Essential Cofactors in Anaerobic Microbial Consortia Used for Bioremediation: Biosynthesis, Function and Regeneration

ESSENTIAL COFACTORS IN ANAEROBIC MICROBIAL CONSORTIA USED FOR BIOREMEDIATION: BIOSYNTHESIS, FUNCTION AND REGENERATION

Po-Hsiang Wang

A thesis submitted in conformity with the requirements

for the degree of Doctor of Philosophy

Department of Chemical Engineering and Applied Chemistry

University of Toronto

Po-Hsiang Wang

Doctor of Philosophy

Department of Chemical Engineering and Applied Chemistry University of Toronto

2018

ABSTRACT

Most microorganisms in nature live in communities and have developed tightly coordinated metabolism via metabolite exchanges. Cofactors are “helper molecules” in all cells, modulating the activity of many enzymes or serving as the electron shuttle. However, biosynthesis of cofactors, such as cobamides, is sometimes accomplished by only one specific phylogenetic group of microorganisms. Therefore, cofactors and their producers play pivotal roles in the functionality, metabolic rates, and population structure of a microbial community. This thesis focuses on three types of cofactors: (i) cobamides, (ii) NADPH, and (iii) prenylated flavin mononucleotide (prFMN) that are involved in anaerobic bioremediation of chlorinated solvents and aromatic pollutants.

Cobamides are a family of cobalt-containing tetrapyrroles involved in biochemical reactions including methyltransfer reactions, isomerizations, and reductive dehalogenation. This thesis reports the identification of a functional cobamide prosthetic group in tetrachloroethene

ii dehalogenases (PceA) of Desulfitobacterium hafniense and Geobacter lovleyi using a tiered blue-native polyacrylamide gel electrophoresis (BN-PAGE) and liquid chromatography-mass spectroscopy (LC-MS) method.

The metabolic annotations of Dehalobacter restrictus (Dhb), a model organism for bioremediation of chlorinated solvents, were experimentally verified. The verified annotations were written into a constraint-based metabolic model, which identified that NADPH regeneration and de novo serine biosynthesis could be bottlenecks in Dhb metabolism. Using an integrated computational/experimental approach, the stringent nutrient requirements of Dhb were characterized. Further experimental analysis on the Dhb-enriched ACT-3 consortium has revealed an interspecies malate-pyruvate shuttle system between Dhb and its syntrophic partner. prFMN is a newly identified cofactor of UbiD reversible aromatic decarboxylases that are involved in ubiquinone biosynthesis, biological decomposition of lignin monomers, and anaerobic biodegradation of aromatic pollutants. We discovered that in Escherichia coli, dimethylallyl-monophosphate (DMAP), the prenyl donor of prFMN, can be produced from either prenol phosphorylation or from dimethylallyl-pyrophosphate dephosphorylation.

In conclusion, this thesis reports biosynthesis and function of new cofactors as well as new mechanisms for reconciling cofactor regeneration in anaerobic microbial communities.

Realizing these metabolic interdependencies generates opportunities to manipulate the microbiomes for better outcomes in bioremediation and industrial biotechnology. Interestingly, the identified interspecies cofactor exchange mechanism also provides insights into how life gradually evolved into complexity.

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ACKNOWLEDGEMENTS

First of all, I would like to express my gratitude to all the people who have helped me in my life during these four and half years, especially my parents. Without their support, I could not make my dream, studying abroad, come true. Life is evanescent and is filled with surprises. I can still clearly remember the day I landed at Pearson International Airport. During my PhD study, I have met and collaborated with so many peers and professors in BioZone and at the University of Tennessee Knoxville, USA. Their help empowered me to finish my PhD and to write this thesis. Second, I want to acknowledge my supervisor, Prof. Elizabeth Edwards, for her unsparing contributions to my PhD study. While being strict, she never set up a hierarchy between us. She significantly improved my English speaking and writing skills, bridged my jumping ideas, and provided the research resources to satisfy my scientific curiosity. Moreover, she helped me realize the importance of communication and brought sophistication to my research. Furthermore, she always managed to expand my academic networks and paved the way for my future career.

Third, I would like to acknowledge my committee members, Prof. Alexander Yakunin and Prof. Emma Master, and departmental examiner Prof. Radhakrishnan Mahadevan for their support and insightful suggestions on my thesis. I also acknowledge Prof. Frank Löffler, at University of Tennessee, USA, for teaching me the use of multiple lines of evidence approach to gain confidence in scientific hypothesis.

Finally, friendship was an essential element in my PhD life. I would like to acknowledge all my friends, especially Dr. Shuiquan Tang, Dr. Fei Luo, Luz A. Puentes, Shen Guo, and Olivia Molenda in the Edwards Lab; Kayla Nemr, Naveen Venayak, and Kevin Correia in the Mahadevan Lab; Dr. Anna Khusnutdinova and Robert Flick in the Yakunin Lab; and Dr. Jun Yan in the Löffler Lab.

TABLE OF CONTENTS

ACKNOWLEDGEMENTS ...... IV

TABLE OF CONTENTS…………………………………………………………………………………………………………….V

LIST OF TABLES ...... VII

LIST OF FIGURES ...... VIII

LIST OF APPENDICES ...... X

LIST OF ABBREVIATIONS ...... XI

CHAPTER 1 - LITERATURE REVIEW AND GENERAL INTRODUCTION ...... 1 1.1 LITERATURE REVIEW ...... 1

1.1.1 ORGANOHALIDE RESPIRATION ...... 2

1.1.2 CORRINOIDS ...... 11

1.1.3 NADPH: REGENERATION AND SHUTTLE SYSTEM ...... 18

1.1.4 PRENYLATED FLAVINS AND REVERSIBLE DECARBOXYLATION OF AROMATICS ...... 21

1.2 RATIONALE AND RESEARCH OBJECTIVES...... 23

1.3 THESIS OUTLINE AND STRUCTURE ...... 25 1.4 STATEMENT OF AUTHORSHIP AND PUBLICATION STATUS ...... 27

CHAPTER 2: IDENTIFICATION OF FUNCTIONAL COBAMIDE PROSTHETIC GROUP IN REDUCTIVE DEHALOGENASES USING BN-PAGE AND LC-MS ...... 31 2.1 ABSTRACT ...... 31 2.2 INTRODUCTION ...... 32 2.3 MATERIALS AND METHODS ...... 34 2.4 RESULTS ...... 37 2.5 CONCLUSION ...... 41

CHAPTER 3: REFINED EXPERIMENTAL ANNOTATION REVEALS CONSERVED CORRINOID AUTOTROPHY IN CHLOROFORM-RESPIRING DEHALOBACTER ISOLATES ...... 43 3.1 ABSTRACT ...... 43 3.2 INTRODUCTION ...... 44 3.3 MATERIALS AND METHODS ...... 47

3.4 RESULTS AND DISCUSSION ...... 50 3.5 IMPLICATION FOR MICROBIAL ECOLOGY ...... 68

CHAPTER 4: INTERSPECIES MALATE-PYRUVATE SHUTTLE DRIVES AMINO ACID EXCHANGE IN ORGANOHALIDE-RESPIRING MICROBIAL COMMUNITIES ...... 70 4.1 ABSTRACT ...... 70 4.2 INTRODUCTION ...... 71 4.3 MATERIALS AND METHODS ...... 73 4.4 RESULTS AND DISCUSSION ...... 78 4.5 IMPLICATION FOR MICROBIAL ECOLOGY ...... 93

CHAPTER 5: BIOSYNTHESIS AND ACTIVITY OF PRENYLATED FMN COFACTORS .... 95 5.1 ABSTRACT ...... 95 5.2 INTRODUCTION ...... 96 5.3 MATERIALS AND METHODS ...... 98 5.4 RESULTS AND DISCUSSION ...... 105 5.5 SIGNIFICANCE ...... 121

CHAPTER 6:SUMMARY, SIGNIFICANCE, AND FUTURE WORK ...... 123 6.1 SUMMARY ...... 123 6.2 IMPLICATION ...... 129 6.3 FUTURE WORKS ...... 130

REFERENCES ...... 132

APPENDICES ...... 149

LIST OF TABLES

Table 2.1. Proteomic analysis of the Desulfitobacterium (Dsf) hafniense strain JH1 proteins associated with different BN-PAGE slices...... 39

Table 3.1. Dechlorinating activity observed in strain CF cell extracts ...... 55

Table 3.2. Enzymatic activities in cell extracts of E. coli and strain CF and promiscuous threonine aldolase activity of purified Dehalobacter serine hydroxymethyltransferase (SerB) ...... 57

Table 4.1. Stepwise curation of genome-scale constraint-based Dehalobacter metabolic model ...... 79

Table 4.2. Number of NADPH molecules required per amino acid synthesized...... 83

Table 6.1. Summary of the function and symbiotic interactions of cobamides, NADPH, and prFMN in literature and proposed in this thesis (in bold)...... 128

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LIST OF FIGURES

Fig. 1.1. Schematic of RDase-mediated reductive chloroform dechlorination with H2 as the electron donor...... 3

Fig. 1.2. The transfer history of cultures used in the research presented in this thesis...... 7

Fig. 1.3. The structure of corrinoids and their lower ligands...... 13

Fig. 1.4. Anaerobic DMB biosynthesis pathway and corresponding biosynthetic genes in Eubacterium limosum...... 16

Fig. 1.5. Reversible interconversion of NADPH and NADP+...... 19

Fig. 1.6. Coenzyme Q8 (ubiquinone) biosynthesis by MEP isoprenoid pathway and UbiX- UbiD pair for Electron Transport Chain-mediated energy conservation...... 22

Fig. 2.1. Identification of purinyl-cobamide as the native cobamide cofactor in Dsf PceA following non-denaturing, gel-electrophoretic separation of Dsf crude protein extracts using BN-PAGE...... 38

Fig. 3.1. Dechlorination over three serial 10% transfers of strain CF...... 51

Fig. 3.2. Microscopic images of Dehalobacter strains...... 52

Fig. 3.3. Schematic of the proposed metabolic map of strain CF...... 56

Fig. 3.4. Determination of intracellular thiamine derivatives in strain CF cells...... 62

Fig. 3.5. Corrinoid production and guided cobalamin synthesis by strain CF...... 64

Fig. 3.6. Heme production by strain CF and its influence on dechlorination...... 67

Fig. 4.1. Alternative serine biosynthesis in Dehalobacter restrictus via threonine...... 81

Fig. 4.2. Resolving redox balancing in Dehalobacter restrictus metabolic model with integration of experimental data...... 84

Fig. 4.3. Organic acid profile in lactate-fed consortium ACT-3 and syntrophy disruption with heme addition...... 90

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Fig. 4.4. Proposed syntrophy and genome streamlining between Dehalobacter restrictus strain CF and the symbiotic Bacteroides sp...... 91

Fig. 5.1. Phylogenetic analysis of the UbiX family...... 107

Fig. 5.2. Recombinant expression and purification of the UbiX-like AF1214...... 109

Fig. 5.3. Screening for DMAP-producing enzymes...... 111

Fig. 5.4. Proposed biochemical and chemical transformations of prFMN...... 116

Fig. 5.5. Activation of Fdc1 by UbiX-bound or protein-free prFMN...... 119

Fig. 5.6. Fdc1 activation by UbiX-bound and free prFMN cofactors...... 120

Fig. 5.7. Schematic overview of the E. coli enzymes involved in prFMN biosynthesis and associated pathways...... 121

Fig. 6.1. Synthesis of this thesis: biodegrader trinity of chlorobenzene (Cl-benzene) in anaerobic groundwater...... 126

LIST OF APPENDICES

Appendix A: Recipe of Optimized Defined Mineral Medium………………………………..150 Appendix B: Sustainable VC Dechlorination by the KB-1 Cultures Independent of Cobalamin Supplementation with Varying pH……………………………………………...……………153 Appendix C: Isolation of Geobacter lovleyi strain KB-1……………………….……………162 Appendix D: Supplemental Information for Chapter 3…………………...…………………..167 Appendix E: Supplemental Information for Chapter 4………………….……………………184 Appendix F: Supplemental Information for Chapter 5………………….……………………185

LIST OF ABBREVIATIONS

1,1-DCA – 1,1-dichloroethane 1,1,1-TCA – 1,1,1-trichloroethane 1,1,2-TCA – 1,1,2-trichloroethane 1,2-DCA – 1,2-dichloroethane a.a. – amino acids ACN – acetonitrile BN-PAGE – blue native polyacrylamide gel electrophoresis cDCE – cis-dichloroethene CA – monochloroethane CF – chloroform DCM – dichloromethane Dhb - Dehalobacter Dhc – Dehalococcoides Dsf – Desulfitobacterium DMAP – dimethylallyl monophosphate DMAPP – dimethylallyl pyrophosphate DMB – 5,6-dimethylbenzimidazole DNAPL – dense, non-aqueous phase liquid ESI-LC-MS – electrospray ionization-liquid chromatography-mass spectroscopy FMN – flavin mononucleotide GC-FID – gas chromatography Geo – Geobacter IPTG – Isopropyl β-D-1-thiogalactopyranoside MeOH – methanol MoCo – molybdopterin cofactor OHRB – organohalide-respiring bacteria PCE – tetrachloroethene (perchloroethene) PCR – polymerase chain reaction prFMN – prenylated FMN qPCR – quantitative PCR

RDase – reductive dehalogenase rdh – reductive dehalogenase homologous gene rRNA – ribosomal RNA genes TCE – trichloroethene UV-Vis – ultraviolet-visible VC – vinyl chloride

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CHAPTER 1 – LITERATURE REVIEW AND GENERAL INTRODUCTION

This chapter includes the literature review addressing the biosynthesis, function and regeneration of three types of cofactors: cobamides, NADPH, and prenylated FMN, which play essential roles in anaerobic microbial communities, especially those specialized in organohalide respiration and anaerobic aromatic degradation (Section 1.1). Based on the literature review and my research interests, four major research objectives were proposed (Section 1.2) and were studied in some details (Chapter 2-5). Finally, this chapter outlines the overall thesis structure (Section

1.3), along with the statement of authorship and publication status of the results reported in this thesis (Section 1.4).

1.1 Literature Review

Interspecies nutrient exchange, or cross-feeding interactions between microbes, describe the complex network of entangled metabolisms within a microbial community that are essential for maintaining the function and stability of a community (Dolfing, 2013; Embree et al., 2015;

Mee et al., 2014). Mineralization of organics is challenging and is highly coordinated in anaerobic microbial communities, serving as a great model to study microbial symbiosis. Cofactors are essential components required by all the organisms, serving as biocatalysts or electron carriers in cells. However, biosynthesis of these “helper molecules”, like cobamides, are often accomplished by only one specific phylogenetic group of microorganisms (Seth and Taga, 2014). Therefore, cofactors and their producers play pivotal roles in the function, metabolic rates, and population composition of a microbial community. The following four sub-sections report: (1) The mechanism of organohalide respiration as well as the applications and microbial ecology of anaerobic microbial communities enriched with organohalide-respiring bacteria (OHRB); (2) The

1 function, biosynthesis and cross-feedings of cobamides in OHRB-enriched microbial communities;

(3) The function and regeneration of NADPH for cellular anabolism and redox homeostasis; and

(4) The biosynthesis of prenylated FMN for anaerobic aromatic de/carboxylation.

1.1.1 Organohalide respiration

Organohalide-respiring bacteria and reductive dehalogenases

Organohalides are halogenated organic molecules, often hydrocarbons, produced anthropogenically, or biologically for signaling, defense, and antagonism (Richardson, 2016). The most abundant organohalides in the atmosphere are chlorinated methanes (~1 ppb v/v)

(Kuznetsova et al., 2006) produced primarily by intestinal fungi of termites in terrestrial ecosystems and by algae in marine ecosystems. The annual global flux of halogenated methanes is estimated to be at least 4 million tons (Cappelletti et al., 2012; Harper, 2000). Organohalide respiration is a kind of microbial respiration process that fulfills global halogen cycle (Schägger and von Jagow, 1991). Some organohalide-respiring bacteria (OHRB) utilize hydrogen, formate, or acetate as the electron donors to reduce organohalides. Moreover, this process must operate under anaerobic conditions (Lee et al., 2012). The enzymes responsible for the carbon-halogen bond cleavage are called reductive dehalogenases (RDases). To date, various RDases with different organohalide substrates have been identified, such as chloroform (CF)-degrading CfrA from Dehalobacter restrictus CF (Fig. 1.1), PCE-degrading PceA from Desulfotobacterium hafniense, and VC-degrading VcrA from Dehalococcoides mccartyi (Furukawa et al., 2005;

Parthasarathy et al., 2015; Tang and Edwards, 2013). RDases are mostly membrane-bound, heterooligomeric and cobamide-dependent proteins (Adrian et al., 2007). Nevertheless, research has shown that the RDase NpRdhA from Nitratireductor pacificus is a cytoplasmic protein and

2 appears oxygen-tolerant (Payne et al., 2015). Most RDase-encoding operons contain at minimum two genes: rdhA and rdhB. rdhA encodes the catalytic domain and rdhB encodes the membrane anchor (Schumacher et al., 1997). The catalytic domain contains two conserved iron-sulfur binding motifs (CXXCXXCXXXCP)2 with two iron-sulfur clusters as well as a cobamide-interacting motif

(DXHXXGSXLGG) (Banerjee and Ragsdale, 2003; Bendtsen et al., 2005; Futagami et al., 2008).

The presence of a twin arginine signal peptide sequence (RRXFXK) at the 5’ end indicates that

RDases are secreted to the periplasm by the twin arginine translocation (TAT) system, that exports proteins to periplasm only after they have been folded into mature form. Accordingly, respiratory

RDases are periplasmic membrane-bound proteins (Maillard et al., 2003).

Fig. 1.1. Schematic of RDase-mediated reductive chloroform dechlorination with H2 as the electron donor. Abbreviations: B12, cobamide; RdhA, catalytic domain of RDase; RdhB, membrane anchor and electron shuttle subunit of RDase.

Due to the relatively slow growth (1-2 day doubling time) and relatively low cell yield of most native RDase producers, a functional heterologous expression system can significantly facilitate the functional characterization of such a group of “delicate” enzymes. Heterologous expression of functioning dehalogenases was demonstrated in Shimwellia blattae, E. coli, and in

Bacillus megaterium. The RDase PceA from Desulfitobacterium hafniense was actively expressed

3 in S. blattae (Men et al., 2014). The activity of recombinant PceA is significantly improved by the co-expression of PceA-folding chaperone PceT and hydroxocobalamin, the catalytic prosthetic group. However, a folding chaperone is not necessarily involved in the maturation of RDase. For example, recombinant apo-VcrA fused with maltose binding protein (enhance solubility) and a

TEV protease cleavage site was firstly expressed in E.coli. Subsequently, the maltose binding protein was cleaved, and actively reconstituted by the addition of hydroxocobalamin, Fe3+, sulfide, and mercaptoethanol (Parthasarathy et al., 2015). Recently, CF-degrading TmrA from D. restrictus was functionally expressed in B. megaterium (Jugder et al., 2018). The authors removed the transmembrane domains and the TAT signal peptide from TmrA, which enabled heterologous expression of the soluble recombinant TmrA. Moreover, the B. megaterium culture expressing

TmrA was first grown under aerobic conditions to accumulate biomass, and was then switched to anaerobic conditions for isopropyl β-D-1-thiogalactopyranoside (IPTG) induction of recombinant

TmrA.

The crystal structure of two RDases were elucidated recently, which enables a deeper understanding of the catalytic mechanism (Bommer et al., 2014; Payne et al., 2015). Recombinant

PceA from Sulfurospirillum multivorans was expressed and purified from its native host, and structurally characterized. The 464 residue homodimeric RDase includes two corrinoid-binding motifs (139-163 and 216 to 323), a “letter-box” substrate entrance (164-215) and iron-sulfur cluster binding motif (324 to 394). The stacking of two TCE molecules, which resembles the steric structure of a PCE molecule, renders the two chlorinated ethenes a common substrate for PceA.

Another RDase NpRdhA from Nitratireductor pacificus strain pht-3B, which dechlorinates ortho- halogenated phenolic compounds, was heterologously expressed and purified from B. megaterium.

The crystal structure reveals a high similarity to the PceA in S. multivorans. Finally, electron

4 paramagnetic resonance (EPR) analysis suggests a direct interaction between the cobalt on cobalamin and a halogenated compound via a halogen-cob(II)alamin complex formation. Electron paramagnetic resonance (EPR) analysis of VcrA has revealed that the Co (II) of the cobalamin cofactor is reduced to Co(I) by the reduced [4Fe-4S] cluster, and the electron is then transferred to

VC to generate a vinyl radical intermediate (Parthasarathy et al., 2015). Together, two catalytic mechanisms were hypothesized, one proceeding via a heterolytic mechanism with the transient formation of a Cob(III)alamin bond or one proceeding via a homolytic mechanism following formation of an aryl radical through electron transfer from Cob(I).

Bioremediation of chlorinated solvent contamination

Some chlorinated hydrocarbons, such as 1,1,1-trichloroethane (1,1,1-TCA), chloroform

(CF) and trichloroethene (TCE), are ubiquitous groundwater contaminants worldwide due to the historical use in degreasing or for dry cleaning and due to poor disposal regulations in the past

(Furukawa et al., 2005). Prior to 1984, direct release into soil was an established disposal practice of chlorinated solvents (Crofts et al., 2014). Owing to their poor water solubility and density greater than water, chlorinated hydrocarbon spills generally sink deep beneath the water table and form a stable hydrophobic layer, or dense non aqueous phase liquid (DNAPL), which gradually diffuses into the groundwater flow and soil (Sleep et al., 2006). In addition to manmade production, some organohalide like chlorinated methanes are also produced naturally by seaweed and termite as well as abiotically produced by disinfection of drinking water (Grostern et al., 2010; Harper,

2000; Laturnus et al., 2002). In addition to the neurotoxicity and carcinogenic activity of chlorinated hydrocarbons, CF and especially 1,1,1-TCA are frequent co-contaminants with TCE because of their use as industrial solvents. CF and 1,1,1-TCA are strong inhibitors of many microbial processes essential for biogeochemical cycles, such as methanogenesis, acetogenesis, or

5 reductive dechlorination of chlorinated ethenes (Bagley et al., 2000; Duhamel et al., 2002).

Furthermore, CF and 1,1,1-TCA are also ozone-depleting substances (Montreal Protocol, 1996).

Introducing dechlorinating microbes, such as Dehalococcoides and Dehalobacter spp., into contaminated groundwater can be an effective remediation approach. For example, KB-1® , a chlorinated ethene-degrading consortium, has been used to inoculate over 700 sites worldwide and achieve successful remediations at many contaminated sites (http://www.siremlab.com/kb-1-kb-

1-plus/). However, chlorinated ethene remediation by OHRB can be limited by the co-existence of 1,1,1-TCA and CF, which can result from (i) the cross inhibition of reductive dechlorination of

Dehalococcoides (chlorinated ethene OHRB) and Dehalobacter (chlorinated alkane OHRB)

(Grostern and Edwards, 2006) and (ii) the inhibition of acetogens and methanogens for nutrient production (carbon sources, electron donors, and essential cofactors) (Duhamel et al., 2002;

Loffler and Edwards, 2006). Due to the issues described above, bioremediation of TCE and 1,1,1-

TCA co-contamination often halts at VC and 1,1-DCA. Unfortunately, 1,1,1-TCA and TCE are found to coexist at over 310 sites in US EPA NPL sites (a search of the NPL database in May,

2006). Therefore, the bioremediation of 1,1,1-TCA and CF in groundwater is of great importance.

In the Edwards lab, the addition of a 1,1,1-TCA- and CF-degrading culture (ACT-3) to KB-1® was found to enhance the dechlorination of both TCE and 1,1,1-TCA (Grostern and Edwards, 2006).

Currently, the combined culture, called KB-1® Plus, has been sold commercially for use at co- contaminated sites.

Interspecies nutrient exchanges in dechlorinating mixed cultures

In a more complicated system like Dehaloccocoides-containing dechlorinating microbial community KB-1, in which external organohalide electron acceptors are present, homoacetogens first ferment methanol to acetate and H2 via acetogenesis (Taga and Walker, 2010). Next, the

6 produced H2 serves as the electron donor for TCE dechlorination to ethene by dechlorinating D. mccartyi and a minor portion of produced acetate serves as the carbon source for D. mccartyi, while the remaining portion serves as the electron donor for methanogens to be fermented to methane via methanogenesis. In addition to the exchanges of electron donors and carbon sources, homoacetogens and methanogens are also known as the producers of cobamides, which are essential cofactors involved in organohalide respiration (Yan et al., 2013; Yan et al., 2012).

Interestingly, many obligate OHRB like Dehalococcoides, Dehalogenimonas, and many

Dehalobacter strains cannot synthesize the required cobamide cofactors for respiration, but rely on the supply from other community members, shaping an exclusive type of cross-feeding interactions in dechlorinating mixed culture (See Section 1.2 for additional details).

Background on the microbes and mixed dechlorinating cultures studied in this thesis

Fig. 1.2. The transfer history of cultures used in the research presented in this thesis. Abbreviations: CF, chloroform; ED, electron donor; EA, electron acceptor; LAC, lactate; EtOH, ethanol; 1,1,1-TCA, 1,1,1- trichloroethane; 1,1-DCA, 1,1-dichloroethane; TCE, trichloroethene; MeOH; methanol; Dhb, Dehalobacter restrictus. Dehalococcoides is the dominant OHRB in the KB-1 mixed culture.

Two mixed cultures, along with their sub-transfers, and 6 pure isolate cultures are relevant to this thesis. Their origins and features are described below and illustrated in Fig. 1.2.

Mixed culture ACT-3 is a 1,1,1-TCA-dechlorinating mixed culture enriched from a contaminated site in northeastern US (Tang and Edwards, 2013). It can dechlorinate several chlorinated alkanes, and contains two major OHRB, Dehalobacter restrictus strains DCA and CF.

The culture is maintained with lactate, methanol, and ethanol as the electron donor and carbon source, while CF, 1,1-DCA, or 1,1,1-TCA is used as the final electron acceptor. A Bacteroides sp. and a Clostridium sp. are the major non-OHRB in the ACT-3 mixed culture (Tang et al., 2012).

Based on the literature and genome annotation (Chen and Wolin, 1981; Macy et al., 1978; Miller,

1978; Tang and Edwards, 2013), the Bacteroides sp. likely ferments lactate, methanol and ethanol to acetate and malate (or fumarate). The Clostridium sp. then further ferments malate (or fumarate) to propionate (Berg et al., 2015; Puentes Jácome and Edwards, 2017). Finally, the methanogens in

ACT-3 convert acetate to methane. During the fermentation, H2 is also produced, and is consumed by Dehalobacter for reductive dechlorination and by methanogens for methane production by reduction of CO2. Additional details on the ACT-3 culture are available in the thesis of a former

Edwards Lab PhD graduate, Dr. Shuiquan Tang (Tang, 2014).

Bacteroidales sp. CF is a Gram-negative fermenter and is the most abundant non-OHRB in the dechlorinating mixed culture ACT-3 that contains >50 mg/L CF and 1,1,1-TCA. Dr.

Shuiquan Tang previously assembled and annotated its whole genome from the metagenomic sequencing data (Tang and Edwards, 2013). Although the microbe has yet to be isolated, it is predicted to ferment L-lactate as well as ethanol, and donate essential nutrients like malate to

Dehalobacter restrictus strain CF (see Chapter 4).

Dehalobacter restrictus strains, a Gram-negative Firmicutes, is the first isolated strain in the type species Dehalobacter restrictus (Holliger et al., 1998). The name “restrictus” reveals its restricted metabolism and fastidious growth conditions (Rupakula et al., 2013; Rupakula et al.,

2015). Strain PER-K23 can dechlorinated PCE via TCE to cDCE using H2 as an electron donor.

Its growth on defined medium requires addition of arginine, histidine, and threonine, along with acetate as the major carbon source as well as cobamides and thiamine as cofactors. Strains CF and

DCA were isolated using undefined medium containing ACT-3 supernatant by Dr. Shuiquan Tang, and their whole genomes have been assembled from metagenomics data (Tang et al., 2012). Strain

CF is capable of dechlorinating CF, 1,1,1-TCA, and 1,1,2-TCA to DCM and 1,1-DCA, and 1,2-

DCA, respectively; strain DCA is capable of dechlorinating 1,1-DCA and 1,1,2-TCA to CA, and

1,2-DCA, respectively.

Mixed culture KB-1 is a mixed culture enriched from soil and groundwater samples of a

TCE-contaminated site in Southern Ontario, Canada. It can dechlorinate several chlorinated ethenes and chlorinated aromatics, and contains many species of OHRB including

Dehalococcoides mccartyi, Dehalobacter restrictus and Geobacter lovleyi (Puentes Jácome and

Edwards, 2017). The culture is maintained with methanol as the sole electron donor and carbon source, while TCE or trichlorobenzene is used as the final electron acceptor. Acetobacterium carbinolicum strain KB-1, Sporomusa silvacetica strain KB-1, and a Methanosarcina sp. are the major non-dechlorinating microbes in KB-1. Based on the literature and genome annotation,

Acetobacterium carbinolicum strain KB-1 and Sporomusa silvacetica strain KB-1 likely ferment methanol to acetate and to H2. Acetate is then fermented by the Methanosarcina sp. to methane and CO2 or oxidized by Geobacter lovleyi to CO2. The H2 is consumed by Dehalococcoides and

Dehalobacter for reductive dechlorination.

Acetobacterium carbinolicum strain KB-1 is a Gram-positive homoacetogen in the dechlorinating mixed culture KB-1. Its genome has been sequenced and partially assembled by former Edwards Lab postdoctoral researcher, Dr. Kirill Krivushin. Based on the genome annotation, it possesses a complete anaerobic cobalamin biosynthesis pathway, and is likely the main cobalamin producer in the KB-1 (See Appendix B).

Geobacter lovleyi strain KB-1 is a Gram-negative δ-Proteobactrium enriched from KB-1 and was collaboratively isolated by current Edwards Lab PhD student, Luz Adriana Puentes, and me. The draft genome and plasmid of the bacterium have been sequenced and assembled by Dr.

Laura Hug and PhD student Olivia Molenda. Briefly, this microbe can dechlorinate PCE to cDCE using acetate as the sole electron donor and carbon source. Alternatively, fumarate can also serve as the electron acceptor.

Sporomusa silvacetica strain KB-1 is a Gram-negative homoacetogen in KB-1 and has been isolated by a former Edwards lab PhD graduate, Dr. Melanie Duhamel. Its draft genome has been partially assembled by Dr. Kirill Krivushin. It can grow on methanol as the sole carbon source and electron donor without the supplement of all vitamins or yeast extract. Previous research has shown that the genus of organisms natively produces [p-cresol]cobamides and phenyl-cobamides.

Nevertheless, it is able to synthesize the cobamides with any lower ligand (>10 µM) amended in medium when betaine is used as the substrate (Mok and Taga, 2013).

Desulfitobacterium hafniense is a Gram-positive Firmicutes obtained from the Löffler group at the University of Tennessee Knoxville, USA. Desulfitobacterium is a close relative of

Dehalobacter within the Peptococaceae, but its metabolism is very versatile, as opposed to the restricted metabolism of Dehalobacter. It can use a variety of electron donors (including H2,

10 carbohydrates, alcohols, and organic acids) and acceptors (including organohalides, nitrate, sulfate, and ferric iron).

1.1.2 Corrinoids

Nomenclature, structure diversity, and function

Corrinoids, such as cobalamin (or vitamin B12), are the essential coenzyme for enzymes involved in biosynthesis of amino acids, DNA, and fatty acids as well as in biogeochemical cycles of carbon, halogen, and heavy metals within all the three domains of life (Mori et al., 2016; Taga and Walker, 2008). Being the largest known non-polymeric natural product (m/z~1300~1500), the structure of a cobamide, the mature and catalytically active form of corrinoid, includes a main tetrapyrrole ring (or cobryic acid), a ribose, a central chelated cobalt ion, two axial ligands (upper and lower ligands), and an aliphatic side chain containing a phosphodiester bond bridging the tetrapyrrole ring and the α-ribazole (the ribonucleoside with lower axial ligand). To date, seventeen cobamides (Fig. 1.3), which are differentiated based on the structure of their lower ligands, have been identified and characterized (Wintermute and Silver, 2010). For example, the lower axial ligand of cobalamin is 5,6-dimethylbenzimidazole (DMB). In a cell, the upper axial ligands of cobamides can be a hydroxyl group, an adenosine, a methyl group, or a histidine residue of an enzyme. In industrial/pharmaceutical production, cobalamin is purified in its cyano-form (cyanide upper ligand) because it is more stable. Recently, nor-vitamin B12, which lacks a methyl group on carbon 176 of other cobamides, has also been identified (Taga and Walker, 2008).

According to literature, each species of microbes only produces a very limited range of cobamides with highly similar structure, and most of time, only one type is produced by a single species. For example, members of the family Veillonellaceae, such as Sporomusa, produce phenyl-

11 cobamide or a [p-cresol]cobamide (Men et al., 2014; Yan et al., 2013); members of the family

Eubacteriaceae, such as Acetobacterium woodii and Eubacterium limosum, produce cobalamin

(Renz et al., 1993; Stupperich et al., 1988). Interestingly, to date, only members in the family

Eubacteriaceae were reported to produce cobalamin under anaerobic conditions. However, the cobamides produced from the same genera of microbes can vary. For example, Clostridium thermoaceticum, Clostridium cochlearium, and Clostridium formicaceticum produce the cobamides with a 5-methoxyl-benzimidazole (factor IIIm), an adenine (pseudo-B12), or a 5- methoxy-6-methylbenzimidazole lower ligand ([5MO6MBZ]cobamide), respectively (Santos et al., 2007; Stupperich et al., 1988; Wurm et al., 1975); some sulfate- and sulfur-metabolizing gram- positive bacteria produce the corrinoid with a 5-methyl-benzimidazole ligand ([5MBZ]cobamide); methanogenic archaea produce cobamides with a 5-hydroxy-benzimidazole (factor III; Fig. 1.3)

(Stupperich et al., 1988).

Compared to Gram-positive Firmicutes which produce diverse benzimidazole lower ligands, Proteobacteria, which are Gram-negative bacteria, seem to harbor a variety of purines as lower ligand. For example, Desulfovibrio vulgaris produces the cobamide with a guanine or a hypoxanthine as the lower ligand (Guimarães et al., 1994); Sulfurospirillum multivorans and

Sulfurospirillum multivorans produces norpseudo-vitamin B12 (Seth and Taga, 2014); Salmonella typhimurium produces the cobamides with an adenine or a 2-methyladenine (2MA) ligand (Keck and Renz, 2000). However, some Proteobacteria also produce benzimidazole ligands such as δ-

Proteobacterium Geobacter lovelyi, which produces factor IIIm (Chapter 2). Also, Firmicutes member Desulfitobacterium hafniense strain Y51 produces the cobamide with an unsubstituted purine as the lower ligand (purinyl-cobamide; Chapter 2). Therefore, the type of cobamides produced by microbes cannot be classified based on phylogenetic origin alone.

Fig. 1.3. The structure of corrinoids and their lower ligands. Abbreviations: Cbl, cobalamin; Cbi, cobinamide; Cby, cobyrinic acid DMB, 5,6-dimethylbenzimidazole; 5HBZ, 5-hydroxybenzimidazole; 5MBZ, 5-methylbenzimidazole; 5MOBZ, 5-methoxy-benzimidazole; 5MO6MBZ, 5-methoxy-6- methylbenzimidazole; 5H6MBZ, 5-hydroxy-6-methylbenzimidazole; 2-MA, 2-methyladenine.

Cobamides are the essential prosthetic group (i.e., the tightly-bound coenzyme in enzymes) for numerous methyltransferases, isomerases, and RDases (Mori et al., 2016). It was reported that

Dehalococcoides mccartyi dechlorinates TCE two times faster with the supplement of sufficient cobalamin (25 µg/L) in growth medium compared to under insufficient conditions (1 µg/L) (Yan et al., 2013). For functional catalysis, different types of enzymes require a specific form of cobamide. For example, it is reported that only cobalamin can efficiently support the VC dechlorination by Dehalococcoides mccartyi (Yan et al., 2015), while the use of other forms of cobamides decrease rates of VC dechlorination, suggesting that DMB is essential for the function of VC-degrading RDases BvcA, VcrA, and TceA in D. mccartyi. Since in situ corrinoid addition is cost prohibitive, amending proper cobamide producers to dechlorinating mixed cultures may accelerate in situ bioremediation of chlorinated solvents. The coordination of the lower ligand with

13 the central cobalt ion is critical to the function of many corrinoid-dependent enzymes. For example, phenyl-cobamides produced by Sporomusa spp. are required for the enzymes participating in methanol and 3,4-dimethyl-benzoate metabolism, while the metabolism of fructose and betaine are not (Mok and Taga, 2013). The phenyl lower ligands cannot form a covalent bond with the central cobalt ion, which is always in “base-off” form, while the secondary amine group on nucleotide types of lower ligands is able to bind to the central cobalt ion, alternating between

“base-on” and “base-off” form. It is also reported that the presence of a methyl group on carbon

176 of the corrin ring facilitates the coordination of the lower ligand with the central cobalt ion on corrinoids (Seth and Taga, 2014). However, the PceA from Sulfurospirillum multivorans harbors a norpseudo-B12 as cofactor (adenine as lower ligand), which is critical to its dechlorination activity, and the amendment of excess amount of DMB in medium causes the production of non- functional PceA (Keller et al., 2014). In addition to the cobalt-ligand coordination, the methyl group on the benzimidazolyl ligand also plays an important role in enzyme’s activity. For example, the dechlorination of VC by Dehalococcoides species relies on the cobamides with a 5-methylated benzimidazolyl ligand (Yan et al., 2015).

