Advertisement
Research Article

Phenotypic Heterogeneity of Genomically-Diverse Isolates of Streptococcus mutans

  • Sara R. Palmer,

    Affiliation: Department of Oral Biology, University of Florida, Gainesville, Florida, United States of America

    X
  • James H. Miller,

    Affiliation: Center for Oral Biology, University of Rochester Medical Center, Rochester, New York, United States of America

    X
  • Jacqueline Abranches,

    Affiliations: Center for Oral Biology, University of Rochester Medical Center, Rochester, New York, United States of America, Department of Microbiology and Immunology, University of Rochester Medical Center, Rochester, New York, United States of America

    X
  • Lin Zeng,

    Affiliation: Department of Oral Biology, University of Florida, Gainesville, Florida, United States of America

    X
  • Tristan Lefebure,

    Affiliations: Université de Lyon, CNRS, Ecologie des Hydrosystèmes Naturels et Anthropisés; Université Lyon, Villeurbanne, France, Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, New York, United States of America

    X
  • Vincent P. Richards,

    Affiliation: Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, New York, United States of America

    X
  • José A. Lemos,

    Affiliations: Center for Oral Biology, University of Rochester Medical Center, Rochester, New York, United States of America, Department of Microbiology and Immunology, University of Rochester Medical Center, Rochester, New York, United States of America

    X
  • Michael J. Stanhope,

    Affiliation: Population Medicine and Diagnostic Sciences, College of Veterinary Medicine, Cornell University, Ithaca, New York, United States of America

    X
  • Robert A. Burne mail

    rburne@dental.ufl.edu

    Affiliation: Department of Oral Biology, University of Florida, Gainesville, Florida, United States of America

    X
  • Published: April 16, 2013
  • DOI: 10.1371/journal.pone.0061358
Corrections

8 Nov 2013: Palmer SR, Miller JH, Abranches J, Zeng L, Lefebure T, et al. (2013) Correction: Phenotypic Heterogeneity of Genomically-Diverse Isolates of Streptococcus mutans. PLoS ONE 8(11): 10.1371/annotation/ffff8cd5-b8fa-4d3c-a993-e5169198f1e6. doi: 10.1371/annotation/ffff8cd5-b8fa-4d3c-a993-e5169198f1e6 | View correction

Abstract

High coverage, whole genome shotgun (WGS) sequencing of 57 geographically- and genetically-diverse isolates of Streptococcus mutans from individuals of known dental caries status was recently completed. Of the 57 sequenced strains, fifteen isolates, were selected based primarily on differences in gene content and phenotypic characteristics known to affect virulence and compared with the reference strain UA159. A high degree of variability in these properties was observed between strains, with a broad spectrum of sensitivities to low pH, oxidative stress (air and paraquat) and exposure to competence stimulating peptide (CSP). Significant differences in autolytic behavior and in biofilm development in glucose or sucrose were also observed. Natural genetic competence varied among isolates, and this was correlated to the presence or absence of competence genes, comCDE and comX, and to bacteriocins. In general strains that lacked the ability to become competent possessed fewer genes for bacteriocins and immunity proteins or contained polymorphic variants of these genes. WGS sequence analysis of the pan-genome revealed, for the first time, components of a Type VII secretion system in several S. mutans strains, as well as two putative ORFs that encode possible collagen binding proteins located upstream of the cnm gene, which is associated with host cell invasiveness. The virulence of these particular strains was assessed in a wax-worm model. This is the first study to combine a comprehensive analysis of key virulence-related phenotypes with extensive genomic analysis of a pathogen that evolved closely with humans. Our analysis highlights the phenotypic diversity of S. mutans isolates and indicates that the species has evolved a variety of adaptive strategies to persist in the human oral cavity and, when conditions are favorable, to initiate disease.

Introduction

The development of dental caries is a complex process that is primarily dependent on the presence of microbial biofilms, the composition and biochemical activity of the biofilm organisms, and the diet of the host; but is also affected by a variety of other factors that include the genetic constitution and behavior of the host, tooth architecture and exposure to fluoride [1][5]. Streptococcus mutans has long been acknowledged as the species of bacteria most closely associated with the initiation of dental caries [6], [7]. More recently, epidemiological [8] and mechanistic evidence for associations of certain sub-groups of S. mutans with cardiovascular disease have emerged [9], [10]. The three key virulence attributes of S. mutans that enable this organism to cause dental caries are the ability to form biofilms on the tooth, mediated by sucrose-dependent and sucrose-independent mechanisms [11]; production of organic acids via metabolism of dietary carbohydrates; and the ability to grow and to continue to produce acids in a low pH environment, known as aciduricity [6], [12]. In addition, the ability of S. mutans to rapidly adapt to environmental stresses appears to be central to its ability to form biofilms, persist in the host, and to compete with other oral bacteria, particularly when conditions are conducive to the development of dental caries [13]. Furthermore, some S. mutans strains are naturally competent for genetic transformation and are able to take up DNA from their environment [14]. Additionally, the competence pathway of S. mutans is linked to the production of bacteriocins, which kill susceptible closely related species, thus eliminating competitors while increasing the genetic material available for homologous recombination [15].

S. mutans is a diverse species of bacteria that can usually be classified into four different serological groups (c, e, f, and k) based on the composition of cell-surface rhamose-glucose polysaccharides [16]. Most strains isolated from the oral cavity (70–80%) are serotype c, with 20% composed of serotype e and 2–5% serotype f or k. However, specimens isolated from heart valves and atheromatous plaques have a higher occurrence of non-serotype c strains, with serotype k in higher proportions (12%) than in the oral cavity [17]. There have been several attempts to correlate carriage of certain genotypes of S. mutans with caries incidence, however there has been no consensus among multiple studies [18][23]. Additionally, it has been reported that there was no correlation between the caries status of an individual and the distribution of 41 putative virulence genes or genetic elements in 33 S. mutans isolates [24]. These authors [24] concluded that the virulence genes they tested might be part of the core genome of S. mutans, hence the lack of diversity in their distribution among strains.

Studies using comparative genomic hybridization (CGH) based on the UA159 genome [25] have shown a high degree of content variation among strains, with some isolates lacking up to 20% of the genes present in the reference strain UA159 [26], [27]. In particular there are variations in the presence and content of a 50-kb genomic island, TnSmu2, that contains genes for non-ribosomal peptide synthases (NRPS), polyketide synthases (PKS), and accessory proteins responsible for biosynthesis of mutanobactin, which appears to augment oxidative stress tolerance [28], [29]. Another study identified 122 sequence types (ST) out of 135 strains isolated from around the world [30] using multi-locus sequence typing (MLST) based on the partial gene sequence of 6 housekeeping genes from S. mutans [31]; further demonstrating the genetic diversity of this species and reinforcing the findings that there is not a consistent correlation of the presence of certain genotypes with geographic location or caries status.

While techniques like CGH and MLST, as well as numerous other genetic fingerprinting studies, have allowed researchers to interrogate genotype distribution and to gain an understanding of species diversity, they do not allow for genome-scale correlations of phenotype and genotype, nor can they facilitate functional genomic studies that are key to dissecting how gene content and context relate to the pathogenic potential of the organisms [13], [32]. This is especially true for an organism like S. mutans, which can become naturally competent and therefore has the potential for rapid genome diversification through lateral gene transfer [33].

Given the clear evidence for substantial genetic diversity in the species S. mutans and to better understand the gene content of this species as a whole (i.e. unique core and dispensable genes), completed draft genomes of 57 geographically- and genetically-diverse isolates of S. mutans were generated and analyzed [34]. Based on the sequence information, 15 isolates with a high degree of diversity in gene content were chosen for further phenotypic characterization and genetic analyses. This study represents the first step toward determining whether it is possible to correlate core and pan-genome composition with specific phenotypic characteristics that are associated with the virulence potential of S. mutans. The knowledge gained from these studies can be used to guide more detailed analysis, e.g. transcriptomic studies and future epidemiologic work, to facilitate methods for the control of S. mutans and other cariogenic bacteria. Further, the baseline information provided here establishes a resource that can be utilized to accelerate progress on S. mutans pathogenesis and control of dental caries, as well as certain systemic diseases associated with S. mutans and closely-related species.

Materials and Methods

Bacterial Strains, Media, and Growth Conditions

Isolates of S. mutans used in this study are listed in Table 1. All strains were stored in 25% glycerol at −80°C and freshly streaked on brain heart infusion (BHI) agar before each experiment. Routine cultures of S. mutans strains were inoculated from a single colony and grown in brain heart infusion (BHI) broth (Difco) at 37°C in a 5% CO2 atmosphere. For biofilm experiments, strains were grown in a semi-defined biofilm medium (BM) [35] supplemented with 20 mM glucose or sucrose. For monitoring of growth, overnight cultures from two separate colonies were sub-cultured 1:25 into fresh medium, grown to mid-exponential phase (OD600 = 0.5), and diluted 1:100 into fresh growth media: BHI pH 7.5; BHI that had been titrated to pH 5.5 with HCl; BHI containing 0.2 µM synthetic competence stimulating peptide (CSP) [36] or BHI containing 25 mM paraquat (methyl viologen; catalog no. M2254; Sigma). Growth was then monitored by dispensing 200 µl of the diluted cultures in duplicate into wells of a Bioscreen C plate with a sterile mineral oil overlay to reduce exposure to oxygen, unless otherwise indicated. Plates were incubated at 37°C for 24 to 48 h in a Bioscreen C lab system (Helsinki, Finland) with readings every 20 min after shaking for 10 sec. Doubling times were calculated as described elsewhere [37] and Student’s t-tests were performed to determine significant differences.

thumbnail

Table 1. Clinical isolates used in this study.

doi:10.1371/journal.pone.0061358.t001

Biofilm Assays

Biofilm development was measured in polystyrene 96-well (flat-bottom) cell culture clusters (Costar.595; Corning Inc., Corning, NY) as previously described [38], with the following modifications. Overnight cultures were sub-cultured 1:25 into fresh BHI and grown to mid-exponential phase (OD600 = 0.5–0.6). Each culture was then sub-cultured 1:100 into BM medium and 200 µl was aliquoted into four replicate wells, followed by incubation at 37°C in a 5% CO2 aerobic atmosphere for 48 h. Culture medium was removed by aspiration and wells were gently washed with 200 µl sterile deionized water. Subsequently, 50 µl of a 0.1% solution of crystal violet dissolved in 99% ethanol was applied to each well and incubated at room temperature for 15 min, followed by removal of the fluid by aspiration. Wells were washed twice with 200 µl of water as before and allowed to air dry. The plates were photographed and the wells were de-stained with 200 µl of an acetone:ethanol solution (2:8) for 30 min at room temperature. The de-staining procedure was repeated and the OD575 of the pooled de-staining solution was measured. Results are representative of duplicate assays. Significant differences were determined using Students t-test.

Autolysis Assay

Autolysis was measured as described elsewhere [39], [40], with the following modifications. Overnight cultures were sub-cultured 1:20 into fresh BHI and grown to late exponential phase (OD600 = 0.7). Cells were collected by centrifugation and washed twice in PBS. Cells were resuspended in autolysis buffer (20 mM potassium phosphate buffer, pH 6.5, 1 M KCl, 1 mM CaCl2, 0.04% sodium azide) to an OD600 of 1.0. The cell suspensions (300 µl) were applied to duplicate wells of a 100-well Bioscreen plate and OD600 was monitored at 44°C every 20 min for 10 h in a Bioscreen C lab system. Triplicate cultures of each strain were used.

