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Originally published In Press as doi:10.1074/jbc.M404417200 on August 18, 2004

J. Biol. Chem., Vol. 279, Issue 44, 45485-45494, October 29, 2004
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Hydroxylamine Assimilation by Rhodobacter capsulatus E1F1

REQUIREMENT OF THE hcp GENE (HYBRID CLUSTER PROTEIN) LOCATED IN THE NITRATE ASSIMILATION nas GENE REGION FOR HYDROXYLAMINE REDUCTION*

Purificación Cabello{ddagger}§, Carmen Pino§, M. Francisca Olmo-Mira§||, Francisco Castillo¶, M. Dolores Roldán¶**, and Conrado Moreno-Vivián¶{ddagger}{ddagger}

From the {ddagger}Departamento de Biología Vegetal, Área de Fisiología Vegetal, Edificio Celestino Mutis, 3a planta, and the Departamento de Bioquímica y Biología Molecular, Edificio Severo Ochoa, 1a planta, Campus Universitario de Rabanales, Universidad de Córdoba, 14071-Córdoba, Spain

Received for publication, April 21, 2004 , and in revised form, August 16, 2004.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Rhodobacter capsulatus E1F1 grows phototrophically with nitrate as nitrogen source. Using primers designed for conserved motifs in bacterial assimilatory nitrate reductases, a 450-bp DNA was amplified by PCR and used for the screening of a genomic library. A cosmid carrying an insert with four SalI fragments of 2.8, 4.1, 4.5, and 5.8 kb was isolated, and DNA sequencing revealed that it contains a nitrate assimilation (nas) gene region, including the hcp gene coding for a hybrid cluster protein (HCP). Expression of hcp is probably regulated by a nitrite-sensitive repressor encoded by the adjacent nsrR gene. A His6-HCP was overproduced in Escherichia coli and purified. HCP contained about 6 iron and 4 labile sulfide atoms per molecule, in agreement with the presence of both [2Fe-2S] and [4Fe-2S-2O] clusters, and showed hydroxylamine reductase activity, forming ammonia in vitro with methyl viologen as reductant. The apparent Km values for NH2OH and methyl viologen were 1 mM and 7 µM, respectively, at the pH and temperature optima (9.3 and 40 °C). The activity was oxygen-sensitive and was inhibited by sulfide and iron reagents. R. capsulatus E1F1 grew phototrophically, but not heterotrophically, with 1 mM NH2OH as nitrogen source, and up to 10 mM NH2OH was taken up by anaerobic resting cells. Ammonium was transiently accumulated in the media, and its assimilation was prevented by L-methionine-D,L-sulfoximine, a glutamine synthetase inhibitor. In addition, hydroxylamine- or nitrite-grown cells showed the higher hydroxylamine reductase activities. However, R. capsulatus B10S, a strain lacking the whole hcp-nas region, did not grow with 1 mM NH2OH. Also, E. coli cells overproducing HCP tolerate hydroxyl-amine better during anaerobic growth. These results suggest that HCP is involved in assimilation of NH2OH, a toxic product that could be formed during nitrate assimilation, probably in the nitrite reduction step.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Bacteria use nitrate as a nitrogen source for growth, as a terminal electron acceptor for anaerobic respiration, or as an electron sink for redox balancing. Three different types of bacterial nitrate-reducing systems have been described: the cytoplasmic assimilatory Nas, the membrane-bound respiratory Nar, and the periplasmic dissimilatory Nap (1, 2). Nitrate assimilation is a key process of the nitrogen cycle that has been an object of biochemical and genetic research in higher plants, fungi, algae, and cyanobacteria, although it has only been scarcely studied in other bacteria. It is firmly established that the assimilatory nitrate-reducing system consists of a nitrate transport system and two metalloenzymes, the assimilatory nitrate and nitrite reductases, which catalyze the stepwise reduction of nitrate to nitrite and ammonium (1–4). Nitrate assimilation is usually regulated by nitrate/nitrite induction and ammonium repression. Genes coding for the assimilatory nitrate-reducing systems are normally clustered and have been cloned in several bacteria. These gene clusters code for regulatory and structural proteins required for uptake and reduction of both nitrate and nitrite (1–3). Only a few strains of purple phototrophic bacteria assimilate nitrate. Rhodobacter capsulatus E1F1 uses nitrate or nitrite as a nitrogen source, but only under photoanaerobic conditions because nitrite or hydroxylamine are toxic for heterotrophic growth (5, 6). The purified assimilatory nitrate reductase of this strain is composed of a 95-kDa catalytic subunit with iron-sulfur and molybdenumbis-molybdopterin guanine dinucleotide cofactors, and a 46-kDa FAD-containing diaphorase (7).

HCPs1 are soluble iron-sulfur proteins first described in the strictly anaerobic sulfate-reducing species Desulfovibrio desulfuricans and Desulfovibrio vulgaris. Initial EPR and Mössbauer spectroscopy suggested the presence of a [6Fe-6S] center, and the protein was termed "prismane" (8–13). However, the crystallographic structures at high resolution of the D. desulfuricans and D. vulgaris HCPs reveal that they do not contain a [6Fe-6S] cluster but instead have two types of iron-sulfur centers: cluster 1, a typical cubane [4Fe-4S] coordinated by four Cys residues located in the N-terminal domain, and cluster 2, a novel hybrid [4Fe-2S-2O] center located at the interface of the three protein domains (14–17). An unusual feature of the hybrid center is that it has four different oxidation states within the biologically relevant range of redox potentials. EPR spectra of the reduced HCP from Escherichia coli indicate the presence of an N-terminal [2Fe-2S] center instead of a [4Fe-4S] cluster (18). The Cys motif for binding this cluster 1 in Desulfovibrio and other strictly anaerobic bacteria has the C-X2-C-X7–8-C-X5-C spacing, whereas in E. coli and other facultative anaerobic bacteria has the sequence C-X2-C-X11-C-X6-C (18, 19).

Genes encoding HCP proteins have been identified in a large number of both strict anaerobic and facultative anaerobic bacteria and Archaea. However, hcp genes are not present in the completely sequenced genomes of some prokaryotes. In E. coli (18), Thiobacillus ferrooxidans (20). and Salmonella enterica (21), a gene coding for a putative NADH oxidoreductase (hcr) is located immediately downstream of the hcp gene, and the HCR protein of E. coli catalyzes reduction of HCP in the presence of NADH (18). However, hcr genes are not found near the hcp genes present in the genomes of other bacteria and Archaea.

Despite the fact that HCP has received a lot of attention due to the unusual properties of its hybrid cluster, the regulation and the physiological function of this protein remain enigmatic. As the E. coli and Morganella morganii HCPs are only detected in cells grown anaerobically with nitrate or nitrite, it has been suggested that HCP have a role in nitrate and/or nitrite respiration (18). A similar increase in the expression of hcp gene was observed during anaerobic growth with nitrate in Shewanella oneidensis (22), and a nitrite-inducible Fnr-dependent promoter was identified upstream of nipAB, the hcp-hcr homologues of S. enterica (21). However, in Clostridium perfringens HCP has been proposed to be involved in the adaptive response to oxidative stress (23). Recently, it has been reported that the E. coli HCP protein exhibits an oxygen-sensitive hydroxylamine reductase activity (24). Similarity of the HCP proteins to the FeNiS-containing carbon monoxide dehydrogenases has also been observed, and interestingly, a variant of Rhodospirillum rubrum CO dehydrogenase carrying a H265V amino acid substitution acquires hydroxylamine reductase activity (25). EPR and crystallographic studies also suggest that HCP could act as a reductase, although its physiological role as hydroxylamine reductase has been questioned (17).

