Detoxification of hydrogen peroxide and expression of catalase genes in Rhodobacter

Tanja Zeller and Gabriele Klug

Institut für Mikrobiologie und Molekularbiologie, University of Giessen, Heinrich-Buff-Ring 26-32, D-35392 Giessen, Germany

Correspondence
Gabriele Klug
Gabriele.Klug{at}mikro.bio.uni-giessen.de


   ABSTRACT
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
The two related facultatively photosynthetic bacteria Rhodobacter sphaeroides and Rhodobacter capsulatus show different sensitivities against peroxide stress. R. sphaeroides is able to tolerate higher concentrations of H2O2 and exhibits higher catalase activity than R. capsulatus. The katE gene of R. sphaeroides and the katG gene of R. capsulatus are strongly induced by H2O2. This induction depends on the presence of the OxyR protein, which is able to bind to the promoter regions of these genes. In addition to katE R. sphaeroides harbours the katC gene, which shows no significant response to H2O2 but is induced in stationary phase.


Abbreviations: ROS, reactive oxygen species


   INTRODUCTION
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
The production of reactive oxygen species (ROS) is an inherent consequence of an aerobic lifestyle for living organisms. Well-balanced defence systems are required to guarantee survival of aerobically living cells by removing ROS (Storz & Zheng, 2000), keeping the cellular environment reduced, and repairing oxidation-generated damage. Most organisms that can grow in the presence of oxygen harbour catalases and/or peroxidases which scavenge the H2O2 that is produced endogenously by the respiratory chain or is generated in the environment. H2O2 can damage enzymes by oxidizing thiol groups and iron-sulfur centres, and by conversion to hydroxyl radicals it can produce mutagenic and lethal lesions (Storz & Imlay, 1999). Two types of catalase, Mn-catalase and haem catalase, are presently known (Ueda et al., 2003), which catalyse the destruction of H2O2 with very high Kcat/Ks values to ensure rapid removal of these reactive molecules. H2O2 detoxification in Escherichia coli is mainly due to two distinct types of catalase. The katG gene encodes a bifunctional catalase hydroperoxidase I (HPI) that harbours both catalase and peroxidase activity. HPI is transcriptionally induced by the positive activator OxyR, which directly senses oxidative stress (González-Flecha & Demple, 1997). The katE gene encodes the monofunctional HPII (which harbours only catalase activity) and its transcription requires a functional rpoS gene, which is activated in stationary-phase cells or upon various types of starvation (González-Flecha & Demple, 1997). OxyR is not involved in the regulation of katE in E. coli (Schellhorn, 1994).

Bacteria of the genus Rhodobacter are facultatively photosynthetic bacteria. At low oxygen tension the formation of photosynthetic complexes is induced, which allows the bacteria to perform anoxygenic photosynthesis if no oxygen is available. As long as oxygen is present Rhodobacter species produce ATP by aerobic respiration. The molecular mechanisms underlying the rapid adaptation to changes in oxygen concentration have been extensively studied in the past (reviewed by Gregor & Klug, 1999, 2002; Zeilstra-Ryalls & Kaplan, 2004). In its natural environment Rhodobacter occasionally needs to tolerate very high oxygen concentrations which are produced by micro-organisms performing oxygenic photosynthesis. While some previous studies have addressed the role of superoxide dismutase and thioredoxins in defence against oxidative stress in Rhodobacter (Cortez et al., 1998; Pasternak et al., 1999; Li et al., 2003a, b), little is known about the role of catalases and their regulation. In this study we have addressed the ability of two Rhodobacter species to detoxify H2O2 and have studied the expression of catalase genes in order to understand better how Rhodobacter can deal with high oxygen concentrations in its environment.


   METHODS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
Bacterial strains, plasmids and growth conditions.
All strains and plasmids used in this study are listed in Table 1.


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Table 1. Bacterial strains and plasmids used in this study

 
E. coli cultures were grown in Luria–Bertani (LB) medium at 37 °C with continuous shaking at 180 r.p.m. Rhodobacter cultures were cultivated at 32 °C with continuous shaking at 140 r.p.m. in a malate minimal salt medium (Drews, 1983). For aerobic growth, 20 ml of culture were incubated in 100 ml baffled Erlenmeyer flasks (about 7·7 mg l–1 dissolved oxygen). For semi-aerobic incubation, 40 ml of culture were shaken in 50 ml Erlenmeyer flasks (about 0·85 mg l–1 dissolved oxygen). Ampicillin (200 µg ml–1), spectinomycin (10 µg ml–1) and tetracycline (20 µg ml–1) were added to the medium as needed.

Construction of oxyR deletion mutants of R. capsulatus and R. sphaeroides.
R. capsulatus strain SBoxyR : : {Omega}Sp and R. sphaeroides strain 2.4.1oxyR : : {Omega}Sp were generated by transferring the suicide plasmids pPHUSB{Delta}oxyR : : {Omega}Sp into R. capsulatus SB1003 and pPHU2.4.1{Delta}oxyR : : {Omega}Sp in R. sphaeroides 2.4.1, respectively, and screening for insertion of the omega ({Omega})-spectinomycin cassette into the chromosome by homologous recombination. Briefly, parts of the oxyR genes of R. capsulatus SB1003 or R. sphaeroides 2.4.1, together with upstream and downstream sequences were amplified by PCR using oligonucleotides SBoxyupEco (5'-GTGTTCGAATTCCCCGCC-3'), SBoxy315Bam (5'-GAAGATAGGATCCAATCGTCG-3') and SBoxy500Bam (5'-GAAGATAGGATCCAATCGTCG-3'), SBoxydownPst (5'-CAGGCGCTGCAGAGGGCG-3') for SB1003 and 2.4.1oxyupEco (5'-CGAATTCTGGTTGTCGGCGATC-3'), 2.4.1oxy290Bam (5'-GGGATCCCGCCCAGATTGAC-3'), 2.4.1oxy600Bam (5'-CGGATCCGCTGCGGCGGTGGCGC-3') and 2.4.1oxydownPst (5'-CCTGCAGGACGGCCGCGTGGA-3') for 2.4.1.

