MIAMI UNIVERSITY

The Graduate School

Certificate for Approving the Dissertation

We hereby approve the Dissertation

of

Tomislav Ticak

Candidate for the Degree

Doctor of Philosophy

______Director Dr. Donald J. Ferguson

______Reader Dr. Gary R. Janssen

______Reader Dr. Natosha L. Finley

______Dr. Annette Bollmann

______Graduate School Representative Dr. Carole Dabney-Smith ABSTRACT

ANOXIC QUATERNARY AMINE UTILIZATION BY ARCHAEA AND BACTERIA THROUGH A NON-L-PYRROLYSINE METHYLTRANSFERASE; INSIGHTS INTO GLOBAL ECOLOGY, HUMAN HEALTH, AND EVOLUTION OF ANAEROBIC SYSTEMS

by Tomislav Ticak

Quaternary amines are compounds which are important for every domain of life and play roles as carbon and nitrogen sources but also are known to act as osmoregulants. One quaternary amine, glycine betaine, is considered a key osmoregulatory compound due to its chemical nature and is often the main intersection of choline and carnitine metabolism, both aerobically and anaerobically. Many organisms have the capability of degrading glycine betaine through oxygenases or dehydrogenases aerobically, but there is little literature related to the fate of glycine betaine in anaerobic systems. Many of the reported anaerobic systems for glycine betaine involve a reductase pathway that leads to the formation of trimethylamine and acetate, which are well established methanogenic precursor compounds in anaerobic environments. However, there exist a few reports of acetogens and methanogens with the capability of converting glycine betaine to dimethylglycine, which is a strict deviation from the aformentioned reductase pathway. This suggests a pathway exists for anaerobic glycine betaine metabolism that has largely gone uncharacterized. We used a series of bioinformatic, biochemical, and physiological experiments to examine carbon metabolism in Desulfitobacterium hafniense strain Y51 and demonstrated its ability to perform this novel mechanism of glycine betaine metabolism. We proposed that non-L-pyrrolysine trimethylamine methyltransferases may act as quaternary amine methyltransferases. As a result of this study; we discovered a theoretical key in explaining the evolution of the glycine betaine and trimethylamine methyltransferases regarding incorporation of L-pyrrolysine. The fact that a quaternary amine (e.g., glycine betaine) may bind into the near identical location of the proposed trimethylamine-pyrrolysine adducts may help us to better understand this widespread superfamily of methyltransferases. By using our knowledge of the glycine betaine methyltransferase, we began to investigate anaerobic communities for the presence of these methyltransferase genes by enrichments with quaternary amines resulting in the discovery of methanogens capable of glycine betaine, choline, and tetramethylammonium metabolism. Genomic analysis of these organisms revealed the presence of glycine betaine and trimethylamine methyltransferase-like genes supporting the hypothesis of quaternary amine demethylation by non-L-pyrrolysine methyltransferases. Our future work now points toward the examination of microbial distribution and physiology for anaerobic quaternary amine utilization in human systems, marine and freshwater environments to determine the evolutionary pressure(s) that may have selected for the advent of L-pyrrolysine. ANOXIC QUATERNARY AMINE UTILIZATION BY ARCHAEA AND BACTERIA THROUGH A NON-L-PYRROLYSINE METHYLTRANSFERASE; INSIGHTS INTO GLOBAL ECOLOGY, HUMAN HEALTH, AND EVOLUTION OF ANAEROBIC SYSTEMS

A Dissertation

Submitted to the Faculty of Miami University in partial fulfillment of the requirements for the degree of Doctor of Philosophy Department of Microbiology

by

Tomislav Ticak Miami University Oxford, OH 2015

Dissertation Director: Donald J. Ferguson, Ph.D. TABLE OF CONTENTS

LIST OF TABLES iii

LIST OF FIGURES iv

LIST OF COMMON ABBREVIATIONS vii

DEDICATION ix

INTRODUCTION 1

CHAPTER 1. A nonpyrrolysine member of the widely distributed trimethylamine 32 methyltransferase family is a glycine betaine methyltransferase

CHAPTER 2. Isolation and characterization of a tetramethylammonium-degrading 78 Methanococcoides strain and a novel glycine betaine-utilizing Methanolobus strain

CHAPTER 3. Analysis of the function of L-Pyrrolysine within the trimethylamine 125 methyltransferase superfamily (COG5598) by comparison to the glycine betaine methyltransferase

APPENDIX A. Cloning and expression of auxiliary genes predicted to play roles 147 in quaternary amine-dependent methylotrophy in Desulfitobacterium hafniense Y51

CONCLUDING REMARKS AND FUTURE DIRECTIONS 153

REFERENCES 164

ii

LIST OF TABLES Table Page 1 Gene-specific qRT-PCR and cloning primers used in this study 42

2 Comparative analysis of several Methanolobus species to B1d 95

3 Comparative analysis of several Methanococcoides species to Q3c 96

4 Primers for the generation of expression vectors in pSpeedET and pDL05c 150

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LIST OF FIGURES Figure Page 1 Structure of base amine molecules and structures of relevant amine compounds 3

2 Overview of pathways of Car and Cho conversion to GB by microbes 5

3 Proposed mechanism of the glycine betaine/sarcosine/glycine reductase system 8

4 Schematic of methylamine dehydrogenase or oxygenases for the formation of 13 formaldehyde prior to downstream pathways

5 Overview of methanogenic pathways 17

6 Modfied vanillate:THF C1 pathway of D. hanfiense strain DCB-2 19

showing oxidation of a CH3 group from a methylated pterin molecule

7 Proposed methylotrophic models for both archaea and bacteria 21

8 Neighbor-joining 16S rRNA phylogenetic tree of the genus Desulfitobacterium 26

9 Proposed formation of the methylamine-Pyl adducts in the methylamine 30 methyltransferase during enzymatic catalysis

10 The genomic context of mttB genes suggests a role in quaternary amine metabolism 36

11 DSY3156 and DSY3157 were purified to near-homogeneity 45

12 MtgA (DSY3157) is a methylCbl:THF methyltransferase 49

13 Growth of D. hafniense Y51 in the presence of glycine betaine 52 (GB) and either fumarate (A) or nitrate (B)

14 Thin-layer chromatographic analysis of D. hafniense culture supernatants 54

15 Hypothetical pathway for the conversion of glycine betaine 56

(GB) to dimethylglycine and CO2 by D. hafniense Y51

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16 DSY3156 is a glycine betaine:cob(I)alamin methyltransferase 60

17 Michaelis–Menten kinetics of recombinant DSY3156 62

18 Stoichiometric demethylation of GB to produce DMG and methylCbl 65

19 Phylogenetic tree of the COG5598 Superfamily 68

20 Proposed functional relationship between MtgB and MttB 71

21 Gel electrophoresis of 16S rRNA and mcrA products 91

22 Maximum likelihood trees showing the phylogenetic position of strains B1d 93 and Q3c in relation to the most closely related organisms, based on the partial 16S rRNA gene sequence (A) or partial McrA amino acid sequence (B)

23 Microscopic examination of strains B1d and Q3c 97

24 Effect of increasing GB or QMA concentrations on the growth of 100 Methanolobus vulcani B1d and Methanococcoides methylutens Q3c

25 Growth curves are presented showing changes in OD600 as well as quaternary 103 amine and methane concentrations over time for strains B1d (A) and Q3c (B)

26 Subsystem profile of methanogen isolates generated with RAST 105

27 Proposed quaternary amine metabolic schema for methanogens 108

28 Genomic context of putative dimethylsulfide operons for B1d (A) and Q3c (B) 110

29 Maximum-likelihood tree of the COG5589 superfamily amended with non-Pyl 112 and Pyl MttBs from strains B1d and Q3

30 Gene neighborhoods of mttiB genes in strains B1d (A) and Q3c (B). 115

31 Proposed pathway of QMA breakdown in archaea and bacteria 121

32 Proposed mechanism of MMA catalysis by MtmB. 128

33 Structural superpositioning of the MtmB and MetH crystal structures. 130

34 Structure of the Escherichia coli ProX transporter with bound GB 133

35 Structural overlay of Pyl’s location within each MtxB MT 138

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36 Proposed positioning of GB into MtgB apo-structure based on 141 Q-Site Finder analysis

37 SDS/PAGE of heterologously expressed D. hafniense Y51 MttB enzymes 151

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LIST OF COMMON ABBREVIATIONS Name Abb. Quaternary amines QA

Trimethylamine TMA

Glycine betaine GB

L-pyrrolysine Pyl

Methyltransferase MT

Choline Cho

Carnitine Car

Dimethylglycine DMG

Sarcosine MMG

Sulfur Reducing Bacteria SRB

Monomethylamine MMA

One-carbon C1

Dimethylethanolamine DMEA

Dimethylamine DMA

Tetramethylammonium QMA

2-mercaptoethanesulfonate CoM

Knock-out KO

Betaine/Choline/Carnitine Transporter BCCT

Amine/Polyamine/Organocation APC

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Cluster of orthologous genes COG

Corrinoid-binding protein CBP

Cobalamin Cbl

Tetrahydrofolate THF / FH4

Horizontal gene transfer HGT

Last Universal Common Ancestor LUCA

Open reading frame ORF

Site-directed mutagenesis SDM

Gas chromatography GC

Reducing equivalents [H]

Brackish media BM

High-salt media HM

2QNE MtgB

1NTH MtmB

Amino acid aa

Base pair bp

Kilodalton kDa

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DEDICATION

I dedicate this work to all those who supported me through these long years of my graduate career. I thank my friends, mentors, co-workers, and family for continually believing in my capabilities as a scientist and as an educator. I finally would like to dedicate this work to all those who follow after me, as I hope this work will provide you the insights to understanding the microbial one-carbon world that captivated me.

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INTRODUCTION

Quaternary amine (QA) utilization by microbes is well-understood for aerobic QA breakdown whereas anoxic QA pathways are scarcely understood. These pathways reside in a number of different microbial taxa inhabiting environments as diverse as terrestrial and marine sediments, the rhizosphere, and human and animal intestines (Bremer, 1983; Heijthuijsen and Hansen, 1989; King, 1984; Oren, 1990; Prell and Poole, 2006; Smith et al., 1988; Wargo and Hogan, 2009; Wargo, 2013; Ziegler et al., 2010). QA molecules are widely distributed in nature providing a source of carbon and nitrogen for microbes and their hosts but also acting as osmoprotectants (Boncompagni et al., 1999; Časaite et al., 2011; Diaz-Sanchez et al., 2012; Flanagan et al., 2010; Fougère and Le Rudulier, 1990; Goldmann et al., 1991; Mandon et al., 2003; Smith et al., 1988; Wargo, 2013; Ziegler et al., 2010). The existing knowledge related to anaerobic QA metabolism typically points to reductive cleavage of the trimethylamine (N,N- dimethylmethanamine; TMA) moiety and Stickland fermentation reactions leading to acetate formation (Andreesen, 1994; Naumann et al., 1983). However, reports in the literature suggested that there exists an entirely different form of anaerobic QA metabolism (Möller et al., 1984; Mueller et al., 1981), which has gone largely unnoticed to date. During the course of studying these QA pathways, other lines of research began looking into the importance of QAs in relation to global ecology and human health (Tang et al., 2013; Wang et al., 2011, 2014; Watkins et al., 2012; 2014). These timely findings further highlighted the importance of understanding how QAs, more specifically glycine betaine (N,N,N-trimethylglycine; GB), influence microbes but also the hosts with whom they associate. The research presented in this dissertation highlights the first biochemical mechanism by which QAs are anaerobically demethylated by both archaea and bacteria but also strives to explain the function of the rare 22nd amino acid, L-pyrrolysine (Pyl) (Hao et al., 2002, 2004). Pyl exists in only a few proteins and is incorporated in the methylamine methyltransferase superfamilies. To date those members of the methylamine methyltransferases lacking the residue have been considered active methylamine enzymes, however, the requirement for this residue in the enzymatic catalysis suggested that these non-Pyl enzymes might in fact be responsible for other substrates other than methylamines, which will be describe in the context of this research.

1

The chemistry of GB QAs are widely distributed compounds in nature and play important roles for humans, plants, and microbes (Bremer, 1983; Flanagan et al., 2010; Heijthuijsen and Hansen, 1989; Oren, 1990; Prell and Poole, 2006; Ziegler et al., 2010). The structure of QAs consists of a positively charged base nitrogen atom bonded by four alkyl or aryl groups that is similar to other simpler amines (e.g., tertiary, secondary, and primary amines) (Fig. 1). There are many relevant QAs of biological interest, but this dissertation focuses specifically on understanding GB metabolism in archaea and bacteria. Typically, GB is remarked as an osmoprotectant for invertebrates, plants, and microbes due to increased flux of this compound under osmotic stress conditions (Boncompagni et al., 1999; Chen et al., 2010; Diaz-Sanchez et al., 2012; Fougère and Le Rudulier, 1990; Goldmann et al., 1991; Keller et al., 1999; Lai et al., 1999; Mou et al., 2007; Oren, 1990; Smith et al., 1988; Ziegler et al., 2010). The presence of this compound is high when compared to abundant molecules like amino acids in marine organisms (King, 1988). At neutral pH, GB is a zwitterion, meaning it contains both a positive and negative charge which has little effect on intracellular enzymatic reactions, protein structure, or influence on secondary metabolites, thus making it a compatible solute (Welsh, 2000; Ziegler et al., 2010). The derivation of GB stems from conversion of other QAs, such as choline (Cho) and carnitine (Car), which are both ubiquitous components of lipid synthesis systems (Boncompagni et al., 1999; Flanagan et al., 2010; Fougère and Le Rudulier, 1990; Kirsch et al., 2010; Kleber, 2006; Mandon et al., 2003; Smith et al., 1988; Wargo and Hogan, 2009; Wargo, 2013) (Fig. 2). Regarding the distribution of GB in the environment, there are few studies currently looking at the distribution of GB in either marine or terrestrial sediments; furthermore, in areas of osmotic stress the levels of GB can vary highly suggesting preferential use of certain compatible solutes other than GB (King, 1988). In anoxic environments, GB is a known contributor to the presence of TMA and acetate (Andreesen, 1994, 2004; Oren, 1990). This provides interesting context for the basis of this work, as the relationship between GB and TMA will be explored in the anaerobic environment to highlight a relationship that has been well known but might share more evolutionary connections than previously thought in the metabolism of these two compounds.

Known microbial systems of GB catabolism

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Fig. 1. Structure of base amine molecules and structures of relevant amine compounds. (A) Base amine structures are represented here showing lone pairs of electrons for primary, secondary, and tertiary amines and a positive charge state for quaternary amine. R1-R4 represent groups that can include either alkyl or aryl groups. (B) Examples of relavent compounds for this work including methylamines and glycine betaine represented below their base model in (A).

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Fig. 2. Overview of pathways of Car and Cho conversion to GB by microbes. The conversion of Car and Cho to GB is represented by both (a) and (b). Reaction (a) involves formation of GB via carnitine dehydrogenases or hypothetical decarboxylation reactions. Reaction (b) leads to GB conversion by dehydrogenases or monooxygenases followed by aldehyde dehydrogenases to convert betaine aldehyde from the previous step to GB. The fate of GB is show with (c) which is a catalyzed by GdrA to form TMA and acetate. (Wood et al., 2010).

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Disparity between aerobic routes of microbial GB metabolism versus anoxic pathways exists in the literature, but through comparison of these systems it is possible to understand GB catabolism as a whole. The aerobic pathway of GB metabolism is distinct as it is performed through sequential demethylation of GB first to dimethylglycine (N,N-dimethyglycine; DMG), sarcosine (N-methylglycine; MMG), and finally to glycine through oxidases and/or dehydrogenases (Chen et al., 2010; Meskys et al., 2001; Sun et al., 2011; Wargo, 2013). These pathways are widespread in micrboes and in some cases provide not just a means of methionine homeostasis but allow GB to be used as a sole carbon source. In contrast, anaerobic GB metabolism has been demonstrated as a straightforward process in which GB behaves as an oxidant for other amino acids via the Stickland reaction (Andreesen, 1994, 2004; Oren, 1990). The key enzyme responsible for this reaction is the betaine/sarcosine/glycine reductase subunit A (GrdA; EC: 1.21.4.2) (Fig. 3) and through nutritional studies revealed that there was a strict requirement for selenite in media for organisms undergoing this physiological process (Andreesen, 1994, 2004; Meyer et al., 1995). This was later attributed to the requirement for the 21st genetically encoded amino acid (Böck et al., 1991), L-selenocysteine, at a conserved site in this enzyme surrounded by two additional cysteine residues (Andreesen, 2004). This process yields acetate and TMA, which leads to acetate utilization by sulfer reducing bacteria (SRB) and TMA utilization by methanogens (King, 1984). These pathways likely lead to high concentrations of TMA and acetate in anaerobic systems so, despite few investigations targeting specific concentration in marine or saline environments, it can be speculated that these compounds are in relatively high abundance environmentally and provide adequate carbon and nitrogen sources. Interestingly, the same GB reductase system is responsible for the formation of monomethylamine (N-methylamine; MMA) and acetate from MMG and breakdown of glycine to ammonia and acetate (Andreesen, 2004). An alternate fate of glycine, if not degraded via Gdr, is attributed to the glycine cleavage system to further generate reducing power, carbon and nitrogen sources (Kikuchi et al., 2008). However, there have been several bacterial reports in the literature from early investigations describing pathways of DMG and MMG formation that deviate from these aforementioned pathways, which only recently have started to become more apparent in archaea (L’Haridon et al., 2014; Watkins et al., 2014). This demonstrates that GB metabolism is not limited to the dogmatic Stickland reaction and that there may be GB catabolic pathways analogous to that of aerobic GB demethylation systems in both archaea and bacteria.

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Fig. 3. Proposed mechanism of the glycine betaine/sarcosine/glycine reductase system. Protein B is classified as the enzymes require for binding either glycine, MMG, or GB to form a carboxymethyl-selnoether bound to its respective protein, in the case of GB, GrdH. All known pathways converge by transferring the carboxymethyl-selnoether to GrdA ortherwise classified as Protein A which can be reduced by Stickland fermentation of amino acids, thioredoxin (Trx), or dithiotritol (DTT). Then GdrA interacts with Protein C which is an energy-conserving step prior to cleavage of the respective molecule. (Andreesen, 2004).

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This drove us to understand these pathways which preside as a sizeable gap in understanding the catabolism of GB anoxically, as there is no known mechanism of DMG, MMG, or glycine production from sequential GB demethylation.

Analysis of anaerobic demethylation of GB The two earliest reports of GB demethylation begin with Eubacterium limosum and Sporomusa ovata that belong to the Firmicutes phylum. These two acetogens, organisms that use two one-carbon (C1) compounds to form acetate, provide two very different strategies and end products from GB catbolism. E. limosum utilizes 7 GB + 2 CO2 = 7 DMG + 1.5 acetate + 1.5 butyrate, in this way, the methyl groups from GB are used to generate both the acetate and butyrate while fixing CO2 (Mueller et al., 1981). Additionally, there was no breakdown of DMG, MMG, or glycine by E. limosum but there was an interesting finding in which the bacteria could demethylate Cho into dimethylethanolamine (N,N-dimethylethanolamine; DMEA), acetate, and butyrate, thus, demonstrating not one but two different pathways of QA demethylation (Mueller et al., 1981). This finding was, to our knowledge, the first to highlight the deviation from TMA formation from either of these QAs in an anaerobic study; however, one earlier report remarked that pure cultures isolated from Bethesda, Maryland were able to grow using solely GB, DMG, or MMG, however, these cultures were not investigated further beyond their initial isolation (Hayward and Stadtman, 1959). Shortly after the discovery of anaerobic demethylation of GB in E. limosum, a study was performed on a newly isolated gram-negative spore former known as S. ovata. The fermentation balance of S. ovata is far more complex than was seen with E. limosum and the fermentation balance can vary widely between experiments (Möller et al., 1984). GB is fermented into DMG, TMA, acetate, and CO2 (Möller et al., 1984). This fermentation balance begins with oxidation of methyl groups from GB to generate reducing equivalents via CO2 formation followed by reductive cleavage of GB to TMA and acetate. In the case of DMG, the balance takes on several more complex aspects, as DMG is converted to GB, MMG, TMA,

MMA, acetate, and CO2. The formation of TMA is attributed to the initial methylation of DMG to GB followed by reductive cleavage to TMA and acetate, while demethylation to MMG and

CO2 formation is followed by cleavage to MMA and acetate. These balances suggest a lack of reductive DMG cleavage, as dimethylamine (N-methylmethanamine; DMA) is never generated in this study or in any other study involving anaerobic GB catabolism. This is further supported

10 by the lack of a DMG reductase in the Gdr system (Andreesen, 1994, 2004; Meyer et al., 1995). At the time of starting this research, these were the only few insights on novel GB anaerobic pathways deviating from Gdr and only until recently did archaeal-based QA pathways, other than tetramethylammonium (N,N,N-trimethylmethanaminium; QMA), begin to emerge (L’Haridon et al., 2014; Tanaka, 1994; Watkins et al., 2012, 2014).

QAs as direct substrates for methanogens Complex amines have long been known as viable methanogenic substrates, as these compounds lead to the appearance of methanogenic forming communities during enrichment studies (Hippe et al., 1979; King, 1984). Studies showed that the breakdown of complex amines to simpler amines and acetate is the direct cause of methane formation in these enrichment studies. This paradigm of complex amine breakdown by SRB and simplar amine utilization by methanogens has been perpetuated since these initial findings but methanogens may also play a role in a more direct usage of QAs in anaerobic systems. Researchers have, in several investigations, examined new methanogenic isolates for their ability to utilize complex amines (Sprenger et al., 2000; Tanaka, 1994; Watkins et al., 2012, 2014). The first reported QA utilizing methanogen was isolated from marine sediment in Tokyo Bay that was capable of demethylating QMA (Tanaka, 1994). This isolate was named Methanococcoides sp. NaT1, which most closely belongs to the order Methanosarcinales, the only group known to grow methylotrophically (Yuchen Liu and Whitman, 2008) until recently (Borrel et al., 2013). The QMA pathway was partially completed by Thauer and co-workers by isolating apparently homogenous native enzymes that catalyzed demethylation of QMA to TMA to generate methyl-coenzyme M

(methyl-2-mercaptoethanesulfonate; CH3-CoM) (Asakawa et al., 1998). More recently, Sass and co-workers discovered several methanogen isolates capable of utilizing QAs such as Cho and GB (Watkins et al., 2012, 2014). This was further supported by L’Haridon and co-workers as they also described a methanogen capable of utilizing QAs (L’Haridon et al., 2014). Both Sass and L’Haridon’s isolates belong to the Methanococcoides cluster like Tanaka’s strain NaT1, suggesting a wide range of metabolic capability in this methanogen clade. These are interesting findings for several reasons: 1) QAs had long been suspected to not lead to methane formation in pure methanogen culture; 2) it increased the list of known methane forming molecules; 3) it represents the usage of QAs as catabolic substrates in the domain Archaea; and 4) it further

11 demonstrates the existence of a unknown set of pathways for QA demethylation rather than reductive cleavage.

Genomic analysis of known anaerobic GB utilizers and their relatives reveals conserved gene neighborhoods Genome based searches of known GB utilizers such as E. limosum and related Firmicutes showed a lack of Gdr genes encoded by these genomes, but one caveat to this is that many of these organisms lacked complete genomic sequences meaning that regions encoding this system may not have been sequenced or assembled properly. Following the assumption that this system is absent, it reiterates the notion that there exists an unknown pathway(s) related to GB catabolism. To determine what the possibility is for GB catabolism anoxically, it was first important to revisit the aerobic GB metabolism pathways. In aerobic GB catabolism, GB is sequentially demethylated through a series of oxygenases and/or dehydrogenases to generate glycine. There are many anoxic organisms in the Firmicutes that encode dehydrogenases that are homologous to methylamine dehydrogenases. In both aerobic and anoxic systems these amine dehydrogenases are responsible for the breakdown of these simple amines to generate reducing power and formaldehyde (Colby and Zatman, 1973; Kim et al., 2001; Liffourrena et al., 2010) (Fig. 4). These predicted amine dehydrogenases may in fact be QA dehydrogenases as the amine recognizing moiety may be conserved in this class of enzymes which bioinformatic tools likely register as a specific motif. An alternate hypothesis for QAs is that there may be a class of MT like the GB:homocysteine MT that is responsible for demethylation of GB to homocysteine, a known C1 carrier compound, to generate methionine (Barra et al., 2006). This enzyme has long been suspected to be the key step in the aerobic pathway of GB demethylation leading to growth for bacteria; however, knock-out (KO) studies of this enzyme in Sinorhizobium mesoloti demonstrated a methionine auxotrophic phenotype which could be corrected by methionine supplementation (Barra et al., 2006). The mutant still demonstrated an ability to grow with GB, further supporting the existence of an unknown GB pathway. To further examine what enzymes might be involved in GB catabolism it was decided that rather than looking for an unknown enzyme pathway(s) it might be more beneficial to look at operon structures encoding predicted GB transporters. It is not surprising that GB utilizers

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Fig. 4. Schematic of methylamine dehydrogenase or oxygenases for the formation of formaldehyde prior to downstream pathways. The following enzymes are used for the breakdown of methylamines either aerobically or anoxically by methylotrophs catalyzed by; (3) a TMA dehydrogenase leading to the formation of DMA, (4) a DMA monooxygenase causing breakdown to MMA, and (5) the MMA dehydrogenase resulting in ammonia production. All of these enzymes lead to the production of formaldehyde which is then oxidized to CO2 or assimilated. (Wood et al., 2010).