Acid Killing Assay

The ability to survive a strong acid challenge was determined as previously described [41], with the following modifications. Briefly, cells from an overnight culture were diluted 1:25 into BHI broth and incubated to OD600 = 0.3 (unadapted) or to OD600 = 0.2 followed by a 2-hour incubation period in BHI broth that had been acidified with HCl to pH 5.0 (adapted). Cells were then collected by centrifugation at 3,800×g at 4°C, resuspended in 0.1 M glycine buffer, pH 7.4, and vortexed for 1 min. In order to disperse cell clumps, as several strains tended to aggregate, cells were sonicated for two 20-second cycles in a sonicating water bath at room temperature and placed on ice between cycles. Before the start of the assay, aliquots of cells were removed and placed on ice. The remaining cells were then pelleted and resuspended in an equal volume of 0.1 M glycine buffer, pH 2.8, and rotated continuously at room temperature. Duplicate aliquots were removed at 15, 30 and 60 min and diluted 1:10 in 10 mM Tris-HCL, pH 8.0, and placed on ice. Once the assay was complete, all aliquots were serially diluted in 10 mM Tris-HCL, pH 8.0, and plated on BHI agar followed by a 48 h incubation at 37°C in a 5% CO2 atmosphere. Percent survival for each time point was determined by dividing the CFUs of each time point by the initial CFUs multiplied by 100. Data represents the average of two separate experiments performed in duplicate.

Genetic Competence Assay

Overnight cultures were sub-cultured 1:20 into fresh BHI and grown to OD600 = 0.125. Synthetic competence stimulating peptide (CSP; [36]) was added to final concentrations of 0, 5, 20, 50, 100, or 200 nM. Cells were returned to a 37°C incubator for 15 min before 0.5 µg of the integration vector pBGE [42] was added. Cells were then incubated for 2.5 h and plated on BHI agar plates containing 10 µg ml−1 erythromycin. Induction of competence was evaluated after 48 h of incubation at 37°C in a 5% CO2 atmosphere. Competence induction was determined by comparing the number of resulting colonies of a particular strain as a function of the concentration of input CSP.

Western Blots

Cells from mid-exponential phase cultures (OD600 = 0.5) grown in BHI, were collected by centrifugation at 3,800×g for 10 min. The culture supernates were filtered through a 0.2 µm syringe filter and proteins from a 700 µl aliquot (standardized by OD600) were precipitated with an equal volume of 20% TCA overnight at −20°C. Precipitated proteins were washed once with 300 µl cold acetone and allowed to air dry before the pellets were resuspended in 50 µl TE (50 mM Tris-HCL, 1 mM EDTA, pH 7.5). The suspension was combined with 50 µl 2X SDS-PAGE sample buffer, boiled for 5 min and centrifuged at top speed for 3 min in a microcentrifuge. Cell-wall associated proteins were extracted by boiling cell pellets (adjusted to the same OD600) in 1X SDS-PAGE sample buffer for 5 min, followed by centrifugation to remove whole cells. Proteins from 25 µl of the culture supernates and cell-wall extracts were separated on 4–8% XT Criterion Tris-acetate gradient gels. Proteins were transferred to nitrocellulose for Colloidal Gold Total Protein Stain (Bio-Rad) or PVDF membranes for Western blotting with a rabbit anti-GtfB/C polyclonal antisera (a kind gift from W. H. Bowen, University of Rochester [43]). Western blots were reacted with a 1:500 dilution of the antisera and developed according to the supplier’s directions using the Amersham ECL Western blot kit.

Galleria Mellonella Virulence Assay

Stationary phase (16 h) S. mutans cultures grown in BHI at 37°C in 5% CO2 were diluted 1:20 into fresh BHI supplemented with 5% horse serum. Cultures were grown to OD600 = 0.6 and placed on ice for at least 30 minutes. Cultures were washed twice with an equal volume of sterile saline solution (0.9% NaCl) and adjusted to approximately 5×107 CFU/ml in sterile saline. Bacterial colony counts on trypticase soy agar (TSA) plates were used to confirm initial inocula. A negative control for infection was prepared using heat-killed OMZ175 (15 min at 75°C). G. mellonella larvae in the 4th–5th instar stages, sorted by weight (200 to 300 mg) and showing no signs of melanization were randomly chosen and kept at 4°C prior to injection. A 10-µl Hamilton syringe was used to inject 5-µl aliquots of bacterial inoculum into the hemocoel of each larva via the last left proleg. After injection, larvae were kept at 37°C under atmospheric conditions and survival was recorded at selected intervals. Kaplan-Meier killing curves were plotted and estimation of differences in survival were compared using the log-rank test. A P value ≤ 0.05 was considered significant. All data was analyzed with GraphPad Prism 5.0 software. In addition to cnm+ and cnm- control strains (OMZ175 and UA159, respectively), a cnm-inactivated mutant strain, OMZ175-cnm, was also included to serve as a control to monitor Cnm dependent larvae death [10].

PCR to Confirm Presence or Absence of Genes

PCR was used to confirm the absence of intact comC, comD and comE genes in Smu56 and Smu57. The following primers were used in a standard PCR using colonies of UA159 (positive control), Smu56 or Smu57 as template; comD, SP01F-2 (5′-GAATGAAGCCTTAATGATACTTT-3′) and SP01R (5′CTATTTTATTATTAGGAGTTGCTTGAATA 3′), comE, SP02F (5′ –ATGATTTCTATTTTTGTATTGGAAGAT-3′) and SP02R (5′-TCATTTTGCTCTCCTTTGATCAGCAAT-3′), comC, SP03F (5′-GGAGTATAAAATGAAAAAAACACTA-3′) and SP03R (5′-GCC-TATCTTATTTTCCCAAAGCTT-3′). To confirm the presence or absence of intact Smu.1008 and Smu.1009 in Smu81, primers SP26F (5′-GTAGAATAAAGTTATGCTAAAGCAA-3′) and SP26R (5′-GATTACGCAACAATCATAGCTGTTT-3′), were used in a standard colony PCR reaction.

Results

Selection of Sequenced Strains for Further Characterization

The strains utilized in this study are described in Table 1. To better understand the scope of possible phenotypes within the species S. mutans, strains from two geographically-diverse collections of clinical isolates [30], [44], [45] were sampled based on genetic diversity determined by WGS sequences [34]. Figure S1 depicts gene content differences between strains based on orthologs recovered across genomes via an all-versus-all BLASTP search combined with clustering using OrthoMCLS [46]. Strains were selected for further phenotypic characterization based on their clustering, the presence or absence of selected non-core genes, and preliminary observations of phenotypic behaviors.

Through genome sequence analysis we were surprised to discover that one of the isolates characterized in this study, Smu77/NV1996 [30], was actually a genetically-engineered derivative of S. mutans V403 containing insertionally-inactivated gtfBC, gtfD and ftf genes, and thus represents strain V1996 originally described by Munro et al. [47]. It has been confirmed that Smu77 is resistant to kanamycin, erythromycin and tetracycline and contains the aphA gene interrupting gtfB-gtfC [47], the tetM gene within the gtfD gene [48], and ermAB inserted within the gene for ftf [49]. V403 is a serotype c strain isolated from human blood at the CDC in Atlanta, GA and contains a 5.6 kb cryptic plasmid (pVA403) [50]. V403 has also been reported to contain the cnm gene and is able to bind type-1 collagen [51].

Stress Tolerance Varies Widely among Genetically-diverse Isolates

Growth curve analysis was performed for the 15 strains under various stress conditions and compared to the well-characterized reference strain UA159. Most strains grew at similar rates and to similar final optical densities under non-stressed conditions (BHI, pH 7.5, with an oil overlay) (Table 2), but Smu56 had an unusually long doubling time (129±4 min) under non-stressed conditions. Although all experiments were conducted with fresh isolates from freezer stocks, we noted that Smu56 was viable on agar plates stored at 4°C for only 2 to 3 days, whereas most of the other isolates remained viable for much longer.

thumbnail

Table 2. Growth characteristics of clinical isolates in different stress conditions.

doi:10.1371/journal.pone.0061358.t002

Aciduricity, the ability to grow and to continue to produce acids at low pH, is an important virulence attribute for S. mutans, and growth rate and final yield at pH 5.5 are considered to be good measures of aciduricity [52]. When strains were cultured in medium that had been acidified to pH 5.5 with HCl, substantial variation in growth characteristics were observed, with mean doubling times ranging from 157 to 328 minutes and maximum yields ranging from OD600 values of 0.25 to 0.61. Strain Smu44 grew at a significantly faster rate at pH 5.5 than strain UA159 (Td 157±7 compared to 172±5, P≤0.05). Smu21 also displayed consistently faster exponential growth than UA159, although the difference was not significant (164±5, P≤0.09). Despite the faster exponential growth of Smu21 and Smu44 at pH 5.5, the cell yields were less than for UA159 (OD600 max 0.43 and 0.53, respectively, compared to 0.60 for UA159).

The ability of Smu21 and Smu44 to survive a low pH challenge (pH 2.8) before and after acid adaptation at pH 5.0 for 2 hours was also compared (Figure 1). Both strains exhibited better survival over time at low pH compared to UA159 when cells had not been previously acid-adapted. In contrast, all strains performed similarly when they were first allowed to adapt to growth at low pH. This finding supports the idea that Smu21 and Smu44 have a greater constitutional resistance to acid stress than UA159, consistent with the growth curve data. When the ATPase activity of un-adapted and acid-adapted cells was compared between these strains, greater ATPase activity was consistently observed in acid-adapted cells compared to un-adapted (data not shown). However, when ATPase activity was compared between these strains no significant differences were seen under the conditions tested.

thumbnail

Figure 1. Acid killing - Percent survival after exposure to pH 2.8.

Cells were grown to mid-exponential phase, washed in 0.1 M glycine buffer pH 7.4 and re-suspended in 0.1 M glycine pH 2.8. Cells were removed at 15-, 30- and 60-minute intervals, serial diluted and plated. For acid adaptation, cells were harvested at early exponential phase and re-suspended in pH 5.0 media and grown for 2 additional hours. Percent survival was determined based on the number of cells at each time point divided by the number of cells before acid challenge, t0. Data represents the average of two separate experiments performed in duplicate.

doi:10.1371/journal.pone.0061358.g001

Oxidative stress tolerance is a critically important factor in the establishment, persistence and ecology of oral bacteria, and thus affects the pathogenic potential of oral biofilms in major ways [53], [54]. Organisms in the oral cavity are transiently exposed to different oxygen levels, and to different types and quantities of reactive oxygen species (ROS) generated in saliva and within oral biofilms [53], [54]. Sensitivity to oxygen was determined by examining growth in a Bioscreen C machine in the presence of air (no mineral oil overlay), revealing major differences in doubling times (86 to 218 min), with cell yields ranging from OD600 values of 0.46 to 0.67 (Table 2, O2 Air ODmax). When some strains (Smu81, Smu86, Smu104 and Smu109) reached stationary phase, a sharp drop in OD600 values occurred; followed by a resumption of growth with cell yields reaching similar values to the maximum yield at 48 hours (Figure S2). For example, strain Smu104 reached a maximum cell yield of 0.57 after about 6 hours of growth, the OD of the culture then declined to 0.29 after 15 hours, followed by regrowth of the culture to attain a final yield of OD600 = 0.62 at 48 h. Multiple other strains did not show this growth-lysis-regrowth cycle, displaying more typical plateaus in stationary phase, or a slow and steady decline in optical density during stationary phase.

