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PHYSIOLOGY AND BIOCHEMISTRY

Molecular characterization of FinR, a novel redox-sensing transcriptional regulator in Pseudomonas putida KT2440

Sujin Yeom, Jinki Yeom, Woojun Park

  • Division of Environmental Science and Ecological Engineering, Korea University, Anam-Dong 5 Ga 136-713, Seoul, Republic of Korea
  • Correspondence
    Woojun Park
    wpark{at}korea.ac.kr

    • Microbiology 2010; 156(5):1487–1496 · https://doi.org/10.1099/mic.0.034181-0

    View at publisher PubMed

    Abstract

    FinR is required for the induction of fpr (ferredoxin-NADP+ reductase) under superoxide stress conditions in Pseudomonas putida. Many proteobacteria harbour FinR homologues in their genome as a putative LysR-type protein. Three cysteine residues (at positions 150, 239 and 289 in P. putida FinR) are conserved in all FinR homologues. When these conserved cysteines, along with two other cysteine residues present in FinR, were individually mutated to serines, the FinR remained active, unlike SoxR and OxyR in Escherichia coli. The results of our in vitro DNA-binding assay with cellular extracts showed that FinR binds directly to the fpr promoter region. In order to identify the FinR functional domain for sensing superoxide stress, we employed random and site-directed mutagenesis of FinR. Among 18 single amino acid mutants, three mutants (T39A, R194A and E225A) abolished fpr induction without any alteration of their DNA-binding ability, whereas other mutants also abrogated their DNA-binding abilities. Interestingly, two mutants (L215P and D51A) appeared to be constitutively active, regardless of superoxide stress conditions. Ferrous iron depletion, ferric iron addition and fdxA (ferredoxin) gene deletion also participate in the regulation of fpr. These data indicate that FinR has unusual residues for redox sensing and that the redox-sensing mechanism of FinR differs from the well-known mechanisms of OxyR and SoxR.

    • ↵†These authors contributed equally to this work.

    • Two supplementary tables and one supplementary figure are available with the online version of this paper.

    Edited by: J. W. B. Moir

    INTRODUCTION

    Reactive byproducts of oxygen are generated continuously within cells during their growth under aerobic conditions (Cabiscol et al., 2000). These byproducts, referred to as reactive oxygen species (ROS), are toxic to cells owing to their ability to damage many macromolecules inside cells, including DNA, lipids and proteins. When certain proteins are exposed to subinhibitory concentrations of ROS, either modification of amino acid side chains or destabilization of cofactors in the regulatory proteins occurs, which leads to alterations in the protein structures (Imlay, 2003; Stadtman, 1992). Cells employ these structural alterations as signals for the activation of their defence systems. In Escherichia coli cells, the SoxR protein, a MerR-type transcriptional factor, activates soxS gene expression in response to superoxide or nitric oxide stresses. SoxS is the global transcriptional regulator that controls a variety of oxidative stress defence proteins, including Zwf (glucose-6-phosphate dehydrogenase) and Fpr (ferredoxin-NADP+ reductase); thus, the products of the induced soxRS regulon function to prevent or repair oxidative damage. SoxR harbours a redox-active (2Fe–2S) centre, the redox status of which can govern the transcriptional activity of SoxR (Demple, 1996; Hidalgo & Demple, 1994; Pomposiello & Demple, 2000, 2001).

    The oxidative stress system has been studied extensively in E. coli and Salmonella typhimurium (Farr & Kogoma, 1991). The genome of Pseudomonas putida KT2440, a model soil micro-organism, has been sequenced (Nelson et al., 2002) but its oxidative stress systems are currently poorly understood. Previously, we reported that the SoxRS regulon does not exist in the genome of P. putida KT2440 and that SoxR does not confer oxidative stress defence (Park et al., 2006). The principal function of SoxR in many other micro-organisms is the control of secondary metabolite production, rather than directing oxidative stress defence (Dietrich et al., 2008). Interestingly, we have identified two important possible redox sensors – FinR and HexR – that may control the expression of the fpr gene and the zwf-1 gene, respectively (Kim et al., 2008; Lee et al., 2006b). These two proteins are critical for the maintenance of proper NADPH levels inside cells. Both of these proteins are under the control of the SoxRS regulon in E. coli (Greenberg et al., 1990; Liochev et al., 1994).

    The OxyR protein, a LysR-type transcriptional activator, is another type of redox sensor, which has been shown to be relevant to cellular defence against hydrogen peroxide stresses in E. coli and S. typhimurium (Christman et al., 1989; Tao et al., 1989). OxyR activation occurring in response to H2O2 is dependent on two cysteine residues (Cys199 and Cys208) (Zheng et al., 1998). OxyR in P. putida controls katA, katB and ahpC, which encode three major peroxide-degrading enzymes (Hishinuma et al., 2006). Thioredoxin reductase (trxB) is also regulated by OxyR, and further study will be required to elucidate the mechanisms underlying this OxyR regulation clearly (Hishinuma et al., 2008). The LysR-type transcriptional regulators are a large family of regulatory proteins in the proteobacteria, and are known to regulate a variety of genes and gene functions. They feature an N-terminal DNA-binding motif of ∼75 aa, and a C-terminal inducer binding domain of ∼225 aa (Schell, 1993). The thiol group in the cysteine residues of many transcriptional factors is the key residue associated with redox-sensing, which has also been fairly well characterized in OxyR (Green & Paget, 2004).

