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Antimicrobial Agents and Chemotherapy, February 2006, p. 542-546, Vol. 50, No. 2
0066-4804/06/$08.00+0     doi:10.1128/AAC.50.2.542-546.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

Influence of Apigenin on gtf Gene Expression in Streptococcus mutans UA159

Hyun Koo,^1* Jennifer Seils,¹ Jacqueline Abranches,² Robert A. Burne,² William H. Bowen,¹ and Robert G. Quivey Jr.¹

Eastman Department of Dentistry and Center for Oral Biology, University of Rochester Medical Center, Rochester, New York,¹ Department of Oral Biology, College of Dentistry, University of Florida, Gainesville, Florida²

Received 12 September 2005/ Returned for modification 2 November 2005/ Accepted 21 November 2005

Top Abstract Introduction Materials and Methods Results and Discussion References

 
Apigenin, a potent inhibitor of glucosyltransferase activity, affects the accumulation of Streptococcus mutans biofilms in vitro by reducing the formation of insoluble glucans and enhancing the soluble glucan content of the polysaccharide matrix. In the present study, we investigated the influence of apigenin on gtfB, gtfC, and gtfD expression in S. mutans UA159. Apigenin (0.1 mM) significantly decreased the expression of gtfB and gtfC mRNA (P < 0.05); in contrast, it increased the expression of gtfD in S. mutans growing in the planktonic state. The protein levels of GTF B, GTF C, and GTF D in culture supernatants were also affected; less GTF B and C were detected, whereas the level of GTF D was significantly elevated (P < 0.05). A similar profile of gtf expression was obtained with biofilms, although an elevated concentration (1 mM) of apigenin was required to elicit the effects. The influence of apigenin on gtf gene expression was independent of any effect on GTF activity, did not involve inhibition of growth or effects on pH, and was not affected by addition of sucrose. The data show that apigenin modulates the genetic expression of virulence factors in S. mutans.

Top Abstract Introduction Materials and Methods Results and Discussion References

 
The ability of Streptococcus mutans to synthesize extracellular glucans is a critical virulence factor involved in the pathogenesis of dental caries in animals and humans (19, 22, 26). This bacterium harbors three distinct gtf genes expressing glucosyltransferase (GTF) activity (17). The gtfB and gtfC genes are in an operon-like arrangement and encode enzymes that produce mostly water-insoluble -(13)-linked glucans, whereas the gtfD gene, which is not linked to the gtfBC locus, encodes an enzyme that synthesizes water-soluble -(16)-linked glucans.

Glucans provide binding sites for, and promote accumulation of, cariogenic streptococci (and other oral microorganisms) on the tooth surface, and they contribute to the establishment of the extracellular polysaccharide matrix, which provides bulk and structural integrity to dental biofilms (known as dental plaque) (4). By preventing glucan production, therapeutic approaches to influencing the formation or virulence of dental biofilms related to caries could be precise and selective and would not necessarily suppress the resident oral flora. Mutant strains of S. mutans defective in the gtf genes, especially gtfB and gtfC, are far less cariogenic than the parent strains in vivo (26). Therefore, agents that affect the expression of gtf genes offer a very attractive route for prevention of dental biofilm-related diseases, such as dental caries.

Apigenin, a nontoxic bioflavonoid ubiquitously found in plant-derived foods and propolis (a honeybee product), is a novel anticaries and antiplaque agent in vitro and in vivo (13, 14, 15). Apigenin effectively disrupted the formation and accumulation of S. mutans UA159 biofilms without killing the organism (15). Analysis of the polysaccharide content of the matrix from a biofilm revealed that apigenin remarkably diminished the amounts of insoluble glucans but increased the total amount of soluble glucans (15). Furthermore, topical application of the compound exhibited cariostatic properties in rats without detectable effects on the viability of either S. mutans or the total cultivable flora population (14). The anticaries effect is related to its exceptional inhibitory effects on GTFs, especially GTF B and C, irrespective of whether the enzymes were exposed to the agent before or after adsorption to a hydroxyapatite (HA) surface (13, 14).