In this work we describe for the first time the capability of R. capsulatus E1F1 to growth with hydroxylamine as the sole nitrogen source under phototrophic conditions, and the cloning of a nitrate assimilation (nas) gene region that includes a hcp gene, which is probably regulated by a nitrite-sensitive repressor encoded by the adjacent nsrR gene. A recombinant His6-tagged HCP protein has been expressed in E. coli, and the hydroxylamine reductase activity of the purified protein has been characterized. Evidence that the HCP protein is responsible for the in vivo hydroxylamine reduction is also presented.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 REFERENCES
 
Bacterial Strains, Plasmids, and Growth Conditions—The bacterial strains and plasmids used in this work are listed in Table I. R. capsulatus was grown at 30 °C on peptone-yeast extract plates or RCV minimal medium (26) under phototrophic (light-anaerobiosis) or heterotrophic (dark-aerobiosis) conditions, as previously described (31). D,L-Malate (4 g liter–1) was routinely used as the carbon source, and glutamate, KNO3, NH4Cl (1 g liter–1 each), KNO2 or NH2OH (1 mM each) were used as the nitrogen sources. E. coli strains were grown aerobically in Luria-Bertani (LB) liquid or solid media at 37 °C (28) or anaerobically in the LB medium with 10 mM KNO3 and 0.4% glycerol.


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  		TABLE I
Bacterial strains and plasmids used in this work

 
Preparation of Cell-free Extracts—Cells were harvested by centrifugation at 20,000 x g for 15 min, washed, and resuspended in 50 mM Tris-HCl buffer (pH 8.0), and broken by cavitation at 90 W for 15 s (3 pulses of 5 s) in a VibraCell sonifier. The extracts were placed into anaerobic tubes under argon atmosphere, centrifuged again at 20,000 x g for 15 min, and the resulting supernatant was kept anaerobically at –20 °C until use.

Analytical Determinations and Enzyme Assays—Cell growth was followed turbidimetrically measuring the absorbance of the cultures at 680 nm for R. capsulatus or at 600 nm for E. coli strains. Ammonium was determined by the phenol-hypochlorite method (32) using the microdiffusion technique as described by Wolfe et al. (24). Hydroxylamine was measured by the pyridoxal 5'-phosphate reaction as previously described (33). Protein was estimated according to Lowry et al. (34) or Bradford (35), using bovine serum albumin as standard. Labile sulfur determination in the purified HCP was performed by reaction with N,N-dimethyl-p-phenylenediamine and FeCl3 following the described methods (36) and using a calibration curve with Na2S·9H2O in NaOH (under nitrogen atmosphere) as standard. Labile iron was determined in samples heated at 80 °C for 10 min by reaction with sodium methasulfite and bathophenanthroline as previously described (37), using a calibration curve with FeSO4·7H2O as standard. Hydroxylamine reductase activity was routinely assayed anaerobically (argon atmosphere) measuring the ammonia generated in the reaction with reduced methyl viologen as artificial electron donor (24). The assay mixture contained, per ml, 100 µmol of CHES (pH 9.3), 10 nmol of EDTA, 0.5 µmol of methyl viologen, 10 µmol of sodium dithionite, 10 µmol of NH2OH, and the adequate amount of cell extract or HCP protein. When required, other electron donors were used, or the assays were carried out in the presence of possible enzyme inhibitors at the indicated concentrations. NADH-dependent hydroxylamine reductase activity in R. capsulatus cell-free extracts was assayed by following NADH oxidation at 340 nm in anaerobic cuvettes containing, per ml, 100 µmol of CHES (pH 9.3), 10 nmol of EDTA, 0.25 µmol of NADH, 10 µmol of NH2OH, and the adequate amount of cell extract. The apparent Km values for NH2OH and methyl viologen were determined by varying the concentrations of these substrates in the assay (from 0 to 50 mM for NH2OH and from 0 to 10 mM for methyl viologen). Determination of the optimal pH for hydroxylamine reductase activity was performed by using either a mixed buffer (MES, MOPS, HEPES, Tris-HCl and CHES, 10 mM each) or a 100 mM CHES buffer at various pH values. The presented data are representative of at least three independent experiments with standard deviations lower than 15%.

DNA Methods—Routine DNA manipulations (DNA isolation, restriction enzyme analysis, agarose gel electrophoresis, cloning procedures, and PCR amplifications) were performed using standard methods (28). Conserved motifs in the bacterial nitrate reductases were used to design nas-specific primers taking into account the preferential codon usage of R. capsulatus. Thus, the amino acid sequences of the assimilatory nitrate reductases of Klebsiella oxytoca (PID 4755082), Bacillus subtilis (PID 2828506), Shewanella frigidimarina (PID 2275099), Oscillatoria chalybea (PID 899356), Synechocystis sp. (PID 1652567), and Synechococcus sp. (PID 397157) were aligned and the conserved motifs TGQPNAMGGRE, GTMTNSERRVTL, and AFRPPPGEARADW (positions 343–353, 466–477, and 480–492 of the Synechocystis sp. protein, respectively) were selected for the synthesis of one specific forward primer (NAS2) and two reverse primers (NAS3a and NAS3b). The amplification reactions were performed as described previously (38), using the NAS2 primer 5'-ACSGGSCAGCCSAACGCSATGGGSGSCCGSGA-3' and either the NAS3a primer 5'-GAYSCGSCGYTCCGAGTTSGTCAYSGTGCC-3' or the NAS3b primer 5'-CCAGKCSGCSCGSGCYTCGGCSGGSGGCGG-3'. For the construction of a genomic library of R. capsulatus E1F1, the Lambda FIX® II/XhoI Partial Fill-In Vector Kit (Stratagene) was used following the instructions of the suppliers, and the total genomic DNA from strain E1F1 was partially digested with Sau3AI. The optimal conditions for obtaining the appropriate insert sizes, from 9 to 23 kb, were 0.06 unit of the enzyme, 20 µg of DNA and 15 min incubation. This Sau3AI-digested DNA was partially filled in with dGTP and dATP using the Klenow polymerase, and then ligated into the vector. DNA was packed using the Gigapack® II Packing Extract Kit (Stratagene) under the conditions recommended by the suppliers. Southern blots and hybridization experiments were carried out with the non-radioactive digoxigenin kit from Roche Applied Science. DNA sequencing was performed automatically using an ABI PRISM 373 sequencer and sequence analysis was carried out with the Genetics Computer Group software package programs and data base searches using the BLAST program (ncbi.nlm.nih.gov/BLAST/). The pCap27 plasmid with the 2.8-kb SalI DNA fragment carrying the hcp and nsrR genes (see Fig. 1), and the different mobilizable suicide vectors (pSUP) indicated in Table I were used to construct an insertion mutant in the hcp gene of R. capsulatus, and the resulting recombinant plasmids were mobilized from E. coli S17-1 into R. capsulatus E1F1S by filter matings (27).



