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Journal of Bacteriology, September 2006, p. 6476-6482, Vol. 188, No. 18
0021-9193/06/$08.00+0     doi:10.1128/JB.00737-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

The hmuQ and hmuD Genes from Bradyrhizobium japonicum Encode Heme-Degrading Enzymes

Sumant Puri and Mark R. O'Brian*

Department of Biochemistry and Witebsky Center for Microbial Pathogenesis and Immunology, State University of New York at Buffalo, Buffalo, New York

Received 22 May 2006/ Accepted 5 July 2006


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ABSTRACT
 
Utilization of heme by bacteria as a nutritional iron source involves the transport of exogenous heme, followed by cleavage of the heme macrocycle to release iron. Bradyrhizobium japonicum can use heme as an iron source, but no heme-degrading oxygenase has been described. Here, bioinformatics analyses of the B. japonicum genome identified two paralogous genes renamed hmuQ (bll7075) and hmuD (bll7423) that encode proteins with weak similarity to the heme-degrading monooxygenase IsdG from Staphylococcus aureus. The hmuQ gene is clustered with known heme transport genes in the genome. Recombinant HmuQ bound heme with a Kd value of 0.8 µM and showed spectral properties consistent with a heme oxygenase. In the presence of a reductant, HmuQ catalyzed the degradation of heme and the formation of biliverdin. The hmuQ and hmuD genes complemented a Corynebacterium ulcerans heme oxygenase mutant in trans for utilization of heme as the sole iron source for growth. Furthermore, homologs of hmuQ and hmuD were identified in many bacterial genera, and the recombinant homolog from Brucella melitensis bound heme and catalyzed its degradation. The findings show that hmuQ and hmuD encode heme oxygenases and indicate that the IsdG family of heme-degrading monooxygenases is not restricted to gram-positive pathogenic bacteria.


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INTRODUCTION
 
Iron is essential for most living organisms. It is required for many cellular processes in catalysis, electron transfer, oxygen metabolism, regulation, and signal transduction. Iron can be a limiting nutrient in aerobic environments because the oxidized (ferric, Fe3+) form is extremely insoluble at physiological pH. In addition, iron is made unavailable in animals by binding to specific proteins as a mechanism to limit access by bacterial pathogens. Bacteria have several ways of acquiring iron from their environment to fulfill nutritional needs. One way is by the secretion of siderophores that bind ferric iron with high affinity, followed by transport of the iron chelate into cells (2, 5). Genes encoding proteins involved in ferric siderophore transport or siderophore synthesis are induced under iron limitation. Another mechanism of acquisition is by the direct transport of ferrous (reduced, Fe2+) iron into cells (10). Finally, numerous bacterial pathogens of animals are able to use host iron proteins or heme proteins as iron sources (9, 26).

Heme is an iron-containing tetrapyrrole that serves as the prosthetic group or active moiety of heme proteins. Over 70% of iron in mammals is incorporated into heme, mostly as hemoglobin (18). Utilization of exogenous heme by bacteria requires binding to the cell surface, followed by transport into cells. Gram-negative bacteria have an outer membrane receptor and periplasmic heme binding protein in addition to cytoplasmic proteins that transport heme into cells (9, 26). Studies of heme transport in Corynebacterium diphtheriae and Staphylococcus aureus identified heme-binding cell wall and membrane proteins in gram-positive bacteria (7, 12).

Once exogenous heme is taken up by bacterial cells, the tetrapyrrole ring is cleaved by heme oxygenase in order to release the iron. The pioneering work on bacterial heme oxygenases was carried out with the proteins from Corynebacterium diphtheriae (20, 30) and Neisseriae meningitidis (16, 22, 33, 34), which have limited sequence similarity to each other and to eukaryotic heme oxygenases but have similar structures overall (11, 21, 22). More recently, a structurally unrelated heme-degrading oxygenase was described in Staphylococcus aureus (25, 31), and it is also found in a limited number of gram-positive bacteria, including Bacillus anthracis (24). Both families of heme degradation enzymes use the same substrates and produce biliverdin, iron, and CO.

