Search tips
Search criteria 


Logo of jbacterPermissionsJournals.ASM.orgJournalJB ArticleJournal InfoAuthorsReviewers
J Bacteriol. 2012 November; 194(21): 5886–5896.
PMCID: PMC3486099

An Extracelluar Protease, SepM, Generates Functional Competence-Stimulating Peptide in Streptococcus mutans UA159


Cell-cell communication in Gram-positive bacteria often depends on the production of extracellular peptides. The cariogenic bacterium Streptococcus mutans employs so-called competence-stimulating peptide (CSP) to stimulate mutacin (bacteriocin) production and competence development through the activation of the ComDE two-component pathway. In S. mutans, CSP is secreted as a 21-residue peptide; however, mass spectrometric analysis of culture supernatant indicates the presence of an 18-residue proteolytically cleaved species. In this study, using a transposon mutagenesis screening, we identified a cell surface protease that is involved in the processing of 21-residue CSP to generate the 18-residue CSP. We named this protease SepM for streptococcal extracellular protease required for mutacin production. We showed that the truncated 18-residue peptide is the biologically active form and that the specific postexport cleavage is a prerequisite to activate the ComDE two-component signal transduction pathway. We also showed that the CSP and the mutacins are exported outside the cell by the same ABC transporter, NlmTE. Our study further confirmed that the ComDE two-component system is absolutely necessary for mutacin production in S. mutans.


Quorum sensing is a primary means of bacterial communication that often uses secreted peptide pheromones to regulate expression of various genes when the bacterial cell density reaches a certain threshold concentration. Numerous cellular processes, such as virulence factor expression, extracellular enzyme production, antibiotic production, biofilm formation, and genetic exchange, are regulated by quorum sensing (25, 38, 48, 54, 55). In streptococci, competence-stimulating peptide (CSP)-mediated quorum sensing is a prerequisite for development of competence that leads to cellular DNA uptake from the surroundings for genetic diversity (41). The Gram-positive pathogen Streptococcus mutans, a primary causative agent of dental caries, employs a well-conserved quorum-sensing system called ComCDE that coordinates the expression of bacteriocins encoding genes and stimulates development of genetic competence (22, 23, 30, 40, 51, 57). The ComCDE-regulated quorum-sensing system has also been implicated in regulation of biofilm formation (30) and stress responses (28) in addition to other cellular processes.

Gram-positive bacteria use ribosomally synthesized peptides as quorum-signaling molecules (31). These peptides are typically translated as prepeptides that undergo processing during export to the extracellular environment (48). In S. mutans UA159, the peptide pheromone CSP is encoded by comC as a prepeptide with a leader sequence containing a conserved double glycine (GG) motif. During secretion through a dedicated ABC transporter complex, the N-terminal leader peptide is cleaved off by the proteolytic activity of the transporter to generate a mature peptide that is 21 residues long (CSP-21) (15, 30, 43). When the extracellular CSP concentration reaches a certain threshold, ComD, a membrane-associated histidine kinase, senses the signal. ComD is then activated by autophosphorylation and subsequently transfers the phosphate group to ComE, a cytoplasmic response regulator (24, 56). The activated ComE then stimulates the expression of various mutacins and mutacin-like genes by directly recognizing a conserved direct-repeat sequence present in the promoter regions; ComE also indirectly activates about 20 early competence-related genes (22, 26, 40, 51). While active CSP has been isolated from the culture supernatant of Streptococcus pneumoniae (18), the sequences of other known CSPs were deduced from the sequence information of the comC locus and the putative GG cleavage site (19). CSP sequences are highly variable among various streptococci, with very little or no sequence similarity with one another. The extracellular concentration of CSP required to stimulate gene expression also varies among streptococci. For example, a comparatively much higher concentration of synthetic CSP is required to exhibit the maximum stimulation in S. mutans than in any other streptococci (44). While maximum response is achieved at nanomolar concentrations of CSP in S. pneumoniae and Streptococcus intermedius (18, 42), S. mutans requires micromolar concentrations for optimum response (23, 44, 51, 58). A recent study suggests that in S. mutans, the CSP-21 peptide undergoes postexport processing at the C-terminal end that creates a more potent 18-residue peptide (CSP-18) that works at a much lower concentration than CSP-21 (41).

Being an oral pathogen, S. mutans leads a biofilm lifestyle in the oral cavity, which harbors more than 700 different species (1). This organism undergoes fierce competition with other bacteria for successful colonization and biofilm formation. This process is aided by the ribosomally synthesized small cationic peptides called mutacins (17). Mutacins are generally divided into the following two main categories (12, 35): the lantibiotics, which contain posttranslationally modified peptides with cyclic structure, forming lanthionine and β-methyllanthionine residues, and nonmodified peptide bacteriocins. Both lantibiotics (mutacin I, II, and III and Smb) (45, 46, 57) and nonlantibiotics (mutacins IV, V, and VI) (16, 20, 56) have been identified in S. mutans. Several studies have suggested that the CSP-mediated quorum-sensing system is involved in production of mutacin IV and V (23, 26, 40, 51) as well as the Smb lantibiotic (41, 57). The sequenced S. mutans strain UA159 encodes at least six other small peptides with a high degree of similarity with bacteriocins, and these genes have also been shown to be upregulated by CSP (23, 26, 40, 51).

The goal of this investigation was to identify regulatory factors that modulate the expression of mutacins. We used a transposon mutagenesis approach to identify a cell surface-associated protease encoded by SMU.518, which cleaves the 21-residue CSP to generate 18-residue CSP. We named this protein SepM for streptococcal extracellular protease required for mutacin production. By using reporter fusion assays, we reported that 18-residue CSP is the active form that induces the ComDE two-component signaling pathway. We also showed that both nonlantibiotic mutacins (NlmAB or NlmC) and CSP are exported outside the cell by the same ABC transporter, NlmTE.


Bacterial strains and growth conditions.

Strains and plasmids used are listed in Table 1. Escherichia coli strain DH5α was grown in Luria-Bertani medium supplemented, when necessary, with ampicillin (Amp; 100 μg/ml), erythromycin (Ery; 300 μg/ml), and kanamycin (Kan; 100 μg/ml). Various streptococci strains were routinely grown in Todd-Hewitt medium (BBL; Becton Dickson) supplemented with 0.2% yeast extract (THY) at 37°C. Lactococcus lactis MG1363 was grown in M17 medium supplemented with 0.5% glucose or THY medium at 30°C. When necessary, Ery (5 to 10 μg/ml), Kan (300 to 500 μg/ml), or spectinomycin (Spt; 300 μg/ml) was added to the sterile growth medium. Bacterial growth was monitored with a Klett-Summerson colorimeter as described previously (7).