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  		FIG. 1.
Physical restriction map and genetic organization of the nas gene region of R. capsulatus E1F1. The physical restriction map of the 17,155-bp nas region of R. capsulatus E1F1 is given for the enzymes ClaI (C), EcoRI (E), EcoRV (V), HindIII (H), KpnI (K), PstI (P), SalI (S), and XhoI (X). The four SalI fragments of about 2.8, 4.1, 4.5, and 5.8 kb isolated from a positive cosmid identified in a genomic library of R. capsulatus E1F1 by hybridization with a 450-bp fragment of the nasA gene as a probe are also indicated. For details, see the text. The location, sizes, and orientation of the genes are symbolized by open arrows. The 2.8-kb SalI fragment described in this work is marked by a black bar.

 
Construction, Expression, and Purification of a His6-tagged HCP Protein—A 1.6-kb DNA fragment with the whole hcp gene was amplified by PCR using the primers HcpF (5'-AGCCTTCCGGATCCATTGCATCCAATG-3', BamHI restriction site underlined) and HcpR (5'-CACAAAGCCACTCCACCTGTCTCG-3'). This fragment was first cloned into pGEM-T and then subcloned, as a SalI-BamHI fragment, into the pQE32 expression vector to give the final construct pHCP1, in which the polyhistidine coding sequence of the vector was fused inframe to the 5' end of the R. capsulatus hcp gene. This plasmid pHCP1 was sequenced to confirm that the fusion was made in the correct reading frame and to check the absence of possible mutations generated by PCR. The E. coli JM109 strain carrying the pHCP1 plasmid was cultured aerobically at 37 °C on a shaker at 250 rpm in conical flasks with LB medium supplemented with ampicillin (100 µg ml–1). When the cultures reached an absorbance at 600 nm of about 0.6, 1 mM isopropyl 1-thio-{beta}-D-galactopyranoside was added to induce HCP expression and the cultures were grown for a further 3 h. After disrupting the cells, 1 mM of the protease inhibitor 4-(2-aminoethyl)benzenesulfonyl fluoride, 50 mM thiocyanate, 0.2 mM NaCl, 10 mM imidazole, and 20% glycerol were also added, and these soluble extracts containing the recombinant HCP protein were loaded onto a nickel-nitrilotriacetic acid-agarose (Qiagen) column. Proteins were eluted with an imidazole gradient (10 to 500 mM), and fractions were kept under argon atmosphere until use.

Western Blots with Anti-His6 Antibodies—For electrophoretic separation, samples were loaded onto polyacrylamide gels, with 14% (w/v) resolving gels and 5% (w/v) stacking gels. These gels were used in Western blot analysis or were stained with Coomassie Brilliant Blue. Western blots to detect the His6-tagged HCP protein were performed by using monoclonal anti-polyhistidine clone his-1 from mouse ascites fluid as primary antibody and anti-mouse IgG alkaline phosphatase conjugate from goat as second antibody.

UV Visible Absortion Spectra—HCP samples were placed in anaerobically sealed serum-stoppered quartz cuvettes and spectra were recorded before and after addition of dithionite, thionine, or hydroxylamine in a DU 7500 (Beckman) spectrophotometer.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 REFERENCES
 
Cloning and Sequence Analysis of the R. capsulatus E1F1 nas Gene Region—To identify and clone the DNA region encoding the assimilatory nitrate reduction system of R. capsulatus E1F1, PCR amplifications were performed using genomic DNA from the strain E1F1 as template and nas-specific primers designed taking into account the Rhodobacter codon usage, as indicated under "Experimental Procedures." DNA fragments of about 400 and 450 bp were amplified using the NAS2/NAS3a and NAS2/NAS3b primer pairs, respectively, and cloned into the pGEM-T vector. These sizes agree with the length of the DNA regions located between the primers in the different genes coding for bacterial assimilatory nitrate reductases. Sequencing of both fragments confirmed that they correspond to the expected internal regions of the R. capsulatus E1F1 nasA gene coding for the catalytic subunit of the nitrate reductase. The 0.45-kb fragment of the nasA gene was also used as a probe for the screening of a genomic library of R. capsulatus E1F1 generated in the lambda Fix® II vector (Stratagene), as described under "Experimental Procedures." Two positive cosmids containing the putative nas gene region of this bacterium were identified. One of them was subjected to restriction analysis and four SalI DNA fragments of 2.8, 4.1, 4.5, and 5.8 kb were present in the corresponding insert (Fig. 1). Cloning and sequencing of these fragments revealed the presence of a 17,155-bp nas gene region that includes putative genes coding for an ABC-type nitrate/nitrite transport system (nasFED), structural genes encoding the nitrate reductase (nasA), and the nitrite reductase subunits (nasB, nirD), regulatory genes (nsrR, nasTS), and genes probably involved in taxis toward nitrate (nnrS) and in the synthesis of the molybdenum-bis-molybdopterin guanine dinucleotide cofactor (moaD) of the nitrate reductase and the siroheme cofactor (cysG) of the nitrite reductase (Fig. 1). The nucleotide sequence data of this 17-kb nas gene region have been deposited in the EMBL, GenBankTM, and DDBI Nucleotide Sequence Databases under the accession number AY273169 [GenBank] .

The 2.8-kb SalI fragment (bases 1 to 2,814) includes 83 nucleotides at the 3' end of the moaD gene involved in the molybdopterin cofactor biosynthesis, a putative hcp gene coding for a hybrid cluster protein, a negative regulatory gene of the rrf2 family (nsrR), and the 5' end of the nasT gene encoding a transcription antitermination protein (Fig. 1). The hcp, nsrR, and nasT genes are preceded by typical Shine-Dalgarno sequences. The predicted hcp gene product is a soluble cytoplasmic protein of 546 amino acid residues with a molecular mass of 57.8 kDa and a calculated pI of 5.7. Sequence comparisons indicated that R. capsulatus HCP is highly similar to the hybrid cluster proteins of several bacteria, including the conserved residues for binding the atypical [4Fe-2S-2O] cluster (Fig. 2A). Similarity to class 2 HCPs of E. coli, Salmonella, and other facultative anaerobic Gram-negative bacteria (50% identity and 63% similarity), all of them including the N-terminal C-X2-C-X11-C-X6-C motif for binding a [2Fe-2S] cluster, was higher than to the class 1 HCPs present in Desulfovibrio and other strictly anaerobic bacteria (43% identity and 55% similarity), which have a different sequence C-X2-C-X7–8-C-X5-C for the coordination of the N-terminal [4Fe-4S] cluster (18, 19). The predicted nsrR gene product is a 140-residue protein with a molecular mass of 15.5 kDa and a calculated pI of 8.9. This protein is homologous to the Nitrosomonas europaea nsrR gene product (39% identity and 55% similarity, Fig. 2B), which is a nitrite-sensitive negative regulator of the expression of nirK, the gene encoding the copper-nitrite reductase (39). BLASTP searches also showed high similarity to putative negative regulatory proteins of the Rrf2 family present in D. vulgaris (40) and other bacteria, such as E. coli and Salmonella (31–38% identity and 50–55% similarity). All of these putative regulatory proteins include the conserved motif C-X6-C-X5-C in the C-terminal end (Fig. 2B).



