Structure of a Mature Dental Plaque Biofilm

Dental plaque (see Figure 23-7) is defined clinically as a structured, resilient yellow-grayish substance that adheres tenaciously to the intraoral hard surfaces, including removable and fixed restorations.²⁹ The tough extracellular matrix makes it impossible to remove plaque by rinsing or the use of sprays. Plaque can thus be differentiated from other deposits that may be found on the tooth surface such as materia alba and calculus. Materia alba refers to soft accumulations of bacteria, food matter, and tissue cells that lack the organized structure of dental plaque and are easily displaced with a water spray. Calculus is a hard deposit that forms by mineralization of dental plaque and is generally covered by a layer of unmineralized plaque (Table 23-3).

TABLE 23-3 Differences Between Tooth Deposits

Dental plaque is composed primarily of microorganisms. One gram of plaque (wet weight) contains approximately 10¹¹ bacteria.^339,360 The number of bacteria in supragingival plaque on a single tooth surface can exceed 10⁹ cells. In a periodontal pocket, counts can range from 10³ bacteria in a healthy crevice to >10⁸ bacteria in a deep pocket. Using highly sensitive molecular techniques for microbial identification, it has been estimated that more than 500 distinct microbial phylotypes can be present as natural inhabitants of dental plaque.¹ In fact, this number may be much greater.

Large-scale pyrosequencing of DNA from plaque populations indicates that there may be as many as 19,000 species-level phylotypes (essentially, distinct species in all but name) in dental plaque.¹⁶⁵

Any individual may harbor 150 or more different species. Nonbacterial microorganisms that are found in plaque include archaea, yeasts, protozoa, and viruses.^62,203

Dental plaque is broadly classified as supragingival or subgingival based on its position on the tooth surface toward the gingival margin.

Supragingival plaque typically demonstrates a stratified organization of a multilayered accumulation of bacterial morphotypes (Figure 23-10). Gram-positive cocci and short rods predominate at the tooth surface, whereas gram-negative rods and filaments, as well as spirochetes, predominate in the outer surface of the mature plaque mass.

Figure 23-10 A, 1-day-old plaque. Microcolonies of plaque bacteria extend perpendicularly away from tooth surfaces. B, Developed supragingival plaque showing overall filamentous nature and microcolonies (arrows) extending perpendicularly away from tooth surface. Saliva-plaque interface shown (S). C, Histologic section of plaque showing nonbacterial components such as white blood cells (arrow) and epithelial cells (asterisk), interspersed among bacteria (B).

(Courtesy Dr. Max Listgarten, Philadelphia, PA.)

In general, the subgingival microbiota differs in composition from the supragingival plaque, primarily because of the local availability of blood products and a low reduction-oxidation (redox) potential, which characterizes the anaerobic environment.

Many periodontopathogens are indeed fastidious strict anaerobes and as such may contribute little to the initiation of disease in shallow gingival pockets. In deep periodontal pockets, however, they find their preferred habitat.

The identification of bacteria within intact dental plaque represents a significant challenge. Most of the previously cited studies on dental plaque microbiota used techniques that involved disruption of the dental plaque matrix followed by microbial culture (Figure 23-11) or culture-independent identification. Techniques have been developed that allow the specific visualization of individual bacteria within mixed populations. In these methods, specific labeling is achieved using nucleic acid probes (fluorescence in situ hybridization [FISH]) or specific antibodies (immunofluorescence). FISH has been applied to identify particular species within plaque samples scraped from periodontal pockets.^109,264,404 With this methodology, it is even possible to image bacteria that have never been cultured in the laboratory such as members of the phylum Synergistetes (Figure 23-12). Supragingival biofilms have proved somewhat easier to visualize than subgingival plaque. Biofilms can be cultured on retrievable enamel chips held in intraoral devices in the mouths of volunteers. The enamel pieces can then be removed and processed for FISH or immunofluorescence. Images of biofilms can be captured by confocal laser scanning microscopy, which produces three-dimensional representations of the biofilm architecture. Labeling with nucleic acid probes requires desiccation of the sample before hybridization, and therefore some structural information may be lost. Nevertheless, this technique has been applied successfully to determine the spatial relationships between Actinomyces naeslundii and Streptococcus spp. in dental plaque.⁷⁷ In these studies, A. naeslundii cells were frequently observed juxtaposed to Streptococcus spp. cells. Many strains of Actinomyces coaggregate with oral streptococci and therefore the spatial organization of these organisms within dental plaque may be influenced by adhesin-receptor interactions.¹⁷⁵ In fact, a protein adhesin of A. naeslundii has been shown to colocalize with a cognate oral streptococcal receptor polysaccharide in in situ biofilms using specific antibody (immunofluorescence) labeling (Figure 23-13).²⁶⁶ These studies clearly demonstrate that the interactions identified between bacteria isolated in the laboratory are relevant to the dental plaque biofilm.

Figure 23-11 A plaque sample grown under aerobic (A) and anaerobic (B) conditions. A comparison of the culture plates shows different bacteria and different colony morphologies.

Figure 23-12 Identification of uncultured Synergistetes in subgingival dental plaque. Plaque was removed from deep (6 to 10 mm) periodontal pockets of volunteers with localized or generalized severe periodontitis. Bacteria were transferred to microscope slides and intracellular DNA was hybridized with nucleic acid probes for all eubacteria (blue), Synergistetes cluster A (red), or a specific member of the Synergistetes cluster A, “3.3/BH007” (green). Synergistetes 3.3/BH007 appear white (blue + red + green) and other Synergistetes cluster A appear purple (blue + red). Synergistetes cluster A were commonly detected in healthy and periodontally diseased sites, even though these organisms have never been cultured in the laboratory. Bar = 5 µm.