Although originally though to be a property associated exclusively with bacterial pathogens of animals, utilization of heme and hemoglobin as iron sources was found in nonpathogenic rhizobial bacteria (14). Rhizobia belong to the {alpha}-Proteobacteria, a taxonomic group comprising numerous members that form close or intracellular associations with higher eukaryotes in a symbiotic or pathogenic context. Rhizobia live as free-living soil organisms or in symbiosis with leguminous plants, where they convert atmospheric nitrogen to ammonia to fulfill the nutritional nitrogen requirement of the plant host.

The genes encoding heme transport systems have been described in Bradyrhizobium japonicum (13) and Rhizobium leguminosarum (28), and they share considerable homology with heme transporters from pathogens. The B. japonicum heme transport gene cluster includes hmuR and hmuT, which encode a TonB-dependent heme receptor and periplasmic heme binding protein, respectively, and hmuUV, which encodes ABC transporters of the inner membrane. It also includes tonB, exbB, and exbD, which encode proteins that transduce energy from the inner to the outer membrane (15).

Despite the ability of B. japonicum to use heme as an iron source, no heme oxygenase has been described, nor has a gene homolog been identified in the annotated genome. In the present study, we identify a gene encoding a heme degradation protein and a paralog that likely has the same function. In addition, homologs of the B. japonicum heme degradation genes are present in many bacterial genera.


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MATERIALS AND METHODS
 
Bacterial strains and growth conditions. Bradyrhizobium japonicum strain I110 was cultured in GSY (glycerol-salts-yeast extract) medium as described previously (8). Escherichia coli strain DH5{alpha} was used for propagation of plasmids and was grown on LB medium containing the appropriate antibiotic. Corynebacterium ulcerans parent strain CU712 and hmuO strain CU29 were kindly provided by M. P. Schmitt (20) and were cultured on heart infusion broth medium (Difco, Detroit, MI) supplemented with 0.2% Tween 80 (HIBTW).

Isolation of hmuQ and hmuD and overexpression of hmuQ. The hmuQ (bll7075) and hmuD (bll7423) genes and flanking DNA were isolated by PCR using genomic DNA from B. japonicum strain I110 as template. Primers were designed to amplify the hmuQ open reading frame (ORF) along with 473 bp and 455 bp of upstream and downstream DNA, respectively. The hmuD ORF was amplified along with 724 bp and 379 bp of upstream and downstream DNA, respectively. The products were ligated into the EcoRV site of pBluescript SK(+), and the DNA sequences were verified. The isolated DNA was used as template for amplification of the ORFs, and NdeI and BamHI restriction sites were added to the forward and reverse primers, respectively, for ligation into the expression vector pET14b.

Overnight cultures of E. coli BL21(DE3)(pLysS) strains harboring pET14b-hmuQ were used to inoculate 1 liter of 2x YT medium (16 g/liter tryptone, 10 g/liter yeast extract, 5 g/liter NaCl, pH 7.0) containing chloramphenicol (25 µg/ml) and ampicillin (200 µg/ml). At mid-log phase, cells were induced by addition of isopropyl-1-thio-ß-D-galactopyranoside to a 0.5 mM final concentration and incubated at 20°C for 4 h with shaking. Cells were harvested by centrifugation at 4,000 x g, washed in phosphate binding buffer (5 mM imidazole, 300 mM NaCl, 50 mM NaH2PO4, pH 8.0), and resuspended in 15 ml of phosphate binding buffer, 1 mM phenylmethylsulfonyl fluoride, and 25.5 µg of aprotinin per 5 g of cells. Cells were disrupted by passage twice through a French pressure cell at 1,200 lb/in2 and clarified by centrifugation at 37,000 x g for 45 min. One milliliter of a 50% Ni-nitrilotriacetic acid slurry (QIAGEN Inc., Valencia, CA) was added to 4 ml of clear lysate and rocked for 60 min at 4°C. The Ni-nitrilotriacetic acid slurry-protein mixture was poured into a column and washed four times with 5 ml of phosphate wash buffer (50 mM NaH2PO4, pH 8.0, 300 mM NaCl, 20 mM imidazole) and once with 5 ml of phosphate wash buffer containing 10% glycerol. Purified His-tagged proteins were eluted with phosphate elution buffer (50 mM NaH2PO4, pH 8.0, 300 mM NaCl, 250 mM imidazole, 10% glycerol). Eluted protein fractions were dialyzed against buffer containing 100 mM NaH2PO4, 600 mM NaCl, 20% glycerol, pH 8.0. The purified proteins were stored at –20°C.