Anaerobic biosynthesis of cobamides and lower benzimidazole ligand

Being the most complex coenzyme known, the anaerobic biosynthesis of corrinoids requires a great number of enzymes and associated genes (~30 genes). Moreover, while the family of cobamide cofactors is essential for nearly all living organisms, their biosynthesis is limited to a subset of prokaryotes like methanogens, acetogens, and plant symbionts (Escalante-Semerena,

2007). Briefly, the anaerobic biosynthesis of corrinoid starts from the transformation of 8 glutamates to 5-aminolevulinates, followed by the formation of a tetrapyrrole structure, which is then ligated to a precorrin-2 after several deamination reactions and a ligation step. The cobalt ion

14 is then chelated to precorrin-2, followed by six amidations and two methylations to form a cobyrinic acid. Finally, an amide bond is formed between cobyrinic acid and an aminopropanol converted from threonine to form a cobinamide (Krasna et al., 1957).

The mechanism of the lower ligand incorporation to cobinamide is performed by the phosphoribosyl transferase CobT, which catalyzes the formation of an α-ribazole by replacing the nicotinamide on nicotinamide mononucleotide with the lower ligand base (Hazra et al., 2013). The

α-ribazole is then linked to the cobinamide to form a complete cobamide via a GTP-dependent phosphorylation, followed by a ligation. Recent studies have shown that the substrate specificity of CobT varies in different bacteria, which depends on the sequence and identity of certain functional residues (Crofts et al., 2013). Moreover, one bacterium can encode multiple types of

CobT homologs with varying substrate specificity. Interestingly, from a test of CobT substrate specificity, 6 out of 7 CobT homologs encoded by microbes from different phyla show a common preference to DMB, which reveals the widespread preference of DMB as the lower axial ligand of corrinoid in the biosphere (Degnan et al., 2014). However, the lower ligand availability in the environment will also affect the type of cobamides taken up by corrinoid-auxotrophic microbes

(Sonnenburg and Sonnenburg, 2014).

The aerobic biosynthetic pathway of the lower ligand, DMB, has been studied in detail.

The base is synthesized from a cannibalization of FMN (McCormick, 1989; Taga et al., 2007).

The reaction requires molecular oxygen and thus cannot operate under anaerobic conditions. As mentioned above, only two isolated anaerobes (Acetobacterium woodii and Eubacterium limosum) have been reported to produce DMB (Renz et al., 1993; Stupperich et al., 1988). Previous research also reported that the building blocks of DMB in Eubacterium limosum are glutamine, glycine, formate, and erythrose, which are also used in purine and tryptophan biosynthesis (Munder et al.,

1992). The two methyl groups of DMB originate from S-adenosyl methionine and are responsible for the sequential methylations from 5HBZ via 5MOBZ and then 5H6MBZ to DMB (Fig. 1.3)

(Schulze et al., 1998).

Fig. 1.4. Anaerobic DMB biosynthesis pathway and corresponding biosynthetic genes in Eubacterium limosum. Adapted from Hazra et al., 2015.

Recently, the genes and intermediates of the anaerobic biosynthesis pathway were characterized (Fig. 1.4) (Hazra et al., 2015; Mehta et al., 2015). The anaerobic biosynthesis of

5HBZ shares the same substrate aminoimidazole ribotide (AIR) used in thiamine biosynthesis. The radical SAM enzyme BzaAB catalyzes the conversion of AIR to 5HBZ. An O-methyltransferase

BzaC then methylates 5HBZ to 5MOBZ, followed by two successive methylations by two radical

SAM methyltransferases BzaDE to form a DMB. The five biosynthetic genes encoding the enzymes have been used to probe the anaerobic benzimidazole producers. The data suggest that bzaE is mostly identified in Eubacteriaceae (Acetobacterium and Eubacterium). The data also predicted that Geobacter lovleyi synthesizes Factor IIIm due to the absence of bzaDE genes in genome.

Interspecies cobamide transfer

Dehalococcoides, Dehalogenimonas, and Dehalobacter spp. are ubiquitous and dominant

OHRB in contaminated groundwater. Their dechlorination relies on the RDases which utilize cobamides as the cofactor (Maillard et al., 2003). However, to date, only few strains of

Dehalobacter spp. have been reported to grow without the supplement of cobalamin, and all isolated Dehalococcoides strains are cobalamin-auxotrophic (Sun et al., 2002). Thus,

Dehalococcoides rely on the corrinoid supply from other community members (Yan et al., 2012).

Nevertheless, Dehalococcoides mccartyi can remodel other cobamides into cobalamin in the presence of DMB (Yi et al., 2012). While Dehalococcoides genomes show an incomplete synthesis pathway for cobalamin, it possesses the btuBCD-F genes to uptake corrinoids at femtomolar concentrations and to replace the undesirable lower ligands into DMB (Kube et al., 2005). The uptake mechanism for corrinoids via transporter BtuBCD-F system has been studied in some details in Gram-negative bacteria (Cardona et al., 2016; McInerney et al., 2009). BtuB is a Ca- dependent transporter located in outer membrane, which transports corrinoids into periplasm, and delivers it to the corrinoid-binding protein BtuF. The transport requires energy input and is provided through interaction with an inner membrane protein TonB. Once the corrinoid binds to

BtuF, it will transport the cofactor to an ABC transporter BtuCD, and translocate it into cytoplasm through an ATP-dependent mechanism. However, the uptake mechanism for the various lower ligands remains unknown (Bradbeer et al., 1986; Escalante-Semerena, 2007; Heller et al., 1985).

Dehalococcoides often exist in contaminated groundwater with other OHRB and non- dechlorinating organisms, mostly homoacetogens and methanogens (Duhamel and Edwards,

2006), which results from the syntrophic relationship in which the coupling of acetogenesis to methanogenesis occurs. Recent studies have shown that this two groups of microbes also donate

17 cobamides to fulfill the nutrient requirements of Dehalococcoides. For example, Löffler group reported that Sporomusa silvacetica strain KB-1 and Methanosarcina barkeri are able to provide the [p-cresol]cobamide and [5HBZ]cobamide to Dehalococcoides mccartyi, respectively (Yan et al., 2013). Pelosinus, a relative of Sporomusa in family Veillonellaceae, was also reported to provide the [p-cresol]cobamide to Dehalococcoides in the enrichments from some TCE- contaminated field sites (Men et al., 2014). However, these community members cannot fully satisfy the cofactor requirement of Dehalococcoides. Recently, Men et al. (2014) discovered the accumulation of free DMB in the supernatant of a lactate-fed TCE-dechlorinating consortium, while [p-cresol]cobamide is the dominant corrinoid in supernatant. It is likely that

Dehalococcoides synthesize cobalamin by remodeling undesirable cobamides with DMB. On the other hand, our current data suggest that Acetobacterium carbinolicum strain KB-1 seems to provide cobalamin to Dehalococcoides in the KB-1 without cobalamin supplementation

(Appendix B).

1.1.3 NADPH: regeneration and shuttle systems

Nicotinamide adenine dinucleotide-2’-phosphate (abbreviated NADP+) is an essential cytosolic cofactor required by all organisms in the biosphere (Morris et al., 2013). It is an electron carrier which shuttles the reducing power to the biosynthesis of amino acids, lipids, and nucleic acids (Wakil et al., 1983). Moreover, it is required for reactive oxygen species (ROS) generation

(Price et al., 2004) and for anti-oxidative mechanisms for most organisms (Famili et al., 2003). In cells, NADP+ undergoes reduction and re-oxidation by NADP-dependent dehydrogenase and

+ reductases, respectively. The reducing power (NADP /NADPH E′0: −320 mV) is temporally conserved in a high-energy hydride bond in the nicotinamide ring (abbreviated NADPH; Fig. 1.5).

+ The λmax for NADP (260 nm) and NADPH (340 nm) are distinct, enabling facile measurements of NADP+-dependent reactions. Nicotinamide adenine dinucleotide (abbreviated NAD+) is another essential cofactor in biological systems which lacks the 2’-phosphate on NADP+. Interestingly, such a subtle difference in structure separates their distinct biological functions. In biological systems, NADH is primarily produced during the catabolism, which is then re-oxidized to NAD+, along with the export of proton by Electron Transport Chain Complex I, or NADH:ubiquinone oxidoreductase.

Fig. 1.5. Reversible interconversion of NADPH and NADP+. The high energy hydride bond on the nicotinamide ring provides reducing power for various biosynthetic reactions. NADPH and NADH are different in that the former contains a 2’-phosphate group. Abbreviations: H-, hydride.

to generate the proton motive force for ATP biosynthesis (O’Brien et al., 2015). By contrast,

NADPH is produced via numerous pathways, and are primarily employed in anabolic reactions.

This is reflected by the ratio value for NADP+/NADPH (0.017-0.95) and NAD+/NADH (3.74-31.3) in cells, suggesting that the former pair is maintained in a reduced state while the latter is maintained in an oxidized state under physiological conditions (Hanemaaijer et al., 2015; Reed et al., 2003). Furthermore, intracellular concentration of NAD+ is significantly higher than that of

NADPH (~9) in E. coli (Hanemaaijer et al., 2015). Therefore, although many dehydrogenases are

19 promiscuous to both NAD+ and NADP+, NAD+ is mostly served as the substrate under physiological conditions.

There are several sources for intracellular NADPH pool (Morris et al., 2013; Roling and van Bodegom, 2014). In heterotrophic microorganisms, NADPH is mostly generated via dehydrogenase reactions in the oxidative pentose pathway and glycolysis; 5,10- methylenetetrahydrofolate dehydrogenase reaction in the folate cycle; isocitrate dehydrogenase reaction in the TCA cycle; the malic enzyme-mediated decarboxylation in the pyruvate-malate shuttle (a.k.a. anaplerotic node or malate shunt), which bridges glycolysis/gluconeogenesis and the

TCA cycle (Manor et al., 2014; Tan et al., 2015). In addition to the reactions involved in the central carbon metabolism, several NADPH-generating enzymes, such as ferredoxin:NADP+ oxidoreductase (Shoaie et al., 2015) and cytosolic NADP+-reducing hydrogenase (Dodd et al.,

2017), are functioning in many autotrophic microorganisms.

A shuttle system also exists in biology for the interconversion of NADH and NADPH, which is accomplished by two families of transhydrogenases, energy-independent soluble transhydrogenase and energy-dependent membrane-bound transhydrogenase (Adrian and Loeffler,

2016). According to generally accepted view, the physiological role for soluble transhydrogenase, mainly found in γ-Proteobacteria, is converting NADPH to NADH, which is supported by the ratio of NAD+/NADH and NADP+/NADPH mentioned above along with thermodynamics of view

(Voordouw et al., 1983). The excessive amount of NADPH produced during heterotrophic growth can be converted to NADH for energy conservation. By contrast, membrane-bound transhydrogenase, mainly found in Eukaryotes and some bacteria, physiologically converts NADH to NADPH (Zhuang et al., 2014) since the direction of reaction under physiological conditions requires energy input. Moreover, the gene encoding membrane-bound transhydrogenase is

20 significantly upregulated in E. coli during the growth on substrates whose metabolism is not directly couple to NADPH generation (Zhuang et al., 2014). As estimated 40% of the NADPH in

E. coli is generated from membrane-bound transhydrogenase, which also serves as the major

NADPH source for amino acid biosynthesis. Together, since NADPH is directly linked to anabolism of an organism, it is reasonable that fast-growing microbes like E. coli possess multiple types of NADPH-generating and NADH-NADPH shuttle systems, which provide abundant supply of reducing power for anabolism and provide flexibility for redox balancing. On the other hand, the growth of organisms with limited NADPH-generating systems may heavily rely on specific

NADPH-generating substrates or direct nutrient supplies from other organisms.

1.1.4 Prenylated flavins and reversible decarboxylation of aromatics

Bio-prenylation is a ubiquitous reaction in the biosynthesis of small molecules as well as in the protein- and membrane-protein interactions toward the three kingdoms of life (Chen et al.,

2017). The prenyl (3-methyl-but-2-en-1-yl) moieties, such as dimethylallyl-pyrophosphate

(DMAPP; C5), geranyl-pyrophosphate (C10), and farnesyl-pyrophosphate (C15), are synthesized through the well-known mevalonate pathway in most Archaea (Meganathan, 2001) and Eukarya

(Lichtenthaler et al., 1997) and through the non-mevalonate pathway in most bacteria (Lange and

Croteau, 1999), which are mainly responsible for the production of (i) hopanoid, the modulator for bacterial membrane permeability and fluidity (Rohmer, 2008), (ii) coenzyme-Q, the component in the ETC Complex I and II (CoQ10) for biological energy conservation (Olson and Rudney, 1983) as well as the modulator for membrane permeability (CoQ8) (Sévin and Sauer, 2014), and (iii) heme A, the component in the ETC Complex IV (Fig. 1.6) (Buhaescu and Izzedine, 2007).

Fig. 1.6. Coenzyme Q8 (ubiquinone) biosynthesis by MEP isoprenoid pathway and UbiX-UbiD pair for Electron Transport Chain-mediated energy conservation. Abbreviations: The biosynthetic enzymes are shown in red. Abbreviations: ATPase, ATP synthase; Q or CoQ8, ubiquinone; cytC, cytochrome C; DMAPP, dimethylallyl-pyrophosphate; FPP, farnesyl-pyrophosphate; GPP, geranyl-pyrophosphate; 4HB, 4-hydroxybenzoate; 2OPPh, 2-octaprenyl-phenol; 3OP-PP, 3-octaprenyl-pyrophosphate; 3OP4HB, 3- octaprenyl-4-hydroxybenzoate; Pyr, pyruvate; G3P, glyceraldehyde-3-phosphate; UbiD, 3-octaprenyl-4- hydroxybenzoate decarboxylase; UbiX, flavin prenyltransferase. Components for electron transport chain: I, NADH dehydrogenase; II, succinate dehydrogenase; III, cytochrome C reductase; IV, cytochrome C oxidase.

Over the past two decades, considerable research efforts have been devoted to study and to metabolically engineer the terpenoid pathway in E. coli (Martin et al., 2003; Zheng et al., 2013),

S. cerevisiae (Shiba et al., 2007), and some plant organisms (Roberts, 2007) for the production of biofuels, polymer-precursors, and terpenoid pharmaceuticals. Prenylated flavonoids are widely distributed in the plant kingdom as both primary and secondary metabolites (Botta et al., 2005).

Recently, prenylated FMN (prFMN) was identified as a cofactor for the aromatic carboxy-lyase

UbiD family, has been identified (Ebenau-Jehle et al., 2017; Lan and Chen, 2016; White et al.,

2015). This cofactor is now known to be involved in (i) the biological decomposition of lignin- derived aromatic monomers (p-coumarate; protocatechuate; ferulate) (Lin et al., 2015; Wilbon et al., 2013), (ii) the biodegradation of aromatic pollutants including phthalate (Ebenau-Jehle et al.,

2017), phenol (Schühle and Fuchs, 2004), and potentially benzene (Abu Laban et al., 2010) and naphthalene (Bergmann et al., 2010), and (iii) the biosynthesis of coenzyme-Q (Cox et al., 1969;

Olson and Rudney, 1983; Payne et al., 2015). Unexpectedly, flavin prenyltransferase UbiX employs a dimethylallyl-monophosphate (DMAP), as opposed to the DMAPP employed by other flavonoid prenyltranferases (Yamamoto et al., 2000), and a fully reduced FMN as the substrates to form the fourth non-aromatic ring on FMN (White et al., 2015). The UbiX family, though sharing similar function with flavin prenyltransferases, are separated to the independent group and shares much more similarity with the decarboxylase UbiD family (White et al., 2015).The EPR analyses and the crystal structure of the air-oxidized purple-colored holo-UbiX revealed a stable prFMN radical form embedded and the prFMN:UbiX complex revealed an absorbance maximum at 550 nm (White et al., 2015). Interestingly, the same study also contended that holo-UbiX serves as the chaperone for the transfer and incorporation of prFMN to UbiD, while an earlier study has shown that the addition of extracted prFMN enabled the Fdc1-mediated decarboxylation of cinnamate and ferulate (Lin et al., 2015; Payne et al., 2015).

1.2 Rationale and Research Objectives

The underlying hypothesis of this thesis is that cofactor cross-feedings underpin community function, and are the key to manipulate the microbiomes for a better performance in agriculture, bioremediation, human health and industrial biotechnology. As described in

Literature Review (Section 1.1), cofactors play an essential role in the functionality and population composition of a microbial community. Therefore, the research presented within this 23 thesis aims to examine the cross-feedings interactions of essential cofactors in anaerobic groundwater microbial communities. Taking advantages of the available mixed cultures in the

Edwards Lab, I aimed to study the metabolic interdependency in organohalide-respiring microbial communities and anaerobic aromatic degrading microbial communities. Using available isolate model organisms, I also managed to elucidate the biosynthesis and functions of the essential cofactors for organohalide respiration (cobamides) and for anaerobic aromatic biotransformation

(prenylated FMN). The work was conducted in BioZone, an interdisciplinary research environment, where I was based in Prof. Elizabeth Edwards’ environmental engineering and microbiology research group, while cooperating with Prof. Alexander Yakunin’s enzymology research group and Prof. Radhakrishnan Mahadevan’s computational biology research group.

With their help, as well as the help of collaborators at the University of Tennessee, I was able to investigate research objectives from different scientific perspectives. The major objectives of this thesis were as follows:

Objective 1. Develop methods to identify and to measure the functional corrinoid cofactors in the corrinoid-dependent reductive dehalogenases using a renovated method combining blue native polyacrylamide gel electrophoresis (BN-PAGE) and liquid chromatography-Mass spectroscopy

(LC-MS).

Objective 2. Experimentally verify the metabolic annotation of Dehalobacter restrictus, the key

OHRB in the ACT-3, using the annotated genome and comparative genomic data from former

Edwards Lab PhD student, Dr. Shuiquan Tang.

Objective 3. Identify essential nutrients exchanges between fermenting and dechlorinating organisms that enable the growth of Dehalobacter using constraint-based metabolic models of the microbial community in conjunction with experimental verification.

Objective 4. Elucidate the biosynthetic origin of prenylated FMN, an essential cofactor for ubiquinone biosynthesis, biological decomposition of lignin monomers, and biodegradation of aromatics under nitrate-reducing conditions.

1.3 Thesis Outline and Structure

Chapter 1: Literature Review and General Introduction (this section also includes research objectives and overall thesis structure.)

Chapter 2: Identification of Functional Cobamide Prosthetic Group in Reductive

Dehalogenase Using BN-PAGE and LC-MS

This chapter addresses Objective 1 and reports the development of a renovated BN-PAGE method that allows rapid characterization of the functional corrinoid cofactors in active RDases from dechlorinating consortia with relatively lower biomass, and the application of the renovated method to characterize the native prosthetic group of Desulfitobacterium PceA and Geobacter

PceA, which are purinyl-cobamide and factor IIIm, respectively. The data present in this chapter has been published in Nature Chemical Biology (See Section 1.4) where I am a co-author.

Chapter 3. Refined Experimental Annotation Reveals Conserved Corrinoid Autotrophy in

Chloroform-Respiring Dehalobacter Isolates

This chapter addresses Objective 2 and reports the experimental verification of the genome annotation of Dehalobacter restrictus strain CF, a microorganism specialized in respiring chlorinated alkanes, especially focused on the annotation in central metabolism and in the biosynthesis of amino acids/cofactors. From multiple lines of evidence, this chapter proposed that chloroform, a strong inhibitor for corrinoid-producing acetogens and methanogens in organohalide-respiring communities, has indirectly facilitated the conservation of corrinoid autotrophy in the chloroform-respiring strains among Dehalobacter restrictus species. The content in this chapter has been published in Frontiers in Microbiology and the ISME Journal (See

Section 1.4).

Chapter 4: Interspecies Malate-Pyruvate Shuttle Drives Amino Acid Exchange in

Organohalide-Respiring Microbial Communities

This Chapter addresses Objective 3 and reports a joint research project that integrates the data from computational biology and experimental biology. An unsolved question in Chapter 3 is the discrepancy that Dehalobacter possesses the genes to synthesize all the amino acids, but it still requires amino acid supplementation for growth. Using the experimentally verified metabolic annotation of Dehalobacter restrictus and available proteomic data, Ms Cleo Ho and Mr. Kevin

Correia in the Mahadevan group developed a constraint-based metabolic model for Dehalobacter.

I then performed several experiments to validate the uncertain constraints, including the confirmation of the gap in classical serine biosynthesis pathway and the defects in NADPH regeneration. The experimentally curated model then accurately predicted malate as the essential carbon source to fuel NADPH pool in Dehalobacter cells. Finally, the time-course malate consumption along with pyruvate production in Dehalobacter restrictus isolate cultures suggests a malate-mediated interspecies malate-pyruvate shuttle in Dehalobacter-enriched consortium. A

26 version of this chapter has been written into a manuscript in preparation for submission to the

ISME Journal (See Section 1.4).

Chapter 5: Biosynthesis and Activity of prenylated FMN Cofactors

This chapter addresses Objective 5 and describes the biosynthetic origins and biochemical characterization of prenylated FMN, a newly identified cofactor involved in ubiquinone biosynthesis, biological decomposition of lignin, and anaerobic biodegradation of aromatics. The results suggest that in E. coli, DMAP, the prenyl donor, can be produced from both prenol phosphorylation and DMAPP dephosphorylation. Using the in vivo DMAP synthesis method, Dr.

Anna Khusnutdinova and I produced sufficient quantity of free prFMN species for biochemical characterization, and subsequently applied them to activated prenylated FMN-dependent decarboxylases Fdc1 in vitro. A version of this chapter has been published in Cell Chemical

Biology (See Section 1.4).

Chapter 6: Summary, Significance and Future Work

This chapter integrates and summarizes the work of this thesis and describes further research directions.

1.4 Statement of Authorship and Publication Status

In this thesis, Chapter 2 is wholly reproduced from a published paper in Nature Chemical

Biology where I am a co-author. Chapters 3 and 5 are partially reproduced from the published papers in the ISME Journal and Cell Chemical Biology, respectively. A version of Chapter 4

27 is in preparation for submission to the ISME Journal. The detailed citations and statement of authorship are provided below:

Chapter 2. Identification of functional cobamide prosthetic group in reductive dehalogenases using BN-PAGE and LC-MS

Jun Yan, Meng Bi, Allen K Bourdon, Abigail T Farmer, Po-Hsiang Wang , Olivia Molenda, Andrew T Quaile, Nannan Jiang , Yi Yang, Yongchao Yin, Burcu Simsir, Shawn R Campagna, Elizabeth A Edwards and Frank E Löffler. (2017) Purinyl-cobamide is a native prosthetic group of reductive dehalogenases. Nature Chemical Biology 14, 8-14

Contributions: In this cooperative project, I performed the tiered BN-PAGE in-gel dehalogenase assays and proteomic analysis.

Chapter 3: Refined experimental annotation reveals conserved corrinoid autotrophy in chloroform-respiring Dehalobacter isolates

This chapter regroups information from two papers. The first was helping complete a genome annotation paper largely carried out by a previous PhD student, Shiquan Tang. The second bears the name of the chapter, where I confirmed annotations experimentally.

Paper 1:

Title: Sister Dehalobacter genomes reveal specialization in organohalide respiration and recent strain differentiation likely driven by chlorinated substrates.

Authors: Shuiquan Tang1, Po-Hsiang Wang1, Steven A. Higgins2, Frank E. Löffler2 and

Elizabeth A. Edwards1.

Affiliations: 1-Department of Chemical Engineering and Applied Chemistry, University of

Toronto, Toronto, Ontario, Canada; 2-Department of Microbiology and Department of Civil and

Environmental Engineering, University of Tennessee, Knoxville, Tennessee, USA.

Contributions: EAE conceptualized the study. ST and PHW designed and performed the experiments. ST, PHW, FEL and EAE wrote the manuscript. All the authors provide conceptual and technical suggestion to the study.

Reference to publication: Frontiers in Microbiology (doi:10.3389/fmicb.2016.00100)

Paper 2:

Title: Refined experimental annotation reveals conserved corrinoid autotrophy in chloroform-respiring Dehalobacter isolates

Authors: Po Hsiang Wang1, ShuiquanTang1, Kayla Nemr1, Robert Flick1, Jun Yan2,

Radhakrishnan Mahadevan1, Alexander F. Yakunin1, Frank E. Löffler, Elizabeth A. Edwards1.

Affiliations: 1-Department of Chemical Engineering and Applied Chemistry, University of

Toronto, Toronto, Ontario, Canada; 2-Department of Microbiology and Department of Civil and

Environmental Engineering, University of Tennessee, Knoxville, Tennessee, USA.

Contributions: PHW and EAE conceptualized the study. PHW and ST designed and performed the experiments. KN heterologous enzyme expression and purification. RF performed cobamide analyses of mass spectrum data from LC-MS. PHW and EAE wrote the manuscript. All the authors provide conceptual and technical suggestion to the study.

Reference to publication: the ISME Journal, 2017, 11, 626-640 (doi:10.1038/ismej.2016.158).

Chapter 4: Interspecies malate-pyruvate shuttle drives amino acid exchange in organohalide-respiring microbial communities

Authors: Po-Hsiang Wang, Kevin Correia, Cleo Ho, Naveen Venayak, Kayla Nemr, Robert

Flick, Elizabeth A. Edwards, Radhakrishnan Mahadevan

Affiliations: Department of Chemical Engineering and Applied Chemistry, University of

Toronto, Toronto, Ontario, Canada.

Contributions: PHW, RM, and EAE conceptualized this study. PHW designed and performed the experiments. CH constructed the draft model. KC curated and finalized the model. NV proposed the interspecies transhydrogenase theory. KM performed the HPLC organic acid analysis. RF performed the LC-MS analysis. PHW, KC, EAE, and RM wrote the manuscript. All the authors provide conceptual and technical suggestion to the study.

In preparation for: the ISME Journal.

Chapter 5: Biosynthesis and Activity of Purified Prenylated FMN

Authors: Po-Hsiang Wang, Anna Khusnutdinova, Fei Luo, Johnny Xiao, Kayla Nemr, Robert

Flick, Greg Brown, Radhakrishnan Mahadevan, Elizabeth A. Edwards and Alexander F.

Yakunin.

Affiliations: Department of Chemical Engineering and Applied Chemistry, University of

Toronto, Toronto, Ontario, Canada.

Contributions: A.F.Y., E.A.E., and P.H.W. conceptualized this study. A.K. and F. L. performed the phylogenetic analysis. A.K. and P.H.W. performed enzymatic assays and cofactor characterization. F.L. and J.X. performed proteomic analysis. P.H.W., and R.F. performed LC-

MS analyses. K.N. and G.B. performed gene inactivation and site-directed mutagenesis, respectively. A.F.Y., A.K., E.A.E., and P.H.W. wrote the manuscript with contributions from all authors.

Reference to publication: Cell Chemical Biology, 2018, 25(5), 560-570 (doi:

10.1016/j.chembiol.2018.02.007.)

CHAPTER 2 – IDENTIFICATION OF FUNCTIONAL COBAMIDE PROSTHETIC

GROUP IN REDUCTIVE DEHALOGENASE USING BN-PAGE AND LC-MS

This chapter is extracted and adapted from a published paper entitled “Purinyl-cobamide is a native prosthetic group of reductive dehalogenase” in Nature Chemical Biology where I am a coauthor. In the study, I performed the BN-PAGE analysis and LC-MS analysis to validate that purinyl-cobamide is the native prosthetic group of Desulfitobacterium tetrachloroethene dehalogenase (PceA). Reproduced with permission from Nature Chemical Biology, Springer

Biology. Nov 06, 2018, DOI:10.1038/nchembio.2512. The manuscript for factor IIIm biosynthesis by Geobacter is in preparation with Dr. Jun Yan in the Löffler group, University of Tennessee

Knoxville.

2.1 Abstract

BN-PAGE is a powerful method to identify functional enzymes from both isolate and mixed cultures. This method can partially separate and enrich the functional enzymes of interests, followed by in-gel enzyme activity assays and proteomic analyses. Since

RDases are highly expressed in organohalide-respiring bacteria (OHRB) and are cobamide-dependent, the RDase-enriched gel slice should also be enriched with cobamides. This chapter reports the in-gel extraction and subsequent LC-MS identification of functional/native cobamide cofactors in the tetrachloroethene

31 dehalogenases PceA based on the established BN-PAGE method, along with tiered proteomic analysis and in-gel RDase activity assays, bringing an extended function of

BN-PAGE. The renovated BN-PAGE method was successfully applied to characterize native cobamide cofactors in the PceA from Desulfitobacterium hafniense and from

Geobacter lovleyi, which are purinyl-cobamide and factor IIIm, respectively. Thus purinyl-cobamide represents the 17th member in the naturally occurring cobamide family.

Moreover, the discovery that purinyl-cobamide serves as the functional cobamide in

Desulfitobacterium PceA assign unsubstituted purine, previously recognized as a xenobiotic, a biological function. Together, the renovated BN-PAGE method provides a facile tool to identify functional cobamide cofactors in cobamide-dependent enzymes, and likely to identify other key cofactors fulfilling the functionality in microbial communities.

2.2 Introduction

Blue native PAGE is a polyacrylamide-based electrophoresis technique enabling partial protein separation without denaturing the proteins (Schägger and von Jagow, 1991). Compared to

SDS-PAGE, BN-PAGE keeps the activity of proteins and the assembly of oligomeric protein complexes, and thus does not allow determining the mass of proteins (Wittig et al., 2006). The advantages of this method include that (i) it does not require large amounts of biomass (~ 10 mg of cell pellets) and (ii) it does not require pure culture. Adrian et al. (2007) successfully applied

BN-PAGE to identify chlorobenzene dehalogenase CbrA, along with tiered dechlorination activity assays and LC-MS-based proteomic analysis. Therefore, this method later has been widely applied to identify functional RDases in either isolate OHRB or dechlorinating mixed cultures (Tang and

Edwards, 2013; Wong et al., 2016). Moreover, BN-PAGE for protein extracts of D. mccartyi and Dehalobacter restrictus strain CF lysed by 1% (w/v) digitonin and bead-beating revealed that reductive dehalogenase activity was present in native gel slices with a molecular mass of protein at ∼ 242 kDa (Tang et al., 2013; Tang and Edwards, 2013). No RDase activity was detected at the gel slice with theoretical molecular mass range of the monomeric RdhA (~ 50 kDa), suggesting that RDase is kept in oligomeric state.

Cobamides are essential cofactor in RDases. Crystal structures of NpRdhA from

Sulfurospirillum multivorans suggested that cobamide cofactor is in the base-off configurations, indicating that the lower ligand is uncoordinated and distant from the catalytic Co center (Payne et al., 2015),suggesting that the lower ligands are not involved in the supernucleophile Co(I)- mediated C-Cl bond cleavage. Nevertheless, the lower ligand structure can significantly impact reductive dechlorination rates and growth of corrinoid-auxotrophic OHRB Dehalococcoides mccartyi (Dhc) strains (Löffler et al., 2013; Yan et al., 2016). Maximum reductive dechlorination activity and Dhc growth were observed with the addition of cobalamin or [5MBZ]cobamide to the medium, while the addition of other cobamides has resulted in hampered or complete loss of dechlorination activity (Yan et al., 2016; Yi et al., 2012). These observations indicated that the nature of the lower ligand can significantly affect the RDase activity.

Members of the genus Desulfitobacterium (Dsf) and Geobacter (Geo) are strictly anaerobic, metabolically versatile bacteria ubiquitous in subsurface environments. Some species also contribute to organohalide respiration, including the ubiquitous groundwater pollutant tetrachloroethene (PCE) (Ding et al., 2014; Sung et al., 2006). Their PCE-dechlorinating RDases

(PceA) have a strict requirement for cobamide as cofactor and dechlorinate PCE to cDCE via TCE

(Suyama et al., 2002); however, the identity of their cobamide cofactors remained yet confirmed.

Both Dsf and Geo can grow on defined medium free of cobamide; genome annotation suggested that OHRB D. hafniense and G. lovleyi possess complete genes in the cobinamide biosynthesis pathway and in lower ligand remodeling (Nonaka et al., 2006). However, the anaerobic DMB biosynthesis pathway, the lower ligand of cobalamin, is completely missing in Dsf and is incomplete in G. lovleyi. Therefore, the cobamide cofactor produced by Dsf is unknown; the cobamide cofactor produced by Geo is predicted to be factor IIIm (Hazra et al., 2015).

Since the functional RDases also contain functional cobamide cofactors, BN-PAGE would serve as a suitable method to enrich and to purify the cobamide cofactor. Also, the Dsf and Geo pure cultures cultivated on defined medium free of cobamide addition must produce their native forms of cobamides. This chapter reports the purification of cobamide cofactors from the BN-

PAGE gel slices enriched with RDase activity, and subsequent cobamide identification using LC-

MS.

2.3 Materials and Methods

Chemicals. Cobalamin (≥98%), dicyanocobinamide (Cbi, ≥93%), 5,6-dimethylbenzimidazole

(DMB) (99%), 5-methylbenzimidazole (5-MBZ) (98%), 5-methoxybenzimidazole (5MOBZ)

(97%), benzimidazole (BZ) (98%) were purchased from Sigma-Aldrich. Other cobamide standards are obtained from the Löffler lab. Concentrations of purified cobamides were determined at 361 nm with a UV-Vis spectrometer using a molar extinction coefficient of 28,060 mole-1 cm-1

(Pratt, 1972).

Cultures. Pure cultures were cultivated in 160-mL glass serum bottles containing 100 mL of bicarbonate (30 mM)-buffered, defined mineral salts medium, Wolin vitamin mix excluding cobalamin and a N2/CO2 (80/20, v/v) headspace. Dsf hafniense strains JH1 cultures were

34 supplemented with pyruvate (10 mM) as fermentable substrate, and PCE (1 mM nominal concentration) as electron acceptor. Geo lovleyi cultures were supplemented with acetate (8 mM) as the electron donor and carbon source; PCE (1 mM nominal concentration) and fumarate (10 mM) were supplemented as electron acceptors.

Corrinoid extraction. To extract corrinoids from proteins separated by BN-PAGE, gel slices were excised and transferred to individual 2-mL plastic tubes containing 1 mL of 90% methanol

(v/v) containing 50 mM acetic acid and 10 mM KCN (final pH 5.5~6) for corrinoid extraction.

The closed tubes were incubated at 60oC for 2 hours to fully dehydrate gel slices and solubilize corrinoids, before the aqueous phase was transferred to new 2-mL plastic tubes and vacuum dried using a rotary evaporator at 45oC for 2 hours. The dry pellets were suspended in 75 L ice-cold ammonium acetate (20 mM, pH 6) and centrifugation at 13,000 x g for 5 min at 4 oC removed insoluble Coomassie Blue dye and gel residuals.

Corrinoid analysis. Corrrinoids were analyzed by ultra-performance liquid chromatography coupled with high-resolution mass spectrometry (UPLC-HRMS). The corrinoids extracted from

BN-PAGE gels were analyzed using an UltiMate 3000 UPLC system in tandem with a high- resolution Exactive Plus Orbitrap mass spectrometer (Thermo Fisher, Waltham, MA, USA) and a

UV-Vis detector set to 361 nm as described (Yan et al., 2016). Corrinoids were separated on a

Hypersil Gold C18 column (Thermo Fisher, Waltham, MA, USA, 1.9 μm pore size, 2.1 mm inner diameter x 50 mm length) at a flow rate of 0.2 mL min-1 at 30 oC using 2.5 mM ammonium acetate in water as eluent A and 100% methanol as eluent B. The mobile phase was 100% eluent A and

0% eluent B for 0.46 min, followed by linear increases to 15% eluent B after 0.81 min, 50% eluent

B after 3.32 min, and 90% eluent B after 5.56 min and a 0.3-min hold before equilibration to initial column conditions. Corrinoid mass spectra were collected in positive ionization mode using an

HESI II electrospray ionization source (Thermo Fisher, Waltham, MA, USA) with a m/z scan ranging from 1,000-1,500 and a resolution of 140,000. The detection limit for this LC-MS method was 2.5 ng of corrinoid.

Blue native polyacrylamide gel electrophoresis (BN-PAGE), enzyme assays and proteomic analysis. Cells were harvested from 100 mL of PCE-grown Dsf hafniense strain JH1 cultures or

Geo lovleyi cultures, and the crude protein extracts were prepared as described (Tang et al., 2013).

BN-PAGE using pre-cast 4 to 16% gradient Bis-Tris gel (Thermo Fisher, Waltham, MA, USA) stained with Coomassie Blue was performed following the NativePAGETM Bis-Tris Gel Protocol

(Tang et al., 2013). Gel lanes were loaded with 25 L of protein standards or crude protein extracts containing approximately 6 g of total protein as estimated with the Bradford assay (Bradford,

1976). Electrophoresis used chilled buffers and a BN-PAGE chamber placed in an ice bath, and separation occurred for 60 min at 150 V and then for another 45 min at 200 V. The blank, ladder and sample lanes were separated and stained according to the fast Coomassie G-250 staining protocol (Thermo Fisher, Waltham, MA, USA). The gel slices were cut based on molecular mass ranges as visualized by the protein standards in the ladder lane. Enzyme assays to detect dechlorinating activity were conducted with Dsf hafniense strain JH1 and Geo lovleyi cell lysate

(positive control) and individual gel slices inside an anoxic chamber (Coy Laboratory, Grass Lake,

MI, USA) as described (Tang et al., 2013). The assays (1 mL) were conducted in 2-mL sealed glass vials containing 100 mM Tris-HCl buffer (pH 7.4) amended with 2 mM titanium citrate, 2 mM methyl viologen, and 2 mM TCE. Each assay mixture was incubated in an anoxic chamber for 24 h and analyzed for cDCE formation using a GC-FID method (Tang et al., 2013). In-gel digestion and MS analysis were performed at the BioZone Mass Spectrometry Facility (University of Toronto, Toronto, Canada) as described using X! Tandem for peptide/protein identification (The

GPM, thegpm.org; version X! Tandem Vengeance (2015.12.15.2)) (Tang et al., 2013). X! Tandem was set up to search for tryptic peptides of the Dsf hafniense strain Y51 genome, all characterized

RDases, other Dsf proteins in the NCBI database, as well as common contaminants such as human keratins and trypsin (total of 33,950 proteins) using a fragment ion mass tolerance of 0.40 Da and a parent ion tolerance of 2.5 Da. Deamidation of asparagine and glutamine, oxidation of methionine and tryptophan, N-terminal ammonia loss, or cyclization of glutamine or glutamamic acid to pyroglutamine or pyroglutamic acid were allowed in X! Tandem as possible peptide modifications. Further validation and refinement was performed using Scaffold software version

4.5.3 (Proteome Software Inc., Portland, OR, USA) with a reverse decoy database to establish a false discovery rate. Peptide identifications were accepted as valid at a Peptide Prophet probability of greater than 95% (Keller et al., 2002). Subsequent protein identifications were considered valid at a Protein Prophet probability of greater than 99% (Nesvizhskii et al., 2003), which were filtered to include those that had at least two unique peptide identifications.

