ABSTRACT
Background
Streptococcus salivarius K12 and M18 have recently gained interest as probiotic organisms for caries control. This study investigated the impact of S. salivarius K12 and M18 supplementation on biofilm virulence in a complex saliva-derived in vitro model compared to treatment with Limosilactobacillus reuteri DSM17938 and no treatment.
Methods
Biofilms were grown under low (0.05%) and high (1%) sucrose conditions and supplemented with S. salivarius K12, S. salivarius M18 or L. reuteri. Microscale pH dynamics were assessed using confocal microscopy-based pH ratiometry. The biofilm matrix composition was analyzed by glucan and extracellular DNA mapping. Microbial community composition was determined by 16S rRNA gene amplicon sequencing.
Results
At 0.05% sucrose, pH was significantly higher in S. salivarius M18 biofilms, but not in S. salivarius K12 biofilms, compared to control. pH in L. reuteri biofilms was significantly lower. At 1% sucrose, no significant effects of probiotic supplementation on biofilm pH were observed. Shifts in the bacterial composition were minor, most notably an increased abundance of lactobacilli in L. reuteri-treated biofilms at 1% sucrose. No significant effects on the biofilm matrix composition were observed.
Conclusions
In conclusion, probiotic supplementation exerted only minor effects on biofilm composition and virulence.
KEYWORDS: Biofilm, biofilm matrix, dental caries, extracellular DNA, glucans, extracellular polymeric substance, pH ratiometry, probiotics
Introduction
Probiotic organisms have emerged as an adjunctive strategy for dental caries control, primarily because of their ability to modulate the oral microbiome and inhibit cariogenic species [1]. Early investigations focused on organisms traditionally associated with gastrointestinal health, such as Lactobacillus spp. and Bifidobacterium spp., which were administered through dairy products or lozenges [2–4]. Although some studies reported promising results, such as reductions in salivary Streptococcus mutans counts and general shifts in the oral microbiota, their clinical relevance has been questioned. The employed species were originally intended for gastrointestinal use and exhibited poor capacity to colonize and persist within dental biofilms, restricting their long-term ecological impact in the oral cavity [3,5–7]. This limitation underscores the importance of selecting organisms that are ecologically compatible with the oral environment and capable of integrating into existing biofilm communities when identifying probiotic candidates for caries prevention. Consequently, research attention has shifted towards microorganisms that are natural constituents of the oral microbiota, particularly Streptococcus spp., which may integrate more effectively into established biofilm communities. Among these, Streptococcus salivarius, originally introduced for respiratory health benefits, has recently gained interest as a potential candidate for caries control [8,9]. Specifically, strains S. salivarius K12 and S. salivarius M18 produce multiple bacteriocins with antimicrobial activity that may suppress cariogenic organisms within dental biofilms [10,11]. In addition, both organisms have been shown to produce urease and dextranase, enzymes that may contribute to reduced biofilm acidity and structural stability, respectively [12].
Clinical evidence regarding the effects of S. salivarius K12 and M18 on oral biofilms remains limited. A small number of clinical trials have evaluated their impact on dental plaque accumulation, with inconsistent results. While two trials reported significant reductions in plaque scores upon daily consumption, another study found no significant differences in plaque formation or the salivary levels of S. mutans [8,13,14]. Importantly, these investigations primarily focused on clinical plaque indices and salivary bacterial counts, without assessing ecological parameters that are directly linked to the caries process. As of today, there is a lack of clinical or laboratory data examining the influence of S. salivarius strains K12 and M18 on biofilm pH dynamics, bacterial composition or biofilm matrix composition. This gap limits our understanding of their potential mechanisms of action within dental biofilms.
