Center for Oral Biology and Department of Microbiology and Immunology, University of Rochester School of Medicine and Dentistry, 601 Elmwood Avenue, Rochester, NY 14642, USA1
Author for correspondence: Robert A. Burne. Tel: +1 352 392 0011. Fax: +1 352 392 7357. e-mail: rburne{at}dental.UFL.EDU
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ABSTRACT |
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Keywords: exopolysaccharides, pathogenesis, glucans, dental caries, dental plaque
Abbreviations: CAT, chloramphenicol acetyltransferase
a Present address: Faculty of Dentistry, University of Toronto, 124 Edward Street, Toronto, Ontario, Canada M5G 1G6.
b Present address: Department of Oral Biology, PO Box 100424, Gainesville, FL 32610-0424, USA.
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INTRODUCTION |
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S. mutans synthesizes glucan polymers from sucrose via the actions of three secreted glucosyltransferases (GTFs), encoded by the gtfB, gtfC and gtfD genes (Kuramitsu, 1993 ). The gtfB and gtfC genes are in an operon-like arrangement and encode enzymes that produce mainly water-insoluble glucans, whereas the gtfD gene, which is not linked to the gtfBC locus, encodes an enzyme that catalyses the formation of a water-soluble glucan. It is the water-insoluble glucans made by GtfBC that play important roles in adhesion and accumulation of the organisms on the tooth surfaces, and in establishing the extracellular polysaccharide matrix that is responsible for the structural integrity of dental biofilms. These polysaccharides are also thought to provide the organisms with a unique microenvironment for their growth, metabolism and survival (Bowden & Hamilton, 1998
; Liljemark & Bloomquist, 1996
; Nakano & Kuramitsu, 1992
; Yamashita et al., 1993
). S. mutans produces a single fructosyltransferase (FTF), the product of the ftf gene, which catalyses the synthesis of high-molecular-mass fructans from sucrose (Ebisu et al., 1975
; Shiroza & Kuramitsu, 1988
). Fructans produced by S. mutans are believed to function primarily as extracellular storage compounds that can be metabolized during periods of nutrient deprivation (Burne et al., 1996
).
A number of studies indicate that the expression of the genes for the exopolysaccharide-synthesizing enzymes of S. mutans is dependent on environmental conditions, including growth rate, pH, carbon source, and whether the organisms are attached to the surfaces (Burne et al., 1997 ; Hudson & Curtiss, 1990
; Kiska & Macrina, 1994
; Wexler et al., 1993
). Also, the use of gtf and ftf gene fusion strains in a continuous-flow biofilm fermenter has shown that organisms growing in thicker, mature (7-d-old) biofilms have higher levels of expression of the gtfBC genes compared with cells grown in suspension or in thinner (2-d-old) biofilms (Burne et al., 1997
; Hudson & Curtiss, 1990
; Kiska & Macrina, 1994
; Wexler et al., 1993
). In contrast, ftf was found to be dramatically down-regulated in 7 d biofilms compared with 2 d biofilms. The specific mechanisms governing regulation of exopolysaccharide synthesis in S. mutans biofilms have yet to be discovered. In this study, we test the hypothesis that pH or changes in carbohydrate source and concentration in biofilms influence expression of gtf and ftf in mature biofilms.
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METHODS |
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Assessment of physical characteristics of S. mutans biofilms.
Biofilm dry weights were determined by mechanically dissociating the biofilms from the slides, collecting the material by centrifugation, washing once with dH2O, lyophilizing the sample and weighing. For measurement of total carbohydrate in biofilms, the lyophilized samples were resuspended in 5 ml dH2O and the anthrone method (Dubois et al., 1956 ) was used to measure total carbohydrate using glucose as the standard. Bacterial viability in biofilms was estimated by direct microscopic enumeration and plate counts with cells prepared as follows. Biofilms were collected, washed once, and resuspended in 5 ml of reduced transport fluid (Loesche & Syed, 1973
) at pH 7·2. The samples were then subjected to a gentle sonication at a setting of 150 W for 20 s to break bacterial chains, and the samples were split into two parts. The cell suspensions for viable cell counts were decimally diluted and directly inoculated by a spiral plating system (Autoplate model 3000; Spiral Biotech) onto BHI agar supplemented with 10 µg erythromycin ml-1. The viable counts were made after the plates were cultivated at 37 °C for 24 h in a 5% CO2 aerobic atmosphere. Microscopic enumeration of bacteria was conducted as described by Koch (1994)
using a PetroffHausser counting chamber. The ratio of viable cell counts to the counts obtained by visual enumeration was expressed as percentage viable cells. The formation and spatial distribution of biofilms on the surfaces were also assessed by using phase-contrast microscopy, as previously described (Burne et al., 1997
; Li et al., 2000
).
In situ measurement of biofilm pH.
