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Characterization and Activity Comparisons of Methanotrophic-Heterotrophic Mixed Cultures Derived from a Landfill Environment

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Title:
Characterization and Activity Comparisons of Methanotrophic-Heterotrophic Mixed Cultures Derived from a Landfill Environment
Creator:
STRATE, JESSICA LORENE ( Author, Primary )
Copyright Date:
2008

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Subjects / Keywords:
Bacteria ( jstor )
Flasks ( jstor )
Heterotrophs ( jstor )
Landfills ( jstor )
Methane ( jstor )
Methylococcaceae ( jstor )
Microbiology ( jstor )
Nutrients ( jstor )
Oxidation ( jstor )
Oxygen ( jstor )

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University of Florida
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University of Florida
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Copyright Jessica Lorene Strate. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
Embargo Date:
5/31/2006
Resource Identifier:
436098720 ( OCLC )

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CHARACTERIZATION AND ACTIVITY COMPARISONS OF METHANOTROPHIC-HETEROTROPHIC MI XED CULTURES DERIVED FROM A LANDFILL ENVIRONMENT By JESSICA LORENE STRATE A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF ENGINEERING UNIVERSITY OF FLORIDA 2005

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Copyright 2005 by Jessica Lorene Strate

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Dedicated to Mary O’Conner Strate.

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ACKNOWLEDGMENTS Many individuals made this master’s thesis possible. First, I would like to thank my supervising committee chair (Dr. Angela Lindner) for her patience, support, and guidance me since I began working with her as an undergraduate 5 years ago. She inspired and motivated me to excel through work hard. I would like to thank the other members of my committee (Dr. Madeline Rasche and Dr. Andrew Ogram) who also guided me. They provided valuable input on our study and graciously allowed me to use their laboratory or facilities to conduct research. I also want to recognize the American Association for University Women who provided financial support to me through the Selected Professions Fellowship (2002-2003) and do so for many women in academics. Special thanks go to Donna Williams and Dr. Henry Aldrich in the Electron Microscopy Lab (Microbiology and Cell Science Department), who graciously gave their time and energy in conducting the scanning electron microscopy work, in addition to their feedback and input in finalizing that work. Special thanks also go to Dr. Begonja Hrsak and Dasa Filipic (Center for Marine and Environmental Research, Zagreb, Croatia) who agreed to help us out, and worked with the samples in our study. I also thank, Dr. Rocco Mancinelli (NASAAmes Research Center, Moffett Field, CA) who kindly donated several journal papers he had written, to assist me with our study. I thank all of the members of the Bioremediation research group (Trisha Howard, Chance Lauderdale, Elizabeth O’Brien, Adriana Pacheco, and Callie Whitfield Babbitt); several students or members of the Environmental Engineering Department (Dr. Matthew Booth, Jennifer iv

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Hobbs McElroy, Terese Gregg, Ruben Kertecz, Christina Ludwig, and Marnie Ward); and other laboratories (Bev Driver) who provided technical or emotional support. I also want to recognize those who have given me special love and support throughout the years, including my parents; family members; friends; and especially my wonderful, supportive, and loving husband, Brad Beach. A final acknowledgement goes to God and all of His glory, because it was through His grace and mercy that I have made it through Our study and could not have done so without relying on His strength. v

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TABLE OF CONTENTS Page ACKNOWLEDGMENTS.................................................................................................iv LIST OF TABLES.............................................................................................................ix LIST OF FIGURES.............................................................................................................x ABSTRACT......................................................................................................................xii CHAPTER 1 INTRODUCTION........................................................................................................1 2 LITERATURE REVIEW.............................................................................................3 Greenhouse Gases and Their Relevancy to Landfills...................................................3 Methane.................................................................................................................3 Methane Sources...................................................................................................4 Methane Sinks.......................................................................................................5 Landfill Emissions of Methane.............................................................................6 An Introduction to Landfills and Landfill Microbiology.............................................8 Landfill Environments...........................................................................................8 Description of a Sanitary Landfill.........................................................................8 Common problems encountered with sanitary landfills.................................9 The need for natural attenuation of landfill environments in Florida..........11 Landfill Microbiology.........................................................................................11 Biological decomposition processes............................................................11 Factors influencing methane oxidation in landfills......................................14 Introduction to methanotrophs: methane-oxidizing bacteria.......................17 Brief introduction to heterotrophic bacteria.................................................26 Significance and Rationale of Project.........................................................................29 3 MATERIALS AND METHODS...............................................................................35 Overview of Materials and Methods..........................................................................35 Stage 1: Isolation of Mixed Cultures from Landfill Samples.....................................35 Sampling Sites.....................................................................................................35 Enrichment and Isolation of Mixed Cultures......................................................36 vi

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Stages 2 and 3: Characterization of Mixed Cultures..................................................37 Heterotroph Isolation...........................................................................................37 Morphological and Phenotypic Characterization................................................37 Cellular Characterization.....................................................................................39 Tentative Identification of Heterotrophs.............................................................39 Growth Studies....................................................................................................40 Determination of growth rates......................................................................40 Analysis of protein content..........................................................................41 sMMO assay.................................................................................................41 Assessment of methane-oxidizers and heterotrophs....................................42 Determination of Methane Oxidation Rates........................................................43 Evaluation of Oxidation Potential.......................................................................45 Stage 4: Assessment of Response to Environmental Stressors..................................47 Initial Methane Concentration.............................................................................47 Temperature.........................................................................................................48 The pH Values.....................................................................................................49 Copper Concentration and Media Strength.........................................................49 4 CHARACTERIZATION AND ACTIVITY COMPARISONS OF METHANOTROPHIC-HETEROTROPHIC MIXED CULTURES DERIVED FROM A LANDFILL ENVIRONMENT..................................................................54 Introduction.................................................................................................................54 Materials and Methods...............................................................................................56 Enrichment and Isolation of Mixed Cultures......................................................56 Morphological and phenotypic studies........................................................57 Scanning electron microscopy.....................................................................58 Growth kinetics and methane oxidation rate parameters.............................58 Degradation potential in the presence of landfill contaminants...................60 Effects of Environmental Stressors.....................................................................62 Results.........................................................................................................................64 Isolation of Mixed Cultures.................................................................................64 Characteristics of the Enriched, Mixed Methanotrophic-Heterotrophic Communities....................................................................................................64 Morphological and phenotypic characteristics.............................................64 Cellular characteristics.................................................................................65 Evaluation of activity based on growth and methane oxidation rate...........66 Degradation potential for landfill contaminants based on oxygen uptake analysis....................................................................................................66 Assessment of Communities’ Response to Environmental Stressors.................67 vii

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Discussion...................................................................................................................69 Characterization of the Enriched, Mixed Methanotrophic-Heterotrophic Communities....................................................................................................69 Morphological and phenotypic characteristics of isolated heterotrophs......69 Cellular characteristics.................................................................................69 Degradation potential of landfill contaminants based on oxygen uptake analysis....................................................................................................70 Assessment of Communities’ Response to Environmental Stressors.................71 Conclusions.................................................................................................................73 5 CONCLUSIONS........................................................................................................83 APPENDIX A CALCULATED CONVERSIONS FOR METHANE OXIDATION RATES FROM PREVIOUS PUBLICATIONS.......................................................................85 B GROWTH MEDIA USED FOR ENRICHMENT.....................................................87 C METHANOTROPHIC ISOLATION PROCESS.......................................................88 D SUPPLEMENTARY HETEROTROPH INFORMATION.......................................94 E EXAMPLES OF LABORATORY EQUIPMENT USED.........................................97 F EXAMPLES OF CALCULATIONS: GROWTH, METHANE DEPLETION AND OXYGEN UPTAKE.........................................................................................98 G SUPPLEMENTARY INFORMATION FOR GROWTH AND METHANE DEPLETION EXPERIMENTS................................................................................105 LIST OF REFERENCES.................................................................................................107 BIOGRAPHICAL SKETCH...........................................................................................117 viii

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LIST OF TABLES Table page 2-1 Characteristics of type I, II, and X methanotrophs..................................................31 4-1 Characteristics of isolated heterotrophs on selected media......................................75 4-2 Comparison of TCE and benzene oxidation potential for mixed cultures...............77 A-1 Summary of methane oxidation rates and their converted units..............................85 B-1 Nitrate mineral salts (NMS) media recipe................................................................87 B-2 Whittenbury trace elements......................................................................................87 B-3 Phosphate stock solution..........................................................................................87 B-4 Vitamin stock solution.............................................................................................87 D-1 Phenotype characteristics of heterotroph populations..............................................96 G-1 Growth and methane depletion by landfill samples at varied methane concentrations.........................................................................................................105 G-2 Growth and methane depletion by landfill samples at varied temperature and pH.................................................................................................106 ix

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LIST OF FIGURES Figure page 2-1 Basic landfill components........................................................................................32 2-2 Methane oxidation pathway in methanotrophs........................................................33 2-3 Aerobic methanotrophic degradation of TCE..........................................................33 2-4 Microbial oxidation of benzene utilizing monooxygenase......................................34 2-5 Microbial oxidation of benzene utilizing dioxygenase............................................34 3-1 Flowchart of methodology used...............................................................................50 3-2 Alachua County Landfill site map...........................................................................51 3-3 Side-arm flask used in determining growth rates.....................................................51 3-4 Mininert system used in methane degradation experiments....................................52 3-5 Oxygen uptake system.............................................................................................53 4-1 Cellular diversity of mixed methanotrophic-heterotrophic communities................76 4-2 Effect of methane concentration on growth rates of mixed cultures........................78 4-3 Effect of methane concentration on methane oxidation rates of mixed cultures. .......................................................................................................78 4-4 Effect of temperature on growth rates of mixed cultures.........................................79 4-5 Effect of temperature on methane oxidation rates of mixed cultures.......................79 4-6 Effect of pH on growth rates of mixed cultures.......................................................80 4-7 Effect of pH on methane oxidation rates of mixed cultures.....................................80 4-8 Effect of copper on growth rates of mixed cultures.................................................81 4-9 Effect of copper on methane oxidation rates of mixed cultures...............................81 x

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4-10 Effect of medium strength on growth rates of mixed cultures grown without the presence of copper.............................................................................................82 4-11 Effect of medium strength on growth rates of mixed cultures grown with the presence of copper....................................................................................................82 C-1 Technique using both solid and liquid media for methanotroph isolation...............93 E-1 Laboratory equipment..............................................................................................97 F-1 Growth curve used in calculating growth rates........................................................98 F-2 Linear regression fit of growth curve data in calculating growth rates....................98 F-3 Sample of spreadsheet used for growth rate calculations based on linear regression curves......................................................................................................99 F-4 Measured methane depletion for mixed culture GW70...........................................99 F-5 Linear regression of data based on methane depletion curves...............................100 F-6 Spreadsheet used for specific methane oxidation rate calculations.......................101 F-7 Spreadsheet used in oxygen uptake calculations....................................................102 F-8 Measured oxygen consumption in the presence of methane, showing inhibition of acetylene during oxygen uptake analysis..........................................103 F-9 Measurement of oxygen consumption in the presence of 1,4 dioxane and NMS medium (a control) and then in the presence of cells...................................103 F-10 Measurement of oxygen consumption in the presence of benzene plus NMS (a control).....................................................................................................104 F-11 Measurement of oxygen consumption in the presence of TCE plus NMS (a control).....................................................................................................104 xi

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Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Engineering CHARACTERIZATION AND ACTIVITY COMPARISONS OF METHANOTROPHIC-HETEROTROPHIC MIXED CULTURES DERIVED FROM A LANDFILL ENVIRONMENT By Jessica Lorene Strate May 2005 Chair: Angela S. Lindner Major Department: Environmental Engineering Sciences Landfills are nutrient-rich environments, providing complex organic compounds to microorganisms for substrate use, thus promoting microbial diversity. Because of the rapid increase in population and the associated disposal of waste in landfills, there is a critical need to better understand the fundamental microbiology of landfills to enable better management methods for rapid degradation of this waste. Our study focused on methane-oxidizing bacteria (methanotrophs), which exist where methane and oxygen are present. Methanotrophs are considered to be ecologically significant because of their ability to oxidize methane (a greenhouse gas) and to co-metabolize various chlorinated aliphatic and aromatic compounds. Our aim was to better understand differences in isolated landfill mixed cultures through characterization of two mixed cultures and activity comparisons (primarily based on rates of growth and methane oxidation), thus providing a foundation for a broader microbial landfill-laboratory study in the future. This was initiated by enrichment of two xii

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stable, mixed methanotrophic-heterotrophic cultures from municipal solid waste (MSW) landfill samples derived from regions of the Alachua County Landfill that differed in age by approximately 5 years. These cultures were characterized using fundamental microbiology techniques. We evaluated their activity based on growth, methane depletion, and oxygen uptake studies. Both cultures contained closely associated methanotroph and heterotroph populations that differ in phenotype and that showed different activity responses to changes in temperature; pH; and concentrations of benzene, trichloroethylene (TCE), methane, copper, and strength of medium. Our study was an initial effort to ultimately enable direct linking of the field to the laboratory by isolating and studying two unique and diverse, mixed methanotrophic-heterotrophic cultures that show potential for contaminant degradation; and that are capable of responding and adapting to extreme changes in environmental conditions based on growth and methane oxidation. Our study addresses the need to link methanotrophic diversity and activity observed in the field with that which is more easily observed in laboratory conditions. The ultimate goal of our study is more rapid, facile prediction of biodegradative activity of microbial populations in landfills, thus allowing better management of these systems. xiii

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CHAPTER 1 INTRODUCTION The growing number of landfills in the United States has created a need for more efficient waste degradation, prevention of methane emissions to the atmosphere, and minimization of releases of toxic compounds. The role of methanotrophs in these processes is not well understood. Because of the difficulty in studying methanotrophic populations directly in the field, there is great need to develop protocols to allow study of field processes in the laboratory. To date, a direct link between methanotrophic diversity and activity in field samples with mixed cultures isolated from field samples has not been made. Doing so would enable more rapid, facile prediction of biodegradative activity of microbial populations in landfills, thus allowing better management of these systems. Our study took initial steps in bridging the gap between field and laboratory studies by isolating mixed methanotrophic-heterotrophic cultures from landfill samples. Overall goals of our study were two-fold: 1) to isolate stable mixed methanotrophic-heterotrophic cultures from the initial soil samples derived from a landfill; and 2) to study these stable, enriched, mixed cultures through characterization and activity experiments, to gain further insight into how these tightly-knit communities behave. After a literature review was conducted to evaluate studies that have involved the study of methanotrophs in landfill environments and of mixed methanotrophic-heterotrophic cultures from a variety of environments, the remaining scope of our study consisted of three major objectives: Objective 1: isolate and phenotypically characterize stable mixed methanotrophic-heterotrophic cultures enriched from landfill soil samples under a single set of enrichment conditions. 1

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2 Objective 2: assess both mixed cultures for activity based on growth, methane depletion and their potential for contaminant degradation. Objective 3: evaluate the response of these mixed cultures to changes in environmental conditions. Our study was intended to serve as a first basis for a broader, more in-depth study of microbial isolates derived from landfill environments. Ultimately, we hope that this and future studies will provide a better understanding of the complex processes that occur in landfill environments; thus enabling more efficient management of these environments, to minimize risk to the environment and public health.

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CHAPTER 2 LITERATURE REVIEW Greenhouse Gases and Their Relevancy to Landfills A greenhouse gas is capable of absorbing the infrared radiation that leads to an accumulation of heat into our atmosphere (Borjesson, 1997). This absorbed (or accumulated) radiation creates an imbalance in the atmosphere, which determines the earth’s climate. As increasing concentrations of greenhouse gases are released into the atmosphere, it is probable that global climate will change due to that increase (Borjesson, 1997). Contributing greenhouse gases include carbon dioxide (CO 2 ), methane (CH 4 ), nitrous oxide (N 2 O), ozone (O 3 ), and chlorofluorocarbons (CFCs). Carbon dioxide and methane were reported to have the highest concentration (358,000 and 1720 ppbv, respectively) of these gases in 1994 (Borjesson, 1997). Sources of carbon dioxide include anthropogenic activity (such as increasing intensive land use, through deforestation; and the use of fossil fuels). However, sources of methane are of growing interest, because methane absorbs infrared irradiation more effectively than carbon dioxide; and its contribution to global warming is estimated to be 26 times greater than that of carbon dioxide (mole for mole) (Hanson and Hanson, 1996). To this end, we reviewed sources and sinks of methane. Methane Methane is one of the most important and abundant greenhouse gases in the atmosphere, with its concentration increasing dramatically over the past 200 years (Mancinelli, 1995; Hanson and Hanson, 1996; Borjesson, 1997; Henckel et al., 2000). 3

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4 Within the last 10 to 15 years, methane concentrations in the atmosphere have increased annually by 1% (Whalen et al., 1990; Kightley et al., 1995; Mosher et al., 1999). Methane plays an active role in atmospheric chemistry and is involved in changes in the chemical composition of the atmosphere (Le Mer and Roger, 2001) through its involvement in photochemical reactions determining concentrations of ozone and hydroxyl radical ( • OH) (Crutzen, 1991). Potential consequences of the increase in methane concentrations and its role in atmospheric chemistry include an increase in tropospheric ozone and a decrease in hydroxyl radical concentration, thus potentially reducing the oxidizing capabilities of the atmosphere (Mosher et al., 1999; Le Mer and Roger, 2001). The residence times of many volatile organic carbon (VOC) compounds (Mosher et al., 1999) and CFCs are predicted to increase because of the atmosphere’s reduced oxidation capacity. As a result, the concentration of greenhouse gases such as ozone, CO, and CO 2 (Le Mer and Roger, 2001) is expected to increase. Because of methane’s critical role in the atmosphere, considerable research has been devoted to investigating its production and consumption. Methane Sources Methane is produced by both natural and human sources. Natural sources include production through the anaerobic decay of organic matter by methanogenic bacteria (Mancinelli, 1995; Le Mer and Roger, 2001) in environments including wetlands, rice paddies, the rumen of cattle, and landfills (Crutzen, 1991). Between 100 and 200 teragrams (Tg) of methane are released into the atmosphere annually from wetlands, representing the largest natural source of methane (Whalen and Reeburgh, 2000; Le Mer and Roger, 2001), Rice paddies, the rumen of cattle, and landfills contribute 20 to 130 Tg per year (Crutzen, 1991; Le Mer and Roger, 2001).

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5 Human activities (such as biomass burning and leaks in natural-gas distribution systems, in addition to leaks in oil and gas wells and landfills) have been increasingly linked to atmospheric methane (Crutzen, 1991; Le Mer and Roger, 2001). It is estimated that biomass burning releases 15 to 45 Tg of methane per year. Leakage in natural-gas distribution systems and wells contribute 55 to 85 Tg yr -1 and landfills release 9 to 70 Tg yr -1 (Crutzen, 1991; Bogner and Spokas, 1993; Bogner et al., 1997a; Boeckx et al., 1996; Christophersen et al., 2000). To stabilize atmospheric methane concentrations, it has been estimated that anthropogenic sources must be reduced by 15 to 20 % (Crutzen, 1991). The most practical way to achieve this goal would be to limit emissions from landfills and the fossil fuel sector (Crutzen, 1991). Methane Sinks The main sink for atmospheric methane is its reaction in the troposphere, through oxidation with the hydroxyl radical (Crutzen, 1991; Le Mer and Roger, 2001). It also reacts in the stratosphere with chlorine generated from CFCs (Le Mer and Roger, 2001). Methane is also eliminated from some of the most important sources (wetlands, rice paddies and landfills) by microbial oxidation (King, 1992; Le Mer and Roger, 2001). As an example, in rice fields, more than 90% of the methane produced anaerobically can be re-oxidized by methanotrophs in the aerobic zones (Le Mer and Roger, 2001). Because of the important role that microorganisms play in mitigating methane release into the atmosphere (Higgins et al., 1980), further research has investigated microbial oxidation of methane in a variety of environments (Whalen et al., 1990; King 1992; Mancinelli, 1995; Whalen and Reeburgh, 2000; Le Mer and Roger, 2001). The microorganisms responsible for this oxidation of methane are known as methanotrophs and are ubiquitous in nature wherever there is a stable source of methane and air (Graham et al., 2002).

