OxyR acts as a transcriptional repressor of hydrogen peroxide-inducible antioxidant genes in Corynebacterium glutamicum R

Authors


Abstract

OxyR, a LysR-type transcriptional regulator, has been established as a redox-responsive activator of antioxidant genes in bacteria. This study shows that OxyR acts as a transcriptional repressor of katA, dps, ftn and cydA in Corynebacterium glutamicum R. katA encodes H2O2-detoxifing enzyme catalase, dps and ftn are implicated in iron homeostasis and cydA encodes a subunit of cytochrome bd oxidase. Quantitative RT-PCR analyses revealed that expression of katA and dps, but not of ftn and cydA, was induced by H2O2. Disruption of the oxyR gene encoding OxyR resulted in a marked increase in katA and dps mRNAs to a level higher than that induced by H2O2, and the oxyR-deficient mutant showed a H2O2-resistant phenotype. This is in contrast to the conventional OxyR-dependent regulatory model. ftn and cydA were also upregulated by oxyR disruption but to a smaller extent. Electrophoretic mobility shift assays revealed that the OxyR protein specifically binds to all four upstream regions of the respective genes under reducing conditions. We observed that the oxidized form of OxyR similarly bound to not only the target promoter regions, but also nonspecific DNA fragments. Based on these findings, we propose that the transcriptional repression by OxyR is alleviated under oxidative stress conditions in a titration mechanism due to the decreased specificity of its DNA-binding activity. DNase I footprinting analyses revealed that the OxyR-binding site in the four target promoters is ~ 50 bp in length and has multiple T-N11-A motifs, a feature of LysR-type transcriptional regulators, but no significant overall sequence conservation.

Abbreviations
EMSA

electrophoretic mobility shift assay

RACE

rapid amplification of cDNA ends

Introduction

Living organisms have evolved diverse antioxidant systems to protect themselves against the harmful effects of reactive oxygen species such as superoxide and hydrogen peroxide (H2O2) that are inevitably generated as by-products during aerobic respiration. In bacteria, OxyR is the best-studied transcriptional regulator of antioxidant genes, with its regulatory mechanism in Escherichia coli serving as a model for redox-responsive control of gene expression [1]. In this model, the LysR-type transcriptional regulator, OxyR, acts as an activator of genes encoding peroxide-detoxifying enzymes such as catalase, peroxidase and alkylhydroperoxide reductase. Oxidation of a cysteine residue on the OxyR protein plays an essential role in its redox-sensing mechanism [2-5], with the intramolecular disulfide bond subsequently formed involved in the conformational changes that allow the protein to bind to its target promoter region for transcriptional activation [3, 4]. OxyR-dependent regulation of primary antioxidant genes is conserved among a variety of bacteria. However, recent genome-wide analyses have expanded the OxyR regulon, members of which seem to be different among bacteria, and have indicated diverse modes of OxyR-dependent regulation of gene expression coordinated with other regulatory pathways [6-12]. In addition, the role of OxyR in the virulence of pathogenic bacteria is of interest given that host cells generate reactive oxygen species as a defense system against invading pathogens [13-15].

By contrast to the E. coli model, OxyR has a dual function, acting as not only an activator of peroxide-scavenging enzymes under oxidative stress conditions, but also a repressor of the same target genes under nonstress conditions in some bacterial species such as Xanthomonas campestris [16-18], Burkholderia pseudomallei [19] and Neisseria meningitidis [20]. Although both the reduced and oxidized forms of OxyR bind to the target promoter regions in X. campestris and B. pseudomallei, it is not clear whether the repression in N. meningitidis occurs through direct OxyR binding to the target promoter or through indirect effects on the basal promoter strength. It is noted that inactivation of oxyR results in increased resistance to H2O2 in Neisseria species including N. meningitidis and N. gonorrhoeae [21]. This is in contrast to a H2O2-sensitive phenotype of an oxyR-deficient mutant in many bacterial species, including E. coli. These findings suggest diverse modes of action of bacterial OxyR even in the regulation of primary antioxidant genes.

Corynebacterium glutamicum is a high-GC Gram-positive bacterium, which has a long history in the industrial production of amino acids, and is expected to be a versatile workhorse in the production of biofuels and biochemicals [22-28]. In addition, this nonpathogenic soil bacterium has received increasing interest as a model organism for the suborder Corynebacterineae, including medically important pathogens such as Corynebacterium diphtheriae and Mycobacterium tuberculosis [29-31]. Recently, it was reported that disruption of oxyR results in increased resistance to H2O2-induced oxidative stress in C. glutamicum and C. diphtheriae [32]. In C. diphtheriae, OxyR acts as a repressor of a catalase gene with a major role in resistance to H2O2. However, OxyR-mediated gene expression showed no response to oxidative stress in vivo, and the in vitro DNA-binding activity of the OxyR protein was not affected by its redox state [32]. In C. glutamicum, no OxyR-dependent gene expression has been reported to date.