2.4 Results

Purinyl-cobamide as the native cobamide cofactor of Dsf PceA

Mature Dsf PceA are in the mass range of 57 kDa (Suyama et al., 2002; Villemur et al.,

2006), but maximum reductive dechlorination activity is associated with bands excised from blue native polyacrylamide gel electrophoresis (BN-PAGE) gels the 242-480 kDa region, presumably due to complex formation with other proteins (Kublik et al., 2016). In our study, Dsf culture was harvested via centrifugation under strict anaerobic condition, the pellet was resuspended in 50 mM

Tris-HCl buffer (pH 7.5) containing 1% digitonin as detergent, and was lysed by bead-beating.

After removing the cell debris by centrifugation, the crude protein extracts were load on BN-PAGE

37 following the established protocol. After the non-denaturing separation for the crude protein extracts of Dsf hafniense strain JH1, the highest TCE-to-cDCE dechlorination activity was observed in gel slice 4, which revealed a clear protein band, while the other five gel slices exhibited no or negligible (<8% of cDCE production than observed with slice # 4) dechlorination activity

(Fig. 2.1A).

Fig. 2.1. Identification of purinyl-cobamide as the native cobamide cofactor in Dsf PceA following non-denaturing, gel-electrophoretic separation of Dsf crude protein extracts using BN-PAGE. (A) TCE-to-cDCE dechlorination activity of different BN-PAGE gel slices measured using dehalogenation enzyme assay. UPLC separation and mass spectrometry analysis of (B) the cobamide cofactor recovered from BN-PAGE gel slice 4 harboring the Dsf PceA and (C) purinyl-cobamide (Cba) standard (0.25 mg/L). Subsequent in-gel extraction recovered corrinoid from gel slice 4 but not from slices 3 and 5, and

LC-MS analysis revealed that the corrinoid eluted from gel slice 4 and the purinyl-cobamide standard (Fig. 2.1B,C) have matching retention times (7.5 min) and comparable m/z values

([m+H]+ = 1329.54). The proteomic analysis of gel slices 3, 4, and 5 confirmed that the Dsf PceA

(WP_011460641.1) was significantly enriched in gel slice 4, along with carbon monoxide dehydrogenase in the Wood-Ljunghahl pathway and FAD-dependent fumarate reductase in the

TCA cycle, which are both corrinoid-independent enzymes (Table 2.1) (Dobbek et al., 2001;

Iverson et al., 1999). However, 5-methyltetrahydrofolate-homocysteine methyltransferase, another

38 corrinoid-dependent protein in the Wood Ljungdahl pathway, is likely associated with the CO dehydrogenase, but is present at a low abundance below the detection limit of the proteomic analysis. However, given the fact that no other cobamide or cobamide precursors was detected in gel slice 4, purinyl-cobamide is unequivocally the native cobamide cofactor for Dsf PceA.

Table 2.1. Proteomic analysis of the Desulfitobacterium (Dsf) hafniense strain JH1 proteins associated with different BN-PAGE slices.

No. of Total Gel distinct spectral Protein description NCBI accession no. slice # peptides counts 1 - - - - 2 - - - - 5 11 Pyruvate ferredoxin oxidoreductase WP_019849101.1 3 5 18 Acetyl-CoA synthase WP_005812973.1 6 8 CO dehydrogenase WP_011459977.1 4 4 6 Tetrachloroethene dehalogenase WP_011460641.1 5 11 FAD fumarate reductase WP_005817128.1 2 6 Pyridoxamine 5’-phosphate oxidase WP_005815111.1 7 10 FAD fumarate reductase WP_005817128.1 5 3 4 Substrate-binding ABC transporter WP_005812007.1 2 2 Peptidoglycan-binding LysM WP_015943955.1 6 - - - -

Factor IIIm as the native cobamide cofactor of Geo lovleyi PceA

The cobamide cofactor for Geo lovleyi was predicted to be factor IIIm due to its genome, along with its plasmid, lacks the bzaDE genes to completely synthesize DMB, the lower ligand of cobalamin (Hazra et al., 2015). However, homology-based sequence analysis cannot accurately annotate the phylogenetically distant homologs. Therefore, in this chapter, we manage to validate this prediction. We cultivated the G. lovleyi cultures in the defined medium free of cobamide, and harvested the cells from a 100 mL culture using the method mentioned for characterization of Dsf cobamide.

Fig. 2.2. Identification of factor IIIm as the native cobamide cofactor in Geo tetrachloroethene dehalogenase (PceA) following non-denaturing, gel-electrophoretic separation of Geo crude protein extracts using BN-PAGE. (A) BN-PAGE gel slices separated for measurements of (B) PceA activity and Factor IIIm quantity. (C) MS spectrum of the extracted factor IIIm from gel slice 3. Abbreviations: CFE, cell-free protein extract. Dechlorination activity was observed in slice 2,3 and 4 (Fig. 2.2A), and slice 3 exhibited highest dechlorination activity, along with a clear protein band at protein mass range ~300 kDa (Fig. 2.2B). Consistent to previous prediction, the cobamide extracted from slice 3 and factor IIIm have a comparable m/z value ([m+H] += 1357.55; Fig. 2.2C).

In the MS spectrum, we did not observe any adduct corresponding to the m/z value of other cobamides, including factor III. Moreover, the amount of Factor IIIm extracted from each slice is proportional to the dechlorination activity from each slice (Fig. 2.2B), suggesting that (i) factor IIIm is the native cobamide cofactor in Geo PceA and (ii) most enzyme-bound cobamides in Geo lovleyi are bound in the RDase.

2.5 Discussion

Purine derivatives are widely distributed in nature, fulfilling many essential function in biological systems (Rosemeyer, 2004). However, unsubstituted purine was never assigned a specific biological function before this study. To date, nebularine (purine nucleoside) is the only known biological compound that contains unsubstituted purine. Nebularine was isolated from

Clitocybe nebularis (Löfgren and Lüning, 1953), Streptomyces yokosukanensis (Nakamura, 1961) and a Microbispora isolate (Cooper et al., 1986). In Streptomyces yokosukanensis, nebularine biosynthesis involves in a reductive deamination of adenosine (Brown and Konuk, 1995), albeit the corresponding enzyme is yet to be identified. Nebularine is a potent antibiotic against various

Mycobacterium species, exhibits cytotoxic effects in cell cultures and plants (Brown and Konuk,

1994; Gordon and Brown, 1956), and is toxic to the schistosomiasis (bilharzia) parasite (el Kouni et al., 1987); however, the biological functions for the hosts have not been resolved. Therefore, the identification of purinyl-cobamide as the native functional cofactor in Dsf PceA suggests that unsubstituted purine for some organisms is an essential compound, not merely a secondary metabolite. Moreover, the discovery of a new member of the cobamide family, purinyl-cobamide expands the number of naturally occurring cobamides to 17.

Geo lovleyi is an unusual member of the Geobacteraceae capable of organohalide respiration. Moreover, it has horizontally acquired a genomic island comprising the bzaC gene encoding an O-methyltransferase for the factor III O-methylation (Wagner et al., 2012). The bzaC is not present in the genomes of other characterized Geo species like Geo sulfurreducens and Geo metallireducens, which are both non-OHRB. As opposed to other factor III-synthesizing species, the horizontally acquired bzaC renders Geo lovleyi a desirable syntrophic partner for Dhc species, supplying factor IIIm to fulfill VC dechlorination to ethane by Dhc (Yan et al., 2012; Yan et al.,

2015). However, it is unclear if Geo PceA also requires the [5-methylated BZ]cobamides as the cofactor as Dhc does. Due to the lack of a system to express Geo PceA heterologously, along with the fact that Geo lovleyi does not possess the Btu system to efficiently uptake environmental cobamides (Wagner et al., 2012), this question remains to be resolved.

In this chapter, BN-PAGE was applied to identify the functional/native cobamide cofactors in Geo and Dsf PceA, which is accomplished with the great sensitivity of UPLC-MS to identify cobamide at sub-micromolar concentrations. Moreover, this newly developed method will not only be applied to identify cobamides, but also to identify other cofactors employed by the key enzymes of interests. For example, in the proteomic analysis of BN-PAGES slices (Table 2.1), several respiratory enzymes like CO dehydrogenase and fumarate reductase employ molybdopterin and

FAD as the cofactor, respectively. However, this approach cannot function when two or more cobamide-dependent enzymes co-exist in the same gel slice. Therefore, a pre-treatment such as partial protein fractionation or cutting the gels into thinner slices may enhance the protein separation in BN-PAGE.

Acknowledgements

We thank J. Maillard, École Polytechnique Fédérale de Lausanne, France, for providing

Dehalobacter restrictus strain PER-K23. We also thank R. F. from the BioZone Mass

Spectrometry facility, Toronto, for LC/MS assistance. This research was supported by the

Natural Science and Engineering Research Council of Canada (NSERC) Industrial Biocatalysis

Network to E.A.E. Y.

CHAPTER 3 – REFINED EXPERIMENTAL ANNOTATION REVEALS

CONSERVED CORRINOID AUTOTROPHY IN CHLOROFORM-RESPIRING

DEHALOBACTER ISOLATES

A version of this chapter is published as “Refined experimental annotation reveals conserved corrinoid autotrophy in chloroform-respiring Dehalobacter isolates”. Reproduced with permission from the ISME Journal, Springer Nature. Copyright © Macmillan Publishers

Limited, part of Springer Nature, the ISME Journal. Nov 29, 2016, DOI: 10.1038/ismej.2016.158.

3.1 Abstract Two novel chlorinated alkane-respiring Dehalobacter restrictus strains CF and DCA were isolated from the same enrichment culture, ACT-3, and characterized. The closed genomes of these highly similar sister strains were previously assembled from metagenomic sequence data and annotated.

The isolation of the strains enabled experimental verification of predicted annotations, particularly focusing on irregularities or predicted gaps in central metabolic pathways and cofactor biosynthesis. Similar to D. restrictus strain PER-K23, strains CF and DCA require arginine, histidine and threonine for growth although the corresponding biosynthesis pathways are predicted to be functional. Using strain CF to experimentally verify annotations, we determined that the predicted defective serine biosynthesis pathway can be rescued with a promiscuous serine hydroxymethyltransferase. Strain CF grew without added thiamine even though the thiamine biosynthesis pathway is predicted to be absent; intracellular thiamine diphosphate, the cofactor of carboxylases in central metabolism, was not detected in cell extracts. Thus strain CF may use amino acids to replenish central metabolites, portending entangled metabolite exchanges in ACT-

3. Consistent with annotation, strain CF possesses a functional corrinoid biosynthesis pathway,

43 demonstrated by increasing corrinoid content during growth and guided cobalamin biosynthesis in corrinoid-free medium. Chloroform toxicity to corrinoid-producing methanogens and acetogens may drive the conservation of corrinoid autotrophy in Dehalobacter strains. Heme detection in strain CF cell extracts suggests the “archaeal” heme biosynthesis pathway also functions in anaerobic Firmicutes. This study reinforces the importance of incorporating enzyme promiscuity and cofactor availability in genome-scale functional predictions and identifies essential nutrient interdependencies in anaerobic dechlorinating microbial communities.

3.2 Introduction

Halogenated organic compounds include a wide array of naturally-occurring and manmade chemicals with moderate to high toxicity. Many make excellent hydrophobic solvents. For example, Chloroform (CF) and 1,1,1-trichloroethane (1,1,1-TCA) are frequent groundwater contaminants because of their widespread industrial use as solvents and historically poor disposal practices (Furukawa et al., 2005). CF is also one of the most abundant organohalides in the atmosphere (estimated annual global flux is from 0.7 to 4 million t) (Cappelletti et al., 2012; Harper,

2000). In fact, CF in the environment is primarily of natural origin (~90%) (McCulloch, 2003), produced by algae in marine systems (Nightingale et al., 1995; Scarratt and Moore, 1999) and by termites in terrestrial systems (Khalil et al., 1990; Laturnus et al., 2002). In addition to neurotoxicity and carcinogenic activity, CF and 1,1,1-TCA are also strong inhibitors of many microbial processes essential for biogeochemical cycling, such as methanogenesis, acetogenesis, and reductive dechlorination of chlorinated ethenes (Bagley et al., 2000; Duhamel et al., 2002), and are ozone-depleting substances (Slaper et al., 1996).

Members of the Dehalobacter genus are anaerobes found in sediments of river and marine ecosystems (Zanaroli et al., 2010) as well as groundwater aquifers (Grostern and Edwards, 2006).

Dehalobacter are specialized in organohalide respiration (Holliger et al., 1998) or organohalide fermentation (Justicia-Leon et al., 2012; Lee et al., 2012), and are active participants in the global halogen cycle. Based on 16S rRNA gene sequences (>99% average nucleotide identity), all characterized strains described to-date belong to the type species Dehalobacter restrictus and use

H2 or formate as an electron donor to reductively dechlorinate a variety of chlorinated and brominated hydrocarbons (Deshpande et al., 2013; Grostern et al., 2010; Grostern and Edwards,

2009; Holliger et al., 1998; Nelson et al., 2014; Sun et al., 2002; Tang and Edwards, 2013; Wang et al., 2014; Yoshida et al., 2009). D. restrictus strains CF and DCA were enriched from a mixed culture referred to as ACT-3 that dechlorinates 1,1,1-TCA via 1,1-dichloroethane (1,1-DCA) to chloroethane (CA) and CF to dichloromethane (DCM). ACT-3 is also used commercially for bioaugmentation (http://sirem-lab.com/products/kb-1). Reductive dehalogenases (RDases) are corrinoid-dependent enzymes that catalyze organohalide dehalogenation (Jugder et al., 2016). In

ACT-3, the dechlorination of CF and 1,1,1-TCA is catalyzed by CfrA (CF reductase subunit A) from strain CF; the dechlorination of 1,1-DCA is catalyzed by DcrA from strain DCA. CfrA and

DcrA were the only two RDases found expressed of the 19 RDase genes in ACT-3 (Tang and

Edwards, 2013).

Since OHRB like Dehalobacter are key players in sustaining the global halogen cycle

(Krzmarzick et al., 2012) and in bioremediation (Jackson, 2004; Jugder et al., 2016; Justicia-Leon et al., 2014; Justicia-Leon et al., 2012), realizing their metabolic and physiological characteristics is of great importance. This characterization enables more accurate activity predictions, as illustrated by a recent study that found that Dehalococcoides, another genus of OHRB, requires a

45 specific form of corrinoid cofactor to degrade the carcinogen vinyl chloride (Yan et al., 2015). The closed genomes of strains CF and DCA were previously assembled from the metagenome of the

ACT-3 mixed culture (Tang et al., 2012) and available under NCBI accession numbers CP003869 and CP003870. These two highly similar genomes (>90% whole genome average nucleotide identity) were then compared to three other available Dehalobacter genomes (strains PER-K23,

E1 and UNSWDHB) to categorize common and unique genome-inferred physiological and metabolic traits (Tang et al., 2016). The genomes differed primarily in the complement of RDases and the completeness of the corrinoid biosynthesis pathway. However, all five genomes harbour the nearly identical complement of genes ascribed to central metabolism (the TCA cycle) and amino acid and cofactor biosynthesis. All strains appear to have the same defects in serine, biotin and thiamine biosynthesis, while they all harbour a complete archaeal heme biosynthesis pathway

(Tang et al., 2016).

Recent advances in bioinformatics and sequencing technology have generated a large number of genomes and metagenomes from a wide variety of environments that are being mined for functional and ecological traits. However, bioinformatic analyses, while powerful, are often insufficient to reliably predict many metabolic features of an organism because of the considerable number of mis-annotations and hypothetical genes in every genome (Sañudo-Wilhelmy et al.,

2014). A relevant example is the recent discovery of the anaerobic biosynthesis genes of 5,6- dimethylbenzimidazole (Hazra et al., 2015), the lower ligand of the best known corrinoid, cobalamin (vitamin B12). The genes encoding 5-hydroxybenzimidazole synthase (bzaABF) were previously wrongly annotated as thiamine biosynthesis genes due to the homology-based annotation (Hazra et al., 2015).

Here we describe the isolation, growth, morphological characteristics, and electron donor and electron acceptor substrate ranges of strains CF and DCA. Guided by previous genomic and transcriptomic analyses (Rupakula et al., 2013; Tang et al., 2016), we experimentally tested specific genomic predictions, particularly where annotations were uncertain or inconsistent with the observed phenotypes. Tang et al. (2016) classified and curated genome annotations of 5

Dehalobacter strains including strains CF and DCA. Since the orthologous proteins from the five strains are highly similar, typically sharing >98% protein sequence identity, strain CF was chosen as an appropriate model organism for pathways common to all sequenced strains. We explored the predicted gaps in the TCA cycle, potential defects in amino acid biosynthesis and were able to refine annotations of genes involved in serine biosynthesis. We also evaluated functionality of biosynthetic pathways for biotin, corrinoid, heme, and thiamine. These experimental efforts verify genome annotations and bring previous proteomic data of ACT-3 and strain PER-K23 cultures

(Rupakula et al., 2013; Tang and Edwards, 2013) into context.

3.3 Materials and Methods

(Supplementary Figures and Tables are shown in Appendix D)

Isolation and growth conditions. For strain isolation, two highly-enriched subcultures,

Dehalobacter-1,1,1-TCA/H2 and Dehalobacter-1,1-DCA/H2, were constructed by feeding transfers from the parent mixed culture ACT-3 (Tang and Edwards, 2013) with either 1,1,1-TCA or 1,1-DCA (2 mM nominal) as the electron acceptor in minimal mineral medium (MM medium)

(Edwards and Grbić-Galić, 1994). H2 was provided as the electron donor (25% in the headspace;

20 mM nominal). Sodium acetate (5 mM) was provided as the carbon source. Specifically, for dilution-to-extinction transfers, inside an anoxic chamber, 7 mL of MM medium, 3 mL of ACT-3

47 mixed culture supernatant (described in Appendix D), sodium acetate (5 mM), 2 µL of neat chlorinated substrate (1,1,1-TCA or 1,1-DCA), and 6.4 mL of H2/CO2 (80% v/v) were added to a

25 mL butyl rubber stopper-sealed and crimped Bellco glass tube. After autoclaving, 10 μL of

1,000x filter-sterilized vitamin solution (Appendix D) and 20 μL of autoclave-sterilized iron sulfide (FeS) slurry (~8 g/L) (Appendix D) were injected into each tube. The enriched subcultures were serially diluted from 10-1 to 10-9. To minimize carryover, inocula were pelleted down to remove supernatant, and were resuspended in an equal volume of sterile MM medium before inoculation. The diluted transfers were incubated at room temperature and, dechlorination was monitored by gas chromatography as described in Appendix D. Growth conditions for experiments evaluating electron donors, electron acceptors, and vitamin requirement were similar to the conditions for these dilution-to-extinction transfers with some modifications (Appendix D).

The average volumetric dechlorination rates of cultures were determined from the total amount of dechlorinated products generated per day in each bottle divided by the culture volume when >90% of the chlorinated substrates were depleted in the three replicates.

Isolate purity tests. Contamination of the isolate cultures with other microorganisms was investigated by making transfers (10-3 dilution) into MM medium described above but excluding the chlorinated substrates (to detect homoacetogens), or replacing chlorinated electron acceptors with 5 mM NaNO3 (for nitrate reducers), 5 mM NaSO4 (for sulfate reducers), a mixture of methanol, ethanol and lactate (for fermenting microbes), or 0.1 g/L yeast extract. These new transfers were incubated at room temperature for 1 month and then examined by restriction fragment length polymorphism (RFLP) (Appendix D) and epifluorescence microcopy with 4′,6- diamidino-2-phenylindole dihydrochloride (DAPI) staining (to look for any growth) using standard protocols as described in Appendix D. Two pairs of specific primers targeting cfrA and

48 dcrA (Appendix D) developed previously (Tang and Edwards, 2013) were used to examine the potential cross-contamination of strains CF and DCA in their isolate cultures.

Enzyme activity assays. Established assays for malate dehydrogenase, succinate dehydrogenase,

NADP-dependent malic enzyme (MAE), and reductive dehalogenase (RDase) activity were conducted using cell extracts prepared from Dehalobacter strain CF and from E. coli strain K-12

(Appendix D). Cell extracts from both cultures were adjusted to a final total protein concentration of 50 µg/mL in each assay. Serine hydroxymethyltransferase (SerB) and threonine aldolase activity assays were conducted using purified recombinant SerB from strain CF (accession number

AFV06891). Cloning, expression and purification of Strain CF SerB is described in Appendix D.

Protein concentrations of cell extracts were determined using the Pierce bicinchoninic acid protein assay kit (Thermo Scientific) with a linear range spanning from 20 to 2,000 µg/mL of protein.

Specific activity is reported as µmol of substrate consumed or product generated per min per mg of protein, as well as in ncat/mg.

Extraction and derivatization of intracellular thiamine species. Thiamine can exist as free and phosphorylated forms within a cell. To measure the concentration of intracellular thiamine species, a protocol involving derivatization into thiochromes was employed (Appendix D) (Leonardi and

Roach, 2004).

Corrinoid purification, measurement, and structure characterization. Corrinoid content in strain CF culture was quantified using a Lactobacillus growth-based B12 assay following established protocols with some modifications (Yan et al., 2013; Yan et al., 2012). For the B12 assay, strain CF cells grown in corrinoid-free medium were transferred twice (10-2 dilution) to exclude any carryover. The third transfer was sampled over time during growth and the samples were assayed using the growth-based Lactobacillus auxotroph assay. In this assay, Lactobacillus

49 growth increased linearly over a corrinoid concentration range from 2 to 20 ng/L (Fig. S1). To determine total corrinoid content in cultures, pH of the samples was adjusted to 5-6 with 3% acetic acid, and the samples were incubated in a boiling water bath for 30 min, followed by centrifugation at 16,000 x g for 15 min at 4ºC. The supernatants were diluted 10-50 times and stored at -20 ºC until use. The corrinoids produced by strain CF were extracted and derivatized following previous established protocols (Yan et al., 2013; Yan et al., 2012), and characterized using ultra performance liquid chromatography-mass spectroscopy (UPLC-MS) (Appendix D).

Analytical procedures. Gas chromatography to measure concentrations of chlorinated substrate analyses, high pressure liquid chromatography (HPLC) for measuring thiamine derivatives and acetaldehyde, and UPLC-MS for biotin, corrinoid, heme, and amino acid detection are described in the Appendix D.

3.4 Results and discussion

Isolation and characterization of Dehalobacter strains

(i) Isolation and verification of purity. Isolation of the strains CF and DCA was achieved via repeated dilution-to-extinction transfers from the original mixed parent culture, ACT-3. The addition of autoclaved sterile mixed-culture supernatant prepared from the ACT-3 parent culture was required to sustain the growth of the 10-9 dilution transfers (i.e., detection of dechlorination activity) (Fig. S2). Nineteen successive 10-9 dilution transfers were performed. To replace mixed culture supernatant in the growth medium for the isolate cultures, a chemically-defined medium

(Appendix D) similar to that used for the isolation of D. restrictus strain PER-K23 (Holliger et al.,

1998) was tested. Cultures of the two isolates were able to deplete 1 mM chlorinated substrate

(1,1,1-TCA or 1,1-DCA) after three successive 10% transfers (v/v) without mixed culture

50 supernatant in MM medium when 0.1 mM of each arginine, histidine, and threonine were provided

(Fig. 3.1A). These three amino acids may not represent the only viable combination required for growth but other combinations were not explored. The transfer cultures of the two isolate strains stopped dechlorinating when any of the three amino acids was excluded from the medium (data not shown). Based on the highly similar 16S rRNA gene sequences of strains CF and DCA to strain PER-K23 (99.3% average nucleotide identity), and their shared carbon source requirement, strains CF and DCA belong to the type species Dehalobacter restrictus.

Fig. 3.1. Dechlorination over three serial 10% transfers of strain CF. Panel (A) cultures grown on chemically defined medium amended with the three vitamins (biotin, cobalamin, and thiamine see Appendix D for exact composition); Panel (B) cultures grown in medium lacking biotin, cobalamin, and thiamine; Panel (C) average dechlorination rates for strain CF cultures grown on medium with different vitamin combinations noted below. Symbols: ♦, 1,1,1-TCA; ◊, 1,1-DCA. Data are mean ± SE of three replicates in each experiment.

Fig. 3.2. Microscopic images of Dehalobacter strains. Epifluorescence microscopic images of strains CF (A) and DCA (B) at 1,000x magnification. (C) Scanning electron microscopic image and (D) transmission electron microscopic image of the strain CF cell. Scale bars are shown in each panel. Note the long rods in Fig. 2B are formed by multiple strain DCA cells lining up end-to-end. Abbreviations: S, the proteinous S- layer; P, the peptidoglycan layer; CM, the cytoplasmic membrane; G, the electron-dense granule. Microscopy suggested no growth in chlorinated substrate-free transfer cultures amended with yeast extract, a mixture of methanol, ethanol, and sodium lactate, or with electron acceptors nitrate, and sulfate. To corroborate the purity of isolates, DNA was extracted from isolate cultures and the 16S rRNA gene was amplified by general bacterial primers 27f and 1541r. Restriction fragment length polymorphism (RFLP) patterns reflected the presence of the only two Dehalobacter 16S rRNA genes in the isolate cultures (Fig. S3ABC). Strain CF cultures did not dechlorinate 1,1-DCA, and strain DCA cultures did not dechlorinate CF and 1,1,1-TCA. Moreover, cfrA was only amplified from strain CF DNA, and dcrA was only amplified from strain DCA DNA (Fig. S3D), confirming that the two Dehalobacter restrictus strains were indeed distinct.

(ii) Morphology. Microscopic images of samples of both isolate cultures stained with DAPI revealed a uniform cellular morphology, further supporting the culture purity (Fig. 3.2A,B).

Consistent with the three previously described Dehalobacter isolates (strains PER-K23, TCA1, and UNSWDHB) (Holliger et al., 1998; Sun et al., 2002; Wong et al., 2016), the cells of strains

CF and DCA are either straight or curved rods (Fig. 3.2A,B) approximately 2~6 µm long with a diameter of 0.4-0.6 µm (Fig. 3.2C). Similar to strain PER-K23 (Holliger et al., 1998), strain CF stains Gram-negative (Fig. S4), while it is phylogenetically Gram-positive. This negative staining may result from the presence of a proteinaceous S-layer on the exterior surface of Dehalobacter cells (Fig. 3.2D) (Holliger et al., 1998). In some strain CF cells, an electron-dense granule was also observed (Fig. 3.2D).

(iii) Electron donors and acceptors. According to annotations, Dehalobacter genomes possess various oxidoreductase genes involved in fermentation (Tang et al., 2016), suggesting that they may be able to use electron donors other than H2 or formate. The only genes for electron-accepting reactions in anaerobic respiration identified in the genomes were RDases. A broad range of electron donors and acceptors were tested to evaluate the substrate spectra of the both strains CF and DCA. Of the many electron donors tested, the two strains were only able to oxidize H2 and formate (Table S1A). The ability of both strains to use formate as the electron donor is consistent with the expression of Dehalobacter formate dehydrogenase (EC 1.2.1.2) in both 1,1,1-TCA- and

1,1,-DCA-fed ACT-3 mixed cultures (Tang and Edwards, 2013). However, the ability to use formate appeared to be conditional on the addition of ACT-3 mixed culture supernatant, yeast extract, or casamino acids to the MM medium. Dechlorination with formate as electron donor was not sustained in medium that was not supplemented (data not shown). This conditional formate utilization may result from the inability to synthesize molybdopterin, the cofactor for formate

53 dehydrogenase. Genome annotation previously revealed that none of the strains possess complete molybdopterin biosynthesis genes (Tang et al., 2016). Therefore, strains CF and DCA may rely on molybdopterin supplementation from other community members in ACT-3 to respire formate.

Unfortunately, the absence of commercially available molybdopterin standards has hampered our attempt to study this further.

We determined that strain CF could use 1,1,2-trichloroethane (1,1,2-TCA) as electron acceptor in addition to 1,1,1-TCA and CF. Strain DCA could also use 1,1,2-TCA in addition to

1,1-DCA (Table S1B). However, the tolerance of strain CF to aqueous CF concentrations ([CF]aq) is much lower (below 0.42 mM) than to [1,1,1-TCA]aq and [1,1,2-TCA]aq (above 1 mM). The mixed culture was much less sensitive to CF, tolerating above 1 mM. No other chlorinated ethene or ethane tested was dechlorinated by the isolates (Table S1B). Because we could amend with higher concentrations, 1,1,1-TCA was used as the electron acceptor for strain CF in experiments to verify genome annotations. The genomes of strains CF, DCA, and UNSWDHB contain a complete operon (pceABCT) in their genomes encoding an ortholog of the characterized tetrachloroethene (PCE) RDase PceA from strain PER-K23 (Tang et al., 2016; Wong et al., 2016)

(>89% amino acid identity). However, none of the strains was found to dechlorinate PCE. This lack of activity may result from the loss of a functional transcription factor to induce PceA expression or from a mutation at key catalytic residues.

To determine if CfrA can also dechlorinate 1,1,2-TCA, we conducted RDase enzyme assays using cell extracts of pure strain CF culture grown on 1,1,1-TCA that is known to express only CfrA. These cell extracts not only dechlorinated 1,1,1-TCA, but also CF and 1,1,2-TCA, confirming substrate promiscuity of CfrA (Table 3.1). Heat-treated cell extracts did not catalyze

1,1,2-TCA dechlorination. Moreover, the fact that strain DCA is able to respire 1,1,2-TCA, and

54 that DcrA is the only RDase detected in 1,1-DCA grown cells (Tang and Edwards, 2013) suggest

DcrA also dechlorinates 1,1,2-TCA. 1,1,2-TCA is therefore a common substrate for four similar

RDases, including CfrA and DcrA described above, TmrA from strain UNSWDHB (Wong et al.,

2016), and CtrA from Desulfitobacterium sp. strain PR (Zhao et al., 2015). These chlorinated alkane-dechlorinating RDases define a protein cluster (ortholog group) based on >90% amino acid identity (Hug and Edwards, 2013; Tang et al., 2016). Interestingly, each strain and corresponding active RDase have slightly different substrate preferences supporting a previous contention that differentiation of organohalide respiring strains is driven by chlorinated substrates (Islam et al.,

2010; Tang et al., 2016).

Experimental verification of genome annotations of strain CF

(i) Missing TCA cycle genes. The TCA cycle in all five sequenced Dehalobacter genomes is predicted to be incomplete because the genes encoding malate dehydrogenase and succinate dehydrogenase were not found (Fig. 3.3, pathways marked with red x). If these genes are truly absent, neither fumarate nor malate could be synthesized. Dehalobacter strains need the TCA cycle

Table 3. 1. Dechlorinating activity observed in strain CF cell extracts Dechlorination activity Substrate Product (nmol/day/mg protein)a

CF DCM 868 ± 60

1,1,1-TCA 1,1-DCA 302 ± 26

1,2-DCA (major) 322 ± 28 1,1,2-TCA VC (minor) 7 ± 0.6 aData are mean ± SE of three replicates in each experiment.

Fig. 3.3. Schematic of the proposed metabolic map of strain CF. The red crosses (X) represent missing genes in the strain CF genome, while a red curved dashed line beside a reaction represents the absence of the essential cofactor. The black arrows represent nutrients imported from medium. Note that the threonine- glycine conversion (highlighted in blue) is catalyzed by promiscuous serine hydroxymethyltransferase (SerB). Abbreviations for cofactors and enzymes involved in the reactions are listed below with full name: CfrA, chloroform reductive dehalogenase; PLP, pyridoxal phosphate; TDP, thiamine diphosphate; PK, pyruvate kinase; PPDK, pyruvate phosphate dikinase; FDH, formate dehydrogenase [molybdopterin]; FHL, formate hydrogen lyase complex [molybdopterin]; ACS, acetate:CoA ligase (AMP-forming); PFL, pyruvate formate-lyase; PFO, pyruvate-flavodoxin oxidoreductase [TDP]; PCT, pyruvate carboxyl transferase [biotin]; MAE, NADP-dependent malic enzyme [NADP]; CS, citrate synthase; AH, aconitate hydratase; ICDH, isocitrate dehydrogenase [NADP]; 2-OGs, 2-oxoglutarate:ferredoxin oxidoreductase [TDP]; SCL, succinyl-CoA ligase; FH, fumarate hydratase; GK, glucokinase; PSP, phosphoserine phosphatase; OAD, oxaloacetate decarboxylase [biotin]; SDH, succinate dehydrogenase[heme]; SDHT, serine dehydratase; MDH, malate dehydrogenase; ICL, isocitrate lyase; MS, malate synthase; TAL, threonine aldolase [PLP]; SerB, serine hydroxymethyltransferase [PLP]. Other abbreviations: CM, cytoplasmic membrane; CW, cell wall; WL pathway, Wood–Ljungdahl pathway; Cyt b, cytochrome b. to synthesize oxaloacetate and 2-oxoglutarate for amino acid biosynthesis (Miflin and Lea, 1977).

The absence of the two enzymes may be associated with the amino acid requirement of

Dehalobacter. Alternatively, a gene annotated as NADP-dependent malic enzyme (MAE; EC

1.1.1.40) is present in all Dehalobacter genomes and may play a role in compensating for missing

TCA cycle genes. If active, this predicted MAE gene product could catalyze pyruvate-malate conversion (Fig. 3.3).

Moreover, the annotated malic enzyme is also expressed in the ACT-3 mixed culture and proteome of strain PER-K23 (Rupakula et al., 2013; Tang and Edwards, 2013). We assayed cell extracts of strain CF and E. coli (positive control) for malate and succinate dehydrogenase as well as MAE activities (Table 3.2). The latter activity was detected in both strain CF (0.06

µmol/min/mg or 1 nkat/mg) and E. coli (0.01 µmol/min/mg or 0.17 nkat/mg), while malate dehydrogenase and succinate dehydrogenase activities were only detected in E. coli cell extracts.

Consistent with annotation, enzyme assays revealed that the activities of malate and succinate dehydrogenase are absent in Dehalobacter genomes. However, the MAE can salvage the synthesis of malate and fumarate from pyruvate (Fig. 3.3).

Table 3. 2. Enzymatic activities in cell extracts of E. coli and strain CF and promiscuous threonine aldolase activity of purified Dehalobacter serine hydroxymethyltransferase (SerB) Specific activity

(µmol/min/mg protein) a Detected Substrate E. coli Strain CF Product Enzymatic activities in cell extracts Malate dehydrogenase Oxaloacetate NAD 1.3 ± 0.2 ND Succinate dehydrogenase Succinate DCPIP 0.03 ± 0.006 ND NADP-dependent malic enzyme Malate NADPH 0.01 ± 0.001 0.06 ± 0.003 Enzymatic activity of purified Dehalobacter SerB SerB L-serine Glycine n/a 3.4 ± 0.2

Threonine aldolase L-threonine Acetaldehyde n/a 0.07 ± 0.003 aData are mean ± SE of three replicates in each experiment.

Abbreviations: NAD, nicotiamide adenine dinucleotide; NADPH, nicotinamide adenine dinucleotide phosphate hydrogen; DCPIP, 2,6-dichlorophenolindophenol; ND, not detected; n/a, not analyzed.

(ii) Serine biosynthesis. Strains CF, DCA, and PER-K23 were found to require arginine, histidine, and threonine for growth, yet the biosynthesis pathways for these three amino acids appear complete in all the five Dehalobacter genomes. Rather, their genomes are missing the gene encoding phosphoserine phosphatase (PSP) (EC 3.1.3.3), the enzymes that catalyzes the last step of the conventional serine biosynthesis pathway (Greenberg and Ichihara, 1957), while other serine salvage pathways are not present (Fig. S5A). A defect in serine biosynthesis should disable biosynthesis of glycine and cysteine (Fig. S5). However, strains CF and DCA are able to grow in a defined medium (supplemented with arginine, histidine, and threonine) (Fig. 3.3) without cysteine, glycine, and serine, suggesting the presence of an alternative mechanism to produce serine, most likely from the three amino acids provided, as explained below.

Threonine can be converted to serine via glycine (Fig. 3.3). However, threonine aldolase

(EC 4.1.2.5), the enzyme catalyzing the threonine-glycine conversion, is absent in the five

Dehalobacter genomes. Interestingly, serine hydroxyl methyltransferase (SerB; EC 2.1.2.1), the enzyme catalyzing glycine-serine conversion, has been reported to possess promiscuous threonine aldolase activity in both E. coli and Methanosarcina barkeri strain Fusaro (Buchenau and Thauer,

2004; Ogawa et al., 2000). SerB is also expressed in the proteome of strain PER-K23 (Rupakula et al., 2013). Furthermore, the SerB of M. barkeri strain Fusaro shares >59% protein sequence identity to the sequences of the five Dehalobacter SHMTs, indicating Dehalobacter may utilize

SerB to synthesize serine from threonine. The lack of a PSP-coding gene can also be rescued by other promiscuous phosphatases in the haloacid dehalogenase (HAD) family. For example, it has been reported that histidinol phosphatase (HisB; EC 3.1.3.15) in histidine biosynthesis can rescue missing PSP in E. coli strain K-12 and Thermococcus onnurineus strain NA1 (Lee et al., 2008;

Patrick et al., 2007; Yip and Matsumura, 2013). However, histidinol phosphatases (HisJ) in

Dehalobacter genomes does not belong to the HAD family (Tang et al., 2016).