    The FinR locus is located alongside the fpr. Fpr mediates a reversible redox reaction between NADP+/NADPH and electron carriers (Carrillo & Ceccarelli, 2003). We have demonstrated that FinR is essential for the upregulation of fpr in response to superoxide stress in P. putida KT2440 (Lee et al., 2006b). Further relevant biochemical data have yet to be collected. FinR has been implicated previously as a LysR family transcription regulator. According to alignments of FinR homologues, three cysteine residues are highly conserved in many proteobacteria. These results compelled us to investigate the activation mechanisms of FinR in response to superoxide stress by using a gel shift assay and reporter fusion using a variety of mutants. In order to obtain insight into possible residues required for activation, we conducted random and site-directed mutagenesis of FinR, and screened for mutants that were incapable of activating transcription.

    METHODS

    Bacterial strains, culture conditions and DNA manipulation.

    The bacterial strains and plasmids utilized in this study are listed in Supplementary Table S1 (available with the online version of this paper). Bacteria were grown at 37 °C (E. coli) or 30 °C (P. putida) in LB medium with vigorous aeration. When required, antibiotics were added at the following concentrations: 200 μg rifampicin ml−1; 100 μg ampicillin ml−1; 50 μg kanamycin ml−1; and 15 μg tetracycline ml−1.

    Cloning procedures and mutant construction.

    The primers utilized in this study are listed in Supplementary Table S2. In order to delete fdxA, a 345 bp fragment of its internal region was amplified via PCR using the FdxA-F and FdxA-R primers. The fragment was cloned into the EcoRI and KpnI cloning sites of the pVIK112 vector (Kalogeraki & Winans, 1997), yielding pVIK-fdxA. This plasmid was then introduced into E. coli S17-1 λ pir. Conjugation was conducted with E. coli S17-1 λ pir (pVIK-fdxA) and P. putida KT2440-R as the donor and recipient, respectively. Then, the pRKPfpr gfp plasmid was introduced via triparental conjugation into ΔfdxA, thereby generating P. putida KT2440 (ΔfdxA/pRKPfpr gfp) (Lee et al., 2006a). The 1096 bp fragment of the finR full region was amplified by PCR using the primers FinR-CP1 and FinR-CP2. The fragment was then cloned into the EcoRI and HindIII cloning sites of the pUC19 vector (Yanisch-Perron et al., 1985), thereby generating pUC19-finR. The constructed plasmid was then introduced into E. coli Top10 and E. coli Top10 (pRKPfpr gfp).

    Mutagenesis.

    The pUC19-finR plasmid was utilized as a PCR template under conditions designed to induce frequent nucleotide misincorporation. The PCR conditions were identical to those described previously, with some modifications (Cadwell & Joyce, 1992). The mutagenic reaction mixtures contained 7 mM MgCI2, 1 U Taq polymerase, and varying concentrations of the four dNTPs (0.2 mM dATP, dGTP and 1.0 mM dCTP, dTTP). The reaction was conducted in a thermal cycler (Eppendorf) for 35 cycles of 94 °C for 45 s denaturation, 60 °C for 45 s annealing, and 72 °C for 45 s extension. Mutated transformants were screened to measure green fluorescent protein (GFP) intensity. Site-directed mutants were constructed via a three step–four primer overlap/extension PCR technique, using Platinum Pfx (Invitrogen) (Ho et al., 1989). The first step was prepared via PCR amplification with the primer pairs FinR-CP1/SDM primer-1 and FinR-CP2/SDM primer-2. Each of the PCR products was annealed at the region overlap and extended to form full-length double-strand mutant DNA. The full-length mutant DNA was amplified using primers FinR-CP1/FinR-CP2.

    Microscopic analysis and quantification of GFP fluorescence.

    Cells were viewed with a Carl Ziess Axio Imager microscope and GFP fluorescence was quantified as described previously (Kim et al., 2007). One fluorescence unit was defined as (fluorescence intensity of cells−fluorescence intensity of PBS buffer)/(OD600 of cells).

    Northern blot analysis.

    Northern blot analysis was conducted as described previously (Kim et al., 2008). Total RNA was isolated from 2 ml exponentially growing cells. Samples of total RNA (10 μg) were loaded. Probe was prepared via PCR amplification from P. putida strain KT2440 with the fpr Pp-hA/fpr Pp-hS primer pair.

    Electrophoretic mobility shift assay (EMSA).

    The EMSA was conducted as described previously (Kim et al., 2008). Exponentially growing cells in LB containing ampicillin were harvested. The Pfpr DNA probe was generated using the fpr pro2/fpr pro3 primer pair. The reaction mixture (15 μl final volume), containing the Pfpr probe, crude extract, loading buffer and dI-dC (0.1 μg) in binding buffer [10 mM Tris, pH 7.5, 2 mM MgCl2, 10 % (v/v) glycerol and 75 mM KCl], was incubated for 20 min at room temperature. The resulting complexes were analysed by electrophoresis on 6 % polyacrylamide gels.

    Oxidative stress sensitivity assay.