In this study, we show that OxyR acts as a repressor of the catalase gene katA and three other genes dps, ftn and cydA in C. glutamicum R. dps and ftn encode iron-binding proteins implicated in the cellular homeostasis of iron. cydA encodes a subunit of cytochrome bd oxidase which has a role in aerobic respiration under low oxygen tension. In vitro DNA-binding analyses reveal nonspecific DNA-binding activity of the oxidized form of the OxyR protein in contrast to the specific binding to the four target promoter regions under reducing conditions. We also show that H2O2 induces the in vivo expression of katA and dps, but not of ftn and cydA. Based on these results, we propose a new model for OxyR-mediated redox-responsive regulatory mechanism.

Results

Involvement of oxyR and katA in H2O2 resistance

Previously, we performed DNA microarray analyses to compare the transcriptional profiles of densely packed C. glutamicum cells under oxygen deprivation with those in exponentially growing cells under aerobic conditions [33]. Under the former conditions applicable for a high-productivity bioprocess, the expression of 161 genes was upregulated. In the course of searching for the transcriptional regulators of these genes, we found that expression of three genes, dps, ftn and cydA, is negatively regulated by OxyR. OxyR is one of the best-studied redox-responsive transcriptional regulators of antioxidant genes in bacteria. Although a H2O2-resistant phenotype of the C. glutamicum oxyR-deficient mutant was recently shown using agar-diffusion growth-inhibition assays [32], the OxyR-dependent gene expression in this bacterium remained unknown. In this study, we examined expression of the dps, ftn and cydA genes in response to H2O2. In addition, expression of katA encoding a catalase supposedly involved in resistance to H2O2 was analyzed.

Prior to the gene expression analyses, effects of H2O2 on aerobic growth of a C. glutamicum wild-type strain, an oxyR-deficient mutant strain (a transposon-insertion mutant oxyR::Tn) and a katA-deficient mutant strain (katA::Tn) were compared (Fig. 1). These strains were cultured in minimal BTM medium with or without H2O2. In the presence of 30 mm H2O2, growth of the wild-type strain was suppressed during the first 3 h of cultivation, but the subsequent growth rate was comparable with that in the absence of H2O2 (Fig. 1A). In the presence of 100 mm H2O2, the wild-type strain hardly grew, whereas the oxyR::Tn strain grew well after a 3-h lag period (Fig. 1B), exhibiting a H2O2-resistant phenotype. By contrast, the katA::Tn strain hardly grew in the presence of a much lower concentration of H2O2 at 100 μm (Fig. 1C), showing a H2O2-sensitive phenotype. We confirmed that disruption of either oxyR or katA had no effect on growth in the absence of H2O2.

Figure 1.

The effects of disruption of oxyR and katA on the growth of C. glutamicum in the presence of H2O2. Wild-type (A) and oxyR::Tn (B) strains were cultured in minimal BTM medium containing 40 mm glucose (■) or the same medium supplemented with H2O2 at 30 mm (○) or 100 mm (△). (C) The wild-type (□ and △) and katA::Tn (● and image_n/febs12312-gra-0001.png) strains were cultured in the presence (△ and image_n/febs12312-gra-0002.png) or absence (□ and ●) of H2O2 at 100 μm. The D610 of the cell culture was monitored. Similar results were obtained from at least two independent cultivations, and representative results are shown.

H2O2-inducible expression of katA and dps

In order to examine gene expression in response to H2O2, C. glutamicum wild-type cells were cultured in minimal BTM medium for 4 h, and the exponentially growing cells were then supplemented with H2O2 at 30 or 100 mm. Changes in gene expression upon treatment with H2O2 for 5 and 15 min were analyzed using quantitative RT-PCR (Fig. 2). The level of katA mRNA increased 10-fold within 5 min of H2O2 supplementation, and then decreased to near the initial level over the subsequent 10 min (Fig. 2A). Transient induction of katA expression was also observed when cells were exposed to 100 mm H2O2, although the induction level was slightly lower than the level induced by 30 mm H2O2 (Fig. 2A), probably because of more severe oxidative damage to the cells. A similar response to H2O2 was observed for the expression level of dps (Fig. 2B). In contrast to the case of katA and dps, H2O2 had little effect on the level of ftn and cydA mRNAs under these conditions (Fig. 2C,D).

Figure 2.

The effects of H2O2 on gene expression. Exponentially growing C. glutamicum wild-type cells cultured in minimal BTM medium containing 40 mm glucose were exposed to H2O2 at 30 or 100 mm. The levels of katA (A), dps (B), ftn (C) and cydA (D) mRNAs in cells upon 0-, 5-, and 15-min exposure to H2O2 were determined by quantitative RT-PCR. The mRNA levels are presented relative to the value obtained from cells before exposure to 30 mm H2O2 (0 min). The values represent the mean results from three independent cultivations, with standard errors.

