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J Bacteriol. Jun 2005; 187(12): 4064–4076.
PMCID: PMC1151735

A VicRK Signal Transduction System in Streptococcus mutans Affects gtfBCD, gbpB, and ftf Expression, Biofilm Formation, and Genetic Competence Development


Bacteria exposed to transient host environments can elicit adaptive responses by triggering the differential expression of genes via two-component signal transduction systems. This study describes the vicRK signal transduction system in Streptococcus mutans. A vicK (putative histidine kinase) deletion mutant (SmuvicK) was isolated. However, a vicR (putative response regulator) null mutation was apparently lethal, since the only transformants isolated after attempted mutagenesis overexpressed all three genes in the vicRKX operon (Smuvic+). Compared with the wild-type UA159 strain, both mutants formed aberrant biofilms. Moreover, the vicK mutant biofilm formed in sucrose-supplemented medium was easily detachable relative to that of the parent. The rate of total dextran formation by this mutant was remarkably reduced compared to the wild type, whereas it was increased in Smuvic+. Based on real-time PCR, Smuvic+ showed increased gtfBCD, gbpB, and ftf expression, while a recombinant VicR fusion protein was shown to bind the promoter regions of the gtfB, gtfC, and ftf genes. Also, transformation efficiency in the presence or absence of the S. mutans competence-stimulating peptide was altered for the vic mutants. In vivo studies conducted using SmuvicK in a specific-pathogen-free rat model resulted in significantly increased smooth-surface dental plaque (Pearson-Filon statistic [PF], <0.001). While the absence of vicK did not alter the incidence of caries, a significant reduction in SmuvicK CFU counts was observed in plaque samples relative to that of the parent (PF, <0.001). Taken together, these findings support involvement of the vicRK signal transduction system in regulating several important physiological processes in S. mutans.

Among the hundreds of bacterial species that colonize and persist in the oral cavity, Streptococcus mutans is among the few species that have been consistently linked with caries formation (31). Under the low-pH conditions that define the plaque environment, S. mutans induces an acid tolerance response that helps it survive (14, 15, 18). Hence, the ability to adapt to and generate acids at the tooth surface allows S. mutans to predominate within carious lesions by reducing the plaque pH to levels that are inhibitory to other oral microbes (34, 47). In addition, the ability of S. mutans to synthesize extracellular polysaccharides that promote formation of the plaque biofilm also contributes to its pathogenicity (35, 47). As a result, investigations to characterize virulence determinants of this oral pathogen have continued to dominate the caries microbiology field for decades.

Two-component signal transduction systems (TCSTS) are among the regulatory networks that are essential for bacterial adaptation, survival, and virulence. These systems function as “molecular switches” to modulate gene expression in response to changes in the external environment (44). Typically, signal transduction is accomplished via two regulatory elements consisting of a membrane-associated histidine kinase and a cytoplasmic response regulator. Upon exposure to an environmental cue, such as pH, osmolarity, or oxidation-reduction potential, the histidine kinase becomes autophosphorylated at a conserved histidine residue. Following the transfer of this phosphate group to a response regulator, the regulator can control the transcription of target genes by binding to their promoter regions. Diverse metabolic processes controlled by TCSTS include chemotaxis, sporulation, quorum sensing, and antibiotic/bacteriocin production in a wide variety of bacteria (3, 8, 30, 43, 45). Previous work conducted in one of our laboratories indicated that genetic competence, biofilm formation, and acid tolerance are mediated by detection of a signal peptide by the comDE TCSTS in S. mutans (27, 28). Genetic competence enables recipient bacteria to inherit heterologous genes that can contribute to the emergence of antibiotic resistance, as well as promote genetic variation that can drive overall fitness and evolution (7).

Analysis of the S. mutans UA159 genome database revealed 13 putative TCSTS and at least one independent response regulator (1, 21). The present study describes an investigation into the S. mutans vicRK signal transduction system that encodes a putative histidine kinase (VicK) and a response regulator (VicR) (Fig. (Fig.1A).1A). This TCSTS in S. mutans was first described as covRS by Lee et al. (GenBank accession number AF393849), after its homolog in Streptococcus pyogenes (26). While this system does bear sequence similarity to the covRS system of S. pyogenes, it appears more closely related to the vicRK TCSTS of S. pyogenes and Streptococcus pneumoniae. To clarify this further, the S. pyogenes covR homolog in S. mutans was named gcrR by Sato et al. (40) and later renamed tarC following its characterization by Idone et al. (21). Figure Figure1B1B clarifies the relationship between various members of these TCSTS families. Based on these observations, it seems more appropriate to refer to this TCSTS as vicRK as previously named in the annotated S. mutans UA159 genome sequence available in the National Center for Biotechnology Information (NCBI) GenBank database (1). In this paper, we chose to henceforth refer to the TCSTS as vicRK in accordance with its designation by Ajdic et al. (1).

FIG. 1.
(A) Genetic map of the vicRKX operon. Using BlastP searches, putative functions were assigned to genes based on high identity scores in the NCBI website. Abbreviations: smc, chromosome segregation SMC protein in Streptococcus agalactiae (accession no. ...

The vicRK TCSTS family is best characterized in S. pneumoniae and in Bacillus subtilis (10, 12, 20, 46). In a study conducted by Wagner et al., overexpression of the vicK gene in S. pneumoniae resulted in a mutant attenuated for virulence in a mouse model (46). Also, a vicK null mutant demonstrated a decrease in transformation efficiency (TE) relative to the wild type by approximately 3 orders of magnitude (46). Efforts to generate a deletion mutation in the S. pneumoniae vicR homolog (vicR), however, proved unsuccessful. In contrast to the recent publication by Lee et al. (26), in this study we claim that vicR inactivation in S. mutans is also lethal. Herein we describe a link between vicRK signal transduction and S. mutans sucrose-dependent adhesion, biofilm formation, and competence development. We also report our assessment of the abilities of an S. mutans vicK-deficient mutant to form plaque and generate caries in a specific-pathogen-free rat model.


