ABSTRACT
During initial dental plaque formation, the ability of a species to grow when others cannot would be advantageous, and enhanced growth through interspecies and intergeneric cooperation could be critical. These characteristics were investigated in three coaggregating early colonizers of the tooth surface (Streptococcus gordoniiDL1, Streptococcus oralis 34, and Actinomyces naeslundii T14V). Area coverage and cell cluster size measurements showed that attachment of A. naeslundii and of S. gordonii to glass flowcells was enhanced by a salivary conditioning film, whereas attachment of S. oralis was hindered. Growth experiments using saliva as the sole carbon and nitrogen source showed that A. naeslundii was unable to grow either in planktonic culture or as a biofilm, whereas S. gordonii grew under both conditions. S. oralis grew planktonically, but to a much lower maximum cell density than did S. gordonii;S. oralis did not grow reproducibly as a biofilm. Thus, only S. gordonii possessed all traits advantageous for growth as a solitary and independent resident of the tooth. Two-species biofilm experiments analyzed by laser confocal microscopy showed that neither S. oralis nor A. naeslundii grew when coaggregated pairwise with S. gordonii. However, both S. oralis and A. naeslundii showed luxuriant, interdigitated growth when paired together in coaggregated microcolonies. Thus, the S. oralis-A. naeslundiipair formed a mutualistic relationship, potentially contact dependent, that allows each to grow where neither could survive alone. S. gordonii, in contrast, neither was hindered by nor benefited from the presence of either of the other strains. The formation of mutually beneficial interactions within the developing biofilm may be essential for certain initial colonizers to be retained during early plaque development, whereas other initial colonizers may be unaffected by neighboring cells on the substratum.
The human oral biofilm dental plaque and the biofilms of periodontal diseases are perhaps the best described of the naturally occurring multispecies prokaryotic communities, at least from the standpoint of species composition; more than 500 known species have been isolated from the mouth (27). A recent assessment has placed the proportion of yet-to-be-cultivated bacteria from subgingival dental plaque at approximately 50% (25)—a sharp contrast to other natural environments, such as soil or marine sediment, where ≥99.9% of the population is estimated to be uncultivated (1, 30). The oral microflora is under enormous pressure to grow as a biofilm. Salivary flow rates (dilution rates) are too high, and the free carbohydrate levels in saliva are too low, for significant multiplication of bacteria in the planktonic state in the mouth (6).
The literature is unclear regarding the ability of oral bacterial monocultures to grow in saliva ex vivo. Several reports exist in which growth of an oral bacterial strain is monitored in amended (glucose-supplemented) saliva (13, 14). However, bacterial consortia can increase in biomass to an optical density at 550 nm of 0.35 in unamended saliva (13). The degree to which single-species oral isolates can maintain growth in unamended saliva is less clear. Growth of an Actinomyces species, but not of three Streptococcus species, in saliva has been reported (14). However, growth in these experiments was assessed turbidometrically (A550), and therefore the initial culture density must have been high (>107 cells ml−1). At this density, the salivary concentration of free glucose (10 to 100 μM [6]) cannot support repeated cell division. Also, when growth did occur, it was not monitored over subsequent transfers and therefore might have been attributable to the physiological state of the inoculum (that is, continuance of cell growth after transfer from nutrient-rich culture medium to saliva) rather than to the ability of the cells to use the endogenous nutrients present in the saliva. Studies of the growth of oral isolates on unamended saliva have been limited because the ability of a mixed oral microflora to survive in vivo does not depend on the presence of any single strain. However, if a bacterial species is to become part of a plaque consortium, it could be advantageous for the bacterium to be capable of growth on the nutrients provided solely by saliva. Such species could grow independently on the tooth surface, a niche in which other species that require additional growth factors are unable to grow. Some of these growth factors could be obtained from the processing of complex salivary glycoproteins by neighboring species in the oral plaque community (5). Thus, in the context of early dental plaque development, while the ability of a bacterium to grow independently on unamended saliva could be an important trait, other bacteria that have developed mutualistic partnerships also flourish.
Another potentially important trait of early colonizers of the tooth surface is the ability to coaggregate (a planktonic phenomenon [10, 16, 24]) or to coadhere (coaggregation at a substratum [3, 8]). This ability of a bacterial strain to bind to another strain fosters a close spatial relationship and, potentially, a close metabolic interaction. Coaggregation interactions are known for the majority of oral genera and are hypothesized to play a role in the four-dimensional (x, y,z, and time) structure of dental plaque (22). Nonrandom spatial organization in multispecies biofilms related to metabolic cooperation between organisms is known from nonoral in vitro model systems (7, 26, 28, 34, 35, 37) and, in nature, from microbial mats (12), but in these systems many if not all of the bacterial species are motile. In a system composed primarily of nonmotile organisms such as the oral microflora, biofilm architecture may be influenced to an even greater degree by metabolic interdependence. Thus, coaggregation and coadherence may play a critical role in community development of nonmotile cells, such as oral bacterial early colonizers.
