Regulation of ciaXRH Operon Expression and Identification of the CiaR Regulon in Streptococcus mutans

ABSTRACT The ciaRH operon in Streptococcus mutans contains 3 contiguous genes, ciaXRH. Unlike the CiaRH system in other streptococci, only the ciaH-null mutant displays defective phenotypes, while the ciaR-null mutant behaves like the wild type. The objective of this study was to determine the mechanism of this unusual property. We demonstrate that the ciaH mutation caused a >20-fold increase in ciaR transcript synthesis. A ciaRH double deletion reversed the ciaH phenotype, suggesting that overexpressed ciaR might be responsible for the observed ciaH phenotypes. When ciaR was forced to be overexpressed by a transcriptional fusion to the ldh promoter in the wild-type background, the same ciaH phenotypes were restored, confirming the involvement of overexpressed ciaR in the ciaH phenotypes. The ciaH mutation and ciaR overexpression also caused transcriptional alterations in 100 genes, with 15 genes upregulated >5-fold. Bioinformatics analysis identified a putative CiaR regulon consisting of 8 genes/operons, including the ciaXRH operon itself, all of which were upregulated. In vitro footprinting on 4 of the 8 promoters revealed a protected region of 26 to 28 bp encompassing two direct repeats, NTTAAG-n5-WTTAAG, 10 bp upstream of the −10 region, indicating direct binding of the CiaR protein to these promoters. Taken together, we conclude that overexpressed CiaR, as a result of either ciaH deletion or forced expression from a constitutive promoter, is a mediator in the CiaH-regulated phenotypes.

Bacterial two-component signal transduction systems (TCS) play important roles in bacterial environmental adaptation, production of virulence factors, self-defense, and biofilm formation (5-8, 11, 17, 26). A typical TCS consists of a membrane-bound histidine kinase (HK) sensor and a cytoplasmic response regulator (RR). Upon receiving a signal, the HK undergoes autophosphorylation. The phosphorylated kinase then transfers the phosphate group to its cognate RR, which then activates or represses its target genes by binding to their promoter regions (24).
The sequenced oral pathogen Streptococcus mutans UA159 contains 14 pairs of TCS and one orphan response regulator (3,16). Of these, the CiaRH TCS has been shown to be a global regulator for multiple stress responses such as biofilm formation, acid tolerance, bacteriocin production, and genetic competence (1,3,22). CiaRH is also widely distributed among other streptococcal species and has been shown to play an essential role in stress resistance and pathogenesis (4,12,23). Recently, studies from our group demonstrated that the S. mutans CiaRH TCS is unique among the CiaRH systems in other streptococci in that it actually contains a third compo-nent, CiaX, which is encoded by the first gene of the operon (10). We further showed that the ciaXRH operon expression is autoregulatory and that the operon expression is repressed by calcium through CiaX (10). Interestingly, when the activity of a ciaH-luc (luciferase) reporter was measured, deletion of ciaH affected only the calcium-dependent repression of operon expression, rather than abolishing the regulation of the operon altogether. This suggested that the ciaXRH operon is likely to be regulated differently from the typical autoregulatory TCS.
Another unique feature of the S. mutans CiaRH TCS is the lack of phenotypes associated with a ciaR mutation. Currently, only mutations in the ciaH sensor kinase have been demonstrated to affect multiple cellular functions (1,22). Based on these findings, it was suggested that CiaR functions independently of the CiaH signaling cascade, possibly as a result of cross talk between the Cia system and another TCS (1). In this study, we sought to explain this unusual finding by examining the regulatory mechanism employed by the S. mutans CiaRH system. We show that CiaR is in fact in the same signaling pathway as CiaH; however, unlike a typical TCS, deletion of ciaH causes overexpression of ciaR. Overexpressed CiaR then acts as a positive regulator for the expression of the ciaXRH operon as well as a variety of other downstream targets and a negative regulator for late competence genes and competenceassociated bacteriocin-like genes (14). Through microarray and bioinformatic analyses, a CiaR regulon was identified, consisting of at least 8 genes/operons that are all positively regulated by CiaR. In vitro DNA footprinting confirmed the putative CiaR binding site identified through bioinformatics analysis. From these results, we were able to construct a unique regulatory scheme of the S. mutans CiaRH TCS.