Overexpression of the Brucella melitensis bmeII0706 gene. The bmeII0706 gene was synthesized commercially by Bio S&T Inc. (Montreal, Canada). NdeI and BamHI restriction sites were added for ligation into pET14b. The AGG codon encoding Arg-71 and the CCC codon encoding Pro-93 were changed to CGC and CCG, respectively, which did not change the amino acid sequence but incorporated preferred codons in E. coli. The bmeII0706 gene was expressed, and the product was purified as described above for hmuQ.

Heme binding. The binding of heme to HmuQ or BmeII0706 was assessed spectrophotometrically using an SLM-Aminco DW-2000 UV-VIS spectrophotometer. Aliquots of hemin (0.5 to 20 µM) were added to both the sample cuvette containing 10 µM protein in 50 mM NaH2PO4, 300 mM NaCl, 10% glycerol, pH 8.0, and the reference cuvette (buffer alone) at 25°C. Scans were recorded in the 300- to 600-nm region of the spectrum 5 min after the addition of hemin (ferric heme hydrochloride). To determine the protein-to-heme stoichiometry, and the dissociation binding constant (Kd), the absorption at 411 nm was plotted versus the heme concentration. The data were analyzed using the Prism program (GraphPad). Hemin was purchased from Frontier Scientific (Logan, UT) and was greater than 95% pure.

Heme degradation assays. Heme degradation was determined spectrophotometrically as the decrease in the heme absorbance in the presence of HmuQ, or BmeII0706, and a reductant. Purified heme oxygenases do not readily release the product biliverdin in the absence of biliverdin reductase (30). Thus, most studies involve single turnover assays, as was done here. Thus, the reaction mixture contained 10 µM HmuQ or BmeII0706 and an initial hemin concentration of 8 µM. Furthermore, the assays were performed in the presence of purified recombinant catalase from bovine liver (Sigma) at a ratio of catalase to hemoprotein of 0.5:1 to prevent coupled oxidation reactions. Heme degradation was measured in the presence of either ascorbate or human NADPH-cytochrome P450 reductase.

(i) Reaction with ascorbate. Ascorbate-dependent degradation of heme was monitored spectrophotometrically as previously described (25). Recombinant protein (10 µM) and hemin (8 µM) in buffer (50 mM NaH2PO4, 300 mM NaCl, 10% glycerol, pH 8.0) were incubated with ascorbic acid (10 mM), and the spectral changes between 325 and 700 nm were recorded every 5 min. Alternatively, the absorption at 408 nm versus the 600 nm reference was monitored.

(ii) Reaction with NADPH-cytochrome P450 reductase. Heme degradation in the presence of recombinant human NADPH-cytochrome P450 reductase (Calbiochem) was carried out similar to that described above for ascorbate, with the following modifications. Cytochrome P450 reductase was added to a 0.3:1 ratio with HmuQ or BmeII0706. Initiation of the reaction was carried out by the addition of NADPH. NADPH was added in 10 µM aliquots and then scanned from 300 to 700 nm 5 min after each increment.

