The bacterial strains, plasmids, and primers used are listed in Table S1. S. mutans UA159 was grown at 37°C under anaerobic conditions (90% N₂, 5% CO₂, 5% H₂) in BHI broth (Difco, Sparks, MD, USA). For biofilm formation, 1% sucrose was included in the BHI medium. Escherichia coli strains were grown in Luria–Bertani (LB) medium (Difco, Sparks, MD, USA). If required, antibiotics and other supplements were used at the following final concentrations: spectinomycin (Spe), 800 μg/ml; kanamycin, 50–100 μg/ml. The competence stimulating peptide (CSP) was synthesized by Life Invitrogen (Shanghai, China).

Biofilms were grown in a flow cell (model FC91; BioSurface Technologies, Corp., Bozeman, MT, USA) with connective tubing draining the effluent into a container as described previously (Fux et al., 2004). Overnight culture of S. mutans was resuspended in BHI–1% sucrose and a 2 ml inoculum [5 × 10⁵colony forming units (CFUs)/ml] was injected into the inoculation port containing polystyrene (PLS) blocks (VWR Scientific, CA, USA). Bacteria were allowed to attach on PLS blocks for 45 min before a continuous flow of 1 ml/min at 37°C was started. The flowthrough was collected, and the dispersed cells were harvested by centrifugation and resuspended in PBS. The amount of dispersed cells was determined by plate counting and presented as the log (CFU/ml). The remaining adherent biofilm on the PLS blocks was stained with Live/Dead BacLight (Molecular Probes, Invitrogen, Carlsbad, CA, USA) and scanned by confocal laser scanning microscope (LSM 710; Zeiss, Jena, Germany), followed with COMSTAT program analysis (Liu et al., 2012).

Biofilms at different dispersal stages were collected, suspended in lysis buffer (30 mM Tris-HCl, 1 mM EDTA, 20 mg/ml lysozyme, pH 8.0) and incubated at 37°C with gentle agitation for 40 min. Total RNA was extracted using an RNeasy Mini kit (Qiagen, Dusseldorf, Germany). The RNA obtained from each sample was quantified using the NanoDrop ND-2000 and the RNA integrity was assessed using standard denaturing agarose gel electrophoresis. For microarray analysis, the S. mutans Whole Genome Oligo Array (8 × 15K; Agilent Technologies) and the Agilent Array platform were employed. The sample preparation and microarray hybridization were performed based on the manufacturer's standard protocols (http://www.agilent.com). Raw data were processed and normalized using GENESPRING GX software. Microarray data have been deposited at NCBI-GEO (accession no. GSE95377). Genes with a fold change >1.5 and a P < 0.05 were selected for further gene expression pattern discovery. The protein–protein interactions for the differentially expressed genes (DEGs) were analyzed through Cytoscape software (Institute of Systems Biology, Seattle, WA, USA). Functional annotation of the DEGs was performed using gene ontology (GO) analysis.

The whole deoC gene was replaced with a Spe-resistance cassette. Approximately 800 bp upstream and downstream, the arms of deoC were PCR amplified. The upstream homology arm contained XhoI at the 5′ end and HindIII at the 3′ end, and the downstream homology arm contained NdeI at the 5′ end and SpeI at the 3′ end. The homology arms were ligated into pFW5 (Podbielski et al., 1996), flanking the Spe-resistance cassette in the multiple cloning site. The resulting plasmid was used to transform S. mutans with exogenous CSP added, and transformants were screened on BHI plates containing 800 μg/ml Spe. The sequences of mutant transformants were confirmed by PCR and DNA sequencing. The complementation construct containing the deoC ORF with the putative promoter was cloned into the MCSI sites of pFW5. The construct was used to transform the S. mutans deoC mutant. Transformants without the antibiotic-resistance phenotype were screened for complementation. The confirmed complemented strain was named pFW5-deoCcom.

