Abstract
Extracellular DNA (eDNA) is a key component of many microbial biofilms including dental plaque. However, the roles of extracellular deoxyribonuclease (DNase) enzymes within biofilms are poorly understood. Streptococcus gordonii is a pioneer colonizer of dental plaque. Here, we identified and characterised SsnA, a cell wall-associated protein responsible for extracellular DNase activity of S. gordonii. The SsnA-mediated extracellular DNase activity of S. gordonii was suppressed following growth in sugars. SsnA was purified as a recombinant protein and shown to be inactive below pH 6.5. SsnA inhibited biofilm formation by Streptococcus mutans in a pH-dependent manner. Further, SsnA inhibited the growth of oral microcosm biofilms in human saliva. However, inhibition was ameliorated by the addition of sucrose. Together, these data indicate that S. gordonii SsnA plays a key role in interspecies competition within oral biofilms. Acidification of the medium through sugar catabolism could be a strategy for cariogenic species such as S. mutans to prevent SsnA-mediated exclusion from biofilms.
Similar content being viewed by others
Introduction
The oral cavity typically harbours between approximately 100 and 300 different bacterial taxa that colonise the surfaces of hard and soft tissues1. The taxonomic composition and distribution of bacteria within oral biofilms are determined by selective adherence processes, competitive and mutualistic interactions between species and the environment provided by the host2,3,4,5. The accumulation of dental plaque, the biofilm formed on the surfaces of teeth, occurs in a spatiotemporally ordered manner6. Pioneer colonisers capable of adhering to the acquired enamel pellicle lay the foundation of dental plaque and provide receptors for the recruitment of secondary colonisers, which eventually leads to the development of complex and dynamic microbial communities7,8. Pioneer colonizers are generally considered commensals or even beneficial for maintaining oral health2,9,10. However, if dental plaque is not properly controlled, dental caries or periodontitis can develop11. These two conditions are responsible for the vast majority of tooth loss worldwide12. Disease is associated with shifts in the composition and biochemical activities of dental plaque that result in a state of dysbiosis13,14. In the case of dental caries, excess frequency and/or amount of dietary sugars intake selects for dysbiotic biofilms enriched in aciduric and acidogenic bacteria that reduce the pH at the tooth surface and drive demineralization of tooth enamel or cementum. Further progression of the lesion results in destruction of the dentin and invasion of bacteria into the pulp chamber15,16.
Pioneer colonisers such as Mitis-group streptococci play important roles in controlling the overgrowth of more acidogenic and cariogenic bacteria such as Streptococcus mutans. For example, Streptococcus gordonii expresses an arginine deaminase (AD) system that counteracts the acidification of the biofilm by generating ammonia and carbon dioxide whilst producing energy for the cells9,17,18,19,20. In the presence of oxygen, S. gordonii and other members of the Mitis group of streptococci including Streptococcus sanguinis produce substantial (millimolar) amounts of hydrogen peroxide (H2O2), which can inhibit the growth of S. mutans21. In addition, it is possible that the pioneer colonisers compete with S. mutans for binding sites within the biofilm. S. mutans attachment is heavily dependent on the presence of sucrose, which is metabolised extracellularly to produce glucans that support adhesion and biofilm colonisation22. In the absence of sucrose, S. mutans adhesion appears to be mediated by extracellular DNA (eDNA)23. Extracellular DNA is also important for the structural integrity of mature S. mutans biofilms since treatment with DNase I results in biofilm dispersal24,25. It has recently been shown that eDNA is present in more complex biofilms including mixed-species dental plaque26,27,28,29,30,31. Treatment of dental plaque, particularly at the early stages of development, with exogenous DNase enzymes significantly reduces the biofilm biomass and selectively depletes certain species that appear to be particularly dependent upon eDNA27,31,32. In addition, eDNA serves as a nutrient source and reservoir for horizontal gene transfer within dental plaque3,16.
Many bacteria produce surface bound or secreted deoxyribonuclease (DNase) enzymes33,34. In biofilms, extracellular DNases serve a variety of functions including dissemination of bacterial cells, eDNA turnover and horizontal gene transfer8,35,36,37,38. In pathogens, extracellular DNase enzymes are often linked to virulence and evasion of host immune responses such as neutrophil extracellular traps (NETs)39,40. A number of oral bacteria including S. gordonii produce extracellular DNase enzymes33,34,41. However, their functions in biofilm maintenance and interspecies interactions are poorly understood. Here, we identified and characterised the enzyme responsible for extracellular DNase activity of S. gordonii. Furthermore, we investigated the effect of this enzyme on development of S. mutans biofilms or more complex mixed-species biofilms formed by human oral bacteria.
Results
Extracellular DNA in S. gordonii and S. mutans biofilms
To determine the extent of eDNA within the matrix of S. gordonii DL1 and S. mutans GS-5 biofilms, eDNA was extracted from monospecies biofilms grown statically for 48 h. A band at high molecular weight (>10 kbp) was observed in extracts isolated from S. mutans biofilms but not in S. gordonii biofilms (Fig. 1a). The size of the fragment was similar to intracellular chromosomal DNA (iDNA) extracted from S. mutans cells. This band was not observed following enzymatic treatment of the extracts with DNase I (Supplementary Fig. 1). NanoDrop spectrophotometry analysis of the extracts showed eDNA represented approximately 19% of the total DNA (iDNA + eDNA) in S. mutans biofilms while it accounted for only 6% of the total DNA in S. gordonii biofilms. The role of eDNA in maintaining the integrity of S. mutans and S. gordonii biofilms was investigated by treating pre-formed S. mutans and S. gordonii biofilms with 5 µg/mL DNase I for 1 h at 37 °C (Fig. 1b, c). Crystal violet staining revealed that DNase I treatment significantly reduced S. mutans biofilm biomass by >90%. By contrast, S. gordonii biofilms were not affected by DNase I treatment. Inclusion of DNase I during biofilm development significantly inhibited the accumulation of S. mutans biofilms but did not affect S. gordonii (Fig. 1d).
a Agarose gel electrophoresis of intracellular and extracellular DNA (iDNA and eDNA) isolated from S. gordonii and S. mutans biofilms. Biofilms were grown for 48 h in multi-well plates and cells were separated from extracellular matrix by centrifugation prior to DNA extraction. The black arrow indicates eDNA. Marker (M) was run on the same gel as the S. gordonii samples but separated from the sample lanes. S. mutans samples were run on a different gel. b Pre-established biofilms of S. gordonii and S. mutans grown for 20 h in BHY medium were treated with 5 μg/mL DNase I for 1 h at 37 °C. Biofilms were stained with crystal violet and biomass was quantified by dissolving the dye and measuring absorbance at 570 nm. c Quantification of crystal violet stained S. gordonii DL1 and S. mutans GS-5 biofilms grown in the presence and absence of 5 μg/mL DNase I. Biofilms were grown anaerobically for 20 h in BHY at 37 °C. S. mutans biofilms but not S. gordonii were inhibited when grown in the presence of 5 μg/mL DNase I. d Quantification of pre-formed S. mutans biofilms treated with 5 μg/mL DNase I for 1 h at 37 °C showed significant reduction by DNase I treatment (dispersal), whereas S. gordonii biofilms were not reduced. Data points are shown for 4–6 independent experiments, bars represent the mean and SD is indicated. The distribution of data passed the Shapiro-Wilk normality test and therefore statistical significance was calculated using Student’s t-test; ***p < 0.001, ****p < 0.0001.
S. gordonii expresses an extracellular DNase enzyme
We hypothesised that low levels of eDNA in S. gordonii biofilms may be due to the production of an extracellular DNase enzyme. Screening the S. gordonii DL1 genome using NCBI Blast identified three genes that potentially encoded proteins with DNase I-type domains. Of these, genes encoding an exonuclease III enzyme and a protein labelled RgfB did not have characteristics of secreted proteins. The third gene (SGO_RS08090; GenBank Accession WP_012130709.1) encoded a putative 779 amino acid protein with an N-terminal secretion signal and a C-terminal LPxTG motif for sortase-mediated anchoring to the cell wall, indicative of an extracellular cell wall-bound enzyme (Fig. 2a). In view of homology to other streptococcal extracellular nucleases including SsnA of Streptococcus suis42, the putative DNase gene of S. gordonii was termed ssnA (Streptococcal Surface Nuclease A). To test whether ssnA gene was responsible for extracellular DNase activity in S. gordonii, ssnA was knocked out and extracellular DNase activity was assessed using DNase Test agar (Fig. 2b). A zone of clearance was visible around colonies of wild-type S. gordonii indicating that these cells produced extracellular DNase. By contrast, no extracellular DNase activity was detected in the ∆ssnA mutant. Genetic complementation of the ssnA gene under its native promoter (S. gordonii ssnAComp) restored the DNase activity (Fig. 2b).
a Schematic diagram of the domain organisation of the protein encoded by the ssnA gene. Positions of predicted domains including a putative signal peptide (SP), an OB-fold, a nuclease domain and an LPxTG cell wall anchoring motif are marked with start and end amino acid residue numbers from the N-terminus. b Extracellular DNase activity was assessed using DNase Test agar. A clear halo around colonies of wild S. gordonii DL1 indicates DNase activity. No halo was visible in S. gordonii ΔssnA, whereas extracellular DNase activity was restored in S. gordonii ssnAComp. c The cell fraction and spent culture media were separated by centrifugation of planktonic cultures of wild type S. gordonii, the ΔssnA mutant and ssnAComp strains, and the nuclease activity of each fraction was determined using a fluorescence based quantitative assay. S. aureus FH7 was included as a control. Extracellular DNase activity of S. gordonii was associated primarily with the cells, whereas S. aureus DNase activity was higher in conditioned medium. Identical volumes of cellular and supernatant fractions were used in analysis. Data points are shown for 3 independent repeats, bars represent the mean and SD is indicated.
