Sebastian Hahnel Angela Wieser Reinhold Lang Martin Rosentritt
Authors’ affiliations: Sebastian Hahnel, Angela Wieser, Reinhold Lang, Martin Rosentritt, Department of Prosthetic Dentistry, Regensburg University Medical Center, Regensburg, Germany Corresponding author: Sebastian Hahnel, DDS, PhD Department of Prosthetic Dentistry, Regensburg University Medical Center, 93042 Regensburg, Germany Tel.: +49 941 944 6059 Fax: +49 941 944 6171 e-mail: [email protected]
Biofilm formation on the surface of modern implant abutment materials
Key words: biomaterials, microbiology, prosthodontics Abstract Objective: To investigate the formation of biofilms on the surface of materials applied for the fabrication of implant abutments. Material and methods: Specimens were prepared from the implant abutment materials titanium, zirconia, and polyetheretherketone (PEEK); specimens made from polymethylmethacrylate (PMMA) were used for reference. All specimens were polished to high gloss using silicon carbide paper; surface roughness was determined using profilometry, and surface free energy was calculated from contact angle measurements. After the simulation of salivary pellicle formation, multispecies biofilm formation was initiated by exposing the specimens to a suspension of Streptococcus gordonii, Streptococcus mutans, Actinomyces naeslundii, and Candida albicans for either 20 or 44 h. Viable microbial biomass adherent to the specimens (n = 10 per material and incubation time) and the percentage of dead microorganisms in the different biofilms (n = 5, accordingly) were determined. Results: Significantly lower surface roughness was identified for PEEK and PMMA than for zirconia and titanium (P < 0.001); surface free energy was significantly lower for zirconia than for PEEK (P = 0.038). Significantly higher viable biomass and a significantly higher percentage of dead microorganisms were identified after 44 h than after 20 h of biofilm formation (P < 0.001, respectively); after 20 h, PEEK surfaces harbored significantly lower viable biomass than the surfaces of the other materials (P < 0.0125). After 44 h, significant differences were identified in the percentage of dead microorganisms organized in the biofilms on the different materials (P = 0.012). Conclusions: Within the limitations of a laboratory study, the results suggest that biofilm formation on the surface of PEEK is equal or lower than on the surface of conventionally applied abutment materials such as zirconia and titanium. However, clinical studies are necessary to corroborate these preliminary results.
Date: Accepted 23 June 2014 To cite this article: Hahnel S, Wieser A, Lang R, Rosentritt M. Biofilm formation on the surface of modern implant abutment materials. Clin. Oral Impl. Res. 00, 2014; 1–5. doi: 10.1111/clr.12454
Dental implants machined from titanium have become the treatment option of choice for the replacement of missing teeth. Although survival rates after 10 years of clinical service range around 90% (Norowski & Bumgardner 2009; Bumgardner et al., 2011), there is still demand for an improvement of osseointegration and maintenance of the perimucosal seal. Peri-implant infections may affect around 10% of implants and 20% of patients after 5–10 years of clinical service (Mombelli et al. 2012); however, as result of the limited scientific evidence available, some researchers estimate that the incidence of peri-implant infections might even be higher (Holmberg et al. 2013). With regard to this issue, the choice of the abutment design and material in prosthetic
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
implant restorations has only recently gained increased attention. With the introduction of individually designed abutments by means of computer-aided design and manufacturing (CAD/CAM), it has become possible to better respond to the individual functional and esthetic demands of each patient. Apart from sole esthetic considerations, the interface between prosthetic superstructure and implant can now be located in an ideal position, which means that if cemented superstructures are employed, an excess of cement can be removed more easily. However, as a result of its increased contact area with the periimplant gum tissues, the abutment has gained a steadily more important role in conserving implant health. As abutment surfaces are usually prone to subgingival biofilm formation,
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Hahnel et al Biofilm formation on abutment materials
which are – in most cases – not regularly removed, it is wishful that materials employed for the fabrication of implant abutments feature low biofilm formation on their surface. Routinely, implant abutments are manufactured from titanium; however, in the recent years, novel materials including ceramics and polymers have been introduced in prosthetic implant dentistry. Previous studies regarding biofilm formation on abutment materials predominantly addressed the impact of titanium surface roughness on biofilm formation rather than the impact of the abutment material itself (Quirynen et al. 1993; Elter et al. 2008). However, with regard to the formation of biofilms on alloys and ceramics, several studies have demonstrated that differences in biofilm formation occur on these materials, with alloys featuring thick biofilms with low and ceramic materials featuring thin biofilms with high viability (Ausschill et al. 2002; Busscher et al. 2010). While zirconia abutments are already frequently used, particularly for esthetic considerations, abutments made from the polymer polyetheretherketone (PEEK) have only recently been introduced in implant dentistry and are concurrently available as abutment for provisional implant restorations for a period of 6 months for a number of different implant brands. PEEK is a polymeric thermoplastic material with numerous favorable properties such as good mechanical properties and chemical inertness (Kern & Lehmann 2012; Fuhrmann et al. 2014), which is usually processed by means of CAD/CAM and pressing techniques. Only recently, PEEK has been suggested as a material for the fabrication of definite three-unit fixed dental prostheses (Stawarczyk et al. 2013), which supports the assumption that PEEK might also be employed for the fabrication of definite abutments not limited to temporary use in due course. To date, there is almost no scientific evidence available in the literature regarding the formation of biofilms on the surface of PEEK. With regard to the increased contact areas of modern individual abutments with periimplant gingival and mucosal tissues, the aim of this laboratory study was to analyze the formation of biofilms on the surface of PEEK in relation to conventionally applied abutment materials such as zirconia and titanium. As the conventional wisdom in dentistry is that polymeric materials yield more plaque on their surface than alloys or ceramics, we hypothesized that biofilm formation on the surface of PEEK is increased in comparison to titanium and zirconia.