To test if Dehalobacter can synthesize serine, the gene for SerB (accession number

AFV06891) from strain CF was cloned, heterologously expressed, and purified (Fig. S5B). Strain

CF SerB revealed both threonine aldolase activity (0.07 µmol/min/mg or 1.2 nkat/mg) and hydroxyl methyltransferase activity (3.4 µmol/min/mg or 57 nkat/mg; Table 3.2), suggesting that strain CF can synthesize serine from threonine via glycine. None of the enzyme-free or substrate- free controls showed any activity. Thus, the presence of a serine salvage mechanism enables serine production from threonine (also from glycine or cysteine) in strain CF. However, this does not explain why strain CF requires threonine as the biosynthesis pathway for threonine is complete.

This discrepancy is further discussed in the section on thiamine section below.

In a recent comparative genomic analysis of >6,000 sequenced bacteria from diverse environments, most Firmicutes were found to be serine auxotrophs based on annotated defects in the serine biosynthesis pathway (Mee et al., 2014), consistent with previous annotation of

Dehalobacter genomes (Tang et al., 2016). However, we have now shown that these ostensible serine auxotrophs can actually synthesize serine using promiscuous enzymes, illustrating the shortcomings of typical bioinformatic analyses without experimental verification. Models constructed based on this comparative genomic analysis may assume that Firmicutes require serine from other community members when in fact they do not. For example, Desulfitobacterium hafniense strain Y51, another organohalide-respiring model organism and a member of Firmicutes, may also synthesize serine using promiscuous SerB because (i) the organism lacks the PSP-coding gene in its genome (Nonaka et al., 2006), and (ii) its SerB (accession number BAE86716) shares

75% protein sequence identity with the homolog in strain CF, and (iii) our data revealed that D. hafniense strain Y51 can grow in defined medium without amino acid supplementation (Bi, 2015).

(iii) Biosynthesis of organic cofactors. The growth reliance of Dehalobacter on the three amino acids can also result from defects in cofactor biosynthesis. It is possible that (i) Dehalobacter cannot synthesize specific cofactors or (ii) Dehalobacter is not able to transform certain cofactor pre-cursors into functional forms. For example, thiamine is converted to thiamine monophosphate or thiamine diphosphate (TDP) intracellularly to be functional (Leonardi and Roach, 2004). In genomes of strains CF and DCA, the biosynthetic pathway for biotin appears to be missing 4/5 genes and for thiamine the pathway is missing 5/9 genes (Tang et al., 2016). To determine if these pathways are truly incomplete, growth of strain CF was tested in the absence of these cofactors.

We also tested the growth of strain CF in the presence and absence of cobalamin since genome annotation predicted a complete corrinoid biosynthesis pathway. Growth of strain CF was sustained in the absence of these three cofactors (biotin, cobalamin, and thiamine) in cultures that were spun down and transferred three successive times (10% v/v inoculum) (Fig. 3.1B). Removal of cobalamin decreased the dechlorination rate by about a factor of 3 (from 30 to 10 µM/day nominal), while exclusion of biotin and thiamine from the medium did not affect the growth rate

(11 µM/day nominal) (Fig. 3.1C). When grown with the three vitamins, the cell yield for strain

CF determined from microscopic cell counts, which is 5.73 (± 1.79) x 1012 (n=9) cells/mol Cl- released, is comparable to the qPCR-determined cell yield of strain CF in CF-fed mixed culture

ACT-3, which is 6 (± 0.03) x 1012 cells/mol Cl- released (Grostern et al., 2010).

Biotin biosynthesis and metabolism. Biotin is considered vital for organisms in all domains of life due to its essential role in fatty acid biosynthesis (cofactor for acetyl-CoA carboxylase) (Lin and Cronan, 2011), yet no known biotin biosynthesis pathway was identified in the five

Dehalobacter genomes even though both strain PER-K23 (Holliger et al., 1998) and strain CF

(Fig. 3.1C) can grow without added biotin. It was reported that biotin-independent malonate decarboxylases (Hoenke et al., 1997) can circumvent the need for biotin in fatty acid biosynthesis; however, a homolog of this gene was not found in the five Dehalobacter genomes. Biotin is also an essential cofactor for pyruvate carboxyltransferase (PCT; Fig. 3.3) that can convert pyruvate to oxaloacetate for amino acid biosynthesis, and the PCT coding-gene is annotated in all the five

Dehalobacter genomes (Tang et al., 2016). Moreover, traces of biotin may be sufficient to support growth. It was reported that as few as 1,000 biotin carboxyl carrier proteins are present in each E. coli cell (Li and Cronan, 1993). Accordingly, medium with a biotin concentration of 0.4 µg/L should be enough to support the growth of an E. coli culture to a cell density of 109 cells/mL.

To determine if biotin is present at sufficient quantities in our medium as a contaminant, we analyzed samples of the concentrated stock solutions used to make the MM medium by LC-

MS. We detected biotin in the 1,000x vitamin stock solution (excluding biotin; referred to as biotin-) at a concentration of 15 µg/L (Fig. S6), indicating that (after 1000x dilution of the stock) the medium may contain ~15 ng/L biotin. This amount is low but may be sufficient to support the growth of strain CF cultures because they only reach a cell density ~108 cells/mL and require biotin for anabolism but not for respiration. Nevertheless, we cannot exclude the possibility that an uncharacterized biotin biosynthesis pathway is functioning in strain CF.

Thiamine biosynthesis and metabolism. The absence of a thiamine biosynthesis pathway in

Dehalobacter genomes is consistent with the observation that the growth of strain PER-K23 required thiamine supplementation (Holliger et al., 1998). In contrast, the growth of strain CF seems not to require thiamine addition (Fig. 3.1C).

Fig. 3.4. Determination of intracellular thiamine derivatives in strain CF cells. Chromatograms from HPLC analysis with fluorescence detection (excitation at 365 nm and emission at 435 nm) of: (A) 1 µM thiamine standard solution; (B) 1 µM thiamine monophosphate; (C) 1 µM thiamine diphosphate; (D) cell extract of E. coli; and (E) cell extract of strain CF. TDP is the cofactor of pyruvate-flavodoxin oxidoreductase and 2-oxoglutarate decarboxylase, two important carboxylases in central metabolism (Fig. 3.3) (Pohl et al., 2002). To test whether strain

CF can synthesize TDP from thiamine, cultures of strain CF and E. coli grown on thiamine- containing medium (25 µg/L) were harvested for intracellular thiamine derivatives analysis. TDP was only detected in E. coli cell extracts, while thiamine was detected in both strain CF and E. coli cell extracts (Fig. 3.4). Interestingly, although a gene annotated as thiamine pyrophosphokinase

(EC 2.7.6.2) is present in the strain PER-K23 genome (Tang et al., 2016), the enzyme was not

62 found to be expressed in the strain PER-K23 proteomes (Rupakula et al., 2013), consistent with the lack of TDP detection.

The lack of TDP indicates that strain CF only uses pyruvate formate-lyase for acetyl-CoA- pyruvate conversion, and not the TDP-dependent pyruvate-flavodoxin oxidoreductase (PFO) (Fig.

3.3). However, it is unclear if an organism could use only pyruvate formate-lyase to assimilate acetate and formate into pyruvate in defined medium. The only relevant study we found in literature did not test the effect of deleting the PFO-like ydbK (Nakayama et al., 2013) in their E. coli strain to exclude its contribution to pyruvate production (Zelcbuch et al., 2016). Moreover, threonine can be converted into pyruvate via serine (Fig. 3.3). Furthermore, strain CF cultures grown on pyruvate as the main carbon source dechlorinated at twice the rate (53 µM/day nominal) of cultures grown on acetate (29 µM/day nominal) (amino acids supplemented in both conditions)

(Fig. S7). Therefore, strain CF likely requires amino acid supplementation for growth because of difficulties in acetyl-CoA-pyruvate conversion, which in turn block synthesis of oxaloacetate and thus 2-oxo-glutarate (precursors for amino acids) (Fig. 3.3) (Miflin and Lea, 1977). Such nutrient requirements for strain CF portend a tightly entangled syntrophy in the ACT-3 microbial community.

Corrinoid biosynthesis. Corrinoids, such as cobalamin, play a crucial role in methionine and ribonucleotide biosynthesis, as well as energy metabolism in organohalide-respiring bacteria

(Maillard et al., 2003; Miller et al., 1998; Sañudo-Wilhelmy et al., 2014). Reductive dechlorination and growth of organohalide-respiring bacteria are usually enhanced with corrinoid supplementation (Yan et al., 2013). It is reported that strains TCA1 (Sun et al., 2002) and

UNSWDHB (Wong et al., 2016) grow without corrinoid supplementation. Strain CF could also grow without corrinoid supplementation (Fig. 3.1B).

Fig. 3.5. Corrinoid production and guided cobalamin synthesis by strain CF. (A) Dechlorination profiles (production of 1,1-DCA from 1,1,1-TCA) and (B) corresponding corrinoid production in cultures grown under different conditions as shown. ●, autoclaved negative control (NC); ■, 1,1,1-TCA-free control (TCA-); ∆, cobalamin- and DMB-free cultures (Cbl-); ▼, corrinoid-free and DMB-containing cultures (DMB+); ○, cobalamin-containing positive control (Cbl+). (C) UPLC-ESI-MS analysis of guided cobalamin synthesis in strain CF culture with DMB supplementation. The chromatograms on top panel shown are extracted ion chromatograms (m/z=1355.57 ± 0.013) and the chemicals are ionized in positive mode. Bottom panel shows mass spectrum. CN-Cbl STD, cyanocobalamin standard. Data are mean ± SE of three replicates in each experiment. Consistent with these observations, genomes of strains CF, DCA (Tang et al., 2016), and

UNSWDHB (Deshpande et al., 2013) all possess a complete cobinamide biosynthesis pathway, while strains PER-K23 (Rupakula et al., 2013) and E1 (Maphosa et al., 2012) do not. However, to date, no direct evidence for de novo corrinoid synthesis by Dehalobacter exists. Thus the corrinoid

64 concentration in strain CF cultures grown in corrinoid-free medium was quantified using a microbial B12 assay (Yan et al., 2013). The significant increase of corrinoid concentration along with dechlorination confirms the ability of strain CF to synthesize corrinoid de novo (Fig. 3.5A-

5B). We attempted to characterize the type of corrinoid using UPLC-ESI-MS, however, none of the known cobamides with a benzimidazolyl or phenyl type lower ligand was detected in the cell extract of strain CF culture grown on corrinoid-free medium based on m/z values and retention times of the known cobamide standards (Table S2). Nevertheless, cobalamin was identified in cell extracts of strain CF culture when 10 µM of the lower ligand compound 5,6- dimethylbenzimidazole (DMB) was supplemented to the corrinoid-free medium (Fig. 3.5C). The guided cobalamin biosynthesis confirmed that strain CF possesses a functional cobinamide biosynthesis pathway to assemble cobalamin at the presence of DMB (Tang et al., 2016). No cobinamide or cobalamin was identified in the commercial DMB and medium used for growth

(Fig. S8). Thus, strain CF produces a cobamide that is yet to be identified, likely a purine species as the lower ligand as reported for strains of Clostridium (adenine) and Desulfovibrio species

(guanine and hypoxanthine) (Guindon and Gascuel, 2003; Hoffmann et al., 2000).

None of the bza genes involved in anaerobic DMB biosynthesis (Hazra et al., 2015) was identified in the five Dehalobacter genomes (Tang et al., 2016), consistent with the observation that no benzimidazole-type cobamide was identified from the strain CF culture. However, when

DMB was provided, strain CF preferentially synthesized cobalamin rather than its native cobamide.

This is in agreement with the discovery that DMB is a universally-preferred substrate for the lower ligand-activating enzyme (CobT) in cobamide biosynthesis (Hazra et al., 2013). Dechlorination rates in DMB-supplemented cultures were comparable to rates in cobalamin-supplemented cultures (Fig. 3.5A), indicating that corrinoid synthesis in strain CF is limited by lower ligand

65 availability but not by the availability of the porphyrin backbone. Furthermore, total corrinoid concentrations are higher in DMB-supplemented cultures than those without DMB (Fig. 3.5B).

The Dehalobacter strains enriched from CF- and 1,1,1-TCA-dechlorinating consortia have retained their ability to synthesize corrinoids, while strains PER-K23 and E1 enriched with other halogenated substrates (PCE and β-hexachlorocyclohexane) do not. Strains PER-K23 and E1 do have partial pathways, but specific functions or sections have been lost, perhaps during enrichment.

Chloroform and 1,1,1-TCA, or methylchloroform, are potent inhibitors of homoacetogens and methanogens, well-known corrinoid producers in anaerobic environments (Stupperich et al., 1988;

Whitman and Wolfe, 1984). In addition, most corrinoid biosynthesis genes are missing from the dominant fermenting microbe, a Bacteroides sp., in ACT-3 (Tang et al., 2013; Tang et al., 2012).

Strain CF may have conserved its corrinoid-synthesizing ability due to environmental corrinoid shortage, while other corrinoid-auxotrophic strains lost the ability as a result of an abundant supply from other community members or enrichment in laboratory medium containing B12. For example, the β-hexachlorocyclohexane-dechlorinating strain E1 has only a partial cobinamide pathway and depends on the syntrophic relationship with a Sedimentibacter sp., which contains a complete cobinamide biosynthesis pathway in its genome (Maphosa et al., 2012). Moreover, PCE- dechlorinating strain PER-K23 possesses complete and functional cobinamide biosynthesis genes, except for a truncated CbiH gene, resulting in its corrinoid auxotrophy (Rupakula et al., 2015).

Thus 1,1,1-TCA and CF may indirectly contribute to the conservation of the corrinoid biosynthesis genes among D. restrictus genomes.

Heme biosynthesis. Heme (heme b) is a cofactor for cytochrome b, a subunit of Hup-type Ni,Fe- hydrogenase complex found in all five Dehalobacter genomes (Tang et al., 2016). The hup-type

Ni,Fe-hydrogenase is expressed during growth of strain PER-K23 (Rupakula et al., 2013) and in

Fig. 3.6. Heme production by strain CF and its influence on dechlorination. (A) Average dechlorination rates of strain CF cultures grown with/out heme supplementation. (B) UPLC-MS analysis of heme b standard (hematin form) and (C) heme in cell extract of strain CF. Chromatograms shown are extracted ion chromatograms (m/z=633.038 ± 0.0063) and the chemicals are ionized in negative mode. The * symbol in Fig. 3.5B is an unknown organic present in strain CF cell extract. Data are mean ± SE of three replicates in each experiment. all ACT-3 subcultures (Tang and Edwards, 2013), although the membrane-bound cytochrome b subunit was not detected, perhaps due to the limit of tryptic digestion-based proteomics. All five

Dehalobacter genomes were found to possess a complete archaeal heme biosynthesis pathway first characterized in the methanogenic archaeon Methanosarcina barkeri (Kuhner et al., 2014; Tang et al., 2016). Recently, another heme biosynthesis pathway in aerobic Firmicutes was characterized (Dailey et al., 2015). However, one of the biosynthesis enzymes, HemY, which converts coproporphyrinogen III to coproporphyrin III is an oxygenase and thus cannot function in strict anaerobic Firmicutes including Dehalobacter. Accordingly, the so-called “archaeal pathway” is still the only characterized pathway for anaerobic heme biosynthesis.

When heme (hematin form; 25 µg/L) was added to the medium, Dehalobacter strain CF dechlorinated faster than without heme addition (Fig. 3.6A). To confirm that the heme biosynthesis

67 pathway is functional in strain CF, cell extracts of culture grown in heme-free medium were analyzed by UPLC-ESI-MS. A peak with a retention time of 1.5 min and an m/z value of 633.0404 comparable to that of an authentic heme standard (m/z=633.0380) was identified (Fig. 3.6B,C), confirming that strain CF is able to synthesize heme, most likely via the identified archaeal pathway. Before this experiment, heme has never been included in the component used to make growth medium for Dehalobacter. Our experiment indicates that this “archaeal pathway” is not only functioning in methanogenic archaea but also in some anaerobic Firmicutes.

3.5 Implications for Microbial Ecology

In this study, experiments were conducted to refine or confirm predicted annotations in a strict anaerobe. These improved annotations enhance the accuracy of protein annotations, not only for Dehalobacter, but also for many other microbes, especially anaerobes, with similar pathways and genes. The study also explained several metabolic interdependencies in Dehalobacter- containing mixed microbial communities. Consistent with curated genome annotations, CF- respiring D. restrictus strains are capable of de novo synthesis of the corrinoid cofactor of RDases, which is in contrast to other D. restrictus strains where this ability has been lost. This study revealed that all sequenced Dehalobacter to date are capable of synthesizing heme (cofactor of

Ni,Fe-hydrogenase) for organohalide respiration. 1,1,1-TCA and CF, as inhibitors of many fermentative or acetogenic corrinoid producers, may indirectly contribute to the conservation of corrinoid biosynthesis genes in Dehalobacter genomes. Experimental verification of genome annotation is a laborious effort, particularly for fastidious microorganisms, yet critical to resolving inconsistencies, discovering new functions, and enabling genome-scale metabolic modelling in microbial communities. This study illustrates two aspects of the interpretation of genome

68 annotation that can improve the correspondence between bioinformatic predictions and the reality:

(i) cofactor availability for corresponding metabolic reactions and (ii) the potential for enzyme promiscuity to rescue apparently missing pathways. These findings reinforce the importance of incorporating cofactor availability for corresponding enzymes in genome-wide function prediction as metabolic reactions require that both functional cofactors and corresponding enzymes be present.

Moreover, biochemical characterization of poorly studied alternative metabolic pathways is urgently required to improve accuracy of genome annotations. Finally, the example of serine salvage by promiscuous SerB in strain CF suggests experimental verification is crucial to verify interpretations drawn from bioinformatic analyses.

Acknowledgements

Support was provided by the Government of Canada through Genome Canada and the

Ontario Genomics Institute (2009-OGI-ABC-1405), the Government of Ontario through the ORF-

GL2 program, and the United States Department of Defense through the Strategic Environmental

Research and Development Program (SERDP). S.T. received awards from the Government of

Ontario through the Ontario Graduate Scholarships in Science and Technology (OGSST) and the

Natural Sciences and Engineering Research Council of Canada (NSERC PGS B). We also acknowledge the BioZone Mass Spectrometry facility for UPLC-ESI-MS analyses as well as Dr.

Jane Howe and Mr. Xu Chen (University of Toronto, Canada) and Dr. Doug Holmyard (Mount

Sinai Hospital, Canada) for SEM and TEM analyses. We are grateful to the generous gift of

Lactobacillus delbrueckii subspecies lactis (ATCC 7831) from Agriculture Research Service

(USDA).

CHAPTER 4 – INTERSPECIES MALATE-PYRUVATE SHUTTLE DRIVES AMINO

ACID EXCHANGE IN ORGANOHALIDE-RESPIRING MICROBIAL

COMMUNITIES

This chapter is the result of a cooperative project with the Mahadevan group in BioZone. In fact, it stems from a proposal for an Ontario Genomics Spark grant that I contributed to writing and that was awarded in 2017. For the genome-scale metabolic model reconstruction, constraint incorporation, and simulation analysis, please refer to Kevin Correia’s forthcoming doctoral thesis.

A version of this chapter is in preparation to be submitted to the ISME Journal.

4.1 Abstract

Most microorganisms in the biosphere live in communities and develop coordinated metabolism by trading metabolites. In this study, we sought to deconstruct the metabolic dependency of organohalide-respiring Dehalobacter restrictus (Dhb), an active participant in global halogen cycle, using a complementary approach of genome-scale metabolic modelling and experimental validation. Dhb possesses a complete set of genes for amino acid biosynthesis yet requires amino acid supplementation (Chapter 3). We reconciled this discrepancy using flux balance analysis with consideration of cofactor availability, enzyme promiscuity, and shared protein expression patterns of several Dhb strains. Experimentally, 13C incorporation assays, growth assays, and metabolite analysis of Dhb strain PER-K23 cultures were applied to add additional model constraints. The model resolved that Dhb’s amino acid dependency results from restricted NADPH regeneration system and diagnosed that malate supplementation can replenish the NADPH pool. Interestingly, we observed unexpected export of amino acids and pyruvate in parallel to malate consumption in strain PER-K23 cultures. Further experiments on Dhb-enriched

70 consortium ACT-3 suggested an interspecies malate-pyruvate shuttle between Dhb and the syntrophic Bacteroides, reminiscent of the mitochondrial malate shunt pathway in eukaryotic cells.

Altogether, this study reveals that redox constraints and metabolic complementarity are the driving forces for amino acid exchange between anaerobic microorganisms.

4.2 Introduction

Prokaryotic microorganisms ubiquitously inhabit all the ecosystems on Earth in close association with one another. The study of microbial communities is gaining importance due to their essential contributions in global element cycling, agriculture, bioremediation, human health and industrial biotechnology (Dolfing, 2013; Embree et al., 2015; Mee et al., 2014; Zhao et al.,

2014). The interactions among microorganisms and their surroundings form phenotypes that can be observed in ecosystems, and are classified into three main categories: syntrophy, cross-feeding, and competition (Seth and Taga, 2014). Several methods have been proposed to elucidate these complex interactions, including microbial co-association network analysis as a function of time and other external factors (Cardona et al., 2016).

Metabolic complementarity is a dominant driving force for microbial mutualism (Mori et al., 2016; Wintermute and Silver, 2010). While mutualism confers robustness to microbial communities, a trade-off is that the coevolved microbes within a niche environment are more susceptible to lose non-essential function via genome streamlining, which can lead to auxotrophies in other environments (McCutcheon and Moran, 2012). As a result, the isolation of microorganisms from their syntrophic partners or natural niches is often challenging, as demonstrated by the scarcity of culturable isolates in the laboratory (~2%) (Wade, 2002). An alternative approach is metabolic modeling based on microbial genomes (Manor et al., 2014;

Roling and van Bodegom, 2014; Tan et al., 2015). Genome-scale constraint-based metabolic models have been increasingly used to elucidate metabolic networks at the community level

(Magnúsdóttir and Thiele, 2018; Zhuang et al., 2011). Nevertheless, physiological information and genome annotation verification for organisms in complex communities are often lacking, which results in the inclusion of misannotated gene and non-gene associated reactions (Suthers et al.,

2009), rendering simulations to be physiologically irrelevant. Integration of laboratory experiments with genome-scale metabolic modeling can significantly improve the accuracy of prediction (Amador-Noguez et al., 2010), as demonstrated in the study of syntrophic amino acid exchange in synthetic E. coli communities (Mee et al., 2014).

Anaerobic organic mineralization requires tightly coupled metabolic coordination between microbial community members due to redox constraints (McInerney et al., 2009; Sieber et al.,

2012). The presence of external electron acceptors enables more complex communities to develop where a variety of microbial populations can coexist. Organohalide-respiring microbial communities is a great model to study metabolic interdependency between microbes, which is inhabited by acetogens, fermenting bacteria, methanogens, sulfate-reducing bacteria, and organohalide-respiring bacteria (OHRB) (Adrian and Loeffler, 2016; Duhamel and Edwards,

2007). Dehalobacter restrictus (Dhb) strains are specialized in respiring a variety of organohalide including the chloromethane species (Holliger et al., 1998; Justicia-Leon et al., 2012; Tang et al.,

2016; van Doesburg et al., 2005; Wang et al., 2014; Wong et al., 2016; Yoshida et al., 2009), the most abundant organohalide species in the atmosphere (Harper, 2000; Laturnus et al., 2002). Dhb isolate cultures, with the exception of strain UNSWDHB, require the addition of either complex of amino acids or parent culture supernatants to support growth (Holliger et al., 1998; Wang et al.,

2016), indicating unexplored nutrient cross-feedings from symbiotic partners in natural habitats,

72 e.g. the Bacteroides sp. CF (Bac) in chloroform-dechlorinating consortium ACT-3 (Tang and

Edwards, 2013). However, comparative genomic analysis and a refined metabolic annotation suggested that Dhb possesses a complete set of genes to synthesize all amino acids, including a salvage pathway to obtain serine from threonine (Tang et al., 2016; Wang et al., 2016).

In this study, we explored Dhb’s metabolic dependency using a genome-scale constraint- based metabolic model. The model was built based on a highly annotated Dhb genome (Wang et al, 2016), along with shared expression patterns in proteomic datasets of Dhb strains PER-K23,

UNSWDHB, DCA and CF (Jugder et al., 2016; Rupakula et al., 2013; Rupakula et al., 2015; Tang and Edwards, 2013). The model simulation considers cofactor availability, enzyme promiscuity, physiological redox conditions, and further model constrains based on experimental values obtained in this study. The model resolved that Dhb’s amino acid dependency results from the restrictions in NADPH regeneration, which can be restored with malate supplementation. The strain PER-K23 cultures grown on the model-resolved medium exhibited an unexpected export of pyruvate, glutamate, and other amino acids in parallel to malate consumption. Further experimental analysis on Dhb strain CF-enriched consortium ACT-3 revealed that Dhb’s specialized malate requirement is likely a consequence of genome streamlining, which was driven by the co-adaption with a glutamate-auxotrophic, malate-producing Bacteroides to accomplish interspecies exchange of reducing equivalents (i.e. NADH and NADPH).

4.2 Materials and Methods

All Chemicals were ordered from Sigma-Aldrich at highest purity available unless specified otherwise.

Microbial cultures and growth conditions. Dhb strain PER-K23 was provided by the Löffler

Lab at University of Tennessee (Knoxville, USA). Escherichia coli strain BL21(DE3) Gold was purchased from New England Biolabs Ltd. Consortium ACT-3 was originally enriched from 1,1,1- trichloroethane-contaminated groundwater in 2001 from a northeastern United States industrial area (Grostern and Edwards, 2006), and a subculture (1.8 L) was adapted to respire chloroform

(Chapter 1.1) (Grostern et al., 2010). E. coli strain BL21 was grown on LB broth except in the

13C incorporation assay. The strain PER-K23 cultures and the consortium ACT-3 are maintained in a FeS-reduced, bicarbonate-based mineral medium described previously (Grostern et al., 2010;

Wang et al., 2016), except that trichloroethene (TCE) was used as the electron acceptor for strain

PER-K23 cultures. The strain PER-K23 growth assay was performed following the established protocol reported in Chapter 3. Briefly, the inoculum (1 mL) from mother cultures was inoculated to sterile rubber stopper-sealed serum bottles (160 mL) containing the mineral medium (99 mL) under a N2/CO2 atmosphere (80%:20%; v/v), along with 5 mL of H2 (80%:20%; v/v) as the electron donor and 1 mM TCE (nominal concentration) as the electron acceptor. Besides the 13C incorporation assay, Acetate (1 mM), malate (2 mM), and/or serine (2 mM) were used as the carbon sources to support strain PER-K23 growth. When TCE is depleted, the cultures were sparged with N2/CO2 to extinct cDCE, and re-feed TCE and H2.

For the time-course LC-MS metabolite analysis of CF-fed ACT-3, multiple feeds of CF was given to deplete remaining electron donor before the analysis, followed by a N2/CO2 sparging to remove remaining chlorinated solvents, and a pH adjustment with bicarbonate to neutral pH.

After that, lactate (1 mM) and 30 µL of CF (0.21 mM nominal concentration) were fed to the culture. The culture was sampled right before and 2 h after substrates were given, and was sampled every two other days. For the ACT-3 syntrophy-decoupling assays, in an anaerobic chamber (Coy),

74 the inoculum (5 mL) from mother cultures were firstly pelleted down using a 10 min centrifugation at 13,000 x g and room temperature. The supernatant was removed, washed by sterile mineral medium twice, and the pellet was resuspended by 5 mL of sterile mineral medium. The suspensions

(0.5 mL each) were inoculated into the sterile rubber stopper-sealed serum bottles (40 mL) containing the mineral medium (19.5 mL) under a N2/CO2 atmosphere (80%:20%; v/v), along with

0.25 mM lactate and/or 0.5 mM malate as the electron donor as well as 0.5 mM CF (nominal concentration) as the electron acceptor. Heme (hematin form) was dissolved in 20 mM anaerobic

NaOH, and was added to the heme+ bottles to a final concentration of 1 mg/L. The sub-transfer cultures were sample for gas chromatography analysis every 6 days and was sampled for LC-MS metabolite analysis at Day 12. After 18 days, the cultures were harvested for DNA extraction and

16S-rRNA-based population analysis.

13C incorporation assay. The 13C incorporation assay was performed following a previous study on amino acid biosynthesis of Dehalococcoides with some modifications (Zhuang et al., 2014).

[3-13C]pyruvate (5 mM), and acetate (1 mM), along with 0.1 mM of arginine, histidine, and threonine, were used as the carbon sources to support growth of strain PER-K23 and E. coli except that PER-K23 cultures were continuously fed with TCE. The E. coli cultures (50 mL) were harvested after an overnight incubation. The strain PER-K23 cultures (200 mL) were harvested after consuming 5 mM TCE and centrifuged at 16,000 x g for 10 min at 4oC. The pellets were washed twice with mineral medium, resuspended by 6N HCl (0.2 mL for Dhb pellets; 1 mL for E. coli pellets), transferred to a 2.2 mL O-ring-capped microcentrifuge tubes and incubated at 100oC for 24 h. After cooling to room temperature, the yellow-colored digests were centrifuged at 16,000 x g for 20 min at 4oC to remove cell debris, and the supernatants were transferred to 2.2 mL microcentrifuge tubes, followed by an overnight incubation at a rotary evaporator at room

75 temperature. The dried pellets were resuspended with 0.1 % formic acid (pH 2.0; 0.2 mL for Dhb pellets; 0.5 mL for E. coli pellets), and stored at -80oC before LC-MS analysis.

DNA sampling and quantitative PCR. Culture DNA was extracted from 2 mL samples. Cells were harvested by centrifugation at 16,000 x g for 10 min at 4°C. Since Dhb cell pellets are easily resuspended, in each tube, most (but not all) of supernatant was gently removed (1.9 mL), and the cell pellets were resuspended using the remaining supernatant (0.1 mL), and the DNA was extracted using the MO BIO PowerSoil® DNA isolation kit following the manufacturer's recommendations. Real-time quantitative polymerase chain reaction (qPCR) assays were performed to track the gene copy numbers of Dhb using specific 16S rRNA gene primers reported previously (F: 5’-GAT TGA CGG TAC CTA ACG AGG-3’; R: 5’-TAC AGT TTC CAA TGC

TTT ACG-3’) (Grostern and Edwards, 2006). The PCR preparation was conducted in a UV-treated

PCR cabinet (ESCO Technologies, Hatboro, PA). The 20 μL reaction mixtures contained 10 μL of 2 × SsoFast EvaGreen® (Bio-Rad, Hercules, CA), forward and reverse primers (0.5 μM each), and 2 μL of template DNA (diluted 1:10). The amplification program included an initial denaturation step at 98°C for 2 min, followed by 39 cycles of 5 s at 98°C and 10 s at the corresponding annealing temperature of each primer set. Quantification was performed using 10 times serial dilutions of plasmid DNA as standards. The plasmid contains a cloned Dhb 16S rRNA gene (Grostern and Edwards 2006). The analyses were conducted using a BIO-RAD CFX96 Touch

Real-Time PCR Detection System and the CFX Manager software. The number of copies per mL of culture was calculated assuming a 100% DNA extraction efficiency.

Enzyme activity assays. The conditions for cell extract preparation and RDase activity assays were described in Chapter 3. O-phosphoserine phosphatase activity assay were conducted following established methods with some modifications (Kuznetsova et al., 2006). Briefly, the

76 assay mixture (0.1 mL) contains O-phosphoserine (2.5 mM), Mg2+ (2.5 mM), Tris-HCl (100 mM; pH 7.5), and cell extracts to a final concentration of 50 µg/mL. The assays were incubated at 30oC for 1 h, and filtered with 3 kDa spin filters. The filtrates were stored at -80oC before LC-MS analysis. Protein concentrations of cell extracts were determined using the standard Bradford assay.

Specific activity is reported as µmol of serine produced per min per mg of total proteins.

Analytical procedures. Chlorinated hydrocarbons were measured by injecting a 0.3 mL headspace sample into a Hewlett-Packard 5890 Series II GC fitted with a GSQ column (30-m-by-

0.53-mm [inner diameter] PLOT column; J&W Scientific, Folsom, CA) as described previously

(Wang et al, 2016).

For the metabolite profile analysis, in an anaerobic chamber (Coy), each culture was harvested (0.2 mL), and gently filtered through a 0.1 µm-pore-size syringe filter (Millipore). The flow-through was collected in a plastic microcentrifuge tube, followed by a centrifugation at

16,000 x g for 10 min at 4oC. The supernatants were stored at -80oC before analysis. The amount of acetate, malate and pyruvate from Dhb strain PER-K23 cultures was determined by HPLC using a ICS5000 system (Thermo scientific) equipped with an Aminex HPX-87H column (BioRad) connected to a UV detector. Each sample (25 µL) was injected onto the column incubated at 35oC, using 5 mM H2SO4 eluent at a flow rate of 0.6 mL/min with the UV wavelength set to 210 nm.

Amino acids and organic acids were detected using Liquid chromatography Electrospray- coupled high resolution mass spectrometry (LC-ESI-HRMS) with a Dionex UHPLC system and a

Q-Exactive mass spectrometer (Thermo Scientific) equipped with a HESI II source (Thermo

Scientific) and a Micro-splitter valve (IDEX Health & Science). System control and data handling were performed using Thermo XCalibur 3.1 software. For amino acid detection, separation by liquid chromatography was conducted on a Luna NH2 column (50mm x 2.0 mm, 5 µm particle

77 size, Phenomenex). LC was performed with 10 uL injections at a flow rate of 0.3 ml/min with a gradient of acetonitrile containing 0.1% formic acid (pH 2) (A), into water containing 0.1% formic acid (B), and a column temperature of 40˚C. The gradient was 0 min, 20% B; 1.5 min, 20% B; 3.5 min, 90% B; 8.0 min, 90% B; 10 min, 20% B; followed by equilibration for 5 min with 20% B.

Data collection was done in positive ionization mode with a m/z scan range of 100-250 with a detection limit of 10 nM; resolution 140,000 at 1 Hz, automatic gain control (AGC) target of 3e6; and a maximum injection time of 200 ms.

For organic acid detection, separation by liquid chromatography was conducted on an

Aminex HPX-87H column (300mm x 7.8 mm, 9 µm particle size, Bio-Rad). LC was performed with 20 uL injections at a flow rate of 0.6 ml/min with water containing 0.1% formic acid (pH 2) under isocratic conditions for 20 min, and a column temperature of 60˚C, with the Micro-splitter set to deliver 200 uL/min to the mass spectrometer. Data collection was done in negative ionization mode with a m/z scan range of 50-250 with a detection limit of 50 nM; resolution 140,000 at 1 Hz, automatic gain control (AGC) target of 3e6; and a maximum injection time of 200 ms.

4.3 Results and Discussion

Summary of experimental data integration with genome-scale metabolic model

The primary Dhb metabolic model was built by Cleo Ho using the annotated Dhb genomes, including 625 genes, 1085 reactions, and 992 metabolites, followed by automated metabolic network reconstruction using the SEED Servers (http://blog.theseed.org/servers/). Subsequently, the primary model was curated with refined annotation in TCA cycle, NADPH regeneration and biosynthesis of amino acid and cofactors (Wang et al., 2016). The gap in classical serine biosynthesis pathway was experimentally confirmed in this study (described in Alternative serine

78 biosynthesis in Dhb via threonine). After that, we modified the model based on five considerations: (i) cofactor availability for the corresponding metabolic reactions; (ii) shared features in available proteomes of strains PER-K23 and UNSWDHB and partial proteomes of consortium ACT-3 that is enriched with Dhb strains CF and DCA; (iii) cellular redox state under given growth conditions; (iv) potential promiscuous enzyme activity to rescue missing pathways; and (v) integration of experimental data from culture growth assays and metabolite profile analysis.

Table 4. 1. Stepwise curation of genome-scale constraint-based Dehalobacter metabolic model

Model curation Rationale Exclude non-gene- and mis- Default Integration of refined genome annotation annotated gene-associated reactions

1 Inactivate formate dehydrogenase Consideration of cofactor availability (MoCo)

2 Inactivate pyruvate formate-lyase Absence in all available Dhb proteomes

Prevent NADPH generation from 3 Absence in all available Dhb proteomes NADP-reducing hydrogenase Prevent NADPH generation from Absence in all available Dhb proteomes 4 ferredoxin-NADP reductase except strain UNSWDHB Set 5,10-methylenetetrahydrofolate 5 reductase reaction monodirectional Consideration of cellular redox homeostasis (CH3-THF→CH2=THF) Inactivate threonine aldolase 6 Consideration of enzyme promiscuity activity when serine is present Set pyruvate synthase reaction 7 monodirectional (acetyl-CoA→ Consideration of cellular redox state pyruvate) Incorporate experimental substrate 8 Integration with experimental data consumption/metabolite production Prevent the export of undetected 9 Integration with experimental data metabolites

Based on these considerations, we applied nine constraints to the model in a stepwise fashion

(Table 4.1) by the following four actions: (a) inactivating targeted reactions, (b) limiting the direction of targeted reactions (set monodirectional), (c) applying experimentally relevant metabolite flux, and (d) disabling the export of undetected metabolites. The resulting model reveals a major defect in NADPH regeneration, which can be restored by malate supplementation through the function of NADP-dependent malic enzyme. Another defect is the predicted FRDred accumulation during H2 utilization, which disables the reverse reaction of FRD-dependent pyruvate synthase for acetyl-CoA biosynthesis, rendering acetate supplementation essential. The following sections describe the experimental validation of these model predictions.