Similar to cells exposed to air, growth in 25 mM paraquat, which can generate superoxide anion, yielded great variation in growth rates and final yields, with one strain (Smu81) unable to initiate growth in medium containing paraquat (Table 2). Unlike for cells growing in the presence of air, however, evidence of the stationary-phase lysis, or lysis and regrowth, was not observed in cells cultured in paraquat. It should also be noted that none of the strains were able to grow in the presence of paraquat unless the wells were overlaid with mineral oil.

Autolysis Varied among Clinical Isolates

Autolysis is a natural process whereby cells undergo lysis in response to an environmental signal; this process is important for biofilm formation, competence development and cell wall turnover [55]. In S. mutans, the peptidoglycan hydrolase AltA [40] has been shown to be a significant contributor to autolysis [56]. Autolysis has also been shown to stimulate eDNA accumulation, which can enhance biofilm formation and alter biofilm architecture [57]. In order to examine autolytic behavior within the sequenced isolates, the change in OD600 over time of late-exponential phase cells that were washed, resuspended in autolysis buffer, and incubated at 44°C was monitored (Figure 2). Smu44 was the most autolytic strain, displaying slightly more lysis than UA159, whereas Smu21 showed slightly less autolysis than UA159. Smu57 and Smu104 were the least autolytic of strains tested, with the remaining strains displaying an intermediate level of autolysis. All strains contained the altA gene, and in most cases the gene sequence was identical to that of UA159. There was no correlation between the autolysis phenotype of strains observed here and the results of growth experiments in the presence of oxygen described above.

thumbnail

Figure 2. Autolysis assay.

Late exponential phase cells were suspended in autolysis buffer and OD600 was monitored over 10 hours. Results are the average of triplicate assays and are represented as percent change in OD600 over time.

doi:10.1371/journal.pone.0061358.g002

Competence Development Correlates with Sensitivity to CSP and the Presence of Bacteriocins and Immunity Proteins

Much attention has been focused recently on the role of peptide-based quorum sensing systems that control bacteriocin production and genetic competence, as well as cell lysis and altruistic cell death. One intensively-studied peptide in S. mutans is competence stimulating peptide (CSP), which is sensed by the ComDE two-component system to activate bacteriocin production and DNA uptake [58]. Low concentrations of CSP (e.g. 30 to 100 nM) are sufficient to stimulate competence [59], whereas high concentrations of CSP (2 µM) can induce death in sub-populations of S. mutans [60], in part due to production of an endogenously-acting bacteriocin, mutacin V (CipB). There is also evidence that the expression of a number of bacteriocins (NlmD, Smu.1906 and CipB) can affect the level of induction of genetic competence through the CSP pathway [61]. To examine the sensitivity to CSP of the various isolates, cells were grown in BHI broth containing 400 nM synthetic CSP. There were some striking differences among the clinical isolates with respect to CSP sensitivity, with certain strains showing no adverse effects when CSP was present (Smu44, Smu56, Smu57, Smu81, and Smu93). With the exception of Smu93, these resistant strains could not be made naturally competent for genetic transformation, even with addition of exogenous CSP. In fact, out of the 15 strains tested, we were unable to obtain transformants in seven isolates when cells were cultured in the presence of 200 nM CSP in BHI medium. In four of these seven strains (Smu56, Smu57, Smu81, Smu104; Table 3), the lack of CSP induced competence could be potentially attributed to the absence of one or more Com-related proteins. Specifically, in Smu56 and Smu57, comCDE are absent (not present in the draft genome and not detectable by PCR), whereas comE is present but truncated in Smu81 (Figure S3). Also in Smu81, comC contains a frameshift that introduces a nonsense mutation (Figure S4) and ComX is truncated in Smu104 due to a nonsense mutation 127-bp into the comX gene (Figure S5). Notably, Smu56, Smu57, Smu81 and Smu104 contained ComRS, but none of these strains yielded transformants when grown in the presence of 600 nM XIP in a chemically-defined medium (data not shown).

thumbnail

Table 3. Mean generation time in BHI with CSP, presence of genetic competence, com genes and bacteriocin distribution among clinical isolates.

doi:10.1371/journal.pone.0061358.t003

In strains Smu69 and Smu93, ComC is predicted to be truncated by 3 amino acids because of an insertion of 18 nucleotides within the comC gene (Figure S6). Surprisingly, Smu69 had a faster generation time when growing in the presence of CSP (104±3 min) than the absence of CSP (114±1 min), while Smu93 was unaffected by the presence of CSP (Td 75±2 v.s. Td 73±1). The distribution of predicted bacteriocins (Smu.1902, Smu.1906, Smu.150, Smu.151, Smu.423 and Smu.1914) and putative immunity proteins (Smu.1909, Smu.1913, Smu.925, and Smu.152) also varied among strains (Table 3). In general, we noted that strains that were not able to become competent for natural transformation contained the genes for fewer bacteriocin and immunity proteins than strains that were competent for natural transformation.

Strain Smu86 was the most sensitive to growth inhibition by CSP, with the greatest lag and longest doubling time of all the strains tested (7 h lag, 240 min Td). To test whether the increased sensitivity to growth inhibition by CSP was correlated with competence development, cells were cultured in various concentrations of CSP (25–700 nM) and growth was compared with the reference strain UA159 under the same conditions (Figure 3A). Competence induction was also measured using various concentrations of CSP (0–200 nM; Figure 3B). The growth of Smu86 was inhibited to a greater extent at lower concentrations of CSP than UA159, such that 700 nM CSP was required to elicit the same degree of growth inhibition in UA159 as was seen with 200 nM CSP in Smu86. A similar effect was seen with competence development, where only one-fourth the amount of CSP was needed to stimulate an increase in transformation in Smu86 (25 nM) as that required for UA159 (100 nM).

thumbnail

Figure 3. Sensitivity to CSP.

Growth sensitivity to CSP correlates with level of competence. A) Growth curves of UA159 and Smu86 in various concentrations of CSP. B) Transformation assay with various concentrations of CSP. The photograph of the plates is representative of duplicate biological replicates.

doi:10.1371/journal.pone.0061358.g003

Biofilm-forming Capacity Varied Widely among Strains

S. mutans encodes three functionally-distinct glucosyltransferase enzymes, GtfB, GtfC, and GtfD, that contribute to various degrees to sucrose-dependent adhesion [11]. GtfD is found primarily in cell-free extracts and forms soluble glucan polymers dominated by α1,6- linked glucose chains, whereas GtfB is primarily cell-associated and catalyzes the synthesis of insoluble glucans composed mainly of α1,3 linkages [62]. GtfC can form both soluble and insoluble glucans and is found cell-associated and in culture supernates. S. mutans also encodes four glucan binding proteins, GbpA, GbpB, GbpC, and GbpD, that impact biofilm formation, as reviewed in [63].

In vitro biofilm development was analyzed after 48 hours of growth in a semi-defined medium containing 20 mM glucose or 20 mM sucrose (Figure 4). As has usually been noted for S. mutans, most strains formed more robust biofilms in sucrose than in glucose. Surprisingly, Smu20 and Smu44 formed more biofilm in glucose than sucrose, and some strains formed biofilms to a similar degree in sucrose and glucose (Smu56 and Smu57). The gtfBCD mutant strain Smu77/V1996 of V403 formed almost no biofilm in either of the tested conditions. The gtfB and gtfC genes are co-transcribed and share a high degree of similarity, so we could not unequivocally ascertain the complete content or organization of these genes in the contigs of all draft genomes. Therefore, to more reliably correlate the results of the biofilm assay with the distribution of Gtf enzymes within the clinical isolates, cell-associated and culture supernatant proteins from mid-exponential phase (BHI) cultures were separated by SDS-PAGE and analyzed by Western blotting using a polyclonal antisera that recognizes both GtfB and GtfC [43]. The results (Figure 5) revealed differences between strains, with 10 strains clearly expressing different amounts of both GtfB and GtfC and 2 strains only expressing one band corresponding to GtfC (Smu69, Smu104). Samples prepared from Smu20 and Smu44 also contained only one band that was recognized by the GtfB/C polyclonal antibody and migrated between GtfB and GtfC, indicating the likelihood that a recombination event between gtfB and gtfC occurred in these strains [64][66]. Such a recombination event may account for the poor biofilm formation by these strains in BM-sucrose medium. Mutant strain Smu77/V1996 is lacking any cell-associated GtfB or GtfC protein (Figure 5A; bottom panel) consistent with the mutations described in [47] within the gtfB-gtfC genes. A ΔgtfBC mutant of UA159 that was constructed and verified in our laboratory (SAB109) [67], was used as a negative control for the Western blot.

thumbnail

Figure 4. Biofilm formation by clinical isolates in semi-defined media with 20 mM glucose or sucrose.

Top panel shows numeric values of crystal violet biofilm assay (OD575 of eluted solution). Statistically significant differences compared to UA159 are indicated, ** = P≤0.001, * = P≤0.05. Bottom panel shows a picture of the crystal violet stained biofilm for each strain in the micro-titer plate wells before de-stain procedure.

doi:10.1371/journal.pone.0061358.g004
thumbnail

Figure 5. Expression of cell-wall associated proteins involved in biofilm formation and sucrose mediated cell clumping.

A) Cell associated protein extracts were from cell pellets harvested from mid-log BHI cultures. Proteins were separated on a 4–8% XT Criterion (BioRad) tris-acetate gel, transferred to a nitrocellulose membrane and stained with BioRad Colloidal Gold Total Protein Stain. For Western blot, proteins were transferred to PVDF membranes and reacted with polyclonal antisera that recognizes both GtfB and GtfC. B) Photos were taken after 48 hours of growth in BHI supplemented with 0.3% sucrose.

doi:10.1371/journal.pone.0061358.g005

Because it is possible for a polyclonal antibody to recognize a non-functional protein, we examined the ability of cells to aggregate in the presence of sucrose in broth cultures (Figure 5), a characteristic that requires GTF activity. Strains Smu20, Smu44, Smu77/V1996 and a ΔgtfBC mutant (SAB109) showed very little sucrose-dependent aggregation (Figure 5), consistent with the biofilm results and the Western blots showing aberrant Gtf production. On the other hand Smu56 and Smu57, both of which formed robust biofilms in both sucrose and glucose, displayed somewhat different phenotypes in this assay. Smu56 did not form aggregates or stick to the sides of glass test tubes, as was seen with other strains, and instead settled to the bottom of the tube. Smu57 formed cell aggregates more similar to the other Gtf-positive strains (Figure 4). Based on the genome sequences, all strains contained the gtfD gene sequence and all except Smu77/V1996 expressed a band at ~160 kDa in culture supernatant preparations corresponding to the known size of GtfD (data not shown). On further examination, Smu77/V1996 contained a truncated form of gtfD, which is the result of an insertion into the BglII site of gtfD of a 4605-bp of sequence containing ORFs 11–14 of transposon Tn916 from Enterococcus faecalis DS16 (Figure S7). This is slightly different from what was described in [48] for V1996, where the gtfD gene was interrupted with a 5.4-kb BamHI digest from pLN2 [68] that contained the tetM gene from Tn916.

