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Applied and Environmental Microbiology, March 2008, p. 1428-1435, Vol. 74, No. 5
0099-2240/08/$08.00+0     doi:10.1128/AEM.02039-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.

Surface Response of Fluorine Polymer-Incorporated Resin Composites to Cariogenic Biofilm Adherence{triangledown} ,{dagger}

Mariko Gyo,1* Toru Nikaido,1 Koichi Okada,3 Junichi Yamauchi,3 Junji Tagami,1,2 and Khairul Matin1,2

Cariology and Operative Dentistry, Department of Restorative Sciences, Graduate School, Tokyo Medical and Dental University, 5-45 Yushima 1-chome, Bunkyo-ku, Tokyo 113-8549, Japan,1 Center of Excellence Program for Frontier Research on Molecular Destruction and Reconstruction of Tooth and Bone, Tokyo Medical and Dental University, 5-45 Yushima 1-chome, Bunkyo-ku, Tokyo 113-8549, Japan,2 Kuraray Medical, Ote Center Building, 1-1-3 Otemachi, Chiyoda-ku, Tokyo 110-0004, Japan3

Received 6 September 2007/ Accepted 30 December 2007


   ABSTRACT
Top
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Experimental resin composites with incorporated polytetrafluoroethylene (PTFE) particles were developed, which theoretically could improve the surface properties of the materials, including the inhibition of bacterial adherence. To assess the surface properties in relation to biofilm formation and detachment, 23.1% (wt/wt) linear PTFE particles (FL-30) and cross-linked PTFE particles (FC-30) were incorporated into pure resin composites. Pure PTFE plates and pure resin composites without PTFE (F-0) were used as control specimens. Sucrose-dependent Streptococcus mutans biofilms were formed on the specimen blocks inside an oral biofilm reactor for various time periods and analyzed with or without application of driving forces. In addition, water contact angles and surface roughness were measured. The water contact angles of FL-30 (61.2°) and FC-30 (65.8°) were larger than that of F-0 (48.5°). The largest contact angle (107°) was detected on pure PTFE plates. However, the surfaces of FL-30, FC-30, and pure PTFE plates were rougher than that of F-0. Although the surface properties of the materials differed in terms of contact angles and roughness, these factors seemed not to affect biofilm formation on the surfaces within 5 h. Pure PTFE plates harbored almost the same amounts of biofilm as F-0. However, when a very strong driving force was applied, it was clear that there were significantly smaller amounts of biofilms retained on pure PTFE plates, which showed contact angles much higher than those of the other materials. Hydrophobicity of the resin composite was improved by incorporation of PTFE fillers. However, surface resistance against biofilm formation was not improved.


   INTRODUCTION
Top
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Restorative materials are among the most common indwelling medical devices used to restore decayed functional tooth structures. Newly designed restorative materials which are emerging as a result of advancements in science and technology are, ideally, expected to be resistant to bacterial adhesion. In the case of nonshedding surfaces, oral pathogens tend to adhere to the surfaces and form biofilms, which often results in worsening of the primary tooth structure and induces caries on adjacent teeth (5, 14). Biofilm formation is a stepwise process initiated by adhesion of planktonic bacteria onto the surface of a tooth or other structures in the oral cavity. This process progresses from colonization and coadhesion, via growth and maturation, to detachment and spread of the microorganisms from the biofilm (11, 32). In biofilm formation, early colonization is considered to be the most important step which depends on the nature of the host surface. It is now well documented that a bacterium adheres directly to the pellicle acquired on the host surface before exhibiting extensive inter- and intrageneric coaggregations (19). For these reasons, the physical and chemical properties of the surface affect the feasibility of bacterial infection (7, 17). Therefore, a novel strategy to prevent biofilm formation would be to physically and chemically modify the surface of a restorative material in order to inhibit bacterial adherence to the surface. Correlations between bacterial adhesion and various surface characteristics (chemical composition, surface energy, surface roughness, and presence of functional groups on the surface) have been intensively investigated in an attempt to reduce bacterial adhesion through surface modification (7, 29).

