Roles of DNA Sequence and Sigma A Factor in Transcription of thevraSR Operon

Antoaneta Belcheva,a Vidhu Verma,b Artyom Korenevsky,a Michael Fridman,b Krishan Kumar,c and Dasantila Golemi-Kotraa,b,c

Departments of Biologya and Chemistry,b York University, Toronto, Ontario, Canada, and Department of Chemistry and Biochemistry, University of North Carolina,Greensboro, North Carolina, USAc

Cell wall damage in Staphylococcus aureus induces a rapid genome-wide response, referred to as the cell wall stress stimulon.This response is mediated by a two-component system, the vancomycin resistance-associated sensor/regulator (VraSR). The re-sponse regulator protein VraR is a transcription factor. Here, we demonstrate that two VraR binding sites in the vraSR operoncontrol region are involved in the regulation of the vraSR operon. The sites are centered at the �60 and �35 nucleotide positionsand are referred to as R1 and R2, respectively. DNase I footprinting and lux operon reporter vector studies showed that both ofthese sites communicate intimately with each other to fine-tune the activity of the vraSR operon. Mutagenesis of the VraR bind-ing sites showed that dimerization of unphosphorylated VraR at R1 is driven by a hierarchy in VraR binding and by the proxim-ity of the two tandem VraR binding sequences at this site. On the other hand, these studies show that the lack of sequence con-servation and the distance between the VraR binding sequences in R2 ensure that VraR is recruited to this site only whenphosphorylated (hence, under stress conditions). Furthermore, we demonstrate that sigma A (SigA) factor is involved in the reg-ulation of the vraSR operon. Our study shows that sigma A factor does not bind to the vraSR operon control region in the ab-sence of VraR, suggesting that VraR may interact directly with this factor.

The two-component vancomycin resistance-associated sensor/regulator (VraSR) signal transduction system of Staphylococ-

cus aureus coordinates the bacterial response to cell wall damageand the disruption of cell wall synthesis (7, 21, 22) caused by abroad class of antimicrobial agents (cell wall active inactivators).Inactivation of vraSR both decreases the resistance of different S.aureus strains to �-lactams and vancomycin and converts the ho-mogeneous oxacillin resistance phenotype into a highly heteroge-neous resistance phenotype (15, 25). In contrast, a constitutivelyactive vraSR operon, caused by a single-point mutation in vraS,leads to increased teicoplanin resistance of clinical glycopeptide-intermediate S. aureus strains (19). These findings corroborate theproposed role of VraSR as a sentinel of cell wall integrity (15).

VraSR is a typical two-component system (TCS) composed ofa histidine kinase (HK) and a response regulator (RR) that canrapidly sense and transduce cell wall stress (1, 22, 25). VraS issuggested to be the only biologically relevant kinase of the RRprotein VraR, and hence, the VraSR system may be the main path-way through which the signal to cell wall stress response is trans-duced (1). A recent study by Galbusera et al. showed that theremoval of the phosphorylation site from vraS prevents the emer-gence of glycopeptide resistance in several S. aureus strains (13).

Sequence alignments of VraR with other response regulatorsindicate that this protein belongs to the NarL/FixJ subfamily ofproteins (1). These proteins use helix-turn-helix motifs to bind toDNA (14). Although the members of this subfamily have highsequence similarity at the C terminus (DNA-binding domain),each member recognizes unique DNA sequences and utilizes dif-ferent regulatory strategies. This is likely due to subtle differencesin the primary structure of the helix-turn-helix motif (32) and/orthe tertiary structure of the active state of the response regulators(14). Such diversity in regulatory schemes makes it challenging topredict the general gene regulation mechanism used in this pro-tein family.

Recently, we showed that VraR binds to the vraSR operon con-

trol region at three possible sites (Fig. 1), referred to as the R1, R2,and R3 sites (2). VraR binding sites centered at positions �60 and�35 were analyzed independently, and in vitro studies showedbinding of VraR to R1 was not affected by phosphorylation ofVraR, while unphosphorylated VraR did not bind to R2. Back-ground expression of the vraSR operon is required for propersynthesis of bacterial cell wall peptidoglycan. However, it is notknown how the basal expression level is achieved. In addition, thesigma factor or factors involved in the VraSR-mediated cell wallstimulon have not been identified.

In this study, we investigated the binding mode of VraR to thevraSR operon control region. We examined the role of DNA se-quence in the regulation scheme of vraSR operon expression andin distinguishing between normal and stress conditions. Further,our analysis of the vraSR promoter suggests involvement of anEscherichia coli �70-like factor in the transcriptional regulation ofthe vraSR operon. Herein, we present our investigation into therole of the S. aureus sigma A (SigA) factor in vraSR operon expres-sion.

MATERIALS AND METHODSGrowth media and chemicals. Chemicals were purchased from Sigma-Aldrich or Fisher Scientific, unless otherwise stated. The Escherichia coliNova Blue strain and the pSTBlue cloning vector were purchased fromNovagen. Restriction enzymes were obtained from either New Eng-land BioLabs or Stratagene. The [�-32P]ATP (3,000 Ci/mmol) and

Received 9 September 2011 Accepted 18 October 2011

Published ahead of print 21 October 2011

Address correspondence to Dasantila Golemi-Kotra, dgkotra@yorku.ca.

Supplemental material for this article may be found at http://jb.asm.org/.

Copyright © 2012, American Society for Microbiology. All Rights Reserved.

doi:10.1128/JB.06143-11

0021-9193/12/$12.00 Journal of Bacteriology p. 61–71 jb.asm.org 61

[�-32P]UTP (3,000 Ci/mmol) were purchased from PerkinElmer. TheE. coli RNA polymerase (RNAP) holoenzyme and the ATP, GTP, CTP,and UTP solutions were obtained from Epicenter Biotechnologies.The murine RNase inhibitor (40 U/�l) was purchased from New Eng-land BioLabs.

Phosphorylation of VraR by acetyl phosphate. Full-length VraR wasexpressed in E. coli strain BL21(DE3) and purified as previously described(1). The phosphorylation of VraR by inorganic phosphate was carried outas described previously (1). Briefly, 50 �M VraR was equilibrated in buffercontaining 50 mM Tris at pH 7.4, 50 mM KCl, and 5 mM MgCl2 (phos-phorylation buffer [PB]). Lithium potassium acetyl phosphate was addedto a final concentration of 50 mM, and the reaction mixture was incubatedfor 60 min at 37°C. The extent of VraR phosphorylation was estimated byhigh-pressure liquid chromatography (HPLC) (Varian Inc.) using a Pro-Sphere HP C4 reverse-phase column (5 �m, 300 Å, 4.6 � 250 mm).Elution of the proteins was carried out on a 40-to-48% linear gradient ofacetonitrile and 0.1% trifluoroacetic acid (TFA) over 40 min at a flow rateof 1 ml/min. The extent of VraR phosphorylation was estimated by assess-ment of the surface areas under the respective peaks. Under these phos-phorylation conditions, 46% VraR was phosphorylated. Here, we refer tothis protein mixture as VraR-P (mixture of phosphorylated and unphos-phorylated VraR in which the phosphorylated species typically comprised46% of the total VraR protein).