The pH of the biofilms was measured as previously detailed (Li et al., 2000 ) using a superminiature, Beetrode pH electrode (model MEPH3L; World Precision Instruments). Briefly, after the cultures reached quasi-steady state (Burne & Chen, 1998
), slides with biofilms were removed from the vessel and placed on end on a paper towel to allow excess medium to be absorbed from the end of the slides. The micro-reference electrode, which was connected through the Bee-Cal adapter and two cables to the pH probe and a standard pH meter, was positioned in the biofilm to be partially immersed in the biomass. In situ measurement of pH was conducted immediately by placing the tip of the pH probe into biofilms and a series of pH readings was recorded from a minimum of 30 different sites selected at random.
Carbohydrate pulsing and biofilm sampling.
Quasi-steady-state biofilms, or biofilms which had been pulsed with either glucose or sucrose (25 mM) immediately after initial biofilm sampling (T0), were used for measurements of chloramphenicol acetyltransferase (CAT) activity expressed from the gtf or ftf promoters. Subsequently, the biofilms were sampled by removing three slides at 15, 30 and 60 min after the carbohydrate pulse. The biofilms were mechanically dissociated from the slides by scraping with a sterile razor blade into 40 ml ice-cold Tris/HCl (10 mM, pH 7·8) and were centrifuged at 8000 g for 10 min at 4 °C. The cell pellets were washed twice and resuspended in 1 ml of the same buffer for the preparation of cell-free lysates. The pH profiles in the liquid phase at each sampling time were recorded by measuring pH in 10 ml of culture fluid.
Preparation of cell-free lysates and CAT assays.
Cell-free lysates were prepared for the analysis of CAT activity as previously described (Chen et al., 1998 ). Briefly, the cell pellets were resuspended in 1 ml Tris buffer in a 2 ml, screw-cap microcentrifuge tube (Sarstedt). One-third volume of pre-chilled (-20 °C) glass beads (0·2 mm mean diameter) was added to the sample, and the cells were homogenized using a Bead Beater (Biospec Products) in four 30 s intervals, with cooling on ice in the intervals. The lysates were centrifuged at 12000 g at 4 °C for 15 min and the supernatant material was collected for CAT assays. Protein concentrations of the cell-free lysates were determined by the method of Bradford (1976)
using a commercially available reagent (Bio-Rad). CAT activity was measured by a spectrophotometric method (Shaw, 1979
) with use of the colorimetric substrate 5,5'-dithio-bis-nitrobenzoic acid (DTNB; Boehringer Mannheim). All assays were performed in triplicate with internal standards containing all reagents except chloramphenicol to account for chloramphenicol-independent reduction of DTNB. One unit of CAT activity was defined as the amount of enzyme needed to catalyse the acetylation of 1 µmol chloramphenicol min-1 (mg protein)-1.
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RESULTS |
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DISCUSSION |
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A number of new findings have emerged from this study. The first is that environmental pH clearly influences the expression of the gtfBC genes of S. mutans growing in biofilms. Biofilms grown without pH control, which had biofilm pH values of around 5·3, consistently had twofold more CAT activity expressed from the gtfBC promoter than SMS102 cultivated with pH control. Moreover, following a carbohydrate pulse, the level of expression of gtfBC in the biofilms without pH control was significantly higher than in biofilms with pH control. Since increased production of GtfBC would likely result in a greater proportion of carbohydrate being incorporated into biofilm polysaccharides, this observation is consistent with the finding that the biofilms formed at low pH were composed of a much greater percentage of carbohydrate than those formed at a more neutral pH value. Also, a previous study (Wexler et al., 1993 ) showed that when S. mutans was grown in steady-state continuous chemostat culture, lowering of the pH from 7 to 6 resulted in about a 50% increase in the expression of the gtfBC genes. Interestingly, the level of expression of gtfBC achieved by suspended populations at steady state or after a sucrose pulse (Wexler et al., 1993
) was lower than the level of expression seen in biofilms at quasi-steady state or after sucrose induction in this study. Thus, one possible explanation for the enhanced gtfBC expression in biofilms may be due to the relatively lower pH values achieved in biofilms (Table 2
), compared to the continuous culture studies, or to the existence of low pH microenvironments in which gtf expression is markedly elevated. A second significant, and possibly related, finding is that gtfBC expression in biofilms was induced by addition of excess glucose to the culture, albeit the level of induction was not as great as that seen with sucrose. There is no logical reason for glucose, which is not a substrate for Gtfs, to act as a specific inducer of gtfBC. Instead, it is more likely that 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. The latter could be signalled by directly sensing carbohydrate flow through the PTS or the glycolytic pathway, or could reflect a response to increased growth rate, although in continuous culture we found gtfBC expression was inversely correlated with how fast the cells were growing (Wexler et al., 1993
). Of note, there does not seem to be a hierarchy of control of expression of gtfBC by carbohydrate and pH, since induction by excess carbohydrate occurs whether the cells are growing with or without pH control. Possibly, then, the effects of pH and carbohydrate are exerted through different pathways. Finally, we cannot exclude the idea that differences in phosphate concentrations in the buffered and unbuffered cultures also had an impact on the results.