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14 would encode GB transporters; however, there are several different types of GB transporters and there exists little knowledge related to the mechanisms of regulation for osmotic stress versus catabolic usage of GB. Briefly, QAs are transported by a subfamily of transporters known as betaine/choline/carnitine transporters (BCCTs) from the amine/polyamine/organocation (APC) superfamily (Casagrande et al., 2008; Jack et al., 2000; Wong et al., 2012). The functional annotation of these transporters is likely conserved by the motif responsible for direct recognition of the TMA moiety found in QAs, similar to what is seen with amine dehydrogenases. Analysis is difficult for many transporters since they have the ability to transport multiple compounds of similar chemical configuration by which QAs are no exception (Ziegler et al., 2010). Furthermore, without in-depth studies relating mechanism of multiple BCCTs and identifying differentiating features of these transporters it makes it difficult to properly predict what may or may not be a GB versus a Cho transporter. This is further complicated by the fact that there are multiple different types of GB transporters that vary highly in function and to some degree structure in different organisms (e.g., ProX, OpuD, and BCCT) (Ziegler et al., 2010). It is still unknown how these different transporters are selected as there are several large factors at play such as the metabolic fate of the molecule, genetic regulation, driving force, and substrate specificity. Anazlying these transporters lead to one striking observation related to the gene neighborhoods of organisms housing BCCTs, which corresponds to the apparence of MTs known to demethylate methylamines in methanogens, which may be the key finding to understanding the QA pathways.

Methylotrophic growth of archaea and bacteria via methyltransferase pathways The appearance of methylamine methyltransferase genes near BCCT genes in over thirty- two different genera of archaea and bacteria suggested that these methyltransferases may in fact be QA methyltransferases. The TMA MTs found near these BCCTs belongs to the cluster of orthologous genes 5598 (COG5598), which is more commonly known as the MttB superfamily. The first MttB enzyme was isolated from Methanosarcina barkeri and is the only class of enzyme known to be a TMA MT (Ferguson and Krzycki, 1997). In M. barkeri, MttB is highly abundant in cells grown using TMA as the sole substrate, along with MttC which is a protein that binds a corrinoid (corrinoid-binding protein; CBP) a derivative of cobalamin (Cbl), and a secondary MT known as CH3-CBP:CoM methyltransferase (MtbA). These enzymes lead to the

15 formation of CH3-CoM, the penultimate step of methanogenesis (Thauer et al., 2008) (Fig. 5). The mineralization of TMA occurs in a series of demethylation reactions by two analogous pathways that lead to the formation of ammonium (Burke and Krzycki, 1997; Ferguson et al., 2000). The breakdown pathway for TMA is comparable to what is seen with S. ovata during growth with GB. The operons containing MttBs were examined further in these organisms which led to the discovery of predicted CBPs and CH3-CBP:pterin MTs in close genomic proximity to the BCCT and mttB genes, providing multiple lines of evidence pointing to the importance of these putative operons. These data are further supported by the work of Tanaka and Thauer when they investigated the mechanism of QMA metabolism in M. methylutens NaT1. In this system

QMA was catabolized by a substrate-specific QMA:CBP MT, CBP, and CH3-CBP:CoM MT (Asakawa et al., 1998). Further study of this organism was suggested but the loss of the strain prior to the advent of inexpensive and easily accessible genomic sequencing led to a gap in understanding what genes were responsible for this pathway. In this manner the evidence of archaeal QA and methylamine systems seems tightly connected; however, in the case for bacteria, the cofactor CoM is typically not present, so an alternate version of the pathway is required in these systems. The typical lack of CoM in bacteria implies that if this pathway operates similarly to archaea, it must utilize another type of C1 carrier compound in place of CoM. By investigating methylotrophic pathways in bacteria it became apparent that work on other Cbl-dependent pathways may yield insights to this system. Demethylation of methyloxylated compounds by bacteria is initiated through a substrate MT to a CBP which acts as a substrate for a secondary MT (Kaufmann et al., 1998; Naidu and Ragsdale, 2001; Studenik et al., 2012). In the case of bacteria, this secondary MT leads to the formation of methyl-tetrahydrofolate (CH3-THF). CH3-

THF is then subsequently oxidized to CO2 and reducing equivalents, through anoxic respiration, to generating ATP and precursors for growth (Studenik et al., 2012) (Fig. 6). The fate of pterin molecules in both archaea and bacteria are strikingly similar as archaea use modified THF molecules to act as C1 carriers in methylotrophic pathway oxidation or reduction (Berg, 2011; Braakman and Smith, 2012). These data allowed us to form a putative pathway scheme in which a non-Pyl MttB enzyme intiates methyltransfer from a non-TMA substrate in a CBP-dependent manner to a pterin carrier by a secondary MT (Fig. 7). The lack of Pyl which has been shown to be critical in

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Fig. 5. Overview of methanogenic pathways. Colors indicate specific methanogneic pathways in each section: red indicating the hydrogenotrophic pathway, blue is the acetoclastic pathway, yellow represents the methylotrophic pathway, and the green demonstrates the oxidative pathway of methanogens. (https://mcb.illinois.edu/faculty/profile/metcalf/).

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Fig. 6. Modfied vanillate:THF C1 pathway of D. hanfniense strain DCB-2 showing oxidation of a CH3 group from a methylated pterin molecule. Vanillate is demethylated by an O-demethylase (MTI) which transfers the methyl group to a CBP at the Co(I) state. The MTII then demethylates the Co(III) state CBP and transfers the CH3 group to THF (FH4). In the lower portion of the figure the formation of CH3-FH4 is then proposed to be oxidized to CO2 and provide reducing equivilants. (Modified from Kaufmann et al., 1998; Studenik et al., 2012).

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Fig. 7. Proposed methylotrophic models for both archaea and bacteria. MT1 stands for the substrate-specific methyltransferase, the cognate partner to the MT1 being a CBP, and MT2 is for the secondary methyltransferase removing a CH3 group from the Co(III) state CBP to a C1- carrier molecule for either archaea (CoM) or bacteria (THF).

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TMA growth or TMA catalysis (Mahapatra et al., 2006; Joseph Krzycki; personal communication) suggested to us that these non-Pyl MttB enzymes are in fact different functional enzymes on the basis of biochemical and genomic data. This hypothesis forces us to determine the functionality of Pyl within the MttB superfamily and the organisms potentially capable of encoding and decoding Pyl genetically.

Genetic encoding and decoding of Pyl in archaea and bacteria The ability to genetically encode and synthesize Pyl is endowed by an operon known as the Pyl biosynthesis operon (Gaston et al., 2011; Krzycki, 2005; Longstaff et al., 2007). The Pyl operon typically contains five genes, named pylTSBCD, while in some organisms the tRNA snythetase (pylS) is broken up into C- and N-terminal encoding genes known as pylSc and pylSn (Jiang and Krzycki, 2012). This broken up form of the tRNA synthetase is suggested to be a product of horizontal gene transfer (HGT) from archaea to bacteria (Borrel et al., 2014; Fournier et al., 2009; Fournier, 2009). Pyl biosynthesis requires the pylBCD genes while decoding and insertion of Pyl into a growing nacent polypeptide chain requires pylTS or pylTScSn (Blight et al., 2004; Jiang and Krzycki, 2012; Longstaff et al., 2007; Srinivasan et al., 2002). Initial experiments were done with a KO of the pylT gene (Mahapatra et al., 2006) that encodes an amber-decoding tRNA that is charged by the PylS aminoacyl tRNA synthase, which is dissimilar to the decoding strategy of L-selenocystine which is the modification of a cysteine residue (Zhang et al., 2005). Later studies were targeted towards the PylBCD proteins and their role in the biosynthesis of Pyl. In summary, one L-lysine molecule is converted by PylB into (3R)-3- methyl-D-ornithine then PylC utilizes ATP to fuse a secondary lysine to the terminal amine generating (3R)-3-methyl-D-ornithyl-Nɛ-lysine. PylD then oxidizes (3R)-3-methyl-D-ornithyl-

Nɛ-lysine to an aldehyde and H2O thereby forming L-pyrrolysine (Gaston et al., 2011) and was additionally supported by analysis of the crystallized PylBCD enzymes (Quitterer et al., 2012a, 2012b; Quitterer et al., 2013). Originally, the Pyl operon was only identified in a clade of methylotrophic methanogens belonging to the order Methanosarcinales. Large-scale genomic surveys have led to an expansion of information leading to the discovery of twelve bacterial species now known to possess the Pyl operon which is suggested for the expansion of carbon utilization through anaerobic TMA metabolism in these bacteria (Prat et al., 2012). To date there has been no direct

23 evidence showing that the MttB enzyme from bacteria is responsible for the TMA utilization, as biochemical tests have never been reported for a non-archaeal MttB enzyme containing Pyl. It was originally hypothesized that Pyl was a late archaeal development that led to the utilization of methylamines in methanogens for competitive advantage, however, the presence of the Pyl operon in bacteria makes this hypothesis more complicated (Gaston et al., 2011). This suggests that either there was some HGT between archaea and bacteria, or that Pyl predates the last universal common ancestor (LUCA) (Borrel et al., 2014; Fournier et al., 2009; Fournier, 2009). Through analyzing the bacteria that possess Pyl it is apparent that archaea and bacteria utilize some differential regulation of the operon (Prat et al., 2012). In methanogenic archaea with Pyl the selection against the amber codon is quite apparent in open reading frames (ORFs) whereas for bacteria the amber codon usage in ORFs is much higher (Prat et al., 2012). The difference in codon usage is strongly suggestive that archaea have deselected for this codon’s appearance in ORFs as the biosynthesis of Pyl may be costly to an organism’s lysine pools if not regulated. Recent studies related to the split of the tRNA synthetase and similarity of the Pyl genes in gut microbes provides strong evidence that there was HGT events that led to gut archaea and bacteria to acquire the Pyl operon (Borrel et al., 2013). TMA has recently been shown to be a key metabolite responsible for heart disease (Tang et al., 2013; Wang et al., 2011, 2014) and this discovery may prove to be quite significant in understanding human health in terms C1 cycling in the intestines, as the apparence of Pyl and Pyl-containing MttBs has long gone unnoticed in these systems.

The proposed function of Pyl in methylamine methyltransferases Considering that there are few organisms with the theoretical ability to produce Pyl, it is hard to justify the significance of this amino acid outside the fact that these organism have continually retained this operon evolutionarily and that it provides some essentiality to their way of life. To understand the requirement for Pyl it is important to look at what enzymes require it and what those proteins provide to the organisms’ life styles. The Pyl residue was first discovered in the crystal structure of the MMA MT (MtmB) from M. barkeri, which was proposed to be a catalytic residue due to alternate conformation changes during crystallization (Hao et al., 2002, 2004). The genes responsible for DMA and TMA MTs, mtbB and mttB, were also found to contain the amber codon for Pyl in members of the order Methanosarcinales (Paul

24 et al., 2000). Despite using nearly identical chemical compounds there is no sequence similarity between the MtxBs but there is structural homology at the tertiary and quaternary level, furthermore, Pyl is located directly in the proposed catalytic site of each of these proteins (Hao et al., 2002, 2004; Krzycki, 2004). Furthermore, these three enzymes are strictly associated with their substrate and have no known promiscuity for other chemical compounds, unlike what is seen with methoxylated MTs in bacterial systems (Studenik et al., 2012). It is hypothesized that

Pyl moves around freely in the catalytic site where it can position and display the CH3 group of the methylamines for supernucleophilic attack by the Co(I) state CBP (Gaston et al., 2011). The current hypothesis is that each of the MtxB proteins evolved independently to incorporate Pyl, supported by the lack of sequence similarity and their specific interactions with their cognate CBPs. Studies from the Krzycki group show that removal of Pyl through site-directed mutation (SDM) (Joseph Krzycki; personal commuincation) or deletion of the pylT (Mahapatra et al., 2006) leads to drastically decreased activity to methylate their respective CBP or grow with methylamines, which further suggests the essentiality of this residue for catalysis. When the MttB superfamily was investigated more extensively, it became apparent that it may be a more complex family than was originally thought, as the majority of the MttB enzymes have no predicted Pyl residue, as found in bona fide MttBs, and the majority of the organisms containing these non-Pyl MttBs lack the Pyl operon. These non-Pyl MttBs are the enzymes most commonly encoded near the BCCT transporters suggesting they may in fact encode non-TMA MTs. This set the stage to investigate an interesting environmental bacterium known as Desulfitobacterium hafniense, which was originally discovered to be the first bacterium to contain the Pyl operon (Srinivasan et al., 2002).

Overview of the genus Desulfitobacterium and initial studies of D. hafniense Y51 The genus Desulfitobacterium is comprised of five species (Fig. 8) belonging to the Firmicutes phylum most of which are isolated from sites contaminated by halogenated organic molecules (organohalides) (Villemur et al., 2006). The genus is typically comprised of curved rods with lateral flagella that sometimes possess the ability to form terminal endospores (Villemur et al., 2006). The optimal growth conditions range in 25 - 38˚C and a pH range of 6.5 – 7.8 (Villemur et al., 2006). They have a wide range of metabolic capability in the types of

25

Fig. 8. Neighbor-joining 16S rRNA phylogenetic tree of the genus Desulfitobacterium. The scale bar indicates number of base pair subtitutions per site, with bootstrap values highlighted at nodes and the Desulfitobaterium genus is highlighted by a vertical bar to the right. (Villemur et al., 2006).

26

27 electron acceptors utilized and vary on what types of organohalides they can remediate (Villemur et al., 2006). In conditions where nitrate, sulfite, metals, or humic substances are not present they have the ability to dehalogenate organohalides as electron acceptors which requires the use of a Cbl cofactor (Smidt and de Vos, 2004). Each member has the capability of generating its own Cbl cofactors but cofactor is required to be in the media of some species to prevent the loss of dehalogenation capability when not grown under organohalide respiring conditions (Reinhold et al., 2012). Cbl is also used in these species during the breakdown of methyoxylated compounds as previously mentioned for methylotrophic growth (Studenik et al., 2012). This genus generated much attention after D. hafniense’s genome was completely sequenced, representing the first bacterial species to possess the Pyl operon (Srinivasan et al., 2002). Genomic investigation also showed the appearance of BCCT transporters near non-Pyl and Pyl mttB genes marking this as a potentially interesting species of bacteria to investigate for QA growth capability. D. hafniense strain Y51 was isolated from an organohalide contaminated site in Japan (Suyama et al., 2001). The strain was shown to perform organohalide respiration at the expense of pyruvate and lactacte as an electron donor. This strain was also reported to have the capability to perform thiosulfate, sulfite, fumarate, nitrate, nitrite, and metal respiration with pyruvate as an electron donor (Suyama et al., 2001). In terms of electron donor capability nothing else was utilized by this strain making it limited in terms of electron donors but a robust user of electron acceptors. The genome revealed that the ability of this organism to perform organohalide respiration was directly caused by a HGT event by a phage, which is excised from the genome under non-organohalide conditions (Futagami et al., 2006; Reinhold et al., 2012). There had been no reports of D. hafniense using other electron donors at the start of our research, marking this organism as an ideal model for testing QA growth via anoxic respiration. Furthermore, in an enzymatic survey of protein structures by the Joint Center for Structural Genomics (JCSG), one of D. hafniense’s non-Pyl MttB enzymes known as DSY3156 was purified and crystalized prior to our study. This structure has, to date, no known associated ligand (PDB:2QNE). We began tests looking into D. hafniense’s QA substrate profile through thiosulfate respiration coupled with Cho, GB, DMG, and MMG which resulted in growth with all these substrates but never to the level measured with pyruvate (data not shown). Preliminary gas chromatography coupled with a flame ionization detector (GC-FID) was used to investigate GB growth for the lack of methylamines and acetate to rule out Stickland fermentation. The results showed that GB growth

28 led to an absence of both methylamine and acetate formation, however, CO2 was generated during these experiments suggesting oxidation of methyl groups from GB and possibly demethylated derivatives (data not shown).

Hypothesis The metabolic capability of D. hafniense Y51 is not unique as Cho and GB metabolism is well established in Firmicutes, however, the lack of methylamine formation and presence of a complete genome sequence provides us with a novel opportunity to explore this putative QA pathway. The genome of D. hafniense Y51 encodes both amine dehydrogenases and TMA MTs causing us to predict that these may be the genes responsible for the formation of CO2 through GB breakdown. The genomic context of BCCT transporters however provides stronger evidence for catalysis through non-Pyl MttB enzymes. We propose here that the non-Pyl MttBs are directly responsible for the demethylation of complex amines as the lack of Pyl hallmarks these as catalytically different enzymes; furthermore, we predict Pyl to form a chemical adduct with TMA in those enzymes containing Pyl. TMA has little opportunity for hydrogen bonding, but through the formation of a chemical adduct with Pyl (Fig, 9), it would gain a higher likelihood for methyl groups presentation to a CBP. In this proposed mechanism, the TMA-Pyl adduct would generate a novel QA structure that may reveal evolutionary insights to the advent of Pyl within the COG5598 superfamily.

29

Fig. 9. Proposed formation of the methylamine-Pyl adducts in the methylamine methyltransferase during enzymatic catalysis. The dashed arrow indicates the incorportation of Pyl in the MtxB enzymes, at the top of the figure Pyl essentially lacks a chemical adduct due to the absence of methylamine but when methylamine is introduced an addcut is formed designated with “*”. Supernucleophilic attack by a Co(I) state CBP facilitates the mobilization of a methyl group and subsequent realease of the bound methylamine from Pyl, shown at the right portion of the figure. Names in the center indicate the protein family where MtmB = MMA MT,

MtbB = DMA MT, and MttB = TMA MT and R1 and R2 indicated the number of methyl groups or hydrogen present for each substrate / MT pairing. (Modified from Gaston et al., 2011).

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31

CHAPTER 1

A nonpyrrolysine member of the widely distributed trimethylamine methyltransferase family is a glycine betaine methyltransferase

Tomislav Ticak, Duncan J. Kountz, Kimberly E. Girosky, Joseph A. Krzycki, and Donald J. Ferguson

Proceedings of the National Academy of Science. 2014. 111(43): E4668-76.

Category Contribution (%)

Design 75

Experimentation 60

Analysis 50

Writing 35

32

Summary

COG5598 comprises a large number of proteins related to MttB, the trimethylamine:corrinoid methyltransferase. MttB has a genetically encoded pyrrolysine residue proposed essential for catalysis. MttB is the only known trimethylamine methyltransferase, yet the great majority of members of COG5598 lack pyrrolysine, leaving the activity of these proteins an open question. Here, we describe the function of one of the nonpyrrolysine members of this large protein family. Three nonpyrrolysine MttB homologs are encoded in Desulfitobacterium hafniense, a Gram-positive strict anaerobe present in both the environment and human intestine. D. hafniense was found capable of growth on glycine betaine with electron acceptors such as nitrate or fumarate, producing dimethylglycine and CO2 as products. Examination of the genome revealed genes for tetrahydrofolate-linked oxidation of a methyl group originating from a methylated corrinoid protein, but no obvious means to carry out corrinoid methylation with glycine betaine. DSY3156, encoding one of the nonpyrrolysine MttB homologs, was up-regulated during growth on glycine betaine. The recombinant DSY3156 protein converts glycine betaine and cob(I)alamin to dimethylglycine and methylcobalamin. To our knowledge, DSY3156 is the first glycine betaine:corrinoid methyltransferase described, and a designation of MtgB is proposed. In addition, DSY3157, an adjacently encoded protein, was shown to be a methylcobalamin:tetrahydrofolate methyltransferase and is designated MtgA. Homologs of MtgB are widely distributed, especially in marine bacterioplankton and nitrogen- fixing plant symbionts. They are also found in multiple members of the human microbiome, and may play a beneficial role in trimethylamine homeostasis, which in recent years has been directly tied to human cardiovascular health.

33

Significance

Pyrrolysine, the 22nd amino acid, is found in few proteins. One, the trimethylamine methyltransferase MttB, forms a small portion of a large family of proteins. Most in this family lack pyrrolysine and have no known activity. We show that one such protein, MtgB, is a glycine betaine methyltransferase, providing functional context that may explain the relationship between family members with and without pyrrolysine. Close relatives of MtgB are encoded in many of the abundant bacteria in the oceans, as well as different microbes undertaking symbioses ranging from plants to humans. This finding implies that MtgB might partake in a widespread and underappreciated pathway of GB metabolism contributing significantly to global carbon and nitrogen cycling as well as human health.

34

Introduction

Quaternary amines, such as glycine betaine (N,N,N-trimethylglycine or GB), carnitine, and choline are abundant in nature, playing wide-ranging roles in the ecology and physiology of microbes, marine organisms, plants, and animals. GB is an important compound in marine or brackish environments in which it acts as a common compatible solute for many prokaryotic and eukaryotic organisms (Heijthuijsen and Hansen, 1989; Oren, 1990; Ziegler et al., 2010). Carnitine can also function as a compatible solute but is especially abundant in animal tissues due to its role in mitochrondrial transport of fatty acids for energy metabolism (Bremer, 1983; Flanagan et al., 2010). Legume plants make various quaternary amines, some of which are found in the rhizosphere (Prell and Poole, 2006). In many environments, choline results from the breakdown of phosphatidylcholine, ensuring choline as an intermediate of phospholipid breakdown in environments as diverse as lake sediments and the human colon (King, 1984; Wang et al., 2011). GB can be produced as an intermediate during choline and carnitine degradation (Diaz- Sanchez et al., 2012; Wargo, 2013). The route of catabolic bacterial degradation of GB differs markedly depending on the presence or absence of oxygen. Aerobically, various oxidases or dehydrogenases can oxidatively demethylate GB to dimethylglycine (DMG), and then to sarcosine (monomethylglycine or MMG) (Časaite et al., 2011; Meskys et al., 2001; Smith et al., 1988; Wargo et al., 2009; Wargo and Hogan, 2009). Anaerobically, catabolic pathways containing GB reductase cleave GB to acetate and trimethylamine (TMA) as excreted products. GB reduction is often considered the sole fate of the compound under anaerobic conditions (Andreesen, 1994; Chen et al., 2010; Naumann et al., 1983). However, there have been sporadic reports in the literature that GB reduction is not the only route of microbial GB catabolism under anoxic conditions. Some sulfidogenic bacteria have been documented to demethylate GB to

DMG while oxidizing methyl groups to CO2 (Heijthuijsen and Hansen, 1989). Acetogenic bacteria have been documented to use GB as a source for methyl groups and reducing equivalents (Möller et al., 1984; Mueller et al., 1981). The pathway of GB catabolism by such organisms has not been elucidated. Recent work has revealed a surprising connection between anaerobic degradation of quaternary amines and human health. Circulating trimethylamine N-oxide (TMAO) has been

35 implicated in atherosclerosis leading to heart disease and stroke (Tang et al., 2013; Wang et al., 2014). The TMAO has been shown to arise from TMA, which is generated by anoxic gut microbial activity from precursors such as choline, carnitine, and GB (Tang et al., 2013; Wang et al., 2011, 2014). If alternative means of quaternary amine degradation exist that do not generate TMA, the composition of gut microbiota between individuals could moderate the risk of heart disease. A putative oxygenase that demethylates GB has been identified (Wargo and Hogan, 2009), but, thus far, no enzymes that could be active under anaerobic conditions. We have now found a source of such an enzyme in Desulfitobacterium hafniense strain Y51, an anaerobically respiring Gram-positive bacterium. One strain of D. hafniense, DP7, was isolated from the human intestine (Van de Pas et al., 2001) and lacks the ability to carry out organohalide respiration, a hallmark of the rest of this species (Villemur et al., 2006). D. hafniense is one of few known microbes that carry the pyl genes to enable the biosynthesis and genetic encoding of pyrrolysine (Pyl), the 22nd amino acid (Hao et al., 2002). Organisms with the pyl genes also carry examples of Pyl-containing methyltransferases (Gaston et al., 2011). D. hafniense has an mttB gene with a pyrrolysine codon (pyl-mttB). Pyl-MttB proteins are the only known gene products to carry out TMA-dependent corrinoid methylation. Interestingly, the genome of D. hafniense Y51 encodes three homologs of mttB that lack pyl codons and are thus predicted to form non-Pyl MttB proteins (Nonaka et al., 2006). Each of these non-Pyl mttB homolog genes is found in close proximity in the chromosome to a gene encoding a member of the betaine/choline/carnitine transporter (BCCT) family as well as genes encoding predicted corrinoid-binding proteins and methylcorrinoid:pterin methyltransferases (Fig. 10). This genomic proximity led us to hypothesize that non-Pyl MttB homologs function as corrinoid-dependent quaternary amine methyltransferases. We found that the DSY3156 gene encoding a non-Pyl MttB of D. hafniense is up-regulated during growth on GB, and the gene product is a GB:corrinoid methyltransferase, an enzymatic activity that has not been previously reported. These results reveal the function of a non-Pyl MttB for the first time, to our knowledge, and also a functional relationship that may underlie the evolutionary history of pyrrolysine and the large MttB superfamily. A large number of homologs of the GB methyltransferase are found encoded in available genomes and metagenomes and indicate a previously underappreciated pathway of quaternary amine oxidation to CO2, as also supported by the function of DSY3157 as

36 a methylcorrinoid:tetrahydrofolate methyltransferase. This pathway is likely present in environments as diverse as the oceans, the rhizosphere, and the human colon.

37

Fig. 10. The genomic context of mttB genes suggests a role in quaternary amine metabolism. The clustering of MttB family member genes (blue) with genes predicted to encode corrinoid-binding proteins (red), methylcorrinoid:pterin methyltransferases (green), and BCCT transporters (purple) suggests a role of non-Pyl MttB enzymes in the methylotrophic metabolism of quaternary amines. Gene names of nearest homologs are located within the indicated genes, with loci designations in the D. hafniense Y51 genome listed below. RamA functions in corrinoid protein activation whereas MtrH is a methylcobalamin:THF methyltransferase. HyuA is a predicted D-phenylhydantoinase. Pyl indicates the position of the amber codon encoding the pyrrolysine residue in DSY4970, a homolog of the bona fide TMA methyltransferase. The small arrows indicate the location of primers used for the qRT-PCR experiment.