Detection of biliverdin as a HmuQ reaction product. Heme degradation reactions were carried out in a final volume of 1 ml using ascorbate as reductant as described above. Following the reaction, 200 µl of glacial acetic acid and 200 µl of 3 M HCl were added to quench the cleavage reaction. Subsequently, the reaction mixture was extracted with 1.5 ml of chloroform. The organic layer was washed three times with 1 ml of distilled water, and the chloroform layer was removed under a stream of nitrogen. The resultant residue was dissolved in 100% methanol. The sample was analyzed by liquid chromatography-mass spectrometry using an LCQ Advantage ion trap instrument (Thermo Electron Corp.). A 25-µl aliquot was applied to a Thermo Hypersil C18 Aquasil column (Keystone Scientific Operations), and the products were eluted with a 40 to 90% linear gradient of acetonitrile in 0.1% formic acid, at a flow rate of 200 µl/min. The gradient was varied over a time period of 10 min from 40% to 90%, held constant at 90% for 5 min, and then varied again from 90% to 40% over the next 10 min. The electrospray needle was operated at 4.5 kV with a capillary temperature of 210°C.

In vivo complementation of a heme oxygenase mutant. Complementation of Corynebacterium ulcerans hmuO strain CU29 was carried out as modified from a previously described protocol (20). hmuQ and hmuD were ligated into the BamHI and XbaI sites of pCM2.6 and introduced into the C. ulcerans heme oxygenase mutant CU29. Strain CU29 harboring pCM2.6 or pCD293, which contains the C. diphtheriae hmuO gene in pCM2.6, were negative and positive controls, respectively. Parent strain CU712(pCM2.6) was also used as a positive control. The cells were grown in liquid HIBTW medium containing 3 µg/ml chloramphenicol to maintain the plasmid. The cells were diluted, and then 5 µl was spotted onto HIBTW agar plates containing 200 µg/ml ethylenediamine-N,N'-bis(2-hydroxyphenylacetic acid) (EDDHA), EDDHA plus 7 µM hemin, or EDDHA plus 1 mM FeSO4. Complementation of the mutant was scored as growth with heme as the sole source of iron.


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RESULTS
 
Identification of candidate heme oxygenase-encoding genes. Bacteria that use exogenous heme as a source of iron must be able to transport heme into the cells and cleave the tetrapyrrole ring to release iron. This latter activity is carried out by heme oxygenases. Bradyrhizobium japonicum and some related rhizobia can use heme as an iron source (13, 14), but no heme oxygenase has been described. BLAST homology searches using known heme oxygenases from the bacteria Pseudomonas aeruginosa, Neisseria meningitidis, and Corynebacterium diphtheriae, the plant Arabidopsis thaliana, and from humans did not reveal a candidate enzyme in the annotated B. japonicum genome. Recently, two structurally similar heme-degrading oxygenases, IsdG and IsdI, were identified in the gram-positive bacterium Staphylococcus aureus (25). IsdG and IsdI are structurally distinct from other known heme oxygenases, and homologs have been reported to be limited to Staphylococcus and the related gram-positive genera Listeria and Bacillus (25). A BLAST search of the B. japonicum annotated genome with S. aureus IsdG revealed two weak homologs encoded by bll7075 and bll7423, with E-values of 0.001 and 0.0002, respectively (Fig. 1A). Despite this weak homology, circumstantial evidence suggested that Bll7075 and Bll7423 may be functionally similar to the S. aureus heme oxygenases. Firstly, Pfam analysis that uses hidden Markov models (3) (http://pfam.wustl.edu/) assigns Bll7075 and Bll7423 to the ABM family of monooxygenases, to which S. aureus IsdG and IsdI also belong. Secondly, transcriptional profiling analysis shows that bll7075 mRNA is strongly up regulated in iron-limited cells (19, 32), which is a condition under which the heme utilization system is induced (13). Finally, bll7075 is clustered in the genome with genes known to be involved in heme transport (13) (Fig. 1B). These genes include the hmuTUV gene cluster, hmuR, exbBD, and tonB. A similar genetic region was also characterized in R. leguminosarum (28), which includes a gene designated orfA that has high homology to bll7075. The bll7423 gene is not clustered near known heme metabolism genes. Thus, we further pursued the analysis of these B. japonicum genes. bll7075 and bll7423 were renamed hmuQ and hmuD, respectively, based on data described below that indicate that they are involved in heme metabolism and on the genomic context of hmuQ with hmuR and hmuTUV.


