Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2019 Feb 1.
Published in final edited form as: Environ Microbiol Rep. 2017 Dec 4;10(1):12–22. doi: 10.1111/1758-2229.12600

Differential oxidative stress tolerance of Streptococcus mutans isolates affects competition in an ecological mixed-species biofilm model

Yuan Liu 1,#, Sara R Palmer 2,#, Hsiaochi Chang 2, Ashton N Combs 2, Robert A Burne 3, Hyun Koo 1
PMCID: PMC5812797  NIHMSID: NIHMS921408  PMID: 29124888

Summary

Streptococcus mutans strongly influences the development of pathogenic biofilms associated with dental caries. Our understanding of S. mutans behavior in biofilms is based on a few well-characterized laboratory strains; however, individual isolates vary widely in genome content and virulence-associated phenotypes, such as biofilm formation and environmental stress sensitivity. Using an ecological biofilm model, we assessed the impact of co-cultivation of several S. mutans isolates with Streptococcus oralis and Actinomyces naeslundii on biofilm composition following exposure to sucrose. The laboratory reference strain S. mutans UA159 and clinical isolates Smu44 (most aciduric), Smu56 (altered biofilm formation) and Smu81 (more sensitive to oxidative stress) were used. Our data revealed S. mutans isolates varied in their ability to compete and become dominant in the biofilm after the addition of sucrose, and this difference correlated with sensitivity to H2O2 produced by S. oralis. Smu81 was particularly sensitive to H2O2 and couldn’t compete with S. oralis in mixed-species biofilm, despite forming robust biofilms on its own. Thus, diminished oxidative stress tolerance in S. mutans isolates can impair their ability to compete in complex biofilms, even in the presence of sucrose, which could influence the progression of a healthy biofilm community to one capable of causing disease.

Keywords: Streptococcus mutans, Streptococcus oralis, ecological mixed-species biofilm, oxidative stress tolerance

Introduction

Dental caries is the most prevalent and costly biofilm-associated oral infectious disease (Kassebaum et al., 2015). The pathogenesis of dental caries is characterized by changes in the biofilm (or plaque) composition triggered by the host diet; transitioning from a microbiota dominated by health-associated commensals to one that contains greater proportions of highly acidogenic and acid tolerant organisms enmeshed in an exopolysaccharide (EPS) rich matrix (Paes Leme et al., 2006; Takahashi and Nyvad, 2011). The human oral cavity is home to a diverse microbiome (Dewhirst, 2010) that constantly interacts with a salivary film (pellicle) formed on the tooth surface, to which a group of early colonizers (mostly streptococci and actinomyces species) adhere with high affinity. Many synergistic and antagonistic interactions between these bacterial species influence the composition of the colonizing community (Kuramitsu et al., 2007; Kuboniwa, 2006). For example, health-associated commensal bacteria, mainly Mitis group strep, can antagonize the growth of potentially harmful species through the production of hydrogen peroxide (H2O2) and/or bacteriocins, thus providing protection to the host against disease (Kuramitsu et al., 2007; Merritt and Qi, 2012; Qi and Kreth, 2017).

The transition from health to disease is driven in large part by the metabolism of dietary sugars (Paes Leme et al., 2006). Frequent consumption of sucrose, in particular, dramatically influences cariogenic biofilm development by providing a substrate for both EPS and organic acid production. The EPS promote bacterial accumulation and cohesion on tooth surfaces, while creating a diffusion-limiting matrix (Bowen and Koo, 2011). The metabolic activity of the bacteria enmeshed within EPS-rich matrices can create localized acidic microenvironments, despite the nearly continuous supply of buffering saliva. The persistence of acidic microenvironments allows acidogenic and acid-tolerant species to out-compete the acid-sensitive commensal bacteria associated with health (Takahashi and Nyvad, 2011; Koo et al., 2013). Furthermore, in transcriptomic studies that measured gene transcripts expressed in natural oral biofilm communities, about 2% of all transcripts identified were associated with oxidative stress tolerance genes, including those involved with detoxifying superoxides and peroxides (Peterson et al., 2014), suggesting that the ability of bacteria to cope with oxidative stress is critical to their survival when residing in biofilms. Once established, cariogenic biofilms increases the potential for demineralization of tooth enamel, leading to the onset and progression of dental caries.

Streptococcus mutans is the bacterium that has been most closely and consistently associated (albeit not exclusively) with the initiation of dental caries (Bowen, 2016). S. mutans lives almost entirely in dental plaque on the tooth surface, where there are dramatic fluctuations in nutrient availability, pH and oxidative stress (Bitoun, et al., 2011). The unique combined abilities of this pathogen to produce EPS and acids, to be acid-tolerant, and to rapidly adapt to environmental stresses are essential for establishment and persistence in biofilms and for competition with other oral bacteria when conditions are conducive to the development of dental caries (Lemos and Burne, 2008). However, diversity exists within the S. mutans species, which can be classified into at least four different serological groups (c, e, f, k) (Nakano and Ooshima, 2009); most strains isolated from the human oral cavity are serotype c (70-80%) (Palmer et al., 2013). In addition, the carriage of particular genotypes of S. mutans may correlate (or not) with the caries status of an individual (Napimoga et al., 2004; Pieralisi et al., 2010; Phattarataratip et al., 2011).

Recently, whole genome sequencing was completed on a large collection of strains of S. mutans from across the globe, providing new insights into the core and accessory genomes of the organism (Cornejo et al., 2013). In this study 73 unique-core genes (found in S. mutans, but absent in its closest relatives) were defined, of which 49% were hypothetical proteins, with the remaining involved in carbohydrate metabolism, acid resistance, adaptation to oxidative stress, metal and peptide translocation, and adhesion to host tissues (Cornejo et al., 2013). In a subsequent study, phenotypic properties directly associated with the ability of S. mutans to establish, persist, and cause disease in humans were demonstrated to vary greatly among these genetically- and geographically-heterogeneous clinical isolates (Palmer et al., 2013). These findings provide evidence that the genomic diversity of the species may translate to substantial variations in their ability to compete with physiologically similar organisms in human oral biofilms. Thus, a critical question to be addressed is whether genomic variability influences the way in which S. mutans interacts and competes with abundant commensal bacteria that dominate oral biofilms in a healthy state. Such information could provide powerful new tools to accurately assess caries risk and to develop new therapies that hinder the cariogenic potential of S. mutans and oral biofilms. In the present study, we compared the ability of four S. mutans isolates that vary considerably in genome composition and key phenotypic characteristics to compete with Streptococcus oralis and Actinomyces naeslundii under cariogenic conditions using an established mixed-species ecological biofilm model.

