Novel dental composite with capability to suppress cariogenic species and promote non-cariogenic species in oral biofilms

Abstract

Recurrent caries often occurs and is a primary reason for the failure of dental composite restorations. The objectives of this study were to: (1) develop a bioactive composite containing dimethylaminohexadecyl methacrylate (DMAHDM), (2) investigate its antibacterial effects and suppression on biofilm growth, and (3) investigate its ability to modulate biofilm species composition for the first time. DMAHDM was incorporated into a composite at mass% of 0%, 0.75%, 1.5%, 2.25% and 3%. A commercial composite Heliomolar served as a comparative control. A biofilm model consisting of Streptococcus mutans (S. mutans), Streptococcus sanguinis (S. sanguinis) and Streptococcus gordonii (S. gordonii) was tested by growing biofilms for 48 h and 72 h on composites. Colony-forming units (CFUs), metabolic activity and live/dead staining were evaluated. Lactic acid and polysaccharide productions were measured to assess biofilm cariogenicity. TaqMan real-time polymerase chain reaction was used to determine the proportion of each species in the biofilm. DMAHDM-containing composite had a strong anti-biofilm function, reducing biofilm CFU by 2–3 orders of magnitude, compared to control composite. Biofilm metabolic activity, lactic acid and polysaccharides were decreased substantially, compared to control (p < 0.05). At 72 h, the cariogenic S. mutans proportion in the biofilm on the composite with 3% DMAHDM was 19.9%. In contrast, an overwhelming S. mutans proportion of 92.2% and 91.2% existed in biofilms on commercial control and 0% DMAHDM, respectively. In conclusion, incorporating DMAHDM into dental composite: (1) yielded potent anti-biofilm properties; (2) modulated the biofilm species composition toward a non-cariogenic tendency. The new DMAHDM composite is promising for applications in a wide range of tooth cavity restorations to modulate oral biofilm species and combat caries.

Introduction

Resin composites are widely used in dentistry due to their excellent esthetics and direct-filling capabilities [1]. However, composites tend to accumulate more biofilms (plaque) and can release bacteria-simulating compounds such as triethylene glycol dimethacrylate (TEGDMA) over time [2–4]. Biofilms can produce acids to cause demineralization in tooth structures, and secondary caries incidence at the tooth-composite margins was found to be 3.4 times higher than that in amalgam restorations [5, 6]. Indeed, secondary caries (recurrent caries) is a frequent occurrence and a main reason for the failure of resin composite restorations [7–10].

Efforts have been made to modify dental resins with antibacterial agents to suppress biofilm acids and secondary caries [11, 12]. With the development of nanotechnology, the application of nanoparticles is evolving quickly to improve dental materials [13–15]. Studies have reported several types of promising antimicrobial nanoparticles including silver [16], ZnO [17], TiO₂ [18], and bioactive glass [19]. In addition, quaternary ammonium methacrylates (QAMs) are a class of cationic compounds with a broad spectrum of antimicrobial effects [11, 12]. The antimicrobial mechanism of QAMs is considered to be “contact-killing” by disrupting bacterial membranes [11]. Compared to release-based biomaterials, QAMs can be copolymerized with the resin matrix to anchor itself into the polymer network with prolonged antimicrobial ability [11, 12]. The first QAM in dental resin was 12-methacryloyloxy dodecyl pyridinium bromide (MDPB), which showed potent antibacterial effects against biofilm growth on resins [20]. Since then, several other QAMs were developed, including quaternary ammonium dimethacrylate (QADM) [21], quaternary ammonium polyethylenimine (QPEI) [22], methacryloxylethyl cetyl dimethyl ammonium chloride (DMAE-CB) [23], and dimethylaminododecyl methacrylate (DMADDM) [24]. Recently, a new antibacterial monomer dimethylaminohexadecyl methacrylate (DMAHDM) was developed and incorporated into dental resins which displayed potent anti-biofilm effects on Streptococcus mutans (S. mutans) biofilms and saliva-based microcosm biofilms [25, 26].