Polytetrafluoroethylene (PTFE) is a fluoropolymer developed by R. J. Plunkett in 1938 (30). It has several unique characteristics: high resistance to chemical regents, low surface energy, tolerance to low and high temperatures, resistance to weathering, low friction wiring, electrical insulation, and slipperiness (30). For the last three decades, PTFE has been extensively used as an implant material or a component of various devices in clinical dentistry, such as in oral surgery, prosthetic dentistry, implantology, and periodontology, by taking advantages of its above-mentioned specific surface characteristics (8, 16, 33). However, conventional PTFE has disadvantages for use as a restorative material because of its poor resistance to abrasion. In 1992, the Japan Atomic Energy Research Institute (JAERI) developed cross-linked PTFE (C PTEF) with gamma-beam irradiation. The cross-linking of PTFE drastically enhanced its resistance to abrasion and deformation(20). In this study, experimental resin composites with incorporated PTFE particles were developed; this theoretically would improve the self-cleaning property (4) of the material, which in turn would inhibit bacterial adherence.

Recently, an oral biofilm reactor (OBR) was developed to study oral biofilm formation on dental materials in vitro by simulating the human oral environment (27, 34). By using the OBR, a study model was established to provide a better understanding of how the surface properties of a dental material could influence biofilm adherence and growth. Therefore, the objectives of this study were to evaluate the surface properties of the C PTFE-incorporated resin composites in relation to biofilm formation and detachment by utilizing the OBR model before clinical trials.


   MATERIALS AND METHODS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Materials used in this study.
The PTFE-incorporated composite resins used in this study are listed in Table 1. The conventional linear PTFE (L PTFE) (Sanplatec, Osaka, Japan) and the C PTFE (Hitachi Cable, Tokyo, Japan) are spherical particles with a 20-µm diameter which were used as filler particles in the experimental resin composites as described below. A conventional pure PTFE plate was used as a control specimen. The compositions of the experimental resin composites are listed in Table 2. The experimental resin composites were mixtures of multifunctional methacrylates, photoinitiator, and microfiller, the composition of which was based on a resin composite (White Coat; Kuraray Medical, Tokyo, Japan). F-0 indicates a pure resin composite without PTFE; 23.1% (wt/wt) L PTFE particles (FL-30) and C PTFE particles (FC-30) were incorporated with F-0.


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  		TABLE 1. PTFEs used in this study

 

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  		TABLE 2. Experimental resin composites and pure PTFE plate used in this study

 
Contact angle measurement.
The experimental resin composites (F-0, FL-30, and FC-30) were dispensed from each syringe onto a glass plate. Square specimens (approximately 20 by 20 by 1.5 mm) were prepared using glass plates, pressed with a clean glass plate, and light cured by using visible light (Optilux 501; Sybron Kerr Corp., CA) from the top and bottom sides with a curing light unit at 600 mW/cm2 for 40 s each. The surface of the sample block was then ground flat using SiC paper up to 1,500 grit and polished with a diamond paste (DP-paste; P Struers, Denmark) down to 1/4-µm particle size under running water. The pure PTFE plate was not polished due to technical difficulties. The specimens were then kept under distilled water for 24 h. The surface was air dried gently prior to contact angle measurements. The relative hydrophobicity of the material surfaces was determined by measuring the contact angle using a drop shape analysis system (FTÅ32; First Ten Ångstroms, Portsmouth, England), which can detect the image of a drop with a charge-coupled device camera, and the captured images were analyzed on a personal computer. Deionized water (3.0 µl) was used as a test fluid for contact angle measurements. The surface of the drop was constantly monitored, and the contact angle was measured just after 20 s, when the droplet was stabilized. The contact angle was defined as the angle between the surface of a dome-shaped drop and that of a sample plate at the contact edge. Six specimens were used for each group.

Surface roughness measurement.
Specimens were prepared according to the procedure described above with minor modifications; resin composites (F-0, FL-30, and FC-30) were inserted into metal molds (approximately 4.0 by 4.0 by 1.5 mm) on a glass plate, pressed with another glass plate to flatten the surface, and light cured from the top and bottom sides for 20 s each. The specimens were then polished down to 0.25 µm with diamond paste. The samples and pure PTFE plate were stored in distilled water at 37°C for 24 h before further use. The surface roughness of the specimens was measured using a confocal laser scanning microscope (VK8510; Keyence, Tokyo, Japan) and was expressed as the average roughness (Ra) value prior to biofilm formation. Four specimens were used for each group.

