Department of Oral Biology, Faculty of Dentistry, University of Manitoba, Winnipeg, Manitoba, Canada
Correspondence
I. R. Hamilton
ihamilt{at}ms.umanitoba.ca
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ABSTRACT |
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INTRODUCTION |
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The enhanced resistance of biofilm cells to adverse environmental conditions and antimicrobial agents, compared to cells grown in liquid culture, has been well established and has led to the concept that surface-associated cells are physiologically distinct from their planktonically grown counterparts (Costerton et al., 1987; Fletcher, 1991
; Hoyle & Costerton, 1991
; Brown & Gilbert, 1993
; Goodman & Marshall, 1995
; Davey & O'Toole, 2000
). Recent research has demonstrated the biofilm-dependent regulation of gene expression and cell-to-cell signalling, including a variety of pathways invoked not only during the initiation of biofilm formation, but also during the succession of the biofilm community (O'Toole & Kolter, 1998
; Christensen et al., 1998
; Prigent-Combaret et al., 1999
; Davey & O'Toole, 2000
; Sauer et al., 2002
). The adherence of oral streptococci to surfaces also results in complex changes in cell physiology, with quorum-sensing mechanisms playing a central role in the characteristics of the resulting biofilms (Loo et al., 2000
; Li et al., 2001a
). Biofilm cell density has been shown to modulate adaptation to acid tolerance at low pH, such that high cell density biofilms were more resistant to lethal pH values than those with lower cell densities (Li et al., 2001b
).
Using a proteomics approach, protein expression in 3 day biofilm cells of S. mutans H7, growing in a biofilm-chemostat with limiting glucose at pH 7·5, was shown to be significantly different from that of the planktonic cells associated with the biofilms in the same chemostat (Svensäter et al., 2001). Of the proteins identified, 13 were unique to biofilm cells, while 9 were only expressed in planktonic cells. Overall, 20 % of the detectable proteins were differentially expressed in biofilm cells compared to planktonic cells, confirming the concept that cells assuming growth on a surface undergo significant changes in physiology. More recently, biofilm cells of the organism were shown to be significantly more acid tolerant than planktonic cells, with glycolytic enzymes in the surface-associated cells up-regulated following a pH change from 7·5 to 5·5, suggesting that their enhanced acid resistance is associated with the maintenance of pH homeostasis (Welin et al., 2003
).
In this study, we were interested in assessing selected physiological properties of biofilm cells of S. mutans BM71 growing over a 5 day period with limiting glucose in a biofilm-chemostat at a constant pH (7·5) in comparison with those properties in associated planktonic cells. Furthermore, to assess the effect of an acid shock typical of natural dental plaque biofilms, physiological tests were also carried out with biofilms subjected to three consecutive days of glucose pulses without pH control that resulted in pH decreases below 5·0 over a 5 h period.
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METHODS |
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Biofilm-chemostat.
Biofilms of S. mutans BM71 were generated in the controlled environment, rod-chemostat system of Bowden (1999) by the general methods previously reported (Li & Bowden, 1994a
). Continuous baseline planktonic growth was achieved in the diluted MADM at pH 7·5 with a dilution rate (D) of 0·1 h-1 (doubling time, tD, 7 h) with glucose limitation. The gas phase was 5 % CO2 in nitrogen and the planktonic cultures were grown for at least 10 mean generations before steady state was considered to have been established. For biofilm formation, sterile epon hydroxyapatite rods (Li & Bowden, 1994b
) were aseptically inserted into either an 8- or 10-rod chemostat head and immersed in steady-state planktonic cultures for periods of 15 days. The growth area on each rod was 3·77 cm2. To maintain consistency of biofilm growth, the same rods were inserted in the same locations in the chemostat head. Chemostats were monitored daily for cell density (Klett readings), pH and medium flow (dilution rate), with periodic samples removed for planktonic cell counts and glucose determinations.
Bacterial sampling and viability.