The amplified PCR fragments were cloned into the EcoRI–BamHI and BamHI–PstI sites of the suicide plasmid pPHU281, respectively, generating plasmids pPHUSB{Delta}oxy and pPHU2.4.1{Delta}oxyR. A 2·0 kb BamHI fragment containing the {Omega}-spectinomycin cassette from pHP45{Omega} was inserted into the BamHI site of pPHUSB{Delta}oxy and pPHU2.4.1{Delta}oxyR to generate pPHUSB{Delta}oxyR : : {Omega}Sp and pPHU2.4.1{Delta}oxyR : : {Omega}Sp, respectively. Both pPHUSB{Delta}oxyR : : {Omega}Sp and pPHU2.4.1{Delta}oxyR : : {Omega}Sp were transferred into E. coli strain SM10, and mobilized from SM10(pPHUSB{Delta}oxyR : : {Omega}Sp) or SM10(pPHU2.4.1{Delta}oxyR : : {Omega}Sp), into either R. capsulatus SB1003 or R. sphaeroides 2.4.1 wild-type strain by diparental conjugation. Conjugants were selected on malate minimal salt agar plates containing 10 µg spectinomycin ml–1. Southern blot analysis of chromosomal DNA was carried out to confirm the double crossover event of the {Omega}-spectinomycin cassette into the Rhodobacter chromosome. By insertion of the {Omega}-spectinomycin cassette, 181 bp of the 915 bp R. capsulatus oxyR gene and 247 bp of the 926 bp R. sphaeroides oxyR gene, respectively, were deleted.

Complementation of the oxyR deletion mutants of R. capsulatus and R. sphaeroides.
For complementation of the oxyR deletion mutant of R. capsulatus, a 1·4 kb PCR fragment containing the entire oxyR gene along with approximately 450 bp of the upstream sequence was amplified using the oligonucleotides SBoxyRup450Eco (5'-GACGTTGTCGGGCCAGGAATTCAGC-3') and SBoxydownKpn (5'-GGGGTACCTGGCCTCGGTCAGATTTG-3'). Following digestion with EcoRI and KpnI, the PCR fragment was cloned into the corresponding sites of pRK415, resulting in plasmid pSBoxyR. For complementation of the oxyR deletion mutant of R. sphaeroides, a 1·2 kb PCR fragment containing the entire oxyR gene along with 100 bp of the upstream sequence was amplified using the oligonucleotides 2.4.1oxyupEco (5'-CGAATTCTGGTTGTCGGCGATC-3') and 2.4.1oxydownPst (5'-CCTGCAGGACGGCCGCGTGGA-3'). After digestion with EcoRI and PstI, the PCR fragment was cloned into the corresponding sites of pRK415 resulting in plasmid p2.4.1oxyR. To complement the oxyR deletion in SB1003 and 2.4.1, respectively, plasmids pSBoxyR and p2.4.1oxyR were transferred into E. coli strain SM10 and conjugated into the SBoxyR or 2.4.1oxyR strains by diparental conjugation.

Enzyme assays.
Rhodobacter cells were disrupted by brief sonication, and crude extract was used for enzyme assays. Protein concentration was determined according to Bradford (1976). For total catalase measurement, 30–300 µg of total protein were diluted to 1 ml with water. H2O2 (0·5 ml of 59 mM H2O2 freshly diluted in 50 mM potassium phosphate buffer, pH 7·0) was added, and the absorbance of the samples at 240 nm was measured every 10 s for 1 min. The initial linear rates were used to calculate the activities. Specific activity of catalase (µM H2O2 decomposed per minute per milligram of total protein) was calculated using an extinction coefficient of 43·6 M–1 cm–1 (Hochman & Shemesh, 1987).

For analysis of H2O2 detoxification, cells were grown exponentially to an OD660 of 0·5 and H2O2 was added to the desired final concentration. Aliquots were drawn at various time points, and the amount of H2O2 was determined by oxidation of Fe2+ in the presence of xylenol orange and measuring the absorbance at 560 nm. The absorbance of standard concentrations of H2O2 served to quantify the amount of H2O2 for each experiment. As previously described by Perelman et al. (2003), the ability to decompose H2O2 is strongly dependent on cell density. Therefore, all of the analyses presented in this study were performed at a fixed cell density.

Measurement of resistance to H2O2.
Exponentially grown cultures of Rhodobacter strains (0·2 ml) were diluted into 2 ml of pre-warmed top agar (0·7 % agar) and layered onto minimal malate agar plates. Filter paper disks were placed on the surface of the agar, and 5 µl of different concentrations (0·001–0·2 M) H2O2 were applied on the filter disk. Zones of inhibition were measured after overnight incubation at 32 °C.

RNA extraction and quantitative real-time RT-PCR analysis.
Rhodobacter strains in exponential phase (OD660=0·5), grown under semi-aerobic conditions, were treated with H2O2 (1 mM final concentration) and at certain time-points cells were collected into centrifugation bottles and pelleted by centrifugation. Total RNA was isolated by the hot phenol method, quantified by spectrophotometric analysis (absorbance at 260 nm), and 60 ng of total RNA were used for quantitative real-time PCR.