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39

Materials and Methods

Bacterium Strain and Growth Conditions. D. hafniense Y51 was a kind gift from Taiki Futagami (Department of Bioscience and Biotechnology, Kyushu University, Fukuoka, Japan) and was routinely cultured with a modified DSMZ 720 medium specified by the Leibniz-Institut DSMZ-Deutsche Sammlung von Mikroorganismen und Zellkulturen GmbH (www.dsmz.de/home.html). The modified DSMZ 720 medium designated as MM1 uses a nonchelated SL-10 trace elements solution and lacks NaHCO3. The pH was adjusted to 7.2, and ultra-high purity nitrogen was the sole gas phase. Potassium phosphate, pH 7.2, was added to the medium at a final concentration of 22 mM after autoclaving. D. hafniense Y51 was cultivated anaerobically in the presence of either 25 mM filter-sterilized sodium pyruvate or 25 mM filter- sterilized glycine betaine monohydrate to act as both an electron donor and carbon source. Cultures were also supplemented with 50 mM sodium fumarate or 50 mM sodium nitrate to act as an external electron acceptor. Complete MM1 was inoculated with 0.5–1.0% (vol/vol) stationary phase cultures and grown statically, in the dark, at 37 °C. All O.D. values were measured at 600 nm. All chemical reagents were obtained from either Fisher Scientific or Sigma- Aldrich with a purity ≥ 98%.

Product Stoichiometry of GB Degradation by D. hafniense. Liquid and gas phase samples were taken shortly after inoculation into a total of 11 mL of MM1 medium in Balch anaerobe tubes (27 mL total volume). MM1 medium lacked sodium sulfide and was supplemented with 3 mM methionine, 3 mM cysteine, and 2 mM DTT. The tubes were also amended with 10 mM GB and 50 mM sodium nitrate. Samples were again taken when the optical density reached ∼0.1 units. Liquid samples were analyzed for GB, DMG, and MMG by TLC as described in Analysis of GB and Demethylation Products. The culture was then acidified by addition of 1 mL of 1 M HCl. Total culture carbon dioxide was then determined from gas-phase measurements using a thermal conductivity detector-equipped model 8A gas chromatograph (Shimadzu Scientific) with a Carbosieve S-II (Sigma-Aldrich) column eluted with a flow rate of 45 mL/min He with injector, detector, and oven temperatures at 120 °C. Cultures incubated without GB did not produce significant amounts of CO2 above background (determined by total CO2, the amount

40 present in T0 acidified controls). TMA and acetate were analyzed by gas chromatography as described previously (Burke and Krzycki, 1995; Kremer et al., 1993).

Analysis of GB and Demethylation Products. The GB, DMG, or MMG present in samples was determined by TLC (Lai et al., 1999). A total of 15 µL of standard or unknown sample was spotted onto Silica Gel 60 plates (Merck-Millipore). Plates were then developed in 80% (vol/vol) phenol in H20, dried, and stained with 0.04% (wt/vol) bromocresol green (titrated to dark green with a 0.1-M NaOH solution). The plates were scanned, and then the entire image was converted to monochrome before densitometry. Standard curves were linear between 25 nmol and 125 nmol of analyte, and the volumes of unknown samples were adjusted to fit the analyte concentration within this range.

RNA Isolation and Quantitative Real-Time PCR. D. hafniense Y51 cultures were harvested at the middle of exponential phase (O.D. 600 circa 0.450 for pyruvate and 0.250 for betaine cultures) and centrifuged aerobically at 7,500 × g at 4 °C for 15 min. RNA was extracted using a modified total RNA isolation protocol for RNAzolRT (Molecular Research Center, Inc.) involving lysing cells with RNAzolRT heated to 95 °C with repeated vortexing for 5 min followed by cooling samples on ice for 5 min. RNA samples were treated for genomic DNA contamination with TURBO DNA-free DNase (Ambion). The RNA quality was evaluated and quantitated with a BioAnalyzer 2100 (Agilent Technologies) using an Agilent RNA 6000 Pico Kit according to the manufacturer’s instructions. Isolated RNA samples had RNA integrity numbers (RINs) in the range of 7.9–9.1. Gene-specific primers (Table 1) were generated using Primer3 (Rozen and Skaletsky, 2000). Each primer was used as a query against the D. hafniense Y51 genome in a BLASTn search to ensure that only a single region of significant sequence identity was detected (Altschul et al., 1990). Each qRT-PCR (10 µL) was performed using isolated total RNA from either pyruvate or GB-grown cultures with the iScript One-Step RT-PCR Kit with SYBR Green (Bio- Rad Laboratories, Inc.). The complete master mix for each PCR containing iScript reverse transcriptase and SYBR Green RT-PCR mix was used as per the manufacturer’s recommendation (Bio-Rad Laboratories, Inc.). Reactions contained 200 nM forward and reverse gene-specific primers and 10 ng of total RNA. Negative control reactions were performed by

41

Table 1. Gene-specific qRT-PCR and cloning primers used in this study

Name Sequence Study rpoB_qRT_F 5’- ACAAGAGGTCTTCATGGGTGAT -3’ This study rpoB_qRT_R 5’- CCGTTTCAAACTCTAACCAAGC -3’ “

DSY3156_qRT_F 5’- AAAAGCCGGTCTTAGTCTCCTT -3’ “

DSY3156_qRT_R 5’- AAGAGATTCCGGCTAAGACCTC -3’ “

DSY3648_qRT_F 5’- AACACAGCTTATGTGGGTCCTT -3’ “

DSY3648_qRT_R 5’- GGGTCTGATAAACCATCTCCAG -3’ “

DSY4970_qRT_F 5’- GGAAGCCTTGGAGATTTTCTCT -3’ “

DSY4970_qRT_R 5’- ATGGAAACACCTTCACCAAAGT -3’ “

DSY4971_qRT_F 5’- GCCGGTTTATATTCAGGATTTG -3’ “

DSY4971_qRT_R 5’- CGATATTCGGTTTATCCGTGTT -3’ “

DSY3149_qRT_F 5’- TTGTCGTATCTGCTGATTCCTG -3’ “

DSY3149_qRT_R 5’- AATGAACAAGAAGGGAAATCCA -3’ “

DSY3154_qRT_F 5’- CCGACTCAGCTACCTACGTTCT -3’ “

DSY3154_qRT_R 5’- CAGCATGATCAGCATAAAAGGA -3’ “

DSY3643_qRT_F 5’- GGGTTTTATGAAAAGTGCAAGC -3’ “

DSY3643_qRT_R 5’- ACCAATACCAAAGGCAACAAAG -3’ “

DSY5010_qRT_F 5’- ATCGGCTGTTCACTGTTCTTCT -3’ “

DSY5010_qRT_R 5’- ATCGGCTGTTCACTGTTCTTCT -3’ “ pSpeedET_DSY3157_F 5’- ctgtacttccagggcATGTTCAAATTTACTGCTCAACAACATG -3’ “ pSpeedET_DSY3157_R 5’- aattaagtcgcgttaGAAAATTTTGAGCAAAGGATGCTCTG -3’ “ pSpeedET_IF 5’- taacgcgacttaattAAACGGTCTCCAGCTTGGCTGTTTTGGC -3’ (Klock and Lesley, 2009) pSpeedET_IR 5’- gccctggaagtacagGTTTTCGTGATGATGATGATGATG -3’ (Klock and Lesley, 2009)

Lowercase nucleotides represent complementary 5′ sequences between the target insert and PIPE cloning vector which allow for the annealing and formation of new plasmids without a subsequent ligation step (Klock and Lesley, 2009).

42 omitting either RNA or gene-specific primers. Three independent sets of samples were done in triplicate for each gene of interest and for the negative controls. The reactions were run on a CFX Connect 5200 (Bio-Rad Laboratories, Inc.) thermocycler in Hard-Shell 96-well plates (Bio-Rad Laboratories, Inc.) using the following conditions: 50 °C for 10 min., 95 °C for 5 min., followed by 34 cycles of 95 °C for 10 s, 55 °C for 30 s, and a plate-reading step before each cycle. A melt- curve analysis was also performed under the following conditions: 95 °C for 10 s, and 65 °C to 95 °C ramp with a step increase of 0.5 °C/5 s followed by a plate read before each incremental step. The Bio-Rad CFX Manager 3.0 (Bio-Rad Laboratories, Inc.) was used to obtain and analyze the qRT-PCR run data to calculate quantification cycles (Cq) of each reaction. Cq values were averaged between triplicate reactions and then statistically analyzed between independent −ΔΔC replicates to obtain 1σ and then normalized to rpoB using the 2 T method (Livak and Schmittgen, 2001).

DSY3156 and DSY3157 Production and Purification. The expression vector for DSY3156, pSpeedET_DSY3156, maintained in E. coli DH5α was obtained from the DNASU plasmid repository (http://dnasu.org/DNASU/). DSY3157 was cloned using the polymerase incomplete primer extension (PIPE) cloning technique (Klock and Lesley, 2009) using the primers shown in Table 1. The clone was made in the EC100 strain of E. coli. Both plasmids were purified from cells grown in Luria–Bertani (LB) supplemented with 50 µg/mL kanamycin (Kan) using a Wizard Plus SV Miniprep kit as per the manufacturer’s directions (Promega). Chemically competent E. coli BL21(DE3) cells (Life Technologies) were transformed with pSpeedET_DSY3156 and pSpeedET_DSY3157, separately, and grown on LB agar supplemented with 50 µg/mL Kan. Five-milliliter starter cultures of E. coli containing each clone were grown overnight at 37 °C with shaking and used for inoculation of 1 L of LB broth supplemented with 50 µg/mL Kan and placed into an incubating shaker at 37 °C rotating at 250 rpm until they reached an O.D. 600 of ∼0.5. The cultures were induced with L-arabinose at a final concentration of 0.1% (wt/vol) for 4 h at 37 °C shaking at 250 rpm. Cells were harvested by centrifugation at 7,500 × g at 4 °C for 15 min and then washed with Buffer A (50 mM sodium phosphate, 500 mM NaCl, and 40 mM imidazole at pH 7.2) and spun at 7,500 × g at 4 °C for 15 min. The wet-cell weight for each culture was ∼6.5 g and was suspended in 10 mL of Buffer A

43 before lysing the cells three times via French Press at 20,000 psi. The cell lysate was then spun at 250,000 × g at 4 °C for 1.5 h, and the soluble fractions were isolated from the insoluble fractions via decanting. The cell lysates were loaded on 1-mL HisTrap columns (GE Healthcare) equilibrated with Buffer A at 0.35 mL/min and then washed with 10 mL of Buffer A before the linear gradient. A 40-mL gradient was applied to the column from 0% to 100% Buffer B (50 mM sodium phosphate, 500 mM NaCl, and 500 mM imidazole at pH 7.2) at 0.75 mL/min. DSY3156 eluted in a single peak between 70 mM and 200 mM imidazole. DSY3157 eluted in a single peak between 160 mM and 260 mM imidazole. The eluted fractions were pooled and quantitated via Bradford reagent (Bradford, 1976). The purity of recombinant DSY3156 and DSY3157 was assessed by SDS-polyacrylamide gel electrophoresis (SDS/PAGE) (Fig. 11). SDS/PAGE was performed in accordance with the method of Laemmli with a Minislab electrophoresis system (Idea Scientific) (Laemmli, 1970).

Spectrophotometric Assay of GB:Cob(I)alamin Methylation. Reaction mixtures were assembled in anaerobic 0.2-cm quartz cuvettes under a 2-psi stream of either H2 (100%) or N2 (100%) gas with the following components: 50 µg of purified recombinant DSY3156 (1.6 μM), 50 mM glycine betaine, 16.25 mM Ti(III) citrate, 1.75 mM hydroxocobalamin in 50 mM Mops, pH 6.5, in a final volume of 0.6 mL. A mixture excluding both hydroxocobalamin and DSY3156 was used to blank the HP 8453 Diode-Array Biochemical Analysis Spectrophotometer (Hewlett- Packard Development Company). Hydroxocobalamin conversion to cob(I)alamin was monitored using ΔA540 and ΔA578 after the addition of hydroxocobalamin. When conversion to cob(I)alamin was complete, the assay was initiated with either recombinant DSY3156 or GB injection. The reactions were performed under dim red light at 37 °C for 10 min with UV-Vis spectra recorded every 30 s. The formation of methylCbl from cob(I)alamin was monitored using Δ540 with an extinction coefficient of 4.4 mM−1 • cm−1 (Kreft and Schink, 1994). MethylCbl was not generated in the absence of recombinant DSY3156 or GB in these assays. Assays with alternative substrates included either 50 mM tetramethylammonium, trimethylamine, dimethylamine, monomethylamine, dimethylglycine, monomethylglycine, γ-butyrobetaine, choline, or carnitine as substrates using solutions first adjusted to pH 6.5. The apparent kinetic

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Fig. 11. DSY3156 and DSY3157 were purified to near-homogeneity. (A) A 10% (wt/vol) acrylamide discontinuous SDS/PAGE gel was loaded with a protein standard (Std) having the molecular masses indicated in kDa to the side. Purified MtgB (DSY3156, 4 μg) was loaded in the adjacent lane. (B) Similarly, in a 12.5% (wt/vol) SDS/PAGE, a protein standard (Std) with indicated molecular masses in kDa was loaded in the left hand lane whereas MtgA (DSY3157, 6 μg) was loaded into the adjacent lane.

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46 parameters of MtgB were investigated by varying the GB concentration from 0.25 mM to 50 mM. Results were repeated in triplicate with three independently purified preparations of recombinant MtgB. The results were analyzed using Prism 6 (GraphPad) and fitted with a nonlinear Michaelis–Menten algorithm. The stoichiometry of GB consumed to DMG and methylCbl produced was determined in assays as described above except containing 18 mM potassium phosphate, pH 7.0, 5 mM Ti(III) citrate, 1.8 mM GB, and 50 μg/mL MtgB in a total volume of 560 μL under an N2 gas phase. MethylCbl formation was followed spectrophotometrically, and the reaction was terminated at a fixed time point by removal of the cuvette contents and application to an aerobic C18 1-mL Sep- Pak (Waters Corp.) that had been previously eluted with 10 mL of 100% methanol and then equilibrated with 10 mL of deionized H2O. The flow-through was collected, then the column was further eluted with a total of 1.4 mL deionized H20. The fractions were concentrated in a rotary evaporator and quantitatively combined before final evaporation to dryness and dissolution in a measured volume of deionized H2O for analysis of GB and DMG by the TLC method described above. The total amount spotted on the plates was varied as necessary for the unknown to be within the linear range of the GB and DMG standard curves.

COG5598 Phylogenetic Tree and Sequence Acquisition. The DSY3156 sequence was used as a query in a BLASTp search of the Microbial Genome database maintained at the National Center for Biotechnology Information (Altschul et al., 1990). Full-length pyrrolysine-containing proteins were identified using the same database but with a tBLASTn using Methanosarcina barkeri MttB as a query. These entire ORFs were translated to include the pyrrolysine residues and then added to the database to give 900 sequences. The larger dataset was reduced with the ElimDupes program (hcv.lanl.gov/content/sequence/ELIMDUPES/elimdupes.html), yielding 515 sequences that were at a maximum 90% similar to each other, except all copies of MttB homologs from D. hafniense Y51 were retained. The truncated dataset was aligned with MUSCLE (Edgar, 2004) using default setting in MEGA6 (Tamura et al., 2013). This dataset was tested for the highest Bayesian information criterion (BIC), corrected Akaike information criterion (AICc), and maximum likelihood values (InL) in MEGA6. Molecular phylogeny was inferred using the maximum likelihood method based on the LG model (Le and Gascuel, 2008) with a discrete gamma distribution used to estimate rates among sites [five categories (+G =

47

3.4371)]. Gaps and missing data were given a 95% cutoff and then partially deleted from the analysis of the 515 sequences. There were a total of 439 residue positions present in the final analysis. The tree is presented with a log likelihood value of −249096.7688 and was initialized using the neighbor-joining method to a pairwise distance matrix calculated from the JTT model (Jones et al., 1992) and was computed for 1,000 bootstrap repetitions. The tree was drawn to scale with the scale bar representing 0.5 substitutions per site. A radiation tree is presented with larger groups being collapsed, with bootstrap percentages being located at nodes where applicable; those values less than 50% are hidden. Groups of interest were colored green and red, and large assignments of phylum or orders that make up groups were clustered together.

Assay of MethylCbl:THF Methylation by DSY3157. Spectral determination of the rate of THF methylation by methylCbl was performed essentially as described by Ragsdale and coworkers (Roberts et al., 1994) using the photodiode array spectrophotometer described above. Briefly, spectra were collected every minute after initiation of the reaction, and the THF-dependent demethylation of methylCbl was monitored at 525 nm using Δε = 8.6 mM−1 • cm−1. Assays were conducted in a 0.2-cm path length cuvette containing 0.58 mL of assay volume at 37 °C with a nitrogen gas phase. The complete assay contained 0.42 mM methylCbl, 0.87 mM THF, 6.7 mM DTT, and 22.4 μg of MtgA (Fig. 12B) in 20 mM potassium phosphate buffer, pH 7.0. The production of methyl-THF was confirmed using HPLC. Aliquots of the spectral assays (20 μL) were injected onto a 250 × 4.6-mm Varian Microsorb MV-100 C18 column on a Dionex UltiMate 3000 HPLC system. The column was eluted isocratically with 7% (vol/vol) acetonitrile in 30 mM potassium phosphate buffer, pH 3.0, at 1 mL/min, and monitored at 272 nm. The standards included THF, which eluted at ∼9.5 min and methyl-THF (Santa Cruz Biotechnology), which eluted at ∼12.5 min in this system.

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Fig. 12. MtgA (DSY3157) is a methylCbl:THF methyltransferase. Spectral and HPLC analysis of the MtgA reaction showed that it catalyzes methylCbl demethylation and production of methyl-THF. Details of both analyses are given in Materials and Methods. (A) To calculate the rate of the reaction, the absorbance spectrum of methylCbl was monitored as it was demethylated by enzyme incubated in the presence of THF. In this assay, the cob(I)alamin product is converted to cob(II)alamin due to the presence of a mixture of oxidized and reduced dithiotreitol. The redox buffer then stabilizes the Co(II) form, allowing rate measurement at 525 nm using the Δɛ between methylCbl and cob(II)alamin. (B) The MtgA reaction rate was obtained from the initial linear portion of the curve measured in the complete reaction. The slight background rate observed in the no-enzyme control we attribute to photolysis by the light beam of the spectrophotometer. The MtgA rate was corrected for contribution by this nonenzymatic reaction. In (C) and (D), a single time point at 45 min was removed from spectral assays conducted as in B. At this point, the reaction had fully ceased as indicated by no further change in the spectrum. The aliquots then were analyzed by reverse-phase HPLC. Reactions incubated in the absence of enzyme (C) revealed only a single large peak that has the same retention time as authentic THF standard whereas analysis of the complete reaction reveals a second peak that elutes at the position of methyl-THF (D). The methyl-THF peak was not detectable in otherwise complete reactions lacking methylCbl or THF. It should be noted that cobalamin derivatives do not elute from the column using the 7% (vol/vol) acetonitrile elution buffer employed.

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50

Results

D. hafniense Uses GB as a Methylotrophic Growth Substrate. D. hafniense has not previously been thought to use quaternary amines as substrates. However, GB supported growth when added to culture medium having an exogenous electron acceptor such as fumarate (Fig. 13A). We noticed a low, but significant growth on fumarate in the absence of GB and therefore also tested inorganic electron acceptors. We found that GB also supported growth with nitrate (Fig. 13B). Continued cultivation of GB cultures was dependent on the presence of both nitrate and GB. Yeast extract was stimulatory to growth but not essential and could not replace GB.

However, CO2 could not replace fumarate or nitrate during growth on GB, suggesting that

CO2 reduction for acetogenesis did not underlie growth on GB. Acetate was also not detected during growth. The dependence on exogenous electron acceptors suggested that GB was serving as an electron donor; therefore, cultures were examined for possible degradation products of GB using TLC. In nitrate cultures at midlog, GB was demethylated to DMG (Fig. 14). We measured the ratio of DMG produced to GB consumed as 0.91 ± 0.18 (n = 5). In keeping with the absence of GB reductase in the genome, we could not detect trimethylamine using gas chromatography in growing cultures. Instead, we found that GB-metabolizing cultures also produced CO2 with a

DMG:CO2 ratio of 1.1 ± 0.16 (n = 6). CO2 was not significantly produced when GB was not added to the medium. These results are consistent with a stoichiometry of ∼1 GB → 1 DMG + 1

CO2. We conclude that D. hafniense is capable of methylotrophic growth in which a methyl group of GB is oxidized to CO2 to provide reducing equivalents for anaerobic respiration. Additionally, we noted in some cultures given lower amounts of GB (<5 mM), a compound comigrating with monomethylglycine in TLC was also detectable in late log cultures, suggesting that under some circumstances GB could be further demethylated to form monomethylglycine. qPCR Analysis of Non-Pyl mttB Genes. Examination of the genome of D. hafniense Y51 revealed a possible pathway for conversion of a methyl group to CO2 on a tetrahydrofolate (THF) cofactor (Fig. 15) but supported no route from GB to methyl-THF. However, the genomic context of the genes encoding three non-Pyl MttBs in D. hafniense supported our hypothesis that these proteins may be quaternary amine:corrinoid methyltransferases. Each non-Pyl MttB is

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Fig. 13. Growth of D. hafniense Y51 in the presence of glycine betaine (GB) and either fumarate (A) or nitrate (B). In (A), the numbers above each growth curve refer to the concentration of glycine betaine in mM at the start of incubation. All cultures also contain 50 mM fumarate, unless indicated. In (B), cultures contained 20 mM glycine betaine or 50 mM nitrate, as indicated.

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Fig. 14. Thin-layer chromatographic analysis of D. hafniense culture supernatants. Lanes were spotted at the origin (Ori.) with aliquots of supernatants from (lane 1) uninoculated tube at

T0; (lane 2) uninoculated culture tube after 190 h incubation at 37 °C; (lane 3) stationary phase culture used to inoculate culture in lane 4; (lane 4) newly inoculated culture, O.D. 600 = 0.0; (lane 5) culture after 83 h incubation at 37 °C, O.D. 600 = 0.12; and (lane 6) culture after 190 h, O.D. 600 = 0.19. Arrows indicate relative migration positions of authentic standards including succinate (Suc.), fumarate (Fum.), GB, and dimethylglycine (DMG).

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Fig. 15. Hypothetical pathway for the conversion of glycine betaine (GB) to dimethylglycine and CO2 by D. hafniense Y51. Three MttB family members were considered as candidates for GB:corrinoid methyltransferases (blue oval), which could act in concert with the adjacently encoded corrinoid protein and methylcorrinoid:tetrahydrofolate (THF) methyltransferase (red and green ovals, respectively) to form three-component systems for the methylation of tetrahydrofolate with GB. The locus designations for each of the three non-Pyl MttB proteins is found above the colored ovals, and the putative cognate corrinoid proteins and corrinoid:THF methyltransferases are listed along beside each MttB protein. The bolded methyltransferase DSY3156 was found to have GB:corrinoid methyltransferase activity whereas DSY3157 was shown to have methylcorrinoid:THF methyltransferase activity. The bottom portion illustrates a hypothetical pathway for the oxidation of the methyl group via folate intermediates as carried out by activities predicted for each of the indicated proteins encoded in the genome. Reducing equivalents ([H]) gained via the oxidation of the methyl group can be passed to the electron transport chain for reduction of the anaerobic electron acceptor. ETC, electron transport chain; FHL, formate-hydrogen lyase complex; Hup, uptake hydrogenase.

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57 encoded in gene clusters that are potential transcriptional units and has, within the unit or nearby, a gene encoding a member of the BCCT family of proteins (Ziegler et al., 2010). Additionally, each gene cluster encodes homologs of a methylotrophic corrinoid protein (Krzycki, 2004) and a methylcorrinoid:THF methyltransferase (Naidu and Ragsdale, 2001; Studenik et al., 2012). Such proteins are part of the systems that initiate catabolism by Gram-positive bacteria such as D. hafniense during growth on methoxylated aromatic compounds (Naidu and Ragsdale, 2001; Studenik et al., 2012). So as to determine whether one of the non-Pyl mttB genes might be involved in GB metabolism, transcript abundance was compared by qPCR in cells grown on either pyruvate or GB. Transcripts of the mttB genes (DSY3156, DSY3648, DSY4970, and DSY4971) as well as those encoding the putative BCCT genes (DSY3149, DSY3154, DSY3643, and DSY5010) were targeted at the locations indicated in Fig. 10. The melt curves indicated the formation of single amplified products for each gene, which was supported by gel electrophoresis. In pyruvate- adapted cultures, each of the transcripts of the mttB homologs was in lower abundance than the housekeeping gene rpoB. This trend is unchanged in GB-grown cells for most mttB transcripts, the exception being DSY3156, which became more abundant than rpoB. Analysis using the −ΔΔC 2 T method (Livak and Schmittgen, 2001) indicated that the DSY3156 transcript is 27- to 58- fold higher in GB-grown cultures relative to pyruvate, and this difference was significant (P ≤ 0.001). Similarly DSY3154, the predicted BCCT gene located near DSY3156, increased above rpoB levels in cells grown on GB relative to pyruvate; with DSY3154 23- to 42-fold higher in GB- versus pyruvate-grown cells, and this result also was significant (P ≤ 0.001). Transcripts encoding the other non-Pyl MttB homologs and BCCT proteins were not elevated to this extent during growth on GB. For example, DSY3648 and DSY3643 were only slightly changed with GB relative to pyruvate, with the DSY3648 having a 0.77- to 2.3-fold difference and DSY3643 having a 1.1- to 2.8-fold increase in transcript abundance compared with pyruvate. The adjacent BCCT gene, DSY3149, showed a significant induction, with GB of 2.2- to 7.3-fold compared with pyruvate (P ≤ 0.001). In contrast, expression of other mttB and BCCT genes was actually lower on GB relative to pyruvate, with DSY4970 ∼0.07- to 0.41-fold lower and DSY4971 0.15- to 0.48-fold lower. This trend is also seen with DSY5010, the BCCT transporter gene nearest DSY4971, which showed a 0.34- to 1.34-fold difference in GB-grown cells relative to those grown on pyruvate. Overall, the expression patterns supported our

58 hypothesis and indicated that non-Pyl mttB gene DSY3156 and BCCT gene DSY3154 could play important roles during growth on GB.