Mutagenesis of the vraSR operon control region. A DNA sequencederived from the vraSR operon control region, which includes nucle-otides �121 to �26, was amplified by PCR using the genomic DNA ofS. aureus Mu50 as a template and the primers Dir_PvraSR, 5=-AGGAATTCGGTCCATTTTAACGACAAAAATTG-3=, and Rev_PvraSR, 5=-CGGGATCCTGAAATGACGCATTGATTGTGTTC-3=, which introduced re-striction sites for EcoRI and BamHI (italicized in the sequences). ThePvraSR promoter was subsequently ligated into the blunt-end vectorpSTBlue-1. The ligation mixtures were introduced into E. coli DH5� byheat shock, and the cells were subsequently plated onto Luria Bertani (LB)agar plates supplemented with 50 �g/ml kanamycin, 0.1 M IPTG(isopropyl-�-D-thiogalactopyranoside), and 20 �g/ml X-Gal (5-bromo-4-chloro-3-indolyl-�-D-galactopyranoside) for white and blue screening.The putative-positive transformants were confirmed by sequencing theplasmid for the presence of the insertion. The pSTblue-1::PvraSR plasmidwas used as a template to generate various PvraSR mutants carrying one ortwo mutations in the essential binding elements from the R1, R2, and R3binding sites. A QuikChange mutagenesis kit obtained from Stratagenewas used for the mutagenesis. The mutagenic primers used in this studyare shown in Table S1 in the supplemental material. For cases in whichtwo nucleotides were substituted, the target mutations were carried out intwo steps. The replaced nucleotides are underlined and highlighted inTable S1.

Cloning, isolation, and purification of S. aureus SigA. The sigA genesequence was amplified from S. aureus RN4220 genomic DNA by PCRusing the following primers: Dir-SigA, 5=-ACGCCATGGCTATGTCTGATAACACAGTT-3=, and Rev-SigA, 5=-ACGAAGCTTTTAATCCATAAAGTCTTTCA-3=. The sigA amplicon was double digested with NcoI and

HindIII and ligated into the pET24d(�) vector. The sequence of sigA wasconfirmed by DNA sequencing. The vector pET24a::sigA was subse-quently introduced to E. coli BL21(DE3). Expression of the sigma A factorwas induced with 1 mM IPTG when the culture reached an optical density(OD) of 0.8. Cells were then shaken for 16 h at 25°C.

Purification of SigA was carried out in two steps. In the first step, SigAwas purified on a DEAE-Sepharose column and eluted using a 0-to-100%linear gradient of buffer B (20 mM Tris, 0.1 mM EDTA, 0.1 mM dithio-threitol [DTT], and 1.0 M NaCl, pH 7.9). The second purification wasdone on a heparin-Sepharose column. Coomassie blue staining of SDS-PAGE indicated that the isolated protein was 90% pure. Protein concen-tration was determined by the Bradford assay.

Electromobility shift assays. The pSTblue-1::PvraSR plasmid carryingthe vraSR operon control region (nucleotides �121 to �26) or the mu-tated sequence (see above) was amplified by PCR using the above-mentioned primers. The DNA-binding activities of VraR and VraR-Pused to target DNA sequences were analyzed with electromobility shiftassays (EMSAs). The double-stranded DNA sequences were labeled withT4 polynucleotide kinase using [�-32P]ATP (3,000 Ci/mmol). Bindingreaction mixtures (20 �l) were prepared in binding buffer (10 mM Tris atpH 7.5, 50 mM KCl, 1 mM DTT) supplemented with 5 mM MgCl2, 0.05%NP-40, 200 ng/�l poly(dI-dC), and 2.5% glycerol. Target DNA (2 ng) wasmixed with protein concentrations ranging from 1 to 20 �M. The reactionmixtures were incubated at 25°C for 30 min and loaded into a 10% nativepolyacrylamide gel. Dried gels were exposed to a phosphor screen andanalyzed with a Typhoon Trio� variable-mode imager (GE HealthCare).The incubation time was experimentally determined as the time requiredfor equilibrium to be reached at the smallest VraR (VraR-P) concentra-tion used in these assays. Each EMSA was repeated at least three times. Thedissociation constants (Kd) were determined from nonlinear regressionanalysis of the experimental data using the following binding equation,where n is the Hill coefficient, Cap is the maximum binding capacity (themaximum number of bound species formed between the DNA and theprotein at the end of the titration assay), obtained from the nonlinearfitting of the data, and [P] is the protein concentration used in the assay:percentage of DNA bound � Cap � [P]n/(Kdn � [P]n).

DNase I footprinting assay. The DNA sequence encompassing nucle-otides �121 to �26 (with respect to the transcription start site) of thevraSR operon control region (PvraSR) was amplified using the followingPCR primers: Dir, 5=-ACGAAGCTTGGTCCGATTTTAACGACAAAAATTG-3=, and Rev, 5=-TGAAATGACGCATT GATTGTGTTC-3=. ForDNase I footprinting, the primer of interest (forward or reverse) was 5=end labeled with T4 polynucleotide kinase in the presence of [�-32P]ATP(3,000 Ci/mmol) and was used to amplify PvraSR. Binding reaction mix-tures (20 �l) were prepared in binding buffer (100 mM Tris at pH 7.5, 500mM KCl, 10 mM DTT) supplemented with 5 mM MgCl2, 0.05% NP-40,200 ng/�l poly(dI-dC), and 2.5% glycerol. The 32P-end-labeled PvraSR (10ng) was mixed with VraR or VraR-P at concentrations ranging from 2 to5 �M. Of note, the VraR-P represented a mixture of phosphorylated andunphosphorylated VraR in which the phosphorylated species typicallycomprised 46% of the total VraR protein. Binding reactions were carriedout at 25°C for 30 min and subsequently subjected to DNase I (2 U) at25°C for 2 min. The digestion reactions were stopped by addition of 50 �lDNase I stop solution (1% SDS, 0.2 M NaCl, 20 mM EDTA at pH 8.0, and0.25 mg/ml tRNA). Digested DNA samples were isolated by phenol-chloroform extraction and ethanol precipitation. Pellets containing DNAwere air dried and then resuspended in formamide containing loadingdye. The samples were heated at 95°C for 3 min and loaded into an 8%polyacrylamide gel containing 7 M urea. The dried gels were exposedto a phosphor screen and scanned using a Typhoon Trio� variable-mode imager. The C and G sequencing reactions for these experimentswere carried out with the same end-labeled oligonucleotide using ter-minator DNA polymerase and acyclic nucleotides GTP and CTP(acyGTP and acyCTP) (16).

FIG 1 VraR binding sites on PvraSR. The VraR binding sequences are under-lined, and the nucleotides proposed to be essential for VraR recognition andbinding are shown in boldface. The transcription starting point is denoted byan asterisk, and the �10 box sequence is double underlined.

Belcheva et al.