To further explore the production of SsnA in S. gordonii, a quantitative fluorescence-based DNase activity assay was employed (Fig. 2c). DNase activity was determined in cell-bound fractions and supernatants. Staphylococcus aureus FH7, which is known to release DNase into the extracellular milieu, was included as a control. In wild-type S. gordonii, DNase activity was approximately 5-fold greater in the cell-associated fraction than spent medium, indicating that the majority of secreted SsnA remains bound to the cell wall. DNase activity was almost undetectable in the cell fraction and spent medium of S. gordonii ΔssnA. Extracellular DNase activity was restored in the complemented strain and was mostly associated with the cellular fraction.
Carbon catabolite repression (CCR) governs expression of SsnA
Initially, the expression and activity of SsnA was monitored through the growth of S. gordonii in BHY (Fig. 3a). No significant difference in cell-associated SsnA activity was detected between 2 h and 25 h (Fig. 3b). Gene expression of ssnA relative to expression of the 16 S rRNA gene was also stable during the exponential growth phase. However, expression decreased during stationary phase and was significantly reduced by >30-fold after 25 h (Fig. 3c).
a Growth curve of S. gordonii grown aerobically in BHY at 37 °C. b DNase activity of cells measured at 2, 4, 6, and 25 h using the quantitative fluorescent assay are shown. Optical density (OD600 nm) measurements were used for normalization. c RNA was extracted from the samples at different times through growth to evaluate ssnA expression using RT-qPCR. Fold change was normalised to 16 S rRNA gene expression. The result of 3 independent experiments and SE are shown for each assay. The distribution of data passed the Shapiro-Wilk normality test. Ordinary one-way ANOVA and Dunnett’s multiple comparisons were performed using GraphPad Prism 9 to determine statistical significance; ****p < 0.0001.
Sequence analysis of the promoter region of ssnA revealed a consensus binding site for carbon catabolite protein A (CcpA), or possibly for the maltose regulator MalR protein (Supplementary Fig. 2a). To determine whether CcpA or MalR play a role in regulation of SsnA expression, ∆ccpA and ∆malR mutants were constructed and extracellular DNase activity was assessed in strains grown in the presence or absence of added sugars (Supplementary Fig. 2b). In Gram-positive bacteria such as S. gordonii, CCR is linked to uptake of sugars through the phosphoenolpyruvate-dependent sugar:phosphotransferase system (PTS). Substrate specificity is determined by the enzyme II (EII) component of the PTS. The S. gordonii EIIMan complex appears to have broad substrate specificity43 and we were unable to identify a sugar that is utilised by S. gordonii and that does not trigger CCR (Lin Zeng, personal communication). Purified SsnA was included as a control and degraded DNA in all conditions tested, as indicated by a zone of clearance on the agar. In wild type S. gordonii, DNA degradation was observed in the absence of added sugars or in galactose, but not in glucose, maltose or sucrose. Interestingly, DNase activity was observed in the ∆ccpA mutant grown in the presence or absence of any of the sugars tested (Supplementary Fig. 2). A genetically complemented strain, S. gordonii ccpAComp, exhibited a similar pattern of DNase activity to the wild type. Deletion of malR did not affect DNase activity suggesting that this repressor is not involved in the regulation of ssnA expression under conditions tested here.
Reduced pH from sugar fermentation is associated with inhibition of SsnA
To quantify the extent of the sugar inhibition on SsnA activity, DNase activity of cells grown in BHY medium or BHY supplemented with different sugars was measured using a fluorescence-based assay (Fig. 4a). S. gordonii DNase activity was reduced by >50% in the presence of glucose, maltose, galactose, fructose or inulin. Streptococci produce acid from sugars and it was possible that the DNase enzyme activity may have been affected by the pH of the cultures. The pH of supernatants from stationary phase S. gordonii BHY cultures was approximately 5.6, whereas the pH of cultures grown with galactose was approximately 4.6 (Fig. 4b). The pH was further reduced to between 4 and 4.2 in cultures grown with glucose, maltose, fructose or inulin. To assess the impact of low pH on DNase activity, BHY was amended to pH 5.5 by the addition of HCl and used to culture S. gordonii cells to stationary phase. The final pH following growth was approximately 4.2 and almost no DNase activity was detected in cells grown in acidified BHY (Fig. 4).
a Extracellular nuclease activity of S. gordonii DL1 grown in BHY or BHY + 2% (w/v) sugars: glucose (BHYG), maltose (BHYM), galactose (BHYGal), fructose (BHYF), or inulin (BHYI). Extracellular DNase activity of S. gordonii cells grown in acidified medium BHY/HCl (BHY amended to pH 5.5 with HCl) is shown separated by a red line. Data points are shown for 3 independent repeats, bars represent the mean and SD is indicated. b The pH of cultures following growth to stationary phase in BHY, BHY + sugars or in BHY/HCl. c S. gordonii DL1 overnight cultures were harvested and re-suspended in buffer at pH 7.1 (blue line), 5.5 (red line) or 4.5 (magenta line). After 2 h (red arrow), cells were harvested and resuspended in PBS (pH 7.1) or PBS (pH 7.1) supplemented with chloramphenicol (Chlm; 12.5 μg/mL; light and dark green lines). DNase activity is shown for cell-associated SsnA. Data points are shown for 3 independent repeats, bars represent the mean and SD is indicated. Optical density (OD600 nm) measurements were used for normalization in each assay.
We hypothesised that the effects of low pH on SsnA activity were not directly due to enzyme inhibition since the assay was performed after washing cells and resuspending in PBS at pH 7.1. To further explore pH-mediated reduction of SsnA activity, S. gordonii DL1 was incubated in neutral or acidic buffers and transferred to pH 7.1 in the presence or absence of the protein synthesis inhibitor chloramphenicol (Fig. 4c). The total DNase activity was lower in cells incubated at pH 4.5 or 5.5 than in cells incubated at pH 7.1. Following a shift to pH 7.1, SsnA activity increased in cells that had been incubated in pH 4.5 buffer. The recovery of SsnA activity was impaired in the presence of chloramphenicol indicating that de novo protein synthesis was required to restore enzyme activity. In cells that had been incubated for 2 h at pH 5.5, SsnA activity increased slightly, but not significantly, following a shift to pH 7.1. In the presence of chloramphenicol, SsnA activity continued to decrease after shifting to pH 7.1 indicating that active enzyme was not being restored.
pH and metal dependence of purified SsnA
To characterise the enzyme activity in more detail, SsnA was purified as a GST-tagged construct. The identity of the purified construct was confirmed by peptide mass fingerprinting (Supplementary Fig. 3). In preliminary experiments, the GST tag was cleaved with thrombin and removed by affinity purification. The enzyme was shown to be active with or without the GST tag (Supplementary Fig. 4). Since removal of the tag resulted in a significant loss of yield of the protein, we used the GST-tagged construct for subsequent experiments. Initially, the effects of pH on DNase activity were investigated (Fig. 5a). The optimum pH was between 7 and 9.5. No activity was detectable below pH 6.5 or above pH 10.5.
a The nuclease activity of purified SsnA was quantified at pH ranging from 4.5 to 11. b Using the quantitative fluorescent assay, the DNase activity of SsnA was tested in the presence of a range of Mn2+, Ca2+, Mg2+ and Zn2+ concentrations. Negative controls without enzyme are indicated. In all panels, points or bars indicate means from 3 independent experiments and SE is shown.
Nucleases are frequently dependent on a divalent metal cofactor44. Inductively coupled plasma mass spectrometry (ICP-MS) analysis of SsnA did not detect any metal ions associated with the purified protein. To determine the preferred metal co-factors for SsnA, the enzymatic activity of SsnA was tested in the presence of Mn2+, Ca2+, Mg2+ or Zn2+ (Fig. 5b). In the absence of added metals, no DNase activity was detected in SsnA preparations. Addition of 10 µM MnCl2, 20 µM CaCl2, 40 µM MgCl2, or 40 µM ZnSO4 resulted in high levels of DNase activity. Increasing the concentration of ZnSO4 to 80 µM led to almost complete inactivation of DNase. DNase activity was also abrogated in high concentrations of MnCl2 (>1.28 mM MnCl2).