DSY3156 Is a Glycine Betaine:Cob(I)alamin Methyltransferase. Proteins encoded adjacent to DSY3156 include a small corrinoid protein homologous to MttC, the cognate corrinoid protein of the pyrrolysine-containing MttB from Methanosarcina spp., suggesting that the DSY3156 protein might act to methylate corrinoid cofactors. If so, this activity would explain the up- regulation of the DSY3156 gene during growth on GB because this function would provide a path to methylate THF and thereby initiate a route to formation of CO2. Several different methyltransferases have been shown to methylate cob(I)alamin not bound to protein (Burke and Krzycki, 1997; Goulding et al., 1997; Sauer and Thauer, 1999), and, therefore, we tested the ability of the recombinant DSY3156 protein (Fig. 11A) to methylate cob(I)alamin with various substrates. Quaternary amines such as carnitine and choline did not serve as substrates, nor did tertiary amines such as dimethylglycine or trimethylamine. Instead, DSY3156 protein carries out a robust methylation of cob(I)alamin in the presence of GB. Fig. 16A depicts the changes in the visible spectrum of cob(I)alamin as it is methylated in a reaction dependent on the presence of both GB and DSY3156. The presence of a clear isosbestic point at 578 nm indicates that cob(II)alamin was not generated by adventitious oxidation to a significant extent during the reaction, and that cob(I)alamin and methylCbl were the only detectable forms of cobalamin in the assay. Therefore, the rate of the MtgB-catalyzed reaction can be quantified by the increase in absorbance at 540 nm (Fig. 16B). No change in rate was detected when the reactions were performed under H2 or N2. DSY3156 carried out the methylation of cob(I)alamin with an −1 −1 apparent Km for GB of 1.96 ± 0.2 mM and a Vmax of 1.49 ± 0.04 µmol • min • mg (n = 3) (Fig. 17).

Stoichiometery of the Glycine Betaine:Cob(I)alamin Methyltransferase Reaction. Our experiments with growing cultures revealed that GB was consumed with the concurrent production of dimethylglycine as a cellular product. The methylation of cob(I)almin with GB suggested that this DMG might be produced directly by DSY3156. Assays were first conducted in which the molar ratio of GB consumed to methylCbl produced was determined as 0.98. In separate experiments, the ratio of produced DMG to methylCbl was measured as 1.04, providing

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Fig. 16. DSY3156 is a glycine betaine:cob(I)alamin methyltransferase. (A) UV-visible spectrum of 1.75 mM cobalamin reduced with Ti(III) citrate showing total change of absorbance vs. wavelength between 480 nm to 740 nm to demonstrate cob(I)alamin to methylcob(III)alamin (methylCbl) conversion in the presence of 5 mM GB and 50 μg of DSY3156. Spectra were gathered at 30-s intervals from 0 min to 9.5 min. Arrows indicate increasing or decreasing absorbance during course of reaction in different parts of spectrum. (B) Change in absorbance at 540 nm and 578 nm over time for an experiment in which the assay contained 50 mM GB, and at the arrow 50 μg of DSY3156 was added to the reaction mixture.

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Fig. 17. Michaelis–Menten kinetics of recombinant DSY3156. Enzyme rates are expressed as μmol • min−1 • mg−1 for the conversion of free cob(I)alamin to methylCbl against varying mM levels (0.250 mM, 0.5 mM, 1 mM, 5 mM, 10 mM, and 50 mM) of glycine betaine. The box Inset indicates Km and Vmax values with 1σ calculated using a nonlinear Michaelis–Menten algorithm in Prism 6 (GraphPad). In this experiment, n = 3 and error bars indicate 1σ from the mean.

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63 strong support for an overall reaction stoichiometry of 1 GB consumed to produce 1 DMG and 1 methylCbl. We confirmed this stoichiometry in an experiment in which GB and both products were measured simultaneously by removal of samples at specific time points for TLC from a spectrophotometrically monitored reaction (Fig. 18). Although GB was consumed, methylCbl and DMG were produced at approximately unit stoichiometry over the course of the reaction.

DSY3157 Is a Methylcorrinoid:Tetrahydrofolate Methyltransferase. The genomic context of DSY3156 suggested that it was the first methyltransferase in a pathway to generate methyl-THF from GB and that the DSY3157 gene may encode the predicted pterin methyltransferase needed to catalyze the second half reaction of this pathway. BLASTp alignment revealed that DSY3157 was homologous to the soluble MtrH subunit of the membrane-bound methyltetrahydromethanopterin:cobalamin methyltransferase complex of methanogens (Hippler and Thauer, 1999). We therefore cloned the DSY3157 gene using the polymerase incomplete primer extension (PIPE) cloning technique (Klock and Lesley, 2009) and heterologously expressed DSY3157 in Escherichia coli BL21 (DE3) to test this hypothesis. The enzyme had an N-terminal hexahistidine tag and was purified to apparent homogeneity by nickel-affinity chromatography (Fig. 11B). The DSY3157 enzyme catalyzed the transfer of methyl groups from methylCbl to THF at an initial rate of 0.64 ± 0.03 µmol • min−1 • mg−1 as measured by the change in UV-visible spectrum (Fig. 12A). The conversion of methylCbl to cob(II)alamin was followed by monitoring the decrease in absorbance at 525 nm (Fig. 12B). This reaction was dependent on the presence of both THF and recombinant DSY3157. Aliquots from spectrally monitored reactions run to apparent completion (45 min) were analyzed by reverse-phase HPLC. Reactions lacking enzyme had only THF eluting as the major peak (Fig. 12C); however, in complete reactions, an additional peak having the retention time noted for methyl-THF was observed (Fig. 12D). In three separate reactions, 0.172 ± 0.003 mM methylCbl (calculated by ΔA525) was consumed whereas 0.19 ± 0.01 mM methyl-THF was produced. No methyl-THF peak was detectable during HPLC analysis of aliquots from assays lacking enzyme, methylCbl, or THF.

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Fig. 18. Stoichiometric demethylation of GB to produce DMG and methylCbl. The methylation of cob(I)alamin was monitored spectrophotometrically, and the conversion of GB to DMG was measured by TLC.

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66

Discussion

The genes encoding members of the MttB protein superfamily are often annotated in genomes and metagenomes as probable TMA:corrinoid methyltransferases. However, only genes encoding one small clade within this large family of proteins (Fig. 19) have the pyrrolysine codon that is a hallmark of bona fide TMA methyltransferases (Krzycki, 2004; Soares et al., 2005). This ambiguity has left the function of the non-Pyl MttB proteins an open question. Here, we show that a non-Pyl MttB is indeed a corrinoid-dependent methyltransferase, but with specificity for GB. To our knowledge, DSY3156 is the first known glycine betaine:corrinoid methyltransferase, and we propose that its gene be designated mtgB. We have also shown that the DSY3157 enzyme encoded adjacently to MtgB functions as a methylCbl:THF methyltransferase, and we propose that its gene be designated mtgA. D. hafniense Y51 is able to grow anaerobically at the expense of GB. Neither TMA nor acetate is produced as a product, consistent with the absence of genes specific for known GB reductases in the genome. This finding suggests that, unlike for many Gram-positive bacteria, GB does not serve as an electron acceptor. Rather, the requirement for an external electron acceptor for growth, the demethylation of GB, and production of stoichiometric CO2 indicates that GB provides a source of electrons for anaerobic respiration. Thus, we propose that D. hafniense grows via anaerobic methylotrophy: i.e., the oxidation of the methyl group provides reducing power for anaerobic respiration. D. hafniense now joins a small group of species known to carry out denitrification at the expense of reduced C1 compounds (Auclair et al., 2010; Chistoserdova et al., 2009). The discoveries of MtgB-catalyzed corrinoid methylation with GB as well as the MtgA- catalyzed methylation of THF from methylCbl provide strong support for a hypothetical pathway of GB:THF methyl transfer, which can lead to oxidation of the methyl group to CO2 via THF intermediates (Fig. 15). We recently obtained preliminary proteomic data from cells grown on GB or pyruvate and found that the enzymes of the oxidative methyl-THF pathway, as well as MtgB, MtgA, and the accompanying corrinoid protein (DSY3155) (Fig. 15), are increased in abundance in GB-grown cells. These findings further support the qPCR data indicating that GB increases the transcript abundance of mtgB (DSY3156) and the putative GB transporter gene (DSY3154).

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Fig. 19. Phylogenetic tree of the COG5598 Superfamily. The evolutionary relations between the COG5598 members, or trimethylamine (TMA) methyltransferases (MttBs), are inferred with a maximum likelihood approach using the LG (Le and Gascuel, 2008) substitution model with a discrete gamma distribution. Green coloring (light green, bacterial; dark green, archaeal) is used to highlight those proteins that are predicted to be L-pyrrolysine containing TMA methyltransferases. DSY3156 (MtgB) is found in the part of the tree colored in red.

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The chemistry of methyl-group transfer between GB and cob(I)alamin is not surprising, given that the quaternary amine is essentially already an activated methyl donor. However, the discovery that a member of the MttB superfamily carries out a GB-dependent methylation of corrinoid reveals a functional rationale that might underlie the evolutionary relationship between Pyl-MttB and its non-Pyl MttB homologs. MtgB lacks pyrrolysine and yet uses GB, a quaternary amine, whereas the pyrrolysine containing MttB uses a tertiary amine, trimethylamine. However, if the TMA methyltransferase MttB functions as has been proposed for other Pyl methylamine methyltransferases (Hao et al., 2002, 2004; Krzycki, 2004), TMA would form an adduct with pyrrolysine before methyl transfer. In other words, pyrrolysine would serve to convert TMA into a quaternary amine, such as used by other MttB superfamily members such as MtgB (Fig. 20). This hypothesis mandates further functional analysis of the superfamily because it remains to be seen whether those MttB family members closest to Pyl-MttBs have substrate specificity for quaternary amines. It is possible that the family has diversified to include quaternary amines, such as choline or carnitine, or tertiary amines such as dimethylethanolamine. The function of these non-Pyl MttBs will provide interesting context for evolutionary biologists who seek to understand the driving forces behind entrance of Pyl into the MttB family of proteins. Homologs of MtgB are found in a large number of species, with concentration in the Firmicutes and the alpha proteobacteria and with fewer examples in the gamma and delta proteobacteria. Homologs of MtgB are particularly well-represented in alpha proteobacteria. Prominent are members of the Rhizobiales family, including well-known organisms such as Sinorhizoium meliloti and Mesorhizobium loti. Rhizobiales are known users of glycine betaine, and a number of species have been shown to acquire it for osmotic control or degrade it as sole carbon and energy source (Boncompagni et al., 1999; Fougère and Le Rudulier, 1990). Bacteroids are noted to uptake GB and then either maintain or consume it depending on external salt concentration (Fougère and Le Rudulier, 1990). Choline is at mM concentrations in legume-root nodules, and bacteroids induce genes to convert it to GB within legumes (Mandon et al., 2003). It has been shown that the pSym plasmids of S. meliloti carry the genes for degradation of a number of quaternary amines (Goldmann et al., 1991), and a homolog of mtgB is carried on one of the pSymA plasmids in this organism. One of these homologs is under quorum-sensing control within S. meliloti 1021 (Gao et al., 2005), supporting a potential role during some stage of plant symbiosis. The betaine:homocysteine methyltransferase was suggested as a first step in the

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Fig. 20. Proposed functional relationship between MtgB and MttB. Both proteins are members of the MttB family, of which MttB is the founding member, and until now has been the only member whose function has been known. In methanogenic archaea, MttB carries out the methylation of a corrinoid protein with trimethylamine (TMA) to supply methyl groups to coenzyme M (CoM) for ultimate conversion to methane and carbon dioxide. TMA has relatively few opportunities for strong H-bonding with the active site of MttB, but the pyrrolysine residue is hypothesized to compensate for this situation by binding TMA to form a quaternary amine that both orients the substrate and activates it for methyl group transfer. By comparison, glycine betaine is already a quaternary amine and, as a permanent cation with a carboxyl group, is able to bind the active site in an activated form for methyl transfer to a corrinoid cofactor.

71

72 demethylation of GB for Rhizobiales (Smith et al., 1988); however, it has now been shown by mutagenesis of S. meliloti that this methyltransferase is likely to be an anabolic enzyme (Barra et al., 2006), leaving the possibility that, at least under anaerobic conditions, MtgB homologs might serve this purpose in Rhizobiales. GB is used as a compatible solute by cyanobacteria and other phototrophic bacteria, as well as by eukaryotic uni- and multicellular algae (Hanson et al., 2014; Keller et al., 1999; Welsh, 2000). Thus, many microbes in marine environments acquire exogenous GB to serve as an osmolyte or degrade it during changing osmotic conditions for use as a source of nitrogen, carbon, and/or energy (Chen, 2012; King, 1984). In this regard, it is of interest that a large number of proteobacterial genera with genomes carrying homologs of mtgB inhabit marine environments. Of these homologs, the largest number of sequences can be attributed to those Rhodobacteriales within the Roseobacter clade. We identified homologs of mtgB that are found in isolated representatives of six subclades (Newton et al., 2010) of this ubiquitous major group of heterotrophs. Species have been isolated from both open-water columns or associated with eukaryotic unicellular or multicellular algae, where an enzyme having capability to use GB might be of advantage. Members of the Roseobacter clade, such as Ruggeria pomeroyi, are known to carry out one-carbon metabolism involving dimethylsulfoniopropionate (Reisch et al., 2011) or TMAO (Lidbury et al., 2014), and the Roseobacter–algal symbiosis is thought to involve C1 metabolism with such compounds (Geng and Belas, 2010). A number of Roseobacter genera carry up to five different GB transporters (Moran et al., 2007) and have been shown to degrade GB in studies with enriched bacterioplankton from coastal regions (Mou et al., 2007). In contrast, members of the widespread SAR11 group such as Pelagibacter ubique, known to metabolize GB (Sun et al., 2011), do not possess homologs of non-Pyl MttBs (MtgB), with the exception of an mtgB homolog identified in the hypervariable region of one SAR11 subclade of P. ubique (Gilbert et al., 2008). In addition, a number of marine gamma proteobacterial genomes have been sequenced that possess mtgB homologs, including many from deep-ocean environments (Swan et al., 2011). Finally, GB was recently found to be demethylated by some marine archaea in the methanogenic genus Methanococcoides (L’Haridon et al., 2014; Watkins et al., 2014). Only the genome of Methanococcoides burtonii has been sequenced and is publically available (Goodchild et al., 2004), but it is the only methanogen presently known to

73 carry a non-Pyl MttB, and it will be interesting to see whether this enzyme plays a role in quaternary amine demethylation. In the last few years, a surprising connection has been made between microbial metabolism of quaternary amines and human health. Gut microbiota produce trimethylamine when given precursors such as carnitine, choline, or GB commonly found in meats, fish, and other foodstuffs (Tang et al., 2013; Wang et al., 2011). The TMA then leaves the intestine and is converted to TMAO by liver monooxygenases. Circulating TMAO correlates strongly with incidence of cardiovascular disease, which has been suggested to act as a trigger for deposition of lipids by foam macrophage, leading to atherosclerosis and subsequent risk for stroke and heart attack. Gnotobiotic mice do not produce TMA from the precursors mentioned above except when given a complement of gut microbiota, implicating microbial metabolism as essential to TMA production (Tang et al., 2013; Wang et al., 2011). Presumably, the anoxic microbial route to TMA includes GB reductase (Andreesen, 2004) and choline-TMA lyase (Craciun and Balskus, 2012), as well as carnitine oxygenase (Lidbury et al., 2014) in the oxic portions of the digestive tract. However, competing microbial metabolism has the potential to mitigate TMA production from quaternary amines within the intestine. For example, depletion of the GB pool by demethylation could lessen the total amount of TMA produced in the intestine. Dimethylglycine, the direct product of glycine betaine, has been identified in serum and urine and has been sometimes attributed to liver activities (Heinzmann et al., 2012; Kirsch et al., 2010). However, we identified, at the National Center for Biotechnology Information and the Joint Genome Institute-Integrated Microbial Genomes, multiple intestinal or fecal inhabitants having MtgB homologs encoded in their sequenced genomes. One of these organisms is D. hafniense DP7 (Van de Pas et al., 2001), a fecal isolate that shares nearly identical mtgB-containing gene clusters with the D. hafniense Y51 used in this study. This finding suggests that microbially mediated demethylation of GB via MtgB is likely to occur in the gut. Recently a novel family of methanogens closely related to the Thermoplasmatales was described whose genomes encode pyl genes and pyrrolysine-dependent TMA, DMA, and MMA methyltransferase for the complete demethylation of TMA (Borrel et al., 2013). It has been suggested that these methanogens might serve to mediate levels of TMA in the intestine (Brugère et al., 2013). With the results we described here for MtgB, it seems possible that a non-Pyl

74 member of the TMA-dependent methyltransferase family found in intestinal bacteria might also affect TMA levels by removing a TMA precursor: that is, glycine betaine. If so, both Pyl and non-Pyl members of the MttB family could together be a means by which the deleterious effects of TMAO on human metabolism might be ameliorated.

75

Supporting Information

SI Materials and Methods. A 100-bp DNA Ladder (Promega) and qRT-PCR reactions for the methyltransferase and BCCT genes were loaded with Blue/Orange 1X Dye (Promega) into a 1.5% (wt/vol) agarose 1X TAE (40 mM Tris, 20 mM acetic acid, and 1 mM EDTA at pH 8.0) (LTE agarose) gel supplemented with 0.5 μg/mL ethidium bromide. The agarose gel was loaded in an Owl A1 Large Gel System (Thermo Scientific) filled with 1X TAE Buffer and run at 100 V using a FisherBiotech Electrophoresis Power Supply (Fisher Scientific) until the dye front was ∼75% through the gel. The gel was imaged using a UVP BioDoc-It Imaging System to visualize the predicted 200-bp amplicon size of each qRT-PCR.

76

Acknowledgements

We thank Dr. Andor Kiss and the Center for Bioinformatics and Functional Genomics at Miami University for the use of the ÄKTA chromatography system, Nanodrop spectrophotometers, Agilent BioAnalyzer, and the Bio-Rad CFX Connect thermocycler. We thank Dr. MeiChin Lai and her laboratory for very helpful discussions and protocols used for thin-layer chromatography of GB and its products. We thank Dr. Kari Green and Dr. Liwen Zhang in the Mass Spectrometry and Proteomics Facility at The Ohio State University for performing preliminary proteomic analysis of cell-free extracts. This work was supported by funding from the US Department of Energy, Office of Science, Office of Basic Energy Sciences through Grant DE-FG0202-91ER200042 (to J.A.K.) and Computational Bridges to Experiments (COMBREX) Subaward 4500000088 (to D.J.F.), as well as internal funding from Miami University.

77

CHAPTER 2

Isolation and characterization of a tetramethylammonium-degrading Methanococcoides strain and a novel glycine betaine-utilizing Methanolobus strain

Tomislav Ticak, Dinesh Hariraju, Margarette Bayron Arcelay, Brock A. Arivett, Steven E. Fiester, and Donald J. Ferguson Jr.

Archieves of Microbiology. 2014. DOI: 10.1007/s00203-014-1043-6

Category Contribution (%)

Design 90

Experimentation 50

Analysis 80

Writing 50

78

Summary

Two novel strains of methanogens were isolated from estuarine sediment with the capability to utilize quaternary amines. Based on the 16S rRNA analysis, strain B1d shared 99 % sequence identity with Methanolobus vulcani PL-12/MT and strain Q3c shared 99 % identity with Methanococcoides sp. PM1 and PM2, but our current isolates display clearly different capabilities of growth on quaternary amines and were isolated based on these capabilities. Strain Q3c was capable of growth on tetramethylammonium and choline, while strain B1d was capable of growth on glycine betaine. Ml. vulcani PL-12/MT was incapable of growth on glycine betaine, indicating an obvious distinction between strains B1d and PL-12/MT. Strain Q3c now represents the only known tetramethylammonium-utilizing methanogen in isolation. Strain B1d is the first quaternary amine-utilizing methanogen from the genus Methanolobus. This study suggests that quaternary amines may serve as ready precursors of biological methane production in marine environments.

79

Introduction

Biological methane production, or methanogenesis, is a topic of considerable global and environmental interest due to the potential of methane to serve as a renewable energy source and its potency as a greenhouse gas (Rothman et al., 2014). Canonical methanogenesis is specific to the domain Archaea, and by studying methanogenesis, we can better understand global carbon and nitrogen cycling. Furthermore, studies related to quaternary amine degradation will give insights into the remediation of toxic compounds, i.e., tetramethylammonium (Asakawa et al., 1998; Tanaka, 1994), or cardiovascular associated health factors such as choline and glycine betaine (Tang et al., 2013; Wang et al., 2011, 2014). Methanogenesis encompasses three defined pathways: hydrogenotrophic, aceticlastic, and methylotrophic (Thauer et al., 2008). Methylotrophic methanogenesis results in the mobilization and transfer of methyl groups from methylated compounds directly to the thiol group of coenzyme M (2-mercaptoethanesulfonic acid; CoM) (Asakawa et al., 1998; Bose et al., 2008; Burke and Krzycki, 1997; Ferguson et al., 2000; Ferguson and Krzycki, 1997; Harms et al., 1995; Sauer et al., 1997; Tallant and Krzycki, 1997), which is the penultimate step of methanogenesis. Species of the order Methanosarcinales are some of the most adaptable methanogens to date with the potential to utilize 14 known substrates: H2/CO2, H2/CO, acetate, methanol, methylamine (MMA), dimethylamine (DMA), trimethylamine (TMA), tetramethylammonium (QMA), dimethyl sulfide, methanethiol, methylmercaptopropionate, glycine betaine (GB), choline, and dimethylethanolamine (DMEA). Methylotrophic methanogenesis is a mechanism by which methanogens can compete with sulfate-reducing bacteria (SRB) in marine sediments (Purdy et al., 2003), as determined by the kinetics and thermodynamics in those environments (Banat et al., 1981; Kristjansson and Schönheit, 1983; Lovley et al., 1982; Winfrey et al., 1977). Additionally, sulfate ions are inhibitory of hydrogenotrophic and aceticlastic methanogenesis in the presence of SRB, while negligible inhibition of methylotrophic pathways from sulfate ions has been observed (García- Maldonado et al., 2012; King, 1984; Oremland et al., 1982). Thus, under sulfate-reducing conditions, only methylotrophic methanogenic pathways are likely to account for significant production of methane. This raises the question of whether there are substrates for methylotrophic methanogens that are yet undiscovered and whether lesser-known substrates may play a larger role in methane production in marine sediments. Prior research showed that co-

80 cultures of methanogens with SRB stimulated methane production with quaternary amines such as choline and glycine betaine; this activity was the direct result of trimethylamine production by fermenters in the co-culture (King, 1984). In this way, the presence of fermenters, methanogens, and SRB could lead to a system of quaternary amine breakdown, which could explain their co- localization in the environment. In 1994, Methanococcoides sp. NaT1, of the order Methanosarcinales, was isolated from Tokyo Bay in Japan (Tanaka, 1994). Strain NaT1 was the first methanogen shown to be capable of utilizing QMA, the simplest quaternary amine, as a direct substrate for methanogenesis. This was also the first demonstration of QMA degradation in an anaerobic system. Furthermore, QMA utilization followed the canonical methylotrophic route, which directly methylates CoM (Asakawa et al., 1998). Unfortunately, viable cultures of strain NaT1 no longer exist, so there are no publicly available QMA-utilizing methanogen strains at this time. Consequently, the elucidation of QMA driven methanogenesis remains incomplete, as no sequenced genomic data are publicly available. This may be primarily due to the fact that strain NaT1 was isolated and described prior to the current age of rapid and inexpensive genome sequencing. A related observation was that novel Methanococcoides sp. strains isolated from marine sediments by growth on methylamine are capable of methanogenesis directly from choline, DMEA, and GB (L’Haridon et al., 2014; Watkins et al., 2012, 2014); however, these strains are not known to degrade QMA. No quaternary amine-degrading methanogens have thus far been identified outside of the genus Methanococcoides. The loss of the QMA-degrading methanogen strain NaT1, combined with our interest in anaerobic quaternary amine catabolism, led us to test for the presence of these methanogens in the environment. As we began this project, we used our GB methyltransferase enzyme from Desulfitobacterium hafniense Y51 (Ticak et al., 2014b) coupled with prior studies of quaternary amine-dependent methanogenesis as a benchmark to formulate a hypothesis that GB and QMA methanogens are present environmentally and that we would be able to isolate different genera capable of this form of growth from brackish estuarine sediments. The significance of this work is threefold: (1) to show that quaternary amine-degrading methanogens inhabit estuarine and oceanic environments in numbers sufficient to allow facile isolation, (2) to obtain a QMA- degrading methanogen strain for further study of this pathway, and (3) to show that quaternary amine-degrading methanogens exist outside of the genus Methanococcoides. These substrates

81 may well contribute more to the global carbon and nitrogen cycles than was previously understood. Furthermore, with the expanding knowledge of quaternary amines and abundance of these compounds in the environment, ecological studies will need to be focused on the amounts of methane formed from these compounds and the abundance of the organisms responsible for these processes.

