INTRODUCTION

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Streptococci formerly classified as Streptococcus sanguis have recently been subclassified into at least five distinct genetic groups. These groups have been assigned the species names S. sangius sensu stricto, Streptococcus gordonii, Streptococcus oralis, Streptococcus mitis, and Streptococcus parasanguis (14, 15) and are collectively called sanguis (group) streptococci. These streptococci are early-colonizing microorganisms in the oral cavity of neonates as well as on adult cleaned tooth surfaces (17). The distribution of these species varies among oral sites and changes as dental plaque matures (6, 23). In contrast, mutans streptococci colonize the oral cavity only after the eruption of teeth (8).

Mutans streptococci and Streptococcus salivarius (29) have multiple glucosyltransferases (GTases) encoded by multiple gtf genes, e.g., gtfB, gtfC, and gtfD, in Streptococcus mutans (10, 16). These enzymes synthesize water-soluble and/or -insoluble glucans from sucrose. They contribute to the development of dental plaque and, eventually, to the initiation of dental caries. Recent studies indicate that adhesive glucan is synthesized from sucrose in concert with these GTases in S. mutans (7).

S. oralis, S. gordonii, and S. sanguis are known to possess GTases and produce extracellular polysaccharide from sucrose (36). However, only a limited number of investigations of GTase from sanguis group streptococci have been performed. Recently, the gene encoding S. gordonii strain Challis GTase (gtfG) has been cloned and sequenced (35), and a regulatory gene, rgg, has been described as a positive transcriptional regulator (30, 31, 33). Similar positive regulatory functions have been identified in the rgg gene of S. pyogenes (3) as well as Lactococcus lactis (27).

Since S. oralis is an earlier colonizer in the oral flora (6, 25, 32), the infection and colonization of mutans streptococci may be affected by the presence of S. oralis, and the glucan synthesized by S. oralis GTase may function as a substratum for adhesion of the bacteria. In addition, the prevalence of sanguis group streptococci was found to be different between caries-active and caries-inactive individuals (24).

In this study, we purified a GTase protein from S. oralis and determined its immunochemical properties and contribution to the sucrose-dependent cellular adherence of S. mutans. In addition, a gene encoding S. oralis GTase (designated gtfR) and its regulatory gene (rgg) were cloned and sequenced.

	MATERIALS AND METHODS

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Bacterial strains and growth media. S. oralis ATCC 10557 was used in most of the experiments. For comparison, S. oralis SK23 and ATCC 9811, S. sanguis ATCC 10556, ST3, and ST7, S. gordonii ATCC 10558, F90A, and SK51, S. mitis SK24 and ATCC 903, S. mutans MT8148, S. sobrinus 6715, and S. salivarius HHT were selected from our culture collection. Organisms were routinely cultured in brain heart infusion (BHI) broth (Difco Laboratories, Detroit, Mich.) or mitis salivarius (MS) agar (Difco). Escherichia coli XL-2 (Stratagene Ltd., Cambridge, United Kingdom) was cultured in Luria-Bertani (LB) medium aerobically. Erythromycin, kanamycin, and ampicillin (Wako Pure Chemicals, Osaka, Japan) were added to LB medium to produce final concentrations of 500, 30, and 100 µg/ml, respectively. Erythromycin (5 µg/ml) and kanamycin (250 µg/ml) were added to MS agar for selection of the S. oralis transformants.

Preparation of glucosyltransferases. S. oralis ATCC 10557 was cultured in 5 liters of dialyzed TTY medium (12) at 37°C to an optical density of 0.8 at 550 nm. The culture supernatant was collected by centrifugation and adjusted to 60% saturation with ammonium sulfate. The precipitate was dissolved in 10 mM sodium phosphate buffer (NaPB) (pH 6.5) and then dialyzed against the same buffer. The crude sample was applied to a Q Sepharose FF (Pharmacia Biotech AB, Uppsala, Sweden) column (bed volume, 10 ml) and eluted with a linear gradient of 0 to 1.0 M NaCl in the same buffer. Active fractions were pooled, dialyzed against 10 mM potassium phosphate buffer (KPB) (pH 6.0), applied to a Bio-Scale CHT10-I column (bed volume, 10 ml; Bio-Rad Laboratories, Hercules, Calif.), and then eluted with a 10 to 500 mM KPB linear gradient.

