This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Chong, P.
Right arrow Articles by Biswas, I.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Chong, P.
Right arrow Articles by Biswas, I.

 Previous Article  |  Next Article 

Journal of Bacteriology, July 2008, p. 4478-4488, Vol. 190, No. 13
0021-9193/08/$08.00+0     doi:10.1128/JB.01961-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Modulation of covR Expression in Streptococcus mutans UA159 {triangledown}

Patrick Chong, Laura Drake, and Indranil Biswas*

Department of Microbiology, University of Kansas Medical Center, Kansas City, Kansas 66160

Received 17 December 2007/ Accepted 27 April 2008


arrow
ABSTRACT
 
The biofilm-forming Streptococcus mutans is a gram-positive bacterium that resides in the human oral cavity and is considered to be the primary etiological agent in the formation of dental caries. The global response regulator CovR, which lacks a cognate sensor kinase, is essential for the pathogenesis and biofilm formation of this bacterium, but it is not clear how covR expression is regulated in S. mutans. In this communication, we present the results of our studies examining various factors that regulate the expression of covR in S. mutans UA159. The results of Southern hybridization and PCR analysis indicated that CovR is an orphan response regulator in various isolates of S. mutans. The transcriptional start site for covR was found to be 221 base pairs upstream of the ATG start codon, and site-directed mutagenesis of the upstream TATAAT box confirmed our findings. The expression of covR is growth phase dependent, with maximal expression observed during exponential-growth phase. While changes to the growth temperature did not significantly affect the expression of covR, increasing the pH or the concentration of Mg2+ in the growth medium leads to an increase in covR expression. The results of semiquantitative reverse transcriptase PCR analysis and in vivo transcriptional-fusion reporter assays indicated that CovR autoregulates its own expression; this was verified by the results of electrophoretic mobility shift assays and DNase I protection assays, which demonstrated direct binding of CovR to the promoter region. Apparently, regulation by Mg2+ and the autoregulation of covR are not linked. A detailed analysis of the regulation of CovR may lead to a better understanding of the pathogenesis of S. mutans, as well as providing further insight into the prevention of dental caries.


arrow
INTRODUCTION
 
The human pathogen Streptococcus mutans is a gram-positive bacterium that resides in the oral cavity and is considered to be the primary etiological agent in the formation of dental caries (26, 35). Dental caries is an infectious and costly disease that results in annual expenditures of billions of dollars in the United States (45). S. mutans uses the dietary carbohydrates ingested by its host to adhere to the tooth surface, forming biofilms, known as dental plaque, which enable this organism to maintain its presence in the oral cavity (9). The pathogenesis of S. mutans is linked to its ability to metabolize a wide range of carbohydrates, which also gives it a competitive edge over other microorganisms found in the oral cavity, since S. mutans is tolerant of the acid that it produces (40, 41, 43). The lactic acid formed through the breakdown of carbohydrates via the glycolytic pathway also leads to a localized drop in pH on the tooth surface, resulting in subsequent demineralization of the enamel and the formation of dental caries (43, 44). S. mutans has also been implicated in infective endocarditis, with more than 14% of viridans-streptococcus-induced endocarditis induced by S. mutans (35, 51). The capacity of S. mutans to persist and to maintain a dominant presence in the human oral cavity is due to its ability to rapidly respond to the ever-changing environment of the oral cavity, which includes fluctuations in the availability of nutrients and essential metal ions, the levels of harmful compounds, oxidative and osmotic stresses, and extremes of temperature and pH (8, 47).

Bacteria possess a variety of methods to sense unexpected changes in their extracellular environment, including secondary metabolites, ions, and regulatory proteins, but two-component signal transduction systems (TCS) are the predominant mechanism used to react and respond to environmental fluxes (2, 52). TCS typically consist of a membrane-bound sensor histidine kinase and a cytoplasmic response regulator. The sensor histidine kinase is composed of two components: an amino-terminal sensor/input domain that detects stimuli from the environment and a cytoplasmic transmitter/histidine phosphotransferase domain that autophosphorylates at a specific histidine residue in response to stimulation of the sensor domain (16, 37, 38). The phosphate group is subsequently transferred to the response regulator, which contains two functional components, a conserved amino-terminal domain, which contains a conserved phosphorylatable aspartate residue, and a variable carboxy-terminal effector domain that is activated upon phosphorylation of the aspartate domain (7, 16). The activated response regulator elicits the appropriate cellular response, usually by acting as a transcriptional regulator for specific DNA target sequences.

CovR/S is one of the most-extensively studied TCS in streptococci, particularly in group A streptococci (GAS) (13, 14, 17, 19). CovR/S regulates approximately 15% of the genes in GAS, including virulence genes, such as the has operon (hyaluronic acid capsule synthesis), ska (streptokinase), sagA (streptolysin S), and speB (cysteine protease) (17, 20, 27, 33). CovS is thought to be the cognate sensor kinase for the phosphorylation or dephosphorylation of CovR (11); environmental stimuli, particularly during periods of stress, are recognized by CovS, which in turn regulates the activity of the global response regulator CovR (13). CovR regulates the expression of common sets of genes in different strains, but the repertoire of genes regulated by CovR may vary between strains (12, 50). Besides GAS, CovR/S is also associated with virulence in group B streptococci (GBS), as well as in group C streptococci (20, 32, 48). Approximately 6% of the genes of GBS are regulated by CovR/S, including various virulence factors, such as beta-hemolysin/cytolysin and CAMP factor (29, 32).

In GAS, the expression of covR is growth phase dependent, with the highest levels of transcription observed during exponential-growth phase (17). CovR of GAS negatively regulates its own transcription by directly binding to the promoter region (17, 25); however, autoregulation of covR is not observed in GBS (32). Expression of covR is also influenced by the nutritional conditions of the extracellular environment; Steiner and Malke (49) reported that amino acid starvation led to the up-regulation of covR transcription by an as-yet-unidentified mechanism. Studies conducted by Gryllos et al. (21, 22) demonstrated that extracellular Mg2+ was an effective stimulant of CovR/S, leading to the discovery of a subset of genes under the transcriptional control of CovR that had not previously been identified. Chaussee et al. (10) reported that Rgg, a transcriptional regulator, positively regulates the expression of the covR/S system. The expression of covR in GAS is also up-regulated by the RocA regulator protein, which is not encoded by the genomes of other groups of streptococci, including S. mutans (4).

S. mutans lacks covS but does express CovR, which also plays key roles related to biofilm formation and virulence by regulating the transcription of genes such as glucosyltransferase genes (gtfB/C) and glucan-binding protein C genes (gbpC); the expression of these genes is essential for adherence to the tooth surface and for cariogenicity (3, 6, 31, 39).

