i

The Pennsylvania State University

The Graduate School

Eberly College of Science

THE ELECTRON TRANSPORT OF -GROWN

METHANOSARCINA ACETIVORANS

A Dissertation in

Biochemistry, Microbiology, and Molecular Biology

by

Mingyu Wang

 2010 Mingyu Wang

Submitted in Partial Fulfillment of the Requirements for the Degree of

Doctor of Philosophy

December 2010

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The dissertation of Mingyu Wang was reviewed and approved* by the following:

James G. Ferry Stanley Person Professor and Director, Center for Microbial Structural Biology Dissertation Advisor Chair of Committee

Sarah E. Ades Associate Professor of and Molecular Biology

Donald A. Bryant Ernest C. Pollard Professor of Biotechnology and Professor of Biochemistry and Molecular Biology

Christopher H. House Associate Professor of Geosciences

Ming Tien Professor of Biochemistry

Scott B. Selleck Professor and Head, Department of Biochemistry and Molecular Biology

*Signatures are on file in the Graduate School

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ABSTRACT The electron transport of marine acetate-utilizing acetivorans was investigated, leading to the first identification and partial characterization of two novel ferredoxin: CoM-S-S-CoB electron transport chains. The study of an Rnf-dependent membrane-bound pathway of aceticlastic M. acetivorans that doesn’t reduce CO2 with H2 characterized members both unique to Rnf-dependent pathway and also in common with Ech-dependent ferredoxin: CoM-S-S-CoB electron transport chain in CO2 utilizing aceticlastic . These include the Rnf complex, cytochrome c, ferredoxin, Cdh, methanophenazine, heterodisulfide reductase and

CoM-S-S-CoB. The purification and phylogenetic analysis of ferredoxin suggested an aceticlastic-specific ferredoxin clade among methanogens. This Rnf-dependent electron transport pathway, shared with non-CO2 utilizing Methanosarcina thermophila is analogous to Ech-dependent electron transport pathway in its location and terminal electron partners but differs from the later in that it lacks the use of hydrogenases and H2.

A novel soluble ferredoxin: CoM-S-S-CoB pathway was identified in M. acetivorans by comparing soluble and membrane-bound ferredoxin: CoM-S-S-CoB oxidoreductase activities and was hypothesized to involve a HdrA: MvhD fusion protein, named Etp.

Etp was heterologously overexpressed in Escherichia coli, purified, reconstituted and partially characterized for the first time, showing heavy iron-sulfur cluster content. The reduction of Etp is linked to the oxidation of ferredoxin mediated by unknown soluble factors. Finally, a model of energy conservation for M. acetivorans was constructed suggesting this soluble electron transport chain serves to bypass certain energy coupling sites and maximize energy conservation efficiency under substrate-limited scenarios. iv

TABLE OF CONTENTS LIST OF FIGURES ...... viii

LIST OF EQUATIONS ...... xii

LIST OF TABLES ...... xiv

ACKNOWLEDGEMENTS ...... xv

Chapter 1 Methanogens and ...... 1

1.1 Methanogenesis and methanogens ...... 1

1.1.1 Methanogenesis ...... 1

1.1.2 of methanogens ...... 2

1.1.3 and biology of methanogens ...... 4

1.2 Methanogenesis from three major biochemical pathways ...... 6

1.2.1 Cofactors utilized in methanogenic pathways ...... 6

1.2.2 H2 + CO2 pathway ...... 9

1.2.3 Methylotrophic pathway ...... 25

1.2.4 Acetate pathway ...... 31

1.3 References ...... 36

Chapter 2 Electron transport and energy conservation in methanogens ...... 69

2.1 Introduction ...... 69

2.2 Methanogenesis coupled ATP synthesis ...... 69

2.3 Membrane-bound electron transport chains of methanogens ...... 72 v

2.3.1 F420: CoM-S-S-CoB pathway ...... 73

2.3.2 H2: CoM-S-S-CoB pathway ...... 74

2.3.3 The acetate-specific ferredoxin: CoM-S-S-CoB electron transport

chain ...... 76

2.4 Electron transport in obligate CO2-reducing methanogens ...... 78

2.5 involved in electron transport chains of methanogens ...... 80

2.5.1 F420 dehydrogenase ...... 80

2.5.2 Membrane bound F420 non-reducing hydrogenase ...... 83

2.5.3 Ech hydrogenase ...... 85

2.5.4 Rnf complex ...... 87

2.6 References ...... 90

Chapter 3 The membrane-bound electron transport chain of acetate-grown

Methanosarcina acetivorans ...... 103

3.1 Abstract ...... 103

3.2 Introduction ...... 104

3.3 Methods and materials ...... 107

3.4 Results...... 111

3.4.1 Purification of the CdhAE component of the CO

dehydrogenase/acetyl-CoA complex (Cdh) of M. acetivorans ...... 111

3.4.2 Properties of the ferredoxin purified from acetate-grown M.

acetivorans ...... 111

3.4.3 Role of ferredoxin in the membrane-bound electron transport chain ... 116 vi

3.4.4 Role of Rnf in the membrane-bound electron transport chain ...... 116

3.4.5 Role of cytochrome c in the membrane-bound electron transport

chain ...... 118

3.4.6 Role of methanophenazine in the membrane-bound electron

transport chain...... 119

3.4.7 Comparative analysis of the M. thermophila ...... 123

3.5 Discussion ...... 125

3.6 Acknowledgements ...... 129

3.7 References ...... 130

Chapter 4 Characterization of a novel electron transport protein from

Methanosarcina acetivorans ...... 139

4.1 Abstract ...... 139

4.2 Introduction ...... 140

4.3 Materials and methods ...... 143

4.4 Results...... 149

4.4.1 Purification and Properties ...... 149

4.4.2 Physiology ...... 152

4.4.3 Phylogeny ...... 160

4.5 Discussion ...... 160

4.6 Acknowledgements ...... 165

4.7 References ...... 165 vii

Chapter 5 Discussion and future directions on the understanding of electron

transport in acetate-grown Methanosarcina acetivorans ...... 171

5.1 A model of energy conservation under fluctuating environmental

substrate concentrations ...... 172

5.2 Future directions on the research of electron transport during acetate

metabolism ...... 176

5.2.1 Membrane-bound electron transport chain ...... 176

5.2.2 Soluble electron transport chain ...... 178

5.3 References ...... 179

Appendix A Alignment of the deduced protein sequences of rnf genes

between Methanosarcina thermophila and Methanosarcina. acetivoran ..... 183

Appendix B Alignment of the deduced protein sequences of vht and vhx

genes between Methanosarcina thermophila and Methanosarcina

acetivorans ...... 187

Appendix C Alignment of the deduced protein sequences of mrp genes

between Methanosarcina thermophila and Methanosarcina acetivorans .... 193

VITA ...... 197

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LIST OF FIGURES

Figure 1-1 The global carbon cycle...... 4

Figure 1-2 Cofactors of methanogens...... 7

Figure 1-3 The H2 + CO2 methanogenesis pathway...... 11

Figure 1-4 Hypothetical Hmd-Mtd cycle...... 19

Figure 1-5 Methanogenesis from methylated compounds...... 27

Figure 1-6 Methyl group transfer from to ...... 28

Figure 1-7 DE-type heterodisulfide reductase...... 30

Figure 1-8 Methanogenesis from acetate...... 31

Figure 2-1 Electron microscopy and structure prediction of ATPase from M.

jannaschii...... 71

Figure 2-2 Tentative model of the F420: CoM-S-S-CoB electron transport pathway.

...... 74

Figure 2-3 Tentative model of the H2: CoM-S-S-CoB electron transport pathway. ... 75 ix

Figure 2-4 Tentative model of the ferredoxin: CoM-S-S-CoB electron transport

pathway in M. barkeri and M. mazei...... 77

Figure 2-5 Model of electron bifurcation at Mvh/Hdr complex for obligate CO2

reducers...... 80

Figure 2-6 Tentative model of F420 dehydrogenase from M. mazei...... 82

Figure 2-7 Tentative model of Ech hydrogenase in M. barkeri...... 86

Figure 2-8 Tentative model of Rnf complex in M. acetivorans...... 88

Figure 3-1 Mass spectrometry of ferredoxin from M. acetivorans...... 113

Figure 3-2 UV-visible absorption spectra of purified ferredoxin...... 114

Figure 3-3 Phylogenetic analysis of ferredoxins...... 114

Figure 3-4 Sequence alignment of ferredoxins from Methanosarcina ...... 115

Figure 3-5 Reduction of ferredoxin by CdhAE...... 115

Figure 3-6 Ferredoxin: CoM-S-S-CoB oxidoreductase activity of membranes...... 117

Figure 3-7 Reduction of RnfB by ferredoxin...... 118

Figure 3-8 Ferredoxin-dependent reduction of membrane-bound cytochrome c...... 120

Figure 3-9 Oxidation of membrane-bound cytochrome c by CoM-S-S-CoB...... 121 x

Figure 3-10 Reduction of 2-hydroxyphenazine and membrane-catalyzed oxidation

dependent on CoM-S-S-CoB...... 121

Figure 3-11 Oxidation of membrane-bound cytochrome c by 2-hydroxyphenazine. .. 122

Figure 3-12 Alignment of rnf gene clusters between M. thermophila and M.

acetivorans...... 124

Figure 3-13 Alignment of vht and vhx gene clusters between M. thermophila and

M. acetivorans ...... 124

Figure 3-14 Alignment of mrp gene clusters between M. thermophila and M.

acetivorans...... 125

Figure 3-15 Comparison of electron transport pathways for M. mazei and M.

barkeri versus M. acetivorans...... 126

Figure 4-1 SDS-PAGE of purified Etp...... 149

Figure 4-2 Alignment of the deduced sequences of etp, hdrA and mvhD...... 151

Figure 4-3 UV-visible absorption spectrum of unreconstituted and reconstituted

Etp...... 152

Figure 4-4 The reduction of Etp requires unknown components in soluble fraction

of acetate-grown M. acetivorans...... 154

Figure 4-5 Ferredoxin-dependent Etp reduction...... 155 xi

Figure 4-6 Membrane fraction of acetate-grown M. acetivorans doesn’t stimulate

the reduction of Etp...... 155

Figure 4-7 The stimulation of Etp of ferredoxin: CoM-S-S-CoB oxidoreductase

activity in soluble fraction of acetate-grown M. acetivorans ...... 156

Figure 4-8 Ferredoxin: CoM-S-S-CoB oxidoreductase activity in the soluble

fraction of acetate-grown M. acetivorans...... 157

Figure 4-9 Ferredoxin: CoM-S-S-CoB oxidoreductase activity in fractions of

acetate-grown M. acetivorans with CO/Cdh as ferredoxin regenerating

system...... 158

Figure 4-10 Ferredoxin: CoM-S-S-CoB oxidoreductase activity in fractions of

acetate-grown M. acetivorans with NADPH/FNR as ferredoxin regenerating

system...... 159

Figure 4-11 Proposed electron transport model in acetate-grown M. acetivorans...... 164

Figure 5-1 Model of electron transport and energy conservation in acetate-grown

M. acetivorans...... 174

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LIST OF EQUATIONS

Equation 1-1 Methanogenesis from H2 + CO2 ...... 9

Equation 1-2 CO2 fixation with ...... 10

Equation 1-3 Formyl group transfer to THMPT...... 10

Equation 1-4 Cyclization of N5-formyl-THMPT...... 10

5 10 Equation 1-5 F420 dependent reduction of N , N -methenyl-THMPT...... 12

5 10 Equation 1-6 H2 dependent reduction of N , N -methenyl-THMPT...... 12

Equation 1-7 Reduction of N5, N10-methylene-THMPT...... 12

Equation 1-8 Spontaneous reversible carboxylation of methanofuran...... 14

Equation 1-9 Methylation of coenzyme M...... 20

Equation 1-10 Release of from methyl coenzyme M...... 22

Equation 1-11 Regeneration of coenzyme M and ...... 24

Equation 1-12 Methanogenesis from methanol...... 26

Equation 1-13 Activation of acetate...... 32

Equation 1-14 Acetyl group transfer to ...... 32 xiii

Equation 1-15 Oxidation of carbonyl group and synthesis of methyl-THSPT...... 32

Equation 1-16 Methyl transfer to coenzyme M...... 32

Equation 1-17 Reduction of methyl group to methane...... 32

Equation 1-18 Regeneration of coenzyme M and coenzyme B...... 32

Equation 1-19 Methanogenesis from acetate...... 33

Equation 5-1 Methanogenesis from acetate...... 172

Equation 5-2 Na+ and H+ translocation by Mrp complex...... 175

xiv

LIST OF TABLES

Table 4-1 Activities catalyzed by cell-free extract, soluble fraction and membrane

fraction of acetate-grown M. acetivorans...... 156

Table 4-2 Comparison of protein concentrations determined by BCA assay and

Bradford assay...... 157 xv

ACKNOWLEDGEMENTS

In this unforgettable seven years at Penn State University, I’ve come across ups and downs in my life and in my research. The support from my adviser, my Committee members, our collaborators, my labmates and my family is the reason how I survived the most difficult time. Here, I would like to sincerely thank my adviser Dr. James Gregory

Ferry for his years’ long guidance in research and also being a good friend of mine. His wisdom in science and life opened up the charming door of scientific research to me, and taught me how to appreciate this beauty. Special thanks go out to my Committee members, Dr. Sarah E. Ades, Dr. Donald A. Bryant, Dr. Christopher H. House and Dr.

Ming Tien. I appreciate their critical opinions and creative ideas along the road of my research, as well as their enormous contribution to the BMMB department. I would like to thank our collaborators Dr. Jean-Francois Tomb for providing critical sequence information of Methanosarcina thermophila and Dr. Jan Keltjens for generously providing CoM-S-S-CoB. I would like to thank all the current and former members of

Ferry lab for their help and support in the past seven years. My parents are my moral support as well as the source of my strength. They are my last line of defense and I know no matter what happens to me, there are two people I can always fall back on unconditionally: my father Chuanfeng Wang and my mother Zhiying Liu. I would like them to know that they are forever my heroes and my gods, that we are eternally connected with a bond nothing can break, and that I will adore them as long as my breath holds. My wife, Bo Zhang, is the other half of me. She took care of me when all my other loves ones were far away. She filled me with the sense of responsibility as a man, xvi and encouraged me to rise from defeat. I would like to thank her for her company and love, in the past, and in the seventy years to come. Lastly, but not least, I would like to thank my son, Ruoyuan ‘Tuantuan’ Wang. Although he’s not old enough to comprehend science yet, I hope one day when he reads his old man’s dissertation, he would understand his mere presence filled his dad with motivation and lit his dad’s heart with eternal light and warmth. 1

Chapter 1

Methanogens and methanogenesis

1.1 Methanogenesis and methanogens

1.1.1 Methanogenesis

Methanogenesis is the process of microorgasms converting simple organic matters such as acetate, formate, CO2, methanol and into methane. The first observation of methanogenesis can be dated back to 1770s. In a letter Alessandro

Volta wrote to his friend the famous ‘Volta experiment’ was described (137). In this experiment, combustible gas can be observed and collected when sediment of lakes was stirred. This combustible gas was later known to be methane (203). It was not until late 1800s when the formation of methane was observed to be related to microorganisms

(9). Although there had been plenty of work done on microbial methanogenesis afterwards, the first pure methanogen culture was not obtained until 1947 when

Methanobacterium formicicum and were isolated and purified

(153). By late 1960s, people started realizing methanogens are metabolic specialists, which only use simple organic matters generally with fewer than 2 carbons while unable to break down more complicated and more common nutrients such as sugars and fatty acids (203). The mechanism of methanogenesis started getting elucidated in early 2 1970s, making methanogens the last major group of microorganisms with its biochemical pathways understood (203).

1.1.2 Ecology of methanogens

Methanogens are widespread in anaerobic environments of a wide range of temperature, salinity and pH, including fresh water and marine sediments, swamps and marshes, landfills, rice paddies, animal intestines, hydrothermal vents and sewer digesters

(52, 201). In these anaerobic environments, methanogens collaborate with members of the Bacteria domain in the conversion of complex organic matters into methane, a critical step of global carbon cycle (Fig. 1-1). During this process, organic matters are first degraded to simple organic compounds such as acetate, methylamines, CO2 and formate with fermentative and H2-producing bacteria. These simple compounds then serve as substrates of methanogens by which methane is produced (52). Methanogenesis is responsible for one billion metric tons of biological methane produced annually, two thirds of which come from the conversion of acetate to methane and the other one third comes from the reduction of CO2 to methane with H2 (188). This huge amount of methane is a potential pollution-free biofuel that human can explore, but at the same time is also an environmental threat. It is reported that methane is a 30 times stronger greenhouse gas than CO2 and accounts for 25% of the total global warming effect (188).

Most of methanogens use one or more of three types of substrates: acetate, CO2 +

H2 and methylated compounds such as methanol, mono-, bi- and trimethylamines.

Some other substrates are also used to a lesser extent. Formate can be used by some 3

CO2 reducers, which oxidize formate to CO2 and then reduces CO2 to methane with the electrons obtained during the oxidation of formate (150). Methanogenesis from H2 + methanol (55), CO (32, 116, 135), dimethylsulfide (132, 176), and some short-chain alcohols (202) have also been reported. Most of these known substrates are simple organic compounds with one or two carbons. Metabolism with more complicated compounds such as amino acids, carbohydrates and fatty acids were never identified with pure methanogens.

The majority of known substrates of methanogenesis are the end metabolism products of fermentative bacteria, suggesting an interspecies interdependence between methanogens and fermentative bacteria. Bacteria break down organic matters into simple organic compounds and H2 while methanogens convert simple organic compounds into methane with electrons derived from H2, favoring thermodynamics for

H2-producing bacteria (205). This close relationship can be demonstrated by an example: a long known and extensively studied ‘pure’ methanogen ‘Methanobacillus omelianskii’ was resolved into two organisms decades after its discovery: the symbiotic bacterium (S organism) and the methanogen Methanobacterium bryanii (25). A H2 transfer between these two organisms was identified, which was believed to be a widespread phenomenon (25).

4

Figure 1-1 The global carbon cycle. Biochemical and chemical processes are both responsible for the cycling of carbon between the aerobic world and anaerobic world. Human activities greatly impacted this cycle by industrial generation of CO2 and biochemically generated methane from agriculture.

It is estimated 40 Tg of methane cannot be oxidized to CO2 and remain in atmosphere annually, accounting for 8% of methane produced globally (73).

1.1.3 Taxonomy and cell biology of methanogens

All known methanogens belong to the kingdom of domain. There are five orders of methanogens: Methanopyrales, Methanococcales,

Methanobacteriales, Methanomicrobiles and . Most members of

Methanopyrales, Methanococcales, Methanobacteriales and Methanomicrobiles 5 conserve energy by reducing CO2 with electrons derived from H2 or formate. A few exceptions exist, such as the Methanosphaera genus of Methanobacteriales order that reduces methanol with H2 (16, 55). Methanosarcinales is the most metabolically diverse order that utilizes methylated compounds and acetate as well as CO2 + H2 as substrates (52). Two genera of this order: and Methanosarcina are the only two known genera that produce methane from acetate (97). Methanosarcina is the only known genus that can use all three major methanogenesis pathways (56).

Methanogens share a lot of similarities in morphology and cellular organization with bacteria. These similarities include the presence of , membrane, circular

DNA molecule carrying genetic information, and the lack of a cell nucleus (170). Like bacteria, multiple morphological types have been found among methanogens, including rod shape of Methanopyrus kandleri (110), coccoid shape of Methanococcus thermolithotrophicus (89) and spiral shape of Methanospirillum hungatei (138). Some

Methanosarcina species form cell aggregates under certain condition (168). Flagella are present in many methanogens such as M. hungatei (53).

Although methanogens share many similarities with bacteria, there are striking differences between the two. Methanogenic cell wall is lack of murein, but sometimes has a psudomurein layer and often has a proteinaceous S layer (170). The membrane of methanogens is comprised of phytanyl ether instead of fatty acid ester lipids (104). The transcriptional machinery is more like that of eukaryotes instead of bacteria, bearing

TATA-binding proteins (141) and an eukaryote-like RNA polymerase (86). 6 1.2 Methanogenesis from three major biochemical pathways

1.2.1 Cofactors utilized in methanogenic pathways

A variety of unique cofactors that do not exist in bacteria are used by methanogens, while many common cofactors used by bacteria are not found in methanogens, examples of which include NADH and NADPH. Figure 1-2 summarizes the structures of the unique cofactors used by methanogens.

F420

F420 is a common electron carrier that functions during methanogenesis from H2 +

CO2 and methylated compounds. It is a two electron carrier that plays a role of

‘common electron currency’ by donating electrons to many enzymes such as

F420-reducing methylenetetrahydromethanopterin dehydrogenase (40, 190). Slight variants of F420 are used in the H2 + CO2 pathway and methanogenesis from methylated compounds, but they showed the same function (100, 186).

Methanophenazine

Methanophenazine (MP) is a quinone-like, highly hydrophobic, membrane-bound electron carrier that was isolated and characterized from membranes of Methanosarcina mazei (3). Because of its extreme hydrophobicity, a water soluble analogue

2-hydroxyphenazine of MP has been used in research of the physiological function of MP 7 (3). It has been shown that MP participates in membrane-bound electron transport chains, serving as the electron shuttle of upstream electron donors to DE type heterodisulfide reductases (3, 91, 130). This electron transport pathway couples transmembrane proton translocation, suggesting MP participates in an ‘MP-loop’ analogous to that of bacterial Q-loop (91).

Figure 1-2 Cofactors of methanogens. This figure is modified from Figure 3 in (52). 8 Methanofuran

Methanofuran (MF) is a small molecule that was first identified as a formyl group carrier during methanogenesis from H2 + CO2 (115). It was later shown that MF serves as a formyl group carrier functioning in the fixation of CO2 by formylmethanofuran dehydrogenase which catalyzes the formylation of MF (99). Formylmethanofuran further passes the formyl group to with a formylmethanofuran: tetrahydromethanopterin formyltransferase, regenerating MF (36).

Tetrahydromethanopterin

Tetrahydromethanopterin (THMPT) is the central one carbon carrier during methanogenesis on all known pathways. A slightly modified variant of THMPT, tetrahydrosarcinapterin (THSPT) is used in methanogenesis from methylated compounds and acetate (33, 100). A series of reactions involving THMPT/THSPT has been identified during methanogenesis, a detailed description of which is present in Section

1.2.2 to 1.2.4 of this chapter.

Coenzyme M

Coenzyme M (HS-CoM) is one of the simplest cofactors in methanogens, but plays a central role in the last step of methanogenesis, serving as the direct methyl group donor to methyl coenzyme M reductase, which releases methane as the product of this 9 methyl transfer reaction. HS-CoM is present in all know methanogens and this reaction is conserved in all methanogenesis pathways (Fig. 1-3, Fig. 1-5, Fig. 1-7).

Coenzyme B

Coenzyme B (HS-CoB) was first isolated as ‘component B’ during studies on methyl coenzyme M reductases, and was shown to be an essential component in this last step of methanogenesis (42). It was later characterized as a small molecule organic compound (133), and was renamed coenzyme B. Catalyzed by methyl coenzyme M reductase, HS-CoB reduces CH3-CoM, releasing methane and forming the heterodisulfide of coenzyme M and coenzyme B, CoM-S-S-CoB, which was also shown to be an essential of methanogenesis (Fig. 1-2) (17).

1.2.2 H2 + CO2 pathway

Overview

Almost all known methanogens can utilize the H2 + CO2 methanogenesis pathway.

This pathway can be summarized in Equation 1-1 and Figure 1-3:

o CO2 + 4H2 → CH4 + 2H2O, ΔG ’ = -131 kJ/mol

Equation 1-1 Methanogenesis from H2 + CO2

10

The conversion of CO2 to CH4 first starts by converting CO2 to CH3-THMPT with the collaboration of multiple reactions, listed below:

Equation 1-2 CO2 fixation with methanofuran. FdR, ferredoxin reduced; FdO, ferredoxin oxidized.

Equation 1-3 Formyl group transfer to THMPT.

Equation 1-4 Cyclization of N5-formyl-THMPT. 11

Figure 1-3 The H2 + CO2 methanogenesis pathway. Abbreviations: MF, methanofuran; FdR, ferredoxin reduced; FdO, ferredoxin oxidized; Fmd, molybdenum-containing formylmethanofuran dehydrogenase; Fwd, tungsten-containing formyl-methanofuran dehydrogenase; THMPT, tetrahydromethanopterin; Ftr, formylmethanofuran: tetrahydromethanopterin formyltransferase; Mch.

5,10-methenyltetrahydromethanopterin cyclohydrolase; Hmd, H2-forming methylenetetrahydromethanopterin dehydrogenase; Mtd, F420-reducing methylenetetrahydromethanopterin dehydrogenase; Mer, methylenetetrahydromethanopterin reductase; Mtr,

N5-methyltetrahydromethanopterin: coenzyme M methyltransferase; Mcr, methyl coenzyme M reductase. 12

5 10 Equation 1-5 F420 dependent reduction of N , N -methenyl-THMPT.

5 10 Equation 1-6 H2 dependent reduction of N , N -methenyl-THMPT.

Equation 1-7 Reduction of N5, N10-methylene-THMPT. 13

CO2 fixation with methanofuran

Equation 1-2 shows the fixation of CO2 catalyzed by formylmethanofuran dehydrogenase (Fmd/Fwd). This complex has been purified from mesophilic M. barkeri (98, 192), thermophilic Methanothermobacter marburgensis (15, 20, 87, 88) and

Methanobacterium wolfei (151, 152), as well as hyperthemophilic M. kandleri (193).

All of these enzymes were identified to contain one molecule of guanine dinucleotide (MGD) as the cofactor (Figure 1-2) (99), with the exception of molybdenum-containing formylmethanofuran dehydrogenase from M. marburgensis. In this enzyme molybdopterin adenine dinucleotide (MAD) and molybdopterin hypoxanthine dinucleotide (MHD) were also identified, showing the same function as

MGD (20). All formylmethanofuran dehydrogenases identified so far are , containing multiple iron sulfur clusters (15, 20, 98, 151, 152, 192).

Although the formylmethanofuran dehydrogenases identified so far share obvious similarities, their metal content and subunit composition vary dramatically. Enzyme purified from M. barkeri is a five subunit complex containing molybdenum in its active site (192). Two isoenzymes from M. marburgensis were identified, one of which is a three-subunit molybdenum-containing enzyme, and the other is a four-subunit tungsten-containing enzyme (87). Both isoenzymes from M. wolfei have a α2β2γ2 configuration. One of them is a molybdenum-containing enzyme and the other is a tungsten-containing enzyme (151, 152). M. kandleri has two tungsten-containing isoenzymes, one of which is selenium dependent (193). The functions of each subunit haven’t been well established, except for FmdB (for molybdenum-containing enzymes) 14 and FwdB (for tungsten-containing enzymes) which were shown to contain MGD binding motif, suggesting their role as the catalytic subunits (87).

The mechanism of reaction detailed in Equation 1-2 is still largely unknown.

The electron donor hasn’t been identified, although F420 has been excluded, leaving ferredoxin as a very promising candidate (98). CO2 molecule has been shown as the

- preferred ‘CO2’ species for formylmethanofuran dehydrogenase instead of HCO3 , suggesting N-carboxymethanofuran as an intermediate (191). This raises the hypothesis that the first step of CO2 fixation is the spontaneous reversible formation of

N-carboxymethanofuran from CO2 and methanofuran, shown in Equation 1-8. This hypothesis was further supported by kinetic studies of the formation of

N-carboxymethanofuran under environmental conditions (10).

Equation 1-8 Spontaneous reversible carboxylation of methanofuran.

Formyl transfer to tetrahydromethanopterin (THMPT)

The transfer of formyl group from methanofuran to THMPT is catalyzed by a formylmethanofuran: tetrahydromethanopterin formyltransferase (Ftr) (Fig. 1-3). Ftr has been purified and characterized from M. barkeri (24, 109), Methanothermobacter 15 thermoautotrophicus (35, 36), Methanothermus fervidus (114) and M. kandleri (22, 164).

None of these proteins show the binding of a cofactor. These proteins are present in equilibrium of monomer, homodimer and homotetramer forms (127, 162). The crystal structures of Ftrs from M. barkeri and M. kandleri have been solved, suggesting a biologically active tetrameric form (47, 127, 161). The co-crystallization of Ftr together with THMPT and N-formylmethanofuran has been achieved and the structures were solved, further elucidating the relationship between Ftr and its substrate in detail (5).

The oligomeric state of Ftr from M. kandleri changes with salt concentration. The biological active dimeric/tetrameric form increases with the increase of salt concentration

(162). The oligomeric states of Ftrs from M. barkeri are constantly dimeric/tetrameric state regardless of salt concentration (127).

Enzymatic mechanism studies of the formyl transfer reaction suggested

N5-formyl-THMPT instead of N10-formyl-THMPT is the product of this reaction (24, 36).

N-furfurylformamide can also serve as a pseudo-substrate for formyl transfer reaction instead of N-formylmethanofuran, although with a much higher Km and a much lower

Vmax than N-formylmethanofuran (21). A ternary-complex mechanism has been suggested for Ftr (22).

The salt tolerance, temperature tolerance and enzymatic activity of different Ftrs underwent intensive studies, leading to a better understanding of heat tolerance for hyperthermophiles. Ftrs from mesophilic M. barkeri and hyperthermophilic M. kandleri reach half-maximal activity at 0.01 M and 1.0 M K2HPO4, respectively (127). The maximum stimulation by salt of M. barkeri and M. kandleri are 5- and >1000-fold respectively (22), agreeing with the observation that M. kandleri has a very high 16 intracellular salt concentration (90). Ftr from M. kandleri denatures above 50 ℃, however, in a high salt environment, it is stable up to 130 ℃ (47). This salt dependent heat tolerance pattern was also found during the research of other proteins of M. kandleri, suggesting a common strategy to combat high heat in this hyperthomophile.

5, 10-Methenyl-THMPT cyclohydrolase

The non-energetic reversible conversion of N5-formyl-THMPT to N5,

N10-methenyl-THMPT (Equation 1-4) is catalyzed by

5,10-methenyltetrahydromethanopterin cyclohydrolase (Mch) (184). This enzyme has been purified and characterized in M. thermoautotrophicus, M. barkeri and M. kandleri

(23, 34, 178). Mch from M. thermoautotrophicus and M. barkeri were shown to be functional dimers (34, 178). Initial studies on the oligomeric state of Mch from M. kandleri suggested it is monomeric (23), but later studies showed its structure contains trimeric binding surface, suggesting it is a functional trimer (62). No prosthetic groups have been identified in any of the known Mchs.

Attempts to constructΔmch deletion strain of M. acetivorans were made but unsuccessful, suggesting mch is essential for M. acetivorans (68). Δmch deletion strain of M. barkeri was constructed, but unable to grown on any substrate other than methanol

+ H2 + CO2 (68). This severe limitation on growth substrate suggests the essential role of Mch in methanogenesis from methanol and from H2 + CO2. The incapability ofΔ mch strain of M. barkeri to grow on acetate is a bit surprising because Mch isn’t involved 17 in methanogenesis from acetate (117), but nonetheless agrees with a hypothesis that Mch is also involved in biosynthesis by participating in the generation of CO2 from methyl-THMPT, while providing reducing equivalents (68).

Methylene-THMPT dehydrogenase

Two enzymes are involved in the reduction of N5, N10-methenyl-THMPT to N5,

10 N -methylene-THMPT: H2-forming methylenetetrahydromethanopterin dehydrogenase

(Hmd, Equation 1-6) and F420-reducing methylenetetrahydromethanopterin dehydrogenase (Mtd, Equation 1-5). They have different molecular characteristics and reductant requirement but perform the same role during methanogenesis (190).

Hmd has been purified and characterized from M. marburgensis, M. kandleri, M. wolfei and M. thermolithotrophicus (78, 125, 129, 206, 207). These proteins are either monomers or homotetramers (78, 125, 206). They are generally oxygen sensitive and repressed when oxygen level is high (190). Although they show a hydrogenase activity, unlike other hydrogenases, they don’t have a significant or iron content (125, 206,

207).

Mtd has been purified and characterized from M. thermoautotrophicus, M. barkeri and M. kandleri. They are either homohexmers or homooctamers and relatively stable in air, possessing no identifiable prosthetic groups (45, 80, 103, 177, 179). The crystal structure of Mtd from M. kandleri has been solved (70, 71, 195).

These two types of methylene-THMPT dehydrogenases use different reductants as electron sources: Hmd uses H2 and Mtd uses F420. Most of methanogens that can 18 grow on H2 + CO2 contain both dehydrogenases at the same time, while members of the order Methanomicrobiales contain only Mtd (103). It was shown that the transcription level and enzymatic activity of Hmd is higher during early growth while Mtd starts taking place of Hmd after mid-log phase (128, 134). The studies of transcription level of hmd and mtd at low- and high-H2 levels revealed that hmd is upregulated when H2 is sufficient and mtd is upregulated when H2 is limiting (84, 134). These observations lead to the hypothesis that Hmd primarily functions in H2-sufficient situations when metabolism is high and the amount of F420 in cells is the limiting factor, while Mtd takes over when H2 is limiting and F420 is no longer the limiting factor (134). This gives the cells a comparative advantage in competing for energy in the environment by providing maximized reductant supply under different situations.

Another potential function of Hmd and Mtd was identified during research on

Ni-limited growth of M. marburgensis (6). The levels of Hmd and Mtd were shown to be upregulated when nickel is limited during cell growth, while F420 reducing hydrogenase (Frh) which catalyzes the reduction of F420 with H2 is downregulated (6).

