Genomic, Proteomic, and Biochemical Analysis of the Organohalide Respiratory Pathway in Desulfitobacterium dehalogenans

Abstract

Desulfitobacterium dehalogenans is able to grow by organohalide respiration using 3-chloro-4-hydroxyphenyl acetate (Cl-OHPA) as an electron acceptor. We used a combination of genome sequencing, biochemical analysis of redox active components, and shotgun proteomics to study elements of the organohalide respiratory electron transport chain. The genome of Desulfitobacterium dehalogenans JW/IU-DC1^T consists of a single circular chromosome of 4,321,753 bp with a GC content of 44.97%. The genome contains 4,252 genes, including six rRNA operons and six predicted reductive dehalogenases. One of the reductive dehalogenases, CprA, is encoded by a well-characterized cprTKZEBACD gene cluster. Redox active components were identified in concentrated suspensions of cells grown on formate and Cl-OHPA or formate and fumarate, using electron paramagnetic resonance (EPR), visible spectroscopy, and high-performance liquid chromatography (HPLC) analysis of membrane extracts. In cell suspensions, these components were reduced upon addition of formate and oxidized after addition of Cl-OHPA, indicating involvement in organohalide respiration. Genome analysis revealed genes that likely encode the identified components of the electron transport chain from formate to fumarate or Cl-OHPA. Data presented here suggest that the first part of the electron transport chain from formate to fumarate or Cl-OHPA is shared. Electrons are channeled from an outward-facing formate dehydrogenase via menaquinones to a fumarate reductase located at the cytoplasmic face of the membrane. When Cl-OHPA is the terminal electron acceptor, electrons are transferred from menaquinones to outward-facing CprA, via an as-yet-unidentified membrane complex, and potentially an extracellular flavoprotein acting as an electron shuttle between the quinol dehydrogenase membrane complex and CprA.

INTRODUCTION

Microorganisms in anoxic environments have the ability to reductively dechlorinate highly chlorinated compounds such as tetrachloroethene (PCE), pentachlorophenol (PCP), and polychlorinated biphenyls (PCBs). Anaerobic bacteria from a phylogenetically broad range of genera, including Desulfitobacterium, Dehalobacter, Geobacter, Sulfurospirillum, and Dehalococcoides, couple reductive dechlorination of chlorinated organic compounds to energy conservation in a respiratory manner, previously referred to as (de)halorespiration and currently referred to as organohalide respiration (1–3). Formate and hydrogen are common electron donors in this process. Since these compounds do not allow substrate-level phosphorylation, energy is most likely conserved by the formation of a proton gradient across the cytoplasmic membrane (1).

The localization of the reductive dehalogenase (RD) and the topology of the electron transport pathway from electron donor to electron acceptor have been a matter of debate. In early models, it was suggested that the electron-donating reaction takes place outside the cell, after which electrons would be transported across the cell membrane to an intracellular RD, leading to formation of a proton gradient without vectorial proton translocation (4–6). In contrast with this suggestion, all RDs of the PceA/CprA type (tetrachloroethene/orthochlorophenol reductive dehalogenase) contain a twin arginine translocation (TAT) signal sequence, which is recognized by the TAT export system, leading to the export of folded proteins with incorporated cofactors across the cell membrane (3, 7). The current view is that the catalytic subunit of RDs is located at the outside of the cytoplasmic membrane (3, 8–11). If the electron-donating and -accepting reactions both occur at the same side of the cell membrane, a proton-translocating mechanism is necessary to establish a proton gradient that can drive ATP synthesis (12, 13). The electron transport chain from electron donor to RD has been proposed to involve both menaquinones and an intermediate electron carrier, as menaquinones are unable to directly reduce RDs (1, 14). These electron-transferring elements have, however, not yet been identified.

The low-GC Gram-positive Desulfitobacterium dehalogenans is able to use a range of terminal electron acceptors, including orthochlorophenols, sulfite, thiosulfate, sulfur, nitrate, and fumarate (15, 16). The catalytic subunit of the key enzyme in 3-chloro-4-hydroxyphenyl acetate (Cl-OHPA) dehalogenation, CprA, was purified and characterized as a corrinoid-containing Fe-S protein (17). Molecular analysis of the cpr gene cluster in D. dehalogenans led to the identification of genes encoding putative regulatory proteins and protein-folding catalysts, whose transcription was specifically induced under organohalide-respiring conditions (18). Research on the electron transport chain from formate to Cl-OHPA has been hampered thus far by the lack of D. dehalogenans whole-genome information. Previously, the genomes of Desulfitobacterium hafniense Y51 and DCB-2 have been published (9, 19). The number of published genomes from Desulfitobacterium spp. is expected to increase as the result of ongoing sequencing projects.

In this study, the genome of D. dehalogenans strain JW/IU-DC1^T was sequenced. The proteomes of cells grown with formate as an electron donor and fumarate or Cl-OHPA as an electron acceptor were compared with those of cells grown fermentatively on pyruvate. Involvement of redox active components in the transfer of electrons from formate to either Cl-OHPA or fumarate was investigated. The redox state of these components was analyzed during their reduction by formate and subsequent oxidation by Cl-OHPA or fumarate.

MATERIALS AND METHODS

Organism and growth conditions.

D. dehalogenans strain JW/IU-DC1^T was cultivated under anoxic conditions at 37°C under a 100% N₂ gas phase. Medium was prepared as described previously (17). Cultures for the enzyme assays were grown in 3-liter bottles containing 2 liters medium supplemented with either 20 mM formate and 20 mM Cl-OHPA (FC) or 20 mM formate and 20 mM fumarate (FF). Cultures for the proteomic analyses were grown in 250-ml bottles containing 100 ml medium supplemented with 20 mM formate and 20 mM Cl-OHPA or fumarate as electron donors/acceptors or with 40 mM pyruvate (P) for fermentative growth. Cultures used for genome sequencing were grown in 1-liter bottles containing 0.5 liter medium supplemented with 40 mM pyruvate.

Genome sequencing and analysis.

For genome sequencing, cells grown fermentatively on pyruvate were harvested from an early-stationary-phase culture by centrifugation and resuspended in TE buffer (10 mM Tris, 20 mM EDTA, pH 8), before DNA was extracted according to the cetyltrimethylammonium bromide (CTAB) protocol recommended by the DOE Joint Genome Institute (JGI) (20). The genome of D. dehalogenans, gold stamp Gc02348, was generated by the JGI using a combination of Illumina and 454 sequencing (21, 22).

