Bacteria encounter reactive oxygen species (ROS) as a consequence of the aerobic life or as an oxidative burst of activated neutrophils during infections. In addition, bacteria are exposed to other redox-active compounds, including hypochloric acid (HOCl) and reactive electrophilic species (RES) such as quinones and aldehydes. These reactive species often target the thiol groups of cysteines in proteins and lead to thiol-disulfide switches in redox-sensing regulators to activate specific detoxification pathways and to restore the redox balance. Here, we review bacterial thiol-based redox sensors that specifically sense ROS, RES and HOCl via thiol-based mechanisms and regulate gene transcription in Gram-positive model bacteria and in human pathogens, such as Staphylococcus aureus and Mycobacterium tuberculosis. We also pay particular attention to emerging widely conserved HOCl-specific redox regulators that have been recently characterized in Escherichia coli. Different mechanisms are used to sense and respond to ROS, RES and HOCl by 1-Cys-type and 2-Cys-type thiol-based redox sensors that include versatile thiol-disulfide switches (OxyR, OhrR, HypR, YodB, NemR, RclR, Spx, RsrA/RshA) or alternative Cys phosphorylations (SarZ, MgrA, SarA), thiol-S-alkylation (QsrR), His-oxidation (PerR) and methionine oxidation (HypT). In pathogenic bacteria, these redox-sensing regulators are often important virulence regulators and required for adapation to the host immune defense.
Bacteria have to cope in their natural environment or during bacterial infection in association with the host immune system to reactive oxygen species (ROS) that are known to cause an oxidative stress response and affect the reduced state of the cytoplasm. ROS are produced in microorganisms as the unavoidable consequence of the aerobic life, by incomplete reduction of molecular oxygen during respiration (Imlay, 2003, 2008, 2013). Beside ROS, bacteria have to cope with many other redox-active compounds, including antimicrobials, antibiotics and environmental xenobiotics, which can act as reactive electrophilic species (RES) and affect the cellular redox status (Marnett et al., 2003; Jacobs and Marnett, 2010). ROS and RES cause specific post-translational thiol-modifications in redox-sensing transcription factors which lead to conformational changes and activate or inactive the transcriptional regulator. As consequence, specific detoxification pathways are up-regulated to destroy the reactive species or to repair the resulting damage (Antelmann and Helmann, 2011; Vazquez-Torres, 2012; Imlay, 2013). With the discovery of the peroxide sensor OxyR of Escherichia coli, it became evident that ROS-sensing by thiol-disulfide switches represents an important regulatory device in bacteria (Zheng et al., 1998; Choi et al., 2001; Kim et al., 2002). However, during the last decade this classical thiol-disulfide-switch model for redox regulation has been expanded by different reversible and irreversible thiol-modifications, such as S-thiolation, Cys phosphorylation or thiol-S-alkylation that are employed by thiol-based redox sensors to regulate expression of specific antioxidant enzymes and virulence mechanisms. In addition to thiol-redox switches, redox sensors can also use methionine oxidation switches, His-oxidation or flavin cofactors, iron and iron-sulfur clusters, heme centers either directly or indirectly for redox-sensing. Here, we review the currently known thiol-based ROS, RES and HOCl-specific redox sensors that have been characterized in Gram-positive model bacteria and human pathogens as well as in E. coli.
Thiol chemistry of ROS, RES and HOCl
Reactive oxygen species (ROS) include superoxide anion (O2·-), hydrogen peroxide (H2O2) and the highly reactive hydroxyl radical (OH·) that are generated during aerobic respiration by the incomplete stepwise reduction of O2 (Imlay, 2003, 2008). The highly toxic hydroxyl radical (OH·) is produced in the Fenton reaction by H2O2 and free ferrous iron (Fe2+) (Imlay, 2003, 2008). Upon infection, the oxidative burst from activated neutrophils generates O2·-, H2O2, nitric oxide (NO) and hypochloric acid (HOCl) with the aim to kill invading pathogenic bacteria (Forman and Torres, 2001; Winterbourn and Kettle, 2012). Reactive electrophilic species include quinones, aldehydes, epoxides, diamide and α,β-unsaturated dicarbonyl compounds that have electron-deficient centers (Antelmann and Helmann, 2011). RES can arise endogenously as secondary reactive intermediates from oxidation products of amino acids, lipids or carbohydrates (Marnett et al., 2003; Rudolph and Freeman, 2009). The dicarbonyl compound methylglyoxal is produced as a byproduct during the glycolysis from triose-phosphate intermediates (Ferguson et al., 1998; Booth et al., 2003; Kalapos, 2008). Formaldehyde is encountered by bacteria as intermediate in the C1-metabolism of methanotrophic and methylotrophic bacteria. Thus, bacteria have evolved redox sensors and conserved detoxification pathways for the natural RES formaldehyde and methylglyoxal.
The thiol group of cysteine is the main target for ROS, RES and HOCl and is subject to reversible and irreversible post-translational thiol-modifications. The thiol group can be reversibly oxidized to protein disulfides or irreversibly overoxidized to sulfinic or sulfonic acids by ROS or S-alkylated by RES (Antelmann and Helmann, 2011). ROS lead first to oxidation of protein thiols to Cys sulfenic acids (R-SOH) that rapidly react further to form intramolecular, intermolecular disulfides or mixed disulfides with LMW thiols (termed as S-thiolations) (Figure 1). Hypochloric acid (HOCl) is a strong two-electron oxidant and chlorinating agent that targets the sulfur-containing amino acids cysteine and methionine with the second-order rate constants of k=3×107m-1s-1 (Hawkins et al., 2003). HOCl first chlorinates the thiol group to form the unstable sulfenylchloride intermediate that reacts further with another thiol group to form disulfides. In the absence of another Cys residue, the chlorinated thiol group is overoxidized very rapidly to sulfinic or sulfonic acids (Hawkins et al., 2003; Gray et al., 2013a) (Figure 1). RES, like quinones, can react with Cys thiols via thiol-disulfide switches or thiol-S-alkylation. During the incomplete one-electron reduction of quinones the highly reactive semiquinone radical is produced that leads to subsequent reduction of O2 and the production of O2·-. The electrophilic reaction of quinones involves the 1,4-reductive Michael-type addition of thiols to quinones (Marnett et al., 2003). Toxic quinones lead to irreversible thiol-S-alkylation and protein aggregation to deplete protein thiols in the proteome in vivo (Liebeke et al., 2008). However, non-toxic quinones cause disulfide formation in RES-sensing redox regulators, such as YodB or NemR to up-regulate quinone detoxification pathways (Chi et al., 2010a; Gray et al., 2013b; Lee et al., 2013).
Thiol-based redox sensors for ROS, RES and HOCl in bacteria
OxyR as thiol-based redox sensor for peroxides and NO in E. coli and Actinomycetes
OxyR is a redox sensor for peroxides and NO in Salmonella Typhimurium and E. coli and was the first discovered redox-sensitive regulator that is activated by a thiol-disulfide-switch model (Storz et al., 1990b; Zheng et al., 1998). OxyR belongs to the LysR-family of transcription factors that acts both as transcriptional activator of peroxide detoxification pathways and repressor of its own transcription and binds as tetramer to operator sequences (Figure 2, Table 1). Gisela Storz has shown that OxyR oxidation occurs by H2O2 at the conserved Cys199 that is oxidized to a sulfenic acid and subsequently forms an intramolecular disulfide with Cys208 in each of the four subunits of the OxyR tetramer (Storz et al., 1990b; Zheng et al., 1998). The OxyR tetramer binds to the operator sequences in the reduced and oxidized forms, but the interaction of reduced and oxidized OxyR with the DNA is different (Toledano et al., 1994). Oxidized OxyR recognizes a motif comprised of four ATAGnt elements spaced at 10 bp intervals and binds to this motif in four adjacent major grooves on one face of the DNA. Reduced OxyR binds to the DNA at two pairs of adjacent major grooves separated by on helical turn. The two modes of binding are essential for OxyR to function as both an activator and a repressor in vivo (Toledano et al., 1994). Oxidized OxyR induces the cooperative binding of the RNAP to activate transcription (Kullik et al., 1995a,b).
|Redox sensor||Organism||Signal||Redox-sensing mechanism||Regulon genes||Regulon functions||References|
|C199*-C208 intramolecular disulfide|
Fe-S cluster assembly
antigen 43 outer membrane
|Storz et al. (1990b)|
Zheng et al. (1998)
Choi et al. (2001)
Kim et al. (2002)
|H2O2||C25* peroxide sensing||katG||Catalase||Domenech et al. (2001)|
Li and He (2012)
|Chen et al. (2008)|
|OxyR||Corynebacterium glutamicum||H2O2||C206* and C215 conserved||katA|
dps, ftn cydABCD
Miniferritins cytochrome bd oxidase ferrochelatase
Regulator of iron-proteins
|Teramoto et al. (2013)|
Milse et al. (2014)
The crystal structures of reduced and oxidized OxyR and thiol-trapping assays confirmed the Cys199-Cys208 disulfide-switch model both in vitro and in vivo. The redox-sensing Cys199 is located in a loop between the α-helix and the β8 strand and is 17 Å apart from C208. OxyR oxidation to the Cys199-Cys208 intramolecular disulfide results in unwinding of the α-helix and movement of the α-helix/β8 loop causing large structural changes in the oligomeric interfaces and relative rotation among the OxyR subunits (Choi et al., 2001; Barford, 2004; Lee et al., 2004). Disulfide formation leads to the rearrangement of the N-terminal DNA-binding domains relative to the DNA to facilitate proper DNA-binding of oxidized OxyR to the four adjacent major grooves to induce the cooperative interactions with the RNAP required for transcriptional activation of peroxide detoxification genes.
However, this thiol-disulfide-switch model was questioned by the group of Jonathan Stamler, because mutational analyses suggested that only Cys199 is required for redox-sensing and transcriptional activation of OxyR. Different post-translational thiol-modifications were introduced at Cys199 of OxyR, including sulfenic acid formation, S-nitrosylation or S-glutathionylation that were sufficient for OxyR activation in vitro (Kim et al., 2002). These different OxyR modifications resulted in different OxyR activation states. The S-glutathionylated OxyR conferred non-cooperative DNA-binding while the sulfenic acid and S-nitrosylated forms of OxyR provoke cooperative binding to the DNA.
