Green-synthesized silver nanoparticles against Streptococcus mutans: antibacterial activity and transcriptomic insights into planktonic and biofilm states

Abstract

Dental caries, one of the most common infectious diseases worldwide, is closely associated with Streptococcus mutans (S. mutans) biofilms that exhibit strong resistance to conventional antimicrobial agents. Herein, a green synthesis of silver nanoparticles (AgNPs) is reported to use extracellular metabolic products derived from Pseudomonas putida KT2440 as natural reducing, capping, and stabilizing agents for antibacterial therapy. The resulting AgNPs possess nanoscale size, negative surface charge, and excellent colloidal stability. These green AgNPs display potent antibacterial and antibiofilm activities against S. mutans, significantly disrupting bacterial membranes, suppressing acidogenicity, and inducing metabolic dysfunction. Biofilm evaluation further revealed a marked reduction in bacterial and extracellular polysaccharide biomass, indicating the collapse of the three-dimensional biofilm structure. To gain deeper insight into the molecular mechanisms of AgNPs-mediated antibacterial activity, RNA sequencing (RNA-seq) was conducted, revealing significant transcriptional reprogramming associated with the inhibition of metabolic and translational processes, disruption of cell wall homeostasis, suppression of virulence gene expression, and perturbation of carbohydrate metabolism. This study presents an environmentally friendly and effective strategy that bridges green nanotechnology with oral microbiology, offering a sustainable approach for caries prevention and biofilm control.

Graphical abstract

1. Introduction

Dental caries remains one of the most prevalent oral diseases worldwide, affecting individuals of all ages [1]. Streptococcus mutans (S. mutans), a Gram-positive bacterium, plays a pivotal role in caries pathogenesis. Thriving in the oral cavity, S. mutans adheres to tooth surfaces, forms biofilms, and metabolizes dietary sugars into acids that demineralize enamel [2,3]. In addition, glucosyltransferases produced by S. mutans catalyze the synthesis of polysaccharides from sucrose, facilitating interspecies and intraspecies aggregation within biofilms and the development of complex microcolonies [4,5]. These polysaccharides not only provide nutrients to enhance metabolic activities within the biofilm matrix but also protect bacterial cells from antimicrobial agents and other environmental assaults [6,7]. Effective management of S. mutans is therefore critical for controlling dental caries and maintaining oral health [8]. Conventional caries management strategies, such as fluoride application and antimicrobial therapies, have markedly reduced disease prevalence but face notable challenges. The extensive extracellular matrix of biofilms acts as a diffusion barrier, limiting the penetration and efficacy of antibiotics, while the overuse of antimicrobials has contributed to the emergence of drug resistance [9,10]. These limitations underscore the urgent need for safe, effective, and environmentally sustainable alternative strategies (see Scheme 1).

Scheme 1.

Schematic presentation of green AgNPs formation (A) and their antimicrobial activity against planktonic S. mutans with transcriptomic analysis (B), as well as antibiofilm activity against S. mutans (C).

Nanotechnology offers a promising avenue to address these challenges, particularly using nanoparticles with antimicrobial properties [11]. Green AgNPs are well known for their strong broad-spectrum antimicrobial properties and ability to disrupt biofilms, making them well suited for targeting S. mutans [12,13]. Compared to antibiotics, AgNPs have a lower potential for inducing resistance. Green AgNPs offer unique advantages such as stability and biocompatibility [14]. The use of bioactive compounds derived from natural sources, including plant extracts and microorganisms, has attracted increasing attention in the green synthesis of nanoparticles, as these biomolecules can enhance biocompatibility while simultaneously improving antimicrobial performance [15,16]. AgNPs produced through such green approaches are typically stabilized by naturally derived reducing and capping agents, which may further confer favorable biological interactions. Owing to these advantages, green-synthesized AgNPs have been increasingly explored for biomedical and oral health applications. In particular, several studies have demonstrated their potent antibacterial activity against the cariogenic bacterium S. mutans, highlighting their potential relevance for dental caries prevention [[17], [18], [19]].

Among green synthesis strategies, bacterial-mediated approaches have gained increasing attention as sustainable routes for producing silver nanoparticles through extracellular processes. In such systems, secreted biomolecules including proteins, carbohydrates, and exopolysaccharides can facilitate silver ion reduction and remain associated with the nanoparticle surface, thereby influencing particle stability and surface chemistry. This synthesis route has been previously established using Pseudomonas putida KT2440 and provides a reproducible platform for generating green AgNPs suitable for mechanistic antibacterial investigations [16].

Although AgNPs are well known for their broad-spectrum antimicrobial properties, their activity against the cariogenic bacterium S. mutans remains insufficiently explored. Moreover, most existing studies primarily report phenotypic outcomes, while the underlying cellular and molecular responses of S. mutans to AgNPs exposure remain poorly understood. Therefore, this study aimed to investigate the antibacterial potential of green AgNPs against S. mutans by integrating physicochemical characterization, multiscale antibacterial and antibiofilm evaluation, and transcriptomic profiling via RNA sequencing (RNA-seq). In the context of dental caries-associated biofilm, this work integrates an established Pseudomonas putida KT2440–based green AgNPs synthesis platform with comprehensive antibiofilm assessment and transcriptome-wide analysis of planktonic S. mutans, providing system-level insights into bacterial responses to nanoparticle exposure.

2. Experimental sections

2.1. Green synthesis of AgNPs

The isolated strain Pseudomonas putida KT2440 was cultured overnight in 100 mL of tryptic soy broth (TSB) at 37 °C with shaking at 120 rpm. The culture was then centrifuged at 8000 rpm for 10 min to remove the cells. The resulting cell-free supernatant was supplemented with 2 mM AgNO₃ and incubated in a shaking flask at 37 °C and 200 rpm for 48 h to facilitate AgNPs formation. The nanoparticles were subsequently purified by centrifuging the reaction mixture at 3000 rpm for 5 min to remove residual debris, followed by a second centrifugation at 14,000 rpm for 15 min. After discarding the supernatant, the pellets were washed several times with distilled water, and the purified AgNPs were resuspended in water for use in subsequent experiments.

