UV-photofunctionalization of a biomimetic coating for dental implants application

Materials Science & Engineering C 110 (2020) 110657

Contents lists available at ScienceDirect

Materials Science & Engineering C journal homepage: www.elsevier.com/locate/msec

UV-photofunctionalization of a biomimetic coating for dental implants application

T

Caroline Dinia, Bruna E. Nagaya, Jairo M. Cordeiroa, Nilson C. da Cruzb, Elidiane C. Rangelb, ⁎ Antônio P. Ricomini-Filhoc, Erica D. de Avilad, Valentim A.R. Barãoa, a

Department of Prosthodontics and Periodontology, Piracicaba Dental School, University of Campinas (UNICAMP), Av. Limeira, 901, Piracicaba, São Paulo 13414-903, Brazil b Institute of Science and Technology, São Paulo State University (UNESP), Av. Três de Março, 511, Sorocaba, São Paulo 18087-180, Brazil c Department of Physiological Science, Piracicaba Dental School, University of Campinas (UNICAMP), Av. Limeira, 901, Piracicaba, São Paulo 13414-903, Brazil d Department of Dental Materials and Prosthodontics, School of Dentistry at Araraquara, São Paulo State University (UNESP), R. Humaitá, 1680, Araraquara, São Paulo 14801-903, Brazil

A R T I C LE I N FO

A B S T R A C T

Keywords: Titanium Ultraviolet rays Dental implants Photofunctionalization Plasma electrolytic oxidation Biofilm

Photofunctionalization mediated by ultraviolet (UV) rays changes the physico-chemical characteristics of titanium (Ti) and improves the biological activity of dental implants. However, the role of UV-mediated photofunctionalization of biofunctional Ti surfaces on the antimicrobial and photocatalytic activity remains unknown and was investigated in this study. Commercially pure titanium (cpTi) discs were divided into four groups: (1) machined samples without UV light application [cpTi UV−]; (2) plasma electrolytic oxidation (PEO) treated samples without UV light application [PEO UV−]; (3) machined samples with UV light application [cpTi UV+]; and (4) PEO-treated samples with UV light application [PEO UV+]. The surfaces were characterized according to their morphology, roughness, crystalline phase, chemical composition and wettability. The photocatalytic activity and proteins adsorption were measured. For the microbiological assay, Streptococcus sanguinis was grown on the disc surfaces for 1 h and 6 h, and the colony forming units and bacterial organization were evaluated. In addition, to confirm the non-cytotoxic effect of PEO UV +, human gingival fibroblast (HGF) cells were cultured in a monolayer onto each material surface and the cells viability and proliferation evaluated by a fluorescent cell staining method. PEO treatment increased the Ti surface roughness and wettability (p < 0.05). Photofunctionalization reduced the hydrocarbon concentration and enhanced human blood plasma proteins and albumin adsorption mainly for the PEO-treated surface (p < 0.05). PEO UV+ also maintained higher wettability values for a longer period and provided microbial reduction at 1 h of bacterial adhesion (p = 0.012 vs. PEO UV-). Photofunctionalization did not increase the photocatalytic activity of Ti (p > 0.05). Confocal microscopy analyses demonstrated that PEO UV+ had no cell damage effect on HGF cells growth even after 24 h of incubation. The photofunctionalization of a biofunctional PEO coating seems to be a promising alternative for dental implants as it increases blood plasma proteins adsorption, reduces initial bacterial adhesion and presents no cytotoxicity effect.

1. Introduction Dental implant rehabilitation has been widespread due to its high survival rate predictability of up to 98% [1]. These results are related to the use of titanium (Ti) and its alloys, which are the main biomaterials used for dental implant manufacturing considering their biocompatibility, corrosion resistance and osseointegration ability [2]. However, infections of peri-implant surrounding soft tissues, defined as mucositis, can occur and have become a challenge, mainly in the early healing

⁎

stage of dental implants, with a prevalence of 46.83% [3–5]. The development of infections during the early healing stage is considered a risk factor for the osseointegration process, inducing higher rates of dental implant early failures [6,3]. The early failures due to bacterial infections are influenced by the surface properties of the biomaterials, such as the chemical composition, surface free energy and surface roughness, which are important determinants of biofilm formation on dental implants [7]. Moreover, the surface properties influence the acquired pellicle formation, which

Corresponding author at: Av. Limeira, 901, Piracicaba, SP 13414-903, Brazil. E-mail address: [email protected] (V.A.R. Barão).

https://doi.org/10.1016/j.msec.2020.110657 Received 25 April 2019; Received in revised form 20 December 2019; Accepted 7 January 2020 0928-4931/ © 2020 Elsevier B.V. All rights reserved.

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

implant surfaces [28]. Therefore, surface modifications and improvements of biological properties are relevant approaches for the success of dental implant therapy. The bone biomimetic coating obtained by PEO is a well-established surface treatment. Likewise, photofunctionalization has demonstrated relevant results for dental implant applications. However, the photocatalytic and antimicrobial activities of photofunctionalized biofunctional surfaces have not been previously tested. Thus, in this study, a biofunctional PEO coating for dental implants was developed, and the influence of UV-mediated photofunctionalization on the physico-chemical characteristics, photocatalytic activity, total human blood plasma proteins and albumin adsorption as well as antimicrobial function and cytotoxicity of Ti material were investigated. It was hypothesized that photofunctionalization of the biofunctional PEO coating would improve the surface characteristics by enhancing proteins adsorption, reducing initial bacterial adhesion and presenting no cytotoxicity effect.

acts as a receptor for bacterial adhesion [8–10]. Streptococcus spp. are initial colonizers that are capable of providing additional sites for secondary colonizer adhesion, favoring biofilm development [11,12]. Approaches to prevent the initial attachment of early colonizers in the initial healing stage are important strategies to obtain a more reliable treatment with dental implants [10]. Another important factor for the dental implants surface is the ability to promote osseointegration. Albumin is a major blood plasma protein, and its adsorption is the first critical step for the establishment of osseointegration, since this protein provides the attachment of functional cells [12]. Albumin regulates cytoplasmatic calcium and cellular proliferation of osteoblasts, accelerating the osseointegration process [13]. The albumin adsorption and other human blood proteins are also influenced by dental implants surface characteristics such as roughness, topography, carbon amounts, covalent bonds and electrostatic status [7,14–16]. For this reason, topographical and chemical surface modifications of Ti material have been studied to enhance the biological properties and antimicrobial activity of dental implants. Plasma electrolytic oxidation (PEO) is a technique that is used to improve the bioactivity, osseointegration and corrosion resistance of dental implants [17,18]. PEO is an electrochemical process in which the samples are immersed in an electrolytic solution with micro plasma discharges on the metal surface [17,19]. PEO coatings allow the formation of micropores, the incorporation of chemical elements such as calcium (Ca) and phosphorous (P) [20] and the creation of titanium dioxide (TiO2) in different crystalline structures as anatase and rutile [21]. The photofunctionalization treatment of dental implants surfaces with ultraviolet (UV) light has also been successfully used because it induces physico-chemical modifications and improves the biological properties of Ti surfaces [22]. Although direct exposure to UV radiation on dental implants after placement has harmful effects [23], pretreatment before implant placement (i.e., photofunctionalization) is an alternative to UV light application that avoids damaging human cells [24,25]. The UV photofunctionalization increases protein adsorption and adhesion, as well as the proliferation, differentiation and mineralization of osteogenic cells in vitro [26,27]. In addition to being hydrophobic after aging, because of hydrocarbon accumulation, Ti becomes superhydrophilic after UV photofunctionalization by the removal of hydrocarbons from the surface, favoring water adsorption by the creation of oxygen vacancies as a result of the reduction of Ti4+ to Ti3+ [24]. When TiO2 surfaces containing crystalline structures are exposed to UV light, a photocatalytic activity can be produced that promotes antimicrobial function [28,29]. The mechanism of the photocatalytic activity comprises the generation of reactive oxygen species %OH, O2%ˉ and H2O2, which are responsible for the outer membrane decomposition of microorganisms [30,31]. Nevertheless, there are few reports regarding the photocatalytic activity on photofunctionalized surfaces, which is a possible strategy to generate reactive oxygen species since the photofunctionalization includes UV light application on dental