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6 Since methanotrophs are an important part of methane oxidation, these microorganisms are discussed in further detail (specifically, in the context of their role in landfill environments). Landfill Emissions of Methane On a global scale, estimates for methane emissions from landfills range between 9 and 70 Tg yr -1 , with the most cited estimate of 40 Tg yr -1 (Bogner and Spokas, 1993; Bogner et al., 1997a; Boeckx et al., 1996; Christophersen et al., 2000). This accounts for 3 to 19% of anthropogenic methane emissions globally, strongly implicating its contribution to global warming in greenhouse gas scenarios (Boeckx et al., 1996; Czepiel et al., 1996; Bogner et al., 1997b; Borjesson et al., 1998; Schuetz et al; 2003). Together, Europe and North America contribute about half of the world’s estimated methane emissions from landfills (Boeckx et al., 1996). In the United States alone, landfills represent the largest anthropogenic source of methane emissions, emitting approximately 37% of total emissions in 1997 (Bogner et al., 1997b; Mosher et al., 1999; US EPA, 1999). It has been suggested that improved landfill management practices could considerably reduce methane emissions (Crutzen, 1991; Borjesson, 1998). One proposed mitigating option is through use of landfill gas extraction and recovery systems. This is very effective at large landfills with high methane generation and is generally regarded as being a superior choice because of the decline in gas recovery efficiency at older and/or smaller landfills with low methane contents (Boeckx et al., 1996; Chrisophersen et al., 2000). However, there are several drawbacks of gas recovery methods, namely only 40 to 60% recovery of the generated landfill gas, high cost and intricacy of design, and limited service life (Humer and Lechner, 1999). The second, more cost-effective option

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7 is enhanced microbial oxidation of methane in landfill cover soils (Borjesson, 1998; Humer and Lechner, 1999; Schuetz and Kjeldsen, 2003; Schuetz et al., 2003). This manipulation of cover soils can possibly provide a complimentary strategy for methane emission control and would be more effective and practical at smaller and older landfill sites where less methane is generated and recovered (Kightley et al., 1995; Boeckx et al., 1996; Bogner et al., 1997a; Christophersen, 2000). The potential for mitigation of methane emissions from landfills through biological oxidation will be discussed further. As landfill gas is transported up through soil layers, it mixes with air where methanotrophs can oxidize the methane present in the landfill gas (Christophersen et al., 2000). It has been shown in laboratory studies that landfill cover soils exposed to elevated methane concentrations are capable of demonstrating a high capacity for methane oxidation by methanotrophs (Christophersen et al., 2000; Schuetz et al., 2003). It has also been reported that, in spite of the large amount of methane generated, methanotrophic oxidation has a strong mitigating effect by reducing landfill methane emissions anywhere from 7 to 50% (Mancinelli and McKay, 1985; Whalen et al., 1990; King, 1992; Jones and Nedwell, 1993; Kightley et al., 1995; Boeckx and Van Cleemput, 1996; Czepiel et al., 1996; Bogner et al., 1997b; Borjesson, 1998; De Visscher et al., 1999). As an example, based on both field and laboratory measurements, methane emissions from a small covered landfill site were compared for their gross and net methane emission rates and it was found that the cover soil, because of its methane oxidizing capacity, had a large mitigating effect on the methane emission (Boeckx et al., 1996). Given this large potential for reduction of methane emitted from landfill cover soils, it has been suggested that further investigation of the community structure in

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8 landfill environments may assist in management practices for the attenuation of methane emissions (Wise et al., 1999). This requires a better understanding of the links between landfill environment and landfill microbiology, specifically addressing how various groups of microorganisms interact, what roles they play and how they respond to varying conditions in landfills. An Introduction to Landfills and Landfill Microbiology Landfill Environments Municipal solid waste (MSW) is most commonly known as “trash” or “garbage” and typically consists of grass clippings, packaging, furniture, clothing, bottles, food waste, newspapers, appliances and batteries (US EPA, 2002). Generation of this waste (or refuse) has steadily increased over the past 35 years, with roughly 55% by weight of refuse generated in the United States was disposed of into sanitary landfills regardless of recycling and composting efforts (Hilger and Barlaz, 2002; US EPA, 2002). In the year 2000, 231.9 million tons of MSW was generated with only 30% of the materials being recovered (US EPA, 2002). Description of a Sanitary Landfill Over the years, sanitary landfills have become highly engineered systems that are designed for several purposes, Contain waste and separate it from the environment, Capture leachate (i.e., water that percolates through the landfill, picking up contaminants) Control gas migration (Hilger and Barlaz, 2002). In selecting a landfill site, the size of a landfill site requires considerable area, ranging from 4.5 to 46.5 hectares (Whalen et al., 1990; Czepiel et al., 1996; Mosher et al., 1999).

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9 Also, the natural hydro-geologic setting is important because it can prevent waste from escaping and enable accurate prediction of waste fate in case leakage does occur. There are several components of a landfill that enable them to fulfill their design purposes. Hilger and Barlaz (2002) give a good description of these basic components in a landfill, which are briefly reviewed here. Figure 2-1 gives an overview of basic landfill components, including a liner system, leachate collection system, drainage layer, and a cover layer. The liner system acts as a protective barrier to minimize leachate migration to the groundwater. Such systems typically consist of a thick clay layer overlaid with a durable, puncture-resistant flexible membrane liner. A drainage layer is placed above the liner to promote collection of leachate. On top of this leachate collection layer, another barrier (such as additional soil, baled waste, tire chips) is installed to protect the system from equipment used to place and compact refuse. Refuse is then placed on top of the protective barrier, where it is typically covered daily to reduce wind-blown waste, contamination of storm water runoff, and the attraction of rodents to the site. Once refuse has reached its maximum design height, a final cover is applied. The final cover includes a layer of low-permeability soil that promotes vegetative growth, designed to minimize storm water infiltration and erosion of the soil cover and to promote evapotranspiration. The major terminal products of MSW decomposition, methane and carbon dioxide (which will be addressed later), are typically released through the cover and collected in wells placed in the refuse to minimize off-site migration. The gas may be vented, flared, or, in some cases, recovered for its energy value. Common problems encountered with sanitary landfills Although the methane generated within sanitary landfills can be in some cases recovered for energy use, it also has been implicated in hazards, such as fires, explosions,

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10 and deaths (Mancinelli et al., 1981; Mancinelli and McKay, 1985). This landfill gas is composed primarily of methane (50 to 60%), but it also contains carbon dioxide and trace gases, such as nitrogen and non-methane hydrocarbons (Czepiel et al., 1996), including aromatics, halogenated hydrocarbons, and organic sulfur compounds (Schuetz and Kjeldsen, 2003; Schuetz et al., 2003). Although the trace components make up a small portion of typical landfill gas, they may exert an unbalanced environmental burden (Schuetz et al., 2003). Of particular concern, carcinogenic emissions of benzene may pose a health threat to workers and local inhabitants (Kjeldsen et al., 1997; Schuetz et al., 2003). Exposure pathways for benzene, a known carcinogen that targets the blood-making system, include inhalation and digestion (Cheremisinoff, 1979). Landfill gas also contains benzene at an average concentration of 2,060 ppbv (Tammemagi, 1999). Another toxic chemical of concern is the commonly found groundwater contaminant, trichloroethylene (TCE). TCE is a suspected carcinogen targeting the central nervous system, kidneys and liver with exposure pathways including inhalation and ingestion (U.S. EPA, 2001). TCE is also found in landfill gas at an average concentration of 2,080 ppbv (Tammemagi, 1999). Another serious environmental and health problem associated with landfills is due to liner system failure and groundwater contamination by leachate. The liner system previously described is under stringent regulations at the state and national level through the implementation of Subtitle D of the Resource Conservation and Recovery Act (Hilger and Barlaz, 2002). Unfortunately, no liner is 100% effective and can fail due to stress cracks, fractures and degradation caused by interactions with household chemicals. When failure occurs, leachate can migrate through the hydro-geologic setting and

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11 infiltrate into groundwater. As the leachate percolates through the landfill, it can accumulate organic and inorganic chemicals, metals and other waste. As a result of the concern for groundwater contamination from leachate, regulatory constraints have enforced minimizing infiltration and keeping moisture content low with a net effect of retarding microbial decomposition (Bogner and Spokas, 1993). The need for natural attenuation of landfill environments in Florida Florida has the fourth largest population in the United States with an average 5-year population growth rate of 2.6% (Intelligence Center, 2004). At this projected growth rate, Florida’s population will exceed other state’s population growth over the next 2 decades (Intelligence Center, 2004). Based on the amount of waste generated per person per day of 4.5 pounds (US EPA, 2002), this will result in a tremendous increase in the amount of MSW and hazardous chemicals disposed of in landfills. With the greater waste inputs into landfills in Florida, an urgent need exists to control these environments for both rapid waste degradation and hazardous chemical attenuation to reduce the risk of exposure to harmful compounds, such as benzene and trichloroethylene, and also contamination to drinking water sources through liner failures. One proposed method for landfill control is the utilization of enhanced biological processes to reduce emissions. This is a complex process, requiring a coordinated effort of several trophic groups of bacteria which are addressed in further detail. Landfill Microbiology Biological decomposition processes The previous description of a landfill environment and the type of material disposed of enables, landfills to be extremely variable and heterogeneous as evidenced by its reported diversity in waste composition (Senior, 1995). The physiochemical

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12 properties (such as temperature and pH) vary dramatically both spatially and temporally (Senior, 1995). Temperature can range from 8 to 25C in landfill cover soils based on seasonal temperatures (Boeckx et al., 1996; Schuetz et al., 2003), whereas landfill gas emitted from within the landfill at depths below the cover soil have been reported as high as 41C (Uz et al., 2003). The optimum temperature for methane production in the mesophilic range has been reported at 41 and 42C (Hilger and Barlaz, 2002). For refuse excavated from within the landfill, reported temperatures vary anywhere from 27 to 64C (Hilger and Barlaz, 2002). The pH values reported for landfill soil samples also varied from 4.4 to 7.6 (Whalen et al., 1990; Kightley et al., 1995; Boeckx and Van Cleemput, 1996; Boeckx et al., 1996; Czepiel et al., 1996; De Visscher et al., 1999; Wise et al., 1999; Christophersen et al., 2000; Schuetz et al., 2003), as did the reported pH range for landfill leachate from 3.7 to 9 (Watson-Craik et al., 1992; Tammemagi, 1999). Total organic carbon (based on a percent organic carbon per weight) ranges from 0.52 to 4.9 (Whalen et al., 1990; Boeckx and Van Cleemput, 1996; Boeckx et al., 1996; Czepiel et al., 1996; Borjesson et al., 1998; De Visscher et al., 1999; Wise et al., 1999; Schuetz et al., 2003). Cover soils from a MSW landfill in Athens, Georgia were shown to have a copper content of 139.22 mg L -1 (Wise et al., 1999), whereas leachate concentrations ranged from < 0.008 to 10.0 mg L -1 (Watson-Craik et al., 1992; Tammemagi, 1999). Finally, the percentage of methane also varies in landfill gas sample sites, reported from 45 to 60.6% (v/v), with carbon dioxide comprising anywhere from 40 to 60% (Bogner et al., 1997a; Tammemagi, 1999; Schuetz et al., 2003; Uz et al., 2003) with the remaining portion comprised of trace

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13 gases, such as benzene, hydrogen sulfide, nitrogen, and chlorinated hydrocarbons (Tammemagi, 1999). This environment in a landfill provides heterogeneity that allows interactions between microorganisms that may not be able to thrive together in a homogenous environment (Senior, 1995). Also, this encourages diversity among microbial types enabling the development of different species within their own niches (Senior, 1995), as demonstrated through the decomposition of MSW to methane. MSW decomposition is a microbially mediated process requiring a coordinated effort of several trophic groups of bacteria (Hilger and Barlaz, 2002). This decomposition process will be reviewed and summarized below based on previously described processes by Hilger and Barlaz (2002). The conversion of MSW into methane by microorganisms does not occur immediately after refuse is placed in a landfill. It can take several months or years in order for the appropriate growth conditions and the required trophic groups of bacteria to become established. In general, refuse decomposition occurs in phases including Aerobic phase (Phase 1) Anaerobic acid phase (Phase 2) Accelerated methane phase (Phase 3) Decelerated methane production phase (Phase 4) In Phase 1 (aerobic phase), both oxygen and nitrate are consumed as electron acceptors, and soluble sugars serve as a carbon source. All trophic groups required for refuse degradation (methanogenesis) are present. Trophic groups include cellulolytic organisms, fatty acid degraders, and methanogens. The pH begins at neutral, but, as an imbalance occurs (among fermentative, acetogenic, and methanogenic activity), the pH decreases with an accumulation of carboxylic acids (Phase 2). During this anaerobic acid

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14 phase, the methanogen population increases, and methane is first produced. There is also some decomposition of refuse during this stage. The accumulated carboxylic acids serve as the main substrate supporting methane production in phase 3 (accelerated methane phase). This phase is characterized by a rapid increase in the rate of methane production, an increase in pH, and a decrease in the concentration of carboxylic acids. During Phase 3, there is little hydrolysis of solid materials, and the populations of cellulytic, fatty acid-degrading (acetogenic) and methanogenic bacteria increase. Finally, during the decelerated methane production phase (Phase 4), methane production not only decreases but also carboxylic acids are depleted, and the pH levels out between 7 and 8. The major terminal products of anaerobic decomposition include methane and carbon dioxide. As methane migrates through soil layers in the landfill, it mixes with air where methanotrophs can further oxidize the methane present to carbon dioxide and water. Because of their significant role in intercepting methane before release into the atmosphere, methanotrophs are discussed in greater detail below with regard to their taxonomy, physiology and biochemistry. It is, however, important to first understand the factors that influence methane oxidation in landfills. Factors influencing methane oxidation in landfills Methane availability. Methane concentration has been reported to have a significant effect on methane oxidation rates in landfill cover soils (Czepiel et al., 1996; Bogner et al., 1997a; Hilger and Barlaz, 2002). When tested under the elevated methane concentrations of 1 to 10 5 ppm (Whalen et al., 1990; Bogner et al., 1997a) and 1-6% initial headspace concentration (Kightley et al., 1995; Visvanathan et al., 1999), landfill cover soils showed excellent capacity for methane oxidation with highest observed rates reported between 10 and 166 g m -2 day -1 (Whalen et al., 1990; Kightley et

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15 al., 1995; Bogner et al., 1997a; Visvanathan et al., 1999). In addition to a high capacity for methane oxidation in cover soils, Bogner et al. (1997b) showed the response of methane oxidation to ambient methane concentrations by measuring the initial methane concentration (0.01 ppm to 100 ppm) in the field over a 30-hour time period during an increase and decrease in available methane, thus indicating a rapid response of methanotrophic populations to changing methane concentrations. It has also been shown that changes in population density of methane-oxidizing bacteria (based on colony forming units of soil samples) are proportional to the concentration of methane within the soil (up to a concentration of 50%), thus indicating that some additional factor(s) in the soil environment may limit the methane-oxidizing population in sanitary landfills (Mancinelli et al., 1981; Mancinelli and McKay, 1985). Other studies have focused on the impacts of temperature and pH on methane oxidation rates, as summarized to follow. Temperature. Czepiel et al. (1996) reported a rapid increase in methane oxidation rates for landfill cover soils (0.2 to 4.4 mol g -1 h -1 ) as temperatures increased from 5 to 36C, with no oxidation at 45C. This trend of increasing methane oxidation rates with corresponding temperature increases up to a maximum has been observed in other landfill cover soil samples with maximum reported rates (see also Appendix A) at temperatures of 20 to 25C (7.5 x 10 -4 mol g -1 h -1 and 6.9 x 10 -4 mol g -1 h -1 , respectively; Boeckx et al., 1996), 25 to 30C (1.5 x 10 -4 and 1.4 x 10 -4 mol g -1 h -1 , respectively; Boeckx and Van Cleemput, 1996), and 31C (0.2 mol g -1 h -1 , Whalen et al, 1990). However, oxidation has been observed at temperatures as low as 5C (0.05 mol g -1 h -1 , Whalen et al., 1990; 0.2 mol g -1 h -1 , Czepiel et al., 1996). Another study focused specifically on methane oxidation at lower temperatures (2 to 15C) of landfill soil in temperate climates

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16 (Christophersen et al., 2000). They found that the oxidation rate increased with increasing temperature, with the observed maximum (1.17 mol g -1 h -1 ) occurring at the highest temperature tested (15C). This rate was higher than previously published rates (0.1 mol g -1 h -1 , Whalen et al., 1990; 0.5 mol g -1 h -1 , Czepiel et al., 1996) at the same temperature. Range of pH tolerance. Although methanotrophic organisms have a relatively wide pH tolerance range (4 to 9, Humer and Lechner, 1999), optimal growth and activity for most methanotrophic organisms occur between pH 6.6 and 6.8, with most strains growing over a pH range of 5.8 to 7.4 (Whittenbury et al., 1970). Methanotrophs are sensitive to acidification in the environment, with a notable decrease (nearly 50%) in methane consumption activity reported in clay soils when pH was lowered from 6.3 to 5.6 and no activity observed at lower pH levels (Le Mer and Roger, 2001). One study, conducted by Wise et al. (1999), reported the distribution of type I versus type II isolated methanotrophs (originating from mildly acidic landfill cover soil) in response to a change in pH from near neutral to 4.8, with type I methanotrophs being favored at all tested levels. However, these authors did not study variation in methane oxidation activity. Currently, there is limited information concerning pH effects on methane oxidation potential in landfill environments. To date, research has focused on working with landfill cover soil samples in the field and in the laboratory, with minimal studies investigating the activity and interactions of methanotrophs with other community members in culture-based experiments. Limited information is available on enriched cultures derived from landfill environments and how characterization information and assessment of the metabolic

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17 activities of the microbial populations directly translate to field activities. To better understand the interactions of these microorganisms in landfill environments, it is necessary to elucidate their roles in landfill communities while studying them in a more controlled setting, working with mixed cultures. Introduction to methanotrophs: methane-oxidizing bacteria Methanotrophic environments. Methanotrophs are aerobic or microaerophilic organisms that oxidize methane to derive energy and carbon for biomass (Lipscomb, 1994; Mancinelli, 1995; Hanson and Hanson, 1996; Graham et al., 2002). These bacteria are ubiquitous; they are prevalent in almost every environmental compartment where there is a stable source of methane and air (Graham et al., 2002). They are found in freshwater rivers and lakes, in marine environments, in soils and sediments, and on plant surfaces (Hanson and Hanson, 1996). As obligate aerobes, they are also abundant in anaerobic-aerobic interfaces where they oxidize methane originating from anaerobes. Examples of locations of anaerobic-aerobic interfaces include wetlands, rice paddies and landfill cover soil (Graham et al., 2002; Hilger and Barlaz, 2002). Also, “high affinity” methanotrophs have been identified and found to subsist on atmospheric methane levels ( 1.7 ppmv) in moist tundra soils (Whalen and Reeburgh, 1990; Graham et al., 2002). Another study identified a psychrophilic strain of methanotrophic bacteria that was isolated from acidic soils in the antarctic, exhibiting optimal growth at temperatures at or below 10C (Omel’chenko et al., 1993; Hanson and Hanson, 1996). These reports show how diverse methanotrophs are in a variety of environments and conditions. Methanotrophs are important in the global carbon cycle and are considered ecologically significant (Graham et al., 2002). Not only do they play a vital role in the