MATERIALS AND METHODS
Bacterial strains, plasmid, and growth conditions. Escherichia coli DH5␣ was grown in Luria broth, and S. mutans UA140 and its derivatives were grown in brain heart infusion (BHI) broth (Difco). For the selection of antibiotic-resistant colonies after genetic transformation, erythromycin (300 g ml Ϫ1 for E. coli or 10 g ml Ϫ1 for S. mutans) or spectinomycin (100 g ml Ϫ1 for E. coli or 1 mg ml Ϫ1 for S. mutans) was added to the medium. Plasmid pDL278 (15), an E. coli-Streptococcus shuttle vector carrying a spectinomycin resistance (spc) gene, was used to overexpress ciaR. For the preparation of cells used in the microarray, overnight cultures of wild-type (WT) and mutant strains were diluted 1:30 in fresh BHI medium containing 0.4% bovine serum albumin (BSA). The cells were grown as a static culture at 37°C with 5% CO 2 and collected by centrifugation when the optical density at 600 nm (OD 600 ) of the culture reached 0.3. Cell pellets were stored at Ϫ80°C until use.
Construction of mutant strains. The S. mutans UA140 ciaH mutant was constructed previously (10). The construction of the ciaRH deletion is described as follows. An approximately 600-bp fragment upstream of ciaR and a 350-bp fragment downstream of ciaH were amplified by PCR using primer pairs ciaRHup-F and ciaRH-up-R and ciaRH-dn-F and ciaRH-dn-R ( Table 1). The upstream and downstream fragments were digested with XbaI and XhoI, respectively, and ligated to a kanamycin resistance cassette released from the vector pBS-kan digested with XbaI and XhoI. The ligation mixture was directly transformed into S. mutans UA140, and transformants were selected on kanamycin plates and confirmed by PCR. To construct the ciaR overexpression mutant, the lactate dehydrogenase (ldh) promoter (19) was used to drive the expression of ciaR. The promoter region of ldh was amplified with primers ldh-F and ldh-R (Table 1), and the full-length ciaR was PCR amplified with primers ciaR-F and ciaR-R (Table 1). Next, the ldh promoter fragment was digested with SacI and BamHI, the ciaR fragment was digested with BamHI and HindIII, and the shuttle plasmid pDL278 was digested with SacI and HindIII. After gel purification, all three fragments were mixed and ligated. The ligation mixture was transformed into E. coli, and the correct recombinant plasmid, pDL-OEciaR, was confirmed by restriction enzyme digestion and sequencing. pDL-OEciaR was then transformed into S. mutans UA140, and the transformants were selected on spectinomycin plates and confirmed by real-time PCR analysis of the ciaR gene expression.
Transformation assay. Genetic competence was determined by a transformation efficiency assay with transforming genomic DNA. Overnight cultures of S. mutans strains were diluted 1:20 in BHI medium containing horse serum (0.4%, vol/vol) and grown to an optical density at 600 nm of ϳ0.3. A 0.3-ml aliquot of the culture was distributed into Eppendorf tubes, and genomic DNA (10 g ml Ϫ1 ) isolated from a UA140 derivative strain carrying a tetracycline resistance marker was added to each culture. After 2 h at 37°C, the cultures were briefly sonicated (Misonix) to break the cell chains and plated on antibiotic-containing BHI agar plates, as well as on nonselective BHI plates. Transformation efficiency was determined after 48 h of incubation and expressed as the percentage of transformants among the total viable recipient cells.
Microarray analysis. (i) RNA extraction. Total RNA was extracted using the FastPrep system (MPBio). Frozen samples were thawed on ice and resuspended in 1.0 ml Trizol. The cell suspension was then transferred to a tube containing prechilled Lysing Matrix B. Cells were subjected to two homogenizations (at a speed of 6.0 M/s; time, 30 s) at a 5-min interval. The mixture was centrifuged at 13,000 ϫ g for 10 min, and the supernatant was transferred to a new 2.0-ml tube. The supernatant was extracted with 0.2 ml chloroform, and the aqueous phase was transferred to a new 2.0-ml tube. RNA was purified using the RiboPure-Bacteria kit (Ambion) following the manufacturer's instructions. After elution, DNase I was added to the samples and incubated at 37°C for 30 min. The samples were further purified with Qiagen RNeasy MinElute spin columns according to the manufacturer's protocol and eluted in RNase-free water. The resulting RNA integrity was confirmed by the presence of clearly defined rRNA bands on agarose gels and later quantified for concentration by OD 260 readings.
