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      A Novel Dental Sealant Containing Dimethylaminohexadecyl Methacrylate Suppresses the Cariogenic Pathogenicity of Streptococcus mutans Biofilms

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          Abstract

          Cariogenic oral biofilms are strongly linked to dental caries around dental sealants. Quaternary ammonium monomers copolymerized with dental resin systems have been increasingly explored for modulation of biofilm growth. Here, we investigated the effect of dimethylaminohexadecyl methacrylate (DMAHDM) on the cariogenic pathogenicity of Streptococcus mutans ( S. mutans) biofilms. DMAHDM at 5 mass% was incorporated into a parental formulation containing 20 mass% nanoparticles of amorphous calcium phosphate (NACP). S. mutans biofilms were grown on the formulations, and biofilm inhibition and virulence properties were assessed. The tolerances to acid stress and hydrogen peroxide stress were also evaluated. Our findings suggest that incorporating 5% DMAHDM into 20% NACP-containing sealants (1) imparts a detrimental biological effect on S. mutans by reducing colony-forming unit counts, metabolic activity and exopolysaccharide synthesis; and (2) reduces overall acid production and tolerance to oxygen stress, two major virulence factors of this microorganism. These results provide a perspective on the value of integrating bioactive restorative materials with traditional caries management approaches in clinical practice. Contact-killing strategies via dental materials aiming to prevent or at least reduce high numbers of cariogenic bacteria may be a promising approach to decrease caries in patients at high risk.

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          The Exopolysaccharide Matrix Modulates the Interaction between 3D Architecture and Virulence of a Mixed-Species Oral Biofilm

          Introduction Virulent biofilms formed on surfaces are prime biological phenomena that are associated with many illnesses and infections in humans, including oral diseases, native valve endocarditis, and a number of nosocomial infections [1]. Human dental plaque is one of the most complex biofilm systems in nature, as it is constantly exposed to environmental challenges and therefore it may serve as a general model to study the biology of biofilms [2]. Dental caries is a highly prevalent and costly biofilm-dependent oral infectious disease, which continues to be a significant cause of emergency room visits and absences from work or school [3]. This ubiquitous disease results from the complex interactions that occur on tooth surfaces between specific oral bacteria, the products produced by these organisms, salivary constituents, and dietary carbohydrates [4]. These interactions modulate the transition from a healthy condition to a disease state (dental caries), which is characterized by the establishment of cariogenic biofilms on the susceptible tooth surface, and eventually leads to cavitation through acid dissolution of enamel [4]. All bacterial biofilms have at least one property in common: the presence of a bacterial-derived matrix that envelops the cells and helps to facilitate the formation of multicellular structures that are firmly attached to a surface [5]. The composition and structure of the biofilm matrix may be as variable as the microbial constituents, are influenced by local environmental conditions, and can change with time [5], [6]. Exopolysaccharides (EPS), proteins, lipids, nucleic acids, lipoteichoic acids, and even lipopolysaccharides have been identified in the matrices of bacterial biofilms [5], [7]–[9]. The matrix is considered essential for the existence of the biofilm lifestyle and full expression of virulence by bacterial pathogens [5]. However, most previous research has been focused on the (bacterial) cellular aspects of biofilm, while the functional roles that matrices play in the expression of virulence have received limited attention. Exopolysaccharides are key components of the matrix in cariogenic oral biofilms, and are recognized virulence factors involved in the pathogenesis of dental caries [4]. In the mouth, a highly diverse microbial community is constantly interacting with a proteinaceous film present on the tooth surface, known as the pellicle. The pellicle, to which a small group of organisms (mostly Streptococci and Actinomyces spp.) can adhere in low numbers [10]–[12], is derived from mammalian and bacterial sources (such as salivary proteins and exoenzymes). Cariogenic organisms such as Streptococcus mutans can also be present in this initial colonizing community, albeit in varying numbers, which is dependent on the host diet [13], [14]. However, environmental changes, such as frequent consumption of sucrose, dramatically influence biofilm development by providing a substrate for EPS production on the pellicle surface [4], [11], [15]. Glucosyltransferases (Gtfs) from S. mutans are constituents of the pellicle and are capable of synthesizing glucans in situ from sucrose [16], [17]. The surface-formed polymers provide bacterial binding sites for subsequent colonization and local accumulation of S. mutans (through several membrane-associated glucan binding proteins) and other organisms [18], [19]. Gtfs also bind many oral bacteria, even those that do not synthesize Gtfs [17], [20], [21], thereby converting them into de facto glucan producers [17]. Thus, a bacterial product in the pellicle and on microbial surfaces provides the early foundation of the EPS-rich matrix (and binding sites) [4], [22], thereby dramatically influencing the trajectory of virulent biofilm formation. In parallel, sucrose can also be fermented to produce acids within biofilms. Other fermentable carbohydrates (e.g. glucose, fructose, and maltose) can also be converted into acids, but rarely serve as a substrate for EPS synthesis [4], [23]. The acidification of the EPS-rich matrix favors the growth of the microorganisms that are tolerant to acidic stress and promotes demineralization of the adjacent tooth enamel, leading to the clinical onset of cavitation [4], [24], [25]. Cariogenic biofilms are therefore comprised of mixed microbiota in vivo bound by a common matrix; all of the microorganisms are usually potent producers of acid and are acid-tolerant, which creates a highly adhesive, cohesive, and acidic milieu. How the matrix mediates the construction of the 3D biofilm architecture and establishes spatial heterogeneities, and why acids accumulate within oral biofilms, remain intriguing mysteries, particularly when there is an abundance of buffering saliva that surrounds the teeth and would theoretically be capable of neutralizing these acids. Matrix constituents, such as EPS, could affect the diffusion of substances in and out of the biofilm, perhaps helping to create a diverse range of microenvironments within the biofilm [26]. However, the knowledge of how polysaccharides are assembled three-dimensionally is limited, particularly due to difficulties in studying the matrix of intact biofilms. The polysaccharides undergo structural modifications resulting from the effects of glucanohydrolases, and may not be evenly or homogeneously distributed within intact plaque-biofilms [4], [27]. Prior investigations that have attempted to characterize the unique microenvironments within biofilms have relied heavily on the use of invasive techniques (e.g. using probes or microsensors) [28], [29], which