2.2. Analytical characterization of AgNPs

The UV–Vis spectrum of the AgNPs was recorded on an Agilent Cary 60 spectrophotometer over the wavelength range of 200–800 nm. Zeta potential measurements were performed with a Nano ZS Zetasizer to assess particle surface charge. Attenuated total reflectance Fourier-transform infrared (ATR-FTIR) spectroscopy was used to identify biomolecules and functional groups involved in nanoparticle reduction and capping. ATR-FTIR spectra were collected on a Bruker Alpha spectrometer equipped with a diamond crystal across 500–4000 cm⁻¹ at a resolution of 4 cm⁻¹. Transmission electron microscopy (TEM) was conducted with an FEI Tecnai T20 G2 operating at 200 kV to examine AgNPs morphology. Samples for TEM were prepared by depositing a drop of the aqueous AgNPs suspension onto a carbon-coated copper grid and air-drying prior to analysis. The chemical composition of the AgNPs was determined by X-ray photoelectron spectroscopy (XPS) using a PHI VersaProbe III with a monochromatized Al Kα (1486.6 eV) X-ray source; spectra were acquired at a 90° take-off angle and processed with CasaXPS software. To quantify AgNPs concentration, single-particle inductively coupled plasma mass spectrometry (sp-ICP-MS) was performed using a PerkinElmer NexION 350D instrument.

2.3. Bacterial strains and culture media

S. mutans UA159, a model cariogenic bacterium, was used to evaluate the antibacterial and anti-cariogenic activity of green AgNPs. S. mutans UA159 was obtained from the American Type Culture Collection and cultured in Brain Heart Infusion (BHI) broth at 37 °C in a 5% CO₂ atmosphere.

2.4. Determination of the minimum inhibitory concentration (MIC)

The minimum inhibitory concentration (MIC) of AgNPs against S. mutans was determined using the broth microdilution method in Brain Heart Infusion (BHI) broth. Overnight cultures of S. mutans were diluted in BHI broth to a final concentration of approximately 5 × 10⁵ CFU/mL in each well of a 96-well microplate. Serial two-fold dilutions of AgNPs were prepared in BHI broth, and the plates were incubated at 37 °C in 5 % CO₂ for 24 h. Bacterial growth was evaluated by measuring the optical density at 600 nm with a microplate reader (Varioskan Lux, Thermo Scientific). MIC assays were performed using three independent biological replicates, and the MIC was defined as the lowest AgNPs concentration that completely inhibited visible bacterial growth after 24 h incubation.

2.5. Antibacterial assay for planktonic cells

Time-kill assays were performed in triplicate to evaluate the growth of S. mutans in response to increasing concentrations of AgNPs. Overnight cultures of S. mutans were diluted in Brain Heart Infusion (BHI) broth to yield an inoculum of approximately 1 × 10⁷ CFU/mL. AgNPs were added at final concentrations corresponding to 0 (control), 1, 2, 4 and 8 × the MIC. Cultures were incubated at 37 °C in 5 % CO₂, and at 0, 2, 4, 8 and 24 h, 100 μL aliquots were used for viability assessment. The samples were serially diluted, plated on agar plates, and incubated at 37 °C under 5% CO₂ for 48 h. Colony counts were then performed to quantify bacterial survival. In addition, during the time-kill experiments described above, the pH of each culture was measured at 0, 2, 4, 8 and 24 h.

To evaluate bacterial viability microscopically, Live/Dead staining was performed on cells harvested after 4 h of incubation from the control, 2 × MIC and 4 × MIC AgNPs treatments. 1 mL of culture from each condition was centrifuged for 5 min at 4 °C, and the pellets were washed twice with sterile phosphate-buffered saline (PBS, 10 mM, pH 7.4). Cells were stained with the Live/Dead BacLight™ bacterial viability kit (Thermo Fisher Scientific) according to the manufacturer's instructions. After incubation for 15 min in the dark at room temperature, 10 μL aliquots of the stained suspension were placed on glass slides and observed immediately under a fluorescence microscope. Images were captured with a performed using a Zeiss fluorescence microscope (Zeiss Axio Imager Z2m, Germany) and processed using ImageJ software to estimate the proportion of live and dead cells.

For ultrastructural analysis, cell pellets from the same conditions (4 h, 2 × MIC, 4 × MIC, and control) were fixed in 2.5 % (v/v) glutaraldehyde at 4 °C overnight. After fixation, cells were washed three times with PBS, dehydrated using a graded ethanol series (30, 40, 50, 60, 70, 80, 90, and 100 %, 10 min each), and dried. The dried samples were sputter-coated with gold using a sputter coater and observed under a scanning electron microscope (SEM, JEOL JSM 6301F) to visualize morphological changes induced by AgNPs.

2.6. Transcriptome analysis

Approximately 1 × 10⁷ bacterial cells with or without AgNPs treatment were harvested by centrifugation at 14,000×g and flash-frozen to minimize mRNA degradation. Total RNA was extracted using the RNeasy Mini Kit (GIAGEN, Germany) according to the manufacturer's instructions. Cells were disrupted using a FastPrep homogenizer (MP Biomedicals, USA), and genomic DNA was removed using DNase I (Invitrogen, USA). RNA concentration and integrity were determined using the Qubit RNA HS Assay Kit (Thermo Fisher Scientific, USA) and an Agilent 2100 Bioanalyzer (Agilent Technologies, USA), respectively. RNA sequencing was performed at the SciLifeLab National Genomics Infrastructure (NGI), Sweden. Sequencing libraries were prepared from 300 ng of total RNA using the Illumina Stranded Total RNA Library Preparation Kit. Sequencing was performed on the Illumina NovaSeq X Plus platform. Three independent biological replicates were performed for each condition.

Sequencing data were processed using standard bioinformatics pipelines. Raw sequencing data were processed with FASTP to remove adapter sequences and low-quality reads, generating clean reads. Clean reads were then mapped to the reference genome using Bowtie2. Gene expression levels were quantified with RSEM. Sample correlation was assessed by calculating correlation coefficients among biological replicates using the R package psych, and principal component analysis (PCA) was performed with the R package gmodels to evaluate overall similarity and clustering relationships among samples. Gene expression levels were normalized using the FPKM method to account for differences in gene length and sequencing depth. Differentially expressed genes (DEGs) were identified with edgeR, using thresholds of fold change ≥2 and false discovery rate (FDR) < 0.05. Identified DEGs were subjected to functional annotation with Gene Ontology (GO) terms and pathway enrichment analysis using the Kyoto Encyclopedia of Genes and Genomes (KEGG) database, with significance defined as FDR <0.05. Gene set enrichment analysis (GSEA) was conducted using the GSEA software (Broad Institute) in combination with the MSigDB database to identify significantly enriched GO terms and KEGG pathways between groups. Briefly, gene expression profiles were ranked using the signal-to-noise ratio method, and enrichment scores (ES) and p-values were calculated under default parameters.

