Genome-wide responses and regulatory mechanisms to thiol-specific electrophiles in Bacillus subtilis

Inauguraldissertation zur Erlangung des akademischen Grades doktor rerum naturalium (Dr. rer. nat.) an der Mathematisch-Naturwissenschaftlichen Fakultät der Ernst-Moritz-Arndt-Universität Greifswald

Vorgelegt von Nguyen Thi Thu Huyen geboren am 30.08.1979 in Haiduong-Vietnam

Greifswald, 2008

Dekan: Prof. Dr. Klaus Fesser

1. Gutachter 1: Prof. Dr. Michael Hecker 2. Gutachter 2: Prof. Dr. Jan Maarten van Dijl

Tag der Promotion: 16. 10. 2008 i

CONTENTS

Summary of the thesis...... ii Part 1: Genome-wide responses and regulatory mechanisms to thiol-specific electrophiles in Bacillus subtilis...... 1 INTRODUCTION...... 2 1. The role of low molecular-weight thiols in the oxidative stress response...... 2 2. Induction of the thiol-specific stress response by quinone-like electrophiles and diamide in B. subtilis.....3 3. Regulation of the thiol-specific stress response in B. subtilis...... 5 3.1. Spx-an activator of genes involved in the maintenance of the thiol-disulfide balance...... 6 3.2. CymR-the repressor of the cysteine metabolism regulon...... 7 3.3. HrcA and CtsR-heat-shock-specific regulators ...... 8 3.4. PerR-the main regulator of the oxidative stress response...... 8 3.5. The MarR-type regulators ...... 10 4. The response of bacteria to the electrophile methylglyoxal (MG) ...... 11 5. The response of bacteria to the electrophile formaldehyde (FA) ...... 16 6. The aim of this part of the thesis ...... 21 Chapter 1: Regulation of quinone detoxification by the thiol stress sensing DUF24/MarR-like repressor, YodB in Bacillus subtilis...... 23 Chapter 2: The MarR/DUF24-type repressor CatR (YvaP) controls the thiol-stress inducible catechol dioxygenase YfiE (CatE) in Bacillus subtilis...... 41 Chapter 3: Genome wide responses to carbonyl electrophiles in Bacillus subtilis: control of the thiol- dependent formaldehyde dehydrogenase AdhA and cysteine proteinase YraA by the novel MerR-family regulator YraB (AdhR)...... 62 GENERAL DISCUSSION...... 96 1. The role of the MarR/DUF24 regulators as sensors for electrophilic quinones and diamide in B. subtilis96 2. The MG and FA stimulons of B. subtilis...... 106 Part 2: Genome-wide responses to manganese limitation in Bacillus subtilis ...... 116 SUMMARY...... 117 INTRODUCTION...... 118 1. Manganese (Mn) transport ...... 118 2. Cellular functions of Mn ...... 119 2.1. Function of Mn in the oxidative stress response ...... 121 2.2. Role of Mn during B. subtilis sporulation and germination ...... 122 2.3. Role of Mn in gene regulation...... 124 2.3.1. MntR as regulator of Mn homeostasis...... 124 2.3.2. Fur-like metalloregulators ...... 125 3. The aim of this part of the thesis ...... 128 MATERIALS AND METHODS ...... 129 1. Bacterial strain and medium...... 129 2. Growth conditions...... 129 3. Scanning electron microscopy (SEM)...... 129 4. 2-D PAGE ...... 130 4.1. Protein extraction ...... 130 4.2. 2-D PAGE and quantitative image analysis ...... 130 5. DNA microarray...... 131 RESULTS AND DISCUSSION ...... 132 1. The impact of Mn on growth behavior...... 132 2. Morphological, proteomic and transcriptomic studies of B. subtilis in response to Mn limitation ...... 132 2.1. Morphology of B. subtilis in response to Mn limitation...... 132 2.2. Proteome and transcriptome analyses of B. subtilis in response to Mn limitation ...... 134 References...... 154 List of publications...... 170 Curriculum vitae ...... 171 Acknowledgements...... 172

Nguyen Thi Thu Huyen ii Summary

Summary of the thesis

The soil-dwelling bacterium Bacillus subtilis is regarded as model organism for functional genomic research of low GC Gram-positive bacteria. Several stress and starvation- responsive stimulons have been extensively studied in B. subtilis using transcriptomic and proteomic approaches. However, there was only little information about the response of B. subtilis to toxic phenolic compounds, which are often found in the soil. The group of Haike Antelmann has monitored the expression profile of B. subtilis after exposure to phenol, salicylic acid, catechol, chromanon and MHQ. Interestingly, proteome and transcriptome analyses showed a strong overlap in the expression profile after exposure to catechol, MHQ and the thiol-reactive electrophile diamide. Later on it was shown that catechol and MHQ are auto-oxidized to electrophilic quinones which target thiols via the thiol-(S)-alkylation reaction leading to aggregation and depletion of LMW thiols and thiol-containing proteins.

The response to electrophilic quinones and diamide is governed by a complex network of transcription factors, including Spx, CtsR, PerR, CymR and the novel MarR-type repressors MhqR (YkvE), YodB and YvaP. The regulatory mechanisms of these novel thiol- stress sensors YodB and YvaP are studied as part of this thesis in collaboration with the group of Peter Zuber (Oregon). YodB negatively regulates the expression of the nitroreductase YodC and the azoreductase YocJ (AzoR1) after exposure to electrophilic quinones and diamide. The azoreductase AzoR1 is a paralog of AzoR2 that is under control of MhqR. Both paralogous azoreductases (AzoR1 and AzoR2) have common functions in quinone and azo- compound reduction to protect cells against the thiol reactivity of electrophiles. The group of Peter Zuber (Oregon) found that DNA binding activity of YodB is directly inhibited by thiol- reactive compounds in vitro. Mass spectrometry approaches suggested that YodB is regulated by a thiol-(S)-alkylation mechanism in response to quinones. Mutational analyses revealed that the conserved Cys6 residue of YodB is required for optimal repression in vivo and in vitro. Recent studies further suggest that YodB is redox-regulated by intersubunit disulfide formation in vivo by diamide.

In addition to the azoreductases, several MhqR-controlled thiol-dependent dioxygenases (MhqA, MhqE and MhqO) confer resistance to quinones. In collaboration with Kazuo Kobayashi (Nara), the YodB-paralogous MarR/DUF24-family regulator, YvaP was identified as repressor of the catechol-2,3-dioxygenase encoding yfiDE (catDE) operon. DNA binding activity of YvaP was also directly inhibited by quinones and diamide in vitro indicating that also YvaP is regulated via post-translational modifications. Mutational analyses showed that the conserved Cys7 is essential for YvaP regulation in vivo and serves as sensor for thiol-reactive compounds. In addition, also the basic amino acids K19, R20 are essential for YvaP repression in vivo as well as conserved basic arginine and lysine residues located in the DNA binding helix-turn-helix (HTH) motif. Non-reducing PAGE analysis

Nguyen Thi Thu Huyen iii Summary suggests the formation of an intersubunit disulfide bond in a YvaP dimer upon treatment with quinones and diamide in vitro.

Together these data show that B. subtilis has evolved a network of three MarR-type repressors YkvE (MhqR), YodB and YvaP that control paralogous quinone resistance determinants including quinone reductases (AzoR1 and AzoR2) and thiol-dependent ring- cleavage enzymes (CatE, MhqA, MhqE and MhqO). These thiol-stress responsive regulators confer resistance and allow adaptation to quinone-like electrophiles and diamide as novel physiological mechanism for Gram-positive bacteria.

Besides quinones, also α,β-unsaturated carbonyls are electrophilic compounds which react via the thiol-(S)-alkylation reaction with thiols. Thus, we were also interested in the response of B. subtilis to the toxic electrophiles methylglyoxal (MG) and formaldehyde (FA). We analyzed the changes in the transcriptome and proteome of B. subtilis after exposure to MG and FA. Like quinone compounds, both MG and FA induce the thiol-specific stress response as revealed by the induction of the CtsR, PerR, Spx, CymR, YodB, YvaP, ArsR and CsoR regulons. Metabolomic approaches confirmed that these reactive aldehydes deplete the cellular thiol pool and thus act like quinones as another class of thiol-reactive electrophiles. Additionally, MG and FA also triggered responses to overcome DNA damage as revealed by the induction of the SOS regulon. Our studies further revealed the specific induction of two FA detoxification pathways regulated by the MarR/DUF24 family repressor HxlR, and the novel MerR/NmlR-type regulator YraB (AdhR). HxlR positively regulates the hxlAB operon encoding the ribulose monophosphate pathway. AdhR positively regulates an adhA-yraA operon that encodes the thiol-dependent formaldehyde dehydrogenase (AdhA) and the DJ1/PfpI-like cysteine proteinase (YraA), and the yraC gene that encodes a γ- carboxymuconolactone decarboxylase. Thus, the AdhR regulon is involved in the detoxification of FA to formate via the formaldehyde dehydrogenase AdhA which catalyzes the cleavage of S-hydroxymethylcysteine adducts. In addition, the cysteine proteinase YraA could be involved in the degradation of S-hydroxymethylcysteine-modified and damaged protein thiols. In collaboration with the group of John Helmann (Ithaca), it was shown that AdhR binds in vitro to a conserved inverted repeat between the -10 and -35 promoter elements upstream of adhA, yraB and yraC. In addition, we showed that the conserved Cys52 of AdhR is essential for aldehyde sensing and activation of adhA-yraA transcription in vivo. Thus, we speculate that redox regulation of AdhR involves thiol-(S)-alkylation of this Cys52 residue by aldehydes as another novel mechanism of bacterial physiology.

Nguyen Thi Thu Huyen 1 Thiol-specific stress response

Part 1

Genome-wide responses and regulatory

mechanisms to thiol-specific electrophiles

in Bacillus subtilis

Supervisor: Prof. Dr. Michael Hecker

Dr. Haike Antelmann

Introduction 2 Thiol-specific stress response

INTRODUCTION

1. The role of low molecular-weight thiols in the oxidative stress response

Low molecular-weight (LMW) thiols play an essential role as a thiol redox buffer in the cell to maintain the reducing state of the cytoplasm which is necessary for regular metabolic activities. Thus, these thiol buffers represent a major biological adaptation that is important for the survival of organisms under various endogenous and exogenous stresses. The simplest LMW thiol is hydrogen sulfide, primarily presents as a bound cofactor in iron- sulfur proteins [Fahey, 2001]. The tripeptide γ-glutamylcysteinylglycine, which is also known as glutathione (GSH) is the most dominant thiol predominantly found in eukaryotic cells [Meister and Anderson, 1983]. In addition to its key role in maintaining the proper oxidation state of protein thiols, GSH plays a critical role in protection against severe stresses. GSH protects against osmotic and acid stresses, toxic drugs like methylglyoxal, chlorine compounds like hypoclorous acid, and oxidative stress such as hydrogen peroxide. GSH is also involved in the regulation of intracellular potassium levels and in preventing the formation of non-native protein disulfides in the cytoplasm [Masip et al., 2006]. In Gram- negative bacteria, GSH has also been implicated in the growth of these organisms on recalcitrant nutrients by serving as a cofactor for enzymes in various degradation pathways [Vuilleumier and Pagni, 2002]. Here, the role of GSH in protection against oxidative stress is discussed in more detail.

Oxidative stress occurs when cells are exposed to elevated levels of ROS such as hydrogen peroxides, alkyl hydroperoxides and hydroxyl radicals [Storz and Imlay, 1999]. S- thiolation is crucial for protection and regulation of thiol-containing proteins during oxidative stress and is frequently achieved by the formation of mixed disulfides with GSH. For example, the formation of mixed disulfides between target proteins and the LMW thiol GSH is a key event in the regulation of eukaryotic proteins, including c-Jun, thioredoxin and glyceraldehyde-3-phosphate dehydrogenase [Klatt et al., 1999a; 1999b; Casagrande et al., 2002; Cotgreave et al., 2002]. S-glutathionylation has also been reported to modulate the activity of E. coli methionine synthase (MetE), phosphoadenosine phosphosulfate reductase (CysH) and the oxidative stress-specific transcription factor OxyR [Kim et al., 2002; Lillig et al., 2003; Hondorp and Matthews, 2004].

The Gram-positive model bacterium B. subtilis lacks detectable levels of GSH and the enzymes required for GSH synthesis [Fahey et al., 1978; Copley and Dhillon, 2002]. Thus, B. subtilis produces different LMW thiols, such as cysteine and CoASH, which appear to function in a similar way like GSH. In a previous thiol-redox proteomic analysis of Hochgräfe and coworkers (2005), it was reported that reversible thiol oxidation occurs in a number of B. subtilis proteins in response to treatment with the thiol-reactive compound diamide. Later on,

Introduction 3 Thiol-specific stress response

Hochgräfe and coworkers (2007) showed that some of these reversible thiol-modifications are mixed disulfides with the LMW thiol cysteine (S-cysteinylated proteins). Thus, S-thiolation represents a general mechanism to protect protein thiols from overoxidation and irreversible damage in B. subtilis and may play a role in redox regulation of several thiol-containing proteins. Moreover, Lee et al. (2007) demonstrated that the organic peroxide-sensing repressor OhrR of B. subtilis is reversibly inactivated by S-thiolation with cysteine, coenzyme A (CoASH) and a novel 396 Da thiol under conditions of oxidative stress provoked by cumene hydroperoxides in vivo. The structure of this novel thiol is currently characterized and it seems to be an abundant LMW thiol consisting of cysteine, maleic acid and a sugar which is also present in Deinococcus (John Helmann, personal communication). It was also shown that overoxidation of cysteine residues in the glyceraldehyde-3-phosphate dehydrogenase to sulfonic acid caused by high level peroxide stress results in protein damage and in the irreversible loss of activity in B. subtilis and Staphylococcus aureus [Weber et al., 2004; Hochgräfe et al., 2005]. Such irreversible overoxidation of cysteine to sulfonic acid has been recently shown also as regulatory mechanism for OhrR in response to the strong oxidant linoleic acid [Soonsanga et al., 2008a]. Thus, depending on the oxidant, OhrR can be either reversibly oxidized or can instead function as a sacrificial regulator.

2. Induction of the thiol-specific stress response by quinone-like electrophiles and diamide in B. subtilis

The soil-dwelling bacterium B. subtilis is exposed to different toxic, antibiotic and antimicrobial compounds which are produced in microbial communities, plants and fungi, or derived from environmental pollutants. The response of B. subtilis to different classes of antibiotics was analyzed using the 2D gel-based proteomic approach [Bandow and Hecker, 2007]. However, there is very little knowledge regarding the response of B. subtilis to aromatic antimicrobial compounds and whether B. subtilis is able to detoxify aromatic compounds. Aromatic compounds originate from both natural and man-made sources, which are present in the environment [Smith, 1990]. Aromatic compounds can have toxic effects or provide carbon sources for bacteria. Many of aromatic compounds are major environmental pollutants that must be removed as these often are toxic to cellular systems [Dagley, 1986]. Recently, using proteomic and transcriptomic approaches, the global expression profile of B. subtilis was monitored after exposure to phenol, salicylic acid, the fungal-related antimicrobial 6-brom-2-vinyl-chroman-4-on (chromanon), catechol and methylhydroquinone (MHQ) by Haike Antelmann´s group [Tam et al., 2006b; Duy et al., 2007a; 2007b]. These analyses revealed the induction of general and specific stress responses and uncovered novel complex thiol-stress responsive regulatory mechanisms that confer resistance to electrophiles.

Phenol is a common environmental pollutant and chaotrope solute that causes water stress in Pseudomonas putida [Hallsworth et al., 2003]. The induction of stress responsive

Introduction 4 Thiol-specific stress response proteins caused by aromatic phenolic compounds has been shown in Pseudomonas putida [Santos et al., 2004], Methylocystis sp. [Uchiyama et al., 1999], Nocardioides sp. [Cho et al., 2000], Acinetobacter radioresistens [Giuffrida et al., 2001], Acinetobacter calcoaceticus [Benndorf et al., 2001], Acinetobacter lwoffii [Kim et al., 2004] and Stenotrophomonas sp. [Ho et al., 2004]. Phenol induced a classical heat-shock expression profile in B. subtilis as revealed by the induction of the HrcA, σB and CtsR heat-shock regulons. In addition, the activation of Spx dependent genes was shown by phenol that is induced under thiol-specific oxidative stress conditions such as diamide. However, whereas the diamide-induced stimulon also includes the oxidative stress-specific PerR regulon, no oxidative stress response was observed in phenol-treated cells. Finally, since phenol induced no further specific marker proteins besides the heat-shock proteomic signature, the mode of action for phenol appears to be protein damage [Tam et al., 2006b].

Phenolic acids such as ferulic, p-coumaric and caffeic acids are present in plant-soil ecosystem. They can be considered as natural toxins, which inhibit the growth and cause specific stress responses in microorganisms that allow adaptation and confer resistance to these compounds [Clausen et al., 1994; de Vries et al., 1997; Lambert et al., 1997]. For example, ferulic acid induces virulence gene expression in Agrobacterium tumefaciens [Kalogeraki et al., 1999] and proton pumps are induced in response to cinnamic acid in Saccharomyces cerevisiae [Chambel et al., 1999]. However, phenolic acids can also serve as the sole carbon-energy source in other bacteria such as Pseudomonas and Acinetobacter calcoaceticus [Lynd et al., 2002]. In B. subtilis, salicylic acid, which is also present in the soil, caused predominantly the induction of σB dependent general stress response, which is not related to the acidic conditions. In addition, both phenolic acid decarboxylase (Pad) systems (PadC and BsdBCD) which are involved in detoxification of phenolic acids confer resistance to salicylic acid [Duy et al., 2007a]. Finally, salicylic acid also caused protein damage by non- native disulfide formation as reflected by induction of the HrcA, CtsR heat-shock regulons and the Spx thiol-stress regulon [Nakano et al., 2003b; Zuber, 2004; Duy et al., 2007a].

Chromanon is derived from the natural antibiotic compounds aposphaerin A and B isolated from the fungus Aposphaeria species, which showed significant activity against methicillin resistant Staphyloccocus aureus strains [Albrecht et al., 2005]. In B. subtilis, chromanon caused predominantly thiol-specific protein damage as indicated by the induction of the HrcA, CtsR and Spx regulons [Duy et al., 2007b]. However, the oxidative stress- specific PerR regulon was not induced by chromanon, which indicates that the mode of action of chromanon differs also from that of diamide. In addition, chromanon treatment resulted in a weak cell wall stress response, as reflected by the induction of the bacitracin resistance gene bcrC and the lia operon [Jordan et al., 2007]. Interestingly, several oxidoreductases were strongly induced by chromanon which might confer resistance to chromanon. For example, the ycnDE, ydgGH and ydgIJ operons encode NAD(P)H-flavin-dependent oxidoreductases,

Introduction 5 Thiol-specific stress response an antibiotic biosynthesis monooxygenase and MarR-type transcriptional regulators [Duy et al., 2007b].

The most interesting expression profile was observed for the phenolic compounds catechol and MHQ, which both induce different ring-cleavage enzymes as well as the thiol- specific oxidative stress response in B. subtilis. Like diamide, catechol caused the common induction of the HrcA, CtsR, Spx, CymR, PerR and Fur regulons which resemble the thiol- specific oxidative stress signature. Diamide is a thiol-reactive oxidant that causes the formation of non-native disulfide bonds in cellular thiol-containing proteins [Leichert et al., 2003]. MHQ is the basic structure of the so-called ganomycins, which were isolated from basidiomycete Ganoderma pfeifferi and exhibited broad-spectrum antimicrobial activity [Mothana et al., 2000]. The expression profile of MHQ was also similar to that of diamide and catechol which indicates that these phenolic compounds act like diamide as thiol-reactive compounds in B. subtilis [Duy et al., 2007b; Antelmann et al., 2008]. Further evidence is presented that different extradiol dioxygenases are specifically induced by catechol, MHQ and diamide exposure which are encoded by the genes yfiE, mhqA, mhqE and mhqO [Duy et al., 2007b]. The results presented in this thesis indicate that these genes could encode novel thiol-dependent dioxygenases that are involved in the specific detoxification of quinone-like electrophiles, which deplete the cellular thiol pool via the thiol-(S)-alkylation reaction [Liebeke et al., 2008]. Such quinones are produced upon autooxidation of catechol and MHQ, which act as thiol-stress specific electrophiles. Thus, the induction of thiol-dependent dioxygenases is coupled to the thiol-specific oxidative stress response in B. subtilis.

In conclusion, the transcriptome and proteome results in response to the phenolic compounds phenol, phenolic acid (salicylic acid) and chromanon indicated predominantly a heat shock-specific proteomic signature and protein damage [Tam et al., 2006b; Duy et al., 2007a; 2007b]. In contrast, electrophilic quinones deplete directly LMW thiol buffers as well as thiol-containing proteins via an alkylation mechanism, which results in a thiol-specific oxidative stress response in B. subtilis [Tam et al., 2006b; Duy et al., 2007a; 2007b; Antelmann et al., 2008].

3. Regulation of the thiol-specific stress response in B. subtilis

Oxidative stress is generated by exposure to elevated levels of ROS, such as superoxide anion, hydrogen peroxide and the highly toxic hydroxyl radical, which can damage proteins, nucleic acids and cell membranes [Storz and Imlay, 1999]. Bacteria adapt to elevated levels of ROS by increasing the expression of defense and repair proteins, which is regulated by ROS responsive transcription factors. Several regulatory factors participate in controlling this complex thiol-specific stress response to electrophilic compounds in B. subtilis, such as Spx, CtsR, PerR, CymR, and novel MarR-type repressors MhqR (YkvE) and

Introduction 6 Thiol-specific stress response

YodB [Leichert et al., 2003; Nakano et al., 2003b; Zuber, 2004; Tam et al., 2006b; Leelakriangsak et al., 2007; Töwe et al., 2007].

3.1. Spx-an activator of genes involved in the maintenance of the thiol-disulfide balance

Spx was originally discovered as suppressor of clpP and clpX mutations, formerly YjbD. Spx is one central regulator of the thiol-specific stress response in B. subtilis which controls genes required for maintenance of the thiol-disulfide homeostasis. For example, Spx is required for the transcriptional activation of thioredoxin (trxA) and thioredoxin reductase (trxB) genes in response to diamide stress [Nakano et al., 2003b; 2005; Zuber, 2004, Smits et al., 2005]. In addition to its role in the regulation of the thiol-disulfide homeostasis, recent studies indicate that Spx functions as a regulator of gene expression in cells undergoing steady-state growth. Spx was found to negatively affect the transcription of genes that function in the utilization of organosulfur compounds as alternative sources of sulfur [Erwin et al., 2005]. Spx, an arsenate reductase (ArsC) family protein, is highly conserved and involved either positively or negatively in various cellular functions in low-GC-content Gram- positive bacteria [Nakano et al., 2001; 2002; Zuber, 2004; Chatterjee et al., 2006; Pamp et al., 2006]. For example, Spx has also been implicated as a regulatory factor for virulence-related functions in Staphylococcus aureus [Pamp et al., 2006] and Listeria monocytogenes [Chatterjee et al., 2006].

In B. subtilis, Spx concentration and activity increase when cells encounter thiol- specific oxidative stress conditions [Nakano et al., 2003b; 2005]. Increases in Spx protein concentration are associated with a variety of stress conditions including heat shock, salt, disulfide, oxidative stress and toxic aromatic compounds [Nakano et al., 2003b; Tam et al., 2006a; 2006b; Duy et al., 2007a; 2007b]. The increase in Spx activity in cells encountering thiol-specific oxidative stress is due in part to a post-transcriptional mechanism of spx control resulting in an increase in Spx concentration [Nakano et al., 2003b]. Regulation by Spx requires interaction with the C-terminal domain of the RNA polymerase α subunit [Nakano et al., 2003a; 2003b]. The spx gene resides in the yjbC-spx dicistronic operon. Transcription of the yjbC-spx operon is driven by five promoters, three (P1, P2, and PB) reside upstream of yjbC and the PM and P3 promoters are located in the yjbC-spx intergenic region. The control of spx expression in response to oxidative stress operates at several levels. Spx protein levels increase independently of spx transcriptional control by the downregulation of ATP-dependent ClpXP-catalyzed proteolysis [Nakano et al., 2002, 2003a; 2003b; Zhang and Zuber, 2007]. Spx activity is under redox control through its CxxC disulfide center [Nakano et al., 2005]. In addition, transcription from the newly discovered P3 promoter of spx is induced by thiol- reactive agents [Leelakriangsak and Zuber, 2007]. The P3 promoter was shown to depend on two transcriptional regulators that target the cis-acting negative control elements of the promoter, the peroxide regulon repressor PerR and the novel MarR-type/DUF24 family

Introduction 7 Thiol-specific stress response repressor YodB [Leelakriangsak et al., 2007; Leelakriangsak and Zuber, 2007]. PerR was previously characterized as a repressor of the peroxide stress regulon [Herbig and Helmann, 2001; Lee and Helmann, 2006a; 2006b]. YodB also controls the divergently transcribed yodC gene, which encodes a nitroreductase and is induced in response to thiol-specific stress conditions [Duy et al., 2007b; Leelakriangsak et al., 2007]. YodB protects a region that includes the P3 –10 and –35 regions, while PerR binds to a region downstream of the P3 transcriptional start site [Leelakriangsak et al., 2007]. The detailed regulatory mechanism of the YodB-repressor was studied in collaboration with the group of Peter Zuber (Oregon), which was also a subject of this thesis.

3.2. CymR-the repressor of the cysteine metabolism regulon

The Cysteine metabolism repressor CymR is a master regulator of sulfur metabolism. The CymR repressor exerts global control over a number of sulfur metabolism operons and is responsible for their repression when the preferred sulfur sources, sulfate and cysteine, are present. CymR appears as a repressor controlling several pathways leading to cysteine formation, including the O-acetylserine-thiol-lyase (CysK), L-cystine transporters (TcyP and TcyJKLMN), sulfonate assimilation (SsuABCD) and the methionine-to-cysteine conversion pathway (YrhA and YrhB). In addition, CymR represses the expression of the ytlI and ydbM genes, and the yxeK, ytmI, and yrrT operons. CymR binds to the promoter regions of these genes, except in the case of ytmI, which is indirectly controlled by CymR via YtlI. O- acetylserine or its spontaneous derivative N-acetylserine modulates CymR dependent binding [Even et al., 2006]. Considering the set of controlled genes, the CymR repressor can be seen as a functional equivalent of the LysR-type activator CysB from E. coli [Kredich, 1992].

Previous studies have established the link between sulfur metabolism and oxidative stress [Leichert et al., 2003; Mostertz et al., 2004; Smits et al., 2005]. A higher expression of several genes involved in sulfur metabolism is observed in a strain depleted of thioredoxin A [Smits et al., 2005; Mostertz et al., 2008] and in response to superoxide and disulfide stresses [Leichert et al., 2003; Mostertz et al., 2004]. In particular, most of the cysteine biosynthesis genes repressed by CymR are induced under these conditions. Cysteine itself has been proposed to be involved in the oxidative stress response in Staphylococcus aureus [Lithgow et al., 2004] and B. subtilis [Duy et al., 2007b]. Free cysteine serves as LMW thiol in B. subtilis and protects protein thiols against irreversible overoxidations, which was shown by the identification of S-cysteinylated proteins in response to diamide treatment [Hochgräfe et al., 2007]. Thus, the derepression of the CymR regulated genes involved in cysteine biosynthesis in response to oxidative is caused by the depletion of the LMW thiol cysteine resulting in an imbalanced thiol-redox state of the cell. As shown recently using metabolomic approaches, cysteine is also depleted after treatment with quinones which is based on the thiol-(S)- alkylation reaction [Liebeke et al., 2008]. Moreover, it could be shown that toxic

Introduction 8 Thiol-specific stress response concentrations of quinones deplete also thiol-containing proteins due to the formation of irreversibly damaged quinone-protein-aggregates, which are not accessible for the 2D gel separation.

3.3. HrcA and CtsR-heat-shock-specific regulators

The adaptation to heat stress is one of the best-characterized physical stress responses of mesophilic bacteria. In B. subtilis, the heat shock response is governed by HrcA, σB, and CtsR [Hecker and Völker, 2001; Helmann et al., 2001]. Of these, the HrcA and CtsR regulons are also induced under thiol-stress specific stress conditions [Tam et al., 2006b; Duy et al., 2007a; 2007b; Antelmann et al., 2008]. The HrcA protein is a transcriptional repressor of class I heat shock genes and regulates the expression of two operons, the complex dnaK operon as well as the groEL-groES operon [Schmidt et al., 1992; Homuth et al., 1997]. This repressor binds to conserved cis-acting regulatory sequences known as CIRCE (Controlling Inverted Repeat of Chaperone Expression) elements [Zuber and Schumann, 1994] and responds specifically to heat induction. The CtsR (Class three stress gene repressor) repressor negatively regulates the expression of class III heat shock genes (clpP, clpE and the clpC operon) by binding to a directly repeated heptanucleotide operator sequence [Helmann et al., 2001; Miethke et al., 2006]. The clp genes encode ATPase subunits (ClpA, ClpB, ClpC, ClpE, ClpX) and a proteolytic subunit, ClpP. ATPase subunits can act alone as molecular chaperones [Wawrzynow et al., 1996] but also associate with the ClpP proteolytic subunit to form an ATP-dependent protease whose substrate specificity is conferred by the ATPase [Gottesman et al., 1997]. The ClpP protease was shown to act in global protein quality control. ClpP is involved in development of competence, motility, thermotolerance, degradative enzyme synthesis and sporulation [Kock et al., 2004]. The ClpC ATPase is the interacting partner of ClpP in many physiologically important processes such as competence gene expression or adaptation to diverse stress conditions [Krüger et al., 1994]. While the importance of ClpC and ClpP has been elucidated for a wide range of cellular adaptation processes, ClpE is essential both for efficient CtsR dependent gene derepression and for derepression during heat stress [Miethke et al., 2006]. Regulation of CtsR activity is controlled by a fine-tuned phosphorylation mechanism including the modulators McsB and McsA [Kirstein et al., 2005]. The HrcA and CtsR regulons were also strongly induced by phenol, chromanon, catechol and MHQ which indicates protein damage by these compounds [Tam et al., 2006b; Duy et al., 2007b].

3.4. PerR-the main regulator of the oxidative stress response

In B. subtilis, protection against oxidative stress is primarily mediated by the induction of oxidative stress-specific proteins, which are controlled by the repressor protein PerR [Chen et al., 1995a; Bsat et al., 1998]. The metalloprotein PerR was identified in the John Helmann's laboratory as a member of the Fur family of metal-dependent regulators [Fuangthong and

Introduction 9 Thiol-specific stress response

Helmann, 2003]. PerR is the major regulator of the peroxide stress response. In B. subtilis, PerR is a dimeric, Zn2+-containing metalloprotein with a regulatory metal-binding site that binds Fe2+ (PerR:Zn,Fe) or Mn2+ (PerR:Zn,Mn) [Lee and Helmann, 2006b]. PerR and PerR- like regulators have been identified in many bacteria including Campylobacter jejuni [van Vliet et al., 1999], Streptomyces reticuli [Ortiz de Orué Lucana and Schrempf, 2000], Streptomyces coelicolor [Hahn et al., 2000], Streptococcus pyogenes [King et al., 2000], Staphylococcus aureus [Horsburgh et al., 2001] and Borrelia burgdorferi [Boylan et al., 2003].

Cells respond to ROS by the induction of a variety of protective enzymes and proteins.

For example, superoxide anions are converted to H2O2 and O2 by superoxide dismutase, whereas catalase, GSH peroxidases and peroxiredoxins reduce and detoxify peroxides. Thiol- disulfide reductases (thioredoxin and glutaredoxin) reduce disulfide bonds within proteins and oxidized LMW thiols. Molecular chaperones are stimulated to mediate the refolding of unfolded and aggregated proteins [Barford, 2004]. Most peroxide regulators (e.g. OxyR and Hsp33) are redox-controlled and use reactive cysteines to sense peroxides which often lead to the formation of either intra- or intermolecular disulfides [Paget and Buttner, 2003; Kiley and Storz, 2004]. The redox-sensitive transcription factor of E. coli OxyR mediates the cellular response to both ROS and reactive nitrogen species (RNS) by controlling the expression of the OxyR regulon, which encodes proteins involved in peroxide metabolism and in maintenance of the thiol redox balance of the cell [Paget and Buttner, 2003]. Redox-reactive cysteine residues in OxyR are required for the reversible induction of gene expression by ROS [Storz et al., 1990a]. Cys199 and Cys208 both were strongly implicated in the ROS response and Cys199 serves as the specific redox sensor of ROS and RNS [Kullik et al., 1995; Barford, 2004]. The E. coli Hsp33 responds to oxidative stress with the rapid activation of its chaperone function [Jakob et al., 1999]. The redox-sensitive cysteine residues are located within a C-terminal cysteine cluster that forms a high-affinity Zn2+-binding motif. The four cysteines of this motif are structurally invariant and are maintained as thiolate species by coordinating Zn2+ with high affinity [Jakob et al., 2000]. Under oxidizing conditions, the cysteine cluster releases Zn2+ and two intramolecular disulfide bonds are formed, with concomitant dimerization of oxidized Hsp33 monomers [Graumann et al., 2001].

In contrast, PerR does not appear to use thiol disulfide chemistry to sense ROS despite the presence of four conserved cysteine residues in the Zn2+-binding domain [Lee and

Helmann, 2006b]. The Helmann group showed that PerR senses H2O2 by metal-catalyzed histidine oxidation. PerR oxidation, mediated by a bound ferrous ion, leads to the incorporation of one oxygen atom into His-37 or His-91, two of the residues that are supposed to co-ordinate the bound Fe2+. This oxidation would lead to iron release and dissociation of 2+ the PerR-DNA adduct. The proposed mechanism relies on the reduction of H2O2 by Fe and the generation of highly reactive hydroxyl radicals (Fenton reaction). By analogy to metal-

Introduction 10 Thiol-specific stress response catalyzed oxidative reactions with histidine-containing enzymes and model peptides [Schoneich, 2000], it was shown that the hydroxyl radical reacts with histidine and generates the 2-oxo-histidinyl radical intermediate, which finally leads to the 2-oxo-histidine. However, PerR responds also to thiol-specific stress conditions such as diamide and quinone-like electrophiles, which deplete protein thiols. Thus, there could be a different mechanism of PerR to sense electrophilic compounds, which most probably involves the conserved Cys residues in the Zn2+-binding domain.

3.5. The MarR-type regulators

In addition to the above mentioned thiol-stress-specific regulons, the recently identified thiol-stress-responsive MarR-type repressors YodB and MhqR (Methylhydroquinone specific repressor, formerly YkvE) were characterized in B. subtilis in this study [Leelakriangsak et al., 2007; 2008; Töwe et al., 2007]. Both YodB and MhqR regulons are induced under thiol-stress conditions and confer resistance to quinone electrophiles by the control of paralogous detoxification enzymes consisting of quinone reductases and thiol-dependent dioxygenases [Tam et al., 2006b; Duy et al., 2007b; Leelakriangsak et al., 2008; Antelmann et al., 2008]. The YodB repressor negatively controls Spx, the nitroreductase YodC and a FMN-dependent NADH-azoreductase AzoR1 in response to thiol-stress condition [Duy et al., 2007b; Leelakriangsak et al., 2007; 2008]. The MhqR repressor negatively controls three thiol-dependent dioxygenases (MhqA, MhqO and MhqE), the nitroreductase MhqN and the azoreductase YvaB (AzoR2) in response to the thiol-reactive electrophiles quinone and diamide [Töwe et al., 2007].

Both YodB and MhqR regulators are members of the MarR family of winged-helix transcription factors that regulate a variety of biological functions in bacteria [Wilkinson and Grove, 2006]. MarR family proteins regulate the expression of resistance genes to multiple antibiotics, aromatic compounds, organic solvents, or oxidative stress agents. Most of these MarR family members are repressors of gene expression [Tropel and van der Meer, 2004]. However, in Rhodopseudomonas the MarR type regulator BadR was shown to active transcription of the benzoate degradation pathway [Egland and Harwood, 1999]. In B. subtilis, the most-studied MarR family repressor is the organic hydroperoxide resistance repressor OhrR that represses ohrA by binding to a 15 bp inverted repeat overlapping the promoter region [Fuangthong et al., 2001; Fuangthong and Helmann, 2002; Hong et al., 2005]. Recent studies have shown that a complex thiolate switch regulates OhrR activity in the presence of organic hydroperoxides. Inactivation of OhrR requires Cys15 oxidation, which leads to either a sulfenamide or a mixed disulfide (S-thiolated) form of OhrR in the presence of cumene hydroperoxide [Lee et al., 2007]. In addition, overoxidation of Cys15 to sulfonic acid leads to relief of OhrR-mediated repression in the presence of the strong oxidant linoleic acid [Soonsanga et al., 2008a].

Introduction 11 Thiol-specific stress response

YodB is highly conserved among low GC Gram-positive bacteria, and there are at several YodB paralogs in the genome of B. subtilis. YodB-like regulators share the conserved N-terminal cysteine in position 5 or 6 followed by a conserved proline residue. The YodB protein contains three cysteine residues, at position 6 (C6), 101 (C101) and 108 (C108). Unlike the two C-terminal Cys residues, C6 is conserved among YodB homologues in low GC Gram-positive bacteria. Cys residues have been shown to be involved in the redox-control of other oxidative stress transcription factors, such as OxyR, Hsp33 of E. coli and OhrR of B. subtilis [Graumann et al., 2001; Hong et al., 2005; Nakano et al., 2005; Lee et al., 2007]. Cysteine residues are strong nucleophiles, which are shown to be conjugated with electrophilic compounds such as p-benzoquinone and hydroquinones to form thiol-S-adducts [Giles et al., 2003; Dennehy et al., 2006]. One cysteine residue is present in MhqR (C128), which is located in the dimerization domain and accessible for thiol-modification according to published structure for B. subtilis OhrR and E. coli MarR [Alekshun et al., 2001; Hong et al., 2005]. In this study, it was shown in collaboration with the group of Peter Zuber that the conserved Cys6 residue of YodB is required for optimal YodB-mediated repression in vivo and in vitro [Leelakriangsak et al., 2008]. In addition, our studies with the YodB-paralogous YvaP-repressor showed that Cys7 is also the sensing cysteine residue and required for repression of the catDE operon. Mass spectrometry analyses identified the formation of Cys6- S-adducts upon treatment of YodB with the thiol-reactive compounds MHQ and catechol in vitro leading to the loss of DNA binding activity of YodB [Leelakriangsak et al., 2008]. Thus, YodB and YvaP are both directly sensing thiol-reactive compounds via the conserved C6 and Cys7 residues, since DNA binding activity is directly inhibited by the thiol-reactive compounds MHQ and diamide in vitro. The reactivity of the cysteines is determined by neighboring basic amino acids which lower the cysteine pKa to generate the reactive cysteine thiolate anion. Stabilization of the active site thiolate anion by neighboring basic amino acids has been shown for the oxidative stress responsive regulators OxyR of E. coli and OhrR of B. subtilis [Choi et al., 2001; Soonsanga et al., 2007]. Moreover, it has been shown also that the thiol-(S)-alkylation reaction via the Michael addition chemistry is favored by adjacent arginine and lysine residues [Dennehy et al., 2006]. In the present study, the basic K19 and R20 amino acid residues are also essential for YvaP repression in vivo.

4. The response of bacteria to the electrophile methylglyoxal (MG)

Another objective of this thesis was further to investigate the global expression profile of B. subtilis in response to natural occurring electrophiles. Thus, we have selected methylglyoxal (MG) since this is as one of the best studied natural electrophiles in enteric bacteria. MG is also referred as pyruvaldehyde, pyruvic aldehyde, 2-oxopropanal, 2- ketoproprion-aldehyde and acetyl-formaldehyde, which is a yellow liquid [Kalapos, 1999]. MG is a naturally occurring toxic electrophile that at high concentrations can cause cell death. MG is produced in all cells, not just bacteria, by different mechanisms (Fig. 1.1) when an

Introduction 12 Thiol-specific stress response imbalance occurs in the metabolism [Murata et al., 1989; Kalapos, 1999; Booth et al., 2003]. MG, a glycolytic intermediate, is produced from triose-phosphate intermediates of phosphorylating glycolysis [Phillips and Thornalley, 1993]. MG is also formed by lipid peroxidation systems [Poli et al., 1985], in the metabolism of acetone [Reichard et al., 1986] and aminoacetone [Lyles and Chalmers, 1992], and during the degradation of DNA [Chaplen et al., 1996]. In addition, MG is ubiquitous in beverages and foods, such as coffee and soy sauce [Kasai et al., 1982; Nagao et al., 1986]. MG synthesis is mediated by enzymes including MG synthase, cytochrome P450 and amine oxidase, which are involved in glycolytic bypass, acetone metabolism and amino acid breakdown, respectively [Kalapos, 1999]. In eukaryotic cells, MG is also generated by nonenzymatic fragmentation of dihydroxyacetone phosphate or glyceraldehyde 3-phophate [Phillips and Thornalley, 1993].

Fig. 1.1. Pathways of MG generation and detoxification. Abbreviations used in the figure: G-6P, glucose 6- phosphate; F-6P, fructose 6-phosphate; F-1,6P2, fructose 1,6-bisphosphate; GA-3P, glyceraldehyde 3-phosphate; DHA-P, dihydroxyacetone-phosphate; Gly-P, glycerolphosphate; GSH, glutathione; PD, propanediol. The enzymes involved in the reactions: (i) methylglyoxal synthase; (ii) after a non-enzymatic reaction between methylglyoxal and GSH glyoxalase I; (iii) glyoxalase II; (iv) ∗-oxoaldehyde dehydrogenase; (v) e.g. methylglyoxal reductase; (vi) D-lactate dehydrogenase; (vii) L-lactate dehydrogenase; (viii) the enzymes of phosphorylating glycolysis; (ix) e.g. lactaldehyde dehydrogenase; (x) acetoacetate decarboxylase and spontaneously, too; (xi) acetone monooxygenase; (xii) acetol monooxygenase; (xiii) amine oxidase. Cofactors are not shown in the flow-chart. Dotted lines represent the ways of non-enzymatic methylglyoxal formation. This figure was adapted from Kalapos, 1999.

The role of MG in cell physiology has been studied since many years. Although known as growth regulator [Együd and Szent-Györgyi, 1966] and to act as a bypass pathway for glycolysis [Hopper and Cooper, 1971], MG is a very toxic compound. It can react with the nucleophilic centers of macromolecules such as DNA and RNA. Moreover, MG inhibits growth by interfering with protein synthesis and consequently prevents initiation of DNA replication [Fraval and McBrien, 1980; Lo et al., 1994]. In addition, MG can exert cytotoxic,

Introduction 13 Thiol-specific stress response cytostatic and mutagenic effects in cells [Inoue and Kimura, 1995; Thornalley, 1996; Kalapos, 1999; Booth et al., 2003].

Furthermore, MG modifies irreversibly amino acids in proteins [Lo et al., 1994]. It has been reported that MG binds and modifies a number of proteins, including numerous intra- and extra-cellular membrane proteins [Shipanova et al., 1997]. Proteins that are shown to be modified by MG are collagen [Bowes and Cater, 1968], ribonuclease [Brock et al., 2007], lysozyme [Kato et al., 1986], bovine serum albumin [Westwood et al., 1994], heat and shock protein 27 (Hsp27) [Sakamoto et al., 2002]. MG modifies proteins through the Maillard reaction, resulting in several characteristic advanced glycation, glycoxidation and lipoxidation end products (AGE/ALEs) which can alter protein structure and functions. MG reacts readily with the basic amino acid side chains of arginine and lysine residues [Gomes et al., 2006; Brock et al., 2007]. MG primarily reacts with arginine residues (via guanidino group) to form Nδ-(5-methyl-4-imidazolon-2-yl)-L-ornithine (5-methylimidazolone) and Nδ-(5-hydro-5- methyl-4-imidazolon-2-yl)-L-ornithine (5-hydro-5-methylimidazolone) [Thornalley, 1996; Uchida et al., 1997; Westwood et al., 1997]. MG reacts with lysine residues (via e-amino group) to generate an MG-derived lysine-lysine cross-link (imidazolysine) and Nε- carboxyethyllysine (CEL) [Nagaraj et al., 1996; Uchida et al., 1997; Westwood et al., 1997]. MG also reacts with the nucleophilic thiol group of cysteine to form hemithioacetal [Thornalley, 1996]. MG modification of proteins and amino acids causes cell damage and is the cause of cellular toxicity [Thornalley, 1996; Oya et al., 1999]. Hipkiss and Chana (1998) showed that carnosine can protect proteins against MG toxicity. Carnosine detoxifies MG thereby inhibiting MG-mediated protein modification.

Because of the cytotoxicity of MG, both eukaryotic and prokaryotic cells have evolved specific detoxification systems and protective mechanisms [Thornalley, 1996; Kalapos, 1999; Booth et al., 2003]. There are two lines of protection from MG toxicity [Kalapos, 1999]. The direct mechanism operates through the glyoxalase system thus detoxifying the electrophile, while an indirect mechanism is also operative by the acidification of the cytoplasm [Ferguson et al., 1995]. In the majority of prokaryotic and eukaryotic cells, the major mechanisms for detoxifying MG is the GSH-dependent glyoxalase I-II system (Fig. 1.2), a ubiquitous thiol- dependent detoxification pathway with D-lactate as the final product [Thornalley, 1990; 1996; 2003; Inoue and Kimura, 1995; Kalapos, 1999]. The glyoxalase system consists of two ubiquitous metalloenzymes and requires catalytic amounts of reduced GSH [Racker, 1951]. The metalloenzymes glyoxalase I (S-lactoylglutathione MG lyase) and glyoxalase II (S-2- hydroxyacylglutathione hydrolase) catalyze the step-wise dismutation of MG into the corresponding 2-hydroxyacids (D-lactate), using GSH as a cofactor [Thornalley, 1990]. Glyoxalase I acts on the adduct of MG and GSH, thus catalyzing the substrate isomeration of hemithioacetal into S-D-lactoylglutathione [Mannervik and Ridderström, 1993]. Subsequently, glyoxalase II converts S-D-lactoylglutathione into D-lactate and GSH [Vander

Introduction 14 Thiol-specific stress response

Jagt, 1993]. Glyoxalase I and II possess mononuclear and binuclear active sites, respectively. These glyoxalase enzymes have been characterized from bacteria to man, revealing largely conserved enzymological and structural characteristics [Thornalley, 2003]. To date, glyoxalase I can be roughly sub-divided into two different groups according to the type of divalent cation bound at the active site [Clugston et al., 1998; Sukdeo et al., 2004]. The first class is Zn2+-dependent and is composed of glyoxalase I from mainly eukaryotic organisms. For example, glyoxalase I from human and yeast [Aronsson et al., 1978] and Plasmodium falciparum [Iozef et al., 2003] prefers Zn2+. The second class is composed of non-Zn2+- dependent (but Ni2+ or Co2+-dependent) glyoxalase I enzymes (mainly prokaryotic and leishmanial species). For example, glyoxalase I from several bacteria such as E. coli [Clugston et al., 1998], Yersinia pestis, Pseudomonas aeruginosa and Neisseria meningitidis [Sukdeo et al., 2004] and from the protozoan parasite Leishmania major [Ariza et al., 2005] is optimally activated in the presence of Ni2+. Glyoxalase II is typically Zn2+-activated, containing Zn2+ and either Fe3+/Fe2+ or Mn2+ at the active site depending on the biological source [O’Young et al., 2007]. In addition, some organisms use another LMW thiol instead of GSH. For example, in the trypanosomatid parasite Leishmania major, trypanothione replaces GSH in the defense against cellular damage caused by MG [Fairlamb et al., 1985]. In this protozoon, the first step in MG detoxification is performed by a trypanothione-dependent glyoxalase I containing a Ni2+ cofactor [Ariza et al., 2006].

Fig. 1.2. MG detoxification via the glyoxalase system. MG reacts spontaneously with GSH to form hemithiolacetal, followed by isomerization to S-D-lactoylglutathione by the glyoxalase I enzyme. S-D- lactoylglutathione is substrate for glyoxalase II enzyme which is converted to D-lactate and free GSH. This figure is adapted from Sukdeo et al. (2004).

Introduction 15 Thiol-specific stress response

Besides the GSH-dependent glyoxalase system, there are GSH-independent pathways for MG detoxification [Misra et al., 1995; 1996]. The glyoxalase III catalyses the GSH- independent conversion of MG into D-lactate in E. coli [Misra et al., 1995]. Secondly, the MG reductases convert MG into lactaldehyde [Murata et al., 1989]. Such aldose and aldehyde reductases, mediating the reduction of MG to acetol and D-lactaldehyde have been reported for E. coli, Saccharomyces cerevisiae, plants and mammals [Inoue and Kimura, 1995; Misra et al., 1996]. Important enzymes for reduction of MG are aldo-keto reductases (AKRs), which catalyze the formation of acetol from MG in the presence of NADPH [Kalapos, 1999]. AKRs are a large superfamily of related proteins that catalyze the NADPH-dependent reduction of various aldehydes and ketones [Jez and Penning, 2001; Ellis, 2002]. The AKR superfamily consists of 14 families, based on their structures and sequences [Jez and Penning, 2001]. Although a detailed mechanism has been revealed for some of the AKRs, the physiological function of most putative members of this superfamily is still unclear. This is based in part on their broad substrate specificity and also because of the difficulty of genetic analysis, as many organisms have multiple genes that encode AKRs. For example, E. coli has six AKR genes and yeast has 14 [Goffeau et al., 1996; Blattner et al., 1997]. The number of AKR genes in cyanobacteria varies greatly. Synechocystis sp. PCC 6803, Synechococcus sp. PCC 6301 and Nostoc punctiforme have four, two and 21 potential AKR genes, respectively. There are four potential AKR genes in Synechococcus sp. PCC 7002, based on its genomic sequence [Xu et al., 2006]. Although some mammalian and eukaryotic AKRs have been characterized, only for a small subset of bacterial AKRs substrate specificities have been analyzed [Ellis, 2002]. For example, Ko et al. (2005) recently showed that AKR plays an important role in vivo in MG detoxification in E. coli. YafB and YqhE have been characterized as 2,5-diketo-gluconate (2,5-DKG) reductases, while E. coli YghZ has been shown to reduce MG and enhance MG resistance when the gene is overproduced. Furthermore, the α-oxoaldehyde dehydrogenase (2- oxoaldehyde:NAD(P)+ oxidoreductase) is another enzyme involved in MG degradation [Inoue and Kimura, 1995].

To date, survival strategies in response to MG challenge is best understood in E. coli which is explained now in more detail. To ensure its survival, the E. coli cell has developed severe protective mechanisms against electrophiles. In E. coli, the GSH-based detoxification pathway is central to survival against MG toxicity. There are at least two levels of GSH- dependent protection against MG: the activation of GSH-gated K+ channels and the detoxification by reaction of MG with GSH, followed by the action of glyoxalase I and II to produce D-lactate and free GSH. These two mechanisms are linked, as the product of glyoxalase I (S-lactoylglutathione) is the major activator of the KefB/KefC system [MacLean et al., 1998]. The activation of the KefB/KefC system results in the rapid loss of potassium from the cell accompanied by a decrease in the intracellular pH (pHi) [Ferguson et al., 1995; Ferguson and Booth, 1998]. The decrease in the pHi protects E. coli cells against the toxic effects of MG. The integration of the glyoxalase system with activation of KefB means that

Introduction 16 Thiol-specific stress response the cell has a “detoxification meter”-the glyoxalase I enzyme generates the activator of the KefB/KefC system and removes its inhibitor, and the glyoxalase II enzyme removes the activator and regenerates the inhibitor. Thus, the pH of the cell during exposure to MG accurately reflects the balance of exposure to MG and the rate of detoxification [Booth et al., 2003].

In contrast to Gram-negative bacteria, little is known about the mechanisms of MG detoxification in Gram-positive bacteria. B. subtilis and other Gram-positive bacteria lack GSH as their major LMW thiol and KefB/KefC activities which have been shown to protect E. coli against MG toxicity [Ferguson et al., 1995]. Bacillus stearothermophilus possesses MG synthase and appears to have evolved or acquired the glyoxalase I-II pathways [Burke and Tempest, 1990]. Enzymes with homologies to glyoxalases are encoded also in the genome of B. subtilis, but the physiological importance is still unknown. Thus, the MG- protective mechanisms remain to be identified in B. subtilis [Ferguson et al., 1998]. However, our transcriptome results in response to quinone-like electrophiles, MG and diamide suggest that metal ion efflux systems (e.g. ArsR, CsoR) could play a role in extrusion of electrophiles in B. subtilis since these are strongly derepressed under thiol-stress conditions provoked by electrophiles [Antelmann et al., 2008]. These could play similar roles as the E. coli KefB/KefC systems in protection against MG toxicity in B. subtilis.

Recently, the role of free radicals in the biochemistry of MG receives attention. The possible involvement of free radicals in reactions between MG and macromolecules was raised by Szent-Györgyi (1968), especially the interaction between protein amines and MG [Kon and Szent-Györgyi, 1973]. MG leads also to production of RNS and organic radicals such as MG radical or cross-linked protein radical which causes protein damage [Kalapos, 2008].

5. The response of bacteria to the electrophile formaldehyde (FA)

Since commercially available MG is often contaminated with up to 30% formaldehyde (FA), we studied also the response of B. subtilis to FA as another example for reactive aldehydes. Reduced one-carbon (C1) compounds, such as methane, methanol and FA, are ubiquitously present in the natural environment. C1 microorganisms, including methanotrophic and methylotrophic bacteria, possess metabolic pathways for detoxification and fixation of these C1-compounds. Because FA is located at the branch point between the assimilation and dissimilation pathways during C1 metabolism in these organisms [Anthony,

1982], it is the key intermediate in the metabolism of C1 compounds. FA is produced from methanol by NAD-dependent methanol dehydrogenase (MDH) in the first stage of the utilization of carbon and energy sources by methylotrophs. Besides the methanol oxidation pathway, FA is also synthesized from methylamine in several species of methylotrophs, which are able to utilize methylamine as the sole source of carbon and energy [Kelly et al., 1993]. In

Introduction 17 Thiol-specific stress response other organisms, FA is produced through the degradation of organic compounds that have methyl or methoxyl groups, e.g. lignin, pectin, etc. [Yurimoto et al., 2005]. FA is a subproduct in the metabolism of histidine, choline and of a number of plant-derived methoxylated aromatic chemicals such as vanillate, veratrate and caffeate in certain microorganisms [Mitsui et al., 2000]. FA has also been found as a subproduct of the metabolism of certain xenobiotic compounds such as the explosive hexahydro-1,3-5-trinitro- 1,3,5-triaine (RDX), glyphosate (N-phosphonomethylglycine), atropine and methenamine, a compound used to treat urnary track infections [Musher and Griffith, 1974; Shinabarger and Braymer, 1986; Long et al., 1997; Fournier et al., 2004].

FA is also an important disinfectant in holding-tank sewages, particularly at alkaline pHs [Sobsey et al., 1974] and is widely used to sterilize laboratory rooms and working surfaces [Taylor et al., 1969]. In addition, FA is widely employed in the biochemistry, biomedicine and pharmacy. In particular, FA is used for isotope-labeling of proteins [Jentoft and Dearborn, 1983]. Most importantly, FA is used as cross-linker in the study of protein- protein or protein-DNA interactions [Kunkel et al., 1981]. FA can inactivate, stabilize or immobilize DNA and proteins. FA reacts with the amino group of the N-terminal amino acid residue and the side-chains of arginine, cysteine, histidine, tyrosine, tryptophan, asparagine, glutamine and lysine (Fig. 1.3) [Orlando et al., 1997; Metz et al., 2004]. FA treatment results in a large variety of chemical modifications in proteins, such as the formation of methylol groups, Schiff’s bases and methylene bridges. However, the formation of modifications depends on various factors, such as the rate of a particular cross-link reaction, the position and local environment of each reactive amino acid in the protein, the pH, the components present in the reaction solution and the reactant concentrations. The conversion of peptides after FA treatment might predict the reaction rate of FA-induced intramolecular cross-links. Especially, intramolecular cross-links in proteins are initiated by the reaction of FA with lysine residues [Metz et al., 2004].

FA is a highly reactive chemical that exerts toxic effect in all organisms from bacteria to humans, due to its nonspecific reactivity with biological macromolecules such as DNA and protein [Feldman, 1973; Vorholt et al., 2000; Liu et al., 2006]. Early studies by Neely (1963a; 1963b; 1963c) revealed that FA has bacteriostatic activity at sublethal concentrations due to its ability to disrupt growth and interfere with methionine biosynthesis. At higher concentrations, FA is a potent antimicrobial agent and has been used to inactivate Bacillus anthracis and B. subtilis spores [Manchee et al., 1994; Sagripanti and Bonifacino, 1996].

Since FA is toxic in all organisms, different mechanisms have evolved to counteract its toxicity. These mechanisms involve fixation through a number of metabolic pathways and detoxification via oxidation to formate and CO2. Four different pathways for FA

Introduction 18 Thiol-specific stress response detoxification are known in bacteria, encoded by unrelated or distantly related genes, which will be briefly described in the following part.

Fig. 1.3. Chemical cross-linking of DNA and proteins by FA. FA is a very reactive dipolar compound in which the carbon atom is the nucleophilic centre. Amino and imino groups of proteins (e.g., the side chains of lysine and arginine) and of nucleic acids (e.g., cytosine) react with FA, leading to the formation of a Schiff’s base (reaction I). This intermediate can react with a second amino group (reaction II) and condenses. (a) FA mediated cross-linking between the side chains of two lysines. (b) Cross-linking between cytosine and lysine. This figure is adapted from Orlando et al. (1997).