Because Frh is a nickel-iron-flavin containing protein, its downregulation can be explained by the lack of metals needed for its synthesis. The upregulation of Hmd and

Mtd under Ni-limited scenarios leads to the hypothesis that Hmd and Mtd work together in forming a Hmd-Mtd cycle, in which Hmd catalyses the reduction of N5,

10 5 N -methenyl-THMPT with H2 while Mtd catalyses the oxidation of N ,

10 N -methenyl-THMPT with F420 (Fig. 1-4). The net outcome of this cycle is the reduction of F420 with H2, the function of F420 reducing hydrogenase. This provides an 19 alternative way of F420 regeneration and serves to keep the F420 pool reduced when nickel is limiting in the environment.

Figure 1-4 Hypothetical Hmd-Mtd cycle.

Methylene-THMPT reductase

5, 10-methylenetetrahydromethanopterin reductase (Mer) catalyses the reduction

5 10 5 of N , N -methylene-THMPT to N -methyl-THMPT with F420 as the electron source

(Equation 1-7). This protein has been purified and characterized from M. thermoautotrophicus, M. barkeri, M. kandleri and M. marburgensis (122-124, 177, 180).

These proteins share similar characteristics including: high F420 specificity, lack of a , oxygen stable and a ternary-complex catalytic mechanism (122-124,

177, 180). Mer from M. thermoautotrophicus, M. marburgensis and M. barkeri were shown to be tetramers (123, 124, 177) while Mer from M. kandleri is an octamer (122).

The crystal structures of Mer from M. marburgensis and M. kandleri, as well as the 20 cocrystalization of Mer from M. barkeri with F420 have been achieved and solved (8,

163).

N5-methyltetrahydromethanopterin: coenzyme M methyltransferase

The N5-methyltetrahydromethanopterin: coenzyme M methyltransferase (Mtr) catalyzes the reaction of methyl group transfer from N5-methyl-THMPT to coenzyme M

(Equation 1-9). It is a corrinoid membrane-bound protein complex (54, 154, 155) containing 5-hydroxybenzimidazolyl cobamide as well as one 4Fe-4S cluster as prosthetic groups (57, 120, 140). Mtr has been purified or partially purified from M. marburgensis, M. thermoautotrophicus and M. mazei (57, 102, 118), showing it has 3 to

7 subunits. However, complete mapping of the mtr transcription unit suggested eight cotranscribing genes, indicating a more complex subunit composition (77, 119). The

Δmtr strain of M. barkeri was constructed and the mutant was unable to grown on acetate, methanol or H2 + CO2, suggesting it is essential in all major methanogenesis pathways

(200).

Equation 1-9 Methylation of coenzyme M.

The mechanism of the methyl transfer reactions has been extensively studied, showing a ping-pong mechanism that involves two steps: 1) The methyl transfer from 21

5 N -methyl-THMPT to Co(I), methylating and oxidizing the cobalamin to CH3-Co(III); 2) the methyl transfer from CH3-Co(III) to coenzyme M, forming methyl coenzyme M, as well as regenerating cobalamin to Co(I) state (58, 199). The stimulatory effects of ions were also studied, showing the methyltransferase activity of Mtr is dependent on sodium ions (13, 58, 118, 199). This leads to the hypothesis and eventual discovery that Mtr is a primary Na+ pump. The methylation of cobalamin was identified as the exergonic step, during which Na+ was hypothesized to be translocated (12, 13). The studies of liposomes incorporating purified Mtr showed a stoichiometry of 1.7 sodium ions translocated per pair of electrons transported, suggesting a 2Na+/2e stoichiometry of this sodium pump (118). It was hypothesized that the Na+ gradient generated with Mtr can

+ + + + be used by Na -ATPases or Na /H antiporters together with A1A0-type H -ATPases to couple ATP synthesis (117).

The function of the subunits of Mtr hasn’t been completely understood except for

MtrA and MtrH. MtrA was shown to be the cobalamin binding subunit that binds cobalamin via a histadine at its binding site (74, 175). Mutagenesis studies identified the specific histadine residue for cobalamin binding (75, 147). MtrH was shown to catalyze the methyl transfer from N5-methyl-THMPT to cobalamin (85).

Methyl coenzyme M reductase

The most important step of methanogenesis, the generation of methane (Equation

1-10), is catalyzed by methyl coenzyme M reductase (Mcr), one of the most abundant protein complexes in methanogens which takes up approximately 10% of cellular protein 22

(79). It catalyzes the reduction of CH3-CoM by HS-CoB to form CoM-S-S-CoB and generate methane (160). This protein has been purified and characterized from M. marburgensis, M. thermoautotrophicus, M. thermophila, M. kandleri and M. barkeri (41,

44, 61, 95, 143), suggesting it is a heterohexamer with a configuration of α2β2γ2. Mcr has been shown to bind two molecule of F430, a nickel-containing porpinoid, for every molecule of holoenzyme (Fig. 1-2) (41). Besides the genes encoding the three subunits

(mcrA, mcrB, mcrG), two extra genes mcrC and mcrD were found to cotranscribe with other mcr genes, forming a mcrBDCGA operon (198). The gene products of mcrC and mcrD were purified and characterized, but a clear function of them and their relationship with Mcr is not clear (44, 157, 172, 173).

Equation 1-10 Release of methane from methyl coenzyme M.

The crystal structures of Mcr in multiple species in multiple states have been solved (46, 61, 159), showing F430 is embedded at the bottom of a 50 Å long hydrophobic channel, forming the active site for CH3-CoM reduction. This active site shows a unique feature that is rarely seen: the presence of five conserved methylated amino acids, a thioglycine, a N-methyl-histadine, a S-methyl cysteine, a 5-(S)-methyl-arginine and a

2-(S)-methyl glutamine (46, 61, 159).

The mechanism of reactions involving Mcr has been the subject of intensive biochemical and spectroscopic studies. It was found that purified Mcr has a very low 23 activity when comparing with activities assayed with cell lysate (43). Various reductants such as B12, Ti(III) and dithiolthreitol were shown to stimulate the Mcr activity and were initially proposed to be a requirement for Mcr activity (7, 43, 145). It was later found out that Mcr can be activated by various means such as photoactivation,

H2-preincubation or sulfide treatment of cells prior to purification and Ti(III) reduction, leading to the eventual understanding that Ni(I)F430 is the active species of F430 and the low potential of reaction system helps maintaining the reduced state of F430 (14, 60, 136,

142).

Addition to the intensively studied Mcr (designated Mcr I), an isozyme of Mcr

(designated Mcr II) has been discovered in Methanothermobacter species, showing similar molecular properties but slightly different catalytic properties (18, 144).

Expression and genetic analysis showed different transcription and expression patterns.

Transcriptional analysis showed the mcr genes encoding Mcr I are transcribed in late log phase to stationary phase, while mrt genes encoding Mcr II are transcribed in early to mid log phase (134, 139). Expression analysis showed a similar pattern, with Mcr I expressed in late log phase to stationary phase while Mcr II expressed in early log phase

(19, 144). These studies suggested a transcription-level regulation of Mcr I and Mcr II expression: Mcr I is expressed when substrate-energy supply is limited and Mcr II is expressed when substrate-energy supply is unlimited (19). 24 ABC type Heterodisulfide reductase

In H2 + CO2 grown methanogens, the regeneration of coenzyme M and coenzyme

B is catalyzed by the ABC type heterodisulfide reductase (HdrABC) (Equation 1-11).

This protein complex has been purified and characterized in M. marburgensis, showing a

α4β4γ4 subunit composition (81, 156). Sequence analysis of the three subunits of

HdrABC suggested it’s a protein complex binding multiple prosthetic groups (82). The sequence of HdrA contains binding motifs for one FAD and four 4Fe-4S clusters. The sequence of HdrC suggests it binds to two 4Fe-4S clusters. Although HdrB doesn’t contain a typical iron sulfur cluster binding motif CX2CX2CX3CP, EPR analysis of E. coli expressed HdrB from M. marburgensis showed the binding of one 4Fe-4S cluster

(72). Zinc binding was also observed in this heterologously produced HdrB (72).

Spectroscopic analysis showed iron sulfur clusters is the active site of HdrABC; however, due to the complicated cofactor content of this protein complex, it’s hard to identify the subunit that binds to the active site 4Fe-4S cluster (37, 38, 126). Sequence analysis suggested a hypothetical fusion protein of HdrB and HdrC is homologous to the active site binding HdrD subunit in DE type heterodisulfide reductases, leading to the hypothesis that HdrB or HdrC is the active site of HdrABC (108). One recent analysis of the cofactor content of HdrB suggested a unique iron sulfur binding motif, termed

CCG, and hypothesized that HdrB contains the active site for heterodisulfide reduction

(72).

CoM-S-S-CoB + H2→CoM-SH + CoB-SH

Equation 1-11 Regeneration of coenzyme M and coenzyme B. 25

In M. marburgensis, HdrABC co-purifies with a F420-non-reducing hydrogenase

Mvh (156). This leads to the hypothesis that HdrABC forms a tightly associated complex with the MvhADG complex in vivo. Mvh is hypothesized to oxidize and provides electrons to HdrABC via the electron transfer between HdrA and MvhD

(171). A recent review hypothesized an electron bifurcation pathway in HdrABC, in which part of the electrons from Mvh are used to reduce CoM-S-S-CoB and part of them are used to reduce ferredoxin (186)

1.2.3 Methylotrophic pathway

Overview

Unlike the H2 + CO2 pathway which is utilized by most methanogens, methanogenesis from methylated compounds only occurs in a handful of methanogens, including members of the order Methanosarcinales and three other species

Methanosphaera cuniculi, Methanosphaera stadtmanae and Methanococcus halophilus

(52, 100). Members of the genus are obligate methylotrophic methanogens that can only grow on methylated compounds (100). A number of methylated compounds are utilized by methanogens as carbon and energy sources, out of which methanol, mono-, bi- and trimethylamine are the most common substrates used by most methylotrophic species. Some methanogens are known to use dimethylsulfide as the carbon and energy source, but this substrate is not commonly used by

(100). 26 The methylotrophic pathway of methanogenesis can be summarized in Equation

1-12 using methanol as the representative substrate (100):

o 4CH3OH → 3CH4 + CO2 + 2H2O , ΔG ’ = -106 kJ/mol

Equation 1-12 Methanogenesis from methanol. Figure 1-5 summarizes the methylotrophic pathway. The initial step of methylotrophic pathway involves a substrate-specific methyltransferase that transfers the methyl groups to coenzyme M, yielding CH3-CoM. Three molecules of CH3-CoM are reduced to

CH4, taking up six electrons, which are provided by one molecule of CH3-CoM oxidized to CO2. The ‘reduction branch’ of the methylotrophic pathway is the same as the reduction of CH3-CoM to CH4 in H2 + CO2 pathway, with a different heterodisulfide reductase. The ‘oxidative branch’ of the methylotrophic pathway is the same as the reduction of CO2 to CH3-CoM in H2 + CO2 pathway in the reverse order. Slight differences between the ‘oxidative branch’ of methylotrophic pathway and H2 + CO2 pathway exist: the lack of a H2-forming methylene-THMPT dehydrogenase and slightly different cofactors used in methylotrophic pathway, such as variants of THMPT, MF and

F420 (100).

Methyltransferase

The first step of methanogenesis with methylated compounds is the transfer of methyl groups to coenzyme M. The enzymes catalyzing this reaction have been 27

Figure 1-5 Methanogenesis from methylated compounds. Abbreviations: MF, methanofuran;

FdR, ferredoxin reduced; FdO, ferredoxin oxidized; Fmd, molybdenum containing formylmethanofuran dehydrogenase; Fwd, tungsten containing formyl-methanofuran dehydrogenase; THSPT, tetrahydrosarcinapterin; Ftr, formylmethanofuran: tetrahydromethanopterin formyltransferase; Mch.

5,10-methenyltetrahydromethanopterin cyclohydrolase; Mtd, F420-reducing methylenetetrahydromethanopterin dehydrogenase; Mer, methylenetetrahydromethanopterin reductase; Mtr,

N5-methyltetrahydromethanopterin:coenzyme M methyltransferase; Mcr, methyl coenzyme M reductase;

MtaA, MtaB, MtbA, MtbB, MtmB, MttB, substrate specific methyltransferases.

extensively studied in M. barkeri. Different but analogous enzymatic systems have been identified in methanogenesis from different methylated compounds. 28 The methanol: coenzyme M methyltransferase system is composed of three proteins termed MtaA, MtaB and MtaC. As is shown in Figure 1-6, a two step methyl transfer reaction has been characterized: first the methyl transfer from methanol to MtaC and then from MtaC to coenzyme M. MtaC serves as an intermediate of methyl transfer from methanol to coenzyme M (148). This protein is a monomeric protein that binds to corrinoid at a 1:1 cofactor/peptide ratio (148). MtaB is a zinc-containing protein that catalyzes the methyl transfer from methanol to MtaC (149). MtaB and MtaC form a protein complex in a α2β configuration (α stands for MtaB) (31, 146). This protein complex is also called MT1 and has been crystallized (69). MtaA is a zinc protein that serves as a methylcobamide: coenzyme M methyltransferase (113). This oxygen stable protein was cloned and heterologously expressed in E. coli (76).

Figure 1-6 Methyl group transfer from methanol to coenzyme M.

Methyl group transfer from methylamines to coenzyme M is catalyzed by analogous but different methyltransferases. They share the same MtaA analog catalyzing methylcobamide: coenzyme M methyltransferase activity, annotated as MtbA

(27, 50, 64, 204). However, different MtaB and MtaC analogues are used for different methylamines: MtmB and MtmC for monomethylamine, MtbB and MtbC for dimethylamine and MttB and MttC for trimethylamine. These methyltransferase 29 systems have been purified and studied in details, suggesting a similar reaction scheme with that of methanol system (28, 48, 49). The methyl transfer from dimethylsulfide to coenzyme M uses a different catalytic scheme comparing with other methylated compounds. Although the corrinoid protein is conserved in dimethylsulfide system, one single protein MtsA catalyzes both the methyl transfer from dimethylsulfide to cobalamin and from cobalamin to coenzyme M (176).

Although cobalt is the most powerful neucleophile in nature, thus fitting the role of methyl carrier perfectly, it is prone to inactivation by oxidants. This leads to the requirement of a reductive reactivation system for cobalamins. In methyltransferase systems, this reactivation system has been well characterized. A methyltransferase activation protein (MAP) has been shown to activate MT1 in and methanol systems with ATP and H2 or ferredoxin (30, 31, 189, 196, 197). Another 4Fe-4S cluster binding protein RamA also showed activation of methylamine MT1 using ATP, but its relative role with MAP is currently unknown (51).

DE type heterodisulfide reductase

Unlike the H2 + CO2 pathway, both methanogenesis from acetate and methylated compounds use a DE type heterodisulfide reductase (HdrDE) for the reduction of

CoM-S-S-CoB. This membrane-bound protein complex has been purified and characterized from methanol-grown M. barkeri and acetate-grown M. thermophila (83,

108, 165). Attempts were made to constructΔhdrED1 mutant of M. acetivorans but unsuccessful, consistent with the theory that HdrDE is the primary heterodisulfide 30 reductase during acetate and methanol metabolism (26). On the other hand, Δ hdrA1C1B1 mutant of M. acetivorans was successfully constructed and showed growth on acetate and methanol, further supporting the theory that HdrABC is not the primary

Hdr for the reduction of CoM-S-S-CoB during growth on acetate or methanol (26).

Purified HdrDE is a heterodimer of two subunits: HdrD and HdrE (Figure 1-7).

HdrD is an iron-sulfur protein shown to bind two 4Fe-4S clusters, and was proposed to be the catalytic subunit of HdrDE (83, 108, 130, 165). HdrE was shown to be a b-type cytochrome binding two and was proposed to be involved in electron transfer (83,

108, 130, 165). Methanophenazine was shown to be the electron donor of HdrDE, and this electron transport was shown to couple transmembrane proton translocation with a

2e/2H+ stoichiometry (3, 11, 91, 165).

Figure 1-7 DE-type heterodisulfide reductase. 31 1.2.4 Acetate pathway

Overview

Although only two genera of methanogens, Methanosarcina and Methanosaeta use acetate as the substrate for methanogenesis, it is surprisingly responsible for two thirds of biologically produced methane globally (167, 185). The majority of the biochemical pathway, summarized in Figure 1-8, is different from those of H2 + CO2 and methylotrophic pathway. In sequence, the following reactions happen (THSPT: tetrahydrosarcinapterin, a THMPT derivative used in methanol and acetate metabolism):

Figure 1-8 Methanogenesis from acetate. Abbreviations: FdO, ferredoxin oxidized; FdR, ferredoxin reduced; Ack, acetate kinase; Pta, phosphotransacetylase; CODH/ACS, carbon monoxide dehydrogenase/acetyl-CoA synthase; Mtr, N5-methyltetrahydromethanopterin: coenzyme M methyltransferase; Mcr, methyl coenzyme M reductase. 32

- 2- CH3COO + ATP → CH3COOPO3 +ADP

Equation 1-13 Activation of acetate.

2- CH3COOPO3 + CoA-SH → CH3COSCoA + Pi

Equation 1-14 Acetyl group transfer to coenzyme A.

CH3COSCoA + THSPT + FdO→ CO2 + CoA-SH + CH3-THSPT + FdR

Equation 1-15 Oxidation of carbonyl group and synthesis of methyl-THSPT.

CH3-THSPT + CoM-SH → CH3-CoM + THSPT

Equation 1-16 Methyl transfer to coenzyme M.

CH3-CoM + CoB → CH4 + CoM-S-S-CoB

Equation 1-17 Reduction of methyl group to methane.

CoM-S-S-CoB + MPH2 → CoM-SH + CoB-SH + MP

Equation 1-18 Regeneration of coenzyme M and coenzyme B. 33 Reactions shown in Equation 1-16, 1-17 and 1-18 are catalyzed by

N5-methyl-THMPT: coenzyme M methyltransferase, methyl coenzyme M reductase and

DE type heterodisulfide reductase, described in section 1.2.2 and section 1.2.3 of this chapter. Reactions shown in Equation 1-13, 1-14, 1-15 are catalyzed by acetate kinase, phosphotransacetylase and carbon monoxide dehydrogenase/acetyl-CoA synthase.

Methanogenesis from acetate can be summarized in Equation 1-19:

- + , o CH3COO +H → CH4 + CO2 ΔG ’ = -36 kJ/mol

Equation 1-19 Methanogenesis from acetate.

As is shown in Equation 1-19, for every molecule of acetate metabolized, the cells can only obtain enough energy for the synthesis of one ATP. This low amount of energy methanogens can conserve during methanogenesis from acetate contributes to the long doubling time and high metabolism turnovers of acetate-metabolizing methanogens.

Acetate kinase

The first step of methanogenesis from acetate is the activation of acetate to acetylphosphate by acetate kinase (Ack). Ack has been purified and characterized from

M. thermophila and M. mazei strain 2-P (4, 112, 187), showing it’s a homodimer that requires Mg2+ for its activity (4). Comparison of protein content of acetate- versus methanol- grown M. thermophila suggested Ack is upregulated in acetate-grown cells 34 (94). It was later confirmed that an upregulation of ack transcription takes place in acetate-grown cells versus methanol-grown cells (166). The crystal structure of Ack from M. thermophila has been solved (29).

Phosphotransacetylase

Activated acetyl group is transferred to coenzyme A by phosphotransacetylase

(Pta). Acetyl-CoA, the substrate of CO dehydrogenase/Acetyl-CoA synthase, is the product of this reaction. Pta was purified and characterized from M. thermophila (112,

121), suggesting it’s a monomeric protein. The upregulation of Pta in acetate- versus methanol- grown M. thermophila was observed, further supporting the role of Pta during acetate metabolism (94). Transcriptional studies showed that pta and ack form an operon, suggesting the coherent effort of Pta and Ack in activating acetate to acetyl-CoA

(166). The crystal structure of Pta from M. thermophila has been elucidated (92, 93).

CO dehydrogenase/Acetyl-CoA synthase

A bifunctional CO dehydrogenase/Acetyl-CoA synthase complex (Cdh) catalyzes the reaction shown in Equation 1-15. During this reaction, ferredoxin is reduced with electrons obtained from the oxidation of carbonyl group in acetyl-CoA. The methyl group of acetyl-CoA is transferred to the THMPT derivative THSPT, forming methyl-THSPT (63). Methyl-THSPT is then involved in a methyl transfer reaction catalyzed by N5-methyl-THMPT: coenzyme M methyltransferase described in section 35 1.2.2. Cdh is one of the most abundant protein complexes in acetate-utilizing methanogens, taking up 5-10% of cellular proteins (107, 183). Comparison of the content of Cdh and the mRNA encoding it in acetate- versus methanol- grown methanogens suggested it’s upregulated in acetate-grown cells (106, 169, 183). This protein complex has also been shown to catalyze the synthesis of acetyl-CoA from one-carbon compounds in both aceticlastic methanogens and non-aceticlastic methanogens including M. barkeri, M. thermophila, M. marburgensis and

Methanococcus maripaludis (2, 101, 111, 158, 174). Acetyl-CoA can then be used as a carbon source for biosynthesis.

Out of acetate-utilizing methanogens, Cdh complex has been purified from M. barkeri, M. thermophila, M. acetivorans and Methanosarcina frisia Gö1 (39, 63, 67, 107,

183, 194). Both M. barkeri and M. thermophila have five subunits αβγδε present in equal stoichiometry and contain Ni, Fe and Co. M. frisia Cdh only has two subunits, but that probably is an artifact due to disassociation of Cdh complex during purification, which happened to M. barkeri Cdh purification in early research as well. The Cdh complex of M. thermophila underwent extensive biochemical studies, showing it can be separated into two subcomplexes: αε and γδ subcomplexes (1). Metal content and spectroscopic analysis of each subcomplex suggested that the αε subcomplex contains a

Ni/4Fe-4S dinucleii and the γδ subcomplex contains a Co/4Fe-4S dinucleii with Co in the form of corrinoid (96, 105, 181, 183). It was suggested that the Ni/4Fe-4S core of the

αε subcomplex catalyzes the oxidation of the carbonyl group in acetyl-CoA, passing electrons to ferredoxin while transferring the methyl group to γδ subcomplex (1, 96, 182).

The γδ subcomplex constitutes a corrinoid binding domain that catalyzes the methyl 36 transfer reaction to THSPT (65, 96, 183). Subunit β is a Ni-containing protein and was suggested to be involved in acetyl-CoA binding and C-C bond activation (59, 66, 131)

1.3 References

1. Abbanat, D. R., and J. G. Ferry. 1991. Resolution of component proteins in an

enzyme complex from Methanosarcina thermophila catalyzing the synthesis or

cleavage of acetyl-CoA. Proc Natl Acad Sci U S A 88:3272-6.

2. Abbanat, D. R., and J. G. Ferry. 1990. Synthesis of acetyl coenzyme A by

carbon monoxide dehydrogenase complex from acetate-grown Methanosarcina

thermophila. J Bacteriol 172:7145-50.

3. Abken, H. J., M. Tietze, J. Brodersen, S. Baumer, U. Beifuss, and U.

Deppenmeier. 1998. Isolation and characterization of methanophenazine and

function of phenazines in membrane-bound electron transport of Methanosarcina

mazei Go1. J Bacteriol 180:2027-32.

4. Aceti, D. J., and J. G. Ferry. 1988. Purification and characterization of acetate

kinase from acetate-grown Methanosarcina thermophila: evidence for regulation

of synthesis. J Biol Chem 263:15444-8.

5. Acharya, P., E. Warkentin, U. Ermler, R. K. Thauer, and S. Shima. 2006.

The structure of formylmethanofuran: tetrahydromethanopterin formyltransferase

in complex with its coenzymes. J Mol Biol 357:870-9.

6. Afting, C., A. Hochheimer, and R. K. Thauer. 1998. Function of H2-forming

methylenetetrahydromethanopterin dehydrogenase from Methanobacterium 37

thermoautotrophicum in coenzyme F420 reduction with H2. Arch Microbiol

169:206-10.

7. Ankel-Fuchs, D., and R. K. Thauer. 1986. Methane formation from

methyl-coenzyme M in a system containing methyl-coenzyme M reductase,

component B and reduced cobalamin. Eur J Biochem 156:171-7.

8. Aufhammer, S. W., E. Warkentin, U. Ermler, C. H. Hagemeier, R. K.

Thauer, and S. Shima. 2005. Crystal structure of

methylenetetrahydromethanopterin reductase (Mer) in complex with coenzyme

F420: Architecture of the F420/FMN binding site of enzymes within the nonprolyl

cis-peptide containing bacterial luciferase family. Protein Sci 14:1840-9.

9. Baker, H. A. 1956. Bacterial Fermentations. Wiley, New York.

10. Bartoschek, S., J. A. Vorholt, R. K. Thauer, B. H. Geierstanger, and C.

Griesinger. 2000. N-carboxymethanofuran (carbamate) formation from

methanofuran and CO2 in methanogenic archaea: thermodynamics and kinetics of

the spontaneous reaction. Eur J Biochem 267:3130-8.

11. Baumer, S., E. Murakami, J. Brodersen, G. Gottschalk, S. W. Ragsdale, and

U. Deppenmeier. 1998. The F420H2: heterodisulfide oxidoreductase system from

Methanosarcina species: 2-Hydroxyphenazine mediates electron transfer from

F420H2 dehydrogenase to heterodisulfide reductase. FEBS Lett 428:295-8.

12. Becher, B., V. Muller, and G. Gottschalk. 1992. The

methyl-tetrahydromethanopterin: coenzyme M methyltransferase of

Methanosarcina Strain Go1 is a primary sodium pump. FEMS Microbiol Lett

91:239-244. 38 13. Becher, B., V. Muller, and G. Gottschalk. 1992.

N5-methyl-tetrahydromethanopterin: coenzyme M methyltransferase of

Methanosarcina strain Go1 is an Na+-translocating membrane protein. J Bacteriol

174:7656-60.

14. Becker, D. F., and S. W. Ragsdale. 1998. Activation of methyl-SCoM reductase

to high specific activity after treatment of whole cells with sodium sulfide.

Biochemistry 37:2639-47.

15. Bertram, P. A., R. A. Schmitz, D. Linder, and R. K. Thauer. 1994. Tungstate

can substitute for molybdate in sustaining growth of Methanobacterium

thermoautotrophicum: identification and characterization of a tungsten isoenzyme

of formylmethanofuran dehydrogenase. Arch Microbiol 161:220-8.

16. Biavati, B., M. Vasta, and J. G. Ferry. 1988. Isolation and characterization of

"Methanosphaera cuniculi" sp. nov. Appl Environ Microbiol 54:768-71.

17. Bobik, T. A., K. D. Olson, K. M. Noll, and R. S. Wolfe. 1987. Evidence that the

heterodisulfide of coenzyme M and 7-mercaptoheptanoylthreonine phosphate is a

product of the methylreductase reaction in Methanobacterium. Biochem Biophys

Res Commun 149:455-60.

18. Bonacker, L. G., S. Baudner, E. Morschel, R. Bocher, and R. K. Thauer.

1993. Properties of the two isoenzymes of methyl-coenzyme M reductase in

Methanobacterium thermoautotrophicum. Eur J Biochem 217:587-95.

19. Bonacker, L. G., S. Baudner, and R. K. Thauer. 1992. Differential expression

of the two methyl-coenzyme M reductases in Methanobacterium 39 thermoautotrophicum as determined immunochemically via isoenzyme-specific

antisera. Eur J Biochem 206:87-92.

20. Borner, G., M. Karrasch, and R. K. Thauer. 1991. Molybdopterin adenine

dinucleotide and molybdopterin hypoxanthine dinucleotide in

formylmethanofuran dehydrogenase from Methanobacterium

thermoautotrophicum (Marburg). FEBS Lett 290:31-4.

21. Breitung, J., G. Borner, M. Karrasch, A. Berkessel, and R. K. Thauer. 1990.

N-furfurylformamide as a pseudo-substrate for formylmethanofuran converting

enzymes from methanogenic bacteria. FEBS Lett 268:257-60.

22. Breitung, J., G. Borner, S. Scholz, D. Linder, K. O. Stetter, and R. K. Thauer.

1992. Salt dependence, kinetic properties and catalytic mechanism of

N-formylmethanofuran: tetrahydromethanopterin formyltransferase from the

extreme thermophile Methanopyrus kandleri. Eur J Biochem 210:971-81.

23. Breitung, J., R. A. Schmitz, K. O. Stetter, and R. K. Thauer. 1991.

N5,N10-methenyltetrahydromethanopterin cyclohydrolase from the extreme

thermophile methanopyrus kandleri: increase of catalytic efficiency (KCat/KM) and

thermostability in the presence of salts. Arch Microbiol 156:517-524.

24. Breitung, J., and R. K. Thauer. 1990. Formylmethanofuran:

tetrahydromethanopterin formyltransferase from Methanosarcina barkeri:

identification of N5-formyltetrahydromethanopterin as the product. FEBS Lett

275:226-30. 40 25. Bryant, M. P., E. A. Wolin, M. J. Wolin, and R. S. Wolfe. 1967.

Methanobacillus omelianskii, a symbiotic association of two species of bacteria.

Arch Mikrobiol 59:20-31.

26. Buan, N. R., and W. W. Metcalf. 2009. Methanogenesis by Methanosarcina

acetivorans involves two structurally and functionally distinct classes of

heterodisulfide reductase. Mol Microbiol.

27. Burke, S. A., and J. A. Krzycki. 1995. Involvement of the "A" isozyme of

methyltransferase II and the 29-kilodalton corrinoid protein in methanogenesis

from monomethylamine. J Bacteriol 177:4410-6.

28. Burke, S. A., and J. A. Krzycki. 1997. Reconstitution of monomethylamine:

coenzyme M methyl transfer with a corrinoid protein and two methyltransferases

purified from Methanosarcina barkeri. J Biol Chem 272:16570-7.

29. Buss, K. A., C. Ingram-Smith, J. G. Ferry, D. A. Sanders, and M. S. Hasson.

1997. Crystallization of acetate kinase from Methanosarcina thermophila and

prediction of its fold. Protein Sci 6:2659-62.

30. Daas, P. J., K. A. Gerrits, J. T. Keltjens, C. van der Drift, and G. D. Vogels.

1993. Involvement of an activation protein in the methanol:

2-mercaptoethanesulfonic acid methyltransferase reaction in Methanosarcina

barkeri. J Bacteriol 175:1278-83.

31. Daas, P. J., W. R. Hagen, J. T. Keltjens, C. van der Drift, and G. D. Vogels.

1996. Activation mechanism of methanol: 5-hydroxybenzimidazolylcobamide

methyltransferase from Methanosarcina barkeri. J Biol Chem 271:22346-51. 41 32. Daniels, L., G. Fuchs, R. K. Thauer, and J. G. Zeikus. 1977. Carbon monoxide

oxidation by methanogenic bacteria. J Bacteriol 132:118-26.

33. DiMarco, A. A., T. A. Bobik, and R. S. Wolfe. 1990. Unusual coenzymes of

methanogenesis. Annu Rev Biochem 59:355-94.

34. DiMarco, A. A., M. I. Donnelly, and R. S. Wolfe. 1986. Purification and

properties of the 5,10-methenyltetrahydromethanopterin cyclohydrolase from

Methanobacterium thermoautotrophicum. J Bacteriol 168:1372-7.

35. DiMarco, A. A., K. A. Sment, J. Konisky, and R. S. Wolfe. 1990. The

formylmethanofuran: tetrahydromethanopterin formyltransferase from

Methanobacterium thermoautotrophicum ΔH: nucleotide sequence and functional

expression of the cloned gene. J Biol Chem 265:472-6.

36. Donnelly, M. I., and R. S. Wolfe. 1986. The role of formylmethanofuran:

tetrahydromethanopterin formyltransferase in methanogenesis from carbon

dioxide. J Biol Chem 261:16653-9.

37. Duin, E. C., C. Bauer, B. Jaun, and R. Hedderich. 2003. Coenzyme M binds to

a [4Fe-4S] cluster in the active site of heterodisulfide reductase as deduced from

EPR studies with the [33S]coenzyme M-treated enzyme. FEBS Lett 538:81-4.

38. Duin, E. C., S. Madadi-Kahkesh, R. Hedderich, M. D. Clay, and M. K.

Johnson. 2002. Heterodisulfide reductase from Methanothermobacter

marburgensis contains an active-site [4Fe-4S] cluster that is directly involved in

mediating heterodisulfide reduction. FEBS Lett 512:263-8.

39. Eggen, R. I., R. van Kranenburg, A. J. Vriesema, A. C. Geerling, M. F.

Verhagen, W. R. Hagen, and W. M. de Vos. 1996. Carbon monoxide 42 dehydrogenase from Methanosarcina frisia Go1: characterization of the enzyme

and the regulated expression of two operon-like cdh gene clusters. J Biol Chem

271:14256-63.

40. Eirich, L. D., G. D. Vogels, and R. S. Wolfe. 1978. Proposed structure for

coenzyme F420 from Methanobacterium. Biochemistry 17:4583-93.

41. Ellefson, W. L., W. B. Whitman, and R. S. Wolfe. 1982. Nickel-containing

factor F430: chromophore of the methylreductase of Methanobacterium. Proc Natl

Acad Sci U S A 79:3707-10.

42. Ellefson, W. L., and R. S. Wolfe. 1980. Role of component C in the

methylreductase system of Methanobacterium. J Biol Chem 255:8388-9.

43. Ellermann, J., R. Hedderich, R. Bocher, and R. K. Thauer. 1988. The final

step in methane formation: investigations with highly purified methyl-CoM

reductase (component C) from Methanobacterium thermoautotrophicum (strain

Marburg). Eur J Biochem 172:669-77.