The dental plaque contains multispecies microbial communities. Caries is caused by not just a single pathogenic species, but by a change in the biofilm from a non-cariogenic composition toward a cariogenic composition [27–29]. The changes in oral environment can trigger a shift in the microflora; for example, frequent sugar intake can enrich acidogenic and aciduric species at the expense of the healthy and less aciduric residents, leading the dental plaque to a more cariogenic composition [27]. Therefore, it would be highly desirable to develop a bioactive dental composite to be able to modulate the biofilms from a cariogenic state toward a non-cariogenic microbial community, thereby to reduce biofilm acids and dental caries. This important direction of research is only getting started; indeed, a literature search revealed only one report on the use of QAM to modulate oral biofilms [30]. That study demonstrated that a resin containing DMADDM was able to suppress the cariogenic S. mutans and promote the growth of non-cariogenic species [30]. To date, besides a single study on biofilm modulation via DMADDM [30], there has been no other report on biofilm modulation via QAM resins. Whether the new monomer DMAHDM can suppress cariogenic species and promote non-cariogenic species in oral biofilms remains to be investigated.

The objectives of this study were to incorporate DMAHDM into a dental composite and to investigate its ability to suppress biofilm growth and modulate biofilm species compositions for the first time. It was hypothesized that: (1) The bioactive resin composite containing DMAHDM would reduce the biofilm growth and decrease the cariogenicity of the biofilm, resulting in less acid production and less biofilm polysaccharides; (2) The composite containing DMAHDM could change the species composition in the biofilm, reducing the cariogenic species percentage and increasing the non-cariogenic species percentage in the biofilm; (3) The cariogenicity and the cariogenic species percentage in the biofilm would be inversely proportional to the DMAHDM concentration in the composite.

2. Material and methods

2.1. Fabrication of composite containing DMAHDM

DMAHDM was synthesized using a modified Menschutkin reaction method, as previously described [31]. In brief, 10 mmol of 2-(dimethylamino) ethyl methacrylate (DMAEMA, Sigma-Aldrich, St. Louis, MO, USA), 10 mmol of 1-bromododecane (BDD, TCI America, Portland, OR, USA), and 3 g of ethanol were added into a vial, then capped and stirred at 70 °C to react for 24 h. After the reaction was complete, the ethanol solvent was removed by evaporation, yielding DMAHDM as a colorless viscous liquid.

Bisphenol glycidyl dimethacrylate (BisGMA, Esstech, Essington, PA, USA) and TEGDMA (Esstech, Essington, PA, USA) were mixed at a mass ratio of 1:1, after which 0.2% camphorquinone and 0.8% ethyl 4-N, N-dimethylaminobenzoate were added to render it light-curable (referred to as BT resin) [32]. DMAHDM was incorporated into BT at a DMAHDM/(BT+DMAHDM) mass fractions of 0%, 2.5%, 5%, 7.5% and 10%. The highest concentration of 10% was chosen without compromising the mechanical property of the resin as previously described [33]. Barium boroaluminosilicate glass particles with a median size of 1.4 μm (Caulk/Dentsply, Milford, DE, USA) were silanized with 4% 3-methacryloxypropyltrimethoxysilane [32, 33]. A mass fraction of 70% glass particles was mixed with 30% BT resin, yielding a cohesive paste [32, 33]. Since the resin mass fraction was 30%, the DMAHDM mass fractions in the composite were 0%, 0.75%, 1.5%, 2.25% and 3% respectively. A commercial composite Heliomolar (Ivoclar, Amherst, NY, USA) was used as a comparative control. It contained 40–200 nm nano-silica and ytterbium-trifluoride at a filler mass fraction of 66.7%. Thus, six groups of composites were tested:

• (1)Commercial composite control: Heliomolar;
• (2)Experimental composite control: 30% BT + 70% glass particles (referred to as “0% DMAHDM”);
• (3)29.25% BT + 0.75% DMAHDM + 70% glass particles (referred to as “0.75% DMAHDM”);
• (4)28.5% BT + 1.5% DMAHDM + 70% glass particles (referred to as “1.5% DMAHDM”);
• (5)27.75% BT + 2.25% DMAHDM + 70% glass particles (referred to as “2.25% DMAHDM”);
• (6)27% BT + 3% DMAHDM + 70% glass particles (referred to as “3% DMAHDM”).

