A Pleiotropic Regulator, Frp, Affects Exopolysaccharide Synthesis, Bio
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《感染与免疫杂志》
Department of Oral Biology, State University of New York, Buffalo, New York 14214
ABSTRACT
Exopolysaccharide synthesis, biofilm formation, and competence are important physiologic functions and virulence factors for Streptococcus mutans. In this study, we report the role of Frp, a transcriptional regulator, on the regulation of these traits crucial to pathogenesis. An Frp-deficient mutant showed decreased transcription of several genes important in virulence, including those encoding fructosyltransferase (Ftf), glucosyltransferase B (GtfB), and GtfC, by reverse transcription and quantitative real-time PCR. Expression of Ftf was decreased in the frp mutant, as assessed by Western blotting as well as by the activity assays. Frp deficiency also inhibited the production of GtfB in the presence of glucose and sucrose as well as the production of GtfC in the presence of glucose. As a consequence of the effects on GtfB and -C, sucrose-induced biofilm formation was decreased in the frp mutant. The expression of competence mediated by the competence-signaling peptide (CSP) system, as assessed by comC gene transcription, was attenuated in the frp mutant. As a result, the transformation efficiency was decreased in the frp mutant but was partially restored by adding synthetic CSP. Transcription of the frp gene was significantly increased in the frp mutant under all conditions tested, indicating that frp transcription is autoregulated. Furthermore, complementation of the frp gene in the frp mutant restored transcription of the affected genes to levels similar to those in the wild-type strain. These results suggest that Frp is a novel pleiotropic effector of multiple cellular functions and is involved in the modulation of exopolysaccharide synthesis, sucrose-dependent biofilm formation, and competence development.
INTRODUCTION
Streptococcus mutans is a primary etiologic agent of tooth decay, one of the most prevalent human infectious diseases. The abilities to adhere to teeth and form biofilms on the tooth surface as well as to metabolize carbohydrates and survive under differential environmental stress are critical to the cariogenicity of this microorganism (5, 14, 15, 25). It is known that S. mutans exhibits a high capacity to utilize dietary sucrose to produce extracellular polysaccharides such as fructans and glucans, which play important roles in attachment and adhesion to the tooth surface and serve as a nutrient reservoir.
In the unique oral environment, S. mutans exists primarily on the tooth surfaces. As a colonizer of the enamel surfaces, S. mutans, along with other dental microorganisms, forms a high-cell-density biofilm, dental plaque. The structure and composition of the plaque are strongly influenced by factors such as the source and availability of nutrients, the ability of bacteria to adapt to fluctuations in environmental conditions, and interactions with other plaque organisms (24). S. mutans mutants deficient in synthesis, catabolism, and binding of extracellular polysaccharides exhibit decreased cariogenicity and altered biofilm-forming capacities (2, 21, 46).
Production of extracellular polysaccharides from dietary carbon sources via glucosyltransferases (Gtfs) (4) and fructosyltransferase (Ftf) (1) contributes to the primary virulence of S. mutans. The adhesive glucans formed by the Gtfs, especially the water-insoluble glucans synthesized by the GtfB and -C enzymes, are significant constituents of dental plaque biofilms which facilitate adherence and accumulation of stable biofilms (13, 30, 37). Ftf synthesizes fructans from extracellular sucrose; these fructans are used primarily as a carbohydrate reservoir (5, 14), and it has been suggested that Ftf and fructans may also promote bacterial adhesion (26). GtfB and -C are encoded by the gtfB and gtfC genes, respectively; these genes are arranged in tandem within the same operon and can be transcribed from a common promoter located 5' of the gtfB gene. In addition, the gtfC gene also has its own promoter (12). The ftf gene is transcribed from its own promoter, which has two inverted repeat sequences (16, 33). The expression of these extracellular sugar metabolism enzymes is subject to change depending upon the availability of carbon sources (12, 26, 35, 45).
Bacterial communication via quorum-sensing signaling systems has been shown to be important in biofilm formation and competence development in streptococci, including S. mutans. The comC gene product, i.e., competence-signaling peptide (CSP) (17), and a two-component signal transduction system appear to play an important role in facilitating the natural transformation of S. mutans bacteria as well as in the formation of biofilms by and the aciduricity of the organisms (6). This quorum-sensing signaling system is encoded by the comCDE genes, corresponding to the CSP precursor, a histidine kinase (receptor for CSP), and a response regulator, respectively (20). Bacteria adapt to environmental changes by modulating the expression levels of the genes involved.
While initially characterizing the ftf gene, Sato and Kuramitsu discovered that there was an open reading frame (ORF3) downstream from the ftf gene with its own promoter transcribed in the opposite orientation to that of the ftf gene (29). Shibata and Kuramitsu (33) more recently reported that ORF3, named frp, encodes a putative protein homologous to DNA binding proteins and likely functions as a transcriptional regulator. By using DNA mobility shift assays, it has been shown that Frp binds to a DNA sequence in the ftf promoter region containing two inverted repeats located upstream of –10 and –35 sequences of the ftf gene (33). It was presumed that the function of Frp is to regulate the transcription of ftf, but this was not directly demonstrated. This communication, therefore, describes the function of Frp in extracellular polysaccharide metabolism, biofilm formation, and genetic competence.
MATERIALS AND METHODS
Bacterial strains and plasmids. The S. mutans strains used in this study were GS5 (9); GS5comC (made by transformation of GS5 with S. mutans NG8comC) (17); LN62, a gtfB mutant (22); and SP2, a spontaneous gtfBC mutant (11). Escherichia coli DH5 [80dlacZM15 recA1 endA1 gyrA96 thi-1 hsdR17 supE44 relA1 deoR (lacZYA-argF)] was utilized as a general cloning host. S. mutans strains were grown in Todd-Hewitt broth (THB; Invitrogen, Carlsbad, CA) and on tryptic soy broth agar (TSA) plates (Difco, Detroit, Mich.), as well as in chemically defined medium (CDM; 0.2% L-glutamic acid, 0.02% L-cysteine, 0.09% L-leucine, 0.1% NH4Cl, 0.25% K2HPO4, 0.25% KH2PO4, 0.4% NaHCO3, 0.12% MgSO4 · 7H2O, 0.002% MnCl2 · 4H2O, 0.002% FeSO4 · 7H2O, 0.06% Na-pyruvate, 0.0001% riboflavin, 0.00005% Ca-pantothenate, 0.0001% nicotinic acid, 0.00001% p-aminobenzoic acid, 0.00005% thiamine-HCl, 0.00001% biotin, 0.0001% pyridoxal-HCl, 0.00001% folic acid) with different carbon sources. E. coli strains were cultured in L broth (Invitrogen), and transformants were selected on L agar plates supplemented with the indicated antibiotics. Plasmids used in this study were pUC119Em (unpublished data), in which an erythromycin resistance gene (34) was cloned into the BamHI site of pUC119 (39), and pRKF (unpublished data), which contains a HindIII fragment of frp and partial upstream sequences.
DNA and RNA manipulations. DNA and RNA isolation, restriction endonuclease digestion, PCR, Southern blotting, ligation, transformation, and other DNA manipulations were carried out as described previously (40). Restriction endonucleases and other DNA-modifying enzymes were obtained from Invitrogen, New England Biolabs, Inc. (Beverly, MA), and Promega Corp. (Madison, WI) and used according to the specifications of the suppliers.
Construction of an S. mutans frp mutant. PvuII/NarI-digested pRKF was treated with the Klenow fragment, self-ligated, and transformed into E. coli. The resultant plasmid, pDfrp, contained a new BamHI site with the deletion of a 447-bp frp fragment 64 bp downstream from the start codon. The BamHI-digested erythromycin resistance gene from pUC119Em was introduced into the BamHI site of pDfrp, and this process yielded the recombinant plasmid pDfrpEm. The linearized pDfrpEm was transformed into S. mutans GS5 by natural transformation (40). The erythromycin-resistant transformants were selected on TSA plates with 5 μg/ml erythromycin. The frp deletion mutation was confirmed by Southern blotting and PCR (data not shown).
Reverse transcription and quantitative real-time PCR. Total RNA was treated with DNase I (Promega) at 37°C for 60 min to remove contaminant DNA from the RNA sample and then reverse transcribed with Superscript II (Invitrogen) at 37°C for 60 min using specific primers (Table 1) for each target sequence according to the supplier's instructions. Quantitative real-time PCR was performed using the iCycler iQ real-time PCR detection system (Bio-Rad, Hercules, CA). The reaction solution, 25 μl, consisted of a pair of specific primers for each target sequence (5 μM), 2.5 μl of cDNA, and 12.5 μl of iQ SYBR green Supermix (Bio-Rad). The conditions for quantitative real-time PCR were as follows: preheating at 95°C for 2 min followed by 40 amplification cycles of 95°C for 30 s, 50 to 55°C for 30 s, and 72°C for 30 s. Internal controls were used for each sample by detecting groEL mRNA, a host-keeping gene in S. mutans. Controls for each primer pair and DNase I-treated RNA sample without reverse transcription were included as negative controls. Standard curves were generated using 10-fold serial dilutions of the RNA standards. All samples were analyzed in triplicate. The mean cDNA copy numbers obtained for each gene were divided by the internal control values to standardize for the mRNAs present in each sample (8). Relative comparisons between corrected values were performed using the analysis of Student's t test.
Preparation of S. mutans extracellular protein and cellular fractions. Overnight 25-ml cultures of S. mutans cells in CDM with 1% sugar (sucrose, glucose, or fructose) were centrifuged (5,000 x g, 15 min at 4°C). The supernatants were precipitated with 75% saturated (NH4)2SO4 with stirring at 4°C for 2 h and then centrifuged at 9,000 x g at 4°C. The pellet was dissolved in 2.5 ml of 0.1 M potassium phosphate buffer (pH 6.0). The solution was then dialyzed in the same buffer overnight at 4°C. The protein concentration was measured by the method of Bradford, and the preparations were stored at –80°C.