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  		FIG. 2.
Comparisons of deduced amino acid sequences of the R. capsulatus hcp and nsrR gene products. A, the predicted amino acid sequence of the R. capsulatus HCP protein (RcHCP) is aligned with the sequences of the HCP class 2 proteins of E. coli (EcHCP, accession number BAA35587 [GenBank] and Salmonella typhimurium (StHCP, accession number AAL19873 [GenBank] , and the HCP class 1 protein of Desulfovibrio desulfuricans (DdHCP, accession number ZP00129319). Identical or similar amino acid residues are indicated in dark gray or light gray, respectively. The N-terminal cysteine motif for binding the iron-sulfur cluster 1 is marked by asterisks above the sequences. B, alignment of the predicted amino acid sequence of the R. capsulatus E1F1 NsrR protein (RcNsrR) and the homologous proteins NsrR from N. europaea (NeNsrR, accession number ZP00002681), and Rrf2 from Desulfovibrio vulgaris (DvRrf2, accession number P33395 [GenBank] ). Identical or similar amino acid residues are shown in dark gray or light gray, respectively.

 
Growth of R. capsulatus E1F1 with Hydroxylamine as a Nitrogen Source and Induction of Hydroxylamine Reductase Activity—The physiological function of the HCP proteins is still unknown, but it has been recently demonstrated that E. coli HCP has hydroxylamine reductase activity (24) and is only expressed during anaerobic growth with nitrate (18). A nitriteinducible HCP has also been described in Salmonella (21). The presence of the hcp gene in the nas gene cluster of R. capsulatus suggests that the HCP of this strain could be involved in assimilation and/or detoxification of NH2OH, which is a toxic compound that may be formed during nitrate/nitrite assimilation. Therefore, we tested if R. capsulatus E1F1 is able to grow in the presence of hydroxylamine. This strain did not grow heterotrophically under dark-aerobic conditions in the presence of hydroxylamine, even at a concentration as low as 0.1 mM (not shown). However, R. capsulatus E1F1 grew well with 1 mM NH2OH as the sole nitrogen source under anaerobic phototrophic conditions, although a long lag-phase (4 days) was observed. Hydroxylamine reductase activity was also detected in the cells (Fig. 3). On the contrary, no phototrophic growth with 1 mM NH2OH was observed after 10 days in R. capsulatus B10S, a strain lacking the hcp gene and the nitrate and nitrite reductase nas genes. The maximal phototrophic growth of the strain R. capsulatus E1F1 was observed after hydroxylamine depletion (not shown), suggesting that ammonium is transitorily accumulated before its assimilation. To test this possibility, cells growing with 1 mM NH2OH were harvested and resuspended in fresh media with 5 mM NH2OH under argon atmosphere, with or without 2 mM L-methionine-D,L-sulfoximine (MSX), a glutamine synthetase inhibitor. A rapid hydroxylamine uptake, with similar rates in both MSX-treated and control cells, was observed and no hydroxylamine was detected in the media after about 6 h (Fig. 4). Resting cells were also able to consume added 10 mM NH2OH after about 8 h (not shown). In the absence of MSX, ammonium was accumulated and assimilated after NH2OH consumption. Slightly higher concentrations of ammonium were accumulated in the MSX-treated cells, but only a low assimilation, probably due to a residual glutamine synthetase activity, was observed after NH2OH depletion. Ammonium assimilation in the absence of MSX was also confirmed by the increase in the protein content of the cells (Fig. 4). On the other hand, hydroxylamine reductase activity was significantly higher in the cells growing with hydroxylamine or nitrite than in ammonium- or glutamate-grown cells, and relatively high activities were also observed in cells grown with nitrate or with both NH2OH and NH+4 (Table II). Also, the NH2OH uptake rates by cells grown in the different nitrogen sources correlated well with the hydroxylamine reductase activities (not shown). We also tested if NADH may act as electron donor for the NH2OH reduction in the cell-free extracts from NH2OH-grown cells, and a slightly lower value for the hydroxylamine reductase activity (19 nmol of NADH oxidized min–1 mg–1) was found compared with the activity assayed by measuring ammonia formation with methyl viologen as artificial electron donor (24.5 nmol of NH3 formed min–1mg–1).



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  		FIG. 3.
Phototrophic growth of R. capsulatus E1F1 with hydroxylamine as nitrogen source and time course of hydroxylamine reductase activity. Cells were grown with 1 mM NH2OH as the sole nitrogen source under light-anaerobiosis conditions. At the indicated times, the bacterial growth was followed by the increase of absorbance at 680 nm (), and the hydroxylamine reductase activity ({blacksquare}) was assayed by measuring ammonium formation with methyl viologen as electron donor, as indicated under "Experimental Procedures."

 



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  		FIG. 4.
Hydroxylamine uptake, ammonium accumulation in the media, and protein content of R. capsulatus E1F1 resting cells with and without MSX. Cells were grown with 1 mM NH2OH as the sole nitrogen source under light-anaerobiosis conditions, harvested, and resuspended in fresh media with 5 mM NH2OH and 2 mM MSX (open symbols and dotted lines) or without MSX (filled symbols and solid lines). At the times indicated in the figure, the concentration of hydroxylamine (circles) and ammonium (squares) in the media were determined as described under "Experimental Procedures." The protein content was also estimated both in the 2 mM MSX-treated cells (open triangles) and the resting cells without MSX (filled triangles).

 


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  		TABLE II
Hydroxylamine reductase activity in R. capsulatus E1F1 cells grown with different nitrogen sources

Cells growing phototrophically with the indicated nitrogen sources were harvested at the end of the exponential phase of growth. The hydroxylamine reductase activity was assayed in the cell-free extracts by measuring the ammonium production with methyl viologen as electron donor.

 
Overproduction and Purification of a His6-tagged HCP Protein—To analyze if the R. capsulatus hcp gene product is a hydroxylamine reductase, a recombinant His6-tagged HCP has been overproduced in E. coli and purified by nickel-nitrilotriacetic acid-agarose affinity chromatography. As described under "Experimental Procedures," the whole hcp gene was amplified by PCR, using primers HcpR and HcpF, and cloned into the pQE32 expression vector to obtain the recombinant plasmid pHCP1, in which the polyhistidine coding sequence of the vector is fused in-frame to the 5' end of the R. capsulatus hcp gene. The E. coli JM109 strain was transformed with this pHCP1 plasmid and the synthesis of the recombinant His6-tagged HCP was induced by addition of 1 mM isopropyl 1-thio-{beta}-D-galactopyranoside. Crude extracts from the induced E. coli (pHCP1) cells were subjected to SDS-PAGE, and Coomassie stain revealed overproduction of a protein with an apparent molecular mass of about 53 kDa, a size slightly smaller albeit in the expected range as predicted from the hcp gene sequence (57.8 kDa). Western blot analysis using anti-polyhistidine antibodies confirmed that this 53-kDa protein corresponds to the His6-tagged recombinant HCP (Fig. 5). To purify the recombinant His6-HCP protein, the cell-free extract was loaded onto a nickel-nitrilotriacetic acid-agarose column and proteins were eluted with an imidazole gradient (10 to 500 mM), as indicated under "Experimental Procedures." Most of the recombinant HCP eluted with a high purity grade at about 200 mM imidazole, although some minor contaminants were still present (Fig. 5). A yield of about 0.83 mg of protein per liter of culture was obtained.



