Open Thesis 17May2011.Pdf

The Pennsylvania State University

The Graduate School

Department of Geosciences

BIOGEOCHEMISTRY OF ISOPRENOID PRODUCTION AND ANAEROBIC

HYDROCARBON BIODEGRADATION

A Dissertation in Geosciences

Katherine S. Dawson

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

August 2011

The dissertation of Katherine S. Dawson was reviewed and approved* by the following:

Jennifer L. Macalady Assistant Professor of Geosciences Dissertation Co-Advisor Co-Chair of Committee

Katherine H. Freeman Professor of Geosciences Dissertation Co-Advisor Co-Chair of Committee

Christopher H. House Associate Professor of Geosciences

John M. Regan Associate Professor of Environmental Engineering

Chris J. Marone Professor of Geosciences Associate Head for Graduate Programs and Research in Geosciences

*Signatures are on file in the Graduate School

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ABSTRACT

This dissertation is an exploration of microbial isoprenoid production and destruction by anaerobic hydrocarbon biodegradation. Isoprenoids are methyl-branched hydrocarbons, and include biomarkers from all three domains of life such as archaeal lipids, hopanoids, and sterols. Isoprenoid production was examined through variation in the molecular structure of archaeal lipids across a hypersaline gradient (Chapter 5). This study identified unsaturated analogues of archaeol in four halophilic archaeal strains and revealed an increase in the percentage of unsaturated lipids with increasing salinity.

Anaerobic isoprenoid biodegradation was examined through the enrichment of bacteria under anaerobic conditions utilizing pristane as a carbon source (Chapter 2). Further analysis of anaerobic degradation utilized 13C-labelled phytane as a stable isotope tracer

(Chapter 3). In both cases, a microbial community dominated by denitrifying Beta- and

Gammaproteobacteria was responsible for the degradation of pristane and phytane.

Environmental anaerobic hydrocarbon degradation was examined through the analysis of in situ microbial communities associated with the transformation of coal to methane

(Chapter 4). Through FISH and 16S rRNA tag pyrosequencing a coal transformation pathway ending in acetoclastic/methylotrophic methanogenesis was identified in the

Cook Inlet Basin, Alaska. These studies demonstrate the microbial impact on hydrocarbon production and alteration, which influences the transition over geologic time scales from biomolecules to biomarkers in the sedimentary record.

TABLE OF CONTENTS

LIST OF FIGURES vii

LIST OF TABLES x

ACKNOWLEDGEMENTS xi

CHAPTER 1: INTRODUCTION

1.1 Isoprenoids 1

1.2 Microbial Hydrocarbon Degradation 2

1.3 Archaeal Membrane Lipids 5

1.4 Biogenic Coal Bed Methane 6

1.5 Organization Of The Thesis 8

1.6 Anticipated Publications From This Work 8

1.7 Figures 10

1.8 References 19

CHAPTER 2: ANAEROBIC BIODEGRADATION OF ISOPRENOID

BIOMARKERS BY A DENITRIFYING MICROCOSM

2.1 Abstract 26

2.2 Introduction 26

2.3 Materials and Methods 28

2.4 Results 32

2.5 Discussion 36 v

2.6 Acknowledgements 39

2.7 Figures and Tables 40

2.8 References 47

CHAPTER 3: STABLE ISOTOPE TRACING OF ANAEROBIC ISOPRENOID

BIODEGRADATION WITH 13C LABELLED PHYTANE

3.1 Abstract 52

3.2 Introduction 52

3.3 Experimental Procedures 54

3.4 Results 59

3.5 Discussion 62

3.6 Conclusions 65

3.7 Acknowledgements 66

3.8 Figures and Tables 67

3.9 References 78

CHAPTER 4: QUANTITATIVE FISH ANALYSIS OF MICROBIAL

CONSORTIA FROM A BIOGENIC GAS FIELD IN THE COOK INLET

BASIN, ALASKA

4.1 Abstract 83

4.2 Introduction 83

4.3 Materials and Methods 85

4.4 Results 88 vi

4.5 Discussion 92

4.6 Conclusions 94

4.7 Figures and Table 96

4.8 References 103

CHAPTER 5: MOLECULAR CHARACTERIZATION OF ARCHAEAL LIPIDS

ACROSS A HYPERSALINE GRADIENT

5.1 Abstract 107

5.2 Introduction 107

5.3 Methods 109

5.4 Results 111

5.5 Discussion 113

5.6 Conclusions 115

5.7 Acknowledgements 116

5.8 Figures and Tables 117

5.9 References 123

CHAPTER 6: CONCULSIONS AND FUTURE DIRECTIONS

6.1 Anaerobic isoprenoid degradation 128

6.2 Coal bed methane biogeochemistry 129

6.3 Halophilic archaeal lipids 130

6.4 References 131 vii

LIST OF FIGURES

Figure 1-1: Examples of isoprenoid biopolymers that may serve as biomarkers 10

Figure 1-2: Proposed mechanisms for anaerobic hydrocarbon degradation 11

Figure 1-3: Membrane glycerolipid structures in the three domains of life 12

Figure 1-4: Structures of archaeal glycerol diphytanyl glycerol tetraethers (GDGT’s) and

diphytanyl glycerol diethers (DGD’s) 13

Figure 1-5: Isotopic values of methane showing the differences between thermogenic and

biogenic methane generated from different pathways 14

Figure 1-6: Phylogenetic tree of 16s rDNA sequences of Firmicutes, Bacteroidetes,

Planctomycetes, Spirochaetes, and other bacterial lineages associated with coal bed

methane 15

Figure 1-7: Phylogenetic tree of 16s rDNA sequences of Betaproteobacteria and

Gammaproteobacteria associated with CBM 16

Figure 1-8: Phylogenetic tree of 16s rDNA sequences of Alphaproteobacteria,

Deltaproteobacteria and Epsilonproteobacteria associated with CBM 17

Figure 1-9: Phylogenetic tree of 16s rDNA sequences of methanogenic archaea

associated with CBM 18

Figure 2-1: FISH micrographs of cells in the denitrifying, pristane-degrading enrichment

culture 40

Figure 2-2: Neighbor-joining phylogenetic tree showing betaproteobacteria from pristane

degrading enrichments 41

Figure 2-3: Neighbor-joining phylogenetic tree showing gammaproteobacteria from

pristane degrading enrichments 42 viii

Figure 2-4: Fraction of bacterial cells hybridizing to specific oligonucleotide probes in

pristane degrading cultures 43

Figure 2-5: Anaerobic consumption of nitrate, and the production of nitrite and

bicarbonate in a denitrifying, pristane degrading enrichment culture 44

Figure 2-6: Anaerobic consumption of nitrate in denitrifying enrichments with pristane,

DGD and GDGT core lipids 45

Figure 3-1: Preparation scheme for 13C-labeled phytane 67

Figure 3-2: Gas chromatogram and mass spectra of saponified 13C-labeled diphytanyl

glycerol diether from Haloferax sulfurifontis SD1 68

Figure 3-3: Gas chromatograms and mass spectra of 13C-phytane before and after SPE

purification 69

Figure 3-4: Loss of nitrate and production of nitrite and bicarbonate in incubation grown

with 13C-labeled phytane as a carbon source 70

13 13 Figure 3-5: Values of δ C for the CO2 produced during incubations on C-labeled

phytane, unlabeled phytane and with no added carbon 71

13 13 Figure 3-6: Keeling plot of C for CO2 produced during incubation on C-labeled

- phytane versus 1/[HCO3 ] 72

Figure 3-7: Percentage of cells hybridizing to specific oligonucleotide probes in phytane

degrading incubations 73

Figure 3-8: FISH micrographs of cells in the denitrifying, phytane-degrading enrichment

culture 74

Figure 3-9: Neighbor-joining phylogenetic tree showing gammaproteobacteria clones in

13C-phytane and pristane degrading enrichments 75 ix

Figure 3-9: Neighbor-joining phylogenetic tree showing betaproteobacteria clones in 13C-

labeled phytane and pristane degrading enrichments 76

Figure 4-1: FISH micrographs depicting major bacterial and archaeal lineages observed

in production water samples from the Cook Inlet Basin 97

Figure 4-2: Microbial communities of the Cook Inlet gas field expressed as a percentage

of DAPI stained cells 98

Figure 4-3: Comparison of the microbial communities of the Cook Inlet gas field

determined by FISH and 16S rRNA tag pyrosequencing 99

Figure 4-4: Carbon and hydrogen isotopic classification of methane of the Cook Inlet

sample sites 100

Figure 4-5: PCA of Cook Inlet basin microbial population data with geochemical data

projected onto the ordination using the ‘envfit’ function 101

Figure 4-6: PCA of microbial population data from five sedimentary basins with

geochemical data projected onto the ordination using the ‘envfit’ function 102

Figure 5-1: Structures of archaeal diphytanyl glycerol diether (DGD) with various chain

lengths, cyclization, unsaturation and hydroxyl substitutions 118

Figure 5-2: Total ion chromatograms of saturated C20-20 and C25-20 DGD 119

Figure 5-3: Partial GC-MS chromatograms of polar lipid extracts of halophilic archaea

showing C20-20 and C25-20 DGDs 120

Figure 5-4: Plot of total unsaturated archaeal C20-20 and C25-20 DGD lipids for four

halophilic archaeal strains 121

Figure 5-5: Average fraction of unsaturated DGDs versus optimal % NaCl (w/v) for four

halophilic archaeal strains 122 x

LIST OF TABLES

Table 2-1: Ratios of isoprenoid peak areas in GC-MS chromatograms 46

Table 2-2: Oligonucleotide probes used in the study of denitrifying, pristane degrading

enrichments 46

Table 3-1: Oligonucleotide probes used in the study of denitrifying, phytane degrading

incubations 77

Table 4-1: Oligonucleotide probes used in the study of coal bed methane production

water 96

Table 5-1: Presence of absence of C20-20 and C25-20 DGDs in halophilic archaea from

various genera 117

ACKNOWLEDGEMENTS

First I would like to thank my advisors Jenn Macalady and Kate Freeman. As mentors your support has made this possible. Thank you both for turning a chemist into a geomicrobiologist and organic geochemist.

Chris House, you have been generous with your time and guidance, which has helped me to develop the ideas behind these projects. Thanks also to Jay Regan whose advice has helped to improve this thesis.

I’ve learned analytical techniques from many people. Many members of the

Macalady, Freeman and House labs have helped scientific discussions and experimental trouble-shooting. Irene Schaperdoth taught me most of the molecular biology skills that I now have. Everything I know about anaerobic culturing started from what Zhidan Zhang showed me. Denny Walizer, technical support doesn’t begin to capture what you do.

Thank you for helping me run or fix every instrument I’ve used.

Funding from the NSF-Penn State Biogeochemical Research Initiative for

Education, NASA-Penn State Astrobiology Research Center, American Chemical Society

Petroleum Research Fund, NASA Astrobiology Institute Directors Discretionary Fund, and ConocoPhillips are all gratefully acknowledged.

To all of my friends, you have kept me laughing and made this experience fun when at times it otherwise would have been a chore.

Vanessa, I look up to you (figuratively as well as literally) and appreciate all of your insights into teaching, life and pop culture.

Most importantly I need to thank my parents, Vicki and Colin. From science experiments in the basement and other corners of the house, to setting me loose to explore the Wissahickon and countless hours of questions, you both deserve so much of the credit for who I am today.

Chapter 1: Introduction

1.1 Isoprenoids

Hydrocarbons are aliphatic, cyclic, aromatic or polycyclic aromatic compounds composed primarily of carbon and hydrogen. Due to the lack of functional groups and the apolarity of carbon-hydrogen bonds, hydrocarbons have low biological and chemical reactivity, which serves to enhance their preservation in sedimentary environments (WIDDEL and RABUS,

2001). Susceptibility of hydrocarbons above C20 to biologic breakdown is additionally decreased due to their hydrophobicity and large molecular size. Isoprenoid hydrocarbons derive from lipids in organisms across the tree of life and are synthesized from branched, 5-carbon isoprene units

(Figure 1-1). Aliphatic isoprenoids such as pristane (C19) and phytane (C20) originate as the side chain of chlorophyll-a, chlorophyll-b, and bacteriochlorophyll-a (GILLAN and JOHNS, 1980) but can also derive from α- and γ-tocopherols (GOOSSENS et al., 1984) and halophilic archaeal membrane lipids (ANDERSON et al., 1977). Other aliphatic, alicyclic, and aromatic C20-C80 isoprenoids are derived from archaeal membrane lipids, bacterial hopanoids, eukaryotic and bacterial sterols, and photosynthetic pigments (ALBAIGES et al., 1985; CHAPPE, 1982;

MOLDOWAN and SEIFERT, 1979). Isoprenoids in these major groups include important biomarkers for many taxonomic groups (BROCKS et al., 2003; DUTKIEWICZ et al., 2006; PETERS et al., 2005), paleoenvironmental conditions such as temperature and salinity (WUCHTER et al.,

2006), and petroleum maturity (PETROV and ABRYUTINA, 1989).

While hydrocarbons in general are recalcitrant due to the need for activation (conversion to oxygenated intermediates) prior to biodegradation, variation exists in susceptibility due to chain-length, branching and cyclization (PETERS et al., 2005). For example, the branched

structure of isoprenoids imparts additional recalcitrance by hindering the β-oxidation pathway

(CANTWELL et al., 1978). The need for repeated demethylation or activation before β-oxidation can proceed reduces the potential metabolic energy yield from isoprenoid substrates, and may explain why their degradation rates in natural environments appear slower than equivalent non- isoprenoid hydrocarbons (HEAD et al., 2006; PETERS et al., 2005). In addition, branching at the chain ends (e.g. pristane) or near the chain ends (e.g. archaeol) may prevent some activation mechanisms due to steric hindrance. Nonetheless, anaerobic isoprenoid biodegradation has been demonstrated in a few laboratory microcosm studies utilizing pristane (BREGNARD et al., 1997;

DAWSON and MACALADY, 2007; GROSSI et al., 2000), which suggests that the hydrocarbon chains of other isoprenoid biomarkers are also susceptible to anaerobic degradation.

1.2 Microbial Hydrocarbon Degradation

Over geologic time scales, microbial degradation controls the global flux of hydrocarbons into long-term storage reservoirs in rocks. Microbial degradation can occur before sediment deposition (e.g. in sinking fecal pellets (TURNER, 1979)), during deposition and early diagenesis

(e.g. in marine sediments (MASSIAS et al., 2003)), or during migration and storage in oil reservoirs or coal deposits (HEAD et al., 2003). Biodegradation of hydrocarbons in the presence of oxygen occurs more rapidly than under anaerobic conditions (LARTER et al., 2006; LARTER et al., 2003; YAMANE et al., 1997). However, given the longer timescales of exposure to microbial agents in anoxic environments, anaerobic microbial activity represents a comparable source of biodegradation (LARTER et al., 2006; LARTER et al., 2003; PETERS et al., 2005).

Anthropogenic pollutants and gaseous hydrocarbons such as methane and propane have been the primary focus of studies on the fate of hydrocarbons in the absence of oxygen (HEAD et

al., 2003; HEIDER, 2007; LOVLEY, 2001; WIDDEL et al., 2006; WIDDEL and RABUS, 2001). In contrast, the biodegradation of isoprenoid hydrocarbons has received little attention to date despite relevance to petroleum characterization, coal bed methane generation, and the geologic record of life on earth. Although they are considered more recalcitrant than equivalent length unbranched alkanes, their anoxic degradation as evidenced by changing hydrocarbon profiles and gas evolution takes place at only slightly slower rates (HEAD et al., 2003). Direct evidence for the microbial role in anaerobic isoprenoid degradation is usually lacking, however, microorganisms are implicated by the simultaneous disappearance of isoprenoids and electron acceptors such as sulfate, iron and nitrate (BONIN et al., 2002; GROSSI et al., 2000; HEAD et al., 2003; MASSIAS et al., 2003).

In oxic environments, molecular oxygen plays multiple roles in aerobic hydrocarbon metabolism. It serves both as a terminal electron acceptor in microbial respiration, and as a required substrate for mono- and dioxygenase enzymes that activate hydrocarbons for subsequent degradation by the -oxidation pathway (LEAHY and COLWELL, 1990; SPORMANN and WIDDEL, 2000; VAN HAMME et al., 2003). Nonetheless, hydrocarbons also degrade under anoxic conditions. Analyses of anoxic oil reservoir materials suggesting microbial degradation are consistent with early reports of sulfate reducing bacteria isolated from oil fields (BASTIN et al., 1926; ROSENFELD, 1947). More recent work has shown that the microbial populations present in oil reservoirs reflect in situ geochemical conditions such as the relative abundances of terminal electron acceptors (ORPHAN et al., 2003). Anaerobic hydrocarbon degradation has been observed to be coupled with nitrate reduction, iron reduction, manganese reduction, sulfate reduction and methanogenesis (AECKERSBERG, 1991; BEAL et al., 2009; COATES et al., 2001;

EHRENREICH et al., 2000; FUKUI, 1999; GRBIC-GALIC and VOGEL, 1987; KNIEMEYER et al., 2007;

SPORMANN and WIDDEL, 2000; WIDDEL et al., 2006).

The past decade of research has revealed that anaerobic hydrocarbon degradation is mediated by microorganisms using novel biochemical pathways (Figure 1-2) (WIDDEL and

RABUS, 2001). Fumarate addition initiates the anaerobic biodegradation of toluene and may also be the activation step in the degradation of several other aromatic compounds (AITKEN et al.,

2004; HEIDER, 2007; SPORMANN and WIDDEL, 2000). For alkylbenzenes, dehydrogenation coupled with hydroxylation has been proposed as an activation step (SPORMANN and WIDDEL,

2000; WIDDEL and RABUS, 2001). For polyaromatic hydrocarbons, carboxylation of the aromatic rings is suggested by the incorporation of labeled CO2 into naphthalene and phenanthrene to form carboxylic acids (AITKEN et al., 2004; HEIDER and FUCHS, 1997; ZHANG and YOUNG,

1997), although methylation is an alternative hypothesis consistent with existing data that has also been suggested (SAFINOWSKI and MECKENSTOCK, 2006).