2. Materials and methods 2.1. Experimental design Commercially pure titanium (cpTi) discs were randomly divided into four different groups: (1) machined samples without UV light application [cpTi UV−]; (2) PEO-treated samples without UV light application [PEO UV−]; (3) machined samples with UV light application [cpTi UV+]; and (4) PEO-treated samples with UV light application [PEO UV+]. The surface roughness (n = 5), surface morphology (n = 1) and crystalline phase (n = 1) were assessed only for the control groups (cpTi UV− and PEO UV−), since these parameters are not influenced by photofunctionalization (pilot study – data not shown). The control (UV−) and experimental groups (UV+) were evaluated by means of wettability (n = 5), chemical composition by energy dispersive spectroscopy (n = 1) and X-ray photoelectron spectroscopy (n = 3 for atomic concentration (%) and n = 1 for XPS spectra), human blood plasma proteins and albumin adsorption (n = 5), photocatalytic activity (n = 3) and microbiological assay (n = 6 in triplicate with two independent repetitions). For the microbiological assay, Streptococcus sanguinis was grown onto the disc surfaces for 1 h and 6 h of bacterial adhesion. The number of colony forming units (CFU) and the bacterial organization were assessed. In addition, a cell culture assay using human gingival fibroblast cells was performed to investigate the surfaces biocompatibility by live/dead fluorescence assay (Fig. 1). 2.2. Preparation of titanium discs and surface treatment CpTi discs [American Society for Testing and Materials (ASTM)—Grade II] (Realum Ind e Com de Metais Puros e Ligas Ltd., São Paulo, SP, Brazil), with a diameter of 10 mm and a thickness of 2 mm, were used. Discs were polished by means of an automatic polisher (EcoMet/AutoMet 250 Pro, Buehler, Lake Bluff, IL, USA) at 250 rpm for

Fig. 1. Schematic diagram of the experimental design. The surface morphology, surface roughness and crystalline phases were assessed only for the control groups cpTi UV− and PEO UV−. 2

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

2.3.5. Energy dispersive spectroscopy (EDS) The chemical composition of the surface was evaluated by energy dispersive spectroscopy (EDS) (JEOL JSM-6010LA, Peabody, Massachusets, USA) operating at 5 kV and 2000× of magnification. Three randomly regions were selected to perform the EDS in small volumes, in the order of 1 μm3 [37].

1 min each with SiC abrasive papers (Nos. #320 and #400) (CarbiMet 2, Buehler, Lake Bluff, IL, USA). Then, the discs were cleaned and degreased with alternate ultrasonic baths in distilled water and 70% propanol for 10 min each and air-dried [17,21,32]. This procedure characterized the machined surface. PEO surface treatment was carried out using a pulsed DC power supply (Plasma Technology, Ltd., Hong Kong, China). The treatment was performed in a steel tank with a cooling system consisting of the temperature of the electrolytic solution maintained at 20 °C. The electrolytic solution was prepared by the dissolution of calcium acetate [Ca (CH3CO2)2] (Dinâmica Ltd., Diadema, SP, Brazil) and glycerophosphate disodium (C3H7Na2O6P) (Dinâmica, Ltd., Diadema, SP, Brazil) in distilled water at a proportion of 0.3 M/0.02 M, respectively. A voltage of 290 V, frequency of 250 Hz and duty cycle of 60% were set, and the treatment was carried out for 10 min. Samples were completely submerged in the electrolytic solution. After being submitted to the PEO treatment, the discs were rinsed in distilled water [17,21]. The UV-mediated photofunctionalization was carried out under ambient conditions for 96 h in a customized apparatus immediately before each analysis [33]. The light source used was UV-C 2 × 15 W (λ = 253.7 nm) (Moran Light, São Paulo, SP, Brazil), which was fixed 7 cm perpendicularly to the discs with an intensity of 2.34 mW/cm2 to achieve disc surface. The light intensity was measured using a photometer (Ophir Optronics Solution Ltd., Jerusalem, Israel) [34,35].

2.3.6. X-ray photoelectron spectroscopy (XPS) The chemical composition of the outmost oxide layer was evaluated by X-ray photoelectron spectroscopy (XPS) (K-alpha X-ray XPS, Thermo Scientific™, Vantaa, Helsíque, Finland) with a spot size of 400 μm and 1000 eV of energy step size [14,18]. Detailed survey scans of the main characteristic peaks (Ti 2p, C 1s, O 1s, Ca 2p and P 2p) were performed. Surface charging was corrected by the C1s binding energy of CeC bond at 284.7 eV, which was used as an internal reference. Any remaining small shift was removed after data collection, by using the spectrum processing tools contained within the Thermo Scientific Avantage Data System. The identification and correction of the peaks were based on the National Institute of Standards and Technology (NIST) database for the Simulation of Electron Spectra for Surface Analysis (SESSA) [38]. 2.3.7. Wettability The wettability properties of the Ti discs were measured by the contact angle approach using 2-μL drops of distilled water and a goniometer (Rame-Hart 100-00; Rame-Hart Instrument Co., Succasunna, NJ, USA) associated with software (DROPimage Standard, Ramé-Hart Instrument Co., Succasunna, NJ, USA). The sessile drop method was used, and 10 measurements were performed for each sample. The image obtained was immediately captured by the device, and the contact angle formed by the liquid on the sample surface was automatically measured [17]. The contact angle was evaluated for the UV− and UV+ groups immediately after termination of the 96 h UV light application. For analysis of the hydrophilic status, the cpTi and PEOtreated surfaces without UV treatment (UV−) and photofunctionalized (UV+) were maintained in dark and dry conditions, and the contact angle was measured for 6 h at 1-h intervals.

2.3. Surface analyses Profilometry, scanning electron microscopy (SEM), laser scanning confocal microscopy (LSCM) and X-ray diffraction (XRD) were used to better understand the differences between the surfaces, but only for the UV− groups since the morphological, roughness and topographic features are not changed by UV light application [26]. Contact angle, energy dispersive spectroscopy (EDS) and X-ray photoelectron spectroscopy (XPS) were also performed to characterize the surface in all groups. For these analyses, the UV+ experimental groups were used immediately after the UV light treatment.

2.4. Photocatalytic activity assay

2.3.1. Scanning electron microscopy (SEM) To observe the topography and morphology of the discs, scanning electron micrographs were acquired in a JEOL JSM-6010LA system (Peabody, Massachusetts, USA) using the secondary electrons detector and 4 kV of beam energy (2000× of magnification) [14].

The photocatalytic potential was measured by the methylene blue (MB) ISO technique [39,40] to understand the photocatalytic activity on photofunctionalized surfaces. The samples in the experimental groups were previously photofunctionalized. The control groups were not exposed to UV light. The specimens were soaked in 2 mL of standard MB solution (P.A.C.I. 52.015, Synth, Diadema, SP, Brazil) and maintained in dark conditions (foil wrapped) for 30 min prior to the test for the pre-adsorption measurement [41]. Afterwards, the discs were transferred to 24-well plates with 2 mL of fresh MB solution (10 μmol/ L) in each well. The MB degradation was analyzed in different periods (15 min, 30 min, 1 h, 2 h, 4 h and 6 h) to observe the survival of the photocatalytic potential. MB degradation was measured spectrophotometrically (DU 800 – Beckman Coulter, Brea, CA, USA) by sampling the solution and returning the sample after measurement of the solution's absorbance at 664 nm [39]. The photocatalytic activity was calculated using the following equation: Photocatalytic activity (%) = [(co − c)/co] × [c1/co] × 100. The concentration of the test solution of MB is represented by co, where c is the concentration of MB in the control and test samples previously photofunctionalized and c1 is the concentration of MB after the pre-adsorption test [29,40].