We have observed that apigenin may affect the production/expression of GTFs by S. mutans; planktonic cells of S. mutans UA159 growing in the presence of 0.1 mM apigenin displayed lower GTF B protein levels in the culture supernatant than the same microorganisms growing in the absence of the agent (H. Koo, J. Seils, B. Schobel, and W. H. Bowen, unpublished data). These observations suggest that apigenin may affect the synthesis of the enzymes in addition to its direct effects on enzyme activity. In the present study, we investigated whether apigenin affects the expression of gtf genes in planktonic cells and biofilms of S. mutans UA159.

Top Abstract Introduction Materials and Methods Results and Discussion References

 
Test agent and bacterial strain. Apigenin was obtained from Extrasynthese Co. (Genay-Sedex, France); the compound was verified by means of high-performance liquid chromatography and gas chromatography-mass spectrometry as standard procedures performed by the company for purity (99%) and authenticity. Apigenin was dissolved in ethanol-dimethyl sulfoxide (DMSO) (1:1, vol/vol) just prior to performance of the assays. For this study, we tested 0.1 mM (against planktonic cells) and 1 mM (against biofilms) of apigenin; these concentrations were chosen based on data from our previously published and unpublished dose-response and animal studies (13, 14, 15).

Streptococcus mutans UA159 (serotype c), a proven virulent cariogenic dental pathogen and the strain selected for genomic sequencing (3), was used in this study. The bacterium, either in planktonic cells or in biofilms, was grown in ultrafiltered (Prep/Scale; Millipore, MA) tryptone-yeast extract broth (2.5% tryptone and 1.5% yeast extract, pH 7.0) with 1% glucose (or 1% sucrose) at 37°C under 5% CO₂ (15).

Treatment of planktonic cells and biofilms. The experimental design is diagramed in Fig. 1. Planktonic cells of S. mutans were grown in the presence of 0.1 mM apigenin (or a vehicle control, consisting of 2.5% ethanol and 0.75% DMSO [vol/vol]) at different culture growth phases (experimental design I); bacterial growth was monitored by measuring optical density at 600 nm (OD₆₀₀). Briefly, S. mutans cells were grown to early-exponential phase (OD₆₀₀, 0.2; pH 6.8; 2 x 10⁸ CFU/ml), and at this point apigenin (or the vehicle control) was added to the culture supernatant. When the culture reached mid-exponential phase (OD₆₀₀, 0.5; pH 6.4; 1.7 x 10⁹ CFU/ml) or late-exponential phase (OD₆₀₀, 1.0; pH 5.7; 2 x 10¹⁰ CFU/ml), the cell suspension was immediately processed for RNA extraction (8). In the second experiment (experimental design II), the planktonic cells were grown to mid- or late-exponential phase. At each of the growth phases, apigenin (or the vehicle control) was added to the cell suspension. After a 1-min (t_{1 min}) or 30-min (t_{30 min}) exposure, the treated bacterial suspension (and control) was immediately processed for RNA extraction.

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FIG. 1. Experimental designs for the real-time RT-PCR assays.

Biofilms of S. mutans were formed on saliva-coated hydroxyapatite disks as described elsewhere (8, 15). S. mutans cells were grown undisturbed in a culture medium containing saliva-coated HA disks mounted vertically for 24 h to allow initial biofilm formation on the surface of HA. The 24-h-old biofilms were exposed for 1 min to either 1 mM apigenin or a vehicle control (25% ethanol and 1.25% DMSO) (experimental design III). The biofilms were either harvested (t_{1 min}) or incubated for an additional 30 min (t_{30 min}) at pH 5.0 or 7.0. The treated biofilms (t_{1 min} or t_{30 min}) were immediately immersed in RNALater according to the manufacturer's protocols (Ambion, Inc., Austin, TX).

Viability and pH determination assays. In order to determine whether the addition of apigenin affects the growth or the acidogenicity of S. mutans, the viability and pH of the culture supernatants or biofilms were determined for each of the experiments. For viability assays, an aliquot of the culture supernatants or of a homogenized (sonicated) biofilm suspension was used for determination of the CFU (14, 15). The potential for drug carryover to produce falsely low viability counts was minimized by dilution of inocula and plating of small volumes of diluted samples (50 µl). The in situ pH of the biofilms was measured by placing the tip of a Beetrode pH electrode (World Precision Instruments, New Haven, CT) into the matrix of the biofilms (18).