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18 reduction of the greenhouse effect by controlling methane release into the atmosphere, but they also have been shown to have applications in bioremediation, or using microorganisms to degrade toxic chemicals (Park et al., 1991; Hanson and Hanson, 1996; Sullivan et al., 1998). Taxonomy and physiology. Bacteria with the ability to use reduced carbon substrates (with no carbon-carbon bonds) as their sole source of carbon and energy are methylotrophs (Anthony, 1982; Lidstrom, 1992). Methanotrophs (a subset of this group) are strictly aerobic, gram-negative and obligate, not using multi-carbon compounds as sources of carbon and energy (Hanson, 1998). Known methanotrophs are catalase and oxidase positive, possess intracytoplasmic membranes, and are able to form resting stages (cysts, exospores, or lipid cysts) under adverse environmental conditions (Anthony, 1982; Graham et al., 2002). These characteristics, as summarized in Table 2-1, in addition to pathways for formaldehyde assimilation, which help categorize methanotrophs into three major groupstypes I, II and X. The enzyme responsible for the conversion of methane to methanol (and ultimately to carbon dioxide) by methanotrophs is a mixed-function oxidase, called methane monooxygenase (MMO) (Lidstrom, 1992). This enzyme exists in two forms, a membrane-bound particulate form (pMMO) and a soluble or cytoplasmic form (sMMO). All methanotrophs can synthesize pMMO, and most type II and X methanotrophs can produce both enzymes depending on a change in oxygen levels and copper concentration (Scott et al., 1981; Stanley et al., 1983; Prior and Dalton, 1985; Leak and Dalton, 1986). Figure 2-2 summarizes the methane oxidation pathways of methanotrophs. Several papers review these pathways, which are summarized here to provide a general overview

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19 of this process (Mancinelli and McKay, 1985; Hanson and Hanson, 1996; Graham et al., 2002). MMO catalyzes the first step in the pathway that converts methane to methanol. This step is highly energy-demanding because it requires two reducing equivalents (in the form of NADH) to split the di-oxygen molecule. One of the oxygen atoms is directly incorporated into methane (yielding methanol), whereas the other atom is reduced to water. Electrons from this first oxidation step are then transferred to the second reaction in which methanol is converted to formaldehyde. Formaldehyde serves a central role in methanotrophic metabolism, as it can be oxidized to formate or assimilated by the cell through the ribulose monophosphate and/or serine pathway. Formate can be further oxidized by the cell to carbon dioxide, which can then be released as a gas into the surrounding environment. While carbon dioxide is a greenhouse gas, its impacts on global warming is approximately 26 times less than those of methane (Hanson and Hanson, 1996). Factors regulating the expression of MMO. Changes in oxygen level and copper concentration affect the expression of the MMO enzyme. Knowledge of the specific factors that regulate the expression of MMO is important because it helps determine what different types of methanotrophs thrive in a given environment. These factors regulating MMO expression are summarized as follows. The copper-to-cell ratio in the growth environment has a significant effect in determining whether most type II and X methanotrophs will express sMMO or pMMO, with low ratios supporting sMMO expression and higher ratios supporting pMMO expression (Graham et al., 2002). This has been demonstrated in several studies, such as Prior and Dalton (1985) who studied type X methanotroph, Methylococcus capsulatus

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20 (Bath). When grown in the presence of methanol in continuous culture, they reported that the activity of pMMO increased as the concentration of copper was increased (0 to 31.5 M) in addition to an increase (85%) in the rate of oxygen consumption in whole cells when copper (18.9 M) was added to the medium in the presence of methane (0.1 mM). Another study also reported that Methylococcus capsulatus (Bath) showed enhanced pMMO activity as the copper concentration increased (0.3 to 20 M) with a corresponding increase in copper detected in the membrane fraction of the cell and decrease in the soluble fraction (Chan et al., 1993). Burrows et al. (1984) also discussed that as the copper concentration increased from 1 M to 5 M, the MMO activity was predominately in the soluble form at lower concentrations and in the particulate form at higher concentrations. Thus to ensure 100% pMMO activity, copper concentrations should be maintained greater than 5 M, with optimal pMMO productivity occurring in batch and continuous culture at 10 M (Park et al., 1992; Shah et al., 1992). Cells expressing pMMO have a greater affinity for methane than cells expressing sMMO. It is hypothesized that this is observed because pMMO uses a higher-potential electron donor (cytochrome c) that is energetically more favorable than the electron donor (NADH) used by sMMO (Hanson and Hanson, 1996; Graham et al., 2002). Cells expressing pMMO produce higher growth yields on methane than cells expressing sMMO (Hanson and Hanson, 1996). Studies have reported that carbon conversion efficiencies were 38% higher for Methylococcus capsulatus (Bath) when grown in continuous culture as the growth medium was amended with increasing amounts of copper (0.79 M to 25.2 M) (Leak and Dalton, 1986; Sullivan et al., 1998). A similar trend has been demonstrated in mixed cultures where batches of a mixed methanotrophic

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21 bacteria culture (obtained from a water treatment facility) showed a 3-fold increase in the maximum methane utilization rate (measured as a kinetic constant) with an increase in copper concentration (0 to 6.3 M) (Boisesen et al., 1993). Initial investigation of MMO activity suggested that oxygen may also play a role in MMO expression (Scott et al., 1981; Stanley et al., 1983; Sullivan, 1998). Scott et al. (1981) studied Methylosinus trichosporium OB3b grown in shake flasks under both oxygen-limited and oxygen-excess conditions (15:1 and 1:5 ratio of CH 4 to O 2 , respectively). Based on transmission electron microscopy (TEM) observations, extensive peripheral intracytoplasmic membranes were observed only under the oxygen-limited conditions, which were also accompanied by substantial particulate MMO activity (in cell-free extracts). Contrastingly, under oxygen-excess conditions, the peripheral intracytoplasmic membranes were limited or absent, with MMO predominately in the soluble form. Similarly, Stanley et al. (1983) demonstrated that over a range of biomass concentrations, sMMO in Methylococcus capsulatus (Bath) was evident at lower cell concentrations under oxygen limitation than under methane limitation in low copper medium (3.2 M). Also, the transition from 100% particulate to 100% soluble activity spans a wider range of biomass concentration under methane limitation than oxygen limitation. Thus, when an oxygen-limited chemostat culture (under low copper concentration) was converted to methane-limitation, a transition from soluble to particulate MMO was observed. Graham et al. (1993) showed that, in a continuous-flow reactor, type II strain Methylosinus trichosporium OB3b was competitively successful over type I strain Methylomicrobium album BG8 when copper and oxygen levels were low and methane

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22 levels high, with the converse being true (type I outcompetes type II) under opposite conditions. A similar trend, showing type I methanotrophs thriving under low methane levels while type II methanotrophs dominated under high methane levels, was observed by Amaral and Knowles (1995). A gel-stabilized system with counter gradients of methane and oxygen was used to grow methanotrophs from wetland and lake sediment. Development of methanotrophic bands were observed at the bottom of the columns where methane was highest and oxygen was in trace amounts. 16S rRNA signature probe hybridization indicated that top and bottom band methanotrophs belonged to type I and II physiological groups, respectively. By converting the flows of oxygen and methane to opposite ends of the columns, the band position of methanotrophic growth reversed, implying that type I strains mostly occurred where methane was lowest, whereas type II strains occurred at the higher methane concentrations. In general, it can be assumed that most type I methanotrophs prefer high oxygen, low methane environments, whereas most type II organisms prefer the converse, particularly with limited copper concentrations. Nutrient availability can also influence methanotroph type prevalence. Vecherskaya et al. (1993) reported that representatives of type I and X methanotrophs (Methylomonas, Methylobacter and Methylococcus) dominated over type II representatives (Methylocystis and Methylosinus) at 61.6% and 38.4%, respectively, in tundra bog soils. These authors suggested this was because of type I and X groups resembling r-strategists, characterized by rapid growth under favorable conditions and low survival under unfavorable conditions. On the contrary, the type II group resembled k-strategists, where growth was slower but survival was better. It was concluded that

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23 environments that allow the most rapid growth of methane-utilizing bacteria favor type I methanotrophs, whereas type II methanotrophs tend to survive better in environments under nutrient deprivation (Hanson and Hanson, 1996). Wise et al. (1999) further supported this in their finding that type II species dominated enrichments where the medium used was diluted five-fold, regardless of initial pH or methane concentration in headspace. This result suggested the ability of type II species to better acquire nutrients when it is a limited source. Alternatively, the higher salt concentration at the full strength medium may also have rendered the type II species at a selective disadvantage. Degradation of aliphatic and aromatic compounds by methanotrophs. The uniqueness of the MMO enzyme to methanotrophic oxidation has already been discussed. In addition to oxidizing methane, MMO catalyzes the fortuitous metabolism of a variety of compounds, including xenobiotic chemicals, found in the environment (Higgins et al., 1980; Hanson and Hanson, 1996). The potential application for the use of methanotrophs to biodegrade xenobiotics, defined as any toxic chemical occurring in quantities that pose a threat to man or nature (Muller, 1992), and to catalyze a number of biotransformations has attracted the interest of the environmental community (Hanson et al., 1992; Hanson and Hanson, 1996). With the development of bioremediation research, it has been shown that the soluble form of MMO has a broader cometabolic substrate range in many methanotrophs than the particulate form (Lidstrom, 1992; Sullivan et al., 1998), including aliphatic, straight-chain, branched alkanes or alkenes up to 8 carbons long, phenols, cyclic and aromatic compounds as well as halogenated hydrocarbons (Lidstrom, 1992; Hanson et al., 1992; DiSpirito et al., 1992). On the other hand, pMMO is more substrate

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24 specific and will oxidize alkanes and alkenes up to 5 carbons long but will not oxidize cyclic or aromatic compounds (DiSpirito et al., 1992). Based on field and laboratory studies, cultures expressing sMMO are capable of degrading TCE at rates higher than those expressing pMMO (Tsien et al., 1989; DiSpirito et al., 1992; Hanson et al., 1992; Lontoh and Semrau, 1998). TCE has already been identified as a constituent in landfill gas emissions, is a suspected carcinogen in humans and persistently exists as a groundwater aquifer contaminant (Alvarez-Cohen and McCarty, 1991; Henry and Grbic-Galic, 1991). It was commonly used in the dry-cleaning of clothes and in industry as a degreasing agent for metal parts (Muller, 1992). Its low degradability in the environment coupled with its suspected carcinogenicity has spurred efforts to find alternative substitutes for TCE usage (Muller, 1992). Studies have reported methanotrophic degradation of TCE (Tsien et al., 1989; Uchiyama et al., 1989; Alvarez-Cohen and McCarty, 1991; Henry and Grbic-Galic, 1991; Alvarez-Cohen et al., 1992; DiSpirito et al., 1992; Koh et al., 1993; Smith et al., 1997; Lontoh and Semrau, 1998; Han et al., 1999) with a brief discussion of its degradation pathway. TCE can be biologically transformed either anaerobically or aerobically. Under anaerobic conditions (methanogenic), the degradation pathway begins with tetrachloroethylene. Tetrachloroethylene (PCE) undergoes reductive dehalogenation, where a halogen group (in this case chlorine) is removed with the addition of electrons and a hydrogen atom (Mohn and Tiedje, 1992), yielding TCE. Further reductive dehalogenation of TCE yields other intermediates, cis-dichloroethylene (cis-DC) and vinyl chloride (VC), with carbon dioxide being the terminal product. A major concern with incomplete degradation of PCE is the toxicity of the intermediate compounds formed, specifically, VC which is a

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25 known carcinogen. However, it has been reported that methanotrophs can completely transform TCE to carbon dioxide (Figure 2-3) with no VC residual (Little et al., 1988; Alvarez-Cohen and McCarty, 1991). As proposed by Little et al. (1988), TCE is broken down by methanotrophs to form TCE epoxide, which then breaks down spontaneously in water to form dichloroacetic acid (DCA), glyoxylic acid, or one-carbon compounds, such as carbon monoxide and formate. As with anaerobic transformation, concern arises over the accumulation of these intermediates, namely DCA, which is likely to be a carcinogen in humans (US EPA, 2004). It has also been noted in other studies that, as methanotrophs transform TCE, it can only do so up to a certain concentration (reported range of 6.0 to 22 mg L -1 ) before toxicity effects are observed (Uchiyama et al., 1989; Alvarez-Cohen and McCarty, 1991; Henry and Grbic-Galic, 1991) or growth of the methanotrophic culture was inhibited (Han et al., 1999). Benzene is known for its carcinogenity and has been detected in landfill gas, as previously described. It is an aromatic compound that is also a common constituent in gasoline, diesel and jet fuel (Muller, 1992). Its degradation has also been studied in detail with a brief summary provided here of the bacterial degradation pathway. This pathway is not specific to methanotrophs, but to microorganisms utilizing the monooxygenase enzyme (Figure 2-4) previously described by Muller (1992), Madigan et al. (2003) and Fritsche and Hofrichter (2004). Degradation begins with initial oxidation of the benzene ring, where MMO adds an oxygen atom onto the aromatic ring, yielding benzene epoxide which is further hydroxylated to benzenediol and ultimately, catechol (Madigan et al., 2003). Methanotrophs have also been suspected to oxidize aromatics by a hydrogen-abstraction pathway that involves direct addition of an oxygen atom to an

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26 aromatic carbon (Dalton et al., 1981: Lindner et al., 2000). Regardless of the pathway followed, the final product is a hydroxylated aromatic compound. Methanotrophs are capable of degrading aromatic compounds (such as benzene); however, few studies have specifically investigated methanotrophic degradation of benzene. Uchiyama et al. (1989) studied a pure, methane-oxidizing bacterium that was isolated from a mixed culture that degraded benzene. At an initial concentration of 10 mg L -1 of benzene, the methanotroph (also tentatively identified as a type II strain), degraded benzene (only 6% over a 10 day period). This same study also reported a 96% removal of TCE to a concentration of 0.35 mg L -1 over 7 days. Colby et al. (1977) reported that sMMO (from Methylococcus capsulatus (Bath)) catalyzed the degradation of a number of compounds (n-alkanes, n-alkenes, ethers, alicyclic and heterocyclic compounds) including aromatic compounds benzene and toluene. These authors showed that sMMO isolated from Methylococcus capsulatus (Bath) was able to oxidize benzene to phenol. Other studies have focused on the ability of methanotrophs to oxidize other aromatic compounds, such as polychlorinated biphenyls (Adriaens and Grbic-Galic, 1994; Lindner et al., 2000), chlorinated phenols (Kostova and Todorova, 2000) and polycyclic aromatic hydrocarbons (Stapleton et al., 1998). Brief introduction to heterotrophic bacteria Bergey’s Manual of Determinative Bacteriology (Holt, 1994) defines a heterotroph as an organism that requires one or more organic compounds and cannot use carbon dioxide as its sole carbon source. This indicates that these bacteria are found in a wide variety of environments. Heterotrophs contain an enzyme known as dioxygenase, which is responsible for attack on aromatic compounds, such as benzene (Rochkind-Dubinsky et al., 1987; Muller, 1992). The dioxygenase enzyme requires molecular oxygen as a

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27 substrate (Rochkind-Dubinsky et al., 1987). When this enzyme is used, it incorporates two atoms of oxygen from an oxygen molecule onto the compound being oxidized. In the case of benzene degradation (Figure 2-5), the dioxygenase enzyme inserts two oxygen atoms onto the aromatic structure yielding an intermediate compound, cis-dihydrodiol (Muller, 1992). This dihydrodiol in then dehydrogenated to yield catechol, which can be cleaved in two ways, either as orthoor metacleavage (Muller, 1992). The orthocleavage produces cis,cis-muconic acid, whereas metacleavage forms 2-hydroxymuconic semialdehyde (Muller, 1992). Ultimately, the final products of degradation are acetyl-CoA and succinate (ortho-cleavage pathway) or pyruvate and acetaldehyde (meta-cleavage pathway). Pseudomonas putida has been shown to follow the aromatic dioxygenase enzyme pathway and has also been identified in aerobic degradation of TCE (Wackett and Gibson, 1988). It was found that toluene dioxygenase was responsible for this degradation of TCE (Wackett and Gibson, 1988). In mixed methanotrophic-heterotrophic cultures, it has been proposed that, after TCE was oxidized by methanotrophs to form glyoxylic acid (or carbon monoxide and formate), heterotrophs further mediated transformation to carbon dioxide (Little et al., 1988). Additionally, it has been hypothesized that heterotrophs in a mixed methanotrophic-heterotrophic culture may have assimilated or detoxified toxic organic compounds accumulated during TCE oxidation, which may have allowed the methanotroph (Methylocystis sp. strain M) to degrade TCE at high concentrations (Uchiyama et al., 1989; Uchiyama et al., 1992). These results indicate that heterotrophic bacteria play an important role in complete TCE degradation by methanotrophs. Other studies have suggested the importance of

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28 methanotrophic-heterotrophic interactions in combined metabolic attack on complex linear alkylbenzenesulphonates (Hrsak et al., 1982), as evidenced by faster and more complete transformation of these compounds by the mixed culture versus the individual community members alone (Hrsak and Begonja, 1998). Also, co-metabolic transformation of monoand dichlorobiphenyls and chlorohydroxybiphenyls was compared for both mixed and pure cultures (Adriaens and Grbic-Galic, 1994). These authors demonstrated that the substrate disappearance in the mixed culture was consistently higher than in the pure culture. Additionally, it was presumed that the heterotrophs present in the mixed culture were responsible for the products resulting from ring cleavage of biphenyl. These results indicated that methanotrophic-heterotrophic interactions may play an important role in the further mineralization of the substrates tested. Specific heterotrophs isolated from mixed methanotrophic-heterotrophic cultures have been identified and partially characterized in previous studies (Uchiyama et al., 1992; Hrsak and Begonja, 1998; Dunfield et al., 1999). Heterotrophs that grew on nutrient agar did not grow with methane, possessed colonies that were white and yellow, were gram negative and oxidase positive, and contained short rod-shaped cells (1.1 to 2.4 m) were identified as Xanothobacter autotrophicus, and in the genera Pseudomonas and Bacillus (Uchiyama et al., 1992). Other isolated heterotrophs varied in colony color from yellow, yellowish-green, white, and translucent, grew on nutrient agar, peptone-yeast extract-glucose agar, and peptone-yeast-extract agar with abundant growth at 30C, and ranged in size from < 1 to 5 mm (Hrsak and Begonja, 1998). Tentative identification of these heterotrophs revealed the genera Blastobacter, Pseudomonas, and Xanthobacter.

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29 Also two unidentified heterotrophic community members showed similarities to prosthecates, but this morphology had not been confirmed with electron microscopy. Other identified heterotrophs isolated from a soil derived mixed culture based on partial 16S rRNA gene sequences included Pseudomona, Variovorax, Bradyrhizobium and Hyphomicrobium (Dunfield et al., 1999). Significance and Rationale of Project In light of the growing number of landfills in the United States, there is a resulting need for more efficient waste degradation, prevention of methane emissions to the atmosphere, and minimization of releases of toxic compounds (Hanson and Hanson, 1996; Kjeldsen et al., 1997). The potential role of methanotrophs in these processes is not well understood. Because of the difficulty in studying methanotrophic populations directly in the field, there is great need to develop protocols to allow study of field processes in the laboratory. To date, a direct link between methanotrophic diversity and activity in field samples with isolated mixed cultures from field samples has not been made. Doing so would facilitate prediction of biodegradative activity of microbial populations in landfills, thus allowing better management of these systems. Based on this motivation, our study took initial steps in bridging the gap between field and laboratory studies by isolating mixed methanotrophic-heterotrophic cultures from landfill samples. These landfill soil samples were initially screened for differences in methanotrophic populations from two different locations in the landfill that differed in age by approximately 4 to 5 years. Results from this initial study indicated a predominance of type II methanotrophs in samples removed from the older region and gave strong indication that the age and thus conditions in a landfill can be directly linked to population diversity (Uz et al., 2003), although linking environmental conditions to

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30 methanotrophic diversity was beyond the scope of this previous project. Our study further builds upon this initial work by comparing the isolated mixed cultures to ultimately allow a broader microbial landfill-laboratory study in the future. The aims of this study were, (i) phenotypically characterize two mixed methanotrophic-heterotrophic cultures isolated from a MSW landfill and (ii) assess the effect of environmental conditions on growth and metabolic activities of each mixed culture. This was accomplished through the following specific objectives. Isolate stable mixed methanotrophic-heterotrophic cultures from both samples under a single set of enrichment conditions. Assess the individual heterotrophic populations in each mixed culture through characterization (colony and cellular morphologies, phenotypic studies). Characterize both mixed cultures in terms of cellular morphology, growth using VIS spectrophotometry, rates of methane depletion, and screening the degradation potential of contaminants benzene and TCE using oxygen uptake analysis. Vary initial methane concentration, temperature, pH and copper concentration to assess these mixed cultures response to changes in their growth and ability to oxidize methane. Vary the concentration of media to assess how these cultures respond (in terms of growth rates) to nutrient deprivation. The intention of our study was to serve as a pivot-point for a broader, more in-depth study of microbial isolates derived from landfill environments. Ultimately, we hope that this and future studies will then provide a better understanding of what complex processes occur in landfill environments that will encourage more efficient management of these environments to minimize environmental damage.