82

Materials and Methods

Growth and maintenance of enrichment and pure cultures. Anoxic sediment from the brackish Southwest Branch Back River in Hampton, Virginia, USA (GPS coordinates: 37.066444, −76.311639), was collected in May 2013, sealed in a polypropylene jar, and shipped overnight at ambient temperature to our laboratory. The jar was transferred to an anaerobic chamber (Coy Laboratory Products, Grass Lake, MI, USA), opened, and allowed to equilibrate overnight at 37 °C. Enrichment cultures were prepared in brackish medium (BM). This medium was identical to Methanosarcina acetivorans high-salt medium except that the concentration of added NaCl was lowered from 400 to 275 mM (Metcalf et al., 1997). BM is a chemically defined mineral salts medium that contains no carbon sources, such as yeast extract or acetate, other than the provided growth substrate. The cultures were grown in 27.2-mL sealed anaerobic Balch tubes with 10 mL of medium and 17.2 mL of headspace. The enrichment cultures were inoculated with sediment inside the anaerobic chamber, resealed with sterile butyl rubber stoppers and crimp seals, and the headspace of each tube was exchanged outside of the anaerobic chamber with filtered CO2/N2 (20/80 %) gas mix on a vacuum-gassing manifold. Each 10 mL of enrichment culture was supplemented with one gram of anoxic sediment, filter sterilized (0.22-μm cellulose acetate filter) quaternary amine substrate (10 mM), vitamins, and streptomycin (200 μg/mL). The cultures were incubated at 37 °C statically in the dark and monitored daily for methane production by gas chromatography (GC). Once the cultures had reached an apparent maximum methane production, 1 % (v/v) of culture was transferred to 10 mL of fresh medium, supplemented as before. Subsequent transfers of the enrichment cultures were done into liquid medium lacking antibiotic. After three transfers in liquid BM, enrichment cultures were diluted 10−4–10−5 and plated on BM containing 1.5 % (wt/v) agar and streptomycin (200 μg/mL) and incubated in anaerobic jars flushed with CO2/H2S/N2 (20/0.1/79.9 %) gas mix inside of the Coy anaerobic chamber at 37 °C. After 1–2 weeks of incubation, the jars were opened, and the colonies were picked to liquid BM. Once grown, methanogenic cultures were subcultured on BM agar containing tetracycline (50 μg/mL) and incubated for 2 weeks as described above, then transferred to liquid BM. These cultures were examined for purity by differential interference contrast (DIC) microscopy, thioglycolate broth, as well as 16S rRNA PCR amplification and sequencing. Each culture was

83 then transferred to BM containing 40 mM trimethylamine (TMA) or 62.5 mM methanol. Each strain was subcultured into BM with kanamycin (50 μg/mL) and ampicillin (100 μg/mL). Strain B1d was provided methanol (62.5 mM) and strain Q3c was provided TMA (40 mM) in the medium. Grown cultures were serially diluted in BM and plated onto BM supplemented with both kanamycin (50 μg/mL) and ampicillin (100 μg/mL) containing 1.5 % (wt/v) agar and incubated as described above. Isolated colonies of each culture were grown and maintained in liquid BM lacking antibiotics and used for identification and further characterization. Both strains were deposited in the Biological Resource Center (NBRC), National Institute of Technology and Evaluation, Chiba, Japan, with the accession numbers NRBC 110459 (strain B1d) and NRBC 110458 (strain Q3c). The strains were also deposited in the RIKEN BioResource Center, Japan Collection of Microorganisms (JCM), with the accession numbers JCM 30225 (strain B1d) and JCM 30226 (strain Q3c). Actively growing Methanolobus vulcani PL-12/MT (DSM 3029) was purchased from the Leibniz Institute DSMZ-German Collection of Microorganisms and Cell Cultures (DSMZ) and grown and maintained in BM supplemented with 62.5 mM methanol and culturing was also attempted in BM supplemented with 40 mM GB.

Microscopic examination. DIC microscopy was performed on wet-mounts of strains B1d and Q3c at stationary phase using an Olympus AX70 microscope at 600X total magnification. The mounts were also observed under fluorescence microscopy with the same microscope with excitation wavelengths of 340–380 nm. Specimens for scanning electron microscopy (SEM) were prepared by using a 1:1 ratio of stationary-phase cultures with fixative (2 % paraformaldehyde, 2.5 % glutaraldehyde in BM lacking PO43−) and allowing them to incubate anaerobically for 30 min at 25 °C. One hundred microliters of fixed cells was pipetted onto poly- L-lysine-coated coverslips and allowed to dry for 30 min. Excess medium was wicked away with a KimWipe™, and each sample was washed four times for 10 min with BM lacking PO43−.

Samples were then ethanol dehydrated, CO2 critical point dried in a Tousimis Samdri-780A critical point dryer (Tousimis, Rockville, MD, USA) and gold sputter coated for 90 s at a current of 15 mA with a Denton gold sputter coater (Denton Vacuum, LLC, Moorestown, NJ, USA) (Fiester et al., 2014). Samples were examined at 25,000X, 50,000X, and 75,000X and an accelerating voltage of five kiloelectronvolts with a Zeiss 35VP scanning electron microscope (Carl Zeiss AG, Oberkochen, Germany).

84

Physiological characterization of the isolates. Both of the isolates were tested for their ability to grow in varying NaCl concentrations by amending the BM with the addition of zero, 0.05, 0.10, 0.275, 0.40, 0.60, 0.90, 1.20, or 1.40 M NaCl. BM with no additional NaCl contains 46 mM Na+ from other sources. The temperature dependence of each strain was measured using BM at 4, 15, 18, 23, 30, 37, 40, and 45 °C. The cultures were grown twice using 1 % (v/v) inocula in triplicate at the respective NaCl and temperature conditions. These cultures contained 40 mM GB for strain B1d and 40 mM QMA for strain Q3c. To test for the ability to utilize different growth substrates, the cultures were transferred to BM containing the respective substrate; GB, dimethylglycine (DMG), choline, DMEA, carnitine, QMA, TMA, DMA, MMA, methanol, acetate, or H2/CO2. All substrates were supplemented at 40 mM except acetate (80 mM), methanol (62.5 mM), and H2/CO2 (80/20 %). Cultures with measureable growth were passaged three times in triplicate; with 1 % (v/v) inocula to ensure that growth was maintained and to limit substrate carryover. The cultures that did not grow were observed for 16 weeks.

Phylogenetic and DNA G+C (mol %) content analysis. Genomic DNA was isolated from strains B1d and Q3c as well as Ml. vulcani PL-12/MT by phenol–chloroform extraction as described previously (Sambrook and Russell, 2001). To obtain G+C (mol %) content for each strain, extracted DNA from B1d and Q3c was prepared by using a Nextera XT DNA Sample Preparation Kit and a MiSeq Reagent Kit version 3 (600 cycles), as per the manufacturer’s instructions (Illumina, San Diego, CA, USA). The data were analyzed using the CLC Genomics Workbench version 7.0.4 (CLC Inc, Aarhus, Denmark). PCR amplifications of archaeal and bacterial 16S rRNA were carried out using purified genomic DNA from each strain and three previously described sets of primers: 109f/ A1492r, 1Af/A1492r, and 27f/1492r in addition to a known methanogenic marker, mcrA, with ME1/ME2 (Webster et al., 2006). The 25 μL of PCR reactions contained 1X Phusion DNA polymerase buffer, 1 U Phusion DNA polymerase (New England Biolabs), 200 μM dNTPs, 0.5 μM of each primer pair, and 100 ng purified genomic DNA quantitated via NanoDrop 2000 UV–Vis spectrophotometer (Thermo Scientific, Wilmington, DE, USA). The PCR reactions were performed by the following protocol: 5 min at 98 °C, followed by 30 cycles of 95 °C for 30 s, 58 °C for 30 s, and 72 °C for 90 s, followed by a final extension step of 10 min at 72 °C. PCR

85 products were cleaned using a Wizard® SV Gel and PCR Clean-Up System (Promega, Madison, WI, USA) and sequenced with PCR amplification primers by using a BigDye Terminator (version 3.1) Cycle Sequencing Kit (Life Technology Corporation, Carlsbad, CA, USA) on an Applied Biosystems 3730xl DNA analyzer (Life Technology Corporation). Sequencing was performed twice after initial isolation from the plates and twice again after final isolation to ensure the sequence validity for the isolates, while Ml. vulcani PL-12/MT was sequenced as above three times using the above primers to replace N’s from the deposited NCBI sequence (GI:343206176). The 16S rRNA sequence data for Q3c (1,330 nt) and B1d (1,330 nt) were used as queries for obtaining closely related sequences from both the GreenGenes database (DeSantis et al., 2006) and NCBI nucleotide BLAST database (Altschul et al., 1990), excluding uncultured sequences. The percent identities to the query sequences were recorded and 28 sequences were obtained for phylogenetic analysis. The 28 sequences were aligned via MUSCLE (Edgar, 2004) and then analyzed for the highest log likelihood value (−4,735.6805) for maximum likelihood analysis in MEGA6 (Tamura et al., 2013). The phylogenetic tree was reiterated 1,000 times using the general time-reversible model (Nei and Kumar, 2000) with a discrete gamma distribution among sites (+G, parameter = 0.1593) with any position less than 95 % site coverage being eliminated leaving 1,309-nt positions in the final analysis. M. acetivorans C2AT (GI:470466243) and Methanimicrococcus blatticola PY-27 (GI:374719707) were used as outgroups that were truncated from the tree. Amplified partial mcrA gene products were sequenced as stated above, and data for B1d and Q3c were used as queries for tBLASTx (Gish and States, 1993) and used to search the non- redundant (nr) NCBI amino acid database. A total of 21 sequences were retrieved and used for the initial phylogenetic analysis. The sequences were aligned with MUSCLE and then truncated to 160 amino acids to eliminate bias of longer sequences. MEGA6 was used to calculate the highest log likelihood value (−1,124.629) for maximum likelihood analysis, which encompasses the Le–Gascuel model (LG model) (Le and Gascuel, 2008). The LG model was used with a discrete gamma distribution among the different sites [5 categories (+G, parameter = 0.3718)] in addition to calculating the potential invariability of individual sites [(+I), 16.7476 %] in MEGA6 (Tamura et al., 2013). All positions that were less than 95 % site coverage were removed from the final analysis prior to tree construction yielding sequences with a final dataset of 160 amino

86 acids. After tree construction, the outgroup sequences consisting of M. acetivorans C2AT (GI:499333927), Methanosarcina mazei (GI:145370893), Methanosarcina thermophila TM-1T (GI:154240554), and Methanosarcina barkeri (GI:499625182) were truncated from the final tree. Non-Pyl MttB sequences were obtained from draft genomes of the Ml. vulcani B1d and Mc. methylutens Q3c genomes described below in Genome annotation and analysis. These non- Pyl MttB sequences were amended to the tree represented in Fig. 19 using the same methods as provided in Chapter 1 – Materials and Methods: COG5598 Phylogenetic Tree and Sequence Acquisition.

Genome annotation and analysis. Ml vulcani B1d and Mc. methylutens Q3c were both submitted to the Rapid Annotation using Subsystem Technology (RAST) database for automatic annotation of each respective partial draft genome sequence (Brettin et al., 2015). The annotated genomes were then compared to their closest sequenced relative such as Ml. vulcani B1d versus Methanolobus psychrophilus R-15T (Chen et al., 2012) and Mc. methylutens Q3c versus Methanococcoides burtonii Ace-MT (Allen et al., 2009) by comparing them manually for specific methanogenic pathways using BLASTp and globally in Seed Viewer version 2.0 (Brettin et al., 2015). The Redhawk supercomputer (redhawk.hpc.miamioh.edu) was then used to analyze and correct any open reading frames (ORFs) with the European Molecular Biology Open Software (EMBOSS) (Rice et al., 2000).

Analytical methods. Methane measurements were performed using an Agilent Technologies model 6,890 N GC using a flame ionization detector (FID) with a ShinCarbon ST 80/100 column. Samples (200 μL) of headspace were aseptically removed from growing cultures using a one milliliter glass, gas-tight Hamilton syringe, fitted with a Mininert™ syringe valve and a sterile 25 gauge disposable needle, and injected into the GC. Methane was quantitated by comparison of the peak area values to standard curves of methane concentration generated at the time of each set of experimental measurements. Quaternary amine (GB and QMA) concentrations were measured from 10 μL of culture samples, diluted 1:10, using the periodide method with standard curves specific to each substrate (Wall et al., 1960). The tertiary amines DMG and TMA were tested to ensure that there was no

87 cross-reactivity in the periodide assay at the relevant concentrations. Methylamine and acetate concentrations in the media were measured by GC as described previously (Burke and Krzycki, 1997; Kremer et al., 1993). A colorimetric assay was used to determine ammonium concentrations by first diluting culture media 1:300 prior to being assayed as described previously (Kandeler and Gerber, 1988).

88

Results

Isolation of strains B1d and Q3c. Previous studies have shown the ability of methanogens to utilize quaternary amines such as QMA, choline, and GB as carbon and energy sources (Tanaka, 1994; Watkins et al., 2012, 2014). The lack of publicly available quaternary amine-degrading methanogenic strains along with our general interest in anaerobic quaternary amine metabolism led us to examine whether quaternary amine-degrading methanogens could be isolated from anaerobic estuarine sediment. We chose to examine sediment from a brackish tidal marsh on the coast of Virginia, USA, in the Southwest Branch Back River as a representation of the metabolic capabilities of organisms in such environments. In order to minimize the bacterial involvement in the degradation of the quaternary amines, we selected for methanogens using a modified medium (BM) optimized for marine methanogen growth and supplemented with antibiotics (200 μg/mL streptomycin). Methanogens have been shown to be resistant to many commonly used antibiotics and have been isolated in the presence of antibiotics in other studies (Kumar et al., 2012; Whitman et al., 2006). The QMA and GB enrichment cultures each showed rapid growth and methanogenesis after 2 days of incubation. Subsequent transfers were plated to obtain isolated colonies. We performed an additional selection step by plating dilutions of the cultures obtained from the initial platings onto BM agar containing tetracycline (50 μg/mL). Analysis of the 16S rRNA sequence of these isolates indicated that the sequence of strain B1d from the GB enrichments appeared to be pure, but strain Q3c from the QMA enrichments still contained bacterial contamination, as evidenced by the amplification of a bacterial 16S rRNA sequence. In order to ensure purity of the methanogen cultures, we grew each isolate in either methanol or TMA in medium containing a cocktail of kanamycin and ampicillin on plates. Pure colonies of both strains were obtained at this step that were round, smooth, light brown, and slightly raised. The B1d colonies were translucent, and the Q3c colonies were opaque. Some colonies of each strain also formed bubbles after extended growth and produced cavities within the agar.

Identification and phylogenetic analysis of the isolates. We discovered the identity of the strains by 16S rRNA and mcrA gene analysis. We amplified a fragment of the 16S rRNA gene with Archaea-specific 109f/A1492r and A1f/ A1492r primer pairs, described previously

89

(Webster et al., 2006), and obtained single products from each primer pair (Fig. 21). We amplified a fragment of the mcrA gene product using the ME1/ME2 primer pair and obtained single products from each isolate. No PCR products were observed using the Bacteria-specific 27f/1492r primer pair for strains B1d and Q3c under the conditions tested. Sequencing of the 16S rRNA and mcrA products resulted in single homogeneous sequences for each of the products. A multiple sequence alignment was performed with available sequences from described species in the literature to compare these isolates to known cultivable strains (Fig. 22). The alignment of the 16S rRNA gene from strain B1d clustered strongly with Ml. vulcani PL-12/MT, but a closer examination of the deposited sequence of Ml. vulcani PL-12/MT revealed many unknown bases in the sequence, causing other Methanolobus strains to appear more similar, such as Methanolobus profundi MobMT (Table 2). We therefore obtained Ml. vulcani PL-12/MT from DSMZ and re-sequenced a 16S rRNA gene fragment generated using the A1f/A1492r primer pair. The newly sequenced fragment from Ml. vulcani PL-12/ MT was 99.1 % identical to that of strain B1d suggesting that B1d is a strain of Ml. vulcani PL-12/MT. Strain Q3c showed approximately 99 % identity to Methanococcoides sp. PM1 and PM2 (Table 3), which belongs to an apparent cluster of Methanococcoides methylutens strains from several independent isolation studies. Within this cluster, Mc. methylutens strains PM1, PM2, AM1, NM1, DM1, and Methanococcoides vulcani SLH33 are capable of choline and DMEA degradation. Mc. vulcani SLH33 and Mc. methylutens PM2 and NM1 are capable of GB degradation. Only strain NaT1 was capable of QMA degradation (L’Haridon et al., 2014; Watkins et al., 2012, 2014). Therefore, strain Q3c was unique in its complement of substrates. Secondary phylogenetic analysis was done with the use of the conserved methanogen-specific marker methyl-coenzyme M reductase subunit A gene (mcrA) analyzed with tBLASTx (Gish and States, 1993). The amino acid sequence of McrA for B1d clustered well within the Methanolobus products (Fig. 22B) but was more divergent from Ml. vulcani PL-12/MT compared to the 16S rRNA tree (Fig. 22A). McrA data from Q3c (Fig. 22B) showed nearly identical sequence similarity to Mc. methylutens strains PM1 and PM2, much like the 16S rRNA analysis (Fig. 22A).

Characterization of the novel quaternary amine-utilizing strains. Microscopic analysis of strain B1d revealed small cells 0.6 ± 0.07 μm in diameter (n = 15) (Fig. 23A, 23C). The B1d cells were coccoid but showed some evidence of a lobed morphology as well. The B1d cells

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Fig. 21. Gel electrophoresis of 16S rRNA and mcrA products. The names above the numerals indicate what organism was used for the specific bands. Archaeal 16S rRNA products are represented by reactions 4, 7, 10, and 13. Bacterial 16S rRNA products are represented by reactions 5, 8, 11, and 14. Products for the mcrA gene are represented by reactions 6, 9, 12, and 15. Control were performed with reactions 1, 2, and 3 which represent Archaeal 16S rRNA, Bacterial 16S rRNA, and mcrA PCR reactions lacking M. acetivorans genomic DNA. The ladder is designated as L and base pair (bp) sizes are indicated to the left of the gel image.

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Fig. 22. Maximum likelihood trees showing the phylogenetic position of strains B1d and Q3c in relation to the most closely related organisms, based on the partial 16S rRNA gene sequence (A) or partial McrA amino acid sequence (B). These trees are based on 1,000 bootstrap repetitions with values equal to or greater than 50 % support being shown. Each tree is drawn to scale with distance being measured in substitutions per site. The numbers shown in parentheses are accession numbers for the indicated sequences.

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Table 2. Comparative analysis of several known Methanolobus species to B1d

Taxa: 1, B1d (this study); 2, Ml. profundi MobMT (Michimaru et al., 2009); 3, Ml. vulcani PL-12/MT (Kadam and Boone, 1995); 4, Ml. tindarius Tindari 3T (König and Stetter, 1982); 5, Ml. taylorii GS- 16T (Oremland and Boone, 1994); 6, Ml. oregonensis WAL1T (Liu et al., 1990); 7, Ml. bombayensis B-1T (Kadam et al., 1994); 8, Ml. psychrophilus R15T (Zhang et al., 2008); 9, Ml. zinderi SD1T (Doerfert et al., 2009)

Characteristic 1 2 3 4 5 6 7 8 9 Habitat Estuarine Saline, deep Sea Marine black Estuarine Saline, Sea Cold Saline, deep sediments subsurface sand sediments sediments sediments alkaline sediments Wetlands subsurface coal aquifer seam Cell Diameter (μm) 0.5 – 0.7 0.9 – 1.2 1.0 – 1.25 0.8 – 1.25 0.5 – 1.0 1.0 – 1.5 1.0 – 1.5 0.9 – 1.2 0.5 – 1.0 Cell Shape Cocci Irregular Cocci Irregular Cocci Irregular Irregular Diplococci Irregular coccoides coccides coccoides coccoides coccoides Flagella – + – + – – – + – Growth conditions Temp (˚C) 18 – 40 9 – 37 13 – 45 10 – 45 11 – 40 20 – 42 22 – 40 0 – 25 25 – 50 Na+ concentration (M) 0.05 – 0.94 0.1 – 1 0.1 – 1.2 0.06 – 1.27 0.2 – 1.20 0.1 – 1.6 0.3 – 2.0 0.05 – 0.8 0.05 – 1.8 DNA G+C content (mol%) 40.2 42.4 39 45.9 40.8 40.9 39.2 44.9 42 16S rRNA gene sequence – 97.9 97.3 96.4 96.5 95.5 97.7 96.7 96.1 identity (%) compared to B1d (99.1)

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Table 3. Comparative analysis of several known Methanococcoides species to Q3c

Taxa: 1, Q3c (this study); 2, Mc. burtonii Ace-MT (Franzmann et al., 1992); 3, Mc. alaskense AK-5T, 4, Mc. alaskense AK-9 (Singh et al., 2005); 5, Mc. methylutens TMA-10T (Sowers and Ferry, 1983); 6, Mc. sp. MM1 (Lyimo et al., 2009); 7, Mc. sp. NaT1 (Tanaka, 1994); 8, Mc. vulcani SLH33T (L’Haridon et al., 2014); 9, Mc. sp. AM1, 10, Mc. sp. NM1, 11, Mc. sp. PM1, 12, Mc. sp. PM2 (Watkins et al., 2012)

Charactistics 1 2 3 4 5 6 7 8 9 10 11 12 Habitat Estuarine Antarctic Marine Marine Marine Mangrove Bay Volcanic Bay Volcanic Bay Bay sediment lake sediment sediment sediment sediment sediment sediment sediment sediment sediment sediment Cell Diameter (μm) 0.9 – 1.3 0.8 – 1.8 1.5 – 2.0 1.0 – 1.5 1.0 0.8 – 2.0 0.5 – 1.2 0.6 – 1.7 N.R. N.R. N.R. N.R. Cell Shape Cocci Cocci Irregular Irregular Irregular Irregular Irregular Irregular N.R. N.R. N.R. N.R. coccoides coccoides coccoides coccoides coccoides coccoides Flagella – + – + – – – + N.R. N.R. N.R. N.R. Growth conditions Temp 15 – 40 1.7 – 24 5 – 25 5 – 25 15 – 35 23 – 35 25 – 40 4 – 35 N.R. N.R. N.R. N.R. Na+ concentration (M) 0.05 – 0.94 0.2 – 0.5 0.1 – 0.7 0.1 – 0.7 0.1 – 1.1 0.1 – 0.6 0.1-0.5 0.25 – 1.0 0.1 – 0.41 0.1 – 0.41 0.1 – 0.41 0.1 – 0.41 DNA G+C content (mol%) 43.3 39.6 41.9 39.5 42 N.R. 45 43.4 N.R N.R N.R N.R 16S rRNA gene sequence – 97.9 98.3 98.3 99.2 98.7 99.0 98.5 99.2 98.9 99.3 99.2 identity compared to Q3c

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Fig. 23. Microscopic examination of strains B1d and Q3c. (A) DIC micrograph of strain B1d at X600 magnification. Scale bar indicates 10 μm. (B) DIC micrograph of strain Q3c at X600 magnification. Scale bar indicates 10 μm. (C) Scanning electron micrograph of strain B1d at X75,000 magnification. Scale bar indicates 200 nm. (D) Scanning electron micrograph of strain Q3c at X75,000 magnification. Scale bar indicates 200 nm. On average, strain B1d measured 0.60 ± 0.07 μm (n = 15) in diameter and strain Q3c measured 1.10 ± 0.14 μm (n = 15) in diameter.

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98 appeared primarily as single cells with some in aggregates, a common characteristic of Methanolobus members (König and Stetter, 1982; Michimaru et al., 2009). Strain Q3c cells were 1.1 ± 0.14 μm in diameter (n = 15) and were also cocci arranged primarily as single cells, of which many had external protrusions resembling pili (Fig. 23B, 23D). All cells of each strain auto-fluoresced when viewed under a fluorescence microscope. No motility was observed in either strain when examined by DIC microscopy, and no flagella were observed in either strain using SEM. Both strains lysed easily with 0.01 % (wt/v) SDS or 0.01 % (v/v) Triton X-100 indicating proteinaceous cell walls. Both strains stained primarily gram-negative, although there was some variability in the reactions of each strain. Growth experiments were conducted with each strain to determine its temperature and salt dependence as well as the substrate range. The results of the temperature and salt experiments are reported in Tables 2 and 3 for strains B1d and Q3c, respectively. The temperature optimum for each strain was approximately 37 °C, and the salt optimum was approximately 300 mM Na+ for each strain. Strain B1d grew well in GB, TMA, and methanol, with slower growth in DMA and MMA. No growth of strain B1d was seen on any other substrate. Ml. vulcani PL-12/ MT does not grow in GB. Strain Q3c grew well in QMA and TMA with slower growth in DMA, MMA, and methanol. Interestingly, choline was found to support growth of strain Q3c as well, although choline-dependent growth was considerably slower than growth on other substrates, as seen with close relatives: strains PM1 and PM2 (Watkins et al., 2012). Subsequent transfers still showed a considerable lag time of 10–15 days prior to measurable growth. Neither strain PM1 nor PM2 is reported to use QMA as a methanogenic substrate. Neither B1d nor Q3c grew on acetate, H2/CO2, formate, carnitine, or DMEA. Strain Q3c did not grow on GB, unlike strain PM2. GC measurement of methylamines in the substrates and media confirmed that growth on QMA, GB, or choline was not due to the presence of contaminating TMA, DMA, or MMA. The lack of growth of strain Q3c on GB and the lack of growth of strain B1d on QMA or choline also supported the finding that no contaminating substrates were found in the substrates confounding the growth results. We tested the dependence of GB on growth of strain B1d and the dependence of QMA on the growth of strain Q3c and showed that no growth was observed in the absence of substrate and increased substrate concentration resulted in increased final OD600 up to the highest examined substrate concentration (80 mM) (Fig. 24). The rate of increase in OD600 did not appear to change

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Fig. 24. Effect of increasing GB or QMA concentrations on the growth of Methanolobus vulcani B1d and Methanococcoides methylutens Q3c. (A) Strain B1d was tested at the indicated concentrations of GB (mM), with or without BES. (B) Strain Q3c was tested at the indicated concentrations of QMA (mM), with or without BES. These graphs represent the average values of triplicate cultures.