Extraction of RNA and real-time RT-PCR. The RNA extraction and reverse transcriptase PCR (RT-PCR) conditions and primers were similar to those described by Fujiwara et al. (10) with some modifications (8). Briefly, crude RNA from either planktonic cells or biofilms of S. mutans was extracted and purified using a QIAGEN RNeasy RNA isolation column followed by digestion with RNase-free DNase I according to the manufacturer's instructions (QIAGEN Sciences, MD). cDNAs were synthesized using an iScript cDNA synthesis kit (Bio-Rad Laboratories, Inc., CA). To check for DNA contamination, purified total RNA without reverse transcriptase served as a negative control. The resulting cDNA and negative control were amplified by a MyiQ real-time PCR detection system with iQ SYBR Green supermix (Bio-Rad Laboratories, Inc., CA). The critical threshold cycle (C_T) was defined as the cycle at which the fluorescence becomes detectable above the background and is inversely proportional to the logarithm of the initial number of template molecules. A standard curve was plotted for each primer set as described elsewhere (10, 27). The standard curves were used to transform C_T values to the relative number of cDNA molecules. Relative expression was calculated by normalizing each gtf gene of the treated cells to the 16S rRNA gene (internal control). These values were then compared to those from the vehicle-treated control to determine the change in gtf gene expression (2, 25).

ELISA and GTF activity assays. The levels of GTF B, C, and D in the culture supernatant of S. mutans growing in the presence of apigenin (see experimental design I in Fig. 1) were detected by enzyme-linked immunosorbent assays (ELISA) using polyclonal anti-GTF B, anti-GTF C, and anti-GTF D rabbit sera as described elsewhere (12, 16). An ELISA plate was coated with the culture supernatant. The plates were washed, blocked, and probed with the anti-GTF sera by use of an ELISA kit (Kirkegaard & Perry, Gaithersburg, MD) as described by Voller et al. (23) and Clark and Engval (9). Controls included samples that contained the culture medium only (negative control) or individual purified GTFs (positive controls). The plates were then developed using the kit substrate according to the manufacturer's protocols. The GTFs were detected at 405 nm using an ELISA plate reader.

The GTF activity in culture supernatants was measured by incorporation of [¹⁴C]glucose from labeled sucrose (New England Nuclear Research Products, Boston, MA) into glucans; the water-soluble and water-insoluble glucans were collected, and levels were determined separately as described elsewhere (13, 15). One unit of enzyme activity was defined as the amount of enzyme needed to incorporate 1 µmol of glucose into glucan over a 4-h reaction period. The total GTF activity for synthesis of water-insoluble and water-soluble glucans was measured.

Statistical analysis. The data from real-time PCR, ELISA, and GTF activity assays were assessed by analysis of variance in the Tukey-Kramer honest standard deviation test for all pairs. Triplicates from at least two separate experiments were used in each of the assays. JMP statistical software, version 3.1 (20), was used to perform the analyses. The level of significance was set at 5%.

Top Abstract Introduction Materials and Methods Results and Discussion References

 
Apigenin at a concentration of 0.1 mM had little effect either on the growth of planktonic cells of S. mutans UA159, even up to the late-exponential phase, or on culture pH values (data not shown). Furthermore, by using real-time PCR, we observed no significant differences in the expression of 16S rRNA between cells grown in the presence of apigenin and those grown with a vehicle control. However, expression of gtfB and gtfC was decreased by more than 50%, whereas the level of gtfD mRNA was increased by 45%, in the mid- and late-exponential phases for S. mutans cells growing in the presence of apigenin (P < 0.05) (Fig. 2). The expression of gtf genes can be induced by sucrose (18, 24); we therefore examined whether addition of 30 mM sucrose affected the ability of apigenin to interfere with their expression profile. The addition of sucrose had no effect on the expression of gtf genes in the presence of apigenin. It is noteworthy that the profile of inhibition of expression of gtf genes is consistent with their arrangement in the S. mutans chromosome: both gtfB and gtfC can be cotranscribed and subjected to the same regulatory mechanisms, whereas gtfD is not linked to the gtfBC loci. Moreover, gtfD is regulated in a manner opposite that of gtfB and gtfC (24), suggesting that the induction of gtfD expression could be a result of inhibition of gtfB and gtfC.