Negative regulation of katA, dps, ftn, and cydA by OxyR

In order to examine the role of OxyR in gene expression, an oxyR inframe deletion mutant (ΔoxyR) was constructed. Moreover, a plasmid carrying the oxyR gene under the control of a constitutive promoter was introduced into the ΔoxyR strain, and growth of the resultant oxyR-complemented strain (ΔoxyR/pCRB1-oxyR) was compared with that of the ΔoxyR and wild-type strains, both of which were transformed with a vector plasmid (ΔoxyR/pCRB1 and WT/pCRB1). We confirmed that ΔoxyR/pCRB1 grew as well as oxyR::Tn described above in the presence of 100 mm H2O2, whereas strains ΔoxyR/pCRB1-oxyR and WT/pCRB1 hardly grew under the same conditions (Fig. S1). All of these strains showed similar growth in the absence of H2O2. These results indicated that C. glutamicum OxyR negatively regulates its cellular defense system against oxidative stress induced by H2O2.

When exponentially growing cells of the WT/pCRB1 strain were exposed to 30 mm H2O2 for 5 min, a marked increase in the level of katA mRNA was observed (Fig. 3A) similar to that in the wild-type strain described above (Fig. 2A). In the absence of H2O2, the level of katA mRNA in the ΔoxyR/pCRB1 strain was higher than that induced by H2O2 in the WT/pCRB1 strain. The expression level of katA in the ΔoxyR/pCRB1 strain remained high upon exposure to H2O2. We confirmed that introduction of the oxyR-expression plasmid into the ΔoxyR strain represses the expression of katA to the wild-type level under nonstress conditions. In this strain, induction by H2O2 was less prominent than that in WT/pCRB1 probably because of the overexpression of oxyR. The expression level of oxyR in ΔoxyR/pCRB1–oxyR was ~ 20-fold higher than that in WT/pCRB1. As expected, H2O2-inducible expression of dps was observed in the WT/pCRB1 strain (Fig 3B). Quantitative RT-PCR analyses revealed that the expression of dps was under the strict negative control of OxyR in a manner similar to that observed for katA (Fig. 3B). Disruption of oxyR also resulted in increased expression of ftn (Fig. 3C) and cydA (Fig. 3D). However, the effects of oxyR inactivation on these two genes, whose expression shows little response to H2O2, were smaller than those on the two H2O2-inducible genes katA and dps.

Figure 3.

The effects of oxyR disruption on gene expression. Exponentially growing C. glutamicum cells cultured in minimal BTM medium containing 40 mm glucose were exposed to H2O2 at 30 mm. The strains used were wild-type (WT) and ΔoxyR, transformed with a plasmid for oxyR expression (pCRB1–oxyR) or a control vector plasmid (pCRB1). The levels of katA (A), dps (B), ftn (C) and cydA (D) mRNAs in cells with (+) or without (−) 5-min exposure to H2O2 were determined by quantitative RT-PCR. The mRNA levels are presented relative to the value obtained from WT/pCRB1 cells exposed to H2O2. The values represent the mean results from three independent cultivations, with standard errors.

In vitro binding of OxyR to the upstream regions of katA, dps, ftn and cydA

In order to examine DNA-binding activity in vitro, the OxyR protein was expressed in E. coli and purified. Electrophoretic mobility shift assays (EMSAs) were carried out with the OxyR protein and the upstream regions of the katA, dps, ftn and cydA genes. The DNA probes used and the results of EMSAs are summarized in Fig. 4. It is known that purified OxyR proteins derived from many bacteria are prone to oxidation. When the DNA-binding reactions were performed in the presence of a high concentration of dithiothreitol (200 mm), the purified C. glutamicum OxyR protein reduced the electrophoretic mobility of the DNA probes (katA-P1, dps-P1, ftn-P1 and cydA-P1) each including the upstream region of these target genes; the amount of an OxyR–DNA complex increased as the concentration of the OxyR protein increased (Fig. 5A–D). By contrast, no binding of OxyR to the promoter region of the cgR_0124 gene, which is a copper-inducible gene [34], was observed (Fig. 5H). These results indicated that OxyR specifically binds to the upstream regions of katA, dps, ftn and cydA under reducing conditions. In the presence of a low concentration of dithiothreitol (0.5 mm), the OxyR protein also bound to the upstream region of katA (Fig. 5E), dps (Fig. S2) and cydA (Fig. 5F). However, significant binding of OxyR to the negative control probe (the cgR_0124 promoter) was unexpectedly observed (Fig. 5G). These results suggested that the oxidized form of OxyR has nonspecific DNA-binding activity.

Figure 4.

Upstream regions of katA, dps, ftn, and cydA on the chromosome of C. glutamicum R. DNA probes used for EMSAs are indicated with the results (+/−: positive/no binding to the reduced form of OxyR) under the corresponding region on the chromosome. The OxyR-binding region determined by DNase I footprinting analyses is indicated by a black box. The transcription start site determined is indicated by a bent arrow.

Figure 5.