(Courtesy of S. R. Vartoukian and W. G. Wade, King’s College London Dental Institute, London. For further details, see Vartoukian SR, Palmer RM, Wade WG: Appl Environ Microbiol 75:3777, 2009.)

Figure 23-13 Visualization of specific bacteria in dental plaque biofilms using specific antisera. Biofilms were developed on the surface of enamel chips held in the mouths of volunteers for 8 hours. Samples were labeled with acridine orange (stains all bacteria green), antibodies against streptococcal cell surface polysaccharides (orange), and antibodies against Actinomyces oris type 1 fimbriae (blue). A. oris cells (some are indicated by white arrowheads) and polysaccharide-bearing streptococci can be seen in juxtaposition with other bacteria, indicating that mixed-species interactions are common in dental plaque biofilms. Bar = 20 µm.

(Courtesy of R. J. Palmer Jr, NIDCR, Bethesda, MD.)

The environmental parameters of the subgingival region differ from those of the supragingival region. The gingival crevice or pocket is bathed by the flow of crevicular fluid, which contains many substances which bacteria may use as nutrients. Host inflammatory cells and mediators are likely to have considerable influence on the establishment and growth of bacteria in the subgingival region. Both morphologic and microbiologic studies of subgingival plaque reveal distinctions between the tooth-associated and soft tissue-associated regions of subgingival plaque (see Figure 23-6, A to C).^209,253

The tooth-associated cervical plaque, adhering to the root cementum, does not markedly differ from that observed in gingivitis. At this location, filamentous microorganisms dominate, but cocci and rods also occur. This plaque is dominated by gram-positive rods and cocci, including S. mitis, S. sanguinis, Actinomyces oris (a new species containing strains that were formerly classified as A. naeslundii), A. naeslundii, and Eubacterium spp. (see Figure 23-1, N). However, in the deeper parts of the pocket, the filamentous organisms become fewer in numbers, and in the apical portion they seem to be virtually absent. Instead, the microbiota is dominated by smaller organisms without a particular orientation.²⁰⁹ The apical border of the plaque mass is separated from the junctional epithelium by a layer of host leukocytes, and the bacterial population of this apical tooth-associated region shows an increased concentration of gram-negative rods (see Figure 23-6, D).

The layers of microorganisms facing the soft tissue lack a definite intermicrobial matrix and contain primarily gram-negative rods and cocci, as well as large numbers of filaments, flagellated rods, and spirochetes. Studies on plaque associated with crevicular epithelial cells indicate a predominance of species such as S. oralis, S. intermedius, Parvimonas micra (formerly Micromonas micra and Peptostreptococcus micros), P. gingivalis, Prevotella intermedia, Tannerella forsythia (see Figure 23-10), and F. nucleatum.^76,79 Host tissue cells (e.g., white blood cells and epithelial cells) may also be found in this region (see Figure 23-10, C). Bacteria are also found within the host tissues, such as in the soft tissues (see Figure 23-3), and within epithelial cells (Figure 23-14), as well as in the dentinal tubules (Figure 23-15).^326,327

Figure 23-14 A to C show images of z-section No. 39 from a stack of 74 0.2-µm z-sections (magnification ×600; the scale bar in A also applies to B and C). Buccal epithelial cells (BEC) in this field were double-labeled with the EUB338 universal probe (A) and the A. actinomycetemcomitans-specific probe (B). The cell in the center of A contained a large mass of brightly fluorescent intracellular bacteria (red arrow). Other cells in the field contained smaller bacterial masses (not marked). B shows that a portion of the large mass labeled with the universal probe also hybridized with the A. actinomycetemcomitans-specific probe (green arrow). Images from A and B were superimposed (C), confirming that bacteria labeled with both probes (yellow arrow) were adjacent to other bacteria labeled only with the universal probe (red arrow). D presents a three-dimensional reconstruction of the same field. Bacteria recognized only by the universal probe are shown in solid red, whereas colocalization of the A. actinomycetemcomitans and universal probes is depicted by a green wireframe over a red interior. Reconstructed BEC surfaces are presented in blue. The red and green colors are muted when bacterial masses are intracellular, and brighter when bacteria appear to project out of the surface. The angle of view was rotated along the z-axis, and the image was zoomed. The large mass that appeared to have a lobular structure in z-section No. 39 was seen to be a cohesive unit containing A. actinomycetemcomitans in direct proximity to other species (red and green arrows).

(From Rudney JD, Chen R, Sedgewick GJ: J Dent Res 84:59-63, 2005.)

Figure 23-15 Scanning electron micrograph of bacteria within dentinal tubules.

The composition of the subgingival plaque depends on the pocket depth. The apical part is more dominated by spirochetes, cocci and rods, whereas in the coronal part more filaments are observed.

The site specificity of plaque is significantly associated with diseases of the periodontium. Marginal plaque, for example, is of prime importance in the initiation and development of gingivitis. Supragingival plaque and tooth-associated subgingival plaque are critical in calculus formation and root caries, whereas tissue-associated subgingival plaque is important in the tissue destruction that characterizes different forms of periodontitis. Biofilms also form on artificial surfaces exposed to the oral environment such as prostheses and implants. A large series of papers compared the microbiota in pockets around teeth with those around implants of partially edentulous patients. The similarities were striking.*

Accumulation of a Dental Plaque Biofilm

The process of plaque formation can be divided into several phases: (1) the formation of the pellicle on the tooth surface, (2) initial adhesion/attachment of bacteria, and (3) colonization/plaque maturation.