Mechanisms for anaerobic activation of both branched and linear alkanes are not as well studied, although several pathways have been suggested. In the 1960’s, alkane activation was proposed to occur via dehydrogenation and the formation of a terminal alkene followed by the hydroxylation of the double bond to generate an alcohol (shown for ethylbenzene in Figure 1-2)

(CHOUTEAU et al., 1962; SENEZ and AZOULAY, 1961; SPORMANN and WIDDEL, 2000). More recent studies have proposed the terminal or subterminal methylation or carboxylation of the alkane chain (shown in Figure 1-2 for naphthalene) (AECKERSBERG et al., 1998; CALLAGHAN et al., 2006). Alternatively, a mechanism involving fumarate addition with a higher energy barrier than for aromatic compounds (33 kJ/mol for secondary carbons, and 51 kJ/mol for primary

carbons) has been demonstrated for n-alkanes with chain lengths from 3 to 16 carbons

(CALLAGHAN et al., 2006; HEIDER, 2007; KNIEMEYER et al., 2007; WIDDEL et al., 2006).

1.3 Archaeal Membrane Lipids

Lipid membranes are a unifying feature of life across the three domains (Figure 1-3) and are a major source of compounds that are eventually preserved in sedimentary rocks and serve as valuable as biomarkers. Membrane lipids are primarily glycerides, amphiphilic compounds composed of nonpolar hydrocarbon chains bound to a polar glycerol head group. In bacteria and eukaryotes, the glyceride is typically an ester-linked fatty acid, while in archaea it is an ether- linked isoprenoid. Additionally, the archaeal glycerol ether is a 2,3-di-O-sn-glycerol in contrast to the sn-1,2 sterochemistry observed in bacteria and eukaryotes (DE ROSA, 1986). These differences in membrane lipid chemistry are one of the defining features of the bacterial-archaeal evolutionary divergence.

Archaea have an evolutionary history equally ancient as bacteria and are ubiquitous on earth, inhabiting environments ranging from human dental plaque to those with extreme salinity, temperature and pH values. Their ether-linked, isoprenoid membrane lipids include two principal structures: glycerol diphytanyl glycerol tetraethers (GDGT’s) that form monolayer lipid membranes and diphytanyl glycerol diethers (DGD’s) that form bilayer lipid membranes. The distribution of DGD versus GDGT membrane lipid composition within the archaeal domain varies between GDGT dominated membranes in acidophiles and exclusively DGD containing membranes in the Halobacteria (KATES, 1977; MACALADY et al., 2004). The general archaeal tetraether structure shown in Figure 1-4 can be tuned in vivo to ambient environmental conditions by the inclusion of cyclic groups, hydroxyl substitutions, or varying degrees of

unsaturation in the isoprenoid chains (WILKES et al., 2003). Archaeal diethers, the primary lipid of halophiles and a major component of methanotroph and methanogen membranes, also exhibit varying degrees of unsaturation (GIBSON et al., 2005; STIEHL et al., 2005), a macrocyclic form

(COMITA et al., 1984) and cyclic group inclusion (STADNITSKAIA et al., 2003). However, the environmental or phylogenetic causes for archaeal diether structural variation are not well understood. Therefore, further studies of these structural variations along with isotopic compositions have the potential to provide information about ancient microbial communities and paleoenvironments.

1.4 Biogenic Coal Bed Methane

Coal a global major energy resource, represented 27% of world energy consumption in

2007 (UNITED STATES ENERGY INFORMATION ADMINISTRATION, 2010). There is strong interest in ways to generate and extract natural gas from coal reserves for which mining is not economic, as well as to reduce the environmental impact of coal utilization. Coal mining is associated with

7% of annual methane production, as gas stored in the pore space is released from coal beds

(STRAPOC, 2011). Carbon and hydrogen isotope values of methane from various coal beds methane indicates both thermogenic (δ13C > -60‰) and biogenic (δ13C < -60‰) sources for this methane (MILKOV and DZOU, 2007; SCHOELL, 1983; STRAPOC et al., 2007; THIELEMANN et al.,

2004) (Figure 1-5). Thermogenic methane results from the cracking of hydrocarbons due to heat and pressure (SCHOELL, 1983), while biogenic methane results from archaeal CO2 reduction, acetate fermentation and conversion of other molecules containing a methyl group to methane

(THAUER, 1998). Coal bed methane (CBM) and other gases generated by thermogenic and

biogenic processes are retained by adsorption to the coal, dissolution in associated water, and storage in fractures and pores (NEDERLOF, 1988).

Within coal beds, biogenic methane production appears to be associated with complex microbial communities, and occurs at higher rates in coal of lower rank or maturity (CATCHESIDE and RALPH, 1999; FAKOUSSA and HOFRICHTER, 1999a; GREEN et al., 2008; KRUGER et al., 2008;

LI et al., 2008; MIDGLEY et al., 2010; SHIMIZU et al., 2007; STRAPOC et al., 2008a). Coal has a complex and heterogeneous structure, containing aromatic, polycyclic aromatic and aliphatic moieties. This structural complexity presents a challenge to microbial degradation, requiring complex associations of bacteria to depolymerize coal and ferment hydrocarbon monomers, yielding CO2, acetate and other substrates that are reduced to methane by methanogenic archaea

(FAKOUSSA and HOFRICHTER, 1999b; HARRIS et al., 2008; OREM et al., 2010; STRAPOC, 2011;

THAUER et al., 2008).

The specific pathway for transforming coal to methane depends upon the microbial community and geochemistry of the sedimentary basin. Enhancing biogenic methane production by in situ microbial communities requires an understanding of naturally occurring microbial associations and their physiological constraints. Several recent studies used both culturing and

16S rRNA cloning to characterize CBM microbial communities (FRY et al., 2009; GREEN et al.,

2008; KRUGER et al., 2008; LI et al., 2008; MIDGLEY et al., 2010; SHIMIZU et al., 2007; STRAPOC et al., 2008b). As part of a review on CBM (STRAPOC, 2011), I compiled the publically available sequences from these studies to show the high diversity of bacteria with considerable contribution from Firmicutes, Spirochetes, Bacteroidetes, and all of the subgroups of

Proteobacteria, as well as archaeal representation from Methanosarcinales, Methanomicrobiales and Methanobacteriales (Figures 1.6-1.9). While the bacterial communities in the studies are

diverse, the archaeal communities show the dominance of either CO2 reducing or methylotrophic methanogens, which is supported by the methane isotope data (Figure 1-5). Factors such as the availability of methyl group substrates, partial pressures of H2 and CO2, and competition for substrates with the bacterial community may control the archaeal diversity and ultimate methanogenic pathway (CORD-RUWISCH et al., 1988; KOTSYURBENKO et al., 2001).

1.5 Organization of the Thesis

This work was written over the course of my doctoral study at Penn State from the Fall of

2005 to present under the supervision of Dr. Jennifer L. Macalady and Dr. Katherine H.

Freeman. The thesis comprises three main topics and four chapters of original research. All chapters have been written as publishable units.

Chapter 2 discusses a study of the anaerobic degradation of isoprenoid compounds including pristane, archaeal DGD and archaeal GDGT core lipids. Chapter 3 describes the preparation and use of 13C labeled phytane as a substrate for stable isotope probing in an anaerobic biodegradation experiment. Chapter 4 compares variation observed in the structure of halophilic archaeal lipids in hypersaline pools of Dead Sea water to variation observed in four

Halobacterial isolates. Chapter 5 presents a quantitative fluorescent in situ hybridization study of a CBM microbial community in the Cook Inlet Basin, Alaska.

1.6 Anticipated Publications from this Work

Chapter 2: Anaerobic biodegradation of the isoprenoid biomarker pristane will be submitted to

Environmental Microbiology in April with co-authors Katherine H. Freeman and Jennifer

L. Macalady. This manuscript was previously submitted to Environmental Microbiology in

an earlier form. Concerns about the connection between nitrate loss and the isoprenoid

substrate have been addressed through the use of 13C labeled phytane.

Chapter 3: Stable isotope tracing of anaerobic biodegradation with 13C labeled phytane will be

submitted to Environmental Science and Technology in May after incorporating material

from Chapter 2 with co-authors Katherine H. Freeman and Jennifer L. Macalady.

Chapter 4: Molecular characterization of archaeal lipids across a hypersaline gradient will be

submitted to Geobiology in April with co-authors Katherine H. Freeman and Jennifer L.

Macalady.

Chapter 5: Quantitative FISH analysis of microbial consortia from a biogenic gas field in the

Cook Inlet Basin, Alaska will be submitted to Applied Environmental Microbiology in

April with co-authors Dariusz Strapoc, Brad Huizinga, Matt Ashby and Jennifer L.

Macalady.

Work from this study contributed directly to a review on the biogeochemistry of CBM

formation, now in press:

Strapoc, D., Mastalerz, M., Dawson, K., Macalady, J. L., Callaghan, A., Wawrik, B., and

Ashby, M. 2011. Biogeochemistry of Coal-Bed Methane. Annual Review of Earth and

Planetary Sciences 39, 617-656.

1.7 Figures

Figure 1-1: Isoprene forms the basic biosynthetic unit of isoprenoid hydrocarbons. Examples of isoprenoid biopolymers that may serve as biomarkers include hopanes, steranes, chlorophyll-a, and archaeal membrane lipids such as archaeol and the glycerol dialkyl glycerol tetraethers

GDGT-0 and GDGT-4.

Figure 1-2: Several mechanisms for anaerobic hydrocarbon degradation have been proposed or experimentally demonstrated. Fumarate addition has been shown to activate a variety of n- alkanes and aromatic compounds. Carboxylation, dehydrogenation plus hydroxylation, methylation, and reverse methanogenesis are additional mechanisms of anaerobic hydrocarbon activation.

Figure 1-3: Membrane glycerolipid structures in the three domains of life. Archaea are unique in that they produce ether-linked, isoprenoid structures. Bacteria and eukaryotes produce ester- linked, fatty acid derived structures. Bacterial lipids with unusual ether linkages and membrane- spanning hydrocarbon chains (diabolic acids) are also shown.

Figure 1-4: Structures of archaeal glycerol diphytanyl glycerol tetraethers (GDGT’s) and diphytanyl glycerol diethers (DGD’s) including structures with cyclopentyl moieties, unsaturation and hydroxyl subsitutions.

Figure 1-5: Plot of carbon and hydrogen isotopic values of methane showing the differences between thermogenic and biogenic methane generated from different pathways. Arrows show the direction of increasing thermal maturity. From Strapoc et al. (2011).

Figure 1-6: Phylogenetic tree of 16S rDNA sequences of Firmicutes, Bacteroidetes,

Planctomycetes, Spirochaetes, and other bacterial lineages associated with coal bed methane from four sedimentary basins: Illinois – red; Powder River – orange; Ishikari – blue; Alberta - purple. The neighbor-joining phylogram was generated in PAUP* v.4b10. Bootstrap values from neighbor-joining and maximum-parsimony (in that order) are shown for each node.

Figure 1-7: Phylogenetic tree of 16S rDNA sequences of Betaproteobacteria and

Gammaproteobacteria associated with coal bed methane from two sedimentary basins Ishikari – blue; Alberta - purple. The neighbor-joining phylogram was generated in PAUP* v.4b10.

Bootstrap values from neighbor-joining and maximum-parsimony (in that order) are shown for each node.

Figure 1-8: Phylogenetic tree of 16S rDNA sequences of Alphaproteobacteria,

Deltaproteobacteria and Epsilonproteobacteria associated with coal bed methane from three sedimentary basins: Illinois – red; Ishikari – blue; Alberta - purple. The neighbor-joining phylogram was generated in PAUP* v.4b10. Bootstrap values from neighbor-joining and maximum-parsimony (in that order) are shown for each node.

Figure 1-9: Phylogenetic tree of 16S rDNA sequences of methanogenic archaea associated with coal bed methane from six sedimentary basins: Illinois – red; Powder River – orange; Ruhr – brown; Ishikari – blue; Cook Inlet – green; Alberta - purple. The neighbor-joining phylogram was generated in PAUP* v.4b10. Bootstrap values from neighbor-joining and maximum- parsimony (in that order) are shown for each node.

1.9 References

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Aitken C. M., Jones D. M., and Larter S. R. (2004) Anaerobic hydrocarbon biodegradation in deep subsurface oil reservoirs. Nature 231, 291-294.

Albaiges J., Borbon J., and Walker I. W. (1985) Petroleum isoprenoid hydrocarbons derived from catagenetic degradation of archaebacterial lipids. Organic Geochemistry 8(4), 293- 297.

Anderson R., Kates M., Baedecker M. J., Kaplan I. R., and Ackman R. G. (1977) The stereoisomeric composition of phytanyl chains in lipids of Dead Sea sediments. Geochimica et Cosmochimica Acta 41(9), 1381-1387, 1389-1390.

Bastin E. S., Greer F. E., Merritt C. A., and Moulton G. (1926) The Presence of Sulphate Reducing Bacteria in Oil Field Waters. Science 63(1618), 21-24.

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25 Chapter 2: Anaerobic biodegradation of the branched hydrocarbon biomarker analog pristane

2.1 Abstract

Isoprenoid hydrocarbons contribute up to 30% by mass of crude oil and include nearly all of the biomarker compounds used in paleoenvironmental and paleoecological reconstructions of earth history. Circumstantial evidence suggests that biodegradation of isoprenoids occurs in anoxic environments such as sediments and oil reservoirs, but no data on the phylogeny of microorganisms responsible for this process have been published to date. Here we describe pristane biodegradation and accompanying loss of nitrate in an activated sludge enrichment.

We followed the evolution of the enrichment community using 16S rDNA clone libraries and fluorescence in situ hybridization (FISH). Clone libraries show a transition from a

Bacteriodales-dominated community to a lower-diversity consortium dominated by relatives of Pseudomonas stutzeri (Gammaproteobacteria) and relatives of a denitrifying

Betaproteobacterium. Consistent with changes in clone libraries, FISH experiments show an increase in cells hybridizing with probes specific for Gamma- and Betaproteobacteria in later subcultures. We derive the pristane consumption reaction stoichiometry from quantitative growth experiments. In addition to pristane, the enrichment community is capable of utilizing archaeal glycerol diether lipids but not glycerol tetraether lipids (GDGTs). The lack of GDGT disapearance suggests that access to terminal or subterminal carbons may be required for the acyclic isoprenoid biodegradation pathway utilized by the enrichment.

2.2 Introduction

Anaerobic hydrocarbon degradation has recently been the subject of reviews, highlighting significant progress in understanding the associated microbiology and

biochemistry (HEAD et al., 2003; HEIDER, 2007; LOVLEY, 2001; SPORMANN and WIDDEL,

2000; WIDDEL et al., 2006). The studies described in these reviews primarily focus on the anaerobic biodegradation of anthropogenic pollutants such as benzene and toluene and gaseous hydrocarbons such as methane and propane. In contrast, the anaerobic biodegradation of both linear and cyclic isoprenoid hydrocarbons has received little attention to date despite the relevance of these compounds to petroleum characterization and the geologic record of life on earth. Isoprenoid hydrocarbons can contribute up to 30% by mass of crude oil deposits

(TISSOT and WELTE, 1984) and include essential biomarkers for taxonomic groups (BROCKS and PEARSON, 2005; DUTKIEWICZ et al., 2006; PETERS et al., 2005), paleoenvironmental conditions such as temperature and salinity (WUCHTER et al., 2006), and petroleum maturity

(PETROV and ABRYUTINA, 1989).

Isoprenoid compounds, derived from polymers of isoprene, are produced by organisms in all three domains of life. Their long-term preservation potential makes isoprenoids useful geologic biomarkers with both taxonomic and paleoenvironmental specificity (BROCKS et al., 2003; DUTKIEWICZ et al., 2006; PETERS et al., 2005). For example, glycerol diether glycerol tetraethers (GDGT’s) are indicative of archaea and show a relation to temperature, salinity and nutrient concentrations (TURICH et al., 2007; WUCHTER et al., 2006).

Although direct evidence for the microbial role in anaerobic isoprenoid degradation is usually lacking, microorganisms are implicated by the simultaneous disappearance of isoprenoids and electron acceptors such as sulfate, iron and nitrate, by gas evolution, and by changing hydrocarbon profiles (BONIN et al., 2002; GROSSI et al., 2000; HEAD et al., 2003; MASSIAS et al., 2003).

The methyl branched structure of isoprenoids imparts additional recalcitrance to lipid hydrocarbons by hindering the beta-oxidation pathway (CANTWELL et al., 1978; SCHAEFFER et al., 1979). In studies of aerobic branched alkane degradation, this hindrance is overcome

through additional oxidative decarboxylation steps (PIRNIK et al., 1974; PIRNIK and

MCKENNA, 1977). The need for repeated demethylation, decarboxylation or alternative activation steps before beta-oxidation can proceed would effectively reduce the microbial metabolic energy yield from isoprenoid substrates, and may explain why their degradation rates in natural environments appear slower than equivalent non-isoprenoid hydrocarbons. In addition, iso-methyl branching at the chain ends (e.g. pristane) or near the chain ends (e.g. archaeol) may effectively prevent some activation mechanisms due to steric hindrance.

Anaerobic isoprenoid biodegradation has been demonstrated in only two laboratory studies utilizing pristane. These studies demonstrate anaerobic pristane degradation by a diesel-fuel contaminated aquifer enrichment under denitrifying conditions (BREGNARD et al.,

1997) and a marine sediment enrichment under sulfate-reducing, methanogenic conditions

(GROSSI et al., 2000). However, neither study provides a phylogenetic description of the associated microbial community. Here we report the results of a denitrifying, pristane degradation enrichment study. Full cycle rRNA methods including 16S rDNA cloning and

FISH revealed the phylogenetic affiliation of the dominant organisms associated with degradation. Quantitative measurements of nitrate loss and carbon dioxide evolution and isoprenoid compound ratios (pristane to phytane (Pr/Ph) and pristane to squalane (Pr/Sq)) are used to propose a biodegradation reaction stoichiometry.

2.3 Materials and Methods

2.3.1 Source of bacteria and cultivation

Activated sludge samples for isoprenoid degrading enrichments were obtained from an activated sludge unit at the Pennsylvania State University wastewater treatment plant. Solid material was pelleted by centrifugation, and the supernatant was used as an inoculum.

Thermoplasma acidophilum (DSMZ 1728) was obtained from the Deutsche Samlung von

Mikroorganismen und Zellkulturen , Braunschweig. Haloferax sulfurifontis strain SD1 was obtained from Lee Krumholz, University of Oklahoma. Thermoplasma acidophilum was grown as described by Langworthy et al. (1972). Haloferax sulfurifontis was grown as described by Elshahed et al. (2004).

