Atrazine Mineralization Potential and Catabolic Gene Detection In

ATRAZINE MINERALIZATION POTENTIAL AND CATABOLIC GENE DETECTION IN

AGRICULTURAL AND WETLAND SITES

DISSERTATION

Presented in Partial Fulfillment of the Requirements of the Degree Doctor of Philosophy

in the Graduate School of The Ohio State University

Kristen Lynn Anderson, B. S.

* * * * *

The Ohio State University

2003

Dissertation Committee: Approved by

Dr. Olli H. Tuovinen, Advisor

Dr. Michael J. Boehm ______

Dr. Mark Morrison Advisor

Dr. Samuel J. Traina Department of Microbiology ABSTRACT

Atrazine (2-chloro-4-ethylamine-6-isopropylamino-1,3,5 triazine) is a commonly applied herbicide in corn fields. Although the fate of atrazine in agricultural systems has been well studied, the environmental fate of atrazine in wetland systems is less well characterized. The majority of research in this area has focused on aerobic mineralization of atrazine, although anaerobic conditions are commonly found in wetland sediments and bulk soils associated with agricultural fields. The hypothesis for this work was that atrazine would be actively mineralized in agricultural and wetland sites. It was further hypothesized that active mineralization in soils could be correlated with the presence of selected genes involved in atrazine metabolism. Soil, sediment, and water samples were obtained from three sites in Ohio. Atrazine mineralization was investigated under aerobic and anaerobic conditions in these samples using a biometer system in

14 which CO2 evolution was correlated with atrazine mineralization. All samples mineralized atrazine under aerobic conditions. Under anaerobic conditions, some external electron acceptor amendments inhibited mineralization, while others enhanced it. The effect on mineralization varied with the sample and season. Attempts were made to amplify some of the genes involved in atrazine mineralization. Community DNA was

ii isolated from each sample and PCR amplification using three primer sets specific for atzA, trzD, and trzN was performed. Although PCR inhibitors were present in many of the samples, trzN and trzD could be successfully amplified from some DNA samples. All

Defiance samples tested positive for trzN with the exception of field B and reservoir water samples. An analog of trzD was amplified from DNA extracted from Defiance and

OSU wetland samples. The atzA gene was not detected in any of the samples. An atrazine mineralizing bacterium was isolated from Piketon soil samples and another was obtained from Defiance corn soil. Characterization of these organisms by 16S rDNA sequencing and FAME analysis indicated that both organisms were Arthrobacter spp.

From these results, it is clear that the potential for atrazine mineralization was identified in agricultural soils and wetland sediments. No correlation could be made between atrazine mineralization and the presence of any of the tested genes involved in atrazine mineralization.

iii

To Matt - for supporting me through all the good times, but especially for being there

through all the bad times.

iv ACKNOWLEDGMENTS

I would first like to thank my advisor, Dr. Olli Tuovinen, for encouraging me and helping me to keep on track. I would also like to thank him for giving me the freedom to pursue avenues that we hadn’t initially considered in this project.

I would also like to thank my committee members, Dr. Sam Traina, Dr. Mike

Boehm, Dr. Mark Morrison, and Dr. Michelle Rondon for their insights into my project and their helpful suggestions.

I would like to thank Mr. Peter Bierman, former extension agent, and Mr. Wayne

Lewis, farm manager, at the Piketon Research and Extension Center for helping me gather the crop and herbicide application information on the Piketon sites.

Dr. Kent Harrison graciously allowed me to use his biological oxidizer and Dr.

Xiaoyu Yang instructed me on how to operate the oxidizer and answered my questions on how it actually worked.

Dr. J. S. Karns generously provided the Escherichia coli construct containing the trzD gene and Dr. M. de Souza provided the E. coli construct containing the atzA gene.

Dr. W. Mulbry provided the pWM221011 plasmid containing the trzN gene and also helped me optimize the PCR reaction to amplify this gene.

v Kevin Wheeler, formerly of the University of Dayton, performed the atzA and trzD

PCR amplification reactions on the OSU sediment samples.

I would like to thank Dr. Mark Radosevich for the FAME analysis on the soil bacterial isolates.

The Defiance Agricultural Research Association, especially Bruce Clevenger, provided me with information on the Defiance site.

I would also extend my thanks to past and present members of the Tuovinen lab for their encouragement and discussions on my project. I would especially like to thank

Dr. Ellen Duffy (nee Ostrofsky) who helped me get started on this project and also provided the MPN calculation program used to determine MPN estimates. Dr.

Duongruitai Nicomrat helped me with the 16 rDNA sequencing and provided many helpful suggestions on DNA extraction and PCR amplification from soil samples.

This research was supported by the United States Department of Agriculture,

National Research Initiative Program (Grant No. 98-35107-6388)

vi VITA

October 23, 1973……………………………Born – Sewickley, PA

1996…………………………………………..B. S. Molecular biology/biotechnology,

Clarion University of Pennsylvania

1997 – present……………………………….Graduate Teaching Associate,

The Ohio State University

PUBLICATIONS

Research Publications

1. Kwak, J., L. A. McCue, K. Trzcianka, and K. E. Kendrick. 2001. Identification and characterization of a developmentally regulated protein, EshA, required for sporogenic hyphal branches in Streptomyces griseus. J. Bacteriol.183:3004- 3015.

2. Anderson, K. L., K. A. Wheeler, J. B. Robinson, and O. H. Tuovinen. 2002. Atrazine mineralization in two wetlands. Wat. Res. 36:4785-4794.

vii FIELDS OF STUDY

Major field: Microbiology

viii TABLE OF CONTENTS

Page

Abstract ...... ii

Dedication ...... iv

Acknowledgements ...... v

Vita ...... vii

List of Tables...... xii

List of Figures...... xiii

Chapters

1. Introduction...... 1

1.1 Atrazine mineralization in agricultural soils...... 2 1.1.1 Aerobic mineralization in surface and subsurface soils ...... 2 1.1.2 Effect of the plant rhizosphere ...... 4 1.1.3 Anaerobic mineralization of atrazine in bulk soil...... 5 1.1.4 Effect of amendments on atrazine mineralization...... 6 1.2 Atrazine mineralization in wetland sediments...... 7 1.2.1 Aerobic mineralization in wetland sediments...... 7 1.2.2 Anaerobic mineralization in wetland sediments...... 9 1.3 Abiotic dechlorination of atrazine...... 10 1.4 Genetic basis of atrazine degradation...... 11 1.4.1 The thc genes...... 11 1.4.2 The atrA gene...... 11 1.4.3 The trz genes: Pseudomonas str. NRRLB-12227 ...... 14 1.4.4 The trz genes: Nocardiodes str. C190 ...... 14 1.4.5 The atz genes...... 15 1.5 Statement of purpose...... 17

2. Evaluation of the atrazine mineralization potential at an agricultural research site by soil biometer studies and PCR amplification of catabolic genes ...... 19

2.1 Introduction...... 19 2.2 Materials and methods...... 22 2.2.1 Site description...... 22 2.2.2 Sampling...... 22 2.2.3 Mineralization studies...... 25 2.2.4 Data analysis...... 27 2.2.5 Mass balances...... 27 2.2.6 DNA isolation...... 28 2.2.7 PCR amplification...... 29 2.2.8 Most Probable Number (MPN) technique...... 31 2.2.9 Enrichment for atrazine-mineralizing organisms ...... 32 2.2.10 16S rDNA sequencing...... 32 2.2.11 FAME analysis...... 33 2.3 Results and discussion...... 33 2.3.1 Mineralization studies - Yearly data...... 33 2.3.1.1 1999 sampling year ...... 33 2.3.1.2 2000 sampling year ...... 38 2.3.1.3 2001 sampling year ...... 50 2.3.1.4 2002 sampling year ...... 55 2.3.2 Mineralization data – Summaries ...... 62 2.3.3 Catabolic marker genes...... 69 2.3.4 Isolate characterization...... 72 2.3.5 Most Probable Number analysis...... 77 2.4 Summary of results ...... 81

3. Atrazine mineralization potential in two wetlands...... 82

3.1 Introduction……...... 82 3.2 Materials and methods...... 86 3.2.1 Sampling...... 86 3.2.2 Mineralization studies...... 89 3.2.3 Data analysis...... 90 3.2.4 Mass balances...... 91 3.2.5 DNA isolation and PCR amplification ...... 91 3.2.6 Southern hybridization analysis...... 93 3.3 Results and discussion...... 94 3.3.1 Mineralization data...... 94 3.3.2 Catabolic gene detection...... 117 3.4 Summary of results ...... 122

4. Determination of atrazine mineralization potential using soil and DNA detection methods in a wetland reservoir subirrigation system...... 124

4.1 Introduction……...... 124 4.2 Materials and methods...... 126 4.2.1 Site description...... 126 4.2.2 Sampling...... 128 4.2.3 Mineralization studies...... 130 4.2.4 Data analysis...... 131 4.2.5 Mass balances...... 131 4.2.6 DNA isolation...... 131 4.2.7 PCR amplification...... 133 4.2.8 Most Probable Number (MPN) technique...... 134 4.2.9 Enrichment for atrazine-mineralizing organisms ...... 135 4.2.10 16S rDNA sequencing...... 136 4.2.11 FAME analysis...... 136 4.3 Results and discussion...... 137 4.3.1 Mineralization data – Yearly data ...... 137 4.3.1.1 1999 sampling year ...... 137 4.3.1.2 2000 sampling year ...... 137 4.3.1.3 2001 sampling year ...... 152 4.3.1.4 2002 sampling year ...... 161 4.3.2 Mineralization data – Summaries ...... 167 4.3.3 PCR amplification...... 173 4.3.4 Isolate characterization...... 177 4.3.5 Most Probable Number (MPN) analysis ...... 179 4.4 Summary of results ...... 184

5. Future directions………...... 186

References………………...... 190

LIST OF TABLES

Table Page

1.1 Atrazine metabolic intermediates and corresponding genes and enzymes...... 13

2.1 Field sampling and atrazine application dates for Piketon, OH sites...... 24

2.2 Oligonucleotide sequences and annealing temperatures used in PCR amplification...... 30

2.3 Kinetic parameters and half-life values for Piketon 2000 samples ...... 45

2.4 Kinetic parameters and half-life values for Piketon 2001 samples ...... 54

2.5 Kinetic parameters and half-life values for Piketon 2002 samples ...... 60

2.6 MPN estimates for CC soils incubated with various carbon sources...... 80

3.1 Kinetic parameters and half-life values for Olentangy wetland 2000 samples...112

4.1 Field sampling and atrazine application dates for Defiance, OH sites...... 129

4.2 Kinetic parameters and half-life values for Defiance 2000 samples ...... 140

4.3 Kinetic parameters and half-life values for Defiance 2001 samples ...... 155

4.4 Kinetic parameters and half-life values for Defiance 2002 samples ...... 166

4.5 MPN estimates for field A samples incubated with various carbon sources...... 183

xii LIST OF FIGURES

Figure Page

1.1 Aerobic atrazine mineralization pathway ...... 12

2.1 Piketon Research and Extension Center site map ...... 23

2.2 Biometer assembly...... 26

2.3 Anaerobic mineralization in August, 1999 CC samples...... 35

2.4 Aerobic and anaerobic mineralization in August, 1999 CR samples ...... 36

2.5 Anaerobic mineralization in August, 1999 CR samples after additional electron acceptor amendment ...... 37

2.6 Aerobic and anaerobic mineralization in August, 1999 RZ samples ...... 39

2.7 Aerobic and anaerobic mineralization in April, 2000 CC samples ...... 40

2.8 Aerobic and anaerobic mineralization in April, 2000 CR samples ...... 41

2.9 Aerobic and anaerobic mineralization in April, 2000 RZ samples...... 43

2.10 Aerobic and anaerobic mineralization in June, 2000 CC samples ...... 44

2.11 Aerobic mineralization of June 2000 CC samples amended with various electron

acceptors ...... 46

2.12 Anaerobic and aerobic mineralization in June, 2000 CR samples ...... 47

2.13 Aerobic mineralization in June, 2000 RZ samples...... 48

xiii 2.14 Aerobic and anaerobic mineralization in September, 2000 CC samples...... 49

2.15 Aerobic and anaerobic mineralization in September, 2000 CR samples...... 51

2.16 Aerobic mineralization in September, 2000 RZ samples...... 52

2.17 Aerobic mineralization in June, 2001 Piketon samples ...... 53

2.18 Aerobic mineralization in June, 2001 Scioto River water samples ...... 56

2.19 Aerobic mineralization in August, 2001 Piketon samples ...... 57

2.20 Aerobic mineralization in August, 2001 Scioto River water samples...... 58

2.21 Aerobic mineralization in June, 2002 Piketon samples ...... 59

2.22 Aerobic mineralization in June, 2002 Scioto River water and sediment samples ...... 61

2.23 Aerobic mineralization in July, 2002 Scioto River water samples...... 63

2.24 Aerobic mineralization in September, 2002 Piketon samples...... 64

2.25 Aerobic mineralization in September, 2002 Scioto River water samples ...... 65

2.26 Crop summary for CC site ...... 66

2.27 Crop summary for CR site ...... 68

2.28 Summary of RZ aerobic mineralization over four years...... 70

2.29 Effect of atrazine application on atrazine mineralization...... 71

2.30 Agarose gel electrophoresis of Piketon genomic DNA amplified with atzA-specific primers...... 73

2.31 Agarose gel electrophoresis of Piketon genomic DNA amplified with trzD-specific primers...... 74

2.32 Agarose gel electrophoresis for detection of PCR inhibitors present in Piketon samples ...... 75

2.33 Agarose gel electrophoresis of P-C 3 genomic DNA amplified with atzA-specific primers...... 76

xiv 2.34 Agarose gel electrophoresis of P-C 3 genomic DNA amplified with trzD-specific primers...... 78

2.35 Agarose gel electrophoresis of P-C 3 genomic DNA amplified with trzN-specific primers...... 79

3.1 Site map for the Olentangy wetland and Cedar Bog ...... 87

3.2 Aerobic and anaerobic mineralization in August, 1999 inlet composite samples ...... 95

3.3 Aerobic and anaerobic mineralization in August, 1999 outlet composite samples ...... 96

3.4 Aerobic mineralization in April, 2000 Olentangy wetland composite and Cedar Bog sediment samples ...... 96

3.5 Aerobic and anaerobic mineralization in April, 2000 inlet composite samples ....98

3.6 Aerobic and anaerobic mineralization in April, 2000 outlet composite samples ...... 100

3.7 Aerobic mineralization in April and August, 2000 sectioned samples ...... 101

3.8 Aerobic and anaerobic mineralization in the April, 2000 0-5 cm inlet section sample ...... 103

3.9 Aerobic and anaerobic mineralization in the April, 2000 5-10 cm inlet section sample ...... 104

3.10 Aerobic and anaerobic mineralization in the April, 2000 10-15 cm inlet section sample ...... 105

3.11 Aerobic and anaerobic mineralization in the April, 2000 0-5 cm outlet section sample ...... 106

3.12 Aerobic and anaerobic mineralization in the April, 2000 5-10 cm outlet section sample ...... 107

3.13 Aerobic and anaerobic mineralization in the April, 2000 10-15 cm outlet section sample ...... 108

3.14 Anaerobic mineralization in August, 2000 inlet section samples ...... 109

3.15 Anaerobic mineralization in August, 2000 outlet section samples...... 110

xv 3.16 Aerobic mineralization in August, 2000 Olentangy River and OSU wetland water samples ...... 115

3.17 Aerobic mineralization in June, 2002 Olentangy River and OSU wetland water samples ...... 116

3.18 PCR amplification and Southern blot analysis of genomic DNA extracted from Olentangy sediment samples using trzD specific primers ...... 118

3.19 Agarose gel electrophoresis of genomic DNA extracted from Olentangy water samples and amplified with atzA specific primers ...... 120

3.20 Agarose gel electrophoresis of genomic DNA extracted from Olentangy water samples and amplified with trzD specific primers ...... 121

4.1 Site map for Defiance, OH...... 127

4.2 Aerobic and anaerobic mineralization in August, 1999 field A samples ...... 138

4.3 Aerobic and anaerobic mineralization in August, 1999 field B samples ...... 139

4.4 Aerobic and anaerobic mineralization in April, 2000 field A samples ...... 141

4.5 Aerobic and anaerobic mineralization in April, 2000 field B samples ...... 143

4.6 Aerobic and anaerobic mineralization in April, 2000 wetland sediment samples ...... 144

4.7 Aerobic and anaerobic mineralization in May, 2000 field A samples...... 145

4.8 Aerobic and anaerobic mineralization in May, 2000 field B samples...... 146

4.9 Aerobic mineralization in May, 2000 wetland sediment samples ...... 148

4.10 Aerobic and anaerobic mineralization in September, 2000 field A samples...... 149

4.11 Aerobic and anaerobic mineralization in September, 2000 field B samples...... 150

4.12 Aerobic mineralization in September, 2000 wetland sediment and reservoir water samples ...... 151

4.13 Aerobic mineralization in June, 2001 agricultural samples...... 153

4.14 Aerobic and anaerobic mineralization in June, 2001 wetland sediment samples ...... 154

xvi 4.15 Aerobic mineralization in June, 2001 wetland and reservoir water samples .....156

4.16 Aerobic mineralization in August, 2001 agricultural samples ...... 158

4.17 Aerobic mineralization in August, 2001 wetland sediment and water and reservoir water samples...... 159

4.18 Aerobic mineralization in June, 2002 agricultural samples...... 160

4.19 Aerobic mineralization in June, 2002 wetland sediment and water and reservoir water samples...... 162

4.20 Aerobic mineralization in July, 2002 wetland sediment and water and reservoir water samples...... 163

4.21 Aerobic mineralization in September, 2002 agricultural samples ...... 164

4.22 Aerobic mineralization in September, 2002 wetland sediment and water and reservoir water samples...... 165

4.23 Crop rotation summary for field A samples...... 168

4.24 Crop rotation summary for field B samples...... 169

4.25 Atrazine application effect on field A and B samples...... 170

4.26 Summary of aerobic wetland sediment mineralization ...... 172

4.27 Agarose gel electrophoresis of Defiance genomic DNA amplified with atzA- specific primers...... 174

4.28 Agarose gel electrophoresis of Defiance genomic DNA amplified with trzN- specific primers...... 175

4.29 Agarose gel electrophoresis of Defiance genomic DNA amplified with trzD- specific primers...... 176

4.30 Agarose gel electrophoresis for detection of PCR inhibitors present in Defiance samples ...... 178

4.31 Agarose gel electrophoresis of C5-2B genomic DNA amplified with atzA-specific primers...... 180

4.32 Agarose gel electrophoresis of C5-2B genomic DNA amplified with trzN-specific primers...... 181

xvii 4.33 Agarose gel electrophoresis of C5-2B genomic DNA amplified with trzD-specific primers...... 182

xviii CHAPTER 1

INTRODUCTION

Atrazine (2-chloro-4-ethylamine-6-isopropylamino-1,3,5 triazine) is a member of the s- triazine herbicide family and is used extensively throughout the world as a photosynthesis inhibitor of broad-leaf weeds. It is applied primarily to corn crops prior to weed emergence. It is considered to be a moderately recalcitrant chemical with half-lives in the range of days to weeks. The U.S. EPA estimated that 74-80 million pounds (23 –

33 million kg) of atrazine was applied in 1999 in the United States alone (17). Atrazine runoff from agricultural fields has been identified as a non-point source pollutant and is a common contaminant in ground water. The U.S. EPA has mandated that atrazine concentrations should not exceed 3 ppb in drinking water. However, no standards exist for the concentration of atrazine in surface or groundwater and concentrations in the range of 3-12 ppb are not uncommon following storm events (http://www.epa.state.oh.us

/ddagw /pestsbw2.pdf). This is a concern because atrazine has recently been identified

1 as an endocrine disruptor in frogs when administered in doses as low as 0.1 ppb, which is well below the maximum contaminant level of 3 ppb (23).

A number of microorganisms have been isolated that have the ability to partially or completely degrade atrazine. Pseudomonas strain ADP was isolated from an atrazine spill site and was one of the first organisms identified with the ability to completely mineralize atrazine (31). Many of the genes specific to atrazine mineralization were first identified in Pseudomonas strain ADP and it is the most commonly studied organism in atrazine mineralization. Ralstonia basilensis (formerly M-91) was identified from an agricultural site with a history of atrazine application (44, 60). More recently, several gram positive organisms have been identified with the ability to mineralize atrazine.

Nocardiodes spp. have been identified which can mineralize atrazine (65), but surprisingly, the atzABC genes were not detected. This was the first report of a pure culture of an atrazine-mineralizing bacterium that did not have these genes (65). Several reports of Arthrobacter spp. with the ability to mineralize atrazine have been published which also lack some or all of the atzABC genes (46, 62). These reports indicate the ability to mineralize atrazine is widespread and that the atzABC genes are not completely conserved across all organisms.

1.1: Atrazine mineralization in agricultural soils

1.1.1 Aerobic mineralization in surface and subsurface soils. Much of the initial research on the fate of atrazine in agricultural soils was focused on biotransformation in surface and subsurface soils. Vanderheyden et al. (66) reported that rapid degradation took place in surface soils with a 10 year history of atrazine application. However,

2 biodegradation of atrazine in subsurface samples was very low for most of the samples although samples which did degrade atrazine did so rapidly following a lag period.

Another study (43) involving samples of surface and subsurface soils indicated that surface soil samples showed a rapid mineralization of atrazine while subsurface degradation was generally weak and spatially variable. Interestingly, soil samples from sites with a history of atrazine application showed a rapid rate and high extent of mineralization whereas samples from a non-history site exhibited slow mineralization rates. This suggests that atrazine application may enhance the mineralization activity of the existing microbial population. Addition of a Ralstonia basilensis culture to soils with a low rate of mineralization caused an increase in mineralization rates (43). These findings suggest that the lack of mineralization in subsurface soils may be due primarily to low numbers of microorganisms rather than low nutrient availability, high atrazine sorption to soil particles or other soil properties. Investigation into the effect of agricultural management practices indicated that fields under conventional and ridge tilling practices did not exhibit spatial variability in the rate of atrazine mineralization, suggesting that the mixing caused by tilling distributes the microbes throughout the agricultural fields (39).

Differences were observed in the rate and extent of mineralization for soils which received an annual application of atrazine (CC site) as compared to soils which received atrazine applications every three years (CR site). These data suggest that repeated atrazine applications enhanced the mineralizing ability of the resident microbial population. High levels of residual atrazine in the CC soils which receive yearly atrazine applications may serve as an inducer of the atrazine mineralizing genes. Lower residual atrazine concentrations observed in the CR site may be insufficient to promote a

3 sustained induction of the atrazine-mineralizing organisms, leading to a lower rate and extent of mineralization in these samples (39).

1.1.2 Effect of the plant rhizosphere. Plant rhizospheres, which exude nutrients into the root-soil interface, have been shown to promote the degradation of certain herbicides, including atrazine (41). Because these areas support diverse microbial populations, the possibility of enhanced biodegradation of pesticides in vegetated areas has sparked interest. Arthur et al. (5) reported higher mineralization in soils which contained Kochia scoparia vegetation as compared to non-vegetated soils. In addition, atrazine was less persistent in rhizosphere soils. A 14C-most-probable-number analysis showed an increase in the number of atrazine degrading organisms in the vegetated soils as compared to the non-vegetated samples (5). These results suggest a positive correlation between the presence of vegetation and the increased degradation of atrazine.

The effect of maize rhizospheres on atrazine mineralization has also been studied. It was determined that the presence of maize increased the microbial biomass regardless of the presence of atrazine (32). Based on Biolog measurements of metabolic diversity, it was found that the presence of the maize rhizosphere as well as atrazine, although to a lesser extent, caused a shift in the metabolic patterns of the microbial population. As has been noted in other studies, the pre-application of atrazine significantly increased the rate of atrazine mineralization (32). A comparison of bulk soil and maize-containing soil, both pretreated with atrazine, showed that the maize- containing soil increased the rate of mineralization, although no significant difference in

4 the extent of mineralization was noted (42). Profiles of the microbial populations based on ribosomal intergenic spacer analysis indicated no large changes in the soil microbial populations after the addition of atrazine (42). This may reflect the presence of a stable microbial population or a lack of sensitivity in the assay which may not be able to detect subtle changes in the populations. Nested quantitative competitive (QC) PCR detection of atzC indicated a spike in the relative amount of atzC detected. This effect was transient and could only be detected in soils pretreated with atrazine (42). These studies indicate that the presence of maize rhizosphere may positively influence atrazine mineralization in agricultural soils.

1.1.3 Anaerobic mineralization of atrazine in bulk soil. Relatively little is known about anaerobic mineralization of atrazine. None of the currently known steps in the atrazine mineralization process requires molecular oxygen, so it is possible that anaerobic mineralization can occur using the same enzymes and catalytic steps found in aerobic mineralization. Both Pseudomonas strain ADP and R. basilensis are capable of anaerobic mineralization of atrazine under denitrifying conditions (11, 28). The only report of anaerobic mineralization in bulk soils showed a low, but measurable loss of atrazine under anaerobic conditions (2). Under anaerobic conditions, surface soils had a half-life of 124 d while subsurface soils had a half-life of 408 d. The rate of mineralization appeared to be most affected by soil depth and redox conditions. Subsurface samples were less able to mineralize atrazine. Moderately reducing conditions in the soil samples seemed to promote atrazine degradation while oxidizing conditions appeared to lengthen the half-life (2).