62 jb.asm.org Journal of Bacteriology

KMnO4 footprinting assay. A DNA sequence spanning between nu-cleotides �126 and �10 of the vraSR promoter region was amplifiedfrom genomic DNA of S. aureus RN4220 using PFU DNA polymerase andthe following primers: DIR, 5=-AGGAATTCGGTCCGATTTTAACGACAAAAATTG, and REV, 5=-GATGATATGTTTGCTTAAAAACTACTT.Binding reaction mixtures containing 10 ng of end-labeled PvraSR, VraR (2and 5 �M), or VraR-P (2 and 5 �M) and E. coli RNAP at a final concen-tration of 40 nM were prepared in 10 mM Tris at pH 7.5, 50 mM KCl, 5mM MgCl2, 0.05% NP-40, 200 ng/�l poly(dI-dC), and 2.5% glycerol. Thecomplexes were incubated at 30°C for 20 min. The samples were treatedwith heparin (50 �g/ml) at 30°C for 10 min. Subsequently, a 3-�l KMnO4

solution (20 mM) was added to the complexes for 30 s, after which thereaction was stopped with 3 �l DTT (200 mM), 0.4 M sodium acetate atpH 8.0, 2.5 mM EDTA, and 60 �g/ml herring sperm DNA. The reactionmixtures were immediately placed on ice for 5 min. The proteins wereremoved by phenol-chloroform extraction, and the DNA was precipitatedin 100% ice-cold ethanol at �70°C for 1 h. The DNA pellets were washedonce with 70% ethanol. The dried DNA pellets were resuspended in 100 �lpiperidine (1 M) and heated at 90°C for 30 min. The piperidine solutionwas completely evaporated, and each DNA sample was dissolved in 20 �lwater. The samples were vortexed, allowed to dry, and subsequentlywashed twice with 20 �l water. Finally, the samples were dissolved informamide containing loading dye, heated at 95°C for 3 min, and imme-diately loaded into an 8% polyacrylamide gel containing 7 M urea. Thedried gels were exposed to a phosphor screen and scanned using a Ty-phoon Trio� variable-mode imager.

Runoff in vitro transcription assay. Single-round transcription byE. coli RNAP was performed using a 612-bp fragment of the vraSRpromoter region (PvraSR

L) as a template in the presence or absence ofVraR-P. The PvraSR

L, which spans the region between nucleotides �426 to�150 of the vraSR promoter region, was amplified by PFU Turbo DNApolymerase and the following primers: Dir_PvraSR

L, 5=-AGGAATTCATGGCATTTGAGAATGCA-3=, and Rev_PvraSR

L, 5=-CGGGATCCACGTTCAACATAGTTCATAAC-3= (EcoRI and BamHI sites are italicized in thesequences). To make the PvraSR

L R1DM3 and PvraSRL R2SM1 mutants, we

used the same mutagenic primers used for the R1DM3 and R2SM1 PvraSR

mutants (see Table S1 in the supplemental material). The in vitro tran-scription reactions were performed as previously described, with somemodifications (10). The DNA template (5 nM) was incubated with VraRor VraR-P (5 �M) in transcription buffer (40 mM Tris at pH 7.5, 150 mMKCl, 10 mM MgCl2, 2 mM DTT, 100 �g/ml bovine serum albumin [BSA],and 0.05% Triton X-100) at 30°C for 20 min. One unit of E. coli RNAP (1�g) was added to the mixture, and the reaction mixtures were incubatedfor another 20 min at 30°C. A control reaction mixture containing onlyRNAP (without the VraR proteins) was prepared similarly. Transcriptionwas initiated by addition of a 3-�l nucleoside triphosphate (NTP) mixcontaining 1 mM ATP, 1 mM CTP, 1 mM GTP, 100 �M UTP, 2 �Ci[�-32P]UTP, 600 �g/�l heparin, and murine RNase inhibitor (40 U/�l).The reaction mixtures were placed at 30°C for 20 min and then terminatedby addition of 5 �l transcription stop buffer (20 mM EDTA, 1% SDS, 1 mgbromophenol blue, 1 mg xylene cyanol, and 90% formamide). The sam-ples were heated at 90°C for 5 min and immediately loaded into an 8%polyacrylamide gel containing 7 M urea. The �X174 DNA/HinfI-dephosphorylated DNA fragments (Promega) labeled with [�-32P]ATP(3,000 Ci/mmol) were used as molecular weight markers. The gels weredried, exposed on a phosphor screen, and visualized with a TyphoonTrio� variable-mode imager. The intensity of the resulting transcripts wasmeasured using ImageJ software (National Institutes of Health).

To investigate whether S. aureus SigA activates transcription specifi-cally from the vraSR promoter, we performed in vitro runoff transcriptionassays. In these experiments, the surrogate RNAP haloenzyme is formedbetween S. aureus SigA and the E. coli RNAP core enzyme using excessSigA (10, 20, and 25 molar excess). The reaction mixtures were preparedas follows. The DNA template (5 nM) was incubated with SigA atdifferent concentrations (2 �M, 4 �M, and 8 �M) and VraR (5 �M) in

the transcription buffer (40 mM Tris at pH 7.5, 150 mM KCl, 10 mMMgCl2, 2 mM DTT, 100 �g/ml BSA, and 0.05% Triton X-100) for 5min at 30°C. One unit of E. coli core polymerase, which lacks the E. colisigma subunit, was added (0.2 �M), and the reaction mixture (10 �l)was incubated for an additional 20 min at 30°C. Control reactionmixtures containing core polymerase but not SigA and VraR wereprepared in a similar way.

Determination of the VraR concentration in S. aureus RN4220.Whole-cell extracts were prepared as follows. An overnight seed culture ofS. aureus strain RN4220 was prepared in 5 ml tryptic soy broth (TSB)medium. A 1-ml aliquot of the seed culture was inoculated in 100 ml TSBmedium, and the cells were grown at 37°C to an OD at 600 nm (OD600) of0.8 and then harvested by low-speed centrifugation (2,500 � g, 10 min).The weight of the resulting pellet was measured, and the cells were resus-pended in 3 ml of 50 mM Tris, pH 7.0, and 5 mM MgCl2. The cells werelysed by incubation with 8 �g/ml lysostaphin for 30 min at 37°C. Subse-quently, DNase I (1 U/ml) was added, and the cells were incubated foranother 10 min at 37°C. The cell content was liberated by sonication. Acocktail of protease inhibitors (Roche) was added, and the cell lysates werestored at �80°C. Preparation of cell extracts from oxacillin-induced cellswas carried out by a similar procedure. At an OD600 of 0.8, the cells wereincubated with 20 �g/ml oxacillin for 20 or 40 min at 37°C, after which thecells were harvested and cell lysates obtained, as described above. Theprotein concentrations in the isolated total cell lysates from uninducedand oxacillin-induced cells were estimated by the Bradford assay (3).

Cell counts in the uninduced and oxacillin-induced cell cultures weredetermined by plating serial dilutions of cells on TSB/agar plates. Theplates were incubated for 12 h at 37°C, and the number of resulting colo-nies was counted.

