Formate dehydrogenases and hydrogenases in syntrophic propionate-oxidizing communities

Gene analysis and transcriptional profiling

Petra Worm Thesis committee

Thesis supervisor Prof. dr. ir. A.J.M. Stams Personal chair at the Laboratory of Microbiology Wageningen University

Thesis co-supervisor Dr. C.M. Plugge Assistant Professor at the Laboratory of Microbiology

Other members Prof. dr. S. C. de Vries, Wageningen University Prof. dr. B. Schink, University of Konstanz, Germany Prof. dr. M. J. Teixeira de Mattos, University of Amsterdam Dr. H. J. M. op den Camp, Radboud University Nijmegen

This research was conduted under the auspices of the Netherlands Research School for the Socio-Economic and Natural Sciences of the Environment (SENSE) Formate dehydrogenases and hydrogenases in syntrophic propionate-oxidizing communities

Gene analysis and transcriptional profiling

Petra Worm

Thesis submitted in fulfilment of the requirements for the degree of doctor at Wageningen University by the authority of the Rector Magnificus Prof. dr. M.J. Kropff in the presence of the Thesis Committee appointed by the Academic Board to be defended in public on Monday 28th of June 2010 at 4 p.m. in the Aula Petra Worm Formate dehydrogenases and hydrogenases in syntrophic propionate- oxidizing communities: gene analysis and transcriptional profiling

PhD Thesis, Wageningen University, Wageningen, the Netherlands (2010) 138 pages. With references, and with summaries in Dutch and English

ISBN 978-90-8585-675-7 TABLE OF CONTENTS

Chapter 1 1 General introduction

Chapter 2 21 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

Chapter 3 37 Growth and substrate dependent transcription of formate dehydrogenase and hydrogenase coding genes in fumaroxidans and Methanospirillum hungatei

Chapter 4 53 Formate dehydrogenase and hydrogenase encoding gene clusters in Syntrophobacter fumaroxidans and Methanospirillum hungatei

Chapter 5 67 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

Chapter 6 83 Decreased activity of a propionate-degrading community in a UASB reactor fed with synthetic medium without molybdenum, and selenium

Chapter 7 95 Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in anaerobic granular sludge deprived of molybdenum, tungsten and selenium

Chapter 8 107 General discussion

Summary 122 Samenvatting 124 SENSE certificate 126 Curriculum vitae 128 Publications 129 Dankwoord 130

CHAPTER 1

GENERAL INTRODUCTION

This chapter was adapted from:

Petra Worm, Nicolai Müller, Caroline M. Plugge, Alfons J. M. Stams, and Bernhard Schink. Syntrophy in methanogenic degradation. (Endo)symbiotic Methanogenic Archaea. Edited by Johannes Hackstein (in press).

1 General introduction

1. Anaerobic degradation of organic compounds In -limited environments such as lake sediments or the lower layers of eutrophic lakes, biomass oxidation is coupled to reduction of electron acceptors such as nitrate,

Mn(IV), Fe(III), sulfate, or CO2 (Zehnder 1978; Schink 1989). The relative importance of these electron acceptors depends on their availability in the respective habitat; most freshwater sediments are rich in iron oxides, and marine sediments are rich in sulfate. Only methanogenesis is independent of external electron acceptors because the methanogenic degradation of biomass is actually a dismutation of organic carbon:

0 C6H12O6 → 3 CH4 + 3 CO2 ∆G ’ = -390 kJ per mol

Whereas aerobic, nitrate-reducing or manganese-reducing typically are able

to degrade polymeric organic compounds via the respective monomers to CO2 in one single cell, the conversion of complex organic molecules by iron reducers or sulfate reducers requires cooperation with fermenting bacteria which feed the respective terminal oxidizers with classical products such as fatty acids, alcohols and others. Methanogenic degradation of organic matter is even more complex and requires cooperation of three different metabolic groups (guilds) of microorganisms, including primary fermenters, secondary fermenters and methanogens (Bryant 1979, Fig. 1). Primary fermenting bacteria are well and have been isolated with all kinds of polymeric or monomeric substrates. Also anaerobic protozoa including flagellates and ciliates can ferment complex organic molecules. Different from iron reducers or sulfate

reducers, methanogenic Archaea use only very few substrates, including hydrogen, CO2,

polymers

1 monomers I 1 fatty acids alcohols succinate, lactate 2 hydrogen II acetate C -compounds 1 5 3 4 III

CH4, CO2

Figure 1. Methanogenic degradation of complex organic matter by cooperation of different metabolic groups. Metabolic groups of organisms involved: primary fermenters (1), secondary fermenters

(2), hydrogen and C1-compounds-using methanogens (3), acetoclastic methanogens (4) and homoacetogenic bacteria (5) (modified after Schink 1997).

2 CHAPTER 1

other C1-compounds, and acetate. Some methanogens can also oxidize isopropanol and ethanol (Widdel et al. 1988). Thus, the fermentation products, such as alcohols, fatty acids, branched-chain fatty acids and aromatic fatty acid residues from partial degradation of amino acids, long -chain fatty acids from lipid hydrolysis, heterocyclic aromatic compounds derived from nucleic acids need to be fermented further to the substrates that methanogens can use (Bryant 1979; Schink 1997; Schink and Stams 2006; McInerney et al. 2008; Stams and Plugge 2009). This is the function of the secondary fermenting bacteria which depend on close cooperation with methanogenic partners.

2. Methanogenesis Methanogenic environments are widely distributed in nature. Wetlands, freshwater sediments, swamps, and digestive tracts of ruminants and insects are environments that produce high amounts of methane. Man-made systems, such as rice paddies and anaerobic bioreactors and landfills are other important sources of methane production. Methanogenic Archaea catalyze the final step in the overall anaerobic degradation of organic material to methane and CO2. One metabolic group of methanogenic Archaea converts CO2 plus hydrogen or formate to methane, while another group uses acetate or . Acetate, the most important intermediate in anaerobic digestion, accounts for approximately two -thirds of all methane produced, while the last third is produced from the reduction of CO2 with electrons derived from the oxidation of hydrogen or formate (Ferry 1992; Liu and Whitman 2008). Currently, only two types of acetoclastic methanogens have been identified: Methanosaeta sp. and Methanosarcina sp.. Methanosarcina is a genus of versatile methanogens, including species capable of growing with different substrates such as acetate, methanol, methylamines and H2/CO2, whereas Methanosaeta sp. use only acetate. Methanosaeta sp. are widely distributed in nature and, because of their high affinity for acetate, they outcompeteMethanosarcina sp. in low-acetate environments (Conklin et al. 2006). Both acetoclastic archaea grow slowly, with doubling times of 1-12 (Methanosaeta) and 0.5-2 (Methanosarcina) days (Jetten et al. 1992). Methanogenic archaea are phylogenetically very diverse. They are classified in five orders (Whitman et al. 2006).

3. Syntrophism The ability to transfer electrons to a partner organism is an important metabolic feature associated with many physiologically diverse microorganisms. This trait is usually referred to as syntrophism. Syntrophism is a special type of symbiosis between two microorganisms in which growth of one organism depends on supply of growth factors or nutrients, or removal of products by a partner organism. Especially among anaerobic microorganisms, cooperation of several metabolic types of bacteria in the feeding chain is a common feature. The mutual dependence can be explained by calculating the changes in Gibbs’ free energy (∆G0’) for the oxidation of e.g. ethanol to hydrogen, 5 CO2 and acetate (Bryant et al. 1967). Under standard conditions with gases at 10 Pa pressure, 1 M concentration of products/substrates, at pH 7.0 and 298 K the Gibbs’ free energy value for ethanol oxidation is positive with +9.6 kJ/reaction (Table 1). This

3 General introduction

indicates that the reaction cannot take place, nor can any microbe gain energy from this oxidation. However, the Gibbs’ free energy becomes negative when the hydrogen

partial pressure (pH2) decreases. This example of interspecies hydrogen transfer is characteristic for the way how organic matter is degraded in methanogenic habitats. Life depends on the availability of energy which is stored inside the cell in the form of ATP. Under physiological conditions, including heat losses, the synthesis of ATP requires 60 – 70 kJ per mol (Thauer et al. 1977). Membrane-bound ATPases couple the hydrolysis or synthesis of ATP to the transport of protons (in some cases also Na+ ions) across the cytoplasmic membrane. Depending on the stoichiometry of the ATPase system in question, the ratio of ions translocated versus ATP synthesized or hydrolyzed may vary

Table 1. Equations and standard free energy changes for relevant reactions described in the chapter (Gibbs’ free energy changes are taken from Thauer et al. 1977)

Table 1a. Equations and free energy changes for secondary fermentation reactions Reaction ΔG0’ kJ/reaction

- + Ethanol + H2O → Acetate + H + 2H2 + 9.6

Propionate- + 2 H2O → Acetate- + CO2 + 3 H2 + 76 + Butyrate- + 2 H2O → 2 Acetate- + H + 2H2 + 48 - + Crotonate- + 2 H2O → 2 Acetate + H + H2 - 6.2

Acetate- + 2 H2O → 2 CO2 + 4 H2 + 96

Table 1b. Equations and free energy changes for reactions of methanogenic archaea. Reaction ΔG0’ kJ/reaction

4 H2 + CO2 → CH4 + 2 H2O - 131 - + 4 Formate + 4H → CH4+ 3 CO2 + 2H2O - 145

4 CO + 2 H2O → CH4 + 3 CO2 - 211 + Acetate- + H → CH4 + CO2 - 35

4 Methanol → 3 CH4 + CO2 + 2 H2O - 106

H2 + Methanol → CH4 + H2O - 113 + - CO2 + H2O → H + HCO3 + 4.8 - + CO2 + H2 → Formate + H - 4.5

Table 1c. Equations and free energy changes for hydrogen-consuming reactions Reaction ΔG0’ kJ/reaction

- Crotonate + H2 → Butyrate - 75 - + CO2 + H2 → Formate + H - 4.5

4 CHAPTER 1 between 3 and 5; in most cases, a ratio of 3 to 4 appears to be justified (Engelbrecht and Junge 1997; Cherepanov et al. 1999). As a consequence, the smallest amount of energy that can still be converted to ATP (and with this into metabolic activity and growth) is equivalent to one third or one fourth of an ATP unit, i.e., in the range of -15 to -20 kJ per mol reaction (Schink 1997; Schink and Stams 2006). However, Nakanishi-Matsui and Futai (2008) recently reported that ATPase can translocated between 3.3 and 4.6 protons per ATP, and with this, the smallest quantum of biologically conservable energy may range from -13 to -18 kJ per mol reaction.

4. Anaerobic oxidation of ethanol The biochemistry of syntrophic oxidation of ethanol, although the oldest syntrophic system known, has still not been elucidated in detail. Early work on the so-called S-organism indicated that ethanol is oxidized via acetaldehyde to acetyl-CoA and further to acetate, including ATP synthesis by acetate kinase (Reddy et al. 1972 a, b, c). This concept was confirmed by similar studies on the ethanol-oxidizing bacteria Pelobacter acetylenicus and P. carbinolicus (Schink 1985; Eichler and Schink 1986). Nonetheless, the energetics of this reaction chain is still unclear. The overall reaction

2 ethanol + CO2 →2 acetate + CH4 yields ∆G0’ = -112 kJ per mol, which leaves a total of about -40 kJ per ethanol oxidation reaction for the syntrophic ethanol oxidizer, indicating that part of the ATP formed by substrate-level phosphorylation has to be invested into reversed electron transport. Oxidation of acetaldehyde to acetyl-CoA (E0’ = -370 mV) can be coupled to hydrogen formation via ferredoxin at a sufficiently low hydrogen pressure. A ferredoxin-like electron carrier has been purified from P. acetylenicus (Kowalski and Schink, unpublished). The energetically difficult reaction is the transfer of electrons from the acetaldehyde/ethanol couple (-196 mV) to hydrogen formation. Such a reaction requires energy input in the form of, e. g., a reversed electron transport, a feature that is common to all syntrophically fermenting bacteria studied so far (Fig. 2). Hydrogen formation from ethanol in crude extracts of P. acetylenicus is stimulated by ATP (Hauschild 1997). Since P. acetylenicus also contains a menaquinone-like electron carrier (Strohm and Schink, unpublished) a basically similar reversed electron transport system as suggested for Syntrophomonas wolfei (discussed in section 1.5) can be anticipated, but experimental evidence has not been provided yet. A comproportionating [FeFe]-hydrogenase as described for Thermotoga maritima (Schut and Adams 2009) could finally release the electrons from NADH and ferredoxin towards proton reduction.T. maritimaferments glucose to acetate,

CO2 and H2 via the Embden-Meyerhof pathway, generating two NADH and four reduced ferredoxins per molecule of glucose. In order to re-oxidize these carriers, a proposed bifurcating [FeFe]-hydrogenase uses electrons from NADH and reduced ferredoxin in a

1:2 ratio to produce H2. Schut and Adams (2009) found genes with sequence similarity to this [FeFe]-hydrogenase in several other microorganisms, including the ethanol- degrading P. carbinolicus, the butyrate-degrading S. wolfei (discussed in section 1.5) and the propionate-degrading Syntrophobacter fumaroxidans (discussed in section 1.6).

5 General introduction

+ substrate H , CO2

reducing equivalents

AT P

⅓-⅔ ATP

+ H acetate H2, formate CO2

reducing reducing equivalents equivalents AT P AT P

+ H2 CH4, CO2 H , CO2 CH4

Figure 2 Conversions performed by a secondary fermenting bacterium (top), a hydrogen- and formate- using methanogen (bottom right), and an acetoclastic methanogen (bottom left).

5. Anaerobic oxidation of butyrate Anaerobic butyrate degraders known to date belong to only two groups of bacteria, the genus Syntrophomonas within the phylum Firmicutes and the genus within the order Syntrophobacterales of the phylum . Fermentation of butyrate to acetate and hydrogen is endergonic (Table 1) and occurs only at very low hydrogen partial pressures, e.g. in the presence of methanogenic archaea (Schink 1997). Syntrophic butyrate oxidizers use only very few substrates. Beyond oxidation of saturated fatty acids in coculture with methanogens, axenic growth is possible only with unsaturated fatty acids such as crotonate (Schink 1997; McInerney et al. 2008). They cannot use external electron acceptors for growth, thus reflecting the high degree of specialization of these bacteria for syntrophic cooperation (Schink 1997). Butyrate is oxidized via ß-oxidation to acetate yielding one mole ATP per mole of butyrate. The reducing equivalents are transferred to flavoenzymes and NAD+ (Wofford et al. 1986). Reoxidation of these electron carriers of a relatively positive redox potential with protons to form hydrogen is energetically difficult. Of course, a low hydrogen partial pressure helps to facilitate those reactions, but no known methanogen is able to maintain a hydrogen partial pressure low enough (10-10 atm) to allow direct proton reduction with these electrons (Thauer and Morris 1984; Schink 1997). Therefore, it was postulated that syntrophic butyrate degraders have to invest energy into a reversed electron transport, thus leaving only a fraction of an ATP for growth of the bacterium (Thauer and Morris 1984). Recently, Müller et al. (2009) showed that an system similar to the comproportionating [FeFe]-hydrogenase of Thermotoga maritima is essential in butyrate oxidation by Syntrophomonas wolfei. The comproportionating [FeFe]-hydrogenase of T. maritima drives the endergonic reduction of protons to

6 CHAPTER 1 hydrogen with NADH by exergonic reduction of another couple of protons with reduced ferredoxin which is produced in pyruvate oxidation during growth on glucose (Schut and Adams 2009). In butyrate oxidation by S. wolfei, no such ferredoxin-reducing reaction is involved. Nonetheless, hydrogen formation from NADH is likely catalyzed by a [FeFe]- hydrogenase homologue in S. wolfei. This reaction is possible already at a hydrogen partial pressure of 10-3 atm (Schink 1997; Müller et al. 2009). Since the enzyme found in S. wolfei is associated with a formate dehydrogenase-like protein analogous to its homologue in Eubacterium acidaminophilum, interspecies electron transfer may occurs via either hydrogen and/or formate, depending on the environmental conditions (Graentzdoerffer et al. 2003; Müller et al. 2009). The thermodynamically most difficult step in butyrate oxidation is the transfer of electons derived from butyryl-CoA oxidation to NAD+ for which a redox potential difference of at least +200 mV has to be overcome (Schink 1997). It was hypothesized that electrons from butyrate oxidation are transferred to quinones in the membrane, and that the reduced quinones are reoxidized with NAD+ (Wallrabenstein and Schink 1994). Such a reaction would require energetization by e.g. a proton gradient, which was found to be essential for hydrogen formation from butyrate by S. wolfei (Wallrabenstein and Schink 1994). The S. wolfei [FeFe]-hydrogenase catalyzes the reduction of quinones with NADH, indicating that, besides forming hydrogen from NADH oxidation, this enzyme also catalyzes the proton gradient-driven endergonic oxidation of quinones with NAD+, (Müller et al. 2009). However, a direct linkage between quinol oxidation and proton translocation has not been demonstrated so far. Another possible mechanism for reversed electron transport during butyrate oxidation was postulated forSyntrophus aciditrophicus based on genome data. Here, an Rnf-complex could oxidize NADH and transfer electrons to ferredoxin, driven by influx of protons or sodium ions into the cell (McInerney et al. 2008). Electrons that arise during butyryl-CoA oxidation could be transferred to components of the membrane where NAD+ is reduced in a similar manner as postulated for S. wolfei (McInerney et al. 2008). With the Rnf-complex, S. aciditrophicus has the potential prerequisites for producing reduced ferredoxin during butyrate degradation, which may drive comproportionating reactions such as NADH oxidation by [FeFe]-hydrogenases, or bifurcating reactions such as butyryl-CoA oxidation by the Bcd/EtfAB complex (Li et al. 2008; Herrmann et al. 2008). In contrast, genes that encode for the Rnf-complex are not present in the genome of S. wolfei, indicating that the pathway of butyrate degradation is different in both organisms and does not include reduced ferredoxin in S. wolfei (Müller et al. 2009).

6. Anaerobic oxidation of propionate All currently identified syntrophic propionate-oxidizing bacteria are affiliated with the class of within the phylum of Proteobacteria (McInerney et al. 2005), or the low G+C Gram-positive bacteria in the class Clostridia within the phylum Firmicutes (Imachi et al. 2002; Plugge et al. 2002; de Bok et al. 2005) (Table 2). Some of the Syntrophobacter sp. are able to use sulfate as the electron acceptor for propionate oxidation (McInerney et al. 2005), and can grow in pure culture by propionate oxidation with sulfate. In addition, they can grow by fermentation of pyruvate or fumarate.

7 General introduction

Smithella propionica is phylogenetically related to the genus Syntrophus (Liu et al. 1999) and lacks the ability to reduce sulfate. S. propionica does not oxidize propionate but ferments it to butyrate plus two acetate, and grows in pure culture on crotonate (de Bok et al. 2001; Liu et al. 1999). These substrates or substrate combinations have been used to obtain axenic cultures of the syntrophs since they bypass the energetically unfavorable steps in propionate oxidation. Pelotomaculum schinkii, however, could not be obtained in pure culture until today, nor could it grow on any other compound but propionate. As such, this organism remains a true obligately syntrophic bacterium (de Bok et al. 2005). All species described to date were isolated from anoxic reactors, indicating the importance of these organisms in those types of systems. The question whether hydrogen or formate is transferred in syntrophic cocultures has been studied in detail in propionate-degrading Syntrophobacter fumaroxidans cocultures (Fig. 3).

Figure 3 Scanning electron micrograph of a syntrophic propionate-degrading coculture of S. fumaroxidans (lemon- or oval-shaped) and M. hungatei (rod- shaped). Micrograph by Jacqueline Donkers, Wageningen EM Center.

Thermodynamic calculations, flux measurements in defined cocultures and enzyme measurements all confirmed that interspecies formate transfer is an essential mechanism in syntrophic propionate degradation in suspended cultures (Dong et al. 1994a; Dong and Stams 1995). The terminal reductases were studied in detail and biochemical evidence for formate transfer was found (De Bok et al. 2002b). Two formate dehydrogenases were isolated and characterized. Both contain tungsten

and one has an unusually high specific activity both in the formate oxidation and CO2 reduction assay (Reda et al. 2008). When syntrophic cocultures of S. fumaroxidans and M. hungatei were grown with limiting amounts of tungsten, the propionate degradation activity decreased. This decrease coincided adequately with decreased formate dehydrogenase activity while the hydrogenase activities remained almost unchanged (Plugge et al. 2009a). In their natural habitats, syntrophically propionate degrading bacteria form mixed microcolonies with methanogens in which interspecies distances are much shorter. Under these conditions, interspecies hydrogen transfer may becomes more important than interspecies formate transfer (Stams and Dong, 1995). In syntrophic propionate-degrading microcolonies, Syntrophobacter-like bacteria are often surrounded by Methanobrevibacter sp., methanogens which can use only hydrogen but not formate (Grotenhuis et al. 1991). Also in thermophilic sludge, interspecies hydrogen transfer appears to be the preferred path of electron transfer (Schmidt and Ahring 1993). In addition, slow propionate degradation was observed, also in very concentrated cell suspensions of S. fumaroxidans and the hydrogen-oxidizing Methanobrevibacter arboriphilus (Dong et al. 1994a). The organisms involved in propionate degradation are genuine specialists in obtaining metabolic energy for growth, since they have to grow

8 CHAPTER 1

References [1; [1] [2] [3] [7] [8] [9] [6] 4-6] [10]

Syntrophic partner used Mh Mh Mh Mh Mh Mf Mh Mh Mt Mt ND - ND + ND ND - - Ethanol - - ND + + ND ND - - Propanol - + + - ND ND + - - Lactate - ND ND ND - - + - + Malate - + + + + + + + + Propionate + - + - ND + - - ND Lactate + Sulfate - - ND - + ND ND ND ND Crotonate - ND ND ND ND - ND ND - Acetate - + + - + + - - + Pyruvate - + - ND ND + - - - Malate - Substrate utilization in co-culture with a syntrophic syntrophic with a co-culture in utilization Substrate partner + + - - + - - - Fumarate +

Propionate + fumarate - ND + + - - - ND ------ND Propionate + nitrate - in pure in pure culture Substrate Substrate utilization utilization + + + + - - - ND Propionate + sulfate +

Growth temperature (range/ Methanothermobacter thermoautotrophicus Methanothermobacter optimum) (ºC) ND 23-40/35 25-45/37 20-40/37 23-40/33 20-37/37 20-48/37 45-65/55 45-62/55 , Mt:

Growth pH (range/optimum) ND 6.0-8.0/7 6.3-7.8/7 5.5-7.7/6.9 6.5-8.0/7.0 6-8/7.0-7.5 6.5-7.5/6.5-7.2 6.2-8.0/7.0-7.3 6.2-8.8/7.0-7.6

DNA G+C content (mol%) ND ND ND ND 60.6 57.3 58.6 52.8 53.7 - - - - - + + + + Spore formation formicicum Metanobacterium - - - - - + + +

Motility ND - - - - - + + -a Gram reaction -a

Cell length (µm) 3-10 3-11 2.2-3.0 1.8-2.2 1.0-4.5 1.7-2.8 2.0-4.0 1.8-2.5 2.0-2.5 1 Cell width (µm) 1 1.0 0.5 1.0-1.2 1.0-1.3 0.6-1.0 0.7-0.8 1.1-1.6 , Mf: hungatei Methanospirillum : Selected characteristics of propionate oxidizing acetogenic bacteria that grow in syntrophy with methanogens. syntrophy in grow that bacteria oxidizing acetogenic of propionate characteristics : Selected

Syntrophobacter Syntrophobacter pfennigii Syntrophobacter Syntrophobacter sulfatireducens Syntrophobacter Pelotomaculum Pelotomaculum thermopropionicum Pelotomaculum Pelotomaculum propionicicum Smithella propionica Desulfotomaculum Desulfotomaculum thermobenzoicum subsp. thermosyntrophicum Syntrophobacter Syntrophobacter fumaroxidans Pelotomaculum schinkii Pelotomaculum Cells stain Gram-negative but the organism has a Gram-positive cell wall ultrastructure. wall ultrastructure. cell has a Gram-positive but the organism Gram-negative Cells stain Table 2 Table Substrate utilization: +, utilized, -, not utilized. ND: not determined or not reported. reported. determined or not ND: not -, not utilized. +, utilized, utilization: Substrate partner: Mh: Syntrophic [1] Harmsen et al. 1998; [2] Wallrabenstein et al. 1995; [3] Chen et al. 2005; [4] Boone and Bryant 1980; [5] Wallrabenstein et al. 1994; et [6] al Liu 1999; et [7] de al. Bok 2005; et 1980; [5] Wallrabenstein al. 1995; et [3] al. Chen 2005; et [4] Boone and Bryant al. [1] Harmsen 1998; et [2] Wallrabenstein al. 2002. et al. 2007; [10] Plugge al. 2002; [9] Imachi et [8] Imachi et a

9 General introduction

under thermodynamically very unfavorable conditions. The standard Gibbs’ free energy

change of the complete degradation of propionate to methane and CO2 is about -60 kJ which is approximately equivalent to the amount of energy needed to produce one mol of ATP. A community of three microorganisms brings about this conversion: one bacterium

that degrades propionate to acetate, CO2, and hydrogen, and two methanogenic archaea, one that cleaves the acetate, and another one that uses hydrogen to reduce

CO2 to methane. The actual energy that is available for each member of the community depends on the in-situ concentrations of substrate, intermediates and products and will vary during growth. Moreover, it depends also on the enzyme repertoire the microbes have. Our model organism S. fumaroxidans degrades propionate via the methylmalonyl- CoA pathway (Fig. 4). One ATP is harvested in the conversion of pyruvate to acetate via substrate level phosphorylation. Reducing equivalents are released at three different redox potentials: Reduced ferredoxin is formed in the conversion of pyruvate to

acetate, while NADH and FADH2 are formed in the oxidation of malate and succinate, respectively. These intracellular redox mediators need to be re-oxidized by reduction 0 of protons or CO2. The oxidation of reduced ferredoxin (E ’ Fd(ox)/Fd(red) = -398 mV) and NADH (E0’ NAD+/NADH = - 320 mV) can be coupled to reduction of protons (E0’ = -414

mV) or CO2 (-432 mV) only if the hydrogen or formate concentration is kept low by methanogens. The disposal of reducing equivalents generated in pyruvate oxidation to acetyl-CoA is rather straightforward because most strictly anaerobic bacteria contain pyruvate: ferredoxin (Chabrière et al. 1999). Here, electrons travel via ferredoxin to hydrogenases or formate dehydrogenases to produce hydrogen or formate, respectively. Hydrogen and formate are scavenged by the methanogens, thus enabling an efficient re-oxidation of the ferredoxin. The oxidations of succinate and malate with protons are endergonic even at a hydrogen partial pressure as low as 1 Pa (the minimum level that can be achieved by methanogens). To drive these reactions, input of metabolic energy via reverse electron transport is required. The mechanism that drives succinate oxidation to fumarate (E°’ = +33 mV) during syntrophic growth may be similar to the mechanism of energy conservation in fumarate respiration by Wolinella succinogenes (Kröger et al. 2002), but operating in reverse. Experimental evidence was obtained that ⅔ ATP is needed to drive this conversion (van Kuijk et al. 1998). As such, the net ATP gain for the bacterium is 1/3 mol ATP per mol of propionate converted. However, until present the molecular mechanisms involved inS. fumaroxidans and other syntrophic propionate oxidizers using the methylmalonyl-CoA pathway are not fully understood. Also the oxidation of malate to oxaloacetate with NAD+ is an endergonic reaction. Nonetheless, the purified of S. fumaroxidans exhibits a very high Km value towards oxaloacetate and NADH and as such the organism may be able to efficiently perform this conversion (van Kuijk and Stams 1996). Still, the exact mechanism of NADH re-oxidation remains unclear.S. fumaroxidans and P. thermopropionicum contain [FeFe]-hydrogenases that are homologues to the comproportionating [FeFe]-hydrogenase of T. maritima. This suggests that NADH and ferredoxin that are generated in the methylmalonyl-CoA pathway are simultaneously re-oxidized with the reduction of protons. These novel bifurcating enzyme complexes may be essential in these syntrophic . Additionally, the Rnf-complex in

10 CHAPTER 1

propionate

CoASH

propionyl-CoA

CO2 methylmalonyl-CoA

CO succinyl-CoA 2 2[H]

ATP CoASH CHOOH succinate DG0`=82 2[H] DG`=53 CHO-THF fumarate ATP

malate CH≡THF DG0`=48 2[H] 2[H] DG`=19 oxaloacetate CH2=THF 2[H]

pyruvate CH3-THF CoASH DG0`=41 CO2 DG`=-12 2[H] CO 2[H] 2 acetyl-CoA

acetate

Figure 4 Methyl malonyl-CoA pathway for propionate degradation (thick arrows). For the -1steps that generate reducing equivalents ([H]), the ∆G°’ and ∆G’ (at 1 Pa hydrogen) are indicated (kJ mol ) (Stams and Plugge, 2009). The pathway of fumarate degradation by S. fumaroxidans is shown in thin arrows (Plugge et al 1993). Tetrahydrofolate is indicated as THF.

11 General introduction

S. fumaroxidans might use the membrane potential to reduce NAD+ with ferredoxin re-oxidation in order to stimulate NADH re-oxidation of the comproportionating [FeFe]- hydrogenase.

7. Electron carrier systems Depending on the type of syntrophic conversion, the carrier system that transfers electrons from the producer to the consumer may vary. The best studied and best accepted electron carrier is hydrogen. However, formate is considered to be an important agent in interspecies electron transfer during propionate conversion as already discussed in section 1.6. Formate can also act as electron carrier in syntrophic butyrate conversion by S. wolfei since this bacterium contains a formate dehydrogenase with high homology to a formate dehydrogenase of Eubacterium acidaminophilum (FdhA-II) that was suggested to play a role also in interspecies formate transfer (Müller et al. 2009). In syntrophic acetone-degrading methanogenic cultures, acetate was identified as the only interspecies carrier compound (Platen and Schink 1987; Platen et al. 1994). In this syntrophic culture, growth and conversion of acetone to acetate proceeded until acetate had accumulated to ~10 mM. Addition of an active acetoclastic methanogen (Methanosaeta sp.) greatly enhanced the acetone degradation rate. In 14 addition, experiments with C-labeled CO2 showed that CO2 is stoichiometrically incorporated into the formed acetate (Platen and Schink 1987). Interspecies electron cycling through sulfur and sulfide has been described for Desulfuromonas acetoxidans in syntrophic cultures with Chlorobium limicola, a phototrophic green-sulfur bacterium (Pfennig and Biebl 1976; Biebl and Pfennig 1978). Acetate oxidation by D. acetoxidans and electron transfer to the phototrophic green sulfur bacterium C. limicola (Biebl and Pfennig 1978) occurred in the presence of small amounts of sulfide (53–92 μM) in the light (Biebl and Pfennig 1978). A similar sulfur-cycle-mediated electron transfer

was described in an artificial coculture which syntrophically oxidized acetate toCO2 with concomitant reduction of nitrate (Kaden et al. 2002). The discovery of bacterial nanowires and identification of presumed electron-transfer components required for electrical conductivity in these pili-like structures provided a novel view on mechanisms involved in interspecies electron transfer (Gorby et al. 2006; Reguera et al. 2005). Pili- like structures have been identified in a number of pure and mixed cultures, and also syntrophic cocultures of propionate-oxidizing Pelotomaculum thermopropionicum and Methanothermobacter thermoautotrophicus produced these pili-like structures. Analysis of the conductive properties of pili indicated that they could transfer electrons between cells of Geobacter sulfurreducens and the surface of Fe(III) oxides (Reguera et al. 2005). These pili were not required for attachment to the insoluble electron acceptor; rather they are interpreted to function as channels for electron transfer to the Fe(III) oxides, extending the electron transfer capabilities of the cells well beyond their outer surface (Reguera et al. 2005). Pili “nanowires” also served as electric conduits to mediate long-range electron transfer across biofilms formed on anode electrodes in microbial fuel cells which could maximize current production per unit of anode surface area (Reguera et al. 2005).

12 CHAPTER 1

Table 3. Phylogenetic groups of [NiFe]-hydrogenases as defined by Vignais and Billoud (2007).

Present in Commonly used gene names Bacteria / Function Archaea

hupSL, hypAB, hypAB, hyaAB, hoxKG, Bacteria Membrane-bound uptake hydrogenase mbhSL, hybOC, hynSL, hysAB

vhtGA, vhoGA Archaea Membrane-bound uptake hydrogenase

hupSL, mbhSL Bacteria Uptake hydrogenase

hypUV, hoxBC Bacteria Hydrogen sensor

frhAG, fruAG, frcAG Archaea F420-reducing hydrogenase

hydDA, shyDA, hyjSL Bacteria NADP-reducing hydrogenase

hyhSL, hydDA, hydδα, cytc3DA, Archaea NADP-reducing hydrogenase shyDA

mvhGA, vhcGA, vhuGA, mvhSL Archaea Methyl viologen reducing hydrogenase

Bidirectional NADP/NAD-reducing hoxYH Bacteria hydrogenase Energy converting membrane associated H hyfIG, hycGE, cooLH, echFE Bacteria 2 evolving hydrogenase Energy converting membrane associated H echFE, ehaNO, ehbMN Archaea 2 evolving hydrogenase

8. Hydrogenases When hydrogen and formate are the interspecies electron carriers, both syntrophs require hydrogenases and formate dehydrogenases for production or utilization of hydrogen and formate, respectively. Hydrogenases, enzymes that catalyze the reversible - + reaction: H2 ↔ 2e + 2H , have been extensively studied. Most hydrogenases are found in microorganisms belonging to the Archaea and the Bacteria domains, and few are present in Eukarya such as protozoans, fungi, algae and mosses. The first classification of hydrogenases was based on the specificity for certain electron donors and acceptors: co-enzyme F420 (hydrogenases of EC class 1.12.99.1), (1.12.2.1), ferredoxins (1.18.99.1), and NAD+ (1.12.1.12). The present classification is based on phylogeny and represents three different classes: [NiFe]-hydrogenases, [FeFe]-hydrogenases and [Fe]-hydrogenases. [Fe]-hydrogenases were previously referred to as metal-free hydrogenases (Lyon et al. 2004) and are restricted to some methanogens (Vignais and

13 General introduction

Billoud 2007). In Methanothermobacter marburgensis the [Fe]-hydrogenase catalyzes + the reversible reduction of methenyl-H4MPT with H2 to methylene-H4MPT and H only

under nickel-limiting conditions when the F420-reducing [NiFe]-hydrogenase is no longer synthesized (Thauer 1998). [NiFe]-hydrogenases however, are well studied in bacteria. The enzyme consists of at least two subunits, an α and β subunit that form a heterodimer with a bimetallic NiFe center in the α-subunit and iron-sulfur clusters in the β-subunit. The iron-sulfur clusters conduct electrons between the reactive core and the physiological electron acceptor or donor. The NiFe center is bound by two pairs of cysteine ligands. Sequence comparisons revealed two conserved regions surrounding these cysteine residues; the L1 and L2 motifs (Vignais and Billoud 2007). These motifs are conserved in each phylogenetic group of [NiFe]-hydrogenases (Table 3). Maturation of [NiFe]-hydrogenases involves at least seven proteins; HypA-F and an endopeptidase. Unlike [NiFe]-hydrogenases, [FeFe]-hydrogenases are composed of one up to four subunits and are found not only in prokaryotes, but also in eukaryotes. The catalytic subunit contains the (H-cluster) that consists of a binuclear [FeFe] center bound to a [4Fe4S] iron-sulfur cluster bridged by a cysteine (Schwab et al. 2006). Assembly of the active site requires HydE and HydG that belong to the radical S-adenosylmethionine (SAM) (Rubach et al. 2005; Sofia 2001).