    Cells were grown overnight in liquid LB medium and subsequently diluted 100-fold in the same medium. After further incubation until the cells reached exponential phase (OD600∼0.5), serially diluted cells were spotted on LB agar with or without paraquat (PQ) (5 μM).

    Nitro blue tetrazolium (NBT) assay.

    The NBT test was conducted as previously described, with some modifications (Becerra et al., 2003). Bacterial cells in the exponential growth phase (OD600∼0.5) grown in LB medium were treated with 0.5 mM PQ. After 1 h exposure to PQ, bacterial suspensions (1 ml) were incubated with 0.5 ml 1 mg NBT ml−1 for 30 min at 30 °C. Then 0.1 ml 0.1 M HCl was added, and the tubes were centrifuged for 10 min at 1500 g. The separated pellets were treated with 0.4 ml DMSO to extract the reduced NBT. Finally, 0.8 ml PBS (pH 7.5) was added, and the growth was measured at 575 nm. Total protein concentrations were measured via Bradford assays. One unit was defined as OD575 (mg protein)−1.

    Homology modelling for FinR structure.

    A homology model of FinR was generated as described previously (Yeom et al., 2009a). The FinR model used the CrgA of Neisseria meningitides (Sainsbury et al., 2009).

    Protein purification.

    FdxA was purified as described previously (Yeom et al., 2009b). All purification steps were performed at 4 °C, using an FPLC system (AKTA FPLC, Unicorn 4.0, Amersham Bioscience). To purify FdxA, E. coli cell pellets containing pET-fdxA were resuspended in buffer A (50 mM Tris/HCl and 1 mM DTT, pH 7.5) and disrupted by sonication. After removal of cell debris by centrifugation at 12 000 g for 30 min, the soluble fraction was loaded onto an anion exchange column (1 ml, DEAE-cellulose, Amersham Bioscience) equilibrated with buffer A, and the proteins were eluted with a 20 ml linear gradient of 0–1 M NaCl in Buffer A (pH 7.5). The fractions (0.5 ml each) were collected and concentrated by ultrafiltration with Centricon (2 ml YM-10, Amicon). The concentrates were applied to a Ni-NTA column (1 ml, His-trap, Amersham Bioscience) equilibrated with binding buffer (20 mM sodium phosphate, 0.5 M NaCl, 40 mM imidazole, pH 7.4) and the proteins were eluted with 15 ml elution buffer (20 mM sodium phosphate, 0.5 M NaCl, 250 mM imidazole, pH 7.4). The fractions were dialysed by ultrafiltration with Centricon (2 ml YM-10, Amicon) and stored at −80 °C in 10 % glycerol. After Ni-NTA purification according to the above methods, the fractions of FdxA were applied to a gel-permeation column (Sephacryl S-200 HR 26/60, Pharmacia) and equilibrated with 50 mM Tris/HCl buffer (pH 7.5) containing 150 mM NaCl and 1 mM DTT. The proteins were eluted from the column with the same buffer at a flow rate of 30 ml h−1. SDS-PAGE was carried out using 15 % polyacrylamide gels to check the level of expression and purification.

    RESULTS

    Amino acid sequence analyses of FinR homologues

    FinR is annotated as a putative LysR-type transcriptional factor and consists of 308 aa; its gene is positioned divergently adjacent to fprA in the P. putida KT2440 genome. Many bacteria have one FinR homologue in their genomes, and its function has not been studied at all except in our two previous studies, which showed that FinR is required for the induction of the fpr gene in response to superoxide and osmotic stresses (Park et al., 2006); however, the mechanism underlying its regulation has yet to be established in detail. Among the well-characterized proteins, the most closely related homologue is CysB, a LysR-type transcriptional regulator which controls the expression of a variety of genes required for sulfur assimilation (Henikoff et al., 1988; Ostrowski & Kredich, 1989). The sequence identity of CysB with FinR is only 23 %. When 30 FinR homologues from the GenBank database were aligned, it appeared that several regions were highly conserved: the N-terminal and central domain regions are more conserved than the C-terminal regions (Fig. 1⇓). FinR from P. putida has 81.5 and 64 % amino acid identity with its homologues Azotobacter vinelandii and Aromatoleum aromaticum EbN1, respectively. The phylogenetic tree constructed via the alignment of the amino acid sequences of FinR homologues from various bacterial lineages demonstrated that the majority of FinR homologues could be detected in the gammaproteobacteria (Supplementary Fig. S1a, available with the online version of this paper). Because the phylogenetic distance from FinR is closer, the finR loci were observed to be similar to P. putida KT2440. The widespread distribution of the FinR homologues among the proteobacteria, with few exceptions, is evidence of their later evolutionary appearance. Many redox sensors harbour conserved cysteine residues to control their regulation (Green & Paget, 2004). Three cysteine residues (at positions 150, 239 and 289 in the P. putida protein) are extremely well conserved, whereas two other cysteines are not conserved (Fig. 1⇓; Supplementary Fig. S1b). We hypothesized that those three cysteine residues might prove important for redox sensing in FinR homologues.

    Figure image not available in archive
    Fig. 1.