2.7. Antibiofilm assay

The ability of AgNPs to inhibit biofilm formation was assessed by the crystal violet (CV) staining method. Overnight culture of S. mutans was diluted 1:100 in BHI broth supplemented with 1 % (w/v) sucrose to obtain an inoculum of approximately 2-5 × 10⁶ CFU/mL and dispensed into sterile 96-well microplates (200 μL per well). AgNPs were added at final concentrations of 0.125, 0.25, 0.5, 1, 2, 4 and 8 × MIC, and the plates were incubated at 37 °C in 5 % CO₂ for 24 h to allow biofilm development. Non-adherent cells were removed by washing the wells gently three times with sterile phosphate-buffered saline (PBS, pH 7.4). Biofilms were stained with 0.1 % crystal violet for 20 min at room temperature, rinsed three times with distilled water and air-dried. The bound stain was solubilized with 33 % acetic acid for 30 min, and absorbance was measured at 595 nm using a multimode microplate reader (Varioskan LUX). Furthermore, SEM was used to visualize the effect of AgNPs on biofilm formation. Briefly, biofilms for SEM analysis were formed on sterile cover glass in the presence of AgNPs (2, 4,8 and 16 × MIC) for 24 h. Biofilms were fixed with glutaraldehyde followed by the dehydration with graded series of ethanol. Dehydrated biofilms were dried and sputter coated with gold before acquiring SEM images.

To assess the effect of AgNPs on established biofilms, sterile cover glass was placed into the wells of a 24-well plate. Each well contained 1 mL of BHI broth supplemented with 1 % (w/v) sucrose and was inoculated with S. mutans as described above. The plates were incubated at 37 °C in 5 % CO₂ for 24 h to allow biofilm formation on the substrates. After biofilm formation, the medium was removed, and the wells were gently washed three times with PBS to remove planktonic cells. Fresh BHI broth (1 mL) containing AgNPs at final concentrations of 1, 2, 4, 8 and 16 × MIC was then added, and the plates were incubated at 37 °C in 5 % CO₂ for 24 h. Biofilms were detached from the cover glass substrates by sonication using a Digital Sonifier (Branson) at 10 % amplitude for 30 s, three cycles, on ice, and the resulting suspensions were collected in 5 mL sterile saline (0.89 % NaCl). The suspensions were serially diluted (ten-fold) in sterile saline and 100 μL aliquots were plated on BHI agar plates. Plates were incubated at 37 °C in 5 % CO₂ for 48 h, and colonies were counted to determine viability of cells. Morphological differences of S. mutans biofilms under AgNPs treatment were examined by SEM. For SEM, biofilms were fixed and dehydrated as described above. The dried biofilms were sputter coated before acquiring SEM images.

Confocal laser scanning microscope was used for three-dimensional visualization of live/dead cells and extracellular polymeric substances (EPS) within the biofilms. EPS was labeled in situ by adding Alexa fluor® 647-labeled dextran conjugate to the culture medium as described previously [20]. After 48 h of incubation, biofilms were exposed to 2.5 μM SYTO 9 green-fluorescent nucleic acid stain for 30 min before imaging. For viability assessment, the biofilms were stained with the Live/Dead BacLight™ bacterial viability kit (SYTO 9 and PI) for 20 min in the dark. Stained samples were mounted on glass slides and imaged using a Nikon Ti-E A1+ confocal laser scanning microscope. Five fields of view were captured for each sample. Biofilm was viewed in a three-dimensional framework using Imaris Viewer 7.0 and the biomass and distribution of bacterial and EPS were quantified using the image analysis software ImageJ COMSTAT.

2.7.1. Intracellular reactive oxygen species (ROS) detection

To assess intracellular reactive oxygen species (ROS), bacterial cultures from the control group and the AgNPs treated groups (2 × MIC and 4 × MIC) were collected after 4 h of incubation. Cells were stained using CellROX Deep Red Sensor (Life Technologies) and DAPI. Briefly, bacterial cells were first incubated with CellROX Deep Red dye for 20 min, followed by washing to remove excess dye. Cells were then counterstained with DAPI for 20 min, washed again, and observed under a fluorescence microscope. Fluorescence images were acquired sequentially, and identical imaging parameters were applied to all groups.

2.7.2. Intracellular ATP quantification

Intracellular ATP levels were quantified using a luciferase-based ATP determination assay (ATP Determination Kit, Thermo Fisher Scientific) according to the manufacturer's instructions. Briefly, equal culture volumes from the control and AgNPs treated groups (0.25 × MIC, 0.5 × MIC and 1 × MIC) were collected after 4 h of incubation. Cells were harvested by centrifugation and lysed to release intracellular ATP. The supernatant was mixed with the luciferase reaction reagent, and luminescence was immediately measured using a microplate reader. Luminescence signals were normalized to the untreated control. All measurements were performed in biological triplicates.

2.7.3. Lactate quantification

Extracellular lactate levels were quantified using the Lactate Assay Kit (MAK570, Sigma-Aldrich) according to the manufacturer's instructions. Briefly, bacterial cultures from the control and AgNPs treated groups (0.5 × MIC, 1 × MIC and 2 × MIC) were incubated for 4 h and 24 h. Equal culture volumes were collected and centrifuged to remove cells, and the resulting supernatants were used for lactate measurement. Absorbance was recorded at 570 nm using a microplate reader, and lactate concentrations were calculated based on a standard curve. All experiments were performed in triplicate.

2.8. Statistical analysis

All experiments were performed in triplicate, and data are expressed as the mean ± standard deviation (SD). Statistical analyses were carried out using one-way analysis of variance (ANOVA) followed by Tukey's post-hoc multiple comparison test. A p-value < 0.05 was deemed statistically significant. The statistically significant differences were indicated by asterisks (*p < 0.05; **p < 0.01; ***p < 0.001).