The DNA degradation activity of S. mutans biofilm supernatant was determined using radical enzyme diffusion assays (Macanovic and Lachmann, 1997). In brief, the culture supernatants of S. mutans biofilms (48–60 h, 60–72 h, 72–84 h, and 84–96 h) were collected by centrifugation at 8,000 g for 15 min, and concentrated in polyethyleneglycol 20,000 (Merck). A 10 μl aliquot of the supernatant was added into the well (diameter = 3 mm) of an agarose plate containing 2 mg/ml salmon sperm DNA, 0.1% Gold View I, 1 mM MgCl₂, and 1 mM CaCl_2.After overnight incubation, 0.05 M EDTA was added to the plate to stop the reaction. The plate was photographed with a UV gel imaging system (FluorChem Q; ALPHA, USA) and the diameters of the circles of hydrolyzed DNA were analyzed with Image-Pro Plus 6.0.

Biofilm dispersal of S. mutans strains was examined as described elsewhere (Kaplan and Fine, 2002). The method employed the microcurrents caused by edge evaporation in an open 100 mm culture dish. S. mutans strains (10³ CFU/ml) were incubated overnight at 37°C under 85% humidity and 5% CO₂ for 24 h in the dish. The medium was discarded, and the plate was stained with crystal violet, followed with washing with distilled water and observation by microscopy.

The S. mutans deoC gene was amplified from genomic DNA using PCR (TaKaRa) with appropriate primers (Table S1) and cloned into the pET28a expression vector to produce recombinant vector pET-deoC. The vector was used to transform E. coli BL21 (DE3) cells. The E. coli cells were grown in LB medium containing 100 μg/ml kanamycin at 37°C for 1 h, followed by plating on an agar plate with 50 μg/ml kanamycin. A single colony was inoculated into LB medium and cultured at 16°C after the addition of 0.3 mM (final concentration) isopropyl-β-D-thiogalactopyranoside to induce protein expression. The overnight culture was collected for recombinant DeoC protein purification as described previously (Gerner et al., 2016). In brief, the cell pellets were resuspended in lysis buffer (20 mM Tris, 500 mMNaCl, pH 8.0), lysed using an Ultrasonic Cell Disruptor (VCX130, 3 mm; Sonics, USA) for 120 × 4 s and cellular debris was removed by centrifugation at 12,000 g for 25 min at 4°C. The recombinant protein was captured on a Ni-IMAC column with loading buffer I (20 mM Tris, 500 mMNaCl, pH 8.0), followed by extensive washing with more than 20 column volumes of loading buffer with 200 mM imidazole. Eluted fractions were collected and separated by SDS-PAGE. Purified proteins were examined by SDS-PAGE and protein concentration was determined using RC and DC Protein Assay (Bio-Rad, CA, USA) by measuring spectrophotometric absorbance at 280 nm.

Enzyme activity was determined by cleavage of 2-deoxyribose-5-phosphate (DR5P). The reaction mixture contained 1 mM DR5P, 0.5 mM NADH, and 10 μl diluted recombinant protein in a 200 μl total volume (You et al., 2012). The catalyzing assay was conducted at different pH (3.0–10.0) and temperature (10°–70°C) values to analyze the effects of pH and temperature on purified protein activities. The decrease in the NADH level was monitored at 340 nm.

The biofilm dispersal ability of recombinant protein was determined as described previously (Minh Tran et al., 2016). S. mutans biofilm was preformed in a 24-well plate and spent medium was aspirated, followed with washing once with PBS. Different concentrations of purified protein were added to the biofilm and plates were incubated at 37°C for 1 h. The remaining biofilm was stained with 0.01% crystal violet, washed twice with PBS buffer, and imaged with a Leica light microscope (Leica Biosystems, Buffalo Grove, IL, USA). To quantify the amount of remaining biofilm, 95% ethanol was added into the wells and the absorbance at 590 nm was determined after 20 min incubation (Minh Tran et al., 2016). Different concentrations of DNase I (Sigma, St Louis, MO, USA) were used as a positive control.