For biofilm cell viability, the rods were removed from the chemostat and rinsed gently in DM prior to the removal of the cells by sonication for 60 s (Li & Bowden, 1994a). Viable cell counts (n=40) were determined by diluting and plating aliquots of the suspension on blood agar in triplicate as previously described (Svensäter et al., 1997
). In addition, during baseline characterization the percentage live cells was determined using the Live/Dead Bacterial Viability method (LIVE/DEAD BacLight, Kit L-7012, Molecular Probes) and averaging the live and dead cell counts in five separate fields (McNeill & Hamilton, 2003
). Planktonic cells were collected from the chemostat sample port and washed twice in DM before aliquots were diluted for counts and assessment of live and dead cells. The results are reported as the means±standard error for a minimum of three separate experiments involving a minimum of 24 rods.
Glucose-pulse experiments.
To assess the effect of acid stress on the physiology of biofilm and planktonic cells, a steady-state chemostat culture was pulsed once a day (9 a.m.) with glucose (50 mM final concentration) for three consecutive days. Immediately before the initial glucose addition, sterile rods were inserted in the chemostat and left there for the 3 day pulse period (3 day biofilms). Immediately before the daily glucose addition, the pH control system was shut off and the pH of the culture was allowed to fall for 5 h before pH control was re-established at pH 7·5. For characterization of the glucose pulsing over the 3 day period, samples were removed daily from the planktonic phase just before the glucose addition and throughout the subsequent 12 h period, to measure culture pH, viable cell counts and the glucose concentration. The planktonic and biofilm cell samples used for metabolic analyses were removed at 9 a.m. on day 4, such that the cells had been exposed to medium at pH 7·5 for 19 h following the third and last glucose pulse. Glucose limitation of the planktonic culture was re-established within 12 h of the initiation of glucose pulsing. All such samples were subjected to dilution and plating to determine the viable cell counts.
Rod incubation chamber.
Metabolic activity of intact cells in biofilms was carried out in a rod incubation chamber (RIC), which consisted of an acrylic cylinder (9 cm highx1·8 cm in diameter) cemented to a flat base with a sampling port, fitted with a serum stopper, located 2 cm from the bottom of the cylinder. The rods were held in the chamber in a nylon cap capable of holding 5 rods. The RICs were located in a 37 °C incubator on a magnetic stirrer, with the medium mixed by a small stirring bar. With the rods inserted, the maximum medium volume was 22 ml, which allowed the removal of a total of 10 ml of medium for analysis with the rods still submerged in the liquid phase. In all cases, the assay medium was added to the RIC and pre-incubated at 37 °C before the rods were removed from the chemostat, gently rinsed in pre-warmed medium, and added to the chamber without disturbing the biofilms.
Glycolytic rate.
The rate of acid formation from glucose was determined with cells in intact biofilms, with dispersed biofilm cells and with planktonic cells. The incubation medium (IB) was sterile 5 mM sodium/potassium phosphate buffer, pH 7·5, with 10 mM glucose as the carbon source. All pH titrations were carried out at 37 °C with a Radiometer Autoburette (model ABU 1a) (Hamilton & Buckley, 1991) and included five titrations of IB prior to each incubation (pre-zero time) to establish the baseline volumes of standardized KOH required to reach pH 7·50. Intact biofilms and cells dispersed from biofilms were assayed on the same day by using five alternate rods for the respective assays. Intact biofilms were gently rinsed in 2·0 ml sterile, pre-warmed buffer and then immediately (zero time) added to the RIC containing 22 ml IB. At 30, 60, 90 and 120 min, 3·0 ml samples were removed from the RIC by a sterile syringe, added to the pH stat reaction chamber and the amount of KOH required to titrate to 7·50 recorded. Following removal of the final sample, the rods were rinsed in RM, and the cells were removed by sonication and plated on blood agar as described previously. To correct for the contribution of shed cells to the overall acid production, the remaining medium in the RIC following the last sample was centrifuged and the cell count determined. The net acid formation was determined at each time point and the results reported as nmol acid formed min-1 (109 cells)-1, with the values corrected for the acid generated by the cells lost from the rods [2·8(±0·7) % of the total]. Each daily assay consisted of one set of five rods, with the results of at least three such daily measurements used for 1, 2, 3 and 5 day biofilms.