3. Result and discussions

Transmission electron microscopy (TEM) images (Fig. 1A) revealed that the green-synthesized AgNPs exhibited diverse morphologies, including spherical, triangular, and polygonal shapes. The particle size distribution ranged from 15 to 50 nm. The average particle diameter of 27.3 nm was determined by statistical analysis of individual AgNPs measured from TEM images, as shown in the particle size distribution histogram in Fig. 1C (upper panel). Additional TEM images are shown in Fig. S1. Atomic force microscopy (AFM) provided complementary topographical information, showing particle heights between 15 and 40 nm, with an average height of 19.1 nm (Fig. 1C, lower). These AFM measurements are consistent with the TEM and further confirm the quasi-spherical or plate-like geometry of the particles (Fig. 1B). The size difference between TEM and AFM arises from intrinsic methodological differences, as TEM predominantly reflects the inorganic silver core, whereas AFM may include contributions from surface-associated organic layers derived from the green synthesis process and tip convolution effects. FTIR spectra of the AgNPs and their biological corona displayed several characteristic absorption bands, as shown in Fig. 1D. A broad band around 3200-3400 cm⁻¹ corresponds to O-H stretching vibrations of phenolic compounds and N-H stretching of amide or amine groups. Peaks at 2900-2950 cm⁻¹ are attributed to the asymmetric stretching of methyl groups (-CH₃) as well as C-H stretching of alkanes or secondary amines. An intense band near 1600–1650 cm⁻¹ indicates C=O (amide I) stretching vibrations of proteins or peptides. Bands at 1500 cm⁻¹ can be assigned to N–H bending and C-N stretching vibrations (amide II region), while the band at 1460 cm⁻¹ corresponds to C-H bending. Additional absorptions in the 1100 cm⁻¹ region arise from C-O and C-O-C stretching vibrations characteristic of carbohydrates or glycosidic linkages [16,21]. Overall, the FTIR results indicate that proteins, amino acids, carbohydrates, and other metabolites participate in the reduction and stabilization of the nanoparticles. These FTIR features collectively indicate the presence of extracellular biomolecules associated with the AgNPs surface. However, the observed functional groups cannot be unambiguously assigned to specific molecular species or linked to distinct functional roles such as exclusive reduction or stabilization. AgNPs formation is therefore interpreted as a cooperative process in which multiple extracellular metabolites contribute to Ag⁺ reduction and subsequently remain associated with the nanoparticle surface, providing stabilization [16]. The UV-Vis absorption spectra of the AgNPs were recorded in the 200-800 nm wavelength range (Fig. 1E). The AgNPs exhibited a distinct optical absorption in the visible region (around 419 nm), indicative of silver nanoparticle formation. Zeta potential analysis showed a value of −23.6 mV for the AgNPs (Fig. S2). The negative zeta potential suggests electrostatic repulsion between particles and further supports the colloidal stability of the nanoparticles in aqueous solution. The survey XPS spectrum of the biosynthesized AgNPs (Fig. S3) confirmed the presence of C, N, Ag, O, Na and Cl elements originating from the nanoparticle core and the bacteria-derived biomolecules adsorbed on the surface. High-resolution XPS of the Ag3d region (Fig. 1F) displayed two intense peaks at binding energies of approximately 368.0 eV and 374.0 eV, corresponding to Ag3d₅/₂ and Ag3d₃/₂, respectively. The spin-orbit splitting of 6.0 eV and the peak positions are characteristic of metallic silver (Ag⁰), indicating that most of the silver in the nanoparticles is present in the reduced state. No significant additional peaks attributable to Ag⁺ were detected, suggesting that the bacterial reduction process efficiently converted Ag ⁺ to Ag⁰ [22]. These results corroborate the UV-Vis and FTIR findings and confirm the successful formation of metallic AgNPs capped with bacterial biomolecules. In this study, no intrinsic antimicrobial activity is specifically attributed to the surface-associated biomolecules themselves; rather, the antibacterial effects observed are primarily ascribed to the AgNPs exposure. Collectively, the results demonstrate that the green synthesis produced stable, morphologically diverse AgNPs with biomolecular capping.

Fig. 1.

Physicochemical characterization of green AgNPs. (A) TEM image of AgNPs, (B) AFM image (scale bar 500 nm), (C) Particle size distribution from TEM (upper) and height distribution from AFM (lower), (D) FTIR spectra, (E) UV-Vis absorption spectrum, and (F) High-resolution XPS Ag3d spectrum of AgNPs.

The MIC of green AgNPs against S. mutans was consistently determined to be 4 μg/mL across all independent replicate experiments. The MIC of green AgNPs against S. mutans determined in this study (4 μg/mL) is lower than MIC values reported for some other green-synthesized AgNPs systems [17,18]. Such variation may arise from differences in nanoparticle surface chemistry, associated biomolecular capping and bacterial strains. Time-killing assay revealed concentration-dependent bactericidal activity, as shown in Fig. 2A. At 1 × MIC and 2 × MIC, AgNPs treatment mainly resulted in growth suppression with only a gradual decline in viable counts. In contrast, exposure to 4 × MIC and 8 × MIC produced a pronounced and rapid reduction in viable cells within the first 4 h, followed by a progressive decrease such that no viable S. mutans cells were detected by 24 h. S. mutans is highly acidogenic; its ability to ferment dietary carbohydrates and lower environmental pH is a central virulence factor in dental caries [23,24]. Since acid production is a key virulence factor of S. mutans, we monitored changes in culture pH to assess the effect of AgNPs treatment on acid production (Fig. 2B). Untreated S. mutans cultures exhibited the expected progressive decrease in pH over time, reflecting normal acid production during metabolism. At 0.5 × MIC, the pH profile was comparable to the control, indicating minimal effects on acidogenic activity. At 1 × MIC, AgNPs delayed the onset of acidification, with only a modest decrease in pH observed by 24 h. At 2 × MIC, the pH remained near the initial value throughout the 24 h incubation, indicating a near-complete inhibition of detectable acid production. Importantly, when considered together with the time-kill data (Fig. 2A), viable CFU were still present under these conditions, demonstrating that the absence of acidification cannot be attributed solely to reduced bacterial numbers. Analysis of net pH changes together with corresponding viable counts (Fig. S4) further supports that AgNPs suppress the acidogenic metabolism of S. mutans beyond effects attributable solely to viability loss.

Fig. 2.

Antibacterial effects of green AgNPs on S. mutans. (A) Time-kill curves of S. mutans exposed to AgNPs at different concentrations. (B) pH changes of culture medium over 24 h under AgNPs treatment. (C) Quantitative analysis of live/dead fluorescence ratios after 4 h exposure to AgNPs at 2 × MIC and 4 × MIC. (D) Live/dead staining images after 4 h exposure to AgNPs at 2 × MIC and 4 × MIC. (E) SEM images of S. mutans cells after 4 h exposure to AgNPs at 2 × MIC and 4 × MIC.