    Alignment of FinR homologues from different organisms. Amino acid sequences (obtained from NCBI) were aligned via clustal w. FinR, P. putida KT2440 (AE015451.1); P. ae, Pseudomonas aeruginosa PAO1 (AE004091.2); A. vi, A. vinelandii DJ (CP001157.1); C. sa, Chromohalobacter salexigens DSM 3043 (CP000285.1); M. fl, Methylobacillus flagellatus KT (CP000284.1); A. ar, Aromatoleum aromaticum EbN1 (CR555306.1); C. ja, Cellvibrio japonicus Ueda107 (CP000934.1). Highly conserved residues in seven FinR homologues (black shading), conserved residues in several FinR homologues (grey shading), conserved cysteine residues in all FinR homologues (asterisks) and the rest of the cysteine residues present in FinR (double dots) are highlighted.

    FinR senses superoxide stress and ferrous iron depletion

    In order to determine whether FinR actually senses superoxide stress, we quantified the fprA promoter activity via transcriptional fusion with GFP, as described in our previous studies (Lee et al., 2006a). We verified that the expression of the fprA promoter is induced by superoxide-generating agents (Fig. 3a⇓). This notion is further supported by the fact that overexpressed SodA from the ectopic plasmid effectively eliminates superoxide, and thereby reduces fprA expression under paraquat treatment conditions (Fig. 2a⇓). These data clearly demonstrated that FinR is activated as the result of superoxide stress.

    Figure image not available in archive
    Fig. 2.

    Analysis of fpr expression depending on superoxide stress and iron depletion. (a) GFP intensity was measured after PQ shock. Quantification of GFP in wild-type [P. putida KT2440 (pRKPfpr gfp)] and SodA [P. putida KT2440 (pRKPfpr gfp/pBBR sodA)]. Samples were treated (black bars) or not (white bars) with 1 mM PQ. Error bars indicate sd of data from three independent experiments. (b) Northern blot analysis of fpr transcription. The mRNA induction level of fpr genes in minimal medium (M9) with 0.5 μM ferric-citrate is shown. All chemicals were applied to the cells for 30 min at 10 μM. C, no treatment; Fe-C, ferric-citrate; Fe-E, ferric-EDTA; and Di, 2′,2′-dipyridyl.

    Recently, we have used Western blot analysis to demonstrate that the FprA protein is also profoundly upregulated under iron stress conditions (Yeom et al., 2009a). In order to confirm the relationship between FinR and iron in the regulation of fprA, we measured the fprA expression levels of both the wild-type and finR mutant (Fig. 2b⇑). In the finR mutant, fprA expression is dramatically reduced under both ferric iron-repleted and -depleted conditions, whereas the wild-type induces fprA expression in response to those iron-stress conditions (Fig. 2b⇑). We concluded that the fprA expression under iron stress conditions might be associated with FinR.

    Expression of FinR–fprA system in a heterologous host, E. coli

    It is desirable to use E. coli to critically assess the residues relevant to FinR activation, owing to the ease with which transformations and gene manipulations can be conducted. The fprA–gfp reporter vector coupled with the plasmid in which FinR is constitutively expressed when introduced into E. coli Top10 (data not shown). When exposed to PQ, Top10 cells harbouring the two plasmids showed responses similar to those of P. putida KT2440 (Fig. 3a⇓). The GFP level in the reporter strains accumulated in cases in which cells were exposed to different PQ concentrations. The greater the concentration of PQ applied, the more fprA expression was induced. GFP expression was also visible by using fluorescence microscopy. Without PQ treatment, the reporter strains did not induce GFP expression in LB media (Fig. 3b⇓), whereas high levels of GFP expression were noted in LB media containing 1 mM PQ (Fig. 3b⇓). These data verified that E. coli harbours all the necessary systems to induce the fpr gene in response to superoxide stress.

    Figure image not available in archive
    Fig. 3.

    Expression of P. putida FinR–fpr system in E. coli. (a) GFP intensity was measured after PQ shock in E. coli Top10 (pRKPfpr gfp/ FinR). Bars indicate sd of data from three independent experiments. (b) Phase contrast (i) and fluorescence (ii) microscopy of Top10 (pRKPfpr gfp/ FinR) grown LB media, and phase contrast (iii) and fluorescence (iv) microscopy of the same cells exposed to 1 mM PQ. (c) In vitro EMSA. Lanes: N, retardation assay of the Pfpr probe with no protein; C, Pfpr probe with cell extract from Top10 harbouring empty vector; P, Pfpr probe with cell extract from Top10 harbouring FinR expression vector. A 4 % polyacrylamide gel was used.

    Binding of FinR to the promoter region of the fpr gene

    The binding activity of FinR to the fprA promoter region has never before been studied. As FinR has been shown to be insoluble in overexpression systems, we utilized cell lysates in further binding studies. If the cell extracts from E. coli Top10 harboured an empty vector, the promoter probes were not complexed with any proteins (Fig. 3c⇑) thereby confirming that there are no other proteins that can bind to the fprA promoter regions in E. coli Top10 strains. The EMSA results showed that FinR binds to the fprA promoter region (Fig. 3c⇑).