5 1.1.4 Effect of amendments on atrazine mineralization. The ability of R. basilensis to mineralize atrazine was reduced when incubated in the presence of ammonium sulfate, nitrate or glutamate (21). These results suggest that the availability of exogenous sources of N might inhibit the atrazine metabolic pathway. Abdelhafid et al. (1) investigated the effect of various C and N sources on atrazine mineralization in history and non-history soils. All N sources inhibited atrazine mineralization in both history and non-history soils. The largest decreases were seen with inorganic N sources such as

Ca(NO3)2 and (NH4)2SO4. Organic N sources such as adenine and arginine showed increasing inhibition of atrazine as the availability of the nitrogen increased. This inhibition may be caused by repression of the atrazine degradation pathway due to availability of more easily utilized N sources. However, inorganic N sources must be added in large quantities in order to decrease mineralization. This suggests that inorganic N may affect the synthesis or activity of one or more of the enzymes involved in mineralization rather than depressing the atrazine degradation pathway (1). In history soils, only the addition of compost and glucose caused decreased atrazine mineralization. Compost increased sorption of atrazine, making it biologically unavailable for degradation (1). Although straw also increased adsorption of atrazine, it did not seem to decrease the availability of atrazine for degradation. In the non-history soil, addition of organic C sources increased the degradation of atrazine for all amendments. Addition of both organic C sources and inorganic N sources decreased the mineralization in both soils as compared to mineralization with the organic C source alone (1). A further study of the effect of organic amendments showed that atrazine mineralization was enhanced by the addition of manure, peat, and cornstalk amendments (35). Addition of sawdust

6 appeared to stimulate a general microbial population and its activity, but failed to increase the degradation of atrazine. This suggests that although the general microbial population was stimulated, the subpopulation responsible for atrazine degradation did not respond in the same way (35).

1.2: Atrazine mineralization in wetland sediments

1.2.1 Aerobic mineralization in wetland sediments. A preliminary study of redwater and blackwater wetland sediments indicated that atrazine was mineralized in small amounts in these sediments (20). Blackwater wetlands are characterized by high organic

C concentrations and low concentrations of dissolved and particulate-associated inorganic ions. Redwater wetlands contain high concentrations of suspended clays and inorganic nutrients. The addition of N as NH4NO3 inhibited the mineralization of atrazine in these wetland sediments (20). Mesocosms designed to evaluate the fate of pesticides in wetland sediments were inoculated with set concentrations of atrazine, which were applied as simulated runoff events (34). Approximately 35 days after atrazine addition, approximately 68% of the 74 µg/l atrazine originally added was transferred or transformed out of the water column. The half-life of atrazine in these mesocosms was

16-21 d. Wetlands which had 147 µg/l atrazine applied transferred or transformed only

35% of the atrazine out of the water column and had a half-life of 46-48 d (34). Two studies performed with sediments from a mountainous wetland indicated that atrazine was removed from wetland sediments under aerobic conditions as determined by HPLC

(26, 27). This wetland is fed by two streams which serve as an agricultural watershed upstream of the wetland. Initial measurements of atrazine degradation indicated that an

7 additional C source such as sucrose was required for significant atrazine mineralization.

Samples which were incubated with atrazine as the sole source of C and N failed to degrade any atrazine. This suggests that atrazine was being degraded through cometabolism with the sucrose. In addition, the effect of exogenous N as NH4NO3 on atrazine removal was investigated. Contrary to the results of Moore et al. (34), no inhibition of atrazine removal was noted (26). However, aerobic mineralization was impacted by the organic C content in the samples. Samples with high organic matter contents were able to degrade more atrazine than samples with low contents.

Cometabolism of the atrazine in conjunction with the organic matter may explain this enhancement (27). In contrast, sediments from a wetland fed from an agricultural catchment showed no measurable atrazine mineralization (30). Biometers incubated with sediments obtained from a constructed wetland used to treat irrigation runoff from a container nursery showed no significant mineralization of atrazine (47). Significant mineralization could only be achieved through the addition of soil from an atrazine-spill site. In unamended wetland sediments, <2% of the added atrazine was mineralized while approximately 30% of the atrazine was mineralized in augmented soils (47).

Further experiments involving the constructed wetland indicated that atrazine was removed from the water column after simulated run-off events (48). However, the primary method of removal was the sorption of atrazine to the wetland sediments rather than mineralization. The low MPN determinations of the numbers of atrazine- mineralizing organisms supported the lack of mineralization (48).

8 1.2.2 Anaerobic mineralization in wetland sediments. A study involving anaerobic mineralization in wetland sediments obtained from a sugar mill wastewater wetland indicated the disappearance of atrazine added to biometers (9). Additional nutrients appeared to enhance atrazine disappearance. Contrary to observed results in other studies, the addition of an N source did not seem to inhibit atrazine degradation (9).

Further study of this wetland indicated that approximately 50% of the added atrazine was degraded within 38 weeks of incubation. Acetic acid enhanced degradation, but examination of the total triazine species present revealed a high concentration of hydroxyatrazine, which may indicate chemical hydrolysis of atrazine rather than microbial degradation. Sodium acetate inhibited atrazine degradation, whereas methanol did not. Only glucose appeared to enhance degradation of atrazine in this system (10).

The effect of redox conditions on anaerobic degradation of atrazine were examined in soils from a swamp-forest wetland. Soils incubated under aerobic conditions showed a rapid degradation of atrazine under oxidizing conditions after 14 d.

Under reducing (anaerobic) conditions, atrazine degradation was much slower with atrazine still detectable after 99 d (12). Anaerobic mineralization in a groundwater fed wetland was showed a loss of less than 2% of the added atrazine which is at the level of radiochemical impurities for atrazine (30). Sediments obtained from a tidal freshwater wetland degraded 50% of the added atrazine in 38 d. Hydroxyatrazine was the major metabolite detected (52). In mountainous wetland sediments, atrazine was degraded more rapidly under anaerobic conditions than aerobic conditions, but only in the presence of added sucrose (99% removed in 15 d). Samples which were incubated with atrazine as the sole C and N source showed a low level of degradation. Addition of

9 NH4NO3 did not seem to inhibit anaerobic mineralization (26). Incubation of these sediments in bioreactors showed that samples with a high organic C content completely removed all added atrazine within 20 days of incubation. However, bioreactors with a low organic matter contents only degraded 19% of the atrazine after 50 days. These results indicate that additional C sources are required for significant atrazine degradation

(27).

1.3 Abiotic dechlorination of atrazine

The rate-limiting steps in the mineralization of atrazine are not known in pure culture systems or in the environment. Because dechlorination of atrazine may be a rate- limiting step in atrazine mineralization, abiotic mechanisms of atrazine dechlorination may be important in providing sufficient levels of hydroxyatrazine to microbial communities which do not have the enzymes required for atrazine dechlorination. Three mechanisms have been suggested for abiotic dechlorination: 1. reductive dechlorination by elemental sulfur or ferrous sulfate (or other Fe(II) salts), 2. photolytic hydrolysis in the presence of organic matter, and 3. low or high pH values (pH <6 or >9). Reductive dechlorination has been found to occur, but at slow rates (16). At low pH values (pH <6), protonation of atrazine allows nucleophilic displacement of the chlorine by water. Iron

(Fe0) is believed to provide the available surface area for the protonation of the triazine ring (16). Photolytic degradation has been reported in aqueous systems, but does not occur at wavelengths greater than 300 nm (Solomon). The rates are higher in the presence of humic acid (58).

10 1.4: Genetic basis of atrazine degradation

The intermediate compounds and genes commonly involved in atrazine metabolism are indicated in Figure 1.1. The intermediates are numbered and the corresponding names are listed in Table 1.1. The genes and their corresponding proteins are also listed in Table 1.1.

1.4.1 The thc genes. Studies involving Rhodococcus str. NI86/21 identified a cluster of genes that were involved in the dealkylation of atrazine to produce deethylatrazine and deisopropylatrazine (37). The thcB gene was cloned and identified as a cytochrome

P450 protein which could degrade both atrazine and s-ethyl dipropylthiocarbamate

(EPTC). Two accessory proteins, thcC and thcD were identified as an associated electron-supply system for the cytochrome P450 system (37). Production of thcE, an alcohol oxidoreductase, is induced in the presence of EPTC as well as s-triazine herbicides. The role of this protein in atrazine degradation is unclear (38) However, the thcBCD genes appear to be induced only by the addition of EPTC and other carbamates. Atrazine or other s-triazines do not seem to induce these genes (54). This suggests that, while the P450 system is able to degrade a wide variety of substrates, the regulation of these genes is linked most strongly to the presence of EPTC and other thiocarbamates.

1.4.2 The atrA gene. Rhodococcus str. TE1 was initially identified as an organism capable of degrading EPTC. Additional testing showed that this strain also had the ability

to degrade atrazine to deethylatrazine andCO deisopropylatrazine.2 + NH3 However,

11 OH Cl Cl atzA atrA, thcB N N N NNN 1 2 4 CH 3 CH 3 H3CH 2CHNN NHCH H3CH 2CHN N NHCH H3CH 2CHN N NH 2

atzB H3C atrA, H3C thcB

Cl OH

N N N N 3

5 NH 3 CH 3 H2NN NHCH HON NHCH

H3N atzC H3C

N N 6

HO N OH

atzD, trzD

OO atzE OO 7 8 C C C C O N NH 2 H2N N NH 2 H

atzF Urease

9 CO2 + NH4 N H3C CH 3

Figure 1.1: Pathway for aerobic mineralization of atrazine. Known genes are indicated next to the reaction catalyzed by that enzyme. Compounds, genes, and enzymes are listed in Table 1.1.

Compound Common name Gene Corresponding enzyme 1 Atrazine N/A N/A 2 Deisopropylatrazine atrA, thcB P450 enzyme system 3 Deethylatrazine atrA, thcB P450 enzyme system 4 Hydroxyatrazine atzA, trzN Atrazine chlorohydrolase 5 N-isopropylammelide atzB Hydroxyatrazine ethylaminohydrolase 6 Cyanuric acid atzC N-isopropylammelide isopropylaminohydrolase 7 Biuret atzD, trzD Cyanuric acid amidohydrolase 8 Allophanate atzE Biuret hydrolase 9 Urea atzF Allophanate hydrolase

Table 1.1: List of the common intermediates in the atrazine metabolic pathway as indicated in Figure 1.1. The genes and enzymes refer to the enzyme which catalyzes the reaction which forms the listed compound and its corresponding gene.

Rhodococcus str. TE1 was not able to use atrazine as the sole source of N (6). The degradation of atrazine strictly required oxygen which indicated that the N-dealkylation was catalyzed by an oxidative mechanism. The gene involved in EPTC degradation, eptA, was co-localized with atrA (atrazine degradation) and their activities could not be separated which suggests that these genes may be the same gene which encodes a protein with multiple functions (53). The evidence of the P450 system identified in

13 Rhodococcus str. NI86/21 that can degrade both EPTC and atrazine, in conjunction with the AtrA protein suggests that nonspecific metabolism of these compounds is not uncommon.

1.4.3 The trz genes: Pseudomonas strain NRRLB-12227. Eaton and Karns (18) described a Pseudomonas sp. that could grow with ammeline as the sole nitrogen source, but was unable to grow on atrazine. Three genes that are involved in this pathway have been identified. The trzB gene has been identified as the ammeline aminohydrolase gene. The TrzB enzyme catalyzes the conversion of ammeline to ammelide. The gene which encodes the ammelide aminohydrolase enzyme has been identified as trzC. This protein converts ammelide to cyanuric acid. The cyanuric acid amidohydrolase (encoded by trzD) is responsible for the ring cleavage of cyanuric acid to form biuret (19). trzCD have also been identified in another strain of Pseudomonas

(strain 12228) as well as Klebsiella pneumoniae 99 (19). Further characterization of

TrzD has shown that it does not require divalent cations for proper function. The cyanuric acid amidohydrolase does not have a high affinity for cyanuric acid and has a high turnover rate. BLAST searches using the DNA and amino acid sequence for TrzD showed no significant homology to any known gene or protein. Because TrzD does not require divalent cations and does not have any sequence homology to AtzABC, it is likely that TrzD belongs to a different family of amidohydrolases (25).

1.4.4 The trz genes: Nocardiodes str. C190. Until 2001, atzA was only known gene capable of catalyzing the transformation of atrazine to hydroxyatrazine. Indeed, de

14 Souza et al. (15) had published a report showing that six isolates capable of mineralizing atrazine all had copies of the atzA gene that were 99% homologous to the atzA gene found in Pseudomonas. str. ADP. Due to the high sequence homology and distribution of the isolates, the authors speculated that the atzA gene had arisen from a single, recent evolutionary event and was very widespread (15). However, Mulbry et al. (36) recently reported the isolation of a Nocardiodes sp. that was able to completely mineralize atrazine, but did not have a copy of the atzA gene. Further investigation revealed the presence of a gene that was named trzN, which has the same chlorohydrolase activity as AtzA, but has no DNA sequence homology. Comparison of the proteins at the amino acid level shows about 30% identity to atzB and about 25% homology to trzA (36). A putative trzN gene has also been identified in an Arthrobacter strain that is also capable of mineralizing atrazine but does not possess the atzA gene.

1.4.5 The atz genes. The first gene identified in this group was atzA which was first isolated from Pseudomonas strain ADP, a novel isolate capable of complete mineralization of atrazine (13). The atzA gene encodes the atrazine chlorohydrolase enzyme which converts atrazine to hydroxyatrazine through a dechlorination reaction.

This protein was found to be 41% homologous to TrzA which catalyzes similar reactions in Pseudomonas sp. strain NRRLB-12227 (14). AtzA has been found to contain Fe(II) at the catalytic site in the enzyme (51). The next gene to be identified was atzB which was located approximately 8 kb downstream of the atzA gene in direct orientation. The hydroxyatrazine ethylaminohydrolase encoded by atzB was found to be involved in the removal of the n-ethylamino side chain from hydroxyatrazine to produce N-

15 isopropylammelide. This gene was found to be only 25% homologous to TrzA and had no significant homology to other genes involved in triazine metabolism (7). The discovery of a gene encoding an N-isopropylammelide isopropylaminohydrolase was reported shortly after atzB. This gene was identified as atzC and was located on the same plasmid as atzAB. The protein converts N-isopropylammelide to cyanuric acid and isopropylamine. What is interesting about this gene is its relative distance from atzAB

(34 and 25 kb, respectively) and the different G+C content. atzC has a G+C content of

44% as compared to 58% in atzA and 61% in atzB. This suggests that atzC was likely acquired from an organism with a different G+C content (33, 49). However, a comparison of the amino acid sequence of all three proteins reveals conserved H-X-H motifs common to an amidohydrolase protein superfamily which includes dihydroorotase, adenine deaminase, and several ureases (49). Further analysis of AtzC showed conservation of key residues of the N-terminal dihydroorotase signature pattern which is responsible for coordinating transition metals and activating the molecule of water responsible for the hydrolytic mechanism. Zinc was revealed to be the transition metal in AtzC by inductively coupled plasma emission spectroscopy (ICP) (57). Prior to the 2001 paper by Martinez et al. (33), TrzD had been the only cyanuric acid amidohydrolase found to be involved in atrazine metabolism. However, trzD gene probes failed to identify a TrzD homolog in Pseudomonas strain ADP. In 2001, a gene was identified which encoded AtzD, a cyanuric acid amidohydrolase with a 58% amino acid identity to TrzD. This protein was shown to have cyanuric acid amidohydrolase activity by the identification of biuret, a breakdown product of cyanuric acid, by HPLC analysis (33). Another open reading frame in the same plasmid was found to have

16 homology to a biuret amidohydrolase. This gene was named atzE and crude extracts of

E. coli cells overproducing this protein has been found to hydrolyze biuret to produce allophanate. The production of allophanate was surprising since it had been thought that biuret would be reduced to urea. However, incubation of crude cell extracts with urea failed to hydrolyze the urea. Based on these results, AtzE was identified as a biuret

+ amidohydrolase. AtzF, the final protein identified converts allophanate to CO2 and NH4

(33).

1.5: Statement of purpose

At the beginning of this study, most of the work in this field was focused on the aerobic fate of atrazine in agricultural sites. The transformation of atrazine in wetland sediments and anaerobic mineralization of atrazine had been poorly studied. Although many genes involved in atrazine metabolism have been identified, very few efforts have been made to characterize microbial populations and corresponding catabolic genes in situ in agricultural or wetland sites. If detection of a particular catabolic gene could be linked to environmental samples which show a high potential for mineralization, then future evaluation of sites could be conducted using genomic DNA extraction and PCR amplification with the corresponding target gene-specific primers. This approach could allow a site with high mineralization potential to be identified in a shorter time period (2-3 d compared to 28 d for a biometer incubation). In addition, the use of radioactive materials could be avoided – otherwise a major obstacle in industrial research settings.

The hypothesis of this research was that wetland and agricultural sites have the potential for atrazine mineralization. In addition, the genes involved in atrazine

17 mineralization would be present in these samples. To investigate this hypothesis, three objectives were formulated. The first objective was to determine the atrazine mineralization potential under aerobic and anaerobic conditions at three sites in Ohio.

The second objective was to determine the presence of selected catabolic genes involved in atrazine degradation at these sites. The final objective was to isolate and characterize microorganisms capable of atrazine mineralization.

18 CHAPTER 2

EVALUATION OF THE ATRAZINE MINERALIZATION POTENTIAL AT AN

AGRICUTURAL RESEARCH SITE BY SOIL BIOMETER STUDIES AND

PCR AMPLIFICATION OF CATABOLIC GENES

2.1 INTRODUCTION

Atrazine has been widely used in agricultural systems to kill broad leaf weeds in corn fields. The study site in this work, the Van Meter farm in Piketon, OH, has had atrazine applied to fields for more than thirty years. Previous studies at this site have characterized the mineralization potential of several field plots at this farm. Radosevich et al. (43) examined depth profiles from soils which received atrazine yearly and soils which did not receive a yearly atrazine application (43). The potential to mineralize [U-

14C-ring]-atrazine and [2-14C-ethyl]-atrazine were examined in history and non-history soils sampled at this site. The mineralization of [U-14C-ring]-atrazine decreased with

19 depth in the site with a history of atrazine application. In the non-history site, mineralization was observed only in surface soil samples. In contrast, the extent of [2-

14C-ethyl]-atrazine mineralized was lower than the observed extent in the [U-14C-ring]- atrazine experiments. This may be due to cellular assimilation of the [2-14C-ethyl]-

14 atrazine into the microbial biomass rather than release as CO2. Inoculation of the soil samples with atrazine-mineralizing Ralstonia basilensis increased the extent of mineralization in soils which did not otherwise mineralize atrazine. Inoculated soils showed rapid mineralization indicating that the unamended soils may lack indigenous atrazine-mineralizing organisms (43).

The effects of crop management practices on atrazine mineralization have also been examined at the Piketon, OH site (39). A plot with continuous corn crops and another plot with a corn-wheat-soybean rotation were selected for this study. Corn plots were plowed using conventional tilling methods, whereas soybeans crops were ridge tilled. Wheat plots were not tilled during planting. Atrazine was applied to corn crops only. A riparian zone was selected which did not receive atrazine directly. The continuous corn (CC) plot showed a high extent of mineralization (approx. 80%), indicating the presence of an active atrazine mineralizing community. For crop rotation

(CR) samples, a lag of approximately 14 d was common prior to active mineralization.

The extent of mineralization was 28-40% in these samples. The riparian zone (RZ) showed lower levels of mineralization, from a low of 4% to a high of 28% (39).

Although the riparian zone did not receive a direct application of atrazine, it may be subject to vernal flooding from the adjacent Scioto River, which may deposit atrazine and atrazine-degrading organisms into the riparian zone. The Scioto River is an

20 agricultural watershed upstream of the Piketon site. Mineralization in CR and RZ samples could be enhanced by preincubating the soils with atrazine for 30 d prior to addition of [U-14C-ring]-atrazine (40). This enhancement suggested that atrazine or its metabolites may serve as inducers of the atrazine mineralization genes. The addition of cyanuric acid to soil samples was also examined in efforts to enhance atrazine mineralization (40). Cyanuric acid was selected as a potential inducer because it is a central intermediate in the atrazine mineralization pathway. Samples were preincubated up to 138 days with cyanuric acid before determining the extent of atrazine mineralization. Cyanuric acid induced atrazine mineralization in CR soils, but failed to induce mineralization in CC and riparian soils (40). DNA was extracted from each of the soils to determine whether trzD, the gene encoding the cyanuric acid amidohydrolase, could be detected. A 2.0 kb fragment of the trzD gene was used as a probe in dot blot hybridizations of soil DNA samples. The hybridization method did not detect the presence of trzD in any of the samples, indicating that trzD was not present or was present at levels below the threshold of detection for dot blot hybridization.

The purpose of this study was to determine the fate of atrazine under anaerobic conditions at the CC, CR and riparian sites with various electron acceptor amendment treatments. In addition, detection of several of the genes involved in atrazine metabolism

(atzA, trzN, and trzD) by PCR amplification was attempted. Isolation and characterization of organisms involved in atrazine metabolism was attempted.

21 2.2 MATERIALS AND METHODS

2.2.1 Site description

The sites sampled were located in the Van Meter farm managed by the Piketon

Research and Extension Center in Piketon, OH (Figure 2.1). This site is used for agricultural research in conjunction with The Ohio State University. The first site is a no- till field which has been planted with corn continuously for more than 30 years (CC). The second agricultural field is a crop rotation of wheat, corn, and soybean (CR). Atrazine is applied yearly to CC and every third year with the corn crop at CR. The riparian zone

(RZ) is located at the western edge of the agricultural fields and is bordered by the

Scioto River which has flooded into the riparian zone in the past. The Scioto River serves as an agricultural watershed upstream of Piketon as well as for the Piketon area.

Table 2.1 lists the sampling dates and atrazine applications at each site.

2.2.2. Sampling

Four subsamples were taken from each site and mixed to form a composite sample.

The top 5-10 cm of soil were sampled for each subsample. For CC, the field was entered approximately 9 meters from the NW corner of the site. For CR, the start site was at the transect which crosses the field. For all sites, the first subsample was taken 9 m from the outer edge of the field and each subsequent sample was 9 m further into the field along the plant row. All samples were taken between the plants. RZ sites were cleared of vegetation prior to sampling. The site is located at 83° 2’ 00” W longitude and 39° 2’ 30”

N latitude.

Figure 2.1: Location of the field sites sampled at the Piketon Research and Extension

Center in Piketon, OH. CC = continuous corn, CR = crop rotation, and RZ = riparian zone

Sampling Samples taken Atrazine Amount of atrazine Field date application applied (kg ha-1) application date (m/d/y) 8/19/99 CC, CR, RZ 5/7/99 1.67 CC 4/6/00 CC, CR, RZ 5/5/00 1.67 CC, CR 6/7/00 CC, CR, RZ 5/5/00 1.67 CC, CR 9/19/00 CC, CR, RZ 5/5/00 1.67 CC, CR 6/1/01 CC, CR, RZ, 5/2/01 1.67 CC Scioto 8/9/01 CC, CR, RZ, 5/2/01 1.67 CC Scioto 6/10/02 CC, CR, RZ, 5/23/02 1.67 CC Scioto 7/22/02 Scioto 5/23/02 1.67 CC 9/29/02 CC, CR, RZ, 5/23/02 1.67 CC Scioto

Table 2.1: Field sampling dates and atrazine application information for the Piketon, OH site. The amount of atrazine applied is indicated in kg of active ingredient applied to each field.

Samples taken from the Scioto River were sampled with a 4 x 140 cm acrylic column. Surface water samples were collected as grab samples. Water samples were concentrated 200 fold by centrifugation at 10,000xg for 20 min in a Beckman J2-21 centrifuge (Beckman Coulter, Fullerton, CA). The pellets were re-suspended in the mineral salts solution described below for the mineralization studies. All samples were stored at 4°C before use.

2.2.3 Mineralization studies

Mineralization of [U-ring-14C]-atrazine under aerobic conditions was measured in biometers, which consisted of 60-ml serum bottles equipped with suspended 2-ml vials

14 to trap evolved CO2 (37) (Fig. 2.2). Each biometer received 5.0 g (dry weight) aliquots of soil samples or 5 ml of water. Each sample was tested in duplicate biometers except where noted. For each experiment two soil samples were autoclaved at 121°C for 20 min and used as a sterile control. Each biometer received 0.065 µmol [U-ring-14C]- atrazine (specific activity 1.54 mCi/mmol; Sigma-Aldrich Co, St. Louis, MO) with a total concentration of 2.8 mg/kg dry weight soils or l for water samples. Trace elements (per liter of 100X stock solution): MgSO4·7H2O, 50 mg; CaSO4, 200 mg; FeSO4·7H2O, 1 g;

MnSO4·H2O, 20 mg; CuSO4, 20 mg; ZnSO4·7H2O, 20 mg; CoSO4·7H2O, 10 mg;

NaMoO4·2H2O, 5 mg; H3BO3, 5 mg; nitrilotriacetate, 3 g) were also added (100 µl of

100X) to fulfill any trace metal requirements. For anaerobic biometers, 1 ml of 71.3 mM

KNO3, Fe(COOH)3, or Na2SO4 were added to provide an excess of electron acceptor beyond what was needed to fully mineralize all of the added atrazine. For samples with no additional electron acceptor, 1 ml of dH2O was added. Biometers were flushed with nitrogen for five minutes after biometer set-up and after each sampling.