To address this lack of evidence, further investigation of the ecological effects of S. salivarius K12 and M18 on dental biofilms is warranted. The present study, therefore, aimed to evaluate the influence of these probiotic strains on biofilm virulence in a complex salivary model of dental biofilm, with particular emphasis on ecological and structural characteristics linked to cariogenic potential. Specifically, we assessed the effect of the two strains on microscale pH by confocal-microscopy-based pH ratiometry, compared to treatment with Limosilactobacillus reuteri DSM 17938 and no treatment. Moreover, we determined the bacterial composition of the biofilms using bacterial 16S rRNA gene sequencing. Lastly, we assessed the effect of probiotic supplementation on the production of glucose-based matrix carbohydrates and the abundance of extracellular DNA (eDNA). Our null hypotheses were that treatment with S. salivarius K12 and M18 had no significant effect on a) the bacterial composition of the biofilms, b) biofilm pH, and c) the production of biofilm matrix components, compared to treatment with L. reuteri DSM 17938 and no treatment.
Materials and methods
Bacterial cultures
Streptococcus salivarius subsp. salivarius K12 (Burgerstein Biotics-O, Burgerstein, Rapperswil-Jona, Switzerland), S. salivarius subsp. salivarius M18 Throat Guard® (BLIS Technologies, Wellington, New Zealand) and Limosilactobacillus reuteri DSM 17938 (Sunstar GUM® PERIOBALANCE®, Gothenburg, Sweden) were cultured from lozenges that were suspended in 3 mL Brain Heart Infusion (BHI, Thermo Fisher Scientific, Roskilde, Denmark) for 1.5 h at 37 °C under anaerobic conditions (10% H2, 10% CO2, 80% N2; Whitley A35 Anaerobic Workstation; Don Whitley Scientific, Bingley, UK). Bacterial suspensions of each strain were plated on Columbia blood agar (CBA, Thermo Fisher Scientific, Roskilde, Denmark) and incubated for 24 h at 37 °C under anaerobic conditions. Then colonies were removed with sterile cotton tip applicators (6 inch; Avaline Medical, Lakewood, USA), transferred to sterile freezing vials (2.0 mL; Sigma-Aldrich, Brøndby, Denmark) and frozen at –80°C in BHI containing 20% glycerol (Thermo Fisher Scientific, Roskilde, Denmark) until experimental use.
Saliva collection and biofilm growth
Stimulated saliva was collected from healthy volunteers without oral diseases, pooled and then used as the inoculum for biofilm growth and as part of the growth medium. The use of pooled stimulated saliva as an inoculum is a well-established approach in in vitro biofilm research, as it provides a standardized and compositionally diverse microbial community representative of the oral microbiota while minimizing site-specific variability [15,16].
Saliva collection was considered exempt from ethical review by the Ethical Committee of Region Midtjylland (1-10-72-109-23). All donors provided written informed consent. Pooled saliva was used both as the inoculum for biofilm formation and as a component of the growth medium to support the establishment of a complex salivary biofilm. Biofilms were grown for 72 h in 96-well plates for microscopy (µ-Plate 96 Well Square, 1.5 Polymer cover slip; Ibidi, Gräfelfing, Germany), under anaerobic conditions (10% H2, 10% CO2, 80% N2; Whitley A35 Anaerobic Workstation), at 37 °C with moderate shaking (120 rpm). Oxygen penetration into biofilms is limited due to diffusion constraints and bacterial respiration, resulting in anaerobic niches within deeper biofilm layers [17,18].
Therefore, anaerobic incubation was chosen to approximate the metabolic conditions of mature dental plaque. The wells were inoculated with 20 µL of whole saliva in 180 µL of medium containing 80% sterile saliva, 20% fetal bovine serum (FBS, Thermo Fisher Scientific, Roskilde, Denmark), and either 0.05% or 1% (w/v) sucrose. A sucrose concentration of 1% was used to mimic a cariogenic environment known to promote extracellular polysaccharide synthesis and acidogenicity in biofilm models [19,20]. A reduced sucrose condition (0.05%) was included to investigate a lower carbohydrate exposure scenario. Biofilms grown for glucan mapping (see below) were supplemented with fluorescently labelled dextran (1 µM; 10,000 MW; Alexa Fluor 647; Thermo Fisher Scientific) in the medium.