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31 Table 2-1. Characteristics of type I, II, and X methanotrophs Characteristic type I type II type X Cell morphology Short rod, some cocci or ellipsoid Rods, crescent or pear shaped Cocci Growth at 45C No No Yes pH (optimum) 7 7 7 Membrane arrangement: bundles of vesicular disks Yes No Yes paired, aligned to cell periphery No Yes No Resting stages formed: exospores No Some strains No cysts Some strains Some strains Some strains RuMP pathway present a Yes No Yes Serine pathway present No Yes Sometimes Major PLFA b 16 18 16 Recognized genera Methylococcus Methylomonas Methylobacter Methylomicrobium Methylocaldum Methylospaera Methylosinus Methylocystis Methylocella Methylococcus a RuMP = ribulose monophosphate cycle b PLFA = phospholipid fatty acids

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32 Figure 2-1. Basic landfill components (Freudenrich, C.C., 2000. How landfills work. How Stuff Works www.howstuffworks.com , last accessed November 2002). Arrows indicate flow of leachate percolating through landfill. A) Groundwater. B) Clay liner. C) Flexible membrane liner. D) Leachate collection pipe. E,F,G) Drainage system composed of gravel, sand, and clay. H) Protective barrier. I) Refuse. J) Cover layer. K) Leachate collection pond.

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33 Serine Pathway (Type II/X)CH4CH3OHHCHOHCOOH CO2 RuMP Pathway (Type I/X)sMMOpMMONADHNAD+CytCredCytCoxCytCoxCytCred2 NAD+2 NADH Figure 2-2. Methane oxidation pathway in methanotrophs. Figure 2-3. Aerobic methanotrophic degradation of TCE (Little et al.,1988; Figure 4, pg.955).

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34 Benzene HHO NADHNAD+Benzene epoxide H2O HOHHOHBenzenediolor trans-dihydrodiol NADHNAD+ OH O H Catechol Figure 2-4. Microbial oxidation of benzene utilizing monooxygenase. The monooxygenase enzyme is employed in the hydroxylation of benzene to catechol in which NADH is an electron donor. Benzene NADHNAD+ HOHHOHcis-dihydrodiol NADHNAD+ OH OHCatechol mo metaortho Figure 2-5. Microbial oxidation of benzene utilizing dioxygenase. Dioxygenase is employed in the hydroxylation of the benzene ring showing both orthoand metapathways.

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CHAPTER 3 MATERIALS AND METHODS Overview of Materials and Methods As shown in Figure 3-1, our study was performed in four broad stages. Stage 1 was the isolation of mixed cultures from landfill samples. Stages 2 and 3 involved the characterization of the two isolated mixed cultures, and Stage 4 consisted of evaluating the response of each mixed culture to environmental effects. Each stage is addressed in the following sections in further detail. Stage 1: Isolation of Mixed Cultures from Landfill Samples Sampling Sites Samples used in our study were taken in late September of 1999 from the Alachua County Southwest Landfill, a lined municipal solid waste landfill containing mixed household and commercial/light industrial waste, located in Archer, FL, USA. This landfill was covered with sandy soil to a depth of approximately 30 to 45 cm. The samples were removed during installation of gas collection wells from two sections of the landfill (Figure 3-2), differing in age by approximately 5 years, with the more recent being 2 years of age (well number GW 70). Samples were referred to as GW 60-13 and GW 70-20 to indicate the well location (60 and 70, respectively) and depth of the sample collected (13 and 20 feet, respectively). Visual observations of the collected samples indicated a mixture of MSW and soil (both sandy and humic). Samples were collected in duplicate and immediately refrigerated. Without delay, replicate samples were manually homogenized by mixing with a sterile spatula in a laminar flow hood before use. 35

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36 Enrichment and Isolation of Mixed Cultures All glassware used was acid-rinsed (6M HNO 3 ) to avoid copper contamination as suggested by Bowman and Sayler (1994) and autoclaved for sterility. Two grams of landfill sample was incubated in a 250 mL Erlenmeyer flask (equipped with a rubber stopper and a glass wool-packed filling tube) containing 30 mL of nitrate mineral salts (NMS) medium (Whittenbury et al., 1970; Appendix B) with FeEDTA, an organic ligand that chelates metal ions (Henry and Grbic`-Galic`, 1990) and doubly deionized water (Barnstead International, Melrose Park, IL, USA). A sufficient amount of copper (10 M CuNO 3 ) previously shown to promote maximal pMMO activity (Shah et al., 1992) was added to the medium to promote pMMO expression in GW70, unless otherwise mentioned. Headspace was removed from the flasks through filling tubes using a vacuum pump apparatus and an equivalent amount of highest purity methane (99.9%, Strate Welding Supply Co., Jacksonville, FL, USA) was added to achieve a methane-to-air ratio of 30:70 for GW60 and 20:80 for GW70 for routine culturing. Flasks were incubated at 30C and shaken at 270 rpm in a reciprocal laboratory shaker (Barnstead International, Melrose Park, IL, USA). After detecting sufficient visible turbidity ( 7 days), transfers to fresh medium were prepared using a 10% inoculum and repeated until a stable culture was detected (weekly transfers over a 3-year period). Stability of the mixed culture populations was assessed upon streaking onto nutrient (Difco, Becton Dickinson and Company, Sparks, MD, USA) and NMS agar (NMS medium and Bacto agar, Becton Dickinson and Company, Sparks, MD, USA) and observing no noticeable changes in colony type and

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37 morphologies. These stable mixed cultures were then subjected to further isolation methods and characterization. Stages 2 and 3: Characterization of Mixed Cultures Heterotroph Isolation Heterotrophs were isolated from both mixed, enriched cultures with ease through the serial dilution and spread plating methods previously described. Nutrient agar plates were used to promote heterotrophic growth and the dilution series yielding the best growth of colonies (30 to 300 colony forming units, CFUs) were selected to isolate individual populations. Individual colonies were inspected under a plate microscope and selected for transfer based on pigmentation, size or shape to isolate each representative population. Plates were stored in plastic sealable containers, inverted at 30C and inspected every 1 to 3 days, with routine swabbing of containers with ethanol to prevent fungal contamination as described by Whittenbury (1970). Transfers were made onto nutrient agar, and, after colonies developed, further transfers were made to isolate pure colonies as indicated by uniform growth on plates. Morphological and Phenotypic Characterization Pure colonies were visually inspected over a 20-day period for morphological characteristics as previously described by Norrell and Messley (1997). Characteristics, such as colony shape and appearance (e.g., convex, flat or umbonate elevation), consistency (e.g., smooth, mucoid, dry), edge (e.g., smooth or irregular), and pigmentation (e.g., white, cream, pink, yellow, brown), were used to describe isolated heterotrophs. The time for full colony development was also monitored, and the size was measured using a stage micrometer on the plate microscope. Growth on nutrient agar was also assessed using a previously described method (Hrsak and Begonja, 1998). If no

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38 visible growth was observed on nutrient plates, a “-“ sign was recorded. Very weak growth (as indicated by faint streaks occurring on less than of the plate) was recorded as “-/-+”; weak growth was defined by distinct streaks growing on less than of the plate and recorded as “-+”; good growth had distinct streaks on of the plate and recorded as “+”; and, finally, abundant growth covered the entire plate and was indicated as “++.” Nutritional requirements of isolated heterotrophs was determined by the ability of isolated colonies to grow on other agars, specifically peptone-yeast extract and tryptic soy (Becton Dickinson and Company, Sparks, MD) and prepared according to the labeled directions. Isolated heterotrophs were also tested for their ability to use methane as a sole carbon and energy source by streaking colonies onto NMS agar plates and incubating them under methane-to-air tensions of 30:70. Growth was assessed and monitored daily over a period of 30 days and recorded using the adapted system. Each isolated heterotroph was then subjected to phenotypic assays to aid in further characterizing each population. Gram staining was performed to classify heterotrophs as either gram negative (-) or gram positive (+) using a Gram Stain Kit (Fisher Diagnostics of Fisher Scientific, Middletown, VA, USA). Catalase tests were performed by placing individual colonies on top of a microscope slide and adding a few drops of hydrogen peroxide with a pasteurized pipet, as previously described by Norrell and Messley (1997). The production of bubbles by individual colonies after exposure to hydrogen peroxide was monitored and recorded as positive, (+) if bubbles were formed. The oxidase assay was performed based on procedures described by Gerhardt et al. (1994). A positive test result (+) was recorded if a blue-purple color developed when individual colonies were

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39 exposed to a 1% solution of tetramethyl-p-phenylenediamine, (no.16020-2, not HCl salt), (Sigma-Aldrich, St. Louis, MO, USA). Cellular Characterization Scanning electron microscopy (SEM) was used to evaluate the cellular components present in the enriched, mixed cultures, GW60 and GW70. Both cultures were grown under the previously described conditions where samples were harvested from the late exponential phase to capture the active proportions of bacteria present. Samples were analyzed in the Electron Microscope Lab in the Microbiology and Cell Science Department at the University of Florida. Samples were fixed with 2% glutaraldehyde for 1.5 hours and then micro-centrifuged at 8,000 x g for 1 minute. After being washed 3 times in 0.1 M cacodylate buffer (pH 7.2), the samples were postfixed in 1% osmium tetroxide (OsO 4 ) over night at 4C. The samples were dehydrated in a graded ethanol series to 100% then washed twice in hexamethyldisilazane (HMDS) for 5 to 10 minutes and allowed to air dry. Samples were then mounted on Alcian blue-coated cover slips on SEM stubs, gold coated in a Denton Desk II sputter coater and examined with a Hitachi S-570 scanning electron microscope (Hitachi Co., Tokyo, Japan). Cellular shapes were determined based on SEM results. An average length and diameter, measured directly from the photographs, were calculated for each distinctive cellular shape (rods, coccobacillus, coccobacillus with tails). Tentative Identification of Heterotrophs Morphological and phenotypic results combined with the cellular shapes and sizes determined using electron microscopy were used to tentatively identify isolated heterotrophs using Bergey’s Manual of Determinative Bacteriology (Holt, 1994), which provides a systematic approach to identify bacteria by classification into a major

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40 category, group, genus and species. Bacteria can be divided into 4 major categories with listings of groups within each major category and a brief description of the kinds of bacteria contained within each group. Most groups in the manual provide one or more tables or keys, indicating characteristics that can be used to differentiate the genera within the group. If extensive diagnostic tests have been performed on the unknown bacteria in question, the manual also provides genus descriptions that are accompanied by one or more tables that allow differentiation of the species contained in the genus. Growth Studies Determination of growth rates Specific growth rates were determined by inoculating a 250 mL nephlos flask (Figure 3-3) for measuring turbidity with a UV-VIS spectrophotometer (Milton Roy Company, Ivyland, PA, USA) set at a 600 nm wavelength. The initial optical density (OD) of each sample was diluted to 0.2 (Brusseau et al., 1990) using NMS medium (either with or without copper, depending on culture) to accurately compare rates. Samples were prepared in triplicate plus one control, which contained no methane. Methane was injected as previously described and incubated under the previously described enrichment conditions. Time-zero measurements were made after inoculation with methane and subsequent absorbance readings were recorded every few hours over the course of several days to create a growth curve. The time required for each culture to reach exponential phase (lag time) was also recorded. Specific growth rates and generation times were calculated using linear fits to the slope of the exponential growth phase of each growth curve obtained using the following equation, 1212xxln t t 1 , where is the specific growth rate (h -1 ), (t 2 -t 1 ) is the duration in hours of the exponential

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41 phase, x 2 is the absorbance reading at the end of the exponential phase at time t 2 , and x 1 is the absorbance reading at the beginning of the exponential phase at time t 1 (Hrsak and Begonja, 1998). Generation times (t gen or doubling time) (Madigan et al., 2003) were calculated as 12genxx301.0t (h) and growth rate constants as gent693.0k (h -1 ). Analysis of protein content At the end of each growth curve, samples were analyzed for protein concentrations using the BioRad protein assay kit with bovine serum albumin (BSA) as a standard (Life Science Research, Hercules, CA) based on methods described in the kit and also previously described by Han et al. (1999). Samples were centrifuged in sterile tubes at 1.94 g for 25 minutes in a J2-HS Beckman centrifuge (Beckman Coulter, Fullerton, CA, USA) and re-suspended using NMS to a wet cell weight concentration of 0.2 g mL -1 . Cells were then digested at 98C for 10 minutes in 5 N NaOH and micro-centrifuged at 6,000 x g for 15 minutes. A standard curve was generated using BSA as described by the BioRad protein assay kit with the unknown solutions serially diluted to achieve final protein concentrations within the linear range of the assay. The amount of protein was determined by measuring the absorbance at 595 nm after the BioRad assay reagent had been added. sMMO assay After cultures were re-suspended to 0.2 g mL -1 , a portion of this sample was reserved for detection of the presence of sMMO using the colorimetric assay developed by Brusseau et al. (1990) and previously described by Lindner et al. (2000). One-mL of cells was added to screw-cap tubes in triplicate plus one heat-killed control (which was

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42 autoclaved and allowed to cool). Laboratory-grade naphthalene crystals (Fisher Chemicals of Fisher Scientific, Fair Lawn, NJ, USA) were added to each tube and allowed to incubate at 30C while shaken at 270 rpm between 1 to 3 hours. Another control was included to test for a false positive result that may be due to heterotrophic transformation of naphthalene and not from sMMO activity. A 10 mL vial, containing 1 mL of re-suspended cells and naphthalene crystals, was capped with a rubber septum and crimped. Four mL of a known MMO inhibitor, acetylene (Strate Welding Supply Co., Jacksonville, FL, USA) was injected through the rubber septa using a syringe and allowed to incubate as well. After incubation, 100 L of freshly prepared 4.21 mM tetrazotized-o-dianisidine was added to each tube. For the acetylene control, o-dianisidine was added using a syringe and injected through the septum. A positive test result for the presence of sMMO was indicated by the rapid formation of a pink-purple color created by the dye complex between napthol and o-dianisidine, whereas a negative result appeared as no color change (indicated by the brown color of tetrazotized-o-dianisidine). In the acetylene control, a negative test result would indicate that MMO was inhibited and would not oxidize naphthalene, also implying that the enzyme activity present in the heterotrophs was not creating a false positive test. Assessment of methane-oxidizers and heterotrophs The proportion of methanotrophs and heterotrophs in both mixed cultures was determined by methods described by Hrsak and Begonja (1998). Once mixed cultures reached late exponential phase, samples were serially diluted and spread plated onto NMS (for methane oxidizers) and nutrient agar (for heterotrophs) as previously described. NMS plates and nutrient plates were also stored in the previously described

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43 conditions and allowed to incubate for 2 to 4 days. Plates containing 30 to 300 CFUs were selected and counted as suggested by Madigan et al. (2003). Determination of Methane Oxidation Rates Methane oxidation rates are commonly used to measure the physiological activity of methane users (Hanson and Hanson, 1996). Specific methane oxidation rates ( CH4 ) were evaluated by GW60 and GW70 for a particular cultivation period (t 2 -t 1 ) where methane oxidation was observed. The equation used is described by Hrsak and Begonja (1998): 12114ttVxMCH where M = total mass of CH 4 in the vial (mg), x = average biomass concentration (mg l -1 ) (determined using the BioRad Assay Kit as previously described), and V 1 = volume of liquid. Experiments were conducted using 60 mL glass bottles sealed with Mininert valves (VICI Precision Sampling Inc., Baton Rouge, LA, USA) (Figure 3-4). These valves allowed ease in sampling using a gas-tight Hamilton syringe (Hamilton Company, Reno, NV, USA) with minimal leakage detected during experiments. Samples were run in triplicate with each bottle containing 10.6 mL of NMS medium and 2.6 mL of liquid culture (calculated based on maintaining 78% volume in headspace and 5:1 ratio of media-to-culture) with adjusted optical density of 0.2. Controls with no cells and 12 mL of NMS medium were included to detect the amount methane lost during experimentation. A calculated volume of headspace was removed from each vial using a hypodermic sterile needle and 10-mL disposable sterile syringe (Fisher Scientific, Fair Lawn, NJ, USA) and replaced with an equivalent volume of methane that was injected through a sterile filter attached to a separate needle and syringe. As an example of headspace volumes, to achieve 30% methane, 14.4 mL was

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44 removed and replaced in the controls and 14.0 mL for the triplicate runs. Parafilm was used to secure the connection between the Mininert screw caps and bottle to prevent leakage. A 3-point calibration curve was created for each set of experiments using standards at 10%, 30% and 50% methane. Standards were created by using 10 mL air sampling bags (SKC Inc., Eighty Four, PA, USA) with the appropriate volume of air removed and replaced with filter-sterilized methane. This was repeated for the three concentrations with each bag gently massaged for several seconds to ensure complete mixing before analysis. Sixty L were removed from each sample and injected into a Shimadzu GC-14A gas chromatograph (GC) (Shimadzu Co., Tokyo, Japan) equipped with a flame ionization detector connected with a Chromatopac C-R501 integrator (Shimadzu Co., Tokyo, Japan). The GC column was stainless steel (0.317 by 41 cm) packed with Porapak Q (80/100 mesh; Waters Associates, Inc., Framington, MA, USA). Nitrogen was used as the carrier gas, and analyses were carried out under the following conditions: column temperature, 50C, and injector and detector temperature, 180C. Standard samples were run in triplicate at each concentration and used to generate a calibration curve in Excel (Microsoft Corporation, USA). Headspace samples from bottles were measured for each sample at time-zero and every few hours thereafter over the course of a couple days or until methane oxidation ceased. At the end of the experiments, protein concentrations were analyzed in addition to sMMO as previously described. Data was imported into spreadsheets created in Excel to calculate methane oxidation rates based on linear regression during the phase of methane oxidation. Any detected losses in the controls were accounted for in the rates.