Generation of antiserum. Antisera were prepared by repeated intramuscular injections of rabbits with the purified GTase from S. oralis ATCC 10557 suspended in Freund's complete adjuvant (Difco) followed by immunization with the antigen suspended in Fruend's incomplete adjuvant (Difco). The antibody to S. oralis GTase was purified from rabbit antiserum by repeated 33% saturation with ammonium sulfate.

Glucan synthesis assay. GTase activity was determined using [glucose-¹⁴C]sucrose with or without primer dextran T10, as described previously (11). Briefly, reaction mixtures composed of GTase, 10 mM [glucose-¹⁴C]sucrose (11.47 GBq/mmol), and 0 or 20 µM dextran T10 in 20 µl of 50 mM KPB (pH 6.0) were incubated for 1 h at 37°C, spotted on a filter paper square (1.5 by 1.5 cm), and dried in air. The filters were washed with methanol or distilled water and then immersed in scintillation fluid to estimate the amount of total [¹⁴C]glucan or water-insoluble [¹⁴C]glucan. Kinetic constants were determined by Lineweaver-Burk analyses of the glucan synthesis rates.

Determination of pI and optimum pH. The pI was determined by analytical isoelectric focusing using a PhastSystem (Pharmacia) with a PhastGel IEF3-9 (Pharmacia). After electrophoresis, the gel was incubated for 1 h at 37°C in 10 mM NaPB (pH 6.5) containing 5% sucrose, 2% Triton X-100, and 0.05% NaN₃. The enzyme activity was visualized by periodic acid-Schiff staining. The optimum pH of GTase was determined by measuring the GTase activity in 50 mM KPB (pH 5.0 to 7.5).

SDS-PAGE and Western blotting. Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) and Western blot analyses were carried out as described previously (9). Briefly, GTase samples and E. coli cells carrying the recombinant plasmid were suspended in SDS gel-loading buffer (26) and boiled for 5 min. Proteins separated by SDS-PAGE were transferred onto a polyvinylidene difluoride membrane (Immobilon; Millipore). After being blocked with 5% bovine serum albumin, the membrane was reacted with the rabbit antibody to S. oralis GTase at 37°C for 1 h, and the antibody which was bound to the protein band(s) was detected by a solid-phase immunoassay.

Effects of S. oralis GTase on the sucrose-dependent adhesion of S. mutans resting cells. S. mutans strain MT8148 cells grown in BHI broth were washed at 0°C with 0.1 M KPB (pH 6.0) containing 0.05% NaN₃. The centrifuged cells were resuspended in the same buffer containing 1% sucrose and then adjusted to an optical density of 1.0 at 550 nm. Aliquots (3 ml) of the cell suspension were mixed with various amount of S. oralis GTase and incubated at 37°C for 18 h at a 30° angle. Next, the culture tubes were vigorously vibrated with a Vortex mixer for 3 s. The degree of cell adhesion was determined by reading the optical density at 550 nm and expressed as the percentage of total cell mass. To assess the adhesion of S. mutans growing cells, the organism was grown at 37°C for 18 h at a 30° angle in BHI broth containing 1% sucrose. The percent adhesion was determined as described above.

Amino acid sequence. S. oralis GTase was subjected to SDS-PAGE and blotted onto ProBlott membranes (Applied Biosystems, Foster City, Calif.). The GTase band was excised from several lanes and subjected to sequencing using an ABI 477A/120A protein sequencer (Applied Biosystems).

DNA manipulations. Restriction enzymes, ligase, and other DNA-modifying enzymes were purchased from New England Biolabs (Beverly, Mass.) or Takara (Kyoto, Japan). Manipulations of DNA with these enzymes were performed as recommended by the manufacturers. All other DNA manipulations were carried out using standard protocols (26).

Chromosomal DNA isolation and Southern blot analysis. Organisms were grown in BHI broth for 18 h at 37°C, collected, and then washed by centrifugation. Cells (750 mg [wet weight]) were suspended in 5 ml of 50 mM NaCl-10 mM Tris-HCl (pH 7.4) and then digested with mutanolysin (0.25 mg/ml; Dainippon Pharmaceutical Co., Osaka, Japan) for 1 h at 50°C, and N-lauroyl sarcosine (final concentration, 1.5%) and EDTA (final concentration, 10 mM) were added to lyse the cells. The lysate was treated with RNase (0.3 mg/ml; Wako) and proteinase K (0.3 mg/ml; Merck, Darmstadt, Germany). The DNA was purified from the cell lysate by phenol and phenol-chloroform extractions and then collected by ethanol precipitation.