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  		FIG. 5.
Purification of the recombinant His6-tagged HCP protein. The recombinant His6-HCP protein was purified by nickel-nitrilotriacetic acid-agarose affinity chromatography, eluted by an imidazole gradient (10–500 mM), and loaded onto a 15% SDS-PAGE for both Coomassie stain (panel A) and Western blots with anti-histidine polyclonal antibodies (panel B), as described under "Experimental Procedures." Protein molecular mass standards: myosin (200 kDa), {beta}-galactosidase (116.25 kDa), phosphorylase b (97 kDa), serum albumin (66.2 kDa), ovalbumin (45 kDa), and carbonic anhydrase (21.5 kDa).

 
Labile Sulfide and Iron Analysis and Spectroscopic Characterization of the His6-HCP—Labile sulfide and iron were analyzed in the purified recombinant protein as described under "Experimental Procedures." The protein contained 4.6 labile sulfide atoms and 5.6 iron atoms per molecule. These values are in agreement with the sequence data suggesting the presence of two iron-sulfur centers, the N-terminal [2Fe-2S] cluster 1 and the hybrid [4Fe-2S-2O] cluster 2. An identical value of 5.6 mol of iron per mol of HCP was reported for the E. coli protein (18), in contrast to the content of 8 mol of iron per mol of HCP as measured by Pereira et al. (19).

The optical absorption spectrum of the recombinant HCP (Fig. 6) was very similar to the spectra of Desulfovibrio and E. coli HCPs (8, 18, 19, 24). The spectrum displayed an aromatic peak at 280 nm, a shoulder at 325 nm, and a peak at 420 nm typical of iron-sulfur proteins (Fig. 6). Values for the molar extinction coefficient at 420 nm within the range from 14 to 28 mM–1 cm–1 were obtained in different samples, which are in agreement with the described values (15–34 mM–1 cm–1) for other purified HCPs (18, 19). The signal at 420 nm was diminished by reduction of the protein with dithionite, but upon addition of hydroxylamine the peak at 420 nm was observed, suggesting the oxidation of the iron-sulfur centers. Similar values for the maximal absorbance at 420 nm in hydroxylamine- and thionine-oxidized samples were obtained, also indicating that the iron-sulfur centers of HCP are fully oxidized upon addition of hydroxylamine (Fig. 6).



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  		FIG. 6.
Optical absorption spectra of the purified His6-HCP protein. The UV-visible spectrum of the purified recombinant protein is shown. Protein content was 0.188 mg ml–1. In the upper box, the dithionite-reduced (line B), hydroxylamine-oxidized (line A), and thionine-oxidized (line C) spectra are shown for the 350–500 nm region. The increase of the absorbance at about 500 nm in the sample treated with thionine is due to the presence of this compound, which has a peak in the 500–600 nm region.

 
Characterization of the Hydroxylamine Reductase Activity of the His6-HCP Protein—The purified recombinant HCP protein showed hydroxylamine reductase activity, forming ammonia in vitro with reduced methyl viologen as electron donor. The specific activity determined was about 3 µmol of NH3 formed min–1 mg–1, which is almost identical to the reported activity for the D. desulfuricans HCP (17), but smaller than the activity described for the E. coli protein (24). Activity showed an optimum pH of 9.3 and an optimum temperature of 40 °C. The HCP protein exhibited hyperbolic saturation kinetics with apparent Km values for hydroxylamine and methyl viologen of 1 mM and 7 µM, respectively, at the temperature and pH optima. The activity was oxygen labile and inhibition by excess reduced methyl viologen was also observed. The hydroxylamine reductase activity was also assayed with different electron donors (Table III). As expected, no activity was found with NADH or NADPH because the purified recombinant His6-HCP lacks the NADH oxidoreductase (HCR), which catalyzes the electron transfer from NADH to the HCP protein (18). However, a low activity was measured with reduced FAD and FMN, suggesting that a flavin-containing diaphorase could be involved in the physiological electron transfer from NADH to the Rhodobacter HCP protein. A low activity was also found with benzyl viologen and bromphenol blue as reductants (Table III). The ability to use reduced bromphenol blue as an electron donor has been described as a characteristic of the bacterial assimilatory nitrate reductases (7).


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  		TABLE III
Hydroxylamine reductase of the purified His6-tagged HCP protein with different electron donors

The hydroxylamine reductase activity of the purified His6-Hcp was assayed by measuring the ammonium production using methyl viologen (MV), benzyl viologen (BV), bromophenol blue (BPB), FAD, FMN, NADH, or NADPH as electron donors (10 mM each). All these compounds, except NADH or NADPH, were reduced by addition of sodium dithionite to the assay mixture.

 
Purified His6-tagged HCP was not affected, or even slightly stimulated, by azide, thiocyanate, and cyanate, but was inhibited by cyanide (Table IV). Curiously, the hydroxylamine reductase activity of the E. coli HCP is greatly stimulated when high cyanide concentrations were present in the assays, although an incubation for 1 h with cyanide irreversibly inactivates the protein (24). The activity of the Rhodobacter HCP was also inhibited by sulfide and iron reagents, such as dithioerythritol, 2,2'-dipyridyl, and o-phenanthroline. The strongest inhibition was caused by this last compound (Table IV).


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  		TABLE IV
Effect of different potential inhibitors on the hydroxylamine reductase activity of the purified His6-tagged HCP protein

The hydroxylamine reductase activity of the purified His6-HCP was assayed by measuring the ammonium production using methyl viologen as electron donor in the presence of the indicated compounds.

 
The HCP Protein Is Responsible for the in Vivo Hydroxylamine Reduction—We tried to obtain an hcp insertion mutant of R. capsulatus E1F1 using different mobilizable suicide vectors, but as it was previously described (27), this E1F1 strain showed extremely low mating frequencies due to DNA restriction phenomena and no mutant strains were obtained. However, as mentioned above, the strain R. capsulatus B10S, which lacks the hcp gene, was unable to grow with hydroxylamine as the sole nitrogen source. This result suggests that the HCP protein is responsible for the capability to assimilate hydroxylamine in the strain R. capsulatus E1F1. In addition, the hydroxylamine reductase activity detected in the crude extracts from R. capsulatus E1F1 cells grown phototrophically with NH2OH as nitrogen source was also inhibited by oxygen and by sulfur and iron reagents, and it showed the same temperature and pH optima as the purified His6-tagged HCP protein, also indicating that the hcp gene product could be responsible of the growth of this strain in media with hydroxylamine. To establish a clear link between the observed hydroxylamine reductase activity of the purified HCP protein and the ability to grow in the presence of NH2OH, we tested if the E. coli cells that are overproducing the R. capsulatus HCP protein grow better with hydroxylamine under anaerobic conditions than the control cells that are not overproducing the recombinant protein. As the hydroxylamine reductase activity is oxygen-sensitive, and the overproduced HCP requires a physiological electron donor to be active, both E. coli cells transformed with the plasmid pHCP1, and the control cells transformed with the pQE32 vector, were grown anaerobically in LB media with 0.4% glycerol and 10 mM KNO3 to induce the expression of the endogenous hcr gene coding for the NADH oxidoreductase that mediates the electron transfer from NADH to the HCP protein. To induce overproduction of the His6-HCP protein, 250 µM isopropyl 1-thio-{beta}-D-galactopyranoside was also added, and different hydroxylamine concentrations (0 to 5 mM) were used. As shown in Fig. 7, bacterial growth in the absence of hydroxylamine was lower in the cells overproducing the HCP protein, as expected because the cell metabolism is focused to synthesis of the recombinant protein. However, the cells overproducing the HCP protein were able to grow better in the presence of 0.5 or 1 mM hydroxylamine than the control cells. Thus, 0.5 mM NH2OH caused no inhibition, and even a slight stimulation, in the bacterial growth of the cells overproducing the HCP, and only a 13% growth inhibition was observed after 9 h with 1 m M NH2OH, whereas significant growth inhibition (25% with 0.5 mM NH2OH and 63% with 1 mM NH2OH after 9 h) was observed in the control cells. No growth was found in the presence of 5 mM NH2OH in both strains.