The best characterized FA detoxification pathway is the GSH-dependent NADH- linked FA dehydrogenase (GSH-FDH) and formyl-GSH hydrolase (FGH) which are found in both prokaryotes and eukaryotes, such as Burkholderia fungorum (Fig. 1.4) [Harms et al., 1996; Lee et al., 2002; Sanghani et al., 2002a; 2002b; Tanaka et al., 2002; Marx et al., 2004]. These enzymes belong to the zinc-containing medium-chain alcohol dehydrogenase (ADH) family [Persson et al., 1991]. Some FDHs that do not require GSH for activity have been characterized in Pseudomonas putida and Burkholderia fungorum [Tanaka et al., 2002; Marx et al., 2004].

Introduction 19 Thiol-specific stress response

Fig. 1.4. Reactions of the three alternative pathways for FA oxidation in Burkholderia fungorum LB400. FlhA, GSH, NAD-linked FDH; FghA, FGH; FdhA, NAD-linked (GSH-independent) FDH; Fae, FA-activating enzyme; MtdB, NAD(P)-linked methylene-H4MPT dehydrogenase; Mch, methenyl-H4MPT cyclohydrolase; Fhc, formyltransferase-hydrolase complex. This figure is adapted from Marx et al. (2004).

Another important pathway for FA detoxification is the ribulose monophosphate

(RuMP) pathway. This pathway is present in many methylotrophs, which can utilize C1 compounds as a sole source of carbon and energy [Hanson and Hanson, 1996]. This pathway is present also in a number of bacterial and archaeal strains [Yanase et al., 1996; Mitsui et al., 2000; Vorholt et al., 2000; Yurimoto et al., 2002; Kato et al., 2006]. The RuMP pathway is responsible for the detoxification of FA in the non-methylotrophic B. subtilis [Yasueda et al., 1999; Yurimoto et al., 2005] and Burkholderia cepacia [Mitsui et al. 2003]. Reactions of the RuMP pathway are shown in Fig. 1.5 which can be divided into three phases. The FA fixation phase involves two reactions that are unique to the RuMP pathway: the condensation of FA with ribulose 5-phosphate and the subsequent isomerization of the product, D-arabino-3- hexulose 6-phosphate, to yield fructose 6-phosphate. These reactions are catalyzed by 3- hexulose-6-phosphate synthase (HPS) and 6-phospho-3-hexuloisomerase (PHI), respectively. The second phase involves cleavage of the hexose phosphate to form two triose phosphates; glyceraldehyde 3-phosphate (GAP) enters the third phase of FA fixation, which consists of rearrangement reactions, while dihydroxyacetone phosphate (DHAP) enters the central pathway for synthesis of cell constituents. The third phase is the rearrangement phase, which includes the reactions that are necessary to regenerate the acceptor of FA fixation, ribulose 5- phosphate [Mitsui et al., 2000]. Although HPS and PHI have been detected in many methylotrophs [Ferenci et al., 1974; Arfman et al., 1990], the genes encoding these enzymes remain largely unknown. The genes encoding HPS were cloned from an obligate methylotroph, Methylomonas aminofaciens 77a [Yanase et al., 1996], as well as facultative methylotrophs, such as Mycobacterium gastri MB19 [Mitsui et al., 2000]. Neither enzyme was thought to exist outside methylotrophs. However, bioinformatics analysis has suggested that hps and phi gene homologs are widespread in bacteria [Yasueda et al., 1999; Mitsui et

Introduction 20 Thiol-specific stress response al., 2000; Yurimoto et al., 2002; 2005]. In B. subtilis, the pathway for FA fixation involves the hxlAB operon including the hps homologous gene hxlA and the phi homologous gene hxlB [Yasueda et al., 1999; Yurimoto et al., 2005]. Interestingly, the hxlAB operon of B. subtilis is positively regulated by a MarR/DUF24-family regulator HxlR, which shares sequence similarity to the YodB repressor [Yurimoto et al., 2005]. Since the mechanism of HxlR activation has not been resolved so far, it is tempting to speculate that also the conserved N- terminal Cys residue of HxlR plays a role in sensing aldehydes. Furthermore, the results presented in this thesis confirm the thiol-reactive mode of action for FA that further supports the global control mechanism of MarR/DUF24-family regulators via the thiol-(S)-alkylation, which is reflective for quinones and α,β-unsaturated carbonyls such as MG and FA.

Fig. 1.5. The RuMP pathway. The carbon from FA is asterisked. Enzymes: HPS, 3-hexulose-6-phosphate synthase; PHI, 6-phospho-3-hexuloisomerase. This figure is adapted from Kato et al. (2006).

The last pathway for FA detoxification involves "archaeal" enzymes and is linked to the archaeal cofactors tetrahydromethanopterin (H4MPT) and methanofuran (MFR) (Fig. 1.6) [Chistoserdova et al., 1998]. This pathway has been identified only in methylotrophic bacteria and has been characterized in detail in one methylotroph, Methylobacterium extorquens AM1 [Vorholt et al., 1998; 2000; Pomper et al., 1999; 2002; Hagemeier et al., 2000]. The initial step in this pathway is the reaction of FA with H4MPT to form methylene-H4MPT. While this reaction can occur spontaneously, as is apparently the case for H4F, a specific FA-activating enzyme (Fae) has been shown to catalyze this condensation, and this enzyme is required for methylotrophic growth [Vorholt et al., 2000]. The resulting methylene-H4MPT is oxidized to methenyl-H4MPT through the action of one of two methylene-H4MPT dehydrogenases, MtdA

Introduction 21 Thiol-specific stress response

and MtdB. MtdA, which also catalyzes the oxidation of methylene-H4F, is strictly NADP dependent [Vorholt et al., 1998], whereas MtdB can use either NAD+ or NADP+ but is specific for methylene-H4MPT [Hagemeier et al., 2000]. Methenyl-H4MPT is converted to formyl-H4MPT by methenyl-H4MPT cyclohydrolase (Mch) [Pomper et al., 1999]. Finally, formate is released from formyl-H4MPT through the action of the formyltransferase/hydrolase complex (Fhc) [Pomper and Vorholt, 2001; Pomper et al., 2002]. Formate can then be oxidized to CO2 by formate dehydrogenase, as for the H4F-linked pathway.

Fig. 1.6. (A) Tetrahydrofolate (H4F)-mediated C1 transfer pathway found in bacteria, some archaea, and eukaryotes and (B) tetrahydromethanopterin/methanofuran (H4MPT/MFR)-mediated C1 transfer pathway found in methanogenic and sulfate-reducing archaea. MH4F DH, methylene H4F dehydrogenase;

MH4MPT DH, methylene H4MPT dehydrogenase; MH4F CH, methenyl H4F cyclohydrolase; MH4MPT CH, methenyl H4MPT cyclohydrolase; FH4FS, formyl H4F synthase; FMFR:H4MPT FT, formyl MFR:H4MPT formyltransferase; FDH, formate dehydrogenase; FMFR DH, formyl MFR dehydrogenase. The two alternative pathways are proposed to operate in Methylobacterium extorquens AM1, except that MH4MPT DH is NAD(P)- linked. The known genes of Methylobacterium extorquens AM1 for both pathways are shown in italics. The figure is adapted from Chistoserdova et al. (1998).

6. The aim of this part of the thesis

B. subtilis is exposed to different toxic, antimicrobial and aromatic compounds in the soil. In several previous studies, the expression profile of B. subtilis was monitored after exposure to phenol, salicylic acid, catechol, chromanon and MHQ using the transcriptomic and proteomic approaches [Tam et al., 2006b, Duy et al., 2007a; 2007b; Töwe et al., 2007]. The results showed the induction of the thiol-specific stress response by catechol and MHQ that is controlled by the novel MarR-type repressors MhqR (YkvE) and YodB [Leelakriangsak et al., 2007; Töwe et al., 2007]. YodB was identified as novel redox- controlled transcriptional regulator which directly senses thiol-reactive compounds. The

Introduction 22 Thiol-specific stress response regulatory mechanism of YodB which involves the conserved Cys6 residue was one subject of this study, which was performed in collaboration with the group of Peter Zuber (Oregon). Furthermore, the YodB-paralogous repressor YvaP was identified by Kazuo Kobayashi as repressor of the catechol-inducible catDE operon. Both, YodB and YvaP share the conserved Cys6 and Cys7 residues. One further goal of this thesis was to analyze the roles of the conserved Cys7 and of the basic arginine and lysine residues of YvaP for regulation of YvaP DNA binding activity in vivo.

We were further interested in the physiological responses of B. subtilis to the toxic carbonyl electrophiles MG and FA. Thus, the changes in the transcriptome and proteome of B. subtilis after exposure to different concentrations of MG and FA were analyzed in this thesis. The results showed that both MG and FA induced also a thiol-specific stress response in B. subtilis. Thus, we compared the transcriptional profiles for all thiol-reactive compounds, including diamide, quinones and aldehydes more specifically using hierarchical gene clustering. We found further that detoxification of FA involves the novel MerR/NmlR-like activator YraB(AdhR) which controls a conserved thiol-dependent formaldehyde dehydrogenase of B. subtilis (AdhA) and the cysteine proteinase YraA. In collaboration with the group of John Helmann (Ithaca), we studied regulation of AdhR in vivo and in vitro, which involves the conserved Cys52 residue of AdhR. Finally, the phenotypes of mutants in genes of the FA-inducible AdhR and HxlR regulons were investigated in this thesis which showed that both contribute to FA detoxification.

YodB 23 Leelakriangsak et al., 2008

Chapter 1

Regulation of quinone detoxification by the thiol stress sensing DUF24/MarR-like repressor, YodB in Bacillus subtilis

Montira Leelakriangsak1, Nguyen Thi Thu Huyen2, Stefanie Töwe2, Nguyen Van Duy2, Dörte Becher2, Michael Hecker2, Haike Antelmann2 and Peter Zuber1

1Depertment of Environmental and Biomolecular Systems, OGI School of Science and Engineering, Oregon Health and Science University, Beaverton, OR, USA 2 Institute of Microbiology, Ernst-Moritz-Arndt-University of Greifswald, F.-L.-Jahn-Str. 15, D-17487 Greifswald, Germany

YodB was characterized as a novel DUF24/MarR-type repressor that controls expression of the regulator Spx, the nitroreductase YodC and the azoreductase AzoR1 (YocJ). H. Antelmann and S. Töwe carried out proteome and transcriptome analyses. The phenotype analyses were performed by N. V. Duy, which revealed that both azoreductases, AzoR1 and AzoR2 confer resistance to catechol, MHQ, benzoquinone and diamide. H. Antelmann and D. Becher identified YodB-Cys-S-adducts using mass spectrometry. DNase I footprinting analyses of YodB to the azoR1 and spx promoters were carried out by M. Leelakriangsak in the group of P. Zuber. The construction of YodB Cys mutants and functional analysis of the role of the Cys residues in regulation of YodB DNA binding activity using lacZ assays and primer extension analysis was performed by M. Leelakriangsak in the group of P. Zuber. My contribution in this paper is the Northern blot and phenotype analyses of YodB Cys point mutants. These analyses revealed that the conserved Cys6 residue of YodB is required for optimal repression of YodB-controlled target genes while substitution of all three Cys residues (Cys6, Cys101 and Cys108) of YodB affects induction of azoR1 transcription.

This chapter is published in Molecular Microbiology (2008) 67(5): 1108-24.

Molecular Microbiology (2008) 67(5), 1108–1124 ᭿ doi:10.1111/j.1365-2958.2008.06110.x 24 First published online 16 January 2008 Regulation of quinone detoxification by the thiol stress sensing DUF24/MarR-like repressor, YodB in Bacillus subtilis

Montira Leelakriangsak,1 Nguyen Thi Thu Huyen,2 to protect cells against the thiol reactivity of Stefanie Töwe,2 Nguyen van Duy,2 Dörte Becher,2 electrophiles. Michael Hecker,2 Haike Antelmann2* and 1 Peter Zuber ** Introduction 1Department of Environmental and Biomolecular Systems, OGI School of Science and Engineering, The Gram-positive bacterium, Bacillus subtilis is exposed Oregon Health and Science University, Beaverton, OR, to a variety of toxic and antimicrobial compounds in the USA. soil, which induce general and specific stress responses 2Institute of Microbiology, Ernst-Moritz-Arndt-University in growing cells. As part of the stress response, specific of Greifswald, F.-L.-Jahn-Str. 15, D-17487 Greifswald, detoxification systems confer resistance to reactive aro- Germany. matic compounds. Catecholic and hydroquinone-like intermediates are formed within the catabolic pathways of many aromatic compounds (Vaillancourt et al., 2006). Summary They are metabolized by specific ring-cleavage dioxy- Recently, we showed that the MarR-type repressor genases. Two classes of ring-cleavage dioxygenases YkvE (MhqR) regulates multiple dioxygenases/ have been identified based on the mode of ring cleavage: glyoxalases, oxidoreductases and the azoreductase intradiol and extradiol dioxygenases, which cleave the encoding yvaB (azoR2) gene in response to thiol- aromatic ring ortho and meta to the hydroxyl substituents specific stress conditions, such as diamide, catechol respectively. Catechol and hydroquinones are auto- and 2-methylhydroquinone (MHQ). Here we report on oxidized to toxic ortho- and para-benzoquinones in the the regulation of the yocJ (azoR1) gene encoding presence of oxygen. Quinones can exert their toxicity another azoreductase by the novel DUF24/MarR-type as oxidants and/or electrophiles (O’Brian, 1991; Monks repressor, YodB after exposure to thiol-reactive et al., 1992; Kumagai et al., 2002). As oxidants, quinones compounds. DNA binding activity of YodB is directly act as catalysts in the generation of reactive oxygen inhibited by thiol-reactive compounds in vitro. Mass species (ROS). As substrates for cellular reductases, spectrometry identified YodB-Cys-S-adducts that are quinones can undergo one-electron reduction to yield formed upon exposure of YodB to MHQ and catechol semi-quinone radical anions that reduce molecular in vitro. This confirms that catechol and MHQ are oxygen, producing ROS such as superoxide anion, and auto-oxidized to toxic ortho- and para-benzoquinones lead to redox cycling reactions. As electrophiles, quinones which act like diamide as thiol-reactive electrophiles. can form S-adducts with cellular thiols through reactions Mutational analyses further showed that the con- involving a 1,4-reductive Michael-type addition. As ortho- served Cys6 residue of YodB is required for optimal and para-benzoquinones have a one-electron reduction repression in vivo and in vitro while substitution of all potential that far exceeds the range for redox-cycling three Cys residues of YodB affects induction of azoR1 reactions, these are primarily electrophilic (Rodriguez transcription. Finally, phenotype analyses revealed et al., 2005). Quinones are also biologically active that both azoreductases, AzoR1 and AzoR2 confer compounds that function as lipid electron carriers in the resistance to catechol, MHQ, 1,4-benzoquinone and electron-transport chain (e.g. ubiquinone, menadione) diamide. Thus, both azoreductases that are controlled (Unden and Bongaerts, 1997). by different regulatory mechanisms have common Previous proteome and transcriptome analyses functions in quinone and azo-compound reduction showed a strong overlap in the response of B. subtilis to catechol, 2-methylhydroquinone (MHQ) and diamide. Diamide is an electrophilic azo-compound that reacts with Accepted 1 January, 2008. For correspondence. *E-mail antelman@ protein thiolates leading to the formation of disulphides uni-greifswald.de; Tel. (+49) 3834 864237; Fax (+49) 3834 864202. **E-mail [email protected]; Tel. (+1) 503 748 7335; Fax or S-thiolated proteins (Leichert et al., 2003; Hochgräfe (+1) 503 748 1464 et al., 2007). Thus, it seems that diamide, catechol and

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd 25 Regulation and function of the YodB repressor 1109 MHQ are common thiol-reactive electrophiles that can play major roles as quinone reductases, that prevent toxic generate mixed disulphides and Cys-S-adducts that lead quinones from undergoing redox-cycling and causing to depletion of cellular protein thiolates. Several regula- depletion of cellular thiolates. tory factors participate in controlling this complex thiol- specific stress response to electrophilic compounds, such as Spx, CtsR, PerR, CymR and the novel MarR- Results type repressors YodB and MhqR (YkvE) (Leichert et al., Microarray and proteomic analyses identified the 2003; Nakano et al., 2003a; Zuber, 2004; Leelakriang- azoreductase-encoding yocJ (azoR1) gene as most sak et al., 2007; Töwe et al., 2007). The Spx protein of strongly derepressed in DyodB mutant cells B. subtilis is a global transcriptional regulator that con- trols genes whose products function in maintenance of The novel MarR-type repressor, YodB was shown to thiol homeostasis under disulphide stress conditions control the expression of spx and that of the (Nakano et al., 2003a; 2005; Zuber, 2004). Regulation nitroreductase-encoding yodC gene in response to thiol by Spx requires interaction with the C-terminal domain of stress conditions in B. subtilis (Duy et al., 2007; Leelakri- the RNA polymerase a subunit (Nakano et al., 2003a,b). angsak et al., 2007). To identify the genes of the YodB Recent studies showed that spx transcription initiation at regulon, B. subtilis wild-type and DyodB mutant strains the P3 promoter located upstream of spx is induced in were subjected to proteome and transcriptome com- response to diamide treatment (Leelakriangsak and parison. Table 1 lists genes that are at least threefold Zuber, 2007; Leelakriangsak et al., 2007). Two negative derepressed in the DyodB mutant according to the transcriptional regulators, PerR and YodB, were discov- microarray hybridization analysis. The most strongly ered to interact directly with and repress transcription derepressed, YodB-controlled gene was identified as initiation from the spx P3 promoter (Leelakriangsak and yocJ, which encodes a putative FMN-dependent NADH- Zuber, 2007). PerR was previously characterized as a azoreductase and is also referred to as AzoR1 accord- repressor of the peroxide stress regulon (Mongkolsuk ing to Swiss-Prot database annotations (Accession and Helmann, 2002). The novel DUF24/MarR-type No. O35022). The very high derepression ratio of azoR1 family transcriptional regulator YodB also controls the transcription in the yodB mutant could be due to the fact divergently transcribed yodC gene which encodes a that azoR1 is subject to regulation by YodB only, in con- putative nitroreductase and is induced in response to trast to spx. The azoR1 (yocJ) transcriptional start site disulphide stress conditions (Duy et al., 2007; Leelakri- was mapped previously and the concentration of azoR1 angsak et al., 2007). Moreover, three cysteine residues transcript was observed to increase after treatment are present in the YodB protein, which could be the with diamide (Nakano et al., 2003a). In addition, AzoR1 targets for the redox-sensitive control of repressor activ- protein synthesis is strongly increased by diamide, cat- ity that responds to thiol-reactive compounds. Thus, echol, MHQ and nitrofurantoin treatment as shown by YodB might function as a thiol-responsive regulator of proteomic analysis (Fig. 1, Table 1). However, AzoR1 B. subtilis. Recently, we have shown that another MarR- protein synthesis was not induced in response to oxida- type repressor MhqR (YkvE) controls three putative tive stress provoked by the addition of H2O2 (Fig. 1). The dioxygenases/glyoxalases (MhqA, MhqO and MhqE), proteomic analyses verified the derepression of azoR1 the nitroreductase MhqN and the azoreductase YvaB and yodC in the DyodB mutant (Fig. 1). The proteomic (AzoR2) in response to thiol-reactive electrophiles. data also confirmed that AzoR1 production is several- These confer resistance to MHQ and catechol in fold more elevated than YodC in the DyodB mutant B. subtilis (Töwe et al., 2007). (Fig. 1, Table 1). Recently, we showed that production of In this study, we have identified a paralogue of azoR2, the paralogous azoreductase, AzoR2 (YvaB), is induced the azoreductase-encoding azoR1 (yocJ ) gene, which is under thiol stress conditions and azoR2 expression is the most strongly derepressed member of the YodB regulated the MarR-type repressor, MhqR (YkvE) (Töwe regulon under thiol-specific stress conditions. Mutational et al., 2007) (Fig. 1). However, in contrast to AzoR1, analyses indicate that the Cys residues of YodB play a AzoR2 protein synthesis was not induced after diamide role in repressor activity and the repressor’s response to or nitrofurantoin stress (Fig. 1). Interestingly, synthesis of inducing phenolic compounds. The thiol-specific quinone MhqR-regulated azoR2 expression was reduced in the reactivity of oxidized MHQ and catechol was confirmed in DyodB mutant and, vice versa, the level of the YodB- vitro using mass spectrometry. Finally, we show that the controlled azoR1 and yodC expression was lower in the paralogous enzymes, AzoR1 and AzoR2 that are regu- DmhqR mutant under control conditions (Fig. 1). In con- lated by YodB and MhqR both contribute to electrophile clusion, the microarray and proteome results show that resistance in B. subtilis. Hence, these azoreductases, YodB negatively controls the expression of azoR1, yodC which are conserved among Gram-positive bacteria could and spx.

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1110 M. Leelakriangsak et al. ᭿ 26 Expression of azoR1 and azoR1-bsrB is induced by diamide, catechol and MHQ and derepressed in the , 2007; DyodB mutant et al. Northern blot analyses were performed to examine the

Function level and size of azoR1 transcript in wild-type and DyodB mutant cells after exposure to MHQ, catechol, diamide 2006; Duy NADH-azoreductase 1 thiol stress-specific regulator and H2O2. The results verified the very strong transcription

et al., of azoR1 in response to catechol, MHQ and diamide

mutant under control conditions stress in wild-type cells and derepression in the DyodB /wt mutant (Fig. 2). Consistent with the proteome results, yodB D

yodB there was no increase in azoR1 transcription in response D cations (Tam ome experiments described in this paper. to oxidative stress caused by H2O2. In addition, the basal a

in two proteome experiments which are induced level of azoR1 mRNA was reduced in the DmhqR mutant,

rom BSORF (http://bacillus.genome.jp). which is also consistent with the proteomic data.

/wt Two different azoR1 transcripts were detected using an mutant. azoR1-specific RNA probe (Fig. 2A and B). The smaller yodB D yodB

2.2 3.7 0.7 kb transcript is most strongly elevated after catechol, D Transcriptome Proteome MHQ and diamide stress and corresponds to an azoR1- specific monocistronic transcript. The larger 1.1 kb RNA species corresponds to the azoR1-bsrB-specific transcript which is only visible in yodB mutant cells and in wild-type cells after catechol, MHQ and diamide treatment. The bsrB gene encodes a highly abundant small 6S RNA that is transcribed during exponential growth and decreases in concentration during stationary phase (Ando et al., 2002; c Barrick et al., 2005). The conserved features of this 6S RNA suggest that it binds RNA polymerase by mimicking the structure of DNA in an open promoter complex, thereby regulating RNA polymerase activity (Wassarman and Proteome (wt) Saecker, 2006; Wassarman, 2007). Synthesis of azoR1- bsrB mRNA in treated wild-type cells was also verified using a bsrB-specific mRNA probe (Fig. 2C). These differ- ent azoR1-specific transcripts observed in the Northern blot suggest that the azoR1-bsrB mRNA might be sub- jected to RNA processing. However, Northern analysis using the bsrB-specific probe indicates that expression of bsrB can be attributed largely to constitutive transcription from its own promoter that resides downstream of the azoR1 coding sequence. Thus, the azoR1-bsrB transcript makes only a minor contribution to bsrB transcription under b thiol stress conditions. Transcription of bsrB as part of the azoR1-bsrB mRNA under thiol stress conditions might result from readthrough past the azoR1 terminator owing to the very high induction of azoR1 transcription.

Transcriptome (wt) The DyodB mutant is resistant to catechol and MHQ stress, which is attributable to derepression of the

Catechol MHQ Dia Catechol MHQ DiamideazoR1 NT gene 5.1 9.2 14 10.1 12.4 3.7 3.8 2.8 3.9 7.3 6.9 7.3 7.6 3.9 3.3 3.5 3.7 NAD(P)H nitroreductase 4.1 5.6 3.4 6.7 7.6 The growth phenotype of the DyodB mutant cells after , 2003). ) 36.7 42 106 66.3 38.4 18 20 12 19 8.9 7.4 5.3 8.5 70.9 82.4 19.6 21 FMN-dependent

Induction of genes after MHQ, catechol, diamide and nitrofurantoin stress in the wild type (wt) and derepression in the addition of the thiol-reactive compounds diamide, cat- et al.

/wt) are listed. The gene function or similarity is derived from the SubtiList database (http://genolist.pasteur.fr/SubtiList/index.html) and f echol and MHQ to exponentially growing cells was The transcriptome data for the induction ratios of YodB-dependent genes after catechol, MHQ and diamide treatments were derived from previous publi The protein synthesis ratios of YodB-dependent proteins after catechol, MHQ, diamide and nitrofurantoin treatments were calculated from two prote All genes with induction ratios of at least threefold in two transcriptome experiments and proteins with protein synthesis ratios of at least twofold analysed. The growth of the DyodB mutant was similar to yodB D c. Leichert b. ( in the wild-type (wt) after 10 min treatment with 0.33 mM MHQ, 2.4 mM catechol, 1mM diamide and 0.1 mM nitrofurantoin (NT) and derepressed in the Table 1. Gene a. azoR1(yocJ yodC spx that of the wild-type after exposure to diamide (data not

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 27 Regulation and function of the YodB repressor 1111

168 DyodBI DmhqRI

cat MHQ Dia NT H2O2 168 168 AzoR2 (YvaB) AzoR1 (YocJ)

YodC

Fig. 1. Proteome analysis of the YodB regulon in B. subtilis. Induction of AzoR1 and YodC synthesis by catechol, MHQ, diamide, nitrofurantoin and H2O2 treatment of B. subtilis wild-type cells; elevated AzoR1 and YodC production in the DyodB mutant and reduced AzoR1 and YodC levels in the DmhqR(ykvE) mutant. B. subtilis wild-type cells grown in minimal medium to an OD500 of 0.4 were pulse-labelled for 5 min each with L-[35S]methionine at control conditions (green image) and in the wild type 10 min after exposure to 2.4 mM catechol (cat),

0.33 mM MHQ, 1 mM diamide (dia), 0.1 mM nitrofurantoin (NT) and 100 mMH2O2 or in the DyodB and DmhqR mutant at control conditions (red image). shown). As reported earlier, the growth of wild-type cells catechol and MHQ resistance. As reported previously, the was inhibited in the presence of 4.8 mM catechol and growth of the DazoR2 mutant was impaired in the pres- 0.66 mM MHQ (Tam et al., 2006; Duy et al., 2007). Inter- ence of 3.6 mM catechol and 0.5 mM MHQ (Töwe et al., estingly, the DyodB mutant could grow in the presence of 2007). Surprisingly, no growth difference was detected for 9.6 mM catechol and 0.83 mM MHQ, suggesting that the azoR1 mutant compared with the wild-type parent YodB-controlled genes, derepressed in the yodB mutant, after exposure to these compounds (data not shown). confer resistance to these phenolic compounds (Fig. 3A). However, deletion of both azoreductases in the Likewise, the DmhqR mutant displayed a catechol and DazoR2DazoR1 double mutant resulted in a cessation of MHQ resistance phenotype as it was able to grow with growth after exposure to 2.4 mM catechol and cellular 12 mM catechol and 1 mM MHQ (Töwe et al., 2007). lysis after treatment with 0.5 mM MHQ (Fig. 3C). Thus, Moreover, DyodBDmhqR double mutant cells showed an the DazoR2DazoR1 double mutant was more sensitive to additive hyper-resistance to catechol and MHQ as growth catechol and MHQ stress compared with the DazoR2 was still possible in the presence of 19.2 mM catechol and single mutant. Moreover, DazoR2DazoR1 mutant cells 2 mM MHQ (Fig. 3B). showed a growth defect in the presence of 1 mM diamide The next step was to determine if the paralogous azore- compared with wild-type cells whereas the DyodB DmhqR ductases AzoR2 and AzoR1, production of which is con- double mutant grew faster in the presence of 1 mM trolled by MhqR and YodB, respectively, contribute to diamide than the wild-type parent (Fig. 3D). These

Fig. 2. Trancriptional organization (A) and YodB transcript analyses of azoR1 (B) and bsrB (C) in the wild type in response to MHQ, catechol, A yocJ bsrB diamide and H2O2 stress and in the DyodB (azoR1) and DmhqR mutants. For Northern blot 6S RNA homologue experiments 5 mg RNA isolated from the azoreductase 1 B. subtilis strains before (co) and 10 min after exposure to 2.4 mM catechol, 0.33 mM MHQ, 168 1 mM diamide and 100 mMH2O2 were applied DmhqR DyodB to each well of the denaturing agarose gel. co co MHQ cat Dia H2O2 co The Northern blots were hybridized with azoR1- (B) and bsrB-specific (C) mRNA 1.1 kb azoR1-bsrB 28.5% probes. The arrows point toward the sizes of B azoR1 0.7 kb 71.5% the azoR1-, azoR1-bsrB- and bsrB-specific transcripts. The percentage amounts of azoR1- and azoR1-bsrB-specific mRNAs are calculated with the ImageJ software and are 168 MHQ 168 cat DyodB shown on the right. co 10’ 20’ 10’ 20’ co 1.1 kb azoR1-bsrB 8.5% C

bsrB 0.2 kb 91.5%

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1112 M. Leelakriangsak et al. ᭿ 28 catechol Fig. 3. The DyodB mutant is resistant to MHQ catechol and MHQ (A), the DyodBDmhqR 10 10 double mutant is hyper-resistant (B) and A yodB yodB the DazoR2DazoR1 double mutant is hyper-sensitive to catechol, MHQ (C) and 1 1 diamide stress (D). B. subtilis wild type (wt), 500 DyodB, DyodBDmhqR, DazoR2 and

OD DazoR2DazoR1 mutant strains were grown 0.1 0.1 in minimal medium to an OD500 of 0.4 and wt 4.8 mM cat wt 0.66 mM MHQ treated with catechol, MHQ or diamide at the yodB 9.6 mM cat yodB 0.83 mM MHQ time points that were set to zero. OD500 of the 0.01 0.01 cultures was then measured at the time -180 -120 -60 0 60 120 -180 -120 -60 0 60 120 intervals indicated. 10 10 B yodB mhqR yodB mhqR 1 1 500

OD 0.1 0.1 wt 4.8 mM cat wt 0.66 mM MHQ yodB mhqR 19.2 mM cat yodB mhqR 2 mM MHQ 0.01 0.01 -180 -120 -60 0 60 120 -180 -120 -60 0 60 120 10 10 C azoR1 azoR2 azoR1 azoR2

1 1 500

OD 0.1 0.1 azoR2 2.4 mM cat azoR2 3.6 mM cat azoR2 0.5 mM MHQ azoR1 azoR2- 2.4 mM cat azoR1 azoR2- 0.5 mM MHQ 0.01 0.01 -180 -120 -60 0 60 120 -180 -120 -60 0 60 120 Time (min) Time (min) diamide D 10

1 500

OD 0.1 wt 1 mM diamide DazoR1DazoR2 1 mM diamide DyodBDmhqR 1 mM diamide 0.01 -180 -120 -60 0 60 120 180 240 Time (min) data confirmed that paralogous azoreductases encoded similarly sensitive to catechol and MHQ as the wild-type by azoR2 and azoR1 are major MhqR- and YodB- strain (data not shown). These results confirmed that the controlled determinants contributing to catechol and resistance of the DyodB mutant was caused by the over- MHQ resistance. The increased sensitivity of the production of AzoR1. DazoR2DazoR1 double mutant towards thiol-reactive compounds compared with the DazoR2 single mutant Diamide, hydrogen peroxide and MHQ impair binding of suggests that AzoR2 could complement AzoR1 deficiency YodB protein to azoR1 promoter DNA with respect to detoxification. To verify that the resistance of the DyodB mutant to To identify the region of the azoR1 promoter DNA required catechol and MHQ was attributable to azoR1 derepres- for transcriptional repression by YodB interaction, DNase sion, the DazoR1 mutation was introduced into the DyodB I footprinting was performed. The DNA that was end- mutant. Notably, the DyodB DazoR1 double mutant was labelled on the top strand of the azoR1 promoter was

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 29 Regulation and function of the YodB repressor 1113 toxic electrophiles was still observed despite the high concentration of YodB used in these experiments. We further determined whether the binding of YodB to the azoR1 promoter is reversed by the addition of diamide,

H2O2 or MHQ in varying concentrations to the footprinting reactions. As shown in lanes 4, 5, 9–11 in Fig. 4A and lanes 4–6 in Fig. 4B, YodB bound poorly to the operator

sequence when diamide, H2O2 or MHQ at levels causing toxicity in wild-type cells was present compared with the reactions that contained YodB alone (lanes 3, 8 in Fig. 4A and lane 3 in Fig. 4B). However, the presence of diamide,

MHQ and H2O2 at concentrations that inhibit YodB–target DNA interaction did not effect DNA binding of CymR repressor to the yrrT promoter (Leelakriangsak et al., 2007; M. Leelakriangsak and P. Zuber, unpublished), showing that these compounds do not have a general effect on protein–DNA interaction. In addition, neither

diamide, H2O2 nor MHQ at the higher concentrations tested in the DNA-protein-binding experiments affected DNase I activity (Fig. 4A, lanes 2 and 7 and Fig. 4B, lane 2). These results confirmed that YodB acts as a repressor by interaction with azoR1 promoter DNA and impairment of YodB–DNA interaction occurs when thiol-reactive com- pounds are present.

Fig. 4. Effect of diamide, hydrogen peroxide (A) and MHQ (B) on Role of Cys residues in repression of azoR1 DNA binding activity of YodB to the azoR1 promoter and YodB box alignments (C). DNaseI footprinting was used to assess the effect transcription by YodB in vivo of diamide, hydrogen peroxide and MHQ on YodB binding to the top strand of the azoR1 promoter. The region protected by YodB is The YodB protein contains three cysteine (Cys or C) resi- indicated by the solid line. The positions relative to dues, one at the N-terminal position 6 (C6), and two at the the transcriptional start site are shown on the left. The 15 bp C-terminal positions 101 (C101) and 108 (C108). The YodB consensus includes a 4-7-4 bp inverted repeat which is protected by YodB in the azoR1 and spx promoters. BSA, bovine N-terminal Cys residue (C6) is conserved among yodB serum albumin. homologues in low GC Gram-positive bacteria, unlike the two C-terminal Cys residues. These Cys residues may be mixed with YodB protein (Leelakriangsak et al., 2007), involved in the redox-sensitive control of YodB activity, as followed by digestion with DNase I. YodB protected a has been shown for Cys residues of other regulatory region from approximately -20 to +10 relative to the tran- factors involved in the oxidative stress response, such as scription start site (Fig. 4A, lanes 3 and 8). Promoter OxyR, Hsp33 and OhrR (Barbirz et al., 2000; Graumann alignments identified putative sites of YodB recognition et al., 2001; Fuangthong et al., 2002; Hong et al., 2005; (YodB boxes) that were uncovered by DNase I footprinting Nakano et al., 2005; Lee et al., 2007). To investigate the experiments of YodB with the spx (Leelakriangsak et al., role of Cys residues in YodB-mediated regulation, each 2007) and azoR1 promoters. YodB boxes contained a was replaced with alanine, thus generating the yodBC6A, common 15 bp consensus sequence of TACTWTTTgW yodBC101A and yodBC108A alleles, along with the WAGTA (Fig. 4C). In previous work, mutations in the spx double mutant yodBC101,108A and triple mutant promoter at positions corresponding to the nucleotides 6 yodBC6,101,108A. All mutant proteins were produced at (T), 7 (T) and 12 (A) of the consensus sequence impaired similar levels as shown by Western blot analysis (Fig. S1). YodB binding (Leelakriangsak et al., 2007). The expression of azoR1–lacZ fusions were examined

As shown previously, YodB binding activity on spx P3 in partial diploid strains bearing the DyodB allele and promoter DNA was impaired in the presence of diamide ecotopically expressed versions of the C-to-A alleles inte- and H2O2 (Leelakriangsak et al., 2007). A fourfold higher grated at the thrC locus (Fig. 5A and B). In yodB mutant concentration of YodB was used to demonstrate specific cells (ORB7106), azoR1–lacZ activity was ~60-fold higher binding to the spx P3 promoter then was used for the yocJ compared with wild-type cells (ORB7094). The azoR1– promoter owing to the lower affinity of YodB for the spx lacZ activity in strains ORB7109 (DyodB/yodBC101A), target sequence. Notably, sensitivity of YodB activity to the ORB7110 (DyodB/yodBC108A) and ORB7111 (DyodB/

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1114 M. Leelakriangsak et al. ᭿ 30

Fig. 5. Role of YodB Cys mutants in regulation of azoR1–lacZ activity (A, B) and azoR1 transcription (C, D, E) in vivo. The yodBC6A mutant shows a catechol resistance phenotype (F). A. Expression of azoR1–lacZ fusion cells containing wild-type yodB (᭜, ORB7094), DyodB mutant (᭿, ORB7106), yodB, DyodB mutant (, ORB7107), yodBC6A (᭹, ORB7108), yodBC101A (¥, ORB7109), yodBC108A (᭝, ORB7110), yodBC101,108A (+, ORB7111) and yodBC6,101,108A (ᮀ, ORB7112) were determined. The cells were grown in DSM, and their b-galactosidase activities were determined. Time zero indicates the mid-log phase. Duplicate experiments from two independently isolated strains of yodB Cys mutants were performed. (B) Expression of azoR1–lacZ fusion cells containing wild-type and yodB Cys mutants but not the DyodB mutant (᭿, ORB7106) were shown. (C, D) Quantitative Northern blot analysis and (E) primer extension analysis of azoR1 transcription using RNA extracted from strains ORB6284 (wild-type), ORB6288 (DyodB mutant), ORB6599 (yodBC6A), ORB6715 (yodBC101A), ORB6716 (yodBC108A), ORB6777 (yodBC101,108A) and ORB7083 (yodBC6,101,108A) (co and 10 min) and after treatment with 2.4 mM catechol (C), 0.33 mM MHQ (M) or 1 mM diamide (10D) for 10 min. For primer extension analysis, labelled primer oSN03-63 (Table S2) was used to examine the level of azoR1 transcript. For the dideoxynucleotide sequencing shown on the left, the nucleotide complementary to the dideoxynucleotide added in each reaction mixture is indicated above the corresponding lane (T′,A′,C′ and G′). (F) For the growth phenotypes, B. subtilis wild type, DyodB, yodBC6A and yodBC6,101,108A strains were grown in minimal medium to an OD500 of 0.4 and treated with 4.8 mM catechol at the time points that were set to zero. OD500 of the cultures was then measured at the time intervals indicated. yodBC101,108A) was similar to the wild-type strain (Fig. 5C and D) and diamide (Fig. 5E). As already shown (ORB7094). However, azoR1–lacZ activity was approxi- in Fig. 2, the level of azoR1 transcript increased after mately eightfold derepressed in the yodBC6A mutant diamide, MHQ or catechol treatment in the wild-type cells under control conditions compared with the untreated wild (Fig. 5C and D, lanes 2–3; Fig. 5E, lane 2). The transcript type. In addition, derepression of azoR1–lacZ activity was concentration of azoR1 was high in the untreated DyodB similar in the yodBC6A and triple yodBC6,101,108A mutant (Fig. 5C and D, lane 4; Fig. 5E, lane 3) compared mutants, suggesting that Cys6 is essential for YodB- with untreated wild-type cells (Fig. 5C and D, lane 1; dependent repression of azoR1 transcription under Fig. 5E, lane 1). Induction of azoR1 transcription in the non-inducing conditions. However, there is still YodB- yodBC101, yodBC108 and yodBC101,108A mutants in dependent repression in the yodBC6A and triple response to catechol, MHQ and diamide was similar to yodBC6,101,108A mutants suggesting that other resi- that of the wild type. However, significantly higher basal dues are required for optimal YodB repressor activity. levels of azoR1 transcripts were observed in the Northern blot analysis and primer extension were per- untreated yodBC6A and triple yodBC6,101,108A mutants formed to determine the level of azoR1 transcription after (Fig. 5C and D, lanes 7 and 10; Fig. 5E, lanes 7 and 15) treatment of yodB Cys-to-Ala mutants with MHQ, catechol compared with untreated wild-type cells (Fig. 5C and D,

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 31 Regulation and function of the YodB repressor 1115 Fig. 6. Reduction of YodBC6A protein binding activity to the spx P3 promoter (A) and azoR1 promoter (B) in vitro. DNase I footprinting was used to assess the binding activity of YodB and YobBC6A protein and the effect of diamide on DNA binding. The concentrations of proteins used were 4 mM (A, lanes 3–8) and 0.5 mM (B, lanes 3–8).The protected region by YodB is indicated by the solid line. The positions relative to the transcriptional start site are shown. The concentrations of diamide used were 1 and 1.5 mM for spx, 125 and 250 mM for azoR1. BSA, bovine serum albumin.

lane 1; Fig. 5E, lane 1), which is consistent with the esis that the Cys residues are at least in part involved b-galactosidase activity data (Fig. 5A and B). Quantifica- YodB repressor activity. tion of the Northern blots revealed ~25-fold derepression To examine YodBC6A protein binding activity to spx and of azoR1 transcription in the DyodB mutant (Fig. 5D). In azoR1 promoter DNA, the YodBC6A protein was purified the untreated yodBC6A and triple yodBC6,101,108A (see Experimental procedures) and used in DNase I foot- mutants, the amounts of azoR1 mRNAs were approxi- printing experiments to compare mutant DNA-binding mately eightfold increased, compared with the untreated protein activity with that of wild-type YodB. YodBC6A wild type (Fig. 5C, lanes 7 and 10). There was a two- to protein exhibited reduced binding activity to the target threefold induction of azoR1 transcription in the yodBC6A DNA when compared with the wild-type YodB protein and yodBC6,101,108A triple mutants after exposure to (Fig. 6A and B, lanes 3 and 6). To further clarify whether catechol and MHQ. But the level of expression in the case the binding of YodBC6A protein was sensitive to a toxic of the triple mutant was lower than the induced level of oxidant, diamide was added to the footprinting reactions. wild-type cells (Fig. 5D and E). Furthermore, primer As had been shown previously, diamide reduced YodB extension analysis of the yodBC6,101,108A triple mutant binding to target DNA (Leelakriangsak et al., 2007). showed little if any induction following diamide treatment Diamide also caused a loss of YodBC6A binding activity to (Fig. 5E, lanes 15 and 16), with a lower level of transcript spx and azoR1 promoter DNA in vitro (Fig. 6A and B, than that observed in reactions of RNA from wild-type lanes 7–8). These data indicated that YodBC6A could still cells treated with diamide. Together, the Northern and function and respond somewhat to thiol-reactive com- primer extension analysis suggests that the Cys residues pounds despite its reduced DNA-binding activity. play a sensory function required for the repressor’s Mass spectrometry analysis of YodB modified in vitro by responsiveness to the inducing compounds. catechol and MHQ The DyodB mutant acquires an increased resistance to catechol and MHQ owing to the upregulation of We next examined whether treatment of YodB with the AzoR1. To confirm that the yodBC6A mutant retains this thiol-reactive compounds catechol and MHQ leads to spe- resistance phenotype owing to weakened repressor cific Cys6 modifications. Purified His6-YodB was cleaved activity, growth experiments were performed in the pres- with Tev (tobacco-etch virus) protease to leave only the ence of different concentrations of catechol. The results tripeptide GAH N-terminal extension of the YodB protein. showed that the yodBC6A single or yodBC6,101,108A This modified YodB protein was incubated for 10 min with triple mutants could grow with 4.8 mM catechol (Fig. 5F). each compound and then digested with trypsin for 2 h. The In contrast, growth of the yodBC101A, yodBC108A and tryptic digest was analysed using FT-ICR-MS (Fourier yodBC101,108A mutants was similar to the wild type transform ion cyclotron resonance MS) in the positive ion and inhibited in the presence of 4.8 mM catechol (data mode. In the untreated YodB sample, the 573.75 Da mass not shown). However, resistance of the yodBC6A single peak corresponding to the double charged T1 peptide or yodBC6,101,108A triple mutants was lower than that (m/z = 1145.49 + 2H+) was found as the most abundant of DyodB mutant cells. These results support the hypoth- peptide, suggesting that Cys6 is in the reduced form

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1116 M. Leelakriangsak et al. ᭿ 32

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 33 Regulation and function of the YodB repressor 1117 (Fig. 7A). However, small amounts of T1 peptides with thiol-specific electrophiles and confers resistance to cat- methionine sulphoxides and the fully oxidized cysteine echol and MHQ in B. subtilis (Töwe et al., 2007). Thus, sulphonic acids were also detected, which could be both enzymes have similar functions related to the detoxi- generated upon protein purification or during sample fication of catechol and MHQ. Catechol and hydroquino- preparation. nes can be auto-oxidized to the corresponding ortho-and The treatment of YodB with 1 mM MHQ resulted in the para-benzoquinones that can act as toxic oxidants and appearance of a new mass peak of 634.76 Da correspond- electrophiles (Fig. 8). The formation of the benzoquinones ing to the double charged T1 peptide with a 122 Da modi- could be detected by the appearance of a red-coloured fication (m/z = 1267.52 + 2H +) (Fig. 7B). This mass peak medium if cells are exposed to growth-inhibitory concen- is representative of the sulphhydryl-bound form of MHQ tration of catechol and MHQ. (m/z = T1 + 122 +2H+) as there is the loss of 2 Da owing to Quinones are enzymatically reduced via two different the phenolic ring-conjugation with the thiolate (-2H+). The pathways: (i) formation of semi-quinone radicals and reac- collision induced dissociation MS/MS analysis of the tive superoxide anions via one-electron reduction (e.g. double charged m/z = 1267.52 peak confirmed that MHQ NADPH-cytochrome P450 reductase, z-crystallin) and (ii) is covalently linked to the Cys6 residue (Fig. S2). formation of hydroquinones in the two-electron reduction The treatment of YodB with 10 mM catechol caused the by NAD(P)H quinone oxidoreductases (Ernster, 1987). formation of a new mass peak of 627.76 Da correspond- The functions of the azoreductases AzoR1 and AzoR2 ing to the double charged T1 peptide with a 108 Da are related to NADH:quinone reductases, also called dia- modification (m/z = 1253.52 + 2H+) (Fig. 7C). This phorases, which catalyse the NAD(P)H-dependent two- mass peak represents the Cys6-S-adduct of catechol electron reductions of quinones, quinoneimines, azodyes (m/z = T1 + 108 + 2H+), which was confirmed by collision and nitrogroups to protect cells against the toxic effects of induced dissociation MS/MS analysis of this peptide free radicals and ROS arising from the one-electron (Fig. S3). The reduced Cys101 and Cys108 containing reductions (Ernster, 1987; Li et al., 1995; Cavelier and 1680.73 Da mass peak was observed in the untreated Amzel, 2001). The azoreductases could function to YodB sample corresponding to the double charged detoxify the quinones back to redox-stable catechol and T9-peptide (m/z = 3359.45 + 2H+) (data not shown). In MHQ to protect cells against the redox-cycling action addition, the disulphide-bound form of the T9 peptide was and reaction as electrophiles targeting protein thiolates detected in the untreated YodB sample as judged from (Fig. 8). This thesis is further supported by the fact that the the loss of 2 Da of this peptide (m/z = 3357.45 + 2H+) DmhqR and DyodB mutants show a four- to fivefold (data not shown). Sulphhydryl-bound forms of MHQ and increased resistance to para-benzoquinone (our unpub- catechol were also detected for this T9 peptide, suggest- lished data). The toxicity of para-benzoquinone is more ing that both MHQ and catechol auto-oxidize to quinones, than 50-fold higher than that of hydroquinone in which react as electrophiles with protein thiolates. B. subtilis, which might explain the thiol-reactive electro- philic nature. It has been further shown, that AzoR2 pos- sesses NADH:DCIP oxidoreductase and azoreductase Discussion activity (Nishiya and Yamamoto, 2007). Azoreductases are flavoenzymes that have the unique ability to catalyse YodB and MhqR affect quinone and azo-compound a wide range of biochemical reactions and have broad detoxification by controlling production of substrate specificity (Fraaije and Mattevi, 2000). Thus, paralogous azoreductases both azoreductases, AzoR1 and AzoR2 could also In this study, we uncovered the azoreductase-encoding function in the reductive cleavage of the electrophile azoR1 gene as the most strongly derepressed member of azo-compound diamide to 1,1′-dimethylurea. Another fla- the YodB regulon. The yodB mutant showed increased voenzyme, the nitroreductase YodC is also regulated by resistance to catechol and MHQ, which could be attrib- YodB, and is a paralogue of MhqN, the gene for which is uted to the function of AzoR1. The YodB-controlled azoR1 controlled by MhqR. Thus, paralogous nitroreductases gene encodes an azoreductase, AzoR1, which is a para- are co-regulated with paralogous azoreductases in logue of AzoR2, specified by the MhqR-regulated azoR2 response to thiol-reactive electrophiles. Nitroreductases gene. Production of both enzymes responds to catalyse the two-electron reduction of nitrogroups in

Fig. 7. Modification of YodB in vitro by MHQ and catechol. Purified YodB (5 mg) was incubated for 10 min with 1 mM MHQ and 10 mM catechol and analysed by FT-ICR-MS in the positive ion mode after tryptic digestion. In the untreated YodB sample, the double charged T1 peptide was mostly detected in the Cys6-reduced form (m/z = 573.75) (A). Methionine sulphoxide modifications of T1 are indicated with an asterisk and cysteine sulphonic acid modification of T1 is indicated by #. After treatment of YodB protein with MHQ, a new peak corresponding to the double charged T1 with Cys6-S-conjugated MHQ was detected (m/z = 634.76) (B). The treatment with catechol resulted in a new mass peak corresponding to the double charged T1 with Cys6-S-conjugated catechol (m/z = 627.76) (C). © 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1118 M. Leelakriangsak et al. ᭿ 34 Proposed mechanism for electrophile detoxification in B. subtilis

O MhqA OH ½O H O OH 2 2 MhqO ring cleavage OH O Catechol + R-SH OH MhqE product NAD+ NADH+H+ + R-SH AzoR1 ortho- AzoR2 benzoquinone S-R

OH O OH MhqA ½O2 H2O MhqO CH CH CH 3 3 + R-SH 3 MhqE ring cleavage MHQ product NAD+ NADH+H+ AzoR1 S-R + R-SH OHAzoR2 O OH Methyl-para-

benzoquinone

O

H3C O CH H diamide N N 3 H C H C N N + 2 R-SH 3 CH

3 N N 3 + R-S-S-R

+ N O 2 NADH+H CH3 H C NH 3 CH AzoR1 O 3 AzoR2 Hydrazine 2 NAD+ H3C CH N 3 NH2 2HN N H3C CH3 O O 1,1'-Dimethylurea

Fig. 8. Thiol-specific reactions of electrophilic compounds and model for the functions of MhqR- und YodB-controlled azoreductases and dioxygenases/glyoxalases in B. subtilis. Catechol and MHQ are oxidized to the corresponding ortho- and para-benzoquinones that cause S-adduct formation via 1,4-reductive addition of thiols to quinones (Michael-type addition) (Rodriguez et al., 2004). The paralogous azoreductases AzoR1 and AzoR2 are major quinone resistance determinants and function in quinone reduction to catechol and MHQ (Cavelier and Amzel, 2001). Diamide is an electrophilic azo-compound that reacts with protein thiolates and causes disulphide bond formation or mixed disulphides with LMW thiols (Leichert et al., 2003; Hochgräfe et al., 2007). AzoR1 and AzoR2 also confer resistance to diamide and function in detoxification of diamide via the reductive cleavage of the azo group, resulting in two molecules 1,1′-Dimethylurea. The paralogous dioxygenases/glyoxalases which are co-regulated with the azoreductases (Töwe et al., 2007) might be involved in the thiol-specific detoxification of the LMW-thiol-S-adducts via dioxygenolytic cleavage of the compounds. Thus, this mechanism of electrophile resistance is coupled to the thiol-specific stress responsive regulatory network. nitroaromatic compounds to nitroso, hydroxylamine inter- Interestingly, neither of these azo- or nitroreductases mediates and finally primary amines (Bryant and DeLuca, was induced by oxidative stress provoked by H2O2 or the 1991). As quinones also represent substrates for nitrore- redox-cycling agent paraquat (our unpublished data) ductases (Nivinskas et al., 2002), YodC and MhqN could supporting the thesis that these enzymes are specifically function in quinone reduction in B. subtilis. There is the involved in quinone and electrophile reduction and not in question: Why have paralogous azo- and nitroreductases the oxidative stress response. As ROS are produced evolved that act upon the same class of substrates, but upon auto-oxidation of MHQ and catechol, this could are separately regulated by YodB and MhqR? Two account for the induction of the PerR-controlled oxidative oxygen-insensitive FMN-containing nitroreductases, NfsA stress response by electrophiles. Other quinone oxi- and NfsB, have been also characterized in Eschericia coli. doreductases such as QorA of Staphylococcus aureus Of these, NfsA uses NADPH as electron donor whereas are produced in response to oxidative stress and these NfsB can use either NADH or NADPH as source of catalyse the one-electron reduction of quinones, thus reduction equivalents (Zenno et al., 1996a,b). Thus, the producing superoxide anions (Maruyama et al., 2003). paralogous nitro- and azoreductases of B. subtilis that Interestingly, QorA is homologous to the quinone oxi- contribute to quinone resistance could use different elec- doreductase YfmJ of B. subtilis. YfmJ protein was also tron donors for quinone reduction. The detailed mecha- induced by MHQ treatment (Duy et al., 2007) and indeed nism of quinone reduction remains to be elucidated, functions in quinone reduction in vitro (A. Maruyama, however. pers. comm.). In addition, production of many other

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 35 Regulation and function of the YodB repressor 1119 NAD(P)H-oxidoreductases responds to MHQ or catechol The Cys residues of YodB function in sensing and might contribute to either the one- thiol-reactive electrophiles or two-electron reduction of quinones in B. subtilis (Duy et al., 2007). In this study, the regulation of the MarR/DUF24-family repressor YodB in response to thiol-specific electrophiles was uncovered in B. subtilis. YodB is highly conserved The mechanism of quinone reduction by nitro- and among low-GC Gram-positive bacteria and several para- azoreductases is coupled to the thiol-specific stress logues are encoded within the B. subtilis genome. YodB- response in B. subtilis like regulators share the conserved N-terminal cysteine in The question arises why regulation of quinone reduction position 5 or 6 followed by a conserved proline residue. by the azoreductases is coupled to the thiol-specific As reported herein, this conserved Cys6 is essential for stress response in B. subtilis. Quinones exert their tox- repression of YodB-controlled target genes in vivo and in icity mainly as electrophiles via the 1,4-reductive nucleo- vitro. Inactivation of Cys6 in a yodBC6A mutant resulted in philic addition to protein thiolates (Michael-type addition) a decreased repression of azoR1 in vivo. Furthermore, leading to thiol-S-adduct formation (Fig. 8). Thus, the conserved Cys6 was found to play a role in regulation quinone and diamide exposure results in depletion of of YodB activity as observed by examining the phenotype protein thiolates and non-protein-low molecular weight of the yodBC6A mutant, which was more resistant to (LMW) thiolates. B. subtilis does not possess glutathione catechol than the wild-type cells. Thus, it could be pos- and instead, the major LMW thiols of B. subtilis are likely sible that the Cys6 residue has a higher reactivity to form cysteine, CoASH, or the recently reported 398 Da bio- the nucleophilic thiolate than the Cys101 and Cys108 thiol (Newton et al., 1996; Gusarov and Nudler, 2005; residues. There was still some repression of YodB- Lee et al., 2007; Hochgräfe et al., 2007). Recently, it has dependent genes in the yodBC6A mutant in vivo as the been shown that diamide triggers S-cysteinylation in vivo basal level of azoR1 transcription was lower in the in a number of B. subtilis cytoplasmic proteins that have yodBC6A mutant compared with the yodB null mutant. catalytic cysteine residues (e.g. GuaB, MetE and PpaC) Transcription of azoR1 was still two- to threefold inducible (Hochgräfe et al., 2007). The formation of mixed disul- in the yodBC6A mutant after exposure to catechol and phides with the LMW thiol cysteine is proposed as a MHQ, suggesting that there might be other sites in the mechanism to protect catalytic cysteine residues against YodB protein that are involved in regulation of YodB oxidation to irreversible sulphonic acid. Thus, the LMW activity. Electrophilic quinone-like compounds could react thiol cysteine could function as a GSH equivalent in with other nucleophilic amino acids present in the YodB detoxification of electrophiles via cysteine-S-adduct for- protein, thereby influencing YodB DNA-binding activity. mation. The CymR regulon which regulates cysteine bio- Interestingly, there are reports that quinone-like electro- synthesis is also derepressed under the conditions of philes can also form chemical adducts on protein nitrogen diamide, catechol and MHQ stress which supports the nucleophiles at the N-terminus and at the basic amino idea of cysteine depletion owing to S-adduct formation acid side-chains of lysine, arginine and histidine residues with electrophiles. (Person et al., 2003; 2005). As several basic lysine and The MarR-type repressor MhqR regulates expression arginine residues are present in the N-terminal part of the of azoR2 and genes encoding multiple dioxygenase/ YodB protein, further mutagenesis studies are required to glyoxalase family enzymes in response to thiol stress resolve this issue. Notably, the C-terminal C101 and C108 conditions (Töwe et al., 2007). These dioxygenases/ residues are not required for YodB repression, but were glyoxalases could be involved in the thiol-dependent ring found also in disulphide bond form in vitro as revealed cleavage of the catechol- and MHQ-S-adducts (Fig. 8). by mass spectrometry. The triple mutant, C6,101,108A, This proposed mechanism of electrophile detoxification is shows reduced repression, but also lower responsiveness similar to the GSH-dependent detoxification of methylgly- to inducers, suggesting the Cys residues of YodB could oxal and other electrophiles (e.g. N-ethylmaleimide) in serve a sensory function by reacting directly with thiol- E. coli which involves glutathione-S-adduct formation with reactive compounds. the electrophiles and detoxification via the glyoxalase Using mass spectrometry we uncovered the formation pathway or release of these GSH-adducts from the cell of Cys-S-adducts upon treatment of YodB with MHQ and (Ferguson et al., 1998, 1999; McLaggan et al., 2000; catechol in vitro confirming the thiol reactivity of the quino- Booth et al., 2003). The results reported herein suggest nes produced upon catechol and MHQ auto-oxidation. that azo- and nitroreductases function in the detoxification Thus, the derepression of YodB in response to diverse of electrophiles whereas dioxygenases/glyoxalase could aromatic compounds might be caused by their common be involved mainly in detoxification of the redox-stable electrophilic nature, which leads to covalent cysteine S-hydroquinone adducts. modifications. Cysteine residues are strong nucleophiles