Interestingly, S-nitrosylation of OxyR occurred specifically under conditions of anaerobic nitrate respiration. Moreover, OxyR controls widespread endogenous protein S-nitrosylation and the expression of a different anaerobic OxyR regulon during nitrate respiration (Seth et al., 2012). The anaerobically OxyR-controlled hcp gene, encoding a hybrid cluster protein, was shown to be specifically activated by S-nitrosylated OxyR. The oxyR mutant showed a growth defect with nitrate and Hcp was required for protection against endogenous nitrosative stress under anaerobic nitrate respiration. This indicates that OxyR can be activated by different post-translational thiol-modifications, including the thiol-disulfide-switch under H2O2 stress and S-nitrosylation under anaerobic nitrate respiration to activate distinct regulons for protection against peroxide and nitrosative stress (Seth et al., 2012).
OxyR is conserved in Gram-negative and Gram-positive bacteria and has been studied in Proteobacteria, Bacteroidetes and Actinomycetes (Chiang and Schellhorn, 2012). In many bacteria, OxyR controls catalases and peroxiredoxins while the size of the OxyR regulon varies. In E. coli, OxyR positively regulates genes for the peroxide scavenging peroxiredoxin (ahpCF) and catalase (katG), the iron-uptake regulator (fur), the miniferritin (dps), the Mn-importer (mntH), the FeS-cluster assembly machinery (sufABCDE), the ferrochelatase for ferrous ion incorporation into heme (hemH), thioredoxins (trxC), glutaredoxins (grxA), glutathione reductase (gor) and the periplasmic sulfenic acid oxidoreductase (dsbG) (Storz et al., 1990a) (Figure 2, Table 1). OxyR is both an activator and repressor and controls negatively its own transcription and that of the genes for the ferric ion reductase (fhuF), the antigen 43 outer membrane protein (flu), the mannonate hydrolase and oxidoreductase (uxuAB), the gluconate permease (gntP) and some unknown function proteins (Zheng et al., 2001). Oxidized OxyR is reduced by the glutaredoxin (GrxA)/GSH/Gor reducing system upon return to non-stress conditions. The OxyR regulon genes confer peroxide resistance in E. coli, but protect cells also against heat, UV, singlet oxygen, lipid peroxides and neutrophil killing (Chiang and Schellhorn, 2012).
OxyR homologs have been studied in Gram-positive Actinomycetes, such as Mycobacteria and Corynebacteria where they control catalases and peroxiredoxins. Interestingly, in Mycobacterium tuberculosis the catalase KatG activates the anti-tuberculosis pro-drug isoniazid (INH) upon treatment of M. tuberculosis infections (Zhang et al., 1992). However, katG expression is not regulated by OxyR and the oxyR gene has acquired several non-sense mutations and is non-functional in M. tuberculosis (Deretic et al., 1997). These non-sense oxyR mutations are conserved among most Mycobacteria, except for Mycobacterium leprae and Mycobacterium avium which encode functional oxyR genes (Sherman et al., 1995). Expression of katG is regulated by the ferric uptake regulator FurA in M. tuberculosis and M. smegmatis (Milano et al., 2001; Pym et al., 2001; Zahrt et al., 2001). The furA gene is located upstream of katG and the furA-katG operon was induced by oxidative stress in a FurA-dependent manner (Milano et al., 2001). However, in a fast-growing Mycobacterium sp. strain JC1 DSM 3803, katG was induced FurA-independently under oxidative stress (Lee et al., 2010). This peroxide-inducible expression of katG was shown to be controlled by the OxyR-homolog OxyS in M. tuberculosis (Domenech et al., 2001; Li and He, 2012). The oxyR homologous oxyS gene is located in the M. tuberculosis cosmid T919 and is highly conserved among Mycobacteria. OxyS is a repressor of katG transcription and overexpression of OxyS resulted in stronger repression of katG transcription and increased susceptibility to H2O2 stress in M. smegmatis (Domenech et al., 2001; Li and He, 2012). The operator sequence for OxyS binding was identified as GC-rich T-N(11)-A motif within the katG promoter region. OxyS has 4 Cys residues: Cys113, Cys124 and Cys293 are in the LysR-substrate-binding domain and Cys25 is located in the N-terminal DNA-binding domain. Cys25 of mycobacterial OxyS is required for peroxide-sensing, which is conserved also among enteric OxyR proteins but absent from OxyR of M. leprae and M. avium (Domenech et al., 2001). The DNA-binding activity of OxyS was inhibited by H2O2in vitro in gel-shift assays while the OxySC25A mutant did not respond to H2O2in vitro and in vivo. These results suggest that OxyS regulation involves oxidation of the single Cys25 in the DNA-binding domain under oxidative stress, but the thiol-modification that inactivates OxyS is unknown (Li and He, 2012).
Similar to mycobacterial OxyS, a 1-Cys-type OxyR redox sensor was characterized in Deinococcus radiodurans that is oxidized at the conserved single Cys210 to a sulfenic acid under peroxide stress. OxyR of Deinococcus radiodurans activates transcription of genes for the catalase (katE), the ferrous iron transporter (feoB) and the iron(III)dicitrate transporter (drb0125), but also operates as repressor of dps and mntH transcription to control antioxidant functions and Mn/Fe ion homeostasis (Chen et al., 2008). This indicates, that 1-Cys-type OxyR homologs are also present in other bacteria, including OxyS of Mycobacteria and OxyR of D. radiodurans that might sense peroxide stress by alternative thiol-modifications similar as has been described for OxyR of E. coli (Kim et al., 2002).
In Corynebacterium glutamicum and Corynebacterium diphtheriae, OxyR is functional as transcriptional repressor of the catalase-encoding gene. In both species, disruption of oxyR led to derepression of the catalase gene that conferred a H2O2 resistance phenotype (Kim and Holmes, 2012; Teramoto et al., 2013). DNaseI-footprinting analyses revealed the OxyR binding region that is ∼50 bp long with multiple T-N11-A motifs but no sequence similarities, characteristic for operators recognized by LysR-type regulators. Reduced OxyR binds specifically to this operator sequence in different OxyR target gene promoters (Teramoto et al., 2013). However, DNA-binding activity of OxyR was not inhibited after oxidation and even non-specific binding of oxidized OxyR was observed. This suggests that alleviation of OxyR repression by peroxides might be the result of decreased strength of its interaction with the DNA. In genome-wide transcriptome analyses, the OxyR regulon of C. glutamicum was characterized after peroxide stress (Milse et al., 2014). OxyR acts as transcriptional repressor and negatively regulates expression of 23 genes that belong to 12 transcriptional units. DNA-binding assays confirmed specific binding of OxyR to the 12 target promoters. In total, the OxyR regulon consists of genes encoding the catalase (katA), two miniferritins that function in iron homeostasis (dps and ftn), cytochrome bd oxidases (cydABCD), the heme biosynthesis enzyme ferrochelatase (hemH), a flavin-monooxygenase (cg1292), the FeS-cluster biosynthesis machinery (suf operon), the proline-ectoine transporter (proP) and several unknown function genes (Table 1) (Milse et al., 2014). In addition, transcriptional regulators were regulated by OxyR, such as oxyR, sufR and ripA. OxyR of C. glutamicum shares with E. coli OxyR the conserved redox-sensing Cys199 and Cys206 residues indicating a similar thiol-disulfide-switch model for OxyR of C. glutamicum.
PerR as Fur-family metal-based peroxide sensor in Firmicutes bacteria
In B. subtilis, the Fur-family protein PerR functions as the main peroxide sensor. PerR is a dimeric repressor that binds to the PerR box (TTATAATNATTATAA) as heptameric 7-1-7 inverted repeat in the promoter region of its target genes (Fuangthong and Helmann, 2003). PerR is inactivated by H2O2 stress, leading to derepression of the PerR regulon genes. The PerR regulon includes the genes for the peroxiredoxin (ahpCF), the catalase (katA), the miniferritin (mrgA), the heme biosynthesis operon (hemAXCDBL), the iron-uptake repressor (fur), and the Zn-uptake system (zosA) (Fuangthong et al., 2002; Helmann et al., 2003) (Figure 3, Table 2). Two overlapping PerR boxes are present in the perR upstream region indicating that PerR is autoregulated (Fuangthong et al., 2002). The derepression of the PerR regulon genes under peroxide stress conditions leads to peroxide resistance as adaptive response in B. subtilis (Faulkner and Helmann, 2011). In S. aureus, the PerR regulon is also peroxide-inducible and includes genes for catalase, peroxiredoxins and bacterioferritin comigratory protein (katA, ahpCF, bcp), two miniferritins (mrgA, ftn) and thioredoxin reductase (trxB) that are required for virulence (Horsburgh et al., 2001). In addition, PerR negatively regulates its own transcription and that of the gene for the ferric uptake regulator (fur). PerR is required for full virulence in a murine skin abscess model of infection. It was further shown that both katA and ahpC have compensatory roles in peroxide resistance and mediate environmental persistence and nasal colonization in S. aureus (Cosgrove et al., 2007).