Table 1
Strains and plasmids used in this study

Transformation assay.

To carry out the transformation assay, S. mutans strains were grown up to the desired growth stage (A595 of 0.2 or unless mentioned) and CSP-21 (200 nM) was added if needed, followed by 10 min incubation at 37°C. A total of 1.0 μg of linear or circular DNA was added to 1.0 ml culture, followed by incubation at 37°C for 2 h and plating onto THY agar plates with or without antibiotics. Transformation efficiencies are expressed as the number of transformants/number of viable cells/μg DNA.

Construction of PnlmA-gus reporter strains.

Plasmid pIB107 was used to construct the Pnlm-gusA reporter strains (8). This plasmid contains a promoterless gusA gene that is flanked by sequences from the SMU.1405 locus for integration by homologous recombination. A fragment carrying the promoter region of the nlmA gene was amplified from the UA159 genomic DNA using the primers EcoRI-SMU-NlmA-F and XhoI-NlmA-IGS-R. The amplified fragment was digested with EcoRI and XhoI and cloned into EcoRI-XhoI-digested pIB107 to generate pIB-D21. Plasmid pIB-D21 (or pIB107 for control) was linearized with BglI and transformed into different S. mutans strains to generate the reporter strains. PCR amplification and DNA sequencing verified the insertion of the promoter fusion construct at the SMU.1405 locus.

Isolation of SMU.518 mutants.

The procedure described by Zhang and Biswas (59) was used to generate insertion mutants of S. mutans. Briefly, IBS-D5 was transformed with pGhost9::ISS1 and transformants were selected on THY agar containing kanamycin and erythromycin at 30°C. An overnight culture was made from a single transformed colony (IBS-D23) at 30°C with erythromycin. Cultures were diluted 100-fold in the same medium, grown for 2 h at 30°C, and then shifted to 37°C for 2.5 h to select for transposition. Insertion mutants were selected on THY agar plates containing erythromycin and X-Gluc (5-bromo-4-chloro-3-indolyl-β-d-galactopyranoside) (200 μg/ml) at 37°C. Approximately 12,000 colonies were checked, and about 22 colonies were found to be white. These colonies were further verified for mutacin production, and mutacin-negative clones were selected for identification of the ISS1 insertion site as described previously (59). Briefly, templates generated by self-ligation of HindIII- or EcoRI-digested chromosomal DNA isolated from the mutants were subjected to inverse PCR by using the primers ISS1-ROUT2 and ISS1FOR4. The PCR products were purified and sequenced with the primer ISS1-R OUT 2 to identify the flanking sequences, which were mapped on the genome of S. mutans UA159 by BLAST search analysis. Three of the insertions were mapped in comC, two each were in comD, comE, nlmT, and SMU.518, and the rest were elsewhere in the genome.

Insertional inactivation of SMU.518.

SMU.518 was disrupted using a Cre-loxP-based method (4). The SMU.518 operon was amplified from S. mutans UA159 chromosomal DNA using the primers SMU.518 5′ F and SMU.518 3′ R and cloned into the pGEM-T Easy cloning vector (Promega) to create pIB-D48. A Kan-resistant cassette with flanking modified loxP sites was amplified from pUC4 ΩKan (39) using the primers lox71-Km-F and lox66-Km-R (for all primers, see Table 2) and cloned into EcoRV-digested pIB-D48 to generate pIB-D50. Plasmid pIB-D50 was linearized with EcoRI and transformed into S. mutans UA159 as previously described (9). The transformants were selected on THY plates containing Kan; one such transformant was selected and named IBS-D20. The Kan cassette was removed from the IBS-D20 chromosome using Cre-loxP-based recombination to create a clean deletion mutant, IBS-D29, as previously described (4). The deleted region was verified by PCR amplification followed by DNA sequencing.

Table 2
Oligonucleotides used in this study

Deletion of comC, nmlTE, and clsAB.

For deletion of comC, nmlTE, and clsAB, an overlapping fusion PCR approach was used as described previously (20). Briefly, ~1-kb upstream and downstream regions were amplified with the appropriate primer pairs (Table 2) for each gene using UA159 chromosomal DNA as a template. A Kan-resistant cassette was amplified from pIB-D38 with the primers NcoI-Kan-F and PstI-Kan-R, and overlapping fusion PCR was carried out using equal amounts of each PCR product with the flanking primers. The amplified fusion products were purified and transformed into S. mutans UA159 as described above to obtain intermediate strains. The Kan cassette was removed from the genome by Cre-loxP-based recombination to generate a clean deletion mutant for each locus. The final constructs were verified by PCR.

Cloning of SMU.518 and SMU.1915 (comC).

SMU.518 with the ribosome binding site was amplified from the genomic DNA of UA159 with high-fidelity AccuTaq DNA polymerase (Sigma) using the primers BamHI SMU.518 F and XhoI SMU.518 R. After digestion with BamHI and XhoI, the fragment was cloned into the BamHI-XhoI-digested vector pIB184Km (20). Positive clones were verified with restriction digestion and named pIB-D52. Similarly, full-length SMU.1915 was amplified with BamHI SMU.1915 F and XhoI SMU.1915 R primers and cloned BamHI-XhoI-digested pIB184Km to generate pIB-D58. An 18-amino-acid version of comC was cloned by the same approach, except the reverse primer was XhoI-18-aa-comC R; the plasmid was named pIB-D57.

RNA isolation and semiquantitative RT-PCR.

Total RNA was isolated from bacterial cultures according to the protocol described earlier (5). Briefly, S. mutans cultures were grown until the desired optical density was reached, and the bacterial cell pellet was resuspended into 5 ml of RNAprotect bacterial reagent (Qiagen). Total RNA was isolated using an RNeasy minikit (Qiagen) with a modified bacterial lysis step. The RNA concentrations were determined by UV spectrophotometer. A Titan one-tube reverse transcription-PCR (RT-PCR) system (Roche) was used as described previously (5). The gyrA gene was used as an internal control to ensure that an equal amount of RNA was being used in all RT-PCRs. The PCR products were separated on 2.0% agarose gel electrophoresis and quantified by Doc-It-LS (UVP) software.

β-Glucuronidase (GusA) assay.

Reporter strains were grown overnight at 37°C, diluted 1:20 in THY broth, and incubated at 37°C until the growth reached an optical density at 595 nm (OD595) of 0.2. CSP (intact CSP or CSPs treated with various strains) or the culture supernatants (40% ammonium sulfate precipitates from overnight cultures) of various strains were added, and cultures were incubated until the OD595 reached 1.00. Gus assays were performed as described previously by Biswas et al. (6), and the values are expressed in Miller units (MU).