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Material and methods Specimen preparation
Standardized disc-shaped specimens with a diameter of 6 mm and a thickness of 1 mm were prepared from machined pure titanium. Specimens with analogous dimensions were manufactured from zirconia (Cercon Base; DeguDent, Hanau, Germany) in accordance with the manufacturer‘s instructions. Specimens from polyetheretherketone were prepared by cutting slabs with a thickness of 1 mm from rods with a diameter of 6 mm (PEEK; Vestakeep i4 R, Evonik, Essen, Germany) with a diamond saw. Specimens made from polymethylmethacrylate (PMMA; Palapress vario; Heraeus Kulzer, Wehrheim, Germany) were used for reference. All specimens were subjected to surface polishing using a standardized procedure with grinding paper (grain 1000/4000; Buehler, Lake Bluff, IL, USA) and an automated polishing machine (Motopol 8; Buehler, D€ usseldorf, Germany). All specimens were subsequently manually polished to high gloss using a polishing paste designed for dental materials (Universal Polishing Paste; Ivoclar Vivadent, Schaan, Liechtenstein). All specimens were stored under lightproof conditions in distilled water for 6 days at 37 1°C for minimizing the impact of residual monomer leakage on cell viability and were subsequently carefully cleaned using ethanol (70%) and applicator brush tips (3M ESPE, Seefeld, Germany) prior to any further processing. Surface characterization Surface roughness
Peak-to-valley surface roughness (Ra) was determined on the surface of five randomly selected specimens for each material including the control group using a profilometric contact surface measurement device (Perthen S6P; Feinpr€ uf-Perthen, G€ ottingen, Germany). A distance of 1.75 mm was measured in three line scans for each sample perpendicular to the expected grinding grooves using a standard diamond tip (tip radius 2 lm, tip angle 90°) and a cut-off level of 0.25. Surface free energy
Contact angles of three liquids differing in hydrophobicity (bidistilled water, diiodomethane, ethylene glycol) were determined using the sessile drop method and a computer-aided contact angle measurement device (OCA 15 plus; DataPhysics Instruments GmbH, Filderstadt, Germany). The
contours of ten drops of each liquid with a drop volume of 0.2 ll were analyzed on the surface of each of three randomly selected specimens for each material including the control group; left and right contact angles were averaged. Surface free energy (SFE) was calculated in accordance with the approach introduced by Owens and co-workers (Owens & Wendt 1969). Data are indicated as total SFE, given by the summation of its disperse and polar contribution. Biofilm formation Saliva preparation
Unstimulated whole saliva was collected directly prior to the experiments by expectoration from one volunteer healthy female donor aged 23 years, who refrained from ingestion and oral hygiene for at least two hours. Saliva was sterilized using single-use filtration devices directly prior the experiments (Vacuflo; Schleicher & Sch€ ull Microscience GmbH, Dassel, Germany; 0.45 and 0.22 lm, successively). Microbiological procedures
The species Candida albicans ATCC 10231, Streptococcus mutans ATCC 25175 (both purchased from the German Collection of Microorganisms and Cell Cultures, Braunschweig, Germany), Actinomyces naeslundii T14V, and Streptococcus gordonii DL1 (both kindly provided by Paul E. Kolenbrander; National Institutes of Health, Bethesda, MD, USA) are routinely maintained in our laboratory with weekly subcultures in Schaedler medium, and long-term storage at 70°C. For work with microbial suspensions, microorganisms were separately grown aerobically in continuously shaked liquid cultures at 37°C in Schaedler medium until the early-stationary phase of growth. Cells were harvested by centrifugation (896 g, 18°C, 5 min; Hettich Rotixa P, Tuttlingen, Germany), washed twice with Dulbecco‘s Phosphate-Buffered Saline (PBS; Sigma-Aldrich, St. Louis, MO, USA), and resuspended in sterile Schaedler medium. The optical density of each microbial suspension was adjusted to 0.3 at 550 nm (Genesys 10-S; Thermo Spectronic, Rochester, NY, USA); subsequently, 1 ml of each microbial suspension was used to produce the mixed species microbial suspension employed for simulation of biofilm formation. Simulation of biofilm formation
Biofilm formation on the various specimens was simulated using a multispecies biofilm
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
Hahnel et al Biofilm formation on abutment materials
model. In brief, specimens were transferred to sterile 48-well cell clusters and pellicle formation was initiated by incubating the sample-containing plates with 1 ml per well of sterile human whole saliva at 37°C in a thermo shaking device (OrbitalShaker; Thermo Forma, Marietta, OH, USA). After 2 h, saliva was removed carefully, and each specimen was incubated with 1 ml of mixed species microbial suspension for 2.5 h. Subsequently, microbial suspensions were carefully removed, and each specimen was incubated with 1 ml of Schaedler medium. After 20 or 44 h of incubation, Schaedler medium was carefully removed and specimens were carefully washed twice with phosphate-buffered saline (PBS). Analysis of biofilm formation
The relative amount of viable microorganisms adherent to the different specimens was analyzed using an MTT-based cell viability assay as described previously (Ionescu et al. 2012; Hahnel et al. 2014). A total of 15 specimens were analyzed for each of the investigated materials and incubation times, including 10 test specimens and five samples that had been employed for control purposes (dye controls). In brief, MTT solution was prepared by dissolving 5 mg/ml thiazolyl blue tetrazolium bromide (Sigma-Aldrich) in PBS; PMS solution was prepared by dissolving 0.3 mg/ml phenazine methosulfate (Sigma-Aldrich) in sterile PBS. MTT reaction solution was prepared by freshly mixing 1 ml of MTT solution, 1 ml of PMS solution, and 8 ml of sterile PBS. Subsequent to biofilm formation and careful rinsing, 200 ll of the MTT reaction solution was added to each sample well and incubated under lightproof conditions at 37°C. After 5 h, the reaction solution was gently removed from each well, and 200 ll of a lysing solution consisting of 10% V/V sodium dodecyl sulfate, and 50% V/V dimethylformamide (both SigmaAldrich) in distilled water was added to each well. After 1 h of further incubation under light-proof conditions at 37°C, 180 ll of the aliquot was transferred to new sterile well plates, and the absorbance correlating with cell viability was measured at a wavelength of 550 nm (Fluostar Optima; bmg Labtech, Offenburg, Germany). For fluorescence microscopic analysis of the ratio between live and dead microorganisms biofilms adherent to the different surfaces, each of five specimens for each material and incubation time were randomly placed in a sterile plastic Petri dish and the biofilms were scraped off the surface of using
sterile cell scrapers. Subsequently, the surface of the specimens and the Petri plate was rinsed with PBS and the pooled washings were adjusted to a final volume of 1 ml, which was finally subjected to intense vortexing. An aliquot (25 ll) of the sonified suspensions for each material and incubation time was transferred onto microscopic slides containing Live/Dead BacLight bacterial viability kit solution for microscopy and quantitative assays (Molecular Probes Inc, Eugene, OR, USA). After incubation in the dark for 15 min, five randomly selected fields with an area of 3.8 9 10 8 m2 were analyzed for each of the five specimens for each material and incubation time using a fluorescence microscope (Zeiss Axio Scope A1 LED; Zeiss, Oberkochen, Germany) to determine the percentage of dead and inactive microorganisms (fluorescent red). Statistical procedures
All statistical analyses were performed using statistical software (IBM SPSS 21.0 for Windows; IBM, Armonk, NY, USA); the level of significance (a) was set to 0.05. One-way analysis of variance was used to analyze surface roughness and SFE of the various abutment materials investigated; the Tukey Kramer post hoc test was employed to highlight significant differences where appropriate. The non-parametric Kruskal–Wallis test was employed to investigate differences in cell viability and the percentage of dead microorganisms in the biofilms, setting the abutment material and incubation time as fixed factors. Subsequent post hoc analysis was performed using the Bonferroni-adjusted Mann–Whitney U-test where appropriate; the adjusted a‘ was set to 0.0125.