We studied the ability of three pairwise coaggregating oral bacterial strains (Actinomyces naeslundii T14V, Streptococcus oralis 34, and Streptococcus gordonii DL1) to grow in planktonic monoculture and as monoculture biofilms on saliva-conditioned glass surfaces, using unamended saliva as the growth medium. We compared the abilities of these strains to attach to saliva-conditioned glass and their abilities to attach to unconditioned glass, and we investigated the formation of coadherent, mixed-species microcolonies that were mediated by their coaggregation properties. The growth of these mixed-species colonies on unamended saliva was monitored over 18 h. Results of these experiments demonstrated species-specific initial adherence characteristics for the three strains, documented the cooperative growth of A. naeslundiiwith S. oralis, showed dominance of growth by S. gordonii, and established a baseline for further, more complex, multispecies, oral biofilm investigations. We show here that coaggregation can offer a distinct growth advantage for certain partner strains in vitro. Thus, coaggregation-dependent growth enhancements might dictate specific spatial organization within dental plaque in vivo.
MATERIALS AND METHODS
Flowcells.Two-track flowcells (parallel plate flow chambers with a working volume of 250 μl/track) were constructed by modification of the method of Palmer and colleagues (21, 31) using a microscope slide as the bottom and a no. 1.5 coverglass as the top. The flowcells were cleaned with 0.1 N HCl overnight, washed with several changes of distilled water over a period of 3 h, and autoclaved on the hard-goods cycle without drying. The flow rate of all liquids through the flowcell was 0.0125 mm s−1 (200 μl min−1); liquids were pumped with an MPL pump (Watson-Marlow Inc., Wilmington, Mass.).
Saliva.Saliva stimulated by mastication of silicone tubing or Parafilm balls was collected on ice from at least six volunteers and pooled. Dithiothreitol (Sigma-Aldrich, St. Louis, Mo.) was added to a 2.5 mM final concentration from a 100× stock solution, and the saliva was gently stirred on ice for 10 min, after which it was centrifuged at 4°C and 30,000 × g for 20 min (13). The clarified saliva supernatant was decanted, 3 volumes of distilled water was added, and the 25% saliva was filtered through a 0.20-μm-pore-size filter and frozen in 40-ml aliquots. Immediately prior to an experiment, the sterile saliva was thawed at 37°C; the slight precipitate was pelleted at 1,430 × g for 5 min, and the clear 25% saliva supernatant was used in experiments.
Bacterial strains.Human oral isolates (10)S. gordonii DL1, S. oralis 34, and A. naeslundii T14V were grown from frozen stocks in brain heart infusion broth (BHI; Difco, Detroit, Mich.) overnight at 37°C in an anaerobic glove box (N2-CO2-H2, 90:5:5). For cultures of green fluorescent protein (GFP)-expressing S. gordonii DL1(pCM18) (17), erythromycin was added at 10 μg ml−1. These starter cultures were then subcultured as described below. Erythromycin was omitted from all subculturing.
Attachment of cells to conditioned or to unconditioned glass surfaces.Starter cultures were transferred (0.3 ml) into 8 ml of fresh anaerobic BHI and grown for 2 h (S. gordonii) or 4 h (other strains) at 37°C as static cultures in screw-cap, glass tubes (16 by 150 mm) in an aerobic incubator to reestablish exponential growth. Cultures were split into three aliquots, and the cells were pelleted at room temperature (3,000 ×g for 6 min) in a centrifuge (model 5415C; Eppendorf, Hamburg, Germany). One aliquot was washed three times in phosphate-buffered saline (PBS) and was resuspended in PBS. The other two of the aliquots were washed three times in 25% saliva. One of these saliva-washed aliquots was resuspended in 25% saliva, and the other was resuspended in PBS. The A600of all cell suspensions was adjusted to 0.05 to 0.06.
All following manipulations were performed in a 34°C incubator. Both tracks of one flowcell were saliva conditioned by injection of 0.5 ml of 25% saliva into each track and static incubation for 30 min; another flowcell was likewise exposed to PBS for 30 min (unconditioned). An inoculum of 0.5 ml of cell suspension was injected into each track, and the flowcell was inverted to allow the cells to settle onto the coverglass.
The following matrix of conditions was established (Fig.1): a conditioned surface that was inoculated with saliva-washed cells (condition CS), a conditioned surface that was inoculated with buffer-washed cells (condition CB), an unconditioned (PBS-coated) surface that was inoculated with saliva-washed cells (condition US), and an unconditioned surface that was inoculated with buffer-washed cells (condition UB). The condition US received the cells that had been washed in saliva but then resuspended in buffer. This final resuspension in buffer was performed to minimize adsorption of unbound salivary components to the unconditioned glass.
Flowcells and inocula used in the matrix of conditions established to investigate the effect of substratum conditioning on bacterial attachment. Two flowcells (a saliva-conditioned flowcell and a PBS-coated flowcell) were used for each strain, and the cells of each strain were treated as shown and injected into the indicated tracks.