Anaerobic cultivation was carried out using a modified multipurpose medium described by Widdel and Bak (1992). Trace element, vitamin and selenate-tungstate solutions were prepared as described. The basal medium was modified to include per liter: NaCl, 1g;

. MgCl2, 0.4g; CaCl2 2H20, 0.1g; Na2SO4, 0.5g; NH4Cl, 0.25g; KH2PO4, 0.2g; KCl, 0.5g;

KNO3, 0.25g. Enrichments were set up in triplicate in 200 ml serum bottles, under anoxic conditions. Bottles were prepared using 30 ml of medium, 10 ml of inoculum, 120mg of

2,6,10,14-tetramethylpentadecane (pristane, 98.9%, Chem Service Inc., West Chester, PA), 2g of ashed sand, a 1.5 atm headspace of N2, and butyl rubber stoppers (Bellco, Vineland, NJ).

Controls included both uninoculated bottles containing pristane and inoculated bottles with no added carbon substrate. The amount of pristane in subcultures was reduced to 10mg and inoculum volume was reduced to 400 ul. Aliquots of the third transfer of the enrichment cultures were preserved at -80°C in 25% glycerol. The experiments described in this chapter were initiated from revived glycerol preserved stocks. One transfer was required to obtain cultures where nitrate loss was only observed in the presence of pristane (glycerol from preservation was fully consumed).

2.3.2 Archaeal lipid extraction

Glycerol tetraethers including caldarchaeol and analogs containing one to five cyclopentane rings were prepared from laboratory cultures of Thermoplasma acidophilum

(SWAIN et al., 1997). Glycerol diethers were obtained from cultures of Haloferax sulfurifontis, which produces membrane lipids dominated by C20-20 glycerol diethers (ELSHAHED et al.,

2004). Lipids were extracted from lyophilized cell pellets with CH3OH/CHCl3/H2O (2:1:0.8).

Core lipids were then prepared from the total lipid extract by strong acid methanolysis (10:1:1

CH3OH/CHCl3/HCl) at 100°C for 1 hr (MACALADY et al., 2004). Pigments were removed from Haloferax sulfurifontis total lipid extracts prior to strong acid methanolysis by an overnight precipitation of diether lipids with several volumes of cold acetone (CHOQUET et al.,

1992).

2.3.3 Monitoring growth

IonPac AS18 column with an isocratic elution program using 45mM KOH at a flow rate of 1 ml/min and an oven temperature of 31°C. Anion concentrations were normalized to chloride.

For dissolved inorganic carbon (DIC) analysis, 1 ml aliquots of culture were take during growth and transferred to ashed 8 ml serum bottles with a 1 atm helium headspace and butyl rubber stoppers. Subsequently, these aliquots were acidified with 2 drops phosphoric acid

(12N). DIC samples were analyzed on an SRI 310 gas chromatograph (SRI Instruments,

Menlo Park, CA) with a TCD detector and a PoraPak Q packed column (3’x1/8‖) at 80°C with a helium carrier phase.

2.3.4 Determination of pristane loss and putative metabolites

After 2-3 months of continuous nitrate loss, one of the culture replicates and one of the sterile control replicates were extracted to determine the loss of pristane and the presence of putative metabolites. Cultures were acidified to pH < 2.0 with HCl (6N) and extracted twice with 20 ml of ethyl acetate (KROPP et al., 2000). The pooled extracts were concentrated by

rotary evaporation and subsequently evaporated to near dryness under a stream of N2. Due to the high concentration of pristane and potentially low concentration of alcohol or acid metabolites, 6 ml Supelclean LC-Si SPE tubes (Supelco, Bellefonte, PA, USA) were used to separate the alkane fraction. Elution from the SPE tubes was as follows: 6 ml hexane; 6 ml ethyl acetate; 6 ml of methanol. The ethyl acetate and methanol fractions were derivatized in

100 ul hexane using 25 ul of N,O-bis(trimethylsilyl)trifluoroacetimide (BSTFA) and 25 ul pyridine at 65°C for 20 min. The hexane fraction and derivatized fractions were run on a

Hewlett-Packard 5972 GC/MS with a DB-5 column (length 30m; inside diameter, 0.25 mm; film thickness, 0.25 μm). The injector temperature was 320°C, and the oven was held at 65°C for 1 min before its temperature was increased by 6°C/min to 320°C and held for 20 min. The limit of detection of pristanic acid, a putative metabolite in this experiment, is 0.01 umol/l

(WANDERS and DURAN, 2008).

2.3.5 Phylogenetic analysis

DNA extraction, PCR amplification and cloning were carried out exactly as described in Macalady et al. (2008). Clones were sequenced at the Penn State University Biotechnology

Center using T3 and T7 plasmid-specific primers. CodonCode Aligner v.1.2.4 (CodonCode

Corp., Dedham, MA, USA) was used to assemble sequences and manually check for ambiguities. Assembled gene sequences were compared against the public databases using

BLAST (ALTSCHUL et al., 1990) and chimera checked with Bellerophon 3 (HUBER et al.,

2004) and CHIMERA_CHECK v.2.7 (COLE et al., 2003). Putative chimeras were excluded from subsequent sequence analysis. Non-chimeric sequences were aligned using the NAST aligner (DESANTIS et al., 2006b), added to an existing alignment in ARB (LUDWIG et al.,

2004), and manually refined. Distance based analysis was performed with DOTUR (SCHLOSS

and HANDELSMAN, 2005) using distance matrices calculated using the online phylogenetic tools available at Greengenes (DESANTIS et al., 2006a).

2.3.6 Nucelotide sequence accession numbers

The 16S rRNA gene sequences determined in this study were submitted to the GenBank database under accession numbers GU222220-GU222257 and GU250847-GU250871.

2.3.7 Fluorescence in situ hybridization

Aliquots of enrichment cultures for FISH were fixed in 3 volumes of 4% (w/v) paraformaldehyde in 1x PBS for 2-3 hours at 4°C and stored in 1:1 PBS/ethanol solution at -

20°C. FISH experiments were carried out as described in Hugenholtz et al. (2001) using probes listed in Table 2. Populations of Betaproteobacteria, Gammaproteobacteria and

Bacteriodales were determined from counts of between 1000-2000 DAPI-stained cells for each sample/probe combination. The CFB719 probe described by Weller et al. (2000) was modified using the probe design tools in the ARB software package to increase specificity for

Bacteriodales clones. The designed oligonucleotide was synthesized and labeled at the 5’ end with the indocarbocyanine dye CY3 (Sigma-Aldrich, St. Louis, MO).

2.4 Results

2.4.1 Anaerobic enrichment of denitrifying bacteria with pristane

We enriched pristane-degrading denitrifying bacteria using an enrichment medium described by Widdel and Bak (1992) with 2.5 mM nitrate and pristane (0.3 % initially, v/v).

. -1 The solubility of pristane is low, estimated as 1.0 mg L at 25°C (EASTCOTT et al., 1988), and thus it was in excess in all experiments, forming a separate phase in enrichments. Activated sludge from a waste water treatment plan (University Park, PA) served as a source of

inoculum. After 6 months, enrichment cultures had consumed approximately 14.5 mM nitrate

(80 μM/day) in comparison with approximately 0.38 mM in a pristane-free control and 0.48 mM in an uninnoculated control containing pristane. Subsequent transfers (1 % inoculum, v/v) show somewhat reduced rates of denitrification, ~70 μM/day.

The most abundant cells in the enrichment cultures have rod-shaped morphology (0.91

μm ± 0.15 μm by 0.44 μm ± 0.06 μm; Fig 1) and occur either as planktonic cells or more commonly in large flocs (10-20 μm diameter). These cells may represent the bacterium responsible for pristane degradation. However, several transfers in liquid medium failed to result in a pure culture. Additionally, an attempt at plating on agar with pristane failed to produce colonies. We therefore attempted to identify the dominant organisms on the basis of

16S rDNA clone libraries and fluorescence in situ hybridization (FISH).

2.4.2 Phylogenetic analysis of the enrichment cultures

A 16S rDNA clone library constructed using universal primers and the pristane degrading enrichment after the first transfer had a relatively low diversity. No archaeal sequences were retrieved. Most clones (75%) were members of a single phylotype in the

Bacteriodales family. Based on BLAST searches and neighbor joining phylogenies constructed using the phylogenetic software ARB, these clones are related to anaerobic fatty acid degraders and environmental clones from organic-rich and/or anoxic environments.

Other clones in the library are closely related to Gamma- and Betaproteobacterial isolates and environmental clones associated with hydrocarbons in organic-rich environments.

We constructed two additional 16S rDNA clone libraries from the fourth transfer of parallel enrichments. These libraries document a decrease in overall diversity as well as a strong shift in the most abundant phylotypes. The Bacteriodales clones abundant in the first clone library became a minor component of the later libraries, representing less than 10% of

the sequenced clones. In contrast, Gamma- and Betaproteobacterial clones rose in abundance.

In particular, the later clone libraries are dominated by two phylotypes: ~30% are related to a denitrifying, poly(3-hydroxybutyrate-co-3-hydroxyvalerate) (PHB-HV) degrading

Betaproteobacteria isolate ( >99% identity) (Figure 2) and ~50% are related to the

Gammaproteobacteria Pseudomonas stutzeri ( >99% identity) (Figure 3).

Oligonucleotide probes specific to the Bacteriodales family (CFB719) (WELLER et al.,

2000), and the Beta and Gamma subclasses of Proteobacteria (Bet42a, Gam42a) (MANZ et al., 1992) were used in FISH experiments to obtain quantitative information about the population structure of the pristane-degrading enrichments (Table 2). Consistent with the decline in the number of clones in the Bacteriodales family, the number of cells hybridizing to CFB719 decreases from 52.7 ± 11.4% to 2.8 ± 1.7%. Meanwhile, cells hybridizing to the oligonucleotide probe GAM42a increase from of 36.7 ± 10.3% to 69.0 ± 23.0% of total bacterial cells. Cells hybridizing to BET42a represent 27.3 ± 15.2% of the fourth transfer.

Changes in the fraction of cells hybridizing to each oligonucleotide probe are given Figure 4.

2.4.3 Quantitative growth experiment

Experiments show the consumption of nitrate concomitant with the production of nitrite and bicarbonate in enrichments containing pristane, but not in inoculated controls with no added carbon substrate. Sulfate concentrations remained constant over the course of the experiment. After forty days of growth, the consumption of 2.0 mM nitrate resulted in the production of 1.9 mM nitrite and 1.1 mM bicarbonate (Figure 5). The stoichiometry implied by these data is discussed below.

Pristane obtained from ChemService (West Chester, PA) had a supplier certified purity of >98.6%. Trace impurities were determined by GC-MS to be phytane and squalane.

The peak areas of pristane, phytane and squalane were determined from total ion

chromatograms in the original pristane as well as a pristane-degrading enrichment and control culture at the end of the growth experiment. The ratios of the isoprenoids and the percent differences between the original pristane and extracted cultures are given in Table 1. The ratio of pristane/phytane and pristane/squalane in the control culture decreased and differed from the original pristane by 6.53 and 5.48% respectively. In the pristane-degrading enrichment, the same ratios decreased and differed from the original pristane by 40.62 and 32.61% respectively. The preference for pristane over the chemically similar phytane and squalane is curious. However, precedence does exist for the utilization of discrete alkane size fractions in hydrocarbon degrading isolates (WIDDEL and RABUS, 2001). While alkane size preference may explain the preference for pristane (C19) over squalane (C30), the observed recalcitrance of phytane (C20) remains an odd result.

2.4.4 Test of archaeal DGD and GDGT core lipids as potential substrates

The ability of pristane-degrading enrichment to degrade geologically relevant biomarkers was tested using archaeal diphytanyl glycerol diether (DGD) and glycerol diphytanyl glycerol tetraether (GDGT) core lipids. Due to the structural similarity of the hydrocarbon chains of these membrane lipids to pristane, we hypothesized that the enrichment should be able to utilize them at a similar rate. Additionally, using DGD and GDGT core lipids allows us to test the inhibitory effect of glycerol ether linkages on isoprenoid chain degradation. Incubations with pristane served as a positive control to verify that transferred cells remained active. The progress was determined by nitrate loss (Figure 6). Incubations with DGD core lipids showed a similar rate of dentrification as the pristane incubations, while incubations with GDGT core lipids showed nitrate loss similar to no-carbon and uninocculated controls.

2.5 Discussion

To our knowledge, this study provides the first description of phylogeny of anaerobic isoprenoid degrading microorganisms. The dominant phylotype in the low-diversity community, representing >70% of the total cells, is closely related to Pseudomonas stutzeri

(>99%). Strains of P. stutzeri are known to be denitrifiers, as well as to have the capability of degrading a broad array of organic compounds including aromatic, polycyclic aromatic and aliphatic hydrocarbons (LALUCAT et al., 2006). Two other phylotypes present in the enrichment could have syntrophic interactions with the P. stutzeri strain. The dominant

Betaproteobacterial clone, which shares more than 99% identity with a PHB-HV degrading isolate (KHAN et al., 2002), may be subsisting on polyhydroxyalkanoates produced by P. stutzeri as storage granules (GUO-QIANG et al., 2001). Although nitrite was produced stoichiometrically with nitrate loss in the quantitative growth experiment, dissimilatory nitrate reduction to ammonium (DNRA) remains a possible alternative metabolism in this pristane degrading community.

Denitrifying bacteria from the Alpha, Beta and Gamma subclasses of Proteobacteria are associated with the anaerobic degradation of alkanes, aromatic and polycyclic aromatic hydrocarbons (HEIDER et al., 1998; SPORMANN and WIDDEL, 2000; WIDDEL et al., 2006;

WIDDEL and RABUS, 2001). Additionally, several previously described strains of

Pseudomonas sp. are associated with both aerobic and anaerobic hydrocarbon degradation. Of particular note is the association of one cluster of the Pseudomonas genus, containing P. alcaligenes, P. citronellolis, P. medocina and P. aeruginosa as well as P. stutzeri,

(YAMAMOTO et al., 2000) with both aerobic and anaerobic degradation of unsaturated isoprenoids and isoprenoic primary alcohols (CANTWELL et al., 1978; HARDER and PROBIAN,

1995).

The stoichiometry of pristane degradation can be hypothesized from the quantitative growth experiment. During the reduction of approximately 80 μmol nitrate to nitrite, 40 μmol

- HCO3 was produced. Assuming a range for carbon assimilation from 20% to 50% (WHITE,

2007) and a cellular composition of C5H7O2N (DRTIL et al., 1995), this represents the consumption of between 3.0 and 4.0 μmol of pristane (10 – 14% of the initial pristane). The combined assimilatory and dissimilatory reactions follow.

For 20% assimilation:

- - - C19H40 + 30.4NO3 + 0.76NH3 + 16.72H2O  15.2HCO3 + 0.76C5H7O2N + 30.4NO2 +

55.2H+

For 50% assimilation:

- - - + C19H40 + 19NO3 + 1.9NH3 + 13.4H2O  9.5HCO3 + 1.9C5H7O2N + 19NO2 + 49.7H

During the growth experiments, we observed that pH consistently increased over time. In the quantitative growth experiment, we measured an increase from pH 6.5 to 7.1 over 39 days as well as an increase to pH 8-9 after several months. The observed pH increase strongly suggests that secondary reactions are occurring in conjunction with pristane oxidation.

Under the conditions of this denitrifying enrichment, pristane biodegradation follows first order kinetics with k = 0.95 y-1 (r2 = 0.76, n=8) to 1.57 y-1 (r2 = 0.77, n = 8). This is slower than isoprenoid degradation rates reported by Grossi et al. (2000) for pristane (k =

3.5y-1), squalene (k = 18.5 y-1) and phytadienes (k = 6.6 y-1), degraded under sulfate reducing conditions. In both denitrifying and sulfate reducing enrichments, rate constants are several orders of magnitude greater than the in situ maximum biodegradation rate constant for mixed hydrocarbons in oil fields (10-4 y-1) where oxidant and nutrient supplies are limited by diffusion (LARTER et al., 2003).

Under anaerobic conditions, fumarate addition (HEIDER, 2007) and carboxylation

(CALLAGHAN et al., 2009) are potential methods of aliphatic hydrocarbon activation. We were

unable to detect fumarate addition products or other isoprenoid acids in our enrichments despite repeated efforts to concentrate the organic acid fraction of the culture extracts.

However, a comparison of results from enrichments in the presence of different isoprenoid substrates provides some insight about the location of the initial attack. The lack of nitrate reduction in the presence of GDGT core lipids and the near equivalent rates of nitrate reduction in the presence of pristane and DGD core lipids suggest that activation occurs at a terminal or subterminal position on the hydrocarbon chain. The ether linkages flanking the isoprenoid chain in GDGT lipids appear to inhibit degradation under the conditions of our enrichments. Further experiments will be required to identify the activation mechanism and biodegradation pathway for the anaerobic biodegradation of pristane and similar compounds by the pristane-degraders enriched in this study.

5 -1 Considering the density of archaea in marine environments (10 ml ) (DELONG and

-1 PACE, 2001) and the concentration of GDGT biomarkers in sediments (μg’s g sediment)

(HUGUET et al., 2009) degradation must be occurring but may be limited to aerobic degradation in the water column and oxic sediment/water interfaces. Reduced preservation efficiency of GDGT’s in oxic surface sediments as compared to anoxic/suboxic sediments with similar or lower mass accumulation rates suggests aerobic GDGT degradation processes are active (SINNINGHE DAMSTÉ et al., 2002). Anaerobic alteration of GDGTs may be limited to recycling of the molecules by archaea as shown by Takano et al. (2010) in marine sediment cores. The combination of energetic constrains from GDGT synthesis, anaerobic ether linkage degradation and lower energy yielding metabolisms like sulfate reduction and methanogenesis in anaerobic settings could encourage reuse rather than degradation of large membrane lipid compounds. Further enrichment experiments will be necessary to determine if inocula from additional environmental sources or electron acceptors other than nitrate and sulfate can overcome the energetic barrier presented by tetraether linkages.

2.6 Acknowledgments

We thank Z Zhang and D Walizer for laboratory assistance. This research was supported by an American Chemical Society Petroleum Research Fund (grant 48445-AC2), and the Penn

State Biogeochemical Research Initiative for Education (BRIE) (NSF grant DGE-9972759).