Detection of fructosyltransferase. Ftf activity was measured as described previously with minor modifications (29). Briefly, 50 μl of protein was added to 50 μl of reaction solution (30 μl H2O, 10 μl of 1 M potassium phosphate buffer, 10 μl of 25% sucrose containing 25 mCi/ml [3H]fructose [sucrose supplied by NENTM Life Science Products, Inc., Boston, MA]) and incubated at 37°C for 1 h, and 1.0 ml of methanol was added into each reaction solution, followed by storage at –20°C for 20 min. The sample was then filtered through glass microfiber filters (Whatman GF/A). The filter membranes were washed, dried, and counted in a liquid scintillation counter (29). Ftf protein levels were detected by Western blotting using anti-Ftf antibody (47).
Detection of glucosyltransferase. Gtf activity was measured as described previously with slight modifications (31). Briefly, 50 μl of protein was added to 50 μl of reaction solution (10 μl H2O, 10 μl of 1 M Na acetate buffer, pH 6.0, 10 μl dextran T10 [1 mg/ml], 20 μl of 0.1% sucrose containing 0.02 μCi/ml [U-14C]glucose [sucrose supplied by NENTM Life Science Products, Inc., Boston, MA]) and incubated at 37°C for 1 h, and 1.0 ml of methanol was added to each reaction solution; the solutions were then placed at –20°C for 20 min. The sample was then filtered through glass microfiber filters (Whatman GF/A). The filter membrane was washed, dried, and counted. The Gtf protein levels were detected by Western blotting using anti-GtfBC antibody (47).
Biofilm formation assay. Biofilm formation was quantified as previously described (41). Flat-bottom polystyrene microtiter plates (96-well Easy Wash enzyme immunoassay-radioimmunoassay plates; Corning Inc., Corning, N.Y.) containing 100 μl of quarter-strength THB-mucin-0.5% sucrose per well were inoculated with S. mutans (1.7 x 105 CFU per well) from a 24-h growth in THB. After 24 h of incubation at 37°C, 25 μl of 1% (wt/vol) crystal violet solution was added to each well. After 15 min, the wells were rinsed three times with 200 μl of distilled water and dried in air. The crystal violet on the abiotic surfaces was solubilized in 95% ethanol, and the optical density at 600 nm was measured. Growth was determined by measuring the turbidities (optical densities at 600 nm) of parallel wells following resuspension of the sessile organisms together with the planktonic cells.
Determination of the transformation efficiency of S. mutans. Bacteria were cultured in THB-10% horse serum overnight at 37°C and then inoculated (1/10) into fresh THB-10% horse serum. Bacteria were incubated for 2 h, and 10 μg/100 μl of chromosomal DNA from the GS5::gtfD mutant (tetracycline resistant) (10) was added. The cells were incubated at 37°C anaerobically for 48 h. The transformation efficiency was determined as the number of transformants divided by the total cells transformed.
Complementation of the frp gene. A 1.0-kb DNA fragment containing the frp coding sequence, as well as upstream and downstream sequences amplified by a pair of primers (frp9, 5'-CGTCTATTTAAAATAATAGGC-3'; frp10, 5'-GTTTAGATCTTTTTGTCTAAC-3'), was cloned into the PvuII site of plasmid pResEm, and the resulting plasmid was transformed into S. mutans strains GS5frpSP and GS5, yielding frp single-crossover complemented strain Cfrp and control wild-type GS5frpEM. The correct DNA integration was confirmed by Southern blotting (data not shown). The S. mutans strains were grown in 1% glucose containing THB, and RNA was extracted and subjected to reverse transcription and quantitative real-time PCR.
RESULTS
Frp positively regulates expression of fructosyltransferase. The frp gene is located immediately downstream of the ftf gene and is transcribed in the opposite orientation (33). Our previous report on the DNA mobility shift assays showed that there are two DNA sequences that Frp binds specifically (33). In order to determine if Frp has any effects on ftf gene expression, an S. mutans GS5 frp-deficient mutant was constructed as described in Materials and Methods. Reverse transcription and quantitative real-time PCR showed that ftf transcription in the frp mutant was significantly decreased from the level seen for GS5 in CDM containing 1% sucrose, glucose, or fructose (Fig. 1A). Transcription of ftf in the frp mutant was only about 10% of that for the wild-type strain in the presence of 1% glucose, compared to frp mutant values of about 30% and 70% of the values for the wild-type strain in CDM containing fructose and CDM containing sucrose, respectively.
The Ftf activity assays were also carried out to determine whether or not the Ftf activity is also altered in the frp mutant (Fig. 1B). Culture supernatant fluids were used as the source of the enzyme, since earlier results showed that the bulk of the activity was present in this fraction (29). The results showed that Ftf activity was affected significantly by the presence of different carbon sources in the medium in both GS5 and the frp mutant. When S. mutans was cultured in the presence of sucrose, the observed activity was significantly decreased from that observed when cells were cultured in the presence of other sugars. To determine if the diminished Ftf activity was due to increased fructanase activity, fructanase activities were determined in the frp mutant and the wild-type strain in the presence of sucrose, and the results showed no significant differences between the two strains in the presence of sucrose (data not shown). The strains grown in the presence of glucose expressed higher levels of Ftf activity than those grown in the presence of fructose. There was a significant decrease in the Ftf activity for the frp mutant compared to the wild-type strain when these strains were grown in the presence of the monosaccharides (P < 0.05).
The protein levels of Ftf were assessed by Western blotting using anti-Ftf serum (Fig. 1C). The results showed that the carbon sources had a significant influence on the expression of the ftf gene in both the wild-type GS5 strain and the frp mutant. Protein levels of Ftf were highest in glucose-containing medium, in parallel with the activity results. There was no detectable Ftf protein in supernatant fluids of cultures of either the frp mutant or the wild-type strain grown in the presence of sucrose, which reflected the low Ftf activity detected in the culture fluids. These results indicated that frp was required for optimal transcription and expression of ftf gene.
Frp positively regulates expression of glucosyltransferase B and C in the presence of glucose. Since S. mutans can utilize extracellular sucrose to produce the exopolysaccharide glucans by GtfB, -C, and -D, we examined the effects of the frp mutation on the transcription of the gtfB, -C, and -D genes by reverse transcription and quantitative real-time PCR (Fig. 2A and B). The transcriptions of both gtfB and -C were decreased in the frp mutant compared to those of the wild-type strain in the presence of glucose. However, in fructose medium, the transcriptions of gtfB and -C in the frp mutant were similar to those in the wild-type strain. Transcription of gtfB in the frp mutant was significantly decreased in the presence of sucrose compared to that in the wild-type strain. In the presence of sucrose, in contrast, the transcription of gtfC in the frp mutant was similar to that in the wild-type strain. The transcription of gtfD was not significantly altered in the frp mutant compared to that in the wild-type strain under these conditions (data not shown).
The Gtf activity assays were also carried out to examine whether the total Gtf activity is altered in the frp-deficient mutant (Fig. 2C). The results showed that Gtf activity in both GS5 and the frp mutant was affected significantly by the presence of different carbon sources in the medium. When S. mutans was cultured in the presence of sucrose, the Gtf activity level was decreased significantly from that seen with culturing in the presence of glucose. Comparing the frp mutant and the wild-type strain, there was an apparent decrease of Gtf activity in the frp mutant relative to the wild-type strain, as suggested also by the transcription analysis.
The expression of the gtfB and -C genes was also examined by Western blotting using anti-GtfBC (Fig. 2D). The results consistently showed decreased protein expression of both GtfB and -C in the frp mutant compared to the wild-type strain in the presence of glucose. In contrast, only gtfB showed decreased expression in the frp mutant compared to that in the wild-type strain in the presence of 1% sucrose. There was also no difference between the frp mutant and the wild-type strain in the expression of the GtfB and -C proteins in the presence of fructose. Therefore, the frp mutation either decreased or had little effect on the expression of gtf genes, depending upon the sugar source for growth.
The frp mutation inhibits sucrose-dependent biofilm formation by S. mutans. In order to evaluate the effects of the frp mutation on biofilm formation by S. mutans, the biofilm formation assay was carried out as described earlier in the presence of different sugars (Fig. 3). The results showed that the frp mutant formed an amount of biofilm less than that formed by the wild-type strain only in the presence of sucrose (P < 0.05). The gtfB mutant, LN62, also showed attenuated biofilm formation compared to the wild-type strain (P < 0.05) as previously described (37). Levels of biofilm formation were similar for the frp mutant, the gtfB mutant, and the wild-type strain in the presence of glucose or fructose. These results indicated that the influence of Frp on sucrose-dependent biofilm formation may be due primarily to its effects on gtfB gene expression.
Frp is involved in quorum sensing via regulation of the expression of the comC gene. By searching the S. mutans genome database (University of Oklahoma), there was one sequence identified in the comC gene promoter region that has approximately 61.3% identity with the inverted repeat sequence in the ftf gene promoter that binds Frp specifically. Reverse transcription and quantitative real-time PCR results showed that transcription of comC was decreased in the frp mutant compared to that in the wild-type strain in THB medium and sucrose-containing media (Fig. 4). However, there was no significant difference between the frp mutant and the wild-type strain in the presence of glucose (Fig. 4). Since the comC product is involved in the capacity of natural transformation (18), the transformation efficiencies of the frp mutant, wild-type GS5, and a comC-deficient mutant with and without CSP were compared (Table 2). Sucrose was used in these assays, since recent results have indicated that transformation in S. mutans is optimal in the presence of this sugar (B.-Y. Wang and H. K. Kuramitsu, unpublished results). The results showed that the transformation efficiency of the frp mutant was about 27-fold lower than that of the wild-type strain, while that of the comC-deficient mutant was about 100-fold lower than that of the wild-type strain in the absence of CSP. In contrast, the addition of 1.0 μg/ml of synthetic CSP increased the transformation efficiencies by about 21-, 29-, and 6.5-fold in the frp mutant, the comC mutant, and the wild-type strain, respectively, over those without CSP. These results indicated that the frp gene also influences transformation by modulating the expression of CSP in S. mutans.