The regulation of covR expression in S. mutans has not been fully elucidated to date (3, 28). In this communication, we report the effects of growth phase and various environmental stimulants on the regulation of the expression of covR. Changes in the growth temperature did not significantly change the expression of covR, but increasing the pH of or concentration of Mg2+ in the growth medium leads to an increase in covR expression. The results of our studies also indicate that CovR represses the expression of its own gene by directly binding to the promoter region of covR.


arrow
MATERIALS AND METHODS
 
Bacterial strains and growth conditions. Escherichia coli strain DH5{alpha} was grown in Luria-Bertani (LB) medium supplemented (when necessary) with ampicillin (50 or 100 µg/ml) or kanamycin (50 µg/ml). S. mutans strain UA159 and other S. mutans isolates were typically grown in Todd-Hewitt medium (BBL; Becton Dickinson) supplemented with 0.2% yeast extract (THY). The pH of the THY medium was routinely adjusted with HCl to 7.2 prior to sterilization. For experiments requiring changes to the pH of the growth medium, sterile 50 mM potassium phosphate-citric acid buffer was added to the sterile THY medium. To analyze the effect of Mg2+ on the expression of covR, S. mutans UA159 was grown in chemically defined medium (CDM) as previously described (34, 36), with MnCl2 omitted and supplemented with various concentrations of MgSO4. The growth of the S. mutans cultures was monitored by using a Klett-Summerson colorimeter with a red filter.

Extraction of RNA from S. mutans strain UA159. RNA, which was used for various experiments, was isolated from cultures of S. mutans strain UA159 grown to mid-exponential phase (70 Klett units) under the growth conditions specified above. The cultures were harvested via centrifugation, resuspended and incubated in RNA protect reagent (Qiagen) at room temperature, and then recentrifuged to form a pellet which was either stored overnight at –20°C or processed immediately for RNA extraction. RNA extraction was performed as described by Biswas et al. (3).

Determination of the stability of the covR transcript of S. mutans. The stability of covR mRNA was determined by using a protocol described by Biswas et al. (3). When the culture reached the desired optical density (OD), rifampin, which inhibits RNA polymerase activity, was immediately added to each culture to prevent the de novo synthesis of RNA. RNA was then collected from aliquots of S. mutans cells at eight time points during a total incubation period of 450 (7.5 h) min at 37°C. At each time point, sodium azide (final concentration, 50 mM) was added to the culture to terminate cellular processes, and then the culture was immediately placed in dry ice. RNA was extracted from each culture as described above, and the quality was evaluated by using an Agilent 2100 Bioanalyzer (Agilent Technologies) according to the manufacturer's protocol.

Four micrograms of RNA from each time point was denatured, loaded onto a 1% agarose gel, and separated via electrophoresis for Northern blot analysis. The separated RNA was then transferred to a Zeta-probe nylon membrane (Bio-Rad) according to the manufacturer's instructions (Northern Max-Gly; Ambion). DNA probes were synthesized via PCR amplification using the primer pairs BamHI-CovR-F and HindIII-CovR-R, with chromosomal DNA of S. mutans UA159 as template. The probes were labeled with [{alpha}-32P]dATP by using a DECAprimeII kit (Ambion) and added to a hybridization tube containing the transfer membrane and ULTRAhyb buffer (Ambion), followed by overnight incubation. Following hybridization, the membrane was washed according to the manufacturer's instructions, dried, exposed to a phosphorimager plate, and analyzed by using a Typhoon phosphorimager (Molecular Dynamics).

Semiquantitative RT-PCR analysis. Semiquantitative reverse transcriptase (RT)-PCR was used to quantify the level of expression of covR under various physiological conditions (i.e., growth temperature, pH, and the presence of Mg2+). S. mutans UA159 was grown to mid-exponential phase under different growth conditions as described above, followed by RNA extraction.

RT-PCR was used to determine the transcriptional organization of the covR locus, following a protocol described by Eran et al. (15). DNA-free RNA was used for the synthesis of cDNA using Moloney murine leukemia virus RT (Ambion) according to the manufacturer's protocol. Chromosomal DNA was purified as previously described (4). The primers used for the various RT-PCRs are listed in Table 1.


View this table:
[in this window]
[in a new window]

  		TABLE 1. List of oligonucleotides used in this study

Construction of the PcovR-gusA reporter strain. β-Glucuronidase (GUS) assays were performed as described by Biswas and Biswas (6). The putative promoter region of covR (PcovR; ~700 bp) was amplified from the chromosomal DNA of S. mutans UA159 by using the primers Bam-GcrR-F4 and Xho-GcrR-R6. This DNA fragment was then cloned into the BamHI/XhoI-digested plasmid pIB107 that contains a promoterless GUS gene (gusA) to create plasmid pIB120. pIB120 was then transferred to wild-type strain UA159 and the isogenic covR-negative strain IBS10, using a protocol described by Biswas and Biswas (6), to generate strains IBS373 and IBS375, respectively, via natural transformation.

For complementation analysis, covR was PCR amplified from chromosomal DNA of wild-type strain UA159 by using the primer pair BamHI-covR-F10 and Eco-sal-covR-R10 (Table 1), digested with BamHI and EcoRI, and cloned into a BamHI/EcoRI-digested pOri23 derivative with its native multiple cloning site replaced with the multiple cloning site from plasmid pASK-43 (I. Biswas, unpublished data). The resulting plasmid, containing covR fused to the 3' end of promoter P23, was designated pIB609 and used to transform reporter strains IBS373 (wild type) and IBS375 (covR negative).

For site-directed mutagenesis of the putative pribnow box, the promoter region of covR was amplified by using the primers GcrR-F6 and GcrR-Fout1 (Table 1). The TATAAT sequence was mutated to TCTAGA, and both the wild-type and mutant PcovR fragments were cloned into pIB107, yielding wild-type and mutant PcovR-gusA transcriptional-fusion reporters in plasmids pIB182 and pIB183, respectively. Plasmids pIB182 and pIB183 were subsequently used for the transformation of UA159 to generate the transcriptional-fusion reporter strains IBS607 and IBS608, respectively. IBS104, which contains the promoterless gusA at the SMU_1405 locus, was used as a negative control.

Determination of the transcriptional start site for covR. Primer extension experiments were performed as described by Biswas and Biswas (6). The primer used for the extension reaction was GcrR-FOUT2 (Table 1).

EMSA and DNase I protection assay. Electrophoretic mobility shift assays (EMSA) and DNase I protection assays were performed to demonstrate in vitro binding of CovR to PcovR. Purified His-tagged CovR was produced as described by Biswas et al. (3). Primers GcrR-F7 and GcrR-F-OUT2 were individually labeled with {gamma}32-ATP and used to synthesize radiolabeled PcovR DNA fragments *F7 (using labeled GcrR-F7) and *OUT2 (using labeled GcrR-F-OUT2), respectively, via PCR amplification of a 355-bp GcrR-F7-GcrR-F-OUT2-amplified DNA fragment. EMSA was then performed as described by Biswas and Biswas (6), except that a 4.4% acrylamide gel was used in place of a 4.8% gel.