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1120 M. Leelakriangsak et al. ᭿ 36 and form redox centres in an emerging group of oxidative (yocJ::pMutin4) strain, which carries an azoR1–lacZ fusion, stress transcription factors such as the E. coli oxidative resulting in a strain designated ORB6734. The yodB mutant stress regulator OxyR and the B. subtilis organic hydrop- bearing an azoR1–lacZ fusion was constructed by transfor- mation of ORB6734 with chromosomal DNA of ORB6208 eroxide repressor OhrR (Fuangthong et al., 2002; Paget (yodB mutant) (Leelakriangsak et al., 2007) to generate and Buttner, 2003; Hong et al., 2005; Lee et al., 2007). All ORB6735. attempts to detect YodB modifications upon treatment The azoR1–lacZ fusion was constructed by transformation with diamide failed using mass spectrometry. We only of JH642 competent cells with plasmid pML78, which carries observed that the peptide containing Cys6 was absent an azoR1–lacZ fusion, resulting in strain ORB7093. pML78 in the MS spectrum after diamide treatment (data not was generated by PCR using primers oSN03-64 and oSN03- shown). Thus, the mechanism of YodB inactivation upon 65. The PCR fragment (-155 to +110 with respect to the diamide treatment is currently unknown. As S-thiolation azoR1 transcription start site) was digested with enzymes EcoRI and BamHI, and then was inserted into plasmid has been shown as one mechanism of OhrR and OxyR pDH32 which was digested with the same enzymes. The regulation (Kim et al., 2002; Lee et al., 2007), modification azoR1 sequence was verified by DNA sequencing. Plasmid of YodB could occur in vivo in response to diamide and pCm::tet was used to transform ORB7093, replacing the Cmr other electrophiles via the formation of mixed disulphides cassette with a Tetr marker to generate strain ORB7094. containing cysteine, CoASH or other LMW thiols. One The yodB disruption strain bearing azoR1–lacZ fusion was goal of our future proteomic studies will be to determine constructed by transformation of ORB7094 with chromo- whether the YodB repressor is regulated by S-thiolation or somal DNA of ORB6208 (Leelakriangsak et al., 2007). The resulting strain was designated ORB7106 (azoR1–lacZ, S-adduct formation by electrophiles in vivo. However, yodB). Diploid yodB strains bearing azoR1–lacZ fusion were detection of S-conjugates with quinone-like electrophiles, constructed as follows. The chromosomal DNA of ORB6606 which are formed in vivo will be difficult using mass spec- (thrC::yodB) (Leelakriangsak et al., 2007), ORB6592 trometry as these are rather unstable modifications and (thrC::yodBC6A), ORB6713 (thrC::yodBC101A), ORB6714 are often lost upon prefractionation procedures. (thrC::yodBC108A), ORB6775 (thrC::yodBC101,108A) and In summary, our studies uncovered the function and ORB7081 (thrC::yodBC6,101,108A) were used to transform regulation of the YodB repressor that controls the expres- ORB7106 (azoR1–lacZ, yodB) to generate ORB7107, ORB7108, ORB7109, ORB7110, ORB7111 and ORB7112 sion of spx, and the genes that encode the azoreductase respectively. The yodB::pMutin4 and azoR2::pMutin4 AzoR1 and the nitroreductase YodC in response to thiol- mutants were constructed during the course of the European specific electrophiles. We show that the conserved Cys6 and Japanese B. subtilis functional analysis project. residue of YodB is required for optimal YodB-mediated Chromosomal DNA of the azoR2::pMutin4 (ST8) and repression in vivo and in vitro and that the three Cys yodB::pMutin4 (ST5) mutants were introduced by trans- residues function in inducer sensing. In addition, we formation into the DazoR1 mutant (ST7) to generate the report that paralogous azoreductases AzoR1 and AzoR2 DazoR1DazoR2 (ST9) and DazoR1DyodB (ST10) double mutants respectively. The DmhqR mutant (TF176) was trans- both confer resistance to catechol and MHQ in B. subtilis. formed with chromosomal DNA of the yodB::pMutin4 (ST5) Hence, we presume that these azoreductases of mutant to generate the DyodBDmhqR double mutant (ST6). B. subtilis function as quinone reductases and constitute The lesions harboured by the mutants were verified by PCR. major detoxification pathways to protect bacteria from When necessary, the following antibiotics were added (at a quinone toxicity. As these azo- and nitroreductases are concentration shown in parenthesis): ampicillin (25 mgml-1), highly conserved among pathogenic Gram-positive bac- chloramphenicol (5 mgml-1), erythromycin plus lincomycin (1 -1 -1 teria, it will be interesting to find the naturally ocurring and 25 mgml respectively), and tetracycline (12.5 mgml ). quinone-like substrates for these enzymes. Construction of yodBC6A, C101A, C108A, C101,108A Experimental procedures and C6,101,108A mutants Bacterial strains and growth conditions Plasmids pML60(C6A), pML65(C101A), pML66(C108A), pML69(C101,108A) and pML76(C6,101,108A) were pro- Bacillus subtilis strains used in this study are derivatives of duced by using PCR mutagenesis. First-round PCR was JH642 and 168 and are listed in Table S1. Cells were culti- performed in two separate reactions with primers oyodB- vated in a shaking water bath at 37°C in Difco sporulation EcoRI with oyodB-C6AR and primers oyodB-C6AF with medium (DSM) for b-galactoisidase assays or TSS minimal oyodB-HisB (Table S2) using JH642 chromosomal DNA as medium and Belitsky minimal medium (Stülke et al., 1993) for template. The PCR products were hybridized and subse- treatments with thiol-reactive compounds. Diamide and quently amplified by a second round of PCR using primers hydrogen peroxide were purchased from Sigma-Aldrich. oyodB-EcoRI and oyodB-HisB. The products from the second MHQ was purchased from Acros and catechol was pur- round PCR were then digested with EcoRI and BamHI chased from Fluka. restriction enzymes and inserted into plasmid pUC19 that The isogenic azoR1 mutant was constructed by transfor- was digested with the same enzymes to generate pML60. mation of JH642 with chromosomal DNA of the azoR1D pML76 was generated by the same procedure as that used

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 37 Regulation and function of the YodB repressor 1121 for construction of pML60, but using pML69 as template. The the protein was purified as described previously (Leelakriang- PCR product (ª 60 bp) that was synthesized using primer sak et al., 2007). oyodB-C101A and oyod-HisB was used as the primer together with oyodB-EcoRI for second round PCR to gener- ate pML65. pML66 was generated by using primers oyodB- Primer extension analysis EcoRI and oyodB-C108A. Primers oyodB-EcoRI and oyodB- Strains ORB6284, ORB6288, ORB6599, ORB6715, C108A (Table S2) were used to produce pML69 using pML65 ORB6716 and ORB6777 were grown at 37°C in TSS as the template. The yodB sequences were verified by DNA medium. Primer extension analysis was performed as sequencing. The plasmids pML60, pML65, pML66, pML69 described previously (Leelakriangsak and Zuber, 2007). and pML76 were digested with EcoRI and BamHI and the Primers oML02-15 and oSN03-63 were used to synthesize released fragments bearing the mutations were then inserted the primer extension products to examine the transcripts of into pDG795 that was digested with the same enzymes spx and azoR1 respectively. to generate pML61, pML67, pML68, pML72 and pML77 respectively. pML61, pML67, pML68, pML72 and pML77 were introduced by transformation with selection for DNase I footprinting erythromycin-lincomycin resistance, into JH642, where yodBC6A, C101A, C108A, C101,108A and C6101,108A frag- A DNA probe for azoR1 (position corresponding -160 to +110 ments integrated into the thrC locus. The resulting strains with respect to transcription start site) was synthesized by were designated ORB6592 (thrC::yodBC6A), ORB6713 PCR amplification using primers oSN6-03–64 and oSN03-65. (thrC::yodBC101A), ORB6714 (thrC::yodBC108A), Dideoxy sequencing ladders were obtained using the Thermo ORB6775 (thrC::yodBC101,108A) and ORB7081 Sequenase cycle sequencing kit (USB) and the PCR product (thrC::yodBC6,101,108A). Chromosomal DNA of ORB6592, specifying azoR1 as template. The same primers used for the ORB6713, ORB6714, ORB6775 and ORB7081 were used footprinting reactions were also used for sequencing. DNase to transform ORB6288 to generate ORB6599, ORB6715, I footprinting reactions were performed as described previ- ORB6716, ORB6777 and ORB7083 respectively. ously (Leelakriangsak et al., 2007).

Transcriptome analysis Proteome analysis

For microarray analysis, the B. subtilis 168 wild-type and Cells grown in minimal medium to an OD500 of 0.4 were isogenic DyodB (TF277) mutants were grown in minimal pulse-labelled for 5 min each with 5 mCi of L-[35S]methionine medium and harvested at OD500 of 0.4. Total RNA was iso- per ml before (control) and 10 min after exposure to 0.33 mM lated by the acid phenol method as described (Majumdar MHQ, 2.4 mM catechol, 1 mM diamide, 0.1 mM nitrofuran- et al., 1991). The generation of fluorescence-labelled cDNA toin and 100 mMH2O2. Preparation of cytoplasmic and hybridization with B. subtilis whole-genome microarrays L-[35S]methionine-labelled proteins and separation by two- (Eurogentec) was performed as described previously (Jürgen dimensional gel electrophoresis using the immobilized pH et al., 2005). Two independent hybridization experiments gradients in the pH range 4–7 was performed as described were performed using RNAs from two independent sets of (Tam et al., 2006). The quantitative image analysis was per- cultures. Genes showing induction ratios of at least threefold formed with the DECODON Delta 2D software (http://www. in two independent experiments were considered as signifi- decodon.com). cantly induced. An independent microarray experiment was conducted using strains ORB6208 (DyodB::cat) and the JH642 parent. Procedures for RNA purification and micro- Northern blot analyses array analysis were described previously (Nakano et al., Northern blots were performed as described (Wetzstein et al., 2003a; Choi et al., 2006). 1992) using RNA isolated from B. subtilis wild-type and DyodB (TF277) mutant cells before (control) as well as 10 Assay of b-galactosidase activity and 20 min after treatment with 2.4 mM catechol or 0.33 mM MHQ. Hybridizations specific for azoR1 and bsrB were Toluene treatment and assays of b-galactosidase activity performed with the digoxigenin-labelled RNA probes synthe- using o-nitrophenyl-b-D-galactopyranoside were performed sized in vitro using T7 RNA polymerase from T7 promoter- as previously described (Nakano et al., 1988; Schrogel and containing PCR products amplified with azoR1-specific Allmansberger, 1997). b-Galactosidase activity is expressed primers (yocJ_for and yocJ_rev) and bsrB-specific primers as Miller Units (Miller, 1972). (bsrB-for and bsrB-rev).

Protein purification FT-ICR-MS analysis

The yodB coding sequence was amplified by PCR using LC-ESI MS experiments were performed using an LTQ (linear primers oyodB-HisNC6A and yodB-HisB. The PCR products ion trap) FTICR mass spectrometer (Thermo Electron, San were digested with NdeI and BamHI restriction enzymes and Jose, CA). The SEQUEST algorithm Version 27.12 (Thermo inserted into pPROEX-1(Life Technologies) digested with the Electron) was used to search for modifications of the YodB same enzymes to generate pML74. Cells were cultured and tryptic peptides. A mass deviation of 0.01 Da for precursor

© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 1122 M. Leelakriangsak et al. ᭿ 38 ions as well as for fragment ions and one missed cleavage vinyl-chroman-4-on reveal different degradation systems site of trypsin were allowed. The search result was filtered involved in the catabolism of aromatic compounds in Bacil- using BioWorks 3.2 (Thermo Electron). A multiple threshold lus subtilis. Proteomics 7: 1391–1408. filter applied at the peptide level consisted of the following Ernster, L. (1987) Dt-diaphorase: a historical review. Chem criteria: (i) peptide sequence length: 7–30 amino acids; (ii) Scr 27A: 1–13. RSp Յ 4; (iii) percentage of ions: 70 (70% of all theoretical b- Ferguson, G.P., Tötemeyer, S., MacLean, M.J., and Booth, and y-ions of a peptide are experimentally found); and (iv) I.R. (1998) Methylglyoxal production in bacteria: suicide or Xcorr versus charge state: 1.90 for singly charged ions, 2.20 survival? Arch Microbiol 170: 209–218. for doubly charged ions and 3.75 for triply charged ions. Ferguson, G.P., VanPatten, S., Bucala, R., and Al-Abed, Y. (1999) Detoxification of methylglyoxal by the nucleophilic bidentate, phenylacylthiazolium bromide. Chem Res Acknowledgements Toxicol 12: 617–622. Fraaije, M.W., and Mattevi, A. (2000) Flavoenzymes: diverse The authors wish to thank C. Lee and A. D. Grossman for catalysts with recurrent features. Trends Biochem Sci 25: technical assistance with microarray experiments; and K. 126–132. Review. Kobayashi and H. Yoshikawa for gifts of B. subtilis strains Fuangthong, M., Herbig, A.F., Bsat, N., and Helmann, J.D. used in the experiments described herein. We thank M. M. (2002) Regulation of the Bacillus subtilis fur and perR Nakano for advice and valuable discussions. Research genes by PerR: not all members of the PerR regulon are reported herein was supported by Grant GM45898 from the peroxide inducible. J Bacteriol 184: 3276–3286. National Institutes of Health, USA; and by grants from the Graumann, J., Lilie, H., Tang, X., Tucker, K.A., Hoffmann, Deutsche Forschungsgemeinschaft, the Bundesministerium J.H., Vijayalakshmi, J., et al. (2001) Activation of the redox- für Bildung und Forschung (BACELL-SysMo 031397A), the regulated molecular chaperone Hsp33 – a two-step Fonds der Chemischen Industrie, the Bildungsministerium mechanism. Structure 9: 377–387. of the country Mecklenburg-Vorpommern and European Gusarov, I., and Nudler, E. (2005) NO-mediated cytoprotec- Union grants BACELL-Health (LSHG-CT-2004-503468) and tion: instant adaptation to oxidative stress in bacteria. Proc BACELL-BaSysBio (LSHG-CT-2006-037469) to M.H., by a Natl Acad Sci USA 102: 13855–13860. scholarship of the ‘Ministry of Education and Training of Hochgräfe, F., Mostertz, J., Pöther, D.C., Becher, D., Vietnam’ (MOET) to N.v.D. Helmann, J.D., and Hecker, M. 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© 2008 The Authors Journal compilation © 2008 Blackwell Publishing Ltd, Molecular Microbiology, 67, 1108–1124 CatR (YvaP) 41 Huyen et al., 2008a

Chapter 2

The MarR/DUF24-type repressor CatR (YvaP) controls the thiol-stress inducible catechol dioxygenase YfiE (CatE) in Bacillus subtilis

Nguyen Thi Thu Huyen1, Britta Möhl1, Franzika Schuster1, Kazuo Kobayashi2, Dörte Becher1, Michael Hecker1 and Haike Antelmann1

1Institute for Microbiology, Ernst-Moritz-Arndt-University of Greifswald, F.-L.-Jahn-Str. 15, D-17487 Greifswald, Germany 2Graduate School of Information Science, Nara Institute of Science & Technology, Takayama 8916-5, Ikoma, Nara 630-0192, Japan

CatR (YvaP) was characterized as a further YodB-paralogous thiol-stress sensing DUF24/MarR-type repressor of the previously characterized yfiDE (catDE) operon in B. subtilis. K. Kabayashi identified the MarR-type regulator YvaP as a repressor of the catDE operon by using transcription factor arrays. B. Möhl was involved in construction and phenotype analysis of the yvaP mutant, overproduction and purification of the YvaP repressor. H. Antelmann carried out DNase I footprinting analyses and gel mobility shift assays of purified YvaP protein to the yfiDE promoter. These revealed that DNA binding activity of YvaP is directly inhibited in the presence of thiol-reactive electrophiles. Mass spectrometry analysis of thiol-modifications caused in YvaP by diamide and quinones were performed by H. Antelmann and D. Becher. F. Schuster performed non-reducing SDS-PAGE analysis, which showed that diamide and quinones cause an intersubunit disulfide bond formation in vitro and in vivo in YvaP. My contribution in this chapter is the construction of yvaP point mutants and the Northern blot analysis of yfiDE transcription in these points mutants to analyze the roles of Cys7 and basic arginine and lysine residues (K19A, R20A, R35A, R49A, R54A, K56A, R66A, R75A and K83A) for regulation of YvaP in vivo.

CatR (YvaP) 42 Huyen et al., 2008a

The MarR/DUF24-type repressor CatR (YvaP) controls the thiol-dependent catechol dioxygenase YfiE (CatE) in Bacillus subtilis

Nguyen Thi Thu Huyen1#, Britta Möhl1#, experiments revealed that YvaP confers Franziska Schuster1, Kazuo Kobayashi2, resistance to catechol, but not to Dörte Becher1, Michael Hecker1, and Haike hydroquinone-like electrophiles. Among Antelmann1* the YodB-paralogs encoded in the B. subtilis genome, YvaP and YodB display 1Institute for Microbiology, Ernst-Moritz- most close sequence similarities and share Arndt-University of Greifswald, F.-L.-Jahn- the conserved N-terminal cysteine in Str. 15, D-17487 Greifswald, Germany position 6 and 7, respectively. DNA binding 2Graduate School of Information Science, experiments showed that YvaP protects Nara Institute of Science & Technology, two 6-1-6 bp inverted repeat sequences Takayama 8916-5, Ikoma, Nara 630-0192, (YvaP-boxes) with the consensus of Japan TTACtTtAaGTAA overlapping the yfiDE promoter region. DNA binding activity of # These authors contributed equally to this YvaP was inhibited by quinone-like work. electrophiles and diamide in vitro suggesting that also YvaP is directly *Corresponding author: Tel. +49-3834- sensing thiol-stress conditions via the 864237, Fax. +49-3834-864202, e-mail: conserved Cys7. Mutational analyses [email protected] showed that the conserved Cys7 is essential for YvaP regulation in vivo and probably Key words: serves as a sensor for thiol-reactive Bacillus subtilis/YvaP/ YodB/ MarR-type compounds. In addition, also the basic regulator/ quinone/ dioxygenase amino acids K19, R20 were shown to be essential for YvaP repression in vivo as SUMMARY well as most other conserved basic arginine and lysine residues located in the DNA The response to electrophilic quinones is binding HTH motif. Mass spectrometry governed by a complex network of thiol- and non-reducing PAGE analysis revealed stress sensing transcription factors. the formation of an intersubunit disulfide Recently, we have found that two MarR- bond in a YvaP dimer upon treatment with type repressors, YodB and YkvE control thiol-reactive compounds in vitro. Together paralogous azoreductases, nitroreductases our data reveal that B. subtilis has evolved and three dioxygenases/glyoxalases, which a network of three MarR-type repressors respond to thiol-reactive electrophiles, YkvE (MhqR), YodB and YvaP that such as catechol, 2-methylhydroquinone control paralogous quinone resistance and diamide. These enzymes confer determinants. resistance to quinone compounds and diamide, and protect cells against the thiol- INTRODUCTION reactivity of electrophiles. In this paper, we have identified a further YodB-paralogous Bacillus subtilis is exposed in the soil to a thiol-stress sensing MarR-type regulator, variety of antimicrobial agents that also YvaP as repressor of the previously include phenolic compounds, which are characterized yfiDE (catDE) operon which produced by other soil bacteria, plants or encodes a catechol-2,3-dioxygenase and a fungi. Catecholic or hydroquinone-like DoxX-like oxidoreductase. Phenotype intermediates are common in the metabolism

CatR (YvaP) 43 Huyen et al., 2008a of nitroaromatic compounds or organic reaction as catalyzed by the thioredoxin- insecticides (Vaillancourt et al., 2006). They thioredoxin reductase system (Leelakriangsak are metabolized by specific ring-cleavage et al., 2008). Thus, YodB was identified as dioxygenases, the interdiol or extradiol novel MarR/DUF24-family repressor which dioxygenases which cleave the aromatic ring senses thiol-reactive compounds via the ortho and meta to the hydroxyl substituents, conserved Cys6 residue which was also respectively. Catechol and hydroquinone-like essential for YodB-dependent repression in compounds can also cause toxic effects in vivo. Together, the MarR-type repressors microorganisms since these can be YodB and MhqR control paralogous autooxidized to ortho- and para- azoreductases (AzoR1 and AzoR2), benzoquinones. Quinones can exert their nitroreductases (YodC and MhqN) and thiol- toxicity as oxidants and/or electrophiles dependent dioxygenases (MhqA, MhqE and (O`Brian, 1991; Monks et al., 1992; Kumagai MhqO) (Töwe et al., 2007; Leelakriangsak et et al., 2002). As oxidants, quinones redox al., 2008; Antelmann et al., 2008). The cycle with their semiquinones producing paralogous azoreductases are major quinone reactive oxygen species such as superoxide resistance determinants and have common anion or hydrogen peroxide. As electrophiles, functions as quinone reductases thus quinones can form S-adducts with cellular preventing toxic quinones from undergoing thiols via the thiol-(S)-alkylation reaction redox-cycling and causing depletion of which is an irreversible thiol-modification. cellular thiols. The paralogous thiol- However, quinones are also biologically dependent dioxygenases (MhqA, MhqE and active compounds that function as lipid MhqO) share similarity to the electron carriers in the electron-transport chlorohydroquinone/hydroquinone 1,2- chain (e.g. ubiquinone, menadione) (Unden dioxygenase of Sphingomonas paucimobilis and Bongaerts, 1997). Thus, quinones are LinE (Miyauchi et al., 1999; Endo et al., naturally occurring thiol-reactive compounds 2005). Also MhqA and MhqO confer that are ubiquitously distributed in bacterial resistance to quinone-like electrophiles and systems. could be involved in the specific ring- Recently, proteomic and transcriptomic cleavage of S-quinone adducts which are approaches revealed that catechol and 2- formed in the detoxification reaction of the methylhydroquinone (MHQ) act like diamide LMW thiol buffer with quinones (Töwe et al., as thiol-reactive compounds in B. subtilis. 2007). Thus, quinone compounds and diamide have Our previous studies identified another thiol- the same mode of action causing depletion of stress inducible yfiDE (catDE) operon, which protein and low molecular weight (LMW) encodes an enzyme with catechol-2,3- thiols. This leads to induction of the complex dioxygenase activity (CatE) and a DoxX-like thiol-specific stress response in B. subtilis oxidoreductase CatD (Tam et al., 2006; Duy which is regulated by Spx, CtsR, PerR, CymR et al., 2007). In this study, we report on the and the novel MarR-type repressors MhqR identification and characterization of the (YkvE) and YodB (Nakano et al., 2003; thiol-stress MarR/DUF24 repressor YvaP, Nakano et al., 2005; Zuber, 2004; which regulates this yfiDE (catDE) operon in Leelakriangsak et al., 2007; Leelakriangsak B. subtilis and is strongly related to the YodB and Zuber, 2007; Töwe et al., 2007; repressor as judged by sequence similarity Leelakriangsak et al., 2008; Antelmann et al., and function. We further show the importance 2008). Recently, we further showed that DNA of the conserved sensing Cys7 and the basic binding activity of YodB is directly regulated amino acids K19, R20, R49A, R54A, K56A, via thiol-(S)-alkylation in response to R75A and K83A for regulation of DNA quinone-like electrophiles in vitro which is an binding activity of YvaP in vivo. Non- irreversible thiol-modification that cannot be reducing PAGE and mass spectrometry reduced using thiol-disulfide exchange analyses revealed the formation of

CatR (YvaP) 44 Huyen et al., 2008a intersubunit disulfide bonds in YvaP dimers downstream yvaO gene and the upstream by quinones and diamide as regulatory yvaQ gene. Homology comparison revealed mechanism in vitro. that YvaP belongs to the MarR/DUF24- family of transcriptional regulators and is RESULTS closely related to YodB. These sequence comparisons showed that the start codon of 1. Identification of YvaP as a repressor of yvaP is wrong annotated and the correct start the catE-lacZ fusion using transcription ATG was found 250 bp further downstream factor (TF) arrays (Fig. S1). In addition, sequencing of the yvaO-yvaP-yvaQ region revealed an error in Previously, we have identified four thiol- the stop codon. The previously annotated stop dependent dioxygenases encoded by mhqA, codon had a wrong inserted guanine residue mhqE, mhqO and yfiE (catE) that all respond at nucleotide position 574 which caused a to thiol-specific stress conditions, such as frame shift. After correction of the diamide, catechol and MHQ (Tam et al., sequencing error, the correct stop codon is 2006; Duy et al., 2007). Moreover, three of repositioned 131 bp upstream of the previous these thiol-dependent dioxygenases (MhqA, stop codon (Fig. S1). Consequently, the MhqE and MhqO) are controlled by the novel corrected yvaP gene consists of 327 bp and MarR-type regulator MhqR (YkvE) (Töwe et encodes a protein of 108 aa which shares 46% al., 2007). In addition, the fourth dioxygenase sequence homology to the YodB repressor CatE was shown to function as catechol-2,3- (Fig. 2). dioxygenase in vitro (Tam et al., 2006). However, the specific regulator of the catDE 2. The ΔyvaP mutant shows a catechol operon was unknown. To identify the resistance phenotype regulator of the yfiDE operon, we have applied the transcription factor/transformation Previous phenotype analyses showed that the array technology (TF array) (Hayashi et al., growth of ΔyfiE deletion mutant was impaired 2006; Kobayashi, 2007). Using this TF array in the presence of 3.6 mM catechol which technology, we have previously identified reduced the growth rate of the wild-type MhqR (YkvE) as a repressor of the thiol- strain only slightly (Tam et al., 2006). Thus, dependent dioxygenase encoding mhqE the yfiDE operon confers resistance to (yodE) (Töwe et al., 2007). Chromosomal catechol. Examination of the growth DNA of each of the 287 transcription factor phenotype of the ΔyvaP mutant after deletion mutants was transformed into the treatment with diamide, catechol and MHQ ΔyfiE mutant which carries a yfiE-lacZ fusion showed that the ΔyvaP mutant was able to due to integration of the non-replicating grow with 9.6 mM catechol, concentrations pMutin4 plasmid. The ΔyvaP deletion mutant which are toxic to wild-type cells (Fig. 1B). showed derepression of the yfiE-lacZ fusion Thus, likewise to the ΔyodB and ΔmhqR on LB plates containing Xgal (5-bromo-4- mutants, also the ΔyvaP mutant showed a chloro-3-indolyl-β-D-galactopyranoside) catechol resistance phenotype. However, in (Fig. 1). Thus, YvaP was identified as a contrast to the ΔmhqR and ΔyodB mutants, candidate repressor, which could be involved the ΔyvaP mutant strain showed no increased in negative regulation of the yfiDE operon in resistance to MHQ and diamide suggesting response to thiol-specific stress conditions. specific detoxification functions of the YvaP- However, when we identified YvaP as a controlled thiol-specific dioxygenase CatE for repressor of the yfiDE operon, we noticed that catechol-like electrophiles. the coding region of the annotated yvaP 3. Sequence comparison of YvaP to YodB sequence according to SubtiList and other MarR/DUF24 family members (http://genolist.pasteur.fr/SubtiList/) overlaps of transcriptional regulators with that of the coding regions of the flanking

CatR (YvaP) 45 Huyen et al., 2008a

Fig. 1. (A) Transcription factor ABarray using a catE-lacZ fusion identifies YvaP as a repressor catE(yfiE)-lacZ-TF array of catDE. The individual colonies are transformants of transcription factor deletion mutants carrying the catE-lacZ ydcD ykvN yvaP fusion which were plated on X- 10 gal plates. The deletion of the ydcN yobV yvaV regulatory gene is indicated. (B) 1 The ΔyvaP mutant is resistant

500 to catechol stress. B. subtilis ydeB yodB yvbF OD wild type (wt) and the ΔyvaP 0.1 wt 4.8 mM cat mutant were grown in minimal ydeP yrhO yvlC ΔyvaP 9.6 mM cat medium to an OD500 of 0.4 and 0.01 treated with 4.8 and 9.6 mM ydhR yrzC ywgC -180 -120 -60 0 60 120 catechol at the time points that Time (min) were set to zero.

YodB was previously characterized as the regulator of the yfiDE operon that shows thiol-stress sensing repressor of the strong sequence homology to YodB (Fig. 2). MarR/DUF24 family which is involved in YvaP also shares the conserved Met-Cys-Pro regulation of spx, yodC and azoR1 residues, which are common for YodB- transcription (Leelakriangsak et al., 2008; paralogs found among the taxon firmicutes. Antelmann et al., 2008). MarR/DUF24-like Cys6 was postulated as sensing Cys for YodB regulators are conserved among low GC in response to thiol-reactive electrophiles as Gram-positive bacteria. These all share at also shown by mutagenesis studies. Since least one conserved Cys residue in the N- YodB and YvaP both respond to thiol-stress terminal part and one or two Cys residues in conditions in vivo and share strong sequence the C-terminal part at less conserved positions similarities, these could be regulated via (Fig. S2). Moreover, in the most closely similar mechanism which was also subject of related MarR/DUF24-like regulators, the this study. conserved N-terminal Cys is preceded by conserved Met or Leu and Pro residues. 4. Transcription of the yfiDE (catDE) Those YodB paralogs are widely distributed operon is induced by MHQ, catechol and among the taxon firmicutes including other diamide in the wild-type strain and Bacillus species, such as B. cereus, B. derepressed in the ΔyvaP mutant anthracis, B. halodurans or B. licheniformis as well as species of the genus Listeria, In previous Northern blot experiments, the Staphylococcus, Enterococci or Clostridia expression and transcriptional organization of (Fig. S2). Moreover, the genome of B. subtilis the yfiDE operon were analyzed (Duy et al., encodes at least 8 MarR/DUF24 family 2007). These data showed that yfiD is regulators, most of which have unknown cotranscribed with the yfiE gene and strongly functions (YybR, YdeP, YkvN, YtcD, YdzF, induced in response to quinone-like HxlR, YodB and YvaP) that are listed in the electrophiles (catechol and MHQ) and also DBTBS database for transcriptional diamide (Fig. 3A). However, in contrast to regulation of B. subtilis (http://dbtbs.hgc.jp/) the MhqR-regulated dioxygenases MhqA, (Fig. 2). We have previously shown that the MhqE and MhqO that have been detected MarR/DUF24 member YodB senses thiol- also at the proteome level, we could not stress conditions such as quinone compounds detect induction of the catechol-2,3- via thiol-(S)-alkylation at the conserved Cys6. dioxygenase CatE in the proteome (Tam et Here, we have identified one further al., 2006; Duy et al., 2007). Northern blot MarR/DUF24 family member, YvaP as a analyses further revealed that the catDE

CatR (YvaP) 46 Huyen et al., 2008a

Fig. S1. Corrected annotation of the yvaP DNA sequence and the translated YvaP protein sequence. The start codon is placed 250 bp downstream of the previously annotated ATG. The stop codon is located 131 bp upstream in relation to the wrongly annotated stop codon after correction of the sequencing error.

operon is derepressed in the ΔyvaP mutant To identify the cis-acting sequences which which suggests that YvaP is a repressor of the function as operator sites for repression by catDE operon (Fig. 3A). However, we YvaP, DNase I footprinting analysis was observed still some induction of catDE performed of purified YvaP protein on the top transcription in the ΔyvaP mutant after and bottom strands of the yfiDE promoter exposure to quinone-generating compounds using a yfiDE promoter DNA probe labeled MHQ and catechol which points to additional with polynucleotidkinase at the 5’ end (Fig. control mechanisms. We next determined the 4A and B). Purified His-tagged YvaP protein transcriptional start site of the yfiDE operon protected a region overlapping the yfiDE using total RNA of wild type and ΔyvaP promoter from positions -56 to +12 relative to mutant cells which were exposed to MHQ the transcription start site. Based on these and catechol stress using primer extension footprinting results, we have searched for experiments (Fig. 3B). The 5’ end of the inverted repeat sequences in the YvaP- yfiDE specific mRNA was mapped at adenine protected regions (Fig. 4B). The YvaP- located 36 bp upstream of the start codon protected regions contain two 6-1-6 bp (Fig. 4A). This transcriptional start site is imperfect inverted repeat sequences that preceded by σA-type promoter sequence. overlap the transcription start point (BS1) and Transcription of yfiDE is strongly initiated the -35 promoter region (BS2) with the from this σA-dependent promoter in response consensus of TTACtTtAaGTAA which was to thiol-specific stress conditions (MHQ, designated as YvaP box (Fig. 4B). This YvaP catechol and diamide). box consensus was used to search the B. To identify other genes that are regulated by subtilis genome for additional YvaP boxes the YvaP repressor, microarray experiments using the “Search Pattern” function of the were performed to compare gene expression SubtiList data base in B. subtilis ΔyvaP mutant cells to that of the (http://genolist.pasteur.fr/SubtiList). wild type strain in minimal medium during However, no additional sequences were the exponential growth phase. However, these detected in the promoter regions of other B. transcriptome comparisons showed that only subtilis genes without mismatches to this the genes of the yfiDE operon were 18-24- YvaP consensus sequence (data not shown). In addition, previous microarray data (Tam et fold reproducible up-regulated in the ΔyvaP al., 2006; Duy et al., 2007) do not show an mutant (data not shown). This suggests that increased transcription of the yvaP gene in the yfiDE operon is the only target for response to thiol-specific stress conditions. repression by YvaP in B. subtilis. These previous results and the lack of the YvaP consensus sequence in the yvaP 5. DNA binding experiments of YvaP to the promoter indicate that yvaP is not subject to yfiDE promoter autoregulation. Thus, we can conclude that

CatR (YvaP) 47 Huyen et al., 2008a

Fig. 2. ClustalW format alignment of MarR/DUF24- family repressors of B. subtilis including YodB, YvaP, YdeP, YybR, YdzF, YtcD, YkvN, HxlR. The Cys residues are marked in red in all YodB paralogs of B. subtilis. Identical amino acid residues are shown by stars and conserved amino acid substitutions are marked by two dots.

the YvaP regulon consists only of the yfiDE could also serve a sensory function for thiol- operon. reactive electrophiles in B. subtilis.

6. DNA binding activity of YvaP is 7. The conserved Cys7 residue of YvaP inhibited by thiol-reactive compounds and serves as an electrophile sensor in vivo H2O2 In a recent study, we showed that mutation of Catechol and MHQ are not directly the conserved Cys6 of YodB affects DNA electrophilic, but become electrophilic after binding activity of YodB in vivo and in vitro auto-oxidation to toxic ortho- and para- to its target promoter. This suggests that Cys6 benzoquinones which form S-adducts with serves a sensory function for quinone-like cellular thiols by a thiol-(S)-alkylation electrophiles. However, repression was not reaction (Leelakriangsak et al., 2008; fully relieved in the YodBCys6 mutant since Antelmann et al., 2008). Previously we have also the Cys6 and triple Cys6, 101, 108A proposed that the YodB repressor might be mutants still responded to quinones. In regulated via thiol-(S)-alkylation by quinone- contrast to YodB, the YvaP protein lacks the like electrophiles in vitro which lead directly C-terminal Cys residues present in YodB and to inhibition of DNA binding activity of shares only the conserved Cys7, which is YodB. Thus, gel mobility shift assays were preceded by conserved Met and Pro residues performed to analyze whether DNA binding (Fig. 2, Fig. S2). Thus, we have investigated activity of YvaP is also directly affected by the role of Cys7 of YvaP in regulation and thiol-reactive compounds or reactive oxygen DNA binding activity. In addition, the species. Two shifted DNA-YvaP binding reactivity of the thiolate anion is often complexes were detected with increasing influenced by neighboring basic arginine and concentrations of YvaP which accounts for lysine residues. Thus, we have further the two operator sites in the yfiDE promoter generated YvaP alleles with point mutations that were protected by YvaP in the DNase I in conserved arginine and lysine residues that footprinting experiments. The gel shift were ectopically placed into the thrC locus analyses further showed that DNA binding into the yvaP mutant. Specifically, we have activity of YvaP to the yfiDE promoters is generated YvaPR9A, K19A, R20A, R35A, directly inhibited in the presence of thiol- R49A, R54A, K56A, R75A and K83A point reactive electrophiles (MHQ, benzoquinone, mutants. Of these conserved basic amino acid diamide) and H2O2 (Fig. 5). This indicates residues R49, K56 and R75 were shown that the conserved Cys7 residue of YvaP before to interact with the DNA (Hong et al.,

CatR (YvaP) 48 Huyen et al., 2008a

Fig. S2. ClustalW format alignment of YodB/YvaP-related MarR/DUF24-family repressors among Gram- positive bacteria. The Cys residues are marked in red in all YodB paralogs conserved among low GC-content Gram- positive bacteria including members of the genus bacillus, Listeria, Clostridia, Enterobacter and Lactobacillus. Identical amino acid residues are shown by stars and conserved amino acid substitutions are marked by two dots.

CatR (YvaP) 49 Huyen et al., 2008a

AB

T A -10 T A C 168 ΔyvaP T A ΔyvaP co dia MHQ Cat co dia MHQ cat G 168 C T T G C A co co M C D T A catDE (yfiDE) C A T T A A G T G

Fig. 3. Northern blot (A) and primer extension analysis (B) of catDE transcription in the wild type in response to MHQ, catechol, diamide treatment and derepression in the ΔyvaP mutant. For Northern blot experiments and primer extension analysis, 5 µg RNA each were applied isolated from the B. subtilis strains before (co) and 10 min after the exposure to 0.33 mM MHQ, 2.4 mM catechol (cat), 1 mM diamide (dia). The arrows point toward the size of the catDE specific transcripts. The 5’ end of the catD specific transcripts is marked by an arrow and the -10 promoter sequences are labeled with a solid line. For dideoxynucleotide sequencing, the dideoxy nucleotide added in each reaction is indicated above the corresponding lane.

2005). All YvaP Cys, Lys and Arg point YvaP since these point mutants showed a mutants were analyzed for DNA binding similar derepression level as the ΔyvaP activity and inducer responsiveness by mutant under control conditions. Only the quinone compounds using yfiDE specific R9A and R35A point mutants had a similar transcriptional analysis in Northern blot basal level of yfiDE transcription as the yvaP experiments (Fig. 6). complemented strain. Like the ΔyvaP mutant, As already shown in Fig. 3, the level of the also the C7A, K19A, R20A, R49A, R54A, yfiDE specific transcript was derepressed in K56A, R75A and K83A point mutants the untreated ΔyvaP mutant but also the showed a similar further induction by thiol- ΔyvaP mutant showed some 1.5-2-fold reactive compounds suggesting other levels induction of yfiDE transcription by catechol of control as already mentioned. In and MHQ stress (Fig. 6). Next, we compared conclusion, the Northern blot results of yfiDE transcription of the yfiDE operon in the transcription in the yvaP point mutants ΔyvaP mutant strain complemented with the revealed that Cys7 has a sensory function in yvaP allele to the yvaP point mutants. The regulation of DNA binding activity of YvaP basal level of yfiDE transcription was about and that the conserved basic arginine and 3-fold higher in the untreated yvaPC7A lysine residues except for R9 and R35 are mutant strain compared to the yvaP also essential for DNA binding activity of complemented strain and showed a similar YvaP. derepression level like the ΔyvaP mutant. This confirms the previous hypothesis that 8. Non-reducing SDS-PAGE and mass Cys7 of YvaP is essential for sensing thiol- spectrometry analyses identified reactive compounds in vivo. In addition, the intermolecular disulfide bond formation in conserved basic amino acids including K19, YvaP dimers in response to quinones and R20, R49, R54, K56, R75A and K83A were diamide in vitro also essential for DNA binding activity of

CatR (YvaP) 50 Huyen et al., 2008a

A C G T - 0.05 0.1 0.2 0.3 0.4 µM YvaP A C G T - - 0.05 0.1 0.2 0.3 0.4 µM YvaP

A B

-54 +1 BS2 -10 YvaP -35 BS1

-35 BS1 YvaP -10

+1 -54 BS2 +10

-35 -10 +1 C TTTTCGTAAAATGTAACAAAATTACTTGACGTAAACAAGTTATGTATATATACTAGCTTACATTAAGTAAGTGATAACATTTATCAAGGAGGATAAA BS1 BS2 BS1 TTACTTGACGTAA BS2 TTACATTAAGTAA Cons TTACtTtAaGTAA YvaP-box Fig. 4. Binding of purified YvaP using DNase I footprinting experiments to the top (A) and bottom strand (B) of the catDE promoter and YvaP box alignments (C). The YvaP-protected sequences in the promoter regions are indicated on the right side (A, B) and labeled by arrows in the sequence alignments (C). The positions relative to the transcriptional start site are shown on the left. Transcription start site (+1) is indicated by asterisk. For dideoxynucleotide sequencing, the dideoxy nucleotide added in each reaction is indicated above the corresponding lane. (C) Regions protected by YvaP are shown in the yfiD promoter sequence grey-shaded including two conserved inverted repeats (YvaP box) which are labeled with BS1 and BS2.

Previously, we have identified specific formation of intersubunit disulfide bonds quinone-S-adducts, which are formed upon (Fig. 7). Interestingly, we observed that treatment of the purified YodB repressor with quinone compounds and diamide lead to catechol and MHQ in vitro (Leelakriangsak et formation of intersubunit disulfides in a 32 al., 2008). Thus, we were interested in the kDa YvaP dimer in vitro. These disulfide- mass spectrometry analysis of the YvaP linked YvaP-dimers can be reduced in the repressor to identify the specific Cys7 presence of DTT to the 16 kDa YvaP modification that prevents repression of the monomers. Thus, our analyses showed that catDE operon. In following, the His-tagged YvaP is regulated by reversible disulfide YvaP protein was treated with quinones bond formation in response to quinones and (catechol, MHQ and BQ) and diamide, and diamide. We further confirmed the formation the modified YvaP protein was analyzed of the intermolecular disulfide bond in YvaP using non-reducing SDS-PAGE for the by FTICR-MS (Fourier transform ion

CatR (YvaP) 51 Huyen et al., 2008a

yfiDE-promoter quinones cause predominantly irreversible thiol-(S)-alkylation and even aggregation

MHQ BQ dia H2O2 with cellular thiol-containing proteins in vivo - - - - 0.5 1 2 0.1 0.2 0.5 1 2 1 10 100 mM - 12 25 50 100 100 100 100 100 100 100 100 100 100 100 100 nM YvaP (Liebeke et al., 2008), the in vivo mechanism for YvaP regulation could also involve thiol- (S)-alkylation which has to be resolved in C2 future studies.

C1 DISCUSSION

F In this study, we have identified a further thiol-stress sensing MarR-type repressor, YvaP which controls the catechol-2,3- Fig. 5. DNA binding activity of YvaP is directly dioxygenase encoding catDE operon. Thus, inhibited by thiol-reactive compounds (MHQ, we rename YvaP herewith as CatR repressor. catechol, diamide, H2O2). Gel mobility shift assays were used to analyze the effect of thiol-reactive CatR is highly similar in sequence and compounds and oxidative stress on DNA-binding function to the YodB repressor (Fig. 2) that activity of YvaP to the catDE promoter. The YvaP controls the azoreductase AzoR1(YocJ), the protein amounts used for the DNA binding reactions nitroreductase (YodC) as well as Spx were 100 nM. The concentration of MHQ, (Leelakriangsak et al., 2008). The CatR benzoquinone (BQ), diamide (dia) and H2O2 used are indicated. controlled target genes were defined as the catDE operon. CatE was previously cyclotron resonance mass spectrometry). In characterized as thiol-dependent catechol 2,3- the untreated YvaP protein, the 548.2 Da dioxygenase which catalyzes the extradiol mass peak corresponds to the double charged cleavage of catechol to 2-hydroxymuconic T1 peptide (m/z=1094.44+2H+) which semialdehydes in vitro. CatD is similar to the showed that the N-terminal Cys7 peptide membrane-bound DoxX- and DoxD-subunit (sequence MNQSEMCPR) is in the reduced of the thiosulfate:quinone oxidoreductase of form (Fig. 7A). Acidianus ambivalens involved in the early The treatment of YvaP with 1mM MHQ, 10 steps of sulfur oxidation (Müller et al., 2004). mM catechol and 1mM BQ resulted in the Thus, CatE is the fourth thiol-dependent disappearance of the reduced T1 peptide and dioxygenase of B. subtilis which is the appearance of three new mass peak of paralogous to the three MhqR-regulated 547.97 Da, 551.97, 555.97 and 559.97 Da dioxygenases MhqA, MhqE and MhqO that (Fig. 7B-D, left panels). Interestingly, the are co-regulated by the MhqR repressor in 547.97 Da peak corresponds to the four-fold response to thiol-stress conditions. In charged disulfide-bounded T1 dimer addition, CatD is paralogous to the MhqR- (m/z=2187.88+4H+). The other newly regulated DoxX-like oxidoreductase MhqP. observed mass peaks corresponded to one, Furthermore, the MarR-type repressors YodB two and three additional oxygen masses of and MhqR control also paralogous 16, 32 and 48 Da which are representative for azoreductases AzoR1 and AzoR2 as well as the formation of 1-3 methionine sulfoxides in paralogous nitroreductases, YodC and MhqN the T1 dimer (m/z=2203.88+4H+, (Töwe et al., 2007; Leelakriangsak et al., m/z=2219.88+4H+, m/z=2235.88+4H+). We 2008; Antelmann et al., 2008). These were unable to find Cys-S-adduct formation paralogous azoreductases are major quinone of YvaP with quinone-compounds as resistance determinants and have common previously shown for the YodB repressor. functions as quinone reductases thus These results show that quinones and diamide preventing toxic quinones from undergoing cause intermolecular disulfide bonds in redox-cycling and causing depletion of YvaP-dimers in vitro. However, since cellular thiols. Thus, both YodB and YvaP

CatR (YvaP) 52 Huyen et al., 2008a

ΔyvaP YvaP C7A R9A K19A R20A R35A R49A R54A K56A R75A K83A

co M co M co M co M co M co M co M co M co M co M co M co M

7 6 5 4 3 2

co M co M co M co M co M co M co M co M co M co M co M co M ΔyvaP YvaP C7A R9A K19A R20A R35AR49A R54A K56A R75A K83A

Fig. 6. Role of YvaP Cys7, Arg and Lys mutants on regulation of catD transcription in vivo. Quantitative Northern blot analysis of catD transcription using RNA extracted from ΔyvaP mutant, and the ΔyvaP mutant complemented with the yvaP allele as well as the yvaPC7A, yvaPR9A, yvaPK19A, yvaPR20A, yvaPR35A, yvaPR49A, yvaPR54A, yvaPK56A, yvaPR75A and yvaPK83A point mutations before (co) and after treatment with 0.5 mM MHQ (M) for 10 min. The diagram shows the quantification of the catDE specific transcript in relation to the control level of the catDE transcript in the ΔyvaP mutant complemented with the yvaP allele (YvaP) which was set to 1. repressors control together azoreductases, of B. subtilis was HxlR, which is an activator nitroreductases, DoxX-like oxidoreductases of the hxlAB operon that encodes the genes and thiol-dependent dioxygenases of which for the ribulose monophosphate pathway paralogous genes are controlled by the MhqR (Yurimoto et al., 2005). The 3-hexulose-6- repressor in B. subtilis. Furthermore, the phosphate-sythase HxlA and the 6-phospho- MarR-type repressors YodB, CatR and MhqR 3-hexuloisomerase HxlB are activated in confer resistance to quinone-like response to formaldehyde stress and are electrophiles. Our previous studies have involved in formaldehyde detoxification. shown weak catechol-2,3-dioxygenase (C- YodB and YvaP act both as transcriptional 2,3-O) activity for the purified CatE protein. repressors in contrast to HxlR which is Future studies should define if CatE is more activated after exposure of cells to specifically involved in the thiol-dependent formaldehyde stress. The ClustalW sequence cleavage of catechol-thiol-S-adducts that is comparison further revealed that the formed with LMW thiols. MarR/DUF24 family members YybR, YdeP Both, YodB and CatR repressors also share and YkvN are closely related to HxlR and the conserved N-terminal Cys6 and Cys7 also share a conserved N-terminal Cys residue, respectively that are essential for residue. In addition, previous transcriptome regulation and sensing of electrophile analyses and our unpublished transcriptional compounds in vivo and in vitro. This N- studies revealed that yybR and ykvN are terminal Cys residue is also conserved among induced by quinone-like electrophiles (Duy et MarR/DUF24 family members in other al., 2007). This indicates that YybR and Gram-positive bacteria, and thus it is YkvN are autoregulated as many other MarR- tempting to speculate that this Cys serves as type regulators (Wilkinson and Groove, conserved sensor for electrophile compounds. 2007). Furthermore, YybR acts as Interestingly, B. subtilis has in total 8 transcriptional activator of another quinone MarR/DUF24-family regulators (YybR, oxidoreductase in response to thiol-reactive YdeP, YkvN, YtcD, YdzF, HxlR, YodB and compounds (our unpublished data). Detailed YvaP), and of these, YodB and YvaP share future work is required to elucidate the the highest degree of homology (Fig. 2). The function and regulation of all these first identified MarR/DUF24-family regulator uncharacterized MarR/DUF24 family

CatR (YvaP) 53 Huyen et al., 2008a

A B -DTT + DTT -DTT + DTT Co 0.5 1 5 0.5 1 5mM Dia kDa Co 0.5 1 5 0.5 1 5mM MHQ kDa

70 70 55 55 40 40 D 35 35 25 25 C 15 15

YvaP+1 mM MHQ C YvaP-control D MNQSEMCPR 100 547. 23 547.97 10 0 95

90 S 95 548.22 MNQSEMCPR 90 85 85 (m/z=1095.44+2H+) 80 S 80 75

75 70 551.97 610. 19 70 65 MNQSEMCPR 65 60 60 55 (m/z=2187.88+4H+) 55 54 8. 7 2 50 50 61 0. 1 9 45 45 40 40 593.16 611. 19 35 35 30 30 54 9. 2 2 57 5. 3 4 25 541.12 25 611. 19 593.16 612.18 20 20 54 2. 8 8 Abundance Relative 555.97 594.16 54 2. 8 8

Relative Abundance Relative 54 3. 5 5 612. 18 15 562.26 15 574. 38 57 5. 3 4 59 4. 1 6 562. 26 59 1. 3 4 595. 16 54 9. 7 2 589. 23 10 10 615.14 56 2. 7 6 576.34 595.16 615.14 587.18 562. 76 616.31 55 9. 7 7 60 2. 8 2 5 550.80 577.81 609.23 616.82 557.10 57 7. 3 1 59 6. 1 6 55 7. 1 0 564.72 571.22 596.16 5 564.72 587. 18 60 9. 4 8 616.82 578. 31 583. 25 60 3. 8 2 618.14 57 2. 2 2 602.23 565.72 578.31 58 4. 9 8 59 7. 2 2 603. 82 618.14 0 0 540 54 5 550 555 560 56 5 570 57 5 580 58 5 590 59 5 600 60 5 610 61 5 620 540 545 55 0 555 56 0 565 570 575 580 585 590 595 600 605 61 0 615 620 m/z m/z Fig. 7. Non-reducing SDS-PAGE analysis reveals intersubunit disulfide bond formation in YvaP by MHQ and diamide in vitro. Purified YvaP protein (1 µg) was incubated for 30 min with 1mM MHQ (A) and 1 mM diamide (B) and subsequently either treated with DTT (+DTT) or left reduced (-DTT) and subjected to non-reducing SDS-PAGE analysis. The bands which are labeled as insets in (A) were tryptic digested and analyzed by FT-ICR-MS in the positive ion mode. In the untreated YvaP sample (band C), the double charged Cys-charged T1 peptide was mostly detected in the reduced form (m/z=548.22). (D) After treatment of YvaP with MHQ and diamide, new peaks appeared corresponding to the four-fold charged disulfide bonded T1 dimer (m/z=647.97) and one, two, three methionine oxidations of this T1 dimer (m/z=651.97; m/z=655.97; m/z=659.97). regulators and their regulons in that thiol- diamide in vivo. In addition, more detailed in stress responsive regulatory network. vitro experiments are required to show if also In this study, we show further that YodB and DNA binding activity of YvaP is DTT- CatR have similar functions by the control of reversible in vitro. Both modes of actions, thiol-stress-inducible genes required for reversible and irreversible modifications are quinone detoxification. However, the specific possible as the regulatory in vivo mechanism mechanisms of YodB and YvaP regulation for YvaP and YodB by quinone-compounds were different in our in vitro experiments. For since these can act as oxidants or the YodB repressor we could identify thiol- electrophiles. Quinones are substrates for (S)-alkylation of the conserved Cys6 residue cellular quinone reductases that generate the by quinone compounds in vitro using mass highly toxic semiquinone anions during the spectrometry which could be responsible for one-electron reduction which in turn reduce YodB regulation by quinone-like molecular oxygen producing reactive oxygen electrophiles. In contrast, the formation of species such as superoxide anion or hydrogen intersubunit disulfides in YvaP-dimers was peroxide. These ROS could lead to reversible shown by exposure to quinones and diamide and irreversible oxidations of protein thiols. which inhibits DNA binding activity of YvaP As electrophiles, quinones can form S- in vitro. Thus, our ongoing studies are adducts with cellular thiols which involves a directed to identify the specific Cys- 1,4-reductive Michael-type addition that is modification which is leads to relief of YvaP- also referred as thiol-(S)-alkylation. Since dependent repression by quinones and benzoquinone has a one-electron reduction

CatR (YvaP) 54 Huyen et al., 2008a potential that far exceeds the range for redox- diamide. The chemistry is different for cycling reactions, these are primarily quinones and diamide leading either to ROS electrophilic as shown in the yeast model formation and thiol-(S)-alkylation for (Rodriguez et al., 2004). Our data support quinones or to disulfide bond formations by that quinones do not lead to increased diamide. Thus, the in vivo mechanisms for reversible thiol-oxidations in cellular protein regulation of the YvaP and YodB repressors thiols. We further showed that quinones react can involve reversible and irreversible thiol- via the irreversible thiol-(S)-alkylation with oxidations or thiol-(S)-alkylation cellular thiol-containing proteins in vivo mechanisms. Recent data indicate that YodB which leads to irreversible aggregation and is redox-controlled via intersubunit disulfide depletion of proteins in the proteome formation in response to diamide in vivo (our (Liebeke et al., 2008). Thus, the in vivo unpublished data). This YodB mechanism is mechanism for YvaP and YodB regulation by similar to the two-Cys sensing mechanism for quinones might involve most likely thiol-(S)- OhrR of Xanthomonas campestris which also alkylation. involves an intersubunit disulfide bond However, the YodB and CatR repressors both between one N-terminal Cys22 of one subunit also respond to diamide stress, which is and one C-terminal Cys127 of the other known to cause disulfide bonds and mixed subunit (Panmanee et al., 2006). We currently disulfides with LMW thiols in B subtilis. This define the nature of the intermolecular mechanism of S-thiolation is similar to the S- disulfide bond, which is formed in the YodB glutathionylation, which is thought to protect dimer upon diamide stress using protein thiols against overoxidation to immunoprecipitation and mass spectrometry. irreversible sulfonic acids (Lee et al., 2007; Thiol-(S)-alkylation was recently discovered Hochgräfe et al., 2007). It has been further as a novel mode of redox-regulation by shown that different organic hydroperoxides electrophilic compounds in human neuronal lead to different mechanisms for regulation of cells. In neuronal cells, electrophiles activate the well-studied OhrR repressor in B. subtilis. the Keap1-Nfr2 pathway with the consequent Inactivation of OhrR by cumene induction of the antioxidant phase-2 enzymes hydroperoxide requires Cys15 oxidation that include also quinone oxidoreductase and which leads either to a sulfenamide in vitro or glutathione S-transferase (Satoh and Lipton, a mixed disulfide (S-thiolated) form of OhrR 2007). Keap1 is an adapter protein for in vivo (Lee et al., 2007). In contrast, lineolic ubiquitinating Nfr2, which drives degradation acid was shown as 100-fold more strongly of this transcription factor. Electrophilic oxidizing peroxide which leads to irreversible compounds react with key cysteine residues sulfonic acid formation and a sacrificial in the Keap1 electrophile sensor to form regulation (Soonsanga et al., 2008). Also adducts, which cause the liberation of Nfr2 other redox-sensitive regulators, such as E. and transport into the nucleus where it binds coli OxyR or Hsp33 were shown to be to antioxidant response (ARE) elements and regulated by specific Cys-modifications in stimulates phase-2 gene transcription. Thus, response to oxidative stress, including thiol- such electrophilic compounds have disulfide switches, S-nitrosylation, S- neuroprotective effects in neuronal cells. glutathionylation, sulfenic acid formation Interestingly, this Keap1 sensor has been (Barbirz et al., 2000; Graumann et al., 2001; shown to be regulated by thiol-disulfide Kim et al., 2001; Barford, 2004). However, in switches in mouse cells and by thiol-(S)- contrast to these typical redox-sensitive alkylation in human cells (Wakabayashi et regulators, the YodB and CatR repressors do al., 2004; Satoh et al., 2007; Satoh and not respond in vivo to oxidative stress but Lipton, 2007). instead, their target genes are specifically This study shows that YvaP has like YodB an derepressed in vivo by electrophilic essential conserved Cys residue that senses compounds, such as quinone compounds and electrophilic compounds. The reactivity of