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FIG. 2. Real-time PCR analysis of gtfB, gtfC, and gtfD gene expression by S. mutans growing in the presence of 0.1 mM apigenin. The mRNA level of each gene in each sample was normalized to that of 16S rRNA. These values were then compared to those from vehicle-treated controls to determine the change (n-fold) in gtf gene expression. Expression in treated cells was normalized to that in vehicle-treated controls (corresponding to an arbitrary value of 1). Data are expressed as means ± standard deviations of triplicates from at least three separate experiments; values marked with an asterisk are significantly different from that for the vehicle control at 5% significance (Tukey test).

The influence of apigenin on gtfBCD transcriptional levels of S. mutans also affected the levels of GTF B, C, and D in the culture supernatant. ELISA using anti-GTF B, anti-GTF C, and anti-GTF D rabbit sera indicated that significantly less GTF B and GTF C, and higher levels of GTF D, were present in the culture supernatant of S. mutans grown in the presence of apigenin than for cells grown with the vehicle control (P < 0.05). The levels of GTF B and C in the culture supernatant of apigenin-grown cells were 31 to 55% lower than those for cells grown with the vehicle control, whereas the level of GTF D was 40 to 48% higher than that for the control. GTF activity in the culture supernatant was also examined. The total amount of insoluble glucan synthesized by culture supernatants of cells grown in the presence of apigenin was remarkably reduced (70 to 80% less than that with the vehicle control [P < 0.05]); in contrast, the total amount of soluble glucan was either unaffected (in the mid-exponential phase) or slightly greater (20% more than that with the vehicle control in the late-exponential phase [P < 0.05]).

Apigenin inhibits the activities of GTF B and C and, less effectively, that of GTF D (14); it also affects the activity of Streptococcus sanguinis GTF and S. mutans fructosyltransferases (15). By affecting the activities of GTF B and C and the expression of their genes, apigenin effectively inhibited insoluble-glucan synthesis. Even though gtfD gene expression is enhanced, apigenin still reduces GTF D enzyme activity. However, the net result is still enhanced production of soluble glucans.

In general, therapeutic agents remain in the mouth for a very short period (30 s to a few minutes). Therefore, we examined whether short-term exposure of S. mutans cells to apigenin at specific growth phases (mid- and late-exponential phase) has any influence on the profile of gtf gene expression. As shown in Fig. 3, the mRNA levels of gtfB and gtfC in S. mutans cells at mid- and late-exponential phase were reduced after a 1-min exposure to 0.1 mM apigenin compared to levels in cells exposed to a vehicle control (P < 0.05); a similar profile of gtfBC gene expression was observed 30 min after exposure to the test agent. In contrast, the expression of gtfD was increased by apigenin treatment 30 min after exposure to the test agent at late-exponential phase (P < 0.05).

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FIG. 3. Influence of apigenin on gtfB, gtfC, and gtfD gene expression by S. mutans UA159 at different growth phases. The mRNA level of each gene in each sample was normalized to that of 16S rRNA. These values were then compared to those from vehicle-treated controls to determine the change (n-fold) in gtf gene expression. Expression in treated cells was normalized to that in vehicle-treated controls (corresponding to an arbitrary value of 1). Data are expressed as means ± standard deviations of triplicates from at least three separate experiments; values marked with an asterisk are significantly different from that for the vehicle control at 5% significance (Tukey test).