For the dispersed biofilm cells, the rods were gently rinsed in 2·0 ml cold, sterile 5 mM buffer and the cells removed from the rods by sonication into the same buffer. The dispersed biofilm suspensions were pooled, washed once in buffer, resuspended in 15 ml buffer and stored in ice until used. Triplicate 10 µl samples were removed, diluted and plated on blood agar for cell counts. For glycolytic rate determinations, 3·0 ml of the cell suspension was equilibrated in the pH stat reaction chamber at 37 °C and the pH adjusted to 7·50. At zero time, 75 µl pre-warmed 0·4 M glucose was added and the volume of standardized KOH required to maintain the pH at 7·50 was recorded over a 10 min period. The reaction was repeated a total of four times with fresh samples of the suspension and the results were reported as for the intact biofilms. Each experiment was repeated at least three times. A similar procedure was followed for planktonic cells. For this, 40 ml suspension was removed from the sample port, and the cells were washed twice, resuspended in 20 ml cold, sterile 5 mM buffer and placed in ice until used. Triplicate aliquots were diluted in DM and plated on blood agar to establish the viable count before being assayed as described above.
Glucose uptake.
Each intact rod biofilm was removed and assayed separately for the uptake of [14C]glucose. Upon removal from the chemostat, the rod was gently rinsed in 2·0 ml sterile, pre-warmed 50 mM phosphate buffer (pH 7·5), added to the RIC cap and immediately (time zero) added to a RIC incubated at 37 °C and containing 15 ml of the same buffer and 300 µl sterile [14C]glucose sufficient to give 5·0x105 c.p.m. ml-1. At 40 s, the rod was rapidly removed and added to 2·0 ml sterile 15 mM NaF to stop the reaction and the cells sonicated from the rods into 2·0 ml DM as previously described. Duplicate 10 µl samples were diluted and plated to determine the cell count and the remainder of the suspension was filtered through 0·45 µm cellulose filters (Nucleopore) as described by Cvitkovitch et al. (1995). The filters were dissolved in 1·0 ml acetone before the addition of 4·0 ml Aquasol (Packard) and their radioactivity measured in a liquid scintillation counter (Beckman). A minimum of six biofilms were assayed for each age of biofilm. Planktonic cells were collected as described above, washed and resuspended in 7·5 ml sterile 50 mM buffer; triplicate 10 µl samples were diluted and plated for cell counts. Triplicate uptake reactions consisted of 2·0 ml cell suspension equilibrated at 37 °C in a pH stat reaction vessel containing a small stirring bar. At time zero, 40 µl [14C]glucose was added, and 0·5 ml samples were removed at 10, 20 and 30 s and added to 0·5 ml 30 mM NaF to stop the reaction. The cells were filtered, washed and their radioactivity measured as described above. The results are reported as nmol glucose taken up min-1 (109 cells)-1.
Macromolecular synthesis.