For specimen preparation, composite disks were made using molds with a diameter of 10 mm and a thickness of 1 mm, which were light-cured (Triad 2000; Dentsply, York, PA, USA) for 1 min on each side [32]. The cured disks were immersed in distilled water at 37 °C and agitated for 24 h to remove any uncured monomers. The disks were sterilized with ethylene oxide (AnproleneAN 74i, Andersen, Haw River, NC, USA) and degassed for 3 days following the manufacturer’s instructions [33].

2.2. Bacteria inoculation and biofilm formation

S. mutans ATCC700610, Streptococcus sanguinis ATCC10556 (S. sanguinis) and Streptococcus gordonii ATCC10558 (S. gordonii) were obtained from American Type Culture Collection (ATCC, Manassas, VA, USA). The experiments were approved by the University of Maryland Baltimore Institutional Review Board. All strains were cultured at 37 °C under aerobic condition (5% CO₂) in brain heart infusion broth (BHI, Difco, Sparks, MD, USA).

For multispecies biofilm formation, composite disks were placed into 24-well plates and the biofilm inoculum was adjusted to 10⁷ colony-forming units (CFU)/mL of S. mutans, 10⁷ CFU/mL of S. sanguinis, and 10⁷ CFU/mL of S. gordonii in 1.5 mL BHI supplied with 1% sucrose for each well [30]. The bacteria concentrations of the inoculum were determined by a spectrophotometer (Genesys 10S, Thermo Scientific, Waltham, MA, USA) based on the OD_600nm versus CFU/mL graph of each strain. The ratio of S. mutans/S. sanguinis/S. gordonii =1:1:1 in the inoculum was further confirmed by quantitative real-time polymerase chain reaction in preliminary experiments (qPCR; data not shown). Biofilms on disks were cultured for 48 h to form relatively mature biofilms, following previous studies [25, 26]. In addition, considering that oral bacteria in microgaps at the tooth-restoration margins can stay for long periods of time and may not be easily removed during tooth-brushing, a longer culture time of 72 h was also tested. During the culture, the composite disks with biofilms were transferred to a new 24-well plate with fresh medium every 24 h.

2.3. Live/dead staining assay

Disks with 48 h or 72 h biofilms were washed twice with phosphate buffered saline (PBS), then stained with Live/Dead Baclight bacterial viability kits as previously described (Molecular Probes, Eugene, OR, USA) [32, 34]. Briefly, 2.5 μM SYTO 9 and 2.5 μM propidium iodide were set on each sample for 15 min. Live bacteria were stained with SYTO 9 to emit a green fluorescence. Dead bacteria with compromised membrane were stained with propidium iodide to emit a red fluorescence. Images were captured with an inverted epifluorescence microscope (TE2000-S, Nikon, Melville, NY, USA). Three disks were used for each group and each disk was tested in five randomly-selected positions, yielding 15 images per group.

2.4. Biofilm CFU counts

Biofilms were cultured for 48 h or 72 h as described above. Six disks of each group at each time point were used for CFU counts. Biofilms were harvested in PBS by scraping and sonication/vortexing (Fisher, Pittsburg, PA, USA) as described in previous studies [33, 35]. The suspensions were serially diluted and spread on BHI plates. After 48 h incubation at 37 °C in 5% CO₂, the colony number was counted and CFU counts were determined.

2.5. MTT assay

3-(4,5-Dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) can be reduced by enzymes in viable cells to form purple color formazan, thus reflecting the metabolic activity of the biofilms [24, 33]. Disks with 48 h and 72 h biofilms were washed twice with PBS and transferred into new 24-well plate. One mL MTT dye (0.5mg/mL MTT in PBS) was added into each well and incubated for 1 h at 37 °C in 5% CO₂ [24, 33]. After that, the disks were transferred into new 24-well plate with 1 mL dimethyl sulfoxide (DMSO) in each well and incubated at room temperature for 20 min to dissolve the formazan crystals. DMSO solution was then transferred into 96-well plate and OD_540nm was determined using a microplate reader (SpectraMax M5, Molecular Devices, Sunnyvale, CA, USA) [24, 33]. Six disks were tested for each group at either 48 h or 72 h, respectively.