9. Formate dehydrogenases - + - Formate dehydrogenases (EC 1.2.1.2.) catalyze the reaction HCOO → CO2 + H + 2e . Aerobic and anaerobic enzymes are biochemically and structurally very distinct (Ferry 1990). Formate dehydrogenases from aerobic bacteria and irreversibly oxidize formate with NAD+ and consist of homodimers without any (Ferry 1990). Aerobic formate dehydrogenases are used for regenerating NADH in enzymatic synthesis of optical active compounds with dehydrogenases and as such contribute to enzymatic processes in p.e. the pharmaceutical chemistry (Tishkov and Popov 2004). In contrast, many formate dehydrogenases from anaerobic bacteria and archaea are reversible and can react with a variety of electron carriers such as ferredoxin, quinone,

coenzyme F420, and NADP (Ferry 1990). Moreover, anaerobic formate dehydrogenases are composed of one, two or three subunits, contain iron-sulfur clusters and are extremely oxygen sensitive (Ferry 1990). The catalytic core contains molybdenum or tungsten bound to cysteine residues, of which one can be replaced by a residue (Groysman and Holm 2007). For the maturation of formate dehydrogenases FDH-D and FDH-E may be involved although the exact mechanism is not yet understood (Schlindwein et al. 1990). Based on sequence comparisons, FDH-D and -E were recognized as Redox Enzyme Maturation Proteins (REMP’s) (Turner et al. 2004). Recently, experiments proved that FdhE specifically binds to full length α-subunits of formate dehydrogenases from E. coli (FdnG and FdoG) (Chan et al. 2009). Formate hydrogen (FHL’s) are enzymes that catalyze the conversion of formate to hydrogen and

CO2. FHL’s are composed of a formate dehydrogenase, hydrogenase and electron transfer subunits which are described for several bacteria such as E. coli, Enterobacter

14 CHAPTER 1 aerogenes and Shewanella oneidensis MR-1 (Meshulam-Simon et al. 2007; Zhao et al. 2009; Bagramyan and Trchounian 2003). The FHL of E. coli is a membrane integrated complex facing the cytoplasm that may be involved in proton translocation (Bagramyan and Trchounian 2003; Trchounian et al. 1997), whereas that of S. oneidensis MR-1 is a periplasmic facing membrane associated complex with a mediating FHL activity (Meshulam-Simon et al. 2007). In bacteria mechanisms exist to translocate proteins across the cytoplasmic membrane. Most proteins travel through the cytoplasmic membrane in an unfolded conformation by the Sec-pathway and are recognized by hydrophobic N-terminal amino acids (Emanuelsson et al. 2007). Proteins such as formate dehydrogenases and hydrogenases, that bind cofactors, are cytoplasmically folded before transported through the membrane by the twin arginine translocation (Tat) pathway (Palmer et al. 2005). These proteins are recognized by the twin arginine (RR) motif in their hydrophobic N-terminus (Bendtsen et al. 2005). Archaea also contain a Tat pathway (Pohlschröder et al. 2005; Hutcheon and Bolhuis, 2003) that recognizes a slightly different (RR) motif (Bagos et al. 2009). Another feature of redox-enzymes such as formate dehydrogenases and hydrogenases is that in some cases the catalytic core contains selenium in the form of an integrated selenocysteine. The selenocysteine is abbreviated as Sec or U, and should not be confused with the above mentioned Sec-pathway. Selenocysteine containing proteins were found only a few decades ago in mammals and bacteria (Chambers et al. 1986; Zinoni et al. 1986) and later on also in archaea (Wilting et al. 1997). Comparative genomics revealed that selenocysteine incorporation is used by 22% of the sequenced bacteria representing mostlydelta Proteobacteria or Firmicutes/ Clostridia and that their proteomes may contain 1-31 selenoproteins (Zhang et al. 2006). Selenocysteine is co-transcriptionally inserted into the protein in response to a UGA codon which can also serve as the signal for termination of translation. Ribosomes recognize UGA as Sec codon by a stem-loop structure in the selenoprotein mRNA (Low and Berry 1996). For bacterial stem-loop structures, a consensus structural model and minimum open reading frame constrains have been determined (Fig. 5) (Zhang and Gladyshev 2005).

10. Alternative substrates for pure cultures, and technical systems to replace methanogens Outside the laboratory, bacterial communities are nearly always composed of a wide variety of species. It is appropriate to consider the relevance of these interspecies interactions to the outcome of activity assays and the cultivability in the laboratory. Defined cultures of syntrophically fermenting bacteria are required for detailed physiological and molecular studies and to understand their significant role in nature. To obtain such cultures, technical systems can be used to replace the methanogenic partner, or alternative substrates can be supplied to bypass the energetically unfavorable steps occurring in syntrophic conversions. The first axenic culture of an obligately syntrophic bacterium was S. wolfei (Beaty et al. 1987). Studies on the butyrate of syntrophic cocultures of S. wolfei and M. hungatei revealed a high activity of ß-oxidation enzymes (Wofford et al. 1986). With this knowledge Beaty and coworkers grewS. wolfei

15 General introduction

FigureZhang, 5 The Y. etconsensus al. Bioinformatics mRNA structure 2005 and 21:2580 minimum-2589; constrains doi:10.1093/bioinformatics/bti400 for bacterial stem-loop structures downstream of the Sec-UGA codon (bold and underlined). The consensus model includes: (i) a 3–14 nt apical loop and a 4–16 bp upper-stem, (ii) at least one guanosine (G) among the first two nucleotides in the apical loop, (iii) a spacing of 16–37 nt between the UGA codon and the apical loop and (iv) minimum free energy (MFE) –7.5 kcal/mol. Minimum ORF constraint includes at least one AUG/GUG codon between the Sec-UGA codon and the first upstream stop codon (Zhang and Gladyshev 2005).

on agar plates containing crotonate as the sole source of carbon and energy. The pure culture obtained dismutated crotonate to butyrate and acetate but exhibited butyrate oxidation only after re-association with a syntrophic partner. Later it was shown that S. wolfei and Syntrophospora bryantii could grow in pure culture on butyrate plus 3-pentenoate (Amos and McInerney 1990; Dong et al. 1994b). Butyrate plus 3-pentenoate were converted to valerate, acetate, and propionate. The first successful axenic culture of a syntrophically propionate-degrading bacterium was obtained from an enrichment culture by inhibiting the methanogens with bromoethanesulfonic acid (an analogue of methyl coenzyme M) and subsequently adding fumarate as external electron acceptor. This allowed to isolate S. fumaroxidans (Stams et al. 1993; Harmsen et al. 1998) and to study the pathway of propionate oxidation (Plugge et al. 1993). Phylogenetically, S. fumaroxidans is very closely related to sulfate-reducing bacteria. Some sulfate-reducing bacteria can alter their metabolism and act as syntrophically fermenting partners if sulfate becomes depleted (see above; Bryant et al. 1977; Scholten et al. 2007; Walker et al. 2009). Although this metabolic flexibility may be helpful for the enrichment and isolation of syntrophic bacteria it can be applied only to already highly enriched syntrophic cultures. A strategy for isolation of syntrophs could be stepwise: from enrichment culture via molecular characterization (16S rRNA based) to a strategic choice of substrate, electron acceptor or unsaturated compound for the isolation of the microorganism. Examples of unsaturated compounds used are fumarate, crotonate,

pentenoate, and benzoate. A cultivation apparatus capable of maintaining very low H2 (<0.01 Pa) pressures by mechanical means was developed by Valentine et al. (2000). This apparatus provided a method to study interspecies hydrogen transfer by externally providing the thermodynamic requirement for very low hydrogen concentrations, thus preventing the need for use of cocultures to study the metabolic pathways. The culture

16 CHAPTER 1 vessel is constructed of glass and operates by sparging a liquid culture with purified gases which remove hydrogen directly as it is produced. The culture device was constructed to decouple the syntrophic relationship in an ethanol-oxidizing methanogenic enrichment culture, allowing ethanol oxidation to dominate the methane production. Moreover, the culture apparatus was successfully used to grow pure cultures of the ethanol-oxidizing, proton-reducing P. acetylenicus (Valentine et al. 2000). This culture apparatus may have a potential to study also other forms of syntrophic metabolism, however, it should be realized that fatty acid oxidation requires hydrogen pressures substantially lower than ethanol oxidation.

11. Spatial organization of syntrophic communities The close cooperation of two metabolically different organisms during syntrophic degradation requires short transport paths between the partners to optimize metabolite transfer, especially at low overall energy yields. The metabolite flux from one organism to the other one is an inverse linear function of the diffusion distance (Schink and Thauer 1988). One should assume, therefore, that optimal transfer is ensured in mixed communities in which the partners are homogeneously mixed. Syntrophic cocultures show a defined tendency to form mixed aggregates also in defined laboratory cultures. However, since the respective partners are different organisms they multiply separately and will form sooner or later nests of genetically identical organisms which compete with each other within the nests and have only limited exchange to the partner nests outside. Likely such communities mix through each other to maintain optimal metabolite transfer at short distances. Microscopic pictures of methanogenic communities in biogas reactors have shown that nests, as described, really do exist within such structures, but that in other areas the partners appear to be fairly well-mixed (Grotenhuis et al. 1991; Fang et al. 1995; Harmsen et al. 1996). It is still an open question how such mixing can be accomplished by organisms that appear to be basically immotile and do not show any means of gliding motility.

12. Granular sludge in UASB reactors Up-flow anaerobic sludge blanket (UASB) technology is a form of anaerobic digester that is used in the treatment of wastewater. It is a sustainable technology applicable for a wide range of different industrial effluents (Lettinga 1995; Verstraete and Vandevivere 1999). Advantages of anaerobic over aerobic treatments are that less microbial biomass is produced and instead of consuming energy, useful energy in the form of biogas is produced (Plugge et al. 2010). Nowadays, this anaerobic digestion is applied worldwide for treatment of wastewater from, paper mill industry (Janssen et al. 2009), alcohol distillery (Zandvoort 2006), fish processing (Chowdhury et al. 2010), poultry and livestock (Sakar et al. 2009), agricultural resources (Ward et al. 2008), domestic sewage (Chernicharo 2006), and more. An important feature of UASB technology is that anaerobic microorganisms are densely packed into granules without the presence of any carrier material. The formation of granules is rather complex and involves physicochemical and biological interactions (Plugge et al. 2009b). The granules settle

17 General introduction

and form a sludge bed which prevents the microbes from being washed out at the top of the reactor (Fig. 6) (Plugge et al. 2009b). In the granules microbial processes; fermentation, acetogenesis and methanogenesis, take place at the same time. Changes in the environmental conditions often causes partially uncoupling of the different processes, which results in the accumulation of organic acids such as propionate and butyrate (Plugge et al. 2010). For instance changes in trace metal concentrations can be critical. Cobalt and nickel were shown to be essential for methanol degradation (Zandvoort et al. 2002; Fermoso et al. 2008). Molybdenum (Mo) and Tungsten (W) were shown to be important for propionate degradation in cocultures ofS. fumaroxidans and methanogenic M. hungatei (Plugge et al. 2009a). As was reviewed by Plugge et al. (2010), several microorganisms have been isolated and their physiological characteristics gave insight into their function in the bioreactor. 16S rRNA gene based molecular techniques gave insight into microbial dynamics and the stability of microbial ecosystems (reviewed by O’Flaherty et al. 2006). Still, more research with current and new molecular techniques is required to fill the knowledge gap related to ecological roles and metal requirement of microbial species in anaerobic bioreactors (O’Flaherty et al. 2006; Plugge et al. 2010). A better understanding of the fundamental microbial processes will improve current anaerobic digesters, and increase application possibilities (O’Flaherty et al. 2006).

13. Outline of this thesis Propionate degradation is a critical step in the overall degradation of organic material in UASB reactors. However, little is known about the molecular mechanisms of hydrogen and formate transfer from propionate oxidizing bacteria to methanogenic archaea. Formate dehydrogenases that are involved in formate transfer require the trace metals W/Mo and in some cases Se for enzymatic functioning. However, the effect of W, Mo and Se limitation on the propionate degrading community of a UASB reactor and on the

biogas

effluent

gas bubble

sludge bed influent

Figure 6 Schematic view of an up-flow anaerobic sludge bed reactor (modified from http://www.uasb. org/ and Plugge et al. 2010). Organic wastewater enters from the bottom, treated water elutes from the top and the produced methane and is collected and serves as biofuel.

18 CHAPTER 1 transcription of formate dehydrogenase coding genes were never examined. Recently the genome sequences of the propionate degrading Syntrophobacter fumaroxidans and the syntrophic methanogenic partner Methanospirillum hungatei became available and provided opportunities for bioinformatic and molecular approaches. The aim of this thesis is to get insight in the molecular mechanisms of hydrogen and formate transfer in pure cultures, syntrophic cocultures and UASB reactor sludge, by gene analysis and molecular techniques. Electron transfer mechanisms in S. fumaroxidans (syntrophic propionate degrader) and Syntrophomonas wolfei (syntrophic butyrate degrader) were studied with biochemical data and gene analysis (Chapter 2). These mechanisms comprise enzyme complexes such as hydrogenases, formate dehydrogenases and an Rnf-complex. We designed RT qPCR for genes coding for the main domains of enzyme complexes involved in electron transfer mechanisms in S. fumaroxidans and M. hungatei (Chapter 3). After qPCR, transcription profiles from cells grown in different conditions were compared. Chapter 4 describes the prediction of gene clusters that code for formate dehydrogenases and hydrogenases with gene analysis and microarray data. The involvement of formate dehydrogenases and hydrogenases in succinate oxidation and fumarate reduction was studied with gene analysis and transcription profiles (Chapter 5). Furthermore, the propionate degrading community of UASB reactor sludge and the effect of tungsten (W), molydenum (Mo) and selenium (Se) depletion from the effluent was analyzed with DGGE and 16S rRNA clone libraries (Chapter 6). Chapter 7 describes how RT qPCR primers (used for pure and defined cocultures in Chapter 2) were used to study transcriptional changes of formate and hydrogen transfer mechanisms in the metal depleted propionate-fed USAB reactor. In Chapter 8 the thesis research is put in a broader perspective, generated questions are discussed and ideas for future research are proposed.

19 20 CHAPTER 2

Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

Nicolai Müller*, Petra Worm*, Bernhard Schink , Alfons J. M. Stams, Caroline M. Plugge (*Both authors contributed equally)

Publication in Environmental Microbiology Reports DOI: 10.1111/j.1758-2229.2010.00147.x.

Abstract In anoxic environments such as swamps, rice fields, and sludge digestors, syntrophic microbial communities are important for decomposition of organic matter toCO2 and CH4. The most difficult step is the fermentative degradation of short-chain fatty acids such as propionate and butyrate. Conversion of these metabolites to acetate,

CO2, formate, and hydrogen is endergonic under standard conditions and occurs only if methanogens keep the concentrations of these intermediate products low. Butyrate and propionate degradation pathways include oxidation steps of comparably high redox potential, i. e., oxidation of butyryl-CoA to crotonyl-CoA and of succinate to fumarate, respectively, that require investment of energy to release the electrons as hydrogen or formate. Although investigated for several decades, the biochemistry of these reactions is still not completely understood. Genome analysis of the butyrate-oxidizing Syntrophomonas wolfei and Syntrophus aciditrophicus and of the propionate-oxidizing Syntrophobacter fumaroxidans and Pelotomaculum thermopropionicum reveals the presence of energy-transforming protein complexes. Recent studies indicated that S. wolfei uses electron-transferring flavoproteins coupled to a menaquinone loop to drive butyryl-CoA oxidation, and that S. fumaroxidans uses a periplasmic formate dehydrogenase, b:quinone oxidoreductases, a menaquinone loop, and a cytoplasmic fumarate reductase to drive energy-dependent succinate oxidation. Furthermore, we propose that homologues of the Thermotoga maritima bifurcating [FeFe]-hydrogenase are involved in NADH oxidation by S. wolfei and S. fumaroxidans to form hydrogen.

21 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

Introduction In anoxic environments such as swamps, rice paddy fields and intestines of higher animals, methanogenic communities are important for decomposition of organic

matter to CO2 and CH4 (Schink and Stams, 2006; McInerney et al. 2008; Stams and Plugge, 2009). Moreover, they are the key biocatalysts in anaerobic bioreactors that are used world-wide to treat industrial wastewaters and solid wastes. Different types of anaerobes have specified metabolic functions in the degradation pathway and depend on metabolite transfer which is called syntrophy (Schink and Stams, 2006). The study of syntrophic cooperation is essential to understand methanogenic conversions in different environments (McInerney et al. 2008). The most difficult step in this degradation is the conversion of short-chain fatty acids such as propionate and butyrate. Under standard

conditions (PH2 of 1 atm, substrate and product concentrations of 1M, temperature

298K), propionate and butyrate oxidation to H2, formate and acetate are endergonic

reactions (Table 1). In anoxic environments, methanogenic Archaea maintain low H2, formate, and acetate concentrations which make propionate and butyrate degradation feasible (Stams and Plugge, 2009). Syntrophic propionate and butyrate oxidations involve energy-dependent reactions that are biochemically not fully understood. However, recently several novel reactions were discussed to perform energy transformation in other bacteria. These reactions will be summarized in this report with respect to their possible implications in syntrophic fatty acid oxidation. Moreover, the genomes of two propionate degraders (Syntrophobacter fumaroxidans and Pelotomaculum thermopropionicum) and two butyrate degraders (Syntrophomonas wolfei and Syntrophus aciditrophicus) have been sequenced (McInerney et al. 2007; Kosaka et al. 2008). Based on genome analysis we propose that novel energy-transforming reactions are involved in syntrophic butyrate and propionate degradation.

Table 1. Standard free reaction enthalpies of fatty acid oxidation and methane production (values calculated from the standard free formation enthalpies of the reactants at a concentration of 1 M, pH 7.0 and T = 25°C according to (Thauer et al. 1977) Reaction ΔG0’ (kJ/reaction)

- - Propionate + 2 H2O --> Acetate + CO2 + 3 H2 eq. 1 + 76.0 - - - + Propionate + 2 H2O + 2 CO2 --> Acetate + 3 HCOO + 3 H eq. 1a + 65.3

- - + Butyrate + 2 H2O --> 2 Acetate + H + 2 H2 eq. 2 + 48.3 - - - + Butyrate + 2 H2O + 2 CO2 --> 2 Acetate + 2 HCOO + 2 H eq. 2a + 38.5

4 H2 + CO2 --> CH4 + 2 H2O eq. 3 -131.7 - + 4 HCOO + 4 H --> CH4 + 3 CO2 + 2 H2O eq. 4 -144.5

- + CH3COO + H --> CH4 + CO2 eq. 5 - 36

22 CHAPTER 2

Known energy-conserving mechanisms in syntrophic butyrate and propionate degradation Butyrate degradation Butyrate oxidizers known to date belong to two groups of bacteria within the family Syntrophomonadaceae and the order Syntrophobacterales. Formerly classified as Clostridia, the members of the family Syntrophomonadaceae have been reassigned to a new family within the order Clostridiales, based on their 16S rRNA sequence (Zhao et al. 1993). Members of this family are Syntrophomonas wolfei, Syntrophomonas bryantii (formerly Syntrophospora bryantii (Wu et al. 2006)), Syntrophomonas erecta, Syntrophomonas curvata, Syntrophomonas zehnderi and Thermosyntropha lipolytica. The second group of syntrophic butyrate degraders belongs to the Syntrophobacterales, an order of the delta-proteobacteria subdivision. Organisms of this group are Syntrophus aciditrophicus and . Several other Syntrophus strains such as Syntrophus gentianae are able to oxidize benzoate or other aromatic compounds syntrophically but these processes are not considered in this article. Remarkably, all these organisms are restricted to the use of saturated or unsaturated fatty acids. Alternative substrates or alternative electron acceptors to grow these bacteria in pure culture have not been found yet for these two groups of butyrate- oxidizing bacteria. In all known butyrate-oxidizing bacteria, the beta-oxidation pathway is used (Wofford et al. 1986; Schink, 1997; McInerney et al. 2007). First, butyrate is activated to butyryl- CoA with acetyl-CoA by a CoA . Further oxidation proceeds via crotonyl- CoA and 3-hydroxybutyryl-CoA to acetoacetyl-CoA which is cleaved to two acetyl-CoA moieties. One of these is invested in butyrate activation, and the other one forms ATP via phosphotransacetylase and acetate kinase (Wofford et al. 1986). Electrons are released in the oxidation of butyryl-CoA to crotonyl-CoA and in the oxidation of 3-hydroxybutyryl- CoA to acetoacetyl-CoA at -250 mV (Gustafson et al. 1986). As the standard midpoint redox potentials of these reducing equivalents are too high for reduction of protons to form H2 (-414 mV (Thauer et al. 1977; Schink, 1997)), it was postulated that an energy- dependent reversed electron transport is required to overcome the redox potential difference (Thauer and Morris, 1984). The partner organism keeps the hydrogen partial pressure low, thus raising the redox potential of proton reduction to a level around -300 mV (Schink, 1997). The butyrate-oxidizing organism has to sacrifice part of the gained ATP to shift electrons to this redox potential, and the remaining ATP can be used for biosynthesis and growth. As such fractional amounts of ATP cannot be provided by substrate level phosphorylation such energy transformations have to be coupled to the cytoplasmic membrane (Thauer and Morris, 1984). Indeed, hydrogen production from butyrate has been shown to be sensitive to the protonophore CCCP and the ATPase inhibitor DCCD, thus providing evidence for participation of a transmembrane proton potential (Wallrabenstein and Schink, 1994). However, the underlying biochemical mechanisms remained enigmatic until the completion of the genome sequence of S. aciditrophicus (McInerney et al. 2007). It was stated that electrons released in butyryl- CoA oxidation are transferred to components of the membrane where they reduce NAD+ to NADH in an endergonic manner, e. g., through an rnf-coded , and the

23 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

necessary energy would be supplied by a sodium ion gradient which in turn is provided by ATP-dependent proton efflux and a sodium/proton antiporter. In S. wolfei, however, a different reaction mechanism has to be active since the genome of this bacterium does not contain rnf genes which will be discussed later in this chapter (Müller et al. 2009).

Propionate degradation Several bacterial strains are known to degrade propionate in syntrophic association with methanogens; Syntrophobacter fumaroxidans, S. wolinii, S. pfennigii, S. sulfatireducens, Pelotomaculum thermopropionicum, P. schinkii, P. propionicicum, Smithella propionica and Desulfotomaculum thermobenzoicum subsp. thermosyntrophicum. These bacteria belong to the Syntrophobacterales, an order of the delta-proteobacteria subdivision, and to the family Peptococcaceae within the order Clostridiales. S. propionica converts propionate through a dismutating pathway to acetate and butyrate after which butyrate is oxidized to acetate (de Bok et al. 2001). All other known syntrophic propionate degraders

oxidize propionate to acetate plus CO2. They use the methylmalonyl-CoA pathway which generates per molecule propionate one ATP via substrate level phosphorylation and three electron pairs by; (i) oxidation of succinate to fumarate (Eo’=+30 mV), (ii) oxidation of malate to oxaloacetate (Eo’=-176 mV), and (iii) pyruvate conversion to acetyl-CoA and o o CO2 (E ’=-470 mV). The latter step can easily be coupled to proton reduction (E ’=-414 o mV) or CO2 (E ’=-432 mV) reduction (Thauer et al. 1977) via ferredoxin, as anaerobic bacteria generally contain pyruvate:ferredoxin oxidoreductases (Chabrière et al. 1999). Oxidation of succinate and malate with proto ns would require hydrogen partial pressures of 10-15 and 10-8 atm, respectively (Schink, 1997). Thauer and Morris (1984) and Schink (1997) proposed a reversed electron transport mechanism. The hydrolysis of ⅔ ATP coupled with a transmembrane import of two protons would make succinate oxidation energetically possible. Later, van Kuijk et al. (1998) proposed that this reaction is analogous to that involved in fumarate respiration by Wolinella succinogenes. This bacterium generates a transmembrane proton gradient by periplasmic hydrogen or formate oxidation coupled to cytoplasmic fumarate reduction via cytochromes and a menaquinone loop (Kröger et al. 2002). In S. fumaroxidans, fumarate reductase and activity are membrane bound. Hydrogenase and formate dehydrogenase activity are found in the periplasmic space loosely attached tothe membrane, and cells contain cytochrome c and b and menaquinone-6 and -7 as possible electron carriers (van Kuijk et al. 1998). S. fumaroxidans appears to gain around ⅔ ATP

per mol fumarate if H2 or formate is oxidized with fumarate. It was suggested that this mechanism in reverse could reduce periplasmic protons with the energy-dependent cytoplasmic succinate oxidation. Malate oxidation to oxaloacetate (Eo’= -176 mV) is coupled to NAD+ reduction (Eo’= -320 mV) (van Kuijk and Stams, 1996). Yet, the exact mechanism of NADH oxidation

and terminal reduction of protons and /or CO2 in S. fumaroxidans remains unclear and deserves further investigation.

24 CHAPTER 2

Mechanisms for energy conservation in anaerobic microorganisms Electron transport phosphorylation Electron transport phosphorylation is the most important energy-conserving mechanism in organisms that reduce external electron acceptors such as oxygen, nitrate, sulfate etc., (Richardson, 2000). In most cases, the membrane-bound NADH dehydrogenase (complex I) oxidizes NADH with quinones in the membrane while translocating protons into the periplasmic space via a transmembrane proton pump (Richardson, 2000). The electrons are transferred further to the respective terminal acceptor via cytochromes. Protons can be translocated to the periplasmic space by at least two possible mechanisms. The first one includes the translocation of protons or sodium ions through the transmembrane proton channel of an NADH dehydrogenase (Richardson, 2000). The second mechanism involves a redox loop and is supposed to be the most common way of proton translocation in bacteria (Richardson, 2000). Here, isoprenoid quinones within the membrane are reduced by the membrane-integral domain of the electron- donating enzyme, together with protons. The reduced quinone diffuses laterally through the membrane to the membrane domain of the accepting enzyme where electrons are transferred to an electron acceptor, and the protons are released at the opposite side of the membrane. An example is the redox loop of the formate dehydrogenase FDH-N coupled to nitrate reductase in (Jormakka et al. 2002). During anaerobic growth, formate is oxidized in the periplasm by FDH-N while protons are transferred to menaquinones, together with the electrons released in formate oxidation. Subsequently, menaquinol is oxidized at the membrane domain of the nitrate reductase, releasing protons to the periplasm while electrons are transferred to nitrate to form nitrite at the cytoplasmic side of the membrane (Jormakka et al. 2002).

The smallest quantum of energy in biology Since only small amounts of chemical energy can be transformed in syntrophic oxidation processes (Table 1), energy has to be efficiently conserved. Thauer et al. (1977) and Schink (1997) calculated that the minimal cost of synthesis of one ATP is 60 kJ per mol. Syntrophically fermenting bacteria such as butyrate and propionate oxidizers have to invest part of their ATP to create a proton gradient. For a long time it was thought that three protons are imported for synthesis of one ATP and thus the smallest quantum of energy that can be converted into ATP is in the range of 20 kJ per mol. However, Nakanishi-Matsui and Futai (2008) documented that the number of protons translocated is determined by the number of membrane-integral c-subunits of the ATP synthase which varies between different microorganisms. ATP synthases of and Enterococcus hirae harbor 10, while Ilyobacter tartaricus, Methanopyrus kandleri and chloroplasts harbor 11, 13 and 14 such subunits, respectively. The authors proposed that with one full rotation of the ATP synthase complex, three ATP are hydrolyzed, and each c-subunit translocates one proton. As a consequence, the number of protons translocated per ATP is between 3.3 and 4.6 and with this, the smallest quantum of biologically conservable energy may range from 13 to 18 kJ per mol reaction. In reversed electron transport as hypothesized for syntrophic butyrate and propionate oxidation, a high number of protons transported per ATP hydrolyzed would

25 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

allow ATP synthesis even at low energy gains. Hoehler et al. (2001) calculated minimal amounts of -10 to -19 kJ per translocated proton for organisms in anoxic methanogenic marine sediments. However, the number of membrane-integral c-subunits in ATP synthases of syntrophic bacteria has not been determined yet. Genome analysis of S. fumaroxidans and S. wolfei indicates the presence of one kind

of ATP synthase in each bacterium; the cytoplasmic F1 domain is encoded by Sfum_2581-

2587 and Swol_2381-2385, and the membrane-integral Fo domain is encoded by

Sfum_1604-1605 and Swol_2387-2388, respectively. The ratio of transcription ofF1 domain coding genes to membrane-integral c-subunit-coding genes might give insight into the number of c-subunits per ATP synthase in S. fumaroxidans and S. wolfei in the future.

Buckel-Thauer Bcd/Etf Most fermenting organisms have to regenerate their NAD+ pool in the absence of external electron acceptors. It was assumed that in clostridia NADH is oxidized with ferredoxin, which in turn is oxidized with protons to form hydrogen. This reaction is endergonic and, until recently, it was not known how hydrogen-producing microorganisms could perform such a reaction while strongly accumulating hydrogen in their environment. Clostridium kluyveri ferments ethanol plus acetate to a mix of butyrate, caproate, and hydrogen. The critical step of NADH oxidation with ferredoxin was recently found to be catalyzed by a Butyryl-CoA dehydrogenase (Bcd) / Electron-transferring Flavoprotein subunit (Etf) complex which couples this endergonic reaction to the exergonic reduction of crotonyl-CoA to butyryl-CoA with NADH (Li et al. 2008). Overall, two NADH molecules are oxidized and one molecule reduced ferredoxin (transferring two electrons) plus one molecule butyryl-CoA are formed (Li et al. 2008) which we refer to as bifurcation. For butyrate-oxidizing bacteria, a reversal of this reaction was suggested, i. e., the endergonic reduction of NAD+ with butyryl-CoA could be driven by the exergonic reduction of another NAD+ with reduced ferredoxin (Herrmann et al. 2008) a reaction which we refer to as confurcation. This mechanism could provide a concept for the reversed electron transport in syntrophic since homologues of this enzyme complex were found in genomes of the syntrophs S. wolfei, S. fumaroxidans, and P. thermopropionicum, but not in S. aciditrophicus.

Confurcating/bifurcating [FeFe]-hydrogenases Apart from the Buckel-Thauer Bcd/Etf-complex, another enzyme with bifurcating / confurcating activity was described recently, the [FeFe]-hydrogenase of Thermotoga

maritima (Schut and Adams, 2009). T. maritima ferments glucose to acetate, CO2

and H2 via the Embden-Meyerhof pathway which generates two NADH and four reduced ferredoxins per molecule of glucose. In order to re-oxidize these carriers, the proposed confurcating [FeFe]-hydrogenase uses simultaneously electrons from NADH and reduced ferredoxin in a 1:2 ratio to produce hydrogen (Schut and Adams, 2009). This hydrogenase could not use either NADH or reduced ferredoxin alone for hydrogen production. Additionally, the authors found genes with sequence similarity to this trimeric [FeFe]-hydrogenase also in other organisms such as S. fumaroxidans, P.

26 CHAPTER 2 thermopropionicum and S. wolfei (Table 2). Remarkably, our gene analysis indicated that some putative [NiFe]-hydrogenases and formate dehydrogenases inS. fumaroxidans, P. thermopropionicum, S. wolfei and S. aciditrophicus contain subunits with iron-sulfur- binding motifs and subunits homologous with the NADH dehydrogenase 51 kDa subunit, which is the NADH-binding subunit of Complex I (Figure 1). This indicates a possible confurcating function for [NiFe]-hydrogenases and formate dehydrogenases as well.

Rnf complex In Rhodobacter capsulatus nitrogen fixation (rnf) genes were found which code for a membrane-bound enzyme complex that is most probably involved in energy transformation (Kumagai et al. 1997). Gene analysis indicated that the encoded products RnfB and RnfC contain iron-sulfur clusters, RnfC contains potential NADH- and FMN- binding sites, and the membrane-bound RnfA, RnfD and RnfE are similar to subunits of the sodium-translocating NADH:quinone oxidoreductase (Kumagai et al. 1997). The authors proposed that this complex translocates protons or sodium ions to drive the endergonic reduction of ferredoxin by NADH oxidation. Analogousrnf genes were found in numerous bacteria such as Haemophilus influenzae, Escherichia coli, Acetobacterium

Figure 1 Schematic representation of putative formate dehydrogenases, hydrogenases, and Rnf- complexes in butyrate-degrading Syntrophomonas wolfei (A) and Syntrophus aciditrophicus (B), and in propionate-degrading Syntrophobacter fumaroxidans (C) and Pelotomaculum thermopropionicum (D). Gene locus tag numbers and α-, β-, and γ-subunits are indicated in small characters, predicted iron- sulfur clusters and metal-binding sites are indicated in capitals.“Formate” represent formate- + H+.

A formate CO2 FDH III Se [4Fe4S] W/Mo [4Fe4S] [4Fe4S] [4Fe4S] a 1825/6 1824

?cytb561 Xred 4TM 0797 4TM 1823 Xox formate Se FDH I W or Mo [4Fe4S] [4Fe4S] [4Fe4S] CO a [4Fe4S] 2 0799/0 b 0798 Se FDH II W/Mo 2formate g 0783 4[4Fe4S] 2Fdox 2Fdred [2Fe2S] [4Fe4S] NADH a 0785/6 b 0784 2CO2

NAD+ + H+ 2Fdox 2Fdred

2formate Se FDH IV W/Mo FeFe g 1829 g 1019 2CO2 3[4Fe4S] 4[4Fe4S] NADH 2H [2Fe2S] 2 [2Fe2S] NADH [4Fe4S] 1017 a1830/1 a [4Fe4S] b1828 b 1018 NAD+ + H+ 4H+ NAD+ + H+

2Fdox 2Fdred 2Fdox 2Fdred

27 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

formate formate CO2 B [4Fe4S] [4Fe4S] Se [4Fe4S] Se [4Fe4S] W/Mo [4Fe4S] CO2 W or Mo [4Fe4S] 2formate- ? Rnf [4Fe4S] [4Fe4S] [4Fe4S] [4Fe4S] 00604 00633 a00602/3 b a 00634/5 b

Ox-ac/For 4TM X X Antiporter 4 TM ox red 00632 Heme 02140 00605 A /E 01661 D 01663 A /E 01659

2formate- Se 4[4Fe4S] W or Mo [2Fe2S] 2[4Fe4S] NADH B 01658 G 01662 2formate- 3[4Fe4S] NADH C 01664 + [2Fe2S] 4H + 2CO2 [4Fe4S] Se a 00629 00630 NAD+ + W/Mo + + H NAD 2Fdox 2Fdred + H+ 4H+ + 2CO 2 4[4Fe4S] NADH 2Fdox 2Fdred [2Fe2S] [4Fe4S] a 02138 b 02139 NAD+ + H+ 2Fdox 2Fdred

2H2 2H2

02221 [4Fe4S] 4H+ 4H+ FeFe NiFe [4Fe4S] 02219 FAD NADH 3[4Fe4S] NADH a 02222 [2Fe2S] [4Fe4S] [2Fe2S] b 01369 02220 a 01370 NAD+ NAD+ + H+ + H+ 2Fd 2Fd 2Fdox 2Fdred ox red

FDH 2 FDH 3 FDH 4 formate formate formate C Hyd 2 H Se 2 Se W W or Mo W or Mo CO2 [4Fe4S] Ni Fe [4Fe4S] CO2 [4Fe4S] a1273/4 2952 a 3510 CO2 a + a 0035/6 2formate 2H 2[4Fe4S] Rnf [4Fe4S] 2[4Fe4S] b 1275 b 2953 b 3511 b 0037 Formate transporter Xred Xox Xred Xox Heme Xred Xox Xred Xox 2707 A /E 2696 D 2698 A /E 2695

4[4Fe4S] formate- formate- [2Fe2S] 2[4Fe4S] G 2697 2formate B 2694 C 2699 Se W or Mo W or Mo Se [2Fe2S]2Fe2S + 4[4Fe4S] 4[4Fe4S] H + CO2 H+ + CO WW g 2703 [2Fe2S] 2 [2Fe2S] 2Fd 2Fd NAD+ NADH 3509 2CO red ox a a 0030/1 2 + H+ 4[4Fe4S]4Fe4S 3[4Fe4S]4Fe4S [2Fe2S] [2Fe2S]2Fe2S 4Fe4S NADH H+ ? 2704 2705/6 b 2Fdox 2Fdred FDH 1 a 2Fdox 2Fdred NAD+ + H+ 2Fd ox 2Fdred

2H2 2H2 Hyd 3 Hyd 5 Hyd 6 0846 NiFe b 2715 g + + + + 4H+ FeFe 4H a 2716 2H H2 2H H2 2H H2 g 2714 3[4Fe4S] NADH NiFe NiFeSe NiFeSe [4Fe4S] 3537 3954 [2Fe2S] 3[4Fe4S] NADH a 2221 a a 0845 a 0844 b [2Fe2S] [4Fe4S] b 3536 3955 2714 b 2222 b NAD+ + H+ 2713 + + NAD + H 2220 3535 3956 2Fd ox 2Fdred

Hyd 1 2Fdox 2Fdred Hyd 4 Xox Xred Xox Xred Xox Xred

28 CHAPTER 2

formate- H+ + CO + D 2 H2 2H Se [4Fe4S] NiFe [4Fe4S] [4Fe4S] [4Fe4S] 2H2 W/Mo a1702 formate [4Fe4S] [4Fe4S] [4Fe4S] b 1713 a 1701 b 1703 a1711/2

Formate transporter 10 TM Xox Xred 10 TM Xox Xred 2651 a1714 a1704

g 2012 2650 3[4Fe4S] formate 2649 [2Fe2S] NADH Se 4[4Fe4S] [4Fe4S] W/Mo [2Fe2S] FeFe b 2011 2647 + CO2 a 2010 NAD 4[4Fe4S] NADH + + + H [2Fe2S] 3[4Fe4S] 2H2 4H a 2645/6 2648 NAD+ + H+ 2Fdox 2Fdred

2H2

g 1379 4H+ FeFe

2[4Fe4S] NADH [2Fe2S] 3[4Fe4S] 1378 a 1377 b NAD+ + H+

2Fdox 2Fdred woodii, and Vibrio alginolyticus, thus suggesting a general and important function for the Rnf complex in energy conservation (Müller et al. 2008; Nakayama et al. 2000; Backiel et al. 2008). Müller et al. (2008) investigated the function of an Rnf complex in the homoacetogenic bacterium A. woodii. Although biochemical proof has not been obtained yet, the authors found that caffeate respiration was coupled to ATP synthesis by a chemiosmotic mechanism with sodium ions as coupling ions, and that ferredoxin:NAD+-oxidoreductase was the only membrane-bound enzyme detected in the pathway of H2 dependent caffeate reduction. They postulated oxidation of ferredoxin with reduction of NAD+ and the export of Na+.