Results and Discussion

The clinical isolates utilized in this study are described in Table 1 and selection of these particular strains was based on specific phenotypic characteristics associated with the virulence potential of S. mutans, as previously described (Palmer et al., 2013). Typically, S. mutans UA159 forms more biofilm in the presence of sucrose than glucose. However, Smu56 was able to form biofilms equally well in the presence of either carbohydrates. Smu44 stood out among S. mutans isolates in aciduricity (ability to grow and continue to produce acids at low pH), which is an important virulence attribute of S. mutans. Smu44 grew at a significantly faster rate compared to UA159 in media acidified to pH 5.5, whereas Smu56 and Smu81 grew more slowly than UA159 in acidic conditions (Palmer et al., 2013). Smu81 was selected based on its enhanced sensitivity to oxidative stress and because it contains the cnm gene associated with the ability of S. mutans to invade epithelial and endothelial cells. This strain is unique among S. mutans isolates studied, as it is unable to initiate growth in the presence of 25 mM paraquat (Palmer et al., 2013), which can generate superoxide anion. Importantly, the ability to tolerate oxidative stress is critical to the establishment, persistence and ecology of oral bacteria, and redox and oxygen levels affect the pathogenic potential of dental plaque in major ways (Bradshaw et al., 1996; Marquis, 1995; Lemos and Burne, 2008). These strains also differ in the number of mutacin (bacteriocin) genes present. Mutacin genes encode peptide antibiotics that are important for the establishment of S. mutans in the complex oral microbiota (Merritt and Qi, 2012), and their expression has been linked to oxidative stress (Ahn et al., 2007). Smu81 lacks mutacin IV (nlmA/B), while Smu56 and Smu44 contain only nlmA and are missing nlmB, as detailed in (Palmer et al., 2013).

Table 1.

S. mutans clinical isolates used in this study.

Accession number Lab ID Strain name Notable characteristics compared to reference strains S. mutans UA159* Host/Sample Origin/MLST Serotype
ASM33925 Smu44 11VS1 More aciduric, forms poor biofilms in sucrose, forms more biofilms in glucose, contains type VII secretion system 30 months, initial stages of oral colonization, caries free Brazil/ST28 C
ASM33933 Smu56 N29 Slow growth, forms equal biofilms in sucrose and glucose, sensitive to oxidative stress Dental plaque UK/SK11 C
ASM33943 Smu81 SF14 Oxidative stress sensitive, cnm+ Supragingival plaque US/ST58 C
Open in a new tab
*

Information adapted from Palmer et al., 2013 PLoS ONE

Differential response of clinical isolates towards other oral species in mixed-species system

S. mutans co-exists and associates with a number of other streptococci and bacterial species in the oral cavity (Jenkinson, 2011; Dewhirst et al., 2010). Analysis of the microbial species composition in an initial colonizing community have shown that the majority (47 to 90%) of cultivable bacteria are non-mutans streptococci, such as S. oralis, and that one-third of the remaining bacteria are Actinomyces species, such as A. naeslundii. In the mixed-species ecological biofilm model, S. mutans is co-cultured with the early colonizers S. oralis and A. naeslundii to form an initial microbial consortium on pellicle-coated hydroxyapatite. Then, sucrose is introduced to induce the transition from a non-virulent microbial community (high levels of S. oralis, few A. naeslundii cells and S. mutans as the least abundant species) to a biofilm characterized by an EPS-rich matrix and acidic microenvironments that favor S. mutans survival (Koo et al., 2010; Xiao et al., 2012) (Fig. S1), mimicking the biochemical and microbiological changes associated with cariogenic biofilms. Using this model, we observed intriguing variation in the ability of the different S. mutans strains to compete with the other species during biofilm development.

Prior to introduction of 1% sucrose (Fig. 1) the bacterial populations on the sHA surface were comprised mainly of S. oralis and some A. naeslundii cells. However, after the introduction of 1% sucrose at 29 h, the proportions of S. mutans UA159 and Smu44 rapidly increased, and the S. mutans isolates became the dominant species in 91 h biofilms (Fig. 1A). While Smu56 displayed a much slower population shift, representing a noticeably lower proportion compared to UA159 at 67 h (Fig. 1B, P<0.001), it eventually became dominant at 91 h, suggesting that Smu56 is slow to adapt to environmental stress (i.e. low pH or increased sucrose). Remarkably, even in the presence of 1% sucrose, Smu81 was never able to overtake the other species in the biofilm, and made up a strikingly lower proportion of the bacteria present at 67 h and 91 h, compared to biofilms formed with the other S. mutans strains (Fig. 1B, P<0.001).

Fig. 1. Mixed-species biofilm formation by clinical isolates in UFTYE.

Fig. 1

Open in a new tab

A) Bacterial species were inoculated in the culture medium supplemented with 0.1% (w/v) sucrose until 29 h for establishment of initial biofilm community. The biofilms were then challenged with an environmental change by introducing 1% (w/v) sucrose. Viable population of S. mutans, S. oralis and A. naeslundii recovered from the biofilms were counted (number of CFU recovered per biofilm) over time and the proportion of each strain at each time point was calculated based on CFU data. B) The percentage of S. mutans after exposure to 1% sucrose at 67 h and 91 h. After the introduction of 1% sucrose at 29 h, the proportions of S. mutans UA159 and Smu44 rapidly increased, and became the dominant species in 91 h biofilms. Smu56 displayed a much slower population shift, representing a noticeably lower proportion compared to UA159 at 67 h. While, Smu81 was never able to overtake the other species, with a strikingly lower proportion of the bacteria present at 67 h and 91 h, compared to biofilms formed with the other S. mutans strains. Results represent the mean ± standard deviation. Statistical analysis was performed using One-way ANOVA with post-hoc Tukey HSD test. Each strain was compared to UA159 at each time point. ** P ≤ 0.001.

