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Differential and Quantitative Analyses of mRNA Expression of Glucosyltransferases from Streptococcus mutans MT8148

¹ Departments of Pedodontics and
² Oral Microbiology, Osaka University Graduate School of Dentistry, 1-8, Yamadaoka, Suita-Osaka, 565-0871, Japan;

*corresponding author, fujiwara{at}dent.osaka-u.ac.jp

	ABSTRACT

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Streptococcus mutans produces three glucosyltransferases, coded by gtfB, gtfC, and gtfD, whose cooperative action is essential for sucrose-dependent cellular adhesion. This cellular adhesion plays an important role in the formation of dental plaque and the initiation of dental caries. Since they bear genetic similarities and are large in size, differentiation of their gene expression has been difficult, and little is known about the dynamic process of gtf expression. Using a real-time reverse-transcription/polymerase chain-reaction, we determined the expression of each gtf. Under various conditions, the relative levels of transcription were gtfB > gtfD > gtfC. Sucrose enhanced gtfD expression, whereas it reduced that of gtfB and gtfC, suggesting the presence of independent promoters. Quantitative analyses demonstrated coincidence between the ratio of expression of each gtf and the ratio previously identified as optimal for sucrose-dependent adhesion in vitro, suggesting that S. mutans produces GTF at an optimal ratio to adhere to the tooth surface.

KEY WORDS: glucosyltransferase • Streptococcus mutans • real-time RT-PCR • gene expression

	INTRODUCTION

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Streptococcus mutans is considered to be one of the most important pathogens in the development of dental caries in humans (Hamada and Slade, 1980; Loesche, 1986). From sucrose, the organism synthesizes adhesive extracellular glucans which mediate the firm attachment of cells to the tooth surface. S. mutans produces three glucosyltransferases (GTFs), GTFB, GTFC, and GTFD (Colby and Russell, 1997), whose cooperative action is essential for cellular adhesion to the tooth surface. Adhesive glucans mediate attachment of bacteria to the tooth surface as well as to other bacteria; thus, they also have an effect on the physiological status of plaque biofilm, and contribute to the cariogenicity of S. mutans.

GTFB and GTFC, which mainly synthesize water-insoluble glucans, are encoded by the gtfB and gtfC genes, respectively, and are located on the cell surface. On the other hand, GTFD, which synthesizes water-soluble glucan, has been found to be released into culture supernatant and is encoded by the gtfD gene (Kuramitsu, 1993). Analyses of these gtf genes have shown that these enzymes are closely related and have common structures (Monchois et al., 1999b), and their products exhibit immunological cross-reactivity (Fujiwara et al., 1992). Thus, it is difficult to differentiate these genes and their products genetically or immunologically. Moreover, since the gtfB and gtfC genes are tandemly arranged in the genome, it seems likely that regulation of the respective genes has strong interaction. Therefore, investigation of the expression of individual gtf genes has been difficult because of the complex interactions involved.

Based on the hypothesis that gtfB and gtfC are members of a single operon, investigations of gtf promoters have been performed with the use of promoterless chloramphenicol acetyltransferase reporter gene fusion strains of S. mutans (Hudson and Curtiss, 1990; Wexler et al., 1993). Thereafter, evidence that both gtfB and gtfC are independently expressed was presented by Smorawinska and Kuramitsu (1995) and Fujiwara et al. (1996). Recently, a difference between the gtfB and gtfC promoters was reported with the use of a plasmid-based luciferase reporter assay (Goodman and Gao, 2000). However, little is known regarding the dynamic process of gtf expression in sucrose-dependent cellular adhesion, and no distinctive transcriptional analysis of these gtf genes in an S. mutans growing cell has been previously reported.

Real-time PCR with SYBR Green double-stranded DNA binding dye offers a sensitive, efficient, and reliable approach for the quantitation of RNA/DNA. The purpose of the present study was to examine the differentiation and quantitation of mRNA expression of the gtf genes of S. mutans MT8148, and also to analyze the effects of growth phase, pH, and sucrose presence on the expression of each.

	MATERIALS & METHODS

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Bacterial Strains and Culture Conditions
S. mutans MT8148 (serotype c), used in the present study, was cultured in Brain heart infusion (BHI) broth (Difco, Detroit, MI, USA). To determine the effect of culture medium pH, we cultured the organism in BHI supplemented with 25 mM phosphate buffer (pH 5.0, 6.0, or 7.0). The organism was also cultured with or without 2% sucrose to an optical density of 0.2 at 550 nm.