PCR. PCR was performed in reaction mixtures containing 50 mM KCl, 10 mM Tris-HCl (pH 8.3), 1.5 mM MgCl₂, 200 µM deoxyribonucleoside triphosphate, 1.0 µM primer, template DNA (<10 ng/µl), and AmpliTaq Gold DNA polymerase (0.025 U/µl; Applied Biolystems). Amplification was performed in a Gene AmpPCR System 2400 apparatus (Perkin-Elmer) as specified by the manufacturer. Degenerate PCR was performed as follows: a preincubation step at 95°C for 9 min followed by 30 cycles of a denaturation step at 94°C for 30 s, a primer-annealing step at 36°C for 30 s, and an extension step at 60°C for 30 s. Long PCR was performed using a TaKaPa LA PCR kit Ver 2.1 (Takara), as recommended by the manufacturer.

Cloning and sequencing of the GTase gene. Two sets of genomic libraries were constructed by cloning EcoRI- or KpnI-digested S. oralis ATCC 10557 chromosomal DNA into plasmid pMW119 (Nippon Gene) or pUC19 (Takara) and then transforming them into competent E. coli. In addition, a vector named pGEM-T Easy (Promega, Madison, Wis.) was used for cloning of the PCR products.

Expression of recombinant GTase. E. coli carrying a recombinant plasmid was grown in LB broth (3 ml) to an optical density of 0.6 at 550 nm. Cells were collected by centrifugation, suspended in 100 µl of 10 mM NaPB (pH 6.0), and disrupted by sonication. The sonic supernatant was then separated and examined for glucan synthesis.

Transformation of S. oralis. S. oralis was subjected to transformation as reported previously (9). Briefly, the recipient organisms were cultured in Todd-Hewitt broth (Difco) supplemented with 10% heat-inactivated horse serum (Gibco, Grand Island, N.Y.) for 18 h. The culture was diluted 1:40 with the broth (10 ml) and then incubated for another 1.5 h at 37°C, and the donor DNA was added to a final concentration of 25 µg/ml. The culture was further incubated for 2 h, concentrated approximately 10-fold by centrifugation, and then spread on MS agar plates containing antibiotics. The plates were incubated in a CO₂ incubator for 2 to 3 days at 37°C, and possible transformant colonies were picked up for further examinations.

Construction of the insertional mutants. Recombinant plasmid pYT303 or pYT311 carrying the 830- or 1,070-bp fragment of the erythromycin resistance gene (erm) from pVA838 (20) or the kanamycin resistance gene (aphA) from transposon Tn1545 (2) was used. A subclone, pTHR8, carrying a 1.5-kb SphI-PstI insert containing the rgg gene was generated from pTH171. A 2.5-kb DNA fragment containing the center portion of the gtfR gene was amplified by PCR and cloned to generate pTH808. pTHR8 or pTH808 was restricted with ApaI or HindIII to be linear at a unique site. The linear plasmid was then blunted and ligated with the erm or aphA cassette to yield pTHR805 or pTH818. After being made linear at the unique PstI site, the plasmid was introduced into S. oralis ATCC 10557 by transformation to allow an allelic exchange.

Statistical analysis. Differences between S. oralis GTase concentrations and S. mutans resting-cell adhesion were determined by analysis of variance with subsequent use of the Tukey-Kramer multiple-comparisons test. Significance levels were taken at P < 0.01.

Nucleotide sequence accession numbers. The nucleotide sequences of the rgg and gtfR gene have been deposited in the DDBJ database under accession no. AB025228.

	RESULTS

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Purification of S. oralis GTase. GTase was purified from the culture supernatant of S. oralis ATCC 10557 by ammonium sulfate precipitation followed by anion-exchange and hydroxylapatite chromatography (Fig. 1). The recovery of the purified GTase preparation was 1.7%, and the degree of purification was 42-fold. The specific activity of the purified enzyme, GTase-R, was 8.0 mU/µg of protein (Table 1). SDS-PAGE of GTase-R gave a single protein band with a molecular mass of 173 kDa. The optimum pH and pI values were 6.5 and 6.3, respectively (Fig. 2). The K_m value was determined to be 2.49 mM. Glucan synthesized by GTase-R from sucrose was largely water soluble (89.7%), and its production was not enhanced in the presence of the primer dextran T10.

