unlike the response of Acetobacter sp to 500 µg/ml of chromium,
Pseudomonas fluorescens showed more tolerance to chromium at that concentration.
126
Studies by Bopp et al. (1983) and Viti et al. (2006) showed that Pseudomonas fluorescens
has the capacity to resist chromate and grow. This report is consistent with our findings.
In this work, we observed that 500ug/ml of chromium in the glyphosate medium
caused a reduction in the growth of Pseudomonas fluorescens. Chromate resistance in
Pseudomonas sp has been shown to be plasmid-associated and transmissible to a restricted
host range (Nriagu and Nieboer, 2007; El.Deeb and Altahli, 2009). Chromium at a
concentration of 500ug/ml caused a significant reduction in the growth of Acetobacter sp.
with an eventual inhibition of the isolate in the medium. Two peaks were observed for the
growth of the organism on chromium Viti et al. (2006) suggest a possible explanation for
this; the sudden drop could be due to toxic shock that chromium had on the isolate causing
cell lysis. Thereafter, leaking nutrients from the lysed cells aided the growth of the more
resistant cells. The overall reduction in the growth of the isolates by chromate might be
explained by the higher requirements for maintenance energy in the presence of the metal
(Giller et al., 1998) or due to cell lysing as an effect of the exposure to heavy metal, as
growth was measured by the optical density. In the first case, the bacteria expended energy
to repair the cell damages caused by the metal toxicity, or because they had to use
alternative enzymatic pathways to adapt to the new environmental conditions (Gikas et al.,
2009).
The effect of increasing concentrations (50, 100, 500 µg/ml) of Zn was examined.
It can be concluded that low concentrations of Zn (50 and 100 µg/ml) did not exhibit any
inhibitory effect on the isolates. However, at low concentration of this metal, the growth
of the isolates was similar as in the control (glyphosate alone). A ten-fold increase in the
concentration of the Zn caused a significant increase in the growth of the isolates. A
probable explanation for this is that Zn is an essential trace element that is important in
forming complexes (such as zinc fingers in DNA) and as a component in cellular enzymes
127
(Spain and Alm, 2003). Zinc is usually accumulated by an unspecific uptake mechanism
that is generally coupled to magnesium.
The drop in pH observed in medium pH 6, 7, 8 and 9 could be attributed to the
nature of glyphosate (it is a weak acid). Glyphosate was added to the medium after the
medium pH has been adjusted and the medium sterilized. The growth of both isolates in
the medium had no significant (P < 0.05) effect on the resultant pH as the resultant pH was
maintained at a certain range after the initial drop in pH.
The amount of glyphosate degraded increased as the isolates increased in growth.
This points to the fact that more of the glyphosate was broken down so as to release the
requisite amount of phosphorus required by the isolates for growth. Glyphosate exists at
the monoanionic state between pH 4.9-6.8 (Franz et al., 1997). Tsui et al. (2005) observed
that at this ionic state, glyphosate is easily degraded because there are more sites on the
herbicide that is open to attack by soil microbes. The result of this work showed that the
optimum pH for the degradation of glyphosate by our test organisms (Acetobacter sp and
P. fluorescens) was 5.8. Although, the pH range for effective degradation of glyphosate by
both isolates was between 5.0-6.8.
Conclusion
In Nigeria, most of the rural settlers are farmers. Adani is a typical example where
over 90% of the adult population engages in rice farming. From preliminary studies,
herbicides are extensively used for the farming process and the farmers due to illiteracy
and impatience, use more than the stipulated quantity of the pesticide per application. This
pesticide persists for long periods of time in the environment, thereby affecting non-target
organisms. This study reports the isolation and identification of two bacterial species,
Pseudomonas fluorescens and Acetobacter sp. that possess a high capacity to utilize
128
glyphosate. To our knowledge, there has not been any report showing the utilization of
glyphosate by the two organisms we used. The abilities of these isolates to utilize
glyphosate effectively are a sure means of removing this compound from the environment.
This study also revealed that their capacity for survival and growth in the presence of high
concentration of this herbicide marks them out as good candidates for the bioremediation
of glyphosate polluted ecosystems.