A comparison of the gene content of the clinical isolates revealed there were differences in other genes that could affect the ability to form biofilm. For example Smu77/V1996 (V403, ΔgtfBCD, Δftf), Smu81, Smu86, Smu98 and Smu109 appear to be missing the gene for the major glucan binding protein, GbpA. With the exception of Smu98, it is noteworthy that these same strains also contain the cnm gene that encodes a protein with a collagen binding domain and an LPXTG motif at its C-terminus for cell-wall anchoring [51], which will be discussed in greater detail below. Furthermore, Smu86 and Smu109 also appear to express less of the cell-surface saliva binding adhesin P1 [69] compared to other strains (Figure 5A, top panel).

Distribution of Two-component Signal Transduction Systems in Draft Genomes of Isolates

The genome of S. mutans UA159 revealed 13 putative two-component signal transduction systems (TCS) and one orphan response regulator (GcrR) [25]. Recently, a 14th TCS (Smu.45/Smu.46) was described in UA159, but this TCS could only be found in 2 of the 13 strains analyzed [70]. Apparent Smu.45/Smu.46 homologs were found in 12 of the 57 strains sequenced here, but were in only one (Smu52) of the 15 isolates analyzed in this study. Biswas et al., 2008 also noted that the histidine kinase (HK) of TCS-5 (Smu.1814) was only found in two of the 13 strains examined in that study. We found that TCS-5 (Smu.1814/Smu.1815) was present in 45% of the strains sequenced and was not identified in 11 of the 15 strains characterized here (Table 4). Another recent study detailed the distribution of 18 TCSs found in eight newly-sequenced mutans streptococci (S. sobrinus DSM20742, S. ratti DSM20564 and 6 S. mutans strains) [71]. Song identified a 15th TCS (ComP/CmpR) in one S. mutans serotype f strain isolated from blood [71]. The sensor HK protein is predicted to contain 10 transmembrane domains, and is classified as a membrane-sensing HK and can be further classified into subgroup (ii) as a quorum-sensing HK with ComD (TCS-13). The response regulator (RR) of this novel HK/RR pair contains a LuxR_C-like DNA-binding HTH domain and was classified as a NarL type RR, which in other bacteria are involved in the control of genes that affect nitrogen fixation, sugar phosphate transport, nitrate and nitrite metabolism, quorum sensing or osmotic stress tolerance [72]. The function of this TCS in S. mutans has yet to be determined. However, a recently sequenced serotype k strain (LJ23) also contains TCS-15 [73]. In our analysis of the genomes of the 57 isolates from across the globe, TCS-15 was found in 16 of the 57 strains, and was distributed across all serotypes. Six of the strains phenotypically characterized in this study (Smu21, Smu77/V1996-V403, Smu81, Smu86, Smu98, Smu109) have TCS-15 (Table 4). While most strains contain TCS-7 (Smu.1037/Smu.1038) of unknown function, Smu81 contained several mutations resulting in frameshifts in both the HK and RR of TCS-7, whereas Smu109 has frameshift mutations in only the RR (Smu.1038). Furthermore, according to the draft genome sequence of Smu81, it appears this strain is also missing TCS-8 (Smu.1009/Smu.1008) [74], however these genes were able to be amplified by PCR using gene specific primers, indicating there is a gap in the sequence for this strain and therefore the presence of mutations within this TCS can not be ruled out. Smu56 and Smu57 are completely missing comDE (TCS-13), which are important for genetic competence, biofilm formation, bacteriocin production, quorum sensing, and stress tolerance [56], [75][77].

thumbnail

Table 4. Absence of two-component systems in clinical isolates.

doi:10.1371/journal.pone.0061358.t004

Identification of Novel Genes and Assessment of their Contribution to Virulence in Galleria Mellonella

The Type VII secretion system, found exclusively in Gram-positive bacteria (reviewed in [78], [79]), is responsible for the secretion of the WXG100 family of effector proteins, ESAT-6/EsxA and CSP-10/EsxB, which are required for the virulence of Mycobacterium tuberculosis [79] and necessary for persistent infection of Staphylococcus aureus [80]. Components of a Type VII secretion system (T7SS) were identified in 8 of the 57 S. mutans strains (see Table S1). These strains contained apparent homologs of EsxA, EsaA, EssA, EsaB, EssB and EssC (FtsK-SpoIIIE-domain), of which EssA, EssB and EssC are minimally required for secretion of EsxA or EsxB in S. aureus [80]. There was no EsxB homolog detected in any of the S. mutans strains, however there is at least one, and as many as four, EsaC-like proteins in those strains that encode a Type VII secretion (See Figure S8 and Table S1). The EsaC protein of S. aureus is a 130-aa soluble effector protein of the T7SS, which is negatively regulated post-transcriptionally by EsaB, and is required for persistent abscess formation during animal infections [81]. Burt et al. 2008 found that expression of EsaC is tightly regulated in several S. aureus strains, with EsaC expression being up-regulated in the presence of serum [81]. The EsaC-like proteins of S. mutans range in size from 102 to 106 aa with a DUF4176 (pfam-domain of unknown function 4176) with very little homology to EsaC from S. aureus (Figure S8). Additionally there was an apparent homolog of the T7SS-associated protein, Lmo0069, of Listeria monocytogenes found in four of the strains with a T7SS (Table S1).

As noted above, a number of strains contained the cnm gene, which encodes a collagen binding protein [51]. Located directly upstream of the cnm gene are two ORFs that encode proteins with collagen binding-like domains, for convenience designated here as cnaB/cbpA/cnm. CbpA and Cnm both contain signal sequences for secretion (SignalP 4.0 [82]), whereas CnaB does not. The cnm gene sequence is incomplete for these strains (due to the repeat sequences at the C-terminus of the cnm gene) and only includes the coding sequence for the first 346 amino acids of the Cnm protein. The 120-kDa product of the cnm gene of S. mutans was first described by Sato et al. [51] and contains a collagen binding domain (pfam05737) preceding a putative B-region consisting of a tandem TTTTE(K/A)P followed by 19 TTTE(A/S/T)P repeats, an architecture common among MSCRAMMs (microbial surface components recognizing adhesive matrix molecules) [83]. The cnm gene was found in 12–20% of clinical isolates of S. mutans [84] and has been implicated in invasion of human coronary artery endothelial cells [9]. It was also shown that invasive strains containing the cnm gene were more virulent in the wax-worm Galleria mellonella [10]. The cnaB gene is predicted to encode a 61-kDa protein with conserved domains for collagen binding (pfam05737) and a Cna protein B-type domain (pfam05738) similar to Cnm, found in collagen binding proteins of Staphylococcus aureus (Figure S9) [85]. cbpA is predicted to encode a 53-kDa protein and also contains a conserved Cna protein B-type domain (pfam05738) (Figure S9) and shares 82% sequence identity with the collagen-binding A precursor protein-like protein from Streptococcus ratti FA-1 (EJN94659). To our knowledge this is the first time the cnaB and cbpA genes have been described in S. mutans.

In a recently sequenced cnm+ serotype k strain LJ23, a gene predicted to encode a protein (containing a signal sequence, collagen binding domain, Cna protein B-type domain and the LPTXG-cell wall anchoring motif) with 75% sequence similarity to CnaB (Figure S10) was described [73]. On further examination of the sequence from S. mutans LJ23, there are two possible open reading frames (ORF1 and ORF2) not annotated in NCBI, with homology to the N-terminus and C-terminus, respectively, of CbpA located upstream of the coding sequence for cnm (Figures S11 and S12). In all the strains sequenced in this study [34], the protein sequence (Figure S13) and gene arrangement of the cnaB, cbpA and cnm genes are almost identical.

In order to determine what role the presence of these genes might play in the virulence of S. mutans, strains that contained the cnaB/cbpB/cnm collagen binding protein genes (Smu77/V1996, Smu81, Smu86 and Smu109/OMZ175) or the genes for the T7SS (Smu26, Smu44, Smu80 and Smu102) were assayed for virulence in the G. mellonella model (Figure 6). For comparison purposes, we also included the type strain UA159 (lacking both cnaB-cbpA-cnm and T7SS) and a cnm mutant of OMZ175 (OMZΔcnm). As previously seen [10], OMZ175 was significantly more virulent than UA159 and inactivation of cnm strongly attenuated the virulence of OMZ175 (data not shown). Along these lines, strains Smu77, Smu81 and Smu86 expressing cnaB-cbpA-cnm were significantly more virulent than UA159 (Figure 6A). Strains Smu77/V1996 and Smu86 (serotypes c and e, respectively) were as virulent as Smu109 (OMZ175), a well-characterized serotype f strain [9], [86][89], whereas Smu81 killed G. mellonella more rapidly than OMZ175. These results support previous findings that cnm is an important virulence factor in the G. mellonella invertebrate model and furthermore, reveals that serotype c strains carrying genes for collagen binding proteins are equally as virulent compared to serotype f strains [9], [10]. On the other hand, strains that contained the genes for the T7SS did not display significantly greater virulence compared with UA159 (Figure 6B).

thumbnail

Figure 6. Kaplan-Meier killing curves of G. mellonella larvae (wax worms) inoculated with S. mutans strains.

Graphs show percent survival of G. mellonella after injection of 5 µl inoculums. CFU/100 µl of each inoculum is provided in legend. A) Smu81, Smu86, Smu77 compared to UA159 (non-virulent) and OMZ175. B) Smu44, Smu26, Smu80 and Smu102 compared to UA159. * Indicates significant difference (P≥0.05) compared to OMZ175.

doi:10.1371/journal.pone.0061358.g006

Discussion

S. mutans is a genetically-diverse species that co-exists in the oral cavity with a number of other streptococci and hundreds of additional species of bacteria from a wide range of taxa [90], [91]. Contributing to its diversity is its ability to take up DNA from the environment allowing this organism to acquire new genes through lateral gene transfer, which may have enhanced the ability of S. mutans to adapt to and survive the selective pressures of the oral cavity as human diets became richer in refined sugars and polysaccharides [34], [92]. In this study, we demonstrate that phenotypic properties that have been directly associated with the ability of S. mutans to establish, persist, and/or cause disease in humans vary considerably among 15 genetically- and geographically-heterogeneous clinical isolates for which high coverage genome sequence is now available. The particular phenotypes analyzed here are complex, and multiple gene products contribute to the manifestation of, for example, growth at low pH or oxidative stress resistance. Importantly, this study, coupled with the genome sequencing information, which is publically available via a newly developed genome browser (http://strep-genome.bscb.cornell.edu), opens the way for evaluating relationships between gene content and virulence potential, and lays the foundation for determining how the core and non-core genomes interact to allow adaptation to the constantly-changing environment of the human oral cavity.

The genome sequences of these S. mutans strains reveal that approximately 1,490 genes constitute the core genome, while the pan-genome contains roughly 3,300 genes [34], which is substantial in comparison to the 1,963 ORFs found in UA159 [25]. The knowledge that so many non-core genes can be found in S. mutans underscores the limitations of using only one genome sequence as a reference for functional genomic studies and highlights the adaptive potential of this species for survival in the oral cavity. Population genetic analysis based on the core genes indicates that a large population expansion of S. mutans took place between 3,000 and 10,000 years ago, coinciding with the adoption of agriculture by humans and the associated expansion of the quantity and type of carbohydrates in the diet [34]. While agriculture and industrialization increased survival and influenced evolution of the human species, it also provided new selective pressures for the organisms in the human oral microbiome, resulting in the evolution of bacteria able to withstand the stresses induced by excess carbohydrates (e.g. acid production), a critical characteristic of S. mutans, while dramatically enhancing the prevalence of human dental caries [93][95].