DNA, RNA and protein synthesis by 1, 2, 3 and 5 day biofilms was assessed by the respective uptake of [3H]thymidine, [3H]uridine and a mixture of 14C-labelled amino acids (Hilliard et al., 1999). Each assay consisted of duplicate rods removed from the chemostat, rinsed gently in MADM medium and immediately immersed in a RIC containing 15 ml pre-warmed MADM growth medium plus 150 µl of the appropriate substrate and incubated at 37 °C for 5 min. DNA synthesis was measured with [methyl-3H]thymidine (70 Ci mmol-1 at 0·5 µCi ml-1; Amersham), RNA synthesis with [3H]uridine (25 Ci mmol-1 at 0·1 µCi ml-1; Amersham) and protein synthesis with a 14C-labelled amino acid mixture (54 mCi atom carbon-1 at 1·0 µCi ml-1; NEN) [1 Ci=37 GBq]. At 5 min, the rods were removed and added to 2·0 ml cold sterile DM and the cells sonicated from the rods. Triplicate 10 µl samples were removed for plate counts and this was followed immediately by the addition of 2·0 ml cold 10 % TCA to the remaining cells. This suspension was filtered through glass microfibre filters (Whatman), washed with 5·0 ml cold 10 % TCA and finally with 5·0 ml cold distilled water. The filters were then dried and their radioactivity measured in 4 ml Aquasol. Planktonic cells were collected from the chemostat sample port, washed, resuspended in MADM, and samples diluted for plate counting as previously described. The reactions were initiated by the addition of 10 µl of the radioactive substrate to tubes containing 1·0 ml pre-warmed cell suspension. For the uptake of [3H]thymidine and [3H]uridine, the reactions in duplicate tubes were terminated at 2 and 5 min by the addition of 10 % TCA followed by filtration as described above. A similar method was used to measure the uptake of 14C-labelled amino acids with the exception that 100 µl cold Casamino acids (5 mg ml-1) was added with the 10 % TCA at the termination of uptake at 2 and 5 min. The results are expressed as radioactivity taken up by cells: pmol min-1 (109 cells)-1.
ATPase and glucose-PTS activity.
H+/ATPase activity was assayed in permeabilized cells by measuring the release of inorganic phosphate from ATP (Belli & Marquis, 1994), while glucose-PTS was assayed in permeabilized cells by the phosphoenolpyruvate (PEP)-dependent uptake of [14C]glucose via the phosphotransferase system (PTS) (Hamilton et al., 1989
). Cells, sonicated from a minimum of 10 rinsed biofilm rods, were pooled in 2·0 ml DM and 10 µl samples removed for cell counts. Planktonic cells from the same chemostat were collected as described above. Both biofilm and planktonic cells were then washed twice, resuspended in cold, sterile permeabilization buffer [75 mM Tris/HCl (pH 7·0),10 mM MgCl2] before being frozen in 1·0 ml aliquots. Permeabilization was carried out immediately prior to each assay and involved thawing a 1·0 ml suspension, adding 50 µl of a toluene/acetone mixture (1 : 9) and vigorously mixing for 5 min with cooling in ice every 60 s. For the ATPase assay, cells were washed twice and resuspended in assay buffer consisting of 50 mM Tris/maleate buffer (pH 6·0) and 20 mM MgCl2. Triplicate reactions (0·5 ml) were initiated by the addition of ATP (5 mM, final concentration) with incubation at 37 °C for 30 min. Controls included cells without ATP and ATP without cells. The reactions were stopped by the addition of 25 µl 2 M HCl and the phosphate released was measured by the FiskeSubbaRow method (Sigma Diagnostics). The results were expressed as nmol Pi released min-1 (109 cells)-1. For the glucose-PTS assay, cells were washed twice and resuspended in assay buffer consisting of 50 mM phosphate buffer (pH 7·5), 4 mM MgCl2, 20 mM NaF and 5 mM 2-mercaptoethanol. Triplicate assays (0·5 ml) of pre-warmed cell suspension plus 4 mM PEP were initiated by the addition of 10 µl [14C]glucose (1·0 µCi ml-1) with incubation at 37 °C for 15 min. Controls included cells and [14C]glucose without PEP. The reactions were stopped by the addition of 4·5 ml ethanolic BaBr2 and the suspensions filtered through Whatman HA filters (0·45 µm), following a 20 min incubation in ice, and counted in Aquasol as described above. The results are reported as nmol glucose taken up min-1 (109 cells)-1.
Analyses.
The glucose concentration in the medium was determined with glucose oxidase by the method of Kingsley & Getchell (1960). Statistical analysis was carried out with the computer program Prism 4 for Macintosh.