Probiotic treatment
Probiotic supplementation to the biofilm was performed at 24 and 48 h to simulate the recommended clinical application regimen of probiotic lozenges, which are typically administered daily over consecutive days [13,14]. Prior to probiotic treatment, S. salivarius subsp. salivarius M18, K12 and L. reuteri DSM 17938 were grown on CBA under anaerobic conditions at 37 °C for 24 h. Two colonies from each plate were suspended in 2 mL BHI and incubated at 37 °C for 4 or 14 h, respectively. Next, bacterial suspensions were centrifuged at 9200xg for 5 min at room temperature. The bacterial pellets were resuspended in 2 mL sterile saline (0.9%) and adjusted to 1 × 107 CFU/mL (OD550 = 0.5 ± 0.05). After 24 h of saliva-derived biofilm growth, 100 µL of the supernatant (spent medium) was removed and replaced with 80 µL of fresh medium and 20 µL of one of the probiotic strains in the presence of 0.05% or 1% (w/v) sucrose. In control biofilms, 100 µL of the medium was replaced with 80 µL of fresh medium and 20 µL of sterile saline. This partial medium exchange was performed to simulate repeated probiotic exposure while preserving the established biofilm structure. The procedure was repeated after 48 h. After 72 h of growth, biofilms were subjected to analysis.
DNA extraction and bacterial 16S rRNA gene sequencing
The biofilms were removed from the 96-well plates by adding 100 µL of Power Bead solution (DNeasy PowerLyzer PowerSoil; Qiagen, Venlo, the Netherlands) to each well. Next, a 20-min incubation at room temperature and 10 min of sonication (40 kHz; Branson 5510 DTH Ultrasonic Cleaner, Urdorf, Switzerland) were performed. The detached biofilm suspension was transferred into a Power Bead tube (DNeasy PowerLyzer PowerSoil; Qiagen), and the volume was adjusted to 750 µL with additional Power Bead solution. DNA extraction was carried out according to the DNeasy PowerLyzer PowerSoil kit instructions. The V3–V4 region of the bacterial 16S rRNA gene was amplified using primers Bac 341F and Bac 805R and prepared for sequencing according to Illumina’s 16S Metagenomic Sequencing Library Preparation protocol [21]. Paired-end sequencing (2 × 300 bp) was performed on an Illumina MiSeq platform with the V3 sequencing kit (Illumina, San Diego, CA, USA).
All analyses were done in R (v4.5.2). Amplicon sequence variants (ASVs) were inferred using DADA2 (v.1.38.1) [22]. The taxonomic assignment was based on the Silva SSU reference database version 138 [23]. DNA extraction blanks and PCR negative controls were included for decontamination using the R package Decontam (v.1.30.0) with the prevalence method and a threshold of 0.1 [24]. Principal coordinate analysis (PCoA) and PERMANOVA were performed on CLR-transformed read counts using vegan (v2.7-2) [25]. Differential abundance analysis between treatment groups was done separately on the 1% sucrose and 0.05% sucrose samples using the R-package ANCOM-BC (v2.12.0) [26].
ASVs that were classified as either Streptococcus or Lactobacillus were BLASTed against the Human Oral Microbiome Database (eHOMD 16S rRNA RefSeq Version 15.22) to obtain a species-level classification [27]. S. salivarius ASV_47 and L. reuteri ASV_6, 12, 14, 15, 16 and 17 were also BLASTed against the NCBI database to verify if they originated from the probiotic strains used in this study.
All experiments were conducted in biological triplicate. The raw sequencing data have been deposited in NCBI under BioProject PRJNA1398072.
