l-Arginine Modifies the Exopolysaccharide Matrix and Thwarts Streptococcus mutans Outgrowth within Mixed-Species Oral Biofilms

ABSTRACT

l-Arginine, a ubiquitous amino acid in human saliva, serves as a substrate for alkali production by arginolytic bacteria. Recently, exogenous l-arginine has been shown to enhance the alkalinogenic potential of oral biofilm and destabilize its microbial community, which might help control dental caries. However, l-arginine exposure may inflict additional changes in the biofilm milieu when bacteria are growing under cariogenic conditions. Here, we investigated how exogenous l-arginine modulates biofilm development using a mixed-species model containing both cariogenic (Streptococcus mutans) and arginolytic (Streptococcus gordonii) bacteria in the presence of sucrose. We observed that 1.5% (wt/vol) l-arginine (also a clinically effective concentration) exposure suppressed the outgrowth of S. mutans, favored S. gordonii dominance, and maintained Actinomyces naeslundii growth within biofilms (versus vehicle control). In parallel, topical l-arginine treatments substantially reduced the amounts of insoluble exopolysaccharides (EPS) by >3-fold, which significantly altered the three-dimensional (3D) architecture of the biofilm. Intriguingly, l-arginine repressed S. mutans genes associated with insoluble EPS (gtfB) and bacteriocin (SMU.150) production, while spxB expression (H₂O₂ production) by S. gordonii increased sharply during biofilm development, which resulted in higher H₂O₂ levels in arginine-treated biofilms. These modifications resulted in a markedly defective EPS matrix and areas devoid of any bacterial clusters (microcolonies) on the apatitic surface, while the in situ pH values at the biofilm-apatite interface were nearly one unit higher in arginine-treated biofilms (versus the vehicle control). Our data reveal new biological properties of l-arginine that impact biofilm matrix assembly and the dynamic microbial interactions associated with pathogenic biofilm development, indicating the multiaction potency of this promising biofilm disruptor.

IMPORTANCE Dental caries is one of the most prevalent and costly infectious diseases worldwide, caused by a biofilm formed on tooth surfaces. Novel strategies that compromise the ability of virulent species to assemble and maintain pathogenic biofilms could be an effective alternative to conventional antimicrobials that indiscriminately kill other oral species, including commensal bacteria. l-Arginine at 1.5% has been shown to be clinically effective in modulating cariogenic biofilms via alkali production by arginolytic bacteria. Using a mixed-species ecological model, we show new mechanisms by which l-arginine disrupts the process of biofilm matrix assembly and the dynamic microbial interactions that are associated with cariogenic biofilm development, without impacting the bacterial viability. These results may aid in the development of enhanced methods to control biofilms using l-arginine.

INTRODUCTION

Biofilms formed on surfaces are highly organized and structured microbial communities enmeshed in an extracellular matrix composed of polymeric substances, such as exopolysaccharides (EPS) (1, 2). Many human infections are caused and/or exacerbated by virulent biofilms, including those occurring in the mouth (3). Among them, dental caries is one of the most prevalent and costly biofilm-dependent infectious diseases worldwide (4). The assembly of caries-producing (cariogenic) biofilms on saliva-coated teeth is a prime example of the consequences arising from interactions between bacteria (and their products) and diet (e.g., sugars) that promotes microbial accumulation as an EPS-rich matrix develops (5). EPS are key matrix components that act as a supportive three-dimensional (3D) scaffold and barrier to diffusion, modulating growth of and providing protection to pathogens (1, 2, 6). A major challenge in biofilm-related diseases is that the embedded bacteria become recalcitrant to antimicrobials, making them difficult to remove without disturbing the normal flora.

In the mouth, a highly diverse microbial community is constantly interacting with the salivary pellicle present on the tooth surface, to which early colonizers, including viridans group streptococci and Actinomyces spp., can adhere and coadhere with other organisms (7, 8). Cariogenic bacteria, such as Streptococcus mutans, can be also present in this initial colonizing community, although in varied numbers, which is dependent on the host diet (9). However, environmental changes, such as frequent sucrose intake, can serve as the catalyst for cariogenic biofilm formation, since this sugar serves as a substrate for the production of both EPS and acids (5). S. mutans can rapidly orchestrate the assembly of virulent biofilm via EPS synthesis by exoenzymes (e.g., glucosyltransferases) bound on the pellicle and on bacterial surfaces (10–12). The EPS formed in situ enhances the local accumulation of bacteria on teeth via glucan-binding mechanisms while enmeshing them in a diffusion-limiting matrix (2, 10, 13–15). This ultimately creates highly adherent and cohesive biofilm architecture that shelters and protects resident organisms (2, 6, 14–16). In parallel, sugars are fermented by bacteria within biofilms, creating acidic microenvironments. The low-pH niches induce EPS synthesis and cause ecological shifts by favoring the growth of acidogenic and acid-tolerant bacteria. Consequently, local acidity ensures cariogenic biofilm accretion and acid dissolution of the adjacent enamel, leading to the onset of caries (17). Thus, novel strategies that compromise the ability of virulent species to assemble and maintain pathogenic biofilms could be alternatives to conventional broad-spectrum antimicrobials, which can indiscriminately kill other oral species, including commensal bacteria.

