Aspects of Sulfur Metabolism of Methane-Producing Archaeon

Home , Acidianus infernus, Desulfurella acetivorans, Desulfurococcaceae, Methanocaldococcus, Methanofuran, Methanogen

ASPECTS OF SULFUR METABOLISM OF METHANE-PRODUCING ARCHAEON

METHANOCOCCUS MARIPALUDIS

FENG LONG

(Under the Direction of William B. Whitman)

ABSTRACT

Sulfur is vital for the growth of all known organisms and is present in a wide variety of metabolites with different physiological functions. Consistent with their obligate anaerobic habitats, most methanogenic Archaea only assimilate sulfide and elemental sulfur as sulfur sources, whereas sulfate and other oxidized sulfur compounds are rarely utilized. The methanogenic lifestyle may have evolved from ~ 3.5 billion years ago, and contemporary methanogens may have preserved some of the metabolic relics which were common in the early

Earth anaerobic lifestyles. Recent studies have revealed multiple novel traits of sulfur assimilation in methanogens. However, many aspects of the sulfur assimilation processes and their regulations remain to be investigated. Thereby, the study of the physiology and biochemistry of the sulfur metabolism in methanogens may provide new insights into the biology of ancient microbial life.

Methanococcus maripaludis is unable to assimilate sulfate as a sulfur sole and does not produce sulfate when grew with sulfide as the sole sulfur source. Nevertheless, Methanococcus maripaludis possesses homologs of proteins involved in the sulfate assimilatory reduction pathway. None of these proteins was functional in the Escherichia coli mutant strains deficient in sulfate assimilation metabolism. These results indicated that the assimilatory sulfate reduction pathway is most unlikely to be present in Methanococcus maripaludis.

When grown with elemental sulfur as the sole sulfur source, Methanococcus maripaludis produced sulfide at about 6 mmol per g cell dry weight per hour. Moreover, adenylyl-sulfate reductase (MMP1681), an enzyme that contains an iron-sulfur cluster, was found to be required for elemental sulfur assimilation. Furthermore, proteomics data indicated that the expression of this protein increased three-fold during growth with elemental sulfur in comparison to growth with sulfide. Together with bioinformatics analysis, a different physiological role of MMP1681 in elemental sulfur assimilation, in addition to its in vitro catalytic function as an adenylyl-sulfate reductase was demonstrated. Although the explicit mechanism by which MMP1681 participating in the elemental sulfur incorporation process remains to be elucidated, this evidence advances our understanding of how methanogen possessing MMP1681 assimilate elemental sulfur into key sulfur intermediates.

INDEX WORDS: Archaea, methanogens, Methanococcus maripaludis, sulfur metabolism,

elemental sulfur, sulfide, sulfate, assimilatory sulfate reduction, iron-sulfur

clusters.

ASPECTS OF SULFUR METABOLISM IN METHANE-PRODUCING ARCHAEON

METHANOCOCCUS MARIPALUDIS

FENG LONG

BS, Nanjing Forestry University, China, 2009

A Dissertation Submitted to the Graduate Faculty of The University of Georgia in Partial

Fulfillment of the Requirements for the Degree

DOCTOR OF PHILOSOPHY

ATHENS, GEORGIA

2017

Feng Long

ASPECTS OF SULFUR METABOLISM IN METHANE-PRODUCING ARCHAEON

METHANOCOCCUS MARIPALUDIS

FENG LONG

Major Professor: William B. Whitman Committee: Diana Downs Jorge C. Escalante-Semerena Ellen L. Neidle

Electronic Version Approved:

Suzanne Barbour Dean of the Graduate School The University of Georgia December 2017

DEDICATION

To my grandparents, Long Yongqiu and Liu Donge, my parents, Long Jianming and

Ouyang Mei, and my husband, Wu Yinghui, for their endless love, patience and support every step of the way.

I am grateful for every sacrifice you have made to provide me with the best life possible. I love you more than words can describe.

ACKNOWLEDGEMENTS

First, I would like to thank my advisor Dr. William B. Whitman, who brought me into the

Microbiology department and coached me on how to be a better scientist. He introduced me to the field of Archaea research and taught me the principles of research. His talent, passion, self- disciple and dedication to his work always inspired me when performing my research. Without his invaluable guidance, support and encouragement both in research and life, I would not have been able to make it this far. I would like to thank my doctoral committee; Dr. Diana Downs, Dr. Jorge

C. Escalante-Semerena, and Dr. Ellen L. Neidle for their constructive suggestions all these years.

My special thanks go to Dr. Yuchen Liu, she was always there to share her valuable research experiences, informative discussions, and great ideas! Many thanks to Dr. Hannah Bullock for her company as a great lab colleague and best friend at UGA. I would like to thank Dr. Zhe Lyu for teaching me many of the techniques used in the laboratory and bioinformatic tools. I would also like to thank Dr. Michael Adams for helping me with data analysis and providing a great deal of insights in sulfur metabolism through my research. Furthermore, I thank many of the past and present members of Whitman lab including Felipe Sarmiento, Joseph S. Wirth, Liangliang Wang,

Warren Crabb, Nana Shao, Tao Wang, Taiwo Akinyemi, Hao Shi, Qiuyuan Huang, Yixuan Zhu,

Suet Yee Chong, Peiying Chang, Ghazal Motakef, Dahyun Ji, Hirel B. Patel, Lola Osiefa,

Courtney Ellison and Courtney Grant Jr. for their help, friendship, and encouragement. I thank Dr.

Wendy A. Dustman, Dr. Anna C. Glasgow Karls and Dr. Jennifer Walker, for their support in my teaching certificate application. I also thank my classmates Ajay Arya, Nicole Laniohan,

Christopher Abin, Julie Stoudenmire, Christopher Cotter and the rest of the Microbiology

Department.

Lastly, I would like to thank my family and all my friends for being a constant source of encouragement, support, and friendship all the way along.

Thank you all for being the best memories in my life!

TABLE OF CONTENTS

Page

ACKNOWLEDGEMENTS ...... v

CHAPTER

1 INTRODUCTION AND LITERATURE REVIEW ...... 1

2 A FLEXIBLE SYSTEM FOR CULTIVATION OF METHANOCOCCUS AND

OTHER FORMATE-UTILIZING METHANOGENS ...... 50

3 AN ADENYLYL-SULFATE REDUCTASE IN METHANOCOCCUS

MARIPALUDIS, CONTAINS AN IRON-SULFUR CLUSTER, AND IS REQUIRED

FOR ELEMENTAL SULFUR ASSIMILATION ...... 71

4 EXPLORING THE USE OF GENOME COMPARISONS OF WILD-TYPE AND

RESISTANT MUTANTS OF METHANOCOCCUS MARIPALUDIS TO IDENTIFY

POTENTIAL TARGETS OF INHIBITORY COMPOUNDS ...... 129

5 CONCLUSIONS...... 156

APPENDICES

A CHAPTER 2 SUPPLEMENTARY INFORMATION ...... 159

B CHAPTER 3 SUPPLEMENTARY INFORMATION ...... 179

C CHAPTER 4 SUPPLEMENTARY INFORMATION ...... 214

vii

CHAPTER 1

INTRODUCTION AND LITERATURE REVIEW

Introduction

Methanoarchaea, or methanogens are strictly anaerobic microorganisms that form methane as a major product of their energy metabolism (1). They serve as key agents catalyzing transformations of organic carbon on earth and are responsible for producing most of the methane found in the Earth’s atmosphere (1). Methanogens originate from the early, anoxic earth and share several unique features (1, 2): (i) they are all Archaea; (ii) they produce methane as the end-product in their anaerobic respiration-methanogenesis, which provides all or most of the energy for supporting their growth; (iii) they utilize a restricted number of substrates for methanogenesis: mainly CO2 + H2 and/or formate (hydrogenotrophic methanogens), methyl-containing C-1 compounds (methylotrophic methanogens) and acetate (aceticlastic methanogens). They do not use sugars, amino acids, or most other common organic substrates; (iv) they can be found in a diverse range of anaerobic environments, including marine sediments, flooded soils, gastrointestinal tracts, anaerobic digesters, landfills, geothermal systems, and heartwood of trees.

Methanogenesis may represent one of the most ancient metabolisms and likely contributed a significant role in the evolution of the Earth’s atmosphere. Although it is unknown when methanogens first appeared on Earth, the oldest evidence of their emergence was 3.5 billion years ago (Ga), 1.1 Ga before oxygen appeared (3). Several lines of evidence have suggested that methanogens retain unique metabolic features that may have been common in the anoxic early

Earth. Frist, methanogenic archaea may have a unique origin. The methanogenesis pathway of 1

producing methane requires at least 25 core genes and more than 20 biochemically characterized proteins involved in the biosynthesis of coenzymes. Genes encoding different subunits of an enzyme are usually clustered together in the genome, but these clusters and genes are scattered around the methanogen genome (4). Phylogenetic analysis has found that these core methanogenic enzymes likely originated (as a whole) from the last ancestor of all methanogens and do not seem to have been horizontally transmitted to other organisms or between the methanogenic classes (5).

Nonetheless, part of the pathways involved in methanogenesis, as well as single genes, may have been transferred across diverse phylogenic lineages (6). Second, several methanogenic and biosynthetic enzymes are O2-sensitive, such as methyl-coenzyme M reductase (7) and pyruvate oxidoreductase (8). It is likely that these enzymes resemble forms that were common on the anoxic

Earth. Lastly, methanogens retain a significant amount of iron-sulfur proteins, as indicated by bioinformatics analysis (9) and biochemical measurements (10). Considering that the anoxic Earth provided an environment rich in iron and sulfide, these conditions may have allowed for spontaneous assembly of iron-sulfur clusters. Presumably, these methanogenic iron-sulfur proteins may represent some of the earliest catalysts. For these reasons, exploring the physiology and biochemistry of methanogens may provide exclusive insights to the biology of ancient microbial life.

1. Evolution of the Sulfur cycle

Sulfur compounds, with versatile valence states ranging from +6 to -2, serve as electron donors and/or elector acceptors in microbial metabolism (11). In addition to the metabolic diversity of sulfur, the most extensively studied microbial sulfur cycles are highlighted into four groups: (i)

The microbial sulfur cycle is known to start with the sulfate reduction pathway, as sulfate is the most prevalent form of sulfur in the environment. Sulfate reduction may be either assimilatory, 2

where the produced sulfide is further incorporated into carbon skeletons of amino acids to form cysteine or homocysteine, or dissimilatory, where sulfate is used as a terminal electron acceptor for anaerobic respiration. Dissimilatory sulfate reduction is carried out by anaerobic prokaryotes, typically Deltaproteobacteria in the domain Bacteria or the hyperthermophilic Archaeoglobus from the domain Archaea. Mostly, sulfate reducers utilize organic compounds as the electron donor, although some species also use H2 (12). As the majority of sulfate reducers cohabitate with other organisms that convert complex organic compounds into small molecules, they normally rely on a complex anaerobic microbial food chain to collect substrates for reduction (Figure 3-1). (ii) In the presence of light, free sulfide can be oxidized to elemental sulfur, sulfate, or thiosulfate by anoxygenic phototrophic bacteria (13) (Table 1-1). (iii) While in the absence of light, non- phototrophic bacteria may oxidize sulfide into sulfate using O2 or nitrate as electron acceptors (13).

(iv) The elemental sulfur disproportion activities have been first studied in homogenized tidal flat sediments (14). In this study, the sediment was amended with elemental sulfur and time course experiments monitored the concentration of metabolic end products, such as elemental sulfur, sulfides, sulfate (14). The production of both sulfate and sulfide was consistent with ongoing sulfur disproportionation (14). Additionally, an increase in organic carbon was observed, suggesting the presence of autotrophic microbes taking up inorganic carbon while they disproportionated elemental sulfur (14). Using this similar method, more bacteria conducting active elemental sulfur disproportionation were enriched from marine and freshwater environments (15). All the isolates and the 16S rRNA gene -tracked members of the enrichment cultures belonged to the delta class of the Proteobacteria phylum (15, 16). Many of the Deltaproteobacteria are also capable of using sulfate, thiosulfate, or sulfite as electron acceptors in a respiratory type of metabolism (Table 1-1).

Surprisingly, Pantoea agglomerans, a facultative anaerobic bacterium from the 3

Gammaproteobacteria was reported to be equally capable of disproportionating elemental sulfur

(17). Regardless, the enzymes involved in sulfur disproportionation have not been elucidated.

Thamdrup et al. (18) have proposed the sulfur disproportionation leads to a theoretical sulfide-to- sulfate ratio of 2:1 in the presence of reactive iron oxides (Table 1-1, equation iv). This stoichiometry starts from an initial 3:1 sulfide-to-sulfate production ratio (Table 1-1, equation i), followed by the reoxidation of some sulfide back to elemental sulfur during the reduction of iron oxides (Eq. 2) and precipitation of sulfide as FeS (Table 1-1, equation iii) (18). While in environments that contain large amounts of oxidants such as Mn oxides, sulfide may be completely oxidized into sulfur or sulfate by Mn oxides (Table 1-1, equation v) (19, 20). This proposed scenario is supported by the report that sulfur disproportionating bacteria in sediment was in the range of 105-106 cells cm-3 in coastal and intertidal sediments (18). In contrast, the abundance of sulfur disproportionating bacteria was less than 102 cells cm-3 in off-shore sediments in the

Skagerrak, a site abundant in Mn oxides (21).

Recent studies have revealed that some of these sulfur metabolisms are quite ancient and can be used to deduce the evolution history of early earth (11, 22-25). Others play pivotal roles in modern metabolism (11, 22-25). On the very early earth, sulfide would be found in volcanism and hydrothermal discharges (26, 27). With access to light in places like terrestrial hot spring settings, such as Yellowstone Park, Iceland and New Zealand, sulfide and its bacterial photooxidation products would also be available (28, 29). Not limited to the surface and hydrosphere, sulfate and elemental sulfur would have been formed via numerous pathways in the atmosphere, thus providing substrates for reductive and disproportionative metabolisms (28, 30, 31). The molecular and geochemical evidence of the antiquity of these sulfur metabolisms is discussed below.

1.1 Molecular evidence for the evolution of sulfate reduction 4

The ability to use sulfate as a terminal electron acceptor for energy conservation constitutes a distinctive feature of certain bacterial lineages and one hyperthermophilic archaeal genus. These microorganisms include Gram-positive staining sporulating bacteria (genus Desulfotomaculum, phylum Firmicutes), Gram-negative staining sulfate-reducing bacteria of the Deltaproteobacterial class (ie., genera Desulfovibrio, Desulfobulbus, Desulfobacter, Desulfobacterium, Desulfococcus,

Desulfosarcina) and the thermophilic Thermodesulfovibrio (phylum Nitrospirae),

Thermodesulfobacterium (phylum Thermodesulfobacteria), and the sulfate-reducing archaeal genus Archaeoglobus (phylum Euryarchaeota) (32). They represent a diverse group of prokaryotes and play indispensable roles in the global cycling of carbon and sulfur. Equally to their contributions to the sulfur cycle, sulfate-reducing microorganisms are also essential regulators of diverse processes in wetland soils, including organic matter turnover, biodegradation of chlorinated aromatic pollutants in anaerobic soils and sediments, and mercury methylation (32).

Phylogenetic analysis based upon 16S rRNA genes suggested Thermodesulfo- bacteriaceae was an early sulfate reducer within the bacterial domain of the Tree of Life (33).

However, sequence comparisons of the core genes in the sulfate reduction process, especially dissimilatory sulfite reductase (dsr), demonstrate that members of the Thermodesulfobacteriaceae gained their dsr genes by lateral gene transfer (LGT) from later-evolved sulfate reducers of the

Deltaproteobacteria (33). Loy et al. (2008) (34), proposed that dsr (dissimilatory sulfite reductase) present in Thermodesulfobacteriaceae and some low-G+C Gram-positive staining bacteria of the phylum Firmicutes, are rooted more deeply than in Thermodesulfobacterium species or members of the Deltaproteobacteria, implying that the dsr gene emerged from early microbial evolution (34).

In addition, the ancient horizontal transfer of dsrAB is found to cross the boundaries of the domains of life, where the euryarchaeotic genus Archaeoglobus possesses a bacterial originated dsrAB (35). 5

Archaeoglobus fulgidus (36) and Archaeoglobus profundus (37) are phylogenetically related to the

Methanosarcina genus. Though not producing methane, they possess some methanogenic cofactors, such as methanofuran, methanopterin, and coenzyme F420, but not coenzyme M, coenzyme B, and coenzyme F430, (38). The other two key enzymes that catalyze the earlier steps in sulfate reduction, ATP sulfurylase (39) and adenosine 5'-phosphosulfate (APS) reductase (40), have also been characterized in Archaeoglobus fulgidus. Thus, this organism contains the complete pathway for dissimilatory sulfate reduction known from Bacteria. A 16S rRNA-based phylogenetic analysis has suggested that apsA, the gene that encodes the α-subunit of the APS reductase, was also subject to a few lateral gene transfers among sulfate-reducing microorganisms (41). However, the pattern of lateral gene transfer between apsA and dsr seems to be different (41). The mechanisms underlying these patterns of lateral gene transfer events are yet to be investigated.

Moreover, a novel, highly active, coenzyme F420-dependent sulfite reductase (Fsr) has been identified in Methanocaldococcus jannaschii (42). Unlike most methanogens,

Methanocaldococcus jannaschii is not sensitive to sulfite and can grow with it as the sole sulfur source. In this hyperthermophilic methanogen, sulfite induces Fsr, which reduces sulfite to sulfide, using reduced F420 (H2F420) as the electron donor (42). Fsr’s primary role may be sulfite detoxification as well as S assimilation (42). The N-terminal half of Fsr is a homolog of H2F420 dehydrogenase (FqoF/FpoF). FqoF/FpoF is the electron input unit of a membrane-bound electron transport system of late-evolving methylotrophic methanogens, such as Methanosarcina mazei, and Archaeoglobus fulgidus (42). The C-terminal half (Fsr-C) has homology to the siroheme- containing dissimilatory sulfite reductases of the archaea and bacteria (DsrA) (42). Only four methanogens possess Fsr, including three other extremophilic methanogens and an uncultured archaeon from a consortium performing anaerobic oxidation of methane (AOM) (42). Each of 6

these methanogens possesses a small putative sulfite reductase with sequence similarity to Fsr-C.

Fsr is then proposed to be the ancestor of F420 dehydrogenases, which constitute the electron input units for membrane-based energy transduction systems of certain late-evolving archaea and dissimilatory sulfite reductases of bacteria and archaea (42). Alternatively, Fsr is possibly formed from lateral gene transfer and gene fusion events (42).

Recently, a more complex whole-genome analysis was carried out to investigate the ancient biogeochemical events using modern genomes (43). In this study, individual gene phylogenies were resolved against organismal phylogenies by constructing models of gene birth, gene transfer, gene duplication and horizontal transfers (43). A very early emergence of a wide-ranging sulfur metabolism is pictured in this study, including sulfate reduction, also with a particular significance for sulfite and thiosulfate transformations (43). Overall, certain sulfur metabolisms are ancient, though the order of the emergence is unclear (11, 43).

1.2 Geochemical evidence for the antiquity of S metabolism

Microbial sulfate reduction fractionates the common stable sulfur isotopes 32S, 33S, 34S and

36S (44-46). In nature, prokaryotes reduce sulfate to sulfide while oxidizing organic substrates or

2- - - - H2 (e.g., SO4 +CH3COO +H2OH2S+2HCO3 +OH ) (47, 48). In the fractionation process of sulfate reduction, the product sulfide is depleted in the minor sulfur isotopes (33S, 34S and 36S) compared to the most abundant isotope 32S (45). Laboratory experiments using pure cultures and natural populations of sulfate-reducing microbes have suggested that sulfides are enriched in 32S

34 34 with fractions (≈ δ Ssulfate - δ Ssulfide) of 2-46 % (47, 49-51), but more commonly around 10 % and 40 % relative to parent sulfate (24, 45, 52). The sulfide formed mostly reacts with metal ions, especially Fe, and is largely precipitated as sedimentary pyrite (FeS2) (53-55). Exploring the patterns and magnitude of these fractionated sulfur isotopes in sulfate and pyrite in sedimentary 7

rocks has been regarded as the most solid diagnostic approaches to probe the early evolution of microbial sulfate reduction (11, 56, 57).

The earliest evidence for the presence of large fractionation characteristic of ancient sulfate reduction are in rocks from 3.49 Ga (billion years ago) quartz-barite deposits of the Dresser

Formation, North Pole, Western Australia (58). In this report, microscopic sulphides in barite crystals show a wide range of δ34S values with large fractionations, maximum of 21.1 % and a mean of 11.6 %, relative to co-existing sulfate (58).

Recent reports on mass-independent sulfur-isotope anomalies (MIF-S) in sedimentary sulfur (sulfide and sulfate) from the Archean (59), coupled with photochemical experiments (28) and atmospheric models (31), have supported the hypothesis that the Archean atmospheric O2 concentration was 10-5 times lower than that of the present atmospheric level (PAL) (60). In this model, prior to 2.4 Ga, low levels of ozone, due to the low concentration of atmospheric O2, permitted the penetration of shorter wavelength radiation into lower-levels of the Earth’s atmosphere (24). This high energy radiation facilitated the photochemical reactions involving the sulfur gaseous species such as volcanic SO2 or SO, yielding subsequent sedimentary preservation of sulfate (Figure 1-1) (11, 44). Consequently, this complex photochemistry interplay generated

33 both a reduced sulfur pool (S8 aerosols, H2S, HS) with a highly positive Δ S and an oxidized

33 36 sulfur pool (sulfate aerosols and SO2) with modestly negative Δ S (25, 28, 60). The Δ S is inversely correlated with Δ33S (25, 28, 60). Several recent sulfur mass-independent fractionation

(MIF) isotopic studies on pyrite sediments of Archean age have documented the presence of microbial sulfate reducers and/or microbial sulfur disproportionators, presumably using substrates generated from photochemical reactions (22, 47, 60-63) (Table 1-1). The latest observation has identified δ33S originating from sulfide reducers in metasediment from the Nuvvuagittuq 8

Greenstone Belt, Canada of over 3.8 Ga (64). This unusual atmospheric record has expanded the

S-isotope pattern back to the end of the Hadean, the earliest eon of Earth’s history (64).

Other evidence for microbial sulfur disproportionation is not related to MIF-S but based on the variation in 33S fractionation produced in sulfur disproportionation and sulfate reduction pathways (65, 66). Johnston et al. (65) argued that sulfur compound disproportionation was an active part of the sulfur cycle by 1.3 Ga and may have represented an onset of a progressive oxygenation of Earth’s surface. Canfield and Teske (67) made a similar argument that atmospheric oxygen increased from lower to higher than 5-18 % PAL between 0.64 Ga and 1.5 Ga based on the magnitude of 34S fractionations.

1.3 Conclusion

Among all elements, sulfur has been a key to understanding the history of Earth’s surface chemistry and the abundance of molecular oxygen in the atmosphere and ocean. The sulfate concentration in the ocean was less than 200 µM before the Great Oxidation Event (GOE) about

2.4 Ga (68). Presumably, before GOE, the influence of microbial life on the environment with limited O2 and sulfur species was more subtle than what’s seen today (69). Following the GOE, the sulfate pool increased in parallel with the O2, leading to the onset of widespread microbial sulfate reduction (23, 70). Undoubtedly, the dynamic sulfur cycle has played a great role in our understanding of the history of early Earth.

2. Sulfur metabolism in methanogens

2.1 Sulfur sources for methanogens

In aerobes, sulfur is incorporated into cells as sulfate, which is reduced to sulfide in an eight-electron reduction pathway (71-73). This assimilatory sulfate reduction pathway is the most common and widespread process in plants and microorganisms. In these organisms, sulfate is first 9

assimilated into the cells by membrane-bound proteins, then activated with ATP to form adenosine-5’-phosphosulfate (APS) and further activated with another ATP to form 3’- phosphoadenylsulfate (PAPS) (Figure 3-1) (71). Using thioredoxin as the electron donor, PAPS is subsequently reduced to sulfite and finally reduced to sulfide with NADPH as the electron donor

(Figure 3-1) (71). Because of the toxicity of sulfide, higher plants and many bacteria only use it as the direct S donor for the biosynthesis of L-cysteine, which serves as a central organic sulfur donor for the biosynthesis of L-methionine, glutathione, Fe-S clusters, CoA, lipoic acid, thiamine, molybdopterin, thionucleotides and other organic sulfur-containing compounds (68, 71, 73-75)

(Figure 1-3). For some processes, cysteine desulfurase further decomposes cysteine to alanine and sulfane sulfur via the formation of an enzyme-bound persulfide intermediate (‒SSH), which then functions as the sulfur donor for the biosynthesis of Fe-S clusters and thionucleotides (30-33). The persulfide group is subsequently incorporated into ubiquitin-like S carrier proteins to form a thiocarboxylate group, which functions as the S donor for thiamine and molybdopterin biosynthesis (Figure 1-3) (34, 35). In light of the limited availability of sulfate on the early earth

(68), the relatively late evolution of sulfate reduction pathway and the significant energy consumption to incorporate sulfate (69, 76, 77), the widespread modern sulfate assimilation process may not have been common in the ancient past (78). Unlike plants and most bacteria, methanogens are generally unable to assimilate sulfate. Instead, all known methanogens can use sulfide or elemental sulfur (S0) as the sole sulfur source (79). Only Methanococcus thermolithotrophicus and Methanobrevibacter ruminantium are able to use sulfate as a sole sulfur source (80, 81). This is not surprising considering that sulfate reducers are commonly present in the habitats of most methanogens and sulfide is readily available (24, 82). Similarly, for methanogen habitats on the interface between aerobic and anaerobic zones, the sulfide formed in 10

the anaerobic zone may be oxidized into S0 in an abiotic or biotic manner and make this alternative sulfur source available (82).

The methanococci usually inhabit environments with high levels of sulfide, and they are normally cultured with 3-5 mM Na2S (10, 83-85). Moreover, the intracellular sulfide concentrations of methanococci may be in the millimolar range due to the free diffusion of non- ionized sulfide across the cellular envelope (10). Consistent with their tolerance of high levels of sulfide, methanococci lack targets for sulfide toxicity, such as α, β-unsaturated fatty acid biosynthetic intermediates and cytochromes (70). Rauch and Perona (86) have recently identified an ApbE-like (ApbE: alternative pyrimidine biosynthesis protein) protein COG2122 responsible for sulfide assimilation and trafficking in Methanosarcina acetivorans. A deletion strain lacking ma1715, the gene encoding COG2122, was unable to grow with low concentrations of sulfide as the sole sulfur source. However, rapid growth is recovered when the deletion mutant was supplemented with cysteine, homocysteine, or cystathionine. Together with other genotype- dependent experiments, COG2122 is suggested to be necessary for efficient biosynthesis of both cysteine and homocysteine, but it is not required for maintaining cellular levels of iron-sulfur clusters or sulfur modification in tRNA (86). Although the physiological role of COG2122 has not been fully characterized, mass spectrometry indicates that it may serve as sulfur trafficking protein to generate persulfide with ambient sulfide and transfer sulfane sulfur to proteins, such as Sep- tRNA:Cys-tRNA synthase (SepCysS), for cysteine and homocysteine biosynthesis (87).

In the anoxic and geothermally-influenced Archaean environment, elemental sulfur may have been formed in variable concentrations by photochemical or other forms of oxidation of volcanic sulfur-bearing gaseous species (11, 44). Microorganisms known to reduce elemental sulfur using H2 or organic substrates as electron donors are widespread among Bacteria and 11

Archaea (88, 89). In contrast to bacterial sulfur-reducers, which are usually mesophilic or moderately thermophilic, archaeal sulfur reducers are all hyperthermophilic (88). Most methanogens can assimilate elemental sulfur as a sulfur source (79). In the presence of H2, many methanogens produce a high amount of sulfide from S0 (79, 82). Although abiotic S0 disproportionation produces a significant amount of sulfide, thermophilic and hyperthermophilic methanogens, Methanopyrus, Methanothermobacter, Methanothermus, and Methanocaldococcus also produce a substantial amount of sulfide (79, 90). Among them, Methanothermococcus thermolithotrophicus, which grows at 65 oC, produces sulfide at the highest rate (around 0.85 mol per mg cell dry wt per hour), which is three-fold higher than the abiotic reaction under the same conditions (17). Meanwhile, mesophilic methanogens also reduce S0 well above the rate of abiotic sulfur disproportionation (50). Fauque et al. have detected sulfur reductase activities in crude extracts of Methanosarcina barkeri, Methanothermococcus thermolithotrophicus and

Methanothermobacter thermoautotrophicus (79, 82). Nevertheless, the enzyme that catalyzes S0 reduction in methanogens has not yet been identified.

The anoxic, early Archaean contains a mixed form of sulfur species, which leads to a variety of sulfur-based life, such as sulfide reducers, sulfide oxidizers, sulfate reducers and sulfur disproportionators. However, the role of these sulfur species in ancient metabolism are still unclear

(89). Thiosulfate or sulfite can use be used as alternative sole sulfur source for certain methanogens, namely Methanocaldococcus jannaschii, Methanothermococcus thermolithotrophicus,

Methanothermobacter thermautotrophicus, Methanobrevibacter ruminantium and

Methanosarcina barkeri (80, 81). Thiosulfate and sulfide reductase activities have been detected in the crude extracts of Methanosarcina barkeri strain 227, Methanothermobacter marburgensis and Methanothermococcus thermolithotrophicus (82). Methanosarcina barkeri strain Fusaro has 12

only thiosulfate but not sulfate or sulfite reductase (81). Two sulfite reductases have been characterized (42, 91, 92). A small siroheme (23 kDa) sulfite reductase, P590, has been isolated from Methanosarcina barkeri DSM 800 (91, 92). P590 sulfite reductase contains one Fe-siroheme and one [4Fe-4S] cluster per polypeptide chain (91, 92). This sulfite reductase reduces sulfite to sulfide in vitro using the artificial electron donor methyl viologen (91, 92). The physiological electron donor and the in vivo role for P590 have not yet been determined (91, 92). A 70 kDa F420- dependent sulfite reductase (Fsr) has been isolated and characterized in Methanocaldococcus jannaschii (42, 93). Purified Fsr reduces sulfite to sulfide using reduced F420 (H2F420) as the electron donor (42). The N-terminal of Fsr contains a F420 dehydrogenase (FpoF or FqoF) domain, and the C-terminal is homologous to DsrA or DsrB subunits of siroheme-containing dissimilatory sulfite reductase (42). Fsr homologs are present in Methanothermobacter thermautotrophicus,

Methanopyrus kandleri, and Methanococcoides burtonii (42, 93). Heterologous expression of Fsr in Methanococcus maripaludis permits this methanogen to take up sulfite as the sole sulfur source, further supporting the view of Fsr engaging in sulfite detoxification and assimilation (93).

Many methanogens are able to utilize several mercaptans (methanethiol, ethanethiol, n- propanethiol, n-butanethiol, methyl sulfide, dimethyl sulfoxide, ethyl sulfide or CS2) and organic sulfides as the sulfur source for growth, though these cultures require a supplemental 20-30 µM sulfide to initiate growth (81). These methanogen species are M. thermolithotrophicus, M. jannaschii and M. thermautotrophicus (81). Presumably, these sulfur compounds are converted to sulfide as the sulfur source to support growth, but the corresponding enzymes and mechanism are yet to be studied (81).

2.2 Biosynthesis of sulfur-containing compounds.

Biosynthesis of Cysteine. In most microorganisms, cysteine biosynthesis and Cys- 13

tRNACys formation are operated in two separated processes by the cysteine synthase and cysteinyl- tRNA synthase (CysRS), respectively (Figure 1-2). However, many methanogens lack the canonical class I CysRS, together with other proteins analogous to those involved in bacterial or eukaryotic cysteine biosynthesis (94). Instead, these methanogens possess a tRNACys-dependent cysteine biosynthetic pathway (95). Cys-tRNACys is produced in a two-step process: first, O- phosphoserine (Sep) is acylated to tRNACys by O-phosphoseryl-tRNA synthetase(SepRS), and then Sep-tRNACys is converted to Cys-tRNACys by a PLP-dependent enzyme: Sep-tRNA:Cys- tRNA synthase (SepCysS) (95). An additional component, SepCysE, serves as a scaffold for

SepRS and SepCys to form the SepRS·SepCysS·tRNACys ternary complex, presumably enabling the substrate channeling of Sep-tRNACys (96). SepCysE it is only present in class I methanogens

(96, 97). For certain methanogens (e.g., Methanococcales and thermophilic Methanobacteriales species) that lack the canonical CysRS, this indirect cysteine coding route is the sole pathway to synthesize Cys-tRNACys (70). Some other methanogens, including all three Methanosarcina species whose genome sequences are available, retain both pathways (98, 99). The redundancy of carrying multiple cysteine biosynthesis routes in these organisms has not been studied.

The coupled biosynthesis and coding of Cys has been proposed to be the original mechanism of Cys-tRNACys formation in the last common ancestor of archaea (LUCA) based on the following observations. First, archaeal CysRS genes are found to have multiple bacterial origins (99, 100), and bacterial CysRS is a highly evolved Cys-specific enzyme using a zinc atom to ensure specificity (101-103). Additionally, several phylogenetic studies suggest that CysRS was absent in LUCA and eukaryotes and was horizontally transferred from bacteria through multiple independent events (99, 104, 105). However, a more recent phylogenetic analysis employing all current available genomic and metagenomic protein sequence had identified the distributions of 14

SepRS and SepCysS across all domains (103). Both SepRS and SepCysS are present in four clades of archaea: Euryarchaeota, DPANN, TACK, and Asgard, and two groups of bacteria: Candidatus

Parcubacteria and Chloroflexi (103). These two bacterial SepRS and SepCysS homologs from uncultured Candidatus Parcubacteria and Chloroflexi bacteria were heterologous expressed in E. coli and shown to successfully charge bacterial tRNACys species with cysteine (103). Homologs of

SepCysE, are found in a few groups of TACK and Asgard archaea, whereas the C-terminally truncated homologs exist fused or genetically coupled with diverse SepCysS species (103). These explorations of the distribution of the SepRS-SepCysS-SepCysE system in diverse archaea and bacteria provide an expanded record of the alternative cysteine-coding system’s contribution in sulfur assimilation and diverse metabolism.

Safro et al. (94) clustered the archaeal proteomes into two groups according to their cysteine content. Interestingly, methanogens and Archaeoglobus possess high cysteine content (an average of ~1.3 % of the entire proteome according to genomic sequences) when compared to non- methanogenic archaea (~0.7 %) (94). Presumably, a reduced cysteine content in non-methanogenic archaeal lineages is correlated with their loss of the tRNA-dependent Cys biosynthesis pathway

(70). Nonetheless, the distribution of SepRS-SepCysS pathway across archaea and bacteria (103) and the selection pressure of retaining either or both Cys-tRNACys charging pathway in those organisms (99) are yet to be elucidated.

A number of fundamental questions still remain concerning the tRNA-dependent cysteine biosynthesis pathway. Foremost, the immediate, physiological sulfur donor for changing Sep- tRNACys to Cys-tRNACys is unclear. SepCysS, the enzyme that catalyzes this step and its mechanism also has not been characterized. In bacteria and eukaryotes, cysteine is the sulfur donor for Fe-S cluster biosynthesis. Many bacteria also use cysteine as a precursor for methionine 15

biosynthesis. However, in Methanococcus maripaludis, the intracellular concentration of free cysteine is 20 µM, 5-10-fold less than commonly seen in bacteria. Moreover, cysteine is not the sulfur donor for Fe-S cluster and homocysteine biosynthesis (10). Interestingly, heterologous expression of Methanocaldococcus jannashii SepCysS in E. coli has determined that a persulfide group (containing a sulfane sulfur) is the proximal sulfur donor for cysteine biosynthesis (106). J.

Perona et al. (87) recently used purified natively-expressed Methanosarcina acetivorans SepCysS in their study. Here, the purified SepCysS was found to be active in single-turnover reactions converting Sep-tRNACys to Cys-tRNACys without exogenously added sulfur (87). Therefore, it is likely that the active-site persulfide group of SepCysS is a direct sulfur donor for the incorporation of cysteine into proteins in Methanosarcina acetivorans (87). Regardless, the sulfane sulfur donor to this cysteine residue still requires further investigations to advance the understanding of the

SepCysS mechanism.

Biosynthesis of homocysteine. In most organisms, sulfur assimilation is initiated with sulfate reduction (71). Sulfate is taken up into the cells and reduced into sulfide through a highly conserved, ATP-dependent pathway (71) (Figure 1-3). The sulfide is incorporated into homocysteine via a direct sulfhydration of homoserine, operated by O-acetylhomoserine sulfhydrylase (OAHS), O-succinylhomoserine sulfhydrylase (OSHS), or O-phosphohomoserine sulfhydrylase (OPHS) (71). Homocysteine later becomes a precursor for methionine and S- adenosylmethionine (SAM) biosynthesis (71). Alternatively, intracellular sulfide can also be incorporated into O-acetylhomoserine to form cysteine, catalyzed by O-acetylserine sulfhydrylase

(OASS) (71). Cysteine and homocysteine are interconverted by transulfuration pathways via the enzyme cystathionine gamma-synthase (CGS) and cystathionine beta-lyase (CBL) (71). Cysteine then serves as a substrate for both CysRS and cysteine desulfurase (CD) (71). The CD liberates 16

the sulfur from free cysteine, forming alanine and an enzyme-bound persulfide (-S-SH) (71). This terminal sulfur from the persulfide group is then relayed as sulfane sulfur for further mobilization and incorporation into iron-sulfur clusters, tRNA and other metabolites (71) (Figure 1-3). However, homologs of most of these canonical proteins are absent from methanogens (107) (Figure 1-2).

Impressively, Methanosarcina acetivorans possesses the majority of genes for homocysteine and cysteine biosynthesis, such as OAHS, metA (homoserine O- succinyltransferase), CD, CysRS, SepRS and SepCysS (6). Proteins involved in these redundant sulfur assimilation and trafficking pathways are suggested to be derived from horizontal gene transfer from bacteria (6). Homocysteine in Methanosarcin acetivorans is synthesized by two parallel pathways (108). First, homologs for metA (homoserine O-succinyltransferase) and OAHS are present in the genome, and homocysteine can be produced from O-acetylhomoserine and sulfide (108). The alternative biosynthetic pathway of homocysteine has been identified recently, performed by two proteins (108, 109). MA1821 is composed of a cystathionine β-synthase (CBS) domain at the C-terminus and a DUF39 domain at the N-terminus (108). MA1822, located immediately at the downstream of MA1821, is a NIL-ferredoxin (108). These two proteins together catalyze a unique reductive condensation of sulfide with aspartate-semialdehyde to produce homocysteine (108, 109). Deletion strains lacking ma1821-22 or oahs alone grew in medium containing sulfide as the sole sulfur source, whereas the triple mutant Δma1821-22Δoahs showed no growth (108). Thus, both of these pathways appear to contribute to homocysteine production in Methanosarcina acetivorans (108). Mass spectrometry analysis on anaerobically purified MA1821 protein has identified high frequency of persulfide formation on the conserved cysteine residues (87). The function of these persulfide groups, if any, is not clear yet.

Many methanogenic archaea lack homologs to the bacterial proteins in homocysteine 17

biosynthesis. An example is Methanococcus maripaludis. In this archaeon, cell extracts produce homocysteine from cystathionine but not from H2S and O-phosphohomoserine, O-acetylhomoserine or O-succinyl-homoserine, suggesting that homocysteine is not formed by direct sulfhydrylation of homoserine derivatives (10) (Figure 1-2). Isotope labeling also demonstrated that cysteine is not the sulfur donor for homocysteine (10). Methanocaldococcus jannashchii also lacks homologs for the bacterial homocysteine biosynthetic genes. However, MJ0100 and MJ0099 are homologous to MA1821 and MA1822, respectively. More importantly, isotopic studies using cell extracts have demonstrated Methanocaldococcus jannashchii can biosynthesize homocysteine from aspartate semialdehyde and hydrogen sulfide. Taken together, these experiments demonstrated that aspartate semialdehyde is a likely precursor of homocysteine in many methanogens.

Biosynthesis of methionine. Protein homologs of critical components in methionine biosynthesis are absent in methanogens. In Methanococcus maripaludis, homocysteine is suggested as an intermediate of the methionine biosynthesis based on a 34S-labeling study (10) and bioinformatics analysis (110). This isotopic feeding experiments in Methanococcus maripaludis further found the sulfur in methionine was acquired from inorganic sulfide but not cysteine (10).

MetE, the enzyme that catalyzes the last step of methionine biosynthesis, is encoded in the methanogenic genomes. MetE methylates homocysteine into methionine, in the absence of cobalamin. The activity of this MetE has been characterized in Methanothermobacterium thermoautotrophicus (111). In this in vitro study, MetE from Methanothermobacter thermoautotrophicus only uses methylcobalamin and methylcobinamide as the methyl donor to produce methionine, but not methyltetrahydrofolate or methyltetrahydromethanopterin (111).

However, methylcobalamin is not considered to be the physiological methyl donor of MetE, 18

because MetE lacks a specific binding site for methylcobalamin (111). In addition, the intracellular concentration of free corrinoids in Methanothermobacter thermoautotrophicus is very low, nearly all of the corrinoids are tightly bound to corrinoid proteins (112). Therefore, the methyl donor of converting homocysteine to methionine is yet to be known.

Biosynthesis of iron-sulfur clusters. Iron-sulfur clusters exist in almost all modern organisms as they are indispensable elements of many proteins, such as redox proteins, S- adenosylmethionine-dependent enzymes, and RNA polymerase (113). In bacteria, three types of machinery are known to biosynthesize iron-sulfur clusters, termed NIF (nitrogen fixation), ISC

(iron sulfur cluster), and SUF (mobilization of sulfur) (113)．The NIF system was first discovered in the nitrogen-fixing bacterium Azotobacter vinelandii (114, 115) and is responsible for the maturation of nitrogenase, whereas the ISC and SUF systems permit the maturation of all other

Fe/S proteins in the cell. E. coli has both ISC and SUF systems for iron-sulfur cluster assembly

(116, 117). In E. coli, SUF system is specifically adapted to synthesize iron-sulfur clusters in extreme conditions, such as oxidative or heavy-metal stress and iron starvation, indicating that

SUF takes the predominate role for iron-sulfur synthesis while ISC is likely the housekeeping iron- sulfur cluster assembly system (116-119). Interestingly, ISC is present in bacteria and most eukaryotes whereas SUF is found in bacteria, archaea, plants, and parasites (120). In eukaryotes,

O2-producing compartments like the chloroplast employ an iron-sulfur machinery similar to the bacterial SUF system, presumably because of its lower sensitivity towards oxygen compared to the ISC system (121). Given that at the initial stage of bacterial infection, properties like acquiring iron and resisting oxidative stress are indispensable, the SUF system is considered to play a critical role in the pathogenicity of bacteria like Mycobacterium tuberculosis (122).