2.7 Figures and Tables

Figure 2-1: FISH micrographs of cells in the denitrifying, pristane-degrading enrichment culture. Bar, 10 μm. All cells have been stained with DAPI (blue) and hybridized with

Bacteria specific probe EUB338 (green). Top row: Cells hybridized with

Gammaproteobacteria specific oligonucleotide probe GAM42a (red). Middle row: Cells hybridized with Betaproteobacteria specific oligonucleotide probe BET42a (red). Bottom row: Cells hybridized with Bacteriodales specific probe CFB719 (red).

Figure 2-2: Neighbor-joining phylogenetic tree showing Betaproteobacteria. Pristane degrading enrichment clones are shown in bold followed by the number of clones of each phylotype. PiaDegr clones are from the 2nd transfer. NP13 and NP6D clones are from two independent 4th transfers. Maximum parsimony (MP) and neighbor-joining (NJ) bootstrap values >50% are shown (MP/NJ). Filled circles indicate nodes that were present in the maximum likelihood phylogeny.

Figure 2-3: Neighbor-joining phylogenetic tree showing Gammaproteobacteria. Pristane degrading enrichment clones are shown in bold followed by the number of clones of each phylotype. PiaDegr clones are from the 2nd transfer. NP13 and NP6D clones are from two independent 4th transfers. Maximum parsimony (MP) and neighbor-joining (NJ) bootstrap values >50% are shown (MP/NJ). Filled circles indicate nodes that were present in the maximum likelihood phylogeny. Putative pristane-metabolizing cells are annotated by ―‖.

Figure 2-4: Fraction of bacterial cells hybridizing to specific oligonucleotide probes as a function of transfer to subcultures.

Figure 2-5: Anaerobic consumption of nitrate (top) and the concomitant production of nitrite

(middle) and bicarbonate (bottom) in a denitrifying, pristane degrading enrichment culture.

Figure 2-6: Anaerobic consumption of nitrate in denitrifying enrichments with pristane and

DGD core lipids. Inoculated cultures containing GDGT core lipids or no added carbon substrate show only minimal loss of nitrate over the same incubation period.

Table 2-1: Ratios of isoprenoid peak areas in GC-MS chromatograms

Sample Pristane/Phytane Pristane/Squalane Phytane/Squalane

Pristane (stock) 249.50 455.21 1.82

Uninoculated Culture 233.72 430.90 1.84

Pristane-Degrading Enrichment 165.26 327.58 1.98

Percent Difference 6.53 5.49 1.09 (Pristane – Uninoculated) Percent Difference 40.62 32.61 8.42 (Pristane – Enrichment)

Table 2-2: Oligonucleotide probes used in this study

Probe Target group Sequence (5’ 3’) Formamide (%) Target site Reference

EUB338a Most Bacteria GCTGCCTCCCGTAGGAGT 0 – 50 16S (338-355) Amann et al. (1995)

EUB338-IIa Planctomycetales GCAGCCACCCGTAGGTGT 0 – 50 16S (338-355) Daims et al. (1999)

EUB338-IIIa Verrucomicrobiales GCTGCCACCCGTAGGTGT 0 – 50 16S (338-355) Daims et al. (1999)

GAM42a Gammaproteobacteria GCCTTCCCACATCGTTT 35 23S (1027-1043) Manz et al. (1992) cGAM42a Competitor GCCTTCCCACTTCGTTT 35 23S (1027-1043) Manz et al (1992)

BET42a Betaproteobacteria GCCTTCCCACTTCGTTT 35 23S (1027-1043) Manz et al. (1992) cBET42a Competitor GCCTTCCCACATCGTTT 35 23S (1027-1043) Manz et al. (1992)

CFB719b Most Bacteriodetes AGCTGCCTTCGCTATCGG 30 16S (719-737) Weller et al. (2000)

aCombined in equimolar amounts to make EUBMIX.

bModified to increase specificity for clones in this study.

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Chapter 3: Stable isotope tracing of anaerobic isoprenoid biodegradation with 13C-labeled phytane

3.1 Abstract

Laboratory incubations of a denitrifying, pristane degrading enrichment were examined using a stable isotope tracer. Anaerobic biodegradation of 13C-labeled phytane was examined by ion chromatography analysis of nitrate loss and nitrite production accompanied by GC-TCD

13 12 quantification of CO2 production and stable isotope analysis to determine the C/ C ratio of

13 CO2. The C-labeled phytane for the incubations was prepared from the membrane lipids of

Haloferax sulfurifontis SD1 and purity was confirmed by GC-MS chromatography. After 82

13 13 days, the CO2 produced in incubations grown with C-labeled phytane was δ C 13.5 ± 7.8, as compared to incubations grown with unlabeled phytane δ13C -35.2 ± 0.5 and no added carbon substrate δ13C -33.9 ± 1.7. Full cycle rRNA analysis of the phytane degrading community indicate the putative phytane-degrading bacteria in the incubations are Gammaproteobacteria with >99% similarity to Pseudomonas stutzeri and Betaproteobacteria in the Comomonadaceae family.

3.2 Introduction

In microbial ecology, stable isotope tracing with isotopically enriched substrates is utilized to identify metabolically active members of a community in the environment or an enrichment culture. This technique depends upon the incorporation of the enriched isotope

(i.e.13C, 2H, 15N, 18O) into cellular components including DNA, rRNA and membrane lipids

(DUMONT and MURRELL, 2005; RADAJEWSKI et al., 2000). Additionally, isotopically labeled

compounds facilitate biodegradation research in the identification of metabolites and transient intermediates (DAVIDOVA et al., 2005; WILKES et al., 2003). While isotopically labeled tracers are powerful tools in microbial ecology, their use is limited by the commercial availability of labeled compounds. For example, studies of anaerobic hydrocarbon biodegradation primarily focus on anthropogenic pollutants such as benzene and toluene or short-chain n-alkanes from methane to hexane (HEIDER, 2007; LOVLEY, 2001; SPORMANN and WIDDEL, 2000; WIDDEL et al.,

2006). Conversely, there are few studies on the anaerobic biodegradation of geologically relevant biomarkers.

Glycerol diether glycerol tetraethers (GDGT’s) and diphytanyl glycerol diethers (DGD’s), are biomarkers indicative of archaea and show a relation to temperature, salinity and nutrient concentrations (TURICH et al., 2007; WUCHTER et al., 2006). GDGT’s and DGD’s derive from archaeal membrane lipids and are characterized by a glyceride that is ether-linked to an isoprenoid chain in contrast to the ester-linked fatty acids of bacteria and eukaryotes. Anaerobic isoprenoid degradation has been demonstrated in two laboratory enrichment studies utilizing pristane (BREGNARD et al., 1997; GROSSI et al., 2000). In chapter 2, we described a third study of a denitrifying pristane degrading enrichment. Full cycle rRNA methods including 16S rRNA cloning and FISH revealed the dominant organisms associated with degradation to be closely related to a denitrifying, poly(3-hydroxybutyrate-co-3-hydroxyvalerate) degrading

Betaproteobacteria and a Gammaproteobacteria closely related to Pseudomonas stutzeri.

Here we provide confirmation of isoprenoid utilization by bacteria enriched in Chapter 2 through growth of incubations on 13C-labeled phytane (2,6,10,14-tetramethylhexadecane) and

13 13 12 measurement of the δ Cbicarbonate produced during bacterial growth. The ratios of C/ C show a clear difference between incubations grown on 13C-labeled phytane as compared to incubations

grown on unlabeled phytane or with no added carbon substrate. Additionally, we describe the preparation of 13C-labeled phytane from the membrane lipids of Haloferax sulfurifontis SD1 through the conversion of the ether-linked hydrocarbon chain to an alkyl iodide followed by reductive dehalogenation to the hydrocarbon as shown in Figure 3-1.

3.3 Experimental Procedures

3.3.1 Source of bacteria and cultivation

Bacteria were enriched from activated sludge from the Pennsylvania State University wastewater treatment plant during a previous study of anaerobic pristane degrading bacteria

(Chapter 2). Anaerobic cultivation was carried out using a modified multipurpose medium described by Widdel and Bak (1992). Trace element, vitamin and selenate-tungstate solutions were prepared as described. The basal medium was modified to reduce sulfate concentration and

. included per liter: NaCl, 1g; MgCl2, 0.4g; CaCl2 2H20, 0.1g; Na2SO4, 0.5g; NH4Cl, 0.25g;

KH2PO4, 0.2g; KCl, 0.5g; KNO3, 0.25. Triplicate incubations in 40 ml serum bottles were set up under anoxic conditions, and 10 μl of phytane was introduced by a gas-tight syringe following sterilization. Controls for the stable isotope tracer experiment included uninoculated bottles with

13C-labeled phytane and inoculated bottles with no added carbon source or with unlabeled phytane (99%, Ultra, Kingstown, RI). Incubations were inoculated from glycerol preserved stocks of the pristane degrading enrichment (1%, v/v). The headspace consisted of 1 atm N2.

3.3.2 Archaeal cultivation

Haloferax sulfurifontis strain SD1 (Lee Kumholz, University of Oklahoma) was grown in the modified liquid medium described by Elshahed et al. (2004) for growth on sugars. The

medium contained per liter 150.0g NaCl; 20.0g MgCl2; 12.0g Hepes; 5.0g K2SO4; 0.1g CaCl2;

0.1g yeast extract; 0.5g D-glucose. Glucose added to cultures was a mixture of 1% U-13C6 (99%,

Cambridge Isotope Laboratories, Andover, MA) and 99% unlabeled glucose for δ13C ≈ 900 ‰.

The expected 13R and 13F for glucose can be calculated from the δ13C of the unlabeled glucose and the mole fraction of labeled glucose. Fractional abundance is defined as 13F =

13C/(13C + 12C) and the carbon isotope ratio is defined as 13R = 13C/12C. Using the relationship,

13 13 13 Ra = RVPDB(δa/1000 +1), we converted the δ C value of unlabeled glucose to an isotopic ratio

13 13 13 where RVPDB = 0.0112373. The isotopic ratio was converted to F using the relationship, F =

13R/(13R + 1). The expected 13F of the added glucose was determined using a mass balance

13 13 13 relation for the labeled and unlabeled glucose, FT = Flabeled χ + Funlabeled(1-χ) where χ is the

13 mole fraction of labeled glucose and Flabeled ≈ 0.99.

3.3.3 Lipid extraction and analysis

Lipids were obtained from lyophilized cell pellets by a modified Bligh-Dyer extraction as described in Macalady et al. (MACALADY et al., 2004). Briefly, the lipids were extracted with

CH3OH/CHCl3/H2O (2:1:0.8). Neutral lipids were removed from Haloferax sulfurifontis total lipid extracts prior to strong acid methanolysis by an overnight precipitation of polar lipids with several volumes of cold acetone (CHOQUET et al., 1992). Core lipids were prepared from the polar lipids by strong acid methanolysis (10:1:1 CH3OH/CHCl3/HCl) at 100°C for 1 hour.

Methanolysed lipids were extracted with hexane/CHCl3 (4:1), dried over Na2SO4 and evaporated to dryness with N2.

The resulting core lipids were derivatized using N,O-bis(trimetylsilyl)trifluoroacetamide

(BSTFA) and pyridine (1:1) in hexane at 65°C for 20 minutes. Silylated core lipids were

analyzed for purity on a Hewlett-Packard 5972 GC/MS with a DB-5 column (length 30m; inside diameter, 0.25 mm; film thickness, 0.25 μm). The injector temperature was 320°C. The oven was held at 65°C for 1 min before increasing the temperature by 6°C/min to 320°C where it was maintained for 20 min.

3.3.4 Preparation and analysis of alkanes from DGD lipids

Phytane was prepared from extracted core diphytanyl glycerol diether (DGD) lipids of

Haloferax sulfurifontis SD1 grown on U-13C6 glucose (99%, Cambridge Isotope Laboratories,

Andover, MA). Preparation of alkanes from archaeal lipids followed a modification of the protocols described by Nisihara et al. (NISHIHARA et al., 1989) and Koga and Morii (KOGA and

MORII, 2006). Briefly, core lipids were refluxed for 2 hours at 100°C with 3 ml 55% hydroiodic acid (EM Scientific, Carson City, NV) to generate phytyl iodide. Subsequently, the alkyl iodides were refluxed for 2 hours at 120°C with 30.0 mg of zinc dust (98%, Alfa Aesar, Ward Hill, MA) and 1 ml acetic acid. This reductive dehalogenation step generated phytane that was purified by silica gel chromatography using Supelclean LC-Si SPE tubes (6 ml, Supelco, Bellefonte, PA).

Prepared phytane was analyzed for purity on a Hewlett-Packard 5972 GC/MS with a DB-

5 column (length 30m; inside diameter, 0.25 mm; film thickness, 0.25 μm). The injector temperature was 320°C. The oven was held at 65°C for 1 min before increasing the temperature by 6°C/min to 320°C where it was maintained for 20 min. The δ13C of enriched phytane was determined via continuous flow (He; 120 ml/min) on a Costech elemental analyzer (EA) by oxidation at 1020ºC over chromium (III) oxide and silvered cobalt (II, III) oxide followed by

º reduction over elemental copper at 650 C. The resulting CO2 was subsequently passed through a

º water trap and then a 5Å molecular sieve GC at 50 C to separate N2 from CO2. CO2 was diluted

with helium in a Conflo III interface/open split prior to analysis. 13C values were measured on a

Thermo Finnigan Delta Plus XP irMS. Measured 13C values were corrected for sample size dependency and then normalized to the VPDB scale with a two-point calibration (COPLEN et al.,

2006). Error was determined by analyzing independent standards across all EA runs. Accuracy is

±0.02‰ (n=54) and precision is ±0.02‰ (n=88; 1σ).

3.3.5 Monitoring growth

Growth in enrichment cultures was monitored using turbidity and changes in the concentrations of terminal electron acceptors. Nitrate, nitrite, chloride and sulfate concentrations were measured on an ICS 2500 Dionex ion chromatography system with an IonPac AS18 column with an isocratic elution program using 30 mM KOH at a flow rate of 1 ml/min and an oven temperature of 31°C. Anion concentrations were normalized to chloride. For dissolved inorganic carbon (DIC) analysis, 1 ml aliquots of culture were taken during growth and transferred to ashed 8 ml serum bottles with a 1 atm helium headspace and butyl rubber stoppers.

Subsequently, these aliquots were acidified with 2 drops phosphoric acid (12N). Concentration of DIC was determined on an SRI 310 gas chromatograph (SRI Instruments, Menlo Park, CA) with a TCD detector and a PoraPak Q packed column (3’x1/8‖) at 50°C with a helium carrier phase.

3.3.6 Stable isotope analysis of CO2

Aliquots of CO2 obtained for DIC analysis were cryogenically distilled to remove water vapor (2-propanol and dry ice) and isolate CO2 from non-condensable gases (liquid nitrogen)

13 12 prior to isotope analysis. The C/ C ratio for the CO2 was measured on a Finnigan MAT 252 dual inlet mass spectrometer.

3.3.7 Phylogenetic analysis

DNA extraction, PCR amplification and cloning were carried out exactly as described in

Macalady et al. (2008). Clones were sequenced at the Penn State University Biotechnology

Center using T3 and T7 plasmid-specific primers. CodonCode Aligner v.1.2.4 (CodonCode

Corp., Dedham, MA, USA) was used to assemble sequences and manually check for ambiguities.

Assembled gene sequences were compared against the public databases using BLAST

(ALTSCHUL et al., 1990) and chimera checked with Bellerophon 3 (HUBER et al., 2004) and

CHIMERA_CHECK v.2.7 (COLE et al., 2003). Putative chimeras were excluded from subsequent sequence analysis. Non-chimeric sequences were aligned using the NAST aligner

(DESANTIS et al., 2006), added to an existing alignment in ARB (LUDWIG et al., 2004), and manually refined.

3.3.8 Nucelotide sequence accession numbers

The 16S rRNA gene sequences determined in this study were submitted to the GenBank database under accession numbers JF834284-JF834313.

3.3.9 Fluorescence in situ hybridization

Aliquots of enrichment cultures for FISH were fixed in 3 volumes of 4% (w/v) paraformaldehyde in 1X phosphate-buffered saline (PBS) for 2-3 hours at 4°C and stored in 1:1

PBS/ethanol solution at -20°C. FISH experiments were carried out as described in Hugenholtz et

al. (2001) using probes listed in Table 3-1. Populations of Betaproteobacteria,

Gammaproteobacteria and Bacteriodales were determined from counts of between 3000-4000

DAPI-stained cells for each sample/probe combination. Oligonucleotides were synthesized and labeled at the 5’ end with the indocarbocyanine dye CY3 (Sigma-Aldrich, St. Louis, MO).

3.4 Results

3.4.1 13C-labeled phytane preparation

Growth of 28 liters of Haloferax sulfurifontis SD1 resulted in 33.4 g lyophilized biomass.

A modified Bligh-Dyer extraction and acid methanolysis of the biomass yielded 69.5 mg (0.21% dry biomass) of total lipid extract and 60.9 mg of saponified material (0.18% dry biomass). After derivatization with BSTFA, analysis by GC-MS revealed the saponified material to be archaeol

(Figure 3-2). Identification of archaeol follows MS fragmentation patterns published by Teixidor and Grimalt (1992) and Steihl et al. (2005). Major MS fragments (m/z) of the archaeol-TMS derivative include: the molecular ion (724 m/z); M – C20H41 (445 m/z); M – C20H41OH (426 m/z);

- M – C20H41OSi(CH3)3 (369 m/z); C20H41 (278 m/z); and CH2CHCH2OSi(CH3)3 (130 m/z).

After treatment of archaeol with hydroiodic acid, an extraction with benzene recovered

50.0 mg phytyl iodide. Initially, the phytyl iodide extract had a distinct purple color, but became faintly yellow following removal of residual iodide by a second extraction with benzene and 5% sodium thiosulfate (PANGANAMALA et al., 1971). Reductive dehalogenation of phytyl iodide with zinc and acetic acid produced 39.8 mg phytane. Analysis by GC-MS showed phytane to be the dominant peak, but also showed the presence of several additional peaks (Figure 3-3).