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45 Evaluation of Oxidation Potential Since it is known that type II methanotrophs expressing sMMO are capable of oxidizing aromatic compounds (Colby et al., 1977; Uchiyama et al., 1989; Hanson and Hanson, 1996; Sullivan et al., 1998; Lindner et al., 2000), GW60 and GW70 cultured without copper and testing positive in the sMMO assay, indicative of type II methanotroph presence, were tested for the oxidation of benzene (a common landfill contaminant) using oxygen uptake experiments as described by Lindner et al. (2000). Since toxic effects of benzene on methanotrophic whole cells have not been studied, concentrations over the range of 5 to 50 mg L -1 (64.1 – 641 M) were tested. Fresh stock solutions of benzene (Fisher Chemicals of Fisher Scientific, Fair Lawn, NJ, USA) were prepared (10 M ) using a carrier solvent 1,4-dioxane (Fisher Chemicals of Fisher Scientific, Fair Lawn, NJ, USA), not found to be oxidized by cells expressing sMMO and had no effect on the oxygen probe (Lindner et al., 2000). Cell suspensions of each culture were prepared for the experiments by harvesting cultures at -log phase by centrifugation at 1.94 g for 25 minutes in a J2-HS Beckman centrifuge (Beckman Coulter, Fullerton, CA, USA). Cells were washed in 5 mL NMS to remove any residual CH 4 , re-centrifuged and re-suspended to a final wet cell weight concentration of 0.2 g/mL. The final cell suspension was stored in an ice bath during the oxygen uptake experiments to preserve activity. A 1.9-mL glass, water-jacketed reactor (Figure 3-5) was used at a constant temperature of 30 o C to measure the rates of oxygen consumption over the tested benzene concentration range. An electrolyte and membrane-covered Clarke-type electrode (Instech Laboratories, Plymouth, MA, USA) was inserted into the reactor using a ground-glass port with two rubber o-rings and was connected to a biological oxygen monitor

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46 (Yellow Springs, Yellow Springs, OH, USA). Monitor output was sent to an A/D converter board (DAS08-PGL, Computer Boards, Mansfield, MA, USA) for data collection using Labtech Notebook software (Wilmington, MA, USA). In all assays, the reaction chamber was filled with an appropriate volume of NMS (totaling 1.9 mL after addition of cells and/or substrate) before the addition of cells or substrate. The electrode was calibrated daily using sodium sulfite (following manufacturer’s instructions) after application of fresh electrolyte and membrane. Runs for each concentration were conducted in triplicate. To verify oxygen uptake was occurring by MMO, acetylene was added to confirm inhibition of the MMO enzyme in the presence of methane. Other controls included runs with cells and 1,4-dioxane, NMS with 1,4-dioxane, NMS with benzene, and all rates were corrected for any observed endogenous metabolism. For each set of triplicate runs (i.e. before each benzene concentration change), 4 mL of methane was slowly bubbled into the reactor chamber with NMS medium and cell suspensions only (1.4 mM methane in solution). The rates of oxygen uptake in the presence of each concentration of benzene were divided by the average rate of oxygen uptake in the presence of methane previously measured, thus normalizing for the decrease in cell activity over time because of the decay of the MMO enzyme (Lindner et al., 2000). Oxygen uptake rates were calculated following the methods presented by Hitchman (1978) based on the following equation, 2%DOCVreactormRate (mol s -1 ), where m = slope of the oxygen uptake curve after addition of either methane or benzene, Vreactor = reactor volume of 1.9 mL, C = concentration of oxygen in water at a specified temperature, or 7.6 g mL -1 for 30C, %DO = the number of divisions measured during calibration of the probe, and 2 converts

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47 g atoms of oxygen to mol of molecular oxygen. Outliers were discarded after performing statistical analysis (Grubb’s Test), and rates were reported with a standard error (Taylor, 1990). It is also known that both the particulate and soluble forms of MMO are capable of degrading trichloroethylene (TCE) and other chlorinated aliphatic compounds (Uchiyama et al., 1989; Brusseau et al., 1990; Henry and Grbic`-Galic`, 1991; Alvarez-Cohen et al., 1992; DiSpirito et al., 1992; Fox et al., 1990; Smith et al., 1997; Lontoh and Semrau, 1998). To this end, we tested GW60 and GW70 for TCE oxidation using oxygen uptake experiments at a concentration of 20 mg L -1 (147.1 M), not found to be toxic in a previous study (Alvarez-Cohen and McCarty, 1991). The same procedures described for benzene were followed for evaluation of TCE using oxygen uptake. Stage 4: Assessment of Response to Environmental Stressors Landfill environments are diverse by nature, and its environmental conditions change over the course of time, especially with seasonal variations (Hilger and Barlaz, 2002). Since these mixed enriched cultures were derived from a landfill, it was important to assess how these cultures would respond to a change in external conditions. Specifically, we were interested in how these mixed cultures respond to stressors, such as a change in methane concentration, temperature, pH, copper concentration and medium strength. All experiments included assessment of growth rates, methane depletion (with one exception), analysis of protein content and sMMO assay as previously described. Initial Methane Concentration A low and high methane concentration (10% and 60 to 70%) were selected to assess how the mixed cultures would respond to the changes in methane concentration that occur as landfill waste is broken down by microorganisms (previously discussed in

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48 Chapter 2). The low methane concentration (10%) represents initial phases when methane is first produced, whereas the high methane concentration (60 to 70%) represents previously reported methane levels detected after methane has been produced over time (Bogner et al., 1997a; Tammemagi, 1999; Schuetz et al., 2003) and is close to methane concentrations detected in the sample wells used in this study (Uz et al., 2003). Growth rate experiments were conducted for both low and high initial methane concentrations with an adjustment in the volume of headspace removed to accurately reach the desired methane concentrations. Several transfers were made for each culture at the varied methane concentrations to ensure growth was occurring before experimentation. The specific rates of methane oxidation were also assessed and calculated for each change in methane concentration. All samples were incubated under conditions previously described. Temperature Temperatures of 6 and 15C were selected as values representative of landfill environments during colder seasons (Boeckx et al., 1996; Schuetz et al., 2003). Also, higher temperatures of 41C and 45C were selected with the former temperature reflecting one of the well temperatures used in this study (Uz et al., 2003) and the latter reflecting temperature growth for Methylococcus capsulatus (Bath) (Whittenbury et al., 1970). Cultures were stored in a rotary shaker (Barnstead International, Melrose Park, IL, USA) at the appropriate experimental temperatures (270 rpm) with turbidity being monitored over a course of 14 to 20 days. Once growth was detected, several transfers were made to ensure stable growth before conducting both growth and methane depletion studies.

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49 The pH Values The pH values selected (4.8 and 8.8) fall within previously reported pH range occurring in a landfill (Whalen et al., 1990; Watson-Craik et al., 1992; Kightley et al., 1995; Boeckx and Van Cleemput, 1996; Boeckx et al., 1996; Czepiel et al., 1996; De Visscher et al., 1999; Tammemagi, 1999; Wise et al., 1999; Christophersen et al., 2000; Schuetz et al., 2003) which represent conditions when the pH changes during the decomposition phases previously described (Chapter 2). The pH of the NMS medium was adjusted to 4.8 based on methods described by Dedysh et al. (1998) using concentrated nitric acid instead of phosphoric acid. To achieve a higher pH (8.8), methods described by Khmelenina et al. (1999), using a sodium carbonate buffer added to the medium, were used to adjust the pH. Cultures were then immediately inoculated with the adjusted pH and allowed to grow for several days at the enrichment methane concentrations, temperature and rotation speed. Several transfers were made before growth and methane depletion studies. Copper Concentration and Media Strength The response to a change in copper concentration was evaluated for these mixed cultures at 0 and 10 M. GW60 and GW70 were grown in NMS medium both with and without copper at the enrichment conditions for methane, temperature and rotation speed. Several transfers were made to allow transition of the MMO enzyme in the presence of copper (either from soluble to particulate or vice versa) before growth and methane depletion studies. To evaluate the response to a nutrient scarce environment, the medium strength was diluted five-fold based on procedures described by Wise et al. (1999). Each of the 4 cultures were grown in the diluted media, allowing several transfers before

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50 measuring growth rates. Growth rates were determined, whereas methane depletion was not. 1. Isolation of Mixed Cultures from Landfill Samples From section 5 years older GW 60 From newer section GW 70NMS 30C 20% CH4 pH = 6.8 270 rpm 2. Characterization of Mixed Culture GW60a. Isolation of Heterotrophs and Phenotypic Studiesb. SEM of Mixed Culturec. Growth Ratesd. Methane Depletion Ratese. sMMO Assay f. Oxygen Uptake Analysis (Benzene, TCE) 3. Characterization of Mixed Culture GW70a. Isolation of Heterotrophs and Phenotypic Studiesb. SEM of Mixed Culturec. Growth Ratesd. Methane Depletion Ratese. sMMO Assay f. Oxygen Uptake Analysis (Benzene, TCE) 4. Environmental Effectsa.Initial Methane Concentrationb.Temperaturec.pHd.Copper Concentratione.Media Strength NMS 30C 30% CH4 pH = 6.8 270 rpm 1. Isolation of Mixed Cultures from Landfill Samples From section 5 years older GW 60 From newer section GW 70NMS 30C 20% CH4 pH = 6.8 270 rpm 2. Characterization of Mixed Culture GW60a. Isolation of Heterotrophs and Phenotypic Studiesb. SEM of Mixed Culturec. Growth Ratesd. Methane Depletion Ratese. sMMO Assay f. Oxygen Uptake Analysis (Benzene, TCE) 3. Characterization of Mixed Culture GW70a. Isolation of Heterotrophs and Phenotypic Studiesb. SEM of Mixed Culturec. Growth Ratesd. Methane Depletion Ratese. sMMO Assay f. Oxygen Uptake Analysis (Benzene, TCE) 4. Environmental Effectsa.Initial Methane Concentrationb.Temperaturec.pHd.Copper Concentratione.Media Strength 1. Isolation of Mixed Cultures from Landfill Samples From section 5 years older GW 60 From newer section GW 70NMS 30C 20% CH4 pH = 6.8 270 rpm 2. Characterization of Mixed Culture GW60a. Isolation of Heterotrophs and Phenotypic Studiesb. SEM of Mixed Culturec. Growth Ratesd. Methane Depletion Ratese. sMMO Assay f. Oxygen Uptake Analysis (Benzene, TCE) 3. Characterization of Mixed Culture GW70a. Isolation of Heterotrophs and Phenotypic Studiesb. SEM of Mixed Culturec. Growth Ratesd. Methane Depletion Ratese. sMMO Assay f. Oxygen Uptake Analysis (Benzene, TCE) 4. Environmental Effectsa.Initial Methane Concentrationb.Temperaturec.pHd.Copper Concentratione.Media Strength NMS 30C 30% CH4 pH = 6.8 270 rpm Figure 3-1. Flowchart of methodology used.

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51 GW 67 GW 68 GW 70 GW 71 GW 65 GW 63 GW 62 GW 58 GW 60 GW 67N Pond GW 67 GW 67 GW 68 GW 68 GW 70 GW 70 GW 71 GW 71 GW 65 GW 65 GW 63 GW 63 GW 62 GW 62 GW 58 GW 58 GW 60 GW 60 GW 67 GW 67N Pond Figure 3-2. Alachua County Landfill site map. Shaded circles indicate the representative sampling locations. Figure 3-3. Side-arm flask used in determining growth rates.

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52 Figure 3-4. Mininert system used in methane degradation experiments (Kjeldsen et al., 1997; Figure 2, pg. 1270).

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53 B A C D Figure 3-5. Oxygen uptake system. A) Overall system with reactor, biological oxygen monitor and monitor containing Labtech Notebook. B) Close up of reactor vessel with oxygen probe, water inflow and outflow. C) Top view of reactor system connected to biological oxygen monitor. D) Schematic of reactor vial.

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CHAPTER 4 CHARACTERIZATION AND ACTIVITY COMPARISONS OF METHANOTROPHIC-HETEROTROPHIC MIXED CULTURES DERIVED FROM A LANDFILL ENVIRONMENT: A MANUSCRIPT TO BE SUBMITTED TO JOURNAL OF APPLIED MICROBIOLOGY Introduction Despite recycling efforts, landfills receive roughly 61% of the 220 million metric tons of waste generated annually in the United States (US EPA, 1999). With this large amount of disposed solid waste, there are a variety of complex organic compounds available to microorganisms for substrate use. As a result, environmental conditions become heterogeneous and, in turn, support the creation of and interaction among microbial populations within their own niches (Senior, 1995). Municipal solid waste (MSW) degradation requires a coordinated effort of several trophic groups of bacteria (Hilger and Barlaz, 2002); however, these tightly linked interactions are not fully understood. Of specific interest in landfills are methanotrophic bacteria, microorganisms that are known to thrive in the oxygen-and methane-rich regions of landfill environments. These microorganisms have been implicated in removing significant quantities of anaerobically generated methane gas before its release to the atmosphere as a greenhouse gas (Mancinelli and McKay, 1985; Whalen et al., 1990; King, 1992; Jones and Nedwell, 1993; Kightley et al., 1995; Boeckx and Van Cleemput, 1996; Czepiel et al., 1996; Bogner et al., 1997b; Borjesson, 1998; De Visscher et al., 1999; Schuetz and Kjeldsen, 2003). Methanotrophs have also been shown to co-metabolically oxidize a variety of toxic compounds, including trichloroethylene (TCE) and other chlorinated aliphatic 54

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55 compounds (Uchiyama et al., 1989; Fox et al., 1990; Bowman et al., 1993; Lontoh and Semrau, 1998; Brusseau et al., 1990; Alvarez-Cohen et al., 1992; DiSpirito et al., 1992; Henry and Grbic-Galic, 1991; Smith et al., 1997; Han et al., 1999), benzene, and chlorinated biphenyls (Colby et al., 1977; Uchiyama et al., 1989; Hanson and Hanson, 1996; Sullivan et al., 1998; Lindner et al., 2000). In light of the rapid population growth and subsequent increase in the number of landfills in the United States, there is a resulting need for more efficient waste degradation, prevention of methane emissions to the atmosphere, and minimization of releases of toxic compounds (Hanson and Hanson, 1996; Kjeldsen et al., 1997). The potential role of methanotrophs in these processes is not well understood. Unfortunately, studying population diversity and activity directly in the field poses difficulty, and methods of directly linking population diversity and activity in landfill environments to that of isolated mixed cultures more easily studied in the laboratory are not well developed. Doing so would enable a more rapid, facile prediction of biodegradative activity of microbial populations in landfills, thus allowing better management of these systems. Previous work has reported that samples removed from two locations of a MSW landfill, differing in age by approximately five years, possessed differences in methanogenic and methanotrophic population diversity (Uz et al., 2003). While intending to provide a methodology for genotypic analysis of landfill samples and not a study linking environmental conditions to methanotrophic diversity, this work did report a predominance of type II methanotrophs in the samples removed from the older region. This result provided strong indication that the age and thus conditions in a landfill can be

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56 directly linked to population diversity. Building upon this previously reported work, our study sought to make first efforts in bridging the gap between the field and laboratory studies by isolating mixed methanotrophic-heterotrophic cultures from these landfill samples and comparing their population diversity, phenotypic characteristics, oxidation potential of toxic chemicals, and activity responses under a variety of conditions. Our specific goal was to first better understand differences in isolated landfill mixed cultures to ultimately allow a broader microbial landfill-laboratory study in the future. Here, we report the isolation of two stable mixed methanotrophic-heterotrophic cultures with significant differences in population phenotypes, potential for contaminant oxidation, and ability to respond to changes in methane concentration, temperature, pH, copper concentration and medium strength. Materials and Methods Enrichment and Isolation of Mixed Cultures The samples used in our study were obtained from a lined MSW landfill as previously described by Uz et al. (2003). All glassware used was acid-rinsed to avoid copper contamination as suggested by Bowman and Sayler (1994). Two grams of soil from each sample was added to a 250-mL Erlenmeyer flask (equipped with a rubber stopper and a glass wool-packed filling tube) containing 30 mL of nitrate mineral salts (NMS) medium (Whittenbury et al., 1970) with a metal chelator, FeEDTA (Henry and Grbic-Galic, 1990) and doubly deionized water (Barnstead International, Melrose Park, IL, USA). A sufficient amount of copper (10 M CuNO 3 ) previously shown to promote maximal pMMO activity (Shah et al., 1992) was added to the medium for GW70 to suppress any sMMO activity. Headspace was removed from flasks using a vacuum pump apparatus and replaced with an equivalent amount of highest purity methane (99.9%,

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57 Strate Welding Supply Co., Jacksonville, FL) to achieve a methane-to-air ratio of 30:70 (for GW60) and 20:80 (for GW70) to stimulate methanotrophic growth (Hanson et al., 1992) for routine culturing. Samples were incubated at 30C and shaken at 270 rpm in a reciprocal laboratory shaker (Barnstead International, Melrose Park, IL, USA). Routine transfers were made using a 10% inoculum once sufficient visible turbidity was detected at the aforementioned mentioned incubation conditions unless otherwise noted. After weekly transfers over the course of 3-years, both liquid cultures appeared stable, based on visual observations on solid agar medium (nutrient agar, Difco, Becton Dickinson and Company, Sparks, MD; NMS medium and Bacto agar, Becton Dickinson and Company, Sparks, MD) serial dilutions and spread plating were performed to isolate individual populations. Heterotrophs were isolated using nutrient agar stored at 30C in plastic sealable containers, whereas NMS plates were stored in air-tight desiccators at 30C under methane-to-air tensions previously described. Both containers were routinely swabbed with ethanol to prevent fungal contamination. Once growth was visually detected using a plate microscope on both solid medium, individual colonies were transferred onto fresh plates with routine transfers made thereafter. Growth on NMS plates was frequently checked for purity by streaking onto nutrient plates. Characterization of Isolated Mixed Cultures Morphological and phenotypic studies Pure, isolated, individual heterotrophs were subjected to visual observations in determining morphological characterization entailing colony size, time to full development, pigmentation, transmittance to light, surface consistency, margin edge, elevation, and shape. Growth on nutrient agar was assessed using an adapted method

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58 (Hrsak and Begonja, 1998). Further determination of nutrient requirements was assessed by growth occurring on different agars, peptone-yeast-extract and tryptic soy (Becton Dickinson and Company, Sparks, MD) and for their ability to use methane as a sole carbon source on NMS agar incubated under methane-to-air tensions of 30:70. Gram staining, catalase and oxidase assays were also performed. Scanning electron microscopy Both cultures were grown to late exponential phase and fixed with 2% glutaraldehyde for 1.5 hours, then micro-centrifuged at 8,000 x g for 1 minute. After being washed 3 times in 0.1 M cacodylate buffer (pH 7.2), samples were postfixed in 1% osmium tetroxide (OsO 4 ) over night at 4C. The samples were dehydrated in a graded ethanol series to 100% then washed twice in hexamethyldisilazane (HMDS) for 5 to 10 minutes and allowed to air dry. Samples were then mounted on Alcian blue-coated cover slips on SEM stubs, gold coated in a Denton Desk II sputter coater and examined with a Hitachi S-570 scanning electron microscope (Hitachi Co., Tokyo, Japan). Growth kinetics and methane oxidation rate parameters Growth curves were prepared and measured using a UV/VIS spectrophotometer (Milton Roy Company, Ivyland, PA, USA) at a 600 nm wavelength. The initial optical density (OD) of each sample was adjusted to 0.2 (Brusseau et al., 1990) using NMS medium (either with or without copper, depending on culture). Samples were prepared in triplicate plus one control, which contained no methane. Time zero measurements were made after inoculation with methane and subsequent absorbance readings were recorded every few hours over the course of several days to create a growth curve. Specific growth rates and generation times were calculated using linear fits to the slope of the exponential growth phase of each growth curve obtained based on the methods described