14 Mineralization of atrazine was monitored as evolution of CO2, which was trapped in 1 ml of 0.5 M KOH. The KOH was collected and replaced at intervals. The alkaline trapping solution was mixed in 10 ml scintillation fluid (Scintiverse BD, Fisher Scientific,

Pittsburgh, PA) in a vial and counted in a scintillation counter. The counting efficiency

Figure 2.2: Disassembled and assembled biometers. Biometers consist of a 60 ml serum bottle with a vial suspended from a rubber stopper. The vial is filled with 0.5 M NaOH.

was determined using an external standard and was determined to be 97%.

Mineralization was monitored at 22±2°C for 28-38 d.

2.2.4 Data analysis

The data were analyzed by calculating the means and standards deviations from replicate biometers except where noted. Cumulative atrazine mineralization was calculated by adding the sample percentage of carbon dioxide evolved to the total from the preceding time course. Half-lives and rate constants of atrazine mineralization were

- determined by fitting the graphs to first-order rate function, expressed as P = Pmax (1-e kt 14 ), where P = observed amount of CO2 evolved (%), Pmax = the maximum extent of

-1 mineralization for that site, t = time (d), and k = rate constant (d ). Half-life (t1/2) values were calculated using their respective rate constants as t1/2 = ln2/k (22).

2.2.5 Mass balances

After the final sampling, the biometers were dried for approximately 24 h at 90°C.

The dried soil samples were crushed to a uniform particle size and 0.25 g samples (dry weight) were used for analysis. The samples were combusted in an oxidizer (Biological

Oxidizer OX-400, R. J. Harvey Instrument Corporation, Patterson, NJ) and the evolved

14 CO2 was trapped in scintillation fluid (Carbon-14 Cocktail, R. J. Harvey Instrument

Corporation). The cocktail was transferred to a vial for liquid scintillation counting. For mass balance estimates, the amount of radioactivity remaining in the sample was added

14 to the cumulative % CO2 from the mineralization studies.

2.2.6 DNA isolation

DNA was isolated from soil and water samples using the QIAquick Stool Sample Kit

(Qiagen, Valencia, CA). Before proceeding with step one of the protocol, 10 g samples of soil were added to 10 ml 0.1% sodium pyrophosphate and incubated at 22±2°C with shaking for 2-3 hours. The samples were then centrifuged at room temperature for 15 min at 800 revolutions/minute. The supernatant was removed into a new tube.

Approximately 1.7 ml of each sample was placed in a microfuge tube and centrifuged at

15,000 revolutions/minute for 10 minutes. The supernatant was poured off and the pellet was resuspended as per Qiagen instructions. Positive controls were isolated from plasmid constructs transformed into Escherichia coli DH5α strains (pMD4 for atzA and pJ204 for trzD) and HB101 (pWM221011 for trzN). Plasmids were isolated using the

Qiagen Minipreparation Kit.

Genomic DNA from pure cultures was isolated using a phenol/chloroform extraction technique. A single colony was transferred to 100 ml of atrazine minimal medium containing 10 mM K2HPO4, 5 mM NaH2PO4, 0.1 mM atrazine, 10 mM sucrose, 5 mM

MgCl2, and 1X mineral salts solution and grown for approximately 4 d at 22±2°C with shaking. Approximately 30 ml of cells were pelleted by centrifugation in a Beckman J2-

21 Centrifuge (Beckman Coulter, Fullerton, CA) at 12,000xg for 10 min. All centrifugation runs were performed at 4°C. The supernatant was poured off and the pellet was resuspended in 11.3 ml TE (10 mMTris-Cl, pH 8.0, 1 mM EDTA), 600 µl 10% SDS, and

3.5 ml 200 mM NaOH. The suspension was incubated at 22°C for 10 min. An equal volume of CHCl3:isoamyl alcohol (24:1) was added and the suspension mixed by inversion. The suspension was centrifuged for 15 min at 12,000xg and the aqueous

layer was removed to a new centrifuge tube. An equal volume of phenol:CHCl3:isoamyl alcohol (25:24:1) was added and the suspension was mixed by inversion. The suspension was centrifuged at 12,000xg for 15 min. The aqueous layer was transferred to a fresh tube and 2 ml of 5M NaCl was added. 0.6 volumes of isopropanol was added and the suspension was mixed gently by inversion. The suspension was centrifuged at

27,000xg for 10 min. The supernatant was poured off and approximately 25 ml of ice- cold 95% ethanol was added. The pellet was centrifuged for 10 minutes at 27,000xg.

The supernatant was poured off and the pellet was allowed to air dry. The pellet was resuspended overnight at 4°C in 100 µl of dH2O.

2.2.7 PCR amplification

Primers for atzA were based on the 528 bp internal sequence of atzA (13). The trzD485 primers were generated using a computer software package (GeneRunner, http://www.generunner.com) and were based on an internal 500 bp sequence of the gene, which was sequenced by Karns (25). The sequences for the primers used in this study are listed in Table 2.2. The PCR mixture (final volume: 50 µl) contained 5-7µl template DNA, 1X PCR buffer, 0.1 mM primers, 200 µM dNTPs, 2.25 mM MgCl2, 10 µl Q solution and 2.5 U HotStarTaq polymerase (Qiagen, Valencia, CA). The genes were amplified in a PTC-200 DNA engine thermocycler (MJ Research, Waltham, MA) using the following cycle: denaturation at 95°C for 15 min, 30 cycles of 94°C for 1.5 min, annealing temperature for 1 min, 72°C for 2 min, and a final extension step at 72°C for

10 min. The annealing temperature for each primer set is listed in Table 2.2. All annealing steps were step down cycles that started 5°C higher than the listed annealing

Primer Primer sequence (5'- 3') Annealing name temperature atzA528F CCATGTGAACCAGATCCT 55°C atzA528R TGAAGCGTCCACATTACC - atzA1F AGGTTGTATTGTGCGGAAGC 62°C atzA1R GTTCAGGGGGAAGAAAGCTC - trzD485F TCGTTCAGGTCAAGTGCC 58°C trzD485R ATCGTCCAGCATCGTGTG - trzD1F AGCGCAAGTTTTTCGAGTTC 60°C trzD1R CGCCGGAGACATAAACCAT - trzN420F CACCAGCACCTGTACGAAGG 55°C trzN420R CATTCGAACCATTCCAAACG -

Table 2.2: Oligonucleotide sequences and annealing temperatures for primers used for

PCR amplification.

temperature and decreased 1°C/cycle to the desired annealing temperature. Nested

PCR reactions were set up as described above except 5 µl of the previous PCR reaction was used as the template. 5 µl of the positive control and 10 µl of all other amplified samples were separated on a 1.0% agarose gel in TAE buffer. The DNA was visualized using a 10 mg/ml solution of ethidium bromide.

The PCR mixture to test for PCR inhibitors (final volume: 50 µl) contained 7 µl soil template DNA, 0.5 µl LC2 DNA (ribosomal DNA), 1X PCR buffer, 25 pmol primers, 200

µM dNTPs, 2.25 mM MgCl2, 10 µl Q solution and 2.5 U HotStarTaq polymerase (Qiagen,

Valencia, CA). The ribosomal gene was amplified in a PTC-200 DNA engine

thermocycler (MJ Research, Waltham, MA) using the following cycle: denaturation at

95°C for 15 min, 30 cycles of 94°C for 30 sec, 55°C for 30 sec, 72°C for 2 min, and a final extension step at 72°C for 7 min. A second round of PCR was performed using the previous reaction as a template. The samples were diluted 1:25 or 1:50 (total dilution over two reactions - 1:250 or 1:500) in the PCR reaction buffer and the PCR cycle was performed as above. The agarose gel was run as described above.

2.2.8 Most Probable Number (MPN) technique

The MPN technique was adapted from the protocol by Radosevich et al. (44).

The basal MPN medium consisted of (per liter): 0.4 g K2HPO4 and 22 mg atrazine. To the basal medium, 10 ml of sodium acetate or other carbon source, 10 ml MgSO4·7H2O, and 10 ml 100X mineral salts were added. The sodium acetate was prepared by adding

1.02 g of sodium acetate to 100 ml dH2O. The MgSO4 was prepared by added 5 g of

MgSO4·7H2O to 100 ml dH2O. The mineral salts solution consisted of 0.1 g FeCl3·6H2O,

0.1 CaCl2·2H2O, 0.001 MnCl2·4H2O, and 0.001 g ZnSO4·7H2O added to 100 ml dH2O and then filter sterilized. Five ml of medium were dispensed into 16x150 mm test tubes.

-1 -9 Samples were diluted 10 to 10 in 50 mM phosphate buffer (0.14 g/L KH2PO4 and 8.53 g/L K2HPO4). 1.0 ml of the last five dilutions was added to six replicate MPN tubes. One replicate from each dilution was autoclaved at 121°C for 20 minutes. This replicate served as a sterile control to determine the amount of atrazine sorbed to the soil during the incubation. The rack of tubes was weighed and incubated in the dark at 22±2°C for

30 d. At the end of this period, the rack was reweighed and lost water was replaced in equal volumes to each tube. A 1 ml sample from each tube was removed and filtered

through a 0.22 µm filter to remove particulate matter. 100 µl of the filtrate was added to

900 µl of HPLC mobile phase (65% acetonitrile, 35% dH2O). Samples were run on C18

5µ reverse phase Adsorbosphere column (Alltech Associates, Deerfield, IL). Pump flow rate was set at 1.0 ml/minute and the UV detector was reading at 220 nm. MPN counts were determined using a Microsoft Excel spreadsheet adapted from Briones and

Reichardt (8).

2.2.9 Enrichment for atrazine-mineralizing organisms

5 g of soil from the fields was added to 100 ml of atrazine minimal media (as described in the genomic DNA extraction) and incubated at 22±2°C with shaking.

Samples (1.0 ml) of the enrichments were removed every two weeks and the amount of atrazine remaining was determined by HPLC as described in the MPN technique. When the atrazine was depleted in the culture, 1 ml of the culture was transferred a fresh flask of atrazine minimal medium. The cultures were transferred approximately eight times.

One loopful of culture was transferred to a plate of atrazine minimal medium with 1.5 g/L agar. Colonies that grew on these plates were recultured several times to produce pure cultures. Once the cultures were determined to be pure, they were transferred to liquid atrazine minimal media cultures and tested for the ability to mineralize atrazine by

HPLC.

2.2.10 16S rDNA sequencing

Genomic DNA was isolated from a pure culture of P-C 3 as described in section 2.5.

A 20 µl sample of this DNA was run on a 0.6% low melting point agarose gel at 60 V for

approximately 1.5 hr. DNA that was 1-10 kb in size were cut from the gel and heated at

70°C for 20 minutes to inactivate DNAses. This DNA was amplified using the 8f and

U968r ribosomal primers (4). These products were cloned using the TOPO TA cloning system (Invitrogen, Carlsbad, CA). These clones were submitted to bi-directional sequencing with an ABI Prism model 377 sequencer (Perkin-Elmer Applied Biosystems,

Foster City, CA). The M13 universal forward and reverse primers which can bind to the pCR2.1 vector were used for sequencing.

2.2.11 FAME analysis

The P-C 3 isolate was identified using fatty acid methyl ester (FAME) analysis. Cultures were grown on TSA plates for 3 to 4 d at 27°C prior to harvest and extraction. Cultures were harvested and lipids were extracted and derivitized according to the standard protocol for cultures grown on solid media (MIDI, Inc., Newark, DE). Total cellular fatty acid methyl esters were analyzed by gas chromatography according to the protocols outlined by MIDI (50, 61). The FAME analysis was performed at the University of

Delaware, courtesy of Dr. Radosevich.

2.3 RESULTS AND DISCUSSION

2.3.1 Mineralization studies-Yearly data

2.3.1.1 1999 sampling year

At the continuous (CC) site, mineralization was measured both aerobically and anaerobically. The extent of aerobic mineralization was 102% indicating that these

biometers may have received more radioactivity than was determined for the total amount added, so these data were not included in Figure 2.3. Under anaerobic conditions, biometers with no additional electron acceptor had the highest extent of mineralization (56%) followed by sulfate, iron, and nitrate (Fig. 2.3). Some studies have shown that the addition of nitrate can inhibit atrazine mineralization (1, 21).

The crop rotation (CR) field in 1999 was planted with wheat and thus no atrazine was applied. The extent of aerobic mineralization in this soil was 58%, closely followed by anaerobic mineralization with sulfate as the additional electron acceptor (Fig. 2.4).

The relatively long lag period exhibited by all anaerobic biometers suggests that there is a shift in the microbial population that allows the mineralization of atrazine under anaerobic conditions. Such a shift may involve changes in gene expression or in the composition of the resident microbial population. Molecular genetic methods such as T-

RFLP (Terminal Restriction Fragment Length Polymorphisms) or microarrays could be used to evaluate such changes.

The addition of iron, nitrate, and no acceptor all had a similar, low extent of mineralization. At day 27, additional iron was added to the iron and no acceptor biometers and additional nitrate was added to the nitrate biometers. The additional electron acceptor did not increase the extent of mineralization for those biometers which already had electron acceptor added (iron and nitrate) but did increase the rate of mineralization in the no acceptor biometers which may indicate that anaerobic mineralization in these biometers was inhibited in the absence of an external electron acceptor (Fig. 2.5). The lack of an increase in the other samples indicated that electron

100 No acceptor 90 Iron Nitrate 80 Sulfate 70

50 evolved (%) 2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.3: Mineralization data for August 1999 for samples taken from the CC site.

Each point represents the mean of duplicate biometers. All samples were incubated under anaerobic conditions and amended with the indicated external electron acceptors.

100

90 No acceptor Iron 80 Nitrate Sulfate 70 Aerobic Sterile reference 60

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.4: Mineralization data for August 1999 for samples taken from the CR site.

Each point represents the mean of duplicate biometers. All samples were incubated under anaerobic conditions except the samples labeled aerobic.

100 No acceptor Iron 80 Nitrate

60 Evolved (%)

2 40 CO 14

0 10203040506070 Time (Days)

Figure 2.5: Anaerobic mineralization of selected August 1999 CR samples following additional electron acceptor amendment. No acceptor and iron samples received additional iron, and nitrate samples received more nitrate. The arrow indicates the time point at which the biometers were amended with additional electron acceptor. Each point represents the mean of duplicate biometers.

acceptor concentrations were not depleted and that the lower extent of mineralization was due to some other factor. The riparian zone only showed substantial mineralization under aerobic conditions (Fig. 2.6). Mineralization under anaerobic conditions was less than 1% for all riparian samples.

2.3.1.2 2000 sampling year

At the CC site, aerobic mineralization reached an extent of 71% (Fig. 2.7).

Mineralization under anaerobic conditions with no additional electron acceptor could not be determined due to the loss of one of the replicates early in the experiment. The addition of sulfate depressed mineralization as compared to 1999 (19% compared to

37%). Conversely, the addition of nitrate seemed to enhance mineralization in 2000

(extent 41%) compared to 1999 in which these samples had the lowest extent of mineralization (8%). Mineralization with the addition of iron increased somewhat over

1999. However, lag periods of at least 10 days were common in both sampling dates.

At the CR site, the extent of mineralization under aerobic conditions was 57%, almost identical to the extent in 1999 (Fig. 2.8). Although 2000 was a corn rotation for

CR, at the time of the April sampling, atrazine had not yet been applied. Therefore, the lack of an increase in the extent of mineralization is not surprising. As seen in the CC site, the addition of sulfate showed the lowest extent of mineralization (28% in 28 d) in contrast to the results from 1999 in which biometers with sulfate as the electron acceptor showed the highest extent of mineralization. The other electron acceptors clustered together with the extent of mineralization reaching approximately 44% in all cases. In the riparian zone, all samples showed a low extent of mineralization in the range of 22-29%

100 Aerobic 90 No acceptor Iron 80 Nitrate 70 Sulfate

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.6: Mineralization data for August 1999 for samples taken from the RZ site. Each point represents the mean of duplicate biometers. All samples were incubated under anaerobic conditions except the samples labeled aerobic.

100 Iron 90 Nitrate Sulfate 80 Aerobic 70

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.7: The mineralization data for CC samples taken in April, 2000. Points plotted

14 represent the mean cumulative CO2 evolved from duplicate biometers. Standard deviations may be smaller than the symbols. All samples were incubated under anaerobic conditions except those samples labeled aerobic.

100

90 No acceptor Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.8: The mineralization data for CR samples taken in April, 2000. Points plotted

(Fig. 2.9). The rate of aerobic mineralization decreased in comparison to 1999, but all anaerobic samples showed a higher extent of mineralization.

In May, CC samples showed an increase in the extent and rate of mineralization as compared to April (Fig. 2.10 and Table 2.3). Mineralization under aerobic and anaerobic conditions, with the exception of no added acceptor, all showed a similar extent of mineralization. Because some of the electron acceptors suppressed mineralization in the past, an experiment was designed to determine if addition of a similar amount of each electron acceptor would inhibit mineralization under aerobic conditions (Fig. 2.11). All samples showed approximately equal rates and extents of mineralization, which suggested that the depression of mineralization is due to some reason other than toxicity from an excess of electron acceptor.

Aerobic mineralization in CR samples increased tremendously over the April samples (Fig. 2.12). The extent of mineralization reached almost 100% in these samples while mineralization under anaerobic conditions with no additional electron acceptor also showed a large increase over April levels. In contrast, the extent of mineralization in the riparian zones was similar to that observed in April (Fig. 2.13). The lack of an increase in this sample may indicate an induction of the microbial populations in CC and CR due to the application of atrazine. If factors such as soil temperature, moisture and organic matter content were equally important, than an increase in riparian mineralization would be expected.

There was little change in the extent of mineralization in September CC samples under aerobic conditions as compared to May (Fig. 2.14). However, the extent of mineralization under anaerobic conditions with no electron acceptor increased over May

100 No acceptor 90 Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%)

2 40 CO

14 30

0 10203040 Time (Days)

Figure 2.9: The mineralization data for RZ samples taken in April, 2000. Points plotted

14 represent the mean cumulative CO2 evolved for duplicate biometers. Standard deviations may be smaller than the symbols. All samples were incubated under anaerobic conditions except those samples labeled aerobic.

100

60 Evolved (%)

2 40 CO 14 No acceptor 20 Iron Nitrate Sulfate Aerobic 0

0 5 10 15 20 25 30

Time (Days)

Figure 2.10: The extent of mineralization for CC samples incubated aerobically and

14 anaerobically. Each point represents the mean cumulative CO2 evolved for duplicate biometers. The standard deviations, if not shown, are smaller than the symbols. CC soils were sampled in June, 2000.

Sample Electron acceptor Incubation Date k (d-1) r2 t (d) site amendment conditions 1/2 CC None Anaerobic Apr-00 0.008 0.9554 88.85 CC Iron Anaerobic Apr-00 0.028 0.9276 24.75 CC Nitrate Anaerobic Apr-00 0.031 0.9842 22.35 CC Sulfate Anaerobic Apr-00 0.014 0.5908 49.50 CC None Aerobic Apr-00 0.095 0.9026 7.29 CR None Anaerobic Apr-00 0.033 0.9752 21.00 CR Iron Anaerobic Apr-00 0.034 0.9753 20.38 CR Nitrate Anaerobic Apr-00 0.036 0.9794 19.25 CR Sulfate Anaerobic Apr-00 0.028 0.8828 24.75 CR None Aerobic Apr-00 0.058 0.9803 11.95 RZ None Anaerobic Apr-00 0.014 0.9188 49.50 RZ Iron Anaerobic Apr-00 0.016 0.9535 43.31 RZ Nitrate Anaerobic Apr-00 0.015 0.9117 46.20 RZ Sulfate Anaerobic Apr-00 0.018 0.9531 38.50 RZ None Aerobic Apr-00 0.019 0.9158 36.47

CC None Aerobic Jun-00 0.169 0.9003 4.10 CC Iron Aerobic Jun-00 0.155 0.8940 4.47 CC Nitrate Aerobic Jun-00 0.155 0.9025 4.47 CC Sulfate Aerobic Jun-00 0.142 0.9076 4.88 CC None Aerobic Jun-00 0.143 0.7476 4.85 CC None Anaerobic Jun-00 0.033 0.8904 21.00 CC Iron Anaerobic Jun-00 0.086 0.9535 8.06 CC Nitrate Anaerobic Jun-00 0.061 0.9188 11.36 CC Sulfate Anaerobic Jun-00 0.084 0.9402 8.25 CR None Anaerobic Jun-00 0.073 0.9267 9.49 CR None Aerobic Jun-00 0.172 0.9433 4.03 RZ None Aerobic Jun-00 0.004 0.8017 173.25

CC None Anaerobic Sep-00 0.100 0.9463 6.93 CC None Aerobic Sep-00 0.202 0.9609 3.43 CR None Anaerobic Sep-00 0.039 0.8552 17.77 CR None Aerobic Sep-00 0.040 0.9173 17.33 RZ None Aerobic Sep-00 0.005 0.7635 138.60

Table 2.3: Kinetic parameters and half-lives for atrazine in soils sampled in 2000.

100

60 Evolved (%)

2 40 CO 14 No acceptor 20 Iron Nitrate Sulfate Aerobic 0

0 5 10 15 20 25 30 Time (Days)

Figure 2.11: Aerobic mineralization of June CC samples spiked with electron acceptor solutions at concentrations used in anaerobic biometers to test for toxicity. Points plotted

14 represent the mean cumulative CO2 evolved from duplicate biometers. Standard deviations may be smaller than the symbols.

100

60 Evolved (%) Evolved 2 40 CO 14

No acceptor Aerobic 0

0 5 10 15 20 25 30 Time (Days)

Figure 2.12: Mineralization data for CR samples obtained in June 2000. Samples were incubated aerobically or anaerobically with no additional electron acceptor. Points

14 plotted represent the mean cumulative CO2 evolved for duplicate biometers. Standard deviations, where not shown, are smaller than the symbols.

100

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.13: Mineralization data for RZ samples obtained in June 2000. Samples were

14 incubated aerobically. Points plotted represent the mean cumulative CO2 evolved for duplicate biometers. Standard deviations may be smaller than the symbols.

100 No acceptor 90 Aerobic 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.14: Mineralization data for CC samples obtained in September 2000. Samples were incubated aerobically or anaerobically with no additional electron acceptor. Points

14 plotted represent the mean cumulative CO2 evolved for duplicate biometers. Standard deviations, where not shown, are smaller than the symbols.

levels (83% compared to 51%). Although the extent of mineralization was similar in the aerobic and anaerobic samples, the rate of mineralization under aerobic conditions was double that of the anaerobic samples (Table 2.3).

The extent of mineralization in CR samples dropped to 64% in September (Fig.

2.15). The rate and extent of mineralization were very similar for the anaerobic and aerobic biometers. There was also a lag period of 3-7 days for both conditions which was longer than what was observed in May. This suggests that the activity of the population in this field had decreased somewhat after the initial burst of activity observed after atrazine application. The riparian zone samples also showed a low extent of mineralization, similar to what was observed in May. This suggested that the microbial population was not actively mineralizing atrazine in the riparian zone (Fig. 2.16).

2.3.1.3 2001 sampling year

The June samples taken for CC showed a slightly lower rate and extent of mineralization in these samples compared to the October samples (Fig. 2.17 and Table

2.4). These samples were taken approximately one month following atrazine application.

Because this soil receives an annual application of atrazine, it was expected that the extent of mineralization in these fields would be similar from year to year. Thus, the variation observed is not likely due to a lack of atrazine in the soils and may be due to other factors such as soil moisture and organic matter content.

The rate and extent of mineralization in CR samples was slightly lower than CC samples. In 2001, soybeans were planted at CR and no atrazine was applied. This may explain the slight decrease in the rate and extent of mineralization. As shown in Figure

100 No acceptor Aerobic 80

60 Evolved (%) Evolved

2 40 CO 14

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.15: Mineralization data for CR samples obtained in September 2000. Samples were incubated aerobically or anaerobically with no additional electron acceptor. Points

14 plotted represent the mean cumulative CO2 evolved for duplicate biometers. Standard deviations, where not shown, are smaller than the symbols.

100

50 Evolved (%) Evolved

2 40 CO

14 30

0 102030 Time (Days)

Figure 2.16: The mineralization data for RZ samples taken in September, 2000. Points

14 plotted represent the mean cumulative CO2 evolved for duplicate biometers. Standard deviations may be smaller than the symbols. The biometers were incubated under aerobic conditions.

100 CC 90 CR RZ 80 Sterile reference 70

50 Evolved (%) Evolved 2 40 CO

14 30

0 0 5 10 15 20 25 30 35 Time (Days)

Figure 2.17: Aerobic mineralization in Piketon soils sampled in June 2001. Points plotted

14 represent the mean cumulative CO2 evolved in duplicate biometers. Standard deviations may be smaller than the symbols.

-1 2 Sample site Incubation Date k (d ) r t1/2 (d) conditions CC Aerobic Jun-01 0.188 0.9632 3.69 CR Aerobic Jun-01 0.055 0.9330 12.60 RZ Aerobic Jun-01 0.035 0.9203 19.80 Scioto water Aerobic Jun-01 0.071 0.7984 9.76

CC Aerobic Aug-01 0.141 0.9651 4.91 CR Aerobic Aug-01 0.018 0.8756 38.50 RZ Aerobic Aug-01 0.012 0.8317 57.75 Scioto water Aerobic Aug-01 0.106 0.6906 6.54

Table 2.4: Kinetic data and calculated half-lives for 2001 Piketon samples.

