JOURNAL OF BACTERIOLOGY,0021-9193/00/$04.0010

June 2000, p. 3097–3103 Vol. 182, No. 11

Copyright © 2000, American Society for Microbiology. All Rights Reserved.

A Membrane-Bound Flavocytochrome c-Sulfide Dehydrogenasefrom the Purple Phototrophic Sulfur Bacterium

Ectothiorhodospira vacuolataVESNA KOSTANJEVECKI,1 ANN BRIGE,1 TERRANCE E. MEYER,2 MICHAEL A. CUSANOVICH,2

YVES GUISEZ,1 AND JOZEF VAN BEEUMEN1*

Laboratory for Protein Biochemistry and Protein Engineering, University of Ghent, 9000 Ghent, Belgium,1

and Department of Biochemistry, University of Arizona, Tucson, Arizona 857212

Received 24 November 1999/Accepted 3 March 2000

The amino acid sequence of Ectothiorhodospira vacuolata cytochrome c-552, isolated from membranes withn-butanol, shows that it is a protein of 77 amino acid residues with a molecular mass of 9,041 Da. It is closelyrelated to the cytochrome subunit of Chlorobium limicola f. sp. thiosulfatophilum flavocytochrome c-sulfidedehydrogenase (FCSD), having 49% identity. These data allowed isolation of a 5.5-kb subgenomic clone whichcontains the cytochrome gene and an adjacent flavoprotein gene as in other species which have an FCSD. Thecytochrome subunit has a signal peptide with a normal cleavage site, but the flavoprotein subunit has a signalsequence which suggests that the mature protein has an N-terminal cysteine, characteristic of a diacylglycerol-modified lipoprotein. The membrane localization of FCSD was confirmed by Western blotting withantibodies raised against Chromatium vinosum FCSD. When aligned according to the three-dimensionalstructure of Chromatium FCSD, all but one of the side chains near the flavin are conserved. These include theCys 42 flavin adenine dinucleotide binding site; the Cys 161-Cys 337 disulfide; Glu 167, which modulates thereactivity with sulfite; and aromatic residues which may function as charge transfer acceptors from theflavin-sulfite adduct (C. vinosum numbering). The genetic context of FCSD is different from that in otherspecies in that flanking genes are not conserved. The transcript is only large enough to encode the two FCSDsubunits. Furthermore, Northern hybridization showed that the production of E. vacuolata FCSD mRNA isregulated by sulfide. All cultures that contained sulfide in the medium had elevated levels of FCSD RNAcompared with cells grown on organics (acetate, malate, or succinate) or thiosulfate alone, consistent with therole of FCSD in sulfide oxidation.

Flavocytochrome c-sulfide dehydrogenase (FCSD) was firstidentified by Bartsch and Kamen (6) in the purple phototro-phic bacterium Chromatium vinosum, recently renamed Allo-chromatium vinosum (21). It was subsequently found in thegreen phototrophic bacterium Chlorobium limicola f. sp. thio-sulfatophilum (7, 30) and in six other species of purple andgreen bacteria, Chromatium gracile (4, 5) and Chromatiumpurpuratum (24), both now assigned to the genus Marichroma-tium (21); Chromatium tepidum (20); Thiocapsa roseopersicina(54); Chlorobium limicola f. sp. thiosulfatophilum (43); andChlorobium phaeobacteroides (16). A membrane-bound formof FCSD was discovered in the nonphototrophic aerobic bac-terium Thiobacillus sp. W5 (50). A gene homologous to theflavoprotein subunit of FCSD was found to be associated withthe genes for thiosulfate oxidation in Paracoccus denitrificans,but the adjacent cytochrome gene was more divergent thanexpected (52). The genome sequence of Aquifex aeolicus con-tains two FCSD flavoprotein genes associated with one for aRieske iron-sulfur protein, and a thiosulfate utilization operonis located elsewhere in the genome (14). Thus, there appears tobe a correlation between the presence of FCSD and sulfurmetabolism.

The Chromatium and Chlorobium flavocytochromes c werefound to have sulfide dehydrogenase activity in vitro (19, 25)

and presumably are involved in sulfur metabolism in vivo. Acommon characteristic of the bacteria in which this enzyme isfound is the ability to utilize reduced sulfur compounds aselectron donors for carbon dioxide fixation. Thus, all species ofgreen and purple sulfur bacteria utilize elemental sulfur andhydrogen sulfide, approximately half the species use thiosul-fate, and a few use sulfite or tetrathionate (10). However,thiosulfate and sulfite are not oxidized by FCSD. Once thegenes for C. vinosum FCSD were cloned and sequenced, theperiplasmic location of the enzyme was established by thepresence of signal sequences (15).

The discovery of a homologous enzyme, sulfide-quinone re-ductase (SQR), in some photosynthetic and nonphotosyntheticsulfur bacteria (2, 39, 40, 41) plus the isolation of theRhodobacter capsulatus SQR gene brought a new perspectiveon the involvement of FCSD and SQR in sulfide oxidation.During the oxidation of sulfide, sulfur globules are depositedeither in the periplasmic space, as in Chromatium species (34),or extracellularly (in Chlorobium and Ectothiorhodospira spe-cies) (18). Thus, sulfide oxidation generally occurs on theperiplasmic side of the cytoplasmic membrane in purple andgreen sulfur bacteria, where FCSD is localized. The location ofthe R. capsulatus SQR was not clear until recently because theprotein appeared to have no N-terminal signal peptide fortranslocation into the periplasmic space (41). It has now beendocumented from gene fusion experiments that R. capsulatusSQR functions on the periplasmic side of the cytoplasmicmembrane, using an unknown mechanism for translocation(38). Genes for FCSD have not been found in R. capsulatus,nor have SQR genes been conclusively demonstrated in either

* Corresponding author. Mailing address: Department of Biochem-istry, Physiology and Microbiology, Laboratory of Protein Biochemis-try and Protein Engineering, University of Ghent, Ledeganckstraat 35,B-9000 Ghent, Belgium. Phone: 32-92645109. Fax: 32-92645338. E-mail: Jozef.vanbeeumen@rug.ac.be.