alter the complex structure and physical properties of the matrix, complicating the interpretation of data. Thus, the precise role of EPS in the formation and structure of acidic environments within biofilms remains largely unexplored [4], [5]. In the present study, we focused on understanding the function of the matrix associated with (1) the construction of the 3D biofilm architecture, and (2) the modulation of the microenvironmental pH in the resultant milieu within a mixed-species system. To achieve these goals, we devised novel approaches by incorporating a fluorophore or fluorescent pH indicator during synthesis of the matrix. This technique allowed us to follow the development of the intricate matrix structure in a sequential fashion and simultaneously map the spatial distribution of pH within the undisturbed 3D biofilm architecture. Results/Discussion Our results illustrate the interaction between the architecture of the biofilm matrix and the heterogeneity of the microenvironment present within mixed-species biofilms. The EPS-rich matrix facilitates the formation of punctate areas of low pH values within intact biofilm, many that are located in close proximity to the apatite-surface. The resultant milieu favors the selection of acidogenic, aciduric, and glucan-binding pathogens (e.g. S. mutans), and creates demineralizing sites at the surface of biofilm attachment, thereby shifting from a non-virulent to a virulent phase. The assembly of the EPS matrix in 3D We examined over time the morphogenesis of mixed-species biofilms on saliva-coated apatitic (sHA) discs. We focused on the development of the architecture in three-dimensions (3D), which is associated with the early accumulation of EPS on the sHA surface and the subsequent construction of an EPS-rich matrix. The formation and the sequence of assembly of the EPS matrix are heavily influenced by the type of sugars included in the test system (Figure 1A). Following the introduction of 1% sucrose, a heterogeneous layer of EPS is formed on the sHA surface, and localized accumulations of microorganisms are observed on the EPS surface after 43 h (Figure 1Ag; see arrows). After 67 h of development (Figure 1Ah), the cells are densely packed with EPS, forming a 3D bacterial structure (termed EPS-microcolony complex; highlighted in Figure 1Ah). After 115 h, the size of these bacterial structures had increased in several dimensions; they became more enmeshed in, and surrounded by, the EPS-matrix as time elapsed (Figure 1Ai). Two distinct locations of EPS-microcolony complexes were observed: those attached to the EPS layer formed on the sHA (Figure 1Ah), and those tethered to the body of the matrix, near the fluid interface (highlighted in Figure 1Ai). These observations, made when sucrose is abundant (1%) in the test system, are in marked contrast from those made when sucrose was limited (0.1%) or was replaced completely by glucose. A thin EPS layer on the sHA surface, and a few small EPS-attached microcolonies (at later stages) were detected in biofilms formed with 0.1% sucrose (Figure 1Ad–f). When 1% glucose was present, neither an EPS layer nor 3D bacterial islets were assembled. Rather, a thin and mostly homogenous accumulation of bacterial cells adherent to the sHA surface was observed (Figure 1Aa–c). The close-up images reveal that the structural organization of the accumulated cells lacking an EPS matrix is based primarily on cell to cell binding (co-aggregation). 10.1371/journal.ppat.1002623.g001 Figure 1 Dynamics of the morphogenesis, 3D architecture development and microbial population shifts of mixed-species biofilms. (A) Representative 3D rendering images of mixed-species biofilms at distinct time points. Images (a–c) represent biofilms formed after the introduction of 1% (w/v) glucose. Images (d–f) represent biofilms formed in the presence of 0.1% (w/v) sucrose. Images (g–i) represent biofilms formed after the introduction of 1% (w/v) sucrose. The EPS channel is depicted in red, while the cells are depicted in green. At the upper left of each panel, the two channels are displayed separately, while the merged image is displayed at the bottom right. Lateral (side) views of each biofilm are displayed at the bottom left, while a magnified (close-up) view of each small box depicted in the merged image is positioned in the upper right corner of each panel. (B) The data are mean values ± s.d. (n = 30). The asterisks (*) indicate that the each of the values (EPS and bacterial biomass) in the 1% sucrose group are significantly different from others (P 0.05). Introduction of 1% glucose (in the absence of sucrose) resulted in a population dominated by S. oralis throughout the biofilm development period (Figure 1B-3; P 80% of the total cells are co-localized with EPS at 115 h (Figure 4B), as an increasing amount of the bulk of biofilm is composed of polysaccharides (congruent with our confocal images). We suggest that individual microcolonies encased in polysaccharides may serve as architectural units that become connected during biofilm construction, forming compartmentalized networks that confer highly heterogeneous yet cohesive environments within the 3D architecture. The adhesion between the bacterial cell and the EPS-matrix may be mediated in part through cell-surface glucan-binding protein C (GbpC) and possibly GbpB, both of which are expressed by S. mutans [4], [43]. Secreted (extracellular) GbpA and GbpD may also bind to and provide cross-linking between glucans on the sHA surface and those on bacterial surfaces [4], [43], thus contributing to the maintenance of the 3D architecture. Furthermore, extracellular DNA (eDNA) may have a structural function in the extracellular matrix. eDNA enhances S. mutans adhesion and surface aggregation [44] and could be incorporated during EPS-rich matrix development by binding to bacterial cells and to exopolysaccharides [23], possibly contributing to the formation of microcolonies. The assembly of EPS-matrix changes the biofilm architecture in 3D while the bacterial populations are dynamically shifted towards dominance of S. mutans, a Gtf-producing, glucan-binding and aciduric/acidogenic pathogen. Concomitantly, EPS-microcolony complexes are formed in specific regions within the biofilms (bound to the sHA surface and tethered to the matrix) via localized cell to matrix adhesions, while other areas are devoid of such structures. This highly heterogeneous architecture and assembly of a variety of bacterial islets, particularly those in close proximity to sHA, may create ecological and environmental niches that are essential for biofilm virulence. Relevance of biofilm-matrix architecture in virulence Acid production alone may not be the key factor in the expression of virulence, but rather how and where acidic microenvironments are formed and maintained within the biofilm. Results from previous studies reveal average pH values in biofilms. In contrast, our data show punctate areas of low pH values that are heterogeneously distributed within complex 3D biofilm architecture. Creation of localized acidic microenvironment The in situ pH mapping of the biofilm shows spatially heterogeneous pH microenvironments throughout the 3D biofilm architecture, despite the presence of buffered neutral pH in the external environment (Figure 5A). The darker areas in the pH profile represent acidic pH values, whereas the brighter areas represent pH values that are close to neutral (indicated by the pH-visualization bar in Figure 5A). 