2.17, riparian samples showed a large increase in the rate and extent of mineralization over the May 2000 sample (46%, k = 0.035 as compared to 17%, k = 0.005). The lag period was also considerably shorter. These data suggest that the riparian zone had been exposed to atrazine, possibly as airborne drift or flooding from the adjacent Scioto

River which has been considered a potential source of inocula for the riparian zone. The

Scioto River serves as an agricultural watershed upstream of the sampling site and atrazine concentrations have been detectable in the water, including samples taken during this study which showed concentrations of 2-4 ppb in 2002. Although the Scioto

River has been suspected as a source of atrazine-mineralizing microorganisms, this had

never been tested. Samples of Scioto River water that had been preconcentrated 200 fold showed an extent of mineralization of 52% (Fig. 2.18). These data indicate that the

Scioto River water was likely a source of atrazine mineralizing organisms in the riparian zone.

In August, the extent of mineralization in CC samples was slightly higher than

June (Fig. 2.19). However, the rate of mineralization was slightly slower with a longer half-life. This suggests that the population in this soil was less active, which may be due to a lower concentration of atrazine in the soil late in the growing season.

The CR samples dropped significantly in comparison to the June samples (Fig.

2.19). The lag period was also much longer than was observed in June (9 d compared to

3 d). The extent of mineralization in the riparian samples dropped by almost half in the

August samples. The rate of mineralization was only a third of that observed in June

(Fig. 2.19 and Table 2.4). The extent of mineralization in concentrated Scioto River samples increased to 89% compared to 52% in June (Fig. 2.20). This may be due to increased microbial activity, increased numbers of microorganisms, or a spike in suspended soils due to a storm event.

2.3.1.4 2002 sampling year

CC samples taken in June showed a similar extent of mineralization to samples analyzed in 2001 while the rate of mineralization decreased (Fig. 2.21 and Table 2.5).

Samples were not taken in the CR field in June to avoid damage to the soybean crop. In the riparian zone, the extent of mineralization had decreased to a low level (9%) with a

100

Evolved (%) Evolved 40 2

CO 30 14 20

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.18: Aerobic mineralization data for concentrated Scioto River samples collected

14 in June 2001. Points represent mean cumulative CO2 evolved in duplicate biometers.

Standard deviations, where not shown, are smaller than the symbols.

100 CC 90 CR RZ 80 CC SR 70

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.19: Mineralization data for Piketon soil samples collected in August 2001. All samples were incubated under aerobic conditions. Points represent mean cumulative

14 CO2 evolved in duplicate biometers. Standard deviations may be smaller than the symbols.

100

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 2.20: Mineralization data for concentrated Scioto river samples collected in

August 2001. The biometers were incubated under aerobic conditions. Points represent

14 mean cumulative CO2 evolved in duplicate biometers. Standard deviations may be smaller than the symbols.

100 CC 90 RZ 80 Sterile reference

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.21: Mineralization data for Piketon soil samples collected in June 2002. All samples were incubated under aerobic conditions. Points represent mean cumulative

14 CO2 evolved in duplicate biometers. Standard deviations may be smaller than the symbols.

Incubation Sample Date k (d-1) r2 t (d) conditions 1/2 CC Aerobic Jun-02 0.109 0.9452 6.36 RZ Aerobic Jun-02 0.002 0.8532 346.50 Scioto sediment Aerobic Jun-02 0.160 0.9343 4.33 Scioto water Aerobic Jun-02 0.107 0.8767 6.48

CC Aerobic Oct-02 0.151 0.9436 4.59 CR Aerobic Oct-02 0.018 0.8511 38.50 RZ Aerobic Oct-02 0.002 0.9503 346.50 Scioto water Aerobic Oct-02 0.098 0.8564 7.07

Table 2.5: Kinetic data and half-lives calculated from mineralization data for 2002

Piketon samples.

lag period of at least 26 days (Fig. 2.21). It is unclear why the activity in the soil would be so low in comparison to the observed extent of mineralization in 2001.

Samples of sediment from the Scioto River bed were obtained and tested for atrazine mineralization. Despite the high sand content, the samples mineralized 86% of the added atrazine (Fig. 2.22). This indicates the presence of an active community in the

Scioto River, which furthers confirms the hypothesis that the river is a source of atrazine mineralizing organisms that would inoculate the riparian zone in times of flood. Water samples were also extremely active, mineralizing 80% of the added atrazine (Fig. 2.22).

100 Sediment 90 Water 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.22: Aerobic mineralization data for Scioto river water and sediment samples

14 collected in June 2002. Points represent mean cumulative CO2 evolved in duplicate biometers. Standard deviations may be smaller than the symbols.

The extent of mineralization in the July Scioto River water samples was comparable to the extent observed in June (Fig. 2.23). However, the lag period was approximately 2 days in June and had increased to 5 days in July. This may be due to a lower concentration of atrazine in the river water. Atrazine concentrations measure in

June showed an atrazine concentration of 3.76 ppb which had dropped to 1.77 ppb in

July. However, due to the nature of the ELISA used to obtain these numbers, it is possible that the concentrations of atrazine in either sample may be artificially high due to antibody cross-reactivity with metabolites or other members of the s-triazine family.

In September, CC samples showed an extent of mineralization similar to the observed mineralization in June (Fig. 2.24). However, the rate of mineralization was higher than observed in June and were similar to values obtained in 2001 (Tables 2.4 and 2.5). The samples taken from the CR field mineralized 39% of the atrazine, similar to values observed in 2001 (35%). The riparian samples had an extremely low extent of mineralization (4%), which was the lowest observed value for any year (Fig. 2.25).

Scioto river water showed a similar extent of mineralization to the other 2002 samples.

The lag period and rate of mineralization were also similar (Fig 2.25 and Table 2.5).

2.3.2 Mineralization data - Summaries

Figure 2.26 shows a summary of CC data collected each June for 3 years. This graph shows no significant difference in the extent of mineralization. The rate of mineralization appears to be slightly faster in 2000 than for the other two years. The similarity in the kinetics of atrazine mineralization in the samples is not unusual for a field that has received an application of atrazine annually for more than 30 years. Several

Evolved (%) 30 2 CO

14 20

0102030 Time (Days)

Figure 2.23: Aerobic mineralization data for Scioto river water samples collected in July

14 2002. Points represent mean cumulative CO2 evolved in duplicate biometers. Standard deviations may be smaller than the symbols.

100 CC CR RZ 80 Sterile reference

60 Evolved (%)

2 40 CO 14

0 5 10 15 20 25 30 Time (Days)

Figure 2.24: Mineralization data for Piketon soil samples collected in September 2002.

All samples were incubated under aerobic conditions. Points plotted represent the mean

14 cumulative CO2 for duplicate biometers. Standard deviations may be smaller than the symbols.

100

60 Evolved (%)

2 40 CO 14

0 5 10 15 20 25 30 Time (Days)

Figure 2.25: Mineralization data for Scioto river water samples collected in September

2002. All samples were incubated under aerobic conditions. Points plotted represent the

14 mean cumulative CO2 for duplicate biometers. Standard deviations may be smaller than the symbols.

100

50 Evolved (%) Evolved

2 40 CO

14 30

20 06/00 10 06/01 06/02 0

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.26: Summary of samples taken over a 3 year period at the CC site. All samples

14 were incubated aerobically. Points plotted represent the mean cumulative CO2 for duplicate biometers. Standard deviations may be smaller than the symbols.

studies have shown an increased level of mineralization activity in soils with a history of atrazine application (39, 67).

The results from the CR field do not show a similar history effect (Fig. 2.27).

Samples taken in 1999 and 2000 show the highest extent of mineralization. However, atrazine was only applied in 2000. It is unclear why the 1999 samples showed such a high rate of mineralization. The field was planted with wheat in 1999 and soybean in

1998 and had not received an application of atrazine for at least two years. The RZ samples also showed a high extent of mineralization in 1999.One possible explanation for the high extent of mineralization in CR and RZ may be aerial drift of atrazine that year, perhaps in higher concentrations than normal. Another possible explanation is the spatial variability In the soil samples. It is possible that certain areas sampled in the CR field had very active populations which mineralized atrazine at unusually fast rates in these years. However, a previous study of this same CR site indicated that there was low spatial variability in a transect of the CR field which included 12 samples taken every

20 m (39). This suggests that spatial variability in atrazine-mineralizing microorganisms in CR is low and does not explain the anomalously high extent of mineralization observed in 1999 and 2000. In contrast, the CR samples taken in 2001 and 2002 showed a lower extent of mineralization. This may indicate a difference in the climatic conditions or soil organic matter or nutrient content. Moreover, crop plant roots may contribute to differences in the rhizosphere and bulk soil although the literature on this topic is sparse.

100 Wheat 8/99 90 Corn 10/00 Soybean 08/01 80 Soybean 09/02

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.27: Summary of samples taken over a 4 year period at the CR site. All samples were incubated aerobically. The crops and sample dates for each year are indicated on

14 the key. Points plotted represent the mean cumulative CO2 for duplicate biometers.

Standard deviations may be smaller than the symbols.

In the riparian zone, the extent of mineralization in samples taken in 1999 and

2001 were four times higher than the extent from samples taken in 2000 and 2002 (Fig.

2.28). Possible explanations may include a fresh inoculation of organisms from the

Scioto River or increased drift of atrazine from the surrounding agricultural fields.

The effect of atrazine application is addressed in Figure 2.29. Figure 2.29A shows CC samples taken approximately one month prior to atrazine application and one month after application. Samples taken in April show an extent of mineralization of 71%

The extent is slightly higher in samples taken after atrazine application (84%). The rate of mineralization in May samples is much faster than in the April samples (k=0.143 d-1 and 0.095 d-1, respectively) while the lack of a lag period indicates that both samples were actively mineralizing atrazine. Again, these results are expected in a soil with a history of atrazine application since atrazine residue persists over time in soil and microbial populations are constantly exposed to atrazine. Figure 2.29B shows atrazine mineralization in CR samples collected during the same sampling period. In samples taken prior to atrazine application, the extent of mineralization was 57%. After atrazine application, the extent of mineralization almost doubled. The rate of mineralization tripled from k=0.058 d-1 in April to 0.172 d-1 in May. The apparent stimulation of atrazine mineralization is more dramatic in the crop rotation samples and may reflect the lack of biologically available atrazine after one or more years without an atrazine application.

2.3.3: Catabolic marker genes

Genomic DNA was extracted from both soil and water from 2002 Piketon samples. No

PCR amplification products were obtained using primers specific to atzA and trzD (Figs.

100 8/99 90 06/00 06/01 80 06/02 70

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 2.28: Summary of samples taken over a 4 year period at the riparian zone. All samples were incubated aerobically. The sample dates for each year are indicated on

14 the key. Points plotted represent the mean cumulative CO2 for duplicate biometers.

Standard deviations may be smaller than the symbols.

70 100 AB

50 Evolved (%) 2 40 CO

14 30

20 71 10 05/00 06/00 0

0 5 10 15 20 25 0 5 10 15 20 25 30

Time (Days)

Figure 2.29: Aerobic mineralization of samples taken pre- and post-atrazine application in 2000. (A) CC (B) CR. Points

14 plotted represent the mean cumulative CO2 for duplicate biometers. Standard deviations may be smaller than the

symbols.

2.30 and 2.31). These samples were tested for inhibitors by adding an equal amount of ribosomal DNA to each sample and then amplifying using ribosomal-specific primers. As indicated in Figure 2.32, samples needed to be diluted 1:250 or 1:500 to eliminate the inhibitors. However, 1:500 dilution was not sufficient to eliminate inhibitors .The extracted DNA sample was faintly brown, suggesting the presence of humic matter as possible inhibitor. Further testing of these samples is needed to confirm the presence or absence of atzA and trzD. The samples could be diluted prior to PCR to eliminate the inhibitory effect. It may also be possible to remove some of the inhibitors using polyvinylpyrrolidone or gel purification. However, further studies on elimination of the

PCR inhibitors were not pursued with Piketon samples because of the emphasis on

Defiance samples.

2.3.4: Isolate characterization

One bacterial culture was enriched from Piketon soils which had the ability to mineralize atrazine completely as shown by HPLC analysis. The organism was subsequently isolated as a pure culture by repeated streaks for isolation on solid atrazine minimal medium. The colonies on atrazine minimal agar were 1 mm in diameter, white, entire, and convex after one week of incubation. The bacterium was gram-positive and could assimilate glucose, mannitol, maltose, gluconate, malate, and citrate as carbon sources. It was also weakly positive for assimilation of mannose, N- acetyl glucosamine and phenyl-acetate. The culture was not capable of reducing nitrate.

Figure 2.33 indicates that this organism does not contain the atzA gene while Figure

2.0 kb

1.0 kb

528 bp 0.5 kb

0.1 kb

Figure 2.30: Agarose gel electrophoresis of PCR products amplified with primers specific for the atzA gene. Lane 1: pMD4 (positive control); Lane 2: No DNA (negative control);

Lane 3: MW markers; Lane 4: CC; Lane 5: CR; Lane 6: RZ; Lane 7: Scioto River.

2.0 kb

1.0 kb

0.5 kb 485 bp

0.1 kb

Figure 2.31: Agarose gel electrophoresis of PCR products amplified with primers specific for the trzD gene. Lane 1: pJK204 (positive control); Lane 2: No DNA (negative control);

Lane 3: CC; Lane 4: CR; Lane 5: RZ; Lane 6: Scioto River; Lane 7: MW markers.

2.0 kb

1.0 kb

0.5 kb

0.1 kb

Figure 2.32: Agarose gel electrophoresis of PCR products amplified with primers specific for 16S ribosomal genes. Ribosomal DNA was added to Piketon DNA samples, diluted

1:250 or 1:500 and amplified with ribosomal DNA specific primers to check for inhibitors.

Lane 1: LC2 (positive control); Lane 2: No DNA (Negative control); Lane 3: MW markers;

Lane 4: CC; Lane 5: CR; Lane 6: RZ; Lane 7: Scioto River.

2.0 kb

1.0 kb

528 bp 0.5 kb

0.1 kb

Figure 2.33: Agarose gel electrophoresis of PCR products amplified with primers specific for the atzA gene. Lane 1: P-C 3; Lane 2: pMD4 (positive control); Lane 3: No DNA

(negative control); Lane 4: MW markers.

2.34 shows that trzD is absent. However, primers specific for trzN amplified a PCR product with a size consistent with trzN suggesting that isolate P-C 3 contains this gene

(Fig. 2.35). The organism was identified as an Arthrobacter sp. by both 16S rDNA sequencing and FAME analysis. The 16s rDNA sequencing indicated 99% homology to

Arthrobacter aurescens (621/627 bp). Arthrobacter ilicis and A. aurescens were the closest matches to this isolate as determined by FAME analysis.

2.3.5: Most Probable Number analysis

A most probable number (MPN) technique was used to determine the concentration of atrazine mineralizing organisms in soil samples. Previous studies have shown that the

MPN for a particular sample can be determined (24, 67). However, the current consensus is that these estimates are likely under-representing the number of atrazine- mineralizing organisms present in soil samples. The standard method uses acetate as a carbon source. Because acetate may not be easily metabolized, it was hypothesized that the use of other carbon sources might improve the number of organisms recovered.

Table 2.6 shows the MPN estimate for CC soils using various carbon sources.

As indicated in Table 2.6, tubes incubated with acetate as the added carbon source had an MPN estimate of 2x101-1x102 cells/g soil dry weight which is low for a soil which actively mineralizes atrazine. Carbon sources such as glucose and sucrose showed lower MPN estimates than acetate suggesting that these more readily metabolized substrates were not ideal for this purpose. Nutrient broth showed an estimate similar to that of acetate. The only C sources which gave a higher MPN

2.0 kb

1.0 kb 1.0 kb

0.5 kb

0.1 kb

Figure 2.34: Agarose gel electrophoresis of PCR products amplified with primers specific for the trzD gene. Lane 1: P-C 3; Lane 2: pJK204 (positive control); Lane 3: No DNA

(negative control); Lane 4: MW markers.

2.0 kb

1.0 kb

0.5 kb 420 bp

0.1 kb

Figure 2.35: Agarose gel electrophoresis of PCR products amplified with primers specific for the trzN gene. Lane 1: pWM221011 (positive control); Lane 2: No DNA (negative control); Lane 3: MW markers; Lane 4: P-C 3.

Sample Carbon source MPN (cells/g soil dry wgt.)

10% Loss 25% Loss 50% Loss CC acetate 1.1x102 4.9x101 2.3x101 CC glucose 3.9x101 3.2x101 2.8x101 CC nutrient broth 1.0x102 3.1x101 3.1x101 CC sucrose 7.7x100 4.4x100 2.0x100 CC soil extract 4.1x103 9.1x102 1.1x102 CC atrazine 1.3x104 2.5x102 2.1x102

Table 2.6: MPN estimates for CC soils incubated with various carbon sources. The 10% loss indicates the MPN estimate for tubes which lost 10% of the added atrazine compared to the sterile controls as measured by HPLC. Similar analyses were performed for 25% and 50% loss.

estimate were soil extract and atrazine. The soil extract might be an interesting alternative to standard carbon sources. By providing an extract of the nutrients from the original environment, it may be possible to increase the recovery of these organisms providing an improved MPN estimate of the existing population. However, none of the carbon sources seem to provide an accurate estimate of the number of atrazine degrading/mineralizing organisms present, based on the extent of mineralization observed from this soil. However, no one has attempted to correlate the amount of atrazine mineralized by a pure culture to the MPN estimate for those cells. It is possible that a high extent of mineralization could be obtained from a small number of cells.

2.4 Summary of results

CC soils showed a consistently high extent of mineralization which indicated that the population of atrazine mineralizing organisms is active and is maintained from year to year in the soil. In CR soils, the population is less active, except in years where atrazine is applied to the fields. When atrazine is applied, the rate and extent of mineralization increases dramatically. In RZ samples, the extent of mineralization is generally low although occasional spikes in atrazine mineralization occur, presumably due to a fresh influx of atrazine or atrazine-mineralizing organisms, either from airborne drift or flooding from the Scioto River. Samples of sediment and water from the Scioto

River show an active mineralizing population which supports the theory that the river is the source of mineralizing organism in the riparian zone which is periodically flooded by the Scioto River. PCR amplification of some of the genes was unsuccessful due to the presence of inhibitors. Dilution or removal of the inhibitors will be required before an assessment of the genes present in these samples can be made. An atrazine mineralizing organism isolated from CC soil was identified as a member of the

Arthrobacter genus of gram positive organisms. PCR amplification of genomic DNA isolated from this organism showed a gene product consistent with the trzN gene.

However, trzD and atzA were not amplified from this organism. MPN analysis of these soils indicated that the number of atrazine degrading organisms were low and seemed inconsistent with the extent of mineralization observed. Different C sources added to the

MPN medium did not seem to increase the detection of atrazine degrading organisms.

CHAPTER 3

ATRAZINE MINERALIZATION POTENTIAL IN TWO WETLANDS

3.1 INTRODUCTION

Atrazine (2-chloro-4- (ethylamino)-6-(isopropylamino)-s-triazine) is a relatively common contaminant in groundwater and sometimes exceeds the maximum contaminant level of 3 µg/l established for drinking water. Surface waters with agricultural watersheds have also been reported to contain detectable levels of atrazine.

The major non-point sources of atrazine in water are subsurface runoff and field tile drainage in agricultural sites. Because atrazine is a regulated compound in drinking water, its attenuation in ground and surface water is an important aspect of the environmental fate of this chemical. Elucidation of abiotic and biological degradative pathways of atrazine in the environment is also crucial in view of its endocrine disruptor effects (23). In soils, the most important mechanism for the attenuation of atrazine involves microbial degradation and mineralization. In surface waters and wetlands,

the transformations of atrazine are poorly understood although these systems receive agrochemicals through surface runoff and drainage.

To date, there is no evidence that native microbial populations in natural or constructed wetlands can actively and substantially mineralize atrazine. Ro and Chung

(45) monitored atrazine biotransformation in spiked wetland sediments. The samples were taken from a diked wetland constructed to treat sugar mill wastewater. The initial concentration of atrazine, 10 mg/l, decreased to <10 µg/l within 10 weeks of aerobic incubation, and subsequent spikes of atrazine were depleted in three weeks. Atrazine concentrations were measured by HPLC and the extent of mineralization could not be established from the biodegradation data. Kao et al. reported atrazine attenuation in the water column and sediments of a natural wetland system that received agricultural runoff following a storm event (26). Parallel mesocosm experiments with wetland sediments indicated accelerated biodegradation of atrazine only in the presence of an external carbon source (sucrose). Mineralization potential was not established (26). Wetland mesocosm studies have demonstrated the biodegradation of atrazine in water columns, with losses of about 70% as determined by gas chromatography, but the extent of mineralization and unavailability due to sorption to sediments remain unknown (34).

A study using sediment and groundwater samples from a freshwater wetland showed that approximately 4.5% of atrazine was mineralized under aerobic conditions and less than 2% under anaerobic conditions after 68 weeks of incubation (30).

DeLaune et al. (12) reported that atrazine was relatively persistent in anaerobic sediments of a swamp-forest wetland that received runoff from adjacent atrazine-treated agricultural fields, but the biodegradation could be enhanced upon introducing aerobic

conditions. Mineralization of atrazine was not investigated in the study. Runes et al. (47) used bioaugmentation with an atrazine-spilled soil to enhance an otherwise slow mineralization of atrazine in sediment microcosm and rhizosome mesocosm samples from a constructed wetland receiving irrigation runoff from a container nursery.

There are several known genes that encode the proteins in the atrazine degradative pathway. The most commonly studied is atzA, which encodes atrazine chlorohydrolase.

A survey of five geographically distinct atrazine mineralizing isolates showed that each isolate contained an atzA gene ≥99% identical to the gene found in the original host,

Pseudomonas ADP (15). Shapir et al. (55) tested soil samples from several agricultural fields that had different atrazine treatment histories. Regardless of previous application history, most samples contained organisms that could mineralize atrazine to a certain extent and the atzA gene could be amplified from these soils (55). Shapir et al. (56) amplified a PCR product with atzA primers from DNA extracted from sand/gravel lysimeters that had been amended with secondary wastewater effluent or partially composted sludge. The data suggested that there was a positive association between the presence of atzA and atrazine mineralization, even if the extent was only 1%.

Another gene of interest in this work is trzD, which encodes cyanuric acid amidohydrolase that catalyzes the ring cleavage of cyanuric acid. This compound is believed to be the last aromatic intermediate in the degradative pathway, but to date the gene and the enzyme has been characterized only in a melamine-degrading

Pseudomonas sp. (25). Data based on hybridization signals indicated that, in agricultural soil samples, the gene trzD was not dominant and its detection did not reflect the potential mineralization of atrazine (40). However, molecular approaches such as PCR

amplification of atzA, trzD, and other genes encoding enzymes of the atrazine degradative pathways have not been applied in the study of atrazine mineralization in wetland systems.

In view of the lack of lack of knowledge on atrazine mineralization in wetlands, the purpose of this study was to evaluate the kinetics of atrazine mineralization in water and sediments samples retrieved from a constructed wetland. It was hypothesized that genes such as atzA and trzD from degradative pathways are associated with active mineralization of atrazine in native microbial populations. This relationship may be consistent only in circumstances that involve the presence of atrazine or metabolites in the wetland and may be analogous to the atrazine history effect described for agricultural soils (40, 67).

The wetland, Olentangy River Wetland Research Park of The Ohio State University

(http://kh465a.ag.ohio-state.edu/ORW.html), receives water from the Olentangy River, which has an upstream agricultural watershed. Detectable levels of atrazine, up to 13

µg/l, have been periodically detected in the Olentangy River and in the wetland water column in the past (http://www.epa.state.oh.us/ddagw/pestsbw2.pdf). The atzA and trzD genes from the mineralization pathways were targeted for PCR amplification and

Southern blotting using DNA extracted from the wetland samples. A natural wetland,

Cedar Bog, fed by water from springs was also selected for this work. This wetland, a nature preserve in Ohio, receives no surface runoff from the agricultural fields in the vicinity.

3.2 MATERIALS AND METHODS

3.2.1 Sampling

Sediment and water samples were retrieved from the sites near the inflow and outflow of the experimental wetland 1 at Olentangy River Wetland Research Park,

Columbus, OH (Fig. 3.1). This site is a constructed wetland that receives Olentangy

River water through the inflow and feeds back to the river via the outflow. The Olentangy

River has an agricultural watershed upstream of the wetland and receives non point source run-off from these fields. The site is located at 83° 1’ 81” W longitude and 40° 1’

59” N latitude and was constructed in 1993. The hydraulic retention time was between 4 and 5 d in 1999. The retention time in the wetland varies with the flow velocity of the

Olentangy River and can be as long as three weeks during periods of low flow of the river. The chemical, physical, and biological characteristics of the Olentangy River

Wetland Research Park have been summarized in several publications and annual reports (http://kh465a.ag.ohio-state.edu/ORW.html).

Olentangy wetland sediments and overlying water were sampled with a 4 x 140 cm acrylic column. Sediment samples were pooled or sectioned to measure the potential for atrazine mineralization. Three replicate sediment samples were mixed to form a composite. Individual sediment cores were removed by plunging from the column sampler and then sectioned into three approximately equal parts (about 5 cm each).