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Chromatium or Chlorobium. On the other hand, disruption ofthe FCSD genes in C. vinosum had no effect on sulfide oxida-tion (35). However, there are two FCSD genes in the C. tepi-dum genome (http://www.tigr.org), suggesting that if there aretwo sets of FCSD genes in C. vinosum, the effects of a knockoutmutation would be negated. Nevertheless, which of the twoenzymes, FCSD or SQR, is the sulfide-oxidizing enzyme inChromatium and Chlorobium remains to be determined.

Ectothiorhodospira species make up a small family of mostlymarine and halophilic purple phototrophic bacteria, all ofwhich utilize sulfide for growth. Unlike Chromatium, but sim-ilar to Chlorobium, elemental sulfur is deposited outside thecells in the growth medium. The soluble electron transfer pro-teins of Ectothiorhodospira are generally similar to those ofChromatium. Thus, they are dominated by HiPIP (high-poten-tial iron-sulfur protein) and cytochrome c9 (26, 27, 31). FCSDhas not been previously reported to occur in this family ofbacteria.

The amino acid sequences of the cytochrome subunits ofChromatium and Chlorobium FCSDs have been determined(47, 48), as has that of the Chromatium flavoprotein subunit(49). The three-dimensional structure of Chromatium FCSDhas also been determined (12). We now report the nucleotidesequence of an Ectothiorhodospira vacuolata FCSD gene andits flanking regions, which shows that FCSD is more wide-spread than previously thought and that the genetic context isnot the same as that in Chromatium. The membrane localiza-tion of the E. vacuolata protein is documented by immunoblot-ting.

MATERIALS AND METHODS

Strains and media. E. vacuolata b1 strain 2111, obtained from the DeutscheSammlung von Mikroorganismen und Zellkulturen (DSMZ) was grown onDSMZ medium 1448, which is a modification of the American Type CultureCollection medium 1410 (30 g of NaCl/liter instead of 140 g/liter). The culturewas grown by anaerobic photosynthesis in light provided by a 40-W tungstenlamp at 30°C. Escherichia coli strains were grown on Luria-Bertani medium (36)supplemented with 100 mg of carbenicillin. Strain XL-1 blue was used as arecipient to detect a-complementation for pUC18 derivatives on Luria-Bertaniplates supplemented with 80 mM IPTG (isopropyl-b-D-thiogalactoside) and 32mg of X-Gal (5-bromo-4-chloro-3-indolyl-b-D-galactoside) per ml.

Isolation and purification of cytochrome c-552. Cells were harvested by cen-trifugation and broken in a Ribi cell fractionator (an automated French press).The membranes were sedimented by ultracentrifugation for 3 h in a Spinco Ti45rotor. They were resuspended in water at 4°C, and an equal volume of n-butanolat 220°C was added with stirring (46). After centrifugation, the top layer con-taining butanol and colored pigments was removed. The underlying water layercontained the protein of interest. Cytochrome c-552 chromatographed withHiPIP on DEAE-cellulose and eluted after cytochrome c4. It appeared to existas both monomer and dimer when chromatographed on Sephadex G-50. Thecombined Sephadex fractions were chromatographed on DEAE-cellulose devel-oped with 20 mM Tris-HCl containing 2.5 mM NaCl, which resolved the cyto-chrome c-552 from a HiPIP isozyme.

Amino acid sequence determination of cytochrome c-552. The covalentlybound heme was removed from the native protein by treatment with HgCl2 in 8M urea–0.1 M HCl at 37°C for 16 h (1). After separation of the apoprotein fromheme and salts by gel filtration (Sephadex G-25, 5% HCOOH), the N-terminalsequence was determined by using a freeze-dried aliquot of 400 pmol of theapoprotein. To identify the cysteine residues of the heme binding site, a 1.3-nmolaliquot of apoprotein was treated with 3-bromopropylamine. The alkylated cys-teine residues could then be detected by sequence analysis (22). A second aliquotof 9.5 nmol of the apoprotein was digested for 3 h with Staphylococcus aureusprotease at pH 4, using an enzyme-to-substrate ratio of 1:40. A third aliquot of9 nmol was subjected to partial acid hydrolysis in 2% formic acid for 2 h at 106°C.The peptides were separated by reversed-phase high-performance liquid chro-matography (SMART system; Pharmacia, Uppsala, Sweden) on a PEPSII col-umn using a gradient of 0.07% trifluoroacetic acid in water (solvent A) and0.05% trifluoroacetic acid in acetonitrile (solvent B). Sequence determinationwas performed on a 477A or 476A pulsed liquid sequenator, with on-line analysisof the phenylthiohydantoin-amino acids on a 120A analyzer (PE Biosystems,Foster City, Calif.). Sequencing reagents were from the same firm. The masses ofthe holoprotein, apoprotein, and peptides were determined by using eitherplasma desorption, electrospray ionization, or matrix-assisted laser desorptionmass spectrometry on a Biopolymer (Uppsala, Sweden) analyzer (BIO-ION), a

BIO-Q triple-quadruple instrument (Micromass, Altrincham, United Kingdom),or a TOF-SPEC SE time-of-flight analyzer (Micromass, Whytenshaw, UnitedKingdom), respectively.