l-Arginine has been shown to potentially modulate cariogenic biofilms. In the oral cavity, arginine, mainly derived from diet and host saliva, is utilized by arginolytic bacteria (e.g., S. gordonii) via the arginine deiminase system (ADS) with end products, such as ammonia (18). Due to the pH raising effect of ammonia, arginine has been regarded as a potential endogenous caries modulator (18–21). Indeed, the levels of free arginine in the saliva of caries-free individuals were significantly higher than those in the saliva of those with caries (22), and a positive correlation between ADS activity in plaque biofilms and absence of caries has been clinically validated in adults (23) and children (24). These findings suggest the possibility of arginine supplementation as a strategy to disrupt pathogenic biofilms. In vitro studies show that exogenous l-arginine can inhibit the adhesion of S. mutans to saliva-coated surfaces (25), influence single-species S. gordonii biofilm formation (26), and destabilize a saliva-derived biofilm community (27). Arginine treatments also increased arcA gene (involved in arginine metabolism) abundance compared to that in untreated biofilms, indicating a potential role for the enrichment of alkali-producing microflora (28).

The available laboratory and clinical evidence supports exogenous l-arginine supplementation at the concentration of 1.5% (wt/vol) as a feasible adjunctive anticaries approach via ADS activity (28–31). However, l-arginine may inflict additional changes in the biofilm milieu under conditions conducive to dental caries. Here, we provide new insights on how topical l-arginine exposure modulates biofilm development using a mixed-species model containing both cariogenic (S. mutans) and arginolytic (S. gordonii) bacteria under sucrose challenge. We found that l-arginine thwarts S. mutans outgrowth and promotes S. gordonii dominance while maintaining A. naeslundii growth within sucrose-grown biofilms. Interestingly, l-arginine represses S. mutans genes associated with insoluble EPS (gtfB) and bacteriocin (SMU.150) production during early biofilm stages. In contrast, spxB expression by S. gordonii increases sharply at later phases of development, resulting in elevated levels of H₂O₂ in arginine-treated biofilms. Together, these effects caused major disruption in the EPS matrix assembly, 3D biofilm architecture, and in situ pH at the biofilm-apatite interface. Our data reveal intriguing and dynamic multifaceted roles of l-arginine as a potent disruptor of both EPS matrix and microbial interactions that are associated with cariogenic biofilm development.

MATERIALS AND METHODS

Bacterial strains and mixed-species ecological biofilm model.

S. mutans UA159 serotype c (ATCC 700610), A. naeslundii ATCC 12104, and S. gordonii DL1 were used in present study. These three strains were selected because S. mutans is a well-established virulent cariogenic bacterium (32). S. gordonii, a pioneer colonizer of dental biofilm, is arginolytic and ADS positive (33), and A. naeslundii is also detected during the early stages of dental biofilm formation (34). Furthermore, S. mutans UA159 and S. gordonii DL1 have been used in multiple laboratories in several in vitro biofilm models, and they are the strain of choice for in vivo (rodent) models of dental caries (35–39). Finally, these strains are well characterized (both genetically and phenotypically) and standardized, which are critical for the reproducibility of the study. All strains were stored at −80°C in tryptic soy broth containing 20% glycerol.

The biofilm method used in this study was designed to mimic the formation of cariogenic biofilms according to the “ecological plaque” concept, as detailed previously (6, 40, 41). Briefly, hydroxyapatite discs (1.25 cm in diameter, surface area of 2.7 ± 0.2 cm²; Clarkson Chromatography Products, Inc., South Williamsport, PA) were coated with filter-sterilized clarified human whole saliva (sHA) (6, 40). S. mutans UA159, A. naeslundii ATCC 12104, and S. gordonii DL1 were grown in ultrafiltered (10-kDa molecular mass cutoff membrane; Prep/Scale, Millipore, MA) buffered tryptone-yeast extract broth (UFTYE; 2.5% tryptone and 1.5% yeast extract [pH 7.0]) with 1% glucose to mid-exponential phase (37°C, 5% CO₂). Each of the bacterial suspensions was then mixed to provide an inoculum with a defined microbial population of S. mutans (10⁴ CFU/ml), S. gordonii (10⁴ CFU/ml), and A. naeslundii (10⁶ CFU/ml). To promote the ecological changes found under cariogenic conditions, we determined that the selected inoculum can promote the microbial shifts that favor S. mutans dominance following the addition of 1% sucrose in a reproducible manner, both in terms of timing of the changes and the number of CFU. The mixed population was inoculated in 2.8 ml of UFTYE containing 0.1% (wt/vol) sucrose and incubated for 19 h to form an initial biofilm community on the sHA surface. The biofilms were transferred to UFTYE containing 1% sucrose to induce environmental changes to simulate a cariogenic challenge from 29 h. The culture medium was changed twice daily (8 a.m. and 6 p.m.) until the end of the experimental period (115 h) (Fig. 1).