Confocal-microscopy-based pH ratiometry
The biofilms were washed twice with sterile saline (0.9%, pH 7) to remove planktonic cells and the spent medium. Then, the biofilm pH response to a 4% (w/V) sucrose challenge was monitored using the ratiometric dye C-SNARF-4 (30 µM; Thermo Fisher Scientific), as described previously. pH was recorded after 15 min (t1) and 45 min (t2) of incubation at 37 °C (Stage Top Incubation System; Ibidi, Gräfelfing, Germany), in six pre-defined microscopic fields of view (FOVs), at the biofilm base and top using a confocal microscope (Zeiss LSM 700, Jena, Germany). A detailed description of imaging parameters, digital image analysis and the calibration of the ratiometric dye is presented in the Supplementary Methods. Experiments were performed in biological triplicates [28,29].
Glucan and eDNA mapping
Glucans were visualized by the incorporation of fluorescently labelled dextran in the growth medium. The biofilms were fixed in 3% paraformaldehyde (3 h, 4 °C), washed twice in phosphate-buffered saline and counterstained with SYTO 41 (10 µM; 10 min; Thermo Fisher Scientific) [26]. For eDNA mapping, fresh biofilms were washed with sterile saline (0.9%) and stained using the membrane-impermeable dye TOTO-1 (2 µM; 10 min; Thermo Fisher Scientific). Bacterial cells were counterstained with SYTO 60 (10 µM; 10 min; Thermo Fisher Scientific) [30].
For each biofilm, z-stacks with three slices spanning the height of the biofilm were acquired in six pre-defined FOVs. Imaging parameters are described in detail in the Supplemental Methods. Images for glucan and eDNA mapping were segmented in the software daime (digital image analysis in microbial ecology, v.2.0), and both the bacterial and the biofilm matrix biovolumes were quantified for each employed matrix stain, as described [15,31]. For eDNA, both total matrix biovolumes and intercellular matrix biovolumes, i.e. biovolumes that did not overlap with bacterial cells, were determined. All experiments were performed in biological triplicates.
Statistical analysis
Differences in biofilm thickness, as recorded in the confocal microscopy stacks, were compared between the different biofilms (control biofilms and biofilms spiked with different probiotic organisms) using Kruskal-Wallis tests.
The effect of probiotic supplementation on biofilm pH was analyzed using linear mixed-effects models, with replicate, FOV (nested in replicate) and biofilm layer (nested in FOV) as random effects, for each sucrose level and time point separately. P-values were adjusted using Holm’s method, after which p-values < 0.05 were considered statistically significant. Pre-specified comparisons of control biofilms versus biofilms spiked with S. salivarius K12, M18, and L. reuteri were examined using Tukey-adjusted contrasts.
The effect of probiotic supplementation and sucrose on the amounts of eDNA and glucan in the biofilms was analyzed with linear mixed-effects models, with replicate and FOV (nested in replicate) as random effects. To approximate normality, outcome variables were logit-transformed. Fixed effects were evaluated at a significance level of α = 0.05. Pre-specified comparisons of control biofilms versus biofilms supplemented with S. salivarius K12, M18 and L. reuteri were examined within each sucrose level using Holm-adjusted contrasts. For these multiple comparisons, adjusted p-values < 0.05 were considered statistically significant. All analyses were conducted in R (version 4.5.2) using the packages lme4, lmerTest and emmeans.
Results
Biofilm formation
Biofilm formation in the model was robust, with no significant differences in average biofilm thickness (31.6 µm ± 6.5 µm SD) between control biofilms and biofilms spiked with probiotic strains (0.05% sucrose: p = 0.74; 1% sucrose: p = 0.46; Table S1; Kruskal-Wallis).
Biofilm composition
Biofilms grown in the presence of 1% sucrose were primarily colonized by streptococci and, to a minor extent, by lactobacilli and Rothia spp., with average abundances of 69.1 ± 16.1% SD, 23.4 ± 19.7% SD and 3.5 ± 1.8% SD, respectively. Biofilms grown with 0.05% sucrose in the medium showed a more diverse composition, with Streptococcus (46.3 ± 15.0%), Veillonella (14.2 ± 4.7%), Fusobacterium (5.9 ± 8.4%), Peptostreptococcus (3.4 ± 5.1%) and Prevotella (2.9 ± 1.6%) as the most abundant genera (Figure 1A).
