Regulatory mechanisms of the disulfide stress response and the role of the bacillithiol redox buffer in Gram-positive bacteria !

!

" I n a u g u r a l d i s s e r t a t i o n

zur

Erlangung des akademischen Grades

doctor rerum naturalium (Dr. rer. nat.)

an der Mathematisch-Naturwissenschaftlichen Fakultät

der

Ernst-Moritz-Arndt-Universität Greifswald

vorgelegt von Bui Khanh Chi geboren am 15.08.1983 in Hanoi, Vietnam

Greifswald, den 16. November 2012" " !" " ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! ! Dekan: Prof. Dr. Klaus Fesser

1. Gutachter: Priv.-Doz. Dr. Haike Antelmann

2. Gutachter: Prof. Dr. Henry A. Claiborne

3. Gutachter:

Tag der Promotion : 01 – 02 – 2013

! "! ! Table of Contents

Chapter 1 Introduction and general conclusion 4

Chapter 2 S-bacillithiolation protects against hypochlorite stress in Bacillus 35 subtilis as revealed by transcriptomics and redox proteomics.

Chapter 3 S-bacillithiolation protects conserved and essential proteins against 57 hypochlorite stress in Firmicutes bacteria.

Chapter 4 Structural insights into the redox-switch mechanism of the 81 MarR/DUF24-family regulator HypR.

Chapter 5 The redox-sensing regulator YodB senses quinones and diamide via 97 a -disulfide switch in Bacillus subtilis.

Chapter 6 The paralogous MarR/DUF24-family repressors YodB and CatR 109 (YvaP) control expression of the catechol dioxygenase CatE in Bacillus subtilis.

Zusammenfassung 121

Eidesstattliche Versicherung 125

Publications and poster presentations 127

Acknowledgements 132

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! ! ! ! ! ! ! ! ! ! Chapter 1

Introduction and general conclusion

Bui Khanh Chi

1. The biological function of Cysteine and its thiol group ...... 5 2. Functions of low molecular weight in bacteria ...... 5 2.1. Biosynthesis and functions of and role of S-glutathionylation ...... 6 2.2. Biosynthesis and functions of and role of MSH-dependent . 9 2.3. Biosynthesis and functions of bacillithiol and role of S-bacillithiolations ...... 11 3. Reactive oxygen, nitrogen and electrophilic species (ROS, RNS, RES) ...... 14 3.1. Sources of ROS, RNS and RES ...... 14 3.2. Reaction of ROS, RNS, RES with protein thiols ...... 16 3.3. Reaction of ROS with Iron-sulfur clusters and DNA damage ...... 17 4. Bacterial defence mechanisms against ROS and RES ...... 18 5. Thiol-based redox sensors for ROS and RES ...... 19 5.1. OxyR as thiol-based redox-sensor of E. coli ...... 20 5.2. PerR as metal-based peroxide sensor of B. subtilis ...... 21 5.3. OhrR as MarR-family thiol-based redox sensor of organic hydroperoxides ..... 22 5.4. MarR/DUF24-family thiol-based sensors of RES ...... 24 5.5. Spx as thiol-based redox sensor for disulfide stress ...... 27 6. Conclusion and future perspectives ...... 28 References ...... 29 !

"! ! ! ! "#$%&'(!)! ! ! Introduction and general conclusion 1. The biological function of Cysteine and its thiol group The amino acid Cysteine (Cys) with its functional thiol group is the most rare amino acid in proteins and plays a key role to determine the structure and functions of proteins. Oxidation of cysteines to disulfides occurs mainly in the extracellular environment or periplasm and links polypeptides to stabilize the protein structure [1]. The biosynthesis of Cys occurs from sulfate in the sulfate assimilation pathway in most bacteria. Cys is used as precursor for the biosynthesis of many sulfur-containing compounds, including methionine, Fe-S clusters, thiamine cofactors, coenzyme A and low molecular weight (LMW) thiols. In most proteins, the thiol group has a pKa above 8 and is present in its protonated form [2]. However, the thiol group of redox-sensitive Cys residues is often highly reactive and present in its de-protonated thiolate anion form (R-S-

) at low pKa values. The reactivity of the thiol group can be determined by surrounding basic amino acids that can lower its pKa value at physiological pH values in the cytoplasm. Furthermore, hydrogen bonds with positively charged amino acids stabilize the thiolate anion. The activities of many redox-sensing enzymes or regulators depends on the reactivity of the thiol group of Cys residues that are mentioned in detail later [2]. Protein thiols are among the most susceptible oxidation-sensitive targets and can be reversibly and irreversibly post-translational modified that has regulatory and metabolic consequences for cellular physiology.

2. Functions of low molecular weight thiols in bacteria Low molecular weight thiols are thiol-containing, non-proteinogenous compounds that are usually smaller than 1 kDa. LMW thiols are often present in millimolar concentrations in the cellular cytoplasm and function to maintain the reduced state of the cytoplasm #$%!&'( This reduced state ensures proper protein functions and helps to avoid damages of cellular components that occur during cellular metabolism or stressful conditions [5]. Life evolved from transition of anaerobic to aerobic conditions and LMW thiols represent a major biological adaptation to oxidative stress conditions as consequence of the aerobic life [6]. The best studied LMW thiol-redox buffer is the tripeptide glutathione (GSH) (Figure 1), predominantly found in eukaryotes and most Gram-negative bacteria [4, 6]. Most Gram-positive bacteria do not produce GSH. The Actinomycetes produce mycothiol

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(MSH) as their LMW thiol-redox buffer (Figure 1) and MSH-deficient mutants are very sensitive to thiol-reactive species and antibiotics that affect the redox balance [7, 8]. In Bacillus megaterium, Bacillus cereus, and Coenzyme A (CoASH) serves as an abundant LMW thiol [9]. Many Firmicutes bacteria, including Bacillus and Staphylococcus species have recently been discovered to utilise bacillithiol (BSH) as their major LMW thiol-redox buffer (Figure 1) [10]. Alternative LMW thiols include also the betaine-histidine derivative ergothioneine present in Actinomycetes and Fungi. For example, ergothioneine was shown to compensate for MSH-deficiency in Mycobacterium smegmatis [11]. There are also several glutathione-derivatives, such as trypanothione

(TSH2) that is the major redox buffer of the protozoa Leishmania and Trypanosoma or the glutathionylspermidine detected in E. coli during the stationary phase [6]. Here, the functions and biosynthesis pathways for the major bacterial redox buffers GSH, MSH and BSH are summarized in the following parts including novel results of our studies about S- bacillithiolations in Firmicutes bacteria. ! ! ! ! ! ! ! ! ! ! ! ! ! ! Figure 1: Structures of low molecular weight (LMW) thiols. Major LMW thiols are glutathione (GSH) in eukaryotes and Gram-negative bacteria, mycothiol (MSH) in Actinomycetes and bacillithiol (BSH) in Firmicutes. Coenzyme A (CoASH) also serves as a LMW thiol-redox buffer in some bacteria and Archaea. The schematic is derived from Antelmann & Hamilton, 2012 [12]. 2.1. Biosynthesis and functions of glutathione and role of S-glutathionylation The best studied redox buffer is the tripeptide y-glutamylcysteinyl-glycine or glutathione (GSH) present in eukaryotes, in Escherichia coli (in concentrations at 3.5 - 6.6 mM), in Gram-negative !- and "-Proteobacteria, and few Firmicutes bacteria, such as

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Streptococcus agalactiae and Listeria monocytogenes [4, 6, 13]. GSH acts as a reducing agent, , and it mediates protein protection by S- glutathionylations. GSH functions in detoxification of xenobiotics, antibiotics, reactive oxygen and nitrogen species (ROS and RNS) and maintains protein thiols in its reduced state [4]. GSH detoxifies xenobiotics, toxic electrophiles and antibiotics by conjugation either spontaneously or by the catalytic activity of GSH-S-. The conjugates are usually excreted from the cell. GSH also serves also as a reservoir for cysteine [14]. The biosynthesis of GSH occurs in two steps (Figure 2A). The !-glutamate cysteine catalyzes the formation of !-glutamylcysteine from L-glutamate and L- cysteine. In the second step, ligation of glycine to !-glutamylcysteine is catalyzed by glutathione synthase. The !-glutamyltranspeptidase is able to hydrolyse the gamma carbon peptide bond of GSH and then transfers the !-glutamyl moiety to amino acids [15]. In Listeria species, Streptococcus agalatiae and other Gram-positive bacteria, a bifunctional fusion protein encoded by gshAB exhibits both !-glutamate cysteine ligase

2 H. Antelmann and C.and J. Hamilton glutathione᭿ synthase activity [16].

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Figure 2: Biosynthesis of GSH (A) and summary of redox regulation of protein disulfides by the TrxAB and Grx/GSH/Gor systems in E. coli (B). A) Biosynthesis of GSH by !-glutamylcysteine ligase (GshA) and glutathione synthetase (GshB). The schematic is derived from Fahey (2012) [6]. B) In the GSH-utilising Gram-negative bacterium E. coli, proteins with intramolecular disulfides are commonly reduced by thioredoxins (TrxA) and S-glutathionylated proteins are reduced by glutaredoxins (Grx), respectively. The NADPH-dependent thioredoxin reductase (TrxB) reduces oxidized TrxA using NADPH as electron source. The oxidized Grx is reduced by GSH to give GSSG, which is then recycled back to GSH by glutathione reductase (Gor) at the expense of NADPH. Evidence for analogous mycoredoxin and bacilliredoxin pathways is now beginning to emerge from studies of MSH- and BSH- utilising Gram-positive bacteria. The schematic is derived from Antelmann & Hamilton, 2012 [12]. Glutathione modulates protein functions by forming mixed disulfides with protein thiols under oxidative stress conditions (S-glutathionylations). S-glutathionylation is a post-translational thiol-modification that acts as protein protection mechanism against

Fig. 1. Structures of low molecular weight (LMW) thiols (A); mechanisms for ROS- and RNS-induced protein S-thiolations (B); summary of redox regulation of protein disulphides by the TrxAB and Grx/GSH/Gor systems" in!E. coli (C); Mrx1 mechanisms for reduction of MSH-mixed disulphides and arsenate! (D). In A–D, reduced proteins/substrates are coloured blue and oxidized are labelled red. A. Major LMW thiols are glutathione (GSH) in eukaryotes and Gram-negative bacteria, mycothiol (MSH) in Actinomycetes and bacillithiol (BSH) in Firmicutes. Coenzyme A also serves as a LMW thiol redox buffer in some bacteria and archaea. B. Cysteine thiols are activated first to reactive thiol intermediates: by ROS to Cys sulphenic acids (R-SOH), by HOCl to sulphenylchlorides (R-SCl), by RNS to S-nitrosothiols (R-SNO). These intermediates then react further with proximal protein thiols to form inter- or intramolecular disulphides or with LMW thiols to form protein RSH-mixed disulphides (S-thiolations) as shown here. C. In the GSH-utilizing Gram-negative bacterium E. coli, proteins with intramolecular disulphides are commonly reduced by thioredoxins (TrxA) and S-glutathionylated proteins are reduced by glutaredoxins (Grx) respectively. The NADPH-dependent thioredoxin reductase (TrxB) reduces oxidized TrxA using NADPH as electron source. The oxidized Grx is reduced by GSH to give GSSG which is then recycled back to GSH by glutathione reductase (Gor) at the expense of NADPH. Evidence for analogous mycoredoxin and bacilliredoxin pathways is now beginning to emerge from studies of MSH- and BSH-utilizing Gram-positive bacteria. D. Reduction of MSH-mixed disulphides and arsenate As(V) by Mrx1 via the MSH/Mtr/NADPH pathway. The S-mycothiolated substrate is attacked by the solvent-exposed Cys14 of Mrx1, resulting in Mrx1–SSM intermediate formation and reduction to protein SH. In the second step, the Mrx1–SSM is reduced by MSH, leading to MSSM formation that is recycled by the mycothione reductase Mtr at expense of NADPH. Arsenate [As(V)] detoxification is catalysed by ArsC1 and ArsC2 that form the As(V)–SM adduct from As(V) and MSH. The arseno-thiol bond is reductively cleaved by Mrx1 resulting in the Mrx1–SSM intermediate that is reduced by the MSH/Mtr/NADPH pathway. role of emerging S-thiolation mechanisms in bacteria is nisms implicated in S-thiolations require activation of Cys summarized, which puts into context the structural and thiols to reactive Cys oxidation intermediates, such as mechanistic studies of mycoredoxin-1 (Mrx1) as eluci- sulphenic acid, S-nitrosyl or sulphenylchloride intermedi- dated in this issue by Van Laer et al. (2012). ates (Fig. 1B) (Hawkins et al., 2003; Gallogly and Mieyal, 2007; Allen and Mieyal, 2012). In eukaryotes, Physiological roles of S-glutathionylation and S-glutathionylation controls redox-sensing transcription S-bacillithiolation in bacteria factors and protects active site Cys thiols against overoxi- dation (Dalle-Donne et al., 2009). S-glutathionylation has Protein S-glutathionylation is the reversible oxidation of been shown to control energy metabolism, protein synthe- protein thiols to mixed disulphides with GSH. The mecha- sis, redox balance, calcium homeostasis, cytoskeletal

© 2012 Blackwell Publishing Ltd, Molecular Microbiology ! ! "#$%&'(!)! ! ! irreversible oxidation and controls numerous physiological processes, such as cellular growth and differentiation, cell cycle progression, transcriptional activity, cytoskeletal function and cellular metabolism [14, 17]. Under oxidative stress, metabolic enzymes, such as methionine synthase, PAPS reductase and glyceraldehyde-3-phosphate dehydrogenase are reversibly inactivated by S-glutathionylation in E. coli [18-20]. The reduction of protein disulfides is catalyzed by the glutaredoxin (Grx)/GSH/GSH reductase (Gor) and/or thioredoxin (Trx)/thioredoxin reductase systems [21, 22] (Figure 2B) [12]. The Trx and Grx systems can functionally substitute for each other and either the Trx or the Grx system is essential for disulfide reduction and viability in E. coli [23]. The Trx/Trx reductase system catalyzes reduction of intermolecular and intramolecular protein disulfides. Trx proteins have a conserved CGPC and are ubiquitous present in all kingdoms of life. TrxR reduces oxidized Trx using NADPH as electron source. The Grx/GSH/Gor system is more specific for de-glutathionylation of protein GSH-mixed disulfides. Several Grx isoforms have been characterized in mammals, plants, yeast, protozoa and bacteria [24, 25]. Grx were first discovered in E. coli [26] where they have important functions as electron donors for ribonucleotide reductase (RNR), adenosine-5`-phosphosulfate (APS) reductase, 3'-phosphoadenosine-5'-phosphosulfate (PAPS) reductase and arsenate reductases [27, 28]. Grx are structurally classified into the classical di-thiol Grxs with a CPTC redox active site and the monothiol Grxs containing a CGPS redox active site [24]. In E. coli, three di-thiol Grx proteins (Grx1, Grx2 and Grx3) and one monothiol protein (Grx4) have been characterized. The de-glutathionylation by Grx enzymes involves thiol-disulfide exchange reactions with GSH via nucleophilic double displacement (ping-pong) mechanisms and occurs via mono- or di-thiol mechanisms. Most di-thiol Grx use monothiol mechanisms that take place in two steps: In the first step, the nucleophilic thiolate anion attacks the S- glutathionylated substrate protein, resulting in reduction of the mixed disulfide and the S- glutathionylated Grx (Grx-SSG) intermediate (Figure 2B). This Grx-SSG intermediate is regenerated by GSH in the second step, leading to glutathione disulfide (GSSG) formation. The glutathione disulfide reductase reduces GSSG at expense of NADPH to restore the GSH/GSSG redox balance [25]. The di-thiol mechanism involves a second active site Cys that forms an intramolecular disulfide to resolve the Grx-SSG intermediate that has been shown for some plant Grx enzymes [29]. However, this di-thiol mechanism

"! ! ! ! "#$%&'(!)! ! ! of Grx is less efficient for protein de-glutathionylation and more involved in the reduction of intermolecular protein disulfides [24, 25]. In mammalian cells, Grx 1 is responsible for most de-glutathionylation activity while Grx2 is less efficient and functions in Fe-S cluster homeostasis. Grx2 is bound in large part as inactive dimer to a Fe2-S2 iron sulfur cluster and it is suggested that monomeric Grx2 dissociates under oxidative stress conditions resulting in enhancement of its de-glutathionylase activity [30, 31]. The crystal structure of poplar GrxC1 revealed that the iron-sulfur cluster of the Grx dimer is coordinated by two active site Cys residues of both monomers and two GSH molecules [32]. Binding of Fe-S-clusters, haeme and metallation of other iron enzymes has been demonstrated for yeast monothiol glutaredoxins that function together with the iron chaperone-like Poly(rC)-binding proteins [33].

2.2. Biosynthesis and functions of mycothiol and role of MSH-dependent enzymes Mycothiol (MSH) consists of N-Acetyl-Cys-GlcN-myoinositol (Figure 1) and is present in millimolar concentrations in high-GC Gram-positive Actinomycetes, such as Streptomycetes and Mycobacteria species [7, 8]. Since the Cys amino and carboxyl groups are blocked, MSH is much less susceptible to autoxidation than free Cys [7]. Biosynthesis of MSH proceeds from myo-inositol-1-phosphate, UDP-GlcNAc and cysteine, and occurs in five steps (Figure 3) [7, 8]. The glycosyltransferase MshA conjugates myo-inositol-1-P and UDP-GlcNAc to GlcNAc-Ins-P. Dephosphorylation of GlcNAc-Ins-P by phosphatase MshA2 generates GlcNAc-Ins, which is the substrate for the deacetylase MshB that removes the of GlcNAc-Ins. MshB is a homolog of the MSH S-conjugate amidase (Mca), and has both deacetylase and amidase activities. The MshC cysteine ligase that is homologous to Cys-tRNA synthase adds a Cys residue to GlcN-Ins. Finally, MshD transfers an acetyl group from acetylCoA to Cys-GlcN-Ins. Deficiency in MshA, MshC and MshD activities results in a lack of MSH biosynthesis. MSH biosynthesis is controlled by the transcriptional factor !R or its homologs in Actinomycetes [34] that is sequestered by the redox-sensitive anti sigma factor RsrA in non-stressed cells. Under disulfide stress conditions, RsrA is oxidized in its Zn- that leads to relief of !R. Free !R transcribes genes required to maintain the thiol-redox homeostasis including the genes for TrxAB and for MSH biosynthesis, such as mshA, mshB, mshC, mshD, and mca [35-37].

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(a) OH OH (i) MshA O ATP O UDPGlcNAc HO HO MshD MshB Cys S-conjugation HO HO CoASAc SM (ii) MshA2 R = H MshC MSH MS-transferases RHN GlcN-Ins HN (phosphatase) OIns OIns O MshC Mca R = Ac HS L-myo-Ins-1-P MshD CO H GlcNAc-Ins NH2 2 export S from Cys-GlcN-Ins GlcN-Ins + NHAc cell

(b) OH OH ATP, PPi Cys AMP S-conjugation HO O N-acetyl HO O BS-transferases?? L- UDP-GlcNA c HO HO BSH SB Glycosyltransferase AcHN (BshB) H N Cysteine O 2 O Ligase (BshA) Bca (BshC) CO2H HO2C CO2H HO2C CO2H S = elect rophilic xenobiotic GlcNAc-Mal GlcN-Mal NH2

Figure 3: Parallels between the pathways for MSH biosynthesis (a) and BSH biosynthesis (b) and detoxification of RES by Mca and Bca. (a) MSH biosynthesis requires the MshA, a dephosphorylation step (MshA2), the deacetylase MshB, the Cys adding enzyme MshC and a final N-acetylation of cysteine (MshD). (b) BSH synthesis occurs in three steps catalyzed by the BshA glycosyltransferase, the N-acetylhydrolase BshB, and BshC as cysteine-adding enzyme. MSH and BSH react with RES under catalysis by S-transferases. The resulting conjugate (MSR) is cleaved by mycothiol S-conjugate amidase (Mca or Bca) to generate the mercapturic acid AcCys that is exported from the cell. The figure is adapted from [38].

MSH mutants are very sensitive to ROS, RNS, alkylating agents, redox cycling agents and antibiotics (rifamycin, streptomycin, erythromycin and azithromycin)[39]. Thus, MSH serves analogous functions to GSH in toxin and oxidant detoxification in Actinomycetes. Mycothiol forms conjugates with xenobiotic compounds either spontaneously or using MSH S-transferases, such as the recently identified DinB superfamily proteins [40]. MSH conjugates are transformed to their mercapturic acid derivatives by MSH-S-conjugate amidase (Mca) before excreted from the cell. Mca is the major detoxification enzyme for antibiotics in Actinomycetes [7, 40]. The alcohol dehydrogenase MscR (MSNO reductase/formaldehyde dehydrogenase) is another MSH- dependent detoxification enzyme required for detoxification of formaldehyde and S- nitrosyl-mycothiol (GSNO) [7]. MSH reacts with formaldehyde producing MS-CH2OH that is converted to formate by MscR. MscR also converts MSNO to MSH sulfinamide

(MSONH2). In addition, MSH functions in the metabolism of 3-hydroxybenzoate and acts as cofactor for maleylyruvate in the isomerization of maleylpyruvate to fumarylpyruvate [41]. MSH is oxidized to MSH disulfide (MSSM) under oxidative stress conditions. The mycothiol disulfide reductase Mtr maintains MSH in its reduced state with expense of NADPH. Recently, it was shown that the MSH/Mtr/NADPH electron pathway provides the reducing power for mycoredoxin-1 (Mrx1) in reduction of mixed MSH- disulfides formed under oxidative stress conditions [12, 42]. NMR time course

"#! ! ! ! "#$%&'(!)! ! ! experiments have also demonstrated the transient S-mycothiolation of the active site Cys14 of oxidized Mrx1 during reduction by the MSH/Mtr/NADPH electron pathway. Thus, future studies are directed to identify S-mycothiolated Mrx1 substrates and the function of MSH in redox-regulation and virulence in Mycobacterium tuberculosis.

2.3. Biosynthesis and functions of bacillithiol and role of S-bacillithiolations Bacillithiol (BSH) is composed of Cys-GlcN-malate and serves as major LMW thiol in most Bacillus and Staphylococcus species as well as in Deinococcus radiodurans [10]. BSH confers resistance to hypochlorite and to the epoxide antibiotic in B. subtilis although it is of 10 - 20-fold lower abundance than GSH in E. coli (Figure 1) [38]. The BSH biosynthesis parallels that of MSH biosynthesis and occurs in three steps (Figure 3). In the first step, UDP-GlcNAc is coupled by a glycosyltransferase BshA to L- malic acid to generate GlcNAc-Mal [38, 43, 44]. Subsequent deacetylation of GlcNAc- Mal is performed by a N-acetyl hydrolase (BshB1). BshB1 has a paralog BshB2 that also has deacetylase activity. The functional redundancy of BshB1 and BshB2 suggests that one of these deacetylases might function as BSH-S-conjugate amidase in detoxification of RES likewise to the mycothiol-S-conjugate amidase Mca [44]. The last step of BSH biosynthesis involves the YllA (BshC) protein that presumably adds Cys to GlcN-Mal intermediate. Interestingly, strains of the S. aureus NCTC8325 lineage harbour natural yllA null mutations and do not produce BSH [38, 45]. The availability of bsh mutant strains provided also insights into the physiological role of BSH. The bshA and bshB1 genes belong to a large operon of seven genes including mgsA encoding methyglyoxal synthase suggesting that BSH could function in methylglyoxal detoxification. The bshB2 and bshC genes belong to two different operons that are Spx-dependently induced by diamide and hypochlorite stress [46, 47]. BSH null mutants are only slightly more sensitive to diamide, methylglyoxal and ROS (paraquat, H2O2) [38]. This suggests that cysteine and/or CoA can replace BSH as redox buffer. However, bsh mutants display most sensitive growth phenotypes towards the antibiotic fosfomycin [38] and the strong oxidant hypochlorite [46]. The sensitivity of bsh mutants towards fosfomycin depends on the fosfomycin resistance protein FosB that requires BSH as cofactor and conjugates BSH to the epoxide antibiotic fosfomycin to open the ring structure for its detoxification (Figure 4). FosB is specific for BSH as thiol cofactor and does only poorly work with Cys and its biochemical activity has been recently demonstrated in various Bacillus and Staphylococcus FosB homologs [48].

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Figure 4: BSH-dependent detoxification of the fosfomycin antibiotic by the FosB S- in Bacillus and !"# Staphylococcus. The figure is adapted from Lamers et al., (2012).

In addition, the DinB superfamily S-transferase YfiT of B. subtilis catalyzed the conjugation of BSH to monochlorobimane in vitro [40]. The yfiT gene is flanked by the yfiS and yfiU genes encoding putative efflux transporters of mercapturic acids produced during electrophile detoxification. BSH also functions in the detoxification of the antibiotic rifamycin in S. aureus Newman and rifamycin exposure resulted in increased BSH biosynthesis [45]. It was suggested that the BshB2 deacetylase and its homolog NWMN_0530 of S. aureus could function as BSH S-conjugate amidases. BSH could function in metal ion homeostasis based on the Cys and malate moiety that may chelate Zn2+ or Cu2+ ions [49]. BSH is further involved in detoxification of toxic metal ions, such 2- as selenite SeO3 [49]. In E. coli, GSH has been shown to play a role in direct detoxification of hypochlorite, leading to the oxidation of GSH to GSSG [4]. Hypochlorite is an active ingredient of household-bleach and is also produced by myeloperoxidase from activated macrophages during the infection process. HOCl reacts very fast with cysteine residues 7 -1 -1 (K2=3x10 M s ) and rapidly lead to irreversible oxidation products, such as sulfinic and sulfonic acids [50]. Our study showed that BSH protects cells against NaOCl toxicity by direct detoxification of the oxidant resulting in oxidized BSH disulfide (BSSB) formation. In addition, we have shown for the first time that BSH forms mixed disulfides with protein thiols, termed as S-bacillithiolations that are analogous to S-glutathionylations in eukaryotes (Figure 5) [46, 51]. S-bacillithiolation protects protein thiols or controls the activity of redox-sensing transcription factors, such as OhrR [46, 52]. S-bacillithiolation of OhrR leads to inactivation of the OhrR repressor and up-regulation of the thiol- dependent OhrA peroxiredoxin to protect cells against CHP and NaOCl [46, 53]. S- bacillithiolation is also widespread among other Firmicutes with 8 common and 29 unique S-bacillithiolated proteins identified in B. subtilis, Bacillus amyloliquefaciens, Bacillus pumilus, Bacillus megaterium and Staphylococcus carnosus [46, 51]. The S- bacillithiolome contains mainly biosynthetic enzymes for amino acids (methionine,

"#! ! ! ! "#$%&'(!)! ! ! cysteine, branched chain and aromatic amino acids), cofactors (thiamine), nucleotides (GTP); as well as translation factors, chaperones, redox and antioxidant proteins [46, 51]. The methionine synthase MetE is the most abundant S-bacillithiolated protein in Bacillus species after NaOCl exposure. S-bacillithiolation of MetE occurs at its Zn- binding active site Cys730, leading to methionine starvation in NaOCl-treated cells [46, 51]. Similarly, methionine auxotrophy is caused by S-glutathionylation of MetE in E. coli after diamide stress [54]. Since formyl methionine is required for initiation of translation, MetE inactivation could stop translation during the time of hypochlorite detoxification. This translation arrest caused by S-bacillithiolation is supported by the strong repression of the stringent response RelA regulon by NaOCl stress, which includes genes encoding ribosomal proteins and translation factors [46].

Figure 5: Physiological roles of S-bacillithiolations in B. subtilis and S-glutathionylations in E. coli. NaOCl leads to S-bacillithiolation of OhrR, MetE, YxjG, PpaC, SerA and YphP in B. subtilis. NaOCl stress causes oxidation of BSH to BSSB and the BSH/BSSB redox ratio is 2-fold decreased [46]. S- bacillithiolation of OhrR inactivates the repressor and causes induction of the OhrA peroxiredoxin that likely function in detoxification. S-bacillithiolation of the methionine synthases MetE and YxjG in its active site Zn-center at Cys730 leads to methionine auxotrophy to stop translation during the time of NaOCl detoxification. MetE is also inactivated by S-glutathionylation of Cys645 in E. coli by diamide stress [54]. S-bacillithiolation of the inorganic PpaC could lead to decreased ATP sulfurylase activity. S-bacillithiolation of the phosphoglycerate dehydrogenase SerA causes decreased serine levels required for cysteine and methionine biosynthesis. The thiol-disulfide isomerase YphP could function as bacilliredoxin in reduction of S-bacillithiolated proteins. This figure is adapted from [46].

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The identification of so many S-bacillithiolated proteins raises questions about the regulation of de-bacillithiolation process. Phylogenomic profiling identified three Trx- like proteins YtxJ, YqiW and YphP as putative bacilliredoxins (Brx) that are present in all BSH-producing bacteria. YtxJ is perhaps a monothiol Brx containing a TCPIS motif, and YphP and YqiW are paralogs of the DUF1094 family!(53% identity) with unusual CXC redox motifs [38]. YphP was shown to display weak disulfide isomerase activity in vitro [55]. Mass spectrometry identified all three putative Brx proteins as S-bacillithiolated at their active sites in B. subtilis and S. carnosus by NaOCl stress [46, 51]. The observed S- bacillithiolation of YphP and YtxJ during NaOCl stress could represent an intermediate in a bacilliredoxin redox pathway. Our preliminary results already suggest that YphP and YqiW function in de-bacillithiolation of MetE-SSB in vitro (our unpublished data). However, thus far the BSSB reductase that reduces BSSB and keeps BSH in a reduced state is unknown in Bacillus species.

3. Reactive oxygen, nitrogen and electrophilic species (ROS, RNS, RES) 3.1. Sources of ROS, RNS and RES The cytoplasm of most cells is a reducing environment and protein thiols are maintained in their reduced state by the above-mentioned LMW thiol-redox buffers and enzymatic thiol-disulfide reducing systems. Reactive oxygen species (ROS), reactive nitrogen species (RNS) and reactive electrophilic species (RES) affect the intracellular redox balance, resulting in thiol-specific stress responses. These reathctive species are generated during respiration or cellular metabolism in bacteria or exposed externally by xenobiotics, antibiotics, as part of the host immune defence or by competing bacteria in microbial communities [56]. ROS are generated by stepwise one-electron-transfer reactions to molecular oxygen during respiration as unavoidable consequence of the ! aerobic life. Incomplete reduction of O2 leads to generation of superoxide anion O2! , hydrogen peroxide H2O2 and the highly reactive hydroxyl radical (HO!) (Figure 6). ! Superoxide anion O2! and H2O2 can be also generated by autoxidation of flavoenzymes ! [57]. Superoxide dismutases (SOD) rapidly convert O2! to H2O2. Reactions of H2O2 with 2+ Fe ion in the Fenton reaction generate the hydroxyl HO!.

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Figure 6: Reactive oxygen species (ROS) are generated during respiration by incomplete reduction of molecular oxygen. ROS include superoxide anion, hydrogen peroxide and the highly reactive hydroxyl radical generated in the Fenton reaction. ! The oxidative burst from activated neutrophils also produces O2! , H2O2, nitric oxide (NO), and hypochlorite (HOCl) with the aim to kill the invading pathogenic bacteria [58, 59]. The neutrophil NADPH oxidase is activated and generates superoxide anion in the phagosomal lumen. Myeloperoxidase is released upon degranulation in millimolar concentrations that can dismutate superoxide to H2O2 and catalyzes efficient conversion of hydrogen peroxide with chloride to hypochlorous acid [60]. Reactive nitrogen species (RNS) are also produced by neutrophils that include for example nitric oxide (NO) produced by nitric oxide synthases (NOS) through the oxidation of L-arginine. ! ! Reaction of NO with O2! generates peroxynitrite (ONOO ). Reactive electrophilic species (RES) species are redox-active compounds that have electron-deficient centres (Figure 7). They include quinones, aldehydes, diamide and !,"-unsaturated dicarbonyl compounds [56].

Figure 7: Reactive electrophilic species (RES) include quinones and aldehydes that have partial positive charges (#+). In quinones and aldehydes the electrons are drawn to carbonyl oxygen. Diamide is an electrophilic azocompound. RES can be xenobiotic compounds or generated as secondary reactive metabolites by oxidation of endogenous biomolecules (amino acids, lipids, carbohydrates). Quinones are also the lipid electron carrier of the respiratory chain and the !,"-unsaturated aldehyde methylglyoxal is produced as by-product of the glycolysis. One electron reduction of quinones results in the highly reactive semiquinone radical that can lead to ! generation of ROS, such as O2! . Quinones react with protein thiols via the S-alkylation chemistry and were shown to deplete protein thiols in the proteome due to irreversible protein aggregation [61]. Important lipid-derived RES produced in eukaryotic cells are malondialdehyde (MDA) and 4-hydroxy-2-nonenal (HNE) [62]. HNE is generated from

"#! ! ! ! "#$%&'(!)! ! ! polyunsaturated fatty acids by a peroxidation chain reaction that is readily triggered by ROS [63]. However, since bacteria lack unsaturated fatty acids in their membrane lipids, the source of lipid peroxidation products is unclear in bacteria.

3.2. Reaction of ROS, RNS, RES with protein thiols ROS, RES and RNS can damage all cellular macromolecules leading to protein and DNA damage, point mutations and cell death [64]. On the other hand, in eukaryotes ROS and RNS have been shown to act as second messengers to modulate signal transduction pathways under sub-lethal oxidative stress conditions. All amino acids can react with ROS, but some side chains are more susceptible than others. The amino acids lysine, arginine, histidine, proline and threonine are sensitive to metal-catalyzed oxidation. Oxidation of lysine and arginine side chains in proteins results in protein carbonylation as marker for protein oxidation [62]. The thiol group of cysteine is the strongest nucleophile, and thus the main target for ROS, RNS and RES. The thiol group can be reversibly oxidized to protein disulfides or irreversibly overoxidized to sulfinic or sulfonic acids by ROS, S-nitrosylated by RNS or S-alkylated by RES [56]. Initially, protein thiols undergo oxidation by ROS to reactive Cys sulfenic acids, which are unstable intermediates (R-SOH) (Figure 8). Cys sulfenic acid rapidly reacts further with other thiols to form intramolecular, intermolecular disulfides or mixed disulfides with the LMW thiols GSH, MSH or BSH, termed as S-thiolations. S-thiolations protect thiol groups against the irreversible oxidation stages of cysteine to the Cys sulfinic

(R-SO2H) and sulfonic acid (R-SO3H).

Figure 8: ROS cause disulfide bond formation in redox-sensing proteins. Reversible thiol-oxidation leads first to a Cys sulfenic acid intermediate (R-SOH) that is unstable and reacts further to form intramolecular and intermolecular disulfides or mixed disulfides with LMW thiols, such as glutathione, bacillithiol, cysteine or CoASH, termed as S-thiolations. Cys sulfinic acids of eukaryotic 2-Cys peroxiredoxins can be reversed by sulfiredoxins that are not present in bacteria [65]. Hyprochlorite (HOCl) is a strong oxidant and chlorinating agent. HOCl targets most strongly sulfur-containing amino acids

"#! ! ! ! "#$%&'(!)! ! ! cysteine and methionine with the second-order rate constants of k = 3.0 x 107 M-1s-1 and k = 3.8 x 107 M-1 s-1, respectively [66]. HOCl reacts with Cys thiols to the unstable sulfenylchloride intermediate that reacts further with proximal thiols to form disulfides or the thiol group is overoxidized in the absence of other thiols. When B. subtilis and other Bacillus species were treated with the strong oxidant HOCl, numerous cytoplasmic proteins were oxidized to S-bacillithiolated proteins as major protection mechanism in Firmicutes [46, 51]. Reversible thiol-modifications caused by RNS, such as nitric oxide (NO) and ! peroxinitrite (ONOO ), include S-nitrosothiol (RS-NO) and S-nitrothiol (RS-NO2) respectively. Alternatively, S-nitrosothiol (e.g. GSNO or MSNO) can be formed by direct reaction of NO with LMW thiols [56]. RES including quinones and aldehydes have two modes of action, an electrophilic and oxidative mode. As electrophiles, quinones and aldehydes react with protein thiols and LMW thiols via the S-alkylation chemistry. In the oxidative mode, quinones are incompletely reduced to semiquinone radical that leads to superoxide anion generation [61]. We found that the MarR/DUF24-family transcriptional regulators HypR, YodB and CatR form intermolecular disulfides upon diamide, hypochlorite and quinone exposure in B. subtilis indicative for the oxidative mode of quinones [67-69]. The S-alkylation chemistry was shown in response to toxic quinone concentrations previously [61]. The lipid-derived RES, including malondialdehyde (MDE) and 4-hydroxy-2-nonenal (HNE) can react with DNA bases, protein thiols leading to damages of cellular membranes.

3.3. Reaction of ROS with Iron-sulfur clusters and DNA damage Further targets for ROS and RNS are iron, zinc and iron-sulfur cluster metal centres of redox enzymes. Many Fe-S-cluster containing proteins are important metabolic enzymes, such as dehydratases, SAM superfamily enzymes and ribonucleotide reductase. Iron-sulfur clusters transfer electrons, bind substrates and participate in catalytic reactions 2+ ! [70]. Solvent-exposed [4Fe-4S] clusters can be attacked by radicals. O2! and H2O2 oxidize the [4Fe-4S]2+ to an unstable [4Fe-4S]3+ intermediate, which is degraded to a [3Fe-4S]+ cluster. This process releases a Fe2+ ion and inactivates the enzyme. NO reacts with Fe-S clusters forming nitrosyl and dinitrosyl adducts. Cysteine residues, which coordinate the cluster, release the damage Fe-S cluster and increase the level of Fe2+ in the cytoplasm [64]. Under oxidative and nitrosative stress, intracellular Fe-level is increased due to the oxidation of Fe2+ in the Ferric uptake repressor Fur that inactivates

"#! ! ! ! "#$%&'(!)! ! !

Fur [71]. High Fe2+ concentration under oxidative stress elevates ROS toxicity by catalysing the Fenton reaction that generates hydroxyl radicals. Hydroxyl radicals react with all biological macromolecules, including proteins, lipids, carbohydrates and nucleotides. Hydroxyl radical can cause oxidation of all four DNA bases and especially guanine bases are oxidized to 8-oxoguanine that causes AT-transversion upon replication and consequently leads to point mutations. 8-oxoguanine is the most common base modification and is often measured as an index of oxidative DNA damage [72].

4. Bacterial defence mechanisms against ROS and RES To maintain the thiol-redox balance, cells employ enzymatic and non-enzymatic antioxidant mechanisms to eliminate ROS, RNS and RES. In response to oxidative stress, cells increase expression of ROS scavenging systems, alter metal homeostasis to reduce free Fe2+, induce expression of thiol-disulfide systems and cysteine biosynthesis to restore the thiol redox balance. LMW thiols coupled with thiol-disulfide reduction systems can detoxify directly the reactive species. Specific antioxidant enzymes ! scavenge ROS, such as O2 and H2O2. Antioxidant enzymes are conserved among aerobic bacteria and eukaryotes. Major ROS scavenging enzymes include the superoxide diutase ! (SOD) that catalyzes the dismutation of O2 to H2O2 as well as catalases and peroxidases that catalyze the conversion of hydroperoxide to H2O and O2 (Figure 9) [73-75]. In E. coli, the manganese- and iron-cofactor-containing SOD (Mn-SOD and Fe-SOD) are present in the cytoplasm whereas the Cu-SOD is active in the periplasm [74]. However, some bacteria only contain Mn-SOD or Fe-SOD. Some obligate aerobic bacteria lack ! SOD, but use instead superoxide reductases as O2 scavengers [74]. The efficient activities of peroxidase and catalase keeps the intracellular H2O2 concentration at approximately 20 nM in E. coli [76]. Peroxiredoxin (Prx) is the primary scavenger of

H2O2 under physiological conditions. There are three subgroups of peroxiredoxins (Prx): typical two-Cys Prx, atypical 2-Cys Prx and 1-Cys Prx [77]. They are classified based on the number and position of Cys residues that participate in catalysis. All Prx have a conserved peroxidatic Cys that is oxidized by H2O2 to Cys sulfenic acid. In one-Cys Prx, Cys sulfenic acid reacts further with a LMW thiol to generate S-thiolated Prx. The typical two-Cys Prx forms an intermolecular disulfide between the peroxidatic Cys and the C- terminal Cys of the other subunit of the dimer. In the atypical two-Cys Prx, an intramolecular disulfide is formed between the peroxidatic Cys and the C-terminal Cys of the same subunit. Prx is reduced either by AhpF or Trx reductase [77]. GSH-utilizing

"#! ! ! ! "#$%&'(!)! ! ! bacteria also have glutathione peroxidase (GPX), which reduces H2O2 to O2 using GSH as electron donor that is oxidized to GSSG (Figure 9) [75]. In eukaryotes, GPX activity depends on the selenocysteine as the active site residue [62]. Gpx appears the primary peroxide scavenger in eukaryotes at low H2O2 concentrations [75]. When intracellular

H2O2 concentration increases, peroxidase activity is reduced, and catalase is strongly induced as main peroxide scavenger for high H2O2 concentrations. The haeme-based enzyme catalase has an extremely high peroxide turnover rate. One molecule of catalase can convert millions molecules of H2O2 to H2O and O2 per minute. There are two classes of catalases: the hydroperoxidase II with catalase activity and the hydroperoxidase I with both catalase and peroxidase activities [78]. Other enzymatic systems specifically detoxify RES, such as quinones, diamide or aldehydes. NAD(P)H-dependent quinone , nitroreductases and azoreductases catalyse the two electron reduction of quinones and diamide to redox stable hydroquinones and dimethyurea derivatives [79]. Thiol-dependent glyoxalases and aldehyde dehydrogenases catalyze the detoxification of reactive aldehydes, such as methylglyoxal and formaldehyde to lactate and formate, respectively [80].

Figure 9: Detoxification systems for ROS. Catalases, peroxiredoxins and glutathione peroxidases are the main peroxide scavenging systems and superoxide dismutase converts superoxide anion to H2O2. 5. Thiol-based redox sensors for ROS and RES ROS and RES are sensed by redox-sensing transcriptional regulators that often have conserved Cys residues that are susceptible to various post-translational thiol- modifications [56]. These lead to conformational changes allowing the transcription factor to activate or repress transcription of the genes for detoxification. This thesis focuses on mechanisms of redox-sensing regulators of the MarR/DUF24-family of B. subtilis that sense ROS and RES and control peroxiredoxins, nitroreductases and azoreductases as detoxification mechanisms. Hence, the next section summarizes the

"#! ! ! ! "#$%&'(!)! ! ! main redox sensors known in bacteria for ROS and RES including the new thiol-based sensors of B. subtilis.

5.1. OxyR as thiol-based peroxide redox-sensor of E. coli The first redox-sensitive transcriptional factor that directly senses ROS by reversible oxidation of reactive cysteine residues was identified as OxyR in Salmonella typhimurium [74]. OxyR belongs to the LysR family of DNA-binding proteins that binds to DNA as tetramer. The Storz group has shown that OxyR is activated in response to

H2O2 stress by a thiol-disulfide switch (Figure 10). H2O2 leads to oxidation of the conserved Cys199 to a sulfenic acid intermediate that reacts further with Cys208 in the same subunit, forming an intramolecular disulfide [81]. Disulfide bond formation causes re-orientation of the N- and C-terminal domains that allows binding of the OxyR tetramer to the promoter regions of the OxyR regulon genes to activate transcription by contact with RNA polymerase [82]. The Stamler group argued this disulfide-switch model and proposed that oxidation of Cys199 alone to sulfenic acid, S-nitrosylation or S- glutathionylation is sufficient to induce conformational changes and to activate OxyR [83]. However, the crystal structure of oxidized OxyR revealed the Cys199-Cys208 disulfide bridge, that was confirmed also in vivo. It is suggested that the flexible loop region 205-216 plays an important role in bringing Cys208 in close proximity with Cys199 for disulfide bond formation [84]. Moreover, subsequent studies of the E. coli stress response elicited by RNS failed to reveal a significant role for OxyR [56]. In E. coli, OxyR positively controls a large regulon of genes with antioxidant functions, including the genes for the H2O2 scavenging AhpCF peroxiredoxin system, the catalase KatG, the iron-regulator Fur, the DNA-binding ferritin-like protein Dps, the cation importer MntH, the FeS cluster assembly SufABCDE machinery and genes that maintain the thiol-redox balance including TrxC, GrxA, Gor and DsbG. Hence, oxidized OxyR is reduced by the glutaredoxin (GrxA)/GSH/Gor reducing system to turn off the OxyR response. The OxyR regulon genes confer peroxide resistance in E. coli, but also protect cells against heat, UV, singlet oxygen, lipid and neutrophil killing [85]. OxyR is widely conserved in many Gram-negative and Gram-positive bacteria and has been studied in Proteobacteria, Bacteroidetes and Actinomycetes [85]. OxyR also controls conserved antioxidant function genes like catalases and alkylhydroperoxide reductases in many of those bacteria while the size of the OxyR regulon varies in different species.

"#! ! ! ! "#$%&'(!)! ! !

Figure 10: The disulfide-switch model for transciptional activation of OxyR in E. coli. OxyR responds to hydrogen peroxide (H2O2) in E. coli and other bacteria. The conserved C199 and C208 residues are essential for redox-sensing activity. C199 is initially oxidized to the sulfenic acid intermediate that rapidly reacts further to form an intramolecular disulfide with Cys208. Oxidized OxyR binds as a tetramer to promoter regions to activate transcription of genes encoding proteins with antioxidant functions including GrxA, which is involved in the reduction oxidized OxyR. The Figure is from [56]. 5.2. PerR as metal-based peroxide sensor of B. subtilis

In B. subtilis, the Fur-family protein PerR, functions as the main peroxide sensor. PerR is a dimeric repressor that binds to a heptameric 7-1-7 inverted repeat (the Per box) in the promoter region of its target genes [86]. Under H2O2 stress, PerR is inactivated leading to derepression of the PerR regulon genes. The PerR regulon genes are functionally similar to the OxyR regulon genes, including the alkyl hydroperoxide reductase ahpCF operon, the catalase katA gene, the dps-homologous mrgA gene, the heme biosynthesis hemAXCDBL operon, fur, and Zn uptake zosA gene [87, 88]. Two overlapping PerR boxes are present in the perR upstream region suggesting that PerR is autoregulated [87]. The derepression of the PerR regulon genes under peroxide stress conditions confers protection to lethal peroxide concentrations as adapative response similar as shown for the OxyR regulon of E. coli [89]. Most bacteria except for Neisseria gonorrhoeae use either OxyR or PerR as peroxide sensor. In N. gonorrhoeae, OxyR controls genes for peroxide detoxification and disulfide reduction, while PerR controls metal ion homeostasis [85]. PerR contains two metal binding sites, an structural Zn2+ binding site coordinated by four cysteine residues in the C-terminal domain and a regulatory Fe2+ or Mn2+ binding site with three histidine and two aspartic acid residues as ligands [90]. Both Mn2+ and Fe2+ bind competitively to the PerR regulatory site, but only iron-bound PerR is sensitive "#! ! ! ! "#$%&'(!)! ! !

2+ to metal-catalyzed oxidation [82]. Exposure to H2O2 leads to oxidation of Fe in the regulatory site by a Fenton reaction and generates HO!. The hydroxyl radical causes oxidation of His37 and His91 to 2-oxo-histidine that leads to dissociation of PerR from the promoter DNA (Figure 11) [90, 91]. Although the oxidized His37 still had affinity for the regulatory metal, no metal binding with other ligands was possible and PerR failed to retain the close conformation for DNA binding [91]. 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 as novel mechanism for peroxide sensing. However, in our genome-wide transcriptional studies we have shown that conditions that induce disulfide stress in B. subtilis, such as diamide and hypochlorite stress also lead to thiol-specific oxidative stress responses and induction of the PerR regulon genes [46, 92]. Since diamide and hypochlorite stress induce disulfide stress and S-thiolations as shown by redox proteomics, it is possible that the Cys residues in the structural Zn site of PerR are oxidized to disulfides leading to inactivation of PerR via a thiol-disulfide redox switch. In agreement, we have identified one disulfide-linked peptide in the Zn binding site of PerR using mass spectrometry in vivo after hypochlorite stress [46].

! Figure 11: Metal-catalyzed histidine oxidation of PerR. PerR has a regulatory Fe2+or Mn2+-binding site (red) and a structural Zn2+-binding site coordinated by four cysteine residues (green). Reaction of PerR-Fe with H2O2 leads to oxidation of His37 and His91 to the 2-oxo-His derivatives that inactivate the PerR repressor under H2O2 stress. 5.3. OhrR as MarR-family thiol-based redox sensor of organic hydroperoxides Organic hydroperoxides (OHP) include cumene hydroperoxide (CHP), tertiar- butyl hydroperoxide, linoleic acid hydroperoxide that can be derived from peroxidation of unsaturated fatty acids of eukaryotic membrane lipids. The main OHP detoxification systems in bacteria are the alkyl hydroperoxide reductase AhpCF and the organic hydroperoxide resistance protein or two-Cys peroxiredoxin Ohr [56, 82]. The thiol- dependent peroxiredoxin Ohr is specifically induced by OHPs in Xanthomonas camperstris and B. subtilis and shown to catalyze the reduction of OHPs to their corresponding alcohols [53, 93]. Ohr-like peroxidases are conserved in many Gram-

""! ! ! ! "#$%&'(!)! ! ! negative and Gram-positive bacteria [94]. B. subtilis has two ohr paralogs: ohrA and ohrB [53]. OhrA is regulated by the redox-sensing transcription regulator OhrR and OhrB belongs to the !B general stress regulon [95]. The OhrR repressor belongs to the MarR (Multiple antibiotic resistance) family of transcription factors. Members of the MarR family are dimers with winged helix-turn- helix DNA-binding domains and control genes that confer resistance to antibiotics, organic solvents, detergents, ROS, and RES [56]. OhrR acts as a dimeric repressor that binds to inverted repeat sequences in the ohrA promoter, thereby inhibiting transcription [53]. OhrR harbours 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 [52] and the two-

Cys-type with more than one Cys residue with the prototype of X. camperstris OhrRXc

(Figure 12) [56, 96]. 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 [96]. 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 [56]. In contrast, the one-Cys type OhrRBs of B. subtilis is oxidized at Cys15 to the sulfenic acid that reacts further to a mix-disulfide with BSH (S-bacillithiolated OhrR) in response to OHPs and hypochlorite stress as shown in our studies [46, 52]. It was further shown that B. subtilis OhrRBs can be 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 [97]. Interestingly, also a one-Cys-type OxyR protein senses peroxide stress in Deinococcus radiodurans, but the exact mechanism has yet to be explored. In this thesis, we show for the first time, that ohrA responds also specifically to NaOCl stress and thus, OhrR is a redox sensor for OHPs and NaOCl [46]. Since hypochlorite is produced by activated macrophages, it could be a more physiologically oxidant for OhrR-like proteins in pathogenic bacteria, such as S. aureus that has SarZ and MgrA as one-Cys-type OhrR homologs. Transcriptome analysis showed a 220-fold up- regulation of ohrA and mass spectrometry identified S-bacillithiolated OhrR as regulatory mechanism for OhrR inactivation. Phenotype analyses showed that the OhrA

"#! ! ! ! "#$%&'(!)! ! ! peroxiredoxin and BSH protect cells against hypochlorite stress since the growth of ohrA and bshA mutants was strongly impaired by NaOCl stress [46]. Thus, we hypothesize that OhrA could be involved in NaOCl detoxification. The oxidation of a related human two- Cys peroxiredoxin by NaOCl via intramolecular disulfide formation was recently shown in endothelial cells and the authors have also further identified S-glutathionylated proteins confirming our redox proteomics results [98, 99].

Figure 12. Redox-sensing mechanisms of the 1-Cys and 2-Cys type OhrR family proteins. OhrR controls the OhrA thiol-dependent peroxidase and contains one conserved reactive Cys15 residue in B. subtilis and three Cys residues (C22, C127 and C131) in X. campestris. The 1-Cys OhrR protein of B. subtilis is initially oxidized by CHP to the Cys sulfenic acid, which further undergoes reversible S- thiolation with BSH in vivo. The 2-Cys OhrR protein of X. campestris is regulated by intersubunit disulfide formation between C22 and C127’ of opposing subunits. This figure is adapted from [56]. 5.4. MarR/DUF24-family thiol-based sensors of RES The MarR/DUF24 family of transcription factors is conserved among Gram- positive bacteria [56]. In Corynebacterium glutamicum, the MarR/DUF24-type QorR was characterized as a transcriptional repressor that senses diamide and H2O2 and controls the quinone oxidoreductase QorA [100]. Inactivation of QorR involves intersubunit disulfide formation between the conserved single Cys17 residues of both subunits [100]. B. subtilis encodes eight paralogous MarR/DUF24-family proteins: HxlR, HypR, YodB, CatR, YdeP, YdzF, YkvN, and YtcD. The regulatory role of four of these regulators was characterized. HxlR was identified as an activator of the formaldehyde-inducible hxlAB operon that encodes two key enzymes in the ribulose monophosphate pathway [101]. The transcription factor HypR positively controls expression of a nitroreductase HypO [69].