Figure 1
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  		FIG. 1. Similarity of HmuQ (Bll7075) and HmuD (Bll7423) to heme-degrading monooxygenases from S. aureus and the genomic context of the B. japonicum genes. (A) Alignment of B. japonicum HmuQ and HmuD, Brucella melitensis BmeII0706, and S. aureus IsdG and IsdI. The asterisk and colon denote conserved amino acid identity and similarity, respectively. (B) Genomic context of B. japonicum hmuQ and hmuD.

HmuQ binds heme. hmuQ and hmuD encode proteins 110 and 107 amino acids in length, respectively, and share 61% identity and 79% similarity to each other. We focused on hmuQ because of its genomic context and its regulation by iron, and also because of problems with purification of HmuD. HmuQ was expressed as a histidyl-tagged protein in E. coli and purified by affinity chromatography (Fig. 2) (see Materials and Methods). The purified protein migrated on sodium dodecyl sulfate-polyacrylamide gel electrophoresis consistent with its calculated molecular mass of 14,190 Da (Fig. 2A).


Figure 2
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  		FIG. 2. Binding of recombinant HmuQ to heme. (A) The hmuQ gene was overexpressed in E. coli as described in the text. Purification fractions were run on 15% sodium dodecyl sulfate-polyacrylamide gel electrophoresis gels. Std, molecular mass standards; FT, flowthrough; W1, W3, and W5, column washes; E1 and E4, elution fractions. (B) Absorption spectra of HmuQ in the presence of increasing concentrations of hemin. The sample and reference cuvettes were titrated with 0, 0.5, 2, 6, and 10 µM heme as described in the text, and the spectrum was recorded after each addition. Ten µM HmuQ was present in the sample cuvette. The arrow represents increasing levels of added heme. (C) An experiment as described for panel B was carried out with numerous heme concentrations. The absorbance at 411 nm was plotted versus the heme concentration. The data shown in panel C are from a single experiment representative of three independent experiments.

The binding of heme to HmuQ was assessed spectrophotometrically by taking advantage of the fact that the spectral properties of heme change when bound to a protein. The spectrum of the HmuQ-heme complex showed a major peak at 411 nm (Fig. 2B), which is consistent with other heme oxygenases and heme binding proteins in general. A slight peak was observed for the protein in the absence of exogenous heme, which was likely due to heme derived from the E. coli cells in which the recombinant protein was synthesized. Titration of a HmuQ solution (10 µM) with increasing amounts of heme allowed us to quantify the heme binding properties (Fig. 2). The data best fit a one-binding-site curve with a Kd of 0.8 µM (Fig. 2C). The absorption increase leveled off at approximately 10 µM heme, showing a 1:1 stoichiometry of heme to protein (Fig. 2C). The affinity for heme was somewhat higher (lower Kd) than was found for the S. aureus heme oxygenases (25) and similar to that reported for the human enzyme (29). The data indicate that HmuQ binds heme in a manner consistent with other known heme oxygenases.

HmuQ catalyzes the degradation of heme. Heme oxygenases catalyze the opening of the heme macrocycle in the presence of an electron donor. Purified heme oxygenases do not readily release the product biliverdin in the absence of biliverdin reductase (30). Thus, most studies involve single turnover assays, as was done here. In addition, the in vivo electron donor to bacterial heme oxygenases is not known, but ascorbate or NADPH-cytochrome P450 reductase may be used in catalysis of the pure enzyme (27, 30, 34).