SsnA disperses S. mutans biofilms under neutral but not acidic conditions
As shown above, S. mutans GS-5 biofilms were sensitive to treatment with DNase I whereas S. gordonii biofilms were not. We hypothesised that SsnA would also disperse pre-formed S. mutans biofilms. Using confocal laser scanning microscopy, S. mutans biofilms were almost completely eliminated by treatment with SsnA (Fig. 6a). Inhibition was found to be dependent on the prevailing pH (Fig. 6b). At pH 4 or 5, biofilms were not removed by SsnA. By contrast, S. mutans biofilms were reduced to <40% of the control biomass (without enzyme treatment) by DNase I at pH 5. At pH 6, both DNase I and SsnA were effective at reducing S. mutans biofilms and biomass was reduced to <40% of the controls. Biofilm removal was even greater at pH 7, and only 10–20% of the biofilm remained following treatment with DNase I or SsnA. Based on these findings we hypothesised that S. gordonii wild-type cells, which express SsnA, could disperse S. mutans GS-5 biofilms. To test this hypothesis, pre-established S. mutans biofilms grown for 20 h were treated with S. gordonii wild-type or ΔssnA cells or with spent culture medium (Supplementary Fig. 5). Treatment with wild-type S. gordonii cells or culture supernatant significantly reduced the S. mutans biofilm biomass to approximately 60% of the control. By contrast, treatment with S. gordonii ΔssnA cells did not reduce S. mutans biofilms. A small, but significant reduction in S. mutans biofilms was observed following treatment with S. gordonii ΔssnA culture supernatant (Supplementary Fig. 5). Overall, these data show that SsnA promotes the removal of S. mutans biofilms by S. gordonii.
a Confocal laser scanning microscopy of pre-established 20 h biofilms of S. mutans treated with PBS (control) or 5 µg/mL SsnA for 1 h at 37 °C. Propidium iodide (20 μM) was used to visualise the cells. Scale bars are 30 µm (left panels) or 20 µm (right panels). b Biomass of pre-established S. mutans biofilms treated with DNase I (5 µg/mL) or SsnA (5 µg/mL) at pH 4, 5, 6, and 7. Crystal violet assay was used for quantification. Data points are shown for 3 independent repeats, bars represent the mean and SD is indicated.
SsnA inhibits the accumulation of oral microcosm biofilms
To determine whether SsnA can control the development of more complex oral biofilms, we employed a dual inlet BioFlux microfluidic system to grow oral microcosm biofilms in the presence or absence of SsnA under the flow of natural saliva as the sole nutrient source. The system was inoculated with unsterilised saliva to seed the channels. Biofilms were cultured for 20 h and images were taken every 10 min (Fig. 7a; Supplementary Movie 1). A clear inhibition of oral microcosm development was observed over the 20 h of growth in the half of the channel in which SsnA was included. Staining of extracellular DNA (eDNA) with Yoyo-1 showed that eDNA was also markedly reduced in the presence of SsnA. The fluorescence signal was quantified over the course of biofilm formation (Fig. 7b). A consistent increase in eDNA levels was observed over 20 h of biofilm development in the top half of the channel (control) whilst in the lower half (SsnA-supplemented), levels of eDNA remained stable throughout the experiment.
a Visualisation of oral microcosm biofilms developed in the channel of BioFlux dual inlet microfluidic system with saliva containing SsnA (5 µg/mL) feeding the bottom half of the channel. Microcosm biofilms were grown for 20 h. Yoyo-1 (2.4 nM) was used to stain eDNA within the biofilm. b Quantification of eDNA accumulation for the SsnA-containing and SsnA-free section of the growth channel over 20 h. Images were taken every 10 min and eDNA amounts were determined using GrayValue within FIJI software. c Visualisation of microcosm biofilms grown in 0.3% sucrose-supplemented natural saliva. SsnA (5 µg/mL) was included in the reservoir feeding the bottom half of the channel. Yoyo-1 (2.4 nM) was included for eDNA staining. d Quantification of eDNA accumulation for the SsnA-containing and SsnA-free section of the growth channel over 20 h of growth in saliva supplemented with 0.3% sucrose. Images were taken every 10 min and eDNA amounts were determined using GrayValue within FIJI software. Mean and SE from 3 independent repeats are shown. Scale bars are 50 µm.
Sucrose fermentation blocks eDNA degradation by SsnA in oral microcosm biofilms
Previous work had shown that sugars lead to a reduction in pH and suppression of SsnA enzyme activity (Fig. 4). To assess the effects of sugar on the control of oral microcosm biofilms by SsnA, biofilms were cultured in the BioFlux system in saliva supplemented with 0.3% sucrose (Fig. 7c; Supplementary Movie 2). Under these conditions, no reduction in biofilm accumulation was observed in the presence of SsnA (Fig. 7c, d). To further explore the impact of sucrose on eDNA in oral microcosm biofilms, Bacillus licheniformis extracellular nuclease NucB was employed since it has previously been shown to control the development of oral microcosm biofilms under static conditions31. In the presence of NucB, biofilm development in saliva was strongly inhibited compared with controls (Fig. 8a, b; Supplementary Movie 3). Similar inhibition was observed in cultures grown in saliva supplemented with 0.3% sucrose (Fig. 8c, d; Supplementary Movie 4). Taken together, the above data indicate that eDNA is important for the development of oral microcosm biofilms and that SsnA is not effective at controlling eDNA-dependent biofilm accumulation in the presence of sucrose.
a Oral microcosm biofilms grown for 20 h in natural saliva and supplemented with NucB in the lower of the microfluidic channel. Yoyo-1 (2.4 nM) dye was used for eDNA visualisation. b eDNA quantified in the two halves of the growth channel over 20 h of growth. eDNA was measured through GrayValue determination in FIJI software for images taken every 10 min. c Sucrose (0.3%) supplemented saliva was used to grow microcosm biofilms and NucB was included in the lower half of the growth channel. d eDNA quantified for the NucB-containing and NucB-free halves of the growth channel over 20 h of growth in sucrose (0.3%) containing saliva. Mean and SE from 3 independent repeats are shown. Scale bars are 50 µm.
Discussion
Here, we identified and characterized SsnA, an extracellular cell wall-associated DNase enzyme produced by S. gordonii. Extracellular DNase enzymes are widely produced by pathogenic species of streptococci and are important virulence factors39,45,46,47. A number of oral commensal streptococci including S. sanguinis, Streptococcus intermedius, Streptococcus oralis, Streptococcus parasanguinis and Streptococcus mitis produce detectable levels of extracellular DNase activity34,41. Of these, only the extracellular DNase of S. sanguinis has been characterised. This enzyme, termed SWAN (Streptococcal Wall-Anchored Nuclease), was shown to mediate escape from neutrophil extracellular traps (NETs)41. There is evidence that S. mutans can also escape from NETs using a deoxyribose aldolase DeoC48. However, extracellular nuclease production by S. mutans is weak or undetectable in DNase activity assays34. Our data have identified a role for S. gordonii SsnA in modulating biofilm formation by competing species such as S. mutans. Further, it was shown that the ability of SsnA to mediate interspecies competition within biofilms is dependent upon the prevailing pH and concentration of sugars. Therefore, it appears that streptococcal extracellular DNases may play important roles in maintaining the stability of dental plaque biofilms.
SsnA was identified by searching the S. gordonii genome for genes encoding proteins with DNase I-like domains. The DNase I family includes a diverse range of nucleases and phosphatases that share a conserved protein fold49. Although our analysis suggests that the majority of SsnA is cell-bound, a mass spectrometry study of the whole secretome of S. gordonii revealed the presence of SsnA (SGO_1651) amongst 107 secreted proteins50. Therefore, it is likely that a proportion of the enzyme is released from the cell surface. Homologues of S. gordonii SsnA are present in a wide range of oral streptococci including S. sanguinis, S. intermedius, S. constellatus, S. anginosus and S. parasanguinis (Supplementary Fig. 6), indicating that the enzyme likely plays important roles in oral biofilms. In addition, more distantly related homologues are present in pathogenic streptococci including S. pyogenes and the important animal pathogens S. suis, S. equi and S. iniae (Supplementary Fig. 6).
The SsnA activity was strongly reduced in the presence of sugars. Supplementation with glucose or the glucose-containing disaccharides led to complete inhibition of extracellular DNase activity. Upstream of the ssnA gene is a CRE sequence motif and regulation was abrogated in a ΔccpA mutant. Hence, it is likely that CcpA acts at the level of gene regulation to control the expression of ssnA in the presence of glucose. Unfortunately, the ΔccpA mutant grew slowly in broth cultures and we were unable to obtain reproducible data from RT-qPCR analysis of ssnA expression in this strain. The regulation of extracellular DNase enzymes by CcpA has been observed in other streptococcal species including S. suis and S. pyogenes51,52. In S. gordonii, CCR is elicited by a broad range of sugars and we were unable to identify a sugar that does not lead to CCR in S. gordonii43. In fact, some sugars such as galactose and lactose elicit complex regulatory responses in S. gordonii that involve multiple regulators in addition to CcpA53. It is interesting that galactose did not reduce SsnA activity in an agar plate assay (Supplementary Fig. 2), whereas it strongly repressed SsnA in broth cultures (Fig. 3). We hypothesise that this may be due to regulation pathways that compensate for galacatose-driven CcpA-mediated repression of ssnA expression on agar plate cultures. Further work will be needed to identify the regulatory pathways that control ssnA expression in biofilms.