2.3.2. Laser scanning confocal microscopy (LSCM) A noncontact 3D Laser Scanning Microscope (LSCM, Keyence model VK-X200 series, Osaka, Osaka, Japan) was used to characterize the 2D and 3D surface topography of samples (150× of magnification). The image processing and average surface area measurements were assessed using VK-Analyzer software (Keyence v 3.3.0.0, Osaka, Osaka, Japan) [14]. 2.3.3. Profilometry The surface roughness was analyzed by means of a profilometer (Dektak 150-d; Veeco, Plainview, NY, USA). The arithmetic (Ra), maximum height (Rt), average peak-to-valley height (Rz), and root mean square average (Rq) were measured at a cut-off of 0.25 mm and speed of 0.05 mm/s for 12 s. Three measurements were performed at different areas of each sample and then averaged [36].

2.5. Protein adsorption assay

2.3.4. X-ray diffraction (XRD) The oxide crystalline phases were analyzed with an X-ray diffractometer (XRD; X'Pert PRO MRD, PANalytical, Almelo, Overijssel, The Netherlands) using CuKα radiation (λ = 1,540,598 Å) at 45 kV and 40 mA, a speed of 0.02°/s, a fixed angle of 2.5° and variation of 20° to 90° [37].

For an understanding of how proteins present in blood serum interact with different surface treatments, adsorption of human blood plasma proteins and bovine serum albumin (Sigma-Aldrich, St. Louis, MO, USA) were investigated separately, and the adsorption was 3

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

forming units per mL (log CFU/mL) [14].

measured by the bicinchoninic acid method (BCA Kit, Sigma-Aldrich, St. Louis, MO, USA) [21,42]. Human plasma from healthy donors was gently provided by Blood Center of University of Campinas (UNICAMP, Brazil). This study was approved by the Local Research and Ethics Committee (60177416.4.0000.5418). Photofunctionalized and nonphotofunctionalized discs were incubated in 24-well culture plates containing 1 mL of human blood plasma or 100 μg/mL of albumin in phosphate-buffered saline (PBS) (Gibco, Life Technologies, Grand Island, NY, USA) with horizontal stirring (75 rpm) at 37 °C. After 2 h of incubation, the protein solution was aspirated, and the discs were washed three times in PBS to remove nonadherent proteins [21]. The discs were transferred to cryogenic tubes containing 1 mL of PBS and sonicated in a Cup Horn (5.5 in. cup, Q500, Qsonica, Newtown, Connecticut, USA) at an amplitude of 80% for 60 s. The discs were then vortexed for 60 s to remove the absorbed protein from the disc surface. For human blood plasma proteins, the solution was diluted 10-fold. In a 96-well microtiter plate, 150 μL of the sonicated suspension was added to 150 μL of a mixture of reagents A and B. The microplate was incubated at 60 °C for 1 h, after which the absorbance of the solution was measured in a microplate spectrophotometer reader (Multiskan, Thermo Scientific, Vantaa, Helsíque, Finland) at a wavelength of 562 nm, according to the manufacturer's recommendations. The protein concentration was calculated based on a standard curve prepared with bovine serum albumin (0.5–30 μg/mL of protein) [21].

2.6.2. Scanning electron microscopy (SEM) The bacterial structure and organization for each time of incubation (1 h and 6 h) was evaluated by SEM (JEOL-JSM-5600LV, Peabody, Massachusets, USA) operating at 15 kV with 5000× magnification. Prior to the SEM analysis, the bacterial was fixed in Karnovsky solution (PBS, pH 7.2) for 2 h and serially dehydrated with ethanol washes (50%, 60%, 70%, 90% and three washes in 100% solution for 5 min each). Then, the discs were dried aseptically and gold-sputtered before SEM imaging [45]. 2.6.3. Cell culture assay Human gingival fibroblast cells (HGF - Rio de Janeiro Cell Bank Code 0089) were grown in low-glucose Dulbecco's modified Eagle medium (DMEM, Sigma Chemical Co., St. Louis, MO) supplemented with 10% fetal bovine serum (FBS, Gibco, Grand Island, NY), 100 IU/ mL penicillin, 100 mg/mL streptomycin (Sigma-Aldrich, St. Louis, MO), and 2 mM L-glutamine (Gibco, Grand Island, NY) in 75 cm2 culture flasks at 37 °C with 5% CO2. HGF cells at 5th passage and grown until reaching 90% confluence were used for further analyzes. Briefly, the cells were washed with phosphate-buffered saline (PBS), recovered using accutase solution (Sigma-Aldrich, St. Louis, MO), and resuspended in PBS. HGF cells were adjusted at a concentration of 1 × 105 cells/well. Then, 5 μM of carboxyfluorescein diacetate succinimidyl ester (CFSE-Cell Proliferation Kit, Life Technology, CA) CFSE solution was added and the cells were incubated for 20 min at 37 °C to reduce the autofluorescence. Stained cells were washed twice with PBS buffer containing 2% FBS to suppress any dye remaining in the solution and resuspended in fresh DMEM at the same concentration. Samples were placed into a 24-well polystyrene plate, and 700 μL of HGF stained cells were seeded directly onto each sample surface. The cells incubated with 9% triton served as negative controls, signaling cell death. Concomitantly, cells cultured onto polystyrene well surfaces with DMEM culture medium represented the growth control (positive control). The plates were incubated for 24 h at 37 °C under 5% CO2 conditions. Fifteen minutes prior to fluorescence analyses, the DMEM supplemented medium was removed and samples were maintained with 100 μg/mL of propidium iodide (PI). The effect of the UV treatment on the viability of HGF cells were evaluated by the green (CFSE) and red (PI) fluorescence signals at 488 and 561 nm, respectively, using a confocal microscopy (Zeiss LSM 800, Jena, Germany). Samples were acquired through 10× dry (Plan NeoFluar NA 0.3 air) objective lens. Images were also taken through 5× as a stack for three-dimensional (3D) reconstruction. The experiment was performed in duplicate for each experimental and control group.

2.6. Biological assay 2.6.1. Bacterial culture conditions and microbiology assay Streptococcus sanguinis strains (IAL 1832) were grown in brain heart infusion (BHI) agar plates (Difco Laboratories, Becton, Dickinson and Company, Le Pont-de-Claix, Auvérnia-Ródano-Alpes, France) under 10% CO2 at 37 °C. Afterwards, the bacterial suspensions were cultured in sterile tubes containing 9 mL of fresh BHI broth overnight under 10% CO2 at 37 °C for the exponential growth phase. Prior to the optical density measurements, the tubes were centrifuged (6000g for 5 min at 4 °C), and the precipitate was resuspended in 0.9% NaCl. The optical density was adjusted to OD = 1.00 ± 0.02 at 600 nm using a spectrophotometer (DU800 Beckman Coulter, Brea, CA, USA), and a final suspension of 107 cells/mL was obtained [14]. This study was approved by the Local Research and Ethics Committee (80684517.1.0000.5418/2017). For the acquired pellicle formation, whole human saliva stimulated by chewing a flexible film (Parafilm M, American Can Co., Neenah, WI, USA) was collected from two volunteers, at least 1.5 h after eating or drinking, after providing their informed consent [43]. The collected saliva was centrifuged (10,000g for 10 min at 4 °C), and the supernatant was filtered through a 0.22-μm membrane filter (K15–1500, Kasvi, São José dos Pinhais, Paraná, Brazil) and used immediately [14]. Before the acquired pellicle formation, discs were sterilized by gamma irradiation (14.5 ± 0.05 kGy; Cobalt-60, Gammacell 220, Atomic Energy of Canada Ltd., Otawa, ON, Canada) [44]. For the UV+ experimental groups, after gamma irradiation, the discs were photofunctionalized for 96 h prior to the beginning of the microbiological assay. Immediately after photofunctionalization, the discs were transferred individually to a well of a sterile 24-well polystyrene cell culture plate, covered with 1 mL of saliva and incubated under agitation for 30 min at 37 °C [45]. After pellicle formation, the discs were transferred to new wells and covered with 200 μL of S. sanguinis cell suspension and 1800 μL of BHI broth. Then, the discs were incubated under 10% CO2 at 37 °C for 1 h and 6 h. For removal of non-adherent cells, discs were washed twice in 0.9% NaCl and then transferred to cryogenic tubes containing 3 mL of 0.9% NaCl. The tube was then sonicated (7 W for 30 s) to disaggregate the bacterial cells, and an aliquot of 0.1 mL was 5-fold serially diluted in 0.9% NaCl and plated in BHI agar. Then, the plates were incubated (10% CO2, at 37 °C for 48 h). After obtaining the counts of colony forming units (CFU), data were expressed as the log of the colony