S. mutans occurs in the mouth primarily in the form of biofilms. This bacterium has the ability to adhere to the tooth surface by several mechanisms, including high-affinity adhesins and binding sites in the glucans (7, 21). The synthesis of glucans is critical both for the adherence of the organisms to the tooth surfaces and for their accumulation and persistence (21). Thus, we assessed whether apigenin influences the expression of gtf genes in early-formed S. mutans biofilms. Although our monospecies biofilm model does not mimic the complex microbial community found in dental plaque, it is advantageous for examining specific actions of test agents on S. mutans physiology and genetics, especially on the glucan-mediated processes involved in the formation of a matrix in a biofilm. An elevated concentration of apigenin (1 mM) was used for biofilms because of their higher biomass densities and in view of previous findings that the biofilms are some 5 to 10 times less sensitive to apigenin than cells in suspensions (H. Koo, J. A. Cury, J. Abranches, R. E. Marquis, and W. H. Bowen, Abstr. 80th Gen. Sess. Int. Assoc. Dent. Res., abstr. 3810, 2002). Apigenin at 1 mM had no detectable effect on the bacterial viability and in situ pH of the biofilms, even as long as 30 min after exposure to the agent (data not shown). Nevertheless, the biofilms of S. mutans UA159 treated with apigenin for 1 min showed significant reductions in the expression of all gtf genes, especially gtfB and gtfC, compared to that in vehicle-treated biofilms (P < 0.05) (Fig. 4). In addition, the treated biofilms were incubated for an additional 30 min in the culture medium at either pH 5.0 or pH 7.0. The expression of gtfB and gtfC remained repressed by apigenin at levels similar to those observed after a 1-min exposure to the agent (Fig. 4). However, the levels of gtfD mRNA in apigenin-treated biofilms were increased compared to those in vehicle-treated biofilms 30 min after exposure, which could explain the elevated soluble-glucan content in the biofilm's matrix (15). In general, effects were similar to those observed with planktonic cells.

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FIG. 4. Influence of apigenin on gtfB, gtfC, and gtfD gene expression in S. mutans UA159 biofilms. The mRNA level of each gene in each sample was normalized to that of 16S rRNA. These values were then compared to those from vehicle-treated controls to determine the change (n-fold) in gtf gene expression. Expression in treated cells was normalized to that in vehicle-treated controls (corresponding to an arbitrary value of 1). Data are expressed as means ± standard deviations of triplicates from at least three separate experiments; values marked with an asterisk are significantly different from that for the vehicle control at 5% significance (Tukey test).

The mechanisms involved in regulation of the genes encoding GTF enzymes appear to be complex and still remain to be fully explored. There are several factors that can influence transcription and translation of the gtf genes, such as carbohydrate availability and source, environmental pH, and growth rate/phase (6, 10, 11, 18, 24). For example, lowering of the pH following addition of carbohydrates, an increase in the availability of metabolizable carbohydrates sensed via the phosphotransferase system (PTS), or production of glycolytic intermediates may regulate the expression of gtf genes. The gtfBC genes are induced in response to acidification of the biofilms or in response to the presence of an excess of a metabolizable carbohydrate (glucose or sucrose) (18). In addition, a cell-density-dependent component could be regulating gtfBC expression (11). It is apparent that the influence of apigenin on gtf gene expression is not a result of an indirect effect on S. mutans physiology, such as growth rate and acid production, or environmental pH and carbohydrate availability. However, the possibility that apigenin affects the PTS (which regulates gtf gene expression) cannot be excluded; inactivation of the gene encoding EIIAB^man of the PTS of S. mutans decreases gtfBC expression (1).

It has been shown that the expression of gtf genes involves RegM, the catabolite-repression regulator (5). Inactivation of regM resulted in dramatic decreases in the levels of expression of chloramphenicol acetyltransferase under the control of the gtfBC promoter. RegM is critical for optimal expression of ftf and gtfBC. Furthermore, LuxS-based signaling appears to be involved in the formation of S. mutans biofilms by mediating the expression of gtfB and gtfC but not gtfD. The expression of gtfB and gtfC by an S. mutans luxS mutant was enhanced relative to that by the parental strain, whereas gtfD expression was unaffected (28). Whether luxS, regM, and the PTS are potential targets for apigenin awaits further investigation.

The effects of apigenin on the expression of genes involved in extracellular polysaccharide synthesis are clearly shown in this study. Our findings regarding the influence of apigenin on the expression of the gtf genes are pertinent to a fuller understanding of the physiology, virulence, and therapeutics of oral streptococci in the mouth. Apigenin is a unique potentially therapeutic compound that affects both the activity and the expression of GTF enzymes without displaying antibacterial effects; these activities may be responsible for the caries-reducing properties of apigenin. The observation that apigenin had an immediate effect on gene expression is encouraging, given the difficulties of maintaining therapeutic agents in the mouth at levels sufficient to gain a therapeutic effect. However, the mechanisms involved in these effects appear to be complex and require exploration. The influence of apigenin on gtf gene expression during the course of biofilm formation and accumulation, and its effects on mature biofilms (5 days old), is being investigated using both monospecies and multispecies models.

