Introduction
Although dental implants are widely used to replace missing teeth without damaging adjacent teeth, they remain susceptible to peri-implant diseases. Peri-implant diseases are inflammatory conditions affecting the tissues surrounding dental implants. These conditions are generally classified into peri-implant mucositis and peri-implantitis according to the presence of supporting bone loss.
The definition of peri-implant disease was proposed in a consensus report from the 1st European Workshop on Periodontology [1]. Peri-implant mucositis was defined as a reversible inflammatory reaction in the soft tissues surrounding a functional implant, whereas peri-implantitis was defined as an inflammatory reaction associated with loss of supporting bone around an implant.
The prevalence of peri-implantitis has been reported to range from 10 to 43% at the implant level and from 20% to more than 56% at the patient level [2,3]. In peri-implantitis, the lesion extends apical to the pocket epithelium and contains large proportions of plasma cells and lymphocytes as well as high numbers of polymorphonuclear cells and macrophages [4]. Multiple factors can contribute to peri-implant disease, including poor oral hygiene, a history of periodontitis, excess cement, smoking, diabetes, absence of keratinized mucosa, implant characteristics, and occlusal overload [5]. However, anaerobic plaque bacteria are considered critical etiologic factors in the development of peri-implant diseases [6].
Various surgical and non-surgical approaches have been proposed for the treatment of peri-implantitis. In particular, implant surface decontamination has been regarded as an essential step for creating conditions favorable for peri-implant tissue stability. However, the predictability of treatment outcomes remains limited [7,8]. One possible reason is that mechanical and chemical decontamination procedures may alter the original implant surface characteristics and leave residual contaminants, thereby influencing subsequent cellular responses.
Previous studies have investigated the effects of mechanical treatments, such as metal and plastic curettes and air abrasive devices, as well as chemical treatments, such as hydrogen peroxide, ethylenediaminetetraacetic acid (EDTA), and citric acid. Titanium surfaces after contamination and decontamination may differ substantially from pristine titanium surfaces.
Surface modifications after treatment, such as changes in surface roughness and residual debris resulting from surface debridement, could affect cell attachment to the surface [9]. Implants may contain both smooth and rough surface regions that interact with surrounding tissues. Cellular responses may also differ depending on the surface characteristics. The response of periodontal ligament-human telomerase reverse transcriptase (PDL-hTERT) cells to each surface differs from that of osteoblasts. PDL-hTERT cells exhibited enhanced spreading and differentiation on smooth surfaces compared with rough surfaces. This pattern contrasts with the response observed in osteoblasts [10].
Stem cells are considered important in peri-implant tissue regeneration because of their capacity for osteogenic differentiation, which may contribute to favorable biological conditions for bone regeneration around implants. Therefore, understanding stem cell responses to decontaminated titanium surfaces may provide insight into biological conditions associated with peri-implant tissue wound healing.
Accordingly, the purpose of this study was to evaluate the effects of various mechanical and chemical decontamination methods on contaminated smooth and rough titanium surfaces. In addition, the viability and osteogenic differentiation of gingiva-derived mesenchymal stem cells (GMSCs) were examined to identify surface conditions that may support cellular responses favorable for peri-implant tissue wound healing under in vitro conditions.
Materials and Methods
1. Bacterial culture on pretreated titanium discs
Saliva was collected from five healthy patients, diluted 1:2 with phosphate-buffered saline (PBS), and filtered through a 0.2-μm filter. Titanium discs were coated with filtered saliva, air-dried, and sterilized under ultraviolet light for 24 hours. Untreated titanium discs were placed one in each well of a 24-well plate, and 1 mL of culture medium (brain heart infusion; Becton, Dickinson and Company) supplemented with 10 μg/mL hemin (Sigma-Aldrich Co.) and 0.2 μg/mL vitamin K (Sigma-Aldrich Co.) was added to each well. Porphyromonas gingivalis (ATCC 33277; American Type Culture Collection) was inoculated at 2 × cells/mL and incubated anaerobically in an atmosphere of 85% N2, 10% H2, and 5% CO2 at 37℃ for 2–3 days.
2. Isolation and culturing of GMSCs
Gingival tissues were collected from healthy patients. The study protocol was reviewed and approved by the Institutional Review Board of Seoul St. Mary’s Hospital, College of Medicine, Catholic University of Korea, Seoul, Republic of Korea (KC11SISI0348), and informed consent was obtained from the patients.