To assess whether the loss of viability was accompanied by membrane damage and morphological alterations, Live/Dead staining and SEM imaging were performed after 4 h exposure to 2 × MIC and 4 × MIC. Live/Dead staining images (Fig. 2D) showed that untreated cells were predominantly stained with SYTO9, indicating intact membranes within the population. Following AgNPs treatment, a marked increase in PI-stained cells was observed, suggesting compromised membrane integrity. The analysis of fluorescence ratios (Fig. 2C) indicated that approximately 40-50 % of cells exhibited membrane damage in the treated groups. CFU counts (Fig. 2A) were used as the primary indicator of antibacterial activity. In contrast, live/dead staining assesses cell membrane integrity at a defined time point. This staining method may detect membrane-intact but non-culturable subpopulations, and thus the results do not directly correspond to population-level growth behavior measured by time-kill assays [25]. SEM imaging provided morphological alterations in cells corresponding to the loss in viability (Fig. 2E). Untreated S. mutans cells displayed smooth, coccoid surfaces with intact division septa. In contrast, after 4 h exposure to AgNPs at 2 × MIC and 4 × MIC, many cells showed obvious surface roughening, wrinkling and partial collapse, with fragments of disrupted cells. The time-kill assay, pH measurements, live/dead staining and SEM consistently showed that AgNPs treatment reduced the viability of S. mutans, strongly suppressed acid production and caused marked ultrastructural alterations in cell morphology compared to control. Although both the green-synthesized AgNPs and the Gram-positive bacterium S. mutans exhibit an overall negative surface charge, electrostatic repulsion does not preclude antibacterial activity. The antimicrobial effects of AgNPs are not solely dependent on direct nanoparticle-cell surface adhesion but are attributed to the release of Ag⁺ ions, which can penetrate the bacterial cell envelope and interact with intracellular targets, leading to enzyme inactivation and metabolic disruption [26,27]. In addition, AgNPs-bacteria interactions are governed by multiple mechanisms beyond electrostatic attraction, including membrane disruption and localized physicochemical interactions at the nano-bio interface, collectively enabling effective antibacterial activity despite like-charge repulsion [28]. To further investigate the molecular basis of these phenotypic changes, we next performed RNA sequencing (RNA-seq) analysis.

Gene expression profiles of S. mutans treated with AgNPs were analyzed and compared with untreated controls. Transcriptomic analysis was performed to characterize system-level transcriptional responses of S. mutans under stress induced by AgNPs. This approach provides insight into the global cellular response under antibacterial pressure. The 2 × MIC and 4 × MIC concentrations were selected to represent increasing levels of antibacterial stress. Transcriptomic analysis was conducted after 4 h of AgNPs exposure to capture transcriptional responses during active antibacterial stress while substantial viable cells were still present (Fig. 2A), thereby reflecting biologically relevant adaptive changes. Comparison of these two stress levels enabled identification of common stress-response pathways as well as transcriptional programs associated with intensified antibacterial pressure. Principal component analysis (PCA) revealed a clear separation between the treatment and the untreated control groups, with no overlap observed among biological replicates (Fig. 3A). PCA reflects the overall variance across the entire transcriptome rather than changes in individual genes. Therefore, the distinct clustering indicates a consistent and global transcriptional reprogramming induced by AgNPs exposure, reflecting systematic differences in gene expression profiles between the two groups. Pearson correlation analysis is shown in Fig. S5. Volcano plots (Fig. 3B) showed that 903 differentially expressed genes (DEGs) were identified in the 2×MIC group compared with the control, including 494 upregulated and 409 downregulated genes. In the 4×MIC group, 742 DEGs were detected, with 422 upregulated and 320 downregulated genes. Venn diagram analysis (Fig. S6) further illustrated the overlap and specificity of DEGs: 684 DEGs were shared by both treatment groups, while 220 were unique to the 2×MIC group and 59 were unique to the 4×MIC group. To further investigate the functional roles of these genes, GO and KEGG enrichment analyses were performed. Top 20 enriched GO terms (Fig. 3C) showed that, at 2 × MIC, differentially expressed genes were broadly enriched in pathways associated with amino acid biosynthesis and diverse metabolic processes, indicating overall metabolic adjustments at the cellular level. In contrast, the 4 × MIC group additionally exhibited enrichment in pathways related to fatty acid metabolism, ATP biosynthesis, and energy-generating processes, suggesting functional associations with membrane lipid metabolism and cellular energy metabolism. Consistently, Top 20 enriched KEGG pathways (Fig. 3D) revealed that genes in the 2 × MIC group were primarily enriched in amino acid biosynthesis, the phosphotransferase system (PTS), and various metabolic pathways, whereas the 4 × MIC group additionally involved fatty acid metabolism and oxidative phosphorylation. At both 2 × MIC and 4 × MIC, KEGG analysis showed significant enrichment of PTS-related pathways. Hpr, one of the crucial components in PTS, was downregulated at both 2 × MIC and 4 × MIC, compared with the untreated control. The PTS serves as the primary carbohydrate uptake system in S. mutans, supplying substrates for glycolysis and acid production. Therefore, it is functionally associated with cariogenicity and provides essential carbon and energy sources for biofilm formation [29,30]. In addition, GO and KEGG analyses in the 4 × MIC group showed enrichment of pathways related to fatty acid and energy metabolism. Fatty acid metabolism in S. mutans is primarily involved in regulating membrane lipid composition and membrane remodeling, which are important for adaptation to acidic environments and maintenance of membrane stability. Previous studies have shown that S. mutans dynamically adjust membrane fatty acid composition to enhance acid tolerance and stress resistance, thereby promoting survival and persistence within mature biofilms [31,32]. Meanwhile, energy-related processes, including ATP biosynthesis and oxidative phosphorylation, support multiple energy-dependent physiological activities such as extracellular polysaccharide synthesis, proton transport, and biofilm matrix maintenance, all of which are closely associated with cariogenicity and biofilm formation. To further characterize the distribution of DEGs, GO and KEGG annotation analyses were also performed. GO classification indicated that most DEGs were assigned to biological process and molecular function categories, with metabolism representing the predominant class in both the 2 × MIC and 4 × MIC groups (Fig. S7). KEGG annotation further showed that metabolic pathways accounted for the largest proportion of affected genes, followed by genetic information processing, environmental information processing, and cellular processes (Fig. S8). These findings indicate that AgNPs exposure induces transcriptional alterations in central metabolism, membrane adaptation, and energy homeostasis, processes that are functionally relevant to cariogenicity and biofilm formation [[33], [34], [35]].

Fig. 3.