2.6. Lactic acid production by biofilms

Disks with 48 h or 72 h biofilms were washed twice with PBS, then immersed in 1.5 mL buffered peptone water (BPW; Sigma-Aldrich) supplemented with 0.2% sucrose and incubated at 37 °C in 5% CO₂ for 3 h [24, 32, 33]. The lactate concentrations in BPW were determined using a lactate dehydrogenase enzymatic method by measuring OD_340nm and compared to that of the lactic acid standard curves, as previously described [24, 32, 33]. Standard curves were prepared with a standard lactic acid (Supelco Analytical, Bellefonte, PA, USA). Six disks were tested for each group at each time period.

2.7. Polysaccharide production of biofilms

Water-insoluble polysaccharides of biofilms were measured using a phenol-sulfuric acid method as previously described [36, 37]. Six disks were tested for each group at 48 h or 72 h. Biofilms on disks were collected by scraping and sonication/vortexing in 2 mL PBS, followed by centrifugation. The precipitate was then washed twice with PBS and resuspended in 200 μL of distilled water. 200 μL of 5% phenol solution and 1 mL of 95–97% sulfuric acid were added followed by incubation at room temperature for 30 min [36, 37]. After mixing by pipetting, 200 μL of the solution was transferred into a 96-well plate and OD_490nm was determined with the microplate reader (SpectraMax M5) [36, 37]. Seven standard glucose concentrations of 0, 50, 100, 150, 200, 250 and 300 mg/mL were used to plot the standard curve of OD_490nm versus polysaccharide concentrations.

2.8. DNA isolation and TaqMan qPCR assay

For DNA isolation, biofilms were collected by scraping and sonication/vortexing into 2 mL PBS, followed by centrifuging and resuspending in 200 μL of lysis solution (Sigma-Aldrich) supplemented with 45 mg/mL lysozyme (Sigma-Aldrich) [38]. The solution was incubated at 37 °C for 1 h to lyse bacteria [38]. After that, DNA was isolated and purified with GenElute Bacterial Genomic DNA Kits (Sigma-Aldrich) following the manufacturer’s protocol. The purity and concentration of DNA was confirmed via NanoDrop 2000 spectrophotometer (Thermo Scientific, Waltham, MA, USA). The DNA was then stored at −20 °C until use.

TaqMan qPCR was used to monitor bacteria composition shifts in the biofilms at 48 h and 72 h. The species-specific primers and probes used for S. mutans, S. sanguinis and S. gordonii are listed in Table 1. Conventional PCR was used to confirm the specificity of the primers. For each qPCR, 20 μL of the reaction mixture contained 10 μL of Premix Ex Taq for Probe qPCR (Takara Bio, Shiga, Japan), 2 μL of DNA template, 0.4 μL of each forward and reverse primer at concentration of 10 μM (final concentration 0.2 μM), 0.8 μL of TaqMan probe at concentration of 10 μM (final concentration 0.4 μM), 0.4 μL of ROX passive reference dye (Takara Bio) and 6 μL of ultra-pure water (Quality Biological, Gaithersburg, MD, USA). qPCR amplifications were performed on a 7900HT Fast Real-Time PCR system (Applied Biosystems) programmed for 3 min at 95 °C, followed by 40 cycles of 10 s at 95 °C, 30 s at 56 °C and 30 s at 72 °C. Fluorescence data were collected at the 72 °C elongation step throughout amplification cycles and the critical threshold cycles (Ct) were analyzed using SDS software (version 2.3, Applied Biosystems).

Table 1.

Primers and probes used in TaqMan qPCR

Primers/Probes	Nucelotide Sequence (5’-3’)	Reference
Primers
S. mutans-f	5’-GCCTACAGCTCAGAGATGCTATTCT-3’	[39]
S. mutans-r	5’-GCCATACACCACTCATGAATTGA-3’	[39]
S. sanguinis-f	5’-GAGCGGATGGCCAATTATATCT-3	[30]
S. sanguinis-r	5’-CCGGATGATGTCGGCAATA-3’	[30]
S. gordonii-f	5’-GGTGTTGTTTGACCCGTTCAG-3’	[40]
S. gordonii-r	5’-AGTCCATCCCACGAGCACAG-3’	[40]
Probes
S. mutans	5’-FAM-TGGAAATGACGGTCGCCGTTATGAA-TAMRA-3’	[39]
S. sanguinis	5’-FAM-TGTTCGGGCTCATGATA-Eclipse-3’	[30]
S. gordonii	5’-FAM-AACCTTGACCCGCTCATTACCAGCTAGTATG-TAMRA-3’	[40]

To establish the correlations between Ct and the bacteria number, standard curves of each strain was plotted. Genomic DNA of known numbers of each strain was extracted and successively 10-fold diluted standing for the corresponding concentration of bacteria from 10⁸ CFU to 10³ CFU, as described in previous studies [30, 39, 40]. The Ct values of the known DNA were determined by qPCR, yielding three standard curves of Ct versus bacteria number for S. mutans, S. sanguinis and S. gordonii, respectively. Based on the standard curves, the numbers of each strain on the disks were determined.