|Redox sensor||Organism||Signal||Redox-sensing mechanism||Regulon genes||Regulon functions||References|
C136-C139 intramolecular disulfide
|Fuangthong et al. (2002)|
Lee and Helmann (2006)
Chi et al. (2011)
|PerR||Staphylococcus aureus||H2O2||His43, His97, His99|
Bacterioferritin comigrating protein
|Horsburgh et al. (2001)|
|His32, His86, His88|
Bacterioferritin comigrating protein
|Hillmann et al. (2009a)|
Hillmann et al. (2009b)
Riebe et al. (2009)
The regulatory mechanism for peroxide-sensing of PerR has been shown by the Helmann group in B. subtilis (Lee and Helmann, 2006). PerR contains two metal binding sites, a structural Zn2+ binding site coordinated by four cysteine residues (Cys96, Cys99, Cys136, Cys139) in the C-terminal domain and a regulatory Fe2+ or Mn2+ binding site with three histidine and two aspartic acid residues as ligands (Lee and Helmann, 2006). Both Mn2+ and Fe2+ bind competitively to the PerR regulatory site, but only iron-bound PerR is sensitive to metal-catalyzed oxidation (Mongkolsuk and Helmann, 2002). Exposure to H2O2 leads to oxidation of Fe2+ in the regulatory site by a Fenton reaction generating HO· which causes oxidation of His37 and His91 to 2-oxo-histidine and inactivation of PerR (Figure 3) (Lee and Helmann, 2006; Traore et al., 2009; Duarte and Latour, 2010). Although the 2-oxo-His37 still had affinity for the regulatory metal, no metal binding with 2-oxo-His91 was possible and PerR fails to retain the close conformation for DNA-binding (Traore et al., 2009; Duarte and Latour, 2010). Thus, in contrast to OxyR which is activated by a thiol-disulfide redox switch, the PerR transcription factor senses peroxide stress by metal-catalyzed histidine oxidation. However, the PerR regulon genes are also induced under disulfide conditions, such as diamide and hypochlorite in B. subtilis (Antelmann et al., 2008; Chi et al., 2011). Thus, the response of PerR to disulfide stress could involve thiol-redox switches in the structural Zn-site of PerR, leading to inactivation of its repressor function (Figure 3). In support of this thiol-based mechanism, an intramolecular C136-C139 disulfide in the Zn-binding site of PerR was identified by mass spectrometry in hypochlorite-stressed B. subtilis cells in vivo (Chi et al., 2011). In addition, a thiol-redox switch was identified as peroxide-sensing mechanism of the Fur-family PerR homolog of Streptomyces coelicolor CatR that controls expression of the catalase gene catA (Hahn et al., 2000a,b). In CatR two CXXC motifs are present that coordinate Zn in the reduced state. In response to peroxide stress, intramolecular disulfides are formed in the Zn-site of CatR that inactivate CatR’s repressor function resulting in derepression of catA. Thus, it is likely that PerR homologs respond also via thiol-based mechanisms under certain disulfide-stress conditions that are different from peroxide stress.
The PerR homolog that senses peroxides and O2 has been identified in the strict anaerobic Gram-positive bacterium Clostridium acetobutylicum as defense mechanism against O2 toxicity (Hillmann et al., 2008). PerR inactivation conferred aerotolerance to C. acetobutylicum, enabled aerobic growth and O2 consumption and conferred resistance to H2O2. The PerR regulon includes genes for the reverse ruberythrins as major peroxidases for H2O2 reduction (rbr3A, rbr3B), the peroxiredoxin (bcp), the thiol peroxidase (tpx), the glutathione peroxidase (bsa2), the glutaredoxin (grx), the flavodoxin (CAC2452), the superoxide-reducing desulfoferrodoxin (dfx), the oxygen-reducing flavodiiron proteins (fprA2), rubredoxins (rd) as intermediates to regenerate the reductases FprA2, Dfx and revRbr, the NADH-dependent rubredoxin oxidoreductase (nror) to provide electrons for rubredoxin reduction, the 2-oxoglutarate ferredoxin oxidoreductase (ofrAB) and the NADPH-dependent non-phosphorylating GapDH (gapN) (Hillmann et al., 2009a,b; Riebe et al., 2009)(Figure 4, Table 2). These PerR regulon genes are all induced under O2 and peroxide stress and function collectively as anaerobic O2 and ROS detoxification pathways to promote the survival of the strict anaerobe C. acetobutylicum under short-time microaerophilic conditions.
The MarR/OhrR-family regulators as sensors of organic hydroperoxides
The MarR/OhrR-family regulators in B. subtilis and Xanthomonas campestris
MarR or Multiple antibiotics resistance-type regulators are characterized by winged helix-turn-helix (HTH) DNA-binding motifs and control genes that confer resistance to antibiotics, organic solvents, detergents, ROS and RES. Several MarR-family members are important for the regulation of virulence (Ellison and Miller, 2006). Among the MarR-family regulators, the MarR/OhrR subfamily responds to organic hydroperoxides (OHP). OHP can be derived from peroxidation of unsaturated fatty acids of eukaryotic membrane lipids. Ohr-like peroxiredoxins catalyze the reduction of OHPs to their corresponding alcohols (Atichartpongkul et al., 2001; Fuangthong et al., 2001). B. subtilis has two ohr paralogs: ohrA and ohrB (Fuangthong et al., 2001). The ohrA gene is regulated by the redox-sensing OhrR repressor and ohrB is controlled by the σB alternative sigma factor in B. subtilis (Völker et al., 1998).
The OhrR repressor acts as a dimeric repressor that binds to inverted repeat sequences in the ohrA promoter, thereby inhibiting transcription (Fuangthong et al., 2001). OhrR harbors a conserved redox-sensing Cys residue in its N-terminal region that senses OHPs via different redox-switch mechanisms. Thiol-oxidation of OhrR results in dissociation of the protein from the operator and derepression of ohrA transcription. Based on the number of Cys residues, the OhrR-family can be divided into two subfamilies: the one-Cys-type with the prototype of B. subtilis OhrRBs (Lee et al., 2007) and the two-Cys-type with the example of X. campestris OhrRXc (Figure 5, Table 3) (Panmanee et al., 2006; Antelmann and Helmann, 2011). In the two-Cys type OhrRxc, Cys22 is oxidized by OHPs to a sulfenic acid intermediate, which reacts further with Cys127 in the opposing subunit forming an intersubunit disulfide (Panmanee et al., 2006). Oxidation inactivates OhrRxc and releases the protein from the promoter DNA. X-ray crystallography reveals that disulfide formation causes a large rotation of the DNA-binding domain that is not compatible with DNA-binding (Newberry et al., 2007; Antelmann and Helmann, 2011). In contrast, the one-Cys type OhrRBs of B. subtilis is oxidized at Cys15 to the sulfenic acid that reacts further to a mixed disulfide with BSH (S-bacillithiolated OhrR) in response to OHPs (Lee et al., 2007) (Figure 5). B. subtilis OhrRBs can be also converted from a one-Cys-type to a two-Cys regulator by introduction of a C-terminal Cys at a position equivalent to Cys127 of OhrRxc (Soonsanga et al., 2008).
|Redox sensor||Organism||Signal||Redox-sensing mechanism||Regulon genes||Regulon functions||References|
|OhrR||Bacillus subtilis||ROOH NaOCl||C15*-SSB||ohrA||2-Cys-peroxiredoxin||Fuangthong et al. (2001)|
Lee et al. (2007)
Chi et al. (2011)
|ohrA||2-Cys-peroxiredoxin||Panmanee et al. (2006)|
Newberry et al. (2007)
hlgABC, pvl,lukED, lukMF-PV
lytM, lytN norA, norB, tetAB
agr, lytRS, arlRS, sarS and sarV
Autolysis factors Multidrug efflux pumps
|Luong et al. (2006)|
Chen et al. (2006)
Chen et al. (2009)
Sun et al. (2012)
argGH, ilvD, lysC, hisC
gntRK, lacD, malA, treC,
isdC, epiEF, lrgB, efb, fib, tcaA
|2-Cys peroxiredoxin pyruvate metabolism|
Amino acid metabolism
Fatty acid synthesis
Cell surface proteins
Drug efflux pumps
|Ballal et al. (2009)|
Chen et al. (2009)
Poor et al. (2009)
Sun et al. (2012)
rot, agr, sarS, sarV, sarT
|Superoxide dismutase |
|Ballal and Manna (2009)|
Ballal and Manna (2010)
Sun et al. (2012)
|Brugarolas et al. (2012)|
|Corynebacterium glutamicum||H2O2||C92* redox-sensitive||rosR|
|Bussmann et al. (2010)|
|Palm et al. (2012)|
|Leelakriangsak et al. (2008)|
Chi et al. (2010a)
Chi et al. (2010b)
|C7*redox-sensitive||catDE||Dioxygenase||Chi et al. (2010b)|
|FMN-dependent quinone |
|Ji et al. (2013)|
|qorA||Quinone reductase||Ehira et al. (2009a)|
We could show that OhrR responds also to NaOCl stress and thus, OhrR is a redox sensor for OHPs and NaOCl (Chi et al., 2011). Transcriptome analysis revealed that the ohrA gene was the most strongly up-regulated gene (220-fold) under NaOCl stress in B. subtilis. Mass spectrometry identified the S-bacillithiolation of OhrR as thiol-redox switch mechanism for OhrR inactivation. Phenotype analyses showed that the OhrA peroxiredoxin and BSH protect cells against hypochlorite stress as the growth of ohrA and bshA mutants was strongly impaired by NaOCl stress (Chi et al., 2011). Thus, we hypothesize that OhrA could be involved in NaOCl detoxification. Because hypochlorite is produced by activated neutrophils, it could be also a more physiologically oxidant for the OhrR-homologs SarZ and MgrA in the human pathogen S. aureus.
The B. subtilis OhrR repressor is redox-controlled by S-bacillithiolation which was recently shown to be reversed by bacilliredoxins (BrxA/B), which function as glutaredoxin-like enzymes in the reduction of BSH mixed protein disulfides to regenerate and re-activate the OhrR repressor in B. subtilisin vitro (Gaballa et al., 2014).