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101 considerably with increased substrate concentration for either strain. The methanogenic inhibitor bromoethanesulfonic acid (BES) completely inhibited the growth of both strains. Strain Q3c cells entered death phase shortly after reaching stationary phase when grown in 80 mM QMA. This could be due to increased ammonia in the medium or change in pH, as ammonium did accumulate during growth and the pH of the culture dropped from an initial pH of 6.8 to approximately 6. Strain Q3c utilized all four methyl groups of QMA without excretion of simpler methylamines during the growth cycle, as evidenced by a complete lack of TMA, DMA, and MMA in the medium during growth (data not shown) and the stoichiometry of mol methane produced per mol of QMA (Fig. 25). We also measured ammonium concentration in the medium at points during the growth of the cultures and found that Q3c cultures showed a net increase of approximately 5 mM ammonium when comparing T0- and stationary-phase cultures. Strain B1d showed a net decrease in ammonium concentration from T0- to stationary-phase of approximately 5 mM. Given that all of the QMA (10 mM) in the strain Q3c culture had been mineralized by the time it reached stationary phase, it would appear that each strain consumed 5 mM ammonium for biosynthesis and growth. Consistent with previously described methylotrophic methanogenic pathways, each strain disproportionated its quaternary amine growth substrate. Strain B1d showed a ratio of 0.71:1 (CH4:GB) (Fig. 25A) and strain Q3c showed a ratio of 2.8:1 (CH4:QMA) which indicates a ratio 0.70:1 (CH4:methyl group), since there are four available methyl groups on QMA (Fig. 25B). These ratios are close to the theoretical 0.75:1 (CH4:methyl group) ratio for methylotrophic methanogenesis.

Analysis of draft B1d and Q3c genomes. The next generation sequencing platform, MiSeq, was used to obtain partially complete genomic information from both strains B1d and Q3c. B1d has a genome size of ~2,920,703 bp, which was contained within 20 contigs with an average size of 100,714 bps with an N50 value of 213,164. Q3c housed a ~2,560,744 bp-chromosome yielding 21 contigs with an average size of 98,490 bps with an N50 value of 307,883. The genomic data analyzed on the CLC genomics workbench was then used for submission to RAST for automated genomic analysis. B1d encoded 2815 sequences that comprised a total of 255 subsystems (Fig. 26A), while Q3c encoded 2496 sequences covering 249 subsystems of which B1d encoded 51

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Fig. 25. Growth curves are presented showing changes in OD600 as well as quaternary amine and methane concentrations over time for strains B1d (A) and Q3c (B). Strains B1d and Q3c disproportionate their quaternary amine substrates to produce approximately 0.75 mol methane per mol methyl group utilized. The error bars indicate one standard deviation from the mean.

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Fig. 26. Subsystem profile of methanogen isolates generated with RAST. (A) B1d systems visualized as a circular graph and (B) Q3c systems visualized as a circular graph with respective numbers of each system in parentheses.

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RNAs and Q3c encoded 59 RNAs which were compared with both RAST and EMBOSS (Fig. 26B). Primary focus was given to cofactor biosynthesis, methylotrophic pathways, and amine/quaternary amine transport for both B1d and Q3c. Corrinoid synthesis was completely accounted for in both strains by the appearance of the cob operon and major pathways of CH3 reduction and oxidation through modified pterin derivatives was also present in both genomes (Fig. 27). Some members of the Methanococcoides and Methanolobus clades encode the ability to utilize dimethylsulfide (Dworkin et al., 2006) and unsurprisingly both genomes encoded this pathway (Fig. 28). The breakdown pathway involves a fused methyltransferase/corrinoid protein which cycles the transfer of methyl groups from dimethylsulfide to CoM (Tallant and Krzycki, 1997). In strain B1d this putative operon also encodes a RamA-like protein previously shown to activate methylamine specific corrinoid-binding proteins (Ferguson et al., 2012), though Q3c did not share the same appearance of a ramA gene in its putative dimethylsulfide operon, further analysis by blasting the B1d RamA amino acid sequence against Q3c suggested that it too encodes a like enzyme which shared approximately 61 % identity with an E-value of zero. Additionally, there was the appearance of an N-methylhydantoinase in both operons of B1d and Q3c, which is also encoded by the M. acetivorans C2AT dimethylsulfide operon (Galagan et al., 2002). Though B1d and Q3c encode these predicted pathways they were never tested for growth on dimethylsulfide which merely suggests they have the potential to catabolize dimethylsulfide. B1d and Q3c both catabolize methylamines and each of their genomes’ encoded TMA, DMA, and MMA enzymes with a predicted Pyl residue in the proposed catalytic site (Hao et al., 2002, 2004; Krzycki, 2004) which has consistently been seen throughout the order Methanosarcinales. Only Mc. burtonii Ace-MT, a DMEA demethylating strain, was reported to have a non-Pyl mttB gene (Allen et al., 2009; Watkins et al., 2012). However, through genomic analysis of our isolates we can expand upon this further as B1d and Q3c join the fold of non-Pyl mttB encoding methanogens. Using MtgB (Ticak et al., 2014b) as a query we discovered that B1d possesses one non-Pyl mttB gene and Q3c encodes three non-Pyl mttB genes. These four non-Pyl MttB sequences were analyzed phylogenetically in the COG5598 superfamily (Fig. 29). The B1d non-Pyl MttB sequence clades strongly with our previously reported GB methyltransferase from D. hafniense strain Y51 (Fig. 29B), which presented the first archaeal

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Fig. 27. Proposed quaternary amine metabolic schema for methanogens. (I) In this route, quaternary amines are demethylated by a three-component enzyme system including a substrate- specific methyltransferase, a corrinoid-binding protein, and a 2-mercaptoethanesulfonate (CoM) methyltransferase, which is supported by Asakawa et al. (1998). (II) Quaternary amines could potentially serve as substrates for a route of direct or indirect methylation of tetrahydrosarcinapterin (H4SPT). (III) The proposed Mtr/Mer bypass proposed by Welander and Metcalf (2008) may also present a route of quaternary amine degradation through the formation of formaldehyde from a quaternary amine dehydrogenase followed subsequently by an enzymatic mediated condensation with H4SPT to yield methylene tetrahydrosarcinapterin

(CH2=H4SPT). Arrows indicate experimentally verified pathways of quaternary amine- dependent methanogenesis. Dashed lines indicate hypothetical pathways of quaternary amine demethylation and electron flow through oxidation and reduction of one-carbon groups are also indicated. Transport of quaternary amines is also proposed via betaine/choline/carnitine transporter (BCCT) or a homologue of the predicted trimethylamine permease (MttP) seen in other sequenced methylamine-utilizing methanogens. CHO–MF, formyl–methanofuran; CHO–

H4SPT, formyl–H4SPT; CH≡H4SPT, methenyl–H4SPT; CH3–H4SPT, methyl–H4SPT; CoB, 7- mercaptoheptanoylthreoninephosphate; CH3–CoM, 2-methylthioethanesulfonate; CoB–CoM, mixed disulfide of CoB and CoM; CO2, carbon dioxide; CH4, methane. Arabic numerals 1, formaldehyde; 2, glycine betaine; 3, choline; 4, tetramethylammonium.

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Fig. 28. Genomic context of putative dimethylsulfide operons for B1d (A) and Q3c (B). Numbers above the vertical lines indicate the contig bps of each locus in both B1d and Q3c. Names under each gene arrow indicate their associated locus name and names inside the gene arrows indicate their respective gene homolog: mtsAB, fused corrinoid-methyltransferase; hyuA, N-methylhydanotinase; groEL, heat-shock 60 kDa chaperonin protein; orf1/orf2; open reading frames with no known functions; ramA, corrinoid-activation protein; mtaA; putative corrinoid- thiol methyltransferase belonging to the uroporphyrinogen III decarboxylase superfamily.

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Fig. 29. Maximum-likelihood tree of the COG5589 superfamily amended with non-Pyl and Pyl MttBs from strains B1d and Q3c. (A) The LG method (Le and Gascuel, 2008) with gamma-distribution with 1000 bootstrap repetitions was used to construct the main phylogenetic tree with green indicating the Pyl clade (dark green = archaeal and light green = bacterial) and red indicating the GB methyltransferase from D. hafniense Y51. (B) B1d’s non-Pyl MttB in relationship to MtgB (DSY3156). (C) The location of Q3c’s non-Pyl MttBs and Pyl MttBs within the Pyl clade. (D) The cluster containing both the non-Pyl MttBs from M. burtonii Ace- MT (Dscole4) and M. methulutens Q3c. The scale bar indicates the numbers of substitutions per amino acid site and GI accession numbers are available for shown sequences with bootstraps not being shown for clarity.

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113 sequence in this cluster of Clostridial-dominated sequences. Q3c provides a more complex story, as two of the three non-Pyl MttBs clade within the Pyl cluster (Fig. 29C), and the remaining one clades with the Mc. burtonii Ace-MT non-Pyl (Fig. 29D). Two of the non-Pyl MttBs represent a stark deviation from what we have seen previously, typically, only the Pyl-encoding MttB enzymes, whether archaeal or bacterial, grouped in an isolated clade. Further findings revealed these non-Pyl MttBs genes being adjacently encoded near predicted TMA corrinoid-binding proteins (Fig. 30), suggesting a function for direct methylation of corrinoid cofactor. It is important to note, however, the majority of these putative operons lacked the appearance of a secondary methyltransferase, which we predict would be essential to the methylation of CoM or a pterin derivative (Fig. 27), so the particular fate of these methylated corrinoids remains an open question. Through analyzing these new non-Pyl mttB genes we discovered adjacent transporters in close genomic proximity to belonging to either the amino acid/polyamine/organocation (APC) transporter (APC) family, predicted choline transporters (licB), or BCCT family of transporters (Fig. 30). The non-Pyl mttB gene from strain B1d was located near several GB transporters (opuDs) with one adjacent to a CBP gene (Fig. 30A). This OpuD transporter possessed the highest amino acid identity (51 %) to predicted GB transporters of facultative anaerobes, Paenibacillus spp., that belong to the Bacillales order. The remaining Pyl mttB genes encoded by B1d and the mttB genes of Q3c had several different transporters located to them ranging from either APC transporter genes or licB genes (Fig. 30B). This is similar to what is seen in M. acetivorans C2AT, as the bona-fide TMA methyltransferases genes are near a licB gene.

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Fig. 30. Gene neighborhoods of mttiB genes in strains B1d (A) and Q3c (B). (A) B1d contains one non-Pyl mttB gene that encodes for a protein with the most similarity to MtgB (DSY3156) from D. hafniense Y51 which is located near a predicted GB transporter, opuD. The non-Pyl mttB is flanked by two different genes encoding corrinoid proteins; mtaC, methanol-like corrinoid binding protein and mttC, TMA-like corrinoid binding protein. (B) Q3c has three invidual operons encoding mttB genes which are colored in light blue with respective corrinoid- binding proteins colored in pink similar to TMA-like corrinoid proteins or mttC. In brown are predicted transporters for either I) choline (licB) or II) general amino acids cations. An activation factor for corrinoids (ramA) is found in one gene neighborhood which was not found in any of the other putative operons. The last Q3c gene block contains a putative thioredoxin reductase (trxB), serine/threonine protein kinase (rioN), RubisCO-like activase (ccbQ), and nitric oxide reductase (norD). Genes of unknown functions are labled as orf in their respective operons. Pyl, indicates the presence of a predicted TAG sequence for the encoding of L-pyrrolysine in MM0055 and MM1229.

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Discussion

The isolation of Ml. vulcani B1d by direct enrichment on GB has expanded the number of methanogenic genera capable of quaternary amine utilization, and the isolation of strain Mc. methylutens Q3c from estuarine sediment implies a more widespread ecology of QMA degradation. Given that methylotrophic substrates, such as methylamines, are commonly referred to as non-competitive substrates in these anaerobic communities, it is interesting to see how quaternary amines may fit into this model. Our laboratory has demonstrated that D. hafniense, an organism known to occupy similar environments to methylotrophic methanogens, is also capable of methylotrophic growth when coupled with fumarate, nitrate, or thiosulfate reduction (Ticak et al., 2014b). This suggests that methylotrophic substrates are not always non-competitive. Furthermore, in a recent study involving microcosm experiments, investigation of non- competitive substrate usage was performed for methanogens and SRB from microbial mats in hypersaline environments. Interestingly, members of the genus Methanococcoides, Methanohalophilus, and Methanolobus were observed in these communities (García-Maldonado et al., 2012). It is plausible that quaternary amines may serve as either competitive or non- competitive substrates in soil sediments; hence, further studies on the ecology of quaternary amine distribution and their anaerobic degradation are warranted. There are relatively few known natural sources of QMA in the environment, with some sources known in marine environments being from the phyla Mollusca, Cnidaria, and Bryozoa (Barceloux, 2008). It is unknown whether any other sources of QMA exist in marine and estuarine environments, but it is a possibility worth examining. It is also interesting to note that the ability of strain Q3c to grow in media lacking added NaCl suggests that this organism may be able to grow in freshwater sediments as well. All previous species of quaternary amine-utilizing methanogens have been isolated from marine or brackish environments, but quaternary amines, especially choline, would exist in freshwater systems as well. Our isolation of strain Q3c allows for a more complete physiological and genetic characterization of QMA-dependent methanogenesis to build upon the work done by the laboratories of Tanaka, Thauer and their coworkers (Asakawa et al., 1998; Tanaka, 1994). GB is an abundant quaternary amine in marine and estuarine environments that often serves as a compatible solute in a variety of organisms to combat the osmotic stress of saline environments (Heijthuijsen and Hansen, 1989; Oren, 1990; Ziegler et al., 2010). GB is known to

117 contribute to methanogenesis by serving as a substrate for fermentative bacteria that cleave it via the Stickland reaction to produce the methanogenic precursors TMA and acetate (Naumann et al., 1983). In this study, we isolated strain B1d based on its ability to grow on GB directly, and the speed at which the enrichment cultures grew suggests that GB may readily serve as a substrate for strain B1d, and possibly other methanogens, in situ. We can infer from previous work by Thauer and coworkers (Asakawa et al., 1998) that demethylation of QMA in strain Q3c proceeds via the methylotrophic pathway analogous to other methylamine breakdown pathways, as shown in Fig. 27. MtqA, MtqB, and MtqC, the enzymes for QMA demethylation were isolated from strain NaT1 and characterized previously and the N-termini were reported (Asakawa et al., 1998). However, we have been unable to identify genes in the NCBI database that could encode MtqABC. Further research in our laboratory currently focusing on the genome sequence of strain Q3c may pinpoint the genes responsible for this pathway. Alternative pathways for QMA degradation in Q3c still remain a possibility since the genes responsible for QMA degradation in any methanogen have yet to be discovered. One could speculate that GB and choline are catabolized through methylotrophic pathways as well. This hypothesis is supported by collaborative work in our laboratory, in which we have shown that a pyrrolysine-lacking homologue of MttB, the TMA methyltransferase from methylotrophic methanogens, functions as a GB methyltransferase in the bacterium D. hafniense Y51 (Ticak, et al., 2014b). This enzyme, MtgB, is a non-pyrrolysine member of a superfamily of enzymes (COG5598) that also includes the pyrrolysine-containing enzymes from both the Bacteria and Archaea. We propose that constituents of this family catalyze a variety of corrinoid-dependent methyl transfer reactions with quaternary and tertiary N-methylated amines as substrates. Another possible route is that demethylation of GB and choline could occur through the action of substrate-specific dehydrogenases that demethylate the substrate and generate formaldehyde and reducing power; the formaldehyde could enzymatically react with

H4SPT to generate methylene–H4SPT (Fig. 27) (Welander and Metcalf, 2005, 2008). However, the methanol bypass pathway was unable to support growth on methanol alone in the absence of the mtr/mer operon in M. barkeri, so this possibility is not likely to account for the growth on quaternary amines. It is possible, however, that an analogous quaternary amine bypass pathway exists in these organisms. A third route is also envisioned in which the direct methylation of

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H4SPT from the substrate is carried out by an unknown enzyme or pathway. Therefore, future work in our laboratory, guided by the genome data from strains B1d and Q3c that we have recently obtained, will be critical in gaining insight into the genes and enzymes responsible for quaternary amine dependent methanogenesis. Through targeted analysis of the B1d and Q3c genomes we have uncovered several sequences that encode non-Pyl MttBs that may act as corrinoid-dependent quaternary amine methytransferase. The lack of archaeal MttB sequences resting near MtgB suggests that this homolog may have been HGT into B1d as this would be likely to increase its carbon profile and provide a net gain evolutionarly (Nelson-Sathi et al., 2014). One can speculate as to how closely clustered MtgB enzymes form Mc. methylutens and Mc. vulcani may be in comparison to B1d. Other evidence for HGT may exist as several GB transporters, OpuDs, also resides around this non-Pyl MttB. The Opu-like transporters are the closest in amino acid identity (~50 %) to transporters from Paenibacillus spp. which have been shown to act positively in the rhizosphere by promoting plant growth (Emmert and Handelsman, 1999; Kokalis-Burelle et al., 2006). Given that the many non-Pyl MttBs are encoded by plant growth promoting rhizobacteria (PGPR), it will be interesting to see if Paenibacillus spp. also encode non-Pyl mttB genes, which might suggest a role for carbon and nitrogen cycling via quaternary amine breakdown in the rhizosphere (Boncompagni et al., 1999; Fougère and Le Rudulier, 1990; Goldmann et al., 1991; Prell and Poole, 2006; Smith et al., 1988). Now that a route to initiating methylated corrinoid from GB is present, the fate of methyl transfer is unknown, as the operon does not seem to encode a secondary methyltransferase. Future studies could be aimed at analyzing the function of the non-Pyl MttB enzyme to determine if it is, in fact, a GB methyltransferase and demonstrating the fate of methylated corrinoid in this strain through transcript analysis compounded with isolation of either native or recombinant pterin/thiol methyltransferases targets. The non-Pyl MttBs of Q3c represent some of the most interesting members of the COG5598 superfamily and may support our hypothesis that non-Pyl are responsible for QMA and choline metabolism; additionally, two non-Pyl MttBs clade closely with bona fide TMA methyltransferases of archaea (Fig. 29C), which has not been previously observed. This mandates further experimentation of enzymes in the Pyl cluster, both archaeal and bacterial, as the only biochemical evidence resides from Pyl MttBs from methanogenic archaea (Gaston et al., 2011). No bacterial Pyl MttB has been shown to initiate corrinoid methylation from TMA but

119 there are reports of bacteria growing on TMA anaerobically via unknown mechanisms (Prat et al., 2012). This opens the question of essentiality of Pyl within the Pyl clade of COG5598, as the appearance of two non-Pyl members in the Pyl clade suggests either Pyl could potentially be replaced by another amino acid or close chemistry to the TMA-Pyl adduct, which we previously proposed (Gaston et al., 2011; Hao et al., 2002, 2004; Krzycki, 2004; Ticak et al., 2014b). If it is not a replacement, it is possible that another small amine, such as QMA, may very well behave as the ligand for these enzymes. Furthermore, one finding that continues to appear in genomic analyses of MttBs is the appearance of a gene neighborhood which consists of a Pyl and non-Pyl MttB encoding genes in tandem as seen with D. hafniense (Fig. 10), which suggests a pathway catabolizing related compounds. Given that QMA is the simplest quaternary amine and most structurally similar to TMA it would not be surprising if this joint system of non-Pyl and Pyl MttBs was responsible for QMA and TMA tandem demethylation for both archaea and bacteria via corrinoid and a C1 carrier (Fig. 31). This pathway may very well explain why there was a persistent association of one single bacterial contaminant within the Q3c enrichment study, which was removed through subcultivating the Q3a enrichment on TMA and different antibiotic selection. However, no reports have ever been documented for anaerobic QMA breakdown by a bacterium but aerobic pathways have been documented (Hampton and Zatman, 1973; Ohara et al., 1990; Urakami et al., 1990).

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Fig. 31. Proposed pathway of QMA breakdown in archaea and bacteria. This model predicts the combined utilization of non-Pyl and Pyl MttB in demethylation QMA, where the non-Pyl methyltransferase would act upon QMA to generate TMA, which in turn, would be demethylated by the Pyl-containing methyltransferase. This would yield a 1:1 QMA:DMA stoichiometry and lead to the formation of methylated corrinoid protein from a Co(I) state. The fate of this methylated corrinoid is proposed to either a) lead to CH3-CoM formation as seen previously in

(Asakawa et al., 1998) or b) lead to the formation of methyl-tetrahydrosarcinapterin (CH3-

H4SPT) / CH3-THF. The parentheses indicate supported work in GB:CH3-THF pathways previously proposed with D. hafniense Y51 (Ticak et al., 2014b).

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Supporting Information

Gel Electrophoresis of amplified 16S rRNA and mcrA products. Ten microliters of amplified PCR product (described in Materials and Methods) with 1X Blue/Orange Loading Dye (Promega, Madison, WI) was directly loaded into a 1.5 % (wt/v) agarose 1X TAE gel (40 mM Tris, 20 mM acetic acid, and 1 mM EDTA at pH 8.0). The TAE agarose gel was placed in an OwlTM A1 Large Gel System (Thermo Scientific, Wilmington, DE) and filled with 1X TAE buffer. A 100-V current was used until the loading dye reached approximately 80 % of the way through the gel and was then imaged as seen in Fig. 21. Band sizes were compared with a 1 kb DNA ladder (New England Biolabs).

123

Acknowledgements

We would like to thank Dr. Richard Edelman and Mr. Matt Duley at the Center for Advanced Microscopy and Imaging at Miami University for helpful assistance with microscopy. We would like to thank Dr. Andor Kiss and Ms. Xiaoyun Deng in the Center for Bioinformatics and Functional Genomics for assistance with instrumentation and DNA sequencing. We thank Dr. Catherine Almquist at Miami University for the use of her GC. We thank Dr. Joe Krzycki for many helpful discussions and for critical review of the manuscript. We would like to thank Mr. Daniel Fleming for gathering the sediment at the Southwest Branch Back River. This project was supported by a National Science Foundation Research Experiences for Undergraduates site grant to Miami University (project DBI-1156703). An award given to Dr. Luis Actis at Miami University from Illumina supported sequencing of the genomes. Miami University, Hamilton, and the Department of Microbiology, Miami University, also provided funding.

124

CHAPTER 3

Analysis of the function of L-Pyrrolysine within the trimethylamine methyltransferase superfamily (COG5598) by comparison to the glycine betaine methyltransferase

Tomislav Ticak, Donald J. Ferguson Jr., and Joseph A. Krzycki

Category Contribution (%)

Design 30

Experimentation 30

Analysis 40

Writing 85

125

Summary

Direct binding studies of mono-, di-, and tri- methylamine (MMA, DMA, and TMA) have not yet been successful for the methyltransferases containing the L-pyrrolysine (Pyl) residue. The location of Pyl is found directly within the cleft of the αβ-TIM barrel of each of the three methylamine enzymes, albeit in different locations within the predicted active sites, thus suggesting a common function of Pyl between these structural homologs. To demonstrate the function of Pyl in these enzymes, more specifically the MttB enzyme, we chose to use the already deposited crystal structure of the glycine betaine (GB) specific MttB homolog lacking Pyl, DSY3156 (PDB:2QNE). The likelihood of MtgB binding GB would support the active site of an MttB enzyme and also act as a model comparing the proposed binding of a Pyl-TMA adduct proposed for catalysis of TMA. Superpositioning models of the proposed Pyl-TMA adduct to the predicted GB binding-pocket via Q-Site Finder in the MtgB structures showed near identical overlap of methyl group positioning for attack by a corrinoid cofactor. This structural and biochemical data may potentially highlight an evolutionary relationship not seen previously for Pyl and non-Pyl enzymes and will be used to model the COG5598 superfamily for future studies.

126

Introduction

Pyl exists in only a few proteins to date; the most notable members belong to the methylamine methyltransferases (MTs) (MtmB, MtbB, and MttB) where Pyl was first discovered (Hao et al., 2004). Studies involving the methylamine methyltransferases suggest that Pyl is required for methylamine metabolism (Mahapatra et al., 2006), however, the use of Pyl in other proteins has rarely been addressed except for Thg1 (Heinemann et al., 2009) where Pyl was considered non-essential. Thus, the function of Pyl-proteins can be interrupted in one of three ways; I) there are different forms of catalysis between non-Pyl and Pyl members, II) members lacking Pyl are dysfunctional or III) Pyl is replaceable in enzymes and thus nonessential. Previous data from our lab demonstrates that a non-Pyl member of the TMA MT superfamily, COG5598, a functional enzyme responsible for GB:cobalamin methylation (Ticak et al., 2014b). GB is well known as a methyl donor (Barra et al., 2006; Chen et al., 2010; Kirsch et al., 2010), but the relationship of a non-Pyl MttB catalyzing this reaction helps support the proposed role of Pyl for the formation of a chemical adduct with methylamines (Gaston et al., 2011; Hao et al., 2002, 2004; Krzycki, 2004; Ticak et al., 2014b). Crystallization of MtmB (Hao et al., 2002, 2004) highlighted a catalytic role for Pyl initially through alternate configurations of the residue under different crystallization conditions (Fig. 32). The role of this enzyme as a cobalamin- dependent methyltransferase was confirmed biochemically (Burke and Krzycki, 1997) and the importance of Pyl was later inferred by structural comparison to another known cobalamin- dependent MT belonging to the domain of methionine synthase (MetH) (Krzycki, 2004). MetH catalyzes the methyl transfer from methyl-THF to homocysteine to generate THF and methionine through a cobalamin cofactor, thus the positioning of residues for cobalamin interaction may appear similar to MtmB (Krzycki, 2004). The role of Pyl in MMA catalysis was examined by comparing the positioning of methyl-THF (MeTHF) within the MetH enzyme (Fig. 33), which revealed a close association of residues responsible for methyl transfer to or from a methylated corrinoid cofactor. If the Pyl residue is also found in the αβ-TIM barrel cleft for the DMA and TMA structures this might also suggest a requirement for Pyl during DMA and TMA catalysis much like MMA. To determine the essentiality of Pyl during methylamine growth, genetic approaches targeting deletion of PylT were performed (Mahapatra et al., 2006) which demonstrated null growth phenotypes when strains were supplied with

127

Fig. 32. Proposed mechanism of MMA catalysis by MtmB. A modified mechanism of MtmB catalysis is proposed here where MMA forms a chemical adduct (I) with Pyl during methyl transfer to a corrinoid-binding protein (II), which illustrates the two conformational changes of Pyl seen during crystallization (I and III). (IV) Involves the removal of ammonia caused by demethylation of MMA prior to the binding of another MMA molecule for adduct formation. (Modified from Hao et al., 2004; Kryzcki, 2004).