2.9. Statistical analysis

Statistical analysis was performed with SPSS, version 22.0 (SPSS Inc., Chicago, IL, USA). One-way analyses of variance (ANOVA) were performed to detect the significant effects of the variables. Tukey’s multiple comparison test was used to compare the means of each of the groups at a p value of 0.05.

3. Results

Typical live/dead images of biofilms at 48 h and 72 h are shown in Fig. 1. The commercial control had mostly green staining of live bacteria. Composites with 0% DMAHDM and 0.75% DMAHDM (not included) had similar images to commercial control. With the DMAHDM content increasing to 1.5% (C, D), 2.25% (E, F), and 3% (G, H), the amount of red staining of compromised bacteria increased in the biofilms.

[1].

Representative live/dead staining images of biofilms: (A, B) commercial control, (C, D) 1.5% DMHADM, (E, F) 2.25% DMHADM, and (G, H) 3% DMHADM. All images had the same magnification as (A). Live bacteria were stained green, and compromised bacteria were stained red.

The CFU results are plotted in Fig. 2 for biofilms on composites at 48 h and 72 h (mean ± sd; n = 6). The commercial control and 0% DMAHDM had similar CFU (p > 0.1). The biofilm CFU on DMAHDM composites were reduced significantly with increasing DMAHDM mass fraction (p < 0.05). The composite with 3% DMAHDM was the most efficient in inhibiting biofilms among all the tested groups (p < 0.05). Compared to control, the CFU in 3% DMAHDM group was reduced by 2–3 orders of magnitude.

[2].

Colony-forming unit (CFU) counts of (A) 48 h biofilms and (B) 72 h biofilms on composites with different DMAHDM mass fractions (mean ± sd; n = 6). Note the log scale for the y-axis. Biofilm CFU counts of 3% DMAHDM group were approximately 2 to 3 orders of magnitude lower than those of controls. Values with dissimilar letters are significantly different from each other (p < 0.05).

The MTT metabolic activity of biofilms on composites is plotted in Fig. 3 (mean ± sd; n = 6). The metabolic activity of commercial control resembled that of the experimental composite with 0% DMAHDM (p > 0.1). A decrease in biofilm metabolic activity was observed with increasing DMAHDM mass fraction (p < 0.05).

[3].

MTT metabolic activity of (A) 48 h biofilms and (B) 72 h biofilms on composites with different DMAHDM mass fractions (mean ± sd; n = 6). The metabolic activity of biofilms decreased with increasing DMAHDM content. Values with dissimilar letters are significantly different from each other (p < 0.05).

Lactic acid productions by biofilms on composites are plotted in Fig. 4 (mean ± sd; n = 6). The acid productions in commercial control and 0% DMAHDM group were similar (p > 0.1). The lactic acid production decreased with increasing DMAHDM mass fraction (p < 0.05).

[4].

Lactic acid production of (A) 48 h biofilms and (B) 72 h biofilms on composites with different DMAHDM mass fractions (mean ± sd; n = 6). Values with dissimilar letters are significantly different from each other (p < 0.05).

The polysaccharides production by biofilms on composites are plotted in Fig. 5 (mean ± sd; n = 6). The commercial control and 0% DMAHDM group had similar polysaccharide amounts (p > 0.1). With the DMADHM mass fraction being increased, the polysaccharide amount decreased remarkably, especially at 3% DMAHDM (p < 0.05).

[5].

Polysaccharides production of (A) 48 h biofilms and (B) 72 h biofilms on composites with different DMAHDM mass fractions (mean ± sd; n = 6). The polysaccharides production of biofilms decreased with increasing DMAHDM content. Values with dissimilar letters are significantly different from each other (p < 0.05).