The activity of SsnA may have also been affected by the pH in the growth medium. Sugar metabolism led to an acidification of the growth medium to pH < 5.0. Incubating cells at similarly low pH led to a rapid loss of extracellular DNase activity that was only restored through de novo protein synthesis. Mammalian DNase I enzymes tend to have a relatively low pH optimum, around 6.5–7, whereas other DNase I family members show a preference for higher pH conditions54. The pH activity range of recombinant SsnA (6.5 to 10.5) appears to be slightly higher than the S. sanguinis homologue SWAN, which is active from pH 5.5 to 9.0, with maximal activity at approximately neutral pH41. Incubation of recombinant SsnA in low-pH conditions did not lead to irreversible inactivation of the enzyme, and activity was fully restored when the enzyme was switched to a higher-pH buffer (Supplementary Fig. 7). Therefore, it appears that the inactivation of cell wall-bound SsnA following growth in sugars or low-pH medium is primarily due to protein turnover rather than direct inhibition of the enzyme. Nevertheless, it should be noted that chloramphenicol will result in the broad shutdown of protein synthesis, which could indirectly impact SsnA activity, for example by influencing the pH within cells and in the immediate environment.
In addition to pH, metal ions are important for the activity of DNase I family enzymes. For example, the activity of S. sanguinis SWAN is enhanced by Mg2+ and Ca2+41. Similarly, we found that either Mg2+ or Ca2+ promoted SsnA activity. This suggests that SsnA can utilise either of these metals, rather than requiring both like SWAN or some other DNase I-family enzymes. At low concentrations, Mn2+ and Zn2+ also appeared to activate SsnA. However, the activity markedly decreased at >64 μM Mn2+ or >40 μM Zn2+. Low concentrations (40 μM) of the chelating agent EDTA were included in these experiments to remove trace metals associated with the enzyme or DNA substrate. It is possible that Zn2+ displaced Mg2+ or Ca2+ from the EDTA, indirectly activating the enzyme. At higher concentrations, Zn2+ in particular appears to be a potent inhibitor of SsnA. This is consistent with other DNase I family enzymes including human DNase I that are inhibited by Zn2+55. On the other hand, we speculate that the decrease in the activity at higher Mn2+ concentrations may be caused by significant binding of Mn2+ to the DNA substrate, interfering with the enzyme’s recognition/interaction. Concentrations of Zn2+ in saliva are in the order of 0.2–1.2 μM, whereas Ca2+ is present at approximately 0.5–2.75 mM and Mg2+ at 7–30 μM56. These concentrations would potentially support activity of SsnA in oral biofilms.
The role of extracellular DNase enzymes in mediating interspecies interactions in streptococcal biofilms has not previously been investigated. Here, we showed that biofilms formed by S. mutans GS-5 are dependent on eDNA for stability. When cultured with NucB DNase enzyme, S. mutans NG8 produced approximately 20% of the biofilm biomass compared with control (NucB absent), whereas S. mutans UA159 was only reduced to approximately 80% of the control biomass (Supplementary Fig. 8), a level that is consistent with other published work24. It is not clear why some strains are more dependent on eDNA than others, although this phenomenon has also been observed for other oral streptococci57. It should be noted that some laboratory cultures of S. mutans GS-5, including our own, carry mutations in pac and gbpC genes, which lead to secretion of the cell surface protein PAc (antigen I/II) and a lack of production of GbpC (glucan-binding protein C)58. Future studies are needed to determine whether PAc and/or GbpC affect the dependence of S. mutans biofilms on eDNA.
The ability of SsnA to disperse static S. mutans GS-5 biofilms was dependent on the pH of the medium. At pH 5.0, SsnA did not disperse biofilms in contrast to DNase I, which retained anti-biofilm activity. Interestingly, at pH 6.0, SsnA and DNase I reduced biofilms to approximately 40% of levels in controls, indicating that both enzymes were active in these conditions. By contrast, recombinant SsnA was not active below pH 6.5 when tested against a fluorescent oligonucleotide probe. It is possible that the environment within biofilms may buffer the low pH59. Alternatively, this could be due to differences in the nature of the DNA substrate. The ability of SsnA to control S. mutans biofilms at approximately pH 6.0 may be important in conditions within the oral cavity since it has been shown that S. mutans biofilms contain less glucan and are more dependent on eDNA when cultured at pH 6.0 than when grown at pH 7.060. Nevertheless, reduction of pH to 5.0 or lower may represent a defensive strategy of S. mutans against SsnA. It is noteworthy that sucrose and starch availability stimulate further eDNA release and acid production by S. mutans leading to establishment of a stable biofilm25. Consistently, growth of S. mutans in mixed-species biofilms that also contain S. gordonii biofilm is enhanced in the presence of sucrose61,62.
The presence of SsnA inhibited growth of mixed-species oral microcosm biofilms under flow of natural human saliva, suggesting that the anti-biofilm activity of SsnA is not specific to S. mutans biofilms. In addition to S. mutans, single species biofilms of several oral microorganisms such as S. intermedius and S. sanguinis have been shown to be sensitive to treatment with DNase enzymes63,64,65. Since oral biofilm development is strongly affected by interspecies cell-cell interactions66, we speculate that exclusion of certain species could have dramatic effects on the accumulation of dental plaque. The deleterious effects of SsnA on mixed-species oral biofilms are consistent with results obtained from treatment of in situ dental plaque grown with DNase I32. Previously, we showed that the presence of NucB DNase during oral microcosm development leads to reduced biomass67. Intriguingly, the microbial diversity in oral biofilms was also affected. The biofilm developed in presence of NucB contained reduced proportions of anaerobic and proteolytic late coloniser species such as Peptostreptococcus, Porphyromonas, and Prevotella31. Further work is required to establish whether SsnA selectively inhibits certain species within the biofilm.
Based on our findings, we speculate that it may be possible to target the expression of streptococcal DNases such as SsnA to control the growth of dental plaque. However, the inhibition of SsnA by low pH may restrict the development of this strategy against cariogenic biofilms. We note that SsnA was not effective at controlling biofilm growth in sucrose-supplemented saliva, possibly due to the production of acid by microorganisms within the biofilm. The production of glucans which can interact with DNA and stabilise the matrix68 may also have contributed to the lack of activity of SsnA in these conditions. Further studies aiming at understanding the regulatory mechanisms governing the production and the activity of SsnA, and effects of oral microbial extracellular DNases on the structure and microbial composition of dental plaque, could lead to development of novel approaches for controlling multispecies biofilms by targeting the extracellular matrix.
Methods
Bacterial strains and growth conditions
Streptococcus gordonii DL1 (Challis), Staphylococcus aureus FH7 and Streptococcus mutans GS-5 (Supplementary Table 1) were routinely cultured in BHY medium (Brain Heart Infusion 37 g/L and 5 g/L yeast extract) at 37 °C anaerobically (10% CO2, 10% H2 and 80% N2 in a Ruskinn BugBox Plus, Ruskinn Technology Ltd, Bridgend, UK). BHY agar (BHY medium plus 15 g/L Bacto-agar) was used as solid medium. When necessary, antibiotics were included as follows: kanamycin (250 μg/mL), spectinomycin (250 μg/mL) and erythromycin (10 µg/mL for chromosomal insertions and 2 µg/mL for plasmids). Escherichia coli was cultured in Lysogeny Broth (Melford Laboratories, Ipswich, UK) at 37 °C with shaking at 200 rpm.
Intracellular and extracellular DNA extraction
Intracellular DNA (iDNA) or extracellular DNA (eDNA) was extracted as described by Kreth et al.21. S. gordonii or S. mutans were cultured overnight in TYEG medium containing 10 g Bacto tryptone, 5 g yeast extract, 3 g K2HPO4 and 2 g glucose per L, adjusted to pH 7.5. A total of 10 µL were transferred to 2 mL fresh TYEG in wells of a 12-well plastic dish and cultures were incubated with gentle rocking (20 rpm) for 72 h with changes of medium after each 24 h. The supernatant was aspirated and wells were washed twice with 1 mL phosphate buffered saline (PBS), pH 7.4. After aspirating the liquid, a further 1.5 mL PBS was added to one well and the biofilm was harvested by gentle scraping with a tissue culture scraper. To provide sufficient biomass for DNA extraction, this sample was transferred to an equivalent well and additional biofilm was scraped. In total, biofilms from four wells were combined for each sample. Cells were harvested by centrifugation at 12,000 g, 4 °C, 30 min and the supernatant (eDNA sample) and pellet (containing cells with iDNA) were stored at −20 °C until further purification.