    Site-directed mutagenesis of conserved cysteine residues

    Cysteines have been shown to be very important residues in many redox-controlling transcriptional factors. They can be oxidized to several different reversible redox states, including disulfide (R-S-S-R′), sulfenic acid (R-SOH) and S-nitrosothiol (R-SNO) (Green & Paget, 2004). In order to determine whether the conserved cysteine residues in FinR are relevant to sensing oxidative stress, we mutated each of the three conserved cysteines to serines; the remaining two cysteines were also mutated via site-directed mutagenesis. The transcription activities of those strains containing mutated FinR were then assayed. It appeared that the C46S, C150S, C169S, C239S and C289S mutant strains exhibited transcription activities similar to those of the strain harbouring wild-type FinR (Fig. 4a⇓). Unlike other redox transcriptional factors, those five cysteines are not involved in FinR activation. It was noted that FinR exhibits a redox-sensing mechanism that differs from OxyR (Zheng et al., 1998).

    Figure image not available in archive
    Fig. 4.

    Characterization of amino acid substitutions. (a) Quantification of GFP expression in E. coli Top10 harbouring wild-type finR and single amino acid mutant finR. PQ (1 mM) was added to cells exponentially grown in LB media. Asterisks indicate finR mutants with impaired redox sensing. Error bars indicate sd of data from three independent experiments. (b) In vitro EMSA. Lanes N–C are the same as in Fig. 3(c). The marked numbers indicate each substituted residue in FinR.

    Characterization of single amino acid mutants

    In order to identify the amino acid residues necessary for FinR activation and DNA binding, we conducted random and site-directed mutagenesis of FinR. Random mutagenesis was accomplished via error-prone PCR, and site-directed mutagenesis was achieved via a three step–four primer overlap/extension PCR technique. No fprA induction was detected in three mutants (T39A, R194A and E225A; Fig. 4a⇑), although their DNA-binding ability to the target gene remained intact (Fig. 4b⇑). Two mutants were constitutively active (L215P and D51A) regardless of oxidative stress conditions (Fig. 4a⇑). The basal level of fprA expression in the L215P mutant was found to be similar to that observed in the wild-type strain following PQ treatment. The transcriptional level of fprA in the L215P mutant increased further upon the addition of PQ. However, the D51A mutant exhibited threefold higher induction levels in both the absence and presence of PQ compared with the wild-type strain. Because five mutants (S35A, K55A, V116E, F204S and L264P) lost their DNA-binding abilities, no fprA induction was detected in these strains (Fig. 4b⇑). Two mutants (L5F, L62P) are partially defective in their DNA binding, but their transcriptional activities remain functional (Fig. 4⇑). Transcriptional activities and DNA-binding abilities of all other mutants were similar to those of wild-type in the absence and presence of PQ (Fig. 4⇑). Interestingly, the D142A mutant showed a greater degree of retardation than other proteins (Fig. 4b⇑), although its transcriptional activity remained the same as the wild-type. This mutant might be altered in its multimerization.

    Involvement of iron–sulfur cofactor-containing FdxA in fprA expression

    Previously, it has been demonstrated that the disruption of FdI, encoding A. vinelandii ferredoxin I (AvFdI), a homologue of FdxA, results in the overexpression of ferredoxin-NADP+ reductase. In the presence of PQ, the fprA activation level in the wild-type strain was identical to that observed in the fdxA mutant. The expression level of fprA was not increased in the fdxA mutant, even after the addition of PQ to the growth medium (Morgan et al., 1988). As these data may shed some light on the issue of fprA regulation, we constructed an fdxA mutant of P. putida KT2440 in an effort to evaluate the effects of fdxA mutation. The expression levels of fprA in both the wild-type and mutant strains were measured in the presence and absence of PQ. The promoter activity of fprA in the mutant was very high, even under normal conditions, compared with the wild-type strain, but fprA expression increased even further upon the addition of PQ (Fig. 5a⇓). The transcriptional fusion data were confirmed via Northern blot analysis (Fig. 5b⇓). We speculated that the intracellular superoxide concentration was increased in the mutant, and that the expression of fprA increased in a dose-dependent manner. Superoxide anion production in the fdxA mutant was measured as described previously (Becerra et al., 2003). Superoxide production in the mutant increased more profoundly under normal growth conditions than in the wild-type strain (Fig. 5c⇓). The pattern of fprA expression in the fdxA mutant correlated with the superoxide anion concentration. Collectively, our results show that FdxA participates indirectly (or directly) in fprA regulation via the production of superoxide inside cells. The results further indicate that fprA regulation is a more complicated issue than had been thought previously. Exponentially grown cells were serially diluted and spotted on LB agar, with and without PQ. The fdxA mutant proved more sensitive than the wild-type and FinR mutant strains (Fig. 5d⇓). To investigate whether FdxA binds to the fprA promoter region, EMSAs were conducted using purified FdxA proteins. Surprisingly, the EMSA results demonstrated that FdxA retards the migration of the DNA probe (Fig. 5e⇓). The explanation for this observation is not straightforward. To examine the relationship between FinR and FdxA on the fprA promoter region, we carried out retardation study using FinR crude extract and FdxA protein. This showed that binding of FdxA to the fprA promoter region is blocked by FinR (Fig. 5f⇓).

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    Fig. 5.