Processing of synthetic CSP and CSP precipitation from culture supernatant.

CSP (21-mer) was commercially synthesized by GenScript (GenScript Corporation) with a purity grade of >95% and was resuspended in dimethyl formamide at a 2.0-mg/ml concentration. Bacterial cultures were grown overnight or to late logarithmic phase at 37°C. Cells were harvested from a 1-ml culture, washed, and resuspended in 100 μl of phosphate-buffered saline (PBS). CSP was added to a final concentration of 120 μM and incubated at 37°C for 1 h. Supernatant containing the CSP was separated from the cells. Processing of CSP was verified by mass spectrometric analysis or by spotting at different concentrations on a THY plate containing X-Gluc that was also seeded with the ΔSMU.518::PnlmA-gus reporter strain.

For CSP isolation from culture supernatants, cultures were grown overnight in a shaker incubator and the cells were removed by passing the culture through a 0.45 μM membrane filter. CSP was precipitated from the supernatant with 40% ammonium sulfate. The pellet was resuspended in 50 mM Tris-Cl (pH 7.4), equal amounts of each sample were added to the actively growing reporter strains (OD595 of 0.2), and Gus activity was measured.

Mass spectrometry.

Treated CSP samples were first desalted on a reverse-phase C4 column (Vydac; 300-Å pore size) with a buffer containing acetonitrile and formic acid on a NanoAcquity chromatographic system (Waters Corp.). Electrospray ionization (ESI) spectra were acquired on a Synapt G2 hybrid quadrupole/ion mobility/Tof mass spectrometer (Waters Corp.). Spectra were acquired at a 9,091-Hz pusher frequency, covering the mass range from 100 to 3,000 μm and accumulating data for 1.5 s per cycle. Time to mass calibration was made with NaI cluster ions acquired under the same conditions. Mass spectra of [Glu1]-fibrinopeptide B were acquired in parallel scans, and doubly charged ions at m/z 785.8426 were used as a lock mass reference. MassLynx (4.1) software was used to acquire and process data.

Bacteriocin assay.

Assays were done at 37°C under microaerophilic conditions according to a previously published protocol (20). Streptococcus gordonii (DL-1, ATCC 10558, and M5) and Streptococcus cristatus (BHT) were used as indicator strains for mutacin IV activity, while Lactococcus lactis MG1363 was used as an indicator for mutacin V. The diameters of clear zones in the lawn of the indicator strain, indicative of bacteriocin activity, were measured.


Identification and characterization of the SMU.518 locus.

To facilitate studies of the regulation of the nlmAB operon, we constructed a PnlmA-gusA reporter strain. The entire promoter region of nlmAB, including the first few codons of the nlmA open reading frame (ORF), was fused to a gusA reporter gene to create a transcriptional fusion. This PnlmA-gusA reporter fusion was then integrated at an ectopic location on the chromosome of strain UA159, and ISS1-mediated random transposon mutagenesis was carried out in the reporter fusion strain. Since our goal was to identify factors that stimulate mutacin IV production, we screened for colonies containing ISS1 for the white-color phenotype by a plate assay using X-Gluc to monitor expression of GusA from the PnlmA-gusA reporter fusion. These colonies were further verified for mutacin production. Two of the ISS1 insertions were mapped on a unique gene, SMU.518, which encodes a 346-residue polypeptide and is highly conserved among Gram-positive bacteria and mycobacteria. SMU.518 contains at least one transmembrane domain (from residue 10 to 26), a eukaryotic-type PDZ domain (from residue 131 to 195), and a C-terminal Lon-like protease (S16) domain (from residue 233 to 314). However, this protein does not contain any canonical ATP binding motifs, such as Walker A or B. We renamed this gene sepM for streptococcal extracellular protease required for mutacin production. SepM appears to be organized in an operon with two other genes, SMU.516 and kdtB (phosphopantetheine adenylyltransferase), involved in biosynthesis of lipopolysaccharide. The ISS1 insertions occurred in the sepM gene at residues 116 and 229. In silico analysis of the intergenic sequence (IGS) upstream of SMU.516 indicates the presence of a strong promoter-like sequence (−10 box [AGTTATAAT] and −35 box [TTGTTA]), suggesting that sepM is cotranscribed with SMU.516 and kdtB. Analysis of the IGS present downstream of SMU.518 suggests a rho-independent terminator sequence (GTTCTAGAATTTCTAGGAC) immediately after the sepM stop codon.

To verify that the observed mutacin-negative phenotype was not due to mutations elsewhere in the genome, we constructed a markerless clean deletion mutation of the SMU.518 locus in the UA159 strain using a Cre-loxP-based system. The SepM deletion mutant, IBS-D29, did not show any significant growth difference compared to wild-type UA159 (data not shown). We then tested bacteriocin production by a deferred antagonism assay (20). As shown in Fig. 1A, inactivation of sepM completely abolished both mutacin IV and V activities. To further confirm that this effect was due to inactivation of the sepM locus, we performed a complementation analysis. A fragment containing the entire sepM locus with the ribosome binding site was cloned into the E. coli-streptococcal shuttle vector pIB184Km (20) to generate the plasmid pIB-D52, and it was introduced into IBS-D29. As shown in Fig. 1A, supplying sepM in trans completely rescued the mutant phenotypes for both mutacin IV and V activities. Thus, our results suggest that sepM is involved in mutacin production in S. mutans UA159.

Fig 1
Expression of mutacins by various S. mutans strains. (A) Deferred antagonism assay for the production of mutacin IV and V. Bacterial cultures were stabbed onto THY agar and incubated overnight at 37°C under microaerophilic conditions. The plates ...

SepM regulates several putative bacteriocin genes at the transcriptional level.

To understand the exact molecular basis of the defective mutacin phenotype in IBS-D29, we tested the expression of mutacin IV (encoded by nlmA and nlmB genes) and V (nlmC) by semiquantitative RT-PCR (sqRT-PCR). As shown in Fig. 1B, inactivation of sepM caused the reduction of nlmA expression below the detection level and the nlmC expression was reduced around 15-fold in comparison to wild-type UA159. To get a better understanding of the regulation of mutacin production by SepM, we also tested expression of two other putative mutacin-encoding genes, SMU.423 and SMU.1906. Surprisingly, we found that both genes were remarkably downregulated in the mutant strain (Fig. 1B). However, complementation of sepM can fully restore the expression of these four genes to the wild-type strain. Taken together, our sqRT-PCR results suggest that SMU.518 functions at the transcriptional level and the presence of sepM activates the transcription of mutacin-encoding genes in S. mutans UA159.