Bacterial strains, plasmids, and media.

The bacterial strains, plasmids, and amplicons used in this study are listed in Table Table1.1. The S. mutans UA159 wild-type strain and its derivatives were routinely maintained on Todd-Hewitt yeast extract (THYE) agar (BBL Becton Dickinson, Cockeysville, MD) containing appropriate antibiotics when needed. Antibiotics used for the mutant strains were erythromycin (10 μg/ml), spectinomycin (1,200 μg/ml), kanamycin (500 μg/ml), and tetracycline (10 μg/ml). All S. mutans cultures were routinely grown as standing cultures at 37°C in a 5% CO2-95% air mixture.

Bacterial strains, plasmids, and amplicons used in this study

Construction of S. mutans vicRK mutants.

We searched the S. mutans UA159 genome database (http://www.genome.ou.edu/smutans.html) for vicR and vicK homologs using the S. pneumoniae RR02 and HK02 amino acid sequences as queries, respectively. To delete the vicRK gene pair in S. mutans UA159, we used a ligation-PCR mutagenesis strategy as previously described (25). The resulting putative histidine kinase mutant was termed SmuvicK, while repeated attempts to generate a vicR null mutant in the NG8 and UA159 backgrounds proved to be futile. In the latter case, all recovered transformants overexpressed the vicRKX genes and were hence designated Smuvic+. The primers used for mutant construction and confirmation are listed in Table Table2.2. Ligation constructs were introduced into S. mutans by competence-stimulating peptide (CSP)-induced natural transformation (28), and transformants resistant to erythromycin were selected to confirm appropriate recombination into the chromosome by PCR, followed by nucleotide sequence analysis. The vicRKX expression in the resulting mutants was monitored using quantitative real-time PCR (rtPCR) and compared with expression in the UA159 wild-type progenitor.

Primers used for PCR-ligation mutagenesis, rtPCR, VicR cloning, and mobility shift experiments

Growth rates.

Growth kinetics were monitored using a Bioscreen microbiology reader (Bioscreen C Labsystems, Helsinki, Finland). Overnight cultures were diluted 20× in fresh THYE and grown to an optical density at 600 nm (OD600) of approximately 0.4 to 0.5. Twenty microliters of mid-log-phase cells for the mutant and wild-type strains were inoculated in triplicate into microtiter plate wells containing 400 μl of THYE. Wells containing uninoculated THYE were used as controls. Using Biolink software (Labsystems), the Bioscreen reader was programmed to monitor OD600s at 37°C every 20 min for 24 h, with moderate shaking every 3 min. OD600 measurements were plotted against time to generate growth curves.

S. mutans biofilm formation.

A modified semidefined minimal medium (SDM) was prepared for biofilm growth experiments as described previously (29, 32). Biofilms were formed in 24-well polystyrene microtiter plates containing 2 ml of medium supplemented with 20 mM glucose or 10 mM sucrose. All wells were inoculated with 20 μl of an overnight cell suspension. In addition to SDM, SmuvicK and UA159 biofilms were also formed in 0.25× THYE that was supplemented with 20 mM glucose or 10 mM sucrose. Following incubation at 37°C and 5% CO2 for 16 h, the broth was gently removed by aspiration and the biofilms photographed directly. To closely examine the architecture of the parent and mutant biofilms, we utilized scanning electron microscopy (SEM) as described previously (28).

RNA preparation and rtPCR analysis.

To measure gtfB, gtfC, and ftf expression, total RNA was isolated from bacterial cultures grown in Tryptone yeast extract broth supplemented with 1% sucrose or 1% glucose. To study vicRK expression, bacterial strains were grown in THYE with or without antibiotics. Overnight cultures were diluted 20× in fresh broth and then grown to mid-logarithmic phase. Cells were harvested by centrifugation and immediately resuspended in Trizol reagent (Invitrogen) prior to RNA isolation using the FastPrep system (Bio 101 Savant) as specified by the manufacturer. To monitor gene expression, total RNA was subjected to DNase treatment and then reverse transcribed using a first-strand cDNA synthesis kit (MBI Fermentas) in accordance with the recommendations of the supplier. Controls for cDNA synthesis included a condition with no RNA template and another without reverse transcriptase. Finally, the single-stranded cDNAs were incorporated into rtPCR experiments using a Cepheid Smart Cycler system (Cepheid, Sunnyvale, CA) and a Quantitect SYBR-Green PCR kit (QIAGEN). Each 25-μl reaction mixture included template cDNA, 25 μM each primer, and 2× SYBR-Green mix (containing SYBR-Green, deoxynucleoside triphosphates, MgCl2, and Hotstar Taq polymerase). For maximum efficiency, rtPCR primers were designed to generate amplicons ranging from 100 to 170 bp in size (Table (Table2).2). Controls for rtPCR included reaction mixtures without template cDNA to effectively rule out the presence of contaminating DNA and/or the formation of primer dimers. The cycling conditions were as follows: 95°C for 15 min for the initial denaturation, followed by 35 to 40 cycles of three steps consisting of denaturation at 94°C for 15 s, primer annealing at the optimal temperature (Table (Table2)2) for 30 s, and primer extension at 72°C for 30 s. For each set of primers, cycle threshold (Ct) values, defined as the first cycle that gave rise to a detectable PCR product above the background, were generated. Known genomic DNA concentrations were used to generate Ct values for specific primer sets. By plotting the DNA concentrations versus the Ct value, standard curves were generated and used to determine relative RNA expression levels for the test gene. Results were normalized against S. mutans gyrA expression that was invariant under the experimental test conditions.