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RESULTS |
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Baseline physiological status
The physiology of biofilm cells was assessed under two sets of conditions: during growth with limiting glucose at pH 7·5 (baseline conditions) and following a period of acid stress that resulted from three daily glucose pulses, each followed by a 5 h period without pH control. The growth at pH 7·5 provided the baseline physiological status of 15 day-old biofilms as a control for the comparisons made between 3 day baseline and glucose-pulsed biofilms. Planktonic cells generated in the chemostat under the same conditions were also included in all assays to provide a comparison to their associated biofilms.
Carbohydrate metabolism, as estimated by the glycolytic rate of biofilm cells, was assessed with intact biofilms and with dispersed cells removed from the biofilms. As seen in Fig. 1, baseline acid formation by intact biofilms decreased with age, with 1 day biofilms 14-fold more active than 5 day biofilms [638±60 vs 46±10 nmol acid formed min-1 (109 cells)-1] (P<0·05). The results were corrected for the number of cells shed during the assay, which was minimal [2·8(±0·7) %]. Interestingly, the activity of the biofilm cells removed from alternate rods was higher than that of the intact biofilms, ranging from 2·1- to 4·7-fold higher for the 1 and 5 day biofilm cells, respectively. In addition, the loss in activity over the 5 day period was only 6-fold [1315±163 (1 day) vs 212±55 (5 days) nmol acid formed min-1 (109 cells)-1]. The glycolytic rate of the planktonic (P) cells associated with the biofilms was 295 nmol acid formed min-1 (109 cells)-1, less than that of the 1 and 2 day intact biofilms.
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DISCUSSION |
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Generally, physiological activity under baseline conditions was highest within the first 3 days and decreased with the 5 day rods as the growth rate declined. Predictably, activity of the associated steady-state planktonic cells under the baseline conditions was higher than that of the slower-growing biofilm cells, except for glucose-PTS and H+/ATPase activity (Table 3 vs Table 4
). The increased glycolytic activity upon dispersal of the cells from the rods (Fig. 1
) would suggest glucose diffusion was limited in the intact biofilms, reducing glucose uptake. Although viable cell numbers on the rods were not high, ranging from 1·1 to 2·2x107 cm-2 over the 5 day period, it is conceivable that cells inside small clusters in the biofilm would be deprived of glucose, causing a lower biofilm glycolytic rate compared to the same cells removed from the rods (Stewart, 2003
). This concept is supported by the fact that the ratio of dispersed/intact cell activity increased progressively with the increase in biofilm cell numbers over the 5 day period, e.g. 2.l, 2·5, 3·2 and 4·6 for the 1, 2, 3 and 5 day biofilms, respectively. S. mutans BM71 is known to form microcolonies readily on glass rods in the same biofilm-chemostat system (Li & Bowden, 1994a
).
The pulsing of the chemostat with glucose in the absence of pH control was initiated to simulate the conditions that dental plaque can be subjected to during dietary consumption of refined sugars (Stephan, 1944; Hamilton, 2000
). The concentration of glucose (50 mM) was chosen to provide a period of acid stress in order to test the physiological response of both biofilm and planktonic cells. That acid stress had been achieved was apparent from the fact that the pH minimum (4·374·40) observed over the 3 day pulse period (Table 2
) was 0·40 pH units below the pH limit for growth of the organism (Hamilton, 1986
), indicating uncoupling of growth and metabolism. This is reflected in the gradual decline in the planktonic viable cell counts over the 3 day pulse period from 44·0 to 9·4 %. Whether the loss of cells from the planktonic phase during the glucose pulse period was due to acid-induced growth inhibition or killing is not known; however, such cells have a poor capacity to induce an ATR that would aid survival at low pH, suggesting inhibition of cellular biosynthesis triggered by cytoplasmic acidification (Hamilton & Buckley, 1991
; McNeill & Hamilton, 2003
). This is supported by the observation that thymidine, uridine and amino acid uptake into glucose-pulsed planktonic cells was inhibited compared to baseline cells. It should be noted that, although the cell counts declined during the 5 h glucose-pulsing period on each of the 3 days, the cell numbers increased to normal or pre-pulse values during the 19 h following the resumption of growth at pH 7·5, the last 12 h of which were under glucose-limited conditions.