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59 by Hrsak and Begonja (1998). Samples were also grown to mid-exponential phase where the proportion of methanotrophs and heterotrophs in both mixed cultures was determined by methods described by Hrsak and Begonja (1998). Experiments for methane oxidation were conducted using 60 mL glass bottles sealed with Mininert valves (VICI Precision Sampling Inc., Baton Rouge, LA). Samples were run in triplicate with each bottle containing NMS media and liquid culture (with adjusted optical density of 0.2) that maintained 78% volume in headspace and 5:1 ratio of media-to-culture. Controls were set up to detect the amount of methane lost during experimentation with NMS media and no cells. A calculated volume of headspace was removed from each vial using a hypodermic needle and syringe and replaced with an equivalent volume of methane that was injected through a sterile filter attached to a separate needle and syringe. A 3-point calibration curve was created for each set of experiments using standards at 10%, 30% and 50% methane. Headspace was removed (60 L) using a gas-tight Hamilton syringe (Hamilton Company, Reno, NV, USA) from each sample and injected into a Shimadzu GC-14A gas chromatograph (GC) equipped with a flame ionization detector connected with a Chromatopac C-R501 integrator. The GC column was stainless steel (ca. 0.317 by 41 cm) packed with Porapak Q (80/100 mesh; Waters Associates, Inc., Framington, MA.). Nitrogen was used as the carrier gas and analyses were carried out under the following conditions: column temperature 50C and injector and detector temperature 180C. Standard samples were run in triplicate at each concentration. Headspace samples from bottles were measured for each sample at time zero and every few hours thereafter over the course of a couple days or until methane oxidation ceased. Specific methane oxidation rates were calculated using linear

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60 regression fits during the phase of methane oxidation as previously described by Hrsak and Begonja (1998). Any detected losses in the controls were accounted for in the rates. Detection of sMMO activity, at the end of each growth and methane oxidation experiment, involved incubation of suspended cultures with naphthalene crystals and subsequent addition of o-dianisidine as modified from Brusseau et al. (1990). Assays were run in triplicate plus a heat-killed control and an acetylene control known to inhibit MMO. Degradation potential in the presence of landfill contaminants Since it is known that type II methanotrophs expressing sMMO are capable of degrading aromatic compounds (Colby et al., 1977; Uchiyama et al., 1989; Hanson and Hanson, 1996; Sullivan et al., 1998; Lindner et al., 2000), GW60 and GW70 cultured without copper and showing sMMO activity, indicative of most type II methanotroph presence, were tested for potential to oxidize a range of concentrations of benzene (a common landfill contaminant) using oxygen uptake experiments (Lindner et al., 2000). The particulate and soluble forms of MMO are capable of degrading trichloroethylene (TCE) and other chlorinated aliphatic compounds (Uchiyama et al., 1989; Brusseau et al., 1990; Fox et al., 1990; Henry and Grbic-Galic, 1991; Alvarez-Cohen et al., 1992; DiSpirito et al., 1992; Bowman et al., 1993; Smith et al., 1997; Lontoh and Semrau, 1998; Han et al., 1999). To this end, we tested GW60 and GW70 expressing both forms of MMO for their oxidation potential with 20 mg L -1 (147.1 M) TCE using oxygen uptake experiments. This concentration was not found to be toxic in a previous study (Alvarez-Cohen and McCarty, 1991). Fresh stock solutions of benzene and TCE (Fisher Chemicals of Fisher Scientific, Fair Lawn, NJ, USA) were prepared (10 M ) using a carrier solvent 1,4-dioxane (Fisher Chemicals of Fisher Scientific, Fair Lawn, NJ, USA)

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61 not found to be oxidized by cells expressing sMMO and had no effect on the oxygen probe (Lindner et al., 2000). Cell suspensions of each culture were prepared for the experiments by harvesting cultures at -log phase by centrifugation at 1.94 g for 25 minutes in a J2-HS Beckman centrifuge (Beckman Coulter, Fullerton, CA, USA). Cells were washed in 5 mL NMS to remove any residual methane, re-centrifuged and re-suspended to a final wet cell weight concentration of 0.2 g mL -1 . The final cell suspension was stored in an ice bath during the oxygen uptake experiments to preserve activity. A 1.9-mL glass, water-jacketed reactor was used at a constant temperature of 30 o C to measure the rates of oxygen consumption over the tested benzene concentration range. An electrolyte and membrane-covered Clarke-type electrode (Instech Laboratories, Plymouth, MA, USA) was inserted into the reactor using a ground-glass port with 2 rubber o-rings and was connected to a biological oxygen monitor (Yellow Springs, Yellow Springs, OH, USA). Monitor output was sent to an A/D converter board (DAS08-PGL, Computer Boards, Mansfield, MA, USA) for data collection using Labtech Notebook software (Wilmington, MA, USA). Runs for each concentration were conducted in triplicate. To verify oxygen uptake was occurring as a result of MMO activity, acetylene controls were used. Other controls included runs with cells and 1,4-dioxane, NMS with 1,4-dioxane, NMS with the substrate (either benzene or TCE), and all rates were corrected for any observed endogenous metabolism. For each set of triplicate runs (i.e. before each change in benzene or TCE concentration), 4 mL of methane was slowly bubbled into the reactor chamber with NMS medium and cell suspensions only (1.4 mM methane in solution). The rates of oxygen uptake in the

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62 presence of each concentration of benzene or TCE were divided by the average rate of oxygen uptake in the presence of methane previously measured, thus normalizing for the decrease in cell activity over time because of the decay of the MMO enzyme (Lindner et al., 2000). Oxygen uptake rates were calculated following the methods presented by Hitchman (1978) with outliers discarded after performing statistical analysis (Grubb’s Test) with a reported standard error (Taylor, 1990). Effects of Environmental Stressors To account for temporal variations in landfills, the effects of change in methane concentration, temperature, pH, copper concentration and medium strength on the mixed cultures’ growth rate and methane depletion potential, and sMMO activity were evaluated. A low and high methane concentration (10% and 60 to 70%) was selected to assess how the mixed cultures would respond to the changes in the range of methane concentrations that can exist in landfill environments as microbial populations develop (Hilger and Barlaz, 2002). The low methane concentration (10%) represents conditions in the initial phases of landfill development when methane is first produced, whereas the high methane concentration (60 to 70%) represents conditions during more advanced stages of landfill development as previously reported (Bogner et al., 1997a; Tammemagi, 1999; Schuetz et al., 2003). This value also closely matches methane concentrations detected in the wells from which samples were removed for this study (Uz et al., 2003). Measurement of growth and methane depletion was conducted under both low and high initial methane concentrations with an adjustment in the volume of headspace removed to accurately reach the desired methane concentrations as previously described. Several transfers were made for each culture at the respective methane concentrations.

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63 Temperatures of 6 and 15C were selected to represent low temperatures occurring in landfill environments during colder seasons (Boeckx et al., 1996; Schuetz et al., 2003). Also, higher temperatures of 41C and 45C were selected, with the former temperature reflecting one of the well temperatures used in this study and the latter reflecting the temperature growth optimum for Methylococcus capsulatus (Bath), a well-characterized type X methanotrophs (Whittenbury et al., 1970). Cultures were incubated in a rotary shaker (Barnstead International, Melrose Park, IL, USA) at the appropriate experimental temperatures (270 rpm). Once turbidity was detected (over 14 to 20 days), several transfers were made to ensure stable growth before conducting both growth and methane depletion studies. The pH values selected (4.8 and 8.8) fall within the range reported to occur in a landfill during decomposition (Whalen et al., 1990; Watson-Craik et al., 1992; Kightley et al., 1995; Boeckx and Van Cleemput, 1996; Boeckx et al., 1996; Czepiel et al., 1996; De Visscher et al., 1999; Tammemagi, 1999; Wise et al., 1999; Christophersen et al., 2000; Hilger and Barlaz, 2002; Schuetz et al., 2003). Methods to adjust pH to acidic and basic levels were those outlined in Dedysh et al. (1998) and Khmelenina et al. (1999). The response to a change in copper concentration was evaluated for both mixed cultures at 0 and 10 M. GW60 and GW70 were grown in NMS medium both with and without copper at the previously described enrichment conditions for methane, temperature and rotation speed. Several transfers were made to allow expression of either form of MMO. To evaluate the response to a nutrient-scarce environment, the medium strength was diluted five-fold based on procedures described by Wise et al.

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64 (1999). Each of the 4 cultures was grown in the diluted medium, allowing several transfers before measuring growth rates. Results Isolation of Mixed Cultures Increased turbidity in the initial landfill liquid cultures was visible after approximately 7 days of incubation. Subsequent routine transfers of the stable cultures showed significant growth in approximately 3 days after inoculation. Heterotrophic growth on nutrient agar plates was rapid, with visible colony formation after 1 day. The mixed culture isolated from landfill sample GW60 originally possessed 4 heterotrophs that were consistently observed on nutrient agar plates. Of note, one of these colonies was eventually “lost” after repeated transfers of the liquid mixed culture over a period of several weeks. The mixed culture isolated from landfill sample GW70 originally possessed 5 heterotrophic colonies, but also one heterotroph was “lost” after repeated transfers. The enriched mixed cultures harvested in the mid-exponential phase also showed that methanotrophs out-numbered heterotrophs based on plate counts. For GW60 grown with copper, methane-oxidizers accounted for 2.3x10 8 cfu mL -1 , and heterotrophic populations were 1.7x10 7 cfu mL -1 . For GW70 grown with copper, methane oxidizers accounted for 1.1x10 10 cfu mL -1 (on NMS with copper agar plates) and heterotrophic populations were 2.5x10 8 cfu mL -1 (on nutrient agar plates). Characteristics of the Enriched, Mixed Methanotrophic-Heterotrophic Communities Morphological and phenotypic characteristics Table 4-1 shows the growth and nutritional characteristics of the isolated heterotrophic populations when cultured on different solid media over an 8-day period.

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65 Colony numbers 2 from GW60 and 4 from GW70 were initially isolated and characterized, but both colonies were lost. Nonetheless, colony number 4 was resilient enough to be subjected to further phenotypic studies before it was lost. Heterotrophs in both mixed cultures showed varying growth ability on peptone-yeast extract and tryptic soy agar, whereas, with the exception of one colony, all isolated colonies grew best on nutrient agar. No colonies grew sufficiently with methane as the sole carbon and energy source. Morphological observations were monitored at 30C over a 20-day period, where full development typically occurred between 3 to 4 days. Both mixed cultures showed diversity in heterotroph size, pigmentation, transmittance to light, surface consistency, margin edge, elevation and shape when grown on nutrient agar (Table 4-1). Each isolated heterotroph was further subjected to phenotypic studies where all heterotrophs tested Gram-negative, catalase and oxidase positive. Cellular characteristics Examination of both mixed methanotrophic-heterotrophic communities by scanning electron microscopy (SEM) revealed a diverse array of cellular shapes and sizes (Figure 1). Cells in greatest number in both cultures possessed a rod-shape with length ranging from 1.99 to 3.53 m and diameter from 0.75 to 1.04 m. Less populated cells were cocci (diameter = 0.74 to 0.97 m), coccobacillus (length = 2.21 to 2.74 m, diameter = 1.18 to 1.21 m) and prosthecate (length = 7.04 to 8.31 m, diameter = 0.89 to 0.94 m) in shape. Sizes represent average ranges that were measured directly from the SEM photographs. A common occurrence in both cultures was the appearance of flattened cells with a concave-shaped center; however, similar observations have been reported by Dedysh et al. (1998) due to sample preparation.

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66 Evaluation of activity based on growth and methane oxidation rate Both cultures tested positive for sMMO activity when cultured without copper and negative when cultured with 10 M copper. At the enrichment conditions, growth and specific methane oxidation rates revealed that when cultured without copper GW60 exhibited a similar growth rate (0.83 0.05 d -1 ), longer generation time (8.7 hrs) and lower rate of methane oxidation (0.14 0.021 d -1 ) than GW70 (0.91 0.21 d -1 , 7.9 hrs, and 0.29 0.09 d -1 , respectively). When cultured with copper, GW60 exhibited a similar growth rate (0.79 0.08 d -1 ), shorter generation time (9.2 hrs) and lower rate of methane oxidation (0.20 0.01 d -1 ) than GW70 (0.69 0.03 d -1 , 10.5 hrs, and 0.44 0.01 d -1 , respectively). Degradation potential for landfill contaminants based on oxygen uptake analysis Over this range (64.1 to 641 M), the highest observed rate of consumption of oxygen in the presence of benzene occurred at 320.5 M (25 mg L -1 ), implying that oxidation of this compound may be occurring. To accurately compare both cultures for degradation potential, GW70 was also grown without copper and evaluated at this same concentration. GW70 showed rates approximately 7 times lower for oxygen consumption than GW60 (Table 4-2). Probe effects were observed at concentrations near or above 641 M (50 mg L -1 ), masking the observed activity. Both cultures, GW60 and GW70, grown in the presence of copper showed no consumption of oxygen in the presence of benzene at the tested concentration of 320.5 M. The highest observed rate of consumption of oxygen in the presence of TCE was for GW60 grown without copper , 0.72 0.01 (Table 4-2). In comparison to the other cultures grown either with or without copper, this rate was more than double the rate than GW60 grown with copper, 2.5 times the rate of GW70 (with copper) and 3 times the

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67 rate of GW70 (without copper). GW70 exhibited slightly higher rates of oxygen consumption in the presence of TCE when grown with copper versus GW70 without. Assessment of Communities’ Response to Environmental Stressors With the change in initial methane concentration, growth rates shown in Figure 4-2 revealed a slightly higher growth rate for GW60 at the lowest methane concentration. At this same methane concentration, GW70 did not consistently grow resulting in large deviations between samples, whereas at 20% methane, growth appeared to be highest overall. Overall, GW60 appeared to have a slightly higher growth rates than GW70. For rates of methane oxidation, Figure 4-3 showed no significant difference between rates for GW60. On the other hand, GW70 showed no methane oxidation at 10%, whereas the highest observed rate occurred at 20% methane (rates were twice as fast than at 60-70% methane). Overall, it appeared that GW70 was more sensitive to a change in methane concentration as evidence by the significant change in its methane oxidation rate. GW70 also demonstrated the highest overall methane oxidation rate in comparison to GW60. GW60 was capable of sustainable growth with a temperature change to 15C but was not able to grow at a 6C or at a higher temperature of 41C (Figure 4-4), whereas the moderate temperature of 30C yielded the highest growth rate. On the other hand, GW70 was able to grow at all three reported temperatures (but not at 6C), where the lowest growth rate occurred at 15C and the highest at 30C. At the moderate temperature, where the highest observed growth rates occurred in both samples, GW60 showed slightly enhanced growth rates in comparison to GW70. Overall, a similar trend was observed in both cultures, where 30C yielded the highest growth rates. In terms of methane oxidation (Figure 4-5), GW60 showed oxidation at both 15 and 30C, with the highest rate occurring at 30C. GW70 demonstrated ability to oxidize methane at all

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68 three reported temperatures with the lowest rate at 15C and comparable rates at both 30 and 41C. Overall, the same trend was observed in both samples where the moderate temperature of 30C yielded the highest methane oxidation rates. Both cultures showed adaptation to a change in pH regime as evidenced by growth occurring at all three pH levels (Figure 4-6). GW60 exhibited the highest observed rates of growth and methane oxidation at a near neutral pH, whereas the lowest growth occurred at a pH of 8.8. Similarly, GW70 also exhibited the highest observed growth rate at a near neutral pH, whereas lowest observed growth was at a pH of 4.8. These results show a clear trend with pH affecting growth rates, with near neutral levels yielding the highest growth rates. A similar trend was observed for rates of methane oxidation (Figure 4-7). Although not as drastic of a change between rates with a change in pH, GW60 exhibited the highest rate at near neutral pH. GW70 showed larger differences in methane oxidation with a change in pH, where near neutral yielded the highest observed rate. Figure 4-8 shows the effect of copper on growth rates. Based on these results, there was no significant difference in growth rates when grown with or without the presence of copper. However, there was a difference in the rates of methane oxidation when grown with or without copper (Figure 4-9). In both cultures, when grown in the presence of copper, there was an increase in the rate of methane oxidation, where GW70 showed the highest overall rate. The effect of medium strength on growth was also studied (Figure 4-10). These results indicate that, when both cultures were grown without copper, the full strength medium yielded higher growth rates. The opposite trend

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69 was observed (Figure 4-11) when both cultures were grown with copper. In this case, the full strength medium resulted in a decrease in growth rate. Discussion Characterization of the Enriched, Mixed Methanotrophic-Heterotrophic Communities Morphological and phenotypic characteristics of isolated heterotrophs Both mixed cultures displayed unique and varied heterotrophic populations based on size, pigmentation and growth on various media types. It was expected that all heterotrophs grew well on the nutrient-rich agar, since they are derived from a nutrient-rich environment. Varied growth with the oligotrophic media (peptone yeast extract and tryptic soy agar) showed the ability of individual heterotrophs to adapt to changes in nutrient supply, with some adapting better to a lower supply (as indicated by abundant growth) than others (showing weak growth). Cellular characteristics Similar shapes of cells were detected in both samples, although the sizes of these shapes varied. One characteristic shape was the coccobacillus with tails (or prosthecates), which appear to resemble Hyphomicrobium or Caulobacter. Hyphomicrobium has been reported in other studies working with enriched, mixed cultures containing methanotrophs (Alvarez-Cohen, 1992; Dedysh et al., 1998; Dunfield et al., 1999; Wise et al., 1999). The common occurrence of this species with methanotrophs leads to a higher probability that this prosthecate-shaped bacterium resembles Hyphomicrobium. It could be hypothesized that a mutualistic or synergistic relationship exists between Hyphomicrobium and methanotrophs since Hyphomicrobium can grow well on methanol (Anthony, 1982) and may be utilizing the methanol

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70 intermediate byproduct produced as methanotrophs convert methane into carbon dioxide. In both samples, the rod-shaped cells dominated in numbers compared to other cell shapes. These cells are expected to be methanotrophs due to the selective enrichment conditions using in culturing the samples. Degradation potential of landfill contaminants based on oxygen uptake analysis Initial screening of these mixed cultures showed the ability of both cultures to oxidize methane. Both cultures tested positive for the presence of sMMO when grown without copper, further supporting the presence of methanotrophs, since negative results were demonstrated with the addition of acetylene, a known MMO inhibitor. Both cultures were screened for their ability to oxidize benzene (an aromatic compound) and/or TCE (a chlorinated aliphatic compound). Under sMMO expression, both cultures showed oxygen uptake activity in the presence of benzene. No activity was observed when the cultures were grown under conditions testing negative for sMMO activity. This behavior is consistent with that of most type II methanotrophs (Colby et al., 1977; Brusseau et al., 1990; Uchiyama et al., 1989; Lindner et al., 2000). Significantly higher rates of oxygen consumption in the presence of benzene were observed in GW60 versus GW70. When both cultures were grown with and without copper, consumption of oxygen in the presence of 20 mg L -1 (147.1 M) TCE was observed. Copper-deprived GW60 appeared to more efficiently oxidize TCE than both GW60 and GW70 grown with copper, which is supported by other studies that have reported faster TCE oxidation rates by sMMO compared to pMMO (Little et al., 1988; Henry and Grbic-Galic, 1991; Sullivan et al., 1998). When comparing rates of TCE oxidation to benzene oxidation, both cultures exhibited higher rates of TCE oxidation than benzene. These results are

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71 supported by a previous report demonstrating more complete degradation of TCE than benzene (Uchiyama et al., 1989). Assessment of Communities’ Response to Environmental Stressors Both mixed cultures responded very differently when cultured under varied methane concentrations in the headspace. GW60 adapted well to the change in initial methane concentration where it grew best at both the lowest and highest methane concentrations tested (10 and 60 to 70%), whereas its rate of methane oxidation was not affected with the change in methane concentration. Contrastingly, GW70 preferred its enrichment condition at 20% methane as evidenced by the highest rates of growth and methane oxidation at this concentration. It is not known whether the poor growth and no observed methane oxidation activity were due to inability of GW70 to adapt to the change in methane concentration or due to experimental error. Overall, in can be inferred from these results that the change in methane concentration is not as significant of a factor effecting growth, whereas it does have some effect on methane oxidation. This was primarily observed in mixed culture GW70 which showed more sensitivity to the change in methane concentration. Temperature had a significant effect on the rates of growth and methane oxidation on these mixed cultures. It would appear overall, that the temperature yielding the highest growth and methane oxidation rates occurred at 30C. This temperature falls within the range of other optimum temperatures previously reported for methane oxidation (Whalen et al., 1990; Visvanathan et al., 1999). Although both cultures displayed the ability to adapt to a 15C temperature decrease, these rates of methane oxidation were much slower (5 to 20 times) as reported by another study that observed a significant decrease methane degradation rates with a 15C temperature decrease