In these three different machineries (NIF, ISC and SUF), cysteine desulfurase (NifS, IscS 19

and SufS), is commonly present to mobilize and transfer the sulfur from cysteine to scaffold proteins (120). Additionally, they all possess scaffold proteins (IscU/ISU, IscA/ISA, NifU, NFU,

SufU and SufA), which act as intermediate assembly sites for iron-sulfur cluster or iron-sulfur cluster precursors before delivery to target proteins (120). Bacterial IscS and IscU homologs can be found in certain euryarchaeaotes. However, many other archaea, including the euryarchaeaotes

Pyrococcus furiosus, Methanocaldococcus jannaschii, and Methanococcus maripaludis, and most crenarchaeaotes, either lack a homologous cysteine desulfurase gene or have no homologs of A- type or U-type scaffold genes (123). Only three proteins of the known iron-sulfur cluster assembly and transport machineries are conserved in all methanogens: SufB, a scaffold protein for iron- sulfur cluster assembly; SufC, an ABC-type ATPase that might facilitate the transfer of iron-sulfur clusters from scaffold proteins to target proteins; and ApbC/Nbp35, a carrier protein that could be involved in iron-sulfur cluster transport (70, 123). Given that methanogen inhabit a sulfide- abundant environment and lack homologs to most of the SUF and other known Fe-S cluster assembly systems, they may utilize novel proteins and inorganic sulfur sources to assemble iron- sulfur clusters.

Biosynthesis of sulfur-containing tRNAs. The occurrence of post-transcriptional tRNA modifications is widespread in all domains of life with very broad diversities. The roles of modified nucleotides in tRNA are demanding and diverse, such as stabilizing tRNA structure, maintain reading frame, identifying elements for the translation machinery by aminoacyl-tRNA synthetases, facilitating ribosomal binding to aminoacylated tRNAs, and proofing codon-anticodon base pairing (124, 125). The commonly known nucleotide positions on tRNA for thiolation are U32,

U33, U34, U37, and U54 (126). These thio-modifications are 2-thiouridine (s2U) derivatives, 4- thiouridine (s4U), 2-thiocytidine (s2C), and 2-methylthioadenosine (ms2A) (126). The function of 20

these modifications is wide-ranging and depends on their positions on the tRNA (126). Presumably, modifications located outside the anticodon loop are often related to stabilize the tRNA structure, while modifications within the anticodon loop are usually associated with improving translational fidelity and efficiency (126).

The biosynthesis of these thionucleosides usually involves a cascade of sulfur-carrier proteins rather than a direct transfer from the ultimate sulfur donor to the substrate (127). This process generally begins with the activation of sulfur from free cysteine by cysteine desulfurase, such as IscS in bacteria (128) and Nfs1 in eukaryotes (129), and the formation of an enzyme-bound persulfide (R-S-SH) (130). The terminal sulfur from the persulfide group is then donated through downstream sulfur carrier proteins to the tRNA thiolation enzymes and finally to the target tRNA nucleosides. It is worthwhile to mention that, besides tRNA thiolation, the carrier protein plus cysteine desulfurase also delivers sulfur to a variety of cofactors and iron-sulfur clusters (131).

This thiolation process is usually classified into two major groups depending on the involvement of iron-sulfur cluster biosynthesis. Because the biosynthesis of s4U8 and s2U34 are the most extensively studied tRNA modifications in methanogens, the following discussion will mainly focus on these two types.

The 4-thiouridine modification at position 8 of tRNA (s4U8) is conserved in Bacteria and

Archaea (132). s4U8 modification has not yet been described in eukaryotes, though the genomic homologs of the prokaryotic s4U8 modification enzyme (ThiI) are found in some eukaryotes (133).

The s4U8 serves as a photosensor of near-ultraviolet (UV) radiation physiologically (134-136).

When exposed to irradiation, s4U8 cross-links with the nearby cytidine at position 13, thus inducing structural changes that prevent tRNA aminoacylation and leading to the accumulation of uncharged tRNA (134-136). This effect mimics amino acid starvation and eventually triggers 21

stringent cellular responses (134-136).

2-Thiouridine (s2U) derivatives are present at the first anticodon base (position 34, U34) in

Gln Lys Glu tRNA UUG, tRNA UUU, and tRNA UUC in all three domains of life. U34 can be hypermodified to a variety of s2U derivatives (xm5s2U), depending upon the organism and the subcellular location

(126). The rigid conformation of xm5s2U in the C3´-endo form (137) guarantees an efficient codon: anticodon base pairing (138, 139). The presence of s2U in the codon: anticodon pair also results in a preference for A-ending codons (140), presumably due to the greater stability of the s2U-A vs. s2U-G pair (141). Moreover, the 2-thio group of xm5s2U acts as an identity element in aminoacylation reactions (142-144), facilitating tRNA binding to the ribosomal A-site (142) and arresting frameshifting during translation (145). Due to the key roles of s2U34 modification in translation, its impaired activities are found to result in a pleiotropic phenotype in yeasts and various diseases in humans (127).

The bacterial s4U8 and s2U34 biosynthesis pathways are iron-sulfur cluster independent and require two proteins for these modifications: IscS and ThiI. In E. coli, the cysteine desulfurase

IscS takes the sulfur from a free L-cysteine and generates a persulfide group (75). This persulfidic sulfur in IscS can then be directly transferred to the ThiI that catalyze s4U8 formation (139, 146).

Alternatively, during s2U34 modification in bacteria, a sulfur relay chain composed of multiple intermediate persulfide carriers (TusA, TusBCD complex and TusE) are used to pass the persulfide group of IscS to the s2U34 thiolation enzyme MnmA (139, 146). However, the biosynthesis of s2U34 modification in the eukaryotic cytosol is very different from that in bacteria. In eukaryotes, cysteine desulfurase Nfs1 mobilizes the sulfur from its persulfide group to an ubiquitin-related modifier (Urm1) via multiple persulfide carriers (147, 148). A thiocarboxylate group is then generated on the carboxyl terminus of Urm1 and likely is the sulfur donor for s2U34 formation 22

(147, 148). The final step enzyme that catalyzes the formation of s2U34 is an enzyme complex designated as Ncs6/Ncs2 in yeasts, Ctu1/Ctu2 in nematodes, and ATPBD3/CTU2 in humans (148-

150). This enzyme complex eventually activates and thiolates the C2 atom of U34.

The methanogenic archaea s4U8 biosynthesis pathway is found to be different from bacteria.

Deletion of ThiI (MMMP1354) in Methanococcus maripaludis eliminated the biosynthesis of s4U8

(151). MMP1354 enzyme can use sulfide as an in vitro sulfur donor for U8 thiolation (151). In vitro assay has also found the Km for sulfide donor is ~ 1 mM, close to the intracellular concentration of free sulfide in methanococci (~ 1-3 mM) (151). Therefore, it is likely sulfide is a physiologically relevant donor for s4U8 formation catalyzed by MMP1354 (151). This hypothesis is supported by the fact that cysteine is not the sulfur donor for iron-sulfur clusters and tRNA thionucleotides, and cysteine desulfurase is absent in most methanogenic genomes. Recent studies have also described the catalytic role of the three conserved cysteinyl residues on MMP1354 (151).

More importantly, a [3Fe-4S] cluster is found on MMP1354 and is essential for the enzyme’s tRNA thiolation activity (127). This suggests that the methanogenic archaea operate the s4U8 biosynthesis in a distinctive way from bacteria, and it is iron-sulfur cluster dependent.

The archaeal s2U34 biosynthesis is proposed to be similar to the eukaryotic cytosolic Ncs6 pathway (126, 127). This assumption is based on several observations (i) the Ncs6 homologs are present in all archaeal genomes (133); (ii) deletion strain lacking Ncs6 homologs (ncsA) in

Lys Haloferax volcanii do not form thiolated tRNA UUU (152); (iii) in Haloferax volcanii (153) and

Methanococcus maripaludis (154), Ncs6 homologs generate complexes with the ubiquitin-like small archaeal modifier protein (SAMP), which has high structural homology to Urm1; (iv) the

Haloferax volcanii E1-like protein UbaA activates SAMP in formation of a thioester intermediate

(155); (v) the deletion of either samp2 or ubaA in Haloferax volcanii abolished thiolated 23

Lys tRNA UUU (156); (vi) the Methanococcus maripaludis Ncs6 homolog possess a [3Fe-4S] cluster

(127). These discoveries indicate that archaeal s2U34 biosynthesis employs an iron-sulfur cluster containing Ncs6 homolog and an activated ubiquitin-like protein.

Taken together, the biosynthetic pathways of sulfur-containing tRNA are very complex and vary greatly between different organisms. Fundamental questions, such as the sulfur donor of the thiolation, the identifications of persulfide-carrier proteins, the reaction mechanisms of thiolation, are still waiting to be investigated.

3. Closing remarks and future prospects

The early, anoxic Earth may form a complex sulfur reservoir with limited sulfate, abundant sulfide and elemental sulfur, and many other sulfur compounds. Well beyond any uncertainty over how these sulfur species engage with each other and their ambient environment, the sulfur cycle is certain to have influenced the evolving chemistry of the Earth’s surface. This is supported by the tremendous records of sulfur’s involvement in a variety of biogeochemical cycles and processes through geologic time. Methanogens that assimilate sulfide and elemental sulfur as sole sulfur sources are ideal model organisms for the study of sulfur metabolism on the early Earth.

They may have evolved unique mechanisms to manage the specificities of sulfur incorporation mediated with sulfide or elemental sulfur under anaerobic conditions (Figure 1-4). Hence, the elucidation of these metabolic traits may shed light on ancient sulfur metabolism and its evolutionary process. In addition, the investigation of sulfur relay in methanococci will guide discovery of novel sulfur metabolic pathways that may be common in other anaerobes and advance our knowledge of sulfur chemistry of life.

References:

1. Hedderich R, Whitman WB. 2006. Physiology and biochemistry of the methane-producing

archaea. Prokaryotes: A Handbook on the biology of bacteria, Vol 2, Third Edition

doi:10.1007/0-387-30742-7_34:1050-1079.

2. Liu Y, Whitman WB. 2008. Metabolic, phylogenetic, and ecological diversity of the

methanogenic archaea. Ann N Y Acad Sci 1125:171-89.

3. Ueno Y, Yamada K, Yoshida N, Maruyama S, Isozaki Y. 2006. Evidence from fluid

inclusions for microbial methanogenesis in the early Archaean era. Nature 440:516-519.

4. Reeve JN, Nolling J, Morgan RM, Smith DR. 1997. Methanogenesis: genes, genomes, and

who's on first? J Bacteriol 179:5975-5986.

5. Bapteste E, Brochier C, Boucher Y. 2005. Higher-level classification of the Archaea:

evolution of methanogenesis and methanogens. Archaea 1:353-63.

6. Galagan JE, Nusbaum C, Roy A, Endrizzi MG, Macdonald P, FitzHugh W, Calvo S,

Engels R, Smirnov S, Atnoor D, Brown A, Allen N, Naylor J, Stange-Thomann N,

DeArellano K, Johnson R, Linton L, McEwan P, McKernan K, Talamas J, Tirrell A, Ye

W, Zimmer A, Barber RD, Cann I, Graham DE, Grahame DA, Guss AM, Hedderich R,

Ingram-Smith C, Kuettner HC, Krzycki JA, Leigh JA, Li W, Liu J, Mukhopadhyay B,

Reeve JN, Smith K, Springer TA, Umayam LA, White O, White RH, Conway de Macario

E, Ferry JG, Jarrell KF, Jing H, Macario AJ, Paulsen I, Pritchett M, Sowers KR, et al. 2002.

The genome of M. acetivorans reveals extensive metabolic and physiological diversity.

Genome Res 12:532-42.

7. McBride BC, Wolfe RS. 1971. A new coenzyme of methyl transfer, coenzyme M.

Biochemistry 10:2317-24.

8. Lin WC, Yang YL, Whitman WB. 2003. The anabolic pyruvate oxidoreductase from

Methanococcus maripaludis. Arch Microbiol 179:444-56.

9. Major TA, Burd H, Whitman WB. 2004. Abundance of 4Fe-4S motifs in the genomes of

methanogens and other prokaryotes. FEMS Microbiol Lett 239:117-23.

10. Liu YC, Sieprawska-Lupa M, Whitman WB, White RH. 2010. Cysteine is not the sulfur

source for iron-sulfur cluster and methionine biosynthesis in the methanogenic archaeon

Methanococcus maripaludis. J Biol Chem 285:31923-31929.

11. Canfield DE, Farquhar J. 2012. The global sulfur cycle. Fundamentals of Geobiology:49-

64.

12. Rabus R, Bruchert V, Amann J, Konneke M. 2002. Physiological response to temperature

changes of the marine, sulfate-reducing bacterium Desulfobacterium autotrophicum.

FEMS Microbiol Ecol 42:409-417.

13. Canfield DE. 2005. The early history of atmospheric oxygen: Homage to Robert A.

Garrels. Annu Rev Earth Planet Sci 33:1-36.

14. Canfield DE, Thamdrup B. 1996. Fate of elemental sulfur in an intertidal sediment. FEMS

Microbiol Ecol 19:95-103.

15. Canfield DE, Thamdrup B, Fleischer S. 1998. Isotope fractionation and sulfur metabolism

by pure and enrichment cultures of elemental sulfur-disproportionating bacteria. Limnol

Oceanogr 43:253-264.

16. Finster K. 2008. Microbiological disproportionation of inorganic sulfur compounds. J

Sulphur Chem 29:281-292. 26

17. Obraztsova AY, Francis CA, Tebo BM. 2002. Sulfur disproportionation by the facultative

anaerobe Pantoea agglomerans SP1 as a mechanism for chromium(VI) reduction.

Geomicrobiol J 19:121-132.

18. Thamdrup B, Finster K, Hansen JW, Bak F. 1993. Bacterial disproportionation of

elemental sulfur coupled to chemical reduction of iron or manganese. Appl Environ

Microbiol 59:101-8.

19. Burdige DJ, Nealson KH. 1986. Chemical and microbiological studies of sulfide-mediated

manganese reduction. Geomicrobiol J 4:361-387.

20. Aller RC, Rude PD. 1988. Complete oxidation of solid-phase sulfides by manganese and

bacteria in anoxic marine-sediments. Geochimica Et Cosmochimica Acta 52:751-765.

21. Canfield DE, Thamdrup B, Hansen JW. 1993. The Anaerobic degradation of organic-

matter in aanish coastal sediments: iron reduction, manganese reduction, and sulfate

reduction. Geochimica Et Cosmochimica Acta 57:3867-3883.

22. Farquhar J, Wu NP, Canfield DE, Oduro H. 2010. Connections between sulfur cycle

evolution, sulfurisotopes, sediments, and base metal sulfide deposits. Economic Geology

105:509-533.

23. Lyons TW, Gill BC. 2010. Ancient sulfur cycling and oxygenation of the early biosphere.

Elements 6:93-99.

24. Johnston DT. 2011. Multiple sulfur isotopes and the evolution of Earth's surface sulfur

cycle. Earth-Science Reviews 106:161-183.

25. Farquhar J, Wing BA. 2003. Multiple sulfur isotopes and the evolution of the atmosphere.

Earth Planet Sci Lett 213:1-13.

26. Kasting JF. 1984. Earth history: the chemical evolution of the atmosphere and oceans.

Science 226:332-3.

27. Canfield DE. 2004. The evolution of the Earth surface sulfur reservoir. American Journal

of Science 304:839-861.

28. Farquhar J, Savarino J, Airieau S, Thiemens MH. 2001. Observation of wavelength-

sensitive mass-independent sulfur isotope effects during SO2 photolysis: Implications for

the early atmosphere. J Geophys Res Planets 106:32829-32839.

29. Canfield DE, Raiswell R. 1999. The evolution of the sulfur cycle. American Journal of

Science 299:697-723.

30. Walker JCG, Brimblecombe P. 1985. Iron and sulfur in the pre-biologic ocean.

Precambrian Res 28:205-222.

31. Pavlov AA, Kasting JF. 2002. Mass-independent fractionation of sulfur isotopes in

Archean sediments: Strong evidence for an anoxic Archean atmosphere. Astrobiology

2:27-41.

32. Dahl C, Friedrich CG. 2008. Microbial sulfur metabolism. Springer.

33. Klein M, Friedrich M, Roger AJ, Hugenholtz P, Fishbain S, Abicht H, Blackall LL, Stahl

DA, Wagner M. 2001. Multiple lateral transfers of dissimilatory sulfite reductase genes

between major lineages of sulfate-reducing prokaryotes. J Bacteriol 183:6028-6035.

34. Loy A, Duller S, Wagner M. 2008. Evolution and ecology of microbes dissimilating sulfur

compounds Insights from siroheme sulfite reductases. Microbial Sulfur Metabolism

doi:Doi 10.1007/978-3-540-72682-1_5:46-59.

35. Dahl C, Kredich NM, Deutzmann R, Truper HG. 1993. Dissimilatory sulphite reductase

from Archaeoglobus fulgidus: physico-chemical properties of the enzyme and cloning,

sequencing and analysis of the reductase genes. J Gen Microbiol 139:1817-28.

36. Mander GJ, Duin EC, Linder D, Stetter KO, Hedderich R. 2002. Purification and

characterization of a membrane-bound enzyme complex from the sulfate-reducing

archaeon Archaeoglobus fulgidus related to heterodisulfide reductase from methanogenic

archaea. Eur J Biochem 269:1895-904.

37. Burggraf S, Jannasch HW, Nicolaus B, Stetter KO. 1990. Archaeoglobus profundus sp.

nov., represents a new species within the sulfate-reducing archaebacteria. Syst Appl

Microbiol 13:24-28.

38. Klenk HP, Clayton RA, Tomb JF, White O, Nelson KE, Ketchum KA, Dodson RJ, Gwinn

M, Hickey EK, Peterson JD, Richardson DL, Kerlavage AR, Graham DE, Kyrpides NC,

Fleischmann RD, Quackenbush J, Lee NH, Sutton GG, Gill S, Kirkness EF, Dougherty

BA, McKenney K, Adams MD, Loftus B, Peterson S, Reich CI, McNeil LK, Badger JH,

Glodek A, Zhou L, Overbeek R, Gocayne JD, Weidman JF, McDonald L, Utterback T,

Cotton MD, Spriggs T, Artiach P, Kaine BP, Sykes SM, Sadow PW, D'Andrea KP,

Bowman C, Fujii C, Garland SA, Mason TM, Olsen GJ, Fraser CM, Smith HO, Woese

CR, et al. 1997. The complete genome sequence of the hyperthermophilic, sulphate-

reducing archaeon Archaeoglobus fulgidus. Nature 390:364-70.

39. Dahl C, Koch H-G, Keuken O, Trüper HG. 1990. Purification and characterization of ATP

sulfurylase from the extremely thermophilic archaebacterial sulfate-reducer,

Archaeoglobus fulgidus. FEMS Microbiol Lett 67:27-32.

40. Speich N, Dahl C, Heisig P, Klein A, Lottspeich F, Stetter KO, Truper HG. 1994.

Adenylylsulphate reductase from the sulphate-reducing archaeon Archaeoglobus fulgidus:

cloning and characterization of the genes and comparison of the enzyme with other iron-

sulphur flavoproteins. Microbiology 140 (Pt 6):1273-84.

41. Friedrich MW. 2002. Phylogenetic analysis reveals multiple lateral transfers of adenosine-

5'-phosphosulfate reductase genes among sulfate-reducing microorganisms. J Bacteriol

184:278-89.

42. Johnson EF, Mukhopadhyay B. 2005. A new type of sulfite reductase, a novel coenzyme

F420-dependent enzyme, from the methanarchaeon Methanocaldococcus jannaschii. J Biol

Chem 280:38776-38786.

43. David LA, Alm EJ. 2011. Rapid evolutionary innovation during an Archaean genetic

expansion. Nature 469:93-96.

44. Kaplan IR, Rittenberg SC. 1964. Microbiological fractionation of sulphur isotopes. J Gen

Microbiol 34:195-212.

45. Canfield DE. 2001. Biogeochemistry of sulfur isotopes. Stable Isotope Geochemistry

43:607-636.

46. Johnston DT, Farquhar J, Summons RE, Shen Y, Kaufman AJ, Masterson AL, Canfield

DE. 2008. Sulfur isotope biogeochemistry of the Proterozoic McArthur Basin. Geochimica

Et Cosmochimica Acta 72:4278-4290.

47. Shen YN, Farquhar J, Masterson A, Kaufman AJ, Buick R. 2009. Evaluating the role of

microbial sulfate reduction in the early Archean using quadruple isotope systematics. Earth

Planet Sci Lett 279:383-391.

48. Postgate JR. 1979. The sulphate-reducing bacteria. CUP Archive. 30

49. Chambers LA, Trudinger PA. 1979. Microbiological fractionation of stable sulfur isotopes

: A review and critique. Geomicrobiol J 1:249-293.

50. Bolliger C, Schroth MH, Bernasconi SM, Kleikemper J, Zeyer J. 2001. Sulfur isotope

fractionation during microbial sulfate reduction by toluene-degrading bacteria.

Geochimica Et Cosmochimica Acta 65:3289-3298.

51. Detmers J, Bruchert V, Habicht KS, Kuever J. 2001. Diversity of sulfur isotope

fractionations by sulfate-reducing prokaryotes. Appl Environ Microbiol 67:888-894.

52. Habicht KS, Canfield DE. 1997. Sulfur isotope fractionation during bacterial sulfate

reduction in organic-rich sediments. Geochimica Et Cosmochimica Acta 61:5351-5361.

53. Berner RA. 1984. Sedimentary Pyrite Formation-An update. Geochimica Et

Cosmochimica Acta 48:605-615.

54. Wilkin RT, Barnes HL. 1996. Pyrite formation by reactions of iron monosulfides with

dissolved inorganic and organic sulfur species. Geochimica Et Cosmochimica Acta

60:4167-4179.

55. Rickard D. 1997. Kinetics of pyrite formation by the H2S oxidation of iron (II) monosulfide

in aqueous solutions between 25 and 125 oC: The rate equation. Geochimica Et

Cosmochimica Acta 61:115-134.

56. Shen YN, Buick R. 2004. The antiquity of microbial sulfate reduction. Earth-Science

Reviews 64:243-272.

57. Schidlowski M, Hayes J, Kaplan I. 1983. Isotopic inferences of ancient biochemistries-

Carbon, sulfur, hydrogen, and nitrogen. in: Earth's earliest biosphere: Its origin and

evolution (J. W. Schopf, ed.). Princeton, NJ, Princeton University Press, p. 149-186.

58. Shen YA, Buick R, Canfield DE. 2001. Isotopic evidence for microbial sulphate reduction

in the early Archaean era. Nature 410:77-81.

59. Farquhar J, Bao HM, Thiemens M. 2000. Atmospheric influence of Earth's earliest sulfur

cycle. Science 289:756-758.

60. Muller E, Philippot P, Rollion-Bard C, Cartigny P. 2016. Multiple sulfur-isotope signatures

in Archean sulfates and their implications for the chemistry and dynamics of the early

atmosphere. Proc Natl Acad Sci U S A 113:7432-7437.

61. Philippot P, Van Zuilen M, Lepot K, Thomazo C, Farquhar J, Van Kranendonk A. 2007.

Elemental-sulfur reducing or disproportionating organisms in a similar to 3,5 Myr-old

seafloor setting. Geochimica Et Cosmochimica Acta 71:A786-A786.

62. Wacey D, McLoughlin N, Whitehouse MJ, Kilburn MR. 2010. Two coexisting sulfur

metabolisms in a ca. 3400 Ma sandstone. Geology 38:1115-1118.

63. Fischer WW, Fike DA, Johnson JE, Raub TD, Guan YB, Kirschvink JL, Eiler JM. 2014.

SQUID-SIMS is a useful approach to uncover primary signals in the Archean sulfur cycle.

Proc Natl Acad Sci U S A 111:5468-5473.

64. Thomassot E, O' Neil J, Francis D, Cartigny P, Wing BA. 2015. Atmospheric record in the

Hadean Eon from multiple sulfur isotope measurements in Nuvvuagittuq Greenstone Belt

(Nunavik, Quebec). Proc Natl Acad Sci U S A 112:707-712.

65. Johnston DT, Wing BA, Farquhar J, Kaufman AJ, Strauss H, Lyons TW, Kah LC, Canfield

DE. 2005. Active microbial sulfur disproportionation in the Mesoproterozoic. Science

310:1477-1479.

66. Johnston DT, Farquhar J, Wing BA, Kaufman A, Canfield DE, Habicht KS. 2005. Multiple

sulfur isotope fractionations in biological systems: A case study with sulfate reducers and

sulfur disproportionators. American Journal of Science 305:645-660.

67. Canfield DE, Teske A. 1996. Late Proterozoic rise in atmospheric oxygen concentration

inferred from phylogenetic and sulphur-isotope studies. Nature 382:127-132.

68. Habicht KS, Gade M, Thamdrup B, Berg P, Canfield DE. 2002. Calibration of sulfate

levels in the Archean ocean. Science 298:2372-2374.

69. Blank CE. 2009. Not so old Archaea: the antiquity of biogeochemical processes in the

archaeal domain of life. Geobiology 7:495-514.

70. Liu YC, Beer LL, Whitman WB. 2012. Methanogens: a window into ancient sulfur

metabolism. Trends Microbiol 20:251-258.

71. Sekowska A, Kung HF, Danchin A. 2000. Sulfur metabolism in Escherichia coli and

related bacteria: facts and fiction. J Mol Microbiol Biotechnol 2:145-77.

72. Takahashi H, Kopriva S, Giordano M, Saito K, Hell R. 2011. Sulfur assimilation in

photosynthetic organisms: Molecular functions and regulations of transporters and

assimilatory enzymes. Annu Rev Plant Biol, Vol 62 62:157-184.

73. Schelle MW, Bertozzi CR. 2006. Sulfate metabolism in mycobacteria. Chembiochem

7:1516-1524.

74. Kessler D. 2006. Enzymatic activation of sulfur for incorporation into biomolecules in

prokaryotes. FEMS Microbiol Rev 30:825-840.

75. Mueller EG. 2006. Trafficking in persulfides: delivering sulfur in biosynthetic pathways.

Nat Chem Biol 2:185-94.

76. Blank CE. 2009. Phylogenomic dating-the relative antiquity of archaeal metabolic and

physiological traits. Astrobiology 9:193-219.

77. Blank CE. 2004. Evolutionary timing of the origins of mesophilic sulphate reduction and

oxygenic photosynthesis: a phylogenomic dating approach. Geobiology 2:1-20.

78. Liu Y, Beer LL, Whitman WB. 2012. Sulfur metabolism in archaea reveals novel

processes. Environ Microbiol 14:2632-44.

79. Stetter KO, Gaag G. 1983. Reduction of molecular sulfur by methanogenic bacteria. Nature

305:309-311.

80. Daniels L, Belay N, Rajagopal BS. 1986. Assimilatory reduction of sulfate and sulfite by

methanogenic bacteria. Appl Environ Microbiol 51:703-709.

81. Rajagopal BS, Daniels L. 1986. Investigation of mercaptans, organic sulfides, and

inorganic sulfur-Compounds as sulfur sources for the growth of methanogenic bacteria.

Current Microbiology 14:137-144.

82. Lefaou A, Rajagopal BS, Daniels L, Fauque G. 1990. Thiosulfate, Polythionates and

elemental sulfur assimilation and reduction in the bacterial world. FEMS Microbiol Ecol

75:351-382.

83. Whitman W, and Jeanthon, C. (ed). 2006. Methanococcales. in the Prokaryotes (Dworkin,

M., Falkow, S., Rosenberg, E., Schleifer, K.-H., and Stackebrandt, E. (Eds).

84. Whitman WB, Ankwanda E, Wolfe RS. 1982. Nutrition and carbon metabolism of

Methanococcus voltae. J Bacteriol 149:852-63.

85. Whitman WB, Shieh J, Sohn S, Caras DS, Premachandran U. 1986. Isolation and

characterization of 22 mesophilic methanococci. Syst Appl Microbiol 7:235-240.

86. Rauch BJ, Perona JJ. 2016. Efficient sulfide assimilation in Methanosarcina acetivorans

is mediated by the MA1715 Protein. J Bacteriol 198:1974-1983.

87. Rauch BJ, Klimek J, David L, Perona JJ. 2017. Persulfide formation mediates cysteine and

homocysteine biosynthesis in Methanosarcina acetivorans. Biochemistry 56:1051-1061.

88. Hedderich R, Klimmek O, Kroger A, Dirmeier R, Keller M, Stetter KO. 1998. Anaerobic

respiration with elemental sulfur and with disulfides. FEMS Microbiol Rev 22:353-381.

89. Philippot P, Van Zuilen M, Lepot K, Thomazo C, Farquhar J, Van Kranendonk MJ. 2007.

Early Archaean microorganisms preferred elemental sulfur, not sulfate. Science 317:1534-

1537.

90. Belkin S, Wirsen CO, Jannasch HW. 1985. Biological and abiological sulfur reduction at

high-temperatures. Appl Environ Microbiol 49:1057-1061.

91. Moura JJG, Moura I, Santos H, Xavier AV, Scandellari M, Legall J. 1982. Isolation of

P590 from Methanosarcina barkeri:Evidence for the presence of sulfite reductase activity.

Biochem Biophys Res Commun 108:1002-1009.

92. Moura I, Lino AR, Moura JJG, Xavier AV, Fauque G, Peck HD, Legall J. 1986. Low-spin

sulfite reductases:A new homologous group of nonheme iron-siroheme proteins in a

naerobic bacteria. Biochem Biophys Res Commun 141:1032-1041.

93. Johnson EF, Mukhopadhyay B. 2008. Coenzyme F420-dependent sulfite reductase enabled

sulfite detoxification and use of sulfite as a sole sulfur source by Methanococcus

maripaludis. Environ Microbiol Rep 74:3591-3595.

94. Klipcan L, Frenkel-Morgenstern M, Safro MG. 2008. Presence of tRNA-dependent

pathways correlates with high cysteine content in methanogenic Archaea. Trends Genet

24:59-63. 35

95. Sauerwald A, Zhu WH, Major TA, Roy H, Palioura S, Jahn D, Whitman WB, Yates JR,

Ibba M, Soll D. 2005. RNA-dependent cysteine biosynthesis in archaea. Science 307:1969-

1972.

96. Liu YC, Nakamura A, Nakazawa Y, Asano N, Ford KA, Hohn MJ, Tanaka I, Yao M, Soll

D. 2014. Ancient translation factor is essential for tRNA-dependent cysteine biosynthesis

in methanogenic archaea. Proc Natl Acad Sci U S A 111:10520-10525.

97. Chen MR, Nakazawa Y, Kubo Y, Asano N, Kato K, Tanaka I, Yao M. 2016.

Crystallographic analysis of a subcomplex of the transsulfursome with tRNA for Cys-

tRNA(Cys) synthesis. Acta Crystallogr F Struct Biol Commun 72:569-572.

98. Hauenstein SI, Perona JJ. 2008. Redundant synthesis of cysteinyl-tRNA(Cys) in

Methanosarcina mazei. J Biol Chem 283:22007-22017.

99. O'Donoghue P, Sethi A, Woese CR, Luthey-Schulten ZA. 2005. The evolutionary history

of Cys-tRNACys formation. Proc Natl Acad Sci U S A 102:19003-19008.

100. Li T, Graham DE, Stathopoulos C, Haney PJ, Kim H, Vothknecht U, Kitabatake M, Hong

KW, Eggertsson G, Curnow AW, Lind W, Celic I, Whitman WB, Soll D. 1999. Cysteinyl-

tRNA formation: the last puzzle of aminoacyl-tRNA synthesis. FEBS Lett 462:302-306.

101. Newberry KJ, Hou YM, Perona JJ. 2002. Structural origins of amino acid selection without

editing by cysteinyl-tRNA synthetase. Embo J 21:2778-2787.

102. Zhang CM, Christian T, Newberry KJ, Perona JJ, Hou YM. 2003. Zinc-mediated amino

acid discrimination in cysteinyl-tRNA synthetase. J Mol Biol 327:911-917.

103. Mukai T, Crnkovic A, Umehara T, Ivanova NN, Kyrpides NC, Soll D. 2017. RNA-

dependent cysteine biosynthesis in Bacteria and Archaea. Mbio 8(3): e00561-17..

104. Farahi K, Pusch GD, Overbeek R, Whitman WB. 2004. Detection of lateral gene transfer

events in the prokaryotic tRNA synthetases by the ratios of evolutionary distances method.

J Mol Evol 58:615-631.

105. Helgadottir S, Rosas-Sandoval G, Soll D, Graham DE. 2007. Biosynthesis of

phosphoserine in the Methanococcales. J Bacteriol 189:575-582.

106. Liu Y, Dos Santos PC, Zhu X, Orlando R, Dean DR, Soll D, Yuan J. 2012. Catalytic

mechanism of Sep-tRNA:Cys-tRNA synthase: sulfur transfer is mediated by disulfide and

persulfide. J Biol Chem 287:5426-33.

107. Bult CJ, White O, Olsen GJ, Zhou L, Fleischmann RD, Sutton GG, Blake JA, FitzGerald

LM, Clayton RA, Gocayne JD, Kerlavage AR, Dougherty BA, Tomb JF, Adams MD,

Reich CI, Overbeek R, Kirkness EF, Weinstock KG, Merrick JM, Glodek A, Scott JL,

Geoghagen NS, Venter JC. 1996. Complete genome sequence of the methanogenic

archaeon, Methanococcus jannaschii. Science 273:1058-73.

108. Rauch BJ, Gustafson A, Perona JJ. 2014. Novel proteins for homocysteine biosynthesis in

anaerobic microorganisms. Mol Microbiol 94:1330-42.

109. Allen KD, Miller DV, Rauch BJ, Perona JJ, White RH. 2015. Homocysteine is

biosynthesized from aspartate semialdehyde and hydrogen sulfide in methanogenic

archaea. Biochemistry 54:3129-32.

110. Gophna U, Bapteste E, Doolittle WF, Biran D, Ron EZ. 2005. Evolutionary plasticity of

methionine biosynthesis. Gene 355:48-57.

111. Schroder I, Thauer RK. 1999. Methylcobalamin:homocysteine methyltransferase from

Methanobacterium thermoautotrophicum. Identification as the metE gene product. Eur J

Biochem 263:789-96. 37

112. Krautler B, Moll J, Thauer RK. 1987. The corrinoid from Methanobacterium-

Thermoautotrophicum (Marburg strain). Spectroscopic structure analysis and

identification as Co beta-cyano-5'-hydroxybenzimidazolyl-cobamide (Factor-III). Eur J

Biochem 162:275-278.

113. Johnson DC, Dean DR, Smith AD, Johnson MK. 2005. Structure, function, and formation

of biological iron-sulfur clusters. Annu Rev Biochem 74:247-281.

114. Jacobson MR, Brigle KE, Bennett LT, Setterquist RA, Wilson MS, Cash VL, Beynon J,

Newton WE, Dean DR. 1989. Physical and genetic map of the major nif gene gluster from

Azotobacter Vinelandii. J Bacteriol 171:1017-1027.

115. Jacobson MR, Cash VL, Weiss MC, Laird NF, Newton WE, Dean DR. 1989. Biochemical

and genetic analysis of the nifUSVWZM Cluster from Azotobacter Vinelandii. Mol Gen

Genet 219:49-57.

116. Zheng M, Wang X, Templeton LJ, Smulski DR, LaRossa RA, Storz G. 2001. DNA

microarray mediated transcriptional profiling of the Escherichia coli response to hydrogen

peroxide. J Bacteriol 183:4562-4570.

117. Takahashi Y, Tokumoto U. 2002. A third bacterial system for the assembly of iron-sulfur

clusters with homologs in archaea and plastids. J Biol Chem 277:28380-28383.

118. Outten FW, Djaman O, Storz G. 2004. A suf operon requirement for Fe-S cluster assembly

during iron starvation in Escherichia coli. Mol Microbiol 52:861-872.

119. Nachin L, Loiseau L, Expert D, Barras F. 2003. SufC: an unorthodox cytoplasmic

ABC/ATPase required for [Fe-S] biogenesis under oxidative stress. Embo Journal 22:427-

437.

120. Fontecave M, Ollagnier-De-Choudens S. 2008. Iron-sulfur cluster biosynthesis in bacteria:

Mechanisms of cluster assembly and transfer. Arch Biochem Biophys 474:226-237.

121. Balk J, Lobreaux S. 2005. Biogenesis of iron-sulfur proteins in plants. Trends Plant Sci

10:324-331.

122. Huet G, Daffe M, Saves I. 2005. Identification of the Mycobacterium tuberculosis SUF

machinery as the exclusive mycobacterial system of [Fe-S] cluster assembly: Evidence for

its implication in the pathogen's survival. J Bacteriol 187:6137-6146.

123. Boyd JM, Drevland RM, Downs DM, Graham DE. 2009. Archaeal ApbC/Nbp35 homologs

function as iron-sulfur cluster carrier proteins. J Bacteriol 191:1490-1497.

124. Björk G. R. (1995). “Biosynthesis and function of modified nucleosides” in tRNA:

structure, biosynthesis, and function eds. Soll D., RajBhandary U. L., editors. (Washington,

DC: ASM Press) 165-205.

125. Curran J. F. (1998). “Modified nucleosides in translation,” in Modification and Editing of

RNA eds.Grosjean H., Benne R., editors. (Washington, DC: ASM Press) 493-516.

126. Cavuzic M, Liu Y. 2017. Biosynthesis of sulfur-containing tRNA modifications: A

comparison of bacterial, archaeal, and eukaryotic pathways. Biomolecules 7.

127. Liu Y, Vinyard DJ, Reesbeck ME, Suzuki T, Manakongtreecheep K, Holland PL, Brudvig

GW, Soll D. 2016. A [3Fe-4S] cluster is required for tRNA thiolation in archaea and

eukaryotes. Proc Natl Acad Sci U S A doi:10.1073/pnas.1615732113.

128. Mihara H, Esaki N. 2002. Bacterial cysteine desulfurases: their function and mechanisms.

Appl Microbiol Biotechnol 60:12-23.

129. Pandey A, Golla R, Yoon H, Dancis A, Pain D. 2012. Persulfide formation on

mitochondrial cysteine desulfurase: enzyme activation by a eukaryote-specific interacting

protein and Fe-S cluster synthesis. Biochem J 448:171-87.

130. Kessler D. 2006. Enzymatic activation of sulfur for incorporation into biomolecules in

prokaryotes. FEMS Microbiol Rev 30:825-40.

131. Black KA, Dos Santos PC. 2015. Shared-intermediates in the biosynthesis of thio-

cofactors: Mechanism and functions of cysteine desulfurases and sulfur acceptors. Biochim

Biophys Acta 1853:1470-80.

132. Bjork GR. 1995. Genetic dissection of synthesis and function of modified nucleosides in

bacterial transfer RNA. Prog Nucleic Acid Res Mol Biol 50:263-338.

133. Kotera M, Bayashi T, Hattori M, Tokimatsu T, Goto S, Mihara H, Kanehisa M. 2010.

Comprehensive genomic analysis of sulfur-relay pathway genes. Genome Inform 24:104-

15.

134. Favre A, Yaniv M, Michelson AM. 1969. The photochemistry of 4-thiouridine in

Val Escherichia coli t-RNA1 . Biochem Biophys Res Commun 37:266-71.

135. Favre A, Michelson AM, Yaniv M. 1971. Photochemistry of 4-thiouridine in Escherichia

Val coli transfer RNA1 . J Mol Biol 58:367-79.

136. Thomas G, Favre A. 1975. 4-Thiouridine as the target for near-ultraviolet light induced

growth delay in Escherichia coli. Biochem Biophys Res Commun 66:1454-61.

137. Durant PC, Bajji AC, Sundaram M, Kumar RK, Davis DR. 2005. Structural effects of

hypermodified nucleosides in the Escherichia coli and human tRNALys anticodon loop: the

effect of nucleosides s2U, mcm5U, mcm5s2U, mnm5s2U, t6A, and ms2t6A. Biochemistry

44:8078-89. 40

138. Bjork GR, Hagervall TG. 2014. Transfer RNA modification: presence, synthesis, and

function. EcoSal Plus 6 (1).

139. Shigi N. 2014. Biosynthesis and functions of sulfur modifications in tRNA. Front Genet

5:67.

140. Alkatib S, Scharff LB, Rogalski M, Fleischmann TT, Matthes A, Seeger S, Schottler MA,

Ruf S, Bock R. 2012. The contributions of wobbling and superwobbling to the reading of

the genetic code. PLoS Genet 8:e1003076.

141. Grosjean H, Westhof E. 2016. An integrated, structure- and energy-based view of the

genetic code. Nucleic Acids Res 44:8020-40.

142. Rodriguez-Hernandez A, Spears JL, Gaston KW, Limbach PA, Gamper H, Hou YM,

Kaiser R, Agris PF, Perona JJ. 2013. Structural and mechanistic basis for enhanced

translational efficiency by 2-thiouridine at the tRNA anticodon wobble position. J Mol Biol

425:3888-906.

143. Sylvers LA, Rogers KC, Shimizu M, Ohtsuka E, Soll D. 1993. A 2-thiouridine derivative

in tRNAGlu is a positive determinant for aminoacylation by Escherichia coli glutamyl-

tRNA synthetase. Biochemistry 32:3836-41.

144. Madore E, Florentz C, Giege R, Sekine S, Yokoyama S, Lapointe J. 1999. Effect of

modified nucleotides on Escherichia coli tRNAGlu structure and on its aminoacylation by

glutamyl-tRNA synthetase. Predominant and distinct roles of the mnm5 and s2

modifications of U34. Eur J Biochem 266:1128-35.

145. Urbonavičius J, Qian Q, Durand JM, Hagervall TG, Björk GR. 2001. Improvement of

reading frame maintenance is a common function for several tRNA modifications. EMBO

J 20:4863-4873. 41

146. El Yacoubi B, Bailly M, de Crecy-Lagard V. 2012. Biosynthesis and function of

posttranscriptional modifications of transfer RNAs. Annu Rev Genet 46:69-95.

147. Leidel S, Pedrioli PG, Bucher T, Brost R, Costanzo M, Schmidt A, Aebersold R, Boone C,

Hofmann K, Peter M. 2009. Ubiquitin-related modifier Urm1 acts as a sulphur carrier in

thiolation of eukaryotic transfer RNA. Nature 458:228.