10.1371/journal.ppat.1002623.g005 Figure 5 Three-dimensional pH mapping of intact mixed-species biofilm. (A) This figure displays representative images of the pH profile throughout the biofilm 3D architecture and the spatial distribution of pH within the selected EPS-microcolony complex in mixed-species biofilms formed after introduction of 1% sucrose. Dark areas are indicative of regions of low pH, while white or light areas are indicative of regions of pH that are close to neutral, as indicated by the scale bar. The EPS channel is depicted in red, while the cells are depicted in green. Arrows indicate various acidic pH regions within the microcolony and at the sHA interface. Red arrows indicate acidic pH regions (i) across the structure and (ii) at the microcolony/sHA interface. Yellow arrows indicate pH close to neutral at the microcolony/fluid phase interface. White arrows indicate the corresponding pH landmarks in the merged panel. (B) This panel shows temporal changes of in situ pH across the EPS-microcolony complex after the biofilm was exposed to sodium phosphate-based buffer (pH 7.0) for a total of 120 min. Environments with low pH values were found only in the interior of the EPS-microcolony complexes within the biofilm (highlighted in Figure 5A). A close-up view of the pH map of these bacterial islets reveals three distinct pH profiles (see arrows; Figure 5A) as follows: (1) regions that display pH values that are close to neutral at the microcolony/fluid phase, (2) a range of acidic “pockets” across the structure (from the sHA surface to the fluid phase interface), and (3) acidic pH at the microcolony/sHA interface. The presence of discrete regions containing low pH (4.5 to 5.5) indicates that the acids accumulated and confined in these particular areas are not rapidly neutralized when incubated in neutral pH (7.0) buffer; more than 120 minutes of exposure are required before neutralization occurs (Figure 5B). The presence of acidic regions at the microcolony-sHA interface has direct implications for cariogenesis. Acids present at the surface of the biofilm attachment increase local demineralization of the hydroxyapatite mineral, which is the main constituent of the tooth enamel [45]. Therefore, cross-sectional images of the biofilm at the sHA surface and pH mapping in the same field of view were acquired to illustrate the spatial distribution of surface-attached microcolonies and pH values (Figure 6A). It is readily apparent that the presence of surface attached-microcolonies (white line) corresponded with acidic pH on the sHA surface (darker areas highlighted with yellow lines). The pH measurements taken by ratiometric imaging analysis show that the pH at the microcolony/sHA interface was below 5.5 (regarded as the critical pH for rapid dissolution of enamel), and the pH value was significantly more acidic compared to areas without such particular 3D structures (highlighted in orange in the pH map image) (P<0.05; Figure 6B). 10.1371/journal.ppat.1002623.g006 Figure 6 In situ pH mapping at the EPS-microcolony complex and sHA interface. (A) This figure shows representative cross sectional images (bird's eye view) at the sHA surface. White marks indicate the areas with surface-attached microcolonies (in the bacteria channel; depicted in green) while yellow marks show the corresponding area in the pH channel; the orange mark indicate an area without surface-attached microcolony. Dark areas are indicative of regions of low pH, while white or light areas are indicative of regions of pH that are close to neutral. (B) This table shows the pH values at the sHA interface for areas with and without surface-attached microcolonies after 30 min incubation in sodium phosphate-based buffer (pH 7.0). The data shown are mean values ± s.d. (n = 90). The asterisks (*) indicate that the differences between the two conditions are statistically significant (P<0.05, Tukey-Krammer HSD). The presence of localized and elevated concentrations of acids at the surface of biofilm attachment has clinical relevance, as it could explain the heterogeneous pattern of carious lesions during the onset of this disease in humans. Dental caries is characterized by the temporal and localized/punctate appearance of regions of demineralization on the tooth enamel under plaque, clinically known as “white spots” [46]. These “white spots” eventually coalesce into a broader lesion, rather than a diffuse and homogeneous layer of erosion. Thus, the appearance of the lesions and the level of demineralization may be associated with the size of the surface-attached EPS-microcolony complexes. Correlation analysis showed that there is a linear relationship between low pH on the sHA surface and the diameter or height of these bacterial islets (Figure 7). 10.1371/journal.ppat.1002623.g007 Figure 7 Relationship of in situ pH at sHA surface with size of surface-attached EPS-microcolony complexes. This figure depicts mixed-species biofilms (67 h) formed after the introduction of 1% (w/v) sucrose, which were processed for measurement of in situ pH at the interface between sHA surface and microcolonies. All the pH values were sorted by (A) the diameter and (B) height (distance from surface to fluid-phase) of the surface-attached microcolonies. The data shown are values from three separate experiments (n = 90). Correlation analysis showed that there is a linear relationship between pH and microcolony diameter (Pearson's test, P<0.0001, r-square 0.259) and height (P<0.0001, r-square 0.367). Biofilms formed with the gtfB null mutant (lacking a 3D matrix and EPS-microcolony complexes) were readily neutralized after incubation in neutral pH buffer and failed to generate any acidic microenvironments (Figure S3). Interestingly, biofilms formed with the gtfC null mutant contained isolated and small EPS-microcolony complexes, which displayed some acidic pockets, although these were neutralized within 30 minutes (Figure S3). Collectively, the data clearly demonstrate that there is a substantial limitation to diffusion, which is associated with the assembly of EPS-rich matrix and the size of EPS-enmeshed microcolonies. Their presence could trap acid at the sHA surface and/or restrict access of neutralizing buffer (e.g. saliva) to areas of low pH within the biofilm. The data are consistent with Tatevossian's findings [47], which show that the concentration of solutes in human dental plaque fluid differs markedly from those observed in saliva. The precise mechanisms involved in limiting diffusion are unclear. It is possible that the presence of glucans produced by oral streptococci limit diffusion of charged ions in and out of the plaque-biofilm, whereas uncharged substances, such as sucrose, may diffuse readily [45], [47], [48]. In contrast, others have noted that EPS has little effect on acid diffusion when using artificial plaque and in vitro modeling [49], [50]. However, none of these studies evaluated intact biofilms, as investigated here. Our data show the importance of how the matrix is assembled three-dimensionally and how it is spatially arranged with the bacterial cells to act as a diffusion-limiting physical barrier. The presence of a high density of acidogenic and aciduric bacteria within these structures can facilitate the conversion of sucrose (and other fermentable carbohydrates) into acids via the glycolytic pathway. Furthermore, degradation of extracellular soluble glucans by S. mutans dextranase (also up-regulated within biofilm) and degradation of intracellular polysaccharides increases the availability of fermentable sugars (e.g. maltose and glucose) [4], [23], [31]. The metabolism of these carbohydrates could also contribute to the acidification of the resultant 3D milieu. The presence of alkali-generating bacteria in biofilms may also influence the local pH [51]. The use of metabolic activity markers could help to elucidate whether acidification is an active process or a secondary consequence of cell numbers (density). It is apparent that the combination of the physical and metabolic factors could, at least in part, explain the spatial distribution of punctate acidic regions in our in vitro biofilms and the low fasting pH values observed in plaque from caries-active individuals. The formation of a heterogeneous acidic environment is biologically relevant, as pH changes could trigger dynamic shifts in the microbial population according to the ecological plaque model [11]. Niches with low pH values would activate the acid stress adaptive responses of particular organisms clustered in these EPS-sheltered acidic pockets, favoring survival/growth of aciduric and glucan-binding pathogens, such as S. mutans (initially in low-numbers) [24], [25]. Local pH concentrations would also affect the spatial distribution of the bacterial species throughout the 3D biofilm architecture. The positioning of the microorganisms and their stress response mechanisms may correlate with their susceptibility to, or affinity for, low pH environments. In addition, persistent low pH niches may also affect the expression of the gtf genes, as expression of gtfBC is induced in response to acidification and the presence of excess metabolizable carbohydrates [52]. These genes would therefore likely be up-regulated locally, coinciding with areas that are undergoing active matrix remodeling and acidogenesis. Creation of protective microenvironments The microorganisms within biofilms display increased resistance to antimicrobials in comparison to their planktonic counterparts [1], [5], [53]. This phenomenon is generally associated with poor antimicrobial penetration (due to the presence of an extracellular matrix), activation of adaptive stress responses (including production of persister cells and induction of quorum sensing mechanisms), and physiological heterogeneity within the biofilm population [26], [54]. However, the importance of spatially heterogeneous environments and spatially complex cell-matrix arrangements in bacterial antimicrobial resistance within biofilms has received limited attention. The construction of bacterial islets that are enmeshed and surrounded by an EPS-rich matrix could create localized areas of microbial resistance to antimicrobials within the biofilm. Therefore, we evaluated the viability of two distinct groups of bacterial cells within the same intact biofilm system in response to the immediate local effects of chlorhexidine (CHX) exposure (by time-lapsed measurements). We examined (1) cells within EPS-microcolony complexes, and (2) cells outside (or not forming) such structures (Figure 8). CHX is a broad spectrum antibacterial agent commonly used in mouthwash (at concentrations between 0.12–0.2%) to suppress the levels of mutans streptococci in the oral cavity [55]. The overlay images of live (in green) and dead (in red) cells after exposure to 0.12% CHX are shown in Figure 8A. Clearly, the inside of the EPS-enmeshed microcolony (Figure 8A-1) contains a greater proportion of live cells than those residing outside the structured communities over time (Figure 8A-2). 10.1371/journal.ppat.1002623.g008 Figure 8 Time-lapse imaging of bacterial viability after exposure of the biofilm to 0.12% (v/v) chlorhexidine (CHX). Panel (A) shows the relative populations of live (SYTO 9-labeled; depicted in green) and dead (propidium iodide-labeled; depicted in red) cells over time in the mixed-species biofilm formed with the parental strain (A-1; cells within EPS-microcolony complex and A-2; cells outside EPS-microcolony complex) and the ΔgtfBC::kan mutant (A-3) after exposure to CHX. Panel (B) shows the survival rate of live cells. The data shown are mean values ± s.d. (n = 6). The asterisks (*) indicate that the differences between the conditions tested (cells within EPS-microcolony complex vs. cells outside EPS-microcolony complex or cells not forming such structures, i.e. biofilm formed with ΔgtfBC::kan) are statistically significant (P<0.05). The survival rate of each group of bacterial cells was also calculated using the confocal images (Figure 8B). The data show that less than 10% of the total cells within the EPS-microcolony complex were killed after 15 min exposure. In contrast, more than 70% of the cells outside the structure were not viable (P<0.05). In the absence of the matrix, cells were highly susceptible to CHX. Mixed-species biofilms formed with the gtfBC null mutant strain were rapidly killed by CHX (Figure 8A-3; Figure 8B). Similarly, bacterial cells that generally do not organize into EPS-microcolony complexes, such as those in biofilms grown in 1% glucose, were also highly susceptible to killing by CHX (60–75% of the cells were dead after 15 min exposure to CHX; data not shown). Chlorhexidine (CHX) is a cationic bisbiguanidine [55]. The presence of negatively charged EPS (possibly due to association with lipoteichoic acids) could retard the diffusion of positively charged CHX molecules into biofilms [48], [53], [56]. However, our data show that antimicrobial resistance is not a general attribute associated with the simple physical presence of EPS, but rather with the way the matrix is assembled with the bacterial cells, which likely restricts access of CHX into EPS-microcolony structures. Furthermore, the cells clustered within EPS-microcolony complexes may develop direct antimicrobial resistance in response to cell density and/or the presence of acidic microenvironments in these bacterial islets [26], [54]. These observations support the concept that antimicrobial resistance is a site-specific property within the greater biofilm architecture, although we were unable to explore the additional possibilities described above. Results from a recent study [57] demonstrated the importance of a structured environment (using fabricated interconnected chambers within a microfluidic device) for rapid evolution of bacterial antibiotic resistance via the creation of steep chemical gradients. Our findings agree with the modeling of Zhang et al. [57], recapitulating spatial heterogeneities in our de facto biofilm system. Simply put, the EPS-matrix creates highly compartmentalized, yet interconnected and cohesive microenvironments. As a result, acidic regions and niches of antimicrobial resistance are created locally, demonstrating the importance of the 3D architecture in the evolution of virulent oral biofilms. Conclusion In this study, we show that the concerted actions of S. mutans-derived GtfB and GtfC, in the presence of sucrose, form a heterogeneous layer of EPS on the sHA surface. The surface-formed EPS promotes local accumulation of bacteria. The continuous production of EPS in situ coats the organisms and they rapidly aggregate, forming densely packed bacterial islets (termed EPS-microcolony complexes). Over time, these 3D structures become interconnected through surrounding EPS. The sequential accumulation of EPS on sHA and bacterial surfaces assembles an intricate and complex 