Autoregulation of Frp. Although our results indicated that Frp acts as a positive regulator in the control of the expression of genes involved in pleiotropic cellular functions, it was also demonstrated that Frp acted as a transcriptional regulator to control its own expression (Fig. 5). Instead of being a positive regulator, Frp is a negative regulator of its own transcription regardless of the carbon source used for growth of the cells. This indicated that frp expression is autoregulated and that this regulation is independent of the sugar source.
Complementation of frp gene. In order to verify that the phenotype of the frp mutant is not the result of polar effects in the construction of the frp mutant, we introduced an intact frp gene into the chromosome of the S. mutans frp mutant. Reverse transcription and quantitative real-time PCR (Fig. 6) showed that, in contrast to the frp mutant, the frp-complemented strain restored the transcription of the ftf, frp, gtfB, and gtfC genes to wild-type levels in the presence of 1% glucose. These results suggested that Frp either directly or indirectly modulates the expression of these genes.
DISCUSSION
Oral bacteria are subject to radical periodic fluctuations in the mouth environment. Due to intermittent eating patterns, vast differences in saliva quantity and composition associated with diurnal rhythms, mechanical forces, and host defense mechanisms, bacteria in plaque are constantly subjected to environmental fluctuations. As a major pathogen involved in dental caries, the ability of S. mutans to optimize its expression of a variety of virulence factors, including extracellular polysaccharide synthesis systems, biofilm formation, and cell-cell signaling, should be essential for the pathogenicity of this bacterium. Since the adaptation to the environment is a complex phenomenon involving coordination of a variety of cellular functions, it is reasonable that bacteria contain a variety of mechanisms to coordinate biochemical changes. Previous studies have indicated several mechanisms for such adaptation in S. mutans. For example, the trigger factor is associated with stress tolerance, competence development, and biofilm formation (44). LuxS-based signaling also affects S. mutans biofilm formation (43, 49). Furthermore, an E11ABman mutation increases the production of Ftf, Gtf, and other gene products (1). Likewise, an HtrA surface protease-deficient mutant is altered in terms of expression of Gtf and Ftf as well as of GbpB, enolase, and glyceraldehyde-3-phosphate dehydrogenase. This mutant also displayed a granular patchy appearance rather than the relatively smooth confluent layer normally seen for the wild type in biofilms (3). Furthermore, the VicRK signal transduction system in S. mutans affects gtfBCD, gbpB, and ftf expression, biofilm formation, and genetic competence development (32). These findings indicate that complex regulatory mechanisms operate in S. mutans. Our results demonstrating that Frp functions as a positive transcriptional regulator involved in exopolysaccharide synthesis and also affects sucrose-dependent biofilm formation and competence development suggest the existence of another novel pleiotropic regulatory system in S. mutans. It is possible that Frp might interact with one or more of the previously mentioned regulatory systems.
Extracellular sugar metabolism has been of interest because of its association with dental caries pathogenesis as a virulence factor of S. mutans. Although ftf and gtfs have distinct genetic organizations and different promoters on the chromosome of S. mutans, they are induced by their common substrate, i.e., sucrose in the diet, and are subject to fluctuation in the presence of different sugars (12, 27, 35, 45). Our results that frp deficiency in S. mutans significantly decreases the transcription of ftf in the presence of sucrose, glucose, and fructose suggest that Frp functions as a positive transcriptional regulator of ftf expression. In addition, the apparent dramatic decrease in Ftf activity as well as in Ftf protein levels in both the wild-type strain and the frp mutant in the culture fluids in the presence of sucrose is likely influenced by the relocation of Ftf to a cell-associated form as a result of insoluble glucan synthesis (29).
It is interesting that the effects of Frp on expression of both gtfB and gtfC genes are somewhat distinct from the effect on ftf expression. In general, Frp appears to function as a positive transcriptional regulator of ftf expression as well as gtfB and gtfC expression, especially when S. mutans is grown in presence of glucose. However, in the presence of sucrose, gtfC expression is primarily Frp independent, in contrast to gtfB and ftf expression. Furthermore, fructose appears to affect ftf expression in the frp mutant much more strongly than it affects the expression of gtfB and gtfC. These results are compatible with the hypothesis that there are multiple regulatory mechanisms involved in the expression of exopolysaccharide-synthetic genes. It will be of interest to determine if the Frp regulatory system interacts with these other systems either directly or indirectly. The difference in sugar effects on gtfB, gtfC, and ftf indicates that different mechanisms may also exist in the regulation of transcription of these genes by Frp, since there is no inverted repeat in the promoter region of the gtfBC gene comparable to that in the ftf promoter region which binds Frp specifically.
Sugar metabolism pathways in S. mutans are also subject to catabolic repression when glucose is present (28, 42). The result that glucose, of all the sugars tested, causes the most significant repression in the frp mutant of the exopolysaccharide synthesis enzymes and of gtfB, gtfC, and ftf relative to the levels of these enzymes and genes found for the wild-type strain suggests that Frp may function, in part, as an antagonist of catabolic repression. The Frp defect may therefore enhance catabolic repression in the presence of glucose. Although other sugars besides glucose may produce intermediates involved in catabolite repression, the present results suggest that glucose is more efficiently metabolized to such intermediates in S. mutans. The results showing that transcription of the comC gene, which is not known to be subject to catabolic repression by glucose, are consistent with this possibility and need to be investigated in detail.
Biofilm formation by S. mutans has been shown to be dependent upon several factors interacting with the oral environment. The expression levels of the ftf, gtfB, and gtfC genes have been shown to increase in biofilm cells (16, 19). Among the different sugars, fructose and glucose have ftf expression effects that are enhanced compared to that of sucrose (26), while both gtfB and gtfC are associated with biofilm formation in the presence of sucrose (38). Fructans produced by Ftf contribute to the virulence of the biofilm by acting as potential binding sites for S. mutans adhesion and also as an extracellular nutrient reservoir for plaque bacteria. In contrast, glucans produced by GtfB and GtfC play major roles in sucrose-dependent biofilm formation by S. mutans (36, 38). GtfC in particular plays a crucial role in sucrose-dependent adhesion and is essential for biofilm formation on smooth surfaces. Thus, the defect in biofilm formation displayed by the frp mutant in the presence of sucrose appears to be primarily due to reduction in GtfB production plus a ftf deficiency in this strain (23). Since Ftf activity is decreased in the frp mutant, the decrease in GtfB activity may result from interference with the normal regulatory effects of Ftf on gtfB expression. Interestingly, the frp mutant is not repressed in comC expression in nonsucrose media and therefore is not defective in sucrose-independent biofilm formation. A recent study indicated that a comC mutant of S. mutans GS5 was defective in this property (50).
Competence development in S. mutans is associated with the quorum-sensing signaling system of this organism (7, 17, 18, 23). The frp mutant was shown to display decreased transcription of the comC gene compared to the wild-type strain in the presence of sucrose. Accordingly, the transformation efficiency of the frp mutant is significantly reduced compared to that of the wild-type strain. Furthermore, the addition of CSP to the cells partially restored transformation efficiency in both the frp mutant and the comC mutant; this suggests a role for Frp in competence development and transformation efficiency. It is not clear why the addition of exogenous CSP to the comC mutant did not allow for full complementation of transformation to the wild-type levels. Such incomplete complementation was also observed for other phenotypic traits of the mutant following exposure to synthetic CSP (data not shown). Other virulence factors regulated by competence (48) may also be attenuated in the frp mutant, and this will be examined in future experiments.
Overall, our results with the frp mutant indicate that Frp has pleiotropic effects on virulence-related cellular functions. Frp appears to be an important regulator of exopolysaccharide synthesis, sucrose-dependent biofilm formation, and competence development. Further investigation of how Frp affects all of these activities at the molecular level will be necessary to evaluate the direct role of Frp in the coordination of these physiological functions. In addition, it will be of interest to determine which environmental cues are used by Frp to control these interactions.
ACKNOWLEDGMENTS
This investigation was supported in part by NIH grant DE03258.
FOOTNOTES
Corresponding author. Mailing address: Department of Oral Biology, State University of New York at Buffalo, 3435 Main Street, Buffalo, NY 14214. Phone: (512) 249-5901. Fax: (716) 829-3942. E-mail: kuramitsu@earthlink.net.
REFERENCES
1. Abranches, J., Y. Y. Chen, and R. A. Burne. 2003. Characterization of Streptococcus mutans strains deficient in EIIABMan of the sugar phosphotransferase system. Appl. Environ. Microbiol. 69:4760-4769.
2. Banas, J. A., and M. M. Vickerman. 2003. Glucan-binding proteins of the oral streptococci. Crit. Rev. Oral Biol. Med. 14:89-99.
3. Biswas, S., and I. Biswas. 2005. Role of HtrA in surface protein expression and biofilm formation by Streptococcus mutans. Infect. Immun. 73:6923-6934.
4. Browngardt, C. M., Z. T. Wen, and R. A. Burne. 2004. RegM is required for optimal fructosyltransferase and glucosyltransferase gene expression in Streptococcus mutans. FEMS Microbiol. Lett. 240:75-79.
5. Burne, R. A. 1998. Oral streptococci... products of their environment. J. Dent. Res. 77:445-452.
6. Cvitkovitch, D. G., Y. H. Li, and R. P. Ellen. 2003. Quorum sensing and biofilm formation in streptococcal infections. J. Clin. Investig. 112:1626-1632.