    Characterization of fdxA. (a) Quantification of GFP expression in wild-type [P. putida wild-type (pRKPfpr gfp)] and ΔfdxA [P. putida (ΔfdxA/pRKPfpr gfp)] cells. Samples were treated (black bars) or not (white bars) with 1 mM PQ. (b) Northern blot analysis of fpr transcription. Total RNA was extracted 1, 5, 10, 30 and 60 min after treatment of cells with 1 mM PQ; 0 min is the normal condition. EtBr, Ethidium bromide. (c) Concentration of intracellular superoxide anion. Samples were treated (black bars) or not (white bars) with 0.5 mM PQ. Error bars indicate sd of data from three independent experiments. (d) Sensitivity assay of P. putida wild-type (WT), finR and fdxA mutant strains to PQ. (e, f) In vitro EMSA. Lanes N and C are the same as in Fig. 3(c). FinR, Pfpr probe with cell extract from E. coli Top10 harbouring the FinR overexpression vector; FdxA, purified ferredoxin protein. The amount of protein (μg) is given below the figures.

    DISCUSSION

    Interestingly, data regarding the fpr regulation system of A. vinelandii suggested the involvement of two proteins: ferredoxin I (FdI, a homologue of FdxA, hereafter referred to as FdxA) and the E1 component of pyruvate dehydrogenase complex (PDHE1). It has been demonstrated that PDHE1 binds to a very specific DNA sequence upstream of the fpr gene, and that FdxA binds specifically to DNA-bound PDHE1 (Regnström et al., 1999). Disruption of FdxA transcriptionally activates the fpr gene to the same extent in response to either the presence or the absence of PQ. A recent report has shown that PDHE1 is not the fpr-promoter-binding protein in vivo and theorizing that the effect of FdxA on fpr expression might occur in an indirect manner (Park et al., 2007). Deletion of fdxA may result in a sizeable superoxide burden on cells that leads to the induction of the fpr gene (Park et al., 2007). This assumption was strongly supported by our finding that the deletion of FdxA increased superoxide content within the cells (Fig. 5c⇑). The data presented here demonstrated that FdxA binds to the promoter regions of the fprA gene (Fig. 5e⇑). Because the molecular mass of Fdx is very small and FdxA is not known to be a DNA-binding protein, it seems likely that FdxA assists in the binding of other DNA-binding protein to the fprA promoter. We cannot rule out the possibility that purified FdxA solution was contaminated by PDEH1 during purification steps. Interestingly, FdxA binding is inhibited by the presence of FinR (Fig. 5f⇑). Involvement of PDHE1 in FinR activation remains to be assessed.

    Among several bacterial redox sensors, OxyR and OhrR utilize mechanisms involving cysteine residue modification. Key cysteine residues in both proteins are oxidized in response to oxidative stress conditions. Bacillus subtilis OhrR is a member of the MarR family transcription regulators (Fuangthong et al., 2001). The amino acid sequence of FinR contains five cysteines. Although all FinR homologues harbour conserved cysteines, they do not appear to be thiol-based sensors (Fig. 4a⇑). In this study, we have demonstrated for the first time that FinR is a specific DNA-binding protein (Fig. 3c⇑). We employed random and site-directed mutagenesis to define the regions of FinR relevant to DNA binding and activation. A model of the structure of FinR was established using the structure of CrgA (Fig. 6⇓). CrgA is a member of the LysR-type transcriptional regulatory family, and amino acid similarity between FinR and CrgA is very low (19.8 %). Nevertheless, CrgA is the closest to FinR in terms of protein structure. The CrgA subunit of the full-length protein comprises an N-terminal DNA-binding domain (residues 1–60), connected via a long linker helix (residues 61–89) to a C-terminal regulatory domain. Three mutants (T39A, R194A and E225A) showed low levels of fprA induction, but their DNA-binding ability remained unaffected (Fig. 4b⇑). It is worth commenting that two sensing-defective mutants (R194A and E225A) are mapped to the sensing domain (Fig. 6⇓). They might be mapped at or near possible redox-active residues. The mutants L215P and D51A significantly activated fpr, even without PQ (Fig. 4a⇑). The D51A mutant is mapped to the helix-turn region in the binding domain (Fig. 6⇓). Based on T39A and D51A, we speculated that changes in the DNA-binding domain could influence the transcriptional activity of FinR. The remaining mutation, L215P, mapped at the sensing domain, might be involved in interactions with RNA polymerase or reductase. This constitutively active FinR might be elucidated further by other similar studies. Constitutively active SoxR could be obtained following the deletion of nine C-terminal amino acids, which might interfere with its interaction with a SoxR reductase (Nunoshiba & Demple, 1994). Among an additional seven mutants, four mutants (L5F, S35A, K55A and L62P) have been mapped to the DNA-binding region consistent with the EMSA results (Figs 4⇑ and 6⇓). The remaining three mutants (V116E, F204S and L264P) are also defective in their DNA binding (Fig. 4b⇑), although they are not mapped to the DNA-binding region. This may be because these mutants have an unstable protein structure.

    Figure image not available in archive
    Fig. 6.

    Modelling the structure of FinR in P. putida. Positions of single amino acid mutations on the FinR structure are shown. The residues found to be important for redox sensing (T39, R194 and E225) are shown in red. Constitutive mutants (D51 and L215) are shown in black. Mutants that result in defective DNA binding are shown in blue.