CatR (YvaP) 55 Huyen et al., 2008a cysteine is often determined by neighboring concentrations: 1 µg/ml erythromycin and 5 basic amino acids which lower the cysteine µg/ml chloramphenicol. The compounds used pKa to generate the reactive cysteine thiolate were 2-methylhydroquinone (Acros), catechol anion. Stabilization of the active site thiolate (Fluka), diamide and H2O2 (Sigma). anion by neighboring basic amino acids has The ΔcatE mutant strain that harbors the been shown for the oxidative stress catE-lacZ fusion was constructed using the responsive regulators OxyR of E. coli and pMutin4 plasmid as described previously OhrR of B. subtilis (Choi et al., 2001; (Tam et al., 2006). The transcription factor Soonsanga et al., 2007). Our YvaP point (TF) deletion mutants were constructed using mutant analyses further showed that also the an overlap-extension PCR technique as conserved K19 and R20 residues were described previously (Kobayashi, 2007). The essential for DNA binding activity of YvaP cat gene was amplified from the plasmid which could determine the reactivity of the pCBB31 by PCR with primers pUC-F and Cys7 thiolate anion. It has been also shown pUC-R. Upstream and downstream regions of that neighboring basic amino acids facilitate each regulator gene were amplified by PCR the thiol-(S)-alkylation of the reactive thiolate using gene-specific primer sets F1/R1 and anion (Dennehy et al., 2006). In addition, all F2/R2 as described previously (Kobayashi, conserved basic lysine and arginine residues 2007). Specifically, the primer sets yvaP- located in the helix-turn-helix (HTH) motif F1/yvaP-R1 and yvaP-F2/yvaP-R2 were used are essential for DNA binding of YvaP as for amplification of the yvaP upstream and shown previously also for the OhrR repressor downstream regions (Tab. S2). The 5’ end of (Hong et al., 2005). primers R1 and F2 are complementary to In conclusion, we have identified a further pUC-R and pUC-F sequences, respectively. thiol-stress responsive MarR/DUF24-family Then, the three PCR fragments were mixed repressor, CatR which regulates the thiol- and used as template for a second PCR dependent catechol-2,3-dioxygnenase CatE in reaction with primers F1 and R2. The B. subtilis and confers resistance to catechol- resultant PCR fragments were used for like electrophiles. The mechanism of YvaP transformation of B. subtilis 168. regulation involved also the conserved sensing Cys7 which has been previously Construction of yvaPC7A, yvaPR9A, shown to be critical for the YodB repressor. yvaPK19A, yvaPR20A, yvaPR35A, Thus, B. subtilis has evolved a complex yvaPR49A, yvaPR54A, yvaPK56A, network of three MarR-like repressors that act yvaPR66A, yvaPR75A and yvaPK83A point in the same detoxification pathway for mutants quinone-related electrophiles in B. subtilis. To amplify the yvaP gene including the Experimental procedures flanking regions, PCR was performed with primers yvaP-EcoRI and yvaP-BamHI (Tab. Bacterial strains and growth conditions S2) using chromosomal DNA of B. subtilis 168 as template. The PCR-product was The bacterial strains used were B. subtilis 168 digested with EcoRI and BamHI restriction (trpC2), ΔyvaP (trpC2,yvaP::Cmr), Δcat enzymes and inserted into pDG795 that was (trpC2,cat::pMutin4,Emr) and yvaP point digested with the same enzymes to generate mutants which are described in Tab. S1. pDGyvaP. The yvaP sequence was verified B. subtilis strains were cultivated under by DNA sequencing. Plasmids pDGyvaPC7A, vigorous agitation at 37°C in Belitsky pDGyvaPR9A, pDGyvaPK19A, minimal medium described previously pDGyvaPR20A, pDGyvaPR35A, (Stülke et al., 1993). Escherichia coli strains pDGyvaPR49A, pDGyvaPR54A, were grown in LB for DNA manipulation. pDGyvaPK56A, pDGyvaPR66A, The antibiotics were used at the following pDGyvaPR75A and pDGyvaPK83A were

CatR (YvaP) 56 Huyen et al., 2008a produced by using the GeneTailor Site- Northern blot experiments Directed mutagenesis System (Invitrogen). All plasmids were verified by DNA Northern blot analyses were performed as sequencing for correct introduction of the described (Wetzstein et al., 1992) using RNA yvaP point mutations. Plasmids isolated from B. subtilis wild-type, ΔyvaP, pDGyvaPC7A, pDGyvaPR9A, yvaPC7A, yvaPR19A, yvaPK19A, yvaPR20A, pDGyvaPK19A, pDGyvaP20A, yvaPR35A, yvaPR49A, yvaPR54A, pDGyvaPR35A, pDGyvaPR49A, yvaPK56A, yvaPR66A, yvaPR75A and pDGyvaPR54A, pDGyvaPK56A, yvaPK83A mutant cells before (control) as pDGyvaPR66A, pDGyvaPR75A and well as 10 min after the treatment with 2.4 pDGyvaPK83A were transformed into the mM catechol or 0.33 mM 2-MHQ, thrC locus in the yvaP mutant strain with respectively. Hybridizations specific for catE selection for erythomycin-lincomysin were performed with the digoxigenin-labeled resistance. The resulting strains were RNA probes synthesized in vitro using T7 designated BM4 (thrC::yvaPC7A), BM5 RNA polymerase from T7 promoter thrC::yvaP), BM6 (thrC::yvaPR9A), BM7 containing internal PCR products of the (thrC::yvaPK19A), BM8 (thrC::yvaPR20A), respective genes using the primer sets as BM9 (thrC::yvaPR35A), BM10 described (Tam et al., 2006). (thrC::yvaPR49A), BM11 (thrC::yvaPR54A), BM12 (thrC::yvaPK56A), BM13 Primer extension experiments (thrC::yvaPR66A), BM14 (thrC::yvaPR75A), BM15 (thrC::yvaPK83A). Primer complementary to the N-terminal encoding region of catD (yfiD-PE2) was 5’- Transcription factor array analysis end labeled using T4 polynucleotide kinase 32 (Roche Diagnostics) and 50 µCi [γ P]-ATP The transcription factor array analysis was (GE Healthcare). Primer extension analysis performed as described previously (Hayashi was performed using the labeled primers as et al., 2006; Kobayashi, 2007) using described previously (Wetzstein et al., 1992). competent cells of the ΔcatE::pMutin4 strain Sequencing of the corresponding promoter which carries a catE-lacZ fusion. regions was performed as described (Sanger et al., 1977) using PCR products as templates Transcriptome analysis containing the promoter region of the respective genes amplified with primer set For microarray analysis, the B. subtilis wild yfiD-FT-for2 and yfiD-FT-PE2 (Tab. S2). type and ΔyvaP mutant were grown in Belitsky minimal medium and harvested at Expression and purification of the YvaP OD500 of 0.4. Total RNA was isolated by the protein acid phenol method as described (Majumdar et al. 1991). Generation of fluorescence- E. coli BL21(DE3)pLysS (Invitrogen) was labeled cDNA and hybridization with B. used for overproduction of the protein. The subtilis whole-genome microarrays yvaP coding sequence was amplified by PCR (Eurogentec) was performed as described using primers yvaP-PRSET-for and yvaP- previously (Jürgen et al., 2005). Two PRSET-rev (Tab. S2). The PCR products independent hybridization experiments were were digested with EcoRI and BamHI performed using RNAs from two independent restriction enzymes and inserted into the experiments. Genes showing induction ratios expression plasmid pRSETA containing the of at least three-fold in two independent codons for six N-terminal histidine residues experiments were considered as significantly that was digested with the same enzymes to induced. generate pRSETyvaP. The yvaP sequence was verified by DNA sequencing. E. coli

CatR (YvaP) 57 Huyen et al., 2008a

BL21(DE3)pLysS carrying pRSETyvaP was EDTA, 5mM MgCl2, 0.5mM DTT, 10% cultured in 1 L LB medium, and IPTG glycerol) in the presence of 0.5 mg of BSA (isopropyl-β-D-thiogalactopyranoside) (final ml-1 and 0.05 mg of poly(dI-dC) ml-1. concentration, 1 mM) was added at the mid- Concentrations of the compounds used for the log phase (OD600 of 0.8). Recombinant His- DNA binding reactions were 1-20 mM MHQ, tagged protein (His6-YfiE) was expressed in 2-50 mM catechol, 1-10 mM diamide, 1-100

E. coli at least 2h after the addition of 1 mM mM H2O2. Samples were separated by 5% IPTG and purified by Ni-nitrilotriacetic acid native polyacrylamide gel electrophoresis in chelate affinity chromatography under native 10 mM Tris, 1 mM EDTA buffer, pH 8 at conditions according to the standard 10°C and constant voltage (150V) for 6 procedures of the manufacturer (Qiagen). The hours. Gels were dried and the radiolabeled purified YvaP protein was extensively bands were visualized using phosphoimaging. dialyzed against buffer B (25 mM Tris-HCl [pH8.0], 100 mM KCl, 0.1 mM EDTA, 1 mM FT-ICR-MS analysis MgCl2, 10% glycerol) and stored at -80°C. LC-ESI MS experiments were performed DNase I footprinting analysis using an LTQ (linear ion trap) FTICR mass spectrometer (Thermo Electron Corp., San Primer yfiD-FT-for2 and yfiD-PE2 were each Jose, CA). The SEQUEST algorithm Version 5’-end labeled using T4 polynucleotide 27.12 (Thermo Electron Corp.) was used to kinase (Roche Diagnostics) and 50 µCi search for modifications of the YvaP tryptic 32 peptides. A mass deviation of 0.01 Da for [γ P]-ATP (GE Healthcare) and purified using ethanol precipitation. DNA probes for precursor ions as well as for fragment ions catE (corresponding to positions -114 to +67 and one missed cleavage site of trypsin were allowed. The search result was filtered using relative to TSS) were synthesized by PCR amplification using one 5’ end-labeled primer BioWorks 3.2 (Thermo Electron Corp.). A multiple threshold filter applied at the peptide (yfiD-FT-for2 or yfiD-FT-PE2) and the corresponding non-labeled primer, level consisted of the following criteria: (a) peptide sequence length: 7–30 amino acids; respectively (Tab. S2). Purified PCR fragments that are labeled at one 5’ end were (b) RSp 4; (c) percentage of ions: 70 (70% of used as sequencing templates. DNA probe all theoretical b- and y-ions of a peptide are experimentally found); (d) Xcorr versus purification and DNase I footprinting was performed as described previously (Töwe et charge state: 1.90 for singly charged ions, al., 2007). 2.20 for doubly charged ions and 3.75 for triply charged ions. DNA gel mobility shift assays ACKNOWLEDGEMENTS DNA fragments containing the catD promoter region were generated by PCR using the We thank the Decodon company for support primer sets yfiD-FT-for2 and yfiD-FT-PE2. with the Decodon Delta 2D software, Approximately 600 pmol of the purified PCR Sebastian Grund for the excellent technical products were end-labeled using T4 assistance and Dirk Albrecht for the mass polynucleotide kinase (Roche Diagnostics) spectrometry analysis. This work was 32 supported by grants from the Deutsche and 50 µCi [γ P]-ATP (GE Healthcare). The Forschungsgemeinschaft, the labeled DNA probes were purified by Bundesministerium für Bildung und precipitation and 20000 cpm of each probe Forschung (BACELL-SysMo 031397A), the was incubated with different amounts of Fonds der Chemischen Industrie, the purified YvaP protein for 30 min at room Bildungsministerium of the country temperature in EMSA-binding buffer (10 mM Mecklenburg-Vorpommern and European Tris-HCl pH 7.5, 100 mM KCl, 0.1mM

CatR (YvaP) 58 Huyen et al., 2008a

Union grants BACELL-Health (LSHG-CT- Schweder, T. (2005) Global expression profiling of 2004-503468) and BACELL-BaSysBio Bacillus subtilis cells during industrial-close fed-batch fermentations with different nitrogen sources. (LSHG-CT-2006-037469) to M.H. Biotechnol Bioeng 92: 277-98. Kim, S.J., Jeong, D.G., Chi, S.W., Lee, J.S., and Ryu, REFERENCES S.E. (2001) Crystal structure of proteolytic fragments of the redox-sensitive Hsp33 with constitutive Antelmann, H., Hecker, M., and Zuber, P. (2008) chaperone activity. Nat Struct Biol 8(5): 459-66. Proteomic signatures uncover thiol-specific Kobayashi, K. (2007) Bacillus subtilis pellicle electrophile resistance mechanisms in Bacillus subtilis. formation proceeds through genetically defined Expert Rev Proteomics 5: 77-90 morphological changes. J Bacteriol 189: 4920-4931. Barbirz, S., Jakob, U., and Glocker, M.O. 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CatR (YvaP) 59 Huyen et al., 2008a thiol/disulphide switch in the global regulator, Spx. and transcriptional regulation in response to electron Mol Microbiol 55: 498-510. acceptors. Biochim Biophys Acta 1320(3): 217-34. Nakano, S., Kuster-Schock, E., Grossman, A.D., and Vaillancourt, F.H., Bolin, J.T., and Eltis, L.D. (2006) Zuber, P. (2003) Spx-dependent global transcriptional The ins and outs of ring-cleaving dioxygenases. Crit control is induced by thiol-specific oxidative stress in Rev Biochem Mol Biol 41: 241-267. Review. Bacillus subtilis. Proc Natl Acad Sci USA 100: 13603- Wakabayashi, N., Dinkova-Kostova, A.T., Holtzclaw, 13608. W.D., Kang, M.I., Kobayashi, A., Yamamoto, M., O`Brian, P.J. (1991) The molecular mechanisms of Kensler, T.W., and Talalay, P. (2004) Protection quinone toxicity. Chem Biol Interact 80: 1-41. against electrophile and oxidant stress by induction of Panmanee, W., Vattanaviboon, P., Poole, L.B., and the phase 2 response: fate of cysteines of the Keap1 Mongkolsuk, S. (2006) Novel organic hydroperoxide- sensor modified by inducers. Pro Natl Acad Sci USA. sensing and responding mechanisms for OhrR, a major 101: 2040-2045. bacterial sensor and regulator of organic Wetzstein, M., Völker, U., Dedio, J., Löbau, S., Zuber, hydroperoxide stress. J Bacteriol 188: 1389-95 U., Schiesswohl, M., Herget, C., Hecker, M., and Rodriguez, C.E., Fukuto, J.M., Taguchi, K., Froines, J., Schumann, W. (1992) Cloning, sequencing, and Cho, A.K. (2005) The interactions of 9,10- molecular analysis of the dnaK locus from Bacillus phenanthrenequinone with glyceraldehyde-3- subtilis. J Bacteriol 174: 3300-3310. phosphate dehydrogenase (GAPDH), a potential site Wilkinson, S.P, and Grove, A. (2006) Ligand- for toxic actions. Chem Biol Interact 155: 97-110. responsive transcriptional regulation by members of Sanger, F., Nicklen, S., and Coulson, A.R. (1977) the MarR family of winged helix proteins. Curr Issues DNA sequencing with chain termination inhibitors. Mol Biol 8: 51-62. Review. Proc Nat Acad Sci USA 74: 5463-5467. Yurimoto, H., Hirai, R., Matsuno, N., Yasueda, H., Satoh, T., and Lipton, S.A. (2007) Redox regulation of Kato, N., and Sakai, Y. (2005) HxlR, a member of the neuronal survival mediated by electrophilic DUF24 protein family, is a DNA-binding protein that compounds. Trends Neurosci 30(1): 37-45. acts as a positive regulator of the formaldehyde- Satoh, T., Kosaka, K., Itoh, K., Kobayashi, A., inducible hxlAB operon in Bacillus subtilis. Mol Yamamoto, M., Shimojo, Y., Kitajima, C., Cui, J., Microbiol 57: 511-519. Kamins, J., Okamoto, S., Izumi, M., Shirasawa, T., and Zuber, P. (2004) Spx-RNA polymerase interaction and Lipton, S.A. (2008) Carnosic acid, a catechol-type global transcriptional control during oxidative stress. J electrophilic compound, protects neurons both in vitro Bacteriol 186: 1911-1918. and in vivo through activation of the Keap1/Nrf2 pathway via S-alkylation of targeted cysteines on Keap1. J Neurochem 104(4): 1116-31. Soonsanga, S., Fuangthong, M., and Helmann, J.D. (2007) Mutational analysis of active site residues essential for sensing of organic hydroperoxides by Bacillus subtilis OhrR. J Bacteriol 189(19): 7069-76. Soonsanga, S., Lee, J.W., and Helmann, J.D. (2008) Oxidant-dependent switching between reversible and sacrificial oxidation pathways for Bacillus subtilis OhrR. Mol Microbiol 68(4): 978-86. Stülke, J., Hanschke, R., and Hecker, M. (1993) Temporal activation of β-glucanase synthesis in Bacillus subtilis is mediated by the GTP pool. J Gen Microbiol 139: 2041-2045. Tam, L.T., Eymann, C., Albrecht, D., Sietmann, R., Schauer, F., Hecker, M., and Antelmann H. (2006) Differential gene expression in response to phenol and catechol reveals different metabolic activities for the degradation of aromatic compounds in Bacillus subtilis. Environ Microbiol 8: 1408-1427. Töwe, S., Leelakriangsak, M., Kobayashi, K., Van Duy, N., Hecker, M., Zuber, P., and Antelmann, H. (2007) The MarR-type repressor MhqR (YkvE) regulates multiple dioxygenases/glyoxalases and an azoreductase which confer resistance to 2- methylhydroquinone and catechol in Bacillus subtilis. Mol Microbiol 66(1): 40-54. Unden, G., and Bongaerts, J. (1997) Alternative respiratory pathways of Escherichia coli: energetics

CatR (YvaP) 60 Huyen et al., 2008a

Tab. S1. B. subtilis strains and plasmids used in this study.

Strain or plasmid Genotype Reference or source Strains 168 trpC2 ΔcatE trpC2catE::pMutin4 Tam et al., 2006 BM1 trpC2 yvaP::cat This study BM4 trpC2 yvaP::cat thrC::yvaPC7A This study BM5 trpC2 yvaP::cat thrC::yvaP This study BM6 trpC2 yvaP::cat thrC::yvaPR9A This study BM7 trpC2 yvaP::cat thrC::yvaPK19A This study BM8 trpC2 yvaP::cat thrC::yvaPR20A This study BM9 trpC2 yvaP::cat thrC::yvaPR35A This study BM10 trpC2 yvaP::cat thrC::yvaPR49A This study BM11 trpC2 yvaP::cat thrC::yvaPR54A This study BM12 trpC2 yvaP::cat thrC::yvaPK56A This study BM14 trpC2 yvaP::cat thrC::yvaPR75A This study BM15 trpC2 yvaP::cat thrC::yvaPK83A This study

Plasmids pDGyvaP yvaP in pDG795 This study pDGyvaR9A yvaR9A in pDG795 This study pDGyvaPK19A yvaK19A in pDG795 This study pDGyvaPR20A yvaPRK20A in pDG795 This study pDGyvaPR35A yvaPR35A in pDG795 This study pDGyvaPR49A yvaPR49A in pDG795 This study pDGyvaPR54A yvaPR54A in pDG795 This study pDGyvaPK56A yvaPK56A in pDG795 This study pDGyvaPR75A yvaPR75A in pDG795 This study pDGyvaPK83A yvaPK83A in pDG795 This study pRSETyvaP yvaP in pRSETA This study

CatR (YvaP) 61 Huyen et al., 2008a

Tab. S2. Oligonucleotide primers used in this study.

Primer Sequence yvaP-F1 TGGCAATATCTCAATAAAGAAG yvaP-R1 GTTATCCGCTCACAATTCAGGACACATTTCTGATTGGTT yvaP-F2 CGTCGTGACTGGGAAAACTTAGGTGAGATTTCTAAATGGG yvaP-R2 GCTGCCTTCTTGATTTAAGTC pUC-F GTTTTCCCAGTCACGACG pUC-R GAATTGTGAGCGGATAAC yvaP-pRSETBamfor ATGCGGGATCCCGCATGAACCAATCAGAAATGTGTCCT yvaP-pRSETHindrev TGCCGGAATTCCGGGTCAAGAAAGGAAGGGTCAATCC yfiD-3 TGTAGGAATTCCCTTATATGAAAGCCAATTTGAG yfiD-4 ATGCGGGATCCCGTTATCACTTACTTAATGTAAGCT yvaPC7A-FOR2 AGTATGAACCAATCAGAAATGGCACCTAGATTTG yvaPC7A-REV2 CATTTCTGATTGGTTCATACTTTTCACCTCG yvaP-R9A-FOR4 ACCAATCAGAAATGTGTCCTGCATTTGAAAAAG yvaP-R9A-REV4 AGGACACATTTCTGATTGGTTCATACTTTT yvaP-K19A-FOR5 AAGCAGTCGACATCTTAAGTGCACGCTGGGTCG yvaP-K19A-REV5 ACTTAAGATGTCGACTGCTTTTTCAAATCT yvaP-R20A-FOR6 CAGTCGACATCTTAAGTAAAGCATGGGTCGCTT yvaP-R20A-REV6 TTTACTTAAGATGTCGACTGCTTTTTCAAA yvaP-R35A-FOR7 AGCTCTTGAACGGGTCGCAGGCATTTAGCGAAA yvaP-R35A-REV7 CTGCGACCCGTTCAAGAGCTGAAATACGAT yvaP-R49A-FOR8 CACTTCCAAATCTAAGCGGCGCAGTTCTGTCAG yvaP-R49A-REV8 GCCGCTTAGATTTGGAAGTGCTGCTTCAAT yvaP-R54A-FOR9 GCGGCAGAGTTCTGTCAGAAGCACTGAAAGAAC yvaP-R54A-REV9 TTCTGACAGAACTCTGCCGCTTAGATTTGG yvaP-K56A-FOR10 GAGTTCTGTCAGAACGGCTGGCAGAACTTGAGC yvaP-K56A-REV10 CAGCCGTTCTGACAGAACTCTGCCGCTTAG yvaP-R66A-FOR11 AGCTTGAAGGAGTAGTGAAGGCAGATGTCATCC yvaP-R66A-REV11 CTTCACTACTCCTTCAAGCTCAAGTTCTTT yvaP-R75A-FOR12 TCATCCCTGAAACTCCGGTTGCAATCGAATATT yvaP-R75A-REV12 AACCGGAGTTTCAGGGATGACATCCCGCTT yvaP-K83A-FOR13 TCGAATATTCATTGACTGATGCAGGAAAGGCAC yvaP-K83A-REV13 ATCAGTCAATGAATATTCGATGCGAACCGG yfiD-FT_for2 CTTATATGAAAGCCAATTTGAG yfiD-PE2 AAGATAATACCTGTAATAACTCT yfiD-prom2-rev CTGCATTGCCCATGAATGG

AdhR (YraB) 62 Huyen et al., 2008b

Chapter 3

Genome wide responses to carbonyl electrophiles in Bacillus subtilis: control of the thiol-dependent formaldehyde dehydrogenase AdhA and cysteine proteinase YraA by the novel MerR-family regulator YraB (AdhR)

Nguyen Thi Thu Huyen1, Warawan Eiamphungporn2, Ulrike Mäder3, Manuel Liebeke4, Michael Hecker1, John D. Helmann2 and Haike Antelmann1

1Institute for Microbiology, 3Interfaculty Institute for Genetics and Functional Genomics and 4Institute for Pharmaceutical Biology, Ernst-Moritz-Arndt-University of Greifswald, D-17487 Greifswald, Germany 2Department of Microbiology, Cornell University, Ithaca, NY 14853-8101, USA

The global response of B. subtilis to electrophilic carbonyls MG and FA was analyzed in this study using proteome and transcriptome experiments. Using 5’ RACE experiments, W. Eiamphungporn in the group of J. Helmann identified the transcription start sites in the adhA, adhR and yraC promoter regions. Gel mobility shift assays were also carried out by W. Eiamphungporn and J. Helmann using purified AdhR to the target promoter regions. In addition, W. Eiamphungporn and J. Helmann constructed different mutants that were used in this study. H. Antelmann carried out proteome analyses and U. Mäder performed microarray experiments. The metabolome analysis was performed by M. Liebeke and M. Lalk. My contribution in this chapter includes RNA preparation for microarray experiments, microarray data analyses, detailed transcriptional analyses of FA and MG inducible genes using Northern blot experiments and phenotype analyses of adhR regulon mutants and the adhR Cys mutant.

AdhR (YraB) 63 Huyen et al., 2008b

Genome-wide responses to carbonyl electrophiles in Bacillus subtilis: control of the thiol-dependent formaldehyde dehydrogenase AdhA and cysteine proteinase YraA by the MerR-family regulator YraB (AdhR)

Nguyen Thi Thu Huyen1#, Warawan FA is mediated by both the hxlAB operon, Eiamphungporn2#, Ulrike Mäder3, Manuel encoding the ribulose monophosphate Liebeke4, Michael Lalk4, Michael Hecker1, pathway for FA fixation, and a thiol- John D. Helmann2* and Haike Antelmann1* dependent formaldehyde dehydrogenase (AdhA) and DJ-1/PfpI-family cysteine 1Institute for Microbiology, 3Interfaculty proteinase (YraA). The adhA-yraA operon Institute for Genetics and Functional and the yraC gene that encodes a γ- 4 Genomics and Institute for Pharmaceutical carboxymuconolactone decarboxylase are Biology, Ernst-Moritz-Arndt-University of positively regulated by the MerR-family Greifswald, D-17487 Greifswald, Germany regulator, YraB (AdhR). The upstream 2 Department of Microbiology, Cornell sequences of adhA, yraC and adhR University, Ithaca, NY 14853-8101, USA contained typical MerR-like promoter sequences with a 7-4-7 inverted repeat # These authors contributed equally to the (CTTAAAG-N4-CTTTAAG) between the - work. 35 and -10 elements. Electrophoretic mobility-shift experiments revealed *Corresponding author: specific binding activity of AdhR to its 1) Tel.+49-3834-864237, Fax.+49-3834- target promoters. Mutational analyses 864202, e-mail: [email protected] showed that activation of adhA-yraA 2) Tel. (+1) 607 255 6570; Fax transcription requires the conserved Cys52 (+1) 607 255 3904, e-mail [email protected] of AdhR in vivo. We speculate that AdhR is redox-regulated via thiol-(S)-alkylation by Running title: Regulation of carbonyl aldehydes and that AdhA and YraA are detoxification by YraB (AdhR) in B. subtilis specifically involved in reduction of aldehydes and degradation or repair of Key words: damaged thiol-containing proteins in B. Bacillus subtilis/thiol stress/aldehydes/AdhR/ subtilis, respectively. AdhA/YraA INTRODUCTION SUMMARY Reactive oxygen species (ROS) are produced Quinones and α,β-unsaturated carbonyls as an unavoidable consequence of the aerobic are naturally occurring electrophilic lifestyle (Imlay, 2002). The oxidants compounds that target cysteine residues generated from incomplete O2 reduction can via thiol-(S)-alkylation. We analyzed the react with cellular components, such as amino global expression profile of Bacillus subtilis acids, carbohydrates or lipids, to produce a to the toxic carbonyls methylglyoxal (MG) range of unstable oxidation products that and formaldehyde (FA). Both carbonyl break down to diffusible electrophiles compounds induce a stress response (Marnett et al., 2003; Farmer and Davoine, characteristic for thiol-reactive 2007). These secondary oxidants include α,β- electrophiles as revealed by the induction unsaturated aldehydes which are of particular of the Spx, CtsR, CymR, PerR, YodB, importance since these have two sites of ArsR, CzrA and CsoR regulons. MG and reactivity, which can lead to the formation of FA triggered also a SOS response which cyclic adducts or cross-links. Thus, indicates DNA damage. Protection against eukaroytic and prokaryotic cells have evolved

AdhR (YraB) 64 Huyen et al., 2008b defense mechanisms to detoxify ROS and crosslinking and aggregation (Liebeke et al., reactive electrophiles, and to repair the 2008). B. subtilis expresses an adaptive resulting damage. response to electrophile-stress that is Redox-sensing transcription factors respond mediated, in part, by two MarR-type to changes in the cellular redox status, caused repressors YodB and MhqR. Inactivation of either directly by ROS or as consequence of these repressors by quinones or diamide leads oxidized reactive intermediates which act as to the expression of quinone or azo- electrophiles. Such redox-sensing proteins compound reductases and thiol-specific often have conserved cysteine residues that dioxygenases. The MarR/DUF24-family serve as a primary target for oxidative or repressor YodB is alkylated in vitro at the covalent modifications. The thiol group of conserved Cys6 residue and this residue was cysteine can undergo either reversible essential for repression also in vivo. modification, such as disulfide bond Besides quinone compounds, α,β-unsaturated formation, or irreversible modifications aldehydes are highly toxic natural including oxidation to cysteine sulfinic or electrophiles, which are generated from sulfonic acids or thiol-(S)-alkylation (Giles et oxidation of amino acids, lipids and al., 2003; Marnett et al., 2003; Barford, 2004; carbohydrates (Marnett et al., 2003; Farmer Satoh and Lipton, 2007). The reaction of and Davoine, 2007). Quinone compounds electrophilic quinones, hydroquinones and and α,β-unsaturated aldehydes both react α,β-unsaturated carbonyls involves a 1,4- with cysteine thiols via the Michael addition reductive nucleophilic addition of thiols to mechanism. Methylglyoxal (MG) is the most quinones (Michael-type addition) (O`Brian, studied toxic natural α-oxoaldehyde and is 1991; Monks et al., 1992; Kumagai et al., produced from triose-phosphate intermediates 2002; Marnett et al., 2003). Thiol-(S)- as a by-product of glycolysis, and also from alkylation can function in redox signaling as amino acids and acetone (Ferguson et al., first discovered in human neuronal cells 1998; Booth et al., 2003; Kalapos, 2008). MG where it regulates transcription in response to reacts with nucleophilic centers of DNA, electrophiles (Satoh and Lipton, 2007). RNA and with the side chains of the amino Electrophiles can protect neurons through acids arginine, lysine and cysteine. The activation of the Keap-Nfr2 pathway resulting reaction of MG with arginine and lysine in the induction of the phase-2 antioxidant residues results in the formation of the widely enzymes which include glutathione reductase studied advanced glycation end-products and quinone oxidoreductase. Neuroprotective (AGE), such as argpyrimidine or MG-cross- electrophiles react with specific cysteine linked lysine dimers (MOLD) (Oya et al., residues in the Cys-rich regulatory Keap1 1999; Bourajjaj et al., 2003). The major protein via thiol-(S)-alkylation which in turn protective mechanism against the deleterious liberates the transcription factor Nfr2. Nfr2 is effects of MG in Escherichia coli is the translocated to the nucleus where it activates spontaneous reaction of MG with glutathione transcription of antioxidant enzymes (GSH) to form the hemithiolacetal, followed (Holtzclaw et al., 2004; Wakabayashi et al., by isomerization to S-lactoylglutathione and 2004; Dinkova-Kostova et al., 2005; Satoh et conversion to D-lactate by the glyoxalase I al., 2006; Satoh and Lipton, 2007). and II enzymes (Ferguson et al., 1998; Booth We have recently discovered a similar redox- et al., 2003) (Fig. 1A). regulated system in the Gram-positive Formaldehyde (FA) is another highly toxic bacterium Bacillus subtilis which regulates carbonyl compound and causes protein- the response to quinone-like electrophiles and protein and protein-DNA cross-links in vivo diamide (Töwe et al., 2007; Antelmann et al., (Orlando et al., 1997). Like MG, FA reacts as 2008; Leelakriangsak et al., 2008). Quinones an electrophile with the side chains of react with thiol-containing proteins in vivo via arginine and lysine. Since FA is a key thiol-(S)-alkylation leading to protein intermediate in the metabolism of C1

AdhR (YraB) 65 Huyen et al., 2008b

CH3 CH3 CH3 Glx-II CH3 Glx-I A +R-SH +H2O C O C O HC OH HC OH

O H C H C OH C O R-SH C O

SR SR OH

Methylglyoxal (MG) Hemithioacetal S-Lactoylthiol Lactate

NAD NADH+H H H H H AdhA +R-SH +H2O B C O HCOH CO C O

H RS HO RS R-SH

Formaldehyde (FA) S-Hydroxy- S-Formylthiol Formate methylthiol

Fig. 1. Thiol-dependent detoxification pathways for methylglyoxal (A) and formaldehyde (B) in bacteria. (A) MG reacts spontaneously with glutathione (GSH) to form hemithiolacetal, followed by isomerization to S- lactoylglutathione by the glyoxalase I enzyme. S-lactoylglutathione is substrate for the glyoxalase II enzyme and is converted to D-lactate and free GSH. This scheme is adapted from Ferguson et al. (1998). (B) In prokaryotes and eukaryotes, FA is detoxified by glutathione-dependent formaldehyde dehydrogenases (FDH) (Harms et al., 1996). FA reacts spontaneously and reversible with GSH to form S-hydroxymethylglutathione. FDH-like enzymes such as class III alcohol dehydrogenases catalyze the NAD-dependent oxidation of S-hydroxymethylglutathione into S- formylglutathione (Jensen et al., 1998). This GSH thiol ester is reversibly hydrolyzed by a S-formylglutathione hydrolase to GSH and formiate. This scheme is adapted from Jensen et al. (1998). compounds in methanotrophic and Kato et al., 2006). 3-Hexulose-6-phosphate methylotrophic bacteria, it is ubiquitously synthase and 6-phospho-3-hexuloisomerase, distributed in the environment. Thus, the key enzymes in the RuMP pathway, are prokaryotes and eukaryotes have evolved encoded by the hxlAB operon in B. subtilis. conserved pathways for FA detoxification. Interestingly, the hxlAB operon is positively These include glutathione-dependent regulated by the MarR/DUF24 family protein formaldehyde dehydrogenases (FDH) which HxlR which is closely related (39% sequence are conserved in all kingdoms of life (Harms identity) to the quinone-responsive YodB et al., 1996). FDH catalyzes the NAD- repressor (Yurimoto et al., 2005; dependent oxidation of S- Leelakriangsak et al., 2008). hydroxymethylglutathione into S- In this study, we have applied genome-wide formylglutathione. This GSH thiol ester is transcriptome and proteome profiling to study reversibly hydrolyzed by a S- the adaptive response of B. subtilis to the formylglutathione hydrolase to GSH and toxic electrophiles MG and FA. Both formate (Jensen et al., 1998) (Fig. 1B). In compounds are strongly thiol-reactive and addition, the ribulose monophosphate they induced a genetic response superficially pathway (RuMP), used for FA fixation in similar to that seen with quinones and methylotrophs, is involved in FA diamide. In addition, FA and MG strongly detoxification in many non-methylotrophs induced the adhA-yraA operon encoding a including B. subtilis (Yurimoto et al., 2005; conserved formaldehyde dehydrogenase and a

AdhR (YraB) 66 Huyen et al., 2008b cysteine proteinase of the DJ1/PfpI family. the RuMP pathway encoded by the hxlAB This operon is controlled by the novel MerR- operon (Yurimoto et al., 2005; Kato et al., family regulator YraB which belongs to a 2006). It is presently not clear if B. subtilis is sub-family of MerR/NmlR-like proteins able to detoxify MG via specific pathways which control adhA-like target genes. related to the glyoxalase system of E. coli. According to this conserved regulation of adh-like genes we rename yraB as adhR. This Global responses to MG and FA are adhA-yraA operon, along with the hxlAB similar to those with other thiol-reactive operon, confers resistance to FA. Finally, we electrophiles show that the conserved Cys52 residue of AdhR is essential for activation of Based on the physiological data reported transcription in response to electrophilic above, we decided to analyze the global carbonyls in vivo. response of B. subtilis to concentrations of FA and MG which are growth-inhibitory. B. RESULTS AND DISCUSSION subtilis was exposed to 1 mM FA, 2.8 mM MG, or 5.6 mM MG, labeled for 5 min with B. subtilis tolerates high concentrations of 35S-methionine and subjected to proteome FA and MG analysis (Fig. 3). The proteome results showed the induction of specific marker Firstly, we tested the tolerance of B. subtilis proteins ClpE, ClpC, CysK, YrhB, YxeP, wild type cells to MG and FA to define YxeK, TrxB, NfrA, YqiG and YugJ by both concentrations which cause growth-inhibition carbonyl compounds. However, these marker but are sub-lethal. Accordingly, B. subtilis proteins are induced only by the higher wild type cells were grown in Belitsky concentration of 5.6 mM MG which reduced minimal medium to an OD500 of 0.4 and also the number of viable counts (Fig. 1B). treated with 0.5-10 mM FA and 1.4-11.2 mM These marker proteins indicate the induction MG. The growth curves and survival ratios of the CtsR, CymR and Spx-regulons in B. for FA and MG are shown in Fig. 2A and 2B, subtilis which were previously shown to be respectively. As noticed for other bacteria, B. induced by thiol-reactive compounds subtilis was able to tolerate high doses of FA diamide, catechol and methylhydroquinone without any killing effect (Fig. 2A). The (MHQ) (Antelmann et al., 2008). Spx is a treatment of B. subtilis cells with 1-2 mM FA global transcriptional regulator that controls caused a decreased growth rate but did not genes that function in maintenance of thiol- affect the number of viable counts. The homeostasis under conditions of electrophile number of viable counts was significantly stress such as the thioredoxin/thioredoxin decreased by 5 mM FA indicating that this reductase encoding trxA and trxB genes concentration is toxic for B. subtilis. (Zuber, 2004). Derepression of spx in In response to MG challenge, the growth rate response to thiol-reactive electrophiles is of wild type cells was slightly reduced by controlled by the oxidative stress specific concentrations of 1.4 mM and inhibited by repressor PerR (Mongkolsuk and Helmann, 2.8 mM, but without loss of viability (Fig. 2002) and the DUF24/MarR-type repressor 2B). However, toxic effects of MG were YodB (Leelakriangsak et al., 2007). CtsR observed after exposure to 5.6 mM MG. The controls the Clp proteases which are involved tolerance level of B. subtilis to FA is in repair or degradation of damaged or comparable to that of Pseudomonas putida, misfolded proteins. The cysteine metabolism Escherichia coli or other bacilli which were repressor CymR controls sulfur metabolism able to withstand similar doses of FA without operons and several pathways leading to killing effects (Roca et al., 2007). This is cysteine formation (Even et al., 2006). consistent with the demonstration that B. Together the induction of the Spx, CtsR and subtilis is able to detoxify FA probably via CymR regulons by thiol-reactive electrophiles

AdhR (YraB) 67 Huyen et al., 2008b

ABFig. 2. Growth curves (A, upper panels) and survival ratios (B, lower panels) of B. Formaldehyde Methylglyoxal subtilis wild type cells in the 10 10 presence of FA (left) and MG (right). B. subtilis wild type cells (nm) 1 (nm) 1 were grown in Belitsky minimal 500 500 medium to an OD500 of 0.4 and OD OD 0.1 0.1 exposed to 0.5, 1, 2, 5 and 10 co 0.5 mM FA 1 mM FA 2 mM FA co 1.4 mM MG mM FA or 1.4, 2.8, 5.6 and 11.2 2.8 mM MG 5.6 mM MG 5 mM FA 10 mM FA 11.2 mM MG mM MG at the time points that 0.01 0.01 -150 -90 -30 30 90 150 210 270 -120 -60 0 60 120 180 240 300 were set to zero. Appropriate 109 109 dilutions were plated for viable 108 108 counts (CFU/ml). 107 107 106 106

105 5 10 co 104 104 1.4 mM MG co 0.5 mM FA 2.8 mM MG 103 1 mM FA 2 mM FA 3 5.6 mM MG 10 11.2 mM MG Viable counts (cfu) Viable counts (cfu) 5 mM FA 10 mM FA 102 102 -150 -90 -30 30 90 150 210 270 -120 -60 0 60 120 180 240 300 Time(min) Time(min) indicates an imbalanced thiol-redox more than 100-fold upregulated in the homeostasis due to depletion of low transcriptome experiments by FA. We also molecular weight (LMW) thiol buffers such noted maximal derepression ratios for almost as cysteine. all CymR regulon genes by FA. For the microarray experiments, we used the Transcriptional induction of selected genes same concentrations as for the proteome and operons controlled by CtsR, Spx and analysis: 1 mM FA, 2.8 and 5.6 mM MG. CymR was verified using Northern blot Upon treatment with 1 mM FA, 256 genes analyses (Fig. 4). were up-regulated and 71 genes were The thiol-reactive nature of MG is further downregulated (Tab. S1 and S2). The supported by the induction of the YodB treatment with 2.8 mM and 5.6 mM MG regulon genes yodC, spx and azoR1 which resulted in induction of 230 genes and encode quinone-resistance determinants. repression of 188 genes (Tab. S1 and S2). All However, the induction of the YodB regulon genes and proteins that were induced by FA genes was lower with MG treatment than and/or MG treatment in three biological previously noted for quinone compounds or replicates are listed in Tab. S1 and sorted diamide. YodB was shown to be redox- according to the known stress regulons in B. regulated via thiol-(S)-alkylation by quinone subtilis. compounds in vitro (Leelakriangsak et al., The microarray results verified that both 2008). In addition, the ykvN and yybR genes carbonyl compounds cause most prominently encode two further MarR/DUF24-family the induction of the CtsR, Spx, CymR and regulators that respond to MG and quinone- PerR regulons in B. subtilis. However, in compounds. Finally, the yfiDE (catDE) accordance with the proteome results, only operon was induced by MG and quinones that the growth-inhibitory concentration of 5.6 encodes a thiol-dependent catechol-2,3- mM MG caused induction of these thiol- dioxygenase and is regulated by another stress responsive regulons. Remarkably, MarR/DUF24 paralog (YvaP) which we exposure of cells to FA induced the CtsR, renamed as CatR in B. subtilis (H. Antelmann Spx and CymR regulons at ratios that exceed and coworkers, unpublished data). Thus, we those observed for other thiol-reactive hypothesize that electrophilic carbonyl compounds such as diamide, catechol and compounds react with regulatory thiols of MHQ (Leichert et al., 2003; Tam et al., 2006; several distinct redox-sensing transcription Duy et al., 2007). For example, clpE was factors.

AdhR (YraB) 68 Huyen et al., 2008b

168_co MrgAC 168_co MrgAC 168_5.6 mM MG 168_1mM FA ClpC ClpC ClpE ClpE ClpE DnaK KatA YjbG YjbG GroEL KatA GroEL AhpF YxeK AhpF YxeK YqiG YacK YrhB YrhB YqiG YxeP YacK YxeP YugJ HemH YugJ HemH TrxB CysK Cah TrxB CysK YfjR NfrA GrpE YfkO YwfI YfjR NfrA YwfI YvyD YdgI HxlA YfkO AhpC ClpP YdgI HxlA AzoR1 AhpC CH3 AzoR1 H CO Tpx MrgA C O MrgA H C O H

Fig. 3. Dual-channel images of the protein synthesis pattern of B. subtilis wild type before (green image) and 10 min after the exposure to 5.6 mM MG (A, red image) or 1 mM FA (B, red image). Cytoplasmic proteins were labeled with L-[35S] methionine and separated by 2D PAGE as described in Materials and Methods. Image analysis of the autoradiograms was performed using the Decodon Delta 2-D software. Proteins that are synthesized at increased levels in response to FA or MG stress in at least two independent experiments are indicated by red labels. Their respective induction ratios are listed in Tab. S1. Spot identification was performed using MALDI-TOF-TOF mass spectrometry from coomassie-stained 2D gels as described in the Methods section.

FA and MG also induced the PerR peroxide- 1997). It is also possible that PerR stress regulon. The transcriptional induction derepression is elicited by the reaction of of katA by MG and FA was verified using aldehydes with functionally important Cys Northern blot experiments (Fig. 4). However, residues in PerR. However, the four Cys in the proteome experiments we observed residues in PerR coordinate a tightly bound induction of MrgA but not KatA, AhpC and zinc ion and are known to be generally non- AhpF. The induction of the PerR regulon by reactive with peroxides (Lee and Helmann, FA and MG could indicate that electrophilic 2006). Thus, it seems unlikely that they carbonyl and quinone compounds produce would react strongly with electrophilic ROS (Mongkolsuk and Helmann, 2002). aldehydes, although this requires further Quinone compounds can undergo redox- testing. cycling reactions leading to the generation of FA and MG also induced the DNA damage semiquinone anions and ROS (O`Brian, responsive SOS (LexA) regulon. DNA 1991). MG has been shown to provoke free damage likely results from the known ability radical generation involving ROS and RNS as of FA and MG to cause protein-DNA cross- well as organic methylglyoxal radicals or links, for example between the amino groups cross-linked methylglyoxal radicals which of lysine and cytosine (Orlando et al., 1997). contribute also to MG toxicity mechanisms in These upregulated SOS regulon genes include eukaryotic cells (Kalapos, 2008). Similarly, the DNA repair and recombination systems FA causes cross-links between protein and uvrB, uvrC and recA and also a DNA DNA which results in the generation of alkylation repair enzyme, yhaZ. Similar secondary electrophiles which in turn could responses of DNA damage-inducible lead to ROS production (Orlando et al.,

AdhR (YraB) 69 Huyen et al., 2008b

MG FA MG FA Co 2.8 5.5 1 mM Co 2.8 5.5 1 mM

2.66 2.3 kb 1.82 clpE 1.52 1.2 kb 1.82 CtsR 1.05 hxlAB HxlR 1.52 1.05

2.66 2.0 kb 4.7 nfrA-ywcH 3.6kb 1.82 yrhE-yrhD 1.52 2.66 1.05 Spx 1.82 0.7 kb nfrA 0.58 0.48 2.66 1.82 2.0 kb 1.82 yrhFG-yrzI 1.52 1.52 1.3 kb CymR 1.0 kb 1.05 cysK 1.05

2.66 2.2 kbarsR-yqcK- ArsR 1.82 arsBC 1.52 1.05 2.66

1.82 1.3 kb PerR 1.52 katA 1.05

Fig. 4. Transcriptional analysis of selected genes that are strongly induced in the microarray analyses by FA and MG. Transcript analysis of clpE, nfrA, cysK, arsB and katA indicates the induction of the CtsR, Spx, CymR, ArsR and PerR-dependent thiol-specific stress responses by FA and MG, which overlaps with the response to diamide and quinone-like electrophiles (Antelmann et al., 2008). Transcription of the HxlR-controlled hxlAB operon is specifically induced by aldehydes. The formate dehydrogenase operon yrhED and the formate transporter operon yrhFG-yrzI respond specifically to MG but not to FA. The arrows point toward the size of the specific transcripts. enzymes were also shown in genome-wide these metal efflux systems by electrophilic expression profiling for FA in P. putida carbonyls is an adventitious consequence of (Roca et al., 2007). the fact that these systems are regulated by As has been shown also for diamide and metal-sensing repressors that have exposed quinones, several cation efflux systems were cysteine residues as part of their metal- strongly up-regulated by carbonyl stress binding sites (Morby et al., 1993; Harvie et including the ArsR-dependent arsR-yqcK- al., 2006; Liu et al., 2007; Chen and He, arsBC operon required for resistance to 2008). As(III), Sb(III), Cd(II) and Ag(I) (Moore et Finally, many genes of the SigmaD regulon al., 2005). The induction of this ars operon that are involved in motility, chemotaxis and by FA and MG stress was also verified in cell wall turnover were up-regulated by Northern blot experiments (Fig. 4). In aldehyde treatment (Serizawa et al., 2004). addition, the CzrA-dependent cation diffusion These include the flgKLM, fliDST and motAB facilitator encoding czcDtrkA operon that operons for assembly of flagella and different responds to Zn(II), Cd(II), Cu, Ni(II) and methyl-accepting chemotaxis genes Co(II) is induced by MG and FA. Finally, the (mcpABC, tlpAB, yfmS, yoaH and yvaQ). The CsoR-regulated copZA operon which is induction of the SigmaD regulon has been activated by Cu(II) (Smaldone and Helmann, observed also in response to quinone 2007) also belongs to the metal ion compounds such as MHQ (Duy et al., 2007). responsive systems that is induced by MG To visualize the overlapping expression of and FA. We speculate that the induction of genes and regulons that respond to all

AdhR (YraB) 70 Huyen et al., 2008b analyzed thiol-reactive compounds, we Carbonyl stress specifically induces the performed a hierarchical clustering analysis HxlR-controlled hxlAB operon and the using transcriptomic data of B. subtilis cells AdhR-regulated adhA-yraA operon exposed to diamide (Leichert et al., 2003), Our cluster analysis revealed also a small MHQ (Duy et al., 2007), catechol (Tam et al., subset of genes that were selectively induced 2006), MG and FA (Fig. 5). For the cluster by both carbonyls (FA and MG) but not by analysis, we have chosen a subset of 271 other thiol-reactive electrophiles. These genes genes which are more than 4-fold induced by include the known members of the HxlR either of these thiol-reactive electrophiles or regulon (Yurimoto et al., 2005; Kato et al., known to be coregulated with these strongly 2006) and genes identified here as controlled induced genes. The complete cluster shows 9 by the AdhR activator protein. major groups of genes which are arranged as One of the major routes for FA detoxification nodes with similar expression profiles. The is the RuMP pathway which is encoded by common induction of the thiol stress specific the hxlAB operon that is positively controlled CtsR, Spx, CymR, ArsR and CzrA regulons HxlR (Yurimoto et al., 2005; Kato et al., as well as the SigmaD regulon by all thiol- 2006). We observed very high induction of reactive electrophiles is shown in nodes 6, 7, this hxlAB operon (20-140-fold) and the 2 and 3. The induction of genes regulated by autoregulatory hxlR gene (3-10-fold) by both PerR, YodB, YvaP (CatR) and MhqR seems MG and FA. The induction of the hxlAB to be more specific for quinones and diamide operon was also verified using Northern blots than for FA and MG (nodes 4 and 10). In (Fig. 4) and the HxlA protein was identified contrast, FA and MG lead to selective as specific marker protein for FA and MG induction of the HxlR, AdhR and LexA stress in the proteome (Fig. 3). regulons (node 5). We also observed very high and specific- Although exposure to FA and MG elicits a induction (10-40-fold) of the adhA gene after genetic response similar in many ways to FA and MG treatment. The adhA gene other thiol-reactive compounds, there are encodes a NADH-dependent alcohol some genes that were induced specifically by dehydrogenase (Tab. S1) similar to MG only. These included the yrhED operon glutathione-dependent formaldehyde which encodes the putative formate dehydrogenases (FDH) which are dehydrogenase YrhE and the yrhFG-yrzI evolutionarily conserved among prokaryotic operon encoding the putative formate and eukaryotic organisms (Harms et al., transporter YrhG. The specific induction of 1996). Furthermore, this class of FDH-like the yrhED and yrhFG-yrzI operons can be enzymes has conserved S-nitrosoglutathione also visualized in node 1 of the cluster (GSNO) reductase activity and functions in S- analysis (Fig. 5). Northern blot analyses using nitrosothiol homeostasis and protects against a yrhG specific showed RNA probe revealed nitrosative stress in E. coli, Saccharomyces two separate transcripts of 1.3 and 2.0 kb cerevisiae and mouse macrophages (Liu et which could indicate that the yrhFG-yrzI al., 2001). In contrast, B. subtilis AdhA was transcript is subject to RNA processing (Fig. induced specifically by FA and MG, but not 4). We suspected that the specific induction under conditions of nitrosative stress (Moore of two operons involved in formate et al., 2004). We hypothesize that AdhA metabolism (yrhED and yrhFG-yrzI) by MG functions in the thiol-dependent could be caused by contamination of detoxification of aldehydes. In addition to commercially available MG with formate adhA, we observed induction of the yraB and (Kalapos, 1999). However, our quantitative yraC genes which are encoded upstream of NMR analysis of the commercially available adhA. MG revealed a composition of 88% MG, The yraC gene was removed from the 1.7% acetic acid, 2.1% methanol and 4.7% SubtiList database (previous accession propandiol but no formate (data not shown). number BG 13778) but is still included in our

AdhR (YraB) 71 Huyen et al., 2008b

MHQ/Cat > MG/FA Thiol stress regulons 4 6 7 2

1

ArsR, 2 CzrA

3 SigmaD

YodB,YvaP, 4 MhqR, PerR 10

HxlR, AdhR, 5 LexA

MG/FA > MHQ/Cat CymR, 6 Spx 5

7 CtsR, Spx

Spx, 8 HrcA

9 10 MhqR

Fig. 5. Hierarchical clustering analysis of gene expression in response to electrophiles in B. subtilis. The treatment with the electrophiles includes diamide (Dia), catechol (Cat), methylhydroquinone (MHQ), formaldehyde (FA) and methylglyoxal (MG). Genes expression data were clustered based on the expression level leading to 9 defined groups (nodes). Nodes enriched for genes that belong to the CtsR, Spx, CymR, PerR, ArsR, CzrA, YodB, CatR (YvaP), MhqR, HxlR and AdhR regulons are shown to the right of the cluster. Red indicates induction and green repression under the specific conditions of electrophile stress (see Experimental procedures for details). microarrays. Since the yraC transcript is annotated as CMD in Methanosarcina strongly induced by aldehydes in the acetivorans had protein disulfide reductase transcriptome, we have reannotated the yraC activity which depends on a CxxC motif gene as 192 bp open reading frame which (Lessner and Ferry, 2007). However, this could encode a putative 64 aa protein product CxxC motif is not conserved in YraC. CMD- with similarity to γ-carboxymuconolactone like domains are also found in the decarboxylases (CMD) (Fig. S1). These alkylhydroperoxide reductase AhpD of CMD domain enzymes are involved in mycobacteria (Bryk et al., 2002). We also protocatechuate catabolism and in some noticed that YraC is much smaller than other bacteria a gene fusion event leads to annotated CMD family enzymes (Fig. S1B), expression of CMD with a hydrolase involved and thus the function of YraC is still in the same pathway (Eulberg et al., 1998). unknown. Recently, it was reported that an enzyme The yraB gene encodes a MerR-family

AdhR (YraB) 72 Huyen et al., 2008b

Fig. S1. (A) Reannotation for the previously from the SubtiList database removed yraC gene of B. subtilis (previous accession BG13778) and (B) Clustal 2.0.8 multiple sequence alignment of YraC with putative gamma carboxymuconolactone decarboxylases. (A) The 192 bp open reading frame and the encoded putative 64 aa protein (YraC) are indicated. The promoter sequences of the yraC gene are underlined and the inverted repeat (AdhR-box) is bold-faced. (B) YraC was aligned with gamma carboxymuconolactone decarboxylase homologs of Lactobacillus fermentum (LACF3), Pediococcus pentosaceus (PEDPA), Clostridium sp. (CLOT), Staphylococcus saprophyticus (STRA3), Enterococcus faecium (ENTFC) and Streptococcus agalactiae (STRA3). regulator related to proteins which often NmlR-like regulators under conditions of NO regulate divergently transcribed adhA-like or FA stress has thus far not been described. genes. Since we here show that YraB regulates induction of adhA, we renamed The MerR/NmlR-type regulator AdhR YraB as AdhR. Interestingly, NmlR, an positively regulates the adhA-yraA operon AdhR-homolog of Neisseria gonorrhoeae, is and yraC in response to electrophilic a redox-controlled regulator of the GSNO aldehydes reductase AdhC, a Cpx-type ATPase and a thioredoxin reductase, and protects against The adhA gene is located upstream of the nitric oxide (NO) stress (Kidd et al., 2005). yraA gene which encodes a member of the NmlR-like regulators form a distinct DJ-1/PfpI-family of proteins, some of which subfamily among the MerR family and all may function as cysteine proteinases (Wei et regulate the expression of adh-like genes al., 2007). Although their functions are not (Kidd et al., 2007; Stroeher et al., 2007). In well understood, there is a wealth of addition, these MerR-type regulators have structural information for DJ-1/PfpI family conserved roles in protection against NO members and structures have been stress also in Haemophilus influenzae and determined for human DJ-1, yeast Hsp31p, Pneumococcus (Kidd et al., 2007; Stroeher et and four E. coli homologs YhbO, SCRP 27a, al., 2007). Unlike other MerR-type YajL and Hsp31. The structures of these regulators, NmlR and AdhR lack the proteins are similar; they all possess a conserved Cys residues present in metal- conserved Cys residue in the nucleophile responsive MerR-type regulators. Instead, elbow motif. Recently, Hsp31p has been these NmlR homologues (including AdhR) all shown to be involved in the protection against share the conserved Cys52 residue (B. subtilis ROS (Skoneczna et al., 2008). Interestingly, numbering) that could play a role in sensing yraA is under complex control. In addition to of aldehydes. However, the detailed its induction as part of the bicistronic adhA- mechanisms of transcriptional regulation for yraA operon (controlled by AdhR), it is also

AdhR (YraB) 73 Huyen et al., 2008b very strongly up-regulated under thiol- under conditions of aldehyde stress as specific stress conditions as part of the Spx revealed by the microarray data. regulon (Nakano et al., 2003). Expression of The transcription start sites of adhA, adhR, yraA by Spx is independent of the upstream and yraC were mapped using 5’ RACE adhA gene (Eiamphungporn and Helmann, experiments (Fig. 6B). The 5’ end of the 2008). adhR-specific mRNA was mapped at an A To study the regulation and transcriptional located 34 bp upstream of the start codon. organization of adhA, we performed Northern The transcription start point for yraC was blot experiments using RNA isolated from B. mapped at G and C located 42 and 43 bp subtilis wild type and adhR mutant cells after upstream of the ATG. Finally, the adhA-yraA exposure to FA and MG. Hybridization was specific message initiates with G or A performed using adhA, yraA and adhR- residues located 82 and 83 bp upstream of the specific RNA probes (Fig. 6A). Two ATG start codon. The -10 and -35 elements transcripts were detected using a yraA- of all three promoter regions were separated specific RNA probe, both of which were by 19 bp which is consistent with the strongly elevated under conditions of elongated spacer regions typical for MerR- aldehyde stress in the wild type. The larger family regulators (Brown et al., 2003; 1.8 kb transcript corresponds to the adhA- Hobman et al., 2005). The conserved IR yraA bicistronic transcript which is absent in sequence overlapped slightly the -35 the adhR mutant. The smaller 0.5 kb promoter region which is typical for MerR- transcript originates from a promoter that type regulators. These conserved binding sites precedes yraA and is induced in both the wild and the MerR-type promoter architecture type and adhR mutant strain. Thus, suggests that adhR, yraC and adhA-yraA transcription of yraA is induced constitute the aldehyde-inducible AdhR monocistronically in a Spx-dependent manner regulon of B. subtilis. and as part of the bicistronical AdhR- controlled adhA-yraA message. This positive AdhR binds specifically to the adhA, adhR regulation of the adhA-yraA operon by AdhR and yraC promoter regions was verified also by using an adhA-specific RNA probe. In this case, only the AdhR- Electrophoretic mobility shift assays (EMSA) dependent up-regulation of the 1.8 kb adhA- were performed using purified His-tagged yraA transcript was detected, but not the yraA AdhR protein and the adhA, adhR and yraC specific monocistronic transcript. Finally, promoter regions. As shown in Fig. 7, high using a adhR-specific RNA probe, the affinity DNA-AdhR complexes were detected autoregulation of adhR was verified by for all target promoter regions, but not for a Northern blot analyses. control DNA fragment from the yoeB Next, we used in silico analyses to identify promoter region. Like other MerR family conserved binding sites for the AdhR members, AdhR binds to its target operator regulator upstream of adhA and adhR. A sequences both with and without specific conserved 7-4-7 inverted repeat with the inducer and addition of MG or FA had little consensus sequence CTTAAAG-N4- effect on the AdhR DNA binding activity. CTTTAAG was identified in the promoter Similarly, the FA-responsive transcriptional regions of adhA and adhR. This consensus activator HxlR (Yurimoto et al., 2005) binds was used to search the genome sequence of B. to its target operator sequences with similar subtilis using the search pattern function of affinities in the absence and presence of FA the SubtiList database. In addition to the in vitro. HxlR shares similarity to the YodB adhA and adhR promoters, this conserved IR repressor and 5 other MarR/DUF24-family sequence was detected upstream of yraC (Fig. members of B. subtilis. We have shown 6B). Transcription of yraC was also induced previously shown that Cys6 of YodB is

AdhR (YraB) 74 Huyen et al., 2008b

168 ΔadhR MG FA MG FA Co 2.8 5.5 1 Co 2.8 5.5 1 2.66 1.8 kb 1.82 adhA-yraA

(A) 1.05

0.58 0.5 kb 0.48 yraA

2.66 1.8 kb 1.82 adhA-yraA

1.05 1.05 0.6 kb 0.58 adhR 0.48

adhR yraC adhA yraA (B)

-35 -10 +1 adhR tgttttgagcttgaCTTAAAGttaaCTTTAAGtgttaccttcacaacAtacaggatag-(N23)-ATG

-35 -10 +1 (C) yraC tattcgggacttgcCTTAAAGcacaCTTTAAActctatacttctccatGCactaaaaac-(N32)-ATG

-35 -10 +1 adhA taaaaaacccttgcCTTAAAGcagaCTTTAAGgtttatcgttatgttGAagacaccata-(N71)-ATG

Fig. 6. Transcript analyses (A), transcriptional organization (B), and promoter alignments of the adhR, yraC and adhA-yraA operons which are regulated by the novel MerR-type transcriptional regulator AdhR in response to carbonyl compounds. (A) For Northern blot experiments, 5 µg RNA each were isolated from the B. subtilis strains before (co) and 10 min after the exposure to 1 mM FA, 2.8 mM and 5.5 mM MG. The Northern blots were hybridized with yraA, adhA, or adhR-specific mRNA probes (shown underlined). The arrows point toward the sizes of the adhA- yraA, yraA and adhR specific transcripts. (B) Gene organization of adhR, yraC and adhA-yraA. Transcriptional start sites are indicated by bent arrows. (C) The transcription start sites of the adhR, yraC and adhA-yraA specific mRNAs were determined by 5’-RACE and are indicated as +1. The -10 and -35 promoter elements are underlined. The conserved inverted repeat sequences are shown in upper case letters. alkylated by quinone-like electrophiles in To address the question whether the vitro (Leelakriangsak et al., 2008) and this conserved Cys52 of AdhR is involved in Cys residue is also present in HxlR. We note regulation of adhA-yraA transcription, we that all AdhR-homologs in the MerR/NmlR- constructed adhR mutant strain that subfamily share a conserved Cys52 residue. ecotopically expressed a functional FLAG- Thus, we analyzed next the role of Cys52 in tagged AdhR as well as the AdhRC52A regulation of adhA-yraA transcription by mutant protein from a xylose-inducible AdhR. promoter (Pxyl) based on the pSWEET plasmid (Bhavsar et al., 2001). The adhR and Transcriptional activation of the adhA- adhRC52A complemented ΔadhR mutant yraA operon requires the conserved Cys52 strains were grown in Belitsky minimal of AdhR medium and expression of the Pxyl promoter was induced by addition of 2% Xylose (Fig. 8A). Since activation of adhA-yraA

AdhR (YraB) 75 Huyen et al., 2008b

Control 10 mM MG 10 mM FA Fig. 7. Binding of purified AdhR to the adhR, yraC and [AdhR] nM 0 5 10 20 40 80 160 0 5 10 20 40 80 160 0 5 10 20 40 80 160 adhA promoters using DNA electrophoretic mobility shift A assay. Increasing amounts of purified AdhR protein was added to the labeled adhR (A), yraC (B) and adhA (C) promoter probes. 10 mM of MG or FA were added B to analyze the effect of these compounds on the DNA binding activity of AdhR to these target promoters. (D) AdhR does not bind to an unrelated promoter C fragment (yoeB).