For purification of eDNA, an equal volume of 25:24:1 phenol:chloroform:isoamyl alcohol was added and mixed by inversion 30–40 times. Following centrifugation at 16,000 g, 4 °C for 5 min, the aqueous phase was collected and the phenol:chloroform:isoamyl alcohol extraction was repeated. After another centrifugation, the aqueous phase was collected and DNA was precipitated by adding one tenth volume 3 M sodium acetate, pH 5.2 and mixing. Two-thirds volume of isopropanol was added and mixed, and eDNA was pelleted by centrifugation at 16,000 g, 4 °C for 10 min. After washing with 1 ml 100% ethanol, the pellet was air dried and resuspended in 20 µL 10 mM Tris-HCl, pH 8.0.
To extract iDNA, cell pellets were resuspended in 150 µL spheroplasting buffer (20 mM Tris-HCl, 10 mM MgCl2 and 26% (w/v) raffinose, adjusted to pH 6.8). To this, 1.5 µL lysozyme from a 250 µg mL−1 stock and 5 µL mutanolysin from a 10,000 U mL−1 stock were added and samples were incubated for 30 min at 37 °C. Cells were lysed by addition of 150 µL T&C Lysis solution containing 1 µL proteinase K (Epicentre MasterPure DNA purification kit, Epicentre Biotechnologies, Madison, WI, USA) and DNA was extracted by following the manufacturer’s instructions.
Identification of putative DNase I family proteins and ssnA promoter region
To identify potential DNase I family homologues in S. gordonii, the DNase I domain superfamily was identified in the Superfamily database (https://supfam.mrc-lmb.cam.ac.uk/SUPERFAMILY/cgi-bin/scop.cgi?sunid=56219). S. gordonii DNase I family members were identified under ‘genome assignments’. The ssnA gene and promoter region was retrieved from the S. gordonii Challis genome (GenBank accession CP000725.1). Putative transcription factor binding sites were identified using PePPER69.
Biofilm inhibition and dispersal assay
Bacteria were inoculated in 200 μL of media in triplicate in polystyrene 96-well plates. Plates were incubated anaerobically at 37 °C for 20 h. When performing biofilm inhibition assays, bovine DNase I or GST-SsnA (5 µg/mL) was included at the point of inoculation. For dispersal assays, enzymes (5 µg/mL) were added to 20 h old biofilms and incubated for a further hour at 37 °C. Biofilm extent was quantified using the crystal violet assay57. Biofilm assays were repeated three times independently. The significance of differences between absorbance readings was determined by Student’s two sample t-test.
Fluorescence based quantitative DNase activity assay
An oligonucleotide probe, 5′ HEX-CCC CGG ATC CAC CCC- BHQ2 3′ (PrimeTime probe Integrated DNA Technologies), was used for quantification of DNase activity as described by Kiedrowski et al.70. The probe (2 μM of the probe in a buffer consisting of 20 mM Tris-HCl pH 8) was added to 25 μL of a nuclease source (cells or purified enzyme) in a well of a 384-well microtiter plate (Greiner Bio-One, Stonehouse, UK) and the rate of fluorescence change (λex: 530 nm, λem: 590 nm) was measured at 37 °C over 30 min using the Synergy HT (BioTek, Swindon, UK) microplate reader.
Mutagenesis and genetic complementation of S. gordonii DL1
Disruption of ssnA was carried out by replacing the chromosomal ssnA with the aad9 spectinomycin resistance determinant. Flanking regions of the ssnA gene were PCR amplified using primers SsnAF1 and SsnAR1 to generate a 484-bp product in the 5′ region of ssnA and SsnAF2 and SsnAR2 to generate a 674-bp product in the 3′ end of ssnA. The aad9 gene (782-bp) was amplified from plasmid pFW5 (Supplementary Table 1) with primers aad9_SsnAF and aad9_SsnAR primers. These PCR reactions were performed with Taq Reddymix (Sigma Aldrich, Gillingham, UK) and cycling conditions of 94 °C, 2 min, 35 cycles of 94 °C for 10 s, 56 °C for 30 s, 68 °C for 90 s, then 68 °C for 7 min and 4 °C hold. The PCR products were mixed in equimolar ratios and amplified in an overlap extension PCR using primers SsnAF1 and SsnAR2 with the Expand Long Range PCR enzyme mix (Sigma Aldrich). Thermocycling was performed as follows: 92 °C, 2 min, 10 cycles of 92 °C, 10 s, 56 °C, 15 s, 68 °C 150 s, 25 cycles of 92 °C, 10 s, 56 °C, 15 s, 68 °C 150 s and increasing by 20 s per cycle, then 68 °C for 7 min and 4 °C hold. The resulting product was used for transformation of S. gordonii DL1. For this, S. gordonii was cultured in a candle jar to early exponential phase (OD600 nm ~ 0.3) in BHY medium supplemented with 1 µL mL-1 fetal calf serum and 0.1% (w/v) glucose (BHY/FCS/G). After a further 60 min at 37 °C, cultures were dispensed in 0.8 mL aliquots and approximately 1 µg of the PCR product was added. After incubation for a further 4 h, cultures were diluted and cultured on solidified BHY medium for 36 h in a candle jar. Successful disruption and replacement of ssnA gene with the aad9 gene was confirmed by DNA sequencing.
Disruption of ccpA was carried out using a similar approach by replacing ccpA with the kanR kanamycin resistance gene. The upstream region of ccpA was amplified with CcpAF1 and CcpAR1 generating a 520-bp fragment and the downstream region of ccpA was amplified using CcpAF2 and CcpAR2. The kanR gene was amplified with KanR_F and KanR_R primers using pK18 vector (Supplementary Table 1) as template. Gibson Assembly Master Mix (New England BioLabs, Ipswich, Massachusetts, USA) was employed for fusion of the upstream and downstream fragments of ccpA to the kanR gene according to the instructions of the manufacturer. The resulting 1,795-bp fragment was used for transformation of S. gordonii cells. Successful disruption and replacement of the ccpA gene with the kanR gene was confirmed by DNA sequencing.
Mutagenesis of the malR gene was achieved by replacing malR with the ermAM erythromycin resistance cassette. Using the primers malRF1 and malRR1, a 471-bp fragment upstream of malR gene was amplified. malRF2 and malRR2 were used to amplify a 421-bp sequence downstream region of malR gene. The malRR1 and malRF2 primers contained 17-bp complementary sequences for ermAMF2 and ermAMR2 primers. After amplifying the ermAM cassette from pVA838 (Supplementary Table 1) the PCR products were stitched together in an overlap extension PCR. The resulting product was used for transformation of S. gordonii DL1.
For gene complementation, plasmid pssnAComp was created by amplification of ssnA using SsnA.compF and SsnA.compR primers and genomic DNA from wild-type S. gordonii DL1 as template. SsnA.compF and SsnA.compR primers included EcoRI and BamHI restriction enzyme sites, respectively. The amplicon was digested with BamHI and EcoRI and ligated into pDL276 vector (Supplementary Table 1) that had also been digested with BamHI and EcoRI (New England BioLabs). Successful construction of pssnAComp was confirmed by sequencing and was used to transform S. gordonii ΔssnA mutant to generate the genetically complemented strain S. gordonii ssnAComp.
Plasmid pccpAComp was generated by replacing the arcR gene in parcRComp71 with the ccpA gene. Primers CcpA_compF and CcpA_compR were used to amplify the ccpA gene using wild-type S. gordonii genomic DNA as template, generating a 1048-bp fragment. Lin-vecF and Lin-vecR primers were used to create linearized vector using parcRComp plasmid as template, producing a 5797-bp linear vector. The In-Fusion HD PCR ligation cloning kit (Clontech Laboratories, Mountain View, California, USA) was employed to fuse the ccpA fragment to the vector backbone generating pccpAComp. The integrity of plasmid pccpAComp was confirmed by sequencing and pccpAComp was used for transformation of S. gordonii ΔccpA to generate S. gordonii ccpAComp.
Genetic manipulations of E. coli
Routine genetic manipulations were carried out using protocols described by Sambrook et al.72 and explained below. Plasmids and primers are described in Supplementary Table 1 and Supplementary Table 2.