Preparation of bacterial suspension.
Suspensions of Streptococcus mutans MT8148 in phosphate-buffered saline (PBS) at an optical density at 500 nm OD500) of 4.0 (approximately 4.0 x 107 CFU/ml) were prepared from 16-h fresh cultures in brain heart infusion broth (Becton Dickinson, Sparks, MD) after washing three times with PBS. The bacteria were stored at 4°C with continuous gentle stirring until use. For growth of the cultivated bacteria, a solution of heart infusion broth (Becton Dickinson, Sparks, MD) with sucrose (1.0% final concentration) was utilized.

Specimen assembly and biofilm formation in the OBR.
Specimen preparation for the OBR was carried out in the same manner as described for specimen preparation for surface roughness measurements. The bacteria were grown to form biofilms on slab surfaces placed inside the water jacket-encircled chamber of the OBR. Two sets of four slabs from each group were positioned on the same holder around a flat-bulb pH electrode using red utility wax (GC, Tokyo, Japan). Therefore, one side of the surface of each slab was subjected to biofilm attachment. The open surfaces of each slab were kept horizontal. A holder bearing the slabs was set on a silicon plug at the bottom of the chamber. Pooled sterile human saliva was then poured onto the slabs, followed by incubation for 30 min in order to obtain a coat of salivary pellicle on the slab surface. The top of the chamber was sealed with a silicon plug so that the chamber itself served as an incubator with a 37°C internal temperature. The top silicon plug was equipped with five stainless steel tubes (21 gauge) which were connected via silicon tubes to computer-controlled roller pumps (EYELA EPC-2000; Tokyo Rika, Tokyo, Japan). The tubes were allotted to supply one of the test solutions: one tube for the S. mutans suspension, two for heart infusion broth with sucrose, and the other two for PBS. All of these solutions were pumped into the chambers at 6 ml per hour so that a solution was continuously supplied to the centers of the slab surfaces. All of these solutions formed a water dome on the surface of the holder which was continuously stirred by the falling drops. When the water dome reached its maximum height, the mixture of excess liquid fell down from the edge of the holder. Two chambers were used at the same time to confirm experimental stability. During the experiment, the pH on the surface of the holder was continuously monitored to maintain suitable conditions for biofilm formation.

Quantitative assessment of the biofilms formed on the specimens.
S. mutans biofilm formation on the specimens in the OBR was quantitatively assessed at two different stages in this study. In order to assess initial plaque formation at early stages (stage I), 5-h S. mutans biofilms were quantified when the specimens were covered by only a thin layer of bacterial colonies. The specimens with artificial biofilms were removed from the holder in the OBR and were rinsed with PBS to remove the planktonic bacteria and loosely attached biofilms. The specimens were then incubated in 1 ml of 0.5 M sodium hydroxide solutions individually using microtubes for 15 min and were vortex agitated for 15 s in order to detach the adhered biofilms from the sample slabs and dissolve the water-insoluble glucan (WIG) matrix. The solution was centrifuged at 5,000 rpm for 10 min in order to separate the dissolved WIG from the bacteria embedded in the biofilms. Each bacterial pellet was resuspended in 500 µl of PBS, and 100 µl of the solution was transferred into 96-well flat-bottom microplates to quantify the bacteria by turbidimetric analysis (OD500) using a Biotrak II Plate reader (Biochrom, Cambridge, United Kingdom). The amount of dissolved WIG was measured by the phenol-H2SO4 method (1). The WIG solution (500 µl) from each sample was disintegrated with phenol-H2SO4, and 200 µl of each was used to estimate the amounts of WIG (µg/ml) using a Biotrak II Plate reader (OD492). In order to obtain a standard glucose curve, 0-, 25-, 50-, 75-, 100-, 150-, and 200-µg/ml solutions of glucose were also assayed by OD492.

The stage II quantitative assessment was performed after 20 h, when biofilms were maturing by condensing much thicker layers onto the specimen surfaces. Each slab was removed from the holder by the same method described above and incubated in 1 ml of cool PBS separately using microtubes. Each slab was then subjected to shaking with a TissueLyser (Qiagen/Retsch, Germany) at 30 Hz for 30 s. The slabs were then transferred carefully from the PBS to 1 ml of 0.5 M sodium hydroxide solution in order to separate WIG from the bacteria in the biofilm, which was considered the "retained biofilm" after shaking. The PBS solutions containing "detached biofilm" were collected and centrifuged at 5,000 rpm for 10 min. The bacterial cells and WIG were then separated by careful transfer from PBS to 1 ml of 0.5 M sodium hydroxide solution. The solution was centrifuged at 5,000 rpm for 10 min in order to separate the dissolved WIG and the bacteria embedded in the biofilms. The amounts of the bacterium and WIG were measured using the procedures described above. The sum of the detached biofilms and retained biofilms (measured separately as the amounts of bacteria and glucan) was considered the total amount of biofilms for each slab. All experiments were repeated three times for reproducibility.