MG FA

0 5 10 0 5 10 mM 160 160 160 160 160 160 [AdhR] nM

D

transcription requires aldehydes, cells were 2005). It has been further shown that treated with 2.8 mM MG or 1 mM FA for 10 repression by NmlR requires Zn(II) and min after xylose addition and the isolated transcription of adhC was activated by the RNA were subjected to adhA-specific thiol reactive compound diamide. Mutation of Northern blot analyses. The Northern blot any of the four conserved Cys residues in results showed that adhA-yraA transcription NmlR resulted in constitutive transcriptional is similar strongly activated by MG and FA in repression and lack of activation of adhC wild type cells and in the adhR-FLAG transcription suggesting that NmlR acts as complemented adhR mutant strain (Fig. 8A). transcriptional repressor that responds to thiol However, no expression of adhA-yraA stress conditions (Kidd et al., 2005). transcription occurred in the adhRC52A mutant in response to aldehyde treatment. The ΔhxlR, ΔadhA and ΔyraAyfkM Western blot analyses confirmed that FLAG- mutants are sensitive to FA stress AdhR and FLAG-AdhRC52A proteins are produced in similar amounts in the adhR and Previous analyses showed that the ΔhxlR adhRC52A complemented ΔadhR mutant deletion mutant was sensitive to FA exposure strains (Fig. 8B). These results indicate that (Yurimoto et al., 2005). To determine to Cys52 is required for the activation of which extent the RuMP pathway and the FA transcription by AdhR, presumably because dehydrogenase pathway confer resistance to aldehydes modify this conserved Cys residue carbonyl compounds, growth experiments via thiol-(S)-alkylation to allosterically were performed using ΔhxlR, ΔadhR, ΔyraA activate AdhR. and ΔadhA mutants. The results showed that In contrast to AdhR, which acts as the growth rates of the ΔhxlR and ΔadhA transcriptional activator for adhA expression, mutants were significantly reduced in the the MerR/NmlR regulator of N. gonorrhoeae presence of 0.5 and 1 mM FA compared to represses transcription of adhC under normal wild type cells (Fig. 9). This indicates that conditions and adhC transcription is both FA detoxification systems contribute to derepressed in the nmlR mutant (Kidd et al.,

AdhR (YraB) 76 Huyen et al., 2008b

Fig. 8. The conserved Cys52 of CU1065 ΔadhR ΔadhR ΔadhR AdhR is essential for the adhR adhRC52A activation of adhA-yraA transcription by aldehydes. (A) Co MG FA Co MG FA Co MG FA Co MG FA For adhA-specific Northern blot analysis, RNA was isolated from 1.8 kb strains before (co) and after adhA-yraA treatment with 2% xylose for 20 A min and subsequently with 2.8 mM MG or 1 mM FA for 10 min. (B) Western blot analysis was performed using Anti-FLAG B antiserums to confirm expression of FLAG-AdhR and FLAG- AdhRC52A proteins. FA resistance of B. subtilis. However, neither protection against reactive aldehydes, of the tested mutants showed an increased presumably by repair or degradation of sensitivity to methylglyoxal (data not shown). proteins with specific thiol-modifications. B. subtilis also possesses another alcohol Previous transcriptome analyses have dehydrogenase encoded by the adhB gene revealed that yraA is most strongly induced in which is a part of the yraGF-adhB-yraED response to thiol-specific stress conditions, operon and shown previously to be under such as diamide, catechol and MHQ (Leichert control of the sporulation sigma factor, σG et al., 2003; Tam et al., 2006; Duy et al., (Steil et al., 2005). However, the yraGF- 2007). Expression of yraA was also inducible adhB-yraED operon is also induced by FA by acid, ethanol and vancomycin and yraA and MG (3-8-fold) in microarray experiments mutant cells were sensitive to acid stress (Tab. S1) which indicates that expression (Thackray and Moir, 2003; Jervis et al., does not require the sporulation sigma factor, 2007). In the case of other thiol-specific stress σG. Thus, we tested the growth of an conditions or acid stress, induction of yraA ΔadhAΔadhB double mutant after exposure to occurs in a Spx-dependent manner FA and MG. The results showed no (Eiamphungporn and Helmann, 2008). Recent differences in the growth of the ΔadhAΔadhB studies further showed that the previously M double mutant compared to the ΔadhA single suggested σ -dependent expression of yraA M mutant in the presence of FA and MG (data is derived from the σ -dependent activation not shown). Thus, adhB does not contribute of spx (Nakano et al., 2003; Eiamphungporn significantly to aldehyde resistance of and Helmann, 2008). growing B. subtilis cells. According to the DJ-1/PfpI-family homology, The yraA gene encodes a putative DJ-1/PfpI- YraA could function in repair or degradation family cysteine proteinase. Phenotype of proteins that are damaged by electrophiles analyses showed that the sensitivity of the and related toxic compounds. However, ΔyraA mutant was similar to the wild type according to previous studies, YraA had no after exposure to FA (data not shown). Since detectable protease or chaperone activity yraA displays sequence similarity to the σB- (Thackray and Moir, 2003). In ongoing dependent yfkM gene, we investigated the studies, we are using proteomic approaches growth phenotype of the ΔyraAΔyfkM double and radioactively-labeled electrophiles to mutant after exposure to FA and MG stresses. identify possible substrates for the YraA and The ΔyraAΔyfkM double mutant showed a YfkM proteases in B. subtilis. Together, our reduced growth rate after treatment with FA results show that B. subtilis encodes multiple and MG compared to the wild type and to the systems involved in resistance to FA: the yraA single mutant (Fig. 9). Thus, both HxlR-regulated RuMP pathway and the putative DJ-1/PfpI-family cysteine AdhR-controlled thiol-dependent proteinases can replace each other in formaldehyde hydrogenase AdhA and

AdhR (YraB) 77 Huyen et al., 2008b

0.5 mM FA 1 mM FA MG monitored growth (Fig. 10B,C). These 1

) medium supplementation experiments -1 0.8 revealed that wild type cells pretreated with 1-2 mM cysteine were protected against the 0.6 toxicity of 2 mM FA and growth was restored 0.4 (Fig. 10B). Also, pretreatment of cells with 3 growth rate (h mM cysteine suppressed the growth 0.2 inhibitory effect of 3 mM MG (Fig. 10C). These experiments showed that equimolar wt wt wt hxlR hxlR amounts of cysteine are able to titrate toxic adhA adhA Δ Δ Δ Δ aldehydes, consistent with the notion that yraAyfkM yraAyfkM yraAyfkM Δ Δ Δ electrophilic aldehydes target cysteine (both Fig. 9. The ΔadhA and ΔhxlR mutants are sensitive free and in proteins) and that cysteine- to FA stress and the ΔyraAΔyfkM mutant is dependent enzymes may function in sensitive to FA and MG. B. subtilis wild type (wt), detoxification. Interestingly, benzoquinone ΔadhA, ΔyraAΔyfkM and ΔhxlR mutant strains were (BQ) is toxic at concentrations of 10 µM and grown in Belitsky minimal medium to an OD500 of 0.4 requires only 20 µM Cys for detoxification. and treated with 0.5 and 1 mM FA or 1.4 mM MG. The growth rates of the strains after exposure to FA Thus, much higher concentrations of cysteine and MG are shown. are needed to titrate FA and MG compared to BQ. We used NMR analysis to test whether candidate cysteine proteinases YraA and cysteine reacts in vitro with FA and MG,. YfkM. Indeed, FA reacts completely with cysteine resulting in a thiazolidine carboxylic acid in The LMW thiol cysteine functions in vitro (Fig. S2A). The reaction of MG with detoxification of FA and MG cysteine leads to generation of the acetylated derivative of the thiazolidine carboxylic acid In previous studies we found that the LMW (Fig. S2B). thiol cysteine is decreased in the metabolome The transcriptomic results revealed DNA after treatment with quinones. In addition, damage by aldehydes which indicates that supplementation of the growth medium with aldehydes also cause cross-links between the extracellular cysteine protects cells against side chains of lysine or arginine and nucleic the lethal effect of electrophilic quinones acids. However, unlike Cys, pretreatment of (Liebeke et al., 2008). This suggests that cells with arginine and lysine did not restore cysteine may function in detoxification of growth after addition of toxic MG or FA electrophiles. concentrations (data not shown). These The results of this study also suggest that results suggest that Cys is the strongest cysteine is an important target for toxic nucleophile which is targeted in vivo by aldehydes. Thus, we measured the aldehydes. intracellular concentration of cysteine in the metabolome after exposure to FA and MG Conclusion using GC/MS analyses. Levels of cysteine were decreased about 5-fold after exposure to In this study, we have analyzed the global 0.5-1 mM FA and 5.5 mM MG in B. subtilis response of B. subtilis to the electrophilic (Fig. 10A). In addition, and in contrast to carbonyls FA and MG. The global expression quinone treatment, other metabolites were not profiles suggest that carbonyls act, like changed by FA and MG exposure. diamide and quinones, as thiol-reactive To determine if extracellular cysteine can compounds which most probably target protect cells against aldehyde toxicity, we cysteine residues via the thiol-(S)-alkylation pretreated cells with different concentrations chemistry. Thus, cysteine is also one of the of cysteine prior to FA and MG challenge and first barriers for electrophilic aldehydes as

AdhR (YraB) 78 Huyen et al., 2008b

ABA BC C

cysteine 1.2 10 10 1 0.8 1 1 0.6 (nm) ratio 500 control 0.4 0.1 FA 2 mM 0.1 control OD FA 2 mM + Cys 1 mM MG 3 mM FA 2 mM + Cys 2 mM MG 3 mM + Cys 3 mM 0.2 FA 2 mM + Cys 5 mM MG 3 mM + Cys 6 mM MG 2.8 mM 2.8 MG co FA 1.0 mM mM 5.5 MG FA 0.5 mM 0.01 0.01 -120 -60 0 60 120 180 240 300 -120 -60 0 60 120 180 240 300 Time (min) Time (min)

Fig. 10. The LMW thiol cysteine protects against aldehyde toxicity in B. subtilis. (A) Changes in the level of cysteine in the metabolome were measured using GC-MS in B. subtilis cellular extracts after exposure to FA and MG as described in Methods. Average values and standard deviations are quantified from at least three independent cultivation experiments and the ratios are related to the untreated control. (B, C) B. subtilis wild type cells were grown in Belitsky minimal medium to an OD of 0.4 and different concentrations of extracellular cysteine were added prior to exposure of FA (B) or MG (C). These experiments indicate that 1 mM cysteine is able to protect against 2 mM FA and 3 mM cysteine titrate 3 mM MG which restored the growth. supported by protection experiments with modifications poorly accessible by mass extracellular cysteine. However, in contrast to spectrometry approaches. quinones and diamide, carbonyl electrophiles display also a unique genetic response which Experimental procedures is governed by the HxlR and AdhR regulators. Both the HxlR and AdhR regulons Bacterial strains and growth conditions confer resistance to FA. AdhR is characterized here as novel regulator of the The bacterial strains used were B. subtilis 168 MerR/NmlR-family which positively controls (trpC2) and CU1065 (trpC2), ΔadhA a conserved thiol-dependent formaldehyde (trpC2,adhA::mlsr), ΔadhAΔadhB dehydrogenase (AdhA) and the putative (trpC2,adhA::mlsradhB::spcr), ΔyraA cysteine protease YraA, both of which protect (trpC2,yraA::cmr), ΔyraAΔyfkM against FA toxicity. Transcriptional (trpC2,yraA::cmryfkM::mlsr), ΔadhR regulation by AdhR requires the conserved (trpC2,adhR::kmr) which are described below. Cys52 residue which we hypothesize is B. subtilis strains were cultivated under alkylated by electrophilic carbonyl vigorous agitation at 37°C in Belitsky compounds leading to activation of the minimal medium described previously AdhR-dependent adhA-yraA, yraC and adhR (Stülke et al., 1993). E. coli strains were operons. Future studies are directed to define grown in LB for DNA manipulation. The the nature of post-translational thiol- antibiotics were used at the following modifications which are caused by aldehydes concentrations: 1 µg/ml erythromycin, 25 in AdhR as well as in cellular proteins in vivo. µg/ml lincomycin, 5 µg/ml chloramphenicol These might involve S- and 10 µg/ml kanamycin. Methylglyoxal was hydroxymethylcysteine modifications or purchased from Sigma Aldrich and protein crosslinks between different thiol- formaldehyde was ordered from Roth. containing proteins by FA. These studies are Gene deletions were generated using long- likely to be challenging since these adducts flanking-homology polymerase chain reaction may, like quinone-(S)-adducts, be unstable (LFH-PCR) as previously described (Mascher et al., 2003). Primers used to amplify the up

AdhR (YraB) 79 Huyen et al., 2008b

Fig. S2. Reaction of FA (A) and MG (B) with cysteine was confirmed in vitro using NMR spectroscopy. FA reacts with cysteine resulting in a thiazolidine carboxylic acid (A). MG reacts with cysteine leading to generation of the acetylated derivative of the thiazolidine carboxylic acid (B). NMR spectra were measured as described in the Methods section. The spectra show the resonance signals for the protons numbered for the substances in the reaction schemes. In particular, the NMR pattern of the CH2 group at position 2” of the thiazolidine ring systems clearly shows the ring closure during the reactions of formaldehyde or methylglyoxal with cysteine, respectively. The multiple signal of the 2’ CH2 group in cysteine is divided to two quartet signals for both protons. A similar pattern was observed for the methylene group 3” in the carboxy-thiazolidine formed by the carbon of FA. These signals are not present in the case of the acetylated thiazolidine resulting from the reaction of MG with cysteine. In that case the newly formed acetyl CH3 group was clearly detected in the spectrum (position 3”).

AdhR (YraB) 80 Huyen et al., 2008b and down fragments are listed in Tab. S3. separation by two-dimensional gel Fragments were amplified using Pfu DNA electrophoresis (2-D PAGE) using the polymerase (Stratagene) and were joined immobilized pH gradients (IPG) in the pH using Expand Long Template PCR System range 4-7 was performed as described (Tam (Roche). Five µl of the LFH-PCR product et al., 2006). The quantitative image analysis was introduced into the desired strain by was performed with the Decodon Delta 2D transformation. Integration and deletion of the software (http://www.decodon.com). Proteins gene were confirmed by PCR. showing an induction of at least two-fold Construction of adhR mutant containing during the 5 min of L-[35S]methionine pulse adhR-FLAG and adhRC52A-FLAG under in two independent experiments were PxylA control was performed using the considered as significantly induced. For pSWEET plasmid (a derivative of pDG364 identification of the proteins by mass containing PxylA promoter and spectrometry, nonradioactive protein samples chloramphenicol resistance gene) (Bhavsar et of 200 µg were separated by 2-D PAGE and al., 2001). To generate pSWEET containing the 2D gels were stained with Colloidal adhR-FLAG, adhR gene which was fused in Coomassie Brilliant Blue (Amersham frame with FLAG epitope tag including the Biosciences). Spot cutting, tryptic digestion predicted RBS site upstream of adhR, but of the proteins and spotting of the resulting lacking the promoter was amplified by PCR peptides onto the MALDI-targets (Voyager using primers 4151/4152 as listed in Tab. S3. DE-STR, PerSeptive Biosystems) were The resulting fragment was digested with performed using the Ettan Spot Handling PacI and BamHI and cloned into pSWEET Workstation (Amersham-Biosciences, that was digested with the same enzymes, Uppsala, Sweden) as described previously placing in adhR-FLAG downstream of the (Eymann et al., 2004). The MALDI-TOF- PxylA promoter. The pSWEET containing TOF measurement of spotted peptide adhRC52A-FLAG was generated by PCR solutions was carried out on a Proteome- mutagenesis. Using pSWEET containing Analyzer 4800 (Applied Biosystems, Foster adhR-FLAG as a template, First-round PCR City, CA, USA) as described previously was performed in two separate reactions with (Eymann et al., 2004). primers 4151/4175 and 4152/4170 (Tab. S3). The PCR products were joined by a second Transcriptome analysis round of PCR using primers 4151/4152. The product was digested with PacI and BamHI For microarray analysis, B. subtilis wild type and cloned into pSWEET that was digested cells were grown in Belitsky minimal with the same enzymes. The resulting medium to OD500 of 0.4 and harvested before plasmids were verified by DNA sequencing and 10 min after exposure to 1 mM FA, 2.8 then digested with PstI and transformed into mM and 5.6 mM MG, respectively. Total adhR::Km where fragments integrated into RNA was isolated by the acid phenol method the amyE locus. The resulting strains were as described (Majumdar et al. 1991). selected by KmrCmr and confirmed by PCR. Generation of fluorescence-labeled cDNA and hybridization with B. subtilis whole- Proteome analysis and mass spectrometry genome microarrays (Eurogentec) was performed as described previously (Jürgen et Cells grown in Belitsky minimal medium to al., 2005). Genes showing induction or an OD500 of 0.4 were pulse-labeled for 5 min repression ratios of at least three-fold in three each with 5 µCi of L-[35S]methionine per ml independent experiments were considered as before (control) and 10 min after exposure to significantly induced. The averages ratios and 1 mM FA, 2.8 mM or 5.6 mM MG. standard deviations for all induced or Preparation of cytoplasmic L- repressed genes are calculated from three [35S]methionine-labeled proteins and independent transcriptome experiments after

AdhR (YraB) 81 Huyen et al., 2008b

10 min of exposure to 2.8 and 5.6 mM MG or internal PCR products of the respective genes 1 mM FA and listed in Tab. S4 and S5 using the primer sets listed in Tab. S3. (http://microbio1.biologie.uni- greifswald.de/publications.html). Western blot analysis

Hierarchical clustering analysis Anti-FLAG polyclonal rabbit antiserums (SIGMA) were used at a dilution of 1:3000 to Clustering of gene expression profiles in analyze the cytoplasmic protein amounts of response to different electrophiles was FLAG-AdhR and FLAG-AdhRC52A proteins performed using Cluster 3.0 (de Hoon et al., produced in the AdhR and AdhRC52A 2004). The transcriptome datasets were complemented adhR mutant strains after derived either from previous publications or induction with 2% Xylose. Fifty micrograms this study and included log2-fold expression of each protein sample were loaded onto a changes 10 min after exposure of B. subtilis 12% SDS-PAGE gel and the Western blot to the electrophilic compounds diamide analysis was performed as described (1mM) (Leichert et al., 2003), MHQ (0.33 previously (Liebeke et al., 2008). mM) (Duy et al., 2007), catechol (2.4 mM) (Tam et al., 2006), FA (1 mM) and MG (2.8 5’ RACE PCR and 5.6 mM). The complete data set of all induced genes under these different The transcription start sites of adhR, yraC and conditions of electrophile treatment were adhA genes were mapped. Total RNA was listed and classified into regulons in Tab. S6 prepared from B. subtilis CU1065 grown (http://microbio1.biologie.uni- aerobically in LB medium with and without greifswald.de/publications.html). After the addition of 2 mM methylglyoxal. The cell hierarchical clustering, the output was cultures were grown to an OD600 of 0.4 and visualized using TreeView (Eisen et al., split into two flasks with equal volume, 1998). For the clustering as shown in Fig. 5, methylglyoxal was added to one flask to a we have used of 271 selected genes that are final concentration of 2 mM and cells were indicative for the thiol-specific stress harvested 20 min after treatment. The RNeasy response in B. subtilis and regulated by CtsR, mini kit (Qiagen) was used as a method to PerR, CymR, Spx, ArsR, CsoR, CzrA, YodB, extract total RNA from the cell lysates. The YvaP (CatR). In addition, the MhqR regulon RNA concentration was quantified by which responds specifically to quinones and NanoDrop spectrophotometer (Nanodrop the AdhR and HxlR regulons that are Tech. Inc., Welmington, DE). The 2 μg of specifically activated by aldehydes are total RNA was used as a template for reverse included in the clustering analysis. transcription using MultiscribeTM Reverse transcriptase (Taqman, Roche) and a gene- Northern blot experiment specific primer (GSP1). The resulting cDNA was purified by gel extraction purification Northern blot analyses were performed as column (Qiagen) and added poly(dC) tail at described (Wetzstein et al., 1992) using RNA their 3’-ends with terminal deoxynucleotidyl isolated from B. subtilis wild-type cells transferase (New England Biolabs). The before (control) and 10 min after the resulting cDNA was amplified by PCR using treatment with 1 mM FA, 2.8 mM and 5.6 the poly-dG primer to anneal at the poly(dC) mM MG, respectively. Hybridizations tail and a second gene-specific primer specific for adhA, yraA, adhR, clpE, nfrA, (GSP2), complementary to a region upstream cysK, katA, yqcK, hxlA, yrhE and yrhG were of the GSP1 primer. PCR products were performed with the digoxigenin-labeled RNA separated by gel electrophoresis and probes synthesized in vitro using T7 RNA sequenced. polymerase from T7 promoter containing

AdhR (YraB) 82 Huyen et al., 2008b

Expression and purification of the AdhR yoeB were generated by PCR. Approximately protein 50 ng of purified PCR products were end- labeled using T4 polynucleotide kinase (New AdhR protein was purified using PrepEaseTM England Biolabs) and 50 μCi of [γ-32P]-ATP. His-Tagged High Yield purification Resin The labeled DNA probes were purified by gel (USB). The adhR gene was PCR amplified extraction purification column (Qiagen) and from B. subtilis chromosomal DNA with 5000 cpm of each probe was incubated with oligonucleotides, designed to engineer an different amount of purified AdhR protein for NdeI site upstream and a HindIII site 5 min at room temperature in EMSA-binding downstream of the adhR gene. The PCR buffer (20 mM Tris-HCl pH 8.0, 50 mM product was cloned into pET16b (Novagen) NaCl, 5% glycerol, 5 μg ml-1 Salmon sperm via the NdeI and HindIII sites. The sequence DNA and 50 μg ml-1 BSA). Add MG or FA of adhR in pET16 was verified by DNA into reactions (Final concentration of these sequencing. E. coli BL21/DE3 containing this compounds was 10 mM) and further incubate plasmid was grown to mid-logarithmic phase for 20 min. Samples were separated by 6% at 37 °C in 1 liter of LB medium and 100 μg native polyacrylamide gel electrophoresis in of ampicillin per ml. Tris-Borate buffer pH 8.0 on ice and constant Isopropylthiogalactopyranoside (IPTG) was voltage (100 V) for 2 h. Gels were dried and added to 1 mM and cells were harvested after the radiolabeled bands were visualized using further incubation for 2 h. After phosphoimaging. centrifugation, pellet was resuspended in 5 ml LEW (Lysis-Equilibration-Wash) buffer (50 Determination of the cysteine content in mM NaH2PO4, 300 mM NaCl pH 8.0) and the metabolome of B. subtilis using GC/MS sonicated. After centrifugation, supernatant was transferred to clean tube and then B. subtilis cells were grown in Belitsky incubated for 15 min at room temperature minimal medium to an OD500 of 0.4 and 2+ with 1 g PrepEase Ni -IDA affinity resin treated with FA (0.5-1 mM) and MG (2.8 and (USB). The protein-bound resin was 5.5 mM) for 30 min. Cells were rapidly transferred to a column and washed collected by the fast filtration method and extensively with LEW buffer. AdhR was intracellular metabolites were analyzed using eluted by adding 10 ml of 50 mM NaH2PO4, gas chromatography/mass spectrometry 300 mM NaCl, and 250 mM imidazole pH (GC/MS) as described previously (Liebeke et 8.0 to a column. Fractions of 1 ml were al., 2008). collected and tested for protein content using Protein Assay Dye (Bio-Rad) before pooling 1H-Nuclear Magnetic Resonance (NMR)- for subsequent purification. The protein was spectroscopy concentrated by Ammonium sulfate precipitation and further purified by A solution of FA or MG was incubated with chromatography on an FPLC Superdex-200 equimolar cysteine for 10 min at 25°C in a column in 10 mM Tris-HCl pH 8.0, 50 mM NMR tube (Norell ST-500). NMR spectra NaCl, 1 mM DTT, 0.1 mM EDTA and 10% were recorded on a Bruker 600.27 MHz glycerol buffer followed by the dialysis into Avance-II spectrometer equipped with a 10 mM Tris-HCl pH 8.0, 50 mM NaCl, 1 mM BACS-60 sample changer and controlled with DTT, 0.1 mM EDTA and 50% glycerol TOPSPIN 2.0 (Bruker, Rheinstetten, buffer and stored at -80°C. Germany). Bruker Icon NMR software was used to automate the NMR data collection. DNA electrophoretic mobility shift assays 1H-NMR spectra were measured with a standard Bruker pulse sequence (noesygppr), DNA fragments containing the promoter solvent presaturation, a sweep width of regions of adhR, yraC, adhA and control 12365 Hz, 64K data points at 298.5K. A total

AdhR (YraB) 83 Huyen et al., 2008b of 2 dummy scans and 256 scans were used to adducts in the development of vascular complications obtain the NMR spectra. All peak positions in diabetes mellitus. Biochem Soc Trans 31: 1400- 1402. were measured relative to the TMSP (1mM Brown, N.L., Stoyanov, J.V., Kidd, S.P., and Hobman, sodium 3-trimethylsilyl-[2,2,3,3-D4]-1- J.L. (2003) The MerR family of transcriptional propionic acid) reference peak set to regulators. FEMS Microbiol Rev 27: 145–163. δ = 0.0 ppm. AMIX® Bruker Biospin was Bryk, R., Lima, C.D., Erdjument-Bromage, H., used for compound identification by Tempst, P., and Nathan, C. (2002) Metabolic enzymes 1 of mycobacteria linked to antioxidant defense by a matching the obtained spectra with a H- thioredoxin-like protein. Science 295: 1073-1077 NMR spectra databank. Chen, P.R., and He, C. (2008) Selective recognition of metal ions by metalloregulatory proteins. Curr Opin ACKNOWLEDGEMENTS Chem Biol 12: 214-221. Review. de Hoon, M.J., Imoto, S., Nolan, J., and Miyano, S. (2004) Open source clustering software. We thank the Decodon company for support Bioinformatics 20:1453-4. with the Decodon Delta 2D software, Dinkova-Kostova, A.T., Holtzclaw, W.D., and Sebastian Grund for the excellent technical Kensler, T.W. (2005) The role of Keap1 in cellular assistance and Dirk Albrecht for the mass protective responses. Chem Res Toxicol 18: 1779- spectrometry analysis. We are grateful to 1791. Review. Duy, N.V., Wolf, C., Mäder, U., Lalk, M., Langer, P., Kazuo Kobayashi for providing the hxlR Lindequist, U., Hecker, M., and Antelmann, H. (2007) mutant strain. This work was supported by Transcriptome and proteome analyses in response to 2- grants from the Deutsche methylhydroquinone and 6-brom-2-vinyl-chroman-4- Forschungsgemeinschaft, the on reveal different degradation systems involved in the Bundesministerium für Bildung und catabolism of aromatic compounds in Bacillus subtilis. Proteomics 7: 1391-1408. Forschung (BACELL-SysMo 031397A), the Eiamphungporn, W., and Helmann, J.D. (2008) The Fonds der Chemischen Industrie, the Bacillus subtilis sigmaM regulon and its contribution Bildungsministerium of the country to cell envelope stress responses. Mol Microbiol 67: Mecklenburg-Vorpommern and European 830-848. Union grants BACELL-Health (LSHG-CT- Eisen, M.B., Spellman, P.T., Brown, P.O., and Botstein, D. (1998) Cluster analysis and display of 2004-503468), BACELL-BaSysBio (LSHG- genome-wide expression patterns. Proc Natl Acad Sci CT-2006-037469) to M.H., and from the USA 95:14863-8. National Science Foundation (grant MCB- Eulberg, D., Lakner, S., Golovleva, L.A., and 0640616) to J.D.H. Schlomann, M. (1998) Characterization of a protocatechuate catabolic gene cluster from Rhodococcus opacus 1CP: evidence for a merged REFERENCES enzyme with 4-carboxymuconolactone- decarboxylating and 3-oxoadipate enol-lactone- Antelmann, H., Hecker, M., and Zuber, P. (2008) hydrolyzing activity. J Bacteriol 180: 1072-1081 Proteomic signatures uncover thiol-specific Even, S., Burguiere, P., Auger, S., Soutourina, O., electrophile resistance mechanisms in Bacillus subtilis. Danchin, A., and Martin-Verstraete, I. (2006) Global Expert Rev Proteomics 5: 77-90 control of cysteine metabolism by CymR in Bacillus Barford, D. (2004) The role of cysteine residues as subtilis. J Bacteriol 188, 2184-2197. redox-sensitive regulatory switches. Curr Opin Struct Eymann, C., Dreisbach, A., Albrecht, D., Bernhardt, J., Biol 14: 679-686. Becher, D., Gentner, S., Tam, L.T., Büttner, K., Bhavsar, A.P., Zhao, X., and Brown, E.D. (2001) Buurmann, G., Scharf, C., Venz, S., Völker, U., and Development and characterization of a xylose- Hecker, M. (2004) A comprehensive proteome map of dependent system for expression of cloned genes in growing Bacillus subtilis cells. Proteomics 4: 2849- Bacillus subtilis: conditional complementation of a 2876. teichoic acid mutant. Appl Environ Microbiol 67: 403- Farmer, E.E., and Davoine, C. (2007) Reactive 410. electrophile species. Curr Opin Plant Biol 10: 380- Booth, I.R., Ferguson, G.P., Miller, S., Li, C., 386. Review. Gunasekera, B., and Kinghorn, S. (2003) Bacterial Ferguson, G.P., Totemeyer, S., MacLean, M.J., and production of methylglyoxal: a survival strategy or Booth, I.R. (1998) Methylglyoxal production in death by misadventure? Biochem Soc Trans 31: 1406- bacteria: suicide or survival? Arch Microbiol 170: 209- 1408. Review. 218. Review Bourajjaj, M., Stehouwer, C.D., van Hinsbergh, V.W., Giles, N.M., Giles, G.I., and Jacob, C. (2003) Multiple and Schalkwijk, C.G. (2003) Role of methylglyoxal

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Moore, C.M., Nakano, M.M., Wang, T., Ye, R.W., and Stroeher, U.H., Kidd, S.P., Stafford, S.L., Jennings, Helmann, J.D. (2004) Response of Bacillus subtilis to M.P., Paton, J.C., and McEwan, A.G. (2007) A nitric oxide and the nitrosating agent sodium pneumococcal MerR-like regulator and S- nitroprusside. J Bacteriol 186(14): 4655-64. nitrosoglutathione reductase are required for systemic Morby, A.P., Turner, J.S., Huckle, J.W., and Robinson, virulence. J Infect Dis 196: 1820-1826. N.J. (1993) SmtB is a metal-dependent repressor of the Stülke, J., Hanschke, R., and Hecker, M. (1993) cyanobacterial metallothionein gene smtA: Temporal activation of β-glucanase synthesis in identification of a Zn inhibited DNA-protein complex. Bacillus subtilis is mediated by the GTP pool. J Gen Nucleic Acids Res 21, 921-925. Microbiol 139: 2041-2045. Nakano, S., Kuster-Schock, E., Grossman, A.D., and Tam, L.T., Eymann, C., Albrecht, D., Sietmann, R., Zuber, P. (2003) Spx-dependent global transcriptional Schauer, F., Hecker, M., and Antelmann H. (2006) control is induced by thiol-specific oxidative stress in Differential gene expression in response to phenol and Bacillus subtilis. Proc Natl Acad Sci USA 100: 13603- catechol reveals different metabolic activities for the 13608. degradation of aromatic compounds in Bacillus O`Brian, P.J. (1991) The molecular mechanisms of subtilis. Environ Microbiol 8: 1408-1427. quinone toxicity. Chem Biol Interact 80: 1-41. Thackray, P.D., and Moir, A. (2003) SigM, an Orlando, V., Strutt, H., and Paro, R.O. (1997) Analysis extracytoplasmic function sigma factor of Bacillus of chromatin structure by in vivo formaldehyde cross- subtilis, is activated in response to cell wall antibiotics, linking. Methods 11: 205-214. ethanol, heat, acid, and superoxide stress. J Bacteriol Oya, T., Hattori, N., Mizuno, Y., Miyata, S., Maeda, 185: 3491-3498. S., Osawa, T., and Uchida, K. (1999) Methylglyoxal Töwe, S., Leelakriangsak, M., Kobayashi, K., Van modification of protein. Chemical and Duy, N., Hecker, M., Zuber, P., and Antelmann, H. immunochemical characterization of methylglyoxal- (2007) The MarR-type repressor MhqR (YkvE) arginine adducts. J Biol Chem 274: 18492-18502. regulates multiple dioxygenases/glyoxalases and an Roca, A., Rodriguez-Herva, J.J., Duque, E., and azoreductase which confer resistance to 2- Ramos, J.L. (2007) Physiological responses of methylhydroquinone and catechol in Bacillus subtilis. Pseudomonas putida to formaldehyde during Mol Microbiol 66(1): 40-54. detoxification. Mol Biotechnol, in press. Wakabayashi, N., Dinkova-Kostova, A.T., Holtzclaw, Satoh, T., Okamoto, S.I., Cui, J., Watanabe, Y., Furuta, W.D., Kang, M.I., Kobayashi, A., Yamamoto, M., K., Suzuki, M., Tohyama, K., and Lipton, S.A. (2006) Kensler, T.W., and Talalay, P. (2004) Protection Activation of the Keap1/Nrf2 pathway for against electrophile and oxidant stress by induction of neuroprotection by electrophilic phase II inducers. the phase 2 response: fate of cysteines of the Keap1 Proc Nat Acad Sci USA. 103: 768-773. sensor modified by inducers. Pro Natl Acad Sci USA Satoh, T., and Lipton, S.A. (2007) Redox regulation of 101: 2040-2045. neuronal survival mediated by electrophilic Wei, Y., Ringe, D., Wilson, M.A., and Ondrechen, compounds. Trends Neurosci 30(1): 37-45. M.J. (2007) Identification of functional subclasses in Serizawa, M., Yamamoto, H., Yamaguchi, H., Fujita, the DJ-1 superfamily proteins. PLoS Comput Biol 3: Y., Kobayashi, K., Ogasawara, N., and Sekiguchi, J. e10. (2004) Systematic analysis of SigD-regulated genes in Wetzstein, M., Völker, U., Dedio, J., Löbau, S., Zuber, Bacillus subtilis by DNA microarray and Northern U., Schiesswohl, M., Herget, C., Hecker, M., and blotting analyses. Gene 329: 125-36. Schumann, W. (1992) Cloning, sequencing, and Skoneczna, A., Miciałkiewicz, A., and Skoneczny, M. molecular analysis of the dnaK locus from Bacillus (2007) Saccharomyces cerevisiae Hsp31p, a stress subtilis. J Bacteriol 174: 3300-3310. response protein conferring protection against reactive Yurimoto, H., Hirai, R., Matsuno, N., Yasueda, H., oxygen species. Free Radic Biol Med 42: 1409-1420 Kato, N., and Sakai, Y. (2005) HxlR, a member of the Smaldone, G.T., and Helmann, J.D. (2007) CsoR DUF24 protein family, is a DNA-binding protein that regulates the copper efflux operon copZA in Bacillus acts as a positive regulator of the formaldehyde- subtilis. Microbiology 153(Pt 12): 4123-8. inducible hxlAB operon in Bacillus subtilis. Mol Steil, L., Serrano, M., Henriques, A.O., and Völker, U. Microbiol 57: 511-519. (2005) Genome-wide analysis of temporally regulated Zuber, P. (2004) Spx-RNA polymerase interaction and and compartment-specific gene expression in global transcriptional control during oxidative stress. J. sporulating cells of Bacillus subtilis. Microbiology Bacteriol 186: 1911-1918. 151: 399-420.

AdhR (YraB) 86 Huyen et al., 2008b

Tab. S1. Induction of genes and proteins in response to MG and FA stresses as revealed by transcriptome and proteome analyses

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM Heat shock regulons HrcA groELS groEL 0.34 1.07 5.00 2.22 4.50 class I heat-shock protein (chaperonin) groES 0.13 0.58 5.98 class I heat-shock protein (chaperonin) (hrcA-grpE- hrcA 5.50 repressor of class I heat-shock genes dnaK- dnaJ) grpE 5.26 heat-shock protein (activation of dnaK) class I heat-shock protein (molecular dnaK 3.52 chaperone) CtsR clpE clpE 2.27 21.86 246.24 12.65 7.84 ATP-dependent Clp protease-like B ATP-dependent Clp protease σ clpP clpP 10.43 proteolytic subunit ATP-dependent Clp protease ATP- clpX clpX 3.29 binding subunit (class III heat-shock protein) B transcriptional repressor of class III σ (ctsR- ctsR 1.28 2.76 9.93 stress genes mcsAB-clpC) mcsA (yacH) 1.26 3.14 11.99 modulator of CtsR repression mcsB (yacI) 1.56 3.47 16.61 modulator of CtsR repression class III stress response-related clpC missing missing missing 5.96 7.16 ATPase class III heat-shock ATP-dependent lonA-ysxC lonA 5.06 protease ysxC 2.98 similar to GTP-binding protein Oxidative stress regulon PerR alkyl hydroperoxide reductase (small ahpCF ahpC 0.56 1.61 1.42 1.02 0.90 subunit) alkyl hydroperoxide reductase (large ahpF 0.51 1.88 1.55 1.66 1.00 subunit) hemAXCDBL hemA 1.65 2.78 1.32 glutamyl-tRNA reductase negative effector of the concentration hemX 1.61 3.54 1.25 of HemA hemC 2.01 3.49 1.30 porphobilinogen deaminase hemD 1.74 3.37 1.39 uroporphyrinogen III cosynthase delta-aminolevulinic acid dehydratase hemB 1.74 2.31 1.26 (porphobilinogen synthase) glutamate-1-semialdehyde 2,1- hemL 1.44 2.31 1.18 aminotransferase katA katA 0.75 10.78 5.93 1.28 1.17 vegetative catalase 1 fur fur (yqkL) 1.42 2.64 2.05 transcriptional repressor of iron uptake metalloregulation DNA-binding stress mrgA mrgA 1.98 18.51 2.22 1.79 1.58 protein metal-dependent repressor of oxidative perR perR (ygaG) 1.90 4.77 2.13 stress P-type metal-transporting ATPase; a zosA zosA (ykvW) 3.37 9.56 3.85 peroxide-induced zinc uptake system SOS regulon LexA dinB dinB 1.77 4.88 3.44 nuclease inhibitor transcriptional repressor of the SOS lexA lexA (dinR) 1.49 5.79 3.37 regulon multifunctional protein involved in ComK recA recA (recE) 2.19 4.41 3.18 homologous recombination and DNA repair (LexA-autocleavage) uvrBA uvrB (uvrA) 1.27 4.37 3.77 excinuclease ABC (subunit B) uvrC uvrC (uvrB) 2.60 4.49 4.49 excinuclease ABC (subunit C) PhoPR vpr vpr 3.59 minor extracellular serine protease xkdA xkdA 1.80 3.54 1.35 PBSX prophage similar to DNA-methyltransferase ydiOP ydiO 5.73 (cytosine-specific) similar to DNA-methyltransferase ydiP 5.40 (cytosine-specific) ComK yneAB-ynzC yneA 2.46 11.13 6.54 unknown yneB 2.08 8.30 5.36 unknown; similar to resolvase ynzC 1.22 1.96 3.05 unknown

AdhR (YraB) 87 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM LexA (continued) yolD-uvrX yolD 1.31 4.08 2.48 unknown uvrX 1.66 3.06 2.88 UV-damage repair protein yorBC yorB 1.37 6.11 3.33 unknown similar to DNA alkylation repair yhaZ 2.66 7.81 4.16 enzyme Thiol stress regulons Spx hemEHY hemE 2.81 2.36 2.77 uroporphyrinogen III decarboxylase hemH 2.33 3.64 7.76 5.74 2.15 ferrochelatase hemY 1.78 2.38 4.19 protoporphyrinogen IX oxidase mecA mecA 2.89 5.84 5.14 negative regulator of competence peptidyl methionine sulfoxide msrA-yppQ msrA (yppP) 1.31 3.17 9.02 reductase similar to peptide methionine sulfoxide yppQ 1.45 2.80 5.99 reductase D FMN-containing NADPH-linked σ nfrA-ywcH nfrA (ywcG) 3.06 7.04 19.19 5.44 3.61 nitro/flavin reductase ywcH 3.52 7.64 10.88 similar to monooxygenase pepF (yjbG) 4.44 4.42 3.29 oligoendopeptidase F homolog tpx tpx (ytgI) 4.88 2.47 thiol peroxidase CtsR, σB trxA trxA 1.72 3.19 8.09 thioredoxin trxB 2.41 3.64 6.35 2.20 2.66 thioredoxin reductase ycnED ycnE 2.78 unknown similar to NADPH-flavin ycnD 3.82 oxidoreductase ydbP ydbP 4.19 similar to thioredoxin yfiR yfjR 1.49 2.41 13.90 1.88 2.65 dehydrogenase precursor nucleoside-diphosphate-sugar yhfK 5.65 epimerase similar to adenine-specific DNA yitW 2.98 methylase yjbH 5.88 dithiol-disulfide isomerase yjbI 3.75 hemoglobin-like protein ykhA 3.98 similar to acyl-CoA hydrolase ymfG 3.33 similar to processing protease ymfH 3.03 similar to processing protease ymzA 2.14 3.11 6.78 unknown yojF 4.48 unknown yojG 1.92 3.56 4.87 similar to LMBE-related protein similar to NADH-dependent flavin yquiG yqiG 4.09 5.63 9.29 5.11 2.03 oxidoreductase similar to NADH-dependent flavin yqjM yqjM 1.67 3.67 8.25 oxidoreductase probable serine/threonine-protein yrzF 2.21 3.58 6.50 kinase probable serine/threonine-protein yrzG 1.81 3.59 5.49 kinase ytdI 3.34 similar to ATP-NAD kinase yuaE 6.09 unknown probable NADH-dependent butanol yugJ yugJ 1.80 3.27 12.54 1.68 2.28 dehydrogenase 1 similar to NADH-dependent butanol yugK yugK 4.95 dehydrogenase yugP 3.12 5.60 9.50 similar to Zn-dependent protease yusX 3.52 similar to oligoendopeptidase yusY 3.42 similar to oligoendopeptidase F similar to ketoacyl-carrier protein yvrD 2.14 3.30 7.95 reductase ywnF ywnF 2.66 7.67 13.46 unknown ywrO ywrO 4.53 similar to NAD(P)H oxidoreductase yxcED yxcE 3.10 6.13 3.69 unknown yxcD 2.63 4.76 3.11 unknown YodB FMN-dependent NADH-azoreductase azoR1(yocJ) 2.42 11.20 2.12 1.19 1.09 1 Spx, global regulator of the oxidative stress yjbCD spx (yjbD) 0.86 5.86 2.75 PerR response MarR-type repressor of yocJ, yodC yodB yodB 1.48 3.34 1.63 and spx yodC yodC 1.31 2.78 2.94 similar to NAD(P)H nitroreductase

AdhR (YraB) 88 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM Others similar to DoxX/ DoxD-like CatR catDE catD (yfiD) 4.77 8.92 oxidoreductases (yfiDE) catE (yfiE) 2.46 4.76 catechol-2,3-dioxygenase glyoxalase/ bleomycin resistance MhqR mhqA mhqA (ykcA) 1.53 2.53 5.53 protein/ dioxygenase 2-Methylhydroquinone (thiol stress) mhqR (ykvE) 6.47 2.07 resistance repressor MhqR ydfQ 5.43 similar to thioredoxin ydgI 3.98 8.79 5.25 8.18 3.22 similar to NADH dehydrogenase similar to transcriptional regulator ydgJ 3.02 6.74 4.37 (MarR family) similar to NAD(P)H-flavin YybR yfkO yfkO 3.04 5.65 4.40 2.88 3.77 oxidoreductase YybR yybR yybR 3.57 7.88 10.03 MarR/DUF24-family regulator YkvN ykvN 4.44 MarR/DUF24-family regulator Cysteine metabolism regulon CymR cysK cysK 1.11 5.93 12.14 2.31 5.48 cysteine synthetase A Spx, cysP operon cysH 1.04 4.02 4.67 phosphoadenosine phosphosulfate S-box cysP (ylnA) 1.04 3.90 5.75 sulfate permease sat (ylnB) 0.73 3.49 7.68 sulfate adenylyltransferase cysC (ylnC) 1.02 3.67 6.46 adenylylsulfate kinase similar to uroporphyrin-III C- ylnD 1.08 3.62 5.76 methyltransferase ylnE 1.25 3.86 7.59 unknown similar to uroporphyrin-III C- ylnF 1.11 3.84 6.20 methyltransferase aliphatic sulfonate ABC transporter Spx ssu operon ssuB (ygaL) 37.57 (binding protein) aliphatic sulfonate ABC transporter ssuA (ygbA) 68.39 (binding lipoprotein) aliphatic sulfonate ABC transporter ssuC (ygaM) 32.41 (permease) ssuD (ygcA) 25.91 aliphatic sulfonate monooxygenase yhcL tcyP (yhcL) 1.30 5.18 3.44 similar to sodium-glutamate symporter similar to transcriptional regulator Spx ytmI operon ytmI 1.27 7.04 24.86 (LysR family) tcyJ (ytmJ) 1.23 10.38 29.30 unknown L-Cystine ABC transporter (binding tcyK (ytmK) 1.79 17.24 38.36 protein) tcyL (ytmL) 1.70 8.72 25.82 L-Cystine ABC transporter (permease) tcyM (ytmM) 2.07 9.52 39.08 L-Cystine ABC transporter (permease) tcyN (ytmN, L-Cystine ABC transporter (ATP- 1.57 5.20 22.46 hisP) binding protein) ytmO 3.88 similar to monooxygenase ytnI 5.58 unknown similar to nitrilotriacetate ytnJ 1.27 3.53 19.65 monooxygenase ribR 1.56 4.37 26.44 riboflavin kinase ytnL (hipO) 1.41 3.47 12.38 similar to aminohydrolase ydbM ydbM 0.88 3.08 8.78 similar to butyryl-CoA dehydrogenase Spx ytlI ytlI 0.85 5.05 17.07 transcriptional regulator, LysR family Spx yxeI operon yxeI 1.14 2.61 7.85 similar to penicillin amidase yxeJ 1.35 1.57 1.55 unknown xenobiotic compound monooxygenase, yxeK 1.63 11.40 36.63 2.18 3.25 DszA family putative acetyltransferase, GNAT yxeL 2.39 14.46 26.76 family amino-acid ABC transporter (binding yxeM 3.37 19.68 26.69 protein) amino-acid ABC transporter yxeN 2.91 18.98 31.87 (permease) amino-acid ABC transporter (ATP- yxeO 3.23 15.87 37.43 binding protein) yxeP 1.24 4.81 7.74 1.57 8.61 peptidase, M20/M25/M40 family yxeQ 2.50 7.70 13.71 unknown Spx yxeR-yxxB yxeR 2.00 4.29 6.20 similar to ethanolamine transporter yxxB 5.23 unknown (yrrT-mtn- yrrT 2.08 12.85 23.78 probable S-adenosylmethionine yrhABC) mtn (yrrU) 1.35 4.91 11.32 methylthioadenosine nucleosidase

AdhR (YraB) 89 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM CymR (continued) yrhA 2.64 28.94 44.64 putative cystathionine ß-synthase yrhB 2.85 29.48 43.19 2.12 5.06 putative cystathionine g-lyase yrhC 1.88 8.43 11.63 unknown Formaldehyde detoxification HxlR hxlBA hxlA (yckG) 25.30 49.26 86.66 6.06 4.85 3-hexulose-6-phosphate synthase hxlB (yckF) 32.28 40.19 79.15 6-phospho-3-hexuloisomerase hxlR (yckH) 2.62 3.46 8.76 positive regulator of hxlAB expression YraB (AdhR) yra operon yraA 8.38 9.61 13.45 similar to general stress protein NADP-dependent alcohol adhA 42.05 52.70 45.04 dehydrogenase similar to transcriptional regulator yraB 9.80 16.76 16.11 (MerR family) yraC 11.88 17.02 16.29 unknown Formate detoxification yrhED yrhD 9.76 21.05 unknown yrhE 13.50 29.10 similar to formate dehydrogenase yrhFG-yrzI yrhG 17.24 32.12 1.38 similar to formate transporter yrzI 18.50 26.64 3.92 unknown Metal ion stress responsive regulons ArsR (arsR-yqcK- arsR (yqcJ) 5.09 32.07 15.45 arsenic resistance operon repressor glyoxalase/ bleomycin resistance arsBC) yqcK 9.77 55.45 39.84 protein/ dioxygenase arsB (yqcL) 11.89 61.89 47.59 extrusion of arsenite arsC (yqcM) 11.37 31.04 29.78 arsenate reductase CsoR heavy metal-transporting CPx-type copZA copA (yvgX) 2.12 8.82 3.33 ATPase; confers resistance to Cu(II) copZ (yvgY) 1.99 14.22 3.81 copper/ cadmium resistance protein B regulator of copper resistance copZA σ csoR (yvgZ) 1.38 3.66 2.38 operon CzrA (YozA) czcD-trkA czcD 13.79 37.26 25.84 cation-efflux system membrane protein trkA 13.71 32.27 28.44 potassium uptake Cell wall responsive regulon YycFG antibiotic-inducible cell wall-associated yoeB 0.62 5.98 6.53 protein that protects cells from autolysis σM yacK 5.58 unknown σW X M putative hydrolase; confers resistance σ , σ yqjL yqjL 1.00 3.21 to paraquat σX yrhH yrhH 3.68 5.72 similar to methyltransferase DegSU yxjJI yxjJ 1.66 4.58 unknown ywaC ywaC 0.99 4.77 similar to GTP-pyrophosphokinase Flagellin/Motility and Chemotaxis control (σD) modulator of CheA activity in response cheV cheV 2.00 6.45 to attractants ComK degR degR 4.06 10.30 degradative enzyme production epr epr 3.42 6.28 3.87 extracellular serine protease hemAT hemAT 1.66 3.37 haem-based aerotactic transducer (yhfV) motility protein A (flagellar motor motAB motA 3.62 9.17 rotation) motility protein B (flagellar motor motB 4.46 11.85 rotation) tlpA-mcpA tlpA 3.64 8.54 methyl-accepting chemotaxis protein mcpA 3.50 9.28 methyl-accepting chemotaxis protein mcpB mcpB 3.13 9.60 methyl-accepting chemotaxis protein tlpB tlpB 5.36 11.04 3.44 methyl-accepting chemotaxis protein tlpC tlpC 2.44 3.03 methyl-accepting chemotaxis protein ybdO ybdO 2.07 4.51 unknown similar to benzaldehyde yfmTS yfmT missing missing missing dehydrogenase

AdhR (YraB) 90 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM D σ (continued) similar to methyl-accepting chemotaxis yfmS 2.70 6.45 protein yjcPQ yjcP 2.81 4.65 unknown yjcQ 2.75 6.58 unknown yjfB yjfB 2.22 4.02 unknown ylqB ylqB 1.83 3.58 unknown similar to methyl-accepting chemotaxis yoaH yoaH 3.17 3.34 protein yscB yscB 3.28 6.25 unknown similar to methyl-accepting chemotaxis yvaQ 3.24 5.63 protein Fur yvyC-fliDST yvyC 1.06 2.10 similar to flagellar protein flagellar hook-associated protein 2 fliD 1.65 4.03 (HAP2) fliS 2.57 5.33 flagellar protein fliT 3.33 4.33 flagellar protein ComK flgKLM yvyF 2.17 3.86 similar to flagellar protein flagellin synthesis regulatory protein operon flgM 1.88 4.38 (anti-sigmaD) yvyG 2.28 7.59 similar to flagellar protein flagellar hook-associated protein 1 flgK 2.43 7.15 (HAP1) flagellar hook-associated protein 3 flgL 3.38 12.13 (HAP3) PyrR regulon pyr operon pyrP 3.68 0.56 0.09 uracil permease pyrB 5.12 0.44 0.07 aspartate carbamoyltransferase pyrC 4.42 0.53 0.29 dihydroorotase carbamoyl-phosphate synthetase pyrAA 5.87 0.57 0.28 (glutaminase subunit) carbamoyl-phosphate synthetase pyrAB 3.67 0.87 1.64 (catalytic subunit) dihydroorotate dehydrogenase pyrK (pyrDII) 3.50 0.74 2.15 (electron transfer subunit) dihydroorotate dehydrogenase pyrD 3.67 0.93 1.53 (catalytic subunit) pyrF 2.61 0.77 3.43 orotidine 5'-phosphate decarboxylase pyrE 3.31 1.05 2.40 orotate phosphoribosyltransferase TnrA regulon ComA pel pel 1.95 3.49 3.00 pectate lyase similar to ribose 5-phosphate ywlFG ywlF 1.73 2.79 5.79 epimerase (pentose phosphate) ywlG 1.67 3.72 5.41 unknown Sporulation control Spo0A yaaDE yaaD 0.63 1.25 0.63 1.92 2.17 similar to superoxide-inducible protein ycgMN ycgM 12.40 similar to proline oxidase similar to 1-pyrroline-5-carboxylate ycgN 7.85 dehydrogenase σH cah cah 18.43 10.15 cephalosporin C deacetylase regulator of the activity of phosphatase CodY rapC-phrC phrC 1.15 2.96 3.06 RapC and competence and sporulation stimulating factor (CSF) response regulator aspartate rapC 1.58 3.70 3.62 phosphatase ypgR ypgR 4.71 unknown RelA, similar to sigma-54 modulating factor B yvyD yvyD 0.69 1.33 3.41 1.52 3.49 σ of Gram-negatives σE D-alanyl-D-alanine carboxypeptidase dacB-spmAB dacB 4.85 2.17 (penicillin-binding protein 5*) spmA 2.46 1.36 spore maturation protein spmB 4.03 2.14 spore maturation protein flhP 2.10 3.63 flagellar hook-basal body protein σG (yraGF- adhB 2.29 3.46 2.80 alcohol dehydrogenase adhB-yraED) yraE 1.99 2.91 2.81 similar to spore coat protein yraD 3.61 5.94 4.12 similar to spore coat protein