Methods used for gene analysis Automatic annotations of genomes from DOE-Joined Genome Institute (IMG-JGI- DOE, 2009) were used to indicate the presence of gene clusters coding for formate dehydrogenases, hydrogenases, Buckel-Thauer Bcd/Etf complexes and Rnf clusters in S. wolfei, S. aciditrophicus, P. thermopropionicum and S. fumaroxidans. N-terminal amino acid sequences of FDH-1 and FDH-2 identified by (de Bok et al. 2003) were used to find corresponding fdh-1 and fdh-2 nucleotide sequences in the genome of S. fumaroxidans. search (Sanger institute, 2009) was used to identify motifs in the amino acid sequences and the TMHMM Server v. 2.0 (DTU, 2009) was used to identify transmembrane helices. With the TatP 1.0 Server twin-arginine translocation

29 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

Table 2. General and genome-based characteristics of butyrate- and propionate-degrading syntrophic bacteria

Butyrate degraders Propionate degraders Syntrophomonas Syntrophus Syntrophobacter Pelotomaculum Bacterial species wolfei subsp. aciditrophicus fumaroxidans thermopropionicum wolfei Cell wall / Gram positive rod Gram negative rod Gram negative rod Gram positive rod morphology Delta Delta Class Clostridia Clostridia proteobacteria proteobacteria Methyl malonyl Pathway β-oxidation β-oxidation Methyl malonyl CoA CoA Genome Genbank CP000448 CP000252 CP000478 AP009389 accession SYN_02138-40 Swol_1829-31 [Se]-FDH Sfum_2703-07 PTH_2645-51 [Se]-FDH (FDH IV) SYN_00629-35 [Se]-FDH (FDH1) [Se]-FDH Genes coding Swol_0783-86 [Se]-FDH Sfum_0844-46 PTH_1377-79 for confurcating [Se]-FDH (FDH II) SYN_02219-22 [FeFe]-Hyd (Hyd1) [FeFe]-Hyd FDH’s and Hyd’sa) Swol_1017-19 [NiFe]-Hyd Sfum_2713-16 PTH_2010-12 [FeFe]-Hyd SYN_01369-70 [NiFe]-Hyd (Hyd4) [FeFe]-Hyd [FeFe]-Hyd Sfum_3510-11 FDH (FDH3) Genes coding SYN_00602-05 Sfum_0035-37 PTH_1711-14 for Tat-motif- Swol_1823-26 [Se]-FDH [Se]-FDH (FDH4) [Se]-FDH containing FDH’s [Se]-FDH (FDH III) SYN_00632-35 Sfum_1273-75 PTH_1701-04 and Hyd’sa,d) [Se]-FDH [Se]-FDH (FDH2) [NiFe]-Hyd Sfum_2952-53 [NiFe]- Hyd (Hyd2) Sfum_3509 FDH Sfum_0030-31 [Se]-FDH Genes coding for Sfum_2220-22 Swol_0797-00 other FDH’s and Not present [NiFe]-Hyd Not present [Se]-FDH (FDH I) Hyd’sa) Sfum_3535-37 [NiFeSe]-Hyd Sfum_3954-56 [NiFeSe]-Hyd Buckel -Thauer Sfum_1371-73 PTH_0015-17 No complete Bcd /Etf Swol_0267-68 Sfum_3929-21 PTH_2000-02 complex complexb) Sfum_3686-88 PTH_2431-33 Etf ABb) Swol_0696-97 SYN_02636-37 Sfum_0106-07 PTH_1552-53 Rnf clusterc) Not present Syn_01658-64 Sfum_2694-99 Not present

Gene locus tag numbers of genes in butyrate and propionate degraders, that show similarity with genes coding for energy-transforming protein complexes such as a) formate dehydrogenases (FDH), Hydrogenases (Hyd), b) Electron-transferring flavoproteins (Etf), Butyryl-CoA dehydrogenase (Bcd) and c)Rhodobacter capsulatus nitrogen fixation (Rnf) complexes. d) Twin arginine translocation (Tat) motives indicate that the corresponding proteins are translocated through the cell membrane.

30 CHAPTER 2

(Tat) motifs in the N-terminus were identified to predict protein localization in the cell (Bendtsen et al. 2005). The incorporation of selenocystein (SeCys) was examined by RNA loop predictions with Mfold version 3.2 (Mathews et al. 1999; Zuker, 2003). The RNA loop predicted in the 50-100 bp region downstream of the UGA-codon was compared with the consensus loop described by Zhang and Gladyshev (2005). Complete amino acid sequences of putative selenocystein-containing formate dehydrogenases were aligned to their homologues with ClustalX 1.81 (Kryukov and Gladyshev, 2004). SeCys incorporation was confirmed when the amino acid sequence aligned with conventional cystein in homologous proteins.

Hypotheses for energy conservation mechanisms in butyrate oxidizers and propionate oxidizers Butyrate oxidation in S. wolfei The electron transport in butyrate oxidation byS. wolfei was studied recently in a classical biochemical approach (Müller et al. 2009). The electron transfer from butyryl-CoA to external electron acceptors was found to be inhibited by trifluoperazine, a compound known to inhibit electron transfer to menaquinone by the respiratory complex I in Mycobacterium tuberculosis (Yano et al. 2006). Trifluoperazine also inhibited electron transfer from NADH to quinone analogues. An NADH-oxidizing enzyme complex was partially purified from the membrane fraction of S. wolfei. This activity was also found in the cytoplasmic fraction, especially after repeated treatment in the French Press cell, indicating that it is only superficially associated with the membrane (Müller et al. 2009). The enzyme complex contained several proteins which were analyzed by peptide mass fingerprinting and were compared via the known gene sequence with redox enzymes found in other bacteria, i. e. an enzyme system similar to the confurcating hydrogenase of T. maritima (Schut and Adams, 2009). Moreover, this hydrogenase homologue of S. wolfei appeared to be associated with homologues of an NADH-dependent formate dehydrogenase of Eubacterium acidaminophilum (Graentzdoerffer et al. 2003), a bacterium which can also grow in syntrophic association with partner organisms (Zindel et al. 1988). This enzyme complex could either act as a confurcating hydrogenase / formate dehydrogenase or as a proton-pumping NADH dehydrogenase (NDH) (Müller et al. 2009). So far, it was not possible to show if this enzyme is directly linked to a transmembrane proton channel (Müller et al. 2009). But even if the enzyme lacks such a channel protons could be transferred via a menaquinone cycle (Figure 2) as described above for E. coli (Jormakka et al. 2002). Whether the hydrogenase found really acts in a bifurcating manner as observed in T. maritima has still to be verified. So far, there is no indication of a ferredoxin-coupled redox reaction in butyrate oxidation bythis bacterium. Our results indicate that the “Buckel-Thauer” reaction, i.e., a bifurcation of electrons from NADH with crotonyl-CoA and oxidized ferredoxin by the Bcd/EtfAB complex of C. kluyveri (Herrmann et al. 2008; Li et al. 2008) is not involved in butyrate oxidation by this bacterium. Until now, the function of etf genes in syntrophic butyrate degraders remains unclear. Possibly, the Bcd/EtfAB complex is expressed when S. wolfei grows by dismutation of crotonate. Whether other butyrate oxidizers, e. g., S. aciditrophicus

31 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

butyrate 2 H+

MKH2 MK - 2e Fdh-II formate + foc 2 H + [FeFe]Hyd CO2 + 2 H + H2 2 H Methanogen 2 H+ Etf EtfH2 NAD+ + H+ NADH NAD+ NADH + H+

butyrate butyryl-CoA crotonyl-CoA 3-hydroxy- acetoacetyl- butyryl-CoA CoA Murein H2O layer CoA-SH acetyl-CoA acetate acetyl-CoA ADP ATP nH+ ATP ase acetate

Cytoplasmic membrane

acetate acetate

Figure 2 Hypothetical energy-transforming mechanisms in the butyrate-degrading Syntrophomonas wolfei. The fatty acids acetate, butyrate and formate represent acetate- + H+, butyrate- + H+ and formate- + H+, respectively. (foc) represents a formate transporter.

or S. buswellii, employ the Bcd/EtfAB complex in butyrate oxidation remains an open question at this time. Another interesting feature of the hydrogenase homologue of S. wolfei is its association with a formate dehydrogenase (Müller et al. 2009). This supports older speculations that electrons from NADH oxidation are released as formate rather than hydrogen. The bacterium might even be able to choose which carrier it prefers, depending on the environmental conditions (Graentzdoerfferet al. 2003), e. g., whether a partner is present which consumes hydrogen, formate or both, and this preference might even differ between different butyrate oxidizers: Coculture experiments with S. bryantii and different partners (Dong et al. 1994b) showed highest growth and substrate conversion rates with Methanospirillum hungatei which uses both hydrogen and formate, whereas cocultures with the mainly formate-oxidizing Methanobacterium formicicum were slower, and there was no growth at all with the only hydrogen- consuming Methanobrevibacter arboriphilus. In contrast, it was shown earlier that S. wolfei grows in the presence of M. arboriphilus, although to a lower extent, indicating that S. wolfei can grow by interspecies hydrogen transfer only and formate plays only a minor role in electron transfer (McInerney et al. 1979, McInerney et al. 1981). Therefore, the electron transport during butyrate oxidation by S. bryantii might be different from that described above for S. wolfei, although the molecular prerequisites might be similar due to the close relatedness of both organisms. The formate dehydrogenase of S. bryantii was found to be membrane-bound and was most likely oriented to the periplasmic space whereas the partly membrane-bound hydrogenase

32 CHAPTER 2 was found in the cytoplasm and showed also activity with NAD+ as electron acceptor (Dong and Stams, 1995). It was assumed that hydrogen is produced inside the cell while formate is produced outside. Additionally, an NADH dehydrogenase reacting with the tetrazolium dye MTT was measured, comparable to the described NADH: quinone oxidoreductase in S. wolfei which also reacts with MTT (Müller et al. 2009), (Müller unpublished data). It is tempting to speculate at this point that the described NADH dehydrogenase of S. wolfei could also be a bifurcating hydrogenase that couples NADH- dependent proton reduction with quinone reduction by another molecule of NADH. One electron pair would then be used to reduce protons to form hydrogen and the other one would be transferred to the external formate dehydrogenase. If hydrogenase and/or NADH dehydrogenase are coupled to the formate dehydrogenase via a quinone- mediated redox loop, two additional protons would be transferred outside the cell. Overall, proton consumption in the cytoplasmic space and proton release at the outside would result in a net proton gradient which in turn could drive menaquinol oxidation with NAD+ or ADP phosphorylation by proton influx. This would require that formate and hydrogen both have to be kept at low concentration to allow the thermodynamically unfavourable reactions of CO2 reduction with quinols and proton reduction with NADH. Although such a membrane-bound and quinone-oxidizing formate dehydrogenase has not yet been measured in S. wolfei, there are indications for such a system in its genome (Swol _0797-Swol_0799) (Table 2).

Although the formate/CO2 couple and the hydrogen/proton couple are both at the same redox potential under physiological conditions, the question remains whether they are really equivalent inside the cell as assumed earlier (Schink, 1997). Cocultures of Moorella sp. strain AMP and Desulfovibrio sp. strain G11 with formate as substrate in coculture with hydrogen-only consuming methanogens converted formate to methane (Dolfing et al. 2008). It was assumed that formate is oxidized outside the cytoplasmic membrane, CO2 and protons are released and electrons are shuttled to a membrane- bound hydrogenase facing the cytoplasm where protons are consumed (Dolfing et al. 2008). Thus, a net positive membrane potential could be formed without direct proton translocation but there is so far no proof that this reaction is coupled to energy conservation.

Propionate oxidation by S. fumaroxidans and P. thermopropionicum The most difficult step in syntrophic propionate oxidation is the oxidation of succinate to fumarate. In the past, succinate dehydrogenases and fumarate reductases have been found to be similar based on their amino acid sequence (Lancaster, 2002). These authors classified fumarate reductases in five groups based on molecular composition. The fumarate reductase of W. succinogenes was classified within the group containing one hydrophobic subunit and two heme-groups (Kröger et al. 2002). Our present gene analyses indicate that not only hydrophobic subunits of the fumarate reductase but also those of formate dehydrogenases of W. succinogenes (formate dehydrogenase delta subunits: WS0027, WS0736 and WS1148) contain heme groups and are homologous to cytochrome b.

33 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

S. fumaroxidans genome analysis revealed the presence of periplasmic formate dehydrogenases and hydrogenases (Table 2) as well as cytoplasmic fumarate reductases (Sfum_4092-4095, Sfum_1998-2000) which lack heme groups and a cytochrome b-like membrane-integrated domain. As such, fumarate reductases of S. fumaroxidans could not be classified within the five types described by Lancaster (2002). Instead, scattered over the genome, three cytochrome b (cytb561; Sfum_0091, cytb5; Sfum_3227 and cytb; Sfum_2932) and three cytochrome c homologous genes (Sfum_0090, Sfum_4047 and Sfum_1148) were found. Moreover, three genes with homology to cytochrome b: quinone oxidoreductases were found (Sfum_0339, Sfum_3009 and Sfum_3051). Cytochrome b and cytochrome b:quinone oxidoreductases possibly function in a similar way as the cytochrome-containing membrane-integrated domains of fumarate reductases, hydrogenases, and formate dehydrogenases of W. succinogenes (Figure 3). Candidates for periplasmic formate or hydrogen oxidation are formate dehydrogenase 2, 3 and 4 and hydrogenase 2 (Figure 3). These proteins may bind periplasmic cytochrome c and hydrophobic cytochrome b for succinate oxidation or fumarate reduction. Probably they also interconvert hydrogen and formate via a cytochrome c network, as proposed previously for the sulfate-reducing Desulfovibrio vulgaris Hildenborough (Heidelberg et

al. 2004). Interconversion of hydrogen plus CO2 and formate by S. fumaroxidans was observed by Dong and Stams (1995) and de Bok et al. (2002b). Formate dehydrogenase 1 of S. fumaroxidans was previously characterized (de Bok et al. 2003). It oxidizes formate with benzyl viologen as artificial electron acceptor, but NAD+ did not support oxidation of formate. Based on the amino acid sequence, this selenocystein-containing formate dehydrogenase is similar to FDHII of S. wolfei and to the NADH-dependent formate dehydrogenase of Eubacterium acidaminophilum (Graentzdoerffer et al. 2003). Whether formate dehydrogenase 1 can couple oxidation of

both NADH and ferredoxin to CO2 reduction in a manner analogous to proton reduction by the confurcating [FeFe]-hydrogenase of T. maritima was never tested. Based on our gene analysis, we hypothesize that hydrogenase 1, hydrogenase 4, and formate dehydrogenase 1 can couple the oxidation of NADH generated from malate oxidation with the oxidation of reduced ferredoxin generated from pyruvate oxidation to produce hydrogen or formate (Table 2, Figure 3). Especially transcription of genes coding for hydrogenase 1 (an [FeFe]-hydrogenase) appears to be up-regulated when metabolic conversions generate NADH and reduced ferredoxin (P. Worm et al. unpublished). In S. fumaroxidans the Rnf-complex might be used to conserve the energy of ferredoxin oxidation with NADH reduction by exporting protons, thus equilibrating theratioof reduced ferredoxin to NADH to 1:1. Compared to S. fumaroxidans, the genome of P. thermopropionicum contains less formate dehydrogenase- and hydrogenase-coding genes (Table 2). However, for each metabolic task, several candidates are present just as in S. fumaroxidans. In order to reoxidize the NADH and reduced ferredoxin that are generated during propionate degradation,P. thermopropionicum likely uses the confurcating formate dehydrogenase (PTH_2645-2649) and the two confurcating [FeFe]-hydrogenases (PTH_1377-1379 and PTH_2010-2012) (Kosaka et al. 2008). The produced formate would be transported through the membrane via a formate transporter (PTH_2651) of which the gene is

34 CHAPTER 2

+ formate CO2 + 2H Hyd 2 nH+ H+ ? FDH 2

- MK - 2e 2e Rnf MKH2 ATP 2H+ 2H+ ase + + FRD NAD + H NADH 2 Fdred 2 Fdox

ADP ATP 2H+

ATP CoA-SH [CO2] ADP succinate fumarate malate oxaloacetate pyruvate succinyl-CoA NAD+ NADH CoA-SH FDH 1 methylmalonyl-CoA 2Fdox Hyd 1 CO2 Hyd 4 2Fd [CO ] red 2 confurcating enzymes propionyl-CoA acetyl-CoA CO2 CoA-SH

propionate acetate formate H2 foc Cytoplasmic membrane

formate propionate H2 acetate

Methanogen

Figure 3 Hypothetical energy-transforming mechanisms in the propionate-degradingSyntrophobacter fumaroxidans. The fatty acids acetate, propionate and formate represent acetate- + H+, propionate- + - + + H and formate + H , respectively. (foc) represents a formate transporter, [CO2] a biotin-bound carboxylic group.

located in the operon coding for the cytoplasmic formate dehydrogenase. The produced hydrogen diffuses through the membrane and is used by the methanogen. Also similar to S. fumaroxidans is the mechanism of reversed electron transfer via fumarate reductase, a menaquinone loop and a periplasmic formate dehydrogenase (PTH_1711-1714) or hydrogenase (PTH_1701-1704) (Kosaka et al. 2008). Cytoplasmic and periplasmic formate dehydrogenases and hydrogenases could be used to interconvert formate and hydrogen. P. thermopropionicum can grow only with a hydrogen-using methanogen as its syntrophic partner (Ishii et al. 2005), however, formate is likely to generate hydrogen. The high number of formate dehydrogenase- and hydrogenase-encoding genes in S. fumaroxidans likely provides S. fumaroxidans with more back-up possibilities when formate and hydrogen concentrations vary according to the activity of the partner methanogen. In contrast to S. fumaroxidans, P. thermopropionicum lacks an Rnf cluster and ferredoxin-reducing hydrogenases and formate dehydrogenases. S. furmaroxidans might use these mechanisms as alternatives to reoxidize NADH and ferredoxin, possibly with the use of an electron potential via the Rnf-cluster, when environmental conditions change.

35 Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms

Concluding remarks Recent biochemical studies and genome analyses indicated that S. wolfei uses electron- transferring flavoproteins coupled to a menaquinone loop to drive endergonic butyryl- CoA oxidation, and S. fumaroxidans uses a periplasmic formate dehydrogenase, cytochrome b:quinone oxidoreductases, a menaquione loop and a cytoplasmic fumarate reductase to drive endergonic succinate oxidation. Furthermore, we propose that confurcating [FeFe]-hydrogenases in S. wolfei and S. fumaroxidans are involved in NADH oxidation to form hydrogen. For both S. wolfei and S. fumaroxidans, a similar function is proposed for a formate dehydrogenase which would result in simultaneous hydrogen and formate transfer from the fermenting bacterium to the hydrogen- and formate-consuming syntrophic partner. S. fumaroxidans and S. wolfei are proposed to produce hydrogen and formate in the cytoplasm. P. thermopropionicum and S. wolfei are proposed to contain a mechanism to convert hydrogen into formate which would allow growth with hydrogen-only using methanogens. These proposed energy- converting mechanisms need biochemical verification. We hypothesize that they are key in syntrophic propionate- and butyrate-degrading communities, as well as in other syntrophic communities.

Acknowledgements The authors were financially supported by the German Research Foundation (DFG) and by the Earth and Life Sciences division (ALW) and Chemical Science division (CW) of the Netherlands Organization for Scientific Research (NWO).

36 CHAPTER 3

Growth and substrate dependent transcription of formate dehydrogenase and hydrogenase coding genes in Syntrophobacter fumaroxidans and Methanospirillum hungatei

Petra Worm, Alfons J. M. Stams, Xu Cheng and Caroline M. Plugge Submitted for publication

Abstract Transcription of genes coding for formate dehydrogenases (fdh’s) and hydrogenases (hyd’s) in Syntrophobacter fumaroxidans and Methanospirillum hungatei was studied in different growth conditions. Under all conditions tested all fdh’s and hyd’s were transcribed. However, transcription levels of the individual fdh’s and hyd’s varied dependent on substrate and growth conditions. Our results show that in syntrophically grown S. fumaroxidans cells, the [FeFe]-hydrogenase (encoded by Sfum_844-46) and formate dehydrogenase 1 (encoded by Sfum_2703-06) are the most important enzymes that confurcate electrons from NADH and ferredoxin to protons and carbon dioxide to produce hydrogen and formate, respectively. The periplasmic formate dehydrogenase 2 (Sfum_1273-75) of S. fumaroxidans is the most important periplasmic proton using enzyme that is involved in cytoplasmic succinate oxidation to fumarate. InM. hungatei, a membrane integrated energy converting [NiFe]-hydrogenase (Mhun_1741-46) is involved in the energy dependent CO2 reduction to formylmethanofuran. The best candidates 5 10 5 10 for F420-dependent N ,N -methyl-H4 MPT and N ,N ,-methylene-H4MPT reduction are the cytoplasmic [NiFe]-hydrogenase and formate dehydrogenase 1. The membrane bound hydrogenase likely is involved in the last step of the methanogenic pathway, the reduction of methyl coenzyme-M to methane. In one of the triplicate cocultures of S. fumaroxidans and M. hungatei, less energy was available for S. fumaroxidans which led to an enhanced transcription of genes coding for the Rnf-complex and several formate dehydrogenases and hydrogenases. The Rnf-complex probably re-oxidized NADH with ferredoxin reduction, which was followed by ferredoxin oxidation by the induced formate dehydrogenases and hydrogenases.

37 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

Introduction In anaerobic environments degradation of complex organic matter to carbon dioxide and methane is performed by consortia of microorganisms that each have their specific metabolic function (McInerney et al. 2008, 1981; Schink and Stams, 2006). Important metabolic intermediates are organic acids such as propionate and butyrate. Their degradation depends on syntrophy between bacteria and methanogenic archaea. Even under optimal growth conditions, the available free energy is low and has to be shared by all partners in the consortia (Schink and Stams, 2006). Syntrophobacter fumaroxidans is a δ-proteobacterium that degrades propionate

in syntrophic association with the2 H and formate-using Methanospirillum hungatei or Methanobacterium formicicum (Harmsen et al. 1998). Cultivation and biochemical experiments indicated that both hydrogen and formate transfer are important interspecies electron transfer mechanisms (de Bok et al. 2002a; de Bok et al. 2002b; Harmsen et al. 1998). Propionate degradation by S. fumaroxidans in the absence

of an electron acceptor, such as sulfate or fumarate, requires low H2 and formate concentrations (1 Pa and 10 µM) in order to gain energy for growth (Schink and Stams,

2006). M. hungatei and M. formicicum are able to maintain such low H2 and formate concentrations. Although syntrophic partners regulate their metabolism to grow together, there are conflicting needs. Growth of the methanogens would be greatly

enhanced if H2 and formate concentrations are high. S. fumaroxidans metabolizes propionate via the methylmalonyl-CoA pathway (Plugge et al. 1993). Electrons are generated in three steps: oxidation of succinate to fumarate, oxidation of malate to oxaloacetate, and oxidation of pyruvate to acetyl-CoA plus carbon dioxide. To remove these reducing equivalents, S. fumaroxidans forms hydrogen and formate that is transferred and further metabolized by the methanogenic partner. In pure culture the bacterium is able to use fumarate and sulfate as electron acceptor (Harmsen et al. 1998). S. fumaroxidans can also grow with hydrogen and fumarate, formate and fumarate and even by fumarate fermentation (Harmsen et al. 1998). Analysis of the genome sequence of S. fumaroxidans revealed the presence of four formate dehydrogenases (FDH1-4), six hydrogenases and one [NiFe]-hydrogenase maturation protein (Müller et al. 2010). The six hydrogenases include a cytoplasmic [FeFe]-hydrogenase (FeHyd), two cytoplasmic [NiFe]-hydrogenases (Frh and Hox), a periplasmic [NiFe]-hydrogenase (Hyn), and two cytoplasmic ferredoxin dependent [NiFeSe]-hydrogenases (Hdr and Fnr) (Coppi, 2005). In addition, a gene cluster coding

for cytoplasmic oriented membrane proteins to interconvert H2 plus carbon dioxide and formate, referred to as a formate hydrogen complex, is present. These genes show similarity with those coding for a formate dehydrogenase (FHL-F), a hydrogenase (FHL-H) and several iron-sulfur cluster binding proteins (Müller et al. 2010). Unlike the FHL of E. coli (Bagramyan, and Trchounian, 2003), genes coding for membrane integrated subunits were lacking in the FHL coding gene cluster of S. fumaroxidans. M. hungatei uses the hydrogen and formate generated by S. fumaroxidans to form methane. In hydrogenotrophic methanogens, electrons are required in four reduction 5 10 steps; the conversion of carbon dioxide to formylmethanofuran, N ,N -methyl-H4MPT 5 10 5 10 5 to N ,N -methylene-H4MPT, N ,N -methylene-H4MPT to N -methyl-H4MPT and

38 CHAPTER 3 methyl coenzyme-M to methane and coenzyme M (Schwörer and Thauer, 1980). In M. hungatei hydrogenases and formate dehydrogenases oxidize hydrogen and formate to supply electrons for the reduction reactions. Analysis of the genome sequence of M. hungatei indicated the presence of five formate dehydrogenases (FDH1-5) and three hydrogenases (IMG-JGI-DOE, 2009). The three hydrogenases include a membrane integrated ion-translocating energy-converting [NiFe]-hydrogenase (Ech), a cytoplasmic

F420-reducing hydrogenase (Frc), and an ion-translocating membrane bound [NiFe]- hydrogenase (Mbh) (Hendrickson and Leigh, 2008; Thauer et al. 2008). The genome of M. formicicum has not been sequenced yet. In this study we aim to understand the relative transcription and differential transcription of genes coding for formate dehydrogenases (fdh’s) and hydrogenases (hyd’s) in S. fumaroxidans and M. hungatei and to get insight into syntrophic interactions between S. fumaroxidans and the methanogens M. hungatei and M. formicicum.

Materials and methods Microorganisms and growth conditions. Syntrophobacter fumaroxidans MPOB (DSM 10017), the methanogenic archaea Methanospirillum hungatei JF−1 (DSM 864) and Methanobacterium formicicum JF-1 (DSM 1535) were used in this study. S. fumaroxidans was either grown in pure culture or in coculture with one of the methanogenic archaea. All cultures were grown at 37°C

TABLE 1. Tested growth conditions of S. fumaroxidans and M. hungatei and generation times from triplicate cultures. Culture Generation times Code Organism or coculture Substrate volume (mL) (days) in 20 mM propionate + PF S. fumaroxidans 50 1.4 1.3 1.4 60 mM fumarate

F S. fumaroxidans 30 mM fumarate 50 3.7 2.9 2.6

30 mM fumarate + HF S. fumaroxidans 50 2.1 2.2 2.2 H2/CO2 (80:20, vol/vol) 30 mM fumarate + FF S. fumaroxidans 50 2.7 1.4 1.4 30 mM formate 30 mM propionate + PS S. fumaroxidans 500 9.3 5.9 7.0 30 mM sulfate S. fumaroxidans + 5.8 4.1 3.8 CoMh 30 mM propionate 500 M. hungatei a(45) (4.7) (6.9) S. fumaroxidans + 8.2 6.8 8.4 CoMf 30 mM propionate 500 M. formicicum b(7.3) (2.9) (1.6) 30 mM formate + 1mM For M. hungatei 500 0.6 0.5 0.4 acetate H /CO (80:20, vol/vol) + H M. hungatei 2 2 500 1.8 2.2 1.7 2 1mM acetate a) The ratio of 16SrRNA from M. hungatei / 16SrRNA from S. fumaroxidans is shown in brackets. b) The ratio of 16SrRNA from M. formicium / 16SrRNA from S. fumaroxidans is shown in brackets.

39 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

in anaerobic liquid medium with the following composition: 3 mM Na2HPO4, 3 mM

KH2PO4, 5.6 mM NH4Cl, 0.68 mM CaCl2, 0.6 mM MgCl2, 5mM NaCl, 48 mM NaHCO3, 4

mM cysteine HCl, 1mM Na2S, 7.5 µM FeCl2, 1 µM H3BO3, 50 µM HCl, 0.5 µM ZnCl2, 0.5 µM

MnCl2, 0.5 µM CoCl2, 0.1 µM NiCl2, 0.1 µM Na2SeO3, 0.1 µM Na2WO4, 0.1 µM Na2MoO4, vitamins (µg.L-1); 0.02 biotin, 0.2 nicotinic acid, 0.5 pyridoxine, 0.1 riboflavin, 0.2 thiamin, 0.1 cyanocobalamin, 0.1 p-aminobenzoic acid, 0.1 pantothenic acid. Microorganisms were cultured in 120 mL or 1 L flasks with 50 mL or 500 mL medium, respectively, and

a headspace of 1.7 atm. N2/CO2 or H2/CO2 (80:20, vol/vol). In order to study differential gene expression, RNA was isolated from S. fumaroxidans and M. hungatei cells, grown under different conditions (Table 1). All cultivations were done in triplicate. Growth was followed by measuring optical density and/or methane formation. Cells were harvested in the logarithmic phase.

RNA extraction and reverse transcription Genes coding for formate dehydrogenases and hydrogenases in the genome of M. hungatei were analyzed with the same bioinformatic tools as were previously described for S. fumaroxidans (Müller et al. 2010). With PRED-SIGNAL (Bagos et al. 2009) twin- arginine translocation (Tat) motifs in the N-termini were identified to predict the cell localization of proteins in M. hungatei cells. Cultures were harvested by centrifugation at 3800 × g at 4°C for 20 min. Cell pellets were resuspended in RNA stabilization solution “RNAlater” (Ambion), incubated at 4°C overnight and stored at –20°C. Stored cells were centrifuged 3800 × g at 4°C for 10 min and the resulting cell pellet was resuspended in 125 µL RNase free water and 375 µL TRIzol Reagent (Invitrogen), transferred to a tube containing 0.1 mm silica beads and homogenized 4.5 m.s-1 for 30 s in a FastPrep- 24 (MP Biomedicals). Tubes were incubated at 20 °C for 5 min and 0.1 mL chloroform was added followed by 15 s manual shaking. After incubation at 20 °C for 5 min, the suspension was centrifuged at 9500 × g at 4 °C for 15 min to separate the aqueous phase from the phenol phase. The aqueous phase with an equal amount of 70 % ethanol was transferred to an RNeasy mini spin column (Qiagen) and on-column DNase digestion with RNase-free DNase I (Qiagen) was performed according to manufacturer’s instructions. RNA was ultimately eluted in 60 µL RNase free water and RNA integrity was confirmed by an Experion RNA StdSens Chip (Bio-Rad) using the Bio-Rad Experion system. Copy DNA was synthesized by SuperScript III reverse transcriptase (Invitrogen) according to manufacturer’s instructions with random decamers (Ambion). To distinguish genomic DNA from cDNA in qPCR, RT-minus controls were prepared in a similar reaction mixture in which SuperScript III was replaced by water.

Primer design and qPCR Primers were designed with Primer Premier 5 software and synthesized by BioLegio or Eurogentec (Table 2A and B). The optimal melting temperature (58°C) of each primer set was experimentally verified with genomic DNA from S. fumaroxidans, M. hungatei, or M. formicicum as template. RT-qPCR was performed with the MyiQ Single color Real- Time PCR detection system (Bio-Rad). Each 25 µL reaction mixture contained 5 µL 20x diluted cDNA, 12.5 µL of 2x QuantiTect SYBR Green PCR mix (Qiagen), and 0.3 µM of

40 CHAPTER 3 each primer. Thermocycling conditions were as follows: 95 °C for 10 min, 40 cycles of 95 °C for 15 s, 58 °C for 30 s and 72 °C for 30 s with fluorescence detection at the end of each extension step. Amplification was immediately followed by a melting program consisting of 95 °C for 1 min, 55 °C for 1 min and a stepwise temperature increase of 0.5 °C per 10 s with fluorescence detection at each temperature transition. Template-minus controls were negative for every primer set and RT-minus controls excluded disturbing amounts of residual genomic DNA. Melting curve peaks for each primer pair indicated that single amplicon sizes were generated.