Genes were identified using Prodigal (23) as part of the Oak Ridge National Laboratory genome annotation pipeline, followed by a round of manual curation using the JGI GenePRIMP pipeline (24). The predicted coding sequences (CDSs) were translated and used to search the National Center for Biotechnology Information (NCBI) nonredundant database and the UniProt, TIGRFam, Pfam, PRIAM, KEGG, COG, and InterPro databases. These data sources were combined to assert a product description for each predicted protein. Noncoding genes and miscellaneous features were predicted using tRNAscan-SE (25), RNAmmer (26), Rfam (27), TMHMM (28), SignalP (29), Pred-tat (30), and SMART (31). The Artemis v14 genome browser was used for visualizing the annotated genome (32).

Cell fractionation and enzyme assays.

Cells were harvested in an anoxic glove box with an N₂/H₂ gas phase in a 95/5% ratio. Late-exponential-phase cultures were harvested by 10 min of centrifugation at 16,000 × g and resuspended to a total volume of 2 ml in buffer A, containing 100 mM potassium phosphate (KPi), pH 7.5, and 1 mM dithiothreitol (DTT). A portion, set aside on ice, was used as a whole-cell suspension. Whole cells were permeabilized by incubation in the presence of 0.1% CTAB for 10 min at 4°C or 0.04% toluene when indicated. A few crystals of DNase I were added to a fraction of the whole-cell suspension, and cells were disrupted by 6 cycles of 30 s of sonication and 30 s of cooling on ice. Undisrupted cells were removed by 5 min of centrifugation at 20,000 × g. An aliquot of the cell extract was kept for enzyme assays. The remaining part was centrifuged for 90 min at 140,000 × g. The supernatant containing the soluble proteins was transferred to a glass vial and stored at 4°C. The pellet containing the membranes was resuspended in buffer A. Soluble and membrane fractions were stored at 4°C under a 100% N₂ gas phase.

The protein concentration of the extracts was determined with the microbiuret method with bovine serum albumin as a standard (33). Formate dehydrogenase (FDH), chlorophenol reductive dehalogenase (CPR), and fumarate reductase (FRD) activities in cell extracts were determined by spectroscopic recording of changes in the reduced methyl viologen concentration at 578 nm as described previously (34). Cl-OHPA and OHPA were measured by high-performance liquid chromatography (HPLC) as described elsewhere (34).

Quinone oxidation/reduction and extraction.

To investigate the possible involvement of menaquinones (MKs) in Cl-OHPA and fumarate respiration, concentrated cell suspensions (FC or FF) were incubated for 10 min in the presence of 10 mM formate. Dithionite was added to cell suspensions prior to addition of fumarate or Cl-OHPA, to ensure total reduction of quinones at the start of the experiment. The reduced MKs became oxidized by addition of Cl-OHPA (10 mM) to formate- and dithionite-reduced formate–Cl-OHPA-grown cells, or when fumarate (10 mM) was added to formate- and dithionite-reduced FF-grown cells. Quinones were extracted in methanol-petroleum ether and separated by HPLC as described previously (35). The detection of quinones was followed spectrophotometrically at 245 nm. The absorption spectrum of vitamin K was used as a control. Instability of (chemically prepared) menaquinol prevented reliable quantification of the quinol extinction coefficient. Therefore, the ratio between the peak areas representing reduced (MKH₂) or oxidized (MK) menaquinones in the HPLC chromatogram was used as an indicator of the degree of quinone reduction.

UV-visible spectroscopic analysis of cell suspensions.

Cultures (2 liters) of D. dehalogenans were grown with formate as an electron donor and either fumarate or Cl-OHPA as an electron acceptor. Cells were harvested and washed with 8 ml buffer containing 50 mM Tris-HCl, pH 8.0, and 0.5 mM DTT and resuspended in 50 mM Tris-HCl, pH 8.0, to a final concentration of 0.36 g (wet weight) of cells ml⁻¹ in 1-ml N₂-flushed cuvettes. All handling was done under anoxic conditions. Ninety microliters of the concentrated cell suspension was resuspended in 0.9 ml of 50 mM Tris-HCl (pH 8.0). After incubation with a reducing or oxidizing substance(s), the suspension was frozen in liquid N₂ and the spectrum (320 to 700 nm) was recorded at 77 K, on a DW-2a spectrophotometer (American Instrument Co.) as described in reference 36. After recording, the baselines were corrected for light scattering by subtracting a straight line between 540 and 580 nm. The corrected spectra were deconvoluted into two Gaussian peaks of variable width, one signal (at 552 nm) representing cytochrome c and another signal (at 561 nm) representing cytochrome b. The spectra were analyzed as described in reference 37. Peak areas were taken to be proportional to the reduction level of the cytochromes. Involvement of cytochromes in electron transfer from formate to fumarate or Cl-OHPA was investigated by recording the changes in their redox state after reduction of the cells with 1 mM formate and reoxidation by addition of 10 mM Cl-OHPA or fumarate.

Protein extraction and proteomic analysis.

Triplicate samples were prepared for proteomic analysis as described before (38), with the modification that the protein samples were run over only a short distance, approximately 1 cm, in the SDS-PAGE gel prior to in-gel trypsin digestion of proteins. This was done to be able to cut and process entire lanes as single samples. Detection, identification, and relative quantification of proteins were done as described before (38). In brief, trypsin-digested peptides were run on a Proxeon LTQ-Orbitrap XL nano-liquid chromatography mass spectrometer (nLC-MS), and all spectral data obtained were analyzed with MaxQuant v.1.3.0.5 (39) using a protein database generated from the D. dehalogenans genome. Filtering and further bioinformatic analysis of the MaxQuant/Andromeda workflow output and the analysis of the abundances of the identified proteins were performed with the PERSEUS v.1.3 module (available at the MaxQuant suite). Only proteins showing a false discovery rate (FDR) of less than 1% were accepted. Reversed hits were deleted from the MaxQuant result table as well as all proteins showing fewer than 2 peptides or fewer than 1 unique peptide or with label-free quantification (LFQ) values of 0 for both sample and control. Zero values for one of the log LFQ columns were replaced by a value of 5.5 (just below the lowest measured value) to make sensible ratio calculations possible. Relative protein quantification of sample to control was conducted with PERSEUS by applying a two-sample t test using the “log LFQ intensity” columns obtained with an FDR threshold set to 0.05 and S0 = 1. S0 = 1 indicates that both the average protein abundance ratio and the t test P value have equal weight to decide whether the protein is significantly different between the two different growth conditions. Whether a protein concentration changed significantly was also determined with PERSEUS using t test P value and the average protein abundance ratio between the different growth conditions.