DNA degradation activity of S. mutans strain supernatants and the purified protein was determined as described previously (Berends et al., 2010). Briefly, the overnight culture supernatant of S. mutans strains was collected by centrifugation at 8,000 g for 15 min, and concentrated in polyethyleneglycol 20,000 (Merck). A 5 μl aliquot of the supernatant or the purified protein was incubated with salmon sperm DNA for 1 h at 37°C. The nuclease reaction was stopped with 12.5 μl 0.33 M EDTA (pH 8.0) and the sample was run in a 1% agarose gel for visual examination of DNA degradation.

Human venous blood was obtained from healthy volunteers according to the recommendations of the local ethical committee (ERC-2015-14) and written informed consent was provided by all the participants. Neutrophils were purified by centrifugation (1,800 g) at room temperature for 30 min with BD Vacutainer CPT™ tubes with sodium citrate, according to the manufacturer's instructions. After centrifugation, the polyester gel was removed and cells in the granulocyte layer were collected, followed by erythrocyte lysis, washing with PBS twice, and resuspension in RPMI 1640 with 10% FBS. The purity of the obtained cells was verified by staining and observation with a microscope.

The neutrophil cell suspension in RPMI 1640 with 10% FBS was seeded on coverslides (13 mm) coated with poly-L-lysine (Sigma) in a 24-well plate (3 × 10⁵ cells per well) and incubated for 30 min to allow the cells to attach to the coverslides. To chemically induce NET formation, cells were treated with 100 nM phorbol 12-myristate 13-acetate (PMA; Sigma) for 4 h. Untreated or PMA-treated neutrophils were exposed to S. mutans UA159 wild type, deoC mutant, or pFW5-deoCcom infection at a multiplicity of infection of 200 (MOI = 200) for 4 h. The effect of commercial DNaseI on the NET formation was also investigated as a control. Cells on the coverslides were fixed with 4% paraformaldehyde for 10 min, followed by treatment with 1% Triton X-100 for 10 min and 5% bovine serum albumin (Sigma) block for 1 h. The cells were then incubated with the primary antibody anti-neutrophil elastase (anti-NE; abcam68672) and a secondary antibody conjugated to HRP (abcam6721) to visualize the NE, and the DNA backbone of NETs was detected with DAPI. Specimens were mounted in anti-fade fluorescence medium (Life Invitrogen) and observed with an oil immersion objective fluorescent microscope (Olympus BX63).

ROS production of human neutrophils was measured by a luminometric assay as described previously (Seper et al., 2013). Briefly, neutrophils (3 × 10⁴cells per well) were seeded in a black 96-well plate, followed with addition of 10 μM2, 7-dichlorodihydrofluorescein diacetate. PMA (100 nM) and the S. mutans strain (MOI = 200) were added to stimulate the formation of NETs, and untreated neutrophils were used as a control. Chemiluminescence was determined every 10 min for 2 h in a POLARstar Omega microplate reader (BMG Labtech, Ortenberg, Germany). The ROS production was shown as the area under the curve or as relative light units (RLU).

NET formation was measured by fluorescence assay as described previously (Juneau et al., 2015). Briefly, neutrophils (3 × 10⁴cells per well) were seeded in a black 96-well plate and cell-impermeable Sytox Green DNA-binding dye (5 μM) was added. Cells were stimulated either with 100 nM PMA or S. mutans strains (MOI = 200), or left untreated, and cells were lysed with 1% Triton X-100 as a 100% control. Fluorescence was monitored every 30 min for a period of 9 h in a POLARstar Omega microplate reader (BMG Labtech, Ortenberg, Germany). As a control for DNA degradation, DNase I (100 U/ml) was added after 5 h to PMA- or S. mutans strain-stimulated neutrophils. NET formation was normalized to total neutrophil DNA obtained by Triton X-100 lysis and presented as percentage of DNA fluorescence compared with the 100% lysis control.