TABLE 2. RT qPCR primers designed for cDNA of target gene transcripts in S. fumaroxidans (A) and M. hungatei (B). Specific genes are listed with corresponding locus tag (IMG-JGI-DOE 2009) and the predicted metal content and cell localization of encoding formate dehydrogenase and hydrogenases. Table B also includes primers targeting cDNA of 16S rRNA fromM. formicicum used for calculations of the ratio of 16S rRNA from M. formicicum and S. fumaroxidans. TABLE 2A Locus tag Metal Primer Gene Localization Primer sequence (5’-3’) (IMG) content name Sfum_2706, W and Se SF_fdh1-fw CGG CGT CCC GTG AGT T fdh-1 Cytoplasm Sfum_2705 * ** SF_fdh1-rv GGC AGG GTG CTC TAC CAG TAT Sfum_1274, W and Se SF_fdh2-fw TGG TGC CAG CAT TCG GTG fdh-2 Periplasm Sfum_1273 * ** SF_fdh2-rv ATG TTC CCC AGG AGC AGC SF_fdh3-fw GGA CAT CTC GCG TTT GGA C fdh-3 Sfum_3509 Periplasm W or Mo SF_fdh3-rv CTC GCC TTG ACT TTC ACC TCT Sfum_0031, W or Mo SF_fdh4-fw CTA CCA TAC GCG GAC CCA G fdh-4 Periplasm Sfum_0030 * and Se SF_fdh4-rv CGA TTT CAC CCG GAC TTT G SF_fehyd-fw CCC GAG GAA TAC GAC GCT fehyd-1 Sfum_0844 Periplasm Fe SF_fehyd-rv TCA CGC CGC CAG AAG C SF_hyn-fw ACG TCG GCA AAG GCA ATA C hyn Sfum_2952 Cytoplasm Ni and Fe SF_hyn-rv GTC GTT CAG ACC CGC TCC SF_frh-fw GTT CGC GGT TGA CTT TGC frh Sfum_2221 Cytoplasm Ni and Fe SF_frh-rv CCT CAG CCT GCC ATC GTA GA SF_hox-fw GGT CTG CTG AGG GTG ATG G hox Sfum_2716 Cytoplasm Ni and Fe SF_hox-rv GGC GTC CGT GCC GTA T Ni, Fe and SF_hdr-fw GGC ACC ACC CAC AAC CTC hdr Sfum_3537 * Cytoplasm Se SF_hdr-rv CGC ACG GCC ATC TCC A Ni, Fe and SF_fnr-fw GCC CGT GGC AGC ATC A fnr Sfum_3954 * Cytoplasm Se SF_fnr-rv GCA GAA TCT TGT CTT CGG AGT G SF_rnfc-fw ACC GAG CAT CTG CCC ATA G rnfC Sfum_2699 Cytoplasm ---- SF_rnfc-rv GAA CCG AAG TGT TGA AGA AGG A Sfum_1795, Mo or W SF_fhlf-fw GCG GGT GCG GGT TCT A fhl-f Cytoplasm Sfum_1796 and Se SF_fhlf-rv CGG TTG AGT CAG ACG ATT GG SF_fhlh-fw AAG GTG GAG CCC AAA GCA fhl-h Sfum_1791 Cytoplasm Ni, Fe SF_fhlh-rv CGG TCC CAG GTT GTG AGT G NiFe-hyd- SF_mat-fw CGG TCT TTT CGG CTC ACA Sfum_4014 Cytoplasm ----- mat SF_mat-rv CGA TAG TTG TCC ACC ACT TCC T Sfum_R0013, SF_16S-fw ACG CTG TAA ACG ATG AGC ACT A 16SrRNA ------Sfum_R0015 SF_16S-rv GAT GTC AAG CCC AGG TAA GGT

* Selenocysteine residues predicted (Müller et al. 2010) **Metal content of the purified formate dehydrogenases as previously described (de Bok et al 2003)

41 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

TABLE 2B Metal Primer Gene Locus tag (IMG) Localization Primer sequence (5’-3’) content name MHfdh1-fw CGC AGC AGC AAT GGG ATA fdh-1 Mhun_1813 Cytoplasm W or Mo MHfdh1-rv CGT AGG ACG GGG TGA GTG A MHfdh2-fw CGG TGA ATG AAG GAA AAC TCT G fdh-2 Mhun_1833 Cytoplasm W or Mo MHfdh2-rv CGG GCT GTG GAT AAA CTG G MHfdh3-fw AAG AAG ACG GTT GAG AAC TAC G fdh-3 Mhun_2021 Cytoplasm W or Mo MHfdh3-rv GTT TCC AGT CAG AAG AGC AAG A MHfdh4-fw GTC ACC GAA CTG ACC ACC G fdh-4 Mhun_2023 Cytoplasm W or Mo MHfdh4-rv GCA CCC ATA TCA CAA GCA CC MHfdh5-fw ACC CTA TCA GCG TTC TCC G fdh-5 Mhun_3238 Cytoplasm W or Mo MHfdh5-rv AAA CTC GTG AGC ATA AGT CCC T MHech-fw CTT CCT GAG CCC ATT CAT CTT ech Mhun_1745 Periplasm Ni and Fe MHech-rv ATA GTC CTT GTA ATC ACG CTT TTC MHfrc-fw ATG CGG GAA TAA TTG AGC G frc Mhun_2332 Cytoplasm Ni and Fe MHfrc-rv ATA CCA CAA ACA CGG GAT GAG MHmbh-fw GCA TCC ACC TAC TCA AAC CAG mbh Mhun_2590 Periplasm Ni and Fe MHmbh-rv TGC CCG ACA ACA GCC AC Mhun_R0001, M. hun Mhun_R0072 MH16S-fw TCG TGC TGA CTG GAA TGT TAT ------16SrRNA Mhun_R0068, MH16S-rv CAG ACT CAT CCT GAA GCG AC Mhun_R0027

M. form MF16S-fw CTT CGG GGT CGT GGC GTA C ***NR_025028 ------16SrRNA MF16S-rv CAG ATT ACT GGC TTG GTG GG

*** Genbank number of the 16S rRNA gene from M. formicicum.

The optimal baseline was calculated by Bio-Rad MyiQ system software version 1.0

and generated threshold cycle (Ct) values. A second arbitrary baseline was manually defined and the amplification efficiency for each individual reaction was calculated from the kinetic curve (Liu and Saint 2002). Gene transcription values were normalized to 16S rRNA levels of the corresponding organism. At least three technical replicates were performed for each biological replicate. Outliers in technical replicates were determined with a Grubbs’ test (Burns et al. 2005).

Results Growth of S. fumaroxidans and M. hungatei on different substrates. Generation times of S. fumaroxidans cells grown in cocultures with M. hungatei (CoMh), in cocultures with M. formicicum (CoMf) and in pure cultures with propionate plus sulfate were higher (4-9 days) compared to other growth conditions tested (1.3-3.7 days; Table 1). All cultures with S. fumaroxidans yielded sufficient RNA to determine transcription levels of genes of interest (Fig. 1), as well as cultures withM. hungatei cells

grown on H2 plus carbon dioxide. M. hungatei cultures on formate contained enough RNA to determine most of the gene transcription levels (Fig. 2). The M. hungatei 16S rRNA / S. fumaroxidans 16S rRNA ratio in CoMh1 was 10 fold higher than in CoMh2 and -3, whereas the generation times of the three cocultures were comparable (Table 1). M. hungatei cells in CoMh1 contained gene transcription levels (relative toM. hungatei 16S rRNA) that were about 10 fold lower than in CoMh2 and -3 (Fig. 2).

42 CHAPTER 3

-2 -2 F D H 1 FeHyd A -3 H -3

-4 -4 -5 -5 -6 FF F HF CoMf CoMh PF PS -6 FF F HF CoMf CoMh PF PS -2 F D H 2 -2 B -3 I -3 Hyn -4 -4 -5 -5 -6 -6 FF F HF CoMf CoMh PF PS FF F HF CoMf CoMh PF PS -2 -2 F D H 3 Frh C -3 J -3 -4 -4 -5 -5 -6 -6 FF F HF CoMf CoMh PF PS FF F HF CoMf CoMh PF PS -2 -2 F D H 4 Hox D -3 K -3 -4 -4 -5 -5 -6 -6 FF F HF CoMf CoMh PF PS FF F HF CoMf CoMh PF PS -2 -2 RnfC Hdr E -3 L -3 -4 -4 -5 -5 -6 -6 FF F HF CoMf CoMh PF PS FF F HF CoMf CoMh PF PS -2 -2 FHL-F Fnr F -3 M -3 -4 -4 -5 -5 -6 -6 FF F HF CoMf CoMh PF PS FF F HF CoMf CoMh PF PS -2 -2 FHL-H NiFe-Hyd-mat G -3 N -3 -4 -4 -5 -5 -6 -6 FF F HF CoMf CoMh PF PS FF F HF CoMf CoMh PF PS

Figure 1 Transcription ofS. fumaroxidans genes coding for formate dehydrogenases (A-D), hydrogenases (H-M), RnfC (E), a [NiFe]-hydrogenase maturation protein (N), and formate dehydrogenase (F), hydrogenase (G) domains of a formate hydrogen lyase. The 10Log relative transcription is displayed on the y-axis and seven growth conditions on the x-axis; formate and fumarate (FF), fumarate only (F), hydrogen and fumarate (HF), S. fumaroxidans coculture with M. formicicum (CoMf), S. fumaroxidans coculture with M. hungatei (CoMh), propionate and fumarate (PF), propionate and sulfate (PS). The black, dark gray and light gray bars represent the three replicate cultures (1, 2 and 3 respectively) for each growth condition.

43 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

-1 FDH 1 -1 FDH 5 A -3 E -3

-5 -5

-7 -7 CoMh For H2 CoMh For H2 -1 -1 FDH 2 Ech B -3 F -3

-5 -5

-7 -7 CoMh For H2 CoMh For H2 -1 -1 FDH 3 Frc C -3 G -3

-5 -5

-7 -7 CoMh For H2 CoMh For H2 -1 -1 FDH 4 Mbh D -3 H -3

-5 -5

-7 -7 CoMh For H2 CoMh For H2

Figure 2 Transcription ofM . hungatei genes coding for formate dehydrogenases (A-E) and hydrogenases (F-H). The 10Log relative transcription is displayed on the y-ax and three growth conditions on the x-ax;

S. fumaroxidans coculture with M. hungatei (CoMh), formate (F), and hydrogen (H2). The black, dark gray and light gray bars represent the three replicate cultures (1, 2 and 3 respectively) for each growth condition.

Transcription of fdh’s and hyd’s in S. fumaroxidans. To examine the influence of growth phase on the gene transcription profile, S. fumaroxidans cells were grown on fumarate plus formate and on fumarate plus hydrogen, and harvested in the lag-, log- and stationary phase of growth. Transcription profiles of hydrogenase and formate dehydrogenase coding genes were similar in all three growth phases (data not shown). Therefore, all other measurements were done with cells in the logarithmic growth phase. The gene coding for the catalytic subunit of the periplasmic FDH2 ofS. fumaroxidans is transcribed in high levels in all tested conditions and transcription is even more up- regulated in pure cultures with propionate plus sulfate, in cells that fermented fumarate

44 CHAPTER 3 and in cells grown syntrophically (Fig.1B). Furthermore, transcription of the gene coding for the active subunit of the cytoplasmic FeHyd is up-regulated in S. fumaroxidans cells degrading propionate with fumarate or sulfate as electron acceptor and in cells degrading propionate in syntrophic association with methanogens (Fig. 1H). Transcription of the gene coding for the cytoplasmic FDH1 is high in all conditions and is even more up-regulated in cells grown syntrophically (Fig. 1A). Transcription of S. fumaroxidans genes coding for the active sites of the FHL-F and -H is higher in cocultures than in pure cultures (Fig. 1F and G). Genes coding for the Rnf-complex (Fig. 1E), the periplasmic Hyn (Fig. 1I), the cytoplasmic Hox (Fig. 1K), FDH3 (Fig. 1C) and FDH4 (Fig.1 D), were up-regulated in CoMh1 whereas transcription of these genes was >10 times lower in CoMh2 and -3, with the exception that FDH4 was also induced in CoMh2. Furthermore, transcription of the gene coding for the main subunit of the periplasmic Hyn is down-regulated in most cocultures (Fig. 1I). The gene coding for the cytoplasmic Frh is up-regulated during slow growth; cells grown in pure culture with propionate plus sulfate and cells grown on propionate in coculture with a methanogenic partner (Fig. 1J). Transcription of genes coding for the cytoplasmic active sites of FDH3 and -4 was low in most conditions tested (Fig. 1B and C). Transcription levels of the [NiFe]-hydrogenase maturation protein were similar in all growth condition tested (Fig. 1N).

Transcription of fdh’s and hyd’s in M. hungatei.

Between M. hungatei cells grown with H2 and carbon dioxide and those grown with formate, no significant differences in transcription of hydrogenase or formate dehydrogenase coding genes were observed (Fig. 2). Among all fdh’s and hyd’s in M. hungatei that contained cofactor F420 binding motifs in their amino acid sequences

(FDH1, -3, -4 and -5 and Frc), the gene coding for the active site of the cytoplasmic F420 reducing Frc is the highest up-regulated one (Fig. 2G). The highest up-regulated FDH with a F420 binding motif, is FDH1 (Fig. 2A). Furthermore, in all tested conditions the gene coding for the main subunit of Ech is higher transcribed than the other putative ion-translocating Mbh (Fig. 2F, H). In the presence of formate, genes coding for Mbh are slightly higher transcribed than in the presence of hydrogen.

Discussion Transcription of fdh’s and hyd’s in S. fumaroxidans cells grown in syntrophy with M. hungatei. When S. fumaroxidans degrades propionate, NADH and reduced ferredoxin are generated during malate- and pyruvate oxidation, respectively (Chabrière et al. 1999; van Kuijk and Stams, 1996) (Fig. 3). Genome analysis indicated that the cytoplasmic FeHyd, FDH1 and Hox are able to confurcate electrons from NADH and reduced ferredoxin to carbon dioxide or protons and generate formate or hydrogen, respectively (Müller et al. 2010). Due to the contribution of ferredoxin, NADH oxidation can be coupled to hydrogen or formate production (Müller et al. 2010). Our results confirm that FeHyd is an important confurcating enzyme; transcription of the gene coding for FeHyd is up-regulated when propionate is degraded. The cytoplasmic formate producing FDH1 was especially induced

45 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

propionate CO2

ATP FHL H2 formate Ech Fdred

succinate F420,red FRD fumarate MK H+ FDH2 CHO-MFR malate formate FeHyd H 2 Frc NADH Hyn FDH1 CH≡H4MPT oxaloacetate FDH1 Fd Hox H2 red - formate FDH5 CH2=H4MPT H F420,red 2 FDH4 Fdred Frh formate- pyruvate FDH3 Hdr CH -H MPT FDH2 3 4 F420,red Fnr CO2+ + H 2e- ? FDH3

acetyl-CoA CH3-S-CoM FDH4 Mbh HS-CoM +HS-CoB NADH + H H+ CoM-S-S-CoB Rnf MPH2 Fdred

acetate S. fumaroxidans M. hungatei CH4

Figure 3 Formate dehydrogenases and hydrogenases in S. fumaroxidans (left) and M. hungatei (right) are divided into functional groups. Transcription levels of genes coding for each target enzyme in the three cocultures of S. fumaroxidans and M. hungatei (CoMh1 in red, -2 in dark gray, -3 in light gray) are depicted in graphs with at the y-ax the 10Log relative transcription levels. In CoMh1, in which S. fumaroxidans retrieves less energy than compared to CoMh2 and -3, the Rnf-cluster and several ferredoxin dependent formate dehydrogenases and hydrogenases are induced as a back-up mechanism to re-oxidize NADH and ferredoxin efficiently. during syntrophic growth (Fig. 1A). The produced formate is transported through the membrane via a formate transporter, whose corresponding gene (Sfum_2707) is part of the FDH1 gene cluster (Müller et al. 2010) (Fig. 4). Sfum_2707 shows sequence similarity with the gene coding for FocA that co-transports protons with the negatively charged formate in E. coli (Suppmann and Sawers, 1994). The excreted formate is used by the methanogenic partner, which explains the importance of FDH1 and its formate transporter during syntrophic growth. It was estimated, by activity measurements in soluble fractions that FDH1 contributes 75% to the total formate dehydrogenase activity in S. fumaroxidans in pure and in cocultures (de Bok et al. 2003). Thus, during syntrophic growth, electrons from NADH and reduced ferredoxin are mainly confurcated via FeHyd and FDH1 to protons and carbon dioxide, respectively. In most cocultures the confurcating Hox was less important (Fig. 1K). The cytoplasmically produced hydrogen and formate can be interconverted by the formate-hydrogen lyase according to the requirements of the methanogen. Likely, therefore transcription of fhl-f and fhl-h is most pronounced during syntrophic growth (Fig. 1F and 1G).

46 CHAPTER 3

H2 Hyd 2 formate Se W FDH 2 + Ni Fe + [4Fe4S] H ? 2H a 2H+ a 2952 + CO 1273/4 2 2[4Fe4S] 2formate Rnf b2953 b1275

MK Formate Heme Xred Xox 2e- 2e- transporter D 2698 1998 2707 A /E 2696 A /E 2695 MKH2 2[4Fe4S] 4[4Fe4S] + + 2H 2H Se [2Fe2S] 2H2 g [2Fe2S] 2[4Fe4S] 2[Fe2S] 2formate 2Fe2S 0846 WW g 2703 FeFe B 2694 G 2697 C 2699 2000 FAD 4H+ + 2CO 4[4Fe4S] 3[4Fe4S]4Fe4S 4H+ 3[4Fe4S] 2 4Fe4S [4Fe4S] 1999 [2Fe2S] [2Fe2S]4Fe4S [2Fe2S] + FRD 2Fe2S 0845 2Fdred 2Fdox NAD NADH a2705/6 b2704 a 0844 b + H+ FDH 1 FeHyd

+ + 2Fdox 2Fdred NAD NADH 2Fdox 2Fdred NAD NADH + H+ + H+

ADP ATP + 2H NiFe b 2715 NAD 2 a 2716 propionate succinate fumarate malate NADH 4H+ g 2714 + + H 3[4Fe4S] oxaloacetate [2Fe2S] [4Fe4S] Hox 2714 2713 2Fdred 2Fdox + 2Fdox 2Fdred NAD NADH acetate acetyl-CoA pyruvate + H+

Figure 4 Methylmalonyl-CoA pathway in S. fumaroxidans and a schematic presentation of putative confurcating formate dehydrogenase (FDH1) and hydrogenases (Fehyd and Hox) and periplasmic formate dehydrogenase (FDH2) and hydrogenase (Hyn). Gene locus tag numbers and α-, β-, and γ-subunits are indicated in italic characters, predicted iron-sulfur clusters and metal-binding sites are indicated in bold.“Formate” represent formate- + H+.

The energetic most difficult step in the methylmalonyl-CoA pathway is the oxidation of succinate to fumarate (Stams and Plugge, 2009). It was previously calculated that 2H+ (which is an investment of ~⅔ ATP) have to be exported via a reversible ATP synthase to generate a proton motive force that allows the oxidation of succinate to fumarate (Schink, 1997; van Kuijk et al. 1998). Growth of S. fumaroxidans on fumarate with hydrogen or formate, yielded ~⅔ ATP and in cells grown with these substrates menaquinones were detected (van Kuijk et al. 1998). This is in analogy with the mechanism in Wolinella succinogenes in which a periplasmic formate dehydrogenase or hydrogenase (generating two periplasmic protons) transfers electrons viaa menaquinone loop to a cytoplasmic fumarate reductase, yielding ~⅔ ATP for growth (Kröger et al. 2002). A similar mechanism is probably involved in reversed direction when the energy dependent succinate oxidation to fumarate is performed (Müller et al. 2010, van Kuijk et al. 1998). The genome of S. fumaroxidans indicates two periplasmic candidates; FDH2 and Hyn that use periplasmic protons, (and subsequently decreasing the proton motive force) for formate or hydrogen production. Our results show that FDH2 is the dominant periplasmic proton user when cells are grown in syntrophic association with methanogens. As such, more formate than hydrogen is produced and consequently transferred to the partner methanogen (Fig. 3, 1B and 1I).

47 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

Further, proteins required for posttranscriptional maturation of [NiFe]-hydrogenases were found to be important in all growth conditions tested which corresponds with the observation that genes coding for [NiFe]-hydrogenases were constitutively transcribed.

Transcription of fdh’s and hyd’s in M. hungatei grown in coculture with S. fumaroxidans. At low hydrogen concentration, the first step of the methanogenic pathway is energy dependent and a membrane potential is used to provide enough reduced ferredoxin to allow reduction of carbon dioxide to formylmethanofuran (Thauer et al. 2008). Membrane integrated energy converting hydrogenases (Ech’s) import protons, cytoplasmically oxidize hydrogen and reduce ferredoxin (Thauer et al. 2008). According to Thauer et al. (2008), Ech’s of Methanomicrobiales contain at least sixteen subunits whereas the Ech of M. barkeri contains only six subunits. In contrast, our analysis of the genome of M. hungatei, that belongs to the Methanomicrobiales, indicates the presence of a membrane bound six subunit Ech (Mhun_1741-46) (Fig. 1 and 3). During syntrophic growth ech was transcribed in low levels to keep interspecies hydrogen concentrations as high as possible. In case formate is the sole electron donor, no hydrogen is available to cytoplasmically oxidize ferredoxin by Ech and the first step in the methanogenesis requires other

enzymes. It was previously shown that in M. hungatei this CO2 reduction is coupled to ferredoxin oxidation (Schwörer and Thauer, 1990). However, its genome indicates the

presence of formate dehydrogenases that reduce cofactor F420 only. When formate is the electron donor, the first step of methanogenesis might thus be coupled to cofactor

F420 oxidation as was reported forMethanococcus maripaludis (Hendrickson et al. 2008) (Fig. 3). 5 10 Candidates for the oxidation of cofactor 420F during N ,N -methyl-H4 MPT reduction 5 10 and N ,N ,-methylene-H4MPT reduction are the cytoplasmic FDH1 (Mhun_1813-14), -3 (Mhun_2020-21), -4 (Mhun_2022-23), -5 (Mhun_3237-38) and Frc (Mhun_2329- 32). Frc is most important because it is transcriptionally up-regulated in all conditions tested. It is partly membrane bound (Choquet and Sprott, 1991) which allows rapid

hydrogen consumption. Thereafter, FDH1 is the preferred420 F -reducing enzyme. FDH1 is co-transcribed with a formate transporter whose corresponding gene (Mhun_ 1811) is located in the FDH1 gene cluster (Fig. 5) and likely therefore is an important enzyme as well. The formate transporter allows fast import of formate and thus more effective

formate oxidation and cofactor F420 reduction. In M. hungatei a methylviologen reducing hydrogenase ADG-HdrABC is involved in

H2 oxidation coupled to the heterodisulfide reduction, the last step of methanogenesis in Methanomicrobiales such as M. hungatei (Liu and Saint, 2002). However, the genome of M. hungatei does not indicate the presence of such an enzyme complex. It also does not contain genes with similarity to a membrane-bound methanophenazine reducing [NiFe]-hydrogenase, which is involved in the build-up of a membrane potential via hydrogen oxidation in M. barkeri (Thauer et al. 2008). Instead, M. hungatei contains genes coding for a membrane bound hydrogenase (Mbh) (Mhun_2579-92) that is likely involved in this last step of the methanogensis pathway (Fig. 3).

48 CHAPTER 3

formate [4Fe4S] + + + + + [4Fe4S] [4Fe4S] formate Na /H Na /H 2H + CO2 a1833 b 1823

FDH 2 Formate ?ox ?red transporter I 2587 B 2580 A 2579 D 2582 H 2586 G 2585 F 2584 E 2583 C 2581 1811 B 1742 M 2591 A 1741

+ C 1743 D 1744 J 2588 K 2589 H2 2H N 2592 E 1745 F 1746 L 2590 formate CO2 F420,ox F420, red NiFe 2331 NiFe [4Fe4S] NiFe [4Fe4S] d +2H+ [4Fe4S] Ech [4Fe4S] Mbh a2332 [4Fe4S] [4Fe4S] [4Fe4S] [4Fe4S] b 2329 g 2330 [4Fe4S] a1813 1814 + + Frh FDH 1 b 2H H2 2H H2 Fdred Fdox+H2 ?red ? ox + H2 F420,ox F420, red

CO2 CHO-MFR CH≡H4MPT F420, red formate CO2 F420,ox F420, red + F420,ox +2H [4Fe4S] CH2=H4MPT [4Fe4S] [4Fe4S] a2021 b 2020 F420, red FDH 3 F formate F F 420,ox CO2 420,ox 420, red + CH4 CH3-S-CoM CH3-H4MPT +2H [4Fe4S] [4Fe4S] [4Fe4S] FDH 4 a 2023 b 2022 CoM-S-S-CoB HS-CoM + HS-CoB formate F F CO2 420,ox 420, red +2H+ [4Fe4S] Mbh [4Fe4S] [4Fe4S] FDH 5 a3238 b 3237

Figure 5 Methanogenesis in M. hungatei and a schematic presentation of putative energy converting hydrogenase (Ech), a membrane bound hydrogenase (Mbh) and F420-reducing formate dehydrogenases (FDH1, FDH3 and FDH4) and hydrogenase (Frh). Gene locus tag numbers and α-, β-, and γ-subunits are indicated in italic characters, predicted iron-sulfur clusters and metal-binding sites are indicated in bold.“Formate” represent formate- + H+.

Metabolic flexibility in cocultures of S. fumaroxidans and M. hungatei. Based on the highest formate and hydrogen concentrations maintained by S. fumaroxidans and the lowest possible formate and hydrogen concentrations maintained by M. hungatei, the formate and hydrogen concentrations in cocultures range from 15 – 24 µM and 2 – 7 Pa respectively (Dong and Stams, 1995; schauer et al. 1982; Seitz et al. 1988). Syntrophic partners regulate their metabolism to grow together, however the methanogens are greatly enhanced if hydrogen and formate concentrations are high. Such metabolic flexibility is suggested by the observation thatM. hungatei cells in CoMh1 contained approximately 10 fold more ribosomal activity thanM. hungatei cells in CoMh2 and -3 (Table 1) for reasons that remain unclear. However, it explains why gene transcription values relative to M. hungatei 16S rRNA were approximately 10 fold lower in CoMh1. As a consequence, in CoMh1 S. fumaroxidans retrieved less energy for growth and it induced transcription of genes coding for the Rnf-complex, formate dehydrogenases and hydrogenases. With the assumption that in CoMh1 the formate and hydrogen concentrations are high, FDH1 and FeHyd will reduce less carbon dioxide and protons and thus re-oxidize less NADH and ferredoxin. This will result in the transcription of the genes coding for the third confurcating enzyme; Hox (Fig. 3 and 1K). Moreover, high NADH may lead to the

49 Growth and substrate dependent transcription of fdh’s and hyd’s in S. fumaroxidans and M. hungatei

induced transcription of genes coding for the Rnf-complex that provides the possibility to use the membrane potential for NADH oxidation coupled to ferredoxin reduction. To re-oxidize the ferredoxin the cytoplasmic formate dehydrogenases (FDH3 and 4) and hydrogenases (Frh, Hdr, and Frh) are induced in CoMh1 (Fig. 3). The genome of S. fumaroxidans indicates the presence of the periplasmic FDH2 and Hyn that are likely involved in the endergonic succinate reduction by using periplasmic protons for formate or hydrogen production, respectively. Our results show that in general FDH2 is the dominant periplasmic terminal reductase in the cells grown in syntrophic association with methanogens. However in CoMh1, when formate and hydrogen concentrations are supposed to be high, S. fumaroxidans will obtain less energy and it results in a lower expression of the periplasmic Hyn (Fig. 3 and 1I). Bacteria adapt to environmental conditions commonly by regulation at the level of transcription (Santos et al. 2008). The genome of S. fumaroxidans contains many response regulators (IMG-JGI-DOE, 2009). In general, two component systems are involved in sensing changes in the external environment. Genes coding for putative regulatory proteins were also found in operons coding for formate dehydrogenases or hydrogenases. The gene Sfum_3508 that is located adjacently to the gene cluster coding for FDH3 contains a DNA-binding motif and shows similarity with the gene coding for TenI. In Bacillus subtilis, this gene regulates the production of extracellular proteases (Toms et al. 2005). The gene Sfum_0033 that is located adjacently to the gene cluster coding for FDH4 contains similarity with the gene coding for a transcriptional regulator, LysR that controls the expression of genes required for carbon dioxide fixation in Xanthobacter flavus (Van Keulen et al. 2003). Furthermore, the genes Sfum_3531-34 and Sfum_3958-61 are located adjacently to the gene clusters coding for Hdr and Fnr, respectively, and comprise a DNA binding subunit with similarity to a mercury regulatory protein, a cysteine-rich subunit and a subunit with NAD+/FAD binding capacity and in case of Sfum_3533 and 34 a selenocysteine. These subunits are most likely used to sense the redox potential and to regulate gene transcription inS. fumaroxidans, as was shown for a redox sensor in E. coli (Bagramyan and Trchounian, 2003). Paralogs of genes coding for these subunits (except for MerR) were found in the formate hydrogen lyase coding operon, which indicates a regulatory function on the FHL as well (Sfum_1801 and Sfum_1794). Furthermore, a regulatory protein binding site is possibly present in the intergenic region between the FDH1 coding gene cluster and the rnf-cluster, which are oriented in reversed direction (IMG-JGI-DOE, 2009). This suggests co-regulation by a bi-directional regulator. Probably more than one regulator is involved, because no correlation of the transcription of the FDH1 operon and the rnf-cluster was observed.

Transcription of fdh’s and hyd’s in S. fumaroxidans grown syntrophically with M. formicicum. S. fumaroxidans grows in suspended cocultures with M. hungatei, whereas it forms granules in syntrophic growth with M. formicicum. It was previously calculated that formate is the preferred interspecies electron carrier in suspended cocultures and that hydrogen might be preferred when interspecies distances are smaller in granules (Stams and Plugge, 2009). However, no increased transcription of genes coding for

50 CHAPTER 3 hydrogenases was observed in S. fumaroxidans cells grown with M. formicicum. Neither increased transcription of genes coding for formate dehydrogenases was observed in S. fumaroxidans cells grown in syntrophic association with M. hungatei. Nonetheless it could very well be that the same formate dehydrogenase and hydrogenase complexes allow different electron fluxes and that the presence of certain formate dehydrogenases or hydrogenases only show the possibilities of electron flows.

Transcription of fdh’s and hyd’s in pure cultures of S. fumaroxidans.

When S. fumaroxidans uses H2 or formate as electron donor and fumarate as electron acceptor, Hyn and FDH2 catalyze periplasmic H2 and formate oxidation, respectively (Fig. 1B and I). In addition, the FHL cytoplasmically interconverts formate and hydrogen that are transported out and into the periplasm via the Foc or by diffusion. This explains why

Hyn is required during growth on formate and FDH2 is required during growth on H2 (Fig. 1B and I). Furthermore, Hyn was down-regulated during syntrophic growth and since it is the only periplasmic hydrogenase, it might be the periplasmic component involved in hydrogen cycling. During syntrophic growth, the periplasmically formed hydrogen would be transferred to the methanogen whereas during non-syntrophic growth an energy conserving hydrogen cycle could exist as was described for Desulfovibrio sp. (Heidelberg et al. 2004; Odom and Peck, 1981a).

Metabolic flexibility in other syntrophic bacteria. In analogy with S. fumaroxidans the genome of the syntrophic fatty acid- and aromatic acid-degrading Syntrophus aciditrophicus contains several fdh’s, hyd’s and an rnf- cluster (McInerney et al. 2007). The genome of the syntrophic propionate degrading Pelotomaculum thermopropionicum contains several fdh’s and hyd’s that were also transcribed in all growth conditions tested (Kato et al. 2009). This illustrates the numerous possibilities for interspecies electron transfer in other syntrophs. The specific environmental adaptation of syntrophic communities needs further exploitation to assess the level of metabolic flexibility of each organism in the community.

Acknowledgements The research was supported by the Research Councils for Earth and Life Sciences (ALW) and Chemical Sciences (CW) with financial aid from the Netherlands Organization for Scientific Research (NWO). We gratefully thank Dr. Douwe van der Veen for technical advice.

51 52 CHAPTER 4

Formate dehydrogenase and hydrogenase encoding gene clusters in Syntrophobacter fumaroxidans and Methanospirillum hungatei.

Petra Worm, Alona Rusakov, Remco Kort, Frank Schuren, Alfons J.M. Stams and Caroline M. Plugge.

Abstract To get insight into the genes required for hydrogen and formate transfer in syntrophic cocultures of Syntrophobacter fumaroxidans and Methanospirillum hungatei, formate dehydrogenase and hydrogenase complexes were predicted from gene clusters present in the genomes of both organisms (Chapter 2). Quantitative PCR primers were designed to target transcripts of genes coding for alpha subunits of formate dehydrogenases and hydrogenases in both organisms (Chapter 3). In this chapter a microarray approach was used to determine co-transcription of genes and to further investigate the predicted gene clusters. Microarray data supported the classification in the functional groups of most formate dehydrogenase-, hydrogenase-, and formate hydrogen lyase-complexes as were proposed in Chapter 3 (Fig. 1). Exceptions are the hydrogenase “Frh” from S. fumaroxidans and formate dehydrogenase 3 from M. hungatei. Frh and FDH3 might be involved in maturation processes of hydrogenases and formate dehydrogenases.

53 Formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei

Introduction The regulation of hydrogen and formate transfer in syntrophic cocultures of Syntrophobacter fumaroxidans and Methanospirillum hungatei were studied. Formate dehydrogenase and hydrogenase coding gene clusters were found in both organisms (Chapter 2 and 3). In addition, the genome of S. fumaroxidans contains an rnf-cluster that could code for the different subunits of an Rnf-complex, and a gene cluster that could code for a formate hydrogen lyase (Chapter 2 and 3). Alpha subunits of formate dehydrogenases and hydrogenases contain the catalytic core of the enzyme and in some cases iron-sulfur clusters. Other subunits were

predicted to contain iron-sulfur clusters, bind NADH or FADH2, translocate protons or integrate into the cytoplasmic membrane. Specific qPCR primers successfully targeted the transcripts of genes coding for the main subunits of formate dehydrogenases and hydrogenases, and indicated that some genes are highly transcribed at all growth conditions and others only at conditions in which less energy is available (Chapter 3). Biochemical characterization showed that the native composition of the cytoplasmic

FDH I, the periplasmic FDH2 and the periplasmic Hyn from S. fumaroxidans are α2β2γ2,

α1β1, and A1B1, respectively (Table 1) (de Bok et al. 2003; de Bok 2002). The F420-reducing hydrogenase of M. hungatei has been purified and appeared to be composed of three subunits α, β, and γ (Choquet and Sprott 1991). To predict co-factor binding, electron flows and cell-localization of the other formate dehydrogenase-, and hydrogenase complexes in S. fumaroxidans and M. hungatei and the Rnf-complex in S. fumaroxidans, we made use of bio-informatic gene and ortholog neighborhood comparisons (IMG- JGI-DOE 2009).

Table 1. Subunit composition of formate dehydrogenases, hydrogenases, and Rnf-complex in S. fumaroxidans and formate dehydrogenases and hydrogenases in M. hungatei, based on previous biochemical experiments (de Bok et al. 2003; de Bok et al. thesis; Choquet and Sprott, 1991), ortholog gene comparison (Müller et al. 2009), and microarray data. Not determined with microarray (ND). S. fumaroxidans M. hungatei Enzyme Subunits Enzyme Subunits

FDH1 α2β2γ2 FDH1 αβ Rnf-complex ABCDEG FDH2 αβ

FDH2 α1β1 FDH3 α FDH3p αβ FDH4 αβ FDH3c α FDH5 αβ FDH4p αβ (NDa) Ech ABCDEF FDH4c α (NDa) Frh αβγ

FeHyd α2β2γ2 Mbh ABCDEFGHIJKLMN

Hyn α1β1 Frh A

Hox E2F2U2Y2H2 Hdr αβδ Fnr AG, Crd FHL-H αβγ, hdrA

FHL-F αβ, hdrA2, Crd2, mvhD ano data from Microarray (ND)

54 CHAPTER 4

Microarray approaches can be used to determine co-transcription of genes and to reveal operons (Xiao et al. 2006). In prokaryotes genes that code for different subunits of the same enzyme, usually are located in one operon. In some cases operons contain additional genes coding for maturation proteins or transcriptional regulators that are involved in post-translational maturation or transcriptional regulation of the corresponding genes. Redox enzymes such as formate dehydrogenases and hydrogenases contain metals and cofactors for catalysis, which require posttranslational maturation. Here, we used a microarray approach withS. fumaroxidans and M. hungatei grown under different conditions to get further support for formate dehydrogenase and hydrogenase operons in both organisms and the rnf-cluster in S. fumaroxidans. This provides information on the co-factor binding, electron flows and cell-localization of the uncharacterized formate dehydrogenase- and hydrogenase complexes in S. fumaroxidans and M. hungatei and the Rnf-complex in S. fumaroxidans. This information contributes to the understanding of the electron flow mechanism in both organisms.