FIG 1.

Biofilm preparation and treatment regimen. The inoculum with a defined microbial population of S. mutans (10⁴ CFU/ml), S. gordonii (10⁴ CFU/ml), and A. naeslundii (10⁶ CFU/ml) was inoculated in 2.8 ml of UFTYE containing 0.1% (wt/vol) sucrose (0 h) and cultured without disturbance during the first 19 h to form an initial biofilm community on the sHA surface. The biofilms were transferred to UFTYE containing 1% sucrose to induce environmental changes simulating a cariogenic challenge from 29 h. The culture medium was changed twice daily (8 a.m. and 6 p.m.) until the end of the experimental period. Biofilm treatments started at 19 h and were carried out at 8 a.m., 1 p.m., and 6 p.m. each day.

Biofilm treatments.

Biofilm treatments started at 19 h and were carried out at 8 a.m., 1 p.m., and 6 p.m. each day, as detailed in Fig. 1. Free l-arginine (Ajinomoto AminoScience LLC, Raleigh, NC) was prepared at concentrations of 0.75%, 1.5%, and 3% (wt/vol) in filter-sterilized aqueous solution, and the pH was adjusted to 8 to 8.5 using phosphoric acid to be congruent with the pH values in arginine-containing formulations (29). Before treatment, biofilms were dip-washed in 2.8 ml of sterile saline solution (0.89% [wt/vol] NaCl) and then transferred into 2.8 ml of 0.75%, 1.5%, and 3% l-arginine or vehicle control (filter-sterilized water with pH adjusted to 8 to 8.5 using NaOH) for 10 min. The discs were shaken to remove excess and then transferred into wells containing culture medium without wash. All the discs were inoculated to assess the biological effects of l-arginine at the specific time points (Fig. 1).

Bacterial cell viability, biomass, and exopolysaccharide assays.

At selected time points (29, 53, 67, 91, and 115 h), biofilms were removed, homogenized via sonication, and subjected to biochemical and microbiological analyses, as detailed previously (6, 40, 42); our sonication procedure does not kill bacterial cells while providing optimum dispersal and maximum recoverable counts. Aliquots of biofilm suspension were serially diluted and plated on blood agar using an automated Eddy Jet Spiral Plater (IUL, SA, Barcelona, Spain). The three species were differentiated by observation of colony morphology combined with microscopic examination of cells from selected colonies, and the total number of viable cells in each of the biofilms was determined by counting CFU (6, 40). The remaining suspension was centrifuged at 5,500 × g for 10 min at 4°C, and the cell pellet was washed twice with water, dried in the dry oven at 105°C for 24 h, and weighed (43). The water soluble EPS (s-EPS), insoluble exopolysaccharides (i-EPS), and intracellular iodophilic polysaccharides (IPS) were extracted and quantified via colorimetric assays (42–44). Furthermore, the pH of the culture medium of treated and untreated biofilms was monitored every 2 h with an Orion pH electrode attached to an Orion 290 A+ pH meter (Thermo Fisher Scientific).

Three-dimensional confocal imaging of biofilms.

The influence of arginine exposure on the 3D biofilm architecture and the spatial distribution of EPS and bacterial biomass within intact biofilms were assessed using our established protocols optimized for confocal biofilm imaging and quantification (6, 40). Briefly, EPS was labeled using 2.5 μM Alexa Fluor 647-labeled dextran conjugate (10 kDa, 647/668 nm; Molecular Probes, Inc., Eugene, OR), while the microbial cells were stained with 2.5 μM SYTO 9 (485/498 nm; Molecular Probes, Inc.) (6). Laser scanning confocal fluorescence imaging of 67-h biofilms was obtained with an Olympus FV 1000 two-photon laser scanning microscope (Olympus, Tokyo, Japan) equipped with a 20× (0.45 numerical aperture) water immersion objective lens. The confocal image series were generated by optical sectioning at each selected position, and the step size of z-series scanning was 2 μm. The confocal images were analyzed via COMSTAT for the quantification of EPS and bacteria within intact biofilms, while the Amira 5.4.1 software (Visage Imaging, San Diego, CA) was used to create 3D renderings of the biofilm architecture (6, 40).