The 3 days of glucose pulsing and concomitant acidification resulted in lower numbers of biofilm cells (51 %) than on the rods under baseline conditions; nevertheless, pulsing did have a stimulatory effect on all of the measured physiological properties of the biofilm cells compared to that under baseline conditions (Table 3). The enhanced glucose uptake and metabolism by the biofilm cells during the 3 day pulsing period is indicative of carbohydrate training, which has been shown to be associated with the emergence and domination of the acidogenic/aciduric microflora in mixtures of oral bacteria (Bradshaw et al., 1989
) and in dental plaque of human subjects (Stephan, 1944
). Furthermore, the increased glycolytic activity is consistent with recent proteomic data showing that the synthesis of glycolytic enzymes in biofilm cells of S. mutans H7 is up-regulated during pH changes from 7·5 to 5·5 compared to control cells maintained at pH 7·5 (Welin et al., 2003
). This was made possible since, unlike the associated planktonic cells, biofilm cells increased the uptake of thymidine, uridine and amino acids into DNA, RNA and protein (Table 3
). These latter results are consistent with the enhanced growth and biofilm formation observed previously with S. mutans BM71 under conditions of glucose excess (Li & Bowden, 1994a
, b
) and concur with the recent observation that biofilm cells possessed a greater capacity to induce an ATR than the associated planktonic cells growing in the same biofilm-chemostat (McNeill & Hamilton, 2003
).
Clearly, our results indicate that biofilm cells, under conditions of acid stress, were more physiologically fit than the associated planktonic cells in the same chemostat. The key physiological trait would appear to be related to the enhanced capacity of biofilm cells to maintain cellular pH homeostasis. Intracellular pH levels in S. mutans are maintained above that of the external environment by proton extrusion via membrane-associated H+/ATPase (Bender et al., 1986) and lactate efflux (Dashper & Reynolds, 1996
), and it is known that the sustained growth of the organism at pH 5·05·2 results in the increased synthesis of lactic dehydrogenase (Wilkins et al., 2002
) and H+/ATPase (Hamilton & Buckley, 1991
). This latter result with planktonic cells of S. mutans Ingbritt demonstrated that growth at progressively lower pH values was correlated to increases in H+/ATPase activity, a result confirmed with both the biofilm (Table 3
) and planktonic (Table 4
) cells of S. mutans BM71 in this study. The maintenance of H+/ATPase activity in the 3- and 5 day biofilm cells under baseline conditions in spite of declining glycolytic activity (Table 1
, Fig. 1
) suggests that sufficient ATP was being generated to drive H+ expulsion from the cell. Furthermore, the significantly higher level of ATPase activity in the baseline biofilm cells compared to that in the planktonic cells indicates that the biofilm environment may be more acidic than the bulk liquid phase, a situation which would account for the high level of ATPase activity in the baseline biofilm cells. The capacity to maintain cellular pH could explain the higher level of macromolecular synthesis during glucose pulsing by biofilm cells and account for the enhanced capacity of such cells to induce an ATR when subjected to a pH change from 7·5 to 5·5 (McNeill & Hamilton, 2003
; Welin et al., 2003
).
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ACKNOWLEDGEMENTS |
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REFERENCES |
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Belli, W. A. & Marquis, R. E. (1994). Catabolite modification of acid tolerance of Streptococcus mutans GS5. Oral Microbiol Immunol 9, 2934.[Medline]
Bender, G. R., Sutton, S. C. & Marquis, R. E. (1986). Acid tolerance, proton permeabilities, and membrane ATPase of oral streptococci. Infect Immun 53, 331338.[Medline]
Bowden, G. H. W. (1991). Which bacteria are cariogenic in humans? In Dental Caries Markers of High and Low Risk Groups and Individuals, vol. 1, pp. 266286. Edited by N. M. Johnson. Cambridge: Cambridge University Press.