148. Noma A, Sakaguchi Y, Suzuki T. 2009. Mechanistic characterization of the sulfur-relay

system for eukaryotic 2-thiouridine biogenesis at tRNA wobble positions. Nucleic Acids

Res 37:1335-52.

149. Dewez M, Bauer F, Dieu M, Raes M, Vandenhaute J, Hermand D. 2008. The conserved

Wobble uridine tRNA thiolase Ctu1-Ctu2 is required to maintain genome integrity. Proc

Natl Acad Sci U S A 105:5459-64.

150. Schlieker CD, Van der Veen AG, Damon JR, Spooner E, Ploegh HL. 2008. A functional

proteomics approach links the ubiquitin-related modifier Urm1 to a tRNA modification

pathway. Proc Natl Acad Sci U S A 105:18255-60.

151. Liu Y, Zhu X, Nakamura A, Orlando R, Soll D, Whitman WB. 2012. Biosynthesis of 4-

thiouridine in tRNA in the methanogenic archaeon Methanococcus maripaludis. J Biol

Chem 287:36683-92.

152. Chavarria NE, Hwang S, Cao S, Fu X, Holman M, Elbanna D, Rodriguez S, Arrington D,

Englert M, Uthandi S. 2014. Archaeal Tuc1/Ncs6 homolog required for wobble uridine

tRNA thiolation is associated with ubiquitin-proteasome, translation, and RNA processing

system homologs. PloS one 9:e99104.

153. Humbard MA, Miranda HV, Lim JM, Krause DJ, Pritz JR, Zhou G, Chen S, Wells L,

Maupin-Furlow JA. 2010. Ubiquitin-like small archaeal modifier proteins (SAMPs) in

Haloferax volcanii. Nature 463:54-60.

154. Liu Y, Long F, Wang L, Soll D, Whitman WB. 2014. The putative tRNA 2-thiouridine

synthetase Ncs6 is an essential sulfur carrier in Methanococcus maripaludis. FEBS Lett

588:873-7.

155. Hepowit NL, de Vera IM, Cao S, Fu X, Wu Y, Uthandi S, Chavarria NE, Englert M, Su D,

Sll D, Kojetin DJ, Maupin-Furlow JA. 2016. Mechanistic insight into protein modification

and sulfur mobilization activities of noncanonical E1 and associated ubiquitin-like proteins

of Archaea. FEBS J 283:3567-3586.

156. Miranda HV, Nembhard N, Su D, Hepowit N, Krause DJ, Pritz JR, Phillips C, Soll D,

Maupin-Furlow JA. 2011. E1- and ubiquitin-like proteins provide a direct link between

protein conjugation and sulfur transfer in archaea. Proc Natl Acad Sci U S A 108:4417-22.

Figure 1-1. Schematic model of a putative Archean sulfur cycle, involved with volcanic activity and related environments of barite deposition. Modified from (22, 60). MIF: mass-independent fractionation.

Figure 1-2. Sulfur assimilation and trafficking pathways in methanococci.

Known and proposed reactions are depicted in black. These include (i) biosynthesis of homocysteine from asparatate semialdehyde and hydrogen sulfide, involving enzyme aspartate kinase, aspartate semialdehyde dehydrogenase (ASD), and homoserine dehydrogenase (HSD)

(108, 109). (ii) cysteine biosynthetic pathways, primarily a tRNA-dependent pathway, using O- phosphoseryl-tRNA synthetase(SepRS) and Sep-tRNA:Cys-tRNA synthase (SepCysS) (95). In addition, certain methanoccocus species also possess either the bacterial or the eukaryotic enzymes for cysteine biosynthesis, they are O-acetyltransferase (SAT) and O-acetylserine sulfhydrylase

(OASS). (iii) methionine biosynthesis from homocysteine using the cobalamin-independent methionine synthase (MetE) (111).

Enzyme homologs, reactions, and pathways that are absent in methanococci are colored in blue.

These are (i) homocysteine production with hydrogen sulfide and O-phosphohomoserine, O- acetyl-homoserine or O-succinyl-homoserine by direct sulfhydrylation of homoserine derivatives, using enzymes homoserine succinyltransferase (HSST), homoserine kinase (HSK), homoserine 45

acetyltransferase (HSAT), O-succinylhomoserine sulfhydrylase (OSHS), cystathionine gamma- synthase (CGS), O-acetylhomoserine sulfhydrylase (OAHS) (10). (ii) Homocysteine formation from cystathionine, with the sulfur donated from cysteine, using enzymes cystathionine gamma- synthase (CGS) and cystathionine gamma-lyase (CGL) (109). (iii) Cysteine desulfurase (CD) decomposes cysteine to alanine and sulfane sulfur via the formation of a protein-bound persulfide intermediate ([Protein]‒S-SH), which then functions as the sulfur donor for the biosynthesis of iron-sulfur containing cofactors, tRNA thiolation (tRNA thiol) and small archaeal modifier protein thiolated-tRNAs (SAMP).

Unestablished reactions and pathways are indicated in dash line. These are (i) The biodegradation of methionine to homocysteine, by forming S-adenosylmethionine (Adomet) and S- adenosylhomocysteine (AdoHcy) as intermediates. Enzymes involved in this pathway are S- adenosylmethionine synthetase (SAMS), AdoMet-dependent methyltransferases (MTase), and S- adenosylhomocysteine hydrolase (SAHH). (ii) The enzyme(s) and step (s) that decomposes cystathionine to cysteine.

Unknown substrates in reactions are colored in red. These include (i) The sulfur donor (R-S-SH) to charge Sep-tRNACys to produce Cys-tRNACys by SepCysS. (ii) The methyl donor to form methionine from homocysteine.

Other abbreviations of enzymes are cysteine-tRNA synthetase (CysRS), cobalamin-dependent methionine synthase (MetH).

Figure 1-3. Sulfur assimilation and trafficking scheme in E. coli and related bacteria. Modified from (70).

Figure 1-4. A putative scheme of the sulfur assimilation and trafficking in methanococci. Modified from (70). Methanococci use sulfide or elemental sulfur as sole sulfur source for growth. For homocysteine biosynthesis, sulfide can be used as the sulfur donor for aspartate (Asa) semialdehyde to form homocysteine (109). Other homocysteine biosynthetic pathways, including the direct sulfhydrylation of homoserine derivatives using sulfide, and the β-cleavage pathway of cystathionine pathway is absent in methanococci (10). For methionine biosynthesis, homocystine is found to be an intermediate. A cobalamin-independent methionine synthase is conserved in methanogens. For cysteine biosynthesis, a persulfide (with one sulfane sulfur) group on Sep- tRNA:Cys-tRNA synthase (SepCysS) is proposed to be the proximal sulfur donor for the conversion of Sep-tRNACys to Cys-tRNACys (98, 106). The process of sulfane sulfur formation from sulfide and sulfane sulfur delivery to SepCysS are not known. For biosynthesis of iron-sulfur clusters and tRNA thiolation, cysteine is not the sulfur donor for iron-sulfur cluster biosynthesis

(10). Sulfide is proposed to be the physiological sulfur donor for the production of 4-thiouridine by ThiI (151). 48

Table 1-1. Some examples of important sulfur transformations. Modified from (11, 14)

Sulfate reduction 2- - SO4 + 2CH2O  H2S + 2HCO3

Sulfite Reduction 2- + - 2SO3 + 3CH2O + H  2H2S + 3HCO3

Sulfur disproportionation 0 2- + (i) 4S + 4H2O  3 H2O + SO4 + 2H + 2+ 0 (ii) H2S + 4H + 2FeOOH  2Fe + S + 4H2O 2+ + (iii) 2H2S + 2Fe  2FeS + 4H 0 2- + (iv) 3S + 2FeOOH  SO4 + 2FeS + 2H 0 2- 2+ - (v) S + 3MnO2 +2H2O  SO4 + 3Mn + 4OH

Thiosulfate disproportionation 2- 2- S2O3 + H2O  H2S + SO4

Phototrophic sulfide oxidation 0 H2S + CO2  CH2O + H2O+ 2S 2- + H2S + 2CO2 + H2O  2CH2O + SO4 + 2H

Phototrophic sulfur oxidation 0 2- + 2S + 3CO2 + 5H2O  3CH2O + 2SO4 + 4H

CHAPTER 2

A FLEXIBLE SYSTEM FOR CULTIVATION OF METHANOCOCCUS AND OTHER

FORMATE-UTILIZING METHANOGENS1

1 Long, F., L.L. Wang, B. Lupa, and W.B. Whitman, 2017. A flexible system for cultivation of

Methanococcus and other formate-utilizing methanogens. Archaea. 2017:12. doi:10.1155/2017/7046026. Reprinted here with permission of the publisher. 50

Abstract

Many hydrogenotrophic methanogens use either H2 or formate as the major electron donor to reduce CO2 for methane production. The conventional cultivation of these organisms uses H2 and

CO2 as the substrate with frequent replenishment of gas during growth. H2 is explosive and requires an expensive gassing system to handle safely. Formate is as an ideal alternative substrate from the standpoints of both economy and safety but leads to large changes in the culture pH during growth.

Here we report that glycylglycine is an inexpensive and nontoxic buffer suitable for growth of

Methanococcus maripaludis and Methanothermococcus okinawensis. This cultivation system is suitable for growth on liquid as well as solid medium in serum bottles. Moreover, it allows cultivation of liter scale cultures without expensive fermentation equipment. This formate cultivation system provides an inexpensive and flexible alternative for the growth of formate- utilizing, hydrogenotrophic methanogens.

Introduction

Methanogens are strictly anaerobic microorganisms belonging to the Euryarchaeota. As a large and diverse group, they are distinguished by their capability to obtain most if not all of their energy for growth from methane production or methanogenesis (1). In general, methanogens only utilize a limited number of substrates for methanogenesis, such as CO2; H2; formate; methyl-group containing compounds such as methylamines, methylsulfides, and methanol; acetate, and a few low molecular weight alcohols. They do not use sugars, amino acids, or most other common organic substrates (2). Most methanogens are hydrogenotrophs that use H2 as the primary electron donor to reduce CO2 to methane. Many hydrogenotrophic methanogens can also use formate as the major electron donor (2). As shown in equation 1, four molecules of sodium formate are oxidized, yielding one molecule of methane and three molecules of CO2.

4HCOONa+2H2OCH4+3CO2+4NaOH (Equation 1)

Because no more than one ATP is formed per mol of CH4 (3), relatively large amounts of formate are required for even modest growth. Growing cells with sodium formate also leads to a significant accumulation of NaOH, which raises the pH of the medium and inhibits growth. For methanococci, the alkaline pH also causes cell lysis and rapid killing (4). As a result, pH control becomes a critical concern when cultivating methanogens with sodium formate.

One solution is to titrate the rise in pH with formic acid during growth in a fermenter (5).

For growth in culture tubes and plates, the medium pH can also be controlled with a built-in formic acid reservoir (4). This cultivation system uses a 6×55-mm acid reservoir containing 200 µL formic acid to stabilize the medium pH (4). As the pH increases, the absorption of formic acid from the headspace also increases, maintaining the pH within levels that support growth. Although

this method allows good growth on formate-containing medium, its requirement for manual dexterity precludes it from routine use. Using formate as substrate has also been established in a chemostat system. Costa et al. (6) used formate to grow M. maripaludis in chemostat for studying the transcriptional regulation. The sodium formate was added at 0.38 M, while the pH was maintained at 6.95 by automatic addition of 10 % (v/v) H2SO4 (6). The cell density and growth rate achieved with either formate or H2/CO2 were the same during chemostat cultivation (6-9).

During growth of M. maripaludis with formate, formate dehydrogenase (Fdh) is the key enzyme for formate utilization. Fdh is encoded by two sets of genes, fdhA1B1 and fdhA2B2 in M. maripaludis (10). Lupa et al. (11) found that mutants with deletions in fdhA1 grew poorly on formate only after an extended lag. In contrast, mutants with deletions in fdhA2 grew nearly the same as wild-type. Because of this and other evidence, Fdh1 was proposed to play a major role in fomate utilization (11).

Over the past decade, many genetic methodologies have been developed in M. maripaludis.

These include: effective selectable genetic markers (12-16), multiple plasmid shuttle vectors (17), high efficiency transformation (18), direct gene replacement mutagenesis (19), markerless gene deletion systems (20), random mutagenesis (21), in vivo transposon mutagenesis (22, 23), reporter genes technologies (24, 25) and chemostat cultivation (6-9). Thus, genetic manipulation of M. maripaludis is easy and effective, and these approaches have become powerful tools to study the metabolism and physiology of multiple Methanococcus species. However, the requirement for H2 growth limits the ability of these genetic tools to be widely applied in laboratories that do not have established systems for handling H2 gas.

Here we report a medium to cultivate the mesophilic, marine species M. maripaludis on formate using glycylglycine buffer as the pH stabilizer. Ordinarily, M. maripaludis is cultivated in aluminum sealed tubes with 5 mL of medium under H2/CO2 mixture (4:1, v/v) at 276 kPa (26).

For comparison, in our formate cultivation system, the pressure is reduced to 104 kPa, allowing use of more inexpensive stoppers. In addition, frequent gas refilling is avoided without greatly sacrificing growth yield. Simple modifications of common glassware also allow liter-scale cultivation using only a gassing station and vacuum pump. In addition, the solid medium has a high plating efficiency suitable for genetic experiments. With minor adjustment in medium composition, the procedure is also suitable for growth of the extreme thermophile

Methanothermococcus okinawensis.

Materials and methods

Strains, media and growth conditions. Methanococcus maripaludis strain S2 was obtained from our laboratory collection (Whitman et al., 1986) (27) and cultured at 37 oC.

Methanothermococcus okinawensis strain IH1 was obtained from Miyazaki and Takai and cultured at 62 oC (28).

Cultures were grown in H2/CO2 medium (McNA, a minimal medium with 10 mM sodium acetate) or formate medium (McF) reduced with 3 mM cysteine hydrochloride. The 5 mL cultures were grown in 28 mL aluminum sealed tubes. For McNA, the tubes were pressurized to 276 kPa with H2/CO2 (4:1, v/v) and refilled with the same gas every 24 hours after inoculation. Detailed protocols for growth on formate are given below (Supplementary A, section S2A). Briefly, McF medium contained 0.4 M sodium formate and was buffered with 0.2 M glycylglycine (pH= 8.0).

Medium was first sparged with N2 to remove most of the O2, 3 mM cysteine chloride was then

added. Tubes were pressurized to 103 kPa with N2/CO2 (4:1, v/v) before autoclaving. Prior to inoculation, 3 mM sodium sulfide was added as the sulfur source.

The buffers tested were obtained from Sigma Chemical Co. and included (with the counter ion): Tricine/NaOH (N-[Tris(hydroxymethyl)methyl]glycine), Bicine/NaOH ( N,N-Bis(2- hydroxyethyl)glycine), Tris/HCl (2-Amino-2-hydroxymethyl-propane-1,3-diol), glycine/NaOH and glycylglycine/NaOH. During formate medium preparation, ingredients were added as listed in the appendices, and the organic buffers were added from stock solutions at pH 7. The concentration of NaCl was adjusted depending upon the amount of sodium formate and sodium in the buffer used so that the final concentration of sodium ion was 0.4 M.

The final medium was also tested for plating (Supplementary A, section S2B) and growth of 1.5 L cultures (Supplementary A, section S2C).

Rapid protocol for preparation of formate medium. After combining the components of McF medium, cysteine was added, and the medium was dispensed into culture tubes on the bench without anaerobic precautions (Supplementary A, section S2D). Without delay, the tubes were sealed with stoppers and aluminum seals. The tubes were then connected to a gassing manifold, and the air was removed by three successive cycles comprising 45 seconds of vacuum followed by 15 seconds of 104 kPa N2 : CO2 (4:1, v/v). After exchanging the gas, the medium was autoclaved for 20 min with rapid exhaust. For the control medium, the medium was dispensed in an anaerobic chamber as described in Supplementary A, section S2A, and the gas was exchanged for three cycles with N2/CO2 (4:1, v/v) prior to autoclaving.

Results

Optimization of the formate medium and growth conditions. To determine if organic buffers were inhibitory for growth, they were added to medium during growth of M. maripaludis on H2/CO2. Because the medium was strongly buffered with bicarbonate and CO2, the buffers did not affect the initial pH. Under these conditions, Tricine was strongly inhibitory (Figure 2-1).

While glycine and Bicine had little effect on cell yield, both increased the lag phase at higher concentrations (data not shown). In contrast, Tris and glycylglycine were not inhibitory and resulted in moderate decreases in the lag phase, presumably by maintaining an optimal pH during the early growth phase (data not shown). Therefore, Tricine and Bicine were omitted from further experiments.

Tris, glycine, and glycylglycine were further tested for their buffering capacity during growth with 200 mM sodium formate. In the presence of 100 mM buffer, the culture reached a maximal absorbance of about 0.4-0.45 after 20 h (Figure 2-2). During the first two days of incubation at 37 oC, all three buffers maintained the medium pH around 7.2-7.6. However, during extended incubations, decreased absorbance and cellular lysis were observed in media buffered with Tris and glycine (Figure 2-2 and data not shown). In contrast, the absorbance of cultures supplemented with glycylglycine remained stable for six days at 37 oC (Figure 2-2). Moreover, in glycylgylcine buffered medium, the culture absorbance did not change for up to six weeks at room temperature, and it was still possible to transfer stock cultures to fresh medium. Cultures in McF medium were also used to prepare -80 oC freezer stocks in 30 % (v/v) glycerol (26, 29), and these cultures retained viability for at least five years.

To reduce the cost of anaerobic medium preparation, the influence of different types of stoppers on growth was also tested. Cultivation on H2/CO2 is usually performed at 276 kPa in 28 mL aluminum sealed tubes. For this reason, thick butyl rubber stoppers (Bellco Glass, Inc.,

Vineland, NJ, cat no. 2048-11800) are commonly used. These stoppers are made to minimize gas leakage and sustain multiple needle stabs during medium preparation, inoculation, and sampling.

As an alternative, butyl rubber grey stoppers (Wheaton Science Products, cat. no.: W224100-202) are much less expensive although thinner. Although these stoppers cannot maintain high pressure, they might be suitable for growth on formate at lower pressure. As shown in Figure 2-2, Wheaton stopper-sealed cultures showed comparable growth profiles and stability, especially in medium supplemented with glycylglycine. In contrast, white precipitates were observed in cultures supplemented with Tris and glycine (data not shown). The composition of the medium resembles that of seawater and contains high levels of divalent cations. During autoclaving, the pH of this medium increases due to the reduced solubility of CO2 at high temperatures. Presumably, these precipitates represent phosphate salts that become insoluble at alkaline pHs. The precipitates were rarely observed following autoclaving with the thicker stoppers, probably because they retained

CO2 better during autoclaving.

In the presence of 100 mM glycylglycine, the growth yields increased with formate concentrations in non-linear fashion and were maximal at 0.6 M. Growth was inhibited with 1 M sodium formate, presumably due to sodium toxicity (data not shown). At high formate concentrations and 100 mM glycylglycine, cells lysed in the stationary phase, presumably due to alkalinization of the medium. Increasing the glyclyglcine concentration to 200 mM with 0.4 M formate was found to be optimal for batch growth. In this condition, the growth rate was similar

to that in H2/CO2 medium. Moreover, the maximum OD600nm of 1.0 was comparable to 1.4 in

H2/CO2 medium (Figure 2-3). Thus, the cellular yields per mole of electron of donor were nearly equivalent. For instance, medium with 0.4 M formate contained about 2 mmol of formate in 5 mL, and the growth yield was about 340 mg dry wt L-1 or 0.85 g dry wt mol-1 of formate. For 5 mL

-1 H2/CO2 cultures with 2.7 mmol of H2, the growth yield was about 400 mg dry wt L or 0.74 g dry

-1 wt mol of H2.

Good growth was also found on formate medium containing 1.0 % (w/v) agar in serum bottles. Details on preparation are given in Supplementary A, section S2B, but it is similar to the protocols described earlier (30, 31). Similar to growth with H2/CO2 medium, isolated colonies appeared after 3 to 5 days of incubation, and the plating efficiency was 100 %.

A simple medium-scale cultivation system for M. maripaludis and M. okinawensis with formate. The modified formate medium was also useful for cultivation of M. maripaludis and M. okinawensis at the liter or medium-scale for the preparation of biomass for enzyme and other studies. For this purpose, a simple cultivation system was developed using common lab glassware and equipment (Supplementary A, section S2C). Comprised largely of a 2 L cultivation bottle, water trap and a gas trap, each assembly supported growth of 1.5 L of culture. During growth, the exhaust line allowed the CH4 and CO2 formed to escape, the water trap prevented backflow of water into the culture, and the gas trap prevented back diffusion of air into the culture bottle. A protocol was also developed to ensure complete reduction of the medium before inoculation

(Supplementary A, section S2C). Although the medium was sparged prior to inoculation, after inoculation no gassing was required, and the system could be easily moved to fume hood, incubator or some other well-ventilated space. With a 2 % (v/v) inoculum, M. maripaludis S2 grew to about

o OD600nm= 0.8 after 15 hours of incubation at 37 C. In the same medium, M. okinawensis IH1 grew to an OD600nm= 0.6 (Figure 2-4). However, reduction of the pH of the glycylglycine buffer stock solution to 6.5, reduced the lag phase of M. okinawensis to 12 h at 62 oC without reducing the yield

(data not shown). For both cultures, the cell yield was around 1 g (wet weight) per L-1.

Rapid preparation of medium without an anaerobic chamber. An anaerobic chamber is often used for preparing medium for methanogens, but it is expensive and occupies a large amount of laboratory space. To determine if the formate medium could be prepared in laboratories with limited anaerobic equipment, it was prepared aerobically, and the gas was exchanged with a vacuum pump and gas line connected to a simple gassing manifold controlled by a three-way ball valve. The system was constructed from standard compression fittings so that its fabrication required little equipment and no special expertise. It was designed so that ten tubes or serum bottles could be prepared at one time. A vacuum pressure gauge was used to monitor the gas. After dispensing the medium aerobically, gassing/vacuum cycles were performed to remove O2 from the medium (Supplementary A, section S2D). Interestingly, growth in medium with even one gassing/vacuum cycle was nearly the same as in conventionally prepared medium (data not shown). Cultures of M. maripaludis are often tolerant to O2, and growth of log phase cultures are unaffected by O2 partial pressures < 20 kPa (unpublished observation). Therefore, it was possible that the large size of inoculum may have protected cells from residual O2. To examine the suitability of this method for small inocula, a most probable number (MPN) experiment was performed in medium prepared with three cycles of gas exchange (Table 2-1). The most probable numbers were 50 and 160 in media prepared by the rapid or standard protocol with an anaerobic

chamber, respectively (32). These high numbers were not significantly different and would only be possible if growth could be initiated by only one or two cells in both media.

This protocol was also suitable for preparation of solid medium and plating for isolation of mutants or other clonal cultures (Supplementary A, section S2D). Agar slabs were formed in serum bottles as described in Supplementary A, section S2B. After growth, single colonies were picked with a syringe needle and transferred to broth under a stream of N2 gas.

Discussion

The medium and culturing system for methanogens developed here attempted to address multiple concerns. First, the reagents and equipment should be accessible to many research laboratories. The replacement of H2 with formate as the major substrate for methanogenesis removed the need for a H2 handling system, reducing the cost as well as increasing the safety of culturing. The cost of medium preparation can be further reduced by using much less expensive septum stoppers. Moreover, a simple gassing manifold was sufficient, and an anaerobic chamber was not needed. These methods are straightforward and do not require extensive training. At the

University of Georgia, this culturing system was widely used by undergraduate students to isolate and cultivate mutants of M. maripaludis. While training is still required, especially for the safe use of syringes and pressurized glassware, many of the elaborate manipulations of the Hungate method

(33) are avoided. For many biological investigations, it is often necessary to generate cultures from single cells as well as generate large amounts of biomass. Both of these are often difficult with fastidious anaerobes. The system developed here had a high plating efficiency, and it was possible to develop cultures from only a few cells. Therefore, it is suitable for the isolation of mutants or other genetic experiments. In addition, it was possible to generate sufficient biomass for enzymatic

assays and other biochemical analyses. The glycylglycine buffer prevented alkalinization of the medium and allowed the cultures to remain viable for several weeks on the bench. The addition of glycerol allowed maintenance of viable cultures for at least five years at -80 oC. Nevertheless, the formate medium allows a similar growth rate and cellular yield as H2/CO2 medium. Moreover, this medium and protocol were adapted by Weimara et al. (34) for a multi-well plate method to screen chemical compound libraries (34). M. maripaludis was grown in 96-well microtiter plates sealed in an AGS AnaeroGen compact bag (Oxoid) and incubated at 37 oC inside an anaerobic

chamber containing 5 % H2, 5 % CO2 and 90 % N2 (34). Therefore, these methods can be readily adapted for a number of experimental approaches.

References:

1. Hedderich R, Whitman WB. 2006. Physiology and biochemistry of the methane-producing

Archaea. Prokaryotes: A Handbook on the Biology of Bacteria, Vol 2, Third Edition

doi:10.1007/0-387-30742-7_34:1050-1079.

2. Liu YC, Whitman WB. 2008. Metabolic, phylogenetic, and ecological diversity of the

methanogenic archaea. Incredible Anaerobes: From Physiology to Genomics to Fuels

1125:171-189.

3. Thauer RK, Jungermann K, Decker K. 1977. Energy conservation in chemotrophic

anaerobic bacteria. Bacteriol Rev 41:100-80.

4. Schauer NL, Whitman WB. 1989. Formate growth and pH control by volatile formic and

acetic acids in batch cultures of methanococci. J Microbiol Methods 10:1-7.

5. Schauer NL, Ferry JG. 1980. Metabolism of formate in Methanobacterium formicicum. J

Bacteriol 142:800-807.

6. Costa KC, Yoon SH, Pan M, Burn JA, Baliga NS, Leigh JA. 2013. E Effects of H2 and

formate on growth yield and regulation of methanogenesis in Methanococcus maripaludis.

J Bacteriol 195:1456-1462.

7. Haydock AK, Porat I, Whitman WB, Leigh JA. 2004. Continuous culture of

Methanococcus maripaludis under defined nutrient conditions. FEMS Microbiol Lett

238:85-91.

8. Hendrickson EL, Haydock AK, Moore BC, Whitman WB, Leigh JA. 2007. Functionally

distinct genes regulated by hydrogen limitation and growth rate in methanogenic Archaea.

Proc Natl Acad Sci, U S A 104:8930-8934.

9. Leigh JA. 2011. G Growth of methanogens under defined hydrogen conditions. Methods

Enzymol 494:111-118.

10. Wood GE, Haydock AK, Leigh JA. 2003. Function and regulation of the formate

dehydrogenase genes of the methanogenic archaeon Methanococcus maripaludis. J

Bacteriol 185:2548-54.

11. Lupa B, Hendrickson EL, Leigh JA, Whitman WB. 2008. Formate-Dependent H2

production by the mesophilic methanogen Methanococcus maripaludis. Appl Environ

Microbiol 74:6584-6590.

12. Beckler GS, Hook LA, Reeve JN. 1984. C Chloramphenicol acetyltransferase should not

provide methanogens with resistance to chloramphenicol. Appl Environ Microbiol 47:868-

869.

13. Böck A, Kandler O. 1985. Antibiotic sensitivity of Archaebacteria. The Bacteria 8:525-

544

14. Gernhardt P, Possot O, Foglino M, Sibold L, Klein A. 1990. Construction of an integration

vector for use in the archaebacterium Methanococcus voltae and expression of a eubacterial

resistance gene. MOL GEN GENET 221:273-279.

15. Weisburg WG, Tanner RS. 1982. Aminoglycoside sensitivity of archaebacteria. FEMS

Microbiol Lett 14:307-310.

16. Franke CA, Rice CM, Strauss JH, Hruby DE. 1985. Neomycin resistance as a dominant

selectable marker for selection and isolation of vaccinia virus recombinants. Molecular and

Cellular Biology 5:1918-1924.

17. Gardner WL, Whitman WB. 1999. Expression vectors for Methanococcus maripaludis:

Overexpression of acetohydroxyacid synthase and beta-galactosidase. Genetics 152:1439-

1447.

18. Tumbula DL, Makula RA, Whitman WB. 1994. Transformation of Methanococcus

maripaludis and identification of a PstI-like restriction system. FEMS Microbiol Lett

121:309-314.

19. Kim JE, Kim KH, Lee SW, Seol W, Shiba K, Kim S. 2000. An elongation factor-

associating domain is inserted into human cysteinyl-tRNA synthetase by alternative

splicing. Nucleic Acids Res 28:2866-2872.

20. Moore BC, Leigh JA. 2005. Markerless mutagenesis in Methanococcus maripaludis

demonstrates roles for alanine dehydrogenase, alanine racemase, and alanine permease. J

Bacteriol 187:972-979.

21. Ladapo J, Whitman WB. 1990. Method for isolation of auxotrophs in the methanogenic

archaebacteria: Role of the acetyl-CoA pathway of autotrophic CO2 fixation in

Methanococcus maripaludis. Proc Natl Acad Sci, U S A 87:5598-5602. 63

22. Porat I, Whitman WB. 2009. Tryptophan auxotrophs were obtained by random transposon

insertions in the Methanococcus maripaludis tryptophan operon. FEMS Microbiol Lett

297:250-254.

23. Sarmiento F, Mrazek J, Whitman WB. 2013. Genome-scale analysis of gene function in

the hydrogenotrophic methanogenic archaeon Methanococcus maripaludis. Proc Natl

Acad Sci, U S A 110:4726-4731.

24. Lie TJ, Leigh JA. 2002. Regulatory response of Methanococcus maripaludis to alanine, an

intermediate nitrogen source. J Bacteriol 184:5301-5306.

25. Lie TJ, Leigh JA. 2007. Genetic screen for regulatory mutations in Methanococcus

maripaludis and its use in identification of induction-deficient mutants of the eulyarchaeal

repressor NrpR. Appl Environ Microbiol 73:6595-6600.

26. Sarmiento BF, Leigh JA, Whitman WB. 2011. Genetic Systems for hydrogenotrophic

methanogens. Methods Enzymol 494:43-73.

27. Whitman WB, Shieh J, Sohn S, Caras DS, Premachandran U. 1986. Isolation and

characterization of 22 mesophilic methanococci. Syst Appl Microbiol 7:235-240.

28. Takai K, Inoue A, Horikoshi K. 2002. Methanothermococcus okinawensis sp. nov., a

thermophilic, methane-producing archaeon isolated from a Western Pacific deep-sea

hydrothermal vent system. Int J Syst Evol Microbiol 52:1089-1095.

29. Tumbula D, Keswani J, Shieh J, Whitman W. 1995. Maintenance of methanogen stock

cultures in glycerol at -70 °C. Archaea-a laboratory manual. Cold Spring Harbor

Laboratory Press, Cold Spring Harbor, NY 85-87.

30. Jones WJ, Whitman WB, Fields RD, Wolfe RS. 1983. Growth and plating efficiency of

methanococci on agar media. Appl Environ Microbiol 46:220-226. 64

31. Uffen RL, Wolfe RS. 1970. Anaerobic growth of purple nonsulfur bacteria under dark

conditions. J Bacteriol 104:462-472.

32. De Man J. 1983. MPN tables, corrected. European Journal of Applied Microbiology and

Biotechnology 17:301-305

33. Bryant MP. 1972. Commentary on the Hungate technique for culture of anaerobic bacteria.

Am J Clin Nutr 25:1324-8.

34. Weimar M, Cheung J, Dey D, McSweeney C, Morrison M, Kobayashi Y, Whitman W,

Carbone V, Schofield L, Ronimus R. 2017. Development of multi-well plate methods using

pure cultures of methanogens to identify new inhibitors for suppressing ruminant methane

emissions. Appl Environ Microbiol:AEM. 00396-17.

Figure 2-1. Effect of selected buffers on growth. Methanococcus maripaludis S2 was grown in

McN (H2/CO2) medium with different concentration of tested buffers. The culture absorbance was determined after one day.

0.6

Blue stoppers 0.5

0.4

0.3

0.2 Tris Glycine GlyGly 0.1

600 0.0

A 0.5 Gray stoppers

0.4

0.3

0.2

0.1

0.0 0 50 100 150 Time [h]

Figure 2-2. Growth of Methanococcus maripaludis S2 with 200 mM formate and 100 mM of Tris, glycine, and glycylglycine buffers. Two kinds of serum bottle stoppers were used. Blue stoppers are thick butyl rubber stoppers (Bellco Glass, Inc., Vineland, NJ, cat. no.: 2048-11800, 704.82

USD/1000). They are commonly used for H2/CO2 medium. Butyl rubber gray stoppers (Wheaton

Science Products, cat. no.: W224100-202, 174.2 USD/1000) were also tested for their durability during M. maripaludis cultivation.

Figure 2-3. Growth of Methanococcus maripaludis S2 in H2/CO2 (●) and formate medium (○).

The inoculum size was 5×104 cells per 5 mL of culture. All values were the averages of five cultures.

Figure 2-4. Growth of Methanococcus maripaludis S2 () and Methanothermococcus okinawensis () in the medium-scale culture system. The inoculum was 1010 cells per 1.5 L of culture. All values are the averages of three cultures.

Table 2-1. Most probable number dilution of Methanococcus maripaludis S2 in medium prepared by the rapid protocola.

Inoculum ( # of cells) Positive No. Negative No.

Three O2 removal cycles 1000 5 0 100 5 0 10 1 4 1 1 4 0.1 0 5 Control 1000 5 0 100 5 0 10 3 2 1 1 4 0.1 0 5 a Three cycles of gas exchange used in preparation of the McF medium as described in

Supplementary A, section S2B. The inoculum was serially diluted into 1000, 100, 10, 1 and 0.1 cells. Growth was monitored for 6 days. When the OD600nm was greater than 0.6, growth was defined as positive. Control medium was prepared in the anaerobic chamber as described in

Supplementary A, section S2A.

CHAPTER 3

AN ADENYLYL-SULFATE REDUCTASE IN METHANOCOCCUS MARIPALUDIS,

CONTAINS AN IRON-SULFUR CLUSTER, AND IS REQUIRED FOR ELEMENTAL

SULFUR ASSIMILATION2

2 Long, F., Y. Liu, M. Cavuzic, J. Amster, E. Duin, R.H. White and W.B. Whitman. An adenylyl- sulfate reductase in Methanococcus maripaludis, contains an iron-sulfur cluster, and is required for elemental sulfur assimilation. To be submitted to Molecular Microbiology.

Abstract Methanococcus maripaludis is known to assimilate elemental sulfur as the sole sulfur source, while the corresponding enzyme(s) involved in this process has not been elucidated. In this study, we demonstrated that MMP1681 from M. maripaludis is required for elemental sulfur incorporation. A M. maripaludis strain with an in-frame deletion of mmp1681was not able to grow with elemental sulfur as sole sulfur source and was severely impaired for growth with sulfide as the sole sulfur source. However, when grown with elemental sulfur as the sole sulfur source, genetic complementation of the mmp1681 deletion or supplementation of the medium with coenzyme M or thiosulfate, restored the growth of the mutant to near wild-type levels. In addition, proteomics data showed that the expression of MMP1681 increased 3.3-fold in during growth on elemental sulfur in comparison to growth on sulfide. These observations suggest that MMP1681 is required for elemental sulfur assimilation in M. maripaludis. In vitro enzymatic assay found that

MMP1681 is an adenylyl-sulfate reductase that catalyzes the sulfite production from adenosine-

5’-phosphosulfate (APS), instead of 3'-phosphoadenosine-5'-phosphosulfate (PAPS). The purified recombinant MMP1681 used glutaredoxin as an electron donor in vitro, showing a Km for APS of

-1 112 µM, at a Vmax of 0.03 µmoL·min per mg protein. In addition, an iron-sulfur cluster has been identified on the anaerobically purified recombinant protein MMP1681. Nonetheless, more genetic and biochemical evidence indicate sulfate assimilatory pathway may be absent in M. maripaludis.

Together with further sequence analysis, a different physiological role of MMP1681 in elemental sulfur assimilation, in addition to its in vitro catalytic function as an adenylyl-sulfate reductase was demonstrated.

Introduction Among the lower oxidation states of sulfur compounds, elemental sulfur is possibly the most widespread sulfur species in the sediments and geological deposits. Mainly formed from biological and chemical oxidation of hydrogen sulfide, elemental sulfur is chemically relatively reactive and mostly stable in the form of eight atom ring that constitutes orthorhombic crystals (S8)

(1). Elemental sulfur melts at 119 oC, and spontaneously decomposes above 80 oC (2). Sulfur dissolves poorly in water (0.16 µmol per liter at 25 oC) (3), and the solubility at higher temperatures is not known. Hence the low solubility of sulfur in water becomes one of the biggest obstacles for its utilization. It is commonly known that pure elemental sulfur, such as rhombic sulfur, cannot directly serve as the true substrate for sulfur reductase (4). Instead, the so-called hydrophilic sulfur is presumably the true substrate. This is the “soluble form” in aqueous medium (Supplementary figure S3-1). It may constitute elemental sulfur together with small amounts of oxo-compounds,

- - such as polythionates ( O3S-Sn-SO3 ) (4). Aqueous suspensions of sulfur, particularly chemically precipitated colloidal sulfur, and sulfur formed by sulfide-oxidizing microorganisms, contain diverse portions of such hydrophilic sulfur (5). Another possible form of solubilized sulfur exist

- in the presence of sulfide is present. In the aqueous solution of sulfide (H2S/HS , pKa1=7.02), the

- S8-ring of elemental sulfur is cleaved by a nucleophilic attack of the HS , producing polysulfide as in the soluble form of sulfur (6-8) (Supplementary figure S3-1). In medium with a pH around 7,

2- 2- the majority of the polysulfide species consists of tetrasulfide (S4 ) and pentasulfide (S5 ) (5).

These two polysulfides interconvert rapidly and are in equilibrium with lower concentrations of other polysulfide species. It is still unknown which of these two polysulfides is the preferred substrate for polysulfide reductase. In Wolinella succinogenes, evidence has shown that polysulfide is the actual electron acceptor in sulfur respiration (9). Alternatively, many bacteria

grow with elemental sulfur in the absence of solubilized form of polysulfide. These microorganisms are anaerobes that disproportionate elemental sulfur in the presence of sulfide- scavenging ferric minerals (10) or aerobes that oxidize extracellular sulfur (5). How these organisms deal with the low solubility of sulfur in water is still not known.

Microorganisms known to reduce elemental sulfur using H2 or organic substrates as electron donors are widespread among Bacteria and Archaea (7, 11). The oxidation of organic substrates in sulfur reducers may be complete, leading to CO2 as an end product (genera Acidianus,

Stygiolobus, Thermoproteus, Pyrobaculum, Igneococcus, Pyrodictium, Wolinella, Desulfuromonas,

Ammonifex, and Desulfurobacterium), or incomplete, leading to acetate as the final product

(archaeal genera Desulfurococcus, Staphylothermus, Hyperthermus, Thermococcus, and

Pyrococcus and the bacterial genera Thermotoga, Thermosipho, and Fervidobacterium,

Supplementary table S3-2) (7). The sulfur reductase (EC 1.97.1.3) is a constitutive enzyme characterized in sulfur-reducing bacteria (Desulfovibrio and Desulfomicrobium species), sulfur- reducing eubacteria (W. succinogenes DSM 1740 and Sulfurospirillum deleyianum DSM 6946), and in the hyperthermophilic archaea Pyrococcus furiosus and Pyrodictium species (12, 13). On the other side, sulfur dispropotionators, mainly from delta class of Proteobacteria, produce sulfate and hydrogen sulfide in a chemolithotrophic process using elemental sulfur as both electron donor and acceptor (14). Regardless, the enzymes involved in sulfur disproportionation have not been elucidated.

Several genera of domain Bacteria and Archaea are able to grow by a dissimilatory reduction of elemental sulfur to sulfide in a respiratory type of metabolism (4). This observation is supported by their growth on H2 and elemental sulfur (11). Most methanogens can assimilate elemental sulfur as a sulfur source (15). In S8, the most stable form of elemental sulfur at room 74

temperature, the sulfur atoms are connected via S-S bonds similar to heterodisulfide. The redox

0 potential of the heterodisulfide/[H-S-CoB+H-S-CoM] couple is very close to that of the S /H2S couple (around -200 mV). Among the metabolism of dissimilatory reduction of elemental sulfur to hydrogen sulfide in sulfur-reducing microorganisms and methanogens, one common feature is that they both live at the expense of S-S bond reduction. Nonetheless, methanogens different from sulfur reducers in that they don’t rely on an external sulfur species as electron acceptor because they can re-oxidize the “reduced sulfur” with CO2 or one of the other methanogenic substrates. In

the presence of H2, many methanogens produce a large amount of sulfide from elemental sulfur

(15, 16). Although abiotic elemental sulfur disproportionation produces significant amounts of sulfide, thermophilic and hyperthermophilic methanogens, such as Methanopyrus,

Methanothermobacter, Methanothermus, and Methanothermococcus also produce a substantial amount of sulfide (2, 15). Among them, Methanothermococcus thermolithotrophicus, which grows at 65 oC, produces sulfide at the highest rate [around 845 µmol per g cell dry weight (wt) per hour], which is three-fold higher than the abiotic reaction under the same conditions (17). Meanwhile, mesophilic methanogens also reduce elemental sulfur well above the rate of abiotic sulfur disproportionation (18). Fauque et al. have detected sulfur reductase activities in cell extracts of

Methanosarcina bakeri, M. thermolithotrophicus and Methanothermobacter thermoautotrophicum (15, 16). Nevertheless, the enzyme that catalyzes elemental sulfur reduction in methanogens has not yet been determined.

Present as the most abundant form of sulfur in nature, sulfate is thermodynamically stable, and its reduction constitutes the basis of the biological sulfur cycle. Archaea, bacteria, fungi, and plants reduce sulfate to sulfide for variant purposes. This sulfate reduction is mainly divided into two forms. The first one is assimilatory sulfate reduction pathway, mainly carried out by aerobes 75

(Figure 3-1A). It is indispensable for cysteine biosynthesis. The other form, sulfate dissimilatory reduction, is carried out by anaerobic prokaryotes. In the absence of oxygen, sulfate is utilized as a terminal electron acceptor for respiration (Figure 3-1B). In the assimilatory pathway, the reduced product, sulfide, is incorporated into the thiol group of cysteine, whereas in the dissimilatory pathway, it is released as a waste product in the form of hydrogen sulfide. The steps in both pathways are similar but still different in details. Frist, sulfate is activated with ATP to form adenosine-5’-phosphosulfate (APS). In plants and sulfate dissimilators, using reduced glutathione

(GSH), thioredoxin (Trx) or glutaredoxin (Grx) as the electron donor, APS is reduced to sulfite by

APS reductase (APR). Sulfite is finally reduced to sulfide with NADPH as the electron donor (19).

