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The Pennsylvania State University

The Graduate School

Department of Geosciences

MOLECULAR AND ISOTOPIC SIGNATURES OF MICROORGANISMS

IN LOW-OXYGEN MARINE ENVIRONMENTS

A Dissertation in

Geosciences

Laurence R. Bird

Submitted in Partial Fulfillment

of the Requirements for the Degree of

Doctor of Philosophy

December 2016

The dissertation of Laurence R. Bird was reviewed and approved by the following:

Katherine H. Freeman Evan Pugh University Professor Department of Geosciences Dissertation Advisor Chair of Committee

Jennifer L. Macalady Associate Professor of Geosciences

Christopher H. House Associate Professor of Geosciences

Squire J. Booker Professor of Chemistry and of Biochemistry and Molecular Biology

Demian M. Saffer Professor of Geosciences Associate Head for Graduate Programs and Research in Geosciences

*Signatures are on file in the Graduate School

Abstract

This dissertation explores the molecular and isotopic signatures of methanotrophic Archaea and the molecular signatures of cyanobacteria in low oxygen environments. Archaeal ANerobic MEthaneotrophs (ANME) oxidize methane in anoxic sediment, and prevent methane, a potent greenhouse gas from reaching the atmosphere. This process is hypothesized to take place via the reversal of methanogenesis based on culture and genetic evidence. Coenzyme F430 is a tetrapyrrole used in the last step of methanogenesis, and likely enables the first step in reverse methanogenesis. Therefore, the presence and concentration of F430 in association with AOM serves as a test for the reverse methanogenesis pathway in sediment.

In chapter 2, F430 was extracted, quantified, and isotopically analyzed in methanotrophic sediment from Hydrate Ridge and the Santa Monica Basin (west coast U.S.A). The greatest amounts of F430 were recovered where sulfide, sulfate, and methane concentration profiles indicate the greatest AOM activity in the sediment. These sediment horizons also contained the highest ANME-2 aggregate counts. F430 was found to be isotopically distinct from methane and archaeal lipids, but similar to dissolved inorganic carbon (DIC). In the Hydrate Ridge and Santa Monica sediment F430 was ~60‰ enriched in 13C relative to archaeol lipids.

In chapter 3, the dual assimilation of methane and DIC is explored with a series of stable isotope labeling experiments using sediment from Hydrate Ridge and the Santa Monica Basin. In experiments using Hydrate Ridge sediments, we observed the 13C label from DIC assimilated into archaeol, while in experiments using Santa Monica Basin sediment the 13C labeled from DIC and methane was assimilated into both F430 and lipids. The amount of DIC assimilated into F430 and lipids ranged from ~50% to 100%, with between 0% to 20% of carbon coming from methane. Due to the amount of labeled methane that is oxidized to DIC we cannot be sure if methane is directly assimilated or first oxidized to DIC. Coenzyme F430 was also only recovered from experiments where methane was added to the headspace, strengthening the link between F430 and methanotrophy.

Little Salt Springs is a sinkhole in Florida where a red biofilm in the euxinic water column produces large amounts of bacterialhopanetetrol (BHT), 2-methyl bacterialhopanetetrol (2-MeBHT) and 2-methyl anhydrobacterialhopanetetrol (2-MeAnhydroBHT). The amount of each BHT produced varies seasonally and between years, with the geochemical cause of this variability unknown. In chapter 4, a red cyanobacteria isolated from this biofilm was cultured under a number of different geochemical conditions in an attempt to identify possible causes for variability in bacteriohopanepolyols (BHP) production. No single geochemical control was identified as amounts of BHT and 2-MeAnhydroBHT were similar in all experiments. Future experiments should explore what effects oxygen concentration, fixed nitrogen species, trace metals, microbial community and combinations of different conditions have on BHP production.

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Table of Contents

List of Figures ...... vi List of Tables ...... viii Acknowledgements ...... x Epigraph ...... xi Chapter 1: Introduction ...... 1 1.1. Anoxic methane oxidation ...... 1 1.2. ANME biochemistry ...... 2 1.3. Hopanoids ...... 4 1.4. Bacteriohopanepolyols ...... 4 1.5. Anticipated publications from this work ...... 6 1.6. Figures and tables ...... 7 1.7. References ...... 14 Chapter 2: Carbon isotopic heterogeneity between ANME biomolecules ...... 19 2.1. Abstract ...... 19 2.2. Introduction ...... 19 2.3. Materials and Methods ...... 21 2.4. Results ...... 28 2.5. Discussion ...... 30 2.6. Conclusions ...... 33 2.7. Acknowledgements ...... 34 2.8. Figures and tables ...... 34 2.9. References ...... 51 Chapter 3: Stable isotope probing of ANME carbon assimilation ...... 55 3.1. Abstract ...... 55 3.2. Introduction ...... 55 3.3. Methods ...... 57 3.4. Results ...... 66 3.5. Discussion ...... 68 3.6. Conclusions ...... 72 3.7. Acknowledgements ...... 72 3.8. Figures and tables ...... 73 3.9. References ...... 93 Chapter 4: Quantifying Bacteriohopanepolyol production in Little Salt Springs cyanobacteria ...... 96 4.1. Abstract ...... 96 4.2. Introduction ...... 96 4.3. Methods ...... 98 4.4. Results ...... 101 4.5. Discussion ...... 102 4.6. Conclusions ...... 104 4.7. Acknowledgements ...... 104 4.8. Figures and tables ...... 105

4.9. References ...... 119 Chapter 5: Research summary ...... 122 5.1. Chapter summaries ...... 122 5.2. Future directions ...... 123 5.3. References ...... 124 Appendix A: F430 abundance and isotope values from the Santa Monica basin ...... 126 A.1. Introduction ...... 126 A.2. Methods ...... 126 A.3. Results ...... 130 A.4. Conclusions ...... 130 A.5. Figures and tables ...... 131 A.6. References ...... 132 Appendix B: Bacteriohopanepolyols through the Little Salt Springs water column ...... 133 B.1. Introduction ...... 133 B.2. Methods ...... 133 B.3. Results ...... 135 B.4. Conclusions ...... 135 B.5. Figures ...... 136 B.6. References ...... 137 Appendix C: F430 Extraction and Purification for quantification and Isotope analysis .. 138 C.1. Extraction ...... 138 C.2. Column chromatography ...... 139 C.3. HPLC Purification and quantification ...... 142 C.4. NANO-EA IRMS ...... 144 C.5. Figures and tables ...... 146 C.6. References ...... 149 Appendix D: Data tables ...... 150

List of Figures

Figure 1-1: FISH image of ANME-2...... 7 Figure 1-2: ANME phylogenetic tree...... 8 Figure 1-3: Structure of co-enzyme F430...... 9 Figure 1-4: Hopene...... 9 Figure 1-5: LCMS response to BHP structures...... 10 Figure 1-6: Red biofilm Bacteriohopanepolyols...... 10 Figure 1-7: BHP polar groups...... 11 Figure 2-1: Sampling localities...... 34 Figure 2-2: LC fraction collection...... 35 Figure 2-3: F430 Uv/vis absorbance spectra...... 36 Figure 2-4: F430 ion spectra...... 37 Figure 2-5: Hydrate ridge 1/n plot...... 38 Figure 2-6: Santa Monica 1/n plot...... 39 Figure 2-7: Hydrate Ridge sediment data ...... 40 Figure 2-8: Santa Monica sediment data ...... 41 Figure 2-9: Hydrate ridge carbon isotope values...... 42 Figure 2-10: Santa Monica basin carbon isotope values ...... 43 Figure 2-11: Mass balance modeling results ...... 44 Figure 2-12: Carbon assimilation diagram...... 45 Figure 3-1: PCKD core...... 73 Figure 3-2: Hydrate Ridge 1/n carbon plot...... 74 Figure 3-3: Hydrate Ridge 1/n nitrogen plot ...... 75 Figure 3-4: Santa Monica 1/n carbon plot...... 76 Figure 3-5: Santa Monica 1/n nitrogen plot...... 77 Figure 3-6: Santa Monica 1/n carbon plot...... 78 Figure 3-7: Santa Monica 1/n nitrogen plot...... 79 Figure 3-8: Hydrogen sulfide for PCKD...... 80 Figure 3-9: δ13C-DIC for PCKD experiments...... 81 Figure 3-10: Santa Monica F430 nitrogen results...... 82 Figure 3-11: Uptake of nitrogen into F430 ...... 83 Figure 3-12: Hydrate Ridge Carbon isotope results...... 84 Figure 3-13: Santa Monica F430 carbon values...... 85

Figure 3-14: Santa Monica F430 carbon and nitrogen isotope data...... 86 Figure 3-15: Santa Monica archaeol carbon values...... 87 Figure 3-16: The carbon assimilation for F430 and lipids...... 88 Figure 3-17: Carbon flow within the ANME cell...... 89 Figure 4-1: Water column geochemistry for Little Salt Springs June 2012...... 105 Figure 4-2: Biofilm and cyanobacterial BHP content...... 106 Figure 4-3: Bacterialhopenetetrol mass spectrum ...... 107 Figure 4-4: 2-methyl bacterialhopenetetrol mass spectrum ...... 107 Figure 4-5: 2-methyl anhydro bacterialhopenetetrol mass spectrum ...... 107 Figure 4-6: BHT MS response ...... 108 Figure 4-7: 2-MeAnhydroBHT MS response ...... 109 Figure 4-8: Results of limited light and shaken experiments ...... 110 Figure 4-9: Results of sulfur species experiments ...... 111 Figure 4-10: Results of high salt experiments...... 112 Figure 4-11: 2-Methyl ratio in culture experiments ...... 113 Figure 4-12: Results of control time series experiments ...... 114 Figure 4-13: Results of no fixed nitrogen time series experiments ...... 115 Figure 4-14: Ratio in control cultures...... 116 Figure 4-15: Ratio in no fixed nitrogen cultures...... 117 Figure A-1: F430 concentration in PC6 ...... 131 Figure B-1: Biofilm BHP composition ...... 136 Figure B-2: Concentration of 2-MeAnhydroBHT in the water column...... 137 Figure C-1: LCMS solvent profile using waters columns...... 146 Figure C-2: LCMS profile using Thermo Hypercarb column...... 147 Figure C-3: Nano EA IRMS system diagram...... 148

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List of Tables

Table 1-1: ANME isotope values...... 12 Table 1-2: BHT quantification...... 13 Table 2-1: Inorganic carbon isotope values for Hydrate Ridge ...... 46 Table 2-2: Inorganic carbon isotope values for Santa Monica Basin ...... 46 Table 2-3: Isotope analytical error...... 47 Table 2-4: Sediment data...... 47 Table 2-5: Hydrate Ridge TAG sequencing results...... 48 Table 2-6: Santa Monica Basin TAG sequencing ...... 48 Table 2-7: Isotope results from Hydrate Ridge ...... 49 Table 2-8: Isotope results from the Santa Monica basin ...... 49 Table 2-9: α and ε values ...... 50 Table 2-10: Model results...... 51 Table 3-1: Labeling Experimental set up...... 89 Table 3-2: Isotope measurement error...... 90 Table 3-3: Hydrate Ridge carbon isotope results...... 90 Table 3-4: Hydrate Ridge TAG sequencing...... 90 Table 3-5: Santa Monica Basin carbon and nitrogen isotope results ...... 91 Table 3-6: Santa Monica TAG sequencing...... 91 Table 3-7: Headspace analysis of PCKD experiments 3 and 4 ...... 92 Table 3-8: Experiment results summary...... 92 Table 3-9: α and ε values ...... 93 Table 4-1: Little Salt Springs cyanobacterium growth conditions ...... 118 Table 4-2: Little Salt Springs cyanobacterium growth conditions ...... 118 Table 4-3: water column, biofilm and cyanobacteria results ...... 119 Table 4-4: experiments results...... 119 Table A-1: F430 concentration data for PC6...... 131 Table A-2: F430 isotope data for PC6...... 132 Table C-1: Solvent profile for first dimension of HPLC chromatography...... 148 Table C-2: Solvent profile for second dimension of HPLC chromatography...... 149 Table D-1: Hydrate Ridge sulfide and sulfate data ...... 150 Table D-2: Hydrate Ridge Aggregate counts, methane and pH data ...... 150 Table D-3: Hydrate Ridge Coenzyme F430 data...... 150

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Table D-4: Hydrate Ridge carbon isotope data...... 151 Table D-5: Santa Monica Basin sulfide, sulfate, ammonium and aggregate counts...... 151 Table D-6: Santa Monica Basin...... 152 Table D-7: Results of Hydrate Ridge Labeling experiments ...... 152 Table D-8: Santa Monica Basin DIC concentration results ...... 153 Table D-9: Santa Monica Basin PCKD δ13C results ...... 154 Table D-10: Santa Monica Basin hydrogen sulfide results ...... 155 Table D-11: Santa Monica Basin experiment 3 and 4 methane results (µM) ...... 156 Table D-12: Santa Monica Basin experiment 3 and 4 methane results (ppm) ...... 156 Table D-13: Santa Monica Basin F430 amounts ...... 156 Table D-14: Santa Monica Basin F430 isotope results ...... 157 Table D-15: Santa Monica Basin Archaeol isotope results ...... 158 Table D-16: Santa Monica Basin newly synthesized F430 amounts ...... 159 Table D-17: Santa Monica Basin newly synthesized Archaeol isotope values ...... 160 Table D-18: BHT and 2-MeAnhydroBHT experiment results ...... 161 Table D-19: Nitrogen experiment results ...... 162 Table D-20: Control experiment time series results ...... 162 Table D-21: 2-Methyl Anhydro bacterialhopanetetrol water column concentration...... 163 Table D-22: BHP concentration in biofilm samples...... 163 Table D-23: Little Salt Springs water column data...... 164 Table D-24: Little Salt Springs water column data...... 164

Acknowledgements

I would like to thank the committee Kate Freeman, Jenn Macalady, Chris House and Squire Booker. I would especially like to thank Kate for helping me develop from a geologist into an isotope/organic geochemist over the past six years. I would also like to thank Victoria Orphan at Caltech who without which the ANME study could not have taken place. Thanks to my former undergraduate advisors Roger Summons, Mark Sephton and Peter Allison who set me off down this path many years ago.

I have had some wonderful lab mates during my time at Penn State who I have learnt a lot from and I would like to thank Sara Lincoln, Heather Graham, Kat Dawson, Heidi Albrecht, Jamie Fulton, Christopher Junium, Colin Carney, and Daniel Jones. I have learnt a lot form Sara Lincoln, Heather Graham, and Kat Dawson over the years and I am eternally grateful for the knowledge, wisdom, and perspective they have provided. I would like to give special thanks to Dennis Walizer who without which much of my lab work could not have been completed and I would have probably been at Penn state well into my nineties.

I would like to thank My Parents Angela and Robert for all their support over these six years and my wonderful girlfriend Sarah Hojjitinia. I thank my friends Moshe Rhodes, Matt Gonzales, Kyle Rybacki, Jamie Brainard Samantha Marquart Brainard, Andrew Chorney, Bradly Guy, Fernando Puente Sánchez and Thomas Jewell who have help me outside of the lab. Usual helping to restore some level of sanity and clarity, which can go a missing after 24 hours in the lab. I cannot put adequately into words what your love and support have meant to me.

I would also like to extend my thanks to those involved with me on the International Geobiology course 2011. This was a fantastic experience that helped me develop as a scientist and an experience I will never forget.

Finally, I would like to thank NASA-Penn State Astrobiology Research Center, American chemical society petroleum Research Fund, Royal Dutch Shell and ConocoPhillips for funding.

Epigraph

Try to learn something about everything and everything about something.

The Right Honorable Thomas Henry Huxley, PRS, FLS, Nature Vol. XLVI p. 658, 1902

Chapter 1: Introduction

1.1. Anoxic methane oxidation Methane, an important fuel for heating, transport and electricity generation, produces less carbon dioxide per energy yield than other fossil fuels (Marland et al., 2003). Since the Kyoto protocol, governments have been exploring policies to encourage the use of natural gas over coal and oil (Apergis and Payne, 2010). Growing demand has stimulated exploitation of unconventional natural gas sources such as methane clathrates, coalbed methane and methanogenic sediments (Administration, 2013, Collett, 2002)

Sedimentary basins along the coast of California and Oregon include numerous sites, among them Hydrate Ridge and the Santa Monica Basin, where natural gas could potentially be explored and produced. In such regions, about half of sedimentary methane is prevented from reaching the atmosphere because it serves as an energy source for anaerobic oxidation of methane (AOM) by Archaea (Knittel and Boetius, 2009). Methane oxidation in these sediments is linked to the reduction of sulfate, nitrate (Haroon et al., 2013), nitrite (Raghoebarsing et al., 2006), iron, or manganese (Beal et al., 2009).

AOM is commonly, but not exclusively, carried out by cell aggregates of Archaeal ANerobic MEthanotrophs (ANME) and sulfate-reducing bacteria (SRB) (figure 1-1) (Boetius et al., 2000, Orphan et al., 2001b). This syntrophic relationship was elegantly documented using fluorescent in situ hybridization with secondary ion mass spectrometry (FISH-SIMS) to trace both 13C and 15N incorporation in natural and enrichment studies into cell biomass (Orphan et al., 2001b, Orphan et al., 2009). Working with natural isotope abundances, numerous studies have illustrated that methane is incorporated into biomass and the biochemical constituents of cells, most notably, membrane lipids (Table 1-1) (Hinrichs et al., 2000, House et al., 2009, Orphan et al., 2001a, Orphan et al., 2001b).

Methanotrophic Archaea comprise three broad phylogenetic lineages: ANME-1 ANME-2 and ANME-3 that are all distantly related to methanogens (Hallam et al., 2003, Lloyd et al., 2006, Orphan et al., 2002). ANME-1 is distantly related to Methanosarcinales and Methanomicrobiales (Hinrichs et al., 1999), while ANME-2 and ANME-3 belong to the Methanosarcinales order (Hallam et al., 2003, Knittel et al., 2005, Lloyd et al., 2006, Orphan et al., 2001a) (figure 1-2). All three groups have been identified and isotopically characterized in sediment from the US western coast (Hydrate Ridge and the Eel River Basin) and Black Sea seeps (Boetius et al., 2000, Orphan et al., 2001b, Reitner et al., 2005, Treude et al., 2007). Although isolation in pure culture for biochemical studies has proven difficult, ANME-1 from the Guaymas basin has been successfully enriched in culture (Holler et al., 2011).

ANME-1, 2 and 3 exhibit a number of distinct characteristics and occupy different ecological niches. ANME-1 cells are rectangular in shape and have been observed as single cells and in monospecific chains or clusters (Knittel et al., 2005, Lösekann et al., 2007, Orphan et al., 2002, Schubert et al., 2006). They are loosely

associated with sulfate-reducing bacteria but have been observed in microbial mats with layers of SRB (Knittel et al., 2005, Lösekann et al., 2007, Orphan et al., 2001a, Treude et al., 2007). ANME-2 and 3 form spherical or shell- like cell aggregates comprised of an ANME core surrounded by SRB (Knittel et al., 2005, Lösekann et al., 2007, Orphan et al., 2002, Schubert et al., 2006, Treude et al., 2007). ANME-1 tend to be more abundant in sulfate- depleted sediments (Yanagawa et al., 2011), hydrothermal environments (Dhillon et al., 2005, Kellermann et al., 2012) and environments with lower oxygen levels, as they are more sensitive to oxygen (Knittel et al., 2005). In contrast, ANME-2 tend to be observed in shallow sediment depths and at higher sulfate concentration (Yanagawa et al., 2011).

1.2. ANME biochemistry A growing body of evidence indicates methanogenesis and methane oxidation take place simultaneously in marine sediments characterized by AOM. Even so, field studies suggest the rate of methane oxidation outpaces methane generation by an order of magnitude or more, as shown by the co-occurrence of methane production and oxidation in Black Sea mats and in sediments from the Cascadia Margin (Treude et al., 2007, Yoshioka et al., 2010). Bertram et al. (2013) recently demonstrated AMNE-1 and, especially, AMNE-2 in enrichment samples (from Black Sea sediments), can co-produce significant amounts of methane simultaneously with methane oxidation, at a production-to-oxidation ratio as high as 1:2. This work also demonstrated that 13C-labeled carbon from C-1 substrates contributed carbon to biomass and membrane lipids (archaeol and hydroxyl-archaeol). Bertram et al., (2013) revealed lipids in the AOM communities preferentially capture acetate and methanol carbon, when available, as well as carbon from bicarbonate. This suggests ANME communities have significant metabolic flexibility, perhaps in response to H2 resources (Bertram et al., 2013), which potentially accounts for the extremely wide range of isotope signatures (~50 ‰) observed for AOM cell clusters in seep settings (House et al., 2009).

Anaerobic methanotrophy is hypothesized to proceed by the reversal of the methanogenesis pathway (Scheller et al., 2010, Zehnder and Brock, 1979). This hypothesis, first proposed by Zehnder and Brock (1979), is supported by culture studies and genetic data (Hallam et al., 2004, Scheller et al., 2010). Hallam et al., (2004) suggest that methane is oxidized to carbon dioxide and reduced by-products, with the assimilation of the reduced products. Alternatively, CO2 assimilation could proceed via the methanogenic pathway, with CO2 incorporated into methylene-tetrahydromethanopterin, which then enters the serine cycle, as in methanogenic Archaea (Angelaccio et al., 2003, Hallam et al., 2004, Taupp et al., 2010). This reaction is catalyzed by serine hydroxymethyltransferase, an enzyme which so far has been reported in all sequenced archaeal genomes, including ANME (Angelaccio et al., 2003).

Culture studies of methanogenic Archaea that can carry out trace oxidation of methane provide supporting evidence for a reversed methanogenesis biochemical pathway. The methanogen Methanosarcina acetovorines was 13 shown to oxidize trace amounts of methane to CO2, as documented by observations that C-labeled methane became incorporated into CO2 (Moran et al., 2005). Studies of Methanothermobacter marburgensis in pure culture

demonstrated the last step in methanogenesis is also the first step in methane oxidation (Scheller et al., 2010). This was documented by the incorporation of 13C-labeled methane into methyl-coenzyme M (2- mercaptoethanesulfonate), which is used in the last step of methane production catalyzed by coenzyme F430 (figure 1-3). Thus, if ANME oxidizes methane by reverse methanogenesis, F430 likely catalyzes the first step.

Genetic evidence from environmental samples provides additional support for reverse methanogenesis AOM. Hallam et al. (2004) found genes that code for the enzymes used in methanogenesis, including for the last step, in ANME-1 and ANME-2 dominated samples from the Eel River Basin. This suggests reverse methanogenesis capability is present among organisms in the sediment, and if the process takes place, signature coenzymes, such as F430, should also be present in the sediment.

Coenzyme F430 is a tetrapyrrole with a nickel center and was first identified by Gunsalus and Wolfe (1978). It is used in the last step of methanogenesis and is likely involved in the first step in the reverse pathway (Hallam et al., 2004, Scheller et al., 2010). Ten modified F430 coenzymes have been identified in methanogens and ANME dominated sediment(Allen et al., 2014, Mayr et al., 2008). These modified F430 may be used in reactions other than methanogenesis and methanotrophy or are adaptations to environmental conditions (Allen et al., 2014). F430 is synthesized from glutamate, which is converted to 5-aminolevulinic acid via glutamyl-tRNA and glutamate- 1-semialdehyde (Friedmann and Thauer, 1986, Gilles et al., 1983, Pfaltz et al., 1987). 5-aminolevulinic acid is then converted to uroporphyrinogen III, the common precursor of tetrapyrroles (Gilles and Thauer, 1983, Pfaltz et al., 1987). Unlike F430, ANME lipids that are synthesized from isoprenoids, are formed from acetyl-CoA via the mevalonate pathway (Goldstein and Brown, 1990, Smit and Mushegian, 2000).

Acetyl-CoA and glutamate could be formed from different sedimentary carbon sources, like dissolved inorganic carbon (DIC) and methane. Potentially this could take place via a different part of the methanogenic pathway operating in different directions. Methane is likely assimilated via a reversal of the last steps of reverse methanogenesis and converted to acetyl-CoA, while DIC may be assimilated via the first steps of methanogenesis, and converted to glutamate. This means that F430 and lipids can be used to test the assimilation of DIC and methane in the sediment due to their synthesis from difference biological precursors. F430 is, therefore, a target for reverse methanogenesis in the sediment and the assimilation of multiple carbon substrates

Chapters 2 and 3 aim to link coenzyme F430 in the sediment to AOM and ANME, something that has not been previously been established (Allen et al., 2014, Mayr et al., 2008). In chapter 2, a link between AOM and F430 in Hydrate Ridge and Santa Monica Basin sediment is established from their concentration profiles. Compound-specific isotope analysis of F430 and lipids reflect likely carbon sources in the sediment. The isotopic heterogeneity observed between lipids and F430 suggests ANME are biochemically flexible and able to assimilate methane carbon into their lipids and carbon from DIC into F430.

Chapter 3 evaluates underlying causes for the isotopic heterogeneity between ANME biomolecules identified in chapter 2 and explores implications for understanding isotopic variability that has been previously observed in House et al. (2009). Stable isotope probing using 13C labeled methane and bicarbonate is used to

explore the assimilation of carbon into F430 and lipids. In Hydrate Ridge sediment, where ANME-1 is more abundant, DIC is shown to be assimilated into lipids with limited production of coenzyme F430. In Santa Monica Basin sediment, where ANME-2 is more abundant, methane and DIC are both assimilated into F430 and lipids. This work has been completed at the Pennsylvania State University under the supervision of Prof. Katherine H. Freeman, in conjunction with Prof. Victoria J. Orphan and Dr. Katherine Dawson at the California Institute of Technology

1.3. Hopanoids Hopanoids (figure 1-4) are a class of pentacyclic compounds first identified in 1969 (Albrecht and Ourisson, 1969) and have been a useful tool in the study of ancient microbial life and the characterization of oil source rocks. Because they are highly resistant to degradation, hopanoids are one of the most common geochemical compounds on the Earth (Ourisson and Albrecht, 1992). Even though hopanoids are present throughout the rock record, the information they provide about the ancient microbial community is limited. Interpretation about the types of ancient microbes are based on the position of a methyl group at the C2 (cyanobacteria) or C3 (methanotrophs and acetogenic bacteria) position (Cvejic et al., 2000, Farrimond et al., 2004, Rohmer et al., 1984, Summons et al., 1999).

2-Methyl hopanoids, found widely in Proterozoic sediments, are conventionally interpreted to represent the presence of ancient cyanobacteria (Summons and Walter, 1990, Summons et al., 1999). This interpretation is based on the high proportion of 2-methyl bacteriohopanepolyols (BHPs) in cultured cyanobacteria and the belief that a cyanobacterial origin can account for the ubiquity of 2-methyl hopanoid across a range of environments and geological ages (Summons et al., 1999, Talbot et al., 2008). Yet, this interpretation was challenged by genetic evidence that less than 10% of all modern bacteria are capable of producing BHPs and all currently known marine cyanobacteria don’t produce 2-methyl BHPs (Pearson et al., 2007, Talbot et al., 2008).

A greater understanding of the function and controls on 2-Methyl BHP production is needed to understand how well 2-Methyl hopanoids serve as a cyanobacteria marker. Analytically this has proved challenging as different BHP structures can have vastly different detection response factors depending on the functional head group (figure 1-5), making quantification challenging. Additionally, culturing studies exploring environmental effects on BHP production and distribution yield different lipid signatures in response to the same test parameters. For example, experiments exploring the effects on N2 fixation using Frankia mycelia, Berry et al. (1993) observed and increase in BHP production, whereas Nalin et al. (2000) observed a decrease.

1.4. Bacteriohopanepolyols BHPs were first identified in 1973 (Förster et al., 1973) in bacteria, and are the biological precursor to geological hopanoids. The BHP structure consists of a C30 triterpenoid pentacyclic hydrocarbon skeleton with a

functional group attached at C22 (figure 1-6) (Talbot et al., 2003). Sixty-three different functional groups have been identified so far. Figure 1-7 illustrates most common forms in cyanobacteria cultures (Talbot et al., 2008). When BHPs are preserved in the rock record, the reactive functional groups are lost, and as a result, interpretations about their sources in past environments are limited to the methyl position.

The function and distribution of BHPs through the bacterial domain is unclear (Fischer and Pearson, 2007). Both gram negative and gram positive bacteria can produce BHPs, but not all bacteria contain the necessary squalene hopene cyclase gene for their production (Pearson and Rusch, 2009, Welander et al., 2010). BHPs aren’t essential for life, even in bacteria that produce them, as demonstrated in knockout gene experiments using Streptomyces and Rhodopseudomona (Seipke and Loria, 2009, Welander et al., 2010). Initially, due to the structural similarity with sterols, it was suggested that they are used to regulate membrane permeability (Kannenberg and Poralla, 1999). Numerous other studies have linked BHP production to membrane function and the physiological status of the bacterial cell (Jahnke et al., 1992, Jahnke et al., 1999, Joyeux et al., 2004, Ourisson et al., 1987, Poralla et al., 1980, Simonin et al., 1996).

BHPs have only been identified in culturable cyanobacteria, methanotrophs, acetic acid bacteria and anaerobic photosynthesizers. While 41 species of cyanobacteria produce BHPs, only 19 of these are able to produce 2-methyl BHPs, the modern precursor of 2-methyl hopanoids, in pure culture (Pearson et al., 2007, Talbot et al., 2008). Further, 2-methyl BHPs have not been observed in modern marine sites with cyanobacteria (Pearson et al., 2007, Talbot et al., 2008). This contradicts the interpretation of 2-methyl hopanoids in Proterozoic marine sediments that are believed to be from a cyanobacterial source. Recently, a 2-methyl BHP producing cyanobacterium in the euxinic waters of a sinkhole, that is chemically analogous to the Proterozoic ocean, has been identified (Hamilton et al., Submitted). Previously, cyanobacteria that produce 2-methyl BHP had only been found in hot springs (Jahnke et al., 2004) and terrestrial soils (Cooke et al., 2008).

The production of BHPs has been explored in a number of culture experiments using different oxygenic phototrophs. BHP production has been shown to vary with a number of different parameters, including temperature, pH, nitrogen species and exposure to ethanol (Berry et al., 1993, Doughty et al., 2009, Poralla et al., 1980, Schmidt et al., 1986). These experiments have yet to identify a reason why modern marine cyanobacteria don’t produce 2- methyl BHPs. Potentially this is due to the limited amount of studies that have quantified BHP structures. Table 1- 2 lists the studies that have quantified BHPs, with only Albrecht (2011), Doughty et al. (2009), and Welander et al. (2009) reporting changes in production in pure culture. Only Albrecht (2011) has fully quantified individual BHP structures, allowing for different structures to be compared against each other. Using the cyanobacteria isolated from Little Salt Springs and with accurate quantification, the geochemical controls on BHP production could be resolved.

The production of BHPs under different geochemical condition is explored using the Little Salt Springs cyanobacteria in chapter 4. Similar amounts of bacteriohopanetetrol (BHT) and 2-methyl anhydro bacteriohopanetetrol (2-MeAnhydro BHT) were identified in the tested geochemical conditions and the control

experiments. The recovered amount of BHT and 2-MeAnhydroBHT were lower than in biofilm samples, with 2- methyl bacteriohopanetetrol and anhydrobacteriohopanetetrol identified in the biofilm not present in the culture experiments. A clear geochemical control on production is not identified and future experiments should explore the effects of oxygen concentration, nitrogen species, trace metals and how combinations of different conditions affect BHP production. This work was completed at the Pennsylvania State University under the supervision of Prof. Katherine H. Freeman and Prof. Jennifer L. Macalady with culture samples supplied by Dr. Trinity Hamilton at the University of Cincinnati.

1.5. Anticipated publications from this work Chapter 2: Carbon Isotopic heterogeneity between ANME biomolecules, will be submitted to Environmental Microbiology with co-authors Jamey M. Fulton, Katherine S. Dawson Victoria J. Orphan and Katherine H. Freeman.

Chapter 3: Stable isotope probing of ANME carbon assimilation, will be submitted to Proceedings of the National Academy of Science with co-authors, Katherine S. Dawson Victoria J. Orphan and Katherine H. Freeman

Chapter 4: Quantifying Bacteriohopanepolyol production in Little Salt Springs cyanobacteria, will be submitted to Organic Geochemistry with co-authors, Trinity Hamilton, Jennifer L. Macalady and Katherine H. Freeman

1.6. Figures and tables

Figure 1-1: FISH image of ANME-2. This image was taken using sediment from the Santa Monica basin, which was used for a natural abundance study in chapter 2 and in incubation experiments using 13C substrates in chapter 3

Figure 1-2: ANME phylogenetic tree. 16S rRNA gene sequences470 tree from Knittel et al. (2005) showing how ANME-1, 2 and 3, in addition to their sub groups are related to each other.

Figure 1-3: Structure of co-enzyme F430. Ten additional F430 based structures have been identified in ANME and in methanogens and are believed to be used in functions other than methanogenesis (Allen et al., 2014).

Figure 1-4: Hopene. Also known as diploptene that has been observed in the rock record.

9.E+08 2-methyl anhydro bacteriohopanetetrol bacteriohopanetetrol 8.E+08 pregandiol Linear (2-methyl anhydro bacteriohopanetetrol) 7.E+08 Linear (bacteriohopanetetrol ) 6.E+08 Linear (pregandiol)

5.E+08

4.E+08

3.E+08 Responce on LCMS on Responce

2.E+08

1.E+08

0.E+00 0 500 1000 1500 2000 ng injected

Figure 1-5: LCMS response to BHP structures. The response of 2-MeAnhydroBHT, BHT and pregenanediol used as a standard in the quantification of BHP compounds. Differences in the response of the two BHP compounds are due to the different polar head groups.