Results
Table 1. Surface roughness (Ra) and surface free energy (mJ/m2) of the different materials investigated in this study. Means and standard deviations are indicated; identical superscript letters within a single column indicate significant differences for a = 0.05 Material
Surface roughness
Zirconia Titanium PEEK PMMA
0.16 0.17 0.04 0.05
(0.02)a,b (0.03)c,d (0.02)a,c (0.03)b,d
Surface free energy 31.72 35.62 42.19 39.14
(4.43)a (4.26) (2.80)a (3.39)
Biofilm formation
Statistical analysis indicated significant differences in cell viability, suggesting significant differences in multispecies biofilm formation between the different materials after 20 h (P < 0.001). Lowest cell viability indicating the lowest amount of adherent viable biomass was identified on the surface of PEEK, which was significantly lower than for zirconia (P < 0.001), titanium (P = 0.004), and PMMA (P < 0.001). Cell viability increased significantly from 20 to 44 h of biofilm formation, indicating a significantly higher amount of viable biomass adherent to the abutment materials after prolonged biofilm formation (P < 0.001). After 44 h, no significant differences in cell viability were identified between the different abutment materials (P = 0.068) (cf. Table 2). Regarding fluorescence microscopic analysis of the percentage of dead microorganisms in the biofilms, significant differences were identified between the biofilms on the various materials after 44 h (P = 0.012) but not after 20 h (P = 0.500). After 44 h, the biofilms on zirconia yielding a significantly higher percentage of dead microorganisms than on PMMA (P = 0.002). Generally, a significantly higher percentage of dead microorganisms was identified in the biofilms after 44 h than after 20 h (P < 0.001).
Surface properties
One-way ANOVA indicated significant differences in surface roughness between the different materials (P < 0.001). Post hoc analysis identified significantly lower surface roughness for specimens made from PEEK and PMMA than for specimens made from titanium and zirconia (P < 0.001) (cf. Table 1). One-way ANOVA indicated significant differences in SFE between the various materials (P = 0.044). Subsequent post hoc analysis suggested that zirconia yielded significantly lower SFE than PEEK (P = 0.038); no significant differences were observed between the other materials.
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
Discussion Within the limitations of an in vitro study, the results of this research suggest rejection of the research hypothesis, supporting the assumption that biofilm formation is not increased on the surface of PEEK in comparison to the conventionally applied implant abutment materials zirconia and titanium. As oral biofilm formation is a very complex process, it is clear that the results gathered in this laboratory approach cannot be completely transferred to a clinical setting. Nevertheless, numerous parameters influencing oral biofilm formation have been simulated,
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Hahnel et al Biofilm formation on abutment materials
Table 2. Relative absorbance values indicating viable biomass in biofilms after 20 and 44 h and percentage of dead microorganisms in the different biofilms on the different materials. Means and standard deviations (SD) as well as medians are indicated; identical superscript letters within the median column for each time indicate no statistically significant differences between the various materials Cell viability
Percentage of dead microorganisms
20 h
44 h
20 h
44 h
Material
Mean (SD)
Median
Mean (SD)
Median
Mean (SD)
Median
Mean (SD)
Median
Zirconia Titanium PEEK PMMA
0.31 0.22 0.11 0.27
0.33a,b 0.23a,c 0.10 0.26b,c
0.36 0.50 0.29 0.47
0.37a,b,c 0.46a,d,e 0.30b,d,f 0.53c,e,f
59.12 52.48 59.92 56.27
55.74a,b,c 52.58a,d,e 58.37b,d,f 56.54c,e,f
74.87 68.82 63.72 60.17
77.01a,b 65.33a,c,d 65.52b,c,e 62.30d,e
(0.09) (0.09) (0.04) (0.09)
including the formation of an acquired salivary pellicle on the surface of the abutment materials and the use of a multi-species biofilm model. To keep the experimental conditions applied reproducible, saliva from a single healthy donor rather was used than pooled saliva from a multitude of persons. With regard to the maturing process of biofilms on dental implant surfaces, the conventional wisdom is that bacteria such as Streptococci and Actinomyces species prevail in the early phases of biofilm formation and prepare the ground for the subsequent adhesion of anaerobic and more pathogenic microorganisms, which are dominant in more mature biofilms after 48 h (Nakazato et al. 1989; Kolenbrander et al. 2010). This circumstance has been taken into consideration by simulating biofilm formation with a mixture of at least four early-colonizing microorganisms for a maximum of 44 h. The multi-species biofilm model included microorganisms such as Actinomyces naeslundii and Streptococcus gordonii, which are commonly counted among the early-colonizing microorganisms in the oral cavity (Periasamy & Kolenbrander 2009); for these two bacteria, Streptococcus mutans and Candida albicans, relevant interaction effects have been identified (Palmer et al. 2001; Pereira-Cenci et al. 2008; Bamford et al. 2009; Kolenbrander et al. 2010), which justifies their inclusion in a multispecies biofilm model. Although PEEK is increasingly employed as abutment material in contemporary implant dentistry, biofilm formation on the surface of PEEK has – to the knowledge of the authors – hardly been addressed. Laboratory studies have shown that biofilms can be reproducibly grown on the surface of PEEK (Williams et al. 2011), and it has been reported in a recent conference abstract that the adhesion and proliferation of oral streptococci is similar on the
(0.13) (0.22) (0.14) (0.23)
(15.70) (12.60) (15.20) (11.65)
surface of PEEK and a conventional resinbased composite (Kolbeck et al. 2013). The only study investigating microbial issues in implants supplied with PEEK abutments identified similar microbial counts and levels of periodontal pathogens in the peri-abutment region of implants supplied with PEEK and titanium healing abutments (Volpe et al. 2008); however, no direct investigation of the biofilms on the surface of the different abutments was performed. Analysis of biofilm formation in this study suggests lower biofilm formation on the surface of PEEK than on the conventionally applied abutment materials titanium and zirconia, which hints that – from a microbiological point of view – PEEK features favorable properties as definite abutment material. The conventional wisdom is that the surface properties of a material have a pronounced influence particularly on the early phases of biofilm formation. For the development of biofilms on the surface of dental implant surfaces, this circumstance has led to the postulation that it is particularly the initial adherence of early-colonizing microorganisms that has to be prevented (Subramani et al. 2009). It has been reported that the surface roughness and SFE of an implant surface have an effect on the initial adherence of microorganisms, suggesting that smooth surfaces and those with low SFE feature less microbial adherence than materials with higher surface roughness or SFE. For titanium surfaces, threshold values for surface roughness (Ra) ranging between 0.088 lm (Rimondini et al. 1997) and 0.2 lm (Bollen et al. 1997) have been identified, suggesting that lower values do not have a significant impact on biofilm formation. However, it is doubtful whether the differences in surface roughness values identified for the materials investigated in this study can be accounted for differences in biofilm formation: although
(11.53) (13.76) (10.71) (10.79)
the surface roughness of titanium and zirconia was significantly higher than the surface roughness of both PEEK and PMMA, biofilm formation did not correlate with these results as significantly more viable biomass was identified on PMMA than on PEEK. Although a relationship between the SFE of implant materials and microbial adherence has been proven in numerous studies (Teughels et al. 2006), other researchers assume that the relation is not as simple. Recent studies suggest that the surface composition and surface topography might impact the formation of biofilms to an even higher level (Ionescu et al. 2012; Hahnel et al. 2014), which might serve as an explanation for the poor correlation between surface properties and biofilm formation observed in this study. With regard to this aspect, future studies might set their focus on analyzing topographical differences between the various abutment materials and attempt to correlate their findings to differences in biofilm formation on the different materials. The results of this trial suggest that biofilm formation on PEEK is not higher than on conventionally applied abutment materials such as titanium and zirconia. Thus, from a microbiological point of view, PEEK abutments might be effectively employed as definite abutment material in implant dentistry. To support the data gathered in this laboratory pilot study, the authors are concurrently investigating the subgingival formation of biofilms on the surface of provisional abutments made from PEEK and titanium in a clinical study.