EMSAs with the OxyR protein and the upstream regions of katA, dps, ftn and cydA. The DNA probes used were katA-P1, dps-P1, ftn-P1, and cydA-P1 (Fig. 4). The promoter region of the cgR_0124 gene was used as a negative control DNA. The 20-μL binding reaction mixture containing dithiothreitol at 200 mm for reduced OxyR (OxyRred) or 0.5 mm for oxidized OxyR (OxyRox), 40 ng of the DNA probe, and the OxyR protein at various concentrations (lane C, no protein; lane 1, 23 ng; lane 2, 46 ng; lane 3, 92 ng) was subjected to electrophoresis on a 6% polyacrylamide gel. (image_n/febs12312-gra-0003.png) Free DNA, (image_n/febs12312-gra-0004.png) major DNA–protein complex.

In order to narrow down the OxyR-binding site, the DNA probes for the katA, dps and cydA upstream regions used in EMSAs shown in Fig. 5 were each split into two DNA fragments (Fig. 4). In the presence of a high concentration of dithiothreitol (200 mm), the OxyR protein bound to the DNA fragment (katA-P2) between −185 and +12 bp with respect to the translation start site of katA, but not to the further upstream region (katA-P3) between −391 and −185 bp (Fig. 6, lanes 1–12). Under reducing conditions, OxyR also bound to the DNA fragment (dps-P2) between −188 and +12 bp with respect to the translation start site of dps, but not to the further upstream region (dps-P3) within the ORF of the upstream cgR_2889 gene (Fig. 6, lanes 13–18). By contrast, binding of OxyR to the cgR_2889 ORF region (dps-P3) as well as to the dps-cgR_2889 intergenic region (dps-P2) was observed in the presence of a low concentration of dithiothreitol (Fig. 6, lanes 19–24), which is consistent with the nonspecific DNA-binding activity of the oxidized form of OxyR shown in Fig. 5. We also confirmed that the reduced OxyR protein binds to the DNA fragment (cydA-P2) between −188 and +12 bp with respect to the translation start site of cydA, but not to the fragment (cydA-P3) between −388 and −189 bp (data not shown).

Figure 6.

EMSAs with the OxyR protein and the two target gene upstream regions each split into two DNA fragments. The DNA probes used were katA-P2 and katA-P3 for the katA upstream region, and dps-P2 and dps-P3 for the dps upstream region (Fig. 4). The 20-μL binding reaction mixture containing dithiothreitol (DTT; 0.5 or 200 mm), 40 ng of the DNA probe and the OxyR protein at various concentrations (lanes 1, 7, 13, 16, 19 and 22, no protein; lanes 2 and 8, 23 ng; lanes 3 and 9, 46 ng; lanes 4 and 10, 92 ng; lanes 5, 11, 14, 17, 20 and 23, 184 ng; lanes 6, 12, 15, 18, 21 and 24, 369 ng) was subjected to electrophoresis on a 6% polyacrylamide gel. (image_n/febs12312-gra-0005.png) Free DNA, (image_n/febs12312-gra-0006.png) major DNA–protein complex.

OxyR-binding site in the upstream regions of katA, dps, ftn and cydA

DNase I footprinting analyses showed that the OxyR protein protected an ~ 50-bp region each upstream of katA, dps, ftn and cydA (Figs 7 and 8). In each case of katA, dps and cydA, both the reduced and oxidized forms of OxyR protected almost the same region, implying that preferential binding of OxyR to its specific target site remained in its oxidized state under the experimental conditions. In the case of ftn, the protection region by the oxidized form of OxyR was less than that by the reduced form, and one of the three hypersensitive bands observed in the DNase I footprinting analysis was significantly intensified in the presence of a low concentration of dithiothreitol (Fig. 7, ftn, upper hypersensitive band). These results suggested that the OxyR protein differentially binds to the ftn upstream region in a redox-responsive manner. No significant sequence similarity was found among the four OxyR-binding sites (Fig. 8). The patterns of the hypersensitive bands observed in the DNase I footprinting analyses were also different among these sites (Figs 7 and 8). It is known that there are multiple T–N11–A motifs in the binding sites of the LysR family transcriptional regulators including OxyR [35]. There are five, four, five and six T–N11–A motifs in the OxyR-binding site upstream of the katA, dps, ftn and cydA genes, respectively (Fig. 8). The transcription start site of katA and dps was determined by the rapid amplification of cDNA ends (RACE) method and the consensus sequence of the −10 region of C. glutamicum SigA-dependent promoters [36] was found (Figs 4 and 8). The OxyR-binding site overlaps the transcription start site of katA and the putative −10 region. No significant consensus sequence of the −35 region was found in the katA promoter. The transcription start site of dps was found to coincide with the adenine of its translation start site annotated (Fig. 8). Leaderless transcripts have been frequently found in C. glutamicum [37]. The OxyR-binding site overlaps the putative −10 and −35 regions of the dps promoter.

Figure 7.