The following primers were designed for gene amplification, katE: katE-A (5'-CTATCCGCTGATCGAGGT-3') and katE-B (5'-GTCGGCATAGGAGAAGAC-3'); katC: katC-A (5'-GGATGCGGCGATGCTAGCCGCCAAC-3') and katC-B (5'-GAGGGTTCCCGCCCAGCGTCGCC-3'); katG: katG-A (5'-GCCTCGGTCGCCGATGTGAT-3') and katG-B (5'-CACCGGCTCCAGCACGTCAA-3'); rpoZ genes: 2.4.1rpoZ-A (5'-TTCGAGCTGGTGATGCT-3') and 2.4.1rpoZ-B (5'-ACTCGATCTGGGTCTGG-3'); SBropZ-A (5'-GATGATCTGCGCGAGCGTCT-3') and SBrpoZ-B (5'-CCTTGCGCGTCCATCAATGC-3').

The rpoZ gene (encoding the {omega}-subunit of RNA-polymerase) of R. capsulatus and R. sphaeroides, respectively, was used to normalize expression values for all other genes. The One-Step RT-PCR kit (Qiagen) was used for reverse-transcription-PCR following the manufacturer's instruction, except that a total volume of 15 µl was used. SYBR Green was used to monitor amplification and to quantify the amount of PCR products using the Rotor-Gene 3000 real-time PCR cycler (LTF). Relative expression of kat and rpoZ mRNA were calculated after the method of Pfaffl (2001).

Expression and isolation of the R. capsulatus OxyR protein.
Oligonucleotides SBoxystartBam (5'-CCCCGGATCCTCTCGATGAAACAGC-3') and SBoxydownKpn (5'-GGGGTACCTGGCCTCGGTCAGATTTG-3'), which hybridize to the 5' and 3' regions of the oxyR gene, respectively, were used to amplify the oxyR coding region. The 950 bp PCR product was digested with BamHI and KpnI and ligated into pQE32 to generate pQEoxyR, which was transformed into E. coli JM109. The correct construct as confirmed by sequencing (using a Genetic Analyser 310 sequencer; ABI) was transformed into E. coli M15(pREP4) for overexpression of His-tagged OxyR. For this purpose M15(pREP4 pQEoxyR) was grown in 500 ml of LB medium to an OD600 of 0·7–0·8 and induced with 1 mM IPTG for 4–5 h at 32 °C. Following harvest, cells were resuspended in lysis buffer (50 mM Tris, pH 7·5; 250 mM NaCl; 3 mM imidazole; 1 µg lysozyme µl–1 and 0·1 mM PMSF) and disrupted by a brief sonication. The lysate was centrifuged at 12 000 r.p.m. for 20 min at 4 °C. The supernatant was loaded onto Ni-NTA agarose and incubated at 4 °C for 4–5 h. Proteins were washed with washing buffer (0·05 M Tris, pH 7·5 and 0·25 M NaCl) containing 20–50 mM imidazole and eluted with imidazole at a concentration between 80 and 100 mM. Aliquots of these fractions were analysed on SDS 15 % polyacrylamide gels, using the buffer system of Laemmli (1970). Fractions containing purified OxyR protein were pooled by using Centricon-10 columns (Amicon) and washed in Z buffer (50 mM HEPES, pH 8·0; 0·5 mM EDTA, pH 8·0; 10 mM MgCl2 and 300 mM KCl).

Gel chromatography (FPLC).
The purified OxyR protein in a 2 ml volume was applied to a Superdex-200 HR 16/60 gel filtration column (Pharmacia Biotech) equilibrated with Z buffer. For reducing conditions 200 mM DTT was added to the Z buffer. The protein was eluted with Z buffer and aliquots of the column fractions were analysed by SDS 10 % PAGE using the buffer system of Laemmli (1970).

Determination of OxyR redox status by using AMS.
For in vitro modification of free thiol groups, 4-acetamido-4'-maleimidylstibene-2',2'-disulfonic acid (AMS; Molecular Probes) was used. The addition of AMS leads to the alkylation of free thiol groups, present in the reduced but not in the oxidized OxyR. The addition of the high-molecular-mass AMS moiety to the reduced but not to the oxidized protein allows separation of the two forms by gel electrophoresis. The purified OxyR protein was first treated with freshly prepared H2O2 (1 mM) or DTT (200 mM) at room temperature for 30 min. The protein was then mixed with 10 % trichloroacetic acid (TCA). Precipitated protein was collected by centrifugation (10 000 r.p.m., 10 min). After complete removal of the supernatant, the pellet was dissolved in a buffer containing 0·1 % SDS; 50 mM Tris/HCl, pH 8·0 and 15 mM AMS (apart from the non-AMS-modified sample which was dissolved in the same buffer without AMS), and incubated for 2 h at 37 °C. The samples were loaded on SDS 10 % PAGE and visualized by silver staining.

Gel mobility-shift assay.
Binding of the OxyR protein to the katG and katE upstream region, respectively, was determined by gel mobility-shift assay. For this, a DNA fragment containing the katG or the katE promoter region was generated by PCR. The following oligonucleotides were used to generate the PCR fragments. katG: katGupBspEI (5'-CGTCCGGACCCGCGGCACCATC-3') and katGStuI (5'-AGGCCTTCATCGCGCCATGCAT-3'); katE: katEupEcoRI (5'-GTAGAATTCCTGCCGCAG-3') and katEPstI (5'-GTCTGCAGTCGGCGCGCCGGCCGT-3'). The PCR fragments with a length of 350 bp were cloned into T-vector (Promega), and isolated from the vector by using enzymes BspEI/StuI and EcoRI/PstI, respectively. The restricted DNA fragments were then radioactively labelled in a fill-in reaction with [{alpha}-32P]dCTP using the Klenow fragment.