(ii) Microarray. For cDNA synthesis, 15 g of total RNA was mixed with 1.25 g random hexamers and heated at 70°C for 10 min for denaturation before being added to the reverse transcription reaction mixture. Reverse transcription was performed in a total volume of 60 l, and the reverse transcription reaction mixture contained 12 l 5ϫ Superscript II buffer (Invitrogen, Carlsbad, CA), 6 l 100 mM dithiothreitol (DTT), 3 l 10 mM deoxynucleoside triphosphate (dNTP), 1.5 l RNaseOUT RNase inhibitor, and 7.5 l Superscript II reverse transcriptase. The reaction mixture was incubated at 42°C for 60 min, and then the reaction was terminated by incubation of the mixture at 70°C for 10 min. RNA in the mixture was then hydrolyzed by adding 15 l 1 N NaOH and incubating the mixture at 65°C for 30 min. The solution was neutralized with 1 N HCl, and the cDNA was then purified using a Qiagen MinElute PCR purification kit. The concentration of cDNA was determined by measuring the absorbance at 260 nm, and 3 g of cDNA was fragmented in a 35-l reaction mixture containing 3.5 l Roche 10ϫ DNase buffer and 7.5 l diluted Roche DNase (0.06 U/l). The fragmentation reaction mixtures were prepared on ice and then incubated at 37°C for 10 min before the reaction was terminated by incubation of the mixture at 98°C for 10 min. For hybridization, 32 l of the fragmented cDNA was labeled with the BioArray terminal labeling kit (Enzo, New York, NY) in a total volume of 50 l containing 10 l 5ϫ buffer, 5 l 10ϫ CoCl 2 , and 2 l terminal deoxynucleotide transferase. The reaction mixture was incubated at 37°C for 60 min. Labeled cDNA was hybridized to a standard 49-format custom GeneChip microarray (Affymetrix, Santa Clara, CA) containing each of the predicted open reading frames (ORFs) of S. mutans UA159 as well as both strands of the intergenic regions. The hybridization and washing procedures were performed similarly as recommended in the GeneChip Expression Analysis Technical Manual and as previously described (2). The GeneChips were scanned at 570 nM using an Affymetrix 7G laser scanner.
(iii) Data analysis. Analysis of signal intensities was performed using the GeneChip operating system software (GCOS), version 1.4, and gene expression data were compared using the GCOS batch analysis function. Normalization procedures were performed directly by the software using a script designed by Affymetrix and provided with the S. mutans custom array. Four data sets were generated for each ciaH mutant and ciaR overexpression strain. The 4 data sets were compared by using the Excel program, and the t test was used to generate the P values with gyrA as a reference. A gene expression change of Ն2-fold and a P value of Ͻ0.05 were used as cutoff values to generate the gene list presented in Table 2.
Real-time RT-PCR. Real-time reverse transcriptase PCR (RT-PCR) was performed to validate the results generated from the microarray analysis. Primers were designed using Applied Biosystems Primer Express 3.0 software, which scans DNA sequences for primers suitable for threshold cycle (⌬⌬C T ) analysis. The primer sequences are listed in Table 1. Cells were cultured under the same conditions as those described for the microarray. Three hundred nanograms of total RNA was reverse transcribed using the manufacturer's protocol for Affinityscript reverse transcriptase (Stratagene, La Jolla, CA). Real-time PCR was performed using an Applied Biosystems 7300 PCR system, and the reaction mixtures were prepared using Applied Biosystems SYBR Green PCR Master Mix. Changes in gene expression were calculated automatically in the Applied Biosystems 7300 System software using the ⌬⌬C T method and are briefly described as follows: ⌬C T ϭ C T (target) Ϫ C T (housekeeping gene); ⌬⌬C T ϭ ⌬C T1 Ϫ ⌬C T2 ; fold changes are calculated as 2 Ϫ⌬⌬CT . The gyrA gene was used as the housekeeping gene reference, and all samples included a no-RT control to assess genomic DNA contamination in the reactions.
Identification of transcription start site by 5 RACE. To locate the transcription start site of the ciaX transcript, 5Ј random amplification of cDNA end (5Ј RACE) experiments were performed using the FirstChoice RLM RACE kit (Ambion) according to the manufacturer's protocol. For the first-round PCR, the 5Ј RACE outer primer and the gene-specific outer reverse primers (Table 1) were used with the RT reaction products as the template. PCR mixtures from the first-round PCR were then subjected to a second round of PCR amplification using the 5Ј RACE inner primer and the gene-specific inner reverse primers ( Table 1). The resulting PCR products were then cloned into the pGEM-T Easy vector (Promega) and sequenced using M13 primers.