Colonization and Plaque Maturation

The primary colonizing bacteria (Table 23-4) adhered to the tooth surface provide new receptors for attachment by other bacteria, in a process known as “coadhesion.”¹⁷⁵ Together with growth of adherent microorganisms, coadhesion leads to the development of microcolonies (Figure 23-16) and eventually to a mature biofilm.

TABLE 23-4 Overview of Primary and Secondary Colonizers in Dental Plaque

Primary colonizers

Streptococcus gordonii Streptococcus intermedius Streptococcus mitis Streptococcus oralis Streptococcus sanguinis Actinomyces gerencseria Actinomyces israelii Actinomyces naeslundii Actinomyces oris Aggregatibacter actinomycetemcomitans serotype a Capnocytophaga gingivalis Capnocytophaga ochracea Capnocytophaga sputigena Eikenella corrodens Actinomyces odontolyticus Veillonella parvula

Secondary colonizers

Campylobacter gracilis Campylobacter rectus Campylobacter showae Eubacterium nodatum Aggregatibacter actinomycetemcomitans serotype b Fusobacterium nucleatum ssp nucleatum Fusobacterium nucleatum ssp vincentii Fusobacterium nucleatum ssp polymorphum Fusobacterium periodonticum Parvimonas micra Prevotella intermedia Prevotella loescheii Prevotella nigrescens Streptococcus constellatus Tannerella forsythia Porphyromonas gingivalis Treponema denticola

Figure 23-16 When a single microorganism adheres to the tooth surface, it can start to multiply and slowly forms a microcolony of daughter cells. These views were taken after plaque formation on a plastic strip (e.g., as shown in Figure 23-26), glued to a tooth surface.

Cell-cell adhesion between genetically distinct oral bacteria also occurs in the fluid phase (i.e., in saliva).

In the laboratory, interactions between genetically distinct cells in suspension result in clumps, or “coaggregates,” that can easily be seen with the naked eye (Figure 23-17).

Figure 23-17 Coaggregation between Streptococcus gordonii DL1 and Actinomyces oris MG1 in vitro. A monoculture of S. gordonii appears uniformly turbid. Microscopically, cells labeled with specific anti-DL1 antibodies (green) are in small chains or clumps. Following addition of A. oris, cells clump together to form macroscopic coaggregates (yellow arrows). Under the microscope, S. gordonii (green) are evenly distributed throughout the coaggregates with A. oris (orange). Bar = 20 µm.

(Image reproduced in part from Jakubovics NS, Gill SR, Iobst SE, et al: J Bacteriol 190:3646, 2008.)

Coaggregation is a direct interaction and is distinct from agglutination, which occurs when cells are stuck together by molecules in solution. At least 18 genera from the oral cavity have shown some form of coaggregation.¹⁷³ All oral bacteria possess surface molecules that foster some sort of cell-cell interaction (Figure 23-18).¹⁷⁴ The initial stages of coaggregation or coadhesion are essentially the same as the first steps involved in bacterial binding to surfaces: bacterial cells come into contact through passive or active transport and bind weakly through nonspecific hydrophobic, electrostatic, and Van der Waals forces.^{78,95,170,174} These steps can be dramatically accelerated in vitro by vigorously mixing dense suspensions of bacterial cells.¹⁷⁵ Strong cell-cell binding is then determined by the presence of adhesin proteins or carbohydrates on one partner, and complementary receptor proteins or carbohydrates on the other. Note that adhesin-receptor interactions are mediated by the fundamental physicochemical forces (hydrophobic, electrostatic, and Van der Waals), but they are highly specific.

Figure 23-18 Diagrammatic representation of initial plaque formation. Early colonizers bind to receptors in the pellicle. Each adherent cell becomes in turn the nascent surface and bridge for additional species (secondary colonizers). The complementary sets of adhesin receptor symbols (example in box) represent the various kinds of coaggregations, as well as the interactions with molecules in the pellicle. The symbol with a stem (adhesin) represents a cellular component that is heat inactivated (cell suspension heated to 85° C for 30 minutes) and sensitive to protease treatment. The cell type exhibiting the complementary symbol (receptor) is insensitive to either treatment. The symbols with a rectangular shape represent lactose-inhibitable coaggregations, others are lactose-noninhibitable.

(Adapted from Kolenbrander PE, London J: J Bacteriol 175:3247, 1993.)

Different species, or even different strains of a single species, have distinct sets of coaggregation partners (see Figure 23-18 online). Fusobacteria coaggregate with all other human oral bacteria while Veillonella spp., Capnocytophaga spp. (see Figure 23-1, Q) and Prevotella spp. bind to streptococci and/or actinomyces.^174,176,421 Each newly accreted cell becomes itself a new surface and therefore may act as a coaggregation bridge to the next potentially accreting cell type that passes by.