"#! ! ! ! "#$%&'(!)! ! !

HypO is strongly induced by NaOCl, diamide and quinone stress. Nitroreductases catalyse the detoxification of nitroaromatic compounds [102]. The nitroreductases YodC and MhqN are induced by quinones and diamide and suggested to function as azo- and quinone reductases [92]. The flavin-containing NAD(P)H oxidoreductase NfrA of Staphylococcus aureus showed weak disulfide reductase activity and was suggested to function as a thiol-disulfide oxidoreductase [103]. The homologous enzyme NfrA1 of B. subtilis exhibits NADH oxidase activity, catalyzing the oxidation of NADH to NAD+, and scavenges high concentrations of H2O2 [104]. This suggests that HypO could function either in detoxification or disulfide reduction. HypR is a two-Cys MarR/DUF24-type regulator with a reactive Cys14 and a second Cys49 that are 8Å apart in the reduced HypR structure (Figure 13) [69]. The redox-sensing Cys14 has a lower pKa of 6.4 and is present in the thiolate anion form under physiological pH in the cytoplasm. A hydrogen-bonding network stabilizes the reactive Cys14 thiolate. Both Cys14 and Cy49 are essential for activation of hypO transcription by disulfide stress. HypR is oxidized and activated by disulfide stress to the Cys14-Cys49' intersubunit disulfide according to the two-Cys redox-switch model [69]. Disulfide bond formation breaks the H-bonds of Cys14 and moves the !4 and !4' helices of HypR ~4 Å towards each other causing small conformational changes between the reduced and oxidized conformations. It is suggested that these structural changes are sufficient for recognition of the hypO operator sites and recruitment of the RNAP to initiate transcription [69]. Future studies are required to resolve the crystal structure of oxidized HypR in complex with the hypO operator DNA.

"#! ! ! ! "#$%&'(!)! ! !

Figure 13: The pocket of the redox-sensitive Cys residues (A,B) and superimposition of the reduced and oxidized MarR/DUF24-family regulator HypR (C). A) Key interactions between !1 and !3' helices including the Cys14Ser and Cys49' residues for reduced HypR. Cys14Ser forms hydrogen bonds with Val16 and Glu17. Cys49' is in a hydrophobic environment formed by Ile52', Trp27', Ile30', Leu31', Gln60C"', Ile48'. The distance between Ser14O" and Cys49'S" that form the intermolecular disulfide bond is 8.48 Å. B) The structure of the Cys14-Cys49' intersubunit disulfide region in oxidized HypR which emphasizes the movement of !4 helices. (C) Superimposition of the oxidized HypR dimer (light and dark blue) and the reduced HypR dimer (light and dark green). The HypRox/red side view is shown and Cys14 and Cys49' are shown as yellow sticks. The Figure is from Palm et al., 2012 [69]. The MarR-type regulators YodB, CatR and MhqR control specific detoxification pathways that confer resistance to quinones and diamide. These regulators control azoreductases (AzoR1 and AzoR2), nitroreductases (YodC and MhqN), and thiol- dependent dioxygenases (CatE, MhqA, MhqE, MhqO) [68, 79, 92, 105]. Azoreductases and nitroreductases reduce quinones and diamide to hydroquinones and dimethylurea, respectively. Dioxygenases catalyse 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 [68, 79]. 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 stress conditions. 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 [67]. Although CatR has only Cys7, it is also inactivated by intersubunit disulfide formation [68]. The reactive Cys7 is essential for CatR to sense quinone and disulfide stress. However, further studies are required to resolve the mechanisms how the repressors CatR and YodB are inactivated by quinones and diamide in vivo.

5.5. Spx as thiol-based redox sensor for disulfide stress The thiol-based sensor Spx is an arsenate reductase (ArsC) family protein with a CxxC redox motif that has no DNA-binding domains and controls the thiol-specific stress response in B. subtilis [92]. Oxidation of Spx by ROS and RES to an intramolecular disulfide allows Spx to interact with the C-terminal domain (CTD) of the ! subunit of the RNA polymerase (RNAP) [47, 64, 106, 107]. The Spx-!CTD-RNAP complex recognizes promoter regions of the Spx regulon genes and thereby activates transcription. Spx positively regulates the expression of genes required for maintenance of the thiol-redox balance including thioredoxin/thioredoxin reductases (trxAB), thiol-dependent peroxidase (tpx), FMN-dependent oxidoreductases (nfrA, yugJ), methionine sulfoxide reductase (msrA), and the cysteine biosynthesis genes (yrrT operon, cysK) [56]. Spx also represses transcription of competence genes [64]. Orthologs of Spx have been identified in many Gram-positive bacteria that play a role in oxidative stress resistance and biofilm formation, such as in S. aureus and S. epidermidis [108, 109]. Expression of Spx is controlled at transcriptional and post-translational levels. Transcription of spx is initiated from at least four different promoters and by three forms of RNAP containing "A, "B and "M [110]. Spx is regulated in response to disulfide by the PerR and YodB repressors, that are inactivated by disulfide bond formation [64]. Spx is post-translationally controlled by proteolysis via the ClpXP proteases. Under non-stress condition, Spx is guided by the adaptor YjbH to the ClpXP system for degradation [111]. ROS or RES oxidize the Zn- binding site of YjbH to an intramolecular disulfides leading to Zn release. Inactivation of the YjbH adapter results in Spx stabilization. The unfoldase ClpX contains a Cys-rich Zn- finger domain that is also inactivated by thiol oxidation. Therefore, Spx activity is controlled at transcriptional and post-translational levels. In summary, different thiol-modifications are implicated in redox sensing of prokaryotic transcription factors depending on the number of redox-active Cys residues and the reactive species. Redox-sensitive regulators with more than one reactive Cys "#! ! ! ! "#$%&'(!)! ! ! residue undergo in most cases reversible inter- and/or intramolecular disulfide linkages, which serve as sensing mechanisms for OxyR, the 2-Cys OhrR family, MexR, OspR, Spx, CprK and CrtJ. In contrast to these classical thiol-disulfide-switches, transcription factors with one redox-active Cys residue are reversibly regulated via S-thiolation with LMW thiol-redox buffers as shown for the one-Cys OhrR. This thesis has contributed to the identification and characterization of novel thiol-based redox sensors of B. subtilis.

6. Conclusion and future perspectives Bacterial cells have to maintain their reduced state of the cytoplasm and many reactive species, such as ROS and RES affect the cellular redox balance. LMW thiols and redox-sensing transcription factors that control specific detoxification pathways for ROS and RES have essential roles to sense changes in the redox balance and to activate defence mechanisms. Responses to ROS and RES stress involve a complex interplay between redox-signalling pathways and changes in enzymatic activities of metabolic and antioxidant enzymes. Using phenotypic, transcriptomic, proteomic, metabolomic and detailed genetic and biochemical analyses, we have characterized important redox-sensors that sense ROS and RES (YodB, CatR, HypR) in B. subtilis. Under redox stress conditions, these transcription factors are either activated or inactivated via posttranslational modification at their redox-sensitive cysteine residues. This leads to activation of genes for RES and ROS detoxification and in protection of proteins by reversible S-bacillithiolation mechanisms. We show that the thiol-redox buffer BSH plays the most important role in protection against the strong oxidant hypochlorite in Firmicutes bacteria. Since pathogenic Firmicutes bacteria have to cope with HOCl during infections, BSH plays likely an important role for virulence in pathogens. Thus, the development of drugs that inhibit BSH biosynthesis could be promising to combat life- threatening bacterial infections.

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!

Chapter 2

S-bacillithiolation protects against hypochlorite stress in Bacillus subtilis as revealed by transcriptomics and redox proteomics

Bui Khanh Chi, Katrin Gronau, Ulrike Mäder, Bernd Hessling, Dörte Becher, and Haike Antelmann*.

Molecular and Cellular Proteomics, 10: M111.009506 (2011).

* corresponding author

! "#! Research

© 2011 by The American Society for Biochemistry and Molecular Biology, Inc. This paper is available on line at http://www.mcponline.org S-Bacillithiolation Protects Against Hypochlorite Stress in Bacillus subtilis as Revealed by Transcriptomics and Redox Proteomics*□S

Bui Khanh Chi‡, Katrin Gronau‡, Ulrike Ma¨ der§, Bernd Hessling‡, Do¨ rte Becher‡, and Haike Antelmann‡¶

Protein S-thiolation is a post-translational thiol-modifica- Molecular & Cellular Proteomics 10: 10.1074/mcp. tion that controls redox-sensing transcription factors and M111.009506, 1–21, 2011. protects active site cysteine residues against irreversible oxidation. In Bacillus subtilis the MarR-type repressor OhrR was shown to sense organic hydroperoxides via Reactive oxygen species (ROS)1 are an unavoidable con- formation of mixed disulfides with the redox buffer bacil- sequence of the aerobic lifestyle of many organisms (1, 2). lithiol (Cys-GlcN-Malate, BSH), termed as S-bacillithiola- ROS can be generated by incomplete reduction of molecular tion. Here we have studied changes in the transcriptome oxygen during the respiratory chain. Pathogenic bacteria en- and redox proteome caused by the strong oxidant hypo- counter ROS, such as hydrogen peroxide (H O ), superoxide chloric acid in B. subtilis. The expression profile of NaOCl 2 2 anion and hypochloric acid as defense of the innate immune stress is indicative of disulfide stress as shown by the induction of the thiol- and oxidative stress-specific Spx, response during host-pathogen interactions. Upon bacterial CtsR, and PerR regulons. Thiol redox proteomics identi- infection, myeloperoxidase is released from activated macro- fied only few cytoplasmic proteins with reversible thiol- phages to produce the strong oxidant hypochloric acid from Ϫ oxidations in response to NaOCl stress that include GapA H2O2 and Cl (3, 4). and MetE. Shotgun-liquid chromatography-tandem MS ROS can damage all cellular macromolecules, such as pro- analyses revealed that GapA, Spx, and PerR are oxidized teins, lipids, carbohydrates, and nucleotides (2, 5). Cells ac- to intramolecular disulfides by NaOCl stress. Further- tivate the expression of antioxidant mechanisms to detoxify more, we identified six S-bacillithiolated proteins in ROS and to repair the damage. The response of bacteria to NaOCl-treated cells, including the OhrR repressor, two H O and organic hydroperoxides (ROOH) involves heme- methionine synthases MetE and YxjG, the inorganic py- 2 2 cofactor containing catalases and thiol-dependent peroxi- rophosphatase PpaC, the 3-D-phosphoglycerate dehy- dases as detoxification mechanisms (5, 6). Peroxidases use drogenase SerA, and the putative bacilliredoxin YphP. S-bacillithiolation of the OhrR repressor leads to up- conserved redox-active cysteine residues that function in re- regulation of the OhrA peroxiredoxin that confers to- duction of peroxides. These oxidative stress defense mech- gether with BSH specific protection against NaOCl. S- anisms are often controlled by redox-sensitive transcription bacillithiolation of MetE, YxjG, PpaC and SerA causes factors that undergo post-translational thiol-modifications hypochlorite-induced methionine starvation as sup- upon challenge with ROS leading either to activation or inac- ported by the induction of the S-box regulon. The mech- tivation of the transcription factors (7). Post-translational thiol- anism of S-glutathionylation of MetE has been de- modifications implicated in redox-sensing mechanisms are scribed in Escherichia coli also leading to enzyme inactivation and methionine auxotrophy. In summary, our 1 The abbreviations used are: ROS, reactive oxygen species; BSH, studies discover an important role of the bacillithiol redox bacillithiol; Brx, bacilliredoxin; CHP, cumene hydroperoxide; Cys, buffer in protection against hypochloric acid by S-bacilli- cysteine; GlcNAc, N-acetyl glucoseamine; GSH, glutathione; GST, thiolation of the redox-sensing regulator OhrR and of glutathione S-transferase; IAM, iodoacetamide; IP, immunoprecipita- four enzymes of the methionine biosynthesis pathway. tion; LMW, low molecular weight; Mal, malate; MSH, mycothiol; Met, methionine; MG, methylglyoxal; MHQ, methylhydroquinone; N5-THF, 5-methyltetrahydrofolate; N5,N10-THF, 5,10-methylenetetrahydrofo-

From the ‡Institute for Microbiology and §Interfaculty Institute for late; PAPS, 3Ј-phosphoadenosine-5Ј-phosphosulfate; PPi, inorganic Genetics and Functional Genomics, Ernst-Moritz-Arndt-University of pyrophosphate; ROOH, organic hydroperoxide; ROH, organic alco- Greifswald, D-17487 Greifswald, Germany hol; RES, reactive electrophilic species; RuMP, ribulose-5-mono- Received March 10, 2011, and in revised form, July 2, 2011 phosphate; THF, tetrahydrofolate; TrxAB, thioredoxin/thioredoxin re- Published, MCP Papers in Press, July 12, 2011, DOI 10.1074/ ductase; RNS, reactive nitrogen species; FOX, ferrous oxidation mcp.M111.009506 xylenol orange.

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–1 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis known as thiol-disulfide redox-switches and include in most tions that contribute to the resistance of B. subtilis to the cases inter- or intramolecular disulfides and mixed disulfides strong oxidant hypochloric acid. Hypochloric acid is the active with low molecular weight (LMW) thiols (S-thiolations). The component of household bleach and widely used as antimi- best studied examples for peroxide-sensing thiol-based tran- crobial disinfectant to clean surfaces. The bactericidal effect scription factors are Escherichia coli OxyR (8–11) and yeast of hypochloric acid has been proposed to involve generation Yap1 transcription factor (7, 12, 13) that are activated by of ROS, such as superoxide anion and hydroxyl radical for- intramolecular disulfide bond formation to induce the antiox- mation in E. coli (27). Recent redox proteomics studies in idant defense mechanisms. E. coli using the OxICAT approach have shown that bleach In Bacillus subtilis, the major detoxification enzymes for causes strong disulfide formation and protein aggregation in a peroxides are catalase (KatA) and alkylhydroperoxide reduc- different set of proteins than H2O2 (28). As defense mecha- tase (AhpCF) that are controlled of the peroxide-sensing Fur nism against NaOCl stress, E. coli uses the redox controlled family metalloregulatory PerR repressor (5). PerR harbors a chaperone Hsp33 that is activated by NaOCl by the formation structural Zn-binding site coordinated by four cysteine resi- of intramolecular disulfides in the Zn-redox switch centers dues and a regulatory Fe or Mn binding site consisting of His resulting in Zn release, oxidative unfolding and dimerization and Asp residues. Inactivation of PerR is caused by Fe- (29). Hsp33 protects cells against NaOCl-induced protein ag- catalyzed oxidation of His37 and His91 to 2-oxohistidine in gregation. The mode of action has been also studied using the regulatory site (5, 14–16). The response to ROOH involves transcriptome analyses in pathogenic E. coli O157:H7 out- the MarR-type repressor OhrR in B. subtilis that is conserved break strains and the food-borne pathogen B. cereus in many other bacteria (6, 7). OhrR-like proteins control a ATCC14579 (30, 31). Both transcriptome analyses suggest a thiol-dependent peroxiredoxin that converts ROOH to organic major oxidative stress response mechanism of NaOCl. Regu- alcohols. OhrR proteins can be devided into the one and lons involved in the biosynthesis of sulfur and sulfur-contain- two-Cys families of redox sensing repressors. The OhrR pro- ing amino acids were up-regulated by NaOCl in both genome- tein of Xanthomonas campestris belongs to the two-Cys fam- wide studies. However, the mode of action of hypochloric ily that is oxidized to a Cys22-Cys127‘ intermolecular disulfide acid has not yet been investigated in B. subtilis. between both subunits of the OhrR dimer (17). One-Cys OhrR We have used transcriptomic and redox proteomic ap- proteins harbor one conserved N-terminal Cys with the pro- proaches coupled with shotgun-LC-MS/MS analyses to ana- totype of B. subtilis OhrR or Staphylococcus aureus SarZ and lyze the mode of action and reversible thiol-modifications by MgrA (7, 18). Cumene hydroperoxide (CHP) leads to Cys15 NaOCl stress in B. subtilis. We discovered that the major oxidation to sulfenic acid that is rapidly oxidized to S-thiolated resistant determinant to NaOCl is the OhrA peroxiredoxin that OhrR containing cysteine or the redox buffer bacillithiol (BSH) conferred specific protection against NaOCl toxicity. More- (19, 20). Thus, B. subtilis OhrR is controlled by S-cysteinyla- over, we identified S-bacillithiolations of the OhrR repressor, tion and S-bacillithiolation as redox-switch mechanism lead- two methionine synthases MetE and YxjG, the inorganic py- ing to inactivation of the OhrR repressor function and dere- rophosphatase PpaC, and the 3-D-phosphoglycerate dehy- pression of ohrA transcription. drogenase SerA as major protection mechanisms against hy- In previous studies, we investigated the global response, pochlorite stress in B. subtilis. post-translational modifications and specific regulatory mechanisms that are induced by reactive electrophilic spe- EXPERIMENTAL PROCEDURES cies (RES) in B. subtilis, such as diamide, quinones, or alde- Bacterial Strains and Growth Conditions—The bacterial strains hydes. RES deplete the cellular redox buffer cysteine leading used were B. subtilis wild-type strains 168 (trpC2), JH642 (trpC2 to induction of the Spx-regulon that controls thiol-disulfide attSP␤), and CU1065 (trpC2 pheA1) and mutant strains ⌬spx r r oxidoreductases (TrxAB) to restore the redox homeostasis (trpC2,spx::neo ) (32), ⌬ohrR (trpC2,ohrR::cm ), ⌬ohrA (trpC2, ohrA::cmr), ⌬sigB (trpC2,sigB::cmr), ⌬perR (trpC2,perR::cmr) (33), (21–23). B. subtilis encodes specific redox-sensing regulators HB9121 (CU1065 trpC2,ohrR::kmr ohrR-FLAG (Spcr) ohrA-cat lacZ of the MarR/DUF24-family that sense RES, but not ROS (7). (Neor)(19),HB2048(CU1065SP␤c2⌬2::Tn917::(ohrA-cat- These include the paralogous repressors YodB and CatR lacZ)ohrR::kan,thrC::pXTohrRC15S)(34), HB11002 (CU1065 trpC2, that are inactivated via intermolecular disulfide formation by bshA::mlsr), and HB11053 (CU1065 trpC2, bshB1:: spcr bshB2::cmr) diamide and quinones resulting in derepression of the (35). B. subtilis strains were cultivated under vigorous agitation at 37 °C in Belitsky minimal medium (BMM) as described previously (36). azoreductase (AzoR1), nitroreductase (YodC), and thiol-de- The antibiotics were used at the following concentrations: 1 ␮g/ml pendent dioxygenase (CatE) catalyzing the detoxification of erythromycin, 25 ␮g/ml lincomycin, 5 ␮g/ml chloramphenicol, 10 the electrophiles (24–26). Other proteins of the MarR/ ␮g/ml kanamycin, and 100 ␮g/ml spectinomycin. Sodium hypochlo- DUF24-family (HxlR) and of the MerR/NmlR-family (AdhR) rite (15% stock solution), diamide, and cumene hydroperoxide were sense specifically aldehydes, such as formaldehyde and purchased from Sigma Aldrich. For NaOCl stress exposure, cells were grown in BMM to an optical methylglyoxal (23). density at 500 nm (OD500) of 0.4 and treated with 50, 75, or 100 ␮M In this study, we were interested in the global response, NaOCl diluted freshly in destilled water. The growth experiments in regulatory mechanisms, and post-translational thiol-modifica- the presence of methionine were performed by addition of 75 ␮M

10.1074/mcp.M111.009506–2 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

methionine either after inoculation of the culture or 30 and 60 min after versibly oxidized proteins in the Coomassie-stained thiol-redox NaOCl stress exposure. proteome was performed manually as described previously (24). In Gene deletions for construction of the ohrA mutant were generated brief, gel pieces were washed 3–5 times with 1 ml of 20 mM (w/v) using long-flanking-homology polymerase chain reaction (LFH-PCR) ammonium bicarbonate, pH 8.0/50% (v/v) acetonitrile (ACN) for 30 as described previously (25). Primers ohrA-F1 (5Ј-TGCAGCTGATTG- min and once with 1 ml 75% ACN for 30 min. Gel pieces were dried AGGATACG-3Ј) and ohrA-F2 (5Ј-GTTATCCGCTCACAATTCGCGGT- and the proteins in-gel digested with 20 ␮g/␮ltrypsindissolvedin CTGATGAAATGACCT-3Ј) were used to amplify the up fragment and water (Promega, Madison, WI) at 37 °C for 16 h. Tryptic peptides primers ohrA-R1 (5Ј-CGTCGTGACTGGGAAAACGGTGTGACGCTG- were eluted with 0.5% (w/v) trifluoroacetic acid/50% (v/v) ACN and CAAGTAAA-5Ј) and ohrA-R2 (5Ј-CCCTTCAATCTCCGAATCAA-3Ј) to 0.5 ␮lofthispeptidesolutionwasspottedontheMALDI-targets. amplify the down fragment, respectively. Fragments were amplified Then, 0.5 ␮lofmatrixsolution(50%(v/v)ACN/0.5%(w/v)trifluo- and joined together with the chloramphenicol cassette using Pfusion roacetic acid) saturated with ␣-cyano-4-hydroxy cinnamic acid was DNA polymerase (Invitrogen, Carlsbad, CA) as described (33). Inte- mixed with the spotted tryptic peptides and dried on the target for gration and deletion of the ohrA gene were confirmed by PCR and by 15 min. Northern blot analysis using digoxigenin-labeled RNA probes of the The matrix-assisted laser desorption ionization/time of flight corresponding gene. (MALDI-TOF)-TOF measurement of spotted peptide solutions was Analysis of NaOCl Concentrations in the Cell Culture Supernatant carried out on a Proteome-Analyzer 4800 (Applied Biosystems, Foster Using the FOX Assay—The concentrations of the remaining NaOCl in City, CA). The spectra were recorded in reflector mode in a mass the culture supernatants were determined using the FOX assay (37). range from 900 to 3700 Da with a focus mass of 2000 Da. For one FOX reagent was prepared by mixing 100 ml FOX I (100 mM sorbitol, main spectrum 25 subspectra with 100 shots per subspectrum were 125 ␮M xylenol orange) and 1 ml FOX II (25 mM ammonious ferrous- accumulated using a random search pattern. If the autolytical frag- ϩ (II)sulfate in 2.5 M H2SO4). It was not possible to measure any NaOCl ment of trypsin with the monoisotopic (MϩH) m/z at 2211.104 concentrations in BMM with tryptophane and glutamate. Thus, cells reached a signal to noise ratio (S/N) of at least 10, an internal cali- were grown to an OD500 of 0.4 in BMM, centrifuged and resuspended bration was automatically performed using this peak for one-point- in BMM without tryptophane and glutamate before the addition of 75 calibration. The peptide search tolerance was 50 ppm but the actual ␮M NaOCl. Samples of 500 ␮l medium were taken at different time RMS value was between 10 and 20 ppm. After calibration the peak points after NaOCl addition, mixed with 500 ␮l FOX reagent and lists were created by using the “peak to mascot” script of the GPS incubated at room temperature for 60 min. The absorbance was Explorer™ software version 3.6 with the following settings: mass measured at 560 nm. Calibration curves were generated using NaOCl range from 900 to 3700 Da; peak density of 50 peaks per range of 200 concentrations in the range from 0 to 100 ␮M diluted in BMM without Da; minimal area of 100 and maximal 200 peaks per protein spot and tryptophane and glutamate. minimal S/N ratio of 6. The peak lists were searched against a Bacillus Thiol Redox Proteome Analysis—The thiol redox proteome analysis subtilis sequence database extracted from UniprotKB release 12.7 was performed as described previously (38) with the modifications as (UniProt Consortium, Nucleic acids research 2007, 35, D193–197) explained (21). Cells were harvested before (control conditions) and using the Mascot search engine version 2.1.04 (Matrix Science Ltd, 10, 20, and 30 min after exposure to 50 ␮M NaOCl stress, resus- London, UK). pended in urea/CHAPS alkylation buffer (8 M urea; 1% CHAPS; 1 mM MALDI-TOF-TOF MS/MS analysis was performed for the three EDTA; 200 mM Tris-HCl pH 8,0; 100 mM iodoacetamide (IAM)), son- strongest peaks of the TOF-spectrum. For one main spectrum 20 icated, alkylated for 20 min in the dark, precipitated with 100% sub-spectra with 125 shots per subspectrum were accumulated us- acetone, washed several times with 80% acetone and dried. After ing a random search pattern. The internal calibration was automati- resolving in urea/CHAPS buffer without IAM, 200 ␮g of the protein cally performed as one-point-calibration if the mono-isotopic arginine extract were reduced with 10 mM Tris-(2-carboxyethyl)-phosphine (MϩH)ϩ m/z at 175.119 or lysine (MϩH)ϩ m/z at 147.107 reached a

(TCEP) and labeled with BODIPY FL C1-IA [N-(4,4-difluoro-5,7-di- S/N of at least 5. The peak lists were created by using the “peak to methyl-4-bora-3a,4a-diaza-s-indacene-3-yl)-methyl)-iodoacetamide] mascot” script of the GPS Explorer™ software version 3.6 with the (Invitrogen, Eugene, OR). The fluorescence-labeled protein extract following settings: mass range from 60 Da to a mass that was 20 Da was separated using 2D PAGE as described (21). The two-dimen- lower than the precursor mass; peak density of 5 peaks per 200 Da; sional (2D) gels were scanned using a Typhoon 9400 variable mode minimal area of 100 and maximal 20 peaks per precursor and a imager (Amersham Biosciences, Freiburg, Germany) for BODIPY- minimal S/N ratio of 5. Peptide mixtures that yielded a mowse score fluorescence and then stained with Colloidal Coomassie for protein of at least 50 in the reflector mode and a sequence coverage of at amounts. Quantitative image analysis was performed with the least 30% that were confirmed by subsequent MS/MS analysis were DECODON Delta 2D software (http://www.decodon.com). regarded as positive identification. The complete Mascot search re- The first alkylation protocol (38) that applied the TCA-precipitation sults including the MS and MS/MS data of all protein identifications step to harvest cells to stop thiol-disulfide exchanges was changed are shown in supplemental Figs. S1A–P. previously (21) for two reasons: (1) The 2D gels were of bad quality Immunoprecipitation (IP) and Nonreducing SDS-PAGE Analysis of and exhibited protein streaking during the isoelectric focusing (IEF) the OhrR-FLAG Protein—The OhrR-FLAG protein expressing B. sub- and (2) GapA was oxidized artificially by this TCA precipitation step tilis strain HB9121 was grown in BMM and treated with 50 ␮M NaOCl under control conditions but not if this TCA step was omitted (38) and at an OD500 of 0.4. Cells were harvested before (control conditions) GapA is an important redox-controlled cytoplasmic marker protein and 15 min after NaOCl-treatment in TE-buffer (10 mM Tris-HCl, pH8; and strongly oxidized in response to quinones and diamide (21, 22). 1mM EDTA) with 100 mM IAM. Cells were sonicated, the protein Because GapA was neither oxidized using our alkylation protocol in extracts obtained after repeated centrifugation and alkylated in the the redox proteome of control cells nor in the LC-MS/MS approach, dark for 20 min. OhrR-FLAG protein was purified by IP using anti- this indicates no artificial thiol-disulfide exchange during our sample FLAG M2-affinity agarose (Invitrogen) according to the instructions of preparations. the manufacturer. The precipitated OhrR-FLAG protein was eluted by Identification of Reversibly Oxidized Proteins in the Thiol-Redox boiling in non-reducing SDS sample buffer (4% SDS; 62.5 mM Tris- Proteome Using Matrix-Assisted Laser Desorption Ionization/Time HCl pH 8.0, glycerol) and separated using 15% nonreducing SDS- Of Flight-TOF Tandem MS (MS/MS)—Tryptic digestion of the re- PAGE. The OhrR-FLAG protein band of the expected size was cut

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–3 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

from the SDS-gel, tryptically digested as described above and ana- the manufacturer’s instructions. Synthesis and purification of fluores- lyzed by LTQ-Orbitrap mass spectrometry. cently labeled cDNA were carried out as descibed (43) with minor LTQ-Orbitrap Velos Mass Spectrometry and Identification of Post- modifications. In detail, 10 ␮g of total RNA were mixed with random translational Thiol-modifications—B. subtilis wild-type and ⌬bshA primers (Promega) and spike-ins (Two-Color RNA Spike-In Kit, Agilent mutant cells were harvested before (control conditions) and 15 min Technologies, Santa Clara, CA). The RNA/primer mixture was incu- after exposure to 50 ␮M NaOCl stress. Cells were resuspended in bated at 70 °C for 10 min followed by 5 min incubation on ice. Then, urea/CHAPS alkylation buffer with 100 mM IAM as described above the following reagents were added: 10 ␮l of 5x First Strand Buffer and sonicated to obtain the alkylated protein extracts. The alkylated (Invitrogen), 5 ␮l of 0.1 M DTT (Invitrogen), 0.5 ␮l of a dNTP mix (10 mM protein extracts were separated using 15% nonreducing SDS-PAGE dATP, dGTP, and dTTP, 2.5 mM dCTP), 1.25 ␮l of Cy3-dCTP or (200 ␮g each per lane) and the complete lanes were cut into 10 gel Cy5-dCTP (GE Healthcare) and 2 ␮l of SuperScript II reverse tran- pieces and digested with trypsin as described above. Peptides eluted scriptase (Invitrogen). The reaction mixture was incubated at 42 °C for from tryptic digests of gel pieces were subjected to a reversed phase 60 min and then heated to 70 °C for 10 min. After 5 min on ice, the column chromatography (self packed C18 column, 100-␮m i. D. x 200 RNA was degraded by incubation with 2 units of RNaseH (Invitrogen) mm) operated on a Easy-nLC II (Thermo Fisher Scientific, Waltham, at room temperature for 30 min. Labeled cDNA was then purified MA). Elution was performed by a binary gradient of buffer A (0.1% using the CyScribe GFX Purification Kit (GE Healthcare). The individ- (v/v) acetic acid) and B (99.9% (v/v) ACN, 0.1% (v/v) acetic acid) over ual samples were labeled with Cy5, whereas the reference pool was a period of 100 min with a flow rate of 300 nl/min. MS and MS/MS labeled with Cy3. 500 ng of Cy5-labeled cDNA and 500 ng of Cy3- data were acquired with the LTQ-Orbitrap-Velos mass spectrometer labeled cDNA were hybridized together to the microarray following (Thermo Fisher Scientific) equipped with a nanoelectrospray ion Agilent’s hybridization, washing and scanning protocol (Two-Color source. The Orbitrap Velos was operated in data-dependent MS/MS Microarray-based Gene Expression Analysis, version 5.5). Data were mode using the lock-mass option for real time recalibration. After a extracted and processed using the Feature Extraction software (ver- survey scan in the Orbitrap (r ϭ 30,000) MS/MS data were recorded sion 10.5, Agilent Technologies). For each gene on a microarray, the for the 20 most intensive precursor ions in the linear ion trap. Singly error-weighted average of the log ratio values of the individual probes charged ions were not taken into account for MS/MS analysis. was calculated using the Rosetta Resolver software (version 7.2.1, Post-translational modifications of proteins were identified by Rosetta Biosoftware). Genes showing induction or repression ratios searching all MS/MS spectra in “dta” format against an B. subtilis of at least threefold in three independent experiments were consid- target-decoy protein sequence database (8294 entries) using Sorcer- ered as significantly induced. The averages ratios and standard de- er™-SEQUEST® (Sequest version 2.7 rev. 11, Thermo Electron in- viations for all induced or repressed genes are calculated from three cluding Scaffold_3_00_02, Proteome Software Inc., Portland, OR). independent transcriptome experiments after 10 min of exposure to The target-decoy database includes the complete proteome set of B. NaOCl stress and listed in supplemental Table S1 and S2. All microar- subtilis 168 (4105 database entries) that was extracted from Uni- ray datasets are available in the GEO database under accession protKB release 12.7 (UniProt Consortium, Nucleic acids research numbers [GSE27637]. Hierarchical Clustering Analysis—Clustering of gene expression 2007, 35, D193–197) (39) and an appended set of 4147 reversed profiles was performed using Cluster 3.0 (44). The transcriptome data sequences and 42 sequences of common laboratory contaminants sets were derived from previous publications and this study and created by BioworksBrowser version 3.2 (Thermo Electron Corp.) included log2-fold expression changes 10 min after exposure of B. according to Elias et al. (40). The Sequest search was carried out subtilis to diamide (1 mM)(45), methylhydroquinone (MHQ) (0.33 mM) considering the following parameter: a parent ion mass tolerance 10 (46), catechol (2.4 mM) (47), formaldehyde (1 mM), methylglyoxal (MG) ppm, fragment ion mass tolerance of 1.00 Da. Up to two tryptic (2.8 and 5.6 mM)(23), and 50 ␮M NaOCl. After hierarchical clustering, miscleavages were allowed. Methionine oxidation (ϩ15.994915 Da) the output was visualized using TreeView (48). For the clustering 630 and cysteine carbamidomethylation (ϩ57.021464 Da) were set as genes were selected that are induced by RES and NaOCl stress in B. variable modifications. Multiple Sequest searches were performed for subtilis (e.g. CtsR, CymR, Spx, PerR, ArsR, CsoR, CzrA, OhrR, YodB, either intramolecular disulfide bonds (Ϫ2.01565 Da), S-cysteinyla- YvaP (CatR), MhqR, LexA, SigmaD, AdhR, and HxlR regulons) tions (ϩ119.004099 Da for C H NO S) or S-bacillithiolations 3 7 2 (supplemental Table S4). (ϩ396.083866 Da for C H N O S) as variable post-translational 13 22 2 10 Northern Blot Experiments—Northern blot analyses were per- cysteine modifications, allowing a maximum of three modifications formed as described (49) using RNA isolated from B. subtilis wild-type per peptide in each Sequest search. cells before (control) and 10 min after treatment with 50 ␮M NaOCl, 1 Proteins were identified by at least two peptides applying a strin- mM diamide and 100 ␮M CHP, respectively. Hybridizations specific for gent SEQUEST filter. Sequest identifications required at least Cn ⌬ ohrA, nfrA, cysK, katA, ohrA, azoR1, catE, and yitJ were performed scores of greater than 0.10 and XCorr scores of greater than 1.9, 2.2, with the digoxigenin-labeled RNA probes synthesized in vitro using T7 3.3, and 3.75 for singly, doubly, triply and quadruply charged pep- RNA polymerase from T7 promoter containing internal PCR products tides. The complete CID MS/MS spectra of the modified Cys-con- of the respective genes using the primer sets described previously taining peptides and the corresponding b and y fragment ion series (23, 25, 35) and the primer pairs yitJ-for, 5Ј CCGAACAGCAGTCTTC- are given in detail in supplemental Figs. S2 and S3. The Sequest CTTC 3Ј and yitJ-T7-rev, 5Ј CTAATACGACTCACTATAGGGAGAC- search results are submitted as “dta” and “out” files to the PRIDE CGTTTTACCGCTTTATCCA 3Ј for yitJ. database (http://www.ebi.ac.uk/pride/) (41) and deposited under the accession numbers 17516–17659. RESULTS Transcriptome Analysis—For microarray analysis, B. subtilis wild- type cells were grown in minimal medium to OD500 of 0.4 and har- Determination of the Growth-inhibitory Concentration of vested before and 10 min after exposure to 50 ␮M NaOCl. Total RNA NaOCl in B. subtilis—At first we determined the concentration was isolated by the acid phenol method as described (42). For tran- that inhibited the growth of B. subtilis wild-type cells. Exposure scriptome analysis, 35 ␮g RNA were DNase-treated using the RNase- Free DNase Set (Qiagen) and purified using the RNA Clean-Up and of cells to 50 ␮M NaOCl stress caused a lag in growth for 60 min Concentration Micro Kit (Norgen). The quality of the RNA preparations and then growth was resumed with a similar growth rate as the was assessed by means of the Agilent 2100 Bioanalyzer according to untreated control (Fig. 1). This growth profile is very similar to

10.1074/mcp.M111.009506–4 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis that of diamide (26) indicating that cells are able to detoxify the tal Table S3 and visualized in the corresponding diagram in oxidant and to repair the protein damage within this time frame. Fig. 2. Transcriptional induction of selected thiol-stress spe- Growth of B. subtilis is completely inhibited by 100 ␮M NaOCl. cific genes by NaOCl, diamide and CHP was verified by Transcriptome Analysis of the NaOCl Stress Response in B. additional Northern blot analyses (Fig. 3). In the following subtilis—To analyze the mode of action of NaOCl in B. subtilis, sections we have sorted the transcriptome datasets into thiol- we conducted genome-wide transcriptome analyses of cells and oxidative stress specific and general stress-induced treated for 10 min with 50 ␮M NaOCl stress. Genes that were regulons. Ͼ3-fold up-regulated and down-regulated by NaOCl were Induction of the CtsR and Spx Regulons by NaOCl Stress is sorted according to the known stress regulons in sup- Indicative of Disulfide Stress—The microarray data showed plemental Tables S1 and S2. The transcriptome data revealed the up-regulation of the thiol- or electrophile-stress specific the significant induction of 430 genes and the repression of CtsR and Spx regulons in B. subtilis. Among the CtsR and Spx 400 genes by NaOCl stress in three biological transcriptome regulons, the clpE (28-fold), trxA and nfrA (40-fold) genes replicates. Representative genes of the most strongly up- displayed the highest induction ratios. The fold-changes of regulated genes and regulons are listed in supplemen- Spx-controlled genes are similar to diamide stress as verified by the Northern blots (Fig. 3). Because the Spx and CtsR regulons are generally induced by electrophiles, the global response to NaOCl is indicative of disulfide stress (23, 50). In contrast, the CymR regulon that functions in Cys biosynthesis (51) was strongly up-regulated by RES, but not by NaOCl stress. Among the CymR regulon genes, only cysK was 3-fold induced by NaOCl stress. This indicates that NaOCl probably does not deplete the pool of the redox buffer cysteine in B. subtilis. Induction of the S-box Regulon by NaOCl Stress Indicates Methionine Starvation—Interestingly, the S-box regulon genes that are regulated by an S-adenosyl methionine riboswitch mechanism were induced by NaOCl stress (supple- mental Table S1, Fig. 3). The S-box regulon is induced by methionine starvation when S-adenosyl methionine levels are FIG. 1. Growth curves in response to sub-lethal concentrations of NaOCl stress in B. subtilis. B. subtilis wild type was grown in low. The S-box regulon gene products function in methionine biosynthesis (52–55). The microarray data showed the induc- minimal medium to an OD500 of 0.4 and exposed to 50, 75, and 100 ␮M of NaOCl indicated by time point zero. tion of the methionine synthases encoding metE (5,3-fold),

FIG. 2. Transcriptome changes of NaOCl-induced regulons (CtsR, Spx, PerR, OhrR, ArsR, CzrA, SigmaD, and SigmaB) and of the pstSCAB operon. Fold-changes are average induction ratios of genes induced in NaOCl-treated cells versus untreated cells calculated from three transcriptome replicates with standard deviations given as errors bars. Shown are only representative genes of each regulon that are more than fivefold induced by NaOCl in supplemental Tables S1 and S3.

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–5 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

structural Zn-binding site as confirmed by the liquid chroma- tography (LC)-MS/MS results (Table I). Induction of the OhrR-controlled OhrA Peroxiredoxin by NaOCl Stress—Interestingly, the ohrA gene was most strongly up-regulated (220-fold) in the transcriptome by NaOCl. The redox-sensing OhrR repressor controls the OhrA peroxire- doxin that confers resistance to CHP (6, 7). The Northern blots show that derepression of ohrA transcription occurs by CHP and NaOCl stress at similar levels (Fig. 3). These data suggest that OhrR is a specific determinant of the response to NaOCl and ROOH. Induction of Selective RES-specific MarR/DUF24 Regulons FIG. 3. Northern blot analysis of selected thiol-stress specific by NaOCl Stress—We have shown that RES are sensed by genes of the Spx, CymR, S-box, PerR, OhrR, and MarR/DUF24 regulons in response to NaOCl, diamide, and CHP stress. Tran- members of the redox-sensing MarR/DUF24 family. The script analysis of nfrA, trxA, and katA indicates the induction of the YodB and CatR repressors are specific sensors for quinones Spx and PerR-regulons by NaOCl and diamide stress. The CymR- and diamide (7, 24–26, 50). The transcriptome and Northern controlled cysK gene is strongly induced by diamide. The S-box blot results showed the induction of the YodB-regulon genes regulon gene yitJ responds most strongly to NaOCl stress. The ohrA azoR1 (10-fold), yodB (3-fold), and yodC (5-fold) by NaOCl. gene is controlled by the OhrR repressor and strongly induced by NaOCl and CHP. The YodB and CatR-controlled azoR1 and catDE The CatR-controlled catDE operon was threefold induced by genes are strongly up-regulated by diamide. Cells were grown to an NaOCl. The inductions of the YodB and CatR regulons by

OD500 of 0.4 and harvested before (control conditions, co) and 10 min NaOCl are much lower as by diamide and quinones (Fig. 3) after exposure to 50 ␮M NaOCl, 100 ␮M CHP, or 1 mM diamide. The confirming the specific roles of the azoreductases and dioxy- RNA isolation and Northern blot hybridization was performed as genases in detoxification of diamide and quinones. In addi- described in the Methods section. The arrows point toward the sizes of the specific transcripts. tion, the genes encoding further DUF24-like redox sensors, yybR (35-fold), ykvN (19-fold), ydzF (6-fold), and ydeP (6-fold) yxjG (2,5-fold), and yxjH genes (2-fold), the metIC operon were induced by NaOCl stress indicating that these respond (3–4-fold) encoding cystathionine ␥-synthase and ␤- probably to disulfide stress via thiol-based redox switches. that are involved in cystathionine and homocysteine biosyn- The formaldehyde-sensing DUF24-type regulator HxlR con- thesis, the yoaD gene (3,6-fold) that encodes a 3-D-phospho- trols the hxlAB operon encoding the enzymes of the ribulose- glycerate dehydrogenase required for serine biosynthesis and 5-monophosphate (RuMP) pathway (58). The hxlAB operon the yitJ gene (7,4-fold) that encodes a bifunctional homocys- was 8-fold induced by NaOCl stress. teine S-methyltransferase in the N-terminal part and 5,10- Induction of Metal Ion Efflux Systems, Motility and Che- methylenetetrahydrofolate (N5,N10-THF) reductase in the motaxis by NaOCl Stress—Besides thiol- and oxidative stress C-terminal part (see Fig. 7). The NAD(P)H-dependent N5,N10- responses, the CzrA, ArsR, and CsoR regulons were strongly THF reductase activity is required to reduce N5,N10-THF to up-regulated by NaOCl stress that control the operons for 5-methyltetrahydrofolate (N5-THF) as methyl-group donor for metal ion efflux systems czcD-trkA (32-fold), arsR-yqcK- the methionine synthase MetE (56). The cysH operon that arsBC (57-fold), and yvgXYZ (3-fold) (59–61). Furthermore, encodes gene products for sulfur assimilation and cysteine the SigmaD regulon genes for flagella assembly, motility and biosynthesis and the metK gene encoding a S-adenosyl me- chemotaxis were strongly induced by NaOCl stress (23, 62). thionine synthetase were repressed in the transcriptome by Metal ion uptake systems and SigmaD regulon genes were NaOCl stress. The cysH operon belongs also to the CymR induced by NaOCl and RES. regulon and is rather induced by cysteine starvation than by Induction of the SigmaB-dependent General Stress Re- methionine starvation (57). The metK gene displayed also a sponse by NaOCl Stress—In contrast to electrophiles, NaOCl drop of expression at later time points upon methionine star- stress caused the 3–15-fold induction of 52 genes belonging vation in previous studies (52). Thus, metK and cysH are not to the SigmaB general stress response regulon. Among the induced by NaOCl stress in contrast to other S-box regulon SigmaB-controlled genes, 20 genes were relatively weakly genes listed above. induced (3-fold) and 32 genes in the range of 5–13-fold. Induction of the PerR-dependent Oxidative Stress Re- Representative genes of the SigmaB regulon include dps sponse by NaOCl Stress—NaOCl caused the induction of the (5-fold), gsiB (8-fold), gspA (13-fold), ydaST (6-fold), yfhK (7- oxidative stress specific PerR regulon genes katA (73-fold) fold), yfkM (6-fold), yflT (13-fold) ohrB (8-fold), ysnF (7-fold), and ahpCF (7–13-fold) (Figs. 2 and 3). The PerR regulon genes and ytxGHJ (6–13-fold). The induction of the SigmaB re- were similar strongly induced by NaOCl and diamide indicat- sponse might be caused as a result of starvation or even ing that strong oxidants that cause disulfide stress lead oxidative stress response caused by hypochloric acid which probably to oxidation of the conserved Cys residues in the requires further studies. It is interesting to note that NaOCl

10.1074/mcp.M111.009506–6 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis triggers the induction of thiol- and oxidative stress responses Thiol-redox Proteomics Identifies GapA, MetE, MtnA, LeuC as well as general stress responses. and PurQ as Reversibly Oxidized Proteins by NaOCl Stress— Other Genes Induced Strongly by NaOCl Stress—Finally, Next, we were interested in the changes in the thiol-redox many genes with unknown functions are up-regulated in the proteome by NaOCl stress to identify proteins with reversible NaOCl transcriptome (supplemental Table S1). The putative thiol modifications. In brief, reduced thiol groups were Naϩ/Hϩ antiporter nhaX (30-fold) and the major facilitator blocked with IAM and reversibly oxidized proteins were re- superfamily encoding ybcLM operon (42-fold) could be in- duced and labeled using the fluorescence dye BODIPY FL volved in sodium efflux. The high induction of the pstSCAB C1-IA (21, 38). The fluorescence-labeled proteins representing operon (30-fold) by NaOCl might be PhoPR-independent be- the disulfide proteome were separated using two-dimensional cause other PhoPR-regulon genes were not induced, such as gels and the fluorescence image (red) overlaid with the Coo- phoA, phoB, and phoD encoding alkaline phosphatases and massie-stained protein amount image (green) (Fig. 5A,5B). phosphodiesterases. Finally, the yhdJ gene encoding the pu- The oxidation ratios were quantified as BODIPY-fluorescence tative GCN5-N-acetyltransferase was 67-fold induced by levels versus protein amount levels of the reversibly oxidized NaOCl. proteins (Fig. 5C). In agreement with previous studies, pro- Repression of the Stringent Controlled RelA-regulon by teins that form disulfides under control conditions include NaOCl Stress—There are 400 genes in the transcriptome Adk, AhpC, AccB, Eno, Tpx, PdhD, GuaB, CysH, CysJ (YvgR), datasets that displayed more than 3-fold decreased expres- LeuC, YceC, and YumC (Fig. 5A)(21, 22). The identities of sion ratios in response to NaOCl stress (supplemen- these oxidized proteins were verified using MALDI-TOF-TOF tal Table S2). Among the repressed genes are the stringent MS/MS (supplemental Figs. S1A–P). Surprisingly, we did not detect a strong increase of revers- controlled RelA regulon genes involved in translation, ATP ible thiol-oxidations in the redox proteome in many cytoplas- generation, cell wall biosynthesis, and turnover. The PurR and mic proteins after treatment of cells with 50 ␮M NaOCl stress. RyrR regulons controlling purine and pyrimidine biosynthesis Instead, NaOCl rather caused specific oxidation of few selec- were repressed by NaOCl as result of the slower growth rate. tive proteins, including GapA, MetE, MtnA, LeuC, and PurQ The osmostress-responsive uptake systems for choline and (Fig. 5B,5C). The glyceraldehyde-3-phosphate dehydrogen- glycine betaine encoded by the opuA, opuB, and opuC oper- ase (GapA) was identified as most strongly oxidized protein ons were strongly repressed by NaOCl stress (63). with a 10-fold increased fluorescence/protein ratio (Fig. 5B, Hierarchical Clustering Analyses for RES and NaOCl Indi- 5C). The redox proteome analysis revealed also strongly in- cates a Disulfide Stress Signature of NaOCl—Next, we per- creased oxidation ratios for the cobalamin-independent me- formed a hierarchical clustering analysis using selected tran- thionine synthase MetE. Furthermore, the methylthioribose-1- scriptome datasets for diamide (45), MHQ (46–47), catechol phosphate isomerase MtnA (YkrS) is oxidized by NaOCl (47), formaldehyde, methylglyoxal (23), and NaOCl. In total, stress that is involved in methionine biosynthesis via the sal- 630 genes were selected for the clustering analysis that vage pathway (see Fig. 7). showed inductions of at least threefold in any of these tran- Identification of Proteins with Intramolecular Disulfides Un- scriptome datasets (supplemental Table S4). The complete der Control and NaOCl Stress Conditions Using Shotgun-LC- cluster can be arranged in 14 major groups of genes which MS/MS Analyses—We were interested to identify proteins share a similar expression profile by electrophiles and NaOCl with reversible thiol-oxidations, including intramolecular (Fig. 4). The cluster analysis confirmed the common induction disulfides, S-cysteinylations, and S-bacillithiolations under of the CtsR, Spx (node 9), ArsR, CzrA, and SigmaD regulons NaOCl stress conditions. We are aware that we exclude the (nodes 11, 12) by RES and NaOCl as indicator for disulfide identifications of intramolecular disulfides of proteins with stress. The strong induction of the CymR regulon by RES but Cys residues that are separated by tryptic digestion result- not by NaOCl is visualized in cluster 13. The quinone and ing in cross-linked intermolecular disulfides after enzymatic diamide-specific MarR/DUF24 regulons controlled by YodB, digestion. Alkylated protein extracts of control and NaOCl- CatR, and MhqR clustered with the PerR regulon in nodes 6, treated cells were separated using nonreducing SDS-PAGE, 8, and 10. The aldehyde-specific HxlR, AdhR, LexA regulons tryptically in-gel digested and analyzed using LTQ-Orbitrap- and the formate uptake system and formate dehydrogenase Velos mass spectrometry as described in the Methods encoding yrhFG and yrhDE operons (23) clustered in nodes 11 section. and 14. The NaOCl-specific inductions of the OhrR and Eight proteins with intramolecular disulfides were identified SigmaB regulons, the ydzF and yfkN genes encoding DUF24- in wild-type proteome samples at control and NaOCl stress family regulators and the pstSCAB operon are shown in clus- conditions, including Adk, CysJ (YvgR), GltA, PdhD, TopA, ters 1, 3, 4. In conclusion, the transcriptome comparisons YutI, YdjI, and Zwf (Table I and supplemental Figs. S2A–2K). showed that NaOCl elicits a Spx, CtsR-, and PerR-controlled Adk, PdhD, and CysJ were detected also in the redox pro- disulfide and oxidative stress response and a selective OhrR- teome as reversibly oxidized proteins under control condi- and SigmaB response in B. subtilis. tions (Fig. 5A). The dihydrolipoamide dehydrogenase E3 sub-

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–7 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

FIG. 4. Hierarchical clustering analysis of RES and NaOCl induced gene expression profiles. Log2-fold changes of gene expression ratios were clustered for 1 mM diamide (Dia), 2.4 mM catechol (Cat), 0.5 mM methylhydroquinone (MHQ), 1 mM formaldehyde, 2.8 mM and 5.6 mM methylglyoxal (MG-2 and MG-5), and 50 ␮M NaOCl stress using the Treeview software. The cluster analysis resulted in 14 different nodes that are enriched for RES and NaOCl-induced regulons (CtsR, Spx, ArsR, CzrA, CsoR, SigmaD), the RES-specific CymR regulon, quinone- responsive regulons (PerR, YodB, CatR (YvaP), MhqR), aldehyde-responsive regulons (HxlR, AdhR, LexA) and NaOCl-specific regulons (SigmaB, OhrR). For the cluster analysis 630 genes were selected as listed in supplemental Table S4 that are induced by RES and NaOCl stress. Red indicates induction and green repression under the stress specific conditions. unit of the pyruvate dehydrogenase complex (PdhD) forms an redox proteome (Fig. 5A) nor during tryptic digestion in control active site Cys47-Cys52 intramolecular disulfide used for oxi- extracts indicating that no oxidation occurred during our sam- dation of the dihydrolipoamide to lipoamide (64). The adenylate ple preparation protocol. The intramolecular C152-C156 disul- kinase Adk and the topoisomerase 1 (TopA) form intramolecular fide bond in GapA‘s catalytic center was detected as triply disulfides in Zn binding sites (Table I). Further proteins with charged peptide YDAANHDVISNASC152(-2)TTNC156LAPFAK intramolecular disulfides include those with Fe-S clusters or at an m/z ϭ 842,0466 (supplemental Fig. S2B). Fe-binding sites, such as the glutamate synthase large subunit The intramolecular disulfides for Spx and PerR were iden- GltA (GOGAT), the sulfite reductase flavoprotein alpha subunit tified only in cell extracts of NaOCl-treated bshA mutant cells. CysJ (YvgR) and the Fe-S cluster assembly protein YutI. The redox-sensing regulator Spx is activated by a thiol-

Three redox-sensitive proteins were identified that are oxi- disulfide redox switch in the N-terminal C10xxC13 motif in dized to intramolecular disulfides only under NaOCl stress response to diamide stress (65, 66). This C10-C13 intramo- conditions, including GapA, PerR, and Spx (Table I). GapA is lecular disulfide was identified as doubly charged peptide most strongly oxidized in response to NaOCl stress in the M1(ϩ16)VTLYTSPSC10(-2)TSC13R at an m/z ϭ 781,8378 in redox proteome (Fig. 5B). GapA was neither oxidized in the NaOCl-treated cells (supplemental Fig. S2F). The peroxide-

10.1074/mcp.M111.009506–8 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–9 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis sensing PerR repressor is inactivated in response to hydrogen phosphoglycerate dehydrogenase SerA (supplemental Fig. peroxide by iron-catalyzed His-oxidation (14). The intramolec- S3E, Table II) and the thiol-disulfide oxidoreductase YphP ular disulfide was detected in the structural Zn-binding site of (supplemental Fig. S3F, Table II). The MetE protein was iden-

PerR as doubly charged peptide LEIYGVC136QEC139(-2)SK at tified as specific target for reversible thiol-oxidation by NaOCl an m/z ϭ 685,3092 (supplemental Fig. S2E). This suggests stress in the redox proteome (Fig. 5B,5C). Moreover, MetE, that PerR senses disulfide stress probably via oxidation of the PpaC and SerA have been shown to be modified by S- Cys residues in the Zn binding site. cysteinylations in response to diamide stress (67). Identification of S-bacillithiolations in OhrR, MetE, YxjG, For MetE we detected the triply charged peptide VPS- PpaC, SerA, and YphP in the Proteome of NaOCl-treated TEEMYNIIVDALAVC719(ϩBSH)PTDR at an m/z ϭ 944.7604 Cells Using Shotgun-LC-MS/MS Analysis—The OhrR repres- and the doubly charged peptide FWVNPDC730(ϩBSH)- sor is inactivated by an S-thiolation mechanism including GLK at an m/z ϭ 787,8297 containing the additional mass of cysteine and BSH in response to CHP (7, 19). Our data BSH (ϩ396 Da) at Cys719 and Cys730 (Fig. 5F,5G; showed that NaOCl stress also inactivates the OhrR repressor supplemental Fig. S3B, Table II). In addition, the triply charged because ohrA transcription was derepressed. Thus, we ana- Cys730 peptide with an S-cysteinylation modification was lyzed whether OhrR forms mixed disulfides with cysteine or identified as peptide FWVNPDC730(ϩCys)GLK at an m/z ϭ BSH in response to NaOCl. Because OhrR is a low abundant 433,1952 containing the additional mass of Cys (ϩ119 Da) regulatory protein, a FLAG-epitope-tagged OhrR protein was (supplemental Fig. S3B; Table II). Interestingly, the CID enriched using immunoprecipitation (IP) from NaOCl-treated MS/MS spectra of the bacillithiolated MetE peptides revealed cells as in previous studies (19). Purified OhrR-FLAG protein a characteristic loss of malate from the precursor ions and was tryptically digested and analyzed by LTQ-Orbitrap MS/MS from abundant y-fragment ions as indicated by the loss of 134 analysis as described in the Methods section. In protein sam- Da. The precursor ions minus 134 Da are most abundant in ples containing OhrR-FLAG protein from NaOCl-treated cells, the MS/MS spectra of the bacillithiolated peptides identified the Cys15-peptide was identified as triply charged peptide at an for MetE, PpaC and SerA. Thus, the bacillithiolated and cys- m/z ϭ 684.98 containing the additional mass of BSH (ϩ396 Da) teinylated Cys730 peptides of MetE showed different MS/MS (supplemental Fig. S3A,TableII).ThisindicatesthatOhrRis inactivated by an S-bacillithiolation mechanism in response to fragment ion spectra (supplemental Fig. S3B). Moreover, the NaOCl stress. To verify that only Cys15 of OhrR is involved in abundance of the precursor ions minus malate leads also to redox-sensing of NaOCl stress, we analyzed ohrA transcription decreased intensities of the b and y ions in the MS/MS spec- in an ohrRC15S mutant (34). The Northern blot results in tra of the bacillithiolated PpaC and SerA peptides supplemental Fig. S3A indicate that ohrA transcription is com- (supplemental Fig. S3D, 3E). pletely abolished in the ohrRC15S mutant. This confirms that For YxjG, we detected the triply charged peptide YVSLD- regulation of ohrA transcription requires only inactivation of the QLC341(ϩIAM)LSPQC346(ϩBSH)GFASTEEGNK at an m/z ϭ redox-sensing Cys15 of OhrR. 981,4183 with the additional mass of 396 Da at Cys346 Next, we searched our LC-MS/MS results obtained from (supplemental Fig. S3C, Table II). YxjG is homolog to the whole proteome tryptic digests of NaOCl-treated cells for C-terminal part of MetE and the essential Zn-binding Cys730 S-thiolation modifications. We identified five cytoplasmic pro- residue of MetE aligns with Cys346 in YxjG. This suggests teins that were modified by S-bacillithiolation, including two that also Cys346 of YxjG could be essential for methionine methionine synthase paralogs MetE and YxjG (Fig. 5F,5G, synthase activity. Thus, homolog active site Cys residues are supplemental Fig. S3B, 3C, Table II), the inorganic pyrophos- S-bacillithiolated in the paralogous methionine synthases phatase PpaC (supplemental Fig. S3D, Table II), the 3-D- MetE and YxjG.