Cloning and purification of CiaR. The ciaR coding sequence was amplified by PCR using Herculase DNA polymerase (Stratagene) and chromosomal DNA derived from S. mutans UA159 with gene-specific primers oSG426 and oSG427 (Table 1). Subsequently, the amplicons were digested with BamHI and EcoRI and ligated into the expression vector pCRT7/NT (Invitrogen) to generate a fusion protein with an enterokinase cleavage site, Xpress epitope, and an aminoterminal 6-histidine tag. The resulting plasmid was introduced into E. coli strain Top10, selected for resistance to ampicillin, and sequenced.
For overexpression, the plasmids containing His-CiaR were transformed into E. coli BL21(DE3)pLysS cells (Invitrogen) and grown in 1 liter of LB with 100 g/ml ampicillin at 37°C with shaking. When the cells reached an OD 600 of 0.3 to 0.5, expression of the fusion protein was induced with 0.3 mM IPTG (isopropyl-␤-D-thiogalactopyranoside) for 3 h. The cells were harvested by centrifugation (5,000 ϫ g, 15 min, 4°C), frozen at Ϫ20°C overnight, defrosted, and resuspended in column buffer (50 mM NaH 2 PO 4 , 0.5 M NaCl, pH 8.0) before 1 mg/ml of lysozyme was added. After incubation on ice for 30 min, 0.5% Sarkosyl was 4670 WU ET AL. J. BACTERIOL.
added, cells were lysed by sonication and any unlysed cells along with other debris were removed by centrifugation (10,000 ϫ g, 20 min, 4°C). The cleared cell lysate was diluted in column buffer to a final Sarkosyl concentration of 0.05%, applied to a 2 ml nickel-nitrilotriacetic acid (Ni-NTA) column bed, preequilibrated with column buffer, and washed with 12 column volumes of column buffer with 50 mM imidazole. The protein was eluted with column buffer containing 500 mM imidazole, and 1-ml fractions were collected. Fractions containing a His-CiaR concentration were determined using the Bio-Rad protein assay (Bio-Rad) using bovine serum albumin as the standard. The purified proteins were stored in 25% glycerol at Ϫ80°C until needed. DNase I footprinting assays. Primers oSG570, oSG572, oSG574, oSG576, and oSG590 (Table 1) were end labeled by incubating 1 M primer with 1 M [␥-32 P]ATP, T4 polynucleotide kinase (Promega), and 1ϫ polynucleotide kinase buffer (Promega) at 37°C for 30 min. Labeled oSG570 primer and unlabeled oSG571 primer were then used to amplify a 131-bp SMU.739 promoter fragment. Labeled oSG572 primer and unlabeled oSG573 primer were used to amplify a 130-bp SMU.239 promoter fragment. Labeled primer oSG574 and unlabeled primer oSG575 were used to amplify a 139-bp SMU.139 promoter fragment. Labeled primer oSG576 and unlabeled primer oSG577 were used to amplify a 131-bp ciaX promoter fragment. As a negative control, the putative binding site of ciaX was randomized with mutations in both direct repeat I (DRI) and DRII. The template DNA for the negative control was generated by subjecting oSG597 and oSG598 (Invitrogen) to a single PCR cycle. These oligonucleotides are 85 bp long, incorporate the mutations in DRI and DRII, and are complementary at the last 30 bp of the 3Ј ends. This double-stranded DNA was used as a template for PCR with oSG576 and oSG577 to amplify a 131-bp fragment. The scrambled ciaX promoter was chosen as the negative control because its wild-type form contains a binding site that most closely matches the putative consensus sequence. The labeled DNA for each substrate was also used as the template for sequencing using the SequiTherm Excel II (Epicentre Biotechnologies) DNA sequencing kit.
