Purification, Characterization, and Molecular Analysis of the Gene Encoding Glucosyltransferase from Streptococcus oralis

Abstract

Streptococcus oralis is a member of the oral streptococcal family and an early-colonizing microorganism in the oral cavity of humans. S. oralis is known to produce glucosyltransferase (GTase), which synthesizes glucans from sucrose. The enzyme was purified chromatographically from a culture supernatant of S. oralis ATCC 10557. The purified enzyme, GTase-R, had a molecular mass of 173 kDa and a pI of 6.3. This enzyme mainly synthesized water-soluble glucans with no primer dependency. The addition of GTase markedly enhanced the sucrose-dependent resting cell adhesion of Streptococcus mutans at a level similar to that found in growing cells of S. mutans. The antibody against GTase-R inhibited the glucan-synthesizing activities of Streptococcus gordonii and Streptococcus sanguis, as well as S. oralis. The N-terminal amino acid sequence of GTase-R exhibited no similarities to known GTase sequences of oral streptococci. Using degenerate PCR primers, an 8.1-kb DNA fragment, carrying the gene (gtfR) coding for GTase-R and its regulator gene (rgg), was cloned and sequenced. Comparison of the deduced amino acid sequence revealed that the rgg genes of S. oralis and S. gordonii exhibited a close similarity. The gtfR gene was found to possess a species-specific nucleotide sequence corresponding to the N-terminal 130 amino acid residues. Insertion of erm or aphA into the rgg or gtfR gene resulted in decreased GTase activity by the organism and changed the colony morphology of these transformants. These results indicate that S. oralis GTase may play an important role in the subsequent colonizing of mutans streptoccoci.

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

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.

GTase samples from other streptococci were obtained from the culture supernatants of test strains by 50% saturation ammonium sulfate precipitation. Cell-associated GTase (CA-GTase) was extracted from centrifuged cells of S. mutans with 8 M urea followed by ammonium sulfate precipitation (11).

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.

Southern blot analysis was carried out as a standard procedure. Briefly, chromosomal DNA from the test organisms was digested with EcoRI, separated by electrophoresis on a 0.8% agarose gel, and transferred onto a nylon membrane (Hybond-N; Amersham, Little Chalfont, United Kingdom). Next, the DNA was cross-linked to the membrane by UV radiation. A 397-bp DNA fragment corresponding to positions 54 to 186 in the deduced amino acid of the gtfR gene was amplified by PCR and used as a probe. The membrane was then hybridized stringently with the ³²P labeled probe.

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.

For a DNA-sequencing template, plasmid DNA and PCR products were prepared using a Wizard Plus Minipreps DNA purification system (Promega) and a Centricon 100 spin column (Millipore, Bedford, Mass.), respectively. The dideoxy dye termination reaction was performed with an ABI PRISM cycle-sequencing kit (Perkin Elmer) in a GeneAmp 2400 thermal cycler. The products were then analyzed using an automated DNA sequencer model 373 (Applied Biosystems). The homology search, multiple-sequence alignment, and phylogenetic tree creation were performed with the BLAST, FASTA, and CLUSTAL W programs on the DDBJ “supernig” computer system.

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

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.

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.

TABLE 1.

Purification of S. oralis GTase

Preparationa step	Total amt of protein (mg)	Total GTase activity (U)	GTase sp act (U/mg)	Recovery (%)	Purification (fold)
Culture supernatant	736	140	0.19	100	1
Ammonium sulfate precipitation	180	117	0.65	84.0	3.4
Q Sepharose fraction	5.5	4.0	0.72	2.9	3.8
CHT-I fraction	0.3	2.4	8.00	1.7	42.0

^aS. oralis ATCC 10557 was grown in 5 liters of dialyzed TTY medium to an optical density of 0.8 at 550 nm. The culture supernatant was concentrated by a 60% saturation of ammonium sulfate. The enzyme fraction was purified on a Q Sepharose FF column followed by a Bio-Scale CHT10-I column. 

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.