2.7. Statistical analyses Statistical analyses were performed with statistical software (IBM SPSS Statistics for Windows, v. 21.0, IBM Corp., Armonk, NY, USA). The Shapiro-Wilk method was applied to test the normality of all response variables. An independent t-test was used to verify the influence of the surface treatment on the surface roughness. Two-way ANOVA was used to test the influence of the surface treatment and photofunctionalization on the wettability, proteins adsorption, photocatalytic activity and colony forming unit counts. To verify the influence of time on the wettability of samples, one-way repeated measures ANOVA was performed. The Tukey HSD test was used as a post hoc technique for multiple comparisons when necessary. A mean difference significant at the 0.05 level was used for all tests. The statistical power was calculated using the software G*Power version 3.1.9.2 (Program written, concept and design by Franz, Universitat Kiel, Germany. Freely available windows application software) (β > 0.9, α = 0.05) considering the main variables of the study (contact angle, protein adsorption and colony forming units). 4

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

Fig. 2. SEM micrographs (2000× magnification, 4 kV, WD = 10 mm, scale bar = 10 μm) and two- and three-dimensional laser scanning confocal microscopy images (150× magnification, scale bar = 10 μm) of the cpTi and PEO-treated surfaces.

3. Results 3.1. Surface characteristics Fig. 2 shows the SEM micrographs of the cpTi and PEO-treated surfaces. The machined surfaces presented longitudinal grooves and a more homogeneous surface due to the polishing process. In contrast, the surfaces treated with PEO were patently modified and exhibited a volcanic appearance and pores. These characteristics can also be observed in the 2D and 3D images obtained by LSCM (Fig. 2). Additionally, the average area (Fig. 2) and surface roughness (Fig. 3) of the PEO-treated surfaces were higher than those observed for the machined surface (p < 0.001) as a result of the microdischarges that occurred in the PEO treatment and the modification of the Ti surface. The X-ray diffraction patterns showed peaks of amorphous Ti for the cpTi group. In contrast, crystalline phases of the anatase (101) peak at 25° and rutile (110) peak at 27° were observed for the PEO-treated group (Fig. 4). The machined and PEO surfaces presented a chemical composition consisting of Ti, O and C. Additionally, the PEO surfaces presented Ca and P incorporation by the surface treatment (Fig. 5).

Fig. 4. X-ray diffraction patterns of the cpTi and PEO-treated surfaces.

Notably, the UV-mediated photofunctionalized groups (cpTi UV+ and PEO UV+) showed a reduction of the C atomic concentration (Table 1). These results observed in the EDS were confirmed by XPS analysis (Fig. 6). The XPS spectrum showed the expected compounds, including the presence of Ti2p, O1s, C1s bands for all groups and Ca2p and P2p for the PEO-treated groups. A decrease in the metallic peaks of Ti2p was observed for the PEO-treated groups due to the deposition of the coating layer and incorporation of Ca and P. The presence of Ti suboxides was identified in the cpTi groups, but it was not observed in PEO-treated surfaces, indicating complete element oxidation by the surface treatment. Despite the reduction of the C concentration, an increase in the peak of the covalent bond between C and O (O-C=O) was observed for the photofunctionalized groups. The detailed relative atomic concentration of each element obtained by XPS analysis is shown in Table 1. The contact angle of the cpTi UV− was higher than the angle observed in the PEO UV− (p < 0.0001), indicating a lower wettability of the nontreated surface. After photofunctionalization (cpTi UV+ and PEO UV+), a decrease in the contact angle and an increase in the

Fig. 3. Roughness data (n = 5) for the cpTi and PEO-treated surfaces. Ra (arithmetic roughness), Rq (root mean square average), Rz (average peak-tovalley height), Rt (maximum height). Bars indicate significant differences between the cpTi and PEO groups for each roughness parameter (p < 0.05; Tukey HSD test). 5

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

Fig. 5. Element peaks obtained by energy dispersive spectroscopy analysis for the cpTi and PEO-treated surfaces. Table 1 Mean and (standard deviation) values of the atomic concentration (%) average values from 3 different areas obtained by energy dispersive spectroscopy (EDS) (n = 1) and X-ray photoelectron spectroscopy (XPS) (n = 3). EDS atomic concentration (%)

XPS atomic concentration (%)

Element

cpTi UV−

cpTi UV+

PEO UV−

PEO UV+

Peak

cpTi UV−

cpTi UV+

PEO UV−

PEO UV+

Ti O C Ca P

90.57(1.60) 4.41(1.48) 5.02(2.29) – –

92.66(3.15) 4.03(0.24) 3.31(1.08) – –

4.75(1.16) 68.94(2.90) 8.62(2.35) 10.45(1.16) 12.00(4.91)

8.64(1.45) 70.91(1.71) 4.91(1.53) 9.00(1.46) 6.55(1.39)

Ti2p O1s C1s Ca2p P2p

19.25(1.28) 48.03(0.44) 30.63(1.75) – –

23.54(1.73) 60.20(2.52) 15.71(2.42) – –

7.66(1.24) 51.33(1.54) 20.02(0.30) 13.40(0.93) 9.46(0.48)

7.47(0.76) 53.93(1.17) 14.67(1.28) 12.95(2.08) 10.15(1.00)

proteins adsorption than the cpTi UV− group (p = 0.006) as well as higher albumin adsorption (p = 0.004) (Fig. 10). In addition, when photofunctionalized, the PEO UV+ group presented a considerable rise in human blood plasma proteins and albumin adsorption with a statistically significant difference when compared to PEO UV− (p = 0.008 and p = 0.001, respectively). Although cpTi UV+ showed a slight increase in proteins adsorption when compared to cpTi UV−, no statistically significant difference was noted between them.

surface hydrophilicity (p < 0.0001) were noted, mainly for cpTi UV+ where contact angle reduction was expressive. No significant difference was observed between cpTi UV+ and PEO UV+ after photofunctionalization (p = 0.230) (Fig. 7). After photofunctionalization, the hydrophilic status of the surfaces was evaluated over time to analyze the wettability degradation of the UV light effect. A control experiment was performed with the nonphotofunctionalized (UV−) surfaces and they tended to maintain the wettability status overtime (up to 4 h) (Fig. 8a). It is interesting to note that the contact angle in the UV+ group was lower than the one observed in the UV− group throughout the 6 h for the cpTi surface. However, for the PEO surface such reduced contact angle in the UV+ group lasted long lower than 3 h. A significant increase in the contact angle for cpTi UV+ was observed 2 h after photofunctionalization. However, up to 6 h of aging, the contact angle of the cpTi UV+ surfaces remained lower than the initial values before photofunctionalization. Conversely, the PEO UV+ surface maintained a lower contact angle, but at 4 h after photofunctionalization, the contact angle was the same as before UV light application (Fig. 8b). In addition, at 6 h after photofunctionalization, the contact angle was similar for both cpTi UV+ and PEO UV+ surfaces (p = 0.817).

3.4. Microbiological assay Fig. 11 shows a significant decrease in the CFU counts for PEO UV+ surfaces when compared to PEO UV− (p = 0.012) after 1 h of bacterial adhesion. Although the photofunctionalization reduced the CFU counts for the cpTi UV+ surface, there was no statistically significant difference compared with cpTi UV− (p = 0.269). The PEO-treated surface exhibited greater bacterial adhesion at 1 h than cpTi under both nonphotofunctionalized (p < 0.001) and photofunctionalized (p = 0.02) conditions. After 6 h of adhesion, no difference was observed between the cpTi and PEO surfaces (p = 0.493), nor between nonphotofunctionalized and photofunctionalized surfaces (p = 0.538). The SEM micrographs (Fig. 12) were made to confirm that there was bacterial colonization on the surfaces at 1 and 6 h of adhesion for all surfaces. In addition, it is possible to observe the arrangement of S. sanguinis as streptococcus chains with a similar distribution for the surfaces. Representative images of S. sanguinis shows short streptococcal chains without multicellular organized structures for the surfaces in the initial bacterial adhesion.