Alternative serine biosynthesis in Dhb via threonine

In Chapter 3, we elucidated Dhb’s ability to synthesize serine from threonine using the promiscuous serine hydroxymethyltransferase (GlyA; EC 2.1.2.1) which possesses threonine aldolase activity and is expressed in all available Dhb proteomes. However, an unsolved question is if the gene for o-phosphoserine phosphatase (SerB; EC 3.1.3.3) (Fig. 4.1A) is indeed truly missing from Dhb genomes. Also, some unspecific phosphatases in Dhb proteomes may dephosphorylate o-phosphoserine to serine. Thus, we examined the growth of strain PER-K23 cultures with only acetate (3 mM) or with acetate and serine (1 mM) as carbon sources.

Dechlorination was only observed in the cultures supplemented with serine (Fig. 4.1B). We then examined potential promiscuous phosphatase activity in Dhb cell lysates. While we observed PceA activity in assays containing Dhb cell lysates, o-phosphoserine dephosphorylation activity was only observed in the assays containing E. coli cell lysates (positive control 2)

SerB SerB

course course

(C)

Time

(B)

n orange. orange. n

A activity was used as a (+)

, phosphoserine; SerA, SerA, phosphoserine; ,

C]pyruvate, C]pyruvate, unlabelled acetate, and

free free control. Pce

C]pyruvate C]pyruvate are shown in red and the three carbons

lysate

K23 K23 using [3

K23 cultures grown on unlabeled pyruvate or [3 or pyruvate unlabeled on grown cultures K23

Schematic Schematic of amino acid biosynthesis (alanine, aspartate,

(A)

) ) control, cell

C]pyruvate as a (+) control. Abbreviations: AC, acetate; AKG, AKG, acetate; AC, Abbreviations: control. (+) a as C]pyruvate

alanine for strain PER strain for alanine

via via threonine.

(iv)

labelled labelled carbons originating from [3

strain strain BL21, respectively. (

K23 cultures grown on defined medium with different combination of carbon sources. sources. carbon of combination different with medium defined on grown cultures K23

- isotopome mass different of abundance Relative

glutamate, and and glutamate,

(D)

C incorporation in amino acids of strain PER

(iii)

E. E. coli

Dehalobacter Dehalobacter restrictus

aspartate, aspartate,

cell lysate. lysate. cell

K23 K23 and

(ii)

Dhb

methylenetetrahydrofolate; FRD, ferredoxin; MAL, malate; OAA, oxaloacetate; SER oxaloacetate; OAA, malate; MAL, ferredoxin; FRD, methylenetetrahydrofolate;

serine, serine,

5,10

(i)

strain BL21 was grown on the same medium with [3 with medium same the on grown was BL21 strain

=THF, =THF,

E. coli E.

lene (TCE) dechlorination by strain PER strain by dechlorination (TCE) lene

. . Alternative serine biosynthesis in

C]pyruvate. C]pyruvate.

ketoglutarate; CH ketoglutarate; Fig. Fig. 4. glutamate, and serine; in dashed boxes) and unlabelled threonine (in blue) as the precursors. i shown The is biosynthesis serine classical in SerB by catalyzed step missing The cyan. in highlighted are pyruvate from derived trichloroethy activity in cell lysates of strain PER of quality the examine to control of M2) and M1, (M0, rs THF aminotransferase; phosphoserine SerC, phosphatase; phosphoserine SerB, dehydrogenase; phosphoglycerate tetrahydrofolate. , 13 81

(Fig. 4.1C). These preliminary results indicate that the classical serine biosynthesis pathway

(Greenberg and Ichihara, 1957) is not present in Dhb.

Subsequently, we used a 13C incorporation approach to elucidate serine biosynthesis in

Dhb. [3-13C]pyruvate or unlabeled pyruvate (5 mM), a precursor of serine in the classical pathway

(Fig. 4.1A) (Grundy and Henkin, 2002), was supplemented to the defined medium (containing 1 mM acetate and 0.1 mM of arginine, histidine, and threonine) to grow strain PER-K23 cultures. If

Dhb synthesizes serine via the classical pathway, the MS analysis of isotopomer distribution of serine in positive mode would reveal an enrichment in relative abundance of the [M1+H] adduct.

In contrast, if Dhb synthesizes serine via threonine, the [M1+H] adduct of serine would remain at natural abundance. After the cultures consumed 5 mM TCE, we harvested the cells and hydrolyzed the cellular proteins for LC-MS analysis of amino acids. The 13C-labeling profiles of alanine, aspartate, and glutamate from [3-13C]pyruvate-fed strain PER-K23 culture revealed a 10% increase in the relative abundance of their [M1+H] adducts (Fig. 4.1Dii-Div; red bars). However, the

[M1+H] adduct of serine from both unlabeled pyruvate- and [3-13C]pyruvate-fed strain PER-K23 cultures are comparable, while that from the [3-13C]pyruvate-fed E. coli culture (positive control) reveals a significant enrichment (~70%) (Fig. 4.1Di), suggesting that Dhb converts the unlabeled threonine to serine. Unlike E. coli, Dhb only uses pyruvate as carbon source but not as electron donor and requires supplementation of other carbon sources for growth. Therefore, the 13C incorporation in Dhb amino acids is not as significant as that in E. coli amino acids, but is distinguishable to the unlabeled amino acids. Altogether, based on genome annotation and multiple lines of experimental evidence, the classical serine biosynthesis pathway is absent in Dhb, suggesting that serine is synthesized using the threonine salvage pathway. Interestingly, this

82 salvage pathway could be a major route for serine biosynthesis in all serB-lacking Firmicutes and

Geobacter spp. (Sung et al., 2006).

Resolving redox balancing in Dhb metabolic model

The absence of classical serine biosynthesis pathway in Dhb can result in many metabolic defects because serine is the donor to the cellular C1 pool and the precursor of purines, methionine, cysteine, glycine, and tryptophan (Fig. 4.2A) (Fan et al., 2014). Recent studies also discovered that serine is an important NADPH source (Fan et al., 2014; Tedeschi et al., 2013). Furthermore, unlike the biosynthesis of other amino acids, the biosynthesis of serine and serine-derived amino acids (cysteine and glycine) through the classical pathway does not require NADPH (Table 4.2)

(Grundy and Henkin, 2002). In contrast, using threonine to synthesize serine would elevate cellular demand of NADPH for protein synthesis by approximately 30% (please refer to Kevin Correia’s thesis), assuming a protein composition resembling Bacillus subtilis, which directed our interest to examine NADPH regeneration systems in Dhb.

Table 4. 2. Number of NADPH molecules required per amino acid synthesized. First number is for classical serine biosynthesis pathway; second number is for alternative pathway from threonine.

Thr Lys His Iso Val Leu Phe Tyr Trp Pro Met Arg Glu 2 3 2 4 4 3 2 2 4/6 3 4/6 4 1 Gln Ser Gly Cys Asp Asn Ala 1 0/2 0/2 0/2 1 1 1

Fig. 4.2. Resolving redox balancing in Dehalobacter restrictus metabolic model with integration of experimental data. (A) Proposed central carbon metabolism and redox balancing system in D. restrictus. The name of enzymes and the missing cofactors (in square bracket) are shown in blue. The red crosses (X) represent the missing genes or missing cofactors in the D. restrictus genomes. NAD(P)+/NAD(P)H are shown in red beside the corresponding metabolic reactions. The black arrows represent the metabolic reactions involved in NADPH and ferredoxin regeneration, while grey arrows represent reactions where they are not involved. The double-headed arrows indicate reversible reactions, and the bigger arrowheads represent the direction of reactions under proposed physiological conditions. (B) Three consecutive 1% dilution transfers of strain PER-K23 cultures grown on defined medium with malate and serine as the carbon sources. H2 and trichloroethene (red ●; TCE) are used as the sole electron donor and the final electron acceptor, respectively. cis-1,2-dichloroethene (blue ●; cDCE) is the dechlorination product. (C) TCE dechlorination (blue ●) and cell growth (brown ■) of the strain PER-K23 cultures grown on the same medium as in in (B). When TCE is depleted, the cultures were purged with H2/CO2 to remove cDCE, and re-feed TCE and H2 (red asterisk). Acetate (1 mM) was fed to the cultures after the fourth feeding. (D) Consumption of malate (●) and serine (■) as well as production of pyruvate (▼) in the strain PER-K23 cultures shown in (C). Metabolite profile of the killed control is shown in black. Abbreviations: CODH, carbon monoxide dehydrogenase; FDH, formate dehydrogenase; MAE, malic enzyme; MTHFD, 5,10- methylenetetrahydrofolate dehydrogenase; MTHFR, 5,10-methylenetetrahydrofolate reductase; MDH, malate dehydrogenase; SDH, succinate dehydrogenase; SerB, phosphoserine phosphatase; SerB, bifunctional serine hydroxymethyltransferase also possessing threonine aldolase activity (THR → GLY); THF, 5,10-tetrahydrofolate; WL pathway, Wood- Ljungdahl pathway.

According to genome annotation, seven potential enzyme reactions can contribute to

+ NADPH regeneration (Figure 4.2A), including a putative FRDred:NADP oxidoreductase (NfnAB;

EC 1.6.1.4); a putative NADP+-reducing hydrogenase (HndABCD; EC 1.12.1.3); isocitrate dehydrogenase (Idh; EC 1.1.1.42), NADP-dependent malic enzyme (MAE; EC 1.1.1.40); molybdopterin (MoCo)-dependent NADP+-specific formate dehydrogenase (FdhAB; EC 1.2.1.43); and 5,10-methylenetetrahydrofolate reductase/ dehydrogenase (MetF; EC 1.5.1.20; FolD; EC

1.5.1.5). Since Dhb cannot synthesize MoCo (Wang et al., 2016), FdhAB cannot function. Also, given that Dhb employs the TCA cycle and folate cycle for anabolism but not for respiration, the generated NADPH from Idh, MetF, and FolD is insufficient to support the anabolism using simple carbon source like acetate. Finally, the soluble HndABCD was not expressed in all the available

Dhb proteomes; NfnAB is only expressed in strain UNSWDHB proteome. Consistently, strain

UNSWDHB was reported to grow on acetate as the sole carbon source (Wong et al., 2016) and was isolated from an acetate/H2-fed enrichment, while strains PER-K23, CF, and DCA were isolated from lactate-fed enrichment cultures (Grostern and Edwards, 2006; Holliger et al., 1993).

Accordingly, the difference in NfnAB expression among Dhb strains is likely a result of niche specialization driven by bacterial epigenetic change (i.e. DNA methylation) responding to environmental conditions (Casadesús and Low, 2006). Moreover, the lack of NfnAB expression in the other Dhb strains indicates the availability of amino acids or the nutrients enabling Dhb to regenerate NADPH in the lactate-fed enrichment cultures.

Nevertheless, the respiration-independent MAE can replenish the NADPH pool with malate supplementation. Also, Dhb genomes possess malate permease (maeP) to uptake malate; the MAE was significantly expressed in all the proteomes of Dhb strains. Thus we examined if malate supplementation can support Dhb growth on the acetate-based medium. Consistently, the

85 strain PER-K23 cultures supplemented with malate showed a three-fold faster dechlorination rate

(60 µM bottle/day) than that of the cultures supplemented with serine (20 µM bottle/day) (Figure

4.1B). Supplementing both malate and serine to the strain PER-K23 cultures resulted in the highest dechlorination rate (100 µM bottle/day) (Figure 4.1B), suggesting that the missing serine pathway and restricted NADPH regeneration system are associated with Dhb’s amino acid dependency.

Unexpected export of pyruvate in Dhb isolate cultures

To examine the hypothesis of restricted NADPH regeneration in Dhb, we performed three consecutive 1% transfers of strain PER-K23 cultures using acetate-free defined medium supplemented with both malate (2 mM) and serine (2 mM). Strain PER-K23 cultures were able to sustainably deplete 1 mM TCE after three consecutive 1% transfers (Figure 4.2B). We then examined long-term growth of strain PER-K23 cultures using this medium. The cultures readily depleted three feedings of 1 mM TCE (Figure 4.2C). qPCR analysis revealed that Dhb cell density in the cultures increased by 100-fold after the consumption of 3 mM TCE (~2 x 107 cells/mL). The cell yield (6.3 ± 0.02 x 1012 cells/mol Cl- released) is in good agreement to the qPCR-determined yield of strains CF and UNSWDHB reported previously (Grostern et al., 2010; Wong et al., 2016).

However, after the fourth feeding of TCE, there was a significant drop in the dechlorination rate

(Figure 4.2C). Before each feed, the cultures were purged with N2/CO2 to remove accumulated cDCE, to adjust the pH, and to replenish the medium with H2. Moreover, the metabolite profile of culture supernatants revealed that most malate and serine (~90%) were not consumed (Figure

4.2D), suggesting that the lagging dechlorination is not due to the shortage of electron donor/carbon sources, accumulation of electron acceptor, or acidic pH. Unexpectedly, the metabolite profile of strain PER-K23 culture supernatants revealed the production of pyruvate in

86 parallel to substrate consumption, likely derived from malate and serine through the function of

MAE and serine deaminase (EC 4.3.1.17) expressed in Dhb proteomes. This unexpected pyruvate export suggests an unfavourable sink for pyruvate in Dhb. Indeed, the missing of malate and succinate dehydrogenase (or fumarate reductase) genes prevents Dhb from fermenting pyruvate to succinate and the absence of a classical serine biosynthesis pathway prevents pyruvate from entering the folate cycle via serine (Figure 4.2A). Furthermore, from a thermodynamic point of view, MAE-mediated malate decarboxylation is unfavorable under standard conditions (malate +

+ o NADP → Pyruvate + CO2 + NADPH; eQuilibrator-estimated ∆rG’ = 14 ± 6.2). Therefore, Dhb likely exports pyruvate to drive the malate-dependent NADPH regeneration.

Deconstruction of Dhb metabolism using complementary flux balance analysis and experimental validation

We then constrained the model with the experimental values of cell yields, malate/serine consumption, and pyruvate export to simulate Dhb metabolism. Interestingly, the resulting model revealed excessive NADPH accumulation through the MAE reaction, unless we allowed (a) the export of other organic acids to prevent NADPH production via Idh or (b) the export of other amino acids to consume NADPH (Figure 4.2A). Consistently, the experimentally measured metabolite profiles of strain PER-K23 culture revealed apparent production of glutamate, valine, glycine, leucine, aspartate, and alanine against the killed control, which is in accordance to the model predictions (Appendix E). When we further applied the experimental values of amino acid export, the resulting model suggested CO export. Since we purged the cultures before each feed,

CO would not be expected to accumulate to cause cell toxicity. Instead, CO production via FRD- dependent CO dehydrogenase (CooS; EC 1.2.7.4) indicates FRDred accumulation. In the model,

FRDred is mainly produced from H2-oxidizing Ech hydrogenases expressed in all available Dhb

87 proteomes (Figure 4.2A). FRDred accumulation will inhibit pyruvate conversion to acetyl-CoA using pyruvate-ferredoxin oxidoreductase while enhancing acetyl-CoA conversion to pyruvate.

Since acetyl-CoA is an essential substrate for citrate synthase in the TCA cycle, the lack of acetyl-

CoA would disable citrate synthesis and prevent pyruvate from entering the TCA cycle for glutamate production. Therefore, the model likely predicted CO and pyruvate export to consume excessive FRDred, enabling acetyl-CoA production. Consistent with this scenario, acetate supplementation (1 mM) to the PER-K23 cultures restored the lagging dechlorination (Figure

4.2C), and was accompanied by a significant production of pyruvate production (0.3 mM) in parallel to malate (0.25 mM) and serine (0.1 mM) consumption, supporting the hypothesis of

FRDred accumulation in Dhb. The PER-K23 cultures were able to deplete remaining TCE and another feeding in 10 days, confirming the model prediction that acetate supplementation can ameliorate the imbalanced redox with H2 as the electron donor. Since the cell yield of Dhb grown on TCE (6.3 x 1012 cells/mol Cl- released) is little, strain PER-K23 cells initially may use the acetate contaminated in the medium components (≤ 100 µM). Altogether, our data suggest that both acetate and malate are required to support the minimal growth of Dhb, and serine supplementation can reduce the metabolic burden in Dhb anabolism (i.e. NADPH cost).

In this study, our integrated approaches have resolved the metabolic dependency of Dhb, explaining previously identified arginine and threonine requirements of strain PER-K23 cultures

(Holliger et al., 1998). Due to the lack of serB, Dhb utilizes threonine to synthesize serine and its derived amino acids. Also, due to the lack of a carbon source-independent NADPH regeneration system, arginine is degraded to malate via fumarate to generate NADPH (Cunin et al., 1986). In contrast, when malate is abundant in the medium, Dhb becomes an amino acid producer, exporting amino acids to consume the excessive NADPH. However, since amino acid synthesis also requires

88 considerable ATP and NADH, Dhb also consumes the conserved energy obtained from organohalide respiration to produce and to export amino acids, as demonstrated by its significantly lower cell yield than that of Geobacter lovleyi grown on TCE (~1 × 1014 cells/mol Cl- released)

(Duhamel and Edwards, 2007). Compared to Dhb, other OHRB possess a more flexible system in

NADPH metabolism. For example, the classical serine biosynthesis pathway is present in chloroethene-respiring Dehalococcoides spp. based on the 13C incorporation experiments, and an incomplete Wood–Ljungdahl pathway can support NADPH regeneration (Zhuang et al., 2014).

PCE-respiring Geobacter lovleyi, while lacking serB, possesses a functional TCA cycle to respire acetate and generate sufficient NADPH using Idh (Galushko and Schink, 2000; Sung et al., 2006).

PCE-respiring Desulfitobacterium hafniense, in contrast to its close relative Dhb, retains the malate dehydrogenase and fumarate reductase genes in the TCA cycle, which allows pyruvate conversion to malate for NADPH regeneration via MAE (Peng et al., 2012).

Interspecies malate-pyruvate shuttle enables intercellular NADH/NADPH exchanges in

Dehalobacter–containing consortium ACT-3

After characterizing Dhb’s metabolic dependency, we wondered if these nutrients are available in Dhb-enriched microbial communities. Acetate is the final respiration product of acetogenic and fermenting bacteria that often co-exist in organohalide-respiring consortia

(Heimann et al., 2006), while the presence of malate or serine has not been reported before.

Therefore, we analyzed the metabolite profile of the CF-fed ACT-3 consortium in parallel to CF dechlorination. After lactate and CF were fed, the time-course metabolite profile revealed malate production (~0.5 µM) in the ACT-3 supernatant along with CF dechlorination to dichloromethane

(Figure 4.3A). Malate was not detected until the added lactate was depleted, and malate was not

Fig. 4.3. Organic acid profile in lactate-fed consortium ACT-3 and syntrophy disruption with heme addition. (A) Time-course organic acid profile of the CF-dechlorinating ACT-3 consortium. (B) Dechlorination profile of lactate-fed ACT-3 sub-transfer cultures grown with and without the addition of heme (1 mg/L). (C) Organic acid profile of ACT-3 sub-transfer cultures at Day 12. Abbreviation: DCM, dichloromethane; LAC, lactate; MAL, malate; PYR, pyruvate. detected in the 1 M lactate stock by LC-MS analysis (< 50 nM). Therefore, malate is a natural substrate for Dhb strain CF in the ACT-3 culture, which explains the presence of malate permease gene in Dhb genome and the consistent MAE expression in the ACT-3 proteomes. However, only trace amounts of amino acids and pyruvate (<100 nM) was present, and their concentration remained unchanged (Figure 4.3A), suggesting the presence of amino acid and pyruvate consumers in the mixed culture ACT-3.

To date, fermentative serine production has only been reported in aerobic methylotrophs

(Kubota et al., 1972; Tani et al., 1978). Nevertheless, fermentative malate (or fumarate) production by Bacteroides spp. has been studied extensively (Chen and Wolin, 1981; Macy et al., 1975; Macy et al., 1978; Miller, 1978). Due to the lack of a heme biosynthesis pathway, Bacteroides spp. only ferment lactate/ethanol to acetate, H2, and malate (or fumarate) (Chen and Wolin, 1981), unless exogenous heme is present to support further malate fermentation to succinate. Coincidently, based on available 16S rRNA pyrotag sequences (accession SRX181448), the most abundant organisms in the lactate-fed, CF-dechlorinating ACT-3 are Dhb strain CF (≥80%) and a Bacteroides strain

(Bac) (≥10%) (Grostern et al., 2010; Grostern and Edwards, 2006; Tang et al., 2012).

Fig. 4.4. Proposed syntrophy and genome streamlining between Dehalobacter restrictus strain CF and the symbiotic Bacteroides sp. (A) Proposed schematic diagram of syntrophic lactate degradation and interspecies malate (MAL)-pyruvate (PYR) shuttle. In the simplified model of ACT-3 consortium, lactate (LAC) is used as the sole electron donor and carbon source, and chloroform (CF) is used as the final electron acceptor. Malate dehydrogenase and fumarate reductase genes are missing from strain CF genome. The heme biosynthesis pathway is missing in the annotated genome of the syntrophic Bacteroides sp. (B) Genome streamlining between the symbiotic Bacteroides sp. (Bac) and Dehalobacter (Dhb) demonstrated by the complementary genes in the TCA cycle and in malate metabolism. Abbreviation: AC, acetate; Ac-CoA, acetyl- CoA; BUT, butyrate; CAP, caproate; DCM, dichloromethane; FUM, fumarate; FRD, ferredoxin; OAA, oxaloacetate; PRO, propionate; SUC, succinate. The closed genome of Bac (accession CP006772) reveals multiple genes of H2-producing hydrogenase (Hyf; EC 1.12.1.4) (Trchounian and Trchounian, 2013) and the genes in the TCA

91 cycle for lactate fermentation to succinate (Tang and Edwards, 2013), but lacks most genes for heme biosynthesis. Moreover, citrate synthase, aconitase, and Idh genes in TCA cycle are missing from the Bac genome, which disrupts Bac to synthesize glutamate (Figure 4.4A).

Since heme is insoluble and was never added to the growth medium of the ACT-3 mixed culture, we propose that lactate is first fermented to acetate, H2, and malate by Bac (Figure 4.4A).

Besides that used for biomass, acetate and malate are taken up by Dhb for NADPH regeneration and amino acid synthesis, while the produced H2 is used for organohalide respiration. In return,

Dhb exports glutamate and other amino acids to deplete the excessive NADPH and to facilitate

Bac growth. As opposed to Dhb isolate cultures, pyruvate did not accumulate in the ACT-3 mixed culture. Therefore, the exported pyruvate from Dhb is likely recycled by Bac, which shapes an intercellular metabolic cycle (Figure 4.4A). This interspecies exchange of malate and pyruvate enables the Dhb-Bac pair to trade reducing equivalents, resembling the function of a transhydrogenase (NADH + NADP+ → NAD+ + NADPH) (Voordouw et al., 1983).

We sought to validate the hypothesis of the interspecies malate-pyruvate exchange. Given that Bac ferments lactate to malate as a result of heme-auxotrophy, the addition of heme to the

ACT-3 should enable further malate fermentation to succinate, disrupting the interspecies malate- pyruvate exchange. Consistent with this hypothesis, 4% dilution transfers of ACT-3 culture supplemented with 1 mg/L heme revealed no CF dechlorination after 12 days under electron donor limited conditions (1 times the required theoretical electron equivalents for dechlorination), while the cultures without heme supplementation dechlorinated 10% of CF and the cultures supplemented with both 0.5 mM malate and heme dechlorinated ~40% CF (Figure 4.3B).

Moreover, the organic acid profile of culture supernatants at Day 12 revealed detectable malate and pyruvate in the cultures without added heme but not in the cultures with heme (Figure 4.3C),

92 indicating that the malate shortage disrupted Dhb growth and the malate-pyruvate exchange.

Nevertheless, when we provided excessive electron donors (3 times the theoretical electron equivalents), all the three cultures depleted CF after 6 days (Figure 4.3B). Altogether, our data support the presence of interspecies malate-pyruvate exchange in Dhb-enriched ACT-3.

4.4 Implications for Microbial Ecology

In this study, the proposed pyruvate-malate exchange between Bac and Dhb resembles the mitochondrial malate-pyruvate shuttle in eukaryotic cells (Liu et al., 2002; MacDonald, 1995). In the malate-pyruvate shuttle, cytoplasmic pyruvate is first transported to mitochondria, and is reduced to malate via TCA cycle. Malate is then exported to cytoplasm via an antiporter, and is decarboxylated to pyruvate by MAE for NADPH regeneration. Although the paring of metabolic partners in nature can be random, the complementary gaps in TCA cycle and in malate metabolism between Dhb and Bac genomes are likely a consequence of genome streamlining driven by the co- adaption (Figure 4.4 B). Consistently, malate dehydrogenase and fumarate reductase genes are present in most Peptococcaceae; citrate synthase, aconitase, and Idh are present in the genomes of all the available Bacteroides isolates. Accordingly, this study has found a relevant example in biology, the interspecies malate-pyruvate shuttle, to support the endosymbiotic hypothesis that mitochondria originated from a mutualistic interaction mediated by organic acids (Searcy, 2003).

In conclusion, the data present in this study reveals that metabolic complementarity and redox constraints are the driving forces for symbiotic amino acid exchange in anaerobic microbial communities. The successful model prediction also justifies our previous argument that the accuracy of metabolic annotation can be greatly improved with the consideration of cofactor availability and enzyme promiscuity. Furthermore, finding that Dhb is an amino acid producer in

93 mixed culture reinforces that the observed physiology of isolate under laboratory conditions may not reflect that in natural niches. Therefore, integration of laboratory experiments with computational modeling offers great opportunities to decipher the metabolic interdependency of the fastidious, or currently unculturable, microorganisms.

Acknowledgements Support was provided by the Government of Ontario through Genome

Ontario SPARK Research Grant. We also acknowledge the BioZone Mass Spectrometry facility for UPLC-ESI-HRMS analyses. We are grateful to the gift of active Dehalobacter restrictus strain

PER-K23 culture from Dr. Jun Yan and Prof. Frank Löffler in University of Tennessee (Knoxville,

USA).

CHAPTER 5 – BIOSYNTHESIS AND ACTIVITY OF PRENYLATED FMN

COFACTORS

2018 Elsevier Inc., Cell Chemical Biology.

5.1 Abstract

Prenylated FMN (prFMN) is a recently discovered cofactor required by the UbiD family of reversible decarboxylases involved in ubiquinone biosynthesis, biological decomposition of lignin monomers, and biotransformation of aromatic compounds. This cofactor is synthesized by UbiX- like prenyltransferases catalyzing the transfer of the dimethylallyl moiety of dimethylallyl- monophosphate (DMAP) to FMN. The origin of DMAP for prFMN biosynthesis and the biochemical properties of free prFMN are unknown. We show that in Escherichia coli cells, DMAP can be produced by phosphorylating prenol using ThiM or dephosphorylating DMAPP using

Nudix hydrolases. We produced 14 active prenyltransferases whose properties enabled the purification and characterization of protein-free forms of prFMN. In vitro assays revealed that the

UbiD-like ferulate decarboxylase (Fdc1) can be activated by free prFMNiminium or C2′- hydroxylated prFMNiminium under both oxidized and reduced conditions. These insights into the biosynthesis and properties of prFMN will facilitate further elucidation of the biochemical diversity of reversible UbiD (de)carboxylases.

5.2 Introduction

The enzyme cofactors flavin mononucleotide (FMN) and flavin adenine dinucleotide (FAD)

95 were discovered more than 80 years ago (Haas, 1938). Flavin-containing enzymes or flavoenzymes account for approximately 1% of cellular proteins and catalyze a plethora of redox reactions with organic substrates (Leys and Scrutton, 2016; Piano et al., 2017; Walsh and

Wencewicz, 2013). In addition to catalyzing common redox reactions in cell metabolism, DNA repair, protein folding and degradation of aromatics, flavoenzymes can catalyze acid/base and nucleophilic/electrophilic reactions (Sobrado, 2012), as well as reactions with mechanisms that involve the formation of highly reactive covalent flavin-substrate intermediates (Teufel et al.,

2016).

Some enzymes use modified flavins, including the most recently discovered prenylated FMN

(prFMN) cofactor in the active site of the Aspergillus niger ferulic acid decarboxylase Fdc1, which is required for the non-oxidative reversible decarboxylation of α,β-unsaturated acids via

1,3-dipolar cycloaddition (Payne et al., 2015). Fdc1 belongs to the widespread family of UbiD- like proteins named after E. coli UbiD involved in the biosynthesis of ubiquinone (Cox et al.,

1969; Payer et al., 2017). Previous genetic studies suggested that ubiD and the associated ubiX are isofunctional genes, encoding redundant aromatic decarboxylases (Zhang and Javor, 2003), although decarboxylase activity had not been demonstrated for both proteins (Gulmezian et al.,

2007; Jacewicz et al., 2013; Kopec et al., 2011; Rangarajan et al., 2004). A 2015 study with the

Saccharomyces cerevisiae UbiX-like Pad1 and UbiD-like Fdc1 revealed that rather than being a decarboxylase, Pad1 actually catalyzes the formation of a novel, diffusible FMN-type cofactor required for decarboxylase activity of Fdc1 (Lin et al., 2015). This work also demonstrated that the S. cerevisiae Fdc1 can be activated in vivo by co-expression with E. coli UbiX. The nature of the FMN modification and UbiX activity were determined by structural and biochemical studies of the UbiD from A. niger, S. cerevisiae, and Candida dubliniensis, as well as the UbiX-like

PA4019 from Pseudomonas aeruginosa (Payne et al., 2015; White et al., 2015). The authors demonstrated that UbiD is responsible for decarboxylase activity, which required a prFMN produced by PA4019.

High resolution crystal structures of UbiD in complex with oxidized prFMN revealed the presence of at least three cofactor forms: prFMN-N5-iminium (prFMNiminium), prFMN ketimine, and C1′-hydroxylated prFMN, and suggested that prFMNiminium represented the catalytically- active species (Payne et al., 2015). PA4019 was found to be a FMN prenyltransferase, which transfers a dimethylallyl moiety from dimethylallyl-monophosphate (DMAP) to the flavin N5 and

C6 atoms, adding a fourth ring to the FMN isoalloxazine group (Fig. 5.1a). This reaction required the reduction of the FMN-DMAP-PA4019 complex by dithionite (DT) followed by re-oxidation by air, producing a stable purple-colored protein with a second absorbance maximum (λ2) at 5nm

(White et al., 2015). However, a recent study revealed that in contrast to bacterial UbiX the S. cerevisiae UbiX-like protein PAD1 prefers dimethylallyl pyrophosphate (DMAPP) as the prenyl group donor, which is a common precursor in isoprenoid biosynthesis (Sachs et al., 2011). Liquid chromatography-mass spectrometry (LC-MS) analysis of small molecule extracts from both reduced and re-oxidized PA4019 complexes showed the presence of fully reduced prFMN

(prFMNred) and prFMN-C4a-radical (prFMNradical), respectively (White et al., 2015).

Decarboxylase activity of the A. niger Fdc1 could be reconstituted by anaerobic incubation with

prFMNred PA4019 containing prFMNred (PA4019 ) followed by re-oxidation by air, but not with

prFMNradical PA4019 , suggesting that UbiD requires prFMNred to be oxidized to the catalytically- active prFMNiminium (Payne et al., 2015). In addition, the Fdc1 reconstitution appeared to be independent of the presence of UbiX, because Fdc1 activity was also observed using protein-free extracts of PA4019prFMNradical (Payne et al., 2015) and holo-Fdc1 (Lin et al., 2015), respectively.

The crystal structure of an Fdc1-prFMN complex (Fdc1prFMN) suggested that the reversible substrate decarboxylation proceeded via 1,3-dipolar cycloaddition of the prFMNiminium azomethine ylide to the α,β-double bond of substrate side chain (Payne et al., 2015), which was also supported by molecular modeling (Tian and Liu, 2017), kinetic isotope effects (Ferguson et al., 2016), and mechanism-based inhibitor analysis (Ferguson et al., 2017).

The remarkable discovery of prFMN structure and associated catalytic mechanism for decarboxylation gives rise to many important questions, including confirmation of FMN prenyltransferase activity in other UbiX-like proteins, the biosynthetic origin of DMAP, as well as the biochemical properties of free prFMN and how these affect interactions with UbiD. In this study, we completed a phylogenetic analysis of over 9,000 UbiX-like proteins and demonstrated

FMN prenyltransferase activity in 14 UbiX-like proteins from different phylogenetic groups. We found that in E. coli, DMAP can be produced by direct phosphorylation of prenol by the kinase

ThiM and by dephosphorylation of DMAPP by the Nudix hydrolases NudF and NudJ. Using optimized reaction conditions, we produced several protein-free prFMN forms, including a red- colored protonated prFMNradical, and demonstrated activation of apo-Fdc1 by free cofactors.

5.3 Materials and methods

(Supplemental Information including figures and tables are available in Appendix F)

Experimental model and subject details. All proteins and plasmids used for in vitro studies were purified from Escherichia coli strain BL21(DE3) Gold as described in the text/resource table. The

E. coli strain BW25113 ΔnudF::Kmr and ΔnudJ::Kmr strains were obtained from the Keio

Collection of single-gene knockouts (Baba et al., 2006).

Phylogenetic analysis. A multiple sequence alignment was generated from 9,043 InterPro family

(IPR004507) sequences using online MAFFT alignment tool. The original dataset was reduced to

5,534 sequences using the Max Align and CD HIT refinement strategies from the same website.

The phylogenetic tree was constructed with the “approximately maximum-likelihood” algorithm provided by FastTree 2.1.5 built in Geneious 8.1.8. The phylogenetic tree was then visualized using Itol.

Gene cloning and mutagenesis. The UbiX genes from different organisms were amplified by

PCR from genomic DNA and cloned into a modified pET15b vector (Novagen) containing an N- terminal His6-tag as described previously (Kuznetsova et al., 2006). Site-directed mutagenesis of the E. coli kinase ThiM was performed using the QuikChangeTM site-directed mutagenesis kit

(Agilent) according to the manufacturer’s protocol, and mutations were verified by DNA sequencing. The plasmids were transformed into E. coli strain BL21 (DE3) Gold as expression host. The E. coli strain BW25113 ΔnudF::Kmr and ΔnudJ::Kmr strains were obtained from the

Keio Collection of single-gene knockouts (Baba et al., 2006). The one-step gene inactivation method of Datsenko and Wanner (Datsenko and Wanner, 2000) was used to create a double

ΔnudJ::Kmr, ΔnudF::Cmr knockout strain (in BW25113) and a thiM deletion strain (ΔthiM::Cmr; in BL21(DE3) Gold)).

Protein expression and purification. For recombinant expression and purification of UbiX-like proteins, the cultures were grown under aerobic conditions at 37 °C in a shaker using Terrific Broth

(200 rpm) to the optical density (OD600 nm) 0.8-1 followed by induction with 0.4 mM isopropyl β-

D-1-thiogalactopyranoside (IPTG) and overnight incubation at 16 °C (for aerobic protein expression). For anaerobic protein expression, after the IPTG induction the cultures were transferred to screw-capped glass bottles and supplemented with Dimethyl sulfoxide (2 mL/L) and

(if indicated) riboflavin (0.2 mM) and/or prenol (1 mM), and incubated overnight at room temperature with constant stirring. Recombinant proteins were purified to at least 95% homogeneity using metal-chelate affinity chromatography on Ni-NTA Superflow resin as previously described (Kuznetsova et al., 2006), and protein purity was analyzed by SDS-PAGE.

The E. coli kinases and phosphohydrolases (Nudix) were expressed using the ASKA Collection clones for expression of E. coli proteins (Kitagawa et al., 2006) and affinity purification according to the manufacturer’s protocol.

Enzyme assays. The E. coli strain BL21(DE3) cell lysates for enzyme assays and proteomics analysis were prepared using the BugBuster® Protein Extraction Reagent according to manufacturer’s protocol (except that 1 mM phenylmethylsulfonyl fluoride was added to the cell suspension for protease inhibition). Purified proteins were at least 95 % pure based on SDS-PAGE analysis. Anaerobic assays were performed in an anaerobic glove box (Coy) with an atmosphere of 10% CO2, 10% H2, and balancing N2.

In vitro FMN prenylation by purified UbiX. The reaction mixtures (110 μL) contained 40 mM

HEPES-Na buffer (pH 7.5), 0.4 M NaCl, 60 μM FMN, 0.1 mM dimethyl-allyl monophosphate

(DMAP), 20 mM K-orthophosphate, and up to 5 mg of cofactor-free (colorless) AF1214. Under anaerobic conditions (in a glove box), the reaction mixture was reduced by the addition of 3 mM sodium dithionite (DT; 2 h under anaerobic conditions at 37 °C), filtered through 3 kDa spin filters, oxidized on air (10-20 min), and the flow-through fraction was used for absorption spectra analysis.

Anaerobic reaction mixtures for DMAP saturation experiments with the AF1214FMN (yellow) contained 100 mM HEPES-Na (pH 7.5), 0.4 M NaCl, 3 mM DT, 0-0.175 mM DMAP, and 0.175 mM AF1214FMN. After 1 h incubation at 37 °C, the reaction mixtures were oxidized on air for 10

100 min followed by absorbance analysis at 550 nm. All enzyme assays were conducted in duplicates or triplicates as indicated on figures.

UbiX-based prenol kinase screens. The colorimetric prenol kinase screen is based on the ability of AF1214 to prenylate reduced FMN using DMAP and retain the oxidized prFMN after the reaction producing a purple-colored protein containing prFMNradical with the λ2 at 550 nm. The screen includes two steps: prenol phosphorylation to DMAP and FMN prenylation. The prenol phosphorylation reaction mixture (0.1 mL) contained 100 mM HEPES-Na buffer (pH 7.5), 0.3 M

NaCl, 10 mM MgCl2, ATP (10 mM for cell lysates and 5 mM for purified proteins), 1 mM polyphosphate, prenol (10 mM for cell lysates and 5 mM for purified proteins), 10 mg/mL cell lysate or 0.1 mg/mL purified protein was incubated for 1 h at 37°C with shaking at 200 rpm. After

1 h incubation at 37°C, the reaction mixtures were supplemented with 0.1-0.3 mg of the reduced

(with 3 mM DT) AF1214FMN, followed by 1 h incubation at 37°C in an anaerobic glove box.