AdhR (YraB) 91 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM σK K E spoIVCB- sporulation mother cell-specific σ GerE, σ spoIIIC 6.05 spoIIIC factor similar to phospho-adenylylsulfate yitBA-yisZ yitB 1.08 1.87 2.18 sulfotransferase yitA 0.99 3.18 4.09 similar to sulfate adenylyltransferase yisZ 1.18 3.13 3.99 similar to adenylylsulfate kinase GerE maturation of the outermost layer of cgeCDE cgeD 3.37 4.76 the spore maturation of the outermost layer of cgeE 3.15 4.30 the spore σB regulon (bmrU-bmr- bmr 2.23 4.26 multidrug-efflux transporter transcriptional activator of the bmrUR bmrR) bmrR 2.06 2.56 operon succinate-semialdehyde GabR gabTD gabD (ycnH) 2.82 5.29 14.06 dehydrogenase (rsbRSTUVW rsbR 3.11 positive regulator of sigma-B activity -sigB-rsbX) rsbS 2.86 antagonist of RsbT positive regulator of sigma-B activity rsbT 2.98 (switch protein/serine-threonine kinase) indirect positive regulator of sigma-B rsbU 2.24 activity (serine phosphatase) yacLMN yacL 4.36 unknown yacM 3.23 unknown yvgN yvgN 7.25 similar to dehydrogenase yacN 2.31 unknown Other functions alsSD alsS 1.50 9.36 alpha-acetolactate synthase (pH6) alsD 2.01 18.93 alpha-acetolactate decarboxylase ampS 0.21 0.35 3.02 aminopeptidase AnsR ansAB ansA 1.44 6.20 L-asparaginase ansB missing missing missing L-aspartase similar to transcriptional regulator cueR (yhdQ) 3.84 (MerR family) (gltX-cysES- cysE 3.98 serine acetyltransferase yazC-yacOP) cysS 5.82 cysteinyl-tRNA synthetase yazC 5.62 unknown similar to tRNA/rRNA yacO 5.57 methyltransferase yacP 4.89 unknown GlpP glpD glpD 1.70 4.60 glycerol-3-phosphate dehydrogenase PurR guaC guaC (yumD) 9.40 19.94 GMP reductase expZ 3.70 7.43 3.66 ATP-binding transport protein ResDE hmp hmp 4.46 flavohemoglobin DegSU ispA ispA 2.85 intracellular serine protease luxS luxS (ytjB) 1.07 4.32 5.35 autoinducer-2 production protein aspartokinase II alpha subunit (aa 1- lysC 4.26 >408) and beta subunit (aa 246->408) ndhF 2.43 3.26 1.04 NADH dehydrogenase (subunit 5) ResDE (nrdI - nrdI (ymaA) 1.98 2.22 2.68 similar to NrdI protein ribonucleoside-diphosphate reductase nrdEF-ymaB) nrdE 2.06 2.51 3.50 (major subunit) ribonucleoside-diphosphate reductase nrdF 2.22 3.26 3.24 (minor subunit) ymaB 2.50 2.95 2.90 unknown para-aminobenzoate synthase (subunit TRAP pabB 1.33 3.71 A) phosphoribosylaminoimidazole PurR pur operon purE 2.31 3.88 0.07 carboxylase I phosphoribosylaminoimidazole purK 2.51 4.35 0.13 carboxylase II purB 1.78 3.71 0.09 adenylosuccinate lyase phosphoribosylaminoimidazole purC 1.56 3.29 0.16 succinocarboxamide synthetase radA (sms) 16.43 DNA repair protein homolog AbrB, senS senS 3.33 transcriptional regulator SenS sodA 0.30 0.94 3.53 superoxide dismutase

AdhR (YraB) 92 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM Other functions (continued) CcpN gapB-speD speD (ytcF) 3.32 2.59 S-adenosylmethionine decarboxylase tdh 2.51 4.65 threonine 3-dehydrogenase truA 3.72 3.19 pseudouridylate synthase I udk 3.37 1.18 uridine kinase yacA 4.69 similar to cell-cycle protein ybaF 4.09 1.43 unknown ybbU 1.44 3.11 3.46 unknown similar to chloramphenicol resistance ybcL ybcL 1.50 6.50 4.18 protein similar to glucosamine-fructose-6- ybcM 2.82 4.71 phosphate aminotransferase ybfQ 2.59 3.75 unknown yceB 4.63 unknown similar to transcriptional regulator yceK yceK 1.72 3.31 (ArsR family) ycgA ycgA 9.65 unknown ycgB ycgB 3.27 unknown ycgL 0.79 3.35 8.68 unknown similar to glutamine ABC transporter yckKJI yckK 1.81 2.65 3.08 (glutamine-binding protein) similar to glutamine ABC transporter yckJ (tcyB) 1.80 3.08 3.59 (permease) similar to glutamine ABC transporter yckI 1.77 2.36 3.87 (ATP-binding protein) similar to transcriptional regulator ycnCB ycnC 3.66 2.68 (TetR/AcrR family) ycnB 2.42 1.02 similar to multidrug resistance protein ydaF 3.01 similar to acetyltransferase ydaQ 3.39 unknown yddJ 3.44 3.77 unknown similar to transcriptional regulator ydeC 3.10 (AraC/XylS family) ydeQ 1.71 3.14 similar to NAD(P)H oxidoreductase similar to transcriptional regulator ydeT 3.15 1.21 (ArsR family) ydhK ydhK 2.56 3.54 unknown ydiM 8.40 unknown ydiN 4.37 unknown similar to mannose-6-phosphate ydhS 3.20 isomerase similar to mannan endo-1,4-beta- ydhT 3.29 mannosidase yebC 3.21 3.20 unknown yebD 3.96 unknown yebE 3.53 2.69 unknown yezD 0.79 3.27 24.21 unknown yfhJ 3.46 6.59 10.30 unknown yflGH yflG 4.04 similar to methionine aminopeptidase yflH 2.73 unknown yfmB 3.41 unknown ygaC 3.70 unknown yhaA 0.31 0.42 4.85 similar to aminoacylase yhbB 3.10 2.95 unknown yhdG yhdG 0.25 0.57 4.32 similar to amino acid transporter yhdP 1.60 3.02 4.18 similar to hemolysin similar to ABC transporter (ATP- yheH 6.69 11.03 3.29 binding protein) similar to ABC transporter (ATP- yheI 6.61 12.31 3.99 binding protein) yhfH 7.20 unknown yhzA 3.22 similar to ribosomal protein S14 yhzB 4.12 unknown yhzC yhzC 5.68 unknown yjbQ 3.25 similar to Na+/H+ antiporter similar to cystathionine gamma- yjcIJ yjcI 0.20 0.52 1.48 synthase yjcJ 0.30 0.80 3.17 similar to cystathionine beta-lyase yjcL 3.95 unknown yjcS 3.90 unknown similar to 3-oxoacyl-acyl-carrier protein yjdA 3.04 reductase

AdhR (YraB) 93 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM Other functions (continued) yjiB 3.03 similar to monooxygenase yjnA 9.10 unknown yjoA 5.00 unknown ykbA 7.79 9.71 similar to amino acid permease similar to Na+-transporting ATP ykrM 3.73 1.94 synthase yktB 3.41 unknown yktC 3.10 similar to inositol monophosphatase ykvS 3.23 unknown ykvY 3.98 similar to Xaa-Pro dipeptidase ykwB 4.52 unknown similar to 2-dehydropantoate 2- ylbQ 3.60 reductase yllA yllA 3.48 unknown ylqG 3.49 3.21 unknown ymaD 4.11 unknown similar to general secretion pathway yobE 3.14 protein similar to sodium-dependent yocS 3.96 transporter yomK 3.91 5.73 1.70 unknown yopJ 1.74 4.52 1.11 unknown yopX 1.21 3.05 1.31 unknown yoqY 1.71 3.04 1.23 unknown similar to 3-oxoacyl- acyl-carrier yoxD 3.48 protein reductase yozE 5.59 unknown a MecA paralog; binds ClpC, affecting ypbH 3.37 both competence and sporulation ypiA 2.20 3.36 5.61 unknown ypiB 4.86 unknown ypiF 3.44 unknown yqaB 5.21 similar to phage-related protein yqbF 3.41 3.88 unknown yqbG 6.41 6.54 unknown yqbH 4.16 5.60 similar to phage-related protein yqbI 2.54 3.77 similar to phage-related protein similar to glycerophosphodiester yqiK 5.52 phosphodiesterase yqiW yqiW 3.06 unknown yqjYZ- yqkB 3.23 unknown yqkABC yqkC 2.24 unknown yqkE yqkE 3.62 unknown yrbC 4.24 similar to spore coat protein yrdR 2.79 3.27 unknown yrhP 3.10 similar to efflux protein yrpC 1.99 3.00 similar to glutamate racemase ysbA 1.33 4.79 unknown yscA 5.36 1.15 0.18 unknown ysfC 2.45 5.22 similar to glycolate oxidase subunit yslB yslB 5.67 unknown ytbD 4.31 4.38 similar to antibiotic resistance protein similar to plant metabolite ytbE 2.89 3.21 4.61 dehydrogenase ytiP ytiP 3.61 2.31 0.34 unknown ytoP 3.05 similar to endo-1,4-beta-glucanase RelA ytzB 4.27 unknown similar to transcriptional regulator RelA ytzE 4.41 (DeoR family) similar to metal dependent yueE 6.34 phosphohydrolase yugS yugS 3.33 6.55 4.25 similar to hemolysin similar to two-component sensor yufM yufM 2.47 3.32 histidine kinase YufM yuiA 0.26 0.67 9.95 unknown yuiB 0.28 0.44 4.87 probable membrane protein yusF 3.52 unknown yuzF 4.06 unknown similar to 3-oxoacyl- acyl-carrier yvaG 2.07 3.43 protein reductase

AdhR (YraB) 94 Huyen et al., 2008b

Transcriptome Proteome Gene Regulon Operon MG MG FM MG FM Function or similarity (Synonym) 2,8 mM 5,6 mM 1 mM 5,6 mM 1 mM Other functions (continued) yvbT 3.13 similar to alkanal monooxygenas yvbV 3.45 6.96 3.27 unknown yvdR 1.41 4.02 similar to molecular chaperone yvdS 2.48 5.49 similar to molecular chaperone similar to TetR/AcrR-type yvdT 4.83 13.54 5.54 transcriptional regulator yviF 1.75 3.33 unknown yvjA-cccB yvjA 2.90 3.43 unknown yvkN 3.62 unknown similar to branched-chain amino acid ywaA 3.52 aminotransferase ywbC 3.00 unknown ComK ywfI ywfI 0.80 1.82 4.29 1.77 3.76 chlorite dismutase ywrF 5.62 unknown ywtD ywtD 4.33 4.63 similar to murein hydrolase similar to universal stress family CcpA bglPH-yxiE yxiE 1.37 3.81 5.02 protein; induced by phosphate starvation yxkD yxkD 2.15 3.38 unknown yxkH yxkH 5.12 5.24 unknown yyaM 4.09 4.84 11.69 unknown similar to transcriptional regulator yyaN 3.77 (MerR family) yyaO yyaO 3.44 unknown

SUPPLEMENTAL TABLE LEGENDS

Tab. S1. Induction of genes and proteins in response to methylglyoxal (MG) and formaldehyde (FA) stress as revealed by transcriptome and proteome analyses. The averages ratios for all induced genes and proteins are calculated from three independent transcriptome experiments and two proteome experiments after 10 min of exposure to 2.8 and 5.6 mM MG or 1 mM FA, respectively. All genes with induction ratios of at least three-fold and proteins with protein synthesis ratios of at least two-fold were listed and classified according to previously described regulons (HrcA, CtsR, Spx, PerR, SOS, CymR, SigmaD, SigmaW, SigmaM etc.). The classification of genes into regulons was performed according to http://dbtbs.hgc.jp. The main regulator of the induced genes is shown in bold-face. The gene function or similarity is derived from the SubtiList database (http://genolist.pasteur.fr/SubtiList/index.html). The complete dataset including ratios for all induced genes in each transcriptome experiment and the calculated average values and standard deviations are online available as Tab. S4 as web-supplement at http://microbio1.biologie.uni-greifswald.de/supplements/Antelmann_2008b/.

Tab. S2. Repression of genes in response to MG and FA stress as revealed by transcriptome analysis. The averages ratios for all induced genes are calculated from three independent transcriptome experiments after 10 min of exposure to 2.8 and 5.6 mM MG or 1 mM FA, respectively. All genes with repression ratios of at least three-fold were listed and classified according to previously described regulons (Fur, CcpA, PurR, Rok etc.). The classification of genes into regulons was performed according to http://dbtbs.hgc.jp. The main regulator of the repressed genes is shown in bold-face. The gene function or similarity is derived from the SubtiList database (http://genolist.pasteur.fr/SubtiList/index.html). The complete dataset including ratios for all repressed genes in each transcriptome experiment and the calculated average values and standard deviations are online available as Tab. S5 as web-supplement at http://microbio1.biologie.uni- greifswald.de/supplements/Antelmann_2008b/.

AdhR (YraB) 95 Huyen et al., 2008b

Tab. S3. Oligonucleotide primers used in this study. Tab. S3 is only online available as web- supplement at http://microbio1.biologie.uni-greifswald.de/supplements/Antelmann_2008b/.

Tab. S4. Induction of genes and proteins in response to MG and FA stress as revealed by transcriptome and proteome analyses. Tab. S4 is only online available as web-supplement at http://microbio1.biologie.uni-greifswald.de/supplements/Antelmann_2008b/. The averages ratios and standard deviations for all induced genes and proteins are calculated from three independent transcriptome experiments and two proteome experiments after 10 min of exposure to 2.8 and 5.6 mM MG or 1 mM FA, respectively. All genes with induction ratios of at least three-fold and proteins with protein synthesis ratios of at least two-fold were listed and classified according to previously described regulons (HrcA, CtsR, Spx, PerR, SOS, CymR, SigmaD, SigmaW, SigmaM etc.). The classification of genes into regulons was performed according to http://dbtbs.hgc.jp. The main regulator of the induced genes is shown in bold-face. The gene function or similarity is derived from the SubtiList database (http://genolist.pasteur.fr/SubtiList/index.html).

Tab. S5. Repression of genes in response to MG and FA stress as revealed by transcriptome analysis. Tab. S5 is only online available as http://microbio1.biologie.uni- greifswald.de/supplements/Antelmann_2008b/. The averages ratios and standard deviations for all repressed genes are calculated from three independent transcriptome experiments after 10 min of exposure to 2.8 and 5.6 mM MG or 1 mM FA, respectively. All genes with repression ratios of at least three-fold were listed and classified according to previously described regulons (Fur, CcpA, PurR, Rok etc.). The classification of genes into regulons was performed according to http://dbtbs.hgc.jp. The main regulator of the repressed genes is shown in bold-face. The gene function or similarity is derived from the SubtiList database (http://genolist.pasteur.fr/SubtiList/index.html).

Tab. S6. Induction of genes in response to the electrophiles diamide (Dia), catechol (Cat), methylhydroquinone (MHQ), methylglyoxal (MG) and formaldehyde (FA) as revealed by transcriptome analyses. Tab. S6 is only online available as web-supplement at http://microbio1.biologie.uni-greifswald.de/supplements/Antelmann_2008b/. The transcriptome datasets were derived either from previous publications or this study and included log2-fold expression changes of average values of two transcriptome experiments 10 min after exposure of B. subtilis to the electrophilic compounds diamide (1mM) (Leichert et al., 2003), methylhydroquinone (MHQ) (0.33 mM) (Duy et al., 2007), catechol (2.4 mM) (Tam et al., 2006), FA (1 mM) and MG (2.8 and 5.6 mM). All induced genes under these different conditions of electrophile treatment were listed and classified into regulons according to http://dbtbs.hgc.jp. These data were used for the hierarchical clustering analysis presented in Fig. 5.

General discussion 96 Thiol-specific stress response

GENERAL DISCUSSION

1. The role of the MarR/DUF24 regulators as sensors for electrophilic quinones and diamide in B. subtilis

All living organisms express molecular systems that enable them to resist a variety of toxic substances and environmental stresses. Exposure to ROS, which includes hydrogen peroxide, superoxide anion, hydroxyl radical and organic hydroperoxides (OHP), involves unique adaptive responses beyond those of general stress. ROS can damage proteins, DNA, lipids and membranes, which often lead to mutations and cell death. Thus, ROS must be removed rapidly to avoid cellular damage [Storz and Imlay, 1999]. To overcome oxidative stress, cells activate the expression of detoxification and repair systems. The cellular response to oxidative stress in bacteria is largely regulated at the transcriptional level [Mongkolsuk and Helmann, 2002]. In several cases, the corresponding regulator also functions as a direct sensor of oxidative stress, an example of the "one-component" regulatory strategy [Ulrich et al., 2005]. Regulatory proteins that sense oxidative stress employ a variety of mechanisms ranging from the oxidation of metal centers, as in SoxR and PerR, to the oxidation of cysteine, as in OxyR and OhrR [Mongkolsuk and Helmann, 2002; Paget and Buttner, 2003]. Due to its nucleophilic character, reactive cysteine residues in transcription factors can be oxidized by ROS leading to (i) disulfide bond formation or mixed disulfides with LMW thiols (S- thiolations) (ii) sulfenic, sulfinic or sulfonic acid formation and (iii) cyclic sulfenamide covalent bonds (Fig. 1.7) [Paget and Buttner, 2003; Barford, 2004]. In addition, cysteine residues can be also conjugated with chemically-diverse electrophilic compounds via thiol- (S)-alkylation [Giles et al., 2003]. Depending on their chemistry, electrophilic compounds react with thiols by different mechanisms. Alkylating agents, such as IAA, aliphatic epoxides and alkyl halides display SN2 electrophilic chemistry. The reaction of quinones, hydroquinones and α,β-unsaturated carbonyls involves a 1,4-reductive nucleophilic or Michael-type addition [Marnett et al., 2003]. Of these modifications, disulfide bonds and sulfenic acids, but neither the higher oxidation states of cysteine (sulfinic and sulfonic acid) nor covalent S-adducts with electrophiles, can be reversibly reduced by thiol-disulfide exchange reactions as catalyzed by the thioredoxin and glutaredoxin systems in bacteria [Holmgren et al., 2005].

General discussion 97 Thiol-specific stress response

d) S-S-R

SH SNO c) S-S- f) RSH SH NO,RNS SH SH HS - O ONOO- SH SNO S g) ROS a) ROS SH S S SOH b) S SH O O S-OH S SH S-OH e) + O SH S SH SH

Fig. 1.7. Chemical modifications to cysteine residues. (a) Sequential oxidation of cysteine by ROS gives rise to sulfenic acid (R-SOH), sulfinic acid (R-SOOH) and sulfonic acid (R-SO3H) derivatives. Alternatively, oxidation by ROS may promote (b) intramolecular cysteine disulfide bond formation within a protein, (c) intermolecular cysteine disulfide bond formation between proteins, or (d) a mixed disulfide between a cysteine- containing protein and a LMW thiol such as GSH or mycothiol. (e) Reduced cysteine residues can become oxidized to disulfides as a result of thiol-disulfide exchange reactions with an oxidized protein (or an oxidized LMW thiol such as oxidized GSH; G-S-S-G). Cysteine residues can also be modified by RNS such as nitric oxide (NO) to yield an S-nitrosothiol (R-SNO; f), and by peroxynitrite (NO3 ) to an S-nitrothiol (R-SNO2; g). This figure is adapted from Paget and Buttner (2003).

E. coli OxyR is one of the first characterized bacterial sensors of ROS. The OxyR regulon includes genes with protective and detoxification functions, such as katG (hydroperoxidase I), ahpCF (alkyl hydroperoxide reductase) and dps (DNA-binding protein). The oxidized OxyR binds its target promoters as a tetramer (Fig. 1.8) [Pomposiello and Demple, 2001]. In addition, OxyR activates transcription of genes that maintain intracellular thiols in their reduced states, including gorA (glutathione reductase), grxA (glutaredoxin) and trxA (thioredoxin 2) [Paget and Buttner, 2003]. OxyR can also be activated by other modifications of the active site thiol, including S-nitrosation, S-glutathionylation, and sulfenic acid formation in vitro [Kim et al., 2002], although the physiological relevance of these modifications remains controversial [Paget and Buttner, 2003].

General discussion 98 Thiol-specific stress response

Fig. 1.8. The oxyR regulon. The OxyR protein is produced constitutively and is oxidized by hydrogen peroxide (H2O2). The oxidized form of OxyR binds to promoter regions of target genes and activates transcription by protein–protein contact with RNA polymerase. OxyR-activated genes have direct and indirect antioxidant functions. Each subunit of the OxyR tetramer contains two cysteine residues that form intramolecular disulfide bonds upon exposure to H2O2. The disulfide bonds are re-reduced by GSH, which in turn is re-reduced by GSH reductase. The expression of the GSH reductase and glutaredoxin genes is under transcriptional control of OxyR, and thus the response is self-regulated. Abbreviation: GSH, glutathione. This figure is adapted from Pomposiello and Demple (2001).

The exposure of B. subtilis to ROS such as H2O2 or organic hydroperoxides (OHPs) causes the induction of an oxidative stress stimulon that is controlled by σB, PerR, Spx and OhrR [Fuangthong et al., 2001; Herbig and Helmann, 2001; Helmann et al., 2003; Nakano et al., 2003b]. The OHP resistance regulator OhrR controls the expression of the organic hydroperoxide resistance gene, ohrA, which encodes a thiol-dependent peroxidase (peroxiredoxin). Recent structural and biochemical studies on OhrA have shown that this enzyme contains alkyl hydroperoxide reductase activity and detoxifies OHP by reducing these peroxides to alcohols [Lesniak et al., 2002; Cussiol et al., 2003]. OhrR negatively regulates expression of ohrA by binding to an inverted repeat overlapping the ohrA promoter region. In contrast, a second organic hydroperoxide resistance gene, ohrB, is induced by general stress conditions and starvation for glucose and phosphate which is under control of σB [Völker et al., 1998; Fuangthong et al., 2001]. OhrR is essential for the response against lipophilic ROS by sensing the presence of OHPs [Sukchawalit et al., 2001; Lesniak et al., 2002]. OhrR homologs are present in both Gram-positive and Gram-negative bacteria such as Xanthomonas campestris [Mongkolsuk et al., 1998], Pseudomonas aeruginosa [Ochsner et

General discussion 99 Thiol-specific stress response al., 2001], Enterococcus faecalis [Rince et al., 2001], Actinobacillus pleuropneumoniae [Shea and Mulks, 2002] and Staphylococcus aureus [Chen et al., 2006].

The B. subtilis OhrR repressor senses OHPs with high sensitivity and high selectivity [Fuangthong et al., 2001; Lee et al., 2007]. Insights into the mechanism of peroxide sensing by OhrR have emerged from genetic and biochemical studies [Fuangthong and Helmann, 2002]. OhrR is a homodimer of 147 amino acid residues per subunit (MW~17 kDa) and belongs to the MarR family of transcription regulators [Fuangthong et al., 2001]. The fold of OhrR is similar to that of all other MarR family members and consists of six α helices and three β strands with topology: α1 (residues 14-36), α2 (residues 40-51), β1 (residues 54-57), α3 (residues 58-64), α4 (residues 68-81), β2 (residues 83-88), β3 (residues 95-101), α5 (residues 102-122) and α6 (residues 130-143). Each subunit is divided into two functional domains: the dimerization domain, which involves the N and C termin, α1, α2, α5, and α6; and the wHTH DNA binding domain, which consist of β1, α3, α4, β2, and β3. Helices α1 and α2 also form an interface between these well-defined functional domains [Hong et al., 2005]. Whereas most OhrR orthologs, like Xanthomonas campestris OhrR, contain two or more Cys residues that may participate in their regulatory cycle [Panmanee et al., 2006], B. subtilis OhrR has only a single reactive Cys residue per monomer [Fuangthong and Helmann, 2002]. Like other well-characterized peroxide sensors, OhrR is a peroxide-sensing regulator in that it relies on an active site cysteine (C15) residue, which is conserved amongst all described OhrR proteins [Fuangthong and Helmann, 2002; Kiley and Storz, 2004]. In addition, the structure of OhrR reveals that C15 is located at the N terminus of helix α1, suggesting that, besides its role in DNA binding, the helix-helix motif plays a key role on the peroxidation-mediated induction mechanism of OhrR (Fig. 1.9a) [Hong et al., 2005]. Moreover, the high sensitivity of the active site cysteine (C15) to peroxidation requires interactions with the opposed three tyrosine residues. This structure of OhrR reveals that the active site C15 residue is within hydrogen-binding distance of two tyrosine residues (Y29 and Y40) from the opposing subunit. Y19 is also critical for peroxide sensing (Fig. 1.9b) [Soonsanga et al., 2007]. Recently it was shown that the 1-Cys B. subtilis OhrR can be converted into a 2-Cys OhrR by introducing another cysteine residue in either position 120 or 124 like the 2-Cys Xanthomonas campestris OhrR protein [Soonsanga et al. 2008b]. Finally, the recent analysis of OhrR inactivation by OHPs cumene hydroperoxide (CHP) and linoleic acid hydroperoxide (LHP) suggests that this regulator may switch between reversible and irreversible inactivation pathways depending on the nature of the oxidant (Fig. 1.10) [Lee et al., 2007; Soonsanga et al., 2008a].

General discussion 100 Thiol-specific stress response

Fig. 1.9. The crystal structure of OhrR. (A) Cysteine residue (C15) of OhrR is critical for organic hydroperoxide sensing. Ribbon diagram of OhrR showing the two monomers (green and pink), with C15 displayed as red van der Waals spheres. The DNA-binding helices are at the bottom. (B) The interaction of C15 with the opposed tyrosine residues (Y19, Y29 and Y40) is also required. Close-up of the OhrR active site illustrates the close proximity of C15 to the Y29' and Y40' residues from the other subunit. Residues are represented by sticks, and carbon, nitrogen, oxygen, and sulfur atoms are gold, blue, red, and yellow, respectively. This figure is adapted from Soonsanga et al. (2007).

RS-SR RS-SR OhrR-SH

RSH 2ROH k3 R‘OOH k5

OhrR-S-SR k1 OhrR-SN

k4 k2 R‘OH H2O H2O RSH OhrR-SOH R‘= H (H2O2) R‘OOH (CHP) k6

R‘OH O OhrR-SO H(SO H) OH 2 3 (LHP)

O OH

Fig. 1.10. Oxidation pathways for B. subtilis OhrR. OhrR first is oxidized to the sulfenic acid form (k1) which retains DNA binding activity [Lee et al., 2007]. The OhrR sulfenate is functionally inactivated by either S- thiolation (k2), protein sulfenamide formation (k4), or overoxidation to the sulfinic and sulfonic acid derivatives (k6). The OhrR mixed disulphide can be rapidly re-activated by disulfide exchange in the presence of reduced thiols such as DTT (k3). The OhrR sulfenamide is slowly re-reduced by DTT (k5) whereas the OhrR sulfinic and sulfonic acid derivatives are not. The figure is adapted from Soonsanga et al. (2008a).

In its natural habitat, the Gram-positive soil-dwelling bacterium B. subtilis is exposed to a variety of toxic and antimicrobial agents that also include phenolic compounds. Catecholic or hydroquinone-like intermediates are common in the metabolism of nitroaromatic compounds or organic insecticides, and thus are often present in the soil [Vaillancourt et al., 2006]. Catechol and hydroquinone-like compounds can also cause toxic effects in microorganisms since these can be autooxidized to the corresponding ortho- and para-benzoquinones. Quinones can exert their toxicity as oxidants and/or electrophiles

General discussion 101 Thiol-specific stress response

[Monks et al., 1992]. As oxidants, quinones redox cycle with their semiquinones producing ROS such as superoxide anion or hydrogen peroxide. As electrophiles, quinones can form S- adducts with cellular thiols via the thiol-(S)-alkylation reaction that leads to aggregation of thiol-containing proteins in vivo as irreversible thiol-modification [Liebeke et al., 2008].

Recently, our previous proteomic and transcriptomic expression profiling of B. subtilis after exposure to catechol and MHQ has led to the identification of novel regulatory mechanisms that govern the thiol-specific stress response, which are conserved among low GC Gram-positive bacteria [Tam et al., 2006b; Duy et al., 2007b]. A strong overlap in the response of B. subtilis to catechol, MHQ and diamide was observed. Diamide is an electrophilic azo-compound that reacts with protein thiolates leading to the formation of disulphides or S-thiolated proteins (Fig. 1.11) [Leichert et al., 2003; Hochgräfe et al., 2007; Leelakriangsak et al., 2008]. In contrast, electrophilic quinones target cysteine via the thiol- (S)-alkylation mechanisms resulting in a direct depletion of thiols. Thus, quinone compounds and diamide have the same mode of action but react via different thiol chemistries and lead to depletion of protein and non-protein LMW thiols. This leads to induction of the complex thiol-specific stress response to electrophilic compounds in B. subtilis, which is regulated by a complex network of thiol-stress sensing transcription factors Spx, CtsR, PerR, CymR [Leichert et al., 2003; Nakano et al., 2003b; 2005; Zuber, 2004; Leelakriangsak et al., 2007; Leelakriangsak and Zuber, 2007]. In addition, our recent studies identified the novel MarR- type repressors MhqR (YkvE), YodB and CatR (YvaP) that confer resistance to diamide and quinones via the control of specific detoxification pathways [Töwe et al., 2007; Leelakriangsak et al., 2008; Antelmann et al., 2008; Huyen et al., 2008a].

Fig. 1.11. Thiol-specific reactions of diamide in B. subtilis. Diamide is an electrophile azo-compound which reacts with protein thiolates and causes non-native disulfide bond formation. AzoR1 and AzoR2 confer also resistance to diamide and functions in detoxification of diamide via the reductive cleavage of the azo group, resulting in two molecules 1,1´-Dimethylurea. This figure is adapted from Leelakriangsak et al., 2008.

The MhqR (YkvE) regulon consists of mhqA, mhqED, mhqNOP and azoR2, which encode thiol-dependent dioxygenases, oxidoreductases, a phospholipase and an azoreductase, which function in reduction of quinones and diamide or in ring cleavage of quinone-S- aggregates [Töwe et al., 2007; Antelmann et al., 2008]. Another thiol-stress-inducible

General discussion 102 Thiol-specific stress response repressor YodB controls Spx, the nitroreductases YodC and the AzoR2 paralogous azoreductase AzoR1 [Duy et al., 2007b; Leelakriangsak et al., 2007; 2008]. A further YodB- paralogous thiol-stress sensing MarR-type regulator, CatR (YvaP) regulates the expression of the thiol-stress inducible yfiDE (catDE) operon, which encodes a DoxX-like oxidoreductase CatD and a catechol-2,3-dioxygenase (CatE) [Tam et al., 2006b; Huyen et al., 2008a]. Together, the MarR-type repressors MhqR, YodB and CatR control paralogous detoxification genes, including azoreductases, nitroreductases, thiol-dependent dioxygenases and DoxX-like oxidoreductases that confer resistance to quinone compounds and/or diamide in B. subtilis [Töwe et al., 2007; Leelakriangsak et al., 2008; Antelmann et al., 2008; Huyen et al., 2008a]. Thus, these novel MarR-type repressors YodB, CatR and MhqR constitute the novel quinone resistance network of B. subtilis (Fig. 1.12).

Fig. 1.12. Thiol-dependent detoxification of quinones (BQ). (1) BQ is detoxified to hydroquinone in the two- electron reduction by NAD(P)H-dependent quinone reductases (AzoR1, AzoR2, YodC and MhqN), which are controlled by YodB and MhqR. (2) The one-electron reduction of BQ generates semiquinone radical, which reduces molecular oxygen to superoxide anion. (3) Conversion of accumulated semiquinone radical to hydroquinone. (4) BQ reacts primarily electrophilic with cellular protein thiols via the Michael addition chemistry (thiol-(S)-alkylation) leading to S-adduct formation. Since BQ has two reactive sites for the nucleophilic addition, it can cross-link different protein thiols in vivo. (5) The paralogous thiol-dependent dioxygenases (MhqA, MhqE, MhqO), which are regulated by MhqR might be involved in the dioxygenolytic cleavage of the thiol-(S)-adducts that are probably exported via metal ion efflux systems. This figure is modified from Liebeke et al. (2008).

Azoreductases are flavin-containing enzymes that have the unique ability to catalyze a wide range of biochemical reactions and have broad substrate specificity [Fraaije and Mattevi, 2000]. These enzymes catalyze the NADH-dependent two-electron reductions of a wide range of substrates, including quinones, quinoneimines, azodyes and nitrogroups, to protect cells

General discussion 103 Thiol-specific stress response against the toxic effects of free radicals and ROS arising from one-electron reductions [Fraaije and Mattevi, 2000; Cavelier and Amzel, 2001]. The paralogous azoreductases, which are conserved among Gram-positive bacteria, are major quinone resistance determinants and have common functions as quinone reductases thus, preventing toxic quinones from undergoing redox-cycling and causing depletion of cellular thiols [Töwe et al., 2007; Antelmann et al., 2008; Leelakriangsak et al., 2008]. Moreover, it has been demonstrated recently that AzoR2 has NADH:DCIP oxidoreductase and azoreductase activity [Nishiya and Yamamoto, 2007]. Therefore, both azoreductases, AzoR1 and AzoR2, could also be involved in the reductive cleavage of the electrophile azo-compound diamide to 1,1´-diamethylurea. In addition, another flavoenzyme, the nitroreductase MhqN and a paralogous nitroreductase YodC also respond to thiol-specific stress [Töwe et al., 2007; Leelakriangsak et al., 2008]. Thus, paralogous nitroreductases are co-regulated with paralogous azoreductases in response to thiol-reactive electrophiles. Nitroreductases catalyze the two-electron reduction of nitrogroups in nitroaromatic compounds to nitroso, hydroxylamide intermediates and finally primary amines [Bryant and DeLuca, 1991]. Since quinones represent substrates for nitroreductases [Nivinskas et al., 2002], YodC and MhqN could also function in quinone reduction in B. subtilis [Leelakriangsak et al., 2008]. Interestingly, neither of these azo- or nitroreductases was induced by oxidative stress provoked by H2O2 or the redox-cycling agent paraquat supporting that these enzymes are specifically involved in quinone and electrophile reduction and not in the oxidative stress response. As ROS are produced upon auto-oxidation of MHQ and catechol, this could account for the induction of the PerR-controlled oxidative stress response by electrophiles.

The dioxygenases are also a major part of the thiol-specific stress response that confers resistance to quinone-like electrophiles. Whereas azoreductases and nitroreductases function in the thiol-independent detoxification of electrophiles, dioxygenases could be involved mainly in detoxification of redox-stable quinone-S-adducts [Leelakriangsak et al., 2008; Antelmann et al., 2008]. Thus, we hypothesize that these dioxygenases are another class of thiol-dependent enzymes such as the thiol-dependent peroxidase OhrA. The MhqR-regulated dioxygenases are conserved also among pathogenic Gram-positives including Bacillus anthracis, Listeria or Staphylococcus. Dioxygenases belongs to the vicinal oxygen chelate (VOC) superfamily that is structurally related proteins, which provide a metal coordination environment with two or three open or readily accessible coordination sites to promote direct electrophilic participation of metal ion in catalysis [Armstrong, 2000]. The paralogous thiol- dependent dioxygenases (MhqA, MhqE and MhqO) share similarity to the chlorohydroquinone/hydroquinone 1,2-dioxygenase of Sphingomonas paucimobilis LinE [Endo et al., 2005]. As mentioned above, MhqA, MhqE and MhqO could be involved in the thiol-dependent ring-cleavage of quinone-S-adducts, which are formed in the detoxification reaction of the LMW thiol buffer with quinones [Töwe et al., 2007]. This is supported by the finding that the fourth thiol stress-induced dioxygenase CatE was previously characterized as

General discussion 104 Thiol-specific stress response catechol-2,3-dioxygenase [Tam et al., 2006b]. Furthermore, another CatR-dependent enzyme CatD is similar to DoxX-like oxidoreductases. This enzyme is similar to the membrane-bound DoxX- and DoxD-subunit of the thiosulfate:quinone oxidoreductase of Acidianus ambivalens involved in the early steps of sulfur oxidation [Müller et al., 2004]. CatD is paralogous to the MhqR-regulated DoxX-like oxidoreductase MhqP [Töwe et al., 2007; Huyen et al., 2008a]. Our proteome and transcriptome profiling also revealed that more than 15 other NAD(P)H- oxidoreductases respond to electrophiles and might contribute to either the one- or two- electron reduction of quinones in B. subtilis [Duy et al., 2007b]. These NAD(P)H- oxidoreductases might be regulated by other YodB paralogs of B. subtilis, as we have already shown for the NAD(P)H-flavin oxidoreductase YfkO that is regulated by the transcriptional MarR/DUF24-type activator YybR [Antelmann et al., 2008; our unpublished data].

In summary, these conserved novel thiol-stress-responsive mechanisms seem to have evolved to detoxify quinone-like electrophiles, which are naturally produced during electron transfer in the respiratory chain [Antelmann et al., 2008]. We have further proposed that DNA binding activity of the thiol-stress sensing YodB repressor could be inhibited via thiol-(S)- alkylation at the conserved Cys6 residue in response to quinone-like electrophiles in vitro. This alkylation of regulatory thiols is an irreversible thiol-modification that cannot be reduced using the thiol-disulfide exchange reaction as catalyzed by the thioredoxin-thioredoxin reductase system [Leelakriangsak et al., 2008]. In addition, we have shown that the conserved Cys6 residue of YodB was essential for repression using site directed mutagenesis in vivo and in vitro. Cys residues have been shown to be involved in the redox-control of other oxidative stress transcription factors, such as OxyR, Hsp33 of E. coli and OhrR of B. subtilis [Graumann et al., 2001; Hong et al., 2005; Nakano et al., 2005; Lee et al., 2007]. Thus, YodB was identified as a novel MarR/DUF24-family repressor, which senses thiol-reactive electrophiles via the conserved Cys6 residue. In addition, mutational analyses also indicated that the Cys101 and Cys108 residues of YodB may have role in the repressor activity and response to quinones and diamide. This triple mutant, Cys6, 101, 108A shows reduced repression, but also lower responsiveness to inducers [Leelakriangsak et al., 2008]. Furthermore, using mass spectrometry, we uncovered the formation of YodB-Cys-S-adducts upon treatment of the purified YodB repressor with MHQ and catechol in vitro suggesting that DNA-binding activity of YodB could be regulated via thiol-(S)-alkylation in vitro. Cysteine residues are strong nucleophiles, which are shown to be conjugated with electrophilic compounds such as p-benzoquinone and hydroquinones to form thiol-S-adducts [Giles et al., 2003; Dennehy et al., 2006]. Thus, the derepression of YodB in response to quinones might be caused by quinone-S-adduction or even aggregation which remains to be shown in vivo [Antelmann et al., 2008]. Moreover, S-thiolation has been shown as one regulatory mechanism for OhrR and OxyR [Kim et al., 2002; Lee et al., 2007]. This mechanism of S-thiolation is similar to the S-glutathionylation, which is thought to protect protein thiols against overoxidation to irreversible sulfonic acids [Lee et al., 2007; Hochgräfe

General discussion 105 Thiol-specific stress response et al., 2007]. Thus, it could be also possible that YodB is regulated in vivo in response to diamide via the formation of mixed disulfides containing cysteine, CoASH or other LMW thiols [Leelakriangsak et al., 2008]. However, YodB is redox-controlled via intersubunit disulfide formation in response to diamide in vivo [F. Schuster, Diploma thesis]. This regulation mechanism is similar to those of OhrR in Xanthomonas campestris, which also involves an intersubunit disulfide bond between one N-terminal Cys22 of one subunit and one C-terminal Cys127 of the other subunit [Panmanee et al., 2006]. We currently defined the nature of the intermolecular disulfide bond, which is formed in the YodB dimer upon diamide stress using immunoprecipitation and mass spectrometry.

The MarR-type repressor MhqR also contains a cysteine residue (Cys128), which is located in dimerization domain and accessible for thiol-modification according to published structures for B. subtilis OhrR and E. coli MarR [Alekshun et al., 2001; Hong et al., 2005]. In contrast to Cys6 in YodB orthologs, Cys128 of MhqR is not conserved among other Gram- positive bacteria. In addition, in contrast to YodB and YvaP, DNA binding activity of MhqR was not directly inhibited by quinones or diamide in vitro. Thus, the signal sensed by MhqR and the mechanisms, which lead to derepression of the MhqR regulon, are still unknown [Töwe et al., 2007; Antelmann et al., 2008].

YodB is highly conserved among low-GC Gram-positive bacteria, and there are several YodB paralogs in the genome of B. subtilis. In total, B. subtilis encodes eight MarR/DUF24-family regulators (YybR, YdeP, YkvN, YtcD, YdzF, HxlR, YodB and YvaP). The first identified MarR/DUF24-family regulator of B. subtilis was HxlR, which is an activator of the hxlAB operon that encodes the genes for the RuMP pathway [Yurimoto et al., 2005]. The 3-hexulose-6-phosphate-sythase HxlA and the 6-phospho-3-hexuloisomerase HxlB are activated in response to FA stress and are involved in FA detoxification. In contrast to HxlR, which is activated after exposure of cells to FA stress, YodB and CatR (YvaP) act both as transcriptional repressors [Leelakriangsak et al., 2008; Huyen et al., 2008a]. YodB paralogs have a conserved cysteine residue in the N-terminal part followed by a conserved proline residue. This N-terminal Cys residue is also conserved among MarR/DUF24 family members in other Gram-positive bacteria, and thus it is tempting to speculate that this Cys serves as conserved sensor for electrophile compounds.

The thiol-stress sensing MarR/DUF24 repressor CatR (YvaP), which regulates the yfiDE (catDE) operon in B. subtilis, is strongly related to YodB as judged by sequence similarity and function. Among the YodB-paralogs encoded in the B. subtilis genome, CatR and YodB display most close sequence similarities. Both YodB and CatR repressors also share the conserved N-terminal Cys6 and Cys7 residue, respectively that is essential for regulation and sensing of electrophile compounds in vivo and in vitro. In addition, our studies showed that the conserved basic amino acids K19, R20 are also essential for repression by

General discussion 106 Thiol-specific stress response

CatR in vivo [Huyen et al., 2008a]. It was shown previously that neighboring basic amino acids facilitate the thiol-(S)-alkylation of the reactive thiolate anion [Dennehy et al., 2006]. In addition, all conserved basic lysine and arginine residues located in the HTH motif are essential for DNA binding of CatR as shown previously also for the OhrR repressor [Hong et al., 2005]. Our results suggest that CatR is regulated via thiol-(S)-alkylation or aggregation at the conserved Cys7 by quinones [Leelakriangsak et al., 2008; Huyen et al., 2008a]. However, non-reducing SDS-PAGE analyses showed intersubunit disulfide bond formation for YvaP by diamide and quinones in vitro [Huyen et al., 2008a]. We are currently raising antibodies against YvaP, which will be used to confirm if regulation of YvaP by diamide occurs via intersubunit disulfide formation in vivo.

Together, our data reveal that B. subtilis has evolved a sophisticated network of three MarR-type repressors YkvE (MhqR), YodB and YvaP (CatR) that control paralogous resistance determinants to thiol-specific quinone compounds. YodB and CatR control of quinone detoxification genes including the azoreductase AzoR1 and the thiol-dependent dioxygenase CatE and the similar paralogous genes are controlled by MhqR. Thus, MhqR, YodB and CatR are novel determinants of this complex network that senses and responds to thiol-reactive electrophiles in B. subtilis [Töwe et al., 2007; Leelakriangsak et al., 2008; Huyen et al., 2008a]. Detailed future work is necessary to elucidate the functions and regulation of all other uncharacterized MarR/DUF24 family regulators and their regulons in the thiol-stress responsive regulatory network.

2. The MG and FA stimulons of B. subtilis

Quinones and α,β-unsaturated aldehydes are naturally occurring electrophilic compounds, which target cysteine residues via the thiol-(S)-alkylation reaction. Quinone compounds and α,β-unsaturated aldehydes display the same adduction chemistry, these react with cysteine thiols via the Michael addition mechanism. Like quinone compounds, α,β- unsaturated aldehydes are highly toxic natural electrophiles, which are generated from oxidation of amino acids, lipids and carbohydrates [Marnett et al., 2003; Farmer and Davoine, 2007].

MG is the most studied toxic natural α-oxoaldehyde, which is inevitably produced from triose-phosphate intermediates as by-product of phosphorylating glycolysis and also from amino acids and acetone [Ferguson et al., 1998; Booth et al., 2003; Kalapos, 2007]. MG reacts with nucleophilic centers of DNA, RNA and with the side chains of the amino acids arginine, lysine and cysteine. The reaction of MG reacting with arginine and lysine residues results in the formation of the widely studied advanced glycation endproducts (AGE), such as argpyrimidine or MG-cross-linked lysine dimers (MOLD) [Bourajjaj et al., 2003]. The major protective mechanism against the deleterious effects of MG in E. coli is the spontaneous

General discussion 107 Thiol-specific stress response reaction of MG with GSH to form hemithiolacetal, followed by isomerization to S- lactoylglutathione and conversion to D-lactate by the glyoxalase I and II enzymes (Fig. 1.13a) [Ferguson et al., 1998; Booth et al., 2003].

CH3 CH3 CH3 Glx-II CH3 A Glx-I +R-SH +H2O C O C O HC OH HC OH

O H C H C OH C O R-SH C O

SR SR OH

Methylglyoxal (MG) Hemithioacetal S-Lactoylthiol Lactate

B NAD NADH+H H H H H AdhA +R-SH +H2O C O HCOH CO C O

H RS HO RS R-SH

Formaldehyde (FA) S-Hydroxy- S-Formylthiol Formate methylthiol

Fig. 1.13. Thiol-dependent detoxification pathways for MG (A) and FA (B) in bacteria. (A) MG reacts spontaneously with GSH to form hemithiolacetal, followed by isomerization to S-lactoylglutathione by the glyoxalase I enzyme. S-lactoylglutathione is substrate for glyoxalase II enzyme, which is converted to D-lactate and free GSH. This scheme is adapted from Ferguson et al. (1998). (B) In prokaryotes and eukaryotes, FA is detoxified by GSH-dependent FA dehydrogenases (FDH) [Harms et al., 1996]. FA reacts spontaneously and reversible with GSH to S-hydroxymethylglutathione. FDH-like enzymes such as class III alcohol dehydrogenases catalyze the NAD-dependent oxidation of S-hydroxymethylglutathione into S- formylglutathione. This GSH thiol ester is reversibly hydrolyzed by a S-formylglutathione hydrolase to GSH and formiate. This scheme is adapted from Jensen et al. (1998).

FA is another highly toxic carbonyl compound that is known to cause protein-protein and protein-DNA cross-links in vivo [Orlando et al., 1997]. Like MG, FA also reacts as electrophile with the basic amino acid side chains of arginine and lysine. Since FA is the key intermediate in the metabolism of C1 compounds in methanotrophic and methylotrophilc bacteria, it is ubiquitously distributed in the environment. Thus, prokaryotes and eukaryotes have evolved conserved detoxification pathways for FA detoxification. These include GSH- dependent FA dehydrogenases (FDH), which are evolutionary conserved in all kingdoms of life [Harms et al., 1996]. FDH-like enzymes catalyze the NAD-dependent oxidation of S- hydroxymethylglutathione, formed between GSH and FA, into S-formylglutathione (Fig. 1.13b). This GSH thiol ester is reversibly hydrolyzed by a S-formylglutathione hydrolase to GSH and formiate [Jensen et al., 1998]. In addition, the RuMP pathway for FA fixation is involved in FA detoxification in many bacteria including also the non-methylotrophic B. subtilis [Yurimoto et al., 2005; Kato et al., 2006]. The 3-hexulose-6-phosphate synthase and 6-phospho-3-hexuloisomerase are key enzymes in the RuMP pathway, which are encoded by the hxlAB operon in B. subtilis. Interestingly, this hxlAB operon is positively regulated by the

General discussion 108 Thiol-specific stress response

MarR/DUF24 family protein HxlR which is close related to the quinone-responsive YodB repressor [Yurimoto et al., 2005; Leelakriangsak et al., 2008].

Since both MG and FA are toxic electrophiles, eukaryotic and prokaryotic cells have evolved defense mechanisms to protect directly by detoxification against electrophile toxicity and indirectly by the repair of the damages. In this main part of the thesis, we have applied genome wide transcriptome and proteome profiling to study the mode of action of the toxic electrophilic compounds MG and FA in B. subtilis. The results showed that both compounds are strong thiol-reactive compounds and triggered a similar cellular response as those of quinone-like electrophiles and diamide [Leichert et al., 2003; Tam et al., 2006b; Duy et al., 2007b; Antelmann et al., 2008]. Both MG and FA induce the thiol stress-specific Spx, CtsR, CymR, YodB, PerR, ArsR, CzrA and CsoR regulons [Huyen et al., 2008b]. For example, the CtsR-regulated clpE gene was more than 100-fold upregulated in the transcriptome experiments by FA. In addition, the microarray data also noted maximal derepression ratios for almost all CymR regulon genes by FA. The CymR-controlled ssu operon was very strongly elevated by FA stress, which encodes an uptake system for aliphatic sulfonate compounds and has been not detected as induced by other thiol-reactive compounds. Altogether, these results indicated an imbalanced thiol-redox homeostasis due to depletion of LMW buffer such as cysteine in response to both compounds which was verified also by the metabolomic approach [Huyen et al., 2008b]. In addition, medium supplementation experiments supported that cysteine protects cells against FA and MG toxicity as found in previous studies for electrophilic quinones [Liebeke et al., 2008].

The thiol-reactive nature of FA and MG is further supported by the induction of the YodB regulon genes yodC, spx and azoR1, which encode quinone resistance determinants. YodB was previously shown as direct sensor for electrophiles, which is redox-regulated via thiol-(S)-alkylation by quinone compounds in vitro [Leelakriangsak et al., 2008]. Thus, it was hypothesized that carbonyl compounds also target regulatory thiols of redox-sensing transcription factors. In addition, the ykvN and yybR genes encoding two further MarR/DUF24-family regulators, respond to MG, FA and quinone-compounds. Finally, the yfiDE (catDE) operon was induced by carbonyls and quinones, which encodes a thiol- dependent catechol-2,3-dioxygenase regulated by the MarR/DUF24 paralog CatR (YvaP) in B. subtilis (see previous section of the general discussion).

Interestingly, the PerR regulon was induced in response to MG and FA. The induction of the PerR-controlled oxidative stress response in B. subtilis under MG and FA stress conditions reflects the ROS producing and electrophilic mode of actions of MG and FA. The induction of the oxidative stress specific PerR regulon genes by FA and MG could indicate that electrophilic carbonyl and quinone compounds both are able to produce ROS [Mongkolsuk and Helmann, 2002]. Quinone compounds can undergo redox-cycling reactions

General discussion 109 Thiol-specific stress response leading to the generation of semiquinone anions and ROS [O´Brien, 1991]. MG has been shown to provoke free radical generation involving ROS and RNS as well as organic MG radicals or cross-linked MG radicals, which contribute also to MG toxicity mechanisms in eukaryotic cells [Kalapos, 2007]. Similarly, FA causes cross-links between proteins and DNA, which results in the generation of secondary electrophiles, which in turn could lead to ROS production [Orlando et al., 1997].

Since FA and MG both also cause protein-DNA cross-links, for example between the amino groups of lysine and cytosine [Orlando et al., 1997], the DNA damage responsive SOS regulon is induced under both carbonyl stress conditions. The induced genes of the SOS regulon include the DNA repair and recombination systems uvrB, uvrC and recA and also a DNA alkylation repair enzyme, yhaZ. The induction of DNA damage-responsive regulons by FA has been shown also in genome-wide expression profiling in Pseudomonas putida [Roca et al., 2007].

Furthermore, different cation efflux systems were strongly up-regulated by all electrophiles such as carbonyls, quinones and diamide. These include the ArsR-dependent arsR-yqcK-arsBC operon required for resistance to As(III), Sb(III), Cd(II) and Ag(I) [Moore et al., 2005]. Additionally, the CsoR-regulated copZA operon which is activated by Cu(II) [Smaldone and Helmann, 2007] is induced by MG and FA. The microarray and proteome results of FA and MG-treated cells indicate that metal ion efflux systems also could function in protection against thiol-reactive electrophiles by extrusion of the toxic electrophiles [Antelmann et al., 2008]. In addition, many genes of the σD regulon that are involved in motility, chemotaxis and cell wall turnover were up-regulated by aldehyde treatment as observed also in response to quinone compounds such as MHQ [Duy et al., 2007b].

However, we noted also many differences in the induction profiles by FA and MG compared to diamide and quinones, which might be linked to specific detoxification pathways. For example, the microarray data indicate the specific induction of two operons by MG that could be involved in formate detoxification. These included the yrhED operon, which encodes the putative formate dehydrogenase YrhE and the yrhFG-yrzI operon encoding the putative formate transporter YrhG. The specific induction of both yrhED and yrhFG-yrzI operons involved in formate detoxification by MG could be caused by contamination of commercially available MG with formate [Kalapos, 1999].

Taken together, our global expression profiling strongly implies that aldehydes represents a specific class of thiol-reactive electrophiles, which target cysteine thiols via the alkylation chemistry, thus depleting the pool of the thiol redox buffer which was also shown using the metabolomic approach.

General discussion 110 Thiol-specific stress response

Finally, the most interesting results derived from the genome-wide study of B. subtilis to MG and FA was the induction of two specific regulons involved in FA detoxification. These are the previously described HxlR regulon and the novel AdhR (YraB) regulon. HxlR is the FA-inducible MarR/DUF24-family repressor, which regulates the hxlA-hxlB operon that encodes enzymes of the RuMP pathway for FA detoxification.

Since FA is the key intermediate in the metabolism of C1 compounds in methanotrophic and methylotrophilic bacteria, it is produced through the degradation of compounds containing methyl or methoxyl groups, e.g., lignin and pectin. Because FA is toxic to all organisms, FA detoxification pathways are found in a wide variety of organisms. These include GSH-dependent FA dehydrogenases (FDH), which are evolutionary conserved in all kingdoms of life [Harms et al., 1996]. FDH-like enzymes catalyze the NAD-dependent oxidation of S-hydroxymethylglutathione, formed between GSH and FA, into S- formylglutathione. This GSH thiol ester is reversibly hydrolyzed by a S-formylglutathione hydrolase to GSH and formiate [Jensen et al., 1998]. In addition, the RuMP pathway for FA fixation is involved in FA detoxification in many bacteria including also the non- methylotrophic B. subtilis [Yurimoto et al., 2005; Kato et al., 2006].

The RuMP pathway, which was originally found in methylotrophic bacteria, are now found in a variety of microorganisms including bacteria and archaea for FA fixation and detoxification [Mitsui et al., 2000; Vorholt et al., 2000; Yurimoto et al., 2002; Kato et al. 2006]. In bacteria, the RuMP pathway functions as a highly efficient system for trapping free FA at relatively low concentration [Ferenci et al., 1974]. In the RuMP pathway, 3-hexulose-6- phosphate synthase (HPS) and 6-phospho-3-hexuloisomerase (PHI) are key enzymes. HPS catalyses the aldol condensation of FA with ribulose 5-phosphate. The subsequent product, D- arabino-3-hexulose 6-phosphate, is isomerized by PHI to form fructose 6-phosphate [Arfman et al., 1990; Yurimoto et al., 2005; Kato et al., 2006]. The fructose 6-bisphosphate formed then converted to fructose 1,6.bisphosphate, which is subsequently metabolized into pyruvate via the glycolytic pathway (cleavage stage). Ribulose 5-phosphate, the acceptor of FA, is regenerated from fructose 6-phosphate and glyceraldehyde 3-phosphate through a combination of transketolase, transaldolase, and several reactions for pentose phosphate isomerization (regeneration stage) [Kato et al., 2006].

Additionally, it was recently demonstrated that the RuMP pathway is responsible for the detoxification of FA in non-methylotrophs during growth on a FA-containing medium, e.g. in Bacillus subtilis [Yasueda et al., 1999]. The HPS–PHI system is advantageous to other FA-eliminating reactions such as the GSH-dependent FA dehydrogenase system. The HPS– PHI catalyzed reaction is an exergonic reaction and does not require any cofactors like GSH and both the substrates and products of the HPS–PHI-catalyzed sequential reaction (e.g.,

General discussion 111 Thiol-specific stress response ribulose 5-phosphate and fructose 6-phosphate) are general metabolites that are essentially present in all organisms [Orita et al., 2007].