However, in sulfate assimilatory pathways, the initial substrate for reduction is not APS, but 3'- phosphoadenosine-5'-phosphosulfate (PAPS), which is formed by an ATP-dependent phosphorylation of APS by APS kinase. The generated PAPS is then reduced into APS by PAPS reductase (PAPR) before the further reduction to sulfite and eventually to sulfide (Figure 3-1A).

Similar to NADH and NADPH, APS and PAPS differ only by a single phosphate group in chemical structure. Both the APR and PAPR proteins possess a reductase domain, including a sulfonucleotide-binding motif and a catalytic motif ([KRT]ECG[LI]H) containing active-site cysteine (20, 21) (Figure 3-2A). Nonetheless, only ~20 % identical amino acid residues are shared among APR and PAPR proteins (20, 21). In addition to the catalytic motif, the APR possess another two motifs, CCXXRKXXPL and SXGCXXCT that are not found in the PAPR (20-26)

(Figure 3-2A). Three of the four Cys residues found in these motifs form into two cysteine pairs, and become the ligands for a [4Fe-4S]2+ binding iron-sulfur clusters (20-26), with only one exception in moss Physcomitrella patens (27). All confirmed APRs, including plant APR, assimilatory bacterial APR and dissimilatory APR from sulfate-reducing bacteria, possess the iron- 76

sulfur cluster as cofactors (20). It is proposed that the reduction of APS is associated with the presence of the iron-sulfur cluster in APR (20) because the loss of the iron-sulfur clusters in several plant APRs and assimilatory bacterial APRs had detrimental effects on the enzyme activity (20-

26). An apparent exception that is not understood is the APR from Catharanthus roseus (28).

Additionally, when the iron-sulfur cluster is removed from the bifunctional APR/PAPR reductase from Bacillus subtilis, reduction of APS but not PAPS is abolished (25). Assimilatory APS reductases are further grouped into two types, those that use thioredoxin (Trx) or glutaredoxin

(Grx) as electron donors (EC 1.8.4.10), and those that use reduced glutathione (EC 1.8.4.9). The

Trx/Grx-dependent form of APR is found in diverse microbial species, whereas the glutathione

(GSH)-dependent form is found in plants and green algae. The ability of the plant-type APR to use

GSH as an election donor seems to be a function of the thioredoxin-like C-terminal domain that resembles Trx and Grx (29, 30). However, this domain is absent in the assimilatory bacterial APRs.

Unlike plants and most bacteria, methanogens are generally unable to assimilate sulfate.

Only M. thermolithotrophicus and Methanobrevibacter ruminantium are able to use sulfate as a sole sulfur source (31, 32). This is not surprising considering that sulfate reducers are commonly present the habitat of most methanogens and sulfide is readily available (16, 33). Similarly, for methanogen habitats on the interface between aerobic and aerobic zones, the sulfide formed in the anaerobic zone may be oxidized into elemental sulfur in an abiotic or biotic manner and make this alternative sulfur source available for methanogens (16). Genomic evidence for the absence of sulfate reduction pathway comes from the fact that a gene encoding the first enzyme in the pathway, sulfate adenylyltransferase (ATP sulfurylase or adenosine diphosphate sulfurylase), is absent in all methanogenic genomes. However, the recent characterization of APS reductase (MJ0973) (34),

PAPS reductase (MJ0066) (35) and sulfite reductase (MJ0870) (36, 37) from Methanocaldococcus 77

jannaschii has called into questioned whether or not methanogens can reduce sulfate. The introduction of the MJ0870 gene into M. maripaludis allowed them to reduce sulfite to sulfide (37), which is a known to inhibit methanogenesis in at least one methanogen (38-40). In Methanococcus maripaludis, MMP1681 and MMP0941 are annotated as sulfunonucleotide reductase, and they are homologs of MJ0973 and MJ0066 respectively. In addition, MMP0078 has been described as a small sulfite reductase, as it represents the C-terminal domain of the MJ0870 protein (41).

However, the enzymatic activities of these enzymes have not been demonstrated.

In this study, a M. maripaludis deletion mutant of mmp1681 cannot grow with elemental sulfur as sole sulfur source. Genetic complementation of the deletion mutant of mmp1681 or supplementation of the medium with coenzyme M or thiosulfate was able to restore the Δmmp1681 mutant’s growth to wild-type level. Moreover, proteomics analysis indicated a three-fold upregulated expression of MMP1681 during growth with elemental sulfur in comparison to growth with sulfide. Purified MMP1681 expressed as a His-tagged recombinant protein in E. coli was able to reduce APS instead of PAPS, while MMP0941 was able to reduce PAPS instead of APS, using glutaredoxin as an electron donor. Additionally, an iron-sulfur cluster has been detected on both recombinant MMP1681 and MMP0941. These observations have brought in the question of whether a sulfate assimilation reduction pathway driven by MMP1681 and/or MMP0941 exist in

M. maripaludis. The answer seemed to be negative according to these two lines of evidence: (i) M. maripaludis doesn’t consume or produce sulfate, at least not at micro-moloar levels, when grown with sulfide as sulfur source; (ii) Putative M. maripaludis sulfate assimilation homologs were unable to complement the corresponding E. coli mutants. Lastly, the evolutionary history of

MMP1681 and its methanogenic homologs compose a new group of sulfonucleotide reductases.

This further supports the hypothesis that MMP1681 delivers distinctive physiological role other 78

than as an APR in vitro.

Methods and materials

Strains, medium and growth condition. Two M. marpaludis strains, S2 and S0001, were used in this study. M. maripaludis S0001 is a mutant derived from wild-type strain S2 by the deletion of the gene encoding hypoxanthine phosphoribosyltransferase (mmp0145) and the addition of the gene encoding the rep gene from the Methanococcus shuttle vector pURB500 (42).

It is frequently used in a markerless mutation system and as a host for shuttle vectors of M. maripaludis (43). S0001 was used to generate the gene deletion mutant strain S211 (Δmmp1681.

Cultures were grown in H2/CO2 medium (McNAA) (44) or formate medium (McFAA) (45). Both types of medium were supplemented with 10 mM sodium acetate and 1 mM alanine for optimal growth of M. maripaludis. The medium was reduced with 2 mM dithiothreitol or 2 mM titanium

(III) citrate (46) as indicated. The 5 mL cultures were grown in 28-mL aluminum seal tubes pressurized to 276 kPa H2/CO2 in McNAA or 103 kPa with N2/CO2 (4:1, v/v) in McFAA. The agar medium was prepared in 70 mL serum bottles with 10 mL of McFAA, supplemented with 1 %

(w/v) agar. Serum bottles were pressurized to 103 kPa with N2/CO2 (4:1, v/v). Before inoculation,

2 mM sodium sulfide or 0.02 g/mL (w/v) elemental sulfur was added as the sulfur source. 2.5 µg/ mL of puromycin and 0.25 mg/mL of 8-azahypoxanthine (zhpt) were supplemented when indicated.

Measurement of inorganic sulfide produced in medium and M. maripaludis cultures with elemental sulfur as the sole sulfur source. When measuring the abiotic sulfide production in McFAA medium in the presence or absence of different chemicals, these medium tubes were incubated overnight at indicated temperature in anaerobic chamber or 37 °C open-air incubator

before inorganic sulfide measurement. When measuring the inorganic sulfide production in M. maripaludis S2 culture, it was grown in 5 mL of McFAA medium to an absorbance of 0.8 at 600 nm after 16 hr or 24 hr of incubation at 37 °C. The supernatant from media or M. maripaludis S2 cultures were collected by centrifugation at 13,000 × g for 15 min at room temperature in the anaerobic chamber.

The inorganic acid-labile sulfide content in McFAA media or M. maripaludis culture broth was determined by an adaptation of the methylene blue method (47-49). The measurement was performed in the anaerobic chamber, which contained an atmosphere of 95 % N2 and 5% H2. All reagents were prepared freshly and spared with 103 kPa N2 for one hour prior to inorganic sulfide measurement. To assay the aqueous sulfide levels, 0.3 mL of 1 % (w/v) zinc acetate and 10 µL of

12 % sodium hydroxide were added to 200 µL of medium or supernatant in microcentrifuge tubes.

After mixing, the solution was incubated for 20 min. Then 0.1 mL of 1 % of N, N- dimethylphenylenediamine in 5 M of hydrochloric acid and 40 µL of 11.5 mM Iron (III) chloride in 1.2 M of hydrochloric acid were added, and the solution was mixed rapidly. The assay solution was incubated for 10 min, and its absorbance was determined at 670 nm. Solutions of sodium sulfide were used as standards. 0.07 M NaOH was used as the solvent for sulfide standards and sample dilutions. To measure the gaseous sulfide levels, 0.05 mL - 1 mL gas sample was taken from medium or culture tubes using a syringe and added into a 4.5 mL aluminum-sealed vial, later steps were the same as measuring the aqueous sulfide. Because the presence of dithiothreitol (DTT)

(50) or sodium thiosulfate (51) was found to interfere with this assay (50), McNAA medium with dithiothreitol was first acidified with 2 M hydrochloric acid (final concentration) to volatilize the aqueous sulfide into gaseous sulfide, which was later measured by the described method (Table 3-

1, Supplementary table S3-3). In addition to the described sulfide measurement method, this acidification method was also used along to examine the abiotic sulfide formation in elemental sulfur medium supplemented with titanium (III) citrate, coenzyme M, thiosulfate, sulfate or sulfite

(Table 3-1, Supplementary table S3-4). The results obtained from both methods were consistent.

Differential 14N-15N labeling of proteins. To prepare differentially 14N-15N labeled proteins, the wild-type strain S2 was grown in McNA medium containing either 4.6 mM unlabeled

[14N] ammonium sulfate or 4.6 mM ≥ 99 % [15N] ammonium sulfate in sidearm bottles, using either 2 mM sodium sulfide or 0.02 g/mL (w/v) elemental sulfur as sulfur source. These bottles were manufactured from 160 mL serum bottles by fusing a 28-mL aluminum seal tube to the side.

They could be pressurized to 138 kPa and contained 10 mL of culture. To prevent contamination with other nitrogen compounds, glassware was immersed in 1 M HCl overnight, rinsed with deionized water, and dried before use. Rubber stoppers were autoclaved for 20 min in 0.5 M NaOH and rinsed thoroughly with deionized water. Cultures were inoculated a day after addition of sulfide and ammonium sulfate to the medium. The inocula were 0.1 mL of cultures grown in the same medium. After inoculation, cultures were initially pressurized to 138 kPa with H2/CO2 gas

(80:20 [v/v]) and repressurized twice daily. When S2 cultures reached an average absorbance at

600 nm of 0.6, one culture of unlabeled S2 grown with sulfide was mixed with one culture of 15N- labeled S2 grown with elemental sulfur to get sample A. Sample B was the mixture of one culture of unlabeled S2 grown with elemental sulfur and one culture of 15N-labeled S2 grown with sulfide.

Cells were harvested by centrifugation at 10,000 × g for 30 min at 4°C. The pellets were suspended in 0.5 mL of 100 mM ammonium bicarbonate buffer (pH 7.8), and 5 µL of 0.5 M phenylmethylsulfonyl fluoride was added to inhibit protease activity. The cells were lysed by

freezing at -20°C. Upon thawing, 10 U of DNase I was added to the cell lysate, and the crude extracts were incubated at room temperature for 1 h. After DNA digestion, the crude extracts were centrifuged at 16,000 × g for 30 min to remove unbroken cells. Because of the carryover of

14 15 NH4Cl from the inocula, the proteins from cultures grown with [ N] ammonium sulfate were

~98 % enriched with [15N].

Quantitative proteomics. Protein digestion, fractionation, and mass spectrometry were performed as described previously (52). The accurate mass of each peptide was obtained from the m/z values of monoisotopic peaks. The number of nitrogen atoms in each peptide was determined from the mass separation between the monoisotopic peaks of the peptide and its 15N-enriched counterpart. The identification of peptides was automated using the software described previously

(52). A measured mass peak was considered identified when it was within 10 ppm of the mass of a predicted tryptic peptide that had the same nitrogen content. A protein (open reading frame

[ORF]) was considered identified when at least one unique peptide, i.e., predicted to be encoded in only one ORF, was identified. Multiple peptide pairs from the same ORF were usually detected because more than one unique peptide was detected for many proteins, and most peptides were detected in multiple matrix-assisted laser desorption/ionization spots. The number of peptide pairs collected per ORF was designated. The relative signal intensity (peak height) of 14N and 15N peptide peaks in each peak pair was used as a measure of relative peptide level. Multiple measurements of one protein were averaged to obtain the relative protein levels in S2. The average relative protein levels for all proteins were normalized to 1. Differential protein levels were regarded as significant if N was >2, the normalized relative protein levels were not between 0.7

and 1.5 (equivalent to >1.5-fold changes), and the 95 % confidence interval for the normalized ratio did not include the value 1.00.

Mutagenesis of mmp1681 in M. maripaludis. Mmp1681 was deleted from the M. maripaludis genome using two plasmids: p5L-U and p5L-D (53). This strategy uses two identical repetitive elements (RE) on a suicide plasmid to enable the homologous recombination for removing the selective markers and eventually leave a short scar to replace the target gene. A 1 kb fragment flanking the downstream (Down) of mmp1681 was amplified from the M. maripaludis genome and cloned into the region between AflII and Xbal on plasmid p5L-D. The region of the

90-kb repetitive element together with the subsequent downstream of mmp1681 was then amplified and cloned into plasmid p5L-U, between Spel and HindIII. The upstream (UP) of mmp1681, a 1 kb fragment, was amplified from the M. maripaludis genome and continuously cloned into this plasmid p5L-U, between Ndel and KpnI. This p5L-U plasmid contains RE,

Methanococcus voltae histone-like protein promoter (PhmvA), hypoxanthine phosphoribosyltransferase (hpt) gene, puromycin N-acetyltransferase (pac) (53), flanked by two multicloning sites for inserting the upstream, RE, and downstream region of the gene to be deleted.

The final deletion fragment containing UP-RE-Marker-RE-Down was amplified from the newly constructed p5L-U plasmid, and transformed into M. maripaludis S0001. To enrich and select transformants with UP-RE-Marker-RE-Down knocked-in fragment fused into the M. maripaludis genome, which happened after the first recombination event, 0.1 mL of transformants was inoculated into McFAA medium in the presence of puromycin. The grown-up culture was transferred twice in the same type of medium to homogenize genome copies with knock-ins. To further remove markers from the genome by the second recombination event, a culture from the

previous step was first grown in McFAA medium without puromycin, then grew in McFAA medium with the 8-azahypoxanthine (zhpt). After growth, this culture was subsequently plated on

McFAA agar medium in the presence of 8-azahypoxanthine. Individual colonies were picked and screened for markerless deletion mutations of mmp1681. Three sets of PCR primers, targeting at pac cassette (p5LU-F1 and p5LU-R1), the deleted fragment of mmp1681 (InnerMMP1681F and

InnerMMP1681R), upstream to downstream of mmp1681 (MMP1681UpF and MMP1681DownR) were used to examine the complete deletion of mmp1681 (Supplementary figure S3-3A to D). This mutant Δmmp1681 strain was named as S211.

For complementation of the Δmmp1681 strain (S211), mmp1681 was cloned into the shuttle vector pMEV4 (54) using primers pMEV4-MMP1681F and pMEV4-MMP1681R

(Supplementary table S3-1). The resulting plasmid, pMEV4-MMP1681, was transformed into the strain S211. The transformants were plated on McFAA agar medium plus puromycin and neomycin. Individual colonies were picked from this agar medium and transferred twice in

McFAA medium in the presence of puromycin and neomycin. To confirm the successful introduction of pMEV4-MMP1681 in S211 strain, plasmids were extracted from these screened colonies respectively and used as template for PCR amplification. Primers pAW5042-F and pAW5042-R (Supplementary table S3-1) were used to amplify the fragment covering the region of promoter Pmcr to mmp1681 in pMEV4-MMP1681 (Supplementary figure S3-3E). These PCR amplification products were further sequenced to ensure that single nucleotide polymorphisms

(SNP) were absent. The complemented strain expressing the wild-type MMP1681 was named as

S212.

Plasmid construction, expression, and purification of recombinant MMP1681 and

MMP0941. Mmp1681 and mmp0941 were PCR amplified from M. maripaludis S2 genomic DNA, digested and inserted between the NdeI and SacI restriction sites on the vector pQE2 (Qiagen), vector. DNA sequences of the cloning site were verified by restriction digestion and sequencing with the pQE2 reverse primer (Qiagen). All of the recombinant protein sequences contained a N- terminal His×6 affinity tag. The resulting plasmids were individually transformed into the E. coli

M15 [pREP4] strain (Qiagen) for expression of recombinant proteins.

For aerobic purification of recombinant proteins, the transformed E. coli cells were grown in 1 L Luria-Bertani (LB) medium supplemented with 50 µg/mL ampicillin and 25 µg/mL kanamycin at 37 °C with shaking until they reached an absorbance at 600 nm of 0.4 – 0.6. Cultures were then induced with 1 mM IPTG (Isopropyl β-D-1-thiogalactopyranoside) and incubated overnight at 30°C. The next day, cultures were harvested by centrifugation at 4,750 × g for 10 min at 4°C. Then the cell pellets were resuspended in 10 mL of lysis buffer, including 30 mM Tris-

HCl (pH= 8.0), 1 mM EDTA and 200 mM NaCl. One pellet of complete EDTA-free Protease

Inhibitor Mixture (Roche) together with a final concentration of 50 µg/mL lysozyme were subsequently added to the cell suspension. This cell suspension was further sonicated for 10 min with pulse on 2 seconds, off 1 second and an amplitude of 50 %. Lysed cells were centrifuged at

15,000 × g for 30 min at 4 °C to remove cell debris. The supernatant was applied to a 1 mL Ni-

NTA column (Thermo Scientific) equilibrated with 10 column volumes (CV) H2O, followed with

10 mL binding buffer containing 30 mM Tris-HCl (pH= 8.0), 500 mM NaCl and 10 mM imidazole.

The proteins were washed with 20 mL washing buffer, including 30 mM Tris-HCl (pH= 8.0), 500 mM NaCl and 30 mM imidazole. After incubating for 2 min, proteins bound to the column were

then eluted with 5 mL of elution buffer, consisting of 30 mM Tris-HCl (pH= 8.0), 500 mM NaCl and 200 mM imidazole.

For anaerobic purification of the recombinant proteins, the transformed E. coli cells were grown in 1 L Luria-Bertani (LB) medium supplemented with 50 µg/mL ampicillin, 25 µg/mL kanamycin and 20 µM ammonium iron (III) citrate. The cultures were grown at 37 °C until it reached an optical density at 600 nm of 0.4–0.75. Cultures were then induced with 200 µM ammonium iron (III) citrate, 30 µM L-methionine and 1 mM IPTG and incubated at 30 °C overnight. The next day, the cultures were harvested by centrifugation at 4,750 × g for 10 min at

4 °C. The following steps were performed in the anaerobic chamber. The cell pellets were resuspended in 5 mL of lysis buffer, including 50 mM sodium HEPES (pH= 8.0), 500 mM NaCl and 5 mM MgCl2. One pellet of complete EDTA-free Protease Inhibitor Mixture (Roche), a final concentration of 50 µg/mL lysozyme and 12.5 U of DNase I were subsequently added to the cell suspension. This cell suspension was further sonicated for 5 min with pulse on 2 seconds, off 1 second and an amplitude of 50 %. The cell lysate was then centrifuged at 15,000 × g for 30 min at

4 °C. The supernatant was applied onto a 0.2-0.4 mL Ni-NTA column (Thermo Scientific) equilibrated with a column volume of double-distilled water, followed with 10 column volumes of binding buffer containing 50 mM sodium HEPES (pH= 8.0), 500 mM NaCl and 20 mM imidazole.

The proteins were then washed with 20 column volumes of washing buffer, including 50 mM sodium HEPES (pH= 8.0), 500 mM NaCl and 50 mM imidazole. After incubating for 2 min, proteins bound to the column were eluted with 5 column volumes of elution buffer consisting of

50 mM sodium HEPES (pH= 8.0), 500 mM NaCl and 300 mM imidazole.

The homogeneity of purified proteins was confirmed by 12 % SDS-PAGE polyacrylamide gel electrophoresis and the Gel Pro Analyzer program version 4.0 from Media Cybernetics, L.P.

(Supplementary figure S3-4). Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-

PAGE) was accomplished using Mini-PROTEAN TGX Precast Gels (Bio-Rad) 12 % polyacrylamide. Gels were stained using Acqua Stain (Bulldog-bio). SDS-PAGE of the denatured

E. coli PAPR, MMP0941 and MMP1681 showed molecular weights of ~ 28, 57, and 51 kDa, respectively, consistent with the molecular weight of E. coli PAPR and the predicted molecular weight of MMP0941 and MMP1681. Proteins concentrations were determined using the BCA

Protein Assay Kit (Pierce).

APR/PAPR enzymatic assays. The APR/PAPR activities were measured as the consumption of NADPH by assaying the decrease of its absorbance of 340 nm using a spectrophotometer. The assays were carried out in 200 µL reactions containing 100 mM Tris-HCl

(pH= 8.5), 500 mM MgSO4, 1 mM EDTA, 1~60 µM APS (Sigma, 102029-95-8) or 1~80 µM

PAPS (Sigma, 109434-21-1), the reductant system and 0.5 µM recombinant protein (55). The reaction was incubated for 30 min at 30 °C (55). The reductant system was composed of 2 µM E. coli thioredoxin (Sigma, 52500-60-4), 100 µM NADPH (Sigma, 100929-71-3), and 100 nM thioredoxin reductase (Sigma, 598502) or 7 mM glutathione, 100 µM NADPH, 0.45 µM glutaredoxin (Sigma, G5298) and 1 unit of glutathione reductase (Sigma, 9001-48-3). The assay was initiated by the addition of APS or PAPS. All kinetic experiments were repeated at least three times. Kinetic data analyses, other calculations, and statistical analyses were performed using

SigmaPlot 12.0 (Systat Software Inc.).

Determination of Iron and UV-visible Spectra. The iron content of the proteins was quantified by using the Quantichrom Iron Assay Kit (BioAssay Systems). UV-visible absorption spectra were recorded on a Nanodrop 2000c spectrophotometer with samples in quartz cuvettes

(optic path = 1 cm) closed with rubber stoppers under anoxic conditions. The iron measurements were performed with purified proteins from three independent cultures. 0.1 M NaOH was used for preparing samples of standards and protein dilutions.

Complementation of the E. coli sulfate assimilation mutants with putative M. maripaludis homologs. The genes that encode putative M. maripaludis sulfur metabolism homologs, mmp0078, mmp0941, mmp1282, mmp1471 and mmp1681 were cloned into vector pQE2 (Qiagen). The E. coli PAPR (CysH) was also cloned into vector pQE2 (Qiagen), this construct served as a positive control for the complementation experiment. The cloning of

MMP0941 and MMP1681 into vector pQE2 was introduced in the earlier method description.

Mmp0078, mmp1282 and mmp1471 were PCR amplified from M. maripaludis genomic DNA, digested and inserted between the KpnI and HindIII restriction sites on the vector pQE2 (Qiagen).

The E. coli PAPR was PCR amplified from E. coli K-12 genomic DNA, digested and inserted between the NdeI and SacI restriction sites on the vector pQE2 (Qiagen), DNA sequences at the cloning site were verified by restriction digest and sequencing with the pQE2 reverse primer

(Qiagen). The corresponding primers were listed in Supplementary table S3-1.

The E. coli mutant strains JW2720 (cysC-), JW2722 (cysD-), JW2732 (cysH-), JW2733

(cysI-), JW2734 (cysJ-) and JW 3331 (cysG-) were recovered from the Keio collection (56)

(Supplementary table S3-1). Each of these six E. coli mutants was transformed with each of the above six plasmids: pQE2-mmp0078, pQE2-mmp00941, pQE2-mmp1282, pQE2-mmp1471,

pQE2-mmp1681, and pQE2-cysH, individually. The transformants were plated on to LB agar medium in the in the presence of 100 µg/mL ampicillin and 25 µg/mL kanamycin. Two colonies were picked from each plate and grew in 5 mL LB broth medium with the same antibiotics. The plasmid of each culture was extracted and digested to confirm the presence of the plasmid construct.

To test if the M. maripaludis protein complement the E. coli mutant, 5 µL of the previous LB broth culture was inoculated into 3 mL M9 medium containing 1 mM isopropyl β-D-1- thiogalactopyranoside 100 µg/mL ampicillin and 25 µg/mL kanamycin, in the presence or the absence of 2mM cysteine. Three replicates were made for each type of medium. These inoculated cultures were then incubated at 30 °C with shaker on for two days. The expression of M. maripaludis protein was confirmed by 12 % SDS-PAGE polyacrylamide gel electrophoresis with the corresponding whole-cell lysate.

Measurement of inorganic sulfate levels in McFAA medium and M. maripaludis culture. M. maripaludis S2 cells were grown in 5 mL of sulfate-depleted McFAA medium. In this sulfate-depleted medium, all sulfate anion contained ingredients from general salts, iron, and trace mineral solutions were replaced with chloride-anion contained chemicals. The inoculum size was

2.5×107 cells, the inoculated culture was incubated 49 hours at 37°C for growth till an absorbance of ~0.6 at 600nm was reached. The supernatant was anaerobically harvested by the following steps.

The whole aluminum sealed culture tube was centrifuged at 5000 × g for 15 min at room temperature. The centrifuged tube was then carefully transferred into the anaerobic chamber for collecting the supernatant. This tube was inverted to allow the supernatant drain to the stopper end of the tube without disturbing the cell pellet. A double-ended needle was used to transfer the supernatant into a new tube without exposure to O2. Because aqueous sulfide dissolved in the

supernatant interferes the later sulfate measurement, 1 mL of hydrogen chloride (final concentration 0.6 M) was added to the supernatant to volatilize the aqueous sulfide into gaseous sulfide. After incubating the tube for one hour, it was transferred outside of the chamber for exchanging the headspace gas with N2. Three gas-exchange cycles, composing of 15 seconds of

103 kPa N2 alternating with 45 seconds of vacuum, was applied on the acidified supernatant tubes.

The tube was pressured with 103 kPa N2 after the end of the last gas-exchange cycle. The tube was then transferred back to the anaerobic chamber, and the supernatant was collected for the subsequent sulfate measurement.

The sulfate was quantified aerobically by the turbidimetric method (57) right after the supernatant was collected. To measure the sulfate content, 0.5 mL of sample and 0.5 mL of conditioning buffer was added to a microcentrifuge tube. Immediately, 0.25 mL conditioning buffer was added, and the solution was mixed gently. The conditioning buffer was made with 0.73

M hydrochloric acid, 2.6 M sodium chloride, 10 % (v/v) glycerol and 20 % (v/v) isopropyl alcohol.

Then, 0.01 g of 20-30 mesh barium chloride (Sigma) was added to the solution. The solution was mixed by vortexing for a few seconds and incubated for 5 min at room temperature, and its absorbance was determined at 420 nm. Solutions of sodium sulfate were used as standards.

Electronic Spectra. The measurement the absorbance of M. maripaludis culture was performed with a Thermo Scientific Genesys 20 spectrophotometer. For sulfide and sulfate content quantifications, the absorbance was recorded on a Pharmacia Biotech Ultrosepcc 3000 spectrophotometer. For APR/PAPR enzymatic assay measurements, the absorbance was recorded on an Agilent Technologies Cary 60 UV-Vis spectrophotometer equipped with a temperature- controlled cell compartment.

Bioinformatics analysis. The sequence analysis was performed with Geneious 10 software (58). The NCBI sequence database was screened with the BLAST software with the protein sequences of MMP1681 AND MMP0941 as the query sequences. The collected sequences were aligned by using the MUSCLE program in Geneious 10 software (58).

Results

Quantitative measurement of inorganic sulfide produced in M. maripaludis grown with elemental sulfur as sole sulfur source. Stetter and Gaag detected sulfide formation in all three orders of methanogens during growth with elemental sulfur as sulfur source (15). In those experiments, 0.1 g elemental sulfur per mL culture was added into exponential phase cultures, which were further incubated for 55 h before measuring the sulfide production (15). Among the

12 tested methanogen species, M. thermolithotrophicus that grows at 65 oC, produced sulfide at the highest rate [~ 845 mmol per g cell dry weight (wt) per hour], while the mesophile

Methanococcus voltae produced sulfide at the rate of ~ 125 mmol per g cell dry wt per hour (15).

Sulfide production in Methanococcus maripaludis was not tested (15).

To test the sulfide production in M. maripaludis that grows with elemental sulfur as sole sulfur source, the abiotic sulfide formation was first examined. Either autoclaving the medium or incubating with dithiothreitol as a reducing agent led to abiotic sulfide production from elemental sulfur, especially the latter which produced 312 µM sulfide (Supplementary Table S3-3). However, when titanium (III) citrate was used as a reducing reagent in the medium, sulfide was not produced from elemental sulfur. When measuring the sulfide production in M. maripaludis cultures grown with elemental sulfur, the medium was pre-autoclaved before adding the elemental sulfur (0.02 g per mL culture) to avoid abiotic sulfide formation from heating. In the presence of elemental

sulfur as the sole sulfur source and titanium (III) citrate as the reducing reagent, a 5 mL M. maripaludis culture with OD=0.8 produced 78 µM aqueous sulfide and 81 µM gaseous sulfide

(Table 3-1), of a total of ~ 6 mmol per g cell dry wt per hour. When using 2 mM dithiothreitol as the reducing reagent, M. maripaludis produced 502 µM aqueous sulfide and 495 µM gaseous sulfide (Table 3-1), which was well about the amount of sulfide formed abiotically in the same medium. Subtracting the abiotic sulfide production, the biotic sulfide production was equaled a total of ~ 6 mmol per g cell dry wt per hour. Presumably, the sulfur-dithiothreitol adducts were a more available sulfur source than elemental sulfur alone.

Quantitative proteomic analysis of protein expression patterns. Methanogenic archaea live in environments with wide ranges of sulfide availability, from μM to mM levels. Even the methanococci, which grow well at mM levels of sulfide common in tidal marshes are also found in zones of lower sulfide (59).The methanococci also do not maintain a significant ΔpH across their cellular membranes, so the internal pH varies with the external pH from pH 6-8 (60). Because the pKa for H2S is close to 7.0, the ionic form of sulfide inside the cell also varies with the external pH. In response to environments with changes in sulfide levels and ionic form, methanococci are expected to regulate the sulfur network to prioritize sulfur availability for biosynthesis of the most important sulfur-containing compounds. If this is true, then the components of the network might be identified by their response to sulfur availability.

To test the premise that sulfur availability affects protein expression in methanococci, experiments examined the proteome following growth with either sulfide or elemental sulfur as the sole sulfur source were performed. As is common, the growth of M. maripaludis on elemental sulfur was slower than that on sulfide (Supplementary figure S3-2), suggesting that sulfur

assimilation and/or other physiological processes are limiting when elemental sulfur is the sulfur source. Proteomic comparisons were carried out with HPLC-MALDI FT-ICR mass spectrometry.

The quantitative analysis was achieved by differential labeling of proteins with 14N- or 15N-labeled ammonium sulfate as the sole nitrogen source during growth (61). The cells were collected at an optical density (600 nm) of 0.6. Proteins encoded by 269 out of the 1,772 ORFs were detected at least once, and 135 proteins were detected by multiple measurements (Supplementary table S3-4).

Several important conclusions could be reached. First, the levels of a number of proteins potentially involved in sulfur assimilation increased in cells grown on elemental sulfur relative to cells grown on sulfide. These proteins included the cysteinyl-tRNA synthetase (1.9-fold) and methionyl-tRNA synthetase (1.6-fold) (Supplementary table S3-4). Although SepRS and SepCysS were not detected in this experiment, expression of the transulfidosome component MMP1217 increased by 1.7-fold. These results suggest that sulfur assimilation enzymes were up-regulated during growth on elemental sulfur, which may represent a sulfur limitation condition and supports the expectation that the sulfur network is regulated. Second, expression of a number of methanogenesis enzymes that have previously been shown to respond to H2 levels and/or growth rate (62) also changed during grown on elemental sulfur. These results are best explained by differences in the growth rates with the different sulfur sources and demonstrate that carefully controlled growth conditions are necessary to make meaningful comparisons of the physiological state of the cells. Third, the expression of a number of proteins of uncertain function also changed

(Table 3-2). For instance, expression of a putative transcription regulator Ptr2 (MMP1137) increased 2.8-fold in cells grown on elemental sulfur. This protein is homologous to the Lrp/AsnC family of bacterial transcriptional regulators. The M. jannaschii homolog has been shown in vitro to activate transcription by recruitment of the TATA-binding protein (63, 64); Nonetheless, the 93

physiological targets of its regulation are unclear. The up-regulation of this protein suggests the possibility that it may be involved in the regulation of sulfur metabolism. The top 10 up-regulated proteins were summarized in table 3-2. Interestingly, among them, expression of MMP1681, increased 3.3-fold in cells grown on elemental sulfur. However, MMP0078, MMP0941,

MMP1282 and MMP1471 were not detected in this experiment.

MMP1681 is required for growth of M. maripaludis with elemental sulfur as the sole sulfur source. The physiological function of the M. maripaludis MMP1681 was investigated by construction and characterization of a ∆mmp1681 mutant strain (S211). An established in-frame markerless strategy was used to construct a single-deletion strain lacking mmp1681. The complete deletion of this gene was demonstrated using PCR analysis (Supplementary figure S3-3).

To study the role of MMP1681 in sulfur metabolism, we first tested the growth requirement of S211 with sulfide as the sole sulfur source. This mutant grew somewhat more slowly than the wild-type strain (Supplementary figure S3-5). Moreover, while sulfate appeared to stimulate the growth of the wild-type, it did not stimulate the growth of the mutant. To further test the mutant’s ability to utilize sulfide as sole sulfur source, S211 was grown in a variety of sulfide concentrations, ranging from 100 µM to 20 mM. With respect to the wild-type strain S2, growth of mutant strain

S211 was largely retarded (Figure 3-3). This was most apparent at higher sulfide concentrations

(15 mM to 20 mM), where 20 mM sulfide didn’t support the growth of mutant strain S211. On the other hand, wild-type S2 strain was able to grow robustly at sulfide between 500 µM to 20 mM, although an attenuated growth yield was observed at 500 µM and 10-20 mM (Figure 3-3).

To examine whether MMP1681 is associated with elemental sulfur assimilation, we also tested the mutant strain S211 for its ability to utilize elemental sulfur as a sole sulfur source.

Strikingly, S211 was unable to grow with elemental sulfur as sulfur source. However, strain S212, which contained the genetic complementation of the Δmmp1681 mutation, grew nearly as well as wild-type (Figure 3-4B). This phenotype implies that MMP1681 may involve in elemental sulfur assimilation with critical functions that convert elemental sulfur to certain compound required for growth. It is possible that elemental sulfur reacts with a thiol form a persulfide before being taken up by the cell. Because essential sulfur-containing amino acids such as cysteine, homocysteine or methionine, were not required for the growth of S211 with sulfide as sole sulfur source, other sulfur species were tested for their ability to restore the growth of S211 with elemental sulfur.

Coenzyme M is an indispensable terminal methyl-carrier in methanogenesis, while 3- mercaptopropionate acid (Supplementary figure S3-6) represents one of its structural analogs.

Although known for its inhibitory effects on M. maripaludis’ growth, sulfite is also proposed to be a precursor of a sulfonate group to coenzyme M biosynthesis (42). It is also equally important to test whether MMP1681 is physiologically related to sulfite production in M. maripaludis.

Therefore, 0.1 mM sulfite supplementation to the elemental sulfur medium was tested. Thiosulfate as a significant electron shuttle, possibly forming a variety of sulfur species via abiotic or biological reactions. The abiotic sulfide formation from elemental sulfur and these tested sulfur- containing compound was also examined. Although 2 mM coenzyme M together with elemental sulfur produced 69 µM aqueous sulfide and 20 µM gaseous sulfide, 0.1 mM coenzyme M, 0.1 mM or 2 mM thiosulfate, sulfate, or sulfite supplementation to elemental sulfur medium did not produce detectable amounts of sulfide (Supplementary table S3-5). Consequently, only coenzyme

M (Figure 3-4C) or thiosulfate (Figure 3-4E) supplementation restored the growth of mutant strain

S211 with elemental sulfur as sole sulfur source, but not with 3-mercaptopropionate acid (Figure

3-4D) or sulfite (Figure 3-4F). 95

All together, these results suggest that in M. maripaludis, MMP1681 is essential to sustain robust physiological functions when sulfide is the sole sulfur source and is required for assimilating elemental sulfur when provided as the sole sulfur source. Additionally, coenzyme M and thiosulfate, may serve as key intermediates within this MMP1681-driven elemental sulfur assimilation process. One tentative interpretation of these observations is that MMP1681 converts elemental sulfur into key sulfur species which serve as a sulfur donor for biosynthesizing the downstream metabolites. Thiosulfate is likely to stand as one of these sulfur species formed from elemental sulfur and may subsequently be incorporated into coenzyme M or other core metabolites.

Substrate specificity and catalytic constant of MMP1681 and MMP0941. Although

MMP1681 and MMP0941 are annotated as PAPR in M. maripaludis, homology search indicated that they are homologous to the characterized APR (MJ0973) (34) and PAPR (MJ0066) (35) from

M. jannaschii, respectively. As a result, these two proteins, together with CysH, a known PAPR from E. coli were heterologously expressed in E. coli, respectively. The substrate specificity of these purified recombinant proteins for either sulfonucleotide was investigated, in combination with glutaredoxin as the electron donor under steady-state conditions. The results are summarized in Table 3-3. Aerobically purified recombinant MMP1681 was catalytically active with APS but

-1 not PAPS, with a Km of 112.1 µM and Vmax of 0.03 µmoL·min per mg protein. However, recombinant MMP0941 was catalytically active with PAPS but not APS, with a Km of 36.5 µM

-1 and Vmax of 0.01 µmoL·min per mg protein. In comparison, the E. coli recombinant PAPR tested

-1 in this study possesses an activity with Km of 31.6 µM and Vmax of 0.04 µmoL·min per mg protein.

Here the Vmax value of the tested E. coli recombinant PAPR is ~100 fold less than that of the same enzyme previously reported (65). This is reasonable because, in this study, only 0.45 µM

glutaredoxin was utilized as the electron donor in the enzymatic assay, comparing to that of 50 µM glutaredoxin used earlier (65). Nonetheless, the Km of this tested E. coli recombinant PAPR was very close to the value previously reported (65).

These findings revealed that MMP1681 and MMP0941 indeed were sulfonucleotide reductase enzymes in vitro. In recombinant MMP0941, its affinity toward PAPS was similar to that in E. coli recombinant PAPR. Additionally, recombinant MMP1681 possessed only half of the affinity for sulfonucleotide comparing to that of E. coli recombinant PAPR, though toward

APS as substrate instead of PAPS. The catalytic efficiency of recombinant MMP0941 (180 M-1·s-

1) was one-third of that in E. coli recombinant PAPR (580 M-1·s-1) determined under the same conditions. In comparison, the catalytic efficiency of recombinant MMP1681 (230 M-1·s-1) was similar to that of the recombinant MMP0941.

MMP1681 and MMP0941 are iron-sulfur cluster proteins. The recombinant MMP1681 and MMP0941 proteins were both colored dark-brown when purified either aerobically or anaerobically. In both proteins, UV-visible absorption spectroscopy showed a maximum absorption for the polypeptide at 280 nm. In addition, there was a broad absorption feature between

320 nm and 500 nm, with a shoulder at 420 nm (Figure 3-5). This observation was similar to that of previously reported APR from P. aeruginosa (24), E. intestinalis (23), B. subtilis (25), Lemna minor (26) and A. thaliana (26), which were known to contain a single [4Fe-4S] cluster. Moreover, non-heme iron analyses of both freshly prepared recombinant proteins indicate the presence of

1.43 ± 0.01 mol of iron per mole of MMP1681 and 2.07 ± 0.03 mol of iron per mole of MMP0941.

These results suggest the occurrence of an iron-sulfur cluster on both MMP1681 and MMP0941.

The stability of recombinant MMP1681 and MMP0941 was subsequently tested by exposing the

anaerobically purified proteins to air for 24 hours. The absorbance of both recombinant proteins decreased upon aerobic conditions, suggesting that the iron-sulfur cluster was oxygen-sensitive

(Supplementary figure S3-7). Moreover, addition of 5 mM sodium dithionite also resulted in a quench of the absorbance in the entire visible region, indicating that the iron-sulfur cluster was redox-active (Figure 3-5). To further characterize the stoichiometry of these iron-sulfur clusters on either recombinant protein, the Electron Paramagnetic Resonance (EPR) spectroscopy analysis is ongoing.

Putative M. maripaludis sulfate assimilation homologs do not complement the corresponding E. coli mutants. To explore if the assimilatory sulfate reduction pathway existed in M. maripaludis, the homologs of involved enzymes were first examined. They are ATP sulfurylase (CysD), ATP kinase (CysC), APS reductase, PAPS reductase (CysH) and sulfite reductase (CysJ, CysG, CysI) (Supplementary figure 3-8). In M. maripaludis, MMP1471 is annotated as a nucleotidyltransferase, but a short piece (~ 40 amino acids) of ATP sulfurylase motif was identified according to the KEGG annotation. MMP1282 is annotated as a nucleotide kinase. MMP1681 and MMP0941 are homologs of the known APR (MJ0941) and PAPR

(MJ0066), respectively. MMP0078 has been described as a small sulfite reductase (41). These M. maripaludis proteins show only partial homology but are highly conserved in the key regions involved in catalysis with known proteins evolved in sulfate metabolism. Hence the role of these

M. maripaludis proteins remained ambiguous, and they might catalyze one or more steps in metabolizing sulfate into sulfide. To test the role of the methanococcal homologs in sulfate assimilation, it was tested if any could complement the corresponding mutants in E. coli.