 
We are grateful to Matt Betzenhauser and Roberta Faustoferri for technical assistance.

This research was supported by USPHS research grant 1R03 DE015441 and DE12236 from the National Institute of Dental and Craniofacial Research, National Institutes of Health.

 
* Corresponding author. Mailing address: University of Rochester Medical Center, Eastman Department of Dentistry and Center for Oral Biology, 625 Elmwood Ave., Box 683, Rochester, NY 14620. Phone: (585) 273-4216. Fax: (585) 276-0190. E-mail: Hyun_Koo{at}urmc.rochester.edu.

Top Abstract Introduction Materials and Methods Results and Discussion References

 

1
1. Abranches, J., C. C. Chen, and R. A. Burne. 2003. Characterization of Streptococcus mutans strains deficient in EIIAB Man of the sugar phosphotransferase system. Appl. Environ. Microbiol. 69:4760-4769.[Abstract/Free Full Text]
2
2. Adur, J., T. Uchide, S. Takizawa, J. Quan, and K. Saida. 2004. Real-time polymerase chain reaction quantification of gene expression levels of murine endothelin-A and endothelin-B receptors: gene expression profiles by the standard curve method. J. Cardiovasc. Pharmacol. 44:S321-S328.[CrossRef][Medline]
3
3. Ajdic, D., W. M. McShan, R. E. McLaughlin, G. Savic, J. Chang, M. B. Carson, C. Primeaux, R. Tian, S. Kenton, H. Jian, S. Lin, Y. Qian, S. Li, H. Zhu, F. Najar, H. Lai, J. White, B. A. Roe, and J. J. Ferretti. 2002. Genome sequence of Streptococcus mutans UA159, a cariogenic dental pathogen. Proc. Natl. Acad. Sci. USA 99:14434-14439.[Abstract/Free Full Text]
4
4. Bowen, W. H. 2002. Do we need to be concerned about dental caries in the coming millennium? Crit. Rev. Oral Biol. Med. 13:126-131.[Abstract/Free Full Text]
5
5. 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.[CrossRef][Medline]
6
6. Burne, R. A., Y. M. Chen, and J. E. C. Penders. 1997. Analysis of gene expression in Streptococcus mutans in biofilms in vitro. Adv. Dent. Res. 11:100-109.[Abstract/Free Full Text]
7
7. Burne, R. A. 1998. Oral streptococci: products of their environment. J. Dent. Res. 77:445-452.[Abstract/Free Full Text]
8
8. Chatfield, C. H., H. Koo, and R. G. Quivey. 2005. The putative lytR ortholog in Streptococcus mutans plays a role in cell division and is growth-rate regulated. Microbiology 151:625-631.[Abstract/Free Full Text]
9
9. Clark, B. R., and E. Engval. 1980. Enzyme immunoassay. CRC Press, Inc., Boca Raton, Fla.
10
10. Fujiwara, T., T. Hoshino, T. Ooshima, and S. Hamada. 2002. Differential and quantitative analyses of mRNA expression of glucosyltransferases from Streptococcus mutans MT8148. J. Dent. Res. 81:109-113.[Abstract/Free Full Text]
11
11. Goodman, S. D., and Q. Gao. 2000. Characterization of the gtfB and gtfC promoters from Streptococcus mutans GS-5. Plasmid 43:85-98.[CrossRef][Medline]
12
12. Hayacibara, M. F., H. Koo, A. M. Vacca Smith, L. K. Kopec, K. Scott-Anne, J. A. Cury, and W. H. Bowen. 2004. The influence of mutanase and dextranase on the production and structure of glucans synthesized by streptococcal glucosyltransferases. Carbohydr. Res. 339:2127-2137.[CrossRef][Medline]
13
13. Koo, H., P. L. Rosalen, J. A. Cury, Y. K. Park, and W. H. Bowen. 2002. Effects of compounds found in propolis on S. mutans growth and on glucosyltransferase activity. Antimicrob. Agents Chemother. 46:1302-1309.[Abstract/Free Full Text]
14