Acknowledgement:
Vestakeep i4 R was kindly provided by Evonik, Essen, Germany.
References Ausschill, T.M., Arweiler, N.B., Brecx, M., Reich, E., Sculean, A. & Netuschil, L. (2002) The effect
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of dental restorative materials on dental biofilm. European Journal of Oral Sciences 110: 48–53.
Bamford, C.V., d‘Mello, A., Nobbs, A.H., Dutton, L.C., Vickerman, M.M. & Jenkinson, H.F. (2009)
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Hahnel et al Biofilm formation on abutment materials
Streptococcus gordonii modulates Candida albicans biofilm formation through intergeneric communication. Infection and Immunity 77: 3696– 3704. Bollen, C.M., Lambrechts, P. & Quirynen, M. (1997) Comparison of surface roughness of oral hard materials to the threshold surface roughness for bacterial plaque retention: a review of the literature. Dental Materials 13: 258–269. Bumgardner, J.D., Adatrow, P., Haggard, W.O. & Norowski, P.A. (2011) Emerging antibacterial biomaterial strategies for the prevention of periimplant infectinos. International Journal of Oral and Maxillofacial Implants 26: 553–560. Busscher, H.J., Rinastiti, M., Siswomihardjo, W. & Van der Mei, H.C. (2010) Biofilm formation on dental restorative and implant materials. Journal of Dental Research 89: 657–665. Elter, C., Heuer, W., Demling, A., Hannig, M., Heidenblut, T., Bach, F.W. & Stiesch-Scholz, M. (2008) Supra- and subgingival biofilm formation on implant abutments with different surface characteristics. International Journal of Oral and Maxillofacial Implants 23: 327–334. Fuhrmann, G., Steiner, M., Freitag-Wolf, S. & Kern, M. (2014) Resin bonding to three types of polyaryletherketones (PAEKs) – Durability and influence of surface conditioning. Dental Materials 30: 357–363. Hahnel, S., Wastl, D., Schneider-Feyrer, S., Giessbibl, F.J., Brambilla, E., Cazzaniga, G. & Ionescu, A. (2014) Streptococcus mutans biofilm formation on experimental resin-based composites in dependence of surface treatment and S-PRG filler particle fraction. Journal of Adhesive Dentistry. doi: 10.3290/j.jad.a31800. [Epub ahead of print] Holmberg, K.V., Abdolhosseini, M., Li, Y., Chen, X., Gorr, S.U. & Aparicio, C. (2013) Bio-inspired stable antimicrobial peptide coatings for dental applications. Acta Biomaterialia 9: 8224–8231. Ionescu, A., Wutscher, E., Brambilla, E., SchneiderFeyrer, S., Giessibl, F.J. & Hahnel, S. (2012) Influ-
ence of surface properties of resin-based composites on in vitro Streptococcus mutans biofilm development. European Journal of Oral Sciences 120: 458–465. Kern, M. & Lehmann, F. (2012) Influence of surface conditioning on bonding to polyetheretherketon (PEEK). Dental Materials 28: 1280–1283. Kolbeck, C., Sereno, M., Rosentritt, M. & Handel, G. (2013) Biofilm formation on polyetheretherketone surfaces and cleaning options. IADR Seattle, No. 2353. Kolenbrander, P.E., Palmer, R.J., Jr, Periasamy, S. & Jabucovics, N.S. (2010) Oral multispecies biofilm development and the key role of cell-cell distance. Nature Reviews Microbiology 8: 471–480. Mombelli, A., M€ uller, N. & Cionca, N. (2012) The epidemiology of peri-implantitis. Clinical Oral Implants Research 23(Suppl. 6): 67–76. Nakazato, G., Tsuchiya, H., Sato, M. & Yamauchi, M. (1989) In vivo plaque formation on implant materials. International Journal of Oral and Maxillofacial Implants 4: 321–326. Norowski, P.A. & Bumgardner, J.D. (2009) Biomaterial and antibiotic strategies for peri-implantitis. Journal of Biomedical Materials Research Part B: Applied Biomaterials 88B: 530–543. Owens, D.K. & Wendt, R.C. (1969) Estimation of the surface free energy polymers. Journal of Applied Polymer Science 13: 1741–1747. Palmer, R.J., Jr, Kazmerzak, K., Hansen, M.C. & Kolenbrander, P.E. (2001) Mutualism versus independence: strategies of mixed-species oral biofilms in vitro using saliva as the sole nutrient source. Infection & Immunity 69: 5794–5804. Pereira-Cenci, T., Deng, D.M., Kranevald, E.A., Manders, E.M., Del Bel Cury, A.A., Ten Cate, J.M. & Crielaard, W. (2008) The effect of Streptococcus mutans and Candida glabrata on Candida albicans biofilms formed on different surfaces. Archives of Oral Biology 53: 755–764. Periasamy, S. & Kolenbrander, P.E. (2009) Mutualistic biofilm communities develop with Porphyro-
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
monas gingivalis and initial, early, and late colonizers of enamel. Journal of Bacteriology 191: 6804–6811. Quirynen, M., Van der Mei, H.C., Bollen, C.M., Schotte, A., Marechal, M., Doornbusch, G.I., Naert, I., Busscher, H.J. & Van Steenberghe, D. (1993) An in vivo study of the influence of the surface roughness of implants on the microbiology of supra- and subgingival plaque. Journal of Dental Research 72: 1304–1309. Rimondini, L., Fare, S., Brambilla, E., Felloni, A., Consonni, C., Brossa, F. & Carrassi, A. (1997) The effect of surface roughness on early in vivo plaque colonization on titanium. Journal of Periodontology 68: 556–562. Stawarczyk, B., Beuer, F., Wimmer, T., Jahn, D., Sener, B., Roos, M. & Schmidlin, P.R. (2013) Polyetheretherketone – a suitable material for fixed dental prostheses? Journal of Biomedical Materials Research Part B: Applied Biomaterials 101: 1209–1216. Subramani, K., Jung, R.E., Molenberg, A. & H€ammerle, C.H.F. (2009) Biofilm on dental implants: a review of the literature. International Journal of Oral and Maxillofacial Implants 24: 616–626. Teughels, W., Van Asche, N., Sliepen, I. & Quirynen, M. (2006) Effect of material characteristics and/or surface topography on biofilm development. Clinical Oral Implants Research 17(Suppl. 2): 68–81. Volpe, S., Verrocchi, D., Andersson, P., Gottlow, J. & Sennerby, L. (2008) Comparison of early bacterial colonization of PEEK and titanium healing abutments using real-time PCR. Applied Osseointegration Research 6: 54–56. Williams, D.L., Woodbury, K.L., Haymond, B.S., Parker, A.E. & Bloebaum, R.D. (2011) A modified CDC biofilm reactor to produce mature biofilms on the surface of PEEK membranes for an in vivo animal model application. Current Microbiology 62: 1657–1663.