VraR was detected using a monoclonal antibody, referred to as anti-VraR (GenScript), by Western blotting. Samples of total cell lysates (1.6mg) from uninduced or oxacillin-induced cells were prepared in a totalvolume of 15 �l. In addition, samples of purified VraR at five differentconcentrations ranging from 1 to 5 �g were prepared at a final volume of15 �l. All samples were mixed with 5 �l 5� SDS loading dye and loadedonto 15% SDS-polyacrylamide gels. The proteins then were electrotrans-ferred onto Hybond-P polyvinylidene difluoride (PVDF) membranes(GE Healthcare) at 48 mA for 45 min at 4°C. The proteins were detectedby immunoblotting with anti-VraR (dilution of 1:10,000), followed byanti-rabbit IgG antibody conjugated to horseradish peroxidase (1:2,500).Proteins were visualized by addition of a 3,3=,5,5=-tetramethylbenzidine(TMB) liquid substrate system (Sigma-Aldrich). Band intensities forVraR standards and the VraR proteins from cell lysates were measured byImageJ software. Subsequently, a standard curve of VraR concentrationswas generated by plotting the intensity of the VraR bands against the VraRconcentration. The concentration of VraR in the cell lysates was deter-mined from the standard curve.

Construction of luxABCDE fusion vectors from variants of thevraSR operon control region. The DNA sequence derived from the vraSRoperon control region, which encompasses nucleotides �121 to �26,was amplified by PCR using the genomic DNA of S. aureus RN4220 asa template and was cloned in the blunt-end vector pSTBlue-1 as pre-viously described (2). The resulting plasmid was named pSTblue-1::PvraSR. The pSTblue-1:PvraSR plasmid was used as a template in theconstruction of the PvraSR mutants using the QuikChange mutagenesiskit (Stratagene). The sequences of the mutagenic primers used areprovided in Table S1 in the supplemental material.

After mutagenesis, the DNA fragments were excised from thepSTBlue-1 fusion plasmids with the restriction enzymes EcoRI andBamHI. These amplicons are here referred to as PvraSRR1C67T,PvraSRR1G62T, PvraSRR1C67T/G62T, PvraSRR1G58T, PvraSRR1G52T,PvraSRR1G58T/C52T, PvraSRR1�1A, PvraSRR1�2A, and PvraSRR2C43A,PvraSRR2C31A, PvraSRR2G25A, PvraSRR2C43A/G38A, PvraSRR2G38A,PvraSRR2C31A/G25A, PvraSRR3G85T/C79T, and PvraSRR3C73T/G69. Thefragments were gel purified and cloned in the pXEN1 vector (12). The

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January 2012 Volume 194 Number 1 jb.asm.org 63

putative positive clones were confirmed to harbor the insertions by se-quencing the plasmid with a reverse primer, 5=-GTAAGCAAAAGTTTCCAAATTTCAT-3=, derived from the luxA sequence, and the forwardprimer was used to amplify the insertion. The pXEN1 plasmid carryingPvraSR is referred to as the lux fusion plasmid (PvraSRR1C67T::lux,PvraSRR1G62T::lux, PvraSRR1G58T::lux, PvraSRR1G52T::lux, PvraSRR1G58T/C52T::lux, PvraSRR1�1A::lux, PvraSRR1�2A::lux, PvraSRR2C43A::lux,PvraSRR2C31A::lux, PvraSRR2G25A::lux, PvraSRR2C43A/G38A::lux,PvraSRR2G38A::lux, PvraSRR2C31A/G25A::lux, PvraSRR3G85T/C79T::lux,and PvraSRR3C73T/G69::lux). Each lux fusion plasmid was introducedinto restriction-deficient S. aureus strain RN4220 by electroporation (29).The transformants were plated on TSB/agar plates supplemented with5 �g/ml chloramphenicol. The putative positive colonies were confirmedby sequencing using the primers listed above. The S. aureus RN4220strain carrying a lux fusion plasmid is referred to as the lux fusion strain,as follows: RN(PvraSR R1C67T::lux), PvraSRR1G62T::lux),RN(PvraSRR1G58T::lux), RN(PvraSRR1G52T::lux), RN(PvraSRR1G58T/C52T::lux), RN(PvraSRR1�1A::lux), RN(PvraSRR1�2A::lux),RN(PvraSRR2C43A::lux), RN(PvraSRR2C31A::lux), RN(PvraSRR2G25A::lux), RN(PvraSRR2C43A/G38A::lux), RN(PvraSRR2C31A/G25A::lux),RN(PvraSRR3G85T/C79T::lux), and RN(PvraSRR3C73T/G69::lux).

The effect that each lux fusion plasmid had on the growth of S. aureuswas investigated by monitoring the bacterial growth profiles for all thestrains. Briefly, lux fusion strains and RN(::lux) were inoculated into TSBand grown overnight with 5 �g/ml chloramphenicol at 37°C. Next, 1%culture was inoculated in fresh TSB supplemented with 5 �g/ml chloram-phenicol. The OD620 of all cultures was measured at 1-h intervals for thenext 5 h.

Measurement of bioluminescence from S. aureus strains. Afterovernight growth, the wild-type strain RN(PvraSR::lux) and the strainsharboring the lux fusion plasmids were diluted into fresh TSB and cul-tured at 37°C with shaking at 200 rpm. The strains were grown to anOD620 of �0.4 and were subsequently subjected to oxacillin at 10 �g/mlfor 1 h. The OD620 value was recorded for all samples. The cell cultureswith higher OD620 values were diluted to achieve the same cell density ineach cell culture. A 300-�l aliquot from each sample (in triplicate) wastransferred to an opaque 96-well Optiplate and analyzed by FluoroskanAscent FL 2.5 (Thermo LabSystems). Bioluminescence was measured im-mediately after dispensing the samples into the plates. The signal wasmeasured for 1 min over a period of 10 min, with integral measurementfor 100 ms at a 1-mm luminescence height. The data points collected over10 min were averaged for each strain, and the oxacillin concentration andthe standard deviations were determined for each sample from three in-dependent measurements. For statistical evaluation, data obtained fromthree experiments were pooled and analyzed using Student’s t tests. Pvalues of less than 0.05 were regarded as significant.

Viability testing of the cultures. Overnight seed cultures of the wild-type strain, lux fusion strains, and RN(::lux) were inoculated in TSB (at1%) containing 5 �g/ml chloramphenicol. When the OD620 of the cellcultures reached �0.4, oxacillin was added to each culture to a final con-centration of 10 �g/ml, and the cell cultures were grown for an additional1 h. The OD620 value of each culture was normalized to those of thesamples with the lowest density. A 5-�l aliquot from each induced anduninduced cell culture was diluted 50,000-fold in TSB to remove the an-tibiotic. Subsequently, a 25-�l aliquot from each diluted cell culture sam-ple was plated on a TSB/agar plate without antibiotics. Cells were allowedto grow at 37°C. These experiments were repeated three times.

RESULTSBinding of VraR and VraR-P to the target DNA, as assessed byEMSA. The binding curves obtained from the EMSA data werefitted using the equation shown above. The determined dissocia-tion constants represent the apparent values as we are investigat-ing the VraR binding to the vraSR promoter region, which hasbeen proposed (2) to harbor more than one binding site (Table 1).