Bowden, G. H. W. (1999). Controlled environment model for accumulation of biofilms of oral bacteria. Methods Enzymol 310, 216224.[Medline]
Bradshaw, F. J., McKee, A. S. & Marsh, P. D. (1989). Effects of carbohydrate pulses and pH on populations shifts within oral microbial communities in vitro. J Dent Res 68, 12981302.[Abstract]
Brown, M. R. W. & Gilbert, P. (1993). Sensitivity of biofilms to antimicrobial agents. J Appl Bacteriol Suppl 74, 8797.
Christensen, B. B., Sternberg, C., Andersen, J. B., Eberl, L., Møller, S., Givskov, M. & Molin, S. (1998). Establishment of new genetic traits in a microbial biofilm community. Appl Environ Microbiol 64, 22472255.
Costerton, J. W., Cheng, K.-J., Geesey, G. G., Ladd, T. I., Nickel, J. D., Dagupta, M. & Marrie, T. J. (1987). Bacterial biofilms in nature and disease. Annu Rev Microbiol 41, 435464.[CrossRef][Medline]
Cvitkovitch, D. G., Boyd, D. A., Thevenot, T. & Hamilton, I. R. (1995). Glucose transport by a mutant of Streptococcus mutans unable to accumulate sugars via the phosphoenolpyruvate phosphotransferase system. J Bacteriol 177, 22512258.[Abstract]
Dashper, S. G. & Reynolds, E. C. (1996). Lactic acid excretion by Streptococcus mutans. Microbiology 142, 3339.
Davey, E. M. & O'Toole, G. (2000). Microbial biofilms: from ecology to molecular genetics. Microbiol Mol Biol Rev 64, 847867.
Fletcher, M. (1991). The physiological activity of bacteria attached to solid surfaces. Adv Microb Physiol 32, 5385.[Medline]
Goodman, A. E. & Marshall, K. C. (1995). Genetic responses of bacteria at surfaces. In Microbial Biofilms, pp. 8098. Edited by J. W. Costerton & H. M. Lappin-Scott. Cambridge: Cambridge University Press.
Hamilton, I. R. (1986). Growth, metabolism and acid production by Streptococcus mutans, In Molecular Microbiology and Immunobiology of Streptococcus mutans, pp. 145155. Edited by S. Hamada, S. M. Michalek, H. Kiyono, L. Menaker & J. R. McGhee. Amsterdam: Elsevier Science Publishers.
Hamilton, I. R. (2000). Ecological basis for dental caries. In Oral Bacterial Ecology. The Molecular Basis, pp. 219274. Edited by H. K. Kuramitsu & R. P. Ellen. Wymondham: Horizon Scientific Press.
Hamilton, I. R. & Bowden, G. H. (2000). Oral Microbiology. In Encyclopedia of Microbiology, vol. 3, pp. 466481. Edited by J. Lederberg. San Diego: Academic Press.
Hamilton, I. R. & Buckley, N. D. (1991). Adaptation by Streptococcus mutans to acid tolerance. Oral Microbiol Immunol 6, 6571.[Medline]
Hamilton, I. R., Gauthier, L., Desjardins, B. & Vadeboncoeur, C. (1989). Concentration-dependent repression of the soluble and membrane components of the Streptococcus mutans phosphoenolpyruvate : sugar phosphotransferase system by glucose. J Bacteriol 171, 29422948.[Medline]
Hilliard, J. J., Goldschmidt, R. M., Licata, L., Baum, E. Z. & Bush, K. (1999). Multiple mechanisms of action for inhibitors of histidine protein kinases from bacterial two-component systems. Antimicrob Agents Chemother 43, 16931699.