When individual isolates were compared in single-species biofilms, Smu44 produced significantly less biofilm, likely due to both reduced EPS and bacterial cells, compared to UA159 (Supplementary Fig. S2), but still became the dominant species within the mixed-species biofilm after 1% sucrose was added. While oxidative stress-sensitive strain Smu81 produced similar EPS to UA159 during single species biofilms (Supplementary Fig. S2), it could not out-compete S. oralis in the mixed-species biofilm. Collectively, these results suggest that the differences seen between the abilities of S. mutans strains to establish and survive in the mixed-species ecological biofilms are due to additional factors other than their capacity to form EPS. The glycosyltransferases (Gtfs) of S. mutans contribute to the accumulation of this pathogen in a mixed-species environment after introduction of sucrose by increasing the biomass and EPS content of the biofilms, triggering the formation of structured microcolonies (Koo et al., 2010). However, variations in Gtf expression among S. mutans strains did not seem to contribute to their survival and dominance at the later stages of the mixed-species biofilm development. Considering the inability of Smu81 to grow in the presence of paraquat (Palmer et al., 2013), it is conceivable that the differences observed between the S. mutans isolates in the mixed-species biofilms may be determined by reduced resistance to oxidative stress, perhaps caused by the H2O2 produced by S. oralis, rather than differences in EPS production and biofilm formation.

Differences in oxidative stress tolerance between S. mutans strains

To determine whether the growth of S. mutans strains is inhibited by S. oralis, or vice versa, a plate-based growth inhibition test was performed. Actively growing S. mutans were spotted next to S. oralis on agar plates and allowed to grow for 48 h. While UA159 was unaffected by S. oralis, the growth of Smu81 was strongly inhibited by the commensal (Fig. 2A). Smu44 and Smu56 were slightly sensitive to growth inhibition by S. oralis, but more resistant than Smu81. Importantly, when the plates were pretreated with catalase, which neutralizes H2O2, Smu81, Smu44 and Smu56 were no longer inhibited by S. oralis (Fig. 2A). It’s also important to note that growth of S. oralis was not inhibited by the S. mutans strains, as would be expected if S. oralis were sensitive to mutacins produced by S. mutans. Thus, Smu81 may aberrantly express, be lacking in, or contain key mutations in, gene products required for H2O2 tolerance. To explore in more detail the differences between the S. mutans isolates to cope with H2O2, growth in the presence of H2O2 was monitored using an automated growth monitoring machine (Bioscreen C, Labsystem). The growth rate, lag time and maximum yield were calculated based on the growth curves. Table 2 shows the results for each S. mutans strain grown under non-stress conditions compared to growth in the presence of H2O2. In the presence of H2O2, UA159, Smu44, and Smu81 all had significantly slower growth rates compared to non-stress conditions, while Smu56’s growth rate was not significantly different (P>0.05). In addition, UA159, Smu44, and Smu56 had similar yields when grown with H2O2 compared to non-stress conditions. Consistent with the sensitivity of Smu81 to growth inhibition by S. oralis, this strain showed the most pronounced decrease in growth yield, and greatest increase in lag time (320 min for Smu81), compared to the other S. mutans strains tested (UA159- 50 min, Smu44- 80 min and Smu56- 310 min). To further define the level of oxidative stress sensitivity among the S. mutans strains, a H2O2 killing assay was completed, where the percent viable cells was determined after exposure to H2O2 for 15, 30, and 45 min. Consistent with data from the growth curve assays and the agar spot test, Smu81 was the most sensitive to H2O2 of the four strains tested. It took ~25 min of exposure for Smu56, and 45 min for UA159 or Smu44, to see the same level of killing as Smu81 experienced after only 15 min exposure to assay conditions (Fig. 2B).

Fig. 2. Susceptibility of S. mutans clinical isolates to growth inhibtion by S. oralis and to killing by hydrogen peroxide.

Fig. 2

Open in a new tab

A) Growth inhibition of S. mutans clinical isolates (Smu44, Smu56, Smu81) in response to S. oralis. S. mutans UA159 is resistant to growth inhibition by S. oralis and is included for comparison (left). Growth inhibition of S. mutans clinical isolates by S. oralis is not observed in the presence of 300 Units of Catalase, which degrades H2O2 (right). Images are representative results of quadruplicate experiments. B) Results of hydrogen peroxide killing assay. Aliquots of cells suspended in glycine buffer (pH 7) containing H2O2 were removed at indicated times (y-axis), and percent survival (x-axis) was determined based on CFU. Assay was performed with triplicate biological samples and repeated one time with similar results. Statistical analysis was performed using Two-way ANOVA with post-hoc Tukey test. Each strain was compared to UA159 at each time point. * P ≤ 0.01, ** P ≤ 0.001, and *** P ≤ 0.0001.

Table 2.

Growth characteristics of different S. mutans strains, grown with or without H2O2.

Growth Condition UA159 Smu44 Smu56 Smu81
Non-stress
Td 49±2 52±6 119±4 60±3
Lag 210 210 300 210
Yield 0.94 0.95 0.82 1.00
250 μM H2O2
Td 55±1‡ 79±2† 135±11 71±3†
Lag 260† 290* 610* 530*
Yield 0.96 0.89 0.86 0.80*
Open in a new tab

Mean growth rate (Td) and growth lag with standard deviation are expressed as min, and were calculated based on growth curves. Yield is the mean maximum OD600 reached during growth. Significance is indicated by

*

= P ≤ 0.001,

†

= P ≤ 0.01, and

‡

= P ≤ 0.05, and compares non-stress growth to growth with H2O2.

While S. mutans does not produce catalase, it contains several pathways that neutralize reactive oxygen species (ROS), including an alkyhydroperoxide reductase AhpCF (Smu.764 and Smu.765), a Dps-like iron binding protein Dpr (Smu.540), a Flavin-based NAD (NADH) oxidase (nox, Smu.1117), and a superoxide dismutase (Sod, Smu.629) (Higuchi et al., 2000; Yamamoto et al., 2000). To further explore the differences in ROS resistance between S. mutans strains, we compared the expression of ahpC (Smu.764), ahpF (Smu.765), dpr (Smu.540), and nox (Smu.1117) by qRT-PCR in cells grown with or without a 15 min exposure to H2O2. While there were differences in expression of oxidative stress genes between the clinical isolates and S. mutans UA159 (Fig. 3A), results were contrary to what we expected: the strains that were more sensitive to H2O2 (Smu56 and Smu81) actually expressed significantly higher levels of aphF and/or aphC compared to the more resistant strains, UA159 and Smu44 (Fig. 3B). The reason for this disparity could be due to differences in expression of other genes not evaluated, or from differences in enzymatic activity due to polymorphisms in the oxidative stress genes themselves. Therefore, the NADH oxidase enzyme activity of the S. mutans strains was determined after a 30 or 45 min exposure to H2O2 and compared to untreated samples (no H2O2) (Fig. 3C). While Smu44 was the only strain that did not see a significant increase in NADH oxidase activity after 30 min, all strains saw a significant increase after the 45 min exposure to H2O2. In addition, despite Smu81 being the most sensitive to oxidative stress, we were surprised to find this strain produced more NADH oxidase activity upon H2O2 treatment than any other S. mutans strains tested, including UA159 (Fig. 3D). Furthermore, there was no correlation between the NADH oxidase activity observed and expression of genes responsible for NADH oxidase activity (nox and aphFC).