Expression and purification of recombinant SsnA
SsnA was cloned as a glutathione S-transferase (GST) fusion construct. The GST tag was cleaved where necessary. The ssnA gene, minus the C-terminal LPxTG cell anchor motif and N-terminal secretion signal domain, was amplified using primers ssnA_Pf7 and ssnA_Pr7, creating a product of 2150 bp. This product was cloned into pGEX-KT plasmid (Supplementary Table 1) and transformed into E. coli DH5α. For transformation, chemically competent E. coli cells were produced by culturing E. coli to early exponential phase (OD600 nm ~ 0.3) in 50 mL LB, harvesting for 5 min at 5000 rpm and suspending the pellet in 20 mL ice cold 50 mM CaCl2. After standing in ice for 20 min, cells were harvested by centrifugation at 5000 rpm, 4 °C for 5 min and suspended in 4 mL ice cold CaCl2. Samples were distributed into 300 µL portions, DNA was added and incubated on ice for 40 min. Cells were heat-shocked at 42 °C for 2 min and plunged into ice. Transformants were recovered by adding 500 µL pre-warmed SOC medium containing 20 g L−1 Bacto tryptone, 5 g L−1 yeast extract, 10 mM NaCl, 2.5 mM KCl, 10 mM MgCl2, 10 mM MgSO4 and 20 mM glucose, and incubating for 1 h at 37 °C, 250 rpm before spreading across selective LB agar plates. pGEX-ssnA plasmid then was transferred to E. coli BL21 (DE3) pLysS (Stratagene, La Jolla, California, USA). Expression of GST-SsnA was induced by 1 mM isopropyl β-D-1-thiogalactopyranoside (Sigma Aldrich) for 4 h at 37 °C. A glutathione Sepharose column was used to elute GST-SsnA in 10 mM glutathione, 50 mM potassium phosphate, pH 7.2, 1 mM DTT (Sigma Aldrich). SDS-PAGE and Coomassie Brilliant Blue staining and zymography were performed to assess the size and activity of purified GST-SsnA.
Cleavage of the GST tag from SsnA
The N-terminal 26 kDa GST protein tag was cleaved by thrombin. Purified SsnA was exchanged from elution buffer to TNC buffer (20 mM Tris-HCl [pH 7.5], 50 mM NaCl, and 1 mM CaCl2) for optimal cleavage of the GST-tag. Samples were incubated for 24 h at room temperature with 1 U bovine thrombin (Sigma) for every 0.025 mg of SsnA. Thrombin was inactivated by the addition of 0.3 mM phenylmethylsulfonyl fluoride (PMSF) and incubated at 37 °C, 15 min.
Biofilm growth in the BioFlux 1000 microfluidic system
For mixed-species microcosm biofilm growth, saliva was collected from healthy volunteers who had not eaten or drunk (except water) in the last hour and had not taken antibiotics in the last 3 weeks. Ethical approval for the collection of saliva was granted by the Newcastle University Faculty of Medical Sciences Ethics Committee (ref. 1853/519/2020). Where needed, saliva was treated with dithiothreitol, precipitates were removed by centrifugation, and saliva was sterilised by filtration to create cell-free saliva (CFS)73. The BioFlux 1000 (Fluxion, San Francisco, CA) was employed to grow mixed-species oral microcosm biofilms. Prior to inoculation, channels of BioFlux dual-flow 24 well plates (Fluxion) were primed by addition of 200 μL of cell-free saliva (CFS) and incubation at room temperature for 20 min. Excess CFS was then removed from the outlet well and replaced with 100 μL of untreated saliva. Flow was initiated from the outlet to inlet at 1.0 dyn/cm2 for 6 s. The plate was then incubated at 37 °C for 40 min to allow for seeding to take place. Each of the inlet wells was then filled with pre-warmed (at 37 °C) CFS and flow was initiated at 0.5 dyn/cm2 at 37 °C for 18 h and images were taken every 10 min. Where required, CFS in the inlet wells was supplemented with 2% sucrose or DNase enzymes. DNase enzymes used were bovine DNase I (Sigma Aldrich), Bacillus licheniformis NucB67 and S. gordonii SsnA. They were added to the inlets at a concentration of 5 μM. To visualise eDNA, 2.4 nM Yoyo-1 (LifeTechnologies) dye was included with CFS in both inlet wells.
Imaging biofilms
Imaging of biofilms grown in the BioFlux 1000 was carried out using the BioFlux 1000Z Imaging Workstation consisting of a Zeiss Axio Observer Z1 microscope fitted with an environmental enclosure. A Hamamatsu camera (ORCA-Flash 4.0 LT: 4.2 megapixels with 6.5 micron pixels) and BioFlux Montage Software (Fluxion) were used for acquisition of images. Images were analysed using ImageJ v.1.48 (National Institutes of Health)74.
Image analysis of biofilms cultured in the BioFlux
Time-lapse images of each channel were converted into a stack. Each stack was auto-thresholded using the ‘Otsu’ algorithm. A rectangle of w = 1000 h = 200 pixels (stock image size 1024 × 1024 pixels) was selected and saved as a region of interest (ROI). The average pixel intensity (mean grey value) was measured for each slice and plotted using SigmaPlot 13.0 software (Systat Software, San Jose, CA, USA).
Visualisation of static biofilms
For visualisation of static biofilms with confocal laser scanning microscopy (CLSM), stained coverslips were rinsed with PBS and inverted onto a PBS-filled rubber frame secured on a microscope slide. Imaging was performed using a Nikon A1R confocal laser scanning microscope fitted with CFI PLAN APO VC objective (Nikon 60x/1.40 Oil). Images were captured with NIS-Elements C (v4.4, Nikon) software and processed using Imaris (v8.2, Bitplane) software.
DNase test agar
The extracellular DNase activity of S. gordonii was assessed using DNase Test agar (Sigma Aldrich, Gillingham, UK). Bacterial cultures were streaked or spotted (10 µL) on DNase Test agar and incubated aerobically for 48 h at 37 °C. The DNA in the DNase Test agar was precipitated by flooding the plate with 1 N HCl.
DNase activity assay of bacterial cultures
To determine the nuclease activity of bacterial cultures, overnight cultures were washed twice with PBS at pH 7.1 and resuspended in PBS to an optical density, OD600 nm = 1. The DNase activity was then measured for 25 µL of the cell suspension using the Synergy HT microplate reader (BioTek). For experiments to assay the recovery of DNase activity following low-pH incubation, overnight cultures of S. gordonii DL1 cells grown in THYE medium at 37 °C were harvested and re-suspended in a pH 4.5 buffer (77.1 g/L ammonium acetate, 70 mL/L glacial acetic acid), a pH 5.5 buffer (96.3 mL of solution I [13.61 g/L KH2PO4 (BDH)] and 3.6 mL of solution II [35.81 g/L Na2HPO4]) and PBS (pH 7.1). Cells were incubated at room temperature for up to 2 h, washed in PBS, resuspended in fresh PBS and 25 µL aliquots were assayed for DNase activity using the fluorescence-based DNase activity assay. After 2 h, cells were harvested and washed in PBS at pH 7.1. Cultures that had been incubated in acidic buffers (pH 4.5 or 5.5) were split into equal portions prior to harvesting. One portion was resuspended in PBS while the other was resuspended in PBS containing 12.5 µg/mL chloramphenicol to inhibit de novo synthesis of proteins75. Cultures were incubated for up to a further 2 h before DNase activity was measured.
When investigating the activity of purified SsnA in the presence of metals, 40 µM EDTA was included in the reactions to chelate residual metal contaminants of the buffer or associated with the DNA substrate prior to addition of the putative metal co-factor. When this assay was employed to determine the optimal pH for SsnA enzyme activity, the following buffers were used: Sodium acetate trihydrate buffer (pH 4.5-5.5), Bis-Tris (pH 6.0-6.5), Tris-HCl (pH 7.0-9.0) and 3-Cyclohexylamino-1-propanesulfonic acid buffer (pH 9.7-11.0) (Sigma Aldrich).
DNase zymography
Zymography was used to assess the activity of purified SsnA with or without the GST tag against double stranded DNA. Enzymes in Laemmli’s buffer were loaded (without boiling) onto 12% SDS-PAGE gels containing 100 μg/mL salmon sperm DNA (Sigma Aldrich). Following electrophoresis, the gel was incubated in an enzyme reactivation buffer (40 mM Tris-HCl, 5 mM CaCl2, 5 mM MgCl2 and 3% delipidated milk powder [Premier International Foods, St. Albans, UK]) for 20 h at 37 °C. The gel was stained with ethidium bromide (0.5 μg mL−1) for 30 min and washed 3 times for 15 min each, in distilled water. The gel was visualised under ultraviolet light using G:BOX transilluminator (Syngene).
RNA extraction and reverse transcription
Four mL of cell culture was mixed with 4 mL RNAlater (Life Technologies), and the tubes were vortex-mixed for 5 s and incubated at 20 °C for 5 min. Following incubation, cells were harvested at 3800 g for 15 min at 20 °C, the supernatant was discarded, and the pellets were frozen at −80 °C for no longer than 72 h and RNA was extracted76. Samples were thawed at 20 °C and resuspended in 100 μL spheroplasting buffer plus mutanolysin at 500 U/mL. Cells were incubated at 37 °C for 5 min. Total RNA was extracted using the Ambion RiboPure Bacteria RNA Purification kit (Life Technologies) according to the manufacturer’s instructions. RNA concentrations were determined using a NanoDrop ND‐1000 Spectrophotometer (Nanodrop Technologies, LLC, Wilmington, DE, USA). The RNA was analysed by gel electrophoresis to ensure the integrity of the RNA. One μg of total RNA from bacterial cells was reverse transcribed using the QuantiTect Reverse Transcription kit (Qiagen, Hilden, Germany) in accordance with the manufacturer’s instructions.