Figure 1.
The effect of probiotic supplementation on bacterial biofilm composition. The 72-h salivary biofilms, grown in the presence of 0.05% or 1% sucrose, were spiked with S. salivarius subsp. salivarius M18, K12 or L. reuteri. The bacterial biofilm composition was determined by next-generation sequencing. (A) Heatmap illustrating the relative abundance of different genera. Biofilms grown in the presence of 0.05% sucrose showed a diverse composition dominated by typical oral genera. The biofilms grown in the presence of 1% sucrose were dominated by streptococci and, to a lesser extent, by lactobacilli. (B) and (C) Principal coordinate analysis (PCoA) showed that the composition of biofilms grown with 1% sucrose was rather homogenous and differed markedly from biofilms grown with 0.05% sucrose (PCoA1). Biofilms spiked with L. reuteri were compositionally different from the other treatment groups (PCoA3). The composition of biofilms grown with 0.05% sucrose differed between replicates but not between treatment groups (PCoA2).
At 1% sucrose, the overall biofilm composition differed between treatment groups and not between replicates (p = 0.001 and p = 0.78, respectively; PERMANOVA). Biofilms spiked with L. reuteri, especially, showed a higher abundance of lactobacilli (54.8 ± 11.3% SD vs. 9.7 ± 0.9% SD) and a lower abundance of streptococci (43.6 ± 10.2% SD vs. 79.7 ± 0.9% SD) than control biofilms (Figure 1A–C). Pairwise contrasts between treatment groups, however, were not significant (p = 0.1; PERMANOVA, post-hoc). At 0.05% sucrose, no differences in the biofilm composition were observed between treatment groups (p = 0.89; PERMANOVA), but the biofilm composition differed between biological replicates (p = 0.001; Figure 1A‒C). The null hypothesis (a) was therefore partially rejected.
None of the ASVs that were present in all treatment groups was differentially abundant in biofilms grown in either 1% or 0.05% sucrose (ANCOM-BC). However, some ASVs with a mean relative abundance of more than 0.1% were completely absent in control biofilms and/or in at least one of the three treatment groups spiked with probiotic organisms. In biofilms grown in the presence of 0.05% sucrose, 14 ASVs were absent in at least one of the four groups, and in biofilms grown with 1% sucrose, 10 ASVs were absent from at least one of the groups (Figure S1). ASV 47 showed 99.8% sequence identity with S. salivarius (BLAST) and was only present in biofilms spiked with S. salivarius M18. ASVs 6, 12, 14, 15 and 17 showed 100% or 99.8% sequence identity with L. reuteri and were only present in biofilms spiked with L. reuteri DSM17938. These ASVs, therefore, likely represent the spiked organisms that colonized the biofilm model.
Biofilm pH
The average pH drops in the model were moderate for biofilms grown in the presence of 0.05% sucrose (6.26 ± 0.09 SD), and steeper for biofilms grown with 1% sucrose (5.44 ± 0.32 SD; Figure 2A). For both growth conditions, average pH drops were most pronounced in the first 15 min after the sucrose challenge (t1: 0.05% sucrose = 6.31 ± 0.08 SD; 1% sucrose = 5.46 ± 0.32 SD; Figure S1). In the following 30 min, pH remained fairly stable (t2: 0.05% sucrose = 6.21 ± 0.1 SD; 1% sucrose = 5.43 ± 0.03 SD; Figure S1). The biofilm pH varied considerably between individual FOVs in a biofilm (0.05% sucrose: maximum range = [6.04; 6.52]; 1% sucrose: maximum range = [5.16; 5.68]. The pH gradients between the biofilm top and bottom, in contrast, were moderate (Figure S2).
Figure 2.