Nucleotide sequence accession number.

The annotated genome sequence has been deposited under GenBank accession number CP003348.

RESULTS

Genome analysis of the orthochlorophenol-respiring D. dehalogenans.

The genome of D. dehalogenans is 4.3 Mbp with a G+C content of 45% and encodes six predicted reductive dehalogenases. The general genome characteristics were determined and compared to the two published Desulfitobacterium hafniense genomes (Table 1) (9, 19). We focused our initial genomic analysis on genes coding for enzymes of metabolic pathways expected to be active under the growth conditions tested, including respiration with formate as an electron donor and fumarate or Cl-OHPA as an electron acceptor (Table 2).

TABLE 1.

General genome features of D. dehalogenans and D. hafniense DCB-2 and Y51

Strain	Size (Mbp)	GC (%)	No. of:
			Total predicted genes	Predicted CDSs	16S rRNA-encoding genes	Predicted Rdha-encoding genes
D. dehalogenans JW/IU-DC1T	4.3	45	4,252	4,142	6	6
D. hafniense DCB-2T	5.3	48	5,042	4,953	5	7
D. hafniense Y51	5.7	47	5,208	5,060	6	1

^aRdh, reductive dehalogenase.

TABLE 2.

Selected proteins potentially involved in the respiration of D. dehalogenans using formate as an electron donor and either fumarate or Cl-OHPA as an electron acceptor^g

Gene	Locus taga			Annotation	TMHb	S. Sq.c
	D. dehalogenans	D. hafniense Y51	D. hafniense DCB-2
fdh1A	3637	3101	4271	Formate dehydrogenase, catalytic	No	Yesd
fdh1B	3636	3100	4270	Formate dehydrogenase, Fe-S	Yes	Yes
fdh1C	3635	3099	4269	Formate dehydrogenase, cytochrome b	Yes	No
fdh1E	3634	3098	4268	Uncharacterized protein involved in formate dehydrogenase formation	No	No
fdh2H	1315	3972	1395	5-Formyltetrahydrofolate cycloligase	No	No
fdh2E	1316	3971	1396	NADH:ubiquinone oxidoreductase, 24-kDa subunit	No	No
fdh2F	1317	3970	1397	NADH:ubiquinone oxidoreductase, 51-kDa subunit	No	No
fdh2A	1318	3969-68	1398	NAD-dependent formate dehydrogenase catalytic subunit	No	No
frdC	616	735	743	Fumarate reductase cytochrome b	Yes	No
frdA	617	736	744	Menaquinone-dependent fumarate reductase, flavoprotein subunit	No	No
frdB	618	737	745	Fumarate reductase iron-sulfur protein	No	No
hydD	2200	1599	2740	Hydrogenase maturation protease	No	No
hydC	2201	1598	2741	Ni/Fe-hydrogenase, b-type cytochrome	Yes	No
hydB	2202	1597	2742	Ni,Fe-hydrogenase I large subunit	No	No
hydA	2203	1596	2743	Ni,Fe-hydrogenase I small subunit	No	Noe
	3649	3114	4283	Ni,Fe-hydrogenase III small subunit	No	No
	3650	3115	4284	Ni,Fe-hydrogenase III large subunit	No	No
	3651	3116	4285	Formate hydrogen lyase subunit 3/multisubunit Na+/H+ antiporter, MnhD subunit	Yes	No
	3652	3117	4286	Hydrogenase 4 membrane component (E)	Yes	No
	3653	3118	4287	Formate hydrogen lyase subunit 4	Yes	No
	3654	3119	4288	Formate hydrogen lyase subunit 3/multisubunit Na+/H+ antiporter, MnhD subunit	Yes	No
cprT	602		732	FKBP-type peptidyl-prolyl cis-trans isomerase (trigger factor)	No	No
cprK	603		733	cAMP-binding protein	No	No
cprZ	604		734	Protein of unknown function (DUF2869)	No	No
cprE	605		735	Chaperonin GroEL	No	No
cprB	606		736	Putative membrane anchor	Yes	No
cprA	607		737	Reductive dehalogenase	No	Yes
cprC	608		738	FMN-binding protein	Yes	Yes
cprD	610		740	Chaperonin GroEL	No	No
	212	285	232	Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FAD)	No	Yesd
	309			Flavocytochrome c (FAD-FMN)	No	Yes
	595			Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FAD)	No	Yesd
	643			Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FMN-FAD)	No	Yes
	646	764	773	Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FAD)	No	No
	650	768	779	Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FAD)	No	Yes
	2005	1391	2507	Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FAD)	No	Yesd
	3368	2816	3960	Succinate dehydrogenase/fumarate reductase flavoprotein subunit (FMN-FAD)	Yesf	Yes
	3673	3139	4309	Flavocytochrome c (FAD-FMN)	No	Yes
nrfA	3613	3065	4234	Formate-dependent nitrite reductase, periplasmic cytochrome c552 subunit	No	Yesd
nrfH	3614	3066	4235	Respiratory nitrite reductase specific menaquinol-cytochrome c reductase	Yes	No
nrfA	3046	2472	3631	Formate-dependent nitrite reductase, periplasmic cytochrome c552 subunit	No	Yes
nrfH	3045	2471	3630	Nitrate/TMAO reductase, membrane-bound tetraheme cytochrome c subunit	Yes	No
fixX	2231	1626	2773	Ferredoxin-like protein	No	No
fixC	2232	1627	2774	Flavin-dependent dehydrogenase	No	No
fixB	2233	1628	2775	Electron transfer flavoprotein, α subunit	No	No
fixA	2234	1629	2776	Electron transfer flavoprotein, β subunit	No	No
menF	1960	517	469	Isochorismate synthases	ND	ND
menD	1961	518	470	2-Succinyl-5-enolpyruvyl-6-hydroxy-3-cyclohexene-1-carboxylic acid synthase	ND	ND
menH	1531	519	471	2-Succinyl-6-hydroxy-2,4-cyclohexadiene-1-carboxylate synthase	ND	ND
menC	1964	522	474	O-Succinylbenzoic acid (OSB) synthetase	ND	ND
menE	1963	521	473	O-Succinylbenzoate-CoA ligase	ND	ND
menB	1962	520	472	Naphthoate synthase (dihydroxynaphthoic acid synthetase)	ND	ND
menI	2535	1906	3066	1,4-Dihydroxy-2-naphthoyl-CoA thioesterase	ND	ND
menA	3397	2871	4028	1,4-Dihydroxy-2-naphthoate octaprenyltransferase	ND	ND
menG	2536	1907	3067	Menaquinone biosynthesis methyltransferases	ND	ND

^aPrefixes to locus tags (not shown) are Desde, dsy, and Dhaf for D. dehalogenans and D. hafniense Y51 and DCB-2, respectively.