DNA techniques. E. vacuolata genomic DNA was isolated by the cetyltrimeth-ylammonium bromide method (3). On the basis of the cytochrome c-552 proteinsequence, a set of degenerate primers was designed. The N-terminal primer,EVM8, had the sequence 59 ATGGCHACHACHTGYTAYG 39, and the C-terminal primer, EVT61, had the sequence 59 AGCTTGATYTCYTCRTCHGTRTA 39. The probe was amplified by PCR using Taq polymerase (AmershamPharmacia Biotech, Uppsala, Sweden) under the following conditions: 95°C, 2min; 5 precycles (94°C, 30 s; 50°C, 60 s; 72°C, 30 s); 30 cycles (94°C, 30 s; 52°C,30 s; 72°C, 60 s); 72°C, 10 min (followed by 4°C). The amplified fragment of 177bp was cloned in the pGEM-T vector (Promega) and labeled via PCR withdioxigenin-dUTP (Roche Molecular Biochemicals). Via Southern hybridization,a BamHI fragment was identified. A BamHI pUC18 library was then constructedand analyzed (36). Detection and identification of transformants were done withthe nonradioactive digoxigenin-DNA detection system (Roche Molecular Bio-chemicals). Double-stranded plasmid DNA was sequenced using dye terminatorcycle sequencing (PE Biosystems). The sequencing was started from both endswith the universal primers M13F and M13R (England Biolabs, Inc., Beverly,Mass.) and was continued with the specific primers 552/3 (59 CGCCACCGTCATGGATC 39) and 552/4 (59 GCTGCCGGCACTGTGTC 39), created on thebasis of the c-552 DNA sequences. New primers were synthesized at approxi-mately 450-nucleotide intervals based on the results of the previous sequencing.

Membrane protein purification and Western blotting. Ten liters of E. vacu-olata culture was harvested in the exponential phase. Membranes were purifiedfrom French press-lysed cells via ultracentrifugation (160,000 3 g 3 h) combiningthe EDTA-lysozyme method (32) and fractionation with 20% (NH4)2SO4. Themembranes were solubilized with 1% Triton X-100–10 mM Tris-HCl and 10 mMEDTA, pH 8 (3-h incubation; 4°C), and insoluble material was removed byultracentrifugation. The supernatant was desalted and adsorbed on a DEAE-Sepharose column. The purification protocol was performed basically as de-scribed in reference 50 with the exception of using the above-mentioned Triton-Tris-EDTA buffer. Proteins were eluted with a linear gradient of 0 to 0.5 M KClin the same buffer. The pooled fractions were desalted, concentrated, and furtherpurified by loading them on a Q-Sepharose Fast Flow column in the above-mentioned buffer and gradient. The purified protein was identified on sodiumdodecyl sulfate (SDS) gels by silver and heme staining. Western blotting wasperformed following the manufacturer’s instructions for the enhanced chemilu-minescence membrane (Amersham Pharmacia Biotech, Roosendaal, The Neth-erlands) and using an antibody against C. vinosum FCSD.

Protein electrophoresis. SDS-polyacrylamide gel electrophoresis (PAGE) wasperformed on vertical 10% polyacrylamide gels (28) stained for protein withCoomassie blue or silver. The gels used for heme staining were incubated for 10min in a 10% trichloroacetic acid solution; the staining itself was performed asdescribed in reference 45. PAGE as well as protein-blotting experiments wereperformed using the Mini Protean equipment (Bio-Rad, Veenendaal, The Neth-erlands).

RNA extraction and Northern analysis. Total RNA from E. vacuolata wasextracted with LiCl (3) and electrophoresed in 2% agarose formaldehyde gels.All RNA samples were treated with RNase-free DNase I (Sigma) for 10 min at37°C. Afterwards, 14 ml of denaturation mix (9.2 ml of formamide, 3 ml offormaldehyde, and 1.8 ml of MOPS buffer (40 mM morpholinopropanesulfonicacid, 10 mM sodium acetate, 2 mM EDTA, pH 7.2) was added to 15 mg of RNA.Denaturation was carried out at 65°C for 5 min, followed by cooling on ice.Before the mixture was loaded onto the gels, 2 ml of loading buffer was added.Digoxigenin-labeled pSPTNeo RNA (Roche Molecular Biochemicals) wastreated as described above and used as a size marker on the gels. The RNA wastransferred by capillary blotting in SSC buffer (3 M NaCl, 300 mM sodiumcitrate, pH 7), fixed to positively charged nylon membranes (Roche MolecularBiochemicals), and fixed by UV irradiation. Prehybridization, hybridization(42°C for 16 h), and chemiluminescent detection were carried out essentially asprescribed by Boehringer (The DIG System User’s Guide for Filter Hybridization,Boehringer Mannheim, Mannheim, Germany, 1993). In order to study the reg-ulation of FCSD by sulfide, it was necessary to use 0.05 mg of cysteine/ml in thegrowth medium to maintain anaerobiosis.

RESULTS AND DISCUSSION

Isolation of cytochrome c-552. The soluble electron transferproteins of E. vacuolata strain b1 (DSM 2111) are very similarto those of Ectothiorhodospira shaposhnikovii (26, 27). Thereare at least two HiPIP isozymes and cytochromes c9, c4, and b5(unpublished work). Following buffer solubilization of theseproteins, we extracted the membrane fraction with butanol toidentify peripheral electron transfer proteins. AdditionalHiPIP and cytochrome c4 were released, along with a smallcytochrome c-552 that cochromatographed with a third HiPIPisozyme from which it was difficult to separate it. (We have also

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found four soluble HiPIP isozymes as well as cytochrome c4 inthe related species Ectothiorhodospira mobilis [unpublished]).In addition, an abundant, high-molecular-weight cytochromec-553 was easily separated by gel filtration. This protein has notbeen further characterized but is likely to be the tetrahemereaction center cytochrome, based upon its general occurrencein purple bacteria and its membrane localization. Cytochromec-552 was purified as described in Materials and Methods.