The F1FO-H+-translocating ATPase is considered to be a major determinant of acid resistance in S. mutans [13]. The activity and pH optima of these ATPase enzymes are strongly correlated with acid tolerance in oral bacteria. For example, strongly aciduric organisms, such as lactobacilli, display much greater activity and lower pH optima for the ATPase than does the acid-sensitive species Streptococcus sanguinis [96]. Of interest, there was substantial variation in the ability of different S. mutans strains to contend with acid stress. In particular, several strains displayed faster doubling times during growth at pH 5.5 and showed increased survival at a killing pH of 2.8, compared to UA159. However, the enhanced acid resistance of these strains was not correlated with ATPase activity, which was similar in all strains and conditions tested, although all strains expressed higher ATPase activity after acid-adaptation. We did not directly examine the pH optima of the enzymes, but the F1Fo -ATPase subunits in all strains were very highly conserved (99.6% identical) with very few polymorphisms within individual subunits. Therefore, differences in membrane proton permeability or other physiological traits may account for the differences in acid tolerance between strains.

Interestingly, oxidative stress resistance showed the greatest spectrum of behaviors among the strains, with one strain (Smu81) displaying an unusually high degree of sensitivity to oxidative stress induced by the superoxide generator paraquat. The reason for paraquat sensitivity in Smu81 remains a mystery, as there are no obvious loci missing (e.g. superoxide dismutase) relevant to those pathways that have already been shown to contribute to oxidative stress resistance in other strains of S. mutans. While the presence of genes can clearly be determined with certainty, this is not necessarily so regarding the absence of genes in draft genome sequences. Thus, Smu81 may be lacking gene product(s) required for paraquat tolerance, or aberrantly expresses or contains key mutations in core genes that compromise its tolerance of paraquat. Transcriptomic studies are planned to address the former hypothesis.

A particularly interesting observation was the growth-lysis-regrowth phenotype displayed by several strains in the presence of air, which could indicate that a subset of isolates have adapted this strategy to cope with the stress induced by exposure to oxygen. S. mutans has several known mechanisms that mediate cell lysis, including bacteriocins (regulated by ComCDE), the autolysin AltA, and the apparent holin:antiholin complexes LrgAB and CidAB, which are regulated by oxygen and are growth phase dependent [39], [97], [98], that could account for the growth-lysis-regrowth phenotype. Furthermore, altruistic behavior of bacteria has been well documented, wherein programmed cell death (PCD) is used by organisms to ensure the survival of a population by sacrificing a sub-population during stress or when resources become limiting [55], [99]. It is therefore conceivable that these systems are regulated differently in some S. mutans strains depending on the genome content, and that the differences seen between strains grown in air are due to differential gene regulation in response to oxygen. Further analysis of the gene content, or of the transcriptomes of these strains in the presence of oxygen, might lead to the discovery of novel mechanisms for oxygen tolerance and alternative gene regulation pathways in S. mutans and other oral streptococci that regulate PCD. Notably, the growth-lysis-regrowth phenotype is probably not induced by superoxide radical alone, as the behavior is not observed in the presence of paraquat. Perhaps endogenous production of hydrogen peroxide [100] or hyper-expression of bacteriocins in response to oxygen [97] acts as a signal to trigger sub-population lysis.

The development of genetic competence in S. mutans is a complex process involving multiple inputs [101][104]. The accumulation of a processed form of the comC gene product (CSP) results in signaling through a phosphorylation cascade via a two-component system (ComD and ComE), which induces expression of bacteriocins and stimulates cells to take up exogenous DNA through mechanisms that are not yet completely understood [15], [59]. More recently, it was discovered that another peptide known as XIP, for sigX-inducing peptide, encoded by the comS gene also plays an important role in comX activation by modulating the DNA binding activity of the transcription factor ComR [105], [106]. Interestingly, comR and comS are found in all 57 strains sequenced, but significant numbers of the sequenced strains either lacked the comCDE genes or contained various mutations that could lead to failure to produce functional ComCDE proteins. There were a number of polymorphisms within the comR gene resulting in variation in the protein sequence (Figure S14), however there was no correlation of these polymorphisms with the competence phenotypes observed. On the other hand, the protein sequence for ComS was 100% identical in all 57 S. mutans strains sequenced. Polymorphisms within the comCDE locus of S. mutans isolates have been noted previously [45], resulting in variability in genetic competence, as well as pleiotropic effects associated with deletion of comDE in different S. mutans strains [101]. Interestingly, about one-fifth of the sequenced isolates encoded a ComC peptide with a C-terminal truncation of 3 amino acids (producing an 18-mer CSP peptide), while one-third of sequenced strains lacked comC all together. Although the presence of the 18-mer allele of CSP has been documented previously [36], [107], it was shown only recently that CSP is processed extracellularly at the C-terminus by a protease (SepM) to yield an 18-mer CSP peptide, and that C-terminal processing can be required for CSP signaling [108]. Currently the implications of the distribution of 21-mer and 18-mer CSP species across strains of S. mutans are unclear. However, these results reveal that both versions of CSP are common in the population of S. mutans, and that competence and competence-related phenotypes are largely independent of the allele of comC, which is consistent with studies detailed elsewhere [108]. Our results extend these findings and further demonstrate that tremendous variation exists among strains in the pathways involved in quorum sensing and genetic competence, and also that natural genetic competence can not be readily induced in a laboratory setting in many strains of S. mutans.

A survey of genes that encode non-lantibiotic bacteriocin and immunity proteins revealed a number of polymorphisms and disparities in the distribution of these genes in the strains described here. Of particular note are differences in the pathway thought to be responsible for CSP-induced lysis, with a significant number of strains missing the bacteriocin CipB/Smu.1914 (Smu21, Smu44, Smu.56, Smu.57, and Smu98) or the immunity protein CipI/Smu.925 (Smu86, Smu98 and Smu109) [60]. Additionally, in two strains (Smu77 and Smu81) there appears to be a recombination between Smu.1914/CipB and Smu.1906 resulting in a hybrid protein that contains the signal sequence of CipB and the bacteriocin sequence from Smu.1906. Furthermore, a number of strains contained mutations within the cipI gene sequence that resulted in either a truncation at the N-terminus of 18 amino acids (Smu20, Smu77, Smu81, Smu93 and Smu104) and/or a truncation of 4 amino acids (Smu20, Smu56, Smu77 and Smu104) or 7 amino acids at the C- terminus (Smu21). Recent work by Dufour [61] showed that the unprocessed CipB bacteriocin contributes to the CSP-dependent increase in transformation efficiency seen in UA159, in which a ΔcipB mutant showed no increase in transformants upon addition of CSP. Transcriptome analysis via microarray of a ΔcipB mutant revealed that CipB acts at the transcriptional level through regulation of ComE, ComR and ComX [61]. This study also found that deletion of cipI encoding the immunity protein to CipB results in increased expression of competence genes and increased transformation efficiency in the absence of exogenously-added CSP [61]. On the other hand, nlmT/Smu.286 and nlmE/Smu.287, required for transport of non-lantibiotic bacteriocins [109] are among the core genes of S. mutans. Considering the variation seen among isolates with respect to the presence or absence of competence genes, bacteriocins and immunity proteins, as well as the ability to become naturally competent, further study of multiple S. mutans strains is warranted in order to determine the roles that these gene products might play in competence and biofilm ecology in the context of different genetic backgrounds. Also, variations within the competence pathway tend to sort with genes related to bacteriocin production, which likely have evolved differently from other more highly conserved competence related genes and may therefore be of interest to the study of interspecies interactions, e.g. antagonism of commensal streptococci. Further still, the presence or absence of these gene products may reflect different adaptive strategies of the various isolates of S. mutans, and may be related to cariogenic potential as the differences could impact biofilm persistence and interactions with other members of the oral microbiome.

There was additional variability in the capacity of strains to form biofilms and on further examination this variation could be partially attributed to differences in Gtf isozyme production. Two strains, Smu20 and Smu44, which formed biofilms relatively poorly in sucrose medium, were found to have a recombination between gtfB and gtfC, as reported for some other strains of S. mutans [64][66]. Of note, there were also several strains lacking gbpA, the absence of which appeared to correlate with the presence of the cnm gene. Notably, Nakano et al. reported that 79% of strains containing the cnm gene lacked the gbpA gene [84]. Recently a new collagen binding protein, termed Cbm, was described in S. mutans clinical isolates [110], however this collagen binding protein is different from those described here. Also, the cbm gene was found in only 2% of strains [110] and does not appear to be present in any of the 57 isolates sequenced by our group. The serotype f strain, OMZ175 (referred to as Smu109 in this study), which has been well characterized for its invasiveness of endothelial cells and virulence in the G. mellonella model [9], [10], along with three additional strains, were found to contain two ORFs upstream of cnm that encode putative proteins with possible collagen binding functions (cnaB/cbpA/cnm). While the incidence of collagen binding proteins within the species S. mutans is relatively low, there seems to be an increased frequency in non-serotype c strains. Here we report the incidence of two serotype c (Smu77/V1996 and Smu81) and one serotype e strain (Smu86) that contain the cnaB/cbpA/cnm genes and show for the first time that they are as virulent in the G. mellonella model as a serotype f strain. Considering the invasive nature of strains that carry the cnm gene, collagen-binding proteins could have implications for systemic diseases caused by S. mutans [17] and further characterization of cnaB and cbpA should be pursued. It is also interesting to speculate that the increased frequency of cnm in serotype f and k strains along with decreased glucan-binding capacity due to a lack of gbpA, could be an indication of adaptation by S. mutans for survival outside the oral cavity, and could therefore represent a lineage of this pathogen that is better adapted to contributing to systemic diseases.

We also report for the first time the presence of a Type VII secretion system (T7SS) in a number of S. mutans clinical isolates. While the role that this pathway may play in the physiology or virulence of S. mutans is unknown at this time, its presence in 8 of 57 strains is an exciting discovery. Strains with a T7SS did not display increased virulence in G. mellonella, which could be due to a lack of expression under assay conditions or perhaps the system contributes to the physiology or virulence in ways that are not relevant in the insect model. Preliminary studies looking at invasion of HCAEC did not indicate increased ability to invade (unpublished data) compared to UA159 and deletion of the esxA gene did not seem to affect growth or stress tolerance in at least two strains tested so far (unpublished data). Similar results were seen in Listeria monocytogenes, where deletion of the ESAT-6/EsxA homolog had no effect on virulence and or growth [111]. However, in L. monocytogenes, only the esxA effector gene was inactivated, and therefore the possibility that other effectors secreted by the T7SS could compensate for its loss remains. Further dissection of the S. mutans T7SS may lead to the discovery of other effector proteins that are secreted through this pathway, which could have implications on survival of the organism in complex biofilms or in the circulatory system of the host.