FIG. 5. ABC. The thiol-redox proteome (red) in comparison to the protein amount image (green) at control conditions (A) and after exposure to 50 ␮M NaOCl (B) in the wild type. Reduced protein thiols in cell extracts were alkylated with IAM followed by reduction of oxidized protein thiols with TCEP and labeling with BODIPY FL C1-IA. Proteins with reversible thiol-modifications in the control and NaOCl redox proteome are labeled in white and newly oxidized proteins in NaOCl-treated cells are labeled in red. The oxidized proteins were identified by MALDI-TOF-TOF MS/MS as shown in detail in supplemental Fig. S1A–P. C, The fluorescence/protein amount ratios are quantified as oxidation ratios at control conditions (control) and 10, 20, and 30 min after exposure to 50 ␮M NaOCl stress as shown in the diagram. DEFG. Close-ups of the main NaOCl-sensitive proteins MetE and GapA in the thiol-redox proteome of the wild type (D) and bshA mutant (E) and CID MS/MS spectra of the S-bacillithiolated Cys719 and Cys730 peptides of MetE (F, G). Figs. D and E show sections of reversibly oxidized proteins after NaOCl stress in the thiol-redox proteome (red) in comparison to the protein amount image (green) at control conditions (co) and 10, 20, and 30 min after exposure to 50 ␮M NaOCl. Figs. F and G show the CID MS/MS spectra of the S-bacillithiolated Cys719-and Cys730-peptides of MetE identified in the wild-type proteome using LTQ-Orbitrap-Velos mass spectrometry as described in the Methods section. The MS/MS spectra show the characteristic abundant precursor ions that have lost malate indicated by parent-134

(HOOC-CH2-CHOH-COOH). The Xcorr and ⌬Cn scores and peptide masses are listed in Table II and the corresponding b and y fragment ion series for the modified peptides are given in detail in supplemental Fig. S3B.

10.1074/mcp.M111.009506–10 Molecular & Cellular Proteomics 10.11 oeua ellrPoemc 10.11 Proteomics Cellular & Molecular

TABLE I Proteins with intramolecular disulfides identified at control and NaOCl stress conditions. Cytoplasmic proteins were harvested in IAM-urea/thiourea-buffer, separated using SDS-PAGE, tryptically digested and analyzed in a LTQ Orbitrap-Velos mass spectrometer as described in the Methods section. Peptides with intramolecular disulfides that showed a mass difference of Cys-2 Da were identified for Adk, GltA, CysJ, PdhD, TopA, YdjI, YutI, and Zwf in control and NaOCl samples. The table includes the m/z of the precursor ions, charges, Xcorr and ⌬Cn scores of the disulfide linked peptides. The complete CID MS/MS spectrum of all peptides and the b and y fragment ion series are given in supplemental Fig. S2A–K as indicated in column SI-Fig.

m/z precursor ⌬Cn SI-Fig. Protein Function Redox-active Cys Disulfide-Peptide ion XCorr-score score Charge

b S2A Adk Adenylate kinase Zn-binding: Cys130,133, IC130 SVC133 (-2)GTTYHLVFNPPK 938,9590 4,6515 0,6794 2 150,153 S2B GapAa,b Glyceraldehyde 3-phosphate Cys152 redox-active catalytic YDAANHDVISNASC (-2) 842,0466 5,8694 0,7600 3 152 in Resistance Hypochlorite Confers S-Bacillithiolation dehydrogenase site TTNC156 LAPFAK S2C GltA Glutamate synthase large subunit 3Fe-4S: C113, 1119, 1124 AC1119 (-2) 674,6567 5,2172 0,7056 3 HLDTC1124 PVGVATQNPELR b S2D PdhD dihydrolipoamide dehydrogenase Cys47 and Cys52 redox-active ATLGGVC47 (-2)LNVGC52 IPSK 765,3950 3,6256 0,5999 2 E3 subunit of pyruvate disulfide as catalytic centre dehydrogenase a S2E PerR Fur-family repressor of the Zn-binding site:Cys96,99, LEIYGVC136 QEC139 (-2)SK 685,3092 2,6948 0,7523 2 peroxide regulon 136,139 a S2F Spx Regulator of the disulfide stress Cys10, 13 redox-active sites M1(ϩ16)VTLYTSPSC10 (-2)TSC13 R 781,8378 2,9770 0,7398 2 response S2G TopA DNA topoisomerase 1 3 Zn-fingers: Cys579,582, 599,605 Cys619,622,641,647 C619 PSC622 (-2)GEGNIVER 631,2697 2,3857 0,7171 2 Cys660,663,680,683 S2H YdjI Unknown Unknown TGEANFC315 PNC318 (-2)GQK 683,7802 2,8661 0,6388 2 S2I YutI Putative nitrogen fixation protein, Cys79 and Cys82 Fe-S-cluster LLGAC79 GSC82 (-2)PSSTITLK 774,8928 4,3033 0,6283 2 Fe-S cluster assembly or repair binding b S2J YvgR (CysJ) Sulfite reductase ͓NADPH͔ Cys427 and Cys431 probably GVC427 (-2)SILC431 AER 524,7494 2,1040 0,5121 2 flavoprotein alpha-component Fe-binding

10.1074/mcp.M111.009506–11 S2K Zwf Glucose-6-phosphate Unknown LDYC407 (-2)SNC410 NDELNTPEAYEK 1109,9501 4,8708 0,8275 2 dehydrogenase a The intramolecular disulfide of GapA was detected in NaOCl-treated wild-type and bsh mutant cells and the disulfides for PerR and Spx were detected in NaOCl-treated bsh mutant cells. b Adk, GapA, PdhD, and YvgR (CysJ) were also identified in the redox proteome as reversible oxidized proteins (Fig. 5A,5B). .subtilis B. 10.1074/mcp.M111.009506–12 in Resistance Hypochlorite Confers S-Bacillithiolation

TABLE II Proteins with S-Bacillithiolations and S-Cysteinylations identified in NaOCl-treated cells. Cytoplasmic proteins of the wild type were harvested in IAM-buffer, separated using SDS-PAGE, tryptically digested and analyzed in a LTQ Orbitrap-Velos™ mass spectrometer as described in the Methods section. Peptides with S-bacillithiolations were identied for OhrR, MetE, YxjG, PpaC, SerA, and YphP indicated by the additional mass of 396 Da. Peptides with S-cysteinylations were identified for MetE and PpaC with a mass difference of 119 Da. The table includes the m/z of the precursor ions, charges, Xcorr, and ⌬Cn scores of the S-thiolated peptides. The complete CID MS/MS spectrum of all peptides and the b and y fragment ion series are given in supplemental Fig. S3A–F as indicated in column SI-Fig. m/z ⌬Cn SI-Fig. Protein Function Redox-active Cys S-thiolated Peptide XCorr-score Charge precursor ion score

S3A OhrR Redox-sensing MarR-type Cys15 redox-sensing LENQLC15 (ϩBSH)FLLYASSR 684,9800 2,9444 0,5816 3 repressor for organic hydroperoxide resistance a S3B MetE Cobalamin-independent Cys 647, 730 essential, Zn-binding VPSTEEMYNIIVDALAVC719 (ϩBSH)PTDR 944,7604 4,9737 0,7545 3 methionine synthase active site in B. sub. Cys719 non-essential, FWVNPDC730 (ϩBSH)GLK 787,8297 2,8014 0,6097 2 S-cysteinylated by diamide in

B. sub. subtilis B. Cys645 non-essential, FWVNPDC730 (ϩCys)GLK 433,1952 3,2330 0,5435 3 S-glutathionylated in E. coli S3C YxjG Cobalamin-independent Cys251, 346 homolog to Zn- YVSLDQLC341 (ϩIAM)LSPQC346 - 981,4173 4,9686 0,7521 3 methionine synthase binding Cys647, 730 in MetE of (ϩBSH)GFASTEEGNK B. sub. oeua ellrPoemc 10.11 Proteomics Cellular & Molecular S3D PpaC inorganic pyrophosphatase Cys158 SPTC158 (ϩBSH)TDQDVAAAK 851,8456 3,4552 0,8838 2 S-cysteinylated by diamide in SPTC158 (ϩCys)TDQDVAAAK 713,3041 3,2977 0,8356 2 B. sub. S3E SerA 3-D-phosphoglycerate Cys410 S-cysteinylated by ISSSESGYDNC410 (ϩBSH)ISVK 992,9086 3,6448 0,8507 2 dehydrogenase diamide in B. sub. S3F YphP Thiol-disulfide isomerase Cys53 redox-active AEGTTLVVVNSVC53 (ϩBSH)GC55 - 815,3752 4,077 0,6036 3 (ϩIAM)AAGLAR a MetE was also identified as reversibly oxidized in the redox proteome after NaOCl stress. The identification of the essential Zn-binding ligands Cys647 and 730 of B. subtilis MetE is based on similarity to the E. coli MetE enzyme and derived from the UniprotKB database entry P80877. S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

The inorganic pyrophosphatase PpaC was S-bacillithio- that the OhrA peroxiredoxin and the BSH redox buffer play lated at Cys158 and the same site was also S-cysteinylated most essential roles in NaOCl detoxification. by diamide (67). For PpaC the Cys158-containing doubly The Growth Defect in Response to NaOCl Stress can be charged peptide SPTC158(ϩBSH)TDQDVAAAK was detected Attributed to Methionine Limitation—The methionine synthase at an m/z ϭ 851,8456 Da with the additional mass of 396 Da MetE catalyzes the final step in the methionine biosynthesis, (supplemental Fig. S3D, Table II). We also detected the doubly the chemically difficult methyl group transfer of methyltetra- charged S-cysteinylated Cys158-containing peptide of PpaC hydrofolate (N5-THF) to homocysteine (70) (Fig. 7). Diamide

SPTC158(ϩCys)TDQDVAAAK at an m/z ϭ 713,3041 indicated stress leads to methionine auxotrophy that is caused by MetE by the additional mass of 119 Da (supplemental Fig. S3D, inactivation via S-cysteinylation in B. subtilis and S-glutathio- Table II). Thus, for MetE and PpaC S-bacillithiolations and nylation in E. coli (38, 67, 70, 71). S-bacillithiolation of the S-cysteinylations were observed. active site Cys residues of MetE and YxjG most likely leads to The phosphoglycerate dehydrogenase SerA was S-bacilli- enzyme inactivation and methionine auxotrophy by NaOCl thiolated at Cys410 in response to NaOCl stress and the same stress. This methionine starvation phenotype was supported site was S-cysteinylated by diamide stress (67). The S-bacil- by the induction of the S-box regulon genes in the transcrip- lithiolated Cys410-peptide ISSSESGYDNC410(ϩBSH)ISVK tome (Fig. 7, supplemental Table S1). Using Northern blot was detected as doubly charged peptide at an m/z ϭ analysis the transcriptional induction of the S-box regulon 992,9086 with the additional mass of 396 Da (supple- gene yitJ was verified during the time of starvation after ex- mental Fig. S3E, Table II). posure to 50 ␮M NaOCl (Fig. 7). Interestingly, the thiol-disulfide isomerase YphP (68) was Furthermore, growth experiments were performed in the identified as S-bacillithiolated at the redox-active Cys53 presence of extracellular methionine to confirm the methio- residue. The triply charged peptide AEGTTLVVVNSVC53- nine starvation phenotype. Cells were treated with of 75 ␮l

(ϩBSH)GC55(ϩIAM)AAGLAR was observed at an m/z ϭ NaOCl and 30 or 60 min later 75 ␮M extracellular methionine 815,3752 with the additional mass of 396 Da at Cys53 and the was added. The growth was resumed immediately after me- carbamidomethylation at Cys55 (supplemental Fig. S3F, thionine addition (Fig. 8A). To control that the added methio- Table II). nine does not simply remove the remaining oxidants from the The OhrA Peroxiredoxin and the BSH Redox Buffer are supernatant, the NaOCl concentrations in the culture super- Essential for Protection Against NaOCl Stress—Next, we an- natants were monitored using the FOX-assay. The results alyzed if the growth of ohrA and ohrR mutant strains is af- showed that 66% of NaOCl is consumed and detoxified by fected by NaOCl treatment. The growth of an ohrA mutant the cells after 30 min and 93% after 60 min supporting that the was strongly impaired by 50 ␮M NaOCl compared with that of added methionine abolished the methionine starvation phe- the wild type whereas the ohrR mutant was more resistant to notype (Fig. 8B). In another experiment, cells were inoculated 75 ␮M NaOCl (Fig. 6). Thus, the ohrA encoded peroxiredoxin in minimal medium containing 75 ␮M methionine, grown to an is a specific determinant of NaOCl resistance in B. subtilis and OD500 of 0.4 and treated with 75 ␮M NaOCl. Cells were able to perhaps involved in detoxification of this strong oxidant. grow without any lag phase in the methionine-supplemented Next, we monitored the growth of bshA and bshB1B2 mu- medium after treatment with 75 mM NaOCl (Fig. 8C). This tants with defects in the BSH biosynthesis enzymes (35). The indicates that NaOCl stress causes methionine auxotrophy growth of NaOCl-treated bshA and bshB1B2 mutants was that can be abolished by methionine addition. strongly impaired compared with the wild type (Fig. 6). While B. subtilis contains about 1 ␮mol/g BSH and 0.58 ␮mol/g the growth of the wild type was resumed 60 min after treat- Cys as redox buffers (35). To analyze whether Cys can replace ment with 50 ␮M NaOCl, the bsh mutants required 180 min to BSH in post-translational modification and MetE inactivation, resume growth. This NaOCl-sensitive phenotype of bsh mu- bshA mutant cells were exposed to 60 ␮M NaOCl and 30 and tants points to a major role of BSH in NaOCl detoxification 60 min later 60 ␮M methionine was added. Methionine was similar as glutathione (GSH) in E. coli (69). able to restore also the growth of bshA mutant cells indicating Finally, we investigated whether the PerR, Spx and SigmaB that the growth defect of bshA mutants could be attributed to regulons confer protection against NaOCl. However, the perR methionine auxotrophy (supplemental Fig. S4A). The tran- mutant that overproduces the catalase KatA and the alkylhy- scriptional induction of the S-box regulon gene yitJ in the droperoxide reductase AhpCF was not resistant to NaOCl bshA mutant by 50 ␮M NaOCl stress supports the methionine stress (data not shown). The spx mutant displayed increased starvation phenotype (Fig. 7). In addition, the redox proteome sensitivities toward NaOCl stress, but the growth was also of NaOCl-treated bshA mutant cells revealed similar MetE impaired under non-stress conditions due to the pleiotropic oxidation ratios as observed for the wild type (Fig. 5E). Using phenotype of the spx mutant (Fig. 6). The growth of the sigB the shotgun LC-MS/MS approach we identified the S-cystei- mutant was only slightly impaired by NaOCl indicating per- nylated Cys730-peptide of MetE in the bshA mutant proteome haps a role of the SigmaB-controlled OhrA-paralog OhrB in (supplemental Fig. S5). These results indicate that Cys can NaOCl detoxification. These results lead to the conclusion replace BSH for post-translational modifications and inactiva-

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–13 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

FIG. 6. The OhrA peroxiredoxin and BSH protect cells against NaOCl toxicity. Growth phenotype of B. subtilis wild type (wt) in comparison to the ⌬ohrR (A), ⌬ohrA (B), ⌬bshA (C), ⌬bshB1,B2 (D), ⌬sigB (E), and ⌬spx (F) mutant strains that were treated with 50 or 75 ␮M

NaOCl at an OD500 of 0.4. tion of MetE. However, the bshA mutant was strongly impaired also in host tissue damage and diseases. The redox proteome in NaOCl detoxification and consumption compared with the in response to NaOCl has been recently studied in E. coli wild type because about 50% of NaOCl was left in the medium using the OxICAT approach and several redox-sensitive pro- after 60 min in the bshA mutant (supplemental Fig. teins could be identified that are sensitive to NaOCl-directed S4B). This indicates that BSH is more efficient in NaOCl detox- reversible oxidation (28, 29). In B. subtilis, only few NaOCl- ification than Cys. sensitive proteins were identified in the 2D gel-based thiol- redox proteome. One reason for the observed differences in DISCUSSION the redox proteome analyses between E. coli and B. subtilis Hypochloric acid is widely used as disinfectant and pro- could be that only abundant cytoplasmic proteins with revers- duced in activated macrophages by the enzyme myeloperoxi- ible thiol-modifications are visualized by the 2D gel-based dase as first defense line upon bacterial infections. Hypochlo- approach (e.g. MetE and GapA) and that the gel-free OxICAT ric acid is beneficial by killing invading bacteria but can result approach is much more sensitive for identification of oxidized

10.1074/mcp.M111.009506–14 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

FIG. 7. Pathways for sulfur assimilation, cysteine and methionine biosynthesis, and induction of S-box regulon genes in response to NaOCl stress. The S-box regulon gene products that are induced in the transcriptome as a result of methionine starvation provoked by the S-bacillithiolation of MetE and YxjG are bold-faced in gray. The transcriptome induction ratios are shown below the gene products. The gene products of the sulfate assimilation pathway (Sat, CysC, CysH, CysJI, CysE) are not induced upon NaOCl stress (except for CysK). The inset shows the transcriptional induction of yitJ in the wild type and ⌬bshA mutant using Northern blot analysis before (co) and at different times (10, 30, 60, 90, and 120 min) after exposure to 50 ␮M NaOCl stress. Abbreviations: APS, adenosine-5Ј-phosphosulfate; PAPS, 3Ј-phosphoad- enosine-5Ј-phosphosulfate; OAS, O-acetylserine; N5,N10-THF, 5,10-methylenetetrahydrofolate; N5 -THF, 5-methyltetrahydrofolate; THF, tetrahydrofolate. This figure was adapted from Tomsic et al., 2008 (52). proteins of lower abundance. However, previous 2Dgel-based GapA protein. This confirms previous results in E. coli using redox proteomics analyses in B. subtilis have shown that only the OxICAT approach suggesting that GapA is oxidized to an toxic peroxide concentrations cause increased reversible intramolacular disulfide by NaOCl stress (28). GapDH activity thiol-oxidations and that only few proteins are oxidized upon was also inactivated by NaOCl in endothelial cells accompa- paraquat stress (38). Moreover, no reversible thiol-modifica- nied by thiol depeletion (74). GapA has been shown as redox- tions could be monitored using the 2D gel-based approach in controlled cytoplasmic enzyme in prokaryotes and yeast and response to nitric oxide (NO) in B. subtilis (72). In contrast is susceptible also for inactivation by S-sulfenylation, S-sul- again, several targets for reversible thiol-modifications were fonylation, S-glutathionylation, S-nitrosylation, and intermo- identified by NO stress in E. coli by the 2D gel-based ap- lecular aggregation in response to ROS and RNS (13). GapA proach (73). Thus, there might be differences in the extents of is also strongly oxidized by diamide and quinones in B. subtilis reversible thiol-modifiations in response to ROS and RNS (21–22). The reversible inactivation of GapA‘s glycolytic activ- between E. coli and B. subtilis. To address this point, more ity in B. subtilis by the oxidative mode of quinones has been sensitive gel-free quantifications of reversibly oxidized pro- shown in previous studies (21). Inactivation of GapA leads teins using the OxICAT approach are required in the future to probably to re-routing of glucose-6-phosphate to the pentose quantify the changes in the redox proteome of B. subtilis more phosphate pathway. The function of the pentose phosphate comprehensively. Nevertheless, we show here that NaOCl pathway is to increase NADPH levels and provide reducing leads to S-bacillithiolation of selective proteins in B. subtilis equivalents for the thioredoxin/thioredoxin reductase system that play essential roles in protection against hypochloric acid to restore the redox balance of the cell (13). toxicity. S-bacillithiolation Protects Metabolic Enzymes Against Irre- GapA is Oxidized to an Intramolecular Disulfide Upon versible Overoxidation—We further demonstrate that six cy- NaOCl Stress—One of the strongest targets that forms intra- toplasmic proteins undergo reversible S-bacillithiolations by molecular disulfides upon NaOCl stress was identified as the NaOCl treatment. Protein S-thiolation is an important post-

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–15 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

FIG. 8. NaOCl stress causes methionine auxotrophy that is abolished by methionine addition. Growth phenotypes of the B. subtilis wild type treated with 75 ␮M NaOCl at an OD500 of 0.4. A, Methionine was added 30 or 60 min after exposure to 75 ␮M NaOCl stress and the growth was resumed. B, The concentrations of consumed NaOCl of wild-type cells were monitored in the culture supernatants after exposure to 75 ␮M NaOCl using the FOX assay. The NaOCl concentrations are given as mean values of three independent experiments with error bars. C,

Methionine (75 ␮M) was added after inoculation to the culture at an OD500 of 0,07 and 75 ␮M NaOCl was added when cells had reached an OD500 of 0.4. translational thiol-modification that occurs in response to inactivated by S-bacillithiolation after CHP challenge (19). oxidative stress in eukaryotic and prokaryotic cells and Here we found that S-bacillithiolation in response to NaOCl controls metabolic processes and redox-sensing transcrip- controls OhrR activity resulting in expression of the OhrA tion factors to counteract the cellular damage. In eu- peroxiredoxin as protection mechanism (Fig. 9). This ex- karyotes, S-glutathionylation is implicated in disease mech- pands to role of the thiol-dependent OhrA-like peroxiredox- anisms, including cell-signaling pathways associated with ins in detoxification of hypochloric acid. OhrA-like peroxire- viral infections and with tumor necrosis factor ␣-induced doxins catalyze the reduction of ROOH to their apoptosis (75). In E. coli the activity of the oxidative stress corresponding alcohols (79). The structure of the Pseu- responsive regulator OxyR is controlled by reversible S- domonas Ohr peroxiredoxin has been resolved, which is a glutathionylation in vitro (11). Moreover, the activities of homodimer consisting of a tightly folded ␣/␤ fold with two several metabolic enzymes, such as glyceraldehyde-3- active site cysteines located at the monomer interface on phosphate dehydrogenase, methionine synthase, and the opposite sites of the molecule (80, 81). The mechanism of PAPS reductase are inhibited by S-glutathionylation in hydroperoxide reduction is similar to the structurally unre- E. coli (13, 70, 76). Protein S-thiolation is thought to protect lated eukaryotic 2-Cys peroxiredoxins that exhibit very high active site Cys residues of essential enzymes against irre- specificity and reaction rates with peroxides (81). The cata- versible overoxidation to sulfonic acids. Hypochloric acid lytic mechanism of peroxiredoxins involves the attack of the shows very fast reaction rates with Cys residues (K2 ϭ 3 ϫ hyproperoxide substrate by the active site Cys thiolate (the 7 Ϫ1 Ϫ1 10 M s )thataresevenordersofmagnitudehigherthan peroxidatic Cys) that is oxidized to a sulfenic acid intermediate measured for peroxides and rapidly lead to irreversible ox- with the release of the alcohol. The peroxiredoxin sulfenic acid idation products, such as sulfinic and sulfonic acids (3, 4, undergoes intersubunit or intramolecular disulfide bond forma- 77, 78). Thus, S-bacillithiolation in B. subtilis serves to pro- tion with the resolving Cys located in the same or another tect active site Cys residues of key metabolic enzymes subunit of the dimer. The enzyme is regenerated by thiol-disul- against irreversible overoxidation by the strong oxidant hy- fide exchange with GSH or protein electron donors (82). Our pochloric acid. results show that OhrA confers specific protection against The OhrA Peroxiredoxin and the BSH Redox Buffer Confer NaOCl. This leads to the question by which mechanism OhrA is Protection Against NaOCl—In B. subtilis the OhrR repressor is able to detoxify hypochloric acid?

10.1074/mcp.M111.009506–16 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

FIG. 9. Proposed defensive mechanisms against hypochloric acid stress in B. subtilis. Exposure of B. subtilis to NaOCl induces an oxidative, disulfide and general stress response (OhrR, Spx, CtsR, PerR, SigmaB). NaOCl leads to S-bacillithiolation of OhrR, MetE, YxjG, PpaC, SerA, and YphP and to intramolecular disulfide formation in GapA. The thiol-disulfide isomerase YphP could function as bacilliredoxin in reduction of S-bacillithiolated proteins. (1) The redox buffer BSH could be directly involved in NaOCl detoxification leading to BSSB formation. (2) S-bacillithiolation of OhrR causes induction of the OhrA peroxiredoxin that is involved in specific NaOCl detoxification. (3)

S-bacillithiolation of the inorganic pyrophosphatase PpaC could lead to decreased ATP sulfurylase activity as the removal of PPi is prevented. (4) S-bacillithiolation of the phosphoglycerate dehydrogenase SerA causes decreased levels of serine that is required for cysteine and methionine biosynthesis. (5) S-bacillithiolation of the active site Cys residues of MetE and YxjG leads to methionine auxotrophy to stop translation during the time of NaOCl detoxification (6) The glyceraldehyde-3-phosphate dehydrogenase GapA is inhibited causing decreased glycolysis. The stars indicate the redox-sensing Cys residues that are S-bacillithiolated in OhrR (Cys15), MetE (Cys730), YxjG (Cys346), or oxidized to an intramolecular disulfide in GapA (Cys152). Abbreviations: APS, adenosine-5Ј-phosphosulfate; N5-THF, 5-methyltetrahydrofo- late; THF, tetrahydrofolate; 3-PG, 3-D-phosphoglycerate, GA-3-P, glyceraldehyde-3-phosphate; 1,3-BPG, 1,3-Bisphosphoglycerate.

The reaction of hypochloric acid with Cys proceeds via Cys726 of E. coli MetE and with Cys620 and Cys704 of Ther- chlorination of the thiol group to form the unstable sulfenyl- motoga maritima MetE (84, 85). MetE is S-bacillithiolated at chloride intermediate that reacts with another thiol group to the non-essential Cys719 and the Zn-binding ligand Cys730 form a disulfide (3, 4). It could be possible that the OhrA and the methionine synthase paralog YxjG is S-bacillithiolated peroxiredoxin is attacked at the peroxidatic Cys by chlorina- at Cys346 that aligns with Cys730 in MetE. In addition, tion, resulting in OhrA sulfenylchloride formation and further Cys730 of MetE is also S-cysteinylated by NaOCl stress in the oxidation to the OhrA inter/intrasubunit disulfide between per- wild type as well as in the bshA mutant. The E. coli MetE oxidatic and resolving Cys residues (Fig. 9). The OhrA disul- protein is S-glutathionylated at the nonessential Cys645 in fide could be regenerated by the BSH redox buffer. response to diamide that is not conserved in B. subtilis MetE In addition, BSH likely plays a direct role in detoxification of (71). In E. coli MetE Cys645 is positioned at the entrance of hypochloric acid as has been shown in E. coli for GSH (69). the active Zn site within a cleft between two ␤8␣8 barrels and The reaction of GSH with hypochloric acid is spontaneous it‘s glutathionylation leads to conformational changes of the and does not involve conjugating enzymes such as glutathi- homocysteine binding active site (70, 85). In B. subtilis MetE one S-transferases (GSTs) (83). GSH is oxidized to GSSG by the non-essential Cys719 is about 20 Å apart from the active hypochloric acid nonenzymatically very efficiently as 1 Mol site Cys730. This indicates that thiol-disulfide exchange be- GSH reacted with 4 Mol hypochloric acid in vitro. Hypochloric tween Cys719 and Cys730 residues is not possible. The acid likely also oxidizes BSH directly to BSSB in B. subtilis. question arises how BSH get‘s access to the active site Zn Four Enzymes of the Methionine Synthesis Pathway (MetE, center in B. subtilis MetE ? Recent structural characterizations YxjG, PpaC, SerA) are S-Bacillithiolated by NaOCl Stress of the T. maritima MetE Zn center in the substrate-free and Leading to Methionine Auxotrophy—Besides OhrR, we iden- homocysteine-bound states have revealed an elastic nature tified four enzymes with S-bacillithiolations in response to of the catalytic Zn center upon substrate binding (84). Homo- NaOCl stress as MetE, YxjG, PpaC, and SerA. The methionine cysteine binding leads to an unexpected inversion of Zn ge- synthase MetE has two Zn-coordinating active-site Cys resi- ometry with displacement of the endogenous Zn ligand dues in positions 647 and 730 that align with Cys643 and Glu642 of MetE and movement of the Zn relative to the protein

Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–17 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis scaffold. It is proposed that this Zn geometry inversion en- isomerase YphP was identified as novel S-bacillithiolated pro- hances the nucleophilic activation of the homocysteine thio- tein. YphP is a member of the DUF1094 family with a con- late that is required for methyl transfer (84). The dynamic served CXC motif and a thioredoxin-like structure (68). It has nature and flexibility of the Zn center could also provide been shown that YphP functions as thiol-disulfide isomerase access for oxidation of the Zn ligand Cys730 by BSH. in vitro, but the specific substrate is unknown. Phylogenetical There are also structural differences between GSH and BSH studies suggest that YphP could be a novel bacilliredoxin that that could explain why BSH has access to the active site Zn co-occurs with the enzymes of the BSH biosynthesis pathway center. GSH consists of the tripeptide L-␥-glutamylcysteinylgly- across different genomes of low GC Gram-positive bacteria cine with the Cys bound N-terminally and C-terminally by glu- (35). Because YphP is S-bacillithiolated at the active site tamate and glycine residues that could preclude the accessabil- Cys53, it is likely that YphP might function in de-bacillithiola- ity of the Cys thiol to the active site in MetE of E. coli.The tion of MetE, YxjG, PpaC and SerA upon return to nonstress structure of BSH was determined as N-cysteinyl-␣-D-gluco- conditions in vivo. Thus, our proteome-wide studies have saminyl L-malate (20) with an N-terminally located Cys that discovered physiological substrates of the methionine biosyn- enables access of the thiol to the active site Zn center of MetE. thesis pathway as targets for S-bacillithiolation by NaOCl and Our transcriptional data and growth assays support the a candidate bacilliredoxin that might catalyze the reduction of methionine starvation phenotype of NaOCl-treated cells that these protein mixed disulfides. is caused by MetE and YxjG inactivation via S-bacillithiolation. Why Causes NaOCl S-Bacillithiolation and Diamide S-Cys- The inactivation of MetE by oxidative stress has important teinylation in the Proteome?—Previous proteome analyses consequences for the cell. The initiation of translation requires revealed that diamide causes S-cysteinylation in six cytoplas- formyl-Met as start methionine and depletion of methionine is mic proteins but no S-bacillithiolations (67). Three of these discussed as checkpoint to stop translation (70). The inacti- S-cysteinylated proteins (MetE, PpaC and SerA) were identi- vation of MetE by S-bacillithiolation could cause inhibition of fied as S-bacillithiolated by NaOCl. This raises the question translation to prevent further protein damage allowing the cell why S-bacillithiolation was not observed by diamide stress? to detoxify the oxidant and to restore the thiol redox homeo- The answer could be the different thiol chemistries of diamide stasis. Another possibility could be that MetE inactivation and NaOCl. Diamide is an electrophilic azocompound that serves to increase cysteine levels. This has been discussed in does not oxidize redox-sensitive Cys residues to reactive response to diamide stress that leads to MetE inactivation via sulfenic acids or sulfenylchloride. This is supported by the fact S-cysteinylation (67). The CymR-controlled cystathionine that diamide stress does not lead to inactivation of the OhrR beta-synthase MccA and cystathionine lyase MccB are in- repressor (Fig. 3), which would require oxidation of the Cys15 volved in the methionine-to cysteine conversion (Mcc) path- thiolate which then undergoes S-bacillithiolation. Instead, di- way (51, 86). The mccAB operon is strongly up-regulated by amide forms directly disulfide bonds between two Cys-thiols diamide stress supporting that homocysteine is converted to via an Michael addition reaction of the electrophilic azogroup cysteine via this Mcc pathway. However, the mccAB operon with two nucleophilic Cys thiol groups (89). Thus, S-bacilli- was not induced by NaOCl stress, indicating that S-bacilli- thiolation requires probably strong oxidants such as NaOCl thiolation of MetE leads to methionine starvation, but not to that oxidize peroxidatic Cys residues first to reactive sulfenic conversion of homocysteine to cysteine. acids or sulfenylchloride. Besides MetE and YxjG, the pyrophosphatase PpaC and Another question is why S-cysteinylation and S-bacillithio- the 3-D-phosphoglycerate dehydrogenase SerA were identi- lation was observed after NaOCl stress ? The observed S- fied as S-bacillithiolated that also function in the methionine cysteinylations could originate from S-bacillithiolations that biosynthesis pathway. PpaC is an essential and conserved could be explained by the BSH biosynthesis pathway. The enzyme that catalyzes the hydrolysis of inorganic pyrophos- pathways for BSH biosynthesis and degradation have been phate (PPi). Pyrophosphate is generated in a number of ATP- identified in B. subtilis and the structures of the L-malic acid driven cellular processes, including also the ATP sulfurylation glycosyltransferase BaBshA and the GlcNAc-Mal deacetylase as first step of the sulfate assimilation pathway (87, 88). Effi- BaBshB have been resolved in B. anthracis (35, 90). There are cient removal of PPi is required to drive the forward direction two GlcNAc-Mal deacetylases in B. subtilis, BshB1 and in these reactions. Thus, inhibition of PpaC activity by S- BshB2 with similar activities. It is postulated that one of these thiolation could further contribute to methionine starvation. deacetylases could function as bacillithiol-S-conjugate ami- SerA is the first enzyme in the L-serine biosynthetic pathway dase (likewise to the Mca mycothiol-S-conjugate amidase in catalyzing the conversion of 3-D-phosphoglycerate to 3-phos- Mycobacteria) to cleave conjugates formed by reaction of phonooxypyruvate with the generation of NADH. Since serine electrophiles with BSH (35, 91, 92). In turn, BshB1 and/or is used as precursor for cysteine biosynthesis, it‘s inactivation BshB2 could also cleave GlcN-Mal from S-bacillithiolated could pronouce the methionine starvation phenotype. proteins leaving S-cysteinylated proteins. Thus, S-cysteiny- The S-Bacillithiolated Disulfide Isomerase YphP Could lated proteins observed after NaOCl stress could originate Function as Putative Bacilliredoxin—The putative disulfide from S-bacillithiolations that are cleaved by BshB in vivo (91).

10.1074/mcp.M111.009506–18 Molecular & Cellular Proteomics 10.11 S-Bacillithiolation Confers Hypochlorite Resistance in B. subtilis

The CID MS/MS spectra of the S-bacillithiolated peptides also 3. Davies, M. J. (2011) Myeloperoxidase-derived oxidation: mechanisms of point to a fragmentation of BSH itself because characteristical biological damage and its prevention. J. Clin. Biochem. Nutr. 48, 8–19 4. Hawkins, C. L., Pattison, D. I., and Davies, M. J. (2003) Hypochlorite- fragment and precursor ions appeared that have lost malate. induced oxidation of amino acids, peptides and proteins. Amino Acids Thus, it is possible that further loss of GlcN in the collision cell 25, 259–274 leads to “artificial” S-cysteinylated peptides as a result of 5. Faulkner, M. J., and Helmann, J. D. (2011) Peroxide stress elicits adaptive changes in bacterial metal ion homeostasis. Antioxid Redox Signal. doi fragmentation. However, the redox buffer cysteine can be 10.1089/ars.2010.3682 also directly used for S-cysteinylations as shown by identifi- 6. Mongkolsuk, S., and Helmann, J. D. 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Molecular & Cellular Proteomics 10.11 10.1074/mcp.M111.009506–21 ! ! "#$%&'(!)! ! ! ! ! ! ! ! ! ! ! ! ! ! ! Chapter 3 ! ! ! ! ! S-Bacillithiolation protects conserved and essential proteins against hypochlorite stress in Firmicutes bacteria

! ! Bui Khanh Chi, Alexandra A. Roberts, Tran Thi Thanh Huyen, Katrin Bäsell, Dörte Becher, Dirk Albrecht, Chris J. Hamilton, and Haike Antelmann*.

Antioxidants and Redox Signaling, doi:10.1089/ars.2012.4686 (2012).

! ! ! ! ! ! ! ! ! ! * corresponding author ! !

! "#! & REDOX SIGNALING Volume 00, Number 00, 2012 ORIGINAL RESEARCH COMMUNICATION ª Mary Ann Liebert, Inc. DOI: 10.1089/ars.2012.4686

S-Bacillithiolation Protects Conserved and Essential Proteins Against Hypochlorite Stress in Firmicutes Bacteria

Bui Khanh Chi,1 Alexandra A. Roberts,2 Tran Thi Thanh Huyen,1 Katrin Ba¨sell,1 Do¨rte Becher,1 Dirk Albrecht,1 Chris J. Hamilton,2 and Haike Antelmann1

Abstract

Aims: Protein S-bacillithiolations are mixed disulfides between protein thiols and the bacillithiol (BSH) redox buffer that occur in response to NaOCl in Bacillus subtilis. We used BSH-specific immunoblots, shotgun liquid chromatography (LC)–tandem mass spectrometry (MS/MS) analysis and redox proteomics to characterize the S- bacillithiolomes of B. subtilis, B. megaterium, B. pumilus, B. amyloliquefaciens, and Staphylococcus carnosus and also measured the BSH/oxidized bacillithiol disulfide (BSSB) redox ratio after NaOCl stress. Results: In total, 54 proteins with characteristic S-bacillithiolation (SSB) sites were identified, including 29 unique proteins and eight proteins conserved in two or more of these bacteria. The methionine synthase MetE is the most abundant S- bacillithiolated protein in Bacillus species after NaOCl exposure. Further, S-bacillithiolated proteins include the translation EF-Tu and aminoacyl-tRNA synthetases (ThrS), the DnaK and GrpE chaperones, the two-Cys peroxiredoxin YkuU, the ferredoxin–NADP+ oxidoreductase YumC, the inorganic pyrophosphatase PpaC, the inosine-5¢-monophosphate dehydrogenase GuaB, proteins involved in thiamine biosynthesis (ThiG and ThiM), queuosine biosynthesis (QueF), biosynthesis of aromatic amino acids (AroA and AroE), serine (SerA), branched-chain amino acids (YwaA), and homocysteine (LuxS and MetI). The thioredoxin-like proteins, YphP and YtxJ, are S-bacillithiolated at their active sites, suggesting a function in the de-bacillithiolation process. S- bacillithiolation is accompanied by a two-fold increase in the BSSB level and a decrease in the BSH/BSSB redox ratio in B. subtilis. Innovation: Many essential and conserved proteins, including the dominant MetE, were identified in the S-bacillithiolome of different Bacillus species and S. carnosus using shotgun-LC-MS/MS analyses. Conclusion: S-bacillithiolation is a widespread redox control mechanism among Firmicutes bacteria that protects conserved metabolic enzymes and essential proteins against overoxidation. Antioxid. Redox Signal. 00, 000–000.

Introduction mechanism, S-glutathionylation must meet several criteria: (i) reversibility, (ii) specificity to active site Cys, (iii) change he reduced cytoplasm is maintained by low-molecular- in protein function/activity, and (iv) induction by ROS or Tweight (LMW) thiol-redox buffers, such as glutathione reactive nitrogen species (RNS). S-glutathionylation also (GSH), present in eukaryotes and in Escherichia coli (34). serves as a form of GSH storage to prevent the export of Redox buffers protect bacteria against reactive oxygen oxidized glutathione disulfide (GSSG) under oxidative stress species (ROS) generated during respiration or produced by (8). Many eukaryotic proteins such as a-ketoglutarate dehy- activated macrophages during infections (1). ROS cause re- drogenase, glyceraldehyde 3-phosphate dehydrogenase, or- versible intramolecular and intermolecular disulfides of nithine d-aminotransferase, pyruvate kinase, heat-specific proteins as well as mixed disulfides between protein thiols chaperones, and regulatory proteins (c-Jun and NF-jB) are and LMW thiols (S-thiolations). In eukaryotes, protein S- reversibly inactivated or activated by S-glutathionylation (8, glutathionylation is the most important post-translational 21). In E. coli, the oxidative stress-responsive regulator OxyR thiol modification caused by ROS, and has been shown to is activated by S-glutathionylation in vitro (22), while the control redox-sensing transcription factors and to protect activities of glyceraldehyde-3-phosphate dehydrogenase, active-site cysteine (Cys) residues of essential metabolic methionine synthase, and 3¢-phosphoadenosine-5¢-phospho- enzymes against irreversible oxidation to Cys-sulfinic or sulfate reductase are inhibited by S-glutathionylation in sulfonic acids (8, 13, 51). As an effective redox control E. coli (3, 19, 29).

1Institute for Microbiology, Ernst-Moritz-Arndt-University of Greifswald, Greifswald, Germany. 2School of Pharmacy, University of East Anglia, Norwich Research Park, Norwich, United Kingdom.

1 2 CHI ET AL.

Results Innovation Defining the conditions of NaOCl stress Recently, we identified six S-bacillithiolated proteins in to induce S-bacillithiolations in the proteome Bacillus subtilis in response to NaOCl stress (7). The me- of B. amyloliquefaciens, B. pumilus, thionine synthase MetE was inactivated by S-bacillithio- B. megaterium, and S. carnosus lation causing methionine auxotrophy. Here, we have studied the conserved S-bacillithiolome among four dif- To describe more comprehensively the S-bacillithiolome, ferent Bacillus species and Staphylococcus carnosus using we focused on four industrially important Bacillus species: B. bacillithiol (BSH)-specific immunoblots, shotgun liquid subtilis 168, B. pumilus SBUG1799, B. amyloliquefaciens FZB42 chromatography–tandem mass spectrometry analysis, and (5), B. megaterium SBUG1152 (36), and S. carnosus TM300 (43). redox proteomics. We identified 54 S-bacillithiolated pro- The B. subtilis, B. pumilus, and B. megaterium strains were able teins, including eight conserved proteins (MetE, AroA, to grow in the Belitsky minimal medium (BMM), and low GuaB, TufA, PpaC, SerA, YphP, and YumC) and 29 unique NaOCl concentrations could be applied to reduce the growth proteins involved in amino acid and cofactor biosynthesis, rate: 75 lM NaOCl for B. subtilis (7), 125 lM for B. pumilus, nucleotide metabolism, translation, protein quality con- and 80 lM for B. megaterium (Supplementary Fig. S1B, C; trol, redox, and antioxidant functions. S-bacillithiolation is Supplementary Data are available online at www.liebertpub accompanied by an increased oxidized bacillithiol dis- .com/ars). For the growth of B. amyloliquefaciens FZB42, we ulfide (BSSB) level and a decreased BSH/BSSB ratio. used the BMM with 0.1% casamino acids, and 275–300 lM Together, our data support a major role of the BSH redox NaOCl showed a growth-inhibitory effect (Supplementary buffer in redox control and protection of conserved and Fig. S1A). For S. carnosus, growth was only optimal in a essential proteins against irreversible oxidation by S-ba- complete medium, and 3.25–3.5 mM NaOCl reduced the cillithiolations in Firmicutes bacteria. growth rate in a Luria Broth (LB) medium (Supplementary Fig. S1D). Since amino acids and peptides present in LB also react with NaOCl, much higher NaOCl concentrations were required to inhibit the growth of S. carnosus in the LB medium. Gram-positive bacteria do not produce GSH. Among the actinomycetes, mycothiol is the predominant LMW thiol MetE is the major target for S-bacillithiolation that serves analogous roles to GSH (20, 39). In low-GC in Bacillus species as revealed by nonreducing Gram-positive bacteria, such as Bacillus and Staphylococcus BSH-specific immunoblot analysis species, bacillithiol (BSH) serves as a redox buffer that was structurally characterized as Cys-GlcN-malate (12, 16, 40). To monitor the extent of S-bacillithiolation after NaOCl Mutants with defects in BSH biosynthesis displayed strong stress, BSH-specific antibodies were used to test protein ex- sensitivities to the epoxide antibiotic fosfomycin and to tracts from NaOCl-treated B. subtilis wild-type and bshA- NaOCl stress (7, 12). In Bacillus subtilis, the redox-sensing mutant cells in nonreducing immunoblot analyses. The BSH MarR-type repressor OhrR forms mixed disulfides with BSH immunoblots showed a very strong 90-kDa band after NaOCl (S-bacillithiolations) in response to organic hydroperoxides stress, corresponding to MetE-SSB (Fig. 1A). The identity of and NaOCl stress (7, 26). S-bacillithiolation of OhrR allevi- MetE was confirmed by LC-MS/MS analysis of the specific ates repression of transcription of the ohrA peroxire- band from nonreducing sodium dodecyl sulfate (SDS) gels doxin gene as a protection mechanism against NaOCl (Supplementary Table S2). S-bacillithiolated MetE, modified toxicity (7). Besides OhrR, we have recently identified by at Cys719 and Cys730, was previously identified as the most liquid chromatography (LC)–tandem mass spectrometry strongly oxidized protein in the redox proteome of B. subtilis (MS/MS) analysis five cytoplasmic proteins with S- (7). We further confirmed S-bacillithiolation of MetE by bacillithiolations after NaOCl treatment (7). Hypochloric acid comparison of extracts from NaOCl-treated B. subtilis cells 7 shows very fast reaction rates with Cys residues (k2 = 3 · 10 grown in the BMM and LB media. In a rich LB medium, MetE M - 1 s - 1) and rapidly leads to irreversible oxidation products, is repressed, and thus the MetE-SSB modification is not visible such as sulfinic and sulfonic acids (9, 15, 42, 50). Thus, S- in the BSH immunoblots (Fig. 1B). The MetE-SSB band was bacillithiolation serves to protect active-site Cys residues of also not visible in NaOCl-exposed bshA-mutant cells or in key metabolic enzymes against overoxidation. S-bacillithiola- reducing BSH immunoblots (Supplementary Fig. S2). Besides tion of the methionine synthase MetE at the active site Cys730 MetE, there were a few weaker bands in the 45–55-kDa range, caused a methionine starvation phenotype in NaOCl-treated which represent other protein BSH-mixed disulfides (protein- cells, presumably to slow down translation while the OhrA SSB), since these are absent in the bshA mutant cells. These peroxiredoxin removes the toxic oxidant (7). BSH immunoblot results show that MetE is the most abun- In this work, we have identified proteome-wide conserved dant S-bacillithiolated protein in NaOCl-treated B. subtilis S-bacillithiolations in various industrially important Bacillus cells. and Staphylococcus species. We used BSH-specific immuno- Furthermore, BSH immunoblots indicate that MetE is also blot analyses, redox proteomic approaches, and shotgun-LC- the major BSH-modified protein in diamide-treated cells MS/MS analyses to identify essential and conserved proteins (Fig. 1C). In addition, many other BSH-modified proteins are in the S-bacillithiolome that have been previously identified in visible after diamide stress, indicating that S-bacillithiolation the S-glutathionylome in eukaryotes. The protein bacillithio- is also a significant thiol modification in diamide-treated cells. lations were accompanied by an increased level of oxidized Next, we analyzed the S-bacillithiolome of NaOCl-treated bacillithiol disulfide (BSSB) and a decreased BSH/BSSB redox cells from B. amyloliquefaciens, B. pumilus, B. megaterium, and ratio in B. subtilis. S. carnosus (Fig. 1D). The protein extracts of all NaOCl-treated S-BACILLITHIOLATION AS THIOL PROTECTION MECHANISM IN FIRMICUTES 3

FIG. 1. NaOCl stress causes widespread S-bacillithiolations in four Bacillus species and Staphylococcus carnosus as shown by nonreducing BSH-specific immunoblot analysis. (A, C) IAM-alkylated protein extracts of B. subtilis wild-type and bshA mutant cells harvested before (co) and at different times after NaOCl (A) or diamide stress (C) were subjected to nonreducing SDS-PAGE and BSH-specific immunoblot analysis. (B) IAM-alkylated protein extracts of B. subtilis wild-type cells exposed to 80 lM NaOCl in the BMM and to 4 mM NaOCl in an LB medium were analyzed by BSH-specific immunoblot analysis. MetE is only expressed in BMM and identified as *MetE-SSB. (D) IAM-alkylated protein extracts of B. subtilis (B. sub), B. pumilus (B. pum), B. amyloliquefaciens (B. amy), B. megaterium (B. meg), and S. carnosus (S. carn) harvested before (co) and 10 min after NaOCl stress were analyzed by BSH-specific immunoblot analysis. In all Bacillus strains, the methionine synthase MetE is the most abundant S-bacillithiolated protein as indicated by the asterisks. BMM, Belitsky minimal medium; BSH, bacillithiol; IAM, iodoacetamide; LB, Luria Broth; PAGE, polyacrylamide gel electrophoresis; SDS, sodium dodecyl sulfate.