Neutrophils (3 × 10⁴ per well) were seeded in a 24-well plate and stimulated with 100 nM PMA (37°C, 5% CO₂) for 4 h to induce NET formation. Cells were divided into three groups, treated with 10 μg/ml cytochalasin D (Sigma) to inhibit phagocytic killing, cytochalasin D, and DNase I (100 U/ml), and the same amount of RPMI 1640 medium as a control, for 15 min. The S. mutans strains were added with an MOI of 200, followed by centrifugation at 800 g for 10 min to get the bacteria in contact with the NETs formed (Berends et al., 2010; Seper et al., 2013). After incubation for 4 h, DNase I was added to the wells not having received DNase I for 20 min to release the NET-entrapped bacteria. Well contents were serially diluted and plated for CFU enumeration. Bacterial survival was expressed as a percentage of the initial inoculum per well (Juneau et al., 2015). To visualize S. mutans entrapment, the co-cultured neutrophils and S. mutans strains on PLS blocks were fixed with 2.5% glutaraldehyde, followed with dehydration in ascending concentrations of ethanol and drying to the critical point. The samples were sputter coated with gold-palladium and observed by scanning electron microscopy (Tran et al., 2016).

To measure gene expression of deoC in S. mutans when entrapped in NETs, S. mutans was collected for RNA extraction and cDNA synthesis was performed with a SuperScript cDNA Synthesis kit (Invitrogen) using 200 ng total RNA. Quantitative real-timePCR (qRT-PCR) was used for quantification of deoC expression with 16S rRNA as the internal control. The reaction program was 95°C for 2 min, followed by 40 cycles of 95°C 15 s and 60°C for 30 s. Threshold cycle (CT) values were determined and data were analyzed with the 2^−ΔΔCT method. S. mutans wild-type strain incubated with salmon sperm DNA (Sigma) was used as the positive control and S. mutans incubated without neutrophils or DNA served as the internal control.

Data analyses were performed using SPSS 13.0, and differences among groups were analyzed by ANOVA, followed by Tukey's multiple comparison tests. Results were shown as mean values ± standard deviation and considered statistically significant at P < 0.05.

The biofilm released bacterial cells during the examined time periods and the amount of bacterial cells was determined every 12 h. As shown in Figure 1A, during the time periods of 48–60 h, more cells were dispersed from biofilm compared with the other time periods (P < 0.05), representing a crucial stage for biofilm dispersal. The biofilm dispersal became steady, with no significant difference in the amount of dispersed cells observed, between 60–72 h and 72–84 h (P > 0.05). During 84–96 h, the amount of dispersed cells decreased significantly (P < 0.05). Corresponding to the dispersal, the biomass of biofilm decreased gradually and reached a steady state after 84 h (Figure 1B).

We used whole-genome microarrays to identify the global transcriptional changes during biofilm dispersal and also to investigate the molecular mechanism regulating biofilm dispersal. Based on the data shown in Figure 1, 48–60 h represented a rapid biofilm dispersal stage and at 96 h the dispersal greatly decreased. Therefore, the transcriptome of 96 vs. 60 h biofilm and 60 vs. 48 h biofilm was compared. A total of 160 genes were identified as differentially expressed, with 26 genes up-regulated (Table S2) and 134 genes down-regulated in the 96 vs. 60 h biofilm group (Table S3). In the 60 vs. 48 h group, 137 genes (30 up-regulated and 107 down-regulated) were differentially expressed, as shown in Tables S4, S5.

Since gene interaction networks are important for various biological processes, we analyzed protein interactions among the identified DEGs. In the 96 vs. 60 h group, the DEGs were clustered in 6 interaction networks (Figure 2). Four of the interaction networks were also predicted in the 60 vs. 48 h biofilm group (Figure 3). It seemed that the DEGs formed a highly linked network, which indicated that the genes in the identified interaction networks might be associated with biofilm dispersal.