Morphological study of the biofilms by SEM.
Initial biofilm attachment on the material surface was observed with a scanning electron microscope (SEM) (JSM-5310LV; JEOL, Tokyo, Japan). S. mutans biofilms were formed for 2 h in the OBR in the same manner as described above. The samples (F-0, FC-30, FL-30, and pure PTFE plate) were removed from the chamber, rinsed with PBS, and fixed in 4% paraformaldehyde with 1% glutaraldehyde in PBS for 1 h. The samples were then rinsed with PBS and deionized water three times. Following these procedures, the samples were dehydrated through a series of ethanol washes(50, 70, 80, 95, and 100%), desiccated, and sputter coated using gold (SC-701AT; Elionix, Tokyo, Japan). The specimens were then observed using an SEM.

In addition, biofilms were also formed using Streptococcus mitis ATCC 6249, one of the commensal bacteria of the oral flora which do not secrete WIG when supplemented with sucrose, on the sample surface, employing the same method as described above. These were assessed by SEM as for S. mutans.

Statistics.
All quantitative data were analyzed using the Statistical Package for the Medical Science (SPSS version 11 for Windows) for statistical procedures. The data for surface roughness (Ra value) were analyzed by one-way analysis of variance and Tukey's honestly significant difference test (P > 0.05), while contact angles and amounts of bacteria/mm2 and of glucan/mm2 were analyzed by one-way analysis of variance and the Dunett C test (P > 0.05).


   RESULTS
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
Contact angle and surface roughness.
The results of contact angle and surface roughness (Ra value) measurements of the material surfaces are summarized in Table 3. Addition of PTFE caused a significant increase in contact angle size in both FL-30 and FC-30 (P < 0.05). However, there were no differences between FL-30 and FC-30 (P > 0.05). The largest contact angle was detected on pure PTFE plates (107°); the other materials which displayed contact angles of less than 60°.


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  		TABLE 3. Contact angles and surface roughnesses of the experimental materials

 
For surface roughness, statistical analysis showed that the Ra values of FL-30, FC-30, and pure PTFE plates were larger than that of F-0 (P < 0.05). However, there was no statistical difference in Ra values between FL-30, FC-30, and pure PTFE plates (P < 0.05).

Quantitative assessment of the S. mutans biofilms. (i) Five-hour biofilms.
Sucrose-dependent S. mutans biofilms were formed on all the sample surfaces in a similar fashion. The amounts of initial biofilm formation were almost identical among the four groups, as shown in Fig. 1. There were no significant differences in either the amounts of bacteria/mm2 or levels of glucan/mm2 among the four groups (P > 0.05). For example, with FL-30 the amount of adhered bacteria was 0.0150 ± 0.00215, and with F-0 it was 0.0165 ± 0.00349. The data are typical of those from three identical experiments using freshly prepared samples each time to ensure reproducibility.


Figure 1
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  		FIG. 1. Amounts of S. mutans biofilm formed on experimental material surfaces following 5 h in an OBR. (a) Amounts of bacteria; (b) amounts of glucan. Error bars indicate standard deviations. Asterisks indicate no significant differences (P > 0.05).

 
(ii) Twenty-hour biofilms.
The amounts of biofilm formation were almost identical among the four groups, as shown in Fig. 2. There were no significant differences in bacterial adherence and biofilm formation even after 20 h of growth. However, when a very strong driving force was applied with a TissueLyser, most of the biofilms were detached from the pure PTFE plate surface. Hence, there were significantly smaller amounts of biofilms retained on pure PTFE plates than on the other materials.


Figure 2
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  		FIG. 2. Amounts of S. mutans biofilm retained on experimental material surfaces when a driving force (i.e., shaking with a TissueLyser at 30 Hz for 30 s) was applied after 20 h of biofilm formation. (a) Amounts of bacteria; (b) amounts of glucan. Error bars are standard deviations of the results from each of four samples. There were significantly smaller amounts of biofilms retained on pure PTFE plates compared to the other materials. Horizontal lines indicate no significant differences (P > 0.05).