Amino acid sequence analysis of the solubilized cytochromec-552. The complete amino acid sequence of the small cyto-chrome c-552 was determined as shown in Fig. 1. Cytochromec-552 contains 77 residues and a single heme binding site nearthe N terminus. The sequence is clearly that of a class I cyto-chrome and is most closely related to the 86-residue cyto-chrome subunit of C. limicola FCSD (49% identity with nointernal insertions or deletions) (48) (Fig. 2). It is 26% iden-tical to the first half, including one gap (a gap is defined as aninsertion or deletion), and 19% identical (with four gaps) tothe second half of the 174-residue diheme cytochrome subunit

of C. vinosum FCSD (47). These results strongly suggested thepresence of an associated flavoprotein subunit in E. vacuolata.

Gene sequence of the FCSD locus. Based upon the aminoacid sequence of cytochrome c-552, we obtained a subgenomicclone of 5.5 kb. The translated cytochrome gene contains a28-residue signal sequence and has a normal Ala-Thr-Ala rec-ognition sequence for cleavage by the signal peptidase (51).The derived mature protein sequence was identical to that ofthe cytochrome sequenced by Edman degradation. Sixteenbases downstream of the cytochrome gene is a 1,290-base openreading frame which encodes a 430-residue protein homolo-gous to the flavoprotein subunit of Chromatium FCSD. Thereis a 33-residue signal peptide which does not have an obviouscleavage site. It thus appears that the leader is not cleaved oris cleaved in front of Cys 25, to which a diacyl glycerol may beattached, as is the case in lipoproteins (33). The flavoproteinsubunit is 50% identical to that of Chromatium, and there areonly six small gaps in the protein sequence alignment (Fig. 3).The FCSD of Thiobacillus sp strain W5 is membrane bound

FIG. 1. Amino acid sequence of the cytochrome subunit of flavocytochrome c from E. vacuolata. The N-terminal sequences of the apoprotein and the modifiedapoprotein are indicated by N-apo and N-mod, respectively. Peptides obtained after cleavage with S. aureus protease or after partial acid hydrolysis are named Ec andAH, respectively. The arrows indicate residues chemically identified during Edman degradation. An asterisk means that the sequence analysis was deliberately stopped.Mass spectrometry was used to determine the molecular weight of each of the peptides (results not shown).

FIG. 2. Alignment of the amino acid sequences of the FCSD E. vacuolata (1) and C. limicola f. sp. thiosulfatophilum (2) monoheme cytochrome subunits (48), withthe diheme cytochrome subunit of C. vinosum (47), first half (3) and second half (4). The amino acids in boldface represent the heme binding site.

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and has a small monoheme cytochrome subunit, suggestingclose similarity to the E. vacuolata protein (50).

The FCSD polypeptide chain is predicted to fold in threedomains (12), which are comparable to those of glutathionereductase. The first domain in the latter protein binds flavin

adenine dinucleotide (FAD), the second domain binds a pyri-dine nucleotide, and the third domain provides the subunitinterface. Although the second domain is present in FCSD, itdoes not interact with pyridine nucleotides because of thepresence of a unique disulfide bond between Cys 161 and Cys

FIG. 3. Alignment of the complete amino acid sequences of the FCSD flavoprotein subunits from E. vacuolata (1), C. vinosum (49) (2), P. denitrificans (52) (partial)(3), and A. aeolicus (14) (starts with position 40) (4). The boxed amino acids represent more than 75% conserved positions.

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337 that blocks access to that side of the FAD. The functionalrole of this domain in FCSD is unknown. SQR apparently hasthe same three domains and the same disulfide found in FCSD.The SQR gene is not closely associated with a cytochrome, butSQR interacts with membrane proteins, presumably wherequinone is reduced, and this binding is likely to be mediated bythe third domain, which in the FCSD flavoprotein binds thecytochrome c subunit.

FCSD is a membrane protein. Since no FCSD was found inthe soluble extracts and only the cytochrome subunit was sol-ubilized with butanol, we attempted to confirm that FCSD waslocalized in the membrane fraction by extraction with the de-tergent Triton X-100. The purification protocol yielded a smallamount of partially purified FCSD. SDS-PAGE of the enzymeshowed two abundant protein bands with molecular masses of46 and 10 kDa. Heme staining revealed that the 10-kDa bandwas the cytochrome c-552 subunit (Fig. 4). The spectra of thedithionite-reduced enzyme showed the characteristic maximafor cytochrome c, 417 (g band), 524 (b band), and 552 (a band)nm, similar to those of the butanol-solubilized cytochrome.The oxidized spectrum did not display shoulders at 450 and 480nm or bleaching with dithionite, which is typical of FCSDflavin. This may be due to modification of the native enzymeduring the treatment with Triton X-100. Immunoblotting ofthe solubilized membrane fraction and of the soluble proteinfraction clearly indicated that the flavoprotein subunit reactedwith an antibody against the C. vinosum flavoprotein. It waspresent in the membrane fraction but not in the soluble frac-tion (Fig. 4B).