129
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APPENDIX I
A. Nutrient Agar (Fluka)
Formula (g/l);
Peptone - 5.0
Beef Extract - 3.0
Sodium Chloride - 8.0
Agar - 12.0
Preparation:
Twenty eight (28) grams of powder was suspended in one litre of deionized water
(dH20). This was brought to boil to dissolved completely and subsequently sterilized by
autoclaving at 1210C for 15 min. after cooling to about 470C the solution was distributed,
about 20 ml each into pre-sterilized Petri dish.
B. Nutrient Broth (Fluka)
Formula (g/l);
Beef Extract - 1.0
Yeast Extract - 2.0
Peptone - 5.0
Sodium Chloride - 5.0
Preparation:
Thirteen (13) grams of powder was dissolved in 1 L of dH20, allowed to dissolve
(10 minutes, swirled to mix, dispensed into final containers and sterilized by autoclaved at
1210C for 15 minutes.
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C. Christensen’s Urea Agar (Oxoid)
Formula (of Agar base) (g/l);
Peptone - 1.0
Glucose - 1.0
Sodium Chloride - 5.0
Di-sodium phosphate - 1.2
Potassium dihydrogen phosphate - 0.8
Phenol red - 0.012
Agar - 15.0
Preparation:
A 2.4 g quantity of the agar base powder was dispersed in 95 ml of dH20 according to the
manufacturer’s specification. This was brought to boil to dissolve completely and
subsequently sterilized by autoclaving at 1150C for 20 minutes. It was cooled to about
500C, 5 ml of sterile 40% urea solution (SR 20) was added aseptically and both were
mixed well by gentle shaking and swirling. The mixture was distributed, (10 ml aliquots)
into sterile Bijou bottles and allowed to set in a slope position.
D. Sulphide Indole Motility Agar (Oxoid)
Formula (g/l);
Tryptone - 20.0
Peptone - 6.1
Ferrous ammonium sulphate - 0.2
Sodium thiosulphate - 0.2
Agar - 3.5
144
Preparation:
Preparation was done as in 1A above using (w/v) concentrations equivalent to 30 g in 1 L
of dH20.
E. Kligler Iron Agar (LAB M)
Formula (g/l);
Dextrose - 1.0
Sodium Chloride - 5.0
Ferric ammonium citrate - 0.5
Sodium Thiosulphate - 0.3
Phenol red - 0.025
Agar No. 2 - 12.0
Preparation:
Forty nine (49) grams of powder was add to 1 litre of dH20 in a two litre flask. This was
brought to boil over a gauze with frequent swirling to prevent burning. It was allowed to
simmer for 3 seconds to dissolve and allowed to cool to 470C before pouring into test
tubes in a 3 cm slant position.
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Appendix II
Preparation of McFarland Nephelometer Standards:
Turbidity standard number
Barium chloride dehydrate (1.175%)
Sulphuric acid (1%)
Corresponding approximate density of bacteria
0.5 0.5 ml 99.5 ml 1 x 108
1 0.1 ml 9.9 ml 3 x 108
2 0.2 ml 9.8 ml 6 x 108
3 0.3 ml 9.7 ml 9 x 108
4 0.4 ml 9.6 ml 12 x 108
5 0.5 ml 9.5 ml 15 x 108
6 0.6 ml 9.4 ml 18 x 108
7 0.7 ml 9.3 ml 21 x 108
8 0.8 ml 9.2 ml 24 x 108
9 0.9 ml 9.1 ml 27 x 108
10 9.0 9.0 ml 30 x 108
(WHO, 2003)
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Appendix III
ANOVA Table for glyphosate utilization pattern of isolates gro wing in medium containing glyphosate as sole phosphorus sou rce Source of variation d.f. s.s. m.s. v.r. F pr. Treatment 6 0.831376 0.138 563 47.33 <.001 Residual 329 0.963230 0.002 928 Total 335 1.794606 Grand mean 0.0593