3D matrix that leads to a highly compartmentalized architecture within an intact mixed-species biofilm. The relevance of the matrix 3D architecture in biofilm virulence is revealed by our novel pH mapping approach, which uses a pH marker incorporated directly into the matrix. We illustrate spatially heterogeneous pH microenvironments manifested by “acidic pockets” found in the interior of the EPS-enmeshed bacterial islets only, despite exposure to a neutral buffer. This observation is clinically relevant in the context of the oral cavity in which saliva at neutral pH is abundant. The low pH niches favor acid-tolerant organisms, ensuring continued localized acid production; an EPS-rich environment promotes accumulation of the glucan-binding pathogen S. mutans. The environmental changes, associated with the matrix architecture, shift the microbial balance from a non-virulent to a virulent condition. Moreover, regions of acidic pH (<5.5) were found only in the areas where surface-attached EPS-enmeshed microcolonies were present on the sHA surface. The presence of an elevated concentration of acids at the apatitic surface favors local demineralization (while preventing remineralization), thereby facilitating development of carious lesions at these sites. Furthermore, the bacterial cells residing within EPS-microcolony complexes throughout the biofilm architecture are locally sheltered against a potent antimicrobial agent (CHX). We demonstrated that S. mutans does not always need to dominate within plaque to influence virulence, because the Gtfs released, even at the time of biofilm initiation, adhere to the pellicle and other organisms, mediating the construction of the EPS-rich matrix. Moreover, the expression of GtfB and GtfC is enhanced in the presence of other oral bacteria. Results from previous studies have linked Gtfs to biofilm formation (using standard genetic and/or simple cell staining assays in mono-species biofilms) and expression of virulence (using rodent models) [4]. Our data illuminate the functional roles of GtfB- and GtfC-derived EPS in the basic biology of biofilms. We show how the assembly of a 3D EPS-matrix architecture helps to create spatial heterogeneities, and facilitates the formation of a variety of structured acidic microenvironments within a mixed-species biofilm. Clearly, the data from our study offer new avenues for further elucidation of how cell to matrix interactions govern the creation of ecological and environmental niches that are essential for expression of virulence by an oral biofilm. The information generated here may pave the way for designing future studies aiming to develop novel methods that better prevent the formation and/or promote the disruption of virulent dental plaque. Furthermore, we introduce novel approaches to study the dynamics of biofilm construction and heterogeneity within the intact 3D architecture. Our observations have broad applicability beyond the mouth, because the imaging techniques introduced could be adapted for use in other biofilm systems in which polymers are formed during matrix development, and varied microenvironments are created. Materials and Methods Bacterial strains Streptococcus mutans UA159 serotype c (ATCC 700610), a proven virulent cariogenic dental pathogen selected for genomic sequencing [58], Actinomyces naeslundii ATCC 12104, and Streptococcus oralis ATCC 35037 were used to generate mixed-species biofilms. We selected S. oralis and A. naeslundii in addition to S. mutans because these three organisms are found in the supragingival plaque of humans [59]. S. oralis binds to the pellicle and is one of the most commonly detected pioneer colonizers of the saliva-coated tooth surface [59], [60] in the absence of sucrose; strain ATCC 35037 also produces soluble glucans from sucrose and is acid-tolerant [60]. A. naeslundii is also detected during the early stages of plaque formation and may be associated with development of root caries; strain ATCC 12104 is acidogenic and produces fructans [61]. Furthermore, gtfB, gtfC, and gtfBC null mutants (ΔgtfB::kan, ΔgtfC::kan, and ΔgtfBC::kan, respectively), constructed from the parental S. mutans UA159 strain, were generated by standard allelic replacement with a non-polar kanamycin resistance marker and verified by DNA sequence analysis. These strains were kindly provided by Dr. Robert A. Burne (Department of Oral Biology, University of Florida, Gainesville, Florida) [22]. The strains were checked via PCR before use. The cultures were stored at −80°C in tryptic soy broth containing 20% glycerol. Mixed-species biofilm model The biofilm method is based on a batch culture approach using saliva-coated hydroxyapatite (sHA) discs, which was designed to mimic the formation of biofilms according to the “ecological plaque-biofilm” concept [11], as recently described by Koo et al. [22]. S. mutans UA159, A. naeslundii ATCC 12104, and S. oralis ATCC 35037 cells were grown in ultrafiltered (10 kDa molecular-weight cut-off membrane; Prep/Scale, Millipore, MA) buffered tryptone-yeast extract broth (UFTYE; 2.5% tryptone and 1.5% yeast extract, pH 7.0) with 1% glucose at 37°C and 5% CO2 to mid-exponential phase; the ultrafiltration process removes high-molecular weight mannans and other polysaccharides that could interfere with our analyses. These bacterial suspensions were then mixed to provide an inoculum with a defined microbial population of S. mutans (102 CFU/ml), A. naeslundii (106 CFU/ml), and S. oralis (107 CFU/ml), which are critical for the reproducibility of our model [22]. Biofilms were formed on hydroxyapatite discs (1.25 cm diameter, surface area of 2.7±0.2 cm2, Clarkson Chromatography Products, Inc., South Williamsport, PA) coated with filter-sterilized clarified whole saliva [22], which were placed in a vertical position using a custom-made disc holder (Figure S4). The mixed population of S. mutans, A. naeslundii, and S. oralis was inoculated in 2.8 ml of UFTYE with 0.1% (w/v) sucrose, and incubated at 37°C and 5% CO2. During the first 19 h, the organisms were grown undisturbed to allow initial biofilm formation. At this point (19 h), the culture medium was replaced with fresh medium and the biofilms were grown until 29 h for establishment of the initial biofilm community [22]. At 29 h of biofilm growth, the biofilms were transferred to UFTYE containing 1% (w/v) sucrose or 1% (w/v) glucose to induce environmental changes to simulate a cariogenic challenge, while an additional set of biofilms was grown in UFTYE with 0.1% sucrose. The culture medium was then replaced twice daily (8 am and 6 pm) until the end of the experimental period (115 h). The pH of spent culture medium was measured daily using a standard pH electrode. The biofilms were analyzed at specific time points (29, 43, 67, 91, and/or 115 h) by means of confocal imaging/fluorescence, biochemical, reverse transcription quantitative PCR (RT-qPCR), and multi-dimensional protein identification technology (MudPIT) assays (see experimental design scheme in Figure S4). Furthermore, mixed-species biofilms were also formed in the presence of S. mutans UA159 mutant strains defective in gtfB and/or gtfC because they are critical for insoluble EPS synthesis and expression of virulence in vivo [4], [41], [62]. The gtfD mutant was not included because it did not have significant effect on matrix formation and biofilm architecture. All the gtf mutants used in this study exhibited similar planktonic growth rates (vs. UA159) as monitored (at OD600 nm) using an automated turbidimetric Bioscreen C MBC system (Oy Growth Curves Ab Ltd, Helsinki, Finland). Laser scanning confocal fluorescence microscopy (LCSFM) imaging of biofilm matrix The sequential assembly of the matrix was determined by directly incorporating a fluorescent marker during synthesis of the extracellular polysaccharide (EPS)-matrix, allowing examination of the three-dimensional (3D) structure within intact biofilms [63], [64]. Briefly, 1 µM Alexa Fluor 647-labeled dextran conjugate (molecular weight, 10 kDa; absorbance/fluorescence emission maxima of 647/668 nm; Molecular Probes, Invitrogen Corp., Carlsbad, CA) was added to the culture medium from the beginning of and during the development of the biofilms. This technique is based on the observation that (fluorescently-labeled) dextran serves as a primer and acceptor for Gtfs (particularly GtfB), and is incorporated into newly formed glucan by the exoenzyme during synthesis of the EPS-matrix over the course of biofilm development [63]; it does not stain the bacterial cells at the concentration used in this study [63]. All the bacterial species in the biofilms were labeled by means of SYTO 9 green fluorescent nucleic acid stain (485/498 nm; Molecular Probes) using standard protocols [63], [64]. The imaging was performed using an Olympus FV 1000 two photon laser scanning microscope (Olympus, Tokyo, Japan) equipped with a 10× (0.45 numerical aperture) or 25× LPlan N (1.05 numerical aperture) water immersion objective lens. The excitation wavelength was 810 nm, and the emission wavelength filter for SYTO 9 was a 495/540 OlyMPFC1 filter, while the filter for Alexa Fluor 647 was an HQ655/40M-2P filter [22]. Each biofilm was scanned at 5 positions randomly selected at the microscope stage [65], and confocal image series were generated by optical sectioning at each of these positions. Three independent biofilm experiments were performed, and 10 image stacks (512×512 pixel for quantification or 1024×1024 pixel for visualization in tagged image file format) were collected for each experiment. Computational analyses of the confocal biofilm images The confocal images were analyzed using software for simultaneous visualization and quantification of EPS and bacterial cells within intact biofilms [64]. Amira 5.0.2 (Mercury Computer Systems Inc., Chelmsford, MS) was used to create 3D renderings of each structural component (EPS and bacteria) of the biofilms for visualization of the morphology and 3D architecture as detailed previously [64], [65]. COMSTAT and newly developed DUOSTAT (both are available as free downloadable software at http://www.imageanalysis.dk) were used for quantitative analysis. COMSTAT and DUOSTAT, written as scripts for MATLAB 5.3 (The Mathworks, Natick, MA, USA) equipped with the Image Processing Toolbox (see details at http://www.imageanalysis.dk), converts pixels from confocal image stacks into numerical values, facilitating quantitative characterization of each structural component within 3D biofilm images [64]. We used COMSTAT to calculate the biomass, number, and size (volume, diameter, and height) of microcolonies, as detailed elsewhere [65]. DUOSTAT was developed to calculate the co-localization of two biofilm components (e.g. EPS and bacterial cells) throughout the 3D biofilm architecture [64]. The software overlays the confocal image data sets (one for total bacteria and another for EPS) from the same biofilm, and calculates the proportion of total bacteria image pixels that are co-localized with EPS [64]. Non-invasive 3D in situ pH measurement and visualization The ability to characterize microenvironments (e.g. spatial pH distribution) within biofilms without disrupting their architectural integrity is a major technical challenge. We used a fluorescent pH indicator, Lysosensor yellow/blue (Molecular Probes), to determine the in situ pH in our biofilm model. Lysosensor yellow/blue is conjugated with dextran (10 kDa MW), which allows the labeled dextran to be directly incorporated into the EPS-matrix [63], as described previously. Once incorporated into the biofilm matrix, the pH values can be measured based on fluorescence intensity ratios of the dual-wavelength fluorophore [66]. Lysosensor yellow/blue exhibits a dual-emission spectral peak (fluorescence emission maxima 452 nm and 521 nm) that is pH dependent [66]. The pH measurements are based on the principle that protonation of the fluorophore shifts its emission spectra, such that increasing ion concentrations increase fluorescence intensity at one wavelength, while decreasing it at the other [67]. The fluorescence intensity of both emission wavelengths and the ratio of fluorescent intensity (I450/I520) within each biofilm image was measured using Image J 1.44 and its calculation tools (e.g. channel 1 divide by channel 2 as detailed in http://rsbweb.nih.gov/ij/download.html). Titration curves of ratios versus pH (ranging from 3.5 to 7.0) were calculated as described previously [67], and were used to convert Lysosensor yellow/blue emission (fluorescence intensity) ratios to pH values. The intensity ratio of both wavelengths serve as a quantitative measure of pH, and Lysosensor yellow/blue is particularly suitable for acidic environments (low sensitivity maximum; pKa∼4.2), as expected in our biofilm model [66]. The experimental protocol and titration curve of ratios versus pH are provided in the supporting information (Protocol S1 and Figure S5). The confocal images of Lysosensor-incorporated biofilms were acquired before and after incubation in Na2HPO4-citric acid buffer at pH 7.0. The images were acquired after 10, 30, 60, and 120 min of incubation in the neutral buffer to evaluate the spatial distribution of pH throughout the intact 3D biofilm architecture. For pH measurement, the ratio of fluorescence intensity of selected areas within each biofilm image were converted to pH values using the titration curves (as described in Protocol S1 and Figure S5) and Image J software (http://www.uhnresearch.ca/facilities/wcif/PDF/ImageJ_Manual.pdf). For visualization of in situ pH distribution within the 3D biofilm architecture, the fluorescence intensity ratios (corresponding to the pH values between 7.0 and 3.5) of all confocal images were reconstructed using Image J, then Amira. The fluorescence intensity was converted into grayscale using the Amira tool-box to correlate with the pH range from 7.0 (white) to 3.5 (black) (Protocol S1). Unlabeled biofilms were also imaged to check whether the autofluorescence of each biological component of the biofilm (i.e. bacterial cells and glucans) would interfere with pH quantification at the wavelengths and laser intensity used in our study; there was no interference with the measurements based on our image analysis. Microbiological analysis of the mixed-species bacterial population The biofilms were removed at specific time points (Figure S4) and homogenized by sonication in sterile 0.89% (w/v) NaCl solution (30 sec pulse at an output of 7W; Branson Sonifier 150, Branson Ultrasonics, Danbury, CT); the sonication procedure does not kill any of the bacterial species used in this study [22]. The homogenized suspension was used to determine the number of viable cells by plating on blood agar using an automated EddyJet Spiral Plater (IUL, SA, Barcelona, Spain) to determine the number of viable cells (total number of colony-forming units per biofilm). The three species were differentiated