Transcriptomic effects of AgNPs treatment on S. mutans. (A) PCA plot of control, 2 × MIC and 4 × MIC samples. (B) Volcano plots of differentially expressed genes (DEGs) for 2 × MIC vs control and 4 × MIC vs control (red: upregulated; orange: downregulated; grey: non-significant). (C) Top 20 enriched biological processes of DEGs based on Gene Ontology (GO) analysis. (D) Top 20 enriched pathways of DEGs based on Kyoto Encyclopedia of Genes and Genomes (KEGG) analysis.

Gene set enrichment analysis (GSEA) further revealed the systemic effects of AgNPs on the S. mutans transcriptome. The global NES heatmap showed that both 2 × MIC and 4 × MIC treatments led to significant negative enrichment of numerous metabolisms, peptidoglycan biosynthesis and translation-related pathways, whereas signal transduction pathways were upregulated (Fig. 4A). Comprehensive GSEA enrichment plots for all significantly affected pathways are provided in Fig. S9, further confirming that metabolic and translational pathways were broadly suppressed, while signal transduction pathways were activated. To further illustrate the enrichment of key pathways, representative GSEA of gene expression profiles are shown in Fig. 4B and C. In both concentrations, glycolysis/gluconeogenesis and ribosome pathways displayed strong negative enrichment, whereas the two-component system showed positive enrichment, indicating suppression of central metabolism and translation accompanied by activation of stress signaling. Aminoacyl-tRNA biosynthesis was also negatively enriched, consistent with ribosome inhibition suggesting the suppression of translation [36]. Δ|NES| comparison revealed that the ribosome pathway exhibited the larger difference between 2 × MIC and 4 × MIC, followed by glycolysis/gluconeogenesis and the two-component system (Fig. S10), providing support for more extensive negative enrichment at 4 × MIC.

Fig. 4.

(A) Gene set enrichment analysis (GSEA) heatmap of KEGG pathways with normalized enrichment scores (NES). (B) GSEA of gene expression profiles with the KEGG database for 2 × MIC vs control. (C) GSEA of gene expression profiles with the KEGG database for 4 × MIC vs control. (D) Integrated schematic illustrating major transcriptional responses of S. mutans to AgNPs exposure. Differentially expressed genes identified by transcriptomic analysis were grouped into functional modules. The inside heatmaps indicate relative gene expression changes in 2 × MIC and 4 × MIC compared with the control.

These pathway-level changes were consistent with the phenotypic alterations. Exposure to AgNPs at 2 × MIC inhibited acid production in S. mutans (Fig. 2B). Correspondingly, glycolysis, a key pathway for carbohydrate utilization and acid production, was markedly downregulated at both concentrations (Fig. 4A). Representative glycolytic genes, including pfkA and eno, exhibited decreased expression in the gene heatmap (Fig. 4D), providing gene-level support for the reduced acidogenic activity. In addition, SEM images (Fig. 2E) revealed the marked morphological alterations and membrane damage after 2 × MIC and 4 × MIC AgNPs treatment. Consistently, pathways associated with cell envelope homeostasis were transcriptionally perturbed, including fatty acid metabolism and peptidoglycan biosynthesis. Representative genes involved in fatty acid biosynthesis (e.g., acc and fab genes) showed decreased expression (Fig. 4D). In terms of cell wall biosynthesis, several penicillin-binding proteins (pbp1a, pbp2b, and pbp3) were downregulated, while pbp2a was upregulated, suggesting that AgNPs exposure may disturb cell wall homeostasis and potentially trigger compensatory responses [37,38]. Importantly, these metabolic and envelope-associated enrichments are interpreted as system-level physiological responses to AgNPs induced cellular and membrane stress rather than evidence of a single primary antibacterial target.

To integrate these transcriptional responses, representative differentially expressed genes were grouped according to functional modules and visualized in a schematic heatmap (Fig. 4D), highlighting coordinated alterations across metabolism, energy production, translation, membrane and cell wall homeostasis, and oxidative stress response modules. Energy metabolism was substantially perturbed. Genes encoding ATP synthase subunits (e.g., atpA and atpD) were downregulated, suggesting impaired ATP generation. Reduced transcription of these energy-related genes implies compromised cellular energy homeostasis, which may further affect multiple ATP-dependent processes, including biosynthesis, metabolism, membrane maintenance, and stress adaptation. In addition, pathways related to translation exhibited strong negative enrichment. Multiple ribosomal proteins (e.g. rpl, rps, and rpm) were consistently downregulated, together with aminoacyl-tRNA biosynthesis genes, indicating global suppression of translational capacity. Such inhibition of the ribosomal machinery is expected to limit protein production and cellular growth, consistent with the overall antibacterial effects of AgNPs [39,40]. In parallel, multiple genes associated with oxidative stress defense were consistently upregulated, including components of the thiol redox system (trxB and gorA), peroxide detoxification enzymes (ahpF), NADH oxidase (nox), the redox-responsive transcriptional regulator (nmlR), and the protein repair enzyme (msrA). These genes are commonly involved in maintaining intracellular redox balance, detoxifying reactive oxygen species, and repairing oxidatively damaged proteins. Their coordinated induction suggests that AgNPs exposure imposes substantial oxidative stress on bacterial cells [[41], [42], [43]].

To further evaluate whether the transcriptional alterations translated into physiological changes, functional assays were performed to assess intracellular oxidative stress, energy metabolism, and acidogenic activity, respectively. Transcriptomic analysis revealed significant upregulation of genes associated with oxidative stress responses in S. mutans following AgNPs exposure (Fig. 4D). To further support the observations at the cellular level, oxidative stress-associated signals were examined using CellROX Deep Red staining. As shown in Fig. 5A, cells exposed to AgNPs (2 × MIC and 4 × MIC) exhibited markedly enhanced CellROX fluorescence compared with the untreated control, suggesting elevated oxidative stress-associated changes upon AgNPs exposure. These results indicate that AgNPs treatment is associated with enhanced oxidative stress-related signaling in S. mutans. Consistent with the downregulation of ATP synthase-related genes, intracellular ATP levels were significantly reduced following AgNPs exposure (Fig. 5B). When normalized to the control, the values of ATP content decreased to 34.4%, 7.3%, and 0.37% at 0.25 × MIC, 0.5 × MIC, and 1 × MIC, respectively, indicating severe impairment of cellular energy metabolism. These results directly support the transcriptomic findings indicating the compromised energy metabolism under AgNPs stress. Since glycolysis is the primary source of lactic acid production in S. mutans, extracellular lactate levels were further quantified. As shown in Fig. 5C, AgNPs treated groups displayed significantly lower lactate concentrations at both 4 h and 24 h, demonstrating suppressed acidogenic activity and reduced glycolytic output. This result agrees with the negative enrichment of glycolysis/gluconeogenesis pathways and decreased expression of representative glycolytic genes observed in the transcriptomic analysis (Fig. 4D).