Bacillus strains showed MetE as the most abundant protein- proteins with disulfides after NaOCl stress, including those SSB, indicating that S-bacillithiolation of MetE is a common with potential SSB sites. The BODIPY-fluorescence image was protection mechanism against oxidative stress among Bacillus overlayed with the Coomassie-stained protein amount species. In B. megaterium, MetE-SSB was observed at a higher image, and the oxidation ratios were calculated versus protein molecular weight and confirmed by LC-MS/MS analysis quantities (Fig. 2A, B and Supplementary Figs. S3–S7 and (Supplementary Table S2). In S. carnosus grown in LB me- Table 1). The reversibly oxidized proteins were identified dium, various protein-SSBs were visible, indicating that using MALDI-TOF/TOF mass spectrometry analysis (Sup- NaOCl also causes S-bacillithiolations in Staphylococcus. plementary Table S1), and the oxidation ratios were used for Because MetE synthesis is strongly repressed in LB medium, hierarchical cluster analyses (Supplementary Figs. S3–S7). MetE-SSB was not detected in S. carnosus using BSH-specific In total, 71 different proteins were identified in all five immunoblots. strains that are constitutively oxidized at either control or NaOCl stress conditions. Several conserved redox-sensitive proteins that coordinate metal ions (Zn, Fe-S-cluster) or form Identification of proteins with S-bacillithiolations disulfide bonds for catalysis were identified in untreated cells, in NaOCl-treated cells of B. subtilis, B. amyloliquefaciens, including the adenylate kinase Adk, the alkyl hydroperoxide B. pumilus, B. megaterium,andS. carnosus using redox reductase small-subunit AhpC, the 2-Cys peroxiredoxins proteomics and shotgun-LC-MS/MS analyses YkuU and YgaF (in B. pumilus), the thiol peroxidase Tpx, the Previously, we have used redox proteomics based on the dihydrolipoamide dehydrogenase E3 subunit of the pyruvate reduction of disulfides and labeling of released thiols with the dehydrogenase complex PdhD, and the sulfite reductase fla- fluorescence dye BODIPY FL C1-IA to identify proteins oxi- voprotein alpha-subunit CysJ (Fig. 2B). The intramolecular dized by NaOCl stress in B. subtilis (7). We applied our redox disulfide bonds were confirmed previously for PdhD, Adk, proteome approach for comparison of B. subtilis, B. amyloli- and CysJ in B. subtilis (7). In B. pumilus, which does not possess quefaciens, B. megaterium, B. pumilus, and S. carnosus to identify the AhpC-AhpF peroxiredoxin/reductase system (14), the 4 CHI ET AL.

FIG. 2. Summary of NaOCl-sensitive proteins (A) and constitutively oxidized proteins (B) in the thiol redox proteomes of four different Bacillus strains and S. carnosus including unique and conserved S-bacillithiolated proteins (E) that were identified by LC-MS/MS analysis (F). (A) Close-ups of conserved NaOCl-sensitive proteins including S-bacillithiolated proteins identified in the redox proteomes of B. subtilis, B. amyloliquefaciens, B. pumilus, B. megaterium, and S. carnosus. Reversibly oxidized proteins in the redox proteome after NaOCl treatment are shown that have been identified with specific and conserved SSB sites (*) using LC-MS/MS analysis. Overlay images represent the redox proteome (red) compared to the Coomassie-stained protein amount image (green) at control conditions (co) and 10 and 30 min after NaOCl exposure (1–3). Column 4 shows the overlay redox proteome control versus redox proteome 30-min NaOCl stress. The identified proteins and their fluorescence/protein quantity ratios are listed in Table 1, and the complete redox proteomes of all strains are given in Supplementary Figures S3–S7. (B) Examples of conserved redox-controlled proteins that are oxidized under control condi- tions to intermolecular or intramolecular disulfides representing thiol-dependent peroxiredoxins (AhpC, YkuU, and YgaF) and thiol peroxidases (Tpx), PdhD, and Adk. Overlay images represent the redox proteome (red) compared to the Coomassie- stained protein amount image (green) of B. subtilis, B. amyloliquefaciens, B. pumilus, B. megaterium, and S. carnosus at control conditions (co) and 10 and 30 min after NaOCl exposure (1–3). Column 4 shows the overlay redox proteome control versus redox proteome 30-min NaOCl stress. The identified proteins and their fluorescence/protein quantity ratios are listed in Table 1, and the complete redox proteomes of all strains are listed in Supplementary Figures S3–S7. (C) Diagonal nonre- ducing/reducing SDS-PAGE confirmed intermolecular disulfides for AhpC homologs in B. subtilis, B. amyloliquefaciens, B.megaterium, S. carnosus and for YkuU in B. pumilus. The IAM-alkylated proteins extracts (100 lg) of NaOCl-treated cells of all strains were separated by 2D nonreducing/reducing diagonal SDS-PAGE analysis as described previously (41). The AhpC homologs are encircled in the diagonal assays of B. subtilis (AhpC-Bsub), B. amyloliquefaciens (AhpC-Bam), S. carnosus (AhpC- Scar), B. megaterium (AhpC-Bmeg), and B. pumilus (YkuU-Bpum). (D) Unique and conserved S-bacillithiolated proteins identified in the redox proteomes of different Bacillus strains and S. carnosus. The number of the identified reversibly oxidized proteins in the redox proteomes of the different strains is shown in brackets. Unique and conserved proteins with identified SSB sites are listed by names that are oxidized in the redox proteome. (E) Unique and conserved S-bacillithiolated proteins identified using LC-MS/MS analysis in different Bacillus strains and S. carnosus. The number of proteins with identified SSB sites after NaOCl stress using LC-MS/MS analysis in the different Bacillus strains and S. carnosus is shown in brackets. Unique and conserved proteins with SSB sites are listed by names. LC, liquid chromatography; MS/MS, tandem mass spectrometry.

YkuU 2-Cys peroxiredoxin is strongly oxidized similar to reducing SDS–polyacrylamide gel electrophoresis (PAGE) AhpC in other Bacillus strains, suggesting that YkuU might analysis (Fig. 2C). Notably, in S. carnosus, the NADH dehy- replace AhpC in peroxide detoxification. The intermolecular drogenase Sca_0546 and the universal stress protein UspA are disulfides of AhpC homologs and YkuU were confirmed for all among the most abundant proteins with strong oxidation Bacillus strains and S. carnosus using diagonal nonreducing/ ratios under control conditions (Supplementary Fig. S7). S-BACILLITHIOLATION AS THIOL PROTECTION MECHANISM IN FIRMICUTES 5

FIG. 2. (Continued).

Exposure of Bacillus strains and S. carnosus to sublethal as the Mn-dependent superoxide dismutase SodA and the NaOCl concentrations caused increased oxidation ratios of Fe-S-cluster assembly ATPase SufC (Supplementary Fig. S6 57 different NaOCl-sensitive proteins (Table 1). The infor- and Fig. 2A). The SufA chaperone was identified with an SSB- mation about the conservations of Cys residues was obtained site at the conserved Cys120 within the Fe-S-cluster site in from the CDD database (www.ncbi.nlm.nih.gov/Structure/ B. amyloliquefaciens (Table 2). The glyceraldehyde-3-phosphate cdd/wrpsb.cgi) (33). Many NaOCl-sensitive proteins harbor dehydrogenase (GapA) was among the most strongly de novo active-site Cys residues that are nucleophilic, catalytically oxidized proteins by NaOCl stress in all redox proteomes. important, bind substrate intermediates or serve as Zn or Fe The redox-active intramolecular disulfide in GapA’s active ligands. These NaOCl-sensitive proteins are associated with a site has been mapped (7, 27). variety of cellular functions, such as ROS scavenging, amino We were interested to identify proteins with NaOCl- acid biosynthesis, fatty acid biosynthesis, purine biosynthesis, sensitive thiols that form SSB sites in Bacillus spp. and S. carnosus. translation, and protein modification/folding factors. Ex- Tryptic digests of protein extracts from control and NaOCl- amples are the methionine sulfoxide reductases (MsrA and treated cells were analyzed for S-bacillithiolations using MsrB) and two ArsC family proteins (SpxA and Sca_0451) Orbitrap-Velos LC-MS/MS analysis. The CID MS/MS spectra that are known to form intramolecular disulfides by oxidative of the BSH-modified peptides show characteristic and abun- stress (7, 38) (Supplementary Fig. S7 and Fig. 2A). In dant malate loss precursor ions (precursor-134 Da) that were B. megaterium, major NaOCl-sensitive proteins were identified used as diagnostic fragment ions to search for BSH-modified Table 1. Identification of Reversibly Oxidized Proteins in the Redox Proteome in Bacillus subtilis, B. amyloliquefaciens, B. pumilus, B. megaterium, and Staphylococcus carnosus

Redox ratios BODIPY fluorescence/protein amount

B. subtilis B. pumilus B. amyloliquefaciens B. megaterium S. carnosus AC-No. Post-translational Proteins co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ Protein function (Uniprot) Function of conserved Cys Cys-modifications

AbrB 2.1 2.4 1.5 Possible transcriptional regulator A8F904 Cys54 essential for DNA- BPUM_0021 binding activity AccB 1.9 3.6 3.2 Biotin carboxyl-carrier protein of P49786 Cys117 conserved acetyl-CoA carboxylase AccD 0.8 2.5 2.8 Acetyl-CoA carboxylase, carboxyl B4AMC5 Cys33,35,52,54 Zn binding transferase, beta-subunit AcnA/CitB 1.2 1.2 1.4 1.3 1.6 1.5 Aconitate hydratase (Aconitase) D5DFZ0/ Cys444, 510, 513 conserved: Homolog Cys404 B9DP54 4Fe-4S-cluster-binding site oxidized by NaOCl in E. coli Adk* 5.4 4.7 4.3 2.3 1.5 1.5 1.8 1.8 1.8 2.2 2.5 4.9 2.6 4.5 2.3 Adenylate kinase F4E1S4 Cys130, 133, 150 conserved, Cys130–133 disulfide; Zn-finger motif Cys150*-S-SB by NaOCl in S. carn AhpC 5.4 2.7 2.9 5.4 3.6 3.8 4.0 3.7 3.8 4.1 4.9 4.8 Alkylhydroperoxide reductase A7ZAJ9 Cys47, 166 conserved; Cys47 Cys47-Cys166 form small-subunit active site catalytic disulfide AldA 0.4 1.3 1.1 Putative aldehyde dehydrogenase B9DKD6 Cys290-conserved active site, nucleophile ArcA 0.7 0.9 0.9 Arginine deiminase 1 B9DKE2 Cys401-conserved amidino- cysteine-intermediate 6 active site ArcC 2.2 1.8 1.4 Carbamate kinase 1 B9DJ94 No Cys conserved ArgG 0.4 2.4 1.9 Argininosuccinate synthase B4AMD7 No Cys conserved Cys187 S-SCys by diamide in Bsub AroA* 0.4 1.1 1.3 0.5 1.2 1.6 0.6 3.4 1.1 0.7 1.0 1.8 Bifunctional 3-deoxy-7- A7Z7R9 Cys126, 259 conserved Cys126*-SSB by NaOCl phosphoheptulonate synthase/ in B. sub., B .pum., chorismate mutase B. amy., S. carn. ArsC (Sca_0451) 1.3 5.4 6.2 Putative ArsC/Spx family protein B9DJH2 Cys10, 13 conserved: form redox-active disulfide bond by oxidative stress BMD_4393 0.9 1.2 0.8 Acetyl-CoA acetyltransferase D5DL95 Active-site Cys90 acetylated by AcetylCoA; Cys381 important for proton exchange CheW 0.3 1.7 2.2 Chemotaxis protein CheW A8FDA9 No Cys conserved CheY 0.6 0.5 0.6 Response regulator CheY A8FD98 No Cys conserved CysH 1.6 1.1 1.6 Phosphoadenosine P94498 Cys120, 121, 203, 206, 229 Cys239 forms mixed phosphosulfate reductase conserved disulfide with Cys32 of Trx in E. coli CysI 3.4 1.7 2.6 Sulfite reductase [NADPH] B4AD94 Cys436, 442, 481, 485 hemoprotein beta-component conserved: 4Fe-4S-cluster- binding site CysJ (BAT_2248) 1.0 1.3 0.9 1.4 1.4 1.6 2.7 2.0 2.0 Sulfite reductase [NADPH] B4AD95 Cys166, 559 conserved Cys428–432 intramole- flavoprotein, alpha-component cular disulfide DapA 0.4 2.3 2.4 Dihydrodipicolinate synthase F4EJJ2 No Cys conserved Dat 2.0 1.2 1.3 d-alanine aminotransferase A7Z2X8 Cys141 close to the active-site Lys144

(continued) Table 1. (Continued) Redox ratios BODIPY fluorescence/protein amount

B. subtilis B. pumilus B. amyloliquefaciens B. megaterium S. carnosus AC-No. Post-translational Proteins co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ Protein function (Uniprot) Function of conserved Cys Cys-modifications

DeoD 4.3 3.7 4.3 Purine nucleoside phosphorylase A7Z5M8 No Cys conserved DeoD-type DhbE 1.7 1.9 1.5 Putative 2,3-dihydroxybenzoate- A7Z8A8 Cys218, 324, 473 conserved AMP ligase DhbE DnaJ 2.6 3.7 3.3 Chaperone protein DnaJ B9DNJ9 Cys147, 150, 204, 207 Zn-finger1; Cys164, 167, 190, 193 Zn-finger2 DnaK* 0.7 0.5 0.6 Chaperone protein DnaK B9DNK0 Cys15 not conserved Cys15*-S-SB by NaOCl in S. carn. Eno 0.7 0.8 0.6 Enolase P37869 No Cys conserved FabH 0.5 1.6 1.5 3-oxoacyl-[acyl-carrier-protein] B9DIT1 Cys112 conserved active site synthase 3 FabZ 4.0 5.3 6.3 (3R)-hydroxymyristoyl-[acyl-car- B9DMG5 Cys94, 134 conserved rier-protein] dehydratase FdhL (Sca_1802) 2.9 3.3 3.0 Putative formate dehydrogenase B9DLU9 Cys37, 48, 51, 63: 2Fe-2S; Cys99, 102, 109: 4Fe-4S; Cys147, 150, 153, 157: 4Fe-4S; Cys190, 193, 196, 200: 4Fe-4S; Cys264, 267,

7 271, 299: 4Fe-4S FolE2 1.0 2.3 2.7 GTP cyclohydrolase B9DKT0 Cys176 conserved, catalytically important site, ligand for Zn2 + FusA 0.5 0.7 0.5 1.0 0.6 0.7 Elongation factor G D5D9Q2 Cys258, 389 conserved GapA 0.8 2.9 3.6 0.5 2.3 1.6 0.4 2.4 2.8 0.4 1.0 1.2 0.7 0.9 0.9 Glyceraldehyde-3-phosphate F4E0S2 Cys152 conserved active site Cys152–156 intramole- dehydrogenase cular disulfide GlmS 0.3 0.7 0.6 Putative glucosamine-fructose-6- B9DMA1 Cys2 catalytic site, forms c- phosphate aminotransferase glutamyl-thioester inter- mediate during Gln hydrolysis GlnA 1.6 2.4 2.2 Glutamine synthetase B9DPA6 Cys96 conserved GltB 5.7 5.6 4.0 Glutamate synthase (NADPH) A8FE17 Cys46, 49, 54, 67, 106, 110, small subunit 114, 307 conserved: Two 4Fe-4S cluster GltX 1.8 1.8 2.6 2.3 1.4 1.7 0.4 0.7 0.8 Glutamyl-tRNA synthetase A7Z0L4 Cys108, 110 conserved, form Zn binding site GlyA 0.7 0.8 0.4 Serine hydroxymethyl transferase A8FIC1 Cys64 conserved Cys66 oxidized by NaOCl in E. coli GlyQS 0.8 0.7 0.8 Glycyl-tRNA synthetase B9DNL1 Cys230, 328, 344, 422 conserved, close to ATP binding sites GrpE* 0.2 2.6 2.8 HSP-70 cofactor protein GrpE B9DNK1 No Cys conserved Cys95*-S-SB by NaOCl in S. carn. GuaA 2.7 1.8 1.3 1.2 0.7 0.9 GMP synthase [glutamine- A8FAH5 Cys85 conserved active site, hydrolyzing] nucleophilic thiolate

(continued) Table 1. (Continued) Redox ratios BODIPY fluorescence/protein amount

B. subtilis B. pumilus B. amyloliquefaciens B. megaterium S. carnosus AC-No. Post-translational Proteins co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ Protein function (Uniprot) Function of conserved Cys Cys-modifications

GuaB* 4.9 3.6 3.8 1.1 2.2 2.1 0.8 2.2 2.5 1.4 2.0 1.7 1.3 3.8 3.6 Inosine-5¢-monophosphate F4EQ59 Cys308 conserved- active Cys308*-S-SB by NaOCl dehydrogenase site, forms thioimidate in B. sub, B. amy., intermediate B. meg. GuaC 3.4 5.2 4.6 GMP reductase B9DP67 Cys174 conserved active site, forms thioimidate intermediate Hit 4.7 3.1 3.5 Cell cycle regulation histidine G0IF34 No Cys conserved triad (Hit) protein Hom 0.6 2.5 3.4 Homoserine dehydrogenase B4AG18 No Cys conserved (BAT_0329) Icd 0.4 1.4 1.3 0.4 0.6 0.6 Isocitrate dehydrogenase [NADP] B4AMB6 Cys118, 411 conserved; Cys118 in active substrate binding site IlvB 1.8 1.6 1.1 Acetolactate synthase, large D5DTM7 No Cys conserved subunit, biosynthetic type IlvC 1.5 1.7 2.0 0.6 0.6 0.6 0.7 1.7 1.6 2.5 2.7 1.7 Ketol-acid reductoisomerase G0IJS5 Cys199, 227 conserved IlvD 2.1 2.5 1.9 Dihydroxy-acid dehydratase D5DER1 Cys122, 189, 192, 195: 4Fe-4S cluster binding IspA 1.9 1.5 1.3 Major intracellular serine protease A7Z3U2 No Cys conserved

8 IspG 5.6 2.1 2.6 4-Hydroxy-3-methylbut-2-en-1-yl F4ESF5 Cys268, 271, 303 conserved: diphosphate synthase 4Fe-4S-cluster-binding site Ldh* 2.7 4.4 4.3 l-lactate dehydrogenase B9DN51 Cys72 conserved Cys72*-S-SB by NaOCl in S. carn. LeuC 1.1 1.3 1.6 4.0 3.4 2.7 2.7 3.4 3.2 3-Isopropylmalate dehydratase B4AM28 Cys346, 406, 409 conserved: large subunit 4Fe-4S cluster binding LuxS* 0.7 2.1 4.5 6.9 2.7 3.2 0.3 2.9 2.7 S-ribosyl homocysteine lyase B4ANL1 Cys84, 126 conserved: Cys84 Cys41*-S-SB by NaOCl (B. pumilus) (LuxS) essential for catalysis; stress in B. pum. Cys126 Fe-binding site LysC 0.6 1.8 1.9 Aspartokinase A8FG00 Cys169 conserved LysC 0.5 1.4 1.3 Aspartokinase B4AM64 No Cys conserved (BAT_1429) MenB 2.2 1.4 1.9 Dihydroxynaphtoic acid B9DQ83 Cys71, 128, 174, 205 synthetase conserved; Cys71 in substrate-binding site MetE* 0.3 1.5 1.4 0.4 1.8 1.6 0.3 1.7 2.0 0.8 1.8 1.2 Methionine synthase A7Z3U1 Cys647, 730 essential-Zinc Cys730*-S-SB; binding active site Cys719*-SSB MetK 0.6 0.6 0.5 0.7 0.6 0.4 S-Adenosylmethionine synthase F4EA25 Cys22, 44 conserved, near active metal (Mg2 + ,K+ )- binding site MsrA 1.7 1.9 2.1 Peptide methionine sulfoxide B9DNX6 Cys10, 13 conserved; Cys 10 Cys10-Cys13 disulfide reductase active site MsrB 2.6 2.2 3.0 Peptide methionine sulfoxide B9DNY9 Cys114-conserved reductase nucleophile active site MtnA 1.7 2.3 2.6 Methylthioribose-1-phosphate O31662 Cys160 conserved, in active isomerase site, transition state stabilizer

(continued) Table 1. (Continued) Redox ratios BODIPY fluorescence/protein amount

B. subtilis B. pumilus B. amyloliquefaciens B. megaterium S. carnosus AC-No. Post-translational Proteins co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ Protein function (Uniprot) Function of conserved Cys Cys-modifications

MtnW 0.4 1.1 1.8 2,3-Diketo-5-methylthiopentyl- A8FCG8 No Cys conserved 1-phosphate enolase MurA 1.4 1.4 1.1 UDP-N-acetylglucosamine 1- D5DM45 Cys117-conserved proton carboxyvinyltransferase 1 donor active site, binds UDP-GlcNAc Ndh (Sca_0546) 0.7 0.6 0.5 4.5 4.2 4.6 2.5 1.4 1.4 NADH dehydrogenase ndh B4AHV4 No Cys conserved NifS (BAT_1509) 3.6 4.7 3.1 Cysteine desulfurase B4ALY7 Cys325-conserved active site OhrA 0.2 6.7 6.7 Organic hydroperoxide resistance B4AEW1 Cys59-,123-conserved Cys Catalytic Cys59–Cys123 (BAT_2845) protein disulfide PdhC 1.2 0.6 0.8 Dihydrolipoyl-lysine-residue B4AEF9 No Cys conserved (BAT_2691) acetyl-transferase component of PDH (E2) PdhD 2.7 1.5 1.3 1.3 0.8 1.0 1.4 1.4 1.2 1.1 0.9 1.1 1.2 1.0 1.2 Dihydrolipoyl dehydrogenase, E3 A7Z482 Cys47-, 52-conserved, Cys47–52 disulfide by component of PDH complex redox-active disulfide NaOCl PdxS 4.2 4.6 4.6 Pyridoxal biosynthesis lyase PdxS B9DKX7 Cys128, 130 conserved PfkA 3.3 3.9 2.8 6-Phosphofructokinase 2 D5DPB6 Cys73, 119 conserved in Bacillus species PheT 2.0 1.6 1.8 Phenylalanyl-tRNA synthetase B9DPU9 Cys65, 79, 120 conserved, in beta-chain tRNA-binding domain

9 PpaC* 2.4 2.7 2.8 1.0 1.5 2.1 3.8 1.7 3.9 2.0 4.2 3.2 Manganese-dependent inorganic A7ZAR2 Cys18, 118, 158 conserved Cys158*-S-SB by NaOCl pyrophosphatase in B. sub. Prs 1.0 0.9 1.0 1.9 1.6 1.6 Ribose-phosphate pyrophospho- A8F918 Cys57, 59, 251, 252 widely kinase conserved PtsI 0.9 0.5 0.8 Phosphoenolpyruvate-protein P23533 Cys364, 503, 556 conserved; phosphotransferase Cys503 proton donor ac- tive site PurB 4.7 2.7 2.2 Adenylosuccinate lyase D5DWW6 Cys31 conserved PurL 1.4 1.5 1.1 Phosphoribosylformylglycinami- A8FAM7 Cys55, 164, 190, 259, 527 dine synthase 2 conserved PurQ 0.7 4.8 4.6 2.0 3.0 3.8 Phosphoribosylformylglycinami- B4AI84 Cys13, 86 conserved, Cys86 dine synthase 1 nucleophilic active site PyrAB 1.4 1.0 0.9 Carbamoyl-phosphate synthase D5DJR1 Cys34, 182, 204, 232, 281, large chain 2 525, 1034 conserved PyrE 0.9 1.3 0.6 Orotate phosphoribosyltransferase B4AE64 No Cys conserved PyrG 2.1 2.3 2.1 CTP synthase A7Z9T4 Cys381 conserved active site PyrH 3.3 2.1 2.3 Putative uridylate kinase B9DPH0 Cys207 conserved QueC (Sca_0360) 3.7 3.8 3.8 Putative ExsB family protein B9DK41 Cys191, 200, 203, 206 con- (7-cyano-7-deazaguanine served Zn binding site synthase) QueF* 0.5 1.2 1.9 1.0 5.0 4.8 1.4 5.4 4.9 NADPH-dependent 7-cyano- B4AEQ3 Cys56 in active site (binds Cys56*-S-SB by NaOCl 7-deazaguanine reductase Zn) in B. sub. Sat 1.2 1.1 1.2 Sulfate adenylyltransferase B4AE60 Cys319, 322, 331 conserved, close to active site that binds APS

(continued) Table 1. (Continued) Redox ratios BODIPY fluorescence/protein amount

B. subtilis B. pumilus B. amyloliquefaciens B. megaterium S. carnosus AC-No. Post-translational Proteins co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ Protein function (Uniprot) Function of conserved Cys Cys-modifications

Sca_1381* 2.9 2.5 2.5 Putative transaldolase B9DN38 Cys200 conserved Cys200*-S-SB by NaOCl in S. carn. Sca_1429 1.2 4.9 5.1 Putative tRNA (cytidine/uridine- B9DMX6 Cys23 conserved 2¢-O-)-methyltransferase Sca_1464 1.9 2.3 2.9 CobB/CobQ-like glutamine ami- B9DMV5 Cys94 conserved dotransferase domain protein nucleophilic active site Sca_2233 2.9 2.2 4.0 Putative short-chain dehydroge- B9DJB3 No Cys conserved nase SecA 0.7 1.2 1.5 Protein subunit P47994 Cys828, 830, 839, 840 conserved Zn-binding site SerA* 1.3 1.5 4.1 1.8 1.1 1.3 0.6 0.6 0.9 Phosphoglycerate dehydrogenase B4AKF2 No Cys conserved Cys410*-S-SB by NaOCl (BMD_2386) in B. sub. SerS 0.6 1.4 2.5 Seryl-tRNA synthetase 2 Q659K7 Cys260, 353 conserved in ATP-binding active site SodA 0.3 2.0 1.9 Superoxide dismutase (Mn) D5DMJ9 Cys not conserved SpxA 1.5 3.4 4.7 Regulatory protein Spx B9DIR2 Cys10, 13 conserved redox-active disulfide Ssb 2.7 3.4 2.5 2.4 1.7 1.5 1.8 2.1 1.8 Single-stranded DNA-binding A8FJE6 No Cys conserved protein 10 SucC 1.2 1.0 0.8 1.3 1.0 1.2 Succinyl-CoA ligase [ADP- F4EIU4 Cys102 in ATP-binding site, forming] subunit-beta Cys193 in Mg/Mn- binding site SufC 0.6 1.3 2.1 FeS assembly ATPase SufC D5DM25 No Cys conserved TenA 1.7 2.1 2.3 0.9 1.0 0.9 Tena/thi-4 family (Thiaminase II) B4AF84 Cys135-conserved catalytic (BAT_2971) active site TenA-2 1.9 1.5 1.4 Thiaminase A8FJ01 Cys135-conserved catalytic BPUM_3574 active site ThiC 2.9 3.4 2.8 Phosphomethylpyrimidine D5DA39 Cys538, 541, 546: 4Fe-4S-S- synthase AdoMet cluster ThiD 2.1 2.1 1.7 0.3 2.3 2.5 Phosphomethylpyrimidine kinase A8FIN1 No Cys conserved ThiG* 0.6 2.5 2.1 Thiazole synthase B4AF80 Cys92 conserved Cys102*-S-SB by NaOCl in B. pum. ThrS 1.1 1.8 1.5 Threonyl-tRNA synthetase B9DNC9 Cys336-conserved, catalytic Zn-binding site TpiA 0.7 0.7 0.6 1.0 0.8 0.8 2.2 2.3 1.8 0.8 0.6 0.6 Triosephosphate isomerase B4AGH8 Cys126 conserved, required for folding and stability Tpx 7.2 4.6 4.1 7.0 3.7 5.8 8.6 4.2 3.3 5.0 6.2 5.6 4.9 5.0 5.8 Probable thiol peroxidase B4AME1 Cys60, 95 conserved active Cys60-Cys95 intra- site molecular disulfide TrxABs /TrxBp 1.5 3.9 3.6 6.9 10.3 7.8 Thioredoxin BAT_3297 not con- B4AP00 Cys62 conserved in served in B. subtilis alkaliphilic Bacilli Tsf 0.5 0.4 0.3 0.6 0.5 0.5 0.4 1.0 1.2 0.8 0.5 0.6 Elongation factor Ts A7Z4S1 Cys22 conserved Tuf* 0.6 0.5 0.5 0.6 1.1 1.2 0.3 0.9 0.8 0.5 0.8 0.7 0.4 0.5 0.5 Elongation factor Tu A7Z0N5 Cys83 conserved, in Cys83*-S-SB by NaOCl GTP-binding site in B. sub, B. meg, B. pum TypA 1.1 0.6 0.6 Putative GTP-binding protein B9DQ05 Cys400 conserved family protein TypA

(continued) Table 1. (Continued) Redox ratios BODIPY fluorescence/protein amount

B. subtilis B. pumilus B. amyloliquefaciens B. megaterium S. carnosus AC-No. Post-translational Proteins co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ co 10¢ 30¢ Protein function (Uniprot) Function of conserved Cys Cys-modifications

UspA (Sca_1315) 1.8 1.3 1.2 Putative universal stress protein B9DNA5 Cys111, 137 conserved in UspA-homologs YceC (TerC) 3.1 3.5 3.8 1.8 2.2 2.7 Tellurium resistance protein A7Z138 No Cys conserved YceD (TerD) 1.8 2.4 3.6 2.0 1.9 1.7 Tellurium resistance protein G0IK82 No Cys conserved YceE (TerE) 1.2 2.0 2.2 Tellurium resistance protein TerE A7Z140 No Cys conserved YdjL (BAT_1651) 1.7 1.4 1.5 Zinc-containing alcohol B4ALJ8 Cys34,37 probably dehydrogenase Zn-binding YerC 9.3 4.2 4.1 Sporulation protein YerC A8FAP4 Cys27, 35 conserved YflT (Sca_0046) 3.2 4.2 2.3 Putative uncharacterized protein B9DM00 No Cys conserved YgaF 6.4 4.7 4.7 Peroxiredoxin A8FB93 Cys45, 50 conserved; Cys45 active site YisY (Sca_0638) 1.5 0.7 1.2 Putative hydrolase of alpha-/ B9DQ91 Cys226 conserved beta-fold family YitJ 1.2 2.4 3.1 Methylenetetrahydrofolate A8FBU4 Cys200, 265, 266 conserved reductase Zn-binding site YjcI (MetI)* 1.3 1.3 1.3 Cystathionine gamma-synthase/ A7Z3I1 Cys338 conserved Cys369*-SSB by NaOCl O-acetylhomoserine (thiol)-lyase in B. pum. YjgF (Sca_0147) 1.2 5.0 4.6 Putative purine regulatory protein B9DLD4 Cys103 conserved, in homo- trimer interaction site YjoA 3.8 2.0 2.6 Uncharacterized protein YjoA A7Z3N1 No Cys conserved 11 YkuU* 6.9 4.7 5.6 Peroxiredoxin YkuU A8FCN5 Cys52, 169 conserved; Cys52 Cys52-Cys169 catalytic active site disulfide bond; Cys169*-SSB YneT 2.2 1.8 1.7 CoA-binding protein A8FDR9 No Cys conserved YodC 4.1 2.7 3.1 Nitroreductase YodC A7Z5L7 Cys153 conserved, close to FMN-binding site YphP* 6.3 2.6 4.2 Thioredoxin-like protein UPF0403 A7Z5T6 Cys53, 55, 144 conserved; Cys53*-SSB by NaOCl family protein Cys53 redox-active site in B. sub. YtxJ (Sca_0389)* 1.1 3.0 5.4 Thioredoxin-like protein DUF2847 B9DJN8 Cys30 conserved (TCPIS Cys30*-SSB by NaOCl family protein motif), Cys30 redox active in S. carn. Cys YvdA (Sca_1457) 1.6 6.1 3.9 Putative carbonic anhydrase B9DMU8 Cys38, 99, 103 conserved catalytic Zn-binding site YxjG* 1.0 1.9 2.2 1.4 2.6 3.0 Methionine synthase homolog A7ZAB5 Cys251, 346 homolog to Cys346*-SSB by NaOCl Zn-binding Cys647, 730 in in B. sub. MetE YybR (HypR) 7.4 6.1 12.0 MarR/DUF24-family regulator; P37486 Cys14 conserved in MarR/ Cys14-Cys49’ intersu- responds to disulfide stress DUF24 family, bunit disulfide by redox-sensitive Cys NaOCl

The proteins were cut from Coomassie-stained 2D-gels, tryptic in-gel-digested, and identified using MALDI-TOF-TOF MS/MS. The table lists the oxidation ratios/protein quantities of all identified reversibly oxidized proteins at control (co) and 10 and 30 min after NaOCl stress as average values of two biological replicate experiments (Exp1 and Exp2), their protein names and functions, Uniprot- accession numbers, and information about conserved Cys residues as revealed by the Conserved Domain Database CDD (33) (http://ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi). The Mascot search results with protein and peptide scores and the oxidation ratios/protein quantities of reversibly oxidized proteins at control (co) and 10 and 30 min after NaOCl stress of two biological replicate experiments (Exp1 and Exp2) with standard deviations are given in Supplementary Table S1. The redox ratios/protein quantities for the B. subtilis proteins were taken from the previous study (7). When a protein is identified in more than one Bacillus species and/or S. carnosus, the accession number is related to one species corresponding to the information about Cys conservation and functions. The asterisk (*) indicates that these proteins have been identified with peptide-SSB sites using LC-MS/MS analysis, and the BSH-modified Cys site is indicated in the last column. Proteins with oxidation/ protein amount ratios that are more than 1.5-fold induced after NaOCl stress are highlighted in gray. BSH, bacillithiol; Cys, cysteine; MS/MS, tandem mass spectrometry. Table 2. Identification of S-Bacillithiolated Proteins in the Proteomes of B. subtilis, B. amyloliquefaciens, B. pumilus, B. megaterium, and S. carnosus Using Liquid Chromatography-MS/MS Analysis

m/z m/z prec-134 Peptide Accession Protein Function Essential Redox-sensing Cys Peptide-SSB or -SSCys sequence precursor (-malate) neutral MW Charge

B. subtilis

O34777 OhrR* Organic hydroperoxide No C15 redox sensing (cons.) (K)LENQLC15( + BSH)FLLYASSR(E) 684.9800 640.6500 2051.9183 3 resistance repressor (control of the OhrA peroxiredoxin) P80877 MetE* 5-methyltetrahydropteroyltri- No C647,730 Zn-binding (R)FWVNPDC730 ( + BSH)GLK(T) 787.8297 720.9900 1573.6449 2 glutamate-homocysteine active site (cons.) (R)VPSTEEMYNIIVDALAVC719 ( + BSH)PTDR(F) 944.7584 900.4600 2831.2533 3 methyltransferase (methio- nine synthase) P42318 YxjG* Methionine synthase homolog No C346 Zn-binding active (R)YVSLDQLC341 ( + IAM)LSPQC346 ( + BSH)GFASTEEGNK(L) 981.4173 936.8900 2941.2301 3 site (cons.) P37487 PpaC* Manganese-dependent Yes C158 cons. (K)SPTC158 ( + BSH)TDQDVAAAK(E) 851.8443 785.1000 1701.6740 2 inorganic pyrophosphatase P35136 SerA* d-3-phosphoglycerate No C410 cons. in Bacillus (K)ISSSESGYDNC410 ( + BSH)ISVK(V) 992.9086 926.0500 1983.8026 2 dehydrogenase (serine species biosynthesis) P39912 AroA Phospho-2-dehydro-3- No C126 cons. (R)FIVGPC126 ( + BSH)AVESYEQVAEVAAAAK(K) 883.4080 839.4800 2647.2022 3 deoxyheptonate aldolase/ chorismate mutase (chorismate biosynthesis) 12 P33166 Tuf Elongation factor Tu Yes C83 in GTP-binding site (R)HYAHVDC83 ( + BSH)PGHADYVK(N) 527.7176 494.4700 2106.8413 4 82–86 (cons.) P21879 GuaB Inosine-5¢-monophosphate Yes C308 active site (cons.) (K)VGIGPGSIC308 ( + BSH)TTR(V) 519.5691 475.0800 1555.6854 3 dehydrogenase (GMP biosynthesis) # P54170 YphP*, UPF0403 protein, No C53 active site (cons.) (K)AEGTTLVVVNSVC53 ( + BSH)GC55 ( + IAM)AAGLAR(P) 815.3758 771.1400 2443.1056 3 thioredoxin-like protein O05268 YumC Ferredoxin—NADP reductase Yes C85 probably active site (K)FDQTIC85( + Cys)LEQAVESVEK(Q) 653.3026 n.d. 1956.8860 3 2 (FAD-dependent pyridine (cons.) nucleotide-disulfide oxidoreductase) P18255 ThrS Threonine—tRNA ligase 1 No C338 Zn-binding; C573 (R)LQC573 ( + BSH)EGLR(V) 607.7535 540.8300 1213.4925 2 not cons. O31678 QueF NADPH-dependent 7-cyano- No C56 active Zn-binding (K)FNC49 ( + IAM)PEFTSLC56( + BSH)PK(T) 613.5820 569.1400 1837.7241 3 7-deazaguanine reductase site (cons.) (tRNA-Queuosine biosynthesis) B. amyloliquefaciens

A7Z3U1 MetE 5-methyltetrahydropteroyltri- No C647,730 Zn-binding (R)FWVNPDC730 ( + BSH)GLK(T) 787.8297 721.0100 1573.6448 2 glutamate-homocysteine active site (cons.) (R)FWVNPDC730 ( + Cys)GLK(T) 433.1948 n.d. 1296.5626 3 methyltransferase (R)VPATEEIYQIIDDALEVC719 ( + BSH)PTDR(F) 962.7668 918.2900 2885.2787 3 (methionine synthase) A7ZAR2 PpaC Manganese-dependent Yes C158 cons. (K)SPTC158 ( + Cys)TEQDIAAAK(E) 727.3190 n.d. 1452.6235 2 inorganic pyrophosphatase

(continued) Table 2. (Continued) m/z m/z prec-134 Peptide Accession Protein Function Essential Redox-sensing Cys Peptide-SSB or -SSCys sequence precursor (-malate) neutral MW Charge

A7Z657 SerA d-3-phosphoglycerate No C410 cons. (K)ISSNESGYDNC410 ( + BSH)ISVK(V) 671.2758 626.8900 2010.8054 3 dehydrogenase (serine biosynthesis) A7Z7R9 AroA Phospho-2-dehydro-3-deoxy- No C126 cons. (R)FIVGPC126 ( + BSH)AVESYEQVAEVAAAAK(K) 883.4087 n.d. 2647.2042 3 heptonate aldolase/choris- matemutase (chorismate biosynthesis) A7Z612 AroE 3-phosphoshikimate No C79 not cons. (K)GIDALC79( + BSH)EPDSLLDVGNSGTTIR(L) 881.4009 836.7600 2641.1808 3 1-carboxyvinyltransferase (K)GIDALC79( + Cys)EPDSLLDVGNSGTTIR(L) 789.0375 n.d. 2364.0906 3 (chorismate biosynthesis) A7Z0D1 GuaB Inosine-5¢-monophosphate Yes C308 active site (cons.) (K)VGIGPGSIC308 ( + BSH)TTR(V) 778.8491 712.0500 1555.6837 2 dehydrogenase (GMP forms thioimidate biosynthesis) intermediate A7Z8C1 YumC Ferredoxin—NADP reductase Yes C85 probably active site (K)FDQTIC85 ( + Cys)LEQAVESVEK(Q) 653.3023 n.d. 1956.8851 3 2(FAD-dependentPyridine (cons.) (K)FDQTIC85 ( + BSH)LEQAVESVEK(Q) 745.6595 701.2400 2233.9568 3 nucleotide-disulfide oxidoreductase) A7Z7N3 ThiI Probable tRNA sulfur No C81 not cons. (K)C81( + BSH)ESKLEDIK(K) 730.8164 n.d. 1459.6183 2 transferase (YtbJ homolog) (Thiamine biosynthesis) A7Z4X4 CotE Spore coat protein E No C113 cons. (K)VLQQPNC113 ( + Cys)LEVTISPNGNK(I) 691.6771 n.d. 2072.0094 3 F4EAC4 SufA Chaperone involved in Fe-S No C102,104,120 cons: Fe-S- (K)NAGTPEEC120 ( + BSH)(-) 608.7031 541.5800 1215.3916 2 13 cluster assembly cluster A7Z1M1 Alr Alanine racemase 1 (cell wall Yes C63 not cons. (K)AALEAGASC63( + BSH)LAVAILDEAISLR(K) 851.7523 807.5500 2552.2351 3 biosynthesis) B. pumilus

B4AEV7 MetE 5-methyltetrahydropteroyltri- No C647,730 Zn-binding (R)FWVNPDC730 ( + BSH)GLK(T) 787.8287 721.0000 1573.6428 2 glutamate-homocysteine active site (cons.) methyltransferase (methionine synthase) A8FIT1 YwaA Branched-chain amino-acid No C104 S-SCys by diamide (R)LC104 ( + BSH)IPQIDTETVLEGLNELIR(I) 889.1032 844.8300 2664.2878 3 aminotransferase (valine, in B. sub. leucine, isoleucine biosynthesis) A8FGB2 AroA Phospho-2-dehydro-3-deoxy- No C126 cons. (R)FIVGPC126 ( + BSH)AVESYEQVAEVAAAAK(Q) 883.4035 n.d. 2647.1887 3 heptonate aldolase/ chorismatemutase (chorismate biosynthesis) B4AN33 Tuf Elongation factor Tu Yes C83 in GTP-binding site (R)HYAHVDC83( + BSH)PGHADYVK(N) 527.7175 494.4800 2106.8408 4 82–86 (cons.) B4AF61 MetI Cystathionine-gamma No C335 cons. (R)IANGVC369 ( + BSH)NK(L) 607.7542 540.7300 1213.4939 2 synthase (CGS) (O-succinylhomoserine (thiol)-lyase) (homocysteine biosynthesis) B4ANL1 LuxS S-ribosylhomocysteine lyase No C84 essential for catalysis (R)FC41( + BSH)QPNK(Q) 566.7175 499.7200 1131.4204 2 (homocysteine biosynthesis) (cons.) C126 Fe- binding site (cons.)

(continued) Table 2. (Continued) m/z m/z prec-134 Peptide Accession Protein Function Essential Redox-sensing Cys Peptide-SSB or -SSCys sequence precursor (-malate) neutral MW Charge

B4AF80 ThiG Thiazole synthase (Thiamine No C92 cons. (K)VEVIGC102 ( + BSH)SR(S) 629.7681 562.9200 1257.5217 2 biosynthesis) B4AFT4 KatX2 Catalase ? No Cys cons. (K)LLAIC461 ( + BSH)NFYR(A) 503.5634 459.1000 1507.6684 3 A8FCN5 YkuU 2-Cys peroxiredoxin ? C52, 169 cons.; C52 active (R)VLQALQTGGLC169 ( + BSH)PANWKPGQK(T) 835.7413 791.4100 2504.2022 3 site; C52–C169 catalytic disulfide A8FHX4 FliW Flagellar assembly factor No C144 not cons. (K)HLLEVASSC144 ( + BSH) 677.7782 610.9300 1353.5418 2 A8FG49 CitZ Citrate (Si)-synthase II No C195 cons. (R)VC195 ( + BSH)VATLSDIYSGVTAAIGALK(G) 816.7327 772.5300 2447.1762 3 (TCA cycle) B. megaterium

D5DIR6 MetE 5-methyltetrahydropteroyltri- No C649,732 Zn-binding (R)ALQVLDPALFWINPDC732 ( + BSH)GLK(T) 837.0709 792.9500 2508.1908 3 glutamate-homocysteine active site methyltransferase (methionine synthase) D5DER3 PpaC Manganese-dependent Yes C158 cons. (K)SPTC158 ( + BSH)TDQDVAAAK(E) 568.2314 523.7300 1701.6723 3 inorganic pyrophosphatase (K)SPTC158 ( + Cys)TDQDVAAAK(E) 713.3030 n.d. 1424.5915 2

D5D924 GuaB Inosine-5¢-monophosphate Yes C308 active site, forms (K)VGIGPGSIC308 ( + BSH)TTR(V) 778.8499 712.1000 1555.6853 2 dehydrogenase (GMP thioimidate 14 biosynthesis) intermediate (cons.) S. carnosus

B9DN54 AroA Phospho-2-dehydro-3- No C83, 125 cons; C83 (K)SFIFGPC125 ( + BSH)SVESQEQVDK(V) 765.9924 721.8800 2294.9553 3 deoxyheptonate aldolase/ catalytic site; C125 chorismate mutase Fe-binding site (chorismate biosynthesis) B9DKV8 Tuf Elongation factor Tu No C82 in GTP-binding site (R)HYAHVDC82( + BSH)PGHADYVK(N) 527.7177 494.5300 2106.8417 4 81–85 B9DM03 GuaB Inosine-5¢-monophosphate No C307 active site, forms (K)VGIGPGSIC307 ( + BSH)TTR(I) 519.5693 475.0700 1555.6862 3 dehydrogenase, (GMP thioimidate biosynthesis) intermediate (cons.) B9DNY0 YphP UPF0403 protein No C54 active site (K)NVGKDETTFVVINSTC54( + BSH)GC56( + IAM)AAGLAR(P) 720.5766 687.4300 2878.2773 4 (thioredoxin-like protein) (CxC-motiv) cons. B9DJN8 YtxJ DUF2847 protein No C30 active site (K)HSNTC30 ( + BSH)PISANAYDQFNK(F) 769.3155 724.9500 2304.9246 3 (thioredoxin-like protein) (TCPI-motiv) cons. B9DNK0 DnaK Chaperone protein No No Cys cons. (K)IIGIDLGTTNSC15( + BSH)VAVLEGDEPK(V) 880.7465 836.5500 2639.2176 3 B9DNK1 GrpE Chaperone protein No No Cys cons. (K)TYQAQC95( + BSH)VLTDILPTIDNIER(A) 901.4220 856.9600 2701.2442 3 B9DN51 Ldh l-lactate dehydrogenase No C72 cons. (K)AGSYEDC72( + BSH)SDADLVVITAGAPQKPGETR(L) 787.3511 754.4000 3145.3684 4 Q9RGS6 ThiM Hydroxyethylthiazole kinase No C258 not cons. (R)IDSDAVAENC258 ( + BSH)NLEEVK(-) 715.6331 671.2800 2143.8775 3 (thiamine biosynthesis) B9DMD5 Sca_1625 Putative aldehyde dehydroge- ?C279activesite(cons.)(K)VYNNTGQVC279 ( + BSH)TAGTR(T) 627.2645 582.8600 1878.7716 3 nase family protein (similar (K)VYNNTGQVC279 ( + Cys)TAGTR(T) 534.9047 n.d. 1601.6922 3 to GbsA betaine aldehyde dehydrogenase) (continued) Table 2. (Continued) m/z m/z prec-134 Peptide Accession Protein Function Essential Redox-sensing Cys Peptide-SSB or -SSCys sequence precursor (-malate) neutral MW Charge

B9DN59 YtpR Similar to Phe-tRNA No C126,167 cons. (K)VNVGNEELQIVC126 ( + BSH)GAPNVEAGQK(V) 888.7393 844.1200 2663.1961 3 synthetase (YtpR homolog) (K)VNVGNEELQIVC126 ( + Cys)GAPNVEAGQK(V) 796.3812 n.d. 2386.1218 3 B9DNG1 Dtd d-tyrosyl-tRNA(Tyr) No No Cys cons. (R)LYEAFNEALC113 ( + Cys)QYGVEVK(T) 698.9884 n.d. 2093.9434 3 deacylase (YrvI homolog) B9DLW2 SceB SceB precursor (SsaA ?C166cons (R)TSSGANYYTAGQC166 ( + BSH)TYYAFDR(A) 879.0096 834.6200 2634.0069 3 homolog) (R)TSSGANYYTAGQC166 ( + Cys)TYYAFDR(A) 786.6523 n.d. 2356.9352 3

B9DN38 Sca_1381 Putative transaldolase ? C200 cons. (R)ELLNVIQADEIGADIITC200 ( + BSH)PSGVISK(I) 998.8192 954.2100 2993.4358 3 B9DQ05 TypA Putative GTP-binding protein No C408 not cons. (R)VQC408 ( + BSH)EVPQENAGAVIESLGQR(K) 841.7123 n.d. 2522.1150 3 family protein (YlaG homolog)

15 B9DML0 Sca_1554 Putative ABC transporter ? No Cys cons. (R)QIENC583 ( + IAM)EAEIEAC590 ( + BSH)EQK(I) 730.6216 686.2800 2188.8431 3 ATP-binding protein B9DM56 Adk Adenylate kinase Yes C130, 133, 150 cons., (K)VDGVC150 ( + BSH)DLDGGK(L) 491.8620 447.3800 1472.5642 3 Zn-finger motif