Flanking genes. Two possible open reading frames werefound adjacent to the E. vacuolata FCSD genes. Upstream, inthe same orientation, there is the 39 end of a gene for a132-residue-long protein which by BLAST search appears tobe homologous to the htrB gene of E. coli. The HtrB genecodes for lauroyl acyltransferase involved in the biosynthesis ofthe outer membrane lipid A (13). It is separated from theFCSD cytochrome subunit gene by 558 bases. The HtrB geneproduct is also a heat shock protein required for cell viability athigh temperature in E. coli and is present in Haemophilus

influenzae (29) as well. Separated by 236 bases downstream ofthe flavoprotein subunit gene, and in the opposite orientation,there is a large open reading frame (orf4) encoding a protein ofat least 567 amino acid residues. It is unclear where it begins,but the initiator codon seems to be a TTG triplet starting atbase 4313, 9 bases downstream of a possible ribosome bindingsite (GGAG). A BLAST search shows that it has a largenumber of homologs. The strongest similarity is to four pro-teins derived from Synechocystis (SLR359, SLL267, SLR1305,and SLR2077) (23), with more than 40% identity. The func-tional roles of these proteins are unknown, but the orf4 productis also related to a lesser extent to diguanylate cyclases andphosphodiesterases. These proteins show the greatest conser-vation in the C-terminal 400 residues, which contain charac-teristic GGDEF and EAL motifs (44). It is remarkable that agene similar to orf4 is located downstream of R. capsulatusSQR in the same opposing orientation as in E. vacuolata. Thiscontext is quite different from that of the FCSD gene in Chro-matium, where it was found that the flavoprotein gene is sep-arated from the cytochrome gene by only 15 nucleotides (15,35). Upstream, a tetraheme cytochrome and a homolog ofankyrin were found. The tetraheme cytochrome is part of amultigene locus in Thiosphaera pantotropha, H. influenzae, andAlcaligenes eutrophus (8, 17, 42) which contains several elec-tron transfer proteins involved in nitrate reduction. Ankyrinserves to bind proteins together and/or to bind them to themembrane (9). Downstream of the Chromatium FCSD gene,there are no genes located in the 446 bases that were se-quenced. Thus, the FCSD gene is probably not part of a mul-tigene operon in Chromatium and E. vacuolata because of thelack of conservation of flanking genes and the small sizes of thetranscripts (see below).

Influence of sulfur compounds on the growth of E. vacuolata.FCSD transcription and expression in E. vacuolata was studiedby examining growth in different media. The best growth wasobtained under photoheterotrophic conditions in DSMZ me-dium combining acetate as a carbon source and sulfide as anelectron donor. These results are in agreement with the reportof Zakharchuk and Ivanovskii (53), who described increasedassimilation of 14C-acetate in E. shaposhnikovii cells in thepresence of thiosulfate, sulfide, and bicarbonate. To study theinfluence of carbon sources on photosynthetic activity, cellswere grown on minimal medium supplemented with acetate,succinate, or malate, all of which supported growth. Removalof sulfide, however, reduced growth compared to that in aphotomixotrophic medium, as did replacing acetate with so-dium malate or succinate. RNA was extracted from these cellsand analyzed by Northern blotting with probes specific for boththe heme and the flavoprotein subunits of FCSD. The size andthe hybridization pattern were the same with both probes. Thetranscript of about 3 kb corresponds to an operon that containsthe cytochrome and flavoprotein genes but no others. TheChromatium FCSD transcript is also just large enough to in-clude the two subunit genes (15). The intensity of the RNAtranscripts increased in cells grown on sulfide or thiosulfateplus CO2 or on sulfide plus acetate. In view of the fact thatsome sulfide can be produced from thiosulfate (10, 11), weconclude that sulfide induces FCSD expression (Fig. 5). In theabsence of sulfide or thiosulfate, FCSD expression is signifi-cantly reduced, confirming the role of sulfide for FCSD induc-tion under anoxygenic photosynthesis.

Functional importance of conserved residues in FCSD pro-teins. Several residues in the flavoprotein were identified fromthe three-dimensional structure as having possible functionalimportance (12). The FAD is covalently bound to Cys 42(Chromatium numbering), and this residue is conserved in E.

FIG. 4. (A) SDS-PAGE profiles of FCSD from E. vacuolata. Lane 1, heme-stained cytochrome c-552; lane 2, Coomassie blue-stained flavoprotein and cy-tochrome c-552 subunits solubilized with Triton X-100 and partially purified asdescribed in the text. (B) Localization of E. vacuolata FCSD in the membraneprotein fraction. The Western blot was performed using C. vinosum flavoproteinsubunit antibodies. Lanes: 1, E. vacuolata soluble protein fraction; 2, E. vacuolatamembrane protein fraction; 3, C. vinosum FCSD (1 mg of native protein).

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vacuolata. Glu 167 is conserved in nearly all the proteins ofthe glutathione reductase family of enzymes (37), includingthe Chromatium and E. vacuolata FCSDs, and is locatednear the N5 position of the FAD. In FCSD, Glu 167 modulatesthe reactivity of the FAD with sulfite when ionized at pH 6 andpresumably has a role to play in the oxidation of sulfide. Thedisulfide Cys 161-Cys 337 is adjacent to the FAD; it is con-served and also modulates the reactivity of the FAD withsulfite when it is cleaved by sulfite above pH 8.5. Incidentally,this disulfide is also present in SQR (38), suggesting that it maybe essential for the oxidation of sulfide. Trp 128, Tyr 306, andTrp 391 are all near the flavin, and any one of them could actas the charge transfer acceptor for the flavin-sulfite adduct.Trp 128 and Trp 391 are conserved, but Tyr 306 is replaced byHis in E. vacuolata. This should affect the redox potential ofthe FAD in a pH-dependent manner because of its location atthe positive N-terminal end of the helix. Thus, FAD may havea higher redox potential at low pH when the His is protonated,provided that the charge on the helix does not prevent Hisprotonation within the physiological range of pH. A higherpotential should make the FAD more reactive with sulfite andother nucleophiles.