For the footprinting assays, each labeled fragment was incubated at room temperature for 30 min with 0 to 2,000 nM His-CiaR in the reaction buffer (25 mM Tris-HCl, pH 7.5, 25 mM KCl, 6.25 mM MgCl 2 , and 10% glycerol) in a reaction volume of 50 l. Fifty microliters of a 5 mM CaCl 2 -10 mM MgCl 2 solution was added to the reaction mixture prior to the DNase I treatment, for 10 min at room temperature. Each substrate was digested with 0.5 units of DNase I (Promega) for 1 min at room temperature. The digestion was terminated by adding 90 l of the stop buffer (200 mM NaCl, 30 mM EDTA, 1% SDS, and 100 g/ml salmon sperm DNA) followed by a phenol chloroform extraction. The   All data were calculated based on 4 data sets generated from independent cultures. A cutoff of Ն2-fold change and a P value of Յ0.05 were used to generate the data set. Entries in bold are those that did not meet the criteria but were included as a comparison with the other data set. All genes presented have hybridization signals significantly higher than the noise (P Յ 0.0001) and are designated as P (present) by the microarray data analysis software GCOS, version 1.4. Numbers in parentheses are ratios obtained by RT-PCR. CoA, coenzyme A; PTS, phosphotransferase. VOL. 192, 2010 CiaRH REGULATION IN S. MUTANS 4673

RESULTS
Deletion of ciaR suppressed the ciaH phenotype. During our previous studies of S. mutans strain UA140, we noticed that a mutation of ciaH resulted in defects in competence, acid tolerance, and bacteriocin production, while a ciaR mutant behaved similarly to the wild type (22). The same phenomenon was subsequently reported in the strain UA159 as well (1). This unexpected result was suggested to arise from cross talk between different TCS (1). To examine this further, we made a ciaRH double mutation and tested its phenotype. As shown in Fig. 1, the ciaRH double mutation reversed the competence deficiency of the ciaH mutant, resulting in a transformation efficiency similar to that of the wild type. The double mutation also suppressed the mutacin I bacteriocin deficiency and acid tolerance phenotype of the ciaH mutant (reference 22 and data not shown). These data suggested that ciaR is likely to be in the same signaling pathway as ciaH; however, unlike a typical twocomponent system, deletion of ciaR alone does not affect the normal function of the cell.
Deletion of ciaH caused overexpression of ciaR. Based on the data presented above, we hypothesized that ciaR may function as a repressor for competence development, mutacin production, and acid tolerance. Interestingly, our previous studies showed that the ciaXRH operon is autoregulatory; however, unlike other autoregulatory TCS operons, deletion of ciaH abolished only its transcriptional repression by calcium (10). Thus, we reasoned that the ciaXRH operon is likely regulated by an uncharacterized mechanism, which may help explain why mutations of ciaH and ciaR did not result in similar phenotypes. To identify the mechanism, we used real-time RT-PCR to quantify the ciaR transcript in the ciaH mutant background. To our surprise, the ciaH mutation increased ciaR gene expression ϳ25-fold (Fig. 2). This result suggested that (i) CiaH is a negative regulator of ciaR and (ii) overexpressed ciaR might be responsible for the observed phenotypes of the ciaH mutant.
Overexpression of ciaR in the wild-type background resulted in the same phenotype as that of the ciaH mutant. To further test whether overexpression of ciaR itself is sufficient to cause the functional changes observed in the ciaH mutant, we cloned the ciaR gene under the control of the constitutively expressed ldh promoter on the shuttle plasmid pDL278. The ciaR overexpression strain was tested for transformability, mutacin pro-duction, and acid tolerance. As shown in Fig. 1, the transformation efficiency in the ciaR overexpression strain was not only reduced compared to that of the wild type but also 2 times lower than that of the ciaH mutant strain. This is probably due to the higher ciaR gene expression in the ciaR-overexpressing strain (2-fold) than in the ciaH mutant strain (data not shown). Similarly, mutacin production was completely abolished in the ciaR overexpression strain, and acid tolerance in the ciaR overexpression strain was even weaker than that in the ciaH mutant strain (data not shown), possibly for the same reason as stated above. As expected, overexpressing ciaR in the ciaRH double mutation background resulted in the same phenotypes (data not shown). Taken together, these results suggested that overexpressed ciaR is an essential mediator of ciaH mutant phenotypes.
The ciaH deletion and ciaR overexpression affected the same set of genes. To further determine the mechanism by which CiaR mediates gene regulation in the CiaRH signaling pathway, we performed microarray analysis of the wild-type, ciaH mutant, and ciaR overexpression strains grown to an OD 600 of 0.3 when natural competence is at its peak level (20). From 4 data sets of each mutant and a cutoff of Ն2-fold expression change and a P value of Յ0.05, a list of 100 genes was generated (Table 2). Among these, 45 genes were upregulated and 55 were downregulated in the ciaH mutant and ciaR overexpression backgrounds compared with the wild type. Comparing the ciaH data set with the ciaR overexpression data set revealed nearly identical gene lists, except for 4 genes, SMU.225c, SMU.528c, SMU.539c, and SMU.574c, which showed a 2-to 3-fold downregulation in the ciaR overexpression strain but no change in the ciaH mutant strain. Three of the four genes also showed a large standard deviation and P values in the ciaH data set, making the assignment difficult to ascertain. In addition, 36 genes did not meet the cutoff in one data set or the other, but the trend of change is consistent in the two data sets.