After 20 min of static incubation, flow was started through the tracks. The condition CS received 25% saliva as the flowing medium; all other conditions received PBS as the flowing medium. After 10 min of flow, the flowcells were returned to the upright position (coverglass on top; attached cells hanging to the underside) and flow was continued for an additional 10 min. A bacterial nucleic acid stain (Syto 59; Molecular Probes, Eugene, Oreg.) (1 μl in 1 ml of PBS) was injected into each track, and three random fields of view in each track were imaged using a TCS 4D laser confocal microscope (Leica LaserTechnik, Heidelberg, Germany). A 40× Plan Apo oil immersion lens (NA 1.0) was used. Biofilms varied in depth from approximately 3 μm (streptococcal strains, condition UB) to approximately 8 μm (A. naeslundii, condition CS). Optical sections (Airy disk setting of 1) were taken at 1-μm intervals, and the resulting optical stack was displayed in a single plane via maximum projection. The maximum projection images were analyzed for area coverage and for cell cluster size using IMAQ Image Builder (National Instruments, Austin, Tex.).
Planktonic growth of bacteria in saliva.Starter cultures of the three strains in BHI were transferred (0.3 ml) to fresh anaerobic BHI (8 ml) and allowed to grow statically for 2 h (S. gordonii) or 4 h (other strains) at 37°C in an aerobic incubator. Cells were pelleted, washed three times in 25% saliva, and resuspended in 25% saliva to give a Klett value of 30 to 50 with a 660-nm filter in a Klett-Summerson colorimeter (Klett Manufacturing Co., Inc., New York, N.Y.). Aliquots of these cell suspensions were diluted 10,000-fold with 25% saliva at 34°C in sterile capped 14-ml polypropylene tubes to obtain the initial saliva culture in a final volume of 3 ml. Aliquots (100 μl) of this initial saliva culture were serially diluted in saliva and plated on BHI agar, the plates were incubated anaerobically at 37°C for 36 h, and colonies were counted to obtain the cell density at 0 h. The remaining initial saliva culture was incubated statically in an aerobic incubator at 34°C, and additional aliquots were removed and processed as described above after 2, 4, and 6 h. Immediately after the 6-h dilution had been plated, 0.3 ml of the remaining initial saliva culture was transferred to fresh 25% saliva (3 ml), and aliquots of this first-transfer culture were processed to yield the 0 h cell density. The first-transfer culture and the initial-saliva culture were then allowed to grow overnight, after which aliquots were processed for plating. Immediately thereafter, an aliquot of the first-transfer saliva culture was transferred (0.3 ml to 3 ml) to fresh 25% saliva to begin the second-transfer culture, and this second-transfer culture was sampled at 0, 2, 4, and 6 h. Thus, cells originally in exponential growth in BHI were washed three times in 25% saliva, and the growth of these cells in 25% saliva was monitored over two additional transfers encompassing 36 h.
Biofilm growth of bacteria in saliva.Starter cultures of the three strains were transferred (0.3 ml) to fresh anaerobic BHI (8 ml) and allowed to grow statically for 2 h (S. gordonii) or 4 h (other strains) at 37°C in an aerobic incubator. Cells were pelleted, washed three times in 25% saliva, and resuspended in 25% saliva to an A600of 0.05 to 0.06. All following manipulations were performed at 34°C in an aerobic incubator. An inoculum of 0.5 ml of the cell suspensions was injected into each track of saliva-conditioned flowcells. The flowcells were inverted and incubated for 20 min statically, after which sterile 25% saliva was pumped through the flowcell for 20 min at 200 μl min−1 to wash out unattached cells. In coculture biofilms, a second strain was injected at this time, allowed to adhere for 20 min, and then washed out as described above. This procedure was carried out with three flowcell tracks (for 0, 4, and 18 h), and after the given periods of salivary flow, a track was stained and observed by confocal microscopy (see below) and then was discarded. The three strains were thus examined for the ability to grow in 25% saliva as a monoculture biofilm and, in the three combinations, as coculture biofilms.
Staining of cells was done by various means in the different experiments. When strains were grown as monocultures, Syto 59 (Molecular Probes) or BacLight Live/Dead (Molecular Probes) was used as recommended by the manufacturer. The S. gordoniiparent strain lacking the GFP-containing plasmid was used in Live/Dead viability assessments. When strains were grown in cocultures, visualization was by primary immunofluorescence with Alexa 568 (Molecular Probes)-conjugated immunoglobulin G of a polyclonal antiserum to S. oralis, with intrinsic GFP fluorescence (forS. gordonii), and with secondary immunofluorescence using Cy2-, Cy3-, or Cy5-conjugated goat anti-mouse immunoglobulin G (Jackson ImmunoResearch, West Grove, Pa.) after reaction with mouse monoclonal antibody A8 (9) to A. naeslundii T14V type 1 fimbriae. The polyclonal antiserum was absorbed against S. gordonii to eliminate cross-reactivity. Immunofluorescence was performed by injecting the primary antibody (500 μl; 10 to 50 μg of protein ml−1 in PBS) and incubating for 10 min, subsequent washout of unbound antibody through injection of 500 μl of PBS, introduction of the secondary antibody (250 μl of PBS containing 2.5 μl of the antibody diluted as recommended by the supplier), and another 10-min binding period. A final wash with PBS preceded confocal microscopy.