In the first experiment, heme degradation catalyzed by HmuQ was measured spectrophotometrically using NADPH-cytochrome P450 reductase as the electron donor (Fig. 3A). In these experiments, NADPH was added to the reaction mixture in 10 µM increments and then scanned from 300 to 700 nm after each addition. The heme peak decreased with successive additions of NADPH. At later titrations, the absorption due to NADPH was observed at 340 nm, presumably because the heme substrate became depleted, and the NADPH was not completely oxidized. Heme degradation did not occur if any component in the reaction mixture was omitted (data not shown).


Figure 3
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  		FIG. 3. HmuQ-dependent degradation of heme. (A) Disappearance of heme using NADPH-cytochrome P450 reductase as the reductant. Ten µM aliquots of NADPH were added to the cuvette containing 10 µM HmuQ, 8 µM heme, and the other reagents as described in the text and were scanned 5 min after each addition. The spectrum at 340 nm observed at later points was due to NADPH. The arrow indicates a decrease in absorption with time. (B) Disappearance of heme using ascorbate as the reductant. Ten mM ascorbate was added at time zero and subsequently scanned at 5-min intervals. The arrow indicates a decrease in absorption with time. (C) The decrease in absorbance at 408 nm was observed within the first 5-min interval. This figure is representative of two independent experiments.

In the second experiment, HmuQ-dependent heme disappearance was measured using 10 mM ascorbate as the reductant (Fig. 3B). In this reaction, heme degraded more rapidly than was observed with NADPH-cytochrome P450, and most of the decrease was complete by 5 min after the addition of ascorbate. To confirm that the reduction was time dependent, we measured degradation within the first 5 min, and the absorption at 408 nm was monitored as a function of time with 600 nm as a reference wavelength. The decrease of heme was approximately linear for the first 4 min and then began to level off (Fig. 3C). No degradation was observed in the absence of recombinant HmuQ (data not shown).

Previous reports indicated that some heme binding proteins can degrade heme by a nonenzymatic mechanism called coupled oxidation (23). Heme degradation by this process involves the generation of peroxide by the heme protein, and thus coupled oxidation is prevented by the presence of catalase. The experiments described above were all carried out in the presence of catalase in a 0.5:1.0 ratio with HmuQ. Similar results were obtained in the absence of catalase (data not shown). Collectively, the findings show that HmuQ catalyzes the enzymatic degradation of heme.

Biliverdin is produced from HmuQ-catalyzed heme degradation. Biliverdin is the final tetrapyrrole product of heme cleavage catalyzed by heme oxygenases. Heme degradation by IsdG or IsdI from S. aureus is accompanied by a very small, broad peak in the 650-nm region of the spectrum, which may be biliverdin (25). This spectral feature was not readily observed for B. japonicum HmuQ (Fig. 3). To address whether biliverdin is produced as a product catalyzed by HmuQ, a heme degradation reaction was carried out as described above using ascorbate as the reductant, and the product was analyzed by liquid chromatography-mass spectrometry (see Materials and Methods). The reaction product had a major atomic peak (m/z) at 583.4, which was identical to the biliverdin standard (data not shown). Nonenzymatic cleavage of heme by coupled oxidation does not result in biliverdin production (17). These data show that biliverdin is produced by heme degradation catalyzed by HmuQ, which is expected for a heme oxygenase.