DNase I footprinting analyses with the OxyR protein and the upstream regions of katA, dps, ftn and cydA. The 20-μL binding reaction mixture containing dithiothreitol (0.5 or 200 mm), 80 ng of the DNA probe and the OxyR protein at various concentrations (lanes 1 and 4, no protein; lanes 2 and 5, 368 ng; lanes 3 and 6, 736 ng) was subjected to DNase I treatment followed by electrophoresis on a 5.5% sequencing gel. The protected region and the hypersensitive band are indicated by a solid line and an arrow, respectively.

Figure 8.

OxyR-binding sites in the katA, dps, ftn and cydA upstream regions. The nucleotide sequences upstream of katA (A), dps (B), ftn (C) and cydA (D) are shown. The translation start codon is boxed. The determined transcription start site (+1) is indicated in bold with a bent arrow. The putative −10 and −35 regions are indicated by asterisks above the sequences. The protection region by OxyR in DNase I footprinting analysis (Fig. 7) is indicated by grey background. The position of the T-N11-A motif is indicated by the T and A along with a dotted line below the sequences. The position of the hypersensitive band in DNase footprinting analysis is indicated by a vertical bar below the sequences.

Redox-dependent changes in the DNA-binding specificity of OxyR

The effects of changes in concentrations of dithiothreitol on the DNA-binding activity of OxyR were examined by EMSAs using specific (katA-P2) and nonspecific (katA-P3) DNA probes for the katA upstream region as shown in Fig. 9. In the presence of dithiothreitol at 0.5 and 50 mm, one or two minor complexes between OxyR and the specific DNA probe (katA-P2) showing electrophoretic mobility lower than that of the major complex were detected probably caused by nonspecific DNA binding of OxyR under these conditions (Fig. 9, upper). These minor complexes disappeared in the presence of higher concentrations of dithiothreitol (100, 150 and 200 mm). However, the amount of a major complex between OxyR and the nonspecific DNA probe (katA-P3) observed in the presence of 0.5 mm dithiothreitol was markedly decreased in the presence of 50 mm dithiothreitol, and supplementation with higher concentrations of dithiothreitol (100, 150 and 200 mm) resulted in disappearance of this complex (Fig. 9, upper). In the presence of 0.5 mm dithiothreitol, addition of H2O2 at 1, 3 and 10 mm barely affected the binding activity of OxyR (Fig. 9, lower), indicating that the OxyR proteins were fully in the oxidized state in the presence of 0.5 mm dithiothreitol. This is consistent with the previous findings that OxyR in other bacteria is prone to oxidation.

Figure 9.

The effects of various concentrations of dithiothreitol and H2O2 on in vitro DNA binding of OxyR. EMSAs were carried out using the OxyR protein and the katA upstream region. The DNA probes used were katA-P2 and katA-P3 (Fig. 4). The 20-μL binding reaction mixture containing 40 ng of the DNA probe and 184 ng of the OxyR protein (+) supplemented with indicated concentrations of dithiothreitol (DTT) and H2O2 was subjected to electrophoresis on a 6% polyacrylamide gel. (image_n/febs12312-gra-0007.png) Free DNA, (image_n/febs12312-gra-0008.png) major DNA–protein complex.

Discussion

In this study, we showed that OxyR acts as a transcriptional repressor of four genes, katA, dps, ftn and cydA in C. glutamicum R. In vitro DNA-binding assays revealed that the OxyR protein specifically binds to their upstream regions under reducing conditions. However, the specificity of its DNA-binding activity is significantly decreased under oxidizing conditions. It should be noted that the amount of the complex between C. glutamicum OxyR and a DNA fragment containing its target site was not largely affected by changes in the redox conditions in EMSAs. Based on these findings, we propose that transcriptional repression by the reduced form of OxyR is alleviated under oxidative stress conditions in a titration mechanism because of the decreased specificity of the DNA-binding activity of OxyR. Among the four genes under the control of OxyR in C. glutamicum R found in this study, we detected H2O2-inducible expression of katA and dps in the wild-type strain. It is likely that their OxyR-mediated expression is derepressed, at least in part, in response to oxidative stress. By contrast, H2O2-dependent induction was hardly observed for expression of ftn and cydA under the conditions used. It is interesting to note that the former two H2O2-inducible genes were upregulated by oxyR disruption to a much greater extent than in the case of the latter two genes. The apparent affinity of the OxyR protein for its target promoters shown in EMSAs does not seem to correlate with the different response to H2O2 in expression of these four target genes in vivo. The difference in response to H2O2 in vivo among these genes may be because of possible different binding modes of the oxidized form of OxyR and/or different additional control of other regulatory systems, as discussed later.