Fig. 3. Expression of oxidative stress genes and NADH oxidase activity in S. mutans clinical isolates.

Fig. 3

Open in a new tab

(A) Expression of dpr, ahpF, ahpC, and nox genes, quantitated by real-time PCR (qRT-PCR), were compared between S. mutans strains treated with or without H2O2. Results represent triplicate biological samples with three technical replicates. Student’s t-test was used to determine significance. * = P ≤ 0.01. (B) Results of statistical analysis comparing expression of oxidative stress genes in each clinical isolate to reference strain UA159. * = P ≤ 0.05, determined by One-way ANOVA and post-hoc Tukey HSD test. * denotes clinical isolates that had significantly different level of expression of indicated genes compared to UA159. (C) NADH oxidase activity produced by each strain after a 30 or 45 min exposure to H2O2 compared to untreated controls (no H2O2). Results represent the mean ± standard error of nmol NADH oxidized min−1 mg−1 protein. * = P ≤ 0.05, determined by Student’s t-test comparing H2O2 treated to untreated samples. Assay results represent triplicate biological samples. (D) Statistical analysis compareing S. mutans isolates to reference strain UA159 grown under the same conditions. Statistical differences indicated by * (P ≤ 0.05) determined by One-way ANOVA and post-hoc Tukey HSD test. * denotes clinical isolates that expressed significantly different level of NADH oxidase activity compared to UA159.

Next, we compared the protein sequences of oxidative stress genes to see if there was any incongruence between strains that could explain the differences in H2O2 sensitivity. Smu.629 (sod) and Smu.764 (ahpC) were 100% conserved among all S. mutans strains; however, Smu81 had two polymorphisms in Smu.765 (ahpF), and one in the iron binding protein dpr (Table 3), both of which contribute to S. mutans resistance to H2O2 (Fujishima, et al., 2013). Further, when the sequences for spxA (Smu.1142c), a global regulator of oxidative stress genes in S. mutans (Kajfasz et al., 2015), was compared, Smu81 was the only strain out of the 57 included in the analysis that harbored mutations in this gene (Table 3). Given the high level of conservation of this gene product among sequenced strains of S. mutans, it is possible that these polymorphisms significantly affect the ability of SpxA to sense oxidative stress and appropriately regulate oxidative stress genes and, therefore, could partially explain why Smu81 is so sensitive to H2O2. SpxA functions as a transcriptional regulator by binding to RNA polymerase, either inhibiting or activating transcription of genes required for adaptation to oxidative stress. Previous work found that Smu.764 (ahpC) and Smu.765 (ahpF) are regulated by SpxA in S. mutans UA159, and that spxA deletion mutants display significantly less NADH oxidase activity than the parental strain (Kajfasz et al., 2015). However, Smu81 had 4-fold greater expression of the Smu.764 gene upon treatment with H2O2, compared to UA159, suggesting that decreased expression of Smu.764 is not the reason for the oxidative stress sensitivity of this strain. Compared to UA159, Smu81 also produced more than twice as much NADH oxidase activity. Therefore, aberrant expression of other oxidative stress genes regulated by SpxA, not evaluated here, may explain our results. Further, the polymorphism in the iron binding protein Dpr, which prevents Fenton reactions, may also be responsible for the sensitivity to H2O2 observed in Smu81. Accordingly, future studies will investigate the complete transcriptomes of the S. mutans strains under oxidative stress to gain additional insights into how genetic differences between strains affect global gene regulation in S. mutans. We will also examine the enzymatic activities of oxidative stress response pathways and activity of the transcriptional regulator SpxA to further dissect the exact mechanism behind the differences in oxidative stress tolerance among S. mutans strains.

Table 3.

S. mutans clinical isolates contain polymorphisms within oxidative stress genes when compared to UA159

Gene Enzyme Function Polymorphisms*
ahpF Nox-1 NADH oxidase Reduces 2NADH + O2 -> H2O2 Smu81 A195 to T; P197 to S
ahpC Alkyl hydroperoxide reductase Deactivates H2O2 100 % identical
dpr Dps-like peroxide resistance gene Fe+ binding protein - prevents Fenton reactions Smu81 T43 to A
nox-2 NADH oxidase H20 forming NADH oxidase Smu81 R130 to C; H324 to Y; T459 to I
Smu56 D233 to N; R430 to H
Smu44 T430 to I
spxA1 Transcriptional regulator Major regulator of oxidative stress genes Smu81 A18 to T; S35 to N; K99 to R
Open in a new tab

Polymorphisms are based on sequence alignments performed using shotgun genome sequence for clinical isolates compared to S. mutans reference strain UA159. NCBI accession numbers for each strain are listed in Table 1. Note: The gene for superoxide dismutase (Smu.620) was 100% identical in all strains.

Caries development is clearly associated with the overgrowth of a few bacterial species, including S. mutans, certain lactobacilli and bifidobacteriae, and correlated with acidification of the biofilm microenvironment and a reduction in overall species diversity (Gross et al., 2012; Caufield et al., 2007; Takahashi and Nyvad, 2011). Interspecies interactions play an important role in this shift. Many beneficial early-colonizers are sensitive to low pH conditions, while the disease-associated mutans streptococci are more acid tolerant, but sensitive to H2O2. Therefore, the ability of commensal streptococci (i.e. S. oralis, S. gordonii, and S. sanguinis) to produce H2O2, which can antagonize cariogenic S. mutans (Kreth et al., 2005), is regarded as a key protective mechanism against competing species in early healthy biofilm communities (Bitoun, et al., 2011; Zhu et al., 2012). However, perturbations in the environment triggered by dietary sugars promote the growth of opportunistic pathogenic species, which can be present in health at low numbers, resulting in changes in cariogenic potential of the biofilm community overall (Nyvad and Kilian, 1990; Takahashi and Nyvad, 2011). As a consequence, the populating species can influence whether the biofilm behaves benignly by restricting potential pathogens from either colonizing or accumulating in sufficient proportions to elicit disease. Although, most S. mutans strains have some capacity to deal with oxidative stress caused by damaging reactive oxygen species (e.g. O2.−) and unfavorable cellular redox potential (Fujishima et al., 2013; Marquis, 1995), Smu81 was distinct from the other strains and displayed an unusually high degree of sensitivity to H2O2, and therefore was unable to dominate the biofilm even in the presence of 1% sucrose. Thus, in order to compete within biofilms, S. mutans must be able to cope with various oxidative stresses (Marquis, 1995), either through enzymatic deactivation, or by associating with other oxidative stress tolerant bacteria, before it can become numerically significant and modify the biofilm milieu to become more pathogenic.