Figure 1-6: Red biofilm Bacteriohopanepolyols. These structure were identified in the cyanobacterial dominated biofilm from Little Salt Springs that is analyzed in chapter 4

Figure 1-7: BHP polar groups. Potential cyanobacterial BHP polar head groups as identified in Talbot et al. (2008). Quantifying numerus BHP structures is difficult as these different polar groups produce different responses

Table 1-1: ANME isotope values. Isotope values reported in previous studies of ANME at anoxic methanotrophic sites

Location Compound δ 13C, ‰ Source Reference Eel River Archaeol -104.1 ANME-2 (Orphan et al., 2001b) Eel River Hydroxyarchaeol -107.6 ANME-2 (Orphan et al., 2001b) Eel River Cell cluster -96 ANME-2 (Orphan et al., 2001b) Eel River Archaeol -101.1 ANME-1/2 (Orphan et al., 2001a) Eel River Archaeol -100.6 ANME-1/2 (Orphan et al., 2001a) Eel River Archaeol -102.6 ANME-1/2 (Orphan et al., 2001a) Eel River Archaeol -102.1 ANME-1/2 (Orphan et al., 2001a) Eel River Hydroxyarchaeol -105.2 ANME-1/2 (Orphan et al., 2001a) Eel River Hydroxyarchaeol -105.8 ANME-1/2 (Orphan et al., 2001a) Eel River Hydroxyarchaeol -105.5 ANME-1/2 (Orphan et al., 2001a) Eel River Hydroxyarchaeol -105.7 ANME-1/2 (Orphan et al., 2001a) Eel River Archaeol -100 - (Hinrichs et al., 2000) Eel River hydroxyarchaeol -106 - (Hinrichs et al., 2000) Eel River Cells -24 to -87 ANME-1 (House et al., 2009) Eel River Cells -18 to -75 ANME-2 (House et al., 2009) Hydrate Ridge Archaeol -114 ANME-1 (Boetius et al., 2000) Hydrate Ridge Hydroxyarchaeol -133 ANME-1 (Boetius et al., 2000) Santa Barbra Basin Archaeol -119 - (Hinrichs et al., 2000) Santa Barbra Basin Hydroxyarchaeol -128 - (Hinrichs et al., 2000) Mediterranean mud Archaeol -76.2 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -40.6 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -63.1 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -84.1 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -81.1 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -57.2 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -95.8 - (Pancost et al., 2000) volcanoes Mediterranean mud Archaeol -89 ANME-1 (Aloisi et al., 2002) volcanoes Mediterranean mud Archaeol -97 ANME-1 (Aloisi et al., 2002) volcanoes Mediterranean mud Hydroxyarchaeol -90 ANME-1 (Aloisi et al., 2002) volcanoes Mediterranean mud Hydroxyarchaeol -97 ANME-1 (Aloisi et al., 2002) volcanoes Twentekanaal Hydroxyarchaeol -67 ANME-2 (Raghoebarsing et al., Netherlands 2006)

Black Sea Archaeol -95.6 ANME-1 (Reitner et al., 2005)

Black Sea Archaeol -87.9 - (Michaelis et al., 2002)

Black Sea Hydroxyarchaeol -90 - (Michaelis et al., 2002)

Black Sea Mat -66.4 ANME-1 (Treude et al., 2007)

Black Sea Mat -72.9 ANME-2 (Treude et al., 2007)

Table 1-2: BHT quantification. Other BHP structures are reported in these studies, but BHT is the only one present in all, allowing comparison between the studies.

Sample µg/g TLE BHP Quantification Reference River 564 BHT Quantitative (Sáenz et al., 2011) River 293 BHT Quantitative (Sáenz et al., 2011) Estuary 318 BHT Quantitative (Sáenz et al., 2011) Green Water 191 BHT Quantitative (Sáenz et al., 2011) Blue water 81 BHT Quantitative (Sáenz et al., 2011) Blue water 98 BHT Quantitative (Sáenz et al., 2011) Pigeon creek sediment 25000 BHT Semi-quantitative (Pearson et al., 2009) Grahams Harbour Sediment 30000 BHT Semi-quantitative (Pearson et al., 2009) R. palustris Chemohetertrophic 3400 BHT Semi-quantitative (Welander et al., Exponential 2009) R. palustris Chemohetertrophic 3000 BHT Semi-quantitative (Welander et al., Stationary 2009) R. palustris Photoheterotrophic 10000 BHT Semi-quantitative (Welander et al., Exponential 2009) R. palustris Photoheterotrophic 8000 BHT Semi-quantitative (Welander et al., Stationary 2009) R. palustris pH5 2000 BHT Semi-quantitative (Welander et al., 2009) R. palustris pH7 3000 BHT Semi-quantitative (Welander et al., 2009) R. palustris pH9 2000 BHT Semi-quantitative (Welander et al., 2009) L. ferrooxidans N source 692 BHT Quantitative (Albrecht, 2011) L. ferrooxidans without N 6934 BHT Quantitative (Albrecht, 2011) source L. ferrooxidans N source 3585 BHT Quantitative (Albrecht, 2011) A.variablis photosynthetic 900 BHT Quantitative (Albrecht, 2011) A.variablis photosynthetic 5500 BHT Quantitative (Albrecht, 2011) A.variablis chemoheterotrophic 4 BHT Quantitative (Albrecht, 2011) A.variablis chemoheterotrophic 400 BHT Quantitative (Albrecht, 2011) Peat sample 10 BHT Quantitative (van Winden et al., 2012)

Peat sample 60 BHT Quantitative (van Winden et al., 2012) N. punctiforme 500 BHT Semi-quantitative (Doughty et al., 2009) N. punctiforme 2200 BHT Semi-quantitative (Doughty et al., 2009) Microbial mat 6664 BHT Quantitative (Blumenberg et al., 2006) Sediment 4920 BHT Quantitative (Blumenberg et al., 2006) Sediment 2331 BHT Quantitative (Blumenberg et al., 2006) Oxic zone 40 BHT Quantitative (Rush et al., 2014) Transition zone 200 BHT Quantitative (Rush et al., 2014) Transition zone 150 BHT Quantitative (Rush et al., 2014) anoxic zone 600 BHT Quantitative (Rush et al., 2014) anoxic zone 100 BHT Quantitative (Rush et al., 2014)

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Chapter 2: Carbon isotopic heterogeneity between ANME biomolecules

2.1. Abstract Microbially mediated anaerobic oxidation of methane (AOM) is an important sink for methane, a potent greenhouse gas. A group of Archaeal ANerobic MEthanotrophs (ANME) facilitate oxidation and is hypothesized to proceed via the reversal of the methanogenesis biochemical pathway (Scheller et al., 2010; Zehnder & Brock 1979). Both natural isotope abundance studies and 13C-labeling experiments have provided insight into the biochemistry of methane oxidation and assimilation (House et al., 2009, Orphan et al., 2001b). Isotope studies and geochemical profiles indicate ANME are metabolically diverse and possess the enzymatic machinery to assimilate carbon via more than one pathway. This plasticity would explain the ~50‰ range in archaeal lipids from ANME dominated sediment that has been observed in the Eel River basin (House et al., 2009, Orphan et al., 2001b). F430 is a tetrapyrrole used in the last step of methanogenesis, and likely enables the first step in reverse methanogenesis (Hallam et al., 2004, Scheller et al., 2010). Therefore, the presence and concentration of F430 in association with AOM serves as a test for the reverse methanogenesis pathway in sediment. Tetrapyrroles and archaeol lipids are formed from different biological precursors (glutamate and acetyl-CoA, respectively (Gilles et al., 1983, Koga and Morii, 2007, Pfaltz et al., 1987)), which could be formed from different sedimentary carbon sources. Isotopic analysis of tetrapyrroles, like F430, and archaeol lipids could be used to determine the assimilation of multiple carbon sources.

In sediment from Hydrate Ridge and the Santa Monica Basin (west coast USA), a link between F430 and AOM is established. The greatest amounts of F430 were recovered where sulfide, sulfate, and methane concentration profiles indicate the greatest AOM activity in the sediment. These sediment horizons also contained the highest ANME-2 aggregate counts. F430 was found to be isotopically distinct from methane and archaeal lipids, but similar to DIC. The ability of ANME to assimilate multiple carbon sources may explain the wide range of isotope signatures (~50‰) measured in these different compounds, as well as more generally among AOM cell clusters in seep settings (House et al., 2009, Orphan et al., 2002). We hypothesize that physiologic versatility also drives the observed carbon-isotopic differences between archaeal lipids and F430.

2.2. Introduction Microbial anaerobic oxidation of methane (AOM) is an important process that limits the release of methane from marine sediments. AOM consumes up to 80% of the sedimentary methane flux each year, which prevents this potent greenhouse gas from reaching the atmosphere (Orphan et al., 2001b). A group of Archaeal ANerobic MEthanotrophs (ANME) facilitate oxidation, either in cell aggregates with sulfate reducing bacteria or as single

cells (Boetius et al., 2000, Hinrichs et al., 1999). Methane oxidation is coupled to the reduction of sulfate, nitrate (Haroon et al., 2013), nitrite (Raghoebarsing et al., 2006), iron, or manganese (Beal et al., 2009).

Both natural isotope abundance studies and 13C-labeling experiments have been used to study ANME assimilate carbon (Hinrichs et al., 2000b, Orphan et al., 2002, Orphan et al., 2001a, Orphan et al., 2001b). ANME assimilate methane carbon into their biomass (House et al., 2009, Orphan et al., 2001b) and into membrane lipids (Hinrichs et al., 2000b, Pancost et al., 2000), which results in both being depleted in 13C in natural environments. Yet, a remarkably wide range in carbon isotope signatures (~50‰) for ANME cell clusters is observed in many seep settings (House et al., 2009, Orphan et al., 2001b). What drives this range in carbon isotope values remains unknown, but the pattern suggests ANME can assimilate multiple carbon sources (Bertram et al., 2013, Kellermann et al., 2012, Wegener et al., 2008).

13C-labeling studies and genetic profiling indicate ANME are metabolically diverse and possess the enzymatic machinery to assimilate carbon via more than one pathway. Bertram et al. (2013) found AOM communities preferentially incorporated acetate and methanol carbon into lipids. Incorporation of dissolved inorganic carbon (DIC) into lipids and biomass has also been shown by Bertram et al. (2013), Kellermann et al. (2012) and Wegener et al. (2008). These studies show that ANME can switch carbon substrate, and are not limited to methane. This plasticity would explain the ~50‰ range in archaeal lipids from ANME dominated sediment that has been observed in the Eel River basin (House et al., 2009, Orphan et al., 2001b).

ANME lipids and tetrapyrroles isotope signatures can help determine their carbon sources, which could shed light on carbon budgets for both cellular biomass and individual components. These compound classes are formed from two different biochemical precursors and pathways, glutamate is the precursor for tetrapyrroles (Gilles et al., 1983, Pfaltz et al., 1987) and acetyl-CoA is the precursor for archaeal lipids like archaeol and hydroxyl- archeaol (Koga and Morii, 2007). Glutamate and acetyl-CoA could derive from different carbon substrates, such as DIC and methane. These substrates could be assimilated via different parts of the methanogenic pathway operating in reverse and forward directions. If so, then the carbon isotopic compositions of tetrapyrroles and lipids should reflect their respective carbon sources and biochemical origins in the sediment. In sediment from Hydrate Ridge and the Santa Monica basin, methane (-70‰ to -62‰) and DIC (-12‰ to -50‰) are both isotopically distinct from each other.

Both ANME-1 and 2 contain all but one of the genes for the methanogenic pathway (Hallam et al., 2004). If methane is oxidized and assimilated via the reversal of methanogenesis, then the coenzyme F430 would be produced by ANME for use in the first step of reverse methanogenesis. Coenzyme F430 is a tetrapyrrole used in the active site of methyl coenzyme M reductase to catalyze the last reaction step, where a methyl group is removed from coenzyme M (CH3-S-CoM) and combined with a hydrogen from coenzyme B (HS-CoB), forming methane

(equation 1) (Scheller et al., 2010, Thauer, 1998). This step is the most energy intensive (ΔG°ʹ = –30 ±10 kJ mol-1 (Scheller et al., 2010)) and therefore acts as a control on the rest of the reverse methanogenic steps.

CH3-S-CoM + HS-CoB ⇌ CH4 + CoM-S-S-CoB (1)

The presence of F430 in association with evidence for AOM provides a test of reverse methanogenesis in the sediment (Allen et al., 2014, Mayr et al., 2008). F430 concentrations peaking where sulfide, sulfate, and methane profiles indicate the greatest amount of AOM activity, would support the use of F430 in AOM. The carbon isotopic composition of F430 and its concentration in relation to ANME in sediments has not been previously measured. By modifying a previously developed method (Mayr et al., 2008) and using nano-scale elemental analysis isotope ratio mass spectrometry (nano-EA/IRMS) (Polissar et al., 2009), concentration and carbon and nitrogen isotope measurements can be made on F430.

We also targeted F430 and ANME lipids extracted from marine sediments for carbon isotope analysis. We show that F430 13C abundance is distinct from that of membrane lipids and methane in ANME-dominated sediment from Hydrate Ridge and the Santa Monica Basin. Using mass balance with fractionation calculations, we estimate the amounts of carbon substrates, such as methane and DIC, that could contribute to the observed isotope values of F430 and ANME lipids.

2.3. Materials and Methods

2.3.1. Shipboard collection, core processing, and sample storage Samples were collected using the ROV Jason II by scientists aboard the R/V Atlantis from Hydrate Ridge, Oregon (cruise 18-10) in September 2011 and from the Santa Monica Basin, California (cruise 26-06) in October 2013 (figure 2-1). Sediment push cores were collected from methane seep environments characterized by either the presence of chemosynthetic clam beds (PC28) or microbial mats (PC4). Core PC28 (Hydrate Ridge 44°N 40.19 125°W 5.88) was recovered from a chemosynthetic clam bed with live Calyptogena clams present in the 0-3 and 3-6 cm horizons. Core PC28 was sectioned in 3 cm intervals immediately after recovery, and stored at 20°C prior to being shipped to Penn State. Core PC4 (Santa Monica Basin 33°N 38.403 118°W 48.025) was penetrated through an orange microbial mat. It was sectioned immediately after recovery into 1 cm intervals for the first 6 cm and then in 3 cm section to a depth of 15 cm. Core sections were stored at 20°C prior to being shipped to Penn State.

Parallel cores taken at both locations (PC20 and PC23 – Hydrate Ridge, OR; PC6 – Santa Monica Basin, CA) were processed for DNA, microscopy, lipid, and pore water chemistry analyses. Immediately after the parallel push cores were sectioned, aliquots of sediment were stored at -80°C for DNA extraction, or were fixed for microscopy by adding 4% paraformaldehyde (PFA) to sediment-seawater mixtures in a 1:1 ratio and incubating at 4°C for 12 hours. Fixed samples were washed with 3x phosphate buffered saline (PBS) and stored in 1:1 3x PBS:ethanol at -20°C. Pore water was collected from 1 to 3 cm sediment intervals under Ar using a pressurized gas sediment squeezer (KC Denmark A/S, Silkeborg, Denmark) (Reeburgh, 1967), and residual sediments were then

stored at -80°C for lipid analysis. Pore water geochemical analysis included anion, cation, and sulfide concentrations, as well as dissolved inorganic carbon (DIC) concentration and stable isotope analysis (see below for details).

2.3.2. DNA extraction and tag sequencing Sediment was stored at -80°C until DNA extraction. DNA was extracted using a MoBio Ultraclean soil kit (MO BIO Laboratories Inc., Carlsbad, CA, USA). Preparation for sequencing of the V4 region of the 16S rRNA gene was carried out according to the Earth Microbiome Project protocol (Caporaso et al., 2012, Caporaso et al., 2011) with modifications as previously described (Case et al., 2015). Raw sequences were generated on an Illumina MiSeq platform at Laragen, Inc. (Los Angeles, CA, USA) and are available in the Sequence Read Archive (PRJNA350854). Sequence data were demultiplexed and processed using a modified version of the QIIME pipeline (Caporaso et al., 2010) as described previously (Mason et al., 2015). Prior to sample comparison, singletons and PCR contaminants were removed, and a 0.01% relative abundance threshold was applied.

2.3.3. Geochemical analysis of pore waters Sulfide dissolved in sediment pore waters was preserved by the immediate precipitation as ZnS through the addition of 0.5M Zn-acetate in a 1:1 ratio with water samples. Concentrations were then determined + + spectrophotometrically by the Cline assay (Cline, 1969). Water samples for anion and cation analysis (Na , NH4 , + 2+ 2+ - - - 2- K , Mg , Ca , formate, acetate, Cl , Br , NO3 , SO4 ) were filtered through a 0.2 µm polyethersulfone (PES) syringe filter and stored at -20°C. After thawing, aliquots were diluted 1:20 with MQ water, and subsequently were analyzed on a dual channel Dionex ICS-2000 ion chromatography system in the Caltech Environmental Analysis Center. Water samples were split and simultaneously separated with cation and anion exchange columns at 0.25 ml min-1 and 30°C. Cations were separated isocratically with 20 mM methanesulfonic acid using an IonPac CS12A column and guard column, and anions were separated isocratically with 20 mM KOH using an IonPac AG19 column and guard column.

Water samples for total DIC measurements were filtered through a 0.2 µm PES filter into He flushed, 12 ml exetainer vials (Labco Ltd, Lampeter, UK) that had been pre-weighed after the addition of 100 µl ~40% phosphoric acid. Samples were stored upright at room temperature. Vials were sampled using a GC-PAL autosampler (CTC Analytics, Zwingen, Switzerland) equipped with a double-holed needle that transferred headspace using a 0.5 ml min-1 continuous flow of He to a 50 µm sample loop prior to separation by a PoraPlotQ fused silica column (25m; i.d. 0.32 mm) at 72°C. CO2 was then introduced to a Delta V Plus IRMS using a ConFlo IV interface (Thermo

Scientific, Bremen, Germany) in the Caltech Stable Isotope Facility. A sample run consisted of 3 reference CO2 gas peaks, 10 replicate sample injections, and 2 final reference CO2 peaks. A concentrated solution of NaHCO3 was used to establish a standard curve for concentration determination by adding a range of volumes to additional exetainer vials, which were interspersed with samples. The concentration of DIC (µM) in samples was determined

2 by comparing the average of the combined mass 44, 45, and 46 CO2 peak areas to the standard curve (n = 20, R = 0.99), was calculated after determining sample volume by re-weighing exetainers. δ13C values were corrected for sample-size dependency and then normalized to the VPDB scale with a two-point calibration (Coplen et al., 2006) using NBS-19 and a previously calibrated laboratory carbonate as internal standards. Accuracy (0.11‰, n=79) was determined by analyzing independent standards as samples and precision (0.42‰, n=10) was determined from NBS- 19.

DIC samples taken for Hydrate Ridge had degassed during storage resulting in the loss of depleted CO2.

Therefore, CO2 from carbonate mineral phases, isolated by acidification of ~20mg of sediment was used to determine carbonate carbon isotope values could be obtained for the Hydrate Ridge sediments. Isotope values were obtained following the method described above for water sample DIC values. The precipitated carbonate δ 13C values were then converted to CO2(g) using the εar-CO2 value for mineral aragonite at 4℃ (2) from Romanek et al. (1992) because aragonite is the dominant carbonate mineral in Hydrate Ridge sediments (Joseph et al., 2013). δ13C values for CO2(aq), DIC, bicarbonate and carbonate ions were then calculated at 4℃ using equations for εaq-CO2, (3)

εHCO3-CO2 (4) and εCO3-CO2 (5). εDIC-CO2 for 5.3℃ used from Zhang et al. (1995) as data on the fCO3 is unavailable for the sediment. Results for Hydrate Ridge are reported in table 2-1 and for the Santa Monica Basin in table 2-2.

εar-CO2 = 13.88 – 0.13 T (℃) (2)

εaq-CO2= 0.013 T (℃) + -2.31 (3)

εHCO3-CO2 = -0.1141 T (℃) + 10.78 (4)

εar-CO2 = -0.052 T (℃) + 7.22 (5)

2.3.4. F430 extraction and separation Extraction and isolation of coenzyme F430 followed the method of Mayr et al. (2008), with additional purification steps to allow for quantification and isotope analysis. Approximately ~30 g of Hydrate Ridge wet sediment and ~10 g of Santa Monica wet sediment were needed to quantify, isolate, and make an isotope measurement on coenzyme F430. Sediment samples were agitated by ultra-sonication probe for 20 minutes in neutral (pH 7) 18.2 W water, and held in an ice bath to keep the temperature at 4OC. Sediment was separated from the extract by centrifugation at 5000 g for 15 minutes. The sediments were extracted twice more in 18.2 W water

adjusted to pH 3 using 0.1% formic acid. The three extracts were combined and neutralized to pH 7.2 using NaOH, in order to precipitate proteins, which were separated and removed by centrifuging the solution at 9000 g for 10 minutes.

Coenzyme F430 was separated from the protein-free supernatant using two-dimensional column chromatography. First, the supernatant was applied to a QAE Sephadex A25 column (1.5cm x 10cm) that had equilibrated with 50 nM Tris/HCl (pH 7.5). After the column was flushed with 4 dead volumes of Tris/HCl, the F430-containing fraction was eluted with 90 ml of 20 nM formic acid. This fraction was then applied to a XAD Amberlite column (1cm x 10cm) which had been flushed with two dead volumes of 10 nM formic acid. The F430 fraction was eluted in 10 ml of 100% methanol. This fraction was dried under nitrogen and stored at -20OC before being further purified via high pressure liquid chromatography (HPLC).

High-pressure liquid chromatography (HPLC) was used to purify F430 sufficiently to enable quantification and isotope analysis. The first HPLC separation employed two Waters spherisorb ODS2 columns (5 ㎛, 4.6 mm x 150 mm) linked together and supplemented by a Phenomenex C18 (3 mm x 4 mm) guard cartridge. Mobile phase A consisted of HPLC-grade water, mobile phase B of 0.1% formic acid and mobile phase C of acetonitrile (HPLC grade). At a flow of 0.5 ml/min, the following gradient was applied: 0 minutes 0% A, 70% B 30%C; 2 minutes 0% A, 70% B 30%C; 4 minutes 0% A 50%B 50%C; 20 minutes 50% A, 0% B, 50% C; 25 minutes 25% A, 0% B, 75% C; 28 minutes 25% A, 0% B, 75% C; 30 minutes 0% A, 70% B, 30% C. The F430 peak eluted at 25 minutes and was collected over a 1.5 to 2 minute window based on the UV/vis detector response at 430 nm. Fractions were dried under nitrogen and re-dissolved in methanol for additional purification.

F430 was separated from a co-eluting molecule that was contributing addition carbon in the Nano EA- IRMS analysis using a Thermo Hypercarb column (5㎛, 100mm x 4.6mm). Mobile phase A consisted of HPLC- grade water, mobile phase B of 0.1% HCl and mobile phase C of acetonitrile (HPLC grade). At a flow of 0.5ml/min the following gradient was applied: 0 minutes 0% A, 70% B 30%C; 2 minutes 0% A, 70% B 30%C; 4 minutes 0% A, 50% B, 50%C; 18 minutes 25% A, 50% B 25%C; 20 minutes 50% A, 0% B, 50% C; 25 minutes 25% A, 0% B, 75% C; 28 minutes 25% A, 0% B, 75% C; 30 minutes 0% A, 70% B, 30% C. Quantification and identification were performed on the first run of sample through the Hypercarb column (figure 2-2), and subsequent runs were collected for nano-EA/IRMS analysis. F430 was identified by UV/vis detection of absorbance at 430 nm (figure 2-3) and confirmed by the m/z 905 ion of the complete F430 structure (figure 2-4). A previously published molar extinction coefficient of 21000 M-1 cm-1 was used to quantify F430 (Ellefson et al., 1982, Whitman and Wolfe, 1980). An Agilent 6300 ion trap with an ESI source was used for mass spectral analysis. Fractions for nano-EA/IRMS were collected at 8 minutes for 20 seconds. Samples were then dried under nitrogen and transferred to Costech tin boats using methanol. Samples were covered and left to dry before loading into autosampler for isotope analysis.

2.3.5. Isotope analyses of F430 Quantities of F430 isolated from environmental samples are typically too small for conventional EA-IRMS (elemental analyzer - isotope ratio mass spectrometry). Instead, we used a nano-scale EA/IRMS technique, developed by Polissar et al. (2009). In this method, the combusted sample is concentrated by cryogenic capture, transferred by a low flow of helium through a capillary gas chromatograph column (J&W scientific GS-

CarbonPLOT 30 m 0.32 mm 1.5 µm) to separate N2 and CO2 peaks before isotope analysis by the IRMS (Thermo- Finnigan Delta Plus). Isotope values for samples at natural abundance are reported in the delta notation (equation 2, in units of permil, ‰), after characterization of standards and accounting for analytical blanks (Polissar et al., 2009).

� = (6)

� = ((� − � )/ �) (7)

Samples are first corrected for background contribution, which are generally sourced from the tin boats (Costech), helium carrier gas, and oxygen combustion gas. These “blank” contributions were determined from multiple measurements of empty tin cups, and by a isotope mass balance (Equation 8), where nblank is blank peak area, nobs is the observed sample peak area, δblank is blank δ values and δobs is the observed δ of the sample.

�� = �� + �� (8)

This can be rearranged to the useful form of a linear relationship:

� = � − �(� − �)/(1/�) (9)

With the true isotope value being the intercept and determined using equation 10:

� = (�� − ��)/(� − �) (10)

The intercept of the relationship is the blank-corrected delta value for the sample, which is calculated using the peak areas of the blank and sample peaks (nblank, nobs), and the observed delta values for the blank and sample analyses (δblank, δobs)(equation 9). Plotting δobs against 1/nobs illustrate the mixing relationship between blank and analayte for both carbon and nitrogen isotope data sets and that δx (the intercept) is the true value (figures 2-5 and 2- 6).

Analytical accuracy was assessed using a suite of isotopic standards, which are analyzed with the samples, and similarly corrected for blank contributions. Three standards, octaethylporphine (Frontier Scientific δ13C - 34.05‰, d15N -12.23‰), L-methionine (Sigma Aldrich δ13C -30.45‰, d15N 0.46‰) and Sucrose (NIST δ13C - 10.45‰) were run in conjunction with the samples to determine isotopic offset. The measured and corrected standard values were regressed against their known values, and the resulting linear equation was used to correct unknown samples for any offset:

� = � �� + �� (11)

Errors were propagated using the sum of squares method, which assumes uncertainties for each correction step are both independent and random (Polissar et al., 2009). For this calculation, uncertainties (σ) for the blank cups (equation 12), standards (equation 13) and offset for all standards (equations 14 and 15) were all used to determine total analytical uncertainty (equation 16)

1 � = (� − � )^2 (12) �

1 � = (� − � )^2 (13) �

� = � − � (14)

1 � = (� − � )^2 (15) �

(�∑) = (�) + (�) + (�) + (�) + (�) (16)

Blank correction uncertainty was simplified and represented by the analytical reproducibility of the blank delta values. Uncertainty in blank analyses were determined from n = 6 analyses of empty cups for each analysis batch. Blank correction uncertainty for Hydrate Ridge was 1‰, and 0.4‰ for the Santa Monica Basin.

Reproducibility in carbon measurements were determined from the standard deviation of blank-corrected octaethylporphine, methionine and sucrose values. Uncertainty in the offset correction was determined from the sum of squares of the offset values determined from equation 15 for each standard. The total analytical uncertainty propagated using the sum of squares (equation 16) for the Hydrate Ridge experiments was 2.9‰, and 1.5‰ for the Santa Monica Basin samples (Table 2-3).

In addition, we checked accuracy and reproducibility for carbon measurements using the methionine standard which was removed from the carbon correction and treated as if an unknown. The difference between the observed, and corrected methionine standard value and the known value was 0.3‰ for Hydrate Ridge, and 0.6‰ for Santa Monica Basin analyses.

2.3.6. Lipid Analysis Sediment samples were extracted using a modified Bligh-Dyer (Bligh and Dyer, 1959), with the first extraction using 250 ml of 4:10:5 water:methanol:dichloromethane followed by two extractions with 4:10:5 2.5% trichloacetic acid:methanol: dichloromethane. Samples were disrupted with a sonicator probe for 20 minutes then centrifuged at 5000 g for 15 minutes. Phase separation of the supernatant was induced with 20 ml of dichloromethane and 20 ml of water, with the organic phase removed. The aqueous phase was extracted with the addition of 20 ml of dichloromethane two additional times, and the pooled organic phases were dried under N2. Cold acetone precipitation was then used to separate the Intact polar lipids (IPL) from the neutral lipids. This was done by bringing the sample up in 2 ml of DCM and then adding 40 ml of ice cold acetone and then storing at -20℃ for 24 hours. The precipitated intact polar lipids were separated by centrifuging at 9000 g for 15 minutes. The separated IPL fraction was treated with acidified methanol for 12 hours and then derivatized with BSTFA and pyridine.

Derivatized samples were analyzed on a gas chromatography mass spectrometer (Thermo trace 1310) to identify lipids in the IPL fraction. Samples were injected using splitless injection on to a fused silica column (Resteck DB-5, 30 m 0.25 mm, 25 µm) with the column heated from 60℃ to 320℃ at 6℃ per minute. Archaeol

and hydroxy-archaeol were identified by comparing to published spectra (Hinrichs et al., 1999, Hinrichs et al., 2000a).

Archaeol and hydroxy-archaeol δ13C were determined by gas chromatography isotope ratio mass spectrometry (GC-IRMS). Samples were separated on a Varian 3400 GC with splitless injection onto a fused silica column (Restek DB-5, 30 m, 0.25 mm, 25 µm). Once separated compounds were combusted to carbon dioxide with a nickel platinum catalyst with 1% oxygen in helium at 1020℃. Stable isotope ratios were measured with a Finnegan Mat 252 with isotope values reported in the delta notation relative to the VPDM scale (equation 3). Isotopic offset was corrected for using Mix B standard (n-C16 to n-C30 alkanes, Arndt Schimmelmann, Indiana University) with standards of n-C38 and n-C41 treated as unknowns to determine error. Precision and accuracy were 0.9‰ and 0.7‰ for Hydrate ridge (n=6) and 0.5‰ and 0.4‰ for Santa Monica Basin samples (n=4).

BSTFA adds three methyl groups to the hydroxyl groups on archaeol and hydroxy-archaeol. The values of these methyl groups was determined by derivatizing, Phatalic acid standard (δ13C -27.21‰ Sigma Aldrich, isotope value confirmed via Arndt Schimmelmann, Indiana University) and using mass balance to determine the isotope value of the BSTFA groups (equations 17 and 18). With the value of the BSTFA groups the true value of the lipids can be determined from the measured value (equation 19). The BSTFA groups have a δ13C -39.75‰ and a standard deviation of 0.5‰ (n=3). Error for the lipids is a combination of accuracy and precision determined from the standards and the standard deviation in the BSTFA group measurement (equation 20). Total analytical uncertainty is 1.3‰ for Hydrate Ridge and 0.8‰ for Santa Monica Basin lipids

�∑ = δ� + �(1 − � ) (17)

� = �∑ − �/(1 − � ) (18)

� = � − ��/(1 − �) (19)

(�∑ ) = (�) + (�) + (�) (20)

2.4. Results At both study sites, trends and maxima in F430 concentrations, ANME aggregate counts, and geochemical profiles reveal a strong link between F430 and methane oxidation (figures 2-7, 2-8). In the Hydrate Ridge sediment,

F430 concentrations peaked at 86 µg /g in the 3-6 cm core section, coinciding with the largest ANME aggregate count (figure 2-7). This section of the core corresponds with maximum rate of methane oxidation, based on the concentration profiles of methane, sulfate and sulfide. Above and below the 3-6 cm section, F430 concentrations dropped to 10 µg/g in 0-3 cm, 40 µg/g in 6-9 cm and 5 µg/g in the 9-12 cm sections (table 2-4). Illumina TAG sequencing results for Hydrate Ridge show that ANME-1 were more abundant than in the Santa Monica Basin core (table 2-5 and 2-6).

Both F430 concentrations and ANME cell counts were higher in the Santa Monica sediment than Hydrate Ridge. The top 2 cm of the Santa Monica core contained 1616 µg/g of F430, more than was recovered throughout the complete Hydrate Ridge core. This section of the core also corresponds to the highest ANME aggregate counts and the maximum rate of methane oxidation inferred based on the profiles of sulfate and sulfide (figure 2-8). F430 concentrations and ANME aggregate counts decreased with depth and a secondary, but minor maximum was observed at 4-5 cm. Concentrations of F430 decreases with greater depths reaching 1 µg/g in the 12-15 cm section (Table 2-4). Beneath this depth, concentrations were below the level of detection. Based on the TAG sequencing data, ANME-2c was the most abundant ANME form, with the greatest number of sequences in the top 2 cm. ANME-2a and 2b were also present in the sediment, with ANME-1a and 1b abundances lower than at Hydrate Ridge (table-2-3).

DIC and methane typically exhibit very distinct δ13C values in marine sediments. At Hydrate Ridge, methane δ13C values ranged -62.4‰ to -70.2‰, indicating a mix of biogenic and thermogenic sources (Nittrouer et al., 2009). Carbonate, a proxy for DIC, is more enriched in 13C, varying from -42.6‰ in the 0-3 cm section to - 16.1‰ in the 12-15 cm section. Methane δ13C values for Santa Monica sediment reported by Stolper et al. (2015) range from -66.7‰ to -70.4‰, and DIC was more enriched in 13C, and ranged from -44.3‰ to -50‰.