The MgrA/SarZ/SarA family of virulence and antibiotic regulators
In S. aureus, two homologs of the MarR/OhrR 1-Cys-type repressor are present, including the MgrA and SarZ global regulators for antibiotic resistance and virulence (Figure 6, Table 3) (Truong-Bolduc et al., 2005, 2008; Kaito et al., 2006; Ballal et al., 2009; Chen et al., 2009; Poor et al., 2009). The Multiple gene regulator MgrA of S. aureus regulates more than 300 genes that are involved in virulence, autolysis, antibiotic resistance, biofilm formation and cell wall biosynthesis (Luong et al., 2006). MgrA controls genes for virulence factors that include enzymes for capsule polysaccharide biosynthesis (cap5(8)-locus), α-toxin (hla), coagulase (coa), protein A (spa), extracellular serine proteases (splABCDEF) and nuclease (nuc). In addition, genes encoding autolysis factors (lytM and lytN), multidrug efflux pumps (norA, norB and tetAB) and regulatory genes (agr, lytRS, arlRS, sarS and sarV) are members of the MgrA regulon (Ingavale et al., 2005; Kaatz et al., 2005; Truong-Bolduc et al., 2005, 2008; Luong et al., 2006) (Table 3). Hence, MgrA confers resistance to the antibiotics fluoroquinone, tetracycline, vancomycin and penicillin. MgrA is also required for virulence in murine abscess, septic arthritis and sepsis models. MgrA shares with OhrRBs the single conserved Cys12 and uses a thiol-based oxidation sensing mechanism to control virulence and antibiotic resistance (Chen et al., 2006). Cys12 can be oxidized by CHP, H2O2 and superoxide anion to Cys-SOH that leads to dissociation of MgrA from the operator DNA in vitro and induction of antibiotic resistance in S. aureusin vivo (Chen et al., 2006, 2009). However, it was shown recently that the DNA-binding activity of MgrA can be also reversibly regulated by cysteine phosphorylation via the eukaryotic-like serine/threonine kinase (Stk1) and phosphatase (Stp1)(Sun et al., 2012). Cys phosphorylation was detected as post-translational modification also in other regulatory proteins, including the MarR-family proteins SarZ and SarA and the cysteine biosynthesis regulator CymR. Moreover, Stk1 was required for full virulence and resistance to the antibiotic vancomycin by controlling Cys-phosporylation of MgrA, SarZ and SarA. Increased Cys phosphorylation of the virulence regulators in a stp1 mutant led to decreased virulence in a mouse abscess model. Interestingly, like the previously shown thiol-oxidation mechanism (Chen et al., 2006, 2009) Cys phosphorylation was also DTT-reversible, but the mechanism is still unknown (Sun et al., 2012).
The second MarR/OhrR-type regulator of S. aureus is SarZ, which is also a pleiotropic virulence regulator (Kaito et al., 2006). SarZ controls the ohr peroxiredoxin, genes involved in virulence, autolysis, cell wall metabolism, antibiotic resistance, intermediary, amino acid, sugar, fatty acid and anaerobic metabolism, such as the pyruvate-formate lyase genes pflA and pflB (Figure 6, Table 3). SarZ is also transcriptionally activated by MgrA (Ballal et al., 2009; Chen et al., 2009). SarZ was shown to use a thiol-based oxidation sensing mechanism via the conserved lone Cys13 residue (Chen et al., 2009). The crystal structure of SarZ was resolved in the reduced, sulfenic acid and mixed disulfide form (Poor et al., 2009). SarZ is oxidized at Cys13 to sulfenic acid that still retains DNA-binding activity. Further oxidation of SarZ with an external synthetic thiol (benzene thiol) leads to S-thiolated SarZ. These mixed SarZ disulfides cause steric clashes that contribute to an allosteric conformational change of the DNA-binding domains and release of SarZ from the operator DNA (Poor et al., 2009).
SarZ is also controlled by Cys phosphorylation via the Stk1/Stp1 kinase/phosphatase pair (Sun et al., 2012). However, it remains yet to be shown if S-bacillithiolation can control DNA-binding activity of SarZ or MgrA in vivo also in S. aureus. Future studies should elucidate if there is a possible cross-talk between Cys phosphorylation and thiol-oxidation in these MarR-family virulence regulators of S. aureus.
The Staphylococcal accessory regulator (SarA) is another global redox-sensing regulator of the MarR-family that contains a single Cys9 residue in the dimer interface. SarA positively regulates many virulence factors including fibronectin and fibrinogen binding proteins (fnb), hemolysins (hla), enterotoxins (sec), toxic shock syndrome toxin 1, and genes involved in biofilm formation (icaRA, bap) (Cheung et al., 2008; Tamber and Cheung, 2009). SarA negatively regulates the transcription of proteases (ssp, aur), protein A (spa), and collagen-binding proteins (cna). Many virulence regulators are members of the SarA-regulon of S. aureus, such as rot, agr, sarS, sarV, sarT resulting in pleiotropic phenotypes of the sarA mutant. Furthermore, SarA was shown to control oxidative stress-related genes, such as superoxide dismutase (sodA) and thioredoxin reductase (trxB) (Ballal and Manna, 2009; Ballal and Manna, 2010) (Table 3). The redox-sensitivity of Cys9 in SarA has been analyzed in the wild type and a SarAC9G mutant in vivo and in vitro, which revealed that oxidation of Cys9 by H2O2 and diamide reduced the DNA-binding activity of SarA to the trxB promoter (Ballal and Manna, 2010). SarA has been also shown to sense oxidative stress by Cys phosphorylation in vitro (Sun et al., 2012). However, the detailed thiol-switch mechanism for SarA redox regulation in vivo has yet to be elucidated.
The MarR/OhrR-family regulators MosR and RosR in Actinomycetes
The MarR/OhrR-family of redox regulators is also conserved among Actinomycetes. In M. tuberculosis the OhrR-family regulator MosR has been characterized as transcriptional repressor and sensor for peroxides that shares 28% sequence identity with B. subtilis OhrR and S. aureus MgrA (Brugarolas et al., 2012). Reduced MosR binds to a specific operator sequence (GTGTAnnTACAC) in its target promoters and represses its own transcription and that of the adjacent rv1050 gene, encoding an exported oxidoreductase of unknown function (Brugarolas et al., 2012). The rv1050 gene was most strongly induced by H2O2 and 352-fold derepressed in the mosR mutant. Rv1050 was also up-regulated during infection in INF-γ-activated macrophages, suggesting a role in the host immune defense (Schnappinger et al., 2003). In addition, arachidonic acid and linoleic acid were found to induce rv1050, indicating that Rv1050 could also function in fatty acid metabolism in macrophages.
MosR contains four Cys residues (Cys10, 12, 96, 147), but only Cys12 is conserved. The Cys10 and Cys12 residues are oxidized to intramolecular disulfides by peroxides and both Cys residues are essential for redox-sensing as C10S and C12S mutant proteins were non-responsive to H2O2 in gel-shift assays (Brugarolas et al., 2012). The structures of reduced and oxidized MosR proteins were resolved to reveal the structural mechanism for the inactivation of MosR’s repressor function upon oxidation. Disulfide formation between Cys10 and Cys12 breaks the hydrogen bond of Cys12 to Asn37′ and causes new hydrogen bonds of Arg16 and Ser41′. This rearrangement of hydrogen bonds results in a movement of α2 which pushes α3<4.5 Å towards α4. This causes rotation of α4 and α4′ that prevent them to fit into consecutive major grooves resulting in the release from the operator DNA. Consequently, MosR oxidation leads to rearrangements of hydrogen bonds, resulting in large conformational changes and MosR dissociation from the DNA (Brugarolas et al., 2012).
Corynebacterium glutamicum encodes the redox-sensitive MarR-type repressor RosR that responds to peroxide stress (Bussmann et al., 2010). RosR controls positively expression of the narKGHJI operon encoding a nitrate/nitrite transporter and the dissimilatory nitrate reductase complex. RosR acts as repressor of its own transcription and represses several genes that encode luciferase-like monooxygenases (cg1848, cg2329, cg3085), flavin-containing monooxygenases (cg3084), FMN reductases (cg1150, cg1850), glutathione-S-transferases (cg1426) and a polyisoprenoid-binding protein (cg1322). The polyisoprenoid-binding protein was important under peroxide stress as the mutant showed increased H2O2 sensitivity (Bussmann et al., 2010). Reduced RosR binds to an 18-bp inverted repeat with the consensus sequence TTGTTGAYRYRTCAACWA in its target promoters. The DNA-binding activity of RosR was inhibited by H2O2 and restored by DTT in gel-shift assays in vitro. RosR contains three Cys residues (Cys64, 92, 151), but only Cys92 is conserved among RosR homologs of other Corynebacteria. Cys92 was most important for redox-sensing because the DNA-binding activity of the C92S mutant was not inhibited by H2O2in vitro. In contrast, other single Cys mutants behaved like the wild type RosR, although in double and triple Cys mutants the DNA-binding activity was also affected in response to H2O2in vitro (Bussmann et al., 2010). This suggests that RosR might be inactivated by formation of inter- or intramolecular disulfides by peroxide stress which remains to be demonstrated.
The MarR/DUF24-family regulators as sensors for RES (quinone, diamide)
The MarR/DUF24-family regulators YodB, CatR, HypR and HxlR of B.subtilis
The MarR/DUF24 family of transcription factors is conserved among Gram-positive bacteria (Antelmann and Helmann, 2011). In C. glutamicum, the MarR/DUF24-type regulator QorR was first characterized as a transcriptional repressor that senses diamide and H2O2 and controls the quinone oxidoreductase QorA (Ehira et al., 2009a). Inactivation of QorR involves intersubunit disulfide formation between the conserved single Cys17 residues of both subunits (Ehira et al., 2009a). B. subtilis encodes eight MarR/DUF24-family proteins: HxlR, HypR, YodB, CatR, YdeP, YdzF, YkvN, and YtcD. HxlR was identified as activator of the formaldehyde-inducible hxlAB operon that encodes enzymes of the ribulose monophosphate pathway (Yurimoto et al., 2005). HypR was characterized as positive regulator of the nitroreductase HypO that is induced by NaOCl, diamide and quinones and confers NaOCl-resistance (Palm et al., 2012) (Table 3). HypR is a two-Cys MarR/DUF24-type regulator with a redox-sensing Cys14 and a second Cys49 that are 8 Å apart in the reduced HypR structure. Both Cys14 and Cys49 are essential for activation of hypO transcription by disulfide stress. HypR is activated by Cys14-Cys49′ intersubunit disulfide disulfide formation under diamide and NaOCl stress. Disulfide bond formation breaks the H-bonds of Cys14 and moves the α4 and α4′ helices of HypR ∼4 Å towards each other (Palm et al., 2012).