    The OxyR of Deinococcus radiodurans, which harbours one conserved cysteine, functions not only as a positive regulator but also as a negative regulator of different classes of genes (Chen et al., 2008). OxyR regulation in other bacteria is more complicated than had previously been anticipated. In a similar fashion, fprA gene regulation might be involved intricately with other factors, and FinR appears to utilize a unique redox-sensing mechanism which differs profoundly from other redox-sensing transcriptional regulators.

    In summary, our findings demonstrate that FinR of P. putida KT2440 performs a crucial role in cellular defences against superoxide and ferrous iron depletion stresses. Additionally, FinR can bind directly to the promoter region of fprA. Ferredoxin may also have some influence on the expression of fprA.

    Acknowledgments

    This work was supported by a grant from the MEST/NRF to the Environment Biotechnology National Core Research Center (grant no. 20090091491), Korea, and a Korea Science and Engineering Foundation grant (R01-2008-000-10697-0) program, South Korea.

    References

    1. Becerra, M. C., Eraso, A. J. & Albesa, I. (2003). Comparison of oxidative stress induced by ciprofloxacin and pyoverdin in bacteria and in leukocytes to evaluate toxicity. Luminescence 18, 334–340.
    2. Cabiscol, E., Tamarit, J. & Ros, J. (2000). Oxidative stress in bacteria and protein damage by reactive oxygen species. Int Microbiol 3, 3–8.
    3. Cadwell, R. C. & Joyce, G. F. (1992). Randomization of genes by PCR mutagenesis. PCR Methods Appl 2, 28–33.
    4. Carrillo, N. & Ceccarelli, E. A. (2003). Open questions in ferredoxin-NADP+ reductase catalytic mechanism. Eur J Biochem 270, 1900–1915.
    5. Chen, H., Xu, G., Zhao, Y., Tian, B., Lu, H., Yu, X., Xu, Z., Ying, N., Hu, S. & Hua, Y. (2008). A novel OxyR sensor and regulator of hydrogen peroxide stress with one cysteine residue in Deinococcus radiodurans. PLoS One 3, e1602
    6. Christman, M. F., Storz, G. & Ames, B. N. (1989). OxyR, a positive regulator of hydrogen peroxide inducible genes in Escherichia coli and Salmonella typhimurium, is homologous to a family of bacterial regulatory proteins. Proc Natl Acad Sci U S A 86, 3484–3488.
    7. Demple, B. (1996). Redox signaling and gene control in the Escherichia coli soxRS oxidative stress regulon. Gene 179, 53–57.
    8. Dietrich, L. E., Teal, T. K., Price-Whelan, A. & Newman, D. K. (2008). Redox-active antibiotics control gene expression and community behavior in divergent bacteria. Science 321, 1203–1206.
    9. Farr, S. B. & Kogoma, T. (1991). Oxidative stress responses in Escherichia coli and Salmonella typhimurium. Microbiol Rev 55, 561–585.
    10. Fuangthong, M., Atichartpongkul, S., Mongkolsuk, S. & Helmann, J. D. (2001). OhrR is a repressor of ohrA, a key organic hydroperoxide resistance determinant in Bacillus subtilis. J Bacteriol 183, 4134–4141.
    11. Green, J. & Paget, M. S. (2004). Bacterial redox sensors. Nat Rev Microbiol 2, 954–966.
    12. Greenberg, J. T., Monach, P., Chou, J. H., Josephy, P. D. & Demple, B. (1990). Positive control of a global antioxidant defense regulon activated by superoxide-generating agents in Escherichia coli. Proc Natl Acad Sci U S A 87, 6181–6185.
    13. Henikoff, S., Haughn, G. W., Calvo, J. M. & Wallace, J. C. (1988). A large family of bacterial activator proteins. Proc Natl Acad Sci U S A 85, 6602–6606.
    14. Hidalgo, E. & Demple, B. (1994). An iron sulfur center essential for transcriptional activation by the redox-sensing SoxR protein. EMBO J 13, 138–146.
    15. Hishinuma, S., Yuki, M., Fujimura, M. & Fukumori, F. (2006). OxyR regulated the expression of two major catalases, KatA and KatB, along with peroxiredoxin, AhpC in Pseudomonas putida. Environ Microbiol 8, 2115–2124.
    16. Hishinuma, S., Ohtsu, I., Fujimura, M. & Fukumori, F. (2008). OxyR is involved in the expression of thioredoxin reductase TrxB in Pseudomonas putida. FEMS Microbiol Lett 289, 138–145.
    17. Ho, S. N., Hunt, H. D., Horton, R. M., Pullen, J. K. & Pease, L. R. (1989). Site-directed mutagenesis by overlap extension using the polymerase chain reaction. Gene 77, 51–59.
    18. Imlay, J. A. (2003). Pathways of oxidative damage. Annu Rev Microbiol 57, 395–418.
    19. Kalogeraki, V. S. & Winans, S. C. (1997). Suicide plasmids containing promoterless reporter genes can simultaneously disrupt and create fusions to genes of diverse bacteria. Gene 188, 69–75.