GO enrichment analysis was then carried out to identify pathways that may regulate S. mutans biofilm dispersal based on the identified interaction networks. In total, 37 GO terms were assigned in the 96 vs. 60 h group, among which the DEGs with GO term translational elongation and with GO term endonuclease activity reached 36.6 and 30%, respectively (Figure 2). Among the 31 GO terms in the 60 vs. 48 h group, the DEGs in GO term deoxyribonuclease activity and endodeoxyribonuclease activity reached 30 and 37.5%, respectively (Figure 3). The overlap of DEGs in GO terms deoxyribonuclease activity and endodeoxyribonuclease activity indicated that nuclease activity might be critical to biofilm dispersal.

To validate the microarray data, we randomly selected 20 genes for expression analysis by qRT-PCR using 16S rRNA as the internal control. As shown in Figure S2, most of the selected genes showed a similar expression trend, which supported a strong level of confidence in our microarray data.

To study whether deoC played a potential role in biofilm dispersal, we examined the DNase activity of S. mutans biofilm supernatants at different stages. As shown in Figure 4A, the supernatant of the 48–60 h period exhibited the highest level of DNase activity characterized by the larger diameter compared with the 72–84 h and 84–96 h periods (P < 0.05), which was in accordance with the dispersal extent of biofilm in Figure 1. We thus speculated that this protein may initiate the biofilm dispersal. To verify this speculation, we constructed a deoC deletion mutant and compared the dispersal ability of the mutant with the wild-type strain through the biofilm dispersal experiment. The deoC deletion mutant was verified by PCR and sequencing (Figure S1). In the dispersal assay, microturbulence in the plates causes a comet-like tail to form as organisms disperse from the nascent microcolony. The dispersal was noted by the comet tail of satellite colonies formed by S. mutans wild type (Figure 4B). However, only a few of the microcolonies exhibited a similar comet tail in the plates containing the deoC mutant. These data strongly suggested that DeoC was involved in the biofilm dispersal of S. mutans.

To confirm DeoC activity, we expressed and purified the DeoC His-tagged recombinant protein (Figure 5A). In order to determine the optimal conditions for the catalytic activity, we quantitatively assayed the effects of pH and temperature on purified DeoC activities. The results showed that purified DeoC was capable of catabolizing DR5P and was optimally active at pH 6.0 (Figure 5B) and 50°C (Figure 5C), but could not degrade DNA independently (Figure 5D). The DNA degradation activity of culture supernatants of S. mutans was also investigated via degradation of salmon sperm DNA. The deoC mutant supernatant demonstrated no DNA degradation activity, indicating that DeoC played an indispensable role in nuclease activity of S. mutans. In contrast, supernatants from the wild-type strain and DNaseI efficiently degraded DNA marker (Figure 5D). Degradation of DNA could be restored in the complemented strain; thus, excluding polar effects of mutagenesis.

To determine whether DeoC could disperse preformed biofilm, we treated biofilm with increasing concentrations of purified DeoC, and commercial DNaseI was used as a positive control. The biofilm was progressively disrupted as the DeoC concentration increased, and a significant decrease in biofilm biomass was observed when the DeoC concentration increased to 1μg/ml (P < 0.05) compared with the untreated control (Figures 5E,F).

The oxidative burst is a key feature for neutrophils, which is involved in signaling processes and microbial killing. Thus, we analyzed the ROS production of neutrophils upon contact with S. mutans wild type, deoC mutant, and the complemented strain. As displayed in Figure 6A, the ROS production reached amaximum at approximately 2 h (MOI = 200) in all the tested strains, which was indicative that S. mutans was recognized by human neutrophils and had the potential to induce NET formation. ROS production during the observed time periods was shown by measuring the area under the curve. No significant difference was observed in the ROS levels between the wildtype, deoC mutant, and the complemented strain (Figure 6B), which indicated that the neutrophil activation potential of S. mutans was independent of DeoC.