Composite samples were taken in April 2000 and August 1999 and 2000. Sectioned sediment samples were taken in April and August 2000. All sediment samples were taken under water-saturated conditions. Water content varied between samples with the

A B

Figure 3.1: Sampling sites at the (A) Olentangy River Wetland Research Park (OSU wetland) and (B) Cedar Bog. The boardwalks that provide access to the sites at the OSU wetland are also indicated. The inflow marks the site where water is pumped into the cell from the Olentangy River and the outflow marks the pipe that allows water to return to the river. Sampling sites are indicated with open squares.

inlet composite sample having a water content of 65% while the outlet composite was

34%. Inlet sections had water contents of 63% (0-5 cm), 47% (5-10 cm), and 51% (10-

15 cm). The outlet section water contents were 50%, 35% and 26% for the same sections. The variability in the water content reflects sediment compaction and clay content. Further chemical or mineralogical analyses were not with the scope of this study.

Wetland water samples were taken in August 2000 and June 2002. Olentangy River samples were taken in October 2001 and June 2002. Surface water samples were collected as grab samples. Initial measurements of atrazine mineralization showed very low activity (<1%). Subsequently, water samples were concentrated 200 fold by centrifugation at 10000xg for 20 min. The pellets were re-suspended in the mineral salts solution for mineralization experiments. No effort was made to standardize the concentrated samples with respect to suspended solids which varied with hydraulic flow conditions and rainfall. All sediment and water samples were stored at 4°C before use.

The Cedar Bog site (Figure 3.1) is an undisturbed alkaline fen in Urbana, OH, fed by springs. Fens are characterized by the accumulation of peat which receives water from surrounding mineral soils. The alkalinity at Cedar Bog is due to Ca,Mg- bicarbonates, causing carbonate precipitation around the springs and downstream from the springs. The site is located at 83° 47.5’ W longitude and 40° 3’ N latitude. This wetland was sampled at two forested sites. For Cedar Bog site 1, the samples were collected from a riparian site about 30 m away from a cornfield. Samples for Cedar Bog site 2 were collected along the banks of a small ditch draining into the bog. The adjacent upstream cornfield had been treated annually with atrazine for at least the past 10 years,

but there is no direct access of surface runoff to the Cedar Bog sampling sites. Five subsamples were taken at each site for a composite in May and June 2000. The top 15 cm of soil was sampled each time. Neither sampling site in the Cedar Bog has received any direct application of atrazine in the past.

3.2.2 Mineralization studies

Mineralization of [U-ring-14C]-atrazine under aerobic conditions was measured in biometers, which consisted of 60-ml serum bottles equipped with suspended 2-ml vials

14 to trap evolved CO2 (39). Each biometer received 5.5 g (wet weight) aliquots of sediment samples or 5 ml of water. Each sample was tested in duplicate biometers. Two inlet composite sediment samples were autoclaved at 121°C for 20 min and used as a sterile control. Each biometer received 0.065 µmol [U-ring-14C]-atrazine (specific activity

1.54 mCi/mmol; Sigma-Aldrich Co, St. Louis, MO) with a total concentration of 2.55 mg atrazine/kg wet weight sediment and 2.8 mg/l for water samples. Trace elements (per liter of 100X stock solution): MgSO4·7H2O, 50 mg; CaSO4, 200 mg; FeSO4·7H2O, 1 g;

MnSO4·H2O, 20 mg; CuSO4, 20 mg; ZnSO4·7H·2O, 20 mg; CoSO4·7H2O, 10 mg;

NaMoO4·2H2O, 5 mg; H3BO3, 5 mg; nitrilotriacetate, 3 g) were also added (100 µl of

100X) to fulfill any trace metal requirements. For the August 1999 anaerobic biometers,

1 ml of 10 mM KNO3, 6 mM Na2SO4, or 1 mM Fe(COOH)3 was added to duplicate biometers. For all other sample dates, 1 ml of 71.3 mM KNO3, Fe(COOH)3, or Na2SO4 were added to provide an excess of electron acceptor beyond what was needed to fully mineralize all of the added atrazine. For samples with no additional electron acceptor, 1 ml of dH2O was added. Biometers were flushed with nitrogen for five minutes after

biometer set-up and after each sampling. Mineralization of atrazine was monitored as

14 evolution of CO2, which was trapped in 1 ml of 0.5 M KOH. The KOH was collected and replaced at intervals. The alkaline trapping solution was mixed in 10 ml scintillation fluid (Scintiverse BD, Fisher Scientific, Pittsburgh, PA) in a vial and counted in a scintillation counter. The counting efficiency was determined using an external standard and was determined to be 97%. Mineralization was monitored for up to 38 d.

3.2.3 Data analysis

The data were analyzed by calculating the means and standards deviations from the replicate biometers. Cumulative atrazine mineralization was calculated by adding the sample percentage of carbon dioxide evolved to the total from the preceding time course. A first-order rate expression was used to compare the time courses of atrazine mineralization between different samples. First-order kinetic parameters are commonly used to assess environmental biodegradation of pesticides, based on the premise that degradation or mineralization is limited by the concentration of the pesticide (59). Thus, the kinetic parameters determined in this study can be compared with those previously reported for various soil and other agricultural and environmental conditions. Half-lives

-kt and rate constants were determined by fitting the data to P = Pmax (1-e ), where P =

14 observed amount of CO2 evolved (%), Pmax = the maximum extent of mineralization,

-1 and t = time (d), and k = rate constant (d ). Half-life (t1/2) values were calculated using

-1 their respective rate constants as t1/2 = ln2 k . While atrazine mineralization could not be adequately described with non-linear first-order rate expression in samples, where the

14 evolution of CO2 remained relatively low, additional kinetic analysis to seek the best- fitting rate expressions were beyond the scope of this study.

3.2.4 Mass balances

After the final sampling, the biometers were dried for approximately 24 h at 90°C.

The dried soil samples were crushed to a uniform particle size and 0.25 g samples (dry weight) were used for analysis. The samples were combusted in an oxidizer (Biological

Oxidizer OX-400, R. J. Harvey Instrument Corporation, Patterson, NJ) and the evolved

14 CO2 was trapped in scintillation fluid (Carbon-14 Cocktail, R. J. Harvey Instrument

Corporation). The cocktail was transferred to a vial for liquid scintillation counting. For mass balance estimates, the amount of radioactivity remaining in the sample was added

14 to the cumulative % CO2 from the mineralization studies.

3.2.5 DNA isolation and PCR amplification

DNA was extracted from 0.5 g sectioned sediment samples using the alternative lysis method in the UltraClean Soil DNA Kit (MoBio Laboratories, Solana Beach, CA).

DNA was stored in 10 mM Tris pH 8 at -20oC. For the water samples, DNA was isolated using the QIAquick Stool Sample Kit (Qiagen, Valencia, CA). Approximately 1.7 ml of each concentrated water sample was placed in a microfuge tube and centrifuged at

15,000 revolutions/minute for 10 minutes. The supernatant was poured off and the pellet was resuspended as per Qiagen instructions. Positive controls were isolated from plasmid constructs transformed into Escherichia coli DHα strains (pMD4 for atzA and pJ204 for trzD) or HB101 (pWM221011 for trzN). The Surzycki et al. (63) protocol or the

Qiagen Minipreparation Kit was used for plasmid DNA isolation.

For the sediment samples, primers for atzA were based on the 528 bp internal sequence of atzA (13). The trzD primers were generated using a computer software package (GeneRunner, http://www.generunner.com) and were based on an internal 500 bp sequence of the gene, which was sequenced by Karns (25). See Table 2.2 for atzA528 and trzD485 primer sequences. The PCR mixture (final volume: 50 µl) contained 0.2 µg template DNA, 1X PCR buffer (30 mM Tricine, 0.1% gelatin, 1.0%

Thesit (polyoxyethylene 9 lauryl ether), and 5 mM β-mercaptoethanol), 1.0 mM primers,

0.55 mM dNTPs, 0.2 mg BSA, 3.0 mM MgCl2, and 3-5 U Taq DNA polymerase. The atzA gene was amplified in a PTC-200 DNA engine thermocycler (MJ Research,

Waltham, MA) using the following cycle: denaturation at 92°C for 5 min, 40 cycles of

92°C for 1.5 min, 55°C for 1 min, 72°C for 2 min, and a final extension step at 72°C for 5 min. The trzD gene was amplified with similar cycles with the exception of the extension step which was performed at 58°C. 10 µl samples of all PCR reactions were separated on a 1.0% agarose gel in TAE buffer. The bands were visualized by soaking the gel for one hour in a 10-4 dilution of SYBR Green 1 DNA stain (Molecular Probes, Eugene, OR) in TAE.

For water samples, nested PCR was performed using the atzA primers listed in

Table 2.2 while the trzD1 primers were used in two sequential PCR reactions. The sequences for the primers used in this study are listed in Table 2.2. The PCR mixture

(final volume: 50 µl) contained 5-7µl template DNA, 1X PCR buffer, 0.1 mM primers, 200

µM dNTPs, 2.25 mM MgCl2, 10 µl Q solution and 2.5 U HotStarTaq polymerase (Qiagen,

Valencia, CA). The genes were amplified in a PTC-200 DNA engine thermocycler (MJ

Research, Waltham, MA) using the following cycle: denaturation at 95°C for 15 min, 30 cycles of 94°C for 1.5 min, annealing temperature for 1 min, 72°C for 2 min, and a final extension step at 72°C for 10 min. The annealing temperature for each primer set is listed in Table 2.2. All atzA annealing reactions were step down cycles that started 5°C higher than the listed annealing temperature and decreased 1°C/cycle to the desired annealing temperature. The nested and sequential PCR reactions were set up as described above except 5 µl of the previous PCR reaction was used as the template. 5

µl of the positive control and 10 µl of all other amplified samples were separated on a

1.0% agarose gels in TAE buffer. The DNA was visualized using a 10 mg/ml solution of ethidium bromide.

The detection limit for atzA and trzD in sediment samples was determined to be approximately 100 copies per PCR reaction using plasmid DNA as the template.

However, the detection limits for atzA and trzD when amplified from sediment DNA samples cannot be defined because of unknown and perhaps variable efficiency of DNA extraction.

3.2.6 Southern hybridization analysis

The DNA probes used in Southern blot hybridizations were prepared by PCR amplification of plasmids pMD4 and pJK204 containing the cloned atzA and trzD genes, respectively. The amplified fragments were subsequently purified using a PCR gene clean kit (Qiagen, Valencia, CA). Approximately 50 ng of the fragment DNA was boiled for 5 min followed by a 5-min incubation on ice to denature the double stranded templates. DNA probes were labeled with fluorescein-dUTP using the Gene Images

Random-prime kit (Amersham Pharmacia Biotech, Piscataway, NJ). PCR products were separated by agarose gel electrophoresis and were transferred to Hybond-N+ membranes. Hybridization and all washes were performed at 58°C. Hybridization signals were detected using an anti-fluorescein antibody conjugated with alkaline phosphatase

(CDP-Star detection kit, Amersham Pharmacia Biotech).

3.3 RESULTS AND DISCUSSION

3.3.1 Mineralization data

The initial survey of the Olentangy wetland sediments under aerobic and anaerobic conditions is shown in Figures 3.2 (inlet) and 3.3 (outlet). The extent of mineralization was low under aerobic conditions with the extent reaching 7% in the inlet and 9% in the outlet after 23 d. The extent of mineralization was even lower under anaerobic conditions with only 3-4% of the added atrazine mineralized. Because the incubation period was relatively short, samples taken in April 2000 were incubated for a longer period (38 d).

As shown in Figure 3.4, the extent of mineralization after 38 d of incubation was approximately 43% at the inlet and 47% at the outlet under aerobic conditions.

14 Autoclaved sediment samples showed no mineralization activity ( CO2 evolution <1%).

Figure 3.5 shows the extent of anaerobic mineralization in the inlet composite sample.

The extent of mineralization was approximately 20% in all cases. This may indicate inhibition of mineralization with the addition of electron acceptors. Interestingly, the outlet composite sample showed an extent of mineralization similar to aerobic levels in the

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 Time (Days)

Figure 3.2: Mineralization data for August, 1999 inlet composite samples under aerobic

14 and anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added for duplicate biometers. Standard deviations may be smaller than the symbols.

100 No acceptor 90 Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 Time (Days)

Figure 3.3: Mineralization data for August, 1999 outlet composite samples under aerobic

100 Olentangy Inlet Olentangy Outlet Sterile Reference 80 Cedar Bog 1 Cedar Bog 2

60 Evolved (%)

2 40 CO 14

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.4: Mineralization data from the April, 2000 Olentangy wetland composite samples and Cedar Bog sediment samples. Standard deviations may be smaller than the points.

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.5: Mineralization data for April, 200 in inlet composite samples under aerobic

14 and anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. Standard deviations may be smaller than the symbols.

absence of an external electron acceptor (Fig. 3.6). All other electron acceptor biometers had an extent of mineralization in the 14-20% range. The lower extent of mineralization with the addition of electron acceptor may be a further indication that the acceptors are depressing the mineralization capabilities of the microbial population. This initial

14 screening of CO2 evolution from the ring-labeled atrazine indicated that an atrazine- mineralizing microbial community was present in both segments of the Olentangy wetland.

In general, atrazine mineralization appeared to approach a sustained linear phase.

However, the rate constant and half-life for the composite inlet sample (April, 2000) indicated a reasonable first-order fit (r2>0.988) suggesting that the rate of mineralization was accelerating into a nonlinear phase. This linear phase may be analogous to an extended lag period preceding accelerated mineralization.

The sediment zone closest to the water interface (0-5 cm) showed consistently higher levels and faster aerobic mineralization than the underlying zones at both the inlet and outlet (Fig. 3.7). These data indicated differences in the distribution of atrazine- mineralizing organisms by depth, with the highest activity located at the sediment-water interface. The surface of the sediment is the initial sink for atrazine in the inflow river water. It is noteworthy that the composite inlet sediment sample yielded a lower extent of mineralization as compared to the individual sediment fractions within a similar period of incubation. This difference may reflect variability such as clay content and cell density and distribution among the core samples retrieved for the composite.

14 Figures 3.7A and 3.7B show the aerobic CO2 evolution data for the inlet sections sampled in April and August, respectively, and Figures 3.7C and 3.7D show the

100 No acceptor 90 Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.6: Mineralization data for April, 200 in outlet composite samples under aerobic

14 and anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. Standard deviations may be smaller than the symbols.

100

100 0-5 cm A B 5-10 cm 10-15 cm 80

Evolved (%) C D 2 CO

14 80

0 0 5 10 15 20 25 30 35 0 5 10 15 20 25 30 35 Time (Days)

Figure 3.7: Mineralization data for the Olentangy wetland sediment sections. (A) April,

2000 Inlet, (B) August, 2000 Inlet, (C) April, 2000 Outlet, (D) August, 2000 Outlet.

101

corresponding data for the outlet. At the inlet, the extent of atrazine mineralization in the

0-5 and 5-10 cm zones were approximately 15% lower in August than April at day 28.

However, the rate of mineralization was faster in the August samples. At the outlet, almost no activity was present at depths of 5-15 cm in August, unlike the April samples

(39% and 21% compared to 1%).

Under anaerobic conditions, the 0-5 and 10-15 cm inlet sections sampled in April have similar results (Figs. 3.8 and 3.10). The addition of electron acceptors appears to enhance the capability for mineralization as compared to anaerobic conditions lacking an external electron acceptor. This is in direct contrast to the patterns observed in the composite samples in which the acceptors appeared to inhibit mineralization. This may reflect differences in the composition of the composite sediments as compared to the sections. In the 5-10 cm section, the addition or lack of electron acceptors did not seem to have an effect on mineralization (Fig. 3.9). In the outlet, there does not seem to be a clear pattern to the electron acceptor usage (Figs. 3.11, 3.12, 3.13). In all cases, the extent of mineralization is the lowest when nitrate is the added electron acceptor. In the

0-5 cm section, biometers with no added acceptor and those with added iron appeared to have a higher extent of mineralization than the aerobic biometers. Firm conclusions cannot be made, however, since these results reflect a single biometer for each treatment.

In August, inlet biometers incubated under anaerobic conditions with no external electron acceptor show an extent of mineralization slightly lower than under aerobic conditions (Figs. 3.7B and 3.14). The outlet samples showed no mineralization under anaerobic conditions (Fig. 3.15). Although this is a contrast to the aerobic biometers

102

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.8: April 2000 mineralization data for 0-5 cm inlet section under aerobic and

14 anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent a single biometer for each treatment.

103

100 No acceptor 90 Iron Sulfate 80 Aerobic 70

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.9: April 2000 mineralization data for 5-10 cm inlet section under aerobic and

14 anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent a single biometer for each treatment.

104

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.10: April 2000 mineralization data for 10-15 cm inlet section under aerobic and

14 anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent a single biometer for each treatment.

105

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.11: April 2000 mineralization data for 0-5 cm outlet section under aerobic and

14 anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent a single biometer for each treatment.

106

100 No acceptor 90 Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.12: April 2000 mineralization data for 5-10 outlet section under aerobic and

14 anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent a single biometer for each treatment.

107

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 3.13: April 2000 mineralization data for 10-15 outlet section under aerobic and

14 anaerobic conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent a single biometer for each treatment.

108

100 0-5 cm 90 5-10 cm 10-15 cm 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 3.14: August 2000 mineralization data for inlet sections under anaerobic

14 conditions. Points plotted represent the mean cumulative % CO2 evolved as a percentage of the total radioactivity added. The data represent duplicate biometers for each treatment. Standard deviations may be smaller than the symbols.

109

100 0-5 cm 90 5-10 cm 10-15 cm 80

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 3.15: August 2000 mineralization data for outlet sections under anaerobic

110

sampled in August 2000, the biometers lacking electron acceptor also had the lowest extent of mineralization in April.

Table 3.1 lists kinetic parameters of atrazine mineralization derived from the first- order rate expression. Half-lives were in the order of 5 to 7 d in the most active samples.

Kinetic parameters were not calculated for samples where atrazine mineralization was low and linear over the experimental time course.

There are complex reasons underlying these differences in the kinetics and extent of mineralization in April and August. During this period, changes took place, for example, in ambient temperature, hydraulic flow rate, suspended solids, plant and microbial community diversity, and sediment organic carbon content. The role of such complex factors in influencing atrazine mineralization has yet to be determined.

Mass balance estimates accounted for 85-105% of the 14C initially added in these experiments. These mass balances revealed no major experimental errors or

14 unaccounted losses of CO2 that would have skewed the calculation of the relative

14 % CO2.

Because the activity was so high in the sediment sections, the potential of wetland water samples was also studied. Atrazine, a pre-emergent herbicide, is normally applied to agricultural soils in Ohio at the end of April or beginning of May. Atrazine would largely be washed from the fields during rain events in the summer. In river water, atrazine is mostly associated with suspended solids which include soil particles from runoff and microorganisms. Biometer experiments with August samples of wetland water column showed no mineralization activity, However, when water samples were first concentrated

111

Electron acceptor Incubation -1 2 Sample site Date k (d ) r t(d) amendment conditions 1/2 Inlet- None Anaerobic Apr-00 0.0085 0.5633 81.53 composite Inlet- Iron Anaerobic Apr-00 0.0079 0.3538 87.72 composite Inlet- Nitrate Anaerobic Apr-00 0.0079 0.3538 87.72 composite Inlet- Sulfate Anaerobic Apr-00 0.0091 0.5942 76.15 composite Inlet- None Aerobic Apr-00 0.0166 0.9883 41.70 composite Outlet- None Anaerobic Apr-00 0.0125 0.8937 55.44 composite Outlet- Iron Anaerobic Apr-00 0.0083 0.8302 83.49 composite Outlet- Nitrate Anaerobic Apr-00 0.0070 0.8761 99.00 composite Outlet- Sulfate Anaerobic Apr-00 0.0098 0.9290 70.71 composite Outlet- None Aerobic Apr-00 0.0137 0.8973 50.60 composite Inlet, 10-15 cm Iron Anaerobic Apr-00 0.0176 0.8909 39.38 Inlet, 10-15 cm Nitrate Anaerobic Apr-00 0.0218 0.8858 31.79 Inlet, 10-15 cm Sulfate Anaerobic Apr-00 0.0245 0.8937 28.29 Inlet, 10-15 cm None Aerobic Apr-00 0.0208 0.9320 33.40 Outlet, 0-5 cm None Anaerobic Apr-00 0.0111 0.8427 62.43 Outlet, 0-5 cm Iron Anaerobic Apr-00 0.0109 0.8364 63.58 Outlet, 0-5 cm Nitrate Anaerobic Apr-00 0.0057 0.6133 121.58 Outlet, 0-5 cm Sulfate Anaerobic Apr-00 0.0078 0.1248 88.85 Outlet, 0-5 cm None Aerobic Apr-00 0.0166 0.8938 41.90 Outlet, 5-10 None Anaerobic Apr-00 0.0099 0.8296 70.00 cm Outlet, 5-10 Iron Anaerobic Apr-00 0.0087 0.2775 79.66 cm Outlet, 5-10 Nitrate Anaerobic Apr-00 0.0042 0.2631 165.00 cm

Continued

Table 3.1: Kinetic parameters and half-lives for atrazine mineralization in OSU wetland and Cedar Bog samples obtained in 2000.

112

Table 3.1 continued

Outlet, 5-10 Sulfate Anaerobic Apr-00 0.0075 0.1824 92.40 cm Outlet, 5-10 None Aerobic Apr-00 0.0142 0.9556 48.80 cm Outlet, 10-15 None Anaerobic Apr-00 0.0096 0.9038 72.19 cm Outlet, 10-15 Iron Anaerobic Apr-00 0.0095 0.8560 72.95 cm Outlet, 10-15 Nitrate Anaerobic Apr-00 0.0044 0.5933 157.50 cm Outlet, 10-15 Sulfate Anaerobic Apr-00 0.0073 0.7717 94.93 cm Outlet, 10-15 None Aerobic Apr-00 0.0083 0.8777 83.70 cm

Cedar Bog 1 None Aerobic May-00 0.0053 0.0000 131.00 Cedar Bog 2 None Aerobic May-00 0.0045 0.0000 154.00

Inlet, 0-5 cm August Anaerobic Aug-00 0.0288 0.7020 24.06 Inlet, 0-5 cm August Aerobic Aug-00 0.1458 0.8554 4.60 Inlet, 5-10 cm None Anaerobic Aug-00 0.0338 0.8469 20.50 Inlet, 5-10 cm None Aerobic Aug-00 0.1044 0.8614 6.30 Inlet, 10-15 cm None Anaerobic Aug-00 0.0252 0.7600 27.50 Inlet, 10-15 cm None Aerobic Aug-00 0.0673 0.8771 10.30 Outlet, 0-5 cm None Anaerobic Aug-00 0.0006 0.9735 1155.00 Outlet, 0-5 cm None Aerobic Aug-00 0.0233 0.8703 29.70 Outlet, 5-10 None Anaerobic Aug-00 0.0006 0.9735 1155.00 cm Outlet, 5-10 None Aerobic Aug-00 0.0005 0.9832 1303.00 cm Outlet, 10-15 None Anaerobic Aug-00 0.0006 0.9735 1155.00 cm Outlet, 10-15 None Aerobic Aug-00 0.0004 0.9286 1734.00 cm Inlet-Water None Aerobic Aug-00 0.0389 0.5888 17.80 Outlet-Water None Aerobic Aug-00 0.0002 0.0990 3084.00

113

14 (200 fold) before CO2 evolution measurements in biometers, atrazine mineralization was readily detected in the suspended solids in the inlet water (Fig. 3.16A).

The Olentangy River was grab sampled about 10 m upstream and 50 m downstream of the wetland inlet and outlet, respectively. The biometer data showed active mineralization in concentrated water samples, and there were no differences in the mineralization activity between the two river sample sites (Fig 3.16B). In comparison, the rates of mineralization differ within the wetland, being faster at the inlet segment (Fig.

3.16A). Fresh water samples obtained in June 2002 show a similar pattern. Figure 3.17A shows the extent of mineralization for concentrated wetland water samples. The inlet shows a high extent of mineralization at 93% while the outlet shows an extent similar to sterile controls. In the Olentangy River, there appears to be no difference in the rate or extent of mineralization (Fig. 3.17B). The observed differences suggest that the capacity for mineralization in the wetland decreases with distance from the inlet. Unlike the river, the Olentangy wetland has low flow conditions (for the 200 m distance between the inlet and the outlet), and sometimes virtual stagnation, which plays a part in the sedimentation of atrazine-mineralizing microorganisms entering the wetland with influent river water.

The mineralization of atrazine was low in Cedar Bog samples (Figure 3.2). When fitted into the first-order rate expression, the time course data for atrazine mineralization were fitted into the first order rate expression but the corresponding r2 values indicated a poor fit (Table 3.1). No further kinetic analysis was undertaken due to a low extent of mineralization (≤13%).

114

100 Inlet A Upstream B Outlet Downstream 80

Evolved (%) 40 2 CO 14 20 115

0 5 10 15 20 25 30 0 5 10 15 20 25 30 35

Time (Days)

Figure 3.16: (A) Atrazine mineralization from concentrated wetland water samples taken in August 2000. Both duplicates

of the inlet water reached a similar Pmax value, but one had a 2-d longer lag period before the onset of mineralization. (B)

Atrazine mineralization for concentrated Olentangy River water. Samples were taken in October 2001. Standard

deviations, if not shown, were smaller than the symbols.