^bTMH, transmembrane helix as predicted with TMHMM v.2.0 (28).

^cS. Sq., predicted signal sequence.

^dSignal sequence detected only after analysis with SignalP in sensitive mode (72).

^eDetected only after analysis with SignalP in sensitive mode and using an alternative start codon.

^fOverlaps with TAT signal sequence.

^gAbbreviations: ND, not determined; FKBP, FK506-binding protein; cAMP, cyclic AMP; TMAO, trimethylamine oxide; CoA, coenzyme A.

The genome of D. dehalogenans contains two predicted formate dehydrogenase (FDH)-encoding gene clusters, fdh₁ABCE and fdh₂AFEH (Table 2). fdh₁ABCE encodes a three-subunit membrane-bound formate dehydrogenase, consisting of a molybdenum-containing catalytic subunit, an Fe-S protein, and a membrane-bound cytochrome b. Both fdh₁A and fdh₁B encode TAT signal sequences, indicating that this complex is located at the outer face of the membrane. The second formate dehydrogenase, fdh₂AFEH, encodes a cytoplasmic NADH-dependent formate dehydrogenase, consisting of a catalytic subunit, two (21- and 51-kDa) subunits involved in oxidation/reduction of NADH/NAD⁺, and a 5-formyltetrahydrofolate cycloligase.

The gene cluster frdCAB encodes a membrane-bound fumarate reductase/succinate dehydrogenase, consisting of a flavin adenine dinucleotide (FAD)-containing catalytic subunit, an Fe-S protein, and a membrane-bound cytochrome b. The genes of that complex do not encode any signal sequences, and the complex is therefore predicted to be located at the cytoplasmic face of the membrane.

We identified four gene clusters predicted to encode membrane-bound hydrogenases, including three NiFe uptake hydrogenases and an energy-conserving hydrogenase (Hyc, Table 2). Two of these hydrogenases have previously been identified as potentially involved in organohalide respiration (40). The first is an uptake hydrogenase, encoded by hydABC (Desde_2201-2203), consisting of a NiFe hydrogenase catalytic subunit, a Fe-S protein, and a membrane-bound cytochrome b (Table 2). The second, a putative energy-conserving hydrogenase (Hyc), is encoded by a six-gene cluster (3649 to 3654), of which four genes code for membrane proteins (Table 2). This cluster has some similarity (22 to 36% amino acid identity) with the formate hydrogen lyase 4 complex-encoding gene cluster (hyfABCDEFGHIR-focB) described for Escherichia coli (41). In fact, the D. dehalogenans hyc operon reported here has previously been described as a hyf operon, based on sequence analysis of transposon insertion sites in organohalide respiration-deficient mutants of this strain (40). Unlike the E. coli-carried hyf operon, the D. dehalogenans hyc operon does not encode a formate dehydrogenase.

The cprTKZEBACD gene cluster encodes the well-characterized Cl-OHPA reductive dehalogenase (CprA, Table 2). The catalytic subunit, CprA, contains a TAT signal peptide and two Fe-S binding domains, and the holoenzyme contains a corrinoid as cofactor. This protein is thought to be anchored to the membrane by CprB, a small protein consisting of 103 amino acids, predicted to contain three transmembrane helices. The remaining genes in this gene cluster encode accessory proteins involved in maturation and regulation. The cprTKZEBACD gene cluster is also present in Desulfitobacterium hafniense DCB-2 and has been characterized in detail in both organisms (17, 18, 42).

The genome encodes a complete menaquinone synthesis pathway (menFDHCEBIAG) (43). We found nine flavoproteins resembling FAD binding proteins present in a wide range of bacteria, often associated with electron transfer or reduction of terminal electron acceptors. Among these are the FAD binding C terminus of the Fe³⁺-induced flavocytochrome c fumarate reductase, Ifc3, from Shewanella frigidimarina NCIMB400 (Table 2), or a cytochrome c-dependent methacrylate reductase from Geobacter sulfurreducens AM-1. However, these predicted flavoproteins from D. dehalogenans, unlike Ifc3, do not contain any N-terminal heme binding CxxCH motifs (data not shown) (44–47). Two of these flavoproteins are annotated as flavocytochrome c, and the remaining seven are annotated as succinate dehydrogenase/fumarate reductases. All nine flavoproteins contain signal sequences, suggesting that they are exported outside the cell. Four of the flavoproteins contain a flavin mononucleotide (FMN) binding domain in addition to the FAD-containing fumarate reductase-like domain (Table 2).

In addition, the genome encodes two predicted cytochrome c nitrite reductases, NrfAH, thought to link the menaquinones with periplasmic nitrite reductase (48). Finally, the genome encodes a cluster of predicted membrane-associated electron-transferring flavodoxin and ferredoxin proteins, resembling FixABCX (Table 2). These proteins are involved in generating highly reduced ferredoxin that acts as an electron donor to the dinitrogenase in the diazotrophic Rhodospirillum rubrum (49, 50). It has been speculated that the products of this gene cluster also may provide low-redox-potential electrons for organohalide respiration (51).

Enzyme activity.

Formate alone does not allow fermentative growth of D. dehalogenans. Addition of fumarate or Cl-OHPA as an electron acceptor led to exponential growth, with doubling times of 8 to 10 h. In the late exponential phase, the FC- and FF-grown cultures reached an optical density at 600 nm (OD₆₀₀) of 0.17 and 0.25, respectively.

Subcellular localization of key enzyme activities.