NET formation was quantified using the fluorescent dye SytoxGreen, which detects extracellular DNA. In the case of neutrophils stimulated with S. mutans strains, the NET level increased over time with a lag-phase of 3 h, while only 1 h lag phase was observed for PMA treated neutrophils (Figure 6C). After 6 h of incubation, the fluorescence increased over time and reached a plateau at 47.54 and 62.24% of the total DNA when the neutrophils were stimulated with deoC mutant and PMA, respectively (Figures 6C,D). Notably, the fluorescence continued to increase after 6 h in the mutant stimulated neutrophils and kept steady after 9.5 h (Figure 6C). The sharp increase in DNA fluorescence from the 7th hour might be attributed to the death of the mutant cells and the subsequent release of DNA, since only 10% of the mutant cells survived after 4 h contact with NETs as revealed later in Figure 8B. However, wild-type-stimulated and complemented-strain-stimulated neutrophils exhibited only a slight increase in fluorescent signal. To confirm that the low fluorescence in the wild-type-stimulated neutrophils was due to nuclease activity, DNaseI was added to the PMA-treated neutrophils as a control. As expected, the fluorescent intensity decreased rapidly upon DNaseI addition (Figures 6D,E).

To visualize NET formation induced by S. mutans, human neutrophils were stimulated with PMA, wild type, deoC mutant, and pFW5-deoCcom. As a positive control, PMA-stimulated neutrophils exhibited web-like structures and the elastase was destroyed and scattered, compared with the intact neutrophils with nuclei and elastase within the cytoplasm in the unstimulated intact neutrophils (Figure 7A). After DNaseI treatment for 4 h, only deformed nuclei and scattered elastase could be detected, without DNA web-like structures. When co-cultured with the wild-type S. mutans or the complemented strain, no intact NETs were observed, characterized by the low intensity of DNA or the absence of DNA in areas of elastase. In addition, large areas of NE surrounded by S. mutans in an intact chain were visible (Figure 7B), suggestive of degradation of DNA in NETs by S. mutans. In comparison to the wild type, the deoC-mutant-stimulated neutrophils were associated with web-like DNA structures and dispersed elastase could be observed (Figure 7A), indicative of the incapability of the deoC mutant to degrade the DNA of the NETs, especially in comparison to the intact chain structure of wild type, the deoC mutant was found to be scattered in the DNA web (Figure 7B).

Scanning electron microscope visualization of neutrophils revealed web-like structures trapping S. mutans wild-type cells after 2 h of co-culture, and the web-like structure disappeared after 4 h. In contrast, the deoC mutant got entrapped within NETs and no degradation of NETs was noted (Figure 8A). Due to antimicrobial effectors present at high concentrations in NETs, it was hypothesized that DeoC enhanced S. mutans survival in the presence of neutrophils. To test this hypothesis, a plate-based NET killing assay was performed. As shown in Figure 8B, a lower survival rate of deoC mutant was observed after incubation with PMA-stimulated neutrophils compared with the wild type and complemented strains (P < 0.05). In the presence of cytochalasin D or cytochalasinD+DNaseI, survival of all the S. mutans strains was increased (P < 0.05) in comparison with the PMA-treated neutrophils, suggestive of the role of phagocytic killing in neutrophils. However, a greater increase in survival of the deoC mutant was observed than of the wild type and the complemented strain, indicative that the DeoC activity of S. mutans was critical for the escape from NET-mediated extracellular killing.

To determine whether gene expression of nucleases was induced in the presence of NETs, qRT-PCR was employed to analyze the expression of deoC in S. mutans. It was found that deoC expression was up-regulated five-fold in S. mutans incubated with PMA-treated neutrophils. A similar induction was observed in S. mutans incubated with commercially available salmon sperm DNA (Figure 8C).

The authors declare that the research was conducted in the absence of any commercial or financial relationships that could be construed as a potential conflict of interest.