The results presented here establish that an FCSD is presentin E. vacuolata, thus further expanding the distribution of thisprotein to a third family of photosynthetic sulfur bacteria.Notably, the genetic contexts of the E. vacuolata and Chroma-tium FCSDs are quite distinct, indicating that the FCSD geneis not part of a multigene operon. Importantly, it is quite clearthat E. vacuolata FCSD is a membrane-bound as opposed to asoluble protein as in other species of green and purple photo-synthetic bacteria; thus, it may be present in other species andnot detected in a soluble form. The E. vacuolata FCSD isclearly regulated by sulfide, although the Chromatium FCSD isnot. This is consistent with the view that FCSD functions invivo as sulfide dehydrogenase in E. vacuolata if not in allspecies of sulfur bacteria in which it is found.

ACKNOWLEDGMENTS

J.V.B. is indebted to the Fonds voor Wetenschappelijk Onderzoek-Vlaanderen for research projects G.0068.96 and G.0054.97. This workwas also supported by grant GM 21277 from the National Institutes ofHealth to M.A.C.

We acknowledge D. Brune for helpful discussions and for correctionof the manuscript.

REFERENCES

1. Ambler, R. P., and M. Wynn. 1973. The amino acid sequences of cyto-chromes c-551 from three species of Pseudomonas. Biochem. J. 131:485–498.

2. Arieli, B., Y. Shahak, D. Taglicht, G. Hauska, and E. Padan. 1994. Purifi-cation and characterization of sulfide-quinone reductase (SQR), a novelenzyme driving anoxygenic photosynthesis in Oscillatoria limnetica. J. Biol.Chem. 269:5705–5711.

3. Ausubel, F. M., R. Bocut, R. E. Kingston, D. D. Moore, J. G. Seidman, J. A.Smith, and K. Strohl (ed.). 1990. Current protocols in molecular biology.Wiley Interscience, New York, N.Y.

4. Bartsch, R. G. 1978. Cytochromes, p. 249–279. In R. K. Clayton and W. R.Sistrom (ed.), The photosynthetic bacteria. Plenum Press, New York, N.Y.

5. Bartsch, R. G. 1991. The distribution of soluble metallo-redox proteins inpurple phototrophic bacteria. Biochim. Biophys. Acta 1058:28–30.

6. Bartsch, R. G., and M. D. Kamen. 1960. Isolation and properties of twosoluble heme proteins in extracts of the photoanaerobe Chromatium. J. Biol.Chem. 235:825–831.

7. Bartsch, R. G., T. E. Meyer, and A. B. Robinson. 1968. Complex c-typecytochromes with bound flavin, p. 443–451. In K. Okunuki, M. D. Kamen,and I. Sekuzu (ed.), Structure and function of cytochromes. University ofTokyo Press, Tokyo, Japan.

8. Berks, B. C., D. J. Richardson, A. Reilly, A. C. Willis, and S. J. Ferguson.1995. The napEDABC gene cluster encoding the periplasmic nitrate reduc-tase system of Thiosphaera pantotropha. Biochem. J. 309:983–992.

9. Bork, P. 1993. Hundreds of ankyrin-like repeats in functionally diverse pro-teins: mobile modules that cross phyla horizontally? Proteins 17:363–374.

10. Brune, D. C. 1989. Sulfur oxidation by phototrophic bacteria. Biochim.Biophys. Acta 973:189–221.

11. Brune, D. C. 1995. Sulfur compounds as photosynthetic electron donors, p.847–870. In R. E. Blankenship, M. T. Madigan, and C. E. Bauer (ed.),Anoxygenic photosynthetic bacteria. Kluwer Academic Publishers, NewYork, N.Y.

12. Chen, Z. W., M. Koh, G. Van Driessche, J. J. Van Beeumen, R. G. Bartsch,T. E. Meyer, M. A. Cusanovich, and F. S. Mathews. 1994. The structure offlavocytochrome c sulfide dehydrogenase from a purple phototrophic bacte-rium. Science 266:430–432.

13. Clementz, T., J. J. Bednarski, and C. R. Raetz. 1996. Function of the htrBhigh temperature requirement gene of Escherichia coli in the acylation oflipid A: HtrB catalyzed incorporation of laurate. J. Biol. Chem. 271:12095–12102.

14. Deckert, G., P. V. Warren, T. Gaasterland, W. G. Young, A. L. Lenox, D. E.Graham, R. Overbeek, M. A. Snead, M. Keller, M. Aujay, R. Huber, R. A.Feldman, J. M. Short, G. J. Olsen, and R. V. Swanson. 1998. The completegenome of the hyperthermophilic bacterium Aquifex aeolicus. Nature 392:353–358.

15. Dolata, M. M., J. J. Van Beeumen, R. P. Ambler, T. E. Meyer, and M. A.Cusanovich. 1993. Nucleotide sequence of the heme subunit of flavocyto-chrome c from the purple phototrophic bacterium, Chromatium vinosum.J. Biol. Chem. 268:14426–14431.

16. Fischer, U. 1989. Characterization of two soluble basic cytochromes isolatedfrom the anoxygenic phototrophic sulfur bacterium Chlorobium phaeobacte-roides. Z. Naturforsch. Sect. C 44:71–76.