3.2. Photocatalytic activity Fig. 9 shows the photocatalytic activity for all groups. The cpTi UV− and cpTi UV+ groups presented no photocatalytic activity. Moreover, although the PEO groups presented the crystalline phases anatase and rutile, no photocatalytic activity was observed even for the PEO UV+ group. Although the PEO UV+ treated groups showed a slight increase in photocatalytic activity, no statistically significant difference was observed compared with cpTi even at the maximum values of 2 h (p = 0.198).

3.5. Cell culture assay Since PEO-treated surface increased the blood plasma proteins adsorption and reduced the initial bacterial adhesion at the early time point, we also investigated, for the first time, the biological safety aspects of the experimental surfaces on mammalian cells. Qualitative assessment by fluorescence microscopy confirmed normal

3.3. Protein adsorption The PEO UV− treated group showed greater human blood plasma 6

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

Fig. 6. XPS spectra of cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+) (n = 1).

characteristics and biological properties of Ti material when PEO treatment is associated with the UV photofunctionalization; these results support the study hypothesis. PEO treatment provides a rougher surface and higher wettability than the machined condition. Furthermore, the UV photofunctionalization enhances the proteins adsorption and hydrophilicity on PEO-treated surfaces, in addition to inducing a reduction of bacterial adhesion at the early stage of biofilm formation with noncytotoxic effect on HGF cells. The surface modifications obtained by PEO and UV-mediated

morphological features and non-effect on cell proliferation at PEOtreated surfaces in comparison to their counterparts. After 24 h of incubation, dead cells were only identified at the negative control, when HGF were exposed to a detergent used to lyse de cells causing complete membrane disruption in culture medium (Fig. 13).

4. Discussion In this study, we have shown an enhancement of the surface 7

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

Fig. 7. a) Contact angle (n = 5) of cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+); b) representative images of contact angle. Bars indicate significant differences between groups (p < 0.05; Tukey HSD test).

inhibition of bacterial adhesion are important strategies to be acquired in the biomedical materials field [47]. Despite the higher wettability of the PEO-treated surfaces, the higher CFU counts of PEO-treated surfaces when compared to the machined condition after 1 h of adhesion can be explained by the increased surface roughness obtained by the surface treatment, which provides a larger surface area for bacterial adhesion [7]. Nevertheless, the surface roughness of dental implant surfaces is believed to be a strong factor influencing the bacterial adhesion [48]. The surface roughness average of PEO-treated group was about 0.48 μm in contrast to 0.18 μm for cpTi which can be a factor influencing mainly the adhesion of bacterial cells. Previous study has suggested an increase in early bacterial adhesion when differences in the surface roughness are above the threshold of 0.2 μm [49]. In addition, rougher surfaces have demonstrated no effect on biofilm amount as the biofilm become

photofunctionalization have an important role for improvement of the surface characteristics of dental implants. Regarding wettability, PEOtreated surfaces present a more hydrophilic surface due to the presence of calcium acetate [46]. In addition, photofunctionalization enhances the wettability by hydrocarbon removal and favorable water adsorption through the excitement of electrons from the valence band to the conduction band and the creation of oxygen vacancies as a result of the reduction of Ti4+ to Ti3+ [24]. Our findings also demonstrate a maintenance of the hydrophilic status for PEO-treated surfaces up to 4 h of aging, but it was lost rapidly for machined surfaces when compared to the biofunctional PEO coating. However, at 6 h after photofunctionalization, the contact angle was the same for both machined and PEO-treated surfaces. It is interesting to note that the acceleration of bone formation and

Fig. 8. a) Hydrophilic status of cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+). The timeline “immediately” means the contact angle measured immediately after the 96 h of UV light application in the UV+ groups, and after samples preparation in the UV− groups. b) Comparison of hydrophilic status degradation of cpTi UV+ and PEO UV+ treated surfaces in dark and dry conditions after 96 h of UV light exposure. Different capital letters indicate statistically significant differences between cpTi and PEO within each period of evaluation (p < 0.05; Tukey HSD test). Different lowercase letters indicate statistically significant differences among the evaluation periods within each surface type (p < 0.05; Tukey HSD test). 8

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

found a link between reducing bacterial adhesion and reducing the ionic strength [59,60]. S. sanguinis carries a net negative surface charge, consequently the reduction in surface charge after PEO photofunctionalization can lead to a lower bacterial attraction to the surface, justifying the lower CFU counts at 1 h of adhesion for PEO UV+ [61]. Despite the absence of photocatalytic activity due to the short lifetime of the reactive oxygen species, which longest lifetime is 2.7 μs [62], the photocatalytic activity and hydroxyl groups generation as well as the surface charge modifications are not neglected for PEO UV+ surface. The rougher surface may also be a factor in the improvement of protein adsorption and the osseointegration process [14]. Protein adsorption is influenced by surface properties including roughness, chemical composition, wettability, van der Waals interactions and hydrogen bonding. Importantly, the photofunctionalization was able to increase albumin adsorption for PEO-treated surfaces. The presence of pores and the rougher surface obtained by the PEO treatment promotes a large number of anchorage sites [14]. In addition, the Ca incorporated by the PEO treatment is known to attract negatively charged proteins, encouraging adsorption [16]. The photofunctionalization also has a positive role in protein adsorption, increasing adsorption on PEOtreated surfaces. The photofunctionalization of TiO2 containing crystalline phases anatase and rutile is known to present photocatalytic activity [29]. Such mechanism includes the generation of reactive oxygen species, oxygen vacancies as well as hydroxyl groups formation [24]. The photocatalytic mechanism and hydroxyl groups generation [53] can be a factor influencing the protein adsorption. Furthermore, the hydroxyl group (˗OH) is an intrinsic factor that has been reported to provide strong interaction with the amino group (˗NH3+) in proteins, regulating the initial protein adsorption and subsequent cell behaviors [63]. Previous study has shown higher amount of bovine serum albumin adsorption on anatase compared to amorphous TiO2 [64]. Thus, the higher proteins adsorption on PEO UV+ can be related to the hydroxyl groups generation. Overall, our findings suggest the presence of important surface modifications of Ti material, which directly influence the biological properties of the surfaces in terms of proteins adsorption and bacterial colonization. Concerning the protein adsorption to biomaterials, it is the first step in the attachment of functional cells [12]. Among blood plasma proteins, albumin is an important blood serum protein that regulates cytoplasmatic Ca and the cellular proliferation of osteoblasts [13]. Consequently, the greater protein adsorption acts to accelerate the osseointegration process. Indeed, the lower counts of bacteria when PEO-treated surfaces are photofunctionalized represent a valuable finding. For instance, bacterial colonization may occur at any time during implant rehabilitation, but it is crucial to avoid microorganism proliferation in the early healing stage [3,10], which can diminish early

Fig. 9. Photocatalytic activity (n = 3) of cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+).

mature [50,51]. Similarly, our results showed higher CFU counts at 1 h of adhesion for the rougher surface represented by PEO, conversely at a later stage of biofilm development (6 h) the PEO-treated groups showed similar bacterial counts compared to cpTi. On the other hand, the differences in the CFU counts among PEO UV− and PEO UV+ were not determined by the surface properties since they present the same morphology, roughness and chemical composition. Considering the results, it is suggested an additional effect of UV photofunctionalization on PEO-treated surfaces that was not likely to occur on the cpTi surface. Previous studies have shown lower surface charges as increases the time of UV light application for submicron and nanoparticles containing crystalline phases anatase and rutile [52,53]. When photofunctionalized, surfaces with photocatalytic potential present Ti4+ ions that combine with electrons and reduce to Ti3+, the photoproduced electron positive holes will oxidize and produce oxygen vacancies in the lattices [54]. Such defect sites and oxygen vacancies enhance the dissociation of water molecules [55]. This process of UV-mediated photofunctionalization on TiO2 surfaces provides the formation of reactive oxygen species %OH, O2%ˉ and H2O2 [30,31]. The hydroxyl groups (%OH) formed on TiO2 surface is the result of water molecules dissociation and the generation of bridging hydroxyls would contribute more negative charges [53]. Once the bridging hydroxyl is acidic (pKa 2.9), it could reduce the overall electropositive charge of PEO-treated surface [53,56]. The electrostatic interaction is based on electric forces between charged molecules and indicates the affinity between a molecule and its adjacent ones [57]. Oppositely charged molecules are attractive and similar charged molecules present a repulsive electrostatic interaction [58]. Thus, the surface charge reduction for PEO-treated surfaces can influence the electrostatic interactions between the surface and bacterial cells. Previous studies have