The MarR-type regulators YodB, CatR and MhqR control specific detoxification pathways that confer resistance to quinones and diamide, such as azoreductases (AzoR1 and AzoR2), nitroreductases (YodC and MhqN), and thiol-dependent dioxygenases (CatE, MhqA, MhqE, MhqO) (Töwe et al., 2007; Antelmann et al., 2008; Leelakriangsak et al., 2008; Chi et al., 2010b). Azoreductases and nitroreductases reduce quinones and diamide to hydroquinones and dimethylurea, respectively (Figure 7, Table 3). Dioxygenases catalyze the ring-cleavage reaction of quinone-S-adducts. The azoreductase AzoR1 is controlled by YodB and expression of the catechol-2,3- dioxygenase CatE and oxidoreductase CatD are regulated by both YodB and CatR (Leelakriangsak et al., 2008; Chi et al., 2010b). The promoter region of the catDE operon contains two inverted repeat sequences overlapping the -35 promoter region (BS1) and the transcription start point (BS2) that are the operator sites for CatR and YodB. Both YodB and CatR are inactivated in response to quinone and diamide. YodB is inactivated by a two-Cys-type redox-switch mechanism and oxidized to intersubunit disulfides between Cys6 of one subunit and Cys101 or Cys108 of the other subunit by diamide and quinones in vivo and in vitro (Chi et al., 2010a). The conserved Cys7 is essential for redox-sensing of quinones and diamide in CatR, but its redox-sensing mechanism has yet to be explored.
The MarR/DUF24-family regulator QsrR of S. aureus
The redox-sensing mechanism of the quinone-sensing MarR/DUF24-family regulator YodB (QsrR) has also been characterized in S. aureus, which shares 38% sequence identity with the YodB repressor of B. subtilis (Ji et al., 2013). QsrR contains the conserved N-terminal Cys5 and two further Cys30 and Cys33 residues. QsrR and YodB control both homologous genes involved in quinone reduction and ring-cleavage that confer resistance to benzoquinone. The QsrR regulon includes genes for the FMN-dependent quinone reductase (SAV0340 or azoR1), the nitroreductase (SAV2033 or yodC), the glyoxalase (SAV0338) and the thiol-dependent dioxygenase (SAV2522 or catE). Hence, QsrR controls homologous quinone reductases and dioxygenases in S. aureus that are controlled by YodB and CatR in B. subtilis (Figure 7, Table 3). The quinone resistance QsrR regulon conferred resistance to killing by macrophages in a phagocytosis assay indicating its crucial role for virulence regulation in S. aureus. The conserved Cys5 of QsrR was shown to sense quinones by a thiol-S-alkylation mechanism. The QsrR structure shares strong similarities with the HypR structure of B. subtilis (Palm et al., 2012; Ji et al., 2013). HypR and QsrR are both dimers that have in common the wHTH motif composed of α3, α4, β1 and β2. The wHTH motifs bind to the major and minor grooves of the DNA double helix and the wing is much larger compared to OhrR-like regulators. To elucidate the structural changes upon quinone-binding at Cys5, the menadione-bound QsrR structure was resolved for the QsrRC30,33S mutant (Ji et al., 2013). Menadione-binding at Cys5 causes a shift in the distance and rotation between the α4 and α4′ helices from 29.9 Å distance with 106° rotation in reduced QsrR to 39.1 Å distance and 117° rotation in the menadione-bound form. These structural changes lead to dissociation of QsrR from the operator DNA. In contrast to QsrR, YodB and HypR sense diamide and quinones by intersubunit disulfide bond formation (Palm et al., 2012). However, the in vivo mechanism of quinone-sensing by wild type QsrR has yet to be explored.
Emerging thiol-based redox sensors for RES (quinones, aldehydes) and HOCl
The TetR-family regulator NemR as redox sensor for RES (N-ethylmaleimide, quinones, aldehydes) and HOCl
The thiol-based TetR-family NemR regulator is conserved across Gram-negative and Gram-positive bacteria and has been characterized in E. coli as redox sensor for RES and HOCl (Gray et al., 2013b; Lee et al., 2013; Ozyamak et al., 2013) (Table 4). NemR negatively controls transcription of the nemRA operon and the gloA gene, which function in detoxification of electrophiles. The nemA gene was previously shown to be strongly induced by thiol-alkylating compounds, such as N-ethylmaleimide (NEM) and iodoacetamide and shown to function as NEM reductase (Umezawa et al., 2008). The FMN-dependent reductase NemA belongs to the old-yellow-enzyme family and has a broad substrate spectrum to reduce several quinones (ubiquinone, menaquinone) and aldehydes, (glyoxal, methylglyoxal and glycolaldehyde) in vitro (Lee et al., 2013). GloA is the glyoxalase-I enzyme involved in methylglyoxal detoxification and was revealed as main methylglyoxal protection mechanism (MacLean et al., 1998). Hence, the NemR repressor responds to quinones and aldehydes like methylglyoxal and is inactivated via thiol-based redox switches, which lead to up-regulation of the nemRA operon and gloA that both confer resistance to methylglyoxal and quinones in E. coli (Lee et al., 2013; Ozyamak et al., 2013). Moreover, the nemRA operon and gloA were most strongly induced by methylglyoxal in a transcriptome analyses supporting the major role of GloA and NemA as protection mechanism (Ozyamak et al., 2013). However, NemR was also shown to sense reactive chlorine species, such as HOCl and N-chlorotaurine, which leads to derepression of nemRA and gloA that both confer HOCl resistance as the nemA and gloA mutants displayed HOCl sensitive phenotypes (Gray et al., 2013b). Exposure of cells to HOCl stress caused increased methylglyoxal production suggesting that detoxification of methylglyoxal is an important bacterial HOCl defense mechanism. It remains to be shown if NemA could also confer HOCl resistance by reduction of reactive chlorines as direct substrates.
|Redox sensor||Organism||Signal||Redox-sensing mechanism||Regulon genes||Regulon functions||References|
quinones and NEM
|Umezawa et al. (2008)|
Gray et al. (2013)
Lee et al. (2013)
Ozyamak et al. (2013)
|Parker et al. (2013)|
disulfide (in vitro)
C4: HypT dodecamer
C150: HypT stability
|Sulfur, Cys and Met |
involved in iron
|Gebendorfer et al. (2012)|
Drazic et al. (2013a)
Drazic et al. (2013b)
NemR was shown to sense reactive electrophiles and HOCl via thiol-based oxidation mechanisms that involve intersubunit disulfides which lead to inactivation of its repressor function. NemR possesses 6 Cys residues, but only Cys106 is conserved among bacteria. However, the Cys106S mutant still responds to HOCl and forms intersubunit disulfides both in vitro and in vivo, suggesting that other Cys residues are involved in redox regulation of NemR which can substitute for the absence of the conserved Cys106 (Gray et al., 2013b). In another study, the glyoxal sensitivity and effect on nemRA expression was analyzed for various NemR Cys double mutants, revealing significant lower responsiveness to glyoxal in the Cys21,116S double mutant both in vivo and in vitro, but no difference to the wild type in any other Cys single and double mutant (Ozyamak et al., 2013). Cys21 is located in the DNA-binding domain and Cys116 is in the dimer interface. Both Cys21 and Cys116 were involved in intersubunit disulfide formation and oligomerization of NemR. This indicates that E. coli NemR is inactivated by intersubunit disulfide formation in response to RES and HOCl to up-regulate quinone and glyoxal detoxification enzymes. This thiol-disulfide-switch model of NemR redox regulation resembles that of other quinone-sensing redox regulators, such as QorR, YodB and HypR suggesting that the alternative thiol-S-alkylation as shown for QsrR in vitro might be rather the exception for thiol-based quinone-sensing regulators.
RclR as AraC-family HOCl-specific thiol-based redox sensor in E. coli
Recently, novel HOCl-specific redox regulators have been discovered in E. coli that are specific for chlorine species, such as HOCl, but do not respond to ROS, electrophiles or other thiol-reactive compounds. RclR (formerly YkgD) is widely conserved among Gram-negative bacteria and Actinobacteria and was characterized as redox-sensing transcriptional activator of the AraC family, which uses a thiol-based oxidation mechanism for redox-sensing of HOCl (Parker et al., 2013). The redox-sensing mechanism of RclR involves both conserved Cys residues, Cys21 and Cys89 which likely form an intramolecular disulfide that stabilizes the active RclR protein in vivo. Both Cys21 and Cys89 residues are required for redox-sensing of the HOCl-response in vivo, while only Cys21 is essential for redox-sensing in vitro. Oxidation of RclR by HOCl leads to specific activation of transcription of the rclABC operon, which is important for survival of HOCl and N-chlorotaurine. Mutants in each single gene of the rclABC operon are sensitive to HOCl suggesting that this operon is an important HOCl-protection determinant (Parker et al., 2013). However, the functions of the RclABC proteins for HOCl-protection are still unknown, which resemble a flavoprotein disulfide reductase, periplasmic protein and possible quinone-binding membrane protein (Table 4).
HypT as LysR-family HOCl-specific Met-oxidation switch in E. coli
The LysR-type regulator HypT has been discovered as another HOCl-specific redox sensor and transcriptional activator of E. coli (Gebendorfer et al., 2012). HypT belongs like OxyR to the LysR-family of transcriptional regulators, which often form dimers or tetramers (OxyR) (Maddocks and Oyston, 2008), but HypT was shown to form unusual large dodecameric ring-like structures in vitro that serve as a storage form of HypT (Drazic et al., 2014). These HypT dodecamers dissociate into smaller oligomers (dimers, tetramers) in the presence of DNA in vitro and tetramers are also found in vivo in HOCl-exposed E. coli cells. Specifically, the HypT tetramers were revealed as activation-competent DNA-binding species of HypT (Gebendorfer et al., 2012; Drazic et al., 2014).