Purification with Supelclean LC-Si SPE tubes resulted in the elution of 22.2 mg phytane (0.07% dry biomass). GC-MS chromatography eluted a single peak which was identified as phytane

based upon comparison to the NIST 08 mass spectra library (Figure 3-3). The phytane purity was determined to be 98.5% based upon peak areas in the GC-MS chromatogram based upon the relative areas of these peaks. The 13C/12C ratio of the enriched phytane was ~700 ‰. After dilution with 12C-phytane (δ13C = -29.0 ‰), phytane added to incubations was ~ 325 ‰ (see equations in 3.3.2).

3.4.2 13C-labeled phytane degradation by a denitrifying enrichment

A previous enrichment of pristane-degrading denitrifying bacteria consumed approximately 70 μM nitrate/day, over the course of three months (Chapter 2). After an initial rapid loss of nitrate, incubations grown on 13C-labeled phytane showed reduced rates of denitrification, 15 μM/day. After the first fifty days of incubation, the rate of nitrate loss in the no carbon added control slowed to 2.5 μM/day. IC and DIC analysis show the production of nitrite and bicarbonate concomitant with the loss of nitrate in all incubations. Sulfate concentrations remained constant over the course of the experiment and no methane production was detected. After 120 days of growth, the consumption of 1.7 mM nitrate resulted in the production of 0.99 mM nitrite and 0.8 mM bicarbonate (2.0 mg) (Figure 3-4). Over the same time, a control experiment with no added carbon showed the consumption of 1.3 mM nitrate and the production of 0.8 mM bicarbonate.

Phytane used in enrichments had a δ13C = -29.0‰ for unlabeled phytane and δ13C ≈

13 325‰ for C-labeled phytane. After 47 days, CO2 produced during bacterial metabolism showed enrichment in the 13C-labeled phytane incubations (δ13C -11.4 ± 8.2‰) relative to the unlabeled phytane (δ13C -17.5 ± 0.7‰) and no added carbon incubations (δ13C -37.4 ± 0.8‰).

After 82 days, 13C-labeled phytane incubations showed further enrichment (δ13C 13.5 ± 7.8‰)

compared to unlabeled phytane (δ13C -35.2 ± 0.5‰) and no added carbon incubations (δ13C -

33.9 ± 1.7‰) (Figure 3-5).

DIC production in incubations with no added carbon suggests a background source of

13 carbon. A Keeling plot (PATAKI et al., 2003) was used to confirm the value of C-labeled phytane as part of a two component mixture for carbon substrate in these enrichments. In Figure

13 - 13 3-6, we plot δ CCO2 against 1/[HCO3 ], where the y-intercept is the value of C-labeled phytane

13 13 and the slope is ( Cbackground – Cphytane)*[background]. From a linear regression the y-intercept was determined to be 336.6‰, which is close to the predicted value of diluted 13C-labeled phytane (325‰).

3.4.3 Phylogenetic analysis

The most abundant cells in the incubations are short chains of rods (1.6 μm ± 0.47 μm by

0.63 μm ± 0.10 μm) occurring in large flocs (~60μm diameter). These cells may represent the bacterium responsible for phytane and pristane degradation. Oligonucleotide probes specific to the Bacteriodales family (CFB1082) (WELLER et al., 2000), and the beta and gamma subclasses of proteobacteria (Bet932, GAM42a) (MANZ et al., 1992) (Table 3-1) were used in FISH experiments to obtain quantitative information about the population structure of the pristane- degrading enrichments (Figure 3-7 and 3-8). The majority of cells (90.8 ± 14.0%) hybridize to the bacterial domain specific probe EUBmix. Betaproteobacteria, 86.8 ± 16.0% of cells, represent the dominant bacterial population. Gammaproteobacteria are present and represent 4.2

± 9.3% of bacterial cells. Cells hybridizing to the Bacteriodales specific probe are absent. A small percentage of cells (2.6 ± 6.8%) in the 13C-labeled phytane incubation hybridize to the

archaeal domain specific probe ARC915. However, no archaea were detected in the unlabeled phytane or no carbon added incubations.

We attempted to characterize the phytane degrading community on the basis of 16S rDNA clone libraries. Based upon BLAST searches and neighbor joining phylogenies constructed using the phylogenetic software ARB the most abundant clones (~70% of clones) were related to the Gammaproteobacteria, Pseudomonas stutzeri (>99% identity) previously identified as the most abundant clones in a library constructed using the same primers from the pristane degrading enrichment (Figure 3-9). An additional 25% of clones shared 97-99.8% identity with Betaproteobacteria isolates in the Comamonadaceae family that were previously identified in the Chapter 2 (Figure 3-10).

3.5 Discussion

This study demonstrates the anaerobic degradation of phytane by a denitrifying microbial community and provides a description of the phylogenies of the putative phytane-degrading microorganisms. A 16S rDNA clone library constructed from the 13C-labeled phytane incubation indicates that the most abundant clones were closely related to the Gammaproteobacteria with >

99% identity to Pseudomonas stutzeri, and closely related to the most abundant clones from a pristane degrading enrichment (Chapter 2). P. stutzeri is a known denitrifying heterotroph with the metabolic capacity to degrade organic compounds including aromatic, polycyclic aromatic and aliphatic hydrocarbons (LALUCAT et al., 2006). However, quantitative FISH analysis revealed that Gammproteobacteria (including relatives of P. stutzeri) make up only 4% of cells in the enrichments. Betaproteobacteria were the dominant phylotype active in the 13C-labeled phytane degrading incubations (86.8% of cells). Because sequence libraries are subject to DNA

extraction, amplification and cloning bias, it is unlikely that relatives of P. stutzeri are the primary phytane-degrading bacteria based upon the low representation of Gammaproteobacteria in the quantitative FISH analysis. The observed decrease in Gammaproteobacteria as compared to the pristane degrading enrichment in Chapter 2 may be due to preservation bias during storage in glycerol and subsequent transfer of cultures.

Additional clones were related to polyhydroxyalkanoate (PHA) degrading Simplicispira isolates within the Betaproteobacteria family Comamonadaceae (GRABOVICH et al., 2006; KHAN et al., 2002). In Chapter 2 we suggested that PHA degrading Betaproteobacteria may have syntrophic interactions with P. stutzeri, which can produce PHA storage granules (GUO-QIANG et al., 2001). Members of the Simplicispira genus have been isolated from wastewater

(GRABOVICH et al., 2006; LU et al., 2007), and identified in denitrifying, chlorobenzene degrading enrichments (NESTLER et al., 2007) and in a nitrate treated oil field (CORNISH

SHARTAU et al.). Based upon FISH counts, as well as their persistence in multiple 16S rDNA clone libraries constructed over the course of enrichment culturing, we propose that the

Simplicispira relatives are responsible for phytane and pristane degradation in these enrichments.

Incubations grown with a 13C-labeled phytane tracer produced bicarbonate as a byproduct of phytane metabolism that was enriched in 13C as compared to incubations grown with unlabeled phytane or no added carbon substrate. Based upon purity (98.5%), approximately 0.33 mg of the prepared 13C-labeled phytane may represent contaminants from the preparation process.

Based upon the purity (99%) of unlabeled phytane an additional 0.28 mg of unlabeled impurity may be present. After addition to incubation bottles, approximately 0.042mg of labeled impurity and 0.035 mg unlabeled impurity may be present in each bottle. The production of 2.0 mg bicarbonate (0.40 mg C) accounts for 5 times the amount of impurity and 10 times the amount of

labeled impurity present in any individual bottle. Thus, we believe that any enrichment observed in the bicarbonate is primarily due to utilization of 13C-labeled phytane and not due to any residual 13C-labeled compounds from the preparation process.

The stoichiometry of phytane degradation can be hypothesized from the quantitative growth experiment. During the reduction of approximately 70 μmol nitrate to 40 μmol nitrite, 34

- μmol HCO3 was produced. Assuming a range for carbon assimilation from 20% to 50% (WHITE,

2007) and a cellular composition of C5H7O2N (DRTIL et al., 1995), this represents the consumption of between 2.1 and 3.3 μmol of phytane (7.5 – 11.8% of the initial phytane). The combined assimilatory and dissimilatory reactions follow.

For 20% assimilation:

- + - - C20H42 + 32NO3 + 0.8NH3 + 6H  16HCO3 + 0.8C5H7O2N + 16NO2 + 8N2 + 14.4H2O

For 50% assimilation:

- - - + C20H42 + 20NO3 + 2NH3  10HCO3 + 2C5H7O2N + 10NO2 + 5N2 + 6H2O + 12H

The accumulation of nitrite in the phytane degrading cultures points to incomplete denitrification of nitrate and nitrite to gaseous nitrogen products. Accumulation of nitrite may be due to the inhibitory effect of nitrate on nitrite reduction (KORNAROS et al., 1996). Additionally, nitrite accumulation and incomplete nitrate loss has been demonstrated during carbon limited

. -1 growth (KORNAROS et al., 1996). Phytane solubility is estimated as 0.3 mg L at 25°C(EASTCOTT et al., 1988). The low solubility of phytane in aqueous solutions likely results in carbon limited conditions in these incubations. Alternatively, dissimilatory nitrate reduction to ammonium

(DNRA) remains a possible alternative metabolism.

Under the conditions of this denitrifying enrichment, phytane biodegradation follows first order kinetics with k = 0.20 y-1 (r2 = 0.71, n=15) to 0.33 y-1 (r2 = 0.71, n = 15). This degradation

rate is less than the rate calculated in Chapter 2 for pristane utilization by this enrichment (k =

0.95y-1 – 1.57 y-1). Both pristane and phytane degradation rates for this enrichment are slower than isoprenoid degradation rates reported by Grossi et al. (2000) for pristane (k = 3.5y-1), squalene (k = 18.5 y-1) and phytadienes (k = 6.6 y-1) by a marine, sulfate reducing enrichment.

Rate constants reported for isoprenoid degradation by both denitrifying and sulfate reducing enrichments are several orders of magnitude greater than the in situ anaerobic biodegradation rate constant for mixed hydrocarbons in oil fields (10-8 y-1) where oxidant and nutrient supplies are limited by diffusion, and an order of magnitude greater than anaerobic biodegradation rates

-1 -2 -1 in anoxic surface sediment (10 – 10 y ) (LARTER et al., 2003).

Isoprenoid hydrocarbons represent between 25-30% by mass of crude oil deposits

(PETROV and ABRYUTINA, 1989; TISSOT and WELTE, 1984). Based upon the degradation rates described above, bacteria targeting solely isoprenoids would degrade this fraction in 20-25 million years in an oil field (LARTER et al., 2003). By contrast, in anoxic surface sediment or in this incubation, a similar fraction would be degraded in 2.5-30 years. The increased availability of nutrients and electron acceptors in laboratory experiments and surface sediment is consistent with the difference in rates and time scales.

3.6 Conclusions

Anaerobic microbial isoprenoid degradation was demonstrated in denitrifying incubations grown with 13C-labeled phytane. Nitrate loss and the production of nitrite and bicarbonate were measured over 120 days. Isotope ratio measurements of bicarbonate indicated 47.4‰ enrichment in incubations grown on 13C-labeled phytane as compared to incubations with no added carbon.

The microbial consortia responsible for degradation includes Betaproteobacteria related to

Simplicispira spp. and Gammaproteobacteria related to Pseudomonas stutzeri. Close relatives of both phylotypes are known denitrifying bacteria associated with hydrocarbon degradation. The first order rate constants for phytane degradation were 0.2-0.33 y-1, within the range anticipated for hydrocarbons in anoxic surface sediments, but much higher than rates estimated for oil reservoirs.

3.7 Acknowledgements

This study was supported by a grant from the American Chemical Society Petroleum Research

Fund (grant 48445-AC2). We would like to thank D. Walizer, I. Schaperdoth and H. Graham for technical assistance.

3.8 Figures and Tables

Figure 3-1: Preparation scheme for 13C-labeled phytane.

Figure 3-2: Total ion chromatogram and mass spectrum of saponified 13C-labeled diphytanyl glycerol diether from Haloferax sulfurifontis SD1.

Figure 3-3: Total ion chromatogram and mass spectra of 13C-labeled phytane before and after

SPE purification.

Figure 3-4: Loss of nitrate and concomitant production of nitrite and bicarbonate in incubations grown with 13C-labeled phytane as a carbon substrate. Data points represent the average of three replicate incubations and error bars are ± 1 standard deviation.

13 13 Figure 3-5: Values of δ C for the CO2 produced during incubations on C-labeled phytane

(black diamonds, solid line), unlabeled phytane (black squares, long dashes) and with no added carbon (black triangles, short-dashes). A line is provided at δ13C = -29.0 ‰ to show the value of unlabeled phytane.

13 13 Figure 3-6: Keeling plot of δ C for CO2 produced during incubations on C-labeled phytane

- versus 1/[HCO3 ].

Figure 3-7: Percentage of cells hybridizing to specific oligonucleotide probes. ARC915 – most

Archaea; GAM42a – Gammaproteobacteria; BET932 – Betaproteobacteria.

Figure 3-8: FISH micrographs of cells in the denitrifying, phytane-degrading enrichment culture.

Bar, 10 μm. All cells have been stained with DAPI and hybridized with Bacteria specific probe

EUB338 (green). Top row: Cells hybridized with Betaproteobacterial specific oligonucleotide probe BET932 (red). Middle row: Cells hybridized with Gammaproteobacterial specific oligonucleotide probe GAM42a (red). Bottom row: Cells hybridized with Bacteriodales specific probe CFB1082 (red).

Figure 3-9: Neighbor-joining phylogenetic tree showing Gammaproteobacteria. 13C-labeled phytane and pristane degrading enrichment clones are shown in bold followed by the number of clones of each phylotype. Maximum parsimony (MP) and neighbor-joining (NJ) bootstrap values >50% are shown (MP/NJ). Putative pristane-metabolizing cells are annotated by ―‖.

Figure 3-10: Neighbor-joining phylogenetic tree showing Betaproteobacteria. 13C-labeled phytane and pristane degrading enrichment clones are shown in bold followed by the number of clones of each phylotype. Maximum parsimony (MP) and neighbor-joining (NJ) bootstrap values >50% are shown (MP/NJ). Putative pristane-metabolizing cells are annotated by ―‖.

Table 3-1: Oligonucleotide probes used in this study

Probe Target group Sequence (5’ 3’) Formamide (%) Target site Reference

EUB338a Most Bacteria GCTGCCTCCCGTAGGAGT 0 – 50 16S (338-355) Amann et al. (1995)

EUB338-IIa Planctomycetales GCAGCCACCCGTAGGTGT 0 – 50 16S (338-355) Daims et al. (1999)

EUB338-IIIa Verrucomicrobiales GCTGCCACCCGTAGGTGT 0 – 50 16S (338-355) Daims et al. (1999)

GAM42a Gammaproteobacteria GCCTTCCCACATCGTTT 35 23S (1027-1043) Manz et al. (1992)

ARC915 Archaea GTGCTCCCCCGCCAATTCCT 20 16S (915-935) Stahl and Amann (1991) cGAM42a Competitor GCCTTCCCACTTCGTTT 35 23S (1027-1043) Manz et al (1992)

BET932 Betaproteobacteria ATCATCCACCGCTTGTGC 45 16S (932-950)

CFB1082 Bacteriodales TGGCACTTAAGCCGACAC 30 16S (1082-1100) Weller et al. (2000) aCombined in equimolar amounts to make EUBMIX.

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82 Chapter 4: Quantitative FISH analysis of microbial consortia from a biogenic gas field in the Cook Inlet Basin, Alaska

4.1 Abstract

Production water samples from a methane-rich gas field in the Cook Inlet of Alaska host microbial communities consistent with microbial methane production from coal. A suite of fluorescence in situ hybridization (FISH) probes, archaeal and bacterial, were designed to target groups identified by 16S rRNA tag pyrosequencing of filter-collected production water communities. Based on FISH experiments, the archaeal community is dominated by the obligate methylotrophic, methanogen genus Methanolobus, as well as the CO2-reducing and acetoclastic methanogen genus Methanosarcina. The bacterial community is comprised of Acetobacterium,

Bacteriodales, and Firmicutes. We observed spatial variation among the samples in the microbial community composition in both the percentage of archaea versus bacteria and the dominant member of the bacterial community. A simplified reaction network beginning with the breakdown of macromolecules, followed by fermentation and methylotrophic and acetoclastic methane production is proposed based upon the microbial community composition and the isotopic signature of methane associated with this Cook Inlet Basin site.

4.2 Introduction

Coal bed methane (CBM) refers to natural gas associated with coal that may be formed by the thermogenic cracking of hydrocarbons or biogenically through the degradation of complex organic matter. This gas may be retained by adsorption to the coal, by storage in associated porous layers and coal fractures, and by dissolution in water (NEDERLOF, 1988). Gas

83 composition, reservoir geochemistry and stable isotopes can be used to discriminate between biogenic and thermogenic sources (FLORES et al., 2008; WHITICAR, 1999). The stable isotope signature of methane can additionally distinguish between the two main pathways of biogenic methane formation, carbonate reduction and methyl group fermentation (WHITICAR, 1986;

WHITICAR, 1999). CBM represents a significant and growing unconventional source of natural gas (COMMITTEE ON MANAGEMENT AND EFFECTS OF COALBED METHANE DEVELOPMENT AND

PRODUCED WATER IN THE WESTERN UNITED STATES et al., 2010). It is important to understand the microbial processes that result in coal conversion to methane, especially those involving in situ microbial communities.

Coal presents a challenge to microbial degradation due to its complex, heterogeneous structure composed of aromatic, polycyclic aromatic and aliphatic moieties (FAKOUSSA and

HOFRICHTER, 1999). Biologic conversion of coal to methane requires a consortium of microorganisms that are capable of depolymerizing coal and fermenting monomers to lower molecular weight compounds such as acetate, H2, CO2, methyl amines and methyl sulfides that can be utilized by methanogenic archaea (CATCHESIDE and RALPH, 1999; FAKOUSSA and

HOFRICHTER, 1999; HARRIS et al., 2008; OREM et al., 2010; STRAPOC et al., 2008). The specific pathway for coal degradation depends upon the microbial community present. Coal beds show a high diversity of associated bacteria, including significant representation by Firmicutes,

Spirochetes, Bacteroidetes, and all subgroups of Proteobacteria (STRAPOC, 2011) (Figures 1-6 to

1-8). Methanogenic archaea identified in coal beds include species of Methanosarcinales,

Methanomicrobiales and Methanobacteriales representing all of the known methanogenic pathways (STRAPOC, 2011) (Figure 1-9).