DNase I protection assays were performed by using the same PcovR-CovR mixtures as were synthesized for EMSA, as described by Biswas and Biswas (6), except that His-tagged CovR was substituted for MBP-CovR.


arrow
RESULTS
 
Genetic analysis of the covR locus in S. mutans UA159. Analysis of the annotated genome sequence of S. mutans strain UA159 indicated that covS is not present in this bacterium, unlike GAS or GBS (Fig. 1A). No genes coding for a potential sensor kinase were found in the immediate vicinity of the covR locus, confirming that CovR is an orphan response regulator lacking a cognate sensor kinase in S. mutans UA159.


Figure 1
View larger version (51K):
[in this window]
[in a new window]

  		FIG. 1. Genomic analysis of the covR locus in S. mutans UA159. (A) Schematic representation of the covR locus in S. mutans UA159 (Smu), GAS, and GBS. ORFs are indicated by the block arrows. (B) Southern hybridization blot of chromosomal DNA from various mutans isolates digested with XmnI. XmnI-digested chromosomal DNA from S. mutans UA159 and other mutans isolates was separated via electrophoresis in 1% agarose, followed by depurination, denaturation, and overnight transfer to a Zeta-probe blotting membrane (Bio-Rad). The transferred DNA was then fixed to the membrane via treatment with UV. A labeled DNA fragment containing the full-length covR sequence derived from UA159 was then used as a probe against the digested DNA and allowed to hybridize overnight. The membrane was then air dried and exposed to a phosphorimager plate. (C) PCR analysis of the intergenic region between covR and gene SMU_1923c in various S. mutans isolates. The positions of the primers used for this analysis (GcrR-Rout1 and GcrR-Rev2) are marked in panel A.

While the presence of covS was not identified in strain UA159, it was not known whether this was also the case for other isolates of S. mutans. To determine if the lack of covS in UA159 was anomalous, Southern hybridization was used to verify whether covS is present in other S. mutans isolates from three different serotypes (c, e, and f) of S. mutans. A radiolabeled 1.75-kb DNA fragment, containing the full-length covR sequence derived from the genome of UA159, was used as a probe against the XmnI-digested chromosomal DNA from a number of S. mutans isolates; the results of the Southern hybridization experiment are shown in Fig. 1B. This probe hybridized with three fragments, 0.5, 1.1, and 6.2 kb in length, of XmnI-digested UA159 chromosomal DNA; with the exception of strain V100, the hybridization pattern observed for all of the isolates was identical with that of UA159, indicating that covS is not present in any of the isolates tested. These findings are further supported by the results of the PCR analysis using primers GcrR-Rout1 and GcrR-Rev2. Amplification of the intergenic region between the 3' end of covR and the 5' end of SMU_1923c from the chromosomal DNA of strain UA149 generated a DNA fragment of 448 bp; this was also observed with various other strains of S. mutans (Fig. 1C). Taken together, the results suggest that CovS is not encoded in S. mutans as it is in GAS and GBS.

Analysis of covR expression in S. mutans. In order to better understand the molecular mechanism of covR expression in S. mutans, primer extension was performed to determine the transcriptional start site within the intergenic region upstream of covR. The transcriptional start site was mapped to a position 221 base pairs upstream of the putative start codon (Fig. 2A). The putative pribnow box (–10, TATAAT) was found 9 base pairs upstream from the transcriptional start site; site-directed mutagenesis of the TATAAT sequence to TCTAGA eliminated the ability of the putative PcovR to function as a promoter when fused to the gusA gene for the PcovR-gusA transcriptional-fusion reporter assays (Fig. 2B). A putative –35 box (CATTGA) was found 34 base pairs from the transcriptional start site box, and a potential ribosome-binding site (AGGAG) was found 8 base pairs upstream of the putative start codon.


Figure 2
View larger version (23K):
[in this window]
[in a new window]

  		FIG. 2. Analysis of covR transcription. (A) Primer extension (PE) was performed as described in Materials and Methods. The putative –35 and –10 sites (capitalized) are indicated. The asterisk indicates the transcriptional start site, which has also been capitalized. nt, nucleotides. (B) GUS assay results demonstrating the importance of the putative –10 TATAAT sequence in the promoter region. Strain IBS104 carries promoterless gusA, IBS607 carries the wild-type –10 sequence, and IBS608 carries a mutant –10 sequence. Error bars show standard deviations. (C) Genetic map of the covR locus in S. mutans UA159 indicating the primers used for the PCR analysis to determine potential linkage of the genes downstream of covR. Bent arrow represents transcriptional start site for covR. Stem-loop structures designate the positions of putative rho-independent terminator sites. (D) Results of Northern hybridization analysis of the covR transcript indicating the presence of several species of transcript. The prominent transcripts are noted by length in kilobases to the left and in panel C. (E) Results of PCR analysis showing linkages of the various genes that are cotranscribed with covR. RNA was used as template to produce cDNA. PCR was then performed on RNA (control), cDNA, and chromosomal DNA, using the primer pairs described in Table 1 and depicted in panel C, to determine which of the genes in the covR locus are cotranscribed with covR.

To determine the stability of the covR transcript, RNA was extracted from aliquots of S. mutans UA159 that were grown to various time points. When cultures reached the desired OD, rifampin was immediately added to each culture to prevent the de novo synthesis of RNA. RNA was then extracted from each sample as previously described. The half-life of the major covR transcript (1.8 kb) was quantified by using Northern hybridization analysis and found to be approximately 2 min (data not shown).

Northern blot analysis to determine the transcriptional organization of the covR locus was then performed using RNA extracted from S. mutans UA159; the results are shown in Fig. 2D. Hybridization of the Northern blot with radiolabeled DNA probe containing the covR open reading frame (ORF) indicated the presence of several transcripts, with a major transcript (~33% of the total) corresponding to a length of 1.8 kb, suggesting that covR and SMU_1923c, which encodes a hypothetical protein, are likely cotranscribed. The other transcripts detected on the Northern blot corresponded to lengths of approximately 1.3, 2.2, 2.5, and 5.8 kb, indicating that other genes further downstream of covR are also cotranscribed but to a lesser degree.

The results of the Northern blot analysis were verified via RT-PCR (Fig. 2E). To determine which of the genes from the putative operon were transcriptionally linked, RT-PCRs were performed with cDNA and chromosomal DNA (control) as template, using various oligonucleotide primers (Table 1) designed to amplify different segments of the covR locus, as depicted in Fig. 2C. The results indicate that the upstream ylnB and covR are not cotranscribed. On the other hand, the results confirm that linkage exists between covR and several downstream genes, including SMU_1923c and dnaB and, potentially, dnaI, pgdA, sapR, dedA, comE, comD, and comC, yielding a potential transcript with a length of approximately 9.1 kb. Analysis of the S. mutans genome at the Comprehensive Microbial Resource website (CMR; http://cmr.jcvi.org/cgi-bin/CMR/CmrHomePage.cgi) predicted the presence of two putative rho-independent terminator sequences at the termini of the putative covR locus. One sequence is found in the intergenic region located between ylnB (SMU_1925c) and covR (SMU_1924c), while the other putative terminator site is located at the intergenic region between comD (SMU_1916) and comC (SMU_1915); no other sites were predicted within the region (Fig. 2C).