RESULTS
Initial attachment of cells to conditioned and unconditioned glass surfaces.Area coverage and average cell cluster area were determined for the three strains in the presence and in the absence of a salivary conditioning film (Fig. 2). For these bacterial strains, attachment followed three strain-dependent patterns. Cells of A. naeslundii attached best to the saliva-conditioned glass, where they also demonstrated the greatest cell cluster area. Prior exposure of the cells to saliva enhanced the attachment of cells to the conditioning film (Fig. 2A; compare condition CS with conditions CB, UB, and US). No clear differences were seen between the other treatments. S. gordonii also attached best to saliva-conditioned glass (Fig. 2B, conditions CB and CS), where it likewise displayed the greatest cell cluster area (Fig. 2B, conditions CB and CS). With S. gordonii, attachment was always greater in the presence of a conditioning film regardless of exposure of the cells to saliva (Fig. 2B; compare conditions CB and CS with conditions UB and US). S. oralis showed a pattern quite different from that of the other two strains: area coverage forS. oralis 34 was always at least twofold higher in the absence of a conditioning film than in the presence of a conditioning film (Fig. 2C; compare conditions UB and US with conditions CB and CS). This effect was independent of exposure of the cells to saliva, and no clear differences in cell cluster area were seen among the various treatments. These experiments were performed twice with the same outcome.
Area coverage (filled bars) and average cell cluster area (open bars) for initial adherent cell clusters of A. naeslundii (A), S. gordonii (B), and S. oralis (C). Error bars show standard deviations based on three samples (the three random fields of view). Two-letter abbreviations are as described in Materials and Methods and as shown in Fig 1. The vertical scale units are the same for both percent area coverage (filled bars) and average cell cluster area (in square micrometers) (open bars).
Planktonic growth in unamended saliva.The three bacterial strains were tested for the ability to grow over three consecutive transfers using unamended saliva as the sole carbon and nitrogen source. These experiments were repeated at least twice with each strain. Results of one of these experiments in which the same saliva collection was used for all three strains are shown in Table1. The cell density of S. gordonii was adjusted to 3.2 × 104 cells ml−1 in the initial transfer from maintenance medium (BHI) into saliva (Table 1, transfer 1, incubation time of 0 h). Over the next 6 h, cell density increased more than 200-fold. After a 10-fold dilution during transfer of the sample (transfer 1, incubation time of 6 h) into fresh saliva (transfer 2, incubation time of 0 h), the two cultures were allowed to grow overnight. Both cultures reached a final density of 3 × 107 to 6 × 107 cells ml−1 (transfer 1, incubation time of 18 h; transfer 2, incubation time of 18 h). A subsequent transfer and 10-fold dilution of a sample from transfer 2 (incubation time of 18 h) yielded a new culture with a density of 1.7 × 106 cells ml−1 (transfer 3, incubation time of 0 h). This culture continued to grow over the next 6 h to a density of 2.1 × 107 cells ml−1(transfer 3, incubation time of 6 h), indicating a doubling time of about 2 h in saliva as the sole carbon and nitrogen source. This pattern of continued growth after all transfers to a cell density of 107 cells ml−1 was characteristic of S. gordonii. In contrast, the only increase in A. naeslundii cell density was seen over the first 6 h after the initial transfer to saliva, during which the cell density increased about twofold. After transfer incorporating a 10-fold dilution (transfer 2, incubation time of 0 h) and overnight incubation of the two cultures (transfer 1, incubation time of 18 h, and transfer 2, incubation time of 18 h), cell density failed to increase and appeared to drop somewhat. After the third transfer, cell density never exceeded the statistically meaningful lower limit of the assay. This pattern of very limited growth only in the initial transfer and subsequent decrease in cell density was typical for A. naeslundii. For S. oralis, results were more variable. The results shown in Table 1represent an example in which growth occurred only in the initial transfer. This example was chosen because the saliva pool used in this experiment was the same as that for the other two strains described above. Cell density increased almost 10-fold between the inoculation time (transfer 1, incubation time of 0 h) and the overnight time point (transfer 1, incubation time of 18 h). However, all subsequent transfers failed to grow. In two other experiments with different saliva pools, growth was maintained over all three transfers with the maximum cell density reaching 105 to 106cells ml−1 (data not shown). Thus, S. oralis was capable of planktonic growth in saliva, but the cell yield was 10- to 100-fold lower than that consistently attained by S. gordonii.