Purification of MBP-VicR.

The vicR coding sequence was amplified by PCR using chromosomal DNA derived from S. mutans UA159 with primers oSG241 and oSG242. Subsequently, the amplicon was digested with HindIII and EcoRI and ligated to maltose-binding protein (MBP) expression vector pMalc2 (New England Biolabs, Beverly, MA). The resulting plasmid, pSG385, was introduced into Escherichia coli strain TB1 and selected for resistance to ampicillin. For overexpression, E. coli strain TB1 cells containing pSG385 were grown in 1 liter of LB supplemented with 2% glucose and 100 μg/ml ampicillin at 37°C with shaking. When cells reached an OD600 of 0.3 to 0.5, expression of the MBP fused to VicR (MBP-VicR) was induced with 0.3 mM isopropyl-β-d-thiogalactopyranoside (IPTG) for 2 h. The cells were harvested by centrifugation (4°C, 5,000 × g, 15 min), resuspended in column buffer (20 mM Tris-HCl, pH 7.4, 200 mM NaCl, 1 mM EDTA), frozen at −20°C overnight, and lysed by sonication. Cells that were not lysed along with other debris were removed by centrifugation (9,000 × g, 20 min, 4°C). The cleared cell lysate was applied to an amylose resin column (New England Biolabs), preequilibrated with column buffer, and then washed with 12 column volumes of column buffer. The protein was eluted with column buffer containing 10 mM maltose. Three-milliliter fractions were collected, and fractions containing MBP-VicR, as determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis, were pooled and concentrated using an Amicon Ultra-15 concentrator (Millipore, Billerica, MA). Briefly, fractions containing MBP-VicR were applied to the filter device and centrifuged at 3,000 × g and 4°C for 15 min. To reduce the salt concentration, the filter was washed with modified column buffer (20 mM Tris-HCl, pH 7.4, 50 mM NaCl, 1 mM EDTA) and the purified protein was concentrated to ~2 ml. The concentration of MBP-VicR was determined by using the Bio-Rad Protein Assay (Bio-Rad, Hercules, CA) using bovine serum albumin as the standard. The purified protein was stored at 4°C or in glycerol at −80°C until needed.

Mobility shift experiments.

The primers oSG181, oSG258, and oSG64 were end labeled by incubating 1 μM primer with 1 μM [γ-32P]ATP, 0.5 μM unlabeled ATP, T4 polynucleotide kinase (Promega, Madison, WI), and 1× polynucleotide kinase buffer at 37°C for 30 min. The labeled oSG181 primer was then used to generate a 194-bp gtfC promoter-containing fragment using oSG137. The labeled oSG258 primer was then used to generate a 215-bp ftf promoter-containing fragment using oSG265. Labeled primer oSG64 and unlabeled primer oSG264 were used to generate a 205-bp gtfB promoter-containing fragment. For the binding reactions, the labeled gtfB (−159 to +36), gtfC wild-type (−89 to +102), and ftf (−141 to +74) promoter-containing fragments were incubated at room temperature for 30 min with 0 to 1,000 nM MBP-VicR, reaction buffer (52.5 mM MOPS pH 7.4, 9.5% glycerol, 50 μM EDTA, 50 μg/ml bovine serum albumin), and 50 ng salmon sperm DNA in a volume of 20 μl. Protein-DNA complexes were separated by nondenaturing gel electrophoresis on 6% acrylamide gels at 10 V/cm for 3 h, dried, and visualized with a phosphorimager.

Extracellular polysaccharide synthesis (EPS).

Cell cultures at mid-logarithmic phase derived from 1:20 dilutions of overnight cultures were centrifuged for 10 min at 4,500 rpm. To measure “released” activity in the culture fluid, the supernatants were filter sterilized and stored at −20°C until further use. EPS assays were conducted by adding 50 μl of a buffer mixture (100 mM sodium acetate buffer [pH 5.5], 7 mM sodium fluoride, 0.02% dextran T-10 [average weight, 10,000]) to 200 μl of cell-free supernatants. Following their incubation at 37°C for 10 min, 0.6 mM [14C]sucrose (11 μCi/μmol) was added to the mixtures, which were then vortexed, and 15 μl of each mixture was spotted in triplicate onto 2.3-cm square Whatman 3 MM filter papers. This procedure was repeated after mixtures were incubated at 37°C for 30 min. Subsequently, the filter papers were washed three times, 15 min each, in methanol using at least 10 ml of solvent. Filter squares were then dried and radioactivity counted using a liquid scintillation counter. Net EPS activity was measured as the difference in counts at t = 30 and t = 0.

Competence assay.

Overnight cultures of SmuvicK, Smuvic+, and their UA159 parent were diluted 20- or 40-fold in prewarmed THYE and incubated at 37°C until an OD600 of approximately 0.3 was reached. Following the incubation period, 1 μg of closed circular plasmid DNA, pDL277, Specr (5), was added and the samples were divided into two aliquots, only one of which was supplemented with synthetic CSP (sCSP) (Hospital for Sick Children Biotechnology Services, Toronto, Ontario, Canada) at a final concentration of 750 ng/ml (28). To study genetic competence in the mutant and parent strains, we performed TE assays as described previously (28).

Examination of the vicK deficient strain, in vivo, for plaque formation and cariogenic potential.