Hoyle, B. D. & Costerton, J. W. (1991). Bacterial resistance to antibiotics: the role of biofilms. Prog Drug Res 37, 91105.[Medline]
Kingsley, G. R. & Getchell, G. (1960). Direct ultra micro glucose oxidase method for the determination of glucose in biological fluids. Clinical Chemistry 6, 466475.[Medline]
Li, Y.-H. & Bowden, G. H. (1994a). Characteristics of accumulation of oral gram-positive bacteria on mucin-conditioned glass surfaces in a model system. Oral Microbiol Immunol 9, 111.[Medline]
Li, Y.-H. & Bowden, G. H. (1994b). The effect of environmental pH and fluoride from the substratum on the development of biofilms of selected oral bacteria. J Dent Res 73, 16151626.[Abstract]
Li, Y.-H., Lau, P. C. Y., Lee, J. H., Ellen, R. P. & Cvitkovitch, D. G. (2001a). Natural genetic transformation of Streptococcus mutans growing in biofilms. J Bacteriol 183, 897908.
Li, Y.-H., Hanna, M. H., Svensäter, G., Ellen, R. P. & Cvitkovitch, D. G. (2001b). Cell density modulates acid adaptation in Streptococcus mutans: implications for survival in biofilms. J Bacteriol 183, 68756884.
Loesche, W. J. (1986). Role of Streptococcus mutans in human dental decay. Microbiol Rev 50, 353380.[Medline]
Loo, C. Y., Corliss, D. A. & Ganeshkumar, N. (2000). Streptococcus gordonii biofilm formation: identification of genes that code for biofilm phenotypes. J Bacteriol 182, 13741382.
McNeill, K. & Hamilton, I. R. (2003). Acid tolerance response of biofilm cells of Streptococcus mutans. FEMS Microbiol Lett 221, 2530.[CrossRef][Medline]
O'Toole, G. A. & Kolter, R. (1998). Flagellar and twitching motility are necessary for Pseudomonas aeruginosa biofilm development. Mol Microbiol 30, 295304.[CrossRef][Medline]
Prigent-Combaret, C., Vidal, O., Dorel, C. & Lejeune, P. (1999). Abiotic surface sensing and biofilm-dependent regulation of gene expression in Escherichia coli. J Bacteriol 181, 59936002.
Sauer, K., Camper, A. K., Ehrlich, G. D., Costerton, J. W. & Davies, D. G. (2002). Pseudomonas aeruginosa displays multiple phenotypes during development as a biofilm. J Bacteriol 184, 11401154.
Stephan, R. M. (1944). Intra-oral hydrogen-ion concentration associated with dental caries activity. J Dent Res 23, 257266.
Stewart, P. S. (2003). Diffusion in biofilms. J Bacteriol 185, 14851491.
Svensäter, G., Larsson, U.-B., Greif, E. C. G., Cvitkovitch, D. G. & Hamilton, I. R. (1997). Acid tolerance response and survival by oral bacteria. Oral Microbiol Immunol 12, 266273.[Medline]
Svensäter, G., Welin, J., Wilkins, J. C., Beighton, D. & Hamilton, I. R. (2001). Protein expression by planktonic and biofilm cells of Streptococcus mutans. FEMS Microbiol Lett 205, 139146.[CrossRef][Medline]
Watanabe, S. & Dawes, C. (1988). The effects of different foods and concentrations of citric acid on the flow rate of whole saliva in man. Arch Oral Biol 33, 15.[Medline]
Welin, J., Wilkins, J. C., Beighton, D., Wrzesinski, K., Fey, S. J., Mose Larsen, P., Hamilton, I. R. & Svensäter, G. (2003). Effect of acid shock on protein expression by biofilm cells of Streptococcus mutans. FEMS Microbiol Lett 227, 287293.[CrossRef][Medline]
Wilkins, J. C., Homer, K. A. & Beighton, D. (2002). Analysis of Streptococcus mutans proteins modulated by culture under acidic conditions. Appl Environ Microbiol 68, 23822390.
Wimpenny, J., Manz, W. & Szewzyk, U. (2000). Heterogeneity in biofilms. FEMS Microbiol Rev 24, 661671.[CrossRef][Medline]
Yamada, T., Igarashi, K. & Mitsutomi, M. (1980). Evaluation of cariogenicity of glycosylsucrose by a new method of measuring pH under human dental plaque in situ. J Dent Res 59, 21572162.
Received 12 May 2003;
revised 22 October 2003;
accepted 8 December 2003.
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