RT-qPCR analysis of ssnA expression
RT-qPCR reaction mixtures were prepared containing 0.25 μL of cDNA samples and QuantiTect SYBR Green mix (Qiagen), 1 μM forward primer qRT-ssnAF and qRT-ssnAR reverse primer (Supplementary Table 2) in a total volume of 20 μL. Reactions were performed in triplicate using QuantStudio 3 (ThermoFisher Scientific, Waltham, Massachusetts, USA) with the following thermocycling program: initial denaturation at 95 °C for 2 min and 40 cycles of amplification at 95 °C for 15 s, 55 °C for 10 s and 72 °C for 30 s. The Comparative CT Method (ΔΔCT) was used to analyze RT-qPCR data, and data were normalized to the 16 S rRNA gene as a reference (Supplementary Table 2, qRT-16S-F and qRT-16S-F). RT-qPCR reactions were validated using melt curve analysis.
Statistical analyses
All graphs were plotted and statistical tests were performed using GraphPad Prism (GraphPad Software, La Jolla California, USA) version 9. Details of statistical tests are given in figure legends.
Reporting summary
Further information on research design is available in the Nature Research Reporting Summary linked to this article.
Data availability
Quantitative data underpinning the figures in the main manuscript are available at the Newcastle University Research Repository (https://doi.org/10.25405/data.ncl.21539340). Other data are available from the corresponding author upon reasonable request.
References
Bik, E. M. et al. Bacterial diversity in the oral cavity of 10 healthy individuals. ISME J. 4, 962–974 (2010).
Huang, R., Li, M. & Gregory, R. L. Bacterial interactions in dental biofilm. Virulence 2, 435–444 (2011).
Jakubovics, N. S. Intermicrobial interactions as a driver for community composition and stratification of oral biofilms. J. Mol. Biol. 427, 3662–3675 (2015).
Hajishengallis, G. & Lamont, R. J. Beyond the red complex and into more complexity: the polymicrobial synergy and dysbiosis (PSD) model of periodontal disease etiology. Mol. Oral. Microbiol. 27, 409–419 (2012).
Diaz, P. I. & Valm, A. M. Microbial interactions in oral communities mediate emergent biofilm properties. J. Dent. Res. 99, 18–25 (2020).
Zijnge, V. et al. Oral biofilm architecture on natural teeth. PLoS ONE 5, e9321 (2010).
Kolenbrander, P. E. et al. Communication among oral bacteria. Microbiol. Mol. Biol. Rev. 66, 486–505 (2002).
Mann, E. E. et al. Modulation of eDNA release and degradation affects Staphylococcus aureus biofilm maturation. PLoS ONE 4, e5822 (2009).
Nascimento, M. M. et al. The effect of arginine on oral biofilm communities. Mol. Oral. Microbiol. 29, 45–54 (2014).
Belda-Ferre, P. et al. The oral metagenome in health and disease. ISME J. 6, 46–56 (2012).
Marsh, P. D. Microbiology of dental plaque biofilms and their role in oral health and caries. Dent. Clin. North Am. 54, 441–454 (2010).
Vos, T. et al. Global, regional, and national incidence, prevalence, and years lived with disability for 328 diseases and injuries for 195 countries, 1990–2016: a systematic analysis for the Global Burden of Disease Study 2016. Lancet 390, 1211–1259 (2017).
Marsh, P. D. Microbial ecology of dental plaque and its significance in health and disease. Adv. Dent. Res. 8, 263–271 (1994).
Marsh, P. D. & Zaura, E. Dental biofilm: ecological interactions in health and disease. J. Clin. Periodontol. 44(Suppl 18), S12–s22 (2017).
Simón-Soro, A. & Mira, A. Solving the etiology of dental caries. Trends Microbiol. 23, 76–82 (2015).
Serrage, H. J., Jepson, M. A., Rostami, N., Jakubovics, N. S. & Nobbs, A. H. Understanding the matrix: the role of extracellular DNA in oral biofilms. Front Oral Health 2, https://doi.org/10.3389/froh.2021.640129 (2021).
Liu, Y.-L., Nascimento, M. & Burne, R. A. Progress toward understanding the contribution of alkali generation in dental biofilms to inhibition of dental caries. Int. J. Oral. Sci. 4, 135–140 (2012).
Jakubovics, N. S. et al. Critical roles of arginine in growth and biofilm development by Streptococcus gordonii. Mol. Microbiol. 97, 281–300 (2015).
Nascimento, M. M. et al. Oral arginine metabolism may decrease the risk for dental caries in children. J. Dent. Res. 92, 604–608 (2013).
Huang, X., Schulte, R. M., Burne, R. A. & Nascimento, M. M. Characterization of the arginolytic microflora provides insights into pH homeostasis in human oral biofilms. Caries Res. 49, 165–176 (2015).
Kreth, J., Zhang, Y. & Herzberg, M. C. Streptococcal antagonism in oral biofilms: Streptococcus sanguinis and Streptococcus gordonii interference with Streptococcus mutans. J. Bacteriol. 190, 4632–4640 (2008).
Firestone, A. R., Schmid, R. & Mühlemann, H. R. Cariogenic effects of cooked wheat starch alone or with sucrose and frequency-controlled feedings in rats. Arch. Oral. Biol. 27, 759–763 (1982).
Liao, S. et al. Streptococcus mutans extracellular DNA is upregulated during growth in biofilms, actively released via membrane vesicles, and influenced by components of the protein secretion machinery. J. Bacteriol. 196, 2355–2366 (2014).
Perry, J. A., Cvitkovitch, D. G. & Lévesque, C. M. Cell death in Streptococcus mutans biofilms: a link between CSP and extracellular DNA. FEMS Microbiol. Lett. 299, 261–266 (2009).
Klein, M. I. et al. Dynamics of Streptococcus mutans transcriptome in response to starch and sucrose during biofilm development. PLoS ONE 5, e13478 (2010).
Muto, Y. & Goto, S. Transformation by extracellular DNA produced by Pseudomonas aeruginosa. FEMS Microbiol. Immunol. 30, 621–628 (1986).
Jakubovics, N. S., Shields, R. C., Rajarajan, N. & Burgess, J. G. Life after death: the critical role of extracellular DNA in microbial biofilms. Lett. Appl. Microbiol. 57, 467–475 (2013).
Whitchurch, C. B., Tolker-Nielsen, T., Ragas, P. C. & Mattick, J. S. Extracellular DNA required for bacterial biofilm formation. Science 295, 1487 (2002).
Flemming, H.-C. & Wingender, J. The biofilm matrix. Nat. Rev. Microbiol. 8, 623–633 (2010).
Dragoš, A. & Kovács, Á. T. The peculiar functions of the bacterial extracellular matrix. Trends Microbiol. 25, 257–266 (2017).
Rostami, N. et al. A critical role for extracellular DNA in dental plaque formation. J. Dent. Res. 96, 208–216 (2017).
Schlafer, S., Meyer, R. L., Dige, I. & Regina, V. R. Extracellular DNA contributes to dental biofilm stability. Caries Res. 51, 436–442 (2017).
Porschen, R. K. & Sonntag, S. Extracellular deoxyribonuclease production by anaerobic bacteria. Appl. Microbiol. 27, 1031–1033 (1974).
Palmer, L. J., Chapple, I. L. C., Wright, H. J., Roberts, A. & Cooper, P. R. Extracellular deoxyribonuclease production by periodontal bacteria. J. Periodontal Res. 47, 439–445 (2012).
Mulcahy, H., Charron-Mazenod, L. & Lewenza, S. Extracellular DNA chelates cations and induces antibiotic resistance in Pseudomonas aeruginosa biofilms. PLoS Pathog. 4, e1000213 (2008).
Mulcahy, H., Charron-Mazenod, L. & Lewenza, S. Pseudomonas aeruginosa produces an extracellular deoxyribonuclease that is required for utilization of DNA as a nutrient source. Environ. Microbiol. 12, 1621–1629 (2010).
Binnenkade, L., Kreienbaum, M. & Thormann, K. M. Characterization of ExeM, an extracellular nuclease of Shewanella oneidensis MR-1. Front. Microbiol. 9, 1761 (2018).
Hannan, S. et al. Transfer of antibiotic resistance by transformation with eDNA within oral biofilms. FEMS Immunol. Med. Microbiol. 59, 345–349 (2010).
Sumby, P. et al. Extracellular deoxyribonuclease made by group A Streptococcus assists pathogenesis by enhancing evasion of the innate immune response. Proc. Natl Acad. Sci. USA 102, 1679–1684 (2005).
Berends, E. T. et al. Nuclease expression by Staphylococcus aureus facilitates escape from neutrophil extracellular traps. J. Innate Immun. 2, 576–586 (2010).
Morita, C. et al. Cell wall-anchored nuclease of Streptococcus sanguinis contributes to escape from neutrophil extracellular trap-mediated bacteriocidal activity. PLoS ONE 9, e103125 (2014).
Fontaine, M. C., Perez-Casal, J. & Willson, P. J. Investigation of a novel DNase of Streptococcus suis Serotype 2. Infect. Immun. 72, 774–781 (2004).