7. Echenique, J. R., S. Chapuy-Regaud, and M. C. Trombe. 2000. Competence regulation by oxygen in Streptococcus pneumoniae: involvement of ciaRH and comCDE. Mol. Microbiol. 36:688-696.
8. Fronhoffs, S., G. Totzke, S. Stier, N. Wernert, M. Rothe, T. Bruning, B. Koch, A. Sachinidis, H. Vetter, and Y. Ko. 2002. A method for the rapid construction of cRNA standard curves in quantitative real-time reverse transcription polymerase chain reaction. Mol. Cell. Probes 16:99-110.
9. Gibbons, R. J., K. S. Berman, P. Knoettner, and B. Kapsimalis. 1966. Dental caries and alveolar bone loss in gnotobiotic rats infected with capsule forming streptococci of human origin. Arch. Oral Biol. 11:549-560.
10. Hanada, N., and H. K. Kuramitsu. 1989. Isolation and characterization of the Streptococcus mutans gtfD gene, coding for primer-dependent soluble glucan synthesis. Infect. Immun. 57:2079-2085.
11. Hiratsuka, K., B. Wang, Y. Sato, and H. Kuramitsu. 1998. Regulation of sucrose-6-phosphate hydrolase activity in Streptococcus mutans: characterization of the scrR gene. Infect. Immun. 66:3736-3743.
12. Kiska, D. L., and F. L. Macrina. 1994. Genetic regulation of fructosyltransferase in Streptococcus mutans. Infect. Immun. 62:1241-1251.
13. Kopec, L. K., A. M. Vacca-Smith, and W. H. Bowen. 1997. Structural aspects of glucans formed in solution and on the surface of hydroxyapatite. Glycobiology 7:929-934.
14. Kuramitsu, H. K. 1993. Virulence factors of mutans streptococci: role of molecular genetics. Crit. Rev. Oral Biol. Med. 4:159-176.
15. Lemos, J. A., J. Abranches, and R. A. Burne. 2005. Responses of cariogenic streptococci to environmental stresses. Curr. Issues Mol. Biol. 7:95-107.
16. Li, Y., and R. A. Burne. 2001. Regulation of the gtfBC and ftf genes of Streptococcus mutans in biofilms in response to pH and carbohydrate. Microbiology 147:2841-2848.
17. Li, Y. H., P. C. Lau, J. H. Lee, R. P. Ellen, and D. G. Cvitkovitch. 2001. Natural genetic transformation of Streptococcus mutans growing in biofilms. J. Bacteriol. 183:897-908.
18. Li, Y. H., N. Tang, M. B. Aspiras, P. C. Lau, J. H. Lee, R. P. Ellen, and D. G. Cvitkovitch. 2002. A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. J. Bacteriol. 184:2699-2708.
19. Mattos-Graner, R. O., M. H. Napimoga, K. Fukushima, M. J. Duncan, and D. J. Smith. 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.
20. Morrison, D. A. 1997. Streptococcal competence for genetic transformation: regulation by peptide pheromones. Microb. Drug Resist. 3:27-37.
21. Munro, C., S. M. Michalek, and F. L. Macrina. 1991. Cariogenicity of Streptococcus mutans V403 glucosyltransferase and fructosyltransferase mutants constructed by allelic exchange. Infect. Immun. 59:2316-2323.
22. Perry, D., and H. K. Kuramitsu. 1989. Genetic linkage among cloned genes of Streptococcus mutans. Infect. Immun. 57:805-809.
23. Petersen, F. C., L. Tao, and A. A. Scheie. 2005. DNA binding-uptake system: a link between cell-to-cell communication and biofilm formation. J. Bacteriol. 187:4392-4400.
24. Quivey, R. G., Jr., W. L. Kuhnert, and K. Hahn. 2000. Adaptation of oral streptococci to low pH. Adv. Microb. Physiol. 42:239-274.
25. Rathsam, C., R. E. Eaton, C. L. Simpson, G. V. Browne, T. Berg, D. W. Harty, and N. A. Jacques. 2005. Up-regulation of competence- but not stress-responsive proteins accompanies an altered metabolic phenotype in Streptococcus mutans biofilms. Microbiology 151:1823-1837.
26. Rozen, R., G. Bachrach, and D. Steinberg. 2004. Effect of carbohydrates on fructosyltransferase expression and distribution in Streptococcus mutans GS-5 biofilms. Carbohydr. Res. 339:2883-2888.
27. Rozen, R., D. Steinberg, and G. Bachrach. 2004. Streptococcus mutans fructosyltransferase interactions with glucans. FEMS Microbiol. Lett. 232:39-43.
28. Saier, M. H., Jr., S. Chauvaux, G. M. Cook, J. Deutscher, I. T. Paulsen, J. Reizer, and J. J. Ye. 1996. Catabolite repression and inducer control in Gram-positive bacteria. Microbiology 142:217-230.
29. Sato, S., and H. K. Kuramitsu. 1986. Isolation and characterization of a fructosyltransferase gene from Streptococcus mutans GS-5. Infect. Immun. 52:166-170.
30. Schilling, K. M., and W. H. Bowen. 1992. Glucans synthesized in situ in experimental salivary pellicle function as specific binding sites for Streptococcus mutans. Infect. Immun. 60:284-295.
31. Schroeder, V. A., S. M. Michalek, and F. L. Macrina. 1989. Biochemical characterization and evaluation of virulence of a fructosyltransferase-deficient mutant of Streptococcus mutans V403. Infect. Immun. 57:3560-3569.
32. Senadheera, M. D., B. Guggenheim, G. A. Spatafora, Y. C. Huang, J. Choi, D. C. Hung, J. S. Treglown, S. D. Goodman, R. P. Ellen, and D. G. Cvitkovitch. 2005. A VicRK signal transduction system in Streptococcus mutans affects gtfBCD, gbpB, and ftf expression, biofilm formation, and genetic competence development. J. Bacteriol. 187:4064-4076.
33. Shibata, Y., and H. K. Kuramitsu. 1996. Identification of the Streptococcus mutans frp gene as a potential regulator of fructosyltransferase expression. FEMS Microbiol. Lett. 140:49-54.
34. Shiroza, T., and H. K. Kuramitsu. 1995. Development of a heterodimer plasmid system for the introduction of heterologous genes into streptococci. Plasmid 34:85-95.
35. Steinberg, D., R. Rozen, M. Bromshteym, B. Zaks, I. Gedalia, and G. Bachrach. 2002. Regulation of fructosyltransferase activity by carbohydrates, in solution and immobilized on hydroxyapatite surfaces. Carbohydr. Res. 337:701-710.
36. Tamesada, M., S. Kawabata, T. Fujiwara, and S. Hamada. 2004. Synergistic effects of streptococcal glucosyltransferases on adhesive biofilm formation. J. Dent. Res. 83:874-879.
37. Tsumori, H., and H. Kuramitsu. 1997. The role of the Streptococcus mutans glucosyltransferases in the sucrose-dependent attachment to smooth surfaces: essential role of the GtfC enzyme. Oral Microbiol. Immunol. 12:274-280.
38. Van Der Ploeg, J. R., and B. Guggenheim. 2004. Deletion of gtfC of Streptococcus mutans has no influence on the composition of a mixed-species in vitro biofilm model of supragingival plaque. Eur. J. Oral Sci. 112:433-438.
39. Vieira, J., and J. Messing. 1987. Production of single-stranded plasmid DNA. Methods Enzymol. 153:3-11.
40. Wang, B., and H. K. Kuramitsu. 2003. Control of enzyme IIscr and sucrose-6-phosphate hydrolase activities in Streptococcus mutans by transcriptional repressor ScrR binding to the cis-active determinants of the scr regulon. J. Bacteriol. 185:5791-5799.
41. Wang, B., and H. K. Kuramitsu. 2005. Inducible antisense RNA expression in the characterization of gene functions in Streptococcus mutans. Infect. Immun. 73:3568-3576.
42. Wen, Z. T., and R. A. Burne. 2002. Analysis of cis- and trans-acting factors involved in regulation of the Streptococcus mutans fructanase gene (fruA). J. Bacteriol. 184:126-133.
43. Wen, Z. T., and R. A. Burne. 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.
44. Wen, Z. T., P. Suntharaligham, D. G. Cvitkovitch, and R. A. Burne. 2005. Trigger factor in Streptococcus mutans is involved in stress tolerance, competence development, and biofilm formation. Infect. Immun. 73:219-225.
45. Wexler, D. L., M. C. Hudson, and R. A. Burne. 1993. Streptococcus mutans fructosyltransferase (ftf) and glucosyltransferase (gtfBC) operon fusion strains in continuous culture. Infect. Immun. 61:1259-1267.
46. Yamashita, Y., W. H. Bowen, R. A. Burne, and H. K. Kuramitsu. 1993. Role of the Streptococcus mutans gtf genes in caries induction in the specific-pathogen-free rat model. Infect. Immun. 61:3811-3817.
47. Yamashita, Y., Y. Tsukioka, Y. Nakano, Y. Shibata, and T. Koga. 1996. Molecular and genetic analysis of multiple changes in the levels of production of virulence factors in a subcultured variant of Streptococcus mutans. FEMS Microbiol. Lett. 144:81-87.
48. Yonezawa, H., and H. K. Kuramitsu. 2005. Genetic analysis of a unique bacteriocin, Smb, produced by Streptococcus mutans GS5. Antimicrob. Agents Chemother. 49:541-548.
49. Yoshida, A., T. Ansai, T. Takehara, and H. K. Kuramitsu. 2005. LuxS-based signaling affects Streptococcus mutans biofilm formation. Appl. Environ. Microbiol. 71:2372-2380.