Many coaggregations between strains of different genera are mediated by lectin-like adhesins (proteins that recognize carbohydrates) and can be inhibited by lactose and other galactosides. The significance of coaggregation in oral colonization has been documented in studies on biofilm formation in vitro as well as in animal model studies.^31,234

Well-characterized interactions of secondary colonizers (see Table 23-4) with early colonizers include the coaggregation of F. nucleatum with S. sanguinis, Prevotella loescheii with A. oris, and Capnocytophaga ochracea with A. oris.^{162,164,416,417,418} Streptococci show intrageneric coaggregation, allowing them to bind to the nascent monolayer of already bound streptococci.^{148,172,259,349}

Secondary colonizers (see Table 23-4), such as Prevotella intermedia, Prevotella loescheii, Capnocytophaga spp., F. nucleatum, and P. gingivalis do not initially colonize clean tooth surfaces but adhere to bacteria already in the plaque mass.¹⁷⁴ The transition from early, supragingival dental plaque to mature plaque growing below the gingival margin involves a shift in the microbial population from primarily gram-positive organisms to high numbers of gram-negative bacteria. Therefore, in the later stages of plaque formation, coaggregation between different gram-negative species is likely to predominate. Examples of these types of interactions are the coaggregation of F. nucleatum with P. gingivalis or Treponema denticola (see Figure 23-1, L).^166,171,176

The concept that coaggregation is important during the formation of oral biofilms opens new perspectives especially for the use of probiotics. Special examples of coaggregations are the corn-cob formation (Figure 23-19), in which streptococci adhere to filaments of Corynebacterium matruchotii or Actinomyces spp., and the test-tube brush composed of filamentous bacteria to which gram-negative rods adhere.^60,209,251

Figure 23-19 Long-standing supragingival plaque near the gingival margin demonstrates “corn cob” arrangement. A central gram-negative filamentous core supports the outer coccal cells, which are firmly attached by interbacterial adherence or coaggregation.

An analysis of more than 13,000 plaque samples, looking for 40 subgingival microorganisms using a DNA-hybridization methodology, defined color-coded “complexes” of periodontal microorganisms that tend to be found together in health or disease. The composition of the different complexes was based on the frequency with which different clusters of microorganisms were recovered, and the complexes were color-coded for easy conceptualization (Figure 23-20).³⁶⁴ Interestingly, the early colonizers are either independent of defined complexes (A. naeslundii, A. oris) or members of the yellow (Streptococcus spp.) or purple complexes (A. odontolyticus (see Figure 23-1, D). The microorganisms primarily considered secondary colonizers fell into the green, orange, or red complexes. The green complex includes Eikenella corrodens, A. actinomycetemcomitans serotype a, and Capnocytophaga spp. The orange complex includes Fusobacterium, Prevotella, and Campylobacter spp. (see Figure 23-1, R). The green and orange complexes include species recognized as pathogens in periodontal and nonperiodontal infections. The red complex consists of P. gingivalis, T. forsythia, and T. denticola. This complex is of particular interest because it is associated with bleeding on probing.³⁶⁴ The existence of complexes of species in plaque is another reflection of bacterial interdependency in the biofilm environment.

Figure 23-20 Diagram of the association among subgingival species The data were derived from 13,261 subgingival plaque samples taken from the mesial aspect of each tooth in 185 adult subjects. Each sample was individually analyzed for the presence of 40 subgingival species using checkerboard DNA-DNA hybridization. Associations were sought among species using cluster analysis and community ordination techniques. The complexes to the left are comprised of species thought to colonize the tooth surface and proliferate at an early stage. The orange complex becomes numerically dominant later and is thought to bridge the early colonizers and the red complex species which become numerically more dominant at late stages in plaque development.

(Adapted from Socransky SS, Haffajee AD, Cugini MA, et al: J Clin Periodontol 25:134,1998.)

Factors Affecting Supragingival Dental Plaque Formation

Clinically, early undisturbed plaque formation on teeth follows an exponential growth curve when measured planimetrically.²⁹⁸ During the first 24 hours starting from a clean tooth surface, plaque growth is negligible from a clinical viewpoint (<3% coverage of the vestibular tooth surface, which is an amount nearly undetectable clinically). This “lag time” is due to the fact that the microbial population must reach a certain size before it can be easily detected by the clinician. During the following 3 days, coverage progresses rapidly to the point where, after 4 days, on average 30% of the total coronal tooth area will be covered with plaque (see Video 23-1: Plaque Growth online).

Video 23-1

Plaque Growth.

This movie shows undisturbed plaque formation on a central incisor after professional plaque removal over a period of 96 hours in the absence of oral hygiene. The plaque was visualized with erythrosin. After professional plaque removal, it takes, even in the absence of oral hygiene, at least 36 hours before plaque becomes detectable. Plaque first develops in irregularities on the tooth surface and at the gingival margin. Once established, the speed of plaque formation increases over time. Nevertheless, even in the absence of oral hygiene, no more than one-fourth of the tooth surface is covered after 96 hours of undisturbed plaque growth.

Several reports have shown that the microbial composition of the dental plaque will change with a shift toward a more anaerobic and a more gram-negative flora, including an influx of fusobacteria, filaments, spiral forms, and spirochetes (Figure 23-21). This was beautifully illustrated in experimental gingivitis studies.^375,387 In this ecologic shift within the biofilm, there is a transition from the early aerobic environment characterized by gram-positive facultative species to a highly oxygen-deprived environment in which gram-negative anaerobic microorganisms predominate. Bacterial growth in older plaque is much slower than in newly formed dental plaque, presumably because nutrients become limiting for much of the plaque biomass.⁴¹⁵

Figure 23-21 Artistic impression of the 1965 published classic experimental proof of the bacterial etiology of gingivitis using the experimental gingivitis model. In plaque-free subjects with clinically non-inflamed gingiva, plaque will slowly develop on the teeth when all mechanical plaque control is stopped. With time, the plaque composition changes. During the first few days, the plaque is mainly composed of gram positive (+) cocci and rods. Later on, the composition is shifted toward more gram-negative species, more rods, filaments, and finally gram-negative (−) spirochetes appear (see morphotypes in black). Within a few days, a mild gingivitis ensues (black line, gingival index according to Löe & Silness, GI=1). From the moment that proper plaque control is re-established (vertical line), the plaque composition returns to the initial situation and the symptoms of gingivitis disappear.