TABLE 2.

Effects of anti-S. oralis ATCC 10557 GTase antibody on the GTase activities of various oral streptococci

GTase origina	GTase activity (dpm)b		% Inhibition
	Without anti-GTase-R	With anti-GTase-R
S. oralis
ATCC 10557	4,930.3 ± 12.8	831.8 ± 14.2	83.1
SK23	4,997.9 ± 49.7	1,065.1 ± 42.2	78.7
ATCC 9811	5,022.0 ± 37.6	1,128.8 ± 36.7	77.5
S. sanguis
ATCC 10556	5,000.0 ± 100.0	1,305.4 ± 59.5	73.9
ST3	5,027.1 ± 25.8	1,244.4 ± 36.1	75.3
ST7	5,053.5 ± 21.8	1,243.8 ± 29.6	75.4
S. gordonii
ATCC 10558	4,975.8 ± 37.9	949.4 ± 23.3	80.9
90A	4,950.2 ± 32.9	1,468.6 ± 30.9	70.3
SK51	4,956.5 ± 40.9	1,007.3 ± 58.9	79.7
S. mutans
MT8148 cell-associated	4,996.4 ± 53.3	4,687.9 ± 63.8	6.2
MT8148 cell-free	4,964.7 ± 72.5	3,431.3 ± 61.6	30.9
S. sobrinus
6715	5,006.9 ± 138.4	3,732.1 ± 194.9	25.5
S. sobrinus
HHT	4,989.9 ± 66.9	3,662.2 ± 76.5	26.6

^aGTase was from the concentrates of test strain culture supernatants, except for the cell-associated GTase from S. mutans. The cell-associated GTase was extracted from the cells of S. mutans with 8 M urea. 

^bThe GTase fraction (1 mU) was reacted with the antibody to GTase-R (32 μg of protein) at 37°C for 30 min or left unreacted. The reaction mixture was then incubated with [glucose-¹⁴C]sucrose at 37°C for 1 h, and the amount of synthesized [¹⁴C]glucan was measured. Data are expressed as means ± standard deviations of triplicate experiments. 

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.

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.

FIG. 4.

Genetic map and cloning strategy of the rgg and gtfR genes.

A sequenced analysis of pTH171 revealed that an open reading frame (ORF) composed of the 861-bp nucleotide was present, and an incomplete reading frame without the termination codon was identified 89 bp downstream of the ORF. This frame encoded the N-terminal amino acid sequence of GTase-R; however, the molecular mass of the recombinant protein expressed by pTH171 was only 128 kDa (Fig. 5). To clone the C-terminal region, the KpnI-digested library was screened and pTH181, carrying a 1.8-kb insert with a termination codon, was obtained. The complete nucleotide sequence of the gtfR gene was determined by reconstruction of the DNA sequences from the inserts of pTH171 and pTH181. To confirm the sequence, primers for amplification of the whole gtfR gene were synthesized and long PCR amplification was performed. The PCR product was cloned into a pGEM-T Easy vector to yield pTH275. The recombinant protein expressed by pTH275 had a molecular mass of 175 kDa and glucan-synthesizing activity (Fig. 5).

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%.

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.

The gtfR gene was composed of 4,728 bp, which encoded a polypeptide composed of 1,575 amino acids with a predicted molecular mass of 177 kDa and a pI of 5.58. The alignment of the deduced amino acid sequence of the gtfR gene and the gtfG gene encoding S. gordonii GTase is shown in Fig. 7. The gtfR gene displayed 79.9% homology to the gtfG gene. The N-terminal sequence from amino acids 55 to 186, deduced from the nucleotide sequence data of gtfR, was completely different from that of other streptococcal species. These findings were supported by Southern hybridization analyses, which indicated that no hybridized bands were detected, except for strains of S. oralis, when the PCR-amplified DNA fragment corresponding to the N-terminal 140 amino acid residues of gtfR was used as a probe (Fig. 8).

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.

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.

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

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 (α/β)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-γ-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.