Specific-pathogen-free, caries-susceptible Osborne-Mendel rats (Center for Dental and Oral Medicine and Cranio-Maxillofacial Surgery, Zurich, Switzerland) were used to investigate in vivo effects of the vicK deletion on smooth-surface dental plaque and smooth and fissure caries, as well as on the establishment of S. mutans in the oral microbiota. Each experimental group consisted of 10 animals. Thirteen days after birth, the animals were transferred to stainless steel screen bottom cages without bedding and fed a finely ground stock diet (diet no. 3433; Provimi Kliba AG). Tap water and food were available ad libitum. On day 20 after birth, the dams were removed and the littermates distributed among the treatment groups (10 rats/group). On days 21 and 22, each rat was infected orally, twice daily, using 200 μl of a heavy bacterial suspension that comprised the parent UA159 strain or the vicK deletion mutant. To support the implantation of these bacteria, all rats received drinking water containing 2% sucrose and 2% glucose during days 20 to 22, as well as low-cariogenic diet 2000a (consisting of 28% skim milk, 15% powdered sucrose, 49% wheat flour, 5% brewer's yeast, 2% Gevral protein, and 1% sodium chloride). On day 23 following the association period, sterilization of the feeding and housing equipment was continued. Necessary precautions were taken to avoid cross-contamination and maintain a clean environment. Five days after association with the test strains, swabs were taken from the oral cavities of five rats per treatment group to confirm that the bacteria had become established. Shortly before the end of the study, oral swabs were taken from all 20 rats to obtain a final indication of the microbial status of the animals.

On day 51 (at the end of the 27-day experimental period), the animals were sacrificed. The upper and lower jaws were dissected and immersed in fixative (10% buffered formalin phosphate) for a minimum of 72 h. Erythrosin-stained maxillary molars were evaluated for plaque extent using a method described previously (39). Smooth-surface carious lesions were scored according to Keyes (22), and mandibular molars were sectioned and scored for fissure caries as specified by König et al. (24). The data were analyzed by two-way analysis of variance and least-significant-difference (LSD) tests using the analysis-of-variance statistics program.

Microbiological analyses of rat samples.

The swabs taken from each animal were immersed in sterile test tubes containing phosphate-buffered saline and thoroughly shaken. An aliquot was used for culture analyses, and the remaining suspension was immediately frozen. Sample dilutions were plated onto Trypticase yeast Columbia blood (TYCB) agar using a spiral dilutor to obtain total floral counts. Dilutions were also plated onto Trypticase yeast agar supplemented with 20% sucrose and bacitracin to enumerate S. mutans bacteria. Both the parental and mutant strains were also plated onto TYCB agar containing 10 μg/ml erythromycin as a control for possible contamination.


Structural organization of the S. mutans vicRKX locus.

The S. mutans vicRKX genes span a region of DNA from 1444056 to 1446908 bp (NCBI) on the minus strand of the S. mutans UA159 chromosome (Fig. (Fig.1A).1A). The vicK coding sequence is 1,350 bp in size and encodes a putative histidine kinase sensor protein with a predicted mass of 51,686 Da (450 amino acids [aa]). A BlastP search revealed that VicK shares high similarity with the S. pyogenes VicK protein (NP_268803) and with a putative histidine kinase in S. pneumoniae (HK02, NP_358699). The vicR coding sequence is 705 bp in size and encodes a putative response regulator protein with a predicted molecular mass of 26,900 Da (235 aa). A BlastP search of VicR revealed that it shared high similarity with the S. pyogenes VicR protein (NP_268803.1) and with other response regulators in S. agalactiae (NP_687734) and S. pneumoniae (NP_606786.1). The vicX coding sequence is 801 bp in size and codes for a hypothetical protein with a predicted mass of 29,680 Da (267 aa). A BlastP search indicated that S. mutans VicX shares amino acid similarity with VicX in S. pneumoniae (NP_345691), VicX in S. pyogenes (NP_268804.1), and a metallo-β-lactamase superfamily protein in S. agalactiae (NP_687736).

Confirmation of the S. mutans vic mutants.

To characterize the putative role of vicR and vicK in S. mutans, we constructed mutations within the vicRK coding sequences. The SmuvicK deletion mutation was confirmed by PCR analysis (results not shown). In contrast, we were unable to isolate a vicR null mutant. Sequence analysis of transformants revealed the presence of an intact vicR gene, as well as chromosomal integration of a VicR fragment that was used to mediate allelic exchange 5′ to the vicRKX operon (Table (Table1).1). Hence, instead of the expected double-crossover event, the presence of the intact vicR gene can be attributed to a Campbell-type crossover event mediated by the circularized a VicR fragment. To assess vicRK-specific expression in SmuvicK and Smuvic+ relative to the wild type, we performed rtPCR experiments, the results of which indicated no vicK-specific expression in the SmuvicK deletion mutant. rtPCR results for vic gene expression are shown in Fig. Fig.22.

FIG. 2.
S. mutans UA159 and Smuvic+ cDNAs were used to study expression of the vicRKX genes using quantitative rtPCR. For each strain, cDNA samples derived from three independent experiments were subjected to amplification using vicRKX-specific primers ...

vic mutants have altered growth rates.

Growth curve analysis of the SmuvicK and Smuvic+ strains revealed altered growth rates compared with that of the UA159 parent strain (Fig. (Fig.3).3). While an overnight culture of UA159 grew as a uniformly turbid cell suspension, SmuvicK and Smuvic+ cells aggregated and accumulated at the bottom of the glass tubes (Fig. (Fig.4).4). Notably, a denser cell aggregate was apparent for the SmuvicK mutant compared with that of Smuvic+. This observation was consistent with growth curves derived from each of three independent experiments that demonstrated a higher growth yield for SmuvicK during stationary phase compared with the wild-type and Smuvic+ strains (Fig. (Fig.3).3). While SmuvicK revealed little difference in the exponential-phase growth rate (mean [min/generation] ± standard error, 59.1 ± 2.6) relative to the wild type (50.1 ± 2), the doubling time of Smuvic+ was greatly increased (78.5 ± 1.5).