To confirm the microarray results, real-time RT-PCR was performed to quantify relative gene expression for a subset of the genes in the microarray data set. RNA was isolated from independent cultures grown under the same conditions as in the microarray, and gene expression was measured in both the ciaH deletion and the ciaR overexpression backgrounds. All tested genes showed a trend of expression consistent with that in the microarray data sets (numbers in parentheses in Table 2), suggesting that the trends in gene expression as measured by microarray are likely to be reflective of the results obtained by RT-PCR.

Identification of a CiaR regulon.
To further investigate the regulatory function of CiaR, we sought to identify the genes directly regulated by CiaR. During previous studies, we identified a putative CiaR binding site upstream of ciaX, based upon the CiaR binding site consensus identified in Streptococcus pneumoniae (9). The consensus sequence is an NTTAAG-n5-WTTAAG direct repeat that is located ϳ10 bp upstream of the Ϫ10 region. Therefore, MotifSearch was used to scan the S. mutans genome for the same sequence pattern. After a list of "hits" was generated, each sequence was manually analyzed for its location (upstream of an open reading frame [ORF]) and the presence of a putative Ϫ10 sequence ϳ10 bp downstream of the second direct repeat (DRII) WTTAAG motif. From these analyses, a total of 8 promoters were identified, including the ciaX promoter (Fig. 3A). The promoters for SMU.40, SMU.139, SMU.648, SMU.739, SMU.1093, SMU.2164, and SMU.1131c (ciaX) have a perfect match to the consensus sequence, and all contain a fairly strong Ϫ10 sequence 9 to 10 bp downstream of DRII. The promoter for SMU.239 is almost identical but has a 1-bp deletion in the spacer. SMU.40, SMU.139, SMU.239, SMU.739, SMU.1093, and ciaX are all highly upregulated in both the ciaH mutant and ciaR overexpression strains (18-to 95-fold, Table 2). In contrast, the expression of SMU.2164 was only moderately upregulated (2-to 3-fold), while the expression of SMU.648 was barely affected. Inspection of the genomic organization of SMU.648 revealed that while it may have its own promoter, its transcription might also be influenced by read-through from the upstream gene SMU.647.
DNA footprint of the CiaR regulon promoters. To further confirm that CiaR indeed binds to the putative CiaR binding site of the CiaR regulon promoters identified by bioinformatics, DNase I footprinting assays were performed on selected promoters (ciaX, SMU.139, SMU.239, and SMU.739) using purified CiaR protein. As shown in Fig. 3B to E, CiaR clearly binds to each of the aforementioned promoter regions. In each case a 26to 28-bp region was protected with the consensus sequence located nearly in the middle of the protected region. More importantly, scrambling the consensus CiaR binding site in ciaX from the wild-type sequence (ATTAAGTCTCTTTTAAG) to the mutant derivative (GAATTATCTCTGATATT) effectively abolished CiaR-DNA protection (Fig. 3E). This further supported the role of the direct repeats as the target for CiaR binding. In addition, RACE PCR was performed to identify the transcription start site of the ciaX promoter. Transcription starts at the A residue 7 bp downstream of the putative Ϫ10 sequence (Fig. 3A), further confirming the authenticity of the ciaX promoter region (data not shown). Taken together, these results indicate that the 8 genes/operons identified by bioinformatic analysis are likely to be directly controlled by CiaR.