The phenotypic characterization of S. mutans strains presented here, in conjunction with the WGS sequences of the 57 isolates [34], will provide a powerful set of resources to the S. mutans community, allowing for the discovery of new genes that contribute to the virulence of this caries pathogen. In addition, much insight into the evolution and phenotypic diversity of the S. mutans species as a whole in the context of genome-scale information has been gained. This study will allow researchers to compare the phenotypes of strains with the presence or absence of certain genes, and to take a more focused approach to functional genomics studies. Furthermore, this study will also provide a foundation for future analysis of core and non-core genome interactions. For example studies are currently underway in our laboratory to dissect the role in stress-tolerance of the “unique core” genes, which are a group of genes that is present in all S. mutans isolates but not in closely-related mutans streptococci [34]. Additionally, the genome sequence data combined with RNAseq will allow us to evaluate how different strains utilize their transcriptomes to cope with environmental stress and quorum sensing, leading to a better overall understanding of gene regulation in the streptococci. While the aim of this study was not to correlate the presence of S. mutans strains and or gene content with caries or virulence, our results can be used to form new hypotheses concerning gene content of the S. mutans species and may lead to a better understanding of how the species S. mutans is involved in the process of caries development, infective endocarditis and cardiovascular disease.

Supporting Information

Figure S1.

Gene content differences between strains based on orthologs recovered across genomes via an all-versus-all BLASTP search combined with clustering using OrthoMCL2 [46].

doi:10.1371/journal.pone.0061358.s001

(TIFF)

Figure S2.

Growth-lysis-growth phenotype of select strains in the presence of oxygen stress (no mineral oil overlay). Duplicate early exponential phase cultures were diluted 1:100 into BHI and inoculated in duplicate wells of a Bioscreen C microtiter plate. Growth was monitored at 37°C every 30 min in a Bioscreen C labsystem for 48 hr.

doi:10.1371/journal.pone.0061358.s002

(TIFF)

Figure S3.

Sequence alignment of comE gene from UA159 and Smu81. The comE gene from Smu81 contains a frameshift at 409 nucleotides stemming from an insertion of two nucleotides, resulting in a mutated ComE protein.

doi:10.1371/journal.pone.0061358.s003

(TIFF)

Figure S4.

Sequence alignment of comC gene from UA159 and Smu81. The comC gene from Smu81 contains an extra adenosine four nucleotides into the DNA sequence, resulting in a frameshift and no ComC protein.

doi:10.1371/journal.pone.0061358.s004

(TIFF)

Figure S5.

Sequence alignment of comX from Smu104 and UA159. The comX gene from Smu104 contains a C to T mutation resulting in a stop codon 26 nucleotides into the comX gene.

doi:10.1371/journal.pone.0061358.s005

(TIFF)

Figure S6.

Sequence alignment of comC gene from UA159 compared to Smu69 and Smu93. The comC gene from Smu69 and Smu93 contain an insertion of 18 nucleotides resulting in a stop codon and a truncated 18-mer ComC peptide.

doi:10.1371/journal.pone.0061358.s006

(TIFF)

Figure S7.

Sequence of inserted 4605 bp DNA within the gtfD gene of Smu77. Blast results revealed that the DNA sequence is from the Enterococcus faecalis DS16 transposon Tn916 corresponding to ORF11-14, containing the gene sequence for TetM (Orf11), TetM leader peptide (Orf12), conjugative transposon protein TcpC (Orf13), and a protein containing a lysozyme domain that cleaves beta, 1–4, linked polysaccharides (Orf14).

doi:10.1371/journal.pone.0061358.s007

(PDF)

Figure S8.

The four EsaC homologs from strain Smu44 compared to EsaC from S. aureus subspecies aureus JH9. A) Protein sequence and size of EsaC homologs and contig location in Smu44. B) ClustalW2 sequence alignment of EsaC homologs.

doi:10.1371/journal.pone.0061358.s008

(TIFF)

Figure S9.

Conserved domains of CnaB, CbpA, and Cnm as determined using NCBI’s CDD [85].

doi:10.1371/journal.pone.0061358.s009

(TIFF)

Figure S10.

ClustalW2 sequence alignment of CnaB protein from Smu81 and CnaB homolog from S. mutans Serotype k strain LJ23.

doi:10.1371/journal.pone.0061358.s010

(TIFF)

Figure S11.

ClustalW2 sequence alignment of CbpA from Smu81 and ORF1 from S. mutans strain LJ23.

doi:10.1371/journal.pone.0061358.s011

(TIFF)

Figure S12.

ClustalW2 sequence alignment of CbpA from Smu81 and ORF2 from S. mutans strain LJ23.

doi:10.1371/journal.pone.0061358.s012

(TIFF)

Figure S13.

ClustalW sequence alignment of CnaB, CbpA and Cnm protein sequence from Smu109, Smu86, Smu81 and Smu77. Signal sequence cleavage site for CbpA and Cnm as predicted by SignalP (version 4.0) is indicated *. LPXTG sequence for cell-wall anchoring is underlined for CnaB and CbpA. The Cnm gene sequence is incomplete.

doi:10.1371/journal.pone.0061358.s013

(PDF)

Figure S14.

ClustalW sequence alignment of ComR protein sequence from all 15 strains. Smu44, Smu56, Smu57, Smu69, Smu81, Smu98 and Smu104 are not competent for natural transformation.

doi:10.1371/journal.pone.0061358.s014

(PDF)

Table S1.

Distribution of Type VII secretion genes among clinical isolates of S. mutans.

doi:10.1371/journal.pone.0061358.s015

(PDF)

Acknowledgments

We would like to thank David Beighton and Renata Mattos-Graner for providing collections of the clinical isolates described in this study. We also want to thank Vince Richards for his assistance analyzing genome sequence data.

Author Contributions

Conceived and designed the experiments: SRP LZ JA JL RAB. Performed the experiments: SRP JA JM LZ. Analyzed the data: SRP JA VR LZ RAB TL. Contributed reagents/materials/analysis tools: JL MJS RAB. Wrote the paper: SRP JL JA MJS RAB.