Tong, H., Zeng, L. & Burne, R. A. The EIIABMan phosphotransferase system permease regulates carbohydrate catabolite repression in Streptococcus gordonii. Appl. Environ. Microbiol. 77, 1957–1965 (2011).
Yang, W. Nucleases: diversity of structure, function and mechanism. Q. Rev. Biophys. 44, 1–93 (2011).
Podbielski, A., Zarges, I., Flosdorff, A. & Weber-Heynemann, J. Molecular characterization of a major serotype M49 group A streptococcal DNase gene (sdaD). Infect. Immun. 64, 5349–5356 (1996).
Sriskandan, S., Unnikrishnan, M., Krausz, T. & Cohen, J. Mitogenic factor (MF) is the major DNase of serotype M89 Streptococcus pyogenes. Microbiology 146, 2785–2792 (2000).
Remmington, A. & Turner, C. E. The DNases of pathogenic Lancefield streptococci. Microbiology 164, 242–250 (2018).
Liu, J. et al. A nuclease from Streptococcus mutans facilitates biofilm dispersal and escape from killing by neutrophil extracellular traps. Front Cell Infect. Microbiol 7, 97 (2017).
Dlakić, M. Functionally unrelated signalling proteins contain a fold similar to Mg2+ dependent endonucleases. Trends Biochem. Sci. 25, 272–273 (2000).
Maddi, A., Haase, E. & Scannapieco, F. Mass spectrometric analysis of whole secretome and amylase-precipitated secretome proteins from Streptococcus gordonii. J. Proteom. Bioinform. 7, 287–295 (2014).
Shelburne, S. A. 3rd et al. A direct link between carbohydrate utilization and virulence in the major human pathogen group A Streptococcus. Proc. Natl Acad. Sci. USA 105, 1698–1703 (2008).
Willenborg, J., de Greeff, A., Jarek, M., Valentin-Weigand, P. & Goethe, R. The CcpA regulon of Streptococcus suis reveals novel insights into the regulation of the streptococcal central carbon metabolism by binding of CcpA to two distinct binding motifs. Mol. Microbiol. 92, 61–83 (2014).
Zeng, L., Martino, N. C. & Burne, R. A. Two gene clusters coordinate galactose and lactose metabolism in Streptococcus gordonii. Appl. Environ. Microbiol. 78, 5597–5605 (2012).
Yasuda, T. et al. A single amino acid substitution can shift the optimum pH of DNase I for enzyme activity: biochemical and molecular analysis of the piscine DNase I family. Biochim. Biophys. Acta 1672, 174–183 (2004).
Jang, D. S. et al. Novel high-throughput deoxyribonuclease 1 assay. J. Biomol. Screen 20, 202–211 (2015).
Jakubovics, N. S. Saliva as the sole nutritional source in the development of multispecies communities in dental plaque. Microbiol Spec 3, https://doi.org/10.1128/microbiolspec.MBP-0013-2014 (2015).
Shields, R. C. et al. Efficacy of a marine bacterial nuclease against biofilm forming microorganisms isolated from chronic rhinosinusitis. PLoS ONE 8, e55339 (2013).
Sato, Y., Okamoto, K. & Kizaki, H. gbpC and pac gene mutations detected in Streptococcus mutans strain GS-5. Oral. Microbiol. Immunol. 17, 263–266 (2002).
Dong, Y., Chen, Y. Y., Snyder, J. A. & Burne, R. A. Isolation and molecular analysis of the gene cluster for the arginine deiminase system from Streptococcus gordonii DL1. Appl. Environ. Microbiol. 68, 5549–5553 (2002).
Kawarai, T., Narisawa, N., Suzuki, Y., Nagasawa, R. & Senpuku, H. Streptococcus mutans biofilm formation is dependent on extracellular DNA in primary low pH conditions. J. Oral. Biosci. 58, 55–61 (2016).
Tsutsumi, K., Maruyama, M., Uchiyama, A. & Shibasaki, K. Characterisation of a sucrose-independent in vitro biofilm model of supragingival plaque. Oral. Dis. 24, 465–475 (2018).
Kreth, J., Zhu, L., Merritt, J., Shi, W. & Qi, F. Role of sucrose in the fitness of Streptococcus mutans. Oral. Microbiol. Immunol. 23, 213–219 (2008).
Petersen, F. C., Tao, L. & Scheie, A. A. DNA binding-uptake system: a link between cell-to-cell communication and biofilm formation. J. Bacteriol. 187, 4392–4400 (2005).
Nur, A. et al. Effects of extracellular DNA and DNA-binding protein on the development of a Streptococcus intermedius biofilm. J. Appl. Microbiol. 115, 260–270 (2013).
Moraes, J. J. et al. Two-component system VicRK regulates functions associated with establishment of Streptococcus sanguinis in biofilms. Infect. Immun. 82, 4941–4951 (2014).
Jakubovics, N. S. Talk of the town: interspecies communication in oral biofilms. Mol. Oral. Microbiol. 25, 4–14 (2010).
Nijland, R., Hall, M. J. & Burgess, J. G. Dispersal of biofilms by secreted, matrix degrading, bacterial DNase. PLoS ONE 5, e15668 (2010).
Rainey, K., Michalek, S. M., Wen, Z. T. & Wu, H. Glycosyltransferase-mediated biofilm matrix dynamics and virulence of Streptococcus mutans. Appl Environ. Microbiol. 85, e02247–02218 (2019).
Podbielski, A., Spellerberg, B., Woischnik, M., Pohl, B. & Lütticken, R. Novel series of plasmid vectors for gene inactivation and expression analysis in group A streptococci (GAS). Gene 177, 137–147 (1996).
Kiedrowski, M. R. et al. Nuclease modulates biofilm formation in community-associated methicillin-resistant Staphylococcus aureus. PLoS One 6, e26714 (2011).
Robinson, J. C. et al. ArcR modulates biofilm formation in the dental plaque colonizer Streptococcus gordonii. Mol. Oral. Microbiol 33, 143–154 (2018).
Sambrook J. R. D. Molecular cloning: a laboratory manual, 3rd ed. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, N.Y. (2001).
Samarian, D. S., Jakubovics, N. S., Luo, T. L. & Rickard, A. H. Use of a high-throughput in vitro microfluidic system to develop oral multi-species biofilms. JoVE, 52467 (2014).
Brown, J. M., Blunk, B., Williams, P. & Hardie, K. R. Microfluidic-based growth and imaging of bacterial biofilms. Bio-Protoc. 9, e3460 (2019).
Gale, E. F. & Folkes, J. P. The assimilation of amino-acids by bacteria. XV. Actions of antibiotics on nucleic acid and protein synthesis in Staphylococcus aureus. J. Biochem. 53, 493–498 (1953).
Mutha, N. V. R. et al. Transcriptional responses of Streptococcus gordonii and Fusobacterium nucleatum to coaggregation. Mol. Oral. Microbiol. 33, 450–464 (2018).
Acknowledgements
This work was funded by an International Association for Dental Research (IADR) Innovation in Oral Care Award (NSJ) and by The Dunhill Medical Trust (RPGF1810\101) (NSJ/AHN/KJW). KJW was funded by the Biotechnology and Biological Sciences Research Council (BB/S006818/1). PhD funding for CL was from the National Institutes of Health (DE016690). We thank Ekaterina Kozhevnikova and Philip Hardy for technical support and the Newcastle University Protein and Proteome Analysis (NUPPA) Core Facility for their assistance in enzyme purification.
Author information
Authors and Affiliations
Contributions
N.S.J., N.R., A.N., A.T., K.J.W., and R.C.S. designed experiments; N.R., R.C.S., S.Y., C.L., J.L.B., P.T., and H.A. conducted experiments; N.S.J., N.R., A.N., A.T., K.J.W., and R.C.S. analysed and interpreted the data; N.R., R.C.S., and N.S.J. drafted the manuscript; all authors critically reviewed the manuscript and approved the final submitted version. N.R. and R.C.S. contributed equally to the work.
Corresponding author
Ethics declarations
Competing interests
Newcastle University holds patent WO2011098579A1 ‘Bacterial deoxyribonuclease compounds and methods for biofilm disruption and prevention’. There are no other competing interests.
Additional information
Publisher’s note Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
Rights and permissions
Open Access This article is licensed under a Creative Commons Attribution 4.0 International License, which permits use, sharing, adaptation, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The images or other third party material in this article are included in the article’s Creative Commons license, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons license and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this license, visit http://creativecommons.org/licenses/by/4.0/.
About this article
Cite this article
Rostami, N., Shields, R.C., Serrage, H.J. et al. Interspecies competition in oral biofilms mediated by Streptococcus gordonii extracellular deoxyribonuclease SsnA. npj Biofilms Microbiomes 8, 96 (2022). https://doi.org/10.1038/s41522-022-00359-z
Received:
Accepted:
Published:
DOI: https://doi.org/10.1038/s41522-022-00359-z
Comments
By submitting a comment you agree to abide by our Terms and Community Guidelines. If you find something abusive or that does not comply with our terms or guidelines please flag it as inappropriate.
