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ORIGINAL ARTICLE
Year : 2011  |  Volume : 1  |  Issue : 1  |  Page : 7-12

Structural analysis of in situ biofilm formation on oral titanium implants


Department of Prosthetic Dentistry and Biomedical Materials Science, Hannover Medical School, Hannover, Germany

Date of Web Publication2-Feb-2011

Correspondence Address:
Sebastian Grade
Department of Prosthetic Dentistry and Biomedical Materials Science, Hannover Medical School, Carl-Neuberg-Strasse 1, 30625 Hannover
Germany
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Source of Support: The German Research Foundation (SFB 599, D8), Conflict of Interest: None


DOI: 10.4103/0974-6781.76425

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   Abstract 

Background: The primary etiologic factor for peri-implant infections is the adhesion of biofilms on oral implant surfaces in the area of soft-tissue penetration. The aim of the present study was to examine in situ biofilm growth directly on implant-abutment surfaces without the use of oral splints and to determine the effect of intraoral abutment localization on biofilm growth.
Materials and Methods: Fifteen titanium healing abutments were inserted in six patients for 14 days. The newly formed supragingival biofilm on the titanium surface of the healing abutments was stained with fluorescent Live/Dead Baclight kit before examination by confocal laser scanning microscopy. The biofilm was scanned in terms of its surface coverage and thickness and different sites.
Results: The results show that the biofilm has a different structure in every patient, with the thickness of the biofilm structure ranging between 0 and 80 ΅m and the surface coverage between 0 and 97% of the abutment surface. There was similar biofilm surface coverage at different intraoral locations, whereas the biofilm was significantly thicker in the mandible as compared to maxillary implant abutments.
Conclusion: The method uniquely describes an effective way to depict biofilm development on implant surfaces in the supra- and sub-gingival regions.

Keywords: Biofilm growth, confocal laser scanning microscope, dental plaque, peri-implantitis


How to cite this article:
Grade S, Heuer W, Strempel J, Stiesch M. Structural analysis of in situ biofilm formation on oral titanium implants. J Dent Implant 2011;1:7-12

How to cite this URL:
Grade S, Heuer W, Strempel J, Stiesch M. Structural analysis of in situ biofilm formation on oral titanium implants. J Dent Implant [serial online] 2011 [cited 2020 Aug 6];1:7-12. Available from: http://www.jdionline.org/text.asp?2011/1/1/7/76425


   Introduction Top


Osseointegrated oral implants provide an effective reconstruction of dentition after tooth loss. [1] Titanium is the most common implant material due to its biocompatibility and the lack of allergic responses. [2] For successful long-term prosthetic rehabilitation, it is necessary to inhibit bacterial biofilm formation on implant surfaces, as these bacterial communities are the main sources of inflammation of the peri-implant mucosa and bone. Untreated peri-implantitis may lead to the loss of dental implants. [3] Treatment of biofilms is difficult, as bacteria in biofilms are more resistant to antibiotics than their representatives in the planktonic phase. To eliminate the biofilm, a 100- to 1000-fold higher antibiotic dose is necessary compared to the treatment of planktonic bacteria. [4]

The first phase of oral biofilm development consists in the formation of a salivary pellicle on all oral surfaces. This coating contains host proteins and glycoproteins which serve as adhesion molecules for several species of bacteria. [5] Streptococci (Streptococcus oralis, Streptococcus mitis) make up a high percentage of these first colonizers. [6],[7] These bacteria provide adhesion points for further bacterial genera, such as Actinomyces or Fusobacterium, which adhere by means of specific binding proteins on their outer cell membrane. [8],[9] In this second phase of biofilm development, the cellular concentration of chemical signals secreted by the colonizing bacteria reaches a critical point at which the cells start to synthesize and express exopolysaccharides (EPS). [10] These macromolecules make up the biofilm matrix, incorporating the bacteria. Provided by its protective matrix, the growing biofilm has high tolerance against the shear forces of the oral fluids and the movement of tongue and jaw. Furthermore, it provides various nutritional substances and is a diffusion barrier for bactericidal compounds. [11] After the development of the EPS matrix, the mature biofilm grows until parts of its cells detach. These can adhere to other regions of the oral cavity and elicit other bacterial colonies. Biofilms on the penetration of implants through the gingiva are the main etiologic factor for peri-implantitis.