References

  1. 1. Kuramitsu HK, He X, Lux R, Anderson MH, Shi W (2007) Interspecies interactions within oral microbial communities. Microbiol Mol Biol Rev 71: 653–670. doi: 10.1128/mmbr.00024-07
  2. 2. Marsh PD (2003) Are dental diseases examples of ecological catastrophes? Microbiology 149: 279–294. doi: 10.1099/mic.0.26082-0
  3. 3. Levine M (2011) Susceptibility to dental caries and the salivary proline-rich proteins. Int J Dent 2011: 953412. doi: 10.1155/2011/953412
  4. 4. Kleinberg I (2002) A mixed-bacteria ecological approach to understanding the role of the oral bacteria in dental caries causation: an alternative to Streptococcus mutans and the specific-plaque hypothesis. Crit Rev Oral Biol Med 13: 108–125. doi: 10.1177/154411130201300202
  5. 5. Pizzo G, Piscopo MR, Pizzo I, Giuliana G (2007) Community water fluoridation and caries prevention: a critical review. Clin Oral Investig 11: 189–193. doi: 10.1007/s00784-007-0111-6
  6. 6. Loesche WJ (1986) Role of Streptococcus mutans in human dental decay. Microbiol Rev 50: 353–380.
  7. 7. Bowden GH (1990) Microbiology of root surface caries in humans. J Dent Res 69: 1205–1210. doi: 10.1177/00220345900690051701
  8. 8. Nakano K, Inaba H, Nomura R, Nemoto H, Takeda M, et al. (2006) Detection of cariogenic Streptococcus mutans in extirpated heart valve and atheromatous plaque specimens. J Clin Microbiol 44: 3313–3317. doi: 10.1128/jcm.00377-06
  9. 9. Abranches J, Zeng L, Belanger M, Rodrigues PH, Simpson-Haidaris PJ, et al. (2009) Invasion of human coronary artery endothelial cells by Streptococcus mutans OMZ175. Oral Microbiol Immunol 24: 141–145. doi: 10.1111/j.1399-302x.2008.00487.x
  10. 10. Abranches J, Miller JH, Martinez AR, Simpson-Haidaris PJ, Burne RA, et al. (2011) The collagen-binding protein Cnm is required for Streptococcus mutans adherence to and intracellular invasion of human coronary artery endothelial cells. Infect Immun 79: 2277–2284. doi: 10.1128/iai.00767-10
  11. 11. Bowen WH, Koo H (2011) Biology of Streptococcus mutans-derived glucosyltransferases: role in extracellular matrix formation of cariogenic biofilms. Caries Res 45: 69–86. doi: 10.1159/000324598
  12. 12. Belli WA, Marquis RE (1991) Adaptation of Streptococcus mutans and Enterococcus hirae to acid stress in continuous culture. Appl Environ Microbiol 57: 1134–1138.
  13. 13. Lemos JA, Burne RA (2008) A model of efficiency: stress tolerance by Streptococcus mutans. Microbiology 154: 3247–3255. doi: 10.1099/mic.0.2008/023770-0
  14. 14. Federle MJ, Morrison DA (2012) One If By Land, Two If By Sea: Signaling to the Ranks With CSP and XIP. Mol Microbiol.
  15. 15. Merritt J, Qi F (2012) The mutacins of Streptococcus mutans: regulation and ecology. Mol Oral Microbiol 27: 57–69. doi: 10.1111/j.2041-1014.2011.00634.x
  16. 16. Nakano K, Ooshima T (2009) Serotype classification of Streptococcus mutans and its detection outside the oral cavity. Future Microbiol 4: 891–902. doi: 10.2217/fmb.09.64
  17. 17. Nakano K, Nomura R, Matsumoto M, Ooshima T (2010) Roles of oral bacteria in cardiovascular diseases–from molecular mechanisms to clinical cases: Cell-surface structures of novel serotype k Streptococcus mutans strains and their correlation to virulence. J Pharmacol Sci 113: 120–125. doi: 10.1254/jphs.09r24fm
  18. 18. Shibata Y, Ozaki K, Seki M, Kawato T, Tanaka H, et al. (2003) Analysis of loci required for determination of serotype antigenicity in Streptococcus mutans and its clinical utilization. J Clin Microbiol 41: 4107–4112. doi: 10.1128/jcm.41.9.4107-4112.2003
  19. 19. Napimoga MH, Kamiya RU, Rosa RT, Rosa EA, Höfling JF, et al. (2004) Genotypic diversity and virulence traits of Streptococcus mutans in caries-free and caries-active individuals. J Med Microbiol 53: 697–703. doi: 10.1099/jmm.0.05512-0
  20. 20. Pieralisi FJ, Rodrigues MR, Segura VG, Maciel SM, Ferreira FB, et al. (2010) Genotypic Diversity of Streptococcus mutans in Caries-Free and Caries-Active Preschool Children. Int J Dent 2010: 824976. doi: 10.1155/2010/824976
  21. 21. Phattarataratip E, Olson B, Broffitt B, Qian F, Brogden KA, et al. (2011) Streptococcus mutans strains recovered from caries-active or caries-free individuals differ in sensitivity to host antimicrobial peptides. Mol Oral Microbiol 26: 187–199. doi: 10.1111/j.2041-1014.2011.00607.x
  22. 22. Kreulen CM, de Soet HJ, Hogeveen R, Veerkamp JS (1997) Streptococcus mutans in children using nursing bottles. ASDC J Dent Child 64: 107–111.
  23. 23. Lembo FL, Longo PL, Ota-Tsuzuki C, Rodrigues CR, Mayer MP (2007) Genotypic and phenotypic analysis of Streptococcus mutans from different oral cavity sites of caries-free and caries-active children. Oral Microbiol Immunol 22: 313–319. doi: 10.1111/j.1399-302x.2007.00361.x
  24. 24. Argimón S, Caufield PW (2011) Distribution of putative virulence genes in Streptococcus mutans strains does not correlate with caries experience. J Clin Microbiol 49: 984–992. doi: 10.1128/jcm.01993-10
  25. 25. Ajdic D, McShan WM, McLaughlin RE, Savic G, Chang J, et al. (2002) Genome sequence of Streptococcus mutans UA159, a cariogenic dental pathogen. Proc Natl Acad Sci U S A 99: 14434–14439. doi: 10.1073/pnas.172501299
  26. 26. Waterhouse JC, Swan DC, Russell RR (2007) Comparative genome hybridization of Streptococcus mutans strains. Oral Microbiol Immunol 22: 103–110. doi: 10.1111/j.1399-302x.2007.00330.x
  27. 27. Zhang L, Foxman B, Drake DR, Srinivasan U, Henderson J, et al. (2009) Comparative whole-genome analysis of Streptococcus mutans isolates within and among individuals of different caries status. Oral Microbiol Immunol 24: 197–203. doi: 10.1111/j.1399-302x.2008.00495.x
  28. 28. Wu C, Cichewicz R, Li Y, Liu J, Roe B, et al. (2010) Genomic island TnSmu2 of Streptococcus mutans harbors a nonribosomal peptide synthetase-polyketide synthase gene cluster responsible for the biosynthesis of pigments involved in oxygen and H2O2 tolerance. Appl Environ Microbiol 76: 5815–5826. doi: 10.1128/aem.03079-09
  29. 29. Waterhouse JC, Russell RR (2006) Dispensable genes and foreign DNA in Streptococcus mutans. Microbiology 152: 1777–1788. doi: 10.1099/mic.0.28647-0
  30. 30. Do T, Gilbert SC, Clark D, Ali F, Fatturi Parolo CC, et al. (2010) Generation of diversity in Streptococcus mutans genes demonstrated by MLST. PLoS One 5: e9073. doi: 10.1371/journal.pone.0009073
  31. 31. Maiden MC, Bygraves JA, Feil E, Morelli G, Russell JE, et al. (1998) Multilocus sequence typing: a portable approach to the identification of clones within populations of pathogenic microorganisms. Proc Natl Acad Sci U S A 95: 3140–3145. doi: 10.1073/pnas.95.6.3140
  32. 32. Burne RA, Zeng L, Ahn SJ, Palmer SR, Liu Y, et al. (2012) Progress dissecting the oral microbiome in caries and health. Adv Dent Res 24: 77–80. doi: 10.1177/0022034512449462
  33. 33. Willems RJ, Hanage WP, Bessen DE, Feil EJ (2011) Population biology of Gram-positive pathogens: high-risk clones for dissemination of antibiotic resistance. FEMS Microbiol Rev 35: 872–900. doi: 10.1111/j.1574-6976.2011.00284.x
  34. 34. Cornejo OE, Lefébure T, Pavinski Bitar PD, Lang P, Richards VP, et al.. (2013) Evolutionary and Population Genomics of the Cavity Causing Bacteria Streptococcus mutans. Mol Biol Evol.
  35. 35. Loo CY, Corliss DA, Ganeshkumar N (2000) Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J Bacteriol 182: 1374–1382. doi: 10.1128/jb.182.5.1374-1382.2000
  36. 36. Li YH, Lau PC, Lee JH, Ellen RP, Cvitkovitch DG (2001) Natural genetic transformation of Streptococcus mutans growing in biofilms. J Bacteriol 183: 897–908. doi: 10.1128/jb.183.3.897-908.2001
  37. 37. Khalichi P, Cvitkovitch DG, Santerre JP (2004) Effect of composite resin biodegradation products on oral streptococcal growth. Biomaterials 25: 5467–5472. doi: 10.1016/j.biomaterials.2003.12.056
  38. 38. Wen ZT, Burne RA (2002) Functional genomics approach to identifying genes required for biofilm development by Streptococcus mutans. Appl Environ Microbiol 68: 1196–1203. doi: 10.1128/aem.68.3.1196-1203.2002
  39. 39. Ahn SJ, Burne RA (2007) Effects of oxygen on biofilm formation and the AtlA autolysin of Streptococcus mutans. J Bacteriol 189: 6293–6302. doi: 10.1128/jb.00546-07
  40. 40. Shibata Y, Kawada M, Nakano Y, Toyoshima K, Yamashita Y (2005) Identification and characterization of an autolysin-encoding gene of Streptococcus mutans. Infect Immun 73: 3512–3520. doi: 10.1128/iai.73.6.3512-3520.2005
  41. 41. Wen ZT, Burne RA (2004) LuxS-mediated signaling in Streptococcus mutans is involved in regulation of acid and oxidative stress tolerance and biofilm formation. J Bacteriol 186: 2682–2691. doi: 10.1128/jb.186.9.2682-2691.2004
  42. 42. Zeng L, Burne RA (2009) Transcriptional regulation of the cellobiose operon of Streptococcus mutans. J Bacteriol 191: 2153–2162. doi: 10.1128/jb.01641-08
  43. 43. Wunder D, Bowen WH (2000) Effects of antibodies to glucosyltransferase on soluble and insolubilized enzymes. Oral Dis 6: 289–296. doi: 10.1111/j.1601-0825.2000.tb00141.x
  44. 44. Klein MI, Flório FM, Pereira AC, Höfling JF, Gonçalves RB (2004) Longitudinal study of transmission, diversity, and stability of Streptococcus mutans and Streptococcus sobrinus genotypes in Brazilian nursery children. J Clin Microbiol 42: 4620–4626. doi: 10.1128/jcm.42.10.4620-4626.2004
  45. 45. Klein MI, Bang S, Flório FM, Höfling JF, Gonçalves RB, et al. (2006) Genetic diversity of competence gene loci in clinical genotypes of Streptococcus mutans. J Clin Microbiol 44: 3015–3020. doi: 10.1128/jcm.02024-05
  46. 46. Li L, Stoeckert CJ, Roos DS (2003) OrthoMCL: identification of ortholog groups for eukaryotic genomes. Genome Res 13: 2178–2189. doi: 10.1101/gr.1224503
  47. 47. Munro C, Michalek SM, Macrina FL (1991) Cariogenicity of Streptococcus mutans V403 glucosyltransferase and fructosyltransferase mutants constructed by allelic exchange. Infect Immun 59: 2316–2323.
  48. 48. Hanada N, Kuramitsu HK (1989) Isolation and characterization of the Streptococcus mutans gtfD gene, coding for primer-dependent soluble glucan synthesis. Infect Immun 57: 2079–2085.
  49. 49. Schroeder VA, Michalek SM, Macrina FL (1989) Biochemical characterization and evaluation of virulence of a fructosyltransferase-deficient mutant of Streptococcus mutans V403. Infect Immun 57: 3560–3569.
  50. 50. Macrina FL, Scott CL (1978) Evidence for a disseminated plasmid in Streptococcus mutans. Infect Immun 20: 296–302.
  51. 51. Sato Y, Okamoto K, Kagami A, Yamamoto Y, Igarashi T, et al. (2004) Streptococcus mutans strains harboring collagen-binding adhesin. J Dent Res 83: 534–539. doi: 10.1177/154405910408300705
  52. 52. Lemos JA, Abranches J, Burne RA (2005) Responses of cariogenic streptococci to environmental stresses. Curr Issues Mol Biol 7: 95–107.
  53. 53. Bradshaw DJ, Marsh PD, Allison C, Schilling KM (1996) Effect of oxygen, inoculum composition and flow rate on development of mixed-culture oral biofilms. Microbiology 142 (Pt 3): 623–629. doi: 10.1099/13500872-142-3-623
  54. 54. Marquis RE (1995) Oxygen metabolism, oxidative stress and acid-base physiology of dental plaque biofilms. J Ind Microbiol 15: 198–207. doi: 10.1007/bf01569826
  55. 55. Rice KC, Bayles KW (2003) Death’s toolbox: examining the molecular components of bacterial programmed cell death. Mol Microbiol 50: 729–738. doi: 10.1046/j.1365-2958.2003.t01-1-03720.x
  56. 56. Ahn SJ, Burne RA (2006) The atlA operon of Streptococcus mutans: role in autolysin maturation and cell surface biogenesis. J Bacteriol 188: 6877–6888. doi: 10.1128/jb.00536-06
  57. 57. Montanaro L, Poggi A, Visai L, Ravaioli S, Campoccia D, et al. (2011) Extracellular DNA in biofilms. Int J Artif Organs 34: 824–831. doi: 10.5301/ijao.5000051