Finally, the reaction mixtures containing AF1214prFMNred were oxidized on air, and the absorbance

FMN prFMNradical at 460 nm (λ2 for UbiX ) and 550 nm (λ2 for UbiX ) were recorded. The (λ550 nm /λ460 nm) ×10 values higher than 0.2 or formation of a purple-colored solution were considered as positive results for the presence of prenol kinase activity. The determination of the DMAP product in cell lysate assays was carried out using LC-MS as described in the MS analysis section.

DMAPP dephosphorylation activity. Assays were performed in 96-well microplates using reaction mixtures (25 μL) containing 100 mM Tris-HCl (pH 8.0; for assays with purified phosphohydrolases) or 100 mM HEPES-Na (pH 7.0; for assays with cell lysates), 0.1-1 mM dimethyl-allyl pyrophosphate (DMAPP), 5 mM MgCl2, 0.5 mM MnCl2, and purified phosphohydrolases (2-10 μg) or cell lysates (25 μg). After 10-20 min incubation at 37 °C, the reaction was stopped by the addition of Malachite Green reagent and the production of

101 orthophosphate was measured based on absorbance at 630 nm (Baykov et al., 1988). The determination of the DMAP product and residual amounts of DMAPP in cell lysate assays was carried out using LC-MS as described in the MS analysis section.

Prenol and thiazole phosphorylation assays. Kinase activities of purified ThiM were assayed spectrophotometrically at 37 °C using a microplate-based enzyme-coupled assay with lactate dehydrogenase and pyruvate kinase as previously described (Huo and Viola, 1996). Reaction mixtures (0.2 mL) contained 100 mM HEPES-Na (pH 7.5), 10 mM MgCl2, 5 mM phosphoenolpyruvate, 0.3 mM NADH, 2 mM ATP, lactate dehydrogenase (2.5 units), pyruvate kinase (1.9 units), 5-ethynyl-4-methyl-1,3-thiazole or prenol as substrate (1-50 mM), and purified protein (5-10 µg). In addition, the kinase reaction product ADP was analyzed using reversed phase chromatography on a Varian ProStar HPLC system equipped with a AXXI-CHROM ODS column

(5-μm-size particles, 4.6 × 250 mm; Cole Scientific) (Chen et al., 1997).

Fdc1 activation and decarboxylation assays. For activation of the purified A. niger apo-Fdc1, the reaction mixture (17 µL) containing 100 mM Na-phosphate buffer (pH 6.0), 200 mM NaCl,

0.25 nmol of PA4019prFMN (or AF1214prFMN, or equimolar amounts of free prFMN forms) was incubated with apo-Fdc1 (0.25 nmol) for 5-60 min under aerobic or anaerobic conditions at room temperature. For Fdc1 activation experiments with orthophosphate saturation, the reaction mixture contained 100 mM MES-K buffer (pH 6.0) and 0-10 mM phosphate. The decarboxylation activity of activated Fdc1 was determined spectrophotometrically at 270 nm as previously described(Payne et al., 2015). The reaction mixture (0.2 mL) contained 100 mM MES-K buffer (pH 6.0), 200 mM

NaCl, 0.2 mM MnCl2, 0.3 mM sodium cinnamate, and 1 µg of Fdc1 (incubation at 30 °C, 20-40 min).

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Extraction of prFMN. Purified UbiX-like proteins (AF1214 or PA4019) were saturated with prFMN in vitro by incubating in a reaction mixture containing 100 mM Na-phosphate buffer (pH

7.0), FMN (0.1-1 mM), and DMAP (1 mM) under anaerobic conditions. After reduction with DT

o (3 mM, 37 C, 1 h), the proteins were oxidized by air (producing prFMNradical), precipitated with ice-cold 50 % (v/v) acetonitrile and washed two times with the same solution. All further purification steps were performed in an anaerobic chamber (Coy) using anaerobic solutions. For the purification of free prFMNradical (Structure B, Fig. 4) from PA4019, the protein was resuspended in 70% acetonitrile and heated for 10 min at 80°C on a shaker (800 rpm). For

prFMN prFMN protonated prFMNradical purification, UbiX-like proteins (AF1214 or PA4019 ) were resuspended in 50 % acetonitrile and acidified with 50-100 mM HCl. The proteins were precipitated by the addition of acetonitrile to a final concentration of 86 % and centrifuged at

13,000 x g for 10 min at room temperature. The solution was either directly used for assays or concentrated using a rotary evaporator until full acetonitrile evaporation. Cofactor preparations were stored frozen in the O-ring-capped plastic tubes at -80 °C.

Determination of prFMN extinction coefficient. For the determination of extinction coefficient of free protonated prFMNradical (C), we used the purified AF1214 saturated with FMN. One half of this sample was used to extract FMN (in triplicate) and determine its concentration using a

-1 -1 calibration curve generated with synthetic FMN and extinction coefficient (10.9 mM cm , λ442 nm). The other half was incubated anaerobically with an excess (6x) of DMAP for the complete transformation of FMN to prFMN and subsequent cofactor extraction (also in triplicate). LC-MS analysis of the AF1214FMN extracts and AF1214prFMN extracts revealed negligible amounts of FMN and riboflavin, respectively, suggesting complete transformation of FMN to prFMN. Thus, we assume that the concentration of FMN in the AF1214FMN extract is equal to that of protonated

103

prFMN prFMNradical in the AF1214 extract. Using serial dilutions of purified protonated prFMNradical and its λ2 at 490 nm, we established a calibration curve and determined the extinction coefficient

-1 -1 of prFMN to be 6.3 mM cm (λ490 nm).

Mass spectrometry. For protein mass spectrometry, the E. coli BL21 (DE3) wild-type and ΔthiM mutant strains grown without or with the addition of 1 mM prenol were used. The strains were grown in duplicate at 37 °C on terrific broth to OD600nm 0.6-0.8, and then incubated overnight at

16 °C. The cells were collected by centrifugation at 10,000 x g for 10 min and lysed using

BugBuster Master Mix as described in manufacturer’s protocol. The lysates were mixed with 2 times SDS loading buffer (50 mM Tris-HCl (pH 6.8), 10 % SDS, 40 % glycerol, 3 mM bromophenol blue, 0.5 M DTT, separated on 15 % SDS-PAGE gels, stained by Coomassie Blue for 2 h and de-stained overnight. The portion of the gel corresponding to molecular mass 10-35 kDa was excised and used for overnight trypsin digestion as described previously (Shevchenko et al., 2006). Trypsin digestion was stopped using 0.1 % TFA and desalted using C-18 OMIX tips

(Agilent) according to the manufacturer’s instructions. Protein samples were resuspended in 0.1 % formic acid and analyzed by mass spectrometry using a Q Exactive MS system (Thermo Scientific) at the BioZone Mass Spectrometry Facility (University of 663 Toronto, Toronto, Canada; https://www.biozone.utoronto.ca/mass_spec/). Raw MS data was processed and analyzed using

X!TandemPipeline (Langella et al., 2016) and is presented in in a separate Excel spreadsheet in

Appendix F.

The LC-MS platform for small molecule analysis contained a Dionex Ultimate 3000 UHPLC system equipped with a photodiode array detector and a Q-Exactive MS equipped with a HESI II source (all from Thermo Scientific). Control of the system and data handling was performed using

Thermo XCalibur 2.2 software and Chromeleon 7.2 software. Separation by liquid

104 chromatography was conducted at 40oC on a Thermo Hypersil Gold C18 column (50 mm x 2.1 mm, 1.9 µm particle size, Thermo Scientific). The pump was run at a flow rate of 0.2 mL/min.

Solvent A was water containing 5 mM ammonium acetate (pH 6.0); solvent B was methanol containing 5 mM ammonium acetate (pH 6.0). For prFMN and FMN analysis, the gradient was 0-

1 min, 2 % B; 1-7 min, linear increase to 90 % B; 7-10.5 min, hold at 90% B; 10.5-11 min, return to initial conditions (2 % B) and hold for 7 mins prior to next sample. For DMAP and DMAPP analysis, the gradient was 0 min, 2 % B; 0-1 min, 2 % B; 1-4 min, 90 % B; 4-7 min, 90 % B; 7-

7.5 min, return to initial conditions (2 % B) and hold for 4.5 mins prior to next sample. The autosampler temperature was maintained at 10 ˚C and the injection volume was 20 µL. Data collection was done in both positive and negative ionization modes with a scan range of m/z 200-

700, and mass resolution of 140,000, Automatic Gain Control target of 3e6 and a maximum injection time of 200 ms.

Data and Software Availability. The phylogenetic tree of 5,534 unique UbiX proteins can be accessed through Itol website (http://itol.embl.de/tree/142150588823901497476579). All other data supporting the findings of this study are available within the paper and its supplementary information files.

5.4 Results and Discussion

Phylogenetic analysis of the UbiX family of FMN prenyltransferases

Most of the over 9,000 UbiX-like sequences (IPR004507, InterPro database) are present in bacterial genomes (>7,800 genes) with the remaining sequences found in archaea (~500 genes), fungi and unicellular algae (>500 genes). Phylogenetic analysis of the UbiX family revealed the presence of at least five major clusters (I-V) (Fig. 5.1b, S1). Some bacterial genomes contain

105 more than one ubiX gene in which case these genes are typically found in separate operons, suggesting involvement in different anabolic or catabolic processes. For example, there are two distinct ubiX genes in Geobacter metallireducens and four in Azoarcus sp. EbN1 (Aromatoleum aromaticum EbN1). In G. metallireducens, the ubiX gene in the phenyl-phosphate carboxylase operon (Q39TU0, Gmet2105) encodes a protein that clusters with UbiX sub-family II, whereas the second ubiX gene (Q39Q70, Gmet3392, 38% sequence identity to Gmet2105) encodes a protein that clusters with UbiX sub-family V (Fig. 5.1b).

Of the four ubiX paralogs in Azoarcus sp. strain EbN1, one (Q5P483) is in the phenyl-phosphate carboxylase operon, two (D5NWH5 and D5NWG7) are in the phthaloyl-CoA decarboxylase operon, and the fourth (Q5P010) appears to be a single gene (Table S1), and each encode proteins that cluster with different UbiX sub-families (I, II, and III) (Fig. 5.1b). UbiX whose genes are co- operonic with ubiD genes encoding (de)carboxylases involved in the degradation of aromatic compounds (benzene, phthalate, phenol, or naphthalene) are located in UbiX sub-families II and

III (Figures 5.1b,c; Table S1). Finding multiple ubiX genes within a single genome with different phylogenies suggests that some may have been horizontally-acquired.

Purification of novel UbiX-like proteins with bound prFMN

We cloned 23 UbiX-like proteins sampled around the phylogenetic tree for recombinant expression in E. coli BL21 (Figures 5.1b,S2). Preliminary expression experiments with the recently characterized PA4019 revealed that aerobically grown E. coli cultures produced colorless

PA4019, whereas a purple-colored protein was purified from cultures that were rendered anaerobic after induction with isopropyl-β-D-1-thiogalactopyranoside (IPTG) and that were also provided with riboflavin (FMN precursor) and prenol, a possible DMAP precursor (Figures

5.2a,S3a).

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Fig. 5.1. Phylogenetic analysis of the UbiX family. (a) Schematic diagram showing UbiX-catalyzed biosynthesis of prFMN and its roles in biological processes. (b) Unrooted phylogenetic tree of the UbiX family showing five UbiX clusters (I-V). Cluster II includes UbiX sequences from known anaerobic aromatic degradation operons, whereas cluster III contains UbiX genes from the phthaloyl-CoA decarboxylase operons. UbiX paralogs from Azoarcus sp. strain EbN1 and G. metallireducens are labelled as ♦ and ▼, respectively. UbiX-like proteins purified in this work including PA4019 and AF1214 are marked with purple circles and labelled. (c) Anaerobic aromatic degradation operons from different organisms containing ubiX (red arrows) and ubiD (green arrows) genes, as well as their substrates and products.

107

These results suggested that prFMN biosynthesis in E. coli cells might be limited by low intracellular concentrations of DMAP. When the 23 cloned UbiX-like proteins were over- expressed using this optimized protocol (anaerobic conditions with the addition of riboflavin and prenol), we were able to purify 14 soluble proteins, of which five showed purple color in the preparations: AF1214 from Archaeoglobus fulgidus, MJ0101 from Methanocaldococcus jannaschii, SMb20135 from Sinorhizobium meliloti, MTH0147 from Methanothermobacter thermoautotrophicus, and PA4019 (Table S2). Absorption spectra confirmed the presence of prFMNradical (λ1 360 nm and λ2 550 nm) in these preparations (data not shown). Additional expression trials with AF1214 revealed that, in contrast to PA4019, the AF1214 preparations purified from either aerobically- or anaerobically-grown cultures without prenol addition had yellow color with absorption spectra typical of FMN-containing proteins with λ1 at 370 nm and

λ2 at 460 nm (Fig. 5.2b, samples I, II, and VI). Aerobic preparations with prenol addition also showed similar spectra (Fig. 5.2b, sample IV). However, anaerobic expression of AF1214 in the presence of prenol produced purple-colored preparations with absorption spectra, indicating the presence of prFMNradical with λ1 at 360 nm and λ2 at 550 nm (Fig. 5.2b, samples VII and VIII).

The addition of riboflavin to these cultures did not alter the results. Therefore, recombinant expression of AF1214prFMN in E. coli seemed to be limited by intracellular DMAP levels, and this limitation could be overcome by the addition of prenol to the growth medium. In addition, AF1214 appeared to have a higher affinity to FMN compared to PA4019 as the former retained FMN even after protein purification from aerobically grown E. coli cultures.

108

Fig. 5.2. Recombinant expression and purification of the UbiX-like AF1214. (a) Schematic diagram showing UbiX expression experiment in E. coli cultures. (b) Vials of AF1214 preparations purified from E. coli cells grown under different conditions as indicated. The results (λ550/λ460) show the mean from experiments performed at least in duplicate (variation ~20%). (c) Absorption spectra of purified AF1214 (depicted in b). In vitro prenylation of FMN by PA4019 and AF1214

PA4019 was shown to bind free FMN in the presence of DMAP, but for FMN prenylation, this complex required reduction by DT followed by re-oxidation by air to produce a stable purple- colored prFMNradical (White et al., 2015). We found that FMN binding by the colorless (FMN- free) AF1214 (Fig. 5.2b; sample V) under anaerobic conditions (reduced by DT) was stimulated by the addition of orthophosphate alone, and was further stimulated by addition of DMAP (Fig.

S4a). Interestingly, both orthophosphate and DMAP showed no stimulation of FMN binding by

AF1214 under aerobic conditions (Fig. S4b). Furthermore, reduction of AF1214FMN (samples I

109 and II in Fig. 5.2b) by DT in the presence of DMAP followed by air oxidation resulted in FMN prenylation and formation of purple-colored AF1214prFMNradical based on absorption spectrum (Fig.

S4c). Substrate saturation experiments with DMAP revealed that FMN prenylation by AF1214FMN

FMN is saturated with 100 μM DMAP with apparent Km 30 M (Fig. S4d). Thus, AF1214 can be used as an indicator protein for DMAP detection based on changes in absorption spectrum following DT reduction and air oxidation. Optimized conditions for in vitro FMN prenylation by

UbiX (orthophosphate added) enabled the assay of the 14 previously purified soluble UbiX-like proteins for FMN prenylation activity. We detected purple color in the preparations of HP1451 from Helicobacter pylori and JGI0011 from Alkaliphilus metalliredigens, indicating formation of prFMNradical that was retained within the enzyme (Table S2). For the remaining proteins, LC-MS analysis of protein-free reaction filtrates confirmed production of prFMNiminium and prFMN-OH by all purified colorless UbiX-like proteins (Fig. S4e), including NBRC0004 from a benzene- degrading Peptococcaceae sp. (Fig. 5.1c; Table S2). This suggests that prFMN is likely involved in the predicted benzene carboxylation reaction catalyzed by the associated putative UbiD-like anaerobic benzene carboxylase, AbcA (Fig. 5.1c). While all purified UbiX-like proteins tested possess FMN prenylation activity, they seem to have different affinities to the reaction product, prFMN (Fig. S4e). Nevertheless, FMN prenylation activity appears to be conserved in all major phylogenetic groups of the UbiX family.

Biosynthesis of DMAP in E. coli cells: prenol and DMAPP as precursors

We sought to investigate the origin of DMAP required by UbiX for prFMN biosynthesis.

Partially purified E. coli cell lysates were reported to phosphorylate prenol to DMAP (Lange and

Croteau, 1999), which was confirmed in the experiments using E. coli BL21 wild-type cell lysates in the presence of ATP and Mg2+ (Fig. S3b). This is consistent with our finding that prenol addition

110 to E. coli cultures stimulated the formation of prFMN by recombinant UbiX-like proteins

(Figures 5.2b,S3a).

Fig. 5.3. Screening for DMAP-producing enzymes. (a) Schematic diagram showing the AF1214-based screen for prenol kinase activity using E. coli cell lysates (1st round), or purified candidate proteins (2nd round, shown in b). (b) Microplate wells showing the results of AF1214-based screening (2nd round) of 16 purified candidate proteins (in duplicate). The purple color of two wells with ThiM suggests the presence of prenol kinase activity in this protein. DMAP: a positive control with 1 mM DMAP added instead of protein. λ550 nm/450 nm: the ratio (multiplied by 10) of reaction mixture absorbance at 550 nm to 450 nm indicating the formation of prFMN (from the produced DMAP). (c) Screening of 16 purified E. coli phosphohydrolases for DMAPP dephosphorylation activity. Production of orthophosphate after incubation of DMAPP (1 mM) with 16 purified E. coli Nudix hydrolases. (d) LC-MS analysis of DMAPP conversion to DMAP by the E. coli NudF.

111

To identify E. coli kinases that can phosphorylate prenol to DMAP, we used the purified

AF1214FMN as a reporter protein to screen E. coli cell lysates over-expressing 98 E. coli kinases known to be active toward hydroxyl-containing substrates, as well as several uncharacterized kinases (Table S4a). As shown in Fig. 5.3a, after IPTG induction of kinase expression, the cell lysates were prepared and incubated for 1 h with prenol, ATP, and purified AF1214FMN (yellow), followed by FMN reduction by DT and re-oxidation by air (Fig. 5.3a). Of the 98 clones screened, at least 16 lysates developed a weak reddish color, which correlated with higher values of the λ550

prFMNradical nm/λ460 nm ratio (0.2 – 1.0), suggesting the formation of AF1214 (Table S4a). The recombinant proteins from the positive clones were affinity purified, and a second AF1214-based screen confirmed the presence of prenol kinase activity in the hydroxyethylthiazole kinase ThiM

(Fig. 5.3b). ThiM, a salvage enzyme in thiamine biosynthesis, has been shown to catalyze the phosphorylation of 4-methyl-5-beta-hydroxyethylthiazole (THZ) (Mizote and Nakayama, 1989), with structural similarity to prenol (Fig. S5a). Alanine replacement mutagenesis confirmed that the prenol kinase activity of ThiM depended on the residues known to be critical for THZ phosphorylation (Fig. S5b) (Dyguda-Kazimierowicz et al., 2007). Prenol addition to E. coli BL21 wild-type cultures had no measurable effect on ThiM protein expression level based on LC-MS analysis (Tables S4,S5), whereas a small decrease in activity in the thiM deletion (ΔthiM::CmR) strain was observed using the PA4019-based assays for prFMN formation, suggesting that additional proteins in E. coli have prenol kinase activity (Fig. S5c). In the presence of ATP and

Mg2+, purified ThiM catalyzed comparable phosphorylation rates both with prenol and THZ (0.18 and 0.19 μmol/min/mg protein, respectively), but had lower affinity for prenol (Km 14 mM) compared to THZ (Km 2.0 mM) (Fig. S5a).

112

Another possible biosynthetic route to DMAP in bacteria is from DMAPP dephosphorylation.

DMAPP is an intermediate in the mevalonate (Meganathan, 2001) or MEP (2-C-methyl-D- erythritol 4-phosphate) (Lichtenthaler et al., 1997) isoprenoid biosynthesis pathways. Our preliminary results indicated that Mg2+ addition stimulated DMAPP and DMAP dephosphorylation in E. coli cell extracts (Figures 5.3c,d). In addition, E. coli cultures over- expressing the Nudix hydrolase NudF (ADP-ribose pyrophosphatase) were reported to dephosphorylate DMAPP to prenol (Zheng et al., 2013). However, the intermediary products of

DMAPP dephosphorylation by purified Nudix enzymes have not yet been analyzed. Therefore, we screened 16 purified E. coli Nudix hydrolases and nucleotide pyrophosphatase RdgB (YggV;

Table S3b) for DMAPP dephosphorylation. These screens revealed high DMAPP dephosphorylation activity in four Nudix proteins including NudB, NudF, NudI, and NudJ, which completely hydrolyzed 1 mM DMAPP after 20 min incubation (Fig. 5.3c). LC-MS analysis of

NudF reaction mixtures revealed the presence of DMAP as the only organic reaction product

(prenol undetected) (Fig. 5.3d). Furthermore, LC-MS analysis of the E. coli BL21 wild-type proteomes showed significant expression levels of NudF and NudJ, whereas the other two Nudix hydrolases were not detected (Tables S4,S5). E. coli NudJ has been shown to dephosphorylate several nucleoside tri- and di-phosphates and thiamine pyrophosphate to corresponding monophosphate products (Lawhorn et al., 2004; Xu et al., 2006). DMAPP saturation experiments with purified NudF and NudJ revealed similar Km values (0.8 mM and 0.5 mM, respectively), but NudJ had two times higher Vmax (0.3 μmol/min/mg protein) than NudF (0.15 μmol/min/mg protein)

(Figures S5d,e). With GDP and ADP-ribose as substrates, these enzymes have been reported to have similar Km values (0.6 mM and 0.1 mM, respectively), but higher Vmax (12-297 μmol/min/mg protein) (Dunn et al., 1999; Xu et al., 2006). Nevertheless, DMAPP dephosphorylation assays

113 with cell lysates from E. coli cells with singly or doubly deleted genes (ΔnudJ::KmR; ΔnudF::KmR; or ΔnudJ::KmR, ΔnudF::CmR) indicated that both enzymes can contribute to the in vivo production of DMAP from DMAPP (Fig. S5f). Thus, our results suggest that in E. coli cells ThiM can produce DMAP via prenol phosphorylation, whereas NudF and NudJ can dephosphorylate

DMAPP to DMAP.

Extraction and biochemical characterization of free prFMN forms

High-temperature extraction of PA4019prFMNradical in 70 % acetonitrile under anaerobic

- conditions and neutral pH released purple-colored prFMNradical, identified by MS as an [M-H] • adduct with m/z = 524.165 in negative-ion mode (form B2 in Figures 5.4a,S6a). However, high- temperature extraction of AF1214prFMNradical under neutral pH did not release free prFMN, likely due to protein thermostability. UV-vis absorption spectrum of free prFMNradical (B2) was similar with that of the UbiX-bound prFMNradical (B1), but its λ2 shifted to 515 nm (Fig. 5.4bi), whereas air exposure resulted in rapid loss of purple color and peak disappearance. This is in contrast with the protein-bound prFMNradical (B1), which was found to be stable under aerobic conditions, suggesting that UbiX protects prFMN from O2.

The addition of 50-100 mM HCl to free prFMNradical (B2) under anaerobic conditions changed its color from purple to red, and the absorption spectrum showed a shift in the λ2 peak from 515 nm to 490 nm (Fig. 5.4bi). MS analysis of this preparation in positive-ion mode revealed a radical cation (M+•) with m/z = 526.182 (Fig. 5.4ci). This peak was interpreted as a molecular cation and not the [M+H]+ adduct as the corresponding anionic adduct ([M-H]-) was not observed in negative-ion mode (Schäfer et al., 2007). This suggests the formation of protonated prFMNradical cation at low pH (C in Fig. 5.4a). Furthermore, C was also obtained from AF1214prFMNradical by acidic extraction (50-100 mM HCl) under anaerobic conditions. To determine the extinction

114 coefficient of free prFMN, we used purified AF1214FMN. Half of this sample was used to extract

-1 -1 FMN and determine its concentration and extinction coefficient (10.9 mM cm , λ442 nm) using a calibration curve prepared with synthetic FMN. The other half was incubated anaerobically with an excess (6x) of DMAP for complete transformation of FMN to prFMN (confirmed by LC-MS analysis; Fig. S7a), and subsequent cofactor extraction. Based on the absorbance of purified prFMN samples at 490 nm (λ2) and assuming all FMN was converted to prFMN, we calculated

-1 -1 the extinction coefficient of protonated prFMNradical (C) to be 6.3 mM cm at λ490 nm (Fig. S7b).

Protonated prFMNradical (C) reduction by DT resulted in discoloration and disappearance of the

+ λ490 nm peak (Fig. 5.4bii), whereas MS analysis of the reduced sample revealed a M cation with m/z = 527.190 (Fig. 5.4cii) corresponding to the monoisotopic mass of the protonated prFMNred cation (A2 in Fig. 5.4a). The λ490 nm peak of the reduced preparation was partially restored by air oxidation for 1 min (Fig. 5.4bii), suggesting that C and A2 can undergo reversible reduction and oxidation.

Air exposure of the partially oxidized preparation for an additional 5 min caused a significant decrease in λ490 nm with a color change from red to yellow. MS analysis of this sample revealed the presence of a mixture of a M+ cation and [M+H]+ adduct with m/z = 526.182 and 525.174 (Fig.

5.4ciii) corresponding to monoisotopic masses of protonated prFMNradical (C) and prFMNiminium

(D in Fig. 5.4a), respectively. After overnight air exposure of C at neutral pH (7), most of this cofactor was oxidized to D, which has been identified as the catalytically relevant prFMN species for Fdc1 (Payne et al., 2015). We found D to be relatively stable under aerobic conditions at neutral pH following overnight incubation at 4°C (retention time 10.5 min in LC-MS; Fig. S6c).

115

Fig. 5.4. Proposed biochemical and chemical transformations of prFMN. (a) Diagram showing the bio- formation of prFMN by UbiX (inside the pale purplish rectangle) and its chemical transformations to other prFMN forms after cofactor extraction and different treatments (acid/alkaline; [H], reduction by DT; [O], oxidation by air). The prFMN forms are labelled as follows: A1, UbiX-bound prFMNred; A2, protonated prFMNred; B1, UbiX-bound prFMNradical; B2, prFMNradical; C, protonated prFMNradical; D, prFMNiminium; E, C1′-hydroxylated prFMN; F, C1′-ene-prFMNiminium; G, C2′-hydroxylated prFMNiminium. For free cofactors, the monoisotopic mass is indicated below their structures, as well as the λ2 peaks and solution color for colored species. (b) Changes in absorption spectra (λ2 peaks) of the UbiX-bound prFMNradical form (form B1) and protonated prFMNradical (form C) to other species: (i) after extraction from UbiX (pH 2 or 7); (ii) after reduction by DT and re-oxidation by air; (iii) after overnight oxidation at different pH at 4°C. (c) MS spectra of form C after (i) extraction from UibX, followed by (ii) reduction by DT and (iii) re-oxidation by air for 5 min. 116

Overnight incubation of D at alkaline pH (10) resulted in sample discoloration (Fig. 5.4biii), and LC-MS analysis identified a major [M+H]+ adduct with retention time 10.0 min and m/z =

543.18, corresponding to the monoisotopic mass of a C1′-hydroxylated prFMN (E in Figures

5.4a,4biii,S6d). Recently, this form was observed in old crystals of the A. niger holo-Fdc1 and acetonitrile extracts of holo-UbiD (Marshall et al., 2017; Payne et al., 2015). However, overnight incubation of C at acidic pH (2) produced a green-colored preparation with λ2 at 740 nm (Fig.

5.4biii). Adjustment of pH (from 2 to 6) resulted in discoloration of the preparation, and a λ2 shift from 740 nm to 760 nm. A photodiode array LC-MS analysis assigned the λ760 nm peak at retention time 10.1 min to an M+ cation with m/z = 523.160 (Figures S6b,e,f), representing the monoisotopic mass of C1′-ene-prFMNiminium cation (F in Fig. 5.4a). This is consistent with low solubility of F in acetonitrile and its high solubility in water, as well as with the long wavelength absorbance peak at 740 nm due to the presence of a highly conjugated π-system. This form appears to be the product of D oxidation with the loss of a C2′-hydride; however, we found that

F can only be produced from C but not from D, suggesting participation of the radical in the acidic oxidation process. Prolonged (2-3 days) storage of either D or F under aerobic conditions resulted in further oxidation of these cofactors. MS analysis of the prFMN extract revealed the presence of two [M+H]+ adducts with m/z = 541.17 and 559.18, corresponding to monoisotopic masses of

C2′-hydroxylated prFMNiminium (G in Fig. 5.4a) and C1′,C2′-dihydroxylated prFMN forms, respectively (H in Fig. S6h). Thus, using purified free prFMN, we confirmed formation of B2, D,

E, F, and G. and proposed two previously unreported forms: C and H (Table S6).

In vitro activation of Fdc1 decarboxylase by free prFMN forms

The inactive apo-Fdc1 has been shown to be readily activated by in vitro anaerobic reconstitution with PA4019prFMNred followed by aerobic oxidation. Decarboxylation activity of the

117 reconstituted holo-Fdc1 was observed only after exposure to air, suggesting that prFMNiminium (D) is the active form of this cofactor (Payne et al., 2015). Since we were able to produce several protein-free prFMN species, we sought to compare them in the Fdc1 activation reaction, as well as to determine the affinity of Fdc1 to free prFMN. First, we observed that purified apo-Fdc1

prFMNradical could be activated in the presence of PA4019 (B1) or by D-preparation (prFMNiminium enriched) following DT reduction and air oxidation (Fig. 5.5a). After 5 min activation, Fdc1 activity was ~25% higher in the presence of the free prFMN cofactor. Surprisingly, we found that

prFMNradical Fdc1 was also activated by prFMN forms PA4019 (B1) or D under aerobic conditions without DT reduction (Fig. 5.5a). The observed Fdc1 activity without DT reduction was significant (1-2.5 U/mg), although it was 2-3 times lower than that after activation following DT reduction. This suggests that Fdc1 can also bind oxidized prFMN cofactors, but appears to have higher affinity to prFMNred. Fdc1 activation under aerobic conditions (without DT reduction) was stimulated by addition of orthophosphate (apparent Km 6.7 ± 1.1 mM) and divalent metal cations

(Mn2+, Ni2+, or Co2+) (Figures 5.5b,c).

Fdc1 activation by AF1214prFMNradical required longer incubation time likely due to higher affinity of AF1214 to prFMN, but after 60 min activation, Fdc1 showed activity comparable to that obtained with PA4019prFMNradical or D-preparation (Fig. 5.6a). With protein-free prFMN, Fdc1 was activated under aerobic conditions by addition of D-preparation and the preparation enriched with C1′-ene-prFMNiminium (F-preparation), whereas the preparation enriched with C1′- hydroxylated prFMN (E) and FMN showed no activation (Fig. 5.6a). In these Fdc1 activation experiments, F-preparation appeared to be more efficient than D-preparation.

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Fig. 5.5. Activation of Fdc1 by UbiX-bound or protein-free prFMN. (a) Fdc1 activity after short incubation with dithionite (DT; 5 min) and the indicated prFMN forms followed by air oxidation and decarboxylation assay with cinnamic acid as substrate; (b) Effect of orthophosphate on Fdc1 activation by prFMNiminium followed by decarboxylation assay with cinnamic acid as substrate; (c) Effect of divalent metal cations (0.2 mM) on Fdc1 activation by prFMNiminium and decarboxylation activity against cinnamic acid. (d) Direct-injection MS spectrum of prFMN cofactor extracted from Fdc1 activated by the green-colored F-preparation. Abbreviation for prFMN forms: B1, PA4019-bound prFMNradical; D, prFMNiminium; F, C1′- ene-prFMNiminium; G, C2′-hydroxylated-prFMNiminium; H, C1′,C2′-dihydroxylated prFMN.

Cofactor saturation experiments revealed that Fdc1 is saturated by micromolar concentrations of D- or F-preparations with apparent Km values 9-19 μM, indicating high affinity of this enzyme to both free species (Fig. 5.6b). Moreover, Fdc1 activated by D- or F-preparations showed different ratios of decarboxylation activities against cinnamic and α-fluoro-cinnamic acids (2 and 4, respectively), suggesting that different cofactors (D and F) were involved in these reactions (Table S7). However, it appears that F has no azomethine ylide required to catalyze the

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1,3-dipolar cycloaddition (Fig. 5.4a). Therefore, we performed LC-MS analysis of the cofactor extracted under strictly anaerobic conditions from Fdc1 activated by the F-preparation.

Fig. 5.6. Fdc1 activation by UbiX-bound and free prFMN cofactors. (a) Decarboxylase activity of Fdc1 after 5 min (grey bars) or 60 min (white bar) incubation with a 4 times molar excess of UbiX-bound (PA4019 or AF1214) or free prFMN forms under aerobic conditions (without DT reduction). Free prFMN forms were produced by overnight aerobic incubation of C at 4oC and at different pH (F, D, and E are the major components of the free prFMN preparations at pH 2, pH 7, and pH 10, respectively). (b) Saturation of purified apo-Fdc1 by the D- and F-preparations, respectively. The upper X axis shows prFMN concentrations, calculated in respect to the original concentration of C in the preparation prior to oxidation, whereas the lower X axis indicates the prFMN-Fdc1 molar ratio. Apparent kinetic parameters (Km and Vmax) for both preparations are also shown on the graph (R2 = 0.94 and 0.96).

These experiments revealed a significant [M+H]+ adduct with m/z = 541.17 (Fig. 5.5d), corresponding to the monoisotopic mass of C2′-hydroxylated prFMNiminium (form G), which possesses azomethine ylide (Fig. 5.4a). Moreover, this prFMN extract also activated apo-Fdc1

(0.1-0.15 U/mg), suggesting that G is likely the functional prFMN form in the F-preparation.

However, the exact molecular structure of G remains to be confirmed. Thus, our results reveal that apo-Fdc1 can be activated by addition of free prFMN cofactors with/out reduction by DT.

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Fig. 5.7. Schematic overview of the E. coli enzymes involved in prFMN biosynthesis and associated pathways. Biosynthetic enzymes are shown in red. The novel link between the E. coli isoprenoid (MEP) pathway and prFMN biosynthesis is shown by yellow arrows. Chemicals and enzymes involved in the reactions are listed below: Q or CoQ8, ubiquinone; cyt C, cytochrome C; DMA, dimethylallyl alcohol; DMAP, dimethylallyl-monophosphate; DMAPP, dimethylallyl-pyrophosphate; FMN, flavin mononucleotide; FPP, farnesyl-pyrophosphate; GPP, geranyl-pyrophosphate; 4HB, 4-hydroxybenzoate; 2OPPh, 2-octaprenyl-phenol; 3OP-PP, 3-octaprenyl-pyrophosphate; 3OP4HB, 3-octaprenyl-4- hydroxybenzoate; UbiD, 3-octaprenyl-4-hydroxybenzoate decarboxylase; UbiX, FMN prenyltransferase; prFMN, prenylated-flavin mononucleotide; ThiM, hydroxyethylthiazole kinase; Components of the electron transport chain (ETC): I, NADH dehydrogenase; II, succinate dehydrogenase; III, cytochrome C reductase; IV, cytochrome C oxidase.

5.5. Significance

UbiX-like proteins from all major phylogenetic groups possess FMN prenylation activity, but exhibit different affinity to prFMN, with some proteins retaining this cofactor after formation.

Using optimized reaction conditions for FMN prenylation, we produced protein-free prFMN and proposed two new species: protonated prFMNradical (C) and C1′,C2′-dihydroxylated prFMN (H).

Fdc1 activation assays with UbiX-bound and free prFMN showed significant enzyme activation under aerobic conditions (without reduction by DT), but anaerobic activation resulted in higher

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Fdc1 activity. These experiments also revealed a high affinity of Fdc1 to prFMN cofactors, with full enzyme saturation observed at micromolar cofactor concentrations. We found that recombinant expression of UbiXprFMN is stimulated by the addition of prenol to E. coli cultures, suggesting that prFMN biosynthesis is limited by intracellular DMAP. Our screens have shown that in E. coli cells,

DMAP can be produced by direct phosphorylation of added prenol by the ThiM kinase, and by dephosphorylation of DMAPP by the Nudix hydrolases NudJ and NudF (Fig. 5.7). Therefore, the biosynthesis of prFMN from DMAPP provides an additional metabolic link between the UbiX-

UbiD pair and isoprenoid biosynthesis pathways, which were already connected through the UbiD- mediated decarboxylation of 3-octaprenyl-4-hydroxybenzoate in ubiquinone biosynthesis (Fig.

5.7). Future studies on prFMN biosynthesis and prFMN-dependent enzymes will likely reveal additional biological roles for this novel cofactor.

Acknowledgements

We thank Dr. David Leys (University of Manchester) and Dr. Alexei Savchenko (University of Toronto) for providing the A. niger Fdc1 and UbiX expression plasmids, respectively. This work was supported by the NSERC Industrial Biocatalysis Strategic Network grant and Genome Canada

(by the Government of Canada through Genome Canada and the Ontario Genomics Institute, 2009-

OGI-ABC-1405). We also acknowledge the help from the BioZone Mass Spectrometry facility

(University of Toronto, Canada) for LC-MS analyses.

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CHAPTER 6 – SUMMARY, SIGNIFICANCE, AND FUTURE WORK

6.1 Summary

The major goal of this thesis was to characterize the biosynthesis and cross-feeding of cofactors in anaerobic microbial communities. In addressing these goals, four objectives were proposed and pursued (reported in 4 chapters), along with a side project (presented in Appendix

B). The major conclusions from each Chapter are summarized below.