Extraction of Total RNA from S. mutans
S. mutans MT8148 was grown at 37°C to an optical density of 1.0 at 550 nm. The cell suspension (1 mL) was inoculated into 100 mL of fresh broth and incubated at 37°C. When the culture reached optical densities of 0.2 (early-exponential phase), 0.5 (mid-exponential phase), and 1.0 (late-exponential phase), the pH of supernatant was measured, and cells were collected by centrifugation from 50, 20, and 10 mL of the culture, respectively. RNA samples were extracted by a hot-phenol method (Shaw and Clewell, 1985) with some modifications. Briefly, after the cells were washed with phosphate-buffered saline (pH 7.4), an equivalent number of cells (determined at an optical density of 1.0 at 550 nm) from each stage was collected by centrifugation. The cells were treated with 0.5 mg/mL lysozyme (Wako, Osaka, Japan) and then frozen in liquid nitrogen. Bacterial pellets were thawed on ice and digested with 100 µg/mL Proteinase K (Merck, Darmstadt, Germany). The lysate was extracted twice with Tris-buffered phenol (pH 8.0) at 65°C for 3 min. After centrifugation, the aqueous phase was collected and re-extracted with acid phenol:chloroform:isoamyl alcohol. The nucleic acid was precipitated by ethanol, and suspended in diethyl pyrocarbonate (DEPC; Sigma, St. Louis, MO, USA) treated water. Crude RNA was further purified by means of an RNeasy RNA isolation column (QIAGEN, Hilden, Germany) with digestion of RNase-free DNaseI (QIAGEN), as recommended by the manufacturer. Purified RNA was extracted from the column with 100 µL of DEPC-treated water and stored at -80°C.

Quantitation of DNA and RNA
DNA and RNA amounts were determined with the use of a PicoGreen double-stranded DNA quantitation kit (Molecular Probes, Eugene, OR, USA) and a RiboGreen RNA quantitation kit (Molecular Probes) with a Fluorometer (RF-5300PC Shimadzu, Kyoto, Japan), respectively.

Primer Design
The outline of real-time quantitative RT-PCR and primers used in this study is shown in Fig. 1. The first-strand synthesis of cDNA was primed by means of gene-specific primers. The primers RT-B1117 (5'-cataaggcgttaatttcccttca-3'), RT-C1195 (5'-cctgtgaagttagcttgctattg-3'), and RT-D1164 (5'-ataggctgtcttatcgctgttgcta-3') were designed corresponding to the 5' region of the genes encoding the catalytic domain from the gtfB (GenBank Acc. No. D88651), gtfC (D88652), and gtfD (D88653) genes of S. mutans MT8148, respectively. The primer sets B442f (5'-agcaatgcagccaatctacaaat-3') and B537r (5'-acgaactttgccgttattgtca-3'), C236f (5'-ctcaaccaaccgccactgtt-3') and C326r (5'-ggtttaacgtcaaaattagctgtattagc-3'), and D434f (5'-cacaggcaaaagctgaattaaca-3') and D514r (5'-gaatggccgctaagtcaacag-3') were designed for PCR amplification, corresponding to the hyper-variable region 580-870 bp upstream of the RT primers. The expected sizes for each PCR product from gtfB, gtfC, and gtfD were 98 bp, 93 bp, and 83 bp, respectively. These primers were designed according to the manufacturer's guidelines and with the help of PrimerExpress software (PE Applied Biosystems, Foster City, CA, USA). The primer set RecA/F1 (5'-ccggaatcttctggtaag-3') and RecA/R1 (5'-ctaattcacctgtacgag-3'), corresponding to the recA gene of S. mutans (Acc. No. M61897), was designed to compare with the expression of the housekeeping gene.

View larger version (42K): [in this window] [in a new window]   Figure 1. Outline of and primers for real-time quantitative RT-PCR. The target for PCR was the 5' one-third region of the gtf genes shown at the top. Each gtf gene-specific primer (RT primer) corresponding to the 5' region of the catalytic domain (A) was used for the first-strand synthesis of cDNA (B). Real-time PCR was performed with the PCR primers corresponding to the hyper-variable region of each gtf gene (C). The expected sizes for each PCR product from gtfB, gtfC, and gtfD were 98 bp, 93 bp, and 83 bp, respectively (D).