44. Ellermann, J., S. Rospert, R. K. Thauer, M. Bokranz, A. Klein, M. Voges,

and A. Berkessel. 1989. Methyl-coenzyme-M reductase from Methanobacterium

thermoautotrophicum (strain Marburg): purity, activity and novel inhibitors. Eur J

Biochem 184:63-8.

45. Enssle, M., C. Zirngibl, D. Linder, and R. K. Thauer. 1991. Coenzyme F420

dependent N5, N10-methylenetetrahydromethanopterin dehydrogenase in methanol

grown Methanosarcina barkeri. Arch Microbiol 155:483-490. 43 46. Ermler, U., W. Grabarse, S. Shima, M. Goubeaud, and R. K. Thauer. 1997.

Crystal structure of methyl-coenzyme M reductase: the key enzyme of biological

methane formation. Science 278:1457-62.

47. Ermler, U., M. Merckel, R. Thauer, and S. Shima. 1997. Formylmethanofuran:

tetrahydromethanopterin formyltransferase from Methanopyrus kandleri - new

insights into salt-dependence and thermostability. Structure 5:635-46.

48. Ferguson, D. J., Jr., N. Gorlatova, D. A. Grahame, and J. A. Krzycki. 2000.

Reconstitution of dimethylamine: coenzyme M methyl transfer with a discrete

corrinoid protein and two methyltransferases purified from Methanosarcina

barkeri. J Biol Chem 275:29053-60.

49. Ferguson, D. J., Jr., and J. A. Krzycki. 1997. Reconstitution of

trimethylamine-dependent coenzyme M methylation with the trimethylamine

corrinoid protein and the isozymes of methyltransferase II from Methanosarcina

barkeri. J Bacteriol 179:846-52.

50. Ferguson, D. J., Jr., J. A. Krzycki, and D. A. Grahame. 1996. Specific roles of

methylcobamide: coenzyme M methyltransferase isozymes in metabolism of

methanol and methylamines in Methanosarcina barkeri. J Biol Chem

271:5189-94.

51. Ferguson, T., J. A. Soares, T. Lienard, G. Gottschalk, and J. A. Krzycki.

2009. RamA, a protein required for reductive activation of corrinoid-dependent

methylamine methyltransferase reactions in methanogenic archaea. J Biol Chem

284:2285-95. 44 52. Ferry, J. G., and K. A. Kastead. 2007. Methanogenesis. In R. Cavicchioli (ed.),

Archaea:Molecular and Cellular Biology. ASM Press, Washington, D.C.

53. Ferry, J. G., P. H. Smith, and R. S. Wolfe. 1974. Methanospirillum, a new

genus of methanogenic bacteria, and characterization of Methanospirillum

hungatii sp. nov. Int J Syst Bacteriol 24.

54. Fischer, R., P. Gartner, A. Yeliseev, and R. K. Thauer. 1992.

N5-methyltetrahydromethanopterin: coenzyme M methyltransferase in

methanogenic archaebacteria is a membrane protein. Arch Microbiol 158:208-17.

55. Fricke, W. F., H. Seedorf, A. Henne, M. Kruer, H. Liesegang, R. Hedderich,

G. Gottschalk, and R. K. Thauer. 2006. The genome sequence of

Methanosphaera stadtmanae reveals why this human intestinal archaeon is

restricted to methanol and H2 for methane formation and ATP synthesis. J

Bacteriol 188:642-58.

56. Galagan, J. E., C. Nusbaum, A. Roy, M. G. Endrizzi, P. Macdonald, W.

FitzHugh, S. Calvo, R. Engels, S. Smirnov, D. Atnoor, A. Brown, N. Allen, J.

Naylor, N. Stange-Thomann, K. DeArellano, R. Johnson, L. Linton, P.

McEwan, K. McKernan, J. Talamas, A. Tirrell, W. Ye, A. Zimmer, R. D.

Barber, I. Cann, D. E. Graham, D. A. Grahame, A. M. Guss, R. Hedderich, C.

Ingram-Smith, H. C. Kuettner, J. A. Krzycki, J. A. Leigh, W. Li, J. Liu, B.

Mukhopadhyay, J. N. Reeve, K. Smith, T. A. Springer, L. A. Umayam, O.

White, R. H. White, E. Conway de Macario, J. G. Ferry, K. F. Jarrell, H.

Jing, A. J. Macario, I. Paulsen, M. Pritchett, K. R. Sowers, R. V. Swanson, S.

H. Zinder, E. Lander, W. W. Metcalf, and B. Birren. 2002. The genome of M. 45 acetivorans reveals extensive metabolic and physiological diversity. Genome Res

12:532-42.

57. Gartner, P., A. Ecker, R. Fischer, D. Linder, G. Fuchs, and R. K. Thauer.

1993. Purification and properties of N5-methyltetrahydromethanopterin:

coenzyme M methyltransferase from Methanobacterium thermoautotrophicum.

Eur J Biochem 213:537-45.

58. Gartner, P., D. S. Weiss, U. Harms, and R. K. Thauer. 1994.

N5-methyltetrahydromethanopterin: coenzyme M methyltransferase from

Methanobacterium thermoautotrophicum: catalytic mechanism and sodium ion

dependence. Eur J Biochem 226:465-72.

59. Gencic, S., and D. A. Grahame. 2003. Nickel in subunit β of the acetyl-CoA

decarbonylase/synthase multienzyme complex in methanogens: catalytic

properties and evidence for a binuclear Ni-Ni site. J Biol Chem 278:6101-10.

60. Goubeaud, M., G. Schreiner, and R. K. Thauer. 1997. Purified

methyl-coenzyme-M reductase is activated when the enzyme-bound coenzyme

F430 is reduced to the nickel(I) oxidation state by titanium(III) citrate. Eur J

Biochem 243:110-4.

61. Grabarse, W., F. Mahlert, S. Shima, R. K. Thauer, and U. Ermler. 2000.

Comparison of three methyl-coenzyme M reductases from phylogenetically

distant organisms: unusual amino acid modification, conservation and adaptation.

J Mol Biol 303:329-44.

62. Grabarse, W., M. Vaupel, J. A. Vorholt, S. Shima, R. K. Thauer, A.

Wittershagen, G. Bourenkov, H. D. Bartunik, and U. Ermler. 1999. The 46 crystal structure of methenyltetrahydromethanopterin cyclohydrolase from the

hyperthermophilic archaeon Methanopyrus kandleri. Structure 7:1257-68.

63. Grahame, D. A. 1991. Catalysis of acetyl-CoA cleavage and

tetrahydrosarcinapterin methylation by a carbon monoxide

dehydrogenase-corrinoid enzyme complex. J Biol Chem 266:22227-33.

64. Grahame, D. A. 1989. Different isozymes of :

2-mercaptoethanesulfonate methyltransferase predominate in methanol- versus

acetate-grown Methanosarcina barkeri. J Biol Chem 264:12890-4.

65. Grahame, D. A. 1993. Substrate and cofactor reactivity of a carbon monoxide

dehydrogenase-corrinoid enzyme complex: stepwise reduction of iron-sulfur and

corrinoid centers, the corrinoid Co2+/1+ midpoint potential, and overall

synthesis of acetyl-CoA. Biochemistry 32:10786-93.

66. Grahame, D. A., and E. DeMoll. 1996. Partial reactions catalyzed by protein

components of the acetyl-CoA decarbonylase synthase enzyme complex from

Methanosarcina barkeri. J Biol Chem 271:8352-8.

67. Grahame, D. A., and T. C. Stadtman. 1987. Carbon monoxide dehydrogenase

from Methanosarcina barkeri: disaggregation, purification, and physicochemical

properties of the enzyme. J Biol Chem 262:3706-12.

68. Guss, A. M., B. Mukhopadhyay, J. K. Zhang, and W. W. Metcalf. 2005.

Genetic analysis of mch mutants in two Methanosarcina species demonstrates

multiple roles for the methanopterin-dependent C-1 oxidation/reduction pathway

and differences in H2 metabolism between closely related species. Mol Microbiol

55:1671-80. 47 69. Hagemeier, C. H., M. Krer, R. K. Thauer, E. Warkentin, and U. Ermler.

2006. Insight into the mechanism of biological methanol activation based on the

crystal structure of the methanol-cobalamin methyltransferase complex. Proc Natl

Acad Sci U S A 103:18917-22.

70. Hagemeier, C. H., S. Shima, R. K. Thauer, G. Bourenkov, H. D. Bartunik,

and U. Ermler. 2003. Coenzyme F420-dependent

methylenetetrahydromethanopterin dehydrogenase (Mtd) from Methanopyrus

kandleri: a methanogenic enzyme with an unusual quarternary structure. J Mol

Biol 332:1047-57.

71. Hagemeier, C. H., S. Shima, E. Warkentin, R. K. Thauer, and U. Ermler.

2003. Coenzyme F420-dependent methylenetetrahydromethanopterin

dehydrogenase from Methanopyrus kandleri: the selenomethionine-labelled and

non-labelled enzyme crystallized in two different forms. Acta Crystallogr D Biol

Crystallogr 59:1653-5.

72. Hamann, N., G. J. Mander, J. E. Shokes, R. A. Scott, M. Bennati, and R.

Hedderich. 2007. A cysteine-rich CCG domain contains a novel [4Fe-4S] cluster

binding motif as deduced from studies with subunit B of heterodisulfide reductase

from Methanothermobacter marburgensis. Biochemistry 46:12875-85.

73. Hanson, R. S., and T. E. Hanson. 1996. Methanotrophic bacteria. Microbiol Rev

60:439-71.

74. Harms, U., and R. K. Thauer. 1996. The corrinoid-containing 23-kDa subunit

MtrA of the energy-conserving N5-methyltetrahydromethanopterin: coenzyme M

methyltransferase complex from Methanobacterium thermoautotrophicum: EPR 48 spectroscopic evidence for a histidine residue as a cobalt ligand of the cobamide.

Eur J Biochem 241:149-54.

75. Harms, U., and R. K. Thauer. 1997. Identification of the active site histidine in

the corrinoid protein MtrA of the energy-conserving methyltransferase complex

from Methanobacterium thermoautotrophicum. Eur J Biochem 250:783-8.

76. Harms, U., and R. K. Thauer. 1996. Methylcobalamin: coenzyme M

methyltransferase isoenzymes MtaA and MtbA from Methanosarcina barkeri:

cloning, sequencing and differential transcription of the encoding genes, and

functional overexpression of the mtaA gene in Escherichia coli. Eur J Biochem

235:653-9.

77. Harms, U., D. S. Weiss, P. Gartner, D. Linder, and R. K. Thauer. 1995. The

energy conserving N5-methyltetrahydromethanopterin:coenzyme M

methyltransferase complex from Methanobacterium thermoautotrophicum is

composed of eight different subunits. Eur J Biochem 228:640-8.

78. Hartmann, G. C., A. R. Klein, M. Linder, and R. K. Thauer. 1996.

5 10 Purification, properties and primary structure of H2-forming N ,N

-methylenetetrahydromethanopterin dehydrogenase from Methanococcus

thermolithotrophicus. Arch Microbiol 165:187-93.

79. Hartzell, P. L., and R. S. Wolfe. 1986. Comparative studies of component C

from the methylreductase system of different methanogens. Syst Appl Microbiol

7:376-382.

80. Hartzell, P. L., G. Zvilius, J. C. Escalante-Semerena, and M. I. Donnelly.

1985. Coenzyme F420 dependence of the methylenetetrahydromethanopterin 49 dehydrogenase of Methanobacterium thermoautotrophicum. Biochem Biophys

Res Commun 133:884-90.

81. Hedderich, R., A. Berkessel, and R. K. Thauer. 1990. Purification and

properties of heterodisulfide reductase from Methanobacterium

thermoautotrophicum (strain Marburg). Eur J Biochem 193:255-61.

82. Hedderich, R., J. Koch, D. Linder, and R. K. Thauer. 1994. The

heterodisulfide reductase from Methanobacterium thermoautotrophicum contains

sequence motifs characteristic of pyridine-nucleotide-dependent thioredoxin

reductases. Eur J Biochem 225:253-61.

83. Heiden, S., R. Hedderich, E. Setzke, and R. K. Thauer. 1994. Purification of a

two-subunit cytochrome-b-containing heterodisulfide reductase from

methanol-grown Methanosarcina barkeri. Eur J Biochem 221:855-61.

84. Hendrickson, E. L., A. K. Haydock, B. C. Moore, W. B. Whitman, and J. A.

Leigh. 2007. Functionally distinct genes regulated by hydrogen limitation and

growth rate in methanogenic Archaea. Proc Natl Acad Sci U S A 104:8930-4.

85. Hippler, B., and R. K. Thauer. 1999. The energy conserving

methyltetrahydromethanopterin: coenzyme M methyltransferase complex from

methanogenic archaea: function of the subunit MtrH. FEBS Lett 449:165-8.

86. Hirata, A., and K. S. Murakami. 2009. Archaeal RNA polymerase. Curr Opin

Struct Biol 19:724-31.

87. Hochheimer, A., D. Linder, R. K. Thauer, and R. Hedderich. 1996. The

molybdenum formylmethanofuran dehydrogenase operon and the tungsten

formylmethanofuran dehydrogenase operon from Methanobacterium 50 thermoautotrophicum: structures and transcriptional regulation. Eur J Biochem

242:156-62.

88. Hochheimer, A., R. A. Schmitz, R. K. Thauer, and R. Hedderich. 1995. The

tungsten formylmethanofuran dehydrogenase from Methanobacterium

thermoautotrophicum contains sequence motifs characteristic for enzymes

containing molybdopterin dinucleotide. Eur J Biochem 234:910-20.

89. Huber, H., M. Thomm, H. Konig, G. Thies, and K. O. Stetter. 1982.

Methanococcus thermolithotrophicus, a novel thermophilic lithotrophic

methanogen. Arch Microbiol 132:47-50.

90. Huber, R., M. Kurr, H. W. Jannasch, and K. O. Stetter. 1989. A novel group

of abyssal methanogenic archaebacteria (Methanopyrus) growing at 110 °C.

Nature 342:833-834.

91. Ide, T., S. Baumer, and U. Deppenmeier. 1999. Energy conservation by the H2:

heterodisulfide oxidoreductase from Methanosarcina mazei Go1: identification of

two proton-translocating segments. J Bacteriol 181:4076-80.

92. Iyer, P. P., S. H. Lawrence, K. B. Luther, K. R. Rajashankar, H. P.

Yennawar, J. G. Ferry, and H. Schindelin. 2004. Crystal structure of

phosphotransacetylase from the methanogenic archaeon Methanosarcina

thermophila. Structure 12:559-67.

93. Iyer, P. P., S. H. Lawrence, H. P. Yennawar, and J. G. Ferry. 2003.

Expression, purification, crystallization and preliminary X-ray analysis of

phosphotransacetylase from Methanosarcina thermophila. Acta Crystallogr D

Biol Crystallogr 59:1517-20. 51 94. Jablonski, P. E., A. A. DiMarco, T. A. Bobik, M. C. Cabell, and J. G. Ferry.

1990. Protein content and enzyme activities in methanol- and acetate-grown

Methanosarcina thermophila. J Bacteriol 172:1271-5.

95. Jablonski, P. E., and J. G. Ferry. 1991. Purification and properties of methyl

coenzyme M methylreductase from acetate-grown Methanosarcina thermophila. J

Bacteriol 173:2481-7.

96. Jablonski, P. E., W. P. Lu, S. W. Ragsdale, and J. G. Ferry. 1993.

Characterization of the metal centers of the corrinoid/iron-sulfur component of the

CO dehydrogenase enzyme complex from Methanosarcina thermophila by EPR

spectroscopy and spectroelectrochemistry. J Biol Chem 268:325-9.

97. Kamagata, Y., H. Kawasaki, H. Oyaizu, K. Nakamura, E. Mikami, G. Endo,

Y. Koga, and K. Yamasato. 1992. Characterization of three thermophilic strains

of ("Methanosaeta") thermophila sp. nov. and rejection of

Methanothrix ("Methanosaeta") thermoacetophila. Int J Syst Bacteriol 42:463-8.

98. Karrasch, M., G. Borner, M. Enssle, and R. K. Thauer. 1989.

Formylmethanofuran dehydrogenase from methanogenic bacteria, a

molybdoenzyme. FEBS Lett 253:226-30.

99. Karrasch, M., G. Borner, and R. K. Thauer. 1990. The molybdenum cofactor

of formylmethanofuran dehydrogenase from Methanosarcina barkeri is a

molybdopterin guanine dinucleotide. FEBS Lett 274:48-52.

100. Keltjens, J. T., and G. D. Vogels. 1993. Conversion of methanol and

methylamines to methane and . In J. G. Ferry (ed.), 52 Methanogenesis: Ecology, Physiology, Biochemistry & Genetics. Chapman &

Hall, New York.

101. Kenealy, W. R., and J. G. Zeikus. 1982. One-carbon metabolism in

methanogens: evidence for synthesis of a two-carbon cellular intermediate and

unification of catabolism and anabolism in Methanosarcina barkeri. J Bacteriol

151:932-41.

102. Kengen, S. W., P. J. Daas, E. F. Duits, J. T. Keltjens, C. van der Drift, and G.

D. Vogels. 1992. Isolation of a 5-hydroxybenzimidazolyl cobamide-containing

enzyme involved in the methyltetrahydromethanopterin: coenzyme M

methyltransferase reaction in Methanobacterium thermoautotrophicum. Biochim

Biophys Acta 1118:249-60.

103. Klein, A. R., J. Koch, K. O. Stetter, and R. K. Thauer. 1993. Two

N5,N10-methylenetetrahydromethanopterin dehydrogenases in the extreme

thermophile Methanopyrus kandleri: characterization of the coenzyme

F420-dependent enzyme. Arch Microbiol 160:186-92.

104. Kletzin, A. 2007. General characteristics and important model organisms. In R.

Cavicchioli (ed.), Archaea: Molecular and Cellular Biology. ASM Press,

Washington, D. C. .

105. Krzycki, J. A., L. E. Mortenson, and R. C. Prince. 1989. Paramagnetic centers

of carbon monoxide dehydrogenase from aceticlastic Methanosarcina barkeri. J

Biol Chem 264:7217-21. 53 106. Krzycki, J. A., R. H. Wolkin, and J. G. Zeikus. 1982. Comparison of unitrophic

and mixotrophic substrate metabolism by acetate-adapted strain of

Methanosarcina barkeri. J Bacteriol 149:247-54.

107. Krzycki, J. A., and J. G. Zeikus. 1984. Characterization and purification of

carbon monoxide dehydrogenase from Methanosarcina barkeri. J Bacteriol

158:231-7.

108. Kunkel, A., M. Vaupel, S. Heim, R. K. Thauer, and R. Hedderich. 1997.

Heterodisulfide reductase from methanol-grown cells of Methanosarcina barkeri

is not a flavoenzyme. Eur J Biochem 244:226-34.

109. Kunow, J., S. Shima, J. A. Vorholt, and R. K. Thauer. 1996. Primary structure

and properties of the formyltransferase from the mesophilic Methanosarcina

barkeri: comparison with the enzymes from thermophilic and hyperthermophilic

methanogens. Arch Microbiol 165:97-105.

110. Kurr, M., R. Huber, H. Konig, H. W. Jannasch, H. Fricke, A. Trincone, J. K.

Kristjansson, and K. O. Stetter. 1991. Methanopyrus kandleri, gen. and sp. nov.

represents a novel group of hyperthermophilic methanogens, Growing at 110 °C.

Arch Microbiol 156:239-247.

111. Lange, S., and G. Fuchs. 1987. Autotrophic synthesis of activated acetic acid

from CO2 in Methanobacterium thermoautotrophicum: synthesis from

tetrahydromethanopterin-bound C1 units and carbon monoxide. Eur J Biochem

163:147-54. 54 112. Latimer, M. T., and J. G. Ferry. 1993. Cloning, sequence analysis, and

hyperexpression of the genes encoding phosphotransacetylase and acetate kinase

from Methanosarcina thermophila. J Bacteriol 175:6822-9.

113. LeClerc, G. M., and D. A. Grahame. 1996. Methylcobamide: coenzyme M

methyltransferase isozymes from Methanosarcina barkeri: physicochemical

characterization, cloning, sequence analysis, and heterologous gene expression. J

Biol Chem 271:18725-31.

114. Lehmacher, A. 1994. Cloning, sequencing and transcript analysis of the gene

encoding formylmethanofuran: tetrahydromethanopterin formyltransferase from

the hyperthermophilic Methanothermus fervidus. Mol Gen Genet 242:73-80.

115. Leigh, J. A., K. L. Rinehart, Jr., and R. S. Wolfe. 1985. Methanofuran (carbon

dioxide reduction factor), a formyl carrier in methane production from carbon

dioxide in Methanobacterium. Biochemistry 24:995-9.

116. Lessner, D. J., L. Li, Q. Li, T. Rejtar, V. P. Andreev, M. Reichlen, K. Hill, J.

J. Moran, B. L. Karger, and J. G. Ferry. 2006. An unconventional pathway for

reduction of CO2 to methane in CO-grown Methanosarcina acetivorans revealed

by proteomics. Proc Natl Acad Sci U S A 103:17921-6.

117. Li, Q., L. Li, T. Rejtar, D. J. Lessner, B. L. Karger, and J. G. Ferry. 2006.

Electron transport in the pathway of acetate conversion to methane in the marine

archaeon Methanosarcina acetivorans. J Bacteriol 188:702-10.

118. Lienard, T., B. Becher, M. Marschall, S. Bowien, and G. Gottschalk. 1996.

Sodium ion translocation by N5-methyltetrahydromethanopterin: coenzyme M 55 methyltransferase from Methanosarcina mazei Go1 reconstituted in ether lipid

liposomes. Eur J Biochem 239:857-64.

119. Lienard, T., and G. Gottschalk. 1998. Cloning, sequencing and expression of

the genes encoding the sodium translocating N5-methyltetrahydromethanopterin:

coenzyme M methyltransferase of the methylotrophic archaeon Methanosarcina

mazei Go1. FEBS Lett 425:204-8.

120. Lu, W. P., B. Becher, G. Gottschalk, and S. W. Ragsdale. 1995. Electron

paramagnetic resonance spectroscopic and electrochemical characterization of the

partially purified N5-methyltetrahydromethanopterin: coenzyme M

methyltransferase from Methanosarcina mazei Go1. J Bacteriol 177:2245-50.

121. Lundie, L. L., Jr., and J. G. Ferry. 1989. Activation of acetate by

Methanosarcina thermophila: purification and characterization of

phosphotransacetylase. J Biol Chem 264:18392-6.

122. Ma, K., D. Linder, K. O. Stetter, and R. K. Thauer. 1991. Purification and

properties of N5,N10-methylenetetrahydromethanopterin reductase (coenzyme

F420-dependent) from the extreme thermophile Methanopyrus kandleri. Arch

Microbiol 155:593-600.

123. Ma, K., and R. K. Thauer. 1990. Purification and properties of N5,

N10-methylenetetrahydromethanopterin reductase from Methanobacterium

thermoautotrophicum (strain Marburg). Eur J Biochem 191:187-93.

124. Ma, K., and R. K. Thauer. 1990. Single step purification of

methylenetetrahydromethanopterin reductase from Methanobacterium 56 thermoautotrophicum by specific binding to blue sepharose CL-6B. FEBS Lett

268:59-62.

125. Ma, K., C. Zirngibl, D. Linder, K. O. Stetter, and R. K. Thauer. 1991. N5,

10 N -methylenetetrahydromethanopterin dehydrogenase (H2-forming) from the

extreme thermophile Methanopyrus kandleri. Arch Microbiol 156:43-8.

126. Madadi-Kahkesh, S., E. C. Duin, S. Heim, S. P. Albracht, M. K. Johnson,

and R. Hedderich. 2001. A paramagnetic species with unique EPR

characteristics in the active site of heterodisulfide reductase from methanogenic

archaea. Eur J Biochem 268:2566-77.

127. Mamat, B., A. Roth, C. Grimm, U. Ermler, C. Tziatzios, D. Schubert, R. K.

Thauer, and S. Shima. 2002. Crystal structures and enzymatic properties of three

formyltransferases from archaea: environmental adaptation and evolutionary

relationship. Protein Sci 11:2168-78.

128. Morgan, R. M., T. D. Pihl, J. Nolling, and J. N. Reeve. 1997. Hydrogen

regulation of growth, growth yields, and methane gene transcription in

Methanobacterium thermoautotrophicum ΔH. J Bacteriol 179:889-98.

129. Mukhopadhyay, B., and L. Daniels. 1989. Aerobic purification of

N5,N10-methylenetetrahydromethanopterin dehydrogenase, separated from

N5,N10-methylenetetrahydromethanopterin cyclohydrolase, from

Methanobacterium thermoautotrophicum strain Marburg. Can J Microbiol

35:499-507.

130. Murakami, E., U. Deppenmeier, and S. W. Ragsdale. 2001. Characterization of

the intramolecular electron transfer pathway from 2-hydroxyphenazine to the 57 heterodisulfide reductase from Methanosarcina thermophila. J Biol Chem

276:2432-9.

131. Murakami, E., and S. W. Ragsdale. 2000. Evidence for intersubunit

communication during acetyl-CoA cleavage by the multienzyme CO

dehydrogenase/acetyl-CoA synthase complex from Methanosarcina thermophila:

evidence that the β subunit catalyzes C-C and C-S bond cleavage. J Biol Chem

275:4699-707.

132. Ni, S., C. R. Woese, H. C. Aldrich, and D. R. Boone. 1994. Transfer of

Methanolobus siciliae to the genus Methanosarcina, naming it Methanosarcina

siciliae, and emendation of the genus Methanosarcina. Int J Syst Bacteriol

44:357-9.

133. Noll, K. M., K. L. Rinehart, Jr., R. S. Tanner, and R. S. Wolfe. 1986.

Structure of component B (7-mercaptoheptanoylthreonine phosphate) of the

methylcoenzyme M methylreductase system of Methanobacterium

thermoautotrophicum. Proc Natl Acad Sci U S A 83:4238-42.

134. Nolling, J., T. D. Pihl, A. Vriesema, and J. N. Reeve. 1995. Organization and

growth phase-dependent transcription of methane genes in two regions of the

Methanobacterium thermoautotrophicum genome. J Bacteriol 177:2460-8.

135. O'Brien, J. M., R. H. Wolkin, T. T. Moench, J. B. Morgan, and J. G. Zeikus.

1984. Association of hydrogen metabolism with unitrophic or mixotrophic growth

of Methanosarcina barkeri on carbon monoxide. J Bacteriol 158:373-5. 58 136. Olson, K. D., C. W. McMahon, and R. S. Wolfe. 1991. Photoactivation of the

2-(methylthio)ethanesulfonic acid reductase from Methanobacterium. Proc Natl

Acad Sci U S A 88:4099-103.

137. Paoloni, C. 1976. Storia del Metano. Tipografia SAPIL, Milano.

138. Patel, G. B., L. A. Roth, L. van den Berg, and D. S. Clark. 1976.

Characterization of a strain of Methanospirillum hungatti. Can J Microbiol

22:1404-10.

139. Pihl, T. D., S. Sharma, and J. N. Reeve. 1994. Growth phase-dependent

transcription of the genes that encode the two methyl coenzyme M reductase

isoenzymes and N5-methyltetrahydromethanopterin:coenzyme M

methyltransferase in Methanobacterium thermoautotrophicum ΔH. J Bacteriol

176:6384-91.

140. Poirot, C. M., S. W. M. Kengen, E. Valk, J. T. Keltjens, C. Vanderdrift, and

G. D. Vogels. 1987. Formation of methylcoenzyme M from formaldehyde by

cell-free extracts of Methanobacterium thermoautotrophicum: evidence for the

Involvement of a corrinoid-containing methyltransferase. FEMS Microbiol Lett

40:7-13.

141. Reichlen, M. J., K. S. Murakami, and J. G. Ferry. Functional analysis of the

three TATA binding protein homologs in Methanosarcina acetivorans. J

Bacteriol 192:1511-7.

142. Rospert, S., R. Bocher, S. P. Albracht, and R. K. Thauer. 1991.

Methyl-coenzyme M reductase preparations with high specific activity from 59

H2-preincubated cells of Methanobacterium thermoautotrophicum. FEBS Lett

291:371-5.

143. Rospert, S., J. Breitung, K. Ma, B. Schworer, C. Zirngibl, R. K. Thauer, D.

Linder, R. Huber, and K. O. Stetter. 1991. Methyl-coenzyme M reductase and

other enzymes involved in methanogenesis from CO2 and H2 in the extreme

thermophile Methanopyrus kandleri. Arch Microbiol 156:49-55.

144. Rospert, S., D. Linder, J. Ellermann, and R. K. Thauer. 1990. Two genetically

distinct methyl-coenzyme M reductases in Methanobacterium

thermoautotrophicum strain Marburg and ΔH. Eur J Biochem 194:871-7.

145. Rouviere, P. E., T. A. Bobik, and R. S. Wolfe. 1988. Reductive activation of the

methyl coenzyme M methylreductase system of Methanobacterium

thermoautotrophicum ΔH. J Bacteriol 170:3946-52.

146. Sauer, K., U. Harms, and R. K. Thauer. 1997. Methanol: coenzyme M

methyltransferase from Methanosarcina barkeri: purification, properties and

encoding genes of the corrinoid protein MT1. Eur J Biochem 243:670-7.

147. Sauer, K., and R. K. Thauer. 1998. His84 rather than His35 is the active site

histidine in the corrinoid protein MtrA of the energy conserving methyltransferase

complex from Methanobacterium thermoautotrophicum. FEBS Lett 436:401-402.

148. Sauer, K., and R. K. Thauer. 1998. Methanol: coenzyme M methyltransferase

from Methanosarcina barkeri: identification of the active-site histidine in the

corrinoid-harboring subunit MtaC by site-directed mutagenesis. Eur J Biochem

253:698-705. 60 149. Sauer, K., and R. K. Thauer. 1997. Methanol: coenzyme M methyltransferase

from Methanosarcina barkeri: zinc dependence and thermodynamics of the

methanol: cob(I)alamin methyltransferase reaction. Eur J Biochem 249:280-5.

150. Schauer, N. L., and J. G. Ferry. 1980. Metabolism of formate in

Methanobacterium formicicum. J Bacteriol 142:800-7.

151. Schmitz, R. A., S. P. Albracht, and R. K. Thauer. 1992. A molybdenum and a

tungsten isoenzyme of formylmethanofuran dehydrogenase in the thermophilic

archaeon Methanobacterium wolfei. Eur J Biochem 209:1013-8.

152. Schmitz, R. A., M. Richter, D. Linder, and R. K. Thauer. 1992. A

tungsten-containing active formylmethanofuran dehydrogenase in the

thermophilic archaeon Methanobacterium wolfei. Eur J Biochem 207:559-65.

153. Schnellen, C. G. T. P. 1947. Onderzoekingen over der methaangisting. Technical

University, Delft.

154. Schulz, H., S. P. J. Albracht, J. M. C. C. Coremans, and G. Fuchs. 1988.

Purification and some properties of the corrinoid-containing membrane protein

from Methanobacterium thermoautotrophicum. Eur J Biochem 171:589-597.

155. Schulz, H., and G. Fuchs. 1986. Cobamide-containing membrane protein

complex in Methanobacterium. FEBS Lett 198:279-282.

156. Setzke, E., R. Hedderich, S. Heiden, and R. K. Thauer. 1994. H2:

heterodisulfide oxidoreductase complex from Methanobacterium

thermoautotrophicum: composition and properties. Eur J Biochem 220:139-48. 61 157. Sherf, B. A., and J. N. Reeve. 1990. Identification of the mcrD gene product and

its association with component C of methyl coenzyme M reductase in

Methanococcus vannielii. J Bacteriol 172:1828-33.

158. Shieh, J., and W. B. Whitman. 1988. Autotrophic acetyl coenzyme A

biosynthesis in Methanococcus maripaludis. J Bacteriol 170:3072-9.

159. Shima, S., M. Goubeaud, D. Vinzenz, R. K. Thauer, and U. Ermler. 1997.

Crystallization and preliminary X-ray diffraction studies of methyl-coenzyme M

reductase from Methanobacterium thermoautotrophicum. J Biochem 121:829-30.

160. Shima, S., and R. K. Thauer. 2005. Methyl-coenzyme M reductase and the

anaerobic oxidation of methane in methanotrophic Archaea. Curr Opin Microbiol

8:643-8.

161. Shima, S., R. K. Thauer, H. Michel, and U. Ermler. 1996. Crystallization and

preliminary X-ray diffraction studies of formylmethanofuran:

tetrahydromethanopterin formyltransferase from Methanopyrus kandleri. Proteins

26:118-20.

162. Shima, S., C. Tziatzios, D. Schubert, H. Fukada, K. Takahashi, U. Ermler,

and R. K. Thauer. 1998. Lyotropic-salt-induced changes in

monomer/dimer/tetramer association equilibrium of formyltransferase from the

hyperthermophilic Methanopyrus kandleri in relation to the activity and

thermostability of the enzyme. Eur J Biochem 258:85-92.

163. Shima, S., E. Warkentin, W. Grabarse, M. Sordel, M. Wicke, R. K. Thauer,

and U. Ermler. 2000. Structure of coenzyme F420 dependent 62 methylenetetrahydromethanopterin reductase from two methanogenic archaea. J

Mol Biol 300:935-50.

164. Shima, S., D. S. Weiss, and R. K. Thauer. 1995. Formylmethanofuran:

tetrahydromethanopterin formyltransferase (Ftr) from the hyperthermophilic

Methanopyrus kandleri: cloning, sequencing and functional expression of the ftr

gene and one-step purification of the enzyme overproduced in Escherichia coli.

Eur J Biochem 230:906-13.

165. Simianu, M., E. Murakami, J. M. Brewer, and S. W. Ragsdale. 1998.

Purification and properties of the - and iron-sulfur-containing heterodisulfide

reductase from Methanosarcina thermophila. Biochemistry 37:10027-39.