Meanwhile, it is equally possible that the M. mariplaudis proteins constitute a new branch of

enzymes using different catalytic mechanisms than the known enzymes. Thus, they may not operate in bacterial host strains. To examine if these M. maripaludis proteins are functional in E. coli mutants deficient in sulfate assimilation. Mmp1471, mmp1282, mmp0941, mmp1681 and mmp0078 was expressed in shuttle vector pQE2, respectively. Each of these five recombinant proteins was then tested whether or not they complement any of ∆cysD, ∆cysC, ∆cysH, ∆cysJ,

∆cysG and ∆cysI E. coli Keio knockout mutants (56). E. coli PAPR was used as a positive control, expressed in the same plasmid pQE2 and performed the same complementation test on the six E. coli mutants. None of the M. maripaludis proteins complemented any of the E. coli mutants deficient in sulfate metabolism, while E. coli PAPR complemented only the ∆cysH mutant. These results indicated that the tested M. maripaludis proteins were not functional in E. coli mutants that are deficient in sulfate assimilation reduction pathway.

M. maripaludis doesn’t produce large amount of sulfate when grown with sulfide as the sole sulfur source. The native habit environment of M. maripaludis, marine sediment, usually contains millimolar concentrations of sulfate. Laboratory medium for cultivating M. maripaludis commonly contains 14 mM sodium sulfate to create a similar environment. To examine if M. maripaludis produces sulfate, the cells were grown under formate medium without sulfate addition with 2 mM sulfide as sulfur source and 2 mM dithiothreitol as the reducing agent. In this sulfate- free formate medium, the growth yield of M. maripaludis was not affected, although the lag phase is slightly elongated compared to the presence of sulfate addition (Supplemental figure S3-5). The sulfate-free medium contained no detectable sulfate (Table 3-5). Meanwhile, the sulfate levels in all medium ingredients was also examined. None of the medium components brought any detectable sulfate into the medium (Supplementary table S3-6). The inoculum contained about 41

µM contaminating sulfate (Table 3-5). Nonetheless, only 39 µM sulfate was detected in the supernatant of a stationary phase culture of M. maripaludis (Table 3-5). These results indicate that

M. maripaludis does not produce large amounts of sulfate during growth.

Sequence analysis of MMP1681 and MMP0941. To evaluate the sequence homolog of

APR and PAPR among Archaea, the sequence of MMP1681 and MMP0941 were used to retrieve related sequences in Archaea from the NCBI database using the BLAST program. Remarkably, almost all Achaea possesses two sulfonucleotide reductases. They were annotated as either an APR and PAPR or both as PAPRs. In the currently characterized APRs and PAPRs from plant, fungi, and bacteria, a highly conserved motif (KRT)ECG(LI)H containing a catalytically active cysteine residue is commonly present. However, this key motif is absent in all archaeal homologs. The homology of these archaeal proteins emerges from a segment of ~ 200 amino acids that were 22-

27 % identical with both plant APRs and E. coli PAPR. This domain is preceded by 150-200 amino acids that lacks homolog to other proteins (Figure 3-2B). These distinctive structural features separate the archaeal sulfonucleotide reductases from that of plant, fungi and bacteria.

To further examine the sequence homolog of the sulfonucleotide reductases within the

Archaea domain, an amino sequence alignment with these proteins from

Euryarchaeota and Crenarchaeota species, together with the in vitro characterized M. maripaludis homologs in this study, and the in vitro characterized M. jannaschii homologs previously (34, 35), were constructed. Interestingly, three conserved cysteine residues were found in all tested archaeal proteins (Figure 3-6). Almost all characterized plant and bacterial APR possesses four conserved cysteine residues near their N-termini. Three of these four cysteine residues form into two cysteine pairs and become the binding sites to iron-sulfur clusters (20-26).

100

In M. jannaschii, MJ0066, a PAPR characterized in vitro, this first conserved cysteine residue (M. jannaschii Cys337), together with another two cysteines (M. jannaschii Cys19 and Cys22) that were not conserved in this sequence analysis, were found to be required for the enzyme activity (35).

On the other hand, this study has also shown that the last two cysteine residues (M. jannaschii

Cys19 and Cys22) were not required PAPR activity in vitro (35). It is possible that these cysteinyl residues participate in the formation of the iron-sulfur clusters detected in the recombinant

MMP1681 and MMP0941.

Discussion

It is generally known that the growth of M. maripaludis on elemental sulfur was slower than that on sulfide. We presented here that M. maripaludis produced sulfide in a total of ~ 6 mmol per g cell dry weight per hour, which was much lower than previously considered. Therefore, it is very likely that the sulfide formation from element sulfur is very slow, which further delays the cells’ growth on elemental sulfur. In this study, we also demonstrated that protein MMP1681 in

M. maripaludis, is required for elemental sulfur assimilation in that organism. A putative role of this protein is to mobilize elemental sulfur into metabolic precursors of coenzyme M. This proposed function is supported by the following experiments. First, proteomics study showed that the expression level of MMP1681 increased 3.3-fold in cells grown on elemental sulfur. Second, markerless deletion of mmp1681 has resulted in the mutant unable to grow with elemental sulfur as the sole source, while complementing MMP1681 back to the mutant restored growth to wild- type levels. Lastly, supplementing coenzyme M or and thiosulfate to the Δmmp1681 culture grown with elemental sulfur as sole sulfur source also fully recovered its growth to wild-type levels.

101

The catalytic function of MMP1681 from M. maripaludis was subsequently studied. In vitro activity of the purified recombinant MMP1681 exhibited an ability to catalyze the production sulfite from APS, but not PAPS. In addition, the enzymatic activity was stimulated by glutaredoxin, but it may not be solely dependent on this electron donor. Thioredoxin, as a more relevant physiological relevant reductant in methanogens (66, 67), is currently being tested as an alternative electron donor in this assay. The affinity of recombinant MMP1681 for APS (Km=112 µM) was highly comparable with that of the Bacillus subtilis (Km=105 µM) enzyme (25) when using glutaredoxin as the reductant. In contrast, the plant APRs, such as in Arabidopsis thaliana (APR2)

(68) and Physcomitrella patens (68), have a twenty-fold higher affinity to APS (Km=6 µM) when using thioredoxin as the reductant. The Vmax of recombinant MMP1681 (Vmax=0.03 µmol per mg per minute) was similar to that in E. coli PAPR tested under similar conditions (Vmax=0.04 µmol per mg per minute). However, the Vmax value of recombinant MMP1681 is about 20-fold less of that in B. subtilis (Vmax=0.63 µmol per mg per minute) (25), and 1500-fold of that in plant APR such as in A. thaliana (APR2) (Vmax=45 µmol per mg per minute) (68), P. patens (Vmax=45 µmol per mg per minute) (68). This is not unexpected because, in the enzymatic assay of this study, only

0.45 µM glutaredoxin was used as reductant, comparing to 180 µM glutaredoxin used in Bacillus subtilis (25), and 0.85 µM thioredoxin used in Arabidopsis thaliana (68) and Physcomitrella patens (68). Because the APR or PAPR from different species have varying efficiencies with commercial thioredoxin and glutaredoxin, where the enzyme activity is in a linear relationship to the concentration of thioredoxin (0.5 µM-50 µM) or glutaredoxin (0.5 µM-50 µM) (25, 55, 65,

68). Interestingly, MMP0941, another sulfonucleotide reductase, also exists in M. maripaludis. In vitro enzymological evidence indicated that this enzyme produces sulfite from PAPS, but not APS.

The affinity for PAPS (Km=36.5 µM) was comparable with that of the E. coli (Km=32.6 µM), M. 102

jannaschii (Km=15.9 µM) (35) and B. subtilis (Km=10.7 µM) PAPRs (25). The Vmax of recombinant

MMP0941 (Vmax=0.01 µmol per mg per minute) was a quarter of that of E. coli PAPR (Vmax=0.04

µmol per mg per minute) tested under the same conditions. On the other hand, it is possible that the presence of iron-sulfur clusters is essential for the enzymatic activity of recombinant

MMP1681 and MMP0941, as seen in plant APRs. Because the iron-sulfur clusters on recombinant

MMP0941 and MMP1681 were lost upon oxygen exposure (Supplementary figure S3-7),

APR/PAPR enzymatic assays with anaerobically purified proteins are currently being performed to collect more kinetics data.

The catalytic function of MMP1681 and MMP0941 in vitro has brought into the question whether a sulfate assimilation pathway occurs in M. maripaludis. That is, if the M. maripaludis produces sulfate from sulfide, via the reverse reaction catalyzed by these two proteins. However, a quantitative measurement of the sulfate presence in M. maripaludis culture indicates this organism doesn’t produce large amounts of sulfate. Nonetheless, it cannot exclude the possibility that the sulfate production in M. maripaludis was too small to be detected. Subsequently, a collection of putative M. maripaluds sulfate assimilation protein homologs were gathered based on BLAST research. They are MMP1471, MMP1282, MMP0941, MMP1681 and MMP0078, which may represent ATP sulfurylase, ATP kinase, PAPR, APR and sulfite reductase, respectively.

However, none of these five proteins was able to complement any E. coli mutants for these activities. This is surprising because even MMP0941 or MMP1681 with sulfonucleotide reductase activity in vitro, cannot complement the E. coli ΔcysH mutant. These two experiments together suggest that the sulfate assimilation pathway may not exist in M. maripaludis. On the other hand, considering the complexity of its habitat environment and unique sulfur metabolic features (69) of

103

M. maripaludis, it may possess a unique set of metabolic intermediates for intracellular sulfur species trafficking comparing to E. coli, thus leading the malfunction of M. maripaludis proteins in E. coli mutants.

The co-existence of both APR and PAPR in M. maripaludis and other methanogens are archaea is very interesting and remains enigmatic. CysH1 from B. subtilis possesses both APR and

PAPR enzymatic activity and contains a [4Fe-4S] cluster (25). But generally most other organisms possess only one sulfonucleotide reductase. Additionally, an iron-sulfur cluster was identified on both MMP1681 and MMP0941. This finding is particularly striking, considering that iron-sulfur clusters are absent in the PAPRs from fungi, yeast, and enteric bacteria, (20-26). In contrast, all sulfonucleotide reductases using APS analyzed to date contain the iron-sulfur cluster, including plant APR, and assimilatory and dissimilatory bacterial APR (20). The function of the iron-sulfur cluster in the reaction mechanism of assimilatory APRs is not known yet, but current evidence suggests that it is likely to be associated with the reduction of APS (20). Therefore, the presence of an iron-sulfur cluster in MMP0941, possessing PAPR activity in vitro, calls to question the physiological function of this protein. Lastly, the sequence analysis revealed that MMP1681,

MMP0941, and their archaeal homologs possess different structural aspects from all known APRs and PAPRs. Hence, it is likely these archaeal proteins operate in novel functions, other than as sulfonucleotide reductase in vitro.

Based on the data collected in this study, we here propose a scheme of elemental sulfur assimilation process in M. maripaludis (Figure 3-7). In this scenario, elemental sulfur reacts with a thiol, such as coenzyme M or thiosulfate, to form a persulfide before being taken up into the cell.

After this, elemental sulfur is assimilated into the M. maripaludis in the form of sulfane sulfur on

104

this persulfide group. This sulfane sulfur is further reduced to sulfide by an unknown electron donor. The sulfide then can be utilized as a sulfur donor for synthesizing other sulfur-containing compounds. The mechanism of MMP1681 remains to be elucidated, one possible role is to convert an unknown substrate into sulfite, which is a required sulfur donor for biosynthesizing coenzyme

Taken together, a few outstanding questions on sulfur metabolism in M. maripaludis remain to be elucidated. For instance: (i) The explicit mechanism by which MMP1681 assimilates elemental sulfur and how it links to coenzyme M biosynthesis; (ii) The physiological role of the sulfate assimilation homologs, including MMP0941, MMP1471, MMP1282, and MMP0078.

Nevertheless, the evidence reported here undoubtedly broadens the perception of how methanogens possessing MMP1681 assimilate elemental sulfur into key sulfur intermediates.

References: 1. Cotton FA, Wilkinson G. 1999. Advanced inorganic chemistry, 6th ed. Wiley, New York.

2. Belkin S, Wirsen CO, Jannasch HW. 1985. Biological and abiological sulfur reduction at

high temperatures. Appl Environ Microbiol 49:1057-1061.

3. Boulegue J. 1978. Solubility of elemental sulfur in water at 298 K. Phosphorus and Sulfur

and the Related Elements 5:127-128.

4. Fauque GD, Barton LL. 2012. Hemoproteins in dissimilatory sulfate- and sulfur-reducing

prokaryotes. Adv Microb Physiol Vol 60 60:1-90.

5. Rabus R, Hansen TA, Widdel F. 2006. Dissimilatory sulfate- and sulfur-reducing

prokaryotes, p 659-768. In Dworkin M, Falkow S, Rosenberg E, Schleifer K-H,

105

Stackebrandt E (ed), The Prokaryotes: Volume 2: Ecophysiology and Biochemistry

doi:10.1007/0-387-30742-7_22. Springer New York, New York, NY.

6. Cammack R, Fauque G, Moura JJG, Legall J. 1984. ESR studies of cytochrome c3 from

Desulfovibrio desulfuricans strain Norway 4: Midpoint potentials of the four haems, and

interactions with ferredoxin and colloidal sulphur. Biochimica Et Biophysica Acta 784:68-

74.

7. Hedderich R, Klimmek O, Kröger A, Dirmeier R, Keller M, Stetter KO. 1998. Anaerobic

respiration with elemental sulfur and with disulfides. FEMS Microbiol Rev 22:353-381.

8. Steudel R, Holdt G, Nagorka R. 1986. Sulfur-compounds. 104. On the autooxidation of

aqueous sodium polysulfide Z. Naturforsch. 41:1519-1522.

9. Kitamura M, Sagara T, Taniguchi M, Ashida M, Ezoe K, Kohno K, Kojima S, Ozawa K,

Akutsu H, Kumagai I, Nakaya T. 1998. Cloning and expression of the gene encoding

flavodoxin from Desulfovibrio vulgaris (Miyazaki F). J Biochem. 123:891-898.

10. Thamdrup B, Finster K, Hansen JW, Bak F. 1993. Bacterial disproportionation of

elemental sulfur coupled to chemical reduction of iron or manganese. Appl Environ

Microbiol 59:101-8.

11. Schauder R, Kroger A. 1993. Bacterial Sulfur Respiration. Arch Microbiol 159:491-497.

12. Fauque GD. 1994. Sulfur reductase from rhiophilic sulfate-reducing bacteria. Inorganic

Microbial Sulfur Metabolism 243:353-367.

13. Fauque GD, Klimmek O, Kroger A. 1994. Sulfur reductases from spirilloid mesophilic

sulfur-reducing eubacteria. Inorganic Microbial Sulfur Metabolism 243:367-383.

14. Finster K. 2008. Microbiological disproportionation of inorganic sulfur compounds. J

Sulphur Chem 29:281-292. 106

15. Stetter KO, Gaag G. 1983. Reduction of molecular sulfur by methanogenic bacteria. Nature

305:309-311.

16. Lefaou A, Rajagopal BS, Daniels L, Fauque G. 1990. Thiosulfate, polythionates and

elemental sulfur assimilation and reduction in the bacterial world. FEMS Microbiol Lett

75:351-382.

17. Obraztsova AY, Francis CA, Tebo BM. 2002. Sulfur disproportionation by the facultative

anaerobe Pantoea agglomerans SP1 as a mechanism for chromium(VI) reduction.

Geomicrobiol J 19:121-132.

18. Bolliger C, Schroth MH, Bernasconi SM, Kleikemper J, Zeyer J. 2001. Sulfur isotope

fractionation during microbial sulfate reduction by toluene-degrading bacteria.

Geochimica Et Cosmochimica Acta 65:3289-3298.

19. Sekowska A, Kung HF, Danchin A. 2000. Sulfur metabolism in Escherichia coli and

related bacteria: facts and fiction. J Mol Microbiol Biotechnol 2:145-77.

20. Kopriva S, Koprivova A. 2004. Plant adenosine 5'-phosphosulphate reductase: the past, the

present, and the future. J Exp Bot 55:1775-83.

21. Kopriva S, Buchert T, Fritz G, Suter M, Benda RD, Schunemann V, Koprivova A,

Schurmann P, Trautwein AX, Kroneck PMH, Brunold C. 2002. The presence of an iron-

sulfur cluster in adenosine 5'-phosphosulfate reductase separates organisms utilizing

adenosine 5'-phosphosulfate and phosphoadenosine 5'-phosphosulfate for sulfate

assimilation. J Biol Chem 277:21786-21791.

22. Carroll KS, Gao H, Chen HY, Leary JA, Bertozzi CR. 2005. Investigation of the iron-sulfur

cluster in Mycobacterium tuberculosis APS reductase: Implications for substrate binding

and catalysis. Biochemistry 44:14647-14657. 107

23. Kim SK, Rahman A, Conover RC, Johnson MK, Mason JT, Gomes V, Hirasawa M, Moore

ML, Leustek T, Knaff DB. 2006. Properties of the cysteine residues and the iron-sulfur

cluster of the assimilatory 5'-adenylyl sulfate reductase from Enteromorpha intestinalis.

Biochemistry 45:5010-5018.

24. Kim SK, Rahman A, Bick JA, Conover RC, Johnson MK, Mason JT, Hirasawa M, Leustek

T, Knaff DB. 2004. Properties of the cysteine residues and iron-sulfur cluster of the

assimilatory 5'-adenylyl sulfate reductase from Pseudomonas aeruginosa. Biochemistry

43:13478-13486.

25. Berndt C, Lillig CH, Wollenberg M, Bill E, Mansilla MC, de Mendoza D, Seidler A,

Schwenn JD. 2004. Characterization and reconstitution of a 4Fe-4S adenylyl

sulfate/phosphoadenylyl sulfate reductase from Bacillus subtilis. J Biol Chem 279:7850-

7855.

26. Kopriva S, Buchert T, Fritz G, Suter M, Weber M, Benda R, Schaller J, Feller U,

Schurmann P, Schunemann V, Trautwein AX, Kroneck PMH, Brunold C. 2001. Plant

adenosine 5'-phospho sulfate reductase is a novel iron-sulfur protein. J Biol Chem

276:42881-42886.

27. Carroll KS, Gao H, Chen H, Leary JA, Bertozzi CR. 2005. Investigation of the iron-sulfur

cluster in Mycobacterium tuberculosis APS reductase: implications for substrate binding

and catalysis. Biochemistry 44:14647-57.

28. Prior A, Uhrig JF, Heins L, Wiesmann A, Lillig CH, Stoltze C, Soll J, Schwenn JD. 1999.

Structural and kinetic properties of adenylyl sulfate reductase from Catharanthus roseus

cell cultures. Biochim Biophys Acta 1430:25-38.

108

29. Bick JA, Aslund F, Chen YC, Leustek T. 1998. Glutaredoxin function for the carboxyl-

terminal domain of the plant-type 5'-adenylylsulfate reductase. Proc Natl Acad Sci U S A

95:8404-8409.

30. Weber M, Suter M, Brunold C, Kopriva S. 2000. Sulfate assimilation in higher plants

characterization of a stable intermediate in the adenosine 5'-phosphosulfate reductase

reaction. Eur J Biochem 267:3647-3653.

31. Daniels L, Belay N, Rajagopal BS. 1986. Assimilatory feduction of sulfate and sulfite by

methanogenic bacteria. Appl Environ Microbiol 51:703-709.

32. Rajagopal BS, Daniels L. 1986. Investigation of mercaptans, organic sulfides, and

inorganic sulfur compounds as sulfur sources for the growth of methanogenic bacteria.

Curr Microbiol 14:137-144.

33. Johnston DT. 2011. Multiple sulfur isotopes and the evolution of Earth's surface sulfur

cycle. Earth-Science Reviews 106:161-183.

34. Lee JS, White E, Kim SG, Schlesinger SR, Lee SY, Kim SK. 2011. Discovery of a novel

adenosine 5'-phosphosulfate (APS) reductase from the methanarcheon

Methanocaldococcus jannaschii. Process Biochem 46:154-161.

35. Cho MK. 2013. Discovery of novel 3'-phosphoadenosine-5'-phosphosulfate (PAPS)

reductase from methanarcheon Methanocaldococcus jannaschii. Master Thesis, Baylor

University.

36. Johnson EF, Mukhopadhyay B. 2005. A new type of sulfite reductase, a novel coenzyme

F420-dependent enzyme, from the methanarchaeon Methanocaldococcus jannaschii. J Biol

Chem 280:38776-38786.

109

37. Johnson EF, Mukhopadhyay B. 2008. Coenzyme F420-dependent sulfite reductase-enabled

sulfite detoxification and use of sulfite as a sole sulfur source by Methanococcus

maripaludis. Appl Environ Microbiol 74:3591-3595.

38. Balderston WL, Payne WJ. 1976. Inhibition of methanogenesis in salt-marsh sediments

and whole-cell suspensions of methanogenic bacteria by nitrogen-oxides. Appl Environ

Microbiol 32:264-269.

39. Becker DF, Ragsdale SW. 1998. Activation of methyl-SCoM reductase to high specific

activity after treatment of whole cells with sodium sulfide. Biochemistry 37:2639-2647.

40. Mahlert F, Bauer C, Jaun B, Thauer RK, Duin EC. 2002. The nickel enzyme methyl-

coenzyme M reductase from methanogenic archaea: In vitro induction of the nickel-based

MCR-ox EPR signals from MCR-red2. J Biol Inorg Chem 7:500-513.

41. Johnson EF, Mukhopadhyay B. 2008. A novel coenzyme F420 dependent sulfite reductase

and a small sulfite reductase in methanogenic archaea. Microbial Sulfur Metabolism

doi:Doi 10.1007/978-3-540-72682-1_16:202-216.

42. Walters AD, Smith SE, Chong JP. 2011. Shuttle vector system for Methanococcus

maripaludis with improved transformation efficiency. Appl Environ Microbiol 77:2549-

51.

43. Moore BC, Leigh JA. 2005. Markerless mutagenesis in Methanococcus maripaludis

demonstrates roles for alanine dehydrogenase, alanine racemase, and alanine permease. J

Bacteriol 187:972-9.

44. Sarmiento F, Leigh JA, Whitman WB. 2011. Genetic systems for hydrogenotrophic

methanogens. Methods Enzymol 494:43-73.

110

45. Long F, Wang L, Lupa B, Whitman WB. 2017. A flexible system for cultivation of

Methanococcus and other formate-utilizing methanogens. Archaea. 2017:12.

doi:10.1155/2017/7046026.

46. Zhou X, Meile L, Kreuzer M, Zeitz JO. 2013. The effect of saturated fatty acids on

methanogenesis and cell viability of Methanobrevibacter ruminantium. Archaea

2013:106916.

47. Liu Y, Sieprawska-Lupa M, Whitman WB, White RH. 2010. Cysteine is not the sulfur

source for iron-sulfur cluster and methionine biosynthesis in the methanogenic archaeon

Methanococcus maripaludis. J Biol Chem 285:31923-9.

48. Fogo JK, Popowsky M. 1949. Spectrophotometric determination of hydrogen sulfide. Anal

Biochem 21:732-734.

49. Beinert H. 1983. Semi-micro methods for analysis of labile sulfide and of labile sulfide

plus sulfane sulfur in unusually stable iron-sulfur proteins. Anal Biochem 131:373-8.

50. Miyamoto R, Otsuguro K, Yamaguchi S, Ito S. 2014. Contribution of cysteine

aminotransferase and mercaptopyruvate sulfurtransferase to hydrogen sulfide production

in peripheral neurons. J Neurochem 130:29-40.

51. Siegel LM. 1965. A Direct microdetermination for sulfide. Anal Biochem 11:126-132.

52. Wong RL, Amster IJ. 2006. Combining low and high mass ion accumulation for enhancing

shotgun proteome analysis by accurate mass measurement. J Am Soc Mass Spectrom

17:205-12.

53. Lyu Z, Chou CW, Shi H, Patel R, Ghebreab R, Duin EC, Whitman WB. Methanogenesis

marker 10 protein required for methylation of the arginine at the active site of methyl-

coenzyme M reductase of Methanococcus maripaludis. (Manuscript). 111

54. Lyu Z, Jain R, Smith P, Fetchko T, Yan Y, Whitman WB. 2016. Engineering the autotroph

Methanococcus maripaludis for geraniol production. ACS Synth Biol 5:577-81.

55. Tsang MLS. 1981. Assimilatory sulfate reduction in Escherichia-Coli : identification of

the alternate cofactor for adenosine 3'-phosphate-5'-phosphosulfate reductase as

glutaredoxin. J Bacteriol 146:1059-1066.

56. Baba T, Ara T, Hasegawa M, Takai Y, Okumura Y, Baba M, Datsenko KA, Tomita M,

Wanner BL, Mori H. 2006. Construction of Escherichia coli K-12 in-frame, single-gene

knockout mutants: the Keio collection. Molecular systems biology 2.

57. Sheen R, Kahler H, Ross E, Betz W, Betz L. 1935. Turbidimetric determination of sulfate

in water. Industrial and Engineering Chemistry, Analytical Edition 7:262-265.

58. Kearse M, Moir R, Wilson A, Stones-Havas S, Cheung M, Sturrock S, Buxton S, Cooper

A, Markowitz S, Duran C, Thierer T, Ashton B, Meintjes P, Drummond A. 2012. Geneious

basic: an integrated and extendable desktop software platform for the organization and

analysis of sequence data. Bioinformatics 28:1647-9.

59. Franklin MJ, Wiebe WJ, Whitman WB. 1988. Populations of methanogenic bacteria in a

georgia salt marsh. Appl Environ Microbiol 54:1151-7.

60. Dybas M, Konisky J. 1992. Energy transduction in the methanogen Methanococcus voltae

is based on a sodium current. J Bacteriol 174:5575-5583.

61. Liu Y, Guo L, Guo R, Wong RL, Hernandez H, Hu J, Chu Y, Amster IJ, Whitman WB,

Huang L. 2009. The Sac10b homolog in Methanococcus maripaludis binds DNA at

specific sites. J Bacteriol 191:2315-2329.

112

62. Hendrickson EL, Haydock AK, Moore BC, Whitman WB, Leigh JA. 2007. Functionally

distinct genes regulated by hydrogen limitation and growth rate in methanogenic archaea.

Proc Natl Acad Sci U S A 104:8930-8934.

63. Ouhammouch M, Dewhurst RE, Hausner W, Thomm M, Geiduschek EP. 2003. Activation

of archaeal transcription by recruitment of the TATA-binding protein. Proc Natl Acad Sci

U S A 100:5097-5102.

64. Ouhammouch M, Langham GE, Hausner W, Simpson AJ, El-Sayed NMA, Geiduschek EP.

2005. Promoter architecture and response to a positive regulator of archaeal transcription.

Mol Microbiol56:625-637.

65. Lillig CH, Prior A, Schwenn JD, Aslund F, Ritz D, Vlamis-Gardikas A, Holmgren A. 1999.

New thioredoxins and glutaredoxins as electron donors of 3'-phosphoadenylylsulfate

reductase. J Biol Chem 274:7695-7698.

66. McCarver AC, Lessner DJ. 2014. Molecular characterization of the thioredoxin system

from Methanosarcina acetivorans. FEBS J 281:4598-4611.

67. Susanti D, Wong JH, Vensel WH, Loganathan U, DeSantis R, Schmitz RA, Balsera M,

Buchanan BB, Mukhopadhyay B. 2014. Thioredoxin targets fundamental processes in a

methane-producing archaeon, Methanocaldococcus jannaschii. Proc Natl Acad Sci U S A

111:2608-2613.

68. Kopriva S, Fritzemeier K, Wiedemann G, Reski R. 2007. The putative moss 3'-

phosphoadenosine-5'-phosphosulfate reductase is a novel form of adenosine-5'-

phosphosulfate reductase without an iron-sulfur cluster. J Biol Chem 282:22930-22938.

69. Liu YC, Beer LL, Whitman WB. 2012. Methanogens: a window into ancient sulfur

metabolism. Trends Microbiol 20:251-258. 113

70. Pereira IAC. 2008. Respiratory Membrane Complexes of Desulfovibrio, p 24-35. In Dahl

C, Friedrich CG (ed), Microbial sulfur metabolism doi:10.1007/978-3-540-72682-1_3.

Springer Berlin Heidelberg, Berlin, Heidelberg.

71. Santos AA, Venceslau SS, Grein F, Leavitt WD, Dahl C, Johnston DT, Pereira IA. 2015.

A protein trisulfide couples dissimilatory sulfate reduction to energy conservation. Science

350:1541-5.

72. Hendrickson EL, Kaul R, Zhou Y, Bovee D, Chapman P, Chung J, Conway de Macario E,

Dodsworth JA, Gillett W, Graham DE, Hackett M, Haydock AK, Kang A, Land ML, Levy

R, Lie TJ, Major TA, Moore BC, Porat I, Palmeiri A, Rouse G, Saenphimmachak C, Soll

D, Van Dien S, Wang T, Whitman WB, Xia Q, Zhang Y, Larimer FW, Olson MV, Leigh

JA. 2004. Complete genome sequence of the genetically tractable hydrogenotrophic

methanogen Methanococcus maripaludis. J Bacteriol 186:6956-69.

114

Figure 3-1. Scheme of assimilatory sulfate reduction and dissimilatory sulfate reduction pathways.

In dissimilatory sulfate reduction pathway, the Qmo complex refers to QmoABC, a-membrane- bound enzyme complex which is involved in electron transfer to the APS reductase (70). In the following step, DsrC serves as the electron donor for the sulfite reduction by sulfite reductase

(DsrAB) (71). The resulting oxidized DsrC is reduced by the membrane Dsr complex, DsrMKJOP

(71).

Abbreviations: APS: Adenosine-5’-phosphosulfate; ATP: Adenosine triphosphate; PPi: pyrophosphate; PAPS: 3'-Phosphoadenosine-5'-phosphosulfate; GSH: reduced glutathione; Grx: glutaredoxin; Trx: thioredoxin (Trx); GSSG: glutathione disulfide. 115

Figure 3-2. Domain organization (A) and cysteine conservation (B) with the sulfonucleotide reductase Family.

A: The composition of domain in sulfonucleotide reductase family including adenosine-5’- phosphosulfate reductase (APR) and 3'-phosphoadenosine-5'-phosphosulfate reductase (PAPR).

APR from higher plants possesses a reductase domain and a unique C-terminal domain with homology to thioredoxin. A conserved motif -CXXC- (brown) at the C-terminal of plant APR is known to responsible for electron transfer. Bacterial APR lacks this unique domain, but shares the cysteine motif –CC-X~80-CXXC- (blue) present in the reductase domain. PAPR lack this cysteine 116

motif as well as the thioredoxin domain. All suflonucletide reductases have a cysteine (red) at the end of the C terminus in the reductase domain. This residue is essential for catalysis.

B: Scheme of conserved cysteine residues on plant APR, assimilatory bacterial APR, PAPR,

MMP1681, MMP0941 and the archaeal homolog of MMP1681 and MMP0941. To collect the archaeal homolog of MMP1681 and MMP0941, their protein sequences were used as query sequences for BLAST searches in the NCBI sequence database, respectively. The resulted homologs in the Archaea domain were then collected and aligned by MUSCLE. A diagram of these archaeal homologs is drawn based on the alignment map. C indicates the presence of a cysteinyl residue. X represents a certain amino acid residue, number after X means the number of amino acids after this X residue. The position of C and X on scale bar correlates to their actual positions on the protein. Reductase domains are colored in yellow, while the thioredoxin-like domains are colored in grey. Cysteinyl residues in plant APR, assimilatory bacterial APR, PAPR are colored according to the description in A. Conserved cysteinyl residues in archaeal homolog of MMP1681 and MMP0941 are colored in purple.

117

Figure 3-3. Growth of S0001(●), S211 ()in McNAA medium reduced with 2 mM dithiothreitol.

The sulfur source, sodium sulfide was variated from 100 μM to 20 mM as indicated. The inoculum size was 1.5×105 cells per 5 mL culture. All values were the averages of three independent cultures.

118

Figure 3-4. Growth of S2 (black), mutant strain S211 (red) and the complementtion strain S212

(blue) in McNAA medium reduced with 3mM dithiothreitol, with either 2 mM sulfide (Figure 3-

4A) or 0.1g elemental sulfur per 5 mL culture (Figure 3-4B to F) as sole sulfur source. The elemental sulfur medium was supplemented with 0.1 mM coenzyme M (Figure 3-4C), or 0.1 mM

3-mercaptopropionate acid (Figure 3-4D), or 0.1 mM sodium thiosulfate (Figure 3-4E) or 0.1 mM sodium sulfite (Figure 3-4F). The inoculum size was 1.5×107 cells per 5 mL culture for cultures in

4A and 2.25×108 cells per 5 mL culture for cultures in figure 3-4B to F. All values were the averages of three independent cultures.

119

Figure 3-5. Absorbance spectrum of anaerobically purified recombinant MMP1681 (A) and

MMP0941 (B) in the absence (black) or presence (red) of 5 mM sodium dithionite. The recombinant MMP1681 was at 75 µM in 50 mM sodium HEPES (pH= 8.0) buffer with 500 mM

NaCl. The recombinant MMP0941 was at 92 µM in 50 mM sodium HEPES (pH= 8.0) buffer with

500 mM NaCl. These two recombinant proteins were over-expressed in E. coli by the pQE2 expression system (Qiagen) with N-terminal 6×His tag and anaerobically purified using gravity flow Ni-NTA column. Images on top right corner represent the actual appearance of anaerobically purified recombinant MMP1681 (A) and MMP0941 (B), respectively.

120

121

Figure 3-6. Sequence alignment of MMP1681, MMP0941 and their homologs in Archaeoglobus fulgidus, Methanothermobacter sp., Methanocaldococcus jannaschii, Methanopyrus kandleri,

Pyrococcus furiosus, Thermoplasmatales archeaon and Staphylothermus marinus. The accession numbers of the protein sequences used are as follows (from top to bottom of the alignment):

KUK07756.1, WP_048095118.1, WP_048176530.1, BAM69678.1, WP_011170885.1,

WP_011171625.1, WP_064496383.1, WP_010870487.1, AAM01836.1, AAM01462.1,

WP_014835526.1, AAL82076.1, WP_015492226.1, WP_015492786.1,

WP_011839296.1. WP_011838700.1. The first protein of each organism is the homolog of

MMP1681, while the second protein of each organism is the homolog of MMP0941. The conserved cysteinyl residues mentioned in the text were indicated with a dot above the sequence.

122

Figure 3-7. A proposed scheme of the sulfur assimilation and trafficking in methanococci.

123

Table 3-1. Inorganic sulfide levels in Methanococcus maripaludis cultures grown with elemental sulfur (S0) as sulfur source.

d d Sample H2S (aq) (μM) H2S(g) (μM per liter culture) Mediuma+TiC+S0 < 1.5 < 1.5 Mediuma +TiC+Inoculum 6.4 ± 1.0 < 1.5 Mediuma +TiC+S0+Inoculum 9.7 ± 0.8 < 1.5 Culture broth following growthb 78.2 ± 11.2 80.5 ± 6.3

Mediuma +DTT+S0 172.0 ± 1.3 125.3 ± 19.2 Mediuma +DTT+Inoculum 8.0 ± 0.8 <1.5 Mediuma +DTT+S0+Inoculum 185.1 ± 13.5 125.7 ± 12.8 Culture broth following growthc 502.1 ± 27.1 494.8 ± 36.5 a McNAA medium was McNA medium supplemented with 1 mM alanine, reduced with 2 mM titanium (III) citrate (TiC) or 2 mM dithiothreitol (DTT), together with or without 0.1 g elemental sulfur per 5 mL medium. McNAA medium reduced with titanium (III) citrate, they were incubated at 37 °C for 24 h before measuring the inorganic sulfide content. McNAA medium reduced with dithiothreitol, they were incubated at 37 °C for 16 hr before measuring the inorganic sulfide content b Culture was grown in 5 mL of McFAA medium (reduced with Tic) to an absorbance of 0.8 at

600 nm after 24 h of incubation at 37 °C. c Culture was grown in 5 mL of McFAA medium (reduced with DTT) to an absorbance of 0.8 at

600 nm after 16 h of incubation at 37 °C. d Values were means ± 1 S.D., obtained from three independent samples and represent inorganic sulfide concentration either in the aqueous phase (aq) or gaseous phase (g).

124

Table 3-2. Changes in expression of certain proteins in the Methanococcus maripaludis S2 determined by quantitative proteomic analysisa.

Protein ratio ORF nd Annotationf Meane SE Up regulatedb MMP1684 1 3.77 N.A. Conserved hypothetical protein H -forming N5, N10-methylene- MMP0127 1 3.50 N.A. 2 tetrahydromethanopterin dehydrogenease MMP1212 1 3.30 N.A. Putative thiolase/acetyl-CoA acetyltransferase MMP1681 1 3.27 N.A. Phosphoadenosine phosphosulfate reductase Formylmethanofuran dehydrogenase subunit erelated MMP0965 1 3.12 N.A. protein Lrp-Like Transcriptional regulatory proteins, AsnC MMP1137 1 2.79 N.A. family MMP0283 1 2.67 N.A. Nucleoside-diphosphate kinase MMP0720 1 2.35 N.A. Prismane MMP1410 1 2.29 N.A. LSU ribosomal protein L24P ATP/GTP-binding site motif A (P-loop):small GTP- MMP1642 1 2.18 N.A. binding protein domain

Down regulatedc MMP1385 2 0.27 1.24 Coenzyme F420-reducing hydrogenase subunit beta MMP0831 1 0.30 N.A. Uroporphyrinogen decarboxylase (URO-D) MMP1297 2 0.32 0.18 Formate dehydrogenase beta subunit F -dependent methylenetetrahydromethanopterin MMP0372 7 0.33 0.64 420 dehydrogenase MMP1496 1 0.33 N.A. Phenylalanyl-tRNA synthetase alpha subunit MMP0853 2 0.33 0.15 Nitrogenase iron protein (nitrogenase component II) MMP1299 4 0.34 0.36 Carbonic anhydrase MMP1573 2 0.35 0.24 Dethiobiotin synthetase MMP1298 7 0.36 0.31 Formate dehydrogenase alpha subunit MMP1382 4 0.37 0.10 Coenzyme F420-reducing hydrogenase subunit alpha MMP0103 1 0.40 N.A. Conserved hypothetical protein MMP0794 1 0.45 N.A. Hypothetical protein MMP0011 2 0.46 1.28 Restriction endonuclease subunit M a The cultures for proteomic analysis were grown in McNA medium. b The top 10 up-regulated ORFs among the 269 peptides detected in the proteomic analysis, in a descending order.

125

c The top 10 down-regulated ORFs among the 269 peptides detected in the proteomic analysis, in a descending order. d Number of proteome measurements for the ORF. e Mean of the ratio of protein levels during growth on elemental sulfur to growth on sulfide. f From reference (72).

126

Table 3-3. Sulfonucleotide reductase assays and kinetic constants. APR and PAPR activities were measured with purified recombinant E. coli PAPR, MMP1681 (from Methanococcus maripaludis) and MMP0941 (from Methanococcus maripaludis) as the consumption of NADPH from varying concentration of APS and PAPS in the presence of glutaredoxin (Grx) and reduced glutathione

(GSH). Values are the means of three independent experiments.

V Recombinant K max k k /K Substrate Electron donor m µmol·mg- cat cat m Protein µM s-1 M-1·s-1 1·min-1 E. coli PAPR PAPS 0.45 µM Grx+7 mM GSH 32.6 0.04 0.02 580 MMP1681 APS 0.45 µM Grx+7 mM GSH 112 0.03 0.03 231 MMP0941 PAPS 0.45 µM Grx+7 mM GSH 37 0.01 0.01 180 MjAPRa APS 1 mM Trx 0.29 0.08 0.09 300,000 MjPAPRb PAPS 1 mM Trx 16 0.09 0.09 5,570 BsAPRc APS 180 µM Grx+10 mM GSH 105 0.63 0.34 3,200 BsPAPRc PAPS 180 µM Grx+10 mM GSH 11 0.71 0.38 35,700 E. coli PAPRd PAPS 50 µM Grx+25 mM GSH 15 5 2.33 155,000 a Activities of MjAPR (from Methanocaldococcus jannaschii) were measured in reference (34). b Activities of MjPAPR (from Methanocaldococcus jannaschii) were measured in reference (35). c Activities of BsAPR and BsPAPR (from Bacillus subtilis) were measured in reference (25). d Activities of E. coli PAPR were measured in reference (65).

127

Table 3-4. The levels of inorganic sulfate in Methanococcus maripaludis cultures grown in sulfate-depleted medium with sulfide as sulfur source.

Sample Sulfate conc. (µM) d Mediuma N.D. a b Medium +2 mM Na2S N.D. Mediuma +Inoculumc 41.1 ± 0.6 a b c Medium +2 mM Na2S +Inoculum 36.1 ± 0.7 Culture broth following growth 39.6 ± 0.9 a Sulfate-free McFAA medium was used to grow M. maripaludis. In this medium, sulfate- containing chemicals in the general salts solution, trace minerals solution and iron solution were replaced with the corresponding chemical in chloride salts. b 2 mM sodium sulfide as sulfur source was added into above medium when cultivating M. maripaludis. c An inoculum of 2.5×107 M. maripaludis cells, in a volume of 0.1 mL, was added in the medium.The inoculum was grown in sulfate-depleted McFAA medium and carried over sodium sulfide of 40 µM into the transferred medium. d Values are the means of three independent samples ± S.D.

N.D.: Not detectable.

128

CHAPTER 4

EXPLORING THE USE OF GENOME COMPARISONS OF WILD-TYPE AND

RESISTANT MUTANTS OF METHANOCOCCUS MARIPALUDIS TO IDENTIFY

POTENTIAL TARGETS OF INHIBITORY COMPOUNDS3

3 Long, F., J. Cheung, W.B. Whitman, R.S. Ronimus and G.M. Cook. To be submitted to Applied and Environmental Microbiology.

129

Abstract

Phenotypic screening is routinely used by the pharmaceutical industry with a high level of success for the high-throughput identification of novel inhibitory compounds to control bacterial pathogens. In the process of developing pharmaceuticals, one of the next steps is to identify the target(s) of the testing compounds for studying the resistant mechanism, thus determining the compound derivatives and their enzyme structures. Methane emissions from enteric fermentation in the ruminant digestive system, primarily operated by methanogenic archaea, are a significant contributor to anthropogenic greenhouse gas emissions. Additionally, methane produced as an end product of enteric fermentation is an energy loss from digested feed intake. To control the methane emissions from ruminants, extensive research in the last decades has been focused on developing viable enteric methane mitigation practices, particularly, using methanogen-specific inhibitors. We report here the utilization of two known inhibitors of methanogenic archaea, neomycin, and chloroform, together with a recently identified inhibitor, echinomycin, to produce resistant mutants of Methanococcus maripaludis S2 and S0001. Whole-genome sequencing at high coverage (>

100-fold) was performed subsequently to investigate the potential targets of these inhibitors at the genomic level. Upon analysis of the whole-genome sequencing data, a list of shared mutations that exist in each type of resistant mutants were collected and some may be associated with the resistant mechanisms. Interestingly, several unexpected mutations were also mapped on the re-sequenced

M. maripaludis wild-type strain S2 and S0001, which provided fundamental genomic information for future studies on these organisms. This work has presented the application of whole-genome sequencing of resistant mutants as a viable approach to decipher the underlying molecular mechanisms of methanogen inhibitors.