Planktonic growth of bacteria in unamended salivaa
Growth of monoculture biofilms in unamended saliva.Figure3 shows the developmental patterns and cellular vitality of monoculture biofilms of the three bacterial strains. Figure 3A shows images of A. naeslundii T14V biofilms taken at 0 h (initial attachment; comparable to data presented in Fig. 2A) and after 4 and 18 h of biofilm growth on unamended 25% saliva. Cells were present primarily as clumps; the initial height of these clumps varied between 2 and 8 μm. The number of clumps may have decreased with time, and cellular vitality (membrane potential as assessed by Live/Dead stain) deteriorated (i.e., staining changed from yellow or green to red). For S. oralis 34 (Fig. 3B), cells were initially present primarily as chains or clumps; these clumps were about the same height as forA. naeslundii, and the number of cells appeared to have decreased with time. Cellular vitality, however, remained consistently high throughout the experiment (most cells were stained green or yellow). S. gordonii DL1 (Fig. 3C) was also present initially as clumps or chains, and growth occurred over the course of the experiment. The height of the biofilm did not change dramatically over time. Instead, the biofilm expanded laterally and regions lacking cells were filled in as opposed to an increase in biomass height. Cellular vitality was initially high (most cells were stained green). However, the proportion of reddish cells increased somewhat at 4 h and at 18 h. These results indicated that neither A. naeslundii nor S. oralis was capable of sustained growth as a biofilm using 25% saliva as the sole nutrient source. In contrast, S. gordonii grew as a biofilm by metabolizing 25% saliva.
Time course (left to right) of biofilm development in saliva-conditioned glass flowcells and assessment of cell vitality using Live/Dead stain. Red cells have impaired membrane activity, whereas green cells have fully functional membranes. (A) A. naeslundii T14V. Biofilm fails to grow, and the cells lose vitality (change from green to red). (B) S. oralis 34. Biofilm fails to grow, but cells retain vitality. (C) S. gordonii DL1. Biofilm grows and an increase in the proportion of red cells is seen at 4 h and at 18 h. One representative maximum-projection image from the set of three randomly selectedx-y stacks (square panels) and rotation of the maximum projection to display the x-z perspective (lower panels) are shown. Dimensions of the regions displayed are 100 μm by 100 μm (x-y perspectives; upper panels) and 100 by 30 μm (x-z perspectives; lower panels). The substratum position in the x-z perspective is indicated by the thin white line.
Growth of coculture biofilms.The ability of the three strains to grow as coculture biofilms using unamended 25% saliva as the sole nutrient source was tested by introducing the inoculum of the first strain, washing out unattached cells, introducing the inoculum of the second strain, and again washing out unattached cells. Biofilms were then imaged immediately (0 h) and after 4 and 18 h of saliva delivery. Figure 4 shows results whenS. gordonii was inoculated first, followed by A. naeslundii. To control for variations in biofilm growth of the individual strains due to variations in saliva composition or in cell vitality, monoculture biofilms using the same inocula and the same saliva growth medium were established concurrently with the coculture biofilms. Images of three randomly selected sites were taken. The monoculture of GFP-expressing S. gordonii DL1 (Fig. 4A) exhibited excellent growth (see also Fig. 3C), indicating that it was able to form a biofilm using 25% saliva as the sole nutrient source. Results with A. naeslundii (Fig. 4B) were as demonstrated previously (Fig. 3A); no growth of this strain was seen. Rather, the biomass initially present decreased by 18 h. When the organisms were allowed to interact in coculture (Fig. 4C to E), the ability to coadhere appeared to contribute to the retention of A. naeslundii. At 0 h (Fig. 4C), A. naeslundii showed a propensity to attach directly to, or in the immediate vicinity of, the clusters of S. gordonii, consistent with coadherence. After 4 h of saliva delivery (Fig. 4D), it was apparent thatS. gordonii biomass had increased regardless of its proximity to A. naeslundii cells. The only discernible biomass change visible for A. naeslundii was that cells which coadhered with streptococci were retained in the biofilm. Thus, the biomass of A. naeslundii did not decrease as markedly as in the monoculture biofilm. After 18 h of saliva delivery (Fig.4E), the situation was much like that after 4 h: S. gordonii biomass had increased somewhat, whereas A. naeslundii had not noticeably grown but remained in those areas where it coadhered with the streptococci. Thus, it appears that whenA. naeslundii was added to an established S. gordonii biofilm, it tended to coadhere with the streptococci, and this adherence appeared to offer the advantage of retention (but not of growth) within the biofilm. S. gordonii, on the other hand, grew much as it did in monoculture and was apparently little influenced by the presence of the actinomyces cells.