The genes encoding these two enzymes were first cloned from an obligate methylotroph, Methylomonas aminofaciens 77a [Yanase et al., 1996]. These genes have also been cloned from a facultative methylotroph, Mycobacterium gastri MB19 [Mitsui et al., 2000] and a thermotolerant methylotroph, Bacillus brevis S1 [Yurimoto et al., 2002]. Genes homologous to HPS and PHI have been found in a variety of non-methylotrophic organism, including bacteria and archaea. In the non-methylotroph, B. subtilis, the hxlA (yckG) and hxlB (yckF) encode for HPS and PHI that are involved in FA detoxification [Yasueda et al., 1999; Yurimoto et al., 2005]. This operon is regulated by the transcriptional activator HxlR (YckH) encoded by hxlR (yckH), which is oriented in the opposite direction relative to hxlA and hxlB and is close related to the quinone-responsive YodB repressor [Yurimoto et al., 2005; Leelakriangsak et al., 2008]. Disruption of either the hxlR gene or the hxlAB operon results in increased sensitivity to FA, suggesting that the RuMP pathway acts as a detoxification system for this compound [Yasueda et al., 1999]. HxlR directly binds to specific DNA sequences within the intergenic region between the hxlAB operon and the hxlR gene. Moreover, this HxlR binding site is necessary for FA-induced expression of the hxlAB operon [Yasueda et al., 1999; Yurimoto et al., 2005]. The HxlR protein belongs to the MarR/DUF24 protein family of unknown function, which is widely distributed in prokaryotes. More than 270 members of this protein family belong to the superfamily of winged helix DNA-binding proteins, which is made up of proteins with a helix-turn-helix motif [Gajiwala and Burley, 2000]. Thus, a DNA-binding protein HxlR, which is now designed as a transcriptional activator of the “helix-turn-helix, HxlR type”, is necessary for FA-induced expression of hxlAB operon. Furthermore, two 25 bp regions (BRH1 and BRH2) in the promoter region of the hxlAB operon have been identified as the binding site for HxlR. However, the details as to how FA modulates hxlAB expression are still unclear. Yurimoto et al. (2005) proposed two possibilities to explain this issue. One possibility, which is similar to the SoxR-type transcriptional activation system in E. coli [Hidalgo and Demple, 1994; Hidalgo et al., 1998; Chander and Demple, 2004], is that a conformational change in HxlR, induced by FA binding, may stimulate the recruitment of RNA polymerase to the hxlAB transcription initiation site. Another possibility to explain FA- and HxlR-dependent induction of the hxlAB operon is that FA may bind to other factors, which in turn activate HxlR. It is tempting to speculate that also the conserved N-terminal Cys residue plays a role in regulation of the DNA binding activity of HxlR. Furthermore, we observed the very high induction of this hxlAB operon (20-140-fold) and the autoregulatory hxlR gene (3-10-fold) by either carbonyl compound, MG and FA. The global thiol-reactive mode of action of FA further supports the control mechanism of MarR/DUF24-family regulators via the thiol-(S)-alkylation at the conserved regulatory Cys residue, which is reflective for quinones and α, β-unsaturated carbonyls such as MG and FA.

General discussion 112 Thiol-specific stress response

The novel FA-inducible MerR/NmlR -type regulator AdhR (YraB) regulates the adhA- yraA operon as thiol-dependent FA detoxification pathway

Besides the hxlAB operon, the microarray results identified also the conserved thiol- dependent FA dehydrogenase-encoding adhA gene, which is cotranscribed with the cysteine protease of the DJ1/PfpI family encoded by yraA and the yraC gene, which encodes a γ- carboxymuconolactone decarboxylase (CMD) as the most strongly induced upon FA and MG treatment. These genes are regulated by a MerR-type regulator, YraB (AdhR) that belongs to a sub-family of NmlR-like proteins, which all control adhA-like target genes. The characterization of AdhR as a novel aldehyde sensing regulator and the definition of the AdhR regulon was one further part of this thesis.

Previous transcriptome analyses have revealed that yraA is also most strongly induced in response to other thiol-specific stress conditions, such as diamide, catechol and MHQ [Leichert et al., 2003; Tam et al., 2006b; Duy et al., 2007b]. Expression of yraA was also inducible by acid, ethanol and vancomycin, and yraA mutant cells were sensitive to acid stress [Thackray and Moir, 2003; Jervis et al., 2007]. In the case of other thiol-specific stress conditions or acid stress, induction of yraA occurs in a Spx-dependent manner [Eiamphungporn and Helmann, 2008]. Recent studies further showed that the previously suggested σM-dependent expression of yraA is derived from the σM-dependent transcriptional activation of spx since yraA belongs to those genes that are most strongly activated by Spx [Nakano et al., 2003b; Eiamphungporn and Helmann, 2008]. Thus, expression of yraA is activated either independently of Spx by AdhR from a promoter upstream adhA gene or Spx- dependently from the yraA specific promoter, which is located in the adhA-yraA integenic region [Eiamphungporn and Helmann, 2008]. This double control of yraA by AdhR and Spx was also shown in our Northern blot analysis.

YraA is similar to the DJ-1/PfpI-family cysteine proteinases that are also evolutionary conserved proteins and of thus far unknown functions [Wei et al., 2007]. This DJ-1/PfpI family includes human DJ-1, yeast Hsp31p, four E. coli homologs YhbO, SCRP 27a, YajL and Hsp31, which all have been crystallized. Recently, Hsp31p has been shown to be involved in the protection against ROS [Skoneczna et al., 2007]. The structures of these proteins are similar. They all possess a conserved Cys residue in the nucleophile elbow motif. According to the DJ-1/PfpI-family homology, YraA could function in repair of thiol- modifications in proteins caused by different electrophiles and related toxic compounds. However, according to previous studies, YraA had no detectable protease or chaperone activity [Thackray and Moir, 2003]. Phenotype analyses showed that the sensitivity of the ∆yraA mutant was similar to the wild type after exposure to FA (data not shown). Since yraA displayed sequence similarity to the σB-dependent yfkM gene, we investigated the growth phenotype of the ∆yraA∆yfkM double mutant after exposure to FA stress. The ∆yraA∆yfkM

General discussion 113 Thiol-specific stress response double mutant was more sensitive to FA than the wild type. Thus, both putative DJ-1/PfpI- family cysteine proteinases can replace each other in FA-detoxification probably by repair of proteins with specific thiol-modifications [Huyen et al., 2008b].

The array data showed also the very high induction (10-40-fold) of the adhA gene after FA and MG treatment which encodes a NADH-dependent alcohol dehydrogenase. AdhA is similar to GSH-dependent FA dehydrogenases (FDH), which are evolutionary conserved among prokaryotic and eukaryotic organisms [Harms et al., 1996]. FDH, a ubiquitous cytosolic enzyme, belongs structurally to the family of alcohol dehydrogenases and is actually homologous with the class III alcohol dehydrogenases, the pyridine nucleotide-dependent enzymes, which catalyzes the transformation of adducts of GSH such as hydroxymethylglutathione or S-nitrosoglutathione (GSNO) [Koivusalo et al., 1989; Kidd et al., 2007]. The true substrate for FDH is not free FA but the hemithioacetal formed nonenzymically from FA and GSH, namely S-hydroxymethylglutathione. FDH catalyzes the oxidation of S-hydroxymethylglutathione into S-formylglutathione in the presence of NAD [Koivusalo et al., 1989]. Furthermore, this class of FDH-like enzymes has conserved GSNO reductase activity and functions in S-nitrosothiol homeostasis and protects against nitrosative stress in E. coli, Saccharomyces cerivisiae and mouse macrophages [Liu et al., 2001]. In contrast, our microarray data showed that this conserved FDH-like enzyme AdhA of B. subtilis responds specifically to FA and MG but is not induced under conditions of nitrosative stress [Moore et al., 2004]. Since our array data further revealed the induction of the thiol- specific stress response, it was hypothesized that AdhA functions in the thiol-dependent detoxification of aldehydes. In addition to adhA, the microarray data showed high inductions of the yraB and yraC genes, which are encoded upstream of adhA [Huyen et al., 2008b].

The yraC gene encodes an enzyme of 64 aa with similarity to γ- carboxymuconolactone decarboxylase (CMD). CMDs are formed by self-association of identical subunits [Parke, 1979]. These enzymes are involved in protocatechuate catabolism and in some bacteria a gene fusion event leads to expression of CMD with a hydrolase involved in the same pathway [Eulberg et al., 1998]. Recently, it was reported that an enzyme annotated as CMD in Methanosarcina acetivorans had protein disulfide reductase activity, which depends on a CxxC motif [Lessner and Ferry, 2007]. However, this CxxC motif is not conserved in YraC. CMD-like domains are also found in the alkylhydroperoxide reductase AhpD of mycobacteria [Bryk et al., 2002]. It could be possible that YraC specifies a S- formylthiol hydrolase activity generating formate and free thiol [Huyen et al., 2008b].

We identified the novel MerR/NmlR-type regulator YraB (AdhR), which regulates itself and controls positively the adhA-yraA operon and yraC in B. subtilis in response to aldehyde compounds. The upstream sequences of adhA, yraC and adhR contained typical MerR-like promoter sequences with a perfect 7-4-7 inverted repeat (CTTAAAG-N4-

General discussion 114 Thiol-specific stress response

CTTTAAG) between the -35 and -10 elements. In collaboration with the group of John Helmann (Ithaca), we have shown the specific binding activity of AdhR to its target promoters. The MerR family of transcription factors is recognized as a diverse collection of regulators involved in a variety of stress responses, although most investigations of MerR-like regulators have been directed towards those, which respond to heavy metal ions [Brown et al., 2003]. For example, MerR family proteins include regulators for metal ion resistance and efflux proteins such as ZntR and ZccR [Brocklehurst et al., 1999; Kidd and Brown, 2003], CueR [Stoyanov et al., 2001], CadR [Lee et al., 2001] and PbrR [Borremans et al., 2001]. An exception is SoxR, which contains a [2Fe-2S] iron-sulfur cluster and senses superoxide [Pomposiello and Demple, 2001]. Moreover, the AdhR-homologous MerR/NmlR-like regulator of Neisseria gonorrhoeae NmlR was described as redox-controlled regulator, which positively controls itself and three other genes encoding a GSNO reductase AdhC, a Cpx-type ATPase (CopA) and a thioredoxin reductase (TrxB). These enzymes are involved in the response to thiol stress and are required for protection of Neisseria gonorrhoeae against oxidants such as nitric oxide (NO) and OHPs [Kidd et al., 2005]. Such NmlR-like regulators form a distinct subfamily among the MerR family and all regulate conserved the expression of adh-like genes (Tab. 1.1) [Kidd et al., 2005; Stroeher et al., 2007]. Furthermore, these MerR- type regulators have conserved roles in protection against NO stress also in Haemophilus influenzae and Pneumococcus [Kidd et al., 2007; Stroeher et al., 2007].

Tab. 1.1. Examples of the NmlR subfamily of MerR regulators. This table is adapted from Kidd et al. (2005).

Gene Bacterial origin Target Function Cysteines

NMA1517 Neisseria meningitidis adhC GSH-dependent FDH 40, 54, 71, 95 HI0186 Haemophilus influenzae adhC GSH-dependent FDH 54, 71, 95 LP3013 Lactobacillus plantarum LP3012 Oxidoreductase 54, 71 OB0785 Oceanobacillus iheyensis OB0786 NADP-dependent alcohol 54, 71 DH DS2436 Desulfitobacterium hafiniesne eutG Fe-dependent alcohol DH 54, 71 yraB Bacillus subtilis adhA NADP-dependent alcohol 52, 71 DH Lmo1478 Listeria monocytogenes Lmo1477 Oxidoreductase 54, 71

Unlike other MerR-type regulators, NmlR and AdhR lack the 4 conserved Cys residues present in metal-responsive MerR-type regulators (Fig. 1.14). Instead, these NmlR homologues including AdhR all share the conserved Cys52 residues that could play a role in redox-sensing of aldehydes. Interestingly, similar observations were made in previous studies for the FA-responsive transcriptional activator HxlR [Yurimoto et al., 2005]. HxlR binds to its target operator sequences with similar affinities in the absence and presence of FA in vitro. HxlR shares similarity to the YodB repressor and five other MarR/DUF24-family members of B. subtilis. We have shown previously shown that Cys6 of YodB is the sensing cysteine

General discussion 115 Thiol-specific stress response residue that is alkylated by quinone-like electrophiles in vitro [Leelakriangsak et al., 2008]. This conserved Cys residue is also present in HxlR and all YraB-homologs among the MerR/NmlR-subfamily share a conserved Cys52 residue. Mutational analyses in collaboration with the group of John Helmann (Ithaca) showed that activation of adhA-yraA transcription requires the conserved Cys52 of AdhR in vivo. We speculate that redox regulation of AdhR involves thiol-(S)-alkylation of this Cys52 residue by aldehydes and that AdhA and YraA are specifically involved in reduction of aldehydes and repair of alkylated thiol-containing proteins in B. subtilis. Furthermore, since aldehydes react with thiols like quinones via the Michael addition chemistry, it is speculated that specific activation of HxlR and AdhR involves a S-hydroxymethyl-modification at the conserved Cys residues. Thus, the next attempts are directed towards the identification of those thiol-modifications in vivo in cellular proteins as well as in the aldehyde-specific regulator AdhR.

Fig. 1.14. Sequence alignment of NmlR and selected MerR regulators. Identity of sequences is indicated with name and accession number. The conserved cysteines in MerR and related proteins are indicated by mC at the top of the panel while conserved cysteines related to NmlR are indicated by nC at the bottom of the panel. This figure is adapted from Kidd et al. (2005).

In summary, the global expression profile supported that MG and FA act like diamide and quinones as another class of thiol-reactive compounds, which most probably target cysteine residues via the thiol-(S)-alkylation chemistry. Thus, cysteine is also one of the first barriers for aldehydes, which was strongly supported by protection experiments with extracellular cysteine. However, in contrast to quinones and diamide, carbonyl electrophiles triggered a unique bacterial response, which is governed by the HxlR and AdhR regulons that control different detoxification pathways for aldehydes. The adhA-yraA operon confers along with the hxlAB operon resistance to FA indicating that both systems are involved in FA detoxification, the HxlR-regulated RuMP pathway and the AdhR-controlled thiol-dependent FA dehydrogenase pathway.

Nguyen Thi Thu Huyen 116 Mn limitation

Part 2

Genome-wide responses to

manganese limitation in Bacillus subtilis

Supervisor: Prof. Dr. Georg Auling

Summary 117 Mn limitation

SUMMARY

The first part of this thesis was aimed to study the growth, morphology and gene expression profile of B. subtilis in response to Mn2+ limitation. The morphology of B. subtilis changed from the normal rod form to longer filaments upon Mn2+ limitation in the presence of 0.1 µM Mn2+. In addition, asymmetric cell division was observed under Mn2+ limitation conditions. The adaptation of B. subtilis to Mn2+ limitation was studied in detail using transcriptome and proteome analyses. Under Mn2+ limitation condition, both approaches revealed the upregulation of the MntR, PerR, CcpN, AhrC, PyrR, CymR, DegSU, SOS, Spo0A and ComK regulons. In addition, these analyses showed also the downregulation of the Fur, CcpA, IolR, PhoPR, TnrA, CodY, PurR, σB, σM, σW, YvrGHb, YycFG, BceRS, σY, ResDE, Rok, AbrB, σH, Spo0A and σE regulons. The highest derepression was observed for the PerR-regulated katA gene indicating an oxidative stress response of B. subtilis under our conditions of Mn2+ limitation. The filamentation of Mn2+ limited B. subtilis cells might indicate a blocked of cell division. This inhibition of cell division might be caused by the observed downregulation of the σH dependent ftsZ and ftsA genes. Furthermore, Mn2+ limitation inhibited sporulation and induced competence development which was indicated by the downregulation of the σH, σE regulons and genes positively regulated by Spo0A, and the upregulation of ComK regulon and genes negatively regulated by Spo0A, respectively.

Introduction 118 Mn limitation

INTRODUCTION

Manganese (Mn) is the third most abundant transition metal in Earth’s crust [Frausto da Silva and Williams, 1991]. The name manganese is appropriately derived from the Greek word mangania, which means magic. However, little was known about the cellular roles of this essential trace element [Jakubovics and Jenkinson, 2001]. Mn2+ is required for the growth and survival of almost all living organisms. In plants, a Mn2+ containing cofactor is at the heart of the oxygen-evolving complex [Dismukes et al., 2001] and in mammals, a Mn2+ dependent superoxide dismutase (MnSOD) helps to protect mitochondria from the byproducts of aerobic metabolism [Weisiger and Fridovich, 1973]. In bacteria, Mn2+ also performs these functions and it plays a significant role in the virulence of several pathogens [Jakubovics and Jenkinson, 2001; Kehres and Maguire, 2003; Zaharik and Finlay, 2004]. Mn2+ participates in several vital metabolic processes. The list of enzymes that rely on Mn2+ for activity is numerous. Mn2+ serves as a cofactor for numerous oxidoreductases, transferases and hydrolases [Christianson, 1997; Yocum and Pecoraro, 1999; Crowley et al., 2000]. In addition, Mn2+ dependent enzymes are involved in nitrogen metabolism, hydrolysis of water in photosystem II and the modulation of some signal transduction pathways. Mn2+ also protects cells from oxidative stress, either as a cofactor for Mn2+ dependent catalase (MnCAT) and MnSOD or via its inherent ability to quench free radical-mediated reactions [Coassin et al., 1992; Inaoka et al., 1999]. At next, current studies about the role of Mn2+ in the physiology of bacteria will be introduced.

1. Manganese (Mn) transport

Despite its contributions to cellular chemistry, Mn2+ becomes toxic at elevated concentrations [Que and Helmann, 2000; Guedon et al., 2003; Kehres and Maguire, 2003]. Thus, the health of the organism requires both the ability to acquire Mn2+ from the environment and the capacity to block uptake when cellular concentrations become excessive. Bacteria maintain Mn2+ homeostasis by tightly regulated expression of Mn2+ transport systems [Jakubovics and Jenkinson, 2001].

The molecular mechanisms of Mn2+ uptake were unknown until Bartsevich and Pakrasi (1995) discovered the mntCAB operon encoding an ATP-dependent transporter in Synechocystis sp. PCC 6803, a cyanobacterium with a high Mn2+ requirement for photosynthesis. Then, ABC transporters for Mn2+ have been identified in Streptococcus pneumoniae, Streptococcus gordonii and several other bacteria [Dintilhac et al., 1997; Kolenbrander et al., 1998]. These transport systems are sufficiently structurally conserved within all the currently sequenced bacterial genomes. At least, three types of Mn2+ import systems have been identified. The unusual transporter from Lactobacillus plantarum is a P- type ATPase (MntA) with high specificity for Mn2+ [Hao et al., 1999]. The other two types of

Introduction 119 Mn limitation

Mn2+ uptake systems belong either to the natural resistance-associated macrophage protein (NRAMP) family or to the ATP-binding cassette (ABC) transporter superfamily.

In Bacillus subtilis, Mn2+ is required as an enzyme cofactor and as a structural component of protein. The cell membrane prevents Mn2+ from passively entering the cytosol. This protective barrier must be passed to allow the cell to meet its Mn2+ requirements. Under Mn2+ limited condition, Mn2+ is internalized by an active transport system (Fig. 2.1). Conversely, when Mn2+ accumulates to excess, systems for Mn2+ sequestration or efflux are induced [Moore and Helmann, 2005]. As shown in Fig. 2.1, Mn2+ uptake is performed by both NRAMP (MntH) and ABC-transporter systems (MntABCD). MntH is a proton dependent Mn2+ importer. In contrast, Mn2+ uptake by the MntABCD system is likely to be driven by ATP hydrolysis [Que and Helmann, 2000].

Fig. 2.1. Schematic diagram of known and putative metal homeostasis systems in B. subtilis. Metal ion uptake (left side, blue) and efflux systems (right side, green) are illustrated. For information on the regulators controlling the Mn2+, Fe2+ and Zn2+ transport systems, see section “Role of Mn in gene regulation”. This figure is adapted from Moore and Helmann (2005).

2. Cellular functions of Mn

Mn2+ metalloenzymes are involved in diverse functions within bacterial cells (Tab. 2.1) [Christianson, 1997; Yocum and Pecoraro, 1999]. Mn2+ and other cations may be interchangeable in the metal-binding sites of many proteins. Most commonly, Mn2+ and Mg2+ are interchangeable because of the similarities between chelate structures of these ions. However, Mn2+ is essential for certain metabolic pathways. For example, oxygenic

Introduction 120 Mn limitation photosynthesis in cyanobacteria requires a tetra Mn2+ cluster present in the reaction centre complex of photosystem II [Yocum and Pecoraro, 1999]. A number of enzymes, such as adenylyl cyclase [Reddy et al., 2001], arginase, superoxide dismutase (SOD), catalase (CAT) [Christianson, 1997], polysaccharide polymerase [Cartee et al., 2000] and protein kinase [Chaba et al., 2002], specifically require Mn2+ for activity. The reduction of ribose to deoxyribose in nucleotides is catalyzed by the Mn2+ dependent ribonucleotide reductase in Gram-positive bacteria [Willing et al., 1988]. Enzymes involved in sugar catabolism are Mn2+ dependent or have Mn2+ dependent isoforms [Dismukes, 1996], including L-fucose isomerase [Seemann and Schulz, 1997] and cellobiose-6-phosphate hydrolase CelF of Escherichia coli [Thompson et al., 1999] and the D-glucosaminate dehydratase of Pseudomonas fluorescens [Iwamoto et al., 1995]. Furthermore, it appears that Mn2+ has an important role in bacterial signal transduction. The E. coli proteins PrpA and PrpB belong to a family of Mn2+ containing serine/threonine protein phosphatases that are present widely in eukaryotes and modulate complex signaling pathways [Barford, 1996]. The biological importance of Mn2+ is not restricted to enzyme-mediated catalysis. For example, Mn2+ can detoxify a variety of reactive oxygen species (ROS). Additionally, non-enzymic Mn2+ is crucial for the proper function of a variety of bacterial products, including secreted antibiotics [Archibald, 1986], and contributes to the stabilization of bacterial cell walls [Doyle, 1989].

Tab. 2.1. Selected functions of Mn (modified from Jakubovics and Jenkinson, 2001, and Kehres and Maguire, 2003).

Process or pathway Enzyme or protein Reference(s) Photosynthesis Mn2+-stabilizing protein (PSII-O) Morgan et al., 1998 Phosphoenolpyruvate synthase Chao et al., 1993 Gluconeogenesis Pyruvate carboxylase Mukhopadhyay et al., 1998 Glycolysis 3-phosphoglycerate mutase Chander et al., 1998 Enolase Intermediary metabolism Kessler and Knappe, 1996; Lin, 1996 Pyruvate kinase L-fucose isomerase Seemann and Schulz, 1997 Sugar metabolism 6-phospho-β-glucosidase Thompson et al., 1999 Phospholipid biosynthesis Lipid phosphotransferase Sueyoshi et al., 2002 Polysaccharide biosynthesis Polysaccharide polymerase Cartee et al., 2000 Protein catabolism Protein kinase Chaba et al., 2002 Arginase Sekowska et al., 2000 Amino acid metabolism Glutamine synthetase Abell et al., 1995 Threonine 3-dehydrogenase Chen et al., 1995b Aromatic acid metabolism Muconate cycloisomerase Neidhart et al., 1990 Peptide cleavage Aminopeptidase P Yocum and Pecoraro, 1999 DNA precursor biosyntheis Ribonucleotide reductase Griepenburg et al., 1998 Ribonuclease HII Ohtani et al., 2000 Nucleic acid degradation Endonuclease IV Hosfield et al., 1999 Polyamine biosynthesis Agmatinase Carvajal et al., 1999 Stringent response (p)ppGpp 3´-pyrophosphohydrolase Rao et al., 1998 Regulation Transcriptional regulator of Mn2+ uptake Que and Helmann, 2000; Horsburgh et al., 2002 Modulation of signal pathway Protein phosphatase (prpA, prpB) Barford, 1996 Mn2+ dependent catalase Whittaker et al., 1999 Oxidative stress defense Mn2+ dependent superoxide dismutase Fridovich, 1995

Introduction 121 Mn limitation

2.1. Function of Mn in the oxidative stress response

Oxidative stress can be defined as an excess of ROS that have strong oxidizing potential for cells [Fridovich, 1998]. ROS originate from partial reduction of molecular oxygen (O2) to superoxide (O2 ), hydrogen peroxide (H2O2) and hydroxyl radical (OH·)

[Storz and Imlay, 1999; Imlay, 2003]. The O2 anion is known to attack enzymes with exposed [4Fe–4S] clusters which results in the release of Fe2+ and loss of enzyme activity [Gardner and

Fridovich, 1991a; 1991b]. Furthermore, O2 directly inhibits the synthesis of cysteine and aromatic amino acids [Imlay, 2006]. By spontaneous dismutation or in the course of its enzymic detoxification, O2 is rapidly converted to H2O2. H2O2 is a weak oxidizing agent known to react with cysteinyl-thiols in proteins by formation of disulfide bonds or sulfonic acid derivatives [Imlay, 2003]. H2O2 is able to generate carbonyl groups in lysine, arginine, threonine and proline residues and to oxidize methionine to methionine sulfoxides [Stadtman,

1993]. Most significantly, H2O2 reacts with reduced Fe ions to form OH·, which in turn oxidizes most cellular compounds at diffusion-limited rates [Henle and Linn, 1997].

ROS are very reactive and can cause damage to macromolecular constituents such as DNA, RNA, proteins and lipids [Storz et al., 1990b]. Therefore, during evolution, organisms either became restricted to niches free from oxygen (anaerobes) or acquired protective mechanisms (aerobes). Most aerobic organisms respond to oxidative stress by the inducible synthesis of a variety of protective enzymes and proteins. Enzymes for protection against ROS include SOD, which catalyzes the dismutation of O2 to O2 and H2O2 [Fridovich, 1995], peroxidase and CAT, which prevent the intracellular accumulation of toxic H2O2 [Hassan and Fridovich, 1978] and alkyl hydroperoxide reductase (Ahp) [Bsat et al., 1996]. Fe2+ is linked to 2+ oxidative damage, primarily through the ability of Fe to reduce H2O2 [Touati, 2000]. One strategy for minimizing oxidative damage therefore involves limiting intracellular Fe2+ content. On the other hand, bacteria growing in the absence of Fe2+ appear to have an absolute requirement for Mn2+ [Jakubovics and Jenkinson, 2001].

Bacteria have acquired elaborate defense mechanisms that co-ordinate Fe2+-sensing and Mn2+-sensing with oxidative stress responses [Bsat et al., 1998; Storz and Imlay, 1999]. While Fe2+ is toxic and can promote OH· radical formation, Mn2+ containing compounds react with O2 , H2O2 and OH·, without generating deleterious free radical species [Cheton and Archibald, 1988]. Mn2+ is essential for the detoxification of ROS in most bacteria, principally activates the activity of MnSOD, MnP and MnCAT (see below). Interestingly, Lactobacillus plantarum is essentially devoid of Fe2+ and heme but may accumulate a remarkably high (35 mM) cytoplasmic concentration of Mn2+ [Archibald, 1986]. It is believed that much of this Mn2+ is “free” in a chemical sense and substitutes functionally for Fe2+ and heme in as yet chemically undefined ways. However, no new mechanistic studies on the chemistry of non-

Introduction 122 Mn limitation enzymatic free radical detoxification have been reported, while participation of Mn2+ in other free radical detoxifying processes has received little attention [Kehres and Maguire, 2003].

Manganese dependent superoxide dismutases (MnSODs)

SODs are metalloenzymes that catalyze the disproportionation of O2 to O2 and H2O2. They are present in nearly all organisms from bacteria to humans [McCord, 1976]. However, not all organisms contain the Mn2+ cofactor dismutase [Keele et al., 1970]. Other forms of SODs are known to contain Fe2+ [Yost and Fridovich, 1973], binuclear Cu2+/Zn2+ [Benov et 2+ al., 1995], or even Ni [Youn et al., 1996]. Resistance to the toxic effects of O2 more usually involves the concerted activities of two or more SOD. E. coli possesses three SOD activities: periplasmic Cu/ZnSOD (SodC) and two cytosolic proteins, FeSOD (SodB) and MnSOD (SodA) [Fridovich, 1995]. Disruption of the sodA and sodB genes in E. coli leads to enhanced sensitivity of cells to O2 and generates nutritional auxotrophies resulting from inactivation of dihydroxy-acid dehydratase by O2 [Kuo et al., 1987]. On the other hand, it appears that most streptococci and enterococci [Niven et al., 1999] and B. subtilis [Inaoka et al., 1999] may produce only MnSOD. In B. subtilis, MnSOD encoded by sodA was found to be associated with the abundant spore coat protein CotG [Henriques et al., 1998]. This enzyme is predicted to protect cells from oxidative stress during growth and sporulation [Inaoka et al., 1998].

Manganese dependent peroxidases and catalases (MnPs and MnCATs)

Manganese dependent peroxidases (MnPs) couple the redox activity of H2O2 to degradation of nutrients such as lignin via oxidation of a Mn2+ bound to a propionate side- chain of the heme group. MnPs have thus far only been identified in lignolytic fungi, such as Phanerochaete chrysosporium [Gold et al., 2000].

CATs are widely distributed enzymes that decompose H2O2 to H2O and O2 as a detoxifying mechanism. Three phylogenetically distinct types of CATs have been described [Loewen et al., 2000]. The enzymes are defined as bifunctional and monofunctional CATs, depending on their abilities to function as peroxidases as well. The third group of CATs embraces the nonheme or pseudocatalases containing Mn2+, instead of Fe2+ heme, in the active site, and therefore is called MnCAT. In B. subtilis, one MnCAT, KatX catalase, was found. This enzyme is specifically produced in the spores and may play a role in the protection of germinating spores against H2O2 [Bagyan et al., 1998].

2.2. Role of Mn during B. subtilis sporulation and germination

Early work in the 1950s demonstrated that Mn2+ is unique amongst the transition metals in promoting both endospore formation and spore germination in Bacillus. The

Introduction 123 Mn limitation requirement for Mn2+ is specific and can not be replaced by the addition of sodium (Na), potassium (K), magnesium (Mg), calcium (Ca), strontium (Sr), cobalt (Co), nickel (Ni), cadmium (Cd), zinc (Zn), copper (Cu), stannum (Sn), cerium (Ce) or low concentration of iron (Fe) [Charney et al., 1951]. Mn2+ appears to be important at several stages in the sporulation and germination (Fig. 2.2). In the switch from symmetric to asymmetric cell division, Mn2+ is directly involved as a cofactor of the SpoIIE serine phosphatase [Schroeter et al., 1999]. This is a bifunctional protein that influences polar septum formation through interactions with FtsZ [King et al., 1999] and that separately activates σF in the forespore by dephosphorylation of SpoIIAA [Kroos et al., 1999]. It is possible that σF activation by SpoIIE may be regulated by Mn2+, since Mn2+ accumulates specifically within the developing forespore [Jakubovics and Jenkinson, 2001].

Fig. 2.2. Roles of Mn2+ in the sporulation and germination. Mn2+ homeostasis is essential for efficient initiation of sporulation in the vegetative cell (1). The switch from medial to polar septum formation (2) is influenced by the Mn2+-requiring SpoIIE protein. Following septation, Mn2+ accumulates in the developing forespore. SpoIIE localizes to both poles of the cell, but exclusively activates σF in the forespore. A drop in pH of around 1 unit accompanying forespore development (3) causes dissociation of Mn2+ from 3-phosphoglycerate mutase (3-PGM), inactivating the enzyme (3-PGMi) and leading to accumulation of the storage molecule 3- phosphoglyceric acid (3-PGA) (4). Production of the inner (shaded) and outer (thick line) layers of the spore coat (5) requires MnSOD. Germination of the mature spore (6) involves a rapid increase in pH, leading to the activation of 3-PGM and the mobilization of 3-PGA reserves. Protease production at this stage is stimulated by Mn2+ and high levels of Mn2+ dependent pyrophosphatase (MnPPiase) are associated with an increase in metabolic activity. Chromosomal DNA is represented by wavy lines. This figure is adapted from Jakubovics and Jenkinson (2001).

Mn2+ also influences spore composition, structure and germination. B. subtilis endospores contain a storage reservoir of 3-PGA, which composes 0.1 to 0.3 % of spore dry weight. It is converted to acetate after initiation of spore germination [Paidhungat and Setlow, 2002]. Accumulation of 3-PGA within the developing forespore occurs following acidification of the forespore compartment with concomitant dissociation of the essential Mn2+ cofactor from 3-PGM (Fig. 2.2). Upon initiation of spore germination, 3-PGM is rapidly reactivated following an increase in intracellular pH and 3-PGA degradation [Chander et al., 1998]. When B. subtilis is grown in a Mn2+ deficient medium, a large pool of 3-PGA accumulates in stationary phase cells and sporulation does not take place [Oh and Freese, 1976]. Furthermore,

Introduction 124 Mn limitation during germination, Mn2+ is not only involved in the activation of 3-PGM but also the activation of proteases and inorganic pyrophosphatase [Kuhn and Ward, 1998].

2.3. Role of Mn in gene regulation

B. subtilis contains at least four distinct metal ion dependent repressors. These are the Mn2+ dependent repressor (MntR, manganese transport regulator), controlling the regulon involved in Mn2+ uptake [Que and Helmann, 2000], and three Fur-like proteins that control regulons involved in peroxide stress (PerR, peroxide stress response repressor), Fe2+ uptake (Fur, ferric uptake regulator) and Zn2+ transport (Zur, zinc uptake repressor) [Chen et al., 1995a; Bsat et al., 1998; Gaballa and Helmann, 1998]. All four proteins require a bound divalent metal ion in order to bind DNA: MntR responds to Mn2+ and Cd2+ [Que and Helmann, 2000; Kliegman et al., 2006], PerR responds to either Fe2+ or Mn2+, Fur responds to Fe2+ and Zur responds to Zn2+ [Herbig and Helmann, 2002].

2.3.1. MntR as regulator of Mn homeostasis

Mn2+ is accumulated from the environment via a high-affinity transport system [Silver and Walderhaug, 1992]. Regulation of Mn2+ transport requires the activity of MntR in B. subtilis [Herbig and Helmann, 2002]. MntR is responsible for Mn2+ homeostasis and is a distant homologue of the diphtheriae toxin repressor (DtxR) [Que and Helmann, 2000; Golynskiy et al., 2006]. DtxR proteins are divalent metal ion dependent transcriptional repressors [Tao et al., 1992]. DtxR contains three domains: an amino-terminal DNA-binding domain, a central metal-binding domain (dimerization domain) and a carboxyl-terminal SH3- like domain [Herbig and Helmann, 2002; Golynskiy et al., 2005]. The SH3 domain in DtxR may regulate repressor activity [Wang et al., 1999] and contribute to additional metal binding [Pohl et al., 1998]. A comparison of MntR with the DtxR family leads to three observations (Fig. 2.3): (a) the DNA-binding recognition helix is not highly conserved, suggesting that DNA recognition may vary between these families; (b) most ligands for metal binding are conserved, suggesting that the position of the metal binding sites is conserved (although not, apparently, the metal selectivity); and (c) the third SH3-like domain is absent from MntR. Another significant difference is that the two metal ions in MntR form a bridged, dinuclear metal active site, which is distinct from the two mononuclear sites found in DtxR [Pohl et al., 1998; Kliegman et al., 2006]. In addition, DtxR is a Fe2+ responsive repressor [Boyd et al., 1990], while MntR is responsive to Mn2+ and Cd2+, both in vivo and in vitro [Que and Helmann, 2000; Kliegman et al., 2006].

Introduction 125 Mn limitation

Fig. 2.3. Ribbon representation of the crystal structures of holo MntR (left) and DtxR (right). Proteins are colored to highlight domains and function: helix-turn-helix DNA binding domain (brown), dimerization domain (red), linking α-helix (green), additional dimerization α-helix (purple, MntR only), and SH3-like domain (gray, DtxR only). Mn2+ and Co2+ ions are shown as spheres in the binding sites of MntR and DtxR, respectively. This figure is adapted from Golynskiy et al. (2005).

MntR is involved in Mn2+ homeostasis through regulation of two Mn2+ transporters, MntABCD and MntH [Que and Helmann, 2000]. When the Mn2+ requirements of the cell are sufficient, MntR binds Mn2+, resulting in DNA binding and repression of the mnt operons (Fig. 2.4). After drop of the Mn2+ levels below those required for sufficiency, the metal ions dissociate from MntR, and then MntR is released from DNA resulting in mnt operon transcription [Golynskiy et al., 2006].

Fig. 2.4. Metal mediated mechanism of MntR. Dimerization domains of MntR are shown as blue ovals and DNA-binding domains as red triangles. Dotted lines are used to represent motions of MntR in solution. Spheres are represented as Mn2+ ions. This figure is adapted from Golynskiy et al. (2007).

2.3.2. Fur-like metalloregulators

The Fur regulator has been best characterized from E. coli (designated FurEC). FurEC is a metalloregulator that binds to conserved operator sequences (Fur boxes) of Fe2+ uptake and 2+ siderophore production genes using Fe as a co-repressor [Escolar et al., 1999]. FurEC has long been implicated as both a positive and negative regulator of gene expression [Lee and 2+ Helmann, 2007]. FurEC is presumed to contain a regulatory site for sensing Fe . In addition, 2+ FurEC contains a tightly bound structural Zn ion that is required for proper protein folding 2+ [Althaus et al., 1999]. In vivo, FurEC represses a large regulon of Fe uptake functions when 2+ Fe is present in excess [Braun, 1997]. FurEC is also the prototype for a large family of

Introduction 126 Mn limitation regulators that sense Fe2+ (Fur), Zn2+ (Zur), Mn2+ (Mur) and peroxide stress (PerR) in E. coli [Hantke, 2001].

In B. subtilis, the Fur-like metalloregulatory proteins include several small, dimeric, DNA-binding proteins that respond to metal ions including Fur itself (Fe2+ sensor), Zur (Zn2+ sensor) and PerR (peroxide sensor) [Bsat et al., 1998; Gaballa and Helmann, 1998; Escolar et al., 1999; Lee and Helmann, 2007]. Like FurEC, these three proteins are composed of two domains. The amino-terminal domain contains a helix-turn-helix motif with an unusual turn, and the carboxyl-terminal domain contains two metal-binding sites and is important for dimerization [Althaus et al., 1999]. Thus, the three Fur-like metalloregulatory proteins (Fur, Zur and PerR) are dimeric DNA-binding repressors, which contain two metal ions per monomer [Herbig and Helmann, 2001; 2002]. One of the metal binding sites co-ordinates a Zn2+ ion that is generally assigned a structural role, whereas the second site binds a regulatory metal (Fe2+ for Fur, Zn2+ for Zur and either Fe 2+or Mn2+ for PerR) required for DNA binding.

Fur functions primarily as a Fe2+ activated repressor of Fe2+ homeostasis controlling both the induction of Fe2+ uptake functions (under Fe2+ limitation), and the expression of Fe2+ storage proteins and Fe2+ utilizing enzymes (under Fe2+ sufficiency) [Baichoo et al., 2002; Lee and Helmann, 2007]. Fur is part of the PerR regulon and not autoregulated [Fuangthong et al., 2002]. Therefore, the fur gene is repressed only by Mn2+ [Chen et al., 1995a; Bsat et al., 1996; 1998; Baichoo et al., 2002]. Fur represses at least twenty operons encoding about forty genes implicated in either siderophore synthesis or Fe2+ uptake [Baichoo and Helmann, 2002; Baichoo et al., 2002]. Fur also regulates positively an iron-sparing response in B. subtilis, which is mediated by a Fur-regulated small RNA (FsrA) and three small basic proteins that down-regulate expression of Fe2+-containing proteins (succinate dehydrogenase or aconitase) under Fe2+ limiting conditions [Gaballa et al., 2008]. It is generally accepted that Fur binds to Fe2+, and thereby, is activated to bind DNA, leading to the repression of Fe2+ uptake functions [Bsat and Helmann, 1999]. In addition, in vitro, numerous other divalent cations can also activate the Fur protein to bind DNA. Among these cations, Mn2+ is a functional co-repressor of the Fur protein [Herbig and Helmann, 2002]. Elevated Mn2+ levels appear to repress the Fur regulon in vivo [Guedon et al., 2003]. However, the same authors proposed alternatively that elevated levels of Mn2+ perturb Fe2+ homeostasis by increasing availability of iron within the cell through displacing iron from a kinetically labile, loosely chelated pool. The nature and location of this pool are not clear. Similarly, recent studies have revealed that Synechocystis sp. PCC6803 accumulates large amounts of Mn2+ in a labile pool associated with the outer membrane [Keren et al., 2002].

In B. subtilis, a Zn2+-sensing Fur paralog, Zur, represses the Zn2+ uptake system in response to Zn2+ sufficiency [Gaballa et al., 2002; Moore and Helmann, 2005]. The Zn2+ uptake system consists of the Zn2+ uptake ABC transporters YcdHI-YceA and a postulated

Introduction 127 Mn limitation low-affinity transport system YciABC [Gaballa et al., 2002; Gaballa and Helmann, 2002]. In addition, the P-type ATPase ZosA is also involved in Zn2+ uptake. However, the zosA gene is not repressed by Zn2+ and is regulated by the peroxide-sensing repressor PerR (see below) [Gaballa and Helmann, 2002]. On the other hand, Zur represses several genes not related to Zn2+ uptake but encoding homologues of ribosomal proteins (rpmGC, yhzA and ytiA) and finally a predicted Zn2+ binding membrane protein ZinT [Panina et al., 2003]. In order to regulate Zn2+ uptake genes, Zur uses Zn2+ as a co-repressor. Zur may contain two Zn2+ sites-a structural site and a low affinity regulatory site [Baichoo and Helmann, 2002; Herbig and Helmann, 2002]. Both Zn2+ and Mn2+ can activate DNA binding in vitro [Gaballa and Helmann, 1998].

PerR is the major regulator of the peroxide stress response [Bsat et al., 1998]. The PerR regulon includes a DNA-binding protein (MrgA) [Chen and Helmann, 1995], the major vegetative catalase (KatA) [Chen et al., 1995a], alkyl hydroperoxidase (AhpCF) [Bsat et al., 1996], enzymes of heme biosynthesis (HemAXCDBL) [Chen et al., 1995a], a Zn2+ uptake P- type ATPase (ZosA) [Gaballa and Helmann, 2002] and both the PerR and Fur metalloregulatory proteins [Fuangthong et al., 2002]. Most PerR-regulated genes are 2+ 2+ derepressed in cells treated with low level of H2O2 or starved for either Fe or Mn . The different genes of the PerR regulon have a divergent metalloregulation. Whereas PerR- mediated repression of most target genes can be elicited by either Mn2+ or Fe2+, repression of perR and fur occurs selectively by Mn2+ [Fuangthong et al., 2002].

In conclusion, Mn2+ affects three of four regulons, namely MntR, Zur, and PerR, directly by its ability to activate DNA binding of their metalloregulatory proteins [Gaballa and Helmann, 1998; Que and Helmann, 2000; Herbig and Helmann, 2001; Kliegman et al., 2006]. The effect of Mn2+ on the Fur regulon is likely to be an indirect effect of Mn2+ via perturbation of Fe2+ homeostasis [Guedon et al., 2003]. Furthermore, since MntR, Fur and PerR respond to an overlapping set of metal ions in vivo, these regulons are intimately interconnected (Fig. 2.5).

Introduction 128 Mn limitation

Fig. 2.5. Schematic diagram illustrating three interconnected metalloregulatory circuits in B. subtilis. PerR is proposed to exist as an inactive, Zn2+ metalloprotein that can be activated for DNA-binding by either Fe2+ or Mn2+ to repress peroxide stress genes. The intracellular levels of Mn2+ and Fe2+ are regulated by the MntR [Que and Helmann, 2000] and Fur [Bsat et al., 1998; Bsat and Helmann, 1999] metalloregulatory proteins, respectively. This figure is adapted from Herbig and Helmann (2001).

3. The aim of this part of the thesis

The Mn2+ requirement of B. subtilis for growth with glucose and for sporulation is well known and has been in the focus of research for a long time. Growth on glycolytic substrates depends on the availability of sufficient intracellular Mn2+ for proper functioning of 3-PGM. This enzyme needs 10 mM Mn2+ in vitro to avoid inactivation [Watabe and Freese, 1979]. However, B. subtilis requires Mn2+ also for non-glycolytic growth. Using fumarate or pyruvate as a substrate, Mohamed et al. (1998) demonstrated through in vivo labeling with nucleic acid precursors a reversible impairment of DNA formation in the absence of Mn2+. Ribonucleotide reduction was also impaired under this condition. A peculiarity is that the Mn2+ depleted cultures finally reached roughly the same turbidity after an initial lag. In summary, the effects of Mn2+ limitation in B. subtilis recall phenomena of unbalanced growth previously reported for corynebacteria [Auling and Follmann, 1994].

Although further evidence for a Mn2+ ribonucleotide reductase in B. subtilis was obtained by Q-band EPR-spectroscopy [Mohamed et al.,1998], the factors contributing to unbalanced growth in this bacterium may be more complex since many enzymes require Mn2+ for proper function [Archibald, 1986]. Using proteome and transcriptome analysis, genome- wide expression profiling of B. subtilis in response to Mn2+ limitation were one goal of this thesis. Understanding of gene regulation of Mn2+ limited cultures was expected in addition to further insight into basic functions of Mn2+ homeostasis and the up-coming protective role of Mn2+.

Materials and Methods 129 Mn limitation

MATERIALS AND METHODS

1. Bacterial strain and medium

Bacterial strain used in this study is B. subtilis 168. The pyruvate minimal (PM) medium was used. The PM medium contains 10 g/l KH2PO4, 10 g/l K2HPO4 x 3H2O, 2 g/l

(NH4)2SO4, 0.5 g/l MgSO4x7H2O, 1 g/l KCl, 2 g/l sodium citrate x 2H2O and 10 g/l sodium pyruvate. Before cultivation, 40 ml of a stock solution of trace elements (500 mg/l EDTA, 200 mg/l FeSO4 x 7H2O, 10 mg/l ZnSO4 x 7H2O, 30 mg H3BO3, 20 mg/l CoCl2 x 6H2O, 1 mg/l

CuCl2 x 2H2O, 2 mg/l NiCl2 x 6H2O and 3 mg/l Na2MoO4 x 2H2O), 9 ml of glutamate stock solution (0.5 M, pH 7) and 2 ml of tryptophan stock solution (0.039 M) were added per liter 2+ PM medium. Mn stock solution (1 mM MnCl2) was added into PM medium depended on the final desired concentration in each experiment.

Only water purified by reversed osmosis (3 modules E-pure D4632240 VAC Barnstead, Dubuque, IA, USA) was used to reduce the level of adventitious Mn2+ contamination.

2. Growth conditions

The effect of different Mn2+ concentrations on growth of B. subtilis 168 was studied in 100 ml PM medium at 37oC, 120 rpm in 500 ml Erlenmeyer flasks throughout using mid-log pre-cultures supplemented with 2 µM Mn2+. Cell growth was followed by measuring the optical density (OD) at 578 nm.

The morphology of B. subtilis 168 under Mn2+ limitation (0.1 µM Mn2+) versus sufficiency (2 µM Mn2+) as a control was studied in 400 ml modified PM medium in 2 liter Erlenmeyer flasks using mid-log inocula from PM medium supplemented with 2 µM Mn2+ and incubating at 37oC, 120 rpm.

For proteome studies by two-dimensional polyacrylamide gel electrophoresis (2- D PAGE) analysis and transcriptome studies, B. subtilis 168 was incubated at 37oC and 180 rpm in 1 liter of PM medium supplemented with 2 µM or 0.1 µM Mn2+ using 5 liter Erlenmeyer flasks. Mid-log pre-cultures supplemented with 2 µM served for inoculation. Cell samples were withdrawn as indicated in Fig. 2.6b.

3. Scanning electron microscopy (SEM)

Cell samples from control and Mn2+ limited cultures were taken at different time points along the growth curve, harvested by membrane filtration. After fixation of samples with 2 % glutaraldehyde in 0.01 M dimethylamine buffer, the samples were dehydrated by gradually increasing the ethanol concentration and by subsequent critical point drying (Balzers,

Materials and Methods 130 Mn limitation

Wiesbaden, Germany). The fixed samples were sputter-coated with gold. Microphotographs were taken at 15 kV with a Zeiss DSM 940 scanning electron microscope (Zeiss, Oberkochen, Germany).

4. 2-D PAGE

Duplicates of 2-D PAGE experiments were performed for each protein preparation from two independent cultivations under the control or Mn2+ limitation condition.

4.1. Protein extraction

Cells grown under the control and Mn2+ limitation, which were taken at different time points along the growth curve, were harvested by centrifugation (8000 rpm, 10 min and 4oC) and washed twice with TE buffer (10 mM Tris, 1 mM EDTA, pH 7.5). Cells were re- suspended in TE buffer and disrupted by a twofold passage through a French press (Minicell, Rochester, NY, USA) at 8000 psi and the cell debris was removed by centrifugation (12000 rpm, 10 min and 4oC). After having cleared the protein extract by an additional centrifugation (15000 rpm, 30 min and 4oC), the protein concentration of supernatant was determined with a commercially available kit (RotiNanoquant) according to Bradford [Bradford, 1976]. The protein solution could be stored at -20oC until further analysis.

4.2. 2-D PAGE and quantitative image analysis

Isoelectric focusing (IEF) was performed with commercially available immobilized pH gradient (IPG) strips in the pH range 4-7 (GE Healthcare, Freiburg, Germany). IPG strips were loaded with 200 µg of crude protein extract by rehydration for 24h, at room temperature in a solution containing 8 M urea, 2 M thiourea, 1 % v/v NP 40, 20 mM DTT and 0.5 % v/v Pharmalyte 3-10. The IEF was performed with the Multiphor II unit (Amersham Pharmacia Biotech) employing the following voltage profile: linear increase from 0 to 500 V for 500 Vh, 500 V for 2500 Vh, linear increase from 500V to 3500 V for 10000 Vh and final phase of 35000 V for 35000 Vh. After consecutive equilibration of the IPG strips in solutions containing 0.35 % dithiothreitol (DTT) and then 4.5 % iodoacetamide (IAA), the separation in the second dimension was done in polyacrylamide gels of 12.5 % acrylamide on the Investigator 2-D electrophoresis system (Genomic Solutions, Chelmsford, MA, USA) with approximately 2 W per gel at 12oC. The resulting 2-D gels were fixed with 40 % v/v ethanol and 10 % v/v acetic acid for 1-2 h and subsequently stained with colloidal CBB [Neuhoff et al., 1988]. The gels were scanned with a HP Scanjet Scanner.

Quantitative image analysis was performed with the Decodon Delta2D software (http://www.decodon.com), which is based on the dual channel image analysis technique pioneered by Bernhardt et al. (1999). Using this software, the 2-D gel images from Mn2+ limitation experiments were aligned to reference images (the control images) by using a warp

Materials and Methods 131 Mn limitation transformation according to the culture harvesting points. To avoid incomplete groups of matching spots, a fused 2-D gel was created for spot detection. For preparing a fusion gel, all 2-D gel images from control and Mn2+ limitation samples from each harvesting point were combined. Spot detection was performed in the fusion gel containing all spots present in any gels of the control and the Mn2+ limitation samples from each harvesting point. The resulting spot shapes were reviewed and manually edited in the fusion gel if necessary. The reviewed spot mask served as a spot detection consensus for all gel images of the control and Mn2+ limitation and was applied to the individual gels to guide the spot detection and quantification. This enables spot quantification in all gels at the same locations resulting in 100 % matching and in a reliable analysis of complete expression profiles. Normalization was performed by calculating the quantity of each single spot in percentage related to the total spot quantity per gel. The values of protein abundance of the Mn2+ limitation sample were divided by the values of the protein abundance of the control sample for determination of protein induction or protein repression. Spots showing an induction of at least two-fold under Mn2+ limitation were designated as significantly induced proteins. The identification of the induced proteins was performed by comparing with the cytosolic protein map of B. subtilis [Buttner et al., 2001; Eymann et al., 2004] or via MALDI-TOF- MS according to Eymann et al. (2004).

5. DNA microarray

Cells grown under the control and Mn2+ limitation, which were taken at different time points along the growth curve, were harvested in half of the sample volume ice killing buffer

(20 mM Tris-HCl, pH 7.5, 5 mM MgCl2, 20 mM NaN3) and then centrifuged for 3 min, 8000 o rpm and 4 C. The supernatants were discarded and the pellets were frozen in liquid N2 and stored at -80oC until RNA preparation. The total RNA isolation was performed according to Eymann et al., 2002. The generation of fluorescence-labeled cDNA (Cy3-labled for control samples and Cy5-labled for Mn2+ limitation samples) and hybridization with B. subtilis whole-genome microarrays (Eurogentec) were performed according to Jürgen et al. (2005). The fluorescence intensity of two dyes were detected by the ScanArray Express scanner (PerkinElmer Life and Analytical Sciences, Rodgau-Jügesheim, Germany) using the ScanArray Express image analysis software. The data analysis was performed via the GeneSpring software (Version 7.3 Agilent Technologies, Waldbronn, Germany). After background subtraction, signal intensity was transformed by dependent LOWESS normalization according to the instruction of the manufacturer. The values of the Cy5-labeled samples (Mn2+ limitation) were divided by the normalized values of the Cy3-labeled samples (control) for determination of gene induction or gene repression. Only genes, which were at least two-fold induced (ratios above 2) or two-fold repressed (ratios below 0.5) were considered as significantly induced or repressed under Mn2+ limitation, respectively. Duplicates of DNA microarray experiments were performed for each RNA preparation from two independent cultivations in each condition.

Results and discussion 132 Mn limitation

RESULTS AND DISCUSSION

1. The impact of Mn on growth behavior

Firstly, the effect of difference concentration of Mn2+ on B. subtilis growth was examined. B. subtilis was grown in PM medium supplemented with 0.05-5µM Mn2+. B. subtilis grew well at 2, 2.5 and 5 µM Mn2+ and growth was limited with 0.05 and 0.1 µM Mn2+ (Fig. 2.6a).

3 10 10 2 a b

1 1 1 3 578nm 500nm 2 OD 0.1 OD 0.1 1

0.01 0.01 0123456789101112 0 1 2 3 4 5 6 7 8 9 10 11 12 13 14 time (h) time (h)

Fig. 2.6. (a) Growth of B. subtilis in PM medium with different Mn2+ concentrations (full circle-5 µM, full square-2.5 µM, full triangle-2 µM, empty circle-without Mn2+, empty square-0.05 µM, empty triangle-0.1 µM). (b) Growth of B. subtilis under control and Mn2+ limited conditions for the proteome (the cultures were taken at the time points 1, 2, 3) and transcriptome (the cultures were taken at the time points 1, 2) analyses (full square-the control, empty square-Mn2+ limitation, arrow- sampling point).

2. Morphological, proteomic and transcriptomic studies of B. subtilis in response to Mn limitation

2.1. Morphology of B. subtilis in response to Mn limitation

As shown in Fig. 2.6a, the growth of B. subtilis was limited with 0.1 µM Mn2+ which was chosen as Mn2+ limitation condition. In addition, 2 µM was chosen as condition that provides sufficiently Mn2+ supply as a control to Mn2+ limited condition. The experiment was designed as follows: B. subtilis 168 was grown with 2 µM Mn2+ medium and the exponential phase culture was transferred either to 2 µM Mn2+ or 0.1 µM Mn2+. Firstly, the change of cell morphology under Mn2+ limitation was observed by SEM (Fig. 2.7). The resulting microphotographs showed that limitation of Mn2+ caused an alteration of cell morphology from the normal rod form of control cultures (Fig. 2.7a, c, e) to longer filaments (Fig. 2.7b, d, f). Under Mn2+ limitation, cells with extreme length up to 15.3 µm (figure not shown) were observed. In addition, asymmetric cell division was noticed in the stationary phase under Mn2+ limitation (Fig. 2.7f). On the basis of these data, two global experimental approaches (proteome and transcriptome analyses) were applied to reveal the changes of gene expression when B. subtilis adapts to Mn2+ limitation.

Results and discussion 133 Mn limitation

Fig. 2.7. Morphology of B. subtilis under control condition (left side) and Mn2+ limitation (right side) from cultures taken from the exponential phase (a and b), transition phase (c and d) and stationary phase (e and f). Cells with asymmetric division are indicated by red arrows.

Results and discussion 134 Mn limitation

2.2. Proteome and transcriptome analyses of B. subtilis in response to Mn limitation

Proteome and transcriptome analyses were performed using samples taken at different time points along the growth curve under control and Mn2+ limitation conditions (Fig. 2.6b). These analyses were performed two times for each sample from two independent cultivations.

Using 2-D PAGE analysis, twenty-five proteins were significantly induced under Mn2+ limitation (Tab. 2.2). Six, fifteen and twenty proteins were significantly induced after 5 h cultivation, 9 h cultivation and 12 h cultivation assigned to the exponential (Fig. 2.8), transition (Fig. 2.9) and stationary phase (Fig. 2.10), respectively. In addition, under Mn2+ limitation, 51 and 181 genes were upregulated (Tab. 2.3) whereas 104 and 282 genes were downregulated (Tab. 2.4) in the exponential and transition phase, respectively.