130

Introduction

Methane is a high potential global warming gas whose concentration in the atmosphere has increased from less than 0.6 to 1.8 ppm since the last century (1). Enteric fermentation of feed by ruminants (cattle, sheep, goats) constitutes one of the main anthropogenic sources of atmospheric methane (2). Ruminants emit about 100 million tons of methane per year, approximately ∼20 % of global methane emissions (3). In the intestine of domestic ruminants, methane is produced by methanogenic archaea mainly using CO2 and H2, then escapes into the atmosphere by eructation and breathing of the animals (1). Moreover, the formation of methane reduces the feed efficacy to livestock, resulting in a loss of 12 % of the gross energy ingested by the animal (4). Over the last decades, a number of viable enteric methane mitigation practices has been extensively studied (5-

9), such as alternative forages (10), animal breeding (11), plant secondary products (12), control of protozoal populations (8, 13), vaccines (14) and alternative electron acceptors to divert H2 from methanogenesis and chemical inhibitors (5, 6, 15). The utilization of methanogen inhibitors as mitigation agents, particularly chemical compounds with inhibitory effects on methanogenic archaea, such as bromochloromethane, 2-bromoethane sulfonate, chloroform, and cyclodextrin, have been extensively studied in various ruminant species recently (2). Long-term in vivo experiments have reported up to 60 % of inhibition on methanogenesis by bromochloromethane

(2, 16). However, the effects of these compounds on animal health, food safety and environmental impact remain a big concern. A recently developed inhibitor, 3-nitrooxypropanol, which targeted the methane-producing enzyme methyl-coenzyme M reductase, inhibited the growth of methanogenic archaea without negatively affecting other non-methanogenic bacteria or the livestock. (1, 17). Other compounds known to inhibit the growth of methanogens are anthraquinones and various plant secondary compounds, such as garlic essential oils and allicin (6, 131

18, 19). However, much less is known about the targets of these compounds or the molecular mechanisms of these interactions.

The methanogenic archaeon Methanococcus maripaludis has been used as an excellent model organism to study methanogen metabolism and physiology due to its rapid and reliable growth, availability of a complete genome sequence and a set of well-developed genetic manipulation strategies (20, 21). Echinomycin, found in streptomycetes, is a cyclic octadepsipeptide antibiotic with two quinoxaline rings that bisintercalate into DNA (22).

Echinomycin is composed of two depsipeptides containing D-serine, L-N-methylvaline, L-N- methylcysteine and L-alanine with a quinoxaline base attached to d-serine (23). The two peptide strands are joined by a thioacetal bridge and two ester linkages between D-serine and L-N- methylvaline (23). Echinomycin showed activity against vancomycin-resistant enterococci (24), hypoxia-inducible factor-1 suppression (25), HIV-1 Tat transactivation inhibition (26), antithrombotic activity (27), and against methicillin-resistant Staphylococus aureus (28). This antibiotic was recently identified as a growth inhibitor of M. maripaludis S2 in the low 2 µM concentration range (29).

Chloroform is a well-known potent inhibitor of methanogens (30). The methane production in rumen fluids was inhibited 50 % within 79 min after the addition of 7.8 µM chloroform (30). In addition, chloroform was found to reduce rumen methane production to the same extent as bromochloromethane, however, with little or no negative influence on rumen fermentation in dairy cows and in vitro (31, 32). Due to these advantages, chloroform together with other halogenated compounds, such as bromoethanesulfonate, have been used as an experimental model in ruminant research in the last decades (9, 31, 32). Chloroform has been proposed to interfere with the transfer of the methyl group from methyl-tetrahydroxymethanopterin (methyl-H4MPT) to coenzyme M 132

(CoM), at the cobamide-dependent methyl-H4MPT:HS-CoM methyltransferase (Mtr) step of the methanogenesis pathway (33, 34). However, the specific mechanism is not known yet.

Neomycin is an aminoglycoside antibiotic that inhibits both the growth of M. maripaludis in vivo and protein synthesis in vitro (35, 36). The minimal inhibitory concentration of neomycin to M. maripaludis was previously determined to be 1.1 mM (36, 37). Moreover, neomycin has already been utilized as a selectable marker in the development of genetic systems in M. maripaludis (37). To deliver neomycin resistance in M. maripaludis, the aminoglycoside phosphotransferase genes APH3’I and APH3’II were cloned under the control of the M. voltae methyl reductase promoter and transformed into M. maripaludis (37).

In this report, the investigation of the target of methanogen inhibitors, neomycin, chloroform or echinomycin were attempted. Spontaneous resistant mutants of echinomycin, chloroform or neomycin were produced from M. maripaludis wild-type strain S2 or S0001. These mutant strains, together with their parental strains, were subjected to whole-genome sequencing using the Illumina sequencing platform. Bioinformatic analysis was performed by using an established computational pipeline to identify mutations relative to the ancestral genome. In echinomycin resistant mutants, two shared mutations were mapped on the intergenic region of M. maripaludis S2 genome. Six mutations, including two that occurred on the intergenic regions and the rest on protein-coding regions, were identified in both chloroform and neomycin resistant mutants. Another two independent mutations, on an unannotated protein (MMP0535) and a sodium:proton antiporter (MMP0707), were found only in neomycin-resistant mutants.

Interestingly, several unexpected polymorphisms were observed in the re-sequenced M. maripaludis parental wild-type strain S2 and S0001. Particularly, an iron-sulfur cluster protein, encoded by mmp1146 and mmp1147, was found to be greatly disrupted by a variety of loss-of- 133

function mutations. Overall, these mutational events may provide key insights into the resistant mechanisms of the tested methanogen inhibitors on the genome-wide scale.

Materials and Methods

Strains, media and growth conditions. Two Methanococcus marpaludis parental strains, the wild-type S2 and S0001, were used in this study. M. maripaludis S0001 is a mutant derived from wild-type strain S2 by the deletion of the gene encoding hypoxanthine phosphoribosyltransferase (MMP0145) and the addition of the gene encoding the rep gene from the Methanococcus shuttle vector pURB500 (38). It is frequently used in a markerless mutation system and as a host for shuttle vectors of M. maripaludis (38, 39). Cultures were grown in

McFAA medium (a formate minimal medium supplemented with 10 mM sodium acetate and 1 mM alanine) reduced with 3 mM dithiothreitol as indicated (40). The 5-mL cultures were grown in 28-mL aluminium seal tubes pressurized to 103 kPa with N2/CO2 (4:1, v/v). The agar medium was prepared in 70 mL serum bottles and 10 ml of McFAA with 1 % (w/v) agar (40). Serum bottles were pressurized to 103 kPa with N2/CO2 (4:1, v/v). Before inoculation, 3 mM sodium sulfide was added as the sulfur source. Echinomycin (>98 % purity, CAS #: 512-64-1), chloroform (>99 % purity) and neomycin (>99 % purity) were purchased from Sigma-Aldrich (USA).

Isolation of M. maripaludis echinomycin-resistant mutants from strains S2 and S0001.

Echinomycin-resistant mutants were generated in strains S2 and S0001 using a serial culture method. The minimum inhibitory concentration (MIC) of echinomycin to Methanococcus maripaludis was first determined in broth culture, and it was 1 µM. For these experiments, an inoculum of 1.5 × 105 cells of strain S2 was challenged with 0.1 µM, 0.2 µM, 0.5 µM, 1 µM, 2

µM, and 5 µM of echinomycin in defined formate medium. Three replicate cultures were used for each concentration. Growth was not observed in any of the tubes when the concentration of 134

echinomycin was > 1.0 µM echinomycin. At concentrations <0.5 µM echinomycin, growth was observed in some but not all of the replicate tubes, which was taken as evidence for selection of spontaneous resistance mutants (Supplementary figure S4-1). To obtain individual echinomycin- resistant mutants, three different inoculum sizes of wild-type M. maripaludis S2 (1.5 × 105, 1.5 ×

106 and 1.5 × 107 cells) were plated onto defined agar medium with three concentrations of echinomycin (0.5 µM, 1 µM, and 2 µM) for nine conditions in total. Isolated colonies were only observed on agar plates with 0.5 µM echinomycin at a frequency of about one in 105. To screen isolated resistant mutants, 20 colonies were picked from these agar cultures and inoculated into defined broth medium containing 1 µM echinomycin. Among these 20-independent resistant mutant lines, six lines were chosen for next step experiments. These six parallel mutant lines were then propagated in 5 mL McFAA medium with 1.0 µM echinomycin. Cultures were transferred every 24 hours by inoculating ~5 % (v/v) of the culture to 5 mL fresh medium, four times in total.

The absorbance of these six mutant lines during each transfer was recorded. Two lines, Ech 25 and

Ech 26, with the highest growth rate, were selected for whole-genome sequencing.

In parallel, echinomycin-resistant mutants were isolated from strain S0001. M. maripaludis S0001 was retrieved in rich medium with 2 % (w/v) casamino acids from a glycerol stock and saved as the parental strain for later resequencing. Three parallel broth cultures, EchB,

EchC, and EchD were inoculated with 106 cells from this seed culture and grown to early stationary phase. Cells, 106, from each lineage were inoculated into defined medium agar bottles with 0.5

µM echinomycin. Isolated colonies were observed at a frequency of about one in 104. Twelve colonies were picked from these agar bottles (four colonies from each lineage) and inoculated into defined broth medium containing 0.5 µM echinomycin for further characterization. These 12- parallel mutant lines were then subcultured in 5 mL McFAA medium with 0.5 µM echinomycin 135

by inoculating 105 cells. The absorbance of these twelve mutant lines during each transfer was recorded. Three mutant lines, Ech B1, Ech C4 and Ech D3 that had the highest growth rate, together with mutant Ech C2 that had the lowest growth rate, were selected for whole-genome sequencing.

Isolation of chloroform and neomycin resistance mutants. This portion of resistant mutant isolating experiments was performed by Dr. Cook’s lab at University of Otago, New

Zealand. The MIC of chloroform and neomycin to M. maripaludis S2 was first determined in broth culture and found to be 1 μM and 68 μM, respectively. When isolating resistant mutants, an inoculum of 1.5 × 105 cells of M. maripaludis S2 cells was challenged with 5 μM, 10 μM and

20μM of chloroform, or 100 μM, 169 μM, 338 μM, and 677 μM of neomycin in McFAA broth medium (40). Replicate cultures were made for each line. In tubes with the highest concentration of chloroform or neomycin, growth was not observed. Growth was observed in some of the tubes with the lower concentration of chloroform or neomycin. 10 % (v/v) of grown-up culture from

McFAA broth medium with 10 µM chloroform or 169 µM neomycin from the previous step was subsequently plated to McFAA agar medium in the presence of 10 uM chloroform or 100 uM neomycin, receptively. Replicates were made for each line. Following steps of propagating the resistant mutations were performed in the anaerobic chamber containing a gas mixture of 4 % H2,

5% CO2, and 91% N2. 40 colonies of chloroform or neomycin mutants were picked and inoculated into an individual well of a 96-well microtiter plate, respectively. Each well contained 320 µL

McFAA broth medium with 10 uM of chloroform or 100 uM of neomycin. 15 grown-up resistant cultures from each antibiotic were pooled together and further transferred three times in 28-mL aluminum seal tubes. Mutant cells were transferred every 24 hr by inoculating ~10 % (v/v) of the culture into 5mL McFAA broth medium with the chloroform or neomycin. Starting with 10 uM 136

of chloroform or 100 uM of neomycin, the dosage was increased to 15 uM of chloroform or 150 uM of neomycin at the second transfer, and to 20 uM of chloroform or 200 uM of neomycin at the third transfer. Four chloroform resistant mutants and four neomycin resistant mutants from the last transfer were selected for whole-genome sequencing.

Whole-genome sequencing. Genomic DNA from resistant mutants and the corresponding parental strains were isolated using Quick-DNA™ Fungal/Bacterial Miniprep Kit from Zymo

Research (CAS #: D6005) or Qiagen DNeasy Blood and Tissue Kit. For the six echinomycin resistant strains and their parental strains, 1 µg of genomic DNA for each strain were sheared into

350 bp fragments by the Covaris E220 Evolution instrument at the Georgia Genomics facility. The genomic DNA library for sequencing was then constructed using NEBNext® Ultra DNA Library

Prep Kit for Illumina (CAS #: E7370). The i7/i5 Illumina indexed primers used for each genomic library were listed in Supplementary Table S4-1. These eight genomic libraries were sequenced by Illumina NextSeq Paired-End sequencing with 25-30 Gb bases being read, operated by Georgia

Genomics facility. For the four chloroform resistant mutants, four neomycin resistant mutants and their parental strain S2, the library preparation and sequencing were carried out by New Zealand

Genomics Limited, New Zealand.

Bioinformatics analysis of genome sequences. Sequencing data were analyzed using

Geneious 10 software (41). The pair-ended readings of each strain were first mapped back to the M. maripaludis S2 genome sequence (21). To look for variant Single Nucleotide Polymorphisms

(SNPs) of each strain, the SNPs were called with the variant frequency of at least 0.1 and a 10-6 maximum variant P-value. By using the “Find Variant SNP” function in Genious 10, the SNPs of each resistant mutant was subtracted from that of their corresponding parental strains, SNPs unique to each resistant mutant were then collected. To exclude those mutations that existed before the 137

antibiotic selection began and sequencing artifacts, these variant SNPs of resistant mutants were manually examined to remove any mutations that also existed in the parental strains. Variant polymorphisms present in echinomycin, chloroform or neomycin resistant mutants were then identified. Gene synteny of some identified proteins was examined using SYNtax (42) and BLAST searches from NCBI database (43). The total number of SNPs of the resistant mutants and the number of variant SNPs from their parental strains were summarized in Supplementary Table S4-

Results

Whole-genome sequencing of M. maripaludis echinomycin-resistant mutants. Six echinomycin-resistant mutants, including two S2-derived mutants and four S0001-derived mutants, were subjected to whole-genome sequencing using Illumina technology. Because M. maripaludis is polypoid and contains about 20-50 copies of the genome per cell (44), it was possible that resistance could have resulted from dominant mutations in a small fraction of the alleles. For that reason, all mutations that occurred in at least 10 % of the sequencing reads were examined.

While numerous mutations were detected in echinomycin- resistant mutants, only two loci contained shared mutations in S2-derived mutants. In the mutant E25, there was a transversion from A to C or A to G at position 475,147 on M. maripaludis genome, with a frequency of 10.4 %.

In the mutant E26, there was a transversion from A to C at position 475,147, and another one from

T to G at 475,141, with frequencies of 13.4 % and 16.8 %, respectively. These two loci were present in a 580 bp intergenic region between mmp0478 and mmp0479. These two genes encode proteins of unknown function. For the S0001-derived mutants, no mutations were identified that were shared by all four mutants. Ten mutations were shared in two or three of these S0001-derived

138

mutants. However, none of these mutational events seemed to be functionally relevant to echinomycin resistance.

Whole-genome sequencing of M. maripaludis chloroform- and neomycin-resistant mutants. Four neomycin-resistant and four chloroform-resistant mutants were derived from the same parental strain, M. maripaludis S2. All these lines were subjected to whole-genome sequencing.

In the chloroform-resistant mutants, six shared mutations, with at least 86 % frequency, were identified (Table 4-1). They occurred at position 107,992 in the intergenic region of M. maripaludis genome, position 1,588,772 in the intergenic region, mmp_rs07920 (tRNA-IIe), mmp007 (geranylgeranylglyceryl phosphate synthase), mmp1115 (transketolase) and mmp1689

(comE). Interestingly, these six mutations were also found in the neomycin-resistant mutants

(Table 4-1 and Table 4-2). The mutations that occurred at the first five loci were identical in both two types of resistant mutants, suggesting that they were derived from the parental strain. However, in mmp1689 (comE), the mutations were different. First, a deletion of nucleotide T at position

1,630,332 was found in both chloroform-resistant mutants and neomycin-resistant mutants.

Second, another three mutational events, an insertion and two transversions, were found in the nearby positions in neomycin-resistant mutants only.

In the neomycin-resistant mutants, a deletion in mmp0535 (hypothetical protein) and a transversion in mmp0707 (sodium: proton antiporter) were also found in all neomycin-resistant mutants. (Table 4-2).

Mutations in re-sequenced M. maripaludis S2 and S0001 strains. Three M. maripaludis wild-type S2 strain and three S0001 strain were re-sequenced in this experiment. They were a M.

139

maripaludis S2 strain as the parental strain of neomycin- and chloroform- resistant mutants; a subculture of this S2 parental strain; another S2 strain as the parental strain for generating echinomycin- resistant mutants; a M. maripaludis S0001 strain as the parental strain for generating echinomycin- resistant mutants; and two S0001 subcultures of this parental strain. The first two

M. maripaludis S2 strains together with their resistant mutants were prepared and sequenced in Dr.

Cook’s lab in New Zealand. The next four M. maripaludis strains S2 and S0001, together with their resistant mutants, were prepared and sequenced in this work in U.S.A.

To look for mutations that exist in all re-sequenced M. maripaludis wild-type strains, variant SNPs with at least 80 % of frequency were collected and compared (Table 4-3, 4-4).

Several mutations that existed in all three re-sequenced S2 genomes were identified, and they all occurred at least 89 % of frequency (Table 4-3). Strikingly, a variety of mutational events, such as transitions, transversions, insertions, and substitutions, were found in mmp1176 and mmp1177, with a frequency of 88.9 % to 100 %. (Figure 4-1). Similar but fewer mutations, with a frequency of 14.7 % to 75.8 %, were also mapped in these two genes in the three re-sequenced S0001 strains

(Table 4-4). The emergence of these mutations in mmp1176 and mmp1177 may have reduced the activities of the enzymes these genes encode. Mmp1176 and mmp1177 are overlapping genes that encode the N-terminal and C-terminal regions of a substrate-binding protein of an iron transport system, respectively. One tentative interpretation of this observation is that mutations in the laboratory strains of M. maripaludis may have accumulated following extended cultivation in medium where iron was readily available. The iron concentration for cultivating M. maripaludis in the laboratory is usually around 25 μM (40), a concentration close to that of this organism’s original habitat in marine sediments (45, 46). However, the real available iron for M. maripaludis in their native environment is unknown. In laboratory cultivated M. maripaludis, iron was 140

constantly provided in medium. Therefore, it was possible that the accumulation of mutations in iron-transport protein MMP1176 and MMP1177 was driven by the selection towards less competition for iron compared to that in the organism’s original habitat. Another two mutations that were found in all re-sequenced S2 and S0001 strains were on mmp1477 and mmp1478. These two genes together encode a cobyrinic acid a,c-diamide synthase. The last mutation that has been found on both re-sequenced S2 and S0001 strains was an A nucleotide deletion on the position of

1,495,465 in an intergenic region. This region is between the cytochrome c (MMP1537) and a gene of unknown function (MMP1158). As these mutations were identical in all S2 and S0001 strains, they were unlikely to be involved in the acquisition of resistance.

Other mutations found only in all re-sequenced S2 strains were the following. a nucleotide

C to T transition on the position of 255,618 from mmp0253, encoding a hypothetical protein. This mutation altered the original amino acid glycine into glutamic acid. Two, a nucleotide changed from C to T on the position of 392,113 of mmp0394, encoding uroporphyrinogen III synthase.

This mutation changed an amino acid from aspartic acid to asparagine. Lastly, a nucleotide changed from A to G on the position of 1,428,254 from mmp1466, encoding a CBS domain- containing signal transduction protein. This mutation changed amino acid from aspartic acid glycine. Because these three mutation events were not independent among all re-sequenced S2 strains, possibly, they were structural variant without any functional disruptions.

In the re-sequenced M. maripaludis S0001, both mmp0145 (hypoxanthine phosphoribosyltransferase) and mmp0680 (uracil phosphoribosyltransferase) were deleted during the construction of this strain from strain S2, as it confirmed in our re-sequencing data (Table 4-

141

4). Other polymorphisms, all identical in these three S0001 strains, were summarized in Table 4-

Discussions

We have presented here a genome-sequence-based analysis of M. maripaludis S2 strains resistant to three inhibitors, echinomycin, chloroform, or neomycin, attempting to explore the resistance mechanisms on a genome-wide scale. No studies have yet used whole-genome sequencing to examine the mutations in resistance mutants of methanogens. Resistant mutants to echinomycin, chloroform or neomycin were first generated by serial transfers in the presence of sub-lethal levels of the inhibitors. Based on the subsequent whole-genome sequencing analysis, a number of SNPs were identified in all resistant mutants.

Echinomycin. In this study, only two shared mutations were identified in S2-derived echinomycin resistant mutants. Considering that these two mutations all took place on the intergenic region on M. maripaludis genome, it was most likely that they were not the cause of the resistance phenotype. However, no shared mutations were found in the S0001-derived resistant mutants. These observations suggested that resistance may have resulted from multiple mutational events. Alternatively, mutations may be dominant and never become abundant in the cells of these polyploid microorganisms. When producing the resistant mutants, the M. maripaludis S2 or S0001 strains were challenged with 0.5 μM echinomycin, which is below the MIC of this compound (1

µM). Isolated S2- derived resistant mutants from this step were further transferred four times in the presence of 1 μM echinomycin. These four passages propagated up to ~ 17 generations. At the same time, S0001-derived resistant mutants were further transferred once in the presence of 0.5

μM echinomycin. That is, up to ~ 13 generations were propagated. Similarly, up to ~ 10

142

generations were propagated for chloroform or neomycin resistant mutants. A former study of antibiotic-resistant mechanisms with E. coli resistant mutants used up to 61 independent lines with up to ~336 generations of propagation (47). A total of 402 independent mutational events were detected in all E. coli resistant mutants (47). Therefore, in future experiments of preparing resistant mutants to study the resistant mechanisms at the genomic level, refinements such as applying a higher dose of inhibitor and increasing the strength of propagating the mutation, should be carefully arranged.

Neomycin and Chloroform. Bacterial resistance to the aminoglycoside antibiotics, such as neomycin, is mostly associated with the expression of modifying enzymes which phosphorylate, adenylate or acetylate these compounds (48). Three types of aminoglycoside-modifying enzymes were reported so far, they are O-phosphotransferase (APHs), O-adenyltranferases (ANTs) and N- acetyltransferases (AACs) (48). However, none of these protein homologs exists in M. maripaludis.

Other resistance mechanisms, such as diminished cell membrane permeability, structural alteration in the ribosomal target of the antibiotic, or extrusion of the aminoglycosides from the cell by efflux pumps, were also proposed (49).

In our study, two shared mutations were identified only in neomycin- resistant mutants: (1) an unannotated protein (MMP0535); (2) a sodium: proton antiporter (MMP0707), which is very interesting as this protein is critical for maintaining pH homeostasis. However, these two mutations should be further validated by PCR or quantitative PCR analysis with the genome of resistant mutants, before any conclusion is drawn.

Three mutations at the protein-coding regions were detected in both neomycin- and chloroform- resistant mutants. These included geranylgeranylglyceryl phosphate synthase

(MMP0007), transketolase (MMP1115) and ComE (MMP1689). Nonetheless, whether these 143

proteins are associated with neomycin and or chloroform resistance in M. maripaludis requires further genetic and biochemical validations.

By combing the generation of resistant mutants using serial transfer techniques with inhibitors under sub-lethal concentrations, together with the subsequent whole-genome sequencing analysis, this work has mapped a viable pipeline of exploring the targets of inhibitors. Although certain experimental details still require further refinements, such as increasing size of the mutation propagation, mutations that potentially associated with the inhibitor-resistant mechanisms were identified. This is a critical step to develop novel microbial inhibitors, after a list of candidates has been collected from phenotypic screening techniques. Ultimately, the goal is to elucidate the underlying molecular mechanisms of resistance, thus determining the structural information of the drug.

References:

1. Duin EC, Wagner T, Shima S, Prakash D, Cronin B, Yanez-Ruiz DR, Duval S, Rumbeli

R, Stemmler RT, Thauer RK, Kindermann M. 2016. Mode of action uncovered for the

specific reduction of methane emissions from ruminants by the small molecule 3-

nitrooxypropanol. Proc Natl Acad Sci U S A. 113(22):6172-7.

2. Gerber PJ, Henderson B, Makkar HPS. 2013. Mitigation of greenhouse gas emissions in

livestock production: A review of technical options for non-CO2 emissions. FAO Animal

Production and Health Paper NO.177.

3. Conrad R. 2009. The global methane cycle: recent advances in understanding the microbial

processes involved. Environ Microbiol Rep 1:285-92.

4. Johnson KA, Johnson DE. 1995. Methane emissions from cattle. J Animal Sci 73:2483-

2492. 144

5. VanNevel CJ, Demeyer DI. 1996. Control of rumen methanogenesis. Environ Monit

Assess 42:73-97.

6. Henderson G, Cook GM, Ronimus RS. 2016. Enzyme- and gene-based approaches for

developing methanogen-specific compounds to control ruminant methane emissions: a

review. Anim Prod Sci doi:https://doi.org/10.1071/AN15757.

7. Hook SE, Wright ADG, McBride BW. 2010. Methanogens: methane producers of the

rumen and mitigation strategies. Archaea doi: 10.1155/2010/945785.

8. Buddle BM, Denis M, Attwood GT, Altermann E, Janssen PH, Ronimus RS, Pinares-

Patino CS, Muetzel S, Wedlock DN. 2011. Strategies to reduce methane emissions from

farmed ruminants grazing on pasture. Vet J 188:11-17.

9. Leahy SC, Kelly WJ, Altermann E, Ronimus RS, Yeoman CJ, Pacheco DM, Li D, Kong

ZH, McTavish S, Sang C, Lambie SC, Janssen PH, Dey D, Attwood GT. 2010. The genome

sequence of the rumen methanogen Methanobrevibacter ruminantium reveals new

possibilities for controlling ruminant methane emissions. Plos One 5.

10. Sun XZ, Henderson G, Cox F, Molano G, Harrison SJ, Luo D, Janssen PH, Pacheco D.

2015. Lambs fed fresh winter forage rape (Brassica napus L.) emit less methane than those

fed perennial ryegrass (Lolium perenne L.), and possible mechanisms behind the

difference. Plos One 10.

11. Eckard RJ, Grainger C, de Klein CAM. 2010. Options for the abatement of methane and

nitrous oxide from ruminant production: A review. Livestock Science 130:47-56.

12. Patra AK, Saxena J. 2010. A new perspective on the use of plant secondary metabolites to

inhibit methanogenesis in the rumen. Phytochemistry 71:1198-1222.

145

13. Hegarty RS. 1999. Reducing rumen methane emissions through elimination of rumen

protozoa. Aust J Agric Res 50:1321-1327.

14. Wedlock DN, Pedersen G, Denis M, Dey D, Janssen PH, Buddle BM. 2010. Development

of a vaccine to mitigate greenhouse gas emissions in agriculture: Vaccination of sheep with

methanogen fractions induces antibodies that block methane production in vitro. N Z Vet

J 58:29-36.

15. Chalupa W. 1980. Chemical control of rumen microbial metabolism. In Ruckebusch Y,

Thivend P (ed), Digestive physiology and metabolism in ruminants: Proceedings of the 5th

International Symposium on Ruminant Physiology, held at Clermont Ferrand. 325-347

doi:10.1007/978-94-011-8067-2_16. Springer Netherlands, Dordrecht.

16. Trei JE, Scott GC, Parish RC. 1972. Influence of methane inhibition on energetic efficiency

of lambs. J Animal Sci 34:510-5.

17. Hristov AN, Oh J, Giallongo F, Frederick TW, Harper MT, Weeks HL, Branco AF, Moate

PJ, Deighton MH, Williams SRO, Kindermann M, Duval S. 2015. An inhibitor persistently

decreased enteric methane emission from dairy cows with no negative effect on milk

production. Proc Natl Acad Sci U S A 112(34):10663-8.

18. GarciaLopez PM, Kung L, Odom JM. 1996. In vitro inhibition of microbial methane

production by 9,10-anthraquinone. J Animal Sci 74:2276-2284.

19. Soliva CR, Amelchanka SL, Duval SM, Kreuzer M. 2011. Ruminal methane inhibition

potential of various pure compounds in comparison with garlic oil as determined with a

rumen simulation technique (Rusitec). Br J Nutr 106:114-122.

20. Whitman WB, Shieh J, Sohn S, Caras DS, Premachandran U. 1986. Isolation and

characterization of 22 mesophilic methanococci. Syst Appl Microbiol 7:235-240. 146

21. Hendrickson EL, Kaul R, Zhou Y, Bovee D, Chapman P, Chung J, de Macario EC,

Dodsworth JA, Gillett W, Graham DE, Hackett M, Haydock AK, Kang A, Land ML, Levy

R, Lie TJ, Major TA, Moore BC, Porat I, Palmeiri A, Rouse G, Saenphimmachak C, Soll

D, Van Dien S, Wang T, Whitman WB, Xia Q, Zhang Y, Larimer FW, Olson MV, Leigh

JA. 2004. Complete genome sequence of the genetically tractable hydrogenotrophic

methanogen Methanococcus maripaludis. J Bacteriol186:6956-6969.

22. Gilbert DE, Vandermarel GA, Vanboom JH, Feigon J. 1989. Unstable hoogsteen base-

pairs adjacent to echinomycin binding-sites within a DNA duplex. Proc Natl Acad Sci U S

A 86:3006-3010.

23. Dell A, Williams DH, Morris HR, Smith GA, Feeney J, Roberts GC. 1975. Structure

revision of the antibiotic echinomycin. J Am Chem Soc 97:2497-502.

24. Kim JB, Lee GS, Kim YB, Kim SK, Kim YH. 2004. In vitro antibacterial activity of

echinomycin and a novel analogue, YK2000, against vancomycin-resistant enterococci. Int

J Antimicrob Agents 24:613-615.

25. Kong DH, Park EJ, Stephen AG, Calvani M, Cardellina JH, Monks A, Fisher RJ,

Shoemaker RH, Melillo G. 2005. Echinomycin, a small-molecule inhibitor of hypoxia-

inducible factor-1 DNA-binding activity. Cancer Research 65:9047-9055.

26. Jayasuriya H, Zink DL, Polishook JD, Bills GF, Dombrowski AW, Genilloud O, Pelaez

FF, Herranz L, Quamina D, Lingham RB, Danzeizen R, Graham PL, Tomassini JE, Singh

SB. 2005. Identification of diverse microbial metabolites as potent inhibitors of HIV-1 tat

transactivation. Chem Biodivers 2:112-122.

147

27. Lee YK, Park JH, Moon HT, Lee DY, Yun JH, Byun Y. 2007. The short-term effects on

restenosis and thrombosis of echinomycin-eluting stents topcoated with a hydrophobic

heparin-containing polymer. Biomaterials 28:1523-1530.

28. Park YS, Shin WS, Kim SK. 2008. In vitro and in vivo activities of echinomycin against

clinical isolates of Staphylococcus aureus. J Antimicrob Chemother 61:163-168.

29. Weimar MR, Cheung J, Dey D, McSweeney C, Morrison M, Kobayashi Y, Whitman WB,

Carbone V, Schofield LR, Ronimus RS, Cook GM. 2017. Development of multiwell-plate

methods using pure cultures of methanogens to identify new inhibitors for suppressing

ruminant methane emissions. Appl Environ Microbiol 83(15). pii: e00396-17.

30. Bauchop T. 1967. Inhibition of rumen methanogenesis by methane analogues. J Bacteriol

94:171-5.

31. Knight T, Ronimus RS, Dey D, Tootill C, Naylor G, Evans P, Molano G, Smith A,

Tavendale M, Pinares-Patiño CS, Clark H. 2011. Chloroform decreases rumen

methanogenesis and methanogen populations without altering rumen function in cattle.

Anim Feed Sci Technol 166-167:101-112.

32. Martinez-Fernandez G, Denman SE, Yang C, Cheung J, Mitsumori M, McSweeney CS.

2016. Methane Inhibition alters the microbial community, hydrogen flow, and

fermentation response in the rumen of cattle. Front Microbiol 7:1122.

33. Gunsalus RP, Wolfe RS. 1978. ATP activation and properties of the methyl coenzyme M

reductase system in Methanobacterium thermoautotrophicum. J Bacteriol 135:851-7.

34. Graham DE, White RH. 2002. Elucidation of methanogenic coenzyme biosyntheses: from

spectroscopy to genomics. Nat Prod Rep 19:133-47.

148

35. Pecher T, Böck A. 1981. In vivo susceptibility of halophilic and methanogenic organisms

to protein synthesis inhibitors. FEMS Microbiol Lett 10:295-297.

36. Weisburg WG, Tanner RS. 1982. Aminoglycoside sensitivity of archaebacteria. FEMS

Microbiol Lett 14:307-310.

37. Argyle JL, Tumbula DL, Leigh JA. 1996. Neomycin resistance as a selectable marker in

Methanococcus maripaludis. Appl Environ Microbiol 62:4233-4237.

38. Walters AD, Smith SE, Chong JPJ. 2011. Shuttle vector system for Methanococcus

maripaludis with improved transformation efficiency. Appl Environ Microbiol 77:2549-

2551.

39. Moore BC, Leigh JA. 2005. Markerless mutagenesis in Methanococcus maripaludis

demonstrates roles for alanine dehydrogenase, alanine racemase, and alanine permease. J

Bacteriol 187:972-979.

40. Long F, Wang L, Lupa B, Whitman WB. 2017. A flexible system for cultivation of

Methanococcus and other formate-utilizing methanogens. Archaea. 2017:12.

doi:10.1155/2017/7046026.

41. Kearse M, Moir R, Wilson A, Stones-Havas S, Cheung M, Sturrock S, Buxton S, Cooper

A, Markowitz S, Duran C, Thierer T, Ashton B, Meintjes P, Drummond A. 2012. Geneious

basic: An integrated and extendable desktop software platform for the organization and

analysis of sequence data. Bioinformatics 28:1647-9.

42. Oberto J. 2013. SyntTax: a web server linking synteny to prokaryotic taxonomy. BMC

Bioinformatics 14.

149

43. Altschul SF, Madden TL, Schaffer AA, Zhang JH, Zhang Z, Miller W, Lipman DJ. 1997.

Gapped BLAST and PSI-BLAST: a new generation of protein database search programs.

Nucleic Acids Res 25:3389-3402.

44. Hildenbrand C, Stock T, Lange C, Rother M, Soppa J. 2011. Genome copy numbers and

gene conversion in methanogenic archaea. J Bacteriol 193:734-743.

45. Dale AW, Nickelsen L, Scholz F, Hensen C, Oschlies A, Wallmann K. 2015. A revised

global estimate of dissolved iron fluxes from marine sediments. Global Biogeochem

Cycles 29:691-707.

46. de Chanvalon AT, Metzger E, Mouret A, Knoery J, Geslin E, Meysman FJR. 2017. Two

dimensional mapping of iron release in marine sediments at submillimetre scale. Mar Chem

191:34-49.

47. Lazar V, Nagy I, Spohn R, Csorgo B, Gyorkei A, Nyerges A, Horvath B, Voros A, Busa-

Fekete R, Hrtyan M, Bogos B, Mehi O, Fekete G, Szappanos B, Kegl B, Papp B, Pal C.

2014. Genome-wide analysis captures the determinants of the antibiotic cross-resistance

interaction network. Nat Commun 5:4352.

48. Wright GD. 1999. Aminoglycoside-modifying enzymes. Curr Opin Microbiol 2:499-503.

49. Azucena E, Mobashery S. 2001. Aminoglycoside-modifying enzymes: mechanisms of

catalytic processes and inhibition. Drug Resist Updat 4:106-17.

150

Figure 4-1. Partial genomic map of the region of position 1,158,991 to 1,160,125 presenting variant SNPs of re-sequenced Methanococcus maripaludis S2 (top three) and S0001 (bottom two) strains mapped back to the S2 genome. The top re-sequenced M. maripaludis S2 strain was the parental strain of the echinomycin-resistant mutants, the second re-sequenced S2 strain was the parental strain of chloroform- resistant mutants and neomycin- resistant mutants, and the third S2 strain was a subculture of the second S2 strain. The first re-sequenced M. maripaludis S0001 was the parental strain of echinomycin-resistant mutants, the second re-sequenced S0001 strain was a subculture of this strain.

Blue rectangles represent the position 1,158,991 to 1,160,125 of M. maripaludis S2 genome (53).

Green bars stand for the genes among this region on M. maripaludis S2 genome.

Orange bars represent variant SNPs of each strain, they were 13, 73, 31, 1, 1 variant SNPs on

MMP1176 from top to bottom, with 88.9 %-99.1 %, 81.8 %-99.8 %, 88.1 %-100 %, 14.7 %, 19.8 % variant frequency, respectively; and 7, 12, 12, 2, 1 variant SNPs on MMP1177 from top to bottom, with 89.5 %-98.6 %, 99.1 % - 99.8 %, 99.5 % - 100 % variant frequency, respectively.

151

Table 4-1. Variant SNPs found in at least three out of four chloroform- resistant mutants.

Amino Variant Polymorphis SNPs Encoded Protein/tRNA acid Protein effect frequency m type change Non-coding region 92.1 % to Transversion N/A N/A at 107,992 b 100 % Non-coding region 91.1 % to Transition N/A N/A at 1588,772 b 100 % MMP_RS07920 b tRNA-IIe 91 % to 100 % Transition N/A N/A Geranylgeranylglyceryl 89.30 % to MMP0007 b Transversion L  H Substitution phosphate synthase 99.9 % 88.20 % to Substitution, MMP1115 ab Transketolase Transition L  P 99.8 % Frame shift 86.20 % to Frame shift, MMP1689 b ComE Deletion N/A 100 % Substitution a SNPs shared by three out of four chloroform- resistant mutants. The rest SNPs were found in all four chloroform-resistant mutants. b SNPs also found in neomycin- resistant mutants.

N/A: not applicable.

152

Table 4-2. Variant SNPs found in at least three out of four neomycin- resistant mutants.

Variant Polymorphism Amino acid Protein SNPs Encoded Protein/tRNA frequency type Change effect Non-coding region 100 % Transversion N/A N/A at 107,992 b Non-coding region 99.3 % to Transition N/A N/A at 1588,772 b 100 % 98.4 % to MMP_RS07920 b tRNA-IIe Transition N/A N/A 100 % Geranylgeranylglyceryl 99.6 % to mmp0007b Transversion LH Substitution phosphate synthase 99.90 % mmp0535 Hypothetical protein 100 % Deletion N/A Frame shift Sodium:proton 11.6 % to Transversion G  C, mmp0707a Substitution antiporter 99.5 % Transition A  V 99.8 % to mmp1115 b Transketolase Transition L  P Substitution 99.90 % Insertion, 11.4 % to Frame shift, mmp1689 b ComE Transversion S  T 100% Substitution Deletion a SNPs shared by three out of four neomycin- resistant mutants. The rest SNPs were found in all four neomycin- resistant mutants. b SNPs also found in chloroform- resistant mutants.

N/A: not applicable.

153

Table 4-3. Variant SNPs, with at least 80 % frequency, found in three re-sequenced M. maripaludis S2 strains.

Amino Variant Polymorphism Protein SNPs Encoded Protein/tRNA acid frequency type change change Intergenic 99.4 % to region at N.A. Deletion N/A N/A 99.9 % 1,495,465* Intergenic 98.1 % to region at N.A. Substitution N/A N/A 100 % 454,886 CoA-binding domain- 99.8 % to mmp0253 Transition GE Substitution containing protein 100 % Uroporphyrinogen III 98.7 % to mmp0394 Transition D  N Substitution synthase 99.8 % Transition, N D, Iron transport system Substitution, 88.9 % to Transversion, G D, mmp1176* substrate-binding protein, Truncation, 100 % Insertion, E  Q N-term half Frame shift Substitution …...* Transition, Iron transport system EN, Substitution, 89.5 % to Transversion, LS, mmp1177* substrate-binding protein, Truncation, 100 % Insertion, FT, C-term half Frame shift Substitution ……* CBS domain-containing 99.4 % to mmp1466 Transition DG Substitution signal transduction protein 100 % Cobyrinic acid a,c-diamide 99.3 % to mmp1477 Deletion N/A Frame shift synthase 100 % Cobyrinic acid a,c-diamide 99.3 % to mmp1478 synthase:cobyrinic acid Deletion N/A Frame shift 100 % a,c-diamide synthase CbiA

* Multiple mutations were identified, in total 86 variant SNPs were found in mmp1176, 19 variant

SNPs were found in mmp1177.

154

Table 4-4. Variant SNPs, with at least 80 % frequencya, found in three re-sequenced

Methanococcus maripaludis S0001 cultures.

Variant Polymorphism Amino Protein SNPs Encoded Protein/tRNA frequency type acid change Intergenic 99.4 % to change N.A. Deletion N/A N/A region at 100 % 1,495,465* Transcription initiation 97.2 % to mmp0041 Substitution V  I Substitution factor IIB 100 % MATE family 98.3 % to mmp0166 Transversion N  I Substitution drug/sodium antiporter 99.2 % 99.2 % to mmp0234 Hypothetical protein Deletion N/A Frame shift 100 % 97.7 % to mmp0697 leucyl-tRNA synthetase Transition I  V Substitution 100 % Carbamoyl-phosphate 98 % to mmp1013 Transversion G  C Substitution synthase large subunit 99 % Stationary phase survival 98.8 % to mmp1051 Deletion N/A. Frame shift protein SurE 100 % 98.3 % to mmp1169 SufBD protein Insertion N/A Frame shift 100 % Iron transport system 14.7 % to mmp1176 a Transversion L  R Substitution substrate-binding protein, 19.8 % Iron Ntransport-term half system 14.7 % to mmp1177a Transversion E  N Substitution substrate-binding protein, 75.8 % Iron transportC-term half system 83.9 % to mmp1181 Transition D  G Substitution binding protein 87.2 % 98.1 % to mmp1305 Hypothetical protein Transition A T Substitution 100 % Cobyrinic acid a,c-diamide 98.8 % to mmp1477 Deletion N/A Frame shift synthase 100 % Cobyrinic acid a,c-diamide 98.8 % to mmp1478 Deletion N/A Frame shift synthase:cobyrinic acid 100 % a,c-diamide synthase CbiA 97.3 % to mmp1593 Hypothetical protein Transversion G  V Substitution 99.1 % 98.6 %to mmp1624 Polyferredoxin Insertion N/A Frame shift 99.3 % a All variant SNPs summarized here occurred at a frequency of at least 80 %, except in mmp1177 and mmp1176. 155

CHAPTER 5

CONCLUSIONS Methanogens are strictly anaerobic microorganisms belonging to the Euryarchaeota. A large and diverse group, they are distinguished by their capability to obtain most if not all of their energy for growth from methane production or methanogenesis. Methanogenesis may represent one of the most ancient metabolisms that originated from the early, anoxic earth. The emergence of O2 around 2.4 billion years ago has resulted in the formation of a large oceanic sulfate pool and the occurrence of widespread microbial sulfate reduction. At the same time, methanogens were restricted to sulfate-depleted anaerobic environmental niches. Contemporary methanogens are still restricted to anaerobic habitats and may have preserved some of the metabolic relics which were common in the early Earth anaerobes. This hypothesis is further supported by the unique aspects of sulfur metabolisms in methanogens. For instance, methanogens do not utilize sulfate as a sulfur source, homologs of many genes common in the bacterial and eukaryotic sulfur metabolism are absent in methanogens, cysteine is not the sulfur donor for iron-sulfur cluster and methionine biosynthesis, and cysteine biosynthesis uses an unusual tRNA-dependent pathway. Therefore, studying the physiology and biochemistry of methanogens may provide new insights to the biology of ancient microbial life.