View larger version (27K): [in this window] [in a new window]   FIG. 1.   Chromatographic purification of GTase-R from S. oralis ATCC 10557. (A) Separation of ammonium sulfate-precipitated GTase (60% saturation) by anion-exchange chromatography on a Q Sepharose FF column (bed volume, 10 ml). GTase was eluted with a linear gradient of 0 to 0.3 M NaCl. (B) Further purification of GTase-R containing fractions from the elution profile shown in panel A on a Bio-Scale CHT10-I column (bed volume, 10 ml). Elution was done with a 10 to 500 mM KPB linear gradient. A280, optical density at 280 nm.

View larger version (37K): [in this window] [in a new window]   FIG. 2.   Physical characteristics of GTase-R. (A) SDS-PAGE of GTase preparations at different stages of purification. Lanes: 1, culture supernatant; 2, ammonium sulfate precipitate; 3, pooled active fractions from Q-ion-exchange chromatography; 4, pooled active fraction from CHT-10 hydroxylapatite chromatography; M, molecular mass markers. (B) Isoelectric focusing-PAGE of GTase-R. Lane: 1, pI markers (3.50 to 8.65); 2, Purified GTase-R visualized by PAS staining. (C) Effect of pH on GTase activity.

Immunological properties of GTase-R. Western blot analyses revealed that the rabbit antibody to GTase-R reacted strongly with GTase preparations from other sanguis streptococci and cell-free GTase (CF-GTase) but not with CA-GTase from S. mutans. Further, the enzyme activity of GTase-R was markedly inhibited by the antibody to GTase-R. The antibody strongly inhibited S. sanguis and S. gordonii GTase, as well as S. mutans CF-GTase. S. sobrinus and S. salivarius GTases were only weakly inhibited, while S. mutans CA-GTase was not affected by the antibody (Table 2). The inhibition of glucan synthesis exhibited a similar pattern to the reactivity when analyzed by Western blotting.

Effects of GTase-R on the adhesion of S. mutans. The effects of GTase-R on the sucrose-dependent adhesion of S. mutans resting cells are shown in Fig. 3. Cells incubated without GTase-R adhered to the glass surface only loosely, and approximately 60% of the cells were easily removed by vibration. However, the addition of a small amount of GTase-R (1 mU/ml) resulted in firm adhesion of the S. mutans cells. This adhesion was as strong as that of the S. mutans cells grown in sucrose-containing medium.

View larger version (24K): [in this window] [in a new window]   FIG. 3.   Effects of GTase-R on the cellular adhesion of S. mutans. S. mutans cells grown in BHI broth were resuspended at 0°C in 0.1 M KPB (pH 6.0) with 0.05% NaN3 containing 1% sucrose. The cell suspension was mixed with increasing amounts of GTase-R and incubated at 37°C for 18 h at a 30° angle (resting cells). As a positive control, S. mutans was grown in BHI broth with 1% sucrose at 37°C for 18 h at a 30° angle (growing cells). Numbers of adhesive cells were then determined. Data are expressed as means and standard deviations of triplicate experiments. The asterisks indicate statistical significance (P < 0.01) from the sucrose value for resting cells incubated in sucrose containing KPB without GTase-R.

Amino acid sequencing of GTase-R. The N-terminal amino acid sequence of GTase-R was determined to be DDVKQVVVQEPATAQTSGPGQQ. This sequence did not show any similarity to other reported sequences, including GTases from S. gordonii and other oral streptococci, by BLAST and FASTA homology searching.

Cloning of the GTase-R gene (gtfR) by PCR with degenerate primers. A schematic diagram of the GTase-R gene and cloning strategy is shown in Fig. 4. PCR was done using the degenerated oligonucleotides corresponding to the N-terminal amino acid sequence VKQVVV (5'GTNAARCARGTNGTNGT3', forward primer) and GPGQQ (5'YTGYTGNCCNGGNCC3', reverse primer). A 60-bp gene fragment was cloned into a pGEM-T Easy vector, resulting in the recombinant plasmid pTHN01. The sequence of the insert of pTHN01 was consistent with the N-terminal amino acid sequence. Then a 60-mer oligonucleotide primer (5'GTAAAGCAGGTTGTAGTTCAAGAACCTGCTACAGCTCAGACTAGTGGTC CCGGTCAGCAA3') was synthesized and used for hybridization. The E. coli transformants carrying the EcoRI-digested insert were screened by colony hybridization with this primer. The recombinant plasmids pTH121 and pTH171, with 1.4- and 6.1-kb S. oralis chromosomal inserts, respectively, were isolated.