84 Recent studies have characterized the indigenous coal bed microbial communities using both culturing and 16S rDNA cloning (FRY et al., 2009; GREEN et al., 2008; KRUGER et al., 2008;

LI et al., 2008; MIDGLEY et al., 2010; SHIMIZU et al., 2007; STRAPOC et al., 2008). These studies provide a glimpse of the natural microbial assemblages associated with coal beds, but may not reflect the relative population sizes of the active bacterial and archaeal populations in situ.

Further study is necessary to understand the pathways for the breakdown of the complex coal matrix to precursors for methanogenesis. Here, we combine 16S rRNA tag pyrosequencing with fluorescent in situ hybridization (FISH) to generate quantitative microbial population data and to visualize associations of bacteria and archaea present in coal bed communities in a biogenic gas field in the Cook Inlet, Alaska.

4.3 Materials and Methods

4.3.1 Sampling site and geochemical analysis

Water samples were obtained from production wells of a biogenic gas field in the northern portion of the Cook Inlet Basin, Alaska. The major natural gas reserves consist of 1.8 x

1011 m3 of shallow, dry gas (99% methane) occurring in sandstones interbedded with coal from

13 Miocene to Pliocene aged rocks in the Kenai group. The δ Cmethane ranges from -63 to -56‰ and is interpreted to have a biogenic origin (CLAYPOOL et al., 1980). Production water samples were analyzed for the concentrations of seven major ions at the Bartlesville Technology Center,

ConocoPhillips. Isotopic analysis of gas samples was performed at Isotech, IL. Gas compositional analysis was performed on a Carle AGC400/AGC100. Values for methane δD were measured on an HP6890 coupled with a Delta V Plus GC-P-III interface. Values for

85 methane δ13C were measured on an HP6809 coupled with a Delta Plus Advantage XL via a GC-

C-II interface.

4.3.2 DNA extraction and sequencing

Production water samples (250-500 ml) were filtered through a 47 mm (0.2-m pore size)

Durapore membrane filter (Millipore, Billerica, MA). Using a sterile scalpel, filters were sliced into 96 equal sized portions and transferred equally into two 2.0 ml screw cap centrifuge tubes containing ceramic beads obtained from CeroGlass (Columbia, TN). The bead-beating matrix consisted of one 4-mm glass bead (GSM-40), 0.75 g 1.4- to 1.6-mm zirconium silicate beads

(SLZ-15), and 1.0 g 0.07- to 0.125-mm zirconium silicate beads (BSLZ-1) in 1 ml phosphate buffer (180 mM sodium phosphate, 18 mM EDTA, pH 8.0). Cells were disrupted in a Fastprep

FP120 instrument as previously described (ASHBY et al., 2007). Total genomic DNA was purified by centrifuging the lysed cells at 13,200 x g for 5 min at 4oC. The supernatants were transferred to 1.5 ml centrifuge tubes and 250 l of 2M potassium acetate pH 5.3 was added. The tubes were mixed by rotating end-over-end and were centrifuged as above. The resulting genomic DNA was purified on QIAprep Plasmid Spin columns (Qiagen, Valencia, CA) according to the manufacturer’s instructions.

A portion of the 16S rRNA gene was amplified using the TX9/1391 primers as previously described (ASHBY et al., 2007). Amplicons were agarose gel purified and quantitated using SYBR green (Invitrogen, Carlsbad, CA). A second round of PCR was performed using fusion primers that incorporated the ‘A’ and ‘B’ 454 pyrosequencing adapters onto the 5’ ends of the TX9/1391 primers, respectively. The forward fusion primer also included variable length barcodes that enabled multiplexing multiple samples into a single 454 sequencing run. These

86 amplicons were PAGE purified and quantitated prior to combining into one composite library.

The resulting library was sequenced using the standard 454 Life Sciences Lib-L emulsion PCR protocol and Titanium chemistry sequencing (MARGULIES et al., 2005). Sequences that passed the instrument QC filters were also subjected to additional filters that required all bases be Q20 or higher and the average of all bases in any read to be Q25 or greater. Furthermore, the TX9 primer was trimmed off of the 5’ end and the sequences were trimmed on the 3’ end at a conserved site distal to the V6 region (ca position 1067, E. coli numbering). The final sequences were approximately 250 bp in length and included the V5 and V6 regions.

4.3.3 Probe design and FISH

Production water samples (250-500 ml) for FISH were collected using 47 mm (pore size

0.2 μm) black nucleopore polycarbonate filters (Whatman, UK) using an in-line filter holder and a luer-lok syringe. Filters were fixed for FISH in 1% (w/v) paraformaldehyde, stored on ice and processed within 48 hours of collection. The filters were washed with 1X phosphate-buffered saline (PBS) and stored in 1:1 PBS/ethanol solution at -20°C. FISH experiments were carried out as described in Macalady et al. (MACALADY et al., 2006). The oligonucleotide probes and formamide concentrations used are given in Table 1. Probes were designed using an ARB database (LUDWIG et al., 2004) as described by Hugenholtz et al. (HUGENHOLTZ et al., 2002) and synthesized and labeled at the 5’ end with fluorescent dyes (CY3, CY5 and FLC) at Sigma-

Aldrich. Microbial populations were determined from counting a minimum of 1000 DAPI- stained cells for each sample/probe combination. Standard deviation was determined among separate slides (n ≥ 3 slides).

87 4.3.4 Phylogenetic analysis

The 16S rRNA sequences of bacteria and archaea used in this study were previously submitted to GenBank by authors of studies describing coal bed methane microbial communities

(GREEN et al., 2008; KRUGER et al., 2008; PENNER et al., 2010; SHIMIZU et al., 2007; STRAPOC et al., 2008). Sequences were aligned using the NAST aligner at greengenes (DESANTIS et al.,

2006). NAST aligned sequences were imported into an ARB database (LUDWIG et al., 2004) containing more than 200,000 full-length sequences. Phylogenetic analyses were performed on sequences exported from ARB with distance and maximum parsimony methods using version

4.0b10 of PAUP* (SWOFFORD, 2000). Neighbor-joining trees (neighbor-joining search, jukes- cantor distance, 2000 bootstrap replicates) were compared against maximum parsimony consensus trees (heuristic search, 2000 bootstrap replicates) (Figures 1-6 to 1-9).

4.3.5 Multivariate Statistics

Principal components analysis (PCA) was performed on microbial population data for three Cook Inlet Basin samples, as well as microbial population data from four previously described basins (GREEN et al., 2008; KRUGER et al., 2008; MIDGLEY et al., 2010; SHIMIZU et al.,

2007) using the R-statistical software package (R DEVELOPMENT CORE TEAM, 2007).

Geochemical data were projected onto the ordination using the ‘envfit’ function (1000 permutations). All data were normalized by the parameter maximum prior to ordination in order to eliminate scaling artifacts.

4.4 Results

4.4.1 Microbial populations

88 A total of twelve FISH probes (Table 1) were used to analyze the microbial community in the Cook Inlet Basin samples CIB-2, CIB-1 and CIB-3. Representative epifluorescence micrographs are shown in Figure 4-1. The ratios of archaea to bacteria, as well as the microbial community composition vary between the three samples (Figure 4-2). CIB-1 is dominated by archaea with 56 ± 10.6 % of cells hybridizing to the ARC915 and 40.3 ± 8.0% of cells hybridizing to EUBmix. CIB-2 displayed a more even distribution of archaea and bacteria with

43.4 ± 7.1% of cells hybridizing to ARC915 and 56.5 ± 9.8% of cells hybridizing to EUBmix.

CIB-3 is a bacterial dominated site with 34.3 ± 14.3 % hybridizing to ARC915 and 64.8 ± 4.9 % of cells hybridizing to EUBmix.

Additional analysis of the archaeal and bacterial populations focused on target groups identified by 16S rRNA tag pyrosequencing. In all three sites, a suite of probes for methanogenic archaea reveals that the genera Methanolobus and Methanosarcina are the most numerous archaea with Methanolobus cells outnumbering Methanosarcina. No cells are observed to hybridize to probes specific for Methanobacteria or Methanomicrobiales. Bacterial communities in the three samples are more variable than archaeal communities. CIB-1 consists of SRB385 hybridizing cells (10.8%), Bacteriodales (8.2%) and Acetobacterium (6.0%). CIB-2 consists of

Bacteriodales (32.9%), Acetobacterium (18.5%) and SRB385 hybridizing cells (11.7%). CIB-3 consists of Acetobacterium (28.0%), Bacteriodales (9.6%) and SRB385 hybridizing cells (6.1%).

Cells hybridizing to the probe SRB385, which targets some Deltaproteobacteria as well as some

Firmicutes, are attributed to Firmicutes rather than Deltaproteobacteria due to the lack of hybridization to Delta495a. Despite evidence for Chloroflexi and Clostridiaceae in the pyrosequencing data, the probes targeting these groups utilized in this study, Chis150 and

CFX1223, do not hybridize to cells in any of the samples. Additionally, ~15% of the CIB-1 and

89 ~20% of the CIB-3 bacterial community do not hybridize to the any of the probes used in this study.

FISH and 16S rRNA tag pyrosequencing, compared in Figure 4-2, show similar results in the phylum level as well as the domain level community composition. 16S rRNA tag pyrosequencing show CIB-1 to be dominated by archaeal sequences (81.0%), while sequences from CIB-2 were more evenly distributed between archaea (44.1%) and bacteria (55.9%) and the

CIB-3 community is dominated by bacterial sequences (86.4%). At the phylum level the archaeal communities of all three samples are primarily composed of Methanomicrobia, which includes the genera Methanosarcina and Methanolobus. The most abundant bacterial phyla are

Bacteroides and Firmicutes (7.1% and 6.3% respectively in CIB-1; 5.6% and 41.1% in CIB-2;

5.7% and 66.8% in CIB-3). The bacterial communities also include several phyla representing less than 2% of all sequences that were not detected by FISH.

Despite intense disruption during sampling and sample preparation, a significant fraction of the active microbial cells are imaged in close spatial associations that have distinctive taxonomic structures. Epifluorescence photomicrographs of CIB-1 reveal a close structural association of archaeal and bacterial cells (Figure 4-1). Short filamentous and rod-shaped bacterial cells intertwine with Methanosarcina and Methanolobus cocci forming large, intact clusters of cells. Based on FISH evidence the rod-shaped cells are a mixture of Bacteriodales and

Acetobacterium, while the filamentous cells are a mixture of SRB385 hybridizing cells and another unidentified lineage. Similar associations of archaeal and bacterial cells are not observed in CIB-2 and CIB-3.

4.4.2 Geochemical analysis

90 Isotopic data are plotted in the manner of Whiticar (WHITICAR, 1986). The carbon and hydrogen isotope values place the methane from these three samples in a zone of biogenic methanogenesis characterized by methyl type fermentation (Figure 4-4).

4.4.3 Multivariate analysis

A principal components analysis (PCA) of Cook Inlet Basin microbial population data is shown in Figure 4-5. Principal component 1 (PC1), which explains 71.1% of the variation in the dataset, is strongly correlated with the ratio of archaea to bacteria and population sizes of

Acetobacterium, Methanolobus, and Methanosarcina. PC2 (28.9%) is strongly correlated with

Bacteroidales population sizes. Geochemical data (blue vectors) are projected onto the PCA using the ‘envfit’ function. P-values (<0.001) show significant correlations between the ordination and pH, acetate, propionate and TDS concentrations, but these parameters are only weakly related to PC1. Although there are positive correlations between calcium and sodium and bacterial populations, and between iron concentration and archaeal populations, the p-values relating these geochemical variables to the PCA are > 0.3, thus not significant.

A principal components analysis performed using population data from Cook Inlet,

Gippsland, Ishikari (Yubari), Powder River and Ruhr Basins is shown in Figure 4-6. The addition of other basins results in the Cook Inlet samples clustering together. Of the additional sites, the Ishikari (Yubari) and Ruhr basin sites are the nearest in ordination space. This grouping is related to the presence of the Methanolobus and Acetobacterium populations in these sites and their absence in the others. PC1, which explains 46.1% of the observed variance, is most strongly correlated with the population sizes of Methanolobus and Methanobacteria. PC2 (34.3%) is most strongly correlated with the population sizes of Methanosarcina and Clostridia. Geochemical

91 data (blue vectors) are projected onto the PCA using the ‘envfit’ function. P-values for the environmental variables (>0.3) show no significant correlations between the available geochemical data and the ordination of microbial population data.

4.5 Discussion

Multivariate analysis of the communities at the three Cook Inlet Basin sites and the multiple basin dataset failed to identify significant correlations between microbial community structure and geochemical data. While not statistically significant, sodium, calcium and iron concentrations correlate with the ratio of bacteria to archaea in the Cook Inlet Basin samples.

Although not within the original scope of this study, a more complete set of geochemical data including variables such as redox potential and nutrient and trace metal concentrations is likely to be a sound investment in future studies where the goal is to understand geochemical controls on microbial community structure and methane production potential.

Isotopic data show the dominant methanogenic pathway at the Cook Inlet Basin study site involves the fermentation of acetate or other methyl group substrates. Carbon and hydrogen isotope values place the methane from the three analyzed wells into the range associated with methyl-type fermentation. Additionally, cells hybridizing to Methanosarcina and Methanolobus genera specific probes dominate the observed archaea. While Methanosarcina species are capable of methanogenesis by acetoclastic, methylotropic and CO2 reduction pathways,

Methanolobus species are obligate methylotrophs (DWORKIN et al., 2006). A major component of the bacterial community is the homoacetogen genus Acetobacterium. Homoacetogens produce acetate from the reduction of carbon dioxide with hydrogen. Competition between

Acetobacterium and CO2 reducing methanogens (Methanobacteria and Methanomicrobiales) for

92 CO2 and H2 as substrates may explain the predominance of an acetoclastic or methylotrophic pathway over a carbonate reduction pathway (KOTSYURBENKO et al., 2001).

The close structural association of methanogens and bacteria in CIB-27 is suggestive of syntrophic interactions. The association of methanogens with acetogens (KOTELNIKOVA and

PEDERSEN, 1998; STRUCHTEMEYER et al., 2005) or with sulfate-reducing bacteria (MOSER et al.,

2005) has previously been reported in methanogenic communities. A study by Shimizu et al.

(2007) revealed clones of Methanolobus, Acetobacterium and Syntrophus in a deep coal seam

(SHIMIZU et al., 2007). In CIB-27, a bacterial community composed of Acetobacterium,

Bacteriodales, and SRB385 hybridizing cells cluster with an archaeal community composed of

Methanolobus and Methanosarcina. The microbial consortia within these associations include constituents capable of macromolecule fermentation (Bacteriodales), degradation of fermentation intermediates (Bacteriodales), acetate generation (Acetobacterium) and methanogenesis (Methanosarcina and Methanolobus).

Through the combination of the 16S rRNA tag pyrosequencing data and the quantitative

FISH data we can begin to develop a simplified mechanism for the breakdown of organic matter in coal at the study site. Species of Bacteriodales are capable of the anaerobic degradation of cellulose, polysaccharides and other complex substrates to simple sugars and other fermentation products (XU et al., 2003). Acetobacterium produce acetate from the fermentation of simple sugars, as well as through homoacetogenesis with H2 and CO2 (BUSCHHORN et al., 1989).

Acetoclastic and methylotrophic methanogenesis by Methanosarcina and Methanolobus finally leads to the production of methane. This coal biodegradation scheme retains the general mechanism of macromolecular breakdown followed by fermentation to simple sugars and methanogenesis precursors proposed by Strapoc et al. (2008) for the Illinois Basin. However, the

93 microbial communities and methanogenic pathway differ in the presence of acetogenic bacteria and acetoclastic and methylotrophic archaea.

Although the community composition and the likely pathway for organic matter degradation for all three Cook Inlet Basin sites are similar, we noted significant variations in the proportions of microbial groups present in spatially separated sites within this single production field. Microenvironments favoring particular microbes may result from heterogeneity in the coal structure, hydrology, geochemistry and trace element concentrations. For example, variations in coal structure likely influence the bacterial community. A higher degree of aromatization and condensation could provide a competitive advantage to bacterial hydrocarbon degraders capable of fragmenting and fermenting polycyclic aromatic hydrocarbons over species capable of fermenting cycloalkyl, carboxyl and methoxy groups. Additionally, more favorable conditions for methanogenic archaea could be provided by higher concentrations of trace elements such as nickel, which is required for coenzyme F430, a component of the key enzyme involved in methanogenesis. Future efforts to stimulate in situ biogenic CBM production will need to consider differences in community composition as well as geochemistry among fields as well as at finer scales.

4.6 Conclusions

Quantitative FISH analyses of a biogenic gas field in the Cook Inlet Basin, Alaska revealed that the archaeal community is dominated by the obligate methylotrophic, methanogen genus Methanolobus, as well as the CO2-reducing and acetoclastic methanogen genus

Methanosarcina. The bacterial community is comprised of Acetobacterium, Bacteriodales, and

Firmicutes. The microbial community composition in combination with methane isotope data

94 indicates that acetate and methyl-group fermentation are the dominant methanogenic pathways in the Cook Inlet Basin. Variation in the microbial community composition within this single production field suggests the presence of microenvironmental compositions favoring particular microbes. Further study of fine scale heterogeneity in the geochemistry, hydrology and coal structure is likely to indicate factors that promote particular microbial populations.