166. Singh-Wissmann, K., and J. G. Ferry. 1995. Transcriptional regulation of the

phosphotransacetylase-encoding and acetate kinase-encoding genes (pta and ack)

from Methanosarcina thermophila. J Bacteriol 177:1699-702.

167. Smith, K. S., and C. Ingram-Smith. 2007. Methanosaeta, the forgotten

methanogen? Trends Microbiol 15:150-5.

168. Sowers, K. R., S. F. Baron, and J. G. Ferry. 1984. Methanosarcina acetivorans

sp. nov., an acetotrophic methane-producing bacterium isolated from marine

sediments. Appl Environ Microbiol 47:971-8.

169. Sowers, K. R., T. T. Thai, and R. P. Gunsalus. 1993. Transcriptional regulation

of the carbon monoxide dehydrogenase gene (cdhA) in Methanosarcina

thermophila. J Biol Chem 268:23172-8. 63 170. Sprott, G. D., and T. J. Beveridge. 1993. Microscopy. In J. G. Ferry (ed.),

Methanogenesis: Ecology, Physiology, Biochemistry & Genetics. Chapman &

Hall, New York.

171. Stojanowic, A., G. J. Mander, E. C. Duin, and R. Hedderich. 2003.

Physiological role of the F420-non-reducing hydrogenase (Mvh) from

Methanothermobacter marburgensis. Arch Microbiol 180:194-203.

172. Stroup, D., and J. N. Reeve. 1993. Association of the mcrD gene product with

methyl coenzyme M reductase in Methanococcus vannielii. Biochim Biophys

Acta 1203:175-83.

173. Stroup, D., and J. N. Reeve. 1993. Identification of the mcrC gene product in

Methanococcus vannielii. FEMS Microbiol Lett 111:129-34.

174. Stupperich, E., K. E. Hammel, G. Fuchs, and R. K. Thauer. 1983. Carbon

monoxide fixation into the carboxyl group of acetyl coenzyme A during

autotrophic growth of Methanobacterium. FEBS Lett 152:21-3.

175. Stupperich, E., A. Juza, M. Hoppert, and F. Mayer. 1993. Cloning, sequencing

and immunological characterization of the corrinoid-containing subunit of the

N5-methyltetrahydromethanopterin: coenzyme-M methyltransferase from

Methanobacterium thermoautotrophicum. Eur J Biochem 217:115-21.

176. Tallant, T. C., L. Paul, and J. A. Krzycki. 2001. The MtsA subunit of the

methylthiol: coenzyme M methyltransferase of Methanosarcina barkeri catalyses

both half-reactions of corrinoid-dependent dimethylsulfide: coenzyme M methyl

transfer. J Biol Chem 276:4485-93. 64 177. te Brommelstroet, B. W., W. J. Geerts, J. T. Keltjens, C. van der Drift, and G.

D. Vogels. 1991. Purification and properties of

5,10-methylenetetrahydromethanopterin dehydrogenase and

5,10-methylenetetrahydromethanopterin reductase, two coenzyme F420-dependent

enzymes, from Methanosarcina barkeri. Biochim Biophys Acta 1079:293-302.

178. te Brommelstroet, B. W., C. M. Hensgens, W. J. Geerts, J. T. Keltjens, C. van

der Drift, and G. D. Vogels. 1990. Purification and properties of

5,10-methenyltetrahydromethanopterin cyclohydrolase from Methanosarcina

barkeri. J Bacteriol 172:564-71.

179. te Brommelstroet, B. W., C. M. Hensgens, J. T. Keltjens, C. van der Drift,

and G. D. Vogels. 1991. Purification and characterization of coenzyme

F420-dependent 5,10-methylenetetrahydromethanopterin dehydrogenase from

Methanobacterium thermoautotrophicum strain ΔH. Biochim Biophys Acta

1073:77-84.

180. te Brommelstroet, B. W., C. M. Hensgens, J. T. Keltjens, C. van der Drift,

and G. D. Vogels. 1990. Purification and properties of

5,10-methylenetetrahydromethanopterin reductase, a coenzyme F420-dependent

enzyme, from Methanobacterium thermoautotrophicum strain ΔH. J Biol Chem

265:1852-7.

181. Terlesky, K. C., M. J. Barber, D. J. Aceti, and J. G. Ferry. 1987. EPR

properties of the Ni-Fe-C center in an enzyme complex with carbon monoxide

dehydrogenase activity from acetate-grown Methanosarcina thermophila:

evidence that acetyl-CoA is a physiological substrate. J Biol Chem 262:15392-5. 65 182. Terlesky, K. C., and J. G. Ferry. 1988. Ferredoxin requirement for electron

transport from the carbon monoxide dehydrogenase complex to a

membrane-bound hydrogenase in acetate-grown Methanosarcina thermophila. J

Biol Chem 263:4075-9.

183. Terlesky, K. C., M. J. Nelson, and J. G. Ferry. 1986. Isolation of an enzyme

complex with carbon monoxide dehydrogenase activity containing corrinoid and

nickel from acetate-grown Methanosarcina thermophila. J Bacteriol 168:1053-8.

184. Thauer, R., Hedderich R., Fischer R. 1993. Reactions and Enzymes involved in

methanogenesis from CO2 and H2. In J. G. Ferry (ed.), Methanogenesis: Ecology,

Physiology, Biochemistry & Genetics Chapman & Hall, New York.

185. Thauer, R. K. 1998. Biochemistry of methanogenesis: a tribute to Marjory

Stephenson. 1998 Marjory Stephenson Prize Lecture. Microbiology 144 ( Pt

9):2377-406.

186. Thauer, R. K., A. K. Kaster, M. Goenrich, M. Schick, T. Hiromoto, and S.

Shima. 2010. Hydrogenases from methanogenic archaea, nickel, a novel cofactor,

and H2 storage. Annu Rev Biochem 79:507-36.

187. Tonouchi, A., Y. Nishizaki, H. Tohyama, and K. Takeda. 2002. Cloning of a

gene encoding acetate kinase from Methanosarcina mazei 2-P isolated from a

Japanese paddy field soil. Curr Microbiol 45:390-3.

188. Tyler, S. C. 1991. The global methane budget. In J. E. Rogers and W. B.

Whitman (ed.), Microbial Production and Consumption of Greenhouse Gases:

Methane, Nitrogen Oxide, and Halomethanes. American Society for

Microbiology, Washington, D. C. 66 189. van der Meijden, P., H. J. Heythuysen, H. T. Sliepenbeek, F. P. Houwen, C.

van der Drift, and G. D. Vogels. 1983. Activation and inactivation of methanol:

2-mercaptoethanesulfonic acid methyltransferase from Methanosarcina barkeri. J

Bacteriol 153:6-11.

190. von Bunau, R., C. Zirngibl, R. K. Thauer, and A. Klein. 1991.

Hydrogen-forming and coenzyme-F420-reducing methylene

tetrahydromethanopterin dehydrogenase are genetically distinct enzymes in

Methanobacterium thermoautotrophicum (Marburg). Eur J Biochem 202:1205-8.

191. Vorholt, J. A., and R. K. Thauer. 1997. The active species of 'CO2' utilized by

formylmethanofuran dehydrogenase from methanogenic Archaea. Eur J Biochem

248:919-24.

192. Vorholt, J. A., M. Vaupel, and R. K. Thauer. 1996. A polyferredoxin with

eight [4Fe-4S] clusters as a subunit of molybdenum formylmethanofuran

dehydrogenase from Methanosarcina barkeri. Eur J Biochem 236:309-17.

193. Vorholt, J. A., M. Vaupel, and R. K. Thauer. 1997. A selenium-dependent and

a selenium-independent formylmethanofuran dehydrogenase and their

transcriptional regulation in the hyperthermophilic Methanopyrus kandleri. Mol

Microbiol 23:1033-42.

194. Wang, M., Suharti, J. F. Tomb, and J. G. Ferry. 2010. The electron transport

chain of acetate-grown Methanosarcina acetivorans. submitted.

195. Warkentin, E., C. H. Hagemeier, S. Shima, R. K. Thauer, and U. Ermler.

2005. The structure of F420-dependent methylenetetrahydromethanopterin

dehydrogenase: a crystallographic 'superstructure' of the 67 selenomethionine-labelled protein crystal structure. Acta Crystallogr D Biol

Crystallogr 61:198-202.

196. Wassenaar, R. W., P. J. Daas, W. J. Geerts, J. T. Keltjens, and C. van der

Drift. 1996. Involvement of methyltransferase-activating protein and

methyltransferase 2 isoenzyme II in methylamine: coenzyme M methyltransferase

reactions in Methanosarcina barkeri Fusaro. J Bacteriol 178:6937-44.

197. Wassenaar, R. W., J. T. Keltjens, C. van der Drift, and G. D. Vogels. 1998.

Purification and characterization of dimethylamine:

5-hydroxybenzimidazolyl-cobamide methyltransferase from Methanosarcina

barkeri Fusaro. Eur J Biochem 253:692-7.

198. Weil, C. F., D. S. Cram, B. A. Sherf, and J. N. Reeve. 1988. Structure and

comparative analysis of the genes encoding component C of methyl coenzyme M

reductase in the extremely thermophilic archaebacterium Methanothermus

fervidus. J Bacteriol 170:4718-26.

199. Weiss, D. S., P. Gartner, and R. K. Thauer. 1994. The energetics and

sodium-ion dependence of N5-methyltetrahydromethanopterin:coenzyme M

methyltransferase studied with cob(I)alamin as methyl acceptor and

methylcob(III)alamin as methyl donor. Eur J Biochem 226:799-809.

200. Welander, P. V., and W. W. Metcalf. 2005. Loss of the mtr operon in

Methanosarcina blocks growth on methanol, but not methanogenesis, and reveals

an unknown methanogenic pathway. Proc Natl Acad Sci U S A 102:10664-9.

201. Whitman, W. B., D. C. Coleman, and W. J. Wiebe. 1998. : the

unseen majority. Proc Natl Acad Sci U S A 95:6578-83. 68 202. Widdel, F. 1986. Growth of Methanogenic Bacteria in Pure Culture with

2-Propanol and Other Alcohols as Hydrogen Donors. Appl Environ Microbiol

51:1056-1062.

203. Wolfe, R. S. 1993. An historical overview of methanogenesis. In J. G. Ferry (ed.),

Methanogenesis, Ecology, Physiology, Biochemistry & Genetics. Chapman &

Hall, New York.

204. Yeliseev, A., P. Gartner, U. Harms, D. Linder, and R. K. Thauer. 1993.

Function of methylcobalamin: coenzyme M methyltransferase isoenzyme II in

Methanosarcina barkeri. Arch Microbiol 159:530-6.

205. Zinder, S. H. 1993. Physiological ecology of methanogens. In J. G. Ferry (ed.),

Methanogenesis. Chapman&Hall, New York, NY.

206. Zirngibl, C., R. Hedderich, and R. K. Thauer. 1990.

N5,N10-methylenetetrahydromethanopterin dehydrogenase from

Methanobacterium thermoautotrophicum has hydrogenase activity. Febs Letters

261:112-116.

207. Zirngibl, C., W. Van Dongen, B. Schworer, R. Von Bunau, M. Richter, A.

Klein, and R. K. Thauer. 1992. H2-forming methylenetetrahydromethanopterin

dehydrogenase, a novel type of hydrogenase without iron-sulfur clusters in

methanogenic archaea. Eur J Biochem 208:511-20. 69

Chapter 2

Electron transport and energy conservation in methanogens

2.1 Introduction

Despite playing a critical role in global carbon cycle, methanogenesis is not an energetically rich pathway. The change of free Gibbs energy of methanogenesis from

H2 + CO2 is merely -131 kJ/mol, 5% of the oxidation of glucose. Acetate pathway that’s responsible for 2/3 of global biological methane output is even poorer (25, 78).

The change of Gibbs free energy of the conversion from acetate to CH4 and CO2 is only

-36 kJ/mol at physiological state, barely enough for the synthesis of one ATP per acetate.

How methanogens survive with these poor substrates makes the study of methanogenic energy conservation an interesting topic of research. The progress of this field will be summarized in this chapter.

2.2 Methanogenesis coupled ATP synthesis

The mechanism of energy conservation has been investigated in both whole cells and inverted membrane vesicles of methanogens. Early studies on whole cells of

Methanosarcina barkeri and Methanothermobacter thermoautotrophicus revealed a proton gradient driven ATP synthesis, suggesting a chemiosmotic model for energy conservation (24, 59). It was further concluded that proton motive force is responsible 70 for 90% of membrane potential in methanol-grown M. barkeri (8). This, together with the observation that the absence of Na+ doesn’t impact ATP synthesis, leads to the conclusion that H+ is the primary coupling ion for ATP synthesis in methanol-grown M. barkeri (9). Studies on inverted washed membrane vesicles of Methanosarcina mazei led to more detailed understanding of energy conservation during methanogenesis from methanol. The coupling of ATP synthesis to methane formation, as well as the coupling of electron transport to proton translocation was identified (21, 63). The stoichiometry of proton translocation and electron transport was later characterized in a H2:

CoM-S-S-CoB electron transport pathway in M. mazei, suggesting a 2H+/2e stoichiometry at heterodisulfide reductase and 2H+/2e stoichiometry at membrane-bound

F420 non-reducing hydrogenase (38)

Although methanogens share a similar approach of energy conservation with bacteria, their ATPases share little sequence similarity. For example, the ATPase from

M. barkeri only has around 20% sequence identity with Escherichia coli F1F0 type

ATPase, but surprisingly has a 50% sequence identity with eukaryotic V1V0 type

ATPases (41). On the other hand, methanogenic ATPases are similar in function with

F1F0 type ATPases, showing a physiological ATP synthase activity. In contrast, V1V0 type ATPases only hydrolyze ATP in vivo. These evidences led to the classification of archaeobacterial A1A0 type ATPases, distinct from bacterial and eukaryotic ATPases

(60).

Purification of A1A0 type ATPases in methanogens has been attempted in a number of organisms, including M. thermoautotrophicus, Methanosaeta thermophila, M. barkeri, M. mazei, Methanococcus voltae and Methanococcus jannaschii (13, 39, 40, 50, 71 56, 85). These proteins vary in subunit compositions, possibly due to dissociation of the complex during purification. The purified ATPase with the highest number of subunits is from M. jannaschii, which has 9 subunits (56). Like that of F1F0 ATPases, A1A0-type

ATPases were suggested to be comprised of two subcomplexes: the hydrophilic catalytic

A1 subcomplex and the hydrophobic A0 subcomplex. The A0 subcomplex is embedded in membrane and can potentially form an ion channel. The structure of ATPase from M. jannaschii was studied with electron microscopy, supporting this theory (Fig. 2-1) (15).

Figure 2-1 Electron microscopy and structure prediction of ATPase from M. jannaschii.

Panel A: Subunit and structure prediction of ATPase. Panel B: Electron microscopic image of ATPase.

Figure is modified from (15).

Early inhibitor studies suggested the presence of two types of ATPases in M.

+ thermoautotrophicus and M. mazei, the H -translocating A1A0 type ATPase and the

+ Na -translocating F1F0 type ATPase (6, 71, 72). Native PAGE of cell lysate of M. 72 thermoautotrophicus further strengthened this conclusion by showing two distinct bands after ATPase activity staining (72). However, later studies suggested only three out of fifty-one archaeal strains sequenced carry genes encoding F1F0 type

ATPases, including M. barkeri Fusaro, M. barkeri MS and Methanosarcina acetivorans, but not M. thermoautotrophicus or M. mazei (67). Further knockout studies showed that

F1F0 type ATPase is dispensable for M. acetivorans, suggesting previous conclusion on

+ the role of a Na -translocating F1F0 type ATPase in methanogens is incorrect and a function of F1F0 type ATPases is not established (67).

2.3 Membrane-bound electron transport chains of methanogens

Research done with inverted membrane vesicles of M. mazei suggested two electron transport chains: a H2: CoM-S-S-CoB electron transport chain and a F420:

CoM-S-S-CoB electron transport chain (19, 21). The characterization of these two systems led to the elucidation of membrane-bound, energy-conserving electron transport during methylotrophic and CO2-reducing methanogenesis. Later research on Ech hydrogenase and Rnf complex enhanced our understanding of electron transport during acetate metabolism, leading to the proposal of a third ferredoxin: CoM-S-S-CoB electron transport pathway (57, 80). 73

2.3.1 F420: CoM-S-S-CoB pathway

Characterization of inverted membrane vesicles of M. mazei showed that the membrane-bound F420: CoM-S-S-CoB oxidoreductase activity is coupled to transmembrane H+ translocation and ATP synthesis (21). The members of this electron transport chain include F420 dehydrogenase (Fpo), methanophenazine and DE-type heterodisulfide reductase (Fig. 2-2). The participation of methanophenazine in this electron transport chain has been investigated with its water soluble analogue

2-hydroxyphenazine (5). It was shown that 2-hydroxyphenazine can be reduced when

F420 was added as the terminal electron donor during the in vitro reconstitution of the F420:

CoM-S-S-CoB pathway and it can be re-oxidized when CoM-S-S-CoB was added as the terminal electron acceptor (5). The reduction and oxidation of methanophenazine form a cycle analogous to that of bacterial Q-loop. Two protons are translocated across membrane when every pair of electrons is transported via this ‘MP loop’ (38). F420 dehydrogenase was shown to be another site of proton translocation with a 2H+/2e stoichiometry (4).

The level of F420 was shown to be very low in acetate-grown methanogens, suggesting this F420: CoM-S-S-CoB pathway is not a major electron transport pathway during methanogenesis from acetate (26). The upregulation of F420 dehydrogenase in methanol- versus acetate-grown M. mazei further supported this suggestion (37). These evidences agree with the hypothesis that the F420: CoM-S-S-CoB electron transport pathway primarily functions during methanogenesis from methylated compounds. 74

Figure 2-2 Tentative model of the F420: CoM-S-S-CoB electron transport pathway.

2.3.2 H2: CoM-S-S-CoB pathway

Much like that of the F420: CoM-S-S-CoB pathway, research done on washed inverted membrane vesicles of M. mazei suggested the electron transport from H2 to

CoM-S-S-CoB couples proton translocation as well as ATP synthesis (64). As shown in

Figure 2-2 and Figure 2-3, both the F420: CoM-S-S-CoB pathway and H2: CoM-S-S-CoB pathway use the same cytochrome b containing DE type heterodisulfide reductase. The participation of methanophenazine in H2: CoM-S-S-CoB electron transport was also experimentally proven (2). A membrane-bound F420 non-reducing hydrogenase (Vho) that oxidizes H2 and passes electrons to methanophenazine was shown to be a member of 75

Figure 2-3 Tentative model of the H2: CoM-S-S-CoB electron transport pathway.

this electron transport chain. This hydrogenase has been purified in multiple systems

(for details, see section 2.5.2 of this chapter), and was shown to be a site of proton translocation with a 2H+/2e stoichiometry (22, 38, 45).

Characterization of Δfpo and Δfrh strains of M. barkeri suggested the H2:

CoM-S-S-CoB electron transport pathway is used by not only H2/ CO2-grown, but also methanol-grown M. barkeri (47) (for details, see section 2.5.1 of this chapter). This discovery led to the hypothesis that the H2: CoM-S-S-CoB electron transport pathway functions in both methanogenesis from methylated compounds and H2 + CO2 in

Methanosarcina species. However, this does not apply to other H2 + CO2 metabolizing methanogens. Little activity of either H2: CoM-S-S-CoB oxidoreductase or F420:

CoM-S-S-CoB oxidoreductase was found in obligate H2 + CO2 utilizing methanogen

Methanococcus thermolithotrophicus, suggesting this methanogen doesn’t use either of 76 the electron transport pathways mentioned above (19). This is consistent with current knowledge of methanogens because DE-type heterodisulfide reductase is a cytochrome b containing protein and few obligate H2 + CO2 utilizing methanogens contain cytochromes

(79). The mechanism of energy conservation for these methanogens is still largely unknown, although Setzke et al. suspected the reduction of CoM-S-S-CoB by soluble

ABC type heterodisulfide reductase may couple proton translocation by an unknown membrane protein (70).

2.3.3 The acetate-specific ferredoxin: CoM-S-S-CoB electron transport chain

A ferredoxin: CoM-S-S-CoB electron transport pathway has been identified in acetate-grown M. mazei and M. barkeri by showing membrane-bound ferredoxin:

CoM-S-S-CoB oxidoreductase activity (57, 81). This electron transport pathway involves a membrane-bound hydrogenase, Ech complex. Δech strains of M. mazei and

M. barkeri have been constructed and they were unable to grow on H2 + CO2 or acetate, but were able to grow on methanol or trimethylamine (58, 81). A reversible ferredoxin reducing hydrogenase activity has been identified with purified Ech (57). It was shown in the same report that Ech mediates an electron transport starting from carbon monoxide dehydrogenase and ferredoxin and ending at H2. This ferredoxin-reducing, H2-evolving activity was also found in Methanosarcina thermophila grown on acetate, further suggesting the involvement of Ech hydrogenase in acetate metabolism (76). A model was constructed from these evidences that Ech participates in electron transport in acetate-grown methanogens by providing H2 to membrane-bound F420 non-reducing 77 hydrogenase which further reduces methanophenazine (57, 81, 83). This model is summarized in Figure 2-4.

Figure 2-4 Tentative model of the ferredoxin: CoM-S-S-CoB electron transport pathway in

M. barkeri and M. mazei.

One unique member of Methanosarcina, M. acetivorans is a marine methanogen which doesn’t carry genes encoding Ech hydrogenase in its genome (55). Proteomic analysis of acetate- versus methanol- grown M. acetivorans suggested a unique membrane-bound Rnf complex is upregulated in acetate-grown cells, leading to the hypothesis that Rnf replaces the role of Ech and F420 non-reducing hydrogenase by providing electrons to methanophenazine (53-55). This hypothesis was supported in later research showing the reduction of RnfB with ferredoxin (80). These studies led to 78 a unique model of electron transport in acetate-grown M. acetivorans. The detailed description of this model can be found in Chapter 3 of this thesis.

2.4 Electron transport in obligate CO2-reducing methanogens

The mechanism of electron transport and energy conservation in obligate

CO2-reducing methanogens is poorly understood. In these organisms, methanophenazine and membrane-bound DE-type heterodisulfide reductases are absent.

This suggests the energy conserving membrane-bound H2: CoM-S-S-CoB electron transport pathway cannot be used by these methanogens. Although membrane-bound hydrogenases Eha/Ehb do exist in these methanogens (77), they were hypothesized to function similarly with Ech hydrogenase, namely by catalyzing the endergonic reduction of ferredoxin (E’≈ -500 mV) by H2 (E0’= -414 mV) (79). The reduced ferredoxin participates in the activation of methanofuran to formylmethanofuran (44). In this hypothesis, Eha/Ehb hydrogenases are not involved in energy conservation, leaving the model of electron transport and energy conservation for obligate CO2 reducers a mystery.

During the purification of the H2: CoM-S-S-CoB oxidoreductase in obligate CO2 reducer Methanothermobacter marburgensis, it was observed that the ABC-type heterodisulfide reductase (HdrABC) copurifies with soluble F420 non-reducing hydrogenase (Mvh) (70, 73). This Mvh/Hdr complex also catalyzes

CoM-S-S-CoB-dependent ferredoxin reduction and ferredoxin-dependent CoM-S-S-CoB reduction by H2 (79). Based on these observations, Rudolf Thauer proposed that the

Mvh/Hdr complex of obligate CO2 reducers catalyzes a bifurcation of electrons from H2 79

(E0’= -414 mV) to both CoM-S-S-CoB and ferredoxin (Fig. 2-5A) during which the endergonic reduction of ferredoxin (E’≈ -500 mV) is coupled to the exergonic reduction of CoM-S-S-CoB (E0’= -140 mV). In this hypothesis, reduced ferredoxin is sufficient for the fixation of CO2 to formylmethanofuran without the need of consumption of membrane potential by membrane-bound hydrogenases, retaining the membrane potential to drive ATP synthesis (79). This hypothesis is supported by a recent discovery that

F420 non-reducing hydrogenase Vhu homologous to Mvh, HdrABC, formate dehydrogenase (Fdh) and Tungsten-containing formylmethanofuran dehydrogenase (Fwd) form a complex in obligate CO2 reducer Methanococcus maripaludis (16). During growth on formate, genes encoding F420 non-reducing hydrogenases are dispensable suggesting a direct electron transfer from Fdh to HdrABC. HdrABC meanwhile catalyzes the reduction of ferredoxin, the electron donor to Fwd whose function is the fixation of CO2 to formylmethanofuran. During growth on H2 plus CO2, deletion of both vhu and vhc encoding F420 non-reducing hydrogenases resulted in impaired growth, suggesting a similar electron bifurcation scheme in which the electrons from H2 are used to reduce both CoM-S-S-CoB and ferredoxin. This model of electron bifurcation (Fig.

2-5B), like that of M. marburgensis, also eliminates the need of a membrane potential consuming hydrogenase for the reduction of ferredoxin. The detailed mechanism of energy conservation is currently unknown and the sites coupling energy conservation still remain to be discovered. 80

Figure 2-5 Model of electron bifurcation at Mvh/Hdr complex for obligate CO2 reducers.

Panel A: model of electron bifurcation for M. marburgensis, figure modified from Figure 5A in (79).

Panel B: model of electron bifurcation for M. maripaludis, figure modified from Figure 3 in (16).

Abbreviations: Mvh, F420 non-reducing hydrogenase; Hdr, heterodisulfide reductase; FdO, ferredoxin oxidized; FdR, ferredoxin reduced; CoM-SH, coenzyme M; CoB-SH, coenzyme B; CoM-S-S-CoB, heterodisulfide of coenzyme M and coenzyme B; MFR, methanofuran; Fwd, tungsten-containing formylmethanofuran dehydrogenase; Vhu, F420 non-reducing hydrogenase; Fdh, formate dehydrogenase.

2.5 Enzymes involved in electron transport chains of methanogens

2.5.1 F420 dehydrogenase

The electron entry of the F420: CoM-S-S-CoB electron transport pathway: F420 dehydrogenase (Fpo) has been purified and characterized from tindarius and M. mazei (1, 30). Both enzymes were shown to be membrane-bound protein complexes that harbor multiple iron sulfur clusters. FAD at the level of 0.2 molecule/protein was identified in Fpo from M. mazei (1). Both Fpo purified were shown to be protein complexes containing five subunits with a 1:1:1:1:1 stoichiometry. 81 Analysis of the genetic structure of fpo genes was performed in M. tindarius and

M. mazei (4, 84). The fpo gene cluster of M. tindarius surprisingly only contains four genes (84). This doesn’t exclude the possibly of other fpo genes because of the lack of information of genomic sequence for this microorganism. The fpo operon of M. mazei, on the other hand, has 12 genes encoding four of the five subunits of Fpo purified, FpoB,

FpoC, FpoD and FpoI (4). The fifth subunit of purified Fpo from M. mazei, FpoF, is encoded by a gene located distant from the fpo operon (4). Hydropathy plots of the remaining eight genes in the fpo operon showed that seven of them, FpoA, FpoH, FpoJ,

FpoK, FpoL, FpoM and FpoN, are membrane-integral proteins possibly encoding a membrane-bound subcomplex. The last gene of fpo operon, fpoO encodes a 2Fe-2S cluster binding protein which is possibly involved in electron transport (4). The predicted configuration of Fpo in M. mazei is presented in Figure 2-6.

Sequence comparison of the fpo genes with other protein complexes showed it is homologous to genes encoding NADH: ubiquinone oxidoreductase (complex I) from mitochondria and bacteria. Sequence analysis showed binding motifs of two 4Fe-4S clusters and 1 FAD in FpoF from M. mazei, suggesting it is the catalytic subunit of Fpo complex. The subunits that are not present in the purified Fpo possibly form a ion channel translocating proton across membrane (Fig. 2-6). This proton translocation coupled to F420 oxidation has been observed in inverted membrane vesicles from M. mazei, using F420 as the electron donor and 2-hydroxyphenzine as the electron acceptor.

A stoichiometry of 2H+/2e was observed at the same time (4). These results also 82 suggest that 2-hydroxyphenazine is the electron acceptor of Fpo, agreeing with the model that methanophenazine shuttles electrons between Fpo and heterodisulfide reductase.

Figure 2-6 Tentative model of F420 dehydrogenase from M. mazei. Figure is modified from (5).

Proteomic and microarray analysis of fpo operon and fpoF in M. acetivorans and

M. mazei showed the upregulation of fpo transcription and translation during growth on methanol comparing with acetate (37, 51, 52). The Δfpo andΔfpoF knockout strains of M. barkeri were constructed, but surprisingly showed normal growth and methanogenesis on methanol (47). Knock out strains of genes encoding another F420 reducing protein, F420 reducing hydrogenase (Frh), didn’t show any impaired growth or methanogenesis (47). However, when a double mutant of frh and fpo or frh and fpoF was constructed, growth and methanogenesis on methanol was severely impaired. 83

These results suggest both the F420: CoM-S-S-CoB pathway and the H2: CoM-S-S-CoB pathway take place in methanol-grown M. mazei. In the later pathway, Frh performs the role of feeding the system H2 with reducing equivalent coming from F420. Both pathways can independently transport electrons coupling proton translocation, but when both of them are blocked, methanogenesis on methanol stops. This, however, does not apply to M. acetivorans and M. tindarius, because they either do not possess hydrogenases or show very little H2 production during methanogenesis (29).

2.5.2 Membrane bound F420 non-reducing hydrogenase

Multiple types of hydrogenases have been identified in methanogens, including membrane-bound Ech hydrogenase, membrane-bound F420 reducing hydrogenase and

F420 non-reducing hydrogenase. The F420 non-reducing hydrogenases are divided into two groups: cytoplasmic F420 non-reducing hydrogenases annotated as Mvh hydrogenases found in obligate H2 + CO2 utilizing methanogens and membrane-bound non-reducing hydrogenases annotated as Vho hydrogenases found in Methanosarcina species. Vho hydrogenases were shown to copurify with DE-type heterodisulfide reductases and form a membrane-bound H2: CoM-S-S-CoB oxidoreductase complex in M. barkeri (36), suggesting its participation in H2: CoM-S-S-CoB electron transport pathway. In this pathway, it was hypothesized that Vho hydrogenase accepts electrons from H2, passes them to methanophenazine, and also couples transmembrane proton translocation at the same time (Fig. 2-3). The coupling of H2: 2-hydroxyphenazine oxidoreductase activity with proton translocation was observed in inverted membrane vesicles of M. mazei, 84 supporting this hypothesis (38). Besides participating in the H2: CoM-S-S-CoB electron transport pathway that functions in both methanol and H2 + CO2 pathways, the involvement of Vho in acetate-specific ferredoxin: CoM-S-S-CoB electron transport pathway has also been observed (82).

Genetic analysis of Methanosarcina species revealed multiple copies of vho gene cluster with high sequence identity between each other (18, 20, 28), named vho, vht and vhx (viologen hydrogenase one, two and unknown). The vht operon is comprised of four genes. The genes encoding the large and small subunits of the hydrogenase are specifically vhtA and vhtG. The vhtC gene encodes a cytochrome b protein that’s hypothesized to be the electron acceptor of the hydrogenase. The vhtD gene encodes a proposed maturation protein and is missing from vho and vhx operons (20, 28). M. mazei possesses all three vho homologue operons while M. barkeri and M. acetivorans are lack of the vho operon (28).

The membrane-bound F420 non-reducing hydrogenases have been purified and characterized from M. mazei and M. barkeri (22, 45). In both cases the protein complexes were shown to have both the large and small subunits, contain NiFe center as the active site and are tightly membrane-bound. More detailed biochemical work, most notably the capability of purified Vho to reduce 2-hydroxyphenzine or cytochrome b is still lacking, leaving the H2: CoM-S-S-CoB electron transport pathway still partly unresolved. 85 2.5.3 Ech hydrogenase

Ech hydrogenase is a membrane-bound [NiFe] hydrogenase that shares sequence similarity with E.coli hydrogenase 3, hydrogenase 4, and CO-induced hydrogenase from

Rhodospirillum rubrum. It is named after these homologous proteins (Ech=E.coli hydrogenase 3-type hydrogenase) (57). Sequence homology has been identified between subunits of Ech hydrogenase from M. barkeri and E.coli NADH: ubiquinone oxidoreductase (Complex I), suggesting similar configuration and subunit arrangement

(48, 57). The ech operon of M. barkeri showed the presence of six genes, encoding the six proteins of EchABCDEF complex (48). Ech hydrogenase has been purified in M. barkeri, confirming this subunit composition (57). Sequence analysis suggested echE encodes the NiFe cluster containing hydrogenase large subunit, echC encodes the iron sulfur cluster containing hydrogenase small subunit, echF encodes a 4Fe-4S cluster containing protein that’s hypothesized as the ferredoxin oxidizing subunit, echA and echB encode two membrane-bound proteins that potentially form a proton channel, and echD encodes a hydrophilic protein with no predicted function (27, 48, 57). A model of Ech hydrogenase is thus constructed, showing a similar architecture with that of Complex I in

E. coli (Figure 2-7).

The physiological role of Ech hydrogenase has been subject to biochemical and genetic studies. An electron transport chain from CO to Cdh, to ferredoxin, to Ech hydrogenase and to H2 has been constructed with purified components, showing a

CO-oxidizing, H2-evolving activity in M. barkeri (57). A similar ferredoxin dependent, 86

Figure 2-7 Tentative model of Ech hydrogenase in M. barkeri. Figure is modified from (49).