View larger version (13K): [in this window] [in a new window]   FIG. 4.   Genetic map and cloning strategy of the rgg and gtfR genes.

View larger version (15K): [in this window] [in a new window]   FIG. 5.   Western blot analysis and glucan synthesis activity of recombinant GTase-R. (A) The recombinant proteins and native GTase-R were separated by SDS-PAGE and blotted onto a polyvinylidene difluoride membrane. The blot was reacted with the antibody to GTase-R. (B) The sonic supernatants of E. coli cells carrying the recombinant plasmid were used for the glucan synthesis assay. GTase activity was determined as the amount of [14C]glucan synthesized from [glucose-14C]sucrose. Data are expressed as means and standard deviations of triplicate experiments.

Nucleotide sequences. The nucleotide sequence determined in this study is shown in Fig. 6. The ORF in pTH171 was composed of 861 bp and encoded a polypeptide of 287 amino acids. This gene exhibited a high degree of homology (74%) to the rgg gene of S. gordonii (accession no. M89776), which has been reported to be a regulatory gene of GTase. We therefore designated it rgg. A multiple alignment of the deduced amino acid sequence of rgg revealed that the rgg gene of S. oralis exhibited a 76% homology to the S. gordonii rgg gene, whereas the homology of S. oralis rgg to the S. pyogenes and L. lactis rgg genes was only 21%.

View larger version (65K): [in this window] [in a new window]   FIG. 6.   DNA sequence and deduced amino acid sequence of the rgg and gtfR genes. The putative promoter (10 and 35) and ribosome binding sites (SD) are shown. The sequence corresponding to the identified N-terminal amino acid sequence is marked. Regions of dyad symmetry are indicated by arrows.

View larger version (83104K): [in this window] [in a new window]   FIG. 7.   Alignment of the deduced amino acid sequences of GtfR and GtfG of S. gordonii (accession no. U12643). Identical amino acid sequences are indicated by shaded boxes.

View larger version (49K): [in this window] [in a new window]   FIG. 8.   Southern blot analysis of chromosomal DNA from various oral streptococci digested with EcoRI. As a probe, the PCR-amplified DNA fragment corresponding to the N-terminal sequence of gtfR was used. Lanes: 1, S. mutans MT8148; 2, S. sobrinus 6715; 3, S. salivarius HHT; 4, S. sanguis ATCC 10556; 5, S. sanguis ST3; 6, S. sanguis ST7; 7, S. oralis ATCC 10557; 8, S. oralis SK23; 9, S. oralis ATCC 9811; 10, S. gordonii ATCC 10558; 11, S. gordonii F90A; 12, S. gordonii SK51; 13, S. mitis SK24; and 14, S. mitis ATCC 903.

Inactivation of the rgg or gtfR gene. Inactivation of the chromosomal rgg or gtfR gene of S. oralis ATCC 10557 was performed by insertion of the erm or aphA gene. The colony morphology and glucan synthesis activity of representative transformants are shown in Fig. 9. The rgg mutant had a flat and dull appearance without the zooglealic zone, while the gtfR mutant had smaller colonies with a more transparent appearance. The GTase activity of both mutants had decreased to about 10% that of the parent strain.

View larger version (37K): [in this window] [in a new window]   FIG. 9.   Colonial morphology and GTase activity of S. oralis ATCC 10557 (A), and its erythromycin-resistant rgg-deficient (B) and kanamycin-resistant gtfR-deficient (C) mutants. GTase activity was determined by the synthesis of [14C]glucan from [glucose-14C]sucrose. Data are expressed as means ± standard deviations of triplicate experiments.

	DISCUSSION

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Several methods of comparing the amino acid sequences of GTases from various oral streptococci have revealed that GTase possesses two functional domains. One is a catalytic domain that is composed of approximately 800 amino acid residues located in the N-terminal region, while the other is a glucan binding domain that includes a large number of repeated units in the C-terminal region. The former contains common putative active-site peptides involved in sucrose hydrolysis (22, 28). Molecular modeling analyses have suggested that the core region contains a cyclically permuted form of the (/)₈-barrel structure (4, 19). Recently, circular dichroism analysis has verified the presence of that structure (21). Direct repeats of the glucan binding domain are also found in the C-terminal region of the ligand binding proteins in some gram-positive organisms, including S. mutans glucan binding protein, Clostridium difficile ToxA and ToxB, and lysins from S. pneumoniae and its bacteriophage (37). Structure-function relationship studies have revealed that GTases synthesizing water-insoluble and/or water-soluble glucans exhibit an almost identical amino acid sequence at the putative active sites. The putative active sites of gtfR are the same as those of gene encoding GTases that synthesize water-soluble glucan (28) (Fig. 7). Deletion of glucan binding domain direct repeats affects the activity and localization of GTase (13), as well as its glucan production (1). Our finding that the recombinant protein of pTH171 lacking the C-terminal region of gtfR did not exhibit GTase activity (Fig. 5) accords with the finding reported by Vickerman et al. (34) using S. gordonii GTase.