Fig. 10. a) Human blood plasma proteins adsorption and b) Albumin adsorption (n = 5) of cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+). Bars indicate significant differences between groups (p < 0.05; Tukey HSD test). 9

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

Fig. 11. Colony forming units (log10 CFU/mL) of S. sanguinis that developed on surfaces after 1 h and 6 h of adhesion. The bar indicates a significant difference between groups (p < 0.05; Tukey HSD test).

characteristics of titanium material, and when associated with biofunctional titanium-treated surfaces, it induces greater proteins adsorption and lower initial bacterial colonization with noncytotoxic effects on HGF cells. From a clinical perspective, UV-mediated photofunctionalization of the titanium biofunctional coating is a promising strategy to improve the implant-host interaction and to reduce oral biofilm-associated disease.

implant failures due to peri-implant infections. Mucositis represents a risk factor for the osseointegration process in early healing stage, and its progression can contribute to the development of peri-implantitis [5,3]. Based on these findings, the physical-chemical surface modifications and enhancement of the biological properties obtained by the association of PEO treatment and photofunctionalization are important findings of our study. Additionally, the greater protein adsorption and lower CFU counts could ensure the prevention of early implant loss, as a result of the increased potential for implant bone contact. Moreover, the initial establishment of bone contact has an imperative role for osseointegration and increased success rates of dental implants. However, further studies are needed to better understand and to improve the applicability of photofunctionalized surfaces with respect to antimicrobial activity as well as to assess the remaining effects of surface charge modifications after UV photofunctionalization to prove this effect on a crystalline PEO surface and its influence on early bacterial adhesion. The clinical behavior of photofunctionalized implants regarding the rates of infection and diseases associated with the biofilm presence must also be assessed, as well as further investigations regarding the influence of UV light intensity, wavelength and exposure time on the antimicrobial effect.

Declaration of competing interest The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper. Acknowledgments This study was financed in part by the Coordenação de Aperfeiçoamento de Pessoal de Nível Superior - Brasil (CAPES) Finance Code 001 and by the São Paulo Research Foundation (FAPESP), Brazil (grant numbers 2016/11470-6 and 2017/01320-0). The authors also thank the Oral Biochemistry Laboratory at Piracicaba Dental School at the University of Campinas (UNICAMP) for the microbiology facility, the Laboratory of Technological Plasmas at the São Paulo State University (UNESP) for the PEO facility and the Brazilian Nanotechnology National Laboratory (LNNano) at the Brazilian Center of Research in Energy and Materials (CNPEM) for the LSCM, XPS and

5. Conclusion Photofunctionalization

modifies

the

physico-chemical

Fig. 12. SEM micrographs of bacterial adhesion formed on cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+) at 5000× magnification (scale bar = 5 μm, 15 kV). The arrows indicate S. sanguinis cells arranged as chains. 10

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

Fig. 13. Fluorescence staining of HGF cells with CFSE (green color) and PI (red color) at 24 h of incubation on cpTi and PEO-treated surfaces without UV treatment (UV−) and photofunctionalized (UV+) at 5× and 10× of magnification (scale bar = 100 μm and 50 μm). Live cells are indicated by the green stained HGF that revealed no changes in elongated cell morphology on each surface. The rounded morphology and red signal of the HGF cells were only observed in the negative control (HGF cells exposed to 9% triton). (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

XRD facilities. [16]

References [1] S.M. Beschnidt, C. Cacaci, K. Dedeoglu, D. Hildebrand, H. Hulla, G. Iglhaut, et al., Implant success and survival rates in daily dental practice: 5-year results of a noninterventional study using CAMLOG SCREW-LINE implants with or without platform-switching abutments, Int J Implant Dent 4 (2018) 33, https://doi.org/10. 1186/s40729-018-0145-3. [2] X. Liu, S. Chen, J.K.H. Tsoi, J.P. Matinlinna, Binary titanium alloys as dental implant materials-a review, Regen Biomater 4 (2017) 315–323, https://doi.org/10. 1093/rb/rbx027. [3] H. Mohajerani, R. Roozbayani, S. Taherian, R. Tabrizi, The risk factors in early failure of dental implants: a retrospective study, J Dent (Shiraz, Iran) 18 (2017) 298–303. [4] C.T. Lee, Y.W. Huang, L. Zhu, R. Weltman, Prevalences of peri-implantitis and periimplant mucositis: systematic review and meta-analysis, J. Dent. 62 (2017) 1–12, https://doi.org/10.1016/j.jdent.2017.04.011. [5] F.O. Costa, S. Takenaka-Martinez, L.O.M. Cota, S.D. Ferreira, G.L.M. Silva, J.E. Costa, Peri-implant disease in subjects with and without preventive maintenance: a 5-year follow-up, J. Clin. Periodontol. 39 (2012) 173–181, https://doi. org/10.1111/j.1600-051X.2011.01819.x. [6] R. Figueiredo, O. Camps-Font, E. Valmaseda-Castellon, C. Gay-Escoda, Risk factors for postoperative infections after dental implant placement: a case-control study, J. Oral Maxillofac. Surg. 73 (2015) 2312–2318, https://doi.org/10.1016/j.joms.2015. 07.025. [7] C. Hannig, M. Hannig, The oral cavity a key system to understand substratum-dependent bioadhesion on solid surfaces in man, Clin. Oral Investig. 13 (2009) 123–139, https://doi.org/10.1007/s00784-008-0243-3. [8] M.M. Fürst, G.E. Salvi, N.P. Lang, G.R. Persson, Bacterial colonization immediately after installation on oral titanium implants, Clin. Oral Implants Res. 18 (2007) 501–508, https://doi.org/10.1111/j.1600-0501.2007.01381.x. [9] I.M.G. Cavalcanti, A.P.R. Filho, S.C. Lucena-ferreira, W.J. Silva, A.F.P. Leme, P.M. Senna, et al., Salivary pellicle composition and multispecies biofilm developed on titanium nitrided by cold plasma, Arch. Oral Biol. 59 (2014) 695–703, https:// doi.org/10.1016/j.archoralbio.2014.04.001. [10] E.D. de Avila, B.P. Lima, T. Sekiya, Y. Torii, T. Ogawa, et al., Effect of UV-photofunctionalization on oral bacterial attachment and biofilm formation to titanium implant material, Biomaterials 67 (2015) 84–92, https://doi.org/10.1016/j. biomaterials.2015.07.030. [11] P.E. Kolenbrander, R.N. Andersen, D.S. Blehert, P.G. Egland, J.S. Foster, R.J. Palmer, Communication among oral bacteria, Microbiol. Mol. Biol. Rev. 66 (2002) 486–505 table of contents https://doi.org/10.1128/mmbr.66.3.486-505. 2002. [12] N. Hori, T. Ueno, H. Minamikawa, F. Iwasa, F. Yoshino, K. Kimoto, et al., Electrostatic control of protein adsorption on UV-photofunctionalized titanium, Acta Biomater. 6 (2010) 4175–4180, https://doi.org/10.1016/j.actbio.2010.05. 006. [13] S.R. Sousa, M. Lamghari, P. Sampaio, P. Moradas-Ferreira, M.A. Barbosa, Osteoblast adhesion and morphology on TiO2 depends on the competitive preadsorption of albumin and fibronectin, J. Biomed. Mater. Res. A 84 (2008) 281–290, https://doi. org/10.1002/jbm.a.31201. [14] J.M. Cordeiro, H.N. Pantaroto, E.M. Paschoaleto, E.C. Rangel, N.C. Cruz, C. Sukotjo, et al., Synthesis of biofunctional coating for a TiZr alloy: surface, electrochemical, and biological characterizations, Appl. Surf. Sci. 452 (2018) 268–278, https://doi. org/10.1016/j.apsusc.2018.05.044. [15] F. Iwasa, N. Hori, T. Ueno, H. Minamikawa, M. Yamada, T. Ogawa, Enhancement of osteoblast adhesion to UV-photofunctionalized titanium via an electrostatic