Semiquantitative RT-PCR was used to quantify the level of expression of covR of S. mutans during various growth phases, as well as during growth under different growth conditions. The level of expression of gyrA was included as an internal control to ensure that equivalent amounts of RNA were being used for the RT-PCR analysis. As shown in Fig. 3A, RNA was extracted from S. mutans at eight different time points over an incubation period of 450 min (7.5 h). The expression of covR was highest in the exponential-growth phase during the first 200 min of growth, with equivalent amounts of covR transcript being produced throughout (Fig. 3B). In contrast, there was a significant decrease in the level of expression of covR during the growth of S. mutans in the stationary phase.


Figure 3
View larger version (37K):
[in this window]
[in a new window]

  		FIG. 3. Growth-phase-dependent expression of covR. (A) RNA samples were extracted from cultures of S. mutans UA159 at the indicated time points. (B) The extracted RNA was subjected to semiquantitative PCR analysis, with the results indicating that expression of covR is maximal during exponential-growth phase. The expression of gyrA was also included as an internal control to ensure that equal amounts of RNA were being analyzed for each time point. Experiments were performed in triplicate, with three independent RNA extractions, and the relevant portions of a representative gel are shown.

The human oral cavity is a dynamic environment that is often in a state of flux. To survive, S. mutans must rapidly adapt to its ever-changing environment. Therefore, it was of great interest to determine whether changes to the growth conditions could influence the expression of covR, whose gene product is essential for regulation of the expression of various virulence-associated genes; the results are shown in Fig. 4. No significant changes in the level of expression of covR were observed when the growth temperature was varied from 28°C to 42°C, although a there was a small increase in expression observed at a growth temperature of 37°C. Similarly, variations in pH, from pH 5.5 to pH 7.0, did not lead to significant changes in the level of expression of covR, although there was an increase in the expression of covR when the cultures were grown in unbuffered THY medium (~pH 7.2). The addition of Mg2+ to the growth medium resulted in a dose-dependent increase in the level of expression of covR, with the highest expression observed when the cultures were grown with 5 or 10 mM MgSO4. Taken together, the results suggest that the expression of covR is dependent on the growth phase, the pH, and the addition of Mg2+ but is relatively insensitive to variations in temperature under the growth conditions tested.


Figure 4
View larger version (23K):
[in this window]
[in a new window]

  		FIG. 4. Regulation of expression of covR under different growth conditions. RNA was extracted from cultures of S. mutans UA159 grown with variations in temperature, pH, and MgSO4 concentration (mM). Semiquantitative PCR was then used to measure the levels of covR expressed under the different growth conditions. The expression of gyrA was also included as an internal control to ensure that equal amounts of RNA were being analyzed for each condition. Experiments were performed in triplicate, with three independent RNA extractions, and the relevant portions of a representative gel are shown. UB, unbuffered.

The expression of covR is autoregulated by CovR. CovR regulates the expression of covR in GAS, but not in GBS (17, 25, 32). To determine whether autoregulation occurs in S. mutans, a mutant strain of UA159 (IBS10) was created which contained an insertional mutation in covR. Semiquantitative RT-PCR was performed to compare the levels of expression of covR in wild-type UA159 and IBS10; the transcription of gyrA was used as an internal control to ensure that equal amounts of RNA were being used during the semiquantitative RT-PCR analysis. As shown in Fig. 5A, the transcription of covR in IBS10 increased approximately 1.7-fold relative to the level in the wild-type strain, suggesting that CovR represses the expression of its own gene. The results of the semiquantitative RT-PCR analyses were verified with in vivo reporter assays in which transcription from PcovR was quantified by measuring the activity of GusA produced from the PcovR-gusA fusion reporter (Fig. 5B). An increase in the GUS activity of IBS375 (covR negative) of approximately 2.1-fold compared to the activity of IBS373 (wild-type) was observed. The transformation of IBS375 with pIB609, which contains the covR gene under the control of promoter P23, restored the GusA activity of the covR mutant strain to the level of its isogenic wild-type parent, while cultures of IBS375 transformed with pOri23 demonstrated levels of GusA activity similar to those in the nontransformed mutant strain. Taken together, the results suggest that CovR acts as a negative regulator of covR expression.


Figure 5
View larger version (34K):
[in this window]
[in a new window]

  		FIG. 5. CovR-dependent regulation of covR expression. (A) Results of semiquantitative RT-PCR analysis of RNA extracted from wild-type (WT) S. mutans UA159 and IBS10 (covR negative). The expression of gyrA was also included as an internal control to ensure that equal amounts of RNA were analyzed. (B) Expression from PcovR was quantified using the PcovR-gusA transcriptional-fusion reporter strains IBS373 (wild type), IBS375 (covR negative), IBS375/pIB609 (plasmid pOri23 derivative containing wild-type covR), and IBS375/pOri23 grown to mid-exponential phase. GUS assays were performed in (at least) triplicate, using three independent preparations of each strain/culture. Error bars show standard deviations. (C) Binding of CovR to the promoter region of covR. EMSA was performed with His-tagged CovR as described in the text. Lanes: 1, no CovR; 2, 9.2 pmol CovR; 3, 12.3 pmol CovR; 4, 18.4 pmol CovR; 5, 24.5 pmol CovR; 6, 36.8 pmol CovR; 7, 49.0 pmol CovR. EMSA experiments were performed at least three times, and a representative gel is shown. B, bound DNA; F, free DNA. (D) DNase I protection assay of the covR promoter. DNase I protection assays were performed with His-tagged CovR as described in the text. Lanes: 1, no CovR; 2, 3.1 pmol CovR; 3, 6.1 pmol CovR; 4, 12.3 pmol CovR; 5, 24.5 pmol CovR; 6, 36.8 pmol CovR; 7, 49.0 pmol CovR; 8, 73.5 pmol CovR. CovR-binding regions are defined with vertical lines adjacent to the sequencing gel, and the region containing the transcription start site is marked with an asterisk. Horizontal lines mark the segments bound by CovR on the schematic diagram below the gel. DNase I protection assays were performed in triplicate, and a representative gel is shown here.

In order to verify that the observed repression of covR was directly attributable to CovR, purified His-tagged CovR was incubated with a radiolabeled, GcrR-F7-GcrR-F-OUT2-amplified 355-bp DNA fragment (0.1 pmol) containing the promoter region of covR, as described in Materials and Methods. The CovR-PcovR mixture was then used for an EMSA to demonstrate the binding of CovR with PcovR; a representative gel is shown in Fig. 5C. The addition of increasing amounts of CovR (0 to 49.0 pmol) resulted in a shift in the mobility of PcovR in the gel, indicating that CovR binds to PcovR. The addition of fivefold-higher amounts (i.e., 0.5 pmol) of unlabeled PrpsL did not inhibit the binding of CovR to PcovR, while the addition of fivefold-higher amounts of unlabeled PcovR inhibited the binding of CovR to the labeled PcovR fragment (data not shown), indicating specificity of CovR for PcovR.