100 AB

60 Evolved (%)

2 40

CO Inlet Upstream 14 Outlet Downstream 20 116

0 5 10 15 20 25 0 5 10 15 20 25 30 35 Time (Days)

Figure 3.17: (A) Atrazine mineralization from concentrated wetland water samples. (B) Atrazine mineralization for

concentrated Olentangy River water. Samples were taken in June 2002. Standard deviations, if not shown, were smaller

than the symbols.

No PCR products were obtained with DNA from the Cedar Bog samples using primers designed for atzA and trzD. Southern hybridization of the PCR gels failed to detect any signals when hybridized with PCR products of authentic atzA and trzD (data courtesy of Kevin A. Wheeler). Since the assay system used in this study was based on

14 CO2 evolution rather than a decrease in the concentration of atrazine, it is possible that atrazine degradation exceeded mineralization in Cedar Bog samples. The lack of PCR products and Southern hybridization signals does not, however, support this possibility, although the possibility that the lack of signal was caused by the presence of inhibitors can not be excluded.

3.3.2 Catabolic gene detection

The PCR amplification and Southern blot hybridization data for the OSU wetland sediment and Cedar Bog samples was kindly provided by Mr. Kevin Wheeler. The atzA gene was not detected in DNA from the Olentangy wetland sediment samples using

PCR with atzA specific primers. However, positive signals were detected when PCR products were resolved by agarose gel electrophoresis and subsequently analyzed by

Southern hybridization with the 523 bp PCR product of atzA (data not shown). These data suggested that atzA was amplified from DNA but the product was present at such low levels that it could not be visualized on the gel. Further attempts were not made to enhance amplification of atzA by varying PCR conditions or by nested PCR. In contrast, the trzD PCR product (500 bp) was detected in all sections by PCR amplification as well as by Southern hybridization (Fig. 3.18).

117

Figure 3.18: (A) PCR products of DNA isolated from wetland samples, amplified with the trzD primers. (B) Southern hybridization of the PCR products with a trzD probe. All DNA samples from the Olentangy wetland sediments are from the sampling in April, 2000.

The Cedar Bog samples were taken in May and June 2000. Lane 1, 100 bp ladder; lane

2, pJK204 (positive control); lane 3, Inlet 10-15 cm; lane 4, Inlet 5-10 cm; lane 5, Inlet 0-

5 cm; lane 6, Outlet 10-15 cm; lane 7, Outlet 5-10 cm; lane 8, Outlet 0-5 cm; lane 9,

Cedar Bog site 1 sampled in May; lane 10, CB1 site sampled in June; lane 11, CB2 site sampled in May; lane 12, Cedar Bog site 2 sampled in June; lane 13, No DNA (negative control); lane 14, pJK204 (positive control). This figure was kindly provided by Mr. Kevin

Wheeler.

118

Water samples were also tested for the presence of these genes. Figure 3.19 shows the PCR products amplified with primers specific to atzA. This may indicate that atzA is not a prominent gene in this environment. In contrast, trzD was amplified from all wetland water samples except the inlet (Fig. 3.20). The presence of multiple bands in this gel is probably due to the sequential PCR reactions performed. It is likely that the amount of nonspecific primer binding was increased by the use of sequential reactions, causing the amplification of the extra bands. This could be avoided in the future by using the more stringent step-down PCR as well as nested primers.

Detection of the atzA gene in the Olentangy wetland sediment samples was ambiguous because its presence could only be confirmed by Southern hybridization.

While the data suggest that Southern hybridization as detected by alkaline phosphatase was more sensitive in detecting PCR products than the SYBR Green 1 used in PCR gels, the actual detection level of Southern hybridization is unknown. The lack of detection of PCR products may be explained by differences in the relative sensitivity of the two techniques. Southern hybridization only requires about 80% homology between the probe and the target DNA sequence to allow binding and detection by the probe. In contrast, the shorter sequence of the PCR primers does not tolerate a high mismatch between sequences. The lack of PCR products may, therefore, be explained a lower level of sequence homology between the primer sequences and the genes present in the samples, which did not compromise Southern probe binding due to the higher tolerance for mismatches.

For trzD, there was good agreement with the detection of the PCR product and its

Southern hybridization signal (Fig. 3.18). The atzA gene encodes a chlorohydrolase

119

2.0 kb

1.0 kb

528 bp 0.5 kb

0.1 kb

Figure 3.19: Agarose gel electrophoresis of PCR products amplified with primers specific for the atzA gene. Lane 1: pMD4 (positive control); Lane 2: No DNA (negative control);

Lane 3: Upstream; Lane 4: Downstream; Lane 5: Inlet; Lane 6: Outlet; Lane 7: MW markers.

120

2.0 kb

1.0 kb 1.0 kb

0.5 kb

0.1 kb

Figure 3.20: Agarose gel electrophoresis of PCR products amplified with primers specific for the trzD gene. Lane 1: pJK204 (positive control); Lane 2: No DNA (negative control);

Lane 3: MW markers; Lane 4: Upstream; Lane 5: Downstream; Lane 6: Inlet; Lane 7:

Outlet.

121

at the beginning of the upper pathway. The trzD gene is involved in the lower pathway and encodes the cyanuric acid amidohydrolase that catalyzes the ring cleavage of the heterocyclic s-atrazine. The PCR product of trzD was detected in DNA in all OSU wetland sediment and most water samples examined (Fig. 3.18A). Because trzD was more prevalent than atzA, it is plausible that trzD and atzA represent genes of different pathways of atrazine metabolism in native bacterial communities. These results indicate that atzA, as originally characterized in Pseudomonas ADP, may not be the dominant chlorohydrolase gene in this wetland system. The recently identified trzN, a triazine hydrolase gene from Nocardiodes soil isolate, is supportive of this conclusion (36). The corresponding amino acid sequence showed insignificant homology to AtzA and the trzN gene could not be amplified using atzA-specific primers (36, 65).

3.4 Summary of results

Aerobic mineralization in inlet and outlet composite samples was approximately equal. However, mineralization in the sections was generally higher in the inlet sections.

In general, the 0-5 cm sections were the most active sediments and the 10-15 cm sections were the least active. Anaerobic mineralization was generally lower than aerobic mineralization levels. The Olentangy River receives agricultural runoff and contains low µg/l levels of atrazine. In this work, the river water was shown to contain atrazine-mineralizing microorganisms, believed to originate from the runoff of the upstream agricultural watershed. The water from the river feeds the Olentangy wetland cell, where the mineralization activity was found to be more pronounced near the inlet

122

rather than the outlet. Thus, the mineralization of atrazine in the wetland can be attributed to the actions of microorganisms introduced from the river inflow as well as by microbial populations already established in the sediment zones. The atzA gene was not amplified from any of the sediment or water samples tested, but could be detected by

Southern blot hybridization. However, trzD was amplified from nearly all sediment and water samples. Cedar Bog samples showed a low extent of mineralization and does not appear to have the atzA or trzD genes present, although inhibition of the PCR reactions by soil components can not be excluded without further tests.

123

CHAPTER 4

DETERMINERATION OF ATRAZINE MINERALIZATION POTENTIAL

USING SOIL AND DNA DETECTION METHODS IN A WETLAND-

RESERVOIR SUBIRRIGATION SYSTEM

4.1 INTRODUCTION

One of the major causes of non-point source pollution in water is surface runoff from agricultural fields. Wetlands have been used to remove pollutants such as nitrogen and phosphorus from runoff. However, using wetlands to remediate herbicides has not been well investigated. Moore et al. (34) reported that a series of wetland mesocosms designed to remediate atrazine-containing runoff removed 34-70% of the atrazine within

35 days. Atrazine was introduced into the wetland mesocosms during simulated storm events. Loss of atrazine in the wetland mesocosms was monitored over time by HPLC.

Although mineralizing activity cannot be directly inferred from these data, the amount of atrazine extracted from plant and sediment samples was below the level of detection.

124

from the water column was likely due to biotic transformation. Two studies involving a mountainous wetland, which receives atrazine-containing runoff from surrounding agricultural areas, indicated that atrazine was removed from the water column both aerobically and anaerobically (26, 27). These studies determined that atrazine transformation was slow in the absence of an added carbon source or in sediments with a low organic matter content. Furthermore, anaerobic conditions increased the extent of atrazine transformation, perhaps due to reductive dechlorination. Larsen et al. (30) reported that atrazine was not mineralized either aerobically or anaerobically in sediments sampled from a groundwater-fed wetland. This finding was somewhat surprising since the source of the water feeding the wetland was from an agricultural catchment area.

The nursery industry is a large and growing source of pesticide- and fertilizer- contaminated water that should be treated before it can be released. Runes et al. determined that sediments sampled from a constructed wetland used to treat nursery irrigation water did not mineralize atrazine (extent <3%) (47). When samples of soils taken from an atrazine spill site were added, the extent of mineralization increased to 25-

30% of the added 14C-ethyl-atrazine. In a further study involving a constructed wetland associated with a nursery, it was determined that 16-24% of the 14C-ethyl-atrazine added was transformed within 7 days (48). Atrazine was introduced into the wetland cells in runoff associated with plant irrigation. The extent of atrazine transformation was related to the runoff frequency and intensity. In light of these studies, the ability of wetlands to mineralize atrazine seems to be erratic.

125

Because wetlands may be important in remediating non-point source pollution in agricultural drainage, a wetland site was selected that was designed to treat agricultural drainage. This site, located in Defiance, Ohio, was constructed using a wetland-reservoir subirrigation system model. In this model, surface runoff from the agricultural fields as well as subsurface tile drainage is routed to the wetland for treatment. The water is maintained in the wetland at a constant depth. If a storm event occurs to raise the water level, the excess water is moved to the connected reservoir for storage. The water contained in the reservoir can then be used to irrigate the field through a subirrigation system (3). Studies of this wetland have shown that it can successfully remove sediments and phosphorus from subirrigation waters, but no effort has been made to characterize the potential for pesticide removal (28). In this study, the mineralization potential in the wetland and reservoir as well as the surrounding agricultural fields was evaluated. DNA was isolated from each of the sites and tested for several genes involved in atrazine mineralization. Isolation of organisms with the potential to mineralize atrazine was also attempted.

4.2 Materials and Methods

4.2.1 Site description

The Defiance site was constructed as a Wetland-Reservoir Subirrigation System

(WRSIS) in which a wetland and reservoir are interconnected and provide drainage as well as subirrigation to the surrounding fields (Fig. 4.1). Two 1.4 ha fields surround a

0.12 ha wetland. The wetland is connected to a 0.16 ha reservoir. During storm events,

126 127

Figure 4.1: Site map of the Defiance, OH wetland-reservoir subirrigation system. Key structures related to the wetland

function are shown in the legend. This map is not drawn to scale.

water draining from the fields is routed to the wetland. Because the water level in the wetland is maintained at a constant level, overflow is pumped to the reservoir for storage. In times of low rainfall, water from the reservoir is used to water the fields through the subirrigation system.

4.2.2 Sampling

Four subsamples were taken from each agricultural field and mixed to form a composite sample. The top 5-10 cm of soil were sampled for each subsample. The start position for sampling was approximately the midpoint of the field. The first subsample was taken approximately 9 m from the edge of the field. Each subsequent sample was taken approximately 9 m from the previous one. The two fields rotate between corn and soybean crops with atrazine being applied with the corn crop only. Table 4.1 lists the atrazine application history and samples obtained from each site.

Wetland sediments and overlying water were sampled with a 4 x 140 cm acrylic column. Three replicate sediment samples were mixed to form a composite. Surface water samples were collected as grab samples. Water samples were used without further processing or they were concentrated 200 fold by centrifugation at 10,000xg in a

Beckman J2-21 centrifuge (Beckman Coulter, Fullerton, CA) and re-suspended in a mineral salts solution as described in section 4.2.3. All samples were stored at 4°C before use.

128

Sampling date Samples taken Atrazine Amount of atrazine Field application applied (kg ha-1) application date (m/d/y) 8/21/99 A, B 5/9/99 1.88 B 4/4/00 A, B, Wetland 5/12/00 1.88 A sediment 5/26/00 A, B, Wetland 5/12/00 1.88 A sediment, wetland and reservoir water 9/19/00 A, B, Wetland 5/12/00 1.88 A sediment, Reservoir water 6/2/01 A, B, Wetland 5/7/01 1.88 B composite, top 4 cm, interface sediments, Wetland water, Reservoir water 8/9/01 A, B, Wetland 5/7/01 1.88 B sediment and water, Reservoir water 6/17/02 A, B, Wetland 5/29/02 1.88 A sediment and water, Reservoir water 7/22/02 Wetland and 5/29/02 1.88 A reservoir water 9/19/02 A, B, Wetland 5/29/02 1.88 A sediment and water, Reservoir water

Table 4.1 Atrazine application and sample information for all Defiance sites. The amount of atrazine applied is listed in kg of active ingredient.

129

4.2.3 Mineralization studies

Mineralization of [U-ring-14C]-atrazine under aerobic conditions was measured in biometers, which consisted of 60-ml serum bottles equipped with suspended 2-ml vials

14 to trap evolved CO2 (36) (Fig. 2.2). Each biometer received 5.0 g (dry weight) aliquots of sediment or soil samples, or 5 ml of water. Each sample was tested in duplicate biometers. For each experiment two soil samples were autoclaved at 121°C for 20 min. and used as a sterile control. Each biometer received 0.065 µmol [U- ring-14C]-atrazine (specific activity 1.54 mCi/mol; Sigma-Aldrich Co, St. Louis, MO) with a total concentration of 2.8 mg/ kg dry weight or l. Trace elements (per liter of 100X stock solution): MgSO4·7H2O, 50 mg; CaSO4, 200 mg; FeSO4·7H2O, 1 g; MnSO4·H2O, 20 mg;

CuSO4, 20 mg; ZnSO4·7H2O, 20 mg; CoSO4·7H2O, 10 mg; NaMoO4·2H2O, 5 mg; H3BO3,

5 mg; nitrilotriacetate, 3 g) was also added (100 µl of 100X) to fulfill any trace metal requirements. For anaerobic biometers, 1 ml of 71.3 mM KNO3, Fe(COOH)3, or Na2SO4 were added to provide an excess of electron acceptor beyond what was needed to fully mineralize all of the added atrazine. For samples with no additional electron acceptor, 1 ml of dH2O was added. Biometers were flushed with nitrogen for five minutes after biometer set-up and after each sampling.

Pittsburgh, PA) in a vial and counted in a scintillation counter. The counting efficiency was determined using an external standard and was determined to be 97%.

Mineralization was monitored at 22±2°C for 25-38 d.

130

4.2.4 Data analysis

-kt fitting the graphs to first-order rate function, expressed as P = Pmax (1-e ), where P =

14 observed amount of CO2 evolved (%), Pmax = the maximum extent of mineralization for

-1 that site, t = time (d), and k = rate constant (d ). Half-life (t1/2) values were calculated using their respective rate constants as t1/2 = ln2/k (22).

4.2.5 Mass balances

After the final sampling, the biometers were dried for approximately 24 h at 90°C.

The dried soil samples were crushed to a uniform particle size and 0.25 g samples (dry weight) were used for analysis. The samples were combusted in an oxidizer (Biological

Oxidizer OX-400, R. J. Harvey Instrument Corporation, Patterson, NJ) and the evolved

14 CO2 was trapped in scintillation fluid (Carbon-14 Cocktail, R. J. Harvey Instrument

Corporation). The cocktail was transferred to a vial for liquid scintillation counting. For mass balance estimates, the amount of radioactivity remaining in the sample was added

14 to the cumulative % CO2 from the mineralization studies.

4.2.6 DNA isolation

DNA was isolated from soil, sediment and water samples using the QIAquick Stool

Sample Kit (Qiagen, Valencia, CA). 10 g samples of soil and sediment were added to 10

131

ml 0.1% sodium pyrophosphate and incubated at 22±2°C with shaking for 2-3 hours.

The samples were then centrifuged at 22±2°C for fifteen minutes at 800 revolutions/minute. The supernatant was removed into a new tube. Approximately 1.7 ml of each soil, sediment, or concentrated water sample was placed in a microfuge tube and centrifuged at 15,000 revolutions/minute for 10 minutes. The supernatant was poured off and the pellet was resuspended as per Qiagen instructions. Positive controls were isolated from plasmid constructs transformed into Escherichia coli DHα strains

(pMD4 for atzA and pJ204 for trzD) or HB101 (pWM221011 for trzN). Plasmids were isolated using the Qiagen Minipreparation Kit.

Genomic DNA from pure cultures was isolated using a phenol/chloroform extraction technique. A single colony was transferred to a 100 ml of atrazine minimal media containing 10 mM K2HPO4, 5 mM NaH2PO4, 0.1 mM atrazine, 10 mM sucrose, 5 mM

MgCl2, and 1X mineral salts solution and grown for approximately 4 d at 22±2°C with shaking. Approximately 30 ml of cells were pelleted by centrifugation in a Beckman J2-

21 Centrifuge (Beckman Coulter, Fullerton, CA) at 12,000xg for 10 minutes. All centrifugation runs were performed at 4°C. The supernatant was poured off and the pellet was resuspended in 11.3 ml TE (10 mM Tris-Cl, pH8.0, 1 mM EDTA), 600 µl 10%

SDS, and 3.5 ml 200 mM NaOH. The suspension was incubated at 22°C for 10 minutes.

An equal volume of CHCl3:isoamyl alcohol (24:1) was added and the suspension mixed by inversion. The suspension was centrifuged for 10 min at 12,000xg and the aqueous layer was removed to a new centrifuge tube. An equal volume of phenol:CHCl3:isoamyl alcohol (25:24:1) was added and the suspension was mixed by inversion. The suspension was centrifuged at 12,000xg for 10 min. The aqueous layer was transferred

132

to a fresh tube and 2 ml of 5M NaCl was added. 0.6 volumes of isopropanol were added and the suspension was mixed gently by inversion. The suspension was centrifuged at

27,000xg for 15 min. The supernatant was poured off and approximately 25 ml of ice- cold 95% ethanol was added. The pellet was centrifuged for 15 min at 27,000xg. The supernatant was poured off and the pellet was allowed to air dry. The pellet was resuspended overnight at 4°C in 100 µl of dH2O.

4.2.7 PCR amplification

10 min. The annealing temperature for each primer set is listed in Table 2.2. All annealing steps were step down cycles that started 5°C higher than the listed annealing temperature and decreased 1°C/cycle to the desired annealing temperature. Nested

PCR reactions were set up as described above except 5 µl of the previous PCR reaction was used as the template. 5 µl of the positive control and 10 µl of all other amplified

133

samples were separated on a 1.0% agarose gel in TAE buffer. The DNA was visualized using a 10 mg/ml solution of ethidium bromide.

The PCR mixture to test for PCR inhibitors (final volume: 50 µl) contained 7µl soil template DNA, 0.5 µl LC2 DNA (ribosomal DNA), 1X PCR buffer, 25 pmol primers, 200

µM dNTPs, 2.25 mM MgCl2, 10 µl Q solution and 2.5 U HotStarTaq polymerase (Qiagen,

Valencia, CA). The ribosomal gene was amplified in a PTC-200 DNA engine thermocycler (MJ Research, Waltham, MA) using the following cycle: denaturation at

95°C for 15 min, 30 cycles of 94°C for 30 sec, 55°C for 30 sec, 72°C for 2 min, and a final extension step at 72°C for 7 min. A second round of PCR was performed using the previous reaction as a template. The samples were diluted 1:10 or 1:25 (total dilution over two reactions - 1:100 or 1:250) in the PCR reaction buffer and the PCR cycle was performed as above. The agarose gel was run as described above.

4.2.8 Most Probable Number (MPN) technique

The MPN technique was adapted from the protocol by Radosevich et al. (43).

1.02 g of sodium acetate to 100 ml dH2O. The MgSO4 was prepared by added 5 g of

MgSO4·7H2O to 100 ml dH2O. The mineral salts solution consists of 0.1 g FeCl3·6H2O,

0.1 CaCl2·2H2O, 0.001 MnCl2·4H2O, and 0.001 g ZnSO4·7H2O added to 100 ml dH2O and then filter sterilized. Five ml of medium were dispensed into 16x150 mm test tubes.

-1 -9 Samples were diluted 10 to 10 in 50 mM phosphate buffer (0.14 g/L KH2PO4 and 8.53

134

g/L K2HPO4). 1.0 ml of each sample was added to six replicate MPN tubes. One replicate from each dilution was autoclaved at 121 °C for 20 minutes. This replicate served as a sterile control to determine the amount of atrazine sorbed to the soil during the incubation. The rack of tubes was weighed and incubated in the dark at 22±2°C for

30 d. At the end of this period, the rack was reweighed and lost water was replaced in equal volumes to each tube. A one ml sample from each tube was removed and filtered through a 0.22 µm filter to remove particulate matter. 100 µl of the filtrate was added to

900 µl of HPLC mobile phase (65% acetonitrile, 35% dH2O). Samples were run on C18

Reichardt (8).

4.2.9 Enrichment for atrazine-mineralizing organisms

5 g of soil or sediment from the fields or wetland were added to 100 ml of atrazine minimal media (as described in the genomic DNA extraction) and incubated at 22±2°C with shaking. Samples (1.0 ml) of the enrichments were removed every two weeks and the amount of atrazine remaining was determined by HPLC as described in the MPN technique. When the atrazine was depleted in the culture, 1 ml of the culture was transferred a fresh flask of atrazine minimal medium. The cultures were transferred approximately eight times. One loopful of culture was transferred to a plate of atrazine minimal medium with 1.5 g/L agar. Colonies that grew on these plates were recultured several times to produce pure cultures. Once the cultures were determined to be pure,

135

they were transferred to liquid atrazine minimal media cultures and tested for the ability to mineralize atrazine by HPLC.

4.2.10 16S rDNA sequencing

Genomic DNA was isolated from a pure culture of C5-2B as described in section

4.2.6. A 20 µl sample of this DNA was run on a 0.6% low melting point agarose gel at 60

V for approximately 1.5 hr. DNA that was 1-10 kb in size was cut from the gel and heated at 70°C for 20 minutes to inactivate DNAses. This DNA was amplified using the

8f and U968r ribosomal primers (4). These products were cloned using the TOPO TA cloning system (Invitrogen, Carlsbad, CA). These clones were submitted to bi-directional sequencing with an ABI Prism model 377 sequencer (Perkin-Elmer Applied Biosystems,

Foster City, CA). The M13 universal forward and reverse primers which can bind to the pCR2.1 vector were used for sequencing.

4.2.11 FAME analysis

The C5-2B isolate was identified using fatty acid methyl ester (FAME) analysis.

Cultures were grown on TSA plates for 3 to 4 d at 27°C prior to harvest and extraction.

Cultures were harvested and lipids were extracted and derivitized according to the standard protocol for cultures grown on solid media (MIDI, Inc., Newark, DE). Total cellular fatty acid methyl esters were analyzed by gas chromatography according to the protocols outlined by MIDI (50, 61). The FAME analysis was performed at the University of Delaware, courtesy of Dr. Radosevich.

136

4.3 RESULTS AND DISCUSSION

4.3.1 Mineralization data-Yearly data

4.3.1.1 1999 sampling year

The extent of aerobic mineralization in field A was 58%, which is lower than the extent of mineralization in field B (73%). This difference may be due to a more active population in field A because it was more recently treated with atrazine (Figs. 4.2 and

4.3). However, the rate of mineralization was approximately equal in both fields, and neither samples had a major lag phase preceding the mineralization.

Mineralization was also examined in anaerobic biometers that received various electron acceptor amendments (nitrate, ferric-iron, or sulfate). In field A, external electron acceptors appeared to suppress the mineralization of atrazine. Biometers lacking an additional electron acceptor mineralized 58% of the added atrazine, while samples with additional electron acceptors mineralized 6-17% of the atrazine. Nitrate has previously been shown to inhibit atrazine mineralization (1, 21). In contrast, electron acceptors did not inhibit mineralization levels in field B samples. The extent of mineralization for all electron acceptor treatments was similar to that measured in aerobic biometers (59-70% compared to 73%).

4.3.1.2 2000 sampling year

In field A, planted with corn, the rate of aerobic mineralization was relatively low

-1 (k = 0.131 d ) in April (Table 4.2). However, the extent of aerobic mineralization was considerably higher than the 1999 samples (70% compared to 58%, Fig. 4.4). Because

137

100

90 No acceptor Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.2: Mineralization data for August 1999 samples taken from Defiance, OH field

A. Biometers were incubated under anaerobic conditions except for data labeled

14 aerobic. Each data point represents the mean cumulative percentage of CO2 evolved from two biometers. Standard deviations where not shown are smaller than the symbols.

138

100 No acceptor 90 Iron Nitrate 80 Sulfate Aerobic 70

50 evolved (%) evolved 2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.3: Mineralization data for August, 1999 samples taken from field B in Defiance,

OH. All samples were incubated under anaerobic conditions except the sample labeled

14 aerobic. The means shown represent the cumulative CO2 evolved as a percentage of the total amount of radioactivity added. Standard deviations where not shown are smaller than the symbols.