First, most methanogens are hydrogenotrophs that use either H2 or formate as the major electron donor to reduce CO2 for methane production. The traditional cultivation of these organisms uses H2 and CO2 as the substrate with frequent replenishment of gas during growth. H2 is explosive and requires an expensive gassing system to handle safely. Formate is as an ideal 156

alternative substrate from the standpoints of both economy and safety but leads to large changes in the culture pH during growth. Glycylglycine was found to be an inexpensive and nontoxic buffer suitable for growth of Methanococcus maripaludis and Methanothermococcus okinawensis. A cultivation system of growing these organisms was constructed, using formate as substrate and glycylglycine as medium buffer. It is suitable for growth on liquid as well as solid medium in serum bottles. Moreover, it supports the cultivation of liter scale cultures without expensive and complex fermentation equipment. This formate cultivation system provides an inexpensive and flexible alternative for the growth of formate-utilizing, hydrogenotrophic methanogens.

Second, methanogens are generally unable to assimilate sulfate, instead, all known methanogens can use sulfide or elemental sulfur as the sole S source. In aerobes, sulfate is incorporated into cells via an assimilatory sulfate reduction pathway using several enzymes.

Interestingly, M. maripaludis, a model organism in hydrogenotrophic methanogens, possesses homologs of these known proteins. These M. maripaludis proteins show only low similarity but are highly conserved in the catalytic residues of known sulfate-reduction proteins. They are ATP sulfurylase (MMP1471), ATP kinase (MMP1282), APS reductase (MMP0941 and MMP1681),

PAPS reductase (MMP0941 and MMP1681) and sulfite reductase (MMP0078). However, none of these M. maripaludis proteins was functional in the E. coli mutant strains deficient in sulfate assimilation metabolism. In addition, M. maripaludis does not produce large amounts of sulfate when grown with sulfide as the sole sulfur source. These results indicated that the assimilatory sulfate reduction pathway is most unlikely to present in methanococci.

Third, all methanogens are known to assimilate elemental sulfur as the sole sulfur source, while the corresponding enzyme(s) involving in this process has not been elucidated yet. When

157

grown with elemental sulfur as the sole sulfur source, M. maripaludis produced sulfide in a total of ~ 6 mmol per g cell dry weight per hour. Moreover, adenylyl-sulfate reductase (MMP1681) was found to be required for elemental sulfur incorporation. A M. maripaludis strain with an in-frame deletion of mmp1681 was unable to grow with elemental sulfur as sole sulfur source and was severely impaired for growth with sulfide as the sole sulfur source. However, when grown with elemental sulfur as sole sulfur source, supplementation of the medium with coenzyme M or thiosulfate restored growth to near wild-type levels. Furthermore, proteomics data showed that the expression of MMP1681 increased 3.3-fold in M. maripaludis grown on elemental sulfur. These observations suggested that MMP1681 is required for elemental sulfur assimilation in M. maripaludis. Together with bioinformatics analysis, a different physiological role of MMP1681 in elemental sulfur assimilation, in addition to its in vitro catalytic function as an adenylyl-sulfate reductase was demonstrated.

Lastly, the application of methanogen-specific inhibitors to mitigate the enteric methane emissions from ruminants was studied. Two known inhibitors of methanogenic archaea, neomycin, and chloroform, together with a recently identified inhibitor, echinomycin, were used to produce resistant mutants of M. maripaludis S2 and S0001. Whole-genome sequencing at high coverage (>

100-fold) was performed subsequently to investigate the potential targets of these inhibitors at the genomic level. Upon analysis of the whole-genome sequencing data, a list of shared mutations for each type of resistant mutants was cataloged. Some of these mutations may be associated with the resistant mechanisms. This work developed a viable pipeline for deciphering the underlying molecular mechanisms of methanogen inhibitors at the genome-wide level.

158

APPENDIX A

CHAPTER 2 SUPPLEMENTARY INFORMATION1

1 Long, F., L.L. Wang, B. Lupa, and W.B. Whitman. 2017. A flexible system for cultivation of

Methanococcus and other formate-utilizing methanogens. Archaea. 2017:12. doi:10.1155/2017/7046026. Reprinted here with permission of the publisher.

159

S2A. Basal medium for formate growth (McF) of Methanococcus maripaludis and

Methanothermococcus okinawensis

(1) Select proper glassware.

This medium contains 0.4 M formate. During growth, methanogens convert the formate to CO2 and CH4 gas and cause an increase in pressure. Thus, for every liter of medium, the cells can produce about 10 liters of gas. As a result, the culture vessels can explode if the headspace is not large enough to prevent an accumulation of gas pressure. For this reason, we recommend that the volume of the headspace be no less than five times the volume of medium in culture vessels that can be pressurized up to 276 kPa. Recommended maximum volumes of media for 28 mL aluminum sealed tubes, 160 mL serum bottles and 1 L bottles are 5 mL, 20 mL, and 100 mL, respectively. When using 1 L bottles, reduce the starting pressure of N2/CO2 to 34 kPa (see below).

(2) Deoxygenate all glassware and equipment prior to use in the anaerobic chamber. Bring culture tubes or bottles, stoppers, funnels and pipettes and all plasticware into the chamber at least one day before making medium to allow O2 to desorb or diffuse out.

(3) Medium composition (Table S2-1)

Combine medium ingredients and sparge with a stream of N2 gas for 60 minutes.

(4) Add 0.05 g cysteine-HCl or DTT per 100 mL and continue sparging for 10 minutes.

(5) Transfer medium to an anaerobic chamber, dispense medium into culture vessels and seal tubes with grey butyl stoppers (Wheaton Science Products, cat. no.: W224100-202) and crimp with aluminum seals (Fisher Scientific, cat. no.: 11-126-12).

(6) After removing from the anaerobic chamber, pressurize tubes and serum bottles to 103 kPa with N2/CO2 (4:1, v/v) and autoclave. The pH after autoclaving should be about 7.7-7.8. For one liter bottles, pressurize to 34 kPa N2/CO2 before autoclaving. Autoclave on gravity cycle (rapid 160

exhaust) for 20 minutes. Always autoclave these bottles secured with metal cylinders or wire baskets to limit flying glass in an explosion.

(7) Prior to inoculation, add 0.1 mL of 2.5 % Na2S·9H2O (w/v) per 5 mL of medium [add

2 mL of 2.5 % Na2S·9H2O (w/v) per 100 mL of medium, if made in 1 liter bottle)]. For routine experiments, add the sulfide immediately before inoculation. For critical experiments, add the sulfide 8-24 h before inoculation.

S2A.1 Modifications for growth of Methanothermococcus okinawensis

For cultivation of M. okinawensis, reduce the pH of the glycylglycine buffer to 6.5 and increase the final concentration from 0.2 M to 0.4 M. The other medium ingredients remain the same.

Medium Composition (See table S2-2)

S2A.2 Preparation of stock solutions

S2A.2.1 Preparation of Glycylglycine solution (1M, pH= 8.0)

Use 20 mL per 100 mL of medium

(i) Dissolve 132 g glycylglycine (Amresco, cat. no.: 556-50-3) with 800 ml ddH2O

(ii) Adjust to pH 8.0 with about 18 mL of NaOH (5 M).

(iii) Adjust volume to 1 liter. Store at 4 ℃ in a tightly sealed bottle.

S2A.2.2 Preparation of General Salts solution

Use 50 mL per 100 mL of medium. Ingredients of general salts solution see Table S2-3.

S2A.2.3 Preparation of Iron Stock solution (3)

Use 0.5 mL per 100 mL of medium

To a small screw top bottle, add 0.2 g of Fe(NH4)2(SO4)2·6H2O. Then add 2 drops of concentrated HCl followed by 100 mL of glass-distilled water. 161

S2A.2.3 Preparation of Trace minerals solution (4)

Use 10 mL per 1 liter of medium. Ingredients of trace minerals solution see Table S2-4.

Dissolve the nitriloacetic acid in about 800 mL of diH2O and adjust the pH to 6.5 with KOH. Add minerals in order, allowing each one to dissolve before adding the next mineral, and adjust pH to

7.0.

S2A.2.4 Preparation of 2.5 % Sodium Sulfide solution

(1) Add 100 mL ddH2O to a flask and mark the water line. To limit the formation of volatile

hydrogen sulfide from sodium sulfide, add one pellet of NaOH (about 0.1 g). Add 10

mL more ddH2O to the flask.

(2) Boil the 110 mL ddH2O while flushing with N2 until the water level reaches the marked

100 mL water line.

(3) Let flask cool while flushing with N2, transfer the flask to the gassing station in the

fume hood. Continue to flush with N2.

(4) While flask is cooling, weigh out slightly more than 2.5 g Na2S·9H2O. Wear gloves

and do the subsequent steps in the fume hood. Clean the sodium sulfide crystal by

briefly rinsing the crystal with diH2O. To do this, swirl the crystal in a small beaker

with some water until it is clean. Blot dry the crystal with paper towel. Re-weigh the

crystal to ensure that the final weight is 90-110 % of the desired weight.

(5) Add the cleaned and weighted sodium sulfide to the cooled flask while flushing with

N2 and mix until partially dissolved.

(6) Stopper the flask, discontinue flushing, and transfer to the anaerobic chamber. Dispense

into 5 mL aliquots in 28 mL aluminum sealed tubes.

(7) Remove tubes from chamber and pressure with N2 at 103 kPa. 162

(8) Autoclave on gravity cycle (rapid exhaust) for 20 minutes.

(9) Store these sodium sulfide tubes in anaerobic chamber. They should remain good for

one to two months. Discard if a precipitant forms.

S2A.2.5 Preparation of 25 % Sodium Sulfide solution

(1) Prepare as above but with 25 g Na2S·9H2O. Dispense into 5 mL aliquots in 28 mL

aluminum sealed tubes. For cultures of 0.5, 1.0, and 1.5 L, use 1.0, 2.0, and 3.0 mL,

respectively.

(2) Store these sodium sulfide tubes in anaerobic chamber. They should remain good for

two months. Discard if a precipitant forms.

S2B. Solid medium for formate growth (McF) of Methanococcus maripaludis

This protocol prepares 1 % agar medium in 70 mL serum bottles for plating M. maripaludis and is modified from Tumbula et al. (1).

(1) Solid medium is prepared with 1 % agar (w/v), adding 10 mL of medium to a 70 mL

serum bottle. Add 0.1 g agar into each individual serum bottle prior to medium

preparation. The remaining ingredients are the same as for the broth.

(2) Transfer the broth medium [without sulfide] and the agar-containing bottles to the

anaerobic chamber, dispense medium into agar bottles, seal the bottles with grey butyl

stoppers (Wheaton Science Products, cat. no.: W224100-202), and crimp with

aluminum seals (Fisher Scientific, cat. no.: 11-126-12).

(3) Remove the sealed bottles from the anaerobic chamber; pressurize each bottle to 103

kPa with N2/CO2 (4:1, v/v) and autoclave. Autoclave on gravity cycle (rapid exhaust)

for 20 minutes. After autoclaving, allow the bottles to cool to touch and add sulfide to

163

a final concentration of 2 mM. Mix gently and allow the agar bottles to cool completely

on their sides for at least five hours. (Overnight solidification works best).

(4) To obtain individual colonies of M. maripaludis, make a serial dilution in broth culture

to about 100 cells/mL. Plate the diluted and undiluted culture (positive control) by

adding 0.5 mL of the culture suspension to agar bottles using 1 mL syringes. Add the

liquid to the agar surface, with the bottle resting on its side. Gently swirl the agar bottle

to allow the culture suspension to become evenly distributed on the agar surface. Rest

the agar bottle on its side for another four hours to allow the culture suspension to be

fully absorbed into the agar. Make sure the agar surface is flat and sitting horizontally,

otherwise the colonies will only form on one side of the agar.

(5) Incubate the plated bottles by standing them upright at 37°C. Hence, the water that

forms during incubation will drain to the bottom of the bottle.

(6) Colonies will appear after three to five days of incubation. Because of the liquid

collecting in the bottom of the bottle, confluent growth will occur on the bottom 1 cm.

Avoid shaking the bottle during incubation to prevent inadvertent inoculation of the

upper portion of the agar.

(7) To pick individual colonies into broth in the anaerobic chamber, add 2 mM sulfide

(sulfide stock preparation see S2A) into tubes of broth 24 hours before use. Open the

stopper of agar bottle in the anaerobic chamber, sterilize the opening with an electric

heating element, insert a 1 mL syringe into the agar bottle, carefully touching the top

of a single colony with the needle. Then stab the needle into a tube of broth, and flush

the syringe with medium two times to introduce some cells into the broth. Full growth

is often observed after 1-2 days. 164

S2C. Medium-scale cultivation system for M. maripaludis and M. okinawensis

This protocol is suitable for growth of 1.5 L cultures on formate medium using glassware modified to allow venting of the gas produced.

S2C.1 Day 1. Medium preparation and degassing:

(1) Preparation of the inoculum. One day before initiating the medium-scale culture,

inoculate 0.3 mL of fresh culture into one or more serum bottles containing 30 mL of

formate medium. Incubate without shaking at the proper temperature.

(2) Dispense 1.5 L McF medium into a modified 2 L storage bottle (Figure S2-1).

(3) To sparge the medium after autoclaving, insert a 2cc interchangeable glass-filter

syringe (Micro-mate, cat. no.: 14-825-1B) with luer-to-tubing connector (Sigma-

Aldrich, 1/16-3/32 in., cat. no.: Z118028) into a butyl rubber stopper (Bellco Glass,

Inc., Vineland, NJ, cat. no.: 2048-11800). To the tip of the inserted needle, attach

approximately 20 cm of thin tubing (Zeus PTFE light wall tubing, P/N: TFT22-NT,

AWG 22, LOT#: 623928). Insert the stopper into the side arm of the culture vessel. The

stopper is not sealed in place so that it can fall out and release any pressure that might

form in case the exhaust lines become clogged.

(4) With the cap of the 2 L bottle loosely attached, autoclave this assembly together with

the gas trap (165 mL serum bottle, sealed with blue butyl rubber stopper and crimped

with aluminum seals), tubing and syringes that connect the medium bottle, gas trap and

water trap.

(5) After autoclaving, allow the sterile medium to cool to at least 90 oC before assembling

the gassing and vent lines. First, begin sparging N2/CO2 (4:1 v/v) through the medium

at a flow rate of ~25 mL/min. At the same time, connect the medium bottle to the gas 165

trap and water trap. Add about 300 mL of water to the water trap. Close the cap to the

2 L bottle to begin gas flow through the exhaust system.

(6) Sparging should exchange the headspace of the bottle at least ten times. At a flow rate

of 25 mL/min, the medium should be sparged for 200 min or about 5 L of gas.

(7) While sparging, add cysteine hydrochloride to a final concentration of 0.5 g l-1. To

prepare this cysteine hydrochloride stock solution, affix a sterile 0.2 µM filter

(WhatmanTM, cat. no.: 6780-2502) to a 5 mL syringe with Luer-Lok tip (BDTM, cat. no.:

309646) with the plunger removed. Add 3 ml ddH2O to the syringe, then add 0.75 g of

cysteine hydrochloride (Sigma, cat. no.: 52-89-1), Swirl the solution gently to fully

dissolve the cysteine. Put a needle (BDTM, 15 G × 3 1/2 in., cat. no.: 511108) onto the

tip of the 0.2 µM filter and quickly inject the freshly made cysteine hydrochloride

solution into the 1.5 L culture bottle through the side-arm stopper.

(8) When the gassing is complete and the medium has cooled to the growth temperature or

below, incubate the entire assembly at the growth temperature, 37 oC for M.

maripaludis or 62 oC for M. okinawensis, overnight.

S2C.2 Day 2. Inoculation and following incubation

(1) The next morning, add sodium sulfide to 2 mM with a syringe through the side-arm. It

is often convenient to add 3 mL of a 25 % (w/v) sterile solution prepared as in S2A.

Continue to incubate the medium at the growth temperature for another six hours to

allow the medium to become fully reduced.

(2) Inoculate the bottle with the 30 mL of culture (approximately OD600nm=0.6) from day

o 1. Overnight or about 15 h, M. maripaludis S2 can grow to an OD600nm=0.8 at 37 C.

166

o Similarly, M. okinawensis IH1 can grow to about OD600nm=0.6 at 62 C.

S2D. Rapid preparation of formate medium for growing Methanococcus maripaludis

This rapid protocol is designed for laboratories that have limited anaerobic equipment. All procedures can be performed without using an anaerobic chamber.

S2D. 1 Liquid medium preparation

(1) On the bench, combine all medium ingredients as described in S2A. Add the cysteine

hydrochloride (0.05 g/100 mL) without sparging the medium shortly before dispensing

the medium into tubes.

(2) Dispense medium into aluminum sealed tubes aerobically. Seal each tube with a grey

butyl stopper (Wheaton Science Products, cat. no.: W224100-202) and crimp with

aluminum seals (Fisher Scientific, cat. no.: 11-126-12).

(3) Bring the sealed tubes to a gassing station as shown in Figure S2-2. Run three gas-

exchange cycles on each tube. Each cycle is composed of 45 s of vacuum followed by

15 s of pressurization at 103 kPa by N2/CO2 (4:1, v/v). At this point, the medium may

still be pink because the resazurin is not reduced. Upon autoclaving, the medium should

become colorless. If using serum bottles with larger volumes, increase the cycle times.

For instance, 160 mL serum bottles, 90 s vacuum and 30 s pressurization is sufficient

for a system with a strong vacuum line.

(4) Autoclave on gravity cycle (rapid exhaust) for 20 min.

S2D. 2 Solid medium preparation

(1) The rapid protocol prepares 1 % agar medium in 70 mL serum bottles.

167

(2) On the bench, combine all medium ingredients as described in S2A, add the cysteine

hydrochloride (0.05 g/100 mL) without sparing the medium but shortly before

dispensing into the bottles.

(3) Solid medium is prepared with 1 % agar (w/v) to 10 mL of medium in a 70 mL serum

bottle. Add 0.1 g agar into each serum bottle prior to medium preparation.

(4) Dispense medium into agar-containing serum bottles aerobically. Seal each bottle with

a grey butyl stopper (Wheaton Science Products, cat. no.: W224100-202) and crimp

with aluminum seals (Fisher Scientific, cat. no.: 11-126-12).

(5) Bring sealed bottle to the gassing station as shown in Figure S2-2. Run three gas-

exchange cycles on each bottle, each cycle is composed of 45 s of vacuum followed by

15 s of pressurization to. 103 kPa by N2/CO2 (4:1, v/v). At this point, the medium may

still be pink because the resazurin is not reduced. Upon autoclaving, the medium will

become colorless.

(6) Autoclave on gravity cycle (rapid exhaust) for 20 min. After autoclaving, allow the

bottles to cool to touch, using the gassing station to anaerobically add sodium sulfide

at a final concentration of 2 mM (for sulfide stock solution preparation, see S2A). Mix

gently and then allow the agar bottles to cool completely on their sides for at least five

hours. (Overnight solidification works best).

S2D. 3 Plating and picking colonies on the bench

The Hungate technique (2) is used to make anaerobic transfers at the gassing station.

Briefly, before a transfer, flame-sterilize a cannula and begin the flow of N2 gas. Flush a sterile syringe with N2 by inserting the needle into the cannula and pumping the syringe three times to remove air. Fill the syringe with N2 gas. After sterilizing the top of the stopper, immediately expel 168

the N2 from the syringe while pushing the needle through the stopper. This procedure removes any air that was introduced into the tip of the needle.

(1) Plating is performed as described in S2B. steps 4-6., but using the gassing station for

all anaerobic transfers.

(2) To pick individual colonies into broth, use the Hungate technique (2). At the gassing

station, remove the aluminum crimp sealed stopper from the agar bottle, and flame the

opening. To keep the serum bottle O2-free, introduce a flow of N2 gas with a sterile N2

gassing cannula (Figure S2-3A). To transfer a colony, use a 1 mL syringe and 22 Ga×1-

inch needle. Bend the needle about 45 degrees by folding the end of the needle against

the needle cap. Be sure that the hole on the tip of the needle is on the bottom side after

the bend (Figure S2-3B). This makes it easier to collect a colony inside the needle tip.

Insert this syringe into the agar bottle, pumping the plunger it a few times to fill the

syringe with N2. With the syringe barrel pulled back about 1 cm, carefully touch the

top of a single colony with the needle (Figure S2-3A). Withdraw the syringe from the

bottle and stab the needle into a tube of broth medium. Flush the syringe two times with

0.2 mL of medium to wash the cells from the needle. Full growth is often observed

after 1-2 days.

References:

1. Tumbula DL, Makula RA, Whitman WB. 1994. Transformation of Methanococcus

maripaludis and identification of a PstI-like restriction system. FEMS Microbiol Lett

121:309-314.

2. Bryant MP. 1972. Commentary on the Hungate technique for culture of anaerobic bacteria.

Am J Clin Nutr 25:1324-8. 169

3. Balch WE, Wolfe RS. 1976. New approach to the cultivation of methanogenic bacteria: 2-

mercaptoethanesulfonic acid (HS-CoM)-dependent growth of Methanobacterium

ruminantium in a pressureized atmosphere. Appl Environ Microbiol 32:781-91.

4. Whitman WB, Shieh J, Sohn S, Caras DS, Premachandran U. 1986. Isolation and

characterization of 22 mesophilic methanococci. Syst Appl Microbiol 7:235-240.

5. Romesser JA, Wolfe RS, Mayer F, Spiess E, Walthermauruschat A. 1979. Methanogenium,

a New genus of marine methanogenic bacteria, and characterization of Methanogenium

cariaci sp. nov. and Methanogenium marisnigri sp. nov. Arch Microbiol 121:147-153.

170

Figure S2-1. Medium scale cultivation system with culture bottle, sparging and exhaust. The cultivation system comprises a 2 L cultivation bottle (A), water trap (B: 160mL serum bottle with a blue butyl stopper and an aluminum seal) and a gas trap (C: 150 mL Erlenmeyer flask). The cultivation bottle is constructed from a from Pyrex bottle (cat. no.: 06-414-1E) with the top of a

28-mm aluminum seal tube attached as a side-arm. The side-arm is sealed with a blue butyl rubber stopper without the aluminum seal (Bellco Glass, Inc., Vineland, NJ, cat. no.: 2048-11800).

Leaving off the aluminum seal allows the stopper to pop off and release any pressure that might form if the exhaust line becomes clogged. The gassing line (D, E, and F) is for sparing the medium after autoclaving and is assembled with F, a 2-cc interchangeable glass-filter syringe barrel (Micro mateTM, cat. no.: 14-825-1B) packed with cotton and sealed with a number 00 rubber stopper

[Balch et al., (3)], D: luer-to-tubing connector (Sigma-Aldrich, 1/16- 3/32 in., cat. no.: Z118028) and E: needle (BDTM, 22 G×1 in., cat. no.: 305155) with thin tubing (ZeusTM PTFE light wall tubing, P/N: TFT22-NT, AWG 22) connected. After sparging, the needle is removed prior to inoculation leaving the tubing in the culture. Other components include G: NalgeneTM tubing (80

PVC tubing-FDA/USPVI, 1/8 ID × 3/16 OD × 1/32 wall, Thermo scientific cat. no.: 8000-0010);

171

H: MonojectTM needle (16 G × 1 1/2", cat. no.: 140394); I: needle (BDTM, 15 G × 3 1/2 in., cat. no.: 511108).

172

Figure S2-2. Gassing manifold for exchanging headspace of culture tubes. (A) Vacuum-pressure gauge (Swagelok, cat. no.: PGI-63C-OC60-LAQX); (B) Three-way valve with Swagelok fittings

(Swagelok, cat. no.: B-43XS4) for connection to sources of O2-free gas and vacuum; (C) Swagelok

Tube Fitting, Union Cross, 1/4 in. (Swagelok, cat. no.: B-400-4); (D) Brass Swagelok Tube Fitting,

Reducer, 1/8 in. x 1/4 in. (Swagelok, cat. no.: B-200-R-4); (E) Thin bore polyethylene tubing, 1/8 in. (Freelin Wade Co. cat. no.: 1A-109-01); (F) PFA Swagelok Tube Fitting, Reducing Union, 1/4 x 1/8 in. (Swagelok, cat. no.: PFA-420-6-2); (G) 1 cc plastic syringe barrel (BDTM, cat. no.:

309659; cut off grip on the end of the barrel to allow insertion into the union); (H) Disposable needle (BDTM, 22 G × 1in., cat. no.: 305155).

173

Figure S2-3. Scheme of picking colonies from agar bottles at the gassing station. (A) A 70 mL serum bottle plated with M. maripaludis S2 forms colonies after 3 days. After opening, a sterile

N2 gassing cannula was introduced (top) to prevent air from entering the bottle and to maintain anaerobiosis. A 1 mL syringe with a 22 G × 1-inch needle was used for transferring the colony

(bottom). To pick the colony more accurately, the needle was bent about 45 degrees by folding the end of the needle against the needle cap. (B) Enlarged image of the 1mL syringe with the bent needle.

174

Table S2-1. Compositiona of basal medium for formate growth (McF) of Methanococcus maripaludis.

Component For tubes For 1 liter bottle For 100 mL For 1000 mL Glass-distilled water 30 mL 300 mL Glycylglycine buffer, 1 M, pH= 8.0 20 mL 200 mL General salts solution 50 mL 500 mL

K2HPO4, 14g/L 1.0 mL 10 mL

Na acetate·3H2O, 136 g/L 1.0 mL 10 mL Trace mineral solution (4) 1.0 mL 10 mL Iron stock solution (3) 0.5 mL 5 mL Resazurin, 0.1 g/100 mL 0.1 mL 1 mL Sodium formate (NaCOOH) 2.7 g 27 g

Sodium bicarbonate (NaHCO3) 0.5 g 5.0 g Casamino acids (for complex medium) 0.5 g 5.0 g Alanine (optional, 100 mM) 1.0 mL 10 mL aMedium components are based upon Balch et al. (3), Romesser et al. (5), and Whitman et al. (4).

175

Table S2-2. Composition of basal medium for formate growth (McF) of Methanothermococcus okinawensis.

Component For tubes For 1 liter bottle For 100 mL For 1000 mL Glass-distilled water 10 mL 100 mL Glycylglycine buffer, 1 M, pH= 6.5 40 mL 400 mL General salts solution 50 mL 500 mL

K2HPO4, 14 g/L 1.0 mL 10 mL

Na acetate·3H2O, 136 g/L 1.0 mL 10 mL Trace mineral solution 1.0 mL 10 mL Iron stock solution 0.5 mL 5 mL Resazurin, 0.1 g/100 mL 0.1 mL 1 mL Sodium formate (NaCOOH) 2.7 g 27 g

Sodium bicarbonate (NaHCO3) 0.5 g 5.0 g Casamino acids (for complex medium) 0.5 g 5.0 g

176

Table S2-3. Composition of general salts solution.

Composition g/L Medium concentration (mM) KCl 0.67 4.5

MgCl2·6H2O 5.50 13.5

MgSO4·7H2O 6.90 14.0

NH4Cl 1.00 9.0

CaCl2·2H2O 0.28 0.95

177

Table S2-4. Composition of trace minerals solution.

Composition g/L Medium concentration (µM) Nitriloacetic acid 1.5 78

MnSO4·2H2O 0.1 5.3

Fe(NH4)2(SO4)2·H2O 0.2 5.1

CoCl2·6H2O 0.1 4.2

ZnSO4·7H2O 0.1 3.5

CuSO4·5H2O 0.01 0.4

NiCl2·6H2O 0.025 1.1

Na2SeO3 0.2 11.6

Na2MoO4·2H2O 0.1 4.1

Na2WO4·2H2O 0.1 3.0

178

APPENDIX B

CHAPTER 3 SUPPLEMENTARY INFORMATION2

Supplementary figure S3-1. Dissimilatory sulfur reduction pathway.

180

Supplemental figure S3-2. Growth of Methanococcus maripaludis S2 on sulfide (black), elemental sulfur (red) or no sulfur source (grey). Cells were grown in McNAA medium that reduced with 2 mM titanium (III) citrate. 2 mM sulfide or 0.02 g/mL (w/v) elemental sulfur was added as the sulfur source. The inoculum size was 107 cells per 5 mL culture. All values were the averages of three independent cultures.

181

Supplemental figure S3-3. Verification of the complete deletion of mmp1681 from

Methanococcus maripaludis genome (A-D) and the incorporation of plasmid pMEV4-MMP1681 into strain S212 (E).

A. DNA fragment representing a portion of ∆mmp1681 strain (S211) genome, suicide plasmid p5L-U DNA and M. maripaludis wild-type S0001 genome. Blue arrows indicate the binding sites for primers MMP1681UpF and MMP1681DownR, which amplify the upstream to downstream of mmp1681. Green arrows indicate the binding sites for primers p5LU-F1 and p5LU-R1, which amplify the pac cassette. Grey arrows represent the binding sites for primers InnerMMP1681F and

InnerMMP1681R, which amplify the deleted fragment of S211 strain. 182

B. PCR amplification of the fragment from upstream to downstream of mmp1681 using primers

MMP1681UpF and MMP1681DownR. Lanes: 1, standard 1-kb ladder (New England Biolab); 2, genomic DNA of the mutant strain S211. 3, DNA of the plasmid p5L-U. 4, genomic DNA of wild- type M. maripaludis S0001.

C. PCR amplification of the pac cassette using primers p5LU-F1 and p5LU-R1. Lanes: 1, standard

1-kb ladder (New England Biolab); 2, genomic DNA of the mutant strain S211. 3, DNA of the plasmid p5L-U. 4, genomic DNA of wild-type M. maripaludis S0001.

D. PCR amplification of an internal portion of mmp1681 using primers InnerMMP1681F and

InnerMMP1681R for a mixture of different proportions of genomic DNA of the mutant strain

∆mmp1681 and wild-type strain S0001 Lanes: 1, standard 1-kb ladder (New England Biolab); 2,

100 % genomic DNA of the mutant strain S211; 3, 99.9 % genomic DNA of strain S211 and 0.1 % of wild-type strain S0001; 4–9, 99.5 %, 99 %, 98 %, 95 %, 90 %, and 0 % of genomic DNA of the mutant strain ∆mmp1681 and the balance genomic DNA of wild-type strain S0001; 10, DNA of the plasmid p5L-U and 11, negative control without template.

E. PCR amplification of the region of promoter Pmcr to mmp1681 on plasmid extracted from S212 genome, using primers pAW5042-F and pAW5042-R. Lanes: 1, standard 1-kb ladder (New

England Biolab); 2, genomic DNA of wild-type strain S0001; 3, plasmid DNA of strain S212.

183

Supplemental figure S3-4. SDS-polyacrylamide gel (12 %) analysis of His-tag purified recombinant E. coli PAPR, MMP0941 and MMP1681. The gel was stained with AcquaStain. Lane

1, Color Prestained Protein Lader, 11-245 kDa (NEB); lane 2, 1 μg of purified recombinant E. coli

PAPR; lane 3, 1 μg of purified recombinant MMP0941; lane 4, 10 μg of purified recombinant

MMP1681.

184

0.7

0.6

0.5

0.4

0.3

0.2 Absorbance Absorbance (600nm) 0.1

0 0 20 40 60 80 100 Time (h) Supplemental figure S3-5. Growth of Methanococcus maripaludis S0001 (circle), S211 strain

(triangle) in McNAA medium with (filled) or without (open) the addition of 14 mM sodium sulfate.

McNAA medium was reduced with 2 mM dithiothreitol and 2 mM sulfide was added as sulfur source. The inoculum size was 6×104 cells per 5 mL culture. All values were the averages of three independent cultures.

185

1.2

0.8

0.6

0.4 Absorbance Absorbance (600nm) 0.2

0 0 10 20 30 40 50 60 Time (h) 3-mercaptopropionate acid 0mM 0.1mM 1mM 5mM 10mM

Supplemental figure S3-6. Growth of Methanococcus maripaludis S2 in McFAA medium with

3-mercaptopropionate acid ranging from 0, 0.1, 1, 5, 10 mM. McFAA medium was reduced with

2 mM dithiothreitol, and 2 mM sulfide was added as sulfur source. The inoculum size was 2×105 cells per 5 mL culture. All values were the averages of three independent cultures.

186

Supplemental figure S3-7. Absorbance spectrum of anaerobically purified recombinant

MMP1681 (A) and MMP0941 (B) upon exposure to air. The recombinant MMP1681 was at 81

µM in 50 mM sodium HEPES (pH= 8.0) buffer with 500 mM NaCl. The recombinant MMP0941 was at 92 µM in 50 mM sodium HEPES (pH= 8.0) buffer with 500 mM NaCl. These two recombinant proteins were over-expressed in E. coli by the pQE2 expression system (Qiagen) with

N-terminal 6×His tag and anaerobically purified using gravity flow Ni-NTA column. Time of exposure to air was 0 h (balck), 1 h (yellow), 3 h (blue) and 24 h (red).

187

Supplemental figure S3-8. Complementing E. coli Keio mutants invovled in the assimilatory sulfate reduction pathway using putative Methanococcus maripaludis protein homologs. E. coli

Keio mutants are highlighed in blue and Methanococcus maripaludis proteins are highlighed in red.

188

Supplementary table S3-1. List of strains, plasmids and primers.

Strains Description Source M. maripaludis S2 Wild-type M. maripaludis Another Wild-type strain, descent from M. maripaludis S2 M.S0001 maripaludis S211 Knockout mutant strain of mmp1681 from M. maripaludis S0001 This work M. maripaludis S212 M. maripaludis strain 211 complemented with pMEV4-mmp1681 This work E. coli M15 (pREP4) E. coli expression strain for vector pQE2 Qiagen E. coli JW2720 E. coli ΔcysC mutant strain from the Keio collection (1) E. coli JW2722 E. coli ΔcysD mutant strain from the Keio collection (1) E. coli JW2732 E. coli ΔcysH mutant strain from the Keio collection (1) E. coli JW2733 E. coli ΔcysI mutant strain from the Keio collection (1) E. coli JW2734 E. coli ΔcysJ mutant strain from the Keio collection (1) E. coli JW3331 E. coli ΔcysG mutant strain from the Keio collection (1) Plasmid P5L-D E. coli cloning vector for deleting M. maripaludis gene (2) P5L-U E. coli cloning vector for deleting M. maripaludis gene (2) pMEV4 Shuttle vector for M. maripaludis (3) pQE2 E. coli expressive vector Qiagen Primer MMP1681DownF 5’ - GCACTTAAGGGCGATGCAAATTTTGTGTCT This work MMP1681DownR 5’ -TGCTCTAGAAAGCTTTTTTAGGGGCTGTTGGCGAT This work MMP1681UpF 5’ - GGAATTCCATATGGGATATGAGCACTTCATGTTAAAAGA This work MMP1681UpR 5’ - CGGGGTACCAGAACTCGAAGAAAATATGGGTGT This work InnerMMP1681F 5’ - TTTAACCCATCTTTCAAAGTAGTCAGG This work InnerMMP1681R 5’ - TTCAGAAGATCAGATTGGAACACTC This work pMEV4-MMP1681F 5’ - GCATCTAGATGTGAGGTGAAATAATGTTTAATGAAGACACAAAATTTGCATC This work pMEV4-MMP1681R 5’ - TCGCTGCAGTTACAGTTGAGTAATTCTTTTTAATCTTTTC p5LU-F1 5’ - CTAGAAATAGGTGAAATGCATGAGC (2) p5LU-R1 5’ - TTACTAAATTATGCTCCTGGTTTTCTTG (2) pAW5042-F 5’ - TCCTTCCTTCTTTCCTGCATACT (3) pAW5042-R 5’ -GTCATGGAAGGTCGTCTCC (3) pQE-Forward 5’ - CCCGAAAAGTGCCACCTG Qiagen pQE-Reverse 5’ - GTTCTGAGGTCATTACTGG Qiagen RecMMP1681F 5’ - CGCCATATGTTTAATGAAGACACAAAATTTGCATC This work RecMMP1681R 5’ - TCGGAGCTCTTACAGTTGAGTAATTCTTTTTAATCTTTTC This work RecMMP0941F 5’ - CGCCATATGAAGACTATTTTAGGAAAAATTC This work RecMMP0941R 5’ - TCGGAGCTCCTATCCAAGCCAGTTTTCATTT This work RecMMP0078F 5’ - CGGGGTACCATGAACAAAAATGAAATAGCCAATCTG This work RecMM0078R 5’ - CCCAAGCTTTTAATAATCTTTTAATTTTTCTTTTAAAGAAC This work RecMMP1282F 5’ - CGGGGTACCATGAAGTTAATCGGAATTACTGGAATGCC This work RecMMP1282R 5’ - CCCAAGCTTTTAATTTACGTTAATTATTTTTTTAAACGTGTTTTC This work RecMMP1283F 5’ - CGGGGTACCATGTCTGTTGCTTTAAAGAAGTTTTTTTCG This work RecMMP1471F 5’ - CGGGGTACCATGGAAGACGAAATAAAATCATTTGAAAAAG This work RecMMP1471R 5’ - CCCAAGCTTTTATTCATGATTATTTATCCCGTTTAAAATTC This work RecMMP1472R 5’ - CCCAAGCTTTTACTTTTTATCCCATAAATCAAGATATTCTTG This work RecCysHF 5’ - CGCCATATGTCCAAACTCGATCTAAACGC This work RecCysHR 5’ -TCGGAGCTCTTACCCTTCGTGTAACCCACAT This work

189

Supplementary table S3-2. Genera of Archaea and bacteria that are able to reduce elemental sulfur to H2S. Modified from (4, 5).

o Topt ( C) pHopt Electron donors Reference Archaea Crenarchaeota: Acidianus 70-90 1.5-2.0 H2 (6) Acidilobus 85 3.8 Yeast extract, beef extract and soya extract (7) Stygiolobus 80 2.5-3.0 H2 (8) Pyrobaculum 102 6.0 H2, peptone, extracts of meat and yeast, bacterial and (9) Thermofilum 85-90 5.0-6.0 Peptidesarchaeal cell homogenates (10, 11) Thermoproteus 85-90 5.0-6.5 H2, peptides, maltose, formate, fumarate, ethanol, (12, 13) Desulfurococcus 85-90 6.0-6.4 Peptides,malate, methanol,starch, pectin, glycogen, glycogen, starch, yeast extract,amylopectin, casein (14, 15) Sulfolobus 80 2-4 Glucose,hydrolysateformamide pentose (16) Igneococcus 90 5.5-6.0 H2 (5) Pyrodictium 105 5.5-6.0 H2 (17, 18) Stetteria 95 6.0 H2 (19) Thermodiscus 88 5.5 H2, yeast extract (12) Thermosphaera 85 6.5 Yeast extract, peptone (20) Thermocladium 75 4 glycogen, starch, gelatin and various proteinaceous (21) Staphylothermus 92 6.5 Peptone,complex compoundsextracts of meat and yeast (22) Hyperthermus 95-107 7.0 Tryptone, peptone (23) Vulcanisaeta 85-90 4.0-4.5 Yeast extract, peptone, beef extract, Casamino acids, (24) Euryarchaeota: gelatin, starch, maltose and malate Pyrococcus 96-100 6.8-7.0 Complex substrates, amino acids, starch, maltose, (25, 26) Thermococcus 75-88 5.8-9.0 Peptides,pyruvate amino acids, sugars, starch, chitin, pyruvate (27, 28) Caldococcus 88 6.4 Peptides (29) Caldisphaera 70-75 3.5-4.0 Starch, glycogen, gelatin, beef extract, yeast extract (30) Thermoplasma 59 1.0-2.0 Extractsand peptone of yeast, meat, and bacteria (31) Methanopyrus 98 6.5 H2 (32) Methanobacterium 37-65 7.0 H2 (32) Methanothermus 88 6.5 H2 (32) Methanococcus 85-90 6.0 H2, formate (32) Bacteria Aquifex 85 6.8 H2, sulfur, thiosulfate (33) Ammonifex 70 7.5 H2 (34) Desulfurobacterium 70 6.0 H2 (35) Desulfuromonas 37 7.5 Acetate, pyruvate, ethanol (36) Desulfuromusa 35 6.5-7.0 Acetate, propionate (37) Desulfurella 55 7.0 Acetate (38, 39) Desulfovibrio 37 7.2 Organic acids, alcohols (40) Fervidobacterium 65-70 6.5-7.0 Sugars, pyruvate, yeast extract (41, 42) Geobacter 35 6.5-7.0 Acetate (43) Pelobacter 37 6.5-7.0 H2, ethanol (44) Shewanella 30 6.5-7.0 Lactate (45) Sulfospirillum 37 6.5-7.5 H2, formate (46, 47) Thermotoga 66-80 6.5-7.5 Sugars, peptone, yeast extract, bacterial and archaeal (48, 49) Thermosipho 70-75 6.5-7.5 Yeastcell homogenates extract, brain heart infusion, peptone, tryptone (50, 51) Wolinella 37 8.5 Tryptone (52)

190

Supplementary table S3-3. The level of inorganic sulfide in McNAA medium with different treatments.

a b b Medium Type H2S (aq) (μM) H2S(g) (μM per liter medium) not autoclaved < 1.5 < 1.5 Medium autoclaved < 1.5 < 1.5 not autoclaved < 1.5 < 1.5 Medium+S0 autoclaved 4 ± 1 32 ± 1 not autoclaved < 1.5 < 1.5 Medium+DTT autoclaved < 1.5 < 1.5 not autoclaved 203 ± 10 178 ± 1 Medium+DTT+S0 autoclaved 163 ± 1 149 ± 1 a McNAA medium was McNA medium supplemented with 1 mM alanine, reduced or not reduced with 3mM dithiothreitol (DTT), together with or without 0.1 g elemental sulfur (S0) per 5mL medium. The medium tubes were autoclaved or not autoclaved to test if heat affected the production of sulfide. All medium tubes were incubated in anaerobic chamber overnight at room temperature before measuring inorganic sulfide content. b Values were means ± 1 S.D., obtained from three independent samples and represent inorganic sulfide concentration either in the aqueous phase (aq) or gaseous phase (g).