The derepression of the OxyR-mediated gene expression in response to H2O2 in C. glutamicum is in contrast to the conventional E. coli model of the OxyR-dependent regulatory system, in which OxyR acts as an activator of antioxidant genes under oxidative stress. This is consistent with the effects of oxyR inactivation on the cellular resistance to oxidative stress. In the E. coli model, an oxyR-deficient mutant shows a H2O2-sensitive phenotype, whereas a C. glutamicum oxyR-deficient mutant shows a H2O2-resistant phenotype as shown in this and recent studies [32]. Corynebacterium belongs to actinobacteria including Streptomyces and Mycobacterium. On the genome of Streptomyces coelicolor, oxyR is located divergently from ahpCD encoding alkylhydroperoxide reductase [38]. This gene organization is conserved in several Mycobacterium species [39]. It has been reported that the expression of ahpCD is positively regulated by OxyR in S. coelicolor [38] and Mycobacterium marinum [40], as well as in the E. coli model, although M. tuberculosis oxyR is a pseudogene with multiple mutations [39]. By contrast, negative control of antioxidant genes by OxyR under nonstress conditions has been reported in limited bacterial species such as Xanthomonas [16], Burkholderia [19] and Neisseria [20], although expression of these genes is directly activated by OxyR in response to oxidative stress. It has been recently reported that C. diphtheriae OxyR directly represses a catalase gene (an orthologue of C. glutamicum katA), but shows no redox-responsive regulatory role in vivo and in vitro [32]. It is noteworthy that the oxyR-ahpCD gene organization found in Streptomyces and Mycobacterium, as described above, is also conserved in C. diphtheriae in contrast to its absence in C. glutamicum. The different compositions of antioxidant systems may be related to the difference in the role of the OxyR-mediated regulatory system between pathogenic and nonpathogenic Corynebacterium species. In this context, diverse redox-responsive regulatory systems have been established in actinobacteria [41], and it is noteworthy that a H2O2-inducible catalase/peroxidase gene is regulated by a Fur-type transcriptional regulator in M. tuberculosis [42, 43] and S. coelicolor [44].

A C. glutamicum OxyR-binding site in each of the promoter regions of katA, dps, ftn and cydA was determined by DNase I footprinting analyses. All of these binding sites are ~ 50 bp in length. Their overall sequences are less conserved, but have multiple T–N11–A motifs. These features are consistent with LysR-type transcriptional regulators including OxyR [35]. We observed that the OxyR-binding site overlaps the transcription start site and −10 region of katA. In the dps promoter, the OxyR-binding site was found to overlap −10 and −35 regions. These results suggest that binding of OxyR to its target site prevents RNA polymerase from interacting with these promoters, resulting in transcriptional repression. The role of the T–N11–A motif, which is differently distributed in the multiple OxyR target promoters, in their promoter activity will be the subject of a future study.

As described earlier, the EMSAs in this study indicated that C. glutamicum OxyR in its oxidized form has nonspecific DNA-binding activity in contrast to the specific binding of its reduced form to its multiple target promoters. However, DNase I footprinting analyses using the oxidized form of OxyR showed that the protein still bound to the target site preferentially in the respective DNA probes used. We detected a difference between the reduced and oxidized forms of OxyR in the extent of the protection region by OxyR in the ftn promoter and also a difference in the intensity of one of the three hypersensitive bands, implying that the DNA-binding property of OxyR changes in a redox-responsive manner. It should be noted that two cysteine residues proposed to form an intramolecular disulfide bond in the activated form of E. coli OxyR are conserved in C. glutamicum OxyR. Elucidation of the role of these cysteine residues in the OxyR function and identification of other critical amino acid residues, if any, will be an interesting subject for future studies.

Corynebacterium glutamicum katA encoding a monofunctional catalase is induced by H2O2 and directly regulated by OxyR, as shown in this study. Disruption of katA resulted in decreased resistance to H2O2, confirming that the H2O2-detoxifing enzyme encoded by this gene plays a pivotal role in protection of the cells against oxidative stress. A homology search of the published genomic sequences of multiple C. glutamicum strains did not detect any other homologues of representative peroxide-detoxifying enzymes such as monofunctional catalase, bifunctional catalase–peroxidase and alkylhydroperoxide reductase. These findings suggest that OxyR is involved in the protection of C. glutamicum cells against H2O2-induced oxidative stress mainly through the regulation of katA expression. In addition, the Dps protein encoded by another H2O2-inducible gene, dps, under the control of OxyR is supposed to have an antioxidant role by its iron-sequestration and/or nonspecific DNA-binding activity [45-48]. It has been reported that dps, along with ftn, is directly regulated by an iron-responsive transcriptional regulator DtxR [49, 50]. They are implicated in the homeostasis of iron which is possibly related to the generation of reactive oxygen species by the Fenton reaction, although their roles have not been characterized in C. glutamicum. It should be noted that katA is indirectly regulated by DtxR via direct control of another transcriptional regulator RipA which is transcriptionally regulated by DtxR [50, 51]. These findings suggest that katA, dps and ftn are coordinately regulated by OxyR and DtxR to efficiently protect cells from reactive oxygen species in response to changes in the related intra-/extracellular conditions. Although dps and ftn are regulated under the direct control of both OxyR and DtxR, no significant response to H2O2 in ftn expression was detected, in contrast to the H2O2-dependent upregulation of dps, implying the differential involvement of an additional unknown regulatory system in the expression of these genes. In addition, expression of cydA, which was shown to be directly regulated by OxyR in this study, was not induced by H2O2. cydA encodes a subunit of cytochrome bd oxidase that has a role with its high affinity for oxygen in aerobic respiration under low oxygen tension [52]. We have previously observed that cydA, along with the downstream genes cydB, cydD and cydC, is significantly upregulated not only under oxygen deprivation in densely packed cells for its application for highly efficient bioprocesses [33], but also under anaerobic growth conditions using nitrate as a terminal electron acceptor [53]. The regulatory mechanism of this putative operon remains unclear and may mask the OxyR-mediated response to H2O2 in cydA expression under the conditions used in this study.