Binding reactions were carried out in a final volume of 20 µl and contained an appropriate amount of protein, [{alpha}-32P]CTP-labelled DNA probe, and binding buffer (10 mM Tris/HCl, pH 8·0; 50 mM NaCl; 1 mM DTT; 1 mM EDTA and 5 % glycerol). Since purified OxyR protein is predominantly in the oxidized form, we added 200 mM DTT to the binding reaction to reduce OxyR. Binding incubations were carried out for 30 min at room temperature (25 °C) before the samples were loaded onto a 4 % polyacrylamide gel in 0·5x Tris Borate EDTA (TBE) buffer and run at 130 V for 2 h.

DNaseI footprinting assay.
A BamHI–StuI DNA fragment containing the katG promoter region was 5'-end labelled at the BamHI site as follows. Plasmid TkatGup (Table 1) was linearized with BamHI, dephosphorylated using alkaline phosphatase (NEB) and labelled with polynucleotide kinase (NEB) and [{gamma}-32P]ATP. The DNA was then digested with StuI and the 361 bp BamHI–StuI fragment was purified from a 6 % non-denaturing polyacrylamide gel. The end-labelled DNA fragment (approx. 10 000 c.p.m.) was incubated with different amounts of purified OxyR protein, using the same reaction conditions as described for the gel mobility-shift assay. After 30 min of incubation at room temperature, 5 µl of a buffer containing 0·01 M CaCl2 and 0·01 M MgCl2 were added. DNaseI (1 µl, 0·1 U µl–1; Promega) was then added for 2 min at room temperature. The reaction was stopped by adding 250 mM EDTA. After phenol/chloroform extraction, the samples were dissolved in formamide dye and loaded onto a 6 % sequencing gel. The DNA fragment containing the katG promoter region was sequenced with the primer SBkatG300seq (5'-GCACCATCGCCGCGCCCAG-3') by the dideoxy chain-termination method (Sanger et al., 1977) using the T7 sequencing kit from USB.


   RESULTS
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ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
R. sphaeroides 2.4.1 is more resistant to H2O2 and shows faster H2O2 detoxification than R. capsulatus SB1003
In order to compare the sensitivity of the two related Rhodobacter strains to H2O2 we performed inhibition zone assays. Fig. 1(a) shows the mean sizes of inhibition zones at the different concentrations of H2O2 which had been applied to the filters. R. sphaeroides 2.4.1 is clearly more resistant to H2O2 at lower concentrations and can tolerate higher concentrations of the agent. While R. capsulatus SB1003 showed significant growth inhibition when 12·5 mM H2O2 was spotted on the filter, R. sphaeroides 2.4.1 only showed inhibition when 25 mM H2O2 was applied. We also monitored the detoxification of H2O2 added to exponentially growing cultures of R. sphaeroides or R. capsulatus (Fig. 1b). Both strains showed faster removal of H2O2 when grown aerobically compared to semi-aerobic growth. While the two Rhodobacter strains showed identical rates of H2O2 detoxification under aerobic conditions, semi-aerobically grown R. sphaeroides cultures were clearly more efficient than semi-aerobic R. capsulatus cultures. Similar results were obtained in the measurement of the basal catalase activity (Table 2). While an activity of about 200 µM decomposed H2O2 min–1 (mg total protein)–1 was determined for aerobically grown R. sphaeroides cells in exponential phase, an activity of only 40 µM decomposed H2O2 min–1 (mg total protein)–1 was determined for R. capsulatus cells grown under identical conditions. In contrast to that, semi-aerobically grown cells of R. sphaeroides showed a reduced catalase activity of about 130 µM decomposed H2O2 min–1 (mg total protein)–1 and cells of R. capsulatus an activity of 10 µM decomposed H2O2 min–1 (mg total protein)–1. These results indicate that H2O2-scavenging enzymes are present in higher amounts or show higher activity under aerobic growth conditions.



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Fig. 1. Effect of H2O2 on wild-type strains of Rhodobacter. (a) Inhibition of growth of SB1003 (open bars) and 2.4.1 (filled bars) by H2O2. The values give the mean of five experiments with maximal deviation. (b) Time-course of H2O2 decomposition by aerobic (solid lines) or semi-aerobic (dashed lines) cultures of wild-type SB1003 ({square}) and 2.4.1 ({blacksquare}). H2O2 (3·5 mM) was added to the cultures at time-point 0. Each point represents the mean of four independent experiments. Error bars indicate maximal deviation.