Archaeol and sn-2-hydroxy-archaeol have previously been recovered from numerous methanotrophic sediments and are widely attributed to ANME (Boetius et al., 2000, Elvert et al., 1999, Hinrichs et al., 1999, Hinrichs et al., 2000b). Intact polar lipid (IPL) archaeol from the Hydrate Ridge core was depleted in 13C (-89‰ and -128‰). Values obtained for hydroxy-archaeol from different Hydrate Ridge cores were also depleted, and ranged from -61‰ to -90‰ (Figure 2-9). Santa Monica IPL-archaeol (-127‰ to -119‰) and IPL-hydroxy-archaeol (-111‰ to -129‰) were also depleted in 13C relative to DIC and methane (figure 2-10). These values are similar to those previously reported for archaeal lipids from the Eel River Basin (Hinrichs et al., 1999, Orphan et al., 2001b), Hydrate Ridge (Boetius et al., 2000), and Black Sea (Treude et al., 2007). The depleted 13C isotope value confirms that mostly carbon from methane has been assimilated into the ANME lipids.

Coenzyme F430 was significantly more enriched in 13C than the lipids (figures 2-9 and 2-10). At Hydrate Ridge, F430 ranged from -25‰ to -21‰ indicating little or no carbon assimilated from methane and that it is likely synthesized from DIC (table 2-7). In the Santa Monica Basin sediment, F430 isotope values range from -67‰ to - 89‰ (Table 2-8) and are more variable and generally lower than at Hydrate Ridge. These data indicate that carbon from both methane and DIC may be incorporated into F430 by ANME, particularly in the Santa Monica sediment.

2.5. Discussion Concentration profiles of sulfide, sulfate, methane and F430 along with ANME cell counts, clearly indicate F430 plays a role in AOM at both study sites. F430 was previously identified in ANME-1 and -2 sediment from the Black Sea, Eckernförde Bay (Baltic Sea), Gauymas Basin, Hydrate Ridge and Mediterranean seeps (Allen et al., 2014, Kruger et al., 2003, Mayr et al., 2008), although these studies did not quantify F430, so a link between F430 and AOM was not established using geochemical profiles. The maximum concentration of F430 is associated with AOM maxima at both sites, and this links AOM to the reversed biochemical path associated with methanotrophy (reversal of equation 1).

Methane conversion to methyl-S-CoM catalyzed by coenzyme F430 is likely the gateway to the rest of reverse methanogenesis, as it is the most energy intensive step, with a ΔG0 value of -30 kJ mol-1 (Scheller et al., 2010). Subsequent steps of methanogenesis should also be able to take place in reverse with methane either assimilated into the cell or fully oxidized to DIC.

Multiple forms of F430 have been reported at these sites depending on the dominant ANME group. Our finding are consistent with previous results from ANME-2 dominated Hydrate Ridge sediment, where F430 was the abundant from (Mayr et al., 2008). Allen et al. (2014) and Mayr et al. (2008) found and identified three additional F430 structures (vinyl-methylthio-F430, methylsulfoxide-F430 and Keto-F430) in ANME-1 sediment from the Guaymas basin. Allen et al. (2014) report additional forms (didehydro-F430 and Vinyl-F430) in ANME-2 from Mediterranean seeps and Hydrate Ridge. It is unclear what function these different forms serve, or if they are used in processes other than AOM (Allen et al., 2014). The vinyl, methylthio, methylsulfoxide, keto and didehydro groups affect the reactivity of the nickel group, and only F430 has been identified as the form that catalyzes anaerobic methane oxidation (Mayr et al., 2008). Consistent with this, in both sample sites, we found only F430 association with AOM.

Previous natural abundance isotope studies of ANME-dominated sediments have traced 13C-depleted methane into biomass and lipids (Boetius et al., 2000, Elvert et al., 1999, Hinrichs et al., 1999, Hinrichs et al., 2000b, Orphan et al., 2001b). Based on previous observations, F430 was therefore expected to be depleted in 13C. The isotope record for lipids is considerably variable (House et al., 2009) and we also expected variability among F430 carbon isotope values.

At both locations, DIC and F430 were substantially enriched in 13C relative to methane and lipids. The large isotopic difference between the ANME biomolecules (figures 2-9 and 2-10) suggests that carbon from both DIC and methane was assimilated and incorporated into cell components, including F430. The lower δ13C values of lipids are consistent with carbon derived from methane, as previous studies have suggested (Boetius et al., 2000, Elvert et al., 1999, Hinrichs et al., 1999, Hinrichs et al., 2000b, Orphan et al., 2001b). Higher δ13C observed for coenzyme F430 suggests a significant proportion of DIC was incorporated into its structure. The wide range in the isotope values of F430 suggests it contains varying amounts of carbon from both DIC and methane, but what causes ANME to vary the assimilation from each source is unclear. Alternatively, both F430 and lipids could be derived

from the same carbon substrate, provided significant differences in isotopic fractionation accompany their respective biochemical synthetic pathways.

To explore the origins of the distinct isotope values of lipids and F430, we developed an isotope mass balance with fractionation model to estimate the proportion of methane and DIC assimilated carbon in lipids and F430. This calculation constrains the percentages of DIC and methane needed to generate the observed isotope 13 13 13 13 signatures (figure 2-11). δ C CH4, δ C DIC, δ Clipid and δ C F430 are all measured for Hydrate Ridge and Santa

Monica Basin sediments. A range of α and ε values (table 2-9; equations 20 and 21) were calculated using the 13 13 13 measured δ C CH4 and δ C DIC and by assuming δ C F430 from 100% DIC formation is -20‰, (similar to the values observed at Hydrate Ridge) and by varying the δ13C value for F430 formed from 100% methane (-70‰, -80‰ or -

90‰) based on observation from the Santa Monica Basin sediment. We derived εlipid/DIC assuming lipids formed 13 from 100% DIC have δ Clipid values of -50‰ -60‰ or -70‰, and for εlipid/methane assuming lipids formed from 100% 13 13 methane have δ Clipid values of -100‰ -110‰ or -120‰. These values were based on the range of δ Clipid observed in both the Hydrate Ridge and Santa Monica Basin sediment (figure 2-9 and 2-10).

(δ + 1) α/ = (20) (δ + 1)

ε/ = α − 1 (21)

Using equations 22 and 23, fCH4 is varied from 100% methane (0% DIC) to 0% methane (100% DIC) 13 13 generating a range of possible δ Clipid and δ C F430. Using figure 2-11 the amounts of methane and DIC that are needed to generate the observed values in Hydrate Ridge and the Santa Monica Basin can be determined.

13 13 13 δ Clipid = fCH4(δ C CH4 αlipid/CH4 + εlipid/CH4) + (1-fCH4)(δ C DIC αlipid/DIC + εlipid/ DIC) (22)

13 13 13 δ C F430 = fCH4(δ C CH4 αF430/CH4 + εF430/CH4) + (1-fCH4)(δ C DIC αF430/DIC+ εF430/ DIC) (23)

1 = fCH4 + fDIC

The calculated results (presented in Table 2-10) suggest F430 from Hydrate Ridge contains 90-95% DIC, whereas F430 from Santa Monica F430 contains 40-50% DIC. Depleted lipid isotope values for both sites can be

accounted for by the assimilation of 60-90% methane. The observed ~60‰ variation in the carbon isotope values of the ANME lipids suggests the amount of DIC assimilated varied between 10 to 40%. For both F430 and lipids, a greater proportion of DIC was incorporated into compounds from Hydrate Ridge sediment.

There is evidence from enrichment cultures for the assimilation of DIC in addition to other C1 species into ANME lipids (Bertram et al., 2013, Kellermann et al., 2012, Lloyd et al., 2011, Wegener et al., 2008). Kellermann et al., (2012) have shown that ANME-1 preferentially incorporate DIC into their lipids instead of methane. In contrast, authors who conduct natural abundance studies generally observe highly depleted values for whole ANME cells and lipids, and therefore they suggest that methane is the only ANME carbon source (Orphan et al., 2002, Orphan et al., 2001b).

Lipids recovered from both sample sites also show a range in δ13C values, particular hydroxy-archeaol from multiple Hydrate Ridge cores (figure 2-9) where the range is ~60‰. This variability is similar to the range in (~50‰) for ANME cell clusters and lipids (House et al., 2009, Orphan et al., 2001b), and we suggest it reflects different proportions of DIC and methane carbon incorporation into the lipids.

Previously published metagenomic analysis of Hydrate Ridge sediment has shown both ANME-1 and 2 possess the genetic code for all but one step of the methanogenic pathway (Hallam et al., 2004). Hallam et al., (2004) proposed ANME use the pathway in both the methanogenic direction and in the reverse methanogenic direction. This suggestion is consistent with our evidence that AOM is associated with F430, and that ANME assimilate both DIC and methane, possibly by using the methanogenic pathway in different directions.

We propose that methane is directly assimilated via reverse methanogenesis. The first step, in which F430 is used, removes a hydrogen from methane and converts the methane carbon into a methyl group on coenzyme M. This methyl group is then transferred to form methyl-H4MPT which can then be converted to acetyl-CoA via carbon monoxide dehydrogenase/acetyl-CoA synthase (Hallam et al., 2004, Taupp et al., 2010). Acetyl-CoA can then be used to synthesize isoprenoids, the building blocks of archaeol and hydroxyl-archaeol via the mevalonate pathway (Goldstein and Brown, 1990, Smit and Mushegian, 2000).

We hypothesize that one or more carbon species from the DIC pool are assimilated via the initial steps of methanogenesis operating in the methanogenic direction. In methanogens, DIC in the form of CO2(aq) is reduced to

Formyl-MF, which is converted to methylene-H4MPT(Hallam et al., 2004, Taupp et al., 2010). The methyl group from methylene-H4MPT is combined with glycine to form serine. Angelaccio et al. (2003) provided evidence that this reaction is catalyzed by serine hydroxymethyltransferase, an enzyme which so far has been reported in all sequenced archaeal genomes, including ANME. Serine is readily converted to glutamate, the simplest basic building block of the tetrapyroles, including F430 (Friedmann and Thauer, 1986, Gilles et al., 1983, Gilles and Thauer, 1983, Angelaccio et al., 2003).

Hallam et al., (2004) and Taupp et al., (2010) have reported that unlike the other enzymes and coenzymes in methanogenesis, the genes for the conversion of methyl-H4MPT to methylene-H4MPT are not present in AOM

associated sediment from the Eel River, Black Sea, Baltic Sea or Hydrate Ridge. This point represents a potential division of the pathway, where one half operates in reverse and one half in the methanogenic direction (figure 2-12), as suggested in Hallam et al., (2004).

If the enzyme that converts of methylene-H4MPT to methyl-H4MPT is present in ANME then the assimilation of DIC and methane may instead be controlled by the availability of electrons. The reduction of DIC should consume electrons, while the oxidation of methane releases elections (figure 2-12). If methane is assimilated into acetyl-CoA and then oxidized to CO2, more electrons would be released to reduce DIC, although this is not likely as it would not be energetically advantageous. Electron flow to other electron acceptors may influence the ratio of methane versus DIC assimilated into biochemical constituents. In addition, complete oxidation of methane could significantly lower the isotopic composition of the DIC pool that is available to ANME. Although less likely, DIC could also be potentially enriched in 13C, if 12C is preferentially assimilated and incorporated into lipids. Reconciling the methane-signal of lipids and the DIC-like signal in F430 will require better constraints on the proportion of methane and DIC assimilated by cells, and fractionation factors between the respective substrates and biochemical compounds.

Operating the methanogenic pathway in two directions allows for both DIC and methane to be assimilated. The assimilation of multiple carbon sources, potentially combined with localized depletions in DIC associated with complete oxidation of methane to CO2, could explain the range in carbon isotope values observed at Hydrate Ridge, the Santa Monica basin and previously (~50‰) for lipids and cells (House et al., 2009, Orphan et al., 2002). Potentially this biochemically plasticity is an adaption to the low energy environment allowing ANME to generate energy and assimilate carbon.

2.6. Conclusions A link between F430 and AOM is established in both the Hydrate Ridge and Santa Monica Basin sediment based on profiles of sulfide, sulfate, methane, F430 and ANME aggregate counts. This strengthens the evidence for ANME oxidizing methane via the reversal of methanogenesis. F430 in both the Hydrate Ridge and Santa Monica Basin sediment is enriched in 13C relative to archaeal lipids. This enrichment is likely due to the assimilation of DIC in addition to methane and indicates that there are two pathways of assimilation. Values for both F430 and the ANME lipids are variable in the sediment, with this variability potentially explained by the assimilation of multiple substrates. Using mass balance with fractionation we can constrain the proportion of DIC and methane assimilation to generate the observed isotope values of F430 and lipids. The ability of ANME to assimilate more than one carbon substrate has been previously demonstrated in labeling experiments (Kellermann et al., 2012, Lloyd et al., 2011). Questions remain regarding what controls the proportion of carbon assimilated from each difference carbon pool and the effect mixing between DIC and oxidized methane may have on ANME isotope values.

2.7. Acknowledgements We thank Denny Walizer, and Clayton Magill for help in the lab. For help with the Nano-EA IRMS we thank Christopher Junium and Pratigya Polissar. Anne Dekas, Stephanie Connon and Jennifer Glass are thanked for sample collection. We also thank Sara Lincoln for constructive comments. This research was funded by Royal Dutch Shell Geosciences Energy Research Facilitation Awards, PSARC and the American Chemical Society petroleum research fund.

2.8. Figures and tables

Figure 2-1: Sampling localities. Hydrate Ridge mud was collected on cruise AT-18-10 in 2011 with Santa Monica mud collected on cruise AT26-06 in 2013

Figure 2-2: LC fraction collection. Coenzyme F430 is collected for isotope analysis by Nano-EA IRMS after the second dimension of HPLC separation. The 430 nm wavelength peak is collected after it passes through the diode array detector, additional identification of F430 is based on the UV/vis spectra and the identification of the 905 molecular ion.

Figure 2-3: F430 Uv/vis absorbance spectra. This spectra for coenzyme F430 matches previously published absorbance spectra from Gunsalus and Wolfe (1978).

Figure 2-4: F430 ion spectra. The 905 molecular ion that is used to identify F430 along with UV/vis absorbance spectra

−10 Cups OEP Methionine Sucrose Data −15 OEP regressionline Methionine regression line Sucrose regression line

−20 permil

−25

−30

−35 0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 1/n

−10 Cups OEP Methionine Sucrose Data −15 OEP regressionline Methionine regression line Sucrose regression line

−20 permil

−25

−30

−35 0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 1/n

Figure 2-5: Hydrate ridge 1/n plot. 1/n plot for coenzyme F430 isolated from the Hydrate Ridge sediment. The regression line from the blank cups represents the real isotope number of F430. The isotopic correction corrects for the blank contribution and isotopic offset using the three standards. The raw F430 carbon isotope numbers are more enriches than expected based on previous lipid and ANME aggregate values (Orphan et al., 2001b)

−10

−20

−30

−40 permil

Cups −50 OEP Methionine Sucrose Data −60 OEP regressionline Methionine regression line Sucrose regression line

−70 0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 1/n

Figure 2-6: Santa Monica 1/n plot. 1/n plot for coenzyme F430 isolated from the Santa Monica Basin sediment. The raw uncorrected values are more depleted than the Hydrate Ridge sediment.

Figure 2-7: Hydrate Ridge sediment data Abundance data for Coenzyme F430 and ANME/DSS-2c aggregate counts show that both peak in the area greatest methane oxidation and sulfate reduction (3-6 cm section)

Figure 2-8: Santa Monica sediment data Abundance data from the Santa Monica Basin for Coenzyme F430 and ANME-2/DSS aggregate counts show that both peak in the area greatest methane oxidation inferred from the sulfate and sulfide profiles.

13C Hydrate Ridge 0

−20

−40

−60

−80

−100

−120

−140

− DIC IPL TOC F430 A other Methane hydroxy − cores archaeol archaeol H

Figure 2-9: Hydrate ridge carbon isotope values. Carbon isotope date for DIC, methane, coenzyme F430, intact polar lipid archaeol and hydroxy-archaeol from Hydrate Ridge. The black lines represent the mean value.

13C Santa Monica Basin −40

−50

−60

−70

−80

−90 13C permil

−100

−110

−120

−130 DIC Methane F430 archaeol hydroxy−archaeol

Figure 2-10: Santa Monica basin carbon isotope values Collected isotope data for the Santa Monica Basin, both archaeol and hydroxy-archaeol are the intact polar lipid forms. The black lines represent the mean value. Methane isotope values come from Stolper et al., (Stolper et al., 2015).

Figure 2-11: Mass balance modeling results Isotope results using equations 13 and 14 to explore the effects the amounts of methane and DIC have on the isotope value of ANME cellular components. For F430 values 100% formation from DIC one value (-20‰) is used for F430 based on the observation from Hydrate Ridge, with three values used for 100% methane formation (-70‰, -80‰ and -90‰) based on the range of values from the Santa Monica Basin sediment. For lipid values three different 100% DIC (-50‰, -60‰ and -70‰) and methane (-100‰, -110‰ and -120‰) values were used based on observation in the Hydrate Ridge and Santa Monica basin sediment (figures 2-9 and 2-10).

Figure 2-12: Carbon assimilation diagram. Adapted from Taupp et al. (2010), the methanogenic path way is shown split into two sections. One operates in the normal direction assimilating CO2(aq) and the other in the reverse direction assimilating methane. MF Methanofuran H4MPT Tetrohydromethanopterin

Table 2-1: Inorganic carbon isotope values for Hydrate Ridge. Carbonate was measured with other carbon species determined using ε from Zhang et al. (1995)

Measured value δ13C 2- - carbonate DIC CO3 HCO3 CO2(aq) CO2 (g) -39 -43 -45 -42 -55 -52 -39 -42 -45 -42 -54 -52 -44 -47 -50 -47 -60 -57 -35 -39 -41 -38 -51 -48 -17 -20 -23 -20 -32 -30 -13 -16 -19 -16 -28 -26

Table 2-2: Inorganic carbon isotope values for Santa Monica Basin. DIC was measured with other carbon species determined using ε from Zhang et al. (1995)

Measured δ13C 2- - DIC CO3 HCO3 CO2(aq) CO2 (g) -49 -52 -48 -61 -59 -50 -53 -49 -62 -60 -46 -49 -45 -58 -56 -44 -47 -43 -56 -54 -45 -48 -44 -57 -55 -45 -48 -44 -57 -55 -46 -49 -45 -58 -56 -47 -50 -46 -59 -57 -46 -49 -45 -58 -56 -46 -49 -45 -58 -56 -46 -49 -45 -58 -56 -46 -49 -45 -58 -56

Table 2-3: Isotope analytical error.

Hydrate Santa Ridge Monica OCTAETHYLPORPHINE 2.3 0.7 SUCROSE 0.3 0.7 METHIONINE 0.2 0.4

σoffset 1.6 1.0

σblank 1.0 0.4

σ∑ 2.9 1.5

Table 2-4: Sediment data. Geochemical data, aggregate counts and amount of F430 per gram of wet sediment for Hydrate Ridge and the Santa Monica Basin.

HYDRATE RIDGE - 2- DEPTH CM HS SO4 Methane wet sediment F430 aggs per ml sed mM mM uM g ug/g x10^6 0-3 1.3 8.7 166 31 11 1.1 3-6 7.7 0.6 636 32 83 2.3 6-9 7.5 0.4 1375 32 38 0.8 9-12 4.7 2.0 1273 32 3 0.4

SANTA MONICA BASIN - 2- DEPTH CM HS mM SO4 mM NH4+ µM wet sediment F430 aggs per ml sed g ug/g x10^7 0-2 8.6 10.7 101.7 10 1616 8.5 2-3 24.9 3.4 42.3 12 909 5.2 3-4 35.0 0.9 16.0 11 40 1.6 4-5 33.1 0.6 33.7 11 118 1.9 5-6 10.8 0.2 38.7 10 0 1.1 6-9 36.7 0.2 52.0 23 14 0.3 9-12 21.0 0.2 65.7 30 1 0.6 12-15 11.6 0.1 62.0 36 0 0.7

Table 2-5: Hydrate Ridge TAG sequencing results. The table shows the number of tags for each ANME group. ANME-1 group is the most abundant in the Hydrate Ridge sediment with limited amounts of ANME-2 and ANME- 3

Hydrate Ridge TAG sequencing Sediment depth Taxon 0-3 3-6 6-9 9-12 ANME-1a 9 62 294 216 ANME-1b 0 59 224 190 ANME-1;Other 2 248 721 188 ANME-2b 2 79 21 2 ANME-2a-2b 3 27 36 11 ANME-2a-2b;Other 0 3 1 0 ANME-2c 2 31 145 20 ANME-3 0 0 0 0

Table 2-6: Santa Monica Basin TAG sequencing. The table shows the number of tags for each ANME group. ANME-2 is more dominant than at hydrate Ridge with most of the population in the upper 2 com of the core.

Santa Monica Basin Sediment depth Taxon 0-1 1-2 2-3 3-4 4-5 5-6 6-9 9-12 12-15

ANME-1a 22 51 62 1 2 3 8 6 17

ANME-1b 3 5 3 1 0 0 0 2 5

ANME-1;Other 3 7 1 0 0 0 0 0 3

ANME-2b 8 18 6 0 0 0 0 0 1

ANME-2a-2b; 43 218 81 16 32 24 45 53 40

ANME-2a-2b;Other 0 1 0 0 0 0 0 0 0

ANME-2c 300 1292 435 101 189 146 142 46 186

ANME-3 15 54 17 7 10 7 16 3 13

Table 2-7: Isotope results from Hydrate Ridge. DIC values were calculated from measured carbonate values using ε values from (Zhang et al., 1995)

HYDRATE RIDGE δ13C F430 IPL-Archaeol IPL Hydroxy Carbonate DIC Methane Archaarol -24 -94 -64 -38 -42 -64 -21 -94 -63 -37 -41 -62 -21 -94 -43 -47 -67 -25 -88 -34 -48 -69 -22 -124 -16 -20 -70 -25 -125 -21

Table 2-8: Isotope results from the Santa Monica basin Methane values taken from Stolper (Stolper et al., 2015)

SANTA MONICA BASIN δ13C F430 IPL-Archaeol IPL Hydroxy Archaarol DIC Methane -90 -127 -112 -49 -70* -84 -119 -130 -50 -70* -83 -46 -67* -77 -44 -67* -78 -45 -67 -45 -46 -47 -46 -46 -46 -46

Table 2-9: α and ε values used to model the assimilation of DIC and methane into F430 and lipids

F430

100% DIC αF430/DIC εF430/DIC 100% Methane αF430/methane εF430/methane Model -20 1.02 20.83 -70 1.00 0.00 A -80 0.99 -10.75 B -90 0.98 -21.51 C Lipid

100% DIC αlipid/DIC εlipid/DIC 100% Methane αlipid/methane εlipid/methane Model -50 0.99 -10.42 -100 0.97 -32.26 D -60 0.98 -20.83 -110 0.96 -43.01 E -70 0.97 -31.25 -120 0.95 -53.76 F

Table 2-10: Model results. Results of the assimilation isotope model based on equations 22 and 23

COENZYME F430 MODEL RESULTS MODEL A B C 13 13 13 % CH4 ASSIMILATED δ C F430 δ C F430 δ C F430 100 -90 -80 -70 90 -83 -74 -65 80 -76 -68 -60 70 -69 -62 -55 60 -62 -56 -50 50 -55 -50 -45 40 -48 -44 -40 30 -41 -38 -35 20 -34 -32 -30 10 -27 -26 -25 0 -20 -20 -20

LIPID MODEL RESULTS MODEL D E F 13 13 13 % CH4 ASSIMILATED δ C Lipid δ C Lipid δ C Lipid 100 -120 -110 -100 90 -115 -105 -95 80 -110 -100 -90 70 -105 -95 -85 60 -100 -90 -80 50 -95 -85 -75 40 -90 -80 -70 30 -85 -75 -65 20 -80 -70 -60 10 -75 -65 -55 0 -70 -60 -50

2.9. References ALLEN, K. D., WEGENER, G. & WHITE, R. H. 2014. Discovery of Multiple Modified F430 Coenzymes in Methanogens and Anaerobic Methanotrophic Archaea Suggests Possible New Roles for F430 in Nature. Applied and Environmental Microbiology, 80, 6403-6412. ANGELACCIO, S., CHIARALUCE, R., CONSALVI, V., BUCHENAU, B. R., GIANGIACOMO, L., BOSSA, F. & CONTESTABILE, R. 2003. Catalytic and Thermodynamic Properties of Tetrahydromethanopterin- dependent Serine Hydroxymethyltransferase from Methanococcus jannaschii. Journal of Biological Chemistry, 278, 41789-41797.

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Chapter 3: Stable isotope probing of ANME carbon assimilation

3.1. Abstract Anaerobic oxidation of methane (AOM) oxidizes over a billion kilograms of methane and is performed by an Archaea group known as ANerobic MEthanotrophs (ANME). Evidence indicates that ANME are metabolically flexible and, in addition to methanotrophy, are able to produce methane as well as assimilate a range of carbon substrates including dissolved inorganic carbon (DIC) (Bertram et al., 2013, Kellermann et al., 2012, Lloyd et al., 2011, Wegener et al., 2008). In chapter 2, we observed coenzyme F430 and ANME lipids were isotopically heterogeneous like methane and DIC in the sediment. This indicates that ANME assimilate different carbon substrates into biochemical components. To test this, we incubated ANME-rich sediments with 13C labeled DIC and methane. Coenzyme F430 was recovered only from experiments where methane was added to the headspace, and not from experiments designed to stimulate methanogenesis. This strengthens the link between F430 in the sediment and its use in AOM. In Hydrate Ridge sediments, we observed that labeled DIC was assimilated into archaeol, while labels from both methane and DIC assimilated into F430 and lipids in the Santa Monica Basin experiments. Amounts of F430 recovered in the Hydrate Ridge experiments were too low for isotope analysis. By dual labeling experiment with 15N ammonium, new F430 production was quantified and used to calculate the percentage of each label assimilated. Newly formed F430 was found to have been synthesized from ~70% to 100% from DIC and 0 to ~20% of the labeled from methane. Assuming the amount of newly formed lipid is the same as F430 we find lipids to have assimilated ~50% to 100% of the DIC label and 0% to ~10% of the methane label. Due to the amount of labeled methane that is oxidized to DIC we cannot be sure that methane is directly assimilated only that some of the label from methane is assimilated, possibly after first being oxidized to DIC. Highly variable isotope data have been observed in nature for cell and lipids of both ANME-1 and 2 (House et al., 2009, Orphan et al., 2002, Orphan et al., 2001b). This can either be explained by the assimilate of methane in addition to DIC or the mixing of oxidized methane and DIC. Given that roughly 50% of the label from methane ends up in the DIC pool it seems likely that ANME first oxidize methane to DIC and then assimilate a mix of DIC and oxidized methane into their biomass.

3.2. Introduction Anaerobic oxidation of methane (AOM) is an important sink for the billion kilograms of methane that are produced annually in marine sediments (Hallam et al., 2004). Geochemical and biological evidence indicates that this process is carried out by a microbial consortium of Archaeal ANerobic MEthanotrophs (ANME), often in partnership with sulfate-reducing bacteria (Orphan et al., 2001b, Orphan et al., 2009). ANME comprise of 3 groups AMNE-1, 2 and 3 with ANME-1 existing as single cells in the sediment and ANME-2 and 3 forming cell aggregates, normally with a sulfate reducer (Hallam et al., 2003, Knittel et al., 2005, Orphan et al., 2001a, Orphan et al., 2002, Treude et al., 2007). ANME oxidize methane to CO2 and assimilate a proportion into their biomass (Lloyd

et al., 2011, Orphan et al., 2002, Wegener et al., 2008). Evidence for the assimilation of methane comes from natural abundance studies that have identified methane carbon assimilation into ANME lipids and cells based on depleted d13C values (Aloisi et al., 2002, Boetius et al., 2000, Hinrichs et al., 2000b, House et al., 2009, Orphan et al., 2001b, Pancost et al., 2000, Raghoebarsing et al., 2006, Reitner et al., 2005).

Evidence indicates that ANME, in addition to oxidizing methane, are also able to produce methane, and are able to assimilate other C-1 sources including dissolved inorganic carbon (DIC) and methanol (Bertram et al., 2013, Kellermann et al., 2012, Lloyd et al., 2011, Wegener et al., 2008) . Labeling experiments using sediment with active AOM show that ANME-1 preferentially incorporate DIC into biomass (Kellermann et al., 2012). This indicates that at least some ANME are not exclusively reliant on the oxidation of methane for cellular carbon. Variable use of two different carbon substrates could explain the wide range of isotope signatures (~50‰) measured in ANME cell clusters in seep settings (House et al., 2009, Orphan et al., 2001b). Dual assimilation of methane and DIC is potentially also responsible for the observed isotopic heterogeneity between F430 and ANME lipids (chapter 2).

Previously, we observed a ~30‰ difference in the carbon isotopic composition of F430 and ANME lipids in sediments from Hydrate Ridge and the Santa Monica Basin. This isotopic difference suggests ANME cells assimilate DIC into F430 and methane into the lipids. Assimilation of carbon from both methane and DIC simultaneously has been observed in ANME-dominated mats from the Black Sea (Treude et al., 2007). Assimilation of multiple carbon sources could explain the previously reported wide range in lipid δ13C values (House et al., 2009). In Chapter 2, coenzyme F430 and archaeal lipids were shown to be isotopically distinct and that their differences reflected isotopic differences between methane and DIC in the sediment. F430 and ANME lipids are formed from different biochemical precursors (F430, glutamate and archaeol, acetyl-CoA(Gilles and Thauer, 1983, Koga and Morii, 2007, Pfaltz et al., 1987)) which can be derived from different substrates. The assimilation of DIC into glutamate by the methanogenic pathway and the serine cycle and methane into acetyl-CoA by the reverse methanogenic pathway could explain the isotopic heterogeneity observed in sediments between lipids and F430. Incubations of ANME with 13C labeled substrates are used to trace assimilation into different biochemical components.

Series of incubations were set up to test separately the assimilation of DIC and methane using isotope labels. Additional experiments constructed without methane tested for methanogenesis and to see if methane is essential for ANME survival in sediment with active AOM. Similar experiments have been successful in exploring biochemical pathways in Archaea, including lipid recycling (Takano et al., 2010). ANME are the main diazotroph 15 15 in AOM sediment, fixing N2 to ammonium (Dekas et al., 2014). Using N labeled ammonium, in addition to the 13C labeled DIC and methane, the amount of newly synthesized F430 can be determined from isotopic analysis of the tetrapyrrole nitrogen.

Incubations used sediment from Hydrate Ridge and the Santa Monica Basin where isotopic differences between ANME biomolecules (F430 and lipids) and carbon substrates (DIC and methane) was previously observed. These sites allow for difference between the ANME groups to be explored as TAG data indicate ANME-

1 is more abundant in Hydrate Ridge and ANME-2 are more abundant in the Santa Monica Basin sediment. Based on observations reported in chapter 2, we expect greater assimilation of 13C DIC in experiments using Hydrate Ridge sediments reflected in the isotopic composition of lipids. This is based on the results of chapter 2, where F430 was more enriched in 13C in the Hydrate Ridge sediment, and previous labeling experiments, where ANME-1 have been shown to assimilate DIC (Kellermann et al., 2012). In contrast, for experiments using the Santa Monica Basin sediment, we expect both methane and DIC to be assimilated and reflected in lipid and F430 isotope data. This is based on the carbon isotope values observed for F430 and lipids in chapter 2 and the larger isotopic range previously observed for ANME-2(Orphan et al., 2002).

3.3. Methods

3.3.1. Shipboard collection, core processing, and sample storage Samples were collected from Hydrate Ridge, Oregon as part of the R/V Atlantis cruise 18-10 in September 2011 and from the Santa Monica Basin, California as part of the R/V Atlantis cruise 26-06 in October 2013. Using the ROV Jason II, sediment push cores were collected from seep environments characterized by the presence of chemosynthetic clam beds (PC20) or microbial mats (PCKD). PC20 (Hydrate Ridge, 44°N 40.19 125°W 5.88) was recovered from a chemosynthetic clam bed with live Calyptogena clams present in the 0-3 and 3-6 cm horizons. PC20 was processed shipboard in 3 cm intervals and stored under Ar at 4°C prior to the combinations of the 0 to 12 cm horizons into a sediment slurry for incubations in October 2013. PCKD (Santa Monica Basin, 33°N 38.403 118°W 48.025) was taken through an orange microbial mat. PCKD was stored as an intact core (figure 3-1) at 4°C prior to extrusion and homogenization into a sediment slurry in October 2015.

Parallel cores taken at both locations (PC28 – Hydrate Ridge, OR; PC4 – Santa Monica Basin, CA) were processed for DNA, microscopy, lipid, and pore water chemistry analyses. Immediately after the parallel push cores were sectioned, aliquots of sediment were stored at -80°C for DNA extraction, or were fixed for microscopy by adding 4% paraformaldehyde (PFA) to sediment-seawater mixtures in a 1:1 ratio and incubating at 4°C for 12 hours. Fixed samples were washed with 3x phosphate buffered saline (PBS) and stored in 1:1 3x PBS:ethanol at -20°C. Pore water was collected from 1 to 3 cm sediment intervals under Ar using a pressurized gas sediment squeezer (KC Denmark A/S, Silkeborg, Denmark) (Reeburgh, 1967), and residual sediments were then stored at -80°C for lipid analysis. Pore water geochemical analysis included anion, cation, and sulfide concentrations, as well as dissolved inorganic carbon (DIC) concentration and stable isotope analysis.