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  		FIG. 7.
Anaerobic growth of E. coli (pQE32) and E. coli (pHCP1) in the presence of hydroxylamine. Overnight cultures of the E. coli cells transformed with the vector pQE32 (control) or with the plasmid pHCP1 were diluted with fresh LB medium. When the cultures reached an absorbance at 600 nm of about 0.5, the cells were used to inoculate LB media with 0.4% glycerol, 10 mM KNO3, and different hydroxylamine concentrations (0 to 5 mM). Anaerobic conditions were achieved by filling completely 10-ml screw-capped tubes with culture medium, and 250 µM isopropyl 1-thio-{beta}-D-galactopyranoside was added to induce expression of the hcp gene. The bacterial growth was followed by measuring the absorbance of the cultures at 600 nm. Control without hydroxylamine ({circ}), 0.5 mM NH2OH (), 1 mM NH2OH ({blacksquare}), 5 mM NH2OH ({blacktriangleup}).

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
REFERENCES
 
In this work we describe the identification and cloning of a 17-kb DNA region of the R. capsulatus E1F1 genome encoding the nitrate assimilation system of this bacterium. The nas gene cluster includes the structural genes coding for the subunits of the enzymes nitrate and nitrite reductases and the ABC-type nitrate/nitrite transporter, regulatory genes, and genes involved in the biosynthesis of the molybdopterin guanine dinucleotide and the siroheme cofactors present in the nitrate reductase and the nitrite reductase, respectively (Fig. 1). Interestingly, a gene (hcp) encoding a putative iron-sulfur hybrid cluster protein (formerly known as prismane) is also present in the R. capsulatus nas gene region, suggesting an involvement of this HCP protein in the nitrate assimilation process. The results of this study confirm those published previously describing that the HCP protein has hydroxylamine reductase activity (24), and provide evidence, for the first time, that HCP may participate in assimilation and/or detoxification of hydroxylamine, a toxic product that could be formed during nitrate assimilation, probably in the nitrite reduction step. Nitric oxide and hydroxylamine are intermediates in the 6-electron reduction of nitrite to ammonia by the cytochrome c nitrite reductase, an enzyme that also reduces added hydroxylamine, although at a lower rate than nitrite (41–45). NO and NH2OH are also intermediates of the assimilatory siroheme-containing nitrite reductases, as recently demonstrated for the ferredoxin-dependent enzyme from spinach (46). Therefore, hydroxylamine may be potentially produced in the nitrite reduction step, although at present there is no experimental evidence for releasing hydroxylamine during in vivo nitrite reduction. If this occurs, it would result in a selective advantage to possess a protein with hydroxylamine reductase activity, as the HCP, to metabolize this toxic compound. It is worth noting that R. capsulatus E1F1 efficiently assimilates nitrate or nitrite under phototrophic (anaerobiosis-light) conditions, but only a very low heterotrophic growth is observed with these compounds, despite that both nitrate and nitrite reductase activities are detectable under aerobiosis (5). Similarly, R. capsulatus E1F1 does not grow heterotrophically in the presence of hydroxylamine, but it is able to grow with 1 mM NH2OH as the sole nitrogen source under anaerobic phototrophic conditions (Fig. 3). However, ammonium generated by NH2OH reduction is not significantly assimilated until NH2OH is almost completely depleted (Fig. 4). Therefore, the hydroxylamine reductase activity could be a detoxification mechanism that generates a product, ammonium, which can be further assimilated. The inhibition of cytochrome oxidase and some Krebs cycle enzymes by nitrite has been invoked to explain the absence of aerobic growth of R. capsulatus E1F1 in media with nitrate or nitrite (6), but the inability to detoxify hydroxylamine due to the anaerobic production of the HCP and the oxygen lability of its hydroxylamine reductase activity could also explain the absence of heterotrophic growth in media with nitrite or hydroxylamine in this bacterium.

HCPs contain two types of iron-sulfur centers, one typical [4Fe-4S] or [2Fe-2S] center in the N-terminal domain (cluster 1) and one unusual hybrid [4Fe-2S-2O] center (cluster 2) at the interface of the three domains of the protein (14–18). Despite the spectroscopic characteristics, the crystal structure of HCPs have been well studied, the possible enzymatic activity and the physiological function of the HCP are still controversial. It has been suggested that HCP has a role in nitrate and/or nitrite respiration (18), and the hydroxylamine reductase activity of the E. coli HCP indicates that this protein may act as a scavenger of potentially toxic NH2OH generated in nitrate metabolism (24). However, this physiological role has been questioned due to the high Km value for NH2OH of HCP and the basic optimum pH for activity (17). Concerning this point, it should be noted that R. capsulatus HCP also exhibits a optimum pH of 9.3 and a relatively high Km value for NH2OH (1 mM), but other purified enzymes involved in the nitrogen metabolism in this bacterium also show high apparent Km for their substrates and basic optima pH in vitro. Thus, the assimilatory nitrate reductase has a optimum pH of 9.5 and an apparent Km for nitrate of 13 mM with reduced bromphenol blue as reductant (7), and arginase shows an optimal pH of 9.0 and an apparent Km value for arginine of 16 mM (47).

Genes coding for HCP proteins have been identified in a large number of both strict anaerobic and facultative anaerobic prokaryotes, but clustering of hcp and nas genes has not been described before. In several facultative bacteria, a gene (hcr) coding for an NADH oxidoreductase, which seems to mediate the electron transfer from NADH to the HCP, is located immediately downstream of the hcp gene (18, 20, 21). However, in the genomes of obligate anaerobes and thermophilic Archaea, hcr genes are not present in the immediate surroundings of the hcp genes (18). This is also the case of R. capsulatus, because no hcr gene was located in the nas gene cluster and the hcp gene is oriented in the opposite direction to the adjacent genes (Fig. 1). However, the NADH-dependent hydroxylamine reductase activity detected in the Rhodobacter cell-free extracts and the fact that the purified HCP protein exhibits a low activity with reduced flavins (Table II), indicates that a flavin-containing NADH oxidoreductase mediates the in vivo electron transfer from NADH to HCP. It is worth noting that the R. capsulatus assimilatory nitrate reductase contains a 46-kDa diaphorase subunit (7), but the gene coding for this NADH-dependent flavoprotein is not present in the cloned 17-kb nas region. Therefore, a hcr-like gene should be located elsewhere in the genome of this bacterium.