In situ pH measurement.

To measure the in situ biofilm pH, we used fluorescent pH indicator LysoSensor yellow/blue (Molecular Probes, Inc.) labeling method, as described previously (6). Briefly, the 67-h biofilms were incubated with LysoSensor yellow/blue-labeled dextran conjugate, and the pH values within intact biofilms were measured based on fluorescence intensity ratios of the dual-wavelength fluorophore. The fluorophore exhibits a dual-emission spectral peak (fluorescence emission maxima, 452 nm and 521 nm), and the ratio between the fluorescence intensity of these two spectral peak is pH dependent within biofilms (6). The fluorescence intensity of both emission wavelengths and the ratio of fluorescent intensity (I₄₅₀/I₅₂₀) within each biofilm image were measured using Image J 1.44 and its calculation tools (http://rsbweb.nih.gov/ij/download.html). The ratios of fluorescence intensity of selected areas within each biofilm image were converted to pH values using the titration curves of ratios versus pH (ranging from 3.5 to 7.0), as detailed previously (6).

RNA isolation and real-time analysis.

Quantitative reverse transcription-PCR (qRT-PCR) was performed to quantify the expression of specific target genes (gtfB and SMU.150 of S. mutans, and spxB and arcA of S. gordonii). Biofilms were harvested immediately after arginine exposure (at 43 h and 53 h) and 4 h after treatment (at 33 h and 47 h) from the same initial inoculum (Fig. 1; also see “Biofilm treatments,” above). RNA was extracted and purified from biofilms using standard protocols optimized for biofilms (42). The RNA integrity numbers (RIN) of purified samples used for qRT-PCR were determined by microcapillary electrophoresis on an Agilent 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA). The cDNA was synthesized from 1 μg of purified RNA (RIN, ≥9) with the Bio-Rad iScript cDNA synthesis kit (Bio-Rad Laboratories, Inc., Hercules, CA), and quantitative amplification conditions using Bio-Rad iTaq Universal SYBR green Supermix and Bio-Rad CFX96 system (Bio-Rad Laboratories, Inc.). The primers used in present study were listed in Table 1. Standard curves for each primer were used to determine the relative number of cDNA molecules, and relative expression was calculated by normalizing to the 16S rRNA gene transcripts (6). The Minimum Information for Publication of Quantitative Real-Time PCR Experiments (MIQE) guidelines were used for quality control of the data generated and for data analysis.

TABLE 1.

Primers used for qRT-PCR in the present study

Primer target	Directiona	Primer sequence
S. gordonii 16S rRNA	F	5′-GCTTGCTACACCATAGACTG-3′
	R	5′-AGCCGTTACCTCACCTAC-3′
S. mutans 16S rRNA	F	5′-ACCAGAAAGGGACGGCTAAC-3′
	R	5′-TAGCCTTTTACTCCAGACTTTCCTG-3′
gtfB	F	5′-AGCAATGCAGCCAATCTACAAAT-3′
	R	5′-ACGAACTTTGCCGTTATTGTCA-3′
SMU.150	F	5′-GAAGGTATCGGGTGGAGAAG-3′
	R	5′-CCCAAGTGCCTACACAATATG-3′
spxB	F	5′-GCGTACATCTCCACTCTTTG-3′
	R	5′-CACCCATGATGTTCCATACTT-3′
arcA	F	5′-GTCTTTGACCTCACCAGAAA-3′
	R	5′-ACTCACGAATAGCCACTTTAG-3′

^aF, forward; R, reverse.

H₂O₂ concentration measurement.

The concentration of H₂O₂ was measured using a horseradish peroxidase (HRP) enzymatic assay (45). Briefly, biofilm was gently washed once, released into glass tubes containing 0.5 ml of reaction mixture (1 mg/ml 3,3′,5,5′-tetramethylbenzidine [TMB] and 1:2,000 dilution of HRP [Invitrogen] in 0.1 M sodium acetate buffer [pH 4.5]), and incubated at room temperature for 15 min. The same volume of H₂SO₄ (2 N, 0.5 ml) was added into tubes for reaction termination. The supernatant was transferred into Eppendorf tubes, centrifuged at 4°C for 1 min, and the absorbance at 450 nm was determined using a plate reader. To determine the concentration of H₂O₂, a standard curve was prepared using a serially diluted H₂O₂ solution, and the data were normalized with dry weight.

Statistical analysis.