FIG. 3.
Growth curves of S. mutans UA159, SmuvicK, and Smuvic+ in the presence of THYE. Each datum point is the average of three independent OD values per sample. The results shown are representative of two other independent experiments conducted with ...
FIG. 4.
Biofilm formation (top) and overnight cultures (bottom) of S. mutans UA159, SmuvicK, and Smuvic+. Significant phenotypic differences were apparent in the mutant biofilms formed in the presence of modified SDM supplemented with glucose compared ...

Vic mutant strains form aberrant biofilms.

Based on biofilms formed on microtiter plates, the mutant biofilms were remarkably different compared with that of the parent (Fig. (Fig.4).4). Specifically, UA159 biofilms presented a smooth and even appearance, whereas SmuvicK and Smuvic+ biofilms appeared rough and clumpy, with the latter seemingly less dense overall. As anticipated, UA159 wild-type cells supplemented with sucrose as the sugar source generated thicker biofilms that were strongly attached to the abiotic surface compared with those generated in glucose-containing medium. In contrast to the wild-type biofilms generated in either glucose or sucrose, growth of SmuvicK in the presence of sucrose gave rise to biofilms that were easily disrupted and only loosely adherent to polystyrene. As shown at the bottom of Fig. Fig.5,5, relative to the wild-type strain, only a fraction of the SmuvicK cells attached to the surface. However, this easily dispersed phenotype was not apparent when SmuvicK biofilms were formed in glucose-supplemented medium (Fig. (Fig.5,5, top). Closer examination of the SmuvicK biofilm architecture derived from growth in SDM supplemented with glucose revealed areas of densely packed cells interspersed among “open” areas devoid of cells (Fig. (Fig.5,5, top). Notably, closer examination (×5,000 and ×50,000 magnifications) of SmuvicK biofilms derived from growth in diluted 0.25× THYE supplemented with sucrose revealed abnormally long chains of cells that were often detached at the cellular junctions. Compared with the wild type, these mutant cells were generally larger and seemingly associated with a rougher surface that we interpreted as thicker deposits of exopolymer (Fig. (Fig.5,5, bottom). In addition, the Smuvic+ biofilm also revealed aggregated cell clusters that were readily apparent at a magnification of ×1,000.

FIG. 5.
(Top) SEM of S. mutans UA159, SmuvicK, and Smuvic+ developed in SDM supplemented with glucose. (Bottom) SEM of S. mutans UA159 and SmuvicK biofilms developed using 0.25× THYE supplemented with 10 mM sucrose.

The vic genes affect S. mutans gtfB, gtfC, gtfD, ftf, and gbpB expression.

Since SmuvicK biofilms grown in a sucrose-containing medium were altered in adherence compared with those grown in a medium supplemented with glucose, we hypothesized that the vic genes may influence S. mutans glucosyltransferase (gtfB, gtfC, and gtfD), fructosyltransferase (ftf), and/or gbpB expression. To test our hypothesis, we performed quantitative rtPCR using cDNAs derived from SmuvicK, Smuvic+, and UA159 and primers specific for the gtfB, gtfC, gtfD, ftf, and gbpB genes. Compared with the wild-type parent, the expression of all these genes was increased in the Smuvic+ mutant (Fig. (Fig.6).6). For example, the gtf genes were upregulated by more than fivefold in a medium supplemented with either glucose or sucrose (Fig. (Fig.6A),6A), whereas ftf and gbpB expression was increased by nearly fourfold and twofold, respectively (Fig. (Fig.6B).6B). In SmuvicK, relative to the wild type, expression of gtfD and gbpB was reduced, whereas gtfC expression remained unchanged (Fig. (Fig.6).6). Notably, in glucose-supplemented medium, expression of gtfB showed 3.7-fold more transcript relative to the parent strain.

FIG. 6.
(A) Expression of gtfBCD genes in S. mutans UA159, SmuvicK, and Smuvic+ strains. Gene expression was monitored by rtPCR using cDNAs derived from glucose (top)- or sucrose (bottom)-supplemented cultures. Results are the average of three independent ...

VicR binds specifically to the promoter regions of gtfB, gtfC, and ftf in vitro.

Overproduction of the VicR transcript in Smuvic+ and the concomitant upregulation of the gtf genes and ftf genes in this mutant suggested that VicR, like many response regulators, might act directly on the promoter regions for these genes. We therefore decided to examine VicR binding to the gtfBC and ftf promoter regions in vitro. We cloned the vicR coding sequence into an expression vector that allowed us to overexpress an amino-terminal MBP-VicR protein fusion. This construct was chosen since similar amino-terminal fusion constructs, like the CovR ortholog in S. pyogenes, were shown to bind indistinguishably from native CovR (11). The MBP-S. mutans VicR fusion was subsequently purified by >90% as revealed by Coomassie blue staining (data not shown) and used in electrophoretic mobility shift assays. As shown at the top of Fig. Fig.7,7, MBP-VicR bound the DNA of the promoter regions of gtfB, gtfC, and ftf (MBP purified from the vector alone failed to shift the gtfB and gtfC promoter regions; data not shown). In electrophoretic mobility shift assay reaction mixtures containing MBP-VicR and gtfC, salmon sperm DNA was added in 100-fold excess by weight without diminution of the shifted complex. There was, however, a hierarchy of binding. MBP-VicR bound with the highest affinity to the gtfC promoter. Although the apparent affinity of MBP-VicR was similar for the ftf and gtfB promoters, as judged by the proportion of DNA shifted, it was also clear that the shift of the gtfB promoter was more distinct and clearly visible, similar to the ftf promoter-containing complex. This suggested a greater site specificity of binding. Although we have yet to ascertain the exact sequence that MBP-VicR recognizes, it is likely that there are conserved sequence determinants within the given regions of all three promoter elements. Dubrac and Msadek recently identified a VicR consensus sequence that seems conserved across many gram-positive genera (9). As shown at the bottom of Fig. Fig.7,7, the sequence TGTWAHNNNNNTGTWAH (where W is A or T and H is A, T, or C) was perfectly conserved in the gtfC promoter, partially conserved in the gtfB promoter (although there are several possible iterations), and found perfectly conserved in the ftf promoter but with the important caveat that there was an additional 10 bp or helical repeat that separated the more conserved hexamers.