Materials and methods Cultivation conditions and RNA isolation Syntrophobacter fumaroxidans MPOB (DSM 10017) was grown in pure culture in medium described previously (Chapter 3). S. fumaroxidans was cultivated in volumes ranging from 500 to 1500 ml, with the added substrates; 20 mM propionate plus 60 mM fumarate, and with 30 mM propionate plus 25 mM sulfate in pure cultures, and 30 mM propionate with Methanospirillum hungatei JF-1 (DSM 864) or Methanobacterium formicicum JF-1 (DSM 1535) in syntrophic cocultures. M. hungatei was grown axenically in the same medium with 1 mM acetate and H2/CO2 (80:20, vol/vol). Growth was followed by measuring optical density, and/or by the formation of sulfide or methane. Cells were harvested anaerobically in the logarithmic phase, or in case of M. hungatei cells grown on H2/CO2, during linear growth (Figure 1). RNA was isolated as described previously (Chapter 3).

Comparative gene analysis Automatic annotations of genomes from DOE-Joined Genome Institute were used to detect the presence of gene clusters coding for formate dehydrogenases, and hydrogenases in S. fumaroxidans and M. hungatei, and an Rnf complex, [NiFe]- hydrogenase maturation proteins and an formate hydrogen lyase in S. fumaroxidans. N-terminal amino acid sequences of FDH-1 and FDH-2 identified by de Boket al. (2003) were used to find corresponding fdh-1 and fdh-2 nucleotide sequences in the genome of S. fumaroxidans. Pfam search (Sanger institute, 2009) was used to identify motifs in the amino acid sequences and the TMHMM Server v. 2.0 (DTU, 2009) was used to identify transmembrane helices. With the TatP 1.0 Server twin-arginine translocation (Tat) motifs in the N-terminus were identified to predict protein localization in S. fumaroxidans cells (Bendtsen et al. 2005). With PRED-SIGNAL (Bagos et al. 2009) Tat- motifs were identified to predict the cell localization of proteins inM. hungatei cells. The incorporation of selenocysteine (SeCys) was examined by RNA loop predictions with Mfold version 3.2 (Mathews et al. 1999; Zuker, 2003). The RNA loop predicted in

55 Formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei

A0.8 B6 0.6 4

0.4

2 optical density 0.2 n methane (mM)

0 0 0 2 4 6 0 5 10 15 20 days days C6 D6 3 4 4 M x10 m

2 2 sulfide n Methane (mM)

0 0 0 10 20 30 0 20 40 days days E4 3

2

n methane (mM) 1

0 0 5 10 days Figure 1 Growth curves of S. fumaroxidans grown on propionate plus fumarate (A), in coculture with M. hungatei on propionate (B), on propionate plus sulfate (C) in coculture with M. formicicum on

propionate and M. hungatei grown on H2 /CO2 (E). the 50-100 bp region downstream of the UGA-codon was compared with the consensus loop described by Zhang and Gladyshev (2005). Complete amino acid sequences of putative selenocysteine-enzymes were aligned to their homologues with ClustalX 1.81 (Kryukov and Gladyshev, 2004). SeCys incorporation was confirmed when the amino acid sequence aligned with a conventional cysteine residue in homologous proteins.

S. fumaroxidans and M. hungatei microarrays The S. fumaroxidans and M. hungatei microarrays were constructed by Roche NimbleGen. Microarray slides contained 4 chambers with 72,000 probes each. An oligonucleotide library of 60-mers covered 4064 and 3139 open reading frames of the S. fumaroxidans (Genbank nr. CP000478) and M. hungatei genomes (GenBank nr. CP000254), respectively. Each open reading frame was covered by 9 different oligonucleotides and was present on the microarray in duplicate.

Labeling of cDNA and hybridization cDNA was synthesized as described in Chapter 3 and labeled as described by Ávila-Pérez et al. (2010). Probes were hybridized to the oligo´s on the array with the NimbleGen

56 CHAPTER 4 hybridization system, image analysis and data processing was performed as described previously (Ávila-Pérez et al. 2010).

Results and discussion Formate dehydrogenase and hydrogenase coding gene clusters in the genomes of S. fumaroxidans and M. hungatei were analyzed. Cell localization and cofactor binding sites were predicted and genes coding for putative transcriptional regulators and hydrogenase and formate dehydrogenase maturation proteins were found (Figure 2).

Formate dehydrogenase coding gene clusters in S. fumaroxidans

Biochemical characterization showed that the FDH1 has a α2β2γ2 structure (de Bok et al. 2003). Genes coding for the alpha, beta and gamma subunits of FDH1 (Sfum_2703- 06) were co-transcribed (Figure 3A). Additionally, the gene coding for the bidirectional formate transporter (foc) (Suppmann and Sawers 1994) (Sfum_2707) is co-transcribed. FDH1 is the most abundant formate dehydrogenase in all conditions and it has high activity in the direction of CO2 reduction (de Bok et al. 2003; Chapter 3). Fast production of formate requires active formate export to prevent intracellular formate accumulation, which explains co-regulation of foc (encoding a formate transporter) with fdhα, -β, and -γ. Upstream of the FDH1 operon an rnf-cluster is located in reversed direction. Co- transcription of rnf-genes A-E and G could not be detected because hybridization duplicates (SH1-SH2) showed a large variation of log ratios of -8 to 0 (Figure 3A). In the intergenic region of the rnf-cluster and the FDH1 operon, palindrome sequences were identified (Figure 2A). This indicates transcriptional regulator binding sites and suggests co-regulation of thernf -cluster and the FDH1 operon. RT qPCR data (Chapter 3) did not show any correlation of transcription of the rnfC and fdhα, suggesting that more than one regulator is involved in transcriptional regulation. Genes coding for the alpha and beta subunit of the periplasmic FDH2 (Sfum_1273- 75) are co-transcribed (Figure 3B) as was expected from biochemical characterization that showed a hetero-dimeric structure (Table 1) (de Bok et al. 2003). FDH3 and FDH4 were not characterized biochemically and their gene clusters both are composed of a cytoplasmic alpha (fdhαc) subunit and a periplasmic alpha (fdhαp) and beta subunit (fdhβ) (Figure 2A). RT qPCR experiments revealed that transcription of genes coding for the cytoplasmic alpha subunits are up-regulated when less energy is available for S. fumaroxidans. Microarray data show that the fdhαc (Sfum_3509), was not co-transcribed with fdhαp (Sfum_3510) and fdhβ (Sfum_3511) of FDH3 (Figure 3C). This suggests that the periplasmic alpha and beta subunits form a hetero-oligomer, and the cytoplasmic FDH3c is differentially expressed dependent on the growth conditions. Up-stream of FDH3c, the genes Sfum_3508 and Sfum_3507 are located that may code for transcriptional regulators. Sfum_3508 contains a DNA binding motif and shows similarity with the gene coding for a regulatory phosphatase TenI of Bacillus subtilis (Toms et al. 2005), and Sfum_3507 is annotated as a kinase. With the applied data filter no signal of the gene coding for the periplasmic alpha subunit of FDH4 (Sfum_0035) was obtained and the hybridization duplicates were

57 Formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei

A

Rnf F rnf C rnf D rnf G rnf A/E rnf A/E rnf B

TTTTTGTGAATTGATTCACTA AAG GATGAAGAGCCAACCTCTTCAGAATCGTGCACGATGCGG

FDH 1 Se fdhγ fdhβ fdhα foc

FDH 2 Se fdhα fdhβ fdhE FDH 3 fdhαc fdhαp fdhβ

FDH 4 Se Se fdhα fdhD fdhαp fdhβ fdhE

FHL-F Se Se Se hdrC crd hdrA mvh d crd hdrA hdrA mvhD fdhβ fdhα

B FDH 1 foc reg prot fdhα fdhβ FDH 2 fdhα fdhβ FDH 3 FDH 4 fdh4α fdh4β fdh3α fdh3β fdhD FDH 5 fdhα fdhβ

Figure 2 Formate dehydrogenase coding gene clusters in S. fumaroxidans (A) and M. hungatei (B) and hydrogenase coding gene clusters in S. fumaroxidans (C ) and M. hungatei (D). Conserved motifs for [2Fe2S]-cluster ( ), [4Fe4S]-cluster ( ), FAD ( F ) and NADH ( ) binding, and twin arginine translocation motif ( ), hydrophobic transmembrane domains ( ), and Selenocysteine coding TGA ( Se ) were also predicted. Abbreviations of gene annotations were used: cysteine rich domain coding gene (crd), formylmethanofuran dehydrogenase coding gene (fmd), hetero reductase (hdr), hypothetical (or Hyp coding genes in NiFe-hydrogenase maturation operon) (hyp), [FeFe]-hydrogenase maturation protein coding genes (hydE/G), putative transcriptional regulator coding genemod ( E), gene coding for a transcription factor that regulates Ni uptake (nikR), pyruvate-formate lyase activating enzyme A coding gene (pflA), and regulatory protein coding gene (reg prot).

58 CHAPTER 4

C

NiFe-Hyd maturation hypA hypB hypF hypD hypE pflA FeHyd fehydα-like fehydβ fehydd fehydβ fehydα hydG hydE ktpA/tpt1 Hyn hynB hynA hynC hynD reg prot

Frh F frhG frhA hyd mat Hox

hoxE hoxF hoxU hoxY hoxH

Hdr F Se Se merR crd hdrA mvhD mvhC mvhA

Fnr F Se Se merR crd hdrA mvhD fnrG fnrA hyp

FHL-H F Se hdrA mvhD mvhC mvhA D Ech echA echB echC echD echE echF Frh frhα mat frhγ frhβ Mbh A B D E F G H I J K L M N variable (Figure 3). Thus, complex formation of formate dehydrogenase alpha and beta subunits could not be predicted and the involvement of a putative regulator coding gene Sfum_0033 and a maturation protease FDHD could not be verified.

Hydrogenase coding gene clusters in S. fumaroxidans The genes Sfum_0844 - 46 of S. fumaroxidans show homology with the genes TM1424 - 26 from Thermotoga maritima that code for the subunits of the heterotrimeric confurcating [FeFe]-hydrogenase (Schut and Adams, 2009). Sfum_0844 - 46 were clearly co-transcribed (Figure 3E) and their gene products probably form a heterotrimeric confurcating [FeFe]-hydrogenase in S. fumaroxidans as well (Table 1). In T. maritima, HydE and HydG are involved in the maturation of the [FeFe]-hydrogenase (Rubach et al.

59 Formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei

A

B

C

D

E

F

G

60 CHAPTER 4

H

I

J

K

L

Figure 3 Expression profiles and operon prediction of gene clusters inS. fumaroxidans coding for FDH1 and Rnf-complex (A), FDH2 (B), FDH3 (C), FDH4 (D), [FeFe]-hyd (E), Hyn (F), Frh (G), Hox (H), Hdr (I), Fnr (J), FHL (K), and Hyp (L). The color coding ranges from blue (log ratio is -6) via black (log ratio is 0) to yellow (log ratio is 6). The blue color represents down-regulated transcription compared to cells grown in syntrophic association with M. hungatei, whereas the yellow color represents up-regulated transcription compared to cells grown in syntrophic association with M. hungatei. The grey color indicates no signal with the filters applied. The range of log ratios of the operons is shown in white. Open reading frames depicted in arrows and Gene ID’s were retrieved from (IMG-JGI-DOE 2009). The columns represent hybridization ratio’s of transcripts from S. fumaroxidans grown on propionate plus fumarate (PF), propionate plus sulfate (PS), on propionate in coculture with M. hungatei (SH) or with M. formicicum (SF).

61 Formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei

2005). Downstream of fehydα, -β, and -γ in the genome of S. fumaroxidans, hydE and G homologues are located which are co-transcribed with two hypothetical genes and a gene similar to a 2’-phoshotransferase (Tpt1) coding gene (Figure 3E). Tpt1 transfers the 2’-phosphate from ligated tRNA to NAD in yeast, but the function of its homologue (KptA) in Bacteria such as E. coli is unclear (Steiger et al. 2001). In S. fumaroxidans the products of this gene cluster are probably involved in maturation of the [FeFe]-hydrogenase. The genes Sfum_0847 and -48 are located upstream of fehydα, -β, and –γ, and show similarity with the [FeFe]-hydrogenase small subunit of D. desulfuricans (Nicolet et al. 1999) (fehydβ) and a [FeFe]-hydrogenase alpha subunit (fehydα-like), respectively. The gene fehydα-like lacks a motif coding for binding of an H-cluster. [FeFe]-hydrogenases require the H-cluster for their catalytic function (Schwab et al. 2006), thus fehydα-like probably does not code for a functional [FeFe]-hydrogenase. The genes fehydβ, and fehydα-like seem to be co-transcribed with a hypothetical gene (Sfum_0849) but the function of an [FeFe]-hydrogenase-like enzyme is not clear. As was expected from biochemical characterization of the hetero-dimeric periplasmic [NiFe]-hydrogenase “Hyn” (de Bok 2002) genes coding for the A and B subunit are co- transcribed (Figure 3F). Downstream of this operon, Sfum_2951 and Sfum_2950 are located and show sequence similarity with a hydrogenase expression / formation protein (HynC) and a hydrogenase assembly chaperone (HynD) of Desulfovibrio gigas, respectively (Casalot and Rousset 2001). The gene hynD was co-transcribed with Sfum_2949 that shows similarity with a regulatory protein. Thus HynD and –E could be transcriptionally regulated by the gene product of Sfum_2949, and probably are involved in maturation of the heterodimeric Hyn.

The genes Sfum_2221 and Sfum_2222 show similarity with cofactor F420-reducing hydrogenase (Frh) A and G subunits of the methanogen Methanopyrus kandleri (Coppi, 2005), respectively.Frh A and G homologues are also located adjacently in the genomes of other bacteria such as Dehalococcoides sp VS and Desulfohalobium retbaenese DSM5692 (IMG-JGI-DOE 2009). However, in S. fumaroxidans frhA and -G were differentially transcribed (Figure 3G). FrhA seems to be involved in [NiFe]-hydrogenase maturation, and FrhG possibly forms a complex with an iron-sulfur containing domain and an frd domain. The genes hoxE, -F, -U, -Y, and -H (2712-16) were predicted to code for a cytoplasmic confurcating [NiFe]-hydrogenase inS. fumaroxidans (Chapter 2 and 3). In cyanobacteria Hox is a heteropentamer (Tamagnini et al. 2007). This may also be the case for Hox in S. fumaroxidans (Table 1). In methanogens, a methyl viologen reducing hydrogenase (Mvh) donates electrons to the heterodisulfide reductase (Hdr) during growth on hydrogen and carbon dioxide (Stojanowic et al. 2003). Genes that show homology with those of Mvh and Hdr subunits are located adjacently in the genome of the methanogen Methanocaldococcus jannaschii, but also in genomes of the bacteria Geobacter sulfurreducens, Desulfovibrio desulfuricans, Chlorobium tepidum (Coppi 2005) and S. fumaroxidans (IMG-JGI-DOE 2009). In S. fumaroxidans mvhA, -C, and -D (Sfum_3535-37) are co-transcribed and thus likely form a trimer (Figure 3I). The adjacent gene hdrA seems to be co-regulated with a DNA binding regulatory protein (MerR) and a cysteine-rich protein (cdr), indicating that

62 CHAPTER 4

HdrA has a sensing or regulatory function in S. fumaroxidans.

The genes Sfum_3954-57 show similarity with the F420-non-reducing hydrogenase subunits (Fnr) A and G and MvhD in Methanothermobacter marburgensis (Stojanowic et al. 2003). In S. fumaroxidans, frhA, G and -D seem to be co-transcribed with the upstream locating genes coding for a hypothetical protein (Sfum_3953) and a putative signal transduction protein (Sfum_3952) and with the downstream located genes coding for a cysteine rich protein (Sfum_3960), a DNA binding regulator (MerR) (Sfum_3961) and a hypothetical protein (Sfum_3962) (Figure 2J). Sfum_3958 is annotated as HdrA, but was not detected in the microarray. Likely, FnrA and G form a hydrogenase complex with MvhD. HdrA, a cysteine rich protein, MerR and the hypothetical protein might be involved in transcriptional regulation. The genes coding for the FHL-H, mvhA, -C, -D, and hdrA (Sfum_1791-94) are located adjacently to those coding for a FHL-F (Sfum_1795-Sfum_1797) whose products likely form a complex that functions as a formate hydrogen lyase (Chapter 2). The FHL-H and FHL-F coding genes are differentially transcribed although differences were small (Figure 3K). The cysteine rich subunits, MvhD and Hdr subunits likely transfer electrons between the FHL-H and FHL-F. The genes coding for the [NiFe]-hydrogenase maturation proteins (hypA, B and F) were co-transcribed (Figure 3L). Genes coding for HypD and E were co-transcribed with Sfum_4016 that was annotated as a pyruvate formate lyase alpha subunit (Casalot et al. 2001). This indicates that this operon is involved in the maturation of [NiFe]- hydrogenases in S. fumaroxidans.

Formate dehydrogenase coding gene clusters in M. hungatei The genes fdhα, a (Mhun_1813-14) are co-transcribed (Figure 4A), indicating that they code for a functional formate dehydrogenase (FDH1). In addition, the gene coding for a bidirectional formate transporter (foc) (Mhun_1811) (Suppmann and Sawers 1994) is co-regulated with genes coding for FDH1. Previous RT qPCR experiments indicated that FDH1 is the highest up-regulated formate dehydrogenase in M. hungatei (Chapter 3). This suggests that cytoplasmic formate provided by Foc is used for formate oxidation by FDH1. The genes coding for the alpha and beta subunits of FDH2 (Mhun_1832-33) are clearly co-transcribed (Figure 4B). Downstream in the reversed direction genes are located that code for formylmethanofuran dehydrogenase (Mhun_1834-35) and heterodisulfide reductase (Mhun_1836-Mhun_1836) that are co-transcribed. Genes coding for FDH 3 and 4 and are located adjacently. The alpha and beta subunit of FDH4 (Mhun_2022-23) are clearly co-transcribed confirming the prediction that their gene products form the FDH4 complex (Figure 4C). Genes coding for the alpha and beta subunit of FDH3 (Mhun_2020-21) however, are not co-transcribed. Microarray data indicate that FDH3 beta is co-expressed with FDHD (Figure 4C), which is a putative FDH maturation protein (Schlindwein et al. 1990). Genes coding for the alpha and beta subunits of FDH5 (Mhun_3237-38) are clearly co-transcribed which suggests that they form heteromers (Table 1) (Figure. 4D).

63 Formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei

Figure 4 Expression profiles and operon prediction of gene clusters in M. hungatei coding for FDH1 (A), FDH2 (B), FDH3 and-4 (C), FDH5(E), Ech (F), Frh (G), and Mbh (H). Color coding ranges from blue (log ratio is -4) via black (log ratio is 0) to yellow (log ratio is 4); the grey color indicates no signal with the filters applied. The range of log ratio’s of the operons are shown in white characters in white squares. Open reading frames depicted in arrows and Gene ID’s were retrieved from (IMG-JGI-DOE 2009). The columns represent hybridization ratio’s of transcripts from M. hungatei grown on hydrogen plus carbon dioxide (H2), or on propionate in coculture with S. fumaroxidans (SH).

64 CHAPTER 4

Hydrogenase coding gene clusters in M. hungatei Genes coding for the subunits A-F of the energy converting hydrogenase (Ech) (Mhun_1741-46) were clearly co-transcribed (Figure 4E) and it is likely that these subunits form a functional complex. As was expected from purification and characterization of a cytoplasmic trimeric

F420-reducing hydrogenase in M. hungatei by Choquet et al. (1991), genes coding for the alpha, beta, and gamma subunits of the F420-reducing hydrogenase (Frh) (Mhun_2329- 32) were co-transcribed (Figure 4F). The trimeric Frh complex is likely formed by the hydrogenase maturation protein encoded by Mhun_2331. Genes coding for the subunits A-N of the membrane bound hydrogenase (Mbh) (Mhun_2579-92) are co-transcribed (Figure 4G), which shows that these subunits are present in a Mbh-complex.

Conclusion Microarray data further supported that in S. fumaroxidans the cytoplasmic confurcating enzymes are either trimeric (FDH1, FeFe-hyd) or pentameric (Hox) enzymes whereas the periplasmic formate dehydrogenases and hydrogenases, such as FDH2, Hyn and the periplasmic FDH3 are heteromers (Table 1). Furthermore, co-transcription showed that the cytoplasmic ferredoxin dependent hydrogenases Hdr and Fnr are both trimers. The catalytic subunit FrhA however does not form a dimer with FrhG, but might be involved in hydrogenase maturation. Furthermore, FHL-H and FLH-F are separate enzymes that may form a formate hydrogen lyase complex via iron-sulfur cluster binding subunits that are co-expressed with FHL-H and FHL-F. The [NiFe]-hydrogenase maturation coding genes are located in two different clusters. In M. hungatei formate dehydrogenases are composed of an alpha and beta subunit. The alpha subunit of FDH3 does not form a heteromer with a beta subunit and thus might function as a monomer. The trimeric conformation of Frh and complex forming of the membrane integrated Ech and Mbh was confirmed. Operon prediction with microarray data supported the predicted composition of formate dehydrogenases and hydrogenases in S. fumaroxidans and M. hungatei (Müller et al. 2009). The functional groups as shown in Fig. 1 of Chapter 3 were confirmed with the exceptions of Frh fromS. fumaroxidans and FDH3 from M. hungatei that were shown to be monomers. Frh might be involved in maturation processes instead.

Acknowledgements The research was supported by the Research Councils for Earth and Life Sciences (ALW) and Chemical Sciences (CW) with financial aid from the Netherlands Organization for Scientific Research (NWO). We gratefully thank Michel Ossendrijver for technical assistance regarding the microarray experiments. We gratefully thank Dr. Stan J. J. Brouns for critically reading this chapter.

65 66 CHAPTER 5

Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

Petra Worm, Bernardine L. M. van Kuijk, Wilfred R. Hagen, Remco Kort, Frank Schuren, Alona Rusakov, Alfons J.M. Stams and Caroline M. Plugge.

Abstract S. fumaroxidans degrades propionate to acetate via the methyl-malonyl-CoA pathway in syntrophic association with hydrogen and formate utilizing methanogens. Oxidation of succinate to fumarate is energetically the most difficult step in this pathway. S. fumaroxidans can also grow on propionate with fumarate and sulfate as electron acceptors. Here we describe the characterization of a purified succinate dehydrogenase in S. fumaroxidans and aim to construct the most probable fumarate reduction and succinate oxidation mechanisms by using gene analysis and transcriptional profiling. For succinate oxidationS. fumaroxidans uses Sdh (encoded by Sfum_1998-00) coupled to the periplasmic hydrogenase (Hyn), and the periplasmic formate dehydrogenase, (FDH2). For fumarate reductionS. fumaroxidans uses both Frd (encoded by Sfum_4092-95) and the reversible Sdh coupled to FDH2. Frd lacks a transmembrane subunit and instead receives electrons from menaquinone via cytochrome bd: quinol oxidases (encoded by Sfum_3008-09 and Sfum_0338-39), and cytochrome b (encoded by Sfum_2932).

Hyn and FDH2 both receive electrons from cytochrome c3 (encoded by Sfum_4047). For electron transfer across the cytoplasmic membrane Hyn requires cytochrome b, whereas FDH2 requires cytochrome cb561 (encoded by Sfum_0090-91).

67 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

Introduction Bacterial succinate:quinone oxidoreductases (SQR) are bidirectional enzymes that function either as succinate:quinone oxidoreductase (succinate dehydrogenase, Sdh) in succinate oxidation or as quinol:fumarate oxidoreductase (fumarate reductase, Frd) in fumarate fermentation (Weingarten et al. 2009; Cecchini et al. 2002; Lemma et al. 1991; Samain et al. 1984). In some cases, fumarate reduction and succinate oxidation are carried out by separate enzymes (Hägerhäll 1997). Sdh’s and Frd’s are undistinguishable similar regarding their amino acid sequences. In addition, the primary structures of Sdh’s and Frd’s are undistinguishable similar to L-aspartate oxidases (LASPO’s) that irreversibly reduce fumarate to succinate coupled to oxidation of L-aspartate to imino- aspartate (Mattivi et al. 1999). Succinate oxidation via a menaquinone to fumarate is the most energy dependent reaction in the methyl-malonyl-CoA pathway to degrade propionate to acetate in S. fumaroxidans. Succinate oxidation via menaquinone is endergonic since the midpoint potential of succinate is more positive (+30 mV) than menaquinone (-80 mV). Therefore, the reaction requires a transmembrane proton gradient to function. Schink (1997) calculated that about ⅔ ATP has to be invested to make this reaction energetically possible at hydrogen partial pressure (1Pa) and formate concentrations (10 µM) that can be maintained by methanogens in a syntrophic community. The cytoplasmic oriented fumarate reductase (Frd) of Wolinella succinogenes receives electrons for fumarate reduction from formate or hydrogen oxidation by a periplasmic formate dehydrogenase or hydrogenase via a menaquinone loop (Hägerhäll, 1997). This mechanism generates ⅔ ATP which corresponds with a proton motive force generation by about two excreted protons (Kröger et al. 2002). Van Kuijk et al. (1998) reported that S. fumaroxidans, when grown on fumarate, yields ⅔ ATP, and the cells contain menaquinones 6, -7 and cytochromes c and b. This suggests a similar fumarate reduction mechanism in S. fumaroxidans when compared to W. succinogenes. The authors also proposed that this mechanism in reversed direction could facilitate the utilization of ⅔ ATP for the energy dependent succinate oxidation (van Kuijk et al. 1998). Generally Frd’s and Sdh’s are composed of a cytoplasmic functional FAD containing subunit, a cytoplasmic subunit that binds three iron-sulfur clusters and a membrane integrated subunit that contains heme groups to transfer electrons to menaquinones (Hägerhäll, 1997). The cytoplasmic domains are highly conserved in prokaryotes. By contrast, the membrane integrated subunit is highly variable. Based on their hydrophobic domain and heme content succinate:quinone oxidoreductases can be classified in 5 types (typeA-E) (Hägerhäll 1997; Lancaster 2002; Hägerhäll and Hederstedt 1996; Hederstedt 1999). The periplasmic formate dehydrogenases and hydrogenases from W. succinogenes contain transmembrane subunits with heme-groups to facilitate electron transfer from formate or hydrogen for menaquinol reduction (Fig. 1). The periplasmic formate dehydrogenase and hydrogenase from S. fumaroxidans, however, lack such transmembrane subunits. Instead the soluble cytochrome c and hydrophobic cytochrome b may facilitate electron transfer to menaquinol. In several sulfate-reducing bacteria, such as Desulfovibrio gigas, Desulfovibrio vulgaris and Desulfomicrobium

68 CHAPTER 5

+ + - 2H H2 CO2+H formate

Mo NiFe 4Fe4S + 4Fe4S 2H 4Fe4S 4Fe4S 2H+ 4Fe4S 4Fe4S 3Fe3S 4Fe4S MKH 2 MKH2

MK MK 6 TM 6 TM 3Fe4S 4Fe4S 2H+ 2Fe2S 2H+ FAD

succinate fumarate + 2H+ Figure 1 The molecular mechanism for fumarate reduction inW. succinogenes (modified from Lancaster et al. 2006). Dotted arrows indicate electron transfer. Heme groups are indicated as diamonds.

norvegicum, a periplasmic hydrogenase binds to the periplasmic soluble cytochrome c3 in order to transfer electrons via transmembrane proteins to finally reduce sulfate in the cytoplasm, which is proposed as a general mechanism for hydrogen cycling in sulfate- reducing bacteria (Odom and Peck 1981a; Aubert et al. 2000; Yahata et al. 2006). Here we describe the characterization and purification of a succinate dehydrogenase in S. fumaroxidans. The fumarate reduction and succinate oxidation mechanisms in S. fumaroxidans are proposed based on gene analysis and transcriptional profiling.

Materials and methods Genome analysis The genome of S. fumaroxidans (Genbank nr. CP000478) was screened for genes with similarity to Frd’s, Sdh’s and LASPO’s coding genes. Retrieved putative alpha and beta subunits were aligned against Frd’s, Sdh’s and LASPO’s from other microorganisms with BLAST (Altschul et al. 1997; Altschul et al. 2005) and Clustal X 2.0.12. The N-terminal amino acid sequence of SdhABC was used to find the corresponding sdhABC gene cluster. Pfam search (Sanger institute, 2009) was used to identify motifs in the amino acid sequences and the TMHMM Server v. 2.0 (DTU, 2009) was used to identify transmembrane helices. With the TatP 1.0 Server twin-arginine translocation (Tat) motifs in the N-terminus were identified to predict protein localization S.in fumaroxidans cells (Bendtsen et al. 2005).

Organisms and cultivation Syntrophobacter fumaroxidans MPOB (DSM 10017) was grown in medium described previously (Chapter 3). Mass cultivation for enzyme purification was done in25-L carboys containing 20 L of medium with 20 mM fumarate as the substrate. For the

69 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

preparation of smaller volumes of cell extracts, the bacterium was grown in 3-L serum bottles containing 1.5 L medium, and 20 mM fumarate, or 20 mM propionate plus 20 mM sulfate as the substrate. For microarray analysis, S. fumaroxidans was cultivated with the substrates; 20 mM propionate plus 60 mM fumarate, and with 30 mM propionate plus 25 mM sulfate in pure cultures, and on 30 mM propionate with Methanospirillum hungatei JF-1 (DSM 864) or Methanobacterium formicicum JF-1 (DSM 1535) in syntrophic cocultures (as described in Chapter 4). M. hungatei was grown axenically in the same medium with 1

mM acetate and H2/CO2 (80:20, vol/vol) (Chapter 4). Growth was followed by measuring optical density, sulfide or methane formation. Cells were harvested anaerobically in the logarithmic phase (Chapter 4) and RNA was isolated as described previously (Chapter 3).

Preparation of cell fractions Cell fractions were prepared under anoxic conditions. Cultures of 1.5 L were centrifuged at 16,000 x g for 10 min. Large culture volumes (20 L) were harvested at a similar speed by continuous-flow centrifugation (Biofuge 28RS, Heraeus Sepatech, Osterode, Germany). Cell pellets were washed with 100 mM Tris-HCl, pH 7.5, suspended in 100 mM Tris-HCl, pH 7.5, containing 250 mM sucrose, 5 mM dithiothreitol (DTT), 20 mM KCl, and 5 mM MgCl2 (buffer A), and disrupted by passing twice through a French pressure cell at 110 MPa. After a DNase incubation of 15 min at room temperature, cell debris was removed by centrifugation for 20 min at 16,000 xg . The supernatant was centrifuged for 1.5 h at 140,000 x g and 4°C to separate the membrane fraction from the soluble fraction. The membrane fraction was suspended in buffer A.

Enzyme and protein assays

Enzyme activities were measured spectrophotometrically at 37 °C in2 N -flushed cuvettes, which were closed with rubber stoppers. One unit of enzyme activity corresponds to the amount of enzyme that catalyzes the conversion of 1 µmol substrate per min. Fumarate reductase was assayed with reduced benzyl viologen (Boonstra et al. 1975), reduced

2,3-dimethyl-1,4-naphtohoquinone (DMNH2) (Möller-Zinkhan and Thauer 1988),

reduced flavin mononucleotide (FMNH2) (Hiraishi 1988) or NADH (Meinhardt and Glass 1994) as electron donor. The activity of fumarate reductase with hydrogen or formate as electron donor was measured according to the methods of Macy et al. (1976). The assay mixture contained 50 mM Tris-HCl (pH 7.6), saturated with hydrogen, 50 mM

MgCl2, 1.8 mM GTP, 1.8 mM phosphoenolpyruvate, 0.075 mM NADH, 1.85 mM acetyl- CoA, 5 U succinyl-CoA synthetase, 10 U pyruvate kinase, 25 U lactate dehydrogenase, and membrane fraction. The reaction was started by addition of fumarate (saturated with hydrogen) to a concentration of 2 mM. The oxidation of NADH was followed at 340 nm and 37 °C. The reduction of fumarate by formate was measured by a small- modified assay as described for fumarate reduction with hydrogen; the Tris-buffer and the fumarate solution were saturated with nitrogen, and formate was added to the assay mixture to a concentration of 10 mM. Succinate dehydrogenase was assayed with 2,6-dichlorophenolindophenol (DCPIP)

70 CHAPTER 5 and phenazine methosulfate (PMS) according to Schirawski and Unden (1995). Protein was determined by the method of Bradford (1976), with bovine serum albumin as the standard.

Purification of SdhABC All purification steps were performed at room temperature under strictly anoxic conditions in an anaerobic chamber with N2/H2 (96:4; vol/vol) as gas phase. Membrane fraction was isolated from 24 g (wet weight) MPOB cells as described above. The membranes were solubilized by the addition of Triton-X100 to a final concentration of 3% (vol/vol). After stirring at 4 °C for 15 h the membrane fraction was centrifuged at 140,000 x g and 4 °C for 90 min. The solubilized enzyme preparation (supernatant) was applied to a Q-Sepharose column (3.2 x 9.3 cm) from Pharmacia Biotech (Roosendaal, the Netherlands) that was equilibrated with 50 mM Tris-HCl, pH 7.5, containing 0.1 % Triton-X100 and 2,5 mM DTT (buffer B). A linear gradient of 0-2 M KCl in buffer B was applied at a flow rate of 1.5 ml per min. SdhABC eluted partially in the unbound protein fraction and partially at 120 mM KCl. The active fractions were pooled, concentrated in an Amicon ultrafiltration cell with a PM30 filter and applied to a hydroxyapatite column (1.2 x 8.8 cm) from BioRad (Utrecht, the Netherlands) that was equilibrated with buffer B. SdhABC was eluted with a linear gradient from 0-250 mM potassium phosphate in buffer B at a potassium phosphate concentration of approximately 125 mM. The fractions with fumarate reductase activity were concentrated in an Amicon ultrafiltration cell with a PM30 filter and applied to a column of Superdex 200 (1.6 x 70.5 cm) from Pharmacia Biotech (Roosendaal, the Netherlands) that was equilibrated with buffer B, containing 250 mM NaCl. Protein was eluted at a flow rate of 0.5 ml per min.

The purified SdhABC was stored under N2 at 4 °C until used.

N-Terminal amino acid sequence determination and electron paramagnetic resonance Protein samples for N-terminal amino acid sequence determination were obtained by electroblotting onto a polyvinylidine difluoride (PVDF) membrane (Immobilan Millipore, Etten-Leur, the Netherlands) after separation by SDS-PAGE. Sequence determination was performed by Eurosequence (Groningen, The Netherlands) by automated Edman degradation with an Applied Biosystem 477A protein sequencer (Warrington, UK). The amino acid derivatives were identified by HPLC (Applied Biosystems 120A, Warrington, UK). Electron paramagnetic resonance (EPR) spectroscopy was carried out on a Bruker 200 D spectrometer with cryogenetics, peripheral equipment, and data acquisition/ manipulation facilities as described previously (Pierik and Hagen 1991).

Microarray For microarray cDNA was prepared as described in Chapter 3. Labelling, image analysis and data processing was performed as described in Chapter 4. Gene transcription in cells grown on propionate plus sulfate, propionate plus fumarate and cells grown in syntrophic association with Methanobacterium formicicum was compared to gene transcription in cells grown in syntrophic association with Methanospirillum hungatei.