It seems quite clear from the GTase antibody inhibition data that there are three groups of organisms with similar levels of inhibition (Table 2). The enzymes from S. oralis, S. sanguis, and S. gordonii strains are all inhibited by 75 to 80%; those from S. sobrinus, S. salivarius, and the cell-free enzyme from S. mutans are all inhibited by 25 to 30%; and the cell-associated enzyme from S. mutans is inhibited by 6%. These differences in inhibition should reflect differences in the nucleotide sequence of GTase gene.

Southern blot analysis indicated that the N-terminal 130 amino acid residues were conserved exclusively in S. oralis (Fig. 8). Thus, this region is thought to be a species-specific sequence for S. oralis. Classification of sanguis streptococci has been difficult. In fact, DNA-DNA hybridization studies have revealed that many strains which had previously been identified phenotypically as S. mitis, S. oralis, or S. sanguis were not correctly classified (5). Moreover, 16S rRNA sequencing analysis has indicated that S. mitis, S. pneumoniae, and S. oralis exhibited >99% homology in nucleotide sequencing (14). Thus, the 5' region of gtfR can be used as a useful probe in PCR amplification for the rapid and exact classification of S. oralis.

S. oralis also possessed the rgg gene immediately upstream of gtfR. The presence of an rgg-like gene in strains of S. oralis and S. sanguis was previously reported (33). Moreover, the deduced amino acid sequences of the rgg gene from S. oralis and S. gordonii were very similar. An S. oralis mutant strain in which the rgg gene was inactivated displayed a soft-colony phenotype and markedly reduced GTase activity (Fig. 9). These results indicate that the rgg gene of S. oralis is a positive transcriptional regulator of gtfR. Similar findings have been reported for S. gordonii (31). However, the putative RNA secondary structures at the junction of the rgg and gtf genes in S. oralis were different from those in S. gordonii. These results clearly indicate that the regulatory mechanism of the rgg gene is different for S. gordonii and S. oralis.

It is interesting that the colony morphologies of our rgg and gtfR mutants were different, even though both mutants exhibited minimal GTase activity (Fig. 9). Recently, rgg-like genes have been identified in some bacterial species; the rgg gene in S. pyogenes positively regulates the expression of cysteine proteinase (3, 18), while that of L. lactis has been claimed to be glutamate--aminobutyrate antiporter and glutamate decarboxylase (27). Those two test strains have no GTase, and it is of interest to know if the rgg-like genes regulated the expression of proteins other than GTase. These findings, coupled with the results of the present study, may suggest that the rgg gene of S. oralis can regulate a gene(s) other than gtfR.

Sucrose-dependent adhesion is an important pathogenic trait of mutans streptococci. S. mutans produces three GTases: GTase-I, GTase-SI, and GTase-S, which are encoded by the gtfB, gtfC, and gtfD genes, respectively. GTase-I is present in association with the cell surface, whereas GTase-S is released extracellularly. GTase-SI can be coextracted from cells by 8 M urea treatment and is more likely to play an important role in cellular adhesion (9). S. mutans adheres firmly to solid surfaces by the cooperative action of these GTases in the presence of sucrose in vivo. However, firm cellular adhesion was obtained only when S. mutans was grown in a sucrose-containing broth medium. In this study, we revealed that S. mutans resting cells firmly adhered to a glass surface in the presence of S. oralis GTase and sucrose (Fig. 3). The maximum adhesion was almost equivalent to that of growing S. mutans cells in terms of adhesion strength and macroscopic features. However, the presence of excess amounts of GTase-R resulted in a surfeit of soluble-glucan synthesis, which in turn may interfere with the cell adhesion of S. mutans and cariogenic dental-plaque formation. These results indicate that S. oralis GTase may play a significant role in the formation of dental plaque in vivo.

Further, the evidence reported here suggests that S. oralis GTase strongly contributes to the establishment of oral bacterial biofilms, and therefore a more precise description of its mechanism should be sought.