 

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Table 2. Catalase activities of wild-type, oxyR mutant strains and oxyR-complemented strains of R. capsulatus and R. sphaeroides grown under aerobic or semi-aerobic conditions

 
The oxyR gene product is required for the H2O2-dependent response
To study the involvement of the global stress regulator OxyR in the H2O2-dependent response, we constructed oxyR deletion mutants of both Rhodobacter strains (see Methods). The deletion of the oxyR gene resulted in higher sensitivities of the mutants after exposure to H2O2 in comparison to the wild-types (Fig. 2a, b). The zones of growth inhibition were significantly larger for the oxyR mutants. The intact oxyR gene of 2.4.1 and SB1003, respectively, located on plasmids p2.4.1oxyR and pSBoxyR could restore the oxyR deletion phenotype (Fig. 2a, b). Moreover, the basal level of total catalase activity was reduced in both oxyR mutant strains (fivefold decrease in 2.4.1oxyR under aerobic and semi-aerobic conditions compared to the wild-type 2.4.1; 18- and 11-fold decrease, respectively, for SBoxyR under aerobic and semi-aerobic conditions compared to the wild-type SB1003) (Table 2). The increase in catalase activity after H2O2 treatment in the oxyR mutants was also reduced (1·3-fold increase in 2.4.1oxyR compared to 1·9-fold increase in 2.4.1 wild-type and 1·3-fold decrease in SBoxyR compared to 4·3-fold increase in SB1003 wild-type) (Table 2). The complemented oxyR mutant strains showed catalase activities that were clearly higher than that of both oxyR mutants, but the activities did not reach wild-type levels. In the complemented strains, the oxyR gene is present in 1–2 copies in trans, which may lead to altered expression compared to the chromosomally located wild-type gene. Both oxyR mutants exhibit kinetics of H2O2 removal which were clearly retarded when compared to the kinetics observed for the isogenic wild-type strains (Figs 2c and 1b). These data reveal an involvement of the oxyR gene product in the response to H2O2. Similar observations regarding the essential role of OxyR in the peroxide response have been made in other bacteria, such as E. coli, Haemophilus influenzae and Agrobacterium (González-Flecha & Demple, 1997; Maciver & Hansen, 1996; Nakjarung et al., 2003). The R. capsulatus mutant SBoxyR was more severely affected in its ability to detoxify H2O2 than the R. sphaeroides oxyR mutant. While 2.4.1oxyR was able to remove 3·5 mM H2O2 from the medium within 5 min, SBoxyR was unable to remove this amount of H2O2 even after 5 min. After addition of 3·5 mM H2O2 the growth of the R. capsulatus oxyR mutant was completely inhibited, while both wild-type strains, as well as the 2.4.1oxyR mutant, showed growth inhibition for a period of 60 min after the addition of H2O2, but were able to grow after that time of adaptation (data not shown).



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Fig. 2. Sensitivity of Rhodobacter oxyR mutants to H2O2. Growth inhibition by H2O2 of (a) R. capsulatus [open bars, wild-type SB1003; filled bars, SBoxyR; hatched bars, complemented mutant SBoxyR(pSBoxyR)] and (b) R. sphaeroides [open bars, wild-type 2.4.1; filled bars, 2.4.1oxyR; hatched bars, complemented mutant 2.4.1oxyR(p2.4.1oxyR)]. The values give the mean of five experiments with maximal deviation. (c) Time-courses of H2O2 decomposition by aerobic (solid lines) or semi-aerobic (dashed lines) oxyR mutant strains SBoxyR ({bullet}) and 2.4.1oxyR ({blacktriangleup}). At time-point 0 3·5 mM H2O2 was added to the cultures. Each point represents the mean of four independent experiments. Error bars indicate maximal deviation.

 
H2O2-dependent katG expression in R. capsulatus and katE gene expression in R. sphaeroides are mediated by the OxyR protein
Analysis of the completed genome of R. sphaeroides 2.4.1. (http://genome.ornl.gov/microbial/rsph) revealed the presence of two genes encoding catalases, RSP2779 and RSP2380. The RSP2779 gene product shows 45 % identity to the E. coli hydroxyperoxidase (HP) II, which is encoded by the E. coli katE gene. The RSP2380 gene product shows 51 % identity to the E. coli HPII protein but even higher identity to the katC gene products of Agrobacterium or Sinorhizobium. Like other KatC proteins it harbours a C-terminal protease domain. The almost completed genome of R. capsulatus SB1003 (data taken from: www.ergo-light.com/ERGO) contains only one gene with significant similarity to a catalase/peroxidase, katG. The katG gene product shows 53 % identity to the E. coli HPI protein, which is encoded by the E. coli katG gene. We investigated the expression of these genes after addition of H2O2 by real-time RT-PCR. The expression of the katE gene of R. sphaeroides as well as expression of the katG gene of R. capsulatus were strongly induced after addition of H2O2 to exponentially growing cultures (Fig. 3a, b). High expression levels were already observed 1 min after the addition of H2O2. These data fit well with the fast kinetics of H2O2 detoxification observed for Rhodobacter cultures. Oxygen itself also exerted an effect on katE and katG gene expression in R. sphaeroides and R. capsulatus, respectively (Fig. 4). The basal expression of both katE and katG were reduced about fivefold under semi-aerobic conditions. No change could be observed for the basal expression of katC when the cells were grown under aerobic or semi-aerobic conditions (Fig. 4).



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Fig. 3. Expression of kat genes in wild-type strains (open bars), oxyR mutant strains (filled bars) and complemented mutant strains (hatched bars). H2O2 was added to exponentially grown cultures to a final concentration of 1 mM, samples were taken at the indicated times. Total RNA was isolated and the level of kat expression was monitored by real-time RT-PCR analysis. (a) Expression of katG in R. capsulatus strains. (b) Expression of katE in R. sphaeroides strains. (c) Expression of katC in R. sphaeroides strains. Values represent the fold change of the kat gene expression compared to that of the control at time-point 0. The data represent the mean of three independent experiments. Error bars indicate maximal deviation.