17. Fleischmann, R. D., M. D. Adams, O. White, R. A. Clayton, E. F. Kirkness,A. R. Kerlavage, C. J. Bult, J. F. Tomb, B. A. Dougherty, and J. M. Merrick.1995. Whole-genome random sequencing and assembly of Haemophilus in-fluenzae Rd. Science 269:496–512.

18. Friedrich, C. G. 1998. Physiology and genetics of sulfur-oxidizing bacteria.Adv. Microb. Physiol. 39:235–289.

19. Fukumori, Y., and T. Yamanaka. 1979. Flavocytochrome c of Chromatiumvinosum. J. Biochem. 85:1405–1414.

20. Garcia-Castillo, M. C., B. S. Lou, M. R. Ondrias, D. E. Robertson, and D. B.Knaff. 1994. Characterization of flavocytochrome c-552 from the thermo-philic photosynthetic bacterium Chromatium tepidum. Arch. Biochem. Bio-phys. 315:262–266.

21. Imhoff, J. F., J. Suling, and R. Petri. 1998. Phylogenetic relationships amongthe Chromateaceae, their taxonomic reclassification and description of thenew genera Allochromatium, Halochromatium, Isochromatium, Marichroma-tium, Thiococcus, Thiohalocapsa, and Thermochromatium. Int. J. Syst. Bac-teriol. 48:1129–1143.

22. Jue, R. A., and H. E. Hale. 1993. Identification of cysteine residues alkylatedwith 3-bromopropylamine by protein sequence analysis. Anal. Biochem. 210:39–44.

23. Kaneko, T., S. Sato, H. Kotani, A. Tanaka, E. Asamizu, Y. Nakamura, N.Miyajima, M. Hirosawa, M. Sugiura, S. Sasamoto, T. Kimura, T. Hosouchi,A. Masuno, A. Muraki, N. Nakazaki, K. Naruo, S. Okumura, S. Shimpo, C.Takeuchi, T. Wada, A. Watanabe, M. Yamada, M. Yasuda, and S. Tabata.1996. Sequence analysis of the genome of the unicellular cyanobacteriumSynechocystis sp. strain PCC6803. II. Sequence determination of the entiregenome and assignment of potential protein-coding regions. DNA Res.3:109–136.

24. Kerfeld, C. A., C. Dhan, M. Hirasawa, S. Kleis-San Francisco, T. O. Yeates,and D. B. Knaff. 1996. Isolation and characterization of soluble electron

FIG. 5. Northern blot of RNA isolated from E. vacuolata. Total RNA wasisolated from a 100-ml culture growing exponentially in minimal Pfennig mediumsupplemented with the following compounds: malate (lane 1), sodium thiosulfate(lane 2), sodium sulfide (lane 3), sodium sulfide and acetate (lane 4), acetate(lane 5), and succinate (lane 6). Fifty micrograms of RNA was loaded per lane.Northern hybridization was performed with a digoxigenin-labeled probe ampli-fied with the fcsd gene template. Note the elevated levels of FCSD transcript inthe cultures containing sulfide.

3102 KOSTANJEVECKI ET AL. J. BACTERIOL.

on May 28, 2018 by guest

http://jb.asm.org/

Dow

nloaded from

transfer proteins from Chromatium purpuratum. Biochemistry 35:7812–7818.25. Kusai, A., and T. Yamanaka. 1973. Cytochrome c (553, Chlorobium thio-

sulfatophilum) is a sulfide-cytochrome c reductase. FEBS Lett. 34:235–237.26. Kusche, W. H., and H. G. Truper. 1984. Iron sulfur proteins of the purple

sulfur bacterium Ectothiorhodospira shaposhnikovii. Arch. Microbiol. 137:266–271.

27. Kusche, W. H., and H. G. Truper. 1984. Cytochromes of the purple sulfurbacterium Ectothiorhodospira shaposhnikovii. Z. Naturforsch. Sect. C 39:894–901.

28. Laemlli, U. K. 1970. Cleavage of structural proteins during the assembly ofthe head of bacteriophage T4. Nature 227:680–685.

29. Lee, N.-G., M. G. Sunshine, J. J. Engstrom, B. W. Gibson, and M. A.Apicella. 1995. Mutation of the htrB locus of Haemophilus influenzae non-typeable strain 2019 is associated with modifications of lipid A and phos-phorylation of the lipo-oligosaccharide. J. Biol. Chem. 270:27151–27159.

30. Meyer, T. E., R. G. Bartsch, M. A. Cusanovich, and J. H. Mathewson. 1968.The cytochromes of Chlorobium thiosulfatophilum. Biochim. Biophys. Acta153:854–861.

31. Meyer, T. E., W. P. Vorkink, G. Tollin, and M. A. Cusanovich. 1985. Chro-matium flavocytochrome c—kinetics of reduction of the heme subunit andthe flavocytochrome mitochondrial cytochrome c complex. Arch. Biochem.Biophys. 236:52–58.

32. Myers, C. R., and J. M. Myers. 1992. Localization of cytochromes to theouter membrane of anaerobically grown Shewanella putrefaciens MR-1. J.Bacteriol. 147:3429–3438.

33. Myers, J. M., and C. R. Myers. 1998. Isolation and sequence of omcA, a geneencoding a decaheme outer membrane cytochrome c of Shewanella putrefa-ciens MR-1, and detection of omcA homologs in other strains of S. putrefa-ciens. Biochim. Biophys. Acta 1373:237–251.

34. Pattaragulwanit, K., D. C. Brune, G. Truper, and C. Dahl. 1998. Moleculargenetic evidence for extracytoplasmic localization of sulfur globules in Chro-matium vinosum. Arch. Microbiol. 169:434–444.