The data are presented as the mean ± one standard deviation (SD). Each of the experiments was repeated at least 3 times. Pairwise comparisons were made between the test and control using Student's t test. The chosen level of significance for all statistical tests in present study was a P value of <0.05.

RESULTS

l-Arginine topical exposure causes microbial shifts preventing S. mutans outgrowth and favoring S. gordonii survival.

We first assessed the influence of l-arginine on the dynamics of microbial population changes within biofilms. The treatment regimen is shown in Fig. 1, and three concentrations (0.75%, 1.5%, and 3%) were tested. As shown in Fig. 2, the introduction of l-arginine caused dramatic changes in the number and proportion (right panels) of viable microbial populations (versus vehicle control). In vehicle-treated biofilms, S. mutans was the dominant species at 29 h and maintained its dominance to the endpoint (115 h), consistent with a recent in vivo study showing that S. mutans outcompetes S. gordonii in a rodent caries model (39). S. mutans was considered the dominant species based on its higher proportion of viable cells relative to others (Fig. 2A, right). In parallel, the viable population of S. gordonii declined, while A. naeslundii was undetectable after 67 h of biofilm development.

FIG 2.

Dynamic changes of microbial composition of biofilms with and without topical l-arginine (l-arg) exposure. Total viable cells for each experimental group are shown as bar graphs, while the proportion of different bacterial species in the treated biofilms is presented in the right graphs (n = 12). In vehicle-treated biofilms, S. mutans (S. m) was the dominant species from 29 h to the endpoint, and the viable population of S. gordonii (S. g) declined while A. naeslundii (A. n) was undetectable after 67 h. In contrast, 1.5% and 3% l-arginine-treated biofilms were characterized by S. gordonii dominance from 53 h and detectable levels of A. naeslundii throughout the experimental period, while the S. mutans viable population sharply declined. l-Arginine at 0.75% showed modest effects on the microbial composition compared to vehicle control or the higher concentrations of l-arginine.

In contrast, 1.5% or 3% l-arginine exposure shifted the microbial populations from S. mutans dominance (at 29 h) to S. gordonii-dominated biofilm between 53 h and 67 h, which were also characterized by detectable levels of A. naeslundii throughout the experimental period. Although the total CFU counts in 1.5% l-arginine-treated biofilms appear similar at 53 h (as the values are in log scale), the proportions of viable S. mutans and S. gordonii cells start to shift at this time point, which becomes more evident at 67 h. This observation indicates that l-arginine requires some time to exert its effects (at least in part via metabolism by arginolytic bacteria, e.g., S. gordonii) and thereby impact the microbial composition. Furthermore, the proportion of S. mutans viable cells population declined sharply in l-arginine-treated biofilms, although this bacterium was able to survive over time; 0.75% l-arginine showed limited effects on the microbial composition compared to vehicle control or higher concentrations of l-arginine. Interestingly, we observed an overall drop in the total number of cells between 53 h and 67 h, particularly in the vehicle control group. The exact reasons are unclear, but it is possible that some of the biofilm detached due to increased biomass accumulation, followed by further regrowth after this time point.

Clearly, l-arginine at 1.5% or 3% (despite brief topical exposures) is capable of inducing major changes in the viable microbial populations and proportions despite the presence of sucrose (a cariogenic stimuli), promoting the transition from a virulent (high levels of S. mutans and small numbers of S. gordonii) to a potentially less-cariogenic (S. gordonii dominated) biofilm. The total viable bacterial cell population in the biofilm was unaffected by l-arginine treatments.

l-Arginine treatments impact the biomass accumulation, polysaccharide content, and 3D architecture of biofilms.

Concomitantly, we also observed that the biomass (dry weight) of the 1.5% and 3% l-arginine-treated biofilms was greatly decreased compared to that of vehicle-treated biofilms at each of the time points of biofilm development (Fig. 3A). It is possible that the reduced biomass is associated with changes in EPS content, as fewer S. mutans cells are present in the l-arginine-treated biofilms. S. mutans produces large amounts of insoluble glucans in the presence of sucrose, forming a matrix that provides bulk and structural integrity to the biofilm while facilitating the creation of localized acidic microenvironments (10). Indeed, the total amount of insoluble EPS in 1.5% and 3% arginine-treated biofilms (115 h) was reduced by ∼3-fold compared to those treated with vehicle (Fig. 3B). Consistent with the microbiological data, 1.5% or 3% l-arginine treatments were equally effective in reducing both the biofilm biomass (dry weight) and the amount of insoluble EPS (versus vehicle control), while the biological effects of 0.75% l-arginine were modest. Based on these findings, we selected 1.5% l-arginine, which has also been shown to be a clinically effective concentration (29–31), for further analyses. Interestingly, the content of soluble EPS and intracellular polysaccharides (IPS) was unaffected (see Fig. S1 in the supplemental material) (despite the reduction of S. mutans), likely due to an increase in the number of S. gordonii cells, which is capable of synthesizing soluble glucans from sucrose, and to produce and store IPS.