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72 (Kjeldsen et al., 1997). Growth and methane oxidation was completely inhibited at 45C for both GW60 and GW70 where this temperature has also been previously reported to inhibit methane oxidation in a landfill cover soil (Visvanathan et al., 1999). Overall, based on these results, temperature plays a significant role in influencing both growth and methane oxidation rates. This also implies that the temperature fluctuations associated with seasonal variations will affect these rates as well. Both cultures clearly demonstrated sensitivity to a change in pH, although both were capable of partially adapting, as evidence by growth and methane oxidation occurring at all tested pH levels. Partial adaptation to acidity for methane consumption has also been reported by Dunfield et al. (1993). Near neutral pH appears to support optimal growth and methane oxidation for both cultures which is expected since most methanotrophs prefer neutral pH for growth (Graham et al., 2002) as do most microorganisms (Caldwell, 1995). These results suggested that these mixed methanotrophic communities are only partially adaptable to a change in pH regime, indicating the significance of pH in affecting growth and methane oxidation rates. The observed shift in sMMO expression with the shift in copper concentration for GW60 and GW70 was expected due to the presence of copper suppressing the sMMO enzyme (Prior and Dalton, 1985; Chan et al., 1993; Sullivan et al., 1998; Graham et al., 2002). This shift in copper appeared to have no affect on growth rates in full strength medium for GW60, but did enhance the growth rate of GW70 with the sMMO expression yielding a higher rate. The shift in sMMO expression (from positive to negative) did give rise to an increase in methane oxidation for both cultures indicating that the presence of copper increases the rate of methane oxidation. This trend is supported by previously

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73 published information with copper enhancing the degradation of methane (Boisesen et al., 1993; Hanson and Hanson, 1996; Sullivan et al., 1998; Graham et al., 2002). This implies that copper contributes to the assimilation of methane in the oxidation pathway leading to higher oxidation rates (Graham et al., 2002). Growth rates were compared at the full versus diluted media to see how this culture responded to an oligotrophic environment. Based on these results, it is implied that enriched methanotrophic cultures grown in the presence of copper grow best under nutrient-limited conditions (diluted media) and are capable of scavenging nutrients more efficiently than those expressing sMMO in similar environments. This further implies that cultures testing negative for sMMO can thrive better under oligotrophic conditions and out-compete sMMO positive dominant cultures. On the other hand, methanotrophic enriched cultures expressing sMMO prefer nutrient rich environments and thrive best at those conditions which has been reported for sMMO preferring higher nutrients (Hanson and Hanson, 1996; Borjesson et al., 1998). Conclusions The broad focus of our study was to take initial steps in bridging the gap between field and laboratory studies by isolating mixed methanotrophic-heterotrophic cultures from landfill samples. This was accomplished through the isolation of stable, enriched, mixed cultures derived from a landfill. Subsequently, these mixed cultures were studied in the laboratory and compared for their phenotypic characteristics, growth and methane depletion rates, potential for hazardous chemical degradation and their response to environmental stressors. Our study revealed that these two mixed enriched methanotrophic-heterotrophic cultures showed diversity in terms of colony morphology, nutrient requirements, phenotypic characteristics and cellular shapes and sizes. They also

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74 demonstrated the ability to use methane and showed potential for oxidizing landfill contaminants. Further investigation of these enriched mixed cultures revealed their ability to respond and adapt to environmental stressors. Specifically, the change in methane concentration did not play as significant of a role in controlling growth and methane oxidation rates, as did temperature and pH. Other factors that did influence activity, although not as significant as temperature and pH, were copper concentration and nutrient availability. It is important to consider these parameters and the associated changes that occur in a landfill since they influence the activity of the microbial populations present. Before we can understand the complex processes naturally occurring in the environment, we have to understand what happens in a more controlled environment. As the conditions change in a landfill, which is common as a landfill ages and with seasonal influences, so do the microbial communities respond and adapt to these changes. By studying characterization, activity and responses, we take one step closer to understanding what may potentially be happening in the natural environment. Our study shows promise for linking environmental conditions occurring in the field to how it translates and effects the microbial populations present, by studying them more easily in the laboratory. Our study serves as a springboard for initiating more in-depth studies of microbial isolates derived from landfill environments. Ultimately, we hope that this and future studies will then provide a better understanding of what complex processes occur in landfill environments that will encourage more efficient management of these environments to minimize environmental damage.

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Table 4-1. Characteristics of isolated heterotrophs on selected media Growth b Culture Morphological Observations a NA PY TSA No. 1 3 mm, white-cream, opaque, glistening, smooth, convex, circular ++ ++ ++ No. 2 < 1 mm, beige-glass, translucent, glistening, irregular, flat, circular Not transferable over time No. 3 1 mm, orange, translucent, glistening, smooth, convex, circular +/++ -+ -+ GW60 No. 4 2 mm, beige-cream, translucent, glistening, smooth, convex, circular ++ ++ ++ No. 1 4 mm, white-cream, translucent, glistening, undulate, flat, irregular ++ ++ ++ No. 2 1-2 mm, brownish-orange, opaque, glistening, smooth, convex, circular ++ +/++ ++ No. 3 < 1 mm, beige-glass, translucent, glistening, smooth, convex, circular -+ ++ + No. 4 < 1 mm, creamy-pink, opaque, glistening, smooth, convex, circular Not transferable over time GW70 No. 5 1 mm, beige-cream, translucent, glistening, smooth, convex, circular ++ -+ -+ 75 NA, Nutrient agar; PY, peptone-yeast-extract agar; TSA, tryptic soy agar. -+ weak growth; + good growth; ++ abundant growth a monitored over a 20-day time period b monitored over an 8-day time period

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76 Flattened appearance with concave center prosthecates Flattened appearance with concave center prosthecates Figure 4-1. Cellular diversity of mixed methanotrophic-heterotrophic communities. A) 3000x magnification, bar represents 5 m, GW60. B) 5000x magnification, bar represents 1 m, GW60. C) 3000x magnification, bar represents 5 m, GW70. D) 2000x magnification, bar represents 5 m, GW70.

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77 Table 4-2. Comparison of TCE and benzene oxidation potential for mixed cultures. Oxygen Uptake Rate ( standard error) a Culture TCE Benzene Without Cu 2+ 0.72 (0.01) 0.30 (0.04) GW60 With Cu 2+ 0.32 (0.06) ND Without Cu 2+ 0.22 (0.001) 0.04 (0.03) GW70 With Cu 2+ 0.27 (0.02) ND a Oxygen uptake rates are normalized to oxygen uptake in the presence of the substrate (TCE at 147.1 M or Benzene at 320.5 M) relative to oxygen uptake in the presence of methane. ND, not detected.

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78 00.20.40.60.811.21.410203060-70Percentage of MethaneRate (d-1) GW60 GW70 Figure 4-2. Effect of methane concentration on growth rates of mixed cultures. Rates were not determined for GW60 at 20% or GW70 at 30% methane. Both cultures were grown in the presence of copper. 00.10.20.30.40.510203060-70Percentage of MethaneRate (d-1) GW60 GW70 Figure 4-3. Effect of methane concentration on methane oxidation rates of mixed cultures. Rates were not determined for GW60 at 20% or GW70 at 30% methane. Both cultures were grown in the presence of copper.

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79 00.10.20.30.40.50.60.70.80.91153041Temperature (oC)Rate (d -1) GW60 GW70 Figure 4-4. Effect of temperature on growth rates of mixed cultures. Growth was not detected for GW60 at 41C. Both cultures were grown in the presence of copper. 00.10.20.30.40.50.6153041Temperature (oC)Rate (d -1) GW60 GW70 Figure 4-5. Effect of temperature on methane oxidation rates of mixed cultures. Growth was not detected for GW60 at 41C. Both cultures were grown in the presence of copper.

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80 00.10.20.30.40.50.60.70.80.914.86.88.8pHRate (d -1) GW60 GW70 Figure 4-6. Effect of pH on growth rates of mixed cultures. Both cultures were grown in the presence of copper. 00.10.20.30.40.54.86.88.8pHRate (d -1) GW60 GW70 Figure 4-7. Effect of pH on methane oxidation rates of mixed cultures. Both cultures were grown in the presence of copper.

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81 00.20.40.60.811.2010Copper Concentration (M)Rate (d-1) GW60 GW70 Figure 4-8. Effect of copper on growth rates of mixed cultures. 00.10.20.30.40.5010Copper Concentration (mM)Rate (d-1) GW60 GW70 Figure 4-9. Effect of copper on methane oxidation rates of mixed cultures.

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82 00.20.40.60.811.20.21Medium StrengthRate (d -1) GW60 GW70 Figure 4-10. Effect of medium strength on growth rates of mixed cultures grown without the presence of copper. Medium diluted five-fold (0.2) and non-diluted, full-strength medium (1.0). 00.20.40.60.811.21.41.60.21Medium StrengthRate (d -1) GW60 GW70 Figure 4-11. Effect of medium strength on growth rates of mixed cultures grown with the presence of copper. Medium diluted five-fold (0.2) and non-diluted, full-strength medium (1.0).

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CHAPTER 5 CONCLUSIONS The broad focus of this research project was to take initial steps in bridging the gap between field and laboratory studies by isolating mixed methanotrophic-heterotrophic cultures from landfill samples. This was accomplished through the isolation of stable, enriched, mixed cultures derived from a landfill. Subsequently, these mixed cultures were studied in the laboratory and compared for their phenotypic characteristics, growth and methane depletion rates, potential for hazardous chemical degradation and their response to environmental stressors. This work revealed that these two mixed enriched methanotrophic-heterotrophic cultures showed diversity in terms of colony morphology, nutrient requirements, phenotypic characteristics and cellular shapes and sizes. They also demonstrated the ability to utilize methane and showed potential for oxidizing landfill contaminants. Further investigation of these enriched mixed cultures revealed their ability to respond and adapt to environmental stressors. Specifically, the change in methane concentration did not play as significant of a role in controlling growth and methane oxidation rates, as did temperature and pH. Other factors that did influence activity, although not as significant as temperature and pH, were copper concentration and nutrient availability. It is important to consider these parameters and the associated changes that occur in a landfill since they influence the activity of the microbial populations present. Because of unreasonable constraints of economics, time or access to sites, understanding the complex processes naturally occurring in the environment must always 83

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84 be gained by initial studies in a more controlled environment. As the conditions change in a landfill, because of aging and seasonal influences, so do the microbial communities respond and adapt to these changes. By studying characterization, activity and responses of cultures isolated form landfill samples, we take one step closer to understanding what may potentially be happening in the natural environment. From two different locations in a landfill, two mixed cultures were enriched from these soil samples which showed diversity and differences in activity. The original soil samples showed the presence of both type I and II methanotrophs with a predominance of type II methanotrophs in GW6), whereas GW70 showed a predominance of type I. Similarly, after 3 years of routine transfers, these stable mixed cultures show similar presence of methanotrophs, where GW60 shows type II predominance and GW70 shows type I, although both types are present in both cultures. These initial results show promise for linking environmental conditions occurring in the field to how it translates and effects the microbial populations present, by studying them more easily in the laboratory and warrants a broader study to assess in a statistically meaningful way, the variations in microbial populations to the diverse conditions in a landfill. Ultimately, we hope that this and future studies will subsequently provide a better understanding of what complex processes occur in landfill environments that will encourage more efficient management of these environments to minimize environmental damage.

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APPENDIX A CALCULATED CONVERSIONS FOR METHANE OXIDATION RATES FROM PREVIOUS PUBLICATIONS Table A-1. Summary of methane oxidation rates and their converted units. Author Reported rate Reported units Converted rate Converted units Comment Whalen et al. (1990) 18 g CH 4 (g-soil) -1 d -1 0.05 mol g -1 h -1 Measured substrate-saturated CH 4 oxidation rate of cover soil at 5C Whalen et al. (1990) 45 g CH 4 (g-soil) -1 d -1 0.1 mol g -1 h -1 Measured substrate-saturated CH 4 oxidation rate of cover soil at 15C Whalen et al. (1990) 75 g CH 4 (g-soil) -1 d -1 0.2 mol g -1 h -1 Measured substrate-saturated CH 4 oxidation rate of cover soil at 31C (max.) Boeckx and Van Cleemput (1996) 2.36 ng CH 4 (g-soil) -1 h -1 1.5 x 10 -4 mol g -1 h -1 Measured CH 4 oxidation rate of cover soil at 25C (max. range) Boeckx and Van Cleemput (1996) 2.19 ng CH 4 (g-soil) -1 h -1 1.4 x 10 -4 mol g -1 h -1 Measured CH 4 oxidation rate of cover soil at 30C (max. range) Boeckx et al. (1996) 12 ng CH 4 (g-soil) -1 h -1 7.5 x 10 -4 mol g -1 h -1 Measured CH 4 oxidation rate of cover soil at 20C (max. range) 85

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86 Table A-1. Continued Author Reported rate Reported units Converted rate Converted units Comment Boeckx et al. (1996) 11 ng CH 4 (g-soil) -1 h -1 6.9 x 10 -4 mol g -1 h -1 Measured CH 4 oxidation rate of cover soil at 25C (max. range) Czepiel et al. (1996) 200 nmol g -1 h -1 0.2 mol g -1 h -1 A generalized model of temperature response of CH 4 oxidation of cover soils at 5C Czepiel et al. (1996) 4400 nmol g -1 h -1 4.4 mol g -1 h -1 A generalized model of temperature response of CH 4 oxidation of cover soils at 36C (max.)

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APPENDIX B GROWTH MEDIA USED FOR ENRICHMENT Table B-1. Nitrate mineral salts (NMS) media recipe Chemical Amount (per liter) MgSO 4 H 2 O 1.0 g KNO 3 1.0 g CaCl 2 0.2 g 3.8 % (w/v) FeEDTA 0.1 mL 0.1 % (w/v) NaMolybdate•4H 2 O 0.5 mL Whittenbury Trace Elements 1.0 mL Phosphate Stock Solution 10.0 mL Vitamin Stock Solution 10.0 mL 10 M CuNO 3 (optional) 40.2 L Table B-2. Whittenbury trace elements Chemical Amount (per liter) FeSO 4 H 2 O 500 mg ZnSO 4 H 2 O 400 mg MnCl 2 H 2 O 20 mg CoCl 2 H 2 O 50 mg NiCl 2 H 2 O 10 mg H 3 BO 3 15 mg EDTA 250 mg Table B-3. Phosphate stock solution Chemical Amount (per liter) KH 2 PO 4 26 g Na 2 HPO 4 33 g Table B-4. Vitamin stock solution Chemical Amount (Per Liter) Biotin 2.0 mg Folic acid 2.0 mg Thiamin•HCl 5.0 mg Ca pantothenate 5.0 mg B 12 0.1 mg Riboflavin 5.0 mg Nicotinamide 5.0 mg 87

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APPENDIX C METHANOTROPHIC ISOLATION PROCESS Methanotrophs have been successfully isolated from a variety of environmental samples using the techniques described by Whittenbury (1970) and Hanson (1998). This methodology was initially used in attempts to isolate methanotrophic bacteria from the enriched mixed cultures derived from the landfill samples. This procedure requires preparation of 10 -1 to 10 -6 dilution series where 1 ml of enrichment culture was added to the first sterilized tube containing 9 ml of NMS medium (without copper for GW60 and with copper for GW70). The tube(s) were vortexed to disperse cells evenly throughout the mixture and 1ml of that mixture was transferred to the next tube (10 -2 ) and the procedure was repeated for each tube up to 10 -6 . Then, 0.1 ml of the vortexed, serially diluted samples were spread plated in triplicate (in addition to duplicate for controls) onto freshly prepared NMS agar plates. Plates were then stored in desiccators at the previously mentioned methane-to-air ratios for GW60 and GW70 at 30C, whereas controls were incubated under an atmosphere containing only air. Visible inspection of plates occurred daily using a plate microscope to detect colony formation with methane as a sole carbon source. Controls were also inspected for growth of chemoautotrophs using CO 2 as a carbon source and numbers of bacteria capable of scavenging carbon from any impurities that might be present in the medium (Mancinelli and McKay, 1985). To ensure transfer of methane oxidizers, small, pinpoint colonies, were selected and transferred to fresh agar plates and re-incubated. This procedure was repeated with frequent checks for purity by streaking isolated colonies onto nutrient agar. Also, 88

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89 throughout this process desiccators were frequently swabbed with ethanol to minimize fungal contamination as previously suggested by Whittenbury et al. (1970). After several weeks of this process, no purified, isolated methane-oxidizing bacteria could be isolated and other techniques for isolation were investigated. Another isolation technique that was developed by Button et al. (1993) based on the dilution culture technique was also used in attempts at methanotrophic isolation. This process was also reported by Wise et al. (1999) who successfully isolated numerically dominant methanotrophs from landfill soil samples using this extinction-to-dilution technique. The theory behind extinction-to-dilution is that in cultures diluted to extinction allow the most abundant organisms to become favored during enrichment over the opportunistic species that tend to outcompete the numerically dominant species under the typical set of enrichment conditions. This technique was primarily used for direct isolation from soil samples, so it was adapted for use with already enriched, mixed cultures and is very similar to the previously described serial dilution and spread plate method. Samples were serially diluted to 10 -10 and incubated under methane-to-air ratios as previously described. The dilution series showing the first signs of growth were repeatedly picked and streaked onto the same medium and incubated under the same conditions in an effort to achieve purity. Purity checks were done by observing growth on plates both with and without methane and streaking onto nutrient agar. Despite repeated streaking, no purified isolates were established using this technique either. Since Type I methanotrophs typically favor low methane, high oxygen conditions, whereas Type II methanotrophs favor the opposite (Graham et al., 2002), methane concentrations were varied for both mixed enriched liquid cultures in further attempts to

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90 isolate pure methanotrophs. The maintenance methane concentrations, previously described, in addition to a higher methane concentration (50%) were used in both liquid cultures and solid media (NMS agar) as shown in Figure C-1. It was also thought that since these samples were originally derived from a nutrient rich environment containing various metals and ions, that tap water could be used in replace of the sterilized double de-ionized water in one set of culturing conditions that may promote favorable conditions for methanotrophic growth. Each culture was subjected to growth in both NMS prepared with tap water and also with the traditional sterilized DDI water at both low and higher methane concentrations. Transfers of liquid cultures were performed as soon as turbidity was evident. Continual streaking onto nutrient agar plates was used to check for purity in liquid cultures. Additionally, NMS agar plates were used for isolation at both low and higher methane concentrations after the mixed cultures were serially diluted and spread plated as previously described by Madsen (1998) with repeated transfers and purity checks performed. Despite these extensive measures utilizing both liquid and solid cultures, no pure, isolated methanotrophs were confirmed. Another previously published technique used a straightforward method for screening methanotrophic colonies for soluble methane monooxygenase activity on solid media (Graham et al., 1992). This was applied to samples derived from sanitary landfill soils and was successful in the isolation of sMMO-bearing methanotrophs. Since GW60 was cultured without copper (supporting the sMMO enzyme expression), this sample was used to test this new method on agar plates. sMMO activity results in the development of a colored complex between 1-napthol, which is formed when sMMO reacts with naphthalene and tetrazoitized o-dianisidine. If sMMO activity is present, colonies will

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91 develop a deep purple color when exposed successively to naphthalene and o-dianisidine. A previously incubated plate under methane that had been serially diluted and spread plated from GW60 liquid culture was used to test this new method. A positive control was also used with Methylomonas trichosporium OB3b, which is a known Type II methanotroph that expresses the sMMO enzyme without copper present. This culture was also serially diluted, spread plated and allowed to grow under methane-to-air tensions of 30:70 (at 30C) for a couple of days to allow full colony development. Following the procedures described by Graham et al. (1992), both cultures grown on NMS agar plates were removed from their desiccators and placed into a sterilized bench top in a laminar flow hood. A few laboratory grade naphthalene crystals were sprinkled into the lids of each plate and stored inverted at room temperature for 15 minutes under atmospheric conditions in the flow hood. The plates were then opened and lightly sprayed with freshly prepared, 5 mg/ml o-dianisidine (tetrazotized; zinc chloride complex; Sigma-Aldrich, St. Louis, MO) for 2-3 seconds. The lid was replaced and the plate stored for 15 minutes in the presence of the dye. A deep purple color developed for the positive control (OB3b) and also for GW60. Subsequent transfers were made of the sMMO positive colonies onto fresh NMS agar plates and incubated under methane. Once growth developed, transfers were made and purity was continually checked on nutrient agar. Despite several weeks of transfers, no pure methanotroph was isolated from the sMMO positive test plates. Since difficulty was experienced in isolating methanotrophs from the enriched mixed cultures, samples were sent to another lab to isolate. A laboratory in Croatia (Dr. Hrsak, Center for Marine and Environmental Research, Zagreb, Croatia) agreed to take

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92 these samples and try to isolate a methanotroph from the mixed community. Despite isolation attempts using dilution and spread plate techniques, no pure methanotroph(s) could be separated from the heterotrophs present in the mixed culture. The conclusion drawn after such extensive measures were taken in attempts to isolate a pure methanotroph was that it was not possible with the techniques used in this study and by another laboratory. The difficulty in separating methanotrophic species from other community members begins to raise questions and a need for an explanation of why and how this is happening. In another study where a mixed, enriched, stable culture showed more successful transformation of LAS compounds than the individual populations alone, it was suggested that this particular community was structured on specific relationships between the methanotroph and heterotrophs (Hrsak and Begonja, 1998). This implied that similar highly stable associations between microbial populations would be commonly found in nature. Because of the difficulty in separating out methanotrophic bacteria from its heterotrophic community members, the results from this study would further support this rationale.