The DNA-binding activity of HypT was activated specifically by HOCl and HypT was required for the survival under HOCl stress conditions in vivo (Gebendorfer et al., 2012). HypT was shown to control positively genes that function in sulfur, Cys and Met biosynthesis (metB, metK, metN, cysH, cysK, cysN, cysPUW, sbp, sufA) while genes of the Fur-regulon are negatively regulated by HypT that function in iron homeostasis (entC, entH, fecABCDE, fecR, fepCD, ryhB, tonB, yncE) (Table 4). As Met is rapidly oxidized by HOCl to methionine sulfoxide (Met-SO), it is suggested that HypT activates Met biosynthesis to restore the pool of reduced Met (Gebendorfer et al., 2012). Interestingly, HypT uses a reversible methionine oxidation switch model for transcriptional activation (Drazic et al., 2013a) while the Cys residues are important for stability and oligomerization of HypT (Drazic et al., 2013b). HypT activation involves oxidation of three Met residues (Met123, 206, 230) to their Met-SO forms which were identified in HOCl-activated HypT in vitro. Mutations of the three Met to glutamines mimics the oxidized Met-SO form of HypT and resulted in constitutively active HypT in vivo, while the Met-to-Ile mutation resulted in inactive HypT as revealed by the transcriptional studies of the target genes and HOCl survival assays (Drazic et al., 2013a). Furthermore, inactivation of oxidized HypT required the Met-SO reductases MsrA and MsrB both in vivo and in vitro, revealing the reversibility of this Met-oxidation switch model for HypT.
Surprisingly, HypT was only activated in vivo in E. coli cells by HOCl stress, but in vitro HypT rapidly lost its DNA-binding activity when treated with HOCl (Drazic et al., 2013b). HypT possesses five non-conserved Cys residues and all Cys residues are required for HypT activity in vivo and HypT stability in vitro. HypT oxidization by HOCl in vitro leads to intermolecular Cys4-Cys4 disulfides resulting in HypT inactivation. Furthermore, Cys150 was required for HypT stability and Cys4 involved in oligomerization of HypT to dodecamers (Drazic et al., 2013b). The thiol-switch model of HypT was suggested as check point in the activation of HypT preventing unwanted HypT interaction with its target promoters under oxidative stress conditions that are not sufficient to activate HypT. In conclusion, the HOCl-specific regulator HypT represents an important HOCl-protection mechanism and is activated by a reversible Met-oxidation switch to up-regulate Met biosynthesis in E. coli. It will be interesting to explore if other bacteria also use Met-oxidation switches as defense mechanisms against HOCl or ROS.
The Spx disulfide-stress redox sensors in Gram-positive Firmicutes bacteria
SpxA and MgsR as paralogous thiol-redox sensors for disulfide and general stress conditions in B. subtilis
The thiol-based redox sensor SpxA is an unusual transcription factor without typical DNA-binding domains that responds to different redox stress conditions in Gram-positive bacteria (Zuber, 2004, 2009; Nakano et al., 2005; Antelmann and Helmann, 2011). SpxA is an arsenate reductase (ArsC) family protein with a CXXC redox switch motif in its N-terminus that is essential for redox-sensing and transcriptional activation. ROS, RES and HOCl lead to oxidation of the CXXC motif to an intramolecular disulfide to activate SpxA (Figure 8). Oxidized SpxA interacts with the C-terminal domain (CTD) of the α subunit of the RNA polymerase (RNAP) to recognize promoter regions of the SpxA regulon genes and thereby activates transcription (Nakano et al., 2003, 2005; Zuber, 2004, 2009). SpxA positively regulates the expression of genes that maintain the thiol-redox balance in B. subtilis, including the genes for thioredoxin/thioredoxin reductases (trxAB), thiol peroxidase (tpx), FMN-dependent oxidoreductases (nfrA, yugJ), methionine sulfoxide reductase (msrA), cysteine biosynthesis and cystine transporters (yrrT operon, cysK, tcyABC) and bacillithiol biosynthesis (bshA, bshB1, bshB2 and bshC) (Zuber, 2009; Antelmann and Helmann, 2011; Gaballa et al., 2013) (Table 5). Genome-wide chromatin immunoprecipitation (ChIP-chip) analysis of RNAP-SpxA complexes revealed 275 genes that are directly controlled by SpxA. An extended -35 box was identified within SpxA-controlled promoters in which the -43/-44 positions correlated with the activation by SpxA (Rochat et al., 2012). Additional targets for SpxA control were identified among the Clp machinery as ATPase subunits (clpX, clpE and clpC) and the proteolytic subunit (clpP) and the adpaptor for ClpXP proteolysis (yjbH). Furthermore, SpxA controls thiol-based redox regulators (hxlR, yodB, yhdQ, yceK) and many other transcription factors that respond to diamide and SpxA was required for the basal level of 32 genes under non-stress conditions (Rochat et al., 2012) (Table 5).
|Redox sensor||Organism||Signal||Redox-sensing mechanism||Regulon genes||Regulon functions||References|
methionine sulfoxide reductase
ATPase subunit of Clp
Protease subunit of Clp
Adpaptor for ClpXP
|Nakano et al. (2003)|
Nakano et al. (2005)
Rochat et al. (2012)
|Reder et al. (2008)|
Reder et al. (2012b)
|Pamp et al. (2006)|
Wang et al. (2010)
Jousselin et al. (2013)
CoASH disulfide reductases
DNA damage repair
Spore outgrowth and germination
Adpaptor for ClpXP
|Barendt et al. (2013)|
|Kajfasz et al. (2010)|
Zheng et al. (2014)
Expression of SpxA is controlled at transcriptional and post-translational levels. Transcription of spxA is initiated from at least four different promoters that are recognized by three forms of RNAP containing σA, σB and σM (Leelakriangsak and Zuber, 2007). SpxA is transcriptionally regulated in response to disulfide stress provoked by diamide and NaOCl by the PerR and YodB repressors. Both PerR and YodB repressors are oxidized under disulfide stress conditions, which inactivates their repressor function leading to derepression of spxA transcription (Zuber, 2009). The YodB repressor is oxidized to intermolecular disulfides between C6 and one of the C-terminal Cys residues leading the spxA, azoR1 and yodC derepression. PerR is oxidized to form intramolecular disulfides in the Zn-binding structural site that possibly lead to PerR inactivation and spxA transcription by diamide and NaOCl. The second level is the post-translational control of SpxA protein stability by proteolysis via the ClpXP proteases. The spxA gene was discovered as Suppressor of clpP and clpX mutations and it was shown that the absence of ClpXP leads to stabilization of SpxA responsible for the pleiotropic phenotypes of clpXP mutants (Nakano et al., 2001). Under non-stress conditions, SpxA is targeted to the ClpXP system with the help of the adaptor YjbH for rapid degradation of SpxA (Nakano et al., 2002; Larsson et al., 2007). The YjbH adaptor protein contains a His-Cys-rich N-terminal region that is oxidized by diamide resulting in loss of YjbH adaptor activity and SpxA stabilization. However, mutational analyses showed that the Cys and His residues of YjbH are not required for SpxA stability in vivo (Chan et al., 2012). Studies with the YjbHGt ortholog of Geobacillus thermodenitrificans without the Cys-His redox motif revealed that the C-terminus of YjbHGt is required for stabilization of SpxA (Chan et al., 2012). The ATPase ClpX contains a Cys-rich Zn finger motif that also functions as redox switch and is oxidized under disulfide-stress conditions (Zhang and Zuber, 2007). Therefore, SpxA of B. subtilis is controlled at multiple levels, at the transcriptional level by PerR and YodB and at post-translational levels via three redox switches in SpxA itself, YjbH and ClpX that together lead to up-regulation, stabilization and activation of SpxA (Zuber, 2009; Antelmann and Helmann, 2011) (Figure 8). The mechanism of how the SpxA-RNAP complex activates transcription is unknown, but a cis-acting element in the -10 promoter sequences of trxB and trxA was identified required for activation of transcription (Reyes and Zuber, 2008; Nakano et al., 2010).
While SpxA interacts with RNAP containing σA, a SpxA paralog MgsR was identified as member of the σB general stress regulon (Reder et al., 2008). Thus, B. subtilis contains Spx paralogs that interact with RNAP holoenzymes containing different sigma factors. MgsR is a Modulator of the σBgeneral stress response and controls a sub-regulon within the σB regulon that functions in protection against secondary oxidative stress caused by ethanol, heat, salt stress (Reder et al., 2008). MgsR controls positively about 50 genes including 18 σB-dependent genes that were up-regulated in the mgsR mutant. Antioxidant and protective functions can be attributed to the genes encoding the manganese-containing catalase (ydbD), the thiol-disulfide oxidoreductase (ykuV) and paralogous short-chain oxidoreductases (ydaD, yhdF, yhxC and yhxD) that are postulated to function in NADPH production as electron source for cellular Trx/TrxR reducing systems (Reder et al., 2008) (Table 5). The mechanism of MgsR control resembles in part that of SpxA and involves a positive autoregulatory loop to increase mgsR transcription, and post-translational control of MgsR stability via ClpXP and ClpCP proteolysis (Reder et al., 2012b). In addition, ethanol stress leads to a redox switch and causes intramolecular disulfides in the CXXC motif of MgsR as activation mechanism. However, the detailed mechanism of MgsR interaction with the RNAP containing σB and activation of transcription remains to be elucidated.