Time course of biofilm development in coculture ofS. gordonii DL1 and A. naeslundii T14V. (A) S. gordonii monoculture control (constitutive GFP expression). (B) A. naeslundii monoculture control (secondary immunofluorescence). (C) Coculture biofilm at 0 h. Inoculation was with S. gordonii (constitutive GFP expression) followed by A. naeslundii (secondary immunofluorescence, red). A. naeslundii cells are frequently located in direct proximity to S. gordoniicells. (D) Coculture biofilm after 4 h of saliva flow. Growth ofS. gordonii is apparent, but no clear change inA. naeslundii biomass is discernible. (E) Coculture biofilm after 18 h of saliva flow. S. gordoniibiofilm is present, whereas A. naeslundii failed to grow. Yellow regions result from superimposition of red and green during the projection process. In all six subpanels of panels A and B and in the left-hand subpanels of panels C, D, and E, one representative maximum-projection image from the set of three randomly selected x-y stacks (square panels) and rotation of the maximum projection to display the x-z perspective (rectangular panels) are shown. Dimensions of the regions displayed are 250 by 250 μm (x-y perspectives; square panels) and 250 by 73 μm (x-z perspectives; rectangular panels) in panels A and B. The image pairs presented in panels C through E are 250 by 250 μm (x-y perspective; left panel) and 83 by 83 μm (x-y perspectives; right panel); the right panel is a 3× zoom of the center portion of the left panel. For thex-z perspectives, the dimensions are 250 by 73 μm (left panel) and 83 by 24 μm (right panel); i.e., the right panel is a 3× zoom of the left panel.
Results for S. gordonii followed by S. oralis are shown in Fig. 5. The monoculture biofilm of S. gordonii (Fig. 5A) behaved as previously described (Fig. 3C and Fig. 4A). The monoculture biofilm of S. oralis(Fig. 5B) showed some growth between 4 and 18 h, in contrast to monoculture biofilm results described above (Fig. 3B). This likely represents batch-to-batch variation in the concentration of particular salivary components. As noted above, such variability was seen in monoculture planktonic growth of S. oralis. However, when behavior of S. oralis was examined in the coculture biofilm performed with the same inoculum and the same saliva (Fig. 5C to E), no marked growth occurred, much the same as described previously for monoculture biofilms (Fig. 3B). However, the biomass did not appear to decrease over the course of the experiment. The behavior of S. oralis shown in Fig. 5B represents the single case in which growth was seen in a monoculture biofilm out of six trials. Although this growth does not represent the typical behavior of S. oralisin the flowcell system, the images in Fig. 5B were used because they were obtained as part of the same experiment for which the coculture data are shown. In contrast, the behavior of S. gordonii in the coculture biofilm was identical to that in monoculture. It grew on saliva and appeared to be unaffected by the presence of S. oralis. Also, as was the case for the S. gordonii-A. naeslundii experiment, it is clear that many S. oraliscells were in contact with S. gordonii cells at 0 h, indicating coadherence of these coaggregating partners.
Time course of biofilm development in coculture ofS. gordonii DL1 and S. oralis 34. (A)S. gordonii monoculture control (constitutive GFP expression). (B) S. oralis monoculture control (primary immunofluorescence, red). (C) Coculture biofilm at 0 h. Inoculation was with S. gordonii (constitutive GFP expression) followed by S. oralis (primary immunofluorescence). S. oralis cells are frequently located in direct proximity to S. gordonii cells. (D) Coculture biofilm after 4 h of saliva flow. Growth of S. gordonii is apparent, but no clear change in S. oralis biomass is discernible. (E) Coculture biofilm after 18 h of saliva flow. S. gordonii biofilm is present, whereas S. oralis failed to grow. In all six subpanels of panels A and B and in the left-hand subpanels of panels C, D, and E, one representative maximum-projection image from the set of three randomly selected x-y stacks (square panels) and rotation of the maximum projection to display the x-zperspective (rectangular panels) are shown. Dimensions of the regions displayed are as in Fig. 4.
In a coculture with A. naeslundii inoculated first followed by S. oralis, neither strain could grow as a monoculture (Fig. 6A and B), reinforcing the typical behavior pattern for these strains in biofilms. However, when the organisms were grown as a coculture, the pattern was dramatically different. As in the previous coculture results, the strain introduced second (in this case, S. oralis) was frequently located in contact with cells of the first strain (in this case, A. naeslundii) (Fig. 6C). After 4 h of saliva flow, some growth of both strains was apparent (Fig. 6D). It appeared that biomass increased to the greatest extent in regions where the two strains were in immediate proximity, suggesting metabolic cooperation. Isolated single-species microcolonies did not appear to increase in size compared with the previous time point. After 18 h of saliva flow, large mixed-species colonies were obvious, and the two cell types were clearly interdigitated within the colonies. Thus, the developmental patterns of these two strains were drastically different when they were grown as a coculture biofilm compared to those observed when they were grown in monoculture. Neither strain appeared capable of growth on saliva as a monoculture, but both strains flourished when together. Furthermore, growth of each strain appeared to be dependent on coadherence to the other strain and may have been contact dependent. Much growth of the mixed-species colonies occurs in the axial dimension, in stark contrast to the lateral growth of monocultureS. gordonii biofilms. Lastly, growth of these two strains in coculture greatly exceeded growth of S. gordonii, whether in monoculture or in coculture. This suggests that the A. naeslundii-S. oralis pair forms a mutualistic relationship within the biofilm and that S. gordonii cannot participate in such a relationship with either of the other two strains.