The inhibition of the hydroxylamine reductase activity by iron and sulfur reagents (Table III), and the UV-visible absorption spectra of the recombinant HCP protein (Fig. 6) indicate the presence of iron-sulfur centers in the protein and support evidence for the involvement of these centers in the NH2OH reduction. The N-terminal Cys motif of the R. capsulatus HCP has the same spacing features as in the E. coli protein and other class 2 HCPs (Fig. 2A). Curiously, the R. capsulatus HCP is more similar to these class 2 proteins than to the putative class 1 HCP from another phototrophic bacteria, Rhodospirillum rubrum. This suggests the presence of a [2Fe-2S] cluster instead of the cubane [4Fe-4S] center present in the class 1 HCPs from the Desulfovibrio species and other bacteria (18, 19). Labile sulfide and iron determinations in the purified His6-tagged HCP are also in agreement with the presence of a [2Fe-2S] cluster in addition to the hybrid [4Fe-2S-2O] cluster.

Very recently, it has been described that the N. europaea nsrR gene negatively regulates the expression of the nirK gene encoding the copper-nitrite reductase in response to nitrite and pH (39). This gene is homologous to the D. vulgaris rrf2 gene encoding a protein with a putative function in negative regulation of gene expression (40). The presence of a nsrR homologous gene adjacent to the hcp gene in R. capsulatus (Fig. 1) suggests that hcp gene expression, and probably also nas gene expression, could be regulated in response to nitrite by this repressor of the Rrf2 family. This is in agreement with the higher hydroxylamine reductase activities and hydroxylamine uptake rates detected in the cells growing with nitrite or hydroxylamine (Table II). Also, in E. coli and Shewanella, HCP is only expressed during anaerobic growth with nitrate or nitrite (18, 22), and recently, two nipAB (nitrite-inducible promoter) genes, which corresponds to hcp-hcr genes, have been identified with a green fluorescent protein-based promoter-trap screen in S. enterica (21). The possible regulatory proteins involved in hcp gene expression have not yet been described in these organisms, but interestingly, all these bacteria contain rrf2/nsrR homologous genes, and all these putative Rrf2/NsrR proteins include a conserved C-X6-C-X5-C motif in the C-terminal end (positions 91–104 of the R. capsulatus sequence, Fig. 2B). It could be speculated that this Cys motif is involved in the binding of an iron-sulfur center that is required for the repressor activity of the protein and that, in the presence of nitrite or hydroxylamine, the iron-sulfur center is degraded resulting in the inactivation of the protein and, therefore, the expression of the genes under its control. The existence of iron-sulfur regulatory proteins controlling gene expression is well documented in the literature (48). Putative –35 and –10 boxes for a {sigma}70-like promoter are present upstream from the hcp gene. Interestingly, a perfect palindromic sequence TGGTATT-N3-AATACCA is found between the putative TATA box and the Shine-Dalgarno sequence of the hcp gene (48 bases upstream the start codon of HCP), and a second highly similar TGGAGTT-N3-GATACCA sequence is also present 85 bases upstream from the HCP start codon. However, if these sequences are putative binding sites for the NsrR repressor or have another role in the control of hcp gene expression is presently unknown because studies on transcriptional regulation of the hcp gene has not yet been carried out.

Several results of this work suggest that the R. capsulatus E1F1 hcp gene product is involved in detoxification and further assimilation of hydroxylamine. Thus, the clustering of hcp and nas genes, the oxygen-sensitive hydroxylamine reductase activity of the HCP protein, which is in agreement with the fact that R. capsulatus E1F1 assimilates NH2OH only under anaerobic phototrophic conditions, the higher hydroxylamine reductase activities found in nitrite- or hydroxylamine-grown cells, in accordance with the possible control of hcp gene expression by the putative nitrite-sensitive repressor NsrR, and the identical biochemical parameters of the hydroxylamine reductase activity of cell extracts and the purified His6-HCP protein, as expected if the hcp gene product is the physiological hydroxylamine reductase, suggest that HCP is involved in hydroxylamine reduction in R. capsulatus E1F1. However, these results do not establish a direct linkage between hydroxylamine assimilation in this bacterium and the hydroxylamine reductase activity of the HCP protein. Although it was not possible to obtain a R. capsulatus E1F1 hcp insertion mutant due to DNA restriction (27), other results presented in this work suggest that HCP is the protein responsible for hydroxylamine reduction both in R. capsulatus and E. coli. First, the strain R. capsulatus B10S was unable to grow phototrophically with 1 mM hydroxylamine as the sole nitrogen source, in contrast to the E1F1 strain. R. capsulatus B10S is also unable to assimilate nitrate (49) because it lacks the whole hcp-nas gene region, as revealed by the genome sequence data (50) and Southern blot experiments using hcp-nas probes (not shown). Second, the anaerobic growth of E. coli in LB media with glycerol and nitrate was significantly inhibited by 0.5–1 mM NH2OH. On the contrary, these hydroxylamine concentrations did not inhibit significantly the growth of the E. coli cells that are overproducing the R. capsulatus HCP protein, despite the fact that bacterial growth in the absence of hydroxylamine was lower than in control cells due to the high cost of the overproduction of the recombinant protein (Fig. 7). The increased resistance to hydroxylamine when HCP is overproduced in E. coli (pHCP1), in comparison with the isogenic E. coli strain transformed with the pQE32 plasmid without the hcp gene, is a demonstration of the potential physiological role of HCP in NH2OH reduction in vivo.

In conclusion, we describe for the first time that a bacterium, R. capsulatus E1F1, is able to assimilate hydroxylamine under phototrophic anaerobic conditions and contains a nitrate assimilation nas gene cluster that includes the hcp gene encoding a hybrid cluster protein. Expression of the hcp gene could be regulated by a putative nitrite-sensitive repressor encoded by the adjacent nsrR gene, thus explaining the increase of the hydroxylamine reductase activity in the R. capsulatus cells grown with nitrite or hydroxylamine. We also present some results indicating that HCP may play a role in assimilation and/or detoxification of hydroxylamine, a toxic compound that can be present in some environments or could be potentially formed during bacterial nitrate and nitrite assimilation.


   FOOTNOTES
 
* This work was supported by Ministerio de Ciencia y Tecnología Grant BMC2002-04126-CO3-03 and Junta de Andalucía Grant CVI 0117, Spain. The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

§ These authors contributed equally to this work. Back

|| Recipient of a fellowship from Ministerio de Ciencia y Tecnología. Back

** Supported by a postdoctoral contract from Junta de Andalucía, Spain. Back

{ddagger}{ddagger} To whom correspondence should be addressed. Tel. and Fax: 34-9-57-218588; E-mail: bb1movic{at}uco.es.