The Hill coefficient was determined from the experimental datafitting to the equation shown above. For all the DNA sequencesanalyzed, the calculated Hill coefficient was bigger than 1, with anaverage value of 2.2 � 0.5, indicating cooperativity between theputative binding sites. The binding capacity of the DNA (“Cap” inequation shown above) depended on the mutation of the DNAand the phosphorylation state of VraR. Overall, mutation of theR1 site not only had an effect on the binding affinity of the VraR toDNA but also decreased the binding capacity for unphosphory-lated VraR to 60 and 70%. Similar observations on the mutationsin the R2 and R3 sites were made, whereby the binding capacitywas decreased to 80%. In contrast, mutation of the target DNA didnot affect the binding capacity for phosphorylated VraR (see Fig.S1 to S5 in the supplemental material).

Mutation of the R1 site at position C67 or C52 affected thebinding of VraR and VraR-P to PvraSR the most, while mutation ofthe G62 or G58 position at this site had little to no effect on thebinding of unphosphorylated and phosphorylated VraR to PvraSR.The double mutations carried out on the R1a and R1b sites con-firm the significance of the C67 and C52 positions in binding ofVraR and VraR-P to PvraSR. Separation of the R1a and R1b sites byinsertion of one to three base pairs decreased the binding affinitiesof VraR and Vra-P to PvraSR by 2.5 to 3-fold (Table 1).

Mutation of the R2 site had the largest effect on the binding ofVraR-P to PVraSR. This was expected, as we showed earlier thatunphosphorylated VraR does not bind to the R2 site (2). In par-ticular, mutation of the R2a site at the C43 position decreasedbinding affinity of phosphorylated VraR by �3-fold. Addition of asecond mutation at this site (from G38 to A) did not reduce thebinding affinity further. Mutation of the R2b site had slightly lessof an effect on the binding of VraR-P to PvraSR; mutations of C31and G25 decreased the binding affinity of VraR-P by 1.8- and2.4-fold, respectively. Double mutation at the R2a (C43A G38A)and R2b (C31A G25A) sites decreased the binding affinity ofVraR-P to PvraSR by 2.8- and 2.5-fold, respectively. Mutation of theR3 site did not affect binding of VraR to PvraSR, with the exception

TABLE 1 Apparent dissociation constants of VraR and VraR-P, withPvraSR variants determined from the EMSA experimentsa

Variant

Apparent Kd (�M)

VraR VraR-Pb

Wild type 4.1 � 0.5 1.7 � 0.1PvraSRR1C67T 7.1 � 0.5 2.2 � 0.4PvraSRR1C67T/G62T (R1DM3) 9.0 � 0.4 5.9 � 0.5PvraSRR1G58T 3.9 � 0.5 3.6 � 0.1PvraSRR1C52T 6.1 � 0.4 4.4 � 0.3PvraSRR1G58T/C52T (R1DM4) 7.5 � 0.4 5.0 � 0.3PvraSRR1�1A 8.8 � 0.4 4.2 � 0.4PvraSRR1�2A 11 � 1.1 4.8 � 0.4PvraSRR1�3A 12.0 � 0.6 5.4 � 0.2PvraSRR2G38A (R2SM1) 4.2 � 0.2 2.8 � 0.3PvraSRR2C43A 5.4 � 0.3 4.9 � 0.1PvraSRR2C43A/G38A 5.2 � 0.3 4.8 � 0.5PvraSRR2C31A 4.2 � 0.2 3.0 � 0.1PvraSRR2G25A 4.4 � 0.2 4.0 � 0.3PvraSRR2C31A/G25A 5.4 � 0.3 4.3 � 0.6PvraSRR3G85T/C79T 3.8 � 0.3 2.1 � 0.4PvraSRR3C73T/G69T 5.4 � 0.3 3.9 � 0.6a The Hill coefficient was determined to be �1 in all the cases.b The phosphorylation level of VraR is 46% in these experiments.

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of double mutations at the C73 and G69 positions, whereby thebinding affinity was reduced by 2-fold.

DNase I footprinting experiments. DNase I footprinting ex-periments provided a sensitive and direct means to explore furtherthe binding mode of VraR and VraR-P to PvraSR. Mutation of theR1a site (C67T G62T) prevented binding of VraR-P not only tothe R1 site but also to the R2 site (Fig. 2)—these experiments werenot carried out with the unphosphorylated VraR protein, as underthese conditions, VraR does not bind to the R2 site (2). Similarobservations were also made with the PvraSR�1A, PvraSR�2A andPvraSR�3A mutants. Herein, we observed a progressive decrease inthe binding affinity of VraR and VraR-P to the R1 site as thedistance between the R1a and R1b sites increased from 1 to 3 extrabase pairs. The decrease in the binding affinity of VraR-P for theR2 site parallels that for the R1 site. In contrast, mutation in the R2site (R2SM1 mutant) affected to a greater degree the binding ofVraR-P to the R2 site than to R1 (Fig. 2). Overall, these observa-tions indicate that there is communication between these twoVraR binding sites, with R1 being essential for binding of VraR tothe R2 site but not vice versa. The effect that mutation of the R1site has on the binding of VraR to this site as well to the R2 site isobserved for both strands; the DNase I footprinting experimentswith R1DM3 and R2SM1 were carried out by 32P end labeling of

the top strand, and the experiments with PvraSR�1A, PvraSR�2A,and PvraSR�3A were carried out by 32P end labeling of the bottomstrand (Fig. 2 and 3).

In vivo experiments using a lux reporter vector. The effectsthat the mutations of the putative VraR binding sites had on thebinding affinity of VraR for the PvraSR were investigated in vivousing a lux reporter vector under stress conditions induced by 10�g/�l oxacillin. The lux reporter vector enables cloning of thetarget operon control region upstream of the lux operon by put-ting expression of the lux operon under the direct control of thetarget DNA sequence (12). Fusion of PvraSR upstream of the luxoperon resulted in the increase of the luminescent signal of S.aureus RN4220 by �200-fold in the presence of 10 �g/ml oxacillin(see Fig. S7 in the supplemental material). The effects that muta-tions on the R1 and R2 sites had on the luminescence signal of S.aureus RN4220 mirrored those observed by EMSA and DNase Ifootprinting (Fig. 4).

Double mutation of the R1a site (C67T G62T) decreased thelux expression by 9-fold, compared to a 1.4-fold decrease observedfor the double mutation of the R1b site. A similar effect from themutation of the R2 site was measured, whereby we observed a15-fold decrease in the luminescence signal as a result of the dou-ble mutation of the R2a site compared to a 1.2-fold decreasecaused by the mutation of the R2b site. Furthermore, the increaseof the distance between the R1a and R1b sites had the highest effecton the lux operon expression; we measured a 72-fold decrease in

FIG 2 DNase I footprint of VraR-P on the PvraSR R1DM3 and R2SM1 variants.The top strands of PvraSR, PvraSR R1DM3, and PvraSR R2SM1 were end labeledwith [�-32P]ATP. The binding reaction mixtures consisted of VraR-P and 10ng DNA. After treatment with DNase I, the resulting DNA fragments wereseparated on an 8% polyacrylamide sequencing gel containing 7 M urea. Thepositions of the nucleotides, as indicated by the numbers, were calibrated withrespect to the transcription start position. WT, wild type.