Southern Blot Analysis
Southern blot analysis of real-time PCR amplicons was carried out as described previously (Fujiwara et al., 2000). After amplification, PCR products were applied to 2.0% agarose gel electrophoresis. To detect the contamination of genomic DNA in the extracted RNA samples, we performed PCR with each gtf-specific primer, using the RNA samples, without an RT reaction as a template. The recombinant plasmids pSK6, pSK16, and pYT104 carrying the gtfB, gtfC, and gtfD genes, respectively (Fujiwara et al., 1998), were digested with Sph I, Sph I, and Pst I, respectively, and then used as positive controls. Probes were amplified with the same primers as in the real-time PCR from chromosomal DNA of S. mutans MT8148. These probes were labeled with ³²P and hybridized in stringent conditions.

Statistical Analysis
Inter-group differences of various factors were estimated by a statistical analysis of variance (ANOVA) for factorial models. Fisher's protected least-significant difference test was used to compare individual groups.

	RESULTS

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All isolated RNA samples contained negligible amounts of double-stranded DNA. The amounts of total RNA and the pH values of each growth phase culture are shown in Fig. 2. The RNA yield was inversely proportional to the growth phase. An equal amount of total RNA (1 µg) from each phase culture was used for quantification of the transcript levels of the gtf genes. Using RT-PCR, we observed no significant difference in the expression of the recA gene from each sample (data not shown).

View larger version (49K): [in this window] [in a new window]   Figure 2. Total RNA and relative quantities of gtfB, gtfC, and gtfD mRNA in the growth phase of S. mutans MT8148. Amounts of extracted total RNA were assessed by means of a RiboGreen RNA quantitation kit with a Fluorometer. Following reverse transcription from 1 µg of total RNA, the amount of each gtf cDNA was determined by real-time PCR. Data are expressed as means and standard deviations of triplicate experiments. Statistical differences (P < 0.05, ANOVA) from early-exponential (*) and mid-exponential (#) phases are indicated.

Each gtf-specific amplification was also confirmed by Southern blot analysis (Fig. 3). The real-time RT-PCR amplicon hybridized with each specific gtf probe; however, no cross-hybridization among the other gtf genes was detected. Further, no gtf-specific PCR product was amplified from the template without an RT reaction.

View larger version (25K): [in this window] [in a new window]   Figure 3. Southern blot analysis of real-time RT-PCR products. PCR products with/without reverse transcription (RT) reaction and recombinant plasmids carrying each gtf gene were separated by 2% agarose gel electrophoresis, then transferred onto nylon membranes. The membranes were hybridized with 32P-labeled gtfB-, gtfC-, and gtfD- specific probes, respectively. Lanes 1, RT-PCR amplicon by gtfB-specific primers; 2, RT-PCR amplicon by gtfC-specific primers; 3, RT-PCR amplicon by gtfD-specific primers; 4, PCR product without RT reaction by gtfB-specific primers; 5, PCR product without RT reaction by gtfC-specific primers; 6, PCR product without RT reaction by gtfD-specific primers; 7, pSK6 carrying gtfB; 8, pSK16 carrying gtfC; and 9, pYT104 carrying gtfD.

View larger version (22K): [in this window] [in a new window]   Figure 4. Effects of pH (A) and sucrose (B) on the expression of gtf genes. S. mutans MT8148 was cultured in BHI broth with 25 mM phosphate buffer (A) with or without 2% sucrose (B) at 37°C to an optical density of 0.2 at 550 nm. Data are expressed as means and standard deviations of triplicate experiments. Statistical differences (P < 0.05, ANOVA) between pH 5 (*), pH 6(#), and pH 7 () (A), and without sucrose (*) (B) are indicated.

	DISCUSSION

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This is the first known report to describe differential and quantitative analyses of transcriptions of S. mutans gtf genes directly from growing cells. Real-time PCR is advantageous for the measurement of levels of specific RNAs of the growing bacteria in terms of sensitivity and specificity. The traditional methods, including Northern blotting, generally require a large amount of RNA. In addition, since the coding region of the gtf genes is approximately 5000 bp, it is difficult to isolate whole-size mRNA thoroughly; therefore, no reliable Northern blot analysis could be performed. However, in the present real-time RT-PCR experiments, each gtf transcription could be detected from a 1200-bp 5' fragment, which is one of the advantages of our real-time RT-PCR system.

Sequence analysis revealed that the S. mutans gtf genes possess highly conserved regions: (i) a 5' signal sequence, (ii) a 5' catalytic domain, and (iii) a 3' glucan binding domain (Monchois et al., 1999b). Since there is a high degree of similarity between the gtf genes, genetic differentiation of each has been difficult. However, it has been reported that there is no conservation of the primary sequence in a portion located approximately 400 bp between the signal and the catalytic region of GTFs from oral streptococci and Leuconostoc mesenteroides (Monchois et al., 1999a). In this study, we compared the multiple alignment of the three gtf genes from S. mutans, and found that this portion was also hyper-variable, which is useful when designing specific primers for differentiation of each gene. Furthermore, primers to initiate RT reactions were also designed based on the specific 5' sequence of the gene encoding the catalytic region (Fig. 1). Using these primer sets, we successfully differentiated gtfB, gtfC, and gtfD by real-time RT-PCR with SYBR Green (Figs. 2, 4), and the specificity of the PCR products was confirmed with Southern blot analysis (Fig. 3).