191

Supplementary table 4. Quantitative proteomic analysis of Methanococcus maripaludis S2.

Protein ORF Na SDc Annotationd ratiob

MMP0008 1 1.25 DNA polymerase, archaeal type II, small subunit

MMP0011 2 0.46 1.28 Restriction endonuclease subunit M

MMP0023 2 1.22 0.02 Conserved hypothetical protein

MMP0030 1 0.69 DNA replication protein; MCM family; putative

MMP0045 1 1.42 Bifunctional short-chain isoprenyl diphosphatesynthase

7,8-didemethyl-8-hydroxy-5-deazariboflavin synthase subunit MMP0057 1 1.47 CofH

F -dependent N5, N10 methylenetetrahydromethanopterin MMP0058 3 0.57 0.10 420 reductase

MMP0059 1 0.79 Conserved hypothetical archaeal protein

MMP0060 2 1.68 0.04 Ribosomal LX protein

MMP0063 2 1.12 0.07 Acetylglutamate kinase

MMP0064 1 1.27 Nitrogen regulatory protein P-II

MMP0066 1 1.10 Nitrogen regulatory protein P-II

MMP0073 6 1.11 0.15 Argininosuccinate synthase

MMP0076 6 0.99 0.12 Conserved hypothetical protein

MMP0080 1 1.42 Glutamate synthase; large subunit; archaeal subunit 1

MMP0083 1 1.16 Conserved archaeal protein

MMP0089 1 0.94 Conserved hypothetical protein

MMP0092 3 1.01 0.26 DNA-directed RNA polymerase subunit F

MMP0103 1 0.40 Conserved hypothetical protein

MMP0112 4 0.87 0.26 Phosphoglycerate mutase

MMP0119 2 1.59 0.03 Biotin-[acetyl-CoA-carboxylase] ligase

MMP0123 1 1.08 Phosphoribosylglycinamide formyltransferase 2

H -forming N5, N10-methylene-tetrahydromethanopterin MMP0127 1 3.50 2 dehydrogenease

192

MMP0140 1 1.72 Conserved hypothetical protein

MMP0154 2 1.19 0.19 Conserved hypothetical archaeal protein

MMP0156 2 0.97 0.03 Ribosomal protein S19E (S16A)

MMP0157 1 0.75 Conserved hypothetical protein

MMP0164 2 1.51 0.41 Sirohydrochlorin cobaltochelatase

MMP0176 6 1.12 0.11 CDC48 cell division cycle protein family member

MMP0177 2 0.90 0.38 Conserved hypothetical archaeal protein

MMP0178 1 1.35 Phosphoribosylformylglycinamidine synthase I

MMP0208 1 0.71 Conserved hypothetical protein

MMP0250 2 1.03 0.04 Conserved hypothetical protein

MMP0251 2 1.09 0.02 Proteasome, subunit alpha

MMP0258 21 1.05 0.18 LSU ribosomal protein L12A

MMP0260 1 1.15 LSU ribosomal protein L1P

MMP0261 2 0.86 0.45 DNA directed RNA polymerase; subunit L

MMP0272 4 0.94 0.18 ABC transporter ATPase subunit

MMP0278 1 0.90 Conserved hypothetical protein with 2 CBS domains

MMP0283 1 2.67 Nucleoside-diphosphate kinase

MMP0297 1 0.97 Translation initiation factor IF-2

MMP0298 2 1.11 0.10 LSU ribosomal protein L15E

MMP0304 3 1.56 0.02 Conserved hypothetical archaeal protein

MMP0308 1 1.91 Conserved hypothetical archaeal protein

MMP0309 1 0.93 Intracellular sulfur reduction related protein

MMP0312 2 0.95 0.00 Conserved hypothetical archaeal protein

MMP0317 2 1.29 0.18 Conserved hypothetical protein

MMP0326 1 1.61 Methionyl-tRNA synthetase

MMP0327 1 1.74 Thymidine phosphorylase

MMP0332 1 1.38 Conserved hypothetical protein

193

2-Hydroxyglutaryl-CoA dehydratase (component D) related MMP0346 1 1.18 protein

MMP0358 1 0.99 Conserved hypothetical archaeal protein

MMP0371 2 1.04 Conserved hypothetical protein

F -dependent MMP0372 7 0.33 0.64 420 methylenetetrahydromethanopterindehydrogenase

MMP0383 18 1.17 0.16 S-layer protein

MMP0386 27 1.43 0.13 Archaeal histone A

MMP0389 1 1.04 Ferredoxin

MMP0397 2 0.82 0.42 Alanyl-tRNA synthetase

MMP0411 1 1.54 Sulfopyruvate decarboxylase subunit alpha

MMP0413 1 0.94 Conserved hypothetical protein

MMP0414 1 1.27 Threonyl-tRNA synthetase

MMP0423 1 0.97 Hypothetical protein

Putative Archaeoglobus fulgidus predicted coding region MMP0437 1 0.58 AF2307

MMP0447 1 0.61 Nitrogenase related protein

Possible Archaeoglobus fulgidus predicted coding region MMP0458 1 0.76 AF1474

MMP0460 1 0.73 Conserved Hypothetical Protein

MMP0478 4 1.18 0.21 Conserved hypothetical protein

MMP0481 1 0.93 Conserved hypothetical protein

MMP0516 3 1.11 0.12 Molybdenum ABC transporter, permease protein

MMP0529 2 1.42 0.01 Sulfate transporter family

MMP0540 2 1.52 0.02 Phosphoribosylaminoimidazole-succinocarboxamidesynthase

MMP0559 1 0.98 Family of unknown function ThiJ/PfpI

MMP0572 9 1.33 0.12 Peptidylprolyl isomerase, FKBP-type

MMP0589 3 1.29 0.19 Conserved hypothetical protein

MMP0593 2 1.87 0.14 Walker type ATPase

194

MMP0615 2 0.54 0.69 Conserved hypothetical protein

MMP0618 1 0.63 Conserved hypothetical protein

MMP0620 5 0.89 0.38 Methyl coenzyme M reductase, component A2

MMP0624 1 1.50 Conserved hypothetical protein

MMP0629 1 1.04 Conserved hypothetical archaeal protein

MMP0633 1 1.51 Rubrerythrin

MMP0641 3 1.28 0.03 50S ribosomal protein L7Ae

MMP0650 1 1.20 Acetolactate synthase

MMP0654 2 1.17 0.14 Ketol-acid reductoisomerase

MMP0658 4 1.27 0.13 MoaA/nifB/pqqE family

MMP0662 3 1.16 0.12 Hypothetical protein

MMP0663 1 0.81 Sodium-independent anion transporter

MMP0667 2 1.01 0.14 SSU Ribosomal protein S2

MMP0669 3 1.31 0.08 SSU ribosomal protein S3AE

MMP0680 1 1.27 Uracil phosphoribosyltransferase

MMP0686 1 0.66 Conserved hypothetical

MMP0687 2 1.19 0.11 Triosephosphate isomerase

MMP0695 1 0.86 Proteasome, subunit beta

MMP0696 1 1.44 Prolyl-tRNA synthetase

MMP0704 3 0.74 0.22 ParA type ATPase

MMP0720 1 2.35 Prismane

Exonuclease VII, large subunit:OB-fold nucleicacid binding MMP0732 1 1.69 domain

MMP0736 1 1.38 Conserved hypothetical archaeal protein

MMP0768 1 0.75 Conserved hypothetical protein

MMP0780 3 1.18 0.14 Conserved hypothetical protein

MMP0794 1 0.45 Hypothetical protein

MMP0806 2 0.81 0.18 Conserved hypothetical protein

195

MMP0831 1 0.30 Uroporphyrinogen decarboxylase (URO-D)

MMP0853 2 0.33 0.15 Nitrogenase iron protein (nitrogenase component II)

MMP0859 2 1.23 0.21 Iron-molybdenum cofactor biosynthesis protein, subunit N

MMP0879 2 1.19 0.07 Seryl-tRNA synthetase

MMP0880 3 1.06 0.19 2-oxosuberate synthase, last step

MMP0885 2 0.52 0.12 TPR repeat:ATP/GTP-binding site motif A(P-loop)

MMP0890 1 0.95 Superfamily II helicase

MMP0893 1 0.75 CTP synthase

MMP0894 4 1.05 0.18 GMP synthase (glutamine-hydrolyzing)

MMP0908 2 0.84 0.12 CBS domain

MMP0916 6 1.68 0.10 Conserved hypothetical protein

MMP0917 1 1.88 Diaminopimelate epimerase

MMP0920 1 1.51 S-adenosyl-L-homocysteine hydrolase

MMP0925 2 0.95 0.27 Chemotaxis protein CheW

MMP0928 1 0.82 Chemotaxis protein CheD

MMP0933 2 1.00 0.17 Chemotaxis protein CheY

MMP0945 1 1.50 Glyceraldehyde-3-phosphate ferredoxin oxidoreductase

MMP0961 7 1.47 0.08 Conserved hypothetical protein

MMP0962 1 0.93 Zinc finger protein

MMP0965 1 3.12 Formylmethanofuran dehydrogenase subunit E related protein

MMP0971 1 1.28 Adenylosuccinate lyase

MMP0988 1 1.43 RNA methyltransferase related protein

MMP0989 1 0.95 DNA topoisomerase VI B

MMP1002 1 0.83 Tryptophan synthase, alpha chain

MMP1009 1 1.67 Dihydroorotase

MMP1011 1 0.87 Glutamyl-tRNA synthetase

Transcription factor CBF/NF-Y/archaealhistone:Histone- MMP1015 2 1.23 0.02 fold/TFIID-TAF/NF-Y domain

196

MMP1016 5 0.98 0.12 Conserved Hypothetical protein with 4 CBSdomains.

MMP1018 3 1.89 0.04 (R)-citramalate synthase

MMP1023 1 1.08 Transcriptional regulator, TetR Family Member

MMP1024 1 0.93 MCM family related protein

MMP1026 14 1.21 0.07 Arginyl-tRNA synthetase

MMP1028 1 1.14 Hypothetic protein

MMP1031 3 1.00 0.34 Adenylate kinase

MMP1032 1 0.72 Replication protein A

MMP1038 1 1.37 A1A0 ATPase, subunit H

MMP1044 3 1.52 0.09 A1A0 ATPase, subunit A

MMP1045 6 1.29 0.08 A1A0 ATPase, subunit B

MMP1047 1 1.67 Conserved hypothetical protein

MMP1053 4 0.80 0.20 Heterodisulfide reductase; subunit B2

MMP1054 3 0.62 0.22 Heterodisulfide reductase; subunit C2

MMP1057 2 1.37 0.07 Hypothetical protein

MMP1060 2 1.83 0.07 Cysteinyl-tRNA synthetase

Aminotransferase (subgroup I), similar to aromatic MMP1072 11 1.22 0.16 aminotransferase

Bacterial transferase hexapeptiderepeat:ADP-glucose MMP1076 1 1.02 pyrophosphorylase

MMP1085 1 1.64 Conserved hypothetical protein

MMP1087 1 0.71 Hypothetical protein

MMP1091 3 0.85 0.59 ADP-glucose pyrophosphorylase

MMP1116 1 1.28 Endoglucanase

MMP1129 2 1.55 0.11 Peptidyl-prolyl cis-trans isomerase, cyclophilintype

MMP1131 1 1.03 peptide chain release factor aRF, subunit 1

MMP1137 1 2.79 Lrp-Like transcriptional regulatory proteins, AsnC family

MMP1149 1 1.19 3-Isopropylmalate dehydratase

197

MMP1158 1 1.50 Conserved hypothetical protein

MMP1164 1 0.75 Heavy-metal transport ATPase related protein

MMP1168 1 1.14 ABC transporter ATP-binding protein

MMP1174 1 1.17 Peroxiredoxin

MMP1175 1 0.85 Putative methyltransferase

MMP1186 1 1.90 ATP-dependent protease LonB

MMP1188 4 0.93 0.22 Conserved hypothetical archaeal protein

MMP1190 1 1.67 Peptidylprolyl isomerase, FKBP-type

MMP1191 6 1.53 0.24 N5, N10-methenyltetrahydromethanopterincyclohydrolase

MMP1203 3 1.23 0.11 Cobalamin (vitamin B12) biosynthesis CbiD protein

MMP1206 4 1.38 0.04 Glutamine synthetase

MMP1212 1 3.30 Putative thiolase/acetyl-CoA acetyltransferase

MMP1217 3 1.37 0.14 Hypothetical protein

Uroporphyrin-III C/tetrapyrrole(Corrin/Porphyrin) MMP1227 2 1.24 0.06 methyltransferase

MMP1235 1 0.80 Molybdopterin biosynthesis MoaE

MMP1238 2 1.35 0.01 TonB-dependent receptor protein:Biotin synthase

Tungsten containing formylmethanofurandehydrogenase, MMP1245 1 1.48 subunit F

Tungsten containing formylmethanofurandehydrogenase, MMP1247 7 0.69 0.30 subunit D

Tungsten containing formylmethanofurandehydrogenase, MMP1248 1 0.76 subunit A

Tungsten containing formylmethanofurandehydrogenase, MMP1249 1 0.72 subunit C

MMP1251 3 1.33 0.25 Conserved Hypothetical Protein with 2 CBSdomains.

MMP1252 2 1.00 0.10 CBS domain

MMP1254 1 0.79 Phosphoribosylaminoimidazole synthetase

MMP1274 1 1.96 Acetyl-CoA synthetase, AMP-forming-related

198

MMP1286 1 1.56 Possible DnaG-type primase

MMP1289 1 0.96 Ribosomal protein L10E

MMP1297 2 0.32 0.18 Formate dehydrogenase beta subunit

MMP1298 7 0.36 0.31 Formate dehydrogenase alpha subunit

MMP1299 4 0.34 0.36 Carbonic anhydrase

MMP1306 3 0.90 0.17 Conserved hypothetical protein

MMP1315 1 1.25 2-oxoglutarate oxidoreductase gamma subunit

MMP1319 1 1.18 SSU ribosomal protein S13

MMP1320 3 1.19 0.09 SSU ribosomal protein S4P (S9E)

MMP1321 2 1.06 0.02 SSU ribosomal protein S11

MMP1327 2 1.08 0.01 DNA-directed RNA polymerase, subunit K

MMP1336 1 0.98 Selenocysteine-specific translation elongation factor

MMP1347 17 1.23 0.03 Basic helix-loop-helix dimerization domain bHLH

MMP1352 1 1.77 Ribose 1,5-bisphosphate isomerase

MMP1360 2 0.87 0.30 RpoH DNA-directed RNA polymerase subunit H

MMP1361 2 1.37 0.10 DNA-directed RNA polymerase subunit B

MMP1368 2 0.92 0.13 SSU ribosomal protein S7P

MMP1369 15 1.11 0.13 Translation elongation factor EF-2

MMP1370 35 1.23 0.17 Translation elongation factor EF-1, subunit alpha

MMP1371 10 0.89 0.19 SSU ribosomal protein S10

MMP1373 3 1.35 0.14 Conserved hypothetical archaeal protein

MMP1382 4 0.37 0.10 Coenzyme F420-reducing hydrogenase subunit alpha

MMP1385 2 0.27 1.24 Coenzyme F420-reducing hydrogenase subunit beta

MMP1391 1 1.27 Aspartate-semialdehyde dehydrogenase

MMP1392 1 1.11 RNA-splicing ligase RtcB

MMP1399 1 0.69 Aspartate/glutamate/uridylate kinase

MMP1401 1 1.23 Translation elongation factor aEF-1 beta

199

MMP1403 2 1.06 0.10 LSU ribosomal protein L22P

MMP1404 6 0.99 0.16 SSU ribosomal protein S3P

MMP1409 1 1.11 LSU ribosomal protein L14P

MMP1410 1 2.29 LSU ribosomal protein L24P

MMP1411 4 0.93 0.29 SSU ribosomal protein S4E

MMP1412 1 1.11 LSU ribosomal protein L5P

MMP1414 2 1.21 0.25 SSU ribosomal protein S8P

MMP1419 2 1.18 0.13 SSU ribosomal protein S5P

MMP1420 4 0.67 1.16 LSU ribosomal protein L30P

MMP1426 1 0.94 Bifunctional dCTP deaminase/dUTP diphosphatase

MMP1431 1 1.36 2-phosphoglycerate kinase

MMP1433 7 1.05 0.25 Ribosomal protein L11

MMP1434 1 0.85 LSU ribosomal protein L24A

MMP1436 2 1.51 0.03 Cell division protein FtsZ

MMP1442 2 1.32 0.12 ArsR family transcriptional regulator

MMP1473 1 0.85 Conserved hypothetical protein

MMP1496 1 0.33 Phenylalanyl-tRNA synthetase alpha subunit

MMP1500 2 1.35 0.23 Cell division protein FtsZ2

MMP1502 1 1.12 Conserved archaeal protein, pyruvateoxidoreductase-associated

PorA, 2-ketoisovalerate ferredoxin oxidoreductase, subunit MMP1505 3 1.87 0.03 alpha

MMP1509 1 0.91 Conserved hypothetical protein

MMP1515 3 1.49 0.10 Chaperonin GroEL (thermosome, HSP60 family)

MMP1537 1 0.52 Cytochrome c, heme-binding site

MMP1542 1 1.19 Haloacid dehalogenase

MMP1543 1 1.18 LSU Ribosomal protein L3P

MMP1544 2 1.19 0.11 LSU Ribosomal protein L4P

MMP1546 4 1.05 0.29 LSU Ribosomal protein L2P

200

MMP1550 1 1.41 NADP oxidoreductase, coenzyme F420-dependent

MMP1551 1 0.63 Signal recognition particle protein SRP54

MMP1555 4 1.68 0.15 Methyl-coenzyme M reductase I, beta subunit

MMP1558 5 1.35 0.15 Methyl-coenzyme M reductase I, gamma subunit

MMP1559 8 1.69 0.05 Methyl-coenzyme M reductase I, alpha subunit

MMP1567 4 1.97 0.08 N5-methyltetrahydromethanopterin:methyltransferase, subunit H

MMP1573 2 0.35 0.24 Dethiobiotin synthetase

Dihydropteroate synthase, DHPS:Dihydropteroatesynthase- MMP1580 3 0.92 0.33 related protein synthase-related protein

MMP1586 12 1.69 0.03 Conserved hypothetical archaeal protein

MMP1588 4 1.03 0.13 D-3-phosphoglycerate dehydrogenase

MMP1592 1 1.76 Tryptophanyl-tRNA synthetase

MMP1593 1 0.66 Conserved hypothetical protein

MMP1608 1 0.92 Conserved hypothetical protein

MMP1611 1 1.35 Conserved hypothetical protein

MMP1614 1 0.55 Histidyl-tRNA synthetase

MMP1616 2 1.48 0.06 Aspartyl-tRNA synthetase

MMP1619 2 1.32 0.07 Molybdopterin molybdenumtransferase MoeA

MMP1635 1 0.78 Thioredoxin

MMP1637 9 0.89 0.21 Conserved hypothetical archaeal protein

MMP1640 1 1.93 Archaeal S-adenosylmethionine synthetase (MAT)

ATP/GTP-binding site motif A (P-loop):small GTP-binding MMP1642 1 2.18 protein domain

MMP1643 4 1.24 0.34 Conserved hypothetical protein

MMP1648 1 1.42 Conserved hypothetical protein

MMP1670 2 0.69 0.07 Flagella accessory protein D

MMP1671 2 0.73 0.14 Flagella accessory protein E

MMP1674 2 1.19 0.05 Flagella accessory protein H

201

MMP1681 1 3.27 Phosphoadenosine phosphosulfate reductase

MMP1684 1 3.77 Conserved hypothetical protein

Tungsten-containing formylmethanofuran dehydrogenase 2 MMP1691 5 0.91 0.15 subunit B

MMP1692 2 0.80 0.12 Polyferredoxin, associated with F420-non-reducing hydrogenase

MMP1694 4 1.46 0.09 F420-non-reducing hydrogenase subunit alpha

MMP1695 4 1.32 0.08 F420-non-reducing hydrogenase subunit

MMP1696 1 0.60 F420-non-reducing hydrogenase subunit delta

MMP1697 7 0.70 0.19 Heterodisulfide reductase, subunit A

MMP1700 2 1.35 0.20 Na+/solute symporter

MMP1707 1 1.61 Translation initiation factor IF-2 subunit alpha a Numbers of peptide pairs measured for the ORF. b Mean of the ratio of protein levels during growth on elemental sulfur to growth on sulfide. c Standard error of the mean or the standard deviation/√N. d From reference (53).

202

Supplementary table S3-5. Inorganic sulfide content in McNAA medium containing elemental sulfur and or other sulfur chemical compounds.

a d d Medium Type H2S (aq) (μM) H2S(g) (μM per liter culture) Medium+0.1 mM CoM < 1.5 < 1.5 Medium +0.1 mM CoM+S0b < 1.5 < 1.5

Medium +2 mM CoM < 1.5 < 1.5 Medium +2 mM CoM+S0c 69 ± 1 20 ± 1

Medium +0.1 mM thiosulfate < 1.5 < 1.5 Medium +0.1 mM thiosulfate +S0b < 1.5 < 1.5 Medium +2 mM thiosulfate < 1.5 < 1.5 Medium +2 mM thiosulfate + S0b < 1.5 < 1.5

Medium +2 mM sulfate < 1.5 < 1.5 Medium +2 mM sulfate + S0b < 1.5 < 1.5 Medium +20 mM sulfate < 1.5 < 1.5 Medium +20 mM sulfate +S0b < 1.5 < 1.5

Medium +2 mM sulfite < 1.5 < 1.5 Medium +2 mM sulfite +S0b < 1.5 < 1.5 Medium +20 mM sulfite < 1.5 < 1.5 Medium +20 mM sulfite +S0b < 1.5 < 1.5 a McNAA medium was autoclaved McNA medium supplemented with 1 mM alanine, no reducing reagent was added. Sterile-filtered sulfur chemical compounds and or 0.1 g elemental sulfur (S0) per 5mL medium was subsequently added into the McNAA medium as indicated. The medium contained sulfur chemical compounds were incubated in the anaerobic chamber at room temperature for one day before measuring the inorganic sulfide content. b The media contained sulfur chemical compounds and elemental sulfur were incubated in the anaerobic chamber at room temperature for one day and seven days before measuring the inorganic sulfide content. The results presented here represent the results of two rounds of inorganic sulfide content measurements, including one measurement with medium tubes that had been incubated for one day, and the other measurement with medium tubes that had been incubated for seven days. 203

c The medium that contained 2 mM coM and 2 % (w/v) elemental sulfur was incubated in the anaerobic chamber at room temperature for one day before measuring the inorganic sulfide content. d Values were means ± 1 S.D., obtained from three independent samples and represent inorganic sulfide amount either in the aqueous phase (aq) or gaseous phase (g).

204

Supplementary table S3-6. The levels of inorganic sulfate in McFAA medium compoentsa.

Samplea Sulfate conc. (µM)g Double-distilled waterb N.D. 80 mM Potassium phosphate dibasic N.D. 80 mM Potassium phosphate dibasic (autoclaved)c N.D. 400 mM Sodium formate N.D. 400 mM Sodium formate (autoclaved) c N.D. Sulfate-free trace minerals stock solutiond N.D. Sulfate-free general salts stock solutiond N.D. 1 M glycylglycine N.D. 25 μM Iron (II) chloride N.D. 1 M sodium acetate N.D. 100 mM Alanine N.D. Sulfate-free mediumce N.D. ce Sulfate-free medium +2 mM Na2S N.D. 40 μM Sodium sulfatef 39.3 ± 0.8 80 μM Sodium sulfatef 78.8 ± 0.9 a The detailed information of McFAA medium can be found in reference (54). b The double-distilled water used to prepare medium and chemical stock solutions. c Autoclaved at gravity cycle (rapid exhaust) for 20 minutes. d Sulfate-containing chemicals in these solutions were replaced with the corresponding chemical with chloride. e Sulfate-depleted medium was made with above described sulfate-depleted medium ingredients. f Sodium sulfate solution used as a positive control for this sulfate measurement experiment. g Values are the means of three independent samples ± S.D.

N.D.: Not detectable 205

References:

1. Baba T, Ara T, Hasegawa M, Takai Y, Okumura Y, Baba M, Datsenko KA, Tomita M,

Wanner BL, Mori H. 2006. Construction of Escherichia coli K‐12 in‐frame, single‐gene

knockout mutants: the Keio collection. Molecular systems biology 2.

2. Lyu Z, Chou CW, Shi H, Patel R, Ghebreab R, Duin EC, Whitman WB. Methanogenesis

marker 10 protein required for methylation of the arginine at the active site of methyl-

coenzyme M reductase of Methanococcus maripaludis. (Manuscript).

3. Lyu Z, Jain R, Smith P, Fetchko T, Yan Y, Whitman WB. 2016. Engineering the autotroph

Methanococcus maripaludis for geraniol production. ACS Synth Biol 5:577-81.

4. Fauque GD, Barton LL. 2012. Hemoproteins in dissimilatory sulfate- and sulfur-reducing

prokaryotes. Adv Microb Physiol 60:1-90.

5. Hedderich R, Klimmek O, Kröger A, Dirmeier R, Keller M, Stetter KO. 1998. Anaerobic

respiration with elemental sulfur and with disulfides. FEMS Microbiol Rev 22:353-381.

6. Segerer A, Neuner A, Kristjansson JK, Stetter KO. 1986. Acidianus infernus gen. nov., sp.

nov., and Acidianus brierleyi Comb. nov.: Facultatively aerobic, extremely acidophilic

thermophilic sulfur-metabolizing archaebacteria. Int J Syst Evol Microbiol 36:559-564.

7. Prokofeva MI, Miroshnichenko ML, Kostrikina NA, Chernyh NA, Kuznetsov BB,

Tourova TP, Bonch-Osmolovskaya EA. 2000. Acidilobus aceticus gen. nov., sp nov., a

novel anaerobic thermoacidophilic archaeon from continental hot vents in Kamchatka. Int

J Syst Evol Microbiol 50:2001-2008.

8. Segerer AH, Trincone A, Gahrtz M, Stetter KO. 1991. Stygiolobus Azoricus gen. nov. sp.

nov. represents a novel genus of anaerobic, extremely thermoacidophilic archaebacteria of

the order Sulfolobales. Int J Syst Evol Microbiol 41:495-501. 206

9. Huber R, Kristjansson JK, Stetter KO. 1987. Pyrobaculum Gen. nov., a new genus of

neutrophilic, rod-shaped archaebacteria from continental solfataras growing optimally at

100 oC. Arch Microbiol 149:95-101.

10. Zillig W, Gierl A, Schreiber G, Wunderl S, Janekovic D, Stetter KO, Klenk HP. 1983. The

archaebacterium Thermofilum pendens represents a novel genus of the thermophilic

anaerobic sulfur respiring Thermoproteales. Syst Appl Microbiol 4:79-87.

11. Stetter KO, Segerer A, Zillig W, Huber G, Fiala G, Huber R, Konig H. 1986. Extremely

thermophilic sulfur-metabolizing archaebacteria. Syst Appl Microbiol 7:393-397.

12. Fischer F, Zillig W, Stetter KO, Schreiber G. 1983. Chemolithoautotrophic metabolism of

anaerobic extremely thermophilic archaebacteria. Nature 301:511-513.

13. Selig M, Schonheit P. 1994. Oxidation of organic compounds to CO2 with Sulfur or

thiosulfate as electron acceptor in the anaerobic hyperthermophilic archaea Thermoproteus

tenax and Pyrobaculum islandicum proceeds via the citric-acid cycle. Arch Microbiol

162:286-294.

14. Bonchosmolovskaya EA, Slesarev AI, Miroshnichenko ML, Svetlichnaya TP, Alekseev

VA. 1988. Characteristics of Desulfurococcus amylolyticus nov. sp.: A new extremely

thermophilic arachaebacterium isolated from thermal springs of Kamchatka and Kunashir

Island. Microbiology 57:78-85.

15. Zillig W, Stetter KO, Prangishvilli D, Schafer W, Wunderl S, Janekovic D, Holz I, Palm

P. 1982. Desulfurococcaceae, the second family of the extremely thermophilic, anaerobic,

sulfur-respiring Thermoproteales. Zbl Bakt Hyg, I Abt Orig C 3:304-317.

16. She Q, Singh RK, Confalonieri F, Zivanovic Y, Allard G, Awayez MJ, Chan-Weiher,

Clausen IG, Curtis BA, De Moors A, Erauso G, Fletcher C, Gordon PMK, Heikamp-de 207

Jong I, Jeffries AC, Kozera CJ, Medina N, Peng X, Thi-Ngoc HP, Redder P, Schenk ME,

Theriault C, Tolstrup N, Charlebois RL, Doolittle WF, Duguet M, Gaasterland T, Garrett

RA, Ragan MA, Sensen CW, Van der Oost J. 2001. The complete genome of the

crenarchaeon Sulfolobus solfataricus P2. Proc Natl Acad Sci U S A 98:7835-7840.

17. Stetter KO, Konig H, Stackebrandt E. 1983. Pyrodictium gen. nov., a new genus of

submarine disc-shaped sulfur reducing archaebacteria growing optimally at 105 oC. Syst

Appl Microbiol 4:535-551.

18. Dirmeier R, Keller M, Frey G, Huber H, Stetter KO. 1998. Purification and properties of

an extremely thermostable membrane-bound sulfur-reducing complex from the

hyperthermophilic Pyrodictium abyssi. Eur J Biochem 252:486-491.

19. Jochimsen B, PeinemannSimon S, Volker H, Stuben D, Botz R, Stoffers P, Dando PR,

Thomm M. 1997. Stetteria hydrogenophila, gen. nov. and sp. nov., a novel mixotrophic

sulfur-dependent crenarchaeote isolated from Milos, Greece. Extremophiles 1:67-73.

20. Huber R, Dyba D, Huber H, Burggraf S, Rachel R. 1998. Sulfur-inhibited Thermosphaera

aggregans sp. nov., a new genus of hyperthermophilic archaea isolated after its prediction

from environmentally derived 16S rRNA sequences. Int J Syst Evol Microbiol 48:31-38.

21. Itoh T, Suzuki K, Nakase T. 1998. Thermocladium modestius gen. nov., sp. nov., a new

genus of rod-shaped, extremely thermophilic crenarchaeote. Int J Syst Evol Microbiol

48:879-887.

22. Fiala G, Stetter KO, Jannasch HW, Langworthy TA, Madon J. 1986. Staphylothermus

Marinus sp. nov. Represents a novel genus of extremely thermophilic submarine

heterotrophic archaebacteria growing up to 98 oC. Syst Appl Microbiol 8:106-113.

208

23. Zillig W, Holz I, Janekovic D, Klenk HP, Imsel E, Trent J, Wunderl S, Forjaz VH,

Coutinho R, Ferreira T. 1990. Hyperthermus butylicus, a hyperthermophilic sulfur-

reducing archaebacterium that ferments peptides. J Bacteriol 172:3959-3965.

24. Itoh T, Suzuki K, Nakase T. 2002. Vulcanisaeta distributa gen. nov., sp. nov., and

Vulcanisaeta souniana sp. nov., novel hyperthermophilic rod-shaped crenarchaeotes

isolated from hot springs in Japan. Int J Syst Evol Microbiol 52:1097-1104.

25. Fiala G, Stetter KO. 1986. Pyrococcus furiosus sp nov. represents a novel genus of marine

heterotrophic archaebacteria growing optimally at 100 oC. Archs Microbiol 145:56-61.

26. Gonzalez JM, Masuchi Y, Robb FT, Ammerman JW, Maeder DL, Yanagibayashi M,

Tamaoka J, Kato C. 1998. Pyrococcus horikoshii sp. nov., a hyperthermophilic archaeon

isolated from a hydrothermal vent at the Okinawa Trough. Extremophiles 2:123-130.

27. Dirmeier R, Keller M, Hafenbradl D, Braun FJ, Rachel R, Burggraf S, Stetter KO. 1998.

Thermococcus acidaminovorans sp. nov., a new hyperthermophilic alkalophilic archaeon

growing on amino acids. Extremophiles 2:109-114.

28. Neuner A, Jannasch HW, Belkin S, Stetter KO. 1990. Thermococcus litoralis sp. nov., a

new species of extremely thermophilic marine archaebacteria. Arch Microbiol 153:205-

207.

29. Svetlichnyi VA, Slesarev AI, Svetlichnaya TP, Zavarzin GA. 1987. Caldococcus litoralis

gen. nov. sp. nov., a new marine, extremely thermophilic, sulfur-reducing archaebacterium.

Microbiology 56:658-664.

30. Itoh T, Suzuki K, Sanchez PC, Nakase T. 2003. Caldisphaera lagunensis gen. nov., sp.

nov., a novel thermoacidophilic crenarchaeote isolated from a hot spring at Mt Maquiling,

Philippines. Int J Syst Evol Microbiol 53:1149-1154. 209

31. Segerer A, Langworthy TA, Stetter KO. 1988. Thermoplasma acidophilum and

Thermoplasma volcanium sp. nov. from Solfatara Fields. Syst Appl Microbiol 10:161-171.

32. Stetter KO, Gaag G. 1983. Reduction of molecular sulfur by methanogenic bacteria. Nature

305:309-311.

33. Huber R, Wilharm T, Huber D, Trincone A, Burggraf S, Konig H, Rachel R, Rockinger I,

Fricke H, Stetter KO. 1992. Aquifex pyrophilus gen. nov., sp. nov. represents a novel group

of marine hyperthermophilic hydrogen oxidizing bacteria. Syst Appl Microbiol 15:340-

351.

34. Huber R, Rossnagel P, Woese CR, Rachel R, Langworthy TA, Stetter KO. 1996. Formation

of ammonium from nitrate during chemolithoautotrophic growth of the extremely

thermophilic bacterium Ammonifex degensii gen. nov., sp. nov. Syst Appl Microbiol 19:40-

49.

35. L'Haridon S, Cilia V, Messner P, Raguenes G, Gambacorta A, Sleytr UB, Prieur D,

Jeanthon C. 1998. Desulfurobacterium thermolithotrophum gen. nov., sp. nov., a novel

autotrophic, sulphur-reducing bacterium isolated from a deep-sea hydrothermal vent. Int J

Syst Bacteriol 48 Pt 3:701-11.

36. Pfennig N, Biebl H. 1976. Desulfuromonas acetoxidans gen. nov. and sp. nov., a new

anaerobic, sulfur-reducing, acetate-oxidizing bacterium. Arch Microbiol 110:3-12.

37. Liesack W, Finster K. 1994. Phylogenetic analysis of five strains of gram-negative,

obligately anaerobic, sulfur-reducing bacteria and description of Desulfuromusa sp. nov.,

including Desulfuromusa kysingii sp. nov., Desulfuromusa bakii sp. nov., and

Desulfuromusa succinoxidans sp. nov., Int J Syst Evol Microbiol 44:753-758.

210

38. Bonchosmolovskaya EA, Sokolova TG, Kostrikina NA, Zavarzin GA. 1990. Desulfurella

acetivorans gen. nov. and sp. nov., a new thermophilic sulfur-reducing eubacterium. Arch

Microbiol 153:151-155.

39. Schmitz RA, Bonchosmolovskaya EA, Thauer RK. 1990. Different mechanisms of acetate

activation in Desulfurella acetivorans and Desulfuromonas acetoxidans. Arch Microbiol

154:274-279.

40. Biebl H, Pfennig N. 1977. Growth of sulfate-reducing bacteria with sulfur as electron-

acceptor. Arch Microbiol 112:115-117.

41. Huber R, Woese CR, Langworthy TA, Kristjansson JK, Stetter KO. 1990.

Fervidobacterium islandicum sp. nov., a new extremely thermophilic eubacterium

belonging to the Thermotogales. Arch Microbiol 154:105-111.

42. Patel BKC, Morgan HW, Daniel RM. 1985. Fervidobacterium nodosum gen. nov.and sp.

nov., a new chemoorganotrophic, caldoactive, anaerobic bacterium. Arch Microbiol

141:63-69.

43. Caccavo F, Lonergan DJ, Lovley DR, Davis M, Stolz JF, Mcinerney MJ. 1994. Geobacter

Sulfurreducens sp. nov., a hydrogen-oxidizing and acetate-oxidizing dissimilatory metal-

reducing microorganism. Appl Environ Microbiol 60:3752-3759.

44. Lovley DR, Phillips EJP, Lonergan DJ, Widman PK. 1995. Fe(III) and S0 reduction by

Pelobacter carbinolicus. Appl Environ Microbiol 61:2132-2138.

45. Moser DP, Nealson KH. 1996. Growth of the facultative anaerobe Shewanella putrefaciens

by elemental sulfur reduction. Appl Environ Microbiol 62:2100-2105.

46. Wolfe RS, Penning N. 1977. Reduction of sulfur by spirillum 5175 and syntrophism with

chlorobium. Appl Environ Microbiol 33:427-33. 211

47. Finster K, Liesack W, Tindall BJ. 1997. Sulfurospirillum arcachonense sp. nov., a new

microaerophilic sulfur-reducing bacterium. Int J Syst Bacteriol 47:1212-7.

48. Windberger E, Huber R, Trincone A, Fricke H, Stetter KO. 1989. Thermotoga thermarum

sp. nov. and Thermotoga neapolitana occurring in african continental solfataric springs.

Arch Microbiol 151:506-512.

49. Huber R, Langworthy TA, Konig H, Thomm M, Woese CR, Sleytr UB, Stetter KO. 1986.

Thermotoga maritima sp. nov. represents a new genus of unique extremely thermophilic

eubacteria growing up to 90 oC. Arch Microbiol 144:324-333.

50. Huber R, Woese CR, Langworthy TA, Fricke H, Stetter KO. 1989. Thermosipho africanus

gen. nov., represents a new genus of thermophilic eubacteria within the Thermotogales.

Syst Appl Microbiol 12:32-37.

51. Antoine E, Cilia V, Meunier JR, Guezennec J, Lesongeur F, Barbier G. 1997. Thermosipho

melanesiensis sp. nov., a new thermophilic anaerobic bacterium belonging to the order

Thermotogales, isolated from deep-sea hydrothermal vents in the southwestern Pacific

Ocean. Int J Syst Bacteriol 47:1118-23.

52. Macy JM, Schroder I, Thauer RK, Kroger A. 1986. Growth the Wolinella succinogenes on

H2S plus fumarate and on formate plus sulfur as energy sources. Arch of Microbiol

144:147-150.

53. Hendrickson EL, Kaul R, Zhou Y, Bovee D, Chapman P, Chung J, Conway de Macario E,

Dodsworth JA, Gillett W, Graham DE, Hackett M, Haydock AK, Kang A, Land ML, Levy

R, Lie TJ, Major TA, Moore BC, Porat I, Palmeiri A, Rouse G, Saenphimmachak C, Soll

D, Van Dien S, Wang T, Whitman WB, Xia Q, Zhang Y, Larimer FW, Olson MV, Leigh

212

JA. 2004. Complete genome sequence of the genetically tractable hydrogenotrophic

methanogen Methanococcus maripaludis. J Bacteriol 186:6956-69.

54. Long F, Wang L, Lupa B, Whitman WB. 2017. A flexible system for cultivation of

Methanococcus and other formate-utilizing methanogens. Archaea. 2017:12.

doi:10.1155/2017/7046026.

213

APPENDIX C

CHAPTER 4 SUPPLEMENTARY INFORMATION3

3 Long, F., J. Cheung, W.B. Whitman, R.S. Ronimus and G.M. Cook. To be submitted to Applied and Environmental Microbiology.

214

1.2

1.0

0.8

0.6

0.4

Absorbance Absorbance (600nm) 0.2

0.0 0 50 100 150 200 Time (h) Echinomycin 0 µM 0.1 µM 0.25 µM

Supplementary figure S4-1. Growth of Methanococcus maripaludis S2 in McFAA medium with echinomycin ranging from 0, 0.1, 0.25 µM. McFAA medium was reduced with 2 mM dithiothreitol, and 2 mM sulfide was added as sulfur source. The inoculum size was 1.5×105 cells per 5 mL culture. All values were the averages of three independent cultures.

215

Supplementary table S4-1. Whole-genome sequencing primers used in echinomycin resistance mutants and their parental strains.

Strain i7 primer i7 sequence i5 primer i5 sequence M. m S2 D701 ATTACTCG D505 AGGCGAA M. m S0001 D702 TCCGGAGA D505 AGGCGAA G EchB1 D706 GAATTCGT D505 AGGCGAA G EchC2 D707 CTGAAGCT D505 AGGCGAA G EchC4 D708 TAATGCGC D505 AGGCGAA G EchD3 D709 CGGCTATG D505 AGGCGAA G Ech25 D710 TCCGCGAA D505 AGGCGAA G Ech26 D711 TCTCGCGC D505 AGGCGAA G

216

Supplementary table S4-2. Number of SNPs identified in echinomycin-, neomycin-, chloroform- resistant mutant strains. The frequency of these SNPs was at least 10 %.

Resistant mutant straina Variant SNPs # from parental strain/Total SNPs # E25 12/85 E26 19/90 Ech B1 71/276 Ech C2 60/261 Ech C4 72/283 Ech D3 65/281 Neo 1 213/508 Neo 2 154/456 Neo 3 205/526 Neo 4 204/509 Chl 1 264/595 Chl 2 169/473 Chl 3 171/500 Chl 4 272/581 a The SNPs of three types of resistant mutant strains were summarized: (i) echinomycin- resistant mutants, E25 and E26, derived from Methanococcus maripaludis S2 (113 SNPs in total); Ech B1,

Ech C2, Ech C4 and Ech D3, derived from Methanococcus maripaludis S0001 (290 SNPs in total).

(ii) Neomycin- resistant mutants, Neo 1, Neo 2, Neo 3 and Neo 4. (iii) Chloroform- resistant mutants, Chl 1, Chl 2, Chl 3 and Chl 4. Both neomycin- resistant mutants and chloroform- resistant mutants were derived from Methanococcus maripaludis S2 (428 SNPs in total).

217