FIG 3.

Biomass and insoluble exopolysaccharide composition of biofilms with and without l-arginine exposure. Shown are dry weight (A) and total amounts (B) of insoluble exopolysaccharides (n = 9). The biomass (dry weight) of the 1.5% and 3% l-arginine-treated biofilms was greatly reduced compared to vehicle-treated biofilms at each of the time points. Water-insoluble EPS in arginine-treated biofilms (115 h) was reduced by ∼3-fold compared to those treated with vehicle control; 0.75% l-arginine showed limited biological effects. Values are significantly different from each other at P < 0.05 (*) and P < 0.01 (**).

Confocal images showed clear alteration in the biofilm 3D architecture by 1.5% l-arginine treatments, showing a defective EPS matrix and areas devoid of bacterial clusters or microcolonies on the sHA surface (Fig. 4). In contrast, vehicle-treated biofilm contains a well-structured EPS matrix, with bacterial clusters covering the entire apatitic surface. Furthermore, the spatial distribution of EPS and bacteria across the biofilm thickness (see selected area in the merged images, Fig. 4) was also compromised in arginine-treated biofilms (versus vehicle control). Although the confocal images show overall less bacterial biomass in arginine-treated biofilms, it is noteworthy that several densely packed microcolonies are still present. Collectively, the data show that topical applications of exogenous l-arginine at a concentration of 1.5% cause major biochemical and structural changes, as S. mutans outgrowth was suppressed while S. gordonii dominance was promoted within treated biofilms.

FIG 4.

Three-dimensional architecture of biofilms with and without 1.5% l-arginine exposure. (A) Representative 3D rendering of 67-h biofilms treated with vehicle control or 1.5% l-arginine. (B) The EPS/cell distribution across the biofilm thickness at the selected area in the merged image in panel A was determined via COMSTAT. A defective EPS matrix and areas devoid of bacterial clusters or microcolonies on the sHA surface were observed in the arginine-treated biofilms. In contrast, vehicle-treated biofilms show a well-structured EPS matrix with bacterial clusters covering the entire apatitic surface.

S. mutans and S. gordonii gene expression profile following l-arginine treatment.

To further understand the interplay between S. mutans and S. gordonii during biofilm development and l-arginine exposure, we examined the expression pattern of specific genes associated with insoluble EPS matrix assembly (gtfB of S. mutans), interspecies interactions (SMU.150 of S. mutans and spxB of S. gordonii), and arginine metabolism (arcA of S. gordonii). The dynamics of gene expression (normalized by 16S rRNA) were analyzed between 29 h and 53 h, because the microbial composition shift occurred during this period; 29-h biofilms were excluded since we could not obtain sufficient biomass for RNA extraction.

The transcription of gtfB by S. mutans was significantly repressed by 1.5% l-arginine exposure at early time points (33 h and 43 h) (Fig. 5), which agrees well with the biochemical and structural data. GtfB is essential for both S. mutans accumulation and EPS-rich matrix development (10). Furthermore, we observed interesting changes in the expression pattern of genes involved in the competitive interactions between S. mutans and S. gordonii. SMU.150 gene expression by S. mutans was severely repressed in arginine-treated biofilm at an early time point (33 h) and to a lesser extent at 43 h, while it was moderately increased at 53 h (Fig. 5). Intriguingly, S. gordonii spxB transcripts sharply increased (>3.5-fold versus vehicle control) at a later time point (53 h) (Fig. 6). SMU.150 is involved in the synthesis of bacteriocin that is capable of inhibiting the growth of non-mutans streptococci (46–48). Conversely, pyruvate oxidase encoded by spxB is largely responsible for H₂O₂ production (47), which can be used by S. gordonii to outcompete S. mutans within mixed-species biofilms (47). To further explore if the levels of H₂O₂ were increased in biofilms treated with l-arginine, we measured the amounts of hydrogen peroxide in the whole biofilms via a colorimetric assay (49). Consistent with the spxB expression profile, a higher H₂O₂ level was found in 53-h biofilms treated with arginine versus vehicle control (Fig. 6).

FIG 5.

l-Arginine (1.5%) exposure disrupts the dynamics of gene expression of S. mutans associated with EPS and bacteriocin (n = 6). SMU.150 and gtfB gene expression by S. mutans decreased in arginine-treated biofilm at early time points (33 h and 43 h), but SMU.150 expression increased at 53 h. Values are significantly different from each other at P < 0.05 (*).