Localization of formate dehydrogenase, chlorophenol reductive dehalogenase, and fumarate reductase activity was determined in FC- and FF-grown cells by in vitro enzyme assays using the membrane-impermeant artificial electron donor methyl viologen (Table 3). The membrane and soluble fractions contained 90 to 100% of the activity that was measured in the cell extracts, except for CPR, for which only 48% of the activity could be recovered, possibly due to partial inactivation or loss of cofactors. Furthermore, 20% and 64% of the total protein from the cell extract were recovered from the membrane and soluble fractions of FC- and FF-grown cells, respectively. The activities measured in the membrane and soluble fractions were used to predict the localization of enzymes.

TABLE 3.

Specific activities of formate dehydrogenase, chlorophenol reductive dehalogenase, and fumarate reductase in different fractions of Desulfitobacterium dehalogenans cells, grown with formate as an electron donor and Cl-OHPA or fumarate as an electron acceptor

Enzymeg	Electron acceptor	Sp acta
		Concentrated cells	Permeabilized cellsb	CFEe	Membrane fraction (%)f	Soluble fraction (%)f
FDH	Cl-OHPAd	14	108	59	189 (60)	973 (39)
	Fumarate	47	133	110	167 (90)	654 (32)
CPR	Cl-OHPA	51	87c	89	234 (48)	7 (0)
FRD	Fumarate	37	151	294	457 (92)	0 (0)
Total protein	FC/FF mg	174/94	174/94	180/96	32/56	4/5

^aSpecific activity is expressed as nanomole of substrate converted per minute per milligram of protein.

^bPermeabilization with 0.1% CTAB.

^cCells were permeabilized with 0.04% toluene to avoid inactivation of CPR by CTAB.

^dData from reference 34.

^eCFE, cell extract.

^fValues in parentheses are the ratios of the total activities of the cell extracts recovered from the membrane and cytoplasm fractions.

^gFDH, formate dehydrogenase; CPR, chlorophenol reductive dehalogenase; FRD, fumarate reductase.

In concentrated cell suspensions of formate–Cl-OHPA-grown cells, an FDH activity of 14 nmol formate min⁻¹ mg protein⁻¹ was measured, which increased 7.7-fold after permeabilization of the cells. In FF-grown cells, the specific FDH activity in concentrated cells was 42% of that observed in cell extract and increased 2.8-fold upon permeabilization of the cells with CTAB. We found for both growth conditions that the specific activity of FDH was higher in the soluble fraction than in the membrane fraction, whereas over 90% of the total activity was recovered from the membrane fraction (Table 3).

FRD was found to be membrane bound, and the activity increased 4-fold when cells were permeabilized with CTAB. CPR activity increased 1.7-fold upon permeabilization with toluene, which was used because CPR was found to be inhibited by CTAB. It should be noted that FRD and FDH activities determined in toluene-permeabilized cells were 4 to 5 times lower than those determined in CTAB-treated cells, indicating that permeabilization with toluene was less efficient or may have an inhibitory effect on these (data not shown). CPR activity was mainly found in the membrane fraction.

These results suggest that FDH is membrane associated. The fact that activity was observed on both sides of the cytoplasmic membrane is in agreement with the finding that the genome encodes two FDHs, including a predicted membrane-bound protein orientated toward the outside and a soluble cytoplasmic protein. Both CPR and FRD activity were found to be membrane bound. FRD was determined to be orientated at the cytoplasmic face of the cell membrane.

Menaquinone.

Quinones were extracted from FC- and FF-grown cells, and their involvement in fumarate and organohalide respiration was investigated. Extracted quinones exhibited a UV absorbance spectrum characteristic for menaquinones (MKs; data not shown). In their oxidized form, the isolated quinones showed two peaks at 245 and 270 nm and a broad absorption centered around 325 nm, whereas the dithionite-reduced samples exhibited a single peak at 245 nm and a broad absorption at 330 nm.

The reduced MK became oxidized by addition of Cl-OHPA (10 mM) to formate and dithionite-reduced formate–Cl-OHPA grown cells, or when fumarate (10 mM) was added to formate- and dithionite-reduced FF-grown cells (Table 4). These findings indicate that MK is involved in electron transport from formate to both Cl-OHPA and fumarate.

TABLE 4.

Ratios of peak areas of reduced and oxidized menaquinone extracted under different conditions from cells of D. dehalogenans grown with formate as electron donor and fumarate or Cl-OHPA as electron acceptor

Conditionb	MKH2/MK ratio for cells grown with:
	Formate-fumarate	Formate–Cl-OHPA
Formate	0.33	1.79
Cl-OHPA	NDa	0.09
Fumarate	0.01	ND
No addition	0.12	0.30

^aND, not determined.

^bCells were incubated for 10 min with 10 mM formate, Cl-OHPA, or fumarate prior to quinone extraction.

UV-visible and electron paramagnetic resonance (EPR) spectroscopic analysis.

The absorption spectra were recorded from concentrated cell suspensions of FF- and FC-grown cultures, to determine the involvement of cytochromes in fumarate and Cl-OHPA respiration. Changes in the redox state of cytochromes were monitored by visible spectrometry at 552 and 561 nm (Fig. 1). The low concentration of cytochrome b in formate–Cl-OHPA cells did not allow us to determine changes in its redox state, whereas the concentrations of cytochromes b and c combined were sufficient to study their involvement in respiration during growth on formate and fumarate (Fig. 1). The maxima between 550 and 560 nm, seen in the absorption spectrum of concentrated cell suspensions of FF- and FC-grown cultures, indicated that the cytochromes were partially reduced (Fig. 1, traces A and F). Upon addition of formate to concentrated cell suspensions, a rapid reduction of the cytochromes could be seen (Fig. 1, traces B and G), which is similar to the level of reduction that was obtained upon addition of dithionite (Fig. 1, traces C and H). Incubation of concentrated formate–Cl-OHPA cells with Cl-OHPA or of concentrated FF cells with fumarate resulted in a decrease of the absorption peaks, indicating that the cytochromes became oxidized (Fig. 1, traces D and I). Upon addition of 10 mM Cl-OHPA to formate-reduced formate–Cl-OHPA cells, the cytochrome c became oxidized (Fig. 1, trace E), although a complete reoxidation of cytochrome c was observed at neither low nor high Cl-OHPA concentrations. A complete reoxidation of both cytochromes b and c was observed when 10 mM fumarate was added to formate-reduced FF cells (Fig. 1, trace J).

FIG 1.