CO-oxidizing, H2-evolving electron transport chain was reconstituted earlier with membrane from acetate-grown M. thermophila, supporting a model of Ech hydrogenase functioning in ferredoxin: CoM-S-S-CoB electron transport during acetate metabolism

(76). TheΔech knockout strains of M. barkeri and M. mazei were constructed and both showed the incapability of growth on acetate, further supporting this role (58, 82). A proton gradient generation coupled to the hydrogenase activity of Ech has been observed, confirming it is a proton channel (11, 58, 83). A different role of Ech hydrogenase during H2 + CO2 metabolism has also been proposed. In this model, Ech catalyses the reversed reaction, generating reduced ferredoxin for the activation of methanofuran in the first step of methanogenesis from CO2 (58). 87 2.5.4 Rnf complex

Rnf complex was first discovered in Rhodobacter capsulatus, a photosynthetic nitrogen fixing bacteria (68), and obtains its name from this organism (Rnf=Rhodobacter nitrogen fixation). Since then, Rnf was identified from more organisms, including non nitrogen fixating bacteria E. coli (43), acetogenic bacteria Acetobacterium woodii (7) and archaea M. acetivorans (55). In nitrogen fixing bacteria, membrane-bound Rnf complex has been shown to participate in nitrogen fixation (17, 23, 42, 66, 68). It was hypothesized that Rnf complexes are involved in electron transport, either mediates an electron flow from NADH to ferredoxin coupled to an influx of sodium ions (42, 61, 68), or accepts electrons from ferredoxin and donates them to an electron acceptor (7, 55, 61).

However, there has been no biochemical evidence for either electron transport pathways before its participation in the ferredoxin: CoM-S-S-CoB electron transport chain was identified in M. acetivorans (80).

Bacterial Rnf complexes are composed of six or seven subunits, Rnf ABCDEGH

(43) or Rnf ABCDEG (7). In M. acetivorans and M. thermophila, two more genes are cotranscribed with the six genes homologous to rnfABCDEG. This leads to the hypothesis that archaeal Rnf contains eight subunits instead of six or seven, a model of which is shown in Figure 2-8 (55). These two additional genes encode a membrane-bound protein and a multi-heme cytochrome c. Evidence supporting the participation of Rnf in the ferredoxin: CoM-S-S-CoB electron transport pathway serving as the electron acceptor of ferredoxin and electron donor of methanophenazine has been observed (80). Two subunits of Rnf, RnfB and RnfG have been heterologously 88 expressed and purified in E. coli (74, 75), suggesting RnfB is an iron-sulfur protein and

RnfG is a flavoprotein. Sequence analysis suggests RnfC is an iron-sulfur protein and potentially participate in electron transport. The remaining subunits RnfA, RnfD, RnfE are membrane-bound subunits that were hypothesized to form an ion channel (55).

Figure 2-8 Tentative model of Rnf complex in M. acetivorans.

The rnf genes of M. acetivorans were previously annotated as nqr in databases

(14, 53, 61). This annotation leads to the hypothesis that Rnf translocates Na+ like Nqr, although no biochemical evidence for this ion translocation has been reported for any Rnf

(61, 69). In bacteria, nqr genes encode a Na+-translocating NADH: quinone oxidoreductase (Na+-Nqr), a protein complex analogous to complex I in that both enzyme complexes are bacterial membrane-bound NADH: ubiquinone oxidoreductases, but 89 shares little sequence homology with complex I and generates a transmembrane Na+ gradient instead of a proton gradient (46). The presence of Na+-Nqr in Bacteria has been identified from a number of microorganisms, including halophiles such as Vibrio alginolyticus (35) and pathogenic bacteria such as Haemophilus influenza and Vibrio cholerae (31, 32), Analysis of the genes encoding Na+-Nqr suggested it is encoded by a six gene operon nqrABCDEF well conserved in all species (33). Na+-Nqr has been purified or partially purified and biochemically characterized from a number of microorganisms including V. alginolyticus, Vibrio harveyi and V. cholerae (3, 10, 35, 62,

65, 86). Detailed biochemical analysis of the subunits of Na+-Nqr suggested it is a protein complex rich in flavins: NqrB has one non-covalently bound riboflavin and one covalently bound FMN (12, 34); NqrC has one covalently bound FMN (34); NqrF has one non-covalently bound FAD and one 2Fe-2S cluster and functions as the entry point of electrons for Na+-Nqr (46).

Despite the similarities between bacterial Rnf and Na+-Nqr in the number of subunits, subcellular location and electron donor, significant differences exist between the two: Rnf was never reported to use quinones as electron acceptors; Rnf in many cases couples an influx of ions for its thermodynamically unfavorable electron transport from

NADH to ferredoxin (68); The cofactor content of Rnf dramatically differs from Na+-Nqr in that Rnf is rich in iron sulfur clusters while Na+-Nqr is rich in flavins. Sequence comparison between subunits of Na+-Nqr from V. cholerae and Rnf from M. acetivorans suggested only the two membrane-bound subunits NqrD and NqrE that do not participate in electron transport share a significant sequence homology with their Rnf counterparts 90 (34% and 39% sequence identity respectively). However, the subunits shown to bind to cofactors or participate in electron transport, NqrA, NqrB, NqrC and NqrF, only showed

3-19% sequence identities with their counterparts. No conservation of active sites was observed in this comparison, except for the conservation of the FMN-binding SGAT motif in RnfG and NqrC. These observations lead to the invalidation of the annotation of nqr for the rnf genes from M. acetivorans.

2.6 References

1. Abken, H. J., and U. Deppenmeier. 1997. Purification and properties of an

F420H2 dehydrogenase from Methanosarcina mazei Go1. Fems Microbiology

Letters 154:231-237.

2. Abken, H. J., M. Tietze, J. Brodersen, S. Baumer, U. Beifuss, and U.

Deppenmeier. 1998. Isolation and characterization of methanophenazine and

function of phenazines in membrane-bound electron transport of Methanosarcina

mazei Go1. J Bacteriol 180:2027-32.

3. Barquera, B., P. Hellwig, W. Zhou, J. E. Morgan, C. C. Hase, K. K. Gosink,

M. Nilges, P. J. Bruesehoff, A. Roth, C. R. Lancaster, and R. B. Gennis. 2002.

Purification and characterization of the recombinant Na+-translocating NADH:

quinone oxidoreductase from Vibrio cholerae. Biochemistry 41:3781-9.

4. Baumer, S., T. Ide, C. Jacobi, A. Johann, G. Gottschalk, and U. Deppenmeier.

2000. The F420H2 dehydrogenase from Methanosarcina mazei is a redox-driven 91 proton pump closely related to NADH dehydrogenases. J Biol Chem

275:17968-73.

5. Baumer, S., E. Murakami, J. Brodersen, G. Gottschalk, S. W. Ragsdale, and

U. Deppenmeier. 1998. The F420H2: heterodisulfide oxidoreductase system from

Methanosarcina species: 2-Hydroxyphenazine mediates electron transfer from

F420H2 dehydrogenase to heterodisulfide reductase. FEBS Lett 428:295-8.

6. Becher, B., and V. Muller. 1994. ΔμNa+ drives the synthesis of ATP via an

ΔμNa+-translocating F1F0-ATP synthase in membrane vesicles of the archaeon

Methanosarcina mazei Go1. J Bacteriol 176:2543-50.

7. Biegel, E., S. Schmidt, and V. Muller. 2009. Genetic, immunological and

biochemical evidence for a Rnf complex in the acetogen Acetobacterium woodii.

Environ Microbiol 11:1438-43.

8. Blaut, M., and G. Gottschalk. 1984. Coupling of ATP synthesis and methane

formation from methanol and molecular hydrogen in Methanosarcina barkeri.

Eur J Biochem 141:217-22.

9. Blaut, M., V. Muller, K. Fiebig, and G. Gottschalk. 1985. Sodium ions and an

energized membrane required by Methanosarcina barkeri for the oxidation of

methanol to the level of formaldehyde. J Bacteriol 164:95-101.

10. Bogachev, A. V., Y. V. Bertsova, B. Barquera, and M. I. Verkhovsky. 2001.

Sodium-dependent steps in the redox reactions of the Na+-motive NADH:

quinone oxidoreductase from Vibrio harveyi. Biochemistry 40:7318-23. 92 11. Bott, M., B. Eikmanns, and R. K. Thauer. 1986. Coupling of carbon monoxide

oxidation to CO2 and H2 with the phosphorylation of ADP in acetate-grown

Methanosarcina barkeri. Eur J Biochem 159:393-8.

12. Casutt, M. S., T. Huber, R. Brunisholz, M. Tao, G. Fritz, and J. Steuber.

Localization and function of the membrane-bound riboflavin in the

Na+-translocating NADH: quinone oxidoreductase (Na+-NQR) from Vibrio

cholerae. J Biol Chem 285:27088-99.

13. Chen, W., and J. Konisky. 1993. Characterization of a membrane-associated

ATPase from Methanococcus voltae, a methanogenic member of the Archaea. J

Bacteriol 175:5677-82.

14. Comprehensive, T., and M. Resource. J. Craig Venter Institute,

.

15. Coskun, U., Y. L. Chaban, A. Lingl, V. Muller, W. Keegstra, E. J. Boekema,

and G. Gruber. 2004. Structure and subunit arrangement of the A-type ATP

synthase complex from the archaeon Methanococcus jannaschii visualized by

electron microscopy. J Biol Chem 279:38644-8.

16. Costa, K. C., P. M. Wong, T. Wang, T. J. Lie, J. A. Dodsworth, I. Swanson, J.

A. Burn, M. Hackett, and J. A. Leigh. 2010. Protein complexing in a

methanogen suggests electron bifurcation and electron delivery from formate to

heterodisulfide reductase. Proc. Natl. Acad. Sci. U S A 107:11050-11055.

17. Curatti, L., C. S. Brown, P. W. Ludden, and L. M. Rubio. 2005. Genes

required for rapid expression of nitrogenase activity in Azotobacter vinelandii.

Proc Natl Acad Sci U S A 102:6291-6. 93 18. Deppenmeier, U. 1995. Different structure and expression of the operons

encoding the membrane-bound hydrogenases from Methanosarcina mazei Go1.

Arch Microbiol 164:370-6.

19. Deppenmeier, U., M. Blaut, and G. Gottschalk. 1991. H2: heterodisulfide

oxidoreductase, a second energy-conserving system in the methanogenic strain

Go1. Arch Microbiol 155:272-277.

20. Deppenmeier, U., M. Blaut, S. Lentes, C. Herzberg, and G. Gottschalk. 1995.

Analysis of the vhoGAC and vhtGAC operons from Methanosarcina mazei strain

Go1, both encoding a membrane-bound hydrogenase and a cytochrome b. Eur J

Biochem 227:261-9.

21. Deppenmeier, U., M. Blaut, A. Mahlmann, and G. Gottschalk. 1990. Reduced

coenzyme F420: heterodisulfide oxidoreductase, a proton-translocating redox

system in methanogenic bacteria. Proc Natl Acad Sci U S A 87:9449-53.

22. Deppenmeier, U., M. Blaut, B. Schmidt, and G. Gottschalk. 1992. Purification

and properties of a F420-nonreactive, membrane-bound hydrogenase from

Methanosarcina strain Go1. Arch Microbiol 157:505-11.

23. Desnoues, N., M. Lin, X. Guo, L. Ma, R. Carreno-Lopez, and C. Elmerich.

2003. Nitrogen fixation genetics and regulation in a Pseudomonas stutzeri strain

associated with rice. Microbiology 149:2251-62.

24. Doddema, H. J., T. J. Hutten, C. van der Drift, and G. D. Vogels. 1978. ATP

hydrolysis and synthesis by the membrane-bound ATP synthetase complex of

Methanobacterium thermoautotrophicum. J Bacteriol 136:19-23. 94 25. Ferry, J. G. 1993. Fermentation of acetate, p. 304-334. In J. G. Ferry (ed.),

Methanogenesis: Ecology, Physiology, Biochemistry & Genetics. Chapman &

Hall, New York.

26. Ferry, J. G. 1992. Methane from acetate. J Bacteriol 174:5489-95.

27. Forzi, L., J. Koch, A. M. Guss, C. G. Radosevich, W. W. Metcalf, and R.

Hedderich. 2005. Assignment of the [4Fe-4S] clusters of Ech hydrogenase from

Methanosarcina barkeri to individual subunits via the characterization of

site-directed mutants. FEBS J 272:4741-53.

28. Guss, A. M., G. Kulkarni, and W. W. Metcalf. 2009. Differences in

hydrogenase gene expression between Methanosarcina acetivorans and

Methanosarcina barkeri. J Bacteriol 191:2826-33.

29. Guss, A. M., B. Mukhopadhyay, J. K. Zhang, and W. W. Metcalf. 2005.

Genetic analysis of mch mutants in two Methanosarcina species demonstrates

multiple roles for the methanopterin-dependent C-1 oxidation/reduction pathway

and differences in H2 metabolism between closely related species. Mol Microbiol

55:1671-80.

30. Haase, P., U. Deppenmeier, M. Blaut, and G. Gottschalk. 1992. Purification

and characterization of F420H2-dehydrogenase from . Eur J

Biochem 203:527-31.

31. Hase, C. C., and J. J. Mekalanos. 1999. Effects of changes in membrane sodium

flux on virulence gene expression in Vibrio cholerae. Proc Natl Acad Sci U S A

96:3183-7. 95 32. Hayashi, M., Y. Nakayama, and T. Unemoto. 1996. Existence of

Na+-translocating NADH-quinone reductase in Haemophilus influenzae. FEBS

Lett 381:174-6.

33. Hayashi, M., Y. Nakayama, and T. Unemoto. 2001. Recent progress in the

Na+-translocating NADH-quinone reductase from the marine Vibrio alginolyticus.

Biochim Biophys Acta 1505:37-44.

34. Hayashi, M., Y. Nakayama, M. Yasui, M. Maeda, K. Furuishi, and T.

Unemoto. 2001. FMN is covalently attached to a threonine residue in the NqrB

and NqrC subunits of Na+-translocating NADH-quinone reductase from Vibrio

alginolyticus. FEBS Lett 488:5-8.

35. Hayashi, M., and T. Unemoto. 1986. FAD and FMN flavoproteins participate in

the sodium-transport respiratory-chain NADH: quinone reductase of a marine

bacterium, Vibrio alginolyticus. FEBS Lett 202:327-330.

36. Heiden, S., R. Hedderich, E. Setzke, and R. K. Thauer. 1993. Purification of a

cytochrome b containing H2: heterodisulfide oxidoreductase complex from

membranes of Methanosarcina barkeri. Eur J Biochem 213:529-35.

37. Hovey, R., S. Lentes, A. Ehrenreich, K. Salmon, K. Saba, G. Gottschalk, R. P.

Gunsalus, and U. Deppenmeier. 2005. DNA microarray analysis of

Methanosarcina mazei Go1 reveals adaptation to different methanogenic

substrates. Mol Genet Genomics 273:225-39.

38. Ide, T., S. Baumer, and U. Deppenmeier. 1999. Energy conservation by the H2:

heterodisulfide oxidoreductase from Methanosarcina mazei Go1: identification of

two proton-translocating segments. J Bacteriol 181:4076-80. 96 39. Inatomi, K. 1986. Characterization and purification of the membrane-bound

ATPase of the archaebacterium Methanosarcina barkeri. J Bacteriol 167:837-41.

40. Inatomi, K., Y. Kamagata, and K. Nakamura. 1993. Membrane ATPase from

the aceticlastic methanogen Methanothrix thermophila. J Bacteriol 175:80-4.

41. Inatomi, K., M. Maeda, and M. Futai. 1989. Dicyclohexylcarbodiimide-binding

protein is a subunit of the Methanosarcina barkeri ATPase complex. Biochem

Biophys Res Commun 162:1585-90.

42. Jeong, H. S., and Y. Jouanneau. 2000. Enhanced nitrogenase activity in strains

of Rhodobacter capsulatus that overexpress the rnf genes. J Bacteriol

182:1208-14.

43. Jouanneau, Y., H. S. Jeong, N. Hugo, C. Meyer, and J. C. Willison. 1998.

Overexpression in Escherichia coli of the rnf genes from Rhodobacter

capsulatus--characterization of two membrane-bound iron-sulfur proteins. Eur J

Biochem 251:54-64.

44. Karrasch, M., G. Borner, M. Enssle, and R. K. Thauer. 1989.

Formylmethanofuran dehydrogenase from methanogenic bacteria, a

molybdoenzyme. FEBS Lett 253:226-30.

45. Kemner, J. M., and J. G. Zeikus. 1994. Purification and characterization of

membrane-bound hydrogenase from Methanosarcina barkeri MS. Arch Microbiol

161:47-54.

46. Kerscher, S., S. Drose, V. Zickermann, and U. Brandt. 2007. The three

families of respiratory NADH dehydrogenases. Results Probl Cell Differ

45:185-222. 97 47. Kulkarni, G., D. M. Kridelbaugh, A. M. Guss, and W. W. Metcalf. 2009.

Hydrogen is a preferred intermediate in the energy-conserving electron transport

chain of Methanosarcina barkeri. Proc Natl Acad Sci U S A 106:15915-20.

48. Kunkel, A., J. A. Vorholt, R. K. Thauer, and R. Hedderich. 1998. An

Escherichia coli hydrogenase-3-type hydrogenase in methanogenic archaea. Eur J

Biochem 252:467-76.

49. Kurkin, S., J. Meuer, J. Koch, R. Hedderich, and S. P. Albracht. 2002. The

membrane-bound [NiFe]-hydrogenase (Ech) from Methanosarcina barkeri:

unusual properties of the iron-sulphur clusters. Eur J Biochem 269:6101-11.

50. Lemker, T., C. Ruppert, H. Stoger, S. Wimmers, and V. Muller. 2001.

Overproduction of a functional A1 ATPase from the archaeon Methanosarcina

mazei Go1 in Escherichia coli. Eur J Biochem 268:3744-50.

51. Lessner, D. J., L. Li, Q. Li, T. Rejtar, V. P. Andreev, M. Reichlen, K. Hill, J.

J. Moran, B. L. Karger, and J. G. Ferry. 2006. An unconventional pathway for

reduction of CO2 to methane in CO-grown Methanosarcina acetivorans revealed

by proteomics. Proc Natl Acad Sci U S A 103:17921-6.

52. Li, L., Q. Li, L. Rohlin, U. Kim, K. Salmon, T. Rejtar, R. P. Gunsalus, B. L.

Karger, and J. G. Ferry. 2007. Quantitative proteomic and microarray analysis

of the archaeon Methanosarcina acetivorans grown with acetate versus methanol.

J Proteome Res 6:759-71.

53. Li, Q., L. Li, T. Rejtar, B. L. Karger, and J. G. Ferry. 2005. Proteome of

Methanosarcina acetivorans part I: an expanded view of the biology of the cell. J

Proteome Res 4:112-28. 98 54. Li, Q., L. Li, T. Rejtar, B. L. Karger, and J. G. Ferry. 2005. Proteome of

Methanosarcina acetivorans part II: comparison of protein levels in acetate- and

methanol-grown cells. J Proteome Res 4:129-35.

55. Li, Q., L. Li, T. Rejtar, D. J. Lessner, B. L. Karger, and J. G. Ferry. 2006.

Electron transport in the pathway of acetate conversion to methane in the marine

archaeon Methanosarcina acetivorans. J Bacteriol 188:702-10.

56. Lingl, A., H. Huber, K. O. Stetter, F. Mayer, J. Kellermann, and V. Muller.

2003. Isolation of a complete A1A0 ATP synthase comprising nine subunits from

the hyperthermophile Methanococcus jannaschii. Extremophiles 7:249-57.

57. Meuer, J., S. Bartoschek, J. Koch, A. Kunkel, and R. Hedderich. 1999.

Purification and catalytic properties of Ech hydrogenase from Methanosarcina

barkeri. Eur J Biochem 265:325-35.

58. Meuer, J., H. C. Kuettner, J. K. Zhang, R. Hedderich, and W. W. Metcalf.

2002. Genetic analysis of the archaeon Methanosarcina barkeri Fusaro reveals a

central role for Ech hydrogenase and ferredoxin in methanogenesis and carbon

fixation. Proc Natl Acad Sci U S A 99:5632-7.

59. Mountfort, D. O. 1978. Evidence from ATP synthesis driven by a proton

gradient in Methanosarcina barkeri. Biochem Biophys Res Commun 85:1346-51.

60. Muller, V., M. Blaut, and G. Gottschalk. 1993. Bioenergetics of

methanogenesis. In J. G. Ferry (ed.), Methanogenesis : Ecology, Physiology,

Biochemistry & Genetics. Chapman & Hall, New York. 99 61. Muller, V., F. Imkamp, E. Biegel, S. Schmidt, and S. Dilling. 2008. Discovery

of a ferredoxin:NAD+-oxidoreductase (Rnf) in Acetobacterium woodii: a novel

potential coupling site in acetogens. Ann N Y Acad Sci 1125:137-46.

62. Nakayama, Y., M. Hayashi, and T. Unemoto. 1998. Identification of six

subunits constituting Na+-translocating NADH-quinone reductase from the

marine Vibrio alginolyticus. FEBS Lett 422:240-2.

63. Peinemann, S., M. Blaut, and G. Gottschalk. 1989. ATP synthesis coupled to

methane formation from methyl-CoM and H2 catalyzed by vesicles of the

methanogenic bacterial strain Go1. Eur J Biochem 186:175-80.

64. Peinemann, S., R. Hedderich, M. Blaut, R. K. Thauer, and G. Gottschalk.

1990. ATP synthesis coupled to electron transfer from H2 to the heterodisulfide of

2-mercaptoethanesulfonate and 7-mercaptoheptanoylthreonine phosphate in

vesicle preparations of the methanogenic bacterium strain Go1. FEBS Lett

263:57-60.

65. Pfenninger-Li, X. D., S. P. Albracht, R. van Belzen, and P. Dimroth. 1996.

NADH: ubiquinone oxidoreductase of Vibrio alginolyticus: purification,

properties, and reconstitution of the Na+ pump. Biochemistry 35:6233-42.

66. Saeki, K., and H. Kumagai. 1998. The rnf gene products in Rhodobacter

capsulatus play an essential role in nitrogen fixation during anaerobic

DMSO-dependent growth in the dark. Arch Microbiol 169:464-7.

67. Saum, R., K. Schlegel, B. Meyer, and V. Muller. 2009. The F1FO ATP synthase

genes in Methanosarcina acetivorans are dispensable for growth and ATP

synthesis. FEMS Microbiol Lett 300:230-6. 100 68. Schmehl, M., A. Jahn, A. Meyer zu Vilsendorf, S. Hennecke, B. Masepohl, M.

Schuppler, M. Marxer, J. Oelze, and W. Klipp. 1993. Identification of a new

class of nitrogen fixation genes in Rhodobacter capsulatus: a putative membrane

complex involved in electron transport to nitrogenase. Mol Gen Genet

241:602-15.

69. Schmidt, S., E. Biegel, and V. Muller. 2009. The ins and outs of Na+

bioenergetics in Acetobacterium woodii. Biochim Biophys Acta 1787:691-6.

70. Setzke, E., R. Hedderich, S. Heiden, and R. K. Thauer. 1994. H2:

heterodisulfide oxidoreductase complex from Methanobacterium

thermoautotrophicum: composition and properties. Eur J Biochem 220:139-48.

71. Smigan, P., A. Majernik, and M. Greksak. 1994. Na+-driven ATP synthesis in

Methanobacterium thermoautotrophicum and its differentiation from H+-driven

ATP synthesis by rhodamine 6G. FEBS Lett 347:190-4.

72. Smigan, P., A. Majernik, P. Polak, I. Hapala, and M. Greksak. 1995. The

presence of H+ and Na+-translocating ATPases in Methanobacterium

thermoautotrophicum and their possible function under alkaline conditions. FEBS

Lett 371:119-22.

73. Stojanowic, A., G. J. Mander, E. C. Duin, and R. Hedderich. 2003.

Physiological role of the F420-non-reducing hydrogenase (Mvh) from

Methanothermobacter marburgensis. Arch Microbiol 180:194-203.

74. Suharti, S. de Vries, and J. G. Ferry. 2010. Characterization of the RnfB

subunit of Rnf complex from the Archaeon Methanosarcina acetivorans.

Submitted. 101 75. Suharti, T. Rejtar, S. de Vries, B. L. Karger, and J. G. Ferry. 2010.

Characterization of a flavin-containing subunit of Rnf complex from the

Archaeon Methanosarcina acetivoras. in preparation.

76. Terlesky, K. C., and J. G. Ferry. 1988. Ferredoxin requirement for electron

transport from the carbon monoxide dehydrogenase complex to a

membrane-bound hydrogenase in acetate-grown Methanosarcina thermophila. J

Biol Chem 263:4075-9.

77. Tersteegen, A., and R. Hedderich. 1999. Methanobacterium

thermoautotrophicum encodes two multisubunit membrane-bound [NiFe]

hydrogenases. Transcription of the operons and sequence analysis of the deduced

proteins. Eur J Biochem 264:930-43.

78. Thauer, R. K. 1998. Biochemistry of methanogenesis: a tribute to Marjory

Stephenson. 1998 Marjory Stephenson Prize Lecture. Microbiology 144 ( Pt

9):2377-406.

79. Thauer, R. K., A. K. Kaster, M. Goenrich, M. Schick, T. Hiromoto, and S.

Shima. 2010. Hydrogenases from methanogenic archaea, nickel, a novel cofactor,

and H2 storage. Annu Rev Biochem 79:507-36.

80. Wang, M., Suharti, J. F. Tomb, and J. G. Ferry. 2010. The electron transport

chain of acetate-grown Methanosarcina acetivorans. submitted.

81. Welte, C., V. Kallnik, M. Grapp, G. Bender, S. Ragsdale, and U.

Deppenmeier. Function of Ech hydrogenase in ferredoxin-dependent,

membrane-bound electron transport in Methanosarcina mazei. J Bacteriol

192:674-8. 102 82. Welte, C., V. Kallnik, M. Grapp, G. Bender, S. Ragsdale, and U.

Deppenmeier. 2010. Function of Ech hydrogenase in ferredoxin-dependent,

membrane-bound electron transport in Methanosarcina mazei. J Bacteriol

192:674-678.

83. Welte, C., C. Kratzer, and U. Deppenmeier. Involvement of Ech hydrogenase

in energy conservation of Methanosarcina mazei. FEBS J.

84. Westenberg, D. J., A. Braune, C. Ruppert, V. Muller, C. Herzberg, G.

Gottschalk, and M. Blaut. 1999. The F420H2-dehydrogenase from Methanolobus

tindarius: cloning of the ffd operon and expression of the genes in Escherichia

coli. FEMS Microbiol Lett 170:389-98.

85. Wilms, R., C. Freiberg, E. Wegerle, I. Meier, F. Mayer, and V. Muller. 1996.

Subunit structure and organization of the genes of the A1A0 ATPase from the

Archaeon Methanosarcina mazei Go1. J Biol Chem 271:18843-52.

86. Zhou, W., Y. V. Bertsova, B. Feng, P. Tsatsos, M. L. Verkhovskaya, R. B.

Gennis, A. V. Bogachev, and B. Barquera. 1999. Sequencing and preliminary

characterization of the Na+-translocating NADH: ubiquinone oxidoreductase from

Vibrio harveyi. Biochemistry 38:16246-52. 103

Chapter 3

The membrane-bound electron transport chain of acetate-grown

Methanosarcina acetivorans

3.1 Abstract

The membrane-bound electron transport chain of acetate-grown Methanosarcina acetivorans was investigated. The CdhAE component of the CO dehydrogenase/acetyl-CoA synthase (Cdh) was purified and shown to reduce a ferredoxin purified using an assay coupling reduction of the ferredoxin to oxidation of CdhAE.

Mass spectrometry analysis of the ferredoxin sequence identified the encoding gene from annotations for nine ferredoxins encoded in the genome. The ferredoxin was shown to belong to a clade that included ferredoxins implicated in the pathway for conversion of acetate to methane. The ferredoxin reduced the purified RnfB subunit of the membrane-bound Rnf complex. Reduction of purified membranes from acetate-grown cells with ferredoxin reduced the membrane-associated multi-heme cytochrome c that was re-oxidized by addition of CoM-S-S-CoB. The ferredoxin also reduced a soluble analog of methanophenazine (2-hydoxyphenazine) that in the presence of membranes was re-oxidized by addition of CoM-S-S-CoB. Addition of 2-hydoxyphenazine re-oxidized cytochrome c of purified membranes that was reduced by ferredoxin. The results support an electron transport chain originating with ferredoxin accepting electrons 104 from Cdh and donating to the membrane-bound Rnf complex. The results also implicate a role for cytochrome c and methanophenazine culminating with reduction of

CoM-S-S-CoB catalyzed by a heterodisulfide reductase.

3.2 Introduction

The decomposition of complex organic matter to methane (biomethanation) in diverse anaerobic habitats of Earth’s biosphere is an important component of the global carbon cycle (3). Biomethanation of renewable plant organic matter is also a viable alternative to fossil fuels (11). The process involves an anaerobic microbial food chain comprised of distinct metabolic groups, the first of which metabolizes the complex organic matter primarily to acetate and also formate or H2 that are growth substrates for two distinct methane-producing groups (methanogens) (27). The methyl group of acetate contributes most of the methane produced in the biomethanation process via the aceticlastic pathway whereas the remainder originates primarily from the reduction of

CO2 with electrons derived from the oxidation of formate or H2 in the CO2-reduction pathway (12, 49). Lesser albeit significant amounts of methane derive from the methyl groups of methanol, methylamines and dimethylsulfide (27).

In the aceticlastic pathway, acetate is activated to acetyl-CoA followed by cleavage into methyl and carbonyl groups catalyzed by the CO dehydrogenase/acetyl-CoA synthase (Cdh) enzyme complex that also oxidizes the latter to CO2 and reduces ferredoxin (46, 47). The methyl group is ultimately transferred to coenzyme M

(HS-CoM) producing CH3-S-CoM that is reductively demethylated to methane with 105 electrons donated by coenzyme B (HS-CoB). The heterodisulfide CoM-S-S-CoB is a product of the demethylation reaction that is reduced to the sulfhydryl forms of the cofactors by heterodisulfide reductase (HdrDE) with electrons originating from reduced ferredoxin through a membrane-bound electron transport chain generating an ion gradient driving ATP synthesis by the proton translocating A1A0-type ATP synthase (36, 38). The reactions and enzymes catalyzing carbon flow through each pathway are well understood and ATP synthesis is clearly via a chemiosmotic mechanism (12, 49). However, the understanding of electron transport coupled to generation of the proton gradient is both incomplete and paradoxical.

A few acetotrophic methanogens such as Methanosarcina barkeri and

Methanosarcina mazei also obtain energy for growth by oxidizing H2 and reducing CO2.

In these species, the membrane-bound Ech hydrogenase complex accepts electrons from ferredoxin (13, 32) translocating protons from the cytoplasm to outside the membrane and producing H2 (31, 50, 51). The production of H2 implies re-oxidation with transfer of electrons to HdrDE. A hypothesis has been advanced wherein H2 is re-oxidized by the membrane-bound F420 non-reducing hydrogenase that transfers electrons to methanophenazine (MP), a quinone-like electron carrier, that donates electrons to HdrDE

(20, 51). The genome of Methanosarcina mazei contains operons encoding three copies of this hydrogenase called vho, vht and vhx (8, 9). Transcription of vho is observed during growth on H2 + CO2 or acetate whereas vht is transcribed during growth with H2 +

CO2 but not with acetate (8, 9) consistent with a role for the Vho hydrogenase during growth with acetate. A function for Vhx hasn’t been established.

Most acetotrophic Methanosarcina species described are unable to oxidize H2 and 106 reduce CO2 to methane (17) which include Methanosarcina acetivorans (42) and

Methanosarcina thermophila (53). Electron transport during growth on acetate is least understood for these species. Although encoding Cdh, the genome of M. acetivorans does not encode an Ech hydrogenase (14, 34). The genome encodes genes (vhtGACD and vhxGAC) homologous to genes encoding the Vho hydrogenase of M. mazei, although these genes are not expressed (17). However, acetate-grown cells synthesize a six-subunit

Rnf complex with high identity to membrane-bound Rnf complexes in the domain

Bacteria hypothesized to couple electron transport to translocation of sodium or protons

(25, 26, 39). The Rnf complex is not encoded in the sequenced genomes of species capable of growth by oxidation of H2 and reduction of CO2 to methane (10, 28). Thus, the Rnf complex of M. acetivorans was hypothesized to accept electrons from ferredoxin and function in the electron transport chain during growth with acetate (26). The Rnf and Cdh complexes from M. acetivorans have not been purified and characterized; however, the Cdh from M. thermophila has been purified and shown to reduce a ferredoxin purified from acetate-grown cells (5, 46, 47). Furthermore, membranes from acetate-grown cells are reported to have robust ferredoxin: CoM-S-S-CoB oxidoreductase activity (35). The MP analog 2-hydroxyphenazine is an electron donor to HdrDE of acetate-grown M. thermophila implicating MP in the electron transport pathway of this species (33). However, the mechanism of electron transport between ferredoxin and MP is unknown.

Herein is described a ferredoxin isolated from acetate-grown M. acetivorans that accepts electrons from Cdh and donates to the RnfB subunit of the Rnf complex supporting a role for Rnf in electron transport. Evidence is also presented for 107 involvement of cytochrome c for which the encoding gene is co-transcribed with genes encoding the Rnf complex (26). Finally, evidence is presented supporting a function for

MP in mediating electron transfer between Rnf and the heterodisulfide reductase.

3.3 Methods and materials

Materials

CoM-S-S-CoB was a kind gift of Dr. Jan Keltjens.

N-dodecyl-β-D-maltopyranoside was purchased from Anatrace, Inc. (Maumee, OH).