Cytoplasmic protein extracts of B. subtilis, B. amyloliquefaciens, B. pumilus, B. megaterium, and S. carnosus were prepared in an urea-IAM-buffer, tryptic in-gel-digested, and analyzed in a LTQ Orbitrap- VelosÔ mass spectrometer as described in the Materials and Methods section. Peptides with S-bacillithiolations were identified by the additional mass of 396 Da at Cys residues and the diagnostic neutral malate loss fragment ions that appeared as abundant ions in the fragment ion MS/MS spectra. Peptides with S-cysteinylation were identified with a mass difference of 119 Da at Cys residues. The table includes the Uniprot accession numbers and protein functions as derived from the UniprotKB database (http://uniprot.org/) and information about conserved Cys residues and Cys functions as revealed by the Conserved Domain Database CDD (34) (http://ncbi.nlm.nih.gov/Structure/cdd/wrpsb.cgi). The table also includes the m/z of the precursor ions, m/z of the precursor malate loss ions ( - 134 Da), and the neutral molecular mass of the BSH-modified peptide and peptide charges. The Xcorr, DCn scores, and mass deviations of the S-bacillithiolated peptides and complete collision- induced diffraction (CID) MS/MS spectrum of all peptides and the b and y fragment ion series are given in Supplementary Figures S8 and S9 as follows: OhrR (S8A), MetE (S8B), YxjG (S8C), PpaC (S8D), SerA (S8E), AroA (S8H), Tuf (S8I), GuaB (S8J), YphP (S8F), YtxJ (S8G), YumC (S8K), ThrS (S8L), QueF (S8M), YwaA (S8N), MetI (S8O), LuxS (S8P), ThiG (S8U), KatX2 (S8S), YkuU (S8T), FliW (S8Q), CitZ (S8R), AroE (S8V), CotE (S8W), SufA (S8X), Alr (S8Y), ThiI (S8Z), DnaK (S9A), GrpE (S9B), Ldh (S9C), ThiM (S9D), Sca_1625 (S9E), YtpR (S9F), Dtd (S9G), SceB (S9H), Sca_1381 (S9I), TypA (S9J), Sca_1554 (S9K), and Adk (S9L). * indicates that these proteins with SSB sites have been identified in the previous study in B. subtilis (7) and are listed here for comparison. Bold texts indicate conserved S- bacillithiolated proteins that are identified in more than two of the four Bacillus species and S. carnosus in this study. #The YphP protein was identified in untreated and NaOCl-treated B. subtilis cells. ? indicates that it is unknown if this protein is essential. n.d. indicates that the malate-loss precursor ion was not determined. IAM, iodoacetamide; LTQ, linear-trap quadrupole; MW, molecular weight. 16 CHI ET AL. peptides and labeled in the MS/MS spectra (Table 2, Sup- EF-Tu, aminoacyl-tRNA synthetases (ThrS, YtpR, and TypA), plementary Tables S2 and Supplementary Figs. S8 and S9) (7). and the DnaK and GrpE chaperones (Fig. 2A, Tables 1 and 2). In our previous study, we found S-bacillithiolation sites for Oxidation of DnaK and GrpE was only observed in the S. car- the thioredoxin-like protein YphP in B. subtilis under control nosus redox proteome, and the SSB sites were mapped at Cys15 conditions (7), but no further SSB sites were identified in and Cys95, respectively (Fig. 2A and Supplementary Figs. S3– Bacillus species and S. carnosus. In response to NaOCl stress, S7 and Table 2). we could identify in total 54 proteins with characteristic SSB Among the redox and antioxidant proteins, the thio- sites, including 29 unique proteins and eight conserved pro- redoxin-like proteins YphP and YtxJ (Sca_0389) were identi- teins with identical SSB sites in two or more Bacillus strains fied with SSB sites at their active-site Cys53 and Cys30, and S. carnosus. The eight proteins with conserved SSB sites respectively, and oxidized in the redox proteome of S. carno- include the methionine synthase (MetE), the inorganic pyr- sus by NaOCl stress (Fig. 2A and Supplementary Figs. S8 and ophosphatase (PpaC), the 3-D-phosphoglycerate dehydroge- S9). The ferredoxin–NADP + oxidoreductase YumC (23) was nase (SerA), the bifunctional 3-deoxy-7-phosphoheptulonate identified with an SSB modification at the active-site Cys85. synthase/chorismate mutase (AroA), the translation elonga- In S. carnosus, the lactate dehydrogenase Ldh and the pu- tion factor EF-Tu (TufA), the inosine 5¢-monophosphate (IMP) tative transaldolase Sca_1381 are constitutively oxidized in dehydrogenase (GuaB), the thioredoxin-like protein (YphP), the redox proteome, and the Cys72-SSB and Cys200-SSB sites, and the ferredoxin–NADP + oxidoreductase (YumC) (Fig. 2A, respectively, were identified by LC-MS/MS (Fig. 2A; Table 2). E and Supplementary Figs. S8 and S9). The aldehyde dehydrogenase AldA and the arginine deimi- The methionine synthase MetE was oxidized most strongly nase ArcA are NaOCl-sensitive proteins in the redox pro- and specifically by NaOCl stress in the redox proteome of all teome of S. carnosus that also possess active-site Cys residues. Bacillus strains (Fig. 2A and Supplementary Figs. S3–S7), and the conserved Cys730-SSB and Cys719-SSB sites were map- NaOCl stress caused a decreased BSH/BSSB redox ped by LC-MS/MS analyses (Supplementary Fig. S8). The ratio in B. subtilis as shown by thiol metabolomics YxjG methionine synthase paralog was only oxidized by In B. subtilis, grown in an LB medium, similar amounts of NaOCl stress in B. subtilis and B. amyloliquefaciens (Supple- BSH (0.6–1 lmol/g dry weight [DW]) and Cys (0.6 lmol/g mentary Figs. S3 and S4). Further, NaOCl-sensitive enzymes DW) have been quantified previously (12, 40). The redox ra- with SSB sites at nonconserved Cys369 and Cys41, respec- tios were calculated previously as *400:1 for BSH/BSSB and tively, include the cystathionine-gamma synthase (MetI) and *120:1 for Cys/Cystine in LB-grown B. subtilis cells (40). We S-ribosylhomocysteine lyase (LuxS) of B. pumilus involved in quantified the change of the BSH/BSSB and Cys/Cystine homocysteine biosynthesis (Supplementary Figs. S8 and S9). redox ratios in B. subtilis cells grown in the BMM after NaOCl The conserved proteins SerA with Cys410-SSB and PpaC with stress (Fig. 4 and Supplementary Fig. S10). The BSH level was Cys158-SSB modifications (7) are constitutively oxidized in 2.2 lmol/g DW in growing cells (OD = 0.4) and increased to the redox proteomes of all strains, suggesting that basal S- 500 2.7 lmol/g DW directly after NaOCl stress and to 3.2 lmol/g thiolation is observed for selected proteins during aerobic DW during postexponential growth (Fig. 4A). BSH synthesis growth (Fig. 2A and Supplementary Figs. S3–S7). The AroA is about two-fold increased by NaOCl stress compared to protein was strongly oxidized in all redox proteomes by control cells. The BSSB level was 0.06 lmol/g DW in growing NaOCl, and the conserved Cys126-SSB site was identi- cells, and NaOCl exposure resulted in a rapid two-fold in- fied by LC-MS/MS analysis (Fig. 2A and Supplementary crease to 0.11 lmol/g DW after 15 min (Fig. 4B). The BSSB Figs. S3–S7 and Table 2). AroA functions in the first step of level decreased within 60 min after NaOCl stress to control chorismate synthesis catalyzing the production of 3-deoxy- levels. The BSH/BSSB redox ratio was calculated as 40–50:1 d-arabino-heptulosonate 7-phosphate (DAHP), and thus its S- during exponential growth, dropped two-fold shortly after bacillithiolation could regulate the biosynthesis of aromatic NaOCl exposure, and recovered to 50–60:1 within 2 h (Fig. 4C, amino acids (Fig. 3). In B. amyloliquefaciens, the AroE protein D). Approximately 3% (0.06 lmol/g DW) of the total cellular that functions in the later steps of chorismate biosynthesis was BSH content was bound as protein-SSB during growth and also identified with a Cys79-SSB site. The essential GuaB pro- increased two-fold after exposure to 120 lM NaOCl (Fig. 4E). tein that functions in guanosine 5¢-phosphate biosynthesis The BSH level is four- to eight-fold higher than Cys, and the showed increased oxidation ratios by NaOCl stress in most levels of Cys (0.4 lmol/g DW), Cystine (0.05 lmol/g DW), Bacillus spp., and the active-site Cys308 was identified as a and the Cys/Cystine redox ratio (*10:1) were not affected by conserved SSB site (Figs. 2A and 3 and Supplementary Figs. S3– NaOCl stress. These data confirm that BSH is used as major S7 and Table 2). Enzymes for thiamine biosynthesis also harbor redox buffer and oxidized to BSSB by NaOCl stress, causing a NaOCl-sensitive thiols, such as the thiazole synthase ThiG in B. decreased BSH/BSSB redox ratio and S-bacillithiolation in pumilus that was identified with a Cys102-SSB site. The hy- B. subtilis cells. droxyethylthiazole kinase ThiM of S. carnosus was identified with a Cys258-SSB modification (Figs. 2A and 3 and Supple- Discussion mentary Figs. S3–S7 and Table 2). The NADPH-dependent 7- cyano-7-deazaguanine reductase QueF was identified with an Previously, we have shown that BSH forms mixed protein SSB site at the active site Zn-binding Cys55, and was NaOCl- disulfides (S-bacillithiolations) with essential catalytic Cys oxidized in the redox proteomes of B. amyloliquefaciens and B. residues of enzymes involved in the methionine biosynthesis pumilus. QueF is involved in the biosynthesis of queuosine, a pathway and with the redox-sensing OhrR repressor after hypermodified base found in the wobble positions of tRNAs NaOCl stress in B. subtilis (7). Thus, S-bacillithiolation acts as a (25). Additional proteins with increased oxidation ratios and redox-switch under hypochlorite stress to induce methionine identified SSB sites include the translation elongation factor starvation and up-regulation of the OhrA peroxiredoxin for S-BACILLITHIOLATION AS THIOL PROTECTION MECHANISM IN FIRMICUTES 17

FIG. 3. Metabolic enzymes with SSB sites were identified within the biosynthetic pathways for methionine, Cys, chorismate, thiamine, GMP, and queuosine. The translation and protein quality control machinery is another target of S- bacillithiolation. The schematics of the metabolic pathways refer to the annotated genes of B. subtilis modified from the SubtiPathways database (http://subtiwiki.uni-goettingen.de/subtipathways.html), and possible inhibitory steps in these pathways are shown by the identified S-bacillithiolated proteins SerA, PpaC, MetE, YxjG, MetI, LuxS (Met, Cys biosynthesis); AroA, AroE (Chorismate biosynthesis); ThiG, ThiM (Thiamine biosynthesis); GuaB and QueF (GMP, queuosine biosynthesis). The schematic for queuosine biosynthesis is adapted from (37), and the Met and Cys biosynthesis pathway is from (48). Cys, cysteine; GMP, guanosine 5¢-phosphate. (To see this illustration in color, the reader is referred to the web version of this article at www.liebertpub.com/ars.) detoxification. In this work, S-bacillithiolation sites in 54 alkylation and affinity purification (e.g., the biotin-switch as- proteins have been identified among four different industri- say) (21, 28, 30, 31). By analogy, large-scale identification and ally important Bacillus species and S. carnosus after exposure quantification of BSH-mixed protein disulfides would require to NaOCl. We applied our previous LC-MS/MS analysis to selective reduction by bacilliredoxins (Brxs), followed by al- directly identify S-bacillithiolation sites and proved the MS/ kylation and purification protocols. We identified possible MS spectra by their characteristic malate loss precursor ions. candidates for putative Brxs (YphP and YtxJ) that were S-ba- The number of S-bacillithiolated proteins is lower than that of cillithiolated at their active-site Cys residues. Current attempts glutathionylated proteins identified in eukaryotes. This dis- are directed to develop Brx-based redox proteomics methods crepancy could be due to the labile nature of the Cys-GlcN-Mal to further characterize and quantify the S-bacillithiolome, and moiety that limits its detection, and also because the amount of to analyze the substrate specificities of these putative Brxs. BSH is much lower (2–3 lmol/g DW) compared to GSH in E. The fluorescence-based redox proteomics approach was coli (19 lmol/g DW) or mycothiol (MSH) in Mycobacterium used to quantify all oxidized proteins, and many conserved smegmatis (40 lmol/g DW) (40). Thus, selective thiol-trapping S-bacillithiolated proteins show increased oxidation ratios and enrichment tools need to be developed to quantify upon NaOCl exposure (e.g., MetE, YxjG, AroA, and ThiG). S-bacillithiolated proteins more comprehensively. To identify However, some conserved S-bacillithiolated proteins like S-glutathionylated proteins in the proteome of eukaryotic or- TufA, GuaB, PpaC, and SerA did not show increased oxida- ganisms, redox proteomic approaches have been developed tion ratios by NaOCl stress in the redox proteomes of most that use glutaredoxin (Grx) for specific de-glutathionylation Bacillus spp., indicating that S-bacillithiolation also occurs of cell extracts, followed by biotin-N-ethylmaleimide (NEM) under nonstress aerobic growth conditions. This confirms 18 CHI ET AL.

FIG. 4. The BSH/BSSB redox ratio and BSH protein-mixed disulfides are changed in B. subtilis after NaOCl exposure. B. subtilis cells grown in BMM were exposed to 120 lM NaOCl stress at OD600 = 0.4 and harvested immediately before (0 min) and 15, 60, and 120 min post-NaOCl stress. The reduced thiol redox buffer (RSH) including BSH and Cys (A), the oxidized thiol redox buffer (RSSR) including BSSB and Cystine (B), and protein BSH contents (E) were measured as described in the Materials and Methods section. The RSH/RSSR ratios are calculated for control cells (C) and NaOCl-treated cells (D) in relation to the growth curve. The time point of NaOCl exposure is indicated by an arrow. The combined growth curves of untreated and NaOCl-treated cells are also shown for comparison (F). Experiments were performed in triplicate, and the error bars are given as standard error of the means values. Representing peaks for BSH, Cys, BSSB, Cystine and protein BSH are shown in the high performance liquid chromatography chromatograms in Supplementary Figure S10. BSSB, oxidized bacillithiol disulfide. results of S-glutathionylations in malaria parasites, human, bined redox proteomics and LC-MS/MS analyses showed a and yeast cells, where the basal level S-thiolation occurs even similar composition of the S-bacillithiolome compared to the under nonstress conditions as a result of aerobic growth (21, S-glutathionylomes described for eukaryotes in oxidatively 24, 30). In Plasmodium falciparum, more than 400 S-glutathio- stressed cells (8, 31). The S-bacillithiolome contains mainly nylated proteins were identified in nonstressed cells (21). The biosynthetic enzymes for amino acids, cofactors, nucleotides, authors further demonstrated in vitro that the activities of as well as translation factors, chaperones, and redox and an- only a few identified metabolic enzymes were inhibited by tioxidant proteins. Most proteins are enzymes involved in S-glutathionylation. The physiological role and verification of amino acid biosynthesis pathways, such as methionine, Cys the predicted basal level S-bacillithiolation of essential pro- (MetE, YxjG, MetI, LuxS, and PpaC), serine (SerA), aromatic teins in our study require further detailed investigations, since amino acids (AroA and AroE), branched chain amino acids we could not identify S-bacillithiolations of these proteins in (YwaA), GTP synthesis (GuaB), queousine biosynthesis untreated cells using LC-MS/MS analysis. Overall, our com- (QueF), and thiamine biosynthesis (ThiG, ThiI, and ThiM). We S-BACILLITHIOLATION AS THIOL PROTECTION MECHANISM IN FIRMICUTES 19 further identified S-bacillithiolated proteins involved in Among the conserved S-bacillithiolated proteins, the translation, such as elongation factor EF-Tu and aminoacyl- chorismate synthase AroA was identified, which catalyzes the tRNA-synthetases; the heat-specific chaperones (DnaK and synthesis of DAHP. Notably, the DAHP synthase Aro4p of GrpE); and redox and antioxidant proteins, including yeast cells was found as a target for the Grx2 and showed thioredoxin-like proteins (YphP and YtxJ), the ferredoxin- increased thiol redox ratios at Cys76 and Cys244 in the grx2 NADP + oxidoreductase (YumC), and peroxiredoxins (YkuU), mutant (35). The essential IMP dehydrogenase GuaB and the which could function in the de-bacillithiolation process or pyrophosphatase PpaC have been previously found to be S- require BSH for their regeneration. glutathionylated in human T-lymphocytes and endothelial The identified S-bacillithiolated proteins include eight cells under diamide stress conditions (11, 30). conserved proteins identified in B. subtilis (MetE, GuaB, PpaC, An important finding of this study is the identification of S- SerA, AroA, Tuf, YphP, and YumC), which could be con- bacillithiolated proteins involved in translation and protein firmed as S-bacillithiolated proteins in other species (Fig. 2D, quality control, such as elongation factor EF-Tu and the DnaK E). Most conserved S-bacillithiolated proteins are abundant and GrpE chaperones. The heat-specific Hsp70 and Hsp90 proteins oxidized in the redox proteomes of Bacillus and Sta- chaperones are well represented in the S-glutathionylomes of phylococcus species (e.g., MetE, Tuf, GuaB, PpaC, SerA, and human and yeast cells (11, 45). The elongation factor TufA AroA) and have conserved SSB peptides (Fig. 2A and Table 2). promotes GTP-dependent binding of aminoacyl-tRNAs to the In addition, we detected 29 unique proteins that were S- A-sites of the ribosome, but also has chaperone functions bacillithiolated only in one Bacillus species or S. carnosus. The (4, 47, 49). The TufA protein was oxidized at Cys256 by most unique proteins with SSB sites were identified in S. NaOCl stress in E. coli, and a tufA mutant was highly sensitive carnosus (12 SSB sites) and B. pumilus (8 SSB sites). These to NaOCl (27). In the Bacillus spp. and S. carnosus TufA pro- proteins are less abundant in the gel-based proteome and teins, we identified the conserved Cys82-SSB modification in were only detected by LC-MS/MS analysis. For example, the GTP-binding site. Since TufA is essential in B. subtilis, its ThiM and ThiG proteins were found as specific targets for S- S-thiolation in oxidatively stressed cells could cause inhibition bacillithiolation in S. carnosus and B. pumilus, respectively. of protein synthesis during oxidative stress as previously Novel S-bacillithiolated proteins were mapped in S. carnosus shown in yeast cells (45). Thus, the fact that amino acid bio- as the putative aldehyde dehydrogenase (Sca_1625), the synthetic enzymes and translation proteins dominate in the transaldolase (Sca_1381), and the lactate dehydrogenases conserved S-bacillithiolome suggests that S-bacillithiolation (Ldh), as well as the SceB precursor. These proteins are not could control protein synthesis during oxidative stress. The conserved in Bacillus species, but homologs are present in the strong repression of the stringent controlled RelA regulon, pathogen S. aureus. Interestingly, the chaperones GrpE and including genes for ribosomal proteins, by NaOCl stress DnaK were S-bacillithiolated in S. carnosus, but Cys residues in our previous microarray analysis further indicates a are absent in the GrpE and DnaK homologs of Bacillus species. decreased protein synthesis that could be caused by S-ba- The detection of some unique SSB-peptides could be also due cillithiolation (7). The activation of the stringent response by to better peptide ionization properties of the tryptic peptides ROS and RNS has also been shown in Salmonella (2, 17). of less-conserved proteins. Specifically, the stringent response regulatory DnaK sup- Using BSH-specific immunoblots, we could demonstrate pressor protein (DksA) was shown to be play a role in con- that MetE is the most abundantly S-bacillithiolated protein trolling GSH biosynthesis and other metabolic pathways in all Bacillus species. MetE is most strongly oxidized by associated with the generation of reducing power under NaOCl stress in the redox proteome of B. subtilis wild-type oxidative stress conditions (17). and bshA mutant cells and is S-cysteinylated in the bshA The next question is how the reversibility of this redox mutant (7). Similar MetE oxidation ratios were calculated mechanism (protein de-bacillithiolation) might be mediated. after 30 and 60 min post NaOCl stress in the wild-type and Redoxins catalyzing the reversible S-thiolation/reduction of bshA mutant cells, but reduction of MetE took significant S-glutathionylated proteins include Grxs, thioredoxins, glu- longer in the bshA mutant (Supplementary Fig. S11). Simi- tathione S-transferases, and GSH disulfide reductases that larly, AroA was S-bacillithiolated in the wild-type cells and were identified in large-scale redox proteomics studies (21). displays similar oxidation ratios in the wild-type and bshA The mechanisms of protein de-glutathionylation by mono- mutant cells within the first hour, but is more slowly re- and dithiol Grx have been studied in plants and mammalian duced in the bshA mutant. This may suggest that systems (13, 52). In both mechanisms, the nucleophilic active- S-cysteinylation can fully compensate for S-bacillithiolation site Cys first attacks the GSH-mixed protein disulfide, re- in the absence of BSH, but the reduction of these mixed sulting in substrate de-glutathionylation and formation of a disulfides occurs with different efficiencies. Grx-SSG intermediate (52). In plant dithiol Grx, this Grx-SSB The physiological importance of MetE inhibition by intermediate is then attacked by a second reactive Cys not S-thiolation has previously been demonstrated. Methionine present in the CXXC motif that leads to Grx intramolecular auxotrophy is one common phenotype in bacteria resulting disulfide formation. This Grx disulfide is finally resolved by from S-thiolation of MetE in B. subtilis and E. coli in response an NADPH-dependent ferredoxin:thioredoxin reductase. In to oxidative stress (7, 19). In all Bacillus species, the conserved, the monothiol mechanism, the Grx-SSG intermediate is re- active-site Cys730 and the nonconserved Cys719 were iden- generated by GSH that is oxidized to GSSG and requires the tified as SSB sites for MetE. The observed S-bacillithiolation of GSH reductase to restore the reduced GSH pool (52). The LuxS and MetI could be required to inhibit homocysteine thioredoxin-like proteins YphP and YtxJ could possibly biosynthesis after NaOCl stress, since homocysteine is toxic function as putative Brxs. YphP is a DUF1094 family protein and accumulates when Met biosynthesis is inhibited, due to whose previously solved crystal structure adopts a thio- MetE-SSB modification (Fig. 3). redoxin-fold, has a conserved C53XC55 motif (with a reduction 20 CHI ET AL. potential of - 130 mV), and was demonstrated to display strain HB11002 (CU1065 trpC2 and bshA::mlsr) (12), B. amy- some weak disulfide isomerase activity (10). Under NaOCl loliquefaciens FZB42 (5), B. pumilus SBUG1799, B. megaterium stress, YphP is S-bacillithiolated at its more solvent-exposed SBUG1152 (36), and S. carnosus TM300 (43). Cultivation was active-site Cys53. The DUF2847 family Brx protein YtxJ is performed as detailed in the Supplementary Materials and another thioredoxin-like protein with a conserved TCPIS Methods section. motif reminiscent of the same redox-active motif found in many monothiol Grxs (18). YtxJ was identified in S. carnosus, Thiol redox proteome analysis and protein S-bacillithiolated in its TCPIS motif. The detection of S- identification using MALDI-TOF-TOF MS/MS bacillithiolated YphP and YtxJ during NaOCl stress could The thiol redox proteome analysis was performed as de- represent the trapping of transient intermediates of Brx- scribed (7). Cells were harvested at control and NaOCl stress mediated de-bacillithiolation pathways. The distribution of conditions, sonicated, alkylated in 8 M urea/1% chaps/ YphP and YtxJ homologs among bacteria encoding BSH bio- 100 mM iodoacetamide (IAM), and acetone-precipitated. The synthesis genes (12) further suggests their roles as Brx can- pellet was resolved in an urea/chaps buffer without IAM, and didates. Our current experiments are directed to further 200 lg of the protein extract was reduced with 10 mM Tris-(2- investigate the functions of these putative Brx proteins in the carboxyethyl)-phosphine and labeled with BODIPY FL C -IA de-bacillithiolation pathway. 1 [N-(4,4-difluoro-5,7-dimethyl-4-bora-3a,4a-diaza-s-indacene- We further analyzed whether the BSH/BSSB redox ratio 3-yl)-methyl)-iodoacetamide] (Invitrogen, Eugene, OR). The is affected by NaOCl stress that could contribute to S- fluorescence-labeled protein extract was separated using 2D bacillithiolation of proteins. In E. coli cells, a four-fold decrease PAGE, scanned for BODIPY fluorescence, and stained with in the GSH/GSSG redox ratio from 370:1 to 87:1 was observed Coomassie as described (7). Quantitative image analysis in menadione-treated cells (46). The two-fold increase of BSSB was performed with DECODON Delta 2D software (www post NaOCl stress is comparable to the increase of GSSG in .decodon.com). The oxidation ratios were calculated as E. coli cells exposed to menadione (46). The increased GSSG fluorescence/protein amount ratios and are listed in Sup- levels result in a decreased GSH/GSSG ratio that could con- plementary Table S1. The log2 oxidation ratios were used for tribute to protein S-glutathionylation spontaneously via thiol– hierarchical clustering analysis using Multi-Experiment disulfide exchange between GSSG and protein thiols (13). Viewer (MeV v.4.8) software (http://mev.tm4.org/) (32). However, this thiol–disulfide exchange is kinetically too slow Tryptic digestion of proteins, peptide spotting, and MALDI- and very unlikely in vivo, since it would require a drop in the TOF-TOF measurement were performed as described in the GSH:GSSG ratio from 100:1 to 1:1 for 50% conversion of Supplementary Materials and Methods section. protein thiols to protein GSH-mixed disulfides (13, 52). In our studies, the BSH:BSSB redox ratio dropped two-fold, con- Linear-trap quadrupole–Orbitrap Velos firming that BSH is oxidized to BSSB by NaOCl. This mass spectrometry BSH:BSSB ratio of 25:1 is unlikely to be sufficient for sponta- neous formation of protein-SSB via a direct thiol–disulfide Bacterial cells were sonicated in an urea/chaps alkylation exchange reaction between protein-SH and BSSB. Instead, S- buffer with 100 mM IAM and separated by 15% nonreducing bacillithiolation first requires oxidation of reactive Cys resi- SDS-PAGE. In-gel tryptic digests and Orbitrap Velos LC-MS/ dues to activated thiol derivatives, such as sulfenic acids (9, MS analysis were performed as described in the Supplemen- 15). NaOCl leads to chlorination of Cys residues to unstable tary Materials and Methods section. S-bacillithiolated pep- sulfenylchloride intermediates that rapidly react further to tides were identified by searching all MS/MS spectra in .dta form disulfides (50). Thus, we suggest that sulfenylchloride format against B. subtilis 168, B. megaterium, B. amyloliquefa- formation at reactive Cys residues by NaOCl and the de- ciens FZB42, B. pumilus, and S. carnosus TM300 target-decoy creased BSH/BSSB redox ratio contribute to the mechanism of protein sequence databases extracted from UniprotKB release S-bacillithiolation in Firmicutes bacteria. 12.7 using SorcererÔ-SEQUESTÒ according to the parameters Finally, the question arises if hypochlorite is a physiologi- and SEQUEST filter criteria as described in the Supplemen- cally relevant oxidant that mediates protein S-bacillithiolation tary Materials and Methods section. in Bacillus and Staphylococcus species. Hypochlorite is encountered by soil bacteria and pathogenic Bacillus and Preparation of keyhole limpet hemocyanin-BSH Staphylococcus species, since it is present in household bleach for generation of BSH-antibodies and other disinfectants, and it is also produced by activated Two milligrams of synthetic BSH (44) was conjugated to macrophages during the infection process by the enzyme 10 mg keyhole limpet hemocyanin (KLH) using the Mal- myeloperoxidase. The observed S-bacillithiolations in soil- eimide-Activated KLH Conjugation Kit (Sigma) according to dwelling Bacillus species after NaOCl stress might mimic the the details described in the Supplementary Materials and protective response to toxic ROS, such as hydroxyl radicals, Methods section. which are generated during respiration, explaining the conservation of some of the protein targets across different Nonreducing BSH-immunoblot analysis species. and nonreducing/reducing diagonal SDS-PAGE Materials and Methods Polyclonal BSH antisera were used at 1:500 dilution, and 25 lg IAM-alkylated protein extracts was used for nonre- Bacterial strains and growth conditions ducing 15% SDS-PAGE and immunoblot analysis as de- The bacterial strains used were B. subtilis wild-type strains scribed (6). Nonreducing/reducing diagonal SDS-PAGE 168 (trpC2) and CU1065 (trpC2 pheA1) and bshA-mutant analysis was performed as previously described (6). S-BACILLITHIOLATION AS THIOL PROTECTION MECHANISM IN FIRMICUTES 21

Quantification of reduced thiol redox buffer (RSH), Structure and function of Bacillus subtilis YphP, a prokaryotic oxidized thiol redox buffer (RSSR), and protein BSH disulfide isomerase with a CXC catalytic motif. Biochemistry 48: 8664–8671, 2009. B. subtilis CU1065 was grown in BMM to an OD600 of 0.44 11. Fratelli M, Demol H, Puype M, Casagrande S, Eberini I, and harvested before and after NaOCl stress for subsequent Salmona M, Bonetto V, Mengozzi M, Duffieux F, Miclet E, RSH, RSSR, and protein RSH quantification as described in Bachi A, Vandekerckhove J, Gianazza E, and Ghezzi P. the Supplementary Materials and Methods section. Identification by redox proteomics of glutathionylated pro- teins in oxidatively stressed human T lymphocytes. Proc Natl Acknowledgments Acad Sci U S A 99: 3505–3510, 2002. We thank the Decodon Company for support with Deco- 12. Gaballa A, Newton GL, Antelmann H, Parsonage D, Upton don Delta 2D software, Frieder Schauer for providing strains H, Rawat M, Claiborne A, Fahey RC, and Helmann JD. B. pumilus SBUG1799 and B. megaterium SBUG1152, and Dana Biosynthesis and functions of bacillithiol, a major low-mo- Clausen for excellent technical assistance. We are very grate- lecular-weight thiol in Bacilli. Proc Natl Acad Sci U S A 107: ful to Ahmed Gaballa and John D. Helmann for discussion of 6482–6486, 2010. 13. Gallogly MM and Mieyal JJ. Mechanisms of reversible pro- the results before publication and for providing the B. subtilis tein glutathionylation in redox signaling and oxidative bshA mutant. This work was supported by grants from the stress. Curr Opin Pharmacol 7: 381–391, 2007. Deutsche Forschungsgemeinschaft (AN746/2-1 and AN746/ 14. 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Abbreviations Used LC liquid chromatography ¼ LMW low molecular weight APS adenosine-5¢-phosphosulfate ¼ ¼ LTQ linear-trap quadrupole BMM Belitsky minimal medium ¼ ¼ Met methionine Brx bacilliredoxin ¼ ¼ MS/MS tandem mass spectrometry BSH bacillithiol ¼ ¼ MW molecular weight BSSB oxidized bacillithiol disulfide ¼ ¼ N5,N10-THF 5,10-methylenetetrahydrofolate CDG 7-carboxy-7-deazaguanine ¼ ¼ N5-THF 5-methyltetrahydrofolate CPH4 6-carboxy-5,6,7,8-tetrahydropterin ¼ ¼ OAS O-acetylserine Cys cysteine ¼ ¼ PAPS 3¢-phosphoadenosine-5¢-phosphosulfate DAHP 3-deoxy-d-arabino-heptulosonate ¼ ¼ PEP phosphoenol pyruvate 7-phosphate ¼ PAGE polyacrylamide gel electrophoresis DW dry weight ¼ ¼ PP inorganic pyrophosphate DxP 1-deoxy-d-xylulose 5-phosphate i ¼ ¼ PPP pentose phosphate pathway EPSP 5-enolpyruvylshikimate-3-phosphate ¼ ¼ preQ0 7-cyano-7-deazaguanine GMP guanosine 5¢-phosphate ¼ ¼ preQ1 7-aminomethyl-7-deazaguanine Grx glutaredoxin ¼ ¼ protein-SSB BSH protein-mixed disulfide GSH glutathione ¼ ¼ PRPP 5-phospho-alpha-d-ribose 1-diphosphate GSSG oxidized glutathione disulfide ¼ ¼ Pyr pyruvate GST glutathione S-transferase ¼ ¼ Q queuosine H2NTP 7,8-dihydroneopterin triphosphate ¼ ¼ RNS reactive nitrogen species HET-P 4-methyl-5-(beta-hydroxyethyl)thiazole ¼ ¼ ROS reactive oxygen species monophosphate ¼ RSH reduced thiol redox buffer HMP 2-methyl-4-amino-5-hydroxymethyl ¼ ¼ RSSR oxidized thiol redox buffer pyrimidine ¼ SAM S-adenosyl methionine HMP-PP 2-methyl-4-amino-5-hydroxymethyl ¼ ¼ SDS sodium dodecyl sulfate pyrimidine pyrophosphate ¼ IAM iodoacetamide THF tetrahydrofolate ¼ ¼ IMP inosine 5¢-phosphate TMP thiamine monophosphate ¼ ¼ KLH keyhole limpet hemocyanin TPP thiamine pyrophosphate ¼ ¼ LB Luria broth XMP xanthosine 5¢-phosphate ¼ ¼ ! ! "#$%&'(!)! ! ! ! ! ! ! ! ! ! ! ! ! ! ! Chapter 4 ! ! ! ! ! Structural insights into the redox-switch mechanism of the MarR/DUF24-type regulator HypR

!

Gottfried J. Palm#, Bui Khanh Chi#, Paul Waack, Katrin Gronau, Dörte Becher, Dirk Albrecht, Winfried Hindrichs, Randy J. Read, and Haike Antelmann*.

Nucleic Acids Research, 40: 4178 – 4192 (2012).

! ! ! ! ! ! ! ! !

# Both authors contributed equally to this work * corresponding author

! "#! 4178–4192 Nucleic Acids Research, 2012, Vol. 40, No. 9 Published online 11 January 2012 doi:10.1093/nar/gkr1316

Structural insights into the redox-switch mechanism of the MarR/DUF24-type regulator HypR Gottfried J. Palm1, Bui Khanh Chi2, Paul Waack1, Katrin Gronau2, Do¨ rte Becher2, Dirk Albrecht2, Winfried Hinrichs1, Randy J. Read3 and Haike Antelmann2,*

1Institute for Biochemistry, 2Institute of Microbiology, Ernst-Moritz-Arndt-University of Greifswald, D-17487 Greifswald, Germany and 3CIMR Haematology, University of Cambridge, Wellcome Trust/MRC Building, Cambridge CB2 0XY, England, UK

Received November 10, 2011; Revised December 21, 2011; Accepted December 22, 2011 Downloaded from ABSTRACT ROS that are produced as defense by the innate immune Bacillus subtilis encodes redox-sensing MarR-type system (1,2). During infection activated macrophages regulators of the OhrR and DUF24-families that release the enzyme myeloperoxidase that utilizes H2O2 to sense organic hydroperoxides, diamide, quinones produce the strong oxidant hypochloric acid (HOCl) to kill pathogenic bacteria (3,4). ROS can further generate or aldehydes via thiol-based redox-switches. In secondary reactive electrophilic species (RES) (5,6). http://nar.oxfordjournals.org/ this article, we characterize the novel redox-sensing Bacteria can sense and respond to ROS and RES by MarR/DUF24-family regulator HypR (YybR) that is expression of dedicated detoxification mechanisms. ROS activated by disulphide stress caused by diamide are sensed by redox-sensitive transcriptional regulators and NaOCl in B. subtilis. HypR controls positively a that undergo thiol-disulphide switches leading to activa- flavin oxidoreductase HypO that confers protection tion or inactivation of the transcription factors (7). The against NaOCl stress. The conserved N-terminal OxyR regulator of Escherichia coli is one of the best Cys14 residue of HypR has a lower pKa of 6.36 and studied bacterial peroxide-sensors. OxyR is activated by intramolecular disulphide formation resulting in transcrip- is essential for activation of hypO transcription by at UB Greifswald on November 16, 2012 disulphide stress. HypR resembles a 2-Cys-type tion of genes with antioxidant functions (8–11). In regulator that is activated by Cys14–Cys49 addition, the redox-controlled chaperone Hsp33 provides 0 specific protection against HOCl-induced protein aggrega- intersubunit disulphide formation. The crystal struc- tion in E. coli (12). Bleach leads to oxidation of the tures of reduced and oxidized HypR proteins were Zn-redox switch centres with subsequent Zn-release, oxi- resolved revealing structural changes of HypR upon dative unfolding, dimerization and activation of Hsp33. oxidation. In reduced HypR a hydrogen-bonding Organic peroxides are sensed by the conserved network stabilizes the reactive Cys14 thiolate that MarR-type repressor OhrR that controls the thiol- ˚ is 8–9 A apart from Cys490. HypR oxidation breaks dependent peroxidase OhrA (7,13). The OhrR family these H-bonds, reorients the monomers and includes one- and two-Cys OhrR-proteins that differ moves the major groove recognition a4 and a40 in their redox-sensing mechanisms. OhrRXc of helices 4A˚ towards each other. This is the first Xanthomonas campestris is the prototype of the two-Cys crystal structure of a redox-sensing MarR/DUF24 family that is oxidized to an intermolecular disulphide family protein in bacteria that is activated by between the opposing OhrR subunits (14,15). One-Cys OhrR proteins harbour one conserved N-terminal Cys NaOCl stress. Since hypochloric acid is released with the prototype of B. subtilis OhrRBs that is oxidized by activated macrophages, related HypR-like regu- to S-bacillithiolated OhrR in response to cumene lators could function to protect pathogens against hydroperoxide and HOCl (7,16,17). Redox-sensing the host immune defense. MarR-type regulators of pathogenic bacteria include the OhrR-paralogs MgrA and SarZ of Staphylococcus aureus as global regulators for antibiotic resistance, virulence and INTRODUCTION anaerobiosis, the multidrug-efflux regulator MexR and Reactive oxygen species (ROS) can be generated as a the oxidative stress response and pigment production byproduct of respiration. Pathogenic bacteria encounter regulator OspR of Pseudomonas aeruginosa (7,13,18–23).

*To whom correspondence should be addressed. Tel: +49 3834 864237; Fax: +49 3834 864202; Email: [email protected]

The authors wish it to be known that, in their opinion, the first two authors should be regarded as joint First Authors.

ß The Author(s) 2012. Published by Oxford University Press. This is an Open Access article distributed under the terms of the Creative Commons Attribution Non-Commercial License (http://creativecommons.org/licenses/ by-nc/3.0), which permits unrestricted non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited. Nucleic Acids Research, 2012, Vol. 40, No. 9 4179

In addition, B. subtilis encodes redox-sensing MarR/ Integration and deletion of the ohrA and hypO genes DUF24-family regulators that sense specifically electro- were confirmed by PCR. philes (diamide, quinones or aldehydes) (7,24–28). The paralogous repressors YodB and CatR are inactivated Construction of the hypRC14S, hypRC49S and via intermolecular disulphide formation by diamide and hypRC14,49S point mutants quinones resulting in derepression of the azoreductase Plasmids pDGhypR, pDGhypRC14S, pDGhypRC49S, (AzoR1), nitroreductase (YodC) and thiol-dependent pDGhypRC14,49S were produced by using PCR muta- dioxygenase (CatE) catalysing the reduction or ring- genesis. Using primers yybR-C14S-for1 and yybR- cleavage of the electrophiles (24,25,27). Other proteins of C14S-rev2 and B. subtilis wild-type chromosomal DNA, the MarR/DUF24 family (HxlR) and MerR/NmlR- the hypR gene was amplified by PCR, the PCR product families (AdhR) sense aldehydes (formaldehyde and digested with EcoRI and BamHI restriction enzymes and methylglyoxal) via conserved Cys residues (26,28–30). inserted into plasmid pDG795 digested with the same However, the genome of B. subtilis encodes MarR/ enzymes to generate pDGhypR. For construction of DUF24 family regulators of unknown functions, pDGhypRC14S, first PCR was performed in two separate including YybR, YdeP, YdzF, YkvN and YtcD reactions using primers yybR-C14S-for1 with yybR-C14S- (Supplementary Figure S1A). rev1 and primers yybR-C14S-for2 with yybR-C14S-rev2 In this study, we characterize the hypochloric acid- (Supplementary Table S2) and B. subtilis 168 chromosom- Downloaded from specific regulator YybR (renamed HypR) as a novel al DNA as template. The PCR products were hybridized MarR/DUF24 transcriptional regulator that positively and amplified by a second PCR using primers yybR-C14S- controls the putative nitroreductase YfkO. HypR resem- for1 and yybR-C14S-rev2. The PCR products were bles a two-Cys-type MarR-type regulator that is activated digested with EcoRI and BamHI and inserted into by Cys14–Cys490 intersubunit disulphide formation. plasmid pDG795 digested with the same enzymes to http://nar.oxfordjournals.org/ We present the crystal structure of HypR under reduced generate pDGhypRC14S. For construction of pDGhyp and oxidized conditions and provide for the first time RC49S and pDGhypRC14,49S, first PCRs were insights into the redox-sensing mechanism of a MarR/ performed in two separate reactions using primers yybR- DUF24 family regulator. C49S-for1 with yybR-C49S-rev1 and primers yybR-C49S- for2 with yybR-C49S-rev2 (Supplementary Table S2) and B. subtilis 168 chromosomal DNA and pDGhypRC14S MATERIALS AND METHODS plasmid DNA as templates, respectively. The PCR products were hybridized and amplified by a second Bacterial strains and growth conditions PCR using primers yybR-C49S-for1 and yybR-C49S- at UB Greifswald on November 16, 2012 The bacterial strains used were B. subtilis 168 (trpC2), rev2. The PCR products from the second PCRs were ÁhypR (trpC2,hypR::Cmr), ÁhypO (trpC2,hypO::Cmr), digested with EcoRI and BamHI and inserted into ÁohrA (trpC2,ohrA::Spcr), ÁhypODohrA (trpC2,hypO:: plasmid pDG795 digested with the same enzymes to Cmr,ohrA::Spcr) and the hypRC14S, hypRC49S, generate plasmids pDGhypRC49S and pDGhyp hypRC14,49S point mutants (Supplementary Table S1). RC14,49S. The plasmids pDGhypRC14S, pDGhyp Bacillus subtilis strains were cultivated under vigorous agi- RC49S and pDGhypRC14,49S were verified by DNA sequencing and transformed into the thrC locus of the tation at 37C in Belitsky minimal medium described pre- viously (31). Escherichia coli strains were grown in LB for B. subtilis ÁhypR mutant. DNA manipulation. The antibiotics were used at the fol- Northern blot experiments lowing concentrations: 1 mg/ml erythromycin, 25 mg/ml lincomycin, 5 mg/ml chloramphenicol, 10 mg/ml kanamy- Northern blot analyses were performed as described (17) cin, 100 mg/ml spectinomycin. The compounds used were using RNA isolated from B. subtilis wild-type cells be- 2-methylhydroquinone (Acros), diamide (diazinedi- fore (control) and 10 min after the treatment with 50 mM carboxylic acid bis(N,N-dimethylamide) (Sigma) and NaOCl, 1 mM diamide and 0.5 mM MHQ. Hybridizations sodium hypochlorite (15% stock solution) (Sigma). specific for hypR and hypO were performed with the Gene deletions for construction of the hypO mutant digoxigenin-labelled RNA probes synthesized in vitro were generated using long-flanking-homology polymerase using T7 RNA polymerase from T7 promoter containing chain reaction (LFH–PCR) as previously described (25). internal PCR products of the respective genes using the Primers yfkO-F1 and yfkO-F2 were used to amplify primer sets yybR-T7for with yybR-T7rev and yfkO-T7for the up fragment and primers yfkO-R1 and yfkO-R2 to with yfkO-T7rev (Supplementary Table S2). amplify the down fragment, respectively (Supplementary Table S2). Fragments were amplified and joined together Primer extension experiments with the chloramphenicol cassette using Pfusion DNA Primer complementary to the N-terminus encoding region polymerase (Invitrogen) as described (32). Plasmid of hypO (yfkO-FT-rev2) and hypR (yybR-C14S-rev1) were r pCm::spec was used to replace the Cm cassette with a 50-end labelled using T4 polynucleotide kinase (Roche r Spec marker to generate strain ÁohrA::Spec (17). The Diagnostics) and 50 mCi [g32P]-ATP (GE Healthcare). ÁohrADhypO double mutant was constructed by trans- Primer extension analysis was performed using the formation of chromosomal DNA of the hypO mutant labelled primers as described previously (25). Sequencing into competent cells of the ÁohrA::Spec mutant strain. of the corresponding promoter regions was performed 4180 Nucleic Acids Research, 2012, Vol. 40, No. 9 using PCR products as templates containing the promoter and the radiolabelled bands were visualized using region of the respective genes amplified with primer set phosphoimaging. yybR-C14S-for1 and yybR-Cys14S-rev1 and yfkO-FT- for2 and yfkO-FT-rev2 (Supplementary Table S2). DNase-I footprinting analysis Primer yfkO-FT-for2 and yfkO-FT-rev2 were each 5 -end Expression and purification of recombinant His-HypR, 0 labelled using T4 polynucleotide kinase (Roche His-HypRC14S and His-HypRC49S proteins Diagnostics) and 50 mCi [g32P]-ATP (GE Healthcare) Escherichia coli BL21(DE3)pLysS (Invitrogen) was used and purified using ethanol precipitation. DNA probes for overproduction of His-tagged HypR, HypRC14S for hypO (corresponding to positions 160 to +79 and HypRC49S proteins. For expression of His-HypR, relative to TSS), were synthesized by PCRÀ amplification His-HypRC14S and His-HypRC49S proteins, the hypR using one 50-end-labelled primer and the correspond- coding sequence was amplified by PCR using primers ing non-labelled primer, respectively (Supplementary yybR-NdeI-pETfor and yybR-BamHI-pETrev and Table S2). Purified PCR fragments that are labelled at chromosomal DNA of the B. subtilis wild-type and the one 50-end were used as sequencing templates. DNA hypRC14S and hypRC49S point mutants as templates probe purification and DNase I footprinting was per- (Supplementary Table S2). The reverse primer includes formed as described previously (25). Downloaded from the codons for six C-terminal histidine residues. The hypR specific PCR products were digested with NdeI In vitro transcription assay and BamHI and inserted into the pET11b expression In vitro transcription assays were performed as described plasmid digested with the same enzymes to generate previously (33) using 0.5 mM HypR, HypRC14S and plasmids pEThypR, pEThypRC14S and pEThypRC49S. HypRC49S proteins which were added to the in vitro tran- The hypR, hypRC14S and hypRC49S mutant sequences http://nar.oxfordjournals.org/ scription reactions. Templates for a 380-bp PCR product were verified by DNA sequencing (Supplementary containing the hypO promoter were generated by Table S2). Escherichia coli BL21(DE3)pLysS carrying PCR with primers yfkO-FT-for2 and yfkO-FT-rev3 the pEThypR, pEThypRC14S and pEThypRC49S expres- (Supplementary Table S2), gel purified and used at sion plamids was cultured in 1 l LB medium, and 1 mM 0.1 pmol in each reaction. Core E. coli RNA polymerase IPTG (isopropyl- -D-thiogalactopyranoside) was added at b [Epicentre Biotechnologies (Madison, WI, USA)] was the mid-log phase (OD of 0.8) for 2 h. Recombinant 600 used at 2 pmol per reaction and combined with 8 pmol His-tagged HypR, HypRC14S and HypRC49S proteins purified SigmaA protein from B. subtilis (molar ratio were purified using PrepEaseTM His-Tagged High Yield 1:4) that was expressed from plasmid pNG590 according at UB Greifswald on November 16, 2012 purification Resin (USB) under native conditions accord- to (34). RNAP holoenzyme formation was allowed for ing to the instructions of the manufacturer. HypR was 15 min on ice prior to addition to the reactions containing eluted in 50 mM NaH PO , 300 mM NaCl, and 250 mM 2 4 HypR protein that was reduced by DTT or oxidized by imidazole pH 8.0. The proteins were further purified by diamide and NaOCl. Transcription reactions were per- anion exchange chromatography (POROS 20 HQ, formed for 15 min at 37 C and terminated by addition Applied Biosystems) with 20 mM Tris–HCl pH 7.5 and a  of stop buffer. Reactions were separated on 8.3 M urea– 0 to 1 M NaCl gradient followed by dialysis into 10 mM 6% polyacrylamide gels and the run-off transcripts Tris–HCl pH8.0, 100 mM NaCl, 50% glycerol (v/v) and visualized by phosphoimaging. stored at 80 C. À  Western blot analysis DNA gel mobility shift assays Anti-HypR polyclonal rabbit antiserum was generated DNA fragments containing the hypO promoter region using purified His-tagged HypR protein. The polyclonal were generated by PCR using the primer sets yfkO- antisera were used for immunoprecipitation experiments FT-for2 and yfkO-FT-rev2 (Supplementary Table S2). and western blot experiments at 1:200 dilution. Protein Approximately 600 pmol of the purified PCR product amounts of 25 g were loaded onto a 15% SDS–PAGE was end-labelled using T4 polynucleotide kinase (Roche m 32 gel and the western blot analysis was performed as Diagnostics) and 50 mCi [ P]-ATP (GE Healthcare). The g described previously (25). labelled hypO-specific promoter probe was purified by ammonium sulphate/ethanol precipitation and 2000 cpm Immunoprecipitation and non-reducing/reducing diagonal was incubated with different amounts of purified HypR SDS–PAGE analysis proteins for 10 min at room temperature in EMSA- binding buffer (10 mM Tris–HCl pH 7.5, 100 mM KCl, Bacillus subtilis cells were treated with 1 mM diamide and 5% glycerol (v/v) in the presence of 50 mg/ml of BSA 50 mM NaOCl and harvested with 50 mM iodoacetamide and 5 mg/ml of Salmon sperm DNA. The concentrations (IAM) to alkylate all reduced thiols. Cells were sonicated of the compounds used for the DNA-binding assays were and the protein extracts obtained after repeated centrifu- 1 mM NaOCl, 1 mM diamide, 1 mM DTT. DNA-binding gation. Immunoprecipitation using HypR-specific reactions were separated by 4% native polyacrylamide gel antibodies was performed with Dynabeads-ProteinA electrophoresis in 10 mM Tris–HCl pH 8, 1 mM EDTA (Invitrogen) according to the instructions of the manufac- buffer, containing 2.5% glycerol at room temperature turer. The precipitated proteins were eluted by boiling and constant voltage (250 V) for 15 min. Gels were dried in non-reducing SDS sample buffer (4% SDS; 62.5 mM Nucleic Acids Research, 2012, Vol. 40, No. 9 4181

Tris–HCl pH 8.0, glycerol). Immunoprecipitated HypR 150 mM NaCl, 20 mM Tris–HCl pH 8.0, 5 mM DTT protein was separated using the non-reducing/reducing and crystallized from 12% PEG 8000 by the hanging diagonal SDS–PAGE analysis as described previously drop vapour diffusion method. Prism-shaped crystals (35) and subjected to HypR-specific western blot analysis. appeared after a few days. Crystals were briefly soaked in 20% PEG 4000, 10% PEG 400 and cryocooled in the Proteome analysis N2-stream at 100 K. Oxidized HypR was crystallized from 35 6% PEG 4000, 0.4 M Li SO in the form of thick hex- Preparation of cytoplasmic L-[ S]methionine-labelled 2 4 proteins from cells treated with 0.5 mM MHQ or 1 mM agonal plates or prisms. Fifteen per cent PEG 8000, diamide and separation by 2D gel electrophoresis (2D– 15% PEG 400 were used as cryoprotectant. Diffraction PAGE) was perfomed as described (36). The image data were collected at beamline BL 14.2 at the synchro- analysis was performed with the DECODON Delta 2D tron BESSY in Berlin, Germany (Table 1). software (http://www.decodon.com). The structure of reduced HypR was solved by molecular replacement (37) using the YtcD structure from B. subtilis MALDI–TOF mass spectrometry of in vitro (PDB entry 2 hzt, 49% sequence identity for 133 amino oxidized HypR protein acids). Refmac5 version 5.5.0110 (38) was used to refine using TLS and no NCS restraints (statistics, see The HypR protein was oxidized with NaOCl and Table 1). One ligand per monomer was identified as Downloaded from alkylated with IAM introducing a mass shift of 57 Da in b-mercaptoethanol. A monomer of reduced HypR was reduced Cys residues. Then HypR protein was reduced used to solve the structure of the oxidized protein by with DTT and alkylated with NEM leading to a mass molecular replacement. Pseudosymmetry of the eight shift of 125 Da in oxidized Cys. HypR protein was monomers in the asymmetric unit about a 2-fold axis on separated by non-reducing SDS–PAGE, digested with a causes nearly perfect twinning of the P65 crystals with http://nar.oxfordjournals.org/ trypsin and the peptides were spotted onto MALDI- apparent P6522 symmetry in the diffraction pattern. To targets (Voyager DE-STR, PerSeptive Biosystems) and reduce the influence of model bias, the density was measured using a Proteome-Analyzer 4800 (Applied improved by non-crystallographic symmetry averaging Biosystems, Foster City, CA, USA) as described (25). over the eight copies in the asymmetric unit, using the RESOLVE density modification algorithm (39) as imple- Orbitrap-mass spectrometry of oxidized HypR mented in Phenix (40). Stained gel-bands of immunoprecipitated HypR harvested Secondary structure elements were assigned by DSSP from wild-type cells or purified His–HypR protein were (41). The dimerization interface was calculated by the tryptic digested. Tryptic peptides were separated and PISA server (42). Figures of structural models were at UB Greifswald on November 16, 2012 measured online by ESI-mass spectrometry using a made with PyMOL (http://www.pymol.org). The struc- nanoACQUITY UPLCTM system (Waters, Milford, tural data were submitted to the PDB database under MA, USA) coupled to an LTQ OrbitrapTM XL mass spec- the PDB entry 4a5n for reduced HypR and 4a5m for trometer (Thermo Fisher Scientific, Waltham, MA, USA) oxidized HypR. as described (17). Post-translational modifications of HypR were identified by searching all MS/MS spectra DTNB assay to determine the pKa of Cys14 and Cys49 against a B. subtilis target-decoy protein sequence residues in His-HypR database extracted from UniprotKB release 12.7 using TM Õ The purified His-HypRC14S and His-HypRC49S mutant Sorcerer -SEQUEST . The Sequest search was carried proteins were used to determine the pK of the Cys14 out considering the following parameter: parent ion a and Cys49 residues by reaction with 5,50-dithiobis- mass tolerance of 10 ppm, fragment ion mass tolerance (2-nitrobenzoic acid) (DTNB) generating 2-nitro-5 of 1.00 Da, Methionine oxidation (+15.99492Da), thiobenzoate. Defined amounts of proteins (4.66 mM cysteine carbamidomethylation (+57.021465Da) and HypRC14S or 4.93 mM HypRC49S) reacted with Cys+ICPSITQR peptide (+914.4873 Da) were set as 10–20 mM DTNB in phosphate/citrate buffer (50 mM variable modifications. Sequest identifications required at phosphate, 50 mM citrate, 100 mM NaCl) at various pH least ÁCn scores >0.10 and XCorr scores more than 2.2, values ranging from pH 6 to 8.5. The pH-dependent rate 3.3 and 3.75 for doubly, triply and quadruply charged of reactions k(pH) of the HypR Cys mutants with DTNB peptides. was measured as time-dependent absorbance change at 410 nm at room temperature using a Cary 50 spectropho- Crystallization, data collection and refinement of reduced tometer (Agilent Technologies, Waldbronn, Germany). HypRC14S and oxidized HypR protein The following equation was used for a first fit of the His-tagged HypRC14S and HypR proteins were purified time-dependent absorbance change by non-linear using Ni-affinity chromatography and anion-exchange regression: k pH t chromatography. The HypRC14S protein was purified A410 t A410,0+ÁA410 1 eÀ ð Þ . The resulting with b-mercaptoethanol and used for crystal structure pH-dependentð Þ¼ rate of reactionsð À k(pH)Þ were used in a determination of reduced HypR. The wild-type HypR second fit to obtain the pKa of the thiol. The first order protein was oxidized with 1 mM diamide before purifica- kinetic constant k(pH) is proportional to the fraction of tion by anion exchange chromatography. Reduced deprotonated thiol, which is pH dependent: const HypRC14S protein was concentrated to 15 mg/ml in k pH pKa thiol pH. ð Þ¼1+10 ð ÞÀ 4182 Nucleic Acids Research, 2012, Vol. 40, No. 9

Table 1. Crystallographic data collection and refinement statisticsa

Reduced HypRC14S Oxidized HypR (+b-Mercaptoethanol) (+Diamide)

PDB entry 4a5n 4a5m Data collection Space group P21 P65 Unit cell parameters a, b, c (A˚ ) 54.93, 68.03, 60.42 84.6, 84.6, 358.6 , , () 90.00, 97.45, 90.00 90.00, 90.00, 120.00 Resolution (A˚ ) 36.46–1.81 (1.90–1.81) 100.00–3.00 (3.18–3.00) Wavelength (A˚ ) 0.918 0.918 Reflections, unique 39025 (5528) 24764 (3457) Multiplicity 2.3 (2.2) 5.5 (4.6) Completeness (%) 96.5 (94.0) 89.8 (77.4) Mean, I/s(I) 8.9 (2.2) 14.9 (1.4) Mosaicity () 1.15 (Scala) 0.31 (XDS) Wilson B-factor (A˚ 2) 23.4 82 R 0.080 (0.531) 0.087 (1.089)

meas Downloaded from Refinement Resolution 59.9–1.81 45.6–3.0 Protein atoms 3408 6408 Solvent / ligand atoms 315/20 25/37 Average B factor (A˚ 2) 37.2 107.9 Rcryst/Rfree 0.183/0.223 0.207/0.250 ˚ r.m.s.d. bond lengths/angles (A, ) 0.013/1.53 0.008/1.73 http://nar.oxfordjournals.org/

aValues in parentheses belong to the highest resolution shell.