1 The abbreviations used are: HCP, hybrid cluster protein; HCR, hybrid cluster protein NADH oxidoreductase; LB, Luria-Bertani broth; MSX, L-methionine-D,L-sulfoximine; CHES, 2-(cyclohexylamino)ethanesulfonic acid; MES, 4-morpholineethanesulfonic acid; MOPS, 4-morpholinepropanesulfonic acid. Back


   ACKNOWLEDGMENTS
 
We thank Dr. David J. Richardson (University of East Anglia, Norwich, UK) for helpful discussions and hospitality.



   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Moreno-Vivián, C., Cabello, P., Martínez-Luque, M., Blasco, R., and Castillo, F. (1999) J. Bacteriol. 181, 6573–6584[Free Full Text]
  2. Richardson, D. J., Berks, B. C., Russell, D. A., Spiro, S., and Taylor, C. J. (2001) Cell. Mol. Life Sci. 58, 165–178[CrossRef][Medline] [Order article via Infotrieve]
  3. Lin, J. T., and Stewart, V. (1998) Adv. Microb. Physiol. 38, 1–30
  4. Guerrero, M. G., Vega, J. M., and Losada, M. (1981) Annu. Rev. Plant Physiol. 32, 169–204
  5. Moreno-Vivian, C., Castillo, F., and Cárdenas, J. (1982) Photosynth. Res. 3, 313–319
  6. Martínez-Luque, M., and Castillo, F. (1991) Arch. Microbiol. 155, 149–152[CrossRef]
  7. Blasco, R., Castillo, F., and Martínez-Luque, M. (1997) FEBS Lett. 414, 45–49[CrossRef][Medline] [Order article via Infotrieve]
  8. Moura, I., Tavares, P., Moura, J. J., Ravi, N., Huynh, B. H., Liu, M. Y., and LeGall, J. (1992) J. Biol. Chem. 267, 4489–4496[Abstract/Free Full Text]
  9. Pierik, A. J., Wolbert, R. B., Mutsaers, P. H., Hagen, W. R., and Veeger, C. (1992) Eur. J. Biochem. 206, 697–704[Medline] [Order article via Infotrieve]
  10. Pierik, A. J., Hagen, W. R., Dunham, W. R., and Sands, R. H. (1992) Eur. J. Biochem. 206, 705–719[Medline] [Order article via Infotrieve]
  11. Stokkermans, J. P., Pierik, A. J., Wolbert, R. B., Hagen, W. R., van Dongen, W. M., and Veeger, C. (1992) Eur. J. Biochem. 208, 435–442[Medline] [Order article via Infotrieve]
  12. Van den Berg, W. A., Stevens, A. A., Verhagen, M. F., van Dongen, W. M., and Hagen, W. R. (1994) Biochim. Biophys. Acta 1206, 240–246[CrossRef][Medline] [Order article via Infotrieve]
  13. Kröckel, M., Trautwein, A. X., Arendsen, A. F., and Hagen, W. R. (1998) Eur. J. Biochem. 251, 454–461[Medline] [Order article via Infotrieve]
  14. Arendsen, A. F., Hadden, J., Card, G., McAlpine, A. S., Bailey, S., Zaitsev, V., Duke, E. H., Lindley, P., Kröckel, M., Trautwein, A. X., Feiters, M. C., Charnock, J. M., Garner, C. D., Marrit, S. J., Thomson, A. J., Kooter, I. M., Johnson, M. K., van den Berg, W. A., van Dongen, W. M., and Hagen, W. R. (1998) J. Biol. Inorg. Chem. 3, 81–95
  15. Cooper, S. J., Garner, C. D., Hagen, W. R., Lindley, P. F., and Bailey, S. (2000) Biochemistry 39, 15044–15054[CrossRef][Medline] [Order article via Infotrieve]
  16. Macedo, S., Mitchell, E. P., Romão, C. V., Cooper, S. J., Coelho, R., Liu, M. Y., Xavier, A. V., LeGall, J., Bailey, S., Garner, C. D., Hagen, W. R., Texeira, M., Carrondo, M. A., and Lindley, P. (2002) J. Biol. Inorg. Chem. 7, 514–525[CrossRef][Medline] [Order article via Infotrieve]
  17. Aragão, D., Macedo, S., Mitchell, E. P., Romão, C. V., Liu, M. Y., Frazão, C., Saraiva, L. M., Xavier, A. V., LeGall, J., van Dongen, W. M., Hagen, W. R., Texeira, M., Carrondo, M. A., and Lindley, P. (2003) J. Biol. Inorg. Chem. 8, 540–548[Medline] [Order article via Infotrieve]
  18. Van den Berg, W. A., Hagen, W. R., and van Dongen, W. M. (2000) Eur. J. Biochem. 267, 666–676[Medline] [Order article via Infotrieve]
  19. Pereira, A. S., Tavares, P., Krebs, C., Huynh, B. H., Rusnak, F., Moura, I., and Moura, J. J. (1999) Biochem. Biophys. Res. Commun. 260, 209–215[CrossRef][Medline] [Order article via Infotrieve]
  20. Dominy, C., Deane, S. M., and Rawlings, D. E. (1997) Microbiology 143, 3123–3136[Abstract]
  21. Kim, C. C., Monack, D., and Falkow, S. (2003) Infect. Immunol. 71, 3196–3205[Abstract/Free Full Text]
  22. Beliaev, A. S., Thompson, D. K., Khare, T., Lim, H., Brandt, C. C., Li, G., Murray, A. E., Heidelberg, J. F., Giometti, C. S., Yates, J., III, Nealson, K. H., Tiedje, J. M., and Zhoui, J. (2002) OMICS J. Integrative Biol. 6, 39–60
  23. Briolat, V., and Reysset, G. (2002) J. Bacteriol. 184, 2333–2343[Abstract/Free Full Text]
  24. Wolfe, M. T., Heo, J., Garavelli, J. S., and Ludden, P. W. (2002) J. Bacteriol. 184, 5898–5902[Abstract/Free Full Text]
  25. Heo, J., Wolfe, M. T., Staples, C. R., and Ludden, P. W. (2002) J. Bacteriol. 184, 5894–5897[Abstract/Free Full Text]
  26. Weaver, P. F., Wall, J. D., and Gest, H. (1975) Arch. Microbiol. 105, 207–216[CrossRef][Medline] [Order article via Infotrieve]
  27. Moreno-Vivián, C., Roldán, M. D., Reyes, F., and Castillo, F. (1994) FEMS Microbiol. Lett. 115, 279–284[CrossRef]
  28. Sambrook, J., Fritsch, E. F., and Maniatis, T. (1989) Molecular Cloning: A Laboratory Manual, 2nd Ed., Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY
  29. Simon, R., Priefer, U., and Pühler, A. (1983) Biotechnology 1, 784–791[CrossRef]
  30. Arnold, W., and Pühler, A. (1988) Gene (Amst.)70, 171–179[CrossRef][Medline] [Order article via Infotrieve]
  31. Reyes, F., Roldán, M. D., Klipp, W., Castillo, F., and Moreno-Vivián, C. (1996) Mol. Microbiol. 19, 1307–1318[Medline] [Order article via Infotrieve]
  32. Solorzano, L. (1969) Limnol. Oceanogr. 14, 799–801
  33. Korpela, T. K., and Mäkelä, M. J. (1981) Anal. Biochem. 110, 251–257[Medline] [Order article via Infotrieve]