The results presented here demonstrate that the considerable genomic heterogeneity and disparate phenotypic behaviors in virulence-related properties translate to clear differences in the way in which various S. mutans isolates interact and compete with common oral commensal bacteria. While much attention has been paid to determining the ecology of dental biofilms in health and disease, simply knowing which species are there tells only part of the story. Understanding intra-species variability, such as virulence characteristics or variations in stress tolerance within a species and its effect on biofilm ecology, are also vital to understanding disease etiology. Approximately 28% of S. mutans strains isolated from dental biofilm are naturally competent and able to take up DNA from their environment (Westergren and Emilson, 1983), which contributes to the genomic and phenotypic plasticity of this species. Based on gene content clustering of the S. mutans strains used in this study, Smu44 and UA159 differed by ~23%, Smu56 and UA159 by ~22%, and Smu81 and UA159 by ~19% (Cornejo et al., 2013), thus small differences in gene content can result is great differences in virulence properties. A previous study, based on the genome sequence of 57 S. mutans isolates, revealed a core genome of 1,490 genes (present in all S. mutans strains), and a pan-genome of 3,296 genes (total number of genes found in all strains) (Cornejo et al., 2013). The S. mutans genome contains ~ 2.0 Mb of DNA and encodes roughly 2,000 genes (Ajdic et al., 2002). Thus, in any single isolate of S. mutans, about 500 genes could be transient or dispensable, and fluctuate based on the particular environment or niche of that isolate; so not all S. mutans isolates are likely to be equally virulent in any particular host.

While, it is understood that inter-species interactions are crucial to community dynamics and the development of polymicrobial diseases, such as dental caries (Mira et al., 2017; Marsh and Zaura, 2017), a better understanding of bacteria-bacteria interactions, particularly phenotypically diverse strains of S. mutans and commensal bacteria, and how these interactions influence the establishment of pathogenic microenvironments, could contribute to the design of novel preventive and therapeutic strategies against human dental caries. Furthermore, understanding how S. mutans promotes genomic heterogeneity and disparate phenotypic behaviors could be exploited to modulate its pathogenicity and perhaps turn-off S. mutans virulence.

Experimental procedures

Mixed-species biofilm model

The mixed-species biofilm model was based on a batch culture approach using saliva-coated hydroxyapatite (sHA) disks designed to mimic the formation of biofilms according to the “ecological plaque-biofilm” concept (Marsh, 2003), as recently described by (Koo et al., 2010; Xiao et al., 2012). Briefly, S. mutans UA159, S. oralis ATCC 35037 and A. naeslundii ATCC 12104 were grown in ultrafiltered (10-kDa-cutoff membrane; Prep/Scale; Millipore, MA) buffered (pH 7.0) tryptone-yeast extract broth (UFTYE) containing 2.5% tryptone and 1.5% yeast extract with 1% (wt/vol) glucose to mid-exponential phase (optical density at 600 nm [OD600] = 0.5). All isolates of S. mutans except Smu56 had similar growth rates, as determined by monitoring OD600 (Spectronic 20D+, Thermo Fisher Scientific Inc., Madison, WI). The bacterial suspensions were mixed to obtain an inoculum containing a defined microbial population consisting of S. mutans (102 CFU/ml), S. oralis (107 CFU/ml), and A. naeslundii (106 CFU/ml). Biofilms were formed on hydroxyapatite disks (diameter, 1.25 cm; Clarkson Chromatography Products, Inc., South Williamsport, PA) coated with filter-sterilized, clarified, human whole saliva, with the discs held in a vertical position using a custom-made holder (Xiao et al., 2012). The mixed population of S. mutans isolates, S. oralis, and A. naeslundii was inoculated in 2.8 ml of UFTYE with 0.1% (wt/vol) sucrose and incubated at 37˚C in a 5% CO2 aerobic atmosphere. During the first 19 h, the cells were grown undisturbed to allow initial biofilm formation. At the 19 h time point, spent media was replaced with fresh culture medium containing 0.1% (wt/vol) sucrose and the biofilms were grown until the 29 h time point to allow establishment of the initial biofilm community (Xiao et al., 2012). The biofilms were then transferred to UFTYE containing 1% (wt/vol) sucrose to induce environmental changes that simulate a cariogenic challenge. The culture medium was then replaced twice daily at 8 am and 6 pm until the end of the experimental period (91 h) (Supplementary Fig. S1). S. mutans single-species biofilm was formed under the same condition as described above. The biofilms were analyzed at specific time points (29, 67 and 91 h) using biochemical and microbiological assays. Results are presented as mean ± standard deviation (SD). The experimental results presented are representative of three separate experiments. One-way ANOVA was performed, followed by post-hoc Tukey HSD test and statistical analysis was performed using the Prism-Graphpad software.

Biochemical and microbiological analyses

The single- and mixed-species biofilms were removed at specific time points and dispersed by sonication in a sterile 0.89% (wt/vol) NaCl solution (30 sec pulse at an output of 7 W; Branson Sonifier 150; Branson Ultrasonics, Danbury, CT); the sonication procedure does not kill any of the bacterial species used in this study (Xiao et al., 2012). The cell suspensions were used to determine the dry weight and for plating on blood agar using an automated EddyJet Spiral Plater (IUL, SA, Barcelona, Spain) to determine the total CFU per biofilm. The three species were differentiated by colony morphology in conjunction with microscopic examination of cells from selected colonies (Guggenheim et al., 2001)

Growth inhibition test

The ability of S. oralis to inhibit growth of S. mutans strains was determined by growth on agar plates, and performed similarly to experiments described by (Huang et al., 2016). For assay, 6 μl of overnight cultures of S. mutans strains and S. oralis were spotted next to one another on BHI agar plates and allowed to grow for 48 h in a 5% CO2 aerobic incubator at 37˚C. Experiments were also performed in which 300 units of catalase (3000 U/ml in 50 mM potassium phosphate buffer, pH 7.0) was applied to agar surface and allowed to air dry before strains were spotted on the plates. The experimental results presented are representative of four separate experiments.