by observation of colony morphology in conjunction with microscopic examination of cells from selected colonies [68]. In addition, the bacterial populations were also assessed by qPCR using species specific primers for S. mutans [69], A. naeslundii, and S. oralis. The A. naeslundii (5- CCTCTGGCTTAACTGGGG -3 and 5- CATTCCACCGCTACACCA -3) and S. oralis (5- CCGCATAAGAGTAGATGTTG -3 and 5- GCCTTGGTGAGCCGTTAC -3) species-specific primers were designed using Beacon Designer 2.0 software (Premier Biosoft International, Palo Alto, CA, USA). Bacterial cell quantification using standard curves based on the genome size of each organism was performed as described previously [69]. The data was expressed as total numbers, and as proportion of each of the bacterial species within the biofilm. S. mutans gene expression analysis using multiplex RT-qPCR We performed RT-qPCR to measure the expression profiles of specific genes directly associated with extracellular polysaccharide matrix development, including gtfB, gtfC, gtfD, fruA, ftf, and dexA. RNA was extracted and purified using protocols optimized for biofilms [70]. The RNA integrity number (RIN) for all of our samples was ≥9.0, as determined by on-chip capillary electrophoresis with the Agilent 2100 Bioanalyzer (Agilent Technologies, Inc., Santa Clara, CA). Briefly, cDNAs were synthesized using 0.5 µg of purified RNA and the BioRad iScript cDNA synthesis kit (Bio-Rad Laboratories, Inc., Hercules, CA). To check for DNA contamination, purified total RNA without reverse transcriptase served as a negative control. The resulting cDNA and negative controls were amplified by a Bio-Rad CFX96 system (Bio-Rad Laboratories, Inc., CA) using specific primers and TaqMan probes (Table S2) and iQ Multiplex Powermix (Bio-Rad Laboratories). A standard curve was plotted for each primer set as detailed elsewhere [71]. The standard curves were used to transform the critical threshold cycle (Ct) values to relative numbers of cDNA molecules. Comparative expression was calculated by normalizing each gene of interest to the 16S rRNA gene, which served as a reference gene [71]. S. mutans protein expression analysis by MudPIT We also examined whether changes in S. mutans gene expression would reflect the levels of their corresponding protein products in biofilms. The S. mutans protein expression profile in whole biofilms was analyzed by MudPIT and specific database search algorithms described previously [72]–[75]. Briefly, biofilms at 67 and 115 h of growth were homogenized by sonication (30 sec pulse at an output of 7W; Branson Sonifier 150) in the presence of protease cocktail inhibitor (Roche). After homogenization, aliquots were taken for bacteria quantification by qPCR and total protein quantification (by acid digestion followed by ninhydrin assay [76]); the protease cocktail inhibitor did not affect the ninhydrin assay as determined experimentally. Total protein was extracted, precipitated, and digested with trypsin, as detailed elsewhere [73], [74]. MS/MS spectra were searched with the ProLuCID algorithm [75] or the Sequest algorithm [77] against an NCBI-RefSeq S. mutans and S. oralis database (01/01/2010) concatenated to a decoy database in which the sequence for each entry in the original database was reversed [73]. All searches were parallelized and performed on a Beowulf computer cluster consisting of 100 1.2 GHz Athlon CPUs. ProLuCID results were assembled and filtered using the DTASelect (version 2.0) program [78]. The false positive rates are estimated by the program from the number and quality of spectral matches to the decoy database. We focused on proteins encoded by S. mutans UA159, and compared biofilms at 67 h- vs. 115 h-old. The level of protein abundance in mixed-species was calculated by normalizing the spectral count to the relative numbers of S. mutans by multiplying the spectral count values by the ratio of S. mutans CFU to the total CFU in the mixed-species biofilms samples. Susceptibility of EPS matrix to mutanase The EPS matrix formed in mixed-species biofilms was examined for susceptibility to mutanase [α-1,3 glucanase; EC 3.2.1.59], produced from Trichoderma harzianum, as detailed elsewhere [39]. The enzyme (Novo Nordisk, Denmark) was kindly provided by Dr. William H. Bowen (Center for Oral Biology, University of Rochester Medical Center). Biofilm was formed on the sHA surface, and the EPS-matrix was labeled with Alexa Fluor 647-labeled dextran conjugate (Molecular Probes) as described in the previous sections. At 91 h of biofilm development, mutanase (2 units of enzyme) were added to the culture media. At 115 h, biofilms were retrieved and the bacteria cells were labeled with SYTO 9 (Molecular Probes). Biofilms were examined using confocal fluorescence imaging, and subjected to AMIRA/COMSTAT analysis. Cell viability within intact biofilms by time-lapsed killing assay The effects of chlorhexidine (CHX) on the viability of biofilm cells immediately after introduction of the antimicrobial agent were assessed by time-lapsed measurements. The intact biofilms (formed with parental UA159 strain or gtfBC null mutant) were dip washed twice in 0.89% NaCl, then transferred to a Petri dish (Diameter 3.5 cm) containing 2.5 µM SYTO 9 (for labeling live-cells; Molecular Probes) and 2.5 µM propidium iodide-PI (for labeling dead-cells; Molecular Probes) in 4 ml of 0.89% NaCl, and incubated for 30 min at room temperature using standard protocols described by Hope and Wilson [53]. After labeling, confocal images were acquired using an Olympus FV 1000 two photon laser scanning microscope with an 25× LPlan N (1.05 numerical aperture) water-immersion objective lens (Olympus). The excitation wavelength was set at 780 nm and the emission filter for SYTO 9 was the 495/540 OlyMPFC1 filter, while PI was the HQ655/40M-2P filter. Immediately after completion of the first scan, and while keeping the biofilm in the same position, 48 µl of 10% (v/v) CHX solution was carefully added to the 4 ml cell culture dish and mixed to yield a final concentration of 0.12% CHX (v/v). CHX does not react with the dyes used in this study [53]. For the time-lapse series, images of the same field of view were taken at 15, 30, 45, and 60 minutes after CHX exposure. The total biomass of live and dead bacterial cells in each series of confocal images was quantified using COMSTAT. The ratio of live to the total bacteria at each time point was calculated, and the survival rate of live cells (relative to live cells prior to exposure to CHX) was plotted. The survival rate was calculated based on the amount of viable cells over time. The initial number of viable cells immediately before the introduction of CHX (time point 0 min; t 0 min.) was considered to be 100%. The number of viable cells after the introduction of CHX was then determined at each time point and the percent-survival rate was determined by comparing to t 0 min. Thus, the influence of the structural organization of the matrix on the killing effects of CHX can be monitored in situ and in real-time. Statistical analysis An exploratory data analysis was performed to determine the most appropriate statistical test; the assumptions of equality of variances and normal distribution of errors were also checked. The data were then analyzed using ANOVA, and the F test was used to test any difference between the groups (different experimental conditions and distinct time-points within same experimental condition). When significant