95 4.7 Figures and Tables

Table 4-1: Oligonucleotide probes used in this study for Fluorescent in situ hybridzation

Target group Probe Sequence 5’ 3’ Formamide (%) Reference

a Most Bacteria EUB338 GCT GCC TCC CGT AGG AGT 0-50 (AMANN et al., 1995)

a Planctomycetales EUB338-II GCA GCC ACC CGT AGG TGT 0-50 (DAIMS et al., 1999)

a Verrucomicrobiales EUB338-III CT GCC ACC CGT AGG TGT 0-50 (DAIMS et al., 1999)

(STAHL and AMANN, Archaea Arc915 GTG CTC CCC CGC CAA TTC CT 20 1991)

Methanosarcinales Sarci551 GAC CCA ATA ATC ACG ATC AC 20 (SORENSEN et al., 1997)

Methanolobus Mlob828 CGC ACC RTC CCA GAC ACC 20 This study

Methanobacteria MB311 ACC TTG TCT CAG GTT CCA TCT CC 30 (CROCETTI et al., 2006)

Methanomicrobiales MG1200b ACC TTG TCT CAG GTT CCA TCT CC 30 (CROCETTI et al., 2006)

Clostridiaceae Chis150 TTA TGC GGT ATT AAT CTY CCT TT 30 (FRANKS et al., 1998)

Acetobacterium Aceto828 CTG AGT CTC CCC AAC ACC 30 This study

Chloroflexi CFX1223 CCA TTG TAG CGT GTG TGT MG 30 (BJORNSSON et al., 2002)

Bacteriodales CFB1082 TGG CAC TTA AGC CGA CAC 30 (WELLER et al., 2000)

Deltaproteobacteria, SRB385 CGG CGT CGC TGC GTC AGG 35 (AMANN et al., 1992) some Firmicutes

Deltaproteobacteria Delta495a AGT TAG CCG GTG CTT CCT 45 (LOY et al., 2002) a Combined in equimolar amounts to make EUBmix.

Figure 4-1: FISH micrographs depicting major bacterial and archaeal lineages observed in production water samples from the Cook Inlet Basin. Probe specificities: EUBmix - all bacteria,

Arc915 -most archaea, Mlob828 - genus Methanolobus, SRB385 - some Firmicutes, SARCI551

– genus Methanosarcina, Aceto828 – genus Acetobacterium.

Figure 4-2: Microbial communities of the Cook Inlet gas field expressed as a percentage of

DAPI stained cells. Bacteria are shown in shades of green and archaea are shown in shades of orange. The values represent the percentage determined from counting a minimum of 1000 DAPI stained cells. Standard deviation, 1.5-16%, is determined among separate slides (n ≥ 3 slides).

Figure 4-3: Comparison of the microbial communities of the Cook Inlet gas field determined by

FISH and 16S rRNA tag pyrosequencing. Shades of orange indicate archaeal lineages and other colors indicate bacterial lineages determined by both techniques.

Figure 4-4: Carbon and hydrogen isotopic classification of methane showing the difference between methanogenesis by thermogenic production, microbial carbonate reduction and microbial methyl-type fermentation. The Cook Inlet wells, shown in red, indicate methane generated by methyl-type fermentation (WHITICAR, 1999).

100

Figure 4-5: PCA of Cook Inlet basin microbial population data (red) with geochemical data (blue) projected onto the ordination using the ‘envfit’ function. All data were normalized by the parameter maximum prior to ordination in order to eliminate scaling artifacts.

101

Figure 4-6: PCA of microbial population data from five sedimentary basins (red) with geochemical data (blue) projected onto the ordination using the ‘envfit’ function. All data were normalized by the parameter maximum prior to ordination in order to eliminate scaling artifacts.

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106 Chapter 5: Molecular characterization of archaeal lipids across a hypersaline gradient

5.1 Abstract

Four halophilic archaeal strains, Halorhabdus utahensis, Natronomonas pharaonis,

Haloferax sulfurifontis and Halobaculum gomorrense, were grown at a range of salinities (10-

30% NaCl, (w/v)). These strains represent four archaeal genera and have a range of salinity optima. Molecular analysis of the DGD structure by GC-MS showed the presence of core lipid structures consistent with unsaturated phytanyl and sesterpanyl hydrocarbon chains. In addition to the presence of unsaturation in the isoprenoid chain, we observed three trends. (1) The percentage of unsaturated DGDs increased with increasing concentration of NaCl in the growth medium. (2) Halophilic archaeal strains with a higher optimal NaCl concentration have a higher percentage of unsaturated DGDs. (3) The presence of C25-20 DGDs occurred in the strains with higher optimal NaCl concentrations, N. pharaonsis and H. utahensis.

5.2 Introduction

Hypersaline water bodies including salterns, soda lakes and the Dead Sea are characterized by NaCl concentrations ranging from 1M to saturation. These environments may also have an alkaline pH, high light exposure (due to shallow water depths) and high concentrations of other ions such as Mg2+. Despite the inhospitable chemical conditions, these ecosystems are populated by diverse communities of halophilic algae, bacteria and archaea

(ELAZARI-VOLCANI, 1943; KAMEKURA, 1998; LEY et al., 2006; OREN, 1999b; WILKANSKY,

1936). Two physiological strategies exist for coping with the osmotic stress imposed by high salinity. Eukaryotes and most halophilic bacteria accumulate high intracellular concentrations of

107 organic solutes to balance the osmotic stress (OREN, 1999a). In constrast, halophilic archaea

+ - + accumulate high intracellular concentrations of K Cl , while excluding Na (OREN, 1999a; VAN

DE VOSSENBERG et al., 1998). In addition, the dialkyl glycerol diether (DGD) lipid structure of halophilic archaea confers enhanced membrane stability and reduced membrane permeability to protons, sodium ions and other solutes (CHOQUET et al., 1992; CHOQUET et al., 1994; ELFERINK et al., 1994; TENCHOV et al., 2006; VAN DE VOSSENBERG et al., 1998).

The general DGD structure (Figure 1) has been identified with modifications including isoprenoid chain lengths between C15-C25, the inclusions of cyclic groups, a macrocyclic structure, hydroxyl substitutions, and varying degrees of unsaturation in the phytanyl chain

(COMITA et al., 1984; GIBSON et al., 2005; KATES, 1977; STADNITSKAIA et al., 2003; STIEHL et al., 2005). Polar and glycolipid composition shows some relationship to taxonomy (ASKER and

OHTA, 2002; ELSHAHED et al., 2004; LATTANZIO et al., 2002; LIZAMA et al., 2002; OREN, 2002;

OREN et al., 2009). However, the factors controlling variation in DGD core lipid composition are less well understood. These variations along with isotopic compositions have the potential to provide information about ancient microbial communities and paleoenvironments.

In previous work, monounsaturated archaeol was identified as an analytical artifact of hydroxyarchaeol extraction (EKIEL and SPROTT, 1992). However, in other studies, isotopic differences between hydroxyarchaeol and the monounsaturated form (BLUMENBERG et al., 2005), as well as the presence of multiply unsaturated archaeol in the isoprenoid chain of

Halobacterium lacusprofundi and Methanococcoides burtonii (FRANZMANN et al., 1988; GIBSON et al., 2005; NICHOLS et al., 2004) cannot be explained by hydroxyl group loss. This strongly suggests that unsaturated forms are not always an artifact of extraction. Adaptation to the cold environment in Antarctic lakes may explain the observed polyunsaturated core lipid structures in

108 H. lacusprofundi and M. burtonii. However, an alternative explanation is needed for the presence of highly unsaturated DGD lipids in isolates from warmer environments (GIBSON et al., 2005;

QIU et al., 1998; STIEHL et al., 2005; UPASANI et al., 1994). We hypothesized that the presence of unsaturated DGDs is a physiological response to the osmotic stress of hypersaline environments.

In order to test this hypothesis, we examined the impact of a salinity gradient on the degree and relative percentage of unsaturated DGDs in the polar and total lipid extracts of four species of halophilic archaea.

5.3 Methods

5.3.1 Microorganisms and culture conditions

Haloferax sulfurifontis strain SD1 (Lee Krumholz, University of Oklahoma) was grown in the liquid medium described by Elshahed et al. (2004), which contained per liter 150.0g NaCl;

20.0g MgCl2; 5.0g K2SO4; 0.1g CaCl2; 5.0g yeast extract. Halorhabdus utahensis (DSMZ 12940) was grown in the liquid medium described by Waino et al. (WAINO et al., 2000), which contained per liter 270.0g NaCl; 0.1g NaBr; 20.0g MgSO4 x 7H2O; 5.0g KCl; 2.0g NH4Cl; 12.0g

Tris-HCl; 0.125g KH2PO4; 0.05g CaCl2 x 2H2O; 5.0mg FeCl2 x 4H2O; 5.0mg MnCl2 x 4H2O;

0.5g yeast extract; 2.0g glucose. Natronomonas pharaonsis (DSMZ 3395) was grown at pH 9.0 in the liquid medium described by Tindall et al. (TINDALL et al., 1984), which contained 200.0g

NaCl; 1.0g KH2PO4; 1.0g NH4Cl; 0.24g MgSO4 x 7H2O; 0.17g CaSO4 x 2H2O; 5.0g yeast extract; 1.0g glucose; 5.0g casamino acids; 5.0g Na2CO3; 1 ml trace element solution (per liter

1.5g FeCl2 x 4H2O; 100.0mg MnCl2 x 4H2O; 70.0mg ZnCl2; 6.0mg H3BO3; 190.0mg CoCl2 x

6H2O; 2.0mg CuCl2x2H2O; 24.0mg NiCl2x6H2O; 36.0mg Na2MoO4 x 2H2O). Halobaculum gomorrense (DSMZ 9297) was grown in the liquid medium described by Oren et al. (OREN et al.,

109 1995), which contained per liter 125.0g NaCl; 160.0g MgCl2 x 6H2O; 5.0g K2SO4; 0.1g CaCl2 x

2H2O; 2.0g soluble starch; 1.0g yeast extract; 1.0g casamino acids. All media were modified for the salinity experiment to contain 100g, 150g, 200g, 250g and 300g per liter NaCl. Cells were harvested in late exponential phase by centrifugation.

5.3.2 Lipid extraction and analysis

Cell pellets were lyophilized prior to lipid extraction by a modified Bligh-Dyer extraction as described in Macalady et al. (2004). Briefly, the lipids were extracted with methanol/chloroform/water (2:1:0.8). DGDs were isolated from extracted pigments through precipitation in cold acetone (CHOQUET et al., 1992). In order to preserve allyl groups, base saponification rather than acid methanolysis was used to prepare core lipids from the intact polar lipid (IPL) (KOGA and MORII, 2006; NISHIHARA et al., 1989; PANGANAMALA et al., 1971).

Briefly, IPL’s were treated with 0.5 N KOH in methanol/water (3:1) at 75°C for 2 hours. After cooling, the saponified material was extracted with hexane/chloroform (4:1), dried over Na2SO4 and evaporated to dryness with N2.

Extracted core lipids were derivatized using N,O-bis(trimetylsilyl)trifluoroacetamide

(BSTFA) and pyridine (1:1) in hexane at 65°C for 20 minutes . Silylated core lipids were analyzed on a Hewlett-Packard 5972 GC/MS with a DB-5 column (length 30 m; inside diameter,

0.25 mm; film thickness, 0.25 μm). The injector temperature was 320°C. The oven was held at

65°C for 1 min before increasing the temperature by 6°C/min to 320°C where it was maintained for 20 min. Peak areas were determined using MSD ChemStation version E.02.00 (Agilent,

Wilmington, DE).

110 5.4 Results

5.4.1 Identification of DGDs

After derivatization with BSTFA, analysis by GC-MS revealed the TLE and saponified material to be DGDs. Identification of archaeol and its unsaturated derivatives followed MS fragmentation patterns published by Teixidor and Grimalt (1992), Teixidor et al. (1993) and

Steihl et al. (2005). Major MS fragments (m/z) of the archaeol-TMS derivative include: the

- molecular ion (724 m/z); M – C20H41 (445 m/z); M – C20H41OH (426 m/z); M –

C20H41OSi(CH3)3 (369 m/z); C20H41 (278 m/z); and CH2CHCH2OSi(CH3)3 (130 m/z). The presence of unsaturation was observable in the shifting of the major ions by 2-8 m/z and later elution times (Figure 5-2). Extracts of N. pharaonsis and H. utahensis showed the presence of

C25-20 DGDs, a sesterpanyl as well as a phytanyl chain. The major MS fragments of the C25-20

DGDs include: the molecular ion (794 m/z); M – C20H41 (515 m/z); M – C25H51OH (497 m/z); M

– C25H51 (445 m/z); C25H51 (348 m/z); C20H41 (278 m/z) and CH2CHCH2OSi(CH3)3 (130 m./z).

Again, the presence of unsaturation was observable in the shifting of the major ions by 2-10 m/z and later elution times (Figure 5-2). The observed MS fragmentation is inconsistent with previously described macrocyclic DGD structures (COMITA et al., 1984; STADNITSKAIA et al.,

2003).

5.4.2 DGDs in halophilic archaeal isolates

GC-MS analysis of saponified archaeal lipids extracts from four halophilic archaeal strains revealed that the dominant compounds are DGDs (Figure 5-3). DGDs included the C20-20 and the C25-20 isoprenoid diethers, as well as their unsaturated analogues. Shifts to lower values in the m/z of molecular ions and majors fragments indicated the presence of 1-5 double bonds in

111 unsaturated DGD analogues. C20-20 DGDs were observed in H. gomorrense, H. sulfurifontis, and

N. pharaonsis. C20-25 DGDs were observed in N. pharaonsis and H. utahensis. The percentage of

DGDs with unsaturation increased with the concentration of NaCl in the growth medium in all strains. Insufficient biomass was produced at 10% (w/v) NaCl for lipid analysis in H. utahensis and N. pharaonsis. Insufficient biomass was produced at 30% (w/v) NaCl for lipid analysis in H. gomorrense.

In order of increasing optimal NaCl concentration, the halophile strains were H. gomorrense (12.5% (w/v)), H. sulfurifontis (15% (w/v)), N. pharaonis (20% (w/v)) and H. utahensis (27.5% (w/v)). In H. gomorrense, at 10% (w/v) NaCl, 6% of DGD’s were unsaturated.

The percentage of unsaturated DGD’s decreased to 2.5% during growth at 15% (w/v) NaCl and increased to 3.5% during growth at 30% (w/v) NaCl. In H. sulfurifontis, at 15% (w/v) NaCl,

8.5% of DGD’s contained unsaturation. The percentage increased to 23.5% at 30% (w/v) NaCl.

In N. pharonsis, at 15%, (w/v) NaCl, 25% of DGD’s contained unsaturation. The percentage increased to 45% at 30% (w/v) NaCl. In H. utahensis, during growth at 15%, (w/v) NaCl, 62% of

DGD’s contained unsaturation. The percentage increased to 87% at 30% (w/v) NaCl. At all salinities H. utahensis contained only C25-20 DGDs with the unsaturated analogues consistently representing the dominant structure. N. pharaonis contained both C20-20 and C25-20 DGDs. H. sulfurifontis and H. gomorrense contained only C20-20 DGDs. Figure 5-4 shows the fraction of unsaturated DGDs in the polar lipid extract as a function of NaCl concentration for the four strains. An additional analysis of the average fraction of unsaturated DGDs shows a strong linear correlation with the strain specific optimal salinity (Figure 5-5). The linear regression describing this correlation is given by equation y = 0.0049 – 0.6027, and the R2 value for the linear regression is 0.9791.

112

5.5 Discussion

The C20-20 or C25-20 isoprenoid chains of DGDs from halophilic archaea include both saturated and unsaturated analogues, as has previously been reported by Stiehl et al. (2005) for

Halobacterium marismortui, Haloferax volcanii and Halorubrum sodomense and by Franzmann et al. (1988) and Gibson et al. (2005) for H. lacusprofundi. Analysis of the core DGD structure of halophilic archaeal isolates from strains of four genera revealed three trends. (1) The percentage of unsaturated DGDs in all of the examined strains increased with increasing concentration of NaCl in the growth medium. (2) The percentage of unsaturated DGDs is higher in halophilic archaeal strains with a higher optimal NaCl concentration than in strains. For example, in H. gomorrense (12.5% (w/v) NaCl) 2.5-6% of DGDs were unsaturated as compared to in 62-87% of DGDs H. utahensis (27% (w/v) NaCl). (3) The presence of C25-20 DGDs occurred in the strains with higher optimal NaCl concentrations, N. pharaonsis and H. utahensis.

The C25-20 structure was the sole core DGD structure found in H.utahensis, the strain with the highest optimal NaCl concentration.

Many studies of novel halophilic isolates report the major lipids present on the basis of thin layer chromatography (TLC) with comparison to previously identified strains and authentic standards. This technique provides good detection of the major polar lipids (i.e. phosphatidylglycerol (PG), phosphatidylglycerophosphate (PGP)). Current studies indicate that the genera Halobacterium, Haloarcula, Haloferax, Halobaculum and Halorubrum contain only the C20-20 DGD structure (ASKER and OHTA, 2002; KAMEKURA and KATES, 1999; OREN et al.,

1995) (Table 5-1). The C25-20 DGD structure has been identified as the sole core lipid structure or in addition to the C20-20 DGD in the genera Halococcus, Natronobacterium, Natronococcus,

113 Natronomonas, Natrialba, Natrinema, Natronorubrum, Haloterrigena and several other isolates with optimal pH > 7.0 (KAMEKURA and KATES, 1999; ROMANO et al., 2007; TINDALL et al., 1984;

XU et al., 2001; XU et al., 1999) (Table 5-1). However, in many other studies the core isoprenoid structure is not reported, or as in the case of the description of Halorhabdus utahensis may be misidentified as the C20-20 DGD (WAINO et al., 2000). In addition to polar and glycolipid identification by TLC, base saponification of archaeal lipid extracts followed by BSTFA derivatization and GC-MS analysis provides unambiguous identification of the core lipid structures.

The strong linear correlation between optimal growth salinity and the amount of unsaturated DGDs (Figure 5-5) suggests that the degree of membrane lipid unsaturation is an important adaptation to specific salinity niches in archaeal halophiles. In addition, in three of the four halophile strains we tested, the fraction of unsaturated DGDs increased above a salinity threshold or in response to increasing salinity in the growth medium. Thus, halophilic archaea may regulate membrane lipid unsaturation in response to environmental salinity changes regardless of their salinity optima. Dannenmuller et al. (2000) showed evidence of increased membrane bilayer stability and a decreased membrane permeability with the incorporation of a single double bonds into each DGD isoprenoid chain. In contrast, synthesis of C20-20 versus C25-20

DGDs appears to be related to taxonomy (Table 5-1). However, in halophile strains containing both core lipid structures, a response to salinity is suggested by an increase in the relative proportions of C25-20 to C20-20 with increasing salinity in Natronococcus occultus (NICOLAUS et al., 1989).