SepM does not affect comC transcription.

All of the four mutacin-encoding genes also carry the highly conserved ComE consensus binding in their promoter region and are controlled by the ComDE two-component signal transduction system (24, 26, 40, 51, 56). Consequently, these genes are also CSP inducible. Our initial studies also revealed that the sepM mutant produces the same mutacin-defective phenotype as comDE mutants and impelled us to hypothesize that the observed phenotypes were due to a defect in CSP expression. To test whether SepM affected the CSP expression, we performed sqRT-PCR of the comC gene and nlmT, which encodes the transporter for mutacins. As shown in Fig. 1B, there was no significant change in comC transcription compared to that for UA159. We also tested the expression of comDE and did not find any significant difference between the wild-type and mutant strains (data not shown). Thus, our results suggest that the expression of CSP-21, its cognate two-component pathway, and the mutacin secretion apparatus was not perturbed in the mutant. The observed mutacin-null phenotype may be due to a potential postexport role of SepM on CSP-21. This notion was further supported by the finding that addition of synthetic CSP-21 was not able to restore the Gus activity in the reporter strain IBS-D37 (ΔsepM::PnlmA-gus), compared to the wild-type (IBS-D5) strain, where Gus activity was significantly stimulated by the CSP-21 addition (Fig. 2A).

Fig 2
Effect of synthetic CSP on PnlmA expression and transformation. (A) GusA activity of the reporter strains. Expression from PnlmA at mid-exponential phase was quantified in the IBS-D5 (PnlmA-gusA) and IBS-D37 (ΔsepM::PnlmA-gus) strains by determining ...

CSP cannot stimulate the competence in the ΔsepM mutant.

In S. mutans, the presence of the ComCDE quorum-sensing system enhances competence development (29, 32, 49), and addition of CSP can significantly augment the transformation efficiency of S. mutans (2, 3, 23, 29, 32, 36, 40, 41, 49). To verify the potential role of SepM in the postexport modulation of CSP, we carried out transformation assays in the presence or absence of synthetic CSP. We found that external addition of CSP-21 did not stimulate transformation in the ΔsepM strain as it did in the UA159 and the complemented strains (Fig. 2B). Nevertheless, the mutant was transformable without external CSP-21 and the efficiency was similar to that of the wild type (Fig. 2B). This basal level of transformation phenotype is the hallmark of the ComDE mutation (2, 29, 32, 36, 40). Thus, this result further substantiates the idea that SepM is somehow involved with the ComDE pathway.

SepM processes synthetic CSP.

SepM is predicted to be a membrane-associated protein, and its ortholog in group A streptococcus (GAS) is localized on the surface (13, 47). Since a putative Lon-like protease domain is encoded within the sequence, we speculated that SepM might be involved in the postexport maturation of CSP-21. Interestingly, CSP has been isolated from the culture supernatant as the CSP-18 peptide, and this truncated peptide is more potent than CSP-21 (41). To investigate if SepM can process CSP, we incubated CSP-21 with the cell pellets of UA159, ΔsepM mutant, and sepM-complemented strains as described in Materials and Methods. The treated samples were then tested for PnlmA-gus reporter fusion induction using the IBS-D37 strain (ΔsepM::PnlmA-gus). Surprisingly, we found that CSP-21 processed by UA159 and the complemented strain was able to produce blue color on a THY-X-Gluc plate containing IBS-D37 at a much lower concentration than the CSP-21 treated with ΔsepM mutant cells, for which blue color was developed to a limited extent only with undiluted CSP-21 (Fig. 3A). The control CSP-21 (incubated with PBS) produced faint blue color on a THY-X-Gluc plate at the highest CSP concentration (100 uM). We also performed a quantitative Gus assay with the CSP-21 that has been treated with different strains. Consistent with our qualitative assay, we found that CSP treated by either UA159 or the sepM-complemented strain was able to induce Pnlm expression at the 50 nM concentration, while CSP treated with the mutant strain or PBS alone failed to induce expression (Fig. 3B). To verify if the Lon-like protease domain is involved in the maturation of CSP-21, we first treated the cells (UA159) with a serine protease inhibitor, 4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride (AEBSF) (37), or proteinase K (to remove the cell surface proteins). These treated cells were incubated with synthetic CSP. We observed that prior treatment of cells with either AEBSF or proteinase K was able to completely abolish the SepM-mediated CSP processing (Fig. 3B and data not shown). Thus, SepM appears to process the synthetic CSP-21 to generate a potent signaling molecule, and the proteolytic activity of SepM is necessary for the observed processing.

Fig 3
Processing of synthetic CSP by S. mutans UA159 derivatives. (A) Expression of PnlmA-gusA (IBS-D37) with the processed CSP samples. Bacterial cells were incubated with 21-residue synthetic CSP at 37°C for an hour. Cells were separated from the ...

SepM cleaves C-terminal three residues at the C terminus of CSP.

Treating CSP-21 with the UA159 cell pellet produces an active CSP that triggers gene expression. Petersen et al. (41) previously identified a truncated derivative, CSP-18, which is more potent in competence development than the full-length CSP. Thus, it is possible that SepM may cleave the full-length CSP-21 to generate the truncated CSP-18 peptide. To confirm this notion, CSP samples that were incubated with various strains were analyzed by matrix-assisted laser desorption ionization–time of flight (MALDI-TOF) mass spectrometry. As shown in Fig. 4, only the 21-residue peptide was identified as the major peak (2,364 Da) in the control CSP sample (treated with PBS). Samples treated with UA159 generated two major peaks: one corresponds to the truncated CSP-18 (2,066-Da) form, and the other corresponds to the full-length CSP-21 form. Several other minor peaks were also present in the sample. In the CSP-21 sample that was treated with ΔsepM mutant cells, the peak corresponding to CSP-18 was conspicuously absent, although several minor peaks were visible. Thus, our mass spectrometry data suggest that SepM cleaves the full-length CSP-21 at the C-terminal end to generate the active CSP-18 derivative that acts as a signaling molecule to activate ComDE.

Fig 4
Mass spectrometric analysis of processed CSP. CSP samples were treated with either UA159 or ΔsepM cells. After removing the cells, the samples were desalted on a reverse-phase C4 column and then subjected to MALDI-TOF ESI mass spectrometry on ...

The functional form of CSP is 18 residues.