FIG 3 DNase I footprints of VraR and VraR-P on the PvraSRR1�1A, -R1�2A,and -R1�3A variants. The bottom strand of each PvraSR variant was end la-beled with [�-32P]ATP and incubated with VraR or VraR-P at 2 or 5 �M. Aftertreatment with DNase I, the resulting DNA fragments were separated on an 8%polyacrylamide sequencing gel containing 7 M urea. The C and G sequencingreactions were calibrated with respect to the transcription start point.

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the luminescence signal for the PvraSR�1A mutant and 102-folddecrease for the PvraSR�2A mutant. In fact, expression of the luxoperon measured for these mutants was below the expression lev-els measured in the case of intact PvraSR and in the absence ofoxacillin (see Fig. S7 in the supplemental material).

In vitro transcription experiments. We used the vraSRoperon control region stretching from nucleotides �462 to �150,referred to as PvraSR

L in the in vitro transcription experiments. Inaddition, taking advantage of the conserved nature of the RNApolymerase (RNAP) among Gram-negative and -positive organ-isms, we used the RNAP from E. coli as a substitute for the RNAPof S. aureus.

Runoff in vitro transcription assays revealed that the E. coliRNAP-�70 complex, referred to as RNAP holoenzyme (RNAPEh),was capable of initiating transcription from PvraSR

L (Fig. 5A). Thedetected transcript product was identified as a 160-nucleotidefragment, which is the expected size because the transcription startposition is located 160 bp from the 3= end of the linear templateDNA used. The same experiments were carried out with the S.aureus SigA factor and E. coli RNAP core enzyme (RNAPEc), andthe results were indistinguishable from those of the experimentswith the RNAPEh (Fig. 5B). In these experiments, the amount oftranscript increased with the increase of the SigA concentration in

FIG 4 In vivo investigation of the role of R1 and R2 in the transcriptionalmechanism of VraR. The bioluminescence signals obtained from S. aureusstrains in the presence of 10 �g/ml oxacillin are depicted. Each strain is repre-sented by the name of the promoter fused upstream of the lux operon in thepXEN1 plasmid. Standard deviations were calculated from three independentexperiments. The viability experiments showed that the effect of oxacillin onthe bacterial growth (number of CFU) was the same for all examined strains(data not shown). RLU, relative light units.

FIG 5 Runoff in vitro transcription assays with E. coli RNAP holoenzyme, using PvraSRL as a template (A), and E. coli RNAP haloenzyme, E. coli RNAP core

enzyme, and S. aureus SigA in the presence of VraR-P, using PvraSRL as a template (B). The SigA concentrations in lanes 2 to 5 are 2, 4, 8, and 8 �M, respectively.

The sizes of the transcription products were determined using the molecular weight marker �X174DNA/HinfI and the 160-bp DNA fragment (lane 1).

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the assay, and removal of the SigA factor or VraR-P from thereaction mixture produced no transcript. Isolation of the SigAfactor was challenging. Hence, we used �70 of E. coli as a substitutefor SigA of S. aureus to investigate VraR interaction with the tran-scription machinery.

The runoff in vitro transcription experiments with RNAPEh

showed that the presence of VraR or VraR-P increased the amountof transcript by 2- and 4-fold, respectively (Fig. 6A). In addition,the in vitro transcription assays exhibited a dose-dependent re-sponse to the VraR-P concentration (Fig. 6B), which reached sat-uration at 6 �M VraR-P. We also investigated the effects of mu-tations in the R1 and R2 sites on the activity of the vraSR operon(Fig. 6C). These studies indicated a clear decrease in the transcrip-tion product when either site was mutated, which is in good agree-ment with the DNase I footprinting experiments (Fig. 2) and invivo studies (Fig. 3).

Specific binding of RNAPEh to PvraSR was confirmed by DNaseI footprinting experiments. These experiments show that RNAPEh

alone binds weakly to PvraSR (Fig. 7). Binding of RNAPEh at the�57 to �15 nucleotide region of PvraSR was associated with theappearance of a hypersensitive site at �24 in both strands and �6in the top strand. The presence of VraR-P enhanced the binding ofthe RNAPE holoenzyme to PvraSR by at least 5-fold, especially inthe region from nucleotides �34 to �24 (Fig. 7A). In addition, thepresence of VraR-P enhanced DNase I cleavage at the �24 site.Furthermore, we investigated binding of SigA to PvraSR by EMSA.These studies showed that SigA alone does not bind to the vraSRpromoter region (Fig. 8A). However, in the presence of VraR and

FIG 6 (A) Role of RNAP haloenzyme and phosphorylation of VraR in the tran-scription of the vraSR operon (the error bars represent the standard deviationscalculated from three independent experiments). (B) Effect of VraR-P concentra-tion on runoff in vitro transcription (error bars represent the standard deviationscalculated from three independent experiments). (C) Role of intact R1 and R2 sitesin transcription of the vraSR operon. The reactions were performed in the pres-ence of VraR-P (6 �M) or VraR (6 �M). The sizes of the transcription productswere determined using the �X174DNA/HinfI molecular weight marker.

FIG 7 DNase I footprint of the RNAP holoenzyme on PvraSR in the presence or absence of VraR or VraR-P. Top strand (A) and bottom strand (B) of32P-end-labeled PvraSR. The binding reaction mixtures included 10 ng DNA and proteins at the indicated concentrations. The regions protected by VraR-P areindicated with solid lines. The regions protected by RNAP are indicated with dashed lines. The region of DNA protected by RNAP in the presence of VraR-P ismarked with a double line. The hypersensitive regions induced by RNAP are indicated by gray arrowheads, and the hypersensitive regions caused by VraR-P areindicated by black arrowheads.

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more so in the presence of VraR-P, the presence of SigA intro-duces a supershift in the EMSA experiments. This could be due tothe binding of SigA to VraR (Fig. 8C).

We performed KMnO4 footprinting experiments to determinethe mechanism by which binding of VraR to PvraSR will benefit thetranscription process. This assay enables investigation of pro-moter melting at the onset of transcription initiation. Potassiumpermanganate reacts specifically with thymine residues in single-stranded DNA regions formed during the formation of the RNAP-open promoter complex (28). We observed that RNAPEh alone isable to form an open promoter complex at PvraSR (Fig. 9; see alsoFig. S8 in the supplemental material). The transition of RNAPEh-PvraSR from a closed promoter complex to an open promoter com-plex was stimulated in the presence of VraR and even more so byVraR-P, in agreement with the in vitro transcription assay results.Mutation of the R1 or R2 site decreased the extent of the promotermelting (Fig. S8).