50. Yoshida, A., and H. K. Kuramitsu. 2002. Multiple Streptococcus mutans genes are involved in biofilm formation. Appl. Environ. Microbiol. 68:6283-6291.(Bing Wang and Howard K. Kuramitsu)
ABSTRACT
Exopolysaccharide synthesis, biofilm formation, and competence are important physiologic functions and virulence factors for Streptococcus mutans. In this study, we report the role of Frp, a transcriptional regulator, on the regulation of these traits crucial to pathogenesis. An Frp-deficient mutant showed decreased transcription of several genes important in virulence, including those encoding fructosyltransferase (Ftf), glucosyltransferase B (GtfB), and GtfC, by reverse transcription and quantitative real-time PCR. Expression of Ftf was decreased in the frp mutant, as assessed by Western blotting as well as by the activity assays. Frp deficiency also inhibited the production of GtfB in the presence of glucose and sucrose as well as the production of GtfC in the presence of glucose. As a consequence of the effects on GtfB and -C, sucrose-induced biofilm formation was decreased in the frp mutant. The expression of competence mediated by the competence-signaling peptide (CSP) system, as assessed by comC gene transcription, was attenuated in the frp mutant. As a result, the transformation efficiency was decreased in the frp mutant but was partially restored by adding synthetic CSP. Transcription of the frp gene was significantly increased in the frp mutant under all conditions tested, indicating that frp transcription is autoregulated. Furthermore, complementation of the frp gene in the frp mutant restored transcription of the affected genes to levels similar to those in the wild-type strain. These results suggest that Frp is a novel pleiotropic effector of multiple cellular functions and is involved in the modulation of exopolysaccharide synthesis, sucrose-dependent biofilm formation, and competence development.
INTRODUCTION
Streptococcus mutans is a primary etiologic agent of tooth decay, one of the most prevalent human infectious diseases. The abilities to adhere to teeth and form biofilms on the tooth surface as well as to metabolize carbohydrates and survive under differential environmental stress are critical to the cariogenicity of this microorganism (5, 14, 15, 25). It is known that S. mutans exhibits a high capacity to utilize dietary sucrose to produce extracellular polysaccharides such as fructans and glucans, which play important roles in attachment and adhesion to the tooth surface and serve as a nutrient reservoir.
In the unique oral environment, S. mutans exists primarily on the tooth surfaces. As a colonizer of the enamel surfaces, S. mutans, along with other dental microorganisms, forms a high-cell-density biofilm, dental plaque. The structure and composition of the plaque are strongly influenced by factors such as the source and availability of nutrients, the ability of bacteria to adapt to fluctuations in environmental conditions, and interactions with other plaque organisms (24). S. mutans mutants deficient in synthesis, catabolism, and binding of extracellular polysaccharides exhibit decreased cariogenicity and altered biofilm-forming capacities (2, 21, 46).
Production of extracellular polysaccharides from dietary carbon sources via glucosyltransferases (Gtfs) (4) and fructosyltransferase (Ftf) (1) contributes to the primary virulence of S. mutans. The adhesive glucans formed by the Gtfs, especially the water-insoluble glucans synthesized by the GtfB and -C enzymes, are significant constituents of dental plaque biofilms which facilitate adherence and accumulation of stable biofilms (13, 30, 37). Ftf synthesizes fructans from extracellular sucrose; these fructans are used primarily as a carbohydrate reservoir (5, 14), and it has been suggested that Ftf and fructans may also promote bacterial adhesion (26). GtfB and -C are encoded by the gtfB and gtfC genes, respectively; these genes are arranged in tandem within the same operon and can be transcribed from a common promoter located 5' of the gtfB gene. In addition, the gtfC gene also has its own promoter (12). The ftf gene is transcribed from its own promoter, which has two inverted repeat sequences (16, 33). The expression of these extracellular sugar metabolism enzymes is subject to change depending upon the availability of carbon sources (12, 26, 35, 45).
Bacterial communication via quorum-sensing signaling systems has been shown to be important in biofilm formation and competence development in streptococci, including S. mutans. The comC gene product, i.e., competence-signaling peptide (CSP) (17), and a two-component signal transduction system appear to play an important role in facilitating the natural transformation of S. mutans bacteria as well as in the formation of biofilms by and the aciduricity of the organisms (6). This quorum-sensing signaling system is encoded by the comCDE genes, corresponding to the CSP precursor, a histidine kinase (receptor for CSP), and a response regulator, respectively (20). Bacteria adapt to environmental changes by modulating the expression levels of the genes involved.
While initially characterizing the ftf gene, Sato and Kuramitsu discovered that there was an open reading frame (ORF3) downstream from the ftf gene with its own promoter transcribed in the opposite orientation to that of the ftf gene (29). Shibata and Kuramitsu (33) more recently reported that ORF3, named frp, encodes a putative protein homologous to DNA binding proteins and likely functions as a transcriptional regulator. By using DNA mobility shift assays, it has been shown that Frp binds to a DNA sequence in the ftf promoter region containing two inverted repeats located upstream of –10 and –35 sequences of the ftf gene (33). It was presumed that the function of Frp is to regulate the transcription of ftf, but this was not directly demonstrated. This communication, therefore, describes the function of Frp in extracellular polysaccharide metabolism, biofilm formation, and genetic competence.
MATERIALS AND METHODS
Bacterial strains and plasmids. The S. mutans strains used in this study were GS5 (9); GS5comC (made by transformation of GS5 with S. mutans NG8comC) (17); LN62, a gtfB mutant (22); and SP2, a spontaneous gtfBC mutant (11). Escherichia coli DH5 [80dlacZM15 recA1 endA1 gyrA96 thi-1 hsdR17 supE44 relA1 deoR (lacZYA-argF)] was utilized as a general cloning host. S. mutans strains were grown in Todd-Hewitt broth (THB; Invitrogen, Carlsbad, CA) and on tryptic soy broth agar (TSA) plates (Difco, Detroit, Mich.), as well as in chemically defined medium (CDM; 0.2% L-glutamic acid, 0.02% L-cysteine, 0.09% L-leucine, 0.1% NH4Cl, 0.25% K2HPO4, 0.25% KH2PO4, 0.4% NaHCO3, 0.12% MgSO4 · 7H2O, 0.002% MnCl2 · 4H2O, 0.002% FeSO4 · 7H2O, 0.06% Na-pyruvate, 0.0001% riboflavin, 0.00005% Ca-pantothenate, 0.0001% nicotinic acid, 0.00001% p-aminobenzoic acid, 0.00005% thiamine-HCl, 0.00001% biotin, 0.0001% pyridoxal-HCl, 0.00001% folic acid) with different carbon sources. E. coli strains were cultured in L broth (Invitrogen), and transformants were selected on L agar plates supplemented with the indicated antibiotics. Plasmids used in this study were pUC119Em (unpublished data), in which an erythromycin resistance gene (34) was cloned into the BamHI site of pUC119 (39), and pRKF (unpublished data), which contains a HindIII fragment of frp and partial upstream sequences.
DNA and RNA manipulations. DNA and RNA isolation, restriction endonuclease digestion, PCR, Southern blotting, ligation, transformation, and other DNA manipulations were carried out as described previously (40). Restriction endonucleases and other DNA-modifying enzymes were obtained from Invitrogen, New England Biolabs, Inc. (Beverly, MA), and Promega Corp. (Madison, WI) and used according to the specifications of the suppliers.
Construction of an S. mutans frp mutant. PvuII/NarI-digested pRKF was treated with the Klenow fragment, self-ligated, and transformed into E. coli. The resultant plasmid, pDfrp, contained a new BamHI site with the deletion of a 447-bp frp fragment 64 bp downstream from the start codon. The BamHI-digested erythromycin resistance gene from pUC119Em was introduced into the BamHI site of pDfrp, and this process yielded the recombinant plasmid pDfrpEm. The linearized pDfrpEm was transformed into S. mutans GS5 by natural transformation (40). The erythromycin-resistant transformants were selected on TSA plates with 5 μg/ml erythromycin. The frp deletion mutation was confirmed by Southern blotting and PCR (data not shown).
Reverse transcription and quantitative real-time PCR. Total RNA was treated with DNase I (Promega) at 37°C for 60 min to remove contaminant DNA from the RNA sample and then reverse transcribed with Superscript II (Invitrogen) at 37°C for 60 min using specific primers (Table 1) for each target sequence according to the supplier's instructions. Quantitative real-time PCR was performed using the iCycler iQ real-time PCR detection system (Bio-Rad, Hercules, CA). The reaction solution, 25 μl, consisted of a pair of specific primers for each target sequence (5 μM), 2.5 μl of cDNA, and 12.5 μl of iQ SYBR green Supermix (Bio-Rad). The conditions for quantitative real-time PCR were as follows: preheating at 95°C for 2 min followed by 40 amplification cycles of 95°C for 30 s, 50 to 55°C for 30 s, and 72°C for 30 s. Internal controls were used for each sample by detecting groEL mRNA, a host-keeping gene in S. mutans. Controls for each primer pair and DNase I-treated RNA sample without reverse transcription were included as negative controls. Standard curves were generated using 10-fold serial dilutions of the RNA standards. All samples were analyzed in triplicate. The mean cDNA copy numbers obtained for each gene were divided by the internal control values to standardize for the mRNAs present in each sample (8). Relative comparisons between corrected values were performed using the analysis of Student's t test.
Preparation of S. mutans extracellular protein and cellular fractions. Overnight 25-ml cultures of S. mutans cells in CDM with 1% sugar (sucrose, glucose, or fructose) were centrifuged (5,000 x g, 15 min at 4°C). The supernatants were precipitated with 75% saturated (NH4)2SO4 with stirring at 4°C for 2 h and then centrifuged at 9,000 x g at 4°C. The pellet was dissolved in 2.5 ml of 0.1 M potassium phosphate buffer (pH 6.0). The solution was then dialyzed in the same buffer overnight at 4°C. The protein concentration was measured by the method of Bradford, and the preparations were stored at –80°C.