Low-temperature cytochrome b and c α-band spectra of a concentrated cell suspension of Desulfitobacterium dehalogenans grown with formate as an electron donor and either Cl-OHPA (traces A to E) or fumarate (traces F to J) as an electron acceptor. Spectra were shifted upward in steps of 0.02 absorbance units to clarify the figure. Traces A and F show the absorption spectra from cells without additions. Traces B and G were recorded after incubation of the cells with 10 mM formate. Traces C and H were recorded after reduction of the sample with dithionite for 8 min. Traces D and I were recorded after incubation of the cells with 1 mM formate and 10 mM Cl-OHPA or fumarate. Traces E and J were recorded after incubation of the cells with 10 mM Cl-OHPA or fumarate.

Electron paramagnetic resonance (EPR) spectra of concentrated formate–Cl-OHPA cells after addition of formate, and after addition of formate and Cl-OHPA, were recorded to obtain information on the involvement of Fe-S clusters and molybdenum. A detailed description of these experiments can be found in Text S1 in the supplemental material.

Comparison of the proteomes of FC-, FF-, and P-grown cells.

Shotgun proteomics was used to identify proteins produced at different levels in cells grown by respiration or fermentation. To this end, we compared the proteomes of FC- and FF-grown cells with those of P-grown cells. Furthermore, we compared proteomes of cells grown by organohalide versus fumarate respiration (FC versus FF). We identified in total 578 different proteins out of a theoretical maximum of 4,142 (see Table S1 in the supplemental material). In total, 104 proteins were present at significantly (P < 0.05) different levels between at least two growth conditions (see Table S1 in the supplemental material). The largest variations were seen between respiratory and fermentative growth, with 79 and 70 proteins varying significantly (P < 0.05) between FC- or FF-grown cells and P-grown cells, whereas only 20 proteins were found to differ significantly between FC- and FF-grown cells (see Table S1 in the supplemental material). For many enzyme complexes, we detected only the non-membrane-embedded components. As extracting and detecting membrane proteins is still a challenging task (52), we assume that the values for the detected components are representative for the entire complex.

The catalytic subunit of the membrane-bound formate dehydrogenase complex, FdhA (Desde_3637), increased 59- and 9-fold in FF- and FC-grown cells compared to P-grown cells, respectively. However, only the first value was found to be statistically significant (P < 0.05) (Fig. 2; see also Table S1 in the supplemental material). This upregulation of FdhA for both respiratory growth conditions compared to fermentative growth indicates that the same formate dehydrogenase is used when Cl-OHPA or fumarate is used as an electron acceptor. In contrast, the cytoplasmic NADH-dependent formate dehydrogenase encoded by fdh₂AFEH was present in equal amounts during both respiratory and fermentative growth, indicating that it is part of other cellular processes, rather than energy conservation by respiration with formate as an electron donor (see Table S1 in the supplemental material).

FIG 2.

Changes in relative protein abundance between growth conditions where formate and either Cl-OHPA or fumarate (20 mM) was used as an electron donor or acceptor or growth by fermentation of pyruvate (40 mM). (A) Formate–Cl-OHPA versus pyruvate. (B) Formate-fumarate versus pyruvate. (C) Formate–Cl-OHPA versus formate-fumarate. The logarithm of the total-intensity-based absolute quantification intensity (IBAQ) is plotted against the logarithm of the total protein abundance ratio, for selected proteins (also Fig. 3). The data points for CprA and the flavoprotein encoded by Desde_3368 are highlighted in the graphs.

Two elements of the membrane-bound fumarate reductase encoded by frdAB (Desde_617-618) were upregulated (10- to 14-fold) under both respiratory growth conditions, suggesting that their synthesis is regulated by redox and energy state, rather than by the presence of the substrate.

All elements encoded by the cprTKZEBACD gene cluster were identified, except the transcriptional regulator CprK and the highly hydrophobic CprB. This gene cluster was, as expected, highly (>270-fold) upregulated when D. dehalogenans was grown with Cl-OHPA (Fig. 2A and C; also see Table S1 in the supplemental material).

Two putative extracellular electron-transferring flavoproteins were present in the proteome. A flavoprotein (Desde_3368) predicted to harbor two flavin-binding domains, an N-terminal FMN and a C-terminal FAD, increased strongly in abundance (>1,000-fold) when D. dehalogenans was grown with Cl-OHPA. This increase is of the same magnitude as that observed for elements of the cpr gene cluster. This suggests that this protein is involved in organohalide respiration, potentially as an electron shuttle between the MKs and CprA. The second identified flavoprotein (Desde_3673), also predicted to harbor two flavin-binding domains, an N-terminal FAD and a C-terminal FMN, was found in large amounts under all growth conditions (Fig. 2; see also Table S1 in the supplemental material).

Interestingly, we identified a protein annotated as cytochrome c nitrite reductase NrfA (Desde_3613), which is the outward-facing component of the membrane-bound NrfAH quinol dehydrogenase/nitrite reductase. Unfortunately, NrfA was not identified in all replica samples, preventing firm conclusions on eventual changes in abundance between growth conditions from being drawn (Fig. 2; see also Table S1 in the supplemental material).

The electron-transferring flavoproteins encoded by fixABCX (Desde_2231-34) were detected under all growth conditions. Their levels, however, did not vary significantly between growth conditions (Fig. 2; see also Table S1 in the supplemental material).

An uptake hydrogenase (HydABC, Desde_2200-2203) and an energy-conserving hydrogenase (Hyc, Desde_3649-3654) have previously been suggested to play a role in organohalide respiration (40). Subunits of the uptake hydrogenase HydABC were upregulated 10- to 18-fold during respiratory growth (FC and FF) compared to growth by fermentation (P). This suggests that expression is regulated by redox and energy state, rather than by the presence of the substrate. In contrast, we did not detect any subunits of the two alternative uptake hydrogenases in the proteome (Fig. 2; see also Table S1 in the supplemental material).

Elements of the energy-conserving hydrogenase (Hyc) were not detected. Four of the six subunits of this complex contain several transmembrane helices and are therefore tightly embedded in the membrane (Table 2). We cannot completely rule out the possibility that this complex is present in D. dehalogenans under the tested growth conditions but has not been extracted from the membrane and thus was not observed in the proteome analysis.

DISCUSSION

Formate-fumarate respiration.