DISCUSSION
In this study we sought to address an unresolved aspect of the ciaXRH TCS in S. mutans: why does a deletion of the ciaH sensor kinase gene cause multiple phenotypes, while a deletion of the response regulator ciaR has no effect? To this end, we made a ciaRH double mutation and showed that ciaR is essential for expression of the ciaH phenotypes (Fig. 1), suggesting that CiaR is indeed in the same signaling pathway as CiaH. We further showed that these ciaH mutant phenotypes are caused by the resulting overexpression of ciaR (Fig. 2). Similarly, overexpressing ciaR in a wild-type or ciaRH double mutant background could reproduce the ciaH phenotypes (Fig. 1), indicating that the overexpression of ciaR alone is sufficient for the observed ciaH phenotypes. Microarray analysis of the ciaH mutant and ciaR overexpression strains identified 100 genes whose expression is altered Ն2-fold, 96 of which showed similar changes in the two mutants ( Table 2). Bioinformatics and DNA footprinting analysis identified 8 genes/operon as the CiaR regulon (Fig. 3), 6 of which are the most highly upregulated among all affected genes. Based on these observations, we propose the following model for the mode of regulation by the CiaRH TCS upon its own promoter and that of the CiaR regulon (Fig. 4). When ciaH is absent, expression of ciaR and the ciaXRH operon is increased. This increase in ciaR gene expression creates a positive feedback loop on the gene expression of the ciaXRH operon itself, as well as on the expression of the CiaR regulon genes. Increased expression of the CiaR regulon genes eventually leads to repression of the late competence genes and the mutacin biosynthesis genes (see below), resulting in diminished competence development and mutacin production, two of the ciaH phenotypes. In the presence of ciaH, the activity of CiaR is negated, resulting in lower-level expression of the CiaR regulon and the development of competence and mutacin production. This model suggests that CiaR activates the transcription of the CiaR regulon genes in the absence of CiaH and that CiaH somehow negatively affects the function of CiaR. This notion is further supported by the following observations. (i) The expression of most of the CiaR regulon genes is increased Ͼ10-fold in the ciaH mutant over that in the wild type (Table 2). This makes it unlikely for CiaR to serve as a repressor for the CiaR regulon genes in the absence of CiaH, for that would result in downregulation instead of upregulation of these genes in the ciaH mutant or ciaR overexpression strains. (ii) Inspection of the promoter region of all CiaR regulon genes found a fairly strong Ϫ10 sequence (Ն4 bp out of the 6-bp consensus sequence) but very poor Ϫ35 region (Յ3 bp out of the 6-bp consensus sequence) (Fig. 3A). This indicates that without the assistance of CiaR, RNA polymerase would bind poorly to the promoter region, which explained the large increase in transcription of the CiaR regulon promoters when ciaR was overexpressed as a result of either a ciaH mutation or transcription from the ldh promoter. Whether CiaH affects CiaR due to phosphorylation or dephosphorylation of CiaR has yet to be determined.
As mentioned above, among the significantly downregulated (5-to 10-fold) genes in the ciaH mutant and ciaR overexpression strains are numerous late competence genes, including the 7 genes in the comY operon (SMU.1980c to SMU.1987c), comEA and comEC (SMU.625 and SMU.626), comFA and comFC (SMU.498 and SMU.499), and dprA (SMU.1001). Based on our previous studies, the transcription of the comY operon correlates with the level of competence (20). Thus, the VOL. 192, 2010 CiaRH REGULATION IN S. MUTANS 4675 competence deficiencies observed in the ciaH mutant and the ciaR overexpression strains are likely attributable to downregulation of the late competence genes. In addition, comX is moderately downregulated in the ciaH mutant (1.6-fold) and ciaR overexpression strain (1.75-fold) detected by both microarray and RT-PCR assays (data not shown). Whether this moderate effect is responsible for the 5-to 10-fold downregulation of the late competence genes cannot be determined at this time. None of the genes upstream of comX in the competence regulation cascade (comC, comD, and comE) was affected, suggesting that regulation of competence by the ciaRH system is at the later steps. Inspection of comX and other late competence genes did not find any putative CiaR binding site in their promoter regions, suggesting that CiaRH regulation of competence is likely an indirect effect rather than an effect of direct regulation by CiaR. The same finding was also reported for S. pneumoniae (18). While the competence phenotype can be attributed to the downregulation of the late competence genes in the ciaH mutant and ciaR overexpression strains, the genes responsible for the mutacin I phenotype cannot be discerned from the microarray data set. This is due to the fact that the microarray chip is designed based on the sequence of strain UA159, which does not harbor the mutacin I gene cluster. Another reason is that mutacin I production requires high cell density such as that of colonies grown on a plate (21). Indeed, when RT-PCR was used to measure mutA (structural gene for mutacin I) gene expression in cultures grown on plates, mutA gene expression was downregulated over 10-fold in both the ciaH mutant and ciaR overexpression strains (data not shown). Thus, overexpressed CiaR is also responsible for downregulation of the mutacin I gene expression in the ciaH mutant strain. Like the late competence genes, neither the mutA nor the mutR (positive regulator for mutA) promoter appears to contain the CiaR binding site, suggesting that CiaR regulation of mutacin I production is likely to be indirect.