139

-1 2 Sample Date k (d ) r t1/2 (d) A-No acceptor Apr-00 0.047 0.9571 14.81 A-Aerobic Apr-00 0.131 0.9763 5.28 A-Iron Apr-00 0.072 0.9812 9.69 A-Nitrate Apr-00 0.105 0.9059 6.60 A-Sulfate Apr-00 0.094 0.9492 7.37 B-No acceptor Apr-00 0.046 0.9799 15.00 B-Aerobic Apr-00 0.095 0.9377 7.26 B-Iron Apr-00 0.054 0.9827 12.76 B-Nitrate Apr-00 0.052 0.9389 13.30 B-Sulfate Apr-00 0.061 0.9337 11.36 Wetland sediment-No acceptor Apr-00 0.010 0.8645 67.94 Wetland sediment-Aerobic Apr-00 0.039 0.8634 18.00 Wetland sediment-Iron Apr-00 0.011 0.8618 65.38 Wetland sediment-Nitrate Apr-00 0.036 0.9557 19.14 Wetland sediment-Sulfate Apr-00 0.017 0.8878 41.75

A-No acceptor May-00 0.047 0.9683 14.74 A-Aerobic May-00 0.188 0.9496 3.69 B-No acceptor May-00 0.063 0.9518 11.00 B-Aerobic May-00 0.235 0.9738 2.95 Wetland sediment-Aerobic May-00 0.038 0.8150 18.24

A-No acceptor Sep-00 0.068 0.8912 10.22 A-Aerobic Sep-00 0.169 0.9507 4.10 B-No acceptor Sep-00 0.040 0.8944 17.50 B-Aerobic Sep-00 0.140 0.9264 4.95 Wetland sediment-Aerobic Sep-00 0.037 0.7971 18.83 Reservoir water-Aerobic Sep-00 0.047 0.7756 14.81

Table 4.2: Kinetic parameters and half-lives calculated for samples obtained in 2000.

140

100 No acceptor 90 Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.4: Mineralization data for field A samples taken in April, 2000. All samples were incubated under anaerobic conditions except for the sample labeled aerobic. All points

14 were determined by taking the mean of two biometers. The CO2 evolved is based on

14 the cumulative evolution of CO2 over time. The standard deviations not shown are smaller than the symbols.

141

atrazine had not yet been applied in April, atrazine application does not account for the large increase in the extent of mineralization.

Mineralization under anaerobic conditions also showed an increase in the extent of mineralization (Fig. 4.4). The highest extent of mineralization was seen with added nitrate, in contrast with the previous experiment. Biometers with added sulfate and iron also showed relatively high levels of mineralization (67 and 61%, respectively).

Anaerobic biometers with no added electron acceptor showed the lowest extent of mineralization. The rate of mineralization for all biometer treatments was similar.

Field B also showed an extent of mineralization under aerobic conditions similar to the previous year (69% compared to 73%) (Fig. 4.5). Anaerobic mineralization was slightly lower than aerobic mineralization. However, the extent of mineralization was similar for all electron acceptors (51-62%). In the wetland, the extent of aerobic mineralization was 62%; however, the rate of mineralization was rather slow, with a lag of approximately 7 d before an increase in atrazine mineralization occurred (Fig. 4.6).

Under anaerobic conditions, nitrate-coupled atrazine mineralization was the highest with

46% mineralized, followed by sulfate, iron, and no added acceptor. The initial rate of mineralization, when nitrate was added, was higher than that of aerobic mineralization, and the lag period shorter (3 d compared to 7 d). The rate of mineralization for other anaerobic conditions was extremely slow and exhibited lag periods of 20-27 days.

May samples taken from the agricultural fields show a high extent of mineralization (84%) under aerobic conditions in both fields (Figs. 4.7 and 4.8). Under anaerobic conditions without the addition of an external electron acceptor, the extent of mineralization was much lower in field A than field B. Wetland sediment samples were

142

100 No acceptor 90 Iron 80 Nitrate Sulfate 70 Aerobic

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.5: Mineralization data for field B samples taken in April, 2000. All samples were incubated under anaerobic conditions except for the sample labeled aerobic. Data points

14 were determined from the cumulative CO2 evolved as a percentage of the total radioactivity added. The standard deviations not shown are smaller than the symbols.

143

100 No acceptor 90 Iron Nitrate 80 Sulfate 70 Aerobic

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 35 40 Time (Days)

Figure 4.6: The mineralization time course for wetland sediment samples taken in April,

2000. Biometers were incubated under anaerobic conditions except for the sample

14 labeled aerobic. All data points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity evolved. Standard deviations, where missing, are smaller than the symbols.

144

100 No acceptor 90 Aerobic 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.7: Mineralization data from May 2000 field A samples. The no acceptor samples were incubated under anaerobic conditions while the aerobic samples were incubated

14 aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

145

100 No acceptor 90 Aerobic 80

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.8: Mineralization data from May 2000 field B samples. The no acceptor samples were incubated under anaerobic conditions while the aerobic samples were incubated

14 aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

146

examined under aerobic conditions only (Fig. 4.9). The extent of mineralization was almost equal to the value observed in April samples (63%). The lag period of 8 d was also similar to the observed lag in April.

In September, the biometer experiments included both aerobic and anaerobic mineralization. The extent of mineralization in both agricultural fields was identical at

81% (Figs. 4.10 and 4.11). The rate of mineralization was also very similar between field

A, with k values of 0.169 d-1 and field B, with k values of 0.139 d-1 (Table 4.2). This may indicate that both fields have an active population of microorganisms capable of mineralizing atrazine, although only field A received an application of atrazine. The rate of atrazine mineralization under anaerobic conditions was higher for field A than for field

B (k = 0.68 d-1 and 0.40 d-1, respectively). However, the extent of mineralization was similar (74% and 65%, respectively).

A comparison of the rate of mineralization from the April, 2000 samples to the

October, 2000 samples shows that the rate and half-life for atrazine in field A decreased

-1 -1 from k = 0.047 d (t1/2 = 15 d) in April to k = 0.068 d (t1/2 = 10 d) in October (Table 4.2).

This shift may be due to the application of atrazine following the April sampling, which may have increased the activity of the microbial population. Field B had a k = 0.046 d-1

-1 (t1/2 = 15 d) in April which decreased slightly to k = 0.040 d (t1/2 = 18 d) by October. This increase in the half-life of atrazine may reflect a decrease in the active population (either numbers of organisms or their activity) due to a lack of selective pressure because field

B was planted with soybeans and did not receive atrazine that year.

In the wetland sediment, the extent of aerobic mineralization was 60%, which is similar to the extent of mineralization in April (Fig. 4.12). The lag period for this sample

147

100

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30

Time (Days)

Figure 4.9: Mineralization data from May 2000 wetland sediment samples which were

14 incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

148

100 No acceptor 90 Aerobic 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.10: Mineralization data from September 2000 field A samples. The no acceptor samples were incubated under anaerobic conditions while the aerobic samples were

14 incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

149

100 No acceptor 90 Aerobic 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.11: Mineralization data from September 2000 field B samples. The no acceptor samples were incubated under anaerobic conditions while the aerobic samples were

14 incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

150

100 Wetland sediment 90 Reservoir water 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 102030 Time (Days)

Figure 4.12: Mineralization data from September 2000 wetland and reservoir samples.

14 All samples were incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

151

was approximately 7 d. Reservoir water samples that were preconcentrated 200-fold prior to testing in biometers showed a high extent of mineralization (70%).

4.3.1.3 2001 Sampling year

In June, agricultural soil samples from field A showed a 57% loss of atrazine from the biometers and a lag period of approximately 3 d (Fig. 4.13). Field B received an application of atrazine in 2001, had a high extent of mineralization (77%), and did not show a lag period. Thus field B had a more active atrazine-mineralizing population, and the genes involved in atrazine mineralization may have already been induced in this sample, as indicated by the lack of a lag phase.

To examine further the location of atrazine-mineralizing organisms in the wetland, three different sediment samples were taken. A composite sediment sample was obtained by mixing four individual subsamples. Other subsamples were obtained in the same way, but only the top 4 cm of sediment were retrieved, excluding the bottom portion of the sediment core. The excluded portion of the sample had a high clay content. Finally, samples were taken of the flocculent, silty material located just at the interface of the sediment bed and the water column. which was identified as the interphase. The rate and extent of mineralization for the composite samples and the top

4 cm were similar (Fig. 4.14 and Table 4.3). These data suggest that the most active portion of the sediment column is located in the top portion of the sediment column. The interphase had a low extent of mineralization. This may be due to a low population of atrazine mineralizing organisms. This may be further supported by examining the rate and extent of mineralization in the water samples which have a similar composition to

152

100

50 Evolved (%) Evolved

2 40 A CO

14 30 B B Sterile Reference 20 A Sterile Reference

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.13: Mineralization data from June 2001 agricultural samples. All samples were

14 incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

153

100 Top 4 cm-Aerobic 90 Top 4 cm-Nitrate Interphase-Aerobic 80 Interphase-Nitrate 70 Composite

50 Evolved (%) Evolved

2 40 CO

14 30

0 1020304050 Time (Days)

Figure 4.14: Mineralization data from June 2001 wetland samples. Samples labeled nitrate were incubated anaerobically with nitrate as an added electron acceptor. All other

14 samples were incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Composite samples represent the mean of duplicated biometers. All other samples represent the mean of five biometers. Standard deviations, where not shown, are smaller than the symbols.

154

-1 2 Sample Date k (d ) r t1/2 (d) A Aerobic Jun-01 0.054 0.9516 12.91 B-Aerobic Jun-01 0.379 0.9656 1.83 Wetland sediment composite-Aerobic Jun-01 0.083 0.9374 8.37 Wetland top 4 cm-Aerobic Jun-01 0.071 0.9346 9.79 Wetland top 4 cm-Nitrate Jun-01 0.008 0.7202 91.18 Wetland interphase-Aerobic Jun-01 0.005 0.8128 141.43 Wetland interphase-Nitrate Jun-01 0.001 0.8679 1386.00 Wetland water Jun-01 0.115 0.9292 6.05 Reservoir water Jun-01 0.046 0.7753 15.03

A Aerobic Aug-01 0.038 0.9637 18.24 B-Aerobic Aug-01 0.277 0.9643 2.50 Wetland sediment-Aerobic Aug-01 0.105 0.9260 6.62 Wetland water Aug-01 0.115 0.8242 6.02 Reservoir water Aug-01 0.142 0.9017 4.90

Table 4.3: Kinetic data and half-lives calculated for the 2001 biometer experiments.

the interphase, but were concentrated before use. Both wetland and reservoir water samples showed a high extent of mineralization (60% and 82% respectively, Fig.4.15) compared to the interphase samples (12%). The rate and extent of mineralization of the interphase and top 4 cm samples under anaerobic conditions was also examined.

Nitrate was added as a terminal electron acceptor in these samples. The top 4 cm samples had an extremely long lag period of 22 d and the extent of mineralization was low (33%). The interphase samples did not have a significant extent of

155

100 Reservoir water 90 Wetland water 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.15: Mineralization data from June 2001 water samples which were preconcentrated before use. All samples were incubated aerobically. Points represent

14 the mean cumulative CO2 evolved as a percentage of the total radioactivity added.

Standard deviations, where not shown, are smaller than the symbols.

156

mineralization. Again this is likely due to a low concentration of atrazine mineralizing organisms in the unconcentrated sample.

The August samples for field A showed a similar extent of mineralization to the samples taken in June (Fig.4.16). In contrast, the extent of mineralization in field B samples was higher by 10% in August. However, the rate of mineralization for both samples was slower than the June samples (Table 4.3).

The wetland sediment sample showed a high extent of mineralization (75%). The rate of mineralization was rapid with a k = 0.105 d-1 and a half-life of 7 d (Fig. 4.17). The lag period in the wetland was only 2 days, an extremely short time in comparison to previous years. Both concentrated water samples showed a high extent of mineralization of 89% (Fig. 4.17). This is much higher than wetland sediment samples. However, a direct comparison between sediment and water samples is impossible since the water samples are concentrated. Because the amount of sediment in the concentrated samples can vary greatly from sample to sample, this seems to have an effect on the rate and extent of mineralization, in contrast to the sediment cores taken from the wetland.

Figure 4.18 shows the rate and extent of mineralization from samples taken from fields A and B. Both fields show a similar extent of mineralization and the rate of mineralization is very close, with field A having a k value of 0.211 d-1 and field B having a k value of 0.225 d-1 (Table 4.4). Although only field A received an application of atrazine, field B is also extremely active. This may reflect the acclimation of the soil to repeated applications of atrazine (e.g. the history effect). However, the population in field B is unlikely to be actively mineralizing atrazine as shown by the 2 day lag period observed.

157

100 A 90 B 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.16: Mineralization data from August 2001 agricultural samples. All samples

14 were incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

158

100 Wetland water 90 Reservoir water Wetland sediment 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 Time (Days)

Figure 4.17: Mineralization data from August 2001 wetland and reservoir samples. Water samples were preconcentrated prior to use. All samples were incubated aerobically.

14 Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

159

100 A 90 B 80

50 Evolved (%) Evolved

2 40 CO

14 30

0 102030 Time (Days)

Figure 4.18: Mineralization data from June 2002 agricultural samples. All samples were

14 incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the points.

160

4.3.1.4 2002 Sampling year

In the wetland sediment, the extent of mineralization is slightly lower than the observed extent of mineralization in 2001 (Fig. 4.19). The lag period is also slightly longer (6 d, 2001 = 2-3 d). Water samples taken from the wetland and reservoir showed a high extent of mineralization, although the reservoir samples were slightly more active than the wetland water sample (Fig. 4.19). Water samples taken in July 2002 showed significant changes as compared to the June samples. The extent of mineralization in the reservoir samples dropped slightly from 85% to 78% (Fig. 4.20). However, the wetland water sample had an extent of mineralization of less than 1%. This may be due to rainfall levels in Defiance. The spring of 2002 was extremely wet. However, a severe drought was experienced during the summer months. The water level in the wetland was lower than normal at the time of sampling and the amount of suspended solids was greatly decreased. This likely caused the decrease in mineralization capabilities. The extent of mineralization in wetland sediments was not significantly different from the

June samples, indicating the potential for atrazine mineralization was still present in the wetland.

Agricultural samples taken in September showed a similar extent of mineralization although the rate of mineralization had decreased slightly (Fig. 4.21,

Table 4.4). This indicates that the population was still actively mineralizing atrazine.

Wetland sediment samples showed a similar extent of mineralization, although the rate was slower and the lag period was longer in September (Fig. 4.22). Reservoir water samples also showed a similar extent of mineralization to the July samples. The most significant change was a recovery of the atrazine mineralizing population in the wetland

161

100 Wetland sediment 90 Wetland water 80 Reservoir water

50 Evolved (%) Evolved

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.19: Mineralization data from June 2002 wetland and reservoir samples. Water samples were preconcentrated prior to use. All samples were incubated aerobically.

14 Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the points.

162

100 Wetland water 90 Reservoir water Wetland sediment 80

50 Evolved (%)

2 40 CO

14 30

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.20: Mineralization data from July 2002 wetland and reservoir samples. Water samples were preconcentrated prior to use. All samples were incubated aerobically.

14 Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the points.

163

100 A 90 B 80

50 Evolved (%) Evolved

2 40

CO 30 14

0 5 10 15 20 25 30 Time (Days)

Figure 4.21: Mineralization data from September 2002 soil samples. All samples were

14 incubated aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

164

100 Wetland sediment 90 Wetland water Reservoir water 80

50 Evolved (%) Evolved

2 40

CO 30 14

0 5 10 15 20 25 30 Time (Days)

Figure 4.22: Mineralization data from September 2002 wetland and reservoir samples.

Water samples were preconcentrated prior to use. All samples were incubated

14 aerobically. Points represent the mean cumulative CO2 evolved as a percentage of the total radioactivity added. Standard deviations, where not shown, are smaller than the symbols.

165

-1 2 Sample Date k (d ) r t1/2 (d) A-Aerobic Jun-02 0.211 0.96643.29 B-Aerobic Jun-02 0.225 0.92163.08 Wetland sediment-Aerobic Jun-02 0.052 0.9029 13.30 Wetland water-Aerobic Jun-02 0.070 0.8185 9.84 Reservoir-Aerobic Jun-02 0.085 0.83198.17

Wetland sediment-Aerobic Jul-02 0.102 0.9199 6.77 Wetland water-Aerobic Jul-02 0.000 0.9391 3465.00 Reservoir-Aerobic Jul-02 0.091 0.83517.63

A-Aerobic Oct-02 0.202 0.94453.43 B-Aerobic Oct-02 0.166 0.92684.18 Wetland sediment-Aerobic Oct-02 0.052 0.8877 13.46 Wetland water-Aerobic Oct-02 0.025 0.8465 27.50 Reservoir-Aerobic Oct-02 0.077 0.79149.02

Table 4.4: Kinetic data and half-lives calculated for 2002 mineralization samples.

water samples (Fig. 4.22). The drought which had been affecting Ohio had somewhat recovered by this time which likely lead to the increased ability to mineralize atrazine in the water column. However, the extent of mineralization had still not recovered to levels seen previously.

166

4.3.2 Mineralization data - Summaries

Because the agricultural fields at Defiance only receive an application of atrazine every two years, it was hypothesized that there would be year-to-year differences in the extent of mineralization in these fields. Figure 4.23 shows the crop summary for field A which shows some clustering in the extent of mineralization based on the crop planted/atrazine application for that year. The years in which soybean was planted showed a lower extent of mineralization while years in which atrazine was applied showed a higher extent of mineralization. However, the rate of mineralization varied each year and did not show a pattern similar to the extent of mineralization.

Figure 4.24 shows the crop summary for field B. These samples do not show the same clustering effect seen in field A. The rate of mineralization does not seem to follow a pattern. This suggests that the microbial population in field B is more acclimated to atrazine than the population in field A. The response in field B may be explained by the history effect in which repeated applications of atrazine seem to enhance the activity of the resident populations (37). If this site was monitored further, it is likely that field A would show a similar increase in the activity of the resident population that would persist even in the absence of atrazine. Another source of variability in the microbial responses may be attributed to the higher organic matter content in field B which was largely forested prior to the site construction. Field A was a meadow and so had a lower organic matter content. The organic matter may be stimulating the atrazine mineralizing organisms in manner similar to the results observed by Abdelhafid et al. (1).

The effect of atrazine application was also studied in these fields (Fig. 4.25). Two samples were taken from both fields; one sample approximately two weeks prior to

167

100

50 Evolved (%) Evolved

2 40 Soybean (08/99) Corn (10/00) CO 30 14 Soybean (08/01) Corn (09/02) 20 Sterile reference 10

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.23: Crop rotation mineralization data for field A. The crops planted each year

14 are listed on the key. Points plotted represent mean cumulative % CO2 evolved as a percentage of the added radioactivity. Standard deviations may be smaller than the symbols.

168

100

60 Evolved (%) Evolved

2 40 CO 14

20 Corn (08/99) Soybean (10/00) Corn (08/01) Soybean (09/02) 0

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.24: Crop rotation mineralization data for field B. The crops planted each year

14 are listed on the key. Points plotted represent mean cumulative % CO2 evolved as a percentage of the added radioactivity. Standard deviations may be smaller than the symbols.

169 Field A Field B

100

60 Evolved (%)

2 40 CO

14 20 170 Pre-application Post-application 0

0 5 10 15 20 25 0 5 10 15 20 25 30

Time (Days)

Figure 4.25: Effect of atrazine application on the extent of mineralization in samples from field A and B. Points plotted

14 represent mean cumulative % CO2 evolved as a percentage of the added radioactivity. Standard deviations may be

smaller than the symbols.

atrazine application and a second sample approximately two weeks after atrazine application. The samples taken prior to atrazine application show a similar extent of mineralization while the rate appears to be slightly faster in field A. The samples taken post-atrazine application show a higher extent of mineralization as compared to the pre- application samples. However, only field A received an application of atrazine that year.

This suggests that atrazine application alone does not cause an increase in the rate or extent of mineralization. Instead this is likely due to a combination of factors which may include soil temperature, moisture content, or soil organic matter content.

Figure 4.26 shows a comparison of sediment samples taken each year and analyzed for mineralization potential. In 1999, a survey of several sites was conducted to determine if any had the potential to mineralize atrazine. These samples underwent a 7 day biometer incubation. The wetland did not mineralize atrazine to a significant extent, so it was assumed that the sediment did not currently have an active microbial population capable of mineralizing atrazine. In 2000, a new sediment sample was obtained from the wetland and analyzed for mineralization activity during a full 28 day time course. This sample mineralized approximately 60% of the added atrazine. These samples also showed a lag period of approximately 8 days, suggesting that the 1999 sample may have shown mineralizing activity if the incubation period had been longer.

The 2001 and 2002 samples show a similar extent of mineralization. However, if the rate of mineralization is examined, the rate has increased from samples taken in 2000. The lag period has also shorted to 3-4 days. Although the lag seen prior to active

171

100

50 Evolved (%) Evolved

2 40 CO

14 30

20 06/99 06/00 10 06/01 06/02 0

0 5 10 15 20 25 30 35 Time (Days)

Figure 4.26: Composite atrazine mineralization data for wetland sediment samples.

14 Points plotted represent mean cumulative % CO2 evolved as a percentage of the added radioactivity. Standard deviations may be smaller than the symbols.

172

mineralization suggests that the wetland may not be removing atrazine from the water, the potential for the wetland to actively remediate atrazine-containing water in the future is high.

4.3.3 PCR amplification

Three genes involved in atrazine mineralization were chosen to determine whether the presence of these genes could indicate atrazine mineralization potential.

The atzA gene was selected because it has been identified in a large number of atrazine-mineralizing isolates. trzN was recently identified from a gram-positive

Nocardiodes sp. It is a functional homolog of atzA although it shows no sequence homology. The final gene, trzD, is commonly associated with atrazine degrading and mineralizing organisms and so was also included in the study. Figure 4.27 shows PCR products from genomic DNA amplified with atzA primers. This figure shows that only the positive control produces a signal of the appropriate size (528 bp). Figure 4.28 shows trzN amplified products. In this gel, products of the appropriate size were amplified from field A (planted with corn) as well as wetland sediment and water samples. No signal was obtained from field B (soybean) or the reservoir. It is apparent from this gel that the primers are not specific solely for the trzN gene. This is likely caused by the sequential amplification of PCR products using the same primer set. Also, because of the variety of organisms present, it is difficult to select primer sets which will only amplify the gene of interest and exclude all other genes present. The same nonspecific amplification problems were also observed with the trzD PCR products shown in Figure 4.29. Only the wetland samples show positive signals for trzD. However, the size of the product

173

2.0 kb

1.0 kb

528 bp 0.5 kb

0.1 kb

Figure 4.27: Agarose gel electrophoresis of PCR products amplified with the atzA gene.

Lane 1: pMD4 (positive control); Lane 2: No DNA (negative control); Lane 3: MW markers; Lane 4: Field A; Lane 5: Field B; Lane 6: Wetland sediment; Lane 7: Wetland water.

174

2.0 kb 1.0 kb

0.5 kb 420 bp

0.1 kb

Figure 4.28: Agarose gel electrophoresis of PCR products amplified with the trzN gene.

Lane 1: pWM221011 (positive control); Lane 2: No DNA (negative control); Lane 3: MW markers; Lane 4: Field A; Lane 5: Field B; Lane 6: Wetland sediment; Lane 7: Wetland water; Lane 8: Reservoir water.

175

2.0 kb

1.0 kb

485 bp 0.5 kb

0.1 kb

Figure 4.29: Agarose gel electrophoresis of PCR products amplified with the trzD gene.

Lane 1: pJK204 (positive control); Lane 2: No DNA (negative control); Lane 3: MW markers; Lane 4: Field A; Lane 5: Field B; Lane 6: Wetland sediment; Lane 7: Wetland water; Lane 8: Reservoir water.

176

amplified from these samples is slightly larger than the expected 485 bp. It is possible that the product amplified is not quite homologous to the trzD gene; therefore sequencing of this gene will be performed in the future.

Because none of the target genes could be amplified from field B DNA samples, the presence of inhibitors was tested for in field B and the other samples from Defiance

(Fig. 4.30). All the samples except field A produced PCR products when diluted 1:100.

DNA from field A gave a positive signal when diluted 1:250 (Fig. 4.30B). However, with the exception of the wetland water, the bands amplified were not the correct size for the ribosomal DNA. The amplification of any DNA indicates that the inhibitors have been eliminated by dilution.

It is unclear why genes could be amplified from the other samples, but not from field B. Fields A and B have been classified as Paulding soils but field B contains a slightly higher clay content. It is possible that field B soil samples had a higher concentration of inhibitors. It is also possible that the number of atrazine mineralizing organisms were lower in field B (which did not receive an application of atrazine), which prevented positive PCR signals for the tested genes. The possibility cannot be excluded that the microbial population present in this field lacked all of the genes selected for this study.