differences were detected, a pairwise comparison was made between all the groups using Tukey-Kramer HSD method to adjust for multiple comparisons. Pearson's test was also used to evaluate the linear correlation between pH value and microcolony height (or diameter). Software JMP version 3.1 (SAS Institute, Cary, NC, USA) was used to perform the statistical analyses. The level of significance was set at 5%. Supporting Information Figure S1 Representative 3D rendering of mixed-species biofilms formed in the presence of S. mutans UA159 mutant strains. (A) ΔgtfB::kan or (B) ΔgtfC::kan after introduction of 1% (w/v) sucrose. (TIF) Click here for additional data file. Figure S2 Comparison of biofilm architecture by S. mutans alone (single-species) or in the presence of other species (mixed-species) after introduction of 1% sucrose. (A) Representative 3D images of biofilms (and microcolonies) formed by S. mutans alone or mixed with other species; (B) number and volume of microcolonies as determined using COMSTAT (n = 10). (TIF) Click here for additional data file. Figure S3 Mapping of in situ pH of intact mixed-species biofilm formed in the presence of S. mutans UA159 mutant strains. (A) ΔgtfB::kan or (B) ΔgtfC::kan after introduction of 1% (w/v) sucrose. (TIF) Click here for additional data file. Figure S4 Overview of the experimental design of the mixed-species biofilm model. (TIF) Click here for additional data file. Figure S5 Titration curve used to convert Lysosensor yellow/blue emission (fluorescence intensity) ratios to pH values. (TIF) Click here for additional data file. Protocol S1 Detailed methodology for 3D in situ pH mapping. (DOC) Click here for additional data file. Table S1 pH of the culture medium during mixed-species biofilm formation. (DOC) Click here for additional data file. Table S2 Primers and TaqMan probes used for multiplex RT-qPCR assays. (DOC) Click here for additional data file.
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            Demineralization–remineralization dynamics in teeth and bone

            Biomineralization is a dynamic, complex, lifelong process by which living organisms control precipitations of inorganic nanocrystals within organic matrices to form unique hybrid biological tissues, for example, enamel, dentin, cementum, and bone. Understanding the process of mineral deposition is important for the development of treatments for mineralization-related diseases and also for the innovation and development of scaffolds. This review provides a thorough overview of the up-to-date information on the theories describing the possible mechanisms and the factors implicated as agonists and antagonists of mineralization. Then, the role of calcium and phosphate ions in the maintenance of teeth and bone health is described. Throughout the life, teeth and bone are at risk of demineralization, with particular emphasis on teeth, due to their anatomical arrangement and location. Teeth are exposed to food, drink, and the microbiota of the mouth; therefore, they have developed a high resistance to localized demineralization that is unmatched by bone. The mechanisms by which demineralization–remineralization process occurs in both teeth and bone and the new therapies/technologies that reverse demineralization or boost remineralization are also scrupulously discussed. Technologies discussed include composites with nano- and micron-sized inorganic minerals that can mimic mechanical properties of the tooth and bone in addition to promoting more natural repair of surrounding tissues. Turning these new technologies to products and practices would improve health care worldwide.
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              Micro- and Nanotopography Sensitive Bacterial Attachment Mechanisms: A Review

              Bacterial attachment to material surfaces can lead to the development of biofilms that cause severe economic and health problems. The outcome of bacterial attachment is determined by a combination of bacterial sensing of material surfaces by the cell and the physicochemical factors in the near-surface environment. This paper offers a systematic review of the effects of surface topography on a range of antifouling mechanisms, with a focus on how topographical scale, from micro- to nanoscale, may influence bacterial sensing of and attachment to material surfaces. A good understanding of these mechanisms can facilitate the development of antifouling surfaces based on surface topography, with applications in various sectors of human life and activity including healthcare, food, and water treatment.
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                Author and article information

                Journal
                Int J Mol Sci
                Int J Mol Sci
                ijms
                International Journal of Molecular Sciences
                MDPI
                1422-0067
                16 July 2019
                July 2019
                : 20
                : 14
                : 3491
                Affiliations
                [1 ]Ph.D Program in Dental Biomedical Sciences, University of Maryland School of Dentistry, Baltimore, MD 21201, USA
                [2 ]Department of Preventive Dental Sciences, College of Dentistry, Imam Abdulrahman Bin Faisal University, Dammam 34212, Saudi Arabia
                [3 ]Medical Microbiology Department, Health Monitoring Centers, Ministry of Health, Jeddah 21176, Saudi Arabia
                [4 ]Department of Restorative Dental Sciences, College of Dentistry, Imam Abdulrahman Bin Faisal University, Dammam 34212, Saudi Arabia
                [5 ]Department of Advanced Oral Sciences and Therapeutics, Division of Biomaterials and Tissue Engineering, University of Maryland School of Dentistry, Baltimore, MD 21201, USA
                [6 ]Biosystems and Biomaterials Division, National Institute of Standards and Technology, Gaithersburg, MD 20899, USA
                [7 ]Department of Endodontics, The Dental College of Georgia, Augusta University, Augusta, GA 30912, USA
                [8 ]Center for Stem Cell Biology and Regenerative Medicine, University of Maryland School of Medicine, Baltimore, MD 21201, USA
                [9 ]University of Maryland Marlene and Stewart Greenebaum Cancer Center, University of Maryland School of Medicine, Baltimore, MD 21201, USA
                [10 ]Division of Operative Dentistry, Department of General Dentistry, University of Maryland School of Dentistry, Baltimore, MD 21201, USA
                Author notes
                [* ]Correspondence: mmelo@ 123456umaryland.edu ; Tel.: +1-410-706-8705
                Author information
                https://orcid.org/0000-0001-8461-3354
                https://orcid.org/0000-0001-6678-7940
                Article
                ijms-20-03491
                10.3390/ijms20143491
                6679354
                31315225
                4400ac50-4ddd-4a8d-a6c8-f12cbc774390
                © 2019 by the authors.

                Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license ( http://creativecommons.org/licenses/by/4.0/).

                History
                : 11 April 2019
                : 10 July 2019
                Categories
                Article

                Molecular biology
                antibacterial agents,biofilm,dental caries,sealant,quaternary ammonium compounds,streptococcus mutans

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