RESULTS The HypR-controlled nitroreductase HypO confers Identification of HypR (YybR) as hypochlorite-sensing resistance to NaOCl stress positive regulator of the oxidoreductase Previous phenotype analyses have shown that the at UB Greifswald on November 16, 2012 HypO (YfkO) OhrR-controlled OhrA peroxiredoxin is a specific deter- Previous transcriptome analyses revealed that the minant of NaOCl detoxification (17). Recently, it was unknown DUF24-type regulator encoded by yybR shown that HypO shows nitroreductase activity in vitro responds strongly to diamide, quinones and hypochlorite (47). We analysed the growth phenotype of ÁhypR and (Figure 1B) (17,43,44). In addition, several FMN- ÁhypO single and ÁhypOÁohrA double mutants to inves- dependent NAD(P)H oxidoreductase-encoding genes tigate whether the HypO nitroreductase contributes to were up-regulated, including yfmJ, ytkL, ywnB, yqiG, NaOCl resistance. No differences in the sensitivities of yqjM, ywrO, ydeQ, yugJ and yfkO (Figure 1A and B) the ÁhypO and ÁhypR single mutants were detected (43). Some of these redox enzymes are members of the compared to the wild-type (Supplementary Figure S3A Spx regulon, such as yqiG and yugJ suggesting that and S3B). However, the growth of the ÁhypOÁohrA these could function in maintenance of the thiol-redox double mutant was significantly more impaired than that homeostasis (45,46). We were interested if the of the ÁohrA single mutant after treatment with 75 mM DUF24-family regulator YybR controls one of these NaOCl (Figure 2 and Supplementary Figure S3C). This oxidoreductases. Using northern blot analysis we found indicates that HypO also provides protection against that the NAD(P)H-flavin oxidoreductase encoded by NaOCl stress in B. subtilis. yfkO was strongly induced in the wild-type but not in the yybR mutant in response to diamide, NaOCl and MHQ stress (Figure 1C). This suggests that yybR positive- Cys14 and Cys49 of HypR are essential for activation ly controls yfkO transcription in response to diamide and of hypO transcription NaOCl stress. In addition, yybR transcription is probably HypR has two Cys residues (Cys14 and Cys49) and Cys14 also strongly autoregulated by thiol-specific stress condi- is conserved among DUF24-family regulators tions. Hence, yybR was renamed as hypochlorite- (Supplementary Figure S1A). We investigated the role of responsive regulator hypR and yfkO was renamed as Cys14 and Cys49 of HypR in regulation of hypO tran- hypO. The northern blot analysis further revealed that scription in hypRC14S, hypRC49S and hypRC14,49 S hypR and hypO are transcribed monocistronically. Using mutants in vivo (Figure 1C). Northern blot analysis primer extension, the 50-ends of the hypO and hypR revealed that hypO transcription was abolished in the specific transcripts were mapped at T and A, respectively, hypRC14S, hypRC49S and hypRC14,49 S mutants by located 31 and 55 bp upstream of the start codon diamide and NaOCl in vivo. These results clearly (Supplementary Figure S2). indicate, that both Cys14 and Cys49 of HypR are essential Nucleic Acids Research, 2012, Vol. 40, No. 9 4183

A TrxB MhqE TrxB CysK CysK YcnD YcnDYfjR YfjR NfrA YwfI YkuQ NfrA YkuQ MrgA-2 YhfK YwfI YhfK MhqN YvyD YvyD HypO ClpP AzoR2 ClpP HypO SodA AzoR2 SodA YodC YodC AzoR1 MhqD AzoR1

control control diamide MHQ

B NaOCl Diamide MHQ Chromanon Function yfkO (hypO) 39,5 30,9 2,8 6,8 3,5 5,1 5,6 6,3 NAD(P)H-flavin oxidoreductase

yybR Downloaded from (hypR) 29,2 44,2 43,8 50,8 8,1 15 3,8 6,3 MarR/DUF24 family regulator

C WT DhypR hypRC14S hypRC49S hypRC14,49S hypR

aiDoC NaOCl QHM Co Dia NaOCl Co Dia NaOCl lCOaNaiDoC Co Dia NaOCl aiDoC NaOCl QHM http://nar.oxfordjournals.org/ hypO

hypR

23S rRNA

16S rRNA at UB Greifswald on November 16, 2012 Figure 1. HypO induction in the proteome (A), transcriptional induction of hypO and hypR in the transcriptome (B) and northern blot analysis of hypO and hypR transcription (C) under thiol-specific stress conditions. (A) Cytoplasmic proteins were labelled with 35S-methionine before (control) and after stress exposure and separated by 2D–PAGE as described (36). Close-ups of the overlay proteome images are shown for the wild-type before (green images) and 10 min after exposure to 1 mM diamide (left, red image) or 0.5 mM MHQ (right, red image). Proteins with increased protein synthesis ratios after MHQ and diamide stress including HypO are labelled that were identified from Coomassie-stained 2D gels as described (36). (B) The values represent fold-changes of hypR and hypO induction ratios in two biological replicates of transcriptome experiments of cells treated with 1 mM diamide, 0.5 mM MHQ, 50 mM NaOCl and 100 mM chromanon according to previously published transcriptome data (17,26,44). (C) Northern blot analysis was performed using RNA isolated from B. subtilis wild-type, the ÁhypR mutant and the ÁhypR mutant complemented with hypR, hypRC14S, hypRC49S and hypRC14,49S before (co) and 10 min after treatment with 0.5 mM MHQ, 1 mM diamide and 50 mM NaOCl. The arrows point toward the hypO and hypR specific transcripts. The methylen-blue stained northern blot is shown below as RNA loading control and the 16S and 23S rRNAs are labelled.

10 for activation of hypO transcription by disulphide stress in vivo. 1 Transcription of hypR is possibly autoinduced in the 500 wild-type and in the hypR complemented ÁhypR mutant OD WT control strain by disulphide stress. However, the hypR-specific 0,1 WT 75µM NaOCl mRNA amounts were also induced in the hypRC14S, ∆ohrA control ∆ohrA 75µM NaOCl hypRC49S and hypRC14,49 S mutants after diamide ∆ohrA∆hypO control stress, although hypR transcription was strongly decreased ∆ohrA∆hypO 75µM NaOCl 0,01 in the hypRC14S and hypRC14,49 S mutants compared to 0 30 60 90 -90 -60 -30 120 150 180 210 240 270 300 330 360 390 the wild-type (Figure 1C). The response of the HypRC49S -150 -120 Time (min) protein to disulphide stress could be caused by Figure 2. The OhrA peroxiredoxin and the nitroreductase HypO S-thiolation of the redox-sensitive Cys14 residue, leading protect cells against hypochlorite toxicity. Growth phenotype of to autoinduction of hypR transcription. It is also possible B. subtilis wild-type (WT), ÁohrA and ÁohrAÁhypO mutant strains that hypR transcription is controlled by another MarR/ that were treated with 75 mM NaOCl at an OD500 of 0.4. The growth DUF24-family paralog (e.g. YdeP or YkvN) in the curves are representives of at least three independent growth experi- ments. Two biological replicates are shown in Supplementary Figure hypRC14S, hypRC49S and hypRC14,49 S mutants S3A–S3C. explaining the response of the hypRCys mutants. 4184 Nucleic Acids Research, 2012, Vol. 40, No. 9

Identification of HypR operator sites and effect reduced HypRC14S and HypRC49S mutant proteins of disulphide stress on DNA-binding activity of with 0.14 and 0.12 mM, respectively and oxidation HypR in vitro caused no significant change in the DNA-binding affinities DNase-I footprinting analysis was performed to identify of the Cys mutant proteins (Supplementary Figure S4A the cis-acting sequences which function as operator sites and S4B). However, we observed a change in the mobility for HypR binding in the hypO upstream region in vitro. of oxidized HypR compared to reduced HypR in the HypR-His protein protected a region upstream of the gel-shift assays which was DTT-reversible (Figure 3B). hypO promoter from positions 53 to 95 relative to In addition, the DNase-I footprinting analysis showed the transcription start site (FigureÀ 3A).À The protected a higher affinity of oxidized HypR protein to the region contains a 7-2-7 bp inverted repeat GTATCAA hypO promoter region indicating an increased AATTGATAC that is also present at positions +24 to DNA-binding activity of oxidized HypR protein in vitro +39 downstream of the hypR promoter (Figure 4). The (Figure 3A). position of this HypO-box confirms the notion that HypR is a positive transcriptional regulator of hypO tran- Transcription of hypO is activated by HypR after diamide scription, but probably represses its own transcription. and NaOCl stress in vitro

Furthermore, we were interested if the DNA-binding Downloaded from activity is affected by diamide and NaOCl in vitro and Next, we analysed whether transcription of hypO is performed gel-shift and DNase-I footprinting analysis of activated by oxidized HypR using an in vitro transcription HypR protein under reduced and oxidized conditions. The assay. In brief, using E. coli RNA polymerase core enzyme gel-shift experiments showed binding of HypR to the (RNAP) and purified SigmaA and HypR proteins from B. hypO operator sites at similar affinities under reduced subtilis we performed the in vitro transcription assay for and oxidized conditions (Figure 3B). The calculated dis- the hypO gene as described previously (33). This assay http://nar.oxfordjournals.org/ sociation constants (Kd) were 0.18 mM for reduced HypR, showed the production of a hypO run-off transcript of 0.14 mM for diamide-oxidized HypR and 0.12 mM for the expected size of 220 bp only when HypR was present NaOCl-oxidized HypR proteins (Supplementary Figure in the in vitro reactions, indicating that HypR activates S4A and S4B). This indicates no significant change in transcription of the hypO gene in vitro (Figure 5). the DNA-binding affinities of reduced and oxidized Transcription of hypO increased 2.5-fold when HypR  HypR proteins. Similar Kd values were calculated for protein was oxidized by diamide and NaOCl treatment at UB Greifswald on November 16, 2012

diamide A DTT B 0.015 0.03 0.060.12 0.25 0.5 1 µM HypR A C G T 0.1 0.2 0.5 1 0.1 0.2 0.5 1 µM HypR - HypR DTT

HypR +1 Dia -10

-35 HypR NaOCl

-53 HypR Dia HypR +DTT

HypRC14S DTT

-95 HypRC49S DTT

Figure 3. DNase-I footprinting experiments (A) and gel-shift experiments (B) of purified HypR protein to the hypO promoter in the presence of DTT, diamide and NaOCl. (A) The HypR-protected operator sequence is indicated at the right side of the DNase-I footprint including an 7-2-7 bp inverted repeat with the sequence GTATCAAAATTGATAC that is labelled by arrows in the sequence alignment in Figure 4. The positions relative to the transcriptional start site are shown on the left. Transcription start site is indicated by (+1). For dideoxynucleotide sequencing, the dideoxy nucleotide added in each reaction is indicated above the corresponding lane. HypR protein was treated with 1 mM DTT or 1 mM diamide prior to the DNA-binding reactions. (B) EMSAs were used to analyse the effect of DTT, 1 mM diamide and 100 mM NaOCl on the DNA-binding activity of purified HypR, HypRC14S and HypRC49S proteins to the labelled hypO promoter probe. The HypR protein amounts used for the DNA-binding reactions are indicated. The EMSA experiments are representives of three replicate experiments. The change in the DNA-binding affinity and dissociation constants (Kd) of reduced and oxidized HypR, HypRC14S and HypRC49S proteins are calculated in Supplementary Figure S4A and B. Nucleic Acids Research, 2012, Vol. 40, No. 9 4185

hypO -95 -53 GGCGCCTGCCTGCGGCAGCGCGCTTTTTTGTTTGGTATCAAAATTGATACTATCCAAC -35 -10 +1 AAAAATGTGCGTACTTTTACTTTTTATCATAATGGCTACAATAGAGATGAGAGTAATC

TAAAGGAGAGGTGTTTTACATG

hypR -35 -10 +1 TAAGAAGCATTTTCTTTGAGCTAAGGAGAAAAGGTGTGTAAAATACTGGGTAGA

TTAGTTGCAAAATGTAATGTAGTATCAAAAAAGGTACTATTGGAGGGTCAAGAATG

hypR +24 GTATCAAAAAAGGTAC +39 Downloaded from hypO -71 GTATCAAAATTGATAC -56

HypR-box Figure 4. Sequence alignment of the HypR-boxes in the hypO and hypR promoter regions. The hypO and hypR promoter sequences ( 10 and 35), the transcription start site (+1) and the ATG start codons are underlined in the hypO and hypR upstream regions. The conservedÀ HypR-boxesÀ http://nar.oxfordjournals.org/ including the inverted repeats are boxed and indicated by arrows. The HypR protected region identified by the DNase-I footprinting analysis is grey shaded.

ABCDHypR HypR HypRC14S HypRC14S RNAP RNAP RNAPRNAP RNAP RNAP RNAP RNAP RNAP RNAP RNAP RNAP RNAP RNAP RNAP RNAP SigA SigA SigA SigA SigA SigA SigA SigA SigA SigA SigA SigA SigA SigA SigA DTT Dia Dia NaOCl NaOCl M M DTT Dia NaOCl M DTT Dia NaOCl DTT Dia Dia at UB Greifswald on November 16, 2012 DTT DTT M DTT

700 700 700 700 600 600 600 600 500 500 500 500

400 400 400 400 300 300 300 300 hypO mRNA 200 200 200 200

100 100 100 100

Figure 5. In vitro transcription analysis of hypO in the presence of RNA polymerase holoenzyme (RNAP) and purified HypR (A and B) and HypRC14S proteins (C and D) treated with DTT, diamide or NaOCl. The reactions in (A) and (C) show increased hypO transcription ratios by oxidized HypR but not by oxidized HypRC14S mutant protein. The reactions in (B) and (D) show reduced hypO transcription ratios by oxidized HypR that is subsequently reduced with DTT, but no change in hypO transcription by oxidized HypRC14S that is subsequently reduced with DTT. The in vitro-transcription analyses of hypO are representives of three replicate experiments and the relative transcription ratios were quantified in Supplementary Figure S4C. RNA size standard was generated using the Perfect RNA marker template mix (Novagen). The hypO specific run-off transcript is labelled at a size of 220 bp.

compared to reactions with DTT-reduced HypR in vitro transcription reactions containing oxidized (Figure 5A and Supplementary Figure S4C). This indi- HypR protein that was subsequently reduced with DTT cates that the increased DNA-binding affinity of (Figure 5B and Supplementary Figure S4C). In contrast, oxidized HypR protein to the hypO promoter observed transcription of hypO was not increased by oxidation of in the DNase-I footprinting analyses, results in increased HypRC14S mutant protein in vitro and also not affected activation of hypO transcription in vitro. In addition, by oxidized HypRC14S protein that was treated subse- hypO specific transcription ratios decreased in the quently with DTT (Figure 5C and D; Supplementary 4186 Nucleic Acids Research, 2012, Vol. 40, No. 9

Figure S4C). These results show that oxidation of HypR disulphide linked to the Cys490-peptide (ICPSITQR) of increases activation of hypO transcription by the RNAP 914.4873 Da (Supplementary Figure S5A and S5B). in vitro. Detailed CID MS/MS analysis of this 4-fold charged peptide with an m/z = 620.315 confirmed that Cys14 is HypR senses diamide and NaOCl by a Cys14–Cys490 linked to Cys49’ in the HypR intersubunit- thiol-disulphide switch disulphide-linked dimer (Figure 6B, Supplementary To analyse the oxidative modification of HypR in vitro, Figure S6A, S6C, S6D and S6E). purified HypR-His-protein was treated with diamide and In addition, a differential Cys-alkylation approach by NaOCl and separated using non-reducing SDS–PAGE. IAM and NEM was used to trap the oxidized HypR is reversibly oxidized to intersubunit disulphides Cys-peptides in HypR (Supplementary Figure S5C–S5F). by diamide and NaOCl stress since it migrates at the Reduced and oxidized HypR proteins were first alkylated size of the HypR dimer upon oxidation (Figure 6A). In with IAM. The proteins were washed by acetone precipi- contrast, the majority of HypRC14S and HypRC49S tation, resolved in 8 M urea, followed by DTT reduction mutant proteins do not form intermolecular disulphides of oxidized thiols and alkylation with NEM. Reduced Cys upon oxidation. residues should then be IAM alkylated and oxidized Cys Using MALDI–TOF–MS, we analysed the residues NEM alkylated. The tryptic Cys14 and Cys49 Downloaded from Cys-containing tryptic peptides of reduced and oxidized peptides of reduced and oxidized HypR proteins were HypR proteins. In the mass spectrum of oxidized HypR analysed by MALDI–TOF–MS. The Cys49 peptide was the mass peak of 2478.0686 Da corresponds to the Cys14- alkylated predominantly with IAM in reduced HypR peptide (EGCPVEFTLDVIGGK) of 1563.7723 Da that is (mass peak 974.50 Da) and with NEM in oxidized HypR http://nar.oxfordjournals.org/

HypR HypRC14S HypRC49S A kDa M co Dia NaOCl co Dia NaOCl co Dia NaOCl 40 HypR- 35 Dimer 25 EGC14PVEFTLDVIGGK (1563.7723 Da)

15 HypR IC49PSITQR (+914.4873 Da) 10 at UB Greifswald on November 16, 2012

B

WT WT ∆hypR C Non-reduced kDa M co Dia co Dia Dia Reduced HypR- 35 HypR HypR HypR HypR Dimer 25 Co NaOCl Dia MHQ

HypR 15

-DTT +DTT D

Figure 6. HypR is oxidized to Cys14–Cys490 intersubunit disulphides in response to diamide and NaOCl in vitro (A and B) and in vivo (C and D). (A) His-HypR protein was treated with 1 mM diamide or 100 mM NaOCl, alkylated with 50 mM IAM and subjected to non-reducing SDS–PAGE analysis. The bands corresponding to the oxidized HypR disulphide dimer were tryptically digested and analysed using LTQ-Orbitrap mass spec- trometry. (B) The Cys14–Cys49’-intermolecular disulphide-containing peptide was observed as quadruply charged precursor ion at an m/z = 620.3151. The CID MS/MS spectrum of this Cys14–Cys490-disulphide peptide is shown with b and y ion fragment ions in red and blue. The detailed MS/MS data for this peptide including the Xcorr, ÁCn scores, precursor ion and neutral molecular masses of the peptide and the fragment ion seria are presented in Supplementary Figure S6. (C) 1D-western blot analysis and (D) 2D-diagonal western blot analysis of immunoprecipitated HypR protein purified from wild-type cells before (co) and after exposure to diamide and NaOCl in the presence of 50 mM IAM using HypR-specific polyclonal antibodies. The HypR-specific intersubunit disulphide is indicated by an arrow that was also visible in non-reducing SDS-gels and analysed using Orbitrap LC–MS/MS. The CID MS/MS spectrum, the Xcorr, ÁCn scores, precursor ion and neutral molecular masses of the peptide and fragment ion seria of this Cys14–Cys490-disulphide peptide of HypR purified from cells in vivo are presented in Supplementary Figure S6. Nucleic Acids Research, 2012, Vol. 40, No. 9 4187

(mass peak 1042.53 Da). The Cys14 peptide was alkylated charged amino acid residues in close vicinity of Cys14 ˚ mostly with NEM in oxidized HypR (mass peak that could lower its pKa value; Lys1010 is >10 A away 1688.87 Da). This differential IAM/NEM thiol-trapping and hydrogen bonded to Tyr330. Instead, the reactivity approach confirms that both Cys14 and Cys49 are revers- of the Cys14 thiolate is increased by helix a1 at whose ibly oxidized in HypR protein in vitro. N-terminal end Cys14 is located (49,50). The helix The oxidation of HypR was analysed in vivo by non- dipole lowers the pKa of Cys14 to allow at physiological reducing western blot analyses (Figure 6C) and the 2D pH values the formation of the reactive, nucleophilic non-reducing/reducing diagonal western blot analysis of thiolate susceptible to oxidation. immunoprecipitated HypR protein harvested from We used the reaction with 5,50-dithiobis-(2-nitrobenzoic NaOCl-, diamide- and MHQ-treated cells (Figure 6D). acid) (DTNB) to analyse the reactivity of both cysteines HypR migrates strongly at the right side of the diagonal at different pH values using His-tagged HypRC14S and under disulphide stress in the diagonal western blot His-HypRC49S mutant proteins (Figure 8). Spectroscopic (Figure 6D) indicating HypR oxidation to intersubunit measurement of the released 2-nitro-5 thiobenzoate allows disulphides in vivo. Detailed CID MS/MS analysis of the determination of the fraction of deprotonated thiol groups immunoprecipitated HypR dimer identified the Cys14– and thus the pKa value. The Cys14 thiol has a more acidic Cys490-disulphide-linked peptide in vivo (Supplementary pKa of 6.36 ± 0.04 (random error calculated from the Figure S6B, S6C, S6F and S6G). second fit) suggesting that Cys14 is present as thiolate Downloaded from anion and a preferred target for oxidation by diamide or Crystal structure of the reduced HypRC14S protein NaOCl (Figure 8B). In contrast, the pKa of Cys49 was To avoid artificial oxidation of HypR during crystalliza- determined as 8.51 ± 0.07 indicating that Cys49 is tion, HypRC14S protein was crystallized in the presence present in its protonated thiol form at physiological pH of b-mercaptoethanol. The structure was solved using mo- values in the cytoplasm (Figure 8A). The observed pKa http://nar.oxfordjournals.org/ lecular replacement and refined to 1.8 A˚ resolution. value of the thiol group of Cys49 is also in accordance Electron density is visible for residues 13–117 of the to its localization in a hydrophobic environment native 125 residues (Figure 7). The asymmetric unit provided by Ile52, Trp27, Ile30, Leu31, Gln60Cg and contains four similar monomers, organized in 2 biological Ile48. The sulphur atoms of Cys14 (presumed to be at dimers, 4 mercaptoethanol and 279 water molecules. The the position of Ser14Og) and Cys490 residues of the ˚ structure was refined to an R of 0.1935 and R of opposing subunits are 8–9 A apart in the HypR dimer cryst free ˚ 0.2336. The overall structure of reduced HypR comprises (9.0, 8.5, 8.0 and 8.3 A in monomers A, B, C and D, a homodimer adopting a triangular shape with respectively). at UB Greifswald on November 16, 2012 non-crystallographic 2-fold symmetry as found in all MarR-family proteins (48). The structure has the follow- Structural changes of HypR upon oxidation to ing secondary structure elements: a1(15–23)–a2(28–35)– intermolecular disulphides a3(42–48)–a4(54–66)–b2(70–75)–b3(81–86)–a5(88–114) For crystallization of oxidized HypR protein, His-HypR (Figure 7A and D). The two monomers associate via a protein was treated with 1 mM diamide for 15 min and the ˚ 2 large dimer interface of 1600 A provided by helices a1, formation of HypR intersubunit disulphides was verified a2, a5 and their symmetry mates a10, a20, a50. Helix a5 is by non-reducing SDS–PAGE. The structure of oxidized much longer than in the OhrR-family proteins exemplified HypR protein could be solved in P65 and P6522 using by SarZ of S. aureus and OhrR of X. campestris molecular replacement with reduced HypR as model, (Supplementary Figure S1B). Pro95 and Gly105 induce and taking twinning into account refined to 3.0 A˚ reso- kinks at positions 93 and 107 in HypR. The dimer inter- lution in the lower symmetry. Eight monomers are in face significantly differs from that in OhrR structures since the asymmetric unit forming four biological dimers. a6 is missing in HypR (Supplementary Figures S1B, S7A, Electron density is visible in all monomers only for S7C and S7D). residues 14–108 (Figure 7B). Both intermolecular Cys14– Each monomer contains a winged helix–turn–helix Cys490 and Cys140–Cys49 disulphide bonds are visible at (wHTH) DNA-binding motif constituted by b2, b3, a3 1.3 s in the 2Fo–Fc maps in three of four biological dimers and a4. Helix a4 is the recognition helix in the of the asymmetric unit (Figure 7G and H). To confirm DNA-binding domain and the anti-parallel b-sheet of b2 that there is useful signal in the data to 3.0 A˚ resolution, and b3 forms the wing. The HTH motif binds in the major we computed the correlation between observed and groove, the wing in the minor groove. The b1-strand, that calculated structure factor amplitudes in resolution is found in OhrR-like proteins between a2 and a3 shells, before the highest resolution data had been used (Supplementary Figure S1B) is not explicitely assigned in structure refinement, as described by Ling et al. (51) by DSSP, but the b2–b3 sheet forms two main chain (Supplementary Figure S8D). After refining the molecular hydrogen bonds to Lys40, a shortened version of b1. replacement solution against data limited to 3.3 A˚ reso- lution, there was a good correlation between observed The reactivity of the redox-sensing Cys14 and calculated structure factors to 3.0 A˚ resolution; in The redox-sensing Cys14 of HypR is located at the the 3.1–3.0 A˚ resolution shell, the correlation was 0.21 N-terminus of helix a1 (Figure 7E and F). The Cys14 and the R-factor was 0.45. thiolate is stabilized by hydrogen bonds with Val16N, Superimposition of the oxidized on the reduced Glu17N and a water molecule. There are no positive monomer shows a similar overall structure (r.m.s.d. on 4188 Nucleic Acids Research, 2012, Vol. 40, No. 9

A HypR reduced B HypR oxidized

NaOCl

HypRred and HypRox

C Downloaded from

α1 α2 α3 α4 1 10 20 30 40 50 60 MSEKKNIYPNKEGCPVEFTLDVIGGKWKGILFYHMIDGKKRFNEFRRICPSITQRMLTLQLRELEA D http://nar.oxfordjournals.org/

β2 β3 α5 70 80 90 100 110 120 DGIVHREVYHQVPPKVEYSLTEFGRTLEPIVLQMKEWGESNRDVLESYRSNGLVKDQQK at UB Greifswald on November 16, 2012 E G

F H

Figure 7. Structures of reduced HypRC14S (A) and oxidized HypR proteins (B), superimposition of reduced and oxidized HypR (C) and secondary structure assignments (D). (A) Reduced HypRC14S dimer with monomers C and D coloured in grey and blue, respectively. Residues Ser14 and Cys49 are shown in yellow as sticks and spheres (Og and Sg, respectively). (B) Oxidized HypR protein with the Cys14–Cys490 intersubunit disulphide with monomers C and D, labelled in grey and blue, respectively. The intermolecular disulphide bond is shown in yellow in monomer C. (C) Superimposition of the oxidized HypR dimer (light and dark blue) and the reduced HypR dimer (light and dark green). The HypRox/red side view is shown and Cys14 and Cys490 are shown as yellow sticks. One monomer of each dimer (on the right, dark blue and dark green) is aligned to visualize the differences in the opposing monomers. (D) The secondary structure elements of HypR are a1(15–23), a2(28–35), a3(42–48), a4(54–66), b2(70–75), b3(81–86), a5(88–114) that are shown as red tubes (a-helices) and green arrows (b-sheets). The redox-sensing Cys14 and the Cys49 are labelled in yellow. (E-H) The pocket of the redox-sensitive Cys residues in reduced and oxidized HypR. (E) Electron density map (2Fo–Fc at 1s in blue) and (F) key interactions between a1 and a30 helices including the Cys14Ser and Cys490 residues for reduced HypR. The atoms are coloured in yellow (carbon), dark blue (nitrogen), red (oxygen) and green (sulphur); water molecules are shown as red spheres and hydrogen bonds are labelled as dashed lines. Cys14Ser is shown in orange forming hydrogen bonds with Val16 and Glu17. Cys490 is in a hydrophobic environment formed by Ile520, Trp270, Ile300, Leu310, Gln60Cg0, Ile480. The distance between Ser14Og and Cys490Sg that form the intermolecular disulphide bond is 8.48 A˚ . (G) Electron density map (2Fo–Fc at 1s in blue) and (H) structure of the Cys14–Cys490 intersubunit disulphide region in oxidized HypR. Helices a30 and a40 on the right are oriented as in Figure 7EF, this emphasizes the movement of a4 on the left. Nucleic Acids Research, 2012, Vol. 40, No. 9 4189

A C 0.04 1 00350.035 Cys49 0.03 pH 6,03 0,1 0.025 pH 6,50 (nm) pH 7,03 ) -1

410 0.02 pH 7,51 0,01 (1/s) (s k A 0.015 pH 7, 71 k pH 8,07 0.01 pH 8,48 0,001 k 0.005 pH 8,21 k fitted 0 0,0001 020406080100 6 6,5 7 7,5 8 8,5 9 9,5 Time (sec) pH B D 0.045 1 Downloaded from 0.04 Cys14 0.035 pH 6,03 0.03 pH 6,28 ) -1 (nm) 0.025 pH 6,5

0,1 http://nar.oxfordjournals.org/

410 pH 6,76 k (s k 0.02 (1/s)k A pH 7,03 0.015 k pH 7,51 0.01 k fitted 0.005 pH 8,07 0,01 0 020406080 100 6 6,5 7 7,5 8 8,5 pH Time (sec) Figure 8. The pH dependence of the reactivity of Cys14 and Cys49 with DTNB. The absorbance at 410 nm at different pH values from 6 to 8 at UB Greifswald on November 16, 2012 resulting from the release of 2-nitro-5 thiobenzoate after reaction of Cys49 of the HypRC14S (A) and of Cys14 of the HypRC49S (B) mutant proteins with DTNB is plotted over the time. (C and D) The kinetic constants (k) of the Cys49 and Cys14 reactions with DTNB are plotted against the pH and fitted resulting in a pKa of 8.51 for Cys49 (C) and a pKa of 6.36 for Cys14 (D).

Cas: 0.9–1.1 A˚ ) (Figure 7A–C, Supplementary Figure crevices of the major groove of the DNA duplex under S8A, S8B and S8C). The most significant local difference oxidized conditions. This shift of the DNA-binding close to these cysteines is the change of the 1 angles of domains is observed in all crystallographically inde- Cys14 from 150 in the reduced to 80 in the oxidized pendent dimers, and visualized in the superimposed struc- form (averages for well determinedÀ cysteines). Cys14Sg tures of reduced and oxidized HypR (Supplementary does not make hydrogen bonds in the oxidized form. Figure S8). The spacing between the a4–a40 helices is The major changes of HypR upon oxidation are in the 30 A˚ in reduced HypR and 27 A˚ in oxidized HypR quaternary structure. If one of the monomers in the (measured at the helix axis between residues Leu61 and dimer is superimposed, the second monomer rotates 8 Arg62). Compared to the related two-Cys-type OhrRXc around an axis in the helix a5–a50 interface, the rotation regulator, HypR oxidation results in only a small hinge (Supplementary Figure S8). The dimer interface is domain domain movement of the DNA major groove rec- thus almost unchanged between helices a5 and a50, but is ognition a4 and a40 helices. compressed between helices a1–a2 and a10–a20. In the reduced form water molecules separate Gly25N and Gly250N; in the oxidized form these residues come close. DISCUSSION The DNA-binding domains move up to 4 A˚ towards each The role of the DUF24 family regulator HypR that other, loops a1–a2, a3, N-terminus of a4 and the wing controls the HypO nitroreductase formed of b2–b3 shift strongly. This is visualized in the change of Ca–Ca0 distances in the reduced compared to The MarR/DUF24 family is conserved among the oxidized conformation (Supplementary Figure S9). Gram-positive bacteria, and members of this family have Since HypR resembles a transcriptional activator, disul- been characterized in Corynebacterium glutamicum and phide bond formation should re-arrange the B. subtilis. QorR of C. glutamicum was characterized as DNA-binding helices a4 and a40 in the opposing a repressor that senses diamide and H2O2 and controls subunits of the dimer to enable binding to consecutive the quinone oxidoreductase QorA (52). Bacillus subtilis 4190 Nucleic Acids Research, 2012, Vol. 40, No. 9 encodes eight DUF24-family proteins (HypR, YodB, thiolate anion was confirmed using the DTNB assay, CatR, HxlR, YdeP, YdzF, YkvN and YtcD) and revealing a lower pKa value of 6.36 that is similar to previous studies revealed that YodB and CatR respond that of Cys15 in OhrRBs (with a pKa of 5.2). In both to diamide and quinones by thiol-based mechanisms OhrRBs and HypR structures, the N-terminal Cys is (7,24–26). Here, we have characterized the novel located at the N-terminus of helix a1 and the positive DUF24-type regulator HypR that is activated by hypo- macrodipole of helix a1 lowers its pKa value. Sequence chlorite and diamide stress by a thiol-disulphide switch. alignments indicate that Cys14 is conserved in DUF24 Recent studies showed that the OhrA peroxiredoxin is a paralogs and is also located at the N-terminus of helix specific determinant of NaOCl resistance (17). HypR posi- a1 in other DUF24 paralogs (Supplementary Figure S1). tively controls the putative nitroreductase HypO that It was shown previously that the conserved Cys6 and Cys7 provides together with OhrA protection against NaOCl residues in YodB and CatR are required for redox-sensing stress in B. subtilis. The nitroreductase HypO has been (24,25). The reactive thiolate anion in OhrR proteins is recently structurally characterized (47). Nitroreductases stabilized by a conserved tyrosine hydrogen-bonding are widely distributed flavoenzymes that catalyse the network (15,23,56). In reduced HypR the Cys14 thiolate NAD(P)H-dependent reduction of nitroaromatic and anion forms hydrogen bonds to Val16N and Glu17N. nitroheterocyclic compounds (53). These enzymes have Interestingly, Glu17 is conserved in DUF24 paralogs sug- broad substrate specificities and can reduce also gesting that a conserved hydrogen-bonding network Downloaded from azocompounds, quinones, flavins and metal ions (Fe3+ stabilizes the reactive Cys thiolate. 2– and CrO4 ). Previous results suggested that the nitroreductases YodC and MhqN could function as azo- Structural changes of HypR upon oxidation to and quinone reductases (27,36,43). However, they could intersubunit disulphides

be also involved in the thiol-specific stress response and http://nar.oxfordjournals.org/ function as flavoprotein disulphide reductases. Indeed, the MexR and OhrRXc sense oxidative stress by intermolecu- lar disulphide bond formation of two Cys residues of flavin-containing NAD(P)H oxidoreductase NfrA of ˚ Staphylococcus aureus showed weak disulphide reductase opposing subunits that are >15 A apart. With this large activity and was suggested to function as a thiol- distance, remarkable re-arrangements are required to disulphide oxidoreductase under disulphide stress (54). bring the Cys residues together for disulphide bond for- The homologous enzyme NfrA1 of B. subtilis exhibits mation. Upon oxidation, OhrRXc undergoes conform- NADH oxidase activity, catalysing the oxidation of ational changes in the a6–a60 helices of the dimer NADH to NAD+, and scavenges high concentrations of interface leading to rigid body rotation of the HTH H O (55). This suggests that HypO could function to motifs and dissociation of OhrRXc from the operator at UB Greifswald on November 16, 2012 2 2 DNA (15). Compared to the distance of 15.5 A˚ between restore the redox balance of the cells by direct detoxifica- ˚ tion of hypochlorite and diamide or by reduction of disul- Cys22 and Cys1270 in OhrRXc, the distance of 8–9 A phide bonds. between Cys14 and Cys490 in HypR is rather small. Thus, small structural changes are sufficient for disulphide bond formation and activation of HypR. Oxidation of the Conservation and structural features of the reactive Cys14 thiolate to sulphenylchloride by hypochlorite could Cys14 pocket of HypR abolish the negative charge on the sulphur and break the HypR represents a two-Cys-type MarR-type regulator hydrogen bonding network. Rotation of the Cys14 side that is activated by Cys14–Cys490 intersubunit disulphide chain about 1 can then bring the sulphur closer to formation as confirmed by mass spectrometry. The Cys490. The subsequent formation of the disulphide pulls DNaseI-footprinting analysis showed that oxidized helices a1 and a2 closer to their counterparts, helices a10 HypR bound with higher affinity to the hypO operator and a20. The move might be supported by the helix dipole DNA and using an in vitro transcription assay we of a1 attracting the Cys490 thiolate. At the dimer’s 2-fold provide evidence for transcriptional activation of RNAP axis Gly25N and Gly250N are bridged by a water molecule by oxidized HypR. The crystallographic analysis of and 6.5 A˚ apart in the reduced form, but only 4 A˚ apart reduced and oxidized HypR provides first structural with no water molecule in the oxidized form. The ionic insights into the reactive Cys14 pocket and the conform- interaction between Asp21Od and Lys260Ne is only ational changes of HypR upon oxidation that could be observed in the oxidized form (distance 3.5 A˚ ), but not required for specific interaction with the operator sites in the reduced form ( 9A˚ ). This interaction could tem- and/or transcription activation by RNAP. Examples for porarily stabilize the oxidized conformation until the di- two-Cys type MarR-family repressors that have been sulphide bond is formed. structurally characterized include MexR of Pseudomonas Disulphide bond formation moves the a4 and a40 helices ˚ aeruginosa and OhrRXc of Xanthomonas campestris of HypR 4Atowards each other causing minor changes (14,15,18,19). Interestingly, while OhrR-proteins and in the spacing of the DNA-binding domains. Similarly, DUF24-family members share only limited sequence simi- small conformational changes between the reduced and larity, the redox-sensing N-terminal Cys residues align oxidized conformations were observed for MexR, that is between OhrR and HypR (Supplementary Figure S1B). inactivated by oxidation to Cys30–Cys620 intersubunit di- Mutational analyses indicate that both Cys residues are sulphides (19). In oxidized MexR, neither the spacing of essential for redox-regulation of HypR consistent with the the major groove binding a4–a40 helices, nor the orienta- two-Cys-type redox-switch model. The reactive Cys14 tion of the dimerization domain was changed. Instead, Nucleic Acids Research, 2012, Vol. 40, No. 9 4191

oxidation causes rigid body rotation of the a20 and a30 FUNDING helices resulting in steric clashes of a20 and the disulphide Deutsche Forschungsgemeinschaft (AN 746/2-1 to H.A.); bond with the DNA backbone that lead to dissociation Wellcome Trust (UK) (grant 082961 to R.J.R.). Funding of the MexR repressor from the operator DNA (19). for open access charge: Deutsche Forschungsgemeinschaft In contrast to the MexR repressor, the DUF24-family (AN 746/2-1). regulator HypR is activated by disulphide bond formation and local domain movements in the wHTH region might Conflict of interest statement. None declared. be sufficient for recognition of the operator sites and recruitment of the RNAP to initiate transcription. We did not measure a significant change of the Kd values of REFERENCES reduced and oxidized HypR proteins in the gel-shift 1. Imlay,J.A. (2003) Pathways of oxidative damage. Annu. Rev. assays. Instead, we observed a different mobility of Microbiol., 57, 395–418. oxidized HypR in the gel-shift assays and higher affinity 2. Imlay,J.A. (2008) Cellular defenses against superoxide and of oxidized HypR to the operator DNA in the DNase-I hydrogen peroxide. Annu. Rev. Biochem., 77, 755–776. footprinting analysis. Oxidized HypR also resulted in 3. Davies,M.J. (2011) Myeloperoxidase-derived oxidation: mechanisms of biological damage and its prevention. J. Clin. 2-fold transcriptional induction of hypO in an in vitro-  Biochem. Nutr., 48, 8–19. Downloaded from transcription assay. Thus, the 4 A˚ movements of the a4 4. Hawkins,C.L., Pattison,D.I. and Davies,M.J. (2003) and a40 major groove recognition helices of HypR upon Hypochlorite-induced oxidation of amino acids, peptides and oxidation suggest a different DNA-binding mode of proteins. Amino Acids, 25, 259–274. 5. Marnett,L.J., Riggins,J.N. and West,J.D. (2003) Endogenous oxidized HypR leading to recruitment of the RNAP to generation of reactive oxidants and electrophiles and their initiate transcription at the hypO promoter. In addition, reactions with DNA and protein. J. Clin. Invest., 111, 583–593. the strong autoinduction of HypR itself by disulphide 6. Rudolph,T.K. and Freeman,B.A. (2009) Transduction of redox http://nar.oxfordjournals.org/ stress suggests that also increased amounts of oxidized signaling by electrophile-protein reactions. Sci. Signal, 2, re7. 7. Antelmann,H. and Helmann,J.D. (2011) Thiol-based redox HypR are required for transcriptional induction of the switches and gene regulation. Antioxid Redox. Signal, 14, hypO gene. We are aware that the structure of oxidized 1049–1063. HypR in the DNA-binding state needs to be resolved to 8. Zheng,M., Aslund,F. and Storz,G. (1998) Activation of the OxyR shed light into the specific interaction of oxidized HypR transcription factor by reversible disulfide bond formation. Science, 279, 1718–1721. with the operator DNA and into the mechanism of tran- 9. Lee,C., Lee,S.M., Mukhopadhyay,P., Kim,S.J., Lee,S.C., scriptional activation. Thus, our ongoing studies are Ahn,W.S., Yu,M.H., Storz,G. and Ryu,S.E. (2004) Redox regulation of OxyR requires specific disulfide bond formation

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! ! Bui Khanh Chi, Dirk Albrecht, Katrin Gronau, Dörte Becher, Michael Hecker, and Haike Antelmann*.

!

Proteomics, 10: 3155 – 3164 (2010).

* corresponding author

! "#! Proteomics 2010, 10,3155–3164 DOI 10.1002/pmic.201000230 3155

RESEARCH ARTICLE The redox-sensing regulator YodB senses quinones and diamide via a thiol-disulfide switch in Bacillus subtilis

Bui Khanh Chi, Dirk Albrecht, Katrin Gronau, Do¨rte Becher, Michael Hecker and Haike Antelmann

Institute for Microbiology, Ernst-Moritz-Arndt-University of Greifswald, Greifswald, Germany

The MarR/DUF24-type repressor YodB controls the azoreductase AzoR1, the nitroreductase Received: April 8, 2010 YodC and the redox-sensing regulator Spx in response to quinones and diamide in Bacillus Revised: May 15, 2010 subtilis. Previously, we showed using a yodBCys6-Ala mutant that the conserved Cys6 Accepted: June 14, 2010 apparently contributes to the DNA-binding activity of YodB in vivo. Here, we present data that mutation of Cys6 to Ser led to a form of the protein that was reduced in redox-sensing in response to diamide and 2-methylhydroquinone (MHQ) in vivo. DNA-binding experiments indicate that YodB is regulated by a reversible thiol-modification in response to diamide and MHQ in vitro. Redox-regulation of YodB involves Cys6-Cys101’ intermolecular disulfide formation by diamide and quinones in vitro. Diagonal Western blot analyses confirm the formation of intersubunit disulfides in YodB in vivo that require the conserved Cys6 and either of the C-terminal Cys101’ or Cys108’ residues. This study reveals a thiol-disulfide switch model of redox-regulation for the YodB repressor to sense electrophilic compounds in vivo.

Keywords: Disulfide bond formation / Electrophiles / Microbiology / Redox-sensing / YodB

1 Introduction proteins lead to changes in the protein structure which convert these regulators into transcriptional activators or Redox-sensing transcription factors respond to changes in inactivate repressors from DNA binding. the cellular redox status caused by reactive oxygen, nitrogen Reactive electrophilic species are redox-active com- or electrophilic species. Central for most redox-sensing pounds that include, for example quinones, aldehydes proteins are nucleophilic cysteine residues that sense and diamide. Quinones are endogenously produced elec- directly the reactive species via post-translational modifica- trophiles that are part of the respiratory chain. In tions. These thiol modifications include, in most cases, addition, these are present in the soil in humic substances reversible disulfide bond formation, such as inter- or intra- [6, 7]. Quinones and diamide react with thiol-containing molecular disulfides and mixed disulfides with low mole- proteins via different mechanisms. As shown in the cular weight thiols (S-thiolations) [1]. In addition, the thiol thiol-redox proteome and using MS, diamide causes group of cysteine can be irreversibly oxidized to sulfinic or reversible thiol modifications, including inter- and intra- sulfonic acids or thiol-(S)-alkylated by electrophilic quinones molecular disulfides and S-cysteinylated proteins [8–10]. and aldehydes [2–5]. Thiol modifications of redox-sensing Quinones have two modes of action, an oxidative and electrophilic mode. Quinones can be reduced incompletely Correspondence: Dr. Haike Antelmann, Institute for Micro- to reactive semi-quinone anions that in turn reduce biology, Emst-Moritz-Arndt-University of Greifswald, F.-L.-Jahn- molecular oxygen to ROS such as superoxide anions Street 15, D-17487 Greifswald, Germany [11–13]. The production of ROS could lead in turn to E-mail: [email protected] reversible thiol modifications. As electrophiles, quinones Fax: 149-3834-864202 form S-adducts with cellular thiols via the irreversible Abbreviations: BSH, bacillithiol; IAM, iodoacetamide; mhq, 2- thiol-(S)-alkylation reaction. Our previous proteome- methylhydroquinone wide studies showed that toxic quinones react with thiol-

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com 3156 B. K. Chi et al. Proteomics 2010, 10, 3155–3164 containing proteins predominantly via the thiol-(S)-alkyla- disulfide between the N-terminal Cys22 of one subunit and tion chemistry leading to protein aggregation and depletion the C-terminal Cys1270 of the other subunit in vitro [28]. from the proteome [14]. However, using thiol-redox YodB has a conserved Cys6 residue and two additional proteomic studies, we found the glyceraldehyde-3-phosphate Cys residues at positions 101 and 108. The Cys6-Ala muta- dehydrogenase (GapA) as target for reversible thiol modifi- tion affected DNA-binding activity of YodB in vivo and cations by the oxidative mode of quinones. Thus, quinones in vitro [17]. We further suggested that Cys6 and the act via both the oxidative and the S-alkylation chemistry C-terminal Cys-residues of YodB are S-alkylated by quinones in vivo with the majority of thiol-containing proteins at toxic in vitro. However, the detailed mechanism of YodB-regula- concentrations. tion by diamide and quinones is unknown and the possi- The depletion of the cellular thiol redox buffer by bility of inter-subunit disulfide formation has been not diamide and quinones causes a general electrophile stress investigated in the previous studies. In this article, we show response in Bacillus subtilis which includes expression of that YodB resembles a two-Cys-type redox-sensing MarR- regulons controlled by Spx, CtsR, PerR and CymR [15–21]. type regulator and is regulated via inter-subunit disulfide B. subtilis is able to develop a specific adaptive stress formation in response to diamide and quinones in vitro and response to quinone-like electrophiles and diamide that is in vivo. Thus, YodB-regulation by diamide and quinones is mediated by the MarR-type repressors, MhqR and YodB. similar to that of the two-Cys-type OhrR repressor of YodB controls most strongly the azoreductase AzoR1 which X. campestris [28]. confers resistance to diamide and quinones [17, 21]. MhqR regulates the expression of the paralogous azoreductase AzoR2, several oxidoreductases, and thiol-dependent dioxy- 2 Materials and methods genases (MhqA, MhqE and MhqO) [17, 20, 21]. The quinone and azoreductases function in reduction of the electrophiles 2.1 Bacterial strains and growth conditions and the thiol-dependent dioxygenases are involved in the specific ring cleavage of the quinones. The bacterial strains used were B. subtilis 168 (trpC2), DyodB The MarR/DUF24-family repressor YodB is a redox- (trpC2,yodB::Cmr), and the yodBC6A, yodBC6S, yodBC101A, sensing repressor that directly senses and responds to yodBC108A and yodBC101,108A point mutants which quinone-like electrophiles and diamide [17]. The mechan- are described (Table 1). B. subtilis strains were cultivated ism of redox-sensing for MarR-type regulators has been under vigorous agitation at 371C in Belitsky minimal thoroughly studied for the OhrR repressor in B. subtilis and medium described previously [33]. Escherichia coli Xanthomonas species [22–30]. OhrR senses and responds to strains were grown in LB for DNA manipulation. The organic hydroperoxides and controls the thiol-dependent antibiotics were used at the following concentrations: 1 mg/ peroxidase OhrA. The OhrR proteins can be divided into the mL erythromycin and 5 mg/mL chloramphenicol. The one-Cys-type and two-Cys-type family [27]. The OhrR compounds used were 2-methylhydroquinone (MHQ, protein of B. subtilis has one reactive Cys15 and is the Acros) and diamide (Sigma). prototype of the one-Cys-type subfamily. Cumene hydro- peroxide leads to Cys15 oxidation to sulfenate which is unstable and reacts further to form S-thiolated OhrR 2.2 Construction of the yodBC6S point mutant containing cysteine or bacillithiol (BSH) in vivo [26, 31]. The OhrR protein of Xanthomonas campestris belongs to the two- Plasmid pDGyodBC6S was produced by using PCR muta- Cys-type family that is inactivated by an inter-subunit genesis. For construction of pDGyodBC6S, first-round PCR

Table 1. Bacillus subtilis strains and plasmids used in this study

Strain or plasmid Genotype

Strains B. subtilis 168 trpC2 yodBC6S trpC2 yodB::cat thrC::yodBC6S This study yodBC6A trpC2 yodB::cat thrC::yodBC6A This study and [17] yodBC101A trpC2 yodB::cat thrC::yodBC101A This study and [17] yodBC108A trpC2 yodB::cat thrC::yodBC108A This study and [17] yodBC101,108A trpC2 yodB::cat thrC::yodBC101,108A This study and [17] yodBC6,101,108A trpC2 yodB::cat thrC::yodBC6,101,108A This study and [17] Plasmids pDG795 [32] pDGyodBC6S yodBC6S in pDG795 This study pML54 YodB with N-terminal His-tag in pPROEX [18]

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Table 2. Oligonucleotide primers used in this study extensively dialyzed against 10 mM Tris-HCl (pH 8.0), 100 mM NaCl, 50% glycerol v/v and stored at 801C. Primer Sequence 50–30 À yodBC6S-for1 CGGAATTCGAGATCGGGGCATTTGTATCA yodBC6S-rev1 TTAGGGGACATCGTATTTCCCAT 2.5 DNA gel mobility shift assays yodBC6S-for2 TACGATGTCCCCTAAAATGGAAT yodBC6S-rev2 CGGGATCCGTGCCGCTTTCCTTATTTCTC DNA fragments containing the azoR1 promoter region azoR-forFT TCGCAGACGTATGAAATCTTG were generated by PCR using the primer sets azoR1-for azoR-revFT AGCTAAGAAAGCTTCGTAAAGT and azoR1-rev (Table 2). Approximately, 600 pmol of the purified PCR product was end labelled using T4 poly- was performed in two separate reactions using primers nucleotide kinase (Roche Diagnostics) and 50 mCi [g32P]- yodBC6Sfor1 with yodBC6Srev1 and primers yodBC6Sfor2 ATP (GE Healthcare). The labelled azoR1-specific promoter with yodBC6Srev2 (Table 2) and B. subtilis 168 chromosomal probe was purified by ammonium sulphate/ethanol preci- DNA as template. The PCR products were hybridized and pitation and 2000 cpm was incubated with different subsequently amplified by a second round of PCR using amounts of purified YodB proteins for 10 min at room primers yodBC6Sfor1 and yodBC6Srev2. The PCR products temperature in EMSA-binding buffer (10 mM Tris-HCl from the second round PCRs were then digested with EcoRI pH 7.5, 100 mM KCl, 5% glycerol v/v) in the presence and BamHI restriction enzymes and inserted into plasmid of 50 mg/mL of BSA and 5 mg/mL of Salmon sperm DNA. pDG795 digested with the same enzymes to generate The concentrations of the compounds used for the DNA- pDGyodBC6S. The plasmid pDGyodBC6S was verified by binding assays were 1 mM MHQ, 1 mM diamide and 1 mM DNA sequencing and introduced into the thrC locus of the DTT. DNA-binding reactions were separated by 4% native B. subtilisDyodB mutant, respectively, by transformation polyacrylamide gel electrophoresis in 10 mM Tris, 1 mM with selection for erythomycin–lincomycin resistance. The EDTA buffer, pH 8 containing 2.5% glycerol at room yodBC6A, yodBC101A, yodBC108A, yodBC101,108A and temperature and constant voltage (250V) for 15 min. Gels yodBC6,101,108A mutants were constructed by transforma- were dried and the radiolabelled bands were visualized tion of chromosomal DNA of strains ORB6599, ORB6715, using phosphoimaging. ORB6716, ORB6777 and ORB7083 [17], respectively, into B. subtilis 168. All strains were verified by PCR using yodB- specific primers and the PCR products were verified by DNA 2.6 Western blot analysis sequencing for correct point mutations. Anti-YodB polyclonal rabbit antiserums were generated using purified His-tagged YodB protein. The antisera were 2.3 Northern blot experiments purified using affinity chromatography with NHS-activated sepharose and the His-tagged YodB-proteins to avoid non- Northern blot analyses were performed as described [34] specific cross-reaction of the antibodies with B. subtilis using RNA isolated from the B. subtilis strains before proteins. The polyclonal antisera were used for immuno- (control) and 10 min after the treatment with 1 mM diamide precipitation experiments and the purified antibodies or 0.5 mM MHQ, respectively. Hybridizations specific for were used at a dilution of 1:200 in the Western blot azoR1 and bsrB were performed with the digoxigenin- experiments. GapA-specific antibodies were used for the labeled RNA probes synthesized in vitro using T7 RNA control Western blot experiments at a dilution of 1:1000. polymerase from T7 promoter containing internal PCR Protein amounts of 25 mg were loaded onto a 14% SDS- products of the respective genes using the primer sets as PAGE gel and the Western blot analysis was performed as described [17]. described previously [14].