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Authors’ affiliations: Sebastian Hahnel, Angela Wieser, Reinhold Lang, Martin Rosentritt, Department of Prosthetic Dentistry, Regensburg University Medical Center, Regensburg, Germany Corresponding author: Sebastian Hahnel, DDS, PhD Department of Prosthetic Dentistry, Regensburg University Medical Center, 93042 Regensburg, Germany Tel.: +49 941 944 6059 Fax: +49 941 944 6171 e-mail: [email protected]
Biofilm formation on the surface of modern implant abutment materials
Key words: biomaterials, microbiology, prosthodontics Abstract Objective: To investigate the formation of biofilms on the surface of materials applied for the fabrication of implant abutments. Material and methods: Specimens were prepared from the implant abutment materials titanium, zirconia, and polyetheretherketone (PEEK); specimens made from polymethylmethacrylate (PMMA) were used for reference. All specimens were polished to high gloss using silicon carbide paper; surface roughness was determined using profilometry, and surface free energy was calculated from contact angle measurements. After the simulation of salivary pellicle formation, multispecies biofilm formation was initiated by exposing the specimens to a suspension of Streptococcus gordonii, Streptococcus mutans, Actinomyces naeslundii, and Candida albicans for either 20 or 44 h. Viable microbial biomass adherent to the specimens (n = 10 per material and incubation time) and the percentage of dead microorganisms in the different biofilms (n = 5, accordingly) were determined. Results: Significantly lower surface roughness was identified for PEEK and PMMA than for zirconia and titanium (P < 0.001); surface free energy was significantly lower for zirconia than for PEEK (P = 0.038). Significantly higher viable biomass and a significantly higher percentage of dead microorganisms were identified after 44 h than after 20 h of biofilm formation (P < 0.001, respectively); after 20 h, PEEK surfaces harbored significantly lower viable biomass than the surfaces of the other materials (P < 0.0125). After 44 h, significant differences were identified in the percentage of dead microorganisms organized in the biofilms on the different materials (P = 0.012). Conclusions: Within the limitations of a laboratory study, the results suggest that biofilm formation on the surface of PEEK is equal or lower than on the surface of conventionally applied abutment materials such as zirconia and titanium. However, clinical studies are necessary to corroborate these preliminary results.
Date: Accepted 23 June 2014 To cite this article: Hahnel S, Wieser A, Lang R, Rosentritt M. Biofilm formation on the surface of modern implant abutment materials. Clin. Oral Impl. Res. 00, 2014; 1–5. doi: 10.1111/clr.12454
Dental implants machined from titanium have become the treatment option of choice for the replacement of missing teeth. Although survival rates after 10 years of clinical service range around 90% (Norowski & Bumgardner 2009; Bumgardner et al., 2011), there is still demand for an improvement of osseointegration and maintenance of the perimucosal seal. Peri-implant infections may affect around 10% of implants and 20% of patients after 5–10 years of clinical service (Mombelli et al. 2012); however, as result of the limited scientific evidence available, some researchers estimate that the incidence of peri-implant infections might even be higher (Holmberg et al. 2013). With regard to this issue, the choice of the abutment design and material in prosthetic
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
implant restorations has only recently gained increased attention. With the introduction of individually designed abutments by means of computer-aided design and manufacturing (CAD/CAM), it has become possible to better respond to the individual functional and esthetic demands of each patient. Apart from sole esthetic considerations, the interface between prosthetic superstructure and implant can now be located in an ideal position, which means that if cemented superstructures are employed, an excess of cement can be removed more easily. However, as a result of its increased contact area with the periimplant gum tissues, the abutment has gained a steadily more important role in conserving implant health. As abutment surfaces are usually prone to subgingival biofilm formation,
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Hahnel et al Biofilm formation on abutment materials
which are – in most cases – not regularly removed, it is wishful that materials employed for the fabrication of implant abutments feature low biofilm formation on their surface. Routinely, implant abutments are manufactured from titanium; however, in the recent years, novel materials including ceramics and polymers have been introduced in prosthetic implant dentistry. Previous studies regarding biofilm formation on abutment materials predominantly addressed the impact of titanium surface roughness on biofilm formation rather than the impact of the abutment material itself (Quirynen et al. 1993; Elter et al. 2008). However, with regard to the formation of biofilms on alloys and ceramics, several studies have demonstrated that differences in biofilm formation occur on these materials, with alloys featuring thick biofilms with low and ceramic materials featuring thin biofilms with high viability (Ausschill et al. 2002; Busscher et al. 2010). While zirconia abutments are already frequently used, particularly for esthetic considerations, abutments made from the polymer polyetheretherketone (PEEK) have only recently been introduced in implant dentistry and are concurrently available as abutment for provisional implant restorations for a period of 6 months for a number of different implant brands. PEEK is a polymeric thermoplastic material with numerous favorable properties such as good mechanical properties and chemical inertness (Kern & Lehmann 2012; Fuhrmann et al. 2014), which is usually processed by means of CAD/CAM and pressing techniques. Only recently, PEEK has been suggested as a material for the fabrication of definite three-unit fixed dental prostheses (Stawarczyk et al. 2013), which supports the assumption that PEEK might also be employed for the fabrication of definite abutments not limited to temporary use in due course. To date, there is almost no scientific evidence available in the literature regarding the formation of biofilms on the surface of PEEK. With regard to the increased contact areas of modern individual abutments with periimplant gingival and mucosal tissues, the aim of this laboratory study was to analyze the formation of biofilms on the surface of PEEK in relation to conventionally applied abutment materials such as zirconia and titanium. As the conventional wisdom in dentistry is that polymeric materials yield more plaque on their surface than alloys or ceramics, we hypothesized that biofilm formation on the surface of PEEK is increased in comparison to titanium and zirconia.