Growth experiments

Mid-exponential phase cultures of each S. mutans strain grown in brain heart infusion (BHI) broth (BBL, BD Difco) were sub-cultured 1:100 into BHI broth or BHI broth supplemented with 250 μM H2O2. A 300 μl aliquot of each culture was dispensed into triplicate wells in a Bioscreen C Honeycomb plate, and overlaid with 50 μl sterile mineral oil to limit oxygen exposure. The OD600 was measured every 30 min for 48 h in a Bioscreen C Automated Microbiology Growth Curve Analysis System (Growth Curves USA, Piscataway, NJ) set to 37˚C, with shaking for 10 sec and a 5 sec pause before each reading. Doubling time (Td) was calculated based on the slope of the line of the logarithmic growth phase using the formula Td = (t2-t1)ln (2)/ln(OD2)-ln(OD1), as described in (Khalichi et al., 2004), where OD600 was graphed on a semi-log plot against time and MATLAB was used to fit to the logarithmic growth phase. Data are based on triplicate biological samples. Statistically significant differences were determined based on Student’s t-test. Lag (the time from inoculation until exponential growth began) and yield (maximum OD reached) are based on growth curve results.

Hydrogen peroxide killing assay

Mid-exponential cells grown in BHI were resuspended in 0.1 M glycine (pH 7) containing 58.8 mM (0.2%) hydrogen peroxide. Aliquots of cell suspension were taken at 0, 15, 30, and 45 min and serially diluted in tryptone phosphate buffer (pH 7.0) containing 1 mg/ml catalase. Percent survival was determined by dividing the CFUs of treated samples by CFU of untreated controls × 100. Assays were performed on triplicate biological samples and repeated twice. Statistical analysis was performed in the GraphPad Prism program using a Two-way ANOVA with post-hoc Tukey HSD test, and compared each clinical isolate at each time point to S. mutans reference strain UA159.

qRT-PCR

Briefly, total RNA was isolated using the RNeasy® Mini Kit (Qiagen) as described previously (Palmer and Burne, 2015), from cultures grown in BHI broth and treated with 200 μM H2O2 and incubated at 37˚C in a 5% CO2 environment. cDNA was synthesized using the Maxima™ H Minus cDNA Synthesis Master Mix and dsDNase (Thermo Scientific™) according to manufacturer’s protocol. qRT-PCR was performed using the QuantStudio 3 Real-Time PCR system (Applied Biosystems™) with the Power-Up™ SYBR™ Green Master Mix (Applied Biosystems™) according to supplier’s directions. Absolute quantification of each gene was determined based on standard curves. Results are presented as mean ± standard error (SE). Experiment was performed with triplicate biological samples. Student’s t-test was used to determine significant differences in expression of oxidative stress sensitive genes between H2O2 treated and untreated samples for each clinical isolate (Figure 3A). One-way ANOVA was also performed, followed by post-hoc Tukey HSD test, to compare the level of expression of the individual genes in the S. mutans clinical isolates to that of reference strain UA159 (Figure 3B). Statistical analysis was performed using the Prism-Graphpad software. Primers used can be found in supplemental information (Supplementary Table S1).

NADH oxidase assay

Whole cell lysates were prepared from mid-exponential cultures treated with 250 μM H2O2, or untreated controls, and incubated for 30 or 45 min. Assay reactions contained 0.1 M K3PO4 buffer (pH 7), 0.3 mM EDTA, 0.16 mM NADH, and cell lysates, and the decrease in absorbance at 340 nm was monitored over 4 min. NADH oxidase activity was calculated by multiplying the change in OD340 over time by the extinction coefficient for NADH at 340 nm (6.22 × 103M−1, cm−1). NADH activity was standardized by protein concentration determined using the Pierce BCA Protein Assay Kit (Thermo Scientific™), with BSA as a standard. Assay was performed with triplicate biological samples. Results represent the mean ± standard error (SE). Student’s t-test was used to determine significant differences in NADH oxidase activity between H2O2 treated and untreated samples for each strain (Figure 3C), and One-way ANOVA with post-hoc Tukey HSD test, was used to test differences between clinical islates and S. mutans reference strain UA159 (Figure 3D). Statistical analysis was performed using Prism-Graphpad software.

Supplementary Material

Supp FigS1
Supp FigS2
Supp info

Acknowledgments

This work was supported in part by the National Institute for Dental and Craniofacial Research (NIDCR) grants DE025220 and DE018023 (HK), DE023833 (SRP) and DE132329 and DE25832 (RAB).

Footnotes

Conflict of interest

The authors declare no conflict of interest.