Determination of the VraR concentration in S. aureusRN4220. Using a monoclonal antibody against VraR, we deter-mined the VraR concentration in S. aureus RN4220 in the absenceor presence of 20 �g/ml oxacillin after 20 and 40 min. These ex-periments showed that concentration of VraR increased from2,830 � 637 molecules (2.3 �M) to 7,265 � 989 molecules (6 �M)and 12,368 � 1097 molecules (10 �M) when exposed to oxacillinfor 20 and 40 min, respectively. The concentration of VraR in theabsence of stress is in agreement with those determined for OmpR(3,500 molecules) (9) and CheY (6,000 molecules) (17). The in-crease in the VraR concentration in our study is in agreement withthe maximum increase in the in vitro transcription product ob-served at 6 �M VraR-P and the reported 3.8-fold increase in the�-galactosidase activity of the PvraSR::lacZ reporter vector when S.aureus was subjected to 1.2 �g/ml oxacillin for 30 min (31).

DISCUSSIONVraR mode of binding to the vraSR operon control region. Re-cently, we showed that VraR binds to its promoter (Fig. 1) (2).Two nucleotides in particular, C and G, were observed to be 4 or 5

residues apart in each VraR binding site. We proposed the puta-tive VraR binding sequence to be ACT(X)nAGT, where n is 2 or 3(2). Each site has two copies of this sequence arranged in tandem.Hence, it was proposed that VraR binds as a dimer to these sites. In

FIG 8 Investigation of SigA binding to PvraSR in the absence of VraR (A), the presence of unphosphorylated VraR (B), and the presence of phosphorylatedVraR (C).

FIG 9 Investigation of the VraR transcriptional activation mechanism byKMnO4 footprinting assays. (A) Formation of the open promoter complex onthe PvraSR, PvraSR R1DM3, and PvraSR R2SM1 sequences in the presence orabsence of VraR or VraR-P. Briefly, each 32P-end-labeled DNA sequence wasincubated with the proteins, and the formed complexes were treated with 20mM KMnO4 for 30 s, after which the modified thymine residues were cleavedby 1 M piperidine at 90°C for 30 min. The samples were loaded onto an 8%sequencing gel.

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this study, we investigated the mode of VraR binding to PvraSR bymutating the C or G residues in these sites to T or A singly or incombination and increasing the distance between the two tandemVraR binding sequences by 1, 2, or 3 bp. The mutations werecarried out in the context of full-length PvraSR (nucleotides �126to �10).

Mutation of C or G at either R1 or R2, but not R3, affectedbinding of VraR to DNA either in vitro or in vivo (Table 1 and Fig.2 and 3; see also Fig. S1 to S4 in the supplemental material). Theseobservations demonstrate that the R1 and R2 sites are the VraRbinding sites in PvraSR. Further, the mutagenesis studies showedthat mutation of C67 (R1a) or C52 (R1b) had the strongest effecton the binding of VraR and VraR-P to PvraSR compared to that ofG62 (R1a) and G58 (R1b). These results suggest that C67 and C52are important for VraR binding to each subsite in R1. They alsoindicate that there is directionality in VraR binding to R1, wherebyVraR binds to DNA in a C-to-G direction (Fig. 1).

Double mutations of C and G in each subsite of R1 affectedVraR binding differently, with C67T G62T mutation causing thehighest decrease in the VraR and VraR-P binding affinities toPvraSR (Table 1). This observation suggests that a hierarchy existsin VraR binding within the R1 site itself, whereby VraR binds firstto the R1a site. The in vitro findings are corroborated by in vivostudies (Fig. 4), which showed that mutation in the R1a site had agreater effect on lux expression (9-fold decrease) than mutation inthe R1b site (1.4-fold decrease). Hierarchical binding was alsoobserved in the R2 site. A single point mutation in the R2a site,C43A, decreased lux expression by 15-fold, as opposed to the 1.2-fold decrease measured with other mutants in this site (Fig. 4). Asdetermined by looking carefully at the R1 and R2 sites, the C andG are separated by 4 bp in the R1a and R2a subsites versus 5 bp inthe R1b and R2b subsites. It is possible that the difference in dis-tance between C and G in each subsite could be the determiningfactor for the observed difference in the binding affinity to VraR.

The observed hierarchy in VraR binding to PvraSR suggests thatoccupation of each VraR binding site in PvraSR could be a two-stepevent (i.e., one molecule of VraR binds first to the Ra subsite, andthen a second molecule of VraR binds to the Rb subsite). Theadvantage of this binding model becomes apparent when consid-ering the close proximity of R1a to R1b (they are separated by asingle base pair) and the directionality in binding imposed bycytosines. Both of these factors could drive the dimerization ofVraR without the need for phosphorylation in the R1 site, which isin agreement with the earlier observations that phosphorylationof VraR did not have a significant effect on the binding of VraR tothe R1 site alone [Kd(VraR)R1 � 4.0 � 0.4 �M; Kd(VraR-P)R1 �2.7 � 0.9 �M] (2). This in turn suggests that unphosphorylatedVraR binds to its promoter and mediates its expression in theabsence of cell wall stress. Our hypothesis is in agreement with themeasured vraSR background expression in the absence of stress(22).

To test the hypothesis that VraR binding to R1 is phosphory-lation independent (i.e., dimerization is driven by close proximityof R1a and R1b), we generated PvraSR mutants in which the dis-tance between the R1a and R1b subsites was increased by insertionof 1, 2, or 3 bp (e.g., ACTAAAGTaaaTGAACATCA; lowercaseletters indicate the inserted nucleotides, and boldface letters indi-cate the nucleotides involved in binding VraR) and R2 was leftintact. The in vivo studies (Fig. 4) indicated that 1- and 2-bp in-sertions between the R1a and R1b subsites had a dramatic effect on

lux expression. We measured 72- and 102-fold decreases in biolu-minescence for PvraSRR1�1A and PvraSRR1�2A, respectively. Inaddition, the background expression level (uninduced strain) wasaffected by these mutations (see Fig. S7 in the supplemental ma-terial). Interestingly, phosphorylation rescued the binding ofVraR to the R1 site by 50% in the case of a single-base-pair inser-tion (Fig. 3) but not in the case of 2- and 3-bp insertions. Thisobservation suggests that a preformed VraR dimer (through phos-phorylation) is able to overcome the increased distance betweenR1a and R1b. In addition, it supports our hypothesis that the closeproximity of these two subsites induces dimerization of unphos-phorylated VraR to R1.

Investigation of PvraSR mutants by DNase I footprinting (Fig. 2and 3) consistently showed that mutation of the R1 site had agreater effect on the binding of VraR to the R2 site than mutationof the R2 site on VraR binding to the R1 site. These observationsindicate that there is strong communication between the R1 andR2 sites in PvraSR and that an intact R1 site is essential for bindingof VraR to the R2 site but not vice versa.