71 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

Results and discussion Fumarate reductase and succinate dehydrogenase coding gene clusters in S. fumaroxidans The genome of S. fumaroxidans contains three gene clusters sdhABC, frdABEF, and sdhAB (Table 1) with sequence similarity to Sdh’s, Frd’s or LASPO’s of other organisms (Table 2). S. fumaroxidans might have the possibility to use separate enzymes for succinate oxidation and fumarate reduction. During growth with propionate plus fumarate, S. fumaroxidans (Fig. 2A) needs an active fumarate reductase, whereas during growth with propionate plus sulfate (Fig. 2B), or during syntrophic growth on propionate (Fig. 2C) an active succinate dehydrogenase is required. In S. fumaroxidans, the gene cluster sdhABC (Table 1) codes for two cytoplasmic subunits (sdhA and B) and a five-trans-membrane (5TM) subunit containing heme (sdhC) which is similar to the type B trans-membrane subunit of W. succinogenes Frd and Bacillus subtilis Sdh (Hägerhäll 1997) (Table 3A). SdhA contains the conserved catalytic core residues H, H-P-T and H (indicated as “a”, “b” and “c” in Fig. 3A respectively). SdhB contains motifs for binding of three iron sulfur clusters, [2Fe2S], [4Fe4S], and [3Fe4S] (Fig. 3B). The gene cluster frdABEF (Table 1) of S. fumaroxidans lacks a gene coding for the trans-membrane subunit and is therefore classified as a type E Frd. The type E succinate:quinone oxidoreductases do not contain heme. They have two hydrophobic subunits SdhE and SdhF and are different from the other four types. Rather, the type E succinate:quinone oxidoreductases are more similar to those in Cyanobacteria (Synechosistis) and heterodisulfide reductases from methanogens such as Methanocaldococcus jannaschii (formerly known as Methanococcus jannaschii) (Lemos et al. 2002). Like other gene clusters coding for E-type Frd’s, frdABEF contains frdE that codes for a cysteine-rich domain involved in cytochrome binding and membrane attachment (Lemos et al. 2002). The function of the protein encoded by frdF is unknown (Lemos et al. 2002). FrdA contains the conserved catalytic residues H-P-T and H (indicated

Table 1. Locus tags (JGI 2009) of genes coding for proteins involved in the fumarate reductase or succinate oxidation mechanism of S. fumaroxidans.

Gene Gene locus tag sdhABC Sfum_1998-00 fdh2 αβ Sfum_1273-75

cytcb561 Sfum_0090-91

cytc3 Sfum_4047 cydAB-1 Sfum_3008-09 cydAB-2 Sfum_0338-39 frdABEF Sfum_4092-95 hyn αβ Sfum_2952-53 cytb5 Sfum_3227 cytb Sfum_2932 sdhAB Sfum_0172-74

72 CHAPTER 5

propionate propionate

CoASH CoASH

propionyl-CoA propionyl-CoA

CO2 CO2 methylmalonyl-CoA methylmalonyl-CoA

succinyl-CoA succinyl-CoA

ATP CoASH ATP CoASH

succinate succinate A 2 succinate B ¾ sulfide 2[H] 4[H] 6[H] fumarate fumarate 2 fumarate

¾ sulfate malate malate

C 2[H] 2[H] methanogen oxaloacetate oxaloacetate

3H 3 formate 2 pyruvate pyruvate CoASH CoASH CO2 CO2 CO 2 2[H] 2[H] 6[H] 6[H] acetyl-CoA acetyl-CoA

acetate acetate Figure 2 Pathway of propionate degradation of S. fumaroxidans grown in pure culture with fumarate (A) or sulfate (B) as electron acceptor or in syntrophic association with M. hungatei or M. formicicum (C) (Plugge et al, 1993).

“b” and “c” in Fig. 3A respectively), but lacks the Histidine residue indicated by “a” in Fig. 3A. The latter conserved histidine residue was found to bind FAD in the succinate reductase of Wolinella succinogenes. Remarkably, alpha subunits of LASPO’s also lack this histidine residue. This combined with the fact that S. fumaroxidans can grow on aspartate (Harmsen et al. 1998), might suggest that frdABEF codes for a LASPO. FrdB does not show high similarity with Frd beta subunits of other organisms (Table 2), but does contain motifs for binding of two iron sulfur clusters, [2Fe2S] and [4Fe4S] (Fig. 3B). The lack of [3Fe4S] may be compensated by iron sulfur clusters present in FrdE and F. The third gene cluster with similarity to succinate:quinone oxidoreductases (sdhAB) lacks a gene coding for a transmembrane domain. SdhA contains the conserved catalytic residues indicated as “a”, “b”, and “c” in Fig. 3A. The SdhB contains a motif for [2Fe2S] iron sulfur cluster binding, but lacks motifs for [4Fe4S] or [3Fe4S] binding (Fig. 3B). Therefore, the required electron transfer via three iron-sulfur clusters cannot occur, meaning that sdhAB does not code for a conventional Sdh, Frd or LASPO (Lancaster 2002).

73 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

Table 2 BLAST results with e-value’s indicating the likeliness of non-similarity. Gene numbers of alpha and beta subunits of Sdh’s, Frd’s and LASPO’s are indicated. Table 2A Alpha and beta subunits of SdhABC, FrdABEF, and SdhAB* from S. fumaroxidans aligned against Sdh’s and Frd’s from other microorganisms.

code Frd’s/Sdh’s SdhA SdhB FrdA FrdB SdhA* SdhB*

TA-A Thermoplasma acidophilum 1e-95 2e-25 1e-42 4 1e-122 8e-29 TA-B DSM, A:TA1001, B:TA1002 BS-A Bacillus subtilis 1e-75 6e-30 3e-44 5e-2 2e-78 7e-15 BS-B A: BSU28440, B: BSU28430 WS-A Wolinella succinogenes 1e-2 4e-80 2e-34 2e-1 3e-98 3e-22 WS-B A:2BS2_A, B:2BS2_B ML-A Micrococcus luteus 4e-85 3e-20 2e-48 2e-3 3e-132 1e-29 ML-B A:Mlut_04830, B:Mlut_04820 EC-A Escherichia coli K12 5e-88 3e-28 1e-44 ≥1 3e-105 2e-25 EC-B A: b4154, B: b0724 SC-A Saccharomyces cerevisiae 4e-87 1e-26 2e-41 ≥1 9e-117 5e-23 SC-B A:EDV12956, B:YLL041C

[TA] Anemüller et al. 1995; [BS] Hederstedt 2002; [WS] Lancaster et al. 1999; [ML] Crowe et al. 1983, [EC] Iverson et al. 1999; [SC] Robinson et al. 1992.

Table 2B A subunits of SdhABC, FrdABEF, and SdhAB* from S. fumaroxidans aligned against Sdh’s and Frd’s from other microorganisms

Code LASPO’s SdhA FrdA SdhA*

Escherichia coli K12 EC-L 3e-40 1e-41 6e-65 NadB, AP_003160 Sulfolobus tokodaii ST-L 5e-33 8e-32 2e-54 ST1196 Pyrococcus horikosii OT-3 PH-L 8e-36 1e-29 3e-45 PH0015 Bacillus subtilis BS-L 3e-36 3e-33 3e-62 BSU27870

[EC] Mattevi 1999; [ST] Sakuraba et al. 2008; [PH] Kawarabayasi et al. 1998, Saunders et al. 2008; [BS] Lemma et al. 1991.

Table 3. Specific activity of FDH2 (de Bok et al. 2003) and Hyn (de Bok thesis 2002). One Unit is defined as 1 µmol ml-1 min-1 oxidized/produced. Specific activity Electron donor / acceptor pH (U/mg) FDH2 formate / BV2+ 8 2700 formate / MV2+ 8 510 formate / MV2+ 7.3 68 + - MV /HCO3 7.3 89 2+ Hyn H2 / BV 8 1600 2+ H2 / MV 8 1100 H2 / AQDS 8 29 MV+ / H+ 7 350

74 CHAPTER 5 Figure 3A

a

b

75 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

c

76 CHAPTER 5

Figure 3B

[2Fe2S]

[4Fe4S]

[3Fe4S]

Figure 3 Alpha (Fig. 3A) and beta (Fig. 3B) subunits of SdhABC (SF-1), FrdABEF (SF-2), and SdhAB (SF-3) from S. fumaroxidans are aligned with amino acid sequences of alpha and beta subunits of fumarate reductases, succinate reductases from other microorganisms: Thermoplasma acidophilum (TA-A), Bacillus subtilis (BS-A and BS-B), Wolinella succinogenes (WS-A and WS-B), Micrococcus luteus (ML-A and ML-B), Escherichia coli K12 (EC-A and EC-B) and Saccharomyces cerevisiae (SC-A and SC-B). In addition, amino acid sequences of alpha subunits of LASPO’s from Escherichia coli K12 (EC-L), Sulfolobus tokadaii (ST-L), Pyrococcus horikosii (PH-L) and Bacillus subtilis (BS-L) were used for alignment. Conserved catalytic core residues are indicated as “a”, “b”, and “c”. Iron sulfur cluster binding motifs are indicated as [2Fe2S], [4Fe4S], and [3Fe4S] (Hägerhäll 1997).

77 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

The genome of S. fumaroxidans contains three genes coding for the hydrophobic

cytochrome b (cytb, cytb561 and cytb5) (Table 1). Two genes coding for cytochrome c

were found; one codes for a cytochrome c and is located in an operon with cytb561

(cytcb561) and the other codes for cytochrome c3. Furthermore, two gene clusters coding for cytochrome bd: quinol oxidase (cydAB-1 and -2) (Table 1) were found, that may be involved in trans-membrane proton transfer.

Purification of the membrane bound fumarate reductase/succinate dehydrogenase from S. fumaroxidans The membrane-bound SdhABC was purified after solubilisation with the detergent Triton X-100. The purified enzyme consisted of three subunits, with a molecular mass of 70.5, 33.5 and 23.5 kDa (Fig. 4). This corresponds with the molecular mass that was calculated from the amino acid sequences of Sfum_1999 (67.4 kDa), Sfum_1998 (28.9 kDa) and Sfum_2000 (25.4 kDa). The N-terminal amino acid sequences of the two largest subunits were determined (MDTFYTDLLXVGAGLAGERVA for the 70.5 kDa subunit and MGRPLKFSVFRFNPLD for the 33.5 kDa subunit) confirming that the enzyme was coded by Sfum_1998-2000. The purified enzyme showed activity both in the fumarate reduction as well as succinate oxidation direction. Frd activity varied from 27 to 86 U/ mg protein and Sdh activity from 0.9 to 1.8 U/mg, in a series of 3 purifications. During the purification process fumarate reductase activity was also detected in the unbound fractions, indicating the presence of other enzymes. Purification of these fractions was unsuccessful. The purified SdhABC had an apparent Km for fumarate of 0.025 mM, measured with 1.6 mM benzylviologen at 37 0C. The yield of the purified enzyme was too low to determine the Km for succinate. Electron paramagnetic resonance (EPR) spectroscopy, revealed the presence of three iron-sulfur clusters; [4Fe4S], [2Fe2S], and [3Fe4S]. Gene analysis of sdhB revealed that these iron-sulfur clusters bind the beta subunit.

Mechanisms for fumarate reduction and succinate oxidation by S. fumaroxidans When S. fumaroxidans cells were grown under conditions where Frd activity is needed

Figure 4 SDS-PAGE of purified SdhABC of S. fumaroxidans. Lane 1 shows the purified reductase and lane 2 the low-molecular-mass marker from Pharmacia (Roosendaal, The Netherlands). The gel was stained with Coomassie Brilliant Blue- R250.

78 CHAPTER 5

(with fumarate as the electron acceptor (Fig. 2A)) frdABEF was up-regulated compared to cells that were grown under conditions where Sdh activity is needed; when sulfate is the electron acceptor (Fig. 2B) or in syntrophic association with M. hungatei or M. formicicum (Fig. 2C) (Fig. 5). This indicates that frdABEF codes for an Frd or LASPO. In all conditions,SdhABC was transcribed in similar levels and thus codes for a reversible Sdh, which was confirmed with the purified enzyme. Cytoplasmic fumarate reduction and succinate oxidation are coupled to periplasmic hydrogenases or formate dehydrogenases (Hägerhäll 1997). Previous RT qPCR experiments (Chapter 3) showed that under conditions where Frd activity is needed, S. fumaroxidans transcribes both hynαβ and fdh2αβ in high levels, whereas under conditions where Sdh activity is required, only fdh2αβ is transcribed in high levels. Thus, fumarate reduction is coupled to both formate oxidation by FDH2 and hydrogen oxidation by Hyn (Fig. 6B), at comparable activities (at pH 8) (Table 3) (de Bok et al.

2003; de Bok 2002). Succinate oxidation is mainly coupled to CO2 reduction by FDH2 (Fig. 6A).

In S. fumaroxidans the gene coding for cytochrome c3 (cytc3) was co-transcribed with fdh2αβ (Fig. 5). This indicates binding of FDH2 to the periplasmic soluble cytochrome c3 (Fig. 6). Whether hynαβ is co-transcribed with cytc3 is less clear (Fig.

5), but cytochrome c3 might also bind to this hydrogenase (Fig. 6B). Furthermore, the operon cytcb was co-transcribed with fdhαβ (Fig. 2) indicating binding of the periplasmic

A 4

2

0

-2

-4 sdhABC fdh2 cytbc cytc3 cyd-1 cyd-2 frdABEF hyn cytb5 cytb sdhAB

B 4

2

0

-2

-4 sdhABC fdh2 cytbc cytc3 cyd-1 cyd-2 frdABEF hyn cytb5 cytb sdhAB

C 4 2 0 -2 -4 sdhABC fdh2 cytbc cytc3 cyd-1 cyd-2 frdABEF hyn cytb5 cytb sdhAB

Figure 5 Transcription of genes coding for enzymes putatively involved in the Sdh and Frd mechanisms in S. fumaroxidans. Cells were grown on propionate with fumarate (A) or sulfate (B) as electron acceptor and in syntrophic association withM. formicicum (C). The x-as indicates the log ratio of transcription in the growth conditions (A-C) compared to gene transcription in cells grown in syntrophic association with M. hungatei. Log ratio’s of genes in each operon are depicted in boxplots.

79 Fumarate reduction and succinate oxidation in Syntrophobacter fumaroxidans

+ - H + CO2 formate

A WSe 4Fe4S α 4Fe4S + 2H 4Fe4S β + 2H Cytc3 Cytc MKH 5TM 2 Cytb

C MK 3Fe4S 2H+ 4Fe4S 2Fe2S 2H+ B FAD A

succinate fumarate + 2H+

H+ + CO formate- 2H+ H 2 B 2 NiFe WSe 4Fe4S 4Fe4S α 4Fe4S α 4Fe4S 2H+ 2H+ 2H+ 2H+ 3Fe3S β 4Fe4S β Cytc3 Cytc3 Cytc MKH MKH 5TM 2 2 Cytb MKH2 Cytb CydB CydA

C MK Cytb MK MK 3Fe4S 4Fe4S + 4Fe4S 2H 4Fe4S 2H+ 4Fe4S 2Fe2S 2Fe2S E 2H+ 2H+ B 4Fe4S FAD B F FAD A A

succinate fumarate succinate fumarate + 2H+ + 2H+

Figure 6 The proposed molecular mechanism for succinate oxidation (A) and fumarate reduction (B) in S. fumaroxidans. Dotted arrows indicate electron transfer. Heme groups are indicated as diamonds.

80 CHAPTER 5

formate dehydrogenase and cytochrome c3 with cytochrome c and a membrane bound menaquinone-binding cytochrome b (Fig. 6A). The genes cytb and cytb5 were co- transcribed with hynαβ (Fig. 5), which indicates binding of the Hyn and cytochrome c3 with cytochromes b and b5 (Fig. 6B). The membrane subunit of Frd from W. succinogenes was shown to facilitate proton transfer through the membrane in the direction of fumarate reduction (Madej et al. 2009). This balances the energy of protons transported through the membrane (Madej et al. 2009). A similar proton translocation function might be the case for SdhC in S. fumaroxidans (Fig. 6B), whereas FrdABEF of S. fumaroxidans instead uses a transmembrane cytochrome bd: quinol oxidase (Fig 1B). In E. coli these cytochrome bd: quinol oxidases, bind heme groups and quinones to transport protons through the cytoplasmic membrane (Borisov et al. 2009). In S. fumaroxidans cydAB-1 and -2 were co-transcribed with frdABEF. Thus, the oxidoreductase complex that binds Frd, might allow for this proton transfer.

Conclusion The most probable fumarate reduction and succinate oxidation mechanisms in S. fumaroxidans were constructed based on gene analysis, transcriptional profiling and enzyme measurements, (Fig. 6A and B). Succinate oxidation in S. fumaroxidans occurs via SdhABC and is mainly coupled to CO2 reduction with FDH2 and only partly to proton reduction with Hyn (Fig. 6A). Fumarate reduction can occur via SdhABC and FrdABEF, and is coupled to simultaneous CO2 reduction with FDH2 and proton reduction with Hyn (Fig. 6B). FrdABC, that lacks a transmembrane subunit, retrieves electrons from menaquinone via cytochrome bd oxidoreductases, and cytochrome b. Hyn and FDH2 both conduct electrons from cytochrome c3. For electron transfer across the cytoplasmic membrane, Hyn requires cytochrome b, whereas FDH2 requires Cytcb561.

Acknowledgements The research was supported by the Research Councils for Earth and Life Sciences (ALW) and Chemical Sciences (CW) with financial aid from the Netherlands Organization for Scientific Research (NWO). We gratefully thank Michel Ossendrijver for technical assistance regarding the microarray experiments.

81 82 CHAPTER 6

Decreased activity of a propionate- degrading community in a UASB reactor fed with synthetic medium without molybdenum, tungsten and selenium

Petra Worm, Fernando G. Fermoso, Piet N. L. Lens and Caroline M. Plugge

Published in Enzyme and Microbial Technology 45 (2009) 139-145

Abstract The composition and dynamics of the propionate degrading community in a propionate- fed up-flow anaerobic sludge bed (UASB) reactor with sludge originating from an alcohol distillery wastewater treating UASB reactor was studied. The rather stable propionate degrading microbial community comprised relatives of propionate degrading Syntrophobacter spp., the hydrogen and formate consuming Methanospirillum hungatei and the acetate consuming Methanosaeta concilii. The effect of the long- term absence of molybdenum, tungsten and selenium from the feed to the UASB reactor on microbial community dynamics and activity was examined. Measurements for metal concentrations of the sludge and specific methanogenic activity tests with supplied molybdenum, tungsten and selenium were found to be unsuitable to detect the potential limitation of the microbial activity of the UASB sludge by these trace metals. During a long-term absence of molybdenum, tungsten and selenium from the feed to the UASB reactor, the methanogenic activity decreased while relatives of Smithella propionica and Pelotomaculum spp. competed with Syntrophobacter spp. for propionate consumption.

83 Decreased activity of a propionate degrading community in a UASB reactor deprived of Mo, W, and Se

Introduction In upflow anaerobic sludge bed (UASB) reactors, complex organic matter is degraded

to methane and CO2 by anaerobic microorganisms with different metabolic capabilities (McInerney et al. 2008). Propionate is an important intermediate in methanogenic degradation of complex organic matter. It often accumulates during failure ofthe wastewater treatment process (Kida et al. 1993; Ariesyady et al. 2007a). Propionate degradation to acetate, carbon dioxide, and hydrogen and/or formate is highly endergonic under standard conditions and only proceeds when other microorganisms, methanogenic archaea in the case of UASB reactors, keep the hydrogen and formate concentration, and to a lesser extent, the acetate concentration low (McInerney et al. 2008). Most propionate oxidizing bacteria and methanogenic archaea contain formate dehydogenases and hydogenases that are likely involved in interspecies formate and hydrogen transfer (de Bok et al. 2002a). The catalytic centres of formate dehydrogenases and hydrogenases contain metal-binding cofactors that are essential for their catalytic function. Formate dehydrogenases usually require molybdenum or tungsten (Hille, 2002), whereas hydrogenases usually require iron and/or nickel (Vignais and Billoud, 2007). In addition, some formate dehydrogenases and hydrogenases also contain selenium Hille, 2002; Vignais and Billoud, 2007). Depletion of molybdenum and tungsten was recently described to decrease formate dehydrogenase activity in defined syntrophic growth of the propionate oxidizing Syntrophobacter fumaroxidans and methanogenic Methanospirillum hungatei (Plugge et al. 2008). Furthermore, Lebuhn et al. (2008) indicated that for long-term mono-digestion of maize silage, molybdenum and selenium should be supplied. The lack of molybdenum, tungsten and selenium in the influent of a biological process could thus result in a decrease of the propionate consumption and ultimately lead to process failure in a UASB reactor. Several syntrophic propionate degrading cultures enriched from UASB reactors and other environments have been described. Syntrophobacter wolinii was the first syntrophic species described (Boone and Bryant, 1980). Other examples are Syntrophobacter pfennigii (Wallrabenstein et al. 1995), S. fumaroxidans (Harmsen et al. 1998) and S. sulfatireducens (Chen et al. 2005). These Syntrophobacter spp. belong to the δ-proteobacteria and are able to couple propionate degradation also to sulfate reduction. Smithella spp. are propionate degraders more related to syntrophic benzoate degraders such as Syntrophus (Liu et al. 1999). Pelotomaculum is a gram- positive spore forming propionate degrader that is not in pure culture yet (de Bok et al. 2005; Imachi et al. 2007). In contrast to Syntrophobacter spp., both Smithella spp. and Pelotomaculum spp. are not able to reduce sulphate (Liu et al. 1999; Imachi et al. 2007). Syntrophic partners that are commonly present in UASB reactors are the methanogenic archaea Methanospirillum hungatei and Methanobacterium formicicum, that consume hydrogen and formate, and species of the genera Methanosaeta and Methanosarcina that consume acetate (Stams et al. 2005). Pelotomaculum propionicicum and Smithella propionica were never found to be dominant species in UASB reactors (Sekiguchi et al. 1999; Liu et al. 2002; Roest et al. 2005). In contrast, in anoxic sediment samples, Pelotomaculum spp. and Smithella

84 CHAPTER 6

Table 1. Composition of trace element solution added to influent and activity tests media.

Compound Element Conc. (µM)

. FeCl2 4H2O Fe (II) 5 . CuCl2 2H2O Cu (II) 0.5

ZnCl2 Zn (II) 0.5 . MnCl2 4H2O Mn (II) 0.5 . NiCl2 6H2O Ni (II) 0.5 . CoCl2 6H2O Co (II) 0.5 . a (NH4)6Mo7O24 4H2O Mo (VI) 0.5 . a Na2SeO4 5H2O Se (VI) 0.5 . a Na2WO4 2H2O W (VI) 0.5

aadded to the reactor influent, only from operation day 249 onwards.

spp. seem to compete more successfully with Syntrophobacter spp. for the substrate propionate and their syntrophic relation withM. hungatei (Luerdes et al. 2004; Chauhan and Ogram, 2006). S. propionica dominated in an anaerobic digester fed with different concentrations of propionate (Aryesyady et al. 2007a). In chemostat experiments it was shown that high dilution rates promotePelotomaculum spp., whereas low dilution rates promote Syntrophobacter spp. (Shigematsu et al. 2006). From these reports can be concluded that UASB reactor conditions are highly selective for Syntrophobacter spp. whereas Pelotomaculum spp., Smithella spp. and Syntrophobacter spp. are able to simultaneously degrade propionate in syntrophy with hydrogen and formate consuming methanogens in other environments. The aim of this study was to analyse the composition of the propionate degrading community in a propionate-fed UASB reactor with sludge originating from an alcohol distillery wastewater treating UASB reactor. Moreover, we examined the effect of the long-term absence of molybdenum, tungsten and selenium from the feed to the UASB reactor on microbial community dynamics and activity.

Materials and methods UASB reactor operation Methanogenic granular sludge was obtained from a lab scale reactor that was inoculated with methanogenic sludge from a full-scale UASB reactor treating alcohol distillery wastewater at Nedalco (Bergen op Zoom, The Netherlands) and was fed with propionate as the sole substrate during 140 days. The wet sludge contained 5.23 (± 0.02) % total suspended solids (TSS) and 4.93 (± 0.03) % volatile suspended solids (VSS). The total suspended solids (TSS) and volatile suspended solid (VSS) concentrations were determined according to Standard Methods (APHA, AWWA, and WEF, 1998). A 3.25 L glass cylindrical UASB reactor was inoculated with 1 litre of wet sludge

85 Decreased activity of a propionate degrading community in a UASB reactor deprived of Mo, W, and Se

and operated in a temperature-controlled room at 30 (±2)°C at a hydraulic retention time (HRT) of 12 h and a superficial liquid upflow velocity of 0.35 m.h-1. For the influent and recycle flow, peristaltic pumps (type 505S, Watson and Marlow, Falmouth, U.K.) were used. The produced biogas was led through a water lock filled with a concentrated

NaOH (15%) solution in order to remove CO2 and H2S. The volume of produced methane was measured with a wet gas meter (Schlumberger Industries Dordrecht, The Netherlands). -1 The UASB reactor contained an initial biomass concentration of 6.38 g VSS.LReactor and was fed with a basal medium consisting of propionate, inorganic macronutrients and a trace element solution dissolved in double distilled water. From day 0 till day 44, the

-1 -1 Table 2. Evolution of the specific methanogenic activity (g CH4-CODg VSS day ) as a function of time, organic source and trace metal content of the medium.

Operation days

0 92 167 182 235 249

No trace metals 0.74 0.37 0.10 0.28 0.09 n.d. Complete trace element 0.60 0.38 0.11 0.27 0.10 0.21 solution except Mo, W, Se Propionate Complete trace element 0.82 0.35 0.10 0.25 n.d. n.d. solution Mo, W and Se solely 0.75 0.33 0.11 0.25 n.d. n.d.

No trace metals 0.33 1.39 0.89 1.18 0.96 n.d. Complete trace element 0.38 1.27 0.99 1.15 0.91 0.67 solution except Mo, W, Se Formate Complete trace element 0.31 1.36 n.d. 1.08 0.85 n.d. solution Mo, W and Se solely 0.39 1.34 0.97 n.d. 0.97 n.d.

No trace metals 0.39 n.d. 0.22 0.26 0.19 n.d. Complete trace element 0.34 0.86 0.23 n.d. 0.21 0.30 solution except Mo, W, Se H /CO 2 2 Complete trace element 0.35 0.79 0.20 n.d. 0.25 n.d. solution Mo, W and Se solely 0.29 0.77 0.20 0.25 0.21 n.d.

No trace metals 0.89 0.85 0.46 0.63 0.59 n.d. Complete trace element 0.97 n.d. 0.42 0.53 0.51 0.48 solution except Mo, W, Se Acetate Complete trace element 0.93 0.70 0.38 0.45 0.53 n.d. solution Mo, W and Se solely 0.91 1.10 0.46 0.44 0.55 n.d.

n.d.: not determined

86 CHAPTER 6 reactor was operated at an organic loading rate (OLR) of 3.3 (± 1.4) g COD.L-1.day-1. From day 44 till day 276 (end of the experiment), the OLR applied was 11.8 (± 2.9) g COD.L-1. day-1. Volatile Fatty Acids (VFA) were determined using gas liquid chromatography as described by Weijma et al. (Weijma et al. 2000). -1 The basal medium contained inorganic macronutrients (in mg.L ); NH4Cl (280),

K2HPO4 (250), MgSO4·7H2O (100) and CaCl2·2H2O (10). To ensure pH stability around pH

7.0, 2.52 g (30 mM) of NaHCO3 was added per litre of basal medium. To avoid precipitation in the storage vessels, the influent was composed of 3 streams: (1) macronutrients and trace element solution without 2K HPO4 and NaHCO3, (2) propionate with NaHCO3 and

K2HPO4 and (3) double distilled water. The composition of the trace elements solution supplied to the UASB reactor is shown in Table 1. During the first 249 days, no molybdenum, tungsten or selenium was supplied. From day 249 till day 276 (end of the experiment), the UASB reactor was supplied with 0.5 µM of molybdate, tungstate and selenate. The metal content of the sludge was measured on days 0, 92, 126, 170, 180, 190 and 240, and determined by Inductively Coupled Plasma - Optical Emission Spectrometer (ICP-OES) (Varian, Australia) after destruction with Aqua regia (mixture of 2.5 ml 65% HNO3 and 7.5 ml 37% HCl) as described by Fermoso et al. (2008). The method quantification limit (MQL) for tungsten, selenium and molybdenum was 20 µg g TSS-1, 10 µg g TSS-1 and 5 µg g TSS-1, respectively.

Specific methanogenic activity test The activity with different substrates involved in propionate degradation (propionate, acetate, H2 and formate) and possible metal limitation of the sludge that developed in the reactor were investigated by specific methanogenic activity (SMA) tests (Table 2). The SMA of the sludge was determined in duplicate at 30 (±2) oC using on-line methane production measurements as described by Zandvoort et al. (2002). Approximately 1 g (wet weight) of granular sludge was transferred to 120-ml serum bottles containing 50 ml of basal medium with the same composition as the reactor basal medium, -1 -1 -1 supplemented with propionate (2 g COD L ), acetate (2 g COD L ), H2 (1 g COD L ) or formate (2 g COD L-1) as the substrate. Data were plotted in a rate versus time curve, using moving average trend lines with an interval of 10 data points. The SMA with each of the substrates was determined on days 0, 92, 167, 182, 235 and 249. The samples were always taken from the same place (the mid-height) in the sludge bed.

DNA extraction, amplification and DGGE Total genomic DNA was extracted from 50 mg granules (wet weight) sampled at day 40, 81, 123, 180, 231, 246 and 276 with the bead-beat and phenol-chloroform based DNA extraction method (van Doesburg et al. 2005). For denaturing gradient gel electrophoresis (DGGE) analysis, PCR was performed as described by Sousa et al. (2007) with the primers “U968-f ” (with GC-clamp) and “L1401-r” for bacterial 16S rRNA genes and primers “A109(T)-f” and “515-r” (with GC-clamp) for archaeal 16S rRNA genes (Table 3). Amplicons were separated by DGGE as described by Zoetendal et al. (2001) and band profiles were compared.

87 Decreased activity of a propionate degrading community in a UASB reactor deprived of Mo, W, and Se

Table 3. Primers used for cloning, sequencing and denaturing gradient gel electrophoresis.

Specificity Primer name Primer sequence (5’-3’) Reference

Bacterial 16S Bact-27-f GTT TGA TCC TGG CTC AG Lane, 1991

Universal 16S Uni1492-r CGG CTA CCT TGT TAC GAC Großkopf et al. 1998

Archaea 4F TCC GGT TGA TCC TGC CRG Kendall et al. 2007

pGEM-T T7 TAA TAC GAC TCA CTA TAG GG Promega, 2007

pGEM-T SP6 GAT TTA GGT GAC ACT ATA G Promega, 2007 CGC CCG GGG CGC GCC CCG GGC GGG GC clamp Muyzer et al. 1993 GCG GGG GCA CGG GGG G Bacterial 16S U968-f AAC GCG AAG AAC CTT AC Nübel et al. 1996

Bacterial 16S L1401-r CGG TGT GTA CAA GAC CC Nübel et al. 1996 Original Großkopf et al. 1998, Archaeal 16S A109(T)-f ACT GCT CAG TAA CAC GT but with the third nucleotide changed into T Universal 16S 515-r ATC GTA TTA CCG CGG CTG CTG GCA C Lane, 1991

Cloning and sequencing DNA from sludge samples of day 40 and day 276 were each used to make bacterial 16S rRNA gene and archaeal 16S rRNA gene clone libraries. For cloning, full length bacterial and archaeal 16S rRNA genes were amplified in a PCR with forward primers, Bact-27-f and 4F respectively and the universal primer Uni1492-r (Table 3) and thermocycling conditions as described by Sousa et al. (2007). PCR amplicons were purified with the Qiaquick PCR purification kit according to manufacturer’s instructions and cloned into E. coli JM109 by using the Promega pGEM-T easy vector system (Promega, Madison, WI). After blue white screening, positive colonies were transferred to 200 µl ampicilin containing LB medium and grown overnight at 37 oC. For each of the four clone libraries 192 colonies were picked. To lyse the cells, 10 µl overnight culture plus 90 µl Tris-EDTA buffer was incubated at 95 oC for 10 min. Pools with 12 clones were prepared and corresponding 16Sr RNA genes were analyzed with DGGE. Four clone pools with archaeal 16S rRNA genes and four clone pools with bacterial 16S rRNA genes that contained the highest band variety were selected and 16S rRNA genes from separate clones were analyzed with DGGE. The 16S rRNA genes with a unique band location after DGGE, were selected. Complete inserts were amplified in a PCR with primers “T7” and “SP6” (Table 3) and thermocycling conditions as described by Sousa et al. (2007). After purification with the Qiaquick PCR purification kit nearly full-length 16S rRNA genes were sequenced by BaseClear (Leiden, The Netherlands). Chromatogram analysis and sequence assembly were performed with DNA Baser v2.11 and phylogenetic affiliation of sequences was examined with BLAST similarity searches.

88 CHAPTER 6

Results Reactor operation During the first 44 days of reactor operation, the VFA-COD degradation was not stable (Figure 1A). Although VFA removal was not complete, the OLR of the reactor was increased to 12 g COD-L-1 day-1 (6 g COD-VFA L-1 in the influent) on day 44 of operation. VFAs were degraded up to 70% during the following 23 days of operation. The VFA effluent concentration increased after day 67, reaching a maximum of 4.7 g COD-VFA L-1 on day 81, corresponding to 60% of the influent propionate concentration on that day (Figure 1A). The VFA in the effluent in that period were composed of on average 60% propionate, 20% acetate and 20% butyrate. From day 81, the VFA effluent concentration decreased to 0 on day 116 and between day 116 and day 140, VFA were absent in the effluent. From day 140 to day 225, the VFA effluent concentrations were highly variable, but not exceeding 3.5 g COD-VFA L-1. The average VFA effluent concentration during this period was 1.5 g COD-VFA L-1 and composed of 90% propionate. At day 225 VFA started to accumulate, reaching a value of 7.4 g COD-VFA L-1 on day 249 and composed of 98% propionate. To investigate if addition of molybdenum, tungsten and selenium could recover the reactor performance, 0.5 µM of molybdenum, tungsten and selenium was supplied to the influent from day 249 onwards. Partially, the VFA effluent concentration decreased up to 1.4 g COD-VFA L-1, with 98% propionate on day 258. A further VFA effluent accumulation was observed reaching a value of 4 g COD-VFA-1 L , of which 96% propionate on day 271.

Figure 1. (A) Evolution of propionate-fed reactor performance with time. Influent COD ( ), effluent COD ( ). Dotted line represents expected VFA-COD in the effluent at operation conditions. (B) Evolution of SMA with propionate ( ) and with the intermediates to methane; formate ( ), H2 ( ) and acetate ( ) with time.

89 Decreased activity of a propionate degrading community in a UASB reactor deprived of Mo, W, and Se

Table 4. Bacterial and archaeal clones sequenced and their similarity with their closest relative. Corresponding DGGE bands are shown in Figure 2.

DGGE Band Clone Closest cultured relative Similarity (%) Accession no. 1 A01 Methanosaeta concilii 99 EU888804 2 A06 Methanosaeta concilii 99 EU888805 3 A08 Methanosaeta concilii 99 EU888806 4 A10 Methanospirillum hungatei 99 EU888808 5 A05 Methanospirillum hungatei 99 EU888809 6 A07 Methanospirillum hungatei 99 EU888810 7 A03 Methanosaeta concilii 99 EU888811 8 A12 Methanosaeta concilii 99 EU888812 9 A09 Methanospirillum hungatei 100 EU888814 10 A11 Methanosaeta concilii 94 EU888815 11 B14 Chlorobium phaeobacteroides 82 EU888816 12 B18 Chlorobium phaeobacteroides 82 EU888817 13 B12 Rubrobacter xylanophilus 82 EU888818 14 B09 Smithella propionica 99* EU888819 15 B01 Pelotomaculum propionicicum 94 EU888820 16 B10 Pelotomaculum propionicicum 94 EU888821 17 B13 Pelotomaculum propionicicum 94 EU888822 18 B03 Clostridium orbiscindens 94 EU888823 19 B04 Thermoanaerobacter pseudethanolicus 96 EU888824 20 B07 Spirochaeta caldaria 89 EU888825 21 B02 Syntrophobacter sulfatireducens 95 EU888826 22 B16 Syntrophobacter sulfatireducens 99 EU888828 * Only a partial sequence (1371bp) of Smithella propionica 16S rRNA gene available in NCBI database.