References

  1. Aas JA, Paster BJ, Stokes LN, Olsen I, Dewhirst FE. Defining the normal bacterial flora of the oral cavity. J Clin Microbiol. 2005;43:5721–5732. doi: 10.1128/JCM.43.11.5721-5732.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Abdallah AM, Gey van Pittius NC, Champion PA, Cox J, Luirink J, Vandenbroucke-Grauls CM, et al. Type VII secretion—mycobacteria show the way. Nat Rev Microbiol. 2007;5:883–891. doi: 10.1038/nrmicro1773. [DOI] [PubMed] [Google Scholar]
  3. Ajdic D, McShan WM, McLaughlin RE, Savic G, Chang J, Carson MB, et al. Genome sequence of Streptococcus mutans UA159, a cariogenic dental pathogen. Proc Natl Acad Sci U S A. 2002;99:14434–14439. doi: 10.1073/pnas.172501299. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Ahn SJ, Wen ZT, Burne RA. Effects of oxygen on virulence traits of Streptococcus mutans. J Bacteriol. 2007;189:8519–8527. doi: 10.1128/JB.01180-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bitoun JP, Nguyen AH, Fan Y, Burne RA, Wen ZT. Transcriptional repressor Rex is involved in regulation of oxidative stress response and biofilm formation by Streptococcus mutans. FEMS Microbiol Lett. 2011;320:110–117. doi: 10.1111/j.1574-6968.2011.02293.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Bowen WH. Do we need to be concerned about dental caries in the coming millennium? Crit Rev Oral Biol Med. 2002;13:126–131. doi: 10.1177/154411130201300203. [DOI] [PubMed] [Google Scholar]
  7. Bowen WH. Dental caries-not just holes in teeth! A perspective. Mol Oral Microbiol. 2016;31:228–233. doi: 10.1111/omi.12132. [DOI] [PubMed] [Google Scholar]
  8. Bowen WH, Koo H. Biology of Streptococcus mutans-derived glucosyltransferases: role in extracellular matrix formation of cariogenic biofilms. Caries Res. 2011;45:69–86. doi: 10.1159/000324598. [DOI] [PMC free article] [PubMed] [Google Scholar]
  9. Bradshaw DJ, Marsh PD, Allison C, Schilling KM. Effect of oxygen, inoculum composition and flow rate on development of mixed-culture oral biofilms. Microbiology. 1996;142:623–629. doi: 10.1099/13500872-142-3-623. [DOI] [PubMed] [Google Scholar]
  10. Burts ML, Williams WA, DeBord K, Missiakas DM. EsxA and EsxB are secreted by an ESAT-6-like system that is required for the pathogenesis of Staphylococcus aureus infections. Proc Natl Acad Sci U S A. 2005;102:1169–1174. doi: 10.1073/pnas.0405620102. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Caufield PW, Li Y, Dasanayake A, Saxena D. Diversity of lactobacilli in the oral cavities of young women with dental caries. Caries Res. 2007;41:2–8. doi: 10.1159/000096099. [DOI] [PMC free article] [PubMed] [Google Scholar]
  12. Clermont O, Couffignal C, Blanco J, Mentré F, Picard B, Denamur E, et al. Two levels of specialization in bacteraemic Escherichia coli strains revealed by their comparison with commensal strains. Epidemiol Infect. 2016;28:1–11. doi: 10.1017/S0950268816003010. [DOI] [PMC free article] [PubMed] [Google Scholar]
  13. Cornejo OE, Lefébure T, Bitar PD, Lang P, Richards VP, Eilertson K, et al. Evolutionary and population genomics of the cavity causing bacteria Streptococcus mutans. Mol Biol Evol. 2013;20:881–893. doi: 10.1093/molbev/mss278. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Derr AM, Faustoferri RC, Betzenhauser MJ, Gonzalez K, Marquis RE, Quivey RG., Jr Mutation of the NADH oxidase gene (nox) reveals an overlap of the oxygen- and acid-mediated stress responses in Streptococcus mutans. Appl Environ Microbiol. 2012;78:1215–1227. doi: 10.1128/AEM.06890-11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  15. Dewhirst FE, Chen T, Izard J, Paster BJ, Tanner AC, Yu WH, et al. The human oral microbiome. J Bacteriol. 2010;192:5002–5017. doi: 10.1128/JB.00542-10. [DOI] [PMC free article] [PubMed] [Google Scholar]
  16. Fujishima K, Kawada-Matsuo M, Oogai Y, Tokuda M, Torii M, Komatsuzawa H. dpr and sod in Streptococcus mutans are involved in coexistence with S. sanguinis, and PerR is associatied with resistance to H2O2. Appl Environ Microbiol. 2013;79:1436–1443. doi: 10.1128/AEM.03306-12. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Gross EL, Beall CJ, Kutsch SR, Firestone ND, Leys EJ, Griffen AL. Beyond Streptococcus mutans: dental caries onset linked to multiple species by 16S rRNA community analysis. PLoS One. 2012;7:e47722. doi: 10.1371/journal.pone.0047722. [DOI] [PMC free article] [PubMed] [Google Scholar]
  18. Guggenheim B, Giersten E, Schupbach P, Shapiro S. Validation of an in vitro biofilm model of supragingival plaque. J Dent Res. 2001;80:363–370. doi: 10.1177/00220345010800011201. [DOI] [PubMed] [Google Scholar]
  19. Han YW. Fusobacterium nucleatum: a commensal-turned pathogen. Curr Opin Microbiol. 2015;23:141–147. doi: 10.1016/j.mib.2014.11.013. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Higuchi M, Yamamoto Y, Kamio Y. Molecular biology of oxygen tolerance in lactic acid bacteria: functions of NADH oxidases and Dpr in oxidative stress. J Biosci Bioeng. 2000;90:484–493. [PubMed] [Google Scholar]
  21. Huang X, Palmer SR, Ahn SJ, Richards VP, Williams ML, Nascimento MM, et al. A highly arginolytic Streptococcus species that potently antagonizes Streptococcus mutans. Appl Environ Microbiol. 2016;82:2187–2201. doi: 10.1128/AEM.03887-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  22. Jenkinson HF. Beyond the oral microbiome. Environ Microbiol. 2011;13:3077–3087. doi: 10.1111/j.1462-2920.2011.02573.x. [DOI] [PubMed] [Google Scholar]
  23. Kajfasz JK, Rivera-Ramos I, Scott-Anne K, Gregoire S, Abranches J, Lemos JA. Transcription of Oxidative Stress Genes Is Directly Activated by SpxA1 and, to a Lesser Extent, by SpxA2 in Streptococcus mutans. J Bacteriol. 2015;197:2160–2170. doi: 10.1128/JB.00118-15. [DOI] [PMC free article] [PubMed] [Google Scholar]
  24. Kassebaum NJ, Barnabé E, Dahiya M, Bhandari B, Murray CJ, Marcenes W. Global burden of untreated caries: a systematic review and metaregression. J Dent Res. 2015;94:650–658. doi: 10.1177/0022034515573272. [DOI] [PubMed] [Google Scholar]
  25. Khalichi J, Cvitkovitch DG, Santerre JP. Effect of composite resin biodegradation products on oral streptococcal growth. Biomaterials. 2004;25:5467–5472. doi: 10.1016/j.biomaterials.2003.12.056. [DOI] [PubMed] [Google Scholar]
  26. Koo H, Falsetta ML, Klein MI. The exopolysaccharide matrix: a virulence determinant of cariogenic biofilm. J Dent Res. 2013;92:1065–1073. doi: 10.1177/0022034513504218. [DOI] [PMC free article] [PubMed] [Google Scholar]