The σB general stress regulon is induced after exposure to heat, salt and ethanol stress which causes non-specific resistance to secondary oxidative stress in B. subtilis (Mols and Abee, 2011; Reder et al., 2012b). The involvement of σB in the protection against secondary oxidative stress has been demonstrated by the role of the σB-dependent miniferritin Dps in stationary-phase-induced peroxide resistance (Antelmann et al., 1997). Moreover, several genes of the primary PerR-, OhrR- and SpxA-controlled oxidative stress response (katA, mrgA, ohrA, spxA) have paralogs within the σB regulon (katE, katX, ydbD, dps, ohrB, mgsR) conferring non-specific secondary oxidative stress resistance (Zuber, 2009). Other redox-stress-related genes like trxA are controlled by both, SpxA and σB or by specific regulators, such as clpC that is controlled by σB and CtsR (Zuber, 2009). Moreover, phenotype screening of 94 mutants in σB-controlled genes identified 62 mutants with increased sensitivity towards paraquat or peroxides (Reder et al., 2012a). Thus, B. subtilis employs PerR, OhrR and SpxA as specific antioxidant control mechanisms to cope with ROS and MgsR and σB for protection against primary and secondary generated oxidative stress (Mols and Abee, 2011; Reder et al., 2012b).
Spx as thiol-redox sensor for oxidative stress in the pathogens Bacillus anthracis, S. aureus and Streptococcus
Spx homologs are highly conserved in low GC Gram-positive bacteria, such as Bacillus, Staphylococcus, Streptococcus, Lactobacillus and Listeria species where they play important functions in the oxidative stress resistance and virulence (Zuber, 2004; Pamp et al., 2006; Kajfasz et al., 2010; Wang et al., 2010). In S. aureus, Spx functions as global regulator of genes that maintain the thiol-redox balance, such as the Trx/TrxR system. The spx mutant displays growth defects and is hypersensitive to peroxide and disulfide-stress, heat shock and osmotic stress (Pamp et al., 2006; Wang et al., 2010). Spx controls biofilm development in S. aureus and S. epidermidis through control of the icaABCD operon. The post-translational proteolytic control of Spx by the ClpXP system is similar like in B. subtilis, shown by the increased stability of Spx in S. aureusclpP and clpX mutants (Pamp et al., 2006). In S. aureus, YjbH functions as adaptor for ClpXP-mediated proteolysis of Spx although there is only 30% sequence identity between YjbH homologs of S. aureus and B. subtilis (Engman et al., 2012). The Cys-His rich N-terminal domain is not conserved in S. aureus YjbH and instead a CxC motif is present at another location. The B. subtilisyjbH mutant could be complemented with S. aureusyjbH to restore the Spx level and diamide susceptability to that of wild type cells (Gohring et al., 2011; Engman et al., 2012). Hence YjbH controls Spx proteolysis like in B. subtilis. The yjbH mutant further shows growth-impaired phenotypes and increased pigmentation (Engman et al., 2012), resistance to oxacillin and other β-lactam antibiotics, glycopeptides and sensitivity to desiccation stress (Charbonnier et al., 2005; Chaibenjawong and Foster, 2011; Gohring et al., 2011). This higher resistance to β-lactam antibiotics might be caused by the higher PBP4 level and increased peptidoglycan cross-linking in yjbH mutants, but the Cys residues were not required for the antibiotic resistant phenotype (Gohring et al., 2011). This indicates that YjbH regulates Spx proteolysis and antibiotic resistance mechanisms in S. aureus. Spx also controls trfA, the B. subtilismecA homolog and trfA mutants are sensitive to oxacillin and glycopeptide antibiotics (Jousselin et al., 2013). Expression of trfA was constitutively up-regulated in glycopeptide-intermediate S. aureus (GISA) derivatives of methicillin-susceptible or methicillin-resistant in S. aureus (MRSA) clinical or laboratory isolates (Jousselin et al., 2013). This indicates that the up-regulation of trfA in the yjbH mutant is responsible for the β-lactam antibiotic resistance. In summary, S. aureus YjbH regulates Spx proteolysis and the Spx-dependent trfA confers antibiotic resistance in S. aureus. The role of YjbH for virulence was shown in Listeria monocytogenes where yjbH and clpX mutants had hypohemolytic phenotypes indicating that YjbH and ClpX are required for the function of the pore-forming toxin listeriolysin as virulence factor (Zemansky et al., 2009).
Bacillus anthracis encodes two spx paralogs, spxA1 and spxA2. Mutants lacking spxA1 displayed increased peroxide sensitivity but only the spxA1spxA2 double mutant was hypersensitive to diamide stress suggesting overlapping roles of SpxA1 and SpxA2 in disulfide-stress resistance (Barendt et al., 2013). Microarray analyses identified many genes involved in the thiol-redox homeostasis that were up-regulated when stabilized protease-resistant forms of SpxA1DD or SpxA2DD were produced. These SpxA1 and SpxA2-controlled disulfide-sress related genes encode for thioredoxins and thioredoxin reductases (trxA, trxB, ytpP), two CoASH disulfide reductases (BA1263, BA0774) to keep CoASH in its reduced state and genes for bacillithiol biosynthesis (bshA, bshB1, bshB2, bshC), bacilliredoxins (ytxJ, yphP) and the putative BSH reductase (ypdA) (Table 5). In addition, the Spx paralogs control genes involved in DNA damage repair (uvrC, uvrD), spore outgrowth and germination (exoA), detoxification of alcohols, aldehydes, and quinones (BA0838, BA2647, BA3438), unknown oxidoreductase functions (BA1951) and adapter for ClpXP proteolysis (yjbH) (Barendt et al., 2013). Furthermore, both Spx paralogs control also their own subregulon. Interestingly, spxA2 was shown to be up-regulated in phagocytosis assays with infected macrophages (Bergman et al., 2007). The expression of spxA2 is negatively controlled by the Rrf2 family regulator SaiR that is conserved among the Bacillus cereus group. The Rrf2-family regulators include also IscR and NsrR with 3 conserved Cys residues that coordinate an FeS-cluster. SaiR shares C89 and C96 with IscR and NsrR and C96 was required for SaiR repressor activity and redox regulation of spxA2. Repression of spxA2 is alleviated under NaOCl stress and in infected macrophages probably by thiol-oxidation of SaiR leading to its dissociation from the spxA2 promoter (Nakano et al., 2014). However, the role of the Cys residues and the possible involvements as ligands are still unknown in the regulation of SaiR.
Streptococci also encode two spx paralogs: spxA and spxB in Streptococcus mutans (Kajfasz et al., 2010); spxA1 and spxA2 in Streptococcus pneumoniae (Turlan et al., 2009), Streptococcus suis (Zheng et al., 2014) and Streptococcus sanguinis (Chen et al., 2012). These Spx paralogs share the conserved redox-sensing CXXC motif and also the Gly residue required for Spx-RNAP interaction. However, only SpxA and SpxA1 contain the RPI motif involved in modulation of the reactivity of the CXXC motif. In SpxB and SpxA2, the arginine is replaced by a serine residue and this may affect the sensory function (Newberry et al., 2005; Turlan et al., 2009; Kajfasz et al., 2010; Chen et al., 2012; Zheng et al., 2014). These Spx proteins are involved in the oxidative stress response and in the virulence of streptococci as demonstrated by the use of murine, rabbit or insect infection models (Kajfasz et al., 2010; Chen et al., 2012; Zheng et al., 2014). Both Spx paralogs control the expression of genes for the thioredoxin reductase (trxB), the peroxide resistance protein (dpr), the superoxide dismutase (sod), the peroxiredoxin (ahpCF), the glutathione reductase (gor), the NADH oxidase (nox) and thiol peroxidase (tpx) in S. mutans and S. suis (Kajfasz et al., 2010; Zheng et al., 2014) (Table 5). In S. sanguinis, SpxA1 regulates the genes for the pyruvate oxidase (spxB) and the NADH oxidase (nox) that are both responsible for the high production of H2O2 (Chen et al., 2012). Thus, the spxA1 mutant is more sensitive to H2O2 but produces also lower amounts of cytoplasmic H2O2 as defense mechanism against bacterial competitors.
ECF sigma factors and their cognate redox-sensitive zinc-associated anti-sigma factors (RsrA/SigR in Streptomyces; RshA/SigH in C. glutamicum and M. tuberculosis)
In Actinomycetes, the disulfide-stress response is controlled by ECF sigma factors and their redox-sensitive cognate zinc-containing anti-sigma factors (ZAS) that share a conserved HX(3)CX(2)C (HCC) motif (Jung et al., 2011). In addition, the region K(33)FEHH(37)FEEC(41)SPC(44)LEK(47) that includes the conserved HCC motif was identified as redox-sensitive determinant in ZAS factors. In another model, alternative Zn-binding sites were identified in redox-sensitive ZAS factors that might increase the susceptibility of zinc-coordinating cysteine residues to oxidation (Heo et al., 2013). Redox-sensitive ZAS factors that are involved in the disulfide-stress response include RsrA of S. coelicolor and its homologs RshA of M. tuberculosis and C. glutamicum. RsrA of S. coelicolor is the best studied ZAS factor that coordinates zinc and sequesters its cognate sigma factor SigR under reducing conditions forming a RsrA/SigR complex (Paget et al., 2001). Three of the seven cysteines in RsrA (C11, C41 and C44) are essential for anti-sigma factor activity and redox-sensing (Kang et al., 1999) (Cherney et al., 2003; Zdanowski et al., 2006). Upon disulfide stress, the redox-sensitive Cys11 and Cys44 form an intramolecular disulfide resulting in zinc release and conformational changes in RsrA that lead to free SigR (Kang et al., 1999; Bae et al., 2004) (Figure 9). Free SigR interacts with the RNAP to direct transcription of the SigR disulfide-stress regulon.