Overall, our investigation of the PvraSR mutants showed thatthe VraR binding site is composed of two tandem arrayedsequences. In the R1 site, the orientation and distance of thesetwo sequences mediates VraR dimerization in a phos-phorylation-independent manner. The observed higher bind-ing affinity of VraR for this site could also serve as a means ofrecruitment of VraR to the less conserved binding site, R2 (i.e.,the presence of VraR bound to the R1 site could serve as a “flag”for the phosphorylated VraR to bind to a site that is less recogniz-able due to its lack of conserved sequence). The implication ofthese findings is as follows. In the absence of stress, the DNAsequence of the vraSR promoter region facilitates binding of un-phosphorylated VraR to enable low-level expression of the vraSRoperon. This event does not require phosphorylation, asdimerization is DNA assisted. In the presence of cell wall stress, thealready bound VraR will serve as the recognition point for phos-phorylated (and dimerized) VraR to bind to the less conserved R2site of the promoter and mediate higher-level expression of thevraSR operon.

Bridge between VraR and the transcription machinery: roleof the SigA factor. It has been generally accepted that VraR regu-lates expression of its own operon. The in vitro transcription stud-ies were used to explore the mechanism through which VraR af-fects the transcription machinery and identify the factor(s)involved in regulation of the vraSR operon.

We used the RNA polymerase (RNAP) from E. coli as a substi-tute for the RNAP of S. aureus, taking advantage of the conservednature of the RNA polymerase among Gram-negative and-positive organisms. Several factors, listed below, suggest that thesigma factor required for the regulation of vraSR operon is similarto E. coli �70. First, the �10 box in PvraSR (Fig. 1), represented by5=-TTATAA-3=, is highly similar to the conserved �10 motif re-ported for E. coli �70 (5=-TATAAT-3=) (8). Second, the putative�35 box (5=-TTCATA-3=) shares significant sequence similaritieswith the �35 motif recognized by E. coli �70 (5=-TTGACA-3=) (8,24). Third, the �10 and �35 boxes in PvraSR are separated by 17nucleotides, a feature that is shared by the E. coli �70-regulatedpromoters (33).

Investigation of the S. aureus genome indicated that this organ-ism has three sigma factors, the primary sigma factor (SigA), thealternate sigma factor (SigB), and SigH, involved in the regulation

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of genetic competence genes (10, 20, 26, 34). Sequence alignmentsof SigA, E. coli �70, and Bacillus subtilis SigA indicate that theyshare large sequence similarities, especially in the two regions im-plicated in anchoring the sigma factor to the DNA and docking itto the polymerase core-binding domain (see Fig. S6 in the supple-mental material). Based on this evidence, we propose that SigA isinvolved in regulation of the vraSR operon. Additional evidencefor this hypothesis comes from the work of Galbusera et al. andBoyle-Vavra et al., which showed that the inactivation of SigB doesnot affect the activity of the vraSR operon (6, 13).

The runoff in vitro transcription assays revealed that the E. coliRNAP holoenzyme (RNAPEh) was capable of initiating transcrip-tion from PvraSR

L (Fig. 5A). Substitution of the SigA factor of S.aureus for the E. coli �70 factor in the RNAP haloenzyme producedindistinguishable results (Fig. 5B). Furthermore, formation of thetranscript product was sensitive to the presence of SigA andVraR-P and the SigA concentration; removal of the SigA factor orVraR-P from the reaction mixture produced no transcript. Theformation of the transcript (in the presence of RNAPEh) also de-pended on the VraR and VraR-P concentrations, reaching a max-imum at 6 �M VraR-P. Incidentally, the in vivo concentration ofVraR in the presence of cell wall stress is 6 �M. These resultsstrongly indicate formation of a specific complex between the sur-rogate RNAP haloenzyme (RNAPEc-SigA) and the PvraSR pro-moter in the presence of VraR; hence, SigA is likely to be involvedin regulation of the vraSR operon. This also serves as evidence thatSigA and E. coli �70 are functionally indistinguishable; hence, theRNAP haloenzyme from E. coli (in complex with �70) is a goodsubstitute for the S. aureus RNAP haloenzyme (in complex withSigA).

The specificity of the RNAPEh interaction with PvraSR was fur-ther corroborated by the DNase I footprinting experiments.RNAPEh alone bound weakly to PvraSR (Fig. 7); however, the pres-ence of VraR-P enhanced the binding of RNAPEh to PvraSR by5-fold, especially in the region encompassing nucleotides �34 to�24. The enhanced interaction of RNAPEh with PvraSR could re-sult from a direct interaction of VraR-P with SigA. Investigation ofthe SigA binding to PvraSR in the presence and absence of VraR-Pindicates that SigA may interact with VraR (Fig. 8). Furthermore,the KMnO4 footprinting experiments revealed that the transitionof RNAPEh-PvraSR from a closed promoter complex to an openpromoter complex was stimulated in the presence of VraR, andeven more so by VraR-P, in agreement with the in vitro transcrip-tion assay results. This stimulation could happen as a result of theestablishment of additional contacts for the RNAP holoenzymeon the promoter through the interaction of VraR-P with SigA. Asimilar role has been suggested for BvgA polymerase binding toptx and fha promoters in Bordetella pertussis (4).

Transcription factors are known to interact with RNAP ho-loenzyme (18). Based on the type of interactions in which they areinvolved, these factors are divided into two classes. Class I factors,such as BvgA (5), Sp0A (30), and OmpR (27), interact with theC-terminal domain of the � subunit of RNAP (�CTD). Class IIfactors, such as PhoB (23) and VanRB (11), have been shown tointeract with the � subunit of RNAP. In this context, it is possiblethat VraR is a class II factor. The overlap between the VraR andSigA binding sites in the vraSR operon control region certainlysupports this hypothesis. Nevertheless, an interaction with theother parts of RNAP (e.g., �CTD) cannot be excluded.

In conclusion, our study demonstrates that VraR binds to two

regions centered at nucleotides �60 (R1) and �35 (R2) in thevraSR operon control region. There is a hierarchy in VraR bindingto these sites. Specifically, the high-affinity and phosphorylation-independent binding site R1 serves to recruit VraR to the vraSRoperon control region under normal growth conditions to medi-ate basal expression of the operon. On the other hand, cell wallantibiotic stress drives binding of phosphorylated VraR to thelow-affinity and phosphorylation-dependent binding site R2,which results in upregulation of the vraSR operon. The phosphor-ylation independence of VraR binding to R1 is driven by the closeproximity of two VraR binding sequences at this site, whereas thedistance between the VraR binding sequences in R2 ensures thatVraR binds to this site only under stress conditions.

The position of the R2 site is quite strategic in PvraSR, as itoverlaps with that of the SigA factor. Our study reveals thatVraR-P mediates tight binding of the RNAP holoenzyme to PvraSR

and enhances the formation of the RNAP open promoter com-plex. It is likely that VraR interacts intimately with the RNAPholoenzyme to extend and stabilize the RNAP contacts with DNAand/or to induce topological changes (e.g., DNA bending) in thepromoter region. Hence, the sequence and location of R2 enable S.aureus to fine-tune the vraSR operon activity under cell wall stress.

ACKNOWLEDGMENTS

We are in debt to David Heinrichs (Western University) for providing thepXEN1 plasmid.

This work was supported by a Discovery Grant to D.G.-K. from theNatural Sciences and Engineering Research Council of Canada and anEarly Researcher Award from the Ontario Ministry of Research andInnovation.

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