DNase I protection assays were performed to determine the specific region of PcovR bound by CovR. The PCR-amplified 355-bp fragment used for EMSA was also used for the DNase I protection assays, as described in Materials and Methods. Analysis of the sequencing gel (Fig. 5D) indicated that CovR binds to the DNA fragment spanning positions –6 to +250 (+1 is the transcriptional start site), encompassing part of the pribnow box and the transcriptional start site described above, as well as the region immediately downstream of the transcriptional start site, ending 30 bp upstream of the start codon. There is a clear increase in the size and intensity of the footprint generated when the amount of CovR incubated with PcovR is increased, consistent with the results of the EMSA analysis.

Thus, the results indicate that CovR autoregulates transcription from its own promoter by binding directly to the promoter region of covR, including the transcriptional start site as well as regions downstream of the transcriptional start site, thereby enabling CovR to repress its own expression. One interesting question was whether Mg2+ had any role in the CovR autoregulation, since the addition of Mg2+ increases the expression of covR, as shown in Fig. 4. The covR mutant IBS10 and its isogenic wild-type parent, UA159, were grown in CDM supplemented with various concentrations of MgSO4 (0.5 to 5.0 mM). As shown in Fig. 6, increases in the level of expression of the mutant covR mRNA coincided with the increase in wild-type covR mRNA, concurrent with increases in the concentration of MgSO4. Given that the mutant CovR is inactive and unable to regulate its own expression, the results suggest that the autoregulation of covR is independent of Mg2+ and that the observed increase in covR expression in the presence of Mg2+ is likely due to an as-yet-unidentified Mg2+-dependent activator of covR.


Figure 6
View larger version (21K):
[in this window]
[in a new window]

  		FIG. 6. Autoregulation of covR is independent of Mg2+. Cultures of wild-type UA159 and covR mutant IBS10 were grown in CDM supplemented with increasing amounts of MgSO4 as described in Materials and Methods. RNA was extracted from each culture and subjected to semiquantitative RT-PCR. Experiments were performed in triplicate, and the relevant portions of a representative gel are shown.


arrow
DISCUSSION
 
CovR is essential for the biofilm formation and pathogenesis of S. mutans, as the transcription of genes encoding various virulence factors, such as glucosyltransferase and glucan-binding protein C, is essential for the adherence of this pathogen to the tooth surface, leading to the colonization of the oral cavity as well as the onset of cariogenicity and tooth decay (3, 6, 28). Although CovR of S. mutans shares significant homology with CovR of GAS (84% similarity and 75% identity [1]), CovR in S. mutans functions primarily as a transcriptional activator (Biswas, unpublished), whereas CovR in GAS is predominantly a transcriptional repressor (20). Furthermore, the regulation of covR from the cariogenic pathogen S. mutans has not been well characterized to date. Given that dental decay is an extremely costly global health problem that affects 60 to 90% of school children, as well as many adults, in industrialized countries (30, 45, 46), it would be of great interest to elucidate the factors leading to the regulation of covR expression and, ultimately, to the development of vaccines to combat the spread of S. mutans and dental caries.

Unlike CovR of GAS or GBS, CovR of S. mutans UA159 is not partnered with a cognate sensor kinase (1). While the gene pair of covR and covS is well conserved and cotranscribed in GAS and GBS, covR is an orphan in the genome of S. mutans UA159 (Fig. 1A). The results of Southern hybridization and PCR analysis indicate that the lack of covS is common to all of the isolates of S. mutans tested (Fig. 1B and C). However, the results of a tBLASTx analysis of the S. pyogenes MGAS315 complete genome sequence (GenBank accession number NC_004070.1) using the 372-bp intergenic-region sequence located between covR and SMU_1923c as query indicated that the 5' (i.e., 3' end of covR) 146 bp of the intergenic-region sequence encodes a truncated TCS histidine kinase (data not shown). Thus, it would appear that a deletion event occurred at some point that removed the cognate sensor kinase of CovR in S. mutans. Cross talk between CovR and other sensor kinases, phosphorylation by a donor such as acetyl phosphate, or activation by other transcriptional activators may substitute for CovS in S. mutans. This is similar to the observations of Dalton and Scott (13), who reported that the activity of CovR was not dependent on the presence of CovS in GAS covS mutants under standard growth conditions.

The results of primer extension analysis indicated that the transcriptional start site of covR was located 221 bp upstream of the putative start codon. This is consistent with the observations of Miller et al. (42), who reported that the transcriptional start site of covR/S in GAS was found 200 bp upstream of the putative start codon. Furthermore, the results of the transcriptional-fusion reporter assays, following site-directed mutagenesis of the pribnow box sequence from TATAAT to TCTAGA, show that GusA activity from the mutant PcovR-gusA reporter strain (IBS608) is not expressed, which is consistent with the lack of activity from the promoterless gusA strain (IBS104); in contrast, GusA activity was only observed from the wild-type PcovR-gusA reporter strain (IBS607), indicating that the targeted TATAAT sequence, found 9 bp upstream of the transcriptional start site, is essential for transcription. While the exact function of this untranslated region (UTR) requires further analysis, we speculate that this region may have a role in the regulation of covR mRNA stability, specifically, in mRNA destabilization. Analysis of the UTR sequence also reveals the presence of short ORFs; in lower eukaryotes, such as Saccharomyces cerevisiae, short ORFs upstream of the target gene encode small peptides that destabilize the mRNA following termination of translation (53, 54). Further studies are required to determine if this is also observed with the ORFs found within the intergenic region preceding covR. There is also ample evidence in the literature that describes various elements found in the UTR of various bacterial genes that regulate mRNA stability; future studies will determine if any such element is present in the UTR of PcovR.

The results of Northern blot analysis indicated that the half-life of the covR transcript of S. mutans was approximately 2 min; this is similar to the observations of Steiner and Malke (48), who reported a half-life of 0.5 min for the covR/S transcript extracted from Streptococcus dysgalactiae. Despite the short half-life, transcripts of covR of S. mutans were detected at high levels during the exponential-growth phase, but not during stationary phase. Similarly, the levels of covR transcript in GAS are also found at maximal levels during mid- to late-exponential growth and decline upon the entry of the cultures into stationary phase (17). Recent data also suggest that CovR is essential for the growth of S. mutans. Mutant cultures containing a near-full deletion of the covR gene did not grow to the same OD as the wild-type culture; complementation of the mutant in trans with the full-length covR gene restored growth to wild-type levels (data not shown). Furthermore, cells containing inactivated covR formed clumps after incubation in THY broth to mid-exponential growth phase, which was not observed in wild-type cultures or in covR mutant strains complemented in trans with covR (data not shown).

The results of Northern blot analysis also indicated the presence of transcripts of various lengths (1.3, 2.2, 2.5, and 5.8 kb), suggesting that several genes downstream of covR may be cotranscribed. Oddly, the length of some of the transcripts suggests incomplete transcription of some of the genes downstream of covR. Genomic analysis at the CMR website did not reveal the presence of rho-independent terminator sites within the genes. One possible explanation for the observed unusual length of the transcripts could be RNA degradation during Northern blot analysis.