14. Koo, H., S. K. Pearson, K. Scott-Anne, J. Abranches, J. A. Cury, P. L. Rosalen, Y. K. Park, R. E. Marquis, and W. H. Bowen. 2003. Effects of apigenin and tt-farnesol on glucosyltransferase activity, biofilm viability and caries development in rats. Oral Microbiol. Immunol. 17:337-343.
15
15. Koo, H., M. F. Hayacibara, B. D. Schobel, J. A. Cury, P. L. Rosalen, Y. K. Park, A. M. Vacca Smith, and W. H. Bowen. 2003. Inhibition of Streptococcus mutans biofilm accumulation and polysaccharide production by apigenin and tt-farnesol. J. Antimicrob. Chemother. 52:782-789.[Abstract/Free Full Text]
16
16. Kopec, L. K., A. M. Vacca Smith, D. Wunder, L. Ng-Evans, and W. H. Bowen. 2002. Influence of antibody on the structure of glucans. Caries Res. 36:108-115.[CrossRef][Medline]
17
17. Kuramitsu, K. K. 1993. Virulence factors of mutans streptococci: role of molecular genetics. Crit. Rev. Oral Biol. Med. 4:159-176.[Abstract/Free Full Text]
18
18. 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.[Abstract/Free Full Text]
19
19. Loesche, W. J. 1986. Role of Streptococcus mutans in human dental decay. Microbiol. Rev. 50:353-380.[Free Full Text]
20
20. SAS Institute, Inc. 1989. JMP user's guide: version 2 of JMP. SAS Institute, Cary, N.C.
21
21. 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.[Abstract/Free Full Text]
22
22. Tanzer, J. M., M. L. Freedman, and R. J. Fitzgerald. 1985. Virulence of mutants defective in glucosyltransferase, dextran-mediated aggregation, or dextranase activity, p. 204-211. In S. E. Mergenhagen and B. Rosan (ed.), Molecular basis of oral microbial adhesion. American Society for Microbiology, Washington, D.C.
23
23. Voller, A., D. E. Bidwell, and A. Barlett. 1979. The enzyme-linked immunosorbent assay (ELISA), a guide with abstracts of microplate applications. Dynatech Laboratories, Inc., Alexandria, Va.
24
24. 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.[Abstract/Free Full Text]
25
25. Winer, J., C. K. S. Jung, I. Shackel, and P. M. Williams. 1999. Development and validation of real-time quantitative reverse transcriptase-polymerase chain reaction for monitoring gene expression in cardiac myocytes in vitro. Anal. Chem. 270:41-49.
26
26. 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.[Abstract/Free Full Text]
27
27. Yoshida, A., and H. K. Kuramitsu. 2002. Streptococcus mutans biofilm formation: utilization of a gtfB promoter-green fluorescent protein (PgtfB::gfp) construct to monitor development. Microbiology 148:3385-3394.[Abstract/Free Full Text]
28
28. 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.[Abstract/Free Full Text]

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Antimicrobial Agents and Chemotherapy, February 2006, p. 542-546, Vol. 50, No. 2
0066-4804/06/$08.00+0     doi:10.1128/AAC.50.2.542-546.2006
Copyright © 2006, American Society for Microbiology. All Rights Reserved.

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• Koo, H., Jeon, J. G. (2009). Naturally Occurring Molecules as Alternative Therapeutic Agents against Cariogenic Biofilms. ADR 21: 63-68 [Full Text]  
• Klein, M. I., Duarte, S., Xiao, J., Mitra, S., Foster, T. H., Koo, H. (2009). Structural and Molecular Basis of the Role of Starch and Sucrose in Streptococcus mutans Biofilm Development. Appl. Environ. Microbiol. 75: 837-841 [Abstract] [Full Text]  
• Koo, H. (2008). Strategies to Enhance the Biological Effects of Fluoride on Dental Biofilms. ADR 20: 17-21 [Full Text]  

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