The genome of D. dehalogenans encodes two FDHs and one FRD. FDH₁ and FRD were found to be upregulated under both respiratory (FC and FF) growth conditions. FDH₁ and FRD from D. dehalogenans resemble the well-characterized FDH and FRD of Wolinella succinogenes (Fig. 2 and 3A; see also Table S1 in the supplemental material) (53), suggesting similar topologies of the electron transport chain from formate to fumarate in the two bacteria. Electrons are channeled from an outward-facing FDH complex containing a Fe-S protein and a cytochrome b integral membrane protein via menaquinones to an FRD complex facing the cytoplasm, thereby creating a proton motive force (53, 54). These findings are supported by both the genome and the localization studies. The genome predicts the presence of both an outward-facing and a cytoplasmic formate dehydrogenase (FDH₁ and FDH₂) (Table 2). In agreement with this, we found FDH activity at both sides of the membrane, whereas FRD activity was localized at the cytoplasmic face of the cell membrane (Table 3; Fig. 3A). The involvement of menaquinones as electron carriers was demonstrated by the observation that they could be reduced with formate and reoxidized with fumarate or Cl-OHPA (Table 4).

FIG 3.

(A and B) Overview of enzyme complexes shown to be, or predicted to be, involved in energy conservation in D. dehalogenans growing with formate as an electron donor and fumarate (A) or Cl-OHPA (B) as an electron acceptor. (C) Putative interaction between the energy-conserving hydrogenase Hyc and uptake hydrogenase HUP. Numbers in parentheses indicate locus tags with the Desde_ prefix removed. Bold arrows show directions of reactions or movements; dashed arrows show potential directions of reactions or movements. FDH, formate dehydrogenase; FRD, fumarate dehydrogenase; NrfAH, cytochrome c quinol dehydrogenase/nitrite reductase; ?, unidentified membrane-bound electron transfer complex; Fix, fix gene cluster products; CprABC, chlorophenol reductase; Hyc, energy-conserving hydrogenase; HUP, uptake hydrogenase.

The second formate dehydrogenase, FDH₂, an NAD-dependent formate dehydrogenase, does not contain any signal sequences or transmembrane domains, indicating a cytoplasmic localization. Proteomic data showed that FDH₂ was present in the cells in equal amounts under all three tested growth conditions (Fig. 2; see also Table S1 in the supplemental material). In the same operon, we identified a gene predicted to encode a 5-formyltetrahydrofolate cycloligase (Desde_1315). Analysis by tBLASTn revealed that this gene organization is commonly found in sequenced genomes of bacteria beyond the Desulfitobacterium genus (data not shown). Taken together, these data suggest that FDH₂ is not part of the electron transport chain from formate to fumarate or Cl-OHPA but rather may be part of the methyl branch of the Wood-Ljungdahl pathway, catalyzing the reduction of CO₂ to formate (for a review, see reference 55). It has previously been demonstrated that D. dehalogenans is capable of CO₂ fixation (34).

Formate–Cl-OHPA respiration.

The electron transport pathway from electron donor to acceptor during organohalide respiration is still poorly understood. Previously, a simple model analogous to formate-fumarate respiration has been proposed (4, 6, 12). According to this, the electron-donating and electron-accepting reactions are orientated toward the periplasmic and cytoplasmic faces, respectively, thereby creating a proton motive force driving ATP synthesis (4, 6, 12). The cprA gene, however, like other reductive dehalogenase-encoding genes, codes for a preprotein with a TAT signal sequence, strongly suggesting that CprA is exported across the cytoplasmic membrane (3, 56). In Desulfitobacterium hafniense strain Y51 and Sulfurospirillum multivorans, PceA is constitutively expressed. In both strains, PceA was found to be present on both sides of the cell membrane when grown with PCE, but PceA was found solely inside the cells when grown without PCE (57, 58). It seems likely that the tightly regulated CprA in Desulfitobacterium dehalogenans also is exported across the cell membrane. The increase in CprA activity after permeabilization would then be due to the presence of folded and active CprA that has not yet been exported.

Our findings suggest that the electron transport chain from FDH₁ to Cl-OHPA consists of menaquinones, likely an extracellular flavoprotein and a membrane complex linking the menaquinones with extracellular respiratory processes (Fig. 3B). This is supported by the finding that menaquinones are oxidized by both Cl-OHPA and fumarate (Table 4). The involvement of menaquinones in the electron transport chain during organohalide respiration has also been shown for the closely related Dehalobacter restrictus (6). The standard redox potential of menaquinones is −74 mV, whereas it has been demonstrated that PceA from Sulfurospirillum multivorans (initially named Dehalospirillum multivorans) requires an electron donor with a standard redox potential below −360 mV (14, 59). Menaquinones are thus unlikely to deliver the electrons directly to the reductive dehalogenase. It has been suggested that strong reducing equivalents, necessary for reducing reductive dehalogenases, are generated by reverse electron flow or electron bifurcation (14, 57, 60). Combining genome and proteomic data, we were able to identify some candidates for the link between the menaquinones and the periplasm-orientated CprA. (Fig. 3B). One of these is NrfAH, a putative membrane-bound cytochrome c-containing quinol dehydrogenase/nitrite reductase. The genome of D. dehalogenans encodes two NrfAH homologues (Table 2), of which one, Desde_3313-14, was identified in the proteomes from all tested growth conditions (Fig. 2 and 3B; see also Table S1 in the supplemental material). Some studies have found NrfH capable of donating electrons to periplasmic electron donors other than NrfA (48, 61), whereas other studies did not find evidence for such a mechanism (62). This may suggest a role for NrfH as an electron hub between the menaquinones and extracellular electron acceptors, analogous to the function of CymA in Shewanella spp. (reference 46 and references therein). Similar to this, it was recently suggested that an NapGH-like quinol dehydrogenase acts as the link between the menaquinones and the tetrachloroethene reductase PceA in Sulfurospirillum multivorans (63). The genome of D. dehalogenans does not encode any CymA homologue. Another candidate electron transport complex is encoded by the fixABCX gene cluster (Desde_2231-2234), the products of which are involved in generation of reductants for nitrogen fixation by reversed electron flow in Rhodospirillum rubrum (49, 50). As the genome of D. dehalogenans does not encode the minimum machinery necessary for nitrogen fixation (64), the FixABCX complex most likely is linked to other cellular processes in this bacterium. Recently, comparative transcriptomics of D. hafniense Y51 (65) and proteomics of D. hafniense TCE1 (51) showed expression of fixABCX to be induced under organohalide-respiring conditions, leading to the speculation that the encoded complex may provide low-redox-potential electrons for organohalide respiration (51). In our proteomic data set, we saw no differences in the abundances of the fixABCX gene products between the tested growth conditions (P < 0.05) (Fig. 2; see also Table S1 in the supplemental material). The lack of differential expression indicates that if FixABCX were part of the organohalide respiratory electron transport chain to CprA, it would also be involved in other cellular processes. Furthermore, FixABCX is located in the cytoplasm and loosely associated with the membrane (49). If the FixABCX complex creates low-potential electron donors needed for CprA activity, an as-yet-unidentified electron carrier must transport these across the membrane to the outward-facing CprA (Fig. 3B).