2-hydroxyphenazine was custom synthesized by Sigma-Aldrich (St. Louis, MO). All other chemicals were purchased from Sigma-Aldrich or VWR International (West

Chester, PA). All chromatography columns, resins and pre-packed columns were purchased from GE Healthcare (Waukesha, WI). The RnfB subunit of the Rnf complex was overproduced in Escherichia coli and purified as described (44).

Preparation of cell extract and membranes

M. acetivorans (42) was cultured with acetate as described previously (43) and the cell paste was frozen at -80C. All solutions were O2-free and manipulations were performed anaerobically in an inert atmosphere. When possible, an anaerobic chamber

(Coy Manufacturing, Ann Arbor, MI) containing 95% N2 and 5% H2 was used. Frozen cells were thawed, re-suspended (1 g wet weight/ml buffer) in 50 mM MOPS buffer (pH 108 6.8) containing 10% (v/v) ethylene glycol and passed twice through a French pressure cell at 6.9 x 103 kPa. The lysate was centrifuged at 7,200 x g for 15 min to pellet cell debris and unbroken cells.

Membranes were purified from the cell extract using a discontinuous sucrose gradient comprised of 2 ml 70% sucrose, 4 ml 30% sucrose and 1.5 ml 20% sucrose contained in 50 mM MOPS buffer (pH 6.8). A 2 ml volume of cell extract was overlaid on the gradient and centrifuged at 200,000 x g for 2 h in a Beckman type 50 Ti rotor.

The brown band containing membranes at the 30% and 70% sucrose interface was collected and stored at -80C until use.

Purification of the αε component of the CO dehydrogenase/acetyl-CoA synthase complex (CdhAE)

Crude cell extract of acetate-grown M. acetivorans was centrifuged at 200,000 × g for 2 h to pellet the membrane fraction. The supernatant solution (200 mg protein) containing the soluble fraction was immediately loaded onto a Q-Sepharose FF column

(50 ml bed volume) equilibrated with 50 mM MOPS (pH 6.8). The column was developed with 500 ml of a 0-1.0 M NaCl linear gradient. Each 10 ml fraction was assayed for CO dehydrogenase activity as previously described (48). The pooled fractions from the peak with the highest specific activity were concentrated 10-fold with a Vivacell 70 protein concentrator equipped with a 10-kDa cut off membrane (Sartorius

Group, Göttingen, Germany). A 1.0 M solution of (NH4)2SO4 contained in 50 mM 109 MOPS (pH 6.8) was added to the concentrated protein solution to final concentration of

900 mM and loaded onto a Phenyl-Sepharose FF (low sub) column (20-ml bed volume) equilibrated with 50 mM MOPS buffer (pH 6.8) containing 1.0 M (NH4)2SO4. The column was developed with 100 ml of a 1.0-0.0 M (NH4)2SO4 decreasing linear gradient.

Fractions from the peak of CO dehydrogenase activity were pooled and concentrated followed by addition of a volume of 50 mM MOPS (pH 6.8) to lower the (NH4)2SO4 concentration to below 100 mM and then loaded on a HiTrap Q-Sepharose HP column (5 ml bed volume) equilibrated with 50 mM MOPS buffer (pH 6.8). The column was developed with 50 ml of a 0-1.0 M NaCl linear gradient. The peak containing CO dehydrogenase activity that eluted at approximately 0.3 M NaCl was collected and stored at -80C until use.

Purification of ferredoxin

Ferredoxin was assayed by coupling CO oxidation by CdhAE to the reduction of

-1 -1 metronidazole and followed by the decrease in A320 (320 = 9300 M cm ) similar to that described previously (46). The reaction mixture (100 µl) contained 100 μM metronidazole and 1-3 µg CdhAE in 50 mM Tris buffer (pH 8.0) to which 1-10 µl of the column fraction was added. The reaction was contained in an anaerobic cuvette flushed with 100% CO. The activity of ferredoxin was determined by the rate of decrease of absorbance at 320 nm, which indicates the reduction of metronidazole. One unit of activity was the amount that reduced 1 µmol of metronidazole per minute. 110 The soluble fraction of cell extract from acetate-grown M. acetivorans was loaded onto a Q-sepharose FF column (20 ml bed volume) equilibrated with 50 mM MOPS (pH

6.8) containing 10% (v/v) ethylene glycol. The column was developed with 200 ml of a

0-1.0 M linear NaCl gradient. The fraction with the highest activity was then diluted with 50 mM MOPS buffer (pH 6.8) containing 10% (v/v) ethylene glycol and loaded on a

Mono Q column (1.7 ml bed volume) to which 10 ml of a 0-1.0 M NaCl linear gradient was applied. The fraction containing ferredoxin that eluted at 600 mM NaCl was loaded on a Sephadex G-75 gel filtration column (100 ml bed volume) and developed with 50 mM MOPS (pH 6.8) containing 10% (v/v) ethylene glycol and 150 mM NaCl. The peak containing the purified ferredoxin was concentrated to A402 > 0.2 with the Vivacell

70 protein concentrator equipped with a 5-kDa cut off membrane and stored at -80C until use. The protein concentration was estimated by the ratio of absorbance at 230 and

260 nm as described (22).

Analytical

All protein concentrations except for ferredoxin were determined by the bicinchoninic acid assay (22) using the reagent from Thermo Scientific, Inc.. The

DTNB assay was performed similarly as previously described (35). The DTNB buffer used in this assay is 25 mM sodium acetate, 1 mM DTNB. All assays in this study were performed anaerobically with vacuum degassed solutions contained in sealed cuvettes with the indicated atmosphere and at room temperature. 111 3.4 Results

3.4.1 Purification of the CdhAE component of the CO dehydrogenase/acetyl-CoA complex (Cdh) of M. acetivorans

The genome of M. acetivorans is annotated with duplicate gene clusters (14), each encoding five subunits of the Cdh complex (CdhABCDE) homologous to the complex characterized from Methanosarcina thermophila with 77-95% sequence identity (15, 29,

30). Previous proteomic analyses identified subunits CdhA, CdhB and CdhC from one cluster (MA1012-16) and CdhA, CdhB CdhC and CdhE from the other (MA3860-65) in acetate-grown cells (25). The methyl viologen-dependent CO dehydrogenase specific activity of the protein purified from acetate-grown cells was 24.1 U/mg, in range of the

23-72 U/mg reported for the αε (CdhAE) sub-complex of Cdh from M. thermophila that reduces ferredoxin (1). Native PAGE showed one primary band. SDS-PAGE identified two bands with molecular masses of 16 kDa and 85 kDa consistent with the predicted values for the α and ε subunits encoded in the genome. Mass spectrometry of these two protein bands identified α and ε subunits encoded by both Cdh gene clusters consistent with proteomic analyses (25).

3.4.2 Properties of the ferredoxin purified from acetate-grown M. acetivorans

The genome of M. acetivorans is annotated with nine genes encoding ferredoxins, however, mass spectrometry analysis of the ferredoxin purified from acetate-grown M. 112 acetivorans detected only one protein identified as the product of MA0431 (Fig. 3-1) (7).

Phylogenetic analysis of the annotated ferredoxins in M. acetivorans and identified ferredoxins from other Methanosarcina species revealed the product of MA0431 is closely related to the 2 x 4Fe-4S ferredoxin that functions in the aceticlastic pathway of

M. thermophila (5, 6, 46, 47) and the ferredoxin up-regulated in acetate- versus methanol-grown Methanosarcina mazei (21). These three ferredoxins contain two

CX2CX2CX3CP motifs typical of 2 x 4Fe-4S ferredoxins and share high identity (Fig. 3-4) within a distinct clade (Fig. 3-3). These observations suggest a specific function for this clade (Fig. 3-3) in the electron transport pathway of acetate-utilizing Methanosarcina species.

The UV-visible absorption spectrum of the as-purified protein was typical of ferredoxins with an absorption maximum at 402 nm that decreased upon reduction with dithionite (Fig. 3-2). The A395/A295 ratio was 0.74 which is in agreement with the

A395/A295 ratio of 0.80 - 0.88 reported for the ferredoxin purified from acetate-grown cells of M. thermophila (47). Purification of the ferredoxin from M. acetivorans was monitored by coupling the oxidation of CO with CdhAE to the ferredoxin-dependent reduction of metronidazole as described for the ferredoxin purified from acetate-grown M. thermophila (46). Figure 3-5 shows CO-dependent reduction of the purified M. acetivorans ferredoxin catalyzed by the CdhAE component of the Cdh complex, suggesting ferredoxin is the electron acceptor of Cdh.

113

Figure 3-1 Mass spectrometry of ferredoxin from M. acetivorans. 114

Figure 3-2 UV-visible absorption spectra of purified ferredoxin. As-purified (), dithionite reduced (…). The protein concentration was 20 µM.

Figure 3-3 Phylogenetic analysis of ferredoxins. The M. mazei and M. acetivorans sequences, labeled with the prefix MA, were derived from the CMR database (7). The M. barkeri (19) and M. thermophila sequences (5) are published. The sequence of the 2 x 4Fe-4S Clostridium pasteurianum ferredoxin is published (16) and the sequence of the 2Fe-2S Spinacia oleracea ferredoxin was obtained from the NCBI database (accession number O04683). The tree was constructed by the neighbor-joining method with the MEGA4 program (45). Bootstrap values are shown at the nodes. Bar, evolutionary distance of 0.2. 115

Figure 3-4 Sequence alignment of ferredoxins from Methanosarcina species. See Figure 3-3 caption for the origins of sequences. MA, Methanosarcina acetivorans; MM, Methanosarcina mazei;

M.t. Methanosarcina thermophila. Motifs predicted to ligate two 4Fe-4S clusters are highlighted. The alignment was performed with ClustalX2 (23).

Figure 3-5 Reduction of ferredoxin by CdhAE. The 70 μl reaction mixture consisted of 2.2 μg

(final concentration 31μg /ml) of CdhAE and 28 μM (final concentration) of ferredoxin contained in 50 mM MOPS buffer (pH 6.8) under 1 atm CO. The reaction was initiated with CdhAE. A, complete reaction mixture. B, reaction mixture minus CdhAE. C, reaction mixture minus ferredoxin. The reduction of ferredoxin was followed by the decrease in absorbance at 402 nm.

116 3.4.3 Role of ferredoxin in the membrane-bound electron transport chain

Opposed to H2/CO2-utilizing strains of Methanosarcina, an Rnf complex is proposed to function in the membrane-bound electron transport chain of M. acetivorans and ferredoxin is the electron donor to the complex (26). Thus, a role for the ferredoxin in the electron transport chain terminating with reduction of CoM-S-S-CoB was investigated in a system containing sucrose gradient-purified membranes and plant ferredoxin-NADP+ reductase (FNR) to regenerate reduced ferredoxin. The

CO-dependent reduction of ferredoxin with Cdh was not used to avoid binding of CO to the high spin hemes in cytochrome c and potentially inhibiting membrane-bound electron transport. The NADPH: CoM-S-S-CoB oxidoreductase activity was monitored by detecting the sulfhydryl groups of HS-CoM and HS-CoB (Fig. 3-6). No significant activity was detected when each component of the reaction mixture was deleted. The dependence of the activity on membranes and the concentration of ferredoxin purified from acetate-grown M. acetivorans indicated a role for the ferredoxin in transfer of electrons from Cdh to the membrane-bound electron transport chain that includes heterodisulfide reductase.

3.4.4 Role of Rnf in the membrane-bound electron transport chain

It has been postulated that the RnfB subunit (MA0664) of the Rnf complex is the point of entry for electrons to the complex donated by ferredoxin (26). Biochemical evidence for this role of RnfB and the Rnf complex in the membrane-bound electron transport chain of acetate-grown M. acetivorans was investigated with RnfB. RnfB was 117 reduced in the presence of CO, CdhAE and ferredoxin (Fig. 3-7). Omission of ferredoxin from the reaction mixture severely reduced the rate suggesting ferredoxin is the electron donor to RnfB and supporting a role for the Rnf complex in the membrane-bound electron transport chain of acetate-grown M. acetivorans.

Figure 3-6 Ferredoxin: CoM-S-S-CoB oxidoreductase activity of membranes. The 100 μl reaction mixture consisted of 20 mM NADPH, 2 μg FNR, the membrane fraction of acetate-grown cells (60

μg protein), 1.1 mM CoM-S-S-CoB and the indicated concentrations of ferredoxin contained in 50 mM

MOPS buffer (pH 6.8). Total thiols were determined by the DTNB assay. Symbols: ()1.2 μM ferredoxin, () 0.6 μM ferredoxin, () 0.3 μM ferredoxin, () minus ferredoxin. 118

Figure 3-7 Reduction of RnfB by ferredoxin. The 80 μl reaction mixture consisted of 18 μg

CdhAE, 1.3 μM ferredoxin and 34 μM RnfB contained in 50 mM MOPS buffer (pH 6.8). The mixture was equilibrated with 1 atm CO and initiated by addition of CdhAE. The reduction of RnfB was followed by the change of absorbance at 420 nm. A, complete reaction mixture; B, minus CdhAE; C, 1 atm N2; D, minus ferredoxin; E, minus RnfB.

3.4.5 Role of cytochrome c in the membrane-bound electron transport chain

Two ORF's (MA0658 and MA0665) are co-transcribed with the six genes

MA0659-0664 homologous to those encoding six-subunit Rnf complexes from the

Bacteria domain (26). ORF MA0658 encodes a multi-heme cytochrome c that dominates the visible absorbance spectrum of membranes from acetate-grown M. acetivorans with the major peak centered at 554 nm (26) (Fig. 3-8). Absorbance at 554 nm increased on incubation of the membrane fraction with the reduced ferredoxin regenerating system comprised of NADPH, H+ and FNR indicating reduction of 119 cytochrome c that was dependent on ferredoxin (Fig. 3-8). Interpretation of these results is complicated by the ability of ferredoxin to directly reduce purified cytochrome c and the uncertain periplasmic versus cytoplasmic location on the purified membranes that potentially contain an unknown fraction of inverted vesicles. Nonetheless, addition of CoM-S-S-CoB oxidized the reduced cytochrome (Fig. 3-9) indicating that it is a component of the membrane-bound electron transport chain terminating with reduction of the heterodisulfide. Oxidation of the cytochrome was incomplete, a result consistent with a periplasmic location reduced by ferredoxin in inverted closed vesicles via the electron transport chain and directly reduced by ferredoxin in closed right-side-out vesicles wherein CoM-S-S-CoB is unable to access the heterodisulfide reductase.

Indeed, the purified ferredoxin reduced bovine heart cytochrome c.

3.4.6 Role of methanophenazine in the membrane-bound electron transport chain

Methanophenazine (MP) is a quinone-like two-electron carrier implicated in mediation of electron transfer to the heterodisulfide reductase of acetate- and

H2/CO2-utilizing Methanosarcina species (33). The soluble analog of MP,

2-hydroxyphenazine, has been used to investigate the role of MP in methanogens (2, 33).

The reduced ferredoxin regenerating system comprised of CO, CdhAE and ferredoxin reduced the MP analog in the presence of membranes from acetate-grown cells that was re-oxidized upon the addition of CoM-S-S-CoB (Fig. 3-10). The re-oxidation generated a total of 293 μM free thiol groups in this representative experiment, a result that is in approximate agreement with the observed oxidation of 106 μM 2-hydroxyphenazine. 120 Assuming a two-electron transfer from the MP analog, 212 μM free thiol groups would be expected. The difference in observed and expected is likely the result of reducing equivalents derived from reduced electron transfer components of the reaction mixture and residual CO after sparging with N2. These results indicate that MP is a component of the membrane-bound electron transport chain terminating with reduction of

CoM-S-S-CoB.

Figure 3-8 Ferredoxin-dependent reduction of membrane-bound cytochrome c. The 100 μl reaction mixture consisted of purified membranes (300 μg protein), the indicated amount of ferredoxin, 1 μg FNR and 1 mM NADPH in 50 mM MOPS (pH 6.8). The reaction was initiated by addition of FNR. The reduction of cytochrome c was followed at 554 nm. Panel A, time-course for the reduction of cytochrome c. Symbols: () 4 μM ferredoxin; (○) 0.2 μM ferredoxin; (□) minus ferredoxin; (∆) minus FNR; (●) minus NADPH. Panel B, reduced minus oxidized spectra recorded at the indicated times after initiation of the reaction containing 4 μM ferredoxin. 121

Figure 3-9 Oxidation of membrane-bound cytochrome c by CoM-S-S-CoB. The reduction of cytochrome c was performed as described in the caption to Figure 3-8. The 100 μl reaction mixture consisted of membranes (420 μg protein), 2 μM ferredoxin, 1 μg FNR and 1 mM NADPH. NADPH was added at time zero and either 0.12 mM (final concentration) CoM-S-S-CoB or an equivalent volume (1 μl) of buffer minus CoM-S-S-CoB was added (arrow). Reduction and oxidation of cytochrome c was monitored by the absorbance at 554 nm. Symbols: () addition of CoM-S-S-CoB at 8 min (arrow); () addition of buffer at 8 min (arrow).

Figure 3-10 Reduction of 2-hydroxyphenazine and membrane-catalyzed oxidation dependent on CoM-S-S-CoB. The 100 μl reaction mixture consisted of membranes (107 μg protein), 4 μM ferredoxin, 100 μM 2-hydroxyphenazine and CdhAE (40 μg) in 50 mM MOPS (pH 6.8) under 1 atm CO. The -1 reduction and oxidation of 2-hydroxyphenazine was followed by the absorbance at 475 nm (ε475 = 2.5 mM cm-1). CdhAE was added to initiate the reduction at time zero. At point A the cuvette was flushed with

100% N2 and 2 μl of MOPS buffer (pH 6.8) was added. At points B and C, 2μl of MOPS buffer (pH 6.8) containing CoM-S-S-CoB was added to the reaction reaching final concentrations of 240 and 480 μM. 122 The results implicating MP and cytochrome c in the membrane-bound electron transport chain suggests electron transfer should occur between these carriers. The MP analog 2-hydroxyphenazine re-oxidized cytochrome c when added to membranes of acetate-grown cells previously reduced with ferredoxin (Fig. 3-11). These results suggest that MP is either directly or indirectly linked to cytochrome c, a result further supporting the participation of MP and cytochrome c in the membrane-bound electron transport chain.

Figure 3-11 Oxidation of membrane-bound cytochrome c by 2-hydroxyphenazine. The 100

μl reaction mixture consisted of membranes (750 μg protein), 4 μM ferredoxin 1 mM NADPH and1 μg

FNR contained in 50 mM MOPS buffer (pH 6.8). The reduction of cytochrome c was initiated by addition of FNR. The reduction and re-oxidation was monitored at 554 nm. When fully reduced, 200

μM 2-hydroxyphenazine (2 μl) was added (arrow). Panel A, time course for the reduction and re-oxidation by 2-hydroxyphenazine added at the arrow. Panel B, reduced minus oxidized UV-visible spectra of membranes before (lower trace) and after (upper trace) addition of 2-hydroxyphenazine. 123 3.4.7 Comparative analysis of the M. thermophila genome

M. thermophila is an acetotrophic Methanosarcina species incapable of growth with

H2/CO2 (52, 53). Analysis of the genomic sequence revealed a gene cluster identical in arrangement and homologous to genes encoding the six subunits of Rnf and multi-heme cytochrome c of M. acetivorans with deduced sequence identities ranging from 86 to 98%

(Fig. 3-12). Alignments of the deduced sequences showed strict conservation of heme-binding, flavin binding and iron-sulfur binding motifs suggesting conserved functions (Appendix A). Although analysis of the genome of M. thermophila revealed homologs of the vhtGACD and vhxGAC genes from M. acetivorans (Fig. 3-13, Appendix

B), not identified were homologs of genes encoding the six-subunit Ech hydrogenase complex of acetotrophic Methanosarcina species capable of growth with H2/CO2.

Although not conclusive, these results are consistent with a role for the Rnf complex and multi-heme cytochrome c in the electron transport pathway of M. thermophila grown with acetate. Furthermore, the genome of M. thermophila contains a gene cluster (Fig.

3-14, Appendix C) homologous to genes encoding the seven subunits of the sodium/proton antiporter (Mrp) that is up-regulated in acetate- versus methanol-grown cells of M. acetivorans and absent in the sequenced genomes of acetotrophic

Methanosarcina species capable of growth with H2/CO2 (25, 26). 124

Figure 3-12 Alignment of rnf gene clusters between M. thermophila and M. acetivorans.

Numbers next to the arrows indicate deduced sequence identity

Figure 3-13 Alignment of vht and vhx gene clusters between M. thermophila and M. acetivorans. Numbers next to the arrows indicate deduced sequence identity.

125

Figure 3-14 Alignment of mrp gene clusters between M. thermophila and M. acetivorans.

Numbers next to the arrows indicate deduced sequence identity.

3.5 Discussion

Methanogens capable of growth via conversion of the methyl group of acetate to methane are divided in the ability to also grow by oxidizing H2 and reducing CO2 to methane. Here we report the first biochemical investigation of electron transport in M. acetivorans, a species incapable of growth with H2/CO2. The results provide the first biochemical evidence supporting a role for an Rnf complex in methanogens and the

Archaea domain, and a role for cytochrome c in the aceticlastic pathway of methanogens.

Figure 3-15 compares membrane-bound electron transport in the aceticlastic pathway for acetate-grown M. acetivorans with that for M. mazei and M. barkeri that in addition are capable of growth with H2/CO2. In both pathways ferredoxin is oxidized and electrons are passed to MP that donates electrons to HdrDE. In H2/CO2-utilizing species, a H2 cycling mechanism has been proposed in which the Ech hydrogenase 126

Figure 3-15 Comparison of electron transport pathways for M. mazei and M. barkeri versus M. acetivorans. Panel A: M. mazei and M. barkeri. Panel B: M. acetivorans. Ech, Ech hydrogenase; Fdr, ferredoxin reduced; Fdo, ferredoxin oxidized; Vho, Vho hydrogenase; MP, methanophenazine; HdrDE, heterodisulfide reductase; CoM-SH, coenzyme M; CoB-SH, coenzyme B; Atp, ATP synthase; MaRnf, Rnf complex from M. acetivorans; Mrp, putative sodium/proton antiporter.

complex accepts electrons from ferredoxin and translocates a proton from the cytoplasm to outside the membrane producing H2 (51). The H2 is then re-oxidized by the

MP-reducing Vho-type hydrogenase providing two additional proton translocations driving ATP synthesis. In contrast, the genome of M. acetivorans does not encode an

Ech hydrogenase (14) and although the genome contains homologs of genes encoding

Vho-type hydrogenases they are not expressed during growth with acetate (17).

Furthermore, biochemical plus genetic evidence suggests H2 is not an obligatory intermediate during growth on acetate (14, 18, 34). Instead, the biochemical results presented here indicate that M. acetivorans evolved an electron transport pathway independent of H2 involving the Rnf complex and a multi-heme cytochrome c that was previously postulated based on up-regulation of the co-transcribed encoding genes in 127 acetate- versus methanol-grown cells (25, 26). Based on the results presented here, it is not possible to conclude that the Rnf complex couples electron transport to the exchange of ions across the cytoplasmic membrane as has been proposed for Rnf homologs from the domain Bacteria (39). Based on up-regulation of the multi-subunit sodium/proton antiporter Mrp in acetate- versus methanol-grown M. acetivorans, it was hypothesized that the Rnf complex pumps sodium ions outside the cytoplasmic membrane that are exchanged for protons by the Mrp complex (26).

Further support for Rnf and cytochrome c involvement in the electron transport of acetotrophic methanogens incapable of growth with H2/CO2 resides in the genome of M. thermophila that contains a gene cluster identical to the Rnf gene cluster of M. acetivorans albeit no evidence for an Ech hydrogenase. Furthermore, the genome of M. thermophila encodes an Mrp complex shown to be up-regulated in acetate- versus methanol-grown M. acetivorans that is not present in Methanosarcina species able to grow with H2/CO2. Interestingly, genes encoding subunits of Rnf, Ech or Mrp are absent in the genome of the acetate-utilizing isolate Methanosaeta thermophila (41) that is also incapable of growth with H2/CO2 suggesting still other alternative electron transport pathways coupled to generation of ion gradients driving ATP synthesis in acetate-utilizing methanogens. The physiological significance of these diverse electron transport pathways is yet to be determined; however, it has been suggested that avoiding

H2 is advantageous to the marine isolate M. acetivorans since sulfate reducing species that dominate this environment outcompete methanogens for H2 potentially disrupting electron transport (26). It is important to note here that although M. acetivorans is incapable of growth with H2/CO2 it synthesizes all of the enzymes necessary for 128 reduction of CO2 to methane and is capable of robust growth via the CO2-reduction pathway albeit with electrons derived from the oxidation of CO (24, 37).

The results reported here showing that ferredoxin donates electrons to RnfB suggest it is the entry point for electrons to the Rnf complex. Indeed, the deduced sequence of

RnfB from M. acetivorans harbors three iron-sulfur cluster binding motifs

(CX2CX2CX3CP) and spectroscopic analyses establish the presence of multiple iron-sulfur clusters in purified RnfB from M. acetivorans consistent with an electron transfer function (44). The electron acceptor for the Rnf complex is unknown, although cytochrome c and MP are candidates. Indeed, the gene encoding the membrane-bound multi-heme cytochrome c that is up-regulated in acetate-grown cells is co-transcribed with the genes encoding the Rnf complex (26). Transcriptomic, proteomic and genetic evidence supports a role for the HdrDE heterodisulfide reductase in acetate-grown M. acetivorans (4, 25). Based on evidence that MP is the direct electron donor to HdrDE in

M. thermophila (33, 40), we propose that MP is also the electron donor to HdrDE in M. acetivorans and contributes to a proton gradient (high outside). Thus, it follows that cytochrome c most likely mediates electron transfer between the Rnf complex and MP consistent with the results reported here showing oxidation of cytochrome c on addition of 2-hydroxyphenazine to ferredoxin-reduced membranes. Based on the proposed function for Rnf complexes in the Bacteria domain, it is postulated that the Rnf complex of M. acetivorans either pumps protons or sodium from the cytoplasm to outside the membrane. The A1A0 ATP synthase is up-regulated in acetate- versus methanol-grown

+ M. acetivorans (26) and it is proposed that H is the coupling ion in the A1A0 ATP synthase of the acetotroph M. mazei (36). These results are consistent with a proton 129 gradient driving ATP synthesis in these species. Further, the putative sodium-translocating F1F0 ATP synthase is dispensable for growth of M. acetivorans with acetate (38). Thus, if sodium is translocated by the Rnf complex, a sodium/proton antiporter must function to couple ion translocation by Rnf to ATP synthesis which is a postulated function of the Mrp complex reported to be up-regulated in acetate- versus methanol-grown M. acetivorans (25, 26).

The sequenced genomes of all Methanosarcina species are annotated with multiple genes encoding ferredoxins including nine in M. acetivorans, a result raising the question of specific roles. The results presented here suggest a specific clade functions as the electron acceptor of the Cdh complex during acetotrophic growth of Methanosarcina species. A phylogenetically distant ferredoxin (MM0760) is up-regulated in methanol- versus acetate-grown M. mazei consistent with evolution of ferredoxins evolved for specific functions in this species (21). The sequence of a ferredoxin purified from an acetate-grown M. barkeri strain has not been reported.

3.6 Acknowledgements

This work was supported by the National Science Foundation. We thank Dr. Jan

Keltjens for generously supplying CoM-S-S-CoB and the Penn State Hershey Core

Research Facilities for mass spectrometry analyses. 130 3.7 References

1. Abbanat, D. R., and J. G. Ferry. 1991. Resolution of component proteins in an

enzyme complex from Methanosarcina thermophila catalyzing the synthesis or

cleavage of acetyl-CoA. Proc. Natl. Acad. Sci. USA 88:3272-3276.

2. Abken, H.-J., M. Tietze, J. Brodersen, S. Baumer, U. Beifuss, and U.

Deppenmeier. 1998. Isolation and characterization of methanophenazine and the

function of phenazines in membrane-bound electron transport of Methanosarcina

mazei Go1. J. Bacteriol. 180:2027-2032.

3. Bloom, A. A., P. I. Palmer, A. Fraser, D. S. Reay, and C. Frankenberg. 2010.

Large-scale controls of methanogenesis inferred from methane and gravity

spaceborne data. Science 327:322-325.

4. Buan, N. R., and W. W. Metcalf. 2010. Methanogenesis by Methanosarcina

acetivorans involves two structurally and functionally distinct classes of

heterodisulfide reductase. Mol. Microbiol. 75:843-853.

5. Clements, A. P., and J. G. Ferry. 1992. Cloning, nucleotide sequence, and

transcriptional analyses of the gene encoding a ferredoxin from Methanosarcina

thermophila. J. Bacteriol. 174:5244-5250.

6. Clements, A. P., L. Kilpatrick, W.-P. Lu, S. W. Ragsdale, and J. G. Ferry. 1994.

Characterization of the iron-sulfur clusters in ferredoxin from acetate-grown

Methanosarcina thermophila. J. Bacteriol. 176:2689-2693.

7. Comprehensive, T., and M. Resource. J. Craig Venter Institute,

. 131 8. Deppenmeier, U. 1995. Different structure and expression of the operons encoding

the membrane-bound hydrogenases from Methanosarcina mazei Gö1. Arch.

Microbiol. 164:370-376.

9. Deppenmeier, U., M. Blaut, S. Lentes, C. Herzberg, and G. Gottschalk. 1995.

Analysis of the vhoGAC and vhtGAC operons from Methanosarcina mazei strain

Go1, both encoding a membrane-bound hydrogenase and a cytochrome b. Eur. J.

Biochem. 227:261-269.

10. Deppenmeier, U., A. Johann, T. Hartsch, R. Merkl, R. A. Schmitz, R.

Martinez-Arias, A. Henne, A. Wiezer, S. Baumer, C. Jacobi, H. Bruggemann,

T. Lienard, A. Christmann, M. Bomeke, S. Steckel, A. Bhattacharyya, A.

Lykidis, R. Overbeek, H. P. Klenk, R. P. Gunsalus, H. J. Fritz, and G.

Gottschalk. 2002. The genome of Methanosarcina mazei: evidence for lateral

gene transfer between Bacteria and Archaea. J. Mol. Microbiol. Biotechnol.

4:453-61.

11. Ferry, J. G. 2008. Acetate-based methane production, p. 155-170. In J. D. Wall, C.

S. Harwood, and A. Demain (ed.), Bioenergy. ASM press, Washington.

12. Ferry, J. G. 2010. How to make a living exhaling methane. Annu. Rev. Microbiol.

64:453-473.

13. Fischer, R., and R. K. Thauer. 1990. Ferredoxin-dependent methane formation

from acetate in cell extracts of Methanosarcina barkeri (strain MS). FEBS Lett.

269:368-372.

14. Galagan, J. E., C. Nusbaum, A. Roy, M. G. Endrizzi, P. Macdonald, W.

FitzHugh, S. Calvo, R. Engels, S. Smirnov, D. Atnoor, A. Brown, N. Allen, J. 132 Naylor, N. Stange-Thomann, K. DeArellano, R. Johnson, L. Linton, P.

McEwan, K. McKernan, J. Talamas, A. Tirrell, W. Ye, A. Zimmer, R. D.

Barber, I. Cann, D. E. Graham, D. A. Grahame, A. M. Guss, R. Hedderich, C.

Ingram-Smith, H. C. Kuettner, J. A. Krzycki, J. A. Leigh, W. Li, J. Liu, B.

Mukhopadhyay, J. N. Reeve, K. Smith, T. A. Springer, L. A. Umayam, O.

White, R. H. White, E. C. de Macario, J. G. Ferry, K. F. Jarrell, H. Jing, A. J.

Macario, I. Paulsen, M. Pritchett, K. R. Sowers, R. V. Swanson, S. H. Zinder,

E. Lander, W. W. Metcalf, and B. Birren. 2002. The genome of M. acetivorans

reveals extensive metabolic and physiological diversity. Genome Res. 12:532-42.

15. Grahame, D. A., S. Gencic, and E. Demoll. 2005. A single operon-encoded form of

the acetyl-CoA decarbonylase/synthase multienzyme complex responsible for

synthesis and cleavage of acetyl-CoA in Methanosarcina thermophila. Arch.

Microbiol. 184:1-9.

16. Graves, M. C., G. T. Mullenbach, and J. C. Rabinowitz. 1985. Cloning and

nucleotide sequence determination of the Clostridium pasteurianum ferredoxin

gene. Proc. Natl. Acad. Sci. USA 82:1653-1657.

17. Guss, A. M., G. Kulkarni, and W. W. Metcalf. 2009. Differences in hydrogenase

gene expression between Methanosarcina acetivorans and Methanosarcina

barkeri. J. Bacteriol. 191:2826-2833.

18. Guss, A. M., B. Mukhopadhyay, J. K. Zhang, and W. W. Metcalf. 2005. Genetic

analysis of mch mutants in two Methanosarcina species demonstrates multiple

roles for the methanopterin-dependent C-1 oxidation/reduction pathway and 133

differences in H2 metabolism between closely related species. Mol. Microbiol.

55:1671-1680.

19. Hausinger, R. P., I. Moura, J. J. G. Moura, A. V. Xavier, M. H. Santos, J.

LeGall, and J. B. Howard. 1982. Amino acid sequence of a 3Fe:3S ferredoxin

from the "archaebacterium" Methanosarcina barkeri (DSM 800). J. Biol. Chem.

257:14192-14197.

20. Hedderich, R., O. Klimmek, A. Kroger, R. Dirmeier, M. Keller, and K. O.

Stetter. 1999. Anaerobic respiration with elemental sulfur and with disulfides.

FEMS Microbiol. Rev. 104:353-381.