FIG. 7.
(Top) Gel mobility shift assays. DNA-binding reaction mixtures were prepared with end-labeled DNA fragments for the 5′-proximal region of gtfB, gtfC, or ftf and purified MBP-VicR. Lanes 1, no protein; lanes 2, 100 nM; lanes 3, 200 nM; lanes 4, ...

The vic mutants affect extracellular polysaccharide formation.

Since the biofilm architecture and adhesive properties are affected by mutagenesis of the vic genes, we sought to gain insight into the synthesis of extracellular polysaccharides in SmuvicK, Smuvic+, and their parent UA159 strain. Based on the results of three independent experiments, cell-free supernatants of SmuvicK cultures resulted in a negative value for the percentage increase in 14C incorporated into dextran, possibly the result of degradation or removal of the newly synthesized dextran during the methanol washes (Fig. (Fig.8).8). In contrast, the rate of extracellular polysaccharide formation by Smuvic+ culture supernatants was increased relative to the wild type.

FIG. 8.
EPS by SmuvicK, Smuvic+, and their UA159 progenitor strain. Results shown are the averages of three independent experiments conducted to monitor the percentage of 14C incorporated into dextran following 30 min of incubation.

The vic mutants have altered transformation efficiencies.

Genetic competence assays indicated that SmuvicK and Smuvic+ have altered TE compared to the UA159 wild-type progenitor strain, thereby supporting a role for the vic gene products in competence development. In the absence of CSP, the TE of Smuvic+ was reduced by approximately 100-fold compared with the wild type, whereas the SmuvicK TE was not remarkably different from that of the wild type (data not shown). While addition of sCSP markedly increased the transformability of the wild-type strain by nearly 1,000-fold, the TEs of SmuvicK and Smuvic+ were only increased by approximately 20- and 88-fold, respectively (Fig. (Fig.9).9). In other words, the percent change in TE (with added CSP) decreased by 60-fold (n = 5) and 13-fold (n = 3) for SmuvicK and Smuvic+, respectively.

FIG. 9.
n-fold increases in transformation efficiencies of S. mutans UA159, SmuvicK, and Smuvic+ strains when supplemented with sCSP. TE of cultures without addition of CSP was set at a user-defined value of 1.0. The abilities of SmuvicK and Smuvic+ ...

SmuvicK is significantly increased in surface plaque formation and reduced in CFU count in vivo.

Compared with the UA159 parent strain, SmuvicK was significantly increased in its ability to form smooth-surface plaque (PF, <0.001; Table Table3)3) in a specific-pathogen-free rat model. However, the development of initial dentinal T lesions, advanced dentinal fissure lesions, and smooth-surface caries lesions were not significantly altered in the vicK-deficient mutant compared with the parent (Table (Table3).3). Although growth of SmuvicK and UA159 in blood agar was significantly different at a confidence interval of 95%, there were no significant differences in the total number of CFU between these strains in samples derived from rats (Table (Table4).4). While there was no visible growth of the parent strain on TYCB plates supplemented with erythromycin, data collected from TYCB plates without erythromycin revealed a significant reduction in CFU counts for SmuvicK compared with the parent strain (PF, <0.001; Table Table44).

Mean (per rat) smooth-surface plaque extent, initial and advanced dentinal fissure lesions, and smooth-surface cariesa
Mean (per rat) total flora on Columbia blood agar plates and CFU counts on TYCB plates containing bacitracin with and without erythromycin


The aim of this study was to investigate the VicRK signal transduction system and its effects on various virulence attributes of S. mutans. Similar to its ortholog in S. pneumoniae, S. mutans VicR seems essential for the viability of this bacterium. Based on our findings, the vic gene products appear to modulate adherence, biofilm formation, and genetic competence development in S. mutans. Notably, they regulate the expression of several virulence-associated genes affecting synthesis and adhesion to polysaccharides, including gtfBCD, ftf, and gbpB. Moreover, studies conducted utilizing the vicK-deficient mutant in specific-pathogen-free rats revealed a significant increase in smooth-surface plaque compared with the wild-type UA159 parent, whereas the incidence of dental caries was not affected.