In order to prevent the first stage of biofilm formation in this region, gingival cells must settle on the implant surface before bacteria are able to adhere to the same area. Therefore, future implant surfaces should suppress the first colonization of bacteria and at the same time provide an advantage for the gingival cells to cover the implant surface first. One promising approach consists in a change of the chemical structure of the implant surface. [12],[13] To develop effective implant surfaces in the supra- and sub-gingival region, characterization of biofilm formation in this region is necessary. Many studies have been conducted to characterize the spatial structure of oral biofilms with confocal laser scanning microscopy (CLSM). [14] The biofilms examined had been developed either in vivo in batch cultures or in vivo on oral splints. Both methods only inadequately reflect the conditions for the biofilm development on oral implants because in vivo conditions orally (with its diversity of biofilm micro-flora) are different from in vivo models and the splint model is confounded by tongue and cheek activity. Extrapolation of results generated by use of these methods on peri-implant structures is not possible.

A recent study with healing abutments inserted during conventional prosthetic treatment shows that this method allows biofilm growth under realistic conditions, including shear forces and nutrients, but no data about biofilm thickness were obtained. [15]

The present study provides a structural analysis of oral biofilm growth in vivo under the aforementioned conditions. Additionally, the impact of the intraoral abutment location on biofilm growth was evaluated.


   Materials and Methods Top


Abutment preparation

Healing abutments (Astra GmbH, Moelndal, Sweden and Straumann AG, Basel, Switzerland) were ground manually on four sides with 1700 grit silicon carbide rotary disks (MD-Pan, Struers GmbH, Willich , Germany) to provide a flat area for the analysis of the surface coverage.

After treatment with a cloth mop and polishing compound containing amorphous silicon carbide (MD-Cloth/DiaPro, Struers GmbH, Germany), the average roughness of the area lied between Ra = 0.05 and 0.2 μm. After cleaning and sterilization, the abutments were inserted onto the osseo-integrated fixtures, where they remained for 14 days.

Patients

Six patients, three men and three women, aged 27-78 years (54 ± 21 years), who were treated at the Department of Prosthetic Dentistry and Biomedical Material Science, Hannover Medical School, Germany, participated in this study. The study protocol was approved by the ethics committee of Hannover Medical School (No. 3791). The examination was performed with the understanding and written consent of each subject.

Criteria for exclusion were a history of periodontitis and probing depth of the remaining dentition being more than 3 mm. All patients were in good systemic health and none had received antibiotic therapy. All participants were permitted to proceed with their regular oral hygiene, but not to use anti-bacterial mouth rinse. In total, 15 titanium abutments were inserted in six patients in different sites. The orientation of the four areas of each abutment was recorded.

Confocal laser scanning microscopy

After 14 days, the healing abutments were removed and prepared for CLSM by fluorescent staining using the Live/Dead BacLight bacterial viability kit (Invitrogen, Carlsbad, USA) containing the dyes Syto 9 (yellow) and propidium iodide (blue) that both stain nucleic acids. Syto 9 can penetrate cell membranes, thereby highlighting both living and dead bacteria, whereas propidium iodide cannot penetrate intact membranes and marks only dead bacteria, thereby reducing the Syto 9 fluorescence.

The abutments were submerged into 1 ml phosphate buffered saline (PBS) and 1 μl of every dye for 25 minutes at 21C. Then, they were examined on a Leica TCS LFSA confocal microscope (Leica Microsystems, Heidelberg, Germany) with an excitation wavelength of 488 nm (He/Ne-Laser). The emitting light of the fluorescent dyes was detected between 495 and 540 nm for Syto 9 and between 595 and 750 nm for propidium iodide. For the determination of the biofilm coverage, the whole polished surface was scanned with a lens of Χ10/0.3 numerical aperture. The thickness of the biofilm was determined by recording 10 slides in vertical scan mode (xz) using a Χ40/0.8 numerical aperture with 30 μm intervals between each vertical scan.