Time course of biofilm development in coculture ofA. naeslundii T14V and S. oralis 34. (A)A. naeslundii monoculture control (Syto 59 staining). (B) S. oralis monoculture control (Syto 59 staining). (C) Coculture biofilm at 0 h. Inoculation was with A. naeslundii (secondary immunofluorescence, green) followed byS. oralis (primary immunofluorescence, red). S. oralis cells are frequently located in direct proximity toA. naeslundii cells. (D) Coculture biofilm after 4 h of saliva flow. Growth of both strains is apparent, especially in mixed-species colonies. Note increased interdigitation of the two cell types within the colonies. (E) Coculture biofilm after 18 h of saliva flow. Marked growth of both strains has occurred. Mixed-species colonies dominate the biomass. In all six subpanels of panels A and B and in the left-hand subpanels of panels C, D, and E, one representative maximum projection image from the set of three randomly selected x-y stacks (square panels) and rotation of the maximum projection to display x-z perspective (rectangular panels) are shown. Dimensions of the regions displayed are as in Fig. 4, except for the right x-z perspective in panel E, which is 83 by 48 μm.
DISCUSSION
Unamended saliva was the sole source of nutrient in our studies of oral bacterial biofilms; no sugars, amino acids, vitamins, or other growth supplements were added. Saliva was pooled to minimize variability that may occur in separate saliva collections from a single individual. At least six individuals contributed saliva for each pool, and the pooled saliva was diluted 1:4 to increase the volume available for the flowcell experiments. Different individuals contributed saliva to the various pools, and pools were chosen at random for use in the experiments. Despite the potential for variability, two of the three strains showed remarkably consistent patterns of growth or no growth as biofilms and as planktonic cells. S. gordonii DL1 consistently exhibited planktonic growth to a density of 2 × 107 cells ml−1 in different saliva pools, and growth as a biofilm always occurred.A. naeslundii T14V consistently failed to grow planktonically in saliva or as a monoculture biofilm. In one saliva pool, S. oralis 34 showed no planktonic growth. In other saliva pools, however, it grew but always to a cell density 10- to 100-fold less than that observed for S. gordonii DL1. As a biofilm, S. oralis 34 exhibited growth in only one of six experiments. Thus, each of the three strains had its own growth characteristics and, while saliva may be variable, our overall results indicate reproducible patterns of bacterial growth and demonstrate that unamended saliva is a useful culture medium for oral bacterial biofilms.
Using saliva as the conditioning film and as the sole nutrient source in vitro offers the opportunity to test hypotheses in flowcell-grown biofilms that may be directly applicable to community development on saliva-conditioned surfaces in vivo. In the present study, each strain had different and characteristic properties of attachment and colonization in biofilms. S. oralis bound better to unconditioned glass, whereas both A. naeslundii and S. gordonii attached better to saliva-conditioned glass. In addition, exposure of A. naeslundii and S. gordonii cells to saliva enhanced their adherence. This may be due to binding of salivary components to the cell surface, which in turn increases binding to the conditioning film or salivary pellicle. Also, A. naeslundii may modify its cell surface in response to salivary exposure, as is the case for S. gordonii, which has been shown to upregulate the gene sspAB encoding the surface-exposed salivary binding protein in response to salivary exposure (15). In addition, S. gordonii but notS. oralis binds salivary alpha-amylase (4, 33), and each of the three strains examined in this report binds to saliva-coated hydroxyapatite, a model for the enamel surface of teeth (8, 18, 36). However, only S. gordonii andA. naeslundii bind to latex beads coated with salivary proline-rich protein (11, 18), indicating that bacteria possess specific binding properties for certain salivary components. Collectively, the known adherence properties of these three strains suggest that A. naeslundii and S. gordonii have multiple and strong interactions with salivary molecules, whereasS. oralis has fewer and weaker interactions. Data presented in the present report support these adherence characteristics, and they emphasize the role of prior exposure of bacteria to saliva in altering adherence to both saliva-conditioned and unconditioned surfaces.
All three organisms coaggregate pairwise when suspended in buffer (10, 20) and in saliva (23). The microscopy data presented in this report extend these observations to include pairwise coadherence with these three organisms in biofilms formed in saliva-conditioned glass flowcells. Each pair exhibited preferential juxtapositioning in initial attachment and growth. Our results are in agreement with those of Bos et al. (3), who reported that the ability to coadhere is important in initial recruitment of oral bacteria from the bulk fluid to the early biofilm in vitro. In the present report single bacterial cells can be clearly identified, the direct interaction between the coaggregation-coadherence partners is more evident, and the consequences of these interactions over longer time periods are described. For example, coadherence of cells may play a role in retention in the biofilm. Without S. gordonii,A. naeslundii could not multiply (Fig. 4B, 18 h) and, in fact, the biofilm biomass seemed to decrease with time. However, in the presence of S. gordonii, A. naeslundiicoadhered and was retained (Fig. 4C to E). Although A. naeslundii did not appear to flourish in this situation, retention may provide an opportunity for later interaction by A. naeslundii and an organism such as S. oralis, with which it can establish a mutually beneficial metabolic collaboration. In fact, S. gordonii may act as a template that physically brings these organisms into proximity with one another. Thus, while coaggregation and coadherence bring cells of different species into contact, contact alone does not necessarily result in enhanced growth of the mixed species community.