35. Reinartz, M., J. Tschape, T. Bruser, H. G. Truper, and C. Dahl. 1998. Sulfideoxidation in the phototrophic sulfur bacterium Chromatium vinosum. Arch.Microbiol. 170:59–68.

36. Sambrook, J., E. F. Fritsch, and T. Maniatis. 1989. Molecular cloning: aLaboratory Manual, 2nd ed. Cold Spring Harbor Laboratory, Cold SpringHarbor, N.Y.

37. Schulz, G. E., and P. A. Karplus. 1987. Enzymes with active-site redox-activedisulfide bonds. Biochem. Soc. Trans. 16:81–84.

38. Schutz, M., I. Maldener, C. Griesbeck, and G. Hauska. 1999. Sulfide-qui-none reductase from Rhodobacter capsulatus: requirement for growth,periplasmic localization, and extension of gene sequence analysis. J. Bacte-riol. 181:6516–6523.

39. Schutz, M., Y. Shahak, E. Padan, and G. Hauska. 1997. Sulfide-quinonereductase from Rhodobacter capsulatus. J. Biol. Chem. 272:9890–9894.

40. Shahak, Y., B. Arieli, E. Padan, and G. Hauska. 1992. Sulfide quinonereductase (SQR) activity in Chlorobium. FEBS Lett. 299:127–130.

41. Shahak, Y., M. Schutz, M. Bronstein, C. Griesbeck, G. Hauska, and E.Padan. 1999. Sulfide-dependent anoxygenic photosynthesis in prokaryotes,

p. 217–228. In G. A. Peschek, W. Loffelhardt, and G. Schmetterer (ed.), Thephototrophic prokaryotes. Kluwer Academic Publishers, New York, N.Y.

42. Siddiqui, R. A., U. Warnecke-Eberz, A. Hengsberger, B. Schneider, S. Kost-ka, and B. Friedrich. 1993. Structure and function of a periplasmic nitratereductase in Alcaligenes eutrophus H16. J. Bacteriol. 175:5867–5876.

43. Steinmetz, M. A., and U. Fischer. 1981. Cytochromes of the non-thiosulfate-utilizing green sulfur bacterium Chlorobium limicola. Arch. Microbiol. 130:31–37.

44. Tal, R., H. C. Wong, R. Calhoon, D. Gelfand, A. L. Fear, G. Volman, R.Mayer, P. Ross, D. Amikam, H. Weinhouse, A. Cohen, S. Sapir, P. Ohana,and M. Benziman. 1998. Three cdg operons control cellular turnover ofcyclic di-GMP in Acetobacter xylinum: genetic organization and occurrenceof conserved domains in isoenzymes. J. Bacteriol. 180:4416–4425.

45. Thomas, P. E., D. Ryan, and W. Ledwin. 1976. An improved staining pro-cedure for the detection of the peroxidase activity of cytochrome P-450 onsodium dodecyl sulfate polyacrylamide gels. Anal. Biochem. 75:168–176.

46. Tissieres, A. 1956. Purification and properties and the specific biologicalactivity of cytochromes c4 and c5 from Azotobacter vinelandii. Biochem. J.64:582–589.

47. Van Beeumen, J. J., H. Demol, B. Samyn, R. G. Bartsch, T. E. Meyer, M. M.Dolata, and M. A. Cusanovich. 1991. Covalent structure of the dihemecytochrome subunit and amino terminal sequence of the flavoprotein subunitof flavocytochrome c from Chromatium vinosum. J. Biol. Chem. 266:12921–12931.

48. Van Beeumen, J. J., S. Van Bun, T. E. Meyer, R. G. Bartsch, and M. A.Cusanovich. 1990. Complete amino acid sequence of the cytochrome subunitand amino-terminal sequence of the flavin subunit of flavocytochrome c(sulfide dehydrogenase) from Chlorobium thiosulfatophilum. J. Biol. Chem.265:9793–9799.

49. Van Driessche, G., M. Koh, Z. W. Chen, F. S. Mathews, T. E. Meyer, R. G.Bartsch, M. A. Cusanovich, and J. J. Van Beeumen. 1996. Covalent structureof the flavoprotein subunit of the flavocytochrome c: sulfide dehydrogenasefrom the purple phototrophic bacterium Chromatium vinosum. Protein Sci.5:1753–1764.

50. Visser, M., G. A. H. de Jong, L. A. Robertson, and J. G. Kuenen. 1997. Anovel membrane-bound flavocytochrome c sulfide dehydrogenase from thecolourless sulfur bacterium Thiobacillus sp. W5. Arch. Microbiol. 167:295–301.

51. Von Heijne, G. 1992. Membrane protein structure prediction, hydrophobicityanalysis and the positive-inside rule. J. Mol. Biol. 225:487–495.

52. Wodara, C., F. Bardischewsky, and C. G. Friedrich. 1997. Cloning andcharacterization of sulfite dehydrogenase, two c-type cytochromes, and aflavoprotein of Paracoccus denitrificans GB17: essential role of sulfite dehy-drogenase in lithotrophic sulfur oxidation. J. Bacteriol. 179:5014–5023.

53. Zakharchuk, L. M., and R. N. Ivanovskii. 1980. Acetate metabolism inEctothiorhodospira shaposhnikovii growing in dark. Mikrobiologiia 49:14–19.

54. Zorin, N. A., and I. N. Gogotov. 1980. Purification and properties of cyto-chrome c-552 from purple sulfur bacterium Thiocapsa roseopersicina. Bio-khimia 45:1134–1138. (English translation.)

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