The effect of probiotic supplementation on biofilm pH. Biofilms, grown in the presence of 0.05% or 1% sucrose (SUC), were spiked with S. salivarius subsp. salivarius M18, K12 or L. reuteri. The pH response to a sucrose challenge (4%) was determined ratiometrically after 45 min (t2). (A) The pH was significantly lower in 1% SUC than in 0.05% SUC biofilms. At 0.05% SUC, a significant effect of S. salivarius M18supplementation on biofilm pH was observed. Boxplots show medians and the interquartile range, with whiskers indicating the standard deviation. (B) Representative images illustrating extracellular biofilm pH. The black areas represent bacterial cells that were removed during image analysis. False colouring was applied for visualization purposes. Scale bars = 20 µm.
For biofilms grown in the presence of 0.05% sucrose, pH differed between treatment groups (t1 and t2: p < 0.001). Pairwise comparisons demonstrated lower pH in biofilms supplemented with L. reuteri compared to control (t1: p = 0.03; t2: p < 0.0001), higher pH in biofilms supplemented with S. salivarius M18 (t1: p = 0.02; t2: p = 0.006), and no significant differences between S. salivarius K12 and control biofilms (t1: p = 0.73; t2: p = 0.88). In biofilms grown with 1% sucrose, no significant pH differences were observed between treatment groups (t1: p = 0.07; t2: p = 0.81). The null hypothesis (b) was therefore partially rejected. Representative images of biofilm pH at t2 are shown in Figure 2B.
Biofilm matrix composition
eDNA volumes in the biofilms were not significantly affected by the amount of sucrose in the medium, nor by the supplementation of probiotic strains or the interaction between both factors (p = 0.09, p = 0.42 and p = 0.44, respectively; Figure 3A). Likewise, pairwise comparisons between control biofilms and biofilms spiked with probiotic strains showed no significant differences in the amount of eDNA for 0.05% and 1% sucrose (Table S2). Similar results were observed for intercellular eDNA biovolumes (Fig. S3). Representative biofilm images are shown in Figure 3A.
Figure 3.
The effect of probiotic supplementation on the biofilm matrix. Biofilms, grown in the presence of 0.05% or 1% sucrose (SUC), were spiked with S. salivarius subsp. salivarius M18, K12 or L. reuteri. The relative amounts of extracellular DNA (eDNA) and matrix glucans, normalized to the bacterial cell volumes, were determined by confocal microscopy. (A) eDNA biovolumes in the biofilms were not significantly affected by the amount of sucrose in the growth medium or by supplementation of the probiotic organisms S. salivarius subsp. salivarius M18, K12 or L. reuteri. Error bars = SD. The lower panels show representative biofilm images. Bacterial cells are shown in red, eDNA is shown in green. Bars = 20 µm. (B) Across all biofilms, glucan biovolumes were significantly higher for biofilms grown with 1% SUC, although an opposite trend was observed for L. reuteri. No significant differences between control biofilms and biofilms spiked with probiotic organisms were observed. Error bars = SD. The lower panels show representative biofilm images. Bacterial cells are displayed in blue, and glucans in red. Bars = 20 µm.
The relative amount of glucans in the biofilms was significantly affected by the concentration of sucrose in the growth medium, with higher amounts at 1% sucrose (0.05% SUC: 13.6%, CI [3.1%, 44.6%]; 1% SUC: 76.1%, CI [38.6%, 94.1%], p = 0.013). Biofilms spiked with L. reuteri showed an opposite, but non-significant trend (p = 0.78; Figure 3B). Neither probiotic supplementation (p = 0.90) nor the interaction between probiotic supplementation and the sucrose concentration in the medium (p = 0.66) had a significant effect on glucan volumes in the biofilms. Pairwise comparisons between control biofilms and biofilms exposed to probiotic organisms showed no significant differences in glucan biovolumes (Table S2). The null hypothesis (c) was therefore not rejected. Representative images are displayed in Figure 3B.