 
Morphological observations.
SEM photographs of the material surfaces before bacterial exposure are shown in Fig. 3a to d. F-0 demonstrated relatively smooth surfaces, while PTFE particles and voids were observed for FL-30 and FC-30. Deep crater-like gaps occasionally formed between PTFE particles and the resin matrix on the polished surfaces. The surfaces of the pure PTFE plates visually appeared to be rougher than those the other samples, showing numerous shallow cutting scratches on the entire surface.


Figure 3
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  		FIG. 3. SEM photomicrographs of the material surfaces. Magnification, x1,000. Material surfaces without bacterial exposure (a to d), with S. mutans biofilms (e to h), and with S. mitis biofilms (i to l) are shown. Plane surfaces with almost no scratches were detected on F-0 specimens (a). Asterisks indicate PTFE fillers and dislodged or broken edges of the PTFE particles observed at the interface of the resin matrix (b and c). The surface of pure PTFE plates was occupied by numerous cutting scratches of SiC paper (d, arrowhead). Large arrows indicate colony clusters with glucan on all specimen surfaces (e to h), and small arrows indicate glucan-free colony clusters (i to l).

 
The typical surface topography after 2 h of initial biofilm formation with S. mutans is shown in Fig. 3e to h. As observed by SEM, formation of S. mutans colonies on the material surfaces was almost identical in all groups. There were also many chains of streptococci free of extracellular polysaccharide matrix, most likely WIG, in all of the samples. The colonies were formed on both PTFE fillers and the resin matrix.

Biofilm formation with S. mitis is shown in Fig. 3i to l. Bacterial colonies and chains of streptococci were observed on all the experimental materials irrespective of the material components, including PTFE. No remarkable differences could be detected in the WIG-free colonies that were uniformly formed on all of the materials.


   DISCUSSION
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
In the present study, the surface properties of the C PTFE incorporated into resin composites were investigated relative to biofilm adherence and formation on the surfaces of the new materials, using an OBR system.

Biofilm formation begins with the attachment of a small number of bacterial cells (11). Cellular attachment is known to be mediated by two major factors, i.e., substrate surface properties and bacteriological properties. Substrate surface factors include surface free energy, surface charge, hydrophobicity, roughness, and surface chemistry. Bacteriological properties include bacterial cell surface free energy, surface charge, and hydrophobicity, in addition to bacterial species and strain differences (6, 13, 26, 31). The growth of biofilms makes adhesion stronger by producing extracellular polysaccharides which are also more protective for the bacterial community. The present results indicated that biofilm formation patterns were almost the same on the surfaces of the new materials as for the conventional PTFE-incorporated resin composites.

Relatively mature biofilms were formed on the sample surfaces after 5 h, which comprised the initial biofilms. Hence, 5-h biofilm adherent surface conditions were not clear enough to detect any topographical differences visually by SEM (data not shown). Nevertheless, SEM photomicrographs of 2-h biofilms were presented in this report to visualize the topographical differences more clearly on four different materials. The colonial clusters of both S. mutans and S. mitis were formed in almost similar patterns on the surfaces of the experimental materials. In the present study, biofilm formation (with or without WIG) on the surfaces was the same, irrespective of the material composition.

Concerning the relative surface properties, the results showed that the contact angles of FL-30 (61.2°) and FC-30 (65.8°) were higher than that of F-0 (48.5°), indicating that hydrophobicity of the materials was improved with incorporation of PTFE fillers in the resin composite. However, the surfaces of the materials were rougher than that of F-0. Under the test conditions used, adaptation between PTFE filler particles and the resin matrix was not sufficient to prevent gaps forming between the two ingredients, resulting in high Ra values. Furthermore, the edges of the fillers chipped off at places, which contributed to the roughness of the surface. When the roughnesses of the matrix area and filler area were measured separately, the Ra value of the filler area was found to be significantly larger than that of the matrix area (data not shown; see Fig. S1 in the supplemental material). It was also found that the Ra values of the FL-30 and FC-30 matrix areas were same as that of F-0. In contrast, pure PTFE plates that appeared rough on SEM photomicrographs actually had shallow stretches with smaller roughness depths visible on the entire surface. Therefore, the Ra value of pure PTFE plates turned out to be almost same as those of FL-30 and FC-30. Perhaps the shallow and symmetric rough surface of the pure PTFE plate did not provide enough room to build a strong base for biofilm formation, unlike the case for FL-30 and FC-30.