 


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Fig. 4. Effect of oxygen tension on the expression of katC, katE and katG. Cultures of strains SB1003 (wild-type R. capsulatus) and 2.4.1 (wild-type R. sphaeroides) were grown under aerobic or semi-aerobic conditions to exponential phase. Total RNA was isolated and expression of kat genes (katG in R. capsulatus and katE and katC in R. sphaeroides) was analysed by real-time RT-PCR. Filled bars represent the fold change of the kat expression under semi-aerobic growth compared to the expression under aerobic conditions (=1; open bars). Values are means and maximal deviation of three independent experiments.

 
The known ability of OxyR to positively regulate the expression of catalases in E. coli (Storz & Zheng, 2000) prompted us to determine whether the OxyR protein of Rhodobacter is also involved in the regulation of the H2O2-dependent expression of catalases. The expression of katE in R. sphaeroides and katG in R. capsulatus were both dependent on OxyR (Fig. 3a, b). Without the addition of H2O2 the expression levels in the oxyR mutant strains were similar to that of the parental wild-type strains. The H2O2-inducibility of kat expression was regained by provision of a functional oxyR gene in trans (Fig. 3a, b). These data imply that the oxyR gene product is important for regulation of the genes encoding katE and katG, respectively, in Rhodobacter.

Regulation of katC gene expression
In contrast to the gene encoding katE of R. sphaeroides, the katC gene showed only a weak response when H2O2 was added to exponentially growing cultures of R. sphaeroides (Fig. 3c). The low H2O2-dependent induction of katC expression in R. sphaeroides was independent of OxyR (Fig. 3c). In contrast to the katE and katG basal expression levels, respectively, oxygen tension had no effect on the basal katC expression level (Fig. 4). To determine whether expression of the katC gene is induced in the stationary phase, as it was reported for Sinorhizobium meliloti (Sigaud et al., 1999), expression of the gene was measured by real-time RT-PCR during growth (Fig. 5a, b). An 11- to 38-fold increase in expression of katC was observed when cells reached stationary phase, suggesting a function of the katC gene product in this growth stage. This finding is consistent with the results of Terzenbach & Blaut (1998), who observed a twofold higher catalase activity in R. sphaeroides cells grown in stationary phase. Expression of katE did not increase after cells entered stationary phase (data not shown).



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Fig. 5. Real-time RT-PCR analysis of katC expression during stationary phase. (a) Bacterial growth (expressed as log OD660) was monitored, samples were taken after 0, 8, 12, 16, 24, 29 and 32 h of growth. (b) Total RNA was isolated and the expression of katC was measured by real-time RT-PCR. Values were calculated as fold change of the expression of katC compared to that of the control at time-point 0. The data represent the mean of three independent experiments. Error bars indicate maximal deviation.

 
Reduced and oxidized OxyR bind to the kat promoter region
In E. coli, OxyR exerts its effect on gene expression by binding to promoter regions of target genes (Kullik et al., 1995). Some promoter regions bind only oxidized or reduced OxyR, other promoter regions can bind both forms, but only the oxidized form activates transcription (Kullik et al., 1995). To understand the regulation of kat genes by various forms of OxyR better, the DNA-binding ability of the Rhodobacter OxyR protein was tested. To do this, the oxyR gene of SB1003 was cloned and overexpressed with an N-terminal His6-tag in E. coli. The isolated protein showed specific binding to the katG promoter region, no matter whether it was in its reduced or oxidized form. However, different types of DNA–protein complexes were formed by reduced and oxidized OxyR (Fig. 6a). At high concentrations, OxyR may bind to the katG promoter as an oligomer leading to the C2 band in the gel-shift experiment.



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Fig. 6. Binding of reduced and air-oxidized OxyR to the kat promoter region. To generate reduced conditions, 200 mM DTT was added to the binding reactions. Ox and red indicate oxidized and reduced OxyR, respectively. (a) Binding of OxyR to the katG promoter region. C1ox, complex 1 of DNA and oxidized OxyR; C1red, complex 1 of DNA and reduced OxyR; C2, complex 2. The following amounts of OxyR were added to the reaction: 10 ng (lanes 1 and 5), 20 ng (lanes 2 and 6), 50 ng (lanes 3 and 7) and 100 ng (lanes 4 and 8). (b) Binding of OxyR to the katE promoter region. C1, complex of DNA and oxidized or reduced OxyR; C2, complex 2. The following amounts of OxyR were added to the reaction: 10 ng (lanes 1 and 5), 20 ng (lanes 2 and 6), 50 ng (lanes 3 and 7), 100 ng (lanes 4 and 8). P represents unbound probe, c represents the control without addition of OxyR. (c) DNaseI footprinting assay with oxidized and reduced (in the presence of 200 mM DTT) OxyR binding to the katG promoter region. The regions protected by OxyR are indicated by the brackets. The heavy brackets indicate the reduced extended footprint of OxyR, the light brackets indicate the shorter footprint of oxidized and reduced OxyR. The positions of the footprints are labelled with respect to the katG translational start site. (d) Sequence of the katG and oxyR promoters. The starts of the katG and the oxyR coding sequences are indicated by arrows. Binding sites of oxidized and reduced OxyR are denoted by black bars.

 
To test whether these different types of DNA–protein complexes are due to different sized DNA–protein complexes or rather to different conformation of the complexes, we performed gel filtration assays with purified protein. The purified protein was loaded on a Superdex-200 gel filtration column, and the fractions were analysed by SDS-PAGE. The oxidized protein eluted predominantly in fractions that most likely correspond to a tetramer of the 32·2 kDa OxyR protein (128·8 kDa) (Fig. 7a). We also examined the size of the protein when the column was loaded and eluted in the presence of 200 mM DTT. As shown by AMS modification, 200 mM DTT reduced the OxyR protein completely (Fig. 7c). The elution profile observed under reducing conditions was identical to that in the absence of DTT (Fig. 7a, b). Protein eluted from the column in presence of DTT still resulted in band C1red in the gel shift (not shown), confirming that OxyR was still reduced. These data indicate that oxidized as well as reduced OxyR binds as a tetramer, but that the conformation of the DNA–protein complexes differ.