The mediators for acid tolerance cannot be determined because many of the genes detected in the microarray have not been characterized with regard to their functions in S. mutans. Of particular interest are those that were upregulated Ͼ20fold, such as SMU.40, the SMU.139 operon, SMU.239, SMU.739, and the SMU.1093 operon. These genes all belong to the putative CiaR regulon (Fig. 3) and thereby are assumed to be directly regulated by CiaR. SMU.40 encodes a 53-aminoacid (aa) peptide with a relE-like toxin domain. It is localized in the same operon as SMU.41, encoding a 33-aa peptide, in strain UA159. This organization is typical of the toxin-antitoxin pairs used by bacteria for plasmid maintenance. SMU.139 en-  codes a putative oxalate decarboxylase protein whose function in S. mutans has not been characterized. SMU.139 appears to be cotranscribed with two downstream genes, SMU.140, encoding a putative glutathione reductase, and SMU.141, encoding a membrane protein with unknown function. SMU.239c encodes a membrane protein with a VanZ-like transporter domain. SMU.739 encodes a large hypothetical protein predicted to be extracellular and cell wall associated. SMU.1093 and SMU.1094 encode ABC transporters with unknown function. Experiments are under way to determine whether any of these genes are involved in the ciaH mutant phenotypes. It is interesting that among the 100 differentially regulated genes only 5 encode transcription regulators (ciaR, ciaH, SMU.927, SMU.928, and SMU.345c). In addition to the ciaRH operon itself, SMU.927 and SMU.928 encode another TCS, which is also upregulated (ϳ2-fold) in the ciaH mutation and ciaR overexpression strains ( Table 2). SMU.927 or SMU.928 resides in the same operon as SMU.926, which encodes a putative GTP pyrophosphokinase family protein. In a previous report, Biswas et al. deleted the histidine kinase gene, SMU.928, and did not find any effect on the various phenotypes that they analyzed, including mutacin production and stress resistance (3). SMU.345c resides on a single gene operon, whose expression is increased ϳ1.5-fold and 3.6-fold in the ciaH mutant and ciaR overexpression strains, respectively (Table 2). SMU.345c has an hxlR-type DNA binding domain, named after the Bacillus subtilis transcription activator, HxlR, which activates the transcription of the hxlAB operon responsible for detoxication of formaldehyde (27). Another group of significantly affected genes encode bacteriocins, bacteriocin-like peptide, and small peptides. A total of 15 genes belong to this group (Table 2). Of the 8 downregulated genes, SMU.150 and SMU.151 are mutacin IV structural genes, SMU.423, SMU.1905, and SMU.1906 encode bacteriocin-like peptides with unknown function. All these genes are known to be coregulated with competence (13,14,25). Among the 7 upregulated genes, 3 (SMU.1862, SMU.1889c, and SMU.1892c) encode bacteriocin-like peptides. Whether the two-component system, the putative transcription regulator, or these bacteriocin-like peptides play any role in the CiaRH TCS-mediated cellular functions awaits further experimentation.
Using a combination of bioinformatics and DNA footprinting analysis, we identified 8 genes/operons that are likely to be directly regulated by CiaR (Fig. 3). Promoters of these genes all share the consensus CiaR binding site, NTTAAG-n5-WTTAAG, upstream of their putative Ϫ10 regions, and all but one (SMU.648) were found to be positively regulated in the ciaH mutant and ciaR overexpression strains ( Table 2). These 8 genes/operons, however, may not represent the entire CiaR regulon in S. mutans, due to the restricted NTTAAG-n5-WTTAAG motif used as the MotifSearch query. For example, the putative transcription regulator SMU.345c was upregulated Ͼ3-fold in the CiaR overexpression strain (Table 2), and its putative promoter region contains the two consensus direct repeats 10 bp upstream of the putative Ϫ10 region, but the two direct repeats are separated by 11 bp instead of the consensus 5 bp. Thus, the CiaR regulon member list is likely more extensive than currently recognized. It is also worth noting that in S. pneumoniae, the CiaR regulon also includes 5 small RNAs (sRNAs) (9). Bioinformatics analysis of the S. mutans UA159 genomic sequence did indeed identify 3 putative small RNAs (IG53, IGR358, and IGR1234) that are homologous to the ones identified in S. pneumoniae; however, none of these putative sRNAs exhibited significant levels of signal above the background in our microarray (data not shown). Thus, whether the CiaR regulon of S. mutans includes small RNAs remains an open question.