Previous studies of archaeal lipid stability showed that liposomes composed of glycerol dialkyl glycerol tetraethers (GDGT’s) provided greater thermal stability and reduced permability

114 as compared to macrocyclic archaeol, which in turn was superior to archaeol (CHOQUET et al.,

1992; CHOQUET et al., 1994; ELFERINK et al., 1994; VAN DE VOSSENBERG et al., 1998). However, a study by Gmajner et al. (2011) examined the physical properties of C25-25 DGDs from the hyperthermophile Aeropyrum pernix and showed comparable thermal stability and permeability of these liposomes compared to those composed of GDGT’s. The C25 sesterpanyl isoprenoid chain is 20% longer than the C20 phytanyl chain, thus membranes composed solely of these longer lipids would be 20% thicker (GMAJNER et al., 2011). Evidence of C25-20 DGDs providing increased membrane stability and decreased permeability in hypersaline conditions is provided by the identification of a novel hydroxylated C25-20 DGD structure in sediments from a saline cold seep (STADNITSKAIA et al., 2008). Stadnitskaia et al (2008) propose that the novel structure may represent a biomarker for methanotrophs at elevated salinity. Membranes composed of C25-

20 DGDs could represent an intermediate level of stability and reduced permeability necessary for growth in certain hypersaline conditions.

5.6 Conclusions

Archaeal DGDs with unsaturation were found in four halophile strains. The unsaturated analogues as a percentage of total DGDs increased with increasing NaCl concentration in the growth medium. Unsaturated DGDs included analogues of both C20-20 and C25-20 DGDs core structures. Increasing unsaturation in halophilic archaeal membrane lipids might represent a physiological response to transient increases in salinity. The presence of C25-20 DGDs is typically associated with haloalkaliphiles. However, here we found C25-20 DGDs as the sole membrane lipid structure in Halorhabdus utahensis, a strain which grows optimally between pH 6.7-7.1.

115 Although the presence or absence C25-20 DGDs appears to be related to taxonomy, the structure may also provide increased membrane stability and decreased permeability.

5.7 Acknowledgements

We thank D. Walizer for technical support. This work was supported by the NASA

Astrobiology Institute (NAI) through a NAI DDF award to C.H. House and by a Pennsylvania

State University Biogeochemistry Program fellowship.

116 5.8 Figures and Tables:

Table 5-1: Presence or absence of C20-20 and C25-20 DGDs in various halophilic archaea genera.

The presence of a core lipid structure is indicated with ‘+’, absence with ‘-‘, and data not

available with ‘n.a’.

Optimal Optimal Genus Species C C Reference NaCl % pH 25-20 20-20 Haloalkalicoccus Haloalkalicoccus tibetensis 20% 9.5-10.0 + + (XUE et al., 2005) Haloarcula Haloarcula argentinensis 15-17.5% n.a - + (IHARA et al., 1997) Haloarcula Haloarcula mukohataei 17.5-20% n.a - + (IHARA et al., 1997) Haloarcula Haloarcula marismortui 20-23% n.a - + (OREN et al., 1990) Halobaculum Halobaculum gomorrense 9-15% 6.0-7.0 - + (OREN et al., 1995) Halobiforma Halobiforma hoaloterrestris 20% 7.5 + + (HEZAYEN et al., 2002) (ASKER and OHTA, Haloferax Haloferax alexandrius 25% 7.2 - + 2002) Haloferax Haloferax larsenii 12-20% 6.5-7.0 - + (XU et al., 2007) Haloferax Haloferax sulfurifontis 15% 6.4-6.8 - + (ELSHAHED et al., 2004) Halopiger Halopiger xanaduensis 25% 7.5-8.0 + + (GUTIERREZ et al., 2007) Halorhabdus Halorhabdus utahensis 27% 6.7-7.1 + - (WAINO et al., 2000) Halorubrum Halorubrum xinjiangense 18-20% 7.0-7.5 - + (FENG et al., 2004) (FRANZMANN et al., Halorubrum Halorubrum lacusprofundi 14.5-20% n.a. - + 1988) Halostagnicola Halostagnicola larsenii 20% 7.0-8.0 + + (CASTILLO et al., 2006b) Haloterrigena Haloterrigena salina 20% 7.5 + + (GUTIERREZ et al., 2008) Haloterrigena Haloterrigena hispanica 20% 7 + + (ROMANO et al., 2007) Halovivax Halovivax asiaticus 20% 7.0-7.5 + + (CASTILLO et al., 2006a) Natrialba Natrialba aegyptiaca 15-17.5% 7.0-8.0 + + (HEZAYEN et al., 2001) Natrialba Natrialba taiwanensis 20% 7.5-7.8 + + (HEZAYEN et al., 2001) Natrialba Natrialba hulunbeirensis 20% 9 + + (XU et al., 2001) Natrialba Natrialba chahannaoensis 15% 9 + + (XU et al., 2001) Natrinema Natrinema versiforme 20-25% 6.5-7.0 + + (XIN et al., 2000) Natrinema Natrinema pellirubrum 20-25% 7.2-7.6 + + (MCGENITY et al., 1998) Natronomonas Natronomonas pharaonis 20% 8.5 + + (TINDALL et al., 1984) Natronorubrum Natronorubrum tibetense 20% 9 + - (XU et al., 1999) Natronorubrum Natronorubum bangense 22.50% 9.5 + - (XU et al., 1999)

117

Figure 5-1: Structures of archaeal dialkyl glycerol diether (DGD) with various chain lengths, cyclization, unsaturation and hydroxyl substitutions.

118

Figure 5-2: Mass spectra of saturated C20-20 DGD and C25-20 DGD.

119

Figure 5-3: Partial GC-MS chromatograms of polar lipid extracts of halophilic archaea grown at strain specific optimal salinity. Peaks eluting between 42 and 45 minutes are C20-20 DGDs. Peaks eluting between 45 and 47.5 minutes are C25-20 DGDs.

120

Figure 5-4: Plot of total unsaturated archaeal C20-20 and C20-25 DGD lipids as a fraction of total polar DGDs in four halophilic archaea. Cultures were grown between 10% and 30% NaCl (w/v).

Halorhabdus utahensis (blue diamonds) makes solely C20-25 DGDs and grows optimally at

27.5% NaCl (w/v). Natronomonas pharaonis (pink squares) makes both C20-20 and C20-25 DGDs and grows optimally at 20% NaCl (w/v). Haloferax sulfurifontis (green triangles) and

Halobaculum gomorrense (orange circles) make solely C20-20 DGDs and grow optimally at 15 %

NaCl (w/v) and 12.5% NaCl (w/v) respectively. Optimal NaCl concentrations are indicated with bars on the graph.

121

Figure 5-5: Average fraction of unsaturated DGDs versus optimal % NaCl (w/v) for four halophilic archaeal strain

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124 Kamekura M. (1998) Diversity of extremely halophilic bacteria. Extremophiles 2, 289-295.

Kamekura M. and Kates M. (1999) Structureal Diversity of Membrane Lipids in Members of Halobacteriaceae. Bioscience, Biotechnology and Biochemistry 63(6), 969-972.

Kates M. (1977) The phytanyl ether-linked polar lipids and isoprenoid neutral lipids of extremely halophilic bacteria. Progress in the Chemistry of Fats and other Lipids 15(4), 301-342.

Koga Y. and Morii H. (2006) Special methods for the analysis of ether lipid structure and metabolism in archaea. Analytical Biochemistry 348(1), 1-14.

Lattanzio V., Corcelli A., Mascolo G., and Oren A. (2002) Presence of two novel cardiolipins in the halophilic archaeal community in the crystallizer brines from the salterns of Margherita di Savoia (Italy) and Eilat (Israel). Extremophiles 6(6), 437-444.

Ley R. E., Harris J. K., Wilcox J., Spear J. R., Miller S. R., Bebout B. M., Maresca J. A., Bryant D. A., Sogin M. L., and Pace N. R. (2006) Unexpected Diversity and Complexity of the Guerrero Negro Hypersaline Microbial Mat. Appl. Environ. Microbiol. 72(5), 3685-3695.

Lizama C., Monteoliva-Sanchez M., Suarez-Garcia A., Rosello-Mora R., Aguilera M., Campos V., and Ramos-Cormenzana A. (2002) Halorubrum tebenquichense sp. nov., a novel halophilic archaeon isolated from the Atacama Saltern, Chile. Int J Syst Evol Microbiol 52(1), 149-155.

McGenity T. J., Gemmell R. T., and Grant W. D. (1998) Proposal of a new halobacterial genus Natrinema gen. nov., with two species Natrinema pellirubrum nom. nov. and Natrinema pallidum nom. nov. Int J Syst Bacteriol 48(4), 1187-1196.

Nichols D. S., Miller M. R., Davies N. W., Goodchild A., Raftery M., and Cavicchioli R. (2004) Cold Adaptation in the Antarctic Archaeon Methanococcoides burtonii Involves Membrane Lipid Unsaturation. J. Bacteriol. 186(24), 8508-8515.

Nicolaus B., Lanzotti V., Trincone A., De Rosa M., Grant W. D., and Gambacorta A. (1989) Glycine betaine and polar lipid composition in halophilic archaebacteria in response to growth in different salt concentration. FEMS microbiology Letters 59, 157-160.

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Oren A. (1999b) Microbiological studies in the Dead Sea: future challenges toward the understanding of life at the limit of salt concentrations. Hydrobiologia 405(0), 1-9.

125 Oren A. (2002) Molecular ecology of extremely halophilic Archaea and Bacteria. FEMS Microbiology Ecology 39(1), 1-7.

Oren A., Arahal D. R., and Ventosa A. (2009) Emended descriptions of genera of the family Halobacteriaceae. Int J Syst Evol Microbiol 59(3), 637-642.

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Chapter 6: Summary and Future Directions

6.1 Anaerobic isoprenoid degradation

Chapters 2 and 3 described the anaerobic degradation of pristane and phytane by a denitrifying enrichment, and showed evidence that the same enrichment can utilize archaeal diphytanyl glycerol diether (DGD) lipids. Anaerobic pristane degradation was previously described for a denitrifying enrichment (BREGNARD et al., 1997) and a sulfate reducing enrichment (GROSSI et al., 2000). However, I provided the first description to date of the taxonomic identities of microbial communities associated with linear isoprenoid degradation. My results indicate that the isoprenoid degrading enrichments are composed of a low diversity community of Beta- and Gammaproteobacteria based upon Fluorescent in situ hybridization

(FISH) and 16S rDNA clone libraries. Through FISH coupled to secondary ion mass spectrometry (SIMS) or DNA-stable isotope probing (SIP) techniques, we could trace

13 12 enrichment in the ratio of C/ C to particular phylotypes in aggregates of cells (ORPHAN, 2009;

ORPHAN et al., 2001) or in extracted DNA (DUMONT and MURRELL, 2005; NEUFELD et al., 2007).

This would provide a conclusive identification of the microorganisms responsible for isoprenoid degradation, as well as provide further support for the assimilation of 13C-phytane by anaerobic bacteria.

Anaerobic hydrocarbon degradation initiates with the activation of the hydrocarbon substrate. Fumarate addition, described in Chapter 1, is the primary mechanism of alkane activation (CALLAGHAN et al., 2006; HEIDER, 2007; KNIEMEYER et al., 2007; WIDDEL et al.,

2006). Analysis of pristane degrading enrichment cultures failed to detect metabolites indicative of fumarate addition or metabolites suggestive of an alternative anaerobic activation mechanism

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for isoprenoids. Further analysis of these cultures by microarray may indicate the presence of genes associated with fumarate addition (GUSCHIN et al., 1997). Additional extractions and identification of metabolites produced from 13C-labelled phytane degradation may indicate activation by a novel mechanism necessitated by the methyl-branched structure of isoprenoids or a modified version of fumarate addition.

6.2 Coal bed methane biogeochemistry

In Chapter 4, I used FISH to examine the microbial community associated with coal bed methane (CBM) production waters in the Cook Inlet, Alaska. FISH identification of the active lineages of bacteria and archaea confirmed the dominant lineages retrieved by 16S rRNA tag pyrosequencing. Additionally, FISH provided a visualization of a potential bacterial-archaeal syntrophic association between archaeal methanogens Methanosarcina spp. and Methanolobus spp. and anaerobic bacteria including Acetobacterium spp. and Bacteriodales spp. The combination of methane isotope values and microbiology suggest methanogenesis via an acetoclastic/methylotrophic pathway. However, variation in the microbial populations across several sampling sites in the same basin indicates that the specific microbial pathway of coal transformation to methane depends upon the geochemistry, hydrology and coal structure present at microenvironments within the gas field. Further analysis of nutrients, trace element concentrations and redox potential will provide a better understanding of the variation in microbial community structure observed in the Cook Inlet samples, as well as in other CBM basins. Additionally, the isolation of the syntrophic association identified in Chapter 4 may reveal the biogeochemical conditions necessary to stimulate similar bacterial-archaeal

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associations for coal transformation, and natural abundance FISH-SIMS may clarify the nature of the syntrophic association.

6.3 Halophilic archaeal lipids

In Chapter 5, I investigated changes in the molecular structure of archaeal DGD lipids in cultures grown at a range of salinities. Unsaturated analogues of archaeol, previously reported by in Dead Sea (STIEHL et al., 2005) and Antarctic halophile isolates (FRANZMANN et al., 1988;

STIEHL et al., 2005), were present in Halorhabdus utahensis, Natronomonas pharaonis,

Haloferax sulfurifontis and Halobaculum gomorrense. The percentage of unsaturated lipids increased as a function of NaCl concentration in growth medium and reflected the optimal salinity for each strain. Additionally, GC-MS analysis of derivatized DGDs provided unambiguous identification of both C20-20 and C25-20 core isoprenoid structures. H. utahensis, previously described as synthesizing C20-20 DGDs (WAINO et al., 2000), was shown to produce

C25-20 DGDs exclusively. My experiments revealed the influence of a NaCl concentration on core

DGD molecular structure. Further analysis of the intact polar lipids produced by these archaeal strains may reveal variation in the polar head group in response to salinity. A comparison of the impact of a salinity gradient utilizing MgCl2 to the results described in Chapter 5 may provide insight into strategies for coping with hypersaline environments like the Dead Sea where there is a dominance of divalent over monovalent cations (OREN, 1999). Further knowledge of changes in archaeal membrane lipid structure in response to salinity will allow for improved interpretation of paleoenvironmental conditions based upon geologically preserved biomarkers derived from DGDs.

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6.4 References

Bregnard T. P., Haner A., Hohener P., and Zeyer J. (1997) Anaerobic Degradation of Pristane in Nitrate-Reducing Microcosms and Enrichment Cultures. Appl. Environ. Microbiol. 63(5), 2077-2081.

Dumont M. G. and Murrell J. C. (2005) Stable isotope probing - linking microbial identity to function. Nat Rev Micro 3(6), 499-504.

Guschin D. Y., Mobarry B. K., Proudnikov D., Stahl D. A., Rittmann B. E., and Mirzabekov A. D. (1997) Oligonucleotide microchips as genosensors for determinative and environmental studies in microbiology. Appl. Environ. Microbiol. 63(6), 2397-2402.

Heider J. (2007) Adding handles to unhandy substrates: anaerobic hydrocarbon activation mechanisms. Current Opinion in Chemical Biology 11, 188-194.

Neufeld J. D., Vohra J., Dumont M. G., Lueders T., Manefield M., Friedrich M. W., and Murrell J. C. (2007) DNA stable-isotope probing. Nat. Protocols 2(4), 860-866.

Oren A. (1999) Microbiological studies in the Dead Sea: future challenges toward the understanding of life at the limit of salt concentrations. Hydrobiologia 405(0), 1-9.

Orphan V. J. (2009) Methods for unveiling cryptic microbial partnerships in nature. Current Opinion in Microbiology 12(3), 231-237.

Orphan V. J., House C. H., Hinrichs K.-U., McKeegan K. D., and DeLong E. F. (2001) Methane- Consuming Archaea Revealed by Directly Coupled Isotopic and Phylogenetic Analysis. Science 293(5529), 484-487.

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Widdel F., Boetius A., and Rabus R. (2006) Anaerobic Biodegradation of Hydrocarbons Including Methane. In The Prokaryotes, pp. 1028-1049.

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CURRICULUM VITAE KATHERINE STRIGARI DAWSON EDUCATION 2011 Ph.D. Geosciences/Biogeochemistry, Pennsylvania State University, August 2011 Title: Biogeochemistry of isoprenoid production and anaerobic hydrocarbon biodegradation 2005 B.A. Chemistry, Goucher College, May 2005 Title: Fluorescence of 4(1)-Pyrrolyl-Pyridine TEACHING AND RESEARCH EXPERIENCE Present Pennsylvania State University, Postdoctoral Researcher 2005-2011 Pennsylvania State University, Research Assistant 2007 ConocoPhillips, Geochemistry Intern 2006 Pennsylvania State University, Teaching Assistant 2002-2003 Goucher College, Teaching Assistant AWARDS AND HONORS 2011 Biogeochemistry Program Fellowship 2010 Chesapeake Energy Scholarship in Geosciences 2009 Hiroshi and Koya Ohmoto Graduate Fellowship in Geosciences 2008/2006 Donald and Mary Tait Scholarship in Microbial Biogeochemistry 2007 John Meachum Hunt Graduate Student Award in Petroleum Geochemistry 2005 Biogeochemical Research Initiative in Education Fellowship PUBLICATIONS JONES, D. S., SCHAPERDOTH, I., STOFFER, T. S., DAWSON, K.S., FREEMAN, K. H., ALBRECHT, H.A., and MACALADY, J. L. Metagenomic, phylogenetic, and culture- based analysis of subsurface extremophile microbial communities. ISME Journal. In press. STRAPOC, D., MASTALERZ, M., DAWSON, K.S., MACALADY, J. L., CALLAGHAN, A., WAWRIK, B., and ASHBY, M. Biogeochemistry of Coal Bed Methane. (2011) Annual Review of Earth and Planetary Science 39, 617-656. ANTICIPATED PUBLICATIONS DAWSON, K.S., SCHAPERDOTH, I., FREEMAN, K.H., and MACALADY, J.L. Stable isotope probing of anaerobic biodegradation with 13C labeled phytane. In prep for ES&T. DAWSON, K.S., STRAPOC, D., HUIZINGA, B., LIDSTROM, U., ASHBY, M. and MACALADY, J.L. Quantitative FISH analysis of a microbial consortia from a biogenic gas field in the Cook Inlet Basin, Alaska. Submitted to AEM. DAWSON, K.S., FREEMAN, K.H. and MACALADY, J.L. Molecular characterization of archaeal lipids across a hypersaline gradient. In prep for Geobiology