The oral cavity is a dynamic environment, such that the pH, temperature, levels of nutrients, metal ion availability, and conditions of osmotic pressure or oxidative stress remain in a state of flux. The expression of covR was not affected significantly by changes in the growth temperature, but there was an increase in the expression of covR concomitant with an increase in the growth medium's pH or Mg2+ content (Fig. 4). The increase in expression at pH 7.2 is not surprising, given that human saliva can be slightly alkaline. Gryllos et al. (21, 22) reported that the presence of Mg2+ in the growth medium stimulated the activity of CovR/S, specifically acting as a stimulant for CovS. In the absence of CovS, it is possible that Mg2+ activates one of the previously identified TCS of S. mutans (5), which subsequently leads to the transcription of covR or the activation of CovR in S. mutans. However, the increase in the expression of covR under in vitro conditions was observed using concentrations of Mg2+ that are significantly higher than physiological conditions; the concentration of Mg2+ in saliva is 0.22 mM, while the concentration of Mg2+ in blood is approximately 1 mM (22, 47). As such, the physiological relevance of Mg2+ for the expression of covR remains to be studied.

Response regulator proteins often regulate the expression of the genes that encode them (17). A strain of S. mutans UA159 containing an insertional mutation in covR (IBS10) produced higher levels of covR transcript (~1.7-fold) than the strains containing the wild-type UA159 (Fig. 5A). Similarly, the results of the transcriptional-fusion (PcovR-gusA) reporter assays indicated that transcriptional activity in the covR mutant IBS375 was increased approximately 2.1-fold in comparison to the level in the wild-type strain, IBS373 (Fig. 5B); complementation in trans with wild-type covR restored the GusA activity to the wild-type level. Taken together, the results indicate that CovR down-regulates transcription from its own promoter. Although the reduction of the covR transcript is less than twofold, it may or may not correlate with the amount of CovR protein present in the cell. More analysis is needed to determine the actual reduction of the level of CovR protein due to autoregulation.

The results described above are supported by the findings from the EMSA and DNase I protection assays, which demonstrate that CovR directly binds to a large region of the DNA sequence containing the promoter of covR (Fig. 5C and D), suggesting that the CovR bound to PcovR could prevent the binding of RNA polymerase to PcovR, thereby inhibiting the expression of covR. The autoregulation of covR was not dependent on the presence of Mg2+, although the addition of Mg2+ did stimulate the expression of covR in both the wild-type and covR mutant strains. These results are not surprising as the homologous protein from GAS also induces direct repression of covR expression (17, 25). This is an important characteristic as it allows the bacteria to rapidly modulate the expression of covR and, in turn, the CovR regulon, in response to various environmental cues. However, since the degree of covR repression was observed to be 2.1-fold, it is not clear how the autoregulation of covR affects the expression of CovR-regulated genes; further studies are required to elucidate the link between gene expression and covR autoregulation.

A consensus binding sequence (CBS), ATTARA, was identified in the promoter sequences regulated by CovR in GAS, including the promoters for covR, has, and dppA (18, 23-25). However, while the ATTARA sequence appeared to be essential for binding under in vitro conditions, the CBS did not appear to be required under in vivo conditions, since expression was not inhibited in vivo after mutation of the CBS (23), such that the physiological relevance of these sequences is not clear. Genomic analysis indicates that similar sequences (i.e., ATTATA) are also found in part of the intergenic region between ylnB and covR that is protected by CovR. Experiments are under way to determine whether these sequences are CBS for CovR binding in S. mutans.

Although we are beginning to develop an understanding of the regulation of covR expression in S. mutans, more studies are required to elucidate the full nature of covR regulation. The results of studies in our laboratory have indicated that covR expression in S. mutans may be associated with the activity of the ClpP protease (Biswas, unpublished), since inactivation of the clpP gene leads to up-regulation of the expression of covR. However, it is not clear if there is a direct link between ClpP and CovR or if ClpP is acting upon another protein that may act as a transcriptional regulator of covR. Further studies will be required to elucidate the relationship between ClpP and the expression of covR. Additional analysis of the various factors that influence the regulation of CovR will give us further insight into the pathogenic mechanisms by which S. mutans causes disease.


arrow
ACKNOWLEDGMENTS
 
We thank Nick Fromm for his invaluable technical assistance.

This publication was made possible in part by an NIDCR grant, DEO16686, awarded to I.B.


arrow
FOOTNOTES
 
* Corresponding author. Mailing address: 3000 Wahl Hall East, University of Kansas Medical Center, 3901 Rainbow Blvd., Kansas City, KS 66160. Phone: (605) 588-7019. Fax: (913) 588-7295. E-mail: ibiswas{at}kumc.edu Back