128

129

Fig. 33. Structural superpositioning of the MtmB and MetH crystal structures. (A) The corrinoid-binding domain of MetH is illustrated with its substrate MeTHF (Evans et al., 2004). (B) The MetH crystal (blue) is directly superimposed upon the MtmB crystal (green) with MeTHF shown in the catalytic site of MetH to directly compare to the positioning of Pyl (designated with a red arrow) within the proposed MtmB active site cleft of the αβ TIM-barrel structure. (Krzycki, 2004).

130

131 methylamines. Biochemical tests performed on multiple methylamine MT mutants in which Pyl was replaced by other amino acids showed drastically lower levels of methylamine catalysis to their cognate corrinoid proteins or free cobalamin cofactor (Longstaff, 2004; Joseph Krzycki; personal communication). In the case of MtmB, this background rate was attributed to low levels of chromosomal transcription and translation of MtmB1/2 enzymes during growth on methanol (MeOH) for the MtmB mutants (Longstaff, 2007). We decided to use MtgB, a homolog of MttB, to model the binding of ligand (e.g., GB) into a member of the COG5598 superfamily. These data would assist in multiple directions: I) to identify key residues for methyl transfer to a corrinoid cofactor; II) act as a tool for comparing the proposed positioning of the TMA-Pyl chemical adduct; and III) use as a model to identify other ligands across the COG5598 superfamily. To identify a putative active site in MtgB we compared conserved sites from the clade containing MtgB along with crystalized cobalamin- dependent methyltransferases (PDB:1Q8J and 2I2X) (Evans et al., 2004; Hagemeier, et al., 2006) and binding studies of QAs in the betaine/choline/carnitine transporter (BCCT) family (Casagrande et al., 2008; Jack et al., 2000; Wong et al., 2012). Previously, structural studies on the BCCT family demonstrated the binding of QAs (e.g., GB, choline, carnitine) through Van Der Waal’s forces and cation-Π bonding of aromatic residues to the methyl groups of the amine moiety (Fig. 34) (Schiefner et al., 2004). Utilizing both modeling and ligand prediction software, a putative active site for GB within 2QNE was proposed that is similar to what is seen in QA- binding BCCTs, near the predicted Pyl-TMA adduct location of MttB (Joseph Kryzcki; personal communication) which may present for the first time an evolutionary relationship between GB and TMA catalysis within the COG5598 superfamily.

132

Fig. 34. Structure of the Escherichia coli ProX transporter with bound GB. The proline- betaine transporter from E. coli was crystalized with GB and reveals the interaction of several aromatic residues (W65, W188, and W140) with the amine moiety and interaction of polar residues (H69, G141, and C142) to the carboxyl group of GB that protrudes outside the cation-Π cage. (Schiefner et al., 2004).

133

134

Materials and Methods

Protein sequence and structure acquisition. Structures of MtmB (PDB:1NTH) and MtgB (PDB:2QNE) were retrieved from the Protein Data Bank (PDB) in pdb format for analysis (http://www.rcsb.org/pdb/home/home.do). Complete amino acid (aa) sequences for MtbB (467aa) (GI:115589808) and MttB (495 aa) (GI:215276695) from Methanosarcina bakeri Fusaro were obtained through access to the National Center for Biotechnology Information (NCBI) (http://www.ncbi.nlm.nih.gov).

Homology modeling. MtbB and MttB sequences were submitted through the Multi-source Protein Structure Threader (MUSTER) program to obtain a template structure for these enzymes (Wu and Zhang, 2008). MtbB was threaded against the deposited PDB entry 1NTH while MttB was threaded to 2QNE. The structure of MtmB and homology models of MtbB and MttB were truncated at both the N- and C-termini in Chimera (Pettersen et al., 2004). N-termini truncations were at THR2-GLN99 (MtmB), ALA2-LYS58 (MtbB), and ALA2-PRO82 (MttB) while C- termini truncations were at THR378-PHE458 (MtmB), GLY373-ALA466 (MtbB), and SER375- MET494 (MttB). An additional loop was removed from the MtbB from residues SER93- VAL123 which entered into the αβ TIM-barrel structure of the enzyme. Only the αC trace is shown with the exception of the Pyl residue in O202 (MtmB) or place holder residue (ALA) for Pyl at O366 (MtbB) and O334 (MttB). Separately, the MtbB and MttB models were superimposed upon the 1NTH structure within ~ 2.0 to 3.0 Å distance cutoff using the Needleman-Wunsch algorithm (Needleman and Wunsch, 1970) with a BLOSUM-62 matrix with a gap extension penalty of one.

Modeling of GB within the 2QNE structure. SwissDock (http://www.swissdock.ch) from the Swiss Institute of Bioinformatics as well as the energy-based method through Q-site finder (http://www.modelling.leeds.ac.uk/qsitefinder/) were both used to analyze the 2QNE structure. Structural sites that corresponded to the interaction of the methanol methyltransferase bound to its cognate corrinoid protein (PDB:2I2X) (Hagemeier et al., 2006), location of methyl-THF in MetH (PDB:1Q8J) (Evans et al., 2004), and the like binding from BCCTs were given the highest

135 weight based on conserved residues in the MtgB containing clade of COG5598 and subsequently modeled in Chimera manually.

136

Results

Pyl resides in different regions of the proposed catalytic sites in the methylamine methyltransferases. There is currently no overall structural comparison between the methylamine methyltransferases as only MtmB has a publicly available crystal structure (PDB:1NTH) (Hao et al., 2002). To understand the role of Pyl in the methylamine enzymes, it may be beneficial to compare these non-homologous proteins through both biochemical and structural studies. Modeling of MttB and MtbB demonstrate that the Pyl residue likely projects into the interior cleft of the αβ TIM-barrel, much like MtmB (Fig. 35A) where methylamines are proposed to bind via chemical adduction to Pyl (Gaston et al., 2011; Hao et al., 2002, 2004; Krzycki, 2004). This site allows for the docking of a cognate corrinoid-binding protein while positioning the cobalamin cofactor for supernucleophilic attack. Interestingly, these three enzymes lack sequence homology but have a conserved structure specific to other cobalamin- dependent methyltransferases such as the cobalamin-dependent portion of MetH (Evans et al., 2004) (Fig. 33). This allowed for the modeling of MtbB by using MtmB as a template (Fig. 35B) or the structural comparison of modeled MttB against MtmB to compare the positioning of Pyl (Fig. 35C). Consistent with previous comparisons between MtmB and MetH, the surrogate Pyl residue in MtbB (Fig. 35B) and MttB (Fig. 35C) projects into the interior cleft of the proposed catalytic site. The structural modeling of MtbB using MtmB as a query template was considered a poor alignment yielding a coverage of 0.854 with a Z-score of 3.281 (Wu and Zhang, 2008). The modeled MtbB structure contained one major structural anomaly: a loop threading through the αβ TIM-barrel that was manually removed. Further remodeling and less stringent restraints on the structural alignment failed to alleviate these errors, suggesting that biophysical data will be required for more accurate analysis of the MtbB structure. MttB was initially modeled to MtgB (PDB:2QNE) which shares ~31% aa identity leading to 0.917 coverage with a Z-score of 17.228. In both cases the overall coverage of structure was high in both data sets but the specificity of sequence confidence along the structure was poor for MtbB/MtmB compared to MttB/MtgB. This is unsurprising as MttB and MtgB are homologus enzymes within the ~30% aa cutoff suggested by Wu and Zhang (2008) for accurate threading. The resulting overlay reveals more

137

Fig. 35. Structural overlay of Pyl’s location within each MtxB MT. (A) Chain A of MtmB from the deposited pdb file, 1NTH, is illustrated to show the direct position of Pyl within the proposed active site cleft. (B) MtbB (blue) was modeled with MUSTER using 1NTH (brown) as a template. (C) MttB (blue) was threaded in MUSTER against 2QNE and then superimposed onto 1NTH (brown). N- and C-termini were trimmed to improve clarity of the active site with the N-termini being represented as red and the C-termini being represented in dark blue. Pyl is labeled in the 1NTH structure (A) and highlighted in green while the proposed location of Pyl in (B) and (C) is also colored green.

138

139 overall global changes in the folds between MttB/MtmB compared to MtbB/MtmB which is likely an artifact caused by use of MtmB as the template for MtbB. Whereas the MttB/MtmB model is a more accurate estimation of structure as the template quality of MttB is of higher quality by modeling it from a crystallized homolog.

Modeling MtgB with GB through structural analysis and bioinformatics. The 2QNE structure of MtgB (Fig. 36) was analyzed for ligand binding with SwissDock and Q-site Finder. This Q-Site Finder generated eight different putative binding sites for GB were curated based on their location relative to that of the methyl-THF domain of MetH (PDB:1Q8J) (Evans et al., 2004) or similar active site relative to the methanol methyltransferases (PDB:2I2X) (Hagemeier et al., 2006). The region selected contained several sites from Q-Site Finder; Y94, G95, L197, N199, R309, H345, and F353 (Fig. 36B). Using these data and SwissDock, GB was modeled into 2QNE, which showed a similar association of aromatic residues responsible for amine coordination present in the ProX structure (Fig. 34) (Schiefner et al., 2004).

140

Fig. 36. Proposed positioning of GB into MtgB apo-structure based on Q-Site Finder analysis. (A) The deposited MtgB pdb structure (2QNE) is represented here as a dimer from studies previously performed by the Joint Center for Structural Genomics. (B) SwissDock/Chimera model of MtgB positioned with GB by Q-site finder predictions of the active site.

141

142

Discussion

The incorporation of Pyl within proteins is an interesting phenomenon and is difficult to analyze as regulation of Pyl biosynthesis must be addressed and examined in those organisms with the capability of producing Pyl. In methanogens, Pyl incorporation is efficient within the methylamine methyltransferases (Longstaff et al., 2007). Through comparing the first Pyl- containing structure, 1NTH, to modeled MtbB and MttB, which demonstrates different spatial orientations of Pyl within the predicted active site cleft of these enzymes. It is interesting to observe that three non-homologous enzymes at the sequence level each incorporate this rare biological amino acid in their polypeptide structure. However, these enzymes are not the only proteins predicted to contain Pyl (Galagan, et al., 2002). The first enzyme outside the methylamine methyltransferases to be confirmed with Pyl was incorporated into an enzyme known as Thg1, a reverse Watson-Crick DNA polymerase, from Methanosarcina acetivorans C2AT (Heinemann et al., 2009). Pyl in Thg1 suggests the continual evolution of the M. acetivorans C2AT genome but also demonstrates the ability of Pyl to be expanded into other proteins through point mutations despite strong selection against the UAG codon in methanogenic archaea (Heinemann et al., 2009). Interestingly, under TMA-growth conditions, proteomic analysis revealed that several Pyl-proteins were produced by a bacterium known as Acetohalobium arabaticum (Prat et al., 2012). No direct biochemical tests were performed on any of these enzymes to address the function of Pyl in A. arabaticum proteins but studies with Thg1 demonstrated no alterations in catalytic activity with or without the Pyl residue (Heinemann et al., 2009; Prat et al., 2012). This suggests that organisms can encode Pyl into enzymes through random genomic drift and that encoding of the residue will likely persist if not deleterious to the function the protein or survivability of the organism. Therefore, the placement and essentiality of Pyl in the methylamine methyltransferases raises questions about what residues are required for catalysis with Pyl and whether those residues are required for the formation the proposed methylamine-Pyl adduct. If a chemical adduct is formed by a respective methylamine and Pyl, what was(were) the driving force(s) that led to the determination of the location of Pyl in these structures? This may be explained by the relationship of non-Pyl and Pyl homologs within each methylamine methyltransferases family and the resulting coordination of corrinoid-binding proteins within the binding pocket of these enzymes. However, failure to

143 confirm ligand bound co-crystals of the methyltransferase has proven difficult thus far for physical detection of the formation of a methylamine-Pyl adduct (Joseph Krzycki; personal communication). We then turned to a homolog in the TMA methyltransferase family for which we had new biochemical data and completed structure. Desulfitobacterium hafniense Y51 encodes four members of the COG5598 superfamily, one of which is highly similar in aa identity to bona fide TMA methyltransferases from methanogenic archaea. The genomic location of the DSY3156 gene (mtgB) and appearance near BCCT genes (DSY3154 and DSY3149) suggested to us that this enzyme may in fact be a cobalamin-dependent quaternary amine methyltransferase rather than a TMA methyltransferase. Additionally, the lack of Pyl within MtgB suggests that a more complex amine would bind in the catalytic site of the enzyme, contrary to what is seen in MttB, which would require Pyl. A small amine compound has few opportunities for hydrogen bonding; therefore, a true MttB would presumably require Pyl. The deposited structure of MtgB, 2QNE, was used for ligand-binding studies with GB using both SwissDock and Q-site Finder. This analysis was guided through biochemical experimentation of MtgB and comparison to known GB-binding transport proteins (Schiefner et al., 2004). We expanded upon this by collaborating with Dr. Bing Hao to crystallize MtgB with its respective substrate (e.g., GB). These data are likely to provide the first major support of structural information within the COG5598 superfamily related to direct binding of an enzyme’s substrate for a member of COG5598. The most important aspect of this finding is that the predicted binding pocket of MtgB aligns closely with the proposed formation of a TMA-Pyl adduct by comparing it to the unpublished apo-MttB crystal (Joseph Krzycki; personal communication). This suggests one of two possibilities in which I) Pyl entered a family of corrinoid-dependant N-methyltransferases to now include small methylamines or II) loss of Pyl led to the utilization of more complex amines. The multiple sequence alignment (MSA) of the COG5598 was analyzed with the understanding of GB activity within MtgB. The majority of the tree contains a putative active site similar to MtgB where there is a conserved tyrosine (Y94) above an invariant arginine (R302) and glycine (G92) with aromatic residues. This suggests that a majority of these enzymes are likely GB methyltransferase or diversified to have specificity for different quaternary amines, as aromatic residues are thought to be key to trapping quaternary amines (Schiefner et al., 2004; Ziegler et al., 2010). This range of substrate binding is observable when analyzing the BCCT

144 superfamily to their substrate. BCCTs typically have the ability to transport multiple different compounds (Ziegler et al., 2010). However, at other times they can be oddly specific (Kappes et al., 1996; Wetzel et al., 2011), based on substrate competition, extra- or intracellular substrate conditions, gene regulation, and metal availability. This makes it difficult to accurately predict what reactions the majority of these non-Pyl MttB enzymes may catalyze since we lack physiological characterization of many organisms under quaternary amine growth conditions, unlike D. hafniense Y51. Many of these COG5598 members may catalyze reactions with different ligands and maybe these enzymes are ligand specific as there has yet to be an example of a methyltransferase from any MtxB family having multiple catalytic functions. Strict analysis of each group in the superfamily should be performed to help unravel the functions of COG5598, as catalytic residues from these experiments may help to determine QA-binding specificity within BCCTs. These data led us to hypothesize that in anoxic environments there would be a close connection of quaternary amine metabolism to increasing numbers of genes encoding non-Pyl MttBs. Efforts were focused on the presence of QA utilization by methanogens in aquatic environments which have recently been reported (L’Haridon et al., 2014; Watkins et al., 2012, 2014). Our study showed that different methanogen strains we had isolated contained numerous different non-Pyl mttB genes in distinct operon structures which was consistent with our predictions (Ticak et al., 2014a). This supports our hypothesis that these non-Pyl MttBs may in fact be different QA methyltransferases. In the course of this analysis we discovered two of the first enzymes lacking Pyl clustered within the Pyl clade, a phenomenon which suggests that either Pyl is replaceable for TMA methyl transfer, the enzymes may catalyze a compound(s) very similar to TMA, or the enzymes are dysfunctional. This raises questions about whether divergent evolution of the methylamine methyltransferases may be occurring. Why would the TMA-Pyl adduct be reiterated in DMA-Pyl and MMA-Pyl if one methyltransferase (e.g., MttB) chemically binds each pairing initially after demethylation? Is this a direct effect of co-evolution with the corrinoid-binding proteins and positioning of Pyl or convergent evolution of Pyl into three different protein families? Future studies must be aimed at a more detailed biochemical examination of the Pyl clade and the physical formation of the methylamine-Pyl adduct must be undertaken to provide more information regarding the evolution of Pyl and its advent into cobalamin-dependent methyltransferases.

145

Acknowledgements

I would like to thank Dr. Joseph A. Krzycki for the critical review and editing of this chapter and for the initial study analysis of the proposed binding site of glycine betaine along with shared knowledge of the requirement for Pyl in methylamine methyltransferases. I would also like to thank Dr. Michael Chan and collaborators for their work on the MttB apo-structure in collaboration with Dr. Joseph A. Krzycki which allowed for the intial comparison of the proposed active sites between MtgB and MttB. I thank the ongoing collaboration with Dr. Bing Hao and Dr. Yunfeng Li on the efforts relating to the crystallization study of MtgB with glycine betaine. Finally, I would like to thank my advisor Dr. Donald J. Ferguson for allowing me to work on this project and continuing with its analysis.

146

APPENDIX A

Cloning and expression of auxiliary genes predicted to play roles in quaternary amine- dependent methylotrophy in Desulfitobacterium hafniense Y51

The genome of D. hafniense strain Y51 contains three operons containing non-Pyl mttB genes, which are all unique in structure (Fig. 10). Culturing studies with glycine betaine (GB) revealed little to no transcript production for the other mttB genes (DSY3648, DSY4970, DSY4971) besides the mtgB gene in comparison to pyruvate grown cells. These data led us to predict that these operons may in fact be used for growth on compounds other than GB. The second operon encoding the DSY3648 gene has a conserved gene neighborhood as seen in methanogens and other SRBs. The third operon encoding both a Pyl (DSY4970) and non-Pyl (DSY4971) genes belongs to a group of operons which are less common among sequenced microbes but still appears in both archaea and bacteria (Fig. 10 and 30). The unique aspect of the third operon is that the predicted betaine/choline/carnitine transporter (BCCT) for that pathway is several genes upstream, which is adjacent to choline TMA-lyase (cutC/cutD) and microcompartment genes (Craciun and Balskus, 2012). It is tempting to speculate if choline is brought in by D. hafniense Y51 through DSY5010 then CutC/CutD splits choline into TMA for use by DSY4970, which could explain the continual presence of the pyl operon in this set of organisms, it is also interesting to note that a relative, D. hafniense DP7, was isolated from human intestines which may play a role in choline, GB, and TMA homeostasis (Van de Pas et al., 2001). Lastly, if the appearance of linked non-Pyl and Pyl mttB genes is seen in multiple organisms, it may speak to the biochemical pathway for which it is responsible. Enzymes for like pathways have a tendency to compartmentalize themselves in similar or close operons; this may suggest a QA similar to TMA such as QMA which may act as a substrate for this non-Pyl MT as we predict non-Pyl MTs to be responsible for methyl transfer reactions using a variety of QAs as substrates and biochemical evidence for QMA methyltransfer is established (Asakawa et al., 1998). We decided to clone each gene in these operons to begin biochemical tests and possibly crystallization of these new enzymes for structural analysis. The CBPs (DSY3647 and DSY4973), pterin methylases (DSY3646 and DSY4972), CBP activase (DSY3152), and non-Pyl

147

APPENDIX A

MTs (DSY3648 and DSY4971) were cloned into pSpeedET via PIPE cloning (Klock and Lesley, 2009) with primers listed in Table 4. Briefly, 100 ng of genomic DNA, 0.5 µM F and R primers for both vector and insert template, 1.5 U Phusion DNA Polymerase (NEB, Inc.), 1x Phusion Buffer, and 200 µM dNTPs under the PCR cycling conditions specified (Klock and Lesley, 2009). The Pyl mttB gene was cloned into pDL03c using standard cloning techniques prior to site-directed mutagenesis (SDM) to remove an internal restriction site (SphI) for downstream subcloning into pDL05c with a QuikChange Lightning Site-directed Mutagenesis Kit (Agilent Technologies, Inc.), per manufactuer’s guidelines. The gene insert from pDL03c was then subcloned into pDL05c using a double digest with SphI for both gene and vector followed by T4 DNA Ligase (NEB, Inc.), per manufactuer’s guidelines. Once in pDL05, the plasmid containing the DSY4970 gene was transformed via PEG-mediated transformation as previously reported for Methanosarcina acetivorans C2AT (Dhungana, 2014) while pSpeedET clones (DSY3152 and DSY3155) were chemically transformed into E. coli BL21(DE3). The pSpeedET derivatives (pSpeedET_DSY3152, pSpeedET_DSY3155, and pSpeedET_DSY4971) and pDL05c derivative (pDL05c_DSY4970) were sequence confirmed by BigDye Terminator and stored as glycerol stocks in E. coli EC100 at -80˚C. The DSY4971 protein was produced by similar methods to the purification of MtgB (Chapter 1 – Materials and Methods: DSY3156 and DSY3157 Production and Purification) while the DSY4970 protein was produced in M. acetivorans C2AT as it is the only genetically amendable strain in our lab which contains the pyl operon for Pyl insertion into a nacent polypeptide. Production of DSY4970 was first attempted at 37 ˚C in HS media (Metcalf et al., 1997) amended with puromycin (200 µg/mL) to maintain selective pressure for individuals containing the plasmid but production of DSY4970 proved unsuccessful. We then tested room temperature (RT) growth versus 37 ˚C growth experiments which showed a more robust production of DSY4970 at RT (data not shown). We concluded that production of DSY4970 may be low from western blot analysis so we performed upscaled growth of the DSY4970 containing strain in 15 L of HS media to increase protein yields. Growth after one month at RT

148

APPENDIX A was halted by dispensing cultures into 250-mL centrifuge tubes and collected by centrifugation at 4 ˚C at 6000 x g. Sonication was used to disrupt the cells in 30 s bursts at 30 % amplitude for 10 min. Both DSY4970 and DSY4971 extracts were clarified by ultracentrifugation for 1.5 h at 250,000 x g to remove insoluble material prior to purification of recombinant enzymes. Both DSY4970 and DSY4971 were purified in the same manner as MtgB (Chapter 1 – Materials and Methods: DSY3156 and DSY3157 Production and Purification), quantitated via Bradford reagent (Bradford, 1976), and visualized for homogeneity with SDS/PAGE gels (Fig. 37) in comparison to MtgB. Free Cbl methylation experiments were performed for both DSY4970 and DSY4971 enzymes as reported for MtgB (Chapter 1 – Materials and Methods: Spectrophotometric Assay of GB:Cob(I)alamin Methylation), which resulted in no measurable methylation activity for either enzyme despite extension of assay times to one hour per substrate which was repeated twice with separate purifications of each enzyme. This suggests a lack of function either at the level of cobalamin-binding, as MTs of this family are often known not to methylate free cobalamin (e.g., bona fide MttB), incorrect substrates were used for these assays, or the enzymes are dysfunctional. To address the first issue, we are attempting to purify each MTs respective cognate corrinoid-binding protein in either E. coli or Shimwellia blattae ((Mac Nelly et al., 2014) to eliminate the chance non-cofactor binding confounding further biochemical analysis to solely focus on activity at the level of substrate. To the second point, we will expand the range of substrates tested during these in vitro assays as only QMA, TMA, and GB were tested. Finally, if there is an issue with recombinant protein production we could begin by removal of hexahistidine tag utilized for affinity chromotrography through the TEV protease site immediately downstream of the hexhistidine tag or change the location and/or type of affinity tag utilized for purification.