Most importantly, the MntR, PerR and Zur regulons [Que and Helmann, 2000; Herbig and Helmann, 2002; Gaballa and Helmann, 2002] were induced by Mn2+ limitation. In B. subtilis, the mntH and mntABCD genes are regulated by MntR and involved in the pathways for Mn2+ uptake (Tab. 2.3). The transcriptome analysis revealed that all five MntR regulated genes (mntABCD and mntH, Mn2+ transport) belonging to two operons were induced by Mn2+ limitation. However, we noticed that induction of the MntR regulon was only at the level of 2- 3 fold, which indicates that the medium and conditions used for Mn2+ limitation were not optimal in our experiments. Moreover, the transcriptome results revealed a high expression level of mntH and lower induction of mntABCD under Mn2+ limitation (Tab. 2.3). This is in agreement to data of Que and Helmann (2000), which showed that MntH and MntABCD are high- and low-affinity systems for Mn2+ uptake. In addition, the transcriptome analysis also showed that mntH was induced more in the exponential phase whereas the mntABCD operon was slightly more induced during the transition phase. According to the report of Eisenstadt et al. (1973), Mn2+ uptake is activated during exponential growth phase, decreased upon stationary phase and increased again early during sporulation. Thus, MntH may be the predominant transport activity during growth, while the ATP-dependent MntABCD system may play the major role in Mn2+ uptake during sporulation.

In B. subtilis, Mn2+ is an effective co-repressor for the metalloregulatory proteins MntR and PerR [Que and Helmann, 2000; Herbig and Helmann, 2001; 2002]. Mn2+ mediates the regulation of PerR [Herbig and Helmann, 2001; 2002] via binding to a regulatory site required for DNA binding. PerR is the major regulator of the inducible peroxide stress response [Bsat et al., 1998]. The DNA microarray analysis revealed that all genes of the PerR regulon [Herbig and Helmann, 2002] were induced under Mn2+ limitation. Of thirteen PerR regulated genes (ahpCF, fur, hemAXCDBL, katA, mrgA, perR and zosA), the highest induction was observed with the katA gene. Similarly, ahpCF and mrgA were strongly induced in the exponential and transition phase. These indicate that cells mainly suffer from oxidative stress

Results and discussion 135 Mn limitation in our Mn2+ limitation experiments that confirms largely the proteome results in which the PerR-regulated proteins KatA, AhpC and AhpF were strongly induced during all growth phases and the highest level of the induction was noticed for KatA. Furthermore, the induction of the regulatory gene fur (ferric uptake regulator) was higher during the exponential growth than in the transition phase. Three genes (hemB, hemC, hemD) involved in heme biosynthesis were induced in the exponential growth and all heme biosynthetic genes (hemLBDCXA) were induced in the transition phase. The Zn2+ uptake gene zosA (PerR regulon) was highly induced only in the exponential phase. These results are in accordance with reports by Herbig and Helmann (2001) and Guedon et al. (2003) that elevated levels of Mn2+ repressed the PerR regulon genes. Thus, Mn2+ mediates regulation of the PerR regulon as suggested by Herbig and Helmann (2001, 2002) and Fuangthong et al. (2002). The transcriptome analysis also showed that the Fur regulon genes dhbACEBF, yclP, yclQ, ydbN, yfkM, ykuNOP, yuiI, and yxeB [Baichoo et al., 2002; Ollinger et al., 2006] were downregulated in response to Mn2+ limitation. In contrast, the Fur regulon was also downregulated by elevated Mn2+ concentrations as published by Guedon et al. (2003). It was suggested that elevated Mn2+ displaces Fe2+ from cellular-binding sites and resulting rise in free Fe2+ levels leads to downregulation of the Fur regulon. Since fur is a member of the PerR regulon, the derepression of fur by Mn2+ limitation could result in the enhanced transcriptional repression of the Fur regulon in our studies.

The induction of the Zur regulated Zn2+ transporter-encoding ycdH gene by Mn2+ limitation was unexpected [Gaballa and Helmann, 1998]. This could indicate a role of YcdH for Mn2+ uptake. Interestingly, Bunai et al. (2004) speculated on a possible function of YcdH in uptake of Mn2+ in addition to Zn2+. Furthermore, Mn2+ limitation also induced the PerR regulon member zosA encoding a third Zn2+ uptake system. Thus, Mn2+ limitation has an impact on two Zn2+ transporters by different regulatory mechanisms. The high induction of zosA in the exponential phase and the slight induction of ycdH in the transition phase might suggest that ycdH and zosA genes function at different growth phases.

Furthermore, the CcpN regulon [Servant et al., 2005], the AhrC regulon [Dennis et al., 2002], the PyrR regulon [Switzer et al., 1999], eight genes of CymR regulon [Even et al., 2006], three genes of the DegSU regulon [Mäder et al., 2002] and six genes of the SOS regulon [Au et al., 2005] were induced under Mn2+ limitation. Three members of the CcpN regulon (gapB, speD and pckA) and the AhrC regulated argCJBD-carAB-argF operon were induced in both phases. Induction of the CcpN regulated gene gapB were also observed throughout growth while that of the AhrC regulated gene argD was observed only in the exponential phase on protein level. In addition, three members of the DegSU regulon (licT, sacX and sacY) and the PyrR dependent pyr operon were induced only in the transition phase. The PyrR regulated protein PyrAA (carbamoyl-phosphate synthetase) was also induced during transition and stationary phases. Moreover, the CymR dependent gene ssuB were

Results and discussion 136 Mn limitation induced only during the exponential growth, while other six genes (cysC, cysH, sat, ylnD, ylnE and ylnF) were induced in the transition phase and only the last one (yhzA) was induced in both phases. Two members of the CymR regulon (Mtn and Sat) were induced during the transition and/or stationary phases. The microarray analysis further revealed the induction of six genes (dinB, recA, tagC, yneA, yneB and ynzC) of the SOS regulon in the transition phase under Mn2+ limited conditions. The induction of recA gene was also found during stationary phase on the protein level. In this context, a report on a Mn2+ dependent ribonucleotide reductase in B. subtilis [Mohamed et al., 1998] is very important. Impairment of DNA precursor biosynthesis by Mn2+ limitation will inevitably cause phenomena of unbalanced growth, e.g., SOS response, inhibition of DNA synthesis and cell division [Elhariry et al., 2004].

Moreover, a subset of the ComK regulon [Berka et al., 2002; Ogura et al., 2002] and genes negatively regulated by Spo0A [Molle et al., 2003; Fujita et al., 2005] were upregulated whereas some genes regulated by σH and σE factors [Britton et al., 2002; Feucht et al., 2003; Steil et al., 2005] and genes positively regulated by Spo0A [Molle et al., 2003; Fujita et al., 2005] were downregulated in the transition phase. Some genes (comEB, nin and ssb) of the ComK regulon were also induced on the protein level under Mn2+ limitation. These data support a clear indication of inhibited sporulation and activated competence development in case less favorable for the sporulation, e.g. here, Mn2+ limitation. Furthermore, the ftsZ gene that plays a central role in bacterial cell division [Harry, 2001] and ftsA gene that is crucial for septum formation [Errington and Daniel, 2002] were downregulated on the mRNA level in the transition phase. The downregulation of these genes in combination with the induction of yneA gene (SOS regulon), the downregulation of genes belonged to the σM, σW, YycFG, YvrGHb and BceRS regulons [Cao et al., 2002; Mascher et al., 2003; Serizawa et al., 2005; Bisicchia et al., 2007; Eiamphungporn and Helmann, 2008] involved in cell wall stress response and the morphology observation (filamentous cell) under Mn2+ limitation suggests that the cell division is presumably blocked under Mn2+ limitation.

Finally, many nitrogen metabolism genes of the TnrA regulon [Yoshida et al., 2003] and about twenty σB dependent genes [Helmann et al., 2001; Petersohn et al., 2001; Price et al., 2001] were downregulated by the transcriptome analysis (Tab. 2.4). In 2003, Guedon and colleagues described that elevated Mn2+ activated the TnrA and σB regulons. They suggested that the TrnA regulator was controlled by Mn2+ dependent enzyme glutamine synthetase. Elevated Mn2+ reduced the sensitivity of glutamine synthetase to feedback inhibitors, and it lead to the observed increase in TnrA activity in their study. They also explained that the activation of the σB might result from the role of Mn2+ as a cofactor for regulatory protein phosphatases. Thus, our finding of the downregulation of TnrA and σB regulons under Mn2+ limitation adds the missing piece to TnrA and σB activation observed in the shift-up approach of Guedon et al. (2003).

Results and discussion 137 Mn limitation

pH 7 pH 4

Control-exponential phase

Mn limitation-exponential phase

KatA

AhpF

GapB GapB

ArgD

GapB

GtaB

AhpC

Fig. 2.8. The exponential-phase cytosolic proteome of B. subtilis under Mn2+ limitation in comparison with control. Proteins synthesized only under control condition appeared green, proteins synthesized only under the Mn2+ limitation appeared red, whereas proteins synthesized under both conditions appeared yellow.

Results and discussion 138 Mn limitation pH 7 pH 4

control-transition phase

Mn limitation-transition phase

InfB

KatA AhpF

GapB GapA

PyrAA

GapB YfhB Mtn

GtaB

YjbV Cmk

ComEB AhpC

Maf

Ssb

Nin

Fig. 2.9. The transition-phase cytosolic proteome of B. subtilis under Mn2+ limitation in comparison with control. Proteins synthesized only under control appeared green, proteins synthesized only under the Mn2+ limitation appeared red, whereas proteins synthesized under both conditions appeared yellow.

Results and discussion 139 Mn limitation

pH 7 pH 4

Control-stationary phase

Mn limitation-stationary phase

DnaK KatA

YvfW MetK AhpF RibA RecA

Sat GapB GapA

YxjG PyrAA YwaA

-GapB

Mtn GtaB

Maf

ComEB AhpC

Nin

AbrB

Fig. 2.10. The stationary-phase cytosolic proteome of B. subtilis under Mn2+ limitation in comparison with control. Proteins synthesized only under control appeared green, proteins synthesized only under the Mn2+ limitation appeared red, whereas proteins synthesized under both conditions appeared yellow.

Results and discussion 140 Mn limitation

Tab. 2.2. Summary of induced proteins under Mn2+ limitation compared to the control as revealed by 2-D PAGE analysis.

Ratio (fold) Protein Regulon Operon Function or similarity (Synonym) Exponential Transition Stationary phase phase phase PerR regulon ahpCF AhpC 14.27 8.70 2.91 alkyl hydroperoxide reductase (small subunit) AhpF 3.43 3.52 4.38 alkyl hydroperoxide reductase (large subunit) CymR katA KatA 61.09 36.65 27.62 vegetative catalase 1 ComK regulon ComEB late competence operon required for DNA comEABC 7.83 2.98 (ComD) binding and uptake nucA-nin Nin (ComJ) 3.07 5.39 inhibitor of the DNA degrading activity of NucA yyaF-rpsF- Ssb 2.50 single-strand DNA-binding protein ssb-rpsR CymR regulon yrrT-mtn- Mtn (YrrU) 3.90 4.30 methylthioadenosine nucleosidase yrhABC cysHP-sat- cysC- Sat (YlnB) 2.20 probable sulfate adenylyltransferase ylnDEF Other functions Spo0A, transcriptional pleiotropic regulator of transition abrB AbrB (CpsX) 3.34 AbrB state genes argCJBD- AhrC ArgD 3.10 N-acetylornithine aminotransferase carAB-argF Cmk (YpfC) 3.44 cytidylate kinase hrcA-grpE- HrcA DnaK 4.84 class I heat-shock protein (molecular chaperone) dnaK-dnaJ cggR-gapA- CggR pgk-tpiA- GapA (Gap) 2.11 2.06 glyceraldehyde-3-phosphate dehydrogenase 1 pgm-eno CcpN gapB-speD GapB 3.98 4.59 7.50 glyceraldehyde-3-phosphate dehydrogenase 2 σB gtaB GtaB 8.18 31.05 22.22 UTP-glucose-1-phosphate uridylyltransferase Maf 6.24 8.55 septum formation metK MetK (MetE) 2.62 S-adenosylmethionine synthetase carbamoyl-phosphate synthetase (glutaminase PyrR pyr operon PyrAA 2.81 4.00 subunit) multifunctional protein involved in homologous LexA, recA RecA (RecE) 2.90 recombination and DNA repair (LexA- ComK autocleavage) ypuE- GTP cyclohydrolase II / 3,4-dihydroxy-2- RibA 2.11 ribDEAHT butanone 4-phosphate synthase yfhB YfhB 2.08 unknown tenAI-goxB- YjbV 2.37 similar to phosphomethylpyrimidine kinase thiSGF-yjbV YvfW 5.38 unknown similar to branched-chain amino acid YwaA 2.57 aminotransferase YxjG 2.84 unknown

The name and the function of the induced proteins derived from the Subtilist database [http://genolist.pasteur.fr/SubtiList/]. All proteins with an induction factor of at least two as revealed by 2-D PAGE analysis were classified according to http://dbtbs.hgc.jp. The help of Dr. Dirk Albrecht (Greifswald) during identification of the induced proteins is acknowledged.

Results and discussion 141 Mn limitation

Tab. 2.3. Upregulated genes under Mn2+ limitation compared to the control as revealed by the DNA microarray analysis. Ratio (fold) Gene Regulon Operon Exponential Transition Function or similarity (Synonym) phase phase MntR regulon manganese ABC transporter mntABCD mntA (ytgA) 2.46 2.68 (membrane protein) manganese ABC transporter mntB (ytgB) 2.24 2.55 (ATP-binding protein) manganese ABC transporter mntC (ytgC) 2.47 2.98 (membrane protein) mntD (ytgD) 2.24 3.05 manganese ABC transporter mntH mntH (ydaR) 4.45 3.71 manganese transporter PerR regulon alkyl hydroperoxide reductase ahpCF ahpC 7.64 5.39 (small subunit) alkyl hydroperoxide reductase ahpF 9.54 6.62 (large subunit) transcriptional repressor of iron fur fur (yqkL) 3.29 2.55 uptake hemAXCDBL hemA 1.57 2.19 glutamyl-tRNA reductase negative effector of the hemX 1.84 2.37 concentration of HemA hemC 2.09 2.63 porphobilinogen deaminase hemD 2.29 2.52 uroporphyrinogen III cosynthase delta-aminolevulinic acid hemB 2.68 2.73 dehydratase glutamate-1-semialdehyde 2,1- hemL (hemK) 1.84 2.56 aminotransferase katA katA 62.04 26.40 vegetative catalase 1 metalloregulation DNA-binding mrgA mrgA 5.66 1.86 stress protein transcriptional repressor of the perR perR (ygaG) 2.25 1.82 peroxide regulon similar to heavy metal- zosA (ykvW) 4.31 1.65 transporting ATPase CcpN regulon glyceraldehyde-3-phosphate gapB-speD gapB 2.11 8.16 dehydrogenase S-adenosylmethionine speD (ytcF) 2.24 6.35 decarboxylase phosphoenolpyruvate pckA pckA (ppc) 2.39 9.10 carboxykinase AhrC regulon argCJBD-carAB- N-acetylglutamate gamma- AbrB argC 4.19 2.49 argF semialdehyde dehydrogenase ornithine acetyltransferase / argJ (argA, argE) 4.09 1.76 amino-acid acetyltransferase N-acetylglutamate 5- argB 5.15 2.77 phosphotransferase N-acetylornithine argD 5.05 2.54 aminotransferase carbamoyl-phosphate carA (cpaA) 4.76 3.37 transferase-arginine (subunit A) carbamoyl-phosphate carB (cpaB) 4.33 3.32 transferase-arginine (subunit B) argF 3.87 3.76 ornithine carbamoyltransferase PyrR regulon carbamoyl-phosphate pyr operon pyrAA 1.00 2.57 synthetase (glutaminase subunit) carbamoyl-phosphate pyrAB 0.88 2.85 synthetase (catalytic subunit) pyrB 1.48 3.60 aspartate carbamoyltransferase pyrC 1.12 2.63 dihydroorotase dihydroorotate dehydrogenase pyrD (pyrDI) 0.97 3.49 (catalytic subunit) orotate pyrE (pyrX) 0.98 4.03 phosphoribosyltransferase orotidine 5'-phosphate pyrF 0.94 3.42 decarboxylase dihydroorotate dehydrogenase pyrK (pyrDII) 0.99 3.73 (electron transfer subunit)

Results and discussion 142 Mn limitation

Ratio (fold) Gene Regulon Operon Exponential Transition Function or similarity (Synonym) phase phase PyrR regulon (continued) pyrP 1.31 2.89 uracil permease transcriptional attenuation of the pyrimidine operon / uracil pyrR 2.37 3.76 phosphoribosyltransferase activity CymR regulon Spx, S-box cysP operon cysC (ylnC) 1.17 2.68 probable adenylylsulfate kinase phosphoadenosine cysH 1.26 2.13 phosphosulfate probable sulfate sat (ylnB) 1.16 2.58 adenylyltransferase similar to uroporphyrin-III C- ylnD 1.17 3.03 methyltransferase ylnE 1.08 2.30 unknown similar to uroporphyrin-III C- ylnF 1.02 2.01 methyltransferase aliphatic sulfonate ABC Spx ssuBACD-ygaN ssuB (ygaL) 2.03 1.93 transporter (binding protein) Spx yhzA 2.31 2.07 similar to ribosomal protein S14 DegSU regulon transcriptional antiterminator required for substrate- CcpA licT-bglS licT 0.97 2.02 dependent induction and catabolite repression of bglPH negative regulatory protein of sacXY sacX (sacS) 1.41 3.04 SacY transcriptional antiterminator involved in positive regulation of sacY (sacS) 1.18 2.61 levansucrase and sucrase synthesis SOS regulon (LexA regulon) dinB dinB 0.82 3.08 nuclease inhibitor multifunctional protein involved in homologous recombination ComK recA recA (recE) 1.15 4.05 and DNA repair (LexA- autocleavage) possibly involved in polyglycerol tagC tagC (dinC) 0.96 6.86 phosphate teichoic acid biosynthesis ComK yneAB-ynzC yneA 0.94 4.97 unknown yneB 0.98 5.64 similar to resolvase ynzC 1.96 4.52 unknown Spo0A regulon transcriptional pleiotropic AbrB abrB abrB (cpsX) 1.33 2.83 regulator of transition state genes cotD cotD 1.55 2.42 spore coat protein (inner) ftsEX ftsE 2.00 2.57 cell-division ATP-binding protein ftsX 1.51 2.67 cell-division protein med-comZ med (yjaW) 0.96 2.13 positive regulator of comK comZ (zjzA) 1.08 2.28 late competence gene ykuJK-ykzF-ykuL- CcpA ykuJ 1.10 2.09 unknown ccpC yvyE-yvhJ yvyE (yvhK) 1.74 2.23 unknown similar to transcriptional yvhJ 1.70 2.74 regulator similar to capsular ywqCD ywqC 1.65 3.00 polysaccharide biosynthesis similar to capsular ywqD 1.42 2.23 polysaccharide biosynthesis ComK regulon σE bdbDC bdbC (yvgU) 0.90 2.74 thiol-disulfide oxidoreductase bdbD (yvgV) 1.04 2.63 thiol-disulfide oxidoreductase late competence protein required for processing and comC comC 0.88 4.53 translocation of ComGC, ComGD, ComGE, ComGG comEABC comEA (comD) 1.09 2.76 exogenous DNA-binding protein late competence operon comEB (comD) 1.13 3.33 required for DNA binding and uptake

Results and discussion 143 Mn limitation

Ratio (fold) Gene Regulon Operon Exponential Transition Function or similarity (Synonym) phase phase ComK regulon (continued) late competence operon comEC (comD) missing missing required for DNA binding and uptake (comGABCDEFG- comGA 0.67 2.91 late competence gene yqzE) comGB 0.64 3.73 DNA transport machinery comGC 0.63 3.57 exogenous DNA-binding comGD 0.67 3.68 DNA transport machinery comGE 0.64 5.11 DNA transport machinery comGF 0.71 6.40 DNA transport machinery comGG 0.80 9.28 DNA transport machinery yqzE 0.69 5.95 unknown AbrB, CodY, Rok, competence transcription factor DegSU, comK comK 0.93 6.01 (CTF) Med, Spo0A inhibitor of the DNA degrading nucA-nin nin (comJ) 0.88 6.18 activity of NucA nucA (comI) 0.83 6.56 membrane-associated nuclease σM radC radC (ysxA) 0.72 3.27 probable DNA repair protein sbcD-yirY-yisB sbcD (yixA) 1.39 2.48 exonuclease SbcD homolog similar to two-component ybdK ybdK 0.81 2.98 sensor histidine kinase [YbdJ] yvrPON yvrP 0.99 3.37 similar to ABC transporter similar to amino acid ABC yvrO 0.89 2.91 transporter (ATP-binding protein) similar to ABC transporter (ATP- yvrN 0.88 2.48 binding protein) (yyaF-rpsF- yyaF 1.25 4.61 similar to GTP-binding protein ssb-rpsR) rpsF 0.90 2.40 ribosomal protein S6 (BS9) single-strand DNA-binding ssb 0.96 2.26 protein rpsR missing missing ribosomal protein S18 Other functions ald ald (spoVN) 2.04 2.57 L-alanine dehydrogenase argG 5.57 4.89 argininosuccinate synthase argH 5.96 4.85 argininosuccinate lyase 3-deoxy-D-arabino- heptulosonate 7-phosphate aroA (aroG) 2.36 9.52 synthase / chorismate mutase- isozyme 3 aroB 1.44 3.37 3-dehydroquinate synthase aroC aroC 1.26 2.24 3-dehydroquinate dehydratase aroF 1.34 3.93 chorismate synthase chorismate mutase (isozymes 1 aroH 1.50 2.95 and 2) aroK aroK (aroI) 2.28 2.11 shikimate kinase cheR 1.38 2.52 MCPs methyltransferase cmk (jofC, ypfC) 1.62 2.55 cytidylate kinase non-essential gene for comER (comD) 0.99 2.75 competence csd (yurW) 2.01 1.47 probable cysteine desulfurase σE cwlJ cwlJ (ycbQ) 1.01 5.83 cell wall hydrolase (sporulation) penicillin-binding protein 5 (D- dacA 1.48 2.77 alanyl-D-alanine carboxypeptidase) CcpA, 6-phosphogluconate gntRKPZ gntZ 1.67 2.28 GntR dehydrogenase inosine-monophosphate guaB (guaA) 1.67 2.80 dehydrogenase PucR guaD guaD (yknA) 1.01 2.18 guanine deaminase CodY ilvD 1.59 2.13 dihydroxy-acid dehydratase σB, σF katX katX 3.08 2.02 major catalase in spores σG lysA 1.34 2.41 diaminopimelate decarboxylase maf 0.78 3.48 septum formation cobalamin-independent metE (metC) 1.19 2.92 methionine synthase S-adenosylmethionine metK metK (metE) 1.26 2.49 synthetase σH mreBCD-minCD mreB (divIVB) 1.00 2.94 cell-shape determining protein mreC 0.95 2.62 cell-shape determining protein

Results and discussion 144 Mn limitation

Ratio (fold) Gene Regulon Operon Exponential Transition Function or similarity (Synonym) phase phase Other functions (continued) Na+ ABC transporter (extrusion) natAB natA 1.62 2.09 (ATP-binding protein) pyruvate dehydrogenase (E1 pdhABCD pdhA (aceA) 1.25 3.22 alpha subunit) pyruvate dehydrogenase (E1 pdhB 1.14 2.90 beta subunit) pyruvate dehydrogenase pdhC 1.28 2.47 (dihydrolipoamide acetyltransferase E2 subunit) pyruvate dehydrogenase / 2- pdhD (aceD, oxoglutarate dehydrogenase 1.18 2.31 citL) (dihydrolipoamide dehydrogenase E3 subunit) phenylalanyl-tRNA synthetase pheST pheS 2.22 4.75 (alpha subunit) phenylalanyl-tRNA synthetase pheT 2.15 4.81 (beta subunit) riboflavin synthase (alpha ypuE-ribDEAHT ribE (ribB) 1.87 2.16 subunit) GTP cyclohydrolase II / 3,4- ribA 1.68 2.10 dihydroxy-2-butanone 4- phosphate synthase transcriptional regulator of AbrB, SenS senS senS 3.13 3.36 extracellular enzyme genes (amyE, aprE, nprE) tenAI-goxB- transcriptional regulator of TenI tenA 1.27 2.36 thiSGF-yjbV extracellular enzyme genes transcriptional activator of tenI 1.29 2.23 extracellular enzyme genes goxB (yjbR) 1.52 2.04 glycine oxidase hydroxyethylthiazole phosphate thiS (yjbS) 1.33 1.96 biosynthesis hydroxyethylthiazole phosphate thiF (yjbU) 1.38 2.34 biosynthesis hydroxyethylthiazole phosphate thiG (yjbT) 1.47 2.87 biosynthesis similar to yjbV 1.53 2.77 phosphomethylpyrimidine kinase D methyl-accepting chemotaxis σ tlpC tlpC 0.95 4.18 protein trpEDCFBA-hisC- TRAP trpE 1.09 2.23 anthranilate synthase tyrA-aroE ycbP 1.06 2.98 unknown similar to toxic cation resistance ycbR ycbR 1.19 4.12 protein similar to ABC transporter Zur ycdHI-yceA ycdH 1.28 2.24 (binding protein) similar to amino acid ABC yckA 0.66 4.52 transporter (permease) similar to amino acid ABC yckB 0.78 5.08 transporter (binding protein) yckC yckC 0.76 5.04 unknown σG yckD yckD 0.98 5.45 unknown similar to transcriptional ydeC 1.37 3.47 regulator (AraC/XylS family) ydeD 1.46 3.60 unknown similar to transcriptional ydeE 1.19 5.79 regulator (AraC/XylS family) similar to transcriptional ydeF 1.28 2.38 regulator (GntR family) / aminotransferase (MocR-like) ydjM ydjM (yzvA) 2.00 3.94 unknown ydzE 1.44 2.16 unknown yfhB yfhB 0.94 2.21 unknown yhdG yhdG 2.24 3.13 similar to amino acid transporter similar to glycerophosphodiester yhdW 1.67 1.29 phosphodiesterase yhdX 2.63 1.61 unknown similar to acetyl-CoA C- yhfS 1.32 2.15 acetyltransferase similar to long-chain fatty-acid- yhfT 1.29 2.13 CoA ligase

Results and discussion 145 Mn limitation

Ratio (fold) Gene Regulon Operon Exponential Transition Function or similarity (Synonym) phase phase Other functions (continued) yhfU 1.57 2.72 similar to biotin biosynthesis yjbL 1.73 2.18 unknown similar to GTP yjbM 1.12 2.20 pyrophosphokinase yjbN 1.16 2.22 unknown yjbO 1.16 2.78 unknown similar to diadenosine yjbP 1.09 2.03 tetraphosphatase similar to cystathionine gamma- yjcIJ yjcI 1.78 2.40 synthase similar to cystathionine beta- yjcJ 1.45 2.20 lyase ykoC 1.76 2.67 unknown similar to cation ABC ykoD 1.79 2.65 transporter (ATP-binding protein) ykoE 1.53 2.88 unknown ykoF 1.53 3.10 unknown ymaC 0.92 3.68 similar to phage-related protein ymaD 1.05 2.42 unknown yoaC 1.49 3.05 similar to xylulokinase similar to phosphoglycerate yoaD 1.78 2.75 dehydrogenase similar to formate yoaE 1.32 3.07 dehydrogenase similar to immunity to yocD 1.28 2.12 bacteriotoxins ypbS 1.55 2.16 unknown yqeN 1.18 3.35 unknown similar to amino acid ABC yqiX 4.33 2.62 transporter (binding protein) similar to amino acid ABC yqiY 2.65 1.82 transporter (permease) similar to amino acid ABC yqiZ 3.04 2.12 transporter (ATP-binding protein) σF yqzG yqzG 0.58 2.47 unknown ysgA ysgA 1.75 2.16 similar to rRNA methylase ytzD 5.60 3.65 unknown yurU 2.06 1.55 unknown yurV 2.00 1.47 similar to NifU protein homolog yurX 2.23 1.62 unknown similar to ABC transporter (ATP- yurY 2.00 1.78 binding protein) similar to cell wall-binding yvcE (yzkA) 3.27 6.45 protein similar to branched-chain amino ywaA ywaA 1.61 2.89 acid aminotransferase yweA yweA 1.06 3.15 unknown ywfM 0.87 2.52 unknown ywfO 1.45 2.94 unknown ywgA ywgA 1.38 2.63 unknown yxiP 1.12 2.48 unknown yxjG yxjG 1.38 2.10 unknown yyaK yyaK 1.86 2.26 unknown

The name and the function of the induced genes are derived from the Subtilist database [http://genolist.pasteur.fr/SubtiList/]. All genes with an induction factor of at least two as revealed by transcriptome analysis were classified according to http://dbtbs.hgc.jp. The help of Dr. Ulrike Mäder (Greifswald) to perform DNA microarray experiments is acknowledged.

Results and discussion 146 Mn limitation

Tab. 2.4. Downregulated genes under Mn2+ limitation compared to the control as revealed by the DNA microarray analysis.

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase Fur regulon 2,3-dihydro-2,3-dihydroxybenzoate dhbACEBF dhbA 0.65 0.25 dehydrogenase dhbC 0.64 0.25 isochorismate synthase 2,3-dihydroxybenzoate-AMP ligase dhbE 0.69 0.27 (enterobactin synthetase component E) dhbB 0.60 0.21 isochorismatase involved in 2,3-dihydroxybenzoate dhbF 0.61 0.32 biosynthesis similar to ferrichrome ABC transporter yclNOPQ yclP 0.70 0.50 (ATP-binding protein) similar to ferrichrome ABC transporter yclQ 0.62 0.39 (binding protein) ydbN ydbN 0.22 0.25 unknown yfkM yfkM 0.54 0.29 unknown ykuNOP ykuN 0.56 0.20 similar to flavodoxin ykuO 0.63 0.15 unknown ykuP 0.69 0.20 similar to flavodoxin yuiI yuiI 1.08 0.40 unknown similar to ABC transporter (binding yxeB yxeB 0.64 0.44 protein) CcpA regulon CodY acsA acsA 0.68 0.37 acetyl-CoA synthetase DegSU amyE amyE 0.53 0.42 alpha-amylase PTS beta-glucoside-specific enzyme bglPH-yxiE bglP 0.59 0.48 IIBCA component bglH 0.62 0.46 beta-glucosidase yxiE 0.28 0.27 unknown citZ-icd-mdh citZ 0.55 0.26 citrate synthase II glpFK glpF 0.43 0.25 glycerol uptake facilitator glpK 0.48 0.30 glycerol kinase similar to cellobiose phosphotransferase ydhO ydhO 0.94 0.22 system enzyme II multiple sugar-binding transport ATP- yxkF-msmX msmX (yxkG) 0.76 0.41 binding protein yxkJ yxkJ 0.64 0.46 similar to metabolite-sodium symport IolR regulon methylmalonate-semialdehyde (mmsA-iolBCDEF mmsA (iolA) 0.75 0.38 dehydrogenase -idh-iolHI-fbaB) iolB 0.58 0.35 myo-inositol catabolism iolC 0.62 0.36 myo-inositol catabolism iolD 0.64 0.35 myo-inositol catabolism iolE 0.76 0.36 myo-inositol catabolism iolF 0.73 0.40 inositol transport protein idh (iolG) 0.58 0.34 myo-inositol 2-dehydrogenase iolH 0.63 0.31 myo-inositol catabolism iolI (yxdH) 0.76 0.33 myo-inositol catabolism fbaB (iolJ) 0.74 0.43 fructose-1,6-bisphosphate aldolase transcriptional repressor of the myo- iolRS iolR 0.92 0.49 inositol catabolism operon iolS (yxbF) 0.69 0.42 myo-inositol catabolism PhoPR regulon vpr vpr 0.73 0.19 minor extracellular serine protease similar to 2',3'-cyclic-nucleotide 2'- yfkN yfkN 0.85 0.46 phosphodiesterase yjdB yjdB 0.32 0.03 unknown

Results and discussion 147 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase TnrA regulon GlnR glnRA glnA 0.84 0.31 glutamine synthetase pel pel 0.38 0.61 pectate lyase CodY, BkdR (ptb-bcd-buk- ptb (yqiS) 1.08 0.30 probable phosphate butyryltransferase lpdV-bkdAABB) bcd (yqiT) 0.94 0.29 leucine dehydrogenase probable branched-chain fatty-acid buk (yqiU) 1.02 0.28 kinase (butyrate kinase) probable branched-chain alpha-keto acid lpdV (yqiV) 1.02 0.32 dehydrogenase E3 subunit (dihydrolipoamide dehydrogenase) branched-chain alpha-keto acid bkdAA dehydrogenase E1 subunit (2- 0.98 0.34 (bfmBAA) oxoisovalerate dehydrogenase alpha subunit) branched-chain alpha-keto acid bkdAB dehydrogenase E1 subunit (2- 0.99 0.36 (bfmBAB) oxoisovalerate dehydrogenase beta subunit) branched-chain alpha-keto acid bkdB 1.11 0.35 dehydrogenase E2 subunit (lipoamide (bfmBB) acyltransferase) CodY regulon (srfAA-srfAB-comS srfAA 0.41 0.55 surfactin synthetase / competence -srfAC-srfAD) srfAB 0.46 0.58 surfactin synthetase / competence assembly link between regulatory comS 0.48 0.48 components of the competence signal transduction pathway srfAC 0.42 0.98 surfactin synthetase / competence srfAD 0.46 0.55 surfactin synthetase / competence σH ureABC ureA 0.70 0.20 urease (gamma subunit) ureB 0.73 0.20 urease (beta subunit) ureC 0.72 0.27 urease (alpha subunit) PurR regulon purEKBCSQLFMNHD purB 0.50 2.06 adenylosuccinate lyase phosphoribosylaminoimidazole purC 0.40 1.39 succinocarboxamide synthetase required for purS (yexA) 0.42 0.84 phosphoribosylformylglycinamidine synthetase activity phosphoribosylformylglycinamidine purQ 0.44 1.82 synthetase I phosphoribosylformylglycinamidine purL 0.43 1.75 synthetase II glutamine phosphoribosylpyrophosphate purF 0.42 1.62 amidotransferase phosphoribosylaminoimidazole purM 0.42 1.35 synthetase phosphoribosylglycinamide purN 0.42 1.34 formyltransferase phosphoribosylaminoimidazole carboxy purH (purJ) 0.46 1.26 formyl formyltransferase / inosine- monophosphate cyclohydrolase purD 0.46 1.45 phosphoribosylglycinamide synthetase xpt-pbuX xpt 0.45 1.61 xanthine phosphoribosyltransferase pbuX 0.48 1.19 xanthine permease σB regulon csbA csbA 0.75 0.35 putative membrane protein σX csbB csbB 0.70 0.35 stress response protein ywmF-csbD csbD (ywmG) 0.41 0.25 sigma-B-controlled gene stress- and starvation-induced gene dps dps 0.49 0.21 controlled by sigma-B gcaD-prs-ctc ctc 0.77 0.28 general stress protein katE-yxiS katE (katB) 0.86 0.45 catalase 2

Results and discussion 148 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase σB regulon (continued) nhaX nhaX (yheK) 0.56 0.21 putative regulatory gene for nhaC negative regulator of sigma-B activity rsbRSTUVW-sigB- rsbW 0.72 0.31 (switch protein/serine kinase, anti-sigma rsbX factor) RNA polymerase general stress sigma sigB 0.78 0.40 factor yacLMN yacL 0.97 0.44 unknown ydaDEFG ydaG 0.81 0.35 similar to general stress protein similar to organic hydroperoxide ohrB ykzA (ohrB) 0.60 0.19 resistance protein yocK yocK 0.61 0.32 similar to general stress protein yoxCB-yoaA yoxC 0.58 0.34 unknown yqgZ yqgZ 0.32 0.22 unknown σH ytxGHJ ytxG 0.61 0.24 similar to general stress protein ytxH 0.78 0.31 similar to general stress protein ytxJ 0.81 0.33 similar to general stress protein yvgO yvgO 0.87 0.27 unknown

H similar to sigma-54 modulating factor of σ , Spo0A yvyD yvyD (yviI) 0.36 0.30 gram-negative bacteria ywjC ywjC 0.26 0.20 unknown Cell wall responsive regulons σM yacK 0.83 0.35 unknown σW yceCDEFGH yceC 0.95 0.35 similar to tellurium resistance protein yceD 1.01 0.29 similar to tellurium resistance protein yceE 1.17 0.30 similar to tellurium resistance protein yceF 1.26 0.35 similar to tellurium resistance protein yceG 1.05 0.35 unknown yceH 1.33 0.30 similar to toxic anion resistance protein pspA-ydjGHI ydjH 0.78 0.30 unknown ydjI 0.86 0.31 unknown σB yfhKLM yfhK 0.41 0.12 similar to cell-division inhibitor yfhL 0.73 0.27 unknown yfhM 0.78 0.30 similar to epoxide hydrolase similar to cell-division protein FtsH yjoB yjoB 0.85 0.23 homolog AbrB yknWXYZ yknW 0.65 0.25 unknown yknX 0.60 0.32 unknown similar to ABC transporter (ATP-binding yknY 0.58 0.17 protein) yknZ 0.72 0.15 unknown yobJ yobJ 0.81 0.29 unknown yozO yozO 1.60 0.24 unknown yqeZ-yqfA-yqfB yqeZ 0.80 0.28 unknown yqfA 0.90 0.27 unknown yqfB 0.90 0.28 unknown σM, σX yqjL yqjL 1.02 0.41 unknown σX yrhH yrhH 0.82 0.29 similar to methyltransferase ysdB ysdB 0.90 0.27 unknown similar to ABC transporter (ATP-binding ythPQ ythP 1.22 0.30 protein) ythQ 1.01 0.30 unknown yuaFGI yuaF 0.70 0.21 unknown yuaG (yuaH) 0.76 0.22 similar to flotillin 1 yuaI 0.79 0.22 unknown

Results and discussion 149 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase σW (continued) AbrB yvlABCD yvlA 0.97 0.34 unknown yvlB 0.91 0.29 unknown yvlC 0.96 0.35 unknown yvlD 0.98 0.27 unknown ywrE ywrE 1.07 0.21 unknown DegSU yxjJI yxjI 0.97 0.42 unknown yxjJ 0.90 0.32 unknown AbrB yxzE yxzE 0.94 0.33 unknown YvrGHb wapA-yxxG wapA 0.36 0.25 cell wall-associated protein precursor yxxG 0.37 0.25 unknown yvrI-yvrH yvrI 1.34 0.35 unknown similar to two-component response yvrH 1.46 0.49 regulator [YvrG] YycFG yoeB 0.52 0.12 unknown BceRS similar to ABC transporter (ATP-binding bceAB ytsC (bceA) 1.08 0.09 protein) ytsD (bceB) 1.14 0.09 similar to ABC transporter (permease) σY regulon sigY-yxlCDEFG sigY (yxlB) 0.93 0.40 RNA polymerase ECF-type sigma factor yxlC 0.97 0.36 unknown yxlD 0.98 0.34 unknown yxlE 1.20 0.29 unknown similar to ABC transporter (ATP-binding yxlF 0.96 0.33 protein) yxlG 1.01 0.40 unknown ResDE regulon

E cytochrome caa3 oxidase (required for σ ctaA ctaA 0.43 0.99 biosynthesis) cytochrome caa3 oxidase (assembly ctaBCDEF ctaB 0.46 0.71 factor) ctaC 0.46 0.52 cytochrome caa3 oxidase (subunit II) ctaD 0.56 0.8 cytochrome caa3 oxidase (subunit I) ctaE 0.45 0.66 cytochrome caa3 oxidase (subunit III) ctaF 0.41 0.61 cytochrome caa3 oxidase (subunit IV) cytochrome bd ubiquinol oxidase CcpA, YdiH cydABCD cydA 0.18 0.47 (subunit I) cytochrome bd ubiquinol oxidase cydB 0.42 0.76 (subunit II) YdiH ldh-lctP ldh (lctE) 0.01 0.01 L-lactate dehydrogenase lctP (ycgC) 0.19 0.18 L-lactate permease TnrA, Fur nasBCDEF nasD 0.43 0.86 assimilatory nitrite reductase (subunit) essential protein similar to cytochrome c PhoPR resABCDE resA 0.41 0.79 biogenesis protein essential protein required for cytochrome resB 0.42 0.71 c synthesis Rok regulon AbrB, ResDE sboAX-albABCDEFG sboA (sbo) 0.12 0.12 subtilosin A sboX missing missing bacteriocin-like product antilisterial bacteriocin (subtilosin) albA (ywiA) 0.45 0.42 production antilisterial bacteriocin (subtilosin) albB (ywhR) 0.69 0.45 production antilisterial bacteriocin (subtilosin) albC (ywhQ) 0.83 0.48 production yydFGHIJ yydF missing missing unknown

Results and discussion 150 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase Rok regulon (continued) yydG 0.34 0.28 unknown yydH 0.34 0.29 unknown similar to ABC transporter (ATP-binding yydI 0.43 0.35 protein) yydJ 0.43 0.45 unknown AbrB regulon CodY dppABCDE dppA 0.81 0.12 D-alanyl-aminopeptidase dipeptide ABC transporter (permease) dppB 0.80 0.30 (sporulation) dipeptide ABC transporter (permease) dppC 0.67 0.27 (sporulation) dipeptide ABC transporter (ATP-binding dppD 0.71 0.26 protein) (sporulation) dipeptide ABC transporter (dipeptide- dppE 0.66 0.25 binding protein) (sporulation) similar to transcriptional regulator (DeoR ycnK ycnK 0.31 1.07 family) (yxbBA-yxnB- yxbB (yxaP) 0.34 0.18 unknown asnH-yxaM) yxbA 0.33 0.18 unknown yxnB 0.39 0.17 unknown asnH (yxaN) 0.37 0.26 asparagine synthetase yxaM 0.36 0.24 similar to antibiotic resistance protein similar to transcriptional regulator (ArsR sdpRI yvbA 0.96 0.11 family) yvaZ 0.83 0.11 unknown yxbCD yxbC (yxaQ) 0.44 0.17 unknown yxbD (yxaR) 0.44 0.36 unknown Spo0A regulon σF, σG, σH dacF-spoIIAAB-sigF spoIIAA 0.62 0.17 anti-anti-sigma factor spoIIAB 0.54 0.12 anti-sigma factor / serine kinase RNA polymerase sporulation forespore- sigF 0.60 0.10 specific (early) sigma factor response regulator aspartate rapA-phrA rapA 0.75 0.16 phosphatase phrA 0.81 0.16 phosphatase (RapA) inhibitor AbrB, PhoPR skf operon skfA (ybcO) 0.18 0.04 sporulation killing factor A skfB (ybcP) 0.15 0.02 similar to subtilosin (antibiotic production) skfC (ybcS) 0.14 0.03 unknown skfD (ybcT) 0.15 0.03 unknown similar to ABC transporter (binding skfE (ybdA) 0.19 0.04 protein) skfF (ybdB) 0.18 0.06 similar to ABC transporter (permease) skfG (ybdD) 0.25 0.06 unknown skfH (ybdE) 0.43 0.16 unknown

H two-component response regulator σ spo0F spo0F 0.70 0.32 involved in the initiation of sporulation F G protease (processing of pro-sigma-E to σ , σ spoIIGA-sigE-sigG spoIIGA 0.31 0.32 active sigma-E) sigE RNA polymerase sporulation mother cell- 0.94 0.39 (spoIIGB) specific (early) sigma factor σH racA ywkC (racA) 0.90 0.28 unknown yqxIJ yqxI (yqdE) 0.36 0.38 unknown yqxJ (yqdF) 0.45 0.31 unknown yqxM-sipW-tasA sipW (yqhE) 0.63 0.45 signal peptidase I translocation-dependent antimicrobial tasA (cotN) 0.36 0.29 spore component σH regulon AbrB, YycFG ftsAZ ftsA 1.20 0.38 cell-division protein (septum formation) cell-division initiation protein (septum ftsZ 1.23 0.40 formation)

Results and discussion 151 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase σH regulon (continued) spo0M spo0M (ygaI) 0.68 0.32 sporulation-control gene AbrB spoVG spoVG 0.71 0.29 required for spore cortex synthesis σE regulon glgBCDAP glgA 0.98 0.42 starch (bacterial glycogen) synthase glgP 0.94 0.46 glycogen phosphorylase UDP-N-acetylglucosamine-N- murE-mraY-murD- acetylmuramyl- SpoIIID spoVE-murG-murB- murG 1.03 0.37 (pentapeptide)pyrophosphoryl- divIB-ylxWX-sbp undecaprenol N-acetylglucosamine transferase UDP-N-acetylenolpyruvoylglucosamine murB 0.83 0.36 reductase yabQ 1.32 0.49 unknown σW, σX, yabMNOPQ-divIC- cell-division initiation protein (septum divIC 1.52 0.32 spoIIID yabR formation) Other functions acpA (acpP) 0.47 0.58 acyl carrier protein bpr (bpf) 0.48 0.58 bacillopeptidase F cccA 0.40 0.42 cytochrome c550 cggR-gapA-pgk-tpiA- CggR cggR (yvbQ) 0.40 0.28 transcriptional repressor of gapA pgm-eno secondary transporter of divalent metal citH citH (yxiQ) 0.69 0.41 ions/citrate complexes csn 0.37 0.30 chitosanase transcriptional repressor of class III CtsR ctsR-mcsAB-clpC ctsR 0.74 0.45 stress genes clpC 0.67 0.47 class III stress response-related ATPase dhaS 0.78 0.24 aldehyde dehydrogenase transcriptional regulator of anaerobic Fnr narK-fnr fnr 0.29 0.70 genes (narK-fnr, narGHJI, lctEP, alsSD) lepA-hemN-hrcA- transcriptional repressor of class I heat- HrcA hrcA 0.94 0.39 grpE-dnaKJ-yqeTUV shock genes radA (sms) 0.86 0.41 DNA repair protein homolog choline ABC transporter (ATP-binding opuBABCD opuBA missing missing protein) choline ABC transporter (membrane opuBB 0.68 0.43 protein) choline ABC transporter (choline-binding opuBC 0.60 0.38 protein) choline ABC transporter (membrane opuBD 0.64 0.44 protein) glycine betaine/carnitine/choline ABC opuCA 0.49 0.07 transporter (ATP-binding protein) glycine betaine/carnitine/choline ABC opuCB 0.74 0.33 transporter (membrane protein) glycine betaine/carnitine/choline ABC opuCC 0.76 0.30 transporter (osmoprotectant-binding protein) glycine betaine/carnitine/choline ABC opuCD 0.75 0.42 transporter (membrane protein) oxdC (yvrK) 0.82 0.11 oxalate decarboxylase transcriptional repressor of sporulation, paiA 0.76 0.33 septation and degradative enzyme genes transcriptional repressor of sporulation paiB (yumE) 0.82 0.28 and degradative enzyme genes pps 0.77 0.42 phosphoenolpyruvate synthase rpoB rpoB 0.82 0.39 RNA polymerase (beta subunit) rpoC 0.62 0.37 RNA polymerase (beta' subunit) transcriptional antiterminator involved in sacT sacT 0.89 0.30 positive regulation of sacA and sacP sdpA (yvaW) 0.76 0.42 unknown sdpB (yvaX) 0.61 0.41 unknown sdpC (yvaY) 0.43 0.19 unknown

Results and discussion 152 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase Other functions (continued) similar to chloramphenicol resistance ybcL ybcL 0.61 0.39 protein similar to two-component response ybdJ 0.62 0.41 regulator [YbdK] ybdG 0.38 0.18 unknown similar to transcriptional regulator ybfP 0.66 0.28 (AraC/XylS family) ybfQ 0.88 0.35 unknown ybyB ybyB 0.49 0.17 unknown similar to transcriptional regulator (LysR ycgK 0.88 0.44 family) ycnJ 0.43 1.29 similar to copper export protein yczC 0.42 0.91 unknown ydcO 1.18 0.48 unknown ydcP 0.94 0.47 similar to transposon protein similar to cellobiose phosphotransferase ydhM 0.92 0.23 system enzyme II similar to cellobiose phosphotransferase ydhN 0.87 0.19 system enzyme II ydhP 0.88 0.23 similar to beta-glucosidase similar to mannose-6-phosphate ydhS 1.00 0.48 isomerase ydzH 0.62 0.40 unknown yebC 1.08 0.30 unknown yebD 0.47 0.54 unknown yerA 0.22 0.35 similar to adenine deaminase yflT yflT 0.38 0.35 unknown yfmG yfmG 0.2 0.09 unknown yfmH yfmH 0.65 0.26 unknown yfmQ yfmQ 0.21 0.68 unknown ygxA 1.05 0.47 unknown similar to 5-oxo-1,2,5-tricarboxilic-3- yisK 0.71 0.42 penten acid decarboxylase yitM 0.44 0.10 unknown yitN 0.75 0.24 unknown yjbCD yjbC 0.66 0.29 unknown yjcE 0.4 0.15 unknown ykfA 0.64 0.23 similar to immunity to bacteriotoxins similar to chloromuconate ykfB 0.65 0.22 cycloisomerase ykfC 0.69 0.25 similar to polysugar degrading enzyme similar to oligopeptide ABC transporter ykfD 0.75 0.22 (permease) yklA yklA 0.42 0.68 unknown similar to transcriptional regulator (MarR ykoM 0.39 0.20 family) ykzI 0.34 0.26 unknown ylbP 0.57 0.30 unknown ymcA 0.86 0.28 unknown yncM 0.77 0.37 unknown yocB 0.91 0.35 unknown yocL 0.96 0.39 unknown yopA 0.88 0.42 unknown yopL 0.64 0.45 unknown yoqM 0.35 0.24 unknown yozB 0.37 0.64 unknown ypiB 0.57 0.20 unknown ypiF 0.65 0.33 unknown

Results and discussion 153 Mn limitation

Ratio (fold) Gene Regulon Operon Function or similarity (Synonym) Exponential Transition phase phase Other functions (continued) yrhP 0.87 0.26 similar to efflux protein yrpD 0.40 0.39 unknown ytkA 0.39 0.62 unknown yuaE 1.10 0.41 unknown yukJ 0.98 0.35 unknown similar to multiple sugar ABC transporter yurJ 0.83 0.20 (ATP-binding protein) similar to glutamine-fructose-6- yurRQPONML yurP 0.35 0.03 phosphate transaminase yurO 0.64 0.09 similar to multiple sugar-binding protein yurN 0.83 0.21 similar to sugar permease yurM 0.93 0.25 similar to sugar permease yurL 0.91 0.20 similar to ribokinase yuxI 0.84 0.37 unknown yvfD 0.77 0.45 similar to serine O-acetyltransferase yvmA 1.15 0.45 similar to multidrug transporter yvmB 0.75 0.39 involved in protein secretion yvmC 0.70 0.34 unknown ywaC 1.01 0.21 similar to GTP-pyrophosphokinase ywbH 0.15 0.49 unknown ywcI 0.85 0.36 unknown ywcJ 0.15 0.38 similar to nitrite transporter ywsB ywsB 0.54 0.27 unknown ywzA ywzA 0.51 0.24 unknown similar to arabinan endo-1,5-alpha-L- yxiA yxiA 0.68 0.40 arabinosidase yxiF 0.47 0.37 unknown yxiG 0.45 0.35 unknown yxiI 0.48 0.28 unknown yxiJ 0.50 0.43 unknown yxiK 0.46 0.43 unknown similar to rhamnogalacturonan yxiM-deaD yxiM 0.53 0.28 acetylesterase deaD 0.72 0.40 ATP-dependent RNA helicase yxlA yxlA 0.90 0.49 similar to purine-cytosine permease yxlH 1.19 0.46 similar to multidrug-efflux transporter yxxF yxxF 0.61 0.45 unknown yxzC 0.49 0.32 unknown yxzG 0.50 0.36 unknown

The name and the function of the repressed genes are derived from the Subtilist database [http://genolist.pasteur.fr/SubtiList/]. All genes with a repression factor of at least two as revealed by transcriptome analysis were classified according to http://dbtbs.hgc.jp. The help of Dr. Ulrike Mäder (Greifswald) to perform DNA microarray experiments is acknowledged.

Nguyen Thi Thu Huyen 154 References

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List of publications

1. Leelakriangsak, M., Huyen, N.T.T., Töwe, S., van Duy, N., Becher, D., Hecker, M., Antelmann, H., and Zuber, P. (2008) Regulation of quinone detoxification by the thiol stress sensing DUF24/MarR-like repressor, YodB in Bacillus subtilis. Mol Microbiol 67(5): 1108-24.

2. Huyen, N.T.T., Möhl, B., Schuster, F., Kobayashi, K., Becher, D., Hecker, M., and Antelmann, H. (2008) The MarR/DUF24-type repressor CatR (YvaP) controls the thiol-stress inducible catechol dioxygenase YfiE (CatE) in Bacillus subtilis. (in preparation).

3. Huyen, N.T.T., Eiamphungporn, W., Mäder U., Liebeke M., Hecker, M., Helmann, J.D., and Antelmann, H. (2008) Genome wide responses to carbonyl electrophiles in Bacillus subtilis: control of the thiol-dependent formaldehyde dehydrogenase AdhA and cysteine proteinase YraA by the MerR-family regulator YraB (AdhR). Mol Microbiol (submitted).

171

Curriculum vitae

Personal Information

Name Nguyen Thi Thu Huyen Date of birth August 30th , 1979 Place of birth Haiduong, Vietnam Nationality Vietnamese Sex Female Marital status Single

Education and work experience

09/1997-06/2001 B.A in Biotechnology (Grade: very good), Faculty of Biology, Hanoi University of Science, Vietnam 07/2001-05/2004 Researcher, Institute of Biotechnology, Vietnamese Academy of Science and Technology, Vietnam 02/2003-02/2004 Student of Diploma Equivalent course through Joint Graduate Education Program held by Hanoi University of Science, Institute of Biotechnology, Vietnam and University of Greifswald, Germany 02/2004 Diploma Equivalent certificate of the University of Greifswald, Germany 05/2004- present Ph.D student, Institute of Microbiology and Molecular Biology, Ernst- Moritz-Arndt University of Greifswald, Germany 05/2004-05/2007 Ph.D student, Institute of Microbiology, Leibniz University of Hannover, Germany 21/06/2004-02/07/2004 Practice course “Labor und Geländemethoden der Gewässerökologie” of Prof. Dr. Heike Schulz-Vogt, Institute of Microbiology, Leibniz University of Hannover, Germany 28/06/2005-30/06/2005 Workshop of “2D-DIGE: two dimensional difference gel electrophoresis” (organizer: GE healthcare), Institute of Microbiology, Leibniz University of Hannover, Germany 31/07/2005-06/08/2005 2nd summer school in proteomic basics “From samples to sequence” (Organizers: Dr. Henning Urlaub, Max-Planck-Institute for Biophysical Chemistry, Göttingen and Dr. Katrin Marcus, Medizinsches Proteom- Centre, Bochum), Kloster Neutift (Brixen/Bressanone, South Tyrol), Italy

172

Acknowledgements

First of all, I would like to express my thankfulness to my supervisors. I gratefully thank Prof. Dr. Michael Hecker, at the Institute of Microbiology, Ernst-Moritz-Arndt- University of Greifswald for giving me a special opportunity to work in the scientific atmosphere at his institute and his kindly supporting the final finance that I could enable to finish my thesis. I also sincerely thank to Prof. Dr. Georg Auling, at Institute of Microbiology, Leibniz University of Hannover, for giving me a nice chance to study in the friendly atmosphere at his institute. Finally, I especially thank my direct supervisor, Dr. Haike Antelmann, at the Institute of Microbiology, Ernst-Moritz-Arndt-University of Greifswald for her numerous ideas, suggestions and advices, excellent guidance, helpful discussion, kindly help and encouragement. Without her support, I could not get such quick, nice and interesting results for my thesis.

Secondly, I am very much thankful to many co-workers for sharing their contribution. I am much obligated to Dr. Ulrike Mäder for her microarray performance. I also would like to express my thankfulness to Dr. Dirk Albrecht and Dr. Dörte Becher for their mass spectrometry work. I would like to thank very much to Britta Möhl and Franzika Schuster for their cooperation in cloning, transcriptional analysis and protein purification. Many thanks come to Dr. Nguyen Van Duy for his phenotype analysis in YodB paper. I also would like to thank Manuel Liebeke for his metabolome work. I gratefully acknowledge Prof. Dr. Peter Zuber and Montira Leelakriangsak (Oregon Health and Science University, Beaverton, OR, USA) for their primer extension experiments, YodB purification and DNase I footprinting analyses. I also greatly appreciate Prof. Dr. John D. Helmann and Warawan Eiamphungporn (Cornell University, Ithaca, NY, USA) for their supporting of strains and their performance of DNA gel mobility shift assay. Finally, many thanks to Dr. Kazuo Kabayashi (Nara Institute of Science and Technology, Nara, Japan) for his transcriptional factor array performance.

Thirdly, I would like to say many thanks to all members in the Institute of Microbiology, Ernst-Moritz-Arndt-University of Greifswald and in Institute of Microbiology, Leibniz University of Hannover for their help and friendly working atmosphere. I gratefully acknowledge the proteome group of Bacillus subtilis for kindly sharing information, helpful advice and experiences. A special thank goes to Dr. Olaf Kniemeyer for kindly training the 2-D PAGE. Many thanks to Armgard Janczikowski and Inge Reupke for their kind help with the physiological works of Mn limitation part.

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Finally, my special appreciation is expressed to Anita Harang, Anke Arelt and Sebastain Grund for their experimental support.

Fourthly, I gratefully appreciate the 3-year financial support of the Ministry of Education and Training of Vietnam (MOET) and all members of the 322 projects enable me to this Ph.D. work. I also would like to thank the German Academic Exchange Service (DAAD) for additional annual financial support. I also highly appreciate all organizers of the Joint Graduate Education Program (JGEP) that I luckily joined, especially Prof. Dr. Le Tran Binh, Prof. Dr. Le Thi Lai, Dr. Luu Lan Huong and last but not least, PD. Dr. Jörn Kasbohm, who supports me the finance for one additional year.

Finally, my mother with her endless love and all friends of mine are the great encouragement for me to fulfill my Ph.D. work. I would like to show the deep gratitude from the bottom of my heart to them!

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

Ferner erkläre ich, daß ich diese Arbeit selbständig verfaßt und keine anderen als die darin angegebenen Hilfsmittel benutzt habe.