Our mass spectrometry data suggest that the functional form of CSP is the 18-residue derivative and not the 21-residue form. To confirm it further, we cloned the CSP-18 derivative (without the C-terminal three residues) and the CSP-21 form (as a control) into the shuttle plasmid pIB184Km. These clones also contain the native signal peptide sequence for secretion outside the cell. These plasmids were introduced into the ΔsepM ΔcomC double mutant IBS-D45 and examined for mutacin production. As expected, the plasmid carrying the nucleotide sequences for the 18-residue form was able to restore the mutacin production, while the plasmid carrying the full-length comC gene was not able to restore the mutacin production in the double mutant IBS-D45 (Fig. 5A).

Fig 5
In vivo complementation with the truncated CSP derivative. (A) Mutacin IV activity was tested in the IBS-D45 (ΔsepM ΔcomC) strain complemented with either full-length (21-residue) or 18-residue CSP from a plasmid. Mutacin assays were performed ...

To further confirm that CSP-18 is the functional form of CSP, we precipitated the culture supernatants (40% ammonium sulfate) from the IBS-D45 strain carrying either the 18- or 21-amino-acid version of comC, added the precipitates to the actively growing culture of the IBS-D46 reporter strain (ΔsepM ΔcomC::PnlmA-gus), and measured the Gus activity. As shown in Fig. 5B, culture supernatants of wild-type UA159 and IBS-D45 with the 18-residue CSP and ΔsepM::SMU.518 showed similar Gus activities. However, IBS-D45 with the 21-residue CSP was not able to restore the Gus activity of the double mutant. Culture supernatants from the ΔnlmTE (see below), ΔcomC, and ΔsepM mutants and the mock (PBS) treatment were used as controls. Taken together, our data suggest that CSP-21 is nonfunctional, and cleavage of three C-terminal residues is required to generate CSP-18, which is the functional form.

CSP is essential for mutacin production.

The results presented above suggest that CSP is the sole regulator for mutacin production in S. mutans. Our original mutagenesis screening also revealed that the ComDE system is essential for mutacin production, since we obtained insertions in both genes (data not shown). This finding is in agreement with a previous report that showed that mutation of any of the components of the ComDE pathway can abolish mutacin production (51). To confirm the involvement of CSP in mutacin production, we constructed a markerless comC deletion mutant and tested it against the indicator bacteria for mutacin IV activity. As expected, no mutacin activity was observed in the ΔcomC mutant, while the mutant carrying a plasmid with comC was able to completely restore the mutacin IV production (Fig. 6A). Furthermore, external addition of synthetic CSP also restored the Gus activity in the ΔcomC strain carrying the PnlmA-gusA reporter fusion (Fig. 6B). Thus, consistent with the previous findings, our results suggest that CSP is essential for induction of mutacin genes in S. mutans.

Fig 6
CSP is essential for mutacin gene expression. (A) Mutacin IV activity of the ΔcomC strain. Assays were performed at least three times, and a representative plate is shown. (B) Expression of PnlmA in the presence (50 nM) or absence of CSP. The ...

NlmTE (ComAB) is the transporter of CSP.

Based on the sequence homology and competence phenotype, it was previously proposed that the ABC transporter CslAB (encoded by SMU.1897 to SMU.1900) is responsible for the secretion CSP in S. mutans (15, 43). Since CSP is essential for mutacin production in S. mutans, a mutation in genes related to the transporter of CSP should abrogate mutacin production. Since we obtained two ISS1 insertions in the nlmTE locus (SMU.286 to SMU.287) that did not express PnlmA-gusA (and generated the mutacin-negative phenotype), we speculated that NlmTE might be the transporter for CSP. To confirm our hypothesis, we deleted either the nlmTE or cslAB locus and assayed for mutacin activity. As expected, we found that deletion of nlmTE caused the complete abrogation of mutacin production, while there was no discernible change in mutacin production when the cslAB locus was deleted (Fig. 7A).

Fig 7
Mutacin production by ABC transporter mutants. (A) Mutacin IV activity of the ΔcomC, ΔnlmTE, and ΔcslAB strains was measured. Antagonism assays were performed as described in Fig. 1, and a representative plate from at least three ...

Since mutation in the transporter responsible for CSP secretion should block the endogenous CSP from secretion to the outside, addition of the culture supernatant from the mutant strain should not induce PnlmA-gusA. To further confirm that NlmTE, and not CslAB, is the transporter of CSP, we precipitated the culture supernatants by 40% ammonium sulfate from both the nlmTE and cslAB mutants, added them to the actively growing cultures of two reporter strains, IBS-D37 (ΔsepM::PnlmA-gusA) and IBS-D40 (ΔcomC::PnlmA-gusA), and measured Gus activity. We also included culture supernatants from UA159 as a positive control and from the ΔcomC and ΔsepM strains as negative controls. As shown in Fig. 7B, addition of culture supernatant from the ΔnlmTE mutant did not stimulate the Gus gene expression in the reporter strains, whereas culture supernatants from the ΔcslAB mutant were able to completely restore the Gus activity. To further validate our hypothesis, we constructed another reporter strain in the ΔnlmTE background (IBS-D41). PnlmA-gusA was not expressed in the IBS-D41 strain. However, PnlmA-gusA was expressed upon addition of the culture supernatant of the ΔcslAB mutant but not that of the ΔnlmTE mutant (Fig. 7B). Furthermore, when we performed a transformation assay with the ΔcomC and ΔnlmTE mutants, the transformation efficiencies were similar in these two strains (Fig. 7C). However, addition of CSP to these strains augmented the transformation efficiency at least 100-fold. Taken together, our studies suggest that CSP, like mutacins, uses the same transporter, NlmTE, for secretion outside the cells.


For successful colonization, S. mutans produces various mutacins to inhibit the growth of other competing species, thereby helping establish a flourishing biofilm in the oral cavity (27). Mutacin-mediated cell killing is also considered a source of nutrients during nutrient-limited conditions (34). This process is highly coordinated and regulated by the CSP-mediated ComDE quorum-sensing signaling pathway (23, 26, 40, 51). Petersen and colleagues (41) recently isolated CSP-18, a truncated form of CSP (lacking the C-terminal three amino acids) that is functionally more potent than CSP-21, from the culture supernatant. However, the exact mechanism by which this truncated CSP-18 is generated in S. mutans was unknown. Here, we set up a screening system to identify putative transcriptional regulators for mutacin gene expression in S. mutans. Our screening yielded a cell surface-associated protease, SepM, which appears to cleave CSP-21 to generate the truncated CSP-18 peptide that acts as a true signaling molecule.