Fig. 6 displays standard curves of Ct value versus bacteria numbers of S. mutans, S. sanguinis and S. gordonii. The coefficient of determination (R²) of each regression line is approximately to 1, revealing a high linear correlation between Ct and bacteria number in all strains. In ideal condition, the qPCR amplification efficiency is 100% showing as an ideal slope of −3.32. In the present study, the slope of each regression line is in range between −3.61 to −3.27, representing a good amplification efficiency from approximately 90% to 100%.

[6].

Standard curves of Ct versus bacteria number. Genomic DNA of known numbers of (A) S. mutans, (B) S. sanguinis and (C) S. gordonii were used for amplifications. Ct is the cycle number at which the threshold fluorescence was reached. The coefficient of determination (R²) of each regression line is approximately to 1, revealing a high linear correlation. The slope of each regression line represents a good amplification efficiency from approximately 90% to 100%.

The results on bacterial composition shift in the multispecies biofilms on composites are plotted in Fig. 7 (mean ± sd; n = 3). After 48 h, the commercial control and 0% DMAHDM group had a relatively high proportion of S. mutans of 73.3% ± 0.7% and 69.2% ± 2.1%, respectively. The S. mutans proportion decreased with increasing mass fraction of DMAHDM, accounting for 14.5% ± 0.5% in 2.25% DMHADM group, and 10.3% ± 2.3% in 3% DMHADM group. The proportion of S. sanguinis increased with increasing DMAHDM content. The proportion of S. gordonii also showed a rising trend with increasing DMAHDM content.

[7].

Bacterial species shift in 48 h and 72 h biofilms on composites (mean ± sd; n = 3). Cariogenic S. mutans had overwhelming proportions in commercial control and 0% DMAHDM group in both 48 h and 72 h biofilms. However, with increasing DMAHDM mass fraction, the proportion of S. mutans decreased sharply, whereas S. sanguinis or S. gordonii achieved a predominant proportion in the biofilms. Values with dissimilar letters are significantly different from each other (p < 0.05).

At 72 h, S. mutans in commercial control and 0% DMAHDM group reached overwhelming proportions of 92.2% ± 0.5% and 91.2% ± 7.5%, respectively. The S. mutans proportion decreased with increasing DMAHDM, accounting for 45.0% ± 3.9% at 2.25% DMHADM, and as low as 19.9% ± 5.2% at 3% DMHADM. On the other hand, the proportions of S. sanguinis increased with increasing DMAHDM content. The proportions of S. gordonii also increased with increase of DMAHDM content.

4. Discussion

Oral biofilm (plaque) produces acids, causing tooth structure demineralization, leading to caries. It is highly desirable for dental composites to be able to modulate the biofilm composition to promote non-cariogenic species and suppress cariogenic species. However, only one report exists on the use of QAM resin to modulate oral biofilms [30]. The present study developed a DMAHDM-containing dental composite and determined its ability to suppress biofilm growth and modulate biofilm species composition for the first time. The DMAHDM composite significantly inhibited the growth of multispecies biofilms and decreased the cariogenicity of the biofilms by reducing acid production and polysaccharide formation. Furthermore, the DMAHDM composite changed the microbial equilibrium from a cariogenic species-dominated (S. mutans) biofilm to a noncariogenic species-dominated (S. sanguinis and S. gordonii) biofilm. These results show that the DMAHDM composite could induce a biofilm species shift from a caries-risk state to a non-cariogenic state.