Cytochromes were analyzed in whole cells grown with Cl-OHPA or fumarate. The shifts in the absorption spectra of reduced cytochromes after addition of either Cl-OHPA or fumarate showed that cytochrome c could be completely oxidized by fumarate and partly by Cl-OHPA (Fig. 1). These findings would be in agreement with the activity of a membrane-bound cytochrome c quinol dehydrogenase, such as NrfAH. The oxidation of menaquinones by fumarate indirectly leads to oxidation of NrfAH, although this complex is not part of the electron transport pathway from formate to fumarate (Fig. 3A). The seemingly less efficient oxidation of cytochrome c with Cl-OHPA might be due to partial loss of extracytoplasmic components of the electron transport chain from cytochrome c to Cl-OHPA, caused by handling of the samples (Fig. 3B).

The genomes of D. dehalogenans and D. hafniense strains Y51 and DCB-2 encode, unlike other metal-oxidizing bacteria, only a limited number of cytochromes c (9, 19), leading to the speculation that Desulfitobacterium spp. utilize electron carriers other than cytochrome c for organohalide respiration (9). Two predicted extracytoplasmic electron-transferring flavoproteins (Desde_3368 and Desde_3673) were identified in the proteome in the present study. These resemble flavocytochromes c, but both lack known consensus heme binding domains (CxxCH) and are not found next to cytochrome c-encoding genes in the genome. One of these flavoproteins (Desde_3368) increased in abundance in the presence of Cl-OHPA by 3 orders of magnitude, an increase similar to that of elements encoded by the cprTKZEBACD gene cluster (Fig. 2; see also Table S1 in the supplemental material). This was a surprising finding, as this type of flavoprotein has not previously been associated with organohalide respiration. Flavoproteins similar to those from D. dehalogenans have been found in a range of bacteria (66). Usually, these are linked with a tetraheme cytochrome c, either directly as a flavocytochrome c fusion protein or as a flavoprotein-cytochrome c enzyme complex (45–47). In Shewanella frigidimarina, a flavocytochrome c, Ifc3, is induced in the presence of Fe³⁺. It was speculated that Ifc3 acts as an electron shuttle between a membrane-bound quinol dehydrogenase, CymA, and extracellular electron acceptors (45). A similar role has been suggested for other flavocytochromes c (46). A recent study on selenite reduction by Shewanella oneidensis MR-1 indicated that a deletion of cymA results in an inability to reduce selenite. Deleting the flavocytochrome c-encoding gene fccA led to lower selenite reduction rates, which seems to be in agreement with a role for flavocytochrome c as an electron shuttle, although it cannot be ruled out that FccA acts as the terminal selenite reductase (67). Methacrylate reduction in Geobacter sulfurreducens AM-1 was shown to be carried out by a periplasmic flavoprotein tetraheme cytochrome c complex (47). The strong increase in abundance of Desde_3368 when the cells were grown in the presence of Cl-OHPA makes this flavoprotein (Desde_3368) a promising candidate for the yet-unidentified electron shuttle between a membrane-bound electron-donating complex and CprA (Fig. 2 and 3B). The genomes of D. dehalogenans and D. hafniense strains DCB-2 and Y51 encode 9, 26, and 30 homologues of these putative electron-transferring flavoproteins, respectively (Table 2) (9, 19). Some of these have been annotated as flavocytochromes c, but all lack heme binding domains. The majority are, however, annotated as fumarate reductase/succinate dehydrogenase or flavoproteins (data not shown), showing that careful curation of automatically generated predictions of protein function is needed after more detailed functional studies.

One (Desde_2200-2203) out of the three uptake hydrogenases (HUP) encoded in the genome of D. dehalogenans increased 10- to 18-fold in abundance during respiratory growth compared to fermentative growth (Fig. 2; see also Table S1 in the supplemental material). This is in line with previous findings of the involvement of hydrogenases in respiratory growth of D. dehalogenans (40). Interestingly, a similar increase in the expression of genes encoding an uptake hydrogenase corresponding to hydABC has been reported for Desulfitobacterium hafniense Y51 under respiratory growth compared to fermentative growth (65). This points toward a more general role in respiratory metabolism rather than a specific role in organohalide respiration, potentially as part of a hydrogen recycling mechanism as described elsewhere for diazotrophs and sulfate reducers (68–70). No elements of the Hyc energy-conserving hydrogenase could be detected in the proteome. This may be due to the presence of transmembrane helices in four of the six subunits, embedding it tightly in the cell membrane (Table 2). We cannot rule out the possibility that the Hyc complex is present in D. dehalogenans under the growth conditions tested. Based on the current data, the exact role of HUP and Hyc cannot be established. It is tempting to speculate that they interact as speculatively shown in Fig. 3C, but whether they play a direct role in the respiratory electron transport or are involved in other metabolic processes as suggested for the organohalide-respiring Dehalococcoides spp. still requires further studies (71).

In this study, we report the full genome sequence of Desulfitobacterium dehalogenans strain JW/IU-DC1^T, which, combined with comparative proteomics and biochemical analysis, allowed us to obtain an improved insight into FF and FC respiration in D. dehalogenans. FF respiration occurs in a way analogous to the well-described mechanism for Wolinella succinogenes (53). Our data show that the first part of the electron transport pathway from FDH to MKs is shared between FF and FC respiration. Although the exact link between the menaquinones and the outward-facing CprA could not be determined, proteome analysis pointed to a possible role of gene products of nrfAH or, alternatively, fixABCX, but this link could also be carried out by other membrane complexes like Hyc, or the products of yet-unidentified gene clusters. Finally, we identified an extracytoplasmic flavoprotein as a candidate for an electron shuttle between a membrane-bound electron-donating complex and the outward-facing CprA. Future investigations such as comparative as well as functional genomics of organohalide-respiring bacteria, including the characterization of knockout mutants, can be expected to further propel our understanding of the energy metabolism of desulfitobacteria and other organohalide-respiring bacteria.