Discussion
Probiotic-based strategies for caries prevention aim to modulate microbial interactions, and thereby biofilm composition and acidogenicity [1]. The present study addressed a critical gap in the literature by investigating the effects of the oral commensal probiotics S. salivarius K12 and M18 on microscale pH, biofilm matrix composition, and microbial community structure in a complex salivary biofilm model, compared to treatment with L. reuteri DSM 17938 and no treatment.
Within the limits of this in vitro study, probiotic supplementation with S. salivarius K12 or M18 only resulted in minor changes in biofilm composition and virulence. The effects of the two strains on the global biofilm composition at the genus level were negligible, both in biofilms grown in the presence of 1% sucrose and 0.05% sucrose (Figure 1). While S. salivarius K12 may not have colonized the biofilms despite repeated spiking, S. salivarius M18 seemed to be present in the model (ASV 47), albeit with low relative abundance (Figure S1). Compared to control biofilms, no significant effects of the two strains on the biofilm matrix composition were observed (Figures 3, S3). The biovolumes of both eDNA and glucans remained largely unchanged; however, in line with previous studies, they exhibited substantial variability among different FOVs within the biofilms, leading to large standard deviations and a reduced statistical power to detect small differences (Table S1) [32,33].
In biofilms grown in the presence of 1% sucrose, no significant pH differences were observed between biofilms spiked with either S. salivarius K12 or M18 and control biofilms. In contrast, a small but significant effect of supplementation with S. salivarius M18 was observed in biofilms grown in the presence of 0.05% sucrose (Figures 2, S2). This effect could potentially derive from the organism’s urease activity, which has the potential to increase biofilm pH through the production of ammonia [34]. Whether the observed findings translate into relevant pH changes in a clinical setting, however, remains to be determined.
The microscale pH is a key functional indicator of biofilm virulence because it reflects localized acid production and the potential for ecological shifts toward cariogenic communities. According to the Integrated Plaque Hypothesis, persistent low-pH microenvironments select for acid-tolerant organisms and promote enamel demineralization, highlighting that even subtle changes in microscale pH can be biologically important [35].
While microscale pH reflects acidogenic potential and cariogenicity, biofilm virulence is multifactorial and may also involve proteolytic activity, metabolic cross-feeding, and competitive interactions within the community. Proteolytic degradation of salivary and host-derived proteins can influence nutrient availability and contribute to ecological shifts in biofilm composition independent of acidogenicity [36,37]. Future studies incorporating proteolytic assays and transcriptomic analyses could therefore provide deeper mechanistic insight into probiotic-mediated modulation of biofilm virulence.
Regarding the predefined null hypotheses, probiotic supplementation did not uniformly affect all assessed parameters of biofilm virulence. While certain strain- and condition-specific effects were observed, particularly under reduced sucrose exposure, the overall impact on biofilm composition, pH dynamics, and matrix-associated components was limited. Therefore, the null hypothesis of no effect could only be partially rejected. These findings suggest that the ecological influence of probiotic supplementation in complex salivary biofilms is strain-dependent and modulated by environmental conditions, rather than resulting in broad alterations of biofilm virulence [10,11]. Our experimental approach was motivated by the inconsistent clinical evidence regarding the efficacy of probiotic S. salivarius strains in reducing dental plaque and biofilm virulence. Thus, clinical trials to date have reported mixed results regarding the effects of S. salivarius K12 and M18 on dental plaque formation. In a randomized double-blind, placebo-controlled trial, Burton et al. reported that daily consumption of S. salivarius M18 for three months reduced dental plaque scores in children [13]. Similarly, a pilot randomized clinical trial observed a significant reduction in plaque accumulation after four weeks of daily S. salivarius K12 lozenge use in healthy adults [8]. In contrast, a randomized, double-blind, placebo-controlled trial suggested that S. salivarius K12 was not effective at improving the plaque index or the salivary levels of S. mutans in adolescent orthodontic patients [14]. These inconsistencies underscore the need for standardized in vitro systems capable of addressing key parameters of biofilm virulence, such as microscale pH and the composition of the biofilm matrix, that are not captured by conventional clinical endpoints. While a previous in vitro study focused on the antibacterial effects of S. salivarius strains K12 and M18 against S. mutans [10], the present study extended this approach by integrating a complex salivary biofilm model with spatially resolved pH analysis.