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93 Enriched Mixed Culture (GW60 or GW70) Solid Media 20 or 30% CH4 NMS agar Transfer as soon as colonies develop. Purity checks on nutrient agar each transfer. 50% CH4 NMS agar Liquid Media 20 or 30% CH4 NMS (tap) NMS (DDI) 50% CH4 NMS (tap) NMS (DDI) Serial Dilution & Spread Plate 10% Inocula Transfers Transfer as soon as turbidity is detected. Purity checks on nutrient agar each transfer. Figure C-1. Technique using both solid and liquid media for methanotroph isolation.

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APPENDIX D SUPPLEMENTARY HETEROTROPH INFORMATION Previous work with the isolated heterotrophs included scanning electron microscopy to determine cellular shapes and sizes as reported in Table D-1 (Cheong, 2000). Heterotrophs were cultured using liquid medium nutrient broth (Becton Dickinson and Company, Sparks, MD, USA) and sent to the Interdisciplinary Center for Biotechnology Research (ICBR) Electron Microscopy Core Lab (EMCL) at the University of Florida. Samples were prepared for scanning electron microscopy by fixing with Trumps fixative (buffered 1% glutaraldehyde, 4% formalin), postfixing with 4% OsO 4 , rinsing with a graded ethanol series and mounting dried samples by sputter coating with a gold/palladium mixture. Prepared samples were then viewed on a Hitachi S-4000 electron microscope (Hitachi Co., Tokyo, Japan). Other previously isolated heterotrophs from a mixed culture (similar to the isolated heterotrophs reported here) belonged to Xanothobacter autotrophicus, Pseudomonas and genus Bacillus (Uchiyama et al., 1992). These heterotrophs grew on nutrient agar, did not grow with methane, were white and yellow, Gram negative, oxidase positive and short rod-shaped (1.1 to 2.4 m). Another study that identified isolated heterotrophs from a soil derived mixed culture using partial 16S rRNA included, Pseudomona pavonecea, Variovorax paradoxus, Bradyrhizobium elkanii and Hyphomicrobium vulgare (Dunfield et al., 1999). Other isolated heterotrophs from mixed cultures that are similar to the heterotrophs reported in this study were diverse as they varied in color from yellowish, white, and translucent, grew on nutrient agar with abundant growth at 30C, 94

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95 and ranged in size from 1 to 5 mm (Hrsak and Begonja, 1998). Tentative identification of these heterotrophs revealed genera, Blastobacter, Pseudomonas, and Xanthobacter. Also 2 unidentified heterotrophic community members showed similarities to prosthecates, but this morphology had not been confirmed with electron microscopy. From these studies alone, it appears that Pseudomonas is a common genus typical of mixed cultures containing methane-oxidizers and may potentially be present in the samples from this study as well.

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96 Table D-1. Phenotype characteristics of heterotroph populations. Culture No. Cellular morphology a Tentative identification of heterotroph 1 Coccobacillus L=0.64 m D=0.42 m Bacillus L=2.42 m D=0.36 m Major category I, group 4, subgroup 4a: Phyllobacterium Afipia Acidovorax 2 Not applicable Not enough information 3 Not applicable Major category I, group 4, subgroup 4a: Pseudomonas Erythrobacter Acidiphilium GW 60 4 Coccobacillus L=1.09 m D=0.50 m Bacillus L=0.87 m D=0.26 m Major category I, group 4, subgroup 4a: Afipia Acidovorax 1 Curved Rod L=0.95 m D=0.25 m Bacillus L=0.94 m D=0.28 m Major category I, group 4, subgroup 4a: Afipia Acidovorax 2 Not applicable Major category I, group 4, subgroup 4a: Pseudomonas Erythrobacter Acidiphilium 3 Not applicable Major category I, group 4, subgroup 4a: Acidovorax Phyllobacterium Pseudomonas Zoogloea 4 Bacillus L=1.16 m D=0.23 m Prosthecates L=1.18 m D=0.16 m Major category I, group 13 : Caulobacter Hyphomicrobium GW 70 5 Not applicable Major category I, group 4, subgroup 4a: Afipia Acidovorax a Length (L) and diameter (D) dimensions are averaged values measured the photographs (Cheong, 2000; Table 3 and Table 4, pg. 34)

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APPENDIX E EXAMPLES OF LABORATORY EQUIPMENT USED ABC Figure E-1. Laboratory equipment A) Desiccator. B) Nutrient plate storage. C) Example of sMMO assay results with acetylene control. 97

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APPENDIX F EXAMPLES OF CALCULATIONS: GROWTH, METHANE DEPLETION AND OXYGEN UPTAKE 00.10.20.30.40.50.60.70.80.9101020304050607080Time (hr)Absorbance @ 600 n m Control Flask #1 Flask #2 Flask #3 Figure F-1. Growth curve used in calculating growth rates. y = 0.0301x 1.4899R2 = 0.9919y = 0.028x 1.4676R2 = 0.9868y = 0.0282x 1.4727R2 = 0.9909-1.6-1.4-1.2-1.0-0.8-0.6-0.4-0.20.0010203040506Time (hr)Log10 Average Absorbance 0 Flask #1 flask #2 flask #3 Linear (flask #2) Linear (flask #3) Linear (Flask #1) Figure F-2. Linear regression fit of growth curve data in calculating growth rates. 98

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99 Results summary Growth rate u = 0.0288 h^-1 u = 0.6912 d ^-1 Doubling time calcs (use linear): Slope of log phase= 0.0288 h^-1 3/4 log phase = 37.5125 time to harvest Generation time, g= 0.301/slope eqn from Brock,pg.142-45 g (doubling time) = 10.45 h Growth rate const., k= 0.693/g eqn from Brock,pg.142-45 k = 0.066 h^-1 Slope flask #1 0.0282 Slope flask #2 0.0301 Slope flask #3 0.028 std. dev. = 0.0011 h^-1 Slope flask #1 0.6768 Slope flask #2 0.7224 Slope flask #3 0.672 std. dev. = 0.0278 d^-1 Figure F-3. Sample of spreadsheet used for growth rate calculations based on linear regression curves. 0.02.04.06.08.010.012.014.016.018.020.001123293544485969768194104Time (hrs)Percentage of Methane Control Flask #1 Flask #2 Flask #3 Figure F-4. Measured methane depletion for mixed culture GW70.

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100 y = -0.0011x + 1.4789R2 = 0.5627y = -0.0149x + 1.5974R2 = 0.9525y = -0.0148x + 1.6464R2 = 0.9333y = -0.0146x + 1.1776R2 = 0.95270.00.20.40.60.81.01.21.41.61.8020406080100120time (hrs)mass of methane in headspace (mg ) Control #1 #2 #3 Linear(Control) Linear (#1) Linear (#2) Linear (#3) Figure F-5. Linear regression of data based on methane depletion curves.

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101 Methane Oxidation Rates* (using linear regression) *no methane oxidation rate pertains to the control flask b/c no cells present **however, the loss of methane from the control must be accounted for Control Slope of delta M (mg) tn-1 tn (h) Corrected M (mg) -0.0011 103.5 -0.11385 Flask #1 Slope of Delta M (mg) tn-1 tn (h) Corrected M (mg) K ch4 (h^-1) K ch4 (d^-1) -0.0149 59 -0.76525 0.01839 0.441 Flask #2 Slope of Delta M (mg) tn-1 tn (h) Corrected M (mg) K ch4 (h^-1) K ch4 (d^-1) -0.0148 59 -0.75935 0.01825 0.438 Flask #3 Slope of Delta M (mg) tn-1 tn (h) Corrected M (mg) K ch4 (h^-1) K ch4 (d^-1) -0.0146 59 -0.74755 0.01797 0.431 Average Kch4 (1,2,3) = 0.01821 hr^-1 std. dev. = 0.0002 Average Kch4 (1,2,3) = 0.437 d^-1 std. dev. = 0.0052 Calculating Mass of Methane in Headspace PV = n RT Calc. for flask P (pressure)= 1.0 atm V (vol. of liquid) = 0.0132 L n (moles of air) = UNKNOWN moles R (ideal gas constant)= 0.083145 L-atm/mol-K T (temperature) = 303 kelvin n (air) = 0.0005 moles of air in flask A % of air was removed from the flasks to be replaced by methane. Take the % of moles of air removed (% methane) and calculate the mass of methane based on its molecular weight M methane (mg)= 16 g/mol *(%methane) * n*1000 Calculating Methane Oxidation Rates (Kch4) See Hrsak and Begonja methods (1998) for eqn source Kch4 = (deltaM/(x*V)* (1/(tn-1 tn) delta M = change in mass of methane in flask (mg) x = avg. biomass concentration (mg/L) V = volume of liquid (L) tn-1 tn = cultivation period (assume log phase) delta M = (change in M for each flask during log phase) x = 53.0 mg/L V = 0.0132 L tn-1 -tn = time frame for methane depletion for each flask h Figure F-6. Spreadsheet used for specific methane oxidation rate calculations.

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102 GW 60,13 1/28/2003 25 ppm Benzene Oxidation Rate Calculations Conversion Factor Eqn (micromol/div) = (Vreactor*C)/(% DO Divisions *2) Where Vreactor = 1.9 mL reactor volume C = 7.6 mg/L of O2 @ 30 deg. C (taken from DO solubility table WW engineering) C = 7.6 microgram atoms O/mL -concentration of O2 in water @ 30 deg. C % DO Div = 100 # of DO divisions (from the calibration curve) Conv.Fact= 0.07213 umol/div Rate eqn (micromol/s) = Mcorrected * Conversion Factor where Mcorrected = Msubstrate Mendogenous or Mmethane (for calculating methane rate) Mmethane = -0.0319 % DO/s (see methane sheet) R^2 meth = 0.989 linear Run #1 Endogenous phase Substrate phase m1 = -0.0137 % DO/sec m2 = -0.0193 % DO/sec R^2 end = 0.8428 linear R^2 sub = 0.9606 linear mcorrected = -0.0056 % DO/sec Rate w/ CH4 = -0.0023 umol/s Rate w/ B= -0.0004 umol/s normalized = 0.175549 Run #2 Endogenous phase Substrate phase m1 = -0.0123 % DO/sec m2 = -0.0154 % DO/sec R^2 endog = 0.7806 linear R^2 substr = 0.9154 linear mcorrected = -0.0031 % DO/sec Rate w/CH4= -0.0023 umol/s Rate w/ B= -0.00022 umol/s normalized = 0.097179 Run #3 Endogenous phase Substrate phase m1 = -0.0139 % DO/sec m2 = -0.0278 % DO/sec R^2 end 0.9119 linear R^2 sub 0.9873 linear mcorrected = -0.0139 % DO/sec Rate w/CH4= -0.0023 umol/s Rate w/ B= -0.001 umol/s normalized = 0.435737 Combined runs Avg. rate = -0.00054 umol/s Rate w/CH4= -0.0023 umol/s Std.dev. = 0.17722 avg. norm. = 0.23615 umol[BEN]/umol[CH4] Figure F-7. Spreadsheet used in oxygen uptake calculations.

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103 y = -0.0107x + 90.618R2 = 0.9226y = -0.0031x + 93.086R2 = 0.2579788082848688909294960200400600800100012001400time (s)DO cells CH4 acetylene Linear (CH4) Linear (cells) Figure F-8. Measured oxygen consumption in the presence of methane, showing inhibition of acetylene during oxygen uptake analysis. 0204060801001201401601802000500100015002000time (s)% DO 1,4 Dioxane + NMS 1,4 Dioxane + Cells Figure F-9. Measurement of oxygen consumption in the presence of 1,4 dioxane and NMS medium (a control) and then in the presence of cells.

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104 y = -0.0051x + 92.46R2 = 0.293350556065707580859095300350400450500550600time (s)DO% BEN+NMS Linear (BEN+NMS) Figure F-10. Measurement of oxygen consumption in the presence of benzene plus NMS (a control). y = -0.005x + 120.03R2 = 0.6452254565851051251450100200300400500600time (s)%DO TCE+NMS Linear (TCE+NMS) Figure F-11. Measurement of oxygen consumption in the presence of TCE plus NMS (a control).

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APPENDIX G SUPPLEMENTARY INFORMATION FOR GROWTH AND METHANE DEPLETION EXPERIMENTS Table G-1. Growth and methane depletion by landfill samples at varied methane concentrations Methane % Culture Growth rate constant (h -1 ) Protein content for growth (mg/L) Protein content for methane (mg/L) GW 60-13 (NMS) 0.061 170 147 GW 60-13 (NMS w. copper) 0.118 135 71 10 % GW 70-20 (NMS w. copper) 0.065 29 NR GW 70-20 (NMS) 0.087 549 368 20 % GW 70-20 (NMS w. copper) 0.066 223 53 GW 60-13 (NMS) 0.079 514 279 30 % GW 60-13 (NMS w. copper) 0.076 127 265 GW 60-13 (NMS) 0.062 557 258 GW 60-13 (NMS w. copper) 0.095 347 107 70 % GW 70-20 (NMS w. copper) 0.052 476 38 All samples cultured at 30C, pH of media = 6.8, 270 rpm NR, no results for this culture: no methane loss was detected in headspace of vials, minimal growth was detected and there was no protein content and thus no methane oxidation rate was calculated nor sMMO assay performed. 105

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106 Table G-2. Growth and methane depletion by landfill samples at varied temperature and pH. Condition varied Culture Growth rate constant (h -1 ) Protein content for growth (mg/L) Protein content for methane (mg/L) GW 60-13 (NMS) a NG NG NG GW 60-13 (NMS w. copper) a 0.071 397 483 12 -15 C GW 70-20 (NMS w. copper) b 0.009 92 457 GW 60-13 (NMS) a NG NG NG GW 60-13 (NMS w. copper) a NG NG NG 41 C GW 70-20 (NMS w. copper) b 0.045 NR 69 GW 60-13 (NMS) c 0.019 139 60 GW 60-13 (NMS w. copper) c 0.021 130 231 pH = 4.8 GW 70-20 (NMS w. copper) d 0.011 118 360 GW 60-13 (NMS) c 0.020 145 112 GW 60-13 (NMS w. copper) c 0.013 167 172 pH = 8.8 GW 70-20 (NMS w. copper) d 0.023 191 206 a Cultured at 30:70 methane-to-air ratio, pH of Media = 6.8, 270 rpm b Cultured at 20:80 methane-to-air ratio, pH of Media = 6.8, 270 rpm c Cultured at 30:70 methane-to-air ratio, 30C, 270 rpm d Cultured at 20:80 methane-to-air ratio, 30C, 270 rpm NG, no growth was detected in samples at these conditions. NR, no results due to insufficient sample size to perform analysis.

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BIOGRAPHICAL SKETCH I began my undergraduate degree at the University of Florida in August of 1996. I initially selected my major in Natural Resource Conservation in the College of Agriculture and Life Sciences through the School of Forestry. This sounded interesting to me because I love the environment and wanted to major in something that would enable me to have a positive impact on the environment and people’s lives. However, as I progressed through that major, I didn’t feel that perfect fit, and began investigating other majors that were available. It was the suggestion of a friend that I look into Environmental Engineering. When I read the major description in the course catalog, I immediately knew that is what I wanted to do. It sounded challenging as it combined science, physics, chemistry, and mathematics, to design systems that helped improve the quality of life while sustaining the environment. I immediately switched my major and worked hard in my undergraduate studies. The coursework was rigorous and the demands were high, but I enjoyed what I learned and persevered. I became involved in various leadership positions as I held an officer position (treasurer) in two different student organizations (Air and Waste Management Association; Florida Water Environment Association) associated directly with my major. I was also involved with the Society of Women Engineers and served as a mentor for incoming freshman. During an organic chemistry course in my junior year, my professor (Dr. Angela Lindner) told our class about a new program initiated to encourage undergraduate students to become involved with research and to learn about the scholarly method. My 117

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118 interest in science and research immediately prompted me to approach my professor and ask her more about the program. She gave me more details and explained a project she had in mind for undergraduates to work on. I applied for the program and became a University Scholars participant. I was very excited to learn more about research and how I could potentially make an impact in this area. Dr. Lindner was an excellent advisor and mentor, who encouraged me to excel and pushed me to be the best that I could. She was very involved with my undergraduate project, and mentored me with a sincere motivation to lead and guide students in academia. I learned much through this program. I was introduced to microbiology and the field of bioremediation, which blends microbiology, science and engineering to clean up the environment through the natural occurring biota. This field fascinated me and I ended up presenting my University Scholars Program research at a symposium and also publishing it in the Journal of Undergraduate Research. Because of this positive experience, I continued to work in the laboratory and perform research as I worked toward my bachelor’s degree. I ended up with the opportunity to travel to places such as Salt Lake City, Utah; and Washington, DC to present my research at conferences (Air and Waste Management Association; American Society for Microbiology) and also at the University of Florida through several poster symposiums. I graduated with my Bachelor of Science in December of 2001 and continued my education by pursuing my master’s degree with a thesis focus in the same department as my undergraduate, with my same advisor (Department of Environmental Engineering Sciences, Dr. Angela Lindner). The laboratory training from my undergraduate work enabled me to transition more smoothly into graduate school. I applied for funding and received both a research

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119 assistantship and also the Selected Professions Fellowship awarded through the American Association for University Women. This organization was wonderful in supporting women pursing degrees in typically male-dominated fields. I met a wonderful group of women in the Gainesville Branch of the association. I also had the opportunity to be a guest speaker in Vero Beach, as part of their monthly chapter meeting. These are just some of the highlights of my academic career here at the University of Florida. This is a great university, providing exciting athletics, plenty of social organizations, nonprofit organizations, intramurals, and numerous majors to choose from. I have enjoyed many opportunities as a student and will always remember this as an overall positive experience. There have also been many challenges, difficulties, and barriers that have taught me so much during this time. I know they will equip me for future challenges that I will face, as I enter the work force in the consulting industry as an environmental engineer.