FIG 6.

l-Arginine (1.5%) exposure promotes H₂O₂ production by S. gordonii. Expression of S. gordonii spxB (n = 6) (A) and H₂O₂ amounts (n = 12) (B) in the treated biofilms were determined. S. gordonii spxB transcripts sharply increased (>3.5-fold versus vehicle control) at a later time point (53 h), and a higher H₂O₂ level was found in 53-h biofilms treated with arginine (versus vehicle control). Values are significantly different from each other at P < 0.05 (*). conc., concentration.

In the oral cavity, arginine is primarily metabolized through the ADS system (18). The genes encoding the ADS are commonly arranged in an operon, and the arcA gene encodes arginine deiminase, a key component of ADS system, which hydrolyzes arginine to generate citrulline and ammonia (19). The expression of arcA was slightly increased, albeit the differences were not statistically significant (see Fig. S2 in the supplemental material); it is possible that strongly arginolytic organisms, such as S. gordonii, do not respond to elevated concentrations of arginine to induce arcA expression. Altogether, topical exposure to 1.5% l-arginine is capable of modulating streptococcal competition by downregulating gtfB and SMU.150 and enhancing spxB expression, which might explain at least in part the biochemical and microbiological changes within treated biofilms.

In situ pH at the biofilm-sHA interface increases following l-arginine treatment.

The deleterious effects of arginine exposure on S. mutans accumulation, insoluble EPS content, and matrix assembly combined with enhancement of S. gordonii growth might affect the pH at the surface of biofilm attachment, which is the hallmark for caries initiation (50, 51). Previous studies have shown the pH-raising capacity of arginine by measuring the pH of the surrounding culture medium (52) or via electrodes placed in the biofilm (28). However, in situ pH measurements at the sHA-biofilm interface without disrupting the integrity of the biofilm 3D architecture have not been reported yet.

Recently, we devised a noninvasive pH mapping approach using a fluorescent pH indicator to determine pH values throughout intact biofilm structure (6). Here, we focused on measuring the pH values in close proximity of the sHA surface (up to 30 μm from the surface), because the maintenance of acidic pH at the biofilm-sHA interface promotes demineralization of the adjacent apatite (50, 51); the critical pH for demineralization of enamel appears to be as low as 5.1 in the plaque fluid, as recently reviewed (51). We found that the average pH values of 1.5% l-arginine-treated biofilms were significantly higher than those from vehicle-treated biofilms (Fig. 7). Strikingly, the differences in the in situ pH values were as high as one full unit (versus vehicle-treated biofilms), particularly between 20 and 30 μm from the sHA surface (Fig. 7A), with pH above 5.7, which might impact the initiation and progression of carious lesions.

FIG 7.

In situ pH measurement at the biofilm-apatite interface. In situ pH values in 67-h biofilm were determined every 2 μm from the sHA surface (A) and averaged based on pH measurements across the biofilm interface (n = 50) (B). The differences in the in situ pH values were as high as one full unit (versus vehicle-treated biofilms), particularly between 20 to 30 μm from the sHA surface. Values are significantly different from each other at P < 0.05 (*).

DISCUSSION

The favorable modification of the microbial composition by l-arginine mediated via ADS activity has been recognized as the primary anticaries mechanism (18, 29), since ADS could generate alkali from arginine to neutralize excessive acid within plaque. Here, we established a mixed-species biofilm model to explore additional mechanisms by which l-arginine impacts the cariogenic properties of biofilms. Although our model does not simulate the episodic dietary intake in humans, it does mimic sucrose-rich exposure conditions found in high caries-active individuals, as well as the biochemical and microbiological changes associated with cariogenic biofilm formation (4, 5). Our data reveal that topical applications of 1.5% l-arginine similar to multiple daily oral exposures disrupt insoluble EPS production, which impairs S. mutans accumulation and further biofilm build-up. At the same time, it modulates interspecies competition that favors S. gordonii growth and dominance. These modifications in turn result in defective EPS matrix assembly, altered biofilm 3D architecture, and significantly higher pH at the biofilm-apatite interface that could attenuate biofilm virulence (cariogenicity) without bacterial killing (53).

Considering the essential roles of EPS in the development and cariogenicity of oral biofilms (1–3, 10), we propose that the action of l-arginine as a disruptor of the process of biofilm matrix assembly is an important additional mechanism for its caries-protective effect. Our data suggest that the repression of gtfB expression at early stages of biofilm formation combined with a rapid decline in the proportion of S. mutans (and its lower abundance during the later stages of biofilm development) might explain the impairment of the EPS matrix assembly over time. Insoluble EPS (glucans) produced by GtfB promote selective binding and accumulation of S. mutans onto apatitic surfaces (10–12). Furthermore, it also enhances cell clustering and microcolony development that facilitate S. mutans persistence within biofilms (2, 6, 13, 16). Thus, the disruption of GtfB-derived EPS would markedly impact the ability of S. mutans to colonize and accumulate within biofilms. As the main producer of insoluble EPS within the oral microbiome (2), such effects could sharply reduce the assembly of the biofilm matrix as well as the biofilm bulk and biomass, as observed in the arginine-treated biofilms.