The hmuQ and hmuD genes complement a heme oxygenase mutant of Corynebacterium ulcerans. A C. ulcerans heme oxygenase mutant cannot use heme as the sole source of iron because heme oxygenase is needed to release the iron from the heme macrocycle (20). To characterize hmuQ and hmuD in vivo, we tested them for their ability to complement the C. ulcerans heme oxygenase mutant strain CU29 for the ability to grow on heme as the sole iron source (Fig. 4). In the presence of the metal chelator EDDHA, none of the strains tested grew in the absence of an exogenous iron source (data not shown), whereas all strains grew when the medium was supplemented with FeSO4 (Fig. 4). Heme supported growth of parent strain CU712 on EDDHA-containing plates, but mutant strain CU29 harboring an empty vector did not grow under those conditions. Introduction of hmuQ or hmuD into strain CU29 on a plasmid complemented the mutant in trans for heme-dependent growth. Similar results were found using the heme oxygenase gene hmuO from C. diphtheriae (Fig. 4). Complemented growth with either of the B. japonicum genes was somewhat less robust than what was observed for the C. diphtheriae hmuO gene but was substantial compared with the empty vector control. The in vivo complementation experiments are in good agreement with the in vitro work and support the conclusion that hmuQ and hmuD encode enzymes that degrade heme to allow for its utilization as a nutritional iron source.


Figure 4
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  		FIG. 4. Complementation of a Corynebacterium ulcerans hmuO mutation in trans with B. japonicum hmuQ or hmuD. Plasmid-borne genes were introduced into the C. ulcerans hmuO mutant strain CU29, and liquid cultures were spotted onto agar medium plates containing EDDHA to chelate iron and supplemented with either ferrous sulfate (Fe) or heme. No growth was observed on plates lacking an exogenous iron source (data not shown). Parent strain CU712 (wt) harboring an empty vector and mutant strain CU29 harboring C. diphtheriae hmuO were used as positive controls. Strain CU29 harboring an empty vector was the negative control. A negative image of the spotted cells is shown to improve the contrast for visualization.

Homologs of hmuQ and hmuD are found in many bacteria. Skaar et al. (25) concluded that homologs of S. aureus isdG and isdI genes are only found in a few related genera of gram-positive bacteria. We searched the nucleotide and protein databases for homologs of HmuQ and HmuD in other prokaryotes using the BLAST algorithm (1). We found homologs in 41 bacteria with at least 45% similarity to the B. japonicum proteins (data not shown). The proteins with the highest similarity were found in the {alpha}-Proteobacteria, to which B. japonicum belongs, but many were also found in other gram-negative proteobacterial groups as well as gram-positive groups, Deinococcus-Thermus, and Chloroflexi (data available upon request).

To address directly whether another hmuQ/hmuD homolog encodes a heme degradation enzyme, the homolog from Brucella melitensis (annotated as bmeII0706 in the genome [6]) was overexpressed in E. coli, and the recombinant protein was characterized (Fig. 5). BmeII0706 bound heme with a Kd value of 1.3 µM (Fig. 5A and B), similar to what was observed for B. japonicum HmuQ (Fig. 2B). In addition, BmeII0706 catalyzed the degradation of heme using NADPH-cytochrome P450 reductase as the electron donor (Fig. 5C). Thus, the hmuQ/hmuD homolog in B. melitensis encodes a heme-degrading enzyme.


Figure 5
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  		FIG. 5. Characterization of recombinant BmeII0706 from Brucella melitensis. (A) Absorption spectra of heme bound to BmeII0706 as a function of the heme concentration. The experiment was carried out as described in the Fig. 2 legend. (B) Absorption of bound heme plotted as a function of the heme concentration. (C) Disappearance of heme using NADPH-cytochrome P450 reductase as the electron donor. The experiment was carried out as described in the Fig. 3 legend.


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DISCUSSION
 
In the present study, we provide evidence that B. japonicum hmuQ, and probably hmuD, encodes heme degradation enzymes. The hmuQ gene is clustered in the genome with heme transport genes (13) and is induced under iron limitation. In addition, HmuQ binds heme and catalyzes heme cleavage to produce biliverdin. Furthermore, both hmuQ and hmuD were able to complement a heme oxygenase mutant of C. ulcerans for growth on heme as an iron source. Thus, both in vitro and in vivo evidence support this conclusion.