    20. Kim, J., Jeon, C. O. & Park, W. (2007). A green fluorescent protein-based whole-cell bioreporter for the detection of phenylacetic acid. J Microbiol Biotechnol 17, 1727–1732.
    21. Kim, J., Jeon, C. O. & Park, W. (2008). Dual regulation of zwf-1 by both 2-keto-3-deoxy-6-phosphogluconate and oxidative stress in Pseudomonas putida. Microbiology 154, 3905–3916.
    22. Lee, Y., Ahn, E., Park, S., Madsen, E. L., Jeon, C. O. & Park, W. (2006a). Construction of a reporter strain Pseudomonas putida for the detection of oxidative stress caused by environmental pollutants. J Microbiol Biotechnol 16, 386–390.
    23. Lee, Y., Peña-Llopis, S., Kang, Y. S., Shin, H. D., Demple, B., Madsen, E. L., Jeon, C. O. & Park, W. (2006b). Expression analysis of the fpr (ferredoxin-NADP+ reductase) gene in Pseudomonas putida KT2440. Biochem Biophys Res Commun 339, 1246–1254.
    24. Liochev, S. I., Hausladen, A., Beyer, W. F., Jr & Fridovich, I. (1994). NADPH: ferredoxin oxidoreductase acts as a paraquat diaphorase and is a member of the soxRS regulon. Proc Natl Acad Sci U S A 91, 1328–1331.
    25. Morgan, T. V., Lundell, D. J. & Burgess, B. K. (1988). Azotobacter vinelandii ferredoxin I: cloning, sequencing, and mutant analysis. J Biol Chem 263, 1370–1375.
    26. Nelson, K. E., Weinel, C., Paulsen, I. T., Dodson, R. J., Hilbert, H., Martins dos Santos, V. A., Fouts, D. E., Gill, S. R., Pop, M. & other authors (2002). Complete genome sequence and comparative analysis of the metabolically versatile Pseudomonas putida KT2440. Environ Microbiol 4, 799–808.
    27. Nunoshiba, T. & Demple, B. (1994). A cluster of constitutive mutations affecting the C-terminus of the redox-sensitive SoxR transcriptional activator. Nucleic Acids Res 22, 2958–2962.
    28. Ostrowski, J. & Kredich, N. M. (1989). Molecular characterization of the cysJIH promoters of Salmonella typhimurium and Escherichia coli: regulation by cysB protein and N-acetyl-l-serine. J Bacteriol 171, 130–140.
    29. Park, W., Peña-Llopis, S., Lee, Y. & Demple, B. (2006). Regulation of superoxide stress in Pseudomonas putida KT2440 is different from the SoxR paradigm in Escherichia coli. Biochem Biophys Res Commun 341, 51–56.
    30. Park, B. S., Kwon, Y. M., Pyla, R., Boyle, J. A. & Jung, Y. S. (2007). E1 component of pyruvate dehydrogenase complex does not regulate the expression of NADPH-ferredoxin reductase in Azotobacter vinelandii. FEMS Microbiol Lett 273, 244–252.
    31. Pomposiello, P. J. & Demple, B. (2000). Identification of SoxS-regulated genes in Salmonella enterica serovar typhimurium. J Bacteriol 182, 23–29.
    32. Pomposiello, P. J. & Demple, B. (2001). Redox-operated genetic switches: the SoxR and OxyR transcription factors. Trends Biotechnol 19, 109–114.
    33. Regnström, K., Sauge-Merle, S., Chen, K. & Burgess, B. K. (1999). In Azotobacter vinelandii, the E1 subunit of the pyruvate dehydrogenase complex binds fpr promoter region DNA and ferredoxin I. Proc Natl Acad Sci U S A 96, 12389–12393.
    34. Sainsbury, S., Lane, L. A., Ren, J., Gilbert, R. J., Saunders, N. J., Robinson, C. V., Stuart, D. I. & Owens, R. J. (2009). The structure of CrgA from Neisseria meningitidis reveals a new octameric assembly state for LysR transcriptional regulators. Nucleic Acids Res 37, 4545–4558.
    35. Schell, M. A. (1993). Molecular biology of the LysR family of transcriptional regulators. Annu Rev Microbiol 47, 597–626.
    36. Stadtman, E. R. (1992). Protein oxidation and aging. Science 257, 1220–1224.
    37. Tao, K., Makino, K., Yonei, S., Nakata, A. & Shinagawa, H. (1989). Molecular cloning and nucleotide sequencing of oxyR, the positive regulatory gene of a regulon for an adaptive response to oxidative stress in Escherichia coli: homologies between OxyR protein and a family of bacterial activator proteins. Mol Gen Genet 218, 371–376.
    38. Yanisch-Perron, C., Vieira, J. & Messing, J. (1985). Improved M13 phage cloning vectors and host strains: nucleotide sequences of the M13mp18 and pUC19 vectors. Gene 33, 103–119.
    39. Yeom, J., Jeon, C. O., Madsen, E. L. & Park, W. (2009a). Ferredoxin-NADP+ reductase from Pseudomonas putida functions as a ferric reductase. J Bacteriol 191, 1472–1479.
    40. Yeom, J., Jeon, C. O., Madsen, E. L. & Park, W. (2009b). In vitro and in vivo interactions of ferredoxin-NADP+ reductases in Pseudomonas putida. J Biochem 145, 481–491.
    41. Zheng, M., Åslund, F. & Storz, G. (1998). Activation of the OxyR transcription factor by reversible disulfide bond formation. Science 279, 1718–1721.