Image analysis of biofilm thickness

Image analysis was performed using Leica LCS lite Software 2.6 (Build 2611537). LCS lite was used to measure the biofilm thickness by determining the distance between the titanium surface and the biofilm-liquid interface on five fixed positions per vertical scan [Figure 1].
Figure 1 :Measurement of biofilm thickness with Leica LCS lite Software (40×); the horizontal line shows the abutment surface. Biofilm thickness was measured at five fixed positions (−50 to +50 ìm); specific values are shown above the vertical line

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Image analysis of biofilm surface coverage

The pictures of the biofilm and the polished surface [[Figure 2]a and b] were used to determine the biofilm surface coverage. Only biofilm on the polished surface was included. For this measurement, the images were placed on top of each other with Adobe Photoshop CS 2 (Adobe Systems Inc., San Jose, USA). The coverage value was determined from the ratio between blue pixels and the number of all pixels within the polished measurement area [Figure 2].
Figure 2 :Graphical analysis of the biofilm surface coverage: (a) selected surface of the biofilm stained with propidium iodide; (b) polished surface of the titanium abutment included in the analysis; (c) montage of picture in a and b, only the biofilm covering the polished area of the titanium surface is used for coverage calculation

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Statistical analysis

For each of the polished areas on each abutment, the percentage of the surface covered by the biofilm as well as the average value of the biofilm thickness measurements were determined.

Surface coverage and thickness were compared according to the orientation of the area (oral/vestibular), to the localization of the implant (anterior teeth/posterior teeth), and to the jaw (upper/lower). The comparisons were performed by a Mann-Whitney U-test using the SPSS statistical package (SPSS Inc., Chicago, USA). Statistical significance was assumed for P ≤ 0.05.


   Results Top


All healing abutments of the six patients were used for analyzing the biofilm thickness and surface coverage. Confocal imaging of the analyzed abutments showed that the oral biofilms were heterogeneous, both within different oral localizations and between patients [Figure 3].
Figure 3 :XYZ display of oral biofilm stained with Live/Dead Baclight Kit (40× resolution) (Imaris Software, Bitplane)

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Biofilm thickness

The mean thickness of all examined biofilms was 10.9 ± 13.1 μm. The mean thickness for mandibular abutments (15.1 ± 16.4 μm) was greater than that for maxillary abutments (7.2 ± 8.4 μm). This difference was statistically significant (P = 0.015). There were no statistically significant differences between the oral and vestibular positions with respect to mean biofilm thickness. Biofilms in the oral position had a mean thickness of 11.9 ± 10.9 μm, compared with 9.8 ± 15.5 μm in the vestibular position. The mean values for anterior (8.1 ± 9.4 μm) and posterior areas (12.3 ± 14.7 μm) showed no statistically significant differences.

Biofilm surface coverage

The surface coverage of the measured dental biofilms ranged between 0 and 95%, with a mean value for coverage of 35.9 ± 28.3%. Comparison of coverage values showed no statistically significant differences for the regional distinctions in the oral cavity [Table 1].
Table 1 :Mean ± SD values of biofilm thickness and surface coverage at different locations in the upper jaw, the lower jaw, the anterior teeth in the upper jaw, the posterior teeth in the upper jaw and for the oral and vestibular positions


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The biofilm on lower jaw abutments showed a mean coverage of 38.9 ± 27.0%. For biofilms on upper jaw abutments, this was 33.2 ± 29.5%. The mean coverage in the oral position was 35.2 ± 22.4%, compared with 36.7 ± 34.2% for vestibular biofilms. The biofilm coverage on anterior teeth was measured as 35.4 ± 27.4%, whereas the value for posterior teeth was 36.2 ± 29.1%.