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Fig. 7. (a) Analytical gel filtration profile of oxidized (thin line) and reduced (heavy line) OxyR protein. The purified OxyR protein was passed through a Superdex-200 HR 16/60 column equilibrated with Z buffer. To generate reducing conditions, 200 mM DTT was added to the Z buffer. The flow rate was 1 ml min–1, and the absorbance of the eluted protein was monitored at 280 nm. (b) The elution fractions were analysed by SDS-PAGE. The elution position of the peak fraction for the standard protein immunoglobulin G (150 kDa) under oxidizing and reducing conditions is indicated. (c) Redox status of OxyR. OxyR protein (125 ng) was reduced or oxidized with 200 mM DTT or 1 mM H2O2. Samples were mixed with 10 % TCA and then treated with AMS. Separation and visualization of OxyR was achieved by SDS-PAGE and silver staining. Control: OxyR without DTT, H2O2 and AMS treatment. The addition of AMS leads to the alkylation of free thiol groups, present in the reduced OxyR. The addition of the high molecular mass AMS moiety to the reduced but not to the oxidized protein allows separation of the two forms by gel electrophoresis.

 
Since the OxyR proteins of both R. capsulatus and R. sphaeroides show a high homology to each other (43 % identity, 56 % similarity), we also tested the binding capacity of the R. capsulatus OxyR protein to the katE promoter region. We observed a binding of both the reduced and oxidized protein to the katE promoter region, but no different DNA–protein complexes were observed (Fig. 6b).

In order to define the binding sites of reduced and oxidized OxyR to the katG promoter region better, we performed footprint analysis. Oxidized OxyR protein bound to AT-rich sequences close to the translational start of oxyR (Fig. 6c, d). When reduced OxyR was applied, the footprint was significantly extended. A long DNA stretch comprising sequences well within the oxyR gene was protected from DNaseI digestion.


   DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
REFERENCES
 
R. sphaeroides and R. capsulatus are related bacteria living in the same habitats. In their natural environment, mostly lakes and ponds, they are exposed to changing light and oxygen conditions. Although the responses of the two species to changes in the environment are similar, some differences can be observed. R. sphaeroides harbours almost no bacteriochlorophyll under high oxygen tension, while R. capsulatus assembles low amounts of photosynthetic complexes. The latter strategy allows a faster adaptation to decreasing oxygen concentrations and a faster switch to anoxygenic photosynthesis when light is present. In this study, we observed a different performance of the two Rhodobacter species under peroxide stress conditions and investigated its molecular basis. Moreover, we could show that environmental stimuli influence the ability of the two Rhodobacter strains to cope with this peroxide stress.

Our results reveal that the two Rhodobacter strains show significant differences in their ability to cope with peroxide stress, which is correlated to strong differences in catalase activities. Both strains showed much faster detoxification of H2O2 under aerobic growth conditions than during growth under low oxygen tension, again correlating to higher catalase activities in both strains when grown under high oxygen tension. The basal expression of genes encoding catalases as determined by real-time RT-PCR was indeed higher under aerobic conditions, in both wild-type strains. Growing at high oxygen tension, bacteria will produce more ROS than during growth at low oxygen tension. Therefore, bacteria adapt to the presence of high oxygen levels by increasing the expression of genes involved in the detoxification of ROS, such as catalases. These higher expression levels result in higher resistance to peroxide stress at high oxygen levels. The assumption that the higher resistance of R. sphaeroides to H2O2 is caused by the presence of two catalases, KatE [encoded by RSP2779 (katE)] and KatC [encoded by RSP2380 (katC)] was not supported by our findings. While the expression of katE was strongly induced by oxygen or peroxide stress, expression of katC remained unaffected (Fig. 3b, c and Fig. 4). The possibility that peroxidases could be responsible for the higher resistance of 2.4.1 can be excluded, since R. sphaeroides does not harbour peroxidase activity (Terzenbach & Blaut, 1998). Hochman & Shemesh (1987), however, observed two different peroxidase activities in R. capsulatus. Thus, the higher resistance of R. sphaeroides to peroxide stress in comparison to R. capsulatus is probably not due to the presence of additional enzymes, but rather to the high activity of KatE. Nevertheless, even the low expression levels of katC might contribute to increase the resistance of R. sphaeroides to H2O2.

A key regulator of the response to H2O2 is the OxyR transcriptional regulator, which induces the expression of antioxidant activities in response to H2O2 stress (Storz & Imlay, 1999). oxyR mutants of both Rhodobacter species were more sensitive to the exposure to H2O2 and showed slower kinetics in the detoxification of this agent compared to the parental strains, confirming a role of OxyR in the oxidative stress response of Rhodobacter. While very little detoxification of H2O2 was observed in strain SBoxyR, the R. sphaeroides mutant 2.4.1oxyR was able to detoxify 3·5 mM H2O2 within 300 s, but with slower kinetics than the isogenic wild-type (Figs 1b and 2c). This indicates that OxyR is more important for defence against H2O2 in R. capsulatus than in R. sphaeroides. Unexpectedly, we found that the oxyR mutation resulted in a decrease of total catalase activity in both Rhodobacter species. This finding is in contrast to observations in E. coli, where the deletion of oxyR did not reduce the basal activity of catalases during exponential growth, but only in cells induced with H2O2 (Visick & Clarke, 1997). The lower levels of catalase activity in strain SBoxyR compared to strain 2.4.1oxyR are in agreement with the very poor detoxification of H2O2 by the R. capsulatus mutant. The addition of H2O2 to semi-aerobically grown wild-type cultures resulted in an induction of catalase activity, whereas no induction was observed in both oxyR mutants. This again is in agreement with findings in other bacterial systems, where OxyR acts as an activator of H2O2-inducible genes. Our observations suggest an effect of OxyR on the catalase activity in unstressed cells as well as an important role of OxyR in the adaptation of Rhodobacter to H2O2.