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Material and methods Specimen preparation
Standardized disc-shaped specimens with a diameter of 6 mm and a thickness of 1 mm were prepared from machined pure titanium. Specimens with analogous dimensions were manufactured from zirconia (Cercon Base; DeguDent, Hanau, Germany) in accordance with the manufacturer‘s instructions. Specimens from polyetheretherketone were prepared by cutting slabs with a thickness of 1 mm from rods with a diameter of 6 mm (PEEK; Vestakeep i4 R, Evonik, Essen, Germany) with a diamond saw. Specimens made from polymethylmethacrylate (PMMA; Palapress vario; Heraeus Kulzer, Wehrheim, Germany) were used for reference. All specimens were subjected to surface polishing using a standardized procedure with grinding paper (grain 1000/4000; Buehler, Lake Bluff, IL, USA) and an automated polishing machine (Motopol 8; Buehler, D€ usseldorf, Germany). All specimens were subsequently manually polished to high gloss using a polishing paste designed for dental materials (Universal Polishing Paste; Ivoclar Vivadent, Schaan, Liechtenstein). All specimens were stored under lightproof conditions in distilled water for 6 days at 37 1°C for minimizing the impact of residual monomer leakage on cell viability and were subsequently carefully cleaned using ethanol (70%) and applicator brush tips (3M ESPE, Seefeld, Germany) prior to any further processing. Surface characterization Surface roughness
Peak-to-valley surface roughness (Ra) was determined on the surface of five randomly selected specimens for each material including the control group using a profilometric contact surface measurement device (Perthen S6P; Feinpr€ uf-Perthen, G€ ottingen, Germany). A distance of 1.75 mm was measured in three line scans for each sample perpendicular to the expected grinding grooves using a standard diamond tip (tip radius 2 lm, tip angle 90°) and a cut-off level of 0.25. Surface free energy
Contact angles of three liquids differing in hydrophobicity (bidistilled water, diiodomethane, ethylene glycol) were determined using the sessile drop method and a computer-aided contact angle measurement device (OCA 15 plus; DataPhysics Instruments GmbH, Filderstadt, Germany). The
contours of ten drops of each liquid with a drop volume of 0.2 ll were analyzed on the surface of each of three randomly selected specimens for each material including the control group; left and right contact angles were averaged. Surface free energy (SFE) was calculated in accordance with the approach introduced by Owens and co-workers (Owens & Wendt 1969). Data are indicated as total SFE, given by the summation of its disperse and polar contribution. Biofilm formation Saliva preparation
Unstimulated whole saliva was collected directly prior to the experiments by expectoration from one volunteer healthy female donor aged 23 years, who refrained from ingestion and oral hygiene for at least two hours. Saliva was sterilized using single-use filtration devices directly prior the experiments (Vacuflo; Schleicher & Sch€ ull Microscience GmbH, Dassel, Germany; 0.45 and 0.22 lm, successively). Microbiological procedures
The species Candida albicans ATCC 10231, Streptococcus mutans ATCC 25175 (both purchased from the German Collection of Microorganisms and Cell Cultures, Braunschweig, Germany), Actinomyces naeslundii T14V, and Streptococcus gordonii DL1 (both kindly provided by Paul E. Kolenbrander; National Institutes of Health, Bethesda, MD, USA) are routinely maintained in our laboratory with weekly subcultures in Schaedler medium, and long-term storage at 70°C. For work with microbial suspensions, microorganisms were separately grown aerobically in continuously shaked liquid cultures at 37°C in Schaedler medium until the early-stationary phase of growth. Cells were harvested by centrifugation (896 g, 18°C, 5 min; Hettich Rotixa P, Tuttlingen, Germany), washed twice with Dulbecco‘s Phosphate-Buffered Saline (PBS; Sigma-Aldrich, St. Louis, MO, USA), and resuspended in sterile Schaedler medium. The optical density of each microbial suspension was adjusted to 0.3 at 550 nm (Genesys 10-S; Thermo Spectronic, Rochester, NY, USA); subsequently, 1 ml of each microbial suspension was used to produce the mixed species microbial suspension employed for simulation of biofilm formation. Simulation of biofilm formation
Biofilm formation on the various specimens was simulated using a multispecies biofilm
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
Hahnel et al Biofilm formation on abutment materials
model. In brief, specimens were transferred to sterile 48-well cell clusters and pellicle formation was initiated by incubating the sample-containing plates with 1 ml per well of sterile human whole saliva at 37°C in a thermo shaking device (OrbitalShaker; Thermo Forma, Marietta, OH, USA). After 2 h, saliva was removed carefully, and each specimen was incubated with 1 ml of mixed species microbial suspension for 2.5 h. Subsequently, microbial suspensions were carefully removed, and each specimen was incubated with 1 ml of Schaedler medium. After 20 or 44 h of incubation, Schaedler medium was carefully removed and specimens were carefully washed twice with phosphate-buffered saline (PBS). Analysis of biofilm formation
The relative amount of viable microorganisms adherent to the different specimens was analyzed using an MTT-based cell viability assay as described previously (Ionescu et al. 2012; Hahnel et al. 2014). A total of 15 specimens were analyzed for each of the investigated materials and incubation times, including 10 test specimens and five samples that had been employed for control purposes (dye controls). In brief, MTT solution was prepared by dissolving 5 mg/ml thiazolyl blue tetrazolium bromide (Sigma-Aldrich) in PBS; PMS solution was prepared by dissolving 0.3 mg/ml phenazine methosulfate (Sigma-Aldrich) in sterile PBS. MTT reaction solution was prepared by freshly mixing 1 ml of MTT solution, 1 ml of PMS solution, and 8 ml of sterile PBS. Subsequent to biofilm formation and careful rinsing, 200 ll of the MTT reaction solution was added to each sample well and incubated under lightproof conditions at 37°C. After 5 h, the reaction solution was gently removed from each well, and 200 ll of a lysing solution consisting of 10% V/V sodium dodecyl sulfate, and 50% V/V dimethylformamide (both SigmaAldrich) in distilled water was added to each well. After 1 h of further incubation under light-proof conditions at 37°C, 180 ll of the aliquot was transferred to new sterile well plates, and the absorbance correlating with cell viability was measured at a wavelength of 550 nm (Fluostar Optima; bmg Labtech, Offenburg, Germany). For fluorescence microscopic analysis of the ratio between live and dead microorganisms biofilms adherent to the different surfaces, each of five specimens for each material and incubation time were randomly placed in a sterile plastic Petri dish and the biofilms were scraped off the surface of using
sterile cell scrapers. Subsequently, the surface of the specimens and the Petri plate was rinsed with PBS and the pooled washings were adjusted to a final volume of 1 ml, which was finally subjected to intense vortexing. An aliquot (25 ll) of the sonified suspensions for each material and incubation time was transferred onto microscopic slides containing Live/Dead BacLight bacterial viability kit solution for microscopy and quantitative assays (Molecular Probes Inc, Eugene, OR, USA). After incubation in the dark for 15 min, five randomly selected fields with an area of 3.8 9 10 8 m2 were analyzed for each of the five specimens for each material and incubation time using a fluorescence microscope (Zeiss Axio Scope A1 LED; Zeiss, Oberkochen, Germany) to determine the percentage of dead and inactive microorganisms (fluorescent red). Statistical procedures
All statistical analyses were performed using statistical software (IBM SPSS 21.0 for Windows; IBM, Armonk, NY, USA); the level of significance (a) was set to 0.05. One-way analysis of variance was used to analyze surface roughness and SFE of the various abutment materials investigated; the Tukey Kramer post hoc test was employed to highlight significant differences where appropriate. The non-parametric Kruskal–Wallis test was employed to investigate differences in cell viability and the percentage of dead microorganisms in the biofilms, setting the abutment material and incubation time as fixed factors. Subsequent post hoc analysis was performed using the Bonferroni-adjusted Mann–Whitney U-test where appropriate; the adjusted a‘ was set to 0.0125.