3.3.2. DNA extraction and tag sequencing Sediment was stored at -80°C until DNA extraction. DNA was extracted using a MoBio Ultraclean soil kit (MO BIO Laboratories Inc., Carlsbad, CA, USA). Preparation for sequencing of the V4 region of the 16S rRNA gene was carried out according to the Earth Microbiome Project protocol (Caporaso et al., 2012, Caporaso et al., 2011) with modifications as previously described (Case et al., 2015). Raw sequences were generated on an Illumina

MiSeq platform at Laragen, Inc. (Los Angeles, CA, USA) and are available in the Sequence Read Archive (PRJNA350854). Sequence data were demultiplexed and processed using a modified version of the QIIME pipeline (Caporaso et al., 2010) as described previously (Mason et al., 2015). Prior to sample comparison, singletons and PCR contaminants were removed, and a 0.01% relative abundance threshold was applied.

3.3.3. Sediment incubations Incubations were constructed following the conditions outlined in table 3-1. Sediment was combined with -1 artificial seawater for a 1:2 v/v sediment slurry. Artificial seawater was N2-sparged and contained (g l ): 24 g NaCl; . . . 5 g MgCl2 6H2O; 1.5 g Na2SO4; 1.31 g CaCl2 2H2O; 0.67 g KCl; 0.1g KBr; 0.027 g H3BO3; 0.027 g SrCl2 6H2O; 0.003 g NaF. Sediment slurry was distributed into 3 parallel bottles per experimental condition, which were further 15 15 15 15 amended as follows: 1 – 5 mM NaHCO3, 0.5 mM NH4Cl (20 at.% N, F 0.2, δ N 68808‰), and a 20% 13 13 13 13 13 13 CH4/80% N2 headspace (25 at.% C, F 0.25, δ C 29000‰); 2 – 5 mM NaHCO3 (3.78 at.% C, F 0.078, δ C 15 15 15 15 2540‰), 0.5 mM NH4Cl (20 at.% N, F 0.2, δ N 68808‰), and a 20% CH4/80% N2 headspace; 3 - 5 mM 15 15 15 15 13 13 13 NaHCO3, 0.5 mM NH4Cl (20 at.% N, F 0.2, δ N 68808‰), 0.5 mM acetate (3.26 at.% C F 0.0326, δ C 13 13 13 1937‰), and a 20% H2/80% CO2 headspace; 4 - 5 mM NaHCO3 (3.78 at.% C, F 0.078, δ C 2540‰), 0.5 mM 15 15 15 15 15 NH4Cl (20 at.% N, F 0.2, δ N 68808‰), and a N2 headspace; 5 - 5 mM NaHCO3, 0.5 mM NH4Cl (20 at.% 15 15 15 N, F 0.2, δ N 68808‰), and a 20% CH4/80% N2 headspace; 6 – autoclaved killed control, 5 mM NaHCO3, 0.5 15 15 15 15 13 13 13 mM NH4Cl (20 at.% N, F 0.2, δ N 68808‰), and a 20% CH4/80% N2 headspace (25 at.% C F 0.25, δ C 29000‰). The % 15N and 13C added to incubations was calculated by isotope mass balance as follows: n n n Ffinal=[( Funlabeled x munlabeled) + ( Flabeled x mlabeled)]/mfinal, where mass (m) was the amount of NH4Cl, CH4, or acetate added to the mixture and at.% = 100 x nF.

The sediment incubations were maintained in the dark, without shaking, and at 4°C. Experimental condition 1 bottles had their seawater replaced at 40 days, and their headspace replaced at 60 days. Bottles were sampled at 2, 33, 75, 100 and 114 days, and samples were preserved for geochemical, microscopy, and DNA analyses. After resuspending the settled sediment, 3 ml of sediment slurry was removed via a gas tight syringe and distributed into a 2 ml microfuge tube and a 1.5 ml cryogenic tube, which was immediately frozen in liquid N2 for later DNA extraction. The sediment slurry was centrifuged for 3 min at 9000 x g. The resulting liquid was preserved for geochemical analysis including anion, cation, and sulfide concentrations, as well as dissolved inorganic carbon (DIC) concentration and stable isotope analysis. The sediment pellet was fixed for microscopy as described above.

3.3.4. Geochemical analysis of pore waters and incubations Sulfide dissolved in sediment pore waters and centrifuged incubation waters was preserved by the immediate precipitation as ZnS through the addition of 0.5M Zn-acetate in a 1:1 ratio with water samples. Concentrations were then determined spectrophotometrically by the Cline assay (Cline, 1969). Water samples for + + + 2+ 2+ - - - 2- anion and cation analysis (Na , NH4 , K , Mg , Ca , formate, acetate, Cl , Br , NO3 , SO4 ) were filtered through a 0.2 µm polyethersulfone (PES) syringe filter and stored at -20°C. After thawing, aliquots were diluted 1:20 with

MQ water, and subsequently were analyzed on a dual channel Dionex ICS-2000 ion chromatography system in the Caltech Environmental Analysis Center. Samples were split and simultaneously separated with cation and anion exchange columns at 0.25 ml min-1 and 30°C. Cations were separated isocratically with 20 mM methanesulfonic acid using an IonPac CS12A column and guard column and anions were separated isocratically with 20 mM KOH using an IonPac AG19 column and guard column.

Scientific, Bremen, Germany) in the Caltech Stable Isotope Facility. Sample runs consisted of 3 reference CO2 gas peaks, 10 replicate sample injections, and 2 final reference CO2 peaks. A concentrated solution of NaHCO3 was used to establish a standard curve for concentration determination by adding a range of volumes to additional exetainer vials, which were interspersed with samples. The concentration of DIC (µmol) in samples was determined by comparing the average of the combined mass 44, 45, and 46 peak areas to the standard curve (n = 20, R2 = 0.99), and calculated after determining sample volume by re-weighing exetainers. δ13C values were corrected for sample- size dependency and then normalized to the VPDB scale with a two-point calibration (Coplen et al., 2006) using NBS-19 and a previously calibrated laboratory carbonate as internal standards. Accuracy (0.11‰, n=79) was determined by analyzing independent standards as samples, and precision (0.42‰, n=10) was determined from NBS-19.

3.3.5. F430 extraction and separation Extraction and isolation of coenzyme F430 followed the separation method of Mayr et al. (2008), as modified in chapter 2, with additional purification steps to allow for quantification and isotope analysis. Approximately ~30 g of Hydrate Ridge and ~10 g of Santa Monica Basin wet sediment were needed to quantify, isolate, and make an isotope measurement on coenzyme F430. Sediment samples were agitated by ultra-sonication probe for 20 minutes in neutral 18.2 Ω water (pH 7), and in an ice bath to keep the temperature at 4OC. Sediment was separated from the extract, by centrifuged at 5000 g for 15 minutes. Sediments were extracted twice more in 18.2 Ω water at pH 3 using 0.1% formic acid. The three extracts were combined and neutralized to pH 7.2 using NaOH, in order to precipitate proteins, which were separated and removed by centrifuging the solution at 9000 g for 10 minutes.

Coenzyme F430 was separated from the protein-free supernatant using two dimensional column chromatography. First, the supernatant was applied to a QAE Sephadex A25 column (1.5 cm x 10 cm) that had equilibrated with 50 nM Tris/HCl (pH 7.5). After the column was flushed with 4 dead volumes of Tris/HCl, the

F430-containing fraction was eluted with 90 ml of 20 nM formic acid. This fraction was then applied to a XAD Amberlite column (1 cm x 10 cm) which had been flushed with two dead volumes of 10 nM formic acid. The F430 fraction was isolated in 10 ml of 100% methanol. This fraction was dried under nitrogen and stored at -20OC before being further purified via HPLC.

High-pressure liquid chromatography (HPLC) was used to purify F430 sufficiently to enable quantification and isotope analysis. The first HPLC separation employed two Waters spherisorb ODS2 columns 5 ㎛ (4.6 mm x 150 mm), linked together and supplemented by a Phenomenex C18 (3mm x 4mm) guard cartridge. Mobile phase A consisted of HPLC-grade water, mobile phase B of 0.1% formic acid and mobile phase C of acetonitrile (HPLC- grade). At a flow of 0.5ml/min, the following gradient was applied: 0 minutes 0% A, 70% B 30%C; 2 minutes 0% A, 70% B 30%C; 4 minutes 0% A 50%B 50%C; 20 minutes 50% A, 0% B, 50% C; 25 minutes 25% A, 0% B, 75% C; 28 minutes 25% A, 0% B, 75% C; 30 minutes 0% A, 70% B, 30% C. The F430 peak eluted at 25 minutes and was collected over a 1.5 minute window based on the UV/vis detector response. Fractions were dried under nitrogen and brought re-dissolved in methanol for additional purification.

To separate F430 from a co-eluting molecule that was contributing extra carbon in the Nano EA-IRMS analysis, a Thermo Hypercarb 5㎛ (100mm x 4.6mm) was used. Mobile phase A consisted of HPLC-grade water, mobile phase B of 0.1% HCl and mobile phase C of acetonitrile (HPLC-grade). At a flow of 0.5ml/min the following gradient was applied: 0 minutes 0% A, 70% B 30%C; 2 minutes 0% A, 70% B 30%C; 4 minutes 0% A, 50% B, 50%C; 18 minutes 25% A, 50% B, 25% C; 20 minutes 50% A, 0% B, 50% C; 25 minutes 25% A, 0% B, 75% C; 28 minutes 25% A, 0% B, 75% C; 30 minutes 0% A, 70% B, 30% C. Quantification and identification were performed on the first run of sample through the Hypercarb column and subsequent runs were collected for nano- EA/IRMS analysis. F430 was identified by UV/vis detection of absorbance in the 430 nm wavelength and confirmed by the m/z 905 ion of the complete F430 structure. A previously published molar extinction coefficient of 21000 M-1 cm-1 was used to quantify F430 (Ellefson et al., 1982, Whitman and Wolfe, 1980). An Agilent 6300 ion trap with an ESI source was used for mass spectral analysis. Fractions for nano-EA/IRMS were collected at 8 minutes for 20 seconds. Samples were then dried under nitrogen and transferred to Costech tin boats using methanol. Samples were covered and left to dry before loading into autosampler for isotope analysis.

3.3.6. Isotope analysis of F430 Quantities of F430 isolated from environmental samples are typically too small for conventional EA-IRMS (elemental analyzer - isotope ratio mass spectrometry). Instead, we used a nano-scale EA/IRMS technique, developed by Polissar et al. (2009). In this method, the combusted sample is concentrated by cryogenic capture, transferred by a low flow of helium through a capillary gas chromatograph column (J&W scientific GS-

� = � = (1)

� = ((� − � )/ �) (2)

Samples are first corrected for background contribution, which are generally sourced from the tin boats (Costech), helium carrier gas, and oxygen combustion gas. These “blank” contributions were determined from multiple measurements of empty tin cups, and by isotope mass balance (Equation 3) where nblank is blank peak area, nobs is the observed sample peak area, δblank is blank δ values and δobs is the observed δ of the sample.

�� = �� + �� (3)

This can be rearranged to the useful form of a linear relationship:

� = � − �(� − �)/(1/�) (4)

With the true isotope value being the intercept and determined using equation 5:

� = (�� − ��)/(� − �) (5)

The intercept of the relationship is the blank-corrected delta value for the sample, which is calculated using the peak areas of the blank and sample peaks (nblank, nobs), and the observed delta values for the blank and sample analyses (δblank , δobs) (equation 5). Plotting δobs against 1/nobs illustrates the mixing relationship between blank and analayte for both carbon and nitrogen isotope data sets and that δx (the intercept) is the true value. Figures 3-2, 3-3, 3-4, 3-5, 3-6 and 3-7 illustrates that the intercept of a regression line between the samples and blanks is the true value of a sample.

Analytical accuracy was assessed using a suite of isotopic standards, which are analyzed with the samples, and similarly corrected for blank contributions. Three standards, octaethylporphine (Frontier Scientific δ13C - 34.05‰, δ15N -12.23‰), L-methionine (Sigma Aldrich δ13C -30.45‰, δ15N 0.46‰) and Sucrose (NIST δ13C - 10.45‰) were run in conjunction with the samples to determine isotopic offset. The measured and corrected standard values were regressed against their known values, and the resulting linear equation was used to correct unknown samples for any offset:

� = � �� + �� (6)

Errors were propagated using the sum of squares method, which assumes uncertainties for each correction step are both independent and random (Polissar et al., 2009). For this calculation, uncertainties (σ) for the blank cups (equation 7), standards (equation 8) and offset (equations 9 and 10) were all used to determine total analytical uncertainty (equation 11)

1 � = (� − � )^2 (7) �

1 � = (� − � )^2 (8) �

� = � − � (9)

1 � = (� − � )^2 (10) �

(�∑) = (�) + (�) + (�) + (�) + (�) (11)

Blank correction uncertainty was simplified and represented by the analytical reproducibility of the blank delta values. Uncertainty in blank analyses was determined from n = 6 analyses of empty cups for each analysis batch. In the Hydrate Ridge experiments, this was 1.3‰ for carbon and 1.4‰ for nitrogen. Santa Monica Basin blank correction uncertainty was 0.6‰ and 1.2‰ for carbon, and 1‰ and 0.4‰ for nitrogen (Table 3-2).

Reproducibility of carbon measurements was determined from the standard deviation of blank-corrected octaethylporphine, methionine and sucrose values. Uncertainty in the offset correction was determined from the sum of squares of the offset values determined from equation 9 for each standard.

The total analytical uncertainty for the Hydrate Ridge experiments is 4.6‰ for carbon and 17.7‰ for nitrogen. In the first run of the Santa Monica Basin samples total analytical uncertainty was 3‰ for carbon and 5.3‰ for nitrogen. In the second run total analytical uncertainty for carbon is 1.7‰ and 4.6‰ for nitrogen.

In addition, we checked accuracy and reproducibility for carbon measurements using the methionine standard which was removed from the carbon correction and corrected as if an unknown. The difference between the observed, and corrected methionine standard value and the known value was 3.7‰ for Hydrate Ridge, and 0.8‰ for the first Santa Monica Basin sample analysis and 0.4‰ for the second.

3.3.7. Lipid Analysis Sediment samples were extracted using a modified Bligh-Dyer (Bligh and Dyer, 1959), with the first extraction using 250 ml of 4:10:5 water:methanol:dichloromethane followed by two extractions with 4:10:5 2.5% trichloroacetic acid:methanol:dichloromethane. Samples were disrupted with a sonicator probe for 20 minutes then centrifuged at 5000 g for 15 minutes. Phase separation of the supernatant was induced with 20 ml of dichloromethane and 20 ml of water, with the organic phase removed. The aqueous phase was extracted with the addition of 20 ml of dichloromethane two additional times and the pooled organic phases were dried under N2. Cold acetone precipitation was then used to separate the intact polar lipids (IPL) from the neutral lipids. This was done by bringing the sample up in 2 ml of DCM and then adding 40 ml of ice-cold acetone and then storing at -20℃for 24 hours. The precipitated intact polar lipids were separated by centrifuging at 9000 g for 15 minutes. The separated IPL fraction was treated with acidified methanol for 12 hours and then derivatized with BSTFA and pyridine.

Derivatized samples were analyzed on a gas chromatography mass spectrometer (Thermo Trace 1310) to identify lipids in the IPL fraction. Samples were injected using splitless injection onto a fused silica column (Restek DB-5, 60 m 0.2 mm, 25 µm) with the column heated from 60℃ to 320℃ at 6℃ per minute. Archaeol and hydroxyarchaeol were identified by comparing to published spectra (Hinrichs et al., 1999, Hinrichs et al., 2000a).

column (Restek DB-5, 30 m, 0.25 mm, 25 µm). Once separated, compounds were combusted to carbon dioxide with a nickel platinum catalyst with 1% oxygen in helium at 1020℃. Stable isotope ratios were measured with a Finnegan Mat 252 with isotope values reported in the delta notation relative to the VPDM scale (equation 3). The isotopic offset was corrected for using Mix B standard (n-C16 to n-C30 alkanes, Arndt Schimmelmann, Indiana University) with standards of n-C38 and n-C41 treated as unknowns to determine the error. Precision and accuracy were 0.2‰ and 0.5‰ for Hydrate ridge (n=4) and 0.7‰ and 0.4‰ for Santa Monica Basin samples (n=13).

BSTFA adds three methyl groups to the hydroxyl groups on archaeol and hydroxy-archaeol. The values of these methyl groups were determined by derivatizing, phthalic acid standard (δ13C -27.21‰ Sigma-Aldrich, isotope value confirmed via Arndt Schimmelmann, Indiana University) and using mass balance to determine the isotope value of the BSTFA groups (equations 12 and 13). With the value of the BSTFA groups, the true value of the lipids can be determined from the measured value (equation 14). The BSTFA groups have a δ13C -39.75‰ and a standard deviation of 0.5‰ (n=3). Error for the lipids is a combination of accuracy and precision determined from the standards and the standard deviation in the BSTFA group measurement (equation 15). Total analytical uncertainty is 0.8 ‰ for Hydrate Ridge and 1‰ for Santa Monica Basin lipids

�∑ = δ � + �(1 − � ) (12)

� = �∑ − � /(1 − � ) (13)

� = � − ��/(1 − �) (14)

(�∑ ) = (�) + (�) + (�) (15)

3.3.8. Estimation of newly synthesized F430 Using the nitrogen isotope value of the experiment and the isotope value of experiment 6, the killed controls, the amount and isotope signature of newly synthesized F430 can be determined. Calculations use F values in order to account for the contribution of the 13C addition to the total carbon inventory of analyzed components (Moran et al., 2005). M represents the absolute abundance of the common isotope (12C and 14N) and M* represents the absolute abundance of the rare isotope (13C and 15N) which are enriched in the experiments. R* is the isotope ratio that is enriched due to the addition of 13C and 15N labels. δ values are first converted to R values with the R* indicates that the rare isotope is enriched (equation 16).

�∗ = + 1 � (16)

R values are converted to F values with the following equations:

∗ � = (17) (∗)

∗ � = (18) ∗

� = (19)

Newly synthesized F430 was determined from the 15N incorporation in experiments relative to the F430 in 15 the killed controls, using the following equations. Where � is the measured N value for F430 from the 15 killed control, �is the N value observed for F430 in the experiments and � is the value of the nitrogen label added to the experiments.

� = 1 − � � + �� (20)

� = � − �� + �� (21)

� = � + �(� − �) (22)

� = (23)

Once the amount of newly synthesized F430 is calculated (�), the fraction of each carbon isotope label assimilated can be determined for each experiment. The 13C incorporation is determined for newly synthesized F430 and reveals the amount of methane or DIC assimilated into the F430 structure. The equations below illustrate that

calculation for methane; similar calculations can also be used to determine the amount of labeled DIC assimilated. We Assumed the proportion of newly synthesized lipid was similar to F430 in order to determine the amount of DIC and methane assimilated into lipids.

� = � − �� = �� + �� (24)

� = (� − 1 − � �) � (25)

() � = (26) ()

3.4. Results In the Hydrate Ridge experiments, the amounts of F430 were recovered (1 to 160 µg/g), were less than had been previously observed in the Hydrate Ridge sediment from chapter 2. The unlabeled control had the highest F430 concentration with 160 µg/g wet sediment (table 3-3). Only trace amounts of F430 were present in the 13C DIC experiments, with not enough recovered for an isotope measurement. TAG sequencing data showed that ANME-1 was the most abundant ANME group in the Hydrate Ridge experiments (table 3-4).

In the Santa Monica basin experiments, substantially more F430 was recovered than in the Hydrate Ridge sample set (table 3-5). F430 amounts ranged from 275 µg/g in experiment 1C (13C methane) to 4059 µg/g in 5B (Unlabeled control). This indicates that production of F430 was greater in Santa Monica Basin experiments, for which profiles of HS- documents active methane oxidation (figure 3-8). δ13C of the DIC in the labeled methane experiments confirms that methane is being oxidizes at it increase to 30,000‰ during the over the course of the experiment (figure 3-9). TAG sequencing data showed that ANME-2 were the more abundant in the Santa Monica Basin experiments than the Hydrate Ridge (table 3-4, 3-6).

In both the Hydrate Ridge and Santa Monica Basin experiment sets, experiment 3 was constructed to stimulate methanogenesis, while experiment 4 was designed to test if methane is essential for ANME to survive. For Hydrate Ridge samples, no F430 was recovered in experiment 3 or 4. Santa Monica Basin sediment experiments 3 and 4 were not extracted for F430 or lipids because the hydrogen sulfide concentration indicated that there was no activity (figure 3-8). Even though ANME can perform methanogenesis (Bertram et al., 2013) none of the 13C from DIC (experiment 3) or acetate (experiment 4) was detected as methane in the headspace (table 3-7). This indicates that F430, produced in experiments 1 and 2, was only used to oxidize methane, and this observation strengthens the theory that methanotrophy takes place via the reversal of methanogenesis.

The Santa Monica Basin experiments contained enough F430 for nitrogen isotope measurements to be made (figure 3-3, 3-5, 3-7). The 15N label from ammonium was picked up in the experiments using 13C methane, 13C DIC, and the unlabeled controls, indicating that new F430 had been synthesized since the start of the experiments (figure 3-10). In the killed control experiments, the average δ15N of F430 was 0‰, indicating that no new F430 had been synthesized. δ15N was variable between the replicate experiments, and for experiments where more F430 was recovered δ15N values were higher (figure 3-10). In the labeled methane experiments, 1A (2419 µg/g, δ15N 206‰) and 1B (1898 µg/g, δ15N 434‰) contained more F430 and it was more enriched in 15N than in experiment 1C, where only 275 µg/g of F430 was recovered with a δ15N of 156‰ (figure 3-11). A similar enrichment pattern was observed in the unlabeled controls with F430 in 5C (4059 µg/g, δ15N 428‰), more abundant and more enriched in 15N than in either 5A (704 µg/g, δ15N 28‰) or 5B (426 µg/g, δ15N 73‰).

In the experiments with Hydrate Ridge sediment, carbon isotope values were obtained for experiments 1A and 5 only, due to the small amount of F430 recovered. The δ13C values of F430 in the labeled methane and the unlabeled control experiments were isotopically similar to each other, -19‰ and -23‰ respectively (figure 3-12). This indicates that the ANME in these experiments did not assimilate carbon from methane. Hydrate Ridge archaeol from the labeled methane experiment was only 20‰ more enriched than the unlabeled control, indicating a limited amount of methane assimilation. Archaeol from labeled DIC experiments had a value of 119‰, indicating DIC was assimilated (table 3-3) into lipids.

In the Santa Monica Basin experiments, for both the labeled methane and DIC experiments, F430 was enriched in 13C compared to both the unlabeled and the killed control (figure 3-13, 3-14). F430 in the labeled methane experiments was, on average, more enriched in 13C (129‰, 21‰ and 82‰) than in the DIC experiments (- 29‰ and -38‰). Average archaeol values for the labeled methane experiment were -37‰ (1A), -41‰ (1B) and - 7‰ (1C), whereas values for the unlabeled controls were -107‰, -115‰ and -96‰, similar to the killed control value, -116‰. Lipids in the labeled DIC experiments were slightly more enriched in 13C (-98‰ -84‰ -100‰) than lipids in the unlabeled and killed controls, and this observation indicates carbon from the DIC pool was assimilated into archaeol (figure 3-15).

Using the nitrogen isotope results, a percentage of newly synthesized F430 was determined for the Santa Monica experiments. Newly synthesized amounts of F430 ranged from 0.2% in 5A to 4% in 1B (table 3-5). The proportion of newly synthesized F430, as determined from the nitrogen isotope values, was greater for DIC than methane. Isotope values and the amount of labeled substrate assimilated were determined using these estimates for the percentage of newly synthesized F430. In the labeled methane experiments, newly synthesized F430 δ13C values range from 2,400‰ (1B) to 12,500‰ (1C) with the assimilation of 5 (1B) to 20 (1C) % of methane. The background DIC value in these experiments become ~30,000‰ (figure 3-9) during the course of the experiment and as a result we cannot be sure that methane is directly assimilate. Methane may first be oxidized to DIC and then assimilated, this cannot be determined as both methane and DIC pools are enriched. Newly synthesized F430 in the 13C DIC experiments ranged from 1,300‰ (2C) to 1,400‰ (2B) and show the assimilation of ~70% from DIC (figure 3-16).

The enrichment of the DIC pool decreases during the course of the experiment from ~1900‰ to ~900‰. As a result, the amount of carbon determined to be assimilated from the DIC pool increases to 100%.

Assuming the amount of newly synthesized archaeol was similar to the amount of newly synthesized F430 in each experiment, the amount of 13C label from DIC and methane assimilated into lipids can be calculated. In the labeled methane experiments, newly synthesized archaeol had δ13C values from 2,900‰ (1A) to 3,800‰ (1C), and indicates that ~20% of the 13C label from methane has been assimilated. Like for F430 due to the amount of 13C label in the DIC pool (figure 3-9) we cannot determine whether the label was directly assimilated or was first oxidized to DIC before being assimilated. Archaeol in the labeled DIC experiments had δ13C value of 600‰ (2C) to 1,400‰ (2B), and reflects the assimilation of 35-70% of carbon from DIC (figure 3-16). Roughly 40% of lipid carbon is either from a source other than DIC and methane or the result of error, potentially underestimation of lipid production, in the calculation. Like for F430 in the labeled DIC experiments when the final isotope value for DIC is used (~900‰) the amount of DIC assimilated increases to 100% for each individual label DIC experiment.

3.5. Discussion The greater abundance of coenzyme F430 in the Santa Monica Basin experiments than the Hydrate Ridge experiments suggests AOM was more active in experiments using the Santa Monica Basin sediment (table 3-8). The incorporation of the 15N label from ammonium into the F430 structure in the Santa Monica Basin experiments confirms this and indicates synthesis of new F430 (figure 3-10, 3-13, 3-14) during the incubation period. Not enough F430 was synthesized in the Hydrate Ridge experiments to confirm new synthesis.

There is environmental and genetic evidence that ANME can perform methanogenesis (Bertram et al., 2013, Hallam et al., 2003). Experiments 3 and 4 without a methane headspace were constructed in an attempt to stimulate methanogenesis in the sediment. The absence of methane accumulation in the headspace during incubation of these experiments showed that methanogenesis did not take place. It also indicated that F430 produced in experiments 1 and 2, with methane in the headspace, and F430 extracted from the sediment in chapter 2 was associated with AOM only. This strengthens the link between AOM and F430 that was previously indicated by abundance profiles and geochemical data in chapter 2.

Even though the Hydrate Ridge experiments had limited growth, they showed that carbon from the DIC pool can be assimilated into lipids in experiments where 13C bicarbonate was added. In experiments with 13C methane, none of the label was assimilated into archaeol (figure 3-12). This is similar to previous labeling experiments with ANME-1, where DIC was preferentially assimilated (Kellermann et al., 2012). Lipids with 13C depleted isotope values have been previously reported for sediment where ANME-1 were abundant (Aloisi et al., 2002, Boetius et al., 2000, House et al., 2009, Orphan et al., 2001b, Reitner et al., 2005, Treude et al., 2007). The low isotopic values for cells and lipids reported in these settings may indicate ANME-1 assimilated highly 13C- depleted DIC in pore water adjacent to cells as a result of the complete oxidation of methane. Alternatively, it could suggest cells exhibit pronounced isotope fractionation during lipid synthesis.

Santa Monica basin experiments show the assimilation of carbon from methane and DIC into coenzyme F430 and lipids (figure 3-13,14,15). The more enriched values of F430 and lipids in the labeled methane experiments suggest that more methane was assimilated. However, this is a result of the high proportion of 13C in the added methane, and when the newly synthesized amount of F430 was accounted for and used to determine the % uptake of methane and DIC, we found more DIC (~70% to 100%) was assimilated than methane (0% to ~20%). Similarly, when we assumed the newly synthesized amounts of lipids is the same as F430, we estimated lipids are derived from of 0% to ~7% methane, ~50% to 100% DIC and when the starting DIC value (~1900‰) is used ~40% from an unknown source.

A significant amount of labeled methane was oxidized to DIC (figure 3-9), and as a result, the isotopic abundances of DIC became highly 13C-enriched. If the initial isotopic composition of methane is used in mass- balance calculations, at most, ~20% (F430) to ~7% (lipids) of methane was assimilated into these compounds. However, because the isotopic label is no longer exclusively associated with methane, we cannot be sure whether methane or DIC was the assimilated substrate, and the results are indeterminate for direct methane uptake. The magnitude of methane oxidized (roughly 50% of the methane label) relative to biomass produced is striking, and illustrates how AOM activity can modify the pore water DIC pool in its immediate surroundings. These results suggest methane carbon may be introduced into biomass only after it is oxidized to DIC and then assimilated.

The δ13C value of DIC decreases during the course of the experiments from ~1900‰ to ~900‰. If the initial, and less enriched value is used in to calculation assimilation, contribution from the DIC pool is ~70% (F430) and ~50% (lipid). When the end value is used (~900‰.) the estimate for both compounds reaches 100%. Since the calculated values represent newly formed amount over the course of the experiment and range of DIC values, 70% (F430) and 50% (lipid) can be viewed as the minimum amount assimilated from DIC, and the likely DIC contribution was likely closer to 100%.

Assimilated amounts of methane and DIC were determined from separate methane or DIC experiments, and by assuming that the proportion of newly synthesized lipids was the same as F430. There is likely a greater amount of newly synthesized archaeol than the co-enzyme, as their production may not scale in a simple fashion. Archaeol is major component of the lipid membrane (Hinrichs et al., 1999), whereas F430 is only present in the active site of methyl-coenzyme M reductase (Hallam et al., 2004, Kruger et al., 2003). However, the influence of greater lipid production should cancel if the greater proportion of new lipids relative to F430 was the same for both experiments. Alternatively, if there could be a lower production of lipids relative to F430, possibly due to recycling of the isoprenoid chains into lipids, a process that has been shown to occur in deep sea Archaea (Takano et al., 2010).

Using the percentages of DIC and methane assimilated the mass balance with fractionation model (equation

29) can be rearranged to determine the relationship between αDIC and αmethane for ANME biomolecules (equations 33 and 34). The slope is dependent on the isotope value of methane and DIC with the intercept dependent on the observed isotope value the fraction assimilated and the delta value of DIC or methane. Using these equation, the

relationship between α that explain the previously reported isotope values in the hydrate Ridge and Santa Monica Basin sediment can be explored.

(δ + 1) α/ = (27) (δ + 1)

�/ = �/ − 1 28

� � = �(�� + � − 1 + �(�� + � − 1 (29)

Where � � = � � ��

−�� − �� + � + � + � � � = (30) � �� + �

−�� − �� + � + � � � = + (31) � �� + � � �� + �

−�� − � � + � + � � � = � ∙ + (32) � �� + � � �� + �

−�(� � − 1) � + � + � � � = � ∙ + 33 � � � + 1 � � � + 1

−�(� � − 1) � + � + � � � = � ∙ + (34) � � � + 1 � � � + 1

By using the average observed lipid values from natural sediments (-100‰ for the Hydrate Ridge and - 120‰ for the Santa Monica basin) sediment, we can use equations 33 and 34 to solve for fractionation factors

associated with lipid production from each carbon substrate (DIC or CH4). For a given observed delta value and a given ratio of uptake from the two substrates, the two alpha values are inversely linked. In other words, as one of the α values increases, the other α value must decrease. Similarly, fractionation factors are linked for F430 from the two carbon substrate if the uptake ratio is set, in order to match calculated values with the observed average F430 (- 65‰) from the Santa Monica Basin sediment.

Our observations from labeled experiments indicate that DIC is directly assimilated into ANME cells, with methane carbon contributing to the DIC pool after it is oxidized. Therefore, the main function of reverse methanogenesis serves is to generate energy or electrons, but not to assimilate carbon and build biomass.

DIC could be assimilated into the ANME cell via the methanogenic machinery that has been previously identified in ANME. In chapter 2 we suggested that the methanogenic pathway is operated in two different directions with the conversion of methylene-H4MPT to methyl-H4MPT being a point that divides the pathway. Based on the evidence of chapter 3, it is likely that methane is oxidized to DIC via the complete reversal of the methanogenic pathway, and potentially generates up to eight electrons per methane molecule for the ANME cell (Taupp et al., 2010). Oxidized methane would then mix with DIC in the surrounding pore water before it is assimilated into the cell as acetyl-CoA using CO dehydrogenase/acetyl-CoA synthase like in methanogens (Hallam et al., 2004, Matschiavelli et al., 2012, Taupp et al., 2010). Figure 3-17 shows the possible flow of carbon within the ANME cell.

Preferential selection of 12C in the formation of isoprenoids would leave the acetyl-CoA that enters other pathways to form glutamate, the basic building block of F430, enriched in 13C. This could explain why F430 was significantly enriched relative to lipids at both sample sites in chapter 2. Evidence from Takano et al. (2010) suggests that some lipid carbon may be derived from recycling of lipids in the environment. The amount of recycling could decrease the relative proportion of acetyl-CoA used to construct lipids, and we might predict that cells with more recycling might have a somewhat lower 13C content in F430.

A wide range in the isotopic composition of ANME lipids and cells have been previously reported in sediment from the Eel River Basin and the Black Sea (House et al., 2009, Orphan et al., 2001b, Treude et al., 2007). ANME- 1 δ13C values ranges from -24‰ to -87‰ with ANME-2 ranging from -18‰ to -75‰ in the Eel River sediment (House et al., 2009). Our labelled experiment results indicate that mainly DIC is assimilated by both ANME 1 and ANME 2 dominated communities. The wide range of isotopic values observed for AOM biomass and lipids suggests the isotopic value of the DIC pool varies substantially. Mixing of oxidized methane into the pore water DIC pool near ANME cells would result in the isotopic value of assimilated DIC being viable and dependent on the rate of oxidation and the amount of mixing.