  27. Koo H, Xiao J, Klein MI, Jeon JG. Exopolysaccharides produced by Streptococcus mutans glucosyltransferases modulate the establishment of microcolonies within multispecies biofilms. J Bacteriol. 2010;192:3024–3032. doi: 10.1128/JB.01649-09. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Kreth J, Merritt J, Shi W, Qi F. Competition and coexistence between Streptococcus mutans and Streptococcus sanguinis in the dental biofilm. J Bacteriol. 2005;187:193–203. doi: 10.1128/JB.187.21.7193-7203.2005. [DOI] [PMC free article] [PubMed] [Google Scholar]
  29. Kuboniwa M, Tribble GD, James CE, Kilic AO, Tao L, Herzberg MC, et al. Streptococcus gordonii utilizes several distinct gene functions to recruit Porphyromonas gingivalis into a mixed community. Mol Microbiol. 2006;60:121–139. doi: 10.1111/j.1365-2958.2006.05099.x. [DOI] [PubMed] [Google Scholar]
  30. Kuramitsu HK, He X, Lux R, Anderson MH, Shi W. Interspecies interactions within oral microbial communities. Microbiol Mol Biol Rev. 2007;71:653–670. doi: 10.1128/MMBR.00024-07. [DOI] [PMC free article] [PubMed] [Google Scholar]
  31. Lemos JA, Burne RA. A model of efficiency: stress tolerance by Streptococcus mutans. Microbiology. 2008;154:3247–3255. doi: 10.1099/mic.0.2008/023770-0. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Marquis RE. Oxygen metabolism, oxidative stress and acid-base physiology of dental plaque biofilms. J Ind Microbiol. 1995;15:198–207. doi: 10.1007/BF01569826. [DOI] [PubMed] [Google Scholar]
  33. Marsh PD. Are dental diseases examples of ecological catastrophes? Microbiology. 2003;149:279–294. doi: 10.1099/mic.0.26082-0. [DOI] [PubMed] [Google Scholar]
  34. Marsh PD, Zaura E. Dental biofilm: ecological interactions in health and disease. J Clin Periodontol. 2017;44:S12–S22. doi: 10.1111/jcpe.12679. [DOI] [PubMed] [Google Scholar]
  35. Merritt J, Qi F. The mutacins of Streptococcus mutans: regulation and ecology. Mol Oral Microbiol. 2012;27:57–69. doi: 10.1111/j.2041-1014.2011.00634.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  36. Mira A, Simon-Soro A, Curtis MA. Role of microbial communities in the pathogenesis of periodontal diseases and caries. J Clin Periodontol. 2017;44:S23–S38. doi: 10.1111/jcpe.12671. [DOI] [PubMed] [Google Scholar]
  37. Nakano K, Ooshima T. Serotype classification of Streptococcus mutans and its detection outside the oral cavity. Future Microbiol. 2009;4:891–902. doi: 10.2217/fmb.09.64. [DOI] [PubMed] [Google Scholar]
  38. Napimoga MH, Kamiya RU, Rosa RT, Rosa EA, Höfling JF, de Oliveira Mattos-Graner R, et al. Genotypic diversity and virulence traits of Streptococcus mutans in caries-free and caries-active individuals. J Med Microbiol. 2004;53:697–703. doi: 10.1099/jmm.0.05512-0. [DOI] [PubMed] [Google Scholar]
  39. Nyvad B, Kilian M. Comparison of the initial streptococcal microflora on dental enamel in caries-active and in caries-inactive individuals. Caries Res. 1990;24:267–272. doi: 10.1159/000261281. [DOI] [PubMed] [Google Scholar]
  40. Paes Leme AF, Koo H, Bellato CM, Bedi G, Cury JA. The role of sucrose in cariogenic dental biofilm formation--new insight. J Dent Res. 2006;85:878–887. doi: 10.1177/154405910608501002. [DOI] [PMC free article] [PubMed] [Google Scholar]
  41. Palmer SR, Miller JH, Abranches J, Zeng L, Lefebure T, Richards VP, et al. Phenotypic heterogeneity of genomically-diverse isolates of Streptococcus mutans. PLoS one. 2013;8:e61358. doi: 10.1371/journal.pone.0061358. [DOI] [PMC free article] [PubMed] [Google Scholar]
  42. Palmer SR, Burne RA. Post-transcriptional regulation by distal Shine-Dalgarno sequences in the grpE-dnaK intergenic region of Streptococcus mutans. Mol Microbiol. 2015;98:302–317. doi: 10.1111/mmi.13122. [DOI] [PMC free article] [PubMed] [Google Scholar]
  43. Peterson SN, Meissner T, Su AI, Snesrud E, Ong AC, Schork NJ, et al. Functional expression of dental plaque microbiota. Front Cell Infect Microbiol. 2014;4:108. doi: 10.3389/fcimb.2014.00108. [DOI] [PMC free article] [PubMed] [Google Scholar]
  44. Pieralisi FJ, Rodrigues MR, Segura VG, Maciel SM, Ferreira FB, Garcia JE, et al. Genotypic Diversity of Streptococcus mutans in Caries-Free and Caries-Active Preschool Children. Int J Dent. 2010;2010:824976. doi: 10.1155/2010/824976. [DOI] [PMC free article] [PubMed] [Google Scholar]
  45. Phattarataratip E, Olson B, Broffitt B, Qian F, Brogden KA, Drake DR, et al. Streptococcus mutans strains recovered from caries‐active or caries‐free individuals differ in sensitivity to host antimicrobial peptides. Mol Oral Microbiol. 2011;26:187–199. doi: 10.1111/j.2041-1014.2011.00607.x. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Qi F, Kreth J. Methods to study antagonistic activities among oral bacteria. Methods Mol Biol. 2017;1537:203–218. doi: 10.1007/978-1-4939-6685-1_12. [DOI] [PubMed] [Google Scholar]
  47. Takahashi N, Nyvad B. The role of bacteria in the caries process: ecological perspectives. J Dent Res. 2011;90:294–303. doi: 10.1177/0022034510379602. [DOI] [PubMed] [Google Scholar]
  48. Westergren G, Emilson CG. Prevalence of transformable Streptococcus mutans in human dental plaque. Infect Immun. 1983;41:1386–1388. doi: 10.1128/iai.41.3.1386-1388.1983. [DOI] [PMC free article] [PubMed] [Google Scholar]
  49. Xiao J, Klein MI, Falsetta ML, Lu B, Delahunty CM, Yates JR, et al. The exopolysaccharide matrix modulates the interaction between 3D architecture and virulence of a mixed-species oral biofilm. PLoS Pathog. 2012;8:e1002623. doi: 10.1371/journal.ppat.1002623. [DOI] [PMC free article] [PubMed] [Google Scholar]
  50. Yamamoto Y, Higuchi M, Poole LB, Kamio Y. Identification of a new gene responsible for the oxygen tolerance in aerobic life of Streptococcus mutans. Biosci Biotechnol Biochem. 2000;64:1106–1109. doi: 10.1271/bbb.64.1106. [DOI] [PubMed] [Google Scholar]
  51. Zhu L, Kreth J. The role of hydrogen peroxide in environmental adaptation of oral microbial communities. Oxid Med Cell Longev. 2012;2012:717843. doi: 10.1155/2012/717843. [DOI] [PMC free article] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Supp FigS1
NIHMS921408-supplement-Supp_FigS1.tif (330KB, tif)
Supp FigS2
NIHMS921408-supplement-Supp_FigS2.tif (155.7KB, tif)
Supp info
NIHMS921408-supplement-Supp_info.docx (16.1KB, docx)

RESOURCES