21. Hovey, R., S. Lentes, A. Ehrenreich, K. Salmon, K. Saba, G. Gottschalk, R. P.

Gunsalus, and U. Deppenmeier. 2005. DNA microarray analysis of

Methanosarcina mazei Go1 reveals adaptation to different methanogenic

substrates. Mol. Genet. Genomics 273:225-239.

22. Kalb, V. F., and R. W. Bernlohr. 1977. A new spectrophotometric assay for protein

in cell extracts. Anal. Biochem. 82:362-371.

23. Larkin, M. A., G. Blackshields, N. P. Brown, R. Chenna, P. A. McGettigan, H.

McWilliam, F. Valentin, I. M. Wallace, A. Wilm, R. Lopez, J. D. Thompson,

T. J. Gibson, and D. G. Higgins. 2007. Clustal W and Clustal X version 2.0.

Bioinformatics 23:2947-2948.

24. Lessner, D. J., L. Li, Q. Li, T. Rejtar, V. P. Andreev, M. Reichlen, K. Hill, J. J.

Moran, B. L. Karger, and J. G. Ferry. 2006. An unconventional pathway for

reduction of CO2 to methane in CO-grown Methanosarcina acetivorans revealed

by proteomics. Proc. Natl. Acad. Sci. U.S.A. 103:17921-17926. 134 25. Li, L., Q. Li, L. Rohlin, U. Kim, K. Salmon, T. Rejtar, R. P. Gunsalus, B. L.

Karger, and J. G. Ferry. 2007. Quantitative proteomic and microarray analysis

of the archaeon Methanosarcina acetivorans grown with acetate versus methanol.

J. Proteome. Res. 6:759-771.

26. Li, Q., L. Li, T. Rejtar, D. J. Lessner, B. L. Karger, and J. G. Ferry. 2006.

Electron transport in the pathway of acetate conversion to methane in the marine

archaeon Methanosarcina acetivorans. J. Bacteriol. 188:702-710.

27. Liu, Y., and W. B. Whitman. 2008. Metabolic, phylogenetic, and ecological

diversity of the methanogenic archaea. Ann. N. Y. Acad. Sci. 1125:171-189.

28. Maeder, D. L., I. Anderson, T. S. Brettin, D. C. Bruce, P. Gilna, C. S. Han, A.

Lapidus, W. W. Metcalf, E. Saunders, R. Tapia, and K. R. Sowers. 2006. The

Methanosarcina barkeri genome: Comparative analysis with Methanosarcina

acetivorans and Methanosarcina mazei reveals extensive rearrangement within

methanosarcinal genomes. J. Bacteriol. 188:7922-7931.

29. Maupin-Furlow, J., and J. G. Ferry. 1996. Characterization of the cdhD and cdhE

genes encoding subunits of the corrinoid iron-sulfur enzyme of the CO

dehydrogenase complex from Methanosarcina thermophila. J. Bacteriol.

178:340-346.

30. Maupin-Furlow, J. A., and J. G. Ferry. 1996. Analysis of the CO

dehydrogenase/acetyl-coenzyme A synthase operon of Methanosarcina

thermophila. J. Bacteriol. 178:6849-6856. 135 31. Meuer, J., S. Bartoschek, J. Koch, A. Kunkel, and R. Hedderich. 1999.

Purification and catalytic properties of Ech hydrogenase from Methanosarcina

barkeri. Eur. J. Biochem. 265:325-335.

32. Meuer, J., H. C. Kuettner, J. K. Zhang, R. Hedderich, and W. W. Metcalf. 2002.

Genetic analysis of the archaeon Methanosarcina barkeri Fusaro reveals a central

role for Ech hydrogenase and ferredoxin in methanogenesis and carbon fixation.

Proc. Natl. Acad. Sci. U. S. A. 99:5632-5637.

33. Murakami, E., U. Deppenmeier, and S. W. Ragsdale. 2001. Characterization of

the intramolecular electron transfer pathway from 2-hydroxyphenazine to the

heterodisulfide reductase from Methanosarcina thermophila. J. Biol. Chem.

276:2432-2439.

34. Nelson, M. J. K., and J. G. Ferry. 1984. Carbon monoxide-dependent methyl

coenzyme M methylreductase in acetotrophic Methanosarcina spp. J. Bacteriol.

160:526-532.

35. Peer, C. W., M. H. Painter, M. E. Rasche, and J. G. Ferry. 1994. Characterization

of a CO:heterodisulfide oxidoreductase system from acetate-grown

Methanosarcina thermophila. J. Bacteriol. 176:6974-6979.

36. Pisa, K. Y., C. Weidner, H. Maischak, H. Kavermann, and V. Muller. 2007. The

+ + coupling ion in the methanoarchaeal ATP synthases: H vs. Na in the A1A0 ATP

synthase from the archaeon Methanosarcina mazei Go1. FEMS Microbiol. Lett.

277:56-63. 136 37. Rother, M., E. Oelgeschlager, and W. M. Metcalf. 2007. Genetic and proteomic

analyses of CO utilization by Methanosarcina acetivorans. Arch. Microbiol.

188:463-472.

38. Saum, R., K. Schlegel, B. Meyer, and V. Muller. 2009. The F1FO ATP synthase

genes in Methanosarcina acetivorans are dispensable for growth and ATP

synthesis. FEMS Microbiol. Lett. 300:230-236.

39. Schmidt, S., E. Biegel, and V. Muller. 2009. The ins and outs of Na+ bioenergetics

in Acetobacterium woodii. Biochim. Biophys. Acta. 1787:691-696.

40. Simianu, M., E. Murakami, J. M. Brewer, and S. W. Ragsdale. 1998. Purification

and properties of the heme- and iron-sulfur- containing heterodisulfide reductase

from Methanosarcina thermophila. Biochemistry 37:10027-10039.

41. Smith, K. S., and C. Ingram-Smith. 2007. Methanosaeta, the forgotten

methanogen? Trends Microbiol. 7:150-155.

42. Sowers, K. R., S. F. Baron, and J. G. Ferry. 1984. Methanosarcina acetivorans sp.

nov., an acetotrophic methane-producing bacterium isolated from marine

sediments. Appl. Environ. Microbiol. 47:971-978.

43. Sowers, K. R., M. J. K. Nelson, and J. G. Ferry. 1984. Growth of acetotrophic,

methane-producing bacteria in a pH auxostat. Curr. Microbiol. 11:227-230.

44. Suharti, S. De Vries, and J. G. Ferry. 2010. Characterization of the RnfB Subunit

of the Rnf Complex from Methanosarcina acetivorans. J. Bacteriol.:submitted.

45. Tamura, K., J. Dudley, M. Nei, and S. Kumar. 2007. MEGA4: molecular

evolutionary genetics analysis (MEGA) software version 4.0. Mol. Biol. Evol.

24:1596-1599. 137 46. Terlesky, K. C., and J. G. Ferry. 1988. Ferredoxin requirement for electron

transport from the carbon monoxide dehydrogenase complex to a

membrane-bound hydrogenase in acetate-grown Methanosarcina thermophila. J.

Biol. Chem. 263:4075-4079.

47. Terlesky, K. C., and J. G. Ferry. 1988. Purification and characterization of a

ferredoxin from acetate-grown Methanosarcina thermophila. J. Biol. Chem.

263:4080-4082.

48. Terlesky, K. C., M. J. K. Nelson, and J. G. Ferry. 1986. Isolation of an enzyme

complex with carbon monoxide dehydrogenase activity containing a corrinoid and

nickel from acetate-grown Methanosarcina thermophila. J. Bacteriol.

168:1053-1058.

49. Thauer, R. K., A. K. Kaster, H. Seedorf, W. Buckel, and R. Hedderich. 2008.

Methanogenic archaea: ecologically relevant differences in energy conservation.

Nat. Rev. Microbiol. 6:579-591.

50. Welte, C., V. Kallnik, M. Grapp, G. Bender, S. Ragsdale, and U. Deppenmeier.

2010. Function of Ech hydrogenase in ferredoxin-dependent membrane-bound

electron transport in Methanosarcina mazei. J. Bacteriol. 192:674-680.

51. Welte, C., C. Kratzer, and U. Deppenmeier. 2010. Involvement of Ech

hydrogenase in energy conservation of Methanosarcina mazei. FEBS J.

277:3396-3403.

52. Zinder, S. H., and R. A. Mah. 1979. Isolation and characterization of a

thermophilic strain of Methanosarcina unable to use H2-CO2 for methanogenesis.

Appl. Environ. Microbiol. 38:996-1008. 138 53. Zinder, S. H., K. R. Sowers, and J. G. Ferry. 1985. Methanosarcina thermophila

sp. nov., a thermophilic, acetotrophic, methane-producing bacterium. Int. J. Syst.

Bacteriol. 35:522-523 139

Chapter 4

Characterization of a novel electron transport protein from

Methanosarcina acetivorans

4.1 Abstract

Comparison of ferredoxin: CoM-S-S-CoB oxidoreductase activity in membrane and soluble fractions of acetate-grown M. acetivorans revealed significant soluble activity, suggesting the presence of a soluble ferredoxin: CoM-S-S-CoB electron transport chain in addition to previously identified membrane-bound ferredoxin: CoM-S-S-CoB electron transport chain. Previous proteomic and genetic evidence suggested a fused protein of heterodisulfide reductase A subunit (HdrA) and F420 non-reducing hydrogenase D subunit

(MvhD), now named Etp, is involved in this soluble electron transport. Etp was overexpressed in Escherichia coli, purified, reconstituted and characterized. This iron-sulfur cluster containing protein shows an activity of accepting electrons from CO mediated by CO dehydrogenase (Cdh), ferredoxin and unknown soluble factors in acetate-grown M. acetivorans, suggesting Etp is a member of a previously unidentified soluble electron transport pathway. The soluble ferredoxin: CoM-S-S-CoB transport 140 chain was further characterized, suggesting the participation of Cdh, ferredoxin,

CoM-S-S-CoB and an unidentified heterodisulfide reductase. Evidence supporting a role of Etp in this electron transport chain was also observed. A model is constructed based on these findings that the acetotroph M. acetivorans utilizes this soluble pathway to bypass certain energy coupling sites in favor of completing metabolism cycle to more energy conservation under substrate-energy stressed situations.

4.2 Introduction

In the CO2 reduction methanogenesis pathway CO2 is reduced to the methyl level with electrons derived from the oxidation of either formate or H2 (27). The methyl group is transferred to HS-CoM producing CH3-S-CoM that is reductively demethylated to methane with electrons donated by HS-CoB (27). As a consequence of this reaction the heterodisulfide CoB-S-S-CoM is formed that is reduced with electrons derived from either formate or H2 to the sulfhydryl forms of the cofactors in a reaction catalyzed by heterodisulfide reductase (Hdr) that is highly exergonic with potential for generation of an ion gradient driving ATP synthesis (11). In obligate CO2 reducing species, the cytoplasmic three-subunit HdrABC is in a tight complex with either the MvhAGD hydrogenase (28) or MvhAGD hydrogenase plus formate dehydrogenase (4). It is proposed that the flavin-containing HdrA subunit accepts electrons from the MvhD 141 subunit that in turn transfers electrons to the catalytic HdrBC subunits (30). It is hypothesized that the flavin of HdrA also donates electrons to a 2 x 4Fe-4S ferredoxin that is the electron donor for the first reaction in CO2 reduction to the methyl level which is an endergonic reaction (28, 30). Thus, flavin based electron bifurcation is postulated to drive the endergonic CO2 reduction reaction by coupling to the exergonic reduction of

CoB-S-S-CoM (29). A similar mechanism is also postulated for the HdrABC/formate dehydrogenase complex (4). However, this hypothesis does not allow generation of the ion gradient that drives ATP synthesis.

In the aceticlastic pathway acetyl-CoA is cleaved by the CO dehydrogenase/acetyl-CoA synthase (Cdh) enzyme complex into methyl and carbonyl groups oxidizing the latter to CO2 and reducing a 2 x 4Fe-4S ferredoxin (18, 31). The methyl group is transferred to HS-CoM. In freshwater species Methanosarcina barkeri and Methanosarcina mazei it is proposed that the reduced ferredoxin donates electrons to a membrane-bound H2-evolving hydrogenase (Ech) generating a proton gradient (high outside) (32, 33). It is further proposed that the H2 is then oxidized by another membrane-bound hydrogenase that delivers electrons to the quinone-like electron carrier methanophenazine (29). The reduced methanophenazine donates electrons to the membrane-bound DE type heterodisulfide reductase, the subunits of which correspond to the HdrBC subunits of the HdrABC type enzyme of obligate CO2-reducing species (2, 10,

21, 22). Reduction and oxidation of methanophenazine further contributes to the proton gradient driving ATP synthesis (11). The same enzymes and reactions function in the 142 marine isolate Methanosarcina acetivorans except the hydrogenases are replaced by the membrane-bound Rnf complex that mediates electron transfer between ferredoxin and methanophenazine and is hypothesized to pump either protons or sodium outside the cytoplasmic membrane that drives ATP synthesis (15, 31).

In an apparent anomaly, the sequenced genomes of all aceticlastic species also contain genes encoding subunits of HdrABC, although biochemical evidence for its presence in cells has not been presented and genetic evidence indicates it is not essential

(2). The sequenced genomes of all aceticlastic species also contain a gene encoding subunits of a HdrA homolog fused to a MvhD homolog that is abundant in acetate-grown

M. acetivorans suggesting a role in the aceticlastic pathway recently confirmed by genetic evidence (2). Repression of the gene encoding the HdrA: MvhD fusion under control of a tetracycline-dependent promoter resulted in a longer lag phase, decreased growth rate and decreased final cell density when cultured with acetate (3). The fusion protein is postulated to form a transcription unit with a polyferredoxin encoding gene and distant from genes encoding HdrBC consistent with it functioning independently from a

HdrABC type of heterodisulfide reductase.

Here we present the overexpression, purification and partial characterization of the

HdrA: MvhD fusion protein from M. acetivorans we name Etp (electron transfer protein) based on results supporting a cytoplasmic electron carrier function linked to ferredoxin. 143

4.3 Materials and methods

Cell growth and materials

Acetate-grown and methanol-grown M. acetivorans (23) were grown in a 10 liter

Microferm fermentor (New Brunswick Scientific, Edison, NJ) as previously described

(24). The cells were collected and harvested by centrifugation at 4,000 rpm for 20 minutes and stored at -80 ℃ until use. CoM-S-S-CoB was custom synthesized by

Sigma-Aldrich Corp. (St. Louis, MO). All chemicals were purchased from

Sigma-Aldrich Corp.. All chromatography columns, resins and pre-packed columns were purchased from GE Healthcare (Waukesha, WI). Cloning vectors and all E. coli strains were purchased from EMD Chemicals Inc. (Gibbstown, NJ). Rainbow marker used in SDS-PAGE analysis was purchased from GE Healthcare. Anti-His (C-term)/AP antibody was purchased from Invitrogen Corporation (Carlsbad, CA). Vivacell 70 protein concentrator and membranes were purchased from Sartorius Group (Göettingen,

Germany). CdhAE and the 2 x 4Fe-4S ferredoxin from acetate-grown M. acetivorans were purified as previously described (31).

Analytical

The protein concentrations of soluble and membrane fractions of M. acetivorans were determined with Bradford assay (Bio-Rad Laboratories, Hercules, CA). All other 144 protein concentrations were determined with bicinchoninic acid assay (Thermo Fisher

Scientific Inc., Waltham, MA). Iron and acid-labile sulfur content of Etp was assayed as previously described (13). The flavin content of Etp was determined following reported protocol (8), except for that UV-visible spectrometry instead of fluorescence spectrometry was used for the determination of flavin content and the HPLC step was omitted. Benzylviologen: CoM-S-S-CoB oxidoreductase activity was assayed as previously described (8), except for an 80 μl reaction system was used and 50 mM MOPS, pH 6.8 was used as reaction buffer. A unit of benzylviologen: CoM-S-S-CoB oxidoreductase activity was defined as the amount of enzyme required to oxidize 1 μmol of benzylviologen per minute. Hydrogenase activity assay was performed as described

(17). Methylviologen was used as the electron acceptor in this assay and its reduction was followed by increase of absorption at 603 nm. DTNB assay was performed as previously described except for a 300 μl reaction system was used in some assays described in this chapter (31).

Overexpression, purification and reconstitution of Etp

All steps starting from breaking cells were performed anaerobically in an anaerobic chamber (Coy Laboratories Products Inc., Grass Lake, MI) under 1 atm atmosphere containing 95% N2 and 5% H2 or Beckman L8-70M ultracentrifuge (Beckman Coulter,

Inc., Brea, CA) using airtight centrifuge tubes to ensure an anaerobic environment. 145

MA2868 encoding Etp was amplified from the genomic DNA of M. acetivorans by PCR and cloned into pET22b vector, generating pMW2. pMW2 was then transformed into

RosettaBlue (DE3) pLacI E. coli cells. Cells were grown at 37 ℃ in LB medium supplemented with 100 μg/ml ampcilin and 30 μg/ml chloramphenicol with shaking at

250 rpm until OD600=1.0. 1 mM isopropyl-β-D-thiogalactopyranoside (IPTG) was then added to culture and temperature was decreased to 16 ℃. The cells were grown overnight, and then harvested and stored at -20 ℃ until use. 120 g of cells were thawn and resuspended in 50 mM MOPS, pH 6.8, 500 mM NaCl, 20 mM imidazole with a 1:1

(w/v) cell/buffer ratio, and were broken by passing through French pressure cell at 6.9 x

103 kPa twice. Cell lysate was centrifuged at 100,000 x g for 30 min to precipitate unbroken cells and cell debris. The supernant was collected and loaded onto a

Ni-Sepharose column (bed volumn 20 ml) and equilibrated by passing 100 ml of 50 mM

MOPS, pH 6.8, 500 mM NaCl, 20 mM imidazole through the column. The column was then washed with 100 ml of 50 mM MOPS, pH 6.8, 500 mM NaCl, 60 mM imidazole.

His-tagged Etp was eluted by passing 100 ml of 50 mM MOPS, pH 6.8, 500 mM NaCl,

250 mM imidazole through the column. The major peak was collected and concentrated with Vivacell 70 protein concentrator equipped with a 10K MWCO membrane (10 kDa cut off). The concentrated protein was diluted with 50 mM MOPS, pH 6.8 until a final concentration of 50 mM NaCl was reached. The diluted protein was loaded on a HiTrap

Q-Sepharose HP column (bed volumn 5 ml). This column was developed by a linear 50 ml 0-1.0 M NaCl gradient. Fractions were collected and examined with SDS-PAGE 146 and western blotting with anti-His(C-term)/AP antibody. Fractions containing a single band at 87 kDa which reacts with anti-His(C-term)/AP antibody were pooled. The purified Etp was reconstituted with iron and sulfur using previously described methods, with and without FAD and FMN (34). The 87 kDa band was cleaved from the gel, digested with trypsin overnight, and analyzed with LC-MS/MS to confirm its content.

Biochemical assays

All biochemical assays in this chapter were performed in an anaerobic chamber under 95% N2, 5% H2 at room temperature.

Ferredoxin: CoM-S-S-CoB oxidoreductase activity of soluble fraction, membrane fraction and cell lysate of acetate-grown M. acetivorans

The cell lysate, membrane fraction and soluble fraction of acetate-grown M. acetivorans were prepared with sucrose gradient ultracentrifugation as previously described (31). The soluble fraction was centrifuged at 200,000 x g for 2 hours with a

Beckman type 50 Ti rotor to precipitate residual membrane. Ferredoxin: CoM-S-S-CoB oxidoreductase activity of soluble fraction, membrane fraction and cell lysate of acetate-grown M. acetivorans were assayed with both Ferredoxin-NADP+ oxidoreductase

(FNR) and Cdh as ferredoxin regenerating systems. In Cdh system, a 200 μl reaction 147 system containing 5 μM ferredoxin, 2.24 μg CdhAE , 320 μM CoM-S-S-CoB and 50 mM

MOPS, pH 6.8 were added to a sealed glass vial with 1 atm 100% CO as atmosphere.

Soluble fraction (75 μg protein) or membrane (178μg protein) or cell lysate (111 μg protein) was added to the system respectively with a Hamilton syringe to initiate reaction.

30 μl of reaction mixture was taken out of the vial every 3 or 4 minutes, and the content of free thiols was assayed with DTNB assay. In FNR system, a 222 μl reaction system containing 18 mM NADPH, 2 μM ferredoxin, 0.9 mM CoM-S-S-CoB, 50 mM MOPS, pH 6.8 and soluble fraction (159 μg protein) or cell lysate (166.8 μg protein) or membrane fraction (106.8 μg protein) were added to a tube. 4 μl FNR (1mg/ml) was added to initiate reaction. 30 μl of reaction mixture was taken out of the tube every 4 minutes, and the content of free thiols was assayed with DTNB assay. 1 unit of ferredoxin: CoM-S-S-CoB oxidoreductase activity was defined as 1 μmol of free thiols generated per minute.

The reduction of Etp and its dependence on ferredoxin

11.3 μM Etp and 3.1 μM ferredoxin in a total volume of 69 μl was added to a rubber stopper sealed cuvette. The reaction system was buffered with 50 mM MOPS, pH 6.8.

The cuvette was flushed with 100% CO, and reaction was initiated by the addition of 1 μl soluble fraction of acetate-grown M. acetivorans (22.1 μg protein). The reduction of

Etp was followed by the decrease of A420. Control experiments were done by omitting 148 each component, and by replacing the soluble fraction with CdhAE of the same amount of total Cdh activity. 0 μM, 1.6 μM, 3.1 μM or 6.3 μM of ferredoxin were used in this assay to test the dependence of this reduction on ferredoxin.

Effect of membrane fraction of acetate-grown M. acetivorans on the reduction of Etp

The reduction of Etpwas done in a 60 μl reaction system containing the following: 50 mM MOPS, pH 6.8, 2.2 μM ferredoxin, 26 μM reconstituted Etp and soluble fraction of acetate-grown M. acetivorans (10.6 μg protein) under 1 atm of 100% CO atmosphere.

The reaction was initiated by the addition of soluble fraction. Two experiments were done in which soluble fraction was omitted or replaced with membrane fraction of acetate-grown M. acetivorans containing 8.9 μg protein. In these two experiments, purified CdhAE with the same amount of total Cdh activity as soluble fraction containing

10.6μg protein was added to initiate reaction. The reduction of Etp was identified by the decrease of A420.

The stimulation of Etp of ferredoxin: CoM-S-S-CoB oxidoreductase activity in soluble fraction of acetate-grown M. acetivorans

To assay if Etp can stimulate the ferredoxin: CoM-S-S-CoB oxidoreductase activity in soluble fraction of acetate-grown M. acetivorans, a 200 μl reaction system was 149 prepared in a rubber stopper sealed cuvette containing the following: 2.2 μM ferredoxin,

0, 4 or 8 μM Etp, 360 μM CoM-S-S-CoB and 50 mM MOPS, pH 6.8. The cuvette was flushed with 100% CO. The reaction was initiated by the addition of 2 μl soluble fraction (44.2 μg protein). The generation of CoM-SH and CoB-SH was followed by assaying the free thiol groups by DTNB assay.

4.4 Results

4.4.1 Purification and Properties

Etp from M. acetivorans was heterologously overexpressed in E. coli and purified to near homogeneity (Fig. 4-1). Mass spectrometry analysis confirmed the purified protein is the product of MA2868. The sequence of HdrA, previously characterized from

Methanothermobacter thermoautotrophicus (9), aligned with N-terminal residues 1-647

Figure 4-1 SDS-PAGE of purified Etp. 150 from the deduced sequence of MA2868 revealing 50% identity and conservation of four

[Fe4S4]-binding motifs and a consensus flavin-binding motif (Fig. 4-2). Alignment of the previously characterized MvhD from M. thermoautotrophicus revealed 49% sequence identity with C-terminal residues 651- 757 of the MA2868 deduced sequence and conservation of a CX2CX25CX24CX4C motif proposed to bind a 2Fe-2S cluster in MvhD

(25) (Fig. 4-2). These analyses predict that Etp is an iron-sulfur flavoprotein. Indeed, the as-purified protein had a UV-visible absorption spectrum with peaks at 330 and 420 nm and a shoulder at 465 nm consistent with the predicted 2Fe-2S cluster. Treatment of the as-purified protein with ferric iron and sulfide increased the absorbance at 420 nm

3-fold suggesting reconstitution of iron-sulfur clusters (Fig. 4-3). Iron and acid-labile sulfur analysis further supported these observations, showing 1.9 ± 0.9 iron and 2.9 ± 0.1 acid-labile sulfur in the as-purified protein and 13.2 ± 1.2 iron and 11.6 ± 0.2 acid-labile sulfur in the reconstituted protein for each molecule of protein. Flavin was not detected and attempts to reconstitute with FAD or FMN failed. Enzymatic activity assays showed purified Etp has little hydrogenase or heterodisulfide reductase activities, suggesting Etp is neither a hydrogenase nor heterodisulfide reductase, although it represents a fusion protein of a subunit of hydrogenase and a subunit of heterodisulfide reductase. 151

Figure 4-2 Alignment of the deduced sequences of etp, hdrA and mvhD. MA2868,

MbarA2589, Mbur0552, Mthe1576 are HdrA: MvhD fusion proteins (Etp) from M. acetivorans, M. barkeri,

Methanococcoides burtonii, Methanosaeta thermophila. HdrA and MvhD are from M. thermoautotrophicus. Protein sequences are from the CMR database (4). Conserved FAD and iron sulfur cluster binding motifs are highlighted. 152

Figure 4-3 UV-visible absorption spectrum of unreconstituted and reconstituted Etp. Panel

A: 11 μM unreconstituted Etp; Panel B: 11μM reconstituted Etp.

4.4.2 Physiology

It is hypothesized that HdrA is the site for reduction of ferredoxin by the

HdrABC/MvhADG complex in obligate CO2-reducing methanogens (29) prompting an examination of the ability of ferredoxin to reduce Etp. A reduced ferredoxin regenerating system was utilized that was comprised of the CO-oxidizing component

(CdhAE) of the CO-dehydrogenase/acetyl-CoA synthase complex (CdhABCDE) and the ferredoxin that accepts electrons from CdhAE (31). Both CdhAE and ferredoxin were purified from acetate-grown M. acetivorans (31). Reduction of Etp was followed by a decrease in absorbance at 420 nm. A significant decrease in absorbance was observed 153 when purified CdhAE was used as the source of Cdh activity, an apparent reduction of iron-sulfur centers in CdhAE. However, when CdhAE was replaced with the soluble fraction from acetate-grown M. acetivorans with the same total Cdh activity, the absorbance change increased 4.8-fold over the background rate with purified CdhAE as the source of Cdh activity and was dependent on the presence of ferredoxin and CO (Fig.

4-4) (Fig. 4-5). The dependence on ferredoxin is further supported by a proportional increase in the rate of Etp reduction with increasing concentrations of ferredoxin (Fig.

4-5). These results indicate that Etp is a component of a ferredoxin-linked soluble electron transport chain in acetate-grown cells originating with Cdh and that ferredoxin is not the direct electron donor to Etp. Membrane fraction of cell-free extract of acetate-grown M. acetivorans was unable to stimulate the rate of reduction when CdhAE was used as the source of Cdh activity, further supporting the hypothesis that additional soluble factors are involved in the reduction of Etp (Fig. 4-6).

The finding that Etp is linked to ferredoxin is consistent with the previously hypothesized role in a soluble ferredoxin: CoM-S-S-CoB oxidoreductase system of acetate-grown M. acetivorans (7) that was further investigated with activities in the membrane and soluble fractions of cell-free extract. The total heterodisulfide reductase activity recovered in the soluble and membrane fractions was 56% and 29% of the extract

(Table 4-1). When assayed with the CO/Cdh ferredoxin regenerating system, the soluble fraction showed ferredoxin: CoM-S-S-CoB oxidoreductase activity (Table 4-1) that was stimulated by the addition of ferredoxin and Etp (Fig. 4-7) (Fig.4-8). 154

Determination of protein concentration in soluble and membrane fractions with both

BCA assay and Bradford assay revealed significant differences (Table 4-2). To eliminate the effect of lipid and lipoproteins, which artificially increase the protein concentration with BCA assay in membranes, Bradford assay was used in protein concentration determination and calculation of specific activity. Approximately one-half of the total ferredoxin: CoM-S-S-CoB oxidoreductase activity in cell extract could be recovered in the soluble fraction (Fig. 4-9) (Table 4-1). The same result was found when activities were assayed with the NADPH/FNR ferredoxin regenerating system (Fig. 4-10) (Table 4-1). These results support a soluble ferredoxin:

CoM-S-S-CoB oxidoreductase system and a role for Etp.

Figure 4-4 The reduction of Etp requires unknown components in soluble fraction of acetate-grown M. acetivorans. In a 70 μl reaction system, the following was added: 11.3 μM Etp, 3.1 μM ferredoxin, 1 μl soluble fraction of acetate-grown M. acetivorans (22.1 μg protein). The reduction of

Etp was followed by the decrease of A420. Various controls were done by deleting each component and replacing soluble fraction with CdhAE of the same total Cdh activity. Symbols: (▲) full assay, (●) no Etp control, (■) no CO control, (△) soluble fraction replaced with CdhAE. 155

Figure 4-5 Ferredoxin-dependent Etp reduction. Panel A: Time-course of the reduction of Etp with different concentrations of ferredoxin. Symbols: (△ ) 6.3 μM ferredoxin, (▲) 3.1 μM ferredoxin, (■)

1.6 μM ferredoxin, (◆) no ferredoxin. Panel B: Correlation between ferredoxin concentration and the reduction rate of Etp .

Figure 4-6 Membrane fraction of acetate-grown M. acetivorans doesn’t stimulate the reduction of Etp. The reduction of Etp with soluble fraction as the source of Cdh is compared with purified CdhAE of the same Cdh activity as the source of Cdh. Membrane fraction was added in the system in which purified CdhAE was used as the source of Cdh activity to see if a stimulation of activity could be observed. Symbols: soluble fraction as the source of Cdh (■), purified CdhAE as the source of

Cdh (Δ), membrane fraction added to the system with purified CdhAE (). 156

ferredoxin: ferredoxin: heterodisulfide heterodisulfide heterodisulfide oxidoreductase oxidoreductase reductase fraction (CO/Cdh) (NADPH/FNR) total specific total specific total specific activity activity activity activity activity activity (mU) (mU/mg) (mU) (mU/mg) (U) (U/mg) extract 2839±300 17.0±1.8 2956±301 17.7±1.8 119 ± 12.8 0.73 ± 0.08 soluble 1579±75 10.6±0.5 1401±164 9.4±1.1 66.5 ± 5.9 0.44±0.04 membrane 193±2 8.4±0.1 223±16 9.7±0.7 34.4 ± 0.6 1.48±0.03

Table 4-1 Activities catalyzed by cell-free extract, soluble fraction and membrane fraction of acetate-grown M. acetivorans.

Figure 4-7 The stimulation of Etp of ferredoxin: CoM-S-S-CoB oxidoreductase activity in soluble fraction of acetate-grown M. acetivorans. The 200 μl reaction mixture contains 2 μl soluble fraction (44.2 μg protein), 2.2 μM ferredoxin, 360 μM CoM-S-S-CoB and 0, 4, or 8 μM Etp. The reaction is followed by the appearance of free thiol groups in system assayed by DTNB assay. Symbols: (▲) 8

μM Etp, (■) 4 μM Etp, (◆) 0 μM Etp.

157

Fractions Assay for protein Soluble Membrane Cell lysate concentration Concentration Total protein Concentration Total protein Concentration Total protein determination (mg/ml) (mg) (mg/ml) (mg) (mg/ml) (mg)

Bradford assay 10.6±1.3 149.5 8.9±0.6 23.1 55.6±4.8 166.8

BCA assay 15.6±0.2 220.0 18.7±0.7 48.6 71.0±1.8 213.0

Table 4-2 Comparison of protein concentrations determined by BCA assay and Bradford

assay. 3 ml of cell lysate of acetate-grown M. acetivorans was separated into soluble fraction and

membrane fraction with sucrose gradient ultracentrifugation. 2.6 ml of membrane fraction and 14.1 ml of

soluble fraction were obtained.

Figure 4-8 Ferredoxin: CoM-S-S-CoB oxidoreductase activity in the soluble fraction of

acetate-grown M. acetivorans. In a 100 μl system, 5 μl of soluble fraction of acetate-grown M.

acetivorans (110.5 μg protein), 5 μM ferredoxin, 1.12 μg CdhAE was added to a rubber stopper sealed

cuvette filled with 100% CO. Controls were performed with every component deleted, resulting in no or

severely impaired activity. Activity is followed by the appearance of free thiols. Symbols: (◆) full

assay with 5 μM ferredoxin, (■) no ferredoxin control. 158

Figure 4-9 Ferredoxin: CoM-S-S-CoB oxidoreductase activity in fractions of acetate-grown

M. acetivorans with CO/Cdh as ferredoxin regenerating system. Panel A: cell-free extract (111 μg protein); Panel B: membrane fraction (178 μg protein); Panel C: soluble fraction (75 μg protein). 159

Figure 4-10 Ferredoxin: CoM-S-S-CoB oxidoreductase activity in fractions of acetate-grown

M. acetivorans with NADPH/FNR as ferredoxin regenerating system. Panel A: cell-free extract

(166.8 μg protein); Panel B: membrane fraction (106.8 μg protein); Panel C: soluble fraction (159 μg protein).

160

4.4.3 Phylogeny

Although a search of the CMR database queried with the deduced sequence of

MA2868 retrieved homologs of Etp in the sequenced genomes of all methane-producing acetotrophic species (Fig. 4-2), none were identified in obligate CO2-reducing species consistent with a role for Etp in the aceticlastic pathway. However, a homolog was identified in Methanococcoides burtonii, the only obligate methylotrophic methanogen for which the genome is sequenced (1). A homolog of Etp was also identified in the sequenced genome of Archaeoglobus fulgidus, a non-methanogenic sulfate-reducing species from the Archaea domain that also encodes homologs of both types of heterodisulfide reductase (16).