The proportion of S. gordonii in dental-plaque streptococci that accumulates during the first 4 h in vivo varies greatly, from 0% in some individuals to 22% in others, whereas S. oralis, S. sanguis, and S. mitis biovar 1 are consistently present in high numbers (29). Some other early colonizers such as S. mitis biovar 2 and S. salivarius are more variable and, thus, similar to S. gordonii (29). Four viridans group streptococci (S. sanguis, S. oralis, S. intermedius, and S. gordonii) and A. naeslundii are 5 of the 10 most numerous cultured bacterial species in samples obtained from subgingival dental plaque from healthy sites, but none of these streptococci are among the 10 most frequently cultured species in samples from sites with gingivitis (27). In addition,S. gordonii is more prevalent in supragingival than in subgingival plaque of periodontally diseased adults (39). Clearly, the species composition of the streptococcal population fluctuates considerably across individuals, according to the periodontal health of the host and according to location in the mouth. In our studies, the apparent biofilm accumulation of independent organisms such as S. gordonii using saliva as the sole carbon and nitrogen source was lower than that of mutualistic organisms such as S. oralis-A. naeslundii (compare Fig. 4E or 5E with Fig. 6E). Thus, low-yield organisms such as S. gordonii would be overshadowed when high-yield mutualistic relationships occur because more cells of the high-yield growers would be present. On the basis of its multifaceted binding to salivary pellicle receptors, we propose that S. gordonii is active as an early colonizer, that it assists in establishment of high-yield mutualistic interactions of other species, and that, as a consequence, it becomes reduced in relative abundance in the rapidly developing plaque.
The synergy exhibited by the A. naeslundii-S. oralisinteraction is of major significance to oral bacterial ecology.S. oralis exhibited some growth as a monoculture biofilm in only one saliva pool, and A. naeslundii never showed growth as a monoculture. However, after coadhesion or coaggregation at the saliva-conditioned surface, a mutualistic interaction developed and both strains grew luxuriantly. Clearly, cooperation between these two genera resulted in enhanced growth on saliva-conditioned surfaces. The oral microbial ecosystem offers an attractive model for the discovery of contact-based communication in bacteria because it has evolved over several hundred millennia of pressure towards biofilm formation. Attachment is mandatory or the organisms are swallowed; metabolic cooperation with other species is mandatory or the organisms starve; and swimming motility (as opposed to gliding motility) may be dangerous because the organism loses its foothold on the substratum, or its hold on organisms bound to the tooth surface, and thereby risks removal from the ecosystem. In the case of the A. naeslundii-S. oralis interaction, it appears that direct contact (as opposed to diffusion-based interactions) between the cell types favors development of the mutualism and the subsequent high-yield growth of the coculture, because isolated single-species colonies in the coculture biofilm tended not to develop as well as the coaggregated colonies (Fig. 6D and E). This apparent requirement for contact is the first example of contact-based intergeneric communication in bacteria. Previous studies have dealt with diffusible signals (2), with cell surface sonicate preparations (38), or with intraspecies contact-based communication (19).
Cooperative growth can have a subsequent effect on developmental processes and architecture in multispecies biofilms. In a natural pesticide-degrading consortium, biofilm microcolony architecture was dependent on the concentration and composition of the carbon sources (37). In a defined two-species model system (28), cells from microcolonies of one species migrated to infiltrate the microcolonies of the other species to attain cooperative growth. With nonmotile bacteria it is convenient to assume that biofilm architecture is controlled primarily through attachment and growth and not by change in position of cells once they have attached. The present study is the first to demonstrate an effect of metabolic interaction on biofilm growth and development in a system composed of nonmotile bacteria under constant nutrient conditions.
The in vitro experimental approach taken in the present work will augment in vivo studies of initial attachment, colonization, and subsequent biofilm development on removable enamel chips placed in the oral cavity of human volunteers (32). Insight gained on bacterial interactions within initial dental plaque and on the succession of bacterial species in maturing dental plaque can lead to noninvasive intervention strategies to prevent formation of pathogenic communities and the subsequent tissue destruction characteristic of periodontal diseases.
ACKNOWLEDGMENTS
We thank J. Cisar (NIDCR, NIH) for antibodies and for many helpful discussions. We thank R. Andersen (NIDCR, NIH) for characterizing the coaggregation properties of the GFP-expressing strain, and we thank J. Cisar and P. Egland (NIDCR, NIH) for comments on the manuscript.
Notes
Editor: R. N. Moore
FOOTNOTES
- Received 1 March 2001.
- Returned for modification 18 April 2001.
- Accepted 25 May 2001.
- Copyright © 2001 American Society for Microbiology