(From Loe H, Theilade E, Jensen SB: Experimental gingivitis in man, J Periodontol 36:177, 1965.)

Topography of Supragingival Plaque

Early plaque formation on teeth follows a typical topographic pattern (Figure 23-22), with initial growth along the gingival margin and from the interdental space (areas protected against shear forces). Later, a further extension in the coronal direction can be observed.^239,302 This pattern may fundamentally change when the tooth surface contains irregularities that offer a favorable growth path (Figure 23-23). Plaque formation can also start from grooves, cracks, perikymata, or pits.

Figure 23-22 Clinical illustration of the typical topography of plaque growth. Initial growth starts along the gingival margins and from the interdental spaces (areas protected against shear forces), to further extend in a coronal direction. This pattern may fundamentally change, for example if the tooth surface contains irregularities (such as those evident in Figure 23-23).

Figure 23-23 Important surface irregularities (crack on central upper incisor, several small pits on canine) are also responsible for the so-called individualized plaque growth pattern.

Scanning electron microscopy studies clearly revealed that the early colonization of the enamel surface starts from surface irregularities in which bacteria escape shear forces, thereby permitting them the time needed to change from reversible to irreversible binding.

By multiplication, the bacteria subsequently spread out from these starting up areas as a relatively even monolayer. Surface irregularities are also responsible for the so-called “individualized” plaque growth pattern (see Figure 23-23), which is reproduced in the absence of optimal oral hygiene.^239,240 This phenomenon illustrates the importance of surface roughness in plaque growth, which should lead to proper clinical treatment options.

Surface Microroughness

Rough intraoral surfaces (e.g., crown margins, implant abutments, and denture bases) accumulate and retain more plaque and calculus in terms of thickness, area, and colony-forming units.²⁸⁹ Ample plaque also reveals an increased maturity/pathogenicity of its bacterial components, characterized by an increased proportion of motile organisms and spirochetes, and/or a denser packing of them (Figures 23-24 and 23-25). Smoothing an intraoral surface decreases the rate of plaque formation. Below a certain surface roughness (R_a < 0.2 µm), however, further smoothing does not result in an additional reduction in plaque formation.^27,291 There seems to be a threshold level for surface roughness (R_a around 0.2 µm), above which bacterial adhesion will be facilitated.²⁶ Although surface free energy and surface roughness are two factors influencing plaque growth, the latter predominates (see Figure 23-24 and 23-25).

Figure 23-24 Photographs showing the clinical impact of surface roughness and surface free energy on de novo plaque formation. Two small strips were glued to the central upper incisors of a patient who refrained from oral hygiene for 3 days. Each strip was divided in two halves, a rough region (R_a 2.0 µm) located mesially, and a smooth region (R_a 0.1 µm) distally located. The left strip was cellulose acetate (medium surface free energy [sfe]: 58 ergcm^-2) and the right strip Teflon (low sfe: 20 ergcm^-2). Plaque was disclosed with 0.5% neutral red solution. The smooth regions show the decrease in biofilm formation because of the low sfe; the rough regions demonstrate the predominance of surface roughness (i.e., more plaque with no difference between the two surfaces), even with different sfe.

(From Quirynen M, Listgarten MA: Clin Oral Implants Res 1:8, 1990.)

Figure 23-25 A, Small plastic strip, divided in half (a rough region, R_a 2.0 µm located mesially and a smooth region R_a 0.1 µm, distally located), had been glued to the central upper incisors of a patient who refrained from oral hygiene for 3 days. B and C, After removal, the strip had been cut in small slices for microscopic evaluation. It is obvious that the rough part (C) contains a thicker plaque layer than the smooth part (B). Arrow shows border between rough and smooth surface.

Spontaneous Tooth Cleaning

Many clinicians still believe that plaque is removed spontaneously from the teeth such as during eating. However, based on the firm attachment between bacteria and surface, this seems unlikely. Even in the occlusal surfaces of the molars, plaque remains, even after chewing fibrous food (carrots, apples, or chips). The inefficiency of the spontaneous plaque removal is neatly illustrated by the clinical pictures in Figure 23-26, taken before and after dinner, starting from 4 days of undisturbed plaque formation. Only negligible differences in plaque extent could be observed.

Figure 23-26 Lower premolars and molars from a dental student who refrained from oral hygiene for 100 hours to evaluate undisturbed plaque formation. A is before dinner, and B is after dinner, eating fibrous food. Nearly no reduction in plaque extension could be observed, illustrating the absence of spontaneous plaque removal.