Although the surface properties of materials differ in terms of contact angle and surface roughness, these factors seem not to affect biofilm formation on the surfaces. It should be noted that pure PTFE plates with contact angles of more than 100° harbored almost the same amounts of biofilms as F-0, which displayed average contact angles of less than 50°.

There is controversy regarding the relationship between the hydrophobicity of a material surface and bacterial adhesion and biofilm formation. Some authors reported that an increase of the hydrophobicity of polymeric biomaterials effectively decreased bacterial adhesion (26, 31). However, some researchers reported that there is increased bacterial adhesion (6, 25), while others have found no significant correlation between hydrophobicity of a material and bacterial adhesion (2, 35). At one stage in the present study, it appeared that there was no significant correlation between the hydrophobicity of the material and biofilm formation. This can be explained by the underwater conditions shown in Fig. 4, which are similar to the intraoral microbial environment and are not favorable for a self-cleaning surface to prevent contamination. Furthermore, under water or under medium supplemented with sucrose, S. mutans formed biofilms under the most favorable conditions, since there were no powerful driving forces functioning during the entire process. Sucrose-dependent biofilms were formed on the surfaces of all the materials in similar patterns that were dependent upon superadherent WIG. Therefore, there were no differences in bacterial adherence and biofilm formation even after 20 h of growth.


Figure 4
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  		FIG. 4. (a) Video pictures of deionized water droplets (3.0 µl) on the surface of F-0 and pure PTFE plates at the time of contact angle measurement. (b) Left, schematic diagram of a chamber of the OBR. Right, enlarged diagram of a specimen holder, showing the experimental conditions during biofilm formation on material surfaces that occurred in an underwater environment. HI, heart infusion broth.

 
However, when a very strong driving force was applied (i.e., shaking with a TissueLyser at 30 Hz for 30 s), it was clear that there were significantly smaller amounts of biofilm retained on pure PTFE plates, which showed a contact angle (107°; a hydrophobic surface) much higher than those of other materials (<70°; a hydrophilic surface). Relatively weak biofilm formation may also result because the pure PTFE plates exhibit waxy (30) or nonwettable surface properties which do not allow the bacteria and WIG to bind strongly to the slippery surfaces compared to other materials.

Similarly, the correlation between the surface roughness of a biomaterial and bacterial adhesion remains to be fully understood. Tanner et al. (36) tested the effect of surface roughness on the adhesion of oral biofilms in vivo. They reported that the roughest material promoted biofilm accumulation significantly more than the other, smoother materials. Other researchers have found similar effects of increased bacterial colonization on rougher biomaterials (3, 37). In contrast, there are reports indicating no difference in bacterial adhesion with roughened surfaces (21, 39). Rough surfaces can be made more hydrophobic, even superhydrophobic, if surface roughness can be altered by proper surface geometry tuning (4).

The results of this study were influenced by the environment for biofilm formation. In this study, the materials were coated with saliva to form a pellicle. Jendresen and Glantz (18) have reported that dental materials, regardless of their original surface properties (high- or low-energy surface) assume the same surface properties as natural tooth surfaces covered with pellicle after only 1 to 2 h in the salivary environment.