[17]

[18]

[19]

[20]

[21]

[22] [23]

[24]

[25]

[26]

[27]

[28]

[29]

[30]

[31]

11

mechanism, Biomaterials 31 (2010) 2717–2727, https://doi.org/10.1016/j. biomaterials.2009.12.024. Y. Mao, J. Yan, L. Wang, W. Dong, Y. Jia, X. Hu, et al., Formation and properties of bioactive barium titanate coatings produced by plasma electrolytic oxidation, Ceram. Int. 44 (2018) 12978–12986, https://doi.org/10.1016/j.ceramint.2018.04. 115. I. da S.V. Marques, V.A.R. Barão, N.C. da Cruz, J.C. Yuan, M.F. Mesquita, A.P. Ricomini-Filho, et al., Electrochemical behavior of bioactive coatings on cp-Ti surface for dental application, Corros. Sci. 100 (2015) 133–146, https://doi.org/10. 1016/j.corsci.2015.07.019. B.E. Nagay, C. Dini, J.M. Cordeiro, A.P. Ricomini-Filho, E.D. de Avila, E.C. Rangel, et al., Visible-light-induced photocatalytic and antibacterial activity of TiO(2) codoped with nitrogen and bismuth: new perspectives to control implant-biofilm-related diseases, ACS Appl. Mater. Interfaces 22 (20) (2019) 18186–18202, https:// doi.org/10.1021/acsami.9b03311 11. F.C. Walsh, R. Wood, F.C. Walsh, C.T.J. Low, R.J.K. Wood, K.T. Stevens, et al., Plasma electrolytic oxidation (PEO) for production of anodised coatings on lightweight metal (Al, Mg, Ti) alloys, Trans. Inst. Met. Finish. 87 (2009) 122–135, https://doi.org/10.1179/174591908X372482. D. Krupa, J. Baszkiewicz, J. Zdunek, J.W. Sobczak, W. Lisowski, J. Smolik, et al., Effect of plasma electrolytic oxidation in the solutions containing Ca, P, Si, Na on the properties of titanium, J Biomed Mater Res B Appl Biomater 100 (2012) 2156–2166, https://doi.org/10.1002/jbm.b.32781. T. Beline, I. da S.V. Marques, A.O. Matos, E.S. Ogawa, A.P. Ricomini-Filho, E.C. Rangel, et al., Production of a biofunctional titanium surface using plasma electrolytic oxidation and glow-discharge plasma for biomedical applications, Biointerphases 11 (2016) 011013, , https://doi.org/10.1116/1.4944061. T. Ogawa, Ultraviolet photofunctionalization of titanium implants, Int. J. Oral Maxillofac. Implants 29 (2014) e95–102, https://doi.org/10.11607/jomi.te47. B. Cortat, C.C.M. Garcia, A. Quinet, A.P. Schuch, K.M. de Lima-Bessa, C.F.M. Menck, The relative roles of DNA damage induced by UVA irradiation in human cells, Photochem Photobiol Sci 12 (2013) 1483–1495, https://doi.org/10.1039/ c3pp50023c. H. Aita, N. Hori, M. Takeuchi, T. Suzuki, M. Yamada, M. Anpo, et al., The effect of ultraviolet functionalization of titanium on integration with bone, Biomaterials 30 (2009) 1015–1025, https://doi.org/10.1016/j.biomaterials.2008.11.004. H. Minamikawa, T. Ikeda, W. Att, Y. Hagiwara, M. Hirota, M. Tabuchi, et al., Photofunctionalization increases the bioactivity and osteoconductivity of the titanium alloy Ti6Al4V, J. Biomed. Mater. Res. A 102 (2014) 3618–3630, https://doi. org/10.1002/jbm.a.35030. M. Takeuchi, K. Sakamoto, G. Martra, S. Coluccia, M. Anpo, Mechanism of photoinduced superhydrophilicity on the TiO2 photocatalyst surface, J. Phys. Chem. B 109 (2005) 15422–15428, https://doi.org/10.1021/jp058075i. W. Att, N. Hori, F. Iwasa, M. Yamada, T. Ueno, T. Ogawa, The effect of UV-photofunctionalization on the time-related bioactivity of titanium and chromium – cobalt alloys, Biomaterials 30 (2009) 4268–4276, https://doi.org/10.1016/j. biomaterials.2009.04.048. A.M. Gallardo-Moreno, M.A. Pacha-Olivenza, M.-C. Fernández-Calderón, C. PérezGiraldo, J.M. Bruque, M.-L. González-Martín, Bactericidal behaviour of Ti6Al4V surfaces after exposure to UV-C light, Biomaterials 31 (2010) 5159–5168, https:// doi.org/10.1016/j.biomaterials.2010.03.005. H.N. Pantaroto, A.P. Ricomini-Filho, M.M. Bertolini, J.H. Dias da Silva, N.F. Azevedo Neto, C. Sukotjo, et al., Antibacterial photocatalytic activity of different crystalline TiO2 phases in oral multispecies biofilm, Dent. Mater. 34 (2018) e182–e195, https://doi.org/10.1016/j.dental.2018.03.011. J.-W. Liou, H.-H. Chang, Bactericidal effects and mechanisms of visible light-responsive titanium dioxide photocatalysts on pathogenic bacteria, Arch. Immunol. Ther. Exp. 60 (2012) 267–275, https://doi.org/10.1007/s00005-012-0178-x. U. Joost, K. Juganson, M. Visnapuu, M. Mortimer, A. Kahru, E. Nõmmiste, et al., Photocatalytic antibacterial activity of nano-TiO2 (anatase)-based thin films: effects

Materials Science & Engineering C 110 (2020) 110657

C. Dini, et al.

[32]

[33]

[34]

[35]

[36]

[37]

[38]

[39]

[40]

[41]

[42]

[43]

[44]

[45]

[46]

[47]