  58. 58. Martin B, Quentin Y, Fichant G, Claverys JP (2006) Independent evolution of competence regulatory cascades in streptococci? Trends Microbiol 14: 339–345. doi: 10.1016/j.tim.2006.06.007
  59. 59. Senadheera D, Cvitkovitch DG (2008) Quorum sensing and biofilm formation by Streptococcus mutans. Adv Exp Med Biol 631: 178–188. doi: 10.1007/978-0-387-78885-2_12
  60. 60. Perry JA, Cvitkovitch DG, Levesque CM (2009) Cell death in Streptococcus mutans biofilms: a link between CSP and extracellular DNA. FEMS Microbiol Lett 299: 261–266. doi: 10.1111/j.1574-6968.2009.01758.x
  61. 61. Dufour D, Cordova M, Cvitkovitch DG, Levesque CM (2011) Regulation of the competence pathway as a novel role associated with a streptococcal bacteriocin. J Bacteriol 193: 6552–6559. doi: 10.1128/jb.05968-11
  62. 62. Tamesada M, Kawabata S, Fujiwara T, Hamada S (2004) Synergistic effects of streptococcal glucosyltransferases on adhesive biofilm formation. J Dent Res 83: 874–879. doi: 10.1177/154405910408301110
  63. 63. Banas JA, Vickerman MM (2003) Glucan-binding proteins of the oral streptococci. Crit Rev Oral Biol Med 14: 89–99. doi: 10.1177/154411130301400203
  64. 64. Mattos-Graner RO, Napimoga MH, Fukushima K, Duncan MJ, Smith DJ (2004) Comparative analysis of Gtf isozyme production and diversity in isolates of Streptococcus mutans with different biofilm growth phenotypes. J Clin Microbiol 42: 4586–4592. doi: 10.1128/jcm.42.10.4586-4592.2004
  65. 65. Narisawa N, Kawarai T, Suzuki N, Sato Y, Ochiai K, et al. (2011) Competence-dependent endogenous DNA rearrangement and uptake of extracellular DNA give a natural variant of Streptococcus mutans without biofilm formation. J Bacteriol 193: 5147–5154. doi: 10.1128/jb.05240-11
  66. 66. Ueda S, Kuramitsu HK (1988) Molecular basis for the spontaneous generation of colonization-defective mutants of Streptococcus mutans. Mol Microbiol 2: 135–140. doi: 10.1111/j.1365-2958.1988.tb00014.x
  67. 67. Koo H, Xiao J, Klein MI, Jeon JG (2010) Exopolysaccharides produced by Streptococcus mutans glucosyltransferases modulate the establishment of microcolonies within multispecies biofilms. J Bacteriol 192: 3024–3032. doi: 10.1128/jb.01649-09
  68. 68. Perry D, Kuramitsu HK (1989) Genetic linkage among cloned genes of Streptococcus mutans. Infect Immun 57: 805–809.
  69. 69. Brady LJ, Maddocks SE, Larson MR, Forsgren N, Persson K, et al. (2010) The changing faces of Streptococcus antigen I/II polypeptide family adhesins. Mol Microbiol 77: 276–286. doi: 10.1111/j.1365-2958.2010.07212.x
  70. 70. Biswas I, Drake L, Erkina D, Biswas S (2008) Involvement of sensor kinases in the stress tolerance response of Streptococcus mutans. J Bacteriol 190: 68–77. doi: 10.1128/jb.00990-07
  71. 71. Song L, Sudhakar P, Wang W, Conrads G, Brock A, et al. (2012) A genome-wide study of two-component signal transduction systems in eight newly sequenced mutans streptococci strains. BMC Genomics 13: 128. doi: 10.1186/1471-2164-13-128
  72. 72. Rodriguez-Moya J, Argandona M, Reina-Bueno M, Nieto JJ, Iglesias-Guerra F, et al. (2010) Involvement of EupR, a response regulator of the NarL/FixJ family, in the control of the uptake of the compatible solutes ectoines by the halophilic bacterium Chromohalobacter salexigens. BMC Microbiol 10: 256. doi: 10.1186/1471-2180-10-256
  73. 73. Aikawa C, Furukawa N, Watanabe T, Minegishi K, Furukawa A, et al. (2012) Complete Genome Sequence of the Serotype k Streptococcus mutans Strain LJ23. J Bacteriol 194: 2754–2755. doi: 10.1128/jb.00350-12
  74. 74. Ouyang J, Tian XL, Versey J, Wishart A, Li YH (2010) The BceABRS four-component system regulates the bacitracin-induced cell envelope stress response in Streptococcus mutans. Antimicrob Agents Chemother 54: 3895–3906. doi: 10.1128/aac.01802-09
  75. 75. Bhagwat SP, Nary J, Burne RA (2001) Effects of mutating putative two-component systems on biofilm formation by Streptococcus mutans UA159. FEMS Microbiol Lett 205: 225–230. doi: 10.1111/j.1574-6968.2001.tb10952.x
  76. 76. Cvitkovitch DG, Li YH, Ellen RP (2003) Quorum sensing and biofilm formation in Streptococcal infections. J Clin Invest 112: 1626–1632. doi: 10.1172/jci20430
  77. 77. van der PloegJR (2005) Regulation of bacteriocin production in Streptococcus mutans by the quorum-sensing system required for development of genetic competence. J Bacteriol 187: 3980–3989. doi: 10.1128/jb.187.12.3980-3989.2005
  78. 78. Schneewind O, Missiakas DM (2012) Protein secretion and surface display in Gram-positive bacteria. Philos Trans R Soc Lond B Biol Sci 367: 1123–1139. doi: 10.1098/rstb.2011.0210
  79. 79. Abdallah AM, Gey van Pittius NC, Champion PA, Cox J, Luirink J, et al. (2007) Type VII secretion–mycobacteria show the way. Nat Rev Microbiol 5: 883–891. doi: 10.1038/nrmicro1773
  80. 80. Burts ML, Williams WA, DeBord K, Missiakas DM (2005) EsxA and EsxB are secreted by an ESAT-6-like system that is required for the pathogenesis of Staphylococcus aureus infections. Proc Natl Acad Sci U S A 102: 1169–1174. doi: 10.1073/pnas.0405620102
  81. 81. Burts ML, DeDent AC, Missiakas DM (2008) EsaC substrate for the ESAT-6 secretion pathway and its role in persistent infections of Staphylococcus aureus. Mol Microbiol 69: 736–746. doi: 10.1111/j.1365-2958.2008.06324.x
  82. 82. Petersen TN, Brunak S, von Heijne G, Nielsen H (2011) SignalP 4.0: discriminating signal peptides from transmembrane regions. Nat Methods 8: 785–786. doi: 10.1038/nmeth.1701
  83. 83. Vengadesan K, Narayana SV (2011) Structural biology of Gram-positive bacterial adhesins. Protein Sci 20: 759–772. doi: 10.1002/pro.613
  84. 84. Nakano K, Nomura R, Taniguchi N, Lapirattanakul J, Kojima A, et al. (2010) Molecular characterization of Streptococcus mutans strains containing the cnm gene encoding a collagen-binding adhesin. Arch Oral Biol 55: 34–39. doi: 10.1016/j.archoralbio.2009.11.008
  85. 85. Marchler-Bauer A, Lu S, Anderson JB, Chitsaz F, Derbyshire MK, et al. (2011) CDD: a Conserved Domain Database for the functional annotation of proteins. Nucleic Acids Res 39: D225–229. doi: 10.1093/nar/gkq1189
  86. 86. Kesavalu L, Lucas AR, Verma RK, Liu L, Dai E, et al. (2012) Increased atherogenesis during Streptococcus mutans infection in ApoE-null mice. J Dent Res 91: 255–260. doi: 10.1177/0022034511435101
  87. 87. Ogier JA, Wachsmann D, Scholler M, Lepoivre Y, Klein JP (1990) Molecular characterization of the gene sr of the saliva interacting protein from Streptococcus mutans OMZ175. Arch Oral Biol 35 Suppl: 25S–31S doi: 10.1016/0003-9969(90)90127-v
  88. 88. Martin V, Kleschyov AL, Scholler M, Klein JP, Beretz A (1997) Induction of NO production by polyosides from the wall of Streptococcus mutans OMZ175 in the rat aorta. Adv Exp Med Biol 418: 669–672. doi: 10.1007/978-1-4899-1825-3_156
  89. 89. Linzer R, Reddy MS, Levine MJ (1987) Structural studies of the serotype-f polysaccharide antigen from Streptococcus mutans OMZ175. Infect Immun 55: 3006–3010.
  90. 90. Jenkinson HF (2011) Beyond the oral microbiome. Environ Microbiol 13: 3077–3087. doi: 10.1111/j.1462-2920.2011.02573.x
  91. 91. Dewhirst FE, Chen T, Izard J, Paster BJ, Tanner AC, et al. (2010) The human oral microbiome. J Bacteriol 192: 5002–5017. doi: 10.1128/jb.00542-10
  92. 92. Marri PR, Hao W, Golding GB (2006) Gene gain and gene loss in streptococcus: is it driven by habitat? Mol Biol Evol 23: 2379–2391. doi: 10.1093/molbev/msl115
  93. 93. Fujita H, Hashimoto H, Shoda S, Suzuki T (2011) Dental caries prevalence as a product of agriculture and subsistence pattern at the Yean-ri site, South Korea. Caries Res 45: 524–531. doi: 10.1159/000331920
  94. 94. Littleton J, Frohlich B (1993) Fish-eaters and farmers: dental pathology in the Arabian Gulf. Am J Phys Anthropol 92: 427–447. doi: 10.1002/ajpa.1330920403
  95. 95. Lukacs JR (1992) Dental paleopathology and agricultural intensification in south Asia: new evidence from Bronze Age Harappa. Am J Phys Anthropol 87: 133–150. doi: 10.1002/ajpa.1330870202
  96. 96. Bender GR, Sutton SV, Marquis RE (1986) Acid tolerance, proton permeabilities, and membrane ATPases of oral streptococci. Infect Immun 53: 331–338.
  97. 97. Ahn SJ, Wen ZT, Burne RA (2007) Effects of oxygen on virulence traits of Streptococcus mutans. J Bacteriol 189: 8519–8527. doi: 10.1128/jb.01180-07
  98. 98. Ahn SJ, Rice KC, Oleas J, Bayles KW, Burne RA (2010) The Streptococcus mutans Cid and Lrg systems modulate virulence traits in response to multiple environmental signals. Microbiology 156: 3136–3147. doi: 10.1099/mic.0.039586-0
  99. 99. Engelberg-Kulka H, Amitai S, Kolodkin-Gal I, Hazan R (2006) Bacterial programmed cell death and multicellular behavior in bacteria. PLoS Genet 2: e135. doi: 10.1371/journal.pgen.0020135
  100. 100. Higuchi M, Yamamoto Y, Poole LB, Shimada M, Sato Y, et al. (1999) Functions of two types of NADH oxidases in energy metabolism and oxidative stress of Streptococcus mutans. J Bacteriol 181: 5940–5947.
  101. 101. Ahn SJ, Wen ZT, Burne RA (2006) Multilevel control of competence development and stress tolerance in Streptococcus mutans UA159. Infect Immun 74: 1631–1642. doi: 10.1128/iai.74.3.1631-1642.2006
  102. 102. Seaton K, Ahn SJ, Sagstetter AM, Burne RA (2011) A transcriptional regulator and ABC transporters link stress tolerance, (p)ppGpp, and genetic competence in Streptococcus mutans. J Bacteriol 193: 862–874. doi: 10.1128/jb.01257-10
  103. 103. Okinaga T, Niu G, Xie Z, Qi F, Merritt J (2010) The hdrRM operon of Streptococcus mutans encodes a novel regulatory system for coordinated competence development and bacteriocin production. J Bacteriol 192: 1844–1852. doi: 10.1128/jb.01667-09
  104. 104. Li YH, Tian XL, Layton G, Norgaard C, Sisson G (2008) Additive attenuation of virulence and cariogenic potential of Streptococcus mutans by simultaneous inactivation of the ComCDE quorum-sensing system and HK/RR11 two-component regulatory system. Microbiology 154: 3256–3265. doi: 10.1099/mic.0.2008/019455-0
  105. 105. Mashburn-Warren L, Morrison DA, Federle MJ (2010) A novel double-tryptophan peptide pheromone controls competence in Streptococcus spp. via an Rgg regulator. Mol Microbiol 78: 589–606. doi: 10.1111/j.1365-2958.2010.07361.x
  106. 106. Lemme A, Gröbe L, Reck M, Tomasch J, Wagner-Döbler I (2011) Subpopulation-specific transcriptome analysis of competence-stimulating-peptide-induced Streptococcus mutans. J Bacteriol 193: 1863–1877. doi: 10.1128/jb.01363-10
  107. 107. Petersen FC, Fimland G, Scheie AA (2006) Purification and functional studies of a potent modified quorum-sensing peptide and a two-peptide bacteriocin in Streptococcus mutans. Mol Microbiol 61: 1322–1334. doi: 10.1111/j.1365-2958.2006.05312.x
  108. 108. Hossain MS, Biswas I (2012) An Extracelluar Protease, SepM, Generates Functional CSP in Streptococcus mutans UA159. J Bacteriol.
  109. 109. Hale JD, Heng NC, Jack RW, Tagg JR (2005) Identification of nlmTE, the locus encoding the ABC transport system required for export of nonlantibiotic mutacins in Streptococcus mutans. J Bacteriol 187: 5036–5039. doi: 10.1128/jb.187.14.5036-5039.2005
  110. 110. Nomura R, Nakano K, Naka S, Nemoto H, Masuda K, et al. (2012) Identification and characterization of a collagen-binding protein, Cbm, in Streptococcus mutans. Mol Oral Microbiol 27: 308–323. doi: 10.1111/j.2041-1014.2012.00649.x
  111. 111. Way SS, Wilson CB (2005) The Mycobacterium tuberculosis ESAT-6 homologue in Listeria monocytogenes is dispensable for growth in vitro and in vivo. Infect Immun 73: 6151–6153. doi: 10.1128/iai.73.9.6151-6153.2005
  112. 112. Macrina FL, Jones KR, Alpert CA, Chassy BM, Michalek SM (1991) Repeated DNA sequence involved in mutations affecting transport of sucrose into Streptococcus mutans V403 via the phosphoenolpyruvate phosphotransferase system. Infect Immun 59: 1535–1543.