QAMs are promising for dental applications including incorporation into composites, primers and adhesives [11, 41]. DMAHDM has a single methacrylate group, a positively charged quaternary amine N⁺ and a long alkyl chain with a chain length of 16. N⁺ can attract negatively charged bacteria cells and disturb the electric balance of cell membrane, leading the bacteria to explode under its own osmotic pressure [33, 36, 42]. In the meanwhile, the long alkyl chain with a chain length of 16 could penetrate bacterial cells, like a needle bursting a balloon [33, 36]. Indeed, recent studies indicated that the quaternary ammonium chain length played an important role in bactericidal effect of QAMs [33, 35, 41]. Regarding the durability of the antibacterial properties, the methacrylate group in DMAHDM allows itself to chemically bond in the resin polymer network, thus displaying long-term and non-release antibacterial effect [32, 43]. As a result, there was no loss in antibacterial properties of DMAHDM-containing composite after 180 days of water-aging [32]. In addition, DMAHDM showed an acceptable and low cytotoxicity matching that of clinically-used dental monomers, as shown in a previous study [35]. The uncured DMAHDM showed less cytotoxicity than BisGMA and similar cytotoxicity to TEGDMA [35]. In regard of the cured resin eluents on cell viability, the DMAHDM-containing group had a similar cell viability to that of a commercial resin control. These results indicate that the incorportion of DMAHDM did not affect the cytotoxicity of resin as compared to clinically-used dental resins [35]. In the present study, to maintain the mechanical properties of the composite, the maximum mass fraction of DMAHDM was 3% according to a previous study [33]. As described in a previous study, the incorporation of up to 3% mass fraction of DMAHDM into the composite did not negatively affect the flexural strength and the elastic modulus as compared to those without DMAHDM [44]. In addition, the incorporation of up to 10% of DMAHDM into an adhesive did not adversely affect the dentin bond strength when compared to that without DMAHDM [25]. In the present study, adding 3% DMAHDM remarkably reduced the metabolic activity and CFU counts of the multispecies biofilms. Compared to control groups, the CFU in 3% DMAHDM group was reduced by 2–3 logs. It should be noted that mature biofilms at 48 h and 72 h were the focus of the present study. This was mainly because that the oral bacteria in the microgaps at the tooth-restoration margins can stay for long periods of time to form mature biofilms, especially in narrow gaps which cannot be cleaned by brushing teeth and are more likely to result in secondary caries. Regarding the early stages of bacteria attachment and biofilm formation, previous studies found that DMAHDM also substantially reduced the early stage of biofilm formation [25,35]. The early attachment of bacteria at 4 h was greatly reduced on DMAHDM-containing dental resin [25,35]. In addition, the early biofilm formation at 24 h was also greatly supressed by the incorporation of DMAHDM [45].

Regarding the cariogenicity of biofilms, acid production by microbial metabolism is long considered to be a key factor leading to the demineralization of teeth [46]. Lactic acid stands for 70% of the organic acids produced in oral biofilm [47]. In the present study, the lactic acid production by biofilms decreased significantly with increasing DMAHDM. Another essential virulence factor of cariogenic biofilm is polysaccharides, including intracellular and extracellular polysaccharides (IPS, EPS) [48–50]. Both IPS and EPS can act as carbohydrate reservoir, allowing the bacteria to continue to produce acid in the absence of exogenous carbohydrates [50, 51]. In addition, EPS serves as a matrix scaffold promoting bacteria colonization and biofilm maturation [52]. Moreover, EPS can act as a barrier to affect the diffusion of substances in and out of the biofilm, which could protect the bacteria from antibacterial agents and thereby improves biofilm resistance [52]. Recent studies revealed that S. mutans was a key contributor in the formation of the EPS matrix in biofilms [49]. Here, we measured the polysaccharides production of the biofilms. We found that the polysaccharides in control groups grew rapidly with the maturation of the biofilms. However, the polysaccharides in 3% DMAHDM group was the lowest at both time points. This could be clinically beneficial, as less polysaccharides indicate less carbohydrate storage to support the biofilm and less barrier to protect the biofilm. As a result, the biofilm would be more vulnerable to environmental challenges such as saliva buffering and fluoride ions. Both the acid production and the polysaccharides results showed that the DMAHDM composite can significantly reduce the virulence and cariogenicity of biofilms.