SepM encodes a Lon-like proteolytic domain at the C-terminal domain and a PDZ domain at the middle region. The Lon protease is a cytoplasmic serine protease consisting of hexameric rings of a single peptide chain carrying the peptidase domain, an AAA+ domain, and a domain with chaperone activity. Lon is essential for maintaining cellular homeostasis by mediating the degradation of nearly half of the abnormal or damaged proteins in E. coli (33). Lon participates in numerous biological processes, such as quorum sensing, motility, biofilm formation, and stress responses (for a recent review, see reference 52). Although Lon is widely distributed in various bacterial kingdoms, some members of the Firmicutes, including streptococci, do not encode a true Lon protease. Whereas most of the Lon proteases characterized so far contain ATP binding motifs and reside in the cytosol, SepM does not possess any ATPase domain and is surface localized. Thus, this SepM protease is a unique class of Lon-like protease present in streptococci.

The active site of Lon is composed of a Ser-Lys dyad and shares no relation to the classical catalytic Ser-His-Asp triad of serine proteases (10). SepM also contains a Ser-Lys dyad (S235 and K280), and when we mutated these two residues to Ala, the protein lost its catalytic activity to process CSP (data not shown), suggesting that SepM is truly a member of the Lon-like protease family. Consistent with this observation, we also found that treating cells with AEBSF inhibits the protein's ability to process CSP. Though it appears that SepM behaves like a Lon protease, several important fundamental differences do exist. For example, Lon degrades a variety of proteins, such as phage λ N protein and CcdA antitoxin, that are either denatured or naturally partially unfolded. Generally, Lon recognizes an N-terminal degradation tag that is rich in nonpolar amino acids (52). Furthermore, Lon can degrade C-terminally SsrA-tagged proteins in E. coli as well as in Mycoplasma (11, 14). Peptide bond hydrolysis is thought to occur in a processive linear manner from the N to the C terminus or vice versa. We have no evidence that suggests that SepM displays a processive protease activity; rather, SepM acts as a site-specific endoprotease and cleaves after the Ala residue at position 18 of the CSP. A recent report by Tian et al. (50) suggests that the Ala18 residue is critical, since deletion or replacement of this residue completely abolished the signaling activity of the peptide. Thus, we speculate that SepM requires the Ala residue either for correct recognition or for optimum cleavage activity. Furthermore, SepM also needs all three residues after Ala for optimum activity, since deletion of the last Lys residue drastically reduces the peptide activity as well (49).

Homologs of SepM have been found in a wide range of Gram-positive organisms, including all streptococci, bacilli, staphylococci, and lactobacilli. Sequence alignment suggests that the protein is highly conserved across the species (data not shown), yet the specificity of the CSP cleavage activity is highly restricted to S. mutans. For example, when synthetic CSP-21 was incubated with the cells from various streptococci, such as Streptococcus agalactiae, it failed to produce a positive signal, suggesting that CSP-21 is either not processed or improperly processed (data not shown). Thus, apart from the conserved Ser-Lys dyad (S235 and K280), other residues of SepM seem to play important roles in determining the substrate specificity. Further biochemical studies are required to fully understand the molecular mechanism of substrate recognition and specificity.

So far, CSP-21 is the only substrate that is recognized and cleaved by the SepM protease. However, it is very likely that SepM might be involved in the maturation of other peptide pheromones produced by S. mutans. Mashburn-Warren et al. (32) recently described a novel competence regulatory peptide (XIP) that is absolutely necessary for competence development. This XIP peptide is produced as a 17-residue propeptide that is processed outside the cell to generate the active seven-residue XIP pheromone (32). Although SepM seems to be the ideal candidate for the XIP processing, SepM is probably not involved in XIP processing. If SepM were responsible for the maturation of XIP, then one would expect the ΔsepM mutant strain to be completely competent deficient, which was not the case (Fig. 2B). SepM might be involved in the inactivation of bacteriocins produced by other bacteria that are present in the dental plaque. In Streptococcus pyogenes, Spy1356, the SepM homolog, has multiple functions. Spy1356 is crucial for surface expression of various proteins, including M protein, and responsible for binding with the extracellular matrix proteins (13). Further studies are necessary to unravel the range of processes regulated by SepM and its homologs in other Gram-positive organisms.

The key question that remains to be answered is why S. mutans employs an additional regulatory mechanism for the generation of the active signaling peptide when other streptococci do not use this strategy. There are several possibilities that may explain the need for the extra three amino acids at the C terminus. First, these terminal amino acids may provide protection against the proteolytic degradation that may occur in the dense multispecies biofilms. Indeed, CSP-21 is degraded by an S. gordonii cell surface protease to provide a selective advantage to S. gordonii in the natural habitat (53). Second, the presence of the three amino acids may confer protection against its own surface proteases. Third, the intact CSP-21 may have additional functions other than participating in quorum signaling. For example, CSP-21 can act as a bacteriocin, and Petersen et al. (41) have shown that CSP-21 can inhibit growth of other streptococci. Furthermore, Jarosz et al. (21) have shown that CSP-21 inhibits the morphological switch from yeast to hypha formation in Candida albicans.

The other important finding that emerged from our study was the identification of NlmTE as the transporter for CSP. S. mutans encodes several ABC transporters, and the CslAB transporter was thought to be responsible for the CSP transport. This is because mutations in the cslAB locus drastically reduce the natural transformation in S. mutans (15, 43). However, the data presented here support that NlmTE and not CslAB is the authentic transporter of CSP. For example, we found that the ΔnlmTE mutant was unable to express PnlmA-gusA. In addition, while the culture supernatant of the ΔnlmTE mutant was not able to trigger the PnlmA-gusA expression, the external addition of CSP was able to restore the PnlmA-gusA expression in the ΔnlmTE mutant (Fig. 8). Furthermore, the ΔnlmTE mutant behaved the same way as the ΔcomC mutant for natural transformation.

Fig 8
Model for the regulation of mutacin production in S. mutans UA159. Competence-stimulating peptide (CSP) is translated as a prepro-CSP (i). During secretion through the ABC transporter, NlmTE, the leader peptide, is cleaved to generate the 21-residue-long ...

Based on our current results and existing evidence on mutacin production, we present a revised model for regulation of mutacin expression by the CSP peptide pheromone in S. mutans (Fig. 8). According to this model, CSP is translated as a 46-residue-long prepropeptide. This 46-residue peptide is transported through the NlmTE transporter, the same transporter that also secretes various mutacins. During transport, the leader sequence is cleaved off after the GG motif (present at position 25) by the proteolytic activity of NlmTE. The exported propeptide CSP further undergoes proteolytic cleavage at the C termini by the cell surface protease SepM. This matured 18-residue CSP peptide acts as a signaling molecule and turns on the ComDE two-component signal transduction pathway, which, in turn, positively regulates expression of various mutacin and mutacin-related genes. An important question that arises from this model is whether the processing of CSP by SepM is coupled with the secretion and whether the NlmTE transporter directly interacts with the SepM protease. Although we have no direct evidence, we believe that these two processes are not coupled. Experiments are under way to unravel the molecular details of CSP-mediated signaling in S. mutans and related streptococci.