In many bacteria, genes encoding hydroperoxidases (HP) are members of the OxyR regulon. Gene expression analysis in oxyR mutant strains revealed that both katG of R. capsulatus and katE of R. sphaeroides are regulated by the OxyR protein. In E. coli the katG gene is strongly induced by H2O2, while the katE gene is not (Schellhorn, 1994). A regulatory effect of OxyR on katG expression has been shown for many bacteria (Loprasert et al., 2003; Ochsner et al., 2000). Our observation of fast kinetics in the induction of kat gene expression by H2O2 as well as the reduction 30 min after addition of H2O2 can be explained by the very fast kinetics of activation of OxyR by H2O2. Aslund et al. (1999) were able to examine the kinetics of OxyR oxidation and reduction in vivo and in vitro. OxyR oxidation by H2O2 was completed within 30 s, and the half-time of deactivation was 10–30 min. Compared to katE and katG expression, respectively, expression of katC was not affected by OxyR in R. sphaeroides. This is in agreement with results for the E. coli katE gene (Schellhorn, 1994). Both the E. coli katE gene (Schellhorn, 1994) and the R. sphaeroides katC gene are induced during stationary phase. The expression of the E. coli katE gene is known to be regulated by the stationary phase sigma factor RpoS, while the regulator of the R. sphaeroides katC gene is presently unknown. So far, no homologue of rpoS has been reported for the {alpha}-subclass of proteobacteria, which includes the Rhodobacter species (Rava et al., 1999; Roop et al., 2003).

The oxyR family is widespread among prokaryotes and nearly all known oxyR genes share overlapping promoters with other genes (Kim & Mayfield, 2000). The majority of genes located adjacent to oxyR are involved in oxidative stress protection, such as aphC, dps and oxyS, and are regulated by OxyR (Nakjarung et al., 2003). In both Rhodobacter species the katE and katG genes, respectively, are located adjacent to the oxyR gene on the chromosome. Both genes are separated by approximately 100 nt (katE/oxyR 101 nt; katG/oxyR 98 nt) of untranslated region and are transcribed divergently. Thus, the oxyR and katE or katG genes, respectively, share a common upstream DNA sequence and may also share cis regulatory elements. This kat/oxyR gene organization is found in many {alpha}-proteobacteria [Rhizobium etli (del Carmen Vargas et al., 2003); Brucella abortus (Kim & Mayfield, 2000); Agrobacterium tumefaciens (Nakjarung et al., 2003)], suggesting a general mechanism of regulation of these genes in {alpha}-proteobacteria.

Since it is reported that the OxyR protein regulates expression of genes by direct binding to the promoter region (reviewed by Schell, 1993), we tested the ability to bind DNA of the R. capsulatus OxyR protein. The OxyR protein showed strong binding to the katG promoter region and our data indicate that different conformations of DNA–protein complexes are formed with oxidized or reduced OxyR. Footprint analysis of the E. coli OxyR protein showed that OxyR binding is different under oxidizing and reducing conditions (Kullik et al., 1995). As described for E. coli (Kullik et al., 1995), the oxidized and reduced forms of R. capsulatus OxyR were predominantly tetrameric as revealed by size exclusion chromatography. As a member of the LysR family of bacterial regulators, OxyR acts as an activator of a regulon of genes and a repressor of its own expression (Storz & Altuvia, 1994). Toledano et al. (1994), looking in E. coli, found that only oxidized OxyR binds katG, ahpC, dps and gorA promoters, whereas both the oxidized and the reduced protein bind the oxyRS promoter. They proposed that, by remodelling its DNA contacts, OxyR can impose opposite regulatory effects on the divergent oxyR and oxyS promoters.

Our results show that oxidized as well as reduced OxyR binds to the katG/oxyR promoter region, but the reduced protein makes extended contacts to the DNA compared to the oxidized protein. Our data also show that katG expression is strongly increased by H2O2, and that H2O2 leads to oxidation of the OxyR protein (Fig. 7c). Without the addition of H2O2, the level of katG expression is similar in the wild-type and the oxyR mutant. Thus OxyR acts as an activator of katG expression in the presence of H2O2. We propose that this activation includes the release of OxyR from binding sites within the oxyR gene. Our findings also suggest that reduced OxyR has little repressing effect on katG expression, which is low in the absence of oxidized OxyR.

Future work will be aimed at the identification of additional OxyR-regulated genes and additional OxyR-binding sites to understand better the OxyR signalling mechanism in Rhodobacter.


   ACKNOWLEDGEMENTS
 
This work was supported by the ‘Deutsche Forschungsgemeinschaft’ (Kl563/16-1), the ‘Fonds der Chemischen Industrie’ and the BMBF.


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RESULTS
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Received 6 May 2004; revised 22 July 2004; accepted 25 July 2004.



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