Interestingly, supplementation with the widely investigated probiotic organism L. reuteri had more pronounced, but negative effects on the biofilm model than treatment with the two S. salivarius strains. Spiking with L. reuteri resulted in substantial colonization of the biofilms and a shift from streptococci towards lactobacilli, especially when biofilms were grown with 1% sucrose. As a result, glucan biovolumes were somewhat reduced, which may reflect that lactobacilli are less efficient producers of extracellular carbohydrates than streptococci and rely more on intracellular carbohydrate storage [38]. When grown with 0.05% sucrose in the medium, the pH in biofilms spiked with L. reuteri was significantly lower than the pH in control biofilms and in biofilms supplemented with the two streptococcal strains (Figure 2). Although the differences were small, these results raise the question whether lactobacilli, which are both potent acid producers and highly aciduric [39,40], are suitable candidates for the probiotic modulation of dental biofilms in a caries context.
The biofilm model employed in this study harboured a variety of typical oral organisms, which allowed assessing the effects of probiotic supplementation in a biologically relevant context [41]. Overall, model reproducibility was high when the medium was supplemented with 1% sucrose, as evidenced by consistent biofilm growth, bacterial composition and pH profiles across replicates (Figures 1 and 2). In contrast, increased variability in the microbial composition was observed between replicates at 0.05% sucrose. This reduced reproducibility likely reflects a greater sensitivity under low-sucrose conditions to minor fluctuations in microbial activity and nutrient availability, highlighting a limitation of the model [42,43].
While our study captured microscale pH changes and biofilm matrix composition, the relatively short experimental period limits extrapolation to long-term biofilm dynamics observed in clinical trials. Further investigation into gene expression and more detailed EPS composition analysis could provide additional mechanistic insight into how probiotic supplementation modulates biofilm virulence. Furthermore, while the employed model is compositionally complex, it does not fully mimic intraoral conditions. The biofilms in the model grew without mechanical shear, salivary flow and exposure to host immune factors. Additionally, the probiotic organisms, but also the sucrose in the growth medium, were supplemented constantly and not as pulses, which may not reflect a clinically relevant dosage. The extrapolation of our findings to a clinical context should therefore be performed with caution.
In conclusion, the present study demonstrated that supplementation with the probiotic organisms
S. salivarius K12 and M18 only had a minor impact on the biofilm composition and virulence in a complex, saliva-based model of dental biofilm. A significant positive effect on biofilm pH was observed for S. salivarius M18, under low-sucrose conditions compared to the other groups.
Supplementary Material
Figure S1.tif
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Table S2_NEW.docx
Supplementary Methods_20251230.docx
Table_S1_NEW.docx
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Acknowledgements
The authors would like to thank Anette Aakjær Thomsen (Aarhus University, Denmark) for substantial laboratory support. The authors would also like to express their sincere thanks to Professor Rikke Louise Meyer (Interdisciplinary Center for Nanoscience, Aarhus University, Denmark) for kindly providing access to the confocal microscope used for image acquisition.
All authors have read and agreed to the published version of the manuscript.
Funding Statement
This work was supported by Aarhus University, Denmark.
Disclosure statement
Elisabeth Reichardt is a co-inventor of a pending patent application related to probiotic strains described in this manuscript (AG1678 PCT BS). The other authors report no competing interests.
Ethical approval
The collection of pooled saliva used in this study was considered exempt from ethical review by the Ethical Committee of Region Midtjylland (1-10-72-109-23). All donors provided written informed consent.
Supplementary material
Supplemental data for this article can be accessed at https://doi.org/10.1080/20002297.2026.2682457.
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