We thank Todd Williams and Nadya Galeva (Mass Spectrometry Lab, University of Kansas) for the mass spectrometry analysis. We thank Mahesh Jakkula for the construction of the CslAB mutant strain and the members of the Biswas laboratory for their insightful comments during the course of this study.

This work was supported in part by an NIDCR grant (DE021664) to I.B.


Published ahead of print 24 August 2012


1. Aas JA, et al. 2008. Bacteria of dental caries in primary and permanent teeth in children and young adults. J. Clin. Microbiol. 46: 1407– 1417 [PMC free article] [PubMed]
2. Ahn SJ, Wen ZT, Burne RA. 2006. Multilevel control of competence development and stress tolerance in Streptococcus mutans UA159. Infect. Immun. 74: 1631– 1642 [PMC free article] [PubMed]
3. Allan E, et al. 2007. Genetic variation in comC, the gene encoding competence-stimulating peptide (CSP) in Streptococcus mutans. FEMS Microbiol. Lett. 268: 47– 51 [PubMed]
4. Banerjee A, Biswas I. 2008. Markerless multiple-gene-deletion system for Streptococcus mutans. Appl. Environ. Microbiol. 74: 2037– 2042 [PMC free article] [PubMed]
5. Biswas I, Drake L, Biswas S. 2007. Regulation of gbpC expression in Streptococcus mutans. J. Bacteriol. 189: 6521– 6531 [PMC free article] [PubMed]
6. Biswas I, Jha JK, Fromm N. 2008. Shuttle expression plasmids for genetic studies in Streptococcus mutans. Microbiology 154: 2275– 2282 [PubMed]
7. Biswas I, Scott JR. 2003. Identification of rocA, a positive regulator of covR expression in the group A streptococcus. J. Bacteriol. 185: 3081– 3090 [PMC free article] [PubMed]
8. Biswas S, Biswas I. 2006. Regulation of the glucosyltransferase (gtfBC) operon by CovR in Streptococcus mutans. J. Bacteriol. 188: 988– 998 [PMC free article] [PubMed]
9. Biswas S, Biswas I. 2005. Role of HtrA in surface protein expression and biofilm formation by Streptococcus mutans. Infect. Immun. 73: 6923– 6934 [PMC free article] [PubMed]
10. Botos I, et al. 2004. The catalytic domain of Escherichia coli Lon protease has a unique fold and a Ser-Lys dyad in the active site. J. Biol. Chem. 279: 8140– 8148 [PubMed]
11. Choy JS, Aung LL, Karzai AW. 2007. Lon protease degrades transfer-messenger RNA-tagged proteins. J. Bacteriol. 189: 6564– 6571 [PMC free article] [PubMed]
12. Cotter PD, Hill C, Ross RP. 2005. Bacteriocins: developing innate immunity for food. Nat. Rev. Microbiol. 3: 777– 788 [PubMed]
13. Fritzer A, et al. 2010. Novel conserved group A streptococcal proteins identified by the antigenome technology as vaccine candidates for a non-M protein-based vaccine. Infect. Immun. 78: 4051– 4067 [PMC free article] [PubMed]
14. Gur E, Sauer RT. 2008. Recognition of misfolded proteins by Lon, a AAA(+) protease. Genes Dev. 22: 2267– 2277 [PubMed]
15. 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 [PMC free article] [PubMed]
16. Hale JD, Ting YT, Jack RW, Tagg JR, Heng NC. 2005. Bacteriocin (mutacin) production by Streptococcus mutans genome sequence reference strain UA159: elucidation of the antimicrobial repertoire by genetic dissection. Appl. Environ. Microbiol. 71: 7613– 7617 [PMC free article] [PubMed]
17. Hamada S, Ooshima T. 1975. Production and properties of bacteriocins (mutacins) from Streptococcus mutans. Arch. Oral Biol. 20: 641– 648 [PubMed]
18. Havarstein LS, Coomaraswamy G, Morrison DA. 1995. An unmodified heptadecapeptide pheromone induces competence for genetic transformation in Streptococcus pneumoniae. Proc. Natl. Acad. Sci. U. S. A. 92: 11140– 11144 [PubMed]
19. Havarstein LS, Hakenbeck R, Gaustad P. 1997. Natural competence in the genus Streptococcus: evidence that streptococci can change pherotype by interspecies recombinational exchanges. J. Bacteriol. 179: 6589– 6594 [PMC free article] [PubMed]
20. Hossain MS, Biswas I. 2011. Mutacins from Streptococcus mutans UA159 are active against multiple streptococcal species. Appl. Environ. Microbiol. 77: 2428– 2434 [PMC free article] [PubMed]
21. Jarosz LM, Deng DM, van der Mei HC, Crielaard W, Krom BP. 2009. Streptococcus mutans competence-stimulating peptide inhibits Candida albicans hypha formation. Eukaryot. Cell 8: 1658– 1664 [PMC free article] [PubMed]
22. Kreth J, et al. 2007. The response regulator ComE in Streptococcus mutans functions both as a transcription activator of mutacin production and repressor of CSP biosynthesis. Microbiology 153: 1799– 1807 [PMC free article] [PubMed]
23. Kreth J, Merritt J, Shi W, Qi F. 2005. Co-ordinated bacteriocin production and competence development: a possible mechanism for taking up DNA from neighbouring species. Mol. Microbiol. 57: 392– 404 [PMC free article] [PubMed]
24. Kreth J, Merritt J, Zhu L, Shi W, Qi F. 2006. Cell density- and ComE-dependent expression of a group of mutacin and mutacin-like genes in Streptococcus mutans. FEMS Microbiol. Lett. 265: 11– 17 [PubMed]
25. Lanigan-Gerdes S, Dooley AN, Faull KF, Lazazzera BA. 2007. Identification of subtilisin, Epr and Vpr as enzymes that produce CSF, an extracellular signalling peptide of Bacillus subtilis. Mol. Microbiol. 65: 1321– 1333 [PubMed]
26. Lemme A, Grobe L, Reck M, Tomasch J, Wagner-Dobler I. 2011. Subpopulation-specific transcriptome analysis of competence-stimulating-peptide-induced Streptococcus mutans. J. Bacteriol. 193: 1863– 1877 [PMC free article] [PubMed]
27. Lemos JA, Abranches J, Burne RA. 2005. Responses of cariogenic streptococci to environmental stresses. Curr. Issues Mol. Biol. 7: 95– 107 [PubMed]