Evolution of SMA over time From day 0 to day 92 of operation, the SMA with acetate as the substrate was constant

and the SMA with H2 and formate as the substrate increased 2- and 3-fold, respectively (Table 2, Figure 1B). However, the SMA with propionate as the substrate decreased -1 -1 around 50% between day 0 (0.74 g CH4-COD g VSS d ) and day 92 (0.35 g CH4-COD g VSS-1 d-1). From day 92 to day 167, the SMA with propionate as the substrate further decreased -1 -1 to 0.10 g CH4-COD g VSS d (Table 2). The SMA with H2, acetate or formate as the

substrate also decreased. From day 167 to day 235, the SMA with H2, propionate or formate followed a slightly decreasing trend, the SMA with acetate as the substrate followed a slightly increasing trend (Figure 1B). From day 235 to day 249, the SMA with propionate as the substrate increased from -1 -1 -1 -1 0.10 g CH4-COD g VSS d to 0.21 g CH4-COD g VSS d , as well as the SMA with H2 as the -1 -1 -1 -1 substrate increased from 0.20 g CH4-COD g VSS d to 0.30 g CH4-COD g VSS d . The SMA with formate as the substrate decreased around 40%. The SMA with each substrate was not influenced by the presence of molybdenum, tungsten and selenium in the medium (Table 2).

90 CHAPTER 6

A 40 81 123 180 231 246 276 B 40 81 123 180 231 246 276

11, 12 13 14 15, 16, 17 1 2 3 4 18, 19, 20 5, 6 21, 22 7, 8, 9 10

Figure 2. DGGE band patterns of (A) Archaea and (B) Bacterial 16S rRNA gene amplicons from seven UASB reactor sludge samples (Day 40, 81, 123, 180, 231, 246 and 276). Numbers indicating DGGE bands correspond to clones listed in Table 4.

Metal content of the methanogenic granular sludge Molybdenum concentrations were constant through all the sludge samples tested (20 ± 2 µg g TSS-1). Tungsten and selenium concentrations were below the method quantification limit (MQL) in all the sludge samples investigated and no wash out of molybdenum from the sludge was observed in the effluent (Data not shown).

Molecular characterization of the propionate degrading community Sludge samples were taken from the UASB reactor over time and were used to characterize the composition and dynamics of the microbial community (Figure 2, Table 4). DGGE analysis revealed that the microbial community was rather stable during a large part of the reactor run (day 40 till day 276) (Figure 2). Within the bacterial population, only minor changes were observed. Clone library analysis revealed that the microbial community comprised micro-organisms affiliating with syntrophic propionate degrading bacteria and methanogenic archaea. Retrieved archaeal 16S rRNA gene sequences were highly similar (99 – 100%) to those of the hydrogen and formate consuming Methanospirillum hungatei (Clone A10, A05, A07 and A09; Table 4) or the acetoclastic Methanosaeta concilii (Clone A01, A06, A08, A03 and A12; Table 4). The 16S rRNA gene sequence of clone A11 was also similar to that of M. concilii, but with much lower similarity (94%). Retrieved bacterial 16S rRNA gene sequences showed high (> 97%) or low (< 97%) similarity with those of the propionate degrading bacteria: Syntrophobacter sulfatireducens (95% and 99% similarity), Smithella propionica (99% similarity) and Pelotomaculum propionicicum (94% similarity) (Table 4). Two bacterial 16S rRNA gene sequences showed some similarity with those of the glucose fermenting, thiosulfate reducing, sporeforming, thermophilic Thermoanaerobacter pseudethanolics (96%) (Onyenwoke et al. 2007) or with those of the quercetin degrading, sporeforming Clostridium orbiscindens (94%) (Winter et al. 1991) (Table 4). Three other bacterial 16S rRNA gene sequences that affiliated with those of uncultured bacteria were only

91 Decreased activity of a propionate degrading community in a UASB reactor deprived of Mo, W, and Se

distantly related (82-89% similarity) to the closest cultured relative. The 16S rRNA gene sequences of clone B14 and B18 showed 82% similarity with those of Chlorobium phaeobacteroides and 16S rRNA gene sequences of clone B07 showed 89% similarity with those of Spirochaeata caldaria (Table 4). In DGGE profiles strong bands (21 and 22), that were dominant in all samples, correspond to Syntrophobacter-like bacteria (Table 4, Figure 2B). During the slightly

decreasing SMA with H2, propionate or formate as the substrate between day 180 and 231, the intensity of the DGGE bands corresponding to S. propionica relatives (Band 16, Table 4, Figure 2) increased. During the increasing SMA with propionate or hydrogen and the decreasing SMA with formate and acetate between days 231 and 246, the intensity of the DGGE bands (Bands 15 and 16 in Figure 2 and Table 4) corresponding to clones distantly related to P. propionicicum increased. Furthermore, increased intensities of DGGE bands (11 and 12 in Figure 2) corresponding to clone B14 and B18, that were distantly related with Chlorobium phaeobacteroides, were observed from day 123 till day 246.

Discussion Molybdenum, tungsten and selenium detection During the UASB reactor run in which molybdenum, tungsten and selenium were depleted from the feed, the propionate degrading activity decreased substantially and minor changes within the propionate degrading community were observed. However, whether a limitation of the UASB sludge for molybdenum, tungsten and selenium developed, could not be measured by the ICP-OES technique and the SMA tests. Because tungsten and selenium amounts in the sludge were below the MQL during the whole reactor run, the ICP-OES technique seemed unsuitable to determine the exact concentration of these trace metals in the reactor. In contrast to tungsten and selenium, molybdenum concentrations in the sludge were detectable with the ICP-OES technique. Because no molybdenum was measured in the effluent it was either recycled by micro-organisms or present in an insoluble, for microorganisms unavailable form. Thus measurements of the molybdenum concentration in the effluent by the ICP-OES technique could neither determine nor exclude molybdenum limitation in the sludge. Retrieved SMA results showed no difference between SMA tests with or without supplied molybdenum, tungsten and selenium. However, metal limitation can still not be excluded, because it was previously described that transcription of genes coding for molybdenum or tungsten containing enzymes can be up-regulated by the presence of these metals (Schemberg et al. 2008; Regulski et al. 2008). Transcriptional regulation might be slower than posttranscriptional regulation, especially in slow growing organisms, such as syntrophic propionate degrading bacteria and methanogenic archaea.

Propionate degrading community in alcohol distillery originating sludge Several publications previously described the propionate degrading microbial community in anaerobic sludge from different locations. Sludge was analyzed from UASB reactors treating wastewater from a sugar refinery (Grotenhuis et al. 1991), a brewery (Liu et al. 2002), a paper mill (Roest et al. 2005), artificial wastewater (Sekiguchi et al. 1999)

92 CHAPTER 6 and from digesters treating domestic wastewater (Ariesyady et al. 2007a; Ariesyady et al. 2007b). Here we present the propionate degrading community in sludge that originated from an alcohol distillery. Similar to previously described UASB sludge communities, the alcohol distillery originating sludge contains relatives of propionate degrading Syntrophobacter spp. and relatives of the acetoclastic Methanosaeta concilii. Different from other UASB sludges (Sekiguchi et al. 1999; Liu et al. 2002; Roest et al. 2005), is that the alcohol distillery originating sludge investigated in this study, contained Methanomicrobiales (M. hungatei relatives) instead of Methanobacteriales, as hydrogen and formate consuming methanogens.

Accumulation of other propionate degraders Although the phylogeny and propionate degrading activity of individual microorganisms in the UASB reactor sludge need to be confirmed with dedicated enrichment studies, we refer in the discussion to the closest cultured relative of the clones. In the UASB reactor competition between Pelotomaculum spp., Smithella spp., and Syntrophobacter spp. was probably challenged. During the reactor run, the SMA with propionate, formate or acetate as the substrate decreased while Syntrophobacter spp. were the dominant propionate degraders. This indicates that Syntrophobacter spp. decreased in activity. As a consequence, Pelotomaculum spp. and Smithella spp. that accumulated in a later stage of the reactor run, likely competed with Syntrophobacter spp. for propionate degradation. Pelotomaculum and some Syntrophobacter spp. (Clone B02) showed low 16S rRNA gene similarities (< 97%) with those of the closest cultured relative, indicating the presence of undiscovered gram positive and gram negative propionate degrading bacteria. The decreased activity of Syntrophobacter spp. and the subsequent accumulation of Smithella spp. and Pelotomaculum spp., that were found to degrade propionate in syntrophic environments other than UASB reactors, might be caused by the long term absence of molybdenum, tungsten and selenium in the UASB reactor feed. Smithella spp. and Pelotomaculum spp. may use formate dehydrogenases and hydrogenases that have characteristics different from those of Syntrophobacter spp. The Pelotomaculum spp. are phylogenetically distant from Syntrophobacter spp. Moreover, S. propionica degrades propionate to acetate and butyrate, the butyrate being degraded further to acetate (de Bok et al. 2001), whereas Syntrophobacter spp. and Pelotomaculum spp. degrade propionate via the methyl malonyl CoA pathway (Plugge et al. 1993; Kosaka et al. 2006; Houwen et al. 1987). Pelotomaculum spp. and S. propionica may need molybdenum for formate dehydrogenase activity whereas Syntrophobacter spp. need tungsten for formate dehydrogenase activity and molybdenum even has an antagonistic effect as was described forSyntrophobacter fumaroxidans (Plugge et al. 2008). Thus, the UASB reactor conditions have changed during a long-term depletion of molybdenum, tungsten and selenium from the feed, in such a way that propionate degraders using formate dehydrogenases and hydrogenases with different characteristics could compete with the dominant propionate degrader.

93 Decreased activity of a propionate degrading community in a UASB reactor deprived of Mo, W, and Se

Acknowledgements The authors would like to thank Hans Heilig for excellent technical advise. We also thank A. J. M. Stams for critically reading the manuscript. The authors were supported by the Research Council for Earth and Life Sciences (ALW) with financial aid from the Netherlands Organization for Scientific Research (NWO). This research was further co- funded by the Marie Curie Excellence Grant “Novel biological engineering processes for heavy metal removal and recovery” (MEXT-CT-2003-509567).

94 CHAPTER 7

Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in anaerobic granular sludge deprived of molybdenum, tungsten and selenium

Petra Worm, Fernando G. Fermoso, Alfons J. M. Stams, Piet N. L. Lens and Caroline M. Plugge

Abstract Formate dehydrogenases and hydrogenases contain molybdenum or tungsten and/ or selenium. These enzymes are crucial for interspecies formate and hydrogen transfer between propionate degrading Syntrophobacter spp. and methanogenic Methanospirillum spp. Here we used reverse transcription of total RNA followed by quantitative PCR (RT qPCR) with specific primers to get insight into interspecies formate and hydrogen transfer. Transcriptional regulation of formate dehydrogenases and hydrogenases in Syntrophobacter and Methanospirillum spp. in a propionate-fed up-flow anaerobic sludge bed (UASB) reactor was examined. In both microorganisms formate dehydrogenase and hydrogenase coding genes (fdh and hyd, respectively) were transcribed simultaneously. During 249 days in which molybdenum, tungsten and selenium were not supplied to the reactor feed, the microbial activity and transcription of fdh and hyd in Syntrophobacter spp. decreased. Transcription of fdh and hyd in Methanospirillum spp. did not decrease, but transcription of fdh increased when after 249 days molybdenum, tungsten and selenium were supplied to the reactor feed. The developed RT-qPCR is a fast technique that can give information about active processes in methanogenic granular sludge and may contribute to predict metal limitation and failure in UASB reactors.

95 Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in slugde deprived of Mo, W, and Se

Introduction The degradation of complex organic matter to methane and carbon dioxide in up-flow anaerobic sludge bed (UASB) reactors is performed by syntrophic microorganisms that are dependent on each others metabolic conversions (Fenske et al. 2005; McInerney et al. 2008). In methanogenic degradation of complex organic matter propionate is an important intermediate, which often accumulates during failure of the wastewater treatment process (Kida et al. 1993; Ariesyady et al. 2007a). Propionate degradation, is thermodynamically unfavourable unless hydrogen and formate levels are kept low by hydrogen / formate utilizing methanogens (McInerney et al. 2008). Several studies indicate that both hydrogen and formate are used as interspecies electron carriers (Stams and Dong, 1995; de Bok et al. 2002b), but the importance of hydrogen versus formate is still not completely understood (Stams et al. 2006; McInerney et al. 2008). Propionate degraders use hydrogenases and formate dehydrogenases to produce hydrogen and formate, which are taken up by methanogens and metabolized by hydrogenases and formate dehydrogenases (de Bok et al. 2002; Nölling and Reeve, 1997). The catalytic centres of these enzymes contain metal-binding cofactors that are essential for their functioning (Vorholt and Thauer, 2002). Formate dehydrogenases contain molybdenum or tungsten and hydrogenases contain nickel and iron or iron only (Vorholt and Thauer, 2002; Vignais and Billoud, 2007). The amino acid sequences of many formate dehydrogenases and hydrogenases contain a selenocysteine that is located in the reactive core of the folded protein (Heider and Böck, 1993). Previously it was shown that when molybdenum, tungsten and selenium were not supplied to a propionate-fed UASB reactor during 249 days, the microbial activity decreased (Worm et al. 2009). However, techniques to adequately measure molybdenum, tungsten and selenium limitation in UASB reactors are lacking. Metal analysis with an Inductively Coupled Plasma - Optical Emission Spectrometer (ICP-OES) and with specific methanogenic activity (SMA) tests with supplied molybdenum, tungsten and selenium were found to be unsuitable to detect the potential limitation of the microbial activity of the UASB sludge by these trace metals (Worm et al. 2009). Syntrophobacter spp. and hydrogen and Methanospirillum spp. are commonly present in UASB reactors (Worm et al. 2009; Roest et al. 2005; Ariesyady et al. 2007a). These microorganisms are able to degrade propionate in syntrophic coculture (Stams and Dong, 1995). When tungsten and molybdenum are depleted in such cocultures, formate dehydrogenase activity of Syntrophobacter fumaroxidans and Methanospirillum hungatei decreases (Plugge et al. 2009a). This indicates that formate dehydrogenase activity is regulated by tungsten and molybdenum in these syntrophic cocultures. The available genome sequences of S. fumaroxidans and M. hungatei allow the use of molecular techniques to study the regulation of formate dehydrogenases and hydrogenases at the transcriptional level. The genome ofS. fumaroxidans indicates the presence of four formate dehydrogenases and six hydrogenases (Müller et al. 2010). The genome of M. hungatei indicates the presence of five formate dehydrogenases and three hydrogenases (Chapter 3). The individual functions of the formate dehydrogenases (FDH’s) and hydrogenases (Hyd’s) in these microorganisms are examined by gene analyses and by transcription analyses with reverse transcription followed by quantitative PCR

96 CHAPTER 7

Table 1 RT-qPCR primers used to specifically determine fdh and hyd transcripts, and 16S rRNA of Syntrophobacter fumaroxidans and Methanospirillum hungatei in pure and syntrophic cocultures (Chapter 3), and of Syntrophobacter spp. and M. hungatei in UASB reactor sludge.

Syntrophobacter spp. Methanospirillum spp.

Gene Metal content Primer name Gene Metal content **) Primer name SF_fdh-fw MH_fdh1-fw fdh1 W and Se *) fdh1 W or Mo SF_fdh1-rv MH_fdh1-rv SF_fdh2-fw MH_fdh2-fw fdh2 W and Se *) fdh2 W or Mo SF_fdh2-rv MH_fdh2-rv SF_fdh3-fw MH_fdh3-fw fdh3 W or Mo fdh3 W or Mo SF_fdh3-rv MH_fdh3-rv W or Mo SF_fdh4-fw MH_fdh4-fw fdh4 fdh4 W or Mo and Se SF_fdh4-rv MH_fdh4-rv SF_fehyd-fw MH_fdh5-fw fehyd Fe fdh5 W or Mo SF_fehyd-rv MH_fdh5-rv SF_hyn-fw MH_ech-fw hyn Ni and Fe ech Ni and Fe SF_hyn-rv MH_ech-rv SF_frh-fw MH_frc-fw frh Ni and Fe frc Ni and Fe SF_frh-rv MH_frc-rv SF_hox-fw MH_mbh-fw hox Ni and Fe mbh Ni and Fe SF_hox-rv MH_mbh-rv SF_hdr-fw MH_16S-fw hdr Ni, Fe and Se 16SrRNA …. SF_hdr-rv MH_16S-rv SF_fnr-fw fnr Ni, Fe and Se SF_fnr-rv SF_16S-fw 16SrRNA …. SF_16S-rv *) FDH1 and -2 from S. fumaroxidans were experimentally determined as tungsten containing formate dehydrogenases (de Bok et al. 2003; Reda et al. 2008). **) The predicted metal content of encoding formate dehydrogenases and hydrogenases is listed (modified after Müller et al. 2010). Fe: Iron, Mo: Molybdenum, Ni: Nickel, Se: Selenium, W: Tungsten.

(RT-qPCR) of cells grown in pure cultures and in syntrophic cocultures (Müller et al. 2010; Chapter 3). The metal content of the formate dehydrogenases and hydrogenases of S. fumaroxidans and M. hungatei was predicted from the amino acid sequence or retrieved from enzyme characterization studies (Table 1) (Müller et al. in press; de Bok et al. 2003).

In this work we performed gene transcription analyses with RT-qPCR to get insight into interspecies formate and hydrogen transfer and transcriptional regulation of FDH’s and Hyd’s in Syntrophobacter and Methanospirillum in a propionate-fed UASB reactor. Additionally we determined the effect of the supply and deprivation of molybdenum, tungsten and selenium to the UASB reactor feed on the transcription of formate dehydrogenase and hydrogenase coding genes (fdh’s and hyd’s respectively) in Syntrophobacter and Methanospirillum.

97 Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in slugde deprived of Mo, W, and Se

Results Primer specificity The SF_16S primer pair perfectly aligned with 16S rRNA genes from Syntrophobacter spp. present in the sludge population and showed much higher (>500 fold) affinity for 16S rRNA genes from Syntrophobacter spp. than for those of other clones retrieved from the UASB reactor (Table 2A). The affinity difference with Chlorobium spp. was even higher (>10,000 fold). Furthermore, all SF_fdh and SF_hyd primer pairs showed low (<82%) sequence identity with any region within genomes of other microbes that affiliated closest to the clones that were present in the UASB reactor sludge (Table 3A). The MH_16S primer pair perfectly aligned with 16S rRNA genes from Methanospirillum spp., whereas it has low (≤ 71%) similarity with 16S rRNA genes from relatives of M. concilii, the only other detectable archaeal species (Table 2B). The MH_16S primer pair showed >106 fold higher affinity for 16S rRNA genes from Methanospirillum spp. than for those of Methanosarcina spp. (Table 2B). Furthermore, all MH_fdh and MH_

Table 2 Specificity of primer pairs, SF_16S from S. fumaroxidans (2A) and MH_16S from M. hungatei (2B) based on sequence similarity and primer affinity with 16S rRNA genes from archaeal and bacterial clones present in the UASB reactor sludge.

Table 2A Sequence Clone Closest cultured relative (similarity %) Accession no. identity % Affinity (%) B02 Syntrophobacter sulfatireducens (95) EU888826 100 99 B16 Syntrophobacter sulfatireducens (99) EU888828 100 100 B09 Smithella propionica (99) EU888819 93 0.188 B07 Spirochaeta caldaria (89) EU888825 91 0.093 B03 Clostridium orbiscindens (94) EU888823 91 < 0.001 B01 Pelotomaculum propionicicum (94) EU888820 88 < 0.001 B10 Pelotomaculum propionicicum (94) EU888821 88 < 0.001 B13 Pelotomaculum propionicicum (94) EU888822 88 0.001 B18 Chlorobium phaeobacteroides (82) EU888817 86 0.001 B14 Chlorobium phaeobacteroides (82) EU888816 86 < 0.001 B04 Thermoanaerobacter pseudethanolicus (96) EU888824 81 < 0.001 B12 Rubrobacter xylanophilus (82) EU888818 79 < 0.001

Table 2B Sequence Clone Closest cultured relative (similarity %) Accession no. Affinity (%) identity % A05 Methanospirillum hungatei (99) EU888809 100 100 A09 Methanospirillum hungatei (100) EU888814 100 73 A10 Methanospirillum hungatei (99) EU888808 100 46 A07 Methanospirillum hungatei (99) EU888810 100 40 A01 Methanosaeta concilii (99) EU888804 71 < 10-5 A06 Methanosaeta concilii (99) EU888805 71 < 10-5 A08 Methanosaeta concilii (99) EU888806 71 < 10-5 A03 Methanosaeta concilii (99) EU888811 71 < 10-5 A12 Methanosaeta concilii (99) EU888812 71 < 10-5 A11 Methanosaeta concilii (94) EU888815 66 < 10-5

98 CHAPTER 7

Table 3 RT-qPCR primer pairs were aligned against genomes of microbes that affiliated closest with the clones that were present in the UASB reactor. Listed are the percentages of sequence identity between the RT-qPCR primer pairs and the most identical part of the bacterial (A) and archaeal (B) genomes.

Table 3A SF_fdh1 SF_fdh2 SF_fdh3 SF_fdh4 SF_fehyd SF_hyn SF_frh SF_hox SF_hdr SF_fnr SF_16S

Syntrophobacter fumaroxidans MPOB 100 100 100 100 100 100 100 100 100 100 100 Chlorobium phaeobacteroides DSM 266 65 67 63 71 74 70 66 71 71 66 77 Chlorobium chlorochromatii CaD3 70 61 58 66 71 65 63 77 68 63 74 Chlorobium limicola DSM 245 65 69 58 63 71 68 71 66 71 82 67 Rubrobacter xylanophilus DSM 9941 68 67 57 61 77 68 60 77 76 58 84 Pelotomaculum thermopropionicum SI 67 70 65 58 71 71 60 80 70 66 75 Clostridium thermocellum ATCC 27405 68 69 55 61 68 76 63 69 68 63 65 Clostridium methylpentosum R2 38 44 35 37 50 38 37 40 44 37 79 Clostridium leptum 753T 49 44 38 39 53 41 42 46 44 39 65

Table 3B MH_fdh1 MH_fdh2 MH_fdh3 MH_fdh4 MH_fdh5 MH_ech MH_frc MH_mbh MH_16S

Methanospirillum hungatei FJ-1 100 100 100 100 100 100 100 100 100 Methanosaeta thermophila PT 60 64 68 63 66 68 60 58 77

hyd primer pairs showed less than 68% sequence identity with any region within the genomic sequence of Methanosaeta thermophila (Table 3). Fdh and hyd transcription in Syntrophobacter spp. The RT-qPCR detection limit was defined as the average of the relative transcription after 40 PCR cycles. For Syntrophobacter spp. in UASB granular sludge this was 2∙10-7 relative transcription units. Transcription levels of fdh’s and hyd’s varied between 10-4 and the detection limit. InSyntrophobacter spp. high transcription levels were observed for fdh1, -2 and –4, whereas fdh3 was not transcribed in detectable amounts (Figure 1A). Moreover, Syntrophobacter spp. transcribed all six hydrogenase coding genes (Figure 1B). Relative transcription levels of fehyd and fnr were close to the detection limit whereas transcription levels offrh and hox were comparable with those of fdh1, -2 and -4. Relative transcription levels of hyn and hdr were somewhat in between.

Fdh and hyd transcription in Methanospirillum spp. The transcription levels of all fdh’s and hyd’s of Methanospirillum spp. were clearly detectable (Figure 1C and D). The detection limit for M. hungatei RNA transcripts in UASB granular sludge was 10-8 relative transcription units. Relative transcription of fdh’s and hyd’s varied between 10-2 and 10-6. Fdh4 was the highest and fdh3 the lowest transcribed formate dehydrogenase coding gene in M. hungatei and frc was the highest transcribed hydrogenase coding gene.

99 Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in slugde deprived of Mo, W, and Se

fehyd fdh1 hyn fdh2 hox 1,E-4 fdh3 1,E-4 fdh4 hdr Detection limit fnr Detection limit 1,E-5 1,E-5

1,E-6 1,E-6 Relative transcription Relative transcription 1,E-7 1,E-7 0 100 200 300 0 100 200 300 Time (days) Time (days)

fdh1 fdh2 ech 1,E-2 fdh3 1,E-2 frc fdh4 fdh5 mbh 1,E-3 1,E-3

1,E-4 1,E-4

1,E-5 1,E-5 Relative transcription Relative transcription transcription Relative 1,E-6 1,E-6 0 100 200 300 0 100 200 300 Time (days) Time (days)

8 Figure 1. Transcription of fdh’s (A) and hyd’s (B) in Syntrophobacter spp. relative to 16S rRNA levels in ) this3 organism. Transcription of fdh’s(C) and hyds (D) in M. hungatei relative to 16S rRNA levels in this 6 *10

organism.-1 This gene transcription was measured during a UASB reactor run of 276 days.

4 Fdh and hyd transcription during the reactor run During2 the reactor run propionate was not degraded completely (Worm et al. 2009). VFA-COD (g*L VFA-COD During0 249 days, while no tungsten, molybdenum and selenium were supplied 0 100 200 300 to the UASB granularTime (days) sludge, decreasing trends of fdh1 and fdh2 transcription in Syntrophobacter spp. were observed (Figure 1A). Addition of molybdenum, tungsten and selenium from day 249 could not recover the decrease of fdh1 and fdh2 transcription

8000 (Figure 1A). Fdh4 transcription seemed rather stable during the reactor run. During the

7000 effluent 249 days of metal deprivation, also decreasing trends were observed for hyn, frh, hox 6000 influent and hdr in Syntrophobacter spp. (Figure 1B). After addition of molybdenum, tungsten 5000 and selenium, no increase of fdh1, -2 or hyn, frh, hox, hdr transcription was observed 4000 (Figure 1 A and B). Transcription levels offehyd-1 and fnr were too close to the detection 3000 limit to observe changes. 2000 Reactor performance During the 249 days of metal deprivation no decreasingUsed transcription trends 1000 were observed for fdh’s and hyd’s in M. hungatei (Figure 1C8000 andeffluent D). Even increasing 0 7000 10 40 81 123 180 231 258 276 trends were measured, especially for fdh3 and to a lesser extent6000 for fdh1, ech, frc, and mbh. Molybdenum, tungsten and selenium addition was accompanied5000 Total bar = Influentwith a slightly increased transcription offdh1 , -2, -3 and -4, whereas transcription4000 ofech , frc and mbh 3000 VFA conc VFA Vak was noteffluent significantly influent ef/1000 inf/1000affected and transcription of fdh5 slightly2000 decreased. 10 0 20 1527,646 1429,439 1,527646 1,429439 1000 40 20 60 1487,03 3247,952 1,48703 3,247952 81 60 101,5 2925,071 7223,634 2,925071 7,223634 0 123 101,5 144 228,8721 5505,437 0,228872 5,505437 180 144 208,5 1335,76 5273,225 1,33576 5,273225 10 40 81 123 180 231100 208,5 256,5 3673,09 6468,51 3,67309 6,46851 258 256,5 261 1399,447 5572,485 1,399447 5,572485 276 267 291 4118,284 5768,664 4,118284 5,768664 Days CHAPTER 7

Discussion Gene transcription analysis in UASB reactor sludge Previously, transcription of genes coding for formate dehydrogenases and hydrogenases in pure and co-cultures of S. fumaroxidans and M. hungatei was analyzed (Chapter 3). The same specific primers were used to analyze transcription of these genesin Syntrophobacter spp. and Methanospirillum spp. in UASB reactor sludge. The closest cultured relative of theSyntrophobacter strains present in the reactor is S. sulfatireducens (95 and 99% sequence identity with clone B2 and B16 respectively) although the second closest relative of the Syntrophobacter strains present in the reactor is S. fumaroxidans (92 and 95% sequence identity with clone B2 and B16 respectively) (Worm et al. 2009). The closest cultured relative of the Methanospirillum strain present in the reactor is M. hungatei (99% sequence identity). In fact primers have to distinguish between specific RNA (16S rRNA and gene transcripts) of Syntrophobacter spp. and Methanospirillum spp. on the one hand and RNA from any other microorganisms in the reactor on the other hand. 16S rRNA of Syntrophobacter spp. and Methanospirillum spp was used as reference RNA, thus high specificity of especially 16S RNA primers is necessary to measure representative gene transcription levels. Based on band intensities after denaturing gradient gel electrophoresis (DGGE) it was shown that Syntrophobacter spp. were the most dominant bacteria during the whole reactor run (Worm et al. 2009). The second dominant group of clones were those related to Chlorobium phaeobacteroides (82%) (Worm et al. 2009). SF_16S primers did not amplify 16S rRNA from the Chlorobium clone in the UASB reactor which confirms the specificity of SF_16S primers for Syntrophobacter spp. 16SrRNA. Other bacterial clones; Smithella propionica (99%) and Pelotomaculum thermopropionicum (94%) became more dominant in the last two samples that were taken from the UASB reactor (days 246 and 276). SF_16S primers did not amplify 16S rRNA from these clones in significant amounts which further confirms the specificity of SF_16S primers for Syntrophobacter spp. 16SrRNA genes. The primers targeting specific fdh and hyd transcripts of Syntrophobacter spp. most likely also do not bind cDNA retrieved from Chlorobium spp. and Pelotomaculum spp. gene transcripts in the UASB reactor, because sequence identity with the genomes of the most related Chlorobium spp. and the only sequenced Pelotomaculum spp. is low. The only archaeal species other than M. hungatei (99%) found in the UASB reactor was the acetate degrading Methanosaeta concilii (94-99%) (Worm et al. 2009). MH_16S primers did not amplify 16S rRNA from these M. concilii clones (Table 2B) which confirms the specificity of MH_16S primers for Methanospirillum spp. 16SrRNA genes. The primers targeting specificfdh and hyd transcripts most likely also do not bind these clones because sequence identity with the genome of only sequenced Methanosaeta spp. is low. FDH’s and Hyd’s of Syntrophobacter spp. in UASB reactor sludge. The fact that fdh’s and hyd’s were transcribed in Syntrophobacter spp. and Methanospirillum spp., not only in defined cocultures as described in Chapter 3, but also in UASB reactor sludge shows that hydrogen and formate are also electron carriers

101 Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in slugde deprived of Mo, W, and Se

in granular UASB reactor sludge. Especially FDH1 and FDH2 are important in S. fumaroxidans grown in syntrophic coculture with M. hungatei (de Bok et al. 2003) which was also observed in Syntrophobacter spp. in syntrophic relation with Methanospirillum spp. in the UASB reactor. FDH3 and -4 seem important only when the energy shared by M. hungatei and S. fumaroxidans in defined syntrophic cocultures goes mainly to M. hungatei, and less energy remains for S. fumaroxidans (Chapter 3). Methanogenic activity decreased during the UASB reactor run (Worm et al. 2009) which coincided with fdh4 transcription in Syntrophobacter spp., whereas fdh3 was not transcribed at all. PCR with SFfdh3 primers and total DNA retrieved from the UASB reactor amplicons were produced (data not shown). This confirms that Syntrophobacter spp. in the UASB reactor contain the gene fdh3, but apparently this gene was not transcribed. During propionate degradation Syntrophobacter spp. generate reduced ferredoxin and NADH (Stams and Plugge, 2009). FeHyd, Hox and FDH1 contain NADH binding motifs and are candidates for the confurcating function of simultaneous ferredoxin and NADH oxidation with proton or carbon dioxide reduction (Müller et al. 2010; Chapter 3). Especially Fehyd and FDH1 were important in defined cocultures ofS. fumaroxidans and M. hungatei (Chapter 3). In Syntrophobacter spp. present in the reactor sludge, Hox and FDH1 were the most important putative confurcating enzymes whereas transcription of the gene coding for Fehyd was down-regulated. The periplasmic Hyn was transcriptionally down-regulated during syntrophic growth with methangens (Worm et al. 2009). Why transcription of hyn in Syntrophobacter spp. decreased in the bio- reactor remains unclear. Frh, Hdr and Fnr are likely cytoplasmic ferredoxin dependent hydrogenases, whose function is to provide an alternative route for the re-oxidation of reducing equivalents generated during propionate degradation. Their importance is similar in S. fumaroxidans grown in pure culture with M. hungatei as in Syntrophobacter spp. during the first half of the UASB reactor run. Both genomes ofSyntrophobacter spp. and Methanospirillum spp. contain a high number of fdh’s and hyd’s, likely for back-up possibilities when less energy is available for growth representing syntrophic flexibility.

Roles of FDH’s and Hyd’s from Methanospirillum spp. in UASB reactor sludge. The relative importance of most hydrogenase and formate dehydrogenase coding genes in Methanospirillum spp. in the UASB reactor sludge was similar as in M. hungatei when grown in syntrophic coculture with S. fumaroxidans (Chapter 3). However, in Methanospirillum spp. in UASB reactor sludge, FDH4 was the most important

cytoplasmic cofactor F420 binding formate dehydrogenase, whereas in M. hungatei grown in suspended coculture with S. fumaroxidans, the most important cytoplasmic

cofactor F420 binding formate dehydrogenase was FDH1 (Chapter 3). Our results indicate that decreased formate and hydrogen production by Syntrophobacter spp. triggered fdh3 transcription in Methanospirillum spp. in order to maintain sufficient formate concentrations for methanogenesis. Similarly, decreased hydrogen concentrations led to increased fdh transcription in Methanothermobacter thermoautotrophicus (Nölling and Reeve, 1997) and Methanococcus maripaludis (Wood et al. 2003).

102 CHAPTER 7

The influence of metal depletion on fdh and hyd transcription Nickel deprivation in a methanol-fed UASB reactor caused a decreased Methanosarcina population coinciding with a decrease in methanol degradation (Fermoso et al. 2008). Similarly, molybdenum, tungsten and selenium deprivation in the propionate-fed UASB reactor likely caused decreased fdh and hyd transcription in the dominant propionate degraders; Syntrophobacter spp. resulting in a decreased propionate degradation (Worm et al. 2009). In several bacteria the presence of metals transcriptionally up-regulates metal- containing enzymes. In Desulfovibrio vulgaris nickel, iron and selenium regulate the transcription of genes coding for [FeFe]-hydrogenase, [NiFe]-hydrogenase and [NiFeSe]-hydrogenase (Valente et al. 2006). In Rhizobium leguminosarum bv. viciae nickel regulates transcription of genes coding for nickel containing hydrogenases (Brito et al. 2008). In many bacteria, molybdenum and tungsten regulate the transcription of genes coding for molybdenum or tungsten containing cofactors via RNA structures called riboswitches (Regulski et al. 2008). By RNA loop prediction with Mfold version 3.2 riboswitches in the up-stream regions of fdh’s in S. fumaroxidans could not be predicted (Mathews et al. 1999; Zuker, 2003; IMG-JGI-DOE, 2009). However molybdenum and tungsten might regulate fdh transcription in theSyntrophobacter spp. via transcriptional regulators such as the putative periplasmic DNA-binding transcriptional regulator of molybdate metabolism, encoded by Sfum_3707 (IMG-JGI-DOE, 2009). Although formate dehydrogenase activity in M. hungatei decreased when molybdenum and tungsten were depleted in syntrophic cocultures with S. fumaroxidans (Plugge et al. 2009a), fdh transcription did not decrease inMethanospirillum spp. in the UASB reactor during the 249 days depletion of these metals. This could be explained by the fact that metal limitation is easier obtained in suspended cocultures than in densely packed granules because complex metal ligands and metal precipitates can be present in the granular matrix (Zandvoort et al. 2006). Moreover, the Methanospirillum spp. may use metal storage proteins that are only effective enough in granular growth. In Azotobacter vinelandii molybdenum storage proteins that guarantee molybdenum dependent nitrogen fixation at molybdenum starvation have been described (Fenske et al. 2005). From day 246 onwards other propionate degraders, Pelotomaculum and Smithella spp. competed with Syntrophobacter spp. (Worm et al. 2009). When at day 249 molybdenum, tungsten and selenium were supplied to the reactor feed (Worm et al. 2009), fdh transcription inSyntrophobacter spp. was not increased. This may have been due to the competition with Pelotomaculum and Smithella spp. The latter bacteria likely use different formate dehydrogenase mechanisms than Syntrophobacter spp. because Pelotomaculum spp. are gram positive and phylogenetically distant from Syntrophobacter spp. (de Bok et al. 2005), and Smithella spp. use a different pathway to degrade propionate compared to Pelotomaculum and Syntrophobacter spp. (Liu et al. 1999). When Mo, W and Se oxianions were added, Pelotomaculum, Smithella and Methanospirillum spp. were active enough to increase fdh transcription in Methanospirillum spp., but not active enough to prevent a decrease in propionate degradation.