During the gingivectomy procedure, gingival tissue was obtained and immediately placed in sterile PBS (Welgene) with 100 U/mL penicillin and 100 μg/mL streptomycin (Sigma-Aldrich Co.) at 4℃. The tissues were de-epithelialized, minced, and digested in alpha-modified, minimal essential medium (α-MEM; Gibco) containing dispase (1 mg/mL; Sigma-Aldrich Co.) and collagenase IV (2 mg/mL; Sigma-Aldrich Co.). Cells were incubated at 37℃ in a humidified incubator with 5% CO2 in air. After 24 hours, the non-adherent cells were washed with PBS, replaced with fresh medium, and fed every 2–3 days.
3. Surface debridement with various instruments
Sandblasted and machined-surface titanium discs measuring 10 mm in diameter and 1 mm in thickness were used. To assess the effects of instrumentation, the discs were divided into 16 groups: (1) ultrasonic scaler with metal tip (Electro Medical Systems), (2) ultrasonic scaler with plastic tip (Electro Medical Systems), (3) ultrasonic scaler with metal tip (Satelec), (4) ultrasonic scaler with carbon tip (Satelec), (5) ultrasonic scaler with titanium tip (Satelec), (6) stainless steel brush (I brush; Neo Biotech), (7) titanium brush (Tigran brush; Tigran), (8) plastic brush (Gingibrush; Neobiotech), (9) air abrasive device (Perio flow; Electro Medical Systems), (10) hydrogen peroxide (3%), (11) citric acid (50%), (12) EDTA (24%), (13) tetracycline (50%), (14) no treatment on pristine discs, (15) no treatment on discs with a pellicle, and (16) untreated bacteria-contaminated discs. Three machined and three sandblasted, large-grit, acid-etched titanium discs were used for each group in each experiment; for Alizarin Red S staining (ARS), cell viability, and alkaline phosphatase (ALP) activity measurements, experiments were conducted independently, and three discs per group within each experiment were used for statistical analysis.
In groups (1)–(5), the surfaces of the titanium discs were instrumented with a total of 40 strokes by a single operator (S.I.L.). The scaler tip was positioned tangentially to the surface, and approximately 30 g of pressure was applied. Back-and-forth movements were performed in the same direction for 40 strokes. For groups (1) and (2), the ultrasonic scaler was used in mode P with a power setting of 3; for groups (3)–(5), a power setting of 3 was used. In groups (6)–(8), rotating brushes were applied to the implant surfaces. The stainless steel brush was used at 1,000 rpm, the titanium brush at 500 rpm, and the plastic brush at 2,000 rpm. The brushes were gently applied to the implant surfaces with a lateral force of 30 N for 20 seconds, followed by saline irrigation for 20 seconds. In group (9), the air abrasive device was applied to the surface for 10 seconds according to the manufacturer’s instructions. In groups (10)–(13), a cotton pellet was soaked in each solution and applied to the surface with 20 reciprocating motions, followed by saline irrigation for 20 seconds.
4. GMSCs seeding and osteogenic differentiation
A total of 5 × 104 GMSCs were seeded onto each treated titanium surface and cultured in osteogenic induction medium (StemPro® Osteogenesis Differentiation Kit; Gibco; Thermo Fisher Scientific, Inc.). The medium was replaced with fresh induction medium every 3–4 days. On day 21, Alizarin Red S staining (Sigma-Aldrich Co.) was performed to detect calcium formation.
5. Determination of cell viability
Cell viability was analyzed on days 4 and 16. WST-8 (2-[2-methoxy-4-nitrophenyl]-3-[4-nitrophenyl]-5-[2,4-disulfophenyl]-2H tetrazolium, monosodium salt) (Cell Counting Kit-8 [CCK]; Dojindo) was added to the cultures, which were then incubated for 1 hours at 37℃. The spectrophotometric absorbance of the samples was measured at 450 nm using a microplate reader (BioTek).
6. ALP activity assay
Cell cultures were maintained in α-MEM or osteogenic induction medium (StemPro® Osteogenesis Differentiation Kit; Gibco) in a humidified incubator at 37℃ with 5% CO2 and harvested on day 14. Cells were detached using trypsin (Gibco; Thermo Fisher Scientific, Inc.) and washed twice with PBS for 1 minute. ALP activity assays were performed using the Alkaline Phosphatase Activity Colorimetric Assay Kit (cat no. K412-500; BioVision, Inc.) according to the manufacturer’s protocol. The cells were resuspended with assay buffer, sonicated at 70–80% intensity for 1 minute at 4℃, and then centrifuged at 13,000 × g for 3 minutes at 4℃ to remove insoluble material. The supernatant was mixed with p-nitrophenylphosphate substrate (provided in the kit) and incubated at 25℃ for 60 minutes. The optical density of the resultant p-nitrophenol at 405 nm was determined spectrophotometrically.