Results
Table 1. Surface roughness (Ra) and surface free energy (mJ/m2) of the different materials investigated in this study. Means and standard deviations are indicated; identical superscript letters within a single column indicate significant differences for a = 0.05 Material
Surface roughness
Zirconia Titanium PEEK PMMA
0.16 0.17 0.04 0.05
(0.02)a,b (0.03)c,d (0.02)a,c (0.03)b,d
Surface free energy 31.72 35.62 42.19 39.14
(4.43)a (4.26) (2.80)a (3.39)
Biofilm formation
Statistical analysis indicated significant differences in cell viability, suggesting significant differences in multispecies biofilm formation between the different materials after 20 h (P < 0.001). Lowest cell viability indicating the lowest amount of adherent viable biomass was identified on the surface of PEEK, which was significantly lower than for zirconia (P < 0.001), titanium (P = 0.004), and PMMA (P < 0.001). Cell viability increased significantly from 20 to 44 h of biofilm formation, indicating a significantly higher amount of viable biomass adherent to the abutment materials after prolonged biofilm formation (P < 0.001). After 44 h, no significant differences in cell viability were identified between the different abutment materials (P = 0.068) (cf. Table 2). Regarding fluorescence microscopic analysis of the percentage of dead microorganisms in the biofilms, significant differences were identified between the biofilms on the various materials after 44 h (P = 0.012) but not after 20 h (P = 0.500). After 44 h, the biofilms on zirconia yielding a significantly higher percentage of dead microorganisms than on PMMA (P = 0.002). Generally, a significantly higher percentage of dead microorganisms was identified in the biofilms after 44 h than after 20 h (P < 0.001).
Surface properties
One-way ANOVA indicated significant differences in surface roughness between the different materials (P < 0.001). Post hoc analysis identified significantly lower surface roughness for specimens made from PEEK and PMMA than for specimens made from titanium and zirconia (P < 0.001) (cf. Table 1). One-way ANOVA indicated significant differences in SFE between the various materials (P = 0.044). Subsequent post hoc analysis suggested that zirconia yielded significantly lower SFE than PEEK (P = 0.038); no significant differences were observed between the other materials.
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
Discussion Within the limitations of an in vitro study, the results of this research suggest rejection of the research hypothesis, supporting the assumption that biofilm formation is not increased on the surface of PEEK in comparison to the conventionally applied implant abutment materials zirconia and titanium. As oral biofilm formation is a very complex process, it is clear that the results gathered in this laboratory approach cannot be completely transferred to a clinical setting. Nevertheless, numerous parameters influencing oral biofilm formation have been simulated,
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Hahnel et al Biofilm formation on abutment materials
Table 2. Relative absorbance values indicating viable biomass in biofilms after 20 and 44 h and percentage of dead microorganisms in the different biofilms on the different materials. Means and standard deviations (SD) as well as medians are indicated; identical superscript letters within the median column for each time indicate no statistically significant differences between the various materials Cell viability
Percentage of dead microorganisms
20 h
44 h
20 h
44 h
Material
Mean (SD)
Median
Mean (SD)
Median
Mean (SD)
Median
Mean (SD)
Median
Zirconia Titanium PEEK PMMA
0.31 0.22 0.11 0.27
0.33a,b 0.23a,c 0.10 0.26b,c
0.36 0.50 0.29 0.47
0.37a,b,c 0.46a,d,e 0.30b,d,f 0.53c,e,f
59.12 52.48 59.92 56.27
55.74a,b,c 52.58a,d,e 58.37b,d,f 56.54c,e,f
74.87 68.82 63.72 60.17
77.01a,b 65.33a,c,d 65.52b,c,e 62.30d,e
(0.09) (0.09) (0.04) (0.09)
including the formation of an acquired salivary pellicle on the surface of the abutment materials and the use of a multi-species biofilm model. To keep the experimental conditions applied reproducible, saliva from a single healthy donor rather was used than pooled saliva from a multitude of persons. With regard to the maturing process of biofilms on dental implant surfaces, the conventional wisdom is that bacteria such as Streptococci and Actinomyces species prevail in the early phases of biofilm formation and prepare the ground for the subsequent adhesion of anaerobic and more pathogenic microorganisms, which are dominant in more mature biofilms after 48 h (Nakazato et al. 1989; Kolenbrander et al. 2010). This circumstance has been taken into consideration by simulating biofilm formation with a mixture of at least four early-colonizing microorganisms for a maximum of 44 h. The multi-species biofilm model included microorganisms such as Actinomyces naeslundii and Streptococcus gordonii, which are commonly counted among the early-colonizing microorganisms in the oral cavity (Periasamy & Kolenbrander 2009); for these two bacteria, Streptococcus mutans and Candida albicans, relevant interaction effects have been identified (Palmer et al. 2001; Pereira-Cenci et al. 2008; Bamford et al. 2009; Kolenbrander et al. 2010), which justifies their inclusion in a multispecies biofilm model. Although PEEK is increasingly employed as abutment material in contemporary implant dentistry, biofilm formation on the surface of PEEK has – to the knowledge of the authors – hardly been addressed. Laboratory studies have shown that biofilms can be reproducibly grown on the surface of PEEK (Williams et al. 2011), and it has been reported in a recent conference abstract that the adhesion and proliferation of oral streptococci is similar on the
(0.13) (0.22) (0.14) (0.23)
(15.70) (12.60) (15.20) (11.65)
surface of PEEK and a conventional resinbased composite (Kolbeck et al. 2013). The only study investigating microbial issues in implants supplied with PEEK abutments identified similar microbial counts and levels of periodontal pathogens in the peri-abutment region of implants supplied with PEEK and titanium healing abutments (Volpe et al. 2008); however, no direct investigation of the biofilms on the surface of the different abutments was performed. Analysis of biofilm formation in this study suggests lower biofilm formation on the surface of PEEK than on the conventionally applied abutment materials titanium and zirconia, which hints that – from a microbiological point of view – PEEK features favorable properties as definite abutment material. The conventional wisdom is that the surface properties of a material have a pronounced influence particularly on the early phases of biofilm formation. For the development of biofilms on the surface of dental implant surfaces, this circumstance has led to the postulation that it is particularly the initial adherence of early-colonizing microorganisms that has to be prevented (Subramani et al. 2009). It has been reported that the surface roughness and SFE of an implant surface have an effect on the initial adherence of microorganisms, suggesting that smooth surfaces and those with low SFE feature less microbial adherence than materials with higher surface roughness or SFE. For titanium surfaces, threshold values for surface roughness (Ra) ranging between 0.088 lm (Rimondini et al. 1997) and 0.2 lm (Bollen et al. 1997) have been identified, suggesting that lower values do not have a significant impact on biofilm formation. However, it is doubtful whether the differences in surface roughness values identified for the materials investigated in this study can be accounted for differences in biofilm formation: although
(11.53) (13.76) (10.71) (10.79)
the surface roughness of titanium and zirconia was significantly higher than the surface roughness of both PEEK and PMMA, biofilm formation did not correlate with these results as significantly more viable biomass was identified on PMMA than on PEEK. Although a relationship between the SFE of implant materials and microbial adherence has been proven in numerous studies (Teughels et al. 2006), other researchers assume that the relation is not as simple. Recent studies suggest that the surface composition and surface topography might impact the formation of biofilms to an even higher level (Ionescu et al. 2012; Hahnel et al. 2014), which might serve as an explanation for the poor correlation between surface properties and biofilm formation observed in this study. With regard to this aspect, future studies might set their focus on analyzing topographical differences between the various abutment materials and attempt to correlate their findings to differences in biofilm formation on the different materials. The results of this trial suggest that biofilm formation on PEEK is not higher than on conventionally applied abutment materials such as titanium and zirconia. Thus, from a microbiological point of view, PEEK abutments might be effectively employed as definite abutment material in implant dentistry. To support the data gathered in this laboratory pilot study, the authors are concurrently investigating the subgingival formation of biofilms on the surface of provisional abutments made from PEEK and titanium in a clinical study.