28. Li YH, Hanna MN, Svensater G, Ellen RP, Cvitkovitch DG. 2001. Cell density modulates acid adaptation in Streptococcus mutans: implications for survival in biofilms. J. Bacteriol. 183: 6875– 6884 [PMC free article] [PubMed]
29. 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 [PMC free article] [PubMed]
30. Li YH, et al. 2002. A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. J. Bacteriol. 184: 2699– 2708 [PMC free article] [PubMed]
31. Lyon GJ, Novick RP. 2004. Peptide signaling in Staphylococcus aureus and other Gram-positive bacteria. Peptides 25: 1389– 1403 [PubMed]
32. 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 [PMC free article] [PubMed]
33. Maurizi MR. 1992. Proteases and protein degradation in Escherichia coli. Experientia 48: 178– 201 [PubMed]
34. Nguyen T, et al. 2009. Genes involved in the repression of mutacin I production in Streptococcus mutans. Microbiology 155: 551– 556 [PMC free article] [PubMed]
35. Nissen-Meyer J, Nes IF. 1997. Ribosomally synthesized antimicrobial peptides: their function, structure, biogenesis, and mechanism of action. Arch. Microbiol. 167: 67– 77 [PubMed]
36. 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 [PMC free article] [PubMed]
37. Overall CM, Blobel CP. 2007. In search of partners: linking extracellular proteases to substrates. Nat. Rev. Mol. Cell Biol. 8: 245– 257 [PubMed]
38. Parsek MR, Greenberg EP. 2005. Sociomicrobiology: the connections between quorum sensing and biofilms. Trends Microbiol. 13: 27– 33 [PubMed]
39. Perez-Casal J, Caparon MG, Scott JR. 1991. Mry, a trans-acting positive regulator of the M protein gene of Streptococcus pyogenes with similarity to the receptor proteins of two-component regulatory systems. J. Bacteriol. 173: 2617– 2624 [PMC free article] [PubMed]
40. Perry JA, Jones MB, Peterson SN, Cvitkovitch DG, Levesque CM. 2009. Peptide alarmone signalling triggers an auto-active bacteriocin necessary for genetic competence. Mol. Microbiol. 72: 905– 917 [PMC free article] [PubMed]
41. 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 [PubMed]
42. Petersen FC, Pecharki D, Scheie AA. 2004. Biofilm mode of growth of Streptococcus intermedius favored by a competence-stimulating signaling peptide. J. Bacteriol. 186: 6327– 6331 [PMC free article] [PubMed]
43. Petersen FC, Scheie AA. 2000. Genetic transformation in Streptococcus mutans requires a peptide secretion-like apparatus. Oral Microbiol. Immunol. 15: 329– 334 [PubMed]
44. Petersen FC, Tao L, Scheie AA. 2005. DNA binding-uptake system: a link between cell-to-cell communication and biofilm formation. J. Bacteriol. 187: 4392– 4400 [PMC free article] [PubMed]
45. Qi F, Chen P, Caufield PW. 1999. Purification of mutacin III from group III Streptococcus mutans UA787 and genetic analyses of mutacin III biosynthesis genes. Appl. Environ. Microbiol. 65: 3880– 3887 [PMC free article] [PubMed]
46. Qi F, Chen P, Caufield PW. 2001. The group I strain of Streptococcus mutans, UA140, produces both the lantibiotic mutacin I and a nonlantibiotic bacteriocin, mutacin IV. Appl. Environ. Microbiol. 67: 15– 21 [PMC free article] [PubMed]
47. Rodriguez-Ortega MJ, et al. 2006. Characterization and identification of vaccine candidate proteins through analysis of the group A Streptococcus surface proteome. Nat. Biotechnol. 24: 191– 197 [PubMed]
48. Sturme MH, et al. 2002. Cell to cell communication by autoinducing peptides in gram-positive bacteria. Antonie Van Leeuwenhoek 81: 233– 243 [PubMed]
49. Syvitski RT, et al. 2007. Structure-activity analysis of quorum-sensing signaling peptides from Streptococcus mutans. J. Bacteriol. 189: 1441– 1450 [PMC free article] [PubMed]
50. Tian X, et al. 2009. A method for structure-activity analysis of quorum-sensing signaling peptides from naturally transformable streptococci. Biol. Proced. Online 11: 207– 226 [PMC free article] [PubMed]
51. van der Ploeg JR. 2005. Regulation of bacteriocin production in Streptococcus mutans by the quorum-sensing system required for development of genetic competence. J. Bacteriol. 187: 3980– 3989 [PMC free article] [PubMed]
52. Van Melderen L, Aertsen A. 2009. Regulation and quality control by Lon-dependent proteolysis. Res. Microbiol. 160: 645– 651 [PubMed]
53. Wang BY, Kuramitsu HK. 2005. Interactions between oral bacteria: inhibition of Streptococcus mutans bacteriocin production by Streptococcus gordonii. Appl. Environ. Microbiol. 71: 354– 362 [PMC free article] [PubMed]
54. Waters CM, Bassler BL. 2005. Quorum sensing: cell-to-cell communication in bacteria. Annu. Rev. Cell Dev. Biol. 21: 319– 346 [PubMed]
55. Winzer K, Williams P. 2001. Quorum sensing and the regulation of virulence gene expression in pathogenic bacteria. Int. J. Med. Microbiol. 291: 131– 143 [PubMed]
56. Xie Z, Okinaga T, Niu G, Qi F, Merritt J. 2010. Identification of a novel bacteriocin regulatory system in Streptococcus mutans. Mol. Microbiol. 78: 1431– 1447 [PMC free article] [PubMed]
57. Yonezawa H, Kuramitsu HK. 2005. Genetic analysis of a unique bacteriocin, Smb, produced by Streptococcus mutans GS5. Antimicrob. Agents Chemother. 49: 541– 548 [PMC free article] [PubMed]
58. Yoshida A, Kuramitsu HK. 2002. Multiple Streptococcus mutans genes are involved in biofilm formation. Appl. Environ. Microbiol. 68: 6283– 6291 [PMC free article] [PubMed]
59. Zhang J, Biswas I. 2009. 3′-Phosphoadenosine-5′-phosphate phosphatase activity is required for superoxide stress tolerance in Streptococcus mutans. J. Bacteriol. 191: 4330– 4340 [PMC free article] [PubMed]

Articles from Journal of Bacteriology are provided here courtesy of American Society for Microbiology (ASM)