4.5 Discussion

Although proteomic and genetic analyses implicate a physiological role for the putative HdrA: MvhD fusion protein in the aceticlastic pathway for methanogenesis (2,

14), the protein had not been characterized and a specific function had not been documented. Here we describe the first biochemical and physiological characterization that showed Etp is a component of an electron transport chain originating with Cdh, the enzyme complex central to the aceticlastic pathway in acetotrophic methanogens that cleaves acetyl-CoA and oxidizes the carbonyl group with transfer of electrons to 161 ferredoxin (7). The results support an electron transfer function in the aceticlastic pathway of M. acetivorans and the rationale for naming the protein Etp (electron transport protein).

A proposed electron acceptor of the HdrA subunit in the HdrABC/MvhADG complex of obligate CO2-reducers is a 2 x 4Fe-4S ferredoxin that donates electrons to the formylmethanofuran dehydrogenase catalyzing the first reductive step in the methanogenic pathway of obligate CO2-reducers (28). The results presented here show that Etp is linked to the 2 x 4Fe-4S ferredoxin from acetate-grown cells, although not the direct electron donor to Etp. The polyferredoxin encoded (MA2867) adjacent to the gene encoding Etp (MA2868) is a candidate electron donor to Etp. The deduced sequence of the polyferredoxin contains 8 [Fe4S4]-binding motifs and is present in acetate-grown M. acetivorans in an amount 12-fold greater than in methanol-grown cells

(14). Furthermore, the juxtaposition of genes encoding the polyferredoxin and Etp is conserved in all sequenced genomes of acetotrophic methanogens except for

Methanosaeta thermophila. In the HdrABC/MvhADG complex of obligate

CO2-reducers the MvhD subunit transfers electrons to the flavin-containing HdrA subunit

(28) which implies that electrons are likely transferred between the HdrA and MvhD domains of Etp, although the entry point of electrons and the electron acceptor of Etp is unknown. The apparent absence of flavin in Etp contrasts with the HdrA subunit of obligate CO2-reducers which contains FAD, the proposed site for reduction of ferredoxin

(28). 162

The results reported here are consistent with a role for Etp in a soluble ferredoxin:

CoM-S-S-CoB oxidoreductase system in the acetotrophic pathway of M. acetivorans.

Based on thermodynamic considerations, a branched electron transport pathway was previously hypothesized that included a soluble component bypassing membrane-bound electron transport allowing flexibility in the generation of ion gradients driving ATP synthesis in response to fluctuating levels of acetate in the environment (7). Although methanogenesis from acetate accounts for the majority of methane produced globally (6),

+ acetate is a relatively poor substrate energetically (CH3COOH + H  CH4 + CO2, G°

 36 kJ), with a change of Gibbs free energy barely enough for the synthesis of one

ATP under standard conditions of equimolar reactants and products. In the native environment the amount of available energy is much lower and fluctuating. Aceticlastic methanogens obtain energy for growth by coupling membrane-bound electron transport from ferredoxin to CoM-S-S-CoB with translocation of ions generating a gradient (high outside) that drives ATP synthesis (5, 11, 20). In freshwater Methanosarcina species, ferredoxin donates electrons to a membrane-bound hydrogenase (Ech) evolving H2 (17,

19, 26) and pumping protons (33). A H2-uptake hydrogenase (Vho) oxidizes H2 and donates electrons to methanophenazine that mediates electron transfer to the HdrDE type of heterodisulfide reductase translocating protons that contribute to the gradient (11).

An additional two protons are translocated by the Vho hydrogenase for a total of four protons translocated by the H2: CoM-S-S-CoB oxidoreductase system (11). In addition, transfer of the methyl group of acetate to HS-CoM by the membrane-bound 163 methyltransferase Mtr (G°  30 kJ) is coupled to translocation of sodium outside the membrane that could potentially drive ATP synthesis in conjunction with a sodium/proton antiporter. However, it is unlikely that the amount of energy available from conversion of acetate to methane (G  36 kJ/mol) can support all three coupling sites requiring a branched electron transport in which one branch bypasses one or more coupling sites, particularly in the native environment where low acetate concentrations limit growth. In support of a soluble branch, a mutant of M. barkeri with the gene encoding the Ech hydrogenase deleted retains a portion of the wild-type ferredoxin:

CoM-S-S-CoB oxidoreductase activity (19). In further support, deletion of the coenzyme F420-dependent Frh hydrogenase and F420 dehydrogenase prevents growth of M. barkeri with acetate suggesting an alternate electron transport pathway involving the soluble electron carrier coenzyme F420 and either of these two enzymes (12).

The same thermodynamic argument for a soluble branch applies to M. acetivorans.

In this marine strain, the coupling sites are the same as for freshwater Methanosarcina except the Ech and Vho hydrogenases are replaced with the membrane-bound Rnf complex that like Ech accepts electrons directly from ferredoxin (31) and is postulated to translocate ions driving ATP synthesis. The results presented here provide a basis for proposing a branched electron transport pathway in which electrons donated to ferredoxin by Cdh are transferred to either Rnf or the soluble electron transport chain involving Etp

(Fig. 4-11). The identity of electron carriers other than Etp and the type of soluble heterodisulfide reductase is unknown. It was previously hypothesized that the soluble 164 electron transport chain joins the membrane-bound electron transport chain either at methanophenazine or HdrDE bypassing one or two coupling sites (7). However, the finding of substantial heterodisulfide reductase and ferredoxin: CoM-S-S-CoB oxidoreductase activity in the soluble fraction suggests either the HdrABC type is present and functions in the soluble fraction or the HdrDE type is loosely associated with the membrane and dissociates during fractionation.

Figure 4-11 Proposed electron transport model in acetate-grown M. acetivorans. Panel A:

Proposed model for electron transport. Panel B: Proposed role of Etp. MaRnf, Rnf complex from M.

acetivorans; Cytc: cytochrome c; fdO, ferredoxin oxidized; fdR, ferredoxin reduced; MP, methanophenazine;

CoM-SH, coenzyme M; CoB-SH, coenzyme B; CoM-S-S-CoB, the heterodisulfide of coenzyme M and

coenzyme B; pfdO, polyferredoxin oxidized; pfdR, polyferredoxin reduced; HdrDE, HdrB2, HdrC2, heterodisulfide reductase DE, subunit B2, subunit C2. 165

4.6 Acknowledgements

This work was supported by the National Science Foundation. We thank the Penn

State Hershey Core Research Facilities for mass spectrometry analyses.

4.7 References

1. Allen, M. A., F. M. Lauro, T. J. Williams, D. Burg, K. S. Siddiqui, D. De

Francisci, K. W. Chong, O. Pilak, H. H. Chew, M. Z. De Maere, L. Ting, M.

Katrib, C. Ng, K. R. Sowers, M. Y. Galperin, I. J. Anderson, N. Ivanova, E.

Dalin, M. Martinez, A. Lapidus, L. Hauser, M. Land, T. Thomas, and R.

Cavicchioli. 2009. The genome sequence of the psychrophilic archaeon,

Methanococcoides burtonii: the role of genome evolution in cold adaptation.

ISME J 3:1012-35.

2. Buan, N. R., and W. W. Metcalf. 2010. Methanogenesis by Methanosarcina

acetivorans involves two structurally and functionally distinct classes of

heterodisulfide reductase. Mol. Microbiol. 75:843-853.

3. Buan, N. R., and W. W. Metcalf. 2009. Methanogenesis by Methanosarcina

acetivorans involves two structurally and functionally distinct classes of

heterodisulfide reductase. Mol Microbiol.

4. Costa, K. C., P. M. Wong, T. Wang, T. J. Lie, J. A. Dodsworth, I. Swanson, J.

A. Burn, M. Hackett, and J. A. Leigh. 2010. Protein complexing in a 166 methanogen suggests electron bifurcation and electron delivery from formate to

heterodisulfide reductase. Proc. Natl. Acad. Sci. U S A 107:11050-11055.

5. Doddema, H. J., T. J. Hutten, C. van der Drift, and G. D. Vogels. 1978. ATP

hydrolysis and synthesis by the membrane-bound ATP synthetase complex of

Methanobacterium thermoautotrophicum. J Bacteriol 136:19-23.

6. Ferry, J. G. 1992. Biochemistry of methanogenesis. Crit Rev Biochem Mol Biol

27:473-503.

7. Ferry, J. G. 2010. How to make a living exhaling methane. Annu. Rev.

Microbiol. 64:453-473.

8. Hedderich, R., A. Berkessel, and R. K. Thauer. 1990. Purification and

properties of heterodisulfide reductase from Methanobacterium

thermoautotrophicum (strain Marburg). Eur J Biochem 193:255-61.

9. Hedderich, R., J. Koch, D. Linder, and R. K. Thauer. 1994. The

heterodisulfide reductase from Methanobacterium thermoautotrophicum contains

sequence motifs characteristic of pyridine-nucleotide-dependent thioredoxin

reductases. Eur J Biochem 225:253-61.

10. Heiden, S., R. Hedderich, E. Setzke, and R. K. Thauer. 1993. Purification of a

cytochrome-b containing H2-heterodisulfide oxidoreductase complex from

membranes of Methanosarcina barkeri. Eur. J. Biochem. 213:529-535.

11. Ide, T., S. Baumer, and U. Deppenmeier. 1999. Energy conservation by the H2:

heterodisulfide oxidoreductase from Methanosarcina mazei Go1: identification of

two proton-translocating segments. J Bacteriol 181:4076-80. 167 12. Kulkarni, G., D. M. Kridelbaugh, A. M. Guss, and W. W. Metcalf. 2009.

Hydrogen is a preferred intermediate in the energy-conserving electron transport

chain of Methanosarcina barkeri. Proc Natl Acad Sci U S A 106:15915-20.

13. Lessner, D. J., and J. G. Ferry. 2007. The archaeon Methanosarcina

acetivorans contains a protein disulfide reductase with an iron-sulfur cluster. J

Bacteriol 189:7475-84.

14. Li, L., Q. Li, L. Rohlin, U. Kim, K. Salmon, T. Rejtar, R. P. Gunsalus, B. L.

Karger, and J. G. Ferry. 2007. Quantitative proteomic and microarray analysis

of the archaeon Methanosarcina acetivorans grown with acetate versus methanol.

J. Proteome. Res. 6:759-771.

15. Li, Q., L. Li, T. Rejtar, D. J. Lessner, B. L. Karger, and J. G. Ferry. 2006.

Electron transport in the pathway of acetate conversion to methane in the marine

archaeon Methanosarcina acetivorans. J. Bacteriol. 188:702-710.

16. Mander, G. J., A. J. Pierik, H. Huber, and R. Hedderich. 2004. Two distinct

heterodisulfide reductase-like enzymes in the sulfate-reducing archaeon

Archaeoglobus profundus. Eur J Biochem 271:1106-16.

17. Meuer, J., S. Bartoschek, J. Koch, A. Kunkel, and R. Hedderich. 1999.

Purification and catalytic properties of Ech hydrogenase from Methanosarcina

barkeri. Eur J Biochem 265:325-35.

18. Meuer, J., H. C. Kuettner, J. K. Zhang, R. Hedderich, and W. W. Metcalf.

2002. Genetic analysis of the archaeon Methanosarcina barkeri Fusaro reveals a

central role for Ech hydrogenase and ferredoxin in methanogenesis and carbon

fixation. Proc. Natl. Acad. Sci. U. S. A. 99:5632-5637. 168 19. Meuer, J., H. C. Kuettner, J. K. Zhang, R. Hedderich, and W. W. Metcalf.

2002. Genetic analysis of the archaeon Methanosarcina barkeri Fusaro reveals a

central role for Ech hydrogenase and ferredoxin in methanogenesis and carbon

fixation. Proc Natl Acad Sci U S A 99:5632-7.

20. Mountfort, D. O. 1978. Evidence from ATP synthesis driven by a proton

gradient in Methanosarcina barkeri. Biochem Biophys Res Commun 85:1346-51.

21. Peer, C. W., M. H. Painter, M. E. Rasche, and J. G. Ferry. 1994.

Characterization of a CO:heterodisulfide oxidoreductase system from

acetate-grown Methanosarcina thermophila. J. Bacteriol. 176:6974-6979.

22. Simianu, M., E. Murakami, J. M. Brewer, and S. W. Ragsdale. 1998.

Purification and properties of the heme- and iron-sulfur- containing

heterodisulfide reductase from Methanosarcina thermophila. Biochemistry

37:10027-10039.

23. Sowers, K. R., S. F. Baron, and J. G. Ferry. 1984. Methanosarcina acetivorans

sp. nov., an acetotrophic methane-producing bacterium isolated from marine

sediments. Appl Environ Microbiol 47:971-8.

24. Sowers, K. R., M. J. K. Nelson, and J. G. Ferry. 1984. Growth of acetotrophic,

methane-producing bacteria in a pH auxostat. Curr. Microbiol. 11:227-230.

25. Stojanowic, A., G. J. Mander, E. C. Duin, and R. Hedderich. 2003.

Physiological role of the F420-non-reducing hydrogenase (Mvh) from

Methanothermobacter marburgensis. Arch Microbiol 180:194-203.

26. Terlesky, K. C., and J. G. Ferry. 1988. Ferredoxin requirement for electron

transport from the carbon monoxide dehydrogenase complex to a 169 membrane-bound hydrogenase in acetate-grown Methanosarcina thermophila. J

Biol Chem 263:4075-9.

27. Thauer, R., Hedderich R., Fischer R. 1993. Reactions and Enzymes involved in

methanogenesis from CO2 and H2. In J. G. Ferry (ed.), Methanogenesis: Ecology,

Physiology, Biochemistry & Genetics Chapman & Hall, New York.

28. Thauer, R. K., A. K. Kaster, M. Goenrich, M. Schick, T. Hiromoto, and S.

Shima. 2010. Hydrogenases from methanogenic archaea, nickel, a novel cofactor,

and H2 storage. Annu. Rev. Biochem. 79:507-536.

29. Thauer, R. K., A. K. Kaster, M. Goenrich, M. Schick, T. Hiromoto, and S.

Shima. 2010. Hydrogenases from methanogenic archaea, nickel, a novel cofactor,

and H2 storage. Annu Rev Biochem 79:507-36.

30. Thauer, R. K., A. K. Kaster, H. Seedorf, W. Buckel, and R. Hedderich. 2008.

Methanogenic archaea: ecologically relevant differences in energy conservation.

Nat. Rev. Microbiol. 6:579-591.

31. Wang, M., Suharti, J. F. Tomb, and J. G. Ferry. 2010. The electron transport

chain of acetate-grown Methanosarcina acetivorans. submitted.

32. Welte, C., V. Kallnik, M. Grapp, G. Bender, S. Ragsdale, and U.

Deppenmeier. 2010. Function of Ech hydrogenase in ferredoxin-dependent

membrane-bound electron transport in Methanosarcina mazei. J. Bacteriol.

192:674-680.

33. Welte, C., C. Kratzer, and U. Deppenmeier. 2010. Involvement of Ech

hydrogenase in energy conservation of Methanosarcina mazei. FEBS J. Epub

ahead of print 170

34. Zhao, T., F. Cruz, and J. G. Ferry. 2001. Iron-sulfur flavoprotein (Isf) from

Methanosarcina thermophila is the prototype of a widely distributed family. J

Bacteriol 183:6225-33. 171

Chapter 5

Discussion and future directions on the understanding of electron

transport in acetate-grown Methanosarcina acetivorans

Although two thirds of biochemically generated methane on earth come from methanogenesis from acetate (7), only nine species of acetate-utilizing methanogens in two genera have been identified so far (4, 6, 13, 20). The mechanism of energy conservation in aceticlastic pathway and the electron transport coupled to energy conservation is still one of the least understood among all methanogenesis pathways.

Due to the very low number of characterized acetate-utilizing species, it is still premature to claim the predominant electron transport pathways acetate-utilizing methanogens use in nature, although those of fresh water methanogens Methanosarcina mazei and

Methanosarcina barkeri are the best characterized as a result of more intensive genetic studies. In this chapter, a model of electron transport and energy conservation in marine methanogen M. acetivorans is constructed and discussed based on knowledge obtained during my thesis research. The future directions and projects following my thesis research are also discussed. 172 5.1 A model of energy conservation under fluctuating environmental substrate concentrations

The presence of two independent electron transport chains in acetate-grown M. acetivorans led to the construction of a unique model of energy conservation (21, 22).

Acetate is the most energy stringent substrate for methanogenesis. Methanogenesis from acetate can only lead to conservation of 36kJ/mol of energy under physiological state, barely enough for the synthesis of one ATP per molecule of acetate (Equation 5-1).

To make this energy starving scenario even worse, the concentration of acetate in natural environments, unlike in a lab environment, is always fluctuating. Because of the low amount of energy M. acetivorans can conserve out of acetate, developing mechanisms to handle this fluctuating concentration of nutrient and maximize energy conservation efficiency, such as developing bypasses of some energy coupling sites, will give it a selective advantage. Depending on environmental substrate level, these bypasses can be turned on and off, conserving more energy when there is more energy to conserve and less energy when there isn’t enough energy to drive more ATP synthesis.

- + , o CH3COO +H → CH4 + CO2 ΔG ’ = -36 kJ/mol

Equation 5-1 Methanogenesis from acetate.

Although the mid-point potential of M. acetivorans ferredoxin is not determined, the mid-point potential of a highly homologous ferredoxin from M. barkeri was reported to 173 be -322 mV at 21℃ (5). It was also reported that the mid-point potential of RnfG is

-129 mV and the reduction of it is a 2 electron transfer (17). Assuming the electron transport from ferredoxin to RnfG is a 2 electron transfer, the change of Gibbs free energy in this electron transfer from ferredoxin to RnfG is ΔGo’=-37.2 kJ/mol.

Assuming the membrane potential is -220 mV, this energy is enough for coupling the translocation of 2H+ for every 2 electrons transported. We thus hypothesize Rnf serves as a proton translocation site that translocates 2 protons for every pair of electrons transferred.

Previous research has established two other energy coupling sites in Methanosarcina species. The membrane-bound methyl transferase Mtr was shown to translocate 2 Na+ for every pair of electrons transported in M. mazei (2, 12). Another energy coupling site, the electron transport from methanophenazine to heterodisulfide reductase in M. mazei couples translocation of 2H＋ for every pair of electrons transported (8). This energy coupling site was also suggested in acetate-grown M. acetivorans (11).

Out of these three energy coupling sites, the Mtr site is mandatory. However, the

Rnf site and the methanophenazine site can be bypassed with the soluble ferredoxin:

CoM-S-S-CoB electron transport pathway described in Chapter 4. This leads to the translocation of 2Na+ + 4H+ for every molecule of acetate metabolized during the ‘fully on mode’, and 2Na+ during the ‘bypass mode’ (Fig. 5-1). 174

Figure 5-1 Model of electron transport and energy conservation in acetate-grown M. acetivorans. Panel A: the ‘fully on’ mode; Panel B: the ‘bypass’ mode.

+ It was shown that the Na -translocating F1F0 type ATPase is dispensable for M. acetivorans, suggesting the physiological coupling ion for ATP synthesis is H+ (15).

The Na+ gradient generated by Mtr can be replaced by H+ gradient with a Mrp complex

(11). Although the stoichiometry of this Na+/H+ antiporter has not been studied in either bacterial Mrp or archaea Mrp, previous research on Mrp-dependent artificial 175 transmembrane potential driven Na+ efflux in Bacillus subtilis suggested an over 1

H+/Na+ stoichiometry with B. subtilis Mrp, which is a homologue of M. acetivorans Mrp

(9, 18). The change of Gibbs free energy in this H+/Na+ antiport was suggested by

Thauer et al. (19), shown in Equation 5-2:

2H+(inside) + 1Na+(outside) → 2H+(outside) + 1Na+(inside), ΔGo’=0 kJ/mol

Equation 5-2 Na+ and H+ translocation by Mrp complex.

Based on these observations, we hypothesize that The Mrp complex in M. acetivorans has a H+/Na+ stoichiometry of 2. If all three coupling sites are used for energy conservation, the ions translocated can be replaced with 8H+, enough for the

+ synthesis of 2 ATP with the H -translocating A1A0 type ATPase (14). Considering the cells need to consume 1 ATP in the activation of acetate by acetate kinase, the cells can gain a net 1 ATP for every molecule of acetate metabolized, agreeing with the maximum amount of energy the cells can conserve from 1 molecule of acetate (Equation 5-1).

However, if the environmental level of acetate is low and cannot support a change of

Gibbs free energy of -36 kJ/mol, the cells can activate the ‘bypass mode’, translocating

2Na+ that can be replaced with 4H+ and support the synthesis of 0 ATP per acetate (1

ATP synthesized by ATPase, 1 ATP used in activation of acetate).

The on and off of the ‘bypass mode’ can be regulated in response to environmental substrate level, leading to the net synthesis of a varied amount (0-1) of ATP per molecule 176 of acetate. M. acetivorans can get a better energy yield out of acetate when its level is high in the environment and a lower energy yield when its level is low in environment, maximizing energy conservation according to environmental substrate-energy level.

This could be the way how M. acetivorans survives with low concentrations of an already energy scarce substrate: acetate.

5.2 Future directions on the research of electron transport during acetate metabolism

5.2.1 Membrane-bound electron transport chain

Although the components of the membrane-bound electron transport chain have been identified, revealing a unique ferredoxin: CoM-S-S-CoB electron transport pathway, a critical question still remains to be answered: the coupling ion of Rnf complex. In

Bacteria, Rnf complexes were hypothesized to be a Na+-translocating protein, but this has never been experimentally confirmed (3, 10, 11, 14). The coupling ion of Rnf complex needs to be addressed by showing a ferredoxin: CoM-S-S-CoB oxidoreductase coupled ion translocation in inverted membrane vesicles of acetate-grown M. acetivorans following an approach previously described during the research of Na+-translocating

N5-methyl-THMPT: coenzyme M methyltransferase of Methanosarcina mazei (2). This investigation may be complicated by the proton translocation at methanophenazine if Rnf is a proton pump. A comparison of ferredoxin: CoM-S-S-CoB oxidoreductase activity 177 coupled versus 2-hydroxyphenazine: CoM-S-S-CoB oxidoreductase activity coupled proton translocation will be needed in this case. The stoichiometry of proton translocated/electron transported for ferredoxin: CoM-S-S-CoB oxidoreductase activity coupled proton translocation should be higher than that of 2-hydroxyphenazine:

CoM-S-S-CoB oxidoreductase activity coupled proton translocation. A similar approach for the investigation of ion translocation in H2: CoM-S-S-CoB electron transport pathway has been reported (8).

Although evidence suggested Rnf is a member of ferredoxin: CoM-S-S-CoB electron transport pathway in M. acetivorans, its biochemical characteristics still remain largely unknown (22). The purification of Rnf from acetate-grown M. acetivorans will lead to the enhancement of our knowledge in the subunit composition, stoichiometry, behavior in liposomes and its physiological electron partners. As a starting point, the purification of this 6-8 subunit membrane-bound complex can follow the strategy used for the purification of Na+-translocating NADH: quinone oxidoreductase (Na+-Nqr) of

Vibrio cholera (1), which is similar to Rnf in its subunit composition and subcellular location. The purification of the subunits of Rnf is also an important project, which will eventually lead to the understanding of the intramolecular electron transport sequence in

Rnf. This project is especially promising because three out of the four subunits participating in electron transport, RnfB (16), RnfG (17) and cytochrome c (unpublished results from Suharti) have already been purified and partially characterized. However, it needs to be pointed out that these projects are not trivial. M. acetivorans grows very slowly on acetate with a relatively low cell yield (maximum yield is 10g/L). To obtain enough protein for biochemical analysis, months, if not years, are needed to get enough 178 cell materials for purification. Overexpression in Escherchia coli is also an option, but because E. coli has an Rnf homologue named Rsx complex (14), this proposed membrane-bound ion pump is likely to be toxic to E. coli.

5.2.2 Soluble electron transport chain

All sequenced Methanosarcina species carry etp and the gene (MA2867) encoding a polyferredoxin cotranscribing with it, suggesting a similar soluble electron transport chain in all known Methanosarcina species. This leads to the hypothesis that a mechanism of bypassing energy coupling sites is a common strategy all acetate-grown methanogens develop to maximize efficiency of energy conservation. It is therefore important to find out if acetate-grown M. barkeri, M. mazei and M. thermophila also carry a soluble ferredoxin: CoM-S-S-CoB oxidoreductase activity. The strict acetate-grown Methanosaeta thermophila carries an etp homologue but not the polyferredoxin encoding gene. It would also be interesting to see if Methanosaeta thermophila develops a similar mechanism of bypassing energy coupling sites during growth on acetate.

More detailed studies are still needed to understand the components and mechanism of the soluble electron transport chain. The immediate electron donor and acceptor of Etp still remain to be identified. It is therefore necessary to purify MA2867 from acetate-grown M. acetivorans and see if it donates electrons to Etp.

Overexpression of this polyferredoxin in E. coli is possible, but to incorporate all of its cofactors (8 x 4Fe-4S clusters) is going to be challenging. The electron acceptor of Etp 179 is suspected to be a heterodisulfide reductase, and the purification of soluble heterodisulfide reductase in acetate-grown M. acetivorans and finding out if Etp can be oxidized by it directly is another interesting project.

More biochemical characterization is needed for Etp. Most importantly, finding out the role of each of its iron sulfur clusters helps us to understand how electron transport happens between Etp and its electron partners, as well as the physiological significance of the MvhD homologue fused to it. Mutagenesis studies of the protein are therefore proposed to knock out the binding motifs of each of the iron-sulfur clusters and

FAD and observe the changes of physiological behavior of these mutants. The properties of these iron-sulfur clusters can also be studied with EPR and Mössbauer spectrometry. The signals of these iron-sulfur clusters will be difficult to distinguish because four of them are predicted to be 4Fe-4S clusters, it is therefore required to construct mutated species of Etp knocking out one or more of the binding motifs for the iron-sulfur clusters and study their spectroscopic characteristics.

5.3 References

1. Barquera, B., P. Hellwig, W. Zhou, J. E. Morgan, C. C. Hase, K. K. Gosink,

M. Nilges, P. J. Bruesehoff, A. Roth, C. R. Lancaster, and R. B. Gennis. 2002.

Purification and characterization of the recombinant Na+-translocating NADH:

quinone oxidoreductase from Vibrio cholerae. Biochemistry 41:3781-9. 180 2. Becher, B., V. Muller, and G. Gottschalk. 1992.

N5-methyl-tetrahydromethanopterin: coenzyme M methyltransferase of

Methanosarcina strain Go1 is an Na+-translocating membrane protein. J Bacteriol

174:7656-60.

3. Biegel, E., S. Schmidt, and V. Muller. 2009. Genetic, immunological and

biochemical evidence for a Rnf complex in the acetogen Acetobacterium woodii.

Environ Microbiol 11:1438-43.

4. Boone, D. R., W. B. Whiteman, and P. Rouviere. 1993. Diversity and

Taxonomy of Methanogens. In J. G. Ferry (ed.), Methanogenesis: Ecology,

Physiology, Biochemistry & Genetics. Chapman & Hall, New York.

5. Daas, P. J., W. R. Hagen, J. T. Keltjens, and G. D. Vogels. 1994.

Characterization and determination of the redox properties of the 2[4Fe-4S]

ferredoxin from Methanosarcina barkeri strain MS. FEBS Lett 356:342-4.

6. Eggen, R. I., R. van Kranenburg, A. J. Vriesema, A. C. Geerling, M. F.

Verhagen, W. R. Hagen, and W. M. de Vos. 1996. Carbon monoxide

dehydrogenase from Methanosarcina frisia Go1: characterization of the enzyme

and the regulated expression of two operon-like cdh gene clusters. J Biol Chem

271:14256-63.

7. Ferry, J. G. 1992. Biochemistry of methanogenesis. Crit Rev Biochem Mol Biol

27:473-503.

8. Ide, T., S. Baumer, and U. Deppenmeier. 1999. Energy conservation by the H2:

heterodisulfide oxidoreductase from Methanosarcina mazei Go1: identification of

two proton-translocating segments. J Bacteriol 181:4076-80. 181 9. Ito, M., A. A. Guffanti, W. Wang, and T. A. Krulwich. 2000. Effects of

nonpolar mutations in each of the seven Bacillus subtilis mrp genes suggest

complex interactions among the gene products in support of Na+ and alkali but

not cholate resistance. J Bacteriol 182:5663-70.

10. Jeong, H. S., and Y. Jouanneau. 2000. Enhanced nitrogenase activity in strains

of Rhodobacter capsulatus that overexpress the rnf genes. J Bacteriol

182:1208-14.

11. Li, Q., L. Li, T. Rejtar, D. J. Lessner, B. L. Karger, and J. G. Ferry. 2006.

Electron transport in the pathway of acetate conversion to methane in the marine

archaeon Methanosarcina acetivorans. J Bacteriol 188:702-10.

12. Lienard, T., B. Becher, M. Marschall, S. Bowien, and G. Gottschalk. 1996.

Sodium ion translocation by N5-methyltetrahydromethanopterin: coenzyme M

methyltransferase from Methanosarcina mazei Go1 reconstituted in ether lipid

liposomes. Eur J Biochem 239:857-64.

13. Ma, K., X. Liu, and X. Dong. 2006. Methanosaeta harundinacea sp. nov., a

novel acetate-scavenging methanogen isolated from a UASB reactor. Int J Syst

Evol Microbiol 56:127-31.

14. Muller, V., F. Imkamp, E. Biegel, S. Schmidt, and S. Dilling. 2008. Discovery

of a ferredoxin:NAD+-oxidoreductase (Rnf) in Acetobacterium woodii: a novel

potential coupling site in acetogens. Ann N Y Acad Sci 1125:137-46.

15. Saum, R., K. Schlegel, B. Meyer, and V. Muller. 2009. The F1FO ATP synthase

genes in Methanosarcina acetivorans are dispensable for growth and ATP

synthesis. FEMS Microbiol Lett 300:230-6. 182 16. Suharti, S. de Vries, and J. G. Ferry. 2010. Characterization of the RnfB

subunit of Rnf complex from the Archaeon Methanosarcina acetivorans.

Submitted.

17. Suharti, T. Rejtar, S. de Vries, B. L. Karger, and J. G. Ferry. 2010.

Characterization of a flavin-containing subunit of Rnf complex from the

Archaeon Methanosarcina acetivoras. in preparation.

18. Swartz, T. H., S. Ikewada, O. Ishikawa, M. Ito, and T. A. Krulwich. 2005.

The Mrp system: a giant among monovalent cation/proton antiporters?

Extremophiles 9:345-54.

19. Thauer, R. K., A. K. Kaster, H. Seedorf, W. Buckel, and R. Hedderich. 2008.

Methanogenic archaea: ecologically relevant differences in energy conservation.

Nat Rev Microbiol 6:579-91.

20. von Klein, D., H. Arab, H. Volker, and M. Thomm. 2002. Methanosarcina

baltica, sp. nov., a novel methanogen isolated from the Gotland Deep of the

Baltic Sea. Extremophiles 6:103-10.

21. Wang, M., and J. G. Ferry. 2010. Characterization of a novel electron transport

protein from Methanosarcina acetivorans. in preparation.

22. Wang, M., Suharti, J. F. Tomb, and J. G. Ferry. 2010. The electron transport

chain of acetate-grown Methanosarcina acetivorans. submitted.

183 Appendix A

Alignment of the deduced protein sequences of rnf genes between

Methanosarcina thermophila and Methanosarcina acetivorans

184

185

186

Alignment of the deduced protein sequences of rnf genes between M. thermophila (Mt) and

M. acetivorans (Ma). Highlighted are: conserved heme binding sites (CXXCH and CXXXCH) in Cyt c, the flavin binding motif (SGAT) in RnfG, and cysteine motifs binding iron-sulfur clusters in RnfC and

RnfB. 187

Appendix B

Alignment of the deduced protein sequences of vht and vhx genes between Methanosarcina thermophila and Methanosarcina acetivorans

188

189

190

191

192

Alignment of the deduced protein sequences of vht and vhx genes between M. thermophila

(Mt) and M. acetivorans (Ma). 193

Appendix C

Alignment of the deduced protein sequences of mrp genes between

Methanosarcina thermophila and Methanosarcina acetivorans

194

195

196

Alignment of the deduced protein sequences of mrp genes between M. thermophila (Mt) and M. acetivorans (Ma). 197

VITA

Mingyu Wang

Education:

August 2003-Present Ph. D. candidate in Biochemistry, Microbiology and Molecular Biology program The Pennsylvania State University, University Park, PA. August 1999-June 2003 B.S. in Biology Peking University, Beijing, People’s Republic of China

Research Experience:

January 2004-Present Graduate research assistant in the Department of Biochemistry and Molecular Biology under the guidance of Dr. James. G. Ferry Thesis: The electron transport of acetate-grown Methanosarcina acetivorans August 2002-June 2003 Undergraduate research assistant in the College of Life Sciences under the guidance of Dr. Binggen Ru

Publications

Wang, M., Suharti, J. F. Tomb, and J. G. Ferry. 2010. The electron transport chain of acetate-grown Methanosarcina acetivorans. submitted.

Wang, M., and J. G. Ferry. 2010. Characterization of a novel electron transport protein from Methanosarcina acetivorans. in preparation.

Doerfert, S. N., M. Reichlen, P. Iyer, M. Wang, and J. G. Ferry. 2009. Methanolobus zinderi sp. nov., a methylotrophic methanogen isolated from a deep subsurface coal seam. Int J Syst Evol Microbiol 59:1064-9