The SigR-regulon consists of more than 100 genes as revealed by DNA microarrays and genome-wide ChIP-chip analyses (Kim et al., 2012). The majority of SigR-controlled genes function in the thiol-redox homeostasis and encode for thioredoxins and thioredoxin reductase (trxAB, trxC), enzymes for mycothiol biosynthesis and recycling (mshA, mca, mtr), putative glutaredoxin-like mycoredoxins (mrxA, mrxB) and methionine sulfoxide reductase (msrA, msrB) (Paget et al., 1998, 2001; Newton and Fahey, 2008; Park and Roe, 2008). Targets for SigR control are also involved in protein quality control (pepN, ssrA, clpP1P2, clpX, clpC, lon), in the prokaryotic ubiquitin-like protein-conjugation and proteasomal degradation pathway (pup, pafD, mpa, prcAB), guanine biosynthesis (guaB, guaB1) and ribosome-associated functions (rpmE, rpmI, rplT, relA, engA, obgE, era, truB, rbfA, infA, infB) (Kallifidas et al., 2010; Kim et al., 2012). The SigR-regulon includes further genes that encode Fe-S assembly components (sufA, sufU), Fe-S containing enzymes (nadA, lipA), biosynthesis enzymes for redox-sensitive sulfur-containing cofactors, such as Fe-S, folate, CoASH and lipoic acid (moeB,coaE, folE, lipA), UV damage repair enzymes (uvrA) and the major housekeeping sigma factor (hrdB) (Figure 9, Table 6). This core SigR-regulon and the SigR consensus sequence are conserved across 42 Actinomycetes revealing a robust adaptation strategy to oxidative and disulfide-stress conditions in different natural environments (Kim et al., 2012).
|Redox sensor||Organism||Signal||Redox-sensing mechanism||Regulon genes||Regulon functions||References|
(ZAS of SigR)
|C11, C41,C44 |
Zn-site of ZAS
pepN, ssrA, clpP1P2, clpX, clpC, lon
pup, mpa, pafD, prcAB
rpmE, rpmI, rplT, relA, engA, obgE, era, truB, rbfA, infA, infB
Mycothiol biosynthesis and recycling
Methionine sulfoxide reductase
Protein quality control
Fe-S assembly machinery
Cofactor synthesis (Fe-S, folate, CoASH, lipoic acid)
Housekeeping sigma factor
UV damage repair
|Paget et al. (1998)|
Paget et al. (2001)
Park and Roe (2008)
Kim et al. (2012)
(ZAS of SigH)
|C23, C53, C56|
Zn-site of ZAS
trxB, trxB1, trxC
sigM, hspR, cglR, sufR, whcA, whcE
Mycothiol biosynthesis and recycling
dsbA-like thiol-disulfide oxidoreductases
methionine sulfoxide reductases
UV damage repair
|Ehira et al. (2009b)|
Busche et al. (2012)
Toyoda et al. (2014)
MSH and TrxA were shown to reduce oxidized RsrA, allowing to switch-off of the SigR-dependent stress response upon return to non-stress conditions (Kang et al., 1999; Park and Roe, 2008). SigR and an unstable SigR′ protein with an N-terminal extension are positively autoregulated under diamide stress (Kim et al., 2009). This unstable SigR′ protein is rapidly degraded by the induced ClpP1/P2 proteases which represents another negative feedback loop.
The ECF sigma factor SigH of M. tuberculosis, an ortholog of SigR, plays an important role in the survival against oxidative stress. SigH is activated upon entry of M. tuberculosis into macrophages and has a key role in the response to oxidative stress as sigH mutants of M. tuberculosis and M. smegmatis are highly sensitive to peroxide stress (Fernandes et al., 1999; Manganelli et al., 2002). SigH is sequestered by its cognate redox-sensitive ZAS factor RshA under reduced conditions. Oxidative stress leads to RshA oxidation and release of SigH to activate transcription of its regulon (Song et al., 2003). SigH controls transcription of approximately 30 genes that include the Trx system, but also other ECF sigma factors (SigE and SigB) (Manganelli et al., 2002). Despite the presence of a ZAS motif, RshA has weak affinity for zinc and the cysteines were shown to coordinate an Fe-S cluster. However, neither the cysteines nor the Fe-S cluster in RshA were essential for RshA/SigH interaction, which is mediated by salt bridge suggesting another regulatory mechanism (Kumar et al., 2012). The serine-threonine kinase PknB that is essential for survival within the host, was shown to phosphorylate both SigH and RshA. Phosphorylation of RshA results in disruption of its interaction with SigH leading to increased activity of SigH (Park et al., 2008). These observations suggest an important role of PknB in mycobacterial adaptation to oxidative stress by activating SigH via RshA phosporylation.
SigE is another important ECF sigma factor required for M. tuberculosis survival under oxidative stress and within infected macrophages (Manganelli et al., 2001, 2004a,b). Its expression is regulated by SigH and the two-component system MprAB, which responds to polyphosphate stress and is implicated in the persistence of M. tuberculosis in vivo (He et al., 2006). SigE activity depends on interaction with its anti-sigma factor RseA, a ZAS protein that senses ROS and disulfide stress. This interaction requires Cys70 and Cys73 of the ZAS motif, which is disrupted under oxidative stress conditions. The release of SigE leads to transcriptional activation of sigB and clgR, that controls the ClpC1/P2 system. SigE regulates also itself in a positive feedback loop (Barik et al., 2010).
In C. glutamicum, the ECF sigma factor SigH is involved in the thiol-specific oxidative stress and heat shock response (Kim et al., 2005; Ehira et al., 2009b). SigH is sequestered by its redox-sensitive ZAS factor RshA, which responds to diamide and NaOCl stress (Busche et al., 2012; Chi et al., 2014). The SigH-regulon includes 83 genes that are up-regulated in the rshA mutant most of which are involved in the thiol-redox balance (Busche et al., 2012). SigH transcribes genes for the Trx/TrxR system (trxB, trxB1, trxC), for MSH biosynthesis and recycling (mshC, mca, mtr), dsbA-like thiol-disulfide oxidoreductases (cg2838, cg2661), methionine sulfoxide reductases (msrA, msrB), phage-associated functions (cg0378), the tRNA-(5-methylaminomethyl-2-thiouridylate)-methyltransferase (cg1397), the ubiquitin-like proteasomal conjugation pathway (pup) and DNA damage repair enzymes (uvrA, uvrD3) (Ehira et al., 2009b; Busche et al., 2012) (Table 6). SigH controls expression of its own sigH-rshA operon and that of genes for other stress-responsive regulators (sigM, hspR, cglR, sufR, whcA and whcE) (Ehira et al., 2009b). ChIP-chip analysis of the rshA mutant identified 75 SigH-dependent promoters that included 39 novel promoters not identified in previous transcriptome analyses (Toyoda et al., 2014). In addition, internal σH-dependent promoters within operons were identified that are involved in the pentose phosphate pathway, riboflavin biosynthesis, and Zn-uptake. This resulted in increased riboflavin production and Zn overload phenotypes in the rshA mutant expanding the roles of the SigH-Regulon in C. glutamicum (Toyoda et al., 2014). In addition to SigH, SigM is also involved in the oxidative stress response. Its transcription is increased after disulfide and peroxide stresses and the sigM mutant displayed a high sensitivity to oxidative stress. SigM controls expression of genes encoding the Trx/TrxR systems, the Fe-S-cluster assembly machinery and the disulfide-stress response (Nakunst et al., 2007). These genome-wide studies have revealed an integrated network of disulfide-stress specific ECF sigma factors that can partially replace each other in functions and maintain together the thiol-redox balance among Actinomycetes. In conclusion, the functions of the SigR/SigH regulons in Actinomycetes are very similar to that of the disulfide-stress responsive Spx regulons among Firmicutes bacteria.
Concluding remarks and future challenges
We have provided an overview about thiol-based redox regulators in Gram-positive bacteria and emerging redox sensors in Gram-negative bacteria that belong to different transcription factor families, including LysR(OxyR,HypT), Fur(PerR), MarR(OhrR/SarA/DUF24), TetR(NemR), AraC(RclR), Spx and ZAS factors (RsrA/RshA). Many of these redox sensors employ conserved cysteine residues for redox-sensing of ROS, HOCl or RES that are characterized by their lower Cys pKa value and hence are in the thiolate anion state and susceptable for versatile post-translational thiol-modifications. The redox-sensing mechanisms of many ROS, HOCl- and RES-sensing redox regulators are often variations of this classical disulfide-switch model of E. coli OxyR. The 1-Cys and 2-Cys-type models of the OhrR-family regulators have also been widely confirmed for the widespread MarR/DUF24-family regulators of Firmicutes bacteria and for NemR and RclR as main sensors for RES and HOCl. However, different in vitro models have shown that thiol-based MarR-type regulators can also be inactivated by Cys phosphorylation and Cys alkylation as new themes of thiol-based redox regulation, yet the reversibility of these modifications still needs to be demonstrated. Another question is to what extent thiol-disulfide switches or alternative Cys phosphorylations contribute to the in vivo model of redox regulation. The burgeoning field of redox proteomics coupled with mass spectrometry and new chemical probes to analyze different types of redox-modifications, such as sulfenic acids, S-glutathionylations, S-bacillithiolations, protein disulfides, S-alkylations, S-sulfhydrations, etc., allows a perspective to analyze global thiol-oxidations in the proteome as well as for specific redox regulators under both normal and stress conditions in vivo (Leonard and Carroll, 2011; Thamsen and Jakob, 2011; Paulsen and Carroll, 2013; Zhang et al., 2014). While significant progress has been made to characterize protection mechanisms and redox sensors for reactive electrophiles (quinones and aldehydes) and reactive chlorines (HOCl), the mechanisms for the specificity of the emerging HOCl and RES-sensing redox regulators still remains a future challenge and requires more structural information.
Thiol-oxidations play also an important role in pathogens as they have to cope with ROS in the defense against the host immune system. The correlation between thiol-switches and regulation of virulence and antibiotic resistance is well established for many redox sensors of important human pathogens, including PerR, SarZ, MgrA, SarA, QsrR and Spx of S. aureus and OxyS, MosR and RshA of M. tuberculosis. Particularly, several Spx paralogs have been characterized among pathogenic Firmicutes bacteria that contribute to virulence and to the ROS defense. While the knowledge about virulence regulation by thiol-based redox switches is emerging, the importance of S-thiolations in redox regulation of virulence functions is still unknown and represents an important challenge for future research in pathogenic bacteria.
This work was supported by grants from the Deutsche Forschungsgemeinschaft (DFG) AN746/3-1 and AN746/4-1 within the DFG priority program SPP1710, by the DFG Research Training Group GRK1947, project [C1] and by an ERC Consolidator grant (GA 615585) MYCOTHIOLOME to H.A.
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