Importantly, S. mutans was selected and an OBR was used to grow biofilms. S. mutans is indigenous to the oral cavity and is considered a principal causative bacterial agent for dental caries. Several characteristics of S. mutans have been implicated in the virulence of this organism in inducing carious lesions in humans (22). These include a proclivity for adhesion to the tooth surface (9); production of WIG with sucrose as a substrate, resulting in subsequent formation of plaque (15); and reduction of the pH by producing acid (10), which causes demineralization of the tooth and leads to the carious lesion. WIG is considered to be critically important in dental plaque formation because it is not dissolved in water and also possesses a marked ability to promote adherence when synthesized de novo on various solid surfaces (15). Initially, attached S. mutans cells become more strongly adherent in the presence of glucans during biofilm formation. It might be noteworthy that to obtain homogenous biofilms with uniformity in biofilm adherence by similarly produced glucan in all experiments and to attain appropriate reproducibility, this study was restricted to single-species biofilms rather than consortium biofilms. The laboratory strain S. mutans MT8148 is known to be capable of producing three major glucosyltransferases (GTFs) (GTFB, GTFC, and GTFD), and their optimum ratio is necessary for sucrose-dependent adherence of the bacteria (28). This would be a logical reason behind the strain being used as sucrose-dependent cariogenic biofilm producer that can also maintain uniformity in pH reduction curves under similar experimental conditions (12), and thus it was used in the present study. However, no differences could be detected when S. mitis adherence was tested on the surfaces of the samples used in this study. S. mitis is one of the major bacteria in the oral indigenous microflora and is not capable of producing glucans, as no gft genes have been reported to be present in any strains of S. mitis (24, 37). Evidently biofilm adherence was enhanced by WIG synthesized by S. mutans, but WIG-free S. mitis colony adhesion onto the material surfaces was also observed. In addition, morphologically no extracellular polysaccharides could be detected around the cells of the S. mitis colonies. Therefore, it can be considered that cell wall proteins or adhesins of S. mitis (38) may play major roles in their initial attachment to material surfaces irrespective of medium composition. Furthermore, strong cell-to-cell adhesion might be involved in the formation of S. mitis biofilms.

Blossey (4) suggested that the self-cleaning properties of superhydrophobic surfaces depend upon the small contact areas of the drops with these surfaces and that limiting wetting surfaces can be achieved by tuning surface geometry. A significant increase in contact angle was achieved by incorporation of PTFE particles into F-0 in these experiments. In the present study, the contact angle of pure PTFE plates was significantly higher than that of F-0, and less surface area was covered by water relative to F-0. Pictures of water drops on the surfaces of experiment materials are shown in Fig. 4a. Both S. mutans and S. mitis adhered or formed biofilms in similar patterns on all materials. However, the question of the relationship between hydrophobicity (>90° contact angle) and biofilm formation still remains unresolved. This anomaly may be relevant to the proposal made by Marmur that surface criteria cannot be applied to underwater superhydrophobicity, since they are meaningless for a plate dipped into a liquid (23). The OBR used in the present study facilitates biofilm adherence studies for comparing the surface properties of different dental materials under similar microbial culture conditions close to those of the oral environment. These environmental conditions appear to mimic an underwater environment. Figure 4b shows that the surfaces of the experimental materials remained completely under water throughout the experiments. This might be the reason why our results show no significant correlation between the surface properties of the material and bacterial accumulation.

The hydrophobicity of the resin composite was improved by incorporation of PTFE fillers. However, the null hypothesis regarding surface resistance against biofilm formation proved to be incorrect in this study. Nevertheless, the pure PTFE plate (Teflon alone), even with its rough surface, proved to be relatively surface cleansed as far as cariogenic biofilm inhibition was concerned. More investigations employing alternative experimental designs with improved surface properties of the PTFE filler-incorporated resin composite material may help further our understanding of the role of surface hydrophobicity in resistance to biofilm formation.


   ACKNOWLEDGMENTS
 
This work was supported by the Center of Excellence (COE) Program for Frontier Research on Molecular Destruction and Reconstruction of Tooth and Bone at Tokyo Medical and Dental University, Tokyo, Japan, and a grant-in-aid from the Japan Society for the Promotion of Science (JSPS; no. 16390544).

We express our gratitude to Howard K. Kuramitsu, State University of New York at Buffalo, for his kind contributions in editing the manuscript. Also, we thank Akihiko Watanabe, Department of Organic Materials, Institute of Biomaterials and Bioengineering, for his contribution in obtaining contact angle data.


   FOOTNOTES
 
* Corresponding author. Mailing address: Cariology and Operative Dentistry, Department of Restorative Sciences, Graduate School, Tokyo Medical and Dental University, 5-45 Yushima 1-chome, Bunkyo-ku, Tokyo 113-8549, Japan. Phone: 81-3-5803-5483. Fax: 81-3-5803-0195. E-mail: gyo2167{at}yahoo.co.jp Back

{triangledown} Published ahead of print on 11 January 2008. Back

{dagger} Supplemental material for this article may be found at http://aem.asm.org/. Back


   REFERENCES
Top
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

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Applied and Environmental Microbiology, March 2008, p. 1428-1435, Vol. 74, No. 5
0099-2240/08/$08.00+0     doi:10.1128/AEM.02039-07
Copyright © 2008, American Society for Microbiology. All Rights Reserved.




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