De Novo Subgingival Plaque Formation

It is technically impossible to record the dynamics of subgingival plaque formation in an established dentition for the simple reason that one cannot sterilize a periodontal pocket. Some early studies, using culturing techniques, examined the changes within the subgingival microbiota during the first week after mechanical debridement and reported an only partial reduction around 3 logs (from 10⁸ bacterial cells to 10⁵ cells), followed by a rapid regrowth towards nearly pretreatment levels (−0.5 log) within 7 days.^111,136,229 The fast recolonization was explained by several factors. A critical review of the effectiveness of subgingival debridement, for instance, revealed that a high proportion of treated tooth surfaces (5% to 80%) still harbored plaque and/or calculus after scaling. These remaining bacteria were considered the primary source for the subgingival recolonization.²⁷⁶ Some pathogens penetrate the soft tissues or the dentinal tubules and eventually escape instrumentation (see Figure 23-15).^4,108,320 In a beagle dog study, Leknes et al studied the extent of subgingival colonization in 6 mm pockets with smooth or rough root surfaces.²⁰¹ They also observed that smooth surfaces harbored significantly less plaque and concluded that subgingival irregularities shelter submerged microorganisms. Moreover, biopsies of the soft tissues showed an elevated proportion of inflammatory cells in the junctional epithelium (and the underlying connective tissue) facing the rough surfaces.²⁰⁰

Finally, the same group reported higher rates of attachment loss around teeth with groves in the root surface.¹⁹⁹

The introduction of oral implants, especially of the two-stage type, provides a new experimental set-up. When the transmucosal part of the implant (the abutment) is inserted on top of the osseointegrated endosseous part, a new “pristine” surface is created on which the intraoral translocation of bacteria can be investigated.²⁹¹ Recent papers demonstrated that a complex subgingival microbiota, including most periodontopathogens, is established within 1 week after abutment insertion. This is followed by a slow increase in the number of periodontopathogens.^101,303,304 Oral implants have also been used as a model to study the impact of surface roughness on subgingival plaque formation.* Smooth abutments (R_a < 0.2 µm) were found to harbor 25 times fewer bacteria than rough ones, with a slightly higher density of coccoid (i.e., nonpathogenic) cells. The subgingival microbiota was also largely dependent on the remaining presence of teeth and the degree of periodontitis in the remaining natural dentition (for review see Quirynen et al²⁹³). These observations highlight the importance of intraoral bacterial translocation for subgingival biofilms.

Metabolism of Dental Plaque Bacteria

The majority of nutrients for dental plaque bacteria originate from saliva or GCF, although the host diet provides an occasional but nevertheless important food supply. The transition from gram-positive to gram-negative microorganisms observed in the structural development of dental plaque is paralleled by a physiologic transition in the developing plaque.

The early colonizers (e.g., Streptococcus and Actinomyces species) use oxygen and lower the redox potential of the environment, which then favors the growth of anaerobic species.^75,408 Many of the gram-positive early colonizers use sugars as an energy source. The bacteria that predominate in mature plaque are anaerobic and asaccharolytic (do not break down sugars), and use amino acids and small peptides as energy sources.²¹⁷

Laboratory studies have demonstrated many metabolic interactions among the different bacteria found in dental plaque (Figure 23-27).

Figure 23-27 Schematic illustration of metabolic interactions among different bacterial species found in plaque, and between the host and plaque bacteria. These interactions are likely to be important to the survival of bacteria in the periodontal environment.

(Based on Carlsson J: Microbiology of plaque associated periodontal disease. In Lindhe J, ed: Textbook of clinical periodontology, Munksgaard, 1983; Munksgaard International Publishers; Grenier D: Infect Immun 60:5298, 1992; Loesche WJ: Periodontics 6:245, 1968; Walden WC, Hentges DJ: Appl Microbiol 30:781, 1975.)

For example, lactate and formate are byproducts of the metabolism of streptococci and actinomyces species and may be used in the metabolism of other plaque microorganisms, including Veillonella spp. and A. actinomycetemcomitans.^41,84

The growth of P. gingivalis is enhanced by metabolic byproducts produced by other microorganisms, such as succinate from C. ochracea and protoheme from Campylobacter rectus (see Figure 23-1, R).^113,114,233 Overall, the total plaque population is more efficient than any one constituent organism at releasing energy from the available substrates.⁴²²

Metabolic interactions occur also between the host and plaque microorganisms.

The bacterial enzymes that degrade host proteins result in the release of ammonia, which may be used by bacteria as a nitrogen source.⁴⁸ Hemin iron from the breakdown of host hemoglobin may be important in the metabolism of P. gingivalis.³³

Increases in steroid hormones are associated with significant increases in the proportions of P. intermedia found in subgingival plaque.¹⁸¹ These nutritional interdependencies are probably critical to the growth and survival of microorganisms in dental plaque and may partly explain the evolution of highly specific structural interactions observed among bacteria in plaque.

Communication Between Biofilm Bacteria

Bacterial cells do not exist in isolation. In a biofilm, bacteria have the capacity to communicate with each other. One example of this is quorum sensing, in which bacteria secrete a signaling molecule that accumulates in the local environment and triggers a response such as a change in the expression of specific genes once they reach a critical threshold concentration. The threshold concentration is reached only at a high-cell density, and therefore bacteria sense that the population has reached a critical mass, or quorum. There is some evidence that intercellular communication can occur after cell-cell contact and in this case, may not involve secreted signaling molecules.¹⁵⁵ Two types of signalling molecules have been detected from dental plaque bacteria: peptides released by gram-positive organisms during growth and a “universal” signal molecule autoinducer 2 (AI-2). Peptide signals are produced by oral streptococci and are recognized by cells of the same strain that produced them. Responses are induced only when a threshold concentration of the peptide is attained, and thus the peptides act as cell density, or quorum, sensors. Local concentrations of signalling molecules may be enhanced in biofilms if the signals become trapped in the biofilm matrix. The streptococcal peptides are known as competence-stimulating peptides, since the major response to these signals is the induction of competence, a physiologic state whereby cells are primed for DNA uptake and incorporation.