2.4 Expression and purification of His-YodB protein 2.7 Immunoprecipitation and nonreducing/ reducing diagonal SDS-PAGE analysis E. coli BL21(DE3)pLysS (Invitrogen) was used for over- production of His-tagged YodB protein. For expression of B. subtilis wild-type cells were treated with 1 mM diamide the YodB protein, plasmid pML54 was used and purification and 1 mM MHQ and cells were harvested in the presence of of His-tagged YodB protein was performed as described 50 mM iodoacetamide (IAM) to alkylate all reduced thiols. previously [17]. Recombinant His-YodB protein that Cells were sonicated and the protein extracts obtained after contains the N-terminal His-tag was purified using repeated centrifugation. Immunoprecipitation of YodB PrepEaseTM His-Tagged High Yield purification Resin using YodB-specific antibodies and protein extracts was (USB) under native conditions according to the instructions performed with Dynabeads-ProteinA (Invitrogen) according of the manufacturer (USB). The purified YodB protein was to the instructions of the manufacturer. The precipitated

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com 3158 B. K. Chi et al. Proteomics 2010, 10, 3155–3164 proteins were eluted by boiling in non-reducing SDS sample one missed cleavage site of trypsin were allowed. The search buffer (4% SDS; 62.5 mM Tris-HCl, pH 8.0, glycerol). result was filtered using BioWorks 3.2 (Thermo Electron). A Immunoprecipitated YodB protein was separated using multiple threshold filter applied at the peptide level consis- the non-reducing/reducing diagonal SDS-PAGE analysis as ted of the following criteria: (i) peptide sequence length: described previously [10]. Proteins were first separated using 7–30 amino acids; (ii) RSp r4; (iii) percentage of ions: 70 non-reducing SDS-PAGE without DTT and the lanes were (70% of all theoretical b- and y-ions of a peptide are cut. The gel lanes were incubated in SDS sample buffer experimentally found); (iv) Xcorr versus charge state: 1.90 for containing 50 mM DTT to reduce all oxidized thiols and singly charged ions, 2.20 for doubly charged ions, and 3.75 subsequently in SDS sample buffer with 100 mM IAM to for triply charged ions. alkylate all newly formed reduced thiols. Bands were posi- tioned horizontally on an SDS-polyacrylamide gel and separated using reducing SDS-PAGE and subjected to 3 Results Western blot analysis. All proteins that migrate along the diagonal represent proteins without disulfide bonds. Those 3.1 Sequence comparison of YodB to MarR/DUF24 proteins that form intermolecular disulfides between two family members of Firmicutes proteins or two subunits run at the right side to the diagonal and proteins with intra-molecular disulfides migrate left to YodB was previously characterized as a redox-sensing the diagonal. repressor of the MarR/DUF24 family which is involved in regulation of spx, yodC and azoR1 transcription [17]. MarR/ DUF24-like regulators are conserved among low GC Gram- 2.8 MS analysis positive bacteria. They are widely distributed among the taxon Firmicutes including other Bacillus species, such as The disulfide-linked YodB dimer was digested with Trypsin B. cereus, B. anthracis, B. halodurans or B. licheniformis as well and AspN manually using the protocol as described for the as species of the genus Listeria, Staphylococcus, Enterococci or Ettan Spot handling workstation. The resulting peptides Clostridia (Fig. 1). Moreover, the genome of B. subtilis were spotted onto MALDI-targets (Voyager DE-STR, encodes at least eight MarR/DUF24 family regulators most PerSeptive Biosystems) and measured using a Proteome- of which have unknown functions (YodB, CatR, HxlR, YdeP, Analyzer 4800 (Applied Biosystems, Foster City, CA, USA). YdzF, YkvN, YtcD and YybR) which are listed in the DBTBS LC-ESI MS experiments were performed using an Orbitrap database for transcriptional regulation of B. subtilis (http:// mass spectrometer (Thermo Electron, San Jose, CA). The dbtbs.hgc.jp/) (Supporting Information Fig. S1). Most of SEQUEST algorithm Version 27.12 was used to search for these share one conserved Cys residue in the N-terminal modifications of the YodB peptides. A mass deviation of part and one or two Cys residues in the C-terminal part at 0.01 Da for precursor ions as well as for fragment ions and less conserved positions. Moreover, in some MarR/DUF24-

Figure 1. ClustalW format alignment of MarR/DUF24-family regulators in Gram-positive bacteria. The alignments of examples for YodB orthologs of B. subtilis (Bsub_yodB), B. amyloliquefaciens (RBAM_019300), B. licheniformis (BL01441), B. pumilus (BPUM_1875), Clostridium botulinum (CLB_0900), B. cereus (BCZK4889), B. halodurans (BH508), Listeria monocytogenes (lmo1220), Enterobacter faecalis (EF1114) and Staphylococcus aureus (SA1925). Identical amino acid residues are shown by stars and conserved amino acid substitutions are marked by two dots. The Cys residues are highlighted in black including the conserved N-terminal Cys residue.

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com Proteomics 2010, 10,3155–3164 3159 like regulators, the conserved N-terminal Cys is preceded by this mutation has affected either the structure or the func- conserved Met or Leu and Pro residues. tion of YodB [17]. Since the C6A mutant repressor retains inducer responsiveness (Figs. 2A and B), Cys6 is not abso- lutely required for sensing electrophiles. Thus, we investi- 3.2 The conserved Cys6 residue is required for gated the role of the conserved Cys6 of YodB again in regulation of YodB in vivo regulation of azoR1 transcription in a yodBC6S mutant in vivo. The basal level of azoR1 transcription was twofold We have previously shown that the conserved Cys6 of YodB increased in the yodBC6S mutant (Figs. 2A and B). This is required for DNA-binding activity of YodB in vivo and suggests that the YodBC6S mutant protein showed in vitro as analyzed in a yodBC6A mutant [17]. However, the decreased DNA-binding activity to the azoR1 promoter yodBC6A mutant showed derepression of azoR1 transcrip- under control conditions in vivo. However, in contrast to the tion in vivo under control conditions which suggests that yodBC6A mutant, the yodBC6S mutant showed no response to diamide in vivo. The response of the yodBC6S mutant was also decreased by MHQ compared with the wild-type A protein, indicating that Cys6 is important for quinone- WT ∆yodB yodBC6S yodBC6A sensing. However, we observed still a twofold derepression Co D M co D M co D M co D M of azoR1 transcription in the yodBC6S mutant to quinones,

azoR1 suggesting additional roles of other amino acid residues in sensing of quinone compounds in vivo. Control Northern blot analysis was performed for bsrB 23S rRNA 16S rRNA which is located downstream of azoR1 and strongly tran- scribed independently of YodB [17]. The bsrB gene is similar transcribed in the yodBC6S mutant and in the wild type B (Supporting Information Fig. S2). Western blot analyses 14 confirmed that the YodBC6S mutant protein is produced at 12 similar protein levels as YodB in the wild type in response to 10 diamide stress (Fig. 2C). We noticed that the YodB protein 8 6 amount is increased in the wild type after diamide treatment 4 transcription ratios transcription which is consistent with the result that yodB expression is 2 subject to negative autoregulatory control [21]. Control

azoR1 Western blots were performed using GapA-specific anti- _D _D _D _M _M _M _co _co _co A S 6A 6S dB 6S dB bodies confirming that similar protein amounts of GapA are 6A dB W WT_D W WT_M WT_co W yod yod yod ∆

∆ produced in the wild type, yodB, yodBC6S and yodBC6A ∆ yodBC6 yodBC6 yodBC6 yodBC6 yodBC6 yodBC6A mutant strains. Furthermore, the Coomassie-stained image of the SDS gel is shown to confirm that equal protein C WT ∆yodB yodBC6S amounts were applied to each lane (Supporting Information kDa Co Dia Co Dia Co Dia Fig. S3). 15 YodB To address the question if the C-terminal Cys residues are involved in YodB-regulation by MHQ in vivo, transcription of Figure 2. The conserved N-terminal Cys6 residue of YodB is azoR1 was analyzed in a triple yodBC6,101,108S mutant. required for redox-sensing in vivo. (A) Northern blot analysis of However, this triple yodBC6,101,108S mutant showed full azoR1 transcription was performed using RNA isolated from derepression of azoR1 transcription under control conditions B. subtilis wild type, yodB, yodBC6S and yodBC6A point mutants indicating that mutation of all three Cys residues leads to a before (co) and 10min after treatment with 0.5mM MHQ (M) and non-functional repressor (data not shown). 1mM diamide (D). The arrow points toward the size of the azoR1 specific transcript. Northern blots were stained with methylene blue before detection to verify that equal amounts of RNA were separated in each lane as indicated by the abundant 16S and 23S 3.3 DNA-binding activity of YodB is reduced by rRNAs. (B) The quantification of azoR1 transcription ratios was diamide and quinones in vitro which is performed from the Northern blot experiments using ImageJ. reversible with DTT The ratios represent middle values of azoR1 transcription rations calculated from three independent experiments. (C) Western blot Previously, we have shown using DNase-I footprinting analysis shows that similar amount of YodB mutant protein is experiments that DNA-binding activity of YodB to the azoR1 produced in the yodBC6S point mutant as in the wild type in promoter DNA is inhibited after treatment with diamide response to diamide stress. Cells were harvested before and 15 min after 1 mM diamide stress and 25 mg of protein extracts and quinones in vitro [17]. Gel mobility shift assays were was applied and subjected to YodB-specific Western blot performed to analyze the change and reversibility in the analysis. DNA-binding affinity of YodB by quinones and diamide

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com 3160 B. K. Chi et al. Proteomics 2010, 10, 3155–3164 in vitro. Under control conditions, YodB bound with a diamide and MHQ, alkylated with IAM and separated using calculated Kd of 1.3 mM to the azoR1 promoter (Fig. 3; non-reducing SDS-PAGE (Figs. 4A and B). YodB forms Supporting Information Fig. S4). DNA-binding activity of DTT-reversible inter-subunit disulfides in response to YodB to the azoR1 promoter was two to fourfold reduced diamide and MHQ stress in vitro. Thus, YodB is oxidized to in the presence of diamide and quinones in vitro. In addi- the disulfide-linked YodB-dimer by both electrophiles. tion, this change in DNA-binding activity of YodB is rever- However, we noticed two closely migrating bands of dimeric sible using reducing agents (DTT) consistent with the YodB protein in response to MHQ treatment which could be hypothesis that a reversible disulfide bond formation is both reduced to the monomeric YodB protein (Figs. 4A and involved in regulation of YodB. B). Using MS, we analyzed the Cys-containing peptides of YodB which are cross-linked in the inter-subunit disulfides. The bands corresponding to the YodB monomer and the 3.4 YodB responds to diamide and quinones via C6- disulfide-linked YodB dimer were digested with Trypsin and C101’ intersubunit disulfide formation in vitro AspN and the resulting peptides measured using MALDI- TOF MS (Figs. 4C and D). In the mass spectrum of the To study the mechanism of diamide and quinone-sensing disulfide-linked YodB dimer, we detected a mass peak by YodB in vitro, purified YodB-protein was treated with of 3260.7878 Da that corresponds to the Cys6-containing T1 peptide (DIPTTENLYFQGAHMGNTMCPK) of 2468.1043 Da that is linked to the Cys101’-containing 0 0.75 1.5 3 6 12 24 M YodB µ peptide DQFCEPG of 794.2978 Da via a disulfide bond indicated by the loss of 2 Da ( 2H1). Detailed MS/MS control À analysis of the double-charged peptide of 1631.2 Da using Orbitrap MS confirmed that the Cys6 peptide is linked to the Cys101’ containing peptide of 794.3 Da (Supporting Infor- mation Fig. S5). Furthermore, we did not detect this 3260.7878 Da mass peak in the reduced YodB monomer + Diamide (Fig. 4C). This 3260.7878 Da mass peak appeared in the spectrum of both close migrating YodB-dimers which are formed after MHQ stress. This indicates that the higher and lower mobility bands represent YodB inter-subunit disulfides that are linked by one and two disulfide bonds between Cys6 and Cys101’ of the opposite subunits. The + Diamide + DTT fact that MHQ causes the formation of inter-subunit disul- fides in YodB indicates that quinones act via the oxidative mode in vitro. We did not found a possible Cys6-Cys108 inter-subunit disulfide with the theoretical mass of 3417.4021 Da containing the C108 peptide DTVCEEEK of 951.3928 Da in the MALDI-TOF spectrum of the YodB dimer (data not shown). This indicates that Cys6-Cys101’ + MHQ disulfide bond formation is the in vitro mechanism for YodB oxidation.

3.5 Diagonal Western blot analysis showed inter- subunit disulfides for YodB by diamide and + MHQ + DTT quinones in vivo

We analyzed the formation of YodB inter-subunit disulfides in vivo using Diagonal Western blot analysis of immuno- precipitated YodB protein that was enriched from wild-type cells and yodBC6S, yodBC101A, yodBC108A and Figure 3. DNA-binding activity of YodB is inhibited by quinones yodBC101,108A mutant cells treated with diamide and and diamide in vitro and can be restored by DTT. EMSAs were MHQ (Fig. 4E). Two close YodB spots were detected at the used to analyze the effect of 1 mM diamide and 1 mM MHQ in the right side of the diagonal under control conditions. Both of absence and presence of 1 mM DTT on the DNA-binding activity of purified YodB protein to the labelled azoR1 promoter probe. these YodB spots at the right side of the diagonal increased The YodB protein amounts used for the DNA-binding reactions strongly in diamide and MHQ-treated wild-type cells. This are indicated. reveals that intermolecular disulfides are formed in vivo in

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A BC100 Dia MHQ 90 co 19.9 80 70 3282.7039 -DTT + DTT -DTT + DTT 60 50 40 kDa co 1 2 5 1 2 mM co 1 2 5 1 2 mM 30

% Intensity 20 35 10 3235.0 3263.2 3291.4 25 D D100 49.8 90 Dia 80 15 70 3282.8091 98.7500 2.7151 3260.7878

C 60 76.7642

50 329 327 40 3292 30 % Intensity 20 10 3237.0 3263.4 3291.8 Mass (m/z)

DIPTTENLYFQGAHMGNTMC PK E Non-Reducing Redu DQFC EPG YodB YodB YodB ∆yodB co Dia MHQ Dia ucing

YodBC6S YodBC101,108A YodBC101A YodBC108A Dia Dia Dia Dia

Figure 4. YodB is regulated by intersubunit disulfide bond formation by diamide and quinones. (A–D) Purified YodB protein was treated with 1–5mM diamide (A) or MHQ (B) and subsequently either reduced with 10mM DTT (1DTT) or left oxidized ( DTT), alkylated with À 50mM IAM and subjected to non-reducing SDS-PAGE analysis. The bands corresponding to the YodB monomer (C) and disulfide-linked YodB dimer (D) were cut, digested with AspN and Trypsin and analyzed by MALDI-TOF MS. The Cys6-Cys101’-containing disulfide-linked peptide that contains parts of the N-terminal fused His-tag was detected as 3260.8 Da mass peak. The MS/MS analysis of this 3260.8 Da mass peak was performed using Orbitrap MS (Supporting Information Fig. S5). (E) Diagonal non-reducing/reducing YodB-specific Western blot analysis of immunoprecipitated YodB protein that was harvested from untreated wild-type cells (co) and from theyodB, yodBC6S, yodBC101A, yodBC108A, yodBC101,108A mutant cells exposed to diamide (Dia) and MHQ in the presence of 50 mM IAM.

YodB in response to diamide and MHQ which is in agree- could be incompletely oxidized YodB linked only by one ment with the in vitro studies. However, the inter-subunit disulfide bond in the YodB dimer. The original positions disulfide form of YodB is more abundant in diamide-treated of both oxidized YodB proteins in the non-reducing cells than after MHQ stress. This suggests that part of YodB SDS-PAGE are visible also at the middle of the diagonal protein could respond in vivo to MHQ by the irreversible at the size of the YodB dimer as shifted to the left side S-alkylation in addition to oxidation which needs to be of the diagonal (left, completely oxidized YodB) and along shown in future studies. The spots representing the the diagonal (right, incompletely oxidized YodB). Both monomeric and inter-subunit disulfides of YodB are not bands of the YodB dimer were also detected in the non- present in diamide-treated yodB mutant cells indicating that reducing SDS-PAGE and Western blot analysis of wild- these are specific for YodB. YodB inter-subunit disulfides type proteins, but were not present in protein extracts of are formed in diamide-treated C101A and C108A mutants, yodB mutant cells confirming that these are specific for but not in the C6S and C101,108A point mutants. Thus, YodB (data not shown). This indicates that the immuno- Cys6 and one of the C-terminal Cys-residues are important precipitated YodB disulfide forms cannot be completely for disulfide bond formation in vivo. This seems to contrast reduced after equilibration of the SDS-gel strip in the the results from the in vitro modification where only the reducing sample buffer before separation using the second C6-C101’ disulfide was detected by MS analysis. It could be SDS-PAGE. possible that the reaction rate of C6 with C108’ is slower We further detected many cross-reacting proteins along than that of C6 with C101’. This indicates that in the wild- the diagonal using the polyclonal YodB-antibodies in the type Cys101 will capture C6 to form disulfides and in its Diagonal Western blot analyses. These result from the absence the C108’ residue will fill the role of C101. YodB-immunoprecipitates separated in the Diagonal-SDS- The nature of these two close YodB spots at the right PAGE that include YodB and the purified YodB-antibodies. side of the diagonal might be explained as follows: The left Thus, the antibodies themselves used for the immunopre- YodB spot could represent the YodB-dimer that is linked cipitation are recognized by the YodB-antibodies used for by two inter-subunit disulfide bonds and the right spot the detection of the Western blot analyses. These antibodies

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com 3162 B. K. Chi et al. Proteomics 2010, 10, 3155–3164 are visible as cross-reacting proteins along the diagonal and Reduced YodB : SH HS at the right side in the upper molecular range in both the SHHS azoR1 wild-type and yodB mutant cells. 108C C108 yodC C C101 -35 101 -10 spx

4 Discussion C6 C6 SH HS 4.1 YodB senses diamide and quinone electrophiles via a thiol-disulfide switch in B. subtilis Diamide Quinone The MarR-type repressors YodB and MhqR control para- logous azoreductases, nitroreductases, dioxygenases and oxidoreductases that confer resistance to diamide and Oxidized YodB : quinone-like electrophiles in B. subtilis [17, 20, 21]. The SH HS SH HS C C paralogous azoreductases and nitroreductases functions as C108 C108 108 108 C C C quinone reductases to reduce toxic quinones to more redox- 101 C101 101 S S 101 S S or stable hydroquinones. In addition, these oxidoreductases S S S S azoR1 function in reduction of the electrophilic azocompound C6 C6 C6 C6 yodC diamide. The paralogous thiol-dependent dioxygenases are spx involved in ring-cleavage of quinone-like electrophiles. -35 -10 Thus, the common functions of the YodB and MhqR regulons are to protect cells against depletion of low mole- Figure 5. Redox-sensing mechanism of the YodB repressor in cular weight thiols and protein thiols by diamide and response to diamide and quinones in B. subtilis.YodBcontrols quinone-like electrophiles. Spx, the nitroreductase YodC and the azoreductase AzoR1. The To react rapidly to those electrophilic compounds, the conserved Cys6 of YodB is required for redox-sensing in vivo. MarR/DUF24 family repressor YodB directly senses thiol- YodB is oxidized by diamide and quinones to Cys6-Cys101’ inter- reactive electrophiles via the conserved Cys6 residue. The subunit disulfides in vitro and to Cys6-Cys101’ or Cys6-Cys108’ inter-subunit disulfides in vivo. YodB oxidation leads to dere- results presented in this article have shown that this Cys6 is pression of transcription of the YodB regulon genes that function required for redox-sensing in vivo. Inducer responsiveness in detoxification of the electrophiles. of the YodBC6S mutant protein was reduced after diamide and quinone stress compared with wild-type YodB. In agreement to the previous studies, this conserved Cys is regulation involves Cys6-Cys101’ inter-subunit disulfide required for DNA-binding activity as derepression is formation in response to diamide in vitro. Using the Diag- observed in both the yodBC6S and the yodBC6A mutants onal Western blot analysis, we identified YodB inter-subunit under control conditions. This mutational analysis suggests disulfides in vivo that involve Cys6 and either of the that the Cys6 residue is important for repressor activity of C-terminal Cys101’ or Cys108’ residues. Thus, the redox- YodB and might interfere with the overall protein confor- sensing mechanism of YodB is similar to that of the two- mation. However, there remains still the question how the Cys-type OhrR repressor of X. campestris. Quinones lead to remaining response of the yodBC6S mutant to quinones can oxidation of YodB via Cys6-Cys101’ inter-subunit disulfide be explained. This question is difficult to address due to the formation in vitro and YodB inter-subunit disulfides were fact that mutation of all three Cys residues (Cys6, 101 and detected in vivo indicating that quinones act, at least in part, 108) either to Ala or Ser results in a non-functional via the oxidative mode. Previously, we suggested that YodB repressor that makes it difficult to investigate the involve- is regulated via S-alkylation by quinones in vitro as we ment of other amino acids in quinone-sensing. We have detected YodB peptides that were S-conjugated by catechol further made exchanges in the amino acids surrounding and MHQ [17]. These alkylated YodB species were detected Cys6 and found that the conserved Pro7 is also required for only after treatment of the YodB protein with quinones DNA-binding activity of YodB (unpublished data). Thus, in vitro and represent modifications that do not affect the current attempts are underway to resolve the structure of regulation of YodB in vitro. This is supported by the finding YodB under reduced and oxidized conditions in the that DNA-binding activity of YodB is regulated by a rever- presence of diamide and quinones that should answer these sible disulfide bond formation excluding the irreversible questions. thiol-(S)-alkylation chemistry as main mechanism. In this study, we have analyzed the mechanism that is Furthermore, the non-reducing PAGE analysis shows used by the YodB repressor to sense diamide and quinones. clearly the formation of reversible inter-subunit disulfides in YodB represents a two-Cys type redox-sensing repressor of YodB by quinones in vitro. Our previous data showed that B. subtilis that is inactivated by a thiol-disulfide switch by toxic quinones act on most cytoplasmic proteins in vivo diamide and quinones in vitro and in vivo (Fig. 5). YodB- mainly via the irreversible thiol-(S)-alkylation mode, causing

& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com Proteomics 2010, 10,3155–3164 3163 depletion of thiol-containing proteins in the proteome [14]. uncharacterized thiol-disulfide oxidoreductases are phylo- It was also revealed that the GapA protein responds to toxic genetically linked to the BSH synthesis enzymes [38]. quinones by disulfide bond formation. However, non-toxic The disulfide reductases that keep BSH in a reduced state concentrations of quinones did not deplete thiol-containing could facilitate reduction of oxidized redox-sensing regula- proteins and might act via the oxidative mode in vivo to tors, such as OhrR, YodB and other MarR/DUF24 family modify redox-sensing proteins. The question arises if the regulators in response to oxidative and electrophile stress signal that is sensed by YodB is ROS instead of electrophilic that is subject of future studies. quinones and diamide. However, according to all the published transcriptome and proteome data, ROS, such as The authors are grateful to John Helmann, Shawn MacLel- H2O2 and paraquat do not lead to induction of the YodB and lan and Ahmed Gaballa for discussion of the results prior to MhqR regulons in vivo [21]. Thus, the YodB sensor is publication. The authors thank especially Peter Zuber and specific for quinones and diamide but not to oxidative stress Montira Leelakriangsak for providing yodB point mutant strains in vivo. and plasmid pML54. The authors further thank Jo¨rg Stulke. for providing GapA antibodies. This work was supported by grants from the Deutsche Forschungsgemeinschaft, the Bundesminis- 4.2 Functions of other MarR/DUF24-type regulators terium fur. Bildung und Forschung (BACELL-SysMo 031397A), in B. subtilis and other Gram-positive bacteria the Fonds der Chemischen Industrie, the Bildungsministerium of the country Mecklenburg-Vorpommern and BACELL-BaSysBio B. subtilis encodes for eight paralogous MarR/DUF24-family (LSHG-CT-2006-037469) to M. H. and by a grant from the regulators (YodB, CatR, HxlR, YdeP, YdzF, YkvN, YtcD and Deutsche Forschungsgemeinschaft AN 746/2-1 to H. A. YybR). The first identified MarR/DUF24-family regulator of B. subtilis was HxlR which is an activator of the formalde- The authors have declared no conflict of interest. hyde-inducible hxlAB operon that encodes the genes for the ribulose monophosphate pathway [5, 35]. Recently, we have shown that reactive aldehydes (formaldehyde and methyl- 5 References gloxal) act as thiol-reactive electrophiles by depletion of the thiol-redox buffer cysteine [5]. Aldehydes cause a general [1] Paget, M. S., Buttner, M. 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Antelmann, unpublished). Neurosci. 2007, 30, 37–45. Recently, the MarR/DUF24-family regulator CgR1435 [5] Nguyen, T. T., Eiamphungporn, W., Mader,. U., Liebeke, M. (QorR) has been characterized as redox-sensing repressor of et al., Genome-wide responses to carbonyl electrophiles in a quinone oxidoreductase in Corynebacterium glutamicum Bacillus subtilis: control of the thiol-dependent formalde- [37]. DNA-binding activity of QorA was inhibited by diamide hyde dehydrogenase AdhA and cysteine proteinase YraA by and H2O2 in vitro. Inactivation of QorR involves inter- the MerR-family regulator YraB (AdhR). Mol. Microbiol. subunit disulfide formation between the conserved single 2009, 71,876–894. Cys17 residues of both subunits. Our ongoing studies are [6] Steinberg, C. E., Meinelt, T., Timofeyev, M. A., Bittner, M., directed to identify the functions and regulatory mechan- Menzel, R., Humic substances. Part 2: interactions with isms of those uncharacterized MarR/DUF24 family regula- organisms. Environ. Sci. Pollut. Res. Int. 2008, 15, 128–135. tors in B. subtilis and we have evidence that these collectively [7] Ratasuk, N., Nanny, M. A., Characterization and quantifica- function as specific sensors for aldehydes, quinones and tion of reversible redox sites in humic substances. Environ. other electrophiles in Gram-positive bacteria. Sci. Technol. 2007, 41, 7844–7850. Since YodB-regulation involves a reversible thiol-disulfide [8] Hochgrafe,. F., Mostertz, J., Albrecht, D., Hecker, M., Fluores- switch in B. subtilis, it will be interesting to identify the cence thiol modification assay: oxidatively modified proteins enzymatic pathways that function in reduction of oxidized in Bacillus subtilis. Mol. Microbiol. 2005, 58, 409–425. YodB in vivo. Recently, BSH, an abundant thiol-redox [9] Hochgrafe,. F., Mostertz, J., Po¨ ther, D. C., Becher, D. et al., buffer in Firmicutes was structurally characterized [31]. S-cysteinylation is a general mechanism for thiol protection The biosynthesis pathway for BSH has parallels to the of Bacillus subtilis proteins after oxidative stress. J. Biol. mycothiol biosynthesis in Actinomycetes and several Chem. 2007, 282, 25981–25985.

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& 2010 WILEY-VCH Verlag GmbH & Co. KGaA, Weinheim www.proteomics-journal.com !

! "#$! ! ! "#$%&'(!)! ! !

! ! ! ! ! ! ! ! ! ! ! Chapter 6 ! ! ! ! ! ! ! The paralogous MarR/DUF24-family repressors YodB and CatR control expression of the catechol dioxygenase CatE in Bacillus subtilis.

! Bui Khanh Chi, Kazuo Kobayashi, Dirk Albrecht, Michael Hecker, and Haike Antelmann*.

! ! Journal of Bacteriology, 192: 4571 – 4581 (2010).

! ! ! ! ! ! ! ! ! ! ! * corresponding author

! "#$!

! JOURNAL OF BACTERIOLOGY, Sept. 2010, p. 4571–4581 Vol. 192, No. 18 0021-9193/10/$12.00 doi:10.1128/JB.00409-10 Copyright © 2010, American Society for Microbiology. All Rights Reserved.

The Paralogous MarR/DUF24-Family Repressors YodB and CatR Control Expression of the Catechol Dioxygenase ᰔ

CatE in Bacillus subtilis † Downloaded from Bui Khanh Chi,1 Kazuo Kobayashi,2 Dirk Albrecht,1 Michael Hecker,1 and Haike Antelmann1* Institute for Microbiology, Ernst-Moritz-Arndt-University of Greifswald, F.-L.-Jahn-Str. 15, D-17487 Greifswald, Germany,1 and Graduate School of Information Science, Nara Institute of Science & Technology, Takayama 8916-5, Ikoma, Nara 630-0192, Japan2

Received 9 April 2010/Accepted 2 July 2010

The redox-sensing MarR/DUF24-type repressor YodB controls expression of the azoreductase AzoR1 and the http://jb.asm.org/ nitroreductase YodC that are involved in detoxification of quinones and diamide in Bacillus subtilis. In the present paper, we identified YodB and its paralog YvaP (CatR) as repressors of the yfiDE (catDE) operon encoding a catechol-2,3-dioxygenase that also contributes to quinone resistance. Inactivation of both CatR and YodB is required for full derepression of catDE transcription. DNA-binding assays and promoter mutagenesis

studies showed that CatR protects two inverted repeats with the consensus sequence TTAC-N5-GTAA over- lapping the ؊35 promoter region (BS1) and the transcriptional start site (TSS) (BS2). The BS1 operator was required for binding of YodB in vitro. CatR and YodB share the conserved N-terminal Cys residue, which is

required for redox sensing of CatR in vivo as shown by Cys-to-Ser mutagenesis. Our data suggest that CatR is on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS modified by intermolecular disulfide formation in response to diamide and quinones in vitro and in vivo. Redox regulation of CatR occurs independently of YodB, and no protein interaction was detected between CatR and YodB in vivo using protein cross-linking and mass spectrometry.

Bacillus subtilis is exposed in the soil to a variety of antimi- to aggregation and depletion of proteins in the proteome (18). crobial agents that include phenolic and quinone-like com- However, we also found the glyceraldehyde-3-phosphate de- pounds which are produced by other soil bacteria, plants, or hydrogenase GapA as a target for reversible thiol oxidation by fungi. In addition, quinone-like compounds are present in hu- quinones in vivo. Thus, quinones act via the oxidative and mic substances of the soil. Quinones are also biologically active electrophilic mode in B. subtilis cells in vivo. compounds that function as lipid electron carriers in the elec- The depletion of the thiol redox buffer by diamide and tron transport chain (e.g., ubiquinone and menadione). Thus, quinones causes a general electrophile stress response in B. quinones are naturally occurring electrophilic compounds that subtilis, which includes expression of regulons controlled by the are ubiquitously distributed in bacterial systems. Spx, CtsR, PerR, and CymR regulators and the MarR-type Proteomic and transcriptomic approaches revealed that cat- repressors MhqR and YodB (1, 4, 15, 16, 21, 22, 31). YodB is echol and methylhydroquinone (MHQ) act like diamide as a MarR/DUF24-family repressor that controls the azoreduc- thiol-reactive electrophiles in B. subtilis. Quinones and diamide tase AzoR1, the nitroreductase YodC, and the redox-sensing deplete the cellular pool of low-molecular-weight (LMW) thi- regulator Spx in response to diamide and quinones. YodB ols via different mechanisms. Diamide leads to an increase in senses quinones and diamide via the conserved Cys6 residue, reversible thiol modifications, such as inter- and intramolecu- which is required for DNA-binding activity and redox sensing lar disulfides and S thiolations (disulfides between proteins and of YodB in vivo (15). In recent studies, we have shown that LMW thiols) (9, 10, 27). Quinones can act as oxidants or YodB functions as a two-Cys-type redox-sensing regulator that electrophiles (13, 20, 25). As oxidants, quinones redox cycle is regulated via intersubunit disulfides between Cys6 and one with their semiquinones, producing reactive oxygen species of the C-terminal Cys residues, Cys101 or Cys108, in response (ROS), such as superoxide anion or hydrogen peroxide (18). to diamide and quinones in vivo (4). Thus, YodB regulation is The production of ROS could lead in turn to reversible thiol similar to that of the two-Cys-type OhrR repressor of Xan- modifications. As electrophiles, quinones can form S adducts thomonas campestris (26). with cellular thiols via the thiol-(S)-alkylation chemistry. Re- Together, the MarR-type repressors YodB and MhqR con- cently, we showed that toxic quinones react mainly via S-alkyl- trol paralogous azoreductases (AzoR1 and AzoR2), nitrore- ation with cellular thiol-containing proteins in vivo, which leads ductases (YodC and MhqN), and thiol-dependent dioxygen- ases (MhqA, MhqE, and MhqO) (1, 15, 31). The paralogous azoreductases are major quinone resistance determinants and * Corresponding author. Mailing address: Institute for Microbiol- have functions as quinone reductases, thus preventing toxic ogy, Ernst-Moritz-Arndt-University of Greifswald, F.-L.-Jahn-Str. 15, quinones from undergoing redox cycling and causing depletion D-17487 Greifswald, Germany. Phone: 49-3834-864237. Fax: 49-3834- of cellular thiols. The paralogous thiol-dependent dioxygen- 864202. E-mail: [email protected]. † Supplemental material for this article may be found at http://jb ases (MhqA, MhqE, and MhqO) could be involved in the .asm.org/. specific ring cleavage of quinone S adducts which are formed in ᰔ Published ahead of print on 16 July 2010. the detoxification reaction of LMW thiols with quinones (31).

4571 4572 CHI ET AL. J. BACTERIOL.

TABLE 1. Bacillus subtilis strains and plasmids used in this study in the regulation of this catDE operon and found that catDE is

Strain, mutation, or Reference negatively controlled by YodB and its paralogous repressor Genotype plasmid or source YvaP (CatR). We further show in this paper that redox regu- lation of catDE expression is mediated by CatR and YodB, Strains B. subtilis 168 trpC2 which are modified by intersubunit disulfide bond formation in ⌬catE trpC2 catE::pMutin4 30 vivo to sense electrophiles.

⌬catR trpC2 catR::cat This study Downloaded from ⌬catR trpC2 catR::spc This study ⌬yodB trpC2 yodB::cat 15 MATERIALS AND METHODS ⌬yodB trpC2 yodB::em 15 Bacterial strains and growth conditions. The bacterial strains used were B. ⌬yodB ⌬catR trpC2 yodB::cat catR::spc This study r ⌬yodB ⌬azoR1 trpC2 yodB::em azoR1::cm 15 subtilis 168 (trpC2) with the following mutations: ⌬catR (trpC2 catR::Cm ), ⌬catR (trpC2 catR::Spcr), ⌬catR ⌬yodB (trpC2 catR::Spcr yodB::Cmr), ⌬yodB (trpC2 catRC7S trpC2 catR::cat thrC::catRC7S This study r r ⌬catR catR-FLAG trpC2 catR::cat This study yodB::Cm ), ⌬yodB (trpC2 yodB::pMutin4, Em ), ⌬yodB ⌬azoR1 (trpC2 azoR1::Cmr yodB::pMutin4, Emr), ⌬catR catR-FLAG (trpC2 catR::Spcr amyE::pSWEETcatR-FLAG r ⌬yodB catR-FLAG trpC2 yodB::em This study amyE::pSWEETcatR-FLAG, Cm ), ⌬yodB catR-FLAG (trpC2 yodB::pMutin4, Emr amyE::pSWEETcatR, Cmr), and the catRC7S point mutant, which is de- amyE::pSWEETcatR-FLAG http://jb.asm.org/ scribed below (Table 1). B. subtilis strains were cultivated under vigorous agita- Plasmids tion conditions at 37°C in Belitsky minimal medium as described previously (29). pSWEET 3 Escherichia coli strains were grown in LB broth for DNA manipulation. The pDG795 7 antibiotics were used at the following concentrations: 3 ␮g/ml erythromycin, 25 pET11b Novagen ␮g lincomycin, 100 ␮g spectinomycin, and 5 ␮g/ml chloramphenicol. The com- pDGcatR catR in pDG795 This study pounds used were 2-methylhydroquinone (Acros), catechol, and diamide pDGcatRC7S catRC7S in pDG795 This study (Sigma). The alkylating agent 4-acetamido-4Ј-maleimidylstilbene-2,2Ј-disulfonic pSWEETcatR-FLAG catR with C-terminal FLAG This study acid (AMS) was purchased from Molecular Probes, Dynabeads protein A was purchased from Invitrogen, and the cross-linking reagent DSP [dithiobis(succin-

tag in pSWEET on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS imidylpropionate)] was purchased from Pierce. pETcatR catR with N-terminal His6 tag This study in pET11b The ⌬catE mutant strain that harbors the catE-lacZ fusion was constructed pML54 YodB with N-terminal His tag 16 using the pMutin4 plasmid as described previously (30). The transcription factor in p-PROEX (TF) deletion mutants were constructed using an overlap-extension PCR tech- nique as described previously (12). The cat gene was amplified from the plasmid pCBB31 by PCR analysis with primers pUC-F and pUC-R. Upstream and down- Our previous studies identified the yfiDE (catDE) operon that stream regions of each regulator gene were amplified by PCR using gene-specific primer sets F1/R1 and F2/R2 as described previously (12). The primer sets is induced by catechol and MHQ and which encodes an en- catR-F1/catR-R1 and catR-F2/catR-R2 were used for amplification of the catR zyme with catechol-2,3-dioxygenase activity (CatE) and a upstream and downstream regions (Table 2). The 5Ј ends of primers R1 and F2 DoxX-like oxidoreductase, CatD (24, 30). We were interested are complementary to pUC-R and pUC-F sequences, respectively. Then, the

TABLE 2. Oligonucleotide primers used in this study Primer Sequence (5Ј–3Ј)a catRC7S-for1...... CGGAATTCTCCTCATAATCCCCTGTTTCTCT catRC7S-rev1 ...... TCTAGGAGACATTTCTGATTGGTT catRC7S-for2...... TCAGAAATGTCTCCTAGATTTGAA catRC7S-rev2 ...... CGGGATCCAGAACCATTTTAGCAATTAGTC catD-forFT ...... CTTATATGAAAGCCAATTTGAG catD-revFT ...... GAAACTTTGATAAACCGTGAAC catD-PE2 ...... AAGATAATACCTGTAATAACTCT catR-F1 ...... TGGCAATATCTCAATAAAGAAG catR-R1...... GTTATCCGCTCACAATTCAGGACACATTTCTGATTGGTT catR-F2 ...... CGTCGTGACTGGGAAAACTTAGGTGAGATTTCTAAATGGG catR-R2...... GCTGCCTTCTTGATTTAAGTC pUC-F ...... GTTTTCCCAGTCACGACG pUC-R ...... GAATTGTGAGCGGATAAC catR-NdeI-pETfor ...... ATATAGAGCATATGCACCACCACCACCACCACATGAACCAATCAGAAATGTGTC catR-BamHI-rev ...... ATAGGATCCTTAGTCAAGAAAGGAAGGGTCAAT T7-promoter-primer ...... TTAATACGACTCACTATAGG T7-terminal-primer...... GCTAGTTATTGCTCAGCGG catD_Ϫ146_for...... CTTATATGAAAGCCAATTTGAG catD_Ϫ80_for...... TTATAGTACTTTTATTTCATT catD_Ϫ60_for...... TTTCGTAAAATGTAACAAAAT catD_Ϫ40_for...... TTACTTGACGTAAACAAGTTA catD_Ϫ60M3_for...... TTTCGTAAAATGTAACAAAAGGGCTTGACGGGGACAAGTT catD_ϩ20_rev ...... AATGTTATCACTTACTTAAT catD_ϩ20M1_rev ...... AATGTTATCACGGGCTTAATGTAAGCTAGTATA catD_ϩ20M2_rev ...... AATGTTATCACGGGCTTAATGGGGGCTAGTATA catD_ϩ1_rev ...... TGTAAGCTAGTATATATACAT catR_FLAGfor...... GTGTTAATTAACGTTATTTGCGTTATAATAG catR_FLAGrev ...... ATAGGATCCTTATTATTTATCATCATCATCTTTATAATCGTCAAGAAAGGAAGGGTCAA

a Underlining indicates mutations in the inverted repeats in the catD promoter region. VOL. 192, 2010 REGULATION OF catDE BY YodB AND CatR (YvaP) 4573 three PCR fragments were mixed and used as a template for a second PCR His-YodB proteins were purified using PrepEase His-tagged high-yield purifica- analysis with primers F1 and R2. The resultant PCR fragments were used for tion resin (USB) under native conditions according to the instructions of the transformation of B. subtilis 168 to generate the ⌬catR::Cm mutant strain. Plas- manufacturer (USB). The purified His-CatR and His-YodB proteins were ex- mid pCm::spec was used to replace the Cmr cassette with a spectinomycin tensively dialyzed against 10 mM Tris-HCl (pH 8.0), 100 mM NaCl, and 50% resistance marker to generate the ⌬catR::Spcr strain. Chromosomal DNA of the glycerol and stored at Ϫ80°C. ⌬catR::Spcr strain was transformed into the ⌬yodB mutant to generate the ⌬yodB DNase I footprinting analysis. Primers catD-forFT and catD-revFT were each ⌬catR double mutant strain. For construction of plasmid pSWEETcatR-FLAG, 5Ј-end labeled using T4 polynucleotide kinase (Roche Diagnostics) and 50 ␮Ci primers catR_FLAGfor and catR_FLAGrev were used to amplify catR, includ- [␥-32P]ATP (GE Healthcare) and purified using ethanol precipitation. DNA Downloaded from ing the ribosomal binding site and a C-terminal FLAG tag sequence, followed by probes for catE (corresponding to positions Ϫ146 to ϩ120 relative to TSS) were two stop codons. The PCR product was digested with PacI and HindIII and synthesized by PCR amplification using one 5Ј-end-labeled primer pair (catD- cloned into the plasmid pSWEET (3) downstream of the xylA promoter that was forFT and catD-revFT) and the corresponding nonlabeled primer pair, respec- digested with the same enzymes. The pSWEETcatR-FLAG plasmid was inte- tively (Table 2). Purified PCR fragments that were labeled at one 5Ј end were grated into the amyE gene of the B. subtilis ⌬catR::spc mutant strain with used as sequencing templates. DNA probe purification and DNase I footprinting selection for chloramphenicol and spectinomycin, resulting in the ⌬catR catR- were performed as described previously (31). FLAG strain. Plasmid pSWEETcatR-FLAG was also integrated into the B. DNA gel mobility shift assays. The DNA fragment containing the catD pro- subtilis ⌬yodB::Em mutant strain that resulted in the ⌬yodB catR-FLAG strain. moter region was generated by PCR using the primer set catD-forFT and catD- Construction of the catRC7S point mutant. To amplify the catR gene including revFT. For catDE promoter mutagenesis and deletion of the upstream CatR the flanking regions, PCR analysis was performed with primers catR-EcoRI and binding region, primers were used to amplify the following regions: from Ϫ146 http://jb.asm.org/ catR-BamHI (Table 2) using chromosomal DNA from B. subtilis 168 as template. to ϩ20 (primers catD_Ϫ146_for and catD_ϩ20_rev); from Ϫ146 to ϩ20, con- The PCR product was digested with EcoRI and BamHI restriction enzymes and taining mutations in the BS2 operator (primers catD_Ϫ146_for and inserted into pDG795 (7), which was digested with the same enzymes to generate catD_ϩ20M1_rev or catD_ϩ20M2_rev); from Ϫ80 to ϩ20 (primers catD_ pDGcatR. The catR sequence was verified by DNA sequencing. Ϫ80_for and catD_ϩ20_rev); from Ϫ60 to ϩ20 (primers catD_Ϫ60_for and Plasmid pDGcatRC7S was produced by using PCR mutagenesis. First-round catD_ϩ20_rev); from Ϫ60 to ϩ20, containing mutations in the BS1 operator PCR analysis was performed using primers catRC7Sfor1 and catRC7Srev1 and (primers catD_Ϫ60M3_for and catD_ϩ20_rev); and from Ϫ40 to ϩ20, relative primers catRC7Sfor2 and catRC7Srev2 (Table 2). The PCR products were to the transcriptional start site (primers catD_Ϫ40_for and catD_ϩ20_rev) (Ta- hybridized and subsequently amplified by a second round of PCR using primers ble 2). Approximately 600 pmol of the purified PCR products was end labeled catRC7Sfor1 and catRC7Srev2. The PCR products from the second-round PCRs using T4 polynucleotide kinase (Roche Diagnostics) and 50 ␮Ci [␥-32P]ATP (GE on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS were then digested with EcoRI and BamHI restriction enzymes and inserted into Healthcare). The labeled catD promoter probes were purified by ammonium plasmid pDG795 digested with the same enzymes to generate plasmid sulfate-ethanol precipitation, and 2,000 cpm of each probe was incubated with pDGcatRC7S. The plasmid pDGcatRC7S was verified by DNA sequencing and different amounts of purified His-CatR and His-YodB proteins for 10 min at introduced into the thrC locus of the B. subtilis ⌬catR mutant by transformation room temperature in electrophoretic gel mobility shift assay (EMSA) binding with selection for chloramphenicol and erythromycin-lincomycin resistance. buffer (10 mM Tris-HCl [pH 7.5], 100 mM KCl, 5% glycerol) in the presence of Transcription factor array analysis. Transcription factor array analysis was 50 ␮g/ml bovine serum albumin (BSA) and 5 ␮g/ml salmon sperm DNA. The performed as described previously (8, 12) using competent cells of the concentrations of the compounds used for the DNA-binding assays were 1 mM ⌬catE::pMutin4 strain which carries a catE-lacZ fusion. MHQ, 1 mM diamide, 10 mM dithiothreitol (DTT), and 5 mM catechol. DNA- Transcriptome analysis. For microarray analysis, the B. subtilis wild type and binding reactions were separated by 4% native polyacrylamide gel electrophore- ⌬catR mutant strain were grown in minimal medium and harvested at an optical sis in 10 mM Tris, 1 mM EDTA buffer, pH 8, containing 2.5% glycerol at room density at 500 nm (OD500) of 0.4. Total RNA was isolated by the acid phenol temperature and constant voltage (250 V) for 15 min. Gels were dried, and the method as described previously (19). Generation of fluorescence-labeled cDNA radiolabeled bands were visualized using phosphorimaging. and hybridization with B. subtilis whole-genome microarrays (Eurogentec) was Western blot analysis. Anti-CatR polyclonal rabbit antiserum was generated performed as described previously (11). Two independent hybridization experi- using purified His-tagged CatR protein. The antiserum was purified using affinity ments were performed using RNAs from two independent experiments. All chromatography with N-hydroxysuccinimide (NHS)-activated Sepharose and the microarray datasets are available in the GEO database under the accession coupled His-CatR protein to avoid nonspecific cross-reaction of the antibodies number GSE22603. with B. subtilis proteins. The CatR antibodies were eluted from the affinity Northern blot experiments. Northern blot analyses were performed as de- column using 100 mM glycine-HCl, pH 3.5, followed by neutralization with 1 M scribed previously (32) using RNA isolated from the B. subtilis strains before Tris-HCl, pH 8.0. The polyclonal antisera were used for immunoprecipitation (control) and 10 min after the treatment with 1 mM diamide, 2.4 mM catechol, experiments, and the purified antibodies were used at a dilution of 1:200 in the or 1 mM MHQ, respectively. Hybridizations were performed with the digoxige- Western blot experiments. Protein amounts of 25 ␮g were loaded onto a 14% nin-labeled catE-specific RNA probe synthesized in vitro using T7 RNA poly- SDS-PAGE gel, and the Western blot analysis was performed as described merase as described previously (30). previously (18). Primer extension experiments. Primer catD-revFT complementary to the N- Immunoprecipitation and nonreducing/reducing diagonal SDS-PAGE analy- terminal encoding region of catD (catD-PE2) was 5Ј-end labeled using T4 sis. B. subtilis wild-type cells were treated with 1 mM diamide and 1 mM MHQ, polynucleotide kinase (Roche Diagnostics) and 50 ␮Ci [␥-32P]ATP (GE Health- and cells were harvested in the presence of 50 mM iodoacetamide (IAM) to care). Primer extension analysis was performed using the labeled primer as alkylate all reduced thiols. Cells were sonicated, and the protein extracts were described previously (32). Sequencing of the corresponding promoter regions obtained after repeated centrifugation. Immunoprecipitation of CatR protein was performed as described previously (28) using PCR products as templates using CatR-specific antibodies and B. subtilis protein extracts was performed with containing the catDE promoter region amplified with primers catD-forFT and Dynabeads protein A (Invitrogen) according to the instructions of the manufac- catD-revFT (Table 2). turer. The precipitated proteins were eluted using nonreducing SDS sample Expression and purification of the His-CatR and His-YodB proteins. Esche- buffer (4% SDS, 62.5 mM Tris-HCl [pH 8.0], glycerol). Protein extracts or richia coli BL21(DE3)pLysS (Invitrogen) was used for overproduction of N- precipitated CatR protein was separated using the nonreducing/reducing diag- terminal His-tagged CatR and YodB proteins. For expression of the YodB onal SDS-PAGE analysis as described previously (17). Proteins were first sepa- protein, plasmid pML54 was used and purification of His-tagged YodB protein rated using nonreducing SDS-PAGE without DTT and the lanes were cut. The was performed as described previously (15). For expression of the His-CatR cut lanes were first incubated in SDS sample buffer containing 50 mM DTT to protein, a PCR product was amplified using primers catR-NdeI-pETfor and reduce all oxidized thiols and subsequently incubated in SDS sample buffer with catR-BamHI-pETrev (Table 2). The forward primer includes a start codon 100 mM IAM to alkylate all newly formed reduced thiols. Bands were positioned followed by the codons for six N-terminal histidine residues. The catR-specific horizontally on an SDS-PAGE gel, separated using reducing SDS-PAGE, and PCR product was digested with the NdeI and BamHI restriction enzymes and subjected to Western blot analysis. All proteins that migrate along the diagonal inserted into the expression plasmid pET11b (Invitrogen) that was digested with represent proteins without disulfide bonds. Those proteins that form intermo- the same enzymes to generate pETcatR. The catR sequence was verified by DNA lecular disulfides between two proteins or two subunits run at the right side of the sequencing using the primers T7-promoter-primer and T7-terminal-primer (Ta- diagonal, and proteins with intramolecular disulfides migrate to the left of the ble 2). E. coli BL21(DE3)pLysS carrying pETcatR and pML54 were cultured in diagonal. 1 liter LB medium, and 1 mM IPTG (isopropyl-␤-D-thiogalactopyranoside) was Analysis of protein interactions of YodB and CatR in vivo. Cells of strains added at the mid-log phase (OD600 of 0.8) for 2 h. Recombinant His-CatR and ⌬catR catR-FLAG and ⌬yodB catR-FLAG were grown in 1 liter Belitsky minimal 4574 CHI ET AL. J. BACTERIOL. Downloaded from http://jb.asm.org/ on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS

FIG. 1. (A) Transcription factor arrays identify CatR as a repressor of the catDE operon. The individual colonies are transformants of transcription factor deletion mutants carrying the catE-lacZ fusion which were plated on X-Gal plates. The deletion of the regulatory gene is indicated. (B to E) Dual control of catDE transcription by the paralogous MarR/DUF24-family repressors CatR and YodB. Northern blot analysis of catDE transcription was performed using RNA isolated from the B. subtilis wild type (WT) and ⌬catR, ⌬yodB, ⌬catR ⌬yodB, and ⌬yodB ⌬azoR1 mutants before (co) and 10 min after treatment with 1 mM MHQ (M), 2 mM MHQ (M2), 2.5 mM MHQ (M3), 2.4 mM catechol (cat), 12 mM catechol (cat2), 1 mM diamide (D), or 2 mM diamide (D2). The arrow points toward the size of the catDE-specific transcript. (E) The quantification of catDE transcription ratios was performed using ImageJ from three independent Northern blot experiments. (F) Primer extension analysis was performed to determine the 5Ј end of the catDE-specific transcript that is marked by an arrow, and the Ϫ10 promoter sequence is labeled with a solid line. For sequencing, the dideoxynucleotide added in each reaction is indicated above the corresponding lane.

medium to an OD500 of 0.2. Production of CatR-FLAG protein was induced by operon, we have applied the transcription factor/transforma- adding 2% xylose to the culture for 90 min. Cells were harvested by centrifuga- tion array technology (TF array) (8, 12). Chromosomal DNA tion, washed twice with reaction buffer (100 mM sodium phosphate [pH 7.5], 150 mM NaCl), and dissolved in 100 ml reaction buffer each. The cross-linking of each of the 287 transcription factor deletion mutants was reagent DSP [dithiobis(succinimidylpropionate)] (Pierce) was added to the cul- transformed into the ⌬catE mutant that carries a catE-lacZ ture at a concentration of 400 ␮M for 30 min. Cells were harvested by centrif- fusion due to integration of the nonreplicating pMutin4 plas- ugation, washed twice with 20 mM Tris-HCl, pH 8.0, and disrupted by sonication. ⌬yvaP CatR-FLAG protein was immunoprecipitated using anti-FLAG M2 affinity aga- mid. The deletion mutant showed derepression of the rose (Sigma) according to the instructions of the manufacturer. CatR-FLAG catE-lacZ fusion on LB plates containing X-Gal (5-bromo-4- protein was eluted, and the protein cross-links were reversed by heating in SDS chloro-3-indolyl-␤-D-galactopyranoside) (Fig. 1A). Thus, YvaP sample loading buffer containing 5% mercaptoethanol. Samples were separated was identified as a putative repressor that is involved in nega- on 15% SDS gels and subjected to CatR-, YodB-, and anti-FLAG-specific West- ern blot analyses. tive regulation of the catDE operon in response to electro- philes. Hence, we renamed YvaP as a putative CatR repressor. We further noted that the catR sequence annotated in the old RESULTS SubtiList database contains sequencing errors that have been Identification of YvaP (CatR) as a repressor of the catE-lacZ corrected in the resequenced genome sequence (2). Homology fusion using TF arrays. To identify the regulator of the catDE comparison revealed that CatR belongs to the MarR/DUF24 VOL. 192, 2010 REGULATION OF catDE BY YodB AND CatR (YvaP) 4575 family of transcriptional regulators and is closely related to YodB (4). The catR gene consists of 327 bp and encodes a protein of 108 amino acids which shares 46% sequence identity to the YodB repressor. Transcription of the catDE operon is negatively controlled by the MarR/DUF24-family paralogs CatR and YodB in re- sponse to diamide and quinones. Northern blot experiments Downloaded from showed that the catDE operon is 13- to 60-fold induced in response to quinone-like electrophiles (catechol and MHQ) and diamide in the wild type (Fig. 1B). The catDE-specific transcript was 10-fold derepressed in the ⌬catR mutant under control conditions, confirming that CatR is a repressor of the catDE operon. However, we still observed a 3- to 6-fold induc- tion of catDE transcription in the ⌬catR mutant after exposure to electrophiles, which points to another redox-sensing regu- http://jb.asm.org/ lator controlling catDE expression. The TF array analysis did not reveal another negative regulator. Since CatR shares strong sequence similarity to the YodB repressor, we analyzed expression of the catDE operon in a ⌬yodB ⌬catR double mutant (Fig. 1B). Interestingly, the ⌬yodB ⌬catR double mu- tant showed 60-fold derepression of catDE transcription, re-

vealing that both YodB and CatR are involved in negative on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS regulation of the catDE operon. Surprisingly, the catDE FIG. 2. The conserved Cys7 residue of CatR is required for redox operon was repressed in the ⌬yodB single mutant (Fig. 1B) sensing in vivo. (A) Northern blot analysis of catDE transcription was under control conditions, suggesting that CatR and YodB performed using RNA isolated from the B. subtilis wild type (WT) and might compete for DNA binding in vivo. Furthermore, the ⌬catR and catRC7S mutant cells before (co) and 10 min after treat- catDE operon was 10- and 20-fold upregulated in the ⌬yodB ment with 1 mM MHQ (M), 2.4 mM catechol (cat), and 1 mM diamide (D). The arrow points toward the size of the catDE-specific transcript. mutant by 1 mM diamide and 3 mM catechol, and no dere- (B) The quantification of the catDE transcription ratios was performed pression was observed by MHQ (Fig. 1B). We thought that this from three independent experiments using ImageJ. (C) Western blot weaker induction of the catDE operon in the ⌬yodB mutant by analysis shows that similar CatR and CatRC7S proteins are produced diamide and catechol could be caused by the increased detox- in the wild type and the catRC7S point mutant, respectively. Cells were harvested before and 15 min after 1 mM of diamide stress, and 25 ␮g ification of the compounds by the azoreductase AzoR1 upregu- of protein extracts was subjected to CatR-specific Western blot anal- lated in the ⌬yodB mutant. Thus, we analyzed catDE transcrip- ysis. tion in the yodB mutant in response to higher sublethal concentrations of the compounds. In fact, a 30-fold and 65-fold derepression of catDE transcription was observed in the ⌬yodB mutant with higher doses of 2.5 mM MHQ and 12 mM cate- (GEO accession no. GSE22603). Thus, the catDE operon is chol, respectively (Fig. 1C). Finally, catDE transcription was the only apparent target for repression by CatR in B. subtilis. analyzed in a ⌬yodB azoR1 double mutant that has lost the The conserved Cys7 residue is required for regulation of resistance phenotype as shown previously (15). The Northern CatR in vivo. YodB was characterized as a redox-sensing re- blot analysis showed similar catDE transcription ratios in wild- pressor of the MarR/DUF24 family (1, 4, 15). CatR and YodB type and ⌬yodB azoR1 double mutant cells in response to 1 share the conserved N-terminal Cys residue, but CatR lacks mM diamide, 1 mM MHQ, and 3 mM catechol (Fig. 1D). This the Cys101 and Cys108 residues of YodB. The conserved Cys6 result indicates that CatR responds independently of YodB to is required for DNA-binding activity and redox sensing of catechol, MHQ, and diamide in vivo. YodB in vivo (4, 15). We analyzed the role of Cys7 of CatR in We next determined the transcriptional start site of the transcriptional regulation of the catDE operon using a catDE operon using total RNA of wild-type and ⌬catR mutant catRC7S mutant in Northern blot experiments (Fig. 2A and B). cells treated with MHQ and catechol stress using primer ex- The basal level of catDE transcription was 3- to 4-fold in- tension experiments (Fig. 1F). The 5Ј end of the catDE-specific creased in the catRC7S mutant compared to that in the wild mRNA was mapped at an adenine located 36 bp upstream of type. Treatment of catRC7S mutant cells with MHQ and di- the start codon (Fig. 1F). This transcriptional start site is pre- amide leads to a 2- to 4-fold derepression of catDE tran- ceded by a ␴A-type promoter sequence. Transcription of catDE scription, but no increased catDE transcript level was de- is strongly initiated from this ␴A-dependent promoter in re- tected after catechol stress (Fig. 2A and B). Thus, the sponse to electrophiles (MHQ, catechol, and diamide). response of the CatRS7S protein is significantly reduced To identify other genes that are regulated by CatR, microar- compared to that of the wild-type CatR protein, which in- ray experiments were performed to compare gene expression dicates that Cys7 is required for redox sensing of CatR in in the B. subtilis ⌬catR mutant and wild-type cells under con- vivo.Westernblotanalysesshowedthatsimilaramountsof trol conditions. However, only the catDE operon was repro- CatR and CatRC7S are produced in the wild type and ducibly upregulated (18- to 24-fold) in the ⌬catR mutant catRC7S mutant, respectively (Fig. 2C). 4576 CHI ET AL. J. BACTERIOL. Downloaded from http://jb.asm.org/

FIG. 3. The ⌬catR and ⌬yodB mutants are resistant and the ⌬catR yodB mutant is hyper-resistant to catechol and MHQ stress. B. subtilis wild on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS type (WT) and the ⌬catR and ⌬catR ⌬yodB mutants were grown in minimal medium to an OD500 of 0.4 and treated with 4.8, 7, 12, and 20 mM catechol and 1, 1.5, and 2.5 mM MHQ at the time points that were set to zero, indicated by an arrow.

⌬catR and ⌬catR ⌬yodB mutants show a catechol and MHQ CatR- and YodB-protected regions in the catDE promoter resistance phenotype. Previous phenotype analyses showed contain two 4-5-4-bp inverted repeat (IR) sequences with the that the growth of the ⌬catE deletion mutant was impaired in consensus of TTAC-N5-GTAA that overlap the Ϫ35 promoter the presence of 3.6 mM catechol, which reduced the growth region (BS1) and the transcription start point (BS2), which are rate of the wild-type strain only slightly (30). Thus, the catDE putative binding sites for CatR and YodB (Fig. 4D). Further operon confers resistance to catechol. Examination of the inspection of the upstream regions of other catDE operon growth phenotype showed that the ⌬catR mutant was able to homologs of related bacilli revealed that the BS2 region and grow with 7 mM catechol and 1 mM MHQ, reflecting a 2-fold the catDE promoter regions are highly conserved also in Ba- increase in quinone resistance compared to that of the wild cillus licheniformis, Bacillus halodurans, and Bacillus amyloliq- type (Fig. 3). The resistance of the ⌬yodB mutant was higher uefaciens (Fig. 5A and B). than that of the ⌬catR mutant since the ⌬yodB mutant was able Mutational analysis of the catDE promoter region reveals to grow with 12 mM catechol and 1.5 mM MHQ. An additive different DNA-binding activities of CatR and YodB to the BS1 increase in these growth phenotypes were observed in the and BS2 operator sites. Next, we analyzed the DNA-binding ⌬yodB ⌬catR double mutant that was resistant even to 20 mM activities of YodB and CatR to the catDE promoter using catechol and 2.5 mM MHQ. However, the ⌬catR mutant strain labeled promoter probes with mutations of the IR sequences. showed no increased resistance to diamide. Finally, also the The EMSAs of CatR bound to the promoter probes containing ⌬yodB ⌬catR double mutant showed a level of diamide resis- regions at Ϫ146 to ϩ20 relative to the TSS are shown in Fig. 6. tance similar to that of the ⌬yodB mutant. These results indi- Mutations of one IR element in BS2 (TTACN5GTAA to TT cate specific detoxification functions of the CatR-controlled ACN5GCCC) (ϩ20M1 probe) decreased binding affinity of thiol-specific dioxygenase CatE for quinone-like electrophiles CatR 4-fold, and deletion of both IR elements in BS2 (TTAC by ring cleavage of the compounds. This function of CatE has N5GTAA to GGGCN5GGGG) (ϩ20M2 probe) abolished already been confirmed in vitro in previous studies (30). CatR binding. Deletions of the upstream binding region to

Identification of overlapping CatR and YodB binding re- Ϫ40 and mutation of both IR elements of BS1 (TTACN5 gions in the catDE promoter. To identify the cis-acting se- GTAA to GGGCN5GGGG) (Ϫ60M3 probe) strongly reduced quences which function as operator sites for repression by CatR binding to the catDE promoter. These results indicate CatR and YodB, DNase I footprinting analysis was performed that both BS1 and BS2 operator sites are essential for binding with purified His-tagged CatR and YodB proteins on the top of CatR in vitro. and bottom strands of the catDE promoter using a 5Ј-end- In contrast, mutations in BS2 did not decrease binding of labeled catDE promoter DNA probe (Fig. 4A to C). Purified YodB to the catDE promoter. However, deletions of the up- CatR protein protected a region overlapping the catDE pro- stream binding region to Ϫ40 and mutation of BS1 abolished moter from positions Ϫ56 to ϩ12 relative to the transcriptional binding of YodB, indicating that BS1 rather than BS2 is es- start site (Fig. 4A and B). Purified YodB protein showed pro- sential for binding of YodB to the catDE promoter in vitro. tection of the same catDE promoter region (Fig. 4C). The Previously, we have proposed TACT-N7-AGTA as the con- VOL. 192, 2010 REGULATION OF catDE BY YodB AND CatR (YvaP) 4577 Downloaded from http://jb.asm.org/ on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS

FIG. 4. DNase I footprinting experiments of purified CatR to the top strand (A) and bottom strand (B) and footprinting experiments of YodB to the top strand of the catDE promoter (C) and CatR/YodB box alignments (D). The CatR- and YodB-protected sequences in the promoter region are indicated to the right (A to C), and the two TTAC-N5-GTAA inverted repeats (BS1 and BS2) are labeled by arrows in the sequence alignment (D). The positions relative to the transcriptional start site are shown to the left. The transcriptional start site (ϩ1) is indicated by an asterisk. For dideoxynucleotide sequencing, the dideoxynucleotide added in each reaction mixture is indicated above the corresponding lane. (D) Regions protected by CatR and YodB are shown with gray shading in the catD promoter sequence, including the inverted repeats as putative CatR boxes, which are labeled by arrows.

FIG. 5. Alignment of the conserved catDE promoter regions and CatR boxes among Bacillus species. (A) The conserved upstream regions of the catDE operons of B. subtilis, B. licheniformis, B. halodurans,andB. amyloliquefaciens were aligned using ClustalW, and the putative CatR boxes are indicated using WebLogo (B). Promoter sequences (Ϫ10 and Ϫ35) and the transcriptional start site are indicated, and the CatR boxes are indicated by arrows. (C) The conserved YodB box identified in the upstream regions of spx, azoR1, yodC,andcatD is displayed using WebLogo. 4578 CHI ET AL. J. BACTERIOL. Downloaded from http://jb.asm.org/ on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS

FIG. 6. DNA-binding experiments of CatR and YodB to the mutated catDE promoter. Electrophoretic gel mobility shift assays (EMSAs) were applied using purified CatR and YodB proteins and catD promoter probes ranging from Ϫ146 to ϩ20 without mutations (ϩ20) and with mutations in one and two repeat elements of BS2 (ϩ20M1 and ϩ20M2), deletions in the upstream binding region (Ϫ60 and Ϫ40), and mutations of the two repeat elements of BS1 (Ϫ60M3).

sensus of YodB binding sites (15). The BS1 site contains the and alkylated either with IAM or AMS. The extracts were sequence TACT-N7-AacA, which is similar to the YodB con- analyzed using CatR-specific Western blot analysis (Fig. 7E). sensus (Fig. 5C). The results showed that AMS-alkylated CatR is shifted under CatR is modified by intersubunit disulfide formation in vitro control conditions compared to the IAM sample, indicating and in vivo. Previously, we showed that the YodB repressor is that Cys7 is in the reduced form. However, diamide, MHQ, regulated by C6-C101Ј intersubunit disulfide formation in vitro. and catechol treatment of CatR shows that most CatR mi- The possibility of intersubunit disulfide bond formation was grates to the size of the IAM samples. This indicates that Cys7 analyzed for His-CatR protein in vitro using nonreducing SDS- of CatR is modified upon exposure to diamide and quinones PAGE and in vivo using diagonal Western blot analysis of inside cells. CatR-specific immunoprecipitates from wild-type cells. CatR Physical interaction of the paralogous DUF24-family re- forms DTT-reducible intermolecular disulfides that are linked pressors CatR and YodB. Next, we analyzed whether the by both Cys7 residues in vitro upon diamide, MHQ, and cate- paralogous MarR/DUF24 proteins YodB and CatR could chol stress (Fig. 7A to C). physically interact in vivo. To monitor complex formation of In the diagonal Western blot analysis, CatR was detected at CatR and YodB in cytoplasmic extracts of B. subtilis in vivo, we the diagonal under control conditions and CatR intersubunit expressed the CatR-FLAG protein under the control of the disulfides appeared to the right of the diagonal after diamide, xylose-inducible xylA promoter in the catR and yodB mutant MHQ, and catechol stress (Fig. 7D). These CatR intersubunit strains. It was verified, using catDE-specific Northern blot disulfides were also detected in the yodB mutant, confirming analysis, that the FLAG-tagged CatR protein was functional in that regulation of CatR occurs independently of YodB (Fig. vivo, as catDE transcription was similar in the ⌬catR catR- 7D). Thus, our data suggest that CatR is modified by intersub- FLAG strain to that in the wild type (see Fig. S1 in the sup- unit disulfide formation in vitro and in vivo. plemental material). Protein cross-linking was performed using To verify that the CatR Cys7 residue is modified in vivo, we the cross-linking reagent DSP, and the CatR-FLAG protein used the large alkylating agent AMS (4-acetamido-4Ј-maleimi- was purified using anti-FLAG affinity agarose from cytoplas- dylstilbene-2,2Ј-disulfonic acid) that results in a mass shift of mic extracts. We used Western blot analysis with YodB- and 500 Da if the Cys residue is free for AMS modification. Cells CatR-specific antibodies and mass spectrometry (MS) to ana- were harvested before and after diamide and MHQ treatment lyze coimmunoprecipitation of YodB and CatR-FLAG pro- VOL. 192, 2010 REGULATION OF catDE BY YodB AND CatR (YvaP) 4579 Downloaded from http://jb.asm.org/ on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS

FIG. 7. CatR forms intersubunit disulfides by diamide and quinones. (A to C) Purified CatR protein was treated with diamide (A), MHQ (B), or catechol (C) and subsequently either reduced with 10 mM DTT (ϩ DTT) or left oxidized (- DTT) and subjected to nonreducing SDS-PAGE analysis. (D) Diagonal nonreducing/reducing CatR-specific Western blot analysis of immunoprecipitated CatR protein that was harvested from wild-type (WT) cells and ⌬catR and ⌬yodB mutant cells before (co) or after exposure to diamide (Dia), MHQ, and catechol. (E) Protein extracts were harvested from wild-type cells under control conditions and after diamide, MHQ, and catechol stress and alkylated either by IAM or AMS that resulted in a mass shift of 500 Da per modified Cys. AMS- and IAM-treated extracts were separated using reducing SDS-PAGE and subjected to CatR-specific Western blot analysis. teins using the anti-FLAG-specific antibodies (Fig. 8). The CatR. However, the signal was also present in the ⌬yodB catR- Western blot analyses revealed strong amounts of precipitated FLAG strain, indicating that the YodB antibodies cross-react CatR-FLAG protein in the CatR-FLAG-expressing strains with the abundant CatR-FLAG protein. This was further con- and no CatR-specific signal using purification from wild-type firmed by mass spectrometry. Both YodB and CatR-FLAG extract. In fact, using YodB-specific Western blot analysis we proteins could be purified using YodB-specific immunoprecipi- also detected a strong signal for YodB at the same size of tation using Dynabeads protein A as verified by matrix-assisted laser desorption ionization–time of flight tandem mass spec- trometry (MALDI-TOF MS/MS) analysis (see Fig. S2C and D in the supplemental material). However, only peptides of the CatR protein could be detected in the MALDI-TOF spectrum of immunoprecipitated CatR-FLAG protein samples (see Fig. S2A and B in the supplemental material). This indicates that CatR and YodB do not form heterodimers in vivo and regulate their target genes independently. This also confirms the North- ern blot results shown in Fig. 1D, which show that CatR re- sponds independently of YodB in repression and regulation of catDE transcription. FIG. 8. Analysis of protein interactions between YodB and CatR in vivo. Soluble fractions were prepared from cells of the wild type (WT) and ⌬catR catR-FLAG and ⌬yodB catR-FLAG strains after protein DISCUSSION cross-linking with DSP according to the description in Materials and Methods. CatR-FLAG protein was immunoprecipitated using anti- In this study, we have identified CatR as another MarR/ FLAG affinity agarose, and the proteins were eluted using reducing DUF24-family repressor that, together with the YodB repres- sample buffer containing 5% mercaptoethanol (IP-CatR-FLAG). The sor, controls the catechol-2,3-dioxygenase-encoding catDE protein extracts were separated in lanes 4 to 6 each. Detection of YodB and CatR was performed using the specific antibodies. Note that operon. Our data show that inactivation of both CatR and the FLAG-tag results in a higher molecular mass of CatR-FLAG YodB is required for full derepression of catDE transcription. compared to CatR detected in wild-type extracts. The DNase I footprinting experiments revealed that both para- 4580 CHI ET AL. J. BACTERIOL. Downloaded from http://jb.asm.org/

FIG. 9. Quinone resistance network of the MhqR, YodB, and CatR regulons of B. subtilis. Three MarR-type repressors, MhqR, YodB, and CatR, respond to diamide and quinone-like electrophiles in B. subtilis. YodB and CatR represent paralogous MarR/DUF24-family repressors that sense electrophiles via the conserved N-terminal Cys residue. The regulatory mechanism of the MhqR repressor is unknown. YodB controls the azoreductase AzoR1, the nitroreductase YodC, and Spx and it also controls together with CatR the DoxX-like oxidoreductase CatD and

thiol-dependent dioxygenase CatE. The MhqR repressor regulates paralogous enzymes of the YodB and CatR regulons, including the azoreduc- on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS tase AzoR2, the nitroreductase MhqN, the DoxX-like oxidoreductase MhqP, and three thiol-dependent dioxygenases, MhqA, MhqE, and MhqO. The operator sequences for binding of MhqR, YodB, and CatR are generated using WebLogo. logs protect the same promoter region upstream of catDE, and DNA-binding activity and redox sensing of YodB in vivo (4, promoter mutagenesis analyses suggest that CatR binds two IR 15). Inducer responsiveness of the CatRC7S mutant protein sequences (BS1 and BS2) whereas only the upstream BS1 is was reduced compared to that of the wild-type CatR protein. needed for binding of YodB. Interestingly, the Northern blot In addition, Cys7 is required for DNA-binding activity of CatR, data (Fig. 1B) indicate that there might be a competition as derepression of catDE transcription is observed in the between both CatR and YodB for binding to the catDE oper- catRC7S mutant under control conditions. This indicates that ator in vivo since the catDE operon was repressed by CatR in mutation of this Cys residue is critical for DNA-binding activity the ⌬yodB mutant under control conditions. However, in and might interfere with the protein structure. This is in agree- ⌬catR and ⌬yodB single mutants we found derepression of ment with studies with the yodBC6S mutant that showed weak catDE transcription by diamide, MHQ, and catechol, suggest- derepression of azoR1 transcription under noninducing condi- ing that YodB and CatR respond independently of each other tions, indicating that this conserved Cys is required for repres- to the electrophiles in vivo. The lower responsiveness of CatR sion of YodB (4, 15). to diamide, MHQ, and catechol in the ⌬yodB mutant was We have further investigated the mechanism by which CatR caused by the quinone resistance phenotype due to derepres- senses and responds to diamide and quinones. The redox- sion of AzoR1. This is supported by the result that catDE sensing mechanisms have been well characterized for MarR- transcription is similarly derepressed in the wild type and in the type repressors of the OhrR family that respond to organic ⌬yodB azoR1 mutant. Furthermore, YodB regulation of azoR1 hydroperoxides. The two-Cys-type OhrR repressor of X. transcription occurs independently of CatR (see Fig. S3 in the campestris is inactivated by cumene hydroperoxide (CHP) via supplemental material). To confirm that both paralogous re- an intersubunit disulfide bond between Cys22 of one subunit pressors CatR and YodB act independently of each other, in and Cys127Ј of the other subunit (26). Inactivation of the one vivo protein cross-linking experiments were performed, fol- Cys-type OhrR protein of B. subtilis by CHP requires Cys15 lowed by affinity purification of FLAG-tagged CatR protein. oxidation to sulfonate that reacts further to a sulfonamide in However, the verification of copurification of possibly interact- vitro or a mixed disulfide (S-thiolated) form of OhrR with ing YodB protein was difficult to address using Western blot cysteine or bacillithiol in vivo (6, 14, 23). Recent data have analyses since the YodB antibodies cross-reacted with the shown that YodB senses quinones and diamide via C6-C101Ј abundant immunoprecipitated CatR-FLAG protein. Using intersubunit disulfides in vitro and resembles the model of the mass spectrometry we could confirm that only the CatR-FLAG two-Cys-type OhrR repressor of X. campestris (4). The data protein was purified from cell extracts using the anti-FLAG presented here suggest that CatR forms intermolecular disul- affinity agarose, and no interacting YodB protein. These re- fides between the lone Cys7 residues of both subunits in re- sults are consistent with the catDE transcription results and sponse to diamide and quinones in vitro and in vivo. We could exclude the possibility of physical interaction of both repres- not detect any S-thiolation modification of immunoprecipi- sors. tated CatR-FLAG protein by cysteine or bacillithiol in vivo Both YodB and CatR repressors share the conserved N- (data not shown). Recently, another one-Cys-type redox-sens- terminal Cys residue, which was shown to be required for ing MarR/DUF24-family regulator CgR1435 (QorR) has been VOL. 192, 2010 REGULATION OF catDE BY YodB AND CatR (YvaP) 4581 characterized that controls a quinone oxidoreductase in Albrecht, M. Hecker, and T. Schweder. 2005. Global expression profiling of Corynebacterium glutamicum Bacillus subtilis cells during industrial-close fed-batch fermentations with (5). Inactivation of QorR by di- different nitrogen sources. Biotechnol. Bioeng. 92:277–298. amide and H2O2 in vitro involves intersubunit disulfide forma- 12. Kobayashi, K. 2007. Bacillus subtilis pellicle formation proceeds through tion between the single Cys17 residues of both subunits. This genetically defined morphological changes. J. Bacteriol. 189:4920–4931. 13. Kumagai, Y., S. Koide, K. Taguchi, A. Endo, Y. Nakai, T. Yoshikawa, and N. confirms our results with CatR that is oxidized to Cys7 inter- Shimojo. 2002. Oxidation of proximal protein sulfhydryls by phenanthraqui- subunit disulfides. none, a component of diesel exhaust particles. Chem. Res. Toxicol. 15:483– 489. In conclusion, in this paper we have expanded the net- Downloaded from 14. Lee, J. W., S. Soonsanga, and J. D. Helmann. 2007. A complex thiolate work of quinone-sensing redox regulators by the identifica- switch regulates the Bacillus subtilis organic peroxide sensor OhrR. Proc. tion of the YodB-paralogous MarR/DUF24-family member Natl. Acad. Sci. U. S. A. 104:8743–8748. CatR as a major regulator of the catechol dioxygenase- 15. Leelakriangsak, M., N. T. Huyen, S. To¨we,N. van Duy, D. Becher, M. Hecker, H. Antelmann, and P. Zuber. 2008. Regulation of quinone detox- encoding catDE operon (Fig. 9). Together YodB, CatR, and ification by the thiol stress sensing DUF24/MarR-like repressor, YodB in MhqR control paralogous azoreductases, nitroreductases, Bacillus subtilis. Mol. Microbiol. 67:1108–1124. 16. Leelakriangsak, M., K. Kobayashi, and P. Zuber. 2007. Dual negative con- DoxX-like oxidoreductases, and thiol-dependent dioxygen- trol of spx transcription initiation from the P3 promoter by repressors PerR ases that collectively function in the detoxification of qui- and YodB in Bacillus subtilis. J. Bacteriol. 189:1736–1744. 17. Leichert, L. I., and U. Jakob. 2006. Global methods to monitor the thiol- nones and diamide by reduction or ring cleavage of these http://jb.asm.org/ disulfide state of proteins in vivo. Antioxid. Redox. Signal. 8:763–772. electrophiles in B. subtilis. 18. Liebeke, M., D. C. Po¨ther,N. van Duy, D. Albrecht, D. Becher, F. Hochgra¨fe, M. Lalk, M. Hecker, and H. Antelmann. 2008. Depletion of thiol-containing ACKNOWLEDGMENTS proteins in response to quinones in Bacillus subtilis. Mol. Microbiol. 69:1513– 1529. We are grateful to John Helmann and Shawn MacLellan for discus- 19. Majumdar, D., Y. J. Avissar, and J. H. Wyche. 1991. Simultaneous and rapid sion of the results prior to publication. We further thank Peter Zuber isolation of bacterial and eukaryotic DNA and RNA: a new approach for isolating DNA. Biotechniques 11:94–101. and Montira Leelakriangsak for providing plasmid pML54. 20. Monks, T. J., R. P. Hanzlik, G. M. Cohen, D. Ross, and D. G. Graham. 1992. This work was supported by grants from the Deutsche Forschungs- Quinone chemistry and toxicity. Toxicol. Appl. Pharmacol. 112:2–16. gemeinschaft, the Bundesministerium fu¨r Bildung und Forschung 21. Nakano, S., K. N. Erwin, M. Ralle, and P. Zuber. 2005. Redox-sensitive on December 29, 2011 by UNIVERSITAETSBIBLIOTHEK GREIFS (BACELL-SysMo 031397A), the Fonds der Chemischen Industrie, the transcriptional control by a thiol/disulphide switch in the global regulator, Bildungsministerium of the country Mecklenburg-Vorpommern and Spx. Mol. Microbiol. 55:498–510. BACELL-BaSysBio (LSHG-CT-2006-037469) (to M.H.), and a grant 22. Nakano, S., E. Kuster-Schock, A. D. Grossman, and P. Zuber. 2003. Spx- from the Deutsche Forschungsgemeinschaft AN 746/2-1 (to H.A.). dependent global transcriptional control is induced by thiol-specific oxidative stress in Bacillus subtilis. Proc. Natl. Acad. Sci. U. S. A. 100:13603–13608. 23. Newton, G. L., M. Rawat, J. J. La Clair, V. K. Jothivasan, T. Budiarto, C. 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zum Thema:

„Regulatory mechanisms of the disulfide stress response and

the role of the bacillithiol redox buffer in Gram-positive bacteria“

vorgelegt von

Bui Khanh Chi

! "#"! Reactive oxygen species (ROS) are generated inside bacteria by incomplete reduction of molecular oxygen during respiration or are generated as part of the host immune defence. ROS can damage all cellular macromolecules and also produce secondary reactive intermediates, like reactive electrophilic species (RES) that include quinones or aldehydes. Low molecular weight (LMW) thiols are small thiol-containing compounds that play essential roles in the defense against ROS and RES in all organisms. The best studied LMW thiol is the tripeptide glutathione (GSH), which is present in millimolar concentrations in the cytoplasm of eukaryotes and Gram-negative bacteria, but is absent in most Gram-positive bacteria. Firmicutes bacteria including Bacillus und Staphylococcus species have been recently discovered to utilize the redox buffer bacillithiol (BSH). LMW thiols function as redox buffers to maintain the reduced state of the cytoplasm. They are involved in the detoxification of ROS and RES and function as co-factors for redox enzymes (e.g. glyoxalases, S-transferases, peroxidases). Under conditions of oxidative stress, LMW thiols also react with protein thiols to form mixed LMW thiol – protein disulfides, termed S-thiolations, as major protection mechanism. Protein S- glutathionylation is the most important post-translational thiol-modification in eukaryotes that protects active site Cys residues against irreversible oxidation to Cys-sulfonic acids. Protein S- glutathionylation functions as redox-switch under oxidative stress and controls the activity of redox sensing transcription factors and metabolic enzymes. Protein S-glutathionylation is also implicated in the pathogenesis of neurodegenerative and cardiovascular diseases in eukaryotes. Investigating the role of BSH in oxidative stress response and ROS-induced S-thiolations in Firmicutes bacteria was one subject of this PhD thesis. Specifically, the regulatory mechanisms and post-translational thiol-modifications in response to NaOCl stress were studied in the model bacterium for low-GC Gram-positive bacteria Bacillus subtilis in chapter 2. Using transcriptomics, redox proteomics, and LTQ-Orbitrap mass spectrometry analysis, changes in gene expression profile and the targets for reversible thiol-modifications after NaOCl stress were analyzed in B. subtilis. The transcriptome profile after NaOCl stress was indicative of disulfide stress and overlapped strongly with the response to diamide. NaOCl stress caused induction of the thiol- and oxidative stress-specific Spx, CtsR, PerR and OhrR regulons. Thiol redox proteomics identified only few NaOCl-sensitive proteins with reversible thiol-oxidations that included GapA, MetE, AroA, MtnA and PurQ. Using mass spectrometry, eleven proteins were identified that were oxidized to mixed BSH protein disulfides (S-bacillithiolated) in B. subtilis cells after NaOCl-exposure. These S-bacillithiolated proteins included the OhrR repressor, two methionine synthases MetE and YxjG, the inorganic pyrophosphatse PpaC, the 3-D-phosphoglycerate dehydrogenase SerA, the bifunctional 3-deoxy-7- phosphoheptulonate synthase/chorismate mutase (AroA), the translation elongation factor EF-Tu (TufA), the inosine-5'-monophosphate dehydrogenase (GuaB), the ferredoxin-NADP+ oxidoreductase (YumC) and the putative bacilliredoxin (YphP). Methionine synthase MetE is the most abundant S- bacillithiolated protein in B. subtilis and other Bacillus species after NaOCl exposure as revealed by BSH-specific immunoblot analyses. S-bacillithiolation of OhrR repressor leads to upregulation of the

! "##! OhrA peroxiredoxin that confers together with BSH specific protection against NaOCl. S- bacillithiolation of MetE, YxjG, PpaC, and SerA causes hypochlorite-induced methionine starvation as supported by the induction of the S-box regulon. S-glutathionylation of MetE described in Escherichia coli also leads to enzyme inactivation and methionine auxotrophy. The putative bacilliredoxin YphP was S-bacillithiolated at its nucleophilic active site Cys suggesting a function in the de-bacillithiolation process. We further investigated the level of BSH and its oxidized BSH disulfide (BSSB) after NaOCl stress by thiol metabolomics analysis in collaboration with Alexandra Roberts and Chris Hamilton (University of East Anglia, Norwich, UK). S-bacillithiolation is accompanied by a 2-fold increase in the BSSB level and a decrease in the BSH/BSSB redox ratio in B. subtilis. To further assess the conservation of targets for S-bacillithiolations in other Firmicutes bacteria, we studied the S-bacillithiolomes of Bacillus megaterium, Bacillus pumilus, Bacillus amyloliquefaciens, and Staphylococcus carnosus under NaOCl stress conditions using redox proteomics and mass spectrometry which is described in chapter 3. In total, 54 S-bacillithiolated proteins were identified, including 29 unique proteins and 8 conserved proteins (MetE, AroA, GuaB, TufA, PpaC, SerA, YphP, YumC) involved in amino acid and cofactor biosynthesis, nucleotide metabolism, translation, protein quality control, redox and antioxidant functions. Together our data support a major role of BSH redox buffer in redox control and thiol protection of conserved and essential proteins against irreversible oxidation by S-bacillithiolations in Firmicutes bacteria. Future studies are underway to identify the function of the conserved bacilliredoxins in the de- bacillithiolation process. In response to ROS and RES, bacteria also activate the expression of antioxidant and detoxification enzymes, such as catalases, peroxidases, thiol-dependent peroxiredoxins and other specific oxidoreductases to detoxify ROS and RES. These defense mechanisms are often controlled by redox-sensitive transcription factors, such as OxyR of Escherichia coli that harbor redox-sensitive Cys residues to sense and respond to ROS or RES. The thiol group of the redox-sensing Cys is the most suseptable target for ROS and RES. It can be reversibly or irreversibly modified leading either to activation or inactivation of the transcription factors. Bacillus subtilis encodes redox-sensing MarR- type regulators belonging to the OhrR and DUF24-families that are conserved among bacteria. While OhrR respond to organic hydroperoxides and NaOCl, the DUF24-family senses electrophiles, such as diamide, quinones or aldehydes via thiol-based redox-switches. Hence, we were further interested in this PhD thesis to study at the molecular and structural level the redox-sensing mechanisms of novel redox-sensing MarR/DUF24-type regulators in B. subtilis. The work is described in chapters 4-6. We have characterized the regulatory mechanisms of HypR, YodB and CatR that sense and respond to hypochlorite, diamide and quinones stress. HypR is the first DUF24-family regulator whose crystal structure was resolved in collaboration with the department for protein biochemistry of Prof. Winfried and Dr. Gottfried Palm at the Ernst-Moritz-Arndt-University of Greifswald (chapter 4). HypR senses

! "#$! specifically disulfide stress and controls positively expression of the flavin oxidoreductase HypO after NaOCl and diamide stress. HypR resembles a 2-Cys-type regulator with a reactive nucleophilic N- terminal Cys14 and a second C-terminal Cys49 that are 8Å apart in the reduced HypR structure. The

Cys14 residue, which is conserved among the DUF24-family, has a lower pKa of 6.36. It is essential for activation of hypO transcription by disulfide stress. HypR is activated by Cys14-Cys49' intersubunit disulfide formation. The crystal structures of reduced and oxidized HypR proteins were resolved revealing structural changes of HypR upon oxidation. In reduced HypR a hydrogen-bonding network stabilizes the reactive Cys14 thiolate that is 8-9 Å apart from Cys49'. Oxidation of Cys14 residue breaks these H-bonds, reorients the monomers and moves the major groove recognition !4 and !4' helices ~4 Å towards each other. It will be interesting to study the functions of HypR-like regulators in pathogens that could have protective functions against the host immune defense. Besides HypR, B. subtilis encodes further MarR/DUF24-family members including the paralogous YodB and CatR repressors that sense quinones and diamide. YodB controls the azoreductase AzoR1, the nitroreductase YodC, and the Spx regulator that is up-regulated by quinones and diamide and confer resistance to these electrophiles (chapter 5). YodB resembles a 2-Cys-type MarR/DUF24-family regulator with three Cys residues (Cys6, Cys101 and Cys108). Using a yodBC6S mutant, it was shown that the conserved Cys6 is essential for redox sensing of diamide and MHQ in vivo. It was demonstrated that YodB is oxidized to Cys6-Cys101' or Cys6-Cys108' intersubunit disulfides in vivo. YodB and its paralog YvaP (CatR) were further identified as repressors of the yfiDE (catDE) operon encoding a catechol-2,3-dioxygenase that also contributes to quinone resistance (chapter 6). Only inactivation of both regulators, YodB and CatR results in full derepression of catDE transcription. DNA-binding assays and promoter mutagenesis showed that CatR protects two inverted repeats as operator sites BS1 and BS2 with the consensus sequence TTAC- N5-GTAA overlapping the -35 promoter region and the transcriptional start site. The BS1 operator was required for binding of YodB in vitro. CatR and YodB share the conserved N-terminal Cys residue, that is required for redox-sensing of CatR in vivo as shown by Cys-to-Ser mutagenesis. Although CatR is a 1-Cys-type regulator, our data showed that CatR also forms intermolecular disulfide in response to diamide and quinones in vitro. Thus, HypR, YodB and CatR are controlled by 2-Cys-type thiol-disulfide redox switches to sense disulfide and RES stress conditions, and to control specific RES detoxification enzymes.

! "#$! ! ! "#$%&'&()*+! ! !

! Eidesstattliche Versicherung

Hiermit erkläre ich, dass diese Arbeit bisher von mir weder an der Mathematisch-Naturwissenschaftlichen Fakultät der Ernst-Moritz-Arndt- Universität Greifswald noch an einer anderen wissenschaftlichen Einrichtung zum Zwecke der Promotion eingereicht wurde.

Ferner erkläre ich, dass ich diese Arbeit selbstständig verfasst und keine anderen als die darin angegebenen Hilfsmittel und Hilfen benutzt und keine Textabschnitte eines Dritten ohne Kennzeichnung übernommen habe.

Greifswald, den 16/11/2012

Bui Khanh Chi

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! "#$! ! ! "#$%&'()&*+,! ! ! List of publications

1. Chi, B.K., Kobayashi, K., Albrecht, D., Hecker, M., and Antelmann, H.* 2010. The paralogous MarR/DUF24-family repressors YodB and CatR (YvaP) control expression of the catechol dioxygenase CatE in Bacillus subtilis. J. Bacteriol. 192: 4571-4581.

2. Chi, B.K., Albrecht, D., Gronau, K., Becher, D. Hecker, M., and Antelmann, H.* 2010. The redox-sensing regulator YodB senses diamide and quinones via a thiol-disulfide switch in Bacillus subtilis. Proteomics 10: 3155-3164.

3. Lima, B.P., Antelmann, H., Gronau, K., Chi, B.K., Becher, D., Brinsmade SR, and Wolfe, A.J.* 2011. Involvement of protein acetylation in glucose-induced transcription of a stress- responsive promoter. Mol. Microbiol. 81: 1190-204.

4. Chi, B.K., Gronau, K., Mäder, U., Hessling, B., Becher, D., and Antelmann, H.* 2011. S- bacillithiolation protects against hypochlorite stress in Bacillus subtilis as revealed by transcriptomics and redox proteomics. Mol. Cell Proteomics 10: M111.009506.

5. Zuber, P.*, Chauhan, S., Pilaka, P., Nakano, M. M., Gurumoorthy, S., Chi, B.K., Antelmann, H., and Mäder, U. 2011. Phenotype enhancement screen of a regulatory spx mutant unveils a rome of the ytpQ gene in the control of iron homeostasis. PLoS One, 6: e25066.

6. Palm, G.#, Chi, B.K.#, Waack, P., Gronau, K., Becher, D., Albrecht, D., Hinrichs, W., Read, R.J. and Antelmann, H.* 2012. Structural insights into the redox-switch mechanism of the MarR/DUF24-family regulator HypR. Nucleic Acid Research 40: 4178-92. (#Both authors contributed equally to this work)

7. Chi, B.K., Roberts, A., Huyen, T.T.T., Gronau, K., Becher, D., Albrecht, D., Hamilton, C. and Antelmann, H.* 2012. S-bacillithiolation protects conserved and essential proteins against hypochlorite stress in Firmicutes bacteria. Antioxid. Redox Signal. doi:10.1089/ars.2012.4686

8. Hu, L., Chi, B.K., Kuhn, M.L., Filippova, E.V., Walker-Peddakotla, A.J., Bäsell, K., Becher, D., Anderson, W.F., Antelmann, H., and Wolfe, A.J.* Phosphorylation and acetylation co-regulate an Escherichia coli transcription factor. J. Biol. Chem., Submitted.

!"#$ ! ! "#$%&'()&*+,! ! ! Poster presentations

1. Bui Khanh Chi, Gottried J. Palm, Katrin Gronau, Dörte Becher, and Haike Antelmann. “Specific control of hypochlorite resistance by the redox-sensing MarR/DUF24-type regulator HypR in Bacillus subtilis”. 4th Congress of European Microbiologists FEMS 2011. Geneve, Switzerland. 2. Bui Khanh Chi, Katrin Gronau, Ulrike Mäder, Bernd Hessling, Dörte Becher, and Haike Antelmann. “S-bacillithiolation protects against hypochlorite stress in Bacillus subtilis as revealed by transcriptomics and proteomics”. 2012 Annual meeting of the VAAM. Tübingen, Germany.

!"#$ ! ! "#$%&'()&*+,! ! ! Authors contributions to the publications

1. Chi, B.K., Kobayashi, K., Albrecht, D., Hecker, M., and Antelmann, H.* 2010. The paralogous MarR/DUF24-family repressors YodB and CatR (YvaP) control expression of the catechol dioxygenase CatE in Bacillus subtilis. J. Bacteriol. 192: 4571-4581.

The authors Haike Antelmann and Bui Khanh Chi designed the experiments, analyzed data and wrote the paper. Haike Antelmann and Michael Hecker supervised this project. Bui Khanh Chi performed genetic, molecular biological and biochemical experiments of this work. Dirk Albrecht performed MALDI-MS/MS analysis. Kazuo Kobayashi provided B. subtilis mutants for this study.

2. Chi, B.K., Albrecht, D., Gronau, K., Becher, D. Hecker, M., and Antelmann, H.* 2010. The redox-sensing regulator YodB senses diamide and quinones via a thiol-disulfide switch in Bacillus subtilis. Proteomics 10: 3155-3164.

Together with Haike Antelmann, Bui Khanh Chi designed the experiments and analyzed data. Haike Antelmann and Michael Hecker supervised this project. Bui Khanh Chi performed genetic, molecular biological and biochemical experiments of this work. She also critically read and appoved the final manuscript. Dirk Albrecht performed MALDI-MS/MS analysis. Katrin Gronau and Dörte Becher performed LTQ Orbitrap LC-MS/MS analysis of YodB protein. Kazuo Kobayashi provided B. subtilis mutants for this study.

3. Lima, B.P., Antelmann, H., Gronau, K., Chi, B.K., Becher, D., Brinsmade SR, and Wolfe, A.J.* 2011. Involvement of protein acetylation in glucose-induced transcription of a stress- responsive promoter. Mol.Microbiol. 81: 1190-204.

Alan J. Wolfe and Bruno P. Lima designed the experiments, analyzed data and wrote the paper. Bruno P. Lima and S.R. Brinsmade performed genetic, molecular biological and biochemical experiments of this work. Haike Antelmann and Bui Khanh Chi made tryptic digestions of acetylated proteins and analyzed LC-MS/MS data. Katrin Gronau and Dörte Becher performed LTQ Orbitrap LC-MS/MS analysis of acetylated peptides.

!"#$ ! ! "#$%&'()&*+,! ! !

4. Chi, B.K., Gronau, K., Mäder, U., Hessling, B., Becher, D., and Antelmann, H.* 2011. S- bacillithiolation protects against hypochlorite stress in Bacillus subtilis as revealed by transcriptomics and redox proteomics. Mol Cell Proteomics 10: M111.009506.

Together with Haike Antelmann, Bui Khanh Chi designed the experiments and analyzed data. Bui Khanh Chi performed genetic, molecular biological, biochemical and redox proteomics experiments of this work. She also critically read and appoved the final manuscript. Katrin Gronau, Bernd Hessling and Dörte Becher performed LTQ Orbitrap LC-MS/MS analysis of S-bacillithiolated proteins. Ulrike Mäder performed transcriptome analysis of B. subtilis for this study.

5. Zuber, P.*, Chauhan, S., Pilaka, P., Nakano, M. M., Gurumoorthy, S., Chi, B. K., Antelmann, H., and Mäder, U. 2011. Phenotype enhancement screen of a regulatory spx mutant unveils a role for the ytpQ gene in the control of iron homeostasis. PLoS One, 6: e25066.

Peter Zuber, S. Chauhan, P. Pilaka, M. M. Nakano and S. Gurumoorthy designed and performed genetic and molecular biological experiments, analyzed data and wrote the paper. Haike Antelmann and Bui Khanh Chi performed proteome analysis, and analyzed proteome and transcriptome data. Ulrike Mäder performed transcriptome analysis of B. subtilis for this study.

6. Palm, G.#, Chi, B.K.#, Waack, P., Gronau, K., Becher, D., Albrecht, D., Hinrichs, W., Read, R.J. and Antelmann, H.* 2012. Structural insights into the redox-switch mechanism of the MarR/DUF24-family regulator HypR. Nucleic Acid Research 40: 4178-92. (#Both authors contributed equally to this work)

Together with Haike Antelmann and Gottried Palm, Bui Khanh Chi designed the experiments and analyzed data. Bui Khanh Chi performed genetic, molecular biological, biochemical and redox proteomics experiments of this work. She also critically read and appoved the final manuscript. Gottfried Palm, Winfried Hinrichs, Randy J. Read and Paul Waack performed protein crystallization and structure refinement of the reduced and oxidized HypR proteins. Dirk Albrecht performed MALDI-MS/MS analysis. Katrin Gronau and Dörte Becher performed LTQ Orbitrap LC-MS/MS analysis of HypR protein.

!"#$ ! ! "#$%&'()&*+,! ! !

7. Chi, B.K., Roberts, A., Huyen, T.T.T., Gronau, K., Becher, D., Albrecht, D., Hamilton, C. and Antelmann, H.* 2012. S-bacillithiolation protects conserved and essential proteins against hypochlorite stress in Firmicutes bacteria. Antioxidants & Redox signaling, doi:10.1089/ars.2012.4686

Together with Haike Antelmann, Bui Khanh Chi designed the experiments and analyzed data. Bui Khanh Chi and T.T.T Huyen performed genetic, molecular biological, biochemical and redox proteomics experiments of this work. Bui Khanh Chi also critically read and appoved the final manuscript. Katrin Bäsell and Dörte Becher performed LTQ Orbitrap LC-MS/MS analysis of S-bacillithiolated proteins. Dirk Albrecht performed MALDI-MS/MS analysis. Alexandra Roberts and Chris J. Hamilton performed thiol metabolome analysis and determined BSH/BSSB ratios in B. subtilis for this study.

8. Hu, L., Chi, B.K., Kuhn, M.L., Filippova, E.V., Walker-Peddakotla, A.J., Bäsell, K., Becher, D., Anderson, W.F., Antelmann, H., and Wolfe, A.J.* Phosphorylation and acetylation co-regulate an Escherichia coli transcription factor. J Biol Chem, Submitted.

Alan J. Wolfe, Linda Hu, A.J. Walker-Peddakotla and Wayne F. Anderson designed the experiments, analyzed data and wrote the paper. Linda Hu, A.J. Walker-Peddakotla, M.L. Kuhn, E.V. Filippova, W.F. Anderson performed genetic, molecular biological and biochemical experiments of this work. Haike Antelmann and Bui Khanh Chi made tryptic digestions of acetylated proteins and analyzed LC-MS/MS data. Katrin Bäsell and Dörte Becher performed LTQ Orbitrap LC-MS/MS analysis of acetylated peptides.

Greifswald, den 16/11/2012 Greifswald, den 16/11/2012

Priv.-Doz. Dr. Haike Antelmann Bui Khanh Chi

!"!# Acknowledgements

I want to thank my supervisor, Privatdozentin Dr. Haike Antelmann for giving me a great opportunity to work on this exciting project funded by the Deutsche Forschungsgemeinschaft. She introduced me into the topic of bacterial redox control mechanisms and taught me many interesting techniques to analyse protein modifications and bacterial physiology. She also gave me the chance to participate in international conferences. I am really grateful for her guidance and support during the time of my PhD thesis in Greifswald. I further thank Prof. Micheal Hecker for giving me the opportunity to work at the Institute of Microbiology with access to state-of-the-art transcriptomics and proteomics techniques. His support and interest in this project is greatly appreciated. I would like to thank the mass spectrometry group of Privatdozentin Dr. Dörte Becher, Dr. Dirk Albrecht, Katrin Bäsell and Bernd Hessling for the mass spectrometry analysis that contributed to the success of this PhD thesis. My specials thank also go to: Dr. Ulrike Mäder (Interfaculty Institute for Genetics and Functional Genomics) for the transcriptomics analysis. Prof. Winfried Hindrichs, Dr. Gottfried Palm, Paul Waack, Birthe Steigemann and Dr. Randy Read (University of Cambridge) for the collaboration with protein crystallization and structural refinement of HypR. Dr. Chris Hamilton and Alexandra Roberts (University of East Anglia) for the collaboration with bacillithiol metabolomics. I am also grateful to Tran Thi Thanh Huyen (Department of Microbiology, Hanoi University of Sciences) for her collaboration with the redox proteome analysis of different Firmicutes bacteria. External collaboration partners for their valuable support with Bacillus subtilis mutants strains and discussions of results, especially the groups of Prof. John Helmann (Cornell University, Ithaca, NY), Prof. Peter Zuber (Oregon Health and Science University, Oregon) and Kazuo Kobayashi (Nara Institute of Science and Technology, Japan). Prof. Frieder Schauer for providing strains Bacillus pumilus SBUG1799 and Bacillus megaterium SBUG1152, and Prof. Kevin Devine (Trinity College Dublin) for the advice on in vitro transcription assays.

! "#$! The Decodon Company for support with Decodon Delta2D software. I want to thank all colleagues and the technical staff at the Institute of Microbiology for their kindness and friendship, especially Dana Clausen for excellent technical assistance, support and for the friendly working atmosphere. I would like to thank Prof. Truong Nam Hai at the Institute for Biotechnology, Vietnam Academy of Science and Technology for supporting me to complete my PhD work. Last but not least, my deep gratitude goes to my family and friends who have given me so much support, encouragement and strength. I am very thankful to know that they will always stand behind me.

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