*** Least significant differences of means (5% leve l) *** l.s.d. 0.02173
147
One-sample T-test for Acetobacter sp. Variate: OD. Sample Size Mean Variance St andard Standard error de viation of mean OD 17 0.1172 0.002230 0.04722 0.01145 95% confidence interval for mean: (0.09296, 0.1415) Test statistic t = 10.24 on 16 d.f. Probability < 0.001
One-sample T-test for P. fluorescens Variate: OD. Sample Size Mean Variance St andard Standard error de viation of mean OD 17 0.1792 0.007275 0.08529 0.02069 95% confidence interval for mean: (0.1353, 0.2230) Test statistic t = 8.66 on 16 d.f. Probability < 0.001
148
ANOVA Table for time course of growth for Acetobacter sp and P. fluorescens Variate: OD
Source of variation d.f. s.s. m.s. v.r. F pr. Treatment 1 0.032612 0.03261 2 6.86 0.013 Residual 32 0.152082 0.00475 3 Total 33 0.184694 ***** Tables of means ***** Grand mean 0.148 Treatment Acetobacter sp P. fluorescens 0.117 0.179 l.s.d. 0.0482
149
ANOVA Table the effect of carbon source on Acetobacter sp
Source of variation d.f. s.s. m.s. v.r. F pr. Time 10 1.204E-02 1.204E-03 778.75 <.001 Carbon_source 3 1.937E-01 6.457E-02 41778.04 <.001 Time.Carbon_source 30 2.471E-02 8.238E-04 533.03 <.001 Residual 88 1.360E-04 1.545E-06 Total 131 2.306E-01 Grand mean 0.03395 Carbon_source Glucose Gly and Glu Glypho sate Gpi 0.01773 0.10000 0.0080 6 0.01000 l.s.d. 0.002017 ANOVA Table for the effect of carbon source on P. fluorescens
Source of variation d.f. s.s. m.s. v.r. F pr. Time 10 2.134E-02 2.134 E-03 1135.88 <.001 Carbon_source 3 1.514E-01 5.048 E-02 26869.52 <.001 Time.Carbon_source 30 3.837E-02 1.279 E-03 680.84 <.001 Residual 88 1.653E-04 1.879 E-06 Total 131 2.113E-01 Variate: OD Grand mean 0.03161 Carbon_source Glucose Gly and Glu Glyph osate Gpi 0.01594 0.09003 0.013 21 0.00727 l.s.d. 0.002224
150
ANOVA Table for the effect of different concentrations of glyp hosate on growth of Acetobacter sp Source of variation d.f. s.s. m.s. v.r. F pr. Time 9 0.19495099 0.02166 122 1568.14 <.001 concentration 4 0.38321916 0.09580 479 6935.67 <.001 Time.concentration 36 0.11484217 0.00319 006 230.94 <.001 Residual 100 0.00138133 0.00001 381 Total 149 0.69439366 Tables of means Variate: OD Grand mean 0.07066 concentration 100mg/ml 250mg/ml 25mg/ml 50mg/m l 7.2mg/ml 0.02950 0.00600 0.10900 0.0647 0 0.14410 l.s.d. 0.006021
ANOVA Table for the effect of different concentrations of glyp hosate on growth of P. fluorescens Source of variation d.f. s.s. m.s. v.r. F pr. Time 9 0.16883907 0.0187 5990 1632.24 <.001 Concentration 4 0.26713691 0.0667 8423 5810.69 <.001 Time.Concentration 36 0.08725003 0.0024 2361 210.87 <.001 Residual 100 0.00114933 0.0000 1149 Total 149 0.52437534 Tables of means Variate: OD Grand mean 0.06138 Concentration 100mg/ml 250mg/ml 25mg/ml 50mg/m l 7.2mg/ml
0.02527 0.00390 0.10300 0.06273 0.11200 l.s.d. 0.005492
151
ANOVA Table for the effect of nitrogen supplementation on grow th of Acetobacter sp
Source of variation d.f. s.s. m.s. v.r. F pr. Time 14 1.49805926 0.1070 0423 4681.31 <.001 Treatment 4 1.16270086 0.2906 7522 12716.69 <.001 Time.Treatment 56 0.32997127 0.0058 9234 257.78 <.001 Residual 150 0.00342867 0.0000 2286 Total 224 2.99416006 ***** Tables of means ***** Variate: OD Grand mean 0.19975 Treatment NH 4Cl (NH 4) 2SO4 Peptone Tryptone Yeast extract 0.12200 0.11869 0.30402 0.206 27 0.24778 l.s.d. 0.007713