Dental plaque is a naturally existing ecosystem composed of various microbial species, and the communities are in a dynamic equilibrium under healthy conditions [27]. Oral diseases happen when changes of environmental factors trigger a deleterious shift in the microbial balance to a pathogenic-associated composition [53]. From the ecological view, since it is impossible to eliminate all bacteria, the maintenance of microbial homeostasis is important for the prevention and treatment of dental caries. Previous studies on DMAHDM mostly treated biofilm as a whole community and seldom considered the possible changes in species composition [25, 26]. However, it is conceivable that DMAHDM, an antibacterial agent, could also affect the equilibrium of the biofilm. Therefore, we examined the competition between cariogenic species and non-cariogenic species with DMAHDM. The selected multispecies biofilm consisted of an acidogenic/aciduric pathogen (S. mutans) and two competitive commensal residents (S. sanguinis and S. gordonii). Both S. sanguinis and S. gordonii are early colonizers and common inhabitants in supragingival plaque [54]. They could antagonize S. mutans by producing hydrogen peroxide [55, 56]. Also, they are alkali-generating species contributing to the neutralization of acids in the biofilm [57]. According to clinical studies, higher proportions of S. sanguinis and S. gordonii in dental plaque are associated with healthy tooth surfaces [55, 58]. On the other hand, a higher proportion of S. mutans indicates acidogenicity and cariogenicity [55, 58]. A rencent clinical study revealed that a greater abundance of S. sanguinis and S. gordonii was detected in caries-free individuals, while larger amounts of S. mutans were detected in caries individuals [59]. Therefore, the competition between these species is associated with dental caires. Indeed, the three-species biofilm model consisting of S. mutans, S. sanguinis and S. gordonii was already used to detect the cariogenic tendency of biofilms in a previous study [30]. The three-species biofilm model could offer a relatively diverse microbial environment mimicking interspecies competitions between cariogenic species and non-cariogenic ones, while being experimentally possible to achieve a relatively high degree of reproducibility between the tests. In the present study, with the maturation of biofilm, the ratio of S. mutans in control groups increased and reached an overwhelming proportion over the other two species in 72 h. In contrast, the ratio of S. mutans in the 3% DMAHDM group did not grow and remained much lower than that of control groups. S. sanguinis and S. gordonii in the 3% DMAHDM group showed predominance over S. mutans. It implies that the addition of DMAHDM in the composite brought the biofilm species balance back toward the favorable non-cariogenic and healthy state. A possible explanation to it is that, along with the development of biofilms, acids tend to accumulate due to the microbial metabolism of carbohydrates and thus avail the dominance of acidogenic/aciduric species. The enrichment of acidogenic/aciduric species will in turn further enhance the acidic environment and suppress the growth of the less aciduric commensal residents. This forms a vicious cycle and feeds on each other, eventually leading to the occurrence of dental caries. However, under the pressure of DMAHDM, the acidogenic/aciduric species seemed to lose its dominant position due to less acid production in biofilms. S. mutans was suppressed in the biofilm, mainly due to (1) the direct antibacterial effect from DMAHDM; (2) the indirect acid inhibitition effect from DMAHDM, which could weaken the competitiveness of S. mutans in the multispecies biofilms; (3) the less-acidic environment could help the non-cariogenic and non-aciduric species to survive and grow. Therefore, the proportion shift in the biofilms occurred in the DMAHDM groups, resulting in a more healthy biofilm development tendency. DMAHDM created an environment in favor of the non-cariogenic species, which would be clinically beneficial to the prevention of caries. It should be noted that the human oral cavity harvests a diverse ecosystem including over 700 species of bacteria [60, 61]. While the present study indicated the exciting potential of microbial species modulation by DMAHDM toward a healthy biofilm composition, it was shown in a three-species biofilm model. Further studies are needed to increase the number of species to investigate the microbial modulation by DMAHDM under conditions more closely mimicking the complex biofilm compositions in vivo.

5. Conclusions

This study incorporated DMAHDM into a dental composite and investigated its ability to suppress biofilm growth and modulate biofilm equilibrium for the first time. Adding DMAHDM endowed the composite with a strong reduction in biofilm growth, CFU, metabolic activity, acid production and polysaccharide production. Furthermore, the addition of DMAHDM caused a species shift in the biofilm, reducing the cariogenic S. mutans proportion from 92.2% on commercial composite, to 19.9% on the composite with 3% DMAHDM. These results indicate that DMAHDM has the potential to modulate the biofilm composition shift from a cariogenic state to a non-cariogenic state. Hence, DMAHDM is promising for applications in preventive and restorative dentistry to combat dental caries.

Highlights.

• Developed a bioactive dental composite containing DMAHDM.

• Composite with 3% DMAHDM achieved greatest reduction in multispecies biofilm.

• Biofilm equilibrium shifted from a cariogenic species-dominated state to a noncariogenic species-dominated state.

• The new composite is promising for cavity restorations to combat caries.