7. ARS for mineralization
On day 21, cells on titanium surfaces were washed twice with PBS for 1 minute, fixed with 70% ethanol at room temperature for 15 minutes, rinsed twice with deionized water, and stained with ARS for 30 minutes at room temperature. To remove nonspecifically bound dye, the cultures were washed 3 times for 2 minutes with deionized water and then with PBS for 15 minutes at room temperature. Bound dye was solubilized in 10 mM sodium phosphate containing 10% cetylpyridinium chloride (Sigma-Aldrich; Merck KGaA) and quantified spectrophotometrically at a wavelength of 560 nm. General flow of the experiment is shown in Fig. 1.
8. Statistical analysis
Data are presented as the mean ± standard deviation. The Shapiro–Wilk test was used to assess normality, and one-way analysis of variance followed by Tukey’s post hoc test was used to determine statistically significant differences among groups. R (version 3.5.3) was used for all statistical tests, and p < 0.05 was considered statistically significant. Pearson’s correlation analysis was performed to assess the relationships among CCK, ALP, and ARS.
Results
1. Determination of cell viability (CCK assay)
Overall, changes in cell viability differed according to surface characteristics. Over time (day 4 to day 16), cell viability decreased on the rough surface in all groups, regardless of bacterial contamination (Fig. 2A). In contrast, on machined surfaces, viability increased over time, particularly in the decontaminated and uncontaminated (control A and C) groups (Fig. 2B); however, this increase was not observed in the untreated contaminated group (control B) (Fig. 2C). These findings suggest that reducing bacterial contamination on machined surfaces may allow GMSCs to resume proliferation.
2. ALP activity
On the rough surface, no significant differences in ALP activity were observed among the experimental and control groups (Fig. 3A). On the machined surface, no significant differences were observed among the groups except for the air abrasive device-treated group and control C (Fig. 3B).
3. ARS for mineralization
On the rough surface, no significant differences in calcium deposition were observed among the groups, including the control (Fig. 4A). On the machined surface, no significant differences were observed among the groups except for the Satelec plastic tip-treated group (Fig. 4B).
4. Correlation between cell viability, ALP activity, and calcium deposition
Correlation patterns differed according to surface characteristics. On the rough surface, no significant correlation (0.03) was observed between CCK (cell viability) and ALP activity; however, a positive correlation (0.38) was observed between CCK and calcium deposition (ARS), indicating that cell viability was associated with the degree of mineralization on rough surfaces (Fig. 5A). On the machined surface, no correlation (0.00) was observed between CCK (cell viability) and ALP activity. In contrast, a negative correlation (−0.35) was identified between ALP activity and calcium deposition (ARS), and no significant relationship (0.08) was observed between cell viability and calcium deposition (Fig. 5B).
Discussion
This study evaluated the effects of various decontamination methods on contaminated rough and machined titanium surfaces and their impact on GMSCs. Cellular responses differed according to surface type.
On rough surfaces, cell viability decreased over time regardless of contamination or treatment, whereas on machined surfaces, viability increased in both decontaminated and uncontaminated groups but not in untreated contaminated controls. These findings suggest that adequate bacterial reduction on machined surfaces may support GMSCs proliferation and create cellular conditions potentially favorable for peri-implant tissue wound healing under in vitro conditions.
Correlation analyses revealed surface-dependent patterns. On rough surfaces, GMSCs viability was positively correlated with calcium deposition, whereas on machined surfaces, ALP activity showed a negative correlation with calcium deposition. These findings indicate that titanium surface morphology may influence the relationship between GMSCs viability and osteogenic mineralization under in vitro conditions.
Although this in vitro model provided valuable insight into GMSCs behavior on decontaminated titanium surfaces, bacterial activity during GMSCs culture could not be fully controlled. Furthermore, the present findings should be interpreted as reflecting cellular and osteogenic responses under controlled experimental conditions rather than direct evidence of re-osseointegration. Future in vivo studies and advanced experimental models are needed to better reproduce clinical conditions and further evaluate biological environments associated with peri-implant wound healing.