3.6. Conclusions The association between F430 and AOM observed in chapter 2 is strengthened by the results of the labeling experiments. F430 is recovered from experiments 1 and 2 which have a methane headspace, while experiments designed to stimulate methanogenesis failed to produce either methane or F430. Assimilation of DIC in addition to the 13C label from methane occurred in experiments using Santa Monica Basin sediment. Due to the label accumulating in the DIC pool in labeled methane experiments we cannot be sure whether methane is directly assimilated or first oxidized to DIC. Newly synthesized amounts of F430 and archaeol indicating significant proportions of each is derived from DIC (50% to 100%), with a limited amount of from methane (0% to 20%). Archaeol results indicate that potentially a source other than DIC or methane may account for ~50% of the carbon assimilated into the molecule. This gap might reflect limitations in the estimates of new lipid production during the incubation period, perhaps due to differences in lipid and coenzyme production rates, and leaves open the suggestion that lipid recycling takes place in ANME as has been previously shown to be important for deep sea Archaea (Takano et al., 2010). DIC appears to be the main carbon source for the ANME cell given the evidence for direct assimilation from the DIC pool in labeled experiment. The function of reverse methanogenesis within the cell may be to generate energy or election and not to assimilate methane. DIC is potentially assimilated into acetyl-CoA using CO dehydrogenase/acetyl-CoA synthase, with the observed isotopic difference between F430 and lipids in chapter 2 due to fractionation of the acetyl-CoA pool. Based on our observation, the variable isotope record that is observed for ANME-1 and ANME-2 is due to the mixing of oxidized methane and DIC. What remains unknown is if there is any direct assimilation of methane or what form DIC is assimilated as (CO2(aq), bicarbonate or carbonate). Changes in the design of the labeling experiment to potential use flow through reactors could help resolve some of these questions.

3.7. Acknowledgements We thank Denny Walizer for help in the lab and with the Nano-EA IRMS. Anne Dekas, Stephanie Connon and Jennifer Glass are thanked for sample collection. Stephanie Connon is thank for help with experimental setup. This research was funded by Royal Dutch Shell Geosciences Energy Research Facilitation Awards, PSARC and the American Chemical Society petroleum research fund.

3.8. Figures and tables

Figure 3-1: PCKD core. This core from the Santa Monica basin that was used in the labeling experiments. The sediment was homogenized with artificial sea water before being added to the incubation bottles.

−5 Cups OEP Methionine Sucrose −10 F430 13C methane experiments F430 control experiments OEP regressionline Methionine regression line Sucrose regression line −15

−20 delta 13C

−25

−30

−35 0 0.2 0.4 0.6 0.8 1 1.2 1.4 1/n Figure 3-2: Hydrate Ridge 1/n carbon plot. F430 from the Hydrate Ridge experiments plots between -20‰ and - 25‰ indicating that no 13C label has been assimilated.

15 Cups OEP Methionine F430 13C methane experiments 10 F430 control experiments OEP regressionline Methionine regression line

0 delta 15N

−5

−10

−15 0 0.005 0.01 0.015 0.02 0.025 0.03 0.035 0.04 0.045 0.05 1/n Figure 3-3: Hydrate Ridge 1/n nitrogen plot. The two enriched sample points indicate that new F430 was being synthesized but due to a high background these values cannot be corrected for

150 Cups OEP Methionine Sucrose 100 F430 13C methane experiments F430 13C bicarbonate experiments F430 control experiments F430 killed control solvent 50 OEP regressionline Methionine regression line Sucrose regression line delta 13C 0

−50

−100 0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 1/n Figure 3-4: Santa Monica 1/n carbon plot. 1/n carbon plot for F430 from the Santa Monica basin experiments, showing 13C enrichment in some samples

140 Cups OEP 120 Methionine F430 13C methane experiments F430 13C bicarbonate experiments 100 F430 control experiments F430 killed control solvent 80 OEP regressionline Methionine regression line

60 delta 15N

−20 0 0.02 0.04 0.06 0.08 0.1 0.12 0.14 1/n Figure 3-5: Santa Monica 1/n nitrogen plot. 1/n nitrogen plot for F430 from the Santa Monica basin experiments., the enriched values show that new F430 has been synthesized

80 Cups OEP 60 Methionine Sucrose F430 13C methane experiments 40 F430 13C bicarbonate experiments F430 killed control 20 solvent OEP regressionline Methionine regression line 0 Sucrose regression line

delta 13C −20

−40

−60

−80

−100 0 0.1 0.2 0.3 0.4 0.5 0.6 0.7 0.8 0.9 1 1/n Figure 3-6: Santa Monica 1/n carbon plot. Second 1/n carbon plot for F430 from the Santa Monica basin experiments, showing 13C enrichment in some samples

140 Cups OEP 120 Methionine F430 13C methane experiments F430 13C bicarbonate experiments 100 F430 killed control solvent OEP regressionline 80 Methionine regression line

60 delta 15N

−20 0 0.02 0.04 0.06 0.08 0.1 0.12 0.14 0.16 0.18 1/n

Figure 3-7: Santa Monica 1/n nitrogen plot. Second 1/n nitrogen plot for F430 from the Santa Monica basin experiments, the enriched values show that new F430 has been synthesized

10 13C Methane (PCKD 1)

9 13C DIC (PCKD 2) Methanogenesis (PCKD 3) 8 No methane headspace (PCKD 4) 7 Control (PCKD 5)

(mM) 5 - HS 4

0 0 20 40 60 80 100 120 Time (days)

Figure 3-8: Hydrogen sulfide for PCKD. Experiments 1,2 and 5 with a methane head space produce hydrogen sulfide indicating methanotrophy was taking place in the experiments. No HS- was generated in experiments 3 and 4 indicating there was not activity.

35,000 13C DIC

methanogeneis 30,000 No methane head space

Control 25,000 Killed control

13C Methane 20,000 DIC -

δ 13C 15,000

10,000

5,000

0 0 20 40 60 80 100 120 time (days)

Figure 3-9: δ13C-DIC for PCKD experiments. In the 13C labeled methane experiments the label from methane is oxidized enriching the DIC pool.

15N F430 PCKD 900

800

700

600

500

400 delta 15N 300

200

100

−100 13C Methane 13C DIC Control Killed control Experiment

Figure 3-10: Santa Monica F430 nitrogen results. Nitrogen isotope results for F430 are enriched relative to the killed control. All experiments had 15N labeled ammonium added to determine the amount of newly synthesized F430. The black lines represent the mean value.

delta 13C and delta 15N for F430 450 Experiment 13C Methane 400 13C DIC Control 350

300

250

200 F430 delta 15N

150

100

0 0 500 1000 1500 2000 2500 3000 3500 4000 4500 F430 ug/g Figure 3-11: Uptake of nitrogen into F430. A higher d15N value is observed when more coenzyme F430 is recovered from samples.

150 Archaeol

100 F430

δ13C 0

-50

-100 13C Methane 13C DIC Control Experiment

Figure 3-12: Hydrate Ridge Carbon isotope results. Carbon isotope values obtained for both F430 and IPL archaeol are shown for Hydrate Ridge experiments

13C F430 PCKD 300

250

200

150

100 delta 13C

−50

−100 13C Methane 13C DIC Control Killed control Experiment

Figure 3-13: Santa Monica F430 carbon values. Carbon isotope results for F430 from the experiments with Santa Monica sediment, both the methane and DIC experiments show some assimilation of the 13C label. The black lines represent the mean value.

delta 13C and delta 15N for F430 Experiment 900 13C Methane 13C DIC 800 Control Killed control 700

600

500

400 delta 15N 300

200

100

−100 −100 −50 0 50 100 150 200 250 300 delta 13C

Figure 3-14: Santa Monica F430 carbon and nitrogen isotope data. F430 data shows that the methane and DIC are enriched relative to the killed controls

13C Archaeol PCKD 0

−20

−40

−60 delta 13C

−80

−100

−120 13C Methane 13C DIC Control Killed control Experiment

Figure 3-15: Santa Monica archaeol carbon values. Carbon isotope results for archaeol from the experiments using the Santa Monica sediment. Experiments with labeled methane are more enriched than those with labeled DIC, unlabeled and the killed control. The black lines represent the mean value.

100% Methane 90% DIC 80% Unknown 70%

60%

50%

40%

30%

20%

10%

0% F430 Lipid

Figure 3-16: The carbon assimilation for F430 and lipids. The assimilated amounts are determined from the newly synthesized F values of F430 and archaeol. Using the starting isotope value for methane (88,000‰) and DIC (1,900‰), F430 is composed of ~70% DIC, with ~30% unaccounted for and Archaeol is composed of ~50% DIC with ~50% unknown and possibly due to lipid recycling (Takano et al., 2010). When the final DIC value (900‰) is used the amount of DIC assimilated into both F430 and lipids increases to 100%

Figure 3-17: Carbon flow within the ANME cell. Methane is not directly assimilated but first oxidized to DIC and then assimilated into the cell

Table 3-1: Labeling Experimental set up. These parameters were used to construct both the Hydrate Ridge and the Santa Monica basin samples

Experiment Methane DIC Acetate No methane Unlabeled Killed 1 2 3 4 5 6 Replicates 3 3 3 3 1 1 Hydrate ridge Replicates 3 3 3 3 3 2 Santa Monica Mud (g) 50 50 50 50 50 50 Total vol (ml) 150 150 150 150 150 150 13 C HCO3 no Yes no Yes no no

H2/CO2 no no yes no no no 13C acetate no no Yes no no no 13C methane Yes no no no no Yes 15 NH4 500 µM 500 µM 500 µM 500 µM 500 µM 500 µM

Table 3-2: Isotope measurement error.

Hydrate Ridge Santa Monica Basin

σ13C σ15N σ13C σ15N σ13C σ15N Octaethylporphine 0.9 12.6 0.9 2.9 0.7 3.5 Sucrose 2.3 0.7 2.0 Methionine 1.4 5.5 0.5 1.8 0.9 1.7

σoffset 23.3 10.9 1.0 3.0 1.5 3.6

σblank 1.3 1.4 0.6 1.0 1.2 0.4

σ∑ 4.6 17.7 1.7 4.6 3.0 5.3

Table 3-3: Hydrate Ridge carbon isotope results.

Hydrate Ridge Experiment µg/g δ13C Archaeol 1A 21 -19 -48 1B 1 2A 3 120 4A Trace 5 160 -23 -77 6 -

Table 3-4: Hydrate Ridge TAG sequencing. The table shows the number of tags for each ANME group. Results are for the Hydrate Ridge experiment in which F430 was recovered

Hydrate Ridge Experiments Taxon 1A 1B 2A 2B 5 ANME-1a 105 141 104 120 79 ANME-1b 32 58 22 36 19 ANME-1; Other 81 250 71 106 65 ANME-2b 6 47 11 11 17 ANME-2a-2b 4 8 2 6 3 ANME-2a-2b; Other 0 1 0 0 0 ANME-2c; 21 54 22 14 4

ANME-3 0 1 0 0 0

Table 3-5: Santa Monica Basin carbon and nitrogen isotope results

Santa Monica Basin F430 Archaeol Experiment µg/g δ15N δ13C % New δ13C New % label δ13C New % label assimilated assimilated 1A 2419 206 129 2 10078 20 -38 2880 6 1B 1898 434 21 4 2427 5 -42 3386 7 1C 275 156 82 1 12517 25 -8 3830 8 2A -99 818 44 2B 3974 358 -29 3 1414 73 -85 1366 70 2C 2578 402 -38 3 1282 66 -100 631 35 5A 704 29 -66 0.2 -108 5B 426 74 -55 1 -115 5C 4059 429 -78 4 -97 6 9017 3 -78 -116

Table 3-6: Santa Monica TAG sequencing. The table shows the number of tags for each ANME group. Results are for the Santa Monica experiment in which F430 was recovered

Taxon 1A 1B 1C 2A 2B 2C 5A 5B 5C ANME-1a 40 135 30 100 114 117 125 98 149 ANME-1b 9 9 18 11 21 25 14 14 17 ANME-2a-2b;Other 0 1 0 0 2 0 0 0 0 ANME-2b 4 23 26 9 14 5 16 8 11 ANME-2a-2b 532 451 681 368 465 238 616 320 297 ANME-2c 2628 3110 3558 1780 1799 1439 2876 1576 1588 ANME-3 339 522 499 333 245 156 289 187 152

Table 3-7: Headspace analysis of PCKD experiments 3 and 4

Headspace methane Experiment uM ppm 3A average 3.69 103 σ 0.30 8 3B average 3.18 89 σ 0.39 11 3C average 3.53 99 σ 0.14 4 4A average 2.05 57 σ 0.15 4 4B average 2.31 65 σ 0.36 10 4C average 1.89 53 σ 0.39 11

Table 3-8: Experiment results summary.

Hydrate Ridge Santa Monica Basin 15N-F430 29‰ to 434‰ Below detection New growth New growth confirmed No new growth confirmed 1-4% Labeled Methane F430 21‰ to 129‰ F430 -19‰ Lipids -42‰ to -8‰ Lipids -48‰ Methane assimilated ~20% Assimilated - Labeled DIC F430 -38‰ to -29‰ F430 - Lipids -100‰ to -85‰ Lipids 119‰ DIC assimilated ~70% assimilated Unlabeled control F430 -78‰ to -55‰ F430 -23‰ Lipids -115‰ to -97‰ Lipids -77‰ No CH4 experiments No growth No growth ANME taxa ANME-2 ANME-1

Table 3-9: α and ε values used to model the assimilation of DIC and methane into F430 and lipids

F430

100% DIC αF430/DIC εF430/DIC 100% Methane αF430/methane εF430/methane Model -40 1 0 -150 0.91 -86.02 A -50 0.99 -10.42 -160 0.90 -96.77 B -60 0.98 -20.83 -170 0.89 -107.53 C Lipid

100% DIC αlipid/DIC εlipid/DIC 100% Methane αlipid/methane εlipid/methane Model -80 0.96 -41.67 -170 0.89 -107.53 D -90 0.95 -52.08 -180 0.88 -118.28 E -100 0.94 -62.50 -190 0.87 -129.03 F

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Chapter 4: Quantifying Bacteriohopanepolyol production in Little Salt Springs cyanobacteria

4.1. Abstract Hopanoids, one of the most ubiquitous geochemical compound in the geologic record, are used to identify ancient microbial life and characterize oil source rocks (Albrecht and Ourisson, 1969, Summons et al., 1999). Chemotaxonomic information comes from the position of a methyl group at C1, 2 or 3. The 2-methyl form conventionally indicates contributions from cyanobacteria (Summons et al., 1999). Yet, this interpretation has been challenged with the identification of other bacteria that produce 2-methyl bacteriohopanepolyols, the modern precursor of 2-methyl hopanoids and the absence of 2-methyl BHP producing cyanobacteria in the modern ocean (Pearson et al., 2007, Rashby et al., 2007, Sáenz et al., 2011b). Little Salt Springs is a marine influenced sinkhole in Florida where a benthic red biofilm produces large amounts of bacterialhopanetetrol (BHT), 2-methyl bacterialhopanetetrol (2-MeBHT) and 2-methyl Anhydrobacterialhopanetetrol (2-MeAnhydroBHT). The amount of each BHT produced is inconsistent and varies between years, with the geochemical cause of this variability unknown. A red cyanobacterium was isolated from the biofilm and used in a series of pure culture experiments to determine the geochemical control on production. The tested geochemical conditions all produce a similar amount of BHT and 2-MeAnhydorBHT and failed to generate 2-MeBHT or Anhydrobacterialhopanetetrol that had previously been identified in the biofilm samples. Only when two conditions were combined (limited light and not shaken) was more BHT and 2-MeAnhydroBHT produced than in the control experiments. The tested conditions don’t identify a single geochemical parameter that controls BHP production. However, the differences between the biofilm and pure culture experiments suggest that oxygen concentration and microbial community may be important in BHP production with these conditions, in addition to others such as fixed nitrogen species and trace metal, ideal targets for future experiments.

4.2. Introduction Hopanoids are abundant through the geologic record and can be used to identify source rocks, oil migration, and depositional environment. Interpretations of their biological origins are based on variations of the hopanoid structure, particular the position of a methyl group (Ourisson and Albrecht, 1992, Seifert and Moldowan, 1980). Insight into ancient microbial communities is based on the position of the methyl group at C-2 or C-3 on the hopanoid ring structure (Farrimond et al., 2004, Rashby et al., 2007, Summons et al., 1999). 2-Methyl hopanoids found widely in Proterozoic sediments, are conventionally interpreted to represent the presence of ancient cyanobacteria based on the high proportion of 2-methyl bacteriohopanepolyols (BHPs) in certain cultured cyanobacteria (Summons and Walter, 1990, Summons et al., 1999). Yet, this interpretation is challenged by genetic evidence that less than 10% of all modern bacteria are capable of producing BHPs, the identification of other bacteria that produce 2-methyl BHPs and no known marine cyanobacteria produce 2-methyl BHPs (Pearson et al., 2007, Rashby et al., 2007, Talbot et al., 2008).

Bacteriohopanepolyols (BHPs), the biochemical precursors to hopanoids, have been identified in cultures of cyanobacteria, methanotrophs, acetic acid bacteria and anaerobic photosynthesizers (Rashby et al., 2007, Rohmer et al., 1984, Summons et al., 1999). Of the 41 cyanobacterial species that produce BHPs, only 19 of these are able to produce 2-methyl BHPs, the biochemical precursor of 2-methyl hopanoids, in pure culture (Pearson et al., 2007, Talbot et al., 2008). Importantly, no marine cyanobacteria have yet been identified that produces any 2-methyl BHPs (Pearson et al., 2007, Talbot et al., 2008). Cyanobacteria belonging to Chroococcales, Nostocales, Oscillatoriales, Prochlorales and Stigonematales orders have been found to produce 2-Methyl BHPs in culture (Jahnke et al., 2004, Summons et al., 1999, Talbot et al., 2008). Production of 2-methyl BHPs is variable between species of the same order, with members of Chroococcales producing between 20 ug/g to 650 ug/g depending on the specific species (Summons et al., 1999). The limited amount of 2-methyl BHP produced by cyanobacteria and the absence of marine cyanobacteria that produces 2-methyl BHP’s cast doubt on the interpretation of 2-methyl hopanoids as a marker for ancient cyanobacteria.

Little Salt Springs, a marine-influenced and stratified sinkhole in Sarasota County, Florida, USA, with molar concentrations of sulfide and oxygen in the photic zone (figure 4-1), analogous to the Proterozoic ocean. Within the sinkhole, a red biofilm is found on sediment surface in the upper 20 m of the water column been found to produce BHPs, including baceriohopanetetrol (BHT), 2-methyl-baceriohopanetetrol (2-MeBHT), anhydrobacteriohopanetetrol (AnhydorBHT) and 2-methyl-anhydro-bacteriohopanetetrol (2-MeAnhydroBHT). This biofilm is dominated by Cyanobacteria, Chlorobi and sulfate-reducing clades of Deltaproteobacteria, with the recovery of a single hpnP gene indicating that they cyanobacteria are the source of the 2-methyl BHP’s in the biofilm (Hamilton et al., Submitted). 2-MeBHT and 2-MeAnhydroBHT are significantly more abundant (~105 µg/g TLE in biofilm samples) than in 2-methyl BHPs in other environmental samples (~50 µg/g TLE) or observed in pure culture species (1 to 650 µg/g TLE) (Sáenz et al., 2011a, Summons et al., 1999). BHPs vary in biofilm samples from different years (figure 4-2), which is likely linked to seasonal changes in the geochemistry of the water column.

The geochemical condition that causes the cyanobacteria in the red biofilm to produce large, but variable amounts of 2-MeBHT and 2-MeAnhydroBHT is currently unknown. Growing the isolated BHP-producing cyanobacteria (Hamilton et al., Submitted) under a variety of different geochemical conditions, could help resolve the discrepancy observed in the biofilm. Previous culturing experiments of other cyanobacteria taxa indicated that a number of different conditions, such as temperature, pH, and growth under anoxic conditions can effect BHP production (Joyeux et al., 2004, Poralla et al., 1980, Rashby et al., 2007, Schmidt et al., 1986). This prior work suggests potentially a number of different geochemical conditions influence the amount and structures of BHPs in the biofilm. Here, we use culture experiments in an effort to identify the condition or conditions that account for BHP variability in the biofilm. Such data may help resolve the discrepancy between the large amount of 2-Methyl hopanoids in ancient marine sediments and their absence in modern oceans (Summons and Walter, 1990, Summons et al., 1999).

The red biofilm is dominated by filamentous cyanobacteria, belong to the Oscillatoriales order, which was isolated to investigate the controls on BHP production (Hamilton et al., Submitted). Other members of the

Oscillatoriales order produce 2-methy BHP forms in amounts significantly less than those of the biofilm (Jahnke et al., 2004, Talbot et al., 2008). Culture experiment using the Little Salt Springs cyanobacteria were set up to explore how salt, sulfide, sulfide with photosystem 2 inhibited, thiosulfate, agitation, available light and lack of a fixed nitrogen source effect BHP production. We find that when in pure culture the cyanobacteria produced less BHT and 2-MeAnhydro BHT than the biofilm. Additionally, 2-MeBHT and anhydorBHT that had been previously identified in samples of the biofilm (Hamilton et al., Submitted) were not present in any of the cultures. No clear geochemical control on BHP production was identified, as most experiments produced similar amounts. Never the less, the results of the light-limited and not shaken experiments suggest that a number of conditions may interact with each other to control BHP production.

4.3. Methods

4.3.1. Field Samples Little Salt Spring is a sinkhole lake located in Sarasota County, FL (lat. 27°04'30"N, long. 82°14'00"W). The sunlit upper basin hosts the red microbial biofilm at the sediment-water interface, which blooms seasonally. Samples of the red biofilm were collected by divers in June of 2012 from the sediment-water interface in the upper basin at a depth of 8.5 m. Biofilm samples were preserved in RNAlater (2 parts per sample volume; Ambion/Applied Biosystems, Foster City, CA, USA) and stored on ice until frozen (within 6 hours).

Water column samples were collected using a 3.8L Van Dorn bottle (Wildlife Supply Company, Yule, FL) deployed from a platform at approximately the center of the sinkhole. Dissolved sulfide concentration was measured immediately in the field once the water sample was returned to the surface, with a portable spectrophotometer (Hach Co., Loveland, CO), using methylene blue for total sulfides (Hach method 690, detection limit ~ 0.2 µM). Water depth, temperature, pH, and dissolved oxygen were measured in situ using a calibrated mulit parameter YSI 6600 sonde probe (Yellow Springs, OH, USA). The temperature of the water column averages ~27 °C and has a salinity of ~3 ppt. Profiles of the water column are presented in figure 4-1

4.3.2. Cultures The control isolate was maintained in BG11 media (Rippka et al., 1979) supplemented with 25 mM HEPES (B-HEPES) and adjusted to pH 7.2 in 125 mL conical flasks at 100 RPM at 28 ºC under a day-night cycle with 100 µmol photons m-2 S-1 supplied by cool white fluorescent lamps. Control samples were harvested after 5, 10, 15, and 20 days. In the not shaken experiment shaking the flask at 100 RPM was omitted from the set up. The light limited experiment 100 µmol photons m-2 S-1 was lowered to 20 mol photons m-2 s-1. The high salt experiment followed the control experiment setup with salinity adjusted to 3.5% to mimic sea water. The high salt, not shaken, light limited and not shaken light limited cultures were harvested after 20 days. The sulfide, DCMU, and thiosulfate experiments were grown until log phase. Cells were then re-suspended in media containing sulfide (50 µM), DCMU (10 µM), or thiosulfate (50 µM) and incubated for 3 hours then harvested (table 4-1).

For growth without fixed nitrogen, nitrate was omitted from the media. Concentration of nitrate and ammonia in the media were monitored daily using a Hach DR 1900 spectrophotometer (Loveland, CO) and the cadmium reduction method and the salicylate method, respectively. The concentration of ammonia was below the detection limit (0.01 mg/L) at all-time points. Nitrate concentration was maintained at ~1 mM by addition of a sterile solution of NaNO3. Growth was monitored with chlorophyll a concentration determined spectrophotometrically using the absorption at 665 nm of a methanol extract and an extinction coefficient of 0.075 ml µg−1 (made from a filtered 2-ml culture subsample) (de Marsac and Houmard, 1988) or protein concentration using the Bradford assay (Bradford, 1976) with bovine serum albumin (Sigma-Aldrich, St. Louis, MO) as the standard. Samples were harvested at 5, 10, 15 and 20 days (table 4-2) and immediately frozen at -20ºC until analyzed.

4.3.3. Extraction Cell pellets were freeze-dried and a biomass weight obtained using a microbalance before being extracted using a modified Bligh-Dyer method as described by Talbot et al. (2007). Samples were submerged in a monophasic solution of 4:10:5 water:methanol:dichloromethane, disrupted with a sonicator bath for 1 h at 400C, shaken at 200 rpm for 1 h, and centrifuged at 9000 rpm for 15 min. The supernatant was removed and the extraction repeated twice. 10 ml of dichloromethane and 10 ml of water were added to the pooled supernatant to induce phase separation, and the organic phase was removed. The aqueous phase was extracted with 10 ml of dichloromethane two additional times, and the pooled total lipid extract (TLE) phases were dried under N2 and weighed.

4.3.4. Acetylation 20% of the sample TLE was transferred to a 2 ml vial and then were spiked with an internal standard of pregnanediol for a concentration of 500 ng per 50 µl injection. The spiked samples were dried under N2 then acetylated using 50 µl of acetic anhydride and 50 µl of pyridine, heated at 60°C for 1 hour as outlined in Blau and Halket (1993). After acetylation samples were dried and re-suspended in 500 µl of methanol for HPLC analysis.

4.3.5. LCMS Analysis BHPs were analyzed using an Agilent 6310 Ion Trap LC/MS system after Talbot et al. (Talbot et al.,

2003). Chromatographic separation was achieved on a Phenomenex Gemini C18 column (5 µm particle size, 150 mm x 3.0 mm i.d.) and 5 µm guard column containing the same solid phase with the following solvent gradient profile: initial 10%A and 90%B to 1%A, 59%B, and 40%C at 25 min; isocratic at this composition until 40 min; and a final ramp returning to 10%A and 90%B at 42 min (where A= water, B= methanol and C= isopropyl alcohol).

1300, and the run divided into 3 segments targeting m/z 285 (0-10 min), 1002 (10-17 min), and 655 (17-50 min). Automatic MSn settings for 2 precursor ions were as follows: absolute threshold 100,000, relative threshold 5%, and fragment amplitude 1.0V. Auto MSn settings for single precursor ions were: absolute threshold of 1,000, relative threshold 5%, and fragment amplitude 1.0V. Ions were excluded from auto MSn after 2 counts and released after 0.5 min. Detection was achieved at an isolation width of 3.0 m/z units and a fragment amplitude of 1.0 V.

4.3.6. Compound identification Compounds were identified by comparison with published MS spectra from Talbot et al. (2003), Talbot et al. (2007). The molecular ions used for identification are as follows: BHT (m/z 655, figure 4-3), 2-Me BHT (m/z 669, figure 4-4), and 2-Me Anhydro BHT (m/z 869, figure 4-5). Methylation at the C-2 position was previously confirmed via oxidative cleavage and GCMS analysis (Albrecht, 2011).

4.3.7. Quantification To quantify the amounts of BHT and 2-Methyl AnhydroBHT in the culture, response curves for known amount of each were produced relative to 500 ng of pregananediol. Standards were purified from cultured biomass using the extraction method previously mentioned. Fractions were collected from the C18 column and then further purified with a second dimension of HPLC using a normal phase silica column (150mm x 30mm, Resteck) and an isocratic flow of 9:1 isopropyl alcohol : hexane. The collected fractions for BHT and 2-Methyl AnhydroBHT were then transferred to weighed 2 ml vials to obtain a mass of each standard (BHT 390 µg +- 10 µg, 2-Methyl AnhydroBHT 350 µg +- 10 µg).

Response factors were produced by running 100 ng, 200 ng, 500 ng, 1000 ng and 2000 ng of standard of each BHP in addition to samples. Peak areas of BHT and 2-MeAnhydroBHT were normalized to the internal standard (500ng pregananediol) in order to account for instrument drift between injections. A linear regression line of the ratio of the known amounts (BHT ng / pregnanediol ng) against the peak area ratio (BHT response / pregnanediol). Unknown amount of BHT and 2-MeAnhydroBHT were determined using the slope and intercept from the linear regression line and to convert the response ratio of the unknown amounts to the ng ratio which is converted to the known amount as the amount of pregenanediol added is known. A regression line is produced for each BHT compound as the two compounds as the response for 2-MeAnhydoBHT is an order of magnitude greater than BHT (figures 4-6 and 4-7). Due to the use of APCI source, which is open to the atmosphere, there is daily variation in the ionization of the standards due to environmental factors such as temperate and humidity. As a result, the linear regression line is produced for every sequence of samples with figures 4-6 and 4-7 showing the daily variation in July 2016 and between a sequence from December 2015. Multiple standards of the same amount are also run to assess variance during a sequence. R2 is used to determine how well the regression line fits the data, with R2 produced for BHT regression lines varying between 0.91 and 0.99 indicating a good fit. Similar R2 values are obtained for 2-MeAnhydoBHT regression lines (0.93 to 0.99) and also indicate a good fit between the data and the

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regression line. Each unknown sample is run and corrected for three times to assess the reproducibility with standard deviation used to assess the variability between each run.

4.4. Results In the biofilm sample, three BHP were identified, BHT, 2-MeBHT and 2-MeAnhydroBHT, but only BHT and 2-MeAnhydroBHT were present in the red cyanobacterium isolated from the Little Salts springs sinkhole (figure 4-2). A green cyanobacterium was also isolated in pure culture but did not produce any BHP compounds. Due to the presence of sediment within the biofilm amounts are reported in µg/g total lipid extract (TLE) (Table 4-3). BHT (110,000 to 130,000 µg/g TLE), 2-MeBHT (120,000 to 130,000 µg/g TLE) and 2-MeAnhydroBHT (60,000 to 150,000 µg/g TLE) were all present in similar amounts and much more abundant that in the pure cultures of the red cyanobacteria (BHT 120 to 540 µg/g TLE, 2-MeAnhydroBHT ~60 µg/g TLE). AnhydroBHT that had been previously identified in samples of the biofilm taken in January 2009 (Albrecht, 2011) was not present in biofilm samples from June 2012 or October 2013.

In only the light-limited not shaken experiments, was more BHT and 2-MeAnhydroBHT produced than in the control experiments (figure 4-8). 4340 to 7050 µg/g biomass of BHT and 1050 to 1400 µg/g biomass of 2- MeAnhydroBHT were recovered compared to 330 to 1040 µg/g biomass of BHT and 40 to 220 µg/g biomass of 2- MeAnhydroBHT in the control experiments. More of BHT and 2-MeAnhydorBHT have been produced, but the 2- methyl ratio is 0.2 similar to 0.3 in the control (table 4-4, figure 4-8).

The thiosulfate and not shaken experiments produced less BHT and 2-MeAnhydroBHT than the control experiments (figure 4-8 and 4-9). No BHT or 2-MeAnhydroBHT were produced in the not shaken experiments with no BHT produced in the thiosulfate experiments. Only 2-MeAnhydroBHT was produced in the thiosulfate experiments (50 to 250 µg/g biomass) in similar amounts to the control experiments (40 to 220 µg/g biomass). As a result of no BHT being produced in the thiosulfate experiments the 2-Methyl ratio is 1, higher than the controls.

The light limited shaken, sulfide, sulfide and DCMU, and high salt experiments all produced BHT and 2- MeAnhydroBHT in similar amounts to the control experiments (figure 4-8, 4-9, 4-10). BHT amounts were 370 to 1550 µg/g biomass in the light limited shaken experiments, 210 to 1150 µg/g biomass in the sulfide experiments, 340 to 1430 µg/g biomass in sulfide and DCMU experiments and 280 to 1080 µg/g biomass in the high salt experiments. Amounts of 2-MeAnhydroBHT were 230 to 750 µg/g biomass in the light limited shaken experiments, 60 to 180 µg/g biomass in the sulfide experiments, 60 to 90 µg/g biomass in sulfide and DCMU experiments and 80 to 270 µg/g biomass in the high salt experiments. The high salt, sulfide and sulfide and DCMU experiments all have 2-methyl ratio that were lower than the controls. In the light limited shaken experiments the ratio was higher than the controls, similar to the light limited not shaken experiments (figure 4-11 and table 4-4).

In the control time series experiments similar amounts of BHT (220 to 520 µg/g biomass) and 2- MeAnhydroBHT (50 to 240 µg/g biomass) were recovered at each time point (5, 10, 15 and 20 days) (figure 4-12).

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There does not appear to be any change in the amounts of BHT or 2-MeAnhydroBHT with growth stage, with similar amounts recovered in the lag phase (day 5, 350 µg/g biomass BHT, 210 µg/g biomass 2-MeAnhydroBHT) and stationary phase (day 20, ~330 µg/g biomass BHT, 120 µg/g biomass 2-MeAnhydroBHT). In the limited nitrogen experiments amounts of BHT recovered at time point 4 (20 days, 530 to 720 µg/g biomass) were found to be significantly different than time point 1 (5 days, 90 to 270 µg/g biomass) and time point 4 (290 to 390 µg/g biomass) in the controls experiments (figure 4-13). These experiments were determined to be significantly different at a 95% confidence level, using a t test. Similar amounts of 2-MeAnhydroBHT (20 to 240 µg/g biomass) were recovered at all 4 time points in the nitrogen and control experiments with a t test showing none of time points were significantly different from each other. In the control experiments the 2-methyl ratio is similar between all 4 time points (figure 4-14). The 2-methyl ratio in the limited nitrogen experiments becomes lower at each subsequent time point (figure 4-15). T test results indicated that the ratios between day 5 and 20 is significantly different at a 95% confidence level. When the t test is performed on the other time points the null hypothesis cannot be rejected, meaning any difference is not statistically significant.