The highly insoluble nature of GtfB-derived glucans is also essential for the scaffolding and structural integrity of the biofilm 3D architecture (1, 2, 6, 16), which directly modulates the surface attachment strength of the biofilm (54). Interestingly, l-arginine appears to affect the biomechanical properties of S. mutans-derived EPS, which attenuated the biofilm adhesion strength (25). In addition to providing bulk and mechanical stability, the EPS matrix also forms a diffusion-limiting milieu that embeds cariogenic bacteria, forming protective niches against antimicrobials (1, 6, 55). Recently, it was shown that the antimicrobial efficacy of cetylpyridinium chloride (CPC) against oral biofilms was enhanced when used in combination with arginine in vitro (27). The authors suggested potential alterations in the EPS matrix, which could facilitate CPC penetration (27); our data certainly support this clinically relevant observation. It is readily apparent that l-arginine effects on EPS might suppress biofilm accumulation while making them more susceptible to treatment or removal.

The results from this study also show that topical exposure of l-arginine affects key modulators involved in competitive interspecies interactions between S. mutans and S. gordonii. The expression of S. mutans SMU.150, which encodes mutacin, was repressed during the early biofilm development stage. Mutacin production by S. mutans is known to inhibit the growth of commensal bacteria, such as S. gordonii (46). Conversely, l-arginine exposure induced S. gordonii H₂O₂ production, which is an effective “chemical weapon” against S. mutans growth in vitro (47). Interestingly, the expression of the mutacin-encoding gene by S. mutans and spxB by S. gordonii are both higher in the arginine-treated biofilms at the later time point. It is possible that S. mutans is upregulating mutacin production in response to H₂O₂ stress caused by the growing numbers of S. gordonii actively expressing spxB in the arginine-treated biofilms, which in turn would help S. mutans persist in this unfavorable milieu. This observation might explain, at least in part, why S. mutans cells are not completely eliminated and survive in the biofilm.

A. naeslundii also benefits from l-arginine supplementation, as this bacterium is detected throughout the experiment period in arginine-treated biofilms while absent after 67 h in control biofilms. Since A. naeslundii has been reported to interact with S. gordonii and promote its growth (56), the enhanced A. naeslundii survival in arginine-treated biofilms may indirectly help S. gordonii compete against S. mutans. Thus, arginine is capable not only of hindering S. mutans colonization and initial accumulation via disruption of EPS-mediated mechanisms but also the competitiveness of this pathogen within sucrose-grown biofilms (although S. mutans appears to survive and possibly fight back over time). We propose that arginine might play an active role in mediating the regulatory pathways associated with the transcription of gtfB, SMU.150, and spxB. Future studies shall elucidate the underlying molecular mechanisms regulating the expression of these genes and their dynamic interactions. It should be noted that the regulation of these genes is complex and linked with multiple pathways, which may reveal additional therapeutic targets.

Ammonia generation via ADS activity can neutralize glycolytic acids and modulate pH homeostasis within biofilms (18). Our data reveal that defective assembly of EPS matrix and microcolony formation as a result of l-arginine exposure may also contribute to elevated in situ pH at the biofilm-sHA interface. Previous studies have shown that EPS-enmeshed microcolonies could trap acid at the sHA surface and/or restrict the access of neutralizing buffer to generate acidic microenvironments locally due to metabolic activity of the densely packed bacteria enmeshed within a diffusion-limiting EPS matrix, particularly at the deeper layers of the biofilm (6, 57, 58). Altogether, it appears that the combination of ADS activity and favorable ecological changes with disruption of EPS/microcolony development may offer a more comprehensive explanation for the biofilm pH-related effects associated with l-arginine exposure. However, additional studies are needed to further understand the role of protonated/deprotonated forms of l-arginine (which varies with pH) in biofilm formation and virulence and validate the biological effects observed here using in vivo and longitudinal clinical studies.

In conclusion, our data reveal intriguing new biological insights into how l-arginine impacts the process of biofilm matrix assembly and the dynamic interspecies competition between a cariogenic oral pathogen and alkali-producing commensal organism, demonstrating the multiaction potency of this readily available biofilm disruptor.

Funding Statement

The funders had no role in the study design, data collection and interpretation, or the decision to submit the work for publication.