{triangledown} Published ahead of print on 9 May 2008. Back


arrow
REFERENCES
 
    1
  1. Ajdic, D., W. M. McShan, R. E. McLaughlin, G. Savic, J. Chang, M. B. Carson, C. Primeaux, R. Tian, S. Kenton, H. Jia, S. Lin, Y. Qian, S. Li, H. Zhu, F. Najar, H. Lai, J. White, B. A. Roe, and J. J. Ferretti. 2002. Genome sequence of Streptococcus mutans UA159, a cariogenic dental pathogen. Proc. Natl. Acad. Sci. USA 99:14434-14439.[Abstract/Free Full Text]
    2. 2
  2. Bijlsma, J. J., and E. A. Groisman. 2003. Making informed decisions: regulatory interactions between two-component systems. Trends Microbiol. 11:359-366.[CrossRef][Medline]
    3. 3
  3. Biswas, I., L. Drake, and S. Biswas. 2007. Regulation of gbpC expression in Streptococcus mutans. J. Bacteriol. 189:6521-6531.[Abstract/Free Full Text]
    4. 4
  4. Biswas, I., and J. R. Scott. 2003. Identification of rocA, a positive regulator of covR expression in the group A Streptococcus. J. Bacteriol. 185:3081-3090.[Abstract/Free Full Text]
    5. 5
  5. Biswas, I., L. Drake, D. Erkina, and S. Biswas. 2008. Involvement of sensor kinases in the stress tolerance response of Streptococcus mutans. J. Bacteriol. 190:68-77.[Abstract/Free Full Text]
    6. 6
  6. Biswas, S., and I. Biswas. 2006. Regulation of the glucosyltransferase (gtfBC) operon by CovR in Streptococcus mutans. J. Bacteriol. 188:988-998.[Abstract/Free Full Text]
    7. 7
  7. Buckler, D. R., G. S. Anand, and A. M. Stock. 2000. Response-regulator phosphorylation and activation: a two-way street? Trends Microbiol. 8:153-156.[CrossRef][Medline]
    8. 8
  8. Carlsson, J. 1997. Bacterial metabolism in dental biofilms. Adv. Dent. Res. 11:75-80.[Abstract/Free Full Text]
    9. 9
  9. Carlsson, J., and I. R. Hamilton. 1994. Metabolic activity of oral bacteria, 2nd ed. Munksgaard, Copenhagen, Denmark.
    10. 10
  10. Chaussee, M. S., G. L. Sylva, D. E. Sturdevant, L. M. Smoot, M. R. Graham, R. O. Watson, and J. M. Musser. 2002. Rgg influences the expression of multiple regulatory loci to coregulate virulence factor expression in Streptococcus pyogenes. Infect. Immun. 70:762-770.[Abstract/Free Full Text]
    11. 11
  11. Churchward, G. 2007. The two faces of Janus: virulence gene regulation by CovR/S in group A streptococci. Mol. Microbiol. 64:34-41.[CrossRef][Medline]
    12. 12
  12. Dalton, T. L., R. I. Hobb, and J. R. Scott. 2006. Analysis of the role of CovR and CovS in the dissemination of Streptococcus pyogenes in invasive skin disease. Microb. Pathog. 40:221-227.[CrossRef][Medline]
    13. 13
  13. Dalton, T. L., and J. R. Scott. 2004. CovS inactivates CovR and is required for growth under conditions of general stress in Streptococcus pyogenes. J. Bacteriol. 186:3928-3937.[Abstract/Free Full Text]
    14. 14
  14. Engleberg, N. C., A. Heath, K. Vardaman, and V. J. DiRita. 2004. Contribution of CsrR-regulated virulence factors to the progress and outcome of murine skin infections by Streptococcus pyogenes. Infect. Immun. 72:623-628.[Abstract/Free Full Text]
    15. 15
  15. Eran, Y., Y. Getter, M. Baruch, I. Belotserkovsky, G. Padalon, I. Mishalian, A. Podbielski, B. Kreikemeyer, and E. Hanski. 2007. Transcriptional regulation of the sil locus by the SilCR signalling peptide and its implications on group A streptococcus virulence. Mol. Microbiol. 63:1209-1222.[CrossRef][Medline]
    16. 16
  16. Fabret, C., V. A. Feher, and J. A. Hoch. 1999. Two-component signal transduction in Bacillus subtilis: how one organism sees its world. J. Bacteriol. 181:1975-1983.[Free Full Text]
    17. 17
  17. Federle, M. J., K. S. McIver, and J. R. Scott. 1999. A response regulator that represses transcription of several virulence operons in the group A streptococcus. J. Bacteriol. 181:3649-3657.[Abstract/Free Full Text]
    18. 18
  18. Federle, M. J., and J. R. Scott. 2002. Identification of binding sites for the group A streptococcal global regulator CovR. Mol. Microbiol. 43:1161-1172.[CrossRef][Medline]
    19. 19
  19. Ferretti, J. J., W. M. McShan, D. Ajdic, D. J. Savic, G. Savic, K. Lyon, C. Primeaux, S. Sezate, A. N. Suvorov, S. Kenton, H. S. Lai, S. P. Lin, Y. Qian, H. G. Jia, F. Z. Najar, Q. Ren, H. Zhu, L. Song, J. White, X. Yuan, S. W. Clifton, B. A. Roe, and R. McLaughlin. 2001. Complete genome sequence of an M1 strain of Streptococcus pyogenes. Proc. Natl. Acad. Sci. USA 98:4658-4663.[Abstract/Free Full Text]
    20. 20
  20. Graham, M. R., L. M. Smoot, C. A. Migliaccio, K. Virtaneva, D. E. Sturdevant, S. F. Porcella, M. J. Federle, G. J. Adams, J. R. Scott, and J. M. Musser. 2002. Virulence control in group A Streptococcus by a two-component gene regulatory system: global expression profiling and in vivo infection modeling. Proc. Natl. Acad. Sci. USA 99:13855-13860.[Abstract/Free Full Text]
    21. 21
  21. Gryllos, I., C. Cywes, M. H. Shearer, M. Cary, R. C. Kennedy, and M. R. Wessels. 2001. Regulation of capsule gene expression by group A Streptococcus during pharyngeal colonization and invasive infection. Mol. Microbiol. 42:61-74.[CrossRef][Medline]
    22. 22
  22. Gryllos, I., J. C. Levin, and M. R. Wessels. 2003. The CsrR/CsrS two-component system of group A Streptococcus responds to environmental Mg2+. Proc. Natl. Acad. Sci. USA 100:4227-4232.[Abstract/Free Full Text]
    23. 23
  23. Gusa, A. A., B. J. Froehlich, D. Desai, V. Stringer, and J. R. Scott. 2007. CovR activation of the dipeptide permease promoter (PdppA) in group A streptococcus. J. Bacteriol. 189:1407-1416.[Abstract/Free Full Text]
    24. 24
  24. Gusa, A. A., J. Gao, V. Stringer, G. Churchward, and J. R. Scott. 2006. Phosphorylation of the group A streptococcal CovR response regulator causes dimerization and promoter-specific recruitment by RNA polymerase. J. Bacteriol. 188:4620-4626.[Abstract/Free Full Text]
    25. 25
  25. Gusa, A. A., and J. R. Scott. 2005. The CovR response regulator of group A streptococcus (GAS) acts directly to repress its own promoter. Mol. Microbiol. 56:1195-1207.[CrossRef][Medline]
    26. 26
  26. Hamada, S., and H. D. Slade. 1980. Biology, immunology, and cariogenicity of Streptococcus mutans. Microbiol. Rev. 44:331-384.[Free Full Text]
    27. 27
  27. Heath, A., V. J. DiRita, N. L. Barg, and N. C. Engleberg. 1999. A two-component regulatory system, CsrR-CsrS, represses expression of three Streptococcus pyogenes virulence factors, hyaluronic acid capsule, streptolysin S, and pyrogenic exotoxin B. Infect. Immun. 67:5298-5305.[Abstract/Free Full Text]
    28. 28
  28. Idone, V., S. Brendtro, R. Gillespie, S. Kocaj, E. Peterson, M. Rendi, W. Warren, S. Michalek, K. Krastel, D. Cvitkovitch, and G. Spatafora. 2003. Effect of an orphan response regulator on Streptococcus mutans sucrose-dependent adherence and cariogenesis. Infect. Immun. 71:4351-4360.[Abstract/Free Full Text]