4.5. Discussion The pure culture experiments using the red cyanobacteria were unable to reproduce the high abundance of BHT or 2-MeAnhydorBHT observed in the biofilm. Further, only BHT and 2-MeAnhydrobBHT was present in culture samples, while BHT, 2-MeBHT, anhydroBHT, 2-MeAnydroBHT and an unknown BHP were all present in the biofilm (figure 4-2). In the high salt, sulfide, sulfide and DCMU and nitrogen time series experiments, BHT and 2-MeAnhydroBHT were present in similar amounts to the controls, with abundance lower in the thiosulfate and not shaken experiments. These conditions can be ruled out as being the source of variability in the biofilm samples as they don’t change BHP production in the pure culture experiments. High salt experiments indicate that exposing the cyanobacteria to salt levels similar to the modern ocean does not inhibit BHP production. Only when grown under limited light conditions and not shaken did the production of both BHT and 2-MeAnhydroBHT increase. No clear geochemical control on BHP production can be identified and our results indicate that production is either controlled by multiple geochemical conditions or by a factor that was not tested by the experimental design.

Although experiments didn’t change BHP production relative to the controls, some of the experimental conditions may still stimulate production in other bacteria, as BHP production by different species is variable. BHP production has been shown to increase at lower pH in cultures of Alicyclobacillus acidocaldarius but not in Rhodopseudomonas palustris (Poralla et al., 1980, Welander et al., 2009). High salt, sulfide, sulfide and DCMU and no fixed nitrogen could affect BHP production in other bacteria, with nitrogen fixation shown to increase production in Frankia mycelia (Berry et al., 1993). However, they can be ruled out as affecting production for the Little Salt Spring cyanobacterium grown under oxic conditions.

Oxygen concentration is one condition that potentially affects BHP production that was not tested. The red biofilm grows in part of the water column (8-14 m) where oxygen levels are below 1 mg/l (figure 4-1). Evidence from the geological record also suggests that oxygen concentration is important for BHP production, as 2-methyl

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hopanoids are abundant in Proterozoic sediments a period of widespread ocean anoxia (Canfield, 1998, Johnston et al., 2009, Summons and Walter, 1990, Summons et al., 1999). The effects of low oxygen concentrations on BHP production could not be tested, as cultures did not grow well without oxygen. Changing the experimental design to periodically stress the cultures with low oxygen conditions could be a way to explore the effects of oxygen concentration on the cyanobacterium. It is unclear why the red cyanobacteria were able to grow under low oxygen conditions in the environment but not in the laboratory. This could be due to the community of other microbes in the biofilm changing the local environment around the red cyanobacteria, allowing it to grow.

Potentially there is a microbial community aspect to BHP production with growth in a biofilm or microbial mat resulting in a greater amount of 2-methyl BHP being produced. In addition to the large amounts of 2-methyl BHP’s being identified in the Little Salt Springs biofilm, 2-methyl BHP’s are observed in microbial mats from Yellowstone hot springs (Jahnke et al., 2004, Summons et al., 1999) and hypersaline lake on Kiritimati (Blumenberg et al., 2013). Possibly the increase in the production of 2-methyl BHPs by the red cyanobacteria in the Little Salt springs biofilm is a response to other microbes in the biofilm with the observed BHP variability between sampling dates due to changes in community composition. BHPs could be a used by the cyanobacteria in the mat to interact with other species in the biofilm.

In the limited nitrogen source time series experiments, no increase in BHP production is observed relative to the control time series experiments. The biofilm, with more abundant BHP’s, grows in part of the column where fixed nitrogen is not limited with both ammonium and nitrate. Previously BHP’s have been inferred to be used in nitrogen fixation, protecting nitrogenase enzymes from oxygen (Berry et al., 1993). The difference between the limited nitrogen and biofilm samples suggest that increase in production in the biofilm is a response to the fixed nitrogen species. This could be tested in future experiments with the cyanobacteria grown with additional nitrate, nitrite, and ammonium. Ammonia has been shown to damage photosystem II in the cyanobacterium Synechocystis (Drath et al., 2008) and to decrease cyanobacteria growth (Dai et al., 2008). Potentially the cyanobacteria in the biofilm produce large amounts of BHT and 2-MeAnhydroBHT to reduce the membrane permeability to ammonium. The production of BHPs in response to ammonium is sported by the abundance of hopanoids in Proterozoic sediments (Summons et al., 1999, Summons and Walter, 1990), a period of time when ammonium is likely to have been abundant due to ocean anoxia (Anbar and Knoll, 2002).

BHPs are thought to be bacterial equivalent of sterols in eukaryotes and used to adjust membrane fluidity and permeability (Kannenberg and Poralla, 1999). Using BHPs to lower membrane permeability is potentially a way the microbial cell protects itself from toxins in the environment (Doughty et al., 2009, Rashby et al., 2007, Sáenz et al., 2012). The large BHP amounts produced in the red biofilm maybe a response of the cyanobacteria to a geochemical condition that is toxic and present in much larger amounts in the Little Salta springs water column than the culture media. Ammonium, metals such as nickel and copper can be toxic to cyanobacteria and oxygen to cyanobacterial nitrogenase (Berry et al., 1993, Dai et al., 2008, Drath et al., 2008, Mann et al., 2002, Martínez-Ruiz and Martínez-Jerónimo, 2016). BHPs may be produced to decrease the permeability of the cyanobacteria membrane to these toxins, protecting the cell in extreme environments.

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The results of the light limited not shaken, not shaken and light limited experiments indicate that when certain conditions are combined they can have an effect on BHP production. Individually not shaking and limiting light do not increase the production of either BHT or 2-MeAnhydroBHT relative to the controls. When they are combined significantly more of BHT and 2-MeAnhydroBHT was produced. This indicates that production cannot be attributed to a single geochemical control, and multiple conditions act together to increase production. Potentially, experiments with both sulfide and high salt could also stimulate the production of BHPs, even though individually similar amounts of BHPs are produced to the controls. To resolve this issue more experiments could be constructed to test individual conditions in combination with each other (e.g. sulfate plus limited light). Additionally, the main geochemical control could be something that was not tested in this experiment set and future experiments could widen the test conditions to include low oxygen, fixed nitrogen species, and metals (nickel and copper).

The absence of 2-MeBHT and AnhydroBHT in the culture samples is useful in determining the source of 2- MeAnhydroBHT. AnhydroBHT has previously been thought to be a diagenetic or analytical artifact when identified previously (Eickhoff et al., 2014, Handley et al., 2010). The presence of 2-MeAnhydroBHT in the pure cultures without 2-MeBHT or AnhydroBHT indicates that it is being produced by the cyanobacteria. AnhydoBHT can be formed from BHT-cyclitol ether and BHT-glucosamine in addition to BHT (Schaeffer et al., 2010), but the absence of the 2-methyl forms of these BHT in the culture experiments indicates that 2-meAnhydroBHT is being produced by the red cyanobacteria and is not a diagenetic or analytical artifact.

4.6. Conclusions In conclusion, the large amounts of 2-meBHT and 2-methyl AnhydroBHT in the Little Salt Springs red biofilm were not reproduced in pure culture experiments. Experiments using the red cyanobacteria isolated from the biofilm produce similar amounts of BHT and 2-meAnhdroBHT except when two conditions were combined (light limited not shaken experiments). A definitive geochemical control on the variability observed between samples of the biofilm cannot be determined from these experiments. Future experiments should explore what effects oxygen concentration, community composition, combinations of different geochemical parameters and additional parameters like nitrogen species and trace metals have on BHP production. Potentially production cannot be linked to a single geochemical control and maybe due a combination of factors. Understanding the variability observed in the biofilm will potentially shed light on their geological source and abundance in Proterozoic marine sediments

4.7. Acknowledgements We thank Denny Walizer for help in the lab and with the LC-MS and Heidi Albrecht for help with compound identification. John Gifford (U. Miami) and divers Steve Koski (U. Miami), and Rick Riera-Gomez (U. Miami) are thanked for their help with sample collection

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4.8. Figures and tables

Figure 4-1: Water column geochemistry for Little Salt Springs June 2012. The light pink area represents the section of the water column that the red biofilm is found in.

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6 Biofilm and Cyanobacteria controls 10 BHT 2−MeBHT AnhydroBHT 5 10 2−MeAnhydroBHT Unknown

4 10

ug/g TLE 3 10

2 10

1 10

control APR Biofilm; 2009 Biofilm 2012 Biofilm 2012 Biofilm 2013 Control DEC

Figure 4-2: Biofilm and cyanobacterial BHP content. Four samples of the Little salt springs biofilm have been collected and analyzed. The 2009 sample was collected in January at a water depth of 8.5 m with abundance data reported in Albrecht (2011), the 2012 samples were collected in June at a depth of 13.7 m. One sample was frozen after collection and the other was air dried which has potentially resulted in the loss of some 2-MeAnhydroBHT. The 2013 sample was collected in October at a water depth of 10 m. Results for the pure culture grown in April and December 2015 are also shown and indicate that there is significantly more of the BHPs in the biofilm.

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Figure 4-3: Bacterialhopenetetrol mass spectrum

Figure 4-4: 2-methyl bacterialhopenetetrol mass spectrum

Figure 4-5: 2-methyl anhydro bacterialhopenetetrol mass spectrum

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0.8 BHT 12-Jul-16

0.7 20-Jul-16 14-Jul-16 y = 0.1565x - 0.0124 9-Dec-15 R² = 0.96234 0.6 Linear (12-Jul-16) y = 0.1517x + 0.0008 Linear (20-Jul-16) R² = 0.95489

0.5 Linear (14-Jul-16)

Linear (9-Dec-15) y = 0.0995x + 0.0202 0.4 R² = 0.93505 Observedratio

0.3

0.2

y = 0.0247x + 0.0118 0.1 R² = 0.91477

0 0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 ratio of known ng amount

Figure 4-6: BHT MS response. The response of the BHT standard varies between sequences meaning that the standard curve has to be produced for each sequence run for accurate quantification

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18 2-Me Anhydro BHT 12-Jul-16 16 20-Jul-16 14-Jul-16 14 9-Dec-15 y = 3.2573x + 0.1798 Linear (12-Jul-16) R² = 0.93633 12 Linear (20-Jul-16) Linear (14-Jul-16) y = 2.4805x - 0.1804 10 Linear (9-Dec-15) R² = 0.9767

8 Observed ratio ratio Observed y = 2.2209x + 0.1342 R² = 0.97521 6

2 y = 0.0889x + 0.0117 R² = 0.95375 0 0 0.5 1 1.5 2 2.5 3 3.5 4 4.5 ratio of known ng amount

Figure 4-7: 2-MeAnhydroBHT MS response. The response of the 2-MeAnhydroBHT standard varies between sequences meaning that the standard curve has to be produced for each sequence run for accurate quantification

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Figure 4-8: Results of limited light and shaken experiments. The amounts of BHT and 2-MeAnhydroBHT are similar to the control in the limited light shaken experiments. In the limited light not shaken experiments more of both BHT and 2-MeAnhydroBHT were present. In the not shaken experiments neither BHT and 2-MeAnhydroBHT were present.

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Figure 4-9: Results of sulfur species experiments. The amounts of BHT and 2-MeAnhydroBHT produced in the sulfur species experiments are similar to the control experiments. Less BHT and 2-MeAnhydroBHT are produced in the thiosulfate experiments.

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Figure 4-10: Results of high salt experiments. The amounts of BHT and 2-MeAnhydroBHT produced in the high salt experiments are similar to the control experiments.

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Figure 4-11: 2-Methyl ratio in culture experiments. Ratios are similar to the control except the thiosulfate experiments where no BHT is produced. n=9 for each time point.

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Figure 4-12: Results of control time series experiments. The amounts of BHT and 2-MeAnhydroBHT are similar at all time points Control experiments growth curve.

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Figure 4-13: Results of no fixed nitrogen time series experiments. The amounts of BHT and 2-MeAnhydroBHT are similar at all time points Nitrogen experiments growth curve.

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Control experiments 1

0.9

0.8

0.7

0.6

0.5

0.4 ratio 2ME/(2ME+BHT)

0.3

0.2

0.1

0 5 10 15 20 days Figure 4-14: Ratio in control cultures. The ratio of 2-MeAnhydroBHT to BHT is consistent between time points in the control culture experiments (n=9 for all time points).

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Nitrogen experiments 1

0.9

0.8

0.7

0.6

0.5

0.4 ratio 2ME/(2ME+BHT)

0.3

0.2

0.1

0 5 10 15 20 days Figure 4-15: Ratio in no fixed nitrogen cultures. The ratio decreases after 5 days and which using a t test is shown to be significantly different than day 20 (n=9 for all time points).

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Table 4-1: Little Salt Springs cyanobacterium growth conditions Sulfide Limited Green High Limited Not Experiment Control Sulfide and Thiosulfate light not control Salt light shaken DCMU shaken LSS Red Green Red Red Red Red Red Red Red cyanobacterium Replicates 6 1 3 3 3 3 3 3 3 25 mM 25 mM 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM Media b-hepes b-hepes hepes hepes hepes hepes hepes hepes b-hepes pH 7.2 7.2 7.2 7.2 7.2 7.2 7.2 7.2 7.2 Temperature 28 28 28 28 28 28 28 28 28 (ºC) Light (µmol photons 100 100 100 100 100 100 20 20 100 m-2 S-1) Shaken 100 100 100 100 100 100 100 0 0 (RPM) 50uM Addition to 3.5% 50uM Sulfide 50 uM - - - - - media salinity Sulfide 10uM thiosulfate DCMU Harvested 20 20 20 20 20 20 20 20 20 (days) BHT BHT BHT BHT BHT BHT 2- BHPs 2- 2- 2- 2- 2- 2- None MeAnhydro None present MeAnhy MeAnhydr MeAnhydr MeAnhydr MeAnhydro MeAnhydr BHT droBHT oBHT oBHT oBHT BHT oBHT

Table 4-2: Little Salt Springs cyanobacterium growth conditions Control Control Control Control Nitrogen Nitrogen Nitrogen Nitrogen Experiment t=1 t=2 t=3 t=4 t=1 t=2 t=3 t=4 LSS Red Red Red Red Red Red Red Red cyanobacterium Replicates 3 3 3 3 3 3 3 3 Harvested 5 10 15 20 5 10 15 20 (days) 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM b- 25 mM b- Media hepes hepes hepes hepes hepes hepes hepes hepes pH 7.2 7.2 7.2 7.2 7.2 7.2 7.2 7.2 Temperature 28 28 28 28 28 28 28 28 (ºC) light (µmol photons m-2 S- 100 100 100 100 100 100 100 100 1) Shaken 100 100 100 100 100 100 100 100 (RPM) Nitrate 1mM 1mM 1mM 1mM Not added Not added Not added Not added

Ammonium 1.5 µg 1.5 µg 1.5 µg 1.5 µg Not added Not added Not added Not added BHT BHT BHT BHT BHT BHT BHT BHT 2- 2- 2- 2- 2- 2- 2- 2- BHPs present MeAnhy MeAnhydro MeAnhydr MeAnhydr MeAnhydr MeAnhydro MeAnhydro MeAnhydro droBHT BHT oBHT oBHT oBHT BHT BHT BHT

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Table 4-3: Water column, biofilm and cyanobacteria results µg/g TLE Sample Depth m BHT 2-Me BHT AnhydoBHT 2-MeAnhydro BHT Unknown Water column 2-30 - - - 2 - Red biofilm January 2009 8.5 100 100 117000 176000 - Red biofilm June 2012 13.7 110000 130000 - 60000 - Red biofilm June 2012 13.7 130000 120000 - 150000 - Red biofilm October 2013 10 - - - 13000 43000 Red cyanobacteria - 330 - - 60 - Green cyanobacteria ------

Table 4-4: Culture experiments results. µg/g biomass Sample Replicates BHT 2-methyl anhydro BHT Ratio Control 6 570 205 0.26 High Salt 3 617 142 0.19 Sulfide 3 716 107 0.13 Sulfide and DCMU 3 732 73 0.09 Thiosulfate 3 4 146 0.97 Limited light 3 813 504 0.38 Limited light not shaken 3 6126 1245 0.17 Not shaken 3 - - -

4.9. References ALBRECHT, H. L. 2011. BACTERIOHOPANEPOLYOLS ACROSS ENVIRONMENTAL GRADIENTS. Ph.D, The Pennsylvania State University. ALBRECHT, P. & OURISSON, G. 1969. Triterpene alcohol isolation from oil shale. Science, 163, 1192-1193. ANBAR, A. D. & KNOLL, A. H. 2002. Proterozoic Ocean Chemistry and Evolution: A Bioinorganic Bridge? Science, 297, 1137-1142. BERRY, A. M., HARRIOTT, O. T., MOREAU, R. A., OSMAN, S. F., BENSON, D. R. & JONES, A. D. 1993. Hopanoid lipids compose the Frankia vesicle envelope, presumptive barrier of oxygen diffusion to nitrogenase. Proceedings of the National Academy of Sciences, 90, 6091-6094. BLAU, K. & HALKET, J. 1993. Handbook of derivatives for chromatography, Wiley New York. BLUMENBERG, M., ARP, G., REITNER, J., SCHNEIDER, D., DANIEL, R. & THIEL, V. 2013. Bacteriohopanepolyols in a stratified cyanobacterial mat from Kiritimati (Christmas Island, Kiribati). Organic Geochemistry, 55, 55-62. BRADFORD, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Analytical Biochemistry, 72, 248-254. CANFIELD, D. E. 1998. A new model for Proterozoic ocean chemistry. Nature, 396. DAI, G., DEBLOIS, C. P., LIU, S., JUNEAU, P. & QIU, B. 2008. Differential sensitivity of five cyanobacterial strains to ammonium toxicity and its inhibitory mechanism on the photosynthesis of rice-field cyanobacterium Ge–Xian–Mi (Nostoc). Aquatic Toxicology, 89, 113-121. DE MARSAC, N. T. & HOUMARD, J. 1988. [34] Complementary chromatic adaptation: Physiological conditions and action spectra. Methods in Enzymology. Academic Press. DOUGHTY, D. M., HUNTER, R. C., SUMMONS, R. E. & NEWMAN, D. K. 2009. 2-Methylhopanoids are maximally produced in akinetes of Nostoc punctiforme: geobiological implications. Geobiology, 7, 524- 532.

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DRATH, M., KLOFT, N., BATSCHAUER, A., MARIN, K., NOVAK, J. & FORCHHAMMER, K. 2008. Ammonia Triggers Photodamage of Photosystem II in the Cyanobacterium Synechocystis sp. Strain PCC 6803. Plant Physiology, 147, 206-215. EICKHOFF, M., BIRGEL, D., TALBOT, H. M., PECKMANN, J. & KAPPLER, A. 2014. Diagenetic degradation products of bacteriohopanepolyols produced by Rhodopseudomonas palustris strain TIE-1. Organic Geochemistry, 68, 31-38. FARRIMOND, P., TALBOT, H., WATSON, D., SCHULZ, L. & WILHELMS, A. 2004. Methylhopanoids: molecular indicators of ancient bacteria and a petroleum correlation tool. . Geochimica et Cosmochimica Acta, 68, 386-3882. HAMILTON, T. L., WELANDER, P. V., ALBRECHT, H. L., FULTON, J. M., SCHAPERDOTH, I., BIRD, L. R., SUMMONS, R. E., FREEMAN, K. H. & MACALADY, J. L. Submitted. Microbial communities and organic biomarkers in a Proterozoic-analog sinkhole environment. Geobiology. HANDLEY, L., TALBOT, H. M., COOKE, M. P., ANDERSON, K. E. & WAGNER, T. 2010. Bacteriohopanepolyols as tracers for continental and marine organic matter supply and phases of enhanced nitrogen cycling on the late Quaternary Congo deep sea fan. Organic Geochemistry, 41, 910-914. JAHNKE, L. L., EMBAYE, T., HOPE, J., TURK, K. A., VAN ZUILEN, M., DES MARAIS, D. J., FARMER, J. D. & SUMMONS, R. E. 2004. Lipid biomarker and carbon isotopic signatures for stromatolite-forming, microbial mat communities and Phormidium cultures from Yellowstone National Park. Geobiology, 2, 31- 47. JOHNSTON, D. T., WOLFE-SIMON, F., PEARSON, A. & KNOLL, A. H. 2009. Anoxygenic photosynthesis modulated Proterozoic oxygen and sustained Earth’s middle age. . Proceedings of the National Academy of Sciences, 106, 16925-16929. JOYEUX, C., FOUCHARD, S., LLOPIZ, P. & NEUNLIST, S. 2004. Influence of the temperature and the growth phase on the hopanoids and fatty acids content of Frateuria aurantia (DSMZ 6220). FEMS Microbiology Ecology, 47, 371-379. KANNENBERG, L. E. & PORALLA, K. 1999. Hopanoid Biosynthesis and Function in Bacteria. Naturwissenschaften, 86, 168-176. MANN, E. L., AHLGREN, N., MOFFETT, J. W. & CHISHOLM, S. W. 2002. Copper Toxicity and Cyanobacteria Ecology in the Sargasso Sea. Limnology and Oceanography, 47, 976-988. MARTÍNEZ-RUIZ, E. B. & MARTÍNEZ-JERÓNIMO, F. 2016. How do toxic metals affect harmful cyanobacteria? An integrative study with a toxigenic strain of Microcystis aeruginosa exposed to nickel stress. Ecotoxicology and Environmental Safety, 133, 36-46. OURISSON, G. & ALBRECHT, P. 1992. Hopanoids. 1. Geohopanoids: the most abundant natural products on Earth? . Accounts of Chemical Research 25, 398-402. PEARSON, A., FLOOD PAGE, S. R., JORGENSON, T. L., FISCHER, W. W. & HIGGINS, M. B. 2007. Novel hopanoid cyclases from the environment. Environmental Microbiology, 9, 2175–2188. PORALLA, K., KANNENBERG, E. & BLUME, A. 1980. A glycolipid containing hopane isolated from the acidophilic, thermophilic bacillus acidocaldarius, has a cholesterol-like function in membranes. FEBS Letters, 113, 107-110. RASHBY, S. E., SESSIONS, A. L., SUMMONS, R. E. & NEWMAN, D. K. 2007. Biosynthesis of 2- methylbacteriohopanepolyols by an anoxygenic phototroph. Proceedings of the National Academy of Sciences USA 104. RIPPKA, R., DERUELLES, J., WATERBURY, J. B., HERDMAN, M. & STANIER, R. Y. 1979. Generic Assignments, Strain Histories and Properties of Pure Cultures of Cyanobacteria. Microbiology, 111, 1-61. ROHMER, M., BOUVIER-NAVE, P. & OURISSON, G. 1984. Distribution of Hopanoid Triterpenes in Prokaryotes. Microbiology, 130, 1137-1150. SÁENZ, J. P., EGLINTON, T. I. & SUMMONS, R. E. 2011a. Abundance and structural diversity of bacteriohopanepolyols in suspended particulate matter along a river to ocean transect. Organic Geochemistry, 42, 774-780. SÁENZ, J. P., WAKEHAM, S. G., EGLINTON, T. I. & SUMMONS, R. E. 2011b. New constraints on the provenance of hopanoids in the marine geologic record: Bacteriohopanepolyols in marine suboxic and anoxic environments. Organic Geochemistry 42, 1351-1362. SÁENZ, J. P., WATERBURY, J. B., EGLINTON, T. I. & SUMMONS, R. E. 2012. Hopanoids in marine cyanobacteria: probing their phylogenetic distribution and biological role. Geobiology, 10, 311-319. SCHAEFFER, P., SCHMITT, G., ADAM, P. & ROHMER, M. 2010. Abiotic formation of 32,35- anhydrobacteriohopanetetrol: A geomimetic approach. Organic Geochemistry, 41, 1005-1008.

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SCHMIDT, A., BRINGER-MEYER, S., PORALLA, K. & SAHM, H. 1986. Effect of alcohols and temperature on the hopanoid content of Zymomonas mobilis. Applied Microbiology and Biotechnology, 25, 32-36. SEIFERT, W. K. & MOLDOWAN, J. M. 1980. The effect of thermal stress on source-rock quality as measured by hopane stereochemistry. Physics and Chemistry of the Earth 12, 229-237. SUMMONS, R. E., JAHNKE, L., HOPE, J. & LOGAN, G. 1999. 2-Methylhopanoids as biomarkers for cyanobacterial oxygenic photosynthesis. Nature, 400. SUMMONS, R. E. & WALTER, M. R. 1990. Molecular fossils and microfossils of prokaryotes and protists from Proterozoic sediments. American Journal of Science, 290A, 212-244. TALBOT, H. M., ROHMER, M. & FARRIMOND, P. 2007. Rapid structural elucidation of composite bacterial hopanoids by atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry. Rapid Communications in Mass Spectrometry 21, 880-892. TALBOT, H. M., SUMMONS, R. E., JAHNKE L. & PAUL FARRIMOND, P. 2003. Characteristic fragmentation of bacteriohopanepolyols during atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry. Rapid Communications in Mass Spectrometry, 17, 2788-2796. TALBOT, H. M., SUMMONS, R. E., JAHNKE, L. L., COCKELL, C. S., ROHMER, M. & FARRIMOND, P. 2008. Cyanobacterial bacteriohopanepolyol signatures from cultures and natural environmental settings. Organic Geochemistry, 39, 232-263. WELANDER, P. V., HUNTER, R. C., ZHANG, L., SESSIONS, A. L., SUMMONS, R. E. & NEWMAN, D. K. 2009. Hopanoids Play a Role in Membrane Integrity and pH Homeostasis in Rhodopseudomonas palustris TIE-1. Journal of Bacteriology, 191, 6145-6156.

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Chapter 5: Research summary

5.1. Chapter summaries In Chapter 2, a link between F430 and AOM was established, supporting the theory that the anoxic oxidation of methane by ANME takes place via the reversal of methanogenesis. This link was based on profiles of sulfide, sulfate, methane, F430 and ANME aggregate counts in Hydrate Ridge and Santa Monica Basin sediment. F430 and ANME were isotopically distinct, with F430 enriched in 13C relative to archaeal lipids. This enrichment was observed in both the Hydrate Ridge and Santa Monica Basin sediment. Potentially this enrichment is due to the assimilation of DIC in addition to methane. Variability was also observed in the archaeol δ13C values for Hydrate Ridge and for F430 and archaeol in Santa Monica Basin sediments. Potentially, variation in the assimilation of multiple carbon substrates accounts for the observed viability both within the two sample sites, and for previously observed isotopic variability at other locations (House et al., 2009, Orphan et al., 2002). The ability of ANME to assimilate multiple carbon substrates has been previously demonstrated in labeling experiments (Kellermann et al., 2012, Lloyd et al., 2011). Using mass balance with fractionation, we constrained the proportions of DIC and methane needed to account for the observed range in isotope values of F430 and lipids. The mass balance calculations show the observed isotope values from both Hydrate Ridge (F430 90-55% DIC, 5-10% methane; archaeol 10-40% DIC, 60-90% methane) and the Santa Monica Basin (F430 40-50%, DIC 50-60% methane; archaeol 10-40% DIC, 60-90% methane) can be generated by the assimilation of both methane and DIC.

In chapter 3, the biochemical flexibility of ANME in the Hydrate Ridge and Santa Monica Basin was further explored with a series of stable isotope labeling experiments. The link between F430 and AOM observed in chapter 2 was strengthened, with F430 recovered only from experiments which had methane in the headspace. The 13C label from both bicarbonate and methane was assimilated into F430 and lipids. When the newly synthesized amount of F430 was determined, we found more DIC (F430 ~70% to 100%, archaeol ~36% to 100%) was assimilated than methane (F430 0% to ~20%, archaeol 0% to ~7%). Due to the amount of labeled methane that enters the DIC pool we cannot determine whether methane is directly assimilated by the ANME. Our results indicate that mostly DIC is assimilated and that methane is assimilated after first being oxidized to DIC. The range in isotope values previously reported for ANME (House et al., 2009) is likely due to mixing between DIC and oxidized methane.

In chapter 4, pure culture experiments using a cyanobacterium isolated from the Little Salt Springs biofilm fail to reproduce the very high amounts or all the types of BHP’s found in the natural biofilm. Large, but variable amounts of bacterialhopanetetrol (BHT), 2-methyl bacterialhopanetetrol (2-MeBHT) and 2-methyl Anhydrobacterialhopanetetrol (2-MeAnhydroBHT) have been identified in the Little Salt Springs biofilm. Using the red cyanobacteria isolated from the biofilm experiments were constructed to identify the geochemical control on BHP production. Pure cultures produced BHT and 2-meAnhdroBHT, with similar amounts recovered in all

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experiments. A definitive geochemical control on the observed variability in the biofilm samples cannot be identified from these experiments, with future experiments needed.

5.2. Future directions

5.2.1. ANME biochemistry Our research established a link between coenzyme-F430 in the sediment and ANME, supporting the theory that methane is oxidized via the reversal of methanogenesis (Hallam et al., 2004, Moran et al., 2005, Scheller et al., 2010, Zehnder and Brock, 1979). To further support the reverse methanogenesis hypothesis other biomolecules used in the last step of methanogenesis that have not been linked to ANME could be targeted for quantification and isotope analysis. Identifying additional componets of the last step would support the reverse methanogenesis hyprothesis and the use of F430 in the sediment to oxidize methane. Additional F430 forms have been identifed in ANME dominated sediment with some belived to be used in processes other than the anoixic oxidation of methane (Allen et al., 2014). Coenzyme M is an ideal target, as if used in methanotrophy it should form methyl-coenzyme M with the methyl group coming from methane (Scheller et al., 2010). If, like F430, coenzyme M is enriched in 13C due to being synthesized from DIC, it should become depleted when methane is oxidized. Other components such as methyl-H4MPT and methylene-H4MPT, which genetic studies indicate are present in the ANME dominated sediment and used in earlier methanogenic steps (Hallam et al., 2004, Taupp et al., 2010), could be targeted in an attempt to follow methane through the reverse methanogenic pathway.

Hallam et al. (2004) identified the genes for all of the methanogenic steps, except for F420-dependent N5,

N10-methylenetetrahydromethanopterin reductase which converts methylene-H4MPT to methyl-H4MPT, in ANME dominated sediment. The biochemical components of these additional steps could be targeted for to determine what additional steps ANME in the sediment are using. To further investigate the assimilation of DIC via the methanogenesis pathway, formylmethanofuran dehydrogenase, which is the first step of methanogenesis, could be targeted for abundance and isotope analysis. Linking it to ANME in the sediment would support the theory proposed in chapter 2 that ANME use part of the methanogenic pathway in the methanogenic direction.

Isotope analysis of F430 and lipids indicates that ANME are metabolically flexible and assimilate DIC, in addition to methane, into their biomass. This metabolically flexibility could be further explored with additional labeled substrates to explore whether ANME can assimilate substrates other than DIC and methane, and if they recycle the hydrocarbon portion of their lipids. Labeled acetate and isoprenoids could be used to explore lipid recycling, something that has previously been identified in Archaea (Takano et al., 2010), and could explain the unknown carbon proportion in chapter 3. ANME have been shown to assimilate a number of other C1 compounds such as methanol (Bertram et al., 2013). Potentially some of the variability in the isotope record could be due to the assimilation of organic carbon substrates from the sediment, which could be explored with labeled compounds.

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The differences in lipid and F430 isotope characteristics observed between Hydrate Ridge and the Santa Monica Basin are potentially due to differences in the portions of AMNE-1 and 2 in the sediment. The more enriched values in the Hydrate Ridge sediment could be due to ANME-1 assimilating more DIC than ANME-2. Potentially, ANME-1 and 2 assimilate methane and DIC via different biochemical pathways, resulting in the observed ~60‰ difference for F430 and lipids between Hydrate Ridge and the Santa Monica basin. This could be resolved with additional stable isotope labeling experiments, once ANME-1,2 and 3 can be grown in pure culture.

5.2.2. Production of bacteriohopanepolyols Future experiments should explore what effects oxygen concentration, community composition, combinations of different geochemical conditions such as, nitrogen species and trace metals have on BHP production. Oxygen concentration is one condition that potentially effects BHP production that was not tested, as cultures did not grow without oxygen. Changing the experimental design to periodically stress the cultures with low oxygen conditions could be way to explore the effects of oxygen. There may also be a microbial community aspect to BHP production with growth in a biofilm or microbial mat resulting in a greater amount of 2-methyl BHP being produced as a response to other microbes. In addition to Little Salt Springs biofilm, 2-methyl BHP are observed in microbial mats from Yellowstone hot springs (Jahnke et al., 2004, Summons et al., 1999) and hypersaline lake on Kiritimati (Blumenberg et al., 2013).

Production is possibly not the result of one geochemical condition but a response to a number of environmental factors. The results of the light limited not shaken, not shaken and light limited experiments indicate that when certain conditions are combined they can increase BHP production. On their own, not shaking and limiting light do not increase the production of either BHT or 2-MeAnhydroBHT when compared to the controls. When they are combined more of BHT and 2-MeAnhydroBHT is produced. To explore this more experiments would have to be constructed testing two condition at the same time (e.g. sulfate and limited light).