Detection of fructosyltransferase. Ftf activity was measured as described previously with minor modifications (29). Briefly, 50 μl of protein was added to 50 μl of reaction solution (30 μl H2O, 10 μl of 1 M potassium phosphate buffer, 10 μl of 25% sucrose containing 25 mCi/ml [3H]fructose [sucrose supplied by NENTM Life Science Products, Inc., Boston, MA]) and incubated at 37°C for 1 h, and 1.0 ml of methanol was added into each reaction solution, followed by storage at –20°C for 20 min. The sample was then filtered through glass microfiber filters (Whatman GF/A). The filter membranes were washed, dried, and counted in a liquid scintillation counter (29). Ftf protein levels were detected by Western blotting using anti-Ftf antibody (47).
Detection of glucosyltransferase. Gtf activity was measured as described previously with slight modifications (31). Briefly, 50 μl of protein was added to 50 μl of reaction solution (10 μl H2O, 10 μl of 1 M Na acetate buffer, pH 6.0, 10 μl dextran T10 [1 mg/ml], 20 μl of 0.1% sucrose containing 0.02 μCi/ml [U-14C]glucose [sucrose supplied by NENTM Life Science Products, Inc., Boston, MA]) and incubated at 37°C for 1 h, and 1.0 ml of methanol was added to each reaction solution; the solutions were then placed at –20°C for 20 min. The sample was then filtered through glass microfiber filters (Whatman GF/A). The filter membrane was washed, dried, and counted. The Gtf protein levels were detected by Western blotting using anti-GtfBC antibody (47).
Biofilm formation assay. Biofilm formation was quantified as previously described (41). Flat-bottom polystyrene microtiter plates (96-well Easy Wash enzyme immunoassay-radioimmunoassay plates; Corning Inc., Corning, N.Y.) containing 100 μl of quarter-strength THB-mucin-0.5% sucrose per well were inoculated with S. mutans (1.7 x 105 CFU per well) from a 24-h growth in THB. After 24 h of incubation at 37°C, 25 μl of 1% (wt/vol) crystal violet solution was added to each well. After 15 min, the wells were rinsed three times with 200 μl of distilled water and dried in air. The crystal violet on the abiotic surfaces was solubilized in 95% ethanol, and the optical density at 600 nm was measured. Growth was determined by measuring the turbidities (optical densities at 600 nm) of parallel wells following resuspension of the sessile organisms together with the planktonic cells.
Determination of the transformation efficiency of S. mutans. Bacteria were cultured in THB-10% horse serum overnight at 37°C and then inoculated (1/10) into fresh THB-10% horse serum. Bacteria were incubated for 2 h, and 10 μg/100 μl of chromosomal DNA from the GS5::gtfD mutant (tetracycline resistant) (10) was added. The cells were incubated at 37°C anaerobically for 48 h. The transformation efficiency was determined as the number of transformants divided by the total cells transformed.
Complementation of the frp gene. A 1.0-kb DNA fragment containing the frp coding sequence, as well as upstream and downstream sequences amplified by a pair of primers (frp9, 5'-CGTCTATTTAAAATAATAGGC-3'; frp10, 5'-GTTTAGATCTTTTTGTCTAAC-3'), was cloned into the PvuII site of plasmid pResEm, and the resulting plasmid was transformed into S. mutans strains GS5frpSP and GS5, yielding frp single-crossover complemented strain Cfrp and control wild-type GS5frpEM. The correct DNA integration was confirmed by Southern blotting (data not shown). The S. mutans strains were grown in 1% glucose containing THB, and RNA was extracted and subjected to reverse transcription and quantitative real-time PCR.
RESULTS
Frp positively regulates expression of fructosyltransferase. The frp gene is located immediately downstream of the ftf gene and is transcribed in the opposite orientation (33). Our previous report on the DNA mobility shift assays showed that there are two DNA sequences that Frp binds specifically (33). In order to determine if Frp has any effects on ftf gene expression, an S. mutans GS5 frp-deficient mutant was constructed as described in Materials and Methods. Reverse transcription and quantitative real-time PCR showed that ftf transcription in the frp mutant was significantly decreased from the level seen for GS5 in CDM containing 1% sucrose, glucose, or fructose (Fig. 1A). Transcription of ftf in the frp mutant was only about 10% of that for the wild-type strain in the presence of 1% glucose, compared to frp mutant values of about 30% and 70% of the values for the wild-type strain in CDM containing fructose and CDM containing sucrose, respectively.
The Ftf activity assays were also carried out to determine whether or not the Ftf activity is also altered in the frp mutant (Fig. 1B). Culture supernatant fluids were used as the source of the enzyme, since earlier results showed that the bulk of the activity was present in this fraction (29). The results showed that Ftf activity was affected significantly by the presence of different carbon sources in the medium in both GS5 and the frp mutant. When S. mutans was cultured in the presence of sucrose, the observed activity was significantly decreased from that observed when cells were cultured in the presence of other sugars. To determine if the diminished Ftf activity was due to increased fructanase activity, fructanase activities were determined in the frp mutant and the wild-type strain in the presence of sucrose, and the results showed no significant differences between the two strains in the presence of sucrose (data not shown). The strains grown in the presence of glucose expressed higher levels of Ftf activity than those grown in the presence of fructose. There was a significant decrease in the Ftf activity for the frp mutant compared to the wild-type strain when these strains were grown in the presence of the monosaccharides (P < 0.05).
The protein levels of Ftf were assessed by Western blotting using anti-Ftf serum (Fig. 1C). The results showed that the carbon sources had a significant influence on the expression of the ftf gene in both the wild-type GS5 strain and the frp mutant. Protein levels of Ftf were highest in glucose-containing medium, in parallel with the activity results. There was no detectable Ftf protein in supernatant fluids of cultures of either the frp mutant or the wild-type strain grown in the presence of sucrose, which reflected the low Ftf activity detected in the culture fluids. These results indicated that frp was required for optimal transcription and expression of ftf gene.
Frp positively regulates expression of glucosyltransferase B and C in the presence of glucose. Since S. mutans can utilize extracellular sucrose to produce the exopolysaccharide glucans by GtfB, -C, and -D, we examined the effects of the frp mutation on the transcription of the gtfB, -C, and -D genes by reverse transcription and quantitative real-time PCR (Fig. 2A and B). The transcriptions of both gtfB and -C were decreased in the frp mutant compared to those of the wild-type strain in the presence of glucose. However, in fructose medium, the transcriptions of gtfB and -C in the frp mutant were similar to those in the wild-type strain. Transcription of gtfB in the frp mutant was significantly decreased in the presence of sucrose compared to that in the wild-type strain. In the presence of sucrose, in contrast, the transcription of gtfC in the frp mutant was similar to that in the wild-type strain. The transcription of gtfD was not significantly altered in the frp mutant compared to that in the wild-type strain under these conditions (data not shown).
The Gtf activity assays were also carried out to examine whether the total Gtf activity is altered in the frp-deficient mutant (Fig. 2C). The results showed that Gtf activity in both GS5 and the frp mutant was affected significantly by the presence of different carbon sources in the medium. When S. mutans was cultured in the presence of sucrose, the Gtf activity level was decreased significantly from that seen with culturing in the presence of glucose. Comparing the frp mutant and the wild-type strain, there was an apparent decrease of Gtf activity in the frp mutant relative to the wild-type strain, as suggested also by the transcription analysis.
The expression of the gtfB and -C genes was also examined by Western blotting using anti-GtfBC (Fig. 2D). The results consistently showed decreased protein expression of both GtfB and -C in the frp mutant compared to the wild-type strain in the presence of glucose. In contrast, only gtfB showed decreased expression in the frp mutant compared to that in the wild-type strain in the presence of 1% sucrose. There was also no difference between the frp mutant and the wild-type strain in the expression of the GtfB and -C proteins in the presence of fructose. Therefore, the frp mutation either decreased or had little effect on the expression of gtf genes, depending upon the sugar source for growth.
The frp mutation inhibits sucrose-dependent biofilm formation by S. mutans. In order to evaluate the effects of the frp mutation on biofilm formation by S. mutans, the biofilm formation assay was carried out as described earlier in the presence of different sugars (Fig. 3). The results showed that the frp mutant formed an amount of biofilm less than that formed by the wild-type strain only in the presence of sucrose (P < 0.05). The gtfB mutant, LN62, also showed attenuated biofilm formation compared to the wild-type strain (P < 0.05) as previously described (37). Levels of biofilm formation were similar for the frp mutant, the gtfB mutant, and the wild-type strain in the presence of glucose or fructose. These results indicated that the influence of Frp on sucrose-dependent biofilm formation may be due primarily to its effects on gtfB gene expression.
Frp is involved in quorum sensing via regulation of the expression of the comC gene. By searching the S. mutans genome database (University of Oklahoma), there was one sequence identified in the comC gene promoter region that has approximately 61.3% identity with the inverted repeat sequence in the ftf gene promoter that binds Frp specifically. Reverse transcription and quantitative real-time PCR results showed that transcription of comC was decreased in the frp mutant compared to that in the wild-type strain in THB medium and sucrose-containing media (Fig. 4). However, there was no significant difference between the frp mutant and the wild-type strain in the presence of glucose (Fig. 4). Since the comC product is involved in the capacity of natural transformation (18), the transformation efficiencies of the frp mutant, wild-type GS5, and a comC-deficient mutant with and without CSP were compared (Table 2). Sucrose was used in these assays, since recent results have indicated that transformation in S. mutans is optimal in the presence of this sugar (B.-Y. Wang and H. K. Kuramitsu, unpublished results). The results showed that the transformation efficiency of the frp mutant was about 27-fold lower than that of the wild-type strain, while that of the comC-deficient mutant was about 100-fold lower than that of the wild-type strain in the absence of CSP. In contrast, the addition of 1.0 μg/ml of synthetic CSP increased the transformation efficiencies by about 21-, 29-, and 6.5-fold in the frp mutant, the comC mutant, and the wild-type strain, respectively, over those without CSP. These results indicated that the frp gene also influences transformation by modulating the expression of CSP in S. mutans.