ANOVA Table for the effect of nitrogen supplementation on grow th of P. fluorescens Source of variation d.f. s.s. m.s. v.r. F pr. Time 14 1.83775613 0.131 26829 8030.26 <.001 Treatment 4 0.97586580 0.243 96645 14924.50 <.001 Time.Treatment 56 0.48900041 0.008 73215 534.18 <.001 Residual 150 0.00245201 0.000 01635 Total 224 3.30507435 ***** Tables of means ***** Variate: OD Grand mean 0.21254 Treatment NH 4Cl (NH 4) 2SO4 Peptone Tryptone Yeast extract 0.11838 0.17122 0.31280 0.215 33 0.24498 l.s.d. 0.006523
152
ANOVA Table for the effect of different concentrations of heav y metals on the growth of Acetobacter sp Source of variation d.f. s.s. m.s. v.r. F pr. Metal 3 0.22497857 0.07499 286 5125.50 <.001 Time 10 1.44128827 0.14412 883 9850.71 <.001 Treatment 2 0.09664791 0.04832 396 3302.78 <.001 Metal.Time 30 0.15772462 0.00525 749 359.33 <.001 Metal.Treatment 6 0.26870348 0.04478 391 3060.83 <.001 Time.Treatment 20 0.02833364 0.00141 668 96.83 <.001 Metal.Time.Treatment 60 0.18213341 0.00303 556 207.47 <.001 Residual 264 0.00386267 0.00001 463 Total 395 2.40367257 ***** Tables of means ***** Variate: OD Grand mean 0.11547 Metal Cadmium Chromium Lead Zinc 0.11430 0.07831 0.14347 0.12578 Treatment 100µg/ml 500µg/ml 50µg/ml 0.10445 0.13756 0.10439 l.s.d. Metal Metal.Time.Tre atment 0.006149 0.001070 ANOVA Table for the effect of different concentrations of heav y metals on the growth of P. fluorescens Source of variation d.f. s.s. m.s. v.r. F pr. Metal 3 0.07994625 0.02664 875 534.19 <.001 Time 10 1.49061996 0.14906 200 2988.03 <.001 Treatment 2 0.05459711 0.02729 855 547.21 <.001 Metal.Time 30 0.11286267 0.00376 209 75.41 <.001 Metal.Treatment 6 0.21515459 0.03585 910 718.82 <.001 Time.Treatment 20 0.02750995 0.00137 550 27.57 <.001 Metal.Time.Treatment 60 0.18473591 0.00307 893 61.72 <.001 Residual 264 0.01317000 0.00004 989 Total 395 2.17859643 ***** Tables of means ***** Variate: OD Grand mean 0.11296 Metal Cadmium Chromium Lead Zinc 0.10041 0.09927 0.11838 0.13378 Treatment 100µg/ml 500µg/ml 50µg/ml 0.10139 0.12906 0.10844 l.s.d. Metal Metal.Time.Treat ment 0.001977 0.011355
153
ANOVA Table for Effect of pH on Glyphosate Degradation by Organism Acetobacter sp
Source of variation d.f. s.s. m.s. v.r. F pr. Time 9 0.3454860 0.0383873 326.99 <.001 pH 5 0.0786586 0.0157317 134.01 <.001 Time.pH 45 0.0354002 0.0007867 6.70 <.001 Residual 120 0.0140873 0.0001174 Total 179 0.4736321 Variate: OD Grand mean 0.10809 pH pH 4 pH 5 pH 6 pH 7 pH 8 pH 9 0.07852 0.13260 0.10498 0.10537 0.13667 0.09040 l.s.d. 0.017516 ANOVA Table for the effect of pH on the degradation of glyphos ate by organism P. fluorescens Source of variation d.f. s.s. m.s. v.r. F pr. Time 9 0.53605445 0.0595616 1 3603.62 <.001 pH 5 0.04604744 0.0092094 9 557.20 <.001 Time.pH 45 0.07319431 0.0016265 4 98.41 <.001 Residual 120 0.00198339 0.0000165 3 Total 179 0.65727959 ***** Tables of means ***** Variate: OD Grand mean 0.11375 pH pH 4 pH 5 pH 6 pH 7 pH 8 pH 9 0.13137 0.12537 0.10140 0.10984 0.12 773 0.08680 l.s.d. 0.006572