103 Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. in slugde deprived of Mo, W, and Se

RT-qPCR is a technique that can give information on the active processes in a UASB reactor and we made a contribution to predict trace element limitation and malfunction in UASB reactors.

Experimental procedures RNA extraction and RT-qPCR The UASB reactor was operated as described previously (Worm et al. 2009). In order to perform reverse transcription followed by RT-qPCR, RNA was extracted from UASB reactor samples taken on operational day 40, 81, 123, 180, 231, 246 and 276. Of each sample three RNA extractions were performed. Granules were separated from the reactor medium, incubated overnight at 4 oC in RNA stabilization solution “RNAlater” from Ambion, and stored at –20 °C. For RNA isolation, 50 mg (wet weight) granules were resuspended in 125 µL RNase free water and 375 µL TRIzol Reagent (Invitrogen). This was transferred to a 0.1 mm silica beads containing tube and homogenized 4.5 m.s-1 for 30 s in a FastPrep-24 (MP Biomedicals). Tubes were incubated at 20 °C for 5 min and 0.1 mL chloroform was added followed by 15 s manual shaking. After incubation at 20 °C for 5 min, the suspension was centrifuged 9500 × g at 4°C for 15 min. The aqueous phase with an equal amount of 70% ethanol was transferred to an RNeasy mini spin column (Qiagen) and on-column DNase digestion with RNase-freeDNase I (Qiagen) was performed according to manufacturers instructions. RNA was ultimately eluted in 60 µL RNase free water and RNA integrity was confirmed by an Experion RNA StdSens Chip (Bio-Rad) using the Bio-Rad Experion system. Copy DNA (cDNA) was synthesized by SuperScript III reverse transcriptase (Invitrogen) according to manufacturers instructions and random decamers (Ambion). To distinguish genomic DNA from cDNA in qPCR, RT minus controls were prepared in a similar reaction mixture in which SuperScript III was replaced by water. qPCR was performed with the method and specific primers that were described previously (Chapter 3).

Primer specificity SF_16S and MH_16S primer pairs (Table 2) were aligned against 12 bacterial and 10 archaeal 16S rRNA genes that were previously determined in the UASB reactor (Worm et al. 2009) with Clustal X 1.81 software. In order to verify primer specificity experimentally, these 16S rRNA genes were amplified in conventional PCRs. Equal amplicon amounts were used as template in RT-qPCR with the SF_16S or MH_16S primer pair and experimental conditions as described above. Primer affinity percentages were calculated by comparison of transcription levels of 16S rRNA genes of clones with the transcription level of 16S rRNA genes of clone B16 (Syntrophobacter sulfatireducens, 99%) in case of amplification with SF_16S, or with the transcription level of 16S rRNA genes of clone A05 (Methanospirillum hungatei, 99%) in case of amplification with MH_16S. With NCBI-BLAST, all SF_fdh and SF_hyd primer pairs (Table 3A) were aligned against the genomes from microbes that affiliate closest with clones determined in the UASB reactor (Worm et al. 2009): Chlorobium phaeobacteroides (Genbank no. CP000492), Chlorobium chlorochromatii(CP000108), Chlorobium limicola (CP001097), Rubrobacter

104 CHAPTER 7 xylanophilus (CP000386), Pelotomaculum thermopropionicum (AP009389), Clostridium thermocellum (CP000568), Clostridium methylpentosum (draft sequence) and Clostridium leptum (draft sequence). MH_fdh and MH_hyd primer pairs (Table 1B) were aligned with the genome of Methanosaeta thermophila (CP000477).

Acknowledgements The authors were supported by the Research Council for Earth and Life Sciences (ALW) with financial aid from the Netherlands Organization for Scientific Research (NWO). This research was further co-funded by the Marie Curie Excellence Grant “Novel biological engineering processes for heavy metal removal and recovery” (MEXT-CT- 2003-509567).

105 106 CHAPTER 8

GENERAL DISCUSSION

Molecular mechanisms involved in interspecies electron transfer The interrelationship of different trophic groups within methanogenic microbial communities is an important subject of study. In methanogenic environments, microorganisms cooperate in a complex process with minimum increments of energy to sustain life. These energy increments are at the lowermost range of energy that can be converted into ATP at all, and with this, these microorganisms are interesting model subjects to study energy starvation on a broader basis. The question arises why nature designed methanogenic degradation in such a modular structure instead of having few types of microorganisms which could convert polymeric substrates all the way down to methane plus CO2. Theoretical considerations suggested that metabolic pathways can be efficient only up to a limited length of reaction chains (Costa et al. 2006). The main bottle neck within the anaerobic degradation reaction chain isthe syntrophic oxidation of propionate and butyrate in which interspecies electron transfer plays an important role. That hydrogen is an interspecies electron carrier is accepted for a long time. Although evidence that formate also plays an important role increased (Stams and Dong 1995; de Bok et al. 2002a), formate transfer is not yet commonly accepted. In this thesis we showed that both hydrogenases and formate dehydrogenases are involved in each electron generating step of the methylmalonyl- CoA pathway of propionate conversion. This shows that hydrogen and formate are both acting as interspecies electron carriers betweenS. fumaroxidans and M. hungatei. With bioinformatic tools we analyzedfdh ´s and hyd´s of other recently sequenced propionate degraders; P. thermopropionicum (Imachi et al. 2002; Kosaka et al. 2008) and butyrate degraders; S. wolfei (IMG-JGI-DOE) and S. aciditrophicus (McInerney et al. 2007) and found similarities with electron carrier systems in S. fumaroxidans (Chapter 2). This indicates that hydrogen and formate both are widely used interspecies electron carriers in syntrophic associations. The flexibility in the syntrophic association of S. fumaroxidans and M. hungatei became clear when they were grown in three separate cocultures and, for unknown reasons, different gene transcription profiles were observed. We found that in one of these cocultures, M. hungatei retrieves more free energy which resulted in less free energy available for S. fumaroxidans. The response of S. fumaroxidans was that it up- regulated transcription of genes coding for additional confurcating and periplasmic electron transferring enzymes and even additional electron transferring routes via the Rnf-complex and ferredoxin dependent hydrogenases and formate dehydrogenases.

107 General discussion

The Rnf-complex utilizes the proton motive force to oxidize NADH coupled to ferredoxin reduction which is followed by ferredoxin oxidation by the up-regulated formate dehydrogenases and hydrogenases. In other syntrophic organisms that contain an rnf gene cluster, such as S. acidotrophicus, the Rnf-complex may utilize proton motive force to reduce the ferredoxin that is generated by the confurcating [FeFe]-hydrogenase or formate dehydrogenase. Syntrophic microorganisms lacking the rnf-cluster, such as P. thermopropionicum might be syntrophicaly less flexible or use other strategies to survive energetically difficult situations.

Application: methanogenesis in bioreactors Methanogenesis in bioreactors is a sustainable technology to produce biogas from organic waste. More than 80 % of the chemical energy in organic waste components is conserved as methane, which in aerobic conversion would have been lost. Anaerobic digestion is applied worldwide to treat organic waste and wastewater from different sources. Presently, much research is done to replace fossil fuels with alternative

sustainable (CO2-neutral) energy sources. Microbial methane formation from waste and wastewater will contribute to this development. From the technological viewpoint it will be important to produce methane at a high rate and to convert all degradable organic compounds to biogas. The proper functioning and structuring of syntrophic communities of anaerobic bacteria and archaea will be important in this respect. A better understanding of the fundamental microbial processes will improve current anaerobic digesters, and increase application possibilities (O’Flaherty et al. 2006). Changes in the environmental conditions often causes partial uncoupling of the different processes, which results in the accumulation of organic acids (such as propionate and butyrate) (Plugge et al. 2009b). For instance changes in trace metal concentrations can be critical. Hydrogenases and formate dehydrogenases are required for hydrogen and formate transfer. Active sites of hydrogenases contain nickel and iron or iron only and in some cases selenium. Active sites of formate dehydrogenases contain tungsten, molybdenum and in some cases selenium. It has previously been shown that molybdenum and tungsten are important for formate dehydrogenase activity in propionate degrading cocultures of S. fumaroxidans and M. hungatei (Plugge et al. 2009a). Furthermore, Lebuhn et al. (2008) indicated that for long-term mono-digestion of maize silage, molybdenum and selenium should be supplied. In this thesis we examined the propionate degrading community in sludge from a UASB reactor that treats wastewater from the alcohol- producing factory NEDALCO in Breda (the Netherlands). We found that this community comprised propionate degrading Syntrophobacter spp., the hydrogen and formate consuming Methanospirillum hungatei and the acetate consuming Methanosaeta concilii. During a long-term absence of molybdenum, tungsten and selenium from the feed of the UASB reactor, the methanogenic activity decreased and relatives of Smithella propionica and Pelotomaculum spp. competed with Syntrophobacter spp. for propionate consumption. However, this could not prevent that the reactor stopped degrading propionate. Changes in the propionate degrading community thus indicate that problems in propionate degradation are about to occur. Determination of 16S rRNA coding genes of specific species with DGGE or qPCR could be a good approach to control

108 CHAPTER 8 a bioreactor. Measurements for metal concentrations in the sludge and specific methanogenic activity tests with supplied molybdenum, tungsten and selenium were not suitable to detect the potential limitation of the microbial activity of the UASB sludge by these trace metals. We developed an RT qPCR method with primers specific for fdh’s and hyd’s of both S. fumaroxidans and M. hungatei. We found that transcription of fdh’s and hyd’s in Syntrophobacter spp. decreased when molybdenum, tungsten and selenium were depleted from the reactor feed. M. hungatei might contain metal storage proteins, because in this methanogen transcription of fdh’s and hyd’s was stable and increased after supplementation of molybdenum, tungsten and selenium. Thus, an RT qPCR approach that targets copied transcripts of fdh’s and hyd’s of Syntrophobacter spp. can predict molybdenum, tungsten and selenium limitation and resulting UASB reactor performance problems. The design of RT qPCR primers was based on the genomic sequence of S. fumaroxidans. We proved that these primers combined with the applied PCR conditions specifically distinguished copied transcripts of other Syntrophobacter species from the rest of the RNA in the bioreactor. This shows that the RT qPCR approach can be very valuable to study the function of specific strains in complex microbial environments in the future, especially because nowadays the number of genomes sequenced increases steadily.

Research requirements and new questions. • The enzyme complexes in S. fumaroxidans that are claimed to confurcate NADH and ferredoxin for hydrogen or formate production (FeHyd, Hox and FDH1), should be purified and assayed for this activity as described by Schut et al. (2009). Although isolation might be laborious with these anaerobic slow growers, purification protocols are available for FDH1 (de Bok et al. 2003) and FeHyd (de Bok 2002b). We found that S. fumaroxidans always transcribes fehyd in high levels, whereas hox transcription is variable.Syntrophobacter spp. in a propionate fed UASB bioreactor, however, transcribed hox in high levels instead of fehyd. If a genetic system for S. fumaroxidans has been developed, it would be very interesting to make hox- and fehyd- mutants to find out if these enzymes can compensate each other. If they do, one can try to grow the hox- mutant under nickel-limiting conditions to investigate if hydrogenase transcription is metal dependent as was shown for hydrogenases in Desulfovibrio vulgaris Hildenborough (Valente et al. 2006). Another interesting topic for further research is the function of the Rnf-complex. The most important question is whether the Rnf-complex can translocate ions. This was never shown experimentally in any microorganism containing an rnf gene cluster. A possible way could be to isolate the putative ion-translocating membrane subunit and use membrane vesicles to study ion transport. • Succinate dehydrogenase and fumarate reductase activities were measured with the reversible Sdh. However, the second candidate; Frd has never been purified and as such could not be tested for its activity. Although transcription of frdABEF was up-regulated when fumarate was the electron acceptor, biochemical assays need to be performed to determine its Sdh, Frd and LASPO activity. Furthermore, the

109 General discussion

binding of cytochromes and cytochrome bd:quinone oxidoreductases to Frd, FDH1 and Hyn, would give more experimental evidence for the proposed mechanism. One could perform on-column affinity experiments to verify binding capacities of the proteins involved. • The first question that arises with respect to syntrophic flexibility is whether the increased amount of 16S rRNA from M. hungatei was due to the increased cell number or to an increased cell length or activity. This can be done by isolation of DNA from the three cocultures and to determine the ratio of S. fumaroxidans / M. hungatei 16S rRNA genes. • The genome of S. fumaroxidans contains operons that show similarity with those coding for butyryl-CoA dehydrogenase/energy-transferring flavoproteins (Bcd/ Etf) in Clostridium kluyveri. In this organism Bcd/Etf catalyses NADH oxidation with ferredoxin coupled to the reduction of crotonyl-CoA to butyryl-CoA (Li et al. 2008). The function of a Bcd/Etf system in S. fumaroxidans is unclear because S. fumaroxidans cannot grow on butyrate or crotonate. • Several hydrogenase and FHL coding operons contain long open reading frames that were annotated as heterodisulfide reductase, whereas the function of such enzymes is only known in methanogenesis. Whether these are remains of former heterologous gene transfer or code proteins with a function in S. fumaroxidans deserves further investigation. • There are indications that pili coding genes (Sfum_1127, Sfum_1129, Sfum_3881, Sfum_3882, Sfum_0117, and Sfum_0126) are up-regulated when S. fumaroxidans was grown in syntrophic association (data not shown). The question is whether these pili are used for attachment or function as conductivenanowires for interspecies electron transfer (Gorby et al. 2006). • The number of c-subunits in the ATP synthase of S. fumaroxidans is not known. The maximum number of protons translocated for each ATP synthesized or hydrolyzed is determined by the number of c-subunits (Nakanishi-Matsui and Futai 2008). In the genome of S. fumaroxidans genes coding for the cytoplasmic F1 part of the complex (Sfum_2587-Sfum_2581) are genomically separated from those coding for the membrane integrated F0 part of the complex (Sfum_1604-Sfum_1605). Sfum_1604 encodes the c-subunit. The c-subunit to ATP synthase ratio would give insight in the minimum energy increment that can be conserved by S. fumaroxidans and other syntrophic bacteria.

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121 Summary

SUMMARY

Many places on earth are without oxygen (anaerobic) such as rice paddy fields, swamps and sediments of freshwater lakes and oceans. When oxygen, nitrate or other electron acceptors are not present, organic material is degraded to carbon dioxide and methane by mixed microbial species that each have their own specific function in degradation. Anaerobic microbial communities are used in anaerobic digesters all over the world to treat organic waste and wastewater. Propionate is one of the most important intermediates in anaerobic digestion. It can only be degraded by propionate oxidizing bacteria when methanogenic archaea keep the concentration of the interspecies electron carriers, hydrogen and formate, low. However, little is known about the molecular mechanism of hydrogen and formate transfer. Hydrogenases are involved in hydrogen transfer and require Fe, Ni and/or Se for catalysis. Formate dehydrogenases that are involved in formate transfer require the trace metals W or Mo and in some cases Se for catalysis. However, the effect of W, Mo and Se limitation on the propionate degrading community of a UASB reactor and the transcription of formate dehydrogenase and hydrogenase encoding genes in this community was never examined. This would give more insight in formate transfer in the propionate degrading community of the UASB reactor and provide a method to study depletion of these metals in the reactor sludge. We used the genome sequences of the propionate degrading Syntrophobacter fumaroxidans and its syntrophic methanogenic partner, Methanospirillum hungatei to study molecular mechanisms of hydrogen and formate transfer in syntrophic cocultures and UASB reactor sludge, by gene analysis and molecular techniques. Gene analysis and microarray data determined formate dehydrogenase and hydrogenase encoding gene clusters in S. fumaroxidans and M. hungatei (Chapter 4). When S. fumaroxidans oxidizes propionate, reducing equivalents are generated by

three intermediate reactions in the form of FADH2, NADH and reduced ferredoxin. We found by gene analysis (Chapter 2) and RT qPCR (Chapter 3) that the genes coding for four formate dehydrogenases, six hydrogenases and one formate hydrogen lyase of S. fumaroxidans and five formate dehydrogenases and three hydrogenases ofM. hungatei were all transcribed during syntrophic and axenic growth. However, the transcription levels were dependent on the growth condition. Comparison of transcription levels also revealed that electrons from ferredoxin and NADH are simultaneously confurcated for hydrogen production by a cytoplasmic [FeFe]-hydrogenase. Moreover, results indicated that during syntrophic growth electrons from ferredoxin and NADH are confurcated to formate via a cytoplasmic formate dehydrogenase (FDH1). During syntrophic growth,

the electrons generated at the level of FADH2, travel via a cytoplasmic oriented succinate dehydrogenase, menaquinones, cytochrome b and c to the periplasmic formate dehydrogenase (FDH2) (Chapter 5). When S. fumaroxidans is grown in pure culture with alternative electron acceptors such as sulfate and fumarate, electrons flow partly to FDH2, and partly to the periplasmic hydrogenase (Hyn). The energy gained from propionate conversion to methane, acetate, and carbon dioxide has to be shared by S. fumaroxidans and M. hungatei. When M. hungatei takes

122 Summary more energy, less energy remains for S. fumaroxidans. In this situationS. fumaroxidans up-regulates transcription of genes coding for an additional cytoplasmic confurcating hydrogenase (Hox) and the periplasmic hydrogenase (Hyn) that is coupled to succinate oxidation. In addition, S. fumaroxidans induces transcription of genes coding for the Rnf-complex and ferredoxin dependent hydrogenases and formate dehydrogenases. This provides the possibility to use the membrane potential for the energy dependent coupling of ferredoxin reduction to NADH oxidation. The designed RT qPCR primers were used in UASB reactor sludge from the alcohol distillery NEDALCO in Bergen op Zoom (Netherlands) to investigate the effect of trace elements depletion. A lab-scale UASB reactor was fed with propionate and synthetic medium without added W, Mo and Se. During the reactor run, Syntrophobacter spp. were the dominant propionate-oxidizers and M. hungatei the dominant hydrogen and formate using methanogen. However, when propionate degradation decreased, two other propionate-oxidizers; Pelotomaculum propionicicum and Smithella propionica became abundant (Chapter 6). RT qPCR showed that in this reactor run the transcription of genes coding for formate dehydrogenases and hydrogenases in S. fumaroxidans decreased while transcription of genes coding for formate dehydrogenases and hydrogenases in M. hungatei were more stable (Chapter 7). This research shows that RT qPCR is a fast technique that can give information on the active processes in a UASB reactor, and that trace element limitation and possible malfunctioning of UASB reactors can be predicted. With this PhD research we gained insight in the molecular mechanisms of hydrogen and formate transfer between S. fumaroxidans an M. hungatei in defined cocultures and in a propionate-fed UASB reactor. This contributes to the understanding of similar molecular mechanisms in other syntrophic microorganisms and may improve the performance of anaerobic digesters in the future.

123 References

SAMENVATTING

Veel plekken op aarde zijn zuurstofloos (anaëroob), zoals moerassen, rijstvelden of de sedimenten in diepe meren en oceanen. Als geen zuurstof, nitraat of andere electronenacceptoren aanwezig zijn, wordt organisch materiaal afgebroken tot kooldioxide en methaan door een mengsel van microbiële soorten die elk een eigen specifieke functie hebben in de afbraak. Wereldwijd worden anaërobe bioreactors gebruikt voor de reiniging van organisch afval en afvalwater. Propionaat is één van de meest belangrijkste tussenproducten in anaërobe afbraak. Het kan alleen worden afgebroken door propionaat-oxiderende bacteriën als methanogene archaea de concentratie van de electronenmediatoren, waterstof en formiaat, laag houden. Er is nog weinig bekend over de moleculaire mechanismen van waterstof en formiaat overdracht. Hydrogenases zijn nodig voor waterstof overdracht en hebben de spore metalen Fe, Ni en/of Se nodig voor katalyse. Formiaat dehydrogenases die betrokken zijn bij formiaat overdracht hebben de spore metalen W of Mo en in sommige gevallen Se nodig voor katalyse. Echter, het effect van W, Mo en Se limitatie op de propionaat afbrekende consortia in een UASB reactor en de transcriptie van genen die coderen voor formiaat dehydrogenases en hydrogenases in deze consortia, is nog niet eerder bestudeerd. Dit zou meer inzicht kunnen geven in formiaat overdracht in de propionaat afbrekende consortia van de UASB reactor en zou een methode kunnen opleveren om limitatie van deze metalen in de reactor materiaal te analyseren. We hebben gebruik gemaakt van de genoomsequenties van de propionaat- afbrekende Syntrophobacter fumaroxidans en de syntrofe methanogene partner, Methanospirillum hungatei om de moleculaire mechanismen van waterstof- en formiaatoverdracht in reincultures, syntrofe cultures en UASB reactor materiaal middels genanalyse en moleculaire technieken te bestuderen. Genclusters die coderen voor formiaat dehydrogenases en hydrogenases in S. fumaroxidans en M. hungatei zijn bepaald met behulp van genanalyse en microarray gegevens (Hoofdstuk 4). Als S. fumaroxidans propionaat oxideert worden reductie-equivalenten

geproduceerd in de vorm van FADH2, NADH en gereduceerd ferredoxine. Wij vonden met genanalyse (Hoofdstuk 2) en RT qPCR (Hoofdstuk 3) dat de genen die coderen voor vier formiaat dehydrogenases, zes hydrogenases en één formiaat waterstof lyase van S. fumaroxidans en vijf formiaat dehydrogenases en drie hydrogenases van M. hungatei allemaal afgeschreven worden in syntroof gegroeide cultures en reincultures. Echter, de hoeveelheid transcripten is afhankelijk van de groeiconditie. Het vergelijken van transcriptie niveaus toonde aan dat elektronen van ferredoxine en NADH tegelijk worden samengebracht (geconfurceerd) om waterstof te vormen met een cytoplasmatisch [FeFe]-hydrogenase. Verder wijzen onze resultaten erop dat tijdens sytrofe groei elektronen van ferredoxine en NADH geconfurceerd worden om formiaat te maken via een cytoplasmatisch formiaat dehydrogenase (FDH1). Tijdens syntrofe

groei gaan elektronen in de vorm van FADH2 via een cytoplasmatisch georiënteerd succinaat dehydrogenase, menaquinonen, cytochroom b en c naar het periplasmatisch formiaat dehydrogenase (FDH2) (Hoofdstuk 5). Als S. fumaroxidans wordt gegroeid

124 References in reincultures met externe elektronacceptoren, zoals sulfaat en fumaraat, gaan de elektronen gedeeltelijk naar FDH2 en gedeeltelijk naar het periplasmatisch hydrogenase (Hyn). De energie die verkregen wordt uit de omzetting van propionaat tot methaan, acetaat en kooldioxide, moet gedeeld worden door S. fumaroxidans en M. hungatei. Als M. hungatei meer energie wegneemt, blijft er minder energie over voorS. fumaroxidans. Dit verhoogt in S. fumaroxidans transcriptie van genen die coderen voor een van de andere cytoplasmatische confurcerende hydrogenases (Hox) en het periplasmatisch hydrogenase (Hyn) dat is gekoppeld aan succinaat oxidatie. Daarnaast induceert S. fumaroxidans genen die coderen voor het Rnf-complex en ferredoxine-afhankelijke hydrogenases en formiaat dehydrogenases. Dit zorgt voor de mogelijkheid om gebruik te maken de membraanpotentiaal die nodig is voor de energieafhankelijke koppeling van ferredoxin reductie en NADH oxidatie. De ontworpen RT qPCR primers zijn gebruikt om te onderzoeken wat het effect was van het niet toevoegen van spore elementen op UASB reactor materiaal van de alcohol distilleerderij NEDALCO in Bergen op Zoom. Een lab-schaal UASB reactor werd gevoed met propionaat en met synthetisch medium zonder toegevoegd W, Mo en Se. Tijdens de reactor run waren Syntrophobacter spp. de dominante propionaatoxideerders en was M. hungatei de dominante waterstof- en formiaat- gebruikende methanogeen. Echter, in de loop van de reactorrun verminderde de propionaatafbraak en begonnen twee andere propionaatoxideerders Pelotomaculum propionicicum en Smithella propionica te groeien. RT qPCR toonde aan dat in deze reactorrun de transcriptie van genen die coderen voor formiaat dehydrogenases en hydrogenases in S. fumaroxidans verminderde terwijl transcriptie van formiaat dehydrogenases en hydrogenases in M. hungatei stabiel bleef (Hoofdstuk 7). Dit onderzoek laat verder zien dat RT qPCR een snelle techniek is die informatie kan geven over de actieve processen in een UASB reactor en dat limitatie van spore elementen en het mogelijk slecht functioneren van UASB reactor kan worden voorspeld. Met dit promotieonderzoek hebben we inzicht gekregen in de moleculaire mechanismen van waterstof- en formiaatoverdracht tussen S. fumaroxidans en M. hungatei in gedefinieerde cocultures en in een UASB reactor. Hierdoor kunnen we vergelijkbare moleculaire mechanismen in andere syntrofe cultures beter begrijpen wat bijvoorbeeld kan bijdragen tot betere anaerobe bioreactors in de toekomst.

125 Sense certificate

126 127 CURRICULUM VITAE

Petra Worm was born on January 29th 1980 in Iserlohn (Germany). After four years of HAVO she started in September 1996 at the MLO (laboratory school) at the Technisch Lyceum Eindhoven. During her first internship, at the Institute for sugar beet research in Bergen op Zoom, she worked on identification and detection of the fungus Rhizoctonia solani under the supervision of dr. ir. Hans Schneider and Ellen Musters-van Oorschot. During her second internship, at Janssen Pharmaceutica in Beerse (Belgium) she screened components for signal transduction by glutamate receptors in hamster ovarium cells under the supervision of dr. Anne Lesage and Ria Wouters. After she received her MLO- diploma “Microbiology”, she started in September 2000 at the HLO (higher laboratory school) at Fontys Hogeschool Eindhoven. During her graduation project, at the Swiss Institute of Allergy and Asthma Research in Davos (Switserland) she worked onthe cloning and characterization of IgE-binding allergenes in wheat under supervision of Prof. Reto Crameri and dr. Michael Weichel. In July 2003 she received her Bachalor diploma “Biology and laboratory research”. In September 2003 she started with the masters Molecular Sciences at Wageningen University. During her graduation project, at the Laboratory of Microbiology, department of Bacterial Genetics at Wageningen University, she worked on the cloning and characterization of enzymes in the pentose pathway of Sulfolobus solfataricus under the supervision of Prof. John van der Oost and Dr. Stan Brouns. During her internship, at BioMérieux at the department Molecular diagnostics in Boxtel she worked on stabilizing the T7-RNA polymerase for optimization of Nucleic Acid Sequence Based Amplification under supervision of Prof. John van der Oost and Dr. Stan Brouns from Wageningen University and Dr. Erik Rump and Dr. Fokke Venema from BioMérieux. In September 2005 she graduated and she started her PhD at the Laboratory of Microbiology, department Microbial Physiology at Wageningen University. The research focussed on the gene analysis and transcription of formate dehydrogenases and hydrogenases in syntrophic propionate-oxidizing communities. Results of this research are described in this thesis.

128 PUBLICATIONS

Worm, P., Fermoso, F.G., Lens, P.N.L., Plugge, C.M. 2009. Decreased activity of a propionate degrading community in a UASB reactor fed with synthetic medium with molybdenum, tungsten and selenium. Enzyme Microb tech 45 139-145 Müller, N.*, Worm, P*., Schink, B., Plugge, C.M., Stams, A.J.M. Syntrophic butyrate and propionate oxidation processes: from genomes to reaction mechanisms. Environ Microbiol Rep. DOI: 10.1111/j.1758-2229.2010.00147.x. (*Both authors contributed equally). Worm, P., Muller, N., Plugge, C.M., Stams, A.J.M., and Schink, B. Syntrophy in methanogenic degradation, Book chapter J. Hackstein (in press). Worm, P., Stams, A.J.M., Cheng, X., Plugge, C.M. Transcription of formate dehydrogenase and hydrogenase coding genes in pure- and cocultures of Syntrophobacter fumaroxidans and Methanospirillum hungatei (Manuscript in preparation) Worm, P., Fermoso, F.G., Stams, A.J.M., Lens, P.N.L., Plugge, C.M. Transcription of fdh and hyd in Syntrophobacter spp. and Methanospirillum spp. Inside anaerobic granular sludge deprived of molybdenum, tungsten and selenium (Manuscript in preparation) Stams, A.J.M., Worm, P., Sousa, D.Z., Alves, M.M., Plugge, C.M. 2010. Syntrophic degradation of fatty acids by methanogenic communities. Book Patrick C. Hallenbeck, Microbial Technologies in Advanced Biofuels Production, Springer-project publication. Brouns, S.J., Barends, T.R., Worm, P., Akerboom, J., Turnbull, A.P., Salmon, L., van der Oost, J. 2008. Structural insight into substrate binding and catalysis of a novel 2-keto-3- deoxy-D-arabinonate dehydratase illustrates common mechanistic features of the FAH superfamily. J Mol Biol. 357-371 Brouns, S.J., Walther, J., Snijders, A.P., van de Werken, H.J., Willemen, H.L., Worm, P., de Vos, M.G., Andersson, A., Lundgren, M., Mazon, H.F., van den Heuvel, R.H., Nilsson, P., Salmon, L., de Vos, W.M., Wright, P.C., Bernander, R., van der Oost, J. 2006. Identification of the missing links in prokaryotic pentose oxidation pathways: evidence for enzyme recruitment. J Biol Chem. 27378-27388

129 DANKWOORD

Vanaf mijn tienerjaren droomde ik van “een eigen onderzoek doen”. Omdat ik geen briljante leerling was, was ik ervan overtuigd dat het bij een droom zou blijven. Maar door het krijgen van kansen, hard werken, en de hulp van velen is het uiteindelijk gekomen tot dit proefschrift. De afgelopen jaren heb ik enorm genoten van het werken in de wetenschap en daarom wil ik iedereen bedanken die daaraan heeft bijgedragen. Op de eerste plaats wil ik mijn dagelijks begeleider en tevens co-promotor Caroline bedanken voor alles. Bedankt voor de dagelijkse begeleiding en het meedenken in het onderzoek. Ik vond het fijn dat ik altijd bij je kon binnen lopen en dat we regelmatig besprekingen hadden. Verder heb je samenwerkingen binnen en buiten Wageningen gestimuleerd wat erg belangrijk is geweest voor het uiteindelijke resultaat. Ook wil ik je bedanken voor het schrijven van het projectvoorstel en ik hoop dat je met mijn bijdrage weer ideeën hebt voor nieuwe projectvoorstellen zodat na mij nog veel aio’s zullen volgen. Natuurlijk wil ik ook Fons mijn promotor bedanken. Ook jij nam altijd tijd voor besprekingen en voor het lezen van manuscripten. Bedankt voor je enthausiasme en je wetenschappelijk meedenken. Het was leuk om mede hierdoor tot nieuwe ideeën te komen over de flexibiliteit van syntrofie. Fernando Gonzalez Fermoso en Piet Lens, I want to thank you for our nice cooperation on the UASB reactor-work. Now, two thesis-chapters are the result of this work; one that has been published already and the other will be ready for submission soon. Dr. Nick Müller and Prof. Bernhard Schink from Konstanz University I would like to thank for our nice cooperation in writing a mini-review and a book-chapter together. Nick, thank you for the inspiring discussions over Skype about electron-transfer in butyrate- and propionate-oxidizing consortia, I very much enjoyed that. All the present and former people in the MicFys-group I want to thank for their nice company; Marjet, Alona, Farrakh, Rozelin, Jeanine, Melike, Heleen, Sander, Bart, Nuria, Miriam, Anne-Meint, Zumi, Frank, Wim van D, Sanne, Ineke, Diana, Flavia, Martin, Teun, Ton, Christian, Zahid, Elsemiek, Urania, Marcel, and everybody I forgot to mention. Also everybody else in Microbiology I want to thank. There was always a nice and helpful atmosphere in the lab. Outside the lab I very much enjoyed the (social) activities such as the yearly labtrips, the PhD trip to California (thank you organizers!), Veluwe-loop (bedankt Mathijs!) and the We-days (thank you Mark L, Mark M, Marco, Katrin, Marcel, Christian en John). It’s a pitty that we were not the number one winners that other year, I still think we deserved it. The SENSE-symposium ‘09 committee members Marjet, Teun, Alona en Urania and the SENSE education director Ad van Dommelen, thank you for the nice cooperation. Without your contributions the symposium wouldn’t have been such a success. Alona, thank you for our regular meetings. The discussions and your questions about interspecies electron transfer helped me to structure my thoughts and to find the gaps in my understanding. Now I wish you lots of luck with your PhD thesis. In the first years of the research it was very important to get suggestions and advise about working with mRNA and performing qPCR. This I got from Douwe and

130 Miguel, many thanks for that! Hans en Ineke wil ik bedanken voor de adviezen omtrent het maken van DNA libraries en het uitvoeren van DGGE. Dankzij jullie hebben deze experimenten vlotjes kunnen verlopen. Wim, als je het mij vraagt ben je redelijk onmisbaar bij Microbiologie. Bedankt voor het oplossen van problemen met computers, lab-apparatuur en andere technische zaken. My one and only student Xu Cheng I want to thank for his contribution to my research and I wish you lots of success in your own PhD research now. Natuurlijk wil ik ook Jannie, Nees, Francis, Anja, Hans en Ton bedanken voor het verzorgen van onmisbare dagelijkse dingen zoals het steriliseren en afwassen van glaswerk, regelen van bestellingen en alle administratieve zaken. Mijn vrienden Karin en Thomas, Geertje, Marcia en Jochem, Maureen, Berina en Maurice jullie bedankt voor jullie steun en voor de gezellige afleiding. Maureen, Berina en Maurice, het was fijn om aio-verhalen met jullie te delen. Mijn familie en schoonfamilie wil ik graag bedanken voor hun steun en interesse in mijn werk en hun begrip als ik geen tijd had voor feestjes of verjaardagen; Pap, Mam, Agnes, Linda en Dennis, Ans, Paul, Casper. Aimée en Wietse jullie heel veel succes met jullie eigen promotie onderzoek. Léon, jou steun was onmisbaar. Bedankt voor je interesse, geduld, je positieve kijk, en voor de praktische ondersteuning thuis. Het was ook leuk dat je af en toe koffie kwam drinken op het lab als je weer gas had geleverd bij Micro. Verder beloof ik dat je de weekenden niet meer met een hoofdtelefoon voor de televisie hoeft door te brengen omdat ik niet gestoord wil worden.

131 The research was supported by the Research Councils for Earth and Life Sciences (ALW) and Chemical Sciences (CW) with financial aid from the Netherlands Organization for Scientific Research (NWO).

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