5.3. References ALLEN, K. D., WEGENER, G. & WHITE, R. H. 2014. Discovery of Multiple Modified F430 Coenzymes in Methanogens and Anaerobic Methanotrophic Archaea Suggests Possible New Roles for F430 in Nature. Applied and Environmental Microbiology, 80, 6403-6412. BERTRAM, S., BLUMENBERG, M., MICHAELIS, W., SIEGERT, M., KRÜGER, M. & SEIFERT, R. 2013. Methanogenic capabilities of ANME-archaea deduced from 13C-labelling approaches. Environmental Microbiologyl, 15, 2384-2393. BLUMENBERG, M., ARP, G., REITNER, J., SCHNEIDER, D., DANIEL, R. & THIEL, V. 2013. Bacteriohopanepolyols in a stratified cyanobacterial mat from Kiritimati (Christmas Island, Kiribati). Organic Geochemistry, 55, 55-62. HALLAM, S. J., PUTNAM, N., PRESTON, C. M., DETTER, J. C., ROKHSAR, D., RICHARDSON , P. M. & DELONG, E. F. 2004. Reverse Methanogenesis: Testing the Hypothesis with Environmental Genomics. Science, 305, 1457-1462.

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HOUSE, C. H., ORPHAN, V. J., TURK, K. A., THOMAS, B., PERNTHALER, A., VRENTAS, J. M. & JOYE, S. B. 2009. Extensive carbon isotopic heterogeneity among methane seep microbiota. Environmental Microbiology, 11, 2207-2215. JAHNKE, L. L., EMBAYE, T., HOPE, J., TURK, K. A., VAN ZUILEN, M., DES MARAIS, D. J., FARMER, J. D. & SUMMONS, R. E. 2004. Lipid biomarker and carbon isotopic signatures for stromatolite-forming, microbial mat communities and Phormidium cultures from Yellowstone National Park. Geobiology, 2, 31- 47. KELLERMANN, M. Y., WEGENER, G., ELVERT, M., YOSHINAGA, M. Y., LIN, Y.-S., HOLLER, T., MOLLAR, X. P., KNITTEL, K. & HINRICHS, K.-U. 2012. Autotrophy as a predominant mode of carbon fixation in anaerobic methane-oxidizing microbial communities. Proceedings of the National Academy of Sciences, 109, 19321-19326. LLOYD, K. G., ALPERIN, M. J. & TESKE, A. 2011. Environmental evidence for net methane production and oxidation in putative ANaerobic MEthanotrophic (ANME) archaea. Environmental Microbiology, 13, 2548-2564. MORAN, J. J., HOUSE, C. H., FREEMAN, K. H. & FERRY, J. G. 2005. Trace methane oxidation studied in several Euryarchaeota under diverse conditions. Archaea, 1, 303-309. ORPHAN, V. J., HOUSE, C., HINRICHS, K.-U., MCKEEGAN, K. & DELONG, E. 2002. Multiple Archaeal Groups Mediate Methane Oxidation in Anoxic Cold Seep Sediments. P Natl Acad Sci Usa, 99, 7663-7668. SCHELLER, S., GOENRICH, M., BOECHER, R., THAUER, R. K. & JAUN, B. 2010. The key nickel enzyme of methanogenesis catalyses the anaerobic oxidation of methane. Nature, 465, 606-608. SUMMONS, R. E., JAHNKE, L., HOPE, J. & LOGAN, G. 1999. 2-Methylhopanoids as biomarkers for cyanobacterial oxygenic photosynthesis. Nature, 400. TAKANO, Y., CHIKARAISHI, Y., OGAWA, N., NOMAKI, H., MORONO, Y., INAGAKI, F., KITAZATO, H., HINRICHS, K. & OHKOUCHI, N. 2010. Sedimentary membrane lipids recycled by deep-sea benthic archaea. Nature Geoscience, 3, 858-861. TAUPP, M., CONSTAN, L. & HALLAM, S. J. 2010. The Biochemistry of Anaerobic Methane Oxidation. In: TIMMIS, K. N. (ed.) Handbook of Hydrocarbon and Lipid Microbiology. Springer Berlin Heidelberg. ZEHNDER, A. J. & BROCK, T. D. 1979. Methane formation and methane oxidation by methanogenic bacteria. J Bacteriol, 137, 420-32.

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Appendix A: F430 abundance and isotope values from the Santa Monica basin

A.1. Introduction In chapter 2 the isotopic composition of F430 was found to vary between different sampling locations. Here we investigate its F430 isotopic composition in a different core taken from the Santa Monica basin to assess local variation in the isotopic value of F430. Carbon Isotope values of lipids have been shown to vary within sample sites (House et al., 2009, Orphan et al., 2001) and variation in the value of F430 is expected within the Santa Monica sediment.

A.2. Methods

A.2.1. Shipboard collection, core processing, and sample storage Samples were collected from Hydrate Ridge, Oregon as part of the R/V Atlantis cruise 18-10 in September 2011 and from the Santa Monica Basin, California as part of the R/V Atlantis cruise 26-06 in October 2013. Using the ROV Jason II, sediment push cores were collected from seep environments characterized by the presence of microbial mats (PC4). PC4 (33°N 38.403 118°W 48.025) was taken through an orange microbial and sectioned into 1 cm section for the first 6 cm and then in 3 cm section to a depth of 15 cm. Core sections were stored at 20°C prior to being shipped to Penn State.

A.2.2. Extraction and separation Extraction and isolation methods followed the separation method of Mayr et al. (2008), as modified in chapter 2, with additional purification steps to allow for quantification and isotope analysis. ~30 g of Hydrate Ridge and ~10 g of Santa Monica Basin wet sediment were needed to quantify, isolate, and make an isotope measurement on coenzyme F430. Sediment samples were agitated by ultra-sonication probe for 20 minutes in neutral 18.2 Ω water (pH 7), and in an ice bath to keep the temperature at 4OC. Sediment was separated from the extract, by centrifuged at 5000 g for 15 minutes. Sediments were extracted twice more in 18.2 Ω water at pH 3 using 0.1% formic acid. The three extracts were combined and neutralized to pH 7.2 using NaOH, in order to precipitate proteins, which were separated and removed by centrifuging the solution at 9000 g for 10 minutes.

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Amberlite column (1 cm x 10 cm) which had been flushed with two dead volumes of 10 nM formic acid. The F430 fraction was isolated in 10 ml of 100% methanol. This fraction was dried under nitrogen and stored at -20OC before being further purified via HPLC.

High pressure liquid chromatography (HPLC) was used to purify F430 sufficiently to enable quantification and isotope analysis. The first HPLC separation employed two Waters spherisorb ODS2 columns 5 ㎛ (4.6 mm x 150 mm), linked together and supplemented by a Phenomenex C18 (3mm x 4mm) guard cartridge. Mobile phase A consisted of HPLC-grade water, mobile phase B of 0.1% formic acid and mobile phase C of acetonitrile (HPLC- grade). At a flow of 0.5ml/min, the following gradient was applied: 0 minutes 0% A, 70% B 30%C; 2 minutes 0% A, 70% B 30%C; 4 minutes 0% A 50% B 50% C; 20 minutes 50% A, 0% B, 50% C; 25 minutes 25% A, 0% B, 75% C; 28 minutes 25% A, 0% B, 75% C; 30 minutes 0% A, 70% B, 30% C. The F430 peak eluted at 25 minutes and was collected over a 1.5-minute window based on the UV/vis detector response. Fractions were dried under nitrogen and brought re-dissolved in methanol for additional purification.

To separate F430 from a co-eluting molecule that was contributing extra carbon in the Nano EA-IRMS analysis, a Thermo Hypercarb 5㎛ (100mm x 4.6mm) was used. Mobile phase A consisted of HPLC-grade water, mobile phase B of 0.1% hydrochloric acid and mobile phase C of acetonitrile (HPLC-grade). At a flow of 0.5ml/min the following gradient was applied: 0 minutes 0% A, 70% B 30%C; 2 minutes 0% A, 70% B 30%C; 4 minutes 0% A, 50% B, 50%C; 18 minutes 25% A, 50% B, 25% C; 20 minutes 50% A, 0% B, 50% C; 25 minutes 25% A, 0% B, 75% C; 28 minutes 25% A, 0% B, 75% C; 30 minutes 0% A, 70% B, 30% C. Quantification and identification were performed on the first run of sample through the Hypercarb column and subsequent runs were collected for nano-EA/IRMS analysis. F430 was identified by UV/vis detection of absorbance in the 430 nm wavelength and confirmed by the m/z 905 ion of the complete F430 structure. A previously published molar extinction coefficient of 21000 M-1 cm-1 was used to quantify F430 (Ellefson et al., 1982, Whitman and Wolfe, 1980). An Agilent 6300 ion trap with an ESI source was used for mass spectral analysis. Fractions for nano- EA/IRMS were collected at 8 minutes for 20 seconds. Samples were then dried under nitrogen and transferred to Costech tin boats using methanol. Samples were covered and left to dry before loading into autosampler for isotope analysis.

A.2.3. Isotope analysis Quantities of F430 isolated from environmental samples are typically too small for conventional EA-IRMS (elemental analyzer - isotope ratio mass spectrometry). Instead, we used a nano-scale EA/IRMS technique, developed by Polissar et al. (2009). In this method, the combusted sample is concentrated by cryogenic capture, transferred by a low flow of helium through a capillary gas chromatograph column (J&W scientific GS-

CarbonPLOT 30 m 0.32 mm 1.5 µm) to separate N2 and CO2 peaks before isotope analysis by the IRMS (Thermo- Finnigan Delta Plus). Isotope values for samples at natural abundance are reported in the delta notation (equation 2), after characterization of standards and accounting for analytical blanks (Polissar et al., 2009).

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� = � = (1)

� = ((� − � )/ �)1000 (2)

Samples are first corrected for background contribution, which are generally sourced from the tin boats (Costech), helium carrier gas, and oxygen combustion gas. These “blank” contributions were determined from multiple measurements of empty tin cups, and by isotope mass balance (Equation 3) where nblank is blank peak area, nobs is the observed sample peak area, δblank is blank δ values and δobs is the observed δ of the sample.

�� = �� + �� (3)

This can be rearranged to the useful form of a linear relationship:

� = � − �(� − �)/(1/�) (4)

With the true isotope value being the intercept and determined using equation 5:

� = (�� − ��)/(� − �) (5)

The intercept of the relationship is the blank-corrected delta value for the sample, which is calculated using the peak areas of the blank and sample peaks (nblank, nobs), and the observed delta values for the blank and sample analyses (δblank , δobs) (equation 5). Plotting δobs against 1/nobs illustrates the mixing relationship between blank and analayte for both carbon and nitrogen isotope data sets and that δx (the intercept) is the true value. Figures 3-2, 3-3, 3-4, 3-5, 3-6 and 3-7 illustrates that the intercept of a regression line between the samples and blanks is the true value of a sample.

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Analytical accuracy was assessed using a suite of isotopic standards, which are analyzed with the samples, and similarly corrected for blank contributions. Three standards, octaethylporphine (Frontier Scientific δ13C - 34.05‰, δ15N -12.23‰), L-methionine (Sigma Aldrich δ13C -30.45‰, δ15N 0.46‰) and Sucrose (NIST δ13C - 10.45‰) were run in conjunction with the samples to determine isotopic offset. The measured and corrected standard values were regressed against their known values, and the resulting linear equation was used to correct unknown samples for any offset:

� = � �� + �� (6)

Errors were propagated using the sum of squares method, which assumes uncertainties for each correction step are both independent and random (Polissar et al., 2009). For this calculation, uncertainties (σ) for the blank cups (equation 7), standards (equation 8) and offset (equations 9 and 10) were all used to determine total analytical uncertainty (equation 11)

1 � = (� − � )^2 (7) �

1 � = (� − � )^2 (8) �

� = � − � (9)

1 � = (� − � )^2 (10) �

(�∑) = (�) + (�) + (�) + (�) + (�) (11)

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Blank correction uncertainty is simplified and represented by the analytical reproducibility of the blank delta values. Uncertainty in blank analyses was determined from n = 6 analyses of empty cups for each analysis batch. Blank correction uncertainty was 0.6‰ for carbon, and 0.8‰ for nitrogen.

Reproducibility of carbon and nitrogen measurements was determined from the standard deviation of blank-corrected octaethylporphine, methionine and sucrose values. Uncertainty in the offset correction was determined from the sum of squares of the offset values determined from equation 9 for each standard.

Reproducibility in carbon measurement were determined from the standard deviation of blank corrected octaethylporphine, methionine and sucrose values. Uncertainty in the offset correction was determined from the sum of squares of the offset values determined from equation 8 for each standard. The total analytical uncertainty propagated using the sum of squares (equation 11) was 3.8‰ for carbon and 3.6‰ for nitrogen.

A.3. Results The concentration of coenzyme F430 in the Santa Monica core is slightly more abundant than in the previously extracted core and peaks at a depth of 3cm instead of 1cm as in the previous core. The carbon isotopic composition of F430 ranges from -70‰ to -44.5‰ indicating that it is more enriched than in the core extracted in chapter 2. Nitrogen isotope data is also obtained from this core and varies from -2.67‰ to 1.17‰.

A.4. Conclusions The increased abundance and more enriched values of suggest a link between production and isotope values. In chapter 2 the abundance of F430 in the Santa Monica sediment was lower and its isotope values were more depleted in 13C. Potentially at higher rates of production more carbon is assimilated into F430 from DIC.

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A.5. Figures and tables

F430 concentration ug/g wet sediment 0 500 1000 1500 2000 2500 0

4 CM

Figure A-1: F430 concentration in PC6. The concentration of F430 peaks at 3 Cm depth but remains high from 3 to 5 cm sediment depth, unlike previous core profiles which show a sharp decrease (see chapter 2).

Table A-1: F430 concentration data for PC6. Depth Wet sediment g F430 µg/g 0-2 13 184 2-4 17 2096 4-6 18 1526 6-8 15 3

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Table A-2: F430 isotope data for PC6. δ15N δ13C

PC6 2-4 -2.37 -47.75 PC6 2-4 II -1.17 -70.52 PC6 4-6 -2.67 -44.91 PC6 4-6 II -2.55 -44.56

A.6. References ELLEFSON, W. L., WHITMAN, W. B. & WOLFE, R. S. 1982. Nickel-containing factor F430: chromophore of the methylreductase of Methanobacterium. Proceedings of the National Academy of Sciences of the United States of America, 79, 3707-3710. HOUSE, C. H., ORPHAN, V. J., TURK, K. A., THOMAS, B., PERNTHALER, A., VRENTAS, J. M. & JOYE, S. B. 2009. Extensive carbon isotopic heterogeneity among methane seep microbiota. Environmental Microbiology, 11, 2207-2215. MAYR, S., LATKOCZY, C., KRÜGER, M., GÜNTHER, D., SHIMA, S., THAUER, R. K., WIDDEL, F. & JAUN, B. 2008. Structure of an F430 Variant from Archaea Associated with Anaerobic Oxidation of Methane. J Am Chem Soc, 130, 10758-10767. ORPHAN, V. J., HOUSE, C. H., HINRICHS, K.-U., MCKEEGAN, K. D. & DELONG, E. F. 2001. Methane- Consuming Archaea Revealed by Directly Coupled Isotopic and Phylogenetic Analysis. Science, 293, 484- 487. POLISSAR, P. J., FULTON, J. M., JUNIUM, C. K., TURICH, C. C. & FREEMAN, K. H. 2009. Measurement of C-13 and N-15 Isotopic Composition on Nanomolar Quantities of C and N. Analytical chemistry, 81, 755- 763. WHITMAN, W. B. & WOLFE, R. S. 1980. Presence of nickel in Factor F430 from Methanobacteriumbryantii. Biochemical and Biophysical Research Communications, 92, 1196-1201.

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Appendix B: Bacteriohopanepolyols through the Little Salt Springs water column

B.1. Introduction The lipid biomarkers 2-methyl hopanoids, found widely in Proterozoic sediments, are conventionally interpreted to represent the presence of ancient cyanobacteria (Summons and Walter, 1990, Summons et al., 1999).This interpretation is based on the high proportion of 2-methyl BHP in cultured cyanobacteria and the belief that a cyanobacterial origin can account for the ubiquity of 2-methyl hopanoid across a range of environments and geological ages (Summons et al., 1999, Talbot et al., 2008). Yet, this interpretation is challenged by genetic evidence that less than 10% of modern bacteria are capable of producing bacteria hopane polyols (BHPs) and all currently known marine cyanobacteria don’t produce 2-methyl BHPs (Pearson et al., 2007, Talbot et al., 2008). Potentially this discrepancy is due to a preservation bias with the desmethyl hopanoids recycled and not exported to the sediment. This can be tested in the water column of Little Salt Springs in Florida a 70 m deep sinkhole with a BHP producing biofilm in the top 10m. Out of the 3 BHP produced by the biofilm (bacteriohopanetetrol, 2-Methyl bacteriohopanetetrol and 2-Methyl Anhydro bacteriohopanetetrol) only 2-Methyl bacteriohopanetetrol is found in the water column.

B.2. Methods

B.2.1. Sampling The water column was sampled in the summer of 2012 and the winter of 2013. Biofilm samples were collected by divers and frozen for return to Penn State. The water column samples were collected via pumping water through 2um filters at 2,5,10,20 and 30 m water depth, with depths determined by using a sonde probe. 60 litters of water was passed through the filter before they were removed from the apparatus and frozen for return to Penn State.

B.2.2. Extraction Cell pellets were freeze-dried and a biomass weight obtained using a microbalance before being extracted using a modified Bligh-Dyer method as described by Talbot et al. (2007). Samples were submerged in a monophasic solution of 4:10:5 water:methanol:dichloromethane, disrupted with a sonicator bath for 1 h at 400C, shaken at 200 rpm for 1 h, and centrifuged at 9000 rpm for 15 min. The supernatant was removed and the extraction repeated twice. 10 ml of dichloromethane and 10 ml of water were added to the pooled supernatant to induce phase separation, and the organic phase was removed. The aqueous phase was extracted with 10 ml of dichloromethane two additional times, and the pooled total lipid extract (TLE) phases were dried under N2 and weighed.

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B.2.3. Acetylation 20% of the sample TLE was transferred to a 2 ml vial and then were spiked with an internal standard of pregnanediol for a concentration of 500 ng per 50 µl injection. The spiked samples were dried under N2 then acetylated using 50 µl of acetic anhydride and 50 µl of pyridine, heated at 60°C for 1 hour as outlined in Blau and Halket (1993). After acetylation samples were dried and re-suspended in 500 µl of methanol for HPLC analysis.

B.2.4. LCMS Analysis BHPs were analyzed using an Agilent 6310 Ion Trap LC/MS system after Talbot et al. (Talbot et al.,

Atmospheric pressure chemical ionization (APCI) source settings were: corona voltage 8000 nA; nebulizer pressure 60 psi; drying gas flow 5 l/min; drying T of 350°C, vaporizer T 490°C. The scanning range was m/z 150- 1300, and the run divided into 3 segments targeting m/z 285 (0-10 min), 1002 (10-17 min), and 655 (17-50 min). Automatic MSn settings for 2 precursor ions were as follows: absolute threshold 100,000, relative threshold 5%, and fragment amplitude 1.0V. Auto MSn settings for single precursor ions were: absolute threshold of 1,000, relative threshold 5%, and fragment amplitude 1.0V. Ions were excluded from auto MSn after 2 counts and released after 0.5 min. Detection was achieved at an isolation width of 3.0 m/z units and a fragment amplitude of 1.0 V.

B.2.5. Compound identification Compounds were identified by comparison with published MS spectra from Talbot et al. (2003), Talbot et al. (2007). The molecular ions used for identification are as follows: BHT (m/z 655), 2-Me BHT (m/z 669), and 2- Me Anhydro BHT (m/z 869). Methylation at the C-2 position was previously confirmed via oxidative cleavage and GCMS analysis (Albrecht, 2011).

B.2.6. Quantification To quantify the amounts of BHT and 2-Methyl AnhydroBHT in the culture, response curves for known amount of each were produced relative to 500 ng of pregananediol. Standards were purified from cultured biomass using the extraction method previously mentioned. Fractions were collected from the C18 column and then further purified with a second dimension of HPLC using a normal phase silica column (150mm x 30mm, Resteck) and an isocratic flow of 9:1 isopropyl alcohol : hexane. The collected fractions for BHT and 2-Methyl AnhydroBHT were then transferred to weighed 2 ml vials to obtain a mass of each standard (BHT 390 µg +- 10 µg, 2-Methyl AnhydroBHT 350 µg +- 10 µg).

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Response factors were produced by running 100 ng, 200 ng, 500 ng, 1000 ng and 2000 ng of standard of each BHP in addition to samples. Peak areas of BHT and 2-MeAnhydroBHT were normalized to the internal standard (500ng pregananediol) in order to account for instrument drift between injections. A linear regression line of the ratio of the known amounts (BHT ng / pregnanediol ng) against the peak area ratio (BHT response / pregnanediol). Unknown amount of BHT and 2-MeAnhydroBHT were determined using the slope and intercept from the linear regression line and to convert the response ration of the unknown amounts to the ng ratio which is converted to the known amount as the amount of pregenanediol added is known. A regression line is produced for each BHT compound as the two compounds as the response for 2-MeAnhydoBHT is an order of magnitude greater than BHT. Due to the use of APCI source which is open to the atmosphere there is daily variation in the ionization of the standards due to environmental factors such as temperate and humidity. As a result, the linear regression line is produced for every sequence of samples. Multiple standards of the same amount are also run to assess variance during a sequence. R2 is used to determine who well the regression line fits the data, with R2 produced for BHT regression lines varying between 0.91 and 0.99 indicating a good fit. Similar R2 values are obtained for 2- MeAnhydoBHT regression lines (0.93 to 0.99) and also indicate a good fit between the data and the regression line. Each unknown sample is run and corrected for three times to assess the reproducibility with standard deviation used to assess the variability between each run.

B.3. Results The biofilm found in the top 10m of the Little Salt Springs sinkhole was found to produce 3 BHP in 2012 (bacteriohopanetetrol, 2-Methyl Bacteriohopanetetrol and 2-Methyl Anhydrobacteriohopanetetrol) and only 2 in 2013 (2-Methyl Anhydrobacteriohopanetetrol and an unknown BHP structure). There is also a greater abundance in 2012 (120000 to 150000 ug/g) than there it in 2013 (2000 to 4000 ug/g) (figure B-1). In the water column filters were analyzed only 2-MeAnhydroBHT is found in concentrations of 3.2 to 0.3 ug/l (figure B-2).

B.4. Conclusions Not all of the BHP produced by the Little Salt Springs biofilm are found in the water column, with only 2- MeAnhydroBHT recovered from the filter samples. As a result, only 2-Methyl anhydrob BHT will be exported to the sediment at the bottom of the sinkhole. If this sediment is later extracted it would have a high ratio of 2-Methyl hopanoid ration. The larger 2-Methyl ratio in the past could be a result of preferential export of 2-Methyl BHPs like in the Little Salt Springs sinkhole, instead of a greater production by cyanobacteria.

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B.5. Figures

4 x 10 Biofilm and Cyanobacteria controls 18 BHT 2−MeBHT 16 2−MeAnhydroBHT Unknown

ug/g TLE 8

0 2012−Frozen 2012−Air dried 2013 Figure B-1: Biofilm BHP composition

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ug / l 0 1 2 3 4 0

15 depthm 20

25 2012 30 2013

Figure B-2: Concentration of 2-MeAnhydroBHT in the water column.

B.6. References BLAU, K. & HALKET, J. 1993. Handbook of derivatives for chromatography, Wiley New York. PEARSON, A., FLOOD PAGE, S. R., JORGENSON, T. L., FISCHER, W. W. & HIGGINS, M. B. 2007. Novel hopanoid cyclases from the environment. Environmental Microbiology, 9, 2175–2188. SUMMONS, R. E., JAHNKE, L., HOPE, J. & LOGAN, G. 1999. 2-Methylhopanoids as biomarkers for cyanobacterial oxygenic photosynthesis. Nature, 400. SUMMONS, R. E. & WALTER, M. R. 1990. Molecular fossils and microfossils of prokaryotes and protists from Proterozoic sediments. American Journal of Science, 290A, 212-244. TALBOT, H. M., ROHMER, M. & FARRIMOND, P. 2007. Rapid structural elucidation of composite bacterial hopanoids by atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry. Rapid Communications in Mass Spectrometry 21, 880-892. TALBOT, H. M., SUMMONS, R. E., JAHNKE L. & PAUL FARRIMOND, P. 2003. Characteristic fragmentation of bacteriohopanepolyols during atmospheric pressure chemical ionisation liquid chromatography/ion trap mass spectrometry. Rapid Communications in Mass Spectrometry, 17, 2788-2796. TALBOT, H. M., SUMMONS, R. E., JAHNKE, L. L., COCKELL, C. S., ROHMER, M. & FARRIMOND, P. 2008. Cyanobacterial bacteriohopanepolyol signatures from cultures and natural environmental settings. Organic Geochemistry, 39, 232-263.

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Appendix C: F430 Extraction and Purification for quantification and Isotope analysis

This method is based on initial work by Dr. Jamey Fulton and Nisha Patel and adapted from Mayr et al. (2008). Using this method F430 can be isolated for quantificationa and isotope analysis (carbon and nitrogen) from sediment and culture samples.

C.1. Extraction

C.1.1. Material o 250 ml polycarbonate bottles for sediment or 50ml teflon tubes for biomass

o Branson sonicator probe

o Centrifuge

o pH meter

C.1.2. Chemicals o Formic acid 98%

o Hydrochloric acid 1%

o Sodium hydroxide 10%

C.1.3. Extraction procedure 1. Weigh out sample into polycarbonate bottle or teflon tube

2. Fill bottle with DI water and adjust if needed to pH 7 with NaOH

3. Vortex into a slurry

4. Sonicate for 20 minutes, 30% duty cycle output control 6, using an ice bath to keep the sample from getting warm. Samples should be kept cold (fridge) before and after sonication.

5. Centrifuge at 12000g (teflon tubes) or 9000g (bottles) for 20 minutes at 4oC to remove sediment from extract.

6. Decant extract into a new bottle and purge the head space with N2 of Argon to remove oxygen.

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7. Re-suspend the sample in DI water and adjust to pH 3 with 98% formic acid (about 3 pipet drops). To successfully re-suspend add a small amount of DI water and vortex, once in a slurry fill the remainder of the bottle and add the formic acid. Be careful with carbonate containing samples as the formic acid can react with the carbonate. Just before sonicating check the pH, it is critical that the next two extractions are close to pH 3

8. Repeat sonication, centrifugation and decanting for a total of 3 extractions

C.1.4. Protein removal This is one of the first stages of purification and involved raising the pH to precipitate out unwanted proteins which would bind to the sephadex and decreasing its separation efficiency

1. Clean the pH probe with DI water before measuring the initial pH of the extract.

2. Adjust extracted solution to ~pH 7 with 10% NaOH. Initial a few drops of 50% NaOH can be added until around pH 5 to 6 be very care full as you don’t want to over shoot. This can be difficult especially with sediment samples where carbonate will buffer the solution. If the pH does rise above pH 7.5 (slightly above pH 7 should be ok) lower the pH to below 3 with formic acid. A brown precipitate should form at the bottom of the bottle.

3. Transfer samples to centrifugal bottles or tubes and centrifuge at 9000 rpm to separate the proteins from the extract.

4. Decant into clean bottles

5. Sample are now ready for sephadex chromatography. If storing overnight purge the headspace with N2 or Argon and refrigerate

C.2. Column chromatography Two stage for column chromatography are used to purify the F430 extract before samples can be run on the HPLC for characterization. Sephadex is initial used to remove unwanted material from the extract and then amberlite to further purify and isolate F430 in methanol.

C.2.1. Sephadex chromatography

Material o Supelco LC column with stopcock 64760-U 300 mm × 25 mm × 22 mm o QAE Sephadex A-25 (GE healthcare 10-0190-01)

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o 50mM Tris/HCl (7.88g Tris/HCl in 1 liter solution; adjust to pH 7.5 with NaOH) o 20 mM formic acid (0.755 ml 98% formic acid in 1 liter) o 2 M NaCl (117g in 1 liter solution) o 0.1M NaOH (20 ml 5 N NaOH in 1 liter) o Recommend making 4 liters Tris/HCl and formic acid § -31.52g Tris/HCl in 4 liters of solution adjust to 7.5 with 5 N NaOH § -3.02 ml 98% formic acid in 4 liters

Procedure 1. Plug the bottom of the columns with Pyrex wool and ash overnight

2. Solvent clean the stopcocks with Methanol, Dichloromethane and Hexane and then attach to column.

3. Added ~20ml of Tris/HCl to remove any air bubbles from the Pyrex wool plug.

4. Stir the sephadex with excess Tris/HCl and pour the slurry in to the column with stopcock open tapping the sides to remove any air bubbles

5. Fill the column with 10cm of sephadex. Dead volume 10cm x 2.5cm = 25cm3. The dead volume changes based on the solvent which passes through it. Dead volumes will be based on the Tris/HCl 10 cm volume

6. Flush the column with 4 dead volumes of 20 mM formic acid

7. Flush the column with 4 dead volumes of Tris/HCl. Check that the effluent is ~pH 7 with pH paper.

8. Add the F430 extract to the column. A yellow band should form at the top of the column (this is not always the case)

9. Flush with 4 dead volumes of Tris/HCl

10. Add 20mM formic acid to the column. The yellow band should start to move through the column.

11. Collect 2 dead volumes as waste (~90ml) Collect 2 dead volumes as sample. If the yellow band is not visible to monitor its progress collect 4 dead volumes. If this yellow band is visible make sure that you collect it.

12. This fraction can then be applied to the Amberlite XAD column.

Cleaning the sephadex

o Flush with 4 dead volumes of 20 mM formic acid

o Flush with 2 daed volumes of 2M NaCl

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o Flush with 2 dead volumes of 0.1M HaOH

o Flush with Tris/HCl until the effluent returns for pH7.5

After the column has been cleaned it can be used again starting from step 8. Sephadex can be stored overnight in the column as long as there is enough Tris/HCl to stop the top of the column drying out. If storing for longer store in 20% ethanol.

C.2.2. Amberlite chromatography

Materials

o 1 cm ID x 30 cm glass column with ~150 ml reservoir plug with Pyrex wool

o Amberlite XAD 1180 (Acros organics, CAS: 9003-69-4)

o 10 mM formic acid (0.377 ml 98% formic acid in liter solution)

o Methanol

o 60 ml Turbovap tube

Cleaning

Using a Buchner funnel rinse the Amberlite with DI water (3 times), methanol (3 times) and then DI water (3 times).

Procedure

1. Add 10 mM formic acid to the column with stopcock open to remove air bubbles from the Pyrex wool

2. Add 10 mM formic acid to the cleaned Amberlite in a beaker and pour it into the column

3. Fill the column with 10 cm with Amberlite (Dead volume 10cm x 1cm = 10 cm3)

4. Flush with 2 dead volumes of 10 mM formic acid. Restrict flow to 1 drop per second with the stopcock

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5. Add F430 fraction from the sephadex column to the column reservoir and pass it through the Amberlite at 1 drop per second.

6. Flush with 2 dead volumes of 10 mM formic acid. Flow 1 drop per second.

7. Elute F430 with methanol at 1 drop per second. The first 1.5ml can be collected as waste. You should see a yellow band move thought the column material, this needs to be collected.

8. Dry down under nitrogen in the turbovap and transfer to a labeled 4 ml vial.

9. Flush head space of 4 ml vial with N2 or Argon and store in -80℃

Methanol and Amberlite react forming gas bubbles within the column meaning the column has to be repacked before being used again. Amberlite is cleaned by rinsing with 2 dead volumes of methanol, Di water and then 10 mM formic acid. The column is repacked by tipping the Amberlite into the reservoir and then back into the column.

C.3. HPLC Purification and quantification

C.3.1. Instruments o LC system with DAD, MS with ESI source and Fraction collector

o Two linked 12 cm Waters C18 columns linked together

o Thermofisher Hypercarb column

C.3.2. Materials o Acetonitrile

o 0.1% formic acid

o 0.1% HCl

o LCMS grade water

o 2 ml Agilent vials

C.3.3. Waters C18 columns procedure 1. Transfer 50% of sample with methanol into an insert in a 2 ml agilent vial

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2. Dry the sample down and suspend the sample in 100㎕ of 75% 0.1% formic acid and 30% acetonitrile

3. Load method (table C-1) and write up sequence

4. Purge lines with starting composition for 10 minutes to purge the column with the starting composition

5. Inject the 100㎕ of sample using the auto sampler

6. Collect the F430 containing fraction which elutes between 22 and 25 minutes (figure C-1). This can be collected by disconnecting the lines or with the fraction collector if the retention times appear to be consistent.

7. Purge the head space with N2 or argon and store below -20℃

Even though F430 contains a number of carboxylic acid groups it does not need to be derivatized. F430 should still be stable at this point, the final dimension of chromatography removes a stabilizing molecule and should be instantly loaded into a tin boat for Nano EA IRMS

C.3.4. Hypercarb column The second dimension of HPLC removes a molecule which was contributing extra carbon in the Nano EA IRMS runs. Removing this molecule makes F430 unstable, with it completely degrading in about an hour. Once F430 is isolated it should be loaded up into a tin boat and put in the auto sampler

1. Transfer 100% of sample with methanol to an insert in a 2 ml Agilent vial

2. Bring sample up to 100 ㎕ with 75% 0.1% HCl and 25% acetonitrile

3. Load method (table C-2), write sequence and create a new data folder with the run date

4. Purge lines with starting composition for 10 minutes to purge the column with the starting composition

5. Inject the 50 ㎕ of sample using the auto sampler for quantification and identification. Quantification is based on the response in the 430 nm wavelength corrected to a known response factor. Identification is based on absorbance in the 430 nm wavelength and the 905 ion in the mass spectra.