Objective 1 (Chapter 2): Identify functional cobamide prosthetic group in RDase using BN-

PAGE and LC-MS

BN-PAGE is an established method for functional characterization of RDases in dechlorinating cultures. In this thesis, an extended function was developed, which employs BN-

PAGE and downstream LC-MS analysis to identify the native prosthetic groups in the Dsf PceA and Geo PceA, which are purinyl-cobamide and Factor IIIm, respectively. Finding purinyl- cobamide as the native prosthetic group of Dsf PceA assigns unsubstituted purine an exclusive biological function and justify its role as a natural product.

Objective 2 (Chapter 3): Experimentally verifying the metabolic annotation of Dehalobacter restrictus, a key organism responsible for the global halogen cycle, especially for biodegradation of chlorinated ethanes and chlorinated aromatics.

Dehalobacter spp. play an essential role in the global halogen cycle. However, its metabolism and essential nutrient requirements remain poorly understood. Based on available genome annotation and proteomic data, the metabolic annotations in central metabolism and biosynthesis for amino acids and cofactors were experimentally verified. This thesis validated its

123 broken TCA cycle and functional biosynthetic pathway for corrinoid and heme, and annotated the functional serine salvage pathway via threonine. CF, a strong inhibitor for corrinoid-producing acetogens and methanogens, may indirectly conserve corrinoid autotrophy in some CF-respiring

Dehalobacter strains. Also, the results of the experimental verification revealed that the accuracy of metabolic annotation can be significantly enhanced by incorporating enzyme promiscuity and cofactor availability.

Objective 3 (Chapter 4): Identifying the essential nutrients enabling Dehalobacter growth using an omics-integrated constraint-based metabolic model.

An unsolved problem of Objective 2 was that Dehalobacter possesses a complete, complementary set of genes for amino acid biosynthesis, but requires amino acid supplementation for growth. Therefore, a constraint-based metabolic model of Dehalobacter spp. was constructed using the annotated genome and available proteomics data. The model considers cofactor availability and enzyme promiscuity. 13C-stable isotope probing, growth assays, and metabolite analysis of a pure Dehalobacter culture were applied to validate the uncertain constraints. The model resolved that the amino acid dependency results from defects in NADPH regeneration and the classical serine biosynthesis pathway, and diagnosed that malate supplementation can replenish the NADPH pool via a NADP-dependent malic enzyme. Interestingly, data from the metabolite analysis and modeling suggest malate consumption along with pyruvate export, portending a mechanism of interspecies malate-pyruvate shuttle enabling intercellular NADH/NADPH exchanges in dechlorinating consortia.

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Objective 4 (Chapter 5): Elucidate the biosynthetic origin of prenylated FMN, an essential cofactor for ubiquinone biosynthesis, biological decomposition of lignin monomers, and biodegradation of aromatics under anaerobic conditions.

Prenylated FMN (prFMN) is involved in the biosynthesis of ubiquinone, bio- decomposition of lignin monomers, and anaerobic biodegradation of aromatic pollutants.

Realizing prFMN biosynthesis is the key to manipulate prFMN-dependent de/carboxylases both in vitro and in vivo. Nevertheless, the biosynthetic origin for DMAP, the prenyl donor of prFMN, and the biochemical properties of the free prFMN species remain unknown. This chapter described that E. coli produces DMAP by phosphorylation of prenol and dephosphorylation of DMAPP, revealing a novel metabolic link between the isoprenoid pathway and ubiquinone biosynthesis.

Using optimized reaction conditions, several protein-free prFMN forms were produced and biochemically characterized, including a red-colored, protonated prFMN radical. These insights into the biosynthesis and properties of prFMN will facilitate future studies on metabolic engineering of lignin and on bioremediation of aromatic pollutants.

Side project (Appendix B): Sustainable and complete TCE dechlorination in

Dehalococcoides-enriched cultures under acidic conditions independent of cobalamin supplementation.

In this cooperative project with Luz Puentes and Olivia Molenda, experiments were conducted to characterize the optimal conditions for complete TCE dechlorination to ethene by

Dehalococcoides-containing KB-1 cultures, independent of cobalamin supplementation. Based on the metagenomic analysis using cobalamin-synthetic genes as biomarkers, we found that a homoacetogen Acetobacterium sp. is the only native cobalamin producer in the KB-1 culture.

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Moreover, using MeOH as the sole electron donor and maintaining the pH at neutral conditions are the key conditions to selectively enrich for the cobalamin-producing Acetobacterium, but not to enrich for the [p-cresol]-producing Sporomusa. Furthermore, it was determined that

Dehalococcoides RDase activity in the KB-1 culture is significantly decreased at acidic pH, and is completely lost below pH 5.0. Finally, the addition of DMB into the KB-1 culture partially enhanced the VC dechlorination and alleviated the drop of pH, which is accomplished by the inhibition of acetogenesis by Sporomusa using methanol and thus enhances methanogenesis.

Synthesis of the discoveries in this thesis

Fig. 6.1. Synthesis of this thesis: biodegrader trinity of chlorobenzene (Cl-benzene) in anaerobic groundwater. (1) Malate-mediated malate-pyruvate shuttle enables cellular redox-balancing in Bacteroides and NADPH regeneration in Dehalobacter. (2) Biosynthesis of cobamide (B12) for RDase- mediated chlorobenzene (Cl-benzene) dechlorination to benzene. (3) Biosynthesis of prenylated FMN (prFMN) by UbiX for AbcA-mediated benzene carboxylation to benzoate, and downstream catabolism to CO2. Abbreviations: DMAP, dimethylallyl monophosphate; FMN, flavin mononucleotide; Lac, lactate; Mal, malate; Pyr, pyruvate; OAA, oxaloacetate. 126

The three projects present in this thesis are seemingly not related; in fact, they are all entangled together in the environment. Some Dehalobacter restrictus strains use chlorinated benzenes in addition to or instead of chlorinated ethenes and ethanes studied here. In groundwater contaminated with chlorobenzenes, one can envisage that lactate-fermenting Bacteroides (Bac),

OHRB Dehalobacter (Dhb), and benzene-fermenting Peptococaceae (Pep) are working in concert

- to completely degrade chlorobenzenes to CO2 and Cl (Fig. 6.1). First, the malate-mediated interspecies malate-pyruvate shuttle enables redox-balancing in Bac and NADPH regeneration in

Dhb for anabolism. Subsequently, Dhb synthesizes cobamides as the prosthetic group of RDase for chlorobenzene dechlorination to benzene. Finally, Pep synthesizes prFMN as the coenzyme of benzene carboxylase AbcA for benzene carboxylation to benzoate, and downstream catabolism to

CO2.

The three cofactors studied in this thesis, cobamides, NADPH and prFMN, each possess distinctive biological functions (Table 6.1), particularly relevant to anaerobic communities.

Nevertheless, the magnificent biological diversity must be underlined by biochemical rules and reveals some unifying features (Berg et al., 2015). One of the shared features for these cofactors is their required amount in cells. When cofactors solely serve for anabolism, the required amount can be extremely low, like the case of the cobamide requirement in Lactobacillus, which is at sub- nanomolar levels (Chapter 3). When the cofactors are involved in respiratory catabolism as the coenzyme, the requirement can rise to sub-micromolar levels, like the case of the cobalamin requirement of Dehaloccocoides (Appendix B). Finally, when the cofactors are directly involved in respiratory reactions and serve as the energy shuttle, like NADPH, the requirement can climb up to sub-millimolar levels (Chapter 4).

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Table 6. 1. Summary of the function and symbiotic interactions of cobamides, NADPH, and prFMN in literature and proposed in this thesis (in bold).

Required Function/ Cofactor Example amount Symbiotic interactions Anabolism Methionine/Lipid biosynthesis PCE dechlorination Catabolism (Chapter 2) Antioxidant Glutathione-cobalaminc Function Lactate fermentation by Detected Metabolic valve Bacteroides; formate concentration (also heme; MoCo) utilization by Dhb (Chapter in KB-1 3) (µg/L; Appendix B) Modulate enzyme Incomplete TCE Cobamide d and in function dechlorination by Dhc VcrA Lactobacillus Acetobacterium- isolate culture (1). Enable growth Dehalococcoides (ng/L; (Appendix B) Acetobacterium-Sporomusa Chapter 3) (2). Inhibit growth Symbiotic (Appendix B) interactions Inhibit acetogenesis/Enhance (3). Adjust pH methanogenesis (Appendix B) Sinorhizobium meliloti (4). Endosymbiosis survival in plant root nodulee Amino acid biosynthesis Anabolism (Chapter 4) Intracellular Catabolism Oxygenasesf Function concentration NADPH Antioxidant Glutathione reductaseg in E. coli (µM)a redox balancing Transhydrogenaseh Symbiotic Interspecies Bacteroides-Dhb (Chapter interactions redox balancing 4) Anabolism Ubiquinone biosynthesisi Intracellular concentration Lignin monomer bio- prFMN Function decomposition; anaerobic (0.2 µM in Catabolism E. coli)b aromatic degradation (Chapter 5) a(Hanemaaijer et al., 2015); bUnpublished data; c(Birch et al., 2009); d(Bi, 2015); e(Taga and Walker, 2010); f(Price et al., 2004); g(Famili et al., 2003); h(Adrian and Loeffler, 2016); i(Marshall et al., 2017).

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The other shared feature for the three cofactors is their biological functions in cells.

Although each cofactor is fundamentally used in different types of enzyme reactions, they are required in cellular anabolism and catabolism. In some challenging biochemical reactions, like methionine synthase-mediated homocysteine methylation, the enzyme complex simultaneously employs NADPH, FMN, and cobamide for catalysis, in which the NADPH is used to reduce FMN to FMNH2, and the FMNH2 is used to generate a co(I)balamin supernucleophile, shuttling the methyl group form methyl-tetrahydrofolate to homocysteine (Wolthers et al., 2007). Interestingly, recent studies have shown that cobamide and NADPH also serve as efficient antioxidants against reactive oxygen species (ROS) in cells. Given that (i) the prFMN precursor riboflavin is a well- known electron scavenger/antioxidant (McCormick, 1989) and (ii) prFMN possesses outstanding redox versatility, prFMN likely also serves as an antioxidant in cells.

6.2 Significance

In this thesis, three main lines of research, along with a side project, were applied to examine how the biosynthesis and interspecies exchanges of cofactors can affect the functionality and population structure of microbial communities. The overarching messages suggest that cofactor cross-feedings from specialized producers can be a common phenomenon in environments, as evidenced by microbes in enriched mixed cultures which often possess incomplete biosynthetic pathways of cofactors like cobamides and MoCo. This nutrient interdependency shapes defined population structure of microbial communities through the selection of suitable symbiotic partners. In addition, cofactor availability channels the pan- metabolomic pathways in a microbiota. Intriguingly, a chemical with a pM to nM intracellular concentration can affect the fate of the metabolites with a million times higher concentration, resembling a bee leading the way for an elephant. Finally, cofactors are well-known modulators

129 in cellular redox homeostasis. Standing on the frontier of microbial symbiosis, this study discovered that anaerobes can overcome the imbalanced redox (NADH/NADPH) via trading organic acids intercellularly, as evidenced by the interspecies malate-pyruvate shuttle in the ACT-

3 consortium. This newfound cross-feeding interaction reflects the outstanding metabolic flexibility of microbial consortia, which is likely the reason that microbial consortia are often more robust than isolated microbes. As a trade-off, these co-adapted symbiotic microbes suffer from genome reduction, sacrificing their ability to live alone, unless a complex of required nutrients, like yeast extracts, are provided. In conclusion, the discoveries described in this thesis are simply the kick-off of the research efforts to explore the great diversity of cofactor cross-feeding interactions in the biosphere. Realizing these metabolic interdependencies is the key to manipulate the microbiomes for better outcomes in bioremediation and industrial biotechnology.

6.3 Future Work

The discoveries in this thesis have generated many new interesting ideas, especially the following:

1. Characterizing the native cobamide structure of Dehalobacter restrictus strain CF.

In Chapter 3, using DMB as the artificial lower ligand, I confirmed that Dehalobacter restrictus strain CF retains corrinoid autotrophy, but the structure of its native cobamide remains unknown. This is due to the slow growth and low cell yield of the Dehalobacter culture.

Nevertheless, the phylogenetic analysis of CobT suggested that Dhb and Dsf CobTs share significant similarity, indicating that Dehalobacter also synthesizes purinyl-cobamide if

Dehalobacter can also synthesize unsubstituted purine. Since accumulating sufficient biomass of

Dehalobacter is challenging, the addition of both unsubstituted purine and cobinamide to the

130 growth medium of Dehalobacter isolate culture likely enhances the yield of cobamide for LC-MS analysis. Another possibility is to use heterologous expression of Dehalobacter CobT to test its activity using unsubstituted purine as the substrate.

2. Elucidate the amino acid exchange between Bacteroides sp. (Bac) and Dehalobacter (Dhb) in the ACT-3 consortium.

An interspecies malate-pyruvate shuttle was annotated in Chapter 4, which enables the

NADPH regeneration in Dhb. However, the amino acid exchanges between Bac and Dhb remain unsolved. Since the closed genome of Bac reveals incomplete pathways for many amino acids,

Dhb likely cross-feeds the required amino acids to Bac, albeit we did not detect amino acids in the

ACT-3 supernatant. A directed method was used to isolate Bac using the medium with nutrients predicted by our model, and characterize its amino acid requirements. After that, a Bac-Dhb coculture can be constructed. Previous attempts to isolate the Bacteroides sp. were not successful, likely due to the lack of essential nutrients for the Bacteroides sp. to grow. Based on the genome annotation, the literature, and this thesis, some C4 organic acids, amino acids, menaquinone, heme, and cobalamin are required to sustain Bacteroides isolate growth. Moreover, since the Bacteroides sp. is a Gram-negative bacterium and is tolerant to high concentrations of CF, adding vancomycin and CF to the growth medium can selectively enrich Bacteroides and inhibit the growth of Gram- positive Dhb and Clostridium sp.

3. Explore the catalytic diversity of prFMN-dependent enzymes.

In Chapter 5, my colleagues and I have developed the method to produce and to measure prFMN for UbiD de/carboxylase activation. The production of sufficient quantity prFMN has enabled future studies on prFMN-dependent enzymes, which is still a nascent field. FMN and FAD

131 have been found to be involved in several enzyme reactions with distinct catalytic mechanisms.

Similarly, prFMN possesses outstanding redox versatility, which can participate in the biochemical reactions of many oxidoreductases and radical enzymes other than UbiD de/carboxylases. An applicable method is to incubate the prFMN cofactors with the cell lysates of model organisms like E. coli, yeast, and Eukaryotic cells, and apply the BN-PAGE method proposed in Chapter 2 to corelate the proteins that bind prFMN.

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Appendix

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Appendix A. Recipe of Optimized Defined Mineral Medium for Isolate Cultures 1. Stock solutions

MM1: Phosphate buffer (100x; 2 mM final concentration)

KH2PO4 27.2 g K2HPO4 34.8 g

Adjust pH to 7.0 by KOH powder. Make up to 1 L with distilled H2O (dH2O).

MM2: Salt solution (100x)

NH4Cl 53.5 g (10 mM final concentration) . CaCl2.6H2O 7.0 g (or 4.79 g CaCl2 2H2O; unknown final concentration due to precipitation after autoclaving) FeCl2.4H2O 2.0 g (0.1 mM final medium concentration)

Make up to 1 L with dH2O. Note that the FeCl2 tends to oxidize and precipitate. The precipitation problem can be minimized by keeping this solution anaerobically (sparge with N2 and kept the bottle positively pressurized).

MM3: Trace Minerals (500x; MM3 is prone to precipitate with phosphate and FeS, so add it right before inoculation)

H3BO3 0.3 g ZnCl 0.1 g Na2MoO4.2H2O 0.1 g NiCl2.6H2O 0.75 g MnCl2.4H2O 1.0 g CuCl2.2H2O 0.1 g CoCl2.6H2O 1.5 g Na2SeO3 0.02 g

Al2(SO4)3.18H2O 0.1 g

Add 1 ml H2SO4 per L to dissolve all components. Make up to 1 L. Autoclave and sparge with N2 to minimize oxygen. Keep it positively pressurized and anaerobic.

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MM4: Magnesium chloride solution (500x; 0.5 mM final concentration)

. MgCl2 6H2O 50.8 g/L

Use this to minimize sulfate reductions. This is normally used for our standard media.

MM5: Redox indicator (1000x; 1 mg/L final concentration)

Resazurin 1 g/L

MM6: Saturated bicarbonate (100x; ~1.3 M for supernatant at room temperature; final concentration 13 mM)

Mix ~ 20 g NaHCO3 in 100 ml dH2O. Pour slurry into narrow-necked 160-ml serum bottle, cover with foil and autoclave. After autoclaving, sparge with N2 for 15 minutes while cooling. Seal with sterile black butyl rubber stopper and crimp. The preparation will have undissolved NaHCO3 in the bottom. Please use the supernatant and do not resuspend the precipitate and keep the bottle positively pressurized.

Solubility of NaHCO3 ~1.3 M at 25oC (https://en.wikipedia.org/wiki/Solubility_table) pH calculation for bicarbonate buffer: https://en.wikipedia.org/wiki/Bicarbonate_buffer_system Under a 20% CO2 head space with balancing N2 The pH in medium with 13 mM NaHCO3 (100x dilution) is 6.68; The pH in medium with 36 mM NaHCO3 is 7.

MM7: Vitamins (10,000x for mixed cultures and 1,000x for isolate cultures; vitamins are easily degraded at room temperature, so add it right before inoculation) Biotin 0.02 g Folic acid 0.02 g Pyridoxine HCl 0.1 g Riboflavin 0.05 g Thiamine 0.05 g Nicotinic acid 0.05 g Pantothenic acid 0.05 g PABA 0.05 g Cyanocobalamin 0.05 g (A cyanocobalamin-free MM7 is available in Freezer) Thioctic (lipoic) acid 0.05 g Coenzyme M 1.0 g (Not necessarily in MM7)

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Make up to 1 L. Adjust pH to 7.0 with 2N NaOH. Store in one or two mL aliquots frozen. Dilute the stock 100x with Milli-Q water. Filter sterilize 100x stock into rubber-stopper-sealed sterile 160-ml serum bottle and sparge with sterile O2-free N2 for 15 minutes. Keep the bottle positively pressurized to maintain the anaerobicity and stored at 4oC.

MM8: Amorphous Ferrous Sulfide (100x, final concentration 0.8 g/L; please refer to Dr. Fei Luo’s method for high quality FeS)

2. Procedure for making 1 L of medium

In a 1 L serum bottle, add: 960 ml ddH2O 10 ml MM1 10 ml MM2 2 ml MM4 1 ml MM5

Heat the water to ~90oC by microwaving (not boiled) before adding to the bottle. Cap the bottle with the gastight cap with silicon stopper in middle. Connect the atmosphere of the serum bottle to a vacuum pump with needle, and vacuum for 1 min (note that a liquid-collecting flack should be set between the vacuum pump and the bottle to prevent the medium going into the pump). After, de-vacuum the bottle with O2-free N2/CO2. Repeat this step twice, and place the bottle on ice bath to cool while keeping sparging with O2-free N2/CO2 for 10 min. After, add:

10 mL MM6 10 mL MM8 (1 mL for isolate culture Add 10 ml of each MM8 and MM6 to the basal medium by syringe (these two solutions are kept anaerobic and sterile in 160-ml serum bottles with crimped black butyl rubber stoppers). Note that only 1 mL of MM8 is required for isolate cultures. The medium (1 L) prepared following this protocol will be fully reduced with 1 mL of MM8. Transfer the bottles to a glove box and wait few days until the FeS and insoluble salts precipitate (skip this step in the medium used for isolate cultures), aliquot the supernatants to the smaller serum bottles used for culture growth. Seal and crimp the bottles followed by autoclaving. When cooling to room temperature, add 100x MM7 and 500x MM3 to the bottles by syringe with sterile filter. Check the final pH (sample a small aliquot out of the bottle and measuring with a portable pH meter). The pH should be around 7.

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Appendix B: Sustainable and complete TCE dechlorination in

Dehaloccocoides-enriched cultures independent of cobalamin supplementation and with varying pH.

Luz A. Puentes Jácome, Po-Hsiang Wang, Olivia Molenda, Ahsanul Islam, Kirill Krivushin and

Elizabeth A. Edwards

A version of this chapter has been written into a manuscript in preparation to be submitted to

Environmental Science & Technology.

Abstract

TCE is a ubiquitous groundwater pollutant due to historical improper disposal. Successful TCE bioremediation with microbial consortia enriched with cobalamin-dependent organohalide respiring bacteria have been demonstrated in many field sites worldwide. However, TCE bioremediation under acidic conditions is challenging and often comes along with the accumulation of a more toxic intermediate VC. Also, cobalamin is mostly included in the medium used to maintain the activity of the dechlorinating cultures in laboratory, which can reduce their performance in the field sites. In this study, we firstly constructed a methanol-fed,

Dehalococcoides (Dhc)-containing mixed culture TCE/M_Vit(-) capable of complete TCE dechlorination and independent of exogenous vitamin amendments. We then identified the native cobalamin producer in TCE/M_Vit(-), and discovered that neutral pH and TCE addition are required to selectively enrich the cobalamin-producing Acetobacterium population. Additionally, our results suggested that an Acetobacterium population with a cell density of 106 cells/mL can provide sufficient cobalamin (~1 µg/L) for complete TCE dechlorination at millimolar levels.

Finally, we identified that the VC accumulation in Dhc-containing cultures under acidic conditions

153 is due to loss-of function of Dehalococcoides RDases. At pH 5.5, Dhc-containing cultures can dechlorinate VC to ethene with a residual dechlorination rate four times lower than at pH 7.0.

Interestingly, although cobalamin amendment cannot enhance the VC dechlorination at pH 5.5, our data suggests that cobalamin can reduce pH drop via inhibiting acetogenesis and boosting methanogenesis. Altogether, this study brings a vitamin-autotrophic dechlorinating culture into play and reflects that pH is the primary index for chloroethene bioremediation in field sites.

Introduction

Groundwater contamination with chlorinated solvents is a worldwide environmental issue

(Bhatt et al., 2007; Field and Sierra-Alvarez, 2004). Bioaugmentation with specialized microbial cultures has been a successful remediation approach at several contaminated sites worldwide

(Major et al., 2002; Scow and Hicks, 2005). These cultures can reductively dechlorinate harmful compounds such as trichloroethene (TCE) via vinyl chloride (VC) to non-toxic ethane; successful dechlorination is dependent on the close coordination of different resident microbial populations.

Often, these cultures harbor methanogenic and acetogenic populations along with dechlorinating populations. The dechlorinating populations are composed of either facultative or obligate organohalide-respiring bacteria (OHRB). The microbial composition of OHRB-enriched cultures is dependent on the growth conditions, such as the choice of electron donors and of carbon sources, pH (Yang et al., 2017), and vitamin supplementation (Men et al., 2014; Men et al., 2017; Yi et al.,

2012). Obligate OHRB, including Dehalococcoides and Dehalobacter, often prosper in communities abundant with H2- and acetate-producing microbes (Richardson, 2016).

In situ vitamin amendments is cost-prohibitive in some developing countries. Ideally, the microbial cultures of bioremediation can be vitamin-autotrophic. In this paper, we report the

154 successful adaption of TCE-fed KB-1 culture grown without vitamin amendments and of VC-fed

KB-1 cultures grown at lower pH (5.5 and 6.0). We firstly studied the optimal growth conditions for the native cobalamin producer to support complete TCE dechlorination by Dehalococcoides.

The BLAST results of the KB-1 pan-genome and 16S rRNA-based microbial population analysis assigned the cobalamin producer to an Acetobacterium carbinolicum strain, which is only enriched when the cultures are fed with chlorinated solvents and are maintained at neutral pH. Also, the cobalamin content in the culture is proportional to cell density of the Acetobacterium population.

Subsequently, we elucidated the reasons for the reduced VC dechlorination rates at low pH, which is due to loss of activity of Dehalococcoides-dechlorinating RDases at below pH 5 but not cobalamin shortage. Nevertheless, we found that cobalamin/DMB can specifically inhibits acetogenesis by acidophilic Sporomusa and thus boosts methanogenesis, preventing further pH drop. These experimental results on microbial enriched cultures brings valuable implications for field site remediation of chloroethenes.

Materials and Methods

Chemicals. Chemical reagents were purchased through Sigma-Aldrich Canada (Oakville, ON,

Canada), Fischer Scientific Canada (Ottawa, ON, Canada), and BioShop (Burlignton, ON,

Canada) at the highest purity available. Gases were purchased from Praxair (Mississauga, ON,

Canada). Phenyl-cobamide was provided by Prof. Frank E. Löffler (University of Tennessee

Knoxville, USA).

Medium preparation. A phosphate buffered pre-reduced mineral medium (Puentes Jacome and

Edwards, 2017) was used to maintain enrichment cultures and set up new experiments, along with amorphous iron (II) sulfide (FeS) slurry (~0.8 g/L) used as the reductant (Wang et al., 2016). If

155 required, filter-sterilized vitamins (as described in Edwards and Grbić-Galić (1994)) were supplied with or without the presence of vitamin cyanocobalamin. The medium was autoclaved and sparged with a gas mix containing 80% N2 and 20% CO2. The pH of the medium was 6.9 ± 0.1. If required, the pH of the medium was adjusted using a 6N HCl solution. Methanol was always used as the electron donor. TCE was fed in solution with methanol and VC (gas) were fed using the gastight syringe. The TCE solution was prepared at a ratio of 1 to 5 electron equivalents of acceptor to donor.

Enrichment cultures. The preparation of the initial TCE-dechlorinating enrichment cultures grown and maintained without the addition of exogenous vitamins is described in chapter 6 of

Islam (2014).(Islam, 2014) These cultures were set up with inoculum from a 4 L TCE-to-ethene- dechlorinating culture derived from the KB-1 parent culture known as TCE/M_parent. After ~ 2 years of enrichment which included the adaptation of the KB-1 microbial community to growth without vitamin amendments, triplicate cultures (100 mL) maintained with vitamin amendments

(controls), without vitamin amendments, and without cobalamin were each scaled-up to 700 mL.

These cultures are referred to as TCE/M_Vit(+), TCE/M_Vit(-), and TCE/M_B12(-). Adaptation of the KB-1 culture to the reductive dechlorination of VC to ethene under non-neutral pH conditions (pH 6 and pH 5.5) is described in (Li, 2012) Triplicate enrichment cultures (50 mL) kept at pH 7 (controls), 6, and 5.5 were scaled up to 500 mL. These cultures are referred to as

VC/M_pH7, VC/M_pH6, and VC/M_pH5.5.

Sporomusa growth assays. In this experiment, for a better measurement of cell density, Na2S (0.2 mM), instead of Fe(II)S, was used as the reductant in the medium to avoid turbidity; resazurin was excluded from the medium to prevent the pink color under aerobic conditions. In an anaerobic chamber with an atmosphere of 10% CO2, 10% H2, and balancing N2, the active Sporomusa culture

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(2 mL; OD600nm ~ 1) were sampled into the 2-mL sterile O-ring-capped plastic microcentrifuge tubes, and pelleted down by centrifugation at 10,000 x g for 10 min at room temperature. The supernatants were removed, and the pellets were washed with sterile cobalamin-free defined medium (1 mL) twice. After, the pellets were resuspended with sterile defined medium containing varying cobalamin concentration (0, 5, 10, 25, 50 µg/L) to a final cell density (OD600 nm) of 0.04, and 5 mL of each was transferred into the sterile 10-mL rubber stopper-sealed glass tubes by sterile plastic syringes and needles. The tubes were then sparged with 20% CO2 and balancing N2 to remove H2, followed by the addition of MeOH to a final concentration of 20 mM. In some assays,

H2 (20 % in head space), formate (10 mM), or lactate (10 mM) was added to the medium. The glass tubes were incubated at 30oC for one day, and 0.2 mL of each culture was sampled, and loaded into the 96 wells optical plate (Falcon). The cell density in each well was determined using a plate reader at OD600nm.

Analytical methods. Chloroethenes, ethene, and methane were quantified using gas chromatography with flame ionization detection (GC-FID) as described in Chapter 3.

Cell-extract dehalogenase activity assay. The cell-extract dehalogenase activity assays were performed as described in Chapter 3 (Wang et al., 2016).

Cobamide extraction. The cobamides were extracted and derivatized following previous established protocols with some modifications (Yan et al., 2012; Yi et al., 2012). Briefly, the cultures were harvested, and pelleted down via continuous centrifugation at 9,000 x g for 20 min at room temperature. The pellets were resuspended with 80% MeOH containing 10 mM KCN, and the pH were adjusted to a pH between 5.0 and 5.5 with 3% acetic acid (v/v), transferred into the 2 mL plastic O-ring-capped microcentrifuge tubes, followed by a one-hour water bath at 80oC with four short vortexing every 15 min. After cooling down to room temperature, the tubes were

157 centrifuged at 13,000 x g for 10 min to remove cell debris. The supernatants were transferred to the 2 mL plastic microcentrifuge tubes, incubated in a rotary evaporator until dried, and resuspended by 0.1 mL Milli-Q water for LC-MS analysis.

Cobamide measurements. Cobamides were detected and quantified using Liquid chromatography coupled mass spectrometry (LC-MS) with an Accela HPLC system and a Q-

Exactive mass spectrometer equipped with HESI II sources (Thermo Scientific) as described in

Chapter 2.

Results and Discussion

Fig. S2.1. The MS spectra of the extracted cobamide from the KB-1 cultures. (A) extracted [p- cresol]cobamide and (B) extracted cobalamin. Both cobamides have been derivatized into cyano-form.

The KB-1 cultures grown at different pH reveal a significant difference in the ratio of cobalamin to [p-cresol]cobamide. In the cultures grown at pH 5.5 where VC accumulates, the dominant acetogen is S. silvaceticans. Consistently, the major type of cobamide identified by LC-MS showed an [m+H]+ adduct with m/z = 1303.54 matching to the monoisotopic mass of [p- cresol]cobamide (Fig. S2.1A). The concentration of [p-cresol]cobamide in these cultures is ~88

µg/L, which is ~20 times higher than the concentration of cobalamin (~4 µg/L; [m+H]+ = 1355.58)

(Fig. S2.1B; S2.2A). In contrast, in the cultures grown at pH 7 where VC dechlorinated completely,

158 the concentration of [p-cresol]cobamide and cobalamin are comparable (~25 µg/L). Therefore, under acidic conditions the Dehalococcoides population in KB-1 likely lost the ability to dechlorinate VC due to the shortage of functional cobamide cofactor, cobalamin, for VcrA. Beside cobalamin shortage, the functional pH range of Dhc-RDases remains unknown. The Dhc-RDase dechlorination activity assays are generally performed at neutral pH (Tang and Edwards, 2013;

Wang et al., 2016). We thus tested dechlorination activity in cell extracts of KB-1 grown under acidic conditions at pH 4.75, 5.0, 5.5, 6.0 and7.0.

Fig. S2.2. Incomplete TCE dechlorination by Dehalococcoides under acidic conditions due to both (A) cobalamin shortage and (B) the loss-of-function of RDases. NC, abiotic control.

Fig. S2.3. Cobalamin specifically inhibits Sporomusa growth on methanol. Sporomusa growth on defined medium with methanol as the only electron donor (unless specified otherwise) and supplemented with 0, 5, 10, 25, or 50 µg of cobalamin per liter of medium. Abbreviations: FA, formate; Lac, lactate.

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The dechlorination activity of the KB-1 cell extracts declined sharply from pH 7.0 to 6.0.

At pH < 5.0, the dechlorination activity was completely lost (Fig. S2.2B), which is consistent with our observations of the pattern of dechlorination VC-dechlorinating KB-1 cultures. Altogether, we finally conclude that the incomplete TCE dechlorination at low pH is related to both the loss of function of VcrA/TceA and the absence of cobalamin producer like Acetobacterium to provide functional cofactor for VcrA. According to our previous data of microbial population in the KB-

1, cobalamin-producing Acetobacterium is the main acetogen under neutral conditions, while

Sporomusa is the main acetogens under acidic conditions. However, Sporomusa can efficiently grow under neutral conditions. This discrepancy indicates a potential strategy for Acetobacterium to compete MeOH with Sporomusa. An earlier study reported that the addition of 10 µM DMB in medium inhibited the growth of Sporomusa on methanol as its methanol methyltransferase requires phenyl-cobamide or [p-cresol]cobamide as prosthetic group (Mok and Taga, 2013;

Stupperich et al., 1992; Stupperich and Konle, 1993). Therefore, cobalamin, the cobamide with

DMB as the lower ligand, likely also inhibits Sporomusa growth on MeOH. Consistently, we observed a reduced Sporomusa growth in the cultures grown on the medium with 25 µg/L cobalamin, and the Sporomusa growth is completely inhibited in the cultures grown on medium with 50 µg/L cobalamin (Fig. S2.3). However, the Sporomusa growth can be restored with the addition of either H2, formate, or lactate, suggesting that the inhibition by cobalamin is only specific to MeOH growth, which requires the phenyl-cobamide specific methanol methyltransferase. Unexpectedly, the use of MeOH as the sole electron donor in the KB-1 cultures indirectly conserves the native cobalamin-producing acetogens, enabling the complete TCE dechlorination by Dhc without external vitamin supplements.

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Fig. S2.4. VC dechlorination (A) and methane production (B) in the KB-1 cultures grown at different pH or with DMB treatment. A second feed of MeOH was given at Day 16. The pH of cultures at Day 20 were also shown on (A).

Moreover, although the addition of DMB cannot enhance the VC dechlorination at pH 5.5 (Fig.

S2.4A), the cultures amended with 10 µM DMB possess a better pH buffering capacity than the cultures without DMB amendment (Fig. S2.4B). Consistently, the methane production in the

DMB-amended culture is much higher than those without DMB amendment. These data reveal an unprecedented effect of cobalamin/DMB amendment: buffering pH drop in dechlorinating microbial communities, which is via the inhibition the acetogenesis by Spormusa using methanol, and thus boosting methanogenesis. This discovery also explains the reason that Acetobacterium- enriched KB-1 cultures have a better pH buffering capacity than the Sporomusa-enriched KB-1 cultures, since the Acetobacterium population is only enriched under neutral conditions.

Interestingly, such a trace concentration (~ 50 nM) of cofactor can contribute to significant influences on the pH shift of microbial cultures.

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Appendix C. Isolation of Geobacter lovleyi Strain KB-1

Purity confirmation

The culture was transferred to solid medium matrix. Single colony was picked and transferred to liquid medium for three successive transfers.

Fig. C1. Cell colonies of the Geobacter lovleyi.

Chemical evidence. Geobacter lovleyi only dechlorinates PCE to cDCE. Methanogen transform acetate to methane and CO2. DHC dechlorinates cDCE to ethene. In the isolate culture grown on acetate as the sole electron donor and PCE as the sole electron acceptor, no methane is produced and PCE is only degraded to cDCE, indicate the absence of methanogen and DHC.

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STD

cDCE TCE

PCE

Ten minutes after inoculation

Methane Ethane

Five days after inoculation

Fig. C2. Dechlorination profile of the Geobacter lovleyi isolate culture.

Microscopic evidence. According to the report of Loeffler lab. G. lovleyi is a Gram-negative bacterium which is rod-shaped. The Gram stain microscopic result reveals the G. lovley isolate culture obtained by Luz and Tommy a uniform rod morphology. While all cells are stained pink, indicating the purity of the isolate culture.

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Fig. C3. Gram stain microscopic image of the Geobacter lovleyi isolate culture.

DNA sequencing evidences. The gDNA of the isolate culture is extracted by PowerSoil kit. The

16S rRNA gene was amplified by general primer 27F and 1541R. The amplicon was sequenced by TCAG using Sanger sequencing technology. The amplicon was also digested by restriction enzyme KpnI to test for purity. The result of Sanger sequencing data reveals 100% sequence identity to Geobacter lovleyi sp. KB-1, and the RFLP result reveal identical result from computer prediction. Also, only the predicted two bands appear on the gel with predicted molecular weight.

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Fig. Fig. C4. Sanger sequencing result and nucleotide BLAST result of 16S rRNA gene PCR amplicon obtained from the Geobacter lovleyi culture.

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Fig. C5. Restriction digestion analysis of PCR products of DNA from the G. lovleyi isolate culture with KpnI.

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Appendix D. Supplementary Information for Chapter 3

Supplementary Materials and Methods

All Chemicals were ordered from Sigma-Aldrich at highest purity available unless specified otherwise.

1.0 Detailed medium recipes

(i) Vitamin solution. The core vitamin solution used in this study contains 2 µg/L folic acid, 5

µg/L α-lipoic acid, 2.5 µg/L nicotinic acid, 2.5 µg/L nicotinamide, 5 µg/L 4-aminobenzoic acid, 5

µg/L pantothenic acid, 2.5 µg/L pyridoxine, 2.5 µg/L pyridoxal amine, 5 µg/L riboflavin. The following vitamins: 20 µg/L biotin, 25 µg/L cyanocobalamin, 25 µg/L thiamine, or 25 µg/L heme

(hematin form) were added to the medium where specified in the main text. Heme was first dissolved in 10 mM NaOH to a concentration of 25 mg/L (1,000-fold stock).

(ii) Mixed culture supernatant (MCS). Autoclaved sterile ACT-3 MCS was added to medium to support the growth of the two Dehalobacter strains in dilution-to-extinction transfers. To prepare

MCS, mixed culture ACT-3 was harvested when the culture was steadily dechlorinating 1,1,1- trichloroethane (1,1,1-TCA) to CA (mono-chloroethane), and was centrifuged at 9,000 ×g at 4 °C for 20 min under anaerobic conditions. The supernatant was then filter-sterilized using a Millipore

0.22 µm filter and stored at -20 °C until use.