Fig. 5.

(A) Representative fluorescence microscopy images of intracellular oxidative stress-associated staining after 4 h exposure to AgNPs (2 × MIC and 4 × MIC). Red, CellROX fluorescence; blue, DAPI-stained cells (scale bar 10 μm). (B) Intracellular ATP levels measured after 4 h exposure to AgNPs using a luciferase-based assay and expressed as relative ATP levels normalized to the untreated control. Data represent mean ± SD (n = 3). (C) Extracellular lactate concentrations measured after 4 h and 24 h exposure to AgNPs. Data represent mean ± SD (n = 3).

Collectively, the transcriptomic and functional analyses provide convergent evidence that AgNPs exposure disrupts multiple core physiological systems in S. mutans. Pathway enrichment and gene-level profiling revealed coordinated suppression of carbohydrate metabolism, ATP generation, and translational machinery, accompanied by perturbation of membrane and cell wall homeostasis and activation of oxidative stress defense responses. Consistently, functional assays demonstrated reduced ATP production, decreased lactate output, and elevated intracellular oxidative stress, confirming these transcriptional alterations at the physiological level. Importantly, these alterations primarily reflect system-level stress responses under relatively strong antibacterial pressure (2 × MIC and 4 × MIC). Given the used concentrations of AgNPs for transcriptomic profiling were higher than MIC, the observed transcriptional changes likely represent global adaptive stress responses rather than specific molecular effects. Together, these findings support a multi-factorial and pleiotropic mode-of-action of AgNPs under defined in vitro conditions, characterized by coordinated disruption of central metabolism, protein synthesis, cell envelope integrity and redox homeostasis. Combined with the known antibacterial mechanisms of AgNPs (including membrane disruption, oxidative stress induction, and interference with energy metabolism and genomic stability), these findings are consistent with the multifaceted inhibitory effects of AgNPs on S. mutans [28,44,45]. It should be noted that transcriptomic analysis in this study was performed on planktonic S. mutans cells to establish a clear baseline for elucidating molecular responses to AgNPs exposure. In contrast, biofilm-associated cells reside in a distinct microenvironment that can give rise to different gene expression profiles. Future studies incorporating biofilm-stage transcriptomic analyses will be valuable for comparing the differences in the responses of S. mutans to AgNPs under planktonic and biofilm conditions.

Biofilm formation is a key virulence trait of S. mutans, conferring enhanced resistance to host defenses and antimicrobials while driving enamel demineralization through robust acid production. This resilience poses a major challenge in caries management. Our transcriptomic analysis revealed that AgNPs exposure downregulated the mutanobactin biosynthetic gene cluster and perturbed central carbon metabolism, suggesting an impairment in EPS synthesis and biofilm capacity. Guided by these findings, we evaluated the antibiofilm effect of AgNPs under sucrose-supplemented conditions to mimic the cariogenic oral environment. AgNPs inhibited S. mutans biofilm formation in a dose-dependent manner. Crystal violet staining showed that low concentrations (0.5-1 μg/mL) had little effect compared with the control, while a significant reduction was observed at 2 μg/mL. At concentrations ≥4 μg/mL, biofilm biomass decreased sharply, and near-complete inhibition was achieved at 16 μg/mL (Fig. 6A). SEM observations supported these findings, revealing dense biofilms in the control group that became progressively looser with increasing AgNPs concentrations. At the concentration of 16 μg/mL, biofilm structure was virtually abolished, with only sparse cells and residual matrix visible (Fig. 6B). Next, we investigated whether silver nanoparticles (AgNPs) could disrupt established biofilms. S. mutans biofilms were allowed to form for 24 h before being treated with an identical concentration range of AgNPs. To ensure biofilm stability during treatment, established biofilms were exposed to AgNPs dispersed in fresh culture medium rather than in a nutrient-free buffer. This approach ensured that the observed results reflected a true antibiofilm effect rather than an artifact of nutrient deprivation, providing more physiological conditions. Biofilm viability and morphology were subsequently assessed by colony-forming unit (CFU) counts and scanning electron microscopy (SEM). CFU counts revealed a significant reduction in viable cells within the biofilms following AgNPs treatment (Fig. 6C). While no significant changes were observed at a concentration of 8 μg/mL compared to the untreated control, treatment with concentrations of 16 μg/mL or higher resulted in a dramatic decrease in biofilm numbers by over an order of magnitude, demonstrating potent bactericidal activity against sessile cells. Notably, the eradication of established biofilms typically requires antimicrobial concentrations higher than the MIC, as the MIC is defined based on planktonic growth inhibition [46]. In addition, the concentration required to eliminate established biofilms was higher than that required to inhibit biofilm formation. Once formed, biofilms are embedded within an EPS matrix that can impede the penetration of nanoparticles and released ions. Biofilm-associated cells often exhibit reduced metabolic activity and include persister-like subpopulations, while pronounced microenvironmental heterogeneity (e.g., gradients of nutrients and oxygen) further contributes to antimicrobial tolerance. Consequently, disrupting mature biofilms generally requires higher antimicrobial concentration than preventing biofilm formation [47,48].

Fig. 6.

The effect of AgNPs on S. mutans biofilm. (A) The effect of AgNPs on S. mutans biofilm formation evaluated using crystal violet (CV) staining. (B) SEM images of biofilms S. mutans formed under various AgNPs concentrations. (C) CFU results of biofilms after treatment with different concentrations of AgNPs. (D) Representative SEM images of S. mutans biofilms after treatment with different concentrations of AgNPs. The white arrows indicate representative regions showing surface irregularities or partial cell deformation. (E) Fluorescent images of S. mutans biofilms treated with different concentrations of AgNPs captured by CLSM (scale bar: 100 μm). Bacterial cells are shown in green and EPS in blue. (F) Evaluation of bacteria and EPS biovolume in S. mutans biofilms. (G) The thickness of bacteria and EPS in biofilms. Different AgNPs concentration ranges were used across panels A-E according to the specific experimental objectives, including biofilm formation inhibition, treatment of established biofilms, and structural visualization.