A great advance was recently made in describing a genome-wide network of transcriptional regulation in C. glutamicum [29, 54], which is of great interest as a fundamental feature of this workhorse for industrial production of various useful compounds and of this nonpathogenic model organism phylogenetically related to human pathogens. Other than the OxyR-mediated regulatory system, multiple global and specific gene-regulatory systems in response to oxidative stress have been characterized in C. glutamicum, such as extracytoplasmic function-type sigma factors SigH [55] and SigM [56] and transcriptional regulators QorR [57], CyeR [58] and RosR [59]. Elucidation of the coordinated and complementary roles of a variety of these regulatory systems will be required to establish the integrated adaptive response of cells exposed to various adverse conditions, either naturally or biotechnologically.

Materials and methods

Bacterial strains and culture conditions

Corynebacterium glutamicum R (JCM 18229) [60] was used as a wild-type strain. Mutant strains deficient in each of oxyR (cgR_1755) and katA (cgR_0322) were obtained from a mutant library constructed by transposon-mediated mutagenesis [61], and are designated here oxyR::Tn and katA::Tn, respectively.

For genetic manipulations, E. coli and C. glutamicum strains were grown as described previously [62].

For analytical purposes, C. glutamicum starter culture was grown aerobically in 10 mL of nutrient-rich A medium [24] containing 1% glucose at 33 °C in a 100-mL test tube overnight. The cells were inoculated in fresh medium at a dilution of 100-fold or higher. Cells were cultured in 100 mL minimal BTM medium [63], each containing 40 mm glucose, at 33 °C in a 500-mL flask. To assess the response to H2O2, the medium was supplemented with H2O2 at the stated concentrations.

Cell growth was monitored by measuring D610 using a spectrophotometer (DU640, Beckman Coulter, Brea, CA, USA).

DNA techniques

Chromosomal and plasmid DNA were prepared from C. glutamicum, and the target DNA regions were amplified by PCR, as described previously [62].

Corynebacterium glutamicum cells were transformed by electroporation as described previously [64]. Escher-ichia coli cells were transformed by the CaCl2 procedure [65].

DNA sequencing was performed with ABI Prism 3100xl Genetic Analyzer (Applied Biosystems, Foster City, CA, USA). DNA sequence data were analyzed with the genetyx program (Software Development, Tokyo, Japan).

Construction of an oxyR inframe deletion mutant and an oxyR-expression plasmid for complementation

To construct an inframe deletion mutant, the upstream and downstream regions of the oxyR gene were amplified using sets of primers summarized in Table S1. The amplicon was fused and cloned into pCRA725 [23], a suicide vector for markerless gene disruption. The resultant plasmid pCRC323 was introduced into C. glutamicum R, and screening for deletion mutants was performed as previously described [23]. Deletion of oxyR was confirmed by PCR.

The region for the oxyR ORF was amplified by PCR using the C. glutamicum R chromosomal DNA as a template and a set of primers with appropriate restriction sites (Table S1; oxyR-Fw and oxyR-Rv). The amplicon was digested with the restriction enzymes, and inserted into the corresponding site downstream of the lac promoter in an E. coli–Corynebacterium shuttle vector pCRB1 [66], yielding pCRC324 for the constitutive expression of oxyR in C. glutamicum R.

Quantitative RT-PCR analysis and rapid amplification of cDNA ends

Total RNA was prepared from C. glutamicum cells using RNeasy minikit and RNAprotect Bacteria reagent (Qiagen, Hilden, Germany), and quantitative RT-PCR analysis was performed using Applied Biosystems 7500 Fast Real-Time PCR System, as described previously [62]. The primers used are listed in Table S2. The relative abundance of the target mRNAs was quantified based on the cycle threshold value. To standardize the results, the relative abundance of 16S rRNA was used as the internal standard.

The 5′-end of mRNA was determined by RACE using primers summarized in Table S3. Using a 5′-Full RACE Core Set (Takara, Osaka, Japan), single-stranded cDNA synthesized from total RNA using the 5′-phosphrylated primer was self-ligated with T4 RNA ligase. The first PCR was performed using inverted primers, and the second PCR was performed using nested inverted primers with an EcoRI site in their 5′-ends. The amplicon was digested with EcoRI and inserted into the corresponding site of a plasmid pHSG398 (Takara). We obtained > 10 clones from E. coli transformed with the resulting plasmid showing the same 5′-end of the mRNA by their sequences determined.