The VicRK signal transduction system in S. mutans was recently mentioned in a study by Bhagwat et al. in which construction of a vicR knockout mutant proved to be futile (4). We experienced the same outcome when repeated attempts to construct an S. mutans vicR null mutation in the UA159 and NG8 wild-type strains resulted in loss of viability. Consistent with this finding is a report by Wagner et al. that indicated an inability to generate a deletion mutation in the S. pneumoniae vicR ortholog (46). Yet, Lee et al. recently published a report claiming to have inactivated the S. mutans vicR gene, called covR in their paper (26). Upon obtaining this mutant and examining the cDNAs generated from its RNA pool using three different primer sets that flank the vicR coding sequence, we noted transcription of the vicR gene at levels that were comparable to that of the parent (results not shown). In fact, if one examines the predicted integration site in the report of Lee et al., it is evident that the map of the locus shows insertion of the mutagenic construct 5′ proximal to the wild-type covR (vicR) gene. During our attempted mutagenesis of vicR, we were unable to demonstrate that vicR is absolutely required for viability. However, it is reasonable to assume that the vicR gene plays a vital role that is essential for the survival of S. mutans under our laboratory conditions. Transformants obtained during the mutagenesis of vicR showed overexpression of the vicRKX genes likely resulting from a promoter duplication caused by a Campbell-type insertion of the circularized VicR fragment. In contrast, S. mutans viability was not affected when vicK was disrupted. Hence, the phenotypic differences that we observed for SmuvicK and Smuvic+, which include adhesion, biofilm formation, and TE, can probably be attributed to their genotypic differences, although intercellular interactions involving cross talk between VicRKX and other signal transduction systems cannot be discounted.

The ability of S. mutans to colonize teeth is paramount to the initiation and progression of dental caries. Among the S. mutans surface-associated proteins that facilitate adherence and colonization are glucosyltransferases (GtfB, GtfC, and GtfD) and a fructosyltransferase (Ftf), which catalyze the cleavage of sucrose to synthesize extracellular glucan and fructan polymers, respectively (2, 16, 17, 19, 38, 41). GtfB and GtfC produce water-insoluble glucans, which function as adhesive molecules that anchor bacteria to the tooth pellicle (13, 36). Oral bacterial aggregation is also mediated by interactions between surface-associated glucan-binding proteins (Gbp) that adhere to glucans, thereby promoting plaque formation (33). Collectively, these enzymes serve an important role in the pathogenicity of S. mutans. For instance, rats harboring S. mutans gtfBCD- or ftf-deficient mutants proved to be hypocariogenic (6, 35, 41, 47). Also, systemic or mucosal immunization with GbpB was shown to induce protective immunity against dental caries in rats, indicating that GbpB may be an important target for the development of caries vaccines (42). In this study, we analyzed biofilms formed by vic mutants using SEM and visual examination of biofilms grown in microtiter plates. In sucrose-supplemented medium, we noticed that biofilms formed using UA159 wild-type cells were thicker and firmly attached to the surface, in contrast to those developed in the presence of glucose. In contrast, SmuvicK biofilms that formed in the presence of sucrose as the sugar source were loosely attached to the abiotic surface and easily disrupted compared with those derived from a glucose-supplemented medium, as well as wild-type biofilms derived from glucose- or sucrose-containing medium. Therefore, our findings support vicRK as a regulator of sucrose-mediated adherence in S. mutans. We henceforth conducted quantitative rtPCR experiments to assess the expression of gtfBCD, ftf, and gbpB in SmuvicK and Smuvic+ cells grown in glucose- and/or sucrose-containing medium. Our results indicated that vicK acts as a positive regulator of ftf, gtfD, and gbpB expression. In the presence of sucrose, increased expression of the gtfBC genes was observed only in the vic-overexpressing mutant. The down-regulation of the ftf, gtfD, and gbpB genes can possibly account for the easily detachable biofilm phenotype of the vicK-deficient mutant as a result of a reduction in its rate of total dextran formation. The EPS assay is indicative of the rate of formation of extracellular glucans and fructans produced by the activity of Gtf proteins and Ftf on sucrose but does not, however, differentiate between soluble and insoluble polymers. Repeated measurements of the rate of dextran formation in SmuvicK resulted in negative values. It is possible that the type of dextran formed by this mutant is degraded or easily disturbed and removed during the methanol washes in the EPS assay protocol. In their publication, Lee et al. reported that CovR (VicR) negatively regulated glucose- and glucuronic acid-containing carbohydrate production (26). Although the CovR (VicR) mutant described by Lee et al. produced a covR transcript, their complementation studies conducted by supplying the mutant with multiple copies of the gene on a plasmid affected the proportion of glucose- and glucuronic acid-containing EPS, suggesting a relationship between this TCSTS and the type and proportion of EPS produced by S. mutans.

Since studying oral bacteria in their natural mode of growth (biofilms) is of enormous significance to understanding pathogenetic mechanisms, we sought to gain insight into the contribution of the vic genes in the formation of S. mutans biofilms. Relative to biofilms formed by the UA159 wild-type parent strain, the mutant biofilms showed altered architecture as judged by visual inspection of biofilms on microtiter plates and by SEM. Specifically, compared with wild-type biofilms that appeared smooth and composed of uniformly distributed streptococcal chains and intracellular spaces, mutant biofilms seemed to clump and to form cellular aggregates. SmuvicK biofilms demonstrated the highest variability, with cellular aggregates emerging from relatively large open areas devoid of cells. Some of the streptococcal chains appeared “curly” (Fig. (Fig.5,5, top), the likely result of aberrant cell division or abnormal carbohydrate polymer deposition at the cell surface. Its chains were unusually long and seemed disconnected at the cell junctions (Fig. (Fig.5,5, bottom), probably contributing to their easily disruptable biofilm phenotype. Similar to SmuvicK, Smuvic+ exhibited cell aggregates that projected outward in the shape of circular mounds from an otherwise evenly distributed biofilm architecture. Recently, Ng et al. demonstrated that the VicRK TCSTS in S. pneumoniae positively regulates expression of PcsB (37). PcsB acts as a cell wall hydrolase, and downregulation of PcsB results in defects in cell separation, synthesis, and morphology (37). Interestingly, the PcsB homolog in S. mutans is GbpB, which is positively regulated by the vic genes. Hence, if gbpB in S. mutans serves a similar function, the long streptococcal chains that seemed disconnected at cell junctions in the vicK-deficient mutant may be possibly caused by the down-regulation of gbpB in this mutant. Additional studies are warranted not only to understand the role of the vicR and vicX genes in regulating the expression of gbpB in S. mutans but also to define the role of gbpB as a cell wall hydrolase in S. mutans.