SEM imaging provided qualitative morphological observations that are consistent with the quantitative biofilm assays (Fig. 6D). In the untreated control group, the biofilm exhibited dense bacterial clusters embedded within a rich EPS matrix, forming a compact three-dimensional structure. In sucrose-supplemented conditions, enhanced EPS production resulted in a more compact and robust biofilm architecture. In contrast, AgNPs treated biofilms showed localized regions with reduced EPS density, accompanied by surface irregularities and partial cell deformation, particularly at higher AgNP concentrations. These morphological features, highlighted by arrows in Fig. 6D, suggest localized surface irregularities and partial deformation of bacterial cells within AgNPs treated biofilms.

To confirm the antibiofilm activity of AgNPs, biofilms were stained with fluorescent markers targeting polysaccharides and nucleic acids and then examined by confocal laser scanning microscopy (Fig. 6E). In the control biofilm, dense green (live bacteria) and blue (EPS) fluorescent signals were evenly distributed, reflecting a mature, dense biofilm structure. However, with increasing AgNPs concentration, both fluorescent signals gradually weakened, and the biofilm structure became thinner, discontinuous, and less dense. To quantitatively assess the extent of bacteria and EPS in the biofilm, the biovolume and thickness of bacterial cells and EPS were measured from confocal z-stack images (Fig. 6F and G). Compared with the untreated control, both bacterial and EPS biomass decreased significantly, and biofilm thickness showed a similar trend, indicating that the three-dimensional biofilm structure collapsed significantly after AgNPs treatment. Fluorescence intensity analysis also revealed that the ratio of bacterial signal intensity relative to EPS was significantly reduced after AgNPs treatment (Fig. S11). To further assess the relative contributions of bacteria and EPS to the total biofilm mass, we calculated the bacterial to EPS ratio based on biomass and thickness. Regarding biovolume, the bacterial/EPS ratio gradually decreased with increasing AgNPs concentration, indicating that bacterial biomass is more sensitive to AgNPs exposure than the EPS matrix (Fig. S12). Our results demonstrate that AgNPs interfere with the structure and function of S. mutans biofilms and possess the potential to diffuse and penetrate biofilm matrices, thereby affecting sessile bacterial cells, although such penetration may be partially hindered by the EPS layer. At present, there is no direct evidence that AgNPs can degrade or physically disrupt preformed EPS networks. Transcriptomic analysis indicated that exposure to AgNPs perturbs bacterial cellular metabolism and translation processes while upregulating defense responses and protein quality-control systems. This disruption of cellular homeostasis likely compromises the ability of S. mutans to maintain biofilm structural stability. By primarily acting on bacterial cells, AgNPs indirectly weaken the synthesis and renewal of extracellular matrix components, thereby impairing the long-term stability and functionality of the biofilm. Collectively, these findings provide a foundation for the development of AgNPs-based strategies as anti-caries and antibiofilm agents. By bridging green nanotechnology with oral microbiology, this work contributes to the rational design of sustainable nanomaterials for caries prevention and biofilm control in oral healthcare applications.

The present study employed a single-species (S. mutans) biofilm model, which does not fully recapitulate the ecological complexity of oral multispecies biofilms. Future studies should therefore extend these findings to polymicrobial oral biofilm models to evaluate the selective antibacterial activity of green-synthesized AgNPs and their potential impact on oral microbial homeostasis. In addition, although reaction conditions were carefully standardized and physicochemical properties were consistent across independent batches, green synthesis inherently involves biologically derived components that may introduce batch-to-batch variability. More detailed molecular characterization of the nanoparticle surface corona would further strengthen reproducibility assessment in future studies. The long-term antibacterial performance of AgNPs is closely associated with silver ion release and diffusion behavior. Systematic characterization of Ag⁺ release kinetics under biologically relevant conditions will therefore be important to better understand sustained antimicrobial efficacy and potential cumulative effects. Although the present study demonstrates the antibacterial and antibiofilm activity of green-synthesized AgNPs against S. mutans, cytotoxicity and biocompatibility evaluations in relevant oral cell types were not included. Previous studies have indicated that the interactions of green-synthesized AgNPs with mammalian cells are strongly dependent on nanoparticle concentration and surface chemistry, underscoring the necessity of comprehensive biosafety assessment prior to dental application [49,50]. Accordingly, future studies should systematically evaluate the effects of AgNPs on oral keratinocytes, gingival fibroblasts, and other host cells. Beyond cellular biocompatibility, the potential effects of AgNPs on dental hard tissues also warrant careful consideration. Prolonged interactions with enamel and dentin may influence surface properties such as microhardness, surface roughness, and mineral balance, as suggested by previous reports [51,52]. Systematic evaluation of enamel and dentin integrity following AgNPs exposure will therefore be necessary to ensure antimicrobial efficacy without compromising tooth structure. Finally, while the present study provides in vitro evidence for the antibacterial and antibiofilm activity of green-synthesized AgNPs against S. mutans, in vivo validation using cariogenic animal models will be required to further evaluate their potential anticaries relevance and biosafety under physiological conditions.

4. Conclusion

We have successfully developed green AgNPs with excellent antibacterial activity against S. mutans. The AgNPs effectively disrupt bacterial biofilms and suppress acidogenicity, while downregulating genes involved in metabolism and translation and upregulating those related to oxidative stress defense and protein quality control systems. Furthermore, the AgNPs disturb bacterial homeostasis and impair EPS turnover, thereby compromising the structural stability of biofilms. This study provides a comprehensive in vitro characterization of the antibacterial and antibiofilm responses of S. mutans to green-synthesized AgNPs. The findings contribute to a system-level understanding of bacterial stress responses under nanoparticle exposure in the context of caries-associated biofilms. Nevertheless, further investigations are warranted to elucidate the regulatory mechanisms of AgNPs against S. mutans and multispecies dental biofilms, as well as to evaluate their efficacy and biosafety in vivo.

CRediT authorship contribution statement

Jian Zhang: Conceptualization, Formal analysis, Investigation, Methodology, Supervision, Validation, Writing – original draft, Writing – review & editing. Priyanka Singh: Data curation, Formal analysis, Investigation, Methodology, Writing – review & editing. Xin Chen: Formal analysis, Investigation, Methodology, Writing – review & editing. Lei Shi: Formal analysis, Investigation, Validation, Writing – review & editing. Zhejian Cao: Formal analysis, Investigation, Methodology, Writing – review & editing. Shadi Rahimi: Investigation, Methodology, Writing – review & editing. Santosh Pandit: Conceptualization, Investigation, Methodology, Supervision, Writing – review & editing. Ivan Mijakovic: Conceptualization, Project administration, Supervision, Writing – review & editing.

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Appendix A. Supplementary data

The following is/are the supplementary data to this article.

Data availability

Data will be made available on request.