   Discussion Top


CLSM is an effective tool to characterize biofilms in their native condition due to the possibility of displaying a three-dimensional image of these bacterial communities. Using software tools, a virtual cut through the biofilm can be performed that allows the measurement of its thickness without affecting the real object. Furthermore, this imaging can be achieved without chemical or physical preparation that would irreversibly damage the biofilm structure. Based on these advantages, CLSM has become a major instrument for microbiological research to analyze biofilms generally, [14],[16] and has been recently used to specifically examine oral biofilm communities. [17],[18],[19],[20]

The use of healing abutments is an effective method for structural analysis of in situ biofilms, which have developed under realistic conditions. The influence of nutrition, peri-implant tissue and shear forces on the location complies with the environmental parameters of the oral cavity. Another important aspect is that probe extraction is painless for the patients participating in the conventional prosthetic treatment. Other studies have utilized experimental appliances which could be inserted reversibly in the mouth. [21],[22],[23],[24] However, these did not provide realistic locations for the brackets studied, as biofilms which developed on these surfaces are located toward the natural teeth. These surfaces were less influenced by the mechanical movement of the masticatory muscles. In the present study, the spatial structure of in vivo dental biofilms was obtained for the first time from healing abutments and analyzed by the combined use of CLSM and software analysis. Earlier studies showed that bacterial adhesion was affected by the properties of the material used, including free surface energy and surface roughness. In order to minimize these influences, the examined abutment areas were polished to provide mean roughness lower than a threshold of Ra = 0.2, a value which is presumed not to affect bacterial adhesion by the above mentioned factors. [25],[26]

The mean values for biofilm thickness were lower than that obtained in other studies, [27],[28],[29],[30],[31] even though the incubation period for in situ biofilm development in the present study (14 days) was considerably longer than that in other studies (between 2 and 7 days). This results from the fact that in this study, the patients were allowed to maintain their normal dental hygiene, whereby the developing biofilm could be damaged by the shear forces of a tooth brush, for instance. In the above mentioned studies, the participants wearing the splints were instructed not to conduct any mechanical cleaning in the oral cavity. Also, the examined surfaces were protected from the shear forces of the tongue and jaw by their orientation within the splint toward the dental surface. Therefore, biofilm thickness in the present study can be assumed to describe a more realistic dental biofilm. Some other studies reported lower values of biofilm thickness due to a different method used to prepare the biofilm for examination, i.e., the biofilm was dried and embedded. [17],[22] The drying process resulted in a 50% loss of biofilm thickness.

In this study, no statistical significant differences in biofilm thickness could be found for the various locations of vestibular/oral positions; the same applies for locations on posterior/anterior teeth. However, a statistically significant thicker biofilm was detected in the mandible in comparison to the maxilla. This difference was probably due to tongue movement, the quantity of saliva or the variable effectiveness of mechanical abrasion during dental hygiene between the upper and lower jaw. In contrast to our results, Arweiler et al, [21] found no statistically significant differences for these areas. This could be attributed to the fact that their study used a removable appliance, which had different geometrical conditions compared to the healing abutments used in the present study. Furthermore, the influence of longer incubation times and continuous oral hygiene without short-term removal of the specimen has to be considered. Comparison of the biofilm surface coverage between the different sites showed no statistically significant differences. It can be assumed that the lateral mechanical forces which were generated by jaw and tongue motion indeed influenced the thickness of the biofilm by abrasion of its superficial layers. However, these forces seemed not to remove all bacteria from the titanium surface if they are protected by the EPS-matrix. Although the mechanical abrasion reduced the biofilm thickness, the remaining bacteria on the surface were the base for a new biofilm with the same surface coverage. Consistent with other authors, our study showed wide individual variability of the measured values explains the high standard deviations of the data. [28],[32],[33] This variability results from the variable composition of sulcular fluid, nutrition and oral bacterial community between individual patients. [34]

In conclusion, this study demonstrated similar biofilm surface coverage at different intraoral locations but significantly higher biofilm thickness on mandibular than maxillary implant abutments. The obtained data may be used in further studies to evaluate the influencing factors for the development of oral biofilms. It has been shown that the described method for analyzing dental biofilms in situ on prosthetic healing abutments is more realistic in displaying bacterial communities under physiological conditions in the oral cavity.

 
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