(iii) Amorphous iron sulfide (FeS) slurry. In an anaerobic chamber, equimolar amounts Na2S

(0.78 g) and FeCl2 (1.27 g)were added to a 160 mL narrow neck serum bottle, and suspended in

100 mL anaerobic Milli-Q water to a final concentration of 100 mM. The bottle was sealed with

167 a butyl rubber stopper and crimped. After sparging with N2 for 20 minutes to get rid of most the

H2S, the black FeS suspension was transferred under anaerobic conditions to a 250 mL polypropylene centrifuge tube (Thermo) and centrifuged at 8,000 x g for 10 min at RT. The supernantant was removed and the slurry resuspended in anaerobic Milli-Q water again. The step was repeated once and the resuspension was transferred back to the serum bottle. The bottle was sealed by a butyl rubber stopper, crimped, and autoclave-sterilized. The final concentration of the

FeS slurry was approximately (~8 g/L).

2.0 Growth conditions for growth assays of electron donors, electron acceptors, and

vitamin requirements

Briefly, 10 mL of minimal mineral medium, main carbon source (5 mM), electron donor (10 mM)

(or 6.4 mL of H2), 1-4 μL of neat chlorinated substrates, and each of L-arginine, L-histidine, and

L-threonine (0.1-0.5 mM) were added to a 25 mL Bellco tube before autoclaving. Acetate, H2, and

1,1,1-TCA or 1,1-DCA were used to grow the isolate cultures in most experiments unless specified otherwise. After autoclaving, 10 µL sterile FeS slurry and vitamin solution (from 1,000-fold stock) were injected to the sealed tubes outside the anaerobic chamber. The serial transfer assays were inoculated with 1 mL of culture (10-1 dilution). The nutrients requirement assays were inoculated with 100 µL of culture (10-2 dilution). To minimize carryover, inocula were pelleted down to remove supernatant, and were resuspended in and equal volume of sterile minimal mineral medium before inoculation. For large-scaled cultivation (1 L) for corrinoid structure characterization by

UPLC-MS and the formate utilization assay, casamino acid (0.5 g/L) (Fisher Scientific) was supplemented to medium.

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3.0 PCR and Restriction fragment length polymorphism (RFLP)

To amplify 16S rRNA gene sequences for RFLP analysis and cfrA/dcrA for purity test, DNA samples from Dehalobacter isolates and mixed culture ACT-3 were extracted using the MoBio

UltraClean® Microbial DNA Isolation Kit. In the PCR reactions, a Thermo PCR master mix was used and the thermocycling program was as follows: (i) for 16S rRNA gene amplification, initial denaturation of 10 min at 95 °C; 40 cycles of 30 s denaturation at 95 °C, 30 s annealing at 50 °C and 2 min extension at 72 °C; and final extension of 10 min at 72 °C; (ii) for cfrA/dcrA amplification, initial denaturation of 10 min at 95 °C; 40 cycles of 30 s denaturation at 95 °C, 30 s annealing at 60 °C and 30 s extension at 72 °C; and final extension of 10 min at 72 °C. General bacterial 16S rRNA primers 27f (5’-AGAGTTTGATCCTGGCTCAG-3’) and 1541r (5’-

AAGGAGGTGATCCAGCCGCA-3’) were used in PCR reactions for RFLP analysis (Löffler et al., 2000). The PCR-amplified 16S rRNA gene product (1 µg) was then digested for 1 h using restriction enzymes NheI-HF and XbaI-HF at 37ºC according to the manufacturer’s standard protocol. Fragments were resolved by electrophoresis for 30 min on 1% agarose gels (100 V).

Specific primers 413f (5’-CCCGAACCTCTAGCACTTGTAG-3’) and 531r (5’-

ACGGCAAAGCTTGC ACGA-3’) for cfrA as well as 424f (5’-AGCACTCAGAGAGC

GTTTTGC-3’) and 533r (5’-CAACGGCCCAGCTTGCAT-3’) for dcrA were used in the PCR reactions for purity test of potential cross-contamination of two Dehalobacter strains in their isolate cultures. Fragments were resolved by electrophoresis for 25 min on 2.5 % agarose gels (90

V).

4.0 Microscopy

For epifluorescence microscopy, 0.1 mL culture samples were centrifuged at 13,000 g for 10 min at room temperature to pellet cells. The pellets were resuspended in 5 µL Milli-Q water and the

169 suspension pipetted onto glass slides for DAPI or Gram-stain. Slides were air-dried, flame-fixed, and stained with DAPI (10 µg/mL) for 10 min. After washing slides to remove stain, a droplet of immersion oil was loaded on each slide and the slides were observed using a epifluorescent microscope (BX 51, Olympus) with a 100x UPlan Apochromat objective, 150 W xenon lamp (Opti

Quip), a blue excitation filter cube (excitation band pass 372 nm; emission barrier filter 456 nm) and 10x focusing eyepiece to check for uniformity of cell morphology (1,000x magnification).

Gram-stain was performed using a commercial kit (Sigma-Aldrich).

Preparation of samples for Scanning and Transmission Electron Microscopy (SEM and TEM) was performed using the service of the Advanced Bioimaging Centre of Mount Sinai Hospital (Ontario,

Canada). For scanning electron microscopy (SEM) observation, the pellets were fixed in 2% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.3) for 2 h and rinsed in buffer. The sample was then dehydrated in an ethanol gradient series and followed by a critical point drying in a Bal-tec

CPD030 critical point dryer. After the drying, the sample was mounted on aluminum stub and coated with gold in a Denton Desk II sputter coater and examined in a Hitachi SU8230 cold FEG

SEM at 5 kV and image processing was performed using the service of Ontario Centre for

Characterization of Advanced Materials (OCCAM) of University of Toronto. Samples for transmission electron microscopy (TEM) were fixed in 2% glutaraldehyde in the same sodium cacodylate buffer, rinsed in buffer, and then post-fixed in 1% osmium tetroxide in buffer. After fixation, samples were dehydrated in an ethanol gradient series followed by a propylene oxide treatment. Subsequently, samples were embedded in EMBed 812 resin, cut on an RMC MT6000 ultramicrotome and stained with uranyl acetate and lead citrate. TEM was carried out using a

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Hitachi HF-3300 at 200 kV and image processing was performed using the service of Ontario

Centre for Characterization of Advanced Materials (OCCAM) of University of Toronto.

5.0 Enzyme activity assays

The preparation of cell extracts was conducted under aerobic conditions except for the RDase activity assays. Fifty milliliters of E. coli strain K-12 culture grown on LB medium or 200 mL of strain CF culture grown on 1,1,1-TCA were harvested, ice chilled, and centrifuged at 9,000 x g for

20 min at 4ºC. The pellets were resuspended in 0.5 mL of 100 mM Tris-HCl buffer (pH 7.5) containing 12.5 mM NaCl, 2.5% glycerol, and 1% digitonin. The suspensions were loaded into a

2 mL O-ring-capped plastic microcentrifuge tube containing 50 μL in volume of 0.1 mm diameter glass beads. To lyse the cell, the tube was vortexed in maximum intensity for 2 min and incubated on ice for 1 min. The step was repeated 4 times. The cell lysates were then centrifuged at 20,000 x g for 15 min at 4ºC to pellet down the cell debris, beads, and unbroken cells.

(i) RDase activity assay. Cell extracts were loaded into 2 mL glass vials which contains 1 mL of

25 mM Tris-HCl buffer (pH 7.5), 2 mM titanium citrate, 2 mM methyl viologen, and 1 mM chlorinated substrates (nominal concentration). The vials were capped, incubated upside down at

RT for 24 h and sampled for GC analysis as previously described (Tang and Edwards, 2013).

(ii) Malate dehydrogenase (MDH) activity assay. The MDH activity assays were conducted following an established protocol (Lin et al., 2015) with some modifications. Twenty five micro liters of cell extracts were loaded into each well of Falcon 96-well microplate which contains 200

µl of 25 mM potassium phosphate buffer (pH 7.5), 1 mM NADH, 0.1 mM EDTA, and 0.1 mM

o MgCl2. The mixtures were first incubated at 30 C for 1 min, and 25 µl of 10 mM sodium oxaloacetate was then loaded into each well to a final concentration of 1 mM. The assays were

171 incubated at 30oC for 10 min. The decrease of absorbance at 340 nm (where NADH absorbs) in each assay was recorded. The assay was calibrated with freshly prepared NADH standards at 340 nm.

(iii) NADP dependent malate enzyme (MAE) activity assay. The conditions of MAE activity assays (Geer et al., 1980) were similar to MDH assay with the following modifications. NADP replaced NADH in the assay buffer. Sodium malate (50 mM) was replaced sodium oxaloacetate to a final concentration of 5 mM in each assay. The increase of absorbance at 340 nm was recorded and the assay was calibrated with freshly prepared NADPH standards at 340 nm.

(iv) Succinate dehydrogenase (SDH) activity assay: The conditions for the SDH activity assays were similar to MDH with the following modifications. Sodium azide (2 mM) (electron transport chain inhibitor), 2,6-dichlorophenolindophenol (DCPIP) (0.2 mM), and phenazine methosulfate

(0.2 mM) replaced NADH in the assay buffer (Kolaj-Robin et al., 2011). Sodium succinate replaced oxaloacetic acid to a final concentration of 1 mM. The decrease of absorbance at 600 nm in each assay was recorded reflecting reduction of DCPIP. The assay was calibrated with freshly prepared DCPIP standards. When oxidized, DCPIP is blue with a maximal absorption at 600 nm; when reduced, DCPIP is colorless.

Serine hydroxymethyltransferase (SerB) and threonine aldolase activity assays were conducted using the purified recombinant SerB from strain CF. Methods for cloning, expression, and purification of SerB are described below and the nucleotide and protein sequences are also provided in Appendix.

(v) SerB activity assay. SerB activity assays were conducted in 0.25 mL reaction mixtures by adding 1 mM L-serine, 1 mM mercaptoethanol, 2 mM tetrahydrofolate (THF), 0.25 mM pyridoxal

172 phosphate (PLP), and 50 µg/mL purified SerB (Buchenau and Thauer, 2004, Ogawa et al., 2000).

Two controls were also prepared, one without enzyme and the other without serine. All mixtures were incubated at 30oC for 75 min. SerB activities were determined by measuring the formation of glycine from L-serine using UPLC-ESI-MS as described below.

(vi) Threonine aldolase activity assay. Threonine aldolase activity assays were conducted in 1 mL reaction mixtures by adding 20 mM L-threonine (Sigma-Aldrich), 50 µM of PLP, 1 mM DTT, and 50 µg of purified SerB to 50 mM phosphate buffer (pH 7.3) (Buchenau and Thauer, 2004,

Ogawa et al., 2000). Two controls were also prepared, one without enzyme and the other without threonine. Threonine aldolase activities were determined by measuring the formation of acetaldehyde from L-threonine using HPLC.

6.0 Cloning of SerB

The SerB gene (locus tag: DCF50_p2888) from Dehalobacter sp. strain CF was cloned into pCOLADUETTM-1 vector (Novagen), between BamHI and PstI restriction sites in the MCS1 cloning site, which encodes an N-terminal His6-tag. SerB was amplified with primers

F_SerB_BamHI (5’-AGTCATTGGATCCAATGGATTACATTCGGAAATATTTAGC-3’,) and

R_SerB_PstI (5’-AGTCATTCTGCAGTTACTTATACAAAGGAAATCTTCC-3’). The standard protocol for Q5® High Fidelity DNA polymerase (New England BioLabs® Inc.) was used, but with an annealing temperature of 58oC and elongation time of 39 seconds. After amplification and purification of the PCR product, 1 µg of amplicon and empty pCOLADUET-1 empty vector were each digested with BamHI-HF (New England BioLabs® Inc.) and PstI-HF (New England

BioLabs® Inc.) by incubating at 37oC for 1 h. After clean-up using a PCR purification kit (Thermo

Scientific), the digested vector was dephosphorylated using Shrimp Alkaline Phosphatase (New

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England BioLabs® Inc.), while the digested amplicon was phosphorylated using T4 polynucleotide kinase (New England BioLabs® Inc.). Ligation was carried out using T4 DNA ligase (New England BioLabs® Inc.) using a 3:1 molar ratio of insert to plasmid and incubating the mixture at 16oC overnight. Chemically-competent E. coli strain DH10β cells were transformed with 5 µL of the ligation mixture. The gene sequence was verified by Sanger sequencing. The sequence verified plasmid was then transformed into the expression host, chemically-competent

E. coli strain BL21 (DE3) Gold (Stratagene). DNA (5’-3’) and protein sequences of recombinant

SerB from strain CF are provided at the end of this file.

7.0 Expression and purification of SerB. E. coli strain BL21 carrying the SerB expression vector was first grown in 10 mL LB medium (BioShop Inc.) with 50 µg/mL of kanamycin, at 37oC overnight. The seed culture was then transferred to 1 L of terrific broth with 50 µg/mL of

o kanamycin in a 2.5 L baffled flask, and grown at 37 C. When the OD600 nm of the culture reached

0.1, the temperature was lowered to 16oC. After another 3 h of shaking, protein expression was induced by the addition of IPTG (to a final concentration of 1 mM) to the culture which continued growing overnight. The cells were harvested at 4oC and resuspended in binding buffer containing

50 µM pyridoxal phosphate (PLP). The resuspended cells were sonicated for 20 min (program 3 s on and 4 s off) in an ice-water bath. The resulting suspension was centrifuged at 58,000 xg for 40 min at 4 oC. The supernatant was loaded onto a Ni-NTA column that was equilibrated with 50 mM

HEPES buffer (pH 7.5) with 0.5 M NaCl, 10% glycerol, 50 µM PLP, and 5 mM imidazole, and washed with 150 mL the same buffer with 30 mM imidazole. The protein is then eluted using the same buffer with 250 mM imidazole and fractions were monitored for protein concentration using standard Bradford assay. The protein purity was verified by SDS-PAGE (Fig. S5).

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8.0 Extraction and derivatization of intracellular thiamine species

In brief, 200 mL of a stationary-phase strain CF culture grown on thiamine-containing (25 µg/L) medium was centrifuged at 9,000 x g for 10 min at 4°C. Cell pellet (50 mg wet weight) or 10 µL of 0.1 mM thiamine standards (to a final concentration of 1 µM) were suspended and diluted in 1 mL of 3.6% perchloric acid (w/v), respectively, followed by a 20 min ice-chilled water bath on a ultrasonic cleaner. The cell lysate was then centrifuged at 16,000 x g for 15 min at 4°C. To derivatize the thiamine species, supernatant (0.5 mL) was mixed with a K3[Fe(CN)6] (12 mM in

3.35 M NaOH) solution (0.1 mL). The mixtures were incubated in the dark at 37°C for 15 min, and then adjusted to pH 7 with 85% phosphoric acid (w/v) for later analysis.

9.0 Analytical procedures

Chlorinated hydrocarbons were measured by injecting a 0.3 mL headspace sample into a Hewlett-

Packard 5890 Series II gas chromatograph (GC) fitted with a GSQ column (30-m-by-0.53-mm

[inner diameter] PLOT column; J&W Scientific, Folsom, CA), as described previously (Grostern and Edwards, 2006). The GC carrier gas pressure was initially 100 kPa, and the oven temperature was programmed to hold at 50°C for 90 s and then increase to 155°C at 30°C/min and then increase to 180°C at 4°C/min and hold for 5 min. Calibration was with external standards.

The amount of acetaldehyde produced from the threonine aldolase activity assay was determined by HPLC using a Dionex Ultimate 3000 system equipped with an Aminex HPX-87H column

(BioRad) connected to a UV detector. Twenty microliters of each sample was injected onto the

175

o column incubated at 60 C, using 5 mM H2SO4 eluent at a flow rate of 0.6 mL/min with the UV wavelength set to 278 nm for acetaldehyde detection.

Thiamine derivatives were detected by high pressure liquid chromatography (HPLC)-

Fluorescence using a Varian Prostar HPLC system consisting of a Model 230 solvent delivery system, a Model 410 AutoSampler, a Model 500 column valve module, and a Model 363

Fluorescence detector (excitation at 365 nm and emission at 435 nm). Separation was performed using a 5 µm Varian Pursuit C18 column (15 cm x 4.6 mm) with a Varian MetaGuard guard column. The mobile phase consisted of two eluants: Mobile phase A was dibasic sodium phosphate

(25 mM, pH 7.0):methanol (90:10, v/v); mobile phase B was dibasic sodium phosphate (25 mM, pH 7.0):methanol (30:70, v/v). To enhance the fluorescence intensity, sodium perchlorate (Sigma-

Aldrich) was added to both solvents to a final concentration of 1 mM. Gradient steps were programmed as follows: 0 min, 0% B; 4.5 min, 13% B; 10 min, 50% B; 15.5 min, 50% B; 19 min,

0% B; 24.5 min, 0% B, with a flow rate of 0.85 mL/min. Injection volume were 20 μL.

Corrinoids and serine were detected and quantified using Liquid chromatography coupled mass spectrometry (LC-MS) with an Accela HPLC system and an Exactive mass spectrometer (Thermo

Scientific), while heme (hematin form) and biotin detection was performed using a Dionex

Ultimate 3000 UPLC and a Q-Exactive mass spectrometer, both equipped with HESI II sources

(Thermo Scientific). System control and data handling were performed using Thermo XCalibur

2.2 software. Separation by liquid chromatography was conducted on a Hypersil Gold C-18 column (50mm x 2.1 mm, 1.9 µm particle size, Thermo Scientific) equipped with a guard column.

LC was performed with 10 uL injections at a flow rate of 0.2 ml/min with a gradient of water containing 2.5 mM ammonium acetate (pH 6.0) (A) into methanol (B), and a column temperature of 30˚C. The gradient was 0 min, 0% B; 0.46 min, 0% B; 0.81 min, 15% B; 3.32 min, 50% B; 4.86

176 min, 90% B; 5.56 min, 90% B; 5.59 min, 0% B; followed by equilibration for 5 min with 0% B.

Corrinoid data collection was done in positive ionization mode with a m/z scan range of 50-150 and 1000-1500 for serine and corrinoid, respectively; resolution 100,000 at 1 Hz, automatic gain control (AGC) target of 5e5; and a maximum injection time of 200 ms. Heme data collection was done in negative ionization mode with a m/z scan range 600-800; resolution 140,000 at 1Hz; AGC target 3e6; and a maximum injection time of 250 ms. Injection volume for heme is 25 µL. Biotin was detected with a different mobile phase with a gradient of water containing 20 mM ammonium acetate and 20mM ammonium hydroxide (A) into acetonitrile (B). The gradient was 0 min, 0% B;

6 min, 100% B; 8 min, 0% B; followed by equilibration for 4 min with 0% B. Biotin data collection was done in positive ionization mode with a m/z scan range of 100-600; resolution 70,000 at 1 Hz, automatic gain control (AGC) target of 3e6; and a maximum injection time of 250 ms. Injection volumes for biotin detection were 20 µL.

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Supplementary Tables

Table S1A. Electron donors tested with strains CF and DCA

Electron donor Dechlorination a Methanol - Ethanol - Formate +b Acetate - Propionate - Pyruvate - Lactate - Glucose -

H2 + a “+” represents the detection of sustained dechlorination of 1,1,1-TCA or 1,1-DCA after two transfers into chemically defined medium. Transfers were made using a 10% dilution. Before inoculation, cells were pelleted down to remove the supernatant, and then were suspended in sterile mineral medium. b Formate utilization by strains CF and DCA requires the supplementation of either ACT-3 mixed culture supernatant, yeast extract, or casamino acids.

Table S1B. Electron acceptors tested with strains CF and DCA

Strain CF Strain DCA e- acceptor Dechlorination Product Dechlorination Product CF + DCM - DCM - - 1,1,1-TCA + 1,1-DCA - 1,1-DCA - + CA 1,2-DCA (major) 1,2-DCA (major) 1,1,2-TCA + VC (minor) + VC (minor) 1,2-DCA - - CA - - PCE - - TCE - - VC - - 1,1-DCE - -

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Supplementary Figures

Fig. S1. Growth of Lactobacillus delbrueckii (ATCC=7831) measured at OD630 nm in response to various concentrations of cobalamin. Data are means ± SE of five replicates in each experiment.

Fig. S2. Typical cumulative dechlorination profiles of 10-9 dilution-to-extinction transfers of strains (A) CF and (B) DCA. The black arrows represent the times when chlorinated substrates we refed. Before adding a new dose of substrates, the cultures were purged with N2/CO2 (80% v/v) to remove dechlorinated products. Symbols: CA, monochloroethane; 1,1-DCA, 1,1-dichloroethane.

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Fig. S3. PCR amplification of 16S rRNA genes and cfrA/dcrA from strains CF and DCA isolate cultures. Restriction digestion analysis with (A) KpnI and (B) XbaI: (a) ACT-3 mixed culture; (b) strain CF isolate grown in medium with autoclaved ACT-3 mixed culture supernatant; (c) strain CF isolate grown on defined medium with complete vitamin supplement; (d) strain CF isolate grown in defined medium without biotin, cobalamin, and thiamine. (C) predicted restriction sites and lengths of the two 16S rRNA gene sequences (5’-3’) and their digested fragments; (D) examination of potential cross-strain contamination in strain CF and DCA DNA using specific primers of cfrA/dcrA. The size of cfrA and dcrA amplicons is 119 bp and 110 bp, respectively. Note that the two bands below 100 bp are primers.

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Fig. S4. Gram-stain microscopic image of Dehalobacter restrictus strain CF cells.

Fig. S5. Serine salvage from threonine via glycine in Dehalobacter restrictus strain CF. (A) Schematic of serine biosynthesis in strain CF; (B) purified recombinant Dehalobacter serine hydroxymethyltransferase (42 KDa). Abbreviations: GT, glycine transaminase; PSP, phosphoserine phosphatase; SDH, serine dehydratase; SPT, serine-pyruvate transaminase; THF, tetrahydrofolate; 5,10- MTHF, 5,10-methylenetetrahydrofolate.

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Fig. S6. MS detection of potential biotin (m/z=245.09) in the stock solutions used to make the defined medium (without biotin added) for strain CF growth (injection volume: 20 µL). The six chromatograms are extracted ion chromatograms (m/z=245.09 ± 0.002); chemicals are ionized in positive mode. The detection limit in the LC-MS method is 1 µg/L (0.02 ng) with a linear range from 5 to 50 µg/L.

Fig. S7. Average dechlorination rates of strain CF grown on acetate or pyruvate as the main carbon source with 0.1 mM of arginine, histidine, and threonine. Symbols: ♦, 1,1-DCA. Data are means ± SE of three replicates in the representative experiment.

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Fig. S8. MS detection of potential cyanocobinamide (CN-Cbi; m/z=1015.49) and cyanocobalamin (CN-Cba; m/z=1355.57) in the DMB standard (m/z=147.09) used for strain CF growth (injection volume: 25 µL). The chromatogram on top panel shown are extracted ion chromatograms and the chemicals are ionized in positive mode.

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Appendix E. Supplementary Information for Chapter 4

Supplementary table. Detected amino acids in the supernatants of the Dhb strain PER-K23 cultures after the consumption of 5 mM TCE. n.d., not detected; n.a., not applicable.

Amino acids Elemental Molecular Concentration STDEV formula weight (µM) Glutamate C5H9O4N 148.0604 32.34594829 1.178797 Valine C5H11O2N 118.086 17.05574161 0.486991

Lysine C6H14N2O2 147.1128 5.73457658 0.088364 Glycine C2H5NO2 76.039 12.13889052 1.308286 Isoleucine/Leucine C6H13NO2 132.1019 19.93066114 0.561976 Phenylanaline C9H11NO2 166.0863 4.204021777 0.081025 Proline C5H9NO2 116.0706 6.753394466 0.408994 Tryptophan C11H12O2N2 205.097 0.791331081 0.032241

Methionine C5H11NO2S 150.0583 2.210257568 0.045601 Glutamine C5H10O3N2 147.0764 2.020619597 0.156998 Aspartic acid C4H7NO4 134.0448 11.90429073 0.497667 Alanine C3H7O2N 90.055 12.87953435 0.180503 Asparagine C4H8N2O3 133.0608 3.849965632 0.165155 Cysteine C3H7NO2S 122.027 4.714690912 0.157711 Histidine C6H9O2N3 156.0768 n.d. n.d. Threonine C4H9O3N 120.0655 3.37604402 0.124791

Tyrosine C9H11NO3 182.0812 n.d. n.d. Arginine C6H14O2N4 175.119 n.d. n.d. Serine C3H7NO3 106.0499 n.a. n.a.

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Appendix F. Supplementary Information for Chapter 5 Figures

Aromatic carboxylase operon

Phthaloyl-CoA decarboxylase operon

Fig. S1. Phylogenetic analysis of the UbiX family. Related to Fig. 1. Rooted phylogenetic tree of over 9,000 UbiX-like sequences (IPR004507, InterPro database) colored by phylum (except for the Proteobacteria that are colored by class). The UbiX sequences from the aromatic carboxylase operons and the phthaloyl-CoA decarboxylase operons are highlighted by red and blue stars, respectively.

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Fig. S2. SDS-PAGE of purified UbiX-like proteins (FMN prenyltransferases). Related to Fig. 1 and Fig.2. 6His-tagged UbiX-like proteins were affinity purified using nickel-chelate chromatography and protein samples were resolved on 15% SDS-PAGE gels followed by Coomassie staining.

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Fig. S3. Purified PA4019 and cell lysate assays for DMAP biosynthesis in E. coli BL21. Related to Fig. 2 and Fig. 3. (a) PA4019 preparations purified from E. coli cells grown with or without prenol addition. The FMN-bound PA4019 shows yellow color in solution, whereas the purple color indicates the presence of prFMNradical in PA4019. MS-based assays with the wild type E. coli BL21 cell lysates for (b) prenol phosphorylation to DMAP; (c) DMAPP dephosphorylation, and (d) DMAP dephosphorylation.

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Fig. S4. FMN binding and prenylation by purified AF1214 and other UbiX proteins. Related to Fig. 2. FMN binding by purified AF1214 under (a) anaerobic (reduced by DT) and (b) aerobic conditions: effect FMN of phosphate (Pi; 20 mM) and DMAP (0.1 mM). (c, d), AF1214 saturation with DMAP measured by following: (c) the formation of prFMN at 550 nm (after dithionite reduction/re-oxidation) or (d) the decrease in FMN absorbance at 388 nm (without dithionite reduction/re-oxidation). (e) LC-MS analysis of FMN prenylation activity of 12 purified UbiX proteins. FMN and reaction products were analyzed in protein-free filtrates using LC-MS as described in Materials and Methods. Some UbiX proteins (PA4019, JGI0011, NBRC0004) showed strong cofactor binding. (-) control: no enzyme addition.

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Fig. S5. Biosynthesis of DMAP in E. coli: biochemical characterization of ThiM, NudF and NudJ. Related to Fig. 3. (a) Thiazole and prenol phosphorylation by purified ThiM. (b) Prenol and thiazole phosphorylation activities of the purified wild-type and mutant ThiM proteins. (c) Prenol phosphorylation activity of cell lysates from the E. coli BL21 wild-type and ΔthiM strains. Enzyme assays were performed using a PA4019-based protocol and two prenol concentrations (0.05 and 1.0 mM) as described in Methods. DMAPP dephosphorylation activity of purified (d) NudF and (e) NudJ. (f) DMAPP dephosphorylation activity of cell lysates from the E. coli BW25113 wild-type and Nudix hydrolase knockout strains (ΔnudF; ΔnudJ; and ΔnudF,nudJ).

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Fig. S6. LC-MS spectra and proposed structures of prFMN forms. Related to Fig. 4. (a) Direct- injection MS spectrum of prFMN-C4a-radical. (b, c, d), LC-ESI-MS extracted ion chromatograms of the preparations obtained from the oxidation of H-prFMNradical at pH 2 (b), pH 7 (c), and pH 10 (d). (e) Direct- injection MS spectrum of F-preparation; (f) Photodiode array (PDA) UV-vis scan of F-preparation during UPLC-MS analysis (pH 6.0). LC-ESI-MS spectra of C2′-hydroxylated prFMNiminium (panel g) and C1′,C2′- dihydroxylated prFMN (panel h). TIC, total ion current spectrum.

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Fig. S7. Development of prFMN measurement methods. Related to Fig. 4, Fig. 5, and Fig. 6. (a) Confirmation of the purity of FMN standard extracted from AF1214FMN and prFMN standard extracted from AF1214prFMN. LC-MS analysis suggested complete FMN prenylation and negligible abiotic dephosphorylation of FMN and prFMN. (b) Extinction coefficient of protonated prFMNradical. ACN, acetonitrile.

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Table S1. UbiX genes from microbial operons related to anaerobic degradation of aromatic compounds (benzene, naphthalene, phenol and phthalate). Related to Fig. 1. UbiX ID Length Associated UbiDs Organisms (Uniprot) (a.a.) D8F9E7 210 Putative naphthalene carboxylase Deltaproteobacterium sp. strain NaphS2

E1YEX1 221 Unknown, gene from Desulfobacterium sp. strain N47 naphthalene-degrading organism E1Y9S2 199 Unknown, gene from Desulfobacterium sp. strain N47 naphthalene-degrading organism D8WWQ0 192 Putative benzene carboxylase Clostridia culture BF (Peptococcaceae)

NBRC0004* 192 Putative benzene carboxylase Benzene-degrading Peptococcaceae sp.

D3S145 163 Putative benzene carboxylase Ferroglobus placidus D8WWQ5 199 Unknown, gene from benzene- Clostridia culture BF degrading organism (Peptococcaceae) D8WWQ7 199 Unknown, gene from benzene- Clostridia culture BF degrading organism (Peptococcaceae) Q39TU0 201 Phenylphosphate carboxylase Geobacter metallireducens B9M8H6 201 Phenylphosphate carboxylase Geobacter daltonii P57767 194 Phenylphosphate carboxylase Thauera aromatica Q5P483 196 Phenylphosphate carboxylase Azoarcus sp. strain EbN1 (a.k.a. Aromatoleum aromaticum)

H0Q4V7 163 Phenylphosphate carboxylase Azoarcus sp. strain KH32C A0A0K1J7P6 200 Phenylphosphate carboxylase Azoarcus sp. strain CIB A0A1N7ASS1 200 Phenylphosphate carboxylase Azoarcus tolulyticus A0A0M0FVG1 194 Phenylphosphate carboxylase Azoarcus sp. strain PA1 Q5NWH5 204 Phthaloyl-CoA decarboxylase Aromatoleum aromaticum Q5NWG7 204 Phthaloyl-CoA decarboxylase Aromatoleum aromaticum A0A0M0FYJ7 199 Phthaloyl-CoA decarboxylase Azoarcus sp. strain PA1 A0A1H5Z7X7 200 Phthaloyl-CoA decarboxylase Thauera chlorobenzoica *NBRC0004 is amplified from a nitrate-reducing benzene-degrading mixed culture (Luo et al. 2014), and the nucleotide sequence is available at IMG with the accession number Ga0197853_11155252.

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Table S2. UbiX proteins purified in this work and analyzed for the presence of prFMN. Related to Fig. 1 and Fig.2.

Protein color FMN Protein UniProt Purified Gene name after in vitro prenylation activity name ID protein color reconstitution (LC-MS)* 1. AF1214 (AF_1214) O29054 purple N/A + 2. APE1647 (APE_1647) Q9YBF0 colorless colorless + 3. BH1651 (BH1651) Q9KCC2 colorless colorless + 4. BSU0364 (BSU0364) P94404 colorless colorless + 5. EC4018 (UbiX) P0AG03 <50% purity N/A N/A 6. HP1451 (HP_1476) O26011 colorless purple + 7. JGI0011 (Amet_4582) A6TWT5 colorless purple + 8. JGI0049 (Clos_0364) A8MLK9 insoluble N/A N/A 9. JGI0070 (CHY_0680) Q3AE99 colorless colorless + 10. MJ0101 (MJ0102) Q57566 purple N/A + 11. MTH0147 (MTH_147) O26250 purple N/A + 12. NBRC0004 N/A N/A colorless colorless + 13. NE0058 (NE0058) Q82Y31 insoluble N/A N/A 14. PA4019 (PA4019) Q9HX08 purple N/A + 15. RP0927 (RPA0930) Q6NB98 insoluble N/A N/A 16. SAV4814 (Saverm_4811) Q82E05 colorless colorless + 17. SC4292 (SCO4492) Q9KYP1 insoluble N/A N/A 18. SM4097 (SMa1285) Q92Z12 <50% purity N/A N/A 19. SM4596 (SMa2219) Q92XP7 colorless colorless + 20. SM4834 (SM_b20135) Q92X27 purple N/A + 21. SSO0437 (SSO0437) Q9Y8K8 <50% purity N/A N/A 22. TA1100 (TA1100) Q9HJ72 <50% purity N/A N/A 23. YST0799 (PAD1) P33751 <50% purity N/A N/A *FMN prenylation activity of purified UbiX proteins was determined by LC-MS analysis of protein-free filtrates and is defined as the observation of (i) prFMNiminium or prFMN-OH or (ii) purple color in the concentrated UbiX proteins. N/A, not applicable.

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Table S3a. Complete list of E. coli proteins screened for prenol phosphorylation activity in this work. Related to Fig. 3.

UniProt UniProt Gene Gene Gene UniProt ID ID ID 1. agaB P42909 37. gatB P37188 73. pykF P0AD61 2. agaV P42904 38. glk P0A6V8 74. rbsK P0A9J6 3. alsK P32718 39. glpK P0A6F3 75. rfaP P25741 4. anmK P77570 40. glvB P69789 76. rfaY P27240 5. apbE P0AB85 41. glxK P77364 77. rhaB P32171 6. araB P08204 42. gntK P46859 78. ribF P0AG40 7. aroK P0A6D7 43. gsk P0AEW6 79. sgcA P39363 8. aroL P0A6E1 44. hldE P76658 80. sgcB P58035 9. ascF P24241 45. idnK P39208 81. srlB P05706 10. bglF P08722 46. ispE P62615 82. srlE P56580 11. chbA P69791 47. kdgK P37647 83. tdk P23331 12. chbB P69795 48. kptA P39380 84. thiD P76422 13. cmtA P69826 49. lpxK P27300 85. thiK P75948 14. cmtB P69824 50. lsrK P77432 86. thiM P76423 15. coaA P0A6I3 51. lyx P37677 87. thrB P00547 16. coaE P0A6I9 52. mak P23917 88. treB P36672 17. cobU P0AE76 53. malX P19642 89. udk P0A8F4 18. crr P69783 54. manX P69797 90. ulaB P69822 19. cysC P0A6J1 55. mngA P54745 91. ulaC P69820 20. dgkA P0ABN1 56. mtlA P00550 92. xylB P09099 21. dgoK P31459 57. murP P77272 93. yadI P36881 22. frlD P45543 58. nadK P0A7B3 94. ydjH P77493 23. fruA P20966 59. nadR P27278 95. yegS P76407 24. fruB P69811 60. nagE P09323 96. yegV P76419 25. fruK P0AEW9 61. nagK P75959 97. yeiI P33020 26. frvA P32155 62. nanK P45425 98. ygcE P55138 27. frvB P32154 63. pdxK P40191 99. yihV P32143 28. frvR P32152 64. pdxY P77150 29. frwB P69816 65. pfkA P0A796 30. frwD P32676 66. pfkB P06999 31. fryA P77439 67. prkB P0AEX5 32. fryB P69808 68. psuK P30235 33. fucK P11553 69. ptsA P32670 34. galK P0A6T3 70. ptsG P69786 35. garK P23524 71. ptsN P69829 36. gatA P69828 72. pykA P21599

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Table S3b. Complete list of E. coli phosphohydrolases screened for DMAPP dephosphorylation activity in this work. Related to Fig. 3.

Gene UniProt ID Gene UniProt ID Gene UniProt ID 1. mutT P08337 7. nudI P32664 13. mutY P65556 2. nudJ P0A776 8. yfcD P37128 14. nudF P77788 3. nudG P0AEI6 9. nudK P43337 15. nudE Q46822

4. nudL P0AFC0 10. nudK P37128 16. nudC Q93K97 5. nudB P17802 11. rppH P45799 17. rdgB P52061 6. gmm P32056 12. idi P52006

Table S5. Mass spectrometry analysis of targeted protein expression in E. coli BL21 wild- type cells grown with and without prenol addition. Related to Fig. 3.

emPAI

Prenol Uniprot Gene Prenol Annotation not ID name added added

Dihydroneopterin triphosphate nudB 0 0 P17802 diphosphatase P0AEI6 nudF ADP-ribose pyrophosphatase 1.5-3 0 P32664 nudI Nucleoside triphosphatase 0 0 P76423 nudJ Phosphatase NudJ 9 0-6 Q93K97 thiM Hydroxyethylthiazole kinase 1.6-8.7 0-6 P0A805* frr Ribosome-recycling factor 11-31 12-15

atpH ATP synthase subunit 0.5-1.4 1-2 P0ABA4*

*The 2 proteins are translated products of housekeeping genes used as a reference for protein expression

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Table S6. Names and structures of prFMN forms proposed in this study. Related to Fig. 4, Fig. 5, and Fig. 6. Label in Short name Complete name Structure Main Text*

a A prFMNred Reduced prFMN

a B prFMNradical prFMN-C4a-radical

Protonated prFMN b C prFMNradical-H -N5-radical

a D prFMNiminium prFMN-N5-iminium

c E prFMN-OH C1'-hydroxylated prFMN

C1'-ene- C1'-ene-prFMN d F prFMNiminium -C4a-iminium cation

C2'-hydroxylated d G prFMNiminium-OH prFMN-N5-iminium

C1',C2'-dihydroxylated b H prFMN-(OH)2 prFMN

*The prFMN structure proposed by aWhite et al., 2015, bthis study, 2015, cPayne et al., and dMarshall et al., 2017, respectively.

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Table S7. Decarboxylation activities of cinnamic acid and α-fluoro-cinnamic acid for Fdc1 activated by different preparations. Related to Fig. 6.

F-preparation D-preparation Fdc1 activated by (green-colored) (yellow-colored) Cinnamic acid (CA) 1.25 ± 0.01 U/mg 0.25 ± 0.01 U/mg α-fluoro-cinnamic acid (FCA) 0.30 ± 0.01 U/mg 0.12±0.001 U/mg Ratio CA/FCA 4 2

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