[48] R. Bürgers, T. Gerlach, S. Hahnel, F. Schwarz, G. Handel, M. Gosau, In vivo and in vitro biofilm formation on two different titanium implant surfaces, Clin. Oral Implants Res. 21 (2) (2010) 156–164, https://doi.org/10.1111/j.1600-0501.2009. 01815.x. [49] C.M. Bollen, P. Lambrechts, M. Quirynen, Comparison of surface roughness of oral hard materials to the threshold surface roughness for bacterial plaque retention: a review of the literature, Dent. Mater. 13 (4) (1997) 258–269, https://doi.org/10. 1016/S0109-5641(97)80038-3. [50] A. Han, X. Li, B. Huang, J.K.H. Tsoi, J.P. Matinlinna, Z. Chen, et al., The effect of titanium implant surface modification on the dynamic process of initial microbial adhesion and biofilm formation, International Journal of Adhesion & Adhesives 69 (2016) 125–132, https://doi.org/10.1016/j.ijadhadh.2016.03.018. [51] A. Al-Ahmad, M. Wiedmann-Al-Ahmad, A. Fackler, M. Follo, E. Hellwig, M. Bachle, et al., In vivo study of the initial bacterial adhesion on different implant materials, Arch. Oral Biol. 58 (2013) 1139–1147, https://doi.org/10.1016/j.archoralbio. 2013.04.011. [52] C. Zhou, Y. Bashirzadeh, T.A. Bernadowski, X. Zhang, UV light− induced aggregation of titania submicron particles, Micromachines (Basel) 7 (11) (2016), https://doi.org/10.3390/mi7110203 pii: E203. 8. [53] J. Sun, L.H. Guo, H. Zhang, L. Zhao, UV irradiation induced transformation of TiO2 nanoparticles in water: aggregation and photoreactivity, Environ Sci Technol 48 (20) (2014) 11962–11968, https://doi.org/10.1021/es502360c 21. [54] N. Sakai, A. Fujishima, T. Watanabe, K. Hashimoto, Enhancement of the photoinduced hydrophilic conversion rate of TiO2 film electrode surfaces by anodic polarization, J. Phys. Chem. B 105 (15) (2001) 3023–3026, https://doi.org/10.1021/ jp003212r. [55] M. Miyauchi, N. Kieda, S. Hishita, T. Mitsuhashi, A. Nakajima, T. Watanabe, et al., Reversible wettability control of TiO2 surface by light irradiation, Surf. Sci. 511 (2002) 401–407, https://doi.org/10.1016/S0039-6028(02)01551-0. [56] H.P. Boehm, Acidic and basic properties of hydroxylated metal oxide surfaces, Discussions of the Faraday Society 52 (1971) 264–275, https://doi.org/10.1039/ DF9715200264. [57] K. Niwa, R. Kuramoto, S. Anandan, Y. Ikuma, Zeta potential and hydrogen production of mesoporous titanium oxide, Procedia Engineering 36 (2012) 62–67, https://doi.org/10.1016/j.proeng.2012.03.011. [58] W. Teughels, N. Van Assche, I. Sliepen, M. Quirynen, Effect of material characteristics and/or surface topography on biofilm development, Clin. Oral Implants Res. 17 (Suppl. 2) (2006) 68–81, https://doi.org/10.1111/j.1600-0501.2006.01353.x. [59] C.H. Bolster, A.L. Mills, G.M. Hornberger, J.S. Herman, Effect of surface coatings, grain size, and ionic strength on the maximum attainable coverage of bacteria on sand surfaces, J. Contam. Hydrol. 50 (3–4) (2001) 287–305, https://doi.org/10. 1016/S0169-7722(01)00106-1. [60] M.C. van Loosdrecht, J. Lyklema, W. Norde, A.J. Zehnder, Bacterial adhesion: a physicochemical approach, Microb. Ecol. 17 (1) (1989) 1–15, https://doi.org/10. 1007/BF02025589. [61] A.H. Weerkamp, H.M. Uyen, H.J. Busscher, Effect of zeta potential and surface energy on bacterial adhesion to uncoated and saliva-coated human enamel and dentin, J. Dent. Res. 67 (12) (1988) 1483–1487, https://doi.org/10.1177/ 00220345880670120801. [62] P. Attri, Y.H. Kim, D.H. Park, J.H. Park, Y.J. Hong, H.S. Uhm, et al., Generation mechanism of hydroxyl radical species and its lifetime prediction during the plasma-initiated ultraviolet (UV) photolysis, Sci. Rep. 5 (2015) 1–8, https://doi. org/10.1038/srep09332. [63] Y. Kang, X. Li, Y. Tu, Q. Wang, H. Agren, On the mechanism of protein adsorption onto hydroxylated and nonhydroxylated TiO2 surfaces, J. Phys. Chem. C 114 (34) (2010) 14496–14502, https://doi.org/10.1021/jp1037156. [64] Y. Hong, M. Yu, J. Lin, K. Cheng, W. Weng, H. Wang, Surface hydroxyl groups direct cellular response on amorphous and anatase TiO2 nanodots, Colloids Surf B Biointerfaces 1 (123) (2014) 68–74, https://doi.org/10.1016/j.colsurfb.2014.08. 030.

on Escherichia coli cells and fatty acids, J. Photochem. Photobiol. B 142 (2015) 178–185, https://doi.org/10.1016/j.jphotobiol.2014.12.010. V.A. Barão, M.T. Mathew, W.G. Assunção, J.C. Yuan, M.A. Wimmer, C. Sukotjo, The role of lipopolysaccharide on the electrochemical behavior of titanium, J. Dent. Res. 90 (2011) 613–618, https://doi.org/10.1177/0022034510396880. N. Harmankaya, K. Igawa, P. Stenlund, A. Palmquist, P. Tengvall, Healing of complement activating Ti implants compared with non-activating Ti in rat tibia, Acta Biomater. 8 (2012) 3532–3540, https://doi.org/10.1016/j.actbio.2012.05. 017. K.-H. Park, J.-Y. Koak, S.-K. Kim, C.-H. Han, S.-J. Heo, The effect of ultraviolet-C irradiation via a bactericidal ultraviolet sterilizer on an anodized titanium implant: a study in rabbits, Int. J. Oral Maxillofac. Implants 28 (2013) 57–66, https://doi. org/10.11607/jomi.2638. S.N. Pleskova, I.S. Golubeva, Y.K. Verevkin, Bactericidal activity of titanium dioxide ultraviolet-induced fi lms, Mater Sci Eng C 59 (2016) 807–817, https://doi.org/10. 1016/j.msec.2015.10.021. L.P. Faverani, W.G. Assunção, P.S.P. de Carvalho, J.C. Yuan, C. Sukotjo, M.T. Mathew, et al., Effects of dextrose and lipopolysaccharide on the corrosion behavior of a Ti-6Al-4V alloy with a smooth surface or treated with double-acidetching, PLoS One 9 (2014) e93377, , https://doi.org/10.1371/journal.pone. 0093377. A.O. Matos, A.P. Ricomini-filho, T. Beline, E.S. Ogawa, B.E. Costa-oliveira, A.A.B. De, et al., Three-species biofilm model onto plasma-treated titanium implant surface, Colloids Surfaces B Biointerfaces 152 (2017) 354–366, https://doi.org/10. 1016/j.colsurfb.2017.01.035. W.S.M. Werner, W. Smekal, C.J. Powell, NIST Database for the simulation of Electron spectra for surface analysis, version 2.0, Standard Reference Data Program Database, 100 Department of Commerce, National Institute of Standards and Technology, Gaithersburg, Maryland, 2014. A. Mills, Environmental an overview of the methylene blue ISO test for assessing the activities of photocatalytic films, Applied Catal B, Environ 128 (2012) 144–149, https://doi.org/10.1016/j.apcatb.2012.01.019. E.S. Loncar, M.M. Radeka, S.B. Petrovi, A.S. Skapin, Determination of the photocatalytic activity of TiO2 coatings on clay roofing tile substrates - methylene blue as model pollutant, Acta Period Technol 220 (2009) 125–133, https://doi.org/10. 2298/APT0940125L. K. Shiraishi, H. Koseki, T. Tsurumoto, K. Baba, M. Naito, H. Shindo, Antibacterial metal implant with a TiO 2 -conferred photocatalytic bactericidal effect against Staphylococcus aureus, Surf. Interface Anal. 41 (2009) 17–22, https://doi.org/10. 1002/sia.2965. S. Choi, W. Jeong, J. Cha, J. Lee, K.-J. Lee, H.-S. Yu, et al., Effect of the ultraviolet light treatment and storage methods on the biological activity of a titanium implant surface, Dent. Mater. 33 (2017) 1426–1435, https://doi.org/10.1016/j.dental. 2017.09.017. B. Guggenheim, E. Giertsen, P. Schüpbach, S. Shapiro, Validation of an in vitro biofilm model of supragingival plaque, J. Dent. Res. 80 (2001) 363–370, https:// doi.org/10.1177/00220345010800011201. I. Soriano, A.Y. Martín, C. Evora, E. Sánchez, Biodegradable implantable fluconazole delivery rods designed for the treatment of fungal osteomyelitis: influence of gamma sterilization, J. Biomed. Mater. Res. A 77 (2006) 632–638, https://doi.org/ 10.1002/jbm.a.30657. T. Pereira-Cenci, D.M. Deng, E.A. Kraneveld, E.M.M. Manders, A.A. Del Bel Cury, J.M. Ten Cate, et al., The effect of Streptococcus mutans and Candida glabrata on Candida albicans biofilms formed on different surfaces, Arch. Oral Biol. 53 (2008) 755–764, https://doi.org/10.1016/j.archoralbio.2008.02.015. S.X. ling, W.Q. liang, W.F. shun, G.S. rong, Effects of electrolytic concentration on properties of micro-arc film on Ti6Al4V alloy, Min. Sci. Technol. 19 (2009) 220–224, https://doi.org/10.1016/S1674-5264(09)60042-9. T. Hanawa, Surface treatment and modification of metals to add biofunction, Dent. Mater. J. 36 (5) (2017) 533–538, https://doi.org/10.4012/dmj.2017-154.

12