Another important finding was that growth of the biofilms under quasi-steady-state conditions with glucose as the limiting carbohydrate did not result in down-regulation of gtf or ftf expression compared to cells growing with sucrose as the limiting carbohydrate. It has been suggested that sucrose is a specific inducer of exopolysaccharide-producing enzymes, but if that were the case, then one might expect higher levels of CAT activity expressed at T0 in cells growing in sucrose (Tables 3 and 5
) versus CAT activities at T0 in biofilms being fed on glucose (Table 4
), which was not the case. It could be that high levels of sucrose are required for induction, although this would not explain why both ftf and gtf can be induced by addition of either glucose or sucrose. These findings emphasize the possibility that lowering of the pH following addition of carbohydrates or that an increase in the availability of carbohydrate, perhaps sensed via the PTS or glycolytic intermediates, are mechanisms by which gtf and ftf gene expression could be regulated. In fact, we now have preliminary evidence (unpublished) from continuous chemostat culture of these strains at different pH values, under carbohydrate-limiting and carbohydrate-excess conditions, that not only pH, but the amount of carbohydrate available to cells is a factor that influences both ftf and gtf gene expression.
The expression pattern of ftf is more complex than that of gtfBC. As with gtf, the expression levels of ftf in quasi-steady-state biofilms are the same regardless of whether glucose or sucrose was the limiting carbohydrate. In contrast, ftf expression in the lower pH biofilms was not enhanced above that in biofilms formed with buffered medium. As seen previously with both suspended (Wexler et al., 1993 ) and adherent (Burne et al., 1997
) populations of S. mutans, sucrose was an efficient inducer of ftf gene expression. In this study, we found that induction by sucrose occurred regardless of whether the cells were growing at the lower or higher pH value, or whether the steady-state biofilms were formed with glucose or sucrose as the limiting carbohydrate. However, addition of glucose to quasi-steady-state biofilms had only a modest effect on induction of ftf. In continuous chemostat culture, increasing the growth rate of strain SMS101, or lowering of the pH from 7 to 6, enhanced ftf transcription (Wexler et al., 1993
), so these factors were likely to have had an effect on ftf transcription after addition of excess carbohydrate. Still, in all cases, the responses of ftf to added carbohydrate, i.e. the rate of induction and the subsequent decline in CAT activity, differed as a function of whether the culture medium was buffered or not. Thus, global factors regulating gene expression in relation to carbohydrate flow or pH may overlap with the regulatory networks that control ftf transcription.
Another notable finding is that expression from the gtfBC promoter in biofilms in this study was higher than that observed in cells grown in continuous chemostat culture (Wexler et al., 1993 ), although not as high as in mature (7 d) biofilms (Burne et al., 1997
). For example, values in the chemostat were reported to range from 0·017 to 0·049 U (mg protein)-1 under steady-state conditions, and levels after sucrose induction were as high as 0·15 U (mg protein)-1. It can be inferred from the results obtained here that the existence of low pH microenvironments and perhaps the presence of extracellular storage polysaccharides that increase the amount of carbohydrate available to subpopulations of cells within the biofilm may account for the apparent stimulation of gtf expression in biofilms (Burne et al., 1997
). Low growth rate microenvironments could also exist and impact gtfBC expression. We also have noted that ftf expression in mature (7 d) biofilms is almost completely repressed. Interestingly, the level of CAT expressed from the ftf gene fusion in quasi-steady-state biofilms (Tables 35
) was about one-third of that in suspended cells in steady-state continuous chemostat culture (Wexler et al., 1993
). Also, following induction of ftf in the chemostat using sucrose, levels of CAT peaked at about 0·45 U (Wexler et al., 1993
), whereas the peak expression noted here was about 0·33 U. So again, the trend is that ftf expression in biofilms is partially repressed compared with suspended populations. Recently, we have also found evidence that growth at pH 5 leads to nearly complete repression of ftf (C. Browngardt & R. A. Burne, unpublished), so low pH in the biofilms may account in part for lowered expression of ftf in biofilms.
Clearly, pH and the source and amount of carbohydrate influence the transcription of the exopolysaccharide machinery of S. mutans in biofilms. However, a number of other factors could also account for altered gtf and ftf expression in mature biofilms. The first is that the availability of other nutrients and oxygen may be altered compared to young biofilms or planktonic cells. Goodman & Gao (2000) have found that gtf expression occurs at specific growth phases and have suggested a possible cell-density-dependent component regulating gtfBC, raising the possibility that intercellular signalling may be involved in regulating gtf expression. Although the results we have collected thus far do not fully support that quorum sensing is a regulator of exopolysaccharide synthesis in S. mutans, a luxS homologue is identifiable in the S. mutans chromosome, and peptide-mediated intercellular signalling is also established in S. mutans (Li et al., 2001
). Therefore, the idea that exopolysaccharide production is regulated by quorum sensing mechanisms cannot yet be excluded. In summary, this study highlights the need for additional research with suspended and adherent oral streptococci to determine whether the changes in the expression patterns of genes of S. mutans growing in biofilms are restricted to spatially isolated subpopulations or to pathways that globally regulate biofilm gene expression in densely packed populations.
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ACKNOWLEDGEMENTS |
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Received 19 April 2001;
revised 4 June 2001;
accepted 8 June 2001.