In some species, such as S. mutans, a small proportion of the cells in a population respond to competence-stimulating peptides by lysing.²⁷⁵ Lysis is considered to be an altruistic behavior that helps to disseminate genetic information throughout the population of S. mutans cells.

In contrast to the strain-specific competence-stimulating peptides, AI-2 is produced and detected by many different bacteria. Detection of AI-2 produces wide-ranging changes in gene expression, in some cases affecting up to one-third of the entire genome.³⁷⁶

Little is known about the specific functions of AI-2 in oral biofilms. However, this molecule has been demonstrated to play a role in mutualistic interactions between S. oralis and A. oris (A. naeslundii).³¹⁵ Thus, in an in vitro model system, neither S. oralis nor A. oris formed biofilms in monoculture. When cultured together, these organisms grew abundantly on surfaces to form thick, confluent biofilms. This mutualistic behavior was only observed when AI-2 was present: disrupting the gene for AI-2 in S. oralis abrogated mutualistic growth. These data demonstrate that AI-2 is produced and sensed by oral bacteria and suggest that interbacterial communication is important for the development of dental plaque.

Quorum sensing therefore appears to play diverse roles in, for example, modulating the expression of genes for antibiotic resistance, encouraging the growth of beneficial species to the biofilm, and discouraging the growth of competitors.

Interactions between Dental Plaque Bacteria

Several clinical studies followed the detection frequency/relative proportion of cariogenic species after periodontal therapy. They all suggest a relative increase in the number, as well as the detection frequency of S. mutans up to 8 months after mechanical debridement.^73,296 In a cross-sectional study, subgingival plaque samples from chronic periodontitis patients were tested for the presence and levels of S. mutans and putative periodontal pathogens.³⁹⁵ They divided the patients into four groups based on the stage of periodontal treatment: untreated, after initial periodontal therapy, maintenance phase without periodontal surgery, and a group of patients after periodontal surgery. The prevalence of mutans streptococci in the four study groups was equivalent. The shift toward a more cariogenic flora observed after initial and surgical periodontal therapy could be explained by a subgingival outgrowth by S. mutans occupying spots that became available after periodontal therapy (e.g., increased number of free adhesion/receptor sites), the creation of a new ecosystem in the subgingival area (a more anaerobic environment, changes in redox potential, pH, and nutrition) that allows/facilitates the growth of S. mutans species, and/or a down growth of S. mutans from the supragingival area in which the species could survive in the saliva. Additionally, Socransky and co-workers showed that the total numbers of P. gingivalis, T. forsythia, and T. denticola are reduced 2 weeks after scaling and root planing, and concomitantly several orange complex organisms are also reduced in number at this point. However, the proportion of S. sanguinis in the population increased 2 weeks after treatment by scaling and root planing in conjunction with azithromycin or metronidazole and 3 months after treatment with doxycycline. Although these observations might be due to an antagonistic relationship between S. sanguinis and periodontal pathogens, they might also reflect the relatively high intrinsic antibiotic resistance of oral streptococci.

There is some evidence from laboratory studies that nonpathogenic organisms in subgingival dental plaque can modify the behavior of periodontal pathogens.

For example, long and short fimbriae of P. gingivalis are required for adhesion and biofilm formation. The expression of long fimbriae is downregulated in the presence of S. cristatus, and short fimbriae are downregulated by S. gordonii, S. mitis, or S. sanguinis.^207,268 Changes in bacterial physiology after transitions from monoculture to mixed-species communities may be quite wide ranging.

Using a proteomics approach to probe the phenotype of P. gingivalis, it has been shown that the expression of almost 500 P. gingivalis proteins is changed in model oral microbial communities containing S. gordonii and F. nucleatum.¹⁸⁶ At present, it is not clear how these changes affect the interaction between P. gingivalis and the host.

In multispecies biofilms in which many bacteria are juxtaposed to cells of different species, interactions between genetically distinct microorganisms can be mutually beneficial (see Figure 23-4, A). However, there are many examples of competitive interactions between different bacteria (Figure 23-28).

Figure 23-28 The growth of Prevotella intermedia (black colonies) is inhibited by the presence of S. mitis, S. salivarius, and S. sanguinis (in the white holes). This is represented by a zone of no growth (halo) around the white holes. This is a typical example of growth inhibition induced by streptococci.

For example, S. mutans produces antimicrobial peptides that have broad activity against bacteria in vitro.¹⁸⁴ Other oral streptococci compete with S. mutans by excreting the strongly oxidizing molecule H₂O₂.¹⁸⁴

In fact, S. oligofermentans can convert lactic acid produced by S. mutans into H₂O₂, which then kills the S. mutans cells.³⁹⁰

In subgingival biofilms, oxygen is scarce and therefore H₂O₂ production by bacteria is low. Oxidizing agents may be more important in the interaction between host and pathogen, since reactive oxygen species are a major component of the neutrophil response to bacteria.

Another example of competitive interaction exists between streptococci and periodontopathogens. S. sanguinis, S. salivarius, and S. mitis have been shown to inhibit hard and soft tissue colonization of A. actinomycetemcomitans (see Video 23-3: Colonization Inhibition online), P. gingivalis, and Prevotella intermedia in vitro.^351,383,397

Video 23-3

Colonization Inhibition.

This movie shows the colonization of a glass surface by a green fluorescent protein labeled A. actinomycetemcomitans (green dots) strain over a 3-hour period. The colonization was visualized in a flow cell mounted on a confocal laser scanning microscope. The left panels show the colonization of a virgin surface while the middle and right panel show the colonization of a glass surface that was coated with either S. mitis or S. sanguinis. It is clearly visible that both streptococci inhibit A. actinomycetemcomitans colonization over time. S. sanguinis was clearly the best inhibitor of A. actinomycetemcomitans colonization.