    29. 29
  29. Jiang, S. M., M. J. Cieslewicz, D. L. Kasper, and M. R. Wessels. 2005. Regulation of virulence by a two-component system in group B streptococcus. J. Bacteriol. 187:1105-1113.[Abstract/Free Full Text]
    30. 30
  30. Kaste, L. M., R. H. Selwitz, R. J. Oldakowski, J. A. Brunelle, D. M. Winn, and L. J. Brown. 1996. Coronal caries in the primary and permanent dentition of children and adolescents 1-17 years of age: United States, 1988-1991. J. Dent. Res. 75(Spec. no.):631-641.[Medline]
    31. 31
  31. Kuramitsu, H. 1993. Virulence factors of mutans streptococci: role of molecular genetics. Crit. Rev. Oral Biol. Med. 4:159-176.[Abstract/Free Full Text]
    32. 32
  32. Lamy, M. C., M. Zouine, J. Fert, M. Vergassola, E. Couve, E. Pellegrini, P. Glaser, F. Kunst, T. Msadek, P. Trieu-Cuot, and C. Poyart. 2004. CovS/CovR of group B streptococcus: a two-component global regulatory system involved in virulence. Mol. Microbiol. 54:1250-1268.[CrossRef][Medline]
    33. 33
  33. Levin, J. C., and M. R. Wessels. 1998. Identification of csrR/csrS, a genetic locus that regulates hyaluronic acid capsule synthesis in group A streptococcus. Mol. Microbiol. 30:209-219.[CrossRef][Medline]
    34. 34
  34. Li, Y. H., P. C. Lau, N. Tang, G. Svensater, R. P. Ellen, and D. G. Cvitkovitch. 2002. Novel two-component regulatory system involved in biofilm formation and acid resistance in Streptococcus mutans. J. Bacteriol. 184:6333-6342.[Abstract/Free Full Text]
    35. 35
  35. Loesche, W. J. 1986. Role of Streptococcus mutans in human dental decay. Microbiol. Rev. 50:353-380.[Free Full Text]
    36. 36
  36. Loo, C. Y., D. A. Corliss, and N. Ganeshkumar. 2000. Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J. Bacteriol. 182:1374-1382.[Abstract/Free Full Text]
    37. 37
  37. Mascher, T. 2006. Intramembrane-sensing histidine kinases: a new family of cell envelope stress sensors in Firmicutes bacteria. FEMS Microbiol. Lett. 264:133-144.[CrossRef][Medline]
    38. 38
  38. Mascher, T., J. D. Helmann, and G. Unden. 2006. Stimulus perception in bacterial signal-transducing histidine kinases. Microbiol. Mol. Biol. Rev. 70:910-938.[Abstract/Free Full Text]
    39. 39
  39. Matsumoto-Nakano, M., K. Fujita, and T. Ooshima. 2007. Comparison of glucan-binding proteins in cariogenicity of Streptococcus mutans. Oral Microbiol. Immunol. 22:30-35.[Medline]
    40. 40
  40. McNeill, K., and I. R. Hamilton. 2003. Acid tolerance response of biofilm cells of Streptococcus mutans. FEMS Microbiol. Lett. 221:25-30.[CrossRef][Medline]
    41. 41
  41. McNeill, K., and I. R. Hamilton. 2004. Effect of acid stress on the physiology of biofilm cells of Streptococcus mutans. Microbiology 150:735-742.[Abstract/Free Full Text]
    42. 42
  42. Miller, A. A., N. C. Engleberg, and V. J. DiRita. 2001. Repression of virulence genes by phosphorylation-dependent oligomerization of CsrR at target promoters in S. pyogenes. Mol. Microbiol. 40:976-990.[CrossRef][Medline]
    43. 43
  43. Mitchell, T. J. 2003. The pathogenesis of streptococcal infections: from tooth decay to meningitis. Nat. Rev. Microbiol. 1:219-230.[CrossRef][Medline]
    44. 44
  44. Napimoga, M. H., J. F. Hofling, M. I. Klein, R. U. Kamiya, and R. B. Goncalves. 2005. Transmission, diversity and virulence factors of Streptococcus mutans genotypes. J. Oral Sci. 47:59-64.[CrossRef][Medline]
    45. 45
  45. Peterson, P. 2003. The world oral health report 2003. Oral Health Programme, Noncommunicable Disease Prevention and Health Promotion, World Health Organization, Geneva, Switzerland.
    46. 46
  46. Pitts, N. B., I. G. Chestnutt, D. Evans, D. White, B. Chadwick, and J. G. Steele. 2006. The dentinal caries experience of children in the United Kingdom, 2003. Br. Dent. J. 200:313-320.[CrossRef][Medline]
    47. 47
  47. Rolerson, E., A. Swick, L. Newlon, C. Palmer, Y. Pan, B. Keeshan, and G. Spatafora. 2006. The SloR/Dlg metalloregulator modulates Streptococcus mutans virulence gene expression. J. Bacteriol. 188:5033-5044.[Abstract/Free Full Text]
    48. 48
  48. Steiner, K., and H. Malke. 2002. Dual control of streptokinase and streptolysin S production by the covRS and fasCAX two-component regulators in Streptococcus dysgalactiae subsp. equisimilis. Infect. Immun. 70:3627-3636.[Abstract/Free Full Text]
    49. 49
  49. Steiner, K., and H. Malke. 2000. Life in protein-rich environments: the relA-independent response of Streptococcus pyogenes to amino acid starvation. Mol. Microbiol. 38:1004-1016.[CrossRef][Medline]
    50. 50
  50. Sumby, P., A. R. Whitney, E. A. Graviss, F. R. DeLeo, and J. M. Musser. 2006. Genome-wide analysis of group A streptococci reveals a mutation that modulates global phenotype and disease specificity. PLoS Pathog. 2:e5.[CrossRef][Medline]
    51. 51
  51. Ullman, R. F., S. J. Miller, M. J. Strampfer, and B. A. Cunha. 1988. Streptococcus mutans endocarditis: report of three cases and review of the literature. Heart Lung 17:209-212.[Medline]
    52. 52
  52. Varughese, K. I. 2002. Molecular recognition of bacterial phosphorelay proteins. Curr. Opin. Microbiol. 5:142-148.[CrossRef][Medline]
    53. 53
  53. Vilela, C., and J. E. McCarthy. 2003. Regulation of fungal gene expression via short open reading frames in the mRNA 5' untranslated region. Mol. Microbiol. 49:859-867.[CrossRef][Medline]
    54. 54
  54. Vilela, C., C. V. Ramirez, B. Linz, C. Rodrigues-Pousada, and J. E. McCarthy. 1999. Posttermination ribosome interactions with the 5'UTR modulate yeast mRNA stability. EMBO J. 18:3139-3152.[CrossRef][Medline]

Journal of Bacteriology, July 2008, p. 4478-4488, Vol. 190, No. 13
0021-9193/08/$08.00+0     doi:10.1128/JB.01961-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.



This article has been cited by other articles:

  • Lemos, J. A., Burne, R. A. (2008). A model of efficiency: stress tolerance by Streptococcus mutans. Microbiology 154: 3247-3255 [Abstract] [Full Text]  

This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Download to citation manager
Right arrowReprints and Permissions
Right arrow Copyright Information
Right arrow Books from ASM Press
Right arrow MicrobeWorld
Citing Articles
Right arrow Citing Articles via HighWire
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Chong, P.
Right arrow Articles by Biswas, I.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Chong, P.
Right arrow Articles by Biswas, I.

Copyright © 2009 by the American Society for Microbiology.   For an alternate route to Journals.ASM.org, visit: http://intl-journals.asm.org | More Info»