In reference to VicR, the results of the in vitro binding studies supported the interaction of MBP-VicR with the promoters of gtfB, gtfC, and ftf. While we were able to demonstrate VicR specificity for these regions, we have yet to identify the specific DNA sequences to which VicR binds. Dubrac and Msadek have described a consensus that accommodates our hierarchy of binding (9). In addition to providing a consensus consisting of a conserved hexamer separated by five nonspecific nucleotides, they demonstrated that at extremely high concentrations, the VicR homolog of Staphylococcus aureus can even bind to a single hexamer. According to our model, each conserved hexamer is recognized by a single VicR monomer and hence binding at a site with properly spaced hexamers, like gtfC, actually occurs as a homodimer. In the case of the ftf promoter sequence, we would argue that the active species is actually a dimer of dimers. Since the hexamers are in direct repeat with the first T's 11 bp apart, it is possible to have cooperative interactions and hence oligomerization, which has been observed for NtrC (48). Since TCSTS response regulators are typically regulated by their cognate histidine kinase, future experiments will need to examine the role of the phosphorylation state of VicR in DNA binding. We do not know the state of phosphorylation of the MBP-VicR fusion protein, nor do we know the effects of the presence of the amino-terminal fusion protein. This leaves us with a conundrum, as the consensus sequences actually overlap the promoters themselves. Since we have seen stimulation of gtfB, gtfC, and ftf transcription in the VicR overproducer, it stands to reason that transcription is enhanced by VicR binding to these promoters. Although the mechanism is unclear, there is a precedent. Lantibiotics are often self-regulated through a TCSTS quorum-sensing system. According to the model, as the lantibiotic concentration increases extracellularly, it binds to its cognate histidine kinase, resulting in phosphorylation of its partnered response regulator. Similar to our observations, the putative response regulator binding sites overlap the promoter regions of select genes just upstream of the −10 region (23). The specifics of this mechanism are unknown but clearly common among bacteria. Future experiments will focus on finer biochemical analysis of the VicR DNA binding interactions, including footprinting experiments that should identify the binding site.

In accordance with SmuvicK aberrant biofilm formation is the variant smooth-surface dental plaque content and CFU count noted for this mutant in vivo relative to the wild type. However, despite an increase in smooth-surface plaque, SmuvicK was not hypercariogenic in a specific-pathogen-free animal model relative to the wild type. One possibility is that the SmuvicK biofilm was easily disrupted, thereby reducing the virulence potential usually associated with increased smooth-surface plaque. Supporting this argument is the diminished SmuvicK viable count observed for this mutant on TYCB agar that was supplemented with bacitracin. Alternatively, one might speculate that the SmuvicK adherence defect was masked in vivo by the presence of other oral microbes that could have “nonspecifically” coaggregated with the mutant, anchoring it to the tooth surface. It is important to note that the main factors that affect cariogenicity include the microbial composition, the diet, and the nature of the polysaccharide matrix, which determines the diffusion properties of plaque. Hence, in reference to polymer production in SmuvicK, excess plaque extent or volume would not necessarily result in hypercariogenicity. Among other phenotypes observed for the vic mutants were alterations in genetic competence development in the presence or absence of CSP. The results of our experiments indicate that the efficiency with which S. mutans can take up foreign DNA is indeed affected by the products of the vic genes. In the absence of CSP, we observed that the TE for Smuvic+ was decreased by approximately 100-fold compared with the parent strain, whereas the TE was not necessarily altered in SmuvicK. The addition of CSP failed to increase the TE of either mutant to levels observed for the wild-type cells (Fig. (Fig.9).9). Previously, we described a comCDE quorum-sensing system in S. mutans that induces genetic competence (28). The comCDE genes encode the precursor CSP (ComC), its sensor protein (ComD), and a cognate response regulator (ComE). The absence of any one of these genes compromises TE. The reliance of the comCDE system on SmuvicK to restore the CSP-dependent wild-type TE levels suggests that the S. mutans vicK-initiated signal transduction system has a distinct regulatory effect on the competence development pathway. Although it is possible that VicK might act as a receptor for CSP in addition to ComD, this needs to be examined directly to test this assumption.

In summary, this work provides significant insight into important regulatory functions of the vicRK signal transduction system in S. mutans. However, more studies are warranted to define the downstream target genes that are regulated by this signaling pathway and the vicX gene. Deciphering the molecular mechanism(s) that underlies the vicRK signaling system in this oral pathogen can foster our understanding of virulence gene regulation in S. mutans and so reveal novel targets for therapeutics directed against S. mutans cariogenicity.


We thank Robert Chernecky for the SEM, Richard Mair and Peter Lau for assistance with bioinformatic analyses, and Song F. Lee for kindly providing the vicR (covR) mutant strain of S. mutans NG8.

This study was supported by NIH grant RO1DE013230 and CIHR grant MT-15431 to D.G.C., NIH grant R01DE013965 to S.D.G., and NIH grant R15DE014854 to G.A.S. D.G.C. is a recipient of a Canada Research Chair. M.D.S. is a CIHR Strategic Training Fellow supported by training grant STP-53877 and a Harron Scholarship.


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