Autoregulation of Frp. Although our results indicated that Frp acts as a positive regulator in the control of the expression of genes involved in pleiotropic cellular functions, it was also demonstrated that Frp acted as a transcriptional regulator to control its own expression (Fig. 5). Instead of being a positive regulator, Frp is a negative regulator of its own transcription regardless of the carbon source used for growth of the cells. This indicated that frp expression is autoregulated and that this regulation is independent of the sugar source.
Complementation of frp gene. In order to verify that the phenotype of the frp mutant is not the result of polar effects in the construction of the frp mutant, we introduced an intact frp gene into the chromosome of the S. mutans frp mutant. Reverse transcription and quantitative real-time PCR (Fig. 6) showed that, in contrast to the frp mutant, the frp-complemented strain restored the transcription of the ftf, frp, gtfB, and gtfC genes to wild-type levels in the presence of 1% glucose. These results suggested that Frp either directly or indirectly modulates the expression of these genes.
DISCUSSION
Oral bacteria are subject to radical periodic fluctuations in the mouth environment. Due to intermittent eating patterns, vast differences in saliva quantity and composition associated with diurnal rhythms, mechanical forces, and host defense mechanisms, bacteria in plaque are constantly subjected to environmental fluctuations. As a major pathogen involved in dental caries, the ability of S. mutans to optimize its expression of a variety of virulence factors, including extracellular polysaccharide synthesis systems, biofilm formation, and cell-cell signaling, should be essential for the pathogenicity of this bacterium. Since the adaptation to the environment is a complex phenomenon involving coordination of a variety of cellular functions, it is reasonable that bacteria contain a variety of mechanisms to coordinate biochemical changes. Previous studies have indicated several mechanisms for such adaptation in S. mutans. For example, the trigger factor is associated with stress tolerance, competence development, and biofilm formation (44). LuxS-based signaling also affects S. mutans biofilm formation (43, 49). Furthermore, an E11ABman mutation increases the production of Ftf, Gtf, and other gene products (1). Likewise, an HtrA surface protease-deficient mutant is altered in terms of expression of Gtf and Ftf as well as of GbpB, enolase, and glyceraldehyde-3-phosphate dehydrogenase. This mutant also displayed a granular patchy appearance rather than the relatively smooth confluent layer normally seen for the wild type in biofilms (3). Furthermore, the VicRK signal transduction system in S. mutans affects gtfBCD, gbpB, and ftf expression, biofilm formation, and genetic competence development (32). These findings indicate that complex regulatory mechanisms operate in S. mutans. Our results demonstrating that Frp functions as a positive transcriptional regulator involved in exopolysaccharide synthesis and also affects sucrose-dependent biofilm formation and competence development suggest the existence of another novel pleiotropic regulatory system in S. mutans. It is possible that Frp might interact with one or more of the previously mentioned regulatory systems.
Extracellular sugar metabolism has been of interest because of its association with dental caries pathogenesis as a virulence factor of S. mutans. Although ftf and gtfs have distinct genetic organizations and different promoters on the chromosome of S. mutans, they are induced by their common substrate, i.e., sucrose in the diet, and are subject to fluctuation in the presence of different sugars (12, 27, 35, 45). Our results that frp deficiency in S. mutans significantly decreases the transcription of ftf in the presence of sucrose, glucose, and fructose suggest that Frp functions as a positive transcriptional regulator of ftf expression. In addition, the apparent dramatic decrease in Ftf activity as well as in Ftf protein levels in both the wild-type strain and the frp mutant in the culture fluids in the presence of sucrose is likely influenced by the relocation of Ftf to a cell-associated form as a result of insoluble glucan synthesis (29).
It is interesting that the effects of Frp on expression of both gtfB and gtfC genes are somewhat distinct from the effect on ftf expression. In general, Frp appears to function as a positive transcriptional regulator of ftf expression as well as gtfB and gtfC expression, especially when S. mutans is grown in presence of glucose. However, in the presence of sucrose, gtfC expression is primarily Frp independent, in contrast to gtfB and ftf expression. Furthermore, fructose appears to affect ftf expression in the frp mutant much more strongly than it affects the expression of gtfB and gtfC. These results are compatible with the hypothesis that there are multiple regulatory mechanisms involved in the expression of exopolysaccharide-synthetic genes. It will be of interest to determine if the Frp regulatory system interacts with these other systems either directly or indirectly. The difference in sugar effects on gtfB, gtfC, and ftf indicates that different mechanisms may also exist in the regulation of transcription of these genes by Frp, since there is no inverted repeat in the promoter region of the gtfBC gene comparable to that in the ftf promoter region which binds Frp specifically.
Sugar metabolism pathways in S. mutans are also subject to catabolic repression when glucose is present (28, 42). The result that glucose, of all the sugars tested, causes the most significant repression in the frp mutant of the exopolysaccharide synthesis enzymes and of gtfB, gtfC, and ftf relative to the levels of these enzymes and genes found for the wild-type strain suggests that Frp may function, in part, as an antagonist of catabolic repression. The Frp defect may therefore enhance catabolic repression in the presence of glucose. Although other sugars besides glucose may produce intermediates involved in catabolite repression, the present results suggest that glucose is more efficiently metabolized to such intermediates in S. mutans. The results showing that transcription of the comC gene, which is not known to be subject to catabolic repression by glucose, are consistent with this possibility and need to be investigated in detail.
Biofilm formation by S. mutans has been shown to be dependent upon several factors interacting with the oral environment. The expression levels of the ftf, gtfB, and gtfC genes have been shown to increase in biofilm cells (16, 19). Among the different sugars, fructose and glucose have ftf expression effects that are enhanced compared to that of sucrose (26), while both gtfB and gtfC are associated with biofilm formation in the presence of sucrose (38). Fructans produced by Ftf contribute to the virulence of the biofilm by acting as potential binding sites for S. mutans adhesion and also as an extracellular nutrient reservoir for plaque bacteria. In contrast, glucans produced by GtfB and GtfC play major roles in sucrose-dependent biofilm formation by S. mutans (36, 38). GtfC in particular plays a crucial role in sucrose-dependent adhesion and is essential for biofilm formation on smooth surfaces. Thus, the defect in biofilm formation displayed by the frp mutant in the presence of sucrose appears to be primarily due to reduction in GtfB production plus a ftf deficiency in this strain (23). Since Ftf activity is decreased in the frp mutant, the decrease in GtfB activity may result from interference with the normal regulatory effects of Ftf on gtfB expression. Interestingly, the frp mutant is not repressed in comC expression in nonsucrose media and therefore is not defective in sucrose-independent biofilm formation. A recent study indicated that a comC mutant of S. mutans GS5 was defective in this property (50).
Competence development in S. mutans is associated with the quorum-sensing signaling system of this organism (7, 17, 18, 23). The frp mutant was shown to display decreased transcription of the comC gene compared to the wild-type strain in the presence of sucrose. Accordingly, the transformation efficiency of the frp mutant is significantly reduced compared to that of the wild-type strain. Furthermore, the addition of CSP to the cells partially restored transformation efficiency in both the frp mutant and the comC mutant; this suggests a role for Frp in competence development and transformation efficiency. It is not clear why the addition of exogenous CSP to the comC mutant did not allow for full complementation of transformation to the wild-type levels. Such incomplete complementation was also observed for other phenotypic traits of the mutant following exposure to synthetic CSP (data not shown). Other virulence factors regulated by competence (48) may also be attenuated in the frp mutant, and this will be examined in future experiments.
Overall, our results with the frp mutant indicate that Frp has pleiotropic effects on virulence-related cellular functions. Frp appears to be an important regulator of exopolysaccharide synthesis, sucrose-dependent biofilm formation, and competence development. Further investigation of how Frp affects all of these activities at the molecular level will be necessary to evaluate the direct role of Frp in the coordination of these physiological functions. In addition, it will be of interest to determine which environmental cues are used by Frp to control these interactions.
ACKNOWLEDGMENTS
This investigation was supported in part by NIH grant DE03258.
FOOTNOTES
Corresponding author. Mailing address: Department of Oral Biology, State University of New York at Buffalo, 3435 Main Street, Buffalo, NY 14214. Phone: (512) 249-5901. Fax: (716) 829-3942. E-mail: kuramitsu@earthlink.net.
REFERENCES
1. Abranches, J., Y. Y. Chen, and R. A. Burne. 2003. Characterization of Streptococcus mutans strains deficient in EIIABMan of the sugar phosphotransferase system. Appl. Environ. Microbiol. 69:4760-4769.
2. Banas, J. A., and M. M. Vickerman. 2003. Glucan-binding proteins of the oral streptococci. Crit. Rev. Oral Biol. Med. 14:89-99.
3. Biswas, S., and I. Biswas. 2005. Role of HtrA in surface protein expression and biofilm formation by Streptococcus mutans. Infect. Immun. 73:6923-6934.
4. Browngardt, C. M., Z. T. Wen, and R. A. Burne. 2004. RegM is required for optimal fructosyltransferase and glucosyltransferase gene expression in Streptococcus mutans. FEMS Microbiol. Lett. 240:75-79.
5. Burne, R. A. 1998. Oral streptococci... products of their environment. J. Dent. Res. 77:445-452.
6. Cvitkovitch, D. G., Y. H. Li, and R. P. Ellen. 2003. Quorum sensing and biofilm formation in streptococcal infections. J. Clin. Investig. 112:1626-1632.
7. Echenique, J. R., S. Chapuy-Regaud, and M. C. Trombe. 2000. Competence regulation by oxygen in Streptococcus pneumoniae: involvement of ciaRH and comCDE. Mol. Microbiol. 36:688-696.