Purification of the OxyR protein expressed in E. coli

A DNA fragment containing the oxyR gene was amplified by PCR using a primer pair (Table S1; oxyR–Ex-F and oxyR–Ex-R). The amplicon was digested with NdeI and EcoRI, and was inserted into the corresponding site of the pET-28a expression vector (Merck KGaA, Darmstadt, Germany). The resulting plasmid, pCRC325, contains the recombinant gene encoding OxyR fused to a His tag sequence at the N-terminus. Escherichia coli BL21(DE3) cells transformed with pCRC325 were grown at 37 °C in 100 mL of Luria–Bertani medium supplemented with kanamycin (50 μg·mL−1). The recombinant gene was expressed in exponentially growing cells (D610 = 0.6) by adding 1 mm isopropyl-β-d-thiogalactooside. After 1 h of incubation, cells were harvested by centrifugation. The His-tagged OxyR protein was extracted and purified by affinity column chromatography using a Ni-NTA Fast Start Kit (Qiagen). The purified protein was loaded onto a gel-filtration column (PD-10 column; GE Healthcare UK, Chalfont ST Giles, UK) equilibrated with buffer containing 20 mm Tris/HCl (pH 7.5), 100 mm NaCl, 5 mm MgCl2, 0.1 mm EDTA and 1 mm dithiothreitol, and was eluted with the same buffer. The His-tagged OxyR protein prepared was used for EMSA.

EMSA

EMSA was performed as described previously [67] with some modifications. The purified OxyR protein at stated concentrations was incubated with 40 ng of a DNA probe in 20 μL of binding buffer containing 20 mm Tris/HCl (pH 7.5), 50 mm NaCl, 2.5 mm MgCl2, 0.45 mm EDTA, 0.5 mm dithiothreitol, 0.05% Nonidet P-40 and 10% glycerol for 30 min at 25 °C. Dithiothreitol and H2O2 were added at the stated concentrations. The binding reaction mixture was subjected to electrophoresis on a 6% polyacrylamide gel containing 5% glycerol in 0.5× TBE electrophoresis buffer, and the DNA probe was detected with SYBR Green.

DNA probes were prepared by PCR using primers summarized in Fig. 4 and Table S4; for the katA upstream region, three sets of primers katA-Pr-F1/R1 (Probe katA-P1), katA-Pr-F2/R1 (katA-P2) and katA-Pr-F1/R2 (katA-P3) were used; for the dps upstream region, three sets of primers dps-Pr-F1/R1 (dps-P1), dps-Pr-F2/R1 (dps-P2) and dps-Pr-F1/R2 (dps-P3) were used; for the ftn upstream region, a set of primers ftn-Pr-F/R (ftn-P1) was used; for the cydA upstream region, three sets of primers cydA-Pr-F1/R1 (cydA-P1), cydA-Pr-F2/cydA-Pr-R1 (cydA-P2), cydA-Pr-F1/cydA-Pr-R2 (cydA-P3) were used; for the cgR_0124 promoter region, a set of primers 0124-Pr-F/R were used.

DNase I footprinting analysis

A labeled DNA fragment was prepared by PCR using primers listed in Table S4: a gene-specific primer (katA-Pr-F2, dps-Pr-F2, ftn-Pr-F and cydA-Pr-F2) and a 5′-IRD700-labeled primer IR-Rv. The length of the DNA probes used is 223, 226, 426 and 226 bp for the promoter region of katA, dps, ftn and cydA, respectively. A plasmid pCRA741 [33] containing each of the gene upstream regions was used as a template. The purified OxyR protein at stated concentrations was incubated with 80 ng of the DNA probe in 20 μL of the binding buffer (as described for EMSAs) for 25 min at room temperature. For analyses of the reduced form of OxyR, dithiothreitol was added at 200 mm. Four microliters of the binding buffer containing 1–2 mU DNase I (Takara), 5 mm MgCl2 and 10 mm CaCl2 were then added and incubated for 1 min, followed by the addition of 2 μL of 325 mm EDTA and subsequent heating at 80 °C for 10 min. The samples mixed with IR2 stop solution (Li-Cor, Lincoln, NE, USA) were heated at 95 °C for 3 min, and separated on a 5.5% KB plus gel matrix (Li-Cor) using a Li-Cor 4300 DNA analyzer. The DNA-sequencing reaction mixtures using the same IRD700-labeled primer and a DYEnamic direct cycle sequencing kit with 7-deaza-dGTP (GE Healthcare) were subjected to the same gel.

Acknowledgements

We thank Crispinus A. Omumasaba (RITE) for critical reading of the manuscript. This work was financially supported in part by the New Energy and Industrial Technology Development Organization (NEDO), Japan.

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