4.3.4 Isolate characterization

One organism (C5-2B) was enriched from Defiance soils which had the ability to mineralize atrazine completely as shown by HPLC analysis. The colonies on atrazine

177 A B

2.0 kb 2.0 kb 1.0 kb

1.0 kb 0.5 kb

0.5 kb 178

0.1 kb 0.1 kb

Figure 4.30: Agarose gel electrophoresis of PCR inhibitor study. Genomic DNA from soil samples was spiked with

ribosomal DNA, diluted either 1:100 (A) or 1:250 (B) and then amplified with ribosomal specific primers. Lane 1: LC2

(positive control); Lane 2: No DNA (negative control); Lane 3: MW markers; Lane 4: Field A; Lane 5: Field B; Lane 6:

Wetland sediment; Lane 7: Wetland water; Lane 9: Reservoir water; (B) Lane 1: Field A; Lane 2: LC2; Lane 3: No DNA;

Lane 4: MW markers.

minimal agar were 1 mm in diameter, white, entire, and convex. The organisms were gram-positive and could assimilate glucose, mannose, maltose, malate, and citrate as carbon sources. They are not capable of nitrate reduction. Figure 4.31 shows an agarose gel with genomic DNA amplified with the atzA gene. This organism does not have the atzA gene as shown by a lack of amplified product. However, the organism does have a PCR product of a size consistent with trzN (Fig. 4.32). Figure 4.33 indicates that the isolate (CE-2B) does not have the trzD gene. This is surprising since a PCR product of a size similar to the trzD gene was amplified from wetland samples. It is possible that there is a gene present which has a sequence homologous to trzD is present in the soil samples which are not present in the isolate. The organism has been identified as an Arthrobacter sp. by both 16S rDNA sequencing and FAME analysis. The

16S rDNA sequencing indicated 99% homology to A. aurescens (627/628 bp). The closest FAME matches were to Arthrobacter ilicis and A. aurescens.

4.3.5 Most Probable Number (MPN) analysis

Although MPN analysis is commonly used to enumerate atrazine-mineralizing organisms, researchers in the field indicate that the MPN estimates obtained using this technique underrepresent the numbers of atrazine-degrading organisms present (22,

61). An attempt was made to improve the MPN analysis for atrazine-degrading organisms by changing the carbon source used to support the organisms. Current protocols use acetate as an additional carbon source. However, this carbon source is not readily metabolized and may be ineffective as a means of encouraging growth. The

179

2.0 kb

1.0 kb 1.0 kb

0.5 kb

0.1 kb

Figure 4.31: Agarose gel electrophoresis of PCR products amplified with primers specific for the atzA gene. Lane 1: pMD4 (positive control); Lane 2: No DNA (negative control);

Lane 3: C5-2B; Lane 4: MW markers.

180

2.0 kb

1.0 kb

0.5 kb 420 bp

0.1 kb

Figure 4.32: Agarose gel electrophoresis of PCR products amplified with trzN primers.

Lane 1: pWM221011 (positive control); Lane 2: No DNA (negative control); Lane 3: MW markers; Lane 4: C5-2B.

181

2.0 kb

1.0 kb 1.0 kb

0.5 kb

0.1 kb

Figure 4.33: Agarose gel electrophoresis of PCR products amplified with trzD primers.

Lane 1: pJK204 (positive control); Lane 2: MW markers; Lane 3: No DNA (negative control); Lane 4: C5-2B.

182

Sample Carbon source MPN (cells/g soil dry wgt.) 10% Loss 25% Loss 50% Loss A (Corn) acetate 3.4x101 2.2x100 1.1x101 A (Corn) glucose 2.0x100 1.1x101 1.1x101 A (Corn) NB 4.4x100 2.2x100 1.1x101 A (Corn) sucrose 8.1x100 6.0x100 6.0x100 A (Corn) soil extract 2.2x100 2.2x100 2.2x100 A (Corn) atrazine 6.6x100 4.4x100 4.4x100

Table 4.5: MPN estimates for samples incubated with various carbon sources. The 10% loss indicates the MPN estimate for tubes which lost 10% of the added atrazine compared to the sterile controls as measured by HPLC. Similar analyses were performed for 25% and 50% loss.

effect of several other carbon sources on MPN estimates are shown in Table 4.5. It does not appear that any of the added carbon sources increased the MPN estimate over that of the acetate tubes. These very low numbers are inconsistent with the high extent of mineralization observed in the soil samples and are in contrast with the results seen for the Piketon soils (Table 2.6). Although the MPN estimates likely do not reflect the numbers of atrazine degrading organisms present, the minimum number of atrazine degrading organisms needed to mineralize a high extent of atrazine has never been evaluated.

183

4.4 Summary of results

The agricultural fields showed varied extents and rates of mineralization according to the field sampled. Field A shows a change in the extent of mineralization based on the crop planted which indicates that the population of atrazine-mineralizing organisms has not fully developed. In contrast, field B shows a similar rate and extent of mineralization regardless of the crop planted. In general, samples with added electron acceptor show a higher rate and extent of mineralization than biometers incubated under anaerobic conditions with no additional electron acceptor. In the wetland sediments, the rate and extent of mineralization appears to be increasing with each subsequent planting season which indicates that the atrazine mineralizing population is becoming more active.

Concentrated water samples from the wetland and reservoir show a high extent of mineralization which indicates that organisms are being transported within the water column. Genomic DNA extracted from soil samples from all sites indicated that trzN is found in all samples except field B. atzA could not be amplified from any sample and trzD was amplified only from wetland samples. All Defiance samples were positive for inhibitors, but they could be diluted out at 1:100 (with the exception of field A), which is the same dilution factor used in the other PCR reactions. This suggests that there is some other reason for the lack of PCR amplification products from field B; e.g., a low concentration of DNA or lack of the tested genes in that community of microorganisms.

C5-2B, an isolate obtained from corn soil, is a member of the Arthrobacter genus and can completely mineralize atrazine as measured by HPLC. This organism has the trzN

184

gene but does not appear to have atzA or trzD. MPN analysis of field A soils showed no improvement of the MPN estimates when various carbon sources were substituted for acetate.

185

CHAPTER 5

FUTURE DIRECTIONS

The goal of this work was to evaluate agricultural and wetland sites for atrazine mineralization potential and the presence of specific atrazine catabolic genes. All three sites, Piketon, Defiance, and OSU wetland, showed atrazine mineralization potential.

However, the atrazine mineralization potential can not be determined solely on the basis of catabolic gene detection. Ultimately, it would be important to develop methods that can be used to promote in situ atrazine biodegradation or mineralization and thereby prevent the entry of atrazine into drinking water supplies. Establishing wetlands to treat agricultural drainage and runoff is one method which may help eliminate or decrease concentrations of atrazine entering receiving surface waters and ground water aquifers.

Detailed knowledge of the catabolic genes involved in atrazine mineralization may allow the design and application of enzymes for atrazine decomposition in drinking water treatment. However, much work remains to bring these ideas into reality.

186

Although some preliminary work was performed using anaerobic conditions, this area is relatively unexplored. Because the addition of electron acceptors seemed to have varied effects depending on the crop and season, further characterization seems warranted. Future experiments could involve characterization of differences in soil chemistry, the effect of crops, or the addition of various soil amendments such as phosphorus and nitrogen. In addition, microbial populations participating in anaerobic mineralization could be evaluated using T-RFLP techniques to detect shifts in the populations exposed to different conditions. Comparisons could be made between aerobic and anaerobic populations amended with various electron acceptors.

Much work can be potentially accomplished with genes involved in atrazine metabolism. Because some of the results obtained in these studies were ambiguous due to inhibitors, optimization of the DNA extraction and PCR amplification is needed for environmental samples including hydric and clay soils and those with a high content of organic matter. Although only three genes were chosen for this study, there are approximately a dozen genes known to be involved in atrazine metabolism. Both community and pure culture DNA could be examined for the presence of known atrazine genes to build a profile of the genes involved in atrazine mineralization in various environments. Other molecular genetic techniques could be used to examine the regulation of the catabolic genes. Reporter genes could be fused to the known catabolic genes allowing research into the expression of these genes under various environmental scenarios. Expression of these genes in a well-defined and simple system such as silica sand–based microcosms (biometers) over a time course provide knowledge on the regulation of various genes in a population of organisms without the complexity of soil-

186

borne factors. Other techniques, such as microarrays could be combined with T-RFLP analysis to provide information on community changes. The expression of the known genes in atrazine metabolism could be monitored in populations incubated under aerobic and anaerobic conditions as well as with the amendment of various electron acceptors.

Sequencing of mRNA from microarrays could also provide information on other proteins whose expression is stimulated or depressed by atrazine application under various conditions.

Wetland design and optimization could be further explored in relation to pesticide attenuation and remediation. Conditions such as sediment chemistry and composition as well as community composition and activity are areas that could be explored further.

This could provide information which will allow for the design of wetlands which are better able to remove pesticides from the water column.

In more global terms, the origin and evolution of the atrazine mineralization genes are unknown. Because atrazine is thought to be a true xenobiotic compound, it is intriguing that so many genes have evolved in the relatively short period of time – about

50 years – that atrazine has been utilized in farming. Investigation into the origin of these genes could be explored by looking at the phylogenetic relationships between the genes, for example, of the amidohydrolase family whose protein structure appears to be conserved among several members of the atz gene family. Another are of exploration would involve lateral transfer of genes or plasmid DNA among bacterial populations in agricultural or wetland sediments. Because all of the currently known atrazine metabolic genes are plasmid-borne, it is plausible that gene transfer occurs in soil and sediment environments, allowing a broad range of microorganisms to participate in atrazine

187

mineralization. Researchers investigating the atzA gene have speculated that this gene is easily transferred as shown by its widespread detection from pure cultures of microorganisms isolated from around the world (14). One possible approach would be to tag one of the genes involved in atrazine mineralization and transform the tagged gene into a subset of atrazine mineralizing organisms. These organisms could be added back to a defined system and then the transfer of the gene could be monitored spatially with the system or by a PCR based method that would allow quantification of the gene.

188

REFERENCES

1. Abdelhafid, R., S. Houot, and E. Barriuso. 2000. Dependence of atrazine degradation on C and N availability in adapted and non-adapted soils. Soil Biol. Biochem. 32:389-401.

2. Accinelli, C., G. Dinelli, A. Vicari, and P. Catizone. 2001. Atrazine and metolachlor degradation in subsoils. Biol. Fert. Soils. 33:495-500.

3. Allred, B. J., L. C. Brown, N. R. Fausey, R. L. Cooper, W. B. Clevenger, G. L. Prill, G. A. La Barge, C. Thronton, and B. J. Czartoski. A water table management approach to enhance crop yields for wetland reservoir subirrigation systems.

4. Amann, R. I. and D. A. Stahl. 1992. Dual staining of natural bacterioplankton with 4', 6-diamino-2-phenylindole and fluorescent oligonucleotide probes targeting kingdom-level 16S rRNA sequences. Appl. Environ. Microbiol. 58:2158-2163.

5. Arthur, E. L., B. S. Perkovich, T. A. Anderson, and J. R. Coats. 2000. Degradation of an atrazine and metolachlor herbicide mixture in pesticide- contaminated soils from two agrochemical dealerships in Iowa. Wat. Air Soil Poll. 119:75-90.

6. Behki, R., E. Topp, W. Dick, and P. Germon. 1993. Metabolism of the herbicide atrazine by Rhodococcus strains. Appl. Environ. Microbiol. 59:1955-1959.

7. Boundy-Mills, K. L., M. L. de Souza, R. T. Mandelbaum, L. P. Wackett, and M. J. Sadowsky. 1997. The atzB gene of Pseudomonas sp. strain ADP encodes the second enzyme of a novel atrazine degradation pathway. Appl. Environ. Microbiol. 63:916-923.

8. Briones, A. M. and W. Reichardt. 1999. Estimating microbial population counts by ‘most probable number’ using Microsoft Excel. J. Microbiol. Meth. 35:157-161.

190

9. Chung, K. H., K. S. Ro, and D. Roy. 1995. Atrazine biotransformation in wetland sediment under different nutrient conditions-I: Anaerobic. J. Environ. Sci. Health. A30:109-120.

10. Chung, K. H., K. S. Ro, and D. Roy. 1996. Fate and enhancement of atrazine biotransformation in anaerobic wetland sediment. Wat. Res. 30:341-346.

11. Crawford, J. J., G. K. Sims, R. L. Mulvaney, and M. Radosevich. 1998. Biodegradation of atrazine under denitrifying conditions. Appl. Microbiol. Biotechnol. 49:618-623.

12. DeLaune, R. D., I. Devai, C. Mulbah and C. W. Lindau. 1997. The influence of soil redox conditions on atrazine degradation in wetlands. Agric. Ecosyst. Environ. 66:41-46.

13. de Souza, M. L., L. P. Wackett, K. L. Boundy-Mills, R. T. Mandelbaum and M. J. Sadowsky. 1995. Cloning, characterization and expression of a gene region from Pseudomonas sp. strain ADP involved in the dechlorination of atrazine. Appl. Environ. Microbiol. 61:3373-3378.

14. de Souza, M. L., M. J. Sadowsky, and L. P. Wackett. 1996. Atrazine chlorohydrolase from Pseudomonas sp. strain ADP: gene sequence, enzyme purification, and protein characterization. J. Bacteriol. 178:4894-4900.

15. de Souza, M. L., J. Seffernick, B. Martinez, M. J. Sadowsky, and L. P. Wackett. 1998. The atrazine catabolism genes atzABC are widespread and highly conserved. J. Bacteriol. 180:1951-1954.

16. Donbek, T., E. Dolan, J. Schultz, and D. Klarup. 2001. Rapid reductive dechlorination of atrazine by zero-valent iron under acidic conditions. Environ. Poll. 111:21-27.

17. Donaldson, D., T. Kiely, and A. Grube. 2002. Pesticides industry sales and usage. U. S. Environmental Protection Agency. [Online.] http://www.epa.gov/oppbead1/pestsales/99pestsales/market_estimates1999.pdf. Accessed February 21, 2003.

18. Eaton, R. W. and J. S. Karns. 1991. Cloning and analysis of s-triazine catabolic genes from Pseudomonas sp. strain NRRLB-12227. J. Bacteriol. 173:1215-1222.

19. Eaton, R. W. and J. S. Karns. 1991. Cloning and comparison of the DNA encoding ammelide aminohydrolase and cyanuric acid amidohydrolase from three s-triazine-degrading bacterial strains. J. Bacteriol. 173:1363-1366.

191

20. Entry, J. A. 1999. Influence of nitrogen on atrazine and 2,4- dichlorophenoxyacetic acid mineralization in blackwater and redwater forested wetland soils. Biol. Fert. Soils. 29:348-353.

21. Gebendinger, N. and M. Radosevich. 1999. Inhibition of atrazine degradation by cyanazine and exogenous nitrogen in bacterial isolate M91-3. Appl. Microbiol. Biotechnol. 51:375-381.

22. Guerin, W. F. and S. A. Boyd. 1992. Differential bioavailability of soil-sorbed naphthalene to two bacterial species. Appl. Environ. Microbiol. 58:1142-1152.

23. Hayes, T. B., A. Collins, M. Lee, M. Mendoza, N. Noriega, A. A. Stuart, and A. Vonk. 2002. Hermaphroditic, demasculinized frogs after exposure to the herbicide atrazine at low ecologically relevant doses. Proc. Natl. Acad. Sci. USA 99:5476-5480.

24. Jayachandran, K., N. B. Stolpe, T. B. Moorman, and P. J. Shea. 1998. Application of 14C-most-probable-number technique to enumerate atrazine- degrading microorganisms in soil. Soil Biol. Biochem. 30:523-529.

25. Karns, J. S. 1999. Gene sequence and properties of an s-triazine ring-cleavage enzyme from Pseudomonas sp. strain NRRLB-12227. Appl. Environ. Microbiol. 65:3512-3517.

26. Kao, C. M., J. Y. Wang, and M. J. Wu. 2001. Evaluation of atrazine removal processes in a wetland. Wat. Sci. Technol. 44:539-544.

27. Kao, C. M., J. Y. Wang, K. F. Chen, H. Y. Lee, and M. J. Wu. 2002. Non-point source pesticide removal by a mountainous wetland. Wat. Sci. Technol. 46:199- 206.

28. Katz, I., C. G. Dosoretz, R. T. Mandelbaum, and M. Green. 2001. Atrazine degradation under denitrifying conditions in continuous culture of Pseudomonas ADP. Wat. Res. 35:3272-3275.

29. Kemerer, J. 2001. M.S. thesis. The Ohio State University, Columbus, OH.

30. Larsen, L., C. Jørgensen and J. Aamand. 2001. Potential mineralization of four herbicides in a ground water-fed wetland area. J. Environ. Qual. 30:24-30.

31. Mandelbaum, R. T., D. L. Allan, and L. P. Wackett. 1995. Isolation and characterization of a Pseudomonas sp. that mineralizes the s-triazine herbicide atrazine. Appl. Environ. Microbiol. 61:1451-1457.

32. Marchand, A., S. Piutti, B. Lagacherie, and G. Soulas. 2002. Atrazine mineralization in bulk soil and maize rhizosphere. Biol. Fertil. Soils. 35:288-292.

192

33. Martinez, B., J. Tomkins, L. P. Wackett, R. Wing, and M. J. Sadowsky. 2001. Complete nucleotide sequence and organization of the atrazine catabolic plasmid pADP-1 from Pseudomonas sp. strain ADP. J. Bacteriol. 183:5684-5697.

34. Moore, M. T., J. H. Rodgers Jr., C. M. Cooper, and S. Smith, Jr. 2000. Constructed wetlands for mitigation of atrazine-associated agricultural runoff. Environ. Poll. 110:393-399.

35. Moorman, T. B., J. K. Cowan, E. L. Arthur, and J. R. Coats. 2001. Organic amendments to enhance herbicide biodegradation in contaminated soils. Biol. Fertil. Soils. 33:541-545.

36. Mulbry, W. W., H. Zhu, S. M. Nour, and E. Topp. 2002. The triazine hydrolase gene trzN from Nocardiodes sp. strain C190: cloning and construction of gene- specific primers. FEMS Microbiol. Lett. 206:75-79.

37. Nagy, I., S. Verheijen, A. De Schrijver, J. Van Damme, P. Proost, G. Schoofs, J. Vanderleyden, and R. De Mot. 1995. Characterization of the Rhodococcus sp. NI86/21 gene encoding alcohol: N,N'-dimethyl-4-nitrosoaniline oxidoreductase inducible by atrazine and thiocarbamate herbicides. Arch. Microbiol. 163:439- 446.

38. Nagy, I., F. Compernolle, K. Ghys, J. Vanderleyden, and R. De Mot. 1995. A single cytochrome P-450 system is involved in the degradation of the herbicides EPTC (s-ethyl dipropylthiocarbamate) and atrazine by Rhodococcus sp. strain NI86/21. Appl. Eviron. Microbiol. 61:2056-2060.

39. Ostrofsky, E. B., S. J. Traina and O. H. Tuovinen. 1997. Variation in atrazine mineralization rates in relation to agricultural management practice. J. Environ. Qual. 26:647-657.

40. Ostrofsky, E. B., J. B. Robinson, S. J. Traina and O. H. Tuovinen. 2001. Effect of cyanuric acid amendment on atrazine mineralization in surface soils and detection of the s-triazine ring-cleavage gene trzD. Soil Biol. Biochem. 33:1539- 1545.

41. Perkovich, B. S., T. A. Anderson, E. L. Kruger, and J. R. Coats. 1996. Enhanced mineralization of [14C]Atrazine in Kochia scoparia rhizospheric soil from a pesticide-contaminated site.

42. Piutti, S., S. Hallet, S. Rousseaux, L. Philippot, G. Soulas, and F. Martin-Laurent. 2002. Accelerated mineralization of atrazine in maize rhizosphere soil. Biol. Fertil. Soils. 36:434-441.

193

43. Radosevich, M., S. J. Traina, and O. H. Tuovinen. 1996. Biodegradation of atrazine in surface soils and subsurface sediments collected from an agricultural research farm. Biodegradation 7:137-149.

44. Radosevich, M., S. J. Traina, Y. Hao, and O. H. Tuovinen. 1995. Degradation and mineralization of atrazine by a soil bacterial isolate. Appl. Environ. Microbiol. 61:297-302.

45. Ro, K. S. and K. H. Chung. 1995. Atrazine biotransformation in wetland sediment under different nutrient conditions-II: Aerobic. J. Environ. Sci. Health. A30:121- 131.

46. Rousseaux, S., A. Hartmann, and G. Soulas. 2001. Isolation and characterization of new gram-negative and gram-positive atrazine degrading bacteria from different French soils. FEMS Microbiol. Ecol. 36:211-222.

47. Runes, H. B., J. J. Jenkins and P. J. Bottomley. 2001. Atrazine degradation by bioaugmented sediment from constructed wetlands. Appl. Microbiol. Biotechnol. 57:427-432.

48. Runes, H. B., J. J. Jenkins, J. A. Moore, P. J. Bottomley, and B. D. Wilson. 2003. Treatment of atrazine in nursery irrigation runoff by a constructed wetland. Wat. Res. 37:539-550.

49. Sadowsky, M. J., Z. Tong, M. de Souza, and L. P. Wackett. 1998. AtzC is a new member of the amidohydrolase protein superfamily and is homologous to other atrazine-metabolizing enzymes. J. Bacteriol. 180:152-158.

50. Sasser, M. 1997. Identification of bacteria by gas chromatography of cellular fatty acids. Technical Note #101, Microbial Identification, Inc., Newark, DE.

51. Seffernick, J. L., H. McTavish, J. P. Osborne, M. L. de Souza, M. J. Sadowsky, and L. P. Wackett. 2002. Atrazine chlorohydrolase from Pseudomonas sp. strain ADP is a metalloenzyme. Biochemistry. 41:14430-14337.

52. Seybold, C. A., W. Mersie, and C. McNamee. 2001. Anaerobic degradation of atrazine and metolachlor and metabolite formation in wetland soil and water microcosms. J. Environ. Qual. 30:1271-1277.

53. Shao, Z. Q. and R. Behki. 1995. Cloning of the genes for degradation of the herbicides EPTC (S-ethyl dipropylthiocarbamate) and atrazine from Rhodococcus sp. strain TE1. Appl. Environ. Microbiol. 61:2061-2065.

54. Shao, Z. Q. and R. Behki. 1996. Characterization of the expression of the thcB gene, coding for a pesticide-degrading cytochrome P-450 in Rhodococcus strains. Appl. Environ. Microbiol. 62:403-407.

194

55. Shapir, N., S. Goux, R. T. Mandelbaum, and L. Pussemier. 2000. The potential of soil microorganisms to mineralize atrazine as predicted by MCH-PCR followed by nested PCR. Can. J. Microbiol. 46:425-432.

56. Shapir, N., R. T. Mandelbaum and P. Fine. 2000. Atrazine mineralization by indigenous and introduced Pseudomonas sp. strain ADP in sand irrigated with municipal wastewater and amended with composted sludge. Soil Biol. Biochem. 32:887-897.

57. Shapir, N., J. P. Osborne, G. Johnson, M. J. Sadowsky, and L. P. Wackett. 2002. Purification, substrate range, and metal center of AtzC: the N-isopropylammelide aminohydrolase involved in bacterial atrazine metabolism. J. Bacteriol. 184:5376- 5384.

58. Solomon, K. R., D. B. Baker, R. P. Richards, K. R. Dixon, S. J. Klaine, T. W. La Point, R. J. Kendall, C. P. Weisskopf, J. M. Giddings, J. P. Giesy, L. W. Hall, Jr., and W. M. Williams. 1996. Ecological risk assessment of atrazine in North American surface waters. Environ. Toxicol. Chem. 15:31-76.

59. Soulas, G. and B. Lagacherie. 2001. Modelling of microbial degradation of pesticides in soils. Biol. Fertil. Soils. 33:551-557.

60. Stamper, D. M., M. Radosevich, K. B. Hallberg, S. J. Traina, and O. H. Tuovinen. 2002. Ralstonia basilensis M91-3, a denitrifying soil bacterium capable of using s-triazines as nitrogen sources. Can. J. Microbiol. 48:1089-1098.

61. Stead, D. E., J. E. Sellwood, J. Wilson, and I. Viney. 1992. Evaluation of a commercial microbial identification system based on fatty acid profiles for rapid accurate identification of plant pathogenic bacteria. J. Appl. Bacteriol. 72:315- 321.

62. Strong, L. C., C. Rosendahl, G. Johnson, M. J. Sadowsky, and L. P. Wackett. 2002. Arthrobacter aurescens TC1 metabolizes diverse s-triazine ring compounds. Appl. Environ. Microbiol. 68:5973-5980.

63. Surzycki, S. J., J. A. Surzycki and R. Wagner. 1997. Techniques in Molecular Biology Workshop: DNA Sequencing, PCR, and RFLP Analysis. Laboratory Manual, University of Indiana, Bloomington, IN.

64. Topp, E., H. Zhu, S. M. Nour, S. Houot, M. Lewis, and D. Cuppels. 2000. Characterization of an atrazine-degrading Pseudaminobacter sp. isolated from Canadian and French agricultural soils. Appl. Environ. Microbiol. 66:2773-2782.

195

65. Topp, E., W. M. Mulbry, H. Zhu, S. M. Nour, and D. Cuppels. 2000. Characterization of s-triazine herbicide metabolism by a Nocardiodes sp. isolated from agricultural soils. Appl. Environ. Microbiol. 66:3134-3141.

66. Vanderheyden, V., P. Debongnie, and L. Pussemier. 1997. Accelerated degradation and mineralization of atrazine in surface and subsurface soil materials. Pestic. Sci. 49:237-242.

67. Yassir, A., B. Lagacherie, S. Houot, and G. Soulas. 1999. Microbial aspects of atrazine biodegradation in relation to history of soil treatment. Pestic. Sci. 55:799-809.

196