8. Fronhoffs, S., G. Totzke, S. Stier, N. Wernert, M. Rothe, T. Bruning, B. Koch, A. Sachinidis, H. Vetter, and Y. Ko. 2002. A method for the rapid construction of cRNA standard curves in quantitative real-time reverse transcription polymerase chain reaction. Mol. Cell. Probes 16:99-110.
9. Gibbons, R. J., K. S. Berman, P. Knoettner, and B. Kapsimalis. 1966. Dental caries and alveolar bone loss in gnotobiotic rats infected with capsule forming streptococci of human origin. Arch. Oral Biol. 11:549-560.
10. Hanada, N., and H. K. Kuramitsu. 1989. Isolation and characterization of the Streptococcus mutans gtfD gene, coding for primer-dependent soluble glucan synthesis. Infect. Immun. 57:2079-2085.
11. Hiratsuka, K., B. Wang, Y. Sato, and H. Kuramitsu. 1998. Regulation of sucrose-6-phosphate hydrolase activity in Streptococcus mutans: characterization of the scrR gene. Infect. Immun. 66:3736-3743.
12. Kiska, D. L., and F. L. Macrina. 1994. Genetic regulation of fructosyltransferase in Streptococcus mutans. Infect. Immun. 62:1241-1251.
13. Kopec, L. K., A. M. Vacca-Smith, and W. H. Bowen. 1997. Structural aspects of glucans formed in solution and on the surface of hydroxyapatite. Glycobiology 7:929-934.
14. Kuramitsu, H. K. 1993. Virulence factors of mutans streptococci: role of molecular genetics. Crit. Rev. Oral Biol. Med. 4:159-176.
15. Lemos, J. A., J. Abranches, and R. A. Burne. 2005. Responses of cariogenic streptococci to environmental stresses. Curr. Issues Mol. Biol. 7:95-107.
16. Li, Y., and R. A. Burne. 2001. Regulation of the gtfBC and ftf genes of Streptococcus mutans in biofilms in response to pH and carbohydrate. Microbiology 147:2841-2848.
17. Li, Y. H., P. C. Lau, J. H. Lee, R. P. Ellen, and D. G. Cvitkovitch. 2001. Natural genetic transformation of Streptococcus mutans growing in biofilms. J. Bacteriol. 183:897-908.
18. Li, Y. H., N. Tang, M. B. Aspiras, P. C. Lau, J. H. Lee, R. P. Ellen, and D. G. Cvitkovitch. 2002. A quorum-sensing signaling system essential for genetic competence in Streptococcus mutans is involved in biofilm formation. J. Bacteriol. 184:2699-2708.
19. Mattos-Graner, R. O., M. H. Napimoga, K. Fukushima, M. J. Duncan, and D. J. Smith. 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.
20. Morrison, D. A. 1997. Streptococcal competence for genetic transformation: regulation by peptide pheromones. Microb. Drug Resist. 3:27-37.
21. Munro, C., S. M. Michalek, and F. L. Macrina. 1991. Cariogenicity of Streptococcus mutans V403 glucosyltransferase and fructosyltransferase mutants constructed by allelic exchange. Infect. Immun. 59:2316-2323.
22. Perry, D., and H. K. Kuramitsu. 1989. Genetic linkage among cloned genes of Streptococcus mutans. Infect. Immun. 57:805-809.
23. Petersen, F. C., L. Tao, and A. A. Scheie. 2005. DNA binding-uptake system: a link between cell-to-cell communication and biofilm formation. J. Bacteriol. 187:4392-4400.
24. Quivey, R. G., Jr., W. L. Kuhnert, and K. Hahn. 2000. Adaptation of oral streptococci to low pH. Adv. Microb. Physiol. 42:239-274.
25. Rathsam, C., R. E. Eaton, C. L. Simpson, G. V. Browne, T. Berg, D. W. Harty, and N. A. Jacques. 2005. Up-regulation of competence- but not stress-responsive proteins accompanies an altered metabolic phenotype in Streptococcus mutans biofilms. Microbiology 151:1823-1837.
26. Rozen, R., G. Bachrach, and D. Steinberg. 2004. Effect of carbohydrates on fructosyltransferase expression and distribution in Streptococcus mutans GS-5 biofilms. Carbohydr. Res. 339:2883-2888.
27. Rozen, R., D. Steinberg, and G. Bachrach. 2004. Streptococcus mutans fructosyltransferase interactions with glucans. FEMS Microbiol. Lett. 232:39-43.
28. Saier, M. H., Jr., S. Chauvaux, G. M. Cook, J. Deutscher, I. T. Paulsen, J. Reizer, and J. J. Ye. 1996. Catabolite repression and inducer control in Gram-positive bacteria. Microbiology 142:217-230.
29. Sato, S., and H. K. Kuramitsu. 1986. Isolation and characterization of a fructosyltransferase gene from Streptococcus mutans GS-5. Infect. Immun. 52:166-170.
30. Schilling, K. M., and W. H. Bowen. 1992. Glucans synthesized in situ in experimental salivary pellicle function as specific binding sites for Streptococcus mutans. Infect. Immun. 60:284-295.
31. Schroeder, V. A., S. M. Michalek, and F. L. Macrina. 1989. Biochemical characterization and evaluation of virulence of a fructosyltransferase-deficient mutant of Streptococcus mutans V403. Infect. Immun. 57:3560-3569.
32. Senadheera, M. D., B. Guggenheim, G. A. Spatafora, Y. C. Huang, J. Choi, D. C. Hung, J. S. Treglown, S. D. Goodman, R. P. Ellen, and D. G. Cvitkovitch. 2005. A VicRK signal transduction system in Streptococcus mutans affects gtfBCD, gbpB, and ftf expression, biofilm formation, and genetic competence development. J. Bacteriol. 187:4064-4076.
33. Shibata, Y., and H. K. Kuramitsu. 1996. Identification of the Streptococcus mutans frp gene as a potential regulator of fructosyltransferase expression. FEMS Microbiol. Lett. 140:49-54.
34. Shiroza, T., and H. K. Kuramitsu. 1995. Development of a heterodimer plasmid system for the introduction of heterologous genes into streptococci. Plasmid 34:85-95.
35. Steinberg, D., R. Rozen, M. Bromshteym, B. Zaks, I. Gedalia, and G. Bachrach. 2002. Regulation of fructosyltransferase activity by carbohydrates, in solution and immobilized on hydroxyapatite surfaces. Carbohydr. Res. 337:701-710.
36. Tamesada, M., S. Kawabata, T. Fujiwara, and S. Hamada. 2004. Synergistic effects of streptococcal glucosyltransferases on adhesive biofilm formation. J. Dent. Res. 83:874-879.
37. Tsumori, H., and H. Kuramitsu. 1997. The role of the Streptococcus mutans glucosyltransferases in the sucrose-dependent attachment to smooth surfaces: essential role of the GtfC enzyme. Oral Microbiol. Immunol. 12:274-280.
38. Van Der Ploeg, J. R., and B. Guggenheim. 2004. Deletion of gtfC of Streptococcus mutans has no influence on the composition of a mixed-species in vitro biofilm model of supragingival plaque. Eur. J. Oral Sci. 112:433-438.
39. Vieira, J., and J. Messing. 1987. Production of single-stranded plasmid DNA. Methods Enzymol. 153:3-11.
40. Wang, B., and H. K. Kuramitsu. 2003. Control of enzyme IIscr and sucrose-6-phosphate hydrolase activities in Streptococcus mutans by transcriptional repressor ScrR binding to the cis-active determinants of the scr regulon. J. Bacteriol. 185:5791-5799.
41. Wang, B., and H. K. Kuramitsu. 2005. Inducible antisense RNA expression in the characterization of gene functions in Streptococcus mutans. Infect. Immun. 73:3568-3576.
42. Wen, Z. T., and R. A. Burne. 2002. Analysis of cis- and trans-acting factors involved in regulation of the Streptococcus mutans fructanase gene (fruA). J. Bacteriol. 184:126-133.
43. Wen, Z. T., and R. A. Burne. 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.
44. Wen, Z. T., P. Suntharaligham, D. G. Cvitkovitch, and R. A. Burne. 2005. Trigger factor in Streptococcus mutans is involved in stress tolerance, competence development, and biofilm formation. Infect. Immun. 73:219-225.
45. Wexler, D. L., M. C. Hudson, and R. A. Burne. 1993. Streptococcus mutans fructosyltransferase (ftf) and glucosyltransferase (gtfBC) operon fusion strains in continuous culture. Infect. Immun. 61:1259-1267.
46. Yamashita, Y., W. H. Bowen, R. A. Burne, and H. K. Kuramitsu. 1993. Role of the Streptococcus mutans gtf genes in caries induction in the specific-pathogen-free rat model. Infect. Immun. 61:3811-3817.
47. Yamashita, Y., Y. Tsukioka, Y. Nakano, Y. Shibata, and T. Koga. 1996. Molecular and genetic analysis of multiple changes in the levels of production of virulence factors in a subcultured variant of Streptococcus mutans. FEMS Microbiol. Lett. 144:81-87.
48. Yonezawa, H., and H. K. Kuramitsu. 2005. Genetic analysis of a unique bacteriocin, Smb, produced by Streptococcus mutans GS5. Antimicrob. Agents Chemother. 49:541-548.
49. Yoshida, A., T. Ansai, T. Takehara, and H. K. Kuramitsu. 2005. LuxS-based signaling affects Streptococcus mutans biofilm formation. Appl. Environ. Microbiol. 71:2372-2380.
50. Yoshida, A., and H. K. Kuramitsu. 2002. Multiple Streptococcus mutans genes are involved in biofilm formation. Appl. Environ. Microbiol. 68:6283-6291.(Bing Wang and Howard K. Kuramitsu)