Previous studies using selective inactivation of the gtf genes have revealed that GTFC plays the most important role in firm sucrose-dependent adhesion (Fukushima et al., 1992; Fujiwara et al., 1996; Tsumori and Kuramitsu, 1997). However, it is necessary to analyze the synthesis of individual GTFs in terms of temporal, quantitative, and qualitative aspects. While heat-killed S. sobrinus cells incubated simultaneously with GTF-Sa, -Sb, and -I in sucrose-containing buffer adhering firmly to a glass surface (Koga et al., 1986), sucrose-dependent firm adhesion of S. mutans cells has been reproduced only in growing cells. Recently, using recombinant GTFs (rGTFs) and resting cells of S. mutans, we successfully reconstituted firm adhesion of the cells, and determined that the adhesive glucan produced by rGTFC in the presence of a constant ratio of rGTFD was essential in the initiation of the sucrose-dependent cellular adhesion of S. mutans. The highest level of sucrose-dependent adherence was found at a ratio of 20 GTFB:1 GTFC:4 GTFD (Ooshima et al., 2001). In the present study, the mRNA expression ratios of gtfB:gtfC:gtfD in the early-, middle-, and late-exponential phases were 16:1:10, 15:1:12, and 69:1:25, respectively, while the expression level of each gtf gene was in general accord with results from our previous study, except for the late-exponential phase. The relative amount of gtfD expression was higher than that of rGTFD in the sucrose-dependent adhesion experiment. This difference may be accounted for by the difference of localization between GTFC and GTFD, since GTFC is cell-associated, and GTFD is found in the culture supernatant. The secreted GTFD could be diluted with secreted saliva; thus, a greater expression of gtfD may be required in the establishment of cellular adhesion in the oral cavity. The prominent production of GTFB seen in the late-exponential phase suggested that the initial adhesion may already have been established by GTFC and GTFD in an earlier stage, and that GTFB may play a role in reinforcing the adhesion by the large amount of insoluble glucan produced by it during this stage.

It is of interest that the effects of growth conditions on the expression patterns of gtfB, gtfC, and gtfD were different. As growth proceeded, the gtfB transcript prominently changed, whereas that of gtfD did not (Fig. 2), while gtfD was expressed in greater amounts under acidic conditions than either gtfB or gtfC (Fig. 4A). Further, when the culture pH was kept constant at 6.0, the expression of gtfB decreased in the late-exponential phase, whereas that of gtfD increased, indicating that the expression of gtfD was more dependent on culture pH than during the growth phase. These results also suggest that the promoters of each gtf gene may be different and confirm previous reports that have noted that both gtfB and gtfC are independently expressed (Smorawinska and Kuramitsu, 1995; Fujiwara et al., 1996). In the presence of sucrose, the expression of gtfB and gtfC mRNA decreased (Fig. 4B); however, investigators using the promoterless chloramphenicol acetyltransferase gene integrated into the chromosomal gtfB gene have reported that the expression of the S. mutans gtfB/C operon was stimulated in the presence of sucrose (Hudson and Curtiss, 1990). On the other hand, a plasmid-based reporter system using a luciferase assay has revealed that the expressions of the upstream regions of gtfB and gtfC remained constant with the presence of sucrose, glucose, and fructose (Goodman and Gao, 2000). The gene that regulates gtf in sanguinis streptococci, rgg, has been reported (Sulavik and Clewell, 1996); however, the gtf regulating gene in S. mutans remains unknown. Therefore, further investigation is required to explain these differences.

In conclusion, a real-time RT-PCR was carried out for differential analyses of mRNA of each gtf gene from S. mutans. The expression ratio of these gtf genes was found to coincide with the optimal ratio that induced sucrose-dependent cellular adhesion.

	ACKNOWLEDGMENTS

 
We thank Dr. Yutaka Terao (Osaka University Graduate School of Dentistry) for providing the recA primers. This work was supported in part by a grant-in-aid from the Ministry of Education, Science and Sports of Japan (11470451 and 13470449).

Received July 30, 2001; Last revision December 6, 2001; Accepted December 12, 2001

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