Surprisingly, when these bacteria were injected in freshly root-planed periodontal pockets in vivo, a significant inhibitory effect was observed on the recolonization of the periodontal pocket by periodontopathogens.³⁸⁴ These observations highlight the importance of investigating nonpathogenic bacteria in periodontal biofilms, in addition to studying the (putative) pathogens, and might open doors for new treatment approaches.

Recent studies also show that these interactions also can influence the host. Although in the study by Teughels and co-workers, no clinical effect was expected, a reduction in bleeding on probing was noted in pockets that received the streptococci.³⁸⁴ At that time, this was explained by the establishment of a more host-compatible microbiota. However, an interaction with the host via the immune system seems an additional interesting hypothesis. Oral streptococci have been shown to modulate bacteria-host interactions involving both F. nucleatum and A. actinomycetemcomitans. Thus S. cristatus and other oral streptococci attenuate the ability of F. nucleatum to stimulate interleukin-8 (IL-8) production by host oral epithelial cells.⁴³⁸ Sliepen and co-workers recently showed similar results for the streptococci that were used in the study of Teughels and co-workers.³⁵²

Another interesting example is the S. gordonii–induced increased expression of a complement resistance protein, ApiA, in A. actinomycetemcomitans, resulting in increased resistance to killing by host serum.³⁰⁹

Biofilms and Antimicrobial Resistance

Bacteria growing in microbial communities adherent to a surface do not “behave” the same way as bacteria growing suspended in a liquid environment (in a planktonic or unattached state). For example, the resistance of bacteria to antimicrobial agents is dramatically increased in the biofilm.^8,65,139,284 Almost without exception, organisms in a biofilm are 1000 to 1500 times more resistant to antibiotics than in their planktonic state. The mechanisms of this increased resistance differ from species to species, from antibiotic to antibiotic, and for biofilm growing in different habitats.

It is generally accepted that the resistance of bacteria to antibiotics is affected by their nutritional status, growth rate, temperature, pH, and prior exposure to subeffective concentrations of antimicrobial agents.^39,40,423 Variations in any of these parameters will thus lead to a varied response to antibiotics within a biofilm. An important mechanism of resistance appears to be the slower rate of growth of bacterial species in a biofilm, which makes them less susceptible to many but not all antibiotics.^18,37,65,431 The biofilm matrix, although not a significant physical barrier to the diffusion of antibiotics, does have certain properties that can retard antibiotic penetration.

For example, strongly charged or chemically highly reactive agents can fail to reach the deeper zones of the biofilm because the biofilm acts as an ion-exchange resin removing such molecules from solution.^106,378

In addition, extracellular enzymes such as β-lactamases, formaldehyde lyase, and formaldehyde dehydrogenase may become trapped and concentrated in the extracellular matrix, thus inactivating some antibiotics (especially positively charged hydrophilic antibiotics).

Some antibiotics, such as the macrolides, which are positively charged but hydrophobic, are unaffected by this process.

Recently, “super-resistant” bacteria were identified within a biofilm. These cells have multidrug resistance pumps that can extrude antimicrobial agents from the cell.³⁷ Since these pumps place the antibiotics outside the outer membrane, the process offers protection against antibiotics that target, for example, cell wall synthesis. The penetration and efficacy of antimicrobials against biofilm bacteria are critical issues for the treatment of periodontal infections.

Antibiotic resistance may be spread through a biofilm by intercellular exchange of DNA. The high density of bacterial cells in a biofilm facilitates the exchange of genetic information among cells of the same species and across species and even genera. Conjugation (the exchange of genes through a direct interbacteria connection formed by a sex pilus), transformation (movement of small pieces of DNA from the environment into the bacterial chromosome), plasmid transfer, and transposon transfer have all been shown to occur in biofilms.

Viruses

Viral diseases of the oral mucosa and the perioral region are often encountered in dental practice. Viruses are important ulcerogenic and tumorigenic agents of the human mouth. The finding of an abundance of mammalian viruses in periodontitis lesions may suggest a role for viruses in more oral diseases than previously recognized.³⁴¹ A more in-depth review can be found in Slots.³⁵⁶

In contrast to most bacteria, viruses replicate only when present within eukaryotic (animals, plants, protists, and fungi) or prokaryotic (bacteria and archaea) cells, and not on their own. The extracellular virion particle ranges in size from 20 nm to 300 nm and consists of either DNA or ribonucleic acid (RNA) contained within a protective protein coat or capsid (Figure 23-30).

Figure 23-30 Electron micrographic picture of herpes simplex virus type 1, enveloped (left) and naked capsids (right). (Magnification ×198,000.)

(From Fields BN, Knipe DM: In Fields BN, eds: Fields’ virology, New York, 1985, Raven Press.)

Some viruses have an additional envelope comprising a lipid bilayer derived from the outer cellular membrane, the internal nuclear membrane, or the endoplasmic reticulum membrane of the infected cell. Four major viral families are associated with the main viral oral diseases of adults, as follows:

TABLE 23-5 Herpesviruses that Can Be Recovered in the Oral Cavity

TABLE 23-6 Papillomaviruses that Can Be Recovered in the Oral Cavity

TABLE 23-7 Picornaviruses that Can Be Recovered in the Oral Cavity

TABLE 23-8 Retroviruses that Can Be Recovered in the Oral Cavity