6. Inject another 50㎕ of the fraction collected sample and collect for Nano-EA IRMS. Depending on the peak size from the identification and quantification run you may need to collect more from the waters column. A response of 20AU is the minimum needed for a Nano-EA run, though at this level problems can be encountered if the background on the Nano-EA are high

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7. Collect the F430 containing fraction which elutes between 7 and 8 minutes (figure C-2). This can be collected by disconnecting the lines or with the fraction collector if the retention times appear to be consistent. Remember to collect blank solvent samples over this same time period to characterize the amount of carbon the background contributes to the sample

8. Dry collected fraction down under nitrogen and transfer with methanol to a clean tin boat.

C.4. NANO-EA IRMS For a detailed description of the Nano EA (figure C-3) see Polissar et al. (2009). Briefly the samples is combusted in the EA, trapped in liquid nitrogen then fed into a low flow of helium. This process concentrates a sample peak which would be diluted by the normal helium flow from the EA.

C.4.1. Material o 6mm x 2.9mm Tin capsules Costech

o Liquid nitrogen

o Standards

o OEP δ13C -34.05, δ15N -12.23

o Sucrose δ13C -10.45

o L-methionine δ13C -30.45, δ15N 0.46

C.4.2. Procedure 1. The tin cups need to be cleaned be soaking them in methanol, DCM and Hexane for 20 minutes and decanting.

2. Transfer the hybercarb collected fractions using methanol to a cleaned tin boat

3. Measure out the (OEP, L-methionine and sucrose) standard with at least 3 different amounts.

4. Load auto sample carousel with samples, standards, blank tin cups and solvent blanks.

5. Vacuum out the auto sample and then purge with helium three times. Depending on when the EA was last used the system may have to be left for 12 hours for the backgrounds to come down. Leave two blank spaces in the auto sampler so two no cups can be run to assess the background levels.

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6. Check the setup of the Delta system. The settings on the conflow should be He 1.5 bar, CO2 2.5 bar and N2 1.54 bar. The EA should be set to micro and plumbed for Nano EA. Also before opening the isolation valve on the auto sample check that the EA is leak tight. Once the EA and conflow are set up switch over from source to waste on the Delta, load the timing power point and run the sequence.

7. Once the sequence is competed create separate .csv files for data, cups, each standard and the solvent back ground. Create a folder with the name and date of the run and place the .csv files and the nanoeacorrectionAAA.m file in the folder and run the script to correct the data.

C.4.3. Backgrounds issues

Most issues with background levels of CO2 or N2 come from leaks in the EA systems or the gas tanks particularly the oxygen tank. Leaks will mostly be around the furnace and auto sampler seals, though they can occur anywhere a connection has been made. Past testing shows that the oxygen tank contributes most of the carbon and nitrogen background. The He tanks will have a small amount of nitrogen with most coming from the EA tank which has the higher flow rate.

145

C.5. Figures and tables

Figure C-1: LCMS solvent profile using waters columns. The figure shows the eluting F430 compound. Identification is based on the 905 extracted ion chromatogram and absorbance at 430 nm. The peak from the waters column is fairly broad due to another molecule eluting at the same time. Also some samples have only shown a small peak or almost no response in the 430 nm but have a very well defined peak when run on the hypercarb column. This is due to other molecules interacting with the F430 and reducing its response. So a fraction should be collected between 20-25 min on the waters column every time as the DAD response at this point is unreliable due to other molecules in the extract.

146

Total ion Chromatogram 50-2200 m/z

0 Extracet ion chromatogram 905.3 m/z Signal intensity 0 UV-vis Chromatogram 430 nm

0 100% Solvent program Elution composition for F430

Acetonitrile

50% Water

0.1% Hydrochloric Acid 0% 5 6 7 8 9 10

Figure C-2: LCMS profile using Thermo Hypercarb column. Solvent profile showing elution of F430 and the 430 UV-vis and 905 extracted ion chromatograph used to identify F430

147

Oxidation Water trap furnace Costech EA VENT

Red to trap reverse to vent to IRMS

Low flow of helium

Reduction furnace Liquid nitrogen trap

GC Column

Reference gases

Delta XP

Conflo 3

Figure C-3: Nano EA IRMS system diagram.

Table C-1: Solvent profile for first dimension of HPLC chromatography. Flow rate 0.5ml/min

Time (minutes) Water % 0.1% Formic acid % Acetonitrile % 2 0 70 30 4 0 50 50 20 50 0 50 25 25 0 75 28 25 0 75 30 0 70 30

148

Table C-2: Solvent profile for second dimension of HPLC chromatography. Flow rate 0.5ml/min

Time (minutes) Water % 0.1% HCl % Acetonitrile % 2 0 70 30 4 0 50 50 18 25 25 50 20 50 0 50 25 25 0 75 28 25 0 75 30 0 70 30

C.6. References MAYR, S., LATKOCZY, C., KRÜGER, M., GÜNTHER, D., SHIMA, S., THAUER, R. K., WIDDEL, F. & JAUN, B. 2008. Structure of an F430 Variant from Archaea Associated with Anaerobic Oxidation of Methane. J Am Chem Soc, 130, 10758-10767. POLISSAR, P. J., FULTON, J. M., JUNIUM, C. K., TURICH, C. C. & FREEMAN, K. H. 2009. Measurement of C-13 and N-15 Isotopic Composition on Nanomolar Quantities of C and N. Analytical chemistry, 81, 755- 763.

149

Appendix D: Data tables

Table D-1: Hydrate Ridge sulfide and sulfate data

Sample Depth (cm) Sulfide (mM) Sulfate (mM) 5116 0-3 1.27 8.65 5117 3-6 7.65 0.56 5118 6-9 7.50 0.37 5119 9-12 4.66 2.00

Table D-2: Hydrate Ridge Aggregate counts, methane and pH data

Sample Depth (cm) Aggregates Methane (µm) pH Per ml 10^6 5048 0-3 1.125 166 7.95 5049 3-6 2.25 636 8 5050 6-9 0.75 1375 8.34 5051 9-12 0.375 1273 8.68 5052 12-15+ 948 8.5

Table D-3: Hydrate Ridge Coenzyme F430 data. Measurements were performed at Pennsylvania State University.

F430 SAMPLE depth (cm) g sediment elution time (min) response mAU µg µg/g 5086 0-3 31.17 8.44 8.02 346.19 11.11 5087 3-6 31.51 8.44 60.50 2611.69 82.89 5088 6-9 31.69 8.44 28.20 1217.29 38.42 5089 9-12 31.99 8.44 2.02 87.19 2.73

150

Table D-4: Hydrate Ridge carbon isotope data. Measurements were performed at Pennsylvania State University.

13 δ C (‰)

Carbonate Methane Total F430 Archaeol Hydroxyl organic carbon -archaeol 0-3 -37.7 -63.5 -26.4 -26.5 -26.7 -94 -88 3-6 -37.4 -62.4 -27.8 -27.7 -28.7 -24 -25 -94 -94 -64 -63

6-9 -42.5 -66.6 -28.3 -28.3 -28.7 -21 -22 -25 -21 -124 -125 9-12 -33.8 -68.7 -24.3 -24.3 -24.3 -21

12-15+ -16.1 -70.2 -23.3 -23.5 -23.5

STDEV 0.3 0.1 0.3 1 1

Table D-5: Santa Monica Basin sulfide, sulfate, ammonium and aggregate counts.

Depth cm HS- mM SO42- mM NH4+ µM Aggs per ml sed (x10^7) 0.5 10.78 19.29 103.33 1.5 8.57 10.69 101.67 3.12 2.5 24.87 3.39 42.33 5.18 3.5 34.95 0.86 16.00 1.58 4.5 33.13 0.64 33.67 1.90 5.5 10.78 0.19 38.67 1.06 7.5 36.66 0.23 52.00 0.32 10.5 21.02 0.21 65.67 0.62 13.5 11.64 0.12 62.00 0.68 16.5 32.55 0.10 74.00 0.46

151

Table D-6: Santa Monica Basin. Measurements were performed at Pennsylvania State University.

CM Amount Peak Response µg µg/g sediment used g Area mAU 0-2 10.0791 1843 377 16293 1616 2-3 12.3804 1278 261 11255 909 3-4 10.8703 46 10 432 40 4-5 10.8319 142 30 1274 118 5-6 9.9384 0 0 6-9 23.0916 37 7 319 14 9-12 29.5166 5 1 43 1 12-15 35.7932 0 0 0 0

Table D-7: Results of Hydrate Ridge Labeling experiments. Measurements were performed at Pennsylvania State University.

Sediment Response δ13C F430 Experiment Condition g mAU µg/sediment (‰) δ13C Archaeol (‰) 13C 1A methane 21.7668 10.96 21 -19 -48 -49 13C 1B methane 22.7652 0.5 0.9 13C bicarbonat 2A e 21.8967 1.61 3 119 117 123 13C bicarbonat 2B e - - 3A - - 4A trace trace 4B - - Unlabled 5 control 17.9161 31.34 75 -23 -88 -65 Killed 6 control - -

152

Table D-8: Santa Monica Basin DIC concentration results

DIC (mM) PCKD Time (days) Condition 2 33 41 75 100 114 1A 9.08 11.74 2.80 8.57 6.79 10.87 1B 9.19 7.10 2.94 9.60 1C 9.21 6.62 2.25 10.36 6.84 11.58 aver 9.16 8.49 2.67 9.51 6.82 11.23 sd 0.07 2.83 0.36 0.90 0.04 0.51 2A 9.17 10.28 17.50 22.31 21.05 2B 9.49 12.63 18.99 22.00 25.05 2C 5.08 12.58 18.64 22.25 24.10 aver 7.91 11.83 18.38 22.18 23.40 sd 2.46 1.34 0.78 0.17 2.09 3A 21.89 25.65 28.49 33.00 32.43 3B 21.23 26.29 34.26 34.09 31.21 3C 23.51 26.89 36.66 34.99 33.64 aver 22.21 26.28 33.13 34.03 32.42 sd 1.18 0.62 4.20 1.00 1.22 4A 9.03 8.48 9.32 9.84 12.02 4B 9.05 8.69 10.16 9.77 11.38 4C 9.35 9.09 9.75 7.86 11.01 aver 9.14 8.75 9.74 9.16 11.47 sd 0.18 0.31 0.42 1.12 0.51 5A 8.81 12.41 14.09 19.90 20.77 5B 9.34 11.97 10.42 23.37 20.85 5C 9.38 12.00 16.64 22.18 20.09 aver 9.18 12.13 13.71 21.82 20.57 sd 0.32 0.25 3.13 1.76 0.42 6A 10.97 14.09

153

Table D-9: Santa Monica Basin PCKD δ13C results

δ13C-DIC PCKD (‰) Time (days) Condition 2 33 41 75 100 114 1A 520.90 16688.12 13486.93 22006.43 30793.11 32842.15 1B 987.74 22119.85 17733.67 25226.72 1C 539.14 16831.15 12767.12 22297.50 30303.13 32266.95 aver 682.60 18546.37 14662.57 23176.88 30548.12 32554.55 sd 264.42 3095.55 2683.89 1781.16 346.47 406.72 2A 1965.63 1278.50 924.19 906.84 793.57 2B 1941.79 1246.89 923.51 905.55 779.06 2C 1961.53 1269.06 945.64 923.42 794.64 aver 1956.32 1264.82 931.11 911.93 789.09 sd 12.75 16.22 12.58 9.97 8.70 3A 29.59 201.49 196.89 197.26 195.54 3B 30.61 224.81 219.76 219.20 218.80 3C 22.07 237.03 232.51 231.53 231.57 aver 27.43 221.11 216.39 216.00 215.31 sd 4.66 18.06 18.05 17.36 18.27 4A 1989.70 1857.41 1717.74 1683.60 1633.01 4B 1872.10 1757.11 1632.58 1597.93 1551.35 4C 1959.02 1834.90 1723.69 1682.73 1648.19 aver 1940.27 1816.47 1691.34 1654.76 1610.85 sd 61.00 52.63 50.97 49.21 52.08 5A -32.44 -37.93 -39.42 -37.95 -40.25 5B -32.05 -37.47 -34.37 -37.79 -39.91 5C -32.31 -37.12 -32.07 -32.80 -39.72 aver -32.26 -37.51 -35.29 -36.18 -39.96 sd 0.20 0.41 3.76 2.93 0.27 6A 1318.33 1258.37

154

Table D-10: Santa Monica Basin hydrogen sulfide results

HS- mM PCKD Time (days) Condition 2 33 75 100 114 1A 0.01 3.21 2.14 4.70 4.59 1B 0.02 4.02 2.42 1C 0.02 3.52 2.55 5.70 5.42 aver 0.02 3.58 2.37 5.20 5.01 sd 0.00 0.41 0.21 0.70 0.58 2A 0.02 2.82 7.79 8.76 6.79 2B 0.02 4.07 8.16 9.18 7.69 2C 0.02 2.84 6.01 9.57 12.49 aver 0.02 3.24 7.32 9.17 8.99 sd 0.00 0.72 1.15 0.41 3.06 3A 0.02 0.02 0.02 0.02 -0.03 3B 0.02 0.02 0.03 0.02 -0.01 3C 0.02 0.02 0.03 0.03 0.01 aver 0.02 0.02 0.03 0.03 -0.01 sd 0.00 0.00 0.01 0.01 0.02 4A 0.02 0.02 0.02 0.02 0.01 4B 0.02 0.02 0.02 0.03 0.02 4C 0.02 0.02 0.03 0.03 0.03 aver 0.02 0.02 0.02 0.03 0.02 sd 0.00 0.00 0.00 0.01 0.01 5A 0.01 3.24 5.67 6.98 6.31 5B 0.02 2.84 7.58 7.55 3.86 5C 0.02 2.87 4.83 7.29 5.79 aver 0.02 2.98 6.03 7.28 5.32 sd 0.01 0.22 1.41 0.29 1.29 6A 0.09 -0.02

155

Table D-11: Santa Monica Basin experiment 3 and 4 methane results (µM)

Methane At 114 days Condition µM µM µM Average stdev 3A 3.48 3.55 4.03 3.69 0.30 3B 2.73 3.35 3.45 3.18 0.39 3C 3.37 3.61 3.60 3.53 0.14 4A 1.99 2.22 1.94 2.05 0.15 4B 2.17 2.03 2.71 2.31 0.36 4C 2.11 2.13 1.44 1.89 0.39

Table D-12: Santa Monica Basin experiment 3 and 4 methane results (ppm)

Methane At 114 days

Condition ppm ppm ppm Average stdev 3A 97.50 99.47 112.78 103.25 8.31 3B 76.57 93.72 96.71 89.00 10.86 3C 94.36 101.17 100.81 98.78 3.83 4A 55.71 62.25 54.29 57.42 4.25 4B 60.79 56.94 75.97 64.57 10.06 4C 58.96 59.59 40.36 52.97 10.92

Table D-13: Santa Monica Basin F430 amounts. Measurements were performed at Pennsylvania State University.

Experiment condition amount used g response mAU µg µg/g 1A 13C methane 10.8863 610 26334 2419 1B 13C methane 19.8798 874 37731 1898 1C 13C methane 10.8163 69 2979 275 2B 13C DIC 10.8836 1002 43257 3974 2C 13C DIC 11.0537 660 28493 2578 5A Unlabled control 11.3925 185.9 8025 704 5B Unlabled control 11.0497 109 4706 426 5C Unlabled control 11.5936 1090 47056 4059 6 Killed control 12.0744 2522 108876 9017

156

Table D-14: Santa Monica Basin F430 isotope results. Measurements were performed at Pennsylvania State University.

Nitrogen (‰) Carbon (‰) Experiment condition Run 1 Run 2 average Run 1 Run 2 average 1A 13C methane 187 97 142 255 67 161 395 146 271 137 58 98 1B 13C methane 249 249 30 14 22 619 619 16 24 20 1C 13C methane 58 57 57 105 47 76 354 354 95 95 2B 13C bicarbonate 313 312 312 -28 -27 -28 562 246 404 -29 -32 -30 2C 13C bicarbonate 280 154 217 -45 -32 -39 356 819 587 -46 -30 -38 5A Unlabeled control 29 29 -48 -48 -85 -85 5B Unlabeled control 74 74 -59 -59 -52 -52 5C Unlabeled control 355 149 252 -90 -90 440 771 605 -67 -67 6 killed control 3.9 1.0 2.5 -62 -94 -78 6 killed control 6.8 -0.5 3.2 -67 -91 -79

157

Table D-15: Santa Monica Basin Archaeol isotope results. Measurements were performed at Pennsylvania State University.

Offset corrected corrected for TMS Average Experiment condition δ 13C (‰) δ 13C (‰) (‰) 1A 13C methane -38 -37 -38 -37 -38 1B 13C methane -32 -51 -31 -52 -42 1C 13C methane -10 -9 -8 -7 -8 2A 13C bicarbonate -92 -98 -96 -102 -99 2B 13C bicarbonate -81 -83 -83 -87 -85 2C 13C bicarbonate -96 -97 -100 -101 -100 5A Unlabled control -105 -101 -110 -105 -108 5B Unlabled control -111 -110 -116 -115 -115 5C Unlabled control -94 -92 -98 -96 -97 6 Killed control -111 -111 -116 -116 -116

158

Table D-16: Santa Monica Basin newly synthesized F430 amounts.

5.4 20.4 24.8 72.5 66.2 assimilated

fDIC

0.2043 0.0538 0.2475 0.7254 0.6615 fCH4 /

2427 1414 1282

10078 12517 new delta

New F 0.1107 0.0371 0.1319 0.0264 0.0250 0.0104 0.0104 0.0099

0.0125 0.0113 0.0120 0.0108 0.0107 0.0104 0.0105 0.0102 0.0103

R 0.0127 0.0115 0.0122 0.0109 0.0108 0.0105 0.0106 0.0104 0.0104

29.0 38.4 66.5 55.3 78.4 78.3 21.0 82.1 ------129.1

average

2.3 4.1 1.4 3.4 3.0 0.2 0.7 3.7

new %

f 0.0227 0.0407 0.0145 0.0336 0.0298 0.0024 0.0067 0.0373

0.0045 0.0052 0.0042 0.0050 0.0048 0.0038 0.0039 0.0051 0.0037

R 0.0046 0.0053 0.0042 0.0050 0.0048 0.0038 0.0039 0.0051 0.0037

2.8 28.6 73.6

242.9 434.0 156.1 358.1 318.1 397.3 average

13C 13C 13C 13C 13C 13C killed killed control control control control

methane methane methane condition Unlabeled Unlabeled Unlabeled Unlabeled bicarbonate bicarbonate

6 1B 1C 2B 2C 5B 5C 1A 5A

Experiment

159

Table D-17: Santa Monica Basin newly synthesized Archaeol isotope values.

From fCH4 / Experiment condition average R Fobs Fnew F new fDIC % 1A 13C methane -38 0.0108 0.0107 0.0406 2767 0.06 6 1B 13C methane -42 0.0108 0.0107 0.0391 2620 0.06 6 1C 13C methane -8 0.0112 0.0110 0.0525 3929 0.09 9 13C 2A -99 0.0101 0.0100 0.0166 503 0.28 bicarbonate 28 13C 2B -85 0.0103 0.0102 0.0221 1007 0.53 bicarbonate 53 13C 2C -100 0.0101 0.0100 0.0161 453 0.26 bicarbonate 26 6 Killed control -116 0.0099 0.0098

160

Table D-18: BHT and 2-MeAnhydroBHT experiment results

Bacteriohopanetetrol 2-Methyl Anhydro Bacteriohopanetetrol

2-Me Sample biomass g ug/g ug/g ug/g average stdev ug/g ug/g ug/g average stdev ratio Control 0.0110 530 710 730 660 110 180 120 270 190 70 0.22 APR2015 1 Control 0.0127 670 570 530 590 70 520 290 420 410 110 0.41 APR2015 2 Control 0.0108 370 250 340 320 70 200 160 210 190 30 0.37 APR2015 3 Control 0.00918 860 1240 1020 1040 190 10 20 100 40 50 0.04 Dec2015 1 Control 0.00119 450 570 420 480 80 40 90 520 220 260 0.31 Dec2015 2 Control 0.00284 570 0 430 330 300 150 170 230 180 40 0.35 Dec2015 3 high salt 1 0.0032 520 470 0 490 40 110 40 0 80 50 0.14 high salt 2 0.00214 920 1060 1260 1080 170 290 280 230 270 30 0.20 High salt 3 0.00449 190 350 300 280 80 50 40 160 80 70 0.22 limited light 0.01758 1980 1290 1390 1550 380 210 210 270 230 30 0.13 shaken 1 limited light 0.01654 520 330 700 520 190 1120 480 640 750 330 0.59 shaken 2 limited light 0.02315 420 330 370 370 50 370 540 710 540 170 0.59 shaken 3 limited light 0.0025 5050 3270 4690 4340 940 1570 1060 1240 1290 260 0.23 not shaken 1 limited light 0.00141 8160 5830 7000 6990 1160 1020 1000 1130 1050 70 0.13 not shaken 2 limited light 0.00422 9300 5030 6810 7050 2150 1440 1440 1330 1400 60 0.17 not shaken 3 Not shaken 1 0.00108 0 0 0 0 0 0 0 0 0 0 - Not shaken 2 0.00136 0 0 0 0 0 0 0 0 0 0 - Not shaken 3 0.00174 0 0 0 0 0 0 0 0 0 0 - Sulfide 1 0.0019 1180 940 1310 1150 190 100 60 90 80 20 0.07 Sulfide 2 0.00595 870 670 850 800 110 150 150 260 180 60 0.18 Sulfide 3 0.00267 0 190 430 210 210 0 0 170 60 100 0.22 Sulfide and 0.00672 440 310 260 340 90 0 0 0 0 0 0.00 DCMU 1 Sulfide and 0.00474 1410 1260 1620 1430 180 0 130 130 90 70 0.06 DCMU 2 Sulfide and 0.00637 440 310 550 430 120 0 90 100 60 50 0.12 DCMU 3 Thiosulfate 1 0.00561 0 0 10 0 10 0 0 140 50 80 1.00 Thiosulfate 2 0.0043 0 0 0 0 0 0 0 0 0 0 Thiosulfate 3 0.00637 0 0 0 0 0 170 150 420 250 150 1.00

161

Table D-19: Nitrogen experiment results

2-Methyl Anhydro Bacteriohopanetetrol Bacteriohopanetetrol 2-Me Sample biomass g µg/g µg/g µg/g average stdev µg/g µg/g µg/g average stdev ratio Nitrogen T1 A 0.0073 0 130 150 90 20 80 50 60 60 10 0.40 Nitrogen T1 B 0.0082 240 300 270 270 30 170 90 130 130 40 0.33 Nitrogen T1 C 0.005 200 140 70 140 70 60 70 100 80 20 0.36 Nitrogen T2 A 0.0095 0 0 0 0 0 60 0 30 30 30 1.00 Nitrogen T2 B 0.0134 520 430 440 460 50 40 0 40 30 20 0.06 Nitrogen T2 C 0.0069 860 500 1230 860 360 250 190 270 240 40 0.22 Nitrogen T3 A 0.0124 480 590 480 510 60 60 20 40 40 20 0.07 Nitrogen T3 B 0.0149 380 450 630 490 130 20 40 90 50 40 0.09 Nitrogen T3 C 0.0108 230 280 350 290 60 150 220 230 200 40 0.41 Nitrogen T4 A 0.0297 630 480 490 530 80 10 20 40 20 20 0.04 Nitrogen T4 B 0.0142 700 810 640 720 90 50 60 80 60 20 0.08 Nitrogen T4 C 0.0187 580 600 700 630 70 70 60 60 60 0 0.09

Table D-20: Control experiment time series results

2-Methyl Anhydro Bacteriohopanetetrol Bacteriohopanetetrol 2-Me Sample biomass g µg/g µg/g µg/g average stdev ug/g ug/g ug/g average stdev ratio Control T0 0.0093 870 540 560 660 180 80 90 110 90 20 0.12

Control T1 A 0.0116 300 190 180 220 70 90 40 60 70 20 0.24 Control T1 B 0.0056 470 530 460 490 40 300 440 320 350 70 0.42 Control T1 C 0.0042 0 0 0 0 0 140 160 250 180 60 1.00 Control T2 A 0.0082 240 260 320 270 40 40 30 70 50 20 0.16 Control T2 B 0.0138 690 340 430 490 180 240 100 130 160 70 0.25 Control T2 C 0.0099 300 560 680 520 190 230 260 230 240 20 0.32

Control T3 A 0.0122 310 290 330 310 20 10 30 80 40 30 0.11

Control T3 B 0.0152 310 210 250 250 50 50 60 110 70 30 0.22 Control T3 C 0.0108 480 400 350 410 70 60 40 30 50 20 0.11 Control T4 A 0.0194 200 360 330 290 90 60 100 70 80 20 0.22 Control T4 B 0.0125 300 310 290 300 10 100 80 80 80 10 0.21 Control T4 C 0.006 320 400 430 390 50 140 170 320 210 90 0.35

162

Table D-21: 2-Methyl Anhydro bacterialhopanetetrol water column concentration. Filter samples were collected in 2012 and 2013.

µg/l Sample Depth (m) µg/l µg/l µg/l stdev average LS12-10 2 1.0 1.1 1.0 1.0 0.05 LS12-11 5 1.5 1.4 1.4 1.4 0.03 LS12-12 10 2.0 1.9 2.0 1.9 0.07 LS12-13 20 0.2 0.2 0.1 0.2 0.04 LS12-14 30 0.2 0.3 0.3 0.3 0.05 LS13-35 2 2.9 2.8 2.6 2.8 0.16 LS13-36 5 3.5 3.0 2.5 3.0 0.50 LS13-37 10 3.1 1.4 1.1 1.9 1.07 LS13-38 20 1.4 1.0 0.8 1.1 0.27 LS13-39 30 2.3 1.0 0.9 1.4 0.79

Table D-22: BHP concentration in biofilm samples. Biofilm samples were collected in 2012 and 2013

Sample Compound µg/g biomass µg/g biomass µg/g biomass µg/g average stdev

LS12-30 BHT 730 730 730 730 3 LS12-30 2-MeBHT 820 630 640 700 110 LS12-30 2-MeAnhydro 720 930 960 870 130 LS13-25 2-MeAnhydro 700 660 620 660 40 LS13-26 Unknown 2020 2490 2030 2180 270

163

Table D-23: Little Salt Springs water column data. Water samples were taken for sulfide, nitrite nitrate and ammonium measurements were made in the field.

NO2- (mg NO3- (mg NH4+ (mg depth (m) µM H2S N/L) N/L) N/L) LS12-1 60.0 42.4 0.0080 0.0000 0.0033 LS12-3 0.1 0.0 0.0010 0.0100 0.0100 LS12-4 53.4 65.5 0.0000 0.0067 0.4033 LS12-5 40.3 53.1 0.0000 0.0100 0.4033 LS12-6 30.0 58.0 0.0013 0.0100 0.3700 LS12-7 20.0 55.7 0.0000 0.0100 0.3567 LS12-8 10.0 57.5 0.0000 0.0100 0.3933 LS12-8 10.0 60.8 LS12-8 10.0 47.3 LS12-9 4.7 28.4 0.0000 0.0033 underrange LS12-9 4.7 25.1 LS12-15 54.9 39.4 LS12-16 60.1 42.1 LS12-17 74.7 23.3

Table D-24: Little Salt Springs water column data. Temperature, pH and dissolved oxygen (DO) were measured in the water column in June 2012

Depth Temp pH DO Conc m C mg/L 0.639 28.11 7.45 1.89 3.121 27.86 7.43 1.85 5.158 27.52 7.37 1.84 5.147 27.31 7.33 1.48 5.151 27.22 7.31 1.18 5.15 27.18 7.3 1.01 5.2 27.16 7.29 0.9 6.254 27.14 7.27 0.81 9.217 27.08 7.26 0.75 10.208 27.04 7.25 0.68 10.241 27.03 7.24 0.65 10.25 27.02 7.24 0.63 10.265 27.02 7.23 0.62

164

10.265 27.02 7.23 0.6 10.266 27.02 7.23 0.6 10.268 27.02 7.23 0.6 10.27 27.02 7.23 0.61 10.291 27.02 7.23 0.61 12.824 27.01 7.23 0.61 15.703 27.01 7.23 0.61 18.526 27.01 7.23 0.6 19.87 27.01 7.23 0.59 19.865 27.01 7.23 0.59 19.862 27.01 7.23 0.59 19.865 27.01 7.23 0.59 19.868 27.01 7.23 0.59 19.869 27.01 7.23 0.59 19.87 27.01 7.24 0.6 19.87 27.01 7.24 0.6 19.87 27.01 7.24 0.6 19.873 27.01 7.24 0.6 20.039 27.01 7.24 0.6 21.79 27.01 7.24 0.6 24.418 27.01 7.24 0.59 26.164 27.01 7.24 0.58 27.326 27.01 7.24 0.58 29.836 27.01 7.24 0.57 29.847 27.01 7.25 0.57 29.84 27.01 7.25 0.57 29.836 27.01 7.25 0.56 29.833 27.01 7.25 0.57 29.832 27.01 7.25 0.57 29.825 27.01 7.25 0.57 29.812 27.01 7.26 0.57 29.797 27.01 7.26 0.56 29.784 27.01 7.26 0.56 29.773 27.01 7.26 0.56 29.761 27.01 7.26 0.56 29.747 27.01 7.26 0.56 29.742 27.01 7.27 0.57 29.75 27.01 7.27 0.57 29.764 27.01 7.27 0.56 29.838 27.01 7.27 0.56

165

31.701 27.01 7.27 0.56 31.956 27.01 7.27 0.55 31.982 27.01 7.27 0.55 32.197 27.01 7.27 0.54 32.432 27.01 7.27 0.53 33.746 27.01 7.27 0.53 36.438 27.01 7.27 0.53 38.859 27.01 7.28 0.53 39.645 27.01 7.28 0.53 39.658 27.01 7.28 0.52 39.661 27.01 7.28 0.52 39.656 27.01 7.28 0.52 39.657 27.01 7.28 0.52 39.663 27.01 7.29 0.52 39.667 27.01 7.29 0.52 39.669 27.01 7.29 0.52 39.671 27.01 7.29 0.52 39.671 27.01 7.29 0.52 39.671 27.01 7.29 0.52 39.671 27.01 7.29 0.52 39.67 27.01 7.29 0.52 39.668 27.01 7.3 0.52 39.667 27.01 7.3 0.51 39.664 27.01 7.3 0.52 39.654 27.01 7.3 0.52 39.645 27.01 7.3 0.53 39.645 27.01 7.3 0.53 39.663 27.01 7.3 0.53 40.508 27.01 7.3 0.53 42.509 27.01 7.3 0.52 45.544 27.01 7.3 0.52 48.523 27.01 7.3 0.5 49.516 27.01 7.3 0.5 49.52 27.01 7.31 0.5 49.513 27.01 7.31 0.51 49.51 27.01 7.31 0.51 49.511 27.01 7.31 0.51 49.51 27.01 7.31 0.51 49.509 27.01 7.31 0.51 49.509 27.01 7.31 0.51

166

49.503 27.01 7.32 0.5 49.494 27.01 7.32 0.5 49.49 27.01 7.32 0.5 49.495 27.01 7.32 0.5 49.516 27.01 7.32 0.51 49.538 27.01 7.32 0.5 49.596 27.01 7.32 0.51 51.769 27.01 7.32 0.5 54.057 27.01 7.32 0.5 56.499 27.02 7.32 0.5 58.602 27.02 7.32 0.5 60.165 27.02 7.32 0.49 62.277 27.02 7.32 0.49 62.915 27.02 7.32 0.49 62.939 27.03 7.29 0.49 62.896 27.04 7.25 0.49 62.608 27.03 7.27 0.5 62.865 27.03 7.29 0.49 61.809 27.03 7.3 0.5

167

Curriculum Vitae Laurence Robert Bird MSci (Hons), ARSM

Education

Year Degree Honors University Location DEC 2016 Ph.D Pennsylvania State University USA 2010 MSci 1st Class Imperial College London UK

Professional Experience

Year Title Company Experience 2015 Researcher Penn State Analysis of Hominid samples by Nano-EA IRMS as 2015 Speaker ExxonMobil Invited to present research on Coenzyme F430 2014 Researcher Penn State and Caltech incubate methanotrophic sediment with 13C substrate to explore carbon assimilation 2013 Researcher Penn State Contributed to successfully funded American Chemical Society petroleum research fund proposal 2012 Researcher Penn State Research into Bacterial Hopane Polyol production by cyanobacteria. 2011 Researcher Penn State and MIT Collaborative project making isotope measurements of Tintinnid microfossils 2011 Researcher Penn State Research into the use of coenzyme F430 by methanotrophs. 2009 Researcher Massachusetts Institute Research into the organic content of Ooids. of Technology

Field Experience

Year Duration Formation & Location Activity 2008 5 Weeks Assynt complexity zone, Independent mapping project Scotland 2011 5 Weeks Yellowstone & Green River, Learnt how to collect microbial samples in Western US the field for community analysis 2012 10 Days Sudbury impact creator, Canada Visit to sites relevant to early life on Earth 2012 1 Week Little salt springs sinkhole, Collection of microbial mat and water Florida column samples 2015 1 Week C-Image Northern Gulf of Sediment core collection and extrusion at Mexico research cruise sea

Awards Shell Geosciences Energy Research Facilities Award - 2011,2012 & 2014 John Meacham Hunt graduate student award in petroleum geochemistry - 2012-2013 ConocoPhillips Graduate Student Fellowship - 2013-2014 Charles E. Knopf, Sr. Memorial Scholarship - 2011 CECG summer fellowship - 2012 Donald B. and Mary E. Tait Scholarship in Microbial Biogeochemistry - 2014

Memberships Associate of the Royal School of Mines, AGU, AAPG, ACS

Publications Summons, R.E., Bird, L.R., Gillespie, A.L., Pruss, S.B., Roberts, M. & Sessions, A.L. (2013) Lipid biomarkers in ooids from different locations and ages: evidence for a common bacterial ﬂora. Geobiology, 11. 5.