HmuQ/HmuD homologs are found in many bacterial genera representing numerous taxonomic groups, and we confirmed that the B. melitensis homolog encodes a heme oxygenase (Fig. 5). Thus, this new class of heme-degrading oxygenases represented by the S. aureus and B. japonicum heme oxygenases is found throughout the {alpha}-Proteobacteria, and probably within numerous taxonomically diverse bacterial genera as well. Although utilization of exogenous heme has not been characterized in most of the bacteria identified from the BLAST search, homologs of the B. japonicum heme transport proteins HmuT/HmuR or the S. aureus heme permease IsdF were found in all of the identified species in which the complete genome sequence has been determined, except for Mycobacterium tuberculosis. These observations suggest that heme degradation enzymes identified in B. japonicum and S. aureus represent a class of protein found in many bacteria that can utilize exogenous heme.

HmuO in Corynebacterium diphtheriae and HemO of Neisseria meningitidis represent the classic heme oxygenase family. We used these proteins to search for homologs in the bacteria identified as having hmuQ/hmuD homologs. Nine of the 41 species contain hmuO or hemO homologs, whereas the remainder do not. This raises the possibility that the hmuQ/hmuD homolog is the only heme oxygenase gene in most of the identified bacteria. The role of multiple heme oxygenases in bacteria that have them appears to correspond with different cellular functions. Several bacteria have a heme oxygenase that provides the biliverdin prosthetic group to the BphP phytochrome in addition to an enzyme for heme iron utilization (4, 27). Heme oxygenase activity is needed for photosynthesis accessory pigment formation, and the hmuO-like gene from the photosynthetic bacterium Rhodopseudomonas palustris is clustered with genes encoding bacteriochlorophyll synthesis proteins. The B. japonicum hmuQ gene is clustered with heme transport genes needed for heme iron utilization (Fig. 1). hmuQ/hmuD homologs are also proximal to heme transport genes in the related {alpha}-proteobacterial species Agrobacterium tumefaciens, Nitrobacter winogradskyi, R. palustris, R. leguminosarum, Mesorhizobium loti, and Sinorhizobium meliloti. Further studies should clarify the diverse roles of heme oxygenases in bacteria.


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ACKNOWLEDGMENTS
 
We thank Michael Schmitt for C. ulcerans strains, the C. diphtheriae hmuO gene, plasmids, and helpful discussions. We also thank Angela Wilks for helpful discussions.

This work was supported by National Institutes of Health grant R01-GM067966 to M.R.O.


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FOOTNOTES
 
* Corresponding author. Mailing address: Department of Biochemistry, 140 Farber Hall, State University of New York at Buffalo, Buffalo, NY 14214. Phone: (716) 829-3200. Fax: (716) 829-2725. E-mail: mrobrian{at}buffalo.edu. Back


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Journal of Bacteriology, September 2006, p. 6476-6482, Vol. 188, No. 18
0021-9193/06/$08.00+0     doi:10.1128/JB.00737-06
Copyright © 2006, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Small, S. K., Puri, S., Sangwan, I., O'Brian, M. R. (2009). Positive Control of Ferric Siderophore Receptor Gene Expression by the Irr Protein in Bradyrhizobium japonicum. J. Bacteriol. 191: 1361-1368 [Abstract] [Full Text]  
  • Lee, W. C., Reniere, M. L., Skaar, E. P., Murphy, M. E. P. (2008). Ruffling of Metalloporphyrins Bound to IsdG and IsdI, Two Heme-degrading Enzymes in Staphylococcus aureus. J. Biol. Chem. 283: 30957-30963 [Abstract] [Full Text]  
  • Sangwan, I., Small, S. K., O'Brian, M. R. (2008). The Bradyrhizobium japonicum Irr Protein Is a Transcriptional Repressor with High-Affinity DNA-Binding Activity. J. Bacteriol. 190: 5172-5177 [Abstract] [Full Text]  
  • Kunkle, C. A., Schmitt, M. P. (2007). Comparative Analysis of hmuO Function and Expression in Corynebacterium Species. J. Bacteriol. 189: 3650-3654 [Abstract] [Full Text]  
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