149

APPENDIX A

Table 4. Primers for the generation of expression vectors in pSpeedET and pDL05c

Name Sequence Study

DSY3152 F (PIPE) 5’- ctgtacttccagggcaTGATCATCAGGATAAAGCATTACGGAG -3’ This study

DSY3152 R (PIPE) 5’- aattaagtcgcgttaGTCTGTTGGCTCCAAATACATACTTCT3’ “

DSY3155 F (PIPE) 5’- ctgtacttccagggcATGATCATGAGTTTATTGGATGAACTG -3’ “

DSY3155 R (PIPE) 5’- aattaagtcgcgttaTTTGCCCAAAATCTTATTAGCAAGTTC -3’ “

DSY3646 F (PIPE) 5’- ctgtacttccagggcaTGGGTTATGTGATGCTAATTATC -3’ “

DSY3646 R (PIPE) 5’- aattaagtcgcgttaGTCTTCGTATAAGCCCTTGC -3’ “

DSY3647 F (PIPE) 5’- ctgtacttccagggcATGAGCGATTTAACGAAGTTAGC -3’ “

DSY3647 R (PIPE) 5’- aattaagtcgcgttaAACCGCCAGTTTCTCGCAC -3’ “

DSY3648 F (PIPE) 5’- ctgtacttccagggcATGCAAAGATATCAGATTCTGTC -3’ “

DSY3648 R (PIPE) 5’- aattaagtcgcgttaTTTCTCTTTATATTTTCTTAAAGCGG -3’ “

DSY4971 F (PIPE) 5’- ctgtacttccagggcATGGCACTTAAATCAAAATTGGAAGTA -3’ “

DSY4971 R (PIPE) 5’- aattaagtcgcgttaCGTCTAACTTTTTGAATATAGGACTT -3’ “

DSY4972 F (PIPE) 5’- ctgtacttccagggcaTGTTAATTGTTGGTGAATTGATCAATACCAGC -3’ “

DSY4972 R (PIPE) 5’- aattaagtcgcgttaACTGCCTTCATACAGCCCATTGCGAT -3’ “

DSY4973 F (PIPE) 5’- ctgtacttccagggcaTGAAAATGAGTGATTTTGCCC -3’ “

DSY4973 R (PIPE) 5’- aattaagtcgcgttaATTCAAAAATCCTTTGGCCATAT -3’ “ pSpeedET_IF 5’- taacgcgacttaattAAACGGTCTCCAGCTTGGCTGTTTTGGC -3’ (Klock and Lesley, 2009) pSpeedET_IR 5’- gccctggaagtacagGTTTTCGTGATGATGATGATGATG -3’ (Klock and Lesley, 2009)

Dh mttb1 pDL03c F 5’- GGCACTCGAGTAGGTGACCAGTCCCAAAATGATTTTAAT “

AAATTAAGGAGGAAATTCATATGgttcaggcgggcgttgc -3’

Dh mttB1 pDL03c R 5’- GCATccgcgggatttattattattattagtggtggtggtggtggtgTCTGTAGTGTG “

CTGCTCCTGGtatcaatgatctggcaactcc -3’

Dh mttB1 NdeI SDM F 5’- agatcatacctatgcccaCatgagaacacagtcccaga -3’ “

Dh mttB1 NdeI SDM R 5’- tctgggactgtgttctcatGtgggcataggtatgatct -3’ “

150

APPENDIX A

Fig. 37. SDS/PAGE of heterologously expressed D. hafniense Y51 MttB enzymes. Protein standards were co-electrophoresed along with MttBs on a discontinuous 15 % Bis-Tris SDS/PAGE gel with sizes highlighted to the left of the gel by the lane marked (L) and the three MttBs (MtgB, DSY4970, and DSY4971) are in lanes 1, 2, and 3 respectivly with the predicted Pyl containing enzyme highlighted with (Pyl). The predicted size for DSY4970 would be ~ 35 kDa if Pyl or another residue was not present in the polypetide chain (i.e. premature termination of the polypeptide at the Pyl position), thus, demonstrating full-length production of DSY4970 which may contain Pyl.

151

APPENDIX A

152

CONCLUDING REMARKS AND FUTURE DIRECTIONS

The molecular mechanism for demethylation of GB and its derivatives in anaerobic microbial physiology has remained unknown since its first observations back in 1959 (Hayward and Stadtman, 1959). Since that time, the importance of QAs has steadily increased in studies as diverse as ecology, compatible solute transport, and human health (Tang et al., 2013; Wang et al., 2011, 2014; Ziegler et al., 2010). This mandated a thorough investigation of what enzymes are responsible for these pathways, which led us to investigate the genomes of hundreds of facultative and obligate anaerobes. In the course of analyzing these microbial genomes, we recognized many of these organisms possessed BCCTs, a realization that was not surprising since GB is an osmoregulatory compound who’s intercellular concentrations could be regulated through either direct transport or through conversion from choline or carnitine (Wood et al., 2010). However, there was a significant frequency of BCCTs near predicted genes for TMA demethylation, which belong to the COG5598 superfamily. These MttBs were originally predicted to be for “methane metabolism” or “TMA demethylation” despite lacking a proposed key residue responsible for TMA catalysis known as Pyl (Ferguson and Krzycki, 1997; Hao et al., 2002, 2004; Kryzcki, 2004; Mahapatra et al., 2006; Paul et al., 2000). Furthermore, the canonical biogenic formation of methane is a process unique to methanogens; thus, the organisms encoding these non-Pyl MttBs were unlikely to be participating in this process as the overwhelming majority of organisms were bacteria. These organisms also lacked known anaerobic GB catabolic enzymes or the betaine/sarcosine/glycine reductase system which is responsible for the formation of acetate, TMA, and MMA through anaerobic fermentation (Andreesen, 1994, 2004; Meyer et al., 1995). This suggested the possibility that these gene neighborhoods may be involved in QA demethylation as two known GB demethylatiing organisms, E. limosum and S. ovata, also contained these conserved gene neighborhoods (Möller et al., 1984; Mueller et al., 1981). We proposed using an undescribed QA-utilizing organism to test our hypothesis and to this end we choose D. hafniense Y51 (Futagami et al., 2006). D. hafniense Y51 is a gram positive, low G+C, Firmicute that was reported for using only a few electron donors but a vast array of electron acceptors (Futagami et al., 2006). This provided us a good candidate to test if a SRB could anaerobically demethylate QAs by respiration; furthermore, unlike E. limnosa and S.

153 ovate, this organism is not an acetogen, meaning that respiration would be directly linked to CO2 formation and production of demethylated products. The growth of D. hafniense Y51 on GB, DMG, MMG, Cho, and TMA (data not shown) suggested to us that this organism was a robust C1-metabolizing member in the environment in which these compounds may be of equal or greater concentrations than pyruvate. We decided to focus our efforts on unraveling the molecular mechanism of GB metabolism in D. hafniense Y51 as it provided the highest growth yields and had similar gene neighborhoods to known GB demethylating acetogens, E. limnosa and S. ovata, suggesting that these three organisms may in fact undergo GB demethylation through homologous pathways.

Anaerobic GB respiration is initiated through a non-Pyl MttB homolog and a pterin- dependent methyltransferase encoded by D. hafniense Y51. Growth, transcript abundance, proteomics, and biochemical investigations of the putative C1 operons were performed on D. hafniense Y51. Growth of D. hafniense Y51 was compared with pyruvate- versus GB-grown cells as pyruvate was unlikely to invoke the upregulation of C1 metabolic genes. This comparison led to identifying the putative operon containing DSY3156 as a candidate for demethylation of GB as both transcript and proteomics correlated significantly when grown on GB compared to pyruvate. Recombinant DSY3156 was produced and experiments revealed that it was in fact not a TMA MT but rather a GB MT (Fig. 16). DSY3156 was designated MtgB (methyltransferase glycine betaine, B) and was tested further to confirm the biological formation of DMG from GB from this enzyme (Fig. 18). We then decided to test the adjacent gene which showed homology to a methanogenic CH3-Cbl:pterin methyltransferase known as MtrH (Hippler and Thauer, 1999). DSY3157 was capable of methylation of THF with CH3-Cbl cofactor and was designated MtgA (methyltransferase glycine betaine, A). Together these two enzymes establish a predicted pathway through a Cbl cofactor leading to the methylation of THF from GB, thus establishing for the first time a molecular mechanism for anaerobic GB demethylation. Furthermore, we established that SRB are in fact likely to be important methylotrophic organisms, as often times they are regarded as non-methylotrophic in nature though they are almost always commonly found with methanogens. D. hafniense Y51 contributes to physiological diversity of microbes by performing a rare event through methylotrophic growth

154 coupled to nitrate reduction (Fig. 13) that is only until recently been investigated (Auclair et al., 2010; Chistoserdova et al., 2009). While studying D. hanfniense Y51 we also discovered two additional operons encoding non-Pyl mttB genes (Fig. 10). Phylogenetically, the DSY3648 protein belongs to the same clade as MtgB while non-Pyl MttB DSY4971 and the Pyl MttB DSY4970 belong to different clades of COG5598. We attempted to purify recombinant DSY3648 but have so far been unsuccessful under the conditions tested. DSY4971 and DSY4970 were recombinantly expressed in either E. coli BL21(DE3) or M. acetivorans C2AT, respectively (Fig. 37). Both DSY4970 and DSY4971 were tested for the ability to methylate free Cbl from QAs or TMA but no methylation of Cbl was observed for these enzymes. We also cloned the genes encoding the remaining CBPs, pterin MTs, and predicted corrinoid reductase into the pSpeedET (Table 4) for further completion and biochemical analysis of these putative enzyme pathways. The discovery of GB demethylation through a non-Pyl MttB highlights the potential of a wide-spread diversity of GB-utilizing organisms ranging from the rhizosphere, to aquatic environments, and to human health. This highlights the necessity for further ecological and biochemical studies on the distribution of QAs and organisms responsible for these pathways. Furthermore, the realization of GB demethylation through MtgB supports our hypothesis that TMA is converted to a quaternary amine during TMA-Pyl adduct formation prior to supernucleophilic attack by a CBP. These data direct us to focus on not only the mechanisms and diversity of the COG5598 superfamily but in the organisms encoding them and the environments they inhabit, by which we can begin to explain the evolutionary pressure(s) that selected for Pyl and mechanisms of QA metabolism.

Novel methanogen isolates demonstrate direct GB-, Cho-, and QMA-driven methanogenesis, providing a like between QA demethylation in archaea and bacteria. Analysis of COG5598 and comparisons of anaerobic pathways of D. hafniense Y51 and methanogenic archaea lead us to predict the likely appearance of GB-utilizing methanogens in anaerobic environments. The Tanaka, Sass, and Laurent laboratories each reported the isolation of QA utilizing methanogens from oceanic sediments (L’Haridon et al., 2014; Tanaka, 1994; Watkins et al., 2014, 2012), which supported our hypothesis that there are GB-utilizing methanogens in this ecological system. We performed enrichment studies of sediments from

155

Southwest Branch Back River located in Virginia to test this hypothesis. Initially, enrichments were performed with Cho, GB, and QMA of which all three yielded methane within one to two days (data not shown). We took measures to try to limit bacterial influences of QA degradation through antibiotic inhibition of SRB that are commonly known to degrade QAs into other methanogenic precursors (e.g., TMA and acetate) (Hippe et al., 1979; King, 1984, 1988; Naumann et al., 1983). Isolation of Cho-utilizing methanogens proved difficult as these isolations either did not grow in liquid culture following growth on plates or involved several different organisms as visualized through DIC microscopy, which did not generate methane, suggesting that enrichments were likely forming TMA via choline TMA-lysase (Craciun and Balskus, 2012). Whereas, isolation of GB- and QMA-utilizing methanogens was simpler, despite the appearance of a common bacterial co-culture under QMA-growth conditions that was eventually removed with subcultivation on TMA and antibiotics. Physiological tests and substrate stoichiometry was performed on the GB-utilizing methanogen, Ml vulcani B1d, and QMA-degrading methanogen, Mc. methylutens Q3c, to confirm demethylation of QAs by these methanogens. To further test our hypothesis of the involvement of COG5598 for QA demethylation systems we began genomic sequencing on both strains of methanogens. This study revealed the appearance of an MttB homolog clustering with MtgB within B1d and several unique non-Pyl MttBs encoded by Q3c. It is interesting to speculate on whether the non-Pyl MttB homolog clustering with MtgB (Fig. 29) from Ml. vulcani B1d is in fact a GBMT. We have begun experiments of cloning this non-Pyl mttB gene into the expression vector pSpeedET to recombinantly produce it in E. coli BL21(DE3). If these efforts prove unsuccessful we will grow large cultures of Ml. vulcani B1d and attempt to purify native enzyme in addition to its cognate CBP while later experiments will be aimed at identifying the MT2 for this putative pathway. In the case of Q3c we later discovered that this strain is actually capable of growth with dimethylethanolamine (DMEA) which was only seen after pre-growth with Cho, however, this may be a product of the primed metabolic machinery from Cho-growth as Cho and DMEA lag times are abysmal when coming from non Cho-derived substrates (Watkins et al., 2012). This finding is interesting in a number of ways, as Mc. burtonii Ace-MT is also capable of growth on DMEA (Watkins et al., 2012). The genome of Mc. burtonii Ace-MT encodes one non-Pyl mttB gene whos product clusters with one Mc. methylutens Q3c’s non-Pyl MttB protein (Fig. 29D), suggesting that these gene products may in

156 fact be DMEA MTs, highlighting our hypothesis that this COG5598 family has diversified to include the catalysis of multiple N-methylamines. Furthermore, the remaining two non-Pyl mttB genes are adjacent to a Pyl mttB gene in addition to genes encoding several conserved MMA and DMA MTs or CBPs. Phylogenetic analysis of these homologs revealed these non-Pyl MttBs clustered with the highly conserved TMA MTs within the archaeal-Pyl cluster (Fig. 29C) despite the absence of Pyl. This mandates a strict analysis of the active site of these enzymes and biochemical tests to confirm if they are in fact TMA MTs, if not, there is a possibility that they may be responsible for demethylation of the closest TMA-like compound, QMA. We will begin this by cloning and trying to produce recombinant enzymes to test for their ability to generate

CH3-Cbl from QMA and TMA.

Distribution of QAs and those microbes responsible for QA-utilization may highlight new ecological and human connections. The presence of a QMA-degrading methanogen is not unheard of but the likelihood of isolating one randomly, based on COG5598s’ functions, highlights an unprecedented series of questions. The common thought of the field was that QMA-degrading methanogens were only predicted to be isolated from QMA contaminated sites, which was never stated by Tanaka (1994). Tanaka’s work led to the discovery of a QMA- degrading methanogen known as Mc. methylutens NaT1, which was later shown to carry out demethylation of QMA, through a series of substrate-specific MTs and CBPs, down to ammonium (Sauer et al., 1997; Tanaka, 1994). Unfortunately, the loss of the strain and people’s interpretation of QMA contamination led to the absence of additional information on the molecular mechanism of QMA demethylation. There are few known biological reservoirs of QMA and even fewer uses of QMA industrially (Barceloux, 2008; Hampton and Zatman, 1973; Ohara et al., 1990; Urakami et al., 1990), so the abundance of QMA environmentally is essentially unknown. Thus, we embarked to determine if a brackish environment would allow for the facile isolation of anaerobic organisms utilizing this compound and to our surprise all QMA- enrichments from this study yielded methane. Interestingly, while we tried to isolate what is now Mc. methylutens Q3c we discovered an incredibly difficult to remove bacterium most related to Desulfovibrio sp. P20 (96% identity by nucleic acid from 1330 nucleotides). Removal of the bacteria by subculturing on TMA suggests that QMA may have acted as a substrate for this bacterium or dependence on the methanogen which would be the first anaerobic bacterium

157 capable of QMA-degradation so we are attempting to isolate this organism currently from our original co-culture. We later discovered that Q3c was not the only methanogen isolated with the capability to use QMA, as conversations with Dr. William Metcalf (personal communication) revealed that his laboratory had also isolated a QMA methanogen some years before from Woods Hole, Massachusetts. This suggests a more widespread distribution and formation of QMA in aquatic environments, which may be generated from either biological or geochemical sources similar to the proposed mechanism of DMG decarboxylation (Wu et al., 2002). The physiology of Q3c to grow well in low saline conditions encouraged us to test for growth of QMA-utilizers in freshwater systems such as rivers and lakes. Currently, we have a series of enrichments in the laboratory producing methane from these environments (data not shown) and are starting isolation studies to unravel the microbes that may be responsible for QMA- degradation in fresh water systems. Originally, studies of Cho-enrichments were unsuccessful as methane was generated but isolation attempts led to non-methanogen or isolate that simily failed to grow once isolated off plates. We determined that the choline TMA-lyase system was likely responsible for the formation of TMA leading to methane production in these enrichments as SRB typically have been seen with these pathways and often times contain predicted Pyl-MttBs. This suggests that TMA may in fact be a competitive substrate for both SRB and methanogens, as the generation of TMA from Cho would theoretically be useable by SRB containing Pyl-MttBs, like D. hafniense Y51 (Fig. 10). There have been no biochemical experiments performed on a bacterial Pyl-MttB until we recombinantly produced and purified DSY4970 and the adjacent DSY4971 from D. hafniense Y51 in M. acetivorans C2AT and E. coli BL21(DE3), respectively (Fig. 37). These experiments were performed in a similar manner to studies on MtgB but revealed a complete lack of enzyme activity (Appendix A). This could be explained by the absence of a CBP for these MTs, as the MttB and MtmB enzymes of Methanosarcina spp. are dependent on their cognate CBPs for methyltransferase activity (Burke and Krzycki, 1997; Ferguson et al., 2000; Ferguson and Krzycki, 1997). If continued attempts fail, we can address substrate binding directly through either isothermal calorimetry or differential radial capillary action of ligand assays (DRaCALA) (Roelofs et al., 2011). We revisited the Cho-enrichments with a new realization that one thing was indeed different between the plates in an anaerobic chamber and culture tubes involving the presence of

158 molecular H2. Hydrogen has been shown to increase the rate of growth and sometimes be a requirement for methanogen isolates, especially those related to gut systems (Dridi et al., 2012; Fricke et al., 2006; Lang et al., 2014; Miller and Wolin, 1983, 1985; Muller et al., 1986;

Sprenger et al., 2007). Molecular H2 acts by directly reducing the heterodisulfide (CoM-H-H- CoB) molecule prior to methane formation, thus, generating more energy production rather than loss of electrons through methyl group oxidation to CO2 (Muller et al., 1986). We realized that the anaerobic chamber we use for cultivation typically contains 2-4 % H2 and this can freely diffuse through the plastic containers we use for plate growth under a reducing atmosphere (Chapter 2 – Materials and Methods: Growth and maintenance of enrichment and pure cultures). We tested Q3c’s ability to perform methyltrophic metabolism under 2, 4, 20, and 80 %

H2, which demonstrated an increased rate of growth and maximum optical density by Q3c for only Cho and DMEA (data not shown). Given that H2 is a key component in anaerobic systems for growth and energy (Boone et al., 1987; Traore et al., 1983; Ushida et al., 1997), it may help to explain the exceedingly long lag times and low rates of growth reported by Sass and our laboratory under Cho- and DMEA-growth conditions (Watkins et al., 2012). To further test this hypothesis, we began by performing these isolation experiments again from the glycerol stocks of Cho-grown enrichments cultures after the fourth passage. These newly isolated cultures yielded methane once isolated to tubes with H2 compared to those lacking it (data not shown) though these cultures have yet been examined for purity, so a mixed culture may be present.

The importance of H2 is not only attributed to ecological environments but also the hindgut of insects, the rumen of animals, and intestines of vertebrates (Dridi et al., 2012; Lang et al., 2014; Miller and Wolin, 1985; Sprenger et al., 2007). Methanogens isolated from these systems are typically shown to respond to increasing partial pressures of H2 (Boone et al., 1987). The bulk of methanogenesis from these environments is attributed to hydrogenotrophic methanogenesis (Kumar et al., 2012; Poulsen et al., 2013; Ushida et al., 1997), however, there are increasing reports regarding the importance and prevalence of less understood methylotrophic methanogens from these same systems. These methanogens form what is known as the seventh clade of methanogenic archaea called Methanomassiliicoccales and only recently are studies being performed in order to isolate and characterize their physiological capabilities (Borrel et al., 2014). This highlights the importance of understanding C1-cycling in the symbiosis of higher organisms within the gut, as amelioration of circulating TMA could lessen

159

TMAO levels generated by the host by direct removal of the TMA precursor thus lessening cardiovascular risk via TMAO (Tang et al., 2013; Wang et al., 2011, 2014). These data suggest that studies should be aimed at which anaerobic gut-associated microbes are capable of utilizing QAs that are cardiovascular risk factors (e.g., Cho, GB, and carnitine). This may reveal how C1- cycling pathways occur in the gut and may also lead to the discovery of more functional COG5598 members.

Initial structural analysis between MttB and MtgB reveal a possible evolutionary link to the advent of Pyl and positioning of the proposed methylamine-Pyl adduct. We began studying the putative active site by using Q-site finder to identify potential residues of interaction based on known catalytic sites of other cobalamin-dependant methyltransferases. The αβ-TIM barrel of MtgB was highlighted in Q-site finder experiments and highlighted several residues predicted for binding; Y94, G95, L197, N199, R309, H345, and F353 which was then manually modeled with SwissDock (Fig. 36B). When these residues were compared to the binding of GB in other proteins, namely BCCTs, the binding of GB and proline betaine were both positioned to interact with nearby aromatic residues for forming an cation-Π cage for the amine moiety of GB while polar residues interacted with the carboxyl portion of the molecule (Fig. 34) (Ziegler et al., 2010). This led us to seek collaboration with the Hao laboratory to attempt crystallization of MtgB with GB. The proposed binding of GB within the MtgB structure highlights perhaps the most important aspect of this work, being that we, for the first time, may obtain evidence to suppor the formation of aTMA-Pyl adduct by comparing GB-bound MtgB to the MttB. These data would suggest that the geometry and chemistry of Pyl may have been selected for based on molecular mimicry of QA binding within this superfamily. A complete analysis of the Pyl clade will have to be performed with both archaeal and bacterial members to test whether all of these enzymes are responsible for TMA methyl transfer which contain Pyl or whether there other molecular mimics acting as substrates for some of these enzymes. The diversity of the active sites in COG5598 can be separated into three main groups: the GB-like, TMA-like, and atypical active sites. The vast majority of the COG5598 superfamily seems to possess many conserved residues like the proposed binding of GB in MtgB (Fig. 36B), suggesting that many members of this tree are using either GB or QA molecules. While the

160

TMA-like portion is highly conserved in relation to known TMA MTs from methanogens, suggesting they may in fact all be responsible for TMA methyl transfer, however, Q3c’s non-Pyl enzymes clade into this branch suggesting either replacement of Pyl or breakdown of another small amine (e.g., QMA). When comparing the mttB genes of these two main active site groups, many of the gene neighborhoods contain CBP genes, pterin MT genes, or CoM methylase-like genes, but there are some that seem to lack these hallmark neighborhoods. Most putative operons around the Pyl mttB cluster have putative genes for amino acid like pathways involving: proline betaine, ornithine, lysine, arginine, or histidine. The most common group belongs to histidine degradation pathways with proline betaine being the second most common; however, many of the genomes for these organisms are in the preliminary draft stage making it difficult to actually ascertain what may truly be the most abundant. If non-Pyl or Pyl MttBs within or around the Pyl- clade perform functions other than TMA methyl transfer, it may be that these enzymes are responsible for the demethylation or methylation of these types of compounds (e.g. lysine, histidine, and arginine). This raises the questions if these pathways may in fact also be essential in anabolic rather than catabolic metabolism. Finally, the most distant active-site branch of COG5598 contains members with an active site unlike either of the previous two systems and genomic information of those surrounding gene neighborhoods is the least complete since much of the data was obtained from non-isolation sequencing projects. This highlights the diversity of COG5598 through a protein level analysis, but we must take these data back to isolation and physiological studies of the organisms which contain these pathways to bring more biological and evolutionary insights into Pyl’s advent into COG5598.

GB-binding may represents a link between GB transport and catabolism. The predicted binding site of GB in MtgB was initially compared to studies of BCCTs (i.e. ProX). The conserved chemistry of binding GB seems to have been duplicated within these two systems through the interactions of cation-Π bonding and Van der Waal’s forces locking down the positively charged amine moiety of GB and like QAs (Schiefner et al., 2004; Ziegler et al., 2010). If we are to predict the specificity of transporters for different QAs, studies involving

COG5598 may prove to be useful in delineating what residues are essential to binding the R4- group of QAs as well as if there is a preferential selection for aromatic aa. However, there are likely more elements to BCCTs that we need to understand outside of substrate binding such as

161 their regulation and protein:protein interactions. Many of the known BCCTs being analyzed are in the context of osmolyte influx to counteract environmental cues such as saline stress; we are interested in understanding catabolism of these compounds, which may work in a slightly different manner i.e., protein:protein interaction with methyltransferases. To highlight this, we have long held the idea that these substrate MTs interact directly with the substrate transporters in both archaea and bacteria. Experiments have been performed in the lab (unpublished data) in relation to genetically knocking-out methylamine transporters of either the APC group (MtmP or MtbP) or the BCCT group (MttP1 and MttP2) (Dhungana, 2014). One proteomic study highlighted the occurrence of MttB associated with the membrane fraction (Williams et al., 2010), suggesting an interaction with some membrane protein(s). KOs of the predicted methylamine permeases have revealed attenuation of growth but not abolishment of substrate transport (Dhungana, 2014); implying that these substrates do not require their specific substrate- transporter but overall growth is more favorable when they are present. The binding affinity or

Km for each substrate transporters becomes an interesting question as attenuation of maximal growth phenotype in some mutants versus WT is apparent. Suggesting that either substrate is entering through cross-reactive transporters at much slower levels due to Km of cross-reacting methylamines or there is lack of proper substrate breakdown rates by disrupting some level of protein:protein interaction. If transport of substrate occurs through protein:protein interactions then the rate of substrate breakdown may occur higher as substrate would feed directly to MTs upon transport while associating with the membrane permease thus continually making a forward gradiant of substrate entry. Whereas, if Km for substrate is the main driver, slower levels of substrate influx could have a direct relationship to slower ammounts of methane production and growth yields. It will be interesting to see if these same things are seen with the BCCTs and non- Pyl MttBs which may help to explain why there is almost always a BCCT uniquely found in each putative operon containing mttB genes.

Concluding Remarks. The appearance of Pyl in three methyltransferase superfamilies suggests that Pyl divergently entered each protein family individually as all three groups have both Pyl and non-Pyl containing members. Dr. Rudolf Thauer once told me that difficult things are invented once in nature and from that they are perpetuated; this seems apt to explain the evolution of Pyl as to date; only these three families are capable of methylamine methyl transfer,

162 suggesting a difficult biochemical mechanism. If Pyl had entered these enzymes individually, then that raises the question, what led to Pyl’s advent into their polypeptide structure? Pyl’s biochemical formation likely predates its insertion into the methylamine MTs. In this way, perhaps the formation of the methylamine-Pyl adduct was acted upon by several different Cbl- dependent MTs as a substrate for microbes and by insertion of this amino acid into these MTs a new chemistry and biological amino acid was formed into established Cbl-dependent MT families by reducing turnover of largely abundant methylamine molecules within these anaerobic environments. In future years, direct physical evidence of the Pyl cluster’s formation of the methylamine-Pyl adduct will be necessary to determine the role and function of Pyl in the evolution of anaerobic microbial metabolism.

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