Acknowledgement:
Vestakeep i4 R was kindly provided by Evonik, Essen, Germany.
References Ausschill, T.M., Arweiler, N.B., Brecx, M., Reich, E., Sculean, A. & Netuschil, L. (2002) The effect
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of dental restorative materials on dental biofilm. European Journal of Oral Sciences 110: 48–53.
Bamford, C.V., d‘Mello, A., Nobbs, A.H., Dutton, L.C., Vickerman, M.M. & Jenkinson, H.F. (2009)
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Hahnel et al Biofilm formation on abutment materials
Streptococcus gordonii modulates Candida albicans biofilm formation through intergeneric communication. Infection and Immunity 77: 3696– 3704. Bollen, C.M., Lambrechts, P. & Quirynen, M. (1997) Comparison of surface roughness of oral hard materials to the threshold surface roughness for bacterial plaque retention: a review of the literature. Dental Materials 13: 258–269. Bumgardner, J.D., Adatrow, P., Haggard, W.O. & Norowski, P.A. (2011) Emerging antibacterial biomaterial strategies for the prevention of periimplant infectinos. International Journal of Oral and Maxillofacial Implants 26: 553–560. Busscher, H.J., Rinastiti, M., Siswomihardjo, W. & Van der Mei, H.C. (2010) Biofilm formation on dental restorative and implant materials. Journal of Dental Research 89: 657–665. Elter, C., Heuer, W., Demling, A., Hannig, M., Heidenblut, T., Bach, F.W. & Stiesch-Scholz, M. (2008) Supra- and subgingival biofilm formation on implant abutments with different surface characteristics. International Journal of Oral and Maxillofacial Implants 23: 327–334. Fuhrmann, G., Steiner, M., Freitag-Wolf, S. & Kern, M. (2014) Resin bonding to three types of polyaryletherketones (PAEKs) – Durability and influence of surface conditioning. Dental Materials 30: 357–363. Hahnel, S., Wastl, D., Schneider-Feyrer, S., Giessbibl, F.J., Brambilla, E., Cazzaniga, G. & Ionescu, A. (2014) Streptococcus mutans biofilm formation on experimental resin-based composites in dependence of surface treatment and S-PRG filler particle fraction. Journal of Adhesive Dentistry. doi: 10.3290/j.jad.a31800. [Epub ahead of print] Holmberg, K.V., Abdolhosseini, M., Li, Y., Chen, X., Gorr, S.U. & Aparicio, C. (2013) Bio-inspired stable antimicrobial peptide coatings for dental applications. Acta Biomaterialia 9: 8224–8231. Ionescu, A., Wutscher, E., Brambilla, E., SchneiderFeyrer, S., Giessibl, F.J. & Hahnel, S. (2012) Influ-
ence of surface properties of resin-based composites on in vitro Streptococcus mutans biofilm development. European Journal of Oral Sciences 120: 458–465. Kern, M. & Lehmann, F. (2012) Influence of surface conditioning on bonding to polyetheretherketon (PEEK). Dental Materials 28: 1280–1283. Kolbeck, C., Sereno, M., Rosentritt, M. & Handel, G. (2013) Biofilm formation on polyetheretherketone surfaces and cleaning options. IADR Seattle, No. 2353. Kolenbrander, P.E., Palmer, R.J., Jr, Periasamy, S. & Jabucovics, N.S. (2010) Oral multispecies biofilm development and the key role of cell-cell distance. Nature Reviews Microbiology 8: 471–480. Mombelli, A., M€ uller, N. & Cionca, N. (2012) The epidemiology of peri-implantitis. Clinical Oral Implants Research 23(Suppl. 6): 67–76. Nakazato, G., Tsuchiya, H., Sato, M. & Yamauchi, M. (1989) In vivo plaque formation on implant materials. International Journal of Oral and Maxillofacial Implants 4: 321–326. Norowski, P.A. & Bumgardner, J.D. (2009) Biomaterial and antibiotic strategies for peri-implantitis. Journal of Biomedical Materials Research Part B: Applied Biomaterials 88B: 530–543. Owens, D.K. & Wendt, R.C. (1969) Estimation of the surface free energy polymers. Journal of Applied Polymer Science 13: 1741–1747. Palmer, R.J., Jr, Kazmerzak, K., Hansen, M.C. & Kolenbrander, P.E. (2001) Mutualism versus independence: strategies of mixed-species oral biofilms in vitro using saliva as the sole nutrient source. Infection & Immunity 69: 5794–5804. Pereira-Cenci, T., Deng, D.M., Kranevald, E.A., Manders, E.M., Del Bel Cury, A.A., Ten Cate, J.M. & Crielaard, W. (2008) The effect of Streptococcus mutans and Candida glabrata on Candida albicans biofilms formed on different surfaces. Archives of Oral Biology 53: 755–764. Periasamy, S. & Kolenbrander, P.E. (2009) Mutualistic biofilm communities develop with Porphyro-
© 2014 John Wiley & Sons A/S. Published by John Wiley & Sons Ltd
monas gingivalis and initial, early, and late colonizers of enamel. Journal of Bacteriology 191: 6804–6811. Quirynen, M., Van der Mei, H.C., Bollen, C.M., Schotte, A., Marechal, M., Doornbusch, G.I., Naert, I., Busscher, H.J. & Van Steenberghe, D. (1993) An in vivo study of the influence of the surface roughness of implants on the microbiology of supra- and subgingival plaque. Journal of Dental Research 72: 1304–1309. Rimondini, L., Fare, S., Brambilla, E., Felloni, A., Consonni, C., Brossa, F. & Carrassi, A. (1997) The effect of surface roughness on early in vivo plaque colonization on titanium. Journal of Periodontology 68: 556–562. Stawarczyk, B., Beuer, F., Wimmer, T., Jahn, D., Sener, B., Roos, M. & Schmidlin, P.R. (2013) Polyetheretherketone – a suitable material for fixed dental prostheses? Journal of Biomedical Materials Research Part B: Applied Biomaterials 101: 1209–1216. Subramani, K., Jung, R.E., Molenberg, A. & H€ammerle, C.H.F. (2009) Biofilm on dental implants: a review of the literature. International Journal of Oral and Maxillofacial Implants 24: 616–626. Teughels, W., Van Asche, N., Sliepen, I. & Quirynen, M. (2006) Effect of material characteristics and/or surface topography on biofilm development. Clinical Oral Implants Research 17(Suppl. 2): 68–81. Volpe, S., Verrocchi, D., Andersson, P., Gottlow, J. & Sennerby, L. (2008) Comparison of early bacterial colonization of PEEK and titanium healing abutments using real-time PCR. Applied Osseointegration Research 6: 54–56. Williams, D.L., Woodbury, K.L., Haymond, B.S., Parker, A.E. & Bloebaum, R.D. (2011) A modified CDC biofilm reactor to produce